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European Molecular Biology Laboratory, Gene Expression Programme, Meyerhofstrasse 1, 69117 Heidelberg, Germany,1 Division of Molecular Genetics (B060), Deutsches Krebsforschungszentrum, INF 280, 69120 Heidelberg, Germany,2 Adolf-Butenandt-Institut, Schillerstrasse 44, 80336 München, Germany,3 Cancer Epigenetics Laboratory, Molecular Pathology Programme, Spanish National Cancer Centre, Melchor Fernández Almagro 3, 28029 Madrid, Spain4
Received 18 March 2005/ Returned for modification 11 April 2005/ Accepted 5 May 2005
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ABSTRACT |
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H2AX foci and have an impaired repair response to ionizing radiation. Taken together, our data show that hMOF is required for histone H4 lysine 16 acetylation in mammalian cells and suggest that hMOF has a role in DNA damage response during cell cycle progression. |
INTRODUCTION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Sequence analysis of HAT proteins reveal that they can be classified in distinct families, with each family having a characteristic substrate specificity (12). GNAT (GCN5-related N-acetyltransferases) family members mainly acetylate lysines on the histone H3 tail. The founding members of the other family, MYST, include Saccharomyces cerevisiae Ybf2p/Sas3p and Sas2p and human MOZ and Tip60 (56). The MYST homology domain is exceptionally well conserved among all family members. This region includes the acetyl coenzyme A binding domain similar to the one found in GNAT acetyltransferases (41) as well as a C2HC-type zinc finger. Recently, the crystal structure of Esa1, an essential yeast HAT, showed that even though the MYST and GNAT family share sequence homology only in motif A, there is high degree of structural conservation in the central core region between the two families (59, 60). The MYST family can be further divided into subgroups based on additional domains present in these proteins. The first subgroup contains proteins with PhD fingers (such as MOZ and MORF), the second subgroup contains proteins with a chromodomain (such as Esa1, dMOF, and Tip60) (41, 56), and a third one (including HBO1) has other known domains such as zinc fingers.
One of the chromodomain-containing members of the MYST family, dMOF, is an integral player in the Drosophila melanogaster dosage compensation process. Dosage compensation ensures that males and females, despite unequal numbers of X chromosomes, express the same amount of X-linked gene products. In Drosophila, this is thought to occur by an approximately twofold transcriptional upregulation of most male X-linked genes. The male X chromosome is coated by dosage compensation complex (DCC), comprised of at least five proteins (dMOF, dMSL1, dMSL2, dMSL3, and dMLE) and two noncoding RNAs (roX1, roX2). Transcriptional upregulation correlates with specific acetylation of histone H4 at lysine 16 (H4K16) on the male X chromosome by dMOF (3, 8, 51). A point mutation in a conserved glycine residue of dMOF that renders the protein enzymatically inactive leads to male-specific lethality (19). Biochemical characterization of the dMOF has shown that it is an RNA-binding protein that acetylates not only histone H4 lysine 16 but also other members of the DCC, namely dMSL1 and dMSL3 (4, 11, 31).
It is remarkable that all the proteins of the Drosophila DCC have been well conserved during evolution (26, 27, 45), even though dosage compensation is brought about by different means in other animal phyla. Based on current evidence, it is reasonable to assume that dosage compensation has evolved independently several times, illustrating an interesting case of convergent evolution (28). How, then, have different dosage compensation mechanisms evolved? Recent data suggest that animals have co-opted evolutionary ancient chromatin-modifying complexes for a new function in dosage compensation. Caenorhabditis elegans dosage compensation is regulated by condensin-like proteins, which are normally involved in chromosome compaction during mitosis (18). Likewise, polycomb proteins that have been implicated in X chromosome inactivation in mammals have an evolutionary older function in repression of homeotic genes during development (35). With the exception of the mammalian dMLE orthologue, the transcriptional coactivator RNA helicase A, the function of the Drosophila DCC gene orthologues in vertebrates remains unclear.
