Lysine 40 acetylation of α-tubulin (Ac-α-tubulin), catalyzed by the acetyltransferase αTAT1, marks stabilized microtubules. Recently, there is growing evidence to suggest crosstalk between the DNA damage response (DDR) and microtubule organization; we therefore investigated whether αTAT1 is involved in the DDR. Following treatment with DNA-damaging agents, increased levels of Ac-α-tubulin were detected. We also observed significant induction of Ac-α-tubulin after depletion of DNA repair proteins, suggesting that αTAT1 is positively regulated in response to DNA damage. Intriguingly, αTAT1 depletion decreased DNA damage-induced replication protein A (RPA) phosphorylation and foci formation. Moreover, DNA damage-induced cell cycle arrest was significantly delayed in αTAT1-depleted cells, indicating defective checkpoint activation. The checkpoint defects seen upon αTAT1 deficiency were restored by expression of wild-type αTAT1, but not by αTAT1-D157N (a catalytically inactive αTAT1), indicating that the role of αTAT1 in the DDR is dependent on enzymatic activity. Furthermore, αTAT1-depleted direct repeat GFP (DR-GFP) U2OS cells had a significant decrease in the frequency of homologous recombination repair. Collectively, our results suggest that αTAT1 may play an essential role in DNA damage checkpoints and DNA repair through its acetyltransferase activity.

Lys40 (K40) acetylation in α-tubulin is catalyzed by the acetyltransferase αTAT1 (also known as Mec17) and is known to be enriched in stable microtubules, such as those comprising the mitotic spindle and cilia (Akella et al., 2010; Kalebic et al., 2013; Janke and Bulinski, 2011; Janke and Montagnac, 2017). Deacetylation of this residue is catalyzed by two enzymes, the NAD-independent deacetylase HDAC6 and the NAD-dependent deacetylase SIRT2 (Hubbert et al., 2002; North et al., 2003). These three enzymes control the level of α-tubulin K40 acetylation in the cells. α-Tubulin K40 acetylation has reported to be involved in microtubule dynamics, cell migration, autophagy, intracellular trafficking and cell adhesion (Janke and Bulinski, 2011; Magiera and Janke, 2014; Perdiz et al., 2011); however, the precise mechanisms through which K40 acetylated α-tubulin regulates those functions still remains to be elucidated.

Depletion of MEC-17 or αTAT1 in Tetrahymena, C. elegans and mice does not noticeably affect their normal growth (Akella et al., 2010; Kalebic et al., 2013; Kim et al., 2013; Topalidou et al., 2012). However, a recent report demonstrated that depletion of αTAT1 in cancer cell lines impairs cell growth and induces mitotic catastrophe (Chien et al., 2016), suggesting that there is a differential requirement of αTAT1 between different cell types. Interestingly, enhanced levels of microtubule acetylation have been observed in head-neck, pancreatic and breast cancers (Boggs et al., 2015; Saba et al., 2014; Seeley et al., 2009). Furthermore, a recent report demonstrated that enhanced α-tubulin K40 acetylation promotes metastatic activity in breast cancer (Boggs et al., 2015; Lee et al., 2018). These observations imply that α-tubulin acetylation may serve as both a diagnostic biomarker and therapeutic target for the therapy of malignant tumors.

Recently, it has been shown that fission yeast with defective DNA repair functions displayed elongated morphology associated with microtubule stabilization (Graml et al., 2014). More recently, it has been demonstrated that microtubule stabilization is required for intracellular trafficking of DNA repair proteins in response to DNA damage (Poruchynsky et al., 2015), revealing a link between the DNA damage response (DDR) and microtubule networks. Given the involvement of α-tubulin K40 acetylation in microtubule structure and stability, we hypothesized that αTAT1 might have function in the DDR, such as cell cycle checkpoint activation and DNA repair. In the present study, we observed that treatment with DNA-damaging agents or depletion of DNA repair genes induced α-tubulin K40 acetylation, suggesting that αTAT1 is positively regulated in response to genotoxic stress. Intriguingly, depletion of αTAT1 significantly decreased DNA damage-induced phosphorylation of replication protein A (RPA) and CHK1 (also known as CHEK1), and delayed DNA damage-induced S and G2/M cell cycle arrest. These findings suggest that αTAT1 is involved in the DDR. We also examined the role of enzymatic activity of αTAT1 in the DDR and observed that checkpoint defects in αTAT1-depleted cells were restored by expression of wild-type αTAT1, but not by αTAT1-D157N (a catalytically inactive αTAT1). Moreover, αTAT1-depleted direct repeat GFP (DR-GFP) U2OS cells showed defective DNA double strand break (DSB)-induced homologous recombination (HR) repair. Taken together, our results suggest that αTAT1 may play an essential role in DNA damage checkpoints and DNA repair through its acetyltransferase activity.

