Abstract
Ataxia-telangiectasia mutated (ATM) is an apical kinase involved in the cellular response to DNA damage in eukaryotes, especially DNA double-strand breaks (DSBs). Upon DSB, ATM is activated through a hierarchy of well-organized cellular processes and machineries, including post-translational modifications (PTMs), the MRE11-RAD50-NBS1 (MRN) complex and chromatin perturbations. ATM activation initiates a cascade of chromatin modifications and nucleosome remodeling that permits the assembly of repair factors that ensure a highly orchestrated response to repair damaged DNA. Numerous studies have tried to elucidate the mechanisms of ATM activation, but how it is activated by DNA damage signals is still unclear. Histone modifications are considered essential for regulating ATM activation: a histone octamer constitutes the nucleosome core and histone tails protrude into the DNA strands to alter the chromatin landscape and DNA accessibility. Here, we summarize how histone modifications regulate ATM activation, with an emphasis on the functional relevance in DNA damage response and repair.
Similar content being viewed by others
Avoid common mistakes on your manuscript.
Introduction
Genome stability and integrity are constantly challenged by exogenous insults (irradiation, UV, chemical regents, etc.) and endogenous damage (metabolic wastes, replication stress, etc.), which lead to DNA damage and may alter the genetic information if left unrepaired. DNA double-strand breaks (DSBs), in which both strands of the DNA duplex are broken, are the most dangerous and deleterious form of DNA damage. DSB repair requires a massive complicated machinery that involves hundreds of proteins that survey mega bases (Mb) of DNA sequence around and flanking the DSB site. In eukaryotic cells, DSBs are mainly repaired through two pathways: non-homologous end joining (NHEJ) and homologous recombination (HR). The pathway type is chosen depending on genomic location of the DSB, cell-cycle stage and genetic background (Sancar et al. 2004). Deficiencies in responding and repairing DSBs have been linked to various human disorders, including cancer (Jackson and Bartek 2009). Uncovering the nature of the DSB response and repair, therefore, is informative for disease prevention and intervention.
Among the hierarchical events in response to DNA DSBs, activation of Ataxia-telangiectasia mutated (ATM) is considered the most critical. ATM deletion or mutation leads to near-complete loss of response to DSBs, as evident in patients with Ataxia-telangiectasia (A-T) and ATM-deficient cells (Savitsky et al. 1995; Barlow et al. 1996; Elson et al. 1996; Xu et al. 1996). Failure to robustly activate ATM also results in an inadequate DNA damage response and thus inappropriate repair of DSBs, even if ATM is genetically intact and functional. This scenario is evident in various genetic disorders, such as Nijmegen Breakage Syndrome (NBS), caused by mutation or deletion of NBS1 (Carney et al. 1998; Varon et al. 1998). As a protein serine/threonine kinase, ATM was first recognized as a key regulator of the DNA damage response when it was linked to A–T, which displays hypersensitivity to irradiation and cancer predisposition. Since then, ATM has been demonstrated to regulate the DNA damage response and DNA repair by phosphorylating various substrates, including p53, histone H2AX, checkpoint kinase 2 (CHK2) (Banin et al. 1998; Canman et al. 1998; Rogakou et al. 1998; Burma et al. 2001; Matsuoka et al. 1998, 2000). Mechanistically, ATM facilitates DSB repair factor loading in three main ways: (i) by creating a platform to recruit and assemble repair factors, (ii) by changing nucleosome accessibility by altering local chromatin states, and (iii) by promoting histone exchange through chromatin remodeling (Sirbu and Cortez 2013). For example, ATM-mediated H2AX phosphorylation at serine 139 (also termed γH2AX) recruits mediator of checkpoint 1 (MDC1), which in turn recruits ATM through a positive feedback loop to amplify ATM signals (Stucki and Jackson 2006). The γH2AX/MDC1 platform also recruits downstream chromatin modifiers, such as the ubiquitin E3 ligases RNF8/168 to foster sequential loading of repair factors (Stucki and Jackson 2006; Mailand et al. 2007; Huen et al. 2007; Kolas et al. 2007).
