Using a model system, we have shown that replicative senescence is accompanied by a 16-fold increase in base substitution and frameshift mutations near a chromosome end. The increase was dependent on error-prone polymerases required for the mutagenic response to DNA lesions that block the replication fork.

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SACCHAROMYCES cerevisiae cells lacking telomerase, a ribonucleoprotein complex required for telomere replication, experience progressive telomere degradation that culminates in replicative senescence (LENDVAY et al. 1996). Deletions that encompass the CAN1 locus located 32 kb from the telomere on the left arm of chromosome V accumulated during senescence (HACKETT et al. 2001; HACKETT and GREIDER 2003) and have been attributed to replication fork stalling (MOTEGI et al. 2006). Since replication fork stalling also generates mutations (QUAH et al. 1980), we investigated the effect of telomere dysfunction on the generation of mutations at the CAN1 locus.

Mutation rate analysis:

We examined the behavior of mutants defective for EST2, which encodes the catalytic subunit of telomerase (COUNTER et al. 1997)(Table 1). We determined the mutation rate at the CAN1 locus using an assay that selects against deletions in serial cultures of wild-type and est2 mutant cells. We observed no significant change from the wild-type rate of CAN1 mutation in est2 cultures before senescence (P = 0.07), a 16-fold increase during senescence (P < 0.0001), and the restoration of wild-type levels upon recovery (P = 0.09) (Figure 1A). No senescence-dependent changes in the mutation rate at the CYH2 locus, located 310 kb from its telomere on the left arm of chromosome VII, were observed in est2 mutant cultures before (P = 0.68), during (P = 1.0), or after replicative senescence (P = 1.0) (Figure 2), suggesting that the mutagenic effect is restricted to telomere-proximal sequences. The can1 mutation spectrum observed for senescent est2 mutant cells was similar to that of wild type (Table 2), suggesting that the mechanism of senescence-dependent mutagenesis in est2 cells may be similar to the mechanism of spontaneous mutagenesis in wild-type cells. Since 50–70% of spontaneous mutagenesis has been attributed to the action of error-prone polymerases (QUAH et al. 1980), we investigated whether they were involved in the mechanism underlying senescence-dependent mutagenesis. We examined the effects of mutations in the RAD30, REV7, and REV1 genes, which are required for error-prone polymerase function in yeast (HARACSKA et al. 2000; JOHNSON et al. 2000; GOODMAN 2002; PRAKASH et al. 2005). Mutation rates in est2 rad30 cultures before (P = 0.32), during (P = 0.34), and after senescence (P = 0.15) were not significantly different from those in est2 cultures (Figure 1, A and B), suggesting that Pol does not contribute to senescence-dependent mutagenesis. In contrast, the rev1 and rev7 mutations completely suppressed senescence-dependent mutagenesis, as the CAN1 mutation rates were not significantly different from those of rev1 and rev7 mutants before (P = 0.46, rev1; P = 0.53, rev7), during (P = 0.84, rev1; P = 0.09, rev7), or after senescence (P = 1.0, rev1; P = 0.2, rev7) (Figure 1, A and B). Southern blot analysis of representative canavanine-resistant mutants collected from senescent cells revealed that all had unrearranged can1 loci, consistent with base substitution and frameshift mutations (D. MEYER and A. BAILIS, unpublished data). These data suggest that Rev1 and DNA polymerase are required for senescence-dependent mutagenesis, perhaps through mutagenic bypass of DNA replication lesions generated during replicative senescence. The minimal effects of the rad30, rev1, and rev7 mutations on the growth, senescence, and recovery of est2 rad30, est2 rev1, and est2 rev7 cells (Figure 3, A and B) suggest that the mutation of telomere-proximal sequences does not contribute to the initiation of or recovery from senescence.

