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Previous Article | Next Article 
Molecular and Cellular Biology, May 2007, p. 3266-3281, Vol. 27, No. 9
0270-7306/07/$08.00+0 doi:10.1128/MCB.01767-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.
Dipartimento di Biotecnologie e Bioscienze, Università di Milano-Bicocca, 20126 Milan, Italy
Received 19 September 2006/ Returned for modification 27 November 2006/ Accepted 20 February 2007
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ABSTRACT |
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INTRODUCTION |
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In eukaryotes, DNA packaging into chromatin, whose basic structural unit consists of two copies of histones H2A, H2B, H3, and H4 around which DNA is wrapped, represents a natural barrier to DNA accessibility and can interfere with DNA replication. This degree of DNA compaction can be influenced by a wide range of covalent and reversible posttranslational modifications of histones (reviewed in reference 63). In particular, a specialized family of enzymes, the histone acetyltransferases (HATs), catalyzes the transfer of a single acetyl group to the 
-amino group of lysine residues located in the histone N-terminal tails. These events neutralize the lysine charge and can alter histone-DNA and nucleosome-nucleosome interactions, as well as chromatin fiber condensation (reviewed in reference 36). Among Saccharomyces cerevisiae HATs, the NuA4 complex acetylates preferentially histones H4 and H2A through its essential catalytic subunit Esa1, while the SAGA complex acetylates primarily histones H3 and H2B through the Gcn5 subunit (reviewed in references 13 and 36). Histone acetylation is reversible, and all eukaryotic genomes encode histone deacetylases (HDACs) that remove the acetyl groups and thereby reestablish the positive histone charge. S. cerevisiae deacetylases include Rpd3, which together with its regulatory subunit Sin3 is involved in deacetylation of H4, H3, H2A, and H2B lysine residues, and the Hda1 complex, which deacetylates only H3 and H2B histones (reviewed in reference 36).
Although the functions of most acetyltransferases and deacetylases in DNA replication and other cellular processes are still unclear, their combined effects in opening chromatin structure may influence DNA replication and DNA repair. In fact, they can make DNA more accessible to replication and repair machineries and/or create binding sites for replication and repair proteins. Consistent with HAT and HDAC functions in DNA metabolism, loss of the S. cerevisiae deacetylase Rpd3 leads to a general increase in histone acetylation near several replication origins that then undergo precocious firing, indicating that the state of histone acetylation that encompasses an origin is an important determinant for replication timing (2, 61). Moreover, cells lacking the NuA4 subunit Yng2, besides being temperature sensitive, are hypersensitive to DNA-damaging agents and delay completion of S phase in the presence of MMS in a checkpoint-dependent manner at the permissive temperature (7). Similarly, the temperature-sensitive esa1 mutants are hypersensitive to genotoxic agents and show a checkpoint-dependent G2 arrest after shift to the nonpermissive temperature (3, 8).
Some evidence suggests that the actions of acetyltransferase and deacetylase enzymes may be influenced by 14-3-3 proteins, a family of highly conserved polypeptides able to bind to a large number of phosphorylated protein ligands, thus regulating diverse biological processes, such as DNA damage checkpoints, cell cycle progression, apoptosis, stress response, cytoskeleton organization, and malignant transformation (reviewed in reference 1). In fact, although the biological significance of these interactions is still unknown, three members of the Xenopus laevis 14-3-3 protein family have been found to copurify with the HAT Hat1 holoenzyme (20). Moreover, two human 14-3-3 isoforms interact with phosphorylated deacetylases HDAC7 (23) and HDAC4 and HDAC5 (16, 62), and these interactions appear to sequester HDAC4, HDAC5, and probably HDAC7 in the cytoplasm (16).
The two S. cerevisiae members of the 14-3-3 family are encoded by the BMH1 and BMH2 genes, whose double disruption is lethal in most laboratory strains, while single bmh1
or bmh2 mutants do not show detectable growth defects (15, 45, 60). We previously isolated several bmh1 alleles, whose presence in the cell as the sole 14-3-3 source causes temperature sensitivity and hypersensitivity to MMS and hydroxyurea (HU) at the permissive temperature (29, 30). Moreover, several bmh1 alleles caused defective resumption of DNA replication after transient HU-induced S-phase arrest and displayed synthetic lethality when combined with mutations affecting the polymerase 
-primase and replication protein A DNA replication complexes (30). This suggests that Bmh proteins are required to properly carry out DNA replication under stress conditions. Since the activities of HAT and HDAC enzymes are critical for DNA metabolism and may be influenced by 14-3-3 proteins, we used the above bmh1 mutants to investigate the functional and physical interactions between yeast 14-3-3 and HAT/HDAC during DNA replication. We show that inactivation of the HAT NuA4 is detrimental to the growth of bmh mutants, which also display a reduced acetylation of H3 and H4 histone tails after shift to the restrictive temperature. Conversely, the lack of the HDAC Rpd3 or Sin3 allows bmh mutants to complete DNA replication after MMS and HU treatments, suggesting a role for 14-3-3 proteins in responding to replication interference through regulation of acetyltransferase and deacetylase functions. Consistent with such a role, we also find that Bmh1 and Bmh2 physically interact with the acetyltransferase Esa1 throughout the cell cycle and with the deacetylase Rpd3 specifically during unperturbed S phase and after HU treatment.
