2023 年 98 巻 2 号 p. 61-72
Homologous recombination (HR) is a highly accurate mechanism for repairing DNA double-strand breaks (DSBs) that arise from various genotoxic insults and blocked replication forks. Defects in HR and unscheduled HR can interfere with other cellular processes such as DNA replication and chromosome segregation, leading to genome instability and cell death. Therefore, the HR process has to be tightly controlled. Protein N-terminal acetylation is one of the most common modifications in eukaryotic organisms. Studies in budding yeast implicate a role for NatB acetyltransferase in HR repair, but precisely how this modification regulates HR repair and genome integrity is unknown. In this study, we show that cells lacking NatB, a dimeric complex composed of Nat3 and Mdm2, are sensitive to the DNA alkylating agent methyl methanesulfonate (MMS), and that overexpression of Rad51 suppresses the MMS sensitivity of nat3Δ cells. Nat3-deficient cells have increased levels of Rad52-yellow fluorescent protein foci and fail to repair DSBs after release from MMS exposure. We also found that Nat3 is required for HR-dependent gene conversion and gene targeting. Importantly, we observed that nat3Δ mutation partially suppressed MMS sensitivity in srs2Δ cells and the synthetic sickness of srs2Δ sgs1Δ cells. Altogether, our results indicate that NatB functions upstream of Srs2 to activate the Rad51-dependent HR pathway for DSB repair.
DNA double-strand breaks (DSBs) can arise from exogenous agents such as ionizing radiation and chemical mutagens, as well as through cellular processes, including metabolic reactions and DNA replication (Mehta and Haber, 2014). In Saccharomyces cerevisiae, homologous recombination (HR) plays a major role in the repair of DSBs, and is mediated via the evolutionarily conserved RAD52 epistasis group genes (Symington, 2002; Krogh and Symington, 2004). HR is initiated by the generation of single-strand DNA (ssDNA), a process that involves nucleolytic degradation of the 5′ ends of DSBs, and the resulting 3′ ssDNA ends are coated by the heterotrimeric complex of replication protein A (RPA), a eukaryotic ssDNA-binding protein, to prevent the formation of secondary structures and protect from degradation by nucleases (Bhat and Cortez, 2018). Rad52 mediates the exchange of RPA for Rad51 on ssDNA, leading to the formation of a Rad51-ssDNA nucleoprotein filament (Sung, 1997; New et al., 1998; Heyer et al., 2010). Rad51-ssDNA filaments then search for homologous sequences and catalyze DNA strand exchange to generate recombination intermediates (Sung and Robberson, 1995; San Filippo et al., 2008). In addition, Rad52 and its paralog Rad59 have functions independent of Rad51, which involve ssDNA annealing activity (Mortensen et al., 1996; Davis and Symington, 2001).
Rad51-dependent DSB repair can be processed by two different sub-pathways: the synthesis-dependent strand-annealing (SDSA) pathway, which generates non-crossover products, and the canonical DSB repair (DSBR) pathway, which generates crossover or non-crossover products (Sung and Klein, 2006; Symington et al., 2014). In budding yeast, most mitotic non-crossovers are thought to be formed via the SDSA pathway, in which the newly synthesized DNA strand is displaced from the D-loop, and then annealed to the complementary strand of the other broken DNA end. Srs2 is a member of the highly conserved UvrD family of helicases that plays important roles in DNA replication, recombination and repair (Niu and Klein, 2017). Previous in vitro studies have shown that Srs2 can disrupt the D-loop by its helicase activity, and that Srs2 translocase activity can efficiently dissociate Rad51 nucleofilaments from ssDNA (Krejci et al., 2003; Veaute et al., 2003; Liu et al., 2017). Both activities are required during DSB repair to disrupt the invading strand from the extended D-loop and/or to remove the Rad51 filament from the second end of the DSB, thus promoting the SDSA pathway. In addition, Srs2 acts as an anti-recombinase that counteracts Rad51 functions to prevent inappropriate recombination during DNA replication, and to inhibit interchromosomal and ectopic recombination (Antony et al., 2009; Burgess et al., 2009; Keyamura et al., 2016). On the other hand, the DSBR pathway includes a double Holliday junction (dHJ) as a repair intermediate linked by four-way DNA junctions, where the displaced strand produced by D-loop extension captures the other DSB end. Sgs1 can dissociate dHJs via helicase-driven branch migration coupled with Top3 and Rmi1 (Ira et al., 2003; Cejka et al., 2010). This so-called dHJ dissolution, which generates non-crossover products, prevents crossovers in the DSBR pathway. Alternatively, dHJs can be resolved to produce crossover or non-crossover products by structure-specific endonucleases, such as the Mus81–Mms4 complex, the Slx1–Slx4 complex and Yen1 (Fricke and Brill, 2003; Ehmsen and Heyer, 2008; Ip et al., 2008; Schwartz and Heyer, 2011). Thus, HR intermediates formed by Rad51 are processed by various helicases and/or nucleases, each of which produces a different outcome. Failure in the resolution of these structures results in the accumulation of toxic recombination intermediates that lead to gross chromosomal rearrangement, aberrant chromosomal segregation and cell death. Therefore, the mechanisms for processing HR intermediates must be tightly regulated to exert their exact functions.