View this table: In this window In a new window   TABLE 1 S. cerevisiae strains used in this study

 

View larger version (9K): In this window In a new window Download PPT slide   FIGURE 1.— REV1 and REV7, but not RAD30, are required to observe increases in CAN1 mutation rate during replicative senescence in est2 mutant cells. (A) CAN1 mutation rate of () wild type, () est2, rev1, (•) rev7, () rad30, and () exo1 mutant cells at the indicated time points. (B) CAN1 mutation rate of () est2 rev1, () est2 rev7, () est2 rad30, () est2 exo1, (x) rad30 rev7, and (*) est2 rad30 rev7 mutant cells at the indicated time points. Spore colonies of the appropriate genotype were obtained from freshly dissected tetrads of ABX1269, ABX1727, and ABX1729 (Table 1) and dispersed in water. Aliquots were removed to determine viability following dilution, plating on to YPD medium, and incubation for 3 days at 30°. The remainder was plated onto synthetic medium lacking arginine and containing 60 µg/ml canavanine, and incubated for 3 days at 30°. The colonies arising on the canavanine plates were counted and replica plated to synthetic medium lacking uracil and the numbers of Ura– and Ura+ colonies were determined after overnight incubation at 30°. CAN1 mutation frequency was determined by dividing the number of Canr Ura+ colonies by the number of viable cells plated for each spore colony. CAN1 mutation rate was determined using the median CAN1 mutation frequency from at least 10 independent trials (LEA and COULSON 1949). Statistical significance was tested by determining the number of trials with each strain that were above and below the group median frequency and then performing 2-analysis and Fisher's exact test. This process was repeated at five successive growth intervals 25 generations apart, using single colonies that arose on the YPD viability plates.

 

View larger version (8K): In this window In a new window Download PPT slide   FIGURE 2.— CYH2 mutation rate does not increase in telomerase-deficient cells during replicative senescence. CYH2 mutation rate of () wild type and () est2 mutants was determined at the indicated time points. Wild-type and est2 mutant spore colonies were obtained from freshly dissected tetrads of ABX1429 (Table 1) and dispersed in water. Aliquots were removed to determine viability following dilution, plating on to YPD medium, and incubation for 3 days at 30°. The remainder was plated onto synthetic medium containing 1 µg/ml cycloheximide and incubated for 3 days at 30°. CYH2 mutation frequency was determined by dividing the number of Cyhr colonies by the number of viable cells plated for each spore colony. CYH2 mutation rate was determined using the median CYH2 mutation frequency from at least 10 independent trials (LEA and COULSON 1949). Statistical significance was tested by determining the number of trials with each strain that were above and below the group median frequency and then performing 2-analysis and Fisher's exact test. This process was repeated at two additional growth intervals 25 generations apart, using single colonies that arose on the YPD viability plates.

 

View this table: In this window In a new window   TABLE 2 CAN1 mutation spectra in wild-type and mutant cells

 

View larger version (9K): In this window In a new window Download PPT slide   FIGURE 3.— Rev1, Rev7, and Rad30 do not significantly affect senescence and subsequent recovery. (A) () Wild type, (•) rev7, () rev1, and () rad30. (B) () est2, () est2 rev7, () est2 rev1, () est2 rad30, (x) rad30 rev7, and (*) est2 rad30 rev7. Serial liquid growth was performed as described previously (HACKETT et al. 2001). During each day of serial liquid growth, hemocytometer counts were performed to determine the number of cell bodies, after which 500 cells were plated to YPD and incubated at 30° for 3 days. Colonies were then counted and divided by 500 to determine plating efficiency. Finally, viability was determined following each day of growth in liquid culture by taking the number of cell bodies and multiplying by the plating efficiency. Results are the mean ±2 SE from at least eight independent samples of each indicated genotype.

