Activation of Mrc1, a mediator of
the replication checkpoint, by
telomere erosion
Nathalie Grandin, Aymeric Bailly and Michel Charbonneau1
IFR128 BioSciences Gerland, UMR CNRS no. 5161, Ecole Normale Sup ´erieure de Lyon, 46 all ´ee d’Italie, 69364 Lyon, France
Background information. In budding yeast, the loss of either telomere sequences (in telomerase-negative cells) or telomere capping (in mutants of two telomere end-protection proteins, Cdc13 and Yku) lead, by distinct pathways, to telomeric senescence. After DNA damage, activation of Rad53, which together with Chk1 represents a protein kinase central to all checkpoint pathways, normally requires Rad9, a checkpoint adaptor.
Results. We report that in telomerase-negative (tlc1) cells, activation of Rad53, although diminished, could still take place in the absence of Rad9. In contrast, Rad9 was essential for Rad53 activation in cells that entered senescence in the presence of functional telomerase, namely in senescent cells bearing mutations in telomere end-protection proteins (cdc13-1 yku70). In telomerase-negative cells deleted for RAD9, Mrc1, another checkpoint adaptor previously implicated in the DNA replication checkpoint, mediated Rad53 activation. Rad9 and Rad53, as well as other DNA damage checkpoint proteins (Mec1, Mec3, Chk1 and Dun1), were required for complete DNA-damage-induced cell-cycle arrest after loss of telomerase function. However, unexpectedly, given the formation of an active Rad53–Mrc1 complex in tlc1 rad9 cells, Mrc1 did not mediate the cell-cycle arrest elicited by telomerase loss. Finally, we report that Rad9, Mrc1, Dun1 and Chk1 are activated by phosphorylation after telomerase inactivation. Conclusions. These results indicate that loss of telomere capping and loss of telomere sequences, both of which provoke telomeric senescence, are perceived as two distinct types of damages. In contrast with the Rad53– Rad9-mediated cell-cycle arrest that functions in a similar way in both types of telomeric senescence, activation of Rad53–Mrc1 might represent a specific response to telomerase inactivation and/or telomere shortening, the functional significance of which has yet to be uncovered.
Introduction
Cells must detect and repair spontaneous and in-duced DNA damage that can potentially compro-mise genome integrity and generate heritable muta-tions (Zhou and Elledge, 2000; Melo and Toczyski, 2002; Rouse and Jackson, 2002). In general, the com-ponents of the checkpoint machinery have been very well conserved during evolution, thus making geneti-cally tractable organisms, such as bacteria and yeast, excellent model systems for studying DNA repair and checkpoint functions. Although the checkpoint pathways are relatively well understood, principally
1To whom correspondence should be addressed (email
Michel.Charbonneau@ens-lyon.fr).
Key words: cell cycle, DNA damage, phosphorylation, Rad53–Rad9, telomeric senescence.
Abbreviations used: CHK, checkpoint kinase; HA, haemagglutinin; HU, hydroxyurea; IP, immunoprecipitation; MMS, methyl methane sulphonate.
in yeast and humans, numerous aspects of them are still unclear due to the complexity of the mechanisms involved. In particular, recent studies in yeast have highlighted the implication of numerous crosstalks between the various components of the checkpoint machinery, which modulate the response depending on the location and intensity of the damage (Melo and Toczyski, 2002; Rouse and Jackson, 2002).
Although the chromosomes are more susceptible to breakage at the replication forks and during chromo-some segregation, damage can also occur at any other time and place within the genome, due to accidental attacks such as by UV light, irradiation or genotoxic agents. Even in the absence of any insult, the ends of linear chromosomes require particular protection, since, would protection be relieved, these ends would be recognized as double-strand breaks by checkpoint proteins and DNA repair proteins (de Lange, 2002;
Chakhparonian and Wellinger, 2003; Ferreira et al., 2004). Some telomeric proteins also protect against inappropriate homologous recombination between telomeric sequences, as telomeres are composed of TG-rich repetitive tracts of DNA sequences, which are prone to homologous recombination (Lundblad, 2002). Due to the necessity to recruit telomerase at their extremities at intervals, telomeres appear to require several levels of protection, since they may remain only partially protected during telomerase re-cruitment (Kelleher et al., 2002; Vega et al., 2003). In the budding yeast Saccharomyces cerevisiae, both telomerase and two complexes of telomeric proteins, namely Cdc13–Stn1–Ten1 and Yku70–Yku80, co-operate in telomere end protection, defining three dis-tinct and complementary pathways (Dubrana et al., 2001; Lustig, 2001). The absence of telomerase (in
Sa. cerevisiae, TLC1, the RNA template, or Est2, the
reverse transcriptase catalytic subunit) inexorably leads to sequence loss as cells divide, followed by telomere shortening and, ultimately, chromosome de-gradation and death by senescence (Lundblad and Szostak, 1989; Lendvay et al., 1996).
The knowledge of how damage is treated at telomere ends by the checkpoint machinery is im-portant for several reasons. First, in humans, telo-mere erosion is a naturally occurring event, due to telomerase inactivation in most tissues, and it is cru-cial to know the status of checkpoint activation in tumour cells that are about to reactivate telomere maintenance functions. In this respect, yeast remark-ably recapitulates both the checkpoint and telomeric functions in humans because they use highly con-served genes and mechanisms (d’Adda di Fagagna et al., 2004). Secondly, in yeast (Grandin et al., 2001; Grandin and Charbonneau, 2003), like in humans (Karlseder et al., 2002), telomeric senescence can either result from telomere erosion (after telomerase inactivation) or from loss of telomere capping (in mu-tants of CDC13 and YKU in yeast and of TRF2 in hu-mans) without telomere erosion, thus representing, from a fundamental point of view, two interesting situations. Finally, recent studies of whole genome expression in yeast cells deleted for telomerase have identified a specific telomerase signature (Nautiyal et al., 2002). Characterizing the checkpoint pathways during telomeric damage, and comparing with those occurring elsewhere in the genome, therefore repres-ents a major issue.