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MATERIALS AND METHODS |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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ade2-1 can1-100 his3-11,15 leu2-3,112 trp1-1 ura3 ssd1), with the exceptions of strains GA2309, GA2824, JDY16, JDY17, RMY200, RMY253, RMY250, and FLY722, whose genotypes and sources are described below.
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mec1 sml1 strain. All the mec1 strains also carried the sml1 allele (Table 1), which suppresses the lethality of MEC1 deletion but not its effects on DNA damage response (65). To generate the mec1-td allele, a 988-bp DNA fragment downstream of the MEC1 start codon was amplified by PCR using yeast genomic DNA as template and oligonucleotides PRP220 (5'-GGG CAA GCT TCC GGG GGG ATG GAA TCA CAC GTC AAA TAT CTT GAC G-3') and PRP221 (5'-CCA TCG ATT TGT GAA AAA TCT GCC CAC TGG TCA ATT-3') as primers. The PCR amplification product was then cloned into the ClaI-HindIII sites of plasmid pPW66R (11) in order to obtain plasmid pML300. Strains DMP4396/6D and DMP4396/2C were obtained by transforming strains W303 and YLL1090, respectively, with NsiI-digested pML300 plasmid.
The esa1-1851 strain (3) was kindly provided by S. J. Kron (University of Chicago, Chicago, IL). Strain DMP4709/1D was a meiotic segregant from a cross between the esa1-1851 and MATa W303 strains. Strain 1081/6D, carrying the deletion of YNG2, was obtained as a meiotic segregant of the diploid strain 1081 kindly provided by S. J. Kron (7).
Strains JAy22 and JAy45, carrying the deletion of the RPD3 or SIN3 gene, respectively, were kindly provided by O. Aparicio (University of Southern California, Los Angeles, CA) (2) and were crossed to MAT W303 to obtain strains DMP4527/17B and DMP4526/5C, respectively. Strains DMP4512/21B and DMP4509/9D were meiotic segregants from a cross between strain YLL1090 and the MAT rpd3 or MAT sin3 strain, respectively. Strains DMP4590/1D and DMP4591/1C were meiotic segregants from crosses between strain YLL1081 and the MAT rpd3 or MAT sin3 strain, respectively. Strains DMP4592/1B and DMP4593/1D were meiotic segregants from crosses between strain YLL1120 and the MAT rpd3 or MAT sin3 strain, respectively.
Strain YLL1738, carrying a fully functional ESA1-MYC18 allele at the ESA1 chromosomal locus, and strain YLL1739, carrying a fully functional RPD3-MYC18 allele at the RPD3 chromosomal locus, were generated by PCR one-step tagging. Strains YLL909 and YLL910, carrying, respectively, fully functional BMH1-HA3 and BMH2-HA3 alleles at their chromosomal loci, and strain YLL1853, carrying a bmh1-280-HA3 allele at the BMH1 chromosomal locus, were generated by PCR one-step tagging after transformation of strains W303 and YLL1090, respectively. The oligonucleotide sequences used to construct the above tagged alleles are available upon request. MAT strains carrying the BMH1-HA3 and BMH2-HA3 alleles at their chromosomal loci were crossed to strains YLL1738 and YLL1739, respectively, to obtain the meiotic segregants DMP4564/3C, DMP4566/1D, DMP4571/3D, and DMP4572/1B. A MAT bmh2 strain carrying the BMH1-HA3 allele at the BMH1 chromosomal locus was crossed to strains YLL1738 and YLL1739 to obtain the meiotic segregants DMP4653/3A and DMP4654/2B, respectively. A MAT bmh2 strain carrying the bmh1-280-HA allele at the BMH1 chromosomal locus was crossed to strains YLL1738 and YLL1739 to obtain the meiotic segregants DMP4648/10B and DMP4649/10B, respectively.
