Peroxiredoxins are a family of antioxidant enzymes critically involved in cellular defense and signaling. Particularly, yeast peroxiredoxin Tsa1p is thought to play a role in the maintenance of genome integrity, but the underlying mechanism is not understood. In this study, we took a genetic approach to investigate the cause of genome instability in tsa1Δ cells. Strong genetic interactions of TSA1 with DNA damage checkpoint components DUN1, SML1, and CRT1 were found when mutant cells were analyzed for either sensitivity to DNA damage or rate of spontaneous base substitutions. An elevation in intracellular dNTP production was observed in tsa1Δ cells. This was associated with constitutive activation of the DNA damage checkpoint as indicated by phosphorylation of Rad9/Rad53p, reduced steady-state amount of Sml1p, and induction of RNR and HUG1 genes. In addition, defects in the DNA damage checkpoint did not modulate intracellular level of reactive oxygen species, but suppressed the mutator phenotype of tsa1Δ cells. On the contrary, overexpression of RNR1 exacerbated this phenotype by increasing dNTP levels. Taken together, our findings uncover a new role of TSA1 in preventing the overproduction of dNTPs, which is a root cause of genome instability.
Peroxiredoxins are a family of antioxidant enzymes highly conserved from yeast to human. Loss of peroxiredoxin in mice can lead to severe anemia and malignant tumors, but the underlying cause is not understood. One way to derive new knowledge of peroxiredoxins is through genetic analysis in yeast. We have shown that loss of peroxiredoxins in yeast is associated with an increase in mutation rates. Here, we demonstrate that this elevation of mutation rates in yeast cells lacking a peroxiredoxin is due to increased production of deoxyribonucleoside triphosphates (dNTPs), the building blocks of DNA. Our findings suggest a new model in which compromised antioxidant defense causes accumulation of damaged DNA and activation of the DNA damage checkpoint. For yeast cells to survive DNA damage, dNTP production is increased to facilitate DNA replication, but at the price of high mutation rates. This new model could lead to a better understanding of human diseases including cancer.
Citation: Tang H-MV, Siu K-L, Wong C-M, Jin D-Y (2009) Loss of Yeast Peroxiredoxin Tsa1p Induces Genome Instability through Activation of the DNA Damage Checkpoint and Elevation of dNTP Levels. PLoS Genet 5(10): e1000697. https://doi.org/10.1371/journal.pgen.1000697
Editor: Orna Cohen-Fix, National Institute of Diabetes and Digestive and Kidney Diseases, United States of America
Received: May 1, 2009; Accepted: September 23, 2009; Published: October 23, 2009
Copyright: © 2009 Tang et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work is supported by Hong Kong Research Grants Council (projects HKU7340/03M and HKU7670/07M). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Peroxiredoxins belong to a family of thiol-specific peroxidases widely and abundantly expressed in most living organisms ,. Through one or more redox-sensitive cysteines, peroxiredoxins not only scavenge reactive oxygen species (ROS) including peroxides and peroxynitrite ,, but also function as an ROS sensor to regulate cell signaling –. For many peroxiredoxins, another level of regulation can be achieved through oligomerization ,,. In addition to their roles in peroxide reduction, peroxiredoxins are also known to possess chaperone activity ,.
Loss-of-function studies in mice implicated an essential role of peroxiredoxins in antioxidant defense and tumor suppression . Particularly, peroxiredoxin 1-knockout mice not only suffered from severe anemia due to oxidative stress, but were also susceptible to several types of malignant tumors . Consistent with this, genome-wide screening revealed that yeast peroxiredoxin TSA1 was a strong suppressor of gross chromosomal rearrangements and spontaneous mutations ,. In addition, a mutator phenotype was observed in yeast cells lacking one or more peroxiredoxins. The phenotype could be rescued by yeast peroxiredoxin Tsa1p or mammalian Prx1, but not by their active-site mutants defective for peroxidase activity ,. In further support of a role of TSA1 in the maintenance of genome stability, many genetic interaction partners of TSA1 identified through synthetic genetic array analysis were components of DNA repair machinery or DNA checkpoints ,. For example, TSA1 was found to interact genetically with REV1/REV3 and OGG1, which are critically involved in translesion synthesis (TLS) and the repair of oxidative DNA damage, respectively ,. However, the exact mechanism by which Tsa1p suppresses genome instability remains to be fully understood.
Intracellular dNTP levels are one important determinant of cellular response to DNA damage . For yeast cells to survive DNA damage, increased dNTP production would be allowed to facilitate replication, but with a trade-off of high spontaneous mutation rate . In other words, abnormally high dNTP levels are causally associated with genome instability ,.
We previously demonstrated that yeast Tsa1p is a house-keeping peroxiredoxin which sufficiently suppressed the mutator phenotype . Although both an aberrantly high level of ROS and an imbalance in free radical contents, which is caused by compensational activation of other antioxidants such as Sod1p , could underlie the mutator phenotype of tsa1Δ cells, additional events subsequent to the disruption of TSA1 might also be influential in the induction of genome instability. In this study we asked whether perturbation of dNTP pools might contribute to the mutator phenotype observed in tsa1Δ cells. We then investigated the cause of dNTP pool expansion. Our findings suggested that constitutive activation of the DNA damage checkpoint and consequent overproduction of dNTPs are the root cause of genome instability in tsa1Δ cells.
Deletion of DUN1, SML1, or CRT1 Modulates Sensitivity of tsa1Δ Cells to DNA Damage
Yeast peroxiredoxin TSA1 was found to be a strong suppressor of mutations and gross chromosomal rearrangements –. In addition, further deletion of another gene involved in DNA repair or DNA checkpoints caused synthetic growth defect or lethality in tsa1Δ cells ,. Bearing these findings in mind, here we sought to dissect the interaction of TSA1 with the DNA damage checkpoint and particularly the machinery of dNTP synthesis, in order to understand the role of TSA1 in the maintenance of genome stability.
We first examined the sensitivity of tsa1Δ cells to various DNA damaging agents. tsa1Δ cells were sensitive to hydroxyurea (HU), 4-nitroquinoline 1-oxide (4NQO) and ultraviolet (UV) irradiation (Figure 1A, lanes 1 and 5; Figure 1B; Figure 1C and Figure 1D, lanes 1 and 2). Re-expression of TSA1 in tsa1Δ cells suppressed the sensitivity phenotype (Figure 1C and Figure 1D, lane 3). This suppression required the catalytic cysteine (Cys47) of Tsa1p, but not the C-terminal cysteine (Cys170), pointing to the importance of the antioxidant property of Tsa1p in the protection against DNA damage (Figure 1C, lanes 3–5).
(A) Spot tests. Ten-fold serial dilutions of strains BY4741 (WT), dun1Δ, sml1Δ, crt1Δ, tsa1Δ, tsa1Δ dun1Δ, tsa1Δ sml1Δ, and tsa1Δ crt1Δ were spotted on YPD medium containing indicated doses of H2O2, HU or 4-NQO. Cells were exposed to the indicated dose of UV after plating. Plates were incubated for 4 days at 30°C. Experiments were repeated for six times and similar results were obtained. (B) Survival curves. Logarithmically growing yeast cells, BY4741 (WT), dun1Δ, sml1Δ, tsa1Δ, tsa1Δ dun1Δ, and tsa1Δ sml1Δ, in YPD were treated with indicated doses of 4-NQO for 90 min before plating on YPD agar. For UV treatment, cells were first plated onto YPD agar followed by UV irradiation at the indicated doses. Plates were incubated for 3 days at 30°C, and then counted for survival. The number of colonies from untreated plates was taken as 100%. Experiments were repeated for three times and similar results were obtained. (C) TSA1 catalytic cysteine mutant cannot complement the HU sensitivity in tsa1Δ cells. Tenfold serial dilutions of the indicated strains transformed with pRS415, pTSA1, pTSA1C47S, or pTSA1C170S were spotted on SC-Leu plates containing 2% glucose and the indicated doses of diamide or HU. Note that cells grew more slowly on SC plates than on YPD plates as shown in (A). (D) Influence of DUN1, SML1, or CRT1 deletion on the sensitivity of sod1Δ cells to DNA damage. (E) Complementation of drug sensitivity in tsa1Δ dun1Δ cells by TSA1 or DUN1.
