Phosphorylation-Regulated Transitions in an Oligomeric State Control the Activity of the Sae2 DNA Repair Enzyme

Abstract

In the DNA damage response, many repair and signaling molecules mobilize rapidly at the sites of DNA double-strand breaks. This network of immediate responses is regulated at the level of posttranslational modifications that control the activation of DNA processing enzymes, protein kinases, and scaffold proteins to coordinate DNA repair and checkpoint signaling. Here we investigated the DNA damage-induced oligomeric transitions of the Sae2 protein, an important enzyme in the initiation of DNA double-strand break repair. Sae2 is a target of multiple phosphorylation events, which we identified and characterized in vivo in the budding yeast Saccharomyces cerevisiae. Both cell cycle-dependent and DNA damage-dependent phosphorylation sites in Sae2 are important for the survival of DNA damage, and the cell cycle-regulated modifications are required to prime the damage-dependent events. We found that Sae2 exists in the form of inactive oligomers that are transiently released into smaller active units by this series of phosphorylations. DNA damage also triggers removal of Sae2 through autophagy and proteasomal degradation, ensuring that active Sae2 is present only transiently in cells. Overall, this analysis provides evidence for a novel type of protein regulation where the activity of an enzyme is controlled dynamically by posttranslational modifications that regulate its solubility and oligomeric state.

INTRODUCTION

DNA double-strand breaks (DSBs) are a deleterious form of DNA damage that must be repaired correctly to avoid mutagenic consequences, including chromosomal rearrangements, deletions, and translocations. Eukaryotic cells use a combination of two broadly defined pathways to repair DSBs: nonhomologous end joining (NHEJ) and homologous recombination (HR) (1). In the budding yeast Saccharomyces cerevisiae, the choice of DSB repair pathway largely depends on the cell cycle phase. Most DSBs detected in the G₁ phase of the cell cycle are repaired by NHEJ, while DSBs detected during the S and G₂ phases are repaired by one of several forms of homologous recombination (2). Cyclin-dependent kinase (CDK) is required for this cell cycle dependency (3, 4), and a few of the targets involved in this process have been identified (2, 5–7).

CDK-mediated phosphorylation of Sae2 is critical for the 5′ strand resection of DSBs (5), a processing event that is a key transition point in the NHEJ versus HR decision. Strand resection is impaired in G₁ phase, in part through inhibition by Ku and other NHEJ factors, but resection occurs efficiently in S and G₂ (8–10). The resection process occurs in two stages: an initiating phase of short-range resection (∼100 to 200 nucleotides) (11) that is promoted and catalyzed by the cooperative activity of Sae2 and the Mre11/Rad50/Xrs2 (MRX) complex and a later phase of extensive resection (up to several kilobases) catalyzed by the redundant activities of Exo1 and Dna2 (12, 13). Dna2, which also acts in Okazaki fragment processing, was also shown to be a target of CDK phosphorylation (6).

Mutation of the CDK target site on Sae2 to alanine (S267A) was previously shown to reduce the rate and extent of DSB resection and to increase the sensitivity of yeast cells to DNA-damaging agents (5), indicating that CDK-dependent phosphorylation of Sae2 is important for cells to repair damage. Mec1/Tel1-mediated phosphorylation of Sae2 after DNA damage was also demonstrated, and mutation of 5 putative SQ/TQ phosphorylation sites in Sae2 increased DNA damage sensitivity and decreased rates of mitotic recombination (14), although it is not known whether these sites are the actual phosphorylation sites in vivo and what effect any of these phosphorylation sites have on Sae2 activities.

We have previously characterized the activities of recombinant Sae2 in vitro, in the form of a maltose-binding protein (MBP) fusion protein expressed and purified from bacteria (15). The recombinant protein is recovered in three different forms (monomer, dimer, and multimer), but these do not show equivalent specific activities. The monomer form shows the highest activity in binding to DNA and cleaving DNA in 5′ flaps and in single-stranded DNA (ssDNA) regions adjacent to hairpin structures, while the dimer is less active and the multimer is inactive. These DNA-binding and nuclease activities are consistent with observations that Sae2 and the MRX complex are essential for the processing of hairpin recombination intermediates in vivo (16–18) and for the removal of 5′ covalent Spo11 conjugates during meiosis (19–21). Recombinant monomeric Sae2 also strongly increases the activity of yeast Exo1 in vitro in a manner that is cooperative with MRX; this activity primarily acts through an increased recruitment of Exo1 to DSB ends (22).

In this study, we investigated the activity of Sae2 in vivo and in vitro to determine how CDK and Tel1 phosphorylation regulates 5′ strand resection and HR through Sae2. We characterized the sites of posttranslational modification through mass spectrometry (MS) and genetic analysis and found that, surprisingly, the phosphorylation events regulate the oligomeric state of the Sae2 protein in a DNA damage-dependent and dynamic manner. We present a model of Sae2 regulation in which the natural insolubility of this protein provides a strong barrier to its activity; however, it is a barrier that can be breached rapidly and reversibly by transient phosphorylation.

MATERIALS AND METHODS

Recombinant protein expression.

Escherichia coli expression constructs for mutant Sae2 were made from pExpGCK566 (15) using QuikChange mutagenesis (Agilent Technologies) according to the manufacturer's instructions. These included S267A (pTP1176), S267E (pTP1172), and S73D/T90D/S249D/T279D/S289D/S267E (5D/S267E; pTP1173), which were transformed into ArticExpress cells (Stratagene) and induced for expression at 13°C overnight. The purification of recombinant MBP-Sae2 and MRX was performed as described previously (15, 22).

Hemagglutinin (HA)-tagged Tel1 protein was purified from the extract of 0.03% methyl methanesulfonate (MMS)-treated yeast cells (KSC1906 MATa-inc ADH4cs::HIS2 ade1 his2 leu2 trp1 ura3 TEL1-HA::TRP1 XRS2-myc::TRP1; a gift from Katsunori Sugimoto). Yeast cells were lysed by blender with dry ice as previously described (23). The lysed cells were dissolved in lysis buffer (25 mM Tris-HCl, pH 7.4, 150 mM sodium chloride, 1 mM EDTA, 10% glycerol, 0.5% NP-40, 1 mM dithiothreitol [DTT]) with 1 mM phenylmethylsulfonyl fluoride and protease inhibitor cocktail (Roche) and then pelleted by centrifugation for 1 h at 100,000 × g at 4°C in a Beckman 70 Ti rotor (Beckman-Coulter) using an Optima L-100 XP ultracentrifuge (Beckman-Coulter). HA-tagged Tel1 protein was then isolated from the supernatant using anti-HA antibody-conjugated agarose beads (Bethyl) and eluted with 0.5 mg/ml HA peptide (AnaSpec).

The isolation of Flag-tagged Sae2 for gel filtration and mass spectrometry analysis was performed as described for Tel1, except that the protein was bound to anti-Flag antibody-conjugated agarose beads (Sigma) and eluted with 0.4 mg/ml 3× Flag peptide (Sigma).

In vitro resection assays.

Resection assays were performed with recombinant Exo1, MRX, Ku, and Sae2 as described previously (22). Reaction mixtures contained linearized 4.5 kb plasmid DNA (0.2 nM), 25 mM MOPS (morpholinepropanesulfonic acid), pH 7.0, 60 mM NaCl, 1 mM DTT, 5 mM MgCl₂, Exo1 (1.2 nM), MRX (3.5 nM), Ku (20 nM), and the Sae2 monomer (fraction number 28) or dimer (fraction number 23) fraction, as indicated in the appropriate figure legends. The reaction mixtures were incubated at 30°C for 60 min, and the reactions were stopped with 0.1% SDS and 10 mM EDTA. Fifty percent of the reaction mixture was reserved for quantitative PCR analysis, while the remainder was separated on a native agarose gel. The gel was stained with SYBR green (Invitrogen), imaged using a Typhoon imager (GE), and then transferred to a nylon membrane with nondenaturing transfer. After UV cross-linking of the DNA to the membrane, it was probed with an RNA probe specific for the 3′ strand of a 1-kb region at one end of the linearized DNA, as described previously (24). The level of ssDNA produced during the reaction was also quantified by real-time PCR, as described previously (22).