Strains RMY200 (MATa ade2-101 his3200 lys2-801 trp1-901 ura3-52 hht1 hhf1::LEU2 hht2 hhf2::HIS3 [CEN4 ARS1 TRP1 HHT2 HHF2]), RMY250 (MATa ade2-101 his3200 lys2-801 trp1-901 ura3-52 hht1 hhf1::LEU2 hht2 hhf2::HIS3 [CEN4 ARS1 TRP1 hht2-K9,14,18,23G HHF2]), and RMY253 (MATa ade2-101 his3200 lys2-801 trp1-901 ura3-52 hht1 hhf1::LEU2 hht2 hhf2::HIS3 [CEN4 ARS1 TRP1 hht2-K9,14,18,23R HHF2]) were kindly provided by M. Grunstein (University of California, Los Angeles, CA) (34). Strain FLY722 (MATa ade2-101 leu2-3,112 lys2-801 trp1-901 ura3-52 hhf1::HIS3 hhf2::TRP1-hhf2-K5,8,12,16R HHT2) was kindly provided by J. K. Tyler (University of Colorado, Aurora, CO) (54). Strain HMY140 (MATa ade2-1 can1-100 his3-11,15 leu2-3,112 trp1-1 ura3-1 hht2-hhf2::kanMX3 hht1-hhf1::LEU2 trp1::hht1 K56R-HHF1::TRP1) was kindly provided by A. Verreault (University of Montreal, Montreal, Canada) (35). Strains JDY16 [MAT his31 leu2-3,112 trp1-289 ura3-52 (hht1-hhf1) (hht2-hhf2) pWZ414-F13 (HHT2 HHF2)] and JDY17 [MAT his31 leu2-3,112 trp1-289 ura3-52 (hht1-hhf1) (hht2-hhf2) pRS414-59 (hht2-S10A HHF2)] were kindly provided by S. Dent (14). Strains GA2309 (MAT ade1 leu2-3,112 lys5 trp1::hisG ura3-52 hml::ADE1 hmr::ADE1 ade3::GALHO INO80-MYC13-kanMX4) and GA2824 (MAT ade1 leu2-3,112 lys5 trp1::hisG ura3-52 hml::ADE1 hmr::ADE1 ade3::GALHO INO80-MYC13-kanMX4 hta1-S129A-loxP hta2-S129A-loxP) were kindly provided by S. Gasser (59).
The accuracy of all gene replacements and integrations was verified by Southern blot analysis or PCR. Standard yeast genetic techniques and media were used according to reference 46. Cells were grown in YEP medium (1% yeast extract, 2% Bacto peptone, 50 mg/liter adenine) supplemented with 2% glucose (YEPD).
Genetic interactions and cell viability.
In Table 2, strains YLL1081, YLL1090, and YLL1120 were crossed to W303-derived strains carrying the esa1-1851 or yng2 mutation concomitantly with the bmh2::KANMX4 allele. All diploid strains were therefore homozygous bmh2/bmh2, and each of them was heterozygous for one bmh1 allele and one acetyltransferase mutation. For each sporulated diploid, spores from 14 to 45 dissected tetrads were scored for the ability to form colonies on YEPD plates at 23°C, and segregation of the bmh1, esa1-1851, and yng2 alleles in the bmh2 viable spores was assayed by complementation tests. Synthetic lethality was inferred when no viable double mutant spores were observed from a diploid and if there was no significant deviation from the 1 parental ditype:4 tetratype:1 nonparental ditype ratio of tetrad types predicted from segregation of two unlinked genes. In Table 3, strains YLL1081, YLL1090, and YLL1120 were crossed to a strain concomitantly carrying the esa1-1851, bmh2::KANMX4, and rpd3::HIS5 mutations. For each sporulated diploid, spores from 26 to 30 dissected tetrads were scored for the ability to form colonies on YEPD plates at 23°C, and segregation of the bmh1, esa1-1851, and rpd3 alleles in the bmh2 viable spores was assayed by complementation tests.
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Immunoprecipitations were performed as described in reference 50. Briefly, protein extracts were prepared in a buffer containing 50 mM Tris, pH 7.5, 50 mM NaCl, 50 mM NaF, 5 mM sodium pyrophosphate, 1 mM 4-(2-aminoethyl)-benzene-sulfonylfluoride (AEBSF), 0.1 mM sodium orthovanadate, 0.2% Triton, and a protease inhibitor cocktail (Boehringer Mannheim). After the addition of 1:1 volume of acid-washed glass beads and breakage, 150 µg of the clarified protein extracts was incubated for 2 h at 4°C with 100 µl of a 50% (vol/vol) protein A Sepharose mixture, covalently linked to 12CA5 monoclonal antibodies. The resins were then washed five times in the same buffer and resuspended in 25 µl of SDS gel loading buffer. Bound proteins were visualized by Western blotting with 12CA5 or 9E10 monoclonal antibody after electrophoresis on a 10% SDS-polyacrylamide gel.
Other techniques. Synchronization experiments were performed as described in reference 41. Flow cytometric DNA analysis was performed on a Becton Dickinson FACScan cell sorter.