The sensitivity of tsa1Δ cells to DNA damage prompted us to investigate further the genetic interactions between TSA1 and components of the DNA damage checkpoint. In light of the finding that TSA1 genetically interacts with DNA damage checkpoint genes DUN1 and SML1 , we chose these two genes and their effector CRT1 for further analysis. Dun1p is a checkpoint kinase that phosphorylates and regulates ribonucleotide reductase (RNR) inhibitor Sml1p . Dun1p also inhibits Crt1p, a transcriptional corepressor of RNR, through phosphorylation ,. Deletion of DUN1, SML1 or CRT1 in tsa1Δ cells exerted a significant impact on their sensitivity to HU, 4NQO and UV irradiation. Loss of DUN1 further sensitized tsa1Δ cells to H2O2, HU, 4NQO and UV (Figure 1A, lanes 1, 2, 5 and 6). In support of the specificity of effect, this sensitization was reversed upon expression of TSA1 or DUN1 in tsa1Δ dun1Δ cells (Figure 1D, lanes 6–8). Conversely, loss of SML1 or CRT1 rescued the sensitivity phenotype of tsa1Δ cells to 4NQO and UV (Figure 1A, lanes 5, 7 and 8). It is noteworthy that such reversion of sensitivity was not observed in cells treated with H2O2 (Figure 1A, lanes 5, 7 and 8), suggesting that the effect might be specific to DNA damaging agents and was not caused directly by ROS. These observations supported the notion that TSA1 interacts specifically with the DNA damage checkpoint in a manner that is not mediated directly through ROS.
Although the sensitivity pattern of the different mutant strains in the spot assay was highly reproducible, a more quantitative comparison of these strains is desired. Hence, survival curves of strains in the presence of 4NQO and UV were also obtained (Figure 1B). Dose-dependent killing of the strains by 4NQO and UV was observed. At all doses tested, the degrees of sensitivity of different strains to either 4NQO or UV were in the same order as shown in the spot assay. In particular, the survival curves indicated a further enhancement of the sensitivity phenotype in tsa1Δ dun1Δ versus tsa1Δ cells and a suppression of sensitivity in tsa1Δ sml1Δ cells (Figure 1B). Collectively, our results demonstrated that the survival of tsa1Δ cells under DNA damage was decreased upon deletion of DUN1, but enhanced when either SML1 or CRT1 was genetically disrupted.
We also compared the phenotypes of tsa1Δ cells and cells lacking Sod1p, another key antioxidant enzyme . In contrast to the genetic interactions observed in tsa1Δ cells, deletion of DUN1, SML1 or CRT1 in sod1Δ cells enhanced its sensitivity to HU and 4NQO (Figure 1E, lanes 6, 7 and 8). Thus, TSA1 and SOD1 interact with DUN1, SML1 and CRT1 through different mechanisms.
Deletion of DUN1, SML1, or CRT1 Modulates Mutator Phenotype of tsa1Δ Cells
We next investigated whether compromising DNA damage checkpoint genes in tsa1Δ cells might also alter their mutator phenotype. In agreement with previous reports –, tsa1Δ cells exhibited high rates of spontaneous mutations in both canavanine-resistant (CANR) and 5FC-resistant (5FCR) assays (Figure 2A, Figure 2B and Figure 2D, groups 1 and 5). On the other hand, deletion of DUN1 did not significantly affect spontaneous mutation rates (Figure 2A, Figure 2B and Figure 2D, groups 1 and 2), whereas loss of SML1 or CRT1 caused a mild increase in CANR mutation rates in WT cells (Figure 2A and Figure 2D, groups 1, 3 and 4). However, the disruption of DUN1, SML1 and CRT1 in tsa1Δ cells modulated the mutator phenotype in opposite directions (Figure 2A and Figure 2B, columns 5–8). Whereas reduction of spontaneous mutation rates was observed in tsa1Δ dun1Δ cells (Figure 2A and Figure 2B, columns 5 and 6), deletion of SML1 or CRT1 in tsa1Δ cells significantly enhanced the mutator phenotype (Figure 2A and Figure 2B, columns 1, 7 and 8). Complementation of the reduction of mutation rate in tsa1Δ dun1Δ cells by re-introduction of DUN1 or TSA1 further verified the specificity of effect (Figure 2C, columns 6 and 8). Thus, the mutation rates of tsa1Δ cells correlated with the activity of the DNA damage checkpoint.
The number of CANR (A) or 5FCR (B) colonies on synthetic complete solid medium either lacking arginine but containing CAN (60 mg/L) or supplemented with 5FC (100 mg/L) was normalized with the total number of viable cells grown on the same solid medium without CAN or 5FC. The relative mutation rate of BY4741 cells (WT) was taken as 1.00. Results represent the average from triplicate analysis of ten independent cultures. (C) Complementation of mutator phenotype in tsa1Δ dun1Δ cells by TSA1 or DUN1. The rates of spontaneous CANR mutation were calculated as in (A). (D) Influence of ROS on mutation rates. Cells of the indicated genotypes logarithmically growing in YPD were subjected to treatment of H2O2 (0.6 mM, 15 min) before plating on synthetic complete solid medium lacking arginine and supplemented with CAN (60 mg/L). The rates of spontaneous CANR mutation were calculated as in (A).
In all individual deletion mutants, tsa1Δ cells displayed the highest mutation rate (Figure 2). We postulated that this might be attributed either directly or indirectly to the elevation of intracellular ROS levels in these cells . If that is the case, challenging the other DNA damage checkpoint mutants with ROS might have an impact on the mutator phenotype. To test this idea, we treated the cells with low-dose H2O2 and assessed the impact on CANR mutation rates. Interestingly, mutation rates increased in WT and dun1Δ cells to comparable levels (Figure 2D, groups 1 and 2). In contrast, a further increase in mutability was observed when SML1 or CRT1 was comprised (Figure 2D, groups 1, 3 and 4). Although the mutation rates of sml1Δ and crt1Δ cells in the presence of H2O2 were still not as high as that of tsa1Δ cells in the absence of H2O2 (Figure 2D, groups 3–5), our results did suggest that ROS could differentially modulate the mutator phenotype of different mutants.
Loss of TSA1 Elevates Cellular dNTP Production
We next investigated the mechanism that underlies the correlation of DNA damage checkpoint activity in tsa1Δ cells with drug sensitivity and mutator phenotype (Figure 1 and Figure 2). DUN1, SML1 and CRT1 are regulators of RNR, the rate-limiting enzyme in dNTP synthesis –. Considered together with the model that elevated dNTP levels are required for surviving DNA damage in yeast at the price of increasing mutation rates , we asked whether the mutator phenotype of tsa1Δ cells would be due to alteration in cellular dNTP production. Thus, we measured dNTP levels in our mutants. Surprisingly, tsa1Δ cells produced significantly more dNTPs than wild type (WT) cells (Figure 3A, groups 1 and 5). The magnitude of dNTP overproduction in tsa1Δ cells was comparable to that in sml1Δ cells (Figure 3A, groups 3 and 5), in which the removal of Sml1p activates RNR leading to the rise in dNTP levels .
(A) Comparison of dNTP levels. Relative dNTP levels were determined in the indicated strains of cells growing logarithmically. (B) Suppression of dNTP pool phenotype by TSA1. Assays were done with WT and tsa1Δ strains transformed with pRS415, pTSA1, or pTSA1C47S. (C) ROS detection. Cells of the indicated strains logarithmically growing in YPD were subjected to treatment with DCF (10 µM, 45 min). Crude extracts of cells were subjected to DCF fluorescence measurement on an F-4500 spectrofluorimeter (Hitachi). The excitation and emission wavelengths were 488 and 520 nm, respectively. The reading of DCF fluorescence was normalized to protein concentration. The fluorescent intensity of BY4741 cells (WT) was taken as 1. Results represent the average from three independent experiments.
