Histone acetyltransferase 1 is dispensable for replication-coupled chromatin assembly but contributes to recover DNA damages created following replication blockage in vertebrate cells

BBRC Biochemical and Biophysical Research Communications 345 (2006) 1547–1557 www.elsevier.com/locate/ybbrc

Histone acetyltransferase 1 is dispensable for replication-coupled chromatin assembly but contributes to recover DNA damages created following replication blockage in vertebrate cells Hirak Kumar Barman a,c, Yasunari Takami a, Tatsuya Ono d, Hitoshi Nishijima d, Fumiyuki Sanematsu a, Kei-ichi Shibahara d, Tatsuo Nakayama a,b,* b

a Section of Biochemistry and Molecular Biology, Department of Medical Sciences, Miyazaki Medical College, Japan Department of Life Science, Frontier Science Research Center, University of Miyazaki, 5200 Kihara, Kiyotake, Miyazaki 889-1692, Japan c Section of Genetics and Biotechnology, CIFA, ICAR, Bhubaneswar, India d Department of Integrated Genetics, National Institute of Genetics, 1111, Yata, Mishima, Shizuoka 411-8540, Japan

Received 8 May 2006 Available online 23 May 2006

Abstract Histone acetyltransferase 1 (HAT1) is implicated for diacetylation of Lys-5 and Lys-12 of newly synthesized histone H4, the biological significance of which remains unclear. To investigate the in vivo role of HAT1, we generated HAT1-deficient DT40 clone (HAT1/). HAT1/ cells exhibited greatly reduced diacetylation levels of Lys-5 and Lys-12, and acetylation level of Lys-5 of cytosolic and chromatin histones H4, respectively. The in vitro nucleosome assembly assay and in vivo MNase digestion assay revealed that HAT1 and diacetylation of Lys-5 and Lys-12 of histone H4 are dispensable for replication-coupled chromatin assembly. HAT1/ cells had mild growth defect, conferring sensitivities to methyl methanesulfonate and camptothecin that enforce replication blocks creating DNA double strand breaks. Such heightened sensitivities were associated with prolonged late-S/G2 phase. These results indicate that HAT1 participates in recovering replication block-mediated DNA damages, probably through chromatin modulation based on acetylation of Lys-5 and Lys-12 of histone H4.  2006 Elsevier Inc. All rights reserved. Keywords: HAT1; H4 acetylation; DNA repair; DT40; Camptothecin; Methyl methanesulfonate

In eukaryotes, acetylation of N-terminal tails of core histones is thought to change chromatin configuration for transcriptional activation. Such acetylation is catalyzed by histone acetyltransferases (HATs), which are classified into two distinct groups (types A and B) based on their subcellular localization and substrate specificity. Type A HATs (e.g., GCN5, PCAF, ESA1, TIP60, etc.) are nuclear Abbreviations: HAT, histone acetyltransferase; MMS, methyl methanesulfonate; CPT, camptothesin; kb, kilo base; SDS, sodium dodecyl sulfate; PAGE, polyacrylamide gel electrophoresis; PBS, phosphate-buffered saline; BSA, bovine serum albumin; PCR, polymerase chain reaction; EtBr, ethidium bromide; UV, ultraviolet; IR, ionizing radiation; SSB, single strand break; DSB, double strand break. * Corresponding author. Fax: +81 985 85 6503. E-mail address: [email protected] (T. Nakayama). 0006-291X/$ - see front matter  2006 Elsevier Inc. All rights reserved. doi:10.1016/j.bbrc.2006.05.079

enzymes. Their crucial roles as transcriptional coactivators by acetylating chromatin bound histones at respective gene loci, as well as participations in replication, repair, and other chromatin related processes, have been established [1–4]. On the other hand, type B HATs are originally purified from cytoplasmic fraction and considered to be involved in the rapid acetylation of newly synthesized cytoplasmic histones H3 and H4 to be imported into nucleus for de novo deposition onto nascent DNA chains [5,6]. Lys residues at 5 and 12 of histone H4 are acetylated in an evolutionarily conserved pattern [7,8], while an acetylation pattern for histone H3 varies from species to species [8,9]. These acetylated histones H3 and H4 are associated with histone chaperones, chromatin assembly factor-1 (CAF-1) and anti-silencing function 1 (ASF1), and loaded