N-terminal (Nt)-acetylation is one of the most abundant protein modifications in eukaryotes (Aksnes et al., 2016). NAT (Nt-acetyltransferase) catalyzes the transfer of an acetyl moiety from acetyl CoA to the Nt α-amino group of a polypeptide. Up to 80–90% of all proteins in humans and 50–70% in yeast are Nt-acetylated (Arnesen et al., 2009), suggesting that Nt-acetylation impacts a range of different cellular processes. Indeed, Nt-acetylation can affect protein function in several ways, such as modulation of protein folding, stability, subcellular localization and protein–protein interactions (Ree et al., 2018). There are seven types of NAT in humans (NatA–NatF and NatH) and five types in budding yeast (NatA–NatE), and their functions are classified according to subunit composition and substrate specificity. Most NATs localize to the cytoplasm, and NatA–NatE are associated with the large ribosomal subunit, where they act co-translationally on nascent peptides emerging at the ribosomal tunnel exit (Aksnes et al., 2019). In budding yeast, NatB (a dimeric complex composed of the Nat3 catalytic subunit and the Mdm20 auxiliary subunit) acetylates the Nt methionine of proteins when it is followed by an aspartate, glutamine or asparagine (Arnold et al., 1999; Polevoda et al., 1999; Polevoda and Sherman, 2003; Van Damme et al., 2012; Yamada et al., 2017). nat3Δ mutants are highly sensitive to a variety of stresses, including temperature, osmotic stress and DNA damages (MMS, hydroxyurea, 4-nitroquinolone, cisplatin and ionizing irradiation) (Polevoda et al., 2003; Caesar et al., 2006; Friedrich et al., 2021), suggesting that NatB-mediated Nt-acetylation is an important regulator of the tolerance to these stresses. Interestingly, the rad56-1 mutation, which displays X-ray-hypersensitive and HR-defective phenotypes, is a single-base-pair deletion at position 639 of the NAT3 gene, which leads to a truncated Nat3 protein (Mathiasen et al., 2013). These observations suggest that NatB plays an important role in HR, although the detailed molecular mechanisms associated with this process are currently unknown.
In this study, we investigated the role of the NatB (Nat3–Mdm20) acetyltransferase in HR-dependent repair of DSBs. We show that loss of Nat3 or its catalytic activity results in DNA damage accumulation during MMS exposure. The loss of Nat3 also yields a reduced frequency of gene conversion and gene targeting. Moreover, we found that deletion of NAT3 partially suppresses the MMS sensitivity of srs2Δ cells and the srs2Δ sgs1Δ lethality. Our data have implications for a role of NatB-mediated Nt-acetylation in control of HR that favors the Srs2-dependent pathway during the repair of MMS-induced DNA damage.
The NatB complex is composed of the catalytic subunit Nat3 and the auxiliary subunit Mdm20, and mediates N-acetylation of proteins beginning with methionine-acidic/hydrophilic N-termini. In the present study, we confirmed a previous report that nat3Δ and mdm20Δ cells are both more sensitive to MMS than wild-type cells (Fig. 1A) (Caesar et al., 2006). His74 in the catalytically active site of Naa20 from Candida albicans corresponds to His77 of S. cerevisiae Nat3 and is completely conserved in NatB catalytic subunits (Kubiak et al., 2013; Hong et al., 2017). We constructed the nat3-H77A mutant, in which His77 in the NatB catalytic site has been substituted by an alanine residue. The nat3-H77A mutant displayed MMS sensitivity comparable to that of nat3Δ (Fig. 1A), indicating that NatB-mediated acetylation plays an important role in the MMS-induced DNA damage response.