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RESULTS |
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To investigate further the role of Bmh proteins in responding to perturbations during DNA replication, we analyzed the effects of a sublethal concentration of MMS on S-phase kinetics of the bmh1-280 mutant, which showed more severe defects than the other bmh1 mutants in recovering from HU (30) (Fig. 1D). The bmh1-280 allele carries a single-base-pair substitution resulting in the amino acid change E136G located just C terminal to the residues that have been reported to interact with phosphorylated peptides (30). It is worthwhile to point out that the bmh1-280 allele is the sole 14-3-3 source in the mutant strain, which carries a BMH2 deletion that was seen not to cause by itself any of the phenotypes described for the mutant (data not shown). As shown in Fig. 1A, both wild-type and bmh1-280 cells completed DNA replication by 60 min after the release into unperturbed cell cycle from -factor G1 arrest, indicating that the bmh1-280 allele did not affect the kinetics of DNA replication under unperturbed conditions. In contrast, bmh1-280 cells were not able to reach 2C DNA content within 4 h after release from -factor in the presence of MMS, whereas similarly treated wild-type cells completed DNA replication within 2 h, despite the S-phase delay caused by checkpoint activation (Fig. 1A). This indicates that Bmh proteins are required to complete DNA replication not only after transient HU-induced S-phase arrest but also in the presence of MMS.
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Figure 1 also shows that Mec1-dependent activation of the S-phase checkpoint contributes to the inability of bmh1-280 cells to complete DNA replication in the presence of MMS or after transient HU-induced S-phase arrest. For the MMS experiment, wild-type and bmh1-280 cell cultures, both carrying the deletion of MEC1, were arrested in G1 with -factor and then released into fresh medium with or without MMS. As shown in Fig. 1A, MMS-treated mec1 bmh1-280 and mec1 cells behaved very similarly to each other and completed S phase within 60 min after -factor release, indicating that the slow S phase in MMS-treated bmh1-280 cells is checkpoint dependent.
Since mec1 cells per se fail to resume DNA replication after transient nucleotide depletion by HU (10), we could not use MEC1 deletion to analyze the effect of Mec1 inactivation in resuming DNA synthesis after transient HU treatment of bmh1-280 cells. We therefore generated strains where Mec1 could be rapidly inactivated at 37°C after HU-induced S-phase arrest, due to replacement of MEC1 with the mec1-td allele, where the MEC1 coding sequence is fused in frame to the sequence encoding a temperature-sensitive degron (11). Wild-type and bmh1-280 cells, expressing either MEC1 or the mec1-td allele, were arrested in HU at 25°C for 210 min and then released into the cell cycle at 25°C or 37°C to induce Mec1 degradation in the mec1-td strains. As shown in Fig. 1C and D, both phosphorylated Rad53 disappearance and completion of DNA replication after release from HU at 37°C occurred faster in bmh1-280 mec1-td cells than in bmh1-280 cells, indicating that the inability of the latter to complete DNA replication after transient HU-induced S-phase arrest depends on Mec1.
Altogether, these data indicate that 14-3-3 proteins are required to respond to replication stress and that their dysfunctions can delay completion of S phase in the presence of sublethal HU or MMS concentrations in a checkpoint-dependent manner.
Functional interactions between 14-3-3 proteins and HATs or HDACs.
HAT and HDAC enzymes are found associated with some Xenopus and human 14-3-3 isoforms (16, 20, 23). Moreover, the lack of S. cerevisiae Yng2, a subunit of the acetyltransferase NuA4 complex that is not essential at 25°C, causes a checkpoint-mediated S-phase arrest after MMS treatment at this temperature (7), while temperature-sensitive alleles altering Esa1, the NuA4 essential catalytic subunit, cause sensitivity to genotoxic agents at the permissive temperature (3). We therefore asked whether defects in acetyltransferase and/or deacetylase activities might account for the S-phase defects of bmh mutants. We first studied the possible genetic interactions between bmh1 mutations and the deletion of YNG2 or a temperature-sensitive mutation in the ESA1 gene. To this purpose, we constructed different homozygous bmh2/bmh2 diploid strains, each of which was heterozygous for one bmh1 allele and for either the esa1-1851 temperature-sensitive allele (3) or yng2 (7). Besides the bmh1-280 allele, which is not temperature sensitive, we analyzed by this means also the temperature-sensitive bmh1-221 and bmh1-266 alleles, which were previously shown to cause defects in resuming DNA synthesis after transient HU treatment at 25°C (30). After tetrad dissection and incubation at 23°C, we did not find any viable meiotic segregants carrying the bmh1-221, bmh1-266, or bmh1-280 allele as the sole 14-3-3 source concomitantly with either esa1-1851 or yng2 (Table 2). Thus, functional 14-3-3 proteins are essential for cell viability when the NuA4 acetyltransferase is impaired, suggesting that they might support NuA4 activity.