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onto newly replicated DNA [10–13]. After completion of nucleosome assembly, acetylated forms of histones H3 and H4 in the nucleosomes are rapidly deacetylated to form mature chromatin [14,15]. However, the exact mechanism and biological significance of such transient acetylation and deacetylation cycle are poorly understood. Among type B HATs, HAT1 is proposed as a candidate enzyme to diacetylate Lys-5 and -12 residues of newly synthesized histone H4 based on its in vitro acetylating abilities at Lys-5 and -12 residues of free H4 protein [6,16–18]. HAT1 is tightly bound with its regulatory subunit HAT2 in yeast and with the homolog p46 (or RbAp48) in vertebrates, which enhances HAT1 activity by facilitating its binding ability as to H4 protein [17,19– 21]. Recently, in yeast another type B HAT, termed HATB3.1, in a GCN5-containing complex with ADA3 but not ADA2, was identified as histone H3 specific [22]. Distinct acetylation pattern of cytosolic histones by type B HATs has been thought to play a certain role in de novo chromatin assembly. However, genetic analyses in yeast have created controversy, since deletion of either HAT1–HAT2 or GCN5 gene resulted in no apparent cell growth deviations [16,17,20,23]. These findings have been explained by functional redundancy between N-terminal tails of histones H3 and H4 [23,24]. In fact, deletion of the N-terminal tail of either histone H3 or H4 is viable, but combined deletions of N-terminal tails of histones H3 and H4 result in defects in both nucleosome assembly and cell growth [23–25]. Moreover, while the HAT1 deletion alone produces no deviated phenotype, combined mutations to HAT1 plus different Lys residues at N-terminal tail of histone H3 cause defects in both telomeric gene silencing and resistance to methyl methanesulfonate (MMS) [26,27]. Such a sensitivity to MMS resulted primarily from a defect in recombinational repair of double strand breaks (DSBs) through ASF1-dependent chromatin assembly [27]. To gain further insight about physiological roles of HAT1 and/or impact of HAT1 mediated acetylation level in vertebrate cells, we established HAT1 deficient DT40 clones, using the gene targeting technique. Analyses of the mutant cells indicated that HAT1 is not essential for cell viability, even though it primarily acetylates Lys-5 and Lys-12 of post-translated cytosolic histone H4. We also presented evidences, suggesting that HAT1 participates in repair of DNA DSBs caused by replication blocking agents, such as camptothecin (CPT) and MMS, in vertebrates. Materials and methods Plasmid constructs. Genomic DNA clone of chicken HAT1 was isolated by screening DT40 kFIX II genomic library [28], using chicken HAT1 cDNA as a probe [21]. To obtain HAT1 disruption constructs, 4.7 kb upstream fragment, carrying exon 3, excised by HindIII/EcoRI double-digestion and 4 kb downstream fragment, bearing exons 7–9, amplified by PCR from the HAT1 genomic clone were inserted into pBluescript vector (Stratagene). Hygromycin B (hyg) and puromycin

(puro) resistance cassettes, driven by b-actin promoter and flanked by loxP sites, were inserted independently between the upstream and downstream arms [29]. Gene targeting by these constructs (pDHAT1loxhyglox and pDHAT1loxpurolox) was expected to exchange exons 4–6 of the HAT1 gene with selection marker genes. The HAT1 tet-responsible expression vector (ptetHA-HAT1) was constructed by inserting HA-tagged fulllength HAT1 cDNA into pUHD10-3 plasmid [30]. To obtain ptTA-bleo construct, a cassette of bleomycin-resistance gene driven by b-actin promoter was cloned into pUHD15-1 plasmid, carrying tet-responsive transactivator gene (tTA) controlled by CMV promoter [31]. Cell culture, transfection, and mutant isolation. Cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum, 1% chicken serum, streptomycin sulfate (10 lg/ml), and penicillin (100 U/ml) at 37 C. Transfection and selection of drug resistant clones were performed as described [29]. To determine growth rate, the cells were counted at indicated times. Flow cytometric analysis. Cells were fixed with 70% ethanol at 20 C, washed with PBS–0.2% BSA, and stained with 5 lg/ml propidium iodide (PI) plus 100 lg/ml RNase in PBS–0.2% BSA. Stained cells were analyzed by FACS Calibur analyzer (Becton–Dickinson, USA). Antibodies. Chicken full-length HAT1 cDNA was cloned into pGEX2TK (Amersham Pharmacia Biotech). GST-tagged fusion proteins were expressed in Escherichia coli BL21 and purified by glutathione-conjugated beads. Purified GST-HAT1 protein was injected into rabbit for immunization. IgG fractions were passed through GST-immobilized beads column, and then anti-HAT1 antibody was further affinity-purified using GST-HAT1-immobilized beads column. Rabbit anti-H3 (Abcam, UK), -H4 (pancreatic) (Upstate, USA), -site specific acetylated Lys of histone H4 (Upstate, USA), -Chk1 and -phosphoserine 345 of Chk1 (Cell Signaling Technology) antibodies, and mouse monoclonals of anti-cH2AX (Upstate, USA) and -a-tubulin antibodies (Sigma, USA) were used. Western blotting. To obtain cellular proteins, cells were suspended in two volumes of hypotonic buffer (HB) (20 mM Hepes, pH7.5, 5 mM KCl, 1.5 mM MgCl2, 0.5 mM dithiothreitol, 5 mM sodium butyrate, and proteinase inhibitor cocktail (Sigma)), homogenized, and centrifuged at 3000 rpm. Supernatant was further centrifuged at 15,000 rpm, and then resultant supernatant was diluted in sodium dodecyl sulfate (SDS) sample buffer (final concentration to 60 mM Tris, pH 6.8, 2% SDS, 100 mM dithiothreitol, and 10% glycerol), followed by boiling, and used as cytosolic fraction. The nuclei pelleted by initial low centrifugation were washed with HB containing 0.1% Triton X-100 once, suspended in SDS sample buffer followed by boiling, and used as nuclear fraction. To obtain total cell extracts, washed cells (2 · 105) were suspended in 50 ll SDS sample buffer and boiled. The protein samples were subjected to SDS–10% or 14% PAGE, electro-transferred onto polyvinylidene difluoride (PVDF) membrane, and blocked with 5% skim milk before incubation with primary antibodies. Antibody binding was detected using corresponding secondary antibodies conjugated with horseradish peroxidase (Dako, Denmark) and Supersignal detection kit (Pierce, USA), and analyzed with a Luminescent Image Analyzer LAS-1000 plus (Fujifilm, Japan). Immunofluorescence microscopy. Cells (2 · 105) were spotted onto glass slides, fixed with 2% paraformaldehyde in PBS for 15 min, and immersed in cold methanol for 30 min. After washing with 0.5% Triton X-100, cells were probed with rabbit anti-HAT1 antibody. Primary antibody was detected by Alexa 488-conjugated goat anti-rabbit secondary antibody (Molecular Probes, Eugene). DNA was counterstained with 4 0 -6-diamidino-2-phenylindole (DAPI) at 0.1 lg/ml. Stained samples on slides were examined under Axiovert M-200 fluorescence microscope (Zeiss, Germany), and then images were captured with cooled CCD camera (ORCA-ER, Hamamatsu, Japan). SV40 DNA replication-coupled nucleosome assembly reaction. SV40 DNA replication-coupled nucleosome assembly reaction was performed as described [32,33]. After completion of DNA replication reaction at 45 min, 50 ll of the reaction mixture was applied to a column of Sepharose CL-4B beads (Pharmacia), followed by centrifugation at 400g for 2 min, and then the effluent, containing semi-purified DNA, was used for a supercoiling assay. The effluent was mixed with CAF-1 purified from HeLa cells expressing FLAG-CAF-1p150, cytosolic extracts from either DT40 or