Nat3 is required for the HR-dependent repair of DSBs. (A) MMS sensitivity of NatB-deficient cells. Cells were cultured in YPDA medium or SC medium lacking Leu. Ten-fold serial dilutions of asynchronous cell cultures were spotted onto MMS-containing YPDA (upper panel) or SC-Leu (lower panel) plates and incubated at 30 ℃ for 3 days. (B) nat3Δ cells are highly sensitive to HO-induced DSBs. Upper panel: schematic representation of the TH1 and TH2 strains. In TH1, an HO cleavage site (HOcs) was inserted into the URA3 locus on chromosome V (ura3-HOcs). A URA3 gene cassette was inserted into the LYS2 locus on chromosome II (lys2::URA3) of TH1 to generate the TH2 strain. Lower panel: cells were cultured in YP-Gly, diluted, and serially spotted onto YP plates containing glucose or galactose. These plates were incubated at 30 ℃ for 3 days. (C) PFGE analysis of chromosomal DNA from MMS-treated cells. Wild-type and nat3Δ cells were mock treated (–) or treated with 0.1% MMS for 0.5 h (+) and released into MMS-free YPDA medium to recover. Samples were taken at the indicated time points after release from MMS treatment. Chromosome DNA was separated by PFGE and detected by staining with SYBR green. (D) Quantitation of cells with Rad52-YFP foci. Cells were treated as in (C). Cells were collected at the indicated times and examined by fluorescence microscopy. Error bars indicate the standard error for three independent experiments. (E) DNA damage checkpoint activation in nat3Δ cells. Wild-type and nat3Δ cells were treated with 0.03% MMS for 1 h and released into MMS-free YPDA medium to recover. Protein extracts were prepared and separated by 6% SDS-PAGE, followed by western blotting with anti-Rad53 antibody.
To explore the link between NatB and DSB repair, we constructed the strains TH1 and TH2, which are derived from the NA14 strain originally constructed by Kupiec and colleagues (Agmon et al., 2009), in which the ura3 gene on chromosome V carries a cleavage site for HO endonuclease (ura3-HOcs) (Fig. 1B). Since TH1 cells lack the homologous sequences required for HR-dependent repair, galactose-induced expression of HO results in a single and persistent DSB at the ura3 locus. As expected, TH1, TH1 rad51Δ and TH1 nat3Δ strains showed severe growth defects in the presence of galactose (Fig. 1B). On the other hand, the TH2 strain contains two URA3 genes: ura3-HOcs on chromosome V and Lys2::URA3 on chromosome II. The second URA3 is the wild-type gene, and therefore has no HO recognition site. The TH2 strain grew normally in galactose-containing medium, but the TH2 rad51Δ strain displayed poor growth (Fig. 1B), suggesting that the HO-induced DSB can be repaired in an HR-dependent manner using an alternative homologous sequence. Similarly, the TH2 nat3Δ strain is more sensitive than the TH2 strain, but not as sensitive as the TH2 rad51Δ strain (Fig. 1B), suggesting a partial defect in DSB repair.
We next examined whether DSB repair occurred in MMS-treated cells by monitoring the appearance of subchromosomal fragments using pulsed-field gel electrophoresis (PFGE). Asynchronously growing wild-type and nat3Δ cells were transiently treated with 0.1% MMS for 30 min, and then allowed to grow in MMS-free YPDA medium. After MMS treatment, all chromosomes were converted into a heterogeneous pool of subchromosomal fragments, as indicated by the appearance of a low-molecular-weight smear band (Fig. 1C). By 4 h after release from MMS treatment, chromosomal DNA bands had reappeared in wild-type cells, indicating that cells had substantially repaired their chromosomal DNA. In nat3Δ cells, much of the chromosomal DNA did not appear as distinct bands even 6 h after MMS was removed (Fig. 1C). We also examined the repair of DSBs by detecting Rad52-yellow fluorescent protein (YFP) focus formation, which correlates with the activation of HR (Alvaro et al., 2007). Cells were treated with 0.1% MMS for 30 min and then released into YPDA medium to recover. In wild-type cells, the number of Rad52-YFP focus-containing cells peaked at 4 h after release from MMS treatment and then gradually decreased (Fig. 1D). In MMS-treated nat3Δ cells, Rad52-YFP focus formation was initially delayed, but thereafter increased markedly up to 8 h after MMS release (Fig. 1D), indicating that HR is impaired during recovery in nat3Δ cells. We speculate that the observed delay in focus formation is due to a delayed generation of the ssDNA region since nat3Δ cells exhibit a slow-growth phenotype. Similar results were obtained when phosphorylation of the checkpoint protein kinase Rad53 was assessed: MMS-induced Rad53 phosphorylation disappeared at 3 h after MMS release in wild-type cells, but not in nat3Δ cells throughout the experiments (Fig. 1E). These data indicated that the NatB acetyltransferase is critical for promoting the HR repair of MMS-induced DSBs.
