Phosphoproteomics Reveals the AMPK Substrate Network in Response to DNA Damage and Histone Acetylation

Abstract

AMP-activated protein kinase (AMPK) is a conserved energy sensor that plays roles in diverse biological processes via phosphorylating various substrates. Emerging studies have demonstrated the regulatory roles of AMPK in DNA repair, but the underlying mechanisms remain to be fully understood. Herein, using mass spectrometry-based proteomic technologies, we systematically investigate the regulatory network of AMPK in DNA damage response (DDR). Our system-wide phosphoproteome study uncovers a variety of newly-identified potential substrates involved in diverse biological processes, whereas our system-wide histone modification analysis reveals a link between AMPK and histone acetylation. Together with these findings, we discover that AMPK promotes apoptosis by phosphorylating apoptosis-stimulating of p53 protein 2 (ASPP2) in an irradiation (IR)-dependent manner and regulates histone acetylation by phosphorylating histone deacetylase 9 (HDAC9) in an IR-independent manner. Besides, we reveal that disrupting the histone acetylation by the bromodomain BRD4 inhibitor JQ-1 enhances the sensitivity of AMPK-deficient cells to IR. Therefore, our study has provided a resource to investigate the interplay between phosphorylation and histone acetylation underlying the regulatory network of AMPK, which could be beneficial to understand the exact role of AMPK in DDR.

Introduction

AMP-activated protein kinase (AMPK) is a heterotrimeric serine/threonine kinase complex with one catalytic subunit α (including α1 and α2) and two regulatory subunits β (including β1 and β2) and γ (including γ1, γ2, and γ3), which contains theoretically 12 different types of heterotrimers. As a central metabolic sensor to restore intercellular energy homeostasis, activation of AMPK switches off anabolic pathways that consume ATP and switches on catabolic pathways that generate ATP. There are two coordinated mechanisms that modulate the activity of AMPK. One is phosphorylation at its conserved Thr172 in the activation loop of AMPKα1 subunit, which is regulated by upstream kinases liver kinase B1 (LKB1) and calmodulin-dependent protein kinase kinase β (CaMKKβ). The other is the binding of ADP/AMP to γ subunit to allosterically activate AMPK by stabilizing it in an active conformation [1]. Once activated, AMPK, in turn, further phosphorylates diverse downstream substrates, such as metabolic enzymes, transcription factors, and co-activators, to balance energy homeostasis either by short-term provocations of metabolic signaling cascades or by long-term regulations of transcription and posttranslational modification [2].

Emerging studies have uncovered the contradictory roles of AMPK in tumor development and cancer therapy. AMPK was reported to be a key mediator contributing to the suppression effect of LKB1 signaling cascades [3,4]. Such a hypothesis is supported by a series of studies from metformin, whose mechanism of action is widely recognized to be associated with AMPK activation [5,6]. Metformin could significantly inhibit tumor growth and improve chemo-sensitivity and radio-sensitivity via AMPK activation in various types of tumors [7–10]. Remarkably, several studies showed the effect of metformin to kill the cancer stem-like cells [11–13], which are considered a major barrier in cancer therapy. Studies from another AMPK activator 5-aminoimidazole-4-carboxamide-1-β-D-ribofuranoside (AICAR) [14] also supported such a hypothesis. One the other hand, other studies suggest that AMPK could be a context-dependent tumor promoter. The biological consequences in which AMPK is supra-physiologically activated by compounds are different from those in which AMPK is physiologically activated by cellular stresses [15]. Indeed, the AMPK energy-sensing pathway supports cells to survive in hypoxic and nutrient-deficient conditions, which is regarded as a typical tumor microenvironment. Supporting such an assumption is that autophagy, which is partially considered to be regulated by AMPK, might provide enough nutrients to support cancer survival by degrading cellular organelles or proteins [16]. In addition, glyceraldehyde-3-phosphate dehydrogenase (GAPDH) generated by fatty acid oxidation, which is up-regulated by AMPK activation, is beneficial to protect cancer cells from oxidative stresses by neutralizing cytotoxic reactive oxygen species (ROS) [17]. Furthermore, increasing phospho-AMPK levels is associated with higher tumor grade in prostate cancer [18]. Therefore, these contradictory findings suggest an urgent need to understand the exact role of AMPK in tumor biology.

The aforementioned puzzling phenomenon could be partially explained by the complicated functional consequences caused by the diverse substrates of AMPK. For example, AMPK-mediated phosphorylation on p53 [19–21] and the mechanistic target of rapamycin complex 1 (mTORC1) [22–24] signaling pathway are essential for activator-induced tumor suppression. In contrast, AMPK could promote tumor growth via other mechanisms, including the induction of mitophagy by phosphorylating ULK1 [16], the up-regulation of stress-induced gene transcription by phosphorylating histone H2B [25], and the promotion of FA oxidation to neutralize oxidative stress [17]. Moreover, new substrates identified in mitochondrial fission [26,27], hippo-YAP signaling pathway [28], and mitosis [22] implied a subtle regulatory role of AMPK in cancer. In addition, the pharmacological effects of two anti-cancer drugs, etoposide and cisplatin, were reported to be partially dependent on AMPK activation [29–31]. Despite these understanding, the detailed mechanism is not yet fully understood how AMPK coordinately regulates these cascades and whether some unknown partners are involved in this process. Indeed, our knowledge of the cancer-associated AMPK substrates is still limited. Therefore, a comprehensive understanding of the substrates of AMPK is a critical step toward understanding the role of AMPK in cancer.

Human tumors share various biological hallmarks acquired during multistep development. One of these hallmarks is genome instability, which is acquired during cancer development and drug resistance [32]. Abnormal or deficient DNA damage response (DDR) results in genomic instability and neoplastic transformation. In recent years, emerging reports suggested that AMPK was involved in DDR, but the regulatory role remained to be fully understood [33–35].