H.K. Barman et al. / Biochemical and Biophysical Research Communications 345 (2006) 1547–1557 HAT1/ cells, topoisomerase I (13 ng), topoisomerase II (100 ng), and histones H2A and H2B (420 ng) purified from HeLa cells. After the reaction was terminated at 45 min, DNA was treated with RNase and proteinase K, and subjected to agarose gel electrophoresis. Details were as described in the previous reports [33,34]. Cytosolic extracts (S100) from DT40 cell lines were prepared as described [35]. Briefly, cells were swollen in the hypotonic buffer (20 mM Hepes, pH 8.0, 5 mM KCl, 1.5 mM MgCl2, 0.1 mM DTT, and protease inhibitors), homogenized by a Dounce homogenizer, and centrifuged at 20,000g for 20 min twice. We used the resultant supernatant as cytosolic extracts S100. Analysis of chromatin structure in vivo. To examine chromatin structure, cells were pulse-labeled with [3H]thymidine (20 lCi/ml) for 5 min and divided into two portions. One was used as pulse-labeled sample, and the other was washed with PBS twice, followed by culturing in medium without [3H]thymidine for 60 min to use as pulse-labeled and chased samples. Each cell sample was washed with cold PBS, suspended in NB (10 mM Tris–HCl, pH 7.5, 0.1 mM EDTA, 2 mM magnesium acetate, 2 mM CaCl2, and 1 mM DTT) plus protease inhibitor cocktail (Sigma) plus 10% sucrose, and incubated in the presence of 0.1% Nonidet P-40 (NP-40) for 5 min on ice. The resultant nuclei were washed with NB twice, suspended at 10 A260/ml (A260 was measured in 2 M NaCl and 5 M urea) in NB, and digested at 37 C for 8 min with 0.0074–0.2 U/ml of micrococcal nuclease (MNase) (Sigma). The reaction was stopped by adding 10 mM EDTA and 0.5% SDS, and then DNA was purified by incubation with 100 lg/ml proteinase K at 37 C for 2 h, followed by phenol–chloroform extraction and ethanol precipitation. DNA was electrophoresed in 1.2% agarose gel and transferred to Hybond N+ membrane. To detect 3 H-labeled DNA, blot was directly exposed to a BAS-TR204S imaging screen specific for tritium and visualized using a Mac BAS-1000 (BAS, Fuji Film, Tokyo, Japan). Colony formation assay. For UV irradiation, cells in PBS were irradiated with various doses (UV cross linker, Stratagene). X-ray irradiation was performed using MBR-1520R radiator set (Hitachi) at 150-kVp, 20 mA, 0.5 mm aluminum and 0.9 mm copper filtrations with a dose rate of 1 Gy/min. Serially diluted cells were plated in duplicate onto six-well plates containing 5 ml/well of 1% methylcellulose (MC) (Sigma–Aldrich, USA) in DMEM supplemented with 15% FBS and 1.5% chicken serum [36,37]. To test sensitivities to MMS (Nacalai Tesque, Japan), camptothecin (Sigma–Aldrich, USA), and etoposide (Sigma–Aldrich, USA), serially diluted cells were plated in MC plates, containing various doses of each drug. Plates were incubated at 37 C for 8–10 days, and resulting visible colonies were counted. Percentage survival was estimated relative to numbers of colonies from untreated cells.