The α-tubulin K40 acetylation is induced following treatment with DNA damaging agents

HeLa cells were treated with camptothecin (CPT), hydroxyurea (HU) and ultraviolet light (UV) to induce DNA damage, and then α-tubulin K40 acetylation was analyzed as an indicator for microtubule stabilization (Fig. 1). An expanded cytoskeleton, increased intensity of α-tubulin K40 acetylation and increased resistance of microtubules to nocodazole treatment were observed in CPT-treated cells when compared to the untreated cells, indicating that DNA damage-induced microtubule stabilization had occurred (Fig. 1A; Fig. S1). Consistent with this, following treatment with CPT and HU, significantly increased levels of α-tubulin K40 acetylation (Ac-α-tubulin) were detected by immunoblotting (Fig. 1B,C). In response to DNA damage, RPA2 phosphorylation (RPA2 Ser33) and H2AX phosphorylation (γH2AX) are indicative of activation of DNA damage checkpoints (Shiloh, 2001; Zou and Elledge, 2003). To further analyze the dynamics of α-tubulin K40 acetylation in response to DNA damage, UV-radiated cells were collected at various time points after treatment (Fig. 1D,E). At 4–6 h after UV exposure, significant increases in α-tubulin K40 acetylation were detected. This was correlated with morphological changes in UV-radiated HeLa cells (data not shown).

Fig. 1.

α-tubulin K40 acetylation is induced by DNA damage. (A) HeLa cells were either left untreated or treated with CPT (100 ng/ml) for 16 h. Cells were immunostained with anti-acetyl α-tubulin antibody. Representative fields are shown. Scale bars: 10 µm. (B) HeLa cells were either left untreated (NT) or treated with either CPT (100 ng/ml) or HU (2 mM) for 16 h. Whole-cell lysates were analyzed by immunoblotting with the indicated antibodies. (C) Quantification of acetylated α-tubulin in B. The ratios of Ac-α-tubulin/α-tubulin were determined by densitometry analysis of the immunoblot bands. (D) HeLa cells were irradiated with UV (30J/m2) and collected at the indicated time points after irradiation. Whole cell lysates were analyzed by immunoblotting with indicated antibodies. (E) Quantification of acetylated α-tubulin in (D). The ratios of Ac-α-tubulin/α-tubulin were determined by densitometry analysis of the immunoblot bands. Values in C and E are means±s.e.m. from three independent experiments. *P<0.05, **P<0.001 (Student's t-test).

Fig. 1.

α-tubulin K40 acetylation is induced by DNA damage. (A) HeLa cells were either left untreated or treated with CPT (100 ng/ml) for 16 h. Cells were immunostained with anti-acetyl α-tubulin antibody. Representative fields are shown. Scale bars: 10 µm. (B) HeLa cells were either left untreated (NT) or treated with either CPT (100 ng/ml) or HU (2 mM) for 16 h. Whole-cell lysates were analyzed by immunoblotting with the indicated antibodies. (C) Quantification of acetylated α-tubulin in B. The ratios of Ac-α-tubulin/α-tubulin were determined by densitometry analysis of the immunoblot bands. (D) HeLa cells were irradiated with UV (30J/m2) and collected at the indicated time points after irradiation. Whole cell lysates were analyzed by immunoblotting with indicated antibodies. (E) Quantification of acetylated α-tubulin in (D). The ratios of Ac-α-tubulin/α-tubulin were determined by densitometry analysis of the immunoblot bands. Values in C and E are means±s.e.m. from three independent experiments. *P<0.05, **P<0.001 (Student's t-test).

The α-tubulin K40 acetylation is induced after depletion of DNA repair-related proteins

It has been shown that fission yeast with defective DNA repair functions display elongated morphology with stable microtubules (Graml et al., 2014). To further investigate the mechanism underlying microtubule stabilization after genotoxic stress, we determined the level of α-tubulin K40 acetylation following depletion of cell cycle control and DNA repair proteins in HeLa cells (Fig. 2). Cells transfected with siRNAs directed against genes encoding cell cycle and checkpoint proteins (ATM, ATR and TopBP1), HR DNA repair proteins (BRCA1, BRCA2 and PALB2), non-homologous end-joining proteins (CtiP, KU70; also known as RBBP8 and XRCC6, respectively), a nucleotide excision repair protein (ERCC1), a mismatch repair protein (MSH2) and a DNA crosslink repair gene (FANCA) were examined for the induction of α-tubulin K40 acetylation by immunofluorescence and immunoblotting. γH2AX immunostaining was used to monitor the amount of spontaneous DNA damage following siRNA transfection (Fig. 2A) and the efficacy of each siRNA was verified by immunoblotting with their specific antibodies (Fig. 2B). Intriguingly, although not all of those depletions caused significant induction of α-tubulin K40 acetylation, depletion of ATM, ATR, BRCA1, CtiP, MSH2 and FANCA led to significantly increased levels of α-tubulin K40 acetylation compared with that seen in control siRNA-treated cells (Fig. 2A,B; Fig. S2). Co-staining with Ac-α-tubulin and γH2AX revealed that induction of Ac-α-tubulin correlated with an increased intensity of γH2AX. The finding that there was significant change in Ac-α-tubulin from other samples could be explained by inadequate knockdown, but it is also possible that an insufficient amount of spontaneous DNA damage was generated after siRNA transfection, which would not be enough to induce Ac-α-tubulin. Taken together, these results indicate that genome instability caused by either DNA-damaging agents or defective DNA repair function induces α-tubulin K40 acetylation, suggesting that αTAT1 is positively regulated in response to genotoxic stress.

Fig. 2.

α-tubulin K40 acetylation is induced following depletion of DNA repair-related proteins. (A,B) HeLa cells were transfected with the indicated siRNAs for 72 h. (A) Cells were co-immunostained with antibodies for acetyl-α-tubulin (green) and γH2AX (red). All images in each antibody group were collected using the same exposure time for the comparison. Representative fields are shown. Scale bars: 10 µm. (B) Whole-cell lysates were analyzed by immunoblotting with indicated antibodies.