To shed further light on the roles of dNTP production in the generation of mutator phenotype, we compared the dNTP levels in other mutant cells. As expected, dun1Δ cells produced less dNTPs than WT cells (Figure 3A, groups 1 and 2), since Dun1p is required for phosphorylation and subsequent removal of the RNR inhibitor Sml1p . Loss of CRT1 was also found to increase cellular dNTP production (Figure 3A, groups 1 and 4), as Crt1p is a transcriptional corepressor of RNRs . However, loss of DUN1 reduced cellular dNTP production in tsa1Δ cells (Figure 3A, groups 5 and 6), whereas deletion of SML1 or CRT1 in tsa1Δ cells further increased dNTP levels (Figure 3A, groups 5, 7 and 8). Noteworthily, increased production of dNTPs in tsa1Δ cells could be fully complemented by Tsa1p, but not by its catalytic cysteine mutant C47S (Figure 3B). Thus, the antioxidant property of Tsa1p was likely required for preventing overproduction of dNTPs.
We then asked whether the reduction of dNTP pools in tsa1Δ dun1Δ cells would be associated with a further drop in intracellular ROS levels in the absence of DUN1. Interestingly, tsa1Δ dun1Δ cells exhibited a higher level of intracellular ROS over WT, tsa1Δ or dun1Δ cells (Figure 3C, columns 1, 2, 5 and 6), suggesting that the mutator phenotype in tsa1Δ and tsa1Δ dun1Δ cells correlates directly with dNTP production, but not generation of ROS. On the other hand, loss of SML1 or CRT1 did not alter the ROS levels in either WT or tsa1Δ cells (Figure 3C, columns 1, 3, 4, 5, 7 and 8). Thus, in addition to the accumulation of ROS, elevation of dNTP production might also contribute to genome instability in tsa1Δ cells.
While deletion of DUN1, SML1 or CRT1 has an impact on dNTP production, they are multifunctional proteins that might also affect other biological processes –. To address this concern, we modulated the production of dNTP more directly by overexpressing RNR1 gene in tsa1Δ and sod1Δ cells. This overexpression has previously been shown to elevate intracellular dNTP levels substantially ,. Indeed, when we induced the expression of Rnr1-3MYCp in WT cells (Figure 4A), the spontaneous mutation rate was increased (Figure 4B, columns 1 and 2). Furthermore, overexpression of RNR1 also exacerbated the mutator phenotype in tsa1Δ and sod1Δ cells (Figure 4B, columns 3–6). This lent additional support to the importance of dNTP overproduction in the induction of genome instability.
BY4741 (WT), sod1Δ, and tsa1Δ cells carrying plasmid pGal-RNR1 were grown in SC-Ura medium supplemented with raffinose (uninduced) or galactose (induced) to mid-log phase. (A) Galactose-induced expression of Rnr1-3MYCp. Western blotting was performed with mouse anti-MYC (Roche) and mouse anti-Pgk1p (Invitrogen) antibodies. (B) Mutation rates. Experiments were carried out as in Figure 2.
If the mutability of tsa1Δ cells is indeed caused by dNTP overproduction, the mutations generated would primarily be base substitutions rather than large deletion and gross chromosomal rearrangements . With this in mind, we examined the types of mutations arisen in tsa1Δ and other mutants (Table 1). We noted that the majority of mutations found in tsa1Δ cells were base substitutions (83.3%) and frameshifts (13.3%). Large deletions were very rare in tsa1Δ cells (3.3%) when compared with WT (13.3%) cells. In addition, all of the mutations found in tsa1Δ sml1Δ cells with high dNTP levels (Figure 3) were base substitutions (90%) and frameshifts (10%), whereas more deletions (13.3%) were detected in tsa1Δ dun1Δ cells (Table 1) with low dNTP concentrations (Figure 3). In keeping with previous findings , relatively more deletions (10%) were also observed in sod1Δ cells (Table 1). Generally, base substitutions were more prevalent in the strain when dNTP levels were high (Figure 3), whereas the incidences of deletions correlated negatively with dNTP concentrations. Therefore, the mutation spectra of tsa1Δ and other strains are consistent with the notion that elevation of dNTP levels is the underlying cause of genome instability in the absence of TSA1.
Loss of TSA1 Activates DNA Damage Checkpoint Leading to Elevation in dNTP Production
Above we demonstrated the elevation of dNTP levels in tsa1Δ cells (Figure 3). In addition, our results also indicated the genetic interaction of TSA1 with DNA checkpoint genes (Figure 1). This led us to further investigate whether elevated production of dNTPs in the absence of TSA1 might be explained by the activation of the DNA damage checkpoint. As a first step, we assessed checkpoint activation by examining the steady-state levels of Rad53p, the yeast ortholog of human CHK2 kinase whose phosphorylation and activation are pivotally involved in the control of checkpoint response to DNA damage ,. Particularly, Rad53p is a master regulator of Dun1p, Sml1p and Crt1p .
In this analysis we included the sod1Δ control strain, in which the effectors of the Mec1p-dependent DNA damage checkpoint were previously shown to be downregulated . Phosphorylated Rad53p species were more evident in tsa1Δ cells compared to WT and sod1Δ cells (Figure 5A, lanes 1, 3 and 5; Figure 5B, lanes 1 and 2; Figure 6A, lanes 1 and 2; and Figure 7A, lanes 1 and 3). This difference became more pronounced in the presence of H2O2 (Figure 5A, lanes 2, 4 and 6; Figure 5B, lanes 5 and 6; Figure 6A, lanes 3 and 4; and Figure 7A, lanes 5 and 7).
(A) Western blot analysis of Rad53p. Cells of WT, sod1Δ, and tsa1Δ strains logarithmically growing in YPD were subjected to treatment with H2O2 (0.8 mM, 30 min). Western blotting was performed with goat anti-Rad53p (Santa-Cruz) and mouse anti-Pgk1p antibodies. Percentages of phosphorylated Rad53p were determined by densitometry and indicated at the bottom of the panel. (B) TSA1 complementation assay. WT and tsa1Δ cells were transformed with pRS415, pTSA1, or pTSA1C47S plasmid. Western blotting was carried out with goat anti-Rad53p, mouse anti-HA (Santa-Cruz), rabbit anti-histone H2A phosphorylated at S129 (γH2A; Abcam), and mouse anti-Pgk1p antibodies. Relative amounts of Sml1-3HAp or γH2A normalized to Pgk1p were determined by densitometry and indicated at the bottom of the panels. (C) Checkpoint activation in different strains.
(A) Western blot analysis of Rad53p. Cells of the indicated strains in W303 background logarithmically growing in YPD were subjected to treatment with H2O2 (0.8 mM, 30 min). (B) Western blot analysis of Rad9p in tsa1Δ and TSA1-complemented strains. A longer exposure (long exp.) of the Rad9p blot was also shown. (C) Semi-quantitative RT–PCR analysis of RNR transcripts. Logarithmically growing cells of the indicated strains in YPD were subjected to treatment with HU (200 mM) at the indicated time points. Total RNA was extracted and 3 µg of RNA was used for cDNA synthesis. PCR was performed to assess the levels of RNR1/2/3/4, HUG1, and ACT1 transcripts. The expected sizes of the PCR product for RNR1, RNR2, RNR3, RNR4, ACT1, and HUG1 are 219, 390, 199, 455, 520, and 190 bp, respectively. Relative levels of RNA determined by densitometry and normalized to the amount of ACT1 transcript were indicated at the bottom of the panels.