Results Generation of HAT1/ cells, their growth kinetics, and subcellular localization of HAT1 To investigate the in vivo role of HAT1, we first isolated chicken genomic DNA for HAT1 by a library screening, designed two gene targeting constructs, and generated heterozygous (HAT1/+) and homozygous (HAT1/) DT40 mutant cells by sequential transfection of the hyg and puro constructs, respectively (Fig. 1A). The mutant clones were isolated and verified by Southern blot analysis showing expected positive bands with outer probe A and internal probe B (Fig. 1B), and the loss of HAT1 protein in these clones was further confirmed by Western blotting (Fig. 2B). The viability of HAT1/ cells validated that HAT1 is dispensable for cell proliferation. Next, we examined the proliferative rate and cell cycle distribution of HAT1/ cells by flow cytometric analysis. The growth rate of HAT1/ cells was slightly retarded

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with slightly enhanced cell death (data not shown), compared with that of wild-type DT40 cells (Fig. 1C). Distribution of the mutants in each cell cycle stage was not significantly different from that of DT40 cells (Fig. 1D). However, the more detailed analyses for the cell cycle progression after synchronization showed a slight delay in late S/G2 phase in HAT1/ cells (see Fig. 5B). Thus, the slower proliferation rate of HAT1/ cells could be explained by the prolonged cell cycle time. Although HAT1 was originally identified from cytosolic extracts of yeast [17,38], Drosophila [7], and human cells [6], it was also isolated from nuclei in several species [19,20,39,40]. To know subcellular localization of HAT1 in DT40 cells, we conducted immunofluorescence imaging, using antibody against GST-HAT1 fusion protein raised by us. As shown in Fig. 1E (upper panel), HAT1 was present predominantly in nuclei and lesser in cytoplasm of DT40 cells, consistent with the previous result obtained based on forced expression of HAT1 in COS7 cells [19], and then as expected, no signal was detected in HAT1/ cells (Fig. 1E; lower panel). HAT1 deficiency reduces diacetylation levels at Lys-5 and Lys-12 of histone H4 To know the in vivo catalytic role of HAT1 for core histone proteins, we compared acetylated states of histone H4 in DT40 and mutant cells by Western blotting, using anti-site specific acetyl antibodies for histone H4. Cells were fractionated into cytosolic and nuclear fractions, which were rich in soluble pre-deposited (de novo) histones and chromatin-bound histones, respectively. Using two types of antibodies which can recognize non-acetylated and acetylated H3 or H4 proteins, respectively, we showed that total amounts of histone H3 or H4 proteins remained constant between wild-type and mutant cells (Fig. 2A). Consistent with previous observations in other organisms [6,8], the acetylation levels of Lys-5 and Lys-12 (K5/K12) residues of histone H4 in the cytosolic fraction were significantly higher than those in the nuclear fraction in DT40 cells (Fig. 2A), indicating that large portions of soluble histone H4 exist as diacetylated (K5/K12) isoform. However, importantly, in HAT1/ cells, the acetylation levels of K5 and K12 residues of histone H4 in the cytosolic fraction were greatly diminished (Fig. 2A), whereas the acetylation levels of Lys-8 and Lys-16 residues (K8/K16) were not changed, suggesting that diacetylation of K5 and K12 of cytosolic histone H4 is dependent on the HAT1 activity in vertebrate cells, and extending earlier implication that HAT1 diacetylates newly synthesized histone H4. Additionally, our results revealed that the acetylation level of Lys-5 residue of nuclear histone H4 was reduced in HAT1/ cells, as compared with that in DT40 cells (Fig. 2A). To confirm these results, we next generated the HAT1/ clone, carrying HA-tagged HAT1 under the control of tet-repressive promoter, HAT1/ ptetHA-HAT1. In the absence

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Fig. 1. Gene targeting of HAT1 locus, growth kinetics, and subcellular localization of HAT1. (A) Schematic representation of two targeting constructs (top), intact (middle) and targeted (bottom) HAT1 alleles. The loxP sequences flanking hyg and puro genes are shown as triangles. Exons are indicated by opened boxes. Probe A or B used is indicated by the horizontal line. Only relevant restriction sites (B, BamHI site) and possible relevant fragments with their lengths in kb are shown. (B) Southern blot analysis of BamHI digested genomic DNAs of wild-type (+/+), HAT1/+ (/+) and HAT1/ (/) clones using outer probe A or internal probe B indicated in (A). (C) Growth curves of DT40 and HAT1/ cells. Data represent means of three independent experiments. (D) Flow cytometric analysis of asynchronously grown DT40 and HAT1/ cells. Cells were fixed and stained with PI to detect total DNA (x-axis, linear scale). (E) Immunofluorescence imaging of HAT1. Antibody for GST-HAT1 fusion protein was used (a,d). (b,e) DAPI stained nuclei, (c,f) merge.

of tet, HA-HAT1 was expressed in this clone at almost similar level to that of HAT1 in DT40 cells, but HA-HAT1 was completely disappeared by day 2 after the addition of tet (Fig. 2B). When examined the acetylation states at particular Lys residues of histone H4 in this clone, in the absence of tet,

the acetylation levels for Lys-5, Lys-8, Lys-12, and Lys-16 residues (K5, K8, K12, and K16) were found to be almost similar to the levels of those in wild-type cells. However, following the tet addition, the acetylation levels of K5 and K12 were greatly reduced in the cytosolic fraction, and the