Fig. 2.

α-tubulin K40 acetylation is induced following depletion of DNA repair-related proteins. (A,B) HeLa cells were transfected with the indicated siRNAs for 72 h. (A) Cells were co-immunostained with antibodies for acetyl-α-tubulin (green) and γH2AX (red). All images in each antibody group were collected using the same exposure time for the comparison. Representative fields are shown. Scale bars: 10 µm. (B) Whole-cell lysates were analyzed by immunoblotting with indicated antibodies.

DNA damage-induced RPA and CHK1 phosphorylation are decreased in αTAT1-depleted cells

Based on our hypothesis that αTAT1 is positively regulated after genotoxic stress, we next explored the role of αTAT1 in the DNA damage signaling (Fig. 3). αTAT1 was depleted using specific siRNAs in HeLa cells. Because there is no reliable antibody for the detection of endogenous αTAT1 in our hands, the level of anti α-tubulin K40 acetylation was used to determine efficiency of αTAT1 depletion. As expected, the levels of α-tubulin K40 acetylation were greatly reduced in the αTAT1-knockdown cells, using either αTAT1 #1 or αTAT1 #2 (Fig. S3).While both siRNAs efficiently knocked down αTAT1, siRNA#1 was used to perform all the knockdown experiments throughout this study. Immunoblotting analyses were performed to address the impact of αTAT1 depletion in DNA damage-induced checkpoint activation. Consistent with a previous report (Chien et al., 2016), the level of γH2AX was slightly higher in αTAT1-depleted cells compared to control cells in the absence of DNA-damaging agents (Fig. 3A). Following treatment with DNA damaging agents, the phosphorylation of CHK1 and RPA2 was significantly decreased in αTAT1-depleted cells. Similar results were observed in two different cell lines, HCT116 (a human colon cancer cell line) and A549 (a human lung carcinoma cell line) (Fig. S4). Next, to further analyze the defect in RPA phosphorylation after αTAT1 depletion, a cytoplasmic and nucleoplasmic fraction and a chromatin-enriched fraction were prepared (Fig. 3B,C). The defective RPA phosphorylation appeared more evident in the chromatin-enriched fraction than in the cytoplasmic and nucleoplasmic fraction. As it has been shown that chromatin-bound RPA phosphorylation is essential for DNA damage signaling and repair (Maréchal and Zou, 2015; Zou and Elledge, 2003), this result suggests that αTAT1 participates in the DDR at least in part through regulating RPA phosphorylation.

Fig. 3.

DNA damage-induced checkpoint activation is defective in αTAT1-depleted cells. (A–C) HeLa cells were transfected with either control siRNA or αTAT1 siRNAs. At 48 h after transfection, cells were either left untreated (NT) or treated with HU (1 mM for 16 h), CPT (100 nM for 16 h) and UV (30 J/m2 for 8 h). (A) Whole-cell lysates were prepared and analyzed by immunoblotting with indicated antibodies. (B,C) Cell pellets collected were fractionated into a soluble fraction (Fr.; cytoplasmic and nucleoplasmic proteins) and chromatin-enriched fraction (chromatin and nuclear matrix proteins). Samples from each fraction were immunoblotted with the indicated antibodies. α-tubulin and lamin B1 were used as loading controls.

Fig. 3.

DNA damage-induced checkpoint activation is defective in αTAT1-depleted cells. (A–C) HeLa cells were transfected with either control siRNA or αTAT1 siRNAs. At 48 h after transfection, cells were either left untreated (NT) or treated with HU (1 mM for 16 h), CPT (100 nM for 16 h) and UV (30 J/m2 for 8 h). (A) Whole-cell lysates were prepared and analyzed by immunoblotting with indicated antibodies. (B,C) Cell pellets collected were fractionated into a soluble fraction (Fr.; cytoplasmic and nucleoplasmic proteins) and chromatin-enriched fraction (chromatin and nuclear matrix proteins). Samples from each fraction were immunoblotted with the indicated antibodies. α-tubulin and lamin B1 were used as loading controls.

DNA damage-induced RPA foci formation is defective in αTAT1-depleted cells

Hyper-phosphorylation of chromatin-bound RPA2 results in the recruitment of other DNA repair proteins to the sites of DNA damage (Maréchal and Zou, 2015; Zou and Elledge, 2003). We next examined RPA recruitment to the DNA damage sites after αTAT1 depletion (Fig. 4). Following treatment with the DSB-inducing agent CPT, there were fewer αTAT1-depleted cells showing RPA1 and RPA2 foci (43% and 40%, respectively) than control cells (90–95%) (Fig. 4A,B). In contrast, there was no significant difference in γH2AX intensity between control and αTAT1-depleted cells (Fig. 4A). In response to UV radiation, a similar reduction was observed (data not shown). This result suggests that αTAT1 is required for DNA damage-induced RPA phosphorylation and foci formation.

Fig. 4.