Recently, a putative human orthologue of Drosophila MOF, hMOF/MYST1, was isolated (34). Like the Drosophila protein, it contains a chromodomain and a MYST family HAT domain. A C-terminal fragment of this protein was shown to possess histone acetyltransferase activity toward histones H3, H2A, and H4 in vitro (34). A human gene hMSL3/MSL3L1 has also been isolated and characterized previously as a candidate gene for several developmental disorders (15, 39). It encodes a protein with significant homology to the Drosophila MSL3 in three distinct regions, including the two chromo-like domains (27).
Intrigued by this evolutionary conservation, we have studied the role of hMOF and hMSL3 proteins in mammalian cells. We report that human MOF possesses acetyltransferase activity on histones and nucleosomes. Interestingly, depletion of hMOF in HeLa cells leads to a dramatic decrease in histone H4 lysine 16 acetylation, while other acetylation sites appear to be unaffected. In addition, the cells show altered nuclear morphology with polylobular nuclei. This striking phenotype can be rescued by treating the affected cells with the histone deacetylase inhibitor trichostatin A (TSA). HeLa cells transfected with hMOF small interfering RNA (siRNA) show proliferation defects and accumulate in the G2/M phase of the cell cycle. We show that the observed G2/M arrest is at least partially caused by activation of the DNA damage response pathway, illustrated by an increased number of ATM pS1981 and H2AX foci in hMOF-depleted cells.
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MATERIALS AND METHODS |
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Antibodies against hMOF-GST and FLAG-hMSL3 fusion proteins were produced in rats and rabbits, respectively. Both antibodies were further affinity purified. To generate hMOF peptide antibodies, rabbits were injected with keyhole limpet hemocyanin-conjugated peptides PERKITRNQKRKHDE and KWAPPKHKQVKLSKK at Eurogentec, Belgium. Antibody was affinity purified with the peptide KWAPPKHKQVKLSKK. Antibody against RNA helicase A was obtained from C.-G. Lee (University of Medicine and Dentistry, New Jersey), MRG15 was obtained from O. Pereira-Smith (University of Texas, San Antonio), RCC1 was obtained from I. Mattaj (EMBL, Heidelberg), lamin A/C and lamin B1 were obtained from H. Herrmann (DKFZ, Heidelberg), and H4K12Ac was obtained from Bryan Turner (University of Birmingham). ß-Tubulin and FLAG (Sigma), ATM pS1981 (Rockland Immunochemicals), H4K16Ac (Chemicon), H3K14Ac and H3K23Ac (Abcam), and H2AX (Upstate) were purchased as indicated.
Coimmunoprecipitation and GST pull-down assays. HeLa nuclear extract was prepared as previously described (16). Approximately 100 µg nuclear protein in HEMG buffer (25 mM HEPES, pH 7.6, 12.5 mM MgCl2, 0.5 mM EDTA, 1 mM dithiothreitol, 0.2 mM phenylmethylsulfonyl fluoride, 10% glycerol) with 100 mM KCl was used in immunoprecipitation. Immunoprecipitates were washed three times with HEMG buffer with 100 mM KCl at room temperature, and bound proteins were eluted with sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) loading buffer.
In vitro GST pull-down assays were performed in HEMG buffer with 200 mM KCl. Briefly, 300 ng of FLAG-hMSL3 or RCC1 was incubated with 1 µg of recombinant hMOF constructs bound to glutathione beads for 1 h at room temperature. After incubation, beads were washed three times, for 5 min each time, with HEMG buffer with 200 mM KCl. Bound proteins were eluted with SDS loading buffer.
Histone acetyltransferase assays. Histone acetyltransferase assays were performed as described earlier (2). Protein (100 ng or indicated amounts) was incubated for 30 min at 30°C in HAT buffer (20 mM Tris-HCl, pH 8.8, 1.5 mM MgCl2, 10 mM NaCl, supplemented with 125 nCi [3H]acetyl-coenzyme A) with 1 µg of recombinant histone octamer or 1 µg of nucleosomal histones assembled by salt exchange. Reactions were either blotted on a hydrophobic p81 paper and scintillation counted or run on 15% SDS-PAGE and Coomassie stained. The signal on SDS-PAGE gels was intensified with Amplify solution (Amersham).