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Fig. 2. Effects of HAT1 deficiency on site-specific acetylation levels of histone H4 and expression of HAT1 protein. (A) Cytosolic and nuclear fractions from indicated cell lines were prepared as described under Materials and methods. Protein samples were subjected to SDS–14% PAGE and analyzed by Western blotting using appropriate antibodies marked at the right side. Middle and right panels: equal amounts of histone H4 proteins from nuclear and cytosolic fractions, respectively, which correspond to 3 · 105 and 3 · 106 cells, were loaded. Left panel: the nuclear fraction corresponding to 1.5 · 106 cells was loaded. HAT1/ptetHAHAT1 clone was obtained by co-transfection of HAT1/ cells with ptetHA-HAT1 and ptTA-bleo plasmids. The + tet indicates culturing cells in the presence of tet for 4 days. (B) Total cell proteins from indicated cell lines were subjected to SDS–10% PAGE, and analyzed by Western blotting, using anti-chicken HAT1 antibody.

acetylation level of K5 was also decreased in the nuclear fraction (Fig. 2A). These results indicate not only that HAT1 is a primary enzyme for diacetylation of K5 and K12 of cytosolic histone H4, but also that it maintains acetylation state for K5 of chromatin-bound histone H4. HAT1 catalyzed diacetylation of K5 and K12 of histone H4 is not necessary for replication-coupled chromatin assembly Replication-coupled chromatin assembly is primarily catalyzed by CAF-1 that is associated with PCNA and acetylated histones H3 and H4 [11,33,41]. However, our results suggested that HAT1 and/or diacetylation of K5 and K12 of histone H4 are not essential for cell viability, suggesting their dispensability for replication-coupled chromatin assembly. To address whether HAT1 and/or the diacetylation of K5 and K12 of de novo histone H4 have any influence on the CAF-1 dependent nucleosome assembly onto replicated DNA, we performed the in vitro two-step chromatin assembly assay [34], using cytosolic extracts (S100) from wild-type or HAT1/ cells as histone sources and semi-purified pre-replicated SV40 plasmid DNA as template, with or without purified CAF-1

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(Fig. 3A). In the presence of CAF-1, both the S100 fractions effectively induced supercoiling of replicated SV40 DNA in a dose dependent manner with almost equal efficiency (Fig. 3A; compare lanes 6–8 vs 12–14). Even in the absence of CAF-1, large amounts of S100 fractions of wild-type and mutant cells could convert a part of replicated DNA into negatively supercoiled form I, possibly due to a trace of CAF-1 activity in the S100 fractions (Fig. 3A, lanes 5 and 11). But the efficiency of supercoil formation was also the same in the presence or absence of HAT1 (Fig. 3A; compare lanes 3–5 vs 9–11). Thus, these results indicate that HAT1 and/or diacetylation of K5 and K12 of histone H4 are not the limiting factor for the CAF-1 dependent nucleosome assembly onto replicated DNA. Next, to examine the replication-coupled nucleosome assembly in the absence of HAT1, we analyzed nascent chromatin structure by labeling DNA with [3H]thymidine, followed by MNase digestion. DT40 and HAT1/ cells were pulse-labeled with [3H]thymidine for 5 min and divided into two portions, one of which was chased in [3H]thymidine free medium for 60 min. Equal numbers of nuclei obtained were subjected to MNase digestion assay. Bulk DNA stained by EtBr showed same sensitivity to MNase digestion in wild-type and HAT1/ cells (Fig. 3B; upper panel), indicating that the loss of HAT1 has insignificant influence on the global chromatin structure. When pulselabeled chromatin was examined, MNase digested chromatins from wild-type and HAT1/ cells showed indistinguishable patterns of characteristic nucleosome ladders (lanes 1–4 vs 6–9 of lower panel of Fig. 3B). Likewise, pulse-labeled and chased chromatins from both cell lines also exhibited same sensitivity to MNase digestion (lanes 11–14 vs 16–19 of lower panel of Fig. 3B). These results highlighted that the HAT1 deficiency has no effects on both the global and nascent chromatin structures. Thus, the in vitro and in vivo results, together, indicate not only that both diacetylated histone H4 at K5 and K12 residues and non-acetylated histone H4 could be deposited onto newly replicated DNA with an equal efficiency, but also that the HAT1 depletion itself does not affect the replication-coupled nascent chromatin assembly process as well as overall chromatin structure. HAT1/ cells are moderately sensitive to CPT and mildly sensitive to MMS To further explore the in vivo function of HAT1 and/or the biological significance of diacetylation of K5 and K12 of histone H4, we examined the effects of several DNA damaging agents on HAT1 deficient cells. HAT1/ cells were mildly sensitive (5-folds) to alkylating agent MMS (Fig. 4A) and moderately sensitive (15-folds) to chemotherapeutic agent CPT, a topoisomerase-I (TopI) inhibitor (Fig. 4B), but not sensitive to UV irradiation, creating single strand breaks (SSBs) (Fig. 4C), which are repaired by nucleotide excision repair (NER) pathway [2,42]. Further, HAT1/ cells did not show any increased