Deletion of NAT3 reduces the HR levelsTo determine the effect of the nat3Δ mutation on HR function, we measured intrachromosomal gene conversion in the wild-type, nat3Δ and rad51Δ strains, using the lys2BA::URA3::lys2BG direct repeat on chromosome II (Hishida et al., 2002). In these strains, Lys+ Ura− and Lys+ Ura+ prototrophy indicate the occurrence of deletional pop-out and gene conversion events, respectively (Fig. 2A). We found that rad51Δ cells exhibited elevated rates of pop-out recombination (Fig. 2B), in agreement with previous reports that pop-out recombinants due to intrachromosomal direct repeat recombination were generated in a Rad51-independent manner (McDonald and Rothstein, 1994; Liefshitz et al., 1995). Compared to wild-type cells, the rate of gene conversion was reduced 10-fold in rad51Δ cells (Fig. 2B). Likewise, nat3Δ cells exhibited elevated rates of pop-out recombination and reduced rates of gene conversion. These results suggest that NatB loss of function engenders partial defects in HR-dependent gene conversion.
Effect of nat3Δ on the frequency of recombination. (A) Schematic representation of intrachromosomal HR assay between tandemly repeated lys2 heteroalleles. (B) Spontaneous intrachromosomal HR at the lys2BA-URA3-lys2BG locus was assayed as described in Materials and Methods. Recombination rates were calculated according to the median method described by Lea and Coulson (1949). *P < 0.05, **P < 0.01.
A linear double-stranded DNA fragment with sequences homologous to a target chromosomal locus can be integrated at the specific locus by HR. To test the effect of Nat3 on HR-dependent gene targeting (GT), the targeting efficiency was measured by the integration frequency of the URA3 cassette into the CAN1 locus. Ura+ colonies were then tested for canavanine resistance to determine whether marker integration was precisely targeted. The GT frequencies were displayed as the ratio of the URA3 marker integration frequency relative to the plasmid transformation frequency. We found that the GT frequency decreased by 23-fold (0.01) and 8.3-fold (0.03) in rad51Δ and nat3Δ cells, respectively, relative to wild-type cells (0.25) (Table 1), consistent with the reduced HR function in nat3Δ cells. Together, these results demonstrate that NatB contributes to HR-dependent processes.
| Strain | IFa ± SE (× 10−4) | TFa ± SE (× 10−4) | GT (IF/TF)b | Fold decreasec |
|---|---|---|---|---|
| WT | 2.0 ± 0.16 | 8.0 ± 2.2 | 0.25 | 1 |
| nat3Δ | 0.01 ± 0.006 | 0.35 ± 0.24 | 0.029 | 8.3 |
| rad51Δ | 0.018 ± 0.016 | 1.6 ± 0.7 | 0.011 | 23 |
To explore the function of NatB in HR-dependent DSB repair, we constructed the nat3Δ rad52Δ, nat3Δ rad52-327 and nat3Δ rad51Δ double mutants, and compared these double mutants with their respective single mutants for growth and viability in the presence of MMS. A separation-of-function allele, rad52-327, encodes a C-terminally truncated Rad52 protein lacking the Rad51-interaction domain, but retains single-strand-annealing activity (Boundy-Mills and Livingston, 1993; Seong et al., 2008). We observed that the rad52Δ and nat3Δ rad52Δ strains are similarly sensitive to MMS and exhibit a similar reduced viability following MMS treatment (Fig. 3A and 3B). In contrast, the nat3Δ rad51Δ and nat3Δ rad52-327 mutants were more sensitive to MMS and had reduced viability compared to their respective single mutants (Fig. 3A and 3B). These results indicate that Nat3 functions mainly in a Rad52-dependent HR pathway in response to MMS-induced DNA damage, and may also regulate a Rad51-HR pathway and/or a pathway independent of Rad51 (see Discussion).
Genetic interactions between NAT3 and HR genes. (A) Strains of the indicated genotype were spotted in ten-fold serial dilutions onto YPDA plates containing the indicated concentrations of MMS, and incubated at 30 ℃ for 3 days. (B) Survival of MMS-treated cells was measured as described in Materials and Methods. Cells grown to early log phase at 30 ℃ were treated with 0.1% MMS for 1 h, plated on YPDA plates at appropriate dilutions, and incubated at 30 ℃ for 3 days. The number of colonies was counted. Survival is expressed as the percentage of the number of colonies obtained after incubation in the absence of MMS, which was set as 100%. Error bars indicate the standard error for three independent experiments.