DNA damage-inducible RPA foci formation is defective in αTAT1-depleted cells. (A,B) HeLa cells were transfected with either control siRNA or αTAT1 siRNAs. At 48 h after transfection, cells were either left untreated (NT) or treated with CPT (100 ng/ml for 16 h) and then immunostained with the indicated antibodies. (A) Representative images of cells treated with CPT are shown. Scale bars: 10 µm. (B) Quantification of the percentage of cells with RPA1 and RPA2 foci shown in A, respectively. Values represent the mean±s.e.m., examined for least 200 nuclei each in three independent experiments. *P<0.05, **P<0.001 (Student's t-test).

Fig. 4.

DNA damage-inducible RPA foci formation is defective in αTAT1-depleted cells. (A,B) HeLa cells were transfected with either control siRNA or αTAT1 siRNAs. At 48 h after transfection, cells were either left untreated (NT) or treated with CPT (100 ng/ml for 16 h) and then immunostained with the indicated antibodies. (A) Representative images of cells treated with CPT are shown. Scale bars: 10 µm. (B) Quantification of the percentage of cells with RPA1 and RPA2 foci shown in A, respectively. Values represent the mean±s.e.m., examined for least 200 nuclei each in three independent experiments. *P<0.05, **P<0.001 (Student's t-test).

αTAT1-depleted cells have defects in cell cycle arrest after DNA damage

The cell cycle checkpoints play an important role in the control system by sensing DNA damage and inducing a cell cycle arrest in response, until the damage is repaired (Sherr, 2000). To further analyze the impact of αTAT1 deficiency in DNA damage-induced checkpoint activation, we examined cell cycle progression of αTAT1-depleted cells after DNA damage (Fig. 5). αTAT1-depleted cells had a lower proportion of cells in the S phase compared to control siRNA-transfected cells in the absence of CPT (Fig. 5A). In the presence of CPT, the majority of control siRNA-treated cells accumulated in late S and G2 phase, but αTAT1-depleted cells were in G1 and early to mid S phase, indicating a significantly delayed cell cycle arrest. A similar result was observed with another DSB-inducing agent bleomycin (data not shown). Consistent with this, CPT-induced RPA2 and CHK1 phosphorylation was largely reduced in αTAT1-depleted cells (Fig. 5B). These results suggest that αTAT1 is required for efficient DNA damage-induced checkpoint activation and cell cycle arrest.

Fig. 5.

αTAT1-depleted cells have defects in cell cycle arrest after DNA damage. (A,B) HeLa cells were transfected with either control siRNA or αTAT1 siRNAs. At 48 h after transfection, cells were either left untreated (NT) or treated with CPT (100 ng/ml for 16 h). (A) For cell cycle analysis, histograms (upper panels) representing PI staining and dot plots (lower panels) demonstrating BrdU labeling are presented. (B) Whole-cell lysates were prepared and analyzed by immunoblotting with indicated antibodies.

Fig. 5.

αTAT1-depleted cells have defects in cell cycle arrest after DNA damage. (A,B) HeLa cells were transfected with either control siRNA or αTAT1 siRNAs. At 48 h after transfection, cells were either left untreated (NT) or treated with CPT (100 ng/ml for 16 h). (A) For cell cycle analysis, histograms (upper panels) representing PI staining and dot plots (lower panels) demonstrating BrdU labeling are presented. (B) Whole-cell lysates were prepared and analyzed by immunoblotting with indicated antibodies.

Wild-type αTAT1, but not αTAT1-D157N, restores defective checkpoint activation in αTAT1-depleted cells

We next examined whether wild-type αTAT1 cDNA can restore checkpoint defects in αTAT1-depleted cells (Fig. 6). Expression of siRNA-resistant wild-type αTAT1 in αTAT1-depleted cells resulted in restoration of the levels of α-tubulin K40 acetylation, RPA2 phosphorylation and CHK1 phosphorylation (Fig. 6A). Furthermore, expression of wild-type αTAT1 was also able to restore delayed S and G2/M arrest in response to CPT in αTAT1-depleted cells (Fig. 6B). These results further suggest that αTAT1 is required for DNA damage-induced checkpoint activation. Next, in order to clarify whether αTAT1 enzyme activity is required in the DDR, the effect of catalytically inactive αTAT1-D157N (Shida et al., 2010) was examined (Fig. 7). In contrast to wild-type αTAT1, αTAT1-D157N failed to restore defective cell cycle checkpoints in αTAT1-depleted cells. These findings suggest that the function of αTAT1 in DNA damage checkpoints is dependent on its acetylation activity.

Fig. 6.

Expression of wild-type αTAT1 restores checkpoint defects in αTAT1-depleted cells. (A,B) HeLa cells stably expressing either empty vector (EV) or siRNA-resistant wild-type αTAT1 cDNA were transfected with the indicated siRNAs. At 48 h after transfection, cells were either left untreated (NT) or treated with CPT (100 ng/ml for 16 h). (A) Whole-cell lysates were prepared and analyzed by immunoblotting with the indicated antibodies. (B) Cells were analyzed by flow cytometry. Representative histograms for cell cycle distribution are presented. The percentages of cells in G1, S, and G2/M phase are indicated.

Fig. 6.

Expression of wild-type αTAT1 restores checkpoint defects in αTAT1-depleted cells. (A,B) HeLa cells stably expressing either empty vector (EV) or siRNA-resistant wild-type αTAT1 cDNA were transfected with the indicated siRNAs. At 48 h after transfection, cells were either left untreated (NT) or treated with CPT (100 ng/ml for 16 h). (A) Whole-cell lysates were prepared and analyzed by immunoblotting with the indicated antibodies. (B) Cells were analyzed by flow cytometry. Representative histograms for cell cycle distribution are presented. The percentages of cells in G1, S, and G2/M phase are indicated.