To address this question, we carried out a system-wide phosphoproteome study by mass spectrometry (MS)-based proteomic technologies. Our results showed that a variety of newly-identified substrates were involved in diverse biological activities, which shed light on a broad and complex regulatory network of AMPK in DDR. In addition, our system-wide histone modification analysis showed that AMPK played a role in modulating global histone acetylation levels. Disrupting the histone acetylation by the bromodomain BRD4 inhibitor JQ-1 enhanced the sensitivity of AMPK-deficient cells to irradiation (IR) via induction of apoptosis. Thus, our study provided an abundant resource to investigate the interplay between phosphorylation and histone acetylation underlying the regulatory network of AMPK, which might be beneficial to understand the exact role of AMPK in tumor biology.

Results

Establishment of AMPKα1/α2-double knockout cell lines by TALENs

We first investigated whether AMPK was involved in DDR. X-ray IR induced cellular DNA damages and activated DDR signaling to repair DNA double strand breaks (DSBs). After a single dose of X-ray IR exposure, both AMPK activation signal phospho-AMPK (Thr172) and its substrate signal phospho-ACC (Thr79) increased as IR dose elevated, suggesting that AMPK was activated in a dose-dependent way in DDR (Figure 1A and B). To investigate the underlying mechanisms of AMPK activation during DDR, we established stable AMPKα1/α2 (two AMPK catalytic isoforms)-double knockout (KO) mouse embryonic fibroblast (MEF) cell lines using Transcription Activator-Like Effector Nuclease (TALEN) technology. TALEN technology is a genome-editing method widely used to generate KO Caenorhabditis elegans, rats, mice, and zebrafishes [36–42]. It is also used in genomic modification of human embryonic stem cells and induced pluripotent stem cells (iPSCs). The establishment method was according to previous reports [39,41,43]. We obtained three stable AMPKα1/α2-KO cell lines (named as 1#, 2#, and 3#). The AMPKα1 and α2 protein expression levels could not be detected by immunoblot, suggesting successful deletion of total AMPKα subunits (Figure 1C). To evaluate whether downstream signaling was also impaired, we treated cells with AMPK activator AICAR. In contrast to wild-type (WT) MEF cells, AICAR treatment did not induce AMPK activation signal phospho-AMPKα (Thr172) and substrate signal phospho-ACC (Thr79) in the three KO cell lines, further suggesting the loss of AMPKα1 and α2 kinase activity (Figure 1C). Besides, we extracted genomic DNA from individual cell lines for sequencing and found nucleotide deletion in leading chain and lagging chain. In addition, our genomic sequencing of these cell lines also showed that nucleotide deletion happened in both leading chain and lagging chain in all three cell lines (Figure S1). In contrast to 3# cell line, TALENs induced non-3 nt-deletion in 1# and 2# cell lines that resulted in absolute target KO in genomic level because of coding frame shift. We next chose 1# and 2# cell lines for further functional assay. To address the issue whether AMPK is involved in DDR, we exposed WT MEF, 1#, and 2# cells to a single dose of X-ray IR. We observed an increasing γH2A.X (phospho-H2A.X Ser139) foci (a marker of DSB measured by immunofluorescence) after IR exposure (Figure 1D and E). Comparable percentage of γH2A.X foci-positive cells at 48 h post-IR in 1# and 2# cell lines suggested that these two cell lines could be functionally equivalent in DDR. Therefore, 1# cell line (defined as AMPKα-KO cells in this study) was selected for further proteome analysis. Meanwhile, compared with WT MEFs, more γH2A.X foci remained at 48 h in 1# and 2# cell lines, suggesting that AMPKα was required to promote DSB repair efficiency. Besides, a prolonged G2 phase arrest was observed in AMPKα-KO cells after IR (WT, 0 h 27.30% to 24 h 25.72%, P = 0.690; KO, 0 h 30.55% to 24 h 40.65%, P = 0.011) (Figure 1F and G). Taken together, these results suggested that the kinase activity of AMPKα is required in DDR, but the underlying regulatory network remains to be understood.

Quantitative phosphoproteomics uncovered broad signaling pathways regulated by AMPK in DDR

To further profile the regulatory networks of AMPK in DDR, we carried out a system-wide phosphoproteome analysis (Figure 2A). Previous studies suggested that more abundant signaling pathways would be activated in response to IR at relatively high doses than at low doses [44,45]. According to the result that AMPK activated and phosphorylated substrates in a dose-dependent manner (Figure 1A and B), we hypothesized that AMPK likely activated more signaling pathways at higher IR doses. Therefore, a relatively high dose (10 Gy) that mimicked the stressed cellular microenvironment in radio-combined cancer therapy was selected in further experiment. The proteome of AMPKα-KO cells was labeled with “heavy” (¹³C₆-Lys and ¹³C₆¹⁵N₄-Arg) amino acids, whereas the proteome of the WT cells was labeled with “light” (¹²C₆-Lys and ¹²C₆¹⁴N₄-Arg) amino acids in cell culture. WT and AMPKα-KO cells were prepared in both basal group and IR group at the same time. After IR, cells from IR group were released for 90 min before harvest to fully activate diverse signaling pathways involved in DDR. Proteins were extracted from the two cell populations and mixed equally for further analysis. Two types of proteases, trypsin and chymotrypsin, were used to improve the sequence coverage of the phosphoproteome as previously reported [46]. Collectively, we identified 11,065 phosphosites with a localization probability higher than 0.75 in the basal and IR groups. First, we verified the reproducibility between replicate experiments in basal (trypsin and chymotrypsin digested) and IR (trypsin and chymotrypsin digested) groups (Figure S2). Scatter plot depicting the correlation between two replicates in each group showed high correlation coefficient values (r = 0.78–0.84), suggesting reliable reproducibility of our phosphoproteomic data. We used a criterion of more than 1.5-fold between basal and IR groups to define those significantly different phosphosites (P < 0.05 by Student’s t-test). A total of 539 down-regulated phosphosites and 389 up-regulated phosphosites were identified in the basal group (Figure 2B; Table S1). Phosphosites of well-known AMPK substrates were found to be significantly down-regulated in AMPKα-KO cells (Figure 2B). Motif analysis showed that the consensus AMPK substrate motif could be enriched in the down-regulated phosphosites in AMPKα-KO cells (Figure S3). Together, these results suggested the high quality of our proteomic data.