We next examined genetic interactions between other HR members and Nat3 for MMS sensitivity. Diploid strains heterozygous for deletion of each gene involved in the HR pathway and for nat3Δ were subjected to meiosis, and double mutants were obtained by tetrad dissection. Notably, the spot assay revealed that the nat3Δ and srs2Δ mutants displayed approximately the same sensitivity to MMS as a nat3Δ srs2Δ double mutant (Fig. 3A). We note that nat3Δ cells may be less sensitive to MMS than is suggested by the spot assay because of its slow-growth phenotype. Indeed, we found that viability of the srs2Δ mutant decreased after MMS exposure compared to the nat3Δ mutant, and the nat3Δ srs2Δ double mutant showed a slight increase in viability compared to the srs2Δ single mutant (Fig. 3B); this indicates that deletion of NAT3 partially suppresses the MMS sensitivity of the srs2Δ strain. On the other hand, deletion of NAT3 in sgs1Δ, mus81Δ, exo1Δ, slx4Δ or rad59Δ mutants had an additive or synergistic effect on MMS sensitivity compared to their respective single mutants (Fig. 3A and 3B). We speculated that this suppressive effect might be particularly noticeable in diploid cells since the srs2Δ diploid cells are much more sensitive to MMS than the corresponding haploid cells (Aboussekhra et al., 1989; Keyamura et al., 2016). To test this, we constructed the srs2Δ diploid counterparts and tested them for MMS sensitivity. As expected, the srs2Δ diploid cells were much more sensitive to MMS (Fig. 4A). Deletion of NAT3 partially suppressed the MMS sensitivity of srs2Δ diploids (Fig. 4A). Similar results were obtained in the liquid assay: deletion of NAT3 suppressed the MMS-induced lethality of srs2Δ diploids (Fig. 4B). Taken together, these data demonstrate that NAT3 and SRS2 are epistatic for the repair of MMS-induced DNA damage and that Nat3 can positively regulate Rad51-dependent repair of DNA damage. If the repair defects in nat3Δ cells are due to the attenuated Rad51-dependent HR pathway, then overexpression of Rad51 should suppress the MMS sensitivity of nat3Δ cells. To test this, we transformed nat3Δ cells with pSC51, which is 2μ multicopy plasmid expressing Rad51, and tested its resistance to MMS exposure. We found that overexpression of Rad51 increased MMS resistance over nat3Δ cells (Fig. 4C). Taken together, these data suggest that Nat3 functions upstream of Srs2, and support the role of NatB as a regulator of the Rad51–Srs2 pathway during HR repair.
NatB functions in an Srs2-dependent HR pathway. (A) Haploid and diploid strains of the indicated genotype were spotted in ten-fold serial dilutions onto YPDA plates containing the indicated concentrations of MMS, and incubated at 30 ℃ for 3 days. (B) MMS sensitivity of wild-type, nat3Δ, srs2Δ and nat3Δ srs2Δ diploids. Survival of MMS-treated cells was determined as described in Fig. 3B and in Materials and Methods. (C) nat3Δ cells were transformed with vector (pRS426) and pSC51. MMS sensitivity was determined on SC-Leu plates in the presence or absence of MMS. Cells were incubated at 30 ℃ for 3 days.
The synthetic lethal interaction between srs2Δ and sgs1Δ can be suppressed by deletion of RAD51 (Gangloff et al., 2000), suggesting that severe growth defects of the srs2Δ sgs1Δ double mutant could be due to toxic recombination intermediates formed by Rad51. To test whether deletion of NAT3 rescues the srs2Δ sgs1Δ lethality, we crossed strains with deletions of each gene to create double and triple mutants by tetrad analysis, and compared their growth rate. We found that while the srs2Δ sgs1Δ double mutant formed no or very small colonies, the srs2Δ sgs1Δ nat3Δ triple mutant formed lager colonies, although they grew more slowly than the wild-type strain (Fig. 5A). In addition, a subset of single, double and triple mutants were grown in YPD liquid medium, and cell density (OD600) of each culture was monitored every 10 min using an OD-Monitor C&T apparatus (Taitec). We found that the srs2Δ sgs1Δ nat3Δ triple mutant grew better than an srs2Δ sgs1Δ double mutant, and its growth rate was comparable to that of the nat3Δ single mutant (Fig. 5B), indicating that loss of Nat3 can suppress the lethality of srs2Δ sgs1Δ cells. Therefore, NatB-dependent Nt-acetylation may contribute to the formation of toxic recombination intermediates in srs2Δ sgs1Δ cells, possibly by promoting the Rad51-dependent HR pathway.