The mechanisms underlying ATM activation remained obscure until it was demonstrated that ATM autophosphorylation at its serine 1981 residue (S1981) was essential (Bakkenist and Kastan 2003). ATM homodimers or multimers in quiescent cells are disrupted into monomers by DNA damage-induced intermolecular phosphorylation, which was later proved to take place at multiple sites and not just S1981 alone (Bakkenist and Kastan 2003; Pellegrini et al. 2006; Kozlov et al. 2006; Daniel et al. 2008; (Kozlov et al. 2011). ATM phosphorylation by other kinases, such as cyclin-dependent kinase 5 (CDK5) and DNA-dependent protein kinase (DNA-PK), is also important for ATM activation (Tian et al. 2009; Zhou et al. 2017). Other post-translational modifications (PTMs), including acetyltransferase TIP60-mediated acetylation and protein phosphatase 2A/2C (PP2A/2C)-mediated dephosphorylation are also crucial ATM regulators (Sun et al. 2005; Goodarzi et al. 2004; Shreeram et al. 2006). The MRE11-RAD50-NBS1 (MRN) complex is another indispensable cellular component in regulating ATM activity. Early observations in patients with NBS and Ataxia-telangiectasia like disease (ATLD) that is caused by MRE11 mutation or deletion, documented a marked similarity in patient phenotypes to patients with A-T, including radio-sensitivity and cancer predisposition (Carney et al. 1998; Varon et al. 1998; Stewart et al. 1999). Later studies revealed that MRN complex functions as the main damage sensor upstream of ATM signaling (Carson et al. 2003). It was further confirmed that MRN is required for both initial and robust ATM activation in vivo and in vitro, by promoting ATM monomerization, recruitment to damage sites and interaction with its substrates (Uziel et al. 2003; Lee and Paull 2004; Difilippantonio et al. 2005; Lee and Paull 2005; Dupre et al. 2006; Falck et al. 2005). Moreover, nuclease activity of MRE11 is also required for ATM activation, probably through generation of ssDNA oligos when processing DNA breaks (Dupre et al. 2008; Jazayeri et al. 2008). In vivo activation of ATM indeed depends on the presence of DNA, as dimeric ATM cannot be activated or autophosphorylated when DNA is absent (Lee and Paull 2005; Dupre et al. 2006). Despite DNA damage-induced ATM activation being regulated by the above-mentioned mechanisms, ATM activation under other circumstances may not necessarily need any of them. For example, oxidative stress activates ATM by forming an intermolecular disulfide bond at cysteine 2991 residue (C2991), which is independent of autophosphorylation or MRN (Guo et al. 2010). In addition, ATM interacting protein (ATMIN) mediates ATM activation upon hypotonic stress through a MRN-independent pathway (Kanu and Behrens 2007).
Chromatin alterations may also participate in regulating ATM activation, as Bakkenist CJ et al. first reported that non-DNA damaging stimuli, such as histone deacetylase (HDAC) inhibitors and sodium chloride, could also potently activate ATM, even without eliciting any DNA breaks (Bakkenist and Kastan 2003). Since this discovery, it was anticipated that chromatin may sense initial DNA damage signals and transduce these signals to ATM through chromatin alterations or modifications. Later studies proved this theory by showing that chromatin modifications, including certain types of histone modifications, are mechanistically and functionally relevant to ATM activation. This finding is expected and rational, as histones are so tightly linked to DNA that histone modifications are known to impact almost all DNA properties and functions, such as replication and transcription. Consequently, histone modifications have also been extensively explored in terms of how they regulate the DNA damage response and DNA repair (Cao et al. 2016). The complexity of histone modifications adds another layer of regulation so that the DNA damage response and repair machinery can be loaded to damaged chromatin in a precise and orchestrated manner. In this review, we detail the mechanisms as to how histone modifications regulate ATM activation. We elaborate the implications of these modifications on the DNA damage response and repair, as well as other cellular functions.
Histone phosphorylation
One of the best-characterized histone modifications involved in ATM activation is γH2AX (Fig. 1), which is among the first substrates phosphorylated by ATM upon DNA damage (Rogakou et al. 1998; Burma et al. 2001). As a variant of the core histone H2A, H2AX comprises ~ 10% of the total H2A in the eukaryotic genome (Fernandez-Capetillo et al. 2004), suggesting that only one in 10 nucleosomes contains H2AX. H2AX phosphorylation occurs at DNA damage sites within a few seconds and spreads over 1 Mb flanking regions (Rogakou et al. 1998), indicating that H2AX undergoes rapid exchange and re-localization in response to DNA damage. H2AX-deficient mice display various DNA-damage-associated phenotypes, including increased sensitivity to radiation, chromosome instability, growth retardation, immune deficiency, and infertility (Celeste et al. 2002, 2003). These phenotypes are largely shared by ATM-deficient mice and A–T patients (Savitsky et al. 1995; Barlow et al. 1996; Elson et al. 1996; Xu et al. 1996), suggesting a critical role for H2AX in the ATM pathway. In addition, H2AX S139 mutation specifically sensitizes mice or cells to DNA DSBs (Celeste et al. 2003).