In all eukaryotes, the DNA replication and DNA damage stress responses are initiated by members of the phosphoinositide 3-kinase family, Mec1 [ATR (ataxia telangiectasia-related) in humans] and its as-sociated Ddc2/ATRIP and, in distinct complexes, by RFC (replication factor C)-like proteins, such as Rad24 (human RAD17), and PCNA-like proteins, such as Rad17/RAD1, Mec3/HUS1 and Ddc1/RAD9 (Zhou and Elledge, 2000; Rouse and Jackson, 2002). Two key downstream checkpoint components are re-presented by the Chk1 [human CHK1 (checkpoint kinase 1)] and Rad53 (human CHK2) protein kinases. Rad53 becomes hyperphosphorylated and activated in response to DNA damage or DNA replication stress. Rad53 activation is critical for halting cell-cycle progression as long as damage is present, for activating DNA repair via Dun1 and, after repli-cation stress, for delaying late origin firing and stabil-izing the replication complex (Osborn et al., 2002). In response to DNA damage, activation of Rad53 not only requires Mec1 but also Rad9, a budding yeast checkpoint adaptor whose phosphorylation, by Mec1, triggers formation of an active Rad53–Rad9 complex that stimulates the autoactivation, in trans, of Rad53 (Vialard et al., 1998; Pellicioli et al., 1999; Gilbert et al., 2001). In response to DNA replica-tion stress, a different adaptor, Mrc1, activates Rad53 (Alcasabas et al., 2001). Rad9 and Mrc1 show struc-ture and sequence similarities and may work through a common mechanism to activate Rad53 (Osborn et al., 2002; Zegerman and Diffley, 2004). Rad9 also associates with Chk1, independent of Rad53, in re-sponse to DNA damage (Sanchez et al., 1999). While
Sa. cerevisiae Rad9 homologues in Schizosaccharo-myces pombe and humans are Crb2 and either MDC1
or 53BP1 respectively, Sa. cerevisiae Mrc1 also has functional homologues in both Sc. pombe, also called Mrc1, and humans as well as in various other ver-tebrates, in which it is known as Claspin (d’Adda di Fagagna et al., 2004). Mrc1 and Claspin are com-ponents of the replisome, at least in budding yeast and Xenopus (Katou et al., 2003; Lee et al., 2003; Osborn and Elledge, 2003). In fission yeast and hu-mans, Mrc1/Claspin’s DNA-binding domain has been identified recently and the corresponding se-quences appear to have been conserved among several fungi and metazoans (Sar et al., 2004; Zhao and Russell, 2004). As noted above, in yeast, upon DNA replication stress, Mrc1 mediates signal transduction
from Rad3–Rad26, in fission yeast (Mec1-Ddc2 in budding yeast), to Cds1, in fission yeast (budding yeast Rad53) (Alcasabas et al., 2001; Tanaka and Russell, 2001). Therefore Mrc1 has at least two func-tions, one as a component of the replication fork and the second one as a component of the checkpoint machinery, functions that can be genetically separ-ated (Osborn and Elledge, 2003). Interestingly, in
Xenopus, Claspin also associates physically with Chk1
(Kumagai and Dunphy, 2000), an event that is re-quired for activation of Chk1 by ATR-ATRIP in response to incomplete DNA replication (Kumagai et al., 2004). Finally, in Xenopus, Claspin plays a role in adaptation to incomplete DNA replication, a func-tion that is accomplished through its inactivafunc-tion after phosphorylation by the Polo-like kinase (Yoo et al., 2004).
Although two recent studies have explored the DNA damage checkpoint response to loss of telo-merase function in Sa. cerevisiae (Enomoto et al., 2002; Ijpma and Greider, 2003), we nevertheless decided to reinvestigate the situation. In the present study, we report that, in telomerase-negative cells, Mec1, Mec3, Chk1 and Dun1, as well as Rad53 and Rad9, but not Tel1 and Mrc1, are required for the cell-cycle arrest induced by telomeric damage. Moreover, we show that, in telomerase-negative cells, a fraction of Rad53 can still activate in the absence of Rad9 and that this activation depends on Mrc1. In contrast, in cdc13
yku70 senescing cells, Rad53 totally failed to
activ-ate in the absence of Rad9. Unexpectedly, although it participated in Rad53 activation, Mrc1 did not mediate the cell-cycle arrest in telomerase-negative cells. Therefore two distinct losses of telomeric func-tions, both leading to telomeric senescence, employ two distinct pathways of DNA damage checkpoint activation. These results suggest the possibility that the putative Rad53–Mrc1 complex may have an ad-ditional checkpoint function distinct from that in DNA-damage-induced cell-cycle arrest and that this function may be related to genome maintenance in response to loss of telomerase function.
Results
In telomerase-negative cells, Rad53 activation still occurs in the absence of Rad9
To study the checkpoint response to telomeric dam-age in the budding yeast Sa. cerevisiae, we focused
our attention on the activation of the Rad53 kinase, which, with Chk1, is central to all checkpoint path-ways. Upon DNA damage, several states of Rad53 phosphorylation can be detected on Western blots as multiple forms of electrophoretically shifted bands (Sanchez et al., 1996; Sun et al., 1996). Using this criterion and treatment of wild-type cells with the DNA-alkylating agent MMS (methyl methane sul-phonate) as a control (Figure 1B), Rad53 was readily activated after inactivation of telomerase, in tlc1 cells (Figure 1A). Senescence is a slow-going process that is initiated after genetic inactivation of telo-merase. In senescing cells, DNA damage provoked by progressive telomere erosion accumulates as cells divide, leading to cell death after approx. 4–5 days (Lundblad and Szostak, 1989; Lendvay et al., 1996). It is important to note that, for technical reasons, pro-tein phosphorylation could not be analysed on a large enough number of cells until day 1 after telomerase inactivation (see the Materials and methods section). Therefore, although senescence occurs immediately after telomerase inactivation, the molecular events documented here cannot be measured until day 1 after telomerase inactivation. This explains the pres-ence of a slight electrophoretic shift of Rad53 in samples gathered on day 1, because, indeed, such cells had already initiated the process of senescence (Figure 1A). The increase in the population of Rad53-shifted bands with time in telomerase-negative cells, as well as in MMS-treated cells (Figures 1A and 1B), suggested that DNA damage accumulates in these cells as telomeres shorten or as treatment with the genotoxic agent persists.
Rad9, a conserved adaptor checkpoint protein, is required for activation of Rad53 in most types of DNA damage (Schwartz et al., 2002 and references therein), including in telomerase-negative cells (Ijpma and Greider, 2003). On rapid analysis, the Rad53 signal of activation on Western blots indeed appeared to be absent from tlc1 rad9 cells (Fig-ure 1C), compared with that in tlc1 RAD9+ cells (Figure 1A). However, closer examination of the Western blots, as well as comparison with the Rad53 signal of activation in cdc13-1 rad9 cells, revealed that a fraction of Rad53 remained shifted up in the absence of Rad9 (Figures 1C and 1D).
We next decided to measure Rad53 activation using a different technique, exploiting the fact that the ability of Rad53 to autophosphorylate in vitro
Figure 1 Activation of Rad53 visualized by IP-Western blotting
(A) Rad53 activation in telomerase-negative cells (tlc1, lanes 2–5, on days 1–4 after telomerase inactivation) was as-sessed by visualizing electrophoretic mobility alterations after Western-blot analysis of immunoprecipitated cell extracts using monoclonal anti-HA antibodies (IP-Western). Note that on day 1, cells had already initiated the process of telomeric senescence (see the Materials and methods section), hence the presence of a Rad53-shifted band at that time followed by the appearance of additional higher shifted bands on days 2–4 as the senescence process continued. A wild-type strain (wt, lane 1), in which basically all of Rad53–HA migrates as a single band, is shown for comparison. (B) A wild-type strain was treated for the indicated periods of time with 0.1% MMS and processed for measurement of Rad53 activation, as described above for (A). (C) Rad53 activation (assessed by IP-Western blotting) due to telomerase inactivation still per-sisted in the absence of Rad9 [tlc1 rad9, lanes 1–4; compare with the levels in wild-type cells, in (A, D), lanes 1], but it was weaker than in tlc1 RAD9+cells (above, in A). Activation of Rad53 in cdc13-1 cells at 34◦C is shown for comparison (lane 6). (D) Rad53 activation, after cdc13-1-induced telomeric DNA damage in a RAD9+ (lanes 2 and 3) or rad9 background (lanes 4 and 5). Rad53–HA from undamaged wild-type (wt) cells is shown as a control (lane 1). The cdc13-1-induced Rad53 activation, at 34◦C, required Rad9 (compare lanes 3
and 5) unlike the tlc1-induced Rad53 activation (C, above). In (A–D), all strains carried endogenous RAD53–HA integrated at the RAD53 locus.