Meanwhile, 1216 down-regulated phosphosites and 169 up-regulated phosphosites were identified in the IR group (Figure 2C and D; Table S1). This result indicated that various protein phosphorylation events could be regulated by AMPK during DDR. Of these significantly down-regulated phosphosites, 471 phosphosites (38%) were also down-regulated (FC < 0.667) in the basal group (Figure 2B). In contrast, 257 phosphosites (21%) remained unchanged (0.8 < FC < 1.2) in the basal group (Figure 2B and D). For example, phosphorylation on Xrcc1 Thr452 was slightly down-regulated in the basal group (log₂ FC = –0.28, P = 0.01), while it was significantly altered after IR (log₂ FC = –0.78, P = 0.005). Therefore, these results suggested that AMPK played additional regulatory roles when cells suffered severe genomic stress, as compared to its routine regulatory roles in basal status.

To gain insight into the possible biological functions of AMPK during DDR, we subjected all significantly down-regulated and up-regulated phosphoproteins identified in IR group to bioinformatic enrichment analysis using GO and KEGG databases by the DAVID bioinformatic tool. In consistent with a prolonged G2 phase arrest observed in AMPKα-KO cells after IR (Figure 1F and G), we discovered that the changed phosphoproteins were significantly enriched in DNA damage repair-associated events, such as cell cycle, DNA repair, and transcription (Figure 2E). Surprisingly, chromatin-associated functions were also highly enriched, such as chromatin regulation and histone modification (including deacetylation, ubiquitination, and methylation), suggesting a potential regulatory role of AMPK in epigenetic modification. Meanwhile, we also discovered a potential crosstalk between DNA damage and RNA processing (both mRNA processing and rRNA processing), in consistent with emerging evidence that mRNA processing factors are involved in DNA damage signal transduction. Theoretically, BCLAF1 was assembled at damage sites via interacting with core spliceosome components while THRAP3 was excluded from damage sites as a consequence of transcription repression [47]. Phosphorylation event was one of factors to regulate the assemble and dissemble processes. The significantly up-regulated phosphosites Ser248 and Ser572 on THRAP3, as well as the significantly down-regulated phosphosite Ser284 on BCLAF1, were observed. Besides, mRNA splicing factor hnRNPUL1, a DNA-end resection regulator, was found to be significantly down-regulated at Ser4 during DDR (Figure 2F). These findings implied that the phosphorylation events on these mRNA splicing factors might be involved in DSB repair. Taken together, bioinformatic enrichment analysis revealed a broad and complicated regulatory role of AMPK involved in diverse signaling pathways.

Bioinformatic analysis identifies ASPP2 as a new substrate of AMPK involved in apoptosis

We next analyzed the significantly changed phosphoproteins in IR group by Motif-X algorithm to predict the preferred motif sequences. It was well established that AMPK phosphorylated diverse substrates at a consensus amino acid sequence motif. Briefly, –5 site and +4 site contained a hydrophobic amino acid like L/M/V/I/F (Φ), while –3 or –4 site contained at least one basic amino acid R/K (β) [48]. Motif enrichment analysis of down-regulated phosphosites in IR group was carried out. Among several enriched motifs proposed by Motif-X algorithm (Figure 3A), the sequences ranked 2nd and 3rd, xxxxRRxSxxxxxxx and xxxxRxxSxxxxxxx, partially matched to the classical AMPK substrate motif ΦxβxxSxxxΦ. This possibly indicated that there existed some potential AMPK substrates not strictly match the classical motif very well.

Among the aforementioned significantly down-regulated phosphosites, we further divided them into two groups, IR-independent substrates and IR-dependent substrates. The IR-dependent substrates (DNA damage-associated substrates) were phosphorylated by AMPK under DDR. These phosphosites were down-regulated in IR group but remained nearly unchanged in basal group (normalized heavy/light ratio, 0 < IR/Basal < 0.8 and 0.8 < Basal < 1.2). In contrast, the IR-independent substrates (general substrate candidates) were phosphorylated by AMPK independent of DDR. These phosphosites were significantly down-regulated in either basal group or IR group (heavy/light ratio < 0.67). To identify direct potential substrates of AMPK, we aligned these phosphosites in both IR-dependent and IR-independent substrates to the AMPK consensus motif. Based upon the criteria, we totally obtained 42 potential substrates (Figure 3B; Table 1).