If the S-phase defects of bmh mutants under stress conditions were due to reduced acetyltransferase activity, inactivation of HDAC might be expected to enhance NuA4 activity and to suppress the HU hypersensitivity of bmh mutants. Remarkably, when deletions of the RPD3 or SIN3 gene, encoding the catalytic or the regulatory subunit of class I HDACs, respectively, were introduced into strains carrying the bmh1-221, bmh1-266, and bmh1-280 alleles as the sole 14-3-3 source, all the bmh rpd3 and bmh sin3 mutants were viable and formed colonies on HU-containing plates more efficiently than the corresponding bmh mutants with functional Rpd3 or Sin3 (Fig. 2). In contrast, neither SIN3 nor RPD3 deletion suppressed the temperature sensitivity of bmh1-221 and bmh1-266 cells (data not shown), indicating that loss of HDAC specifically compensates for the loss of Bmh with respect to HU hypersensitivity.
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combinations. In fact, when we analyzed the viability of spores from different homozygous bmh2/bmh2 diploid strains, each of which was also heterozygous for one bmh1 allele and for the esa1-1851 and rpd3 alleles, all the rpd3 bmh esa1-1851 meiotic segregants were viable (Table 3), and the same was observed in a similar analysis for the rpd3 bmh yng2 combination (data not shown).
Inactivation of Rpd3 or Sin3 can partially restore the ability of bmh mutants to replicate DNA after genotoxic treatment.
As shown in Fig. 3, when wild type and bmh mutants, with or without the deletion of either RPD3 or SIN3, were first arrested in HU and then released into fresh medium, bmh1-221 rpd3, bmh1-266 rpd3, and bmh1-280 rpd3 cells completed DNA replication after release from the HU block earlier than similarly treated bmh1-221, bmh1-266, and bmh1-280 cells with functional Rpd3 (Fig. 3A). Moreover, Rad53 phosphorylated forms disappeared faster in rpd3 bmh1-221, rpd3 bmh1-266, and rpd3 bmh1-280 cells than in bmh1-221, bmh1-266, and bmh1-280 cells after release from HU (Fig. 3B). We obtained very similar results by performing the same analysis with the SIN3 deletion (data not shown). Thus, RPD3 or SIN3 deletion might partially suppress the HU hypersensitivity of bmh cells by relieving their S-phase defects after genotoxic treatment.
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mutants to replicate DNA in the presence of HU or MMS correlated with switching off of the DNA damage checkpoint. In fact, the amount of phosphorylated Rad53 in bmh1-221 rpd3, bmh1-266 rpd3, and bmh1-280 rpd3 cells released in HU or MMS was lower than in similarly treated bmh1-221, bmh1-266, and bmh1-280 cells, where high levels of Rad53 phosphorylation persisted for at least 300 min (Fig. 4C). Altogether these data indicate that Rpd3 or Sin3 inactivation can suppress the S-phase defects of bmh mutants treated with drugs that interfere with replication fork progression, supporting the hypothesis that defects in acetyltransferase activity may account for the inability of these bmh mutants to properly carry out DNA replication under perturbed conditions.
NuA4 acetyltransferase inactivation leads to persistent Rad53 phosphorylation and defective DNA replication under genotoxic treatments.
If acetyltransferase activity plays a role in responding to replication stress, inactivation of the Esa1 or Yng2 subunit of the NuA4 complex might be expected to impair cells' ability to properly carry out DNA replication in the presence of the drugs. As shown in Fig. 5A, cells carrying either the esa1-1851 or yng2 temperature-sensitive allele showed a reduced ability to form colonies compared to wild type when spotted on HU- and MMS-containing plates at 25°C.
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cells were arrested in G1 with -factor and then released from the pheromone block at 25°C, most esa1-1851 and yng2 cells reached 2C DNA content only 240 and 180 min after release in HU and MMS, respectively, whereas HU- or MMS-treated wild-type cells completed DNA replication within 150 and 60 min, respectively (Fig. 5B). Moreover, similarly to bmh mutants, phosphorylated Rad53 remained at high levels for at least 300 min after -factor release in both HU- and MMS-treated esa1-1851 and yng2 cells, whereas it decreased about 120 min after release in similarly treated wild-type cells (Fig. 5C and D). Thus, in agreement with previous studies showing that yng2 cells undergo checkpoint-dependent delay of S-phase completion in the presence of MMS (7), these data indicate that the NuA4 complex has a role in sustaining DNA replication under perturbed conditions.
Interestingly, although Rad53 phosphorylation persisted longer than in wild type in both esa1-1851 and yng2 cells treated with HU, its amount was lower than in wild-type cells when these cells initiated DNA replication in the presence of HU (time, 30 min) (Fig. 5C). Since S-phase checkpoint activation by HU requires replication origin firing (56), whose timing is regulated by histone acetylation (2, 61), defective histone acetylation in esa1-1851 and yng2 mutants may reduce the number of fired replication origins, thus impairing prompt checkpoint activation.
bmh mutants are defective in histone H3 and H4 acetylation. To verify whether 14-3-3 proteins support NuA4-dependent acetyltransferase activity, we analyzed acetylation of histone H4-K5, which is NuA4 dependent (8, 52, 58) (Fig. 6), and of histone H3-K14, which can be acetylated by NuA4 at least in vitro (8, 52). Although Esa1 was known not to be required for acetylation of histone H3-K56 (40) (Fig. 6), which occurs in newly synthesized histone H3 molecules incorporated into chromosomes during S phase (35), we also monitored this acetylation event, as yeast mutant cells that were unable to acetylate H3-K56 were shown to have S-phase defects in the presence of HU (35).