Fig. 7.

The acetyltransferase activity of αTAT1 is required to restore checkpoint defects in αTAT1-depleted cells. (A) HeLa cells stably expressing either empty vector (EV) or siRNA-resistant αTAT1-D157N cDNA were transfected with the indicated siRNAs. At 48 h after transfection, cells were treated with CPT (100 ng/ml for 16 h). Whole-cell lysates were prepared and analyzed by immunoblotting with indicated antibodies. (B) HeLa cells stably expressing either empty vector (EV) or siRNA-resistant αTAT1-D157N cDNA were transfected with the indicated siRNAs. At 48 h after transfection, cells were either left untreated (NT) or treated with CPT (100 ng/ml for 16 h). Then, cells were analyzed by flow cytometry. Representative histograms for cell cycle distribution are presented. The percentages of cells in G1, S, and G2/M phase are indicated.

Fig. 7.

The acetyltransferase activity of αTAT1 is required to restore checkpoint defects in αTAT1-depleted cells. (A) HeLa cells stably expressing either empty vector (EV) or siRNA-resistant αTAT1-D157N cDNA were transfected with the indicated siRNAs. At 48 h after transfection, cells were treated with CPT (100 ng/ml for 16 h). Whole-cell lysates were prepared and analyzed by immunoblotting with indicated antibodies. (B) HeLa cells stably expressing either empty vector (EV) or siRNA-resistant αTAT1-D157N cDNA were transfected with the indicated siRNAs. At 48 h after transfection, cells were either left untreated (NT) or treated with CPT (100 ng/ml for 16 h). Then, cells were analyzed by flow cytometry. Representative histograms for cell cycle distribution are presented. The percentages of cells in G1, S, and G2/M phase are indicated.

αTAT1-depleted cells have severe defects in DSB-induced HR-mediated repair

When cells encounter DNA damage in the form of DSBs, the recruitment of RPA to the DNA damage sites and hyper-phosphorylation is an important initial step for HR-mediated repair (Krejci et al., 2012; Maréchal and Zou, 2015; Shao et al., 1999). In response to CPT treatment, we observed severe defects in RPA phosphorylation and foci formation in αTAT1-depleted cells (Figs 3 and 4). Furthermore, we found that the frequency of cells with BRCA1 foci was significantly lower in αTAT1-depleted cells (30%) than control cells (70%) after CPT treatment (Fig. 8A,B). As BRCA1 has been shown to have essential role in HR-mediated repair (Isono et al., 2017; Moynahan et al., 1999; Snouwaert et al., 1999), we next investigated the influence of αTAT1 knockdown in DSB repair by HR using a DR-GFP assay, in which reconstitution of a defective GFP gene is dependent upon HR-mediated repair of an introduced DSB (Weinstock et al., 2006) (Fig. 8C–E). The efficiency of HR-mediated DSB repair in αTAT1-depleted cells was reduced by ∼50% compared to control siRNA-transfected cells, indicating a defective HR-mediated repair in αTAT1-depleted cells. To further investigate the effect of α-tubulin K40 acetylation in HR-mediated repair, we measured the HR frequency under conditions in which α-tubulin K40 acetylation is enhanced, in contrast to αTAT1 depletion. Since HDAC6 is known as a major α-tubulin deacetylase (Hubbert et al., 2002), HDAC6 depletion was examined for this purpose (Fig. 8C). As expected, significantly increased level of α-tubulin K40 acetylation was observed after HDAC6 depletion, whereas, a significant reduction in HR frequency was detected (Fig. 8D,E). Furthermore, cells depleted of both αTAT1 and HDAC6 showed similar reduction of HR frequency when compared with αTAT1 siRNA alone. Given that HDAC inhibitors are known to suppress the expression of HR repair genes (Groselj et al., 2013; Kotian et al., 2011), defective HR-mediated repair in HDAC6-depleted cells could be caused by multiple mechanisms. Thus, we still need to clarify the relationship between the level of α-tubulin K40 acetylation and HR efficiency. Overall, these results suggest that αTAT1 may be required for efficient HR-mediated DSB repair after DNA damage.

Fig. 8.

αTAT1-depleted U2OS/DRGFP cells shows decreased HR efficiency. (A,B) HeLa cells were transfected with either control siRNA or αTAT1 siRNAs. At 48 h after transfection, cells were either left untreated (NT) or treated with CPT (100 ng/ml for 16 h) and then immunostained with anti-BRCA1 antibody. (A) Representative images of cells treated with CPT are shown. Scale bar: 10 µm. (B) Quantification of cells with BRCA1 foci shown in A. Values represent the mean±s.e.m., examined for at least 200 nuclei each in three independent experiments. *P<0.05 (Student's t-test). (C–E) DR-GFP/U2OS cells were transfected with indicated siRNAs. At 24 h after siRNA transfection, DR-GFP/U2OS cells were transfected with I-SceI plasmid (pCBASce) to induce DSBs and further incubated for 48 h. (C) The efficacy of each siRNA was verified by immunoblotting with their specific antibodies. (D) Representative flow cytometry plots of GFP+ cells are shown. The percentages of GFP+ cells are indicated. (E) Quantification of the data shown in D. Values were normalized for the transfection efficiency and are displayed as mean±s.e.m. GFP+ frequencies relative to that of control siRNA-treated cells are from five independent experiments. (F) A proposed model for the role of αTAT1 in the DDR. DNA damage could positively regulate αTAT1 by an unknown mechanism, leading to increased K40 acetylation levels of α-tubulin and other substrate(s) if they exist. K40 acetylation of α-tubulin might be partly responsible for DNA damage-induced microtubule stabilization, which, in turn, could promote the DDR, such as checkpoint activation and HR-mediated repair. Furthermore, acetylation of unknown substrates might be involved in this process. The other branch, depicting an acetylation-independent pathway (dotted line) is unlikely involved in the DDR on the basis of our results (Figs 6 and 7). See Discussion for details.