To further confirm the reliability of our results, we randomly selected one of the substrates, apoptosis-stimulating of p53 protein 2 (ASPP2) for further validation. Ser479 on ASPP2 was considered as a DNA damage-associated substrate because the normalized IR/basal ratio of Ser479 on ASPP2 was 0.78 (normalized heavy/light ratio, Basal = 0.95, IR = 0.74). p53 was a well-established tumor suppressor involved in both cell cycle arrest and apoptosis. ASPP2 was initially identified as a binding partner of p53 and required in p53-mediated apoptosis [49,50]. We first investigated whether protein interaction existed between AMPK and ASPP2. Plasmids were synchronically introduced into 293T cells to overexpress Flag-ASPP2 and Myc-AMPKβ1 for 48 h, and the whole cell lysates were harvested for exogenous immunoprecipitation (IP) with the anti-Flag antibody. Immunoblot showed interaction between AMPKβ1 and ASPP2 (Figure S4). The endogenous protein–protein interaction was further confirmed in 293T cells in IP assay using AMPKβ1 antibody (Figure 3C). Bioinformatic analysis revealed that the amino acid sequence surrounding Ser479 on ASPP2 matched the AMPK substrate motif and kept conserved during species evolution (Figure 3D). Therefore, we investigated whether AMPK could phosphorylate ASPP2 at Ser479 by in vitro kinase assay using [³²P]-labeled ATP. Truncated ASPP2-(1-765 aa)-WT and mutant ASPP2-(1-765 aa)-S479A (serine converted to non-phosphorylatable alanine) that mimicked phosphonull status were purified from Escherichia coli and incubated with AMPKα2β2γ2 complex in kinase reaction buffer for 1 h. AMPKα2β2γ2 complex with no kinase activity was purified from E. coli. but retained catalytic activity after incubation with upstream kinase CAMKKβ. Robust [³²P] signal was induced by activated-AMPKα2β2γ2 complex on truncated ASPP2-(1-765 aa)-WT but strongly attenuated on mutant ASPP2-(1-765 aa)-S479A, suggesting that AMPK phosphorylated ASPP2 dominantly at Ser479 (Figure 3E). However, weak [³²P] signal detected on mutant ASPP2-(1-765 aa)-S479A also indicated that other unknown AMPK phosphosites on ASPP2 remained to be further investigated. We then investigated whether Ser479 phosphorylation was involved in apoptosis. HeLa cells were transfected with full-length ASPP2-WT or mutant ASPP2-S479A plasmids for 24 h. Apoptotic cells were labeled with Annexin V and PI dyes, and their fluorescence intensity was measured by flow cytometry. Compared to the control cells transfected with the vector only, overexpression of full-length ASPP2-WT significantly induced an increase in apoptosis (1.67 folds; P < 0.05, ANOVA); overexpression of mutant ASPP2-S479A appeared to increase the percentage of apoptotic cells, while the increase was not significant. Although there seemed a difference between the cells overexpressing full-length ASPP2-WT and those overexpressing mutant ASPP2-S479A, while the difference was not significant (Figure 3F). These results suggest that AMPK-mediated Ser479 phosphorylation partially contributed to ASPP2-induced apoptosis, but other AMPK phosphorylation sites on ASPP2 may also play roles in this process or there exist other unrecognized regulatory mechanisms.

Histone modification analysis characterized AMPK as a regulator of histone acetylation

Since chromatin-associated functions were highly enriched in bioinformatic analysis, we hypothesized that AMPK was involved in histone posttranslational modifications and epigenetic regulation. To address this question, we carried out a system-wide analysis to investigate the histone modification by MS in both non-IR basal status and IR stress status (Figure 4A). The heavy-labeled AMPKα-KO cells and light-labeled WT cells were mixed at a 1:1 ratio, and core histones (H2A, H2B, H3, and H4) were extracted and separated with SDS-PAGE. Histones were in-gel digested with trypsin into peptides and then subjected to MS analysis. The modified peptides were checked manually and quantified based on peak area integral according to the previous report [51]. Interestingly, AMPKα1/α2 KO resulted in significant up-regulation of global histone acetylation levels (Figure 4B; Table S2). Immunoblot assay also suggested increased acetylation levels at H2BK12, H3K18, and H4K16 in AMPKα1/α2-KO cell lines 1# and 2# (Figure 4C). These data suggest that AMPK plays a negative regulatory role in global histone acetylation.

To find out underlying regulation mechanism, we chose another potential substrate histone deacetylase 9 (HDAC9) from our phosphoproteome data for investigation. The amino acid sequence surrounding Ser239 of HDAC9 matched the AMPK substrate motif and was evolutionarily conserved (Figure 4D). Plasmids were synchronically introduced into 293T cells to overexpress Flag-HDAC9 and Myc-AMPKα2 for 48 h, and the whole cell lysates were subjected to exogenous IP assay using the anti-Flag antibody. AMPKα2 was detected using anti-Myc antibody in Flag-HDAC9 immunoprecipitated complex, suggested that AMPKα2 interacted with HDAC9 (Figure S5). Endogenous IP assay conducted in 293T cells further confirmed a reciprocal interaction between AMPKα2 and HDAC9 using anti-AMPKα2 and anti-HDAC9 antibodies (Figure 4E). Additionally, the exogenous fusion protein Flag-HDAC9 was enriched and extracted by anti-Flag antibody from 293T cells and incubated with activated or non-activated AMPKα2β2γ2 complex in kinase reaction buffer for 1 h. Phosphorylation signal (as measured by pan phospho-AMPK-substrate antibody) was detected on HDAC9 when incubated with activated-AMPKα2β2γ2 complex in the presence of CAMKKβ (Figure 4F). Taken together, HDAC9 was newly identified as a substrate of AMPK. Afterward, two plasmids expressing HDAC9-WT and phosphonull mutant HDAC9-S239A were constructed and introduced into 293T cells. Compared with negative control, overexpression of HDAC9-WT enhanced H2BK12 acetylation and H3K18 acetylation. In contrast, overexpression of phosphonull mutant HDAC9-S239A enhanced higher H2BK12 acetylation level but had no impact on H3K18 acetylation level (Figure 4G). Therefore, these results suggested that HDAC9 was involved in balance of histone acetylation while Ser239 phosphorylation on HDAC9 played a negative role in regulation of H2BK12 acetylation. Since there was no significant difference in HDAC9-Ser239 phosphorylation between basal and IR groups, AMPK likely phosphorylated HDAC9 at Ser239 in an IR-independent manner.