The poor growth of srs2Δ sgs1Δ is partially suppressed by deletion of NAT3. (A) Tetrads from heterozygous diploids for srs2Δ, sgs1Δ and nat3Δ were dissected and grown on YPDA plates at 30 ℃ for 4 days. Circles indicate srs2 sgs1 mutants. Squares indicate nat3 srs2 sgs1 mutants. (B) Growth curve of the mutant strains. Cells were grown in liquid YPDA to early logarithmic phase at 30 ℃ and then diluted to 1 × 105 cells/ml. Cell growth was monitored by measuring the optical density at 600 nm every 10 min.
Nt-acetylation contributes to diverse physiological roles, including cell proliferation, protein homeostasis and stress responses (Ree et al., 2018). In this study, we showed that yeast strains with deletion of NAT3 were highly sensitive to MMS- and HO-induced DSBs. Furthermore, Rad52-YFP foci accumulated in nat3Δ cells, which exhibited a defect in DSB repair after release from MMS exposure. These results demonstrate that NatB-mediated Nt-acetylation plays an important role in promoting the HR-dependent repair of DSBs.
Our data indicate that NAT3 is a member of the RAD52 epistasis group genes and that Nat3 is required for both HR-dependent gene conversion and gene targeting, consistent with a previous study showing that the X-ray-sensitive rad56-1 allele maps to the NAT3 gene (Mathiasen et al., 2013). In addition, we found that deletion of NAT3 partially suppressed the sensitivity of srs2Δ cells to MMS, and also rescued the severe growth defect of srs2Δ sgs1Δ cells. These suppressive effects appear to be related to reduced commitment of the Rad51 pathway in repairing DNA damage, because MMS-induced lethality in srs2Δ and srs2Δ sgs1Δ cells is due to improper recombination and/or the accumulation of toxic recombination intermediates through the action of Rad51. This is also consistent with data showing that Rad51 overexpression suppresses the MMS hypersensitivity of nat3Δ cells. We therefore hypothesize that the Nat3 defect affects the activity of Rad51 itself and/or its associated factors, which results in the increase of unrepaired damages due to the limited Rad51-dependent HR pathway. In these situations, some damage can be processed by a Rad51-independent pathway, but other lesions remain unrepaired and cause cell death, which may be responsible for MMS sensitivity and defective HR in nat3Δ cells. Despite this, we note that nat3Δ rad51Δ double mutants are much more sensitive to MMS than the rad51Δ single mutant. This appears to contradict the above conclusion that Nat3 regulates the Rad51-dependent HR pathway. Although we are currently unable to explain this result, we hypothesize that Nat3 deficiency exacerbates the MMS sensitivity of rad51Δ cells possibly through a dominant negative effect. For instance, in nat3Δ rad51Δ cells, unacetylated forms of proteins that would normally promote the Rad51-HR pathway when acetylated may instead impede alternative repair functions, resulting in an increase in unrepaired damages.
We found that NatB functions upstream of Srs2 during Rad51-dependent HR repair, implicating Nt-acetylation as a positive regulator of HR. Previous proteomic studies have identified many substrates of Nt-acetylation mediated by NatB, but none of these proteins are reported as being directly involved in HR (Arnesen et al., 2009; Van Damme et al., 2012). Bioinformatics analysis based on the amino acid sequence of the N-terminal region predicted that Rad52 and Mus81 are candidate NatB substrates, although they were not experimentally characterized (Yamada et al., 2017). In this context, rad52D2A and mus81E2A mutants, which have mutations in the NatB-mediated Nt-acetylation consensus sequence, showed similar sensitivity to MMS as the wild-type strain (data not shown). Thus, a failure to Nt-acetylate these substrates does not appear to affect MMS sensitivity, suggesting that as-yet-unidentified substrates are involved in the regulation of the HR pathway. However, we cannot rule out the possibility that multiple Nt-acetylation substrates, including Rad52 and Mus81, impact HR-dependent repair in a combined manner. Recent studies have demonstrated that NAT activity can be highly regulated through transcriptional mechanisms and post-translational modifications of proteins (Rathore et al., 2016; Ree et al., 2018), suggesting that the level of Nt-acetylation varies with each substrate and in response to different environments. Therefore, it will be critical in future studies to identify the NatB substrates in response to MMS exposure, as this will provide further insight into the regulatory mechanisms that govern HR repair.