correlates with its state of activation in vivo (Pellicioli et al., 1999). IP (immunoprecipitation)-kinase assay (see the Materials and methods section) allowed us to visualize the in vitro autophosphorylation signal of Rad53 in telomerase-negative cells (Figure 2A). Since measuring Rad53 autophosphorylation using such an IP-kinase assay has been less reported in the litera-ture than the in vitro renaturation assay (Pellicioli et al., 1999), we first used MMS as a DNA-damaging agent to validate our assay. The Rad53 autophos-phorylation signal progressively increased with time in MMS-treated cells, as expected (Figure 2A, upper panel). Although in these IP-kinase assays we used strains containing the same integrated RAD53-HA (where HA stands for haemagglutinin) construct as the strains used for the Western blots shown above, we were concerned by the presence of two slightly labelled bands of Rad53 autophosphorylation in undamaged wild-type cells (Figure 2A, upper panel, lane 2). We interpreted this as resulting from the higher sensitivity of the in vitro assay compared with
in vivo activation of Rad53 visualized on Western
blots, but nevertheless performed several controls. First, we determined that strains bearing the inte-grated RAD53–HA construct were not hypersensi-tive to MMS (results not shown). However, to elim-inate the possibility of more subtle defects, we next constructed RAD53/RAD53–HA diploid strains. Although Rad53 function was intact due to the untagged copy of RAD53, the two slightly labelled bands were still present (Figure 2B, upper panel, lane 5) just like in the RAD53–HA strain (lane 4). Next, to determine whether these two bands rep-resented an actual Rad53 signal of phosphorylation or, rather, a background signal, or else a phosphoryl-ation signal conferred by a contaminating kinase, we used a kinase-deficient version of RAD53–HA. The two slightly labelled bands were completely absent from the strains bearing the HA-tagged
rad53 K227A (Lys227→ Ala) allele integrated at the RAD53 chromosomal locus (Figure 2C, upper panel). Therefore we conclude that the RAD53–HA strains used to measure Rad53 activation are fully functional, as established previously (Emili, 1998; Emili et al., 2001), and also that the slightly
Figure 2 Activation of Rad53 visualized by IP-kinase assays
(A) A wild-type strain was treated for the indicated periods of time with 0.1% MMS (lanes 2–6). Lane 1 displays the level of Rad53 activation in telomerase-negative cells (tlc1) on day 3 after inactivation of telomerase for comparison with MMS treatment. In the IP-kinase assay (upper panel), cell extracts were immunoprecipitated with a monoclonal anti-HA antibody and processed for autophosphorylation assays as described in the Materials and methods section. Assessment of Rad53 activation by IP-Western blotting (as described in the legend to Figure 1) was performed in parallel on the same samples (lower panel). (B) Rad53 activation [assessed by IP-kinase assay, upper panel, or by IP-Western blotting (on the same samples), lower panel, as described above in A] in diploid strains bearing one copy of endogenous RAD53–HA and the
wild-type (untagged) copy of RAD53 (lanes 2, 3 and 5) and homozygous for cdc13-1 (lanes 2 and 3) or for CDC13 (lane 5) or in haploid strains bearing integrated RAD53–HA (lane 4) or untagged RAD53 (lane 1). See text for interpretation. Cells cor-responding to lanes 1, 4 and 5 were grown at 30◦C, whereas cells corresponding to lanes 2 and 3, that are temperature-sensitive due to the cdc13-1 mutation, were grown at either 25 or 34◦C. (C) Rad53 activation in telomerase-negative cells (tlc1, lane 1 in lower panel) and, in both upper (IP-kinase as-say) and lower (IP-Western blotting, on the same samples) panels, in wild-type cells (wt) bearing an integrated copy of
RAD53–HA, in wild-type cells bearing an integrated copy
of the kinase-deficient rad53 K227A–HA mutant allele and treated with 0.1% MMS for 2 h (MMS) and in cdc13-1 mutant cells, at either 25 or 34◦C, also bearing integrated rad53 K227A–HA.
labelled bands detected in undamaged cells are true Rad53-dependent phosphorylation signals. We infer that Rad53 is endowed with a constitutive kinase activity throughout the cell cycle and that this faint activity is further amplified in in vitro autophosphorylation assays.
Figure 3(A) shows that, using the IP-kinase as-say described above, there was still a Rad53 signal of activation in telomerase-negative cells deleted for
RAD9. These results thus confirm the actual
exist-ence of the weak signal seen with the IP-Western-blot assay (Figure 1C). Two other examples of the presence of a Rad53 signal of autophosphorylation in tlc1 rad9 cells are shown in Figures 3(B) and 5(B). Therefore, in telomerase-negative cells, unlike with other types of DNA damage (see for in-stance Figure 3C), Rad9 participates in Rad53 activ-ation, but yet another component also participates in Rad53 activation, at least when Rad9 is absent. To document further Rad53 activation in telomerase-negative (tlc1) cells, we next performed Rad53–HA IP-kinase assays on strains in which a particular checkpoint gene among those constituting the whole DNA damage checkpoint cascade had been genetic-ally inactivated. Rad53 perceives signals from DNA damage sensors and transmits them to downstream effectors after functional interactions with the so-called adaptors (Zhou and Elledge, 2000). Mec1– Ddc2 and Mec3–Ddc1–Rad17 represent two dis-tinct complexes that are loaded on to damaged DNA (Rouse and Jackson, 2002). Both complexes were
Figure 3 Telomerase loss activates Rad53 in a partially Rad9-independent manner
(A) Rad53 activation (in vitro autophosphorylation assays after IP with anti-HA antibody) in telomerase-negative cells (tlc1) deleted (tlc1 rad9, lower panel) or not (tlc1, upper panel) for RAD9 and carrying an HA-tagged chromosomal copy of RAD53. Numbers represent days following initiation of senescence at 30◦C. Note that on day 1, telomeres have already started to erode (results not shown; see the Materials and methods section). The Rad53 signal of activation on day 1 is therefore already elevated compared with that in non-senescing cells (wt). (B) Requirement of the Rad53 signal of activation for various checkpoint proteins after telomerase inactivation (tlc1 cells). Rad53 autophosphorylation signals (IP-kinase assays as in A, above) were measured in double mutants of the indicated relevant genotype. See text for in-terpretation. (C) Strains of the indicated relevant genotypes, carrying integrated RAD53–HA, were incubated at 34◦C (a re-strictive temperature for growth for the cdc13-1 mutation) for 2 h and Rad53 kinase signals were recorded.
required for activation of Rad53 during telomeric senescence (Figure 3B). Tel1, an ATM (ataxia telangiectasia)-mutated kinase of the same family as Mec1, was not required for Rad53 activation (Fig-ure 3B). However, we note that Rad53 was less active in tlc1 tel1 cells than in tlc1 TEL1+ cells (Fig-ure 3B). Rad53 IP-kinase assays on cdc13-1 strains in
the same mutation backgrounds as for tlc1 cells (Figure 3B) were performed for comparison (Fig-ure 3C). Importantly, in cdc13-1 cells, which suffer a well-documented case of DNA damage (Garvik et al., 1995), Rad53 activation was totally absent when
RAD9 had been deleted, thus highlighting the
par-ticularity of Rad53 activation in telomerase-negative cells deleted for RAD9. Incidentally, we noted that
cdc13-1 tel1 cells exhibited substantial activation of
Rad53 even at 25◦C (Figure 3C), which we interpret as being due to a synthetic interaction between the two mutations (results not shown).