Taken together, quantitative phosphoproteome and histone modification analyses uncovered subtle crosstalk between AMPK-mediated phospho-regulation and histone acetylation.

JQ-1 synergizing with an AMPK inhibitor sensitizes cell apoptosis to IR

Histone acetylation had an impact on localized chromatin structures and played important roles in initiation and transduction of DNA damage signaling. BRD4, a member of the bromodomain and extraterminal (BET) family that binds acetylated histones H3 and H4, and regulates gene expression [52–54]. In addition to transcriptional regulation, BRD4 is essential in DDR and mediates the recruitment of chromatin-based repair proteins. Once sensing genome-wide DNA breaks, enhanced H4 acetylation led to BRD4 accumulation at breaks [51,55]. Since increased acetylation level at H3K14ac and global H4 acetylation level was observed in AMPKα-KO cells in basal group, but the tendency slightly attenuated after IR (Figure 4B), we hypothesized a potential linkage between AMPK and BRD4 during DDR. JQ-1, a BET inhibitor that impeded BRD4-acetylated lysine interaction, displayed anti-tumor activity in various types of cancer [56,57]. Interestingly, JQ-1 treatment made AMPKα-KO cells more sensitive to IR than WT cells (Figure 5A). Similarly, we found JQ-1 treatment together with Compound C (an AMPK inhibitor) improved the sensitivity of M059J glioma cells to IR, especially in a relatively high dose (Figure 5B). It was suggested that acetylation of histones H3K14, H4K12, and H4K16 were important for BRD4 binding [58]. In consistency, JQ-1 reduced acetylation of H3K14 and H3K18 in both WT and AMPKα-KO cells. However, quite different effects of JQ-1 on the acetylation of H3K23 and H2BK12 were observed in WT and AMPKα-KO cells. JQ-1 treatment had no effect on H3K23 acetylation in WT cells but reduced H3K23 acetylation in AMPKα-KO cells. In contrast, JQ-1 treatment did not change H2BK12 acetylation in AMPKα-KO cells but reduced H2BK12 acetylation in WT cells (Figure 5C and D). Notably, JQ-1 reduced histone acetylation before IR, with no obvious changes observed after IR, suggesting that JQ-1 regulated histone acetylation in an IR-independent manner. Since WT and AMPKα-KO cells showed an opposite trend on H2BK12 and H3K23 acetylation after JQ-1 treatment, we hypothesized that AMPK signaling and BRD4 signaling coordinately regulated H3K23 and H2BK12 acetylation. Next, we investigated how the JQ-1-induced basal changes in histone acetylation finally resulted in lower cell viability under IR stress. Herein, we investigated whether repair signaling activated normally by JQ-1 treatment resulted in enhanced phospho-KAP-1(Ser824) signals and prolonged γH2A.X signals sustained in AMPKα-KO cells, but without changes in phospho-p53(Ser15) (Figure 5E). Therefore, we hypothesized that JQ-1 treatment affected the chromatin relaxation by disrupting the orchestrated regulation of histone acetylation during DDR. Accumulated unrepaired DSB ends, as indicated by enhanced γH2A.X signals, would induce apoptosis. Consistently, we observed that JQ-1 treatment induced a higher proportion of apoptotic cells in the AMPKα-KO cells than in WT cells (Figure 5F). Thus, we concluded that AMPKα1/α2 KO disrupted the orchestrated histone acetylation, which promoted the pro-apoptotic effect induced by JQ-1 treatment during DDR. Taken together, inhibition of AMPK activity improved the anti-tumor efficacy of JQ-1 via disrupting the balance in histone acetylation.

Discussion

AMPK is an essential eukaryotic energy sensor to coordinate metabolism and other biological activities. LKB1, one of the upstream kinases of AMPK, was a well-defined tumor suppressor [59]. However, fully understanding the exact role of AMPK seemed challenging. Previous studies suggested that AMPK, unlike its upstream kinase, played contradictory roles in tumor progression and therapy. AMPK activation regulated diverse downstream signaling pathways, together with complicated tumor microenvironments, which led to context-dependent cell fates. In one hand, AMPK phosphorylated ULK1 and promoted cell autophagy, to maintain cell survival in conditions of hypoxia and low nutrition [16,60]. On the other hand, AMPK negatively regulated TSC-mTOR axis and inhibited cell outgrowth under the circumstance of energy deprivation [61–63]. It seemed that cell fate resulted from AMPK activation mainly depended on comprehensive signaling network within heterogenic tumor cells. The intercellular signaling network in types of tumors was balanced and maintained by both extrinsic stimulators like hypoxia and growth factors, as well as intrinsic features like gene signatures and transcriptional regulation [64,65]. Genome instability and mutation was defined as one of the hallmarks in various types of cancer [66]. Abnormal DNA repair, especially dysfunctional DSB repair, has an impact on the genetic fidelity and chromatin structure and consequently leads to genome instability and mutation [32,67]. Previous studies suggested that AMPK was involved in DNA-damaging agent induced cell apoptosis, but the underlying mechanism remained to be further understood [34,68,69]. Meanwhile, several studies shed light on the linkage between AMPK and DDR. DDR is a critical process to activate accurate repair pathways and other biological processes to coordinately repair damages. Therefore, a comprehensive understanding of the AMPK’s role in DDR is a critical step toward understanding the exact role of AMPK in tumor biology.

To address the question, we designed a study to investigate whether novel AMPK substrates were involved in DDR. We carried out a system-wide phosphoproteomic study and global histone posttranscriptional modification analysis. Bioinformatic analysis of significantly altered phosphoproteins enriched DNA damage repair-associated events and chromatin-associated functions, suggesting complicated regulatory network of AMPK in DDR. After comparing these altered phosphosites to classic AMPK substrates, we totally obtained 42 potential substrates of AMPK from basal group and IR group. This result suggested that although IR might dramatically induce phosphorylation changes, AMPK likely had limited direct impact on DNA-associated signaling pathways.