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cells compared to wild type. Very similar results were obtained by monitoring H4 acetylation using antibodies recognizing acetylated H4-K5, -8, -12, and -16 (data not shown). Because the Esa1 catalytic subunit of NuA4 is essential for cell viability at any temperature (8, 52), while Yng2 becomes essential only at 37°C (25) (Fig. 5A), it was unlikely that the bmh, esa1-1851, and yng2 mutations under analysis could dramatically affect NuA4 activity at 25°C. Thus, since also the bmh1-221 and bmh1-266 alleles cause cell lethality at 37°C (30), we repeated the same analysis after keeping all cell cultures for 3 h at the restrictive temperature (37°C) to completely inactivate 14-3-3 when possible. Besides bmh1-221 and bmh1-266, we analyzed by this means also the temperature-sensitive bmh1-103 and bmh1-342 mutants that were previously shown to slow down resumption of DNA replication after transient HU-induced S-phase arrest (30). Under these conditions, H4-K5 and H3-K14 acetylation was consistently diminished in all the temperature-sensitive bmh mutants compared to wild type, indicating that inactivation of 14-3-3 leads to reduced NuA4-dependent acetylation events (Fig. 6A and B). The temperature-resistant bmh1-280 mutant did not show any detectable decrease in these acetylation events at 37°C, while, as expected, inactivation of the NuA4 complex by high temperature in esa1-1851 and yng2 mutants reduced H4-K5 and H3-K14 acetylation compared to that in similarly treated wild-type cells (Fig. 6A and B) and did not alter H3-K56 acetylation. The latter was not defective in any bmh mutant at any temperatures (Fig. 6A), indicating that 14-3-3 proteins are not required for this NuA4-independent acetylation event. Because histone acetylation is involved in the repair of double-strand breaks (DSBs) introduced by site-specific nucleases (3, 21, 44, 54), we asked whether alterations in histone residues acetylated by NuA4 may impair DNA replication under stress conditions. However, yeast strains carrying replacements of the acetylatable H3 lysines 9, 14, 18, and 23 by glycine (H3-K9,14,18,23G) or arginine (H3-K9,14,18,23R) or of the acetylatable H4 lysines 5, 8, 12, and 16 by arginine (H4-K5,8,12,16R) showed a very limited HU sensitivity (Fig. 7A) and were not defective in resuming DNA replication after transient HU-induced S-phase arrest (Fig. 7B).
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Bmh1 and Bmh2 physically interact with Esa1 and Rpd3. Based on the data described above and on the notion that three Xenopus 14-3-3 isoforms are found associated with the Hat1 acetylase (20), while two mammalian 14-3-3 isoforms physically interact with the HDAC4, HDAC5, and HDAC7 deacetylases (16, 23, 62), we asked whether S. cerevisiae Bmh1 could physically interact with the NuA4 and/or Rpd3/Sin3 complexes. To this end, we generated strains concomitantly expressing fully functional Esa1-Myc18- and Bmh1-HA3- or Rpd3-Myc18- and Bmh1-HA3-tagged proteins from the corresponding endogenous promoters. Antihemagglutinin (anti-HA) antibodies were then used to immunoprecipitate Bmh1-HA3 from cell extracts that were prepared from cells collected either during unperturbed exponential growth and S phase or after HU treatment or G1 or G2 arrest. As shown in Fig. 8A, Esa1-Myc18 was specifically recognized by the anti-MYC antibodies in the Bmh1-HA3 immunoprecipitates prepared from all the above cell cycle stages. Also Rpd3-Myc18 was specifically recognized by the anti-MYC antibodies in the appropriate Bmh1-HA3 immunoprecipitates (Fig. 8B) but only when they were prepared from S-phase or HU-treated cells. When anti-Myc antibodies were then used to immunoprecipitate Esa1-Myc18 from exponentially growing cells (Fig. 8C) or Rpd3-Myc18 from HU-treated cells (Fig. 8D), Bmh1-HA3 was specifically recognized by the anti-HA antibodies in both the Esa1-Myc and Rpd3-Myc immunoprecipitates. The observed Esa1-Bmh1 and Rpd3-Bmh1 interactions were specific, since we failed to detect Esa1-Myc18 or Rpd3-Myc18 in anti-HA immunoprecipitates from cell extracts lacking the Bmh1-HA3 protein (Fig. 8A and B) or Bmh1-HA3 in anti-Myc immunoprecipitates from cell extracts lacking the Esa1-Myc18 (Fig. 8C) or Rpd3-Myc18 (Fig. 8D) proteins. We obtained very similar results also when the above coimmunoprecipitation experiments were performed in the absence of Bmh2 (data not shown). Thus, Bmh1 physically interacts in vivo with both Esa1 and Rpd3, although we still do not know whether these interactions are direct. Whereas equivalent amounts of Bmh1 and Esa1 could be coimmunoprecipitated in G1, G2, S, and HU-treated cells, indicating that this interaction is not cell cycle regulated, Bmh1-Rpd3 interaction seems to occur only in S phase and to increase after HU treatment.