Regulation of ATM activation by γH2AX. ATM is recruited to DSB sites by MRN complex and activated. The phosphorylation of H2AX (γH2AX) by ATM is recognized by the BRCT domain of MDC1 and the phosphorylation of MDC1 by CK2 is recognized by NBS1 component of MRN complex. This promotes the recruitment of another MRN complex, which in turn recruits and activates more ATM molecules for another cycle. This positive feedback loop leads to the spreading of DNA damage-induced ATM signaling along the DSB-flanking chromatin, thus promoting the robust and full ATM activation
Although γH2AX is dispensable for initially recognizing DNA breaks (Celeste et al. 2003), it amplifies ATM-dependent phosphorylation signals through a positive feedback loop. Mechanistically, γH2AX recruits MDC1 via a direct interaction with its breast cancer C-terminal (BRCT) domain (Stucki et al. 2005; Lou et al. 2006). MDC1 phosphorylation by casein kinase 2 (CK2) is then recognized by NBS1 and promotes MRN complex recruitment and retention, which in turn recruits and activates ATM. This process leads to the stabilization and amplification of DNA damage-induced phosphorylation signals (Spycher et al. 2008; Melander et al. 2008; Wu et al. 2008; Chapman and Jackson 2008). Despite the established role of H2AX phosphorylation in ATM activation, a H2A variant found in Arabidopsis (H2A.W.7) is primarily confined to heterochromatin, and is also phosphorylated by ATM and required for an ATM-dependent DNA damage response in heterochromatic regions (Lorkovic et al. 2017). Moreover, H2AX phosphorylation at tyrosine 142 (T142), which is regulated by the WSTF kinase and rapidly dephosphorylated by EYA protein phosphatase upon DNA damage, is also critical for an ATM-dependent DNA damage response (Xiao et al. 2009; Cook et al. 2009; Krishnan et al. 2009). H2AX phosphorylation at S139 and T142 coordinates the DNA damage response by recruiting Microcephalin (MCPH1) (Singh et al. 2012), suggesting a crosstalk between these H2AX phosphorylation events.
In addition to H2A and its variants, phosphorylation of other histones is also involved in ATM signaling. For example, in budding yeast, histone H2B is phosphorylated at threonine 129 (T129) in a Mec1 (yeast homolog of ATR)-dependent and Tel1 (yeast homolog of ATM)-dependent manner (Lee et al. 2014). H2B phosphorylation shows a similar pattern to γH2AX, despite differences in telomeric regions (Lee et al. 2014), suggesting that H2B phosphorylation may also be an active regulator of the DNA damage response. Early studies showed that histone H1 and H3 are instantaneously dephosphorylated upon DNA damage, whereas H1 dephosphorylation is dependent on ATM (Guo et al. 1999, 2000). Although direct evidence is still lacking, these dynamic modifications may have an indirect influence on the DNA damage response. H3 serine 10 (S10) dephosphorylation in G1 phase correlates with γH2AX upon DNA damage (Sharma et al. 2015, 2015). As H3S10 phosphorylation is critical in chromatin condensation (Johansen and Johansen 2006), these data suggest its potential involvement in ATM activation during cell-cycle progression. In addition, H3 phosphorylation at threonine 11 (T11) by checkpoint kinase 1 (CHK1) is also a DNA damage responsive phosphor-mark and DNA damage induces rapid dephosphorylation of H3T11 by protein phosphatase 1γ (PP1γ) (Shimada et al. 2008, 2010). H3T11 phosphorylation is prevented by H3S10 phosphorylation through an intramolecular autoinhibitory feedback mechanism (Liokatis et al. 2012), indicating a dual role for H3S10 phosphorylation in the DNA damage response.
Histone methylation
Chromatin is marked at multiple sites by histone methylation, which carries distinct epigenetic information and coordinates a wide range of cellular functions. Histone methylation is regulated by a group of lysine methyltransferases (KMTs) and demethylases (KDMs), which have a high specificity to target different lysine residues. Various histone methylation sites are dynamically altered, either globally or locally upon DNA damage (Chen and Zhu 2016). One of the key histone methylation sites involved in ATM signaling is H3 lysine 9 (H3K9) (Fig. 2), which is initially identified as an essential factor in gene repression and heterochromatin formation. H3K9 trimethylation (H3K9me3) recruits a repressive complex containing heterochromatin protein 1 (HP1) (Bannister et al. 2001; Nielsen et al. 2001; Lachner et al. 2001; Nielsen et al. 2002; Jacobs and Khorasanizadeh 2002), whereas loss of H3K9me3 or its enzymes results in chromatin relaxation and genome instability (Peng and Karpen 2009; Wang et al. 2013). In response to DNA damage, SUV39H1, a major KMT for H3K9me3, is recruited to DSB sites by chromatin poly-ADP-ribosylation (PARylation), which promotes chromatin compaction through H3K9me3 and the KAP1/HP1/SUV39H1 complex (Ayrapetov et al. 2014). TIP60, which is recruited by MRN in the form of ATM-FOXO3a-TIP60 complex (Tsai et al. 2008; Sun et al. 2009; Adamowicz et al. 2016), is then activated by H3K9me3 via its chromodomain and activates ATM by acetylating lysine 3016 (K3016) (Sun et al. 2005, 2009). These studies demonstrate that H3K9me3 and transient chromatin condensation is required for ATM activation. Nevertheless, chromatin needs to be open to facilitate the expansion of DNA damage signals to distal regions and the recruitment of downstream repair factors. After the initial wave of DNA damage sensing and transducing, activated ATM phosphorylates KAP1 and removes the KAP1/HP1/SUV39H1 complex (Ayrapetov et al. 2014; Ziv et al. 2006), which forms a negative feedback loop to achieve ATM-dependent chromatin decompaction. In addition, KDM4B and KDM4D, which are responsible for demethylating H3K9me2/3, are recruited to DNA damage sites in a PARP1-dependent manner (Young et al. 2013; Khoury-Haddad et al. 2014), indicating an active demethylation of H3K9me2/3 after ATM activation. More importantly, KDM4D is required for efficient ATM activation and recruitment (Khoury-Haddad et al. 2014), and oncogenic stress induced heterochromatin retains DDR signaling (Di Micco et al. 2011), further suggesting that dynamic H3K9 methylation and demethylation are indispensable for timely and orderly ATM activation.