Similar results concerning the activation of Rad53, either in the RAD9+ or rad9 background, were obtained when the est1, est2 or est3 mutations (Lendvay et al., 1996) were used in place of tlc1 to inactivate telomerase (results not shown).
Mrc1 activates Rad53 in telomerase-negative cells deleted for RAD9
Mrc1, another adaptor checkpoint protein, has been shown to functionally interact with Rad53 (Alcasabas et al., 2001). We reasoned that, in telomerase-negative cells deleted for RAD9, Rad53 might interact with Mrc1 to build up the activation of the kinase measured under such conditions (Fig-ure 3A). In agreement with this hypothesis, activ-ation of Rad53 was totally lost in tlc1 rad9 mrc1 cells (overexpressing RNR1 to rescue the lethality in-duced by the combination of rad9 mrc1 muta-tions; Alcasabas et al., 2001) (Figure 4A). Therefore the fraction of Rad53 activation that is observed in
tlc1 rad9 cells (Figures 3A and 3B) depends on
the presence of Mrc1 in the cell.
The above results suggested that both Rad53– Rad9 and Rad53–Mrc1 were activated during telomeric senescence. The participation of Mrc1 in the activation of Rad53 can only be assessed in the absence of Rad9, as seen above. Indeed, assessment of Rad53 kinase autophosphorylation signals in anti-HA immunoprecipitates from cells harbouring endo-genous HA–Mrc1 did not yield any detectable signal (results not shown). Similarly, HA–Rad9 immuno-precipitated from telomerase-negative cells did not contain any detectable Rad53 autophosphorylation signal (results not shown). Therefore, to assess the participation of Rad9 in the activation of Rad53 dur-ing telomeric senescence, we performed Rad53 auto-phosphorylation assays in cells deleted for MRC1. As
Figure 4 Rad53 activation in telomerase-negative cells relies both on Mrc1 and Rad9
(A) Rad53 activation relies on Mrc1 in telomerase-negative cells deleted for RAD9. Rad53 activation [assessed by in vitro autophosphorylation assays (IP-kinase assay), lower panel, or by electrophoretic shift up (IP-Western blotting), upper panel, from the same samples] was measured in tlc1 rad9 mrc1 triple mutants on day 3 after telomerase inactivation (lanes 3),
cdc13-1 rad9 mrc1 triple mutants grown at 34◦C for 2 h (lanes 6) or rad9 mrc1 double mutants, treated with 0.1% MMS for 2 h (lanes 5) or left untreated (lanes 4), surviving ow-ing to the overexpression of RNR1 under the control of the inducible GAL1 promoter (Alcasabas et al., 2001). All strains also carried an integrated copy of RAD53–HA for IP with an anti-HA antibody. Rad53–HA activation is also shown for wild-type cells, either left untreated (lanes 1) or treated for 2 h with 0.1% MMS (lanes 2). In all rad9 mrc1 strains, the Rad53 kinase signal was zero (lower panel), thus showing that no other protein, besides Rad9 and Mrc1, is capable of activating the Rad53 kinase during telomeric damage. Moreover, as ex-pected, in these strains, Rad53–HA was nevertheless phos-phorylated (assessed by visualizing the electrophoretic shift up, upper panel) due to the genotoxic agent MMS, the
cdc13-1 mutation or telomerase inactivation signalling
pre-sumably through Mec1, in agreement with previous find-ings. Note that the slower migrating band of Rad53–HA (the activated form) is also present in untreated rad9 mrc1 GAL-RNR1 cells (upper panel, lane 4), the mrc1 mutation representing in that case a damage by itself (results not shown; Alcasabas et al., 2001). (B) Rad53 activation relies on Rad9 in telomerase-negative cells deleted for MRC1. Rad53 activ-ation was assessed by in vitro autophosphorylactiv-ation assays (IP-kinase assay), as above in (A), in tlc1 mrc1 (lanes 6–9) or tlc1 mutants (lanes 2–5). Measurement in wild-type cells (wt, lane 1) is shown as a control. Since mrc1 rad9 cells cannot activate Rad53, as seen above in (A), the signals recorded here in the tlc1 mrc1 cells are provided by Rad9.
shown in Figure 4(B), Rad53 activation also took place in tlc1 mrc1 cells.
Rad53–Mrc1 is not activated when telomeric senescence results from telomere uncapping in the presence of functional telomerase
To know more about the activation of Rad53–Mrc1 in telomerase-negative cells, we took advantage of the fact that telomeric senescence can be induced in the presence of functional telomerase. Inactivating telomere end-protection proteins to induce telomeric senescence in the presence of functional telomerase has also been exploited in human cells (Karlseder et al., 2002; d’Adda di Fagagna et al., 2003). In hu-mans, this can be achieved by expressing a dominant-negative allele of TRF2 (Karlseder et al., 2002). In budding yeast, the combination of cdc13-1 and
yku70 mutations (at 25◦C) provoke telomere un-capping-induced senescence without prior telomere erosion (Grandin and Charbonneau, 2003). Such a phenomenon probably results from the fact that Cdc13 and Yku70 control distinct and com-plementary pathways of telomere end protection (Nugent et al., 1998; Polotnianka et al., 1998). Tel-omere uncapping-induced senescence in the cdc13-1
yku70 double mutant at 25◦C displayed kinet-ics different from that induced after telomerase loss, but nevertheless also eventually produced post-senescence survivors (Grandin and Charbonneau, 2003). High signals of Rad53 activation were present in cdc13-1 yku70 senescing cells at 25◦C from the time the strain was sporulated (Figure 5A, top panel, left lane) and, incidentally, after homo-logous recombination-based post-senescence survival had occurred (Figure 5A, upper panel, right lane). In these cells, Rad53 activation entirely depended on both Mec3 and Rad9 (Figure 5A, middle and bottom panels), as well as on Mec1 (results not shown). This was therefore in contrast with the Rad9-independent Rad53 activation in telomerase-negative senescing cells (Figure 3). Therefore these two distinct path-ways of telomeric senescence (resulting either from telomere erosion or from telomere uncapping) in-duce two different sequences of checkpoint activation events. Moreover, these results indicate that the sig-nal activating the checkpoint machinery in response to telomere erosion is unlikely to merely represent concomitant loss of attachment of Cdc13 and Yku to eroding telomere ends.
Figure 5 Rad53 activation triggered by telomere uncapping-induced senescence requires Rad9
(A) Rad53 activation was measured either during telomeric senescence (left lane of each of the three panels) or after post-senescence survival (by homologous recombination) has occurred (right lanes for all three panels) in cdc13-1 yku70 (top panel), cdc13-1 yku70 mec3 (middle panel) and
cdc13-1 yku70 rad9 (bottom panel) mutant cells after IP
of the genomic copy product of RAD53–HA with HA anti-body and in vitro autophosphorylation assays (IP-kinase as-says). (B) cdc13-1 tlc1 rad9 (lane 2) and yku70 tlc1
rad9 (lane 4) mutants, as well as their RAD9+counterparts (lanes 1 and 3) (all at day 1 after induction of senescence by genetic inactivation of telomerase due to the tlc1 mutation), exhibit Rad53 activation (in vitro autophosphorylation assays after IP of chromosomal Rad53–HA with anti-HA antibody) during telomeric senescence, just like tlc1 rad9 mutants (lanes 6–9) at the indicated time (in days) after telomerase in-activation. Rad53 activation in wild-type RAD53–HA cells is shown for comparison (lane 5). (C) Rad53 activation (meas-ured as above) in yku70, yku70 mec3 and yku70 rad9 mutant cells and wild-type cells (wt) as a control, incub-ated for 2 h at 37◦C, a temperature that induces a severe growth defect in cells bearing the yku70 mutation (results not shown).
