Involvement of Human MOF in ATM Function

Yeast two-hybrid screen.

The procedure for the construction of hMOF or its fragments is the same as that used for ATM and TRF1 and has been described previously (30, 33, 45, 48). The yeast two-hybrid screen was performed by the transformation of yeast strain Y153 containing plasmid pHM677 (Y153/pHM677) with a cDNA library from human embryonic cells (Clontech). Primary transformants were analyzed for interaction with ATM by selection for histidine prototrophy on dropout plates supplemented with 20 mM 3-aminotriazole and for the expression of β-galactosidase (β-gal) as determined by filter lift tests. Plasmids encoding putative binding partners of ATM were then isolated from double-positive yeast clones and retransformed into Y153/pHM677 in order to confirm the interactions. Positive clones after retransformation were subjected to automated sequencing and homology searches using a BLAST (tBLASTn) search of the GenBank human database. Several putative cellular binding partners of ATM were identified. Most of them were isolated in several copies. To quantify the interaction between ATM and hMOF, interactions between hMOF (wild type and mutated) and ATM fragments in liquid β-gal were assayed (Fig. 1) according to the previously described procedure (12, 30).

Cell culture and derivation of cell lines.

AT221JE-TpEBS7 (A-T [ataxia-telangiectasia] fibroblast with empty vector only), AT221JE-TpEBS7-YZ5 (A-T fibroblast complemented with ATM cDNA), GM5849, and 293 cells were maintained and transfected with plasmids as described previously (45). Complementary DNAs encoding wild or mutant hMOF (ΔhMOF) were cloned into the mammalian expression vector pIND(SP1)/Neo (Invitrogen, Carlsbad, CA) as described previously (18, 45). Mutant forms of hMOF were created by using a PCR approach with appropriate primer pair combinations. ATMΔLZ was generated as described previously (10). The wild or mutant forms tagged to GFP, HA, or Flag were cloned into the pIND(SP1)/Neo vector. The final constructs were verified by DNA sequencing. An adenoviral construct expressing hMOF (Ad.hMOF) was constructed as described previously (44). hMOF small interfering RNA (siRNA) and control Luc siRNA were obtained from Dharmacone Research (Lafayette, CO). We also used the hMOF siRNA vector pRNA-U6.1/Neo (GeneScript).

Western analysis, immunoprecipitation, HAT, and ATM kinase assays.

Cell lysates were prepared according to the previously described procedure (41). Polyclonal hMOF antibody raised in rabbit has been described previously (34). Immunoblotting and detections for hMOF and ATM were done according to a previously described procedure (41, 45). For immunoprecipitation (IP), cells were broken in lysis buffer as described previously (41). Lysates were precleaned with purified immunoglobulin G and protein A/G beads. Proteins were immunoprecipitated with specific antibodies, and immunoprecipitants were washed with lysis buffer as described previously (41). An ATM kinase assay was performed as described previously (41). A histone acetyltransferase (HAT) assay was performed according to the manufacturer's instructions (Upstate Biotechnology).

Assay for ionizing radiation sensitivity.

Cells were plated in 35-mm dishes. The cell count was determined using a Coulter counter. Cells were plated as single cells into 60-mm dishes in 5 ml of media, incubated for 6 h, and subsequently exposed to IR. The actual amounts of cells per dish were chosen to ensure that about 50 colonies would survive a particular radiation dose treatment. Cells were exposed to IR in the dose range of 0 to 8 Gy at room temperature. Cells were incubated for 12 or more days and were fixed in methanol:acetic acid (3:1) prior to staining with crystal violet. Only colonies containing >50 cells were counted.

Metaphase chromosome spreads were prepared by procedures described previously (35). Giemsa-stained chromosomes of metaphase spreads were analyzed for chromosome end-to-end associations. G₁ phase-specific chromosomal aberrations were assessed as described previously (37). Briefly, cells in plateau phase were irradiated with 3 Gy, allowed to incubate for 24 h, and subcultured, and metaphases were collected. Chromosome spreads were prepared by the procedure described previously (35). The categories of G₁-type asymmetrical chromosome aberrations scored included dicentrics, centric rings, interstitial deletions/acentric rings, and terminal deletions.

S-phase-specific chromosomal aberrations were analyzed at metaphase. Exponentially growing cells were treated with 2 Gy of gamma radiation, and mitotic cells were collected 3 to 6 h after irradiation. Both chromosome- as well as chromatid-type aberrations were scored. For G₂ phase-specific chromosomal aberrations, cells in exponential phase were irradiated with 1 Gy, and metaphases were collected at 45 and 90 min following irradiation and examined for chromatid breaks and gaps per metaphase as described previously (33).

Neutral filter elution.

The level of DNA DSBs in cells was estimated as described previously (40). Cells were labeled for a period of one-and-one-half doublings and then were washed free of radioactive medium and reincubated in nonradioactive medium for an additional 3 h prior to the experiment. Cells were exposed to IR and incubated at 37°C for different times postirradiation. Cells were layered onto polycarbonated filters, lysed, and eluted under neutral conditions (pH 9.6) as described previously (40). Relative elution (RE) was calculated as RE = −log(FI/Fc), where FI and Fc are the fractions of DNA left on the filter for the irradiated (I) and unirradiated (c) cells when 75% of the internal standard DNA is left on the filter.