Regulation of ATM activation by H3K9 trimethylation. In response to DNA damage, KAP1/HP1/SUV39H1 complex is recruited to DSB sites by PARP1-dependent chromatin PARylation. SUV39H1 catalyzes H3K9me3, which recruits more SUV39H1 complex through interaction with HP1, thus promoting H3K9me3 spreading in a positive feedback loop. The H3K9me3 activates TIP60 HAT activity and TIP60 phosphorylation by c-Abl promotes its interaction with H3K9me3. The activated TIP60 then promotes ATM activation through acetylation-dependent autophosphorylation. The activated ATM further promotes KAP1 phosphorylation and disassociation from chromatin, and KDM4B/D recruited by chromatin PARylation promotes the demethylation of H3K9, indicating the highly dynamic behaviour of H3K9me3 in ATM orderly activation
H3 lysine 36 (H3K36) methylation is another important histone mark that has been linked to ATM activation. H3K36 dimethylation (H3K36me2) is induced at DNA damage sites and potentiates NBS1 and Ku70 recruitment; K36 mutation impairs the enrichment of these factors and compromises DSB repair (Fnu et al. 2011; Cao et al. 2016). As NBS1 is a key component of the ATM regulatory machinery, it is expected that H3K36me2 may modulate ATM activation. Indeed, we showed that KDM2A, the major KDM for H3K36me2, is phosphorylated by ATM and displaced from DSBs. This process promotes H3K36me2 and MRN complex recruitment, forming a positive feedback loop to achieve robust ATM activation (Cao et al. 2016). Contrary to H3K36me2, H3K36 trimethylation (H3K36me3) is not induced by DNA damage, yet it is still actively involved in the DNA damage response and DNA repair. Depletion or mutation of SETD2, a major H3K36me3 KMT, diminishes ATM activation, p53-mediated checkpoint activation, loading of HR factors and DSB repair (Pfister et al. 2014; Carvalho et al. 2014; Jha and Strahl 2014). H3K36me3 also recruits several readers that may be implicated in ATM activation. For example, plant homeodomain finger protein 1 (PHF1) recruitment via an interaction between its Tudor domain and H3K36me3 is required for its loading onto DNA damage sites (Hong et al. 2008; Musselman et al. 2012). The fact that PHF1 is involved in DNA damage repair and increases nucleosome accessibility raises the possibility that H3K36me3 may direct ATM signaling via PHF1 (Hong et al. 2008; Musselman et al. 2013). Moreover, crosstalk between H3K36me3 and H4K16 acetylation results in TIP60 recruitment through lens epithelium-derived growth factor p75 splicing variant (LEDGF), an H3K36me3 reader (Li and Wang 2017). The established role of TIP60 in ATM activation also prompts us to speculate a more direct link between H3K36me3 and ATM activation.
Beside H3K9 and H3K36 methylation, many other histone sites that are methylated are also potential modulators of the DNA damage response and ATM activation. For example, H4K20 methylation is induced by MMSET in an ATM-dependent manner, which is important for loading p53-binding protein 1 (53BP1), a key factor involved in DSB repair (Pei et al. 2011). Recent data showed that DSB-induced small RNAs (diRNAs)-dependent recruitment of MMSET and TIP60 potentiates a more flexible chromatin through H4K20 methylation and H4K16 acetylation (Wang and Goldstein 2016). MMSET and H4K20 methylation, however, does not affect ATM-dependent γH2AX formation in these settings (Pei et al. 2011; Wang and Goldstein 2016), suggesting a specific role of H4K20 methylation in the DNA damage response. A similar methylation site is H3 lysine 79 (H3K79), which is not induced by DNA damage, but is required for 53BP1 recruitment (Huyen et al. 2004; Giannattasio et al. 2005; Wysocki et al. 2005). The fact that 53BP1 interacts with the MRN complex and can promote MRN-dependent ATM activation (Lee et al. 2010), however, indicates a possible feedback loop between these histone modifications and ATM signaling through 53BP1. DNA damage induces accumulation of H3K27 trimethylation (H3K27me3) and enhancer of zeste homolog 2 (EZH2), the KMT for H3K27me3, via a PARP1-dependent pathway (Chou et al. 2010). In addition, as EZH2 regulates the cellular response to different DNA damage inducers, its deletion sensitizes cells to DNA damage (Wu et al. 2011; Sha et al. 2016). Moreover, ATM regulates EZH2 stability through phosphorylation, and ATM deficiency leads to elevated H3K27me3 levels in A–T neural cells (Li et al. 2013). Whereas H3K27me3 patterns correlate with deregulated neural genes in A–T brains, EZH2 knockdown rescues the neurological abnormalities (Li et al. 2013), suggesting a direct role for EZH2 and H3K27me3 in ATM function in the neural system. A recent study further confirmed that EZH2 regulates ATM activation and its subsequent involvement in lipid metabolism and the DNA damage response in glioblastoma (Ahmad et al. 2017).