Two hypotheses, at least, arose to explain the lack of activation of Rad53 in cdc13-1 yku70 rad9 sen-escing cells but not in telomerase-negative (tlc1)
rad9 cells. First, in tlc1 rad9 cells, Cdc13 and/or
Yku might mediate the recruitment of Mrc1. Sec-ondly, in cdc13-1 yku70 rad9 senescing cells, the presence of telomerase might prevent activation of Rad53–Mrc1. To determine which one of these two hypotheses was correct, we set out to measure Rad53 activation after having simultaneously inactivated both telomere protection systems and, to this end, we constructed the cdc13-1 tlc1 rad9
RAD53-HA and yku70 tlc1 rad9 RAD53-HA triple
mutants. Rad53 activation signals were found to increase both in cdc13-1 tlc1 rad9 cells and in
yku70 tlc1 rad9 cells, just like in tlc1 rad9
cells (Figure 5B). Therefore, in the Rad53–Mrc1 ac-tivation function, telomerase loss somehow domin-ates over loss of telomere capping when both are simultaneously inflicted. In other words, deleting telomerase activates Rad53–Mrc1 even though the presence of the cdc13-1 mutation makes that Rad9 should be required to activate Rad53, at least at 34◦C. Note that the yku70 mutation on its own also ac-tivates Rad53 at 37◦C, as previously noted (Teo and Jackson, 2001), an event that necessitates Rad9 (Fig-ure 5C), just like with cdc13-1 (Fig(Fig-ure 3C). Unfortu-nately, we could not test whether telomerase was still dominant over simultaneous loss of Cdc13 and Yku70 as we could not derive the cdc13-1 yku70
tlc1 rad9 RAD53-HA strain. These experiments
suggest that it was the presence of telomerase rather than the loss of protection by Cdc13 and Yku that prevented activation of Rad53 (by Mrc1) in cdc13-1
yku70 rad9 senescing cells.
Rad53 and Rad9, but not Mrc1, are required for telomerase loss-induced cell-cycle arrest
The primary goals of checkpoint activation are, first, to halt cell-cycle progression in response to damage, and, secondly, to activate the expression of DNA re-pair genes. To evaluate the respective roles of the Rad53–Rad9 and Rad53–Mrc1 functional complexes in cell-cycle arrest upon loss of telomerase function, we analysed the cell proliferation status of various mutants under the microscope. Figure 6(A) shows that on day 5 after telomerase inactivation (tlc1), time at which virtually all tlc1 cells had com-pletely stopped dividing, tlc1 cells deleted for either
Figure 6 Representative morphologies of telomerase-negative cells tlc1 (A) and cdc13-1 cells (B) in various checkpoint mutation backgrounds, as indicated, at 30◦C (A) or at the indicated temperature (B)
(A) Cells were fixed every day after induction of telomeric senescence. Note that day 1 was estimated to correspond to approx. 40 generations of growth after effective inactivation of telomerase (the time of sporulation of the diploid), whereas days 2, 3, 4 and 5 corresponded to approx. 55, 70, 85 and 100 generations after telomerase inactivation respectively (see the Materials and methods section). In parallel, cell-cycle stages were determined at the time of crisis in tlc1 cells (day 5). (B) Cells were fixed 2 h after incubation at the indicated temperature. (A, B) Scale bars, 10 µm. (C) Deleting MRC1 in cdc13-1 yku70 senescing cells did not suppress the damage-induced cell-cycle arrest (compare top and middle panels). RAD9-deleted strains provided a positive control (bottom panel). All strains were grown at 25◦C. Scale bar, 10 µm.
RAD53 or RAD9 still continued to actively divide.
In contrast, tlc1 mrc1 cells had also stopped di-viding (Figure 6A). Although this result is some-what biased by the existence of a synthetic growth defect between the tlc1 and mrc1 mutations (Fig-ure 6A; N. Grandin and M. Charbonneau, unpub-lished work), it is nevertheless clear that the absence of Mrc1 does not lead to the bypass of the DNA-damage-induced cell-cycle arrest. Therefore Mrc1 forms a functional complex with Rad53 that does not appear to impinge on cell-cycle progression, unlike the Rad53–Rad9 complex. Similar to the situation found in tlc1 mrc1 cells (Figure 6A), cell-cycle ar-rest accompanying telomere uncapping-induced sen-escence required Rad9 but not Mrc1 (Figure 6C). We noted that the cdc13-1 and mrc1 mutations ex-hibited a severe synthetic growth defect, at 25◦C, similar to that seen when the cdc13-1 and yku70 mutations were combined (Figure 6C; N. Grandin and M. Charbonneau, unpublished work).
These analyses also revealed that besides the case of RAD53, RAD9 and MRC1, MEC1, MEC3, CHK1 and DUN1 (like RAD53 and RAD9), but not TEL1 (like MRC1), were required for the complete cell-cycle arrest of telomerase-negative cells (Figure 6A) or of cdc13-1 cells at restrictive temperature (Fig-ure 6B). Deletion of either MEC1 or MEC3 was among the more potent in inhibiting cell-cycle arrest (Figures 6A and 6B), as expected from the fact that the Mec1 and Mec3 complexes control all downstream checkpoint events. Similarly, deletion of RAD9 re-sulted in an almost complete lack of cell-cycle arrest in response to telomerase inactivation, an effect lar-ger than deletion of either RAD53 or CHK1 alone (Figures 6A and 6B). This was because Rad9 parti-cipates in both of two parallel checkpoint pathways, one controlled by Rad53 and the other one by Chk1 (Sanchez et al., 1999).
Rad9, Mrc1, Dun1 and Chk1 are activated after telomerase inactivation
To document further the activation of the DNA damage checkpoint machinery during telomeric sen-escence, we monitored the activation of epitope-tagged copies of Rad9, Mrc1, Dun1 and Chk1 in-tegrated at their respective locus and therefore ex-pressed at endogenous levels. For all four proteins, activation can be conveniently assessed by visual-izing on Western blots an electrophoretic
mobil-Figure 7 Activation of Dun1, Rad9 and Mrc1 during telomerase loss-induced senescence
Electrophoretic mobility shifts reflecting phosphorylation of 2 HA–Rad9, 2 HA–Mrc1 and 2 HA–Dun1 upon activation were detected during telomeric senescence resulting from tel-omerase inactivation (in tlc1 cells). cdc13-1 2 HA-RAD9 and
cdc13-1 2 HA-DUN1 cells grown at the restrictive
temperat-ure of 34◦C for 2 h and wild-type 2 HA-MRC1 cells treated for 2 h with 0.2 M HU were used as controls. Phosphoryl-ation of Rad9, Mrc1 and Dun1 (expressed from chromosomal copies of 2 HA–RAD9, 2 HA–MRC1 and 2 HA–DUN1 respect-ively) was assessed by Western blotting after IP with anti-HA antibody (IP-Western).
ity shift corresponding to phosphorylation induced by DNA damage or DNA replication stress (Allen et al., 1994; Vialard et al., 1998; Sanchez et al., 1999; Alcasabas et al., 2001). Figure 7 shows the activation of HA–Rad9, HA–Mrc1 and HA–Dun1 after loss of telomerase function. Chk1 was also found to be activ-ated in telomerase-negative cells (results not shown). These results confirm the implication of these check-point proteins in telomeric senescence uncovered by genetic analysis (Figure 6A) by showing that they are indeed activated by phosphorylation concomitantly with their action on cell-cycle progression.