Oligonucleotide cleavage assay.

Nuclease assays were performed with [α-³²P]cordycepin-labeled oligonucleotide TP3835 (5′-CTG CAG GGT TTT TGT TCC AGT CTG TAG CAC CAT GCC TAC CTG ACA GTC CGA CAC ATC GGA CTG TCA GGT AGG CAT G-3′). DNA substrates (0.125 nM) were incubated with Sae2 in nuclease buffer (25 mM MOPS, pH 7.0, 65 mM NaCl, 1 mM DTT, 5 mM MgCl₂, 0.1 mg/ml bovine serum albumin) in LoBind tubes (Fisher) at 30°C for 2 h. Reactions were stopped by adding 2 μl of stop solution (0.5% SDS, 50 mM EDTA, pH 8.0, 5 μM TP2622 oligonucleotide), and the reaction mixtures were lyophilized, resuspended in formamide loading buffer, and resolved on a 10% acrylamide–6 M urea–20% formamide gel at a constant wattage (40 W) for 2.5 h. Gels were analyzed by use of a phosphorimager (GE).

Yeast strains used in this study.

The wild-type and sae2 deletion strains used in complementation assays whose results are shown in Fig. 2B and C and Fig. 7A and B were BY4741 and the sae2::kanMX derivative (25), with pRS313 used for complementation as the vector control (26). The wild-type and sae2 deletion strains used in immunofluorescence and solubility assays whose results are shown in Fig. 3A and B, 5, 6, and 7D and E were BY4741 and the sae2::kanMX derivative (25), with pRS425 (27) used for complementation as the vector control. The yellow fluorescent protein (YFP)-SAE2 strain used for the assays whose results are presented in Fig. 2D and 3C and D, and in Fig. S2 in the supplemental material was W4249-5C (MATa ADE2 bar1::LEU2 trp1-1 LYS2 RAD5 SAE2-4Ala-YFP) (28). Genomic mutations at the YFP-SAE2 locus in this strain were made via a 2-step PCR-based method (29), generating strains with the following SAE2 mutations: S267A (TP3503), S267E (TP3495), S267A/S249A/S278A/T279A (A3A; TP5880), S134A/S267A/S249A/S278A/T279A (2A3A; TP5881), and S134E/S267E/S249D/S278D/T279D (2E3D; TP5882). Mutant alleles were fully sequenced at both steps.

FIG 2.

Mutation of Sae2 phosphorylation sites increases the sensitivity of yeast cells to DNA damage. (A) Yeast cells expressing Flag-tagged Sae2 (wild-type or S267A mutant) were exposed to mock or MMS treatment (0.03% MMS for 4 h), and Sae2 was isolated by immunoprecipitation and analyzed by mass spectrometry. The diagram shows all identified phosphorylation and acetylation sites. Also see Table S1 in the supplemental material. (B) Sae2 was expressed from a low-copy-number plasmid under the control of the native Sae2 promoter in sae2Δ yeast cells. Fivefold serial dilutions of cells expressing the indicated Sae2 alleles were plated on nonselective medium (untreated) or medium containing camptothecin or MMS. 2A, S267A/S134A; 3A, S249A/S278A/T279A; A3A, S267A/S249A/S278A/T279A; 2A3A, 2A and 3A mutations combined; E3D, S267E/S249D/S278D/T279D; 2E3D, E3D/S134E. Also see Fig. S2 in the supplemental material. (C) Protein extracts from yeast cells, as described for panel B (without DNA damage), were analyzed by SDS-PAGE and protein blotting with anti-Sae2 and anti-Adh1 antibodies. Vec, vector. (D) Selected Sae2 mutations were introduced into the genomic locus of a YFP-tagged wild-type Sae2 strain. Fivefold serial dilutions of cells were plated as described for panel B.

FIG 7.

Sae2 is degraded during DNA damage through autophagy and the proteasome. (A) Sae2 was expressed from a low-copy-number plasmid under the control of the native Sae2 promoter in sae2Δ or sae2Δ atg1Δ yeast cells. Fivefold serial dilutions of cells expressing the indicated Sae2 alleles were plated on medium containing MMS. QQ, K239Q/K266Q. (B) sae2Δ cells from the assay whose results are shown in panel A were analyzed for total Sae2 protein, as described in the legend to Fig. 2C. (C) L25P Sae2 was isolated before and after DNA damage, normalized for Sae2 levels, and analyzed by protein blotting with anti-phospho-SQ/TQ and anti-Sae2 antibodies, as described in the legend to Fig. 4A. (D) atg1Δ yeast cells expressing wild-type Sae2 protein from a high-copy-number plasmid were synchronized in G₁ phase with α-factor, released into medium containing 0.03% MMS, and analyzed for Sae2 solubility and cell cycle progression, as described in the legend to Fig. 6. (E) atg1Δ yeast cells expressing wild-type Sae2 protein were treated as described for panel B, except that cells were released into medium containing both 0.03% MMS and 75 μM MG132 (MG). (F) Model of Sae2 regulation during the DNA damage response: Sae2 exists in a range of multimeric complexes, some of which are phosphorylated at S267 and possibly also S134 during S and G₂ phases of the cell cycle (green). This phosphorylation and the presence of DNA double-strand breaks promote further Tel1-dependent phosphorylation (red), which initiates the transient disruption of Sae2 multimers into smaller units that are active in promoting DSB resection. DNA damage also promotes acetylation (yellow) through an unknown mechanism, which then stimulates degradation of Sae2 by the autophagy pathway and by the proteasome.

FIG 3.

Mutation of Sae2 phosphorylation sites impairs the localization of Sae2 to DNA breaks. (A) Yeast cells expressing Sae2 from a low-copy-number plasmid were synchronized in G₁ with α-factor and released from the block in the presence of 0.03% MMS. Cells were collected at 0, 20, 40, and 60 min after release and fixed with formaldehyde. Sae2 was imaged using anti-Flag primary antibody and Alexa Fluor 488 secondary antibody. Representative images from the 40-min time point are shown. Arrows, Sae foci. See also Fig. S4 in the supplemental material. (B) Yeast cells were prepared as described in the legend to panel A but were analyzed using an ImageStream^X (Ammis) automated microscope (5,000 cells per strain) to score for Sae2 foci. The percentages of cells with foci are shown for each strain at different time points. (C) Foci of YFP-Sae2 expressed from the chromosomal locus were analyzed as described previously (28) with either no treatment or 40 Gy ionizing radiation (IR). Representative images are shown. Arrowheads, Sae2 foci. (D) Cells containing YFP-Sae2 foci were quantified, and standard errors are plotted (200 to 300 cells per condition).

FIG 5.

Phosphorylation affects the size distribution of Sae2 protein in vivo. (A to D) Yeast cells were grown in the presence or absence of MMS (0.03% MMS for 4 h), and Flag-tagged Sae2 (expressed from a high-copy-number vector) was isolated by immunoprecipitation and then separated by gel filtration. Fractions were analyzed by protein blotting with anti-Sae2 antibody (left). (E and F) Flag-tagged Sae2 expressed from a low-copy-number vector (E) or from the chromosome (F) was isolated from cells grown in the presence or absence of MMS as described for panel A and separated by gel filtration. A dot blot was used for the assay whose results are shown in panel F to analyze the level of Sae2 protein in each sample. The Sae2 concentrations in each fraction were quantitated and are shown as a percentage of the total (right). Examples shown in the figures are representative of several trials.

FIG 6.