Histone acetylation
Histone acetylation is dynamically balanced by histone acetyltransferases (HATs) and HDACs. While some HDAC inhibitors have been approved to treat certain types of malignancies (Li and Zhu 2014), they are also DNA damage inducers and regulators (Namdar et al. 2010; Lee et al. 2010; Robert et al. 2011; Wang et al. 2012). The observation that HDAC inhibitors can potently activate ATM leads to the presumption that histone acetylation may also be a bona fide regulator of ATM activation (Bakkenist and Kastan 2003). Different from the mechanisms employed by histone phosphorylation and methylation, which act through the recruitment of specific readers, histone acetylation mainly alters local chromatin states and accessibility to connect chromatin alterations and ATM activation. Kim YC et al. first demonstrated that the nucleosome-binding protein high mobility group N1 (HMGN1) optimizes ATM activation by regulating its interaction with chromatin through histone H3 at lysine 14 (H3K14) acetylation (Kim et al. 2009). The researchers found that exposing cells to an HDAC inhibitor upregulated H3K14 acetylation and bypassed the requirement of HMGN1 for efficient ATM activation (Kim et al. 2009). These data suggest that H3K14 acetylation may regulate ATM chromatin retention and activation.
Another important histone acetylation mark associated with ATM activation is acetylation of histone H4 at lysine 16 (H4K16). H4K16 acetylation is a predominant factor controlling chromatin structure and nucleosome mobilization (Shogren-Knaak et al. 2006), and is required for ATM activation. Depletion of the acetyltransferases responsible for H4K16 acetylation, including MOF and TIP60, leads to profound defects in ATM activation and the DNA damage response (Sun et al. 2005; Gupta et al. 2005; Sharma et al. 2010). TIP60 dictates ATM activation via direct modification of its C terminus, whereas MOF regulates ATM activity through H4K16 acetylation. MOF interacts with ATM: MOF deletion results in defective ATM autophosphorylation and the ATM-dependent DNA damage response (Gupta et al. 2005). Mice with MOF-deficiency in Purkinje cells display a neurological disorder similar to A-T patients (Kumar et al. 2011), suggesting a functional synergy between MOF and ATM in the DNA damage response. Zmpste24 null mice, with defective MOF function and hypoacetylation at H4K16, also exhibit an impaired response to DNA damage and cellular senescence: this phenotype can be rescued by MOF reintroduction and exposure to HDAC inhibitors (Krishnan et al. 2011). Nuclear c-Abl tyrosine kinase, which induces chromatin structural changes through H4K16 hypoacetylation, mediates ATM activation through phosphorylation of TIP60, thus coupling chromatin sensing to ATM signaling (Aoyama et al. 2011; Kaidi and Jackson 2013). The E3 ligases RNF8 and CHFR synergistically regulate ATM activation through histone ubiquitination and H4K16 acetylation (Wu et al. 2011). DNMT1-associated protein 1 (DMAP1), a member of the TIP60-p400 complex, is required for efficient ATM activation in response to DNA damage and hypotonic stress through regulation of H4K16 acetylation (Penicud and Behrens 2014). These reports strongly support that H4K16 acetylation is a pivotal factor involved in the onset and amplification of ATM signaling, mainly through its role in reshaping the local chromatin environment.