Discussion
Mrc1 activates Rad53 after loss of telomerase function but not after loss of telomere capping
In the present study, we show that in budding yeast cells experiencing telomeric senescence due to
telomerase loss, Rad53 activation is partially in-dependent of Rad9. Assessment of Rad53 activation in telomerase-negative cells simultaneously deleted for RAD9 and MRC1 further established that Mrc1 was responsible for the observed Rad9-independent Rad53 signal. Although the DNA damage response to telomere dysfunction begins to be unravelled, there remain many gaps to be filled in (d’Adda di Fagagna et al., 2004). At genome level, the transcription re-sponse to telomeric senescence shares many features with the response to double-strand breaks at other places within the genome, but yet exhibits a few dif-ferences with it (Nautiyal et al., 2002). At the level of the DNA damage itself, the response to telomeric damage globally involves the same checkpoint com-ponents as those involved in response to a double-strand DNA break (d’Adda di Fagagna et al., 2003; Ijpma and Greider, 2003), with a difference, however, concerning the participation of Rad53 and Rad9 in cell-cycle arrest (Enomoto et al., 2002).
Although previous studies have implicated a num-ber of checkpoint proteins in the response to inactiv-ation of telomerase (Enomoto et al., 2002; Nautiyal et al., 2002; d’Adda di Fagagna et al., 2003; Ijpma and Greider, 2003), the present one is the first to im-plicate Mrc1 in that response. The idea that Mrc1 represents the protein that activates Rad53 in telomerase-negative cells deleted for RAD9 was not unexpected. Indeed, Mrc1 has been previously shown to be required for Rad53 activation during DNA rep-lication stress and Rad53 activation was found to be totally absent from HU (hydroxyurea)-treated rad9
mrc1 double mutants (Alcasabas et al., 2001). It is to
be noted that in telomerase-negative cells, Rad9 also contributed to Rad53 activation, as Rad53 activation took place in tlc1 mrc1 cells (Figure 4B). Therefore it is probable that both a Rad53–Rad9 complex and a Rad53–Mrc1 complex are activated during telomere erosion. By Western-blot analysis the contribution of Rad9 to Rad53 activation in telomerase-negative cells appeared to be major (Figure 1C). By the auto-phosphorylation assay, the Rad53–Rad9 signal of ac-tivation (in tlc1 mrc1 cells) appeared to be larger than the Rad53-Mrc1 signal in tlc1 rad9 cells (Figures 3A, 3B, 4B and 5B). However, obviously, the exact participation of Mrc1 and Rad9 in Rad53 activation in terms of number of Rad53 molecules present in each complex, as well as the kinetics of activation of these complexes cannot be accurately
known at the moment. This is particularly critical as, although a physical association between endo-genous 2 HA–Rad9 and Rad53-9 Myc could be readily detected in telomerase-negative cells, inter-actions between 2 HA–Mrc1 and Rad53-9 Myc were not observed in senescent cells (N. Grandin and M. Charbonneau, unpublished work). In fact, although Rad53 was found to phosphorylate Mrc1
in vitro, physical interactions between the two
pro-teins could not be detected in vivo in HU-treated cells (Osborn and Elledge, 2003), whereas, in Sc. pombe, Mrc1–Cds1 interactions could be detected only by two-hybrid method (Tanaka and Russell, 2001).
It is probable that two distinct types of DNA insult activate the putative Rad53–Rad9 and Rad53–Mrc1 complexes during telomere erosion. Indeed, telomeric senescence resulting from inactivation of telomerase can be regarded as provoking both a DNA rep-lication stress and DNA damage. Thus telomere erosion resulting from telomerase inactivation re-moves the telomere end-protection proteins, and un-protected telomeric DNA eventually takes the form of double-strand breaks. Moreover, synthesis of the telomeric lagging strand is coupled with telomere replication (Vega et al., 2003), and in this respect, the absence of telomerase may provoke a DNA repli-cation stress. Alternatively, the absence of telomerase might generate hyper-recombinogenic structures and activate DNA replication as a DNA repair mech-anism (Chakhparonian and Wellinger, 2003; Vega et al., 2003). However, in this respect, inactivation of Cdc13, in cdc13-1 cells, or of Yku can also generate uncontrolled telomeric recombination (Polotnianka et al., 1998; Grandin et al., 2001). Therefore the activation of both Rad53–Rad9 and Rad53–Mrc1 after loss of telomerase may reflect these two distinct types of damage.
Unlike in HU-induced checkpoint activation in which Mrc1 has been implicated in cell-cycle arrest (Alcasabas et al., 2001), we found that Mrc1 was not involved in telomerase loss-induced cell-cycle arrest (Figure 6A). Defining the activation of the check-point adaptors Rad9 and Mrc1 is not an easy task because of the choice of the criteria to be taken into account. Thus, on the basis of electrophoretic mo-bility shift, both Rad9 and Mrc1 were found to be activated during DNA replication stress or DNA damage, including that induced by telomeric sen-escence (Alcasabas et al., 2001; Toh and Lowndes,
2003; the present study). Previous results support the conclusion that Mrc1, and Rad9 as well, are phos-phorylated by the phosphoinositide kinases Mec1 and Tel1 (Vialard et al., 1998; Alcasabas et al., 2001; Osborn and Elledge, 2003). Therefore phosphoryl-ation of Mrc1 presumably represents the first step in the activation of the putative Rad53–Mrc1 complex but can, in some instances, also represent the last step. For instance, Mrc1 was phosphorylated during
cdc13-1-induced DNA damage but nevertheless did
not participate in the concomitantly induced cell-cycle arrest (Alcasabas et al., 2001). If Rad53 kinase activity is to be taken as a criterion for activation, Mrc1 did not mediate Rad53 activation during telo-mere uncapping-induced telomeric senescence, like in all documented types of DNA damage (Toh and Lowndes, 2003; the present study). However, in both HU-induced checkpoint activation (Alcasabas et al., 2001; Katou et al., 2003; Osborn and Elledge, 2003) and telomerase loss-induced checkpoint activation (present study), Mrc1 was responsible for activating Rad53 kinase activity. Another criterion for the ac-tivation of Rad9 and Mrc1 is their requirement for DNA-damage-induced cell-cycle arrest. In this re-gister, it is noticeable that Mrc1 is required for cell-cycle arrest in HU-treated cells (Alcasabas et al., 2001), but not in telomerase-negative cells (the pre-sent study). In conclusion, in response to DNA replication stress (HU treatment), Mrc1 was phos-phorylated, activated Rad53 kinase activity and con-trolled cell-cycle progression (Alcasabas et al., 2001; Osborn and Elledge, 2003). On the other hand, in telomerase-negative cells, Mrc1 was phosphorylated, activated Rad53 kinase activity, but was not required for cell-cycle arrest.