Phosphorylation of Sae2 increases its solubility. (A to D) Yeast cells expressing Sae2 wild-type or mutant proteins from a high-copy-number plasmid were synchronized in G₁ phase with α-factor and released from the block in the presence of 0.03% MMS. Cells were collected before (time zero) and every 20 min after the release and lysed under native conditions, and the soluble and insoluble proteins were separated by centrifugation. Both fractions were analyzed by protein blotting for Sae2 protein and quantified (the signal was normalized to the ADH1 levels). (Left) Relative amounts of soluble and insoluble Sae2 protein as percentages of the total; (center) percentage of soluble Sae2 at each time point; (right) FACS analysis of yeast cells using SYTOX green. The approximate positions of 1n and 2n peaks are indicated. (E) Yeast cells expressing wild-type Sae2 from a low-copy-number plasmid were treated and analyzed as described for panels A to D, except that the soluble and insoluble protein lysates were analyzed by mass spectrometry. Quantitation of two Sae2 peptides relative to that of peptides from ADH1 is shown. Examples shown in the figures are representative of several trials.

Yeast Sae2 expression constructs.

The S. cerevisiae wild-type SAE2 gene was cloned into the low-copy-number pRS313 vector (26) under the control of the native SAE2 promoter with a 2× Flag tag at the N terminus (cloning details are available upon request) to create pTP1496. Mutant alleles of SAE2 were made from pTP1496 by QuikChange mutagenesis (Agilent Technologies) to create S267A (pTP1402), S134A (pTP2409), S134A/S267A (2A; pTP2450), S267A/S249A/S278A/T279A (A3A; pTP2331), S134A/S267A/S249A/S278A/T279A (2A3A; pTP2452), S267E/S249D/S278D/T279D (E3D; pTP2408), S134E/S267E/S249D/S278D/T279D (2E3D; pTP2512), S252A (pTP2322), and K239Q/K266Q (QQ; pTP2344) constructs. A high-copy-number vector containing the wild-type SAE2 gene with a 2× Flag tag in pRS425 (27) (Flag-SAE2/2μ) (30) was a gift from John Petrini. Mutant sae2 alleles were made in this plasmid to generate forms with S267A (pTP1598), S267A/S249A/S278A/T279A (A3A; pTP2370), S134A/S267A/S249A/S278A/T279A (2A3A; pTP2467), S267E/S249D/S278D/T279D (E3D; pTP2384), and S134E/S267E/S249D/S278D/T279D (2E3D; pTP2513).

Protein expression analysis in yeast.

Yeast cells (25 optical density at 600 nm [OD₆₀₀] units) were collected and lysed as previously described (31). Cells were first washed with 20% trichloroacetic acid (TCA) and then lysed in 200 μl of 20% TCA by glass bead vortexing. Glass beads were washed twice with 100 μl of 5% TCA. The resulting extracts were combined. After centrifugation, the pellet was washed twice with water and then dissolved with 1× SDS sample loading buffer by vortexing and heating to 100°C for 5 min. The extract was finally clarified by centrifugation again and subjected to protein blotting with anti-Sae2 antibody (a polyclonal antibody custom-made in mouse; Precision Antibody), followed by anti-yeast Adh1 antibody (Abcam). For estimations of wild-type Sae2 concentrations in cells with genomic and low-copy-number plasmid expression, yeast cells were lysed in lysis buffer (see “Recombinant protein expression” above) and vortexed in the presence of glass beads (20 times for 1 min each time). After removal of insoluble material by centrifugation at 20,800 × g for 20 min at 4°C, the Sae2 protein was isolated with anti-Flag-agarose beads, and the eluted Sae2 was compared with a titration of known amounts of MBP-Sae2 using quantitative Western blotting (Licor Odyssey), as well as with TCA-precipitated material from high-copy-number expression strains.

MS. (i) Posttranslational modifications.

Flag-tagged Sae2 was isolated from 12,000 OD₆₀₀ units yeast cells as described above under “Recombinant protein expression” and separated using 12% SDS-PAGE, followed by staining with Coomassie blue. The Sae2 band was excised from the gel and sequentially washed with 25 mM ammonium bicarbonate, acetonitrile, and 10 mM dithiothreitol at 60°C and 50 mM iodoacetamide at room temperature. Trypsin and elastase digestion was performed at 37°C for 4 h. After quenching with formic acid, the supernatant was analyzed. Each gel digest was analyzed by nano-liquid chromatography (LC)/tandem MS (MS/MS) with a Waters NanoAcquity high-pressure liquid chromatography system interfaced to a Thermo Scientific (San Jose, CA) LTQ Orbitrap Velos MS by MS Bioworks, LLC (Ann Arbor, MI). Peptides were loaded on a trapping column and eluted over a 75-μm analytical column at 350 nl/min; both columns were packed with Jupiter Proteo resin (Phenomenex). The mass spectrometer was operated in data-dependent mode, with MS performed in the Orbitrap at a resolution of 60,000 full width at half maximum and MS/MS performed in the LTQ MS. The 15 most abundant ions were selected for MS/MS. Data were searched using a local copy of Mascot with the following parameters: trypsin or no enzyme (for chymotrypsin, elastase, and pepsin); the Saccharomyces Genome Database (forward and reverse appended with common contaminants and recombinant SAE2 sequence); a carbamidomethyl (C) fixed modification; variable modifications of oxidation (M), acetyl (N terminus; K), pyroglutamic acid (N terminus; Q), deamidation (N and Q), and phosphorylation (S, T, and Y); monoisotopic mass values; a peptide mass tolerance of 10 ppm; a fragment mass tolerance of 0.8 Da; and a maximum number of missed cleavages of 2. Mascot DAT files were parsed into Scaffold (version 3) software (Proteome Software, Portland, OR) for validation, with filtering used to create a nonredundant list per sample. Data were filtered using a minimum protein value of 99%, a minimum peptide value of 50% (Prophet scores), and a requirement for at least two unique peptides per protein. Modified peptides with scores greater than or equal to 85% (peptide Prophet scores) were identified. Within a given peptide, modification site assignments were assessed in Scaffold PTM (version 2.1.0) software using the A score and also through manual validation for low-confidence site localization. The Scaffold A-score localization probability was greater than 50% for all mutations except S149 and T151, which were not distinguishable, and S244 and S249, which were difficult to distinguish from S252. Top-scoring sites are reported, with the scores reported in Table S1 in the supplemental material.

(ii) Sae2 quantification.

Data-dependent and dose-response LC-oxpSRM analyses were both performed on a Thermo Fisher Scientific LTQ-Orbitrap Elite mass spectrometer (San Jose, CA) equipped with an ultra-high-pressure Dionex Ultimate 3000 RSLC nano-LC system (Sunnyvale, CA) with buffer A (0.1% [vol/vol] formic acid in water) and buffer B (0.1% [vol/vol] formic acid in acetonitrile). Peptides were concentrated onto an in-house-packed 100-nm (inner diameter) by 2-cm C₁₈ column (3 μm, 100 Å; Magic C₁₈; Michrom Bioresources Inc.) and then separated on a 75-nm (inner diameter) by 50-cm C₁₈ fused-silica column. Liquid chromatography was performed using a gradient of 5 to 40% buffer B over 200 min. For Fourier transform MS1/ion trap MS2-targeted experiments, one scan cycle included an MS1 scan (m/z 300 to 1,700) at a resolution of 60,000, followed by MS2 on selected Sae2 peptides. Peptide/protein identification was performed with the Proteome Discoverer (version 1.3) program embedded with the SEQUEST program (Thermo Fisher Scientific, San Jose, CA). The search parameters used were as follows: two missed cleavages, fixed modifications on cysteine carbamidomethylation, variable modifications on oxidized methionine, a 10-ppm precursor tolerance, and a 0.8-Da MS/MS tolerance were permitted. Peptide identifications were filtered using Percolator, where a 5% false discovery rate was applied. For skyline quantitation, a spectrum library was created from the Proteome Discoverer result to determine the extracted ion current peak area for the full-scan MS1 filtering feature in Skyline software for the 5 transitions for each targeted peptide. A spectral library was created from the Proteome Discoverer result. Skyline (version 1.4) was applied for chromatogram extraction from the MS1 precursor of an Adh1 peptide (GVIFYESHGK) and targeted MS/MS spectra of Sae2 peptides (DNFLFDFNTNPLTK, EQLNQIVDDGCFFWSDK) for peak area calculation. The MS1 precursor isotopic import filter was set to a count of three, (M, M + 1, and M + 2) at a resolution of 60,000. Similarly, the extracted ion count peak areas for MS2 fragment ions were summed for 5 transitions for each targeted peptide.