Corresponding to the fact that chromatin-associated functions were enriched in phosphoproteomic study, histone modification analysis also suggested that AMPK negatively regulated global histone acetylation. Therefore, an integrative analysis of two studies implicated a crosstalk between AMPK-mediated phospho-regulation and histone acetylation. Based on this result, we further identified HDAC9, a member of Class IIa histone deacetylase family, as a new substrate phosphorylated by AMPK in an IR-independent manner. Overexpression of WT HDAC9 in cells increased the acetylation level at H2BK12 and H3K18, and the non-phosphorylatable mutant HDAC9-S239A further promoted the increase of H2BK12 acetylation. Thus, we concluded that AMPK-mediated phosphorylation on HDAC9 might be required in balancing the histone acetylation at H2BK12, but the mechanism remained to be further investigated.

It was worth noting that AMPK-mediated phosphorylation at Ser479 of ASPP2 partially contributed to ASPP2-induced apoptosis in an IR-dependent manner. p53 plays distinct roles via different co-factors in DDR, including induction of cell cycle arrest or apoptosis. ASPP2 is the co-factor of p53 required in spontaneous induction of apoptosis and cooperates with p53 to suppress tumor growth [70,71]. Previous studies suggested that a high level of ASPP2 sensitized cells to IR and DNA-damaging agents [72,73]. Overexpression of non-phosphorylatable mutant ASPP2-S479A moderately impaired apoptosis, suggesting that AMPK promoted apoptosis during DDR by phosphorylating ASPP2 at Ser479. The accumulation of unrepaired DNA damages activated apoptosis signaling pathway, thus avoiding hereditary of abnormal genetic information in mitosis. Taken together, AMPK-ASPP2 axis promoted apoptosis to prevent cells from genomic instability caused by sustained damages during DDR. Further study might focus on whether the phosphorylation site influenced protein interaction of ASPP2 complex and its correlation with cancer incidence or drug resistance.

Finally, our findings might reveal a crosstalk between AMPK activity and histone acetylation in tumor biology. Indeed, disturbing the histone acetylation by BRD4 inhibitor JQ-1 enhanced the sensitivity of cells to IR via induction of apoptosis. It was worth noting that except for H3K14ac, JQ-1 also influenced acetylation at H2BK12, H3K23, and H3K18. Additionally, AMPKα1/α2-deficient cells were more sensitive than WT cells to JQ-1 treatment. These results suggested the potential role of AMPK in cancer therapy. First, AMPK activity might be used as a biomarker to predict the therapeutic response to acetyltransferase inhibitors or deacetylase inhibitors. Second, selective AMPK inhibitor might be useful to improve the sensitivity of tumor cells to radio-therapy or target-therapy.

Therefore, our study provided a source of AMPK-associated phosphorylation network and histone acetylation events, which might be helpful to understand the role of AMPK in DDR.

Materials and methods

Antibodies and compounds

The primary antibodies Phospho-Histone H2A.X (Ser139) (Catalog No. 2577), Phospho-p53 (Ser15) (Catalog No. 9284), Phospho-AMPKα (Thr172) (Catalog No. 2535), Phospho-Acetyl-CoA Carboxylase (Ser79) (Catalog No. 3661), Acetyl-CoA Carboxylase (Catalog No. 3662), AMPKα (Catalog No. 2532), AMPKα2 (Catalog No. 2757), AMPKβ1 (Catalog No. 12063), AMPKβ2 (Catalog No. 4148), GAPDH (Catalog No. 2118), and Myc-Tag (9B11) (Catalog No. 2276) were bought from Cell Signaling Technology (Danvers, MA). The primary antibodies Phospho-KAP1 (Ser824) (Catalog No. ab70369), H2B-Acetyl-K12 (Catalog No. ab61228), H4-Acetyl-K16 (Catalog No. ab109463), H3-Acetyl-K14 (Catalog No. ab82501), H3-Acetyl-K18 (Catalog No. ab1191), H3-Acetyl-K23 (Catalog No. ab61234), and 53BP2 (Catalog No. ab236448) were bought from Abcam (Cambridge, UK). The primary antibody HDAC9 (Catalog No. MA5-32820) was bought from ThermoFisher Scientific (Waltham, MA). The DYKDDDDK Tag (Catalog No. 018-22783) was bought from Wako (Osaka, Japan). The antibody β-actin (Catalog No. AM1021B) was bought from Abgent (San Diego, CA).

Secondary antibody Alexa Fluor 555 labeled donkey anti-rabbit IgG (Catalog No. A31572) and Hoechst 33342 Solution (Catalog No. 62249) were bought from ThermoFisher Scientific. The secondary peroxidase AffiniPure Goat Anti-Mouse IgG (H+L) (Catalog No. 115-035-003) and Peroxidase AffiniPure Rabbit Anti-Goat IgG (H+L) (Catalog No. 305-035-003) were bought from Jackson ImmunoResearch Laboratories (West Grove, PA). Chemiluminescent detection was completed with enhanced chemiluminescence (ECL) Western blotting reagents (Catalog No. RPN2236, GE Healthcare, Chicago, IL).

The compound JQ-1 (Catalog No. S7110) was bought from Selleck (Houston, TX). The AMPK activator AICAR (Catalog No. A9978) and AMPK inhibitor (Catalog No. 171260) were bought from Sigma (St. Louis, MO) and Millipore (Burlington, MA), respectively.