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Since defects in acetyltransferase activity might partially account for the S-phase defects of HU- and MMS-treated bmh cells, we asked whether the Bmh1 mutant variants were defective in the above interactions. We could perform this analysis only on the Bmh1-280 variant, whose level is similar to that of the wild type (Fig. 8G and H), because all the other bmh1 mutations under analysis caused a reduction in Bmh1 level in whole-cell extracts compared to wild type. Because the phenotypes of bmh1 mutants become apparent only in the absence of Bmh2, we constructed bmh2 strains concomitantly expressing Esa1-Myc18- and Bmh1-280-HA3- or Rpd3-Myc18- and Bmh1-280-HA3-tagged proteins from their endogenous promoters. As shown in Fig. 8G, similar amounts of Bmh1-HA3 and Bmh1-280-HA3 were immunoprecipitated by anti-HA antibodies from crude extracts prepared from both exponentially growing and HU-treated cells, whereas the amount of Esa1-Myc18 that was recognized by the anti-MYC antibodies was much lower in the Bmh1-280-HA3 immunoprecipitates than in the Bmh1-HA immunoprecipitates. Conversely, similar amounts of Rpd3-Myc18 were specifically recognized by the anti-MYC antibodies in both Bmh1-HA and Bmh1-280-HA immunoprecipitates from HU-treated cells (Fig. 8H). Therefore, the Bmh1-280 variant appears to be specifically defective in the physical interaction with the Esa1 acetyltransferase.
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DISCUSSION |
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Consistent with a function of 14-3-3 proteins in ensuring acetylation events, all the analyzed bmh1 mutations cause synthetic lethality when combined with either a temperature-sensitive esa1-1851 allele or the deletion of YNG2. Moreover, acetylation of histone H3-K14 and histone H4-K5, which are both dependent on NuA4 (8, 52, 58), is reduced in the temperature-sensitive bmh1-103, bmh1-221, bmh1-266, and bmh1-342 mutants after shift to the nonpermissive temperature. Finally, both Bmh1 and Bmh2 physically interact, directly or indirectly, with the HAT Esa1 and with the HDAC Rpd3, whereas the Bmh1-280 variant is defective in its interaction with Esa1, but not with Rpd3, suggesting that NuA4 activity might be affected by this 14-3-3 variant, although we could not detect a significant decrease in H3-K14 or H4-K5 acetylation in bmh1-280 whole-cell extract.
Remarkably, whereas Bmh1-Esa1 physical interaction was detectable during the whole cell cycle, the Bmh1-Rpd3 interaction appears to take place only during S phase, either unperturbed or, to a major extent, after HU treatment. Based on these data and on the observation that inactivation of the HAT NuA4 is detrimental to the growth of bmh mutants, while loss of the HDAC Rpd3/Sin3 function suppresses the S-phase defects of HU- or MMS-treated bmh mutants and their synthetic lethality with esa1-1851 and yng2 mutants, we propose that 14-3-3 proteins modulate the turnover of acetyl groups by promoting NuA4 activity constitutively and by inactivating the deacetylase Rpd3/Sin3 complex during DNA replication or in response to replication interference. This 14-3-3-mediated HDAC inhibition during S phase could increase acetylation of unknown target proteins at sites of ongoing replication. This may in turn facilitate replication fork progression on damaged DNA and/or resumption of stalled forks when the dNTP pool is limiting, by creating a favorable chromatin environment and/or binding sites for DNA replication factors.
Interestingly, a similar function in sustaining replicative stresses by inhibiting HDAC enzymes has been proposed for the DNA damage checkpoints. In fact, the lack of Rpd3/Sin3 deacetylase has been shown to suppress the hypersensitivity to DNA-damaging agents of yeast checkpoint-deficient mutants (49). Moreover, Hst3, which together with Hst4 is required to deacetylate H3-K56 (5, 32), is downregulated upon DNA damage in a Mec1- and Rad53-dependent manner (32).