Gel filtration.

The Flag-Sae2 protein was isolated as described above under “Recombinant protein expression,” and the eluate from the anti-Flag antibody resin was separated by gel filtration using a Superdex200 column (GE) as previously described (15). Fractions after the exclusion volume (fractions 16 to 36) were tested for Sae2 protein concentration by protein blotting or dot blotting and quantitated using a LiCor Odyssey system. Examples shown in the figures are representative of several trials.

Yeast cell immunofluorescence staining.

Twenty-five OD₆₀₀ units yeast cells expressing wild-type or mutant Sae2 in a low-copy-number plasmid were collected, washed with water, and resuspended in medium containing 0.167 mg/ml α-factor (GenScript). After 2 h incubation, cells were collected, washed with water, and released into medium containing 0.03% MMS. Five OD₆₀₀ units of cells was collected at 0, 20, 40, and 60 min after release and then fixed with 3% formaldehyde for 30 min. Fixed cells were digested with 0.5 U/ml zymolase (Zymo Research) at 30°C for 2 h and then incubated with 1:2,000-diluted monoclonal anti-Flag M2 antibody (Sigma) at room temperature for 1 h, followed by 1:1,000-diluted Alexa Fluor 488–donkey anti-mouse (Invitrogen) secondary antibody at room temperature for another 30 min. After washing with phosphate-buffered saline (PBS), cells were stained with 1 μg/ml DAPI (4′,6-diamidino-2-phenylindole) in PBS solution for 5 min.

Automated microscopy with an ImageStream system.

Ten OD₆₀₀ units of yeast cells expressing wild-type or mutant Sae2 in a low-copy-number plasmid was collected at 0, 20, 40, and 60 min after release from α-factor into medium containing 0.03% MMS and then fixed as described above. The immunofluorescence staining steps were also the same as those described above. The stained cells were then analyzed by ImageStream^X microscopy (Amnis) with a 405-nm laser to detect the DAPI signal and a 488-nm laser to detect the Alexa Fluor 488 signal. About 5,000 cells were counted from each sample. The resulting data were analyzed by the use of IDEAS software (Amnis). The data were first compensated for by using data from cells stained with DAPI only or Alexa Fluor 488 only and then gated on focused cells, followed by single cells and fully digested cells. The resulting cell population (>500) was then scored for focus formation with the spot-counting function of IDEAS by comparison to a training set of cells (>50) with defined Sae2 foci.

Fluorescence microscopy.

Sae2-YFP (W4249-5C), Sae2-S267A-YFP (TP3502), and Sae2-S267E-YFP (TP3495) mutants were grown in 5 ml SC medium (46) plus 100 mg/liter adenine at 23°C overnight, either harvested for microscopy or exposed to 40 Gy ionizing radiation (Gammacell 220 cobalt-60 irradiator) by centrifugation, and embedded in 1.4% low-melting-point agar. Images were captured under a ×100 magnification oil immersion objective (numerical aperture, 1.46) on a Leica DM5500B upright microscope illuminated with a 100-W mercury arc lamp and a high-efficiency YFP filter cube (Leica Microsystems). The images were captured with a Hamamatsu Orca AG cooled digital charge-coupled-device camera operated by Volocity software (Improvision). Stacks of 11 0.3-μm sections were captured using the following channels and exposure times: differential inference contrast (DIC) and 12 ms and YFP and 2,800 ms. Approximately 200 to 300 cells were analyzed for each strain, and standard errors were plotted. Images were processed and enhanced identically using Volocity software.

In vitro kinase assays.

The dimer form of wild-type Sae2 protein was incubated with 2.5 nM Tel1, 10 nM MRX, and/or 10 ng DNA (1-kb-plus DNA ladder; Invitrogen) in the presence of 1 mM ATP, 5 mM MgCl₂, 50 mM KCl, 1 mM DTT, 50 mM HEPES buffer (pH 7.5), and 10% glycerol in a volume of 40 μl at 30°C for 90 min. Reaction products were analyzed by Western blotting with anti-phospho-SQ/TQ antibody (Cell Signaling), followed by anti-Sae2 antibody (custom antibody; Precision). To analyze the CDK phosphorylation of Sae2, the dimer form of the wild-type or S267A mutant Sae2 protein was incubated with human CDK2-cyclin A (New England BioLabs) and [γ-³²P]ATP in the presence of 50 mM Mg²⁺, 50 mM MOPS, pH 7.2, 4 mM EDTA, 10 mM EGTA, 0.5 mM DTT, and 25 mM β-glycerophosphate. After 40 min incubation at 37°C, reaction products were separated by 12% SDS-PAGE and detected by phosphorimager analysis. For the two-step in vitro kinase assay, Sae2 protein was first incubated with CDK at 37°C for 30 min and then incubated with Tel1, MRX, and/or DNA (as indicated in Fig. 4) at 30°C for an additional 90 min. Reaction products were separated by 6% SDS-PAGE in the presence of 50 μM MnCl₂ and 25 μM Phos-tag reagent (NARD Institute) in the separating gel to accentuate the differences in charge induced by phosphorylation and then transferred to a polyvinylidene difluoride (PVDF) membrane and subjected to Western blotting with anti-phospho-SQ/TQ antibody, followed by anti-Sae2 antibody.

FIG 4.

Phosphorylation of Sae2 by CDK primes Sae2 for Tel1 phosphorylation. (A) Cells expressing Flag-tagged wild-type (WT), S267A/S134A mutant (2A), or S267E/S134E mutant (2E) Sae2 protein were treated with MMS (0.03% for 4 h), and Sae2 was isolated by immunoprecipitation and analyzed by quantitative Western blotting with anti-phospho-SQ/TQ (p-SQ/TQ) and anti-Sae2 antibodies. The ratio of the phospho-SQ/TQ signal to the Sae2 signal was normalized to 1 for the wild-type protein with MMS treatment, and the corresponding ratios for the other samples relative to that for the wild type are shown below each lane. (B) Recombinant wild-type MBP-Sae2 protein was incubated with purified Tel1, MRX, and/or DNA, as indicated, in the presence of 1 mM ATP and 5 mM Mg²⁺ at 30°C for 90 min. Reaction products were analyzed by protein blotting as described for panel A. (C) Recombinant wild-type or S267A MBP-Sae2 protein was incubated with human CDK2-cyclin A at 37°C and [γ-³²P]ATP (P-ATP), as indicated. Reaction products were separated by 12% SDS-PAGE and analyzed by use of a phosphorimager. (D) Recombinant wild-type MBP-Sae2 protein was first incubated with CDK at 37°C for 30 min and then incubated with Tel1, MRX, and/or DNA, as indicated, at 30°C for an additional 90 min. Reaction products were separated by 6% SDS-PAGE in gels containing Mn²⁺ and Phos-tag reagent (NARD Institute) to accentuate the differences in charge induced by phosphorylation and then transferred to a PVDF membrane and analyzed by protein blotting, as described for panel A. (E) Kinase assays were performed as described for panel D with wild-type, S267A, S267E, and 5D/S267E proteins and analyzed by protein blotting with anti-phospho-SQ/TQ and anti-Sae2 antibodies, as indicated.

Sae2 solubility assay.