Cell culture and stable isotope labeling by amino acids in cell culture labeling

WT MEFs and AMPKα1/α2-KO MEF cell lines were cultured in DMEM with light lysine (¹²C₆¹⁴N₂-Lys) and arginine (¹²C₆¹⁴N₄-Arg), or heavy lysine (¹³C₆¹⁴N₂-Lys) and arginine (¹³C₆¹⁵N₄-Arg), respectively. The proteome labeling efficiency of heavy isotopic amino acids was >98%, as determined by MS analysis. The HeLa and 293T cells were cultured in DMEM growth medium (Catalog No. 12100061, Gibco, Carlsbad, CA) supplemented with 10% FBS (Catalog No. 10099, Gibco). The M059J cells were grown in a medium containing a 1:1 mixture of DMEM and F12 medium (Catalog No. 11330057, Gibco) supplemented with 10% FBS. All cells were cultured at 37 °C with 5% CO₂.

Protein lysate preparation and in-solution digestion

Cells were harvested and washed with pre-cold phosphate-buffered saline (PBS). Then, cells were lysed with lysis buffer (8 M Urea in 100 mM NH₄HCO₃) and subjected to sonication. After sonication, the lysates were clarified by centrifugation at 21,130 g for 10 min. Equal amounts of WT and AMPKα1/α2-KO MEF cell lysate were mixed. The cell lysate mixture was reduced by 5 mM dithiothreitol (Catalog No. D0632, Sigma) at 56 °C for 30 min. Then 15 mM iodoacetamide (Catalog No. I6125, Sigma) was added to alkylate the sulfhydryl groups. The extra iodoacetamide was eliminated using 30 mM cysteine. The protein extract was digested with trypsin (Catalog No. V5280, Promega, Madison, WI) (trypsin:protein = 1:50) at 37 °C for 12 h. For complete digesting, additional trypsin (trypsin:protein = 1:100) was added for another 4 h. Same amount of protein extract was digested with chymotrypsin (Catalog No. 11418467001, Roche, Basel, CHE) (chymotrypsin:protein = 1:50) at 25 °C for 18 h. Both tryptic and chymotryptic peptides were desalted through Waters SepPak C18 cartridges (Catalog No. WAT054960, Waters, Milford, MA), vacuum-dried, and stored at −80 °C for further analysis.

Histone extraction and in-gel digestion

Histone extraction was carried out as previously published [74,75]. The isolated cells were lysed with extraction buffer (10 mM HEPES, 10 mM KCl, 1.5 mM MgCl₂, 0.5% NP-40, 1× protease inhibitor mixture). Lysate was centrifuged at 1000 g at 4 °C. Then, the pellets were washed and resuspended in 0.2 M H₂SO₄ overnight at 4 °C. The mixture was clarified by centrifugation, and the supernatant was collected for trichloroacetic acid precipitation. The precipitate was washed with pre-cold acetone several times. The precipitate was dried completely at room temperature and then dissolved in water for SDS-PAGE separation. Bands of histones (H1, H2A, H2B, H3, and H4) were excised and subjected to in-gel digestion. The tryptic peptides were analyzed by LC-MS/MS. After MS analysis, histone modifications of H2A, H2B, H3, and H4 were quantified by examined the peak area of modified peptide ions.

Phosphopeptide enrichment

Phosphopeptide enrichment was carried out as previously published [51]. In brief, tryptic peptides were dissolved in loading buffer (6% TFA, 80% ACN, 1 M lactic acid), and then incubated with titanium dioxide beads (GL Sciences, Japan) at room temperature at a proportion of peptide: TiO₂ = 4:1. The titanium dioxide beads were then washed with loading buffer for three times, wash buffer A (0.5% TFA, 30% ACN) for one time and wash buffer B (0.5% TFA, 80% ACN) for two times. The phosphopeptides were eluted from the beads with 15% NH₃H₂O and separated into six fractions.

Nano-HPLC-MS/MS analysis

The peptides were analyzed by an EASY-nLC 1000 system (ThermoFisher Scientific) connected to an Orbitrap Q-Exactive mass spectrometer (ThermoFisher Scientific). Peptides were eluted from a reverse-phase C18 column (75 μm ID, 3 μm particle size, Dikma Technologies Inc., CA) with a 70 min gradient of 7%-to-80% buffer B (90% acetonitrile, 10% H₂O, 0.1% formic acid) at a flow rate of 300 nl/min. Full MS spectra with an m/z range of 350–1300 were acquired with a resolution of 70,000 at m/z = 200 in profile mode. The AGC targets were 1.0e6 for full scan and 1.0e6 for MS/MS scan, respectively. Fragmentation of the 16 most intense precursor ions occurred at the HCD collision cell with a normalized collision energy of 28%, and tandem MS were obtained with a resolution of 17,500 at m/z = 200. Dynamic exclusion duration was set as 60 s.

MS data analysis

MS/MS data were processed using MaxQuant software (version 1.5.3.2). Mus musculus database from UniProt (release 2018_10_13, 53,781 entries) with a reversed decoy database was used for data processing. For database searching, trypsin was set as the specific enzyme and the maximum number of missed cleavages was fixed at 2. Carbamidomethylation of cysteine residues was set as a fixed modification; oxidation of methionine and protein N-terminal acetylation were set as variable modifications. Phosphorylation of serine, threonine, and tyrosine was set as variable modification for phosphosite analysis. Precursor mass tolerance for MaxQuant analysis was set to 4.5 ppm and MS/MS tolerance was set to 20 ppm. FDR thresholds for protein, peptide, and modification sites were all set as 1%.

Bioinformatic analysis

DAVID bioinformatics functional annotation tool was used to identify enriched GO and KEGG pathway terms. Mus musculus genome was used as background in DAVID functional annotation analysis. The significance of fold enrichment was calculated using P < 0.05. Gene sets with P < 0.05 were visualized in enrichment maps using EnrichmentMapApp and Cytoscape. Motif-X was used to identify phosphorylation motifs present in significantly changed phosphoproteins.