Other histone residues are also acetylated and implicated in the ATM-dependent DNA damage response. For example, TIP60-mediated H2AX acetylation at lysine 5 permits its ubiquitination, which further stimulates histone exchange and the DNA damage response (Ikura et al. 2007). In addition, H2AX acetylation is required for NBS1 turnover at DNA damage sites, which may restrict γH2AX expansion to undamaged regions (Ikura et al. 2015). These data suggest an indirect role for H2AX acetylation in fine-tuning ATM activation. In yeast, histone H4 acetylation at multiple sites, including lysines 5, 12 and 91, is also required for a proper DNA damage response, as mutating these lysine residues causes defective γH2AX domain formation and poor cell survival upon DNA damage (Ge et al. 2013). Histone H3 lysine 56 (H3K56) acetylation, which marks newly synthesized histones, is induced by DNA damage and shows a perfect co-localization with γH2AX (Das et al. 2009; Vempati et al. 2010), indicating a positive role for H3K56 acetylation in the DNA damage response. Later studies have demonstrated that H3K56 is specifically deacetylated at DNA damage sites by the instantly recruited histone deacetylases, HDAC1/2 and SIRT6 (Miller et al. 2010; Toiber et al. 2013), suggesting a dynamic nature of DNA damage-induced H3K56 acetylation. H3K9 acetylation similarly decreases upon DNA damage and constitutive H3K9 hyperacetylation impairs ATM activation in embryonic stem cells (Tjeertes et al. 2009; Meyer et al. 2016). p53-dependent H3K9 and H3K27 acetylation at sub-telomeric regions prevents γH2AX signal accumulation at telomeres (Tutton et al. 2016), suggesting a potential role for histone acetylation in restraining the ATM-dependent DNA damage response in specific chromosomal regions. Recently, we showed that acetylation of linker histone H1, a less-characterized chromatin mark, may also participate in regulating chromatin structure and genome stability in the DNA damage response (Li et al. 2018). Together, these studies show that histone acetylation is mainly implicated in ATM activation through altering local or general chromatin structure, and eventually controls a wide aspect of the DNA damage response and DNA repair.
Histone ADP-ribosylation
ADP-ribosylation is mediated by a group of poly-(ADP-ribose)-polymerases (PARPs), with PARP1 being the most prominent and best characterized. Unlike phosphorylation, methylation or acetylation, ADP-ribosylation is the transfer of one ADP-ribose moiety (mono-ADP-ribosylation) or polymeric ADP-ribose chains (poly-ADP-ribosylation, PARylation) to acceptor proteins (Hottiger 2015). Known acceptors in eukaryotic cells include lysine, arginine, glutamate, aspartate, cysteine, serine and asparagine residues, but the specificity remains ambiguous (Messner and Hottiger 2011; Rosenthal and Hottiger 2014). ADP-ribosylation is reversible via the action of ADP-ribose removing enzymes and tends to be more labile and transient than other small covalent modifications, which only last for a short time in vivo (Hottiger 2015; Koch-Nolte et al. 2008). The fact that only a small fraction (~ 4%) of histones is ADP-ribosylated adds up the complexity of dissecting the functions of site-directed histone ADP-ribosylation in DNA damage and repair (Boulikas 1989). Most previous studies have focused on the modifying enzymes PARP1 and PAR glycohydrolase (PARG). PARP1 is one of the earliest sensors of DNA breaks, being activated within seconds of detecting damage (Polo and Jackson 2011). PARP1 deletion or inhibition results in hypersensitivity to DNA damage inducers and compromised ATM activation (de Murcia et al. 1997; Bryant et al. 2005; Haince et al. 2007). PARP1-dependent chromatin decondensation is also one of the earliest events in the DNA damage response, which may license the spreading of phosphorylation signals, as chromatin compaction restricts the intensity of DNA damage signaling (Murga et al. 2007). PARG, the principal enzyme to hydrolyse PAR chains, is also recruited to DNA damage sites (Mortusewicz et al. 2011). PARG deletion or inhibition enhances γH2AX foci formation, p53 activation, and sensitizes cells to DNA damage (Shirai et al. 2013; Gravells et al. 2017, 2018), indicating a positive link between PARG and ATM activation.
The primary histone ADP-ribosylation targets upon DNA damage are histone H2A and H3, with a minor amount of other histones, such as H1 and H4 (Messner and Hottiger 2011; Jungmichel et al. 2013). Although information of site specificity is limited, recent studies using a series of proteomic screening methods have unraveled several sets of ADP-ribosylation targets under stressed conditions and revealed that serine is the major target on histones upon DNA damage (Leidecker et al. 2016; Palazzo et al. 2018; Fontana et al. 2017). Very recently, we identified that PARylation of linker histone H1.2 at serine 188 (S188) mediates H1.2 dynamics and is required for ATM activation upon DNA damage (Li et al. 2018) (Fig. 3). Mechanistically, H1.2 inhibits ATM activation and recruitment through a direct interaction and an MRN-dependent mechanism (Li et al. 2018). In response to DNA damage, H1.2 is PARylated, which allows its dissociation from chromatin and permits the interaction between ATM and MRN, thus facilitating robust ATM activation (Li et al. 2018). Therefore, we provided the first evidence of site-directed histone ADP-ribosylation in regulating ATM activity and the DNA damage response. While more information is still emerging, explorations of other histone ADP-ribosylation sites in DNA damage and crosstalk with other PTMs are of particular interest.