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Fig. 3. HAT1 deficiency has no influence on replication-coupled chromatin assembly. (A) CAF-1 dependent in vitro two-step nucleosome assembly assay. The SV40 plasmid DNA replication was performed in the presence of [32P]-dATP and HeLa cell extract. Using replicated SV40 DNA, nucleosome assembly reaction was performed with or without recombinant CAF-1 and S100 extracts obtained from DT40 or HAT1/ cells (lanes 3–14). The reaction products were deproteinized and resolved in 1.0% agarose gel. Upper panel shows the radiolabeled DNA in autoradiogram, and lower panel shows bulk DNA stained with EtBr. Positions of form I, form II, and replication intermediates or unresolved catenates (*) are indicated at the left. (B) MNase digestion assay of chromatin of HAT1 deficient cells. Pulse-labeled DT40 and HAT1/ cells with [3H]thymidine (pulse), and pulse-labeled and chased DT40 and HAT1/ cells (chase) were used. Nuclei were prepared soon after pulse-labeling or after 60 min chase period. Same numbers of isolated nuclei were treated with MNase at 0.2, 0.068, 0.022, 0.0074, and 0 U/ml. Purified DNAs were resolved in 1.2% agarose gel, stained with EtBr (upper panel), transferred onto Hybond N+ membrane, and autoradiographed (lower panel). Positions of nuclease-resistant mono, di, and tri-nucleosomal DNA fragments (right) and markers of 100-bp DNA ladder (left) are indicated.

sensitivity to IR irradiation (Fig. 4D) and etoposide (VP16), a topoisomerase-II (TopII) inhibitor (Fig. 4E). Both of IR-irradiation and VP-16 create DSBs, which are preferentially repaired by non-homologous end joining (NHEJ) pathway in DT40 cells [37,43]. The heightened sensitivity of HAT1/ cells towards CPT and MMS was suppressed by the introduction of ptetHA-HAT1 in the absence of tet, while in the presence of tet, with diminishing the HA-HAT1 expression (Fig. 2B), the cells became sensitive to these two agents (Fig. 5A and B), validating the increase in sensitivity to CPT and MMS was related to the lack of the HAT1 function. MMS has pleiotropic effects generating variety of lesions of SSBs and DSBs [44]. CPT inhibits religation step of Top-I activity, resulting in single-strand DNA nicks [45]. Following replication, these SSBs are converted into DSBs [46]. Therefore, we consider that HAT1 and/or diacetylation of K5 and K12 of histone H4 contribute to cellular tolerance to DSBs, particularly those induced by replication blocks.

HAT1/ cells exhibit functional S and G2 phase checkpoint It is possible that the increased lethality to MMS and CPT of HAT1/ cells could be due to defects in the checkpoint function in response to DNA damages [47,48]. To assess this possibility, we first examined the cell cycle distribution after treatment with MMS or CPT of asynchronous wild-type and HAT1/ cells. Upon exposure to MMS (15 lg/ml), both the wild-type and mutant cells showed S phase delay at 4 h, accumulation in S phase at 12 h, and finally G2 phase arrest by 24 h (left panel of Fig. 5A). Likewise, CPT exposure (40 nM) induced evident S phase delay and G2 arrest during 4–8 h in both cell lines (right panel of Fig. 5A). These results suggest that HAT1/ cells should retain functional checkpoint responses of S and G2 phases. Moreover, HAT1/ cells treated with CPT showed stronger G2/M phase arrest after 8–12 h, which was more prominent at 24 h, while DT40 cells resumed cell cycle progression at these time points (right panel of Fig. 5A).

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Fig. 4. Sensitivity of DT40 and HAT1/ cells to DNA-damaging agents. Colonogenic survival curves of DT40 and HAT1/ cells exposed to MMS (A), CPT (B), UV light (C), IR (D), and VP-16 (E). Serially diluted cells (50, 500, and 5000 cells) were plated in duplicate in MC media containing various doses of either MMS or CPT or VP-16 as described under Materials and methods. Cells in PBS and DMEM were irradiated with UV-light and X-ray, respectively, before plating into MC media. (A,B) Include data for HAT1/ ptetHA-HAT1 clone in the presence or absence of tet, indicating the HAT1dependent MMS and CPT lethalities. For tet treatment, HAT1/ ptetHA-HAT1 cells were cultured in the presence of tet for 2 days before plating into MC media containing tet. Data shown are means of three independent experiments.

These results were further confirmed by analyzing G1-synchronized cells. As shown in Fig. 5B, after release from the block at G1 phase with mimosine, the CPT treatment led to the delay in late S/G2/M phase in both cell lines, and the delay was more pronounced in HAT1/ cells. The mitotic index estimation suggested that such G2/M phase delay should be due to arrest at G2 phase (data not shown). Thus, HAT1/ cells seemed to be blocked at G2 phase stronger than DT40 cells, when exposed to CPT. Since Chk1 phosphorylation is necessary for the S phase specific checkpoint activation in response to DNA damage [47], we investigated the status of Chk1 phosphorylation in DT40

and HAT1/ cells. Western blot analyses showed that the MMS treatment efficiently induced Chk1 phosphorylation in a time-dependent manner in both cell lines, but the induction level was comparatively lower in HAT/ cells. On the other hand, Chk1 activation mediated by CPT at 40 nM was not detected (Fig. 5C), but Chk1 phosphorylation could be seen in both cell lines treated with a higher dose (>100 nM) of CPT (data not shown). We next assessed phosphorylation of histone H2AX (cH2AX), which is one of the earliest responses to sites of DSBs [49]. The MMS treatment drastically increased the level of cH2AX in both cell lines (Fig. 5C). As in the