Fig. 8.

αTAT1-depleted U2OS/DRGFP cells shows decreased HR efficiency. (A,B) HeLa cells were transfected with either control siRNA or αTAT1 siRNAs. At 48 h after transfection, cells were either left untreated (NT) or treated with CPT (100 ng/ml for 16 h) and then immunostained with anti-BRCA1 antibody. (A) Representative images of cells treated with CPT are shown. Scale bar: 10 µm. (B) Quantification of cells with BRCA1 foci shown in A. Values represent the mean±s.e.m., examined for at least 200 nuclei each in three independent experiments. *P<0.05 (Student's t-test). (C–E) DR-GFP/U2OS cells were transfected with indicated siRNAs. At 24 h after siRNA transfection, DR-GFP/U2OS cells were transfected with I-SceI plasmid (pCBASce) to induce DSBs and further incubated for 48 h. (C) The efficacy of each siRNA was verified by immunoblotting with their specific antibodies. (D) Representative flow cytometry plots of GFP+ cells are shown. The percentages of GFP+ cells are indicated. (E) Quantification of the data shown in D. Values were normalized for the transfection efficiency and are displayed as mean±s.e.m. GFP+ frequencies relative to that of control siRNA-treated cells are from five independent experiments. (F) A proposed model for the role of αTAT1 in the DDR. DNA damage could positively regulate αTAT1 by an unknown mechanism, leading to increased K40 acetylation levels of α-tubulin and other substrate(s) if they exist. K40 acetylation of α-tubulin might be partly responsible for DNA damage-induced microtubule stabilization, which, in turn, could promote the DDR, such as checkpoint activation and HR-mediated repair. Furthermore, acetylation of unknown substrates might be involved in this process. The other branch, depicting an acetylation-independent pathway (dotted line) is unlikely involved in the DDR on the basis of our results (Figs 6 and 7). See Discussion for details.

In the present study, we investigated the potential role of αTAT1 in the DDR. Depletion of DNA repair-related proteins significantly increased the level of α-tubulin K40 acetylation, suggesting that induction of genome instability promoted microtubule stabilization. After depletion of αTAT1, DNA damage-induced RPA hyper-phosphorylation and nuclear foci formation were largely impaired. In addition, CPT-induced cell cycle arrest was delayed in αTAT1-depleted cells. Furthermore, αTAT1-depleted DR-GFP U2OS cells showed defective DSB-induced HR-mediated repair. Although our findings are limited in terms of the detailed mechanisms through which αTAT1 affects the DDR, these results support our hypothesis that αTAT1 may be required for efficient checkpoint activation and DNA repair.

The K40 acetylation status of α-tubulin is defined by the opposing activities of tubulin acetyl transferases, αTAT1 and lysine deacetylases, namely HDAC6 and SIRT2 (Hubbert et al., 2002; Janke and Bulinski, 2011; North et al., 2003). Unlike HDAC6 and SIRT2, which have multiple substrates, α-tubulin is known as the major substrate of αTAT1 (Akella et al., 2010; Kalebic et al., 2013). In this study, we have focused on positive regulation of αTAT1 in response to DNA damage, but it would be also conceivable that suppressed HDAC6 or SIRT2 could increase the level of α-tubulin K40 acetylation following DNA damage. However, this seems unlikely because it has been recently shown that de-acetylation by HDAC6 and SIRT2 is required for efficient DNA repair (Zhang et al., 2016, 2019). Therefore we speculate that induction of α-tubulin K40 acetylation in response to DNA damage might be primarily caused by αTAT1 rather than inhibition of either HDAC6 or SIRT2.

How is the catalytic activity of αTAT1 regulated upon DNA damage?

In response to DNA damage, activated ATR and ATM phosphorylate numerous DDR and repair proteins at single or multiple Ser/Thr-Gln (S/T-Q) sites (Shiloh, 2001). Although human αTAT1 only contains a single SQ/TQ motif at Ser160, it will be interesting to see whether αTAT1 is phosphorylated by ATM and/or ATR, and whether this modification enhances the catalytic activity of αTAT1. Previously it has been demonstrated that AMP-activated protein kinase (AMPK) enhances αTAT1 activity by means of phosphorylation upon osmotic or oxidative stress (Mackeh et al., 2014). Most recently, it has been shown that TGF-β-activated kinase 1 (TAK1; also known as MAP3K7), activated through a diverse set of intra- and extracellular stresses, directly phosphorylates αTAT1 at Ser237, which causes an increase its catalytic activity (Shah et al., 2018). Therefore, it would be interesting to examine whether Ser237 in αTAT1 is phosphorylated following DNA damage. Alternatively, αTAT1 contains multiple lysine residues close to the N-terminus of αTAT1. Considering that auto-acetylation increases the acetylation activity of αTAT1 (Kalebic et al., 2013), it would be possible that αTAT1 is acetylated by other acetyltransferases engaged in the DDR and increase its catalytic activity (Bharti and Brosh, 2016; Sun et al., 2005; Zhao et al., 2017).