(A) Western blot analysis of Rad53p and Sml1-3HAp. Cells of the indicated strains in W303 background logarithmically growing in YPD were subjected to treatment with H2O2 (0.8 mM, 30 min). Western blotting was performed as in Figure 5B. (B) Spot assay. Ten-fold serial dilutions of the indicated strains were spotted on YPD medium containing the indicated doses of H2O2 or HU. Some cells were exposed to UV after plating. (C) RAD53 mutation suppresses mutator phenotype in tsa1Δ cells. Mutation rates were calculated as in Figure 2B. (D) ROS detection. DCF fluorescence was measured as in Figure 3C.
The steady-state levels of phosphorylated Rad53p in DNA damage checkpoint mutants were also compared. Whereas deletion of DUN1 triggered phosphorylation of Rad53p (Figure 5C, lanes 1 and 2), an observable increase in phosphorylated Rad53p species was not found in sml1Δ or crt1Δ cells (Figure 5C, lanes 1, 3 and 4). Notably, loss of DUN1 in tsa1Δ cells further enhanced the activation of Rad53p (Figure 5C, lanes 5 and 6), whereas tsa1Δ sml1Δ and tsa1Δ crt1Δ cells had similar levels of phosphorylated Rad53p compared to tsa1Δ cells (Figure 5C, lanes 5, 7 and 8).
In addition to Rad53p, we also checked for the status of Rad9p, a more upstream transducer in the DNA damage checkpoint pathway . Rad9-13MYCp was found to be activated in tsa1Δ cells (Figure 6B, lanes 1, 2, 4 and 5) and this activation could not be reversed by the C47S mutant of Tsa1p (Figure 6B, lanes 2, 3, 5 and 6). As a marker for DNA double-strand breaks (DSBs) , the level of γH2A was also found to be elevated in tsa1Δ cells as compared to WT (Figure 5B, lanes 1 and 2; Figure 6A, lanes 1 and 2; and Figure 6B, lanes 1 and 2). This agrees with a recent report that tsa1Δ cells displayed an increased number of Rad52-YFP foci indicative of DNA damage . The levels of γH2A in other DNA damage checkpoint mutant cells were also examined. Among dun1Δ, sml1Δ and crt1Δ cells, an elevation in γH2A level was only found in dun1Δ cells (Figure 5C, lane 2). In addition, disruption of DUN1, SML1 or CRT1 in tsa1Δ cells did not affect γH2A levels significantly (Figure 5C, lanes 5–8). Noteworthily, although phosphorylated Rad53p and γH2A were abundant in dun1Δ and tsa1Δ dun1Δ cells (Figure 5C, lanes 2 and 6), their mutation rates remained low (Figure 2A and Figure 2B, columns 2 and 6) plausibly due to the low levels of dNTPs (Figure 3A, groups 2 and 6). In other words, elevation of dNTP levels might be the direct cause of genome instability.
Consistent with the activation of Rad53p and Rad9p, the levels of Rad53p target Sml1-3HAp were diminished in tsa1Δ cells in the presence (Figure 5B, lanes 5 and 6) and absence of H2O2 (Figure 5B, lanes 1 and 2). Importantly, all of the above changes in tsa1Δ cells could be fully complemented by TSA1 (Figure 5B, lanes 2 and 3), but not by its C47S mutant (Figure 5B, lanes 2 and 4). On the other hand, in agreement with previous findings , we did not observe a significant change of Sml1p level in sod1Δ cells (data not shown).
RNR is an important downstream effector of the DNA damage checkpoint which mediates the production of dNTPs ,. Since the expression of RNR genes is transcriptionally activated in response to DNA damage , we used semi-quantitative RT-PCR to determine the relative levels of RNR1/2/3/4 transcripts in the presence and absence of HU. In this analysis we included an additional control termed HUG1, a target of Mec1p induced highly by DNA damage . As shown in Figure 6C, RNR transcripts were induced to higher levels in tsa1Δ cells than in WT and sod1Δ cells. The induction of RNR1 and RNR3 was greatest in both untreated and HU-treated tsa1Δ cells. The level of HUG1 transcript was also elevated in tsa1Δ cells and this was more pronounced in the presence of HU (Figure 6C). In sharp contrast, sod1Δ cells treated with HU showed a lower magnitude of induction of RNR1, RNR3 and HUG1 mRNAs (Figure 6C). These results obtained from sod1Δ cells were generally consistent with previous findings . Thus, the pattern of RNR induction in tsa1Δ cells was not ascribed to a general effect caused by the lack of any antioxidant enzyme, but was highly specific. Collectively, our results suggested that loss of TSA1 induces the activation of the DNA damage checkpoint leading to the induction of RNR and consequent overproduction of dNTPs.
Mutation of Rad53p Suppresses Genome Instability in tsa1Δ Cells
If activation of the DNA damage checkpoint in tsa1Δ cells is really important to the generation of genome instability caused by dNTP overproduction, genetic disruption of the checkpoint would be able to reverse the mutator phenotype of tsa1Δ cells.
To test this hypothesis, we employed a RAD53 mutant termed rad53AA, in which both T354 and T358 in the activation loop of Rad53p had been replaced by alanine, thereby abrogating the autophosphorylation activity in response to DNA damage . This defective rad53AA allele similar to rad53-11 is thought to be associated with reduced dNTP production due to high abundance of Sml1p ,. Thus, we set out to characterize the phenotypes of tsa1Δ cells carrying the rad53AA allele.
As documented previously , TSA1 rad53AA cells exhibited lower levels of phosphorylated Rad53p and higher abundance of Sml1-3HAp than TSA1 RAD53 cells (Figure 7A, lanes 1, 2, 5 and 6). In response to H2O2, γH2A was induced to higher levels in all mutant cells (Figure 7A, lanes 5–8 compared to lanes 1–4). Notably, both tsa1Δ RAD53 and tsa1Δ rad53AA cells showed similar basal levels of γH2A (Figure 7A, lanes 3 and 4). Although stronger Rad53p activation was observed in tsa1Δ rad53AA cells, a more pronounced Sml1-3HAp protein band was seen (Figure 7A, lane 4 compared to lane 3), suggestive of a defective DNA damage checkpoint.
We next characterized the sensitivity of these mutants towards H2O2, UV and HU. TSA1 rad53AA cells were sensitive to HU (Figure 7B, lanes 1 and 2) as previously described ; while tsa1Δ RAD53 cells were sensitive to H2O2, UV and HU (Figure 7B, lanes 1 and 3) similar to tsa1Δ cells in BY4741 background (Figure 1, lanes 1 and 5). Resembling tsa1Δ dun1Δ cells in BY4741, tsa1Δ rad53AA cells in W303 background displayed further sensitivity to H2O2 and UV when compared to tsa1Δ RAD53 cells (Figure 7B, lanes 3 and 4).
We then looked at the effect of a defective DNA damage checkpoint on genome instability in tsa1Δ cells. Intriguingly, tsa1Δ rad53AA cells displayed a significantly reduced (∼50%) rate of spontaneous 5FCR mutations over tsa1Δ RAD53 cells (Figure 7C). On the other hand, both tsa1Δ rad53AA and tsa1Δ RAD53 cells had high levels of intracellular ROS over WT cells as measured by DCF fluorescence (Figure 7D). These observations suggested that rad53AA mutation can suppress the mutator phenotype in tsa1Δ cells without affecting cellular redox environment. This generally agrees with the phenotypes of tsa1Δ dun1Δ cells (Figure 1 and Figure 2), lending further support to the concept that intracellular dNTP levels are an important determinant in the induction of genome instability in tsa1Δ cells.