Several mechanisms can be envisaged for the 14-3-3-dependent regulation of acetylation events. By binding to phosphorylated target proteins, 14-3-3 proteins can activate/repress their enzymatic activities, modulate their compartmental localization, and/or promote/inhibit protein modifications and interactions (37, 64). Since human 14-3-3 isoforms have been proposed to prevent HDAC4 and HDAC5 nuclear localization (16, 62), their yeast Bmh orthologs might prevent the action of Rpd3 and Sin3 deacetylase by modulating their nuclear localization. However, we could not detect any differences in Rpd3 localization during the cell cycle or after HU treatment either in wild type or in bmh mutants (data not shown), suggesting that other mechanisms are likely to operate. On the other hand, Bmh proteins may target the NuA4 acetyltransferase to the replication forks by interacting with phosphorylated histones and/or nonhistone proteins. Consistent with this possibility, human 14-3-3 isoforms are able to bind to a peptide representing the serine 10-phosphorylated H3 tail (33), which is known to be associated with increased acetylation of H3-K14 (6, 24). Moreover, NuA4 has been found to interact in vitro with a peptide representing the C terminus of phosphorylated histone H2A (12). However, replacing the H3 serine 10 or the H2A serine 129 with alanine does not seem to impair the ability of yeast cells to resume DNA replication after transient HU-induced S-phase arrest. Thus, if 14-3-3 proteins respond to replication perturbations by interacting with phosphorylated histones, either the analyzed residues are not involved in this regulation or they are recognized in a combinatorial fashion with other still-unknown histone-modified residues. Alternatively, since some DNA replication proteins are found as potential 14-3-3 interactors (43), 14-3-3 proteins might regulate targets other than histones.
HATs and HDACs may be envisaged to modulate DNA synthesis under perturbed conditions by different ways. For instance, NuA4-dependent histone acetylation may aid in replication fork progression on damaged DNA and/or in resumption of the stalled replication forks by loosening chromatin compaction or by creating appropriate binding surfaces for protein targets. In addition, because histone acetylation is involved in the repair of DSBs introduced by site-specific nucleases (3, 21, 44, 54), NuA4-dependent histone acetylation may facilitate the repair of chromosome breakages that arise when DNA replication occurs on a damaged template or when the dNTP pool is limited. Indeed, point mutations or truncations eliminating the H3 or H4 N-terminal tail have been shown to render cells sensitive to DSB-inducing agents (3, 44, 54). However, we found that replacement of the known acetylatable lysines in the histone H3 or H4 N-terminal tail with glycine or arginine does not significantly affect either HU sensitivity or resumption of DNA replication after transient HU-induced S-phase arrest. This observation, together with the finding that esa1-1851 and yng2 cells displayed S-phase defects at 25°C without showing a decrease in histone H3 and H4 acetylation on whole-cell extracts, suggests that NuA4 may exert its functions during S phase by acetylating histone lysine residues different from those that we analyzed and/or that these acetylated residues might have redundant functions. Alternatively, NuA4 can regulate DNA replication by acetylating target proteins other than histones.
Interestingly, the Arp4 subunit of the NuA4 complex that binds directly to the checkpoint-dependent phosphorylated histone H2A (12) is also part of two ATP-dependent nucleosome remodeling complexes, Swr1 and Ino80. Since Ino80 was isolated from extracts of HeLa cells using 14-3-3 affinity chromatography (43) and is thought to promote severe disruption of histone-DNA interactions at DSBs (57), this raises the possibility that a localized histone eviction by the checkpoint-mediated association of NuA4 at the stalled forks might facilitate the action of DNA repair and replication proteins.
In any case, it is important to point out that, although HATs are able to acetylate histones, an increasing number of nonhistone proteins are being recognized as substrates for HATs, suggesting that HATs may regulate DNA replication under stress conditions by acetylating nonhistone proteins. For example, the HAT Hbo1 interacts with the replication factors Mcm2 and Orc1, which are involved in the early stage of DNA replication, namely, origin recognition and initiation complex formation (4, 18, 19). Moreover, mammalian PCNA, a DNA polymerase sliding clamp involved in DNA replication and repair, coimmunoprecipitates with HDAC1 enzymes and is acetylated in vivo (38). Finally, the human HAT p300 acetylates DNA polymerase ß, which is implicated in the base excision repair pathway, and this acetylation is important to support polymerase ß function in DNA repair (17).
Given the importance of 14-3-3 proteins and HAT and HDAC enzymes in preventing cancer development, the full understanding of the detailed molecular mechanism by which they interact in regulating the response to perturbations in DNA replication will be a challenging subject for future work.
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ACKNOWLEDGMENTS |
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This work was supported by grants from Associazione Italiana per la Ricerca sul Cancro, Fondazione CARIPLO, and Cofinanziamento 2005 Ministero dell'Istruzione, dell'Università e della Ricerca/Università di Milano-Bicocca to M.P.L. F.L. was supported by a fellowship from Fondazione Italiana per la Ricerca sul Cancro.
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FOOTNOTES |
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Published ahead of print on 5 March 2007.
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