How does αTAT1 affect checkpoint activation?

The precise role of α-tubulin K40 acetylation in microtubule stability has long remained controversial. However, recent reports provide new evidence in support of structural stability, as acetylation appears to protect long-lived MTs against mechanical ageing and resistance to mechanical breakage (Portran et al., 2017; Xu et al., 2017). Furthermore, Poruchynsky et al. have demonstrated that microtubule stabilization is required for intracellular trafficking of DNA repair proteins in response to DNA damage (Poruchynsky et al., 2015), revealing a link between the DDR and microtubule networks. Therefore, there would be several possibilities for how αTAT1 could be involved in the DDR. First, DNA damage-induced α-tubulin K40 acetylation may contribute to microtubule stabilization that, in turn, promotes intracellular transport of DNA repair proteins to the nucleus, resulting in increased RPA phosphorylation and foci formation. Second, it would be also conceivable that αTAT1 may have another substrate than α-tubulin, and acetylation of this alternative substrate is required for checkpoint activation and maintenance (Fig. 8F). Recent studies have shown that many of DDR-related proteins including ATM, RPA1, TopBP1 and ATRIP are regulated by acetylation/deacetylation (Bharti and Brosh, 2016; Liu et al., 2014; Wang et al., 2014; Zhang et al., 2016; Zhao et al., 2017). Thus, it would be interesting to investigate the functional link between αTAT1 and these proteins. Finally, the checkpoint defects seen upon αTAT1 deficiency could be an indirect effect of global cellular defects by lack of α-tubulin K40 acetylation.

It has been recently shown that metastatic breast cancer cells have a high level of α-tubulin K40 acetylation and reduction of its α-tubulin K40 acetylation significantly inhibits migration (Boggs et al., 2015), suggesting that αTAT1 inhibition could represent a new therapeutic strategy for metastatic breast cancer. In this study, we showed that αTAT1 downregulation impaired DNA damage-induced checkpoint activation and HR-mediated DSB repair. Although the detailed mechanisms by which αTAT1 affects the DRR remain to be elucidated, our results suggest that αTAT1 may be required for efficient checkpoint activation and DNA repair. We believe, therefore, the finding in this study has an important implication for the use of αTAT1 inhibitors in cancer therapies that enhance cytotoxicity to DNA damaging anti-cancer drugs.

Cell culture

HeLa, DR-GFP U2OS and A549 (ATCC, CCL-185) cells were grown in DMEM (Dulbecco's modified Eagle's medium; Invitrogen) supplemented with 10% (v/v) FBS (fetal bovine serum; Hyclone). HCT116 cells (ATCC, CCL 247) were grown in McCoy's 5A medium plus 10% FBS.

Antibodies

Antibodies used in this study were as follows: anti-α-tubulin (1:4000; T5168) and anti-Flag M2 (1:1000; F3165) antibodies (Sigma-Aldrich, St. Louis, MO); anti-acetyl-α-tubulin (α-tubulin K40 acetylation; 1:500; #5335), anti-CHK1 S345 (1:500; #2341), anti-CHK2 T68 (1:500; #2661), anti-ATR (1:1000; #2790), anti-HDAC6 (1:500; #7558) and anti-RPA1 (1:500; #2267) antibodies (Cell Signaling Technology, Inc., MA); anti-ATM (1:1000; A300-299A), anti-ATRIP (1:1000; A300-095A), anti-TopBP1 (1:1000; A300-111A), anti-RPA2 S33 (1:1000; A300-246A), anti-RPA2 S4/S8 (1:1000; A300-245A), anti-PALB2 (1:1000; A301-246A), and anti-Ctip (1:500; A300-488A) antibodies (Bethyl Laboratories, Inc., Montgomery, TX); anti-RPA2 (1:500; NA19L), anti-BRCA1 (1:200; OP92) and anti-BRCA2 (1:200; OP95) antibodies (EMD Millipore, Temecula, CA); anti-KU70 (1:1000; N3H10; Neomarkers, Inc., Fremont, CA); anti-γH2AX (1:1000; #05-636; Upstate Biotechnology, Lake Placid, NY); anti-MSH2 (1:200; sc494), anti-ERCC1 (1:200; sc17809) and anti-Lamin B1 (1:200; sc374015) antibodies (Santa Cruz Biotechnology, Santa Cruz, CA); and anti-FANCA (1:500; kindly provided by the Fanconi Anemia Research Fund, Eugene, OR; www.fanconi.org).