Here, we provided the first evidence that loss of yeast peroxiredoxin TSA1 causes genome instability through constitutive activation of the DNA damage checkpoint leading to overproduction of intracellular dNTPs. There are two salient points in our work. First, we demonstrated the elevation of dNTP levels in tsa1Δ cells and its direct correlation with the mutator phenotype (Figure 2, Figure 3, Figure 4). Second, we demonstrated the activation of the DNA damage checkpoint in tsa1Δ cells in relation to elevated production of dNTPs (Figure 1, Figure 5, Figure 6, Figure 7). Our findings suggested a new model for the role of peroxiredoxins in the maintenance of genome integrity, which has implications in the understanding of human diseases including cancer.
In agreement with our findings on the accumulation of γH2A and activation of the DNA damage checkpoint in tsa1Δ cells, several lines of evidence in the literature supported the role of Tsa1p and other peroxiredoxins in the protection of cells against DNA damage. First, tsa1Δ cells produce significantly more ROS , which cause DNA and protein damage ,,. Second, loss of TSA1 results in increased formation of Rad52-YFP foci, an indicator of DNA DSBs . Third, tsa1Δ cells are highly sensitive to the functional state of DNA repair and checkpoints . In particular, tsa1Δ is synthetically lethal with rad51Δ mutation, indicating that the viability of rad51Δ cells deficient in recombination repair requires TSA1 function . Finally, human peroxiredoxins have been implicated in cellular defense against oxidative DNA lesions . In this context, the activation of the DNA damage checkpoint in tsa1Δ cells demonstrated in our study highlights the pivotal roles of the checkpoint in cell survival and provides an explanation for the synthetic lethality seen in various double deletion mutants involving TSA1 and another DNA repair or checkpoint gene .
Deletion of TSA1 in yeast cells has previously been shown to result in both a mutator phenotype and an increase in gross chromosomal rearrangements ,,. Although the causes and origin of gross chromosomal rearrangements remain poorly understood, oxygen metabolism and ROS production are implicated in the prevalence of these rearrangements in tsa1Δ cells . Noteworthily, base substitution, but not chromosomal rearrangement, was the predominant type of mutation found in our analysis of mutation rates (Table 1). Thus, the major type of genome instability analyzed in our study is an increased rate of point mutations, but not gross chromosomal rearrangements involving more complex alterations such as translocations, large deletions and amplifications.
Our findings point to a role of dNTP levels in determining the mutation rate of tsa1Δ cells. Strong genetic interactions between TSA1 and four RNR regulators DUN1, SML1, CRT1 and RAD53 were observed in the context of sensitivity to DNA damage (Figure 1 and Figure 7), spontaneous mutability (Figure 2 and Figure 7) and dNTP production (Figure 3). Although the catalytic cysteine of Tsa1p is required for the suppression of mutator phenotype, the mutability of tsa1Δ cells correlated directly with dNTP concentrations (Figure 2, Figure 3, Figure 4), but not with high ROS levels (Figure 3 and Figure 7). One plausible explanation is that the loss of TSA1 might cause accumulation of both ROS  and DNA damage (Figure 5). This activates the DNA damage checkpoint through Rad53p, Rad9p and Sml1p (Figure 5, Figure 6, Figure 7) leading to transcriptional activation of RNR genes (Figure 6) and elevated production of dNTPs (Figure 3). Once at high dNTP levels, replicative and TLS polymerases by-pass DNA lesions more efficiently to promote survival, but only at the price of increasing mutation rates ,. This model implicates dNTP pool expansion as the major culprit in the induction of genome instability in tsa1Δ cells. Indeed, reducing dNTP levels without affecting ROS production was sufficient to reverse the mutator phenotype of tsa1Δ cells (Figure 3). In particular, tsa1Δ dun1Δ cells have high levels of ROS (Figure 3C), phosphorylated Rad53p (Figure 5C) and γH2A (Figure 5C). However, these cells showed a low mutation rate (Figure 2C) because the dNTP levels were also low (Figure 3A). On the contrary, increasing dNTP levels by overexpressing RNR1 aggravated the mutator phenotype (Figure 4). Furthermore, point mutations but not deletions were predominantly found in tsa1Δ cells (Table 1), implicating a role for dNTP overproduction in compromising genome stability. In further support of this model, compromise of TLS polymerases also suppressed CANR mutations in tsa1Δ cells .
We found that the levels of dNTPs in tsa1Δ cells were as high as those in sml1Δ cells (Figure 3A). This finding revealed an unexpected role of TSA1 in the maintenance of dNTP pools in eukaryotic cells. We further observed transcriptional activation of RNR genes in tsa1Δ cells (Figure 6), which could be mediated through the activation of Rad53p checkpoint. Although this might provide an explanation for the overproduction of dNTPs, exactly how Tsa1p is mechanistically involved in regulating RNR expression remains to be further investigated.
Consistent with previous findings , elevation of intracellular dNTPs over a particular threshold level by overexpressing Rnr1p can sufficiently induce a mutator phenotype (Figure 4). Plausibly, the dNTP levels in tsa1Δ sml1Δ and tsa1Δ crt1Δ cells might have reached the threshold level causing a dramatically increased mutation rate (Figure 2 and Figure 3). When the elevation of dNTP levels have not reached the threshold as in the case of sml1Δ, crt1Δ and tsa1Δ cells, accumulation of intracellular ROS might serve to trigger or aggravate the mutator phenotype. In tsa1Δ cells, ROS levels were constantly high (Figure 3C) causing severe DNA damage (Figure 5). In contrast, ROS levels were low (Figure 3C) and DNA damage was not detected (Figure 5C) in sml1Δ or crt1Δ cells. This might explain the higher mutation rate in tsa1Δ cells versus sml1Δ or crt1Δ cells (Figure 2). Further exacerbation of the mutator phenotype of sml1Δ and crt1Δ cells by ROS such as H2O2 (Figure 2D) lent some credence to this model.
We demonstrated the requirement of the catalytic cysteine for the ability of Tsa1p to modulate dNTP production (Figure 3). Through irreversible hyperoxidation, this residue can act as a redox sensor, which triggers the switch of peroxiredoxin from peroxidase to chaperone activity under stress ,. In this connection, it would be of great interest to understand whether and how the chaperone activity of Tsa1p might be involved in the regulation of dNTP production.
Activation of Rad53p by upstream kinase Mec1p requires adaptor proteins Rad9p and Mrc1p ,. We noted that Rad53p phosphorylation was dramatically increased in tsa1Δ versus WT cells (Figure 6A and Figure 7A). In contrast, the increase in Rad9p phosphorylation in the absence of TSA1 was less pronounced (Figure 6B). Although additional experiments are required to investigate the cause of this difference between Rad53p and Rad9p, one possibility is that the deletion of TSA1 might exert a stronger effect on Mrc1p activity.
Hypermutability or genome instability is a hallmark of cancer . Mammalian Prx1 is a candidate tumor suppressor gene ,. Because peroxiredoxins are highly evolutionarily conserved proteins, an understanding of the mechanism by which yeast Tsa1p protects cell from genome instability might derive novel insight into the tumor suppressive role of Prx1 in mammalian cells. Our work demonstrates the importance of high dNTP levels in the mutability of tsa1Δ cells. Further analysis of dNTP concentrations of Prx1-null mouse cells will reveal whether increased production of dNTPs might be a general mechanism for the generation of genome instability in higher eukaryotes.
Materials and Methods
Strains and Plasmids
S. cerevisiae strains BY4741  and W303-1a, and their isogenic strains (Table 2) were used. All knockout mutants were constructed by one-step gene deletion method . Primers were listed in Table 3. Expression vector for DUN1 was derived from pRS415. Expression plasmids for TSA1 and its mutants have been described .
Plasmid pGal-RNR1 kindly provided by Dr. Stephen Elledge has also been described previously .
Measurement of Mutation Rates, dNTP, and ROS Levels
Rates of spontaneous forward mutations to confer CANR or 5FCR were measured as described ,. Spectra of CANR mutations were determined by DNA sequencing. Ten independent cultures were analyzed in each experiment. Cell extracts were prepared and dNTP levels were measured with Klenow enzyme and [3H] labeled dATP or dTTP (PerkinElmer) as described . Standard curves were used to estimate the cellular dNTP levels. Three independent cultures were analyzed in each experiment. Intracellular ROS levels were measured by fluorimetry using DCF (Molecular Probes) as described ,.