Recently, d’Adda di Fagagna et al. (2003) ana-lysed the checkpoint response to telomeric sen-escence in human cells using a system in which a dominant-negative allele of TRF2 produces telomere uncapping. This system allows to achieve telomeric senescence in the presence of functional telomerase and normal telomere length (Karlseder et al., 2002; Smogorzewska and de Lange, 2002), the same object-ive as that attained in the present study using cdc13-1
yku70-induced senescence, a system previously
de-scribed in detail (Grandin and Charbonneau, 2003). No difference between the two types of human te-lomeric senescence was observed in the activation of several checkpoint proteins, including CHK2
(bud-ding yeast Rad53) or recruitment to foci of DNA damage (d’Adda di Fagagna et al., 2003). However, it should be noted that the yeast and human systems are not completely equivalent as cdc13-1 yku70 cells have extensive single-stranded telomere regions (Garvik et al., 1995; Polotnianka et al., 1998) that have not been reported in TRF2 mutant cells. More-over, telomeres are shorter in yeast than in humans, possibly accounting for different mechanisms in these two telomere erosion-independent senescence pro-cesses. To our knowledge, the participation of human MDC1 or 53BP1 (potential homologues of budding yeast Rad9) and Claspin (yeast Mrc1) in the activ-ation of CHK2 has not been documented. It would be important, by further analysis of human cells or other systems, to know whether the activation of Rad53 kinase activity by Mrc1 after telomerase loss in budding yeast has been conserved during evolution. In telomerase-negative cells, activation of Rad53 by Mrc1 might play a novel role in a pathway of main-tenance of genome stability, which is yet to be de-scribed. An attractive possibility is that telomerase might be required to stabilize the replication forks (when they reach the telomeres) and that, in cells lack-ing telomerase, a checkpoint specifically senslack-ing this defect would activate Rad53–Mrc1, which would then be required to stabilize the stalled forks. Our results on senescing cells suffering a double telo-meric defect (loss of telomerase plus inactivation of either CDC13 or YKU70; Figure 5B) may suggest a ‘dominant’ effect of telomerase presence at telomeres over loss of function of Cdc13 or Yku70. One could imagine that telomerase might physically interact with Mrc1 in order to couple late DNA replication and telomere replication and that, therefore, loss of telomerase would inadvertently ‘unmask’ Mrc1, thus activating Mrc1-dependent Rad53 kinase activity. Preliminary experiments have failed to detect such interactions; however, these may exist under particu-lar conditions, at faint levels or at particuparticu-lar moments of the cell cycle.
Implication of checkpoint proteins in telomeric senescence-induced cell-cycle arrest
The present study confirms the conclusion reached by three recent studies (two in yeast and one in hu-mans) that short telomeres are recognized as DNA damage and signal G2/M arrest (Enomoto et al., 2002; d’Adda di Fagagna et al., 2003; Ijpma and
Greider, 2003). The pattern of cell-cycle arrest de-scribed here for telomerase-negative cells (Figure 6A) is very similar to that described previously (Ijpma and Greider, 2003). We note that, in both studies, at the time of crisis there were still a substantial number of telomerase-negative cells arrested in mitosis. This raises the question of whether telomere erosion might also activate a spindle checkpoint, as previously noted in cdc13-1 cells suffering telomeric DNA damage (Wang et al., 2000; Maringele and Lydall, 2002). Such a possibility has been only partially answered, as Ijpma and Greider (2003) reported that inhi-bition of the Mad2-dependent spindle checkpoint did not alter the cell-cycle stages in telomerase-negative cells. However, it should be noted that inactivation of Bub2, which is essential for the spindle misorienta-tion checkpoint, but not of Mad2, which funcmisorienta-tions in the spindle assembly/integrity checkpoint, resulted in enforcement of cell-cycle progression after cdc13-1-induced telomeric DNA damage (Wang et al., 2000; Maringele and Lydall, 2002).
We also note that the participation of Rad53, Chk1 and Dun1 in the enforcement of senescence-induced cell-cycle arrest was only partial (partial cell-cycle ar-rest in tlc1 checkpoint double mutants). This con-firms that, in this situation too, Rad53–Rad9–Dun1 and Chk1–Rad9 provide two parallel pathways for halting cell-cycle progression in response to DNA damage (Sanchez et al., 1999). We also find here that activation of most of the major checkpoint pro-teins described to date, with the noticeable exceptions of Tel1 and Mrc1, is required for the enforcement of the cell-cycle arrest induced by telomeric senescence, thus confirming previous results (Enomoto et al., 2002; Ijpma and Greider, 2003). However, in our hands, both Rad9 and Rad53 were required for the complete cell-cycle arrest of telomerase-negative cells, in contradiction with the study by Enomoto et al. (2002), but in agreement with that by Ijpma and Greider (2003).
It is not yet clear what advantages activation of the checkpoint proteins confers on the senescing cells in the short term. In fact, the damage in these cells is such that homologous recombination-based post-senescence survival (Lundblad and Szostak, 1989; Lundblad and Blackburn, 1993) will take place whether the checkpoint machinery is functional or not. Indeed, telomerase-negative cells were found to undergo senescence and post-senescence events
with similar kinetics whether the checkpoint path-way had been inactivated or not (N. Grandin and M. Charbonneau, unpublished work). However, it should be noted that genetic inactivation of Mec1 or Tel1 led to a drastic change in the choice of survival type (type I and II have been distinguished on the basis of the nature of the homologous recombination substrate; Lundblad and Blackburn, 1993; Teng and Zakian, 1999), both Mec1 and Tel1 mediating type II recombination (Tsai et al., 2002). In the same vein, DNA damage checkpoint proteins have been found to have a function in activating DNA repair through Rad55 (Bashkirov et al., 2000). Therefore both treat-ment of the DNA lesions by DNA repair proteins and activation of the latter by the checkpoint proteins might be important for orchestrating survival under the optimal conditions. In this respect, the exonu-clease Exo1, which has been shown to process dam-aged DNA at telomeres (Maringele and Lydall, 2002), was also found to control the selection for homologous recombination type during post-senescence survival (Bertuch and Lundblad, 2004; Maringele and Lydall, 2004).
In conclusion, the present data further document the processes of recognition of the DNA damage gen-erated by the loss of telomerase function in an im-portant genetic model system. Perfect knowledge of these pathways is crucial not only for fundamental research but also for better understanding cell-cycle deregulation in tumour cells. Indeed, in human aging cells, these checkpoint mechanisms are of particular importance for coping with telomeric DNA dam-age that frequently occurs due to natural telomere erosion in somatic tissues lacking active telomerase. The present data documenting the existence in telomerase-negative cells of an active Rad53–Mrc1 complex that is not used for cell-cycle arrest also sug-gest that new functions for checkpoint proteins most probably await discovery.