Regulation of ATM activation by linker histone H1.2 PARylation and destabilization. In the absence of DNA damage, the chromatin-binding H1.2 prevents ATM recruitment and activation through the competing interaction with MRN complex. Upon DNA damage, H1.2 is PARylated by PARP1 and disassociated from chromatin for degradation, thereby permitting the recruitment and activation of ATM by MRN complex and DNA breaks. The activated ATM is amplified by an ATM-MDC1-MRN positive feedback loop, leading to efficient DNA damage response and repair
Histone ubiquitination
Similar to ADP-ribosylation, ubiquitination involves the covalent conjugation of a conserved 76-residue polypeptide (mono-ubiquitination) or polypeptide chains (poly-ubiquitination) to target proteins. Ubiquitination is sequentially catalyzed by a group of enzymes including: an E1 activating enzyme, an E2 conjugating enzyme and an E3 ligase that specifically transfers the ubiquitin to substrates, which may eventually alter protein function or stability (Zheng and Shabek 2017). As versatile as ubiquitination may be, histone ubiquitination is extensively involved in chromatin-based cellular processes, such as the DNA damage response and repair. As a crucial part of the hierarchical chromatin signaling network guiding the DNA damage response and repair, histone ubiquitination facilitates the enrichment of several key repair factors, such as 53BP1, onto damaged chromatin (Uckelmann and Sixma 2017). For example, H2A/H2AX ubiquitination at lysine 13/15 (K13/15) by RNF168 is required for repair factor loading, including 53BP1, which is a specific reader of the H2A K15 ubiquitination mark (Mailand et al. 2007; Huen et al. 2007; Kolas et al. 2007; Mattiroli et al. 2012; Wilson et al. 2016). Later studies have shown that RNF168 recruitment is dependent on RNF8-mediated ubiquitination of linker histone H1, which is recognized by RNF168 through its ubiquitin-dependent DSB recruitment module (UDM) (Thorslund et al. 2015). Nevertheless, a very recent study showed that lethal (Savitsky et al. 1995) malignant brain tumor-like protein 2 (L3MBTL2), rather than histone H1, is involved in the sequential RNF8 and RNF168 recruitment and ubiquitin signal amplification in response to DNA damage (Nowsheen et al. 2018).
Although generally believed to function downstream of phosphorylation, histone ubiquitination is also reported to impact on ATM full activation through certain de-ubiquitination feedback loops. For example, USP22 is a member of the de-ubiquitination module of the Spt-Ada-Gcn5-acetyltransferase (SAGA) complex, and is required for γH2AX foci formation through H2B de-ubiquitination at lysine 120 (K120) (Ramachandran et al. 2016). As H2B K120 ubiquitination is dependent on ATM-mediated phosphorylation of the RNF20/RNF40 heterodimer (Moyal et al. 2011), these studies suggest a possible feedback loop centered on H2B ubiquitination that regulates ATM activation. More recently, we, together with other researchers, found that accumulation of p62, an autophagic cargo protein, inhibits RNF168 activity and leads to impaired H2A ubiquitination, which eventually disrupts ATM activation and signaling upon DNA damage (Walker et al. 2017; Wang et al. 2016).
Other modifications
Various other types of histone modifications have also been implicated in the DNA damage response and DNA repair. For example, ubiquitin-like modifications (UBLylations), which conjugate ubiquitin-like proteins to target acceptors, including small ubiquitin-like modifiers (SUMOs) and neural precursor cell expressed, developmentally downregulated 8 (NEDD8), have been identified in histones in the context of DNA damage (Wang et al. 2017). NEDDylation of histone H4 and H2A differentially impacts RNF168-dependent H2A ubiquitination, which alters the DNA damage response and repair (Ma et al. 2013; Li et al. 2014). Although more details are needed, SUMOylation orchestrates the loading of histone-modifying enzymes onto chromatin during the DNA damage response (Hendriks et al. 2015; Huang et al. 2016), which might regulate ATM activation through crosstalk with other histone modifications. Specifically, the yeast H2A variant, H2A.Z-2 is SUMOylated upon DNA damage and facilitates its exchange at damage sites (Fukuto et al. 2018), with a possible involvement in reshaping the local chromatin state and thus promoting the DNA damage response.
Histone acylation is a less well-characterized histone mark that is extensively implicated in metabolic regulation (Sabari et al. 2017), but is also involved in the DNA damage response and repair. For example, sirtuin 7 (SIRT7), a class III HDAC, regulates chromatin compaction and the DNA damage response by desuccinylating core histones at DNA damage sites (Li et al. 2016). Histone succinylation alters chromatin structure and nucleosome dynamics in a similar manner to acetylation (Sabari et al. 2017; Jing et al. 2018), thus it may also guide ATM activation in a similar way. Histone crotonylation, although not yet implicated in DNA damage, is also regulated and interpreted in an acetylation-resembled way and acts in combination with other histone modifications (Tan et al. 2011; Sabari et al. 2015; Zhao et al. 2016; Li et al. 2016; Andrews et al. 2016; (Xiong et al. 2016), which may be engaged in DNA damage response like acetylation. In addition, DNA damage-induced O-GlcNAcylation of H2A at serine 40 (S40) interacts with acetylated H2AZ and γH2AX at the initial phase of the DNA damage response, and is required for the accumulation of DNA damage response and repair factors (Hayakawa et al. 2017).