Total RNA was extracted by phenol/freeze RNA preparation method as described . For RT-PCR, 3 µg of total RNA was used for cDNA synthesis. Semi-quantitative PCR was performed and optimized to ensure that the amplification was in the linear range. PCR primers were listed in Table 3.
Western blot analysis was performed essentially as described . Yeast cells were harvested by centrifugation, followed by trichloroacetic acid extraction with the help of glass beads.
We thank Drs. Stephen Elledge, Mark Longtine, and Achille Pellicioli for reagents; Dr. Christopher Mathews for advice in dNTP measurement; and members of Jin laboratory for critical reading of manuscript.
Conceived and designed the experiments: HMVT KLS CMW DYJ. Performed the experiments: HMVT KLS CMW. Analyzed the data: HMVT CMW DYJ. Wrote the paper: HMVT CMW DYJ.
- 1. Rhee SG, Chae HZ, Kim K (2005) Peroxiredoxins: a historical overview and speculative preview of novel mechanisms and emerging concepts in cell signaling. Free Radic Biol Med 38: 1543–1552.SG RheeHZ ChaeK. Kim2005Peroxiredoxins: a historical overview and speculative preview of novel mechanisms and emerging concepts in cell signaling.Free Radic Biol Med3815431552
- 2. Fourquet S, Huang ME, D'Autreaux B, Toledano MB (2008) The dual functions of thiol-based peroxidases in H2O2 scavenging and signaling. Antioxid Redox Signal 10: 1565–1576.S. FourquetME HuangB. D'AutreauxMB Toledano2008The dual functions of thiol-based peroxidases in H2O2 scavenging and signaling.Antioxid Redox Signal1015651576
- 3. Bryk R, Griffin P, Nathan C (2000) Peroxynitrite reductase activity of bacterial peroxiredoxins. Nature 407: 211–215.R. BrykP. GriffinC. Nathan2000Peroxynitrite reductase activity of bacterial peroxiredoxins.Nature407211215
- 4. Wong CM, Zhou Y, Ng RWM, Kung HF, Jin DY (2002) Cooperation of yeast peroxiredoxins Tsa1p and Tsa2p in the cellular defense against oxidative and nitrosative stress. J Biol Chem 277: 5385–5394.CM WongY. ZhouRWM NgHF KungDY Jin2002Cooperation of yeast peroxiredoxins Tsa1p and Tsa2p in the cellular defense against oxidative and nitrosative stress.J Biol Chem27753855394
- 5. Jin DY, Chae HZ, Rhee SG, Jeang KT (1997) Regulatory role for a novel human thioredoxin peroxidase in NF-κB activation. J Biol Chem 272: 30952–30961.DY JinHZ ChaeSG RheeKT Jeang1997Regulatory role for a novel human thioredoxin peroxidase in NF-κB activation.J Biol Chem2723095230961
- 6. Delaunay A, Pflieger D, Barrault MB, Vinh J, Toledano MB (2002) A thiol peroxidase is an H2O2 receptor and redox-transducer in gene activation. Cell 111: 471–481.A. DelaunayD. PfliegerMB BarraultJ. VinhMB Toledano2002A thiol peroxidase is an H2O2 receptor and redox-transducer in gene activation.Cell111471481
- 7. Biteau B, Labarre J, Toledano MB (2003) ATP-dependent reduction of cysteine-sulphinic acid by S. cerevisiae sulphiredoxin. Nature 425: 980–984.B. BiteauJ. LabarreMB Toledano2003ATP-dependent reduction of cysteine-sulphinic acid by S. cerevisiae sulphiredoxin.Nature425980984
- 8. Woo HA, Chae HZ, Hwang SC, Yang KS, Kang SW, et al. (2003) Reversing the inactivation of peroxiredoxins caused by cysteine sulfinic acid formation. Science 300: 653–656.HA WooHZ ChaeSC HwangKS YangSW Kang2003Reversing the inactivation of peroxiredoxins caused by cysteine sulfinic acid formation.Science300653656
- 9. Wood ZA, Poole LB, Karplus PA (2003) Peroxiredoxin evolution and the regulation of hydrogen peroxide signaling. Science 300: 650–653.ZA WoodLB PoolePA Karplus2003Peroxiredoxin evolution and the regulation of hydrogen peroxide signaling.Science300650653
- 10. Veal EA, Findlay VJ, Day AM, Bozonet SM, Evans JM, et al. (2004) A 2-Cys peroxiredoxin regulates peroxide-induced oxidation and activation of a stress-activated MAP kinase. Mol Cell 15: 129–139.EA VealVJ FindlayAM DaySM BozonetJM Evans2004A 2-Cys peroxiredoxin regulates peroxide-induced oxidation and activation of a stress-activated MAP kinase.Mol Cell15129139
- 11. Choi MH, Lee IK, Kim GW, Kim BU, Han YH, et al. (2005) Regulation of PDGF signalling and vascular remodelling by peroxiredoxin II. Nature 435: 347–353.MH ChoiIK LeeGW KimBU KimYH Han2005Regulation of PDGF signalling and vascular remodelling by peroxiredoxin II.Nature435347353
- 12. Lim JC, Choi HI, Park YS, Nam HW, Woo HA, et al. (2008) Irreversible oxidation of the active-site cysteine of peroxiredoxin to cysteine sulfonic acid for enhanced molecular chaperone activity. J Biol Chem 283: 28873–28880.JC LimHI ChoiYS ParkHW NamHA Woo2008Irreversible oxidation of the active-site cysteine of peroxiredoxin to cysteine sulfonic acid for enhanced molecular chaperone activity.J Biol Chem2832887328880
- 13. Jang HH, Lee KO, Chi YH, Jung BG, Park SK, et al. (2004) Two enzymes in one; two yeast peroxiredoxins display oxidative stress-dependent switching from a peroxidase to a molecular chaperone function. Cell 117: 625–635.HH JangKO LeeYH ChiBG JungSK Park2004Two enzymes in one; two yeast peroxiredoxins display oxidative stress-dependent switching from a peroxidase to a molecular chaperone function.Cell117625635
- 14. Kümin A, Schäfer M, Epp N, Bugnon P, Born-Berclaz C, et al. (2007) Peroxiredoxin 6 is required for blood vessel integrity in wounded skin. J Cell Biol 179: 747–760.A. KüminM. SchäferN. EppP. BugnonC. Born-Berclaz2007Peroxiredoxin 6 is required for blood vessel integrity in wounded skin.J Cell Biol179747760
- 15. Neumann CA, HKrause DSH, HCarman CVH, Das S, HDubey DPH, et al. (2003) Essential role for the peroxiredoxin Prdx1 in erythrocyte antioxidant defence and tumour suppression. Nature 424: 561–565.CA NeumannDSH HKrauseCVH HCarmanS. DasDPH HDubey2003Essential role for the peroxiredoxin Prdx1 in erythrocyte antioxidant defence and tumour suppression.Nature424561565
- 16. Huang ME, Rio AG, Nicolas A, Kolodner RD (2003) A genomewide screen in Saccharomyces cerevisiae for genes that suppress the accumulation of mutations. Proc Natl Acad Sci U S A 100: 11529–11534.ME HuangAG RioA. NicolasRD Kolodner2003A genomewide screen in Saccharomyces cerevisiae for genes that suppress the accumulation of mutations.Proc Natl Acad Sci U S A1001152911534
- 17. Smith S, Hwang JY, HBanerjee SH, HMajeed AH, HGupta AH, et al. (2004) Mutator genes for suppression of gross chromosomal rearrangements identified by a genome-wide screening in Saccharomyces cerevisiae. Proc Natl Acad Sci U S A 101: 9039–9044.S. SmithJY HwangSH HBanerjeeAH HMajeedAH HGupta2004Mutator genes for suppression of gross chromosomal rearrangements identified by a genome-wide screening in Saccharomyces cerevisiae.Proc Natl Acad Sci U S A10190399044