 
Exo1 has been suggested to be necessary for destabilizing the CAN1 locus during senescence by promoting exonucleolytic degradation from the telomere (HACKETT and GREIDER 2003). We observed no significant change in the rates of CAN1 mutation in est2 exo1 cultures before (P = 0.89) or after senescence (P = 1.0) from that observed in exo1 cultures, but did see a significant 10-fold increase during senescence (P < 0.0001) (Figure 1, A and B). These data suggest that Exo1-dependent nucleolytic degradation is not required to observe senescence-dependent increases in CAN1 mutation rate.

GCR analysis:

In addition to mutagenesis, we also examined the rate of gross chromosomal rearrangement (GCR) in est2 mutants grown serially over time. GCR is defined as an event that leads to the simultaneous loss of CAN1 and a URA3 marker inserted at the HXT13 locus that lies between CAN1 and the telomere on the left end of chromosome V (CHEN and KOLODNER 1999). The GCR rate was only 2-fold over that in wild type before senescence, 383-fold over that in wild type during senescence, and decreased to wild-type levels upon recovery from senescence (Table 3), all consistent with previously published reports (HACKETT et al. 2001; MYUNG et al. 2001; PENNANEACH and KOLODNER 2004).

View this table: In this window In a new window   TABLE 3 Control of senescence-dependent gross chromosomal rearrangement

 
Consistent with CAN1 mutagenesis REV1 and REV7 were found to be important in determining the GCR rate during replicative senescence. GCR rates in est2 rev1 and est2 rev7 mutants were only 2- to 4-fold higher during senescence than in rev1 and rev7 mutants (Table 3) that did not undergo senescence (Figure 3A). These results are consistent with Pol and Rev1 being required for senescence-dependent GCR. Strikingly, Rad30 was found to be required to suppress GCR as the rate in est2 rad30 cells was 37-fold over that in wild type before senescence, 2673-fold over that in wild type during senescence, and at wild-type levels upon recovery (Table 3). Interestingly, this increase is nearly completely dependent on Rev7 as the GCR rate in senescent est2 rad30 rev7 cells was <2-fold greater than that in the est2 rev7 mutant (Table 3). Therefore, like senescence-dependent mutagenesis at the CAN1 locus, senescence-dependent GCR requires Rev1 and Pol .

Our data suggest that Rev1- and Pol -dependent mutations and GCR at telomere proximal loci are an important consequence of telomere dysfunction. The involvement of Pol , Pol , and Rev1 suggests that the postreplication repair machinery (MINESINGER and JINKS-ROBERTSON 2005) may be responding to the failure of bidirectional DNA replication in the region (PLOSKY and WOODGATE 2004; LEHMANN 2005). Strathern and colleagues have reported the involvement of Pol in generating mutations associated with the recombinational repair of an enzyme-catalyzed double-strand break (RATTRAY et al. 2002), suggesting that a similar process may be involved in generating mutations at sequences lying near the uncapped telomeres of senescent cells (DUBOIS et al. 2002). However, such a mechanism is unlikely to involve extensive exonucleolytic degradation from the telomere as loss of Exo1, a factor involved in the degradation of uncapped telomeres (HACKETT and GREIDER 2003), does not reduce senescence-dependent mutation (Figure 1B) or GCR (Table 3). Further, this mechanism would likely require homologous recombination between sister chromatids, as the CAN1 gene is a unique sequence in the genome. Perhaps similar forces underlie some of the increased genome instability and cancer in somatic cells of elderly people (LENGAUER et al. 1998; JOHNSON et al. 1999).

We thank D. Gottschling, J. McDonald, R. Woodgate, and V. Lundblad for supplying strains and plasmids. We also thank J. Termini, T. O'Connor, R.-J. Lin, and the members of the Bailis lab for productive discussions. Our thanks also go to several anonymous reviewers for their valuable comments. This work was supported by funds from the Beckman Research Institute of the City of Hope, the National Institutes of Health (GM057484 to A.B.), and the Department of Defense Breast Cancer Research Program (W81XWH0410407 to D.M.).

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Communicating editor: E. ALANI

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