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Fig. 5. Effects of MMS and CPT on cell cycle progression. (A) Flow cytometric analyses of asynchronized cells harvested at indicated times after MMS (left) and CPT (right) treatments. (B) Flow cytometric analyses of synchronized cells. Cells were synchronized at G1/S boundary by sequential treatments of 0.5 lg/ml nocodazole for 4 h and 0.8 mM mimosine for 14 h. After release from the mimosine arrest, CPT (40 nM) was added to cultures, and then cells were harvested at 2 h intervals. (C) Cells were cultured in the presence of MMS (15 lg/ml) or CPT (40 nM) for indicated time periods, and whole cell extracts were prepared, followed by Western blotting, using antibodies against phosphorylated Ser 345 of Chk1 (p-Chk1-ser345), phosphorylated H2AX (cH2AX), and a-tublin as a loading control.

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case of Chk1 phosphorylation, the CPT treatment at 40 nM induced cH2AX at a very low level (Fig. 5C), whereas a higher dose (>100 nM) of CPT led to effective increase in cH2AX in both cell lines (data not shown). Thus, DNA damages after exposures to these two agents and/or immediate follow-up response to the resultant lesions were essentially same between DT40 and HAT1/ cells. These results, together, imply that the increased lethality of HAT1/ cells to MMS or CPT should be due to the accumulation of replication block induced DSBs that were hard to be repaired, but not to a catastrophic defect of checkpoint function. The relatively prolonged G2 arrest of CPT treated HAT1/ cells is presumably attributed to the persistence of extensive unrepaired DNA lesions. Discussion HAT1 has long been thought to function in acetylation of K5 and K12 residues at N-terminal domain of histone H4 that is probably linked with chromatin assembly process [5,11,17]. However, the compelling in vivo evidence has not been elucidated so far. We reported herein the first genetic evidence regarding the in vivo role of HAT1 in acetylation of newly synthesized histone H4 in vertebrate cells, i.e., markedly reduced diacetylation levels of K5 and K12 of histone H4 in the cytoplasm, which are dispensable for cell viability. Further, the lack of HAT1 and/or diacetylation of K5 and K12 of histone H4 did not much deviate chromatin assembly process for newly replicated DNA in vitro and in vivo, and then overall chromatin structure also remained unchanged in HAT1 deficient cells. These observations are mostly consistent with previous results that the N-terminal domains of histones H3 and H4 share redundant functions in nucleosome assembly in Saccharomyces cerevisiae [24,25] and are dispensable for CAF-1 mediated nucleosome assembly onto newly replicated DNA in vitro [34]. However, a slightly slower growth rate accompanied with a merely elevated spontaneous cell death of HAT1/ cells suggests that HAT1 and/or diacetylation of K5 and K12 of histone H4 might have some impact on growth kinetics of DT40 cells. In fact, we found that HAT1/ cells conferred elevated sensitivities to genotoxic agents CPT and MMS, as compared to wild-type cells, indicating that the degree of replicational interference by these two agents, especially by CPT, is relatively more in HAT1/ cells than in DT40 cells. On the other hand, the wild-type and mutant cells were equally resistant to UV, IR, and VP-16 exposures. Based on these mutagenspecific damage sensitivities of HAT1 deficient cells, such lethality is imposed specifically by DNA damage arising from DNA replication arrest. Since hypersensitivity to MMS and CPT has been reported to be correlated to defects in checkpoint response and/or DNA damage signaling [48,50], we determined whether HAT1 deficient cells properly activate checkpoint response after MMS and CPT exposures. Although HAT1/ cells exhibited slightly