Yeast cells expressing Sae2 from high-copy-number (see Fig. 6A to D) or low-copy-number (see Fig. 6E) plasmids were synchronized with α-factor and then released into medium with or without 0.03% MMS, as described above. Samples were collected from the release every 20 min and lysed by glass bead vortexing in extract buffer (100 mM Tris-HCl, pH 8.0, 250 mM ammonium sulfate, 1 mM EDTA, 10% glycerol) with protease inhibitor cocktail (Roche). The cell lysates were centrifuged at 20,800 × g for 20 min at 4°C to separate the supernatant from the pellet. Sae2 protein was extracted from the pellet with extract buffer containing 1% SDS by vortexing and heating to 100°C for 5 min, followed by centrifugation. Both supernatant and pellet extracts were analyzed by Western blotting with anti-Sae2 antibody, followed by anti-ADH antibody. The Sae2 levels on each blot were normalized to the ADH levels for quantification. Examples shown in the appropriate figures are representative of several trials.

Yeast flow cytometry analysis.

One-half OD₆₀₀ unit yeast cells was collected and fixed with 70% ethanol at 4°C overnight with rotation. After fixation, cells were washed, sonicated (three times for 1 s each time), and incubated with 0.25 mg/ml RNase A at 37°C overnight. Cells were then incubated with 5 mg/ml pepsin at 37°C for 15 min and then sonicated again (three times for 1 s each time). Next, cells were stained with 2 μM SYTOX green (Invitrogen) at room temperature for 30 min and analyzed by BD Fortessa flow cytometry with a 488-nm laser.

Targeted mass spectrometry sample preparation.

One hundred OD₆₀₀ units of yeast cells was collected and lysed by a protocol similar to that described above for the solubility assay, except that the pellet fraction was dissolved in extract buffer without SDS or heating. Samples from the supernatant or pellet were incubated with 55 mM iodoacetamide in the dark at room temperature for 30 min, followed by trypsin digestion (2 μg trypsin for 1 to 2 mg/ml total protein) at 37°C for 4 h. Digestion was stopped by adding 1% formic acid. Samples were cleaned up by use of a Pierce C₁₈ spin column (Thermo Fisher Scientific) and then a 50,000-molecular-weight-cutoff Microcon filter (Millipore).

RESULTS

Mutation of phosphorylation sites in Sae2 alters the multimer-monomer distribution.

We have previously analyzed the activities of recombinant Sae2 in vitro as a maltose-binding protein (MBP) and histidine-tagged fusion protein expressed in E. coli, where the recombinant protein elutes as three distinct forms from gel filtration: a multimer, a dimer, and a monomer (15). The monomeric form of Sae2 exhibits the highest specific activity in nuclease assays in vitro, with the dimeric form showing ∼10-fold less activity than the monomer and the oligomeric form showing essentially no activity (Fig. 1A). Although this result suggests that phosphorylation is not required for enzymatic activity since the E. coli-produced protein is not phosphorylated, we mutated the S267 residue to either an alanine or a glutamic acid and found that these changes affected the relative amounts of protein recovered in the multimeric peak (Fig. 1B). Further, mutation of all five putative Tel1 phosphorylation sites (14) to aspartic acid together with S267E (5D/S267E) caused the distribution of Sae2 to change dramatically: the multimer peak nearly disappeared, and the monomer-size protein species increased in abundance (quantified in Fig. 1C).

FIG 1.

Mutation of phosphorylation sites in recombinant Sae2 protein alters the multimer-monomer distribution. (A) Nuclease assays in vitro with recombinant Sae2 monomers, dimers, and oligomers purified by gel filtration. Resection assays included a 3′ ³²P-labeled hairpin DNA substrate, as shown, and 0.5, 2, or 8 nM Sae2. Reaction products were separated by denaturing polyacrylamide gel electrophoresis and analyzed by use of a phosphorimager. nt, nucleotides. (B) E. coli-expressed recombinant wild-type or mutant Sae2 proteins (S267A, S267E, S73D/T90D/S249D/T279D/S289D/S267E [5D/S267E]) containing 6× histidine and MBP affinity tags for purification were analyzed by gel filtration and monitored by determination of the UV absorbance at 280 nm. (C) Quantitation of the data in panel B showing the total absorbance in each peak. The peaks were defined as oligomer (fraction numbers 16 to 19; 8 to 9.5 ml), dimer (fraction numbers 20 to 26; 10 to 13 ml), and monomer (fraction numbers 27 to 29; 13.5 to 14.5 ml). (D) The monomer form of recombinant wild-type (WT) or mutant Sae2 protein (or the dimer form of the wild-type protein) was used in an in vitro DNA resection assay with yeast Exo1 (yExo1) (1.2 nM), MRX (3.5 nM), Ku (20 nM), or Sae2 (2.5 nM monomer, 2.5 and 25 nM dimer), as indicated. The degradation of the 5′ strand at one end of the DNA was analyzed by SYBR green staining for total double-stranded DNA (top) or by quantitative PCR with primers located 29 bp or 1,025 bp from the end (bottom).

Analysis of the activities of the monomer forms of wild-type, S267A, and 5D/S267E recombinant proteins showed that they are all active in endonuclease activity (see Fig. S1 in the supplemental material). The proteins were also tested in an in vitro resection assay with MRX and Exo1, and the degradation of the 5′ strand at one end of the DNA was analyzed by quantitative PCR as previously described (22). Under these conditions where the Ku heterodimer is present, Exo1 is strongly stimulated by MRX and by Sae2 (22). Interestingly, the S267A mutant protein appeared to be much less active than the wild-type protein in the stimulation of Exo1 in vitro when the concentrations of MRX and Exo1 were limiting (Fig. 1D). The wild-type dimer protein showed a lower specific activity in this assay, as in the nuclease assay, but still exhibited a stimulatory effect on Exo1 in the presence of Ku and MRX.

Sae2 phosphorylation is essential for the survival of DNA damage in vivo.

From the analysis of MBP-Sae2 proteins, we observed that mutations in residues predicted to be phosphorylation targets strongly affected oligomeric distributions and also had separate effects on resection. To address how phosphorylation and other modifications affect Sae2 activity in a natural context, we sought to identify those sites using Sae2 protein isolated from S. cerevisiae. Flag-tagged wild-type or S267A mutant Sae2 proteins were immunoprecipitated from yeast cells treated with or without the DNA alkylating agent methyl methanesulfonate (MMS) and then analyzed by mass spectrometry. As summarized in Fig. 2A, many sites were phosphorylated, with additional sites found only in the MMS-treated wild-type sample. Within this group there were two SQ/TQ sites, S249 and T279, which are likely to be Tel1 targets. Phosphorylation on some sites (S21, S134, S244, S249, S278, and T279) was not observed in an S267A mutant, indicating that phosphorylation of these sites is dependent on CDK phosphorylation of S267. Two lysines on Sae2, K239 and K266, were also acetylated after DNA damage.

On the basis of the analysis of posttranslational modifications, individual mutations or combinations of mutations were introduced into a low-copy-number plasmid containing the wild-type Sae2 gene under the control of the native Sae2 promoter and tested for complementation of sae2Δ in DNA damage sensitivity tests (representative data are shown in Fig. 2B) and protein expression (Fig. 2C). Single mutation of most of the sites did not increase the sensitivity of yeast cells to camptothecin (CPT) or MMS (data not shown), except for the known S267A mutation. However, the combination of S267A and S134A (2A) mutations together strongly increased DNA damage sensitivity, while the combination of S267E and S134E (2E) fully complemented the damage sensitivity of sae2Δ (Fig. 2B; see Fig. S2 in the supplemental material). Considering that S134 is an SP site, it is possible that S134 is also targeted by CDK. Many different combinations of SQ/TQ site mutations were also tested, but the highest sensitivity to DNA damage was observed with mutation of S134 and S267 combined with three C-terminal sites, S249, S278A, and T279A (2A3A). In contrast, the phosphomimic version of these mutations (2E3D) showed intermediate levels of growth in the presence of CPT and MMS. Phosphorylation of the three most important non-CDK sites (S249, S278, and T279) was not observed by mass spectrometry in an S267A mutant, yet the phenotype of the 2A3A mutant-expressing strain was clearly more severe than that of either the S267A or S267A/S134A (2A) mutant alone. This result suggests that there is likely some low-level phosphorylation of S249/S278/T279 in the absence of CDK modification. Overall, however, the survival assays showed that the phosphorylation of the S134, S267, S249, S278, and T279 residues is very important for the function of Sae2 in response to DNA damage.