- 18. Wong CM, Siu KL, Jin DY (2004) Peroxiredoxin-null yeast cells are hypersensitive to oxidative stress and are genomically unstable. J Biol Chem 279: 23207–23213.CM WongKL SiuDY Jin2004Peroxiredoxin-null yeast cells are hypersensitive to oxidative stress and are genomically unstable.J Biol Chem2792320723213
- 19. Iraqui I, Faye G, Ragu S, Masurel-Heneman A, Kolodner RD, et al. (2008) Human peroxiredoxin PrxI is an orthologue of yeast Tsa1, capable of suppressing genome instability in Saccharomyces cerevisiae. Cancer Res 68: 1055–1063.I. IraquiG. FayeS. RaguA. Masurel-HenemanRD Kolodner2008Human peroxiredoxin PrxI is an orthologue of yeast Tsa1, capable of suppressing genome instability in Saccharomyces cerevisiae.Cancer Res6810551063
- 20. Tong AH, Lesage G, Bader GD, Ding H, Xu H, et al. (2004) Global mapping of the yeast genetic interaction network. Science 303: 808–813.AH TongG. LesageGD BaderH. DingH. Xu2004Global mapping of the yeast genetic interaction network.Science303808813
- 21. Pan X, Ye P, Yuan DS, Wang X, Bader JS, et al. (2006) A DNA integrity network in the yeast Saccharomyces cerevisiae. Cell 124: 1069–1081.X. PanP. YeDS YuanX. WangJS Bader2006A DNA integrity network in the yeast Saccharomyces cerevisiae.Cell12410691081
- 22. Huang ME, Kolodner RD (2005) A biological network in Saccharomyces cerevisiae prevents the deleterious effects of endogenous oxidative DNA damage. Mol Cell 17: 709–720.ME HuangRD Kolodner2005A biological network in Saccharomyces cerevisiae prevents the deleterious effects of endogenous oxidative DNA damage.Mol Cell17709720
- 23. Ragu S, Faye G, Iraqui I, Masurel-Heneman A, Kolodner RD, et al. (2007) Oxygen metabolism and reactive oxygen species cause chromosomal rearrangements and cell death. Proc Natl Acad Sci U S A 104: 9747–9752.S. RaguG. FayeI. IraquiA. Masurel-HenemanRD Kolodner2007Oxygen metabolism and reactive oxygen species cause chromosomal rearrangements and cell death.Proc Natl Acad Sci U S A10497479752
- 24. Chabes A, Georgieva B, Domkin V, Zhao X, Rothstein R, et al. (2003) Survival of DNA damage in yeast directly depends on increased dNTP levels allowed by relaxed feedback inhibition of ribonucleotide reductase. Cell 112: 391–401.A. ChabesB. GeorgievaV. DomkinX. ZhaoR. Rothstein2003Survival of DNA damage in yeast directly depends on increased dNTP levels allowed by relaxed feedback inhibition of ribonucleotide reductase.Cell112391401
- 25. Sabouri N, Viberg J, Goyal DK, Johansson E, Chabes A (2008) Evidence for lesion bypass by yeast replicative DNA polymerases during DNA damage. Nucleic Acids Res 36: 5660–5667.N. SabouriJ. VibergDK GoyalE. JohanssonA. Chabes2008Evidence for lesion bypass by yeast replicative DNA polymerases during DNA damage.Nucleic Acids Res3656605667
- 26. Chabes A, Stillman B (2007) Constitutively high dNTP concentration inhibits cell cycle progression and the DNA damage checkpoint in yeast Saccharomyces cerevisiae. Proc Natl Acad Sci U S A 104: 1183–1188.A. ChabesB. Stillman2007Constitutively high dNTP concentration inhibits cell cycle progression and the DNA damage checkpoint in yeast Saccharomyces cerevisiae.Proc Natl Acad Sci U S A10411831188
- 27. Ogusucu R, Rettori D, Netto LE, Augusto O (2009) Superoxide dismutase 1-mediated production of ethanol- and DNA-derived radicals in yeasts challenged with hydrogen peroxide: Molecular insights into the genome instability of peroxiredoxin-null strains. J Biol Chem 284: 5546–5556.R. OgusucuD. RettoriLE NettoO. Augusto2009Superoxide dismutase 1-mediated production of ethanol- and DNA-derived radicals in yeasts challenged with hydrogen peroxide: Molecular insights into the genome instability of peroxiredoxin-null strains.J Biol Chem28455465556
- 28. Zhao X, Rothstein R (2002) The Dun1 checkpoint kinase phosphorylates and regulates the ribonucleotide reductase inhibitor Sml1. Proc Natl Acad Sci U S A 99: 3746–3751.X. ZhaoR. Rothstein2002The Dun1 checkpoint kinase phosphorylates and regulates the ribonucleotide reductase inhibitor Sml1.Proc Natl Acad Sci U S A9937463751
- 29. Huang M, Zhou Z, Elledge SJ (1998) The DNA replication and damage checkpoint pathways induce transcription by inhibition of the Crt1 repressor. Cell 94: 595–605.M. HuangZ. ZhouSJ Elledge1998The DNA replication and damage checkpoint pathways induce transcription by inhibition of the Crt1 repressor.Cell94595605
- 30. Zaim J, Speina E, Kierzek AM (2005) Identification of new genes regulated by the Crt1 transcription factor, an effector of the DNA damage checkpoint pathway in Saccharomyces cerevisiae. J Biol Chem 280: 28–37.J. ZaimE. SpeinaAM Kierzek2005Identification of new genes regulated by the Crt1 transcription factor, an effector of the DNA damage checkpoint pathway in Saccharomyces cerevisiae.J Biol Chem2802837
- 31. Carter CD, Kitchen LE, Au WC, Babic CM, Basrai MA (2005) Loss of SOD1 and LYS7 sensitizes Saccharomyces cerevisiae to hydroxyurea and DNA damage agents and downregulates MEC1 pathway effectors. Mol Cell Biol 25: 10273–10285.CD CarterLE KitchenWC AuCM BabicMA Basrai2005Loss of SOD1 and LYS7 sensitizes Saccharomyces cerevisiae to hydroxyurea and DNA damage agents and downregulates MEC1 pathway effectors.Mol Cell Biol251027310285
- 32. Elledge SJ, Zhou Z, Allen JB, Navas TA (1993) DNA damage and cell cycle regulation of ribonucleotide reductase. Bioessays 15: 333–339.SJ ElledgeZ. ZhouJB AllenTA Navas1993DNA damage and cell cycle regulation of ribonucleotide reductase.Bioessays15333339
- 33. Gon S, Beckwith J (2006) Ribonucleotide reductases: influence of environment on synthesis and activity. Antioxid Redox Signal 8: 773–780.S. GonJ. Beckwith2006Ribonucleotide reductases: influence of environment on synthesis and activity.Antioxid Redox Signal8773780
- 34. Nordlund P, Reichard P (2006) Ribonucleotide reductases. Annu Rev Biochem 75: 681–706.P. NordlundP. Reichard2006Ribonucleotide reductases.Annu Rev Biochem75681706
- 35. Sanchez Y, Desany BA, Jones WJ, Liu Q, Wang B, et al. (1996) Regulation of RAD53 by the ATM-like kinases MEC1 and TEL1 in yeast cell cycle checkpoint pathways. Science 271: 357–360.Y. SanchezBA DesanyWJ JonesQ. LiuB. Wang1996Regulation of RAD53 by the ATM-like kinases MEC1 and TEL1 in yeast cell cycle checkpoint pathways.Science271357360