Materials and methods Strains and plasmids
All mutant strains used in the present study were backcrossed at least five times against the BF264-15D genetic background used in our laboratory (Grandin et al., 1997) before experiment-ation. The tel1::KanMX4 strain was purchased from Research Genetics (Huntsville, AL, U.S.A.) and the mrc1::KanMX4, dun1::KanMX4, chk1::KanMX4, yku70::KanMX4, est1:: KanMX4/EST1, est2::KanMX4/EST2 and est3::KanMX4/EST3 strains were obtained from Euroscarf (Frankfurt, Germany). The
mec1::TRP1 sml1::KanMX4 and rad53::TRP1 sml1::KanMX4 mutant strains were obtained from the Hartwell and Emili labor-atories (Emili, 1998; Emili et al., 2001). The tlc1::LEU2 strain was from the Gottschling laboratory (Singer and Gottschling, 1994). The rad9::LEU2 and cdc13-1 strains were from the Hartwell laboratory (Garvik et al., 1995). The mec3::TRP1 strain was from the Lucchini laboratory (Longhese et al., 1996). The yku70/hdf1::URA3 strain was from the Petes laboratory (Porter et al., 1996). The tlc1::TRP1, tlc1::URA3, rad9::TRP1 and rad9::URA3 strains were obtained by disruption of the LEU2 marker by the TRP1 or URA3 marker in the backcrossed tlc1::LEU2 and rad9::LEU2 strains respectively. Yeast cells were grown in YEPD medium (1% yeast extract, 2% Bacto-peptone, 2% glucose, 0.005% adenine and 0.005% uracil) at 30◦C unless otherwise indicated. The integrative TRP1-based pRS304 RAD53-2 HA construct, a gift from A. Emili (Banting and Best Department of Medical Research, CH Best Institute, Toronto, ON, Canada) (Emili, 1998; Emili et al., 2001), was used as such and also subcloned into YIp211 (integrative URA3 plasmid). A truncated form of the rad53 K227A mutant (kinase-deficient) allele was constructed in an integrative YIp211 plasmid, after PCR amplification from YIp5-rad53 K227A (Bashkirov et al., 2003), a gift from W.-D. Heyer (Division of Biological Sciences, University of California at Davis, Davis, CA, U.S.A.) and expressed from the RAD53 genomic locus. RNR1 open reading frame plus 120 bp of post-stop sequences was cloned in a YEp112 (2 µm, episomal, TRP1-based) plasmid in which it was under the control of the inducible GAL1 promoter. Each truncated epitope-tagged DNA construct was integrated at its corresponding genomic locus after digesting with a restriction enzyme unique to the gene in question. Details of the cloning procedures are available upon request.
IP, immunoblotting and measurement of Rad53 autophosphorylation
In all experiments, proteins were immunoprecipitated from yeast cell extracts and separated by electrophoresis (see below). Rad53 activation was assessed by in vitro autophosphorylation (Pellicioli et al., 1999) on 50 ml of yeast cell culture/sample after IP with anti-HA antibody and separation by electrophoresis (IP-kinase assays). All strains analysed harboured a unique chro-mosomal copy of RAD53-tagged in 3with 2 HA epitope (see above). Pellets from cultivated cells were subjected to mech-anical disruption (in the presence of 425–600 µm glass beads, Sigma) in lysis buffer [50 mM Tris/HCl, pH 7.5, 140 mM NaCl, 1 mM EDTA, 1% Triton X-100, 0.1% sodium deoxycholate and phosphatase and protease inhibitors (Roche, Lewes, East Sussex, U.K.)], followed by centrifugation in Eppendorf tubes at 12 000 g for 10 min at 4◦C to clarify the soluble fraction. Lysates were then used for IP with mouse monoclonal anti-HA raw as-cites fluid (16B12, BabCO, Richmond, CA, U.S.A.; Eurogentec, Angers, France). Washed protein-bound Protein A–Sepharose beads (Amersham Biosciences) were incubated in 25 µl of re-action mixture containing 50 mM Tris/HCl (pH 7.5), 5 mM MgCl2, 1 mM dithiothreitol and 0.5 µCi/sample [γ -32P]ATP
(>7000 Ci/mmol; Amersham Biosciences, Orsay, France). Im-munoprecipitates were resolved on polyacrylamide gels before transfer on to a nitrocellulose membrane. For immunoblot ana-lysis, a mouse anti-HA monoclonal 12CA5 antibody (Roche) was used on gel electrophoresis-separated proteins after IP
on 100 ml cultures (IP-Western blots). HA-tagged proteins were then detected by chemifluorescence via fluorescein-labelled anti-mouse secondary antibodies and a tertiary anti-fluorescein alkaline phosphatase conjugate (ECF Western blotting kit, Amersham Biosciences) and signals were analysed using a Storm phosphoimager (Amersham Biosciences).
Analysis of the kinetics of senescence/survival and of growth rates
Telomerase-negative cells undergo senescence in 75–100 gener-ations (Lendvay et al., 1996). In all kinetics, senescence of tlc1 or cdc13-1 yku70 mutant cells, and of their derivatives, was initiated after the tlc1/TLC1+ or cdc13-1 yku70/CDC13+ YKU70+diploids were induced to sporulate. Because the spores needed approx. 1 day to germinate, followed by approx. 2 days of growth to obtain enough cells from each spore, selection of the genotypes of the spores could not be started until 3 days after sporulation. The time of re-streak of the spores on to the selec-tion plates was counted as day 0. Approximately 24 h later, the first analysed cells (referred to as samples of day 1 of senescence in the Figures) were taken from the selection plates and grown overnight in liquid YEPD in order to obtain enough material for protein cell extract and DNA genomic preparations. Cells were harvested when in exponential growth phase. Meanwhile, cells from the original selection plates were re-streaked on to YEPD plates every day in order to cover the entire period of telomeric senescence. The occurrence of senescence was determined both from the appearance of post-senescence survivors on the plates and from the analysis of telomere structure by Southern blotting with a telomeric probe, as described previously (Grandin et al., 1997). Approx. 40 generations elapsed between the time the spores germinated and the time the cells were harvested for pre-paration of the first sample (day 1), including the 2 day growth on the sporulation plate before selection and the last overnight growth in liquid YEPD. Days 2, 3, 4 and 5 corresponded to approx. 55, 70, 85 and 100 generations of growth respectively following sporulation, hence, effective telomerase inactivation. Crisis, the time at which all cells had basically stopped dividing that is shortly followed by post-senescence survival, occurred between days 4 and 5, depending on the strains. Unfortunately, it is not possible in Sa. cerevisiae, unlike in Sc. pombe for instance, to perform mass-spore selection, thus eliminating the possi-bility to analyse at time points earlier than day 1 molecular events normally necessitating a minimal amount of cells without requiring colony formation.
For cell morphology analysis, cells were withdrawn at inter-vals from liquid cultures and fixed with formaldehyde. Cells were then observed in a BX50 Olympus light microscope, using a×40 lens and Nomarski optics, and photographs were taken using an Olympus digital camera.
For determination of cell-cycle stages, cells were fixed with ethanol before staining with propidium iodide. For each time point, approx. 150 cell bodies were counted under a fluorescence microscope.
Acknowledgments
We thank A. Emili, W.-D. Heyer, L. Hartwell (De-partment of Genetics, University of Washington,
Seattle, WA, U.S.A.), T. Petes (Department of Bio-logy and Curriculum in Genetics and Molecular Biology, University of North Carolina at Chapel Hill, Chapel Hill, NC, U.S.A.), G. Lucchini (Di-partimento di Biotecnologie e Bioscienze, Universita degli Studi di Milano-Bicocca, Milan, Italy) and D. Gottschling (Fred Hutchinson Cancer Research Center, Seattle, WA, U.S.A.) for the gifts of strains and plasmids and C. Damon for technical assistance. This work was supported by grants from the ‘As-sociation pour la Recherche contre le Cancer’ and the ‘Comit´e D´epartemental de la Savoie de la Ligue Nationale contre le Cancer’.
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Received 9 December 2004/3 March 2005; accepted 10 March 2005