To further confirm the phenotype, we introduced some of the representative mutations into the genomic Sae2 locus of a YFP-tagged wild-type Sae2 strain. In this context, the S267A mutant showed a more severe defect in response to CPT than MMS, and the phenotype was generally much more severe than the phenotype with plasmid expression, as was the 2E3D mutant phenotype (Fig. 2D). Similar results were observed with ionizing radiation (see Fig. S3 in the supplemental material).

Mutation of Sae2 phosphorylation sites impairs Sae2 localization to DNA damage sites.

Similar to the MRX complex, Sae2 localizes to DNA break sites very rapidly following DNA damage (28). To determine if the phosphorylation of Sae2 alters its localization, yeast cells expressing Flag-tagged wild-type or mutant Sae2 alleles on low-copy-number plasmids were analyzed for focus formation using immunofluorescence staining with anti-Flag antibody. Cells were first synchronized in G₁ phase with α-factor and then released into medium containing 0.03% MMS. Samples were collected before (time zero) or 20, 40, or 60 min after release and then fixed for fluorescence-activated cell sorting (FACS) and immunofluorescence staining (representative images from each of the strains at the 40-min time point are shown in Fig. 3A; see also Fig. S4 in the supplemental material). To quantify the foci, the cells were also imaged and analyzed using an automated microscope, which counted approximately 5,000 cells at each time point and scored for focus formation using a set of training images for comparison (Fig. 3B). These results show that Sae2 foci are initially present in wild-type cells but transiently decrease upon DNA damage and then increase during further damage exposure. Fewer cells expressing the 2A3A and 2E3D mutants than wild-type cells exhibited Sae2 foci, and the percentage for cells expressing the 2A3A mutant failed to increase above the initial percentage of cells showing foci.

We also examined the wild type and the S267A and S267E mutants expressed from the chromosome as YFP fusions and found that the wild-type protein increased focus formation in response to ionizing radiation, but the S267A and S267E mutant strains exhibited reduced responses to DNA damage (Fig. 3C and D). The S267A-expressing strain showed a higher basal level of focus-containing cells than the wild type did (P ≤ 0.005), and this level decreased slightly, although not significantly, after ionizing radiation treatment.

Phosphorylation of Sae2 by CDK primes Sae2 for Tel1 phosphorylation.

The analysis of Sae2 modifications showed that CDK-mediated phosphorylation is required for other phosphorylation events, some of which were at Tel1 consensus sites. To test if CDK phosphorylation promotes the SQ/TQ phosphorylation of Sae2 after DNA damage in vivo, Flag-tagged Sae2 (wild-type, S267A/S134A [2A], or S267E/S134E [2E] mutants) was immunoprecipitated from yeast cells treated with 0.03% MMS. As shown in Fig. 4A, yeast cells with the mutant 2A or 2E Sae2 plasmids had a significantly lower level of phospho-SQ/TQ signal than the wild type, indicating that CDK phosphorylation increases the SQ/TQ phosphorylation of Sae2 after DNA damage in vivo.

An in vitro kinase assay with both CDK and Tel1 could address the question of whether CDK phosphorylation directly primes Sae2 for Tel1 phosphorylation or whether this is an indirect effect. To reconstitute Tel1 activity in vitro, HA-tagged Tel1 protein was purified from extracts of MMS-treated yeast cells by HA antibody-conjugated agarose beads. Recombinant wild-type MBP-Sae2 protein was incubated with Tel1, the MRX complex, and DNA, and the reaction products were analyzed by protein blotting with phospho-SQ/TQ antibody. As shown in Fig. 4B, Sae2 was phosphorylated by Tel1 in vitro, requiring the presence of both MRX and DNA, identical to our previous observations with human ATM and MRN (32).

To investigate the role of CDK in this process, we used recombinant human CDK2-cyclin A to phosphorylate wild-type or S267A Sae2 in an in vitro kinase assay in the presence of [³²P]ATP and monitored phosphorylation with ³²P incorporation (Fig. 4C). Wild-type Sae2 was phosphorylated by CDK, while the S267A mutant showed a greatly reduced signal, confirming that S267 is the major CDK phosphorylation site on Sae2. Low-level phosphorylation was still observed with the S267A mutant, however, suggesting that other CDK target sites exist on Sae2.

To determine if CDK phosphorylation affects the efficiency of subsequent Tel1 phosphorylation, a two-step kinase assay was performed in which wild-type Sae2 protein was incubated first with CDK and then with Tel1, MRX, and DNA. Reaction products were separated by SDS-PAGE in the presence of Phos-tag reagent to accentuate the differences in charge induced by phosphorylation (33) and then analyzed by protein blotting for phospho-SQ/TQ residues and for total Sae2 (Fig. 4D). Sae2 phosphorylation by Tel1 (measured from the phospho-SQ/TQ signal) increased significantly after preincubation with CDK, which shows that CDK phosphorylation directly primes Sae2 for Tel1 phosphorylation. The phospho-SQ/TQ signal appeared at a much higher position than the anti-Sae2 signal, likely indicating multiple phosphorylation events. We also tested the S267A and S267E mutants in this reaction and confirmed that Tel1 phosphorylation of both of these was significantly reduced in comparison to that for the wild-type protein (Fig. 4E). Taken together, these results suggest that Sae2 is phosphorylated by CDK primarily at S267 and that this modification primes Sae2 for Tel1 phosphorylation in the presence of MRX and linear DNA ends.

Phosphorylation affects the size distribution of Sae2 in vivo.

A previous study suggested that the majority of Sae2 exists as a multimer form in vivo (30), and our analysis of recombinant MBP-Sae2 in vitro suggested a relationship between phosphorylation and the equilibrium between different multimeric forms (Fig. 1). To investigate whether a change in the oligomeric state of Sae2 occurs after DNA damage in vivo, we expressed Flag-tagged wild-type Sae2 protein in S. cerevisiae from a high-copy-number vector and isolated the soluble protein by immunoprecipitation either before or after MMS treatment. The protein was separated by gel filtration, and fractions were analyzed by protein blotting with anti-Sae2 antibody, as shown in Fig. 5. Wild-type Sae2 clearly shows a DNA damage-dependent change in the overall size distribution of the protein (Fig. 5A), such that smaller forms become more prevalent with damage treatment. Unlike the distinct monomer/dimer/multimer forms of MBP-Sae2 expressed and purified from E. coli, the non-MBP-tagged protein isolated from yeast appeared in a continual gradient of apparent molecular weight/size. Comparison of this result with the distribution of a molecular weight standard suggested that fractions 18 and 19 contain Sae2 in a multimeric form or in a large protein complex with other proteins, while fractions 25 to 27 and 29 to 31 are expected to contain dimer and monomer Sae2, respectively.

Analysis of Sae2 proteins with mutations in the phosphorylation sites showed that the S267A mutant Sae2 appeared to be slightly smaller than the wild-type protein but changed less with DNA damage (Fig. 5B), and the S134A/S267A/S249A/S278A/T279A (2A3A) mutant did not change at all with damage (Fig. 5C). In contrast, the S134E/S267E/S249D/S278D/T279D (2E3D) mutant exhibited a much smaller size that was approximately similar to that of the expected dimer and also did not change after DNA damage (Fig. 5D). Similar results were obtained with the A3A and E3D mutants, which lack the mutation at S134 (see Fig. S5 in the supplemental material).