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reduced responses to Chk1 and H2AX phosphorylations after MMS and CPT treatments, they were certainly arrested in S and G2 phases. Such arrest was more pronounced in mutant cells than in wild-type cells, and hence it could be excluded that the sensitivity of HAT1/ cells was accounted for by a defect in the checkpoint function. Possibly, prolonged G2 arrest of HAT1/ cells caused by MMS and CPT treatments could be simply explained by the presence of unrepairable DNA lesions. Soon after replication, nucleosomes behind the fork are highly acetylated, by loading with de novo acetylated histones. Such acetylated nucleosomal environment should be favorable for repairing DNA damages arising from replication stress, since open chromatin structure allows easy access for chromatin remodeling factors and subsequent recruitment of repair related proteins, and some acetylated marks of deposition related histones might be preferential recruitment sites for these proteins. However, in HAT1 deficient cells, the nucleosomal environment just behind the stalled fork may be unfavorable, because of the absence of deposition-related diacetylated K5 and K12 of histone H4. To circumvent this, presumably, recruitments of other HAT(s) to the stalled sites are required, and if so, this additional step might extend available times required for DNA repair, leading to the prolonged delay in G2 phase. There are several other possible involvements of HAT1 in repair process. One is that the HAT1 deficiency could cause impaired expression of repair-related genes, since acetylation of histone H4 has been implicated for positive transcription regulation and the acetylation level of K5 of chromatinbound histone H4 was lowered in HAT1/ mutant cells (Fig. 2A). However, this was unlikely, because we could find no transcription alterations in known repair-related genes tested (data not shown). The other possibility is that HAT1 plays an important role for restoration of chromatin after replication-mediated repair process. In S. cerevisiae HAT1/HAT2 contribute to telomeric gene silencing and repair of MMS-induced DNA damage, and then these functions are related to HIF1, which associates with HAT1/ HAT2 in nuclei to form a complex, exhibiting histone deposition activity [40]. In addition, HAT1-p48 and NASP, a human counterpart of HIF1, have been shown to coexist with histone H3.1- and H3.3-containing multi-protein-complexes, which possess replication-coupled and -uncoupled histone deposition activities, respectively [51]. If these multi-protein complexes function in chromatin restoration after replication-mediated DNA damage, the presence of HAT1 in such complexes provides a clue that its catalytic activity may modulate histone deposition activity after completion of repair process in higher eukaryotes. We showed that in vertebrate cells the HAT1-depletion alone could lead to the impairment of cellular tolerances, particularly for DNA damage caused by replication stall, mostly due to impaired repair-related chromatin configuration. Interestingly, in S. cerevisiae the effect of the HAT1depletion was only evident, when combined with additional mutations within N-terminal tail of histone H3 [27]. This

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difference may represent the complexity in regulation of chromatin assembly/modulation during or after replication and repair processes in higher eukaryotes more than in lower eukaryotes. The HAT1 deficient DT40 cell line will be of very useful to ascertain how it participates in replication-mediated repair process, by introducing combined mutations into other repair-associated genes. Acknowledgments We thank Ms. N. Yamamoto-Nagamatsu for technical assistance. This work is partly supported by 21st Century COE Program (Life Science) and Grant-in-Aid for Scientific Research from MEXT, and CREST from JST of Japan. H.K. Barman is a recipient of Monbukagakusho Scholarship. References [1] Y. Nakatani, Histone acetylases-versatile players, Genes Cells 6 (2001) 79–86. [2] C.L. Peterson, J. Cote, Cellular machineries for chromosomal DNA repair, Genes Dev. 18 (2004) 602–616. [3] A. Verger, M. Crossley, Chromatin modifiers in transcription and DNA repair, Cell. Mol. Life Sci. 61 (2004) 2154–2162. [4] H. van Attikum, S.M. Gasser, The histone code at DNA breaks: a guide to repair? Nat. Rev. Mol. Cell. Biol. 6 (2005) 757–765. [5] P.D. Kaufman, Nucleosome assembly: the CAF and the HAT, Curr. Opin. Cell Biol. 8 (1996) 369–373. [6] L. Chang, S.S. Loranger, C. Mizzen, S.G. Ernst, C.D. Allis, A.T. Annunziato, Histones in transit: cytosolic histone complexes and diacetylation of H4 during nucleosome assembly in human cells, Biochemistry 36 (1997) 469–480. [7] R.E. Sobel, R.G. Cook, C.D. Allis, Non-random acetylation of histone H4 by a cytoplasmic histone acetyltransferase as determined by novel methodology, J. Biol. Chem. 269 (1994) 18576–18582. [8] R.E. Sobel, R.G. Cook, C.A. Perry, A.T. Annunziato, C.D. Allis, Conservation of deposition-related acetylation sites in newly synthesized histones H3 and H4, Proc. Natl. Acad. Sci. USA 92 (1995) 1237–1241. [9] M.H. Kuo, J.E. Brownell, R.E. Sobel, T.A. Ranalli, R.G. Cook, D.G. Edmondson, S.Y. Roth, C.D. Allis, Transcription-linked acetylation by Gcn5p of histones H3 and H4 at specific lysines, Nature 383 (1996) 269–272. [10] P.D. Kaufman, R. Kobayashi, N. Kessler, B. Stillman, The p150 and p60 subunits of chromatin assembly factor I: a molecular link between newly synthesized histones and DNA replication, Cell 81 (1995) 1105–1114. [11] A. Verreault, P.D. Kaufman, R. Kobayashi, B. Stillman, Nucleosome assembly by a complex of CAF-1 and acetylated histones H3/H4, Cell 87 (1996) 95–104. [12] J.K. Tyler, C.R. Adams, S.R. Chen, R. Kobayashi, R.T. Kamakaka, J.T. Kadonaga, The RCAF complex mediates chromatin assembly during DNA replication and repair, Nature 402 (1999) 555–560. [13] J.A. Mello, H.H. Sillje, D.M. Roche, D.B. Kirschner, E.A. Nigg, G. Almouzni, Human Asf1 and CAF-1 interact and synergize in a repaircoupled nucleosome assembly pathway, EMBO Rep. 3 (2002) 329–334. [14] A.T. Annunziato, R.L. Seale, Histone deacetylation is required for the maturation of newly replicated chromatin, J. Biol. Chem. 258 (1983) 12675–12684. [15] A. Loyola, G. Almouzni, Histone chaperones, a supporting role in the limelight, Biochim. Biophys. Acta 1677 (2004) 3–11.

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