Conclusions and perspectives
Histones are subjected to numerous types of modification that regulate different aspects of cellular functions. The reversible and dynamic nature of histone modifications, with an upsurge in identifying novel modifications and their regulators, weaves a massive and multilayered molecular network to spatio-temporally regulate cellular responses in distinct contexts. In response to DNA damage, histone modifications protect genetic integrity and genomic stability through various mechanisms. Overall, we can conclude that ATM signaling is regulated by histone modifications to achieve three main aims (Table 1): (i) to create a chromatin mark (such as γH2AX and H3K9me3) to directly anchor or alienate ATM or its regulators; (ii) to reshape local chromatin structure (such as via most histone acylation marks) such that it indirectly affects ATM activation or recruitment; (iii) to alter ATM interactions (such as via H1 PARylation) with its regulators. While some histone modifications dictate ATM activation alone, it is noteworthy that most of them act and crosstalk with other modifications, which may elicit diverse effects than when operating alone. It is therefore more reasonable to consider histone modifications as combinations or clusters in the context of chromatin, rather than as a single epigenetic mark.
Our knowledge in histone modifications has grown exponentially over the past few years thanks to the development of high-throughput proteomic technologies. A major hinderance to delineate a clear and sequential effect of histone modifications in the DNA damage response is the lack of site-specific information, especially for non-canonical modifications, such as UBLylation and PARylation. Even if the modification site is known, it is usually challenging to create a site-directed mutation in mammals because mammalian cells express multiple copies of histone genes. In addition, mutagenesis assays can be pleiotropic and artificial, and thus may not represent the true function of the modification under analysis. Therefore, most studies have focused on the modifiers and specific readers of histone modifications. Chemical ligation or synthesis of modified histone is also informative for biochemical studies, but this strategy is largely limited to in vitro analysis (Holt and Muir 2015). Novel strategies or methodologies are now warranted to better illuminate the precise functions of site-specific histone modifications.
Although much is known about ATM, how ATM activation is triggered upon DNA damage, as well as upon many other stresses, remains elusive. One of the main obstacles is a lack of structural details of ATM and its regulating complexes. Recent studies have illustrated the structures of yeast Tel1 and human ATM at a relatively low resolution but these still provide some additional mechanistic insights into ATM activation (Wang et al. 2016; Baretic et al. 2017; Lau et al. 2016). A higher resolution of ATM structure, which will reveal its conformational changes upon stress or binding with its regulators, is now necessary for a more comprehensive understanding as to how ATM is activated. More importantly, DNA damage-induced activation of ATM relies on its binding to chromatin, which is dynamically regulated by histone modifications. Another critical issue that remains to be addressed is how ATM becomes disassembled and deactivated when DNA damage repair is completed. Previous articles showed that PP2A and PP5, two phosphatases, both regulated ATM activity in vivo. PP2A interacts with ATM in undamaged cells and irradiation induces PP2A dissociation from ATM, resulting in loss of its phosphatase activity (Goodarzi et al. 2004). Unlike PP2A, PP5 interacts with ATM in a DNA damage inducible manner and a catalytically active PP5 mutant inhibits ATM activation (Ali et al. 2004). Interestingly, Shreeram et al. found that another phosphatase wild-type p53-induced phosphatase (Wip1) overexpression was sufficient to downregulate the activation of ATM through dephosphorylating ATM Ser1981 as cells repaired damaged DNA (Shreeram et al. 2006). Moreover, recently our group reported that the histone deacetylase SIRT7-mediated deacetylation was essential for dephosphorylation and deactivation of ATM (Tang et al. 2019). We propose a novel model of ATM deactivation regulated by SIRT7-mediated deacetylation and subsequent by WIP1-mediated dephosphorylation. It is reasonable that the inactivation of ATM should follow a well-organized process and this process needs for more experimentation. Questions that remain include what is the consequence of persistent activation of ATM, what are the serial molecular events that trigger ATM removal and deactivation, does histone modification participate in this process, and are there any other modifications of ATM involved besides dephosphorylation and deacetylation. All these questions need to be answered with delicate studies.
As we have learnt more about the physiological roles of ATM, it has been proposed to be an apical regulator of the cellular response to stresses as much as it is a central kinase in DNA damage (Shiloh and Ziv 2013). Various stresses activate ATM, including oxidative stress, hypotonic stress, hypoxia, insulin, nitrosative stress (Bakkenist and Kastan 2003; Guo et al. 2010; Tripathi et al. 2013; Gibson et al. 2005; Yang and Kastan 2000), but how ATM is activated in these settings remains largely unknown. Insights into the regulatory mechanisms of ATM activation will greatly extend our perception of ATM as a broader sensor of stresses and not just