DNA damage induction

For UV treatment, the cells were washed with PBS and irradiated using UVC crosslinker (Stratagene, La Jolla, CA) at the indicated dose for the indicated times. For drug treatment, the cells were incubated with various DNA-damaging agents and inhibitors at the indicated concentrations and times. Camptothecin (CPT), hydroxyurea (HU) and bleomycin were purchased from Sigma-Aldrich.

siRNA transfection

siRNAs were synthesized by Qiagen (USA) and Bioneer (Korea). The sense sequence of αTAT1 #1 and #2 siRNAs are 5′-GGGAAACUCACCAGAACGAdTdT-3′ and 5′-CGAGAACUCUUCCAGUAUAdTdT-3′, respectively. While both siRNAs efficiently knockdown αTAT1, siRNA#1 was used to perform all the knockdown experiments throughout this study. Cells were transfected with 5 nmol siRNAs using RNAiMAX (Invitrogen, Carlsbad, CA), according to the manufacturer's instructions. Other siRNA target sequences used in this study are listed in Table S1. Negative control siRNA is designed by Bioneer.

Cell fractionation

For whole-cell extracts, cell pellets were directly lysed with SDS sample buffer. For biochemical cell fractionation (Kim et al., 2008), cells were first lysed with 0.1% Triton X-100 CSK buffers (10 mM PIPES pH 6.8, 100 mM NaCl, 300 mM sucrose, MgCl2, 1 mM EGTA, 1 mM EDTA, 1 mM PMSF, 1 μg/ml leupeptin, 1 μg/ml aprotinin, 50 mM NaF, 0.1 mM NaVO4, and 0.1% Triton X-100) for 5 min on ice. After centrifugation (1000 g for 5 min), the supernatant (cytoplasmic and nucleoplasmic proteins) was isolated. The final pellet (chromatin and nuclear matrix proteins) was resuspended with the same buffer.

Immunofluorescence

Cells grown on the coverslips were fixed with 3.7% paraformaldehyde (PFA; Sigma-Aldrich) or ice-cold methanol, followed by permeabilization with 0.5% Triton X-100. Cells were blocked with 5% BSA in PBS and were incubated with indicated primary antibodies [anti-acetyl α-tubulin (1:400), anti-α-tubulin (1:3000), anti-γH2AX (1:1000), anti-RPA1 (1:500), anti-RPA2 (1:1000) and anti-BRCA1 (1:200) antibodies]. Alexa Fluor 488 and Alexa Fluor 568-conjugated secondary antibodies (Invitrogen) were used at a dilution of 1:500. After counterstaining with 4′,6-diamidino-2-phenylindole (DAPI; Sigma-Aldrich), coverslips were mounted with ProLong Gold antifade (Thermo Fisher Scientific), and analyzed by fluorescence microscopy.

Cell cycle analysis

Samples were collected over the indicated time points and fixed in 70% ethanol overnight. For propidium iodide (PI) staining, fixed cells were treated with RNase for 20 min before addition of 5 µg/ml PI (Sigma-Aldrich). For BrdU staining, cells were pulsed with BrdU (10uM for 1 h), fixed, and stained according to the manufacturer's instructions (BD Pharmingen, Heidelberg, Germany). Flow cytometric analysis was then performed using a FACSCalibur flow cytometer (Becton Dickinson, Oxford, UK).

Constructs and retroviral infection

Wild-type αTAT1 cDNA and D157N αTAT1 cDNA were derived from pEF5B-FRT-GFP-wt-αTAT1 (Addgene #27099) and pEF5B-FRT-GFP-D157N-αTAT1 (Addgene #27100), respectively. The αTAT1 cDNAs were sub-cloned into pMSCV-FLAG retroviral vector. To generate a siRNA-resistant construct, silent point mutations in the αTAT1 siRNA #1-targeting sequence were generated by using the QuikChange site-directed mutagenesis kit (Stratagene, La Jolla, CA) and verified by sequencing. Retroviral vectors were co-transfected with pVSV-G and pGag-Pol into 293T-17 cells for virus production. Virus was collected 48 h after transfection and subsequently used to infect HeLa cells. At 2 days after viral infection, cells were selected in 1 μg/ml puromycin (Invitrogen, Carlsbad, CA, USA) for establishing stable clones.

Homologous recombination assays

HR assays were performed as described previously (Kim et al., 2009). DR-GFP U2OS cells were seeded onto 12-well plates in triplicate and then transfected with the indicated siRNAs. At 24 h after transfection, the cells were then transfected with either I-SceI plasmid or GFP plasmid. After another 48 h, the cells were harvested and analyzed for GFP by fluorescence-activated cell sorting (FACS) analysis. In each experiment, the percentage of green (GFP+) cells was measured in triplicate samples. Values were normalized for the transfection efficiency and were displayed as mean±s.e.m. GFP+ frequencies relative to that of control siRNA-treated cells.

Statistical analysis

All data are presented as mean±s.e.m. Statistical significance between two data sets were compared by use of a Student's two-tailed t-test.

We are grateful to Seok-Won Jang for his involvement in the initial stages of the project.

Author contributions

Conceptualization: J.M.K.; Methodology: J.M.K.; Validation: J.M.K.; Formal analysis: N.M.R., J.M.K.; Investigation: N.M.R., J.M.K.; Data curation: J.M.K.; Writing - original draft: N.M.R., J.M.K.; Writing - review & editing: J.M.K.; Supervision: J.M.K.; Funding acquisition: J.M.K.

Funding

This work was supported by Basic science Research Program through the National Research Foundation of Korea funded by the Ministry of Education (2017R1A1A2054983).

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Competing interests

The authors declare no competing or financial interests.

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