In vitro kinase assay and autoradiography

Briefly, the recombinant AMPKα2β2γ2 complex (200 nM) was pre-incubated with 20 nM CAMKKβ to be fully activated in reaction buffer (5 mM MgCl₂, 20 mM Tris-HCl, 8 nM ATP, 1 mM DTT) at 37 °C for 1 h. Then 200 nM substrate was incubated with 20 nM activated AMPK in reaction buffer at 37 °C for 1 h, in which ATP was replaced by [³²P]-labeled ATP (Catalog No. BLU002250UC, Perkin Elmer, Waltham, MA). Afterward, the reaction mixture was terminated by SDS loading buffer and subjected to immunoblot analysis. Then SDS-PAGE was sealed with photographic film together for 12 h before the film was fixed.

Co-immunoprecipitation

Interested plasmids were introduced into 293T cells for 48 h, then 293T cells overexpressing targeted proteins were lysed by buffer A (Catalog No. P0013B, Beyotime Biotechnology, Shanghai, China) (supplemented with 10 mM NaF, 1 mM Na₂VO_3, and protease inhibitor cocktails). The lysates were centrifuged at 10,000 r/min, 4 °C for 10 min. The separated supernatant was divided into three fractions and respectively incubated with negative IgG or reciprocal antibodies at 4 °C overnight. Then pre-processed protein A agarose (Catalog No. P3476, Sigma) was added to mixture. After another 2-h incubation, the bead-antibody-protein complex was isolated from mixture by centrifuging at 1000 r/min, 4 ℃ for 5 min. To remove the non-specific binding proteins, the bead-antibody-protein complex was washed with pre-cold PBS buffer for three times before the samples were subjected to immunoblot assay.

Cell viability assay

Both WT and AMPKα-KO MEF cells (5000 per well) were pre-treated with 250 nM JQ-1 for 12 h, followed by exposure to increasing doses of X-ray (0, 2, 4, and 8 Gy).

M059J cells (7500 per well) were pre-treated with 250 nM JQ-1 and 1 μM Compound C for 12 h, followed by exposure to increasing doses of X-ray (0, 2, 4, and 8 Gy). After 72-h recovery, the cells were subjected to MTS assay according to the manufacturer’s instructions. 10 μl per well of MTS/PMS (20:1, Promega) solution was added to each well containing 100 μl of culture medium, followed by a gentle shake. After incubation at 37 °C under 5% CO₂ for 4 h, the absorbance of the solutions was measured at 490 nm, using an M5 microplate reader (Molecular Device, San Jose, CA).

Cell cycle assay

Both WT and AMPKα-KO MEF cells (7.5 × 10⁴/well) were exposed to a single dose of 8 Gy X-ray. After 24-h recovery, cells were sampled by EDTA-free trypsin and 75% ethanol. Fixed cells were then incubated with 50 μg/ml propidium iodide (Catalog No. P4170, Sigma) and 100 μg/ml RNaseA (Catalog No. ST576, Beyotime Biotechnology) for 15 min at room temperature. The mean fluorescence intensity of DNA content was recorded by Flow cytometer (NovoCyte D2060R, San Diego, CA) by PE channel. 10,000 events per sample were collected and analyzed by Software NoveExpress.

Cell apoptosis assay

The cell apoptosis assay was performed by Annexin V-FITC/PI apoptosis detection kit (Catalog No. KGA108, KeyGEN BioTECH, Nanjing, China) according to the manufacturer’s instructions. Briefly, 2 × 10⁵ cells were prepared according to experimental design. At the sample point, cells were digested with EDTA-free trypsin and incubated with staining solution (500 μl Detection Buffer supplemented with 5 μl PI and 5 μl Annexin-V) for 15 min at room temperature. The fluorescence intensity was recorded by Flow cytometer (NovoCyte D2060R) by FITC and PE channel after fluorescence compensation deduction. 15,000 events per sample were collected and analyzed. Total apoptotic cells were FITC⁺PI⁻ cells plus FITC⁻PI⁺ cells. Data are presented as a densitometric ratio change normalized to the negative control.

Data availability

All MS raw data have been deposited to the ProteomeXchange Consortium via the iProX partner repository (ProteomeXchange: PXD039113; iProX: IPX0001446000), which are publicly accessible at http://proteomecentral.proteomexchange.org and https://www.iprox.cn, respectively.

CRediT author statement

Yuejing Jiang: Conceptualization, Methodology, Validation, Formal analysis, Investigation, Data curation, Writing - original draft, Writing - review & editing, Visualization. Xiaoji Cong: Methodology, Software, Data curation, Formal analysis, Investigation, Data curation, Writing - original draft, Writing - review & editing, Visualization. Shangwen Jiang: Methodology, Software, Formal analysis, Investigation. Ying Dong: Formal analysis, Investigation. Lei Zhao: Software, Investigation. Yi Zang: Conceptualization, Resources, Writing - original draft, Writing - review & editing, Supervision, Project administration, Funding acquisition. Minjia Tan: Conceptualization, Resources, Writing - original draft, Writing - review & editing, Supervision, Project administration, Funding acquisition. Jia Li: Conceptualization, Resources, Supervision, Project administration, Funding acquisition. All authors have read and approved the final manuscript.

Competing interests

The authors declare no competing interests.

Supplementary material

Supplementary data to this article can be found online at https://doi.org/10.1016/j.gpb.2020.09.003.

Acknowledgments

This study was supported by the National Natural Science Foundation of China (Grant Nos. 81872888, 81821005, 81673489, and 31871414), the Special Project on Precision Medicine under the National Key R&D Program (Grant No. 2017YFC0906600), the Shanghai Science and Technology Development Funds, China (Grant No. 19JC1416300), the Key New Drug Creation and Manufacturing Program of China (Grant Nos. 2018ZX09711002-004 and 2018ZX09711002-007), and the KC Wong Education Foundation.

References

Author notes