Yeast strains used in this study are listed in Table 2. All strains are derivatives of BY4741 (Open Biosystems) unless noted otherwise. Standard procedures were used for strain construction (Amberg et al., 2005). To create TH1 and TH2 strains, the NA14 haploid strain (Agmon et al., 2009), a derivative of W303-1A, was mated to the W303-1B RAD5+ haploid strain, and the resultant diploids were sporulated and dissected by tetrad analysis. The TH0 haploid (MATa-inc) strain was subsequently selected according to auxotrophic markers (ade2−, URA3+ and LYS2+). TH0 cells were then plated on SC medium containing 1 mg/ml 5-fluoroorotic acid (FOA) to delete the intrachromosomal URA3 repeat. 5-FOA-resistant and canavanine-sensitive colonies were selected to create the TH1 strain. A wild-type URA3 cassette was integrated into the LYS2 locus of the TH1 strain to construct TH2. Plasmid pSC51 was constructed by subcloning the BamHI fragment containing the RAD51 gene from YEP-51 (Shinohara et al., 1992) into a 2μ vector, pRS426 (Stratagene).
| Strain | Genotype | Source |
|---|---|---|
| BY4741 | MATa leu2Δ0 ura3Δ0 his3Δ1 met15Δ0 | Open Biosystems |
| BY4742 | MATα leu2Δ0 ura3Δ0 his3Δ1 lys2Δ0 | Open Biosystems |
| BY4743 | MATa/α BY4741/BY4742 | Open Biosystems |
| NST002 | BY4741 nat3Δ::HIS3 | This study |
| NST003 | BY4741 rad51Δ::KANMX | This study |
| NST004 | BY4741 rad52Δ::KANMX | This study |
| NST005 | BY4741 rad52-327_URA3 | This study |
| NST006 | BY4741 sgs1Δ::KANMX | This study |
| NST007 | BY4741 exo1Δ::KANMX | This study |
| NST008 | BY4741 srs2Δ::KANMX | This study |
| NST009 | BY4741 srs2Δ::URA3 | This study |
| NST010 | BY4741 mus81Δ::KANMX | This study |
| NST011 | BY4741 slx4Δ::KANMX | This study |
| NST012 | BY4741 rad59Δ::KANMX | This study |
| NST013 | BY4741 mdm20Δ::KANMX | This study |
| NST021 | BY4741 nat3Δ::HIS3 rad51Δ::KANMX | This study |
| NST022 | BY4741 nat3Δ::HIS3 rad52Δ::KANMX | This study |
| NST023 | BY4741 nat3Δ::HIS3 rad52-327_URA3 | This study |
| NST024 | BY4741 nat3Δ::HIS3 sgs1Δ::KANMX | This study |
| NST025 | BY4741 nat3Δ::HIS3 exo1Δ::KANMX | This study |
| NST026 | BY4741 nat3Δ::HIS3 srs2Δ::KANMX | This study |
| NST027 | BY4741 nat3Δ::HIS3 mus81Δ::KANMX | This study |
| NST028 | BY4741 nat3Δ::HIS3 slx4Δ::KANMX | This study |
| NST029 | BY4741 nat3Δ::HIS3 rad59Δ::KANMX | This study |
| NST030 | BY4741 nat3Δ::HIS3 srs2Δ::URA3 sgs1Δ::KANMX | This study |
| NST031 | BY4743 nat3Δ::HIS3/nat3Δ::KANMX | This study |
| NST032 | BY4743 srs2Δ::URA3/srs2Δ::KANMX | This study |
| NST033 | BY4743 nat3Δ::HIS3/nat3Δ::HIS3 srs2Δ::3KANMX/srs2Δ::KANMX | This study |
| NST034 | BY4741 RAD52-YFP_URA3 | This study |
| NST035 | BY4741 nat3Δ::KANMX RAD52-YFP_URA3 | This study |
| NST036 | BY4741 lys2BA-URA3-lys2BG | This study |
| NST037 | BY4741 lys2BA-URA3-lys2BG nat3Δ::KANMX | This study |
| NST038 | BY4741 lys2BA-URA3-lys2BG rad51Δ::KANMX | This study |
| NA14 | Mata-inc ura3-HOcs lys2::ura3-HOcs-inc ade3::GALHO ade2-1 leu2-3,112 his3-11,15 trp1-1 can1-100 | (Agmon et al., 2009) |
| TH1 | Mata-inc ura3-HOcs ade3::GALHO ade2-1 leu2-3,112 his3-11,15 trp1-1 can1-100 | This study |
| TH2 | Mata-inc ura3-HOcs lys2::URA3 ade3::GALHO ade2-1 leu2-3,112 his3-11,15 trp1-1 can1-100 | This study |
Cells were grown in yeast extract-peptone-dextrose medium containing 0.01% adenine sulfate (YPDA) at 30 ℃. Synthetic complete (SC) medium was used in strain selection and recombination frequency analysis. For the cell growth assay, overnight cultures were diluted to 1 × 105 cells/ml and cell growth was monitored by measuring the optical density at 600 nm every 10 min using an OD-Monitor C&T instrument (Taitec).