The size distribution of Sae2 oligomers is likely determined by the level of expression in vivo, so we also isolated Flag-tagged Sae2 from cells expressing the protein from a low-copy-number vector (Fig. 5E). This protein also exhibited a transition in size distribution that was visible after DNA damage treatment; thus, the change in size is not dependent on high-level expression of the protein. With low-copy-number expression, the initial size of the oligomers was clearly smaller than that with high-copy-number expression (compare Fig. 5A to E), but the appearance of even smaller forms in fractions 25 to 29 was visible only with damage treatment. Lastly, wild-type Sae2 was recovered from yeast cells expressing the protein from the chromosomal locus, as shown in Fig. 5F. In this case, a dot blot of the gel filtration fractions was necessary to analyze the lower overall level of the Sae2 antibody signal. The pattern of Sae2 distribution also changed dramatically with DNA damage treatment in this case, with the nearly complete loss of the large oligomers and accumulation of the monomer and dimer forms. Overall, we conclude from this set of experiments that DNA damage promotes the release of Sae2 from a multimer form to a range of smaller forms and that the extent of this release is affected by phosphorylation. The absolute size of the multimer is dependent on the expression level, as would be expected, but there is a transition to the monomer and dimer forms that is damage dependent.

Phosphorylation of Sae2 increases its solubility.

Sae2 protein isolated by immunoprecipitation primarily came from the soluble fraction of the yeast cell extract, but a large majority of Sae2 protein was left in the insoluble pellet (data not shown). To examine the dynamics of Sae2 in both the insoluble and soluble fractions, an assay was developed to extract and separate these pools of Sae2 protein from yeast cells. Cells expressing Flag-tagged Sae2 were first synchronized into G₁ phase with α-factor and then released into medium with or without 0.03% MMS. Samples were collected every 20 min after the release for both solubility assays and cell cycle analysis by flow cytometry. The percentage of soluble and insoluble Sae2 protein was calculated at each time point in comparison to the amount of an ADH1 control protein, which did not change over this time course. The results in Fig. 6 show the soluble and insoluble pools as percentages of the total amount of protein (left column) and the amount of soluble protein only (middle column), while the cell cycle progression of each culture is shown in the right column. The pool of soluble wild-type Sae2 increased 20 min after release and was more apparent in samples with MMS treatment (Fig. 6A and B, middle). In contrast, this increase was absent with the 2A3A and 2E3D mutants (Fig. 6C and D, middle). In addition, the 2A3A mutation resulted in much less soluble Sae2, but the 2E3D mutation made it much more soluble. These experiments were performed with Sae2 expressed from a high-copy-number plasmid in order to have enough protein for detection by blotting, but a similar experiment was also performed with wild-type Sae2 expressed from a low-copy-number vector to rule out the possibility that overexpression was responsible for the Sae2 multimerization. In this experiment (Fig. 6E), targeted mass spectrometry analysis was used instead of protein blotting to determine the relative amount of Sae2 in each fraction. Results from two Sae2 peptides indicated increased levels of Sae2 in the soluble fraction after DNA damage and decreased levels in the insoluble fraction, further confirming our finding that the solubility of Sae2 increases after DNA damage. Lastly, soluble Flag-Sae2 was isolated from cells expressing Sae2 in a single copy from the chromosome; this analysis showed that the amount of soluble Sae2 increased by 1.6- ± 0.13-fold following DNA damage treatment (calculated by quantitative Western blotting of Flag-Sae2 isolated from cells before versus after treatment with 0.03% MMS as the average of 3 trials with the standard deviation).

One prediction of the damage-induced solubility model is that smaller forms of Sae2 should be active. We tested an allele of Sae2 identified in a previous study that was reported to be deficient for self-interaction, L25P (30), which we confirmed (see Fig. S6 in the supplemental material), but found that it fails to complement a null strain for DNA damage survival (Fig. 7A). The levels of this mutant are low relative to those of the wild-type protein, however (Fig. 7B), and we also found that L25P fails to be phosphorylated by Tel1 after DNA damage treatment (Fig. 7C). Thus, oligomerization of Sae2 is essential for its phosphorylation, and monomerization is clearly not sufficient for functional activity.

Sae2 is degraded after DNA damage through autophagy and proteasome degradation.

Autophagy is the preferred pathway for the degradation of some protein aggregates (34), and a previous study showed that Sae2 was acetylated and degraded through the autophagy pathway after DNA damage (35), but acetylation sites were not identified. We observed two damage-dependent acetylation sites on Sae2, K239 and K266 (Fig. 2A), one of which was immediately adjacent to the S267 CDK phosphorylation site. Individual or combined mutations of the acetylated residues to arginine to block acetylation did not have any effect on DNA damage survival (data not shown); however, mutation of both residues to glutamine to mimic acetylation generated a growth defect even in the absence of damage (Fig. 7A), which correlated with a significantly increased level of Sae2 protein (Fig. 7B). This effect was observed in both a wild-type and an autophagy-deficient (atg1) strain background. Interestingly, the S252A mutant that was deficient in one of the DNA damage-dependent phosphorylations identified by mass spectrometry (Fig. 2A) exhibited increased resistance to DNA damage compared to that of the wild-type parent, an effect that was not observed in the atg1 deletion strain (Fig. 7A).

A consistent feature of Sae2 dynamics during DNA replication in the presence of DNA damage is the transient loss of protein observed particularly at the 20-min and 40-min time points (Fig. 6). To determine if this reduction in total protein is dependent on autophagy, Sae2 solubility assays were performed as described in the legend to Fig. 6 but with the atg1 deletion strain (Fig. 7D). These results showed that the transient loss of protein was not seen in the absence of a functional autophagy pathway. The effect of proteasome-mediated degradation was also assessed using MG132 in the atg1 strain, and in this case, a transient increase in total Sae2 was observed (Fig. 7E). These results suggest that both autophagy and the proteasome contribute to the degradation of Sae2 multimeric complexes.

DISCUSSION

The Sae2 protein is an important component of the machinery that initiates DNA double-strand break resection in budding yeast (12, 13) and is the target of CDK phosphorylation, which limits 5′ strand resection to the S and G₂ phases of the cell cycle (5). In this study, we identified the phosphorylation events that occur on Sae2 in vivo and determined that the CDK modifications prime further modification by Mec1/Tel1 kinases that are essential for Sae2 activities in DNA damage survival. On the basis of our analysis of recombinant Sae2 in vitro and the properties of Sae2 in budding yeast, we propose that one of the primary functions of Sae2 phosphorylation is to transiently disrupt Sae2 from large, oligomeric, inactive forms into smaller active forms that promote DNA end resection and homologous recombination (see the model in Fig. 7F). The Sae2 that is released from the larger structures is also rapidly degraded through a combination of autophagy and proteasome-mediated pathways. Overall, this analysis provides evidence that posttranslational modifications are regulators of oligomerization and solubility, such that an inherently insoluble protein can be mobilized rapidly and reversibly to perform its functions.

Extensive phosphorylation of Sae2 upon DNA damage.

Our analysis of posttranslational modifications of Sae2 in vivo showed a much larger set of phosphorylation events than was anticipated. Many of these are not at SQ/TQ Mec1/Tel1 consensus sites; however, two of the functionally important sites match the consensus sequence and are also strongly dependent on the CDK phosphorylation of S267, which was previously shown to be important for DSB resection in vivo (5). We observed a hypomorphic phenotype with the S267A mutant in DNA damage sensitivity but a nearly null phenotype when S267, S134, and three of the other sites in the C terminus of the protein (S249, S278, and T279) were also mutated. These results suggest that there must be residual DNA damage-induced phosphorylation of these C-terminal sites in the absence of S267 phosphorylation. In contrast, the S267E mutant grew in the presence of CPT and MMS similarly to the wild-type strain, and the 2E3D phosphomimic allele exhibited an intermediate sensitivity when expressed from a low-copy-number vector.

Phosphorylation of Sae2 regulates its oligomeric state.