- 36. Pellicioli A, Lucca C, Liberi G, Marini F, Lopes M, et al. (1999) Activation of Rad53 kinase in response to DNA damage and its effect in modulating phosphorylation of the lagging strand DNA polymerase. EMBO J 18: 6561–6572.A. PellicioliC. LuccaG. LiberiF. MariniM. Lopes1999Activation of Rad53 kinase in response to DNA damage and its effect in modulating phosphorylation of the lagging strand DNA polymerase.EMBO J1865616572
- 37. Sweeney FD, Yang F, Chi A, Shabanowitz J, Hunt DF, et al. (2005) Saccharomyces cerevisiae Rad9 acts as a Mec1 adaptor to allow Rad53 activation. Curr Biol 15: 1364–1375.FD SweeneyF. YangA. ChiJ. ShabanowitzDF Hunt2005Saccharomyces cerevisiae Rad9 acts as a Mec1 adaptor to allow Rad53 activation.Curr Biol1513641375
- 38. Fillingham J, Keogh MC, Krogan NJ (2006) γH2AX and its role in DNA double-strand break repair. Biochem Cell Biol 84: 568–577.J. FillinghamMC KeoghNJ Krogan2006γH2AX and its role in DNA double-strand break repair.Biochem Cell Biol84568577
- 39. Basrai MA, Velculescu VE, Kinzler KW, Hieter P (1999) NORF5/HUG1 is a component of the MEC1-mediated checkpoint response to DNA damage and replication arrest in Saccharomyces cerevisiae. Mol Cell Biol 19: 7041–7049.MA BasraiVE VelculescuKW KinzlerP. Hieter1999NORF5/HUG1 is a component of the MEC1-mediated checkpoint response to DNA damage and replication arrest in Saccharomyces cerevisiae.Mol Cell Biol1970417049
- 40. Fiorani S, Mimun G, Caleca L, Piccini D, Pellicioli A (2008) Characterization of the activation domain of the Rad53 checkpoint kinase. Cell Cycle 7: 493–499.S. FioraniG. MimunL. CalecaD. PicciniA. Pellicioli2008Characterization of the activation domain of the Rad53 checkpoint kinase.Cell Cycle7493499
- 41. Koc A, Merrill GF (2007) Checkpoint deficient rad53-11 yeast cannot accumulate dNTPs in response to DNA damage. Biochem Biophys Res Commun 353: 527–530.A. KocGF Merrill2007Checkpoint deficient rad53-11 yeast cannot accumulate dNTPs in response to DNA damage.Biochem Biophys Res Commun353527530
- 42. Beckman KB, Ames BN (1997) Oxidative decay of DNA. J Biol Chem 272: 19633–19636.KB BeckmanBN Ames1997Oxidative decay of DNA.J Biol Chem2721963319636
- 43. Finkel T, Holbrook NJ (2000) Oxidants, oxidative stress and the biology of ageing. Nature 408: 239–247.T. FinkelNJ Holbrook2000Oxidants, oxidative stress and the biology of ageing.Nature408239247
- 44. Iraqui I, Kienda G, Soeur J, Faye G, Baldacci G, et al. (2009) Peroxiredoxin Tsa1 is the key peroxidase suppressing genome instability and protecting against cell death in Saccharomyces cerevisiae. PLoS Genet 5: e1000524.I. IraquiG. KiendaJ. SoeurG. FayeG. Baldacci2009Peroxiredoxin Tsa1 is the key peroxidase suppressing genome instability and protecting against cell death in Saccharomyces cerevisiae.PLoS Genet5e1000524
- 45. Graves JA, Metukuri M, Scott D, Rothermund K, Prochownik EV (2009) Regulation of reactive oxygen species homeostasis by peroxiredoxins and c-Myc. J Biol Chem 284: 6520–6529.JA GravesM. MetukuriD. ScottK. RothermundEV Prochownik2009Regulation of reactive oxygen species homeostasis by peroxiredoxins and c-Myc.J Biol Chem28465206529
- 46. Alcasabas AA, Osborn AJ, Bachant J, Hu F, Werler PJ, et al. (2001) Mrc1 transduces signals of DNA replication stress to activate Rad53. Nat Cell Biol 3: 958–965.AA AlcasabasAJ OsbornJ. BachantF. HuPJ Werler2001Mrc1 transduces signals of DNA replication stress to activate Rad53.Nat Cell Biol3958965
- 47. Lee SJ, Duong JK, Stern DF (2004) A Ddc2-Rad53 fusion protein can bypass the requirements for RAD9 and MRC1 in Rad53 activation. Mol Biol Cell 15: 5443–5455.SJ LeeJK DuongDF Stern2004A Ddc2-Rad53 fusion protein can bypass the requirements for RAD9 and MRC1 in Rad53 activation.Mol Biol Cell1554435455
- 48. Loeb LA, Bielas JH, Beckman RA (2008) Cancers exhibit a mutator phenotype: clinical implications. Cancer Res 68: 3551–3557.LA LoebJH BielasRA Beckman2008Cancers exhibit a mutator phenotype: clinical implications.Cancer Res6835513557
- 49. Cao J, Schulte J, Knight A, Leslie NR, Zagozdzon A, et al. (2009) Prdx1 inhibits tumorigenesis via regulating PTEN/AKT activity. EMBO J 28: 1505–1517.J. CaoJ. SchulteA. KnightNR LeslieA. Zagozdzon2009Prdx1 inhibits tumorigenesis via regulating PTEN/AKT activity.EMBO J2815051517
- 50. Winzeler EA, Shoemaker DD, Astromoff A, Liang H, Anderson K, et al. (1999) Functional characterization of the S. cerevisiae genome by gene deletion and parallel analysis. Science 285: 901–906.EA WinzelerDD ShoemakerA. AstromoffH. LiangK. Anderson1999Functional characterization of the S. cerevisiae genome by gene deletion and parallel analysis.Science285901906
- 51. Longtine MS, McKenzie A 3rd, Demarini DJ, Shah NG, Wach A, et al. (1998) Additional modules for versatile and economical PCR-based gene deletion and modification in Saccharomyces cerevisiae. Yeast 14: 953–961.MS LongtineA. McKenzie 3rdDJ DemariniNG ShahA. Wach1998Additional modules for versatile and economical PCR-based gene deletion and modification in Saccharomyces cerevisiae.Yeast14953961
- 52. Huang M, Elledge SJ (1997) Identification of RNR4, encoding a second essential small subunit of ribonucleotide reductase in Saccharomyces cerevisiae. Mol Cell Biol 17: 6105–6113.M. HuangSJ Elledge1997Identification of RNR4, encoding a second essential small subunit of ribonucleotide reductase in Saccharomyces cerevisiae.Mol Cell Biol1761056113
- 53. Lis ET, O'Neill BM, Gil-Lamaignere C, Chin JK, Romesberg FE (2008) Identification of pathways controlling DNA damage induced mutation in Saccharomyces cerevisiae. DNA Repair 7: 801–810.ET LisBM O'NeillC. Gil-LamaignereJK ChinFE Romesberg2008Identification of pathways controlling DNA damage induced mutation in Saccharomyces cerevisiae.DNA Repair7801810
- 54. North TW, Bestwick RK, Mathews CK (1980) Detection of activities that interfere with the enzymatic assay of deoxyribonucleoside 5′-triphosphates. J Biol Chem 255: 6640–6645.TW NorthRK BestwickCK Mathews1980Detection of activities that interfere with the enzymatic assay of deoxyribonucleoside 5′-triphosphates.J Biol Chem25566406645
- 55. Schmitt ME, Brown TA, Trumpower BL (1990) A rapid and simple method for preparation of RNA from Saccharomyces cerevisiae. Nucleic Acids Res 18: 3091–3092.ME SchmittTA BrownBL Trumpower1990A rapid and simple method for preparation of RNA from Saccharomyces cerevisiae.Nucleic Acids Res1830913092