Solid and liquid assays for MMS sensitivityFor solid spotting assays, cells were grown to mid-log phase in YPDA medium. Serial dilutions of the culture were spotted onto YPDA or YPDA + MMS plates, and plates were photographed after 2 or 3 days. In HO-endonuclease-induced strains, each strain was incubated in YP-Gly medium (YP; 1% yeast extract, 2% Bacto Peptone) supplemented with 3% glycerol, and spotted onto YPG plates (YP + 2% galactose). For liquid viability assays, overnight cultures were diluted and allowed to grow at 30 ℃ for 3 h. Cells were washed twice with sterile water and were then treated for 1 h with 0.1% MMS in phosphate-buffered saline (PBS, pH 7.5) at 30 ℃. Aliquots (0.5 ml) were mixed with 0.5 ml of 10% sodium thiosulfate to inactivate the MMS. The cells were then spread onto YPDA plates at appropriate dilutions. After 3 or 4 days, the number of colonies was then counted on each plate. For each strain, percent viability of each strain was expressed as the number of colonies obtained relative to that obtained for the control (no MMS) samples. All data points represent the mean from at least three independent experiments.
Mitotic recombination frequencyIntrachromosomal recombination was monitored in haploid strains containing lys2BA::URA3::lys2BG alleles. Recombination frequencies were determined as described previously (Hishida et al., 2002; Hayashi et al., 2018). Strains were streaked out on solid YPDA medium and grown at 30 ℃ for 2 days. Five colonies from each strain were scraped, suspended in YPDA medium, grown overnight, and plated at the appropriate dilution to determine the total cell number on YPDA plates, and the number of recombinants on selection plates (SC-Lys or SC-Lys, Ura). Recombination rates were calculated according to the median method described by Lea and Coulson (1949). The average was determined for at least three independent experiments.
Gene targeting (GT) frequencyFor the GT assay, the targeting efficiency was measured by the integration of the deletion cassette, which has 50mer sequences homologous to the upstream or downstream regions of CAN1 at both ends, into the CAN1 locus. To construct the deletion cassette, the URA3 gene was amplified from pRS306 by PCR using outer primers consisting of 25 bp of sequence homologous to the URA3 cassette and 50 bp of sequence flanking either the 5′ or 3′ end of the CAN1 gene. Ura+ colonies were then tested for canavanine resistance to determine whether marker integration was precisely targeted. The efficiency of DNA transfer into yeast cells was determined simultaneously by measuring transformation frequency with plasmids carrying the URA3 gene. Yeast cells were transformed using the lithium acetate procedure, with either 400 ng of a URA3 cassette or 400 ng of the plasmid pRS416. Cells were then plated at the appropriate dilution to determine the total cell number on YPDA plates, and the number of transformants on selection plates (SC-Ura). Relative efficiency of GT was expressed as the ratio of CAN1-targeted integration relative to the ratio of transformation with pRS416.
Pulsed-field gel electrophoresis (PFGE) analysisCells grown to early log phase at 30 ℃ were treated with 0.1% MMS for 30 min, washed with 5% sodium thiosulfate, and released into YPD. Samples were taken at the indicated times and chromosome DNA embedded in agarose plugs was prepared as described previously (Keyamura et al., 2016). The yeast chromosomes were separated with CHEF-Mapper XA (Bio-Rad) in 0.8% agarose with 0.5×TBE buffer and stained using ethidium bromide or SYBR Green I (Life Technologies). Gel images were acquired with an LAS4000 mini system (GE Healthcare).
Fluorescence microscopy and analysis of Rad52-YFP localizationCells were treated as in the PFGE analysis. Samples were collected at various time points and resuspended in PBS. Fluorescence microscopy was performed on a Zeiss Axioplan2. Images were visualized using the Lumina Vision imaging software program (Mitani Corporation). More than 100 individual cells were scored for each strain. Data were obtained for at least three independent sets of experiments.
Yeast cell extract and western blottingCells were treated with 0.03% MMS for 1 h, and then washed with 5% sodium thiosulfate and released into YPDA. Yeast cell extracts were prepared from yeast cultures using the trichloroacetic acid method, as described previously (Hishida et al., 2006). Protein samples were separated using SDS-polyacrylamide gels, and proteins were transferred to polyvinylidene difluoride membranes. Rad53 was detected with anti-Rad53 (Santa Cruz) antibody.
We thank members of the Hishida laboratory for discussions and technical assistance. This work was was supported by JSPS KAKENHI Grant Numbers JP17K07290 and JP23114007 (to T. H.).