Our initial analysis of phosphomimic forms of MBP-Sae2 expressed in E. coli suggested that phosphorylation of Sae2 affects its oligomeric state. We confirmed this notion with Sae2 expressed without MBP in yeast and found that phosphorylation affects Sae2 solubility, as measured by two indices: the amount of protein observed in the soluble pool and the apparent size of soluble complexes measured by gel filtration. The increase in soluble Sae2 occurs in a DNA damage-dependent way and is very rapid upon release of cells into S phase. Analysis of the nonphosphorylatable or phosphomimetic mutations suggested that these sites control Sae2 changes in solubility, with the alanine mutations largely blocking the transitions to more soluble forms and the phosphomimetic mutations promoting their formation.

The absolute concentration of a protein is clearly important when considering solubility and oligomerization. We found that the expression level of Sae2 is extremely low in wild-type cells, much lower than the estimate previously published in a global expression study (36). Our analysis of Sae2 levels in yeast cells using quantitative protein blotting showed that there are ∼100 molecules of the protein per cell when expressed from the chromosome, ∼280 when expressed from a low-copy-number CEN plasmid, and ∼2,000 when expressed from a high-copy-number 2μ plasmid. The measurements of Sae2 solubility by protein blotting required use of high-copy-number expression strains, but analysis of Sae2 levels in a low-copy-number expression strain by mass spectrometry also confirmed transitions in solubility during DNA damage exposure (Fig. 6), as did the analysis of the amount of soluble Sae2 after DNA damage when it is expressed from the chromosome. In addition, the changes in apparent size distribution measured by gel filtration were also observed with protein isolated from low-copy-number and chromosomal expression strains (Fig. 5). Thus, the effects on solubility are not limited to strains with high-copy-number Sae2 expression. In fact, the mutations in phosphorylated residues consistently have a much more dramatic effect when expressed at a low copy number or from the chromosome than when expressed at a high copy number, suggesting that the endogenous pool of Sae2 protein is strongly dependent on these modifications. It is possible that the ∼100 molecules of Sae2 that are present in normal cells are more dependent on the modifications because a minimum level of active Sae2 must be reached for efficient DNA repair. A high-copy-number expression strain generates higher levels of insoluble protein (and larger soluble complexes) than strains with endogenous levels of Sae2, but it also generates a larger pool of smaller oligomers, in theory making the protein less dependent on the phosphorylation-induced transitions.

The observation that L25P Sae2 exists in yeast as a monomer, similar to its behavior when expressed in E. coli (30) (see Fig. S5 in the supplemental material; data not shown), indicates that Sae2-Sae2 interactions are important for oligomeric complex formation. Nevertheless, we do not know if the complexes are homogeneous in yeast. It is also possible that Sae2 phosphorylation releases it from a large oligomer that includes other proteins but requires Sae2 dimerization for binding.

The apparent size of DNA damage-induced Sae2 complexes in yeast is closer to that of a dimer than to that of a monomer (with the caveat that gel filtration is affected by shape as well as by protein size). We previously showed that monomeric MBP-Sae2 exhibits a higher specific activity than dimeric complexes (15) and also showed that here (Fig. 1); however, dimeric Sae2 is also significantly more active than oligomeric complexes. On the basis of the evidence that we have from Sae2 in yeast and from in vitro phosphorylation of oligomeric Sae2 complexes in vitro, it is possible that the active form of Sae2 is a complex with the approximate size of a dimer.

In addition to the effects on oligomeric transitions, there are also likely to be effects of phosphorylation on the functions of Sae2. This is evident from the observation that the proteins with phosphomimic mutations (for instance, the 2E3D mutant) are more soluble and form smaller complexes yet do not fully complement the functional defects seen in the sae2-null strain. These properties may relate to the interactions between phosphorylated Sae2 and DNA or between Sae2 and other protein complexes found at the break site. We also note that the S267A protein exhibits obvious defects in promoting resection in vitro compared to the wild-type or phosphomimic form, even though the wild-type MBP-Sae2 protein made in E. coli is not phosphorylated (Fig. 1). This may indicate a subtle difference in conformation with this mutation that affects its interactions with other proteins, since it behaves similarly to wild-type protein in nuclease assays (see Fig. S1 in the supplemental material). A conformational defect in the S267A protein may also underlie its phenotype relative to that of S267E, a difference that may not be very evident in focus recruitment kinetics but very obvious in DNA damage survival assays over a longer time frame.

The dynamics of Sae2 during DNA damage are reminiscent of those of the yeast Rad9 protein, a checkpoint protein that was shown to exist as a large, ∼850-kDa protein complex in normally growing cells that converts to a smaller ∼560-kDa complex after DNA damage exposure (37). Rad9 is not an enzymatic component of the DNA repair machinery but is responsible for recruiting and activating Rad53, an important checkpoint kinase. In this case, hyperphosphorylation of Rad9 has been shown to correlate with conversion of the larger complex to the smaller complex and binding of the Rad53 protein (37). DNA damage-induced transitions between inactive and active oligomeric forms may be a common feature of regulation in the DNA damage response, considering that many of the initial events occur very rapidly in the absence of de novo gene expression and are reversibly regulated by posttranslational modifications.

DNA damage-induced degradation of Sae2.

The appearance of more Sae2 in the soluble fraction occurs concomitantly with the degradation of ∼20% of the protein during replication in the presence of DNA damage. This damage-induced loss of protein takes place primarily through autophagy, as seen in an atg1 deletion strain, although inhibition of proteasome function further increases Sae2 protein levels. The previously reported involvement of autophagy in Sae2 degradation (35) is consistent with our evidence for large oligomeric complexes since this pathway is primarily responsible for removal of damaged organelles and large aggregates (34). Here we identified two lysine residues as targets for the acetylation that signals Sae2 to the autophagy machinery (K239 and K266). Preventing phosphorylation of Sae2 at S267 by mutation did not block either its acetylation or its degradation (data not shown); thus, the degradation is not strictly dependent on the CDK-induced phosphorylation events, even though it is damage induced. At this point, it is not known which enzymes acetylate or deacetylate Sae2, although this is certainly an interesting area for future study.

Posttranslational modifications as regulators of protein oligomeric transitions.

It is clear that all polypeptides have some propensity to aggregate, although the amino acid sequence of most proteins, particularly abundant factors, evolves away from this outcome (38, 39). Single amino acid mutations often dramatically decrease the solubility of proteins, in some cases with consequences for disease (40). Aggregation is usually viewed as an unfavorable activity, particularly because the accumulation of aggregated proteins is toxic during stress conditions and aging (41), and is also associated with unfolded or misfolded proteins that are destined for degradation. However, there are notable instances where the self-association of a polypeptide is beneficial: for instance, to form functional intracellular structures such as stress granules, to create new extracellular fibers that mediate interactions with the environment, or to promote programmed cell death (42). Transitions between different homooligomeric states have also been noted as a versatile mechanism of enzymatic regulation in many biological contexts (43).

Here we propose that the oligomerization potential of Sae2 has evolved in order that the potentially damaging effects of this endonuclease are not present when it is in large inactive complexes yet can be released rapidly without the need for de novo expression. The data do not support the idea that Sae2 is simply misfolded and segregated into nonfunctional aggregates but, rather, support the idea that functional Sae2 can be liberated from these large complexes. A similar situation may exist with human CtIP, despite the low amino acid conservation between these functional orthologs. CtIP plays a role in DSB resection very similar to that of Sae2 in budding yeast and is already known to be modified by CDK and by ATM/ATR (44). Interestingly, the prolyl isomerase Pin1 was shown to regulate the degradation of CtIP through ubiquitination in human cells (45), but it is unknown whether a similar relationship might exist with Sae2 in budding yeast. The example shown in this study with Sae2 may be a model for many other enzymes in the DNA damage response or in other cellular processes, where the availability of a protein is controlled reversibly at the posttranslational level by altering its oligomeric state.