IR enhances hMOF-mediated acetylation of histone H4 at K16.

To test whether IR exposure alters hMOF levels or activity in the presence or absence of functional ATM, cells in exponential phase were irradiated, and hMOF protein levels were determined at increasing intervals postirradiation. Exposure to IR altered neither hMOF protein levels (Fig. 4A) nor its H4 peptide acetylation activity (Fig. 4B), irrespective of whether cells expressed ATM or not. Like its ortholog in Drosophila (1, 49) and in contrast to yeast Esa1p or mammalian Tip60, hMOF acetylates histone H4 uniquely at lysine 16 (E. R. Smith and J. C. Lucchesi, unpublished observation). Using an antibody specific for human H4 acetylated at K16, we detected an increase in the acetylation of histone H4 following IR exposure without any detectable change in overall histone H4 levels (Fig. 5B). Cells with (293) and without (GM 5849) functional ATM did not show any difference in histone acetylation, suggesting that K16 acetylation of histone H4 by hMOF is independent of ATM function (Fig. 5B). To determine whether the increased K16 acetylation of histone H4 after IR exposure was due specifically to hMOF activity, cells either expressing mutant hMOF (ΔMOF; fragment 1 of Fig. 1A) or with hMOF knockdown were analyzed by Western blotting for IR-induced acetylation of histone H4 at K16. In comparison to the levels in the control, only cells expressing ΔhMOF (Fig. 1A) or cells with hMOF knockdown (Fig. 5C) had decreased acetylation of histone H4 at K16 after IR exposure (Fig. 5B, D), irrespective of the presence of ATM; cells overexpressing full-length hMOF (Fig. 5A) had higher levels of IR-induced K16 acetylation of histone H4 than control cells did (Fig. 5B). These results suggest that IR exposure enhances hMOF-dependent acetylation of histone H4 at K16 independently of ATM function.

hMOF inactivation abrogates ATM activation.

We tested whether the inactivation of hMOF has any influence on IR-induced ATM activation. The expression of ΔhMOF abrogated ATM autophosphorylation in irradiated cells (Fig. 6A), whereas the expression of hMOF fragments 2 to 4 had little effect on ATM autophosphorylation (data not shown). To determine whether the hMOF association influences ATM autophosphorylation, cells expressing HA-tagged wild-type hMOF and ΔhMOF were irradiated and examined for ATM Ser1981 phosphorylation (Fig. 6B). Interestingly, ATM complexed with full-length hMOF displayed IR-induced ATM phosphorylation (Fig. 6Ba), whereas no such phosphorylation was observed in ATM associated with the ΔhMOF deletion mutant (Fig. 6Bb). The influence of hMOF on IR-induced ATM autophosphorylation is consistent with the influence of hMOF on ATM monomerization (Fig. 6C). Furthermore, cells expressing ΔhMOF or cells with hMOF knockdown also had greatly reduced IR-induced ATM kinase activity, as determined by an in vitro kinase assay (Fig. 6D) (41).

To determine whether an increase in the acetylation of histone H4 at K16 alone could influence ATM activation, cells were treated with either 100 nM of the deacetylase inhibitor trichostatin A (TSA) or 5 mM of sodium butyrate (NaB) for 4 h. TSA and NaB enhanced the level of acetylated histone H4 at K16, with a slight effect, in the case of TSA, on ATM autophosphorylation but not on ATM kinase activity (Fig. 6E). The high levels of IR-induced autophosphorylation of ATM and ATM kinase activity were unaffected by the TSA-enriched acetylation of histone H4 at K16. These results suggest that IR-induced acetylation of histone H4 at K16 by hMOF along with DNA breaks induced by IR exposure may induce changes in the chromatin structure and that such chromatin changes may be transduced by hMOF to ATM for its activation. We also tested whether hMOF could influence ATM activation in a nonchromatin context. To address this issue, we exploited the in vitro system for MRN-dependent substrate phosphorylation by ATM as described previously (29). Our analysis revealed that hMOF does not alter ATM function in vitro (Fig. 7), supporting the argument that hMOF may act as a transducer of chromatin structural alterations to ATM after DNA damage.

hMOF influences ATM-mediated phosphorylation of downstream effectors.

In mammalian cells, in response to DNA damage, ATM phosphorylates various proteins that control G₁, S, and G₂ cell cycle arrest (13, 36, 46). We examined whether the inactivation of hMOF influences ATM-mediated IR-induced phosphorylation on Chk2 in 293 cells. ATM phosphorylates Chk2 on Thr-68 in response to DNA damage (31), which activates Chk2 to phosphorylate substrates including p53 and Cdc25 (32). Cells (293) expressing ΔhMOF as well as cells with hMOF knockdown had reduced IR-induced phosphorylation of Chk2 at Thr-68 compared to control cells (Fig. 8). These results support the argument that hMOF function influences ATM-dependent phosphorylation of its downstream DNA damage response effectors.

hMOF influences genomic stability and IR response.