Mass spectrometry. For the in vivo analysis of modified histones, histone bands were modified in gel using propionic anhydride or D6 acetic anhydride and digested with trypsin (6, 7, 38, 49). Matrix-assisted laser desorption ionization (MALDI) spectra were recorded on a Voyager STR instrument (PE-Sciex). For mass spectrometry (MS)/MS analysis, collision-induced decay spectra were recorded on a Q-STAR XL instrument (PE-Sciex) with manually adjusted collision energies. Fragment spectra were interpreted manually.
Correlation of confocal laser scanning and electron microscopy. Ultrastructural investigation by electron microscopy was correlated to observations by laser scanning microscopy as described in reference 40. Briefly, cells grown on gridded coverslips (Cellocate; Eppendorf AG, Germany) were fixed for 1 h on ice in a mixture of 4% freshly prepared formaldehyde, 1% glutaraldehyde (electron microscopy grade; Sigma), 1 mM MgCl2, and 100 mM sodium phosphate buffer, pH 6.8, rinsed in buffer, and embedded in Vectashield fluorescence mounting medium (Vector Laboratories) for observation by confocal laser scanning microscopy (LSM). Following LSM investigation, the coverslips were rinsed in buffer, postfixed in 1% buffered OsO4 for 30 min at room temperature, dehydrated in aqueous ethanol, and embedded in epoxy resin (Epon 812; Sigma). The coverslips were subsequently removed under liquid nitrogen depicting the negative imprint of the Cellocate grid. Ultrathin sections at a nominal thickness of 70 nm were prepared from the grid region of interest, poststained in 2% uranyl acetate and aqueous lead citrate, and observed in a Philips 410 transmission electron microscope.
Cell culture and transfection. HeLa and HepG2 cells were grown in Dulbecco's modified Eagle's medium supplemented with 10% fetal calf serum, penicillin, streptomycin (Invitrogen), and L-glutamine (Invitrogen). Synthetic siRNAs (hMOF-1, GUGAUCCAGUCUCGAGUGA; hMOF-2, AAAGACCAUAAGAUUUAUU; hMOF-3, CAAGAUCACUCGCAACCAA; hMSL3-1, CGGUUAGUGAAACUUCCAU; hMSL3-2, AAAGGUGACUUCGUCUAAA; control, CACGTACGCGGAATACTTCG, sense strand) were purchased from MWG Biotech. Approximately 1.5 x 105 cells were transfected with Oligofectamine (Invitrogen) with a 60 nM final concentration of siRNA complexes. TSA was added to a final concentration of 25 ng/ml for 72 h, and 2 mM caffeine was added for 24 h where indicated.
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RESULTS |
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Loss of hMOF abolishes H4 lysine 16 acetylation in vivo. To study the function of the hMOF protein in vivo, we used RNA interference in HeLa and HepG2 cells. Using three independent synthetic siRNAs against the hMOF coding region, we were able to specifically reduce the levels of hMOF protein down to 10% of its original level in HeLa cells (Fig. 2A, upper panel, and data not shown). To determine which lysine residues would be targets of acetylation by hMOF in vivo, we isolated endogenous histones by acid extraction from cells treated with either control siRNA or hMOF-specific siRNA. The histones were separated by SDS-PAGE, and subsequently, Western blot analysis was performed with antibodies against acetylated histones. Ponceau S staining of the membranes revealed equal loading of histones. Interestingly, the level of histone H4 lysine 16 acetylation (H4K16) in hMOF-depleted cells was severely reduced in comparison to the control cells (Fig. 2A, right and middle panels). In contrast, there was no significant difference in acetylation of H3K14, H3K23, or H4K12, although these lysines were targets of recombinant hMOF in vitro (Fig. 2A and data not shown). Although we did not analyze H4K5 or H4K8 acetylation status by Western blotting, mass spectrometric analysis of endogenous histones did not reveal significant changes (see below).
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As the Western blot results with acetylation-specific antibodies suggested a strong effect of hMOF ablation on the acetylation of H4K16 (Fig. 2A), we performed an MS/MS analysis on the monoacetylated/propionylated H4 peptide that carries amino acids 4 to 17 of the H4 tail. By comparing the relative abundance of the expected fragment ions for the nonacetylated (propionylated) and the acetylated peptides (Fig. 2D), we concluded that in HeLa cells, the major histone H4 acetylation sites are K16 and K12 (85%). There is, however, a marked difference in the ratio of K12/K16 acetylation between cells depleted for hMOF and control cells. Fragment ions containing only K16 (m/z of 530.35 for the acetylated form and m/z of 544.35 for the unacetylated, propionylated form) show an approximately sevenfold-lower appearance of the acetylated form in hMOF knockdown cells compared to control cells (Fig. 2E, second panel). This strong decrease in acetylation is partly rescued by an increase in the acetylation of lysines 5, 8, and 12 (compare the peak height of for acetylated versus nonacetylated in the top, middle, and bottom panels). This shift also explains the moderate effect of hMOF depletion on overall monoacetylation levels despite its strong impact on K16 acetylation.
hMOF and hMSL3 interact directly in vitro and in vivo. It has previously been shown in Drosophila that dMOF and dMSL3 can interact in vitro and that this interaction leads to acetylation of dMSL3 in vivo and in vitro (11). These observations led us to test whether the interaction is conserved between hMOF and hMSL3 in mammalian cells. We performed coimmunoprecipitation experiments with hMOF antibody or corresponding preimmune serum, as a negative control, from nuclear extracts prepared from HeLa cells. The coimmunoprecipitated samples were analyzed by SDS-PAGE followed by Western blot analysis. The results of these experiments show that hMSL3 coimmunoprecipitates with hMOF (Fig. 3A, compare lanes 1 and 2). Interaction was also detected when coimmunoprecipitation experiments were performed with anti-hMSL3 or using nuclear extracts from HEK293 cells (data not shown). It is important to note that we could observe two isoforms of hMSL3 (hMSL3a and hMSL3c) coimmunoprecipitating with hMOF. Based on bioinformatics analysis and GenBank, these isoforms most likely represent two splice variants of the hMSL3 gene.
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The experiment above demonstrates that interaction between hMOF and hMSL3 is conserved in mammalian cells. It does not address, however, whether this interaction is direct or mediated by another unknown component. To address this issue, we performed in vitro GST pull-down assays with GST-hMOF fusion derivatives and full-length hMSL3. Another nuclear protein, RCC1, was used as a control. Following binding, the beads were washed and the bound fraction was analyzed by Western blotting. The blots were first probed with hMSL3 or RCC1 antisera and subsequently with a GST antibody to control for equal loading of hMOF-GST constructs. Consistent with our observations in vivo, hMOF and hMSL3 interacted directly in vitro (Fig. 3B, top panel, lane 1). An interaction could still be detected with a deletion derivate of hMOF containing the MYST domain (lane 3 and 4), but no interaction was detected with the derivative containing only the chromodomain of hMOF (lane 2). The C2HC-type zinc finger that is embedded in the catalytic domain of hMOF was not required for interaction (Fig. 3B, lane 4). hMSL3 interaction with the C terminus of hMOF was specific, since another chromatin-associated protein, RCC1, did not interact with hMOF in the same experimental conditions (Fig. 3B, bottom panel). In summary, we can demonstrate a direct interaction between hMOF and hMSL3. We also found that the C terminus of hMOF mediates this interaction.
hMOF- and hMSL3-depleted cells show nuclear morphology defects. Apart from severe reduction of lysine 16 acetylation, another striking feature of hMOF knockdown cells was their abnormal nuclear structure. About 36 h after hMOF siRNA transfection, cells started to undergo dramatic changes in nuclear morphology and to form multiple lobes (Fig. 4B). The phenotype ranged from nuclei showing one extra lobe to cells with 6 to 8 lobes in the nucleus. This phenotype was observed with three independent hMOF siRNAs tested, and on average, 25 to 30% of the cells treated with hMOF siRNA showed severe morphological defects (see below). Interestingly, hMSL3-depleted cells also showed similar defects, albeit with a lower frequency, suggesting that the two proteins function in the same pathway (Fig. 4B and see below). Knockdown of hMSL3 in HeLa cells was effective with two independent siRNAs tested, as shown in Fig. 4A. Similar results were obtained when hMOF and hMSL3 were depleted in HepG2 cells, excluding a cell type-specific effect (Fig. 4B and data not shown). The polylobular phenotype was also observed in HeLa cells stably expressing enzymatically inactive epitope-tagged hMOF (HA-2x FLAG-hMOF G327D), whereas a cell line stably transfected with a wild-type hMOF construct (HA-2x FLAG-hMOF) had a normal morphological appearance (Fig. 5B).
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It has previously been reported that depletion of nuclear pore complex component RanBP2 leads to changes in nuclear morphology (42). Since the nuclei appear polylobulated after completion of mitosis, we wanted to address whether this defect appears due to abnormal nuclear envelope reassembly. For this purpose, we immunostained hMOF siRNA-treated and control cells with mAB414 monoclonal antibody that recognizes several nucleoporins. However, we observed no major defects in the nuclear pore distribution (Fig. 4D). Similarly, the cytoskeleton of hMOF knockdown cells appeared normal, as judged by phalloidin (F-actin) and ß-tubulin staining (data not shown). We have also tested differences in distribution of nucleoli (fibrillarin), heterochromatin (HP1
, H3K9 methylation, H3K27 methylation), and centromeres (CREST autoimmune serum) by immunofluorescence but have not been able to find any differences between hMOF knockdown and control cells (data not shown).
Another component influencing nuclear architecture is the nuclear lamina. Loss or overexpression of lamins, which constitute the nuclear lamina, has been reported to cause defects in nuclear structure (9, 20). We therefore tested whether the distribution or levels of nuclear lamina proteins, namely lamin A/C and lamin B1, were affected in hMOF siRNA-treated cells. Remarkably, we could observe a modest but consistent enrichment of lamin B1 and lamin A/C, typically in the constriction between the lobes (Fig. 4D; see below). The difference between hMOF-depleted and control cells was mainly in the distribution of lamins, as there was no significant change in lamin B1 or lamin A/C protein levels upon hMOF depletion (data not shown).
Ultrastructural investigation by electron microscopy revealed that the nucleoli were normal in shape and size and showed their characteristic stacked composition in fibrillar center, fibrillar component, and granular component, indicating normal physiological activity in hMOF-depleted versus control cells. Nuclear pores appeared as in normal control cells. In contrast, the lamina underneath the nuclear envelope was more discrete and regular in structure than in control cells. This is consistent with observations made above with enriched lamin staining.
Major structural reorganization was, however, seen at the cytoplasmic side of the folds which separate the nuclear lobules. The folding process appears to be more complicated than a simple indentation of the nuclear envelope from the periphery to the center. At the front of the fold, the substrate-facing side (bottom) of the nuclear envelope forms stacks of wrinkles in an oblique direction to the main fold (Fig. 4E). This front region of folds is densely packed with vesicles and filaments, suggesting that folds are formed also by an active engagement of the cytoplasm. In summary, we did not observe gross changes in nuclear organization in hMOF-depleted cells.
However, it remains possible that subtle changes in chromatin acetylation levels in cells or changes in acetylation of an unknown substrate may contribute to the observed nuclear deformations. We reasoned that if this was the case, changing the balance of acetylation in the cell might rescue the polylobular phenotype. To address this issue, we first transfected HeLa cells with a control siRNA or hMOF siRNA, and 24 h after transfection, we added trichostatin A (25 ng/ml), a potent histone deacetylase inhibitor, to the growth medium for another 72 h. This was followed by Western blot analysis to assess the knockdown efficiency, assay for cell growth, and observation of the nuclear morphology in these cells. hMOF was efficiently knocked down in untreated and TSA-treated cells (data not shown). Interestingly, whereas control cells proliferated significantly more slowly in the presence of TSA, hMOF knockdown cells grew slightly better in the presence than in the absence of TSA (Fig. 5A). TSA did not, however, fully restore normal growth of hMOF knockdown cells. More importantly, nuclear morphology was restored in TSA-treated hMOF knockdown cells. Similar results were obtained with all hMOF siRNAs tested (Fig. 5B). We further confirmed that in both HeLa and HepG2 cells, polylobulation correlates with loss of hMOF, which can also be tracked with an H4K16Ac-specific antibody (Fig. 5C).
However, TSA treatment did not lead to an increase in H4K16 acetylation in hMOF-depleted cells (data not shown), suggesting that this modification is not directly linked to nuclear shape changes and that hMOF has other cellular targets (see Discussion).
Cell cycle checkpoint activation in hMOF-depleted HeLa cells. We also observed that hMOF siRNA-treated HeLa cells showed proliferation defects in comparison to the control siRNA-treated cells (Fig. 6A). hMOF-depleted cells were negative for Trypan blue staining, indicating that the primary cause of growth arrest was not due to apoptosis or necrosis (data not shown). Next, we analyzed the control siRNA and hMOF siRNA-treated populations with flow cytometry to observe changes in cell cycle profiles. We found that hMOF siRNA-treated cells accumulate in the G2/M phase of the cell cycle in comparison to the control cells (Fig. 6B). Furthermore, fluorescence-activated cell sorter (FACS) analysis also revealed that the cells treated with hMOF siRNA were approximately 10% larger than the control cells in all phases of the cell cycle (data not shown).
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We reasoned that activation of either of these checkpoints could explain the accumulation of hMOF-depleted cells in G2/M. First, we tested activation of p38 in hMOF-depleted and control cells but detected no difference in p38 phosphorylation (data not shown).
Next, we tested whether the arrest in hMOF knockdown cells involved the ATM/ATR pathway by using two inhibitors of these cascades, caffeine and wortmannin. It is important to note that both caffeine and wortmannin can inhibit activation of both ATM and ATR, but the 50% inhibitory concentration of both drugs for ATR is an order of magnitude higher than for ATM (46, 47). Therefore, to distinguish between these two possibilities, we used 2 mM caffeine or 2 µM wortmannin, doses that should primarily inhibit ATM. At 72 h after transfection, hMOF-depleted and control cells were treated with caffeine or wortmannin for 24 h or left untreated. Subsequently, cells were analyzed by FACS to determine cell cycle profiles. Both caffeine and wortmannin restored the cell cycle profile of hMOF-depleted cells to almost that of control cells (Fig. 6C and data not shown).
To examine whether hMOF-depleted cells had activated the ATM-dependent checkpoint, we immunostained hMOF knockdown and control cells with an antibody against phosphorylated ATM (pS1981). In control cells, about 20% of cells had one or several ATMp foci. However, the number of cells with ATMp foci after depletion of hMOF was consistently higher, about 40%, with both siRNAs tested (Fig. 6D). Both the percentage of the cells with phospho-ATM foci and the average number of foci was increased upon hMOF knockdown. These foci colocalized with H2AX foci (data not shown), suggesting that they represent double-stranded breaks (DSBs) occurring during the normal cell cycle.
Hypoacetylation of histones could render DNA susceptible to breakage, thereby activating the checkpoint, or alternatively, it could impair repair of normally occurring DSBs. In both cases, the result would be an increased number of ATMp and H2AX foci. To distinguish between these possibilities, we treated the hMOF-depleted and control cells with ionizing radiation (1 Gy) and performed a time course study to assess DNA repair kinetics by counting the presence of ATMp and H2AX foci. hMOF-depleted cells consistently showed a significant delay in kinetics of DNA repair as observed by the presence of more ATMp foci in comparison to control cells (Fig. 6E). Similar results were obtained with a higher dose of 6 Gy and with HepG2 cells. These results suggest that hMOF-depleted and thus H4K16-hypoacetylated HeLa cells activate the G2/M checkpoint due to a delay in the kinetics of the DNA repair process of hypoacetylated chromatin, as opposed to hypoacetylation leading to more DNA damage without having an effect on the repair process itself.
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DISCUSSION |
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Role of hMOF in histone H4 lysine 16 acetylation in vivo. We detected a significant decrease in H4K16 acetylation in hMOF knockdown cells, whereas acetylation of other lysines (H4K12, H3K14, and H3K23) remained mostly unaffected, although these lysines are acetylated by recombinant hMOF in vitro. Endogenous mammalian MSL complex therefore appears to be specific for H4K16, or alternatively, there could be more redundancy between HATs acetylating H3K14, H3K23, and H4K12 in mammalian cells. Purification of the human MSL complex should shed more light on this issue in the future.
The dramatic decrease in H4K16 acetylation in hMOF knockdown cells suggests that hMOF is the major H4K16 acetyltransferase in mammals. To our knowledge, this is the first report connecting a mammalian histone acetyltransferase to a specific lysine residue in vivo. In mammals, lysine 16 is clearly the most abundant acetylation site on histone H4 (32, 53, 55). In bulk chromatin preparations, nearly all acetylated histone H4 molecules are acetylated at H4K16, be it mono-, di-, tri-, or tetra-acetylated H4 (32, 53, 55). It will be interesting to study whether localization of hMOF coincides with H4K16 acetylation patterns on promoters and/or coding regions previously documented in mammalian cells.
It was originally suggested that histone acetylation is a hierarchical process, where one modification is required for subsequent acetylation (55). H4K16, being the most commonly acetylated residue, would be placed on top of the cascade. However, we could detect no reduction in H4K5, H4K8, or H4K12 acetylation in hMOF knockdown cells. On the contrary, hMOF-depleted cells appeared to compensate for the loss of K16 acetylation by higher levels of acetylation on other lysine residues on the H4 tail. This result clearly implies that K16 acetylation is not a prerequisite for subsequent acetylation.
Conserved interaction between hMOF and hMSL3. We found that hMOF and hMSL3 coimmunoprecipitate in vivo in HeLa cells as well as HEK293 cells. Furthermore, we detect a direct interaction between hMOF and hMSL3 in vitro. We have also mapped the sites of interaction to the C-terminal MYST domain. A similar nuclear morphology phenotype in hMOF- and hMSL3-depleted cells corroborates the finding that the two proteins function in the same complex.
The human orthologue of dMLE, RNA helicase A, did not interact with hMOF or hMSL3 under the conditions used. dMLE seems to interact with other members of the Drosophila DCC only transiently (14), suggesting that it is not a stable component of the complex. It is possible that dMLE is a later addition to the DCC, perhaps reflecting the role of noncoding RNAs as functional units of the Drosophila complex. This would be consistent with the results of this study.
hMOF is involved in the maintenance of nuclear structure in HeLa cells. We observed that the nuclei of both hMOF and hMSL3 knockdown cells appear polylobulated. By using live cell imaging, we were able to show that, in hMOF-depleted cells, these defects appear during nuclear envelope reassembly in late telophase. We did not observe any lagging chromosomes in mitosis in these cells, indicating the defects are not due to missegregation of chromosomes. However, lamin A/C and lamin B1 were redistributed to the concave surfaces of invaginations. Consistent with this observation, we could detect thickened nuclear lamina structures in hMOF-depleted cells by electron microscopy. Previous studies have shown that overexpression of full-length or dominant-negative forms of lamins leads to nuclear structure aberrations (9, 20). It was recently shown that expression of Chk tyrosine kinase in the nucleus leads to a strikingly similar polylobular phenotype, including redistribution of lamin B1 and a high concentration of microtubules around the invaginations (33). However, Chk-dependent nuclear shape change was mitosis independent, whereas we could only see morphological changes following mitosis. It is therefore likely that there are multiple mechanisms maintaining nuclear shape.
Acetylation appears to be involved in the process, since expression of an enzymatically inactive hMOF induced changes in nuclear morphology similar to those induced by hMOF knockdown. Consistently, polylobulation of HeLa cells was rescued upon treatment with TSA, a potent histone deacetylase inhibitor. It is, however, unlikely that H4K16 acetylation plays a direct role in nuclear morphology changes. First, TSA treatment of hMOF-depleted cells did not lead to an increase in H4K16 acetylation. Second, hMSL3 depletion had no impact on bulk K16 acetylation despite a similar phenotype (data not shown). Third, the histone deacetylases known to deacetylate H4K16 belong to the TSA-insensitive SIR family (22, 57). It is thus conceivable that hMOF also has other TSA-sensitive cellular targets that remain to be identified.
hMOF, H4K16 acetylation, and DNA damage response pathway.
We also observed that the depletion of hMOF affected cell proliferation in HeLa cells. This effect was not due to apoptosis or necrosis, but we found that the cells were enriched at the G2/M phase of cell cycle. In an effort to understand whether a checkpoint cascade is activated in hMOF-depleted cells, we observed that the G2/M defect could be suppressed by inhibiting checkpoint response with low doses of caffeine. Low doses of caffeine were used to rule out the involvement of DNA-PK and the ATR pathway. Consistent with this observation, we found no defects in UV-induced DNA damage response in hMOF siRNA-treated cells (data not shown). In addition to caffeine, we also observed that wortmannin had a similar effect on the cell cycle profile (data not shown). These results strongly suggest that hMOF-depleted cells have activated the DNA damage pathway. Decreasing cellular hMOF levels led to increased phosphorylation of serine 1981 of ATM and H2AX, hallmarks of activation of this pathway (5). hMOF knockdown cells do not seem to be more susceptible to DNA damage. However, the kinetics of DNA repair after irradiation is slowed down in these cells. Thus, the most likely explanation for the activation of the checkpoint is that hMOF-depleted cells fail to efficiently repair DSBs that occur during the cell cycle, thus delaying the progression to mitosis.
Previously, acetylation of histone H4 has been linked to cell cycle progression through G2/M in yeast (29, 30). Yeast cells with four mutant lysines (K5Q, K8Q, K12Q, and K16Q) in the H4 tail accumulate in G2/M in a RAD9-dependent manner (29, 30). RAD9 is a sensor protein that arrests cell cycle progression if cells have accumulated DNA damage (58). Interestingly, yeast Esa1p is required for cell cycle progression, and rad9 can also suppress the esa1 phenotype (13, 50). Esa1p is a MYST family acetyltransferase closely related to hMOF. It is the catalytic subunit of the NuA4 histone acetyltransferase complex that also contains Eaf3p, the yeast ortholog of hMSL3. Another striking phenotype of esa1 mutant cells is accumulation of visible changes in chromatin structure (13). hMOF-depleted cells accumulate similarly in G2/M, in a caffeine-sensitive manner, and show changes in nuclear shape, suggesting that the involvement of histone H4 acetylation in cell cycle progression and/or DNA repair is evolutionarily conserved from yeast to mammals.
Recently, several studies have shown the involvement of histone methylation and acetylation in DSB repair (21, 23, 25, 37, 44). Intriguingly, acetylation of histones has been shown to increase the activity of DNA-PK in a nucleosomal context in vitro (37). Furthermore, it has been shown that within 1 to 2 kb of a defined DSB, very little H2AX can be detected (48) and that H4K16 is generally hypoacetylated in fission yeast (23). These studies suggest that there is a connection between H4K16 acetylation, DNA-PK activity, and H2AX phosphorylation in DSB repair. If this aspect of DSB repair is evolutionarily conserved to mammals, it would have interesting implications for the role of hMOF in DSB repair.
H4K16 acetylation and cancer. It is remarkable that lysine 16 acetylation by the MOF enzyme has been evolutionarily used for various purposes. In Drosophila, it correlates with increased X-linked gene transcription (1), while in yeast, H4K16 acetylation by orthologous SAS2 regulates heterochromatin spreading (52) and, perhaps surprisingly, negatively correlates with gene transcription (24). It appears that mammals have adopted histone H4 lysine 16 acetylation to be one of the most abundant histone modifications.
Recently, it was shown that loss of acetylation at H4K16 is a common hallmark of human cancer (17). A variety of human tumor cell lines and primary tumors show significant hypoacetylation at H4K16 and hypo(tri)methylation at H4K20. Remarkably, reduction in H4K16 acetylation correlates with tumor progression. It was also shown that hMOF is not associated with hypomethylated repetitive D4Z4 sequences in the HL60 tumor cell line, in contrast to wild-type lymphocytes, where these elements are normally methylated.
We have shown that hMOF is the major H4K16-specific histone acetyltransferase in human cell lines. If this is the case for most tissues, it implies a significant role for hMOF activity in tumorigenesis. Clearly, future studies need to address the role of hMOF and H4K16 acetylation in healthy and cancer tissues.
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ACKNOWLEDGMENTS |
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This work partially funded by DFG Transregio 5. S.R. is a recipient of EMBO and HFSP postdoctoral fellowships.
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FOOTNOTES |
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