Cells derived from A-T individuals exhibit genomic instability and are hypersensitive to IR and radiomimetic drugs (36, 46). A-T fibroblasts also show premature senescence. Telomere loss and chromosome end-to-end associations are frequently seen in A-T cells (36, 42). These cells exhibited approximately 0.39 chromosome end-to-end associations per metaphase, whereas the parental cells displayed 0.12 chromosome end-to-end associations per metaphase (Table 1). The increase in chromosome end-to-end association seen in cells expressing ΔhMOF is similar to that in cells with hMOF knockdown. In contrast, the overexpression of hMOF had no effect on the frequency of chromosome end associations. Since chromosome end-to-end associations may lead to anaphase bridge formation, the same cells were analyzed for anaphase bridges. Cells expressing the ΔhMOF and cells with hMOF knockdown displayed about threefold higher frequencies of anaphase bridges compared to parental cells (Table 1). Chromosome end associations and bridges can induce spontaneous chromosome as well as chromatid breaks. Cells expressing ΔhMOF and cells with hMOF knockdown displayed higher frequencies of chromatid as well as chromosome breaks when compared to parental cells (Table 1). Once again, no significant change in chromosomal break frequencies was observed in cells overexpressing wild-type hMOF (Table 1).

Consistent with the differences in spontaneous genomic damage, ATM^+/+ cells expressing ΔhMOF and cells with hMOF knockdown exhibited modest enhancements of cell killing in response to IR exposure compared to parental cells, while ATM^−/− cells expressing ΔhMOF did not (Fig. 9). However, cells overexpressing hMOF exhibited decreased IR sensitivity for cell killing (Fig. 9). Thus, cells expressing ΔhMOF and cells with hMOF knockdown display karyotypic instability and decreased cell survival after IR exposure. All of these cellular phenotypes could be linked with defective chromosomal repair, as has been reported for cells deficient for ATM (36).

One way to address whether DNA repair is affected by hMOF function is to compare cell cycle stage-specific chromosomal aberrations in cells with and without expression of the ΔhMOF according to the previously described procedure (45). Cells expressing ΔhMOF exhibited a significant increase in residual IR-induced G₁ chromosomal aberrations seen at metaphases (Fig. 10A). To determine whether defective repair can be documented in cells expressing ΔhMOF in phases of the cell cycle other than G₁, S-phase-specific chromosome aberrations in such cells were evaluated. Cells expressing ΔhMOF displayed higher frequencies of S-phase-specific chromatid and chromosomal aberrations per metaphase when compared to parental cells (Fig. 10B). Similar results were obtained when G₂-phase-specific chromosome aberrations were evaluated (Fig. 10C). The overexpression of hMOF resulted in a modest, although not statistically significant, decrease in residual chromosome aberrations in all the three phases of the cell cycle (Fig. 10). In addition, we used a previously described biochemical approach (38) to determine the influence of hMOF on DNA DSB repair after IR exposure. As illustrated in Fig. 11, cells expressing ΔhMOF (fragment 1 in Fig. 1A) have higher amounts of residual DNA DSBs, and cells overexpressing hMOF have relatively less residual DNA DSBs, than control cells. The observation that the expression of ΔhMOF (fragment 1 in Fig. 1A) results in a higher frequency of chromosome aberrations irrespective of which cell cycle phase was analyzed supports the idea that the inactivation of hMOF reduces the cell's capacity for global DNA repair.

A characteristic hallmark of A-T cells is the loss of the DNA-damage-induced cell cycle checkpoint (24, 36, 46). Cells expressing ΔhMOF displayed neither a significant increase in G₁ phase cells nor a decrease in S-phase entry after IR treatment compared to control cells that showed a significant decrease in S-phase entry, indicating G₁ arrest (Fig. 12A). The loss of the G₁ checkpoint seen in cells expressing ΔhMOF is similar to that in A-T cells (53), indicating that this phenotype is mimicked in cells expressing ΔhMOF. In addition to the G₁ checkpoint, IR causes a transient inhibition of DNA replication. Cells deficient in ATM function exhibit radioresistant DNA synthesis, an indicator of the loss of the S-phase checkpoint. Cells expressing ΔhMOF showed decreased inhibition of DNA synthesis, compared to the parental cells, following IR exposure (Fig. 12B). A-T cells also show defective G₂/M checkpoints after DNA damage, and ATM signaling has been shown to be required for the G₂/M checkpoint in response to IR (54). The efficiency of G₂ checkpoint control was evaluated by measuring the proportion of cells in mitosis (mitotic index) after IR exposure (11). Cells expressing ΔhMOF exhibited a decrease in mitotic index of approximately 25%, while the decrease in the parental control cells was approximately 68% (Fig. 12C). These results suggest that the inactivation of hMOF does lead to a defective G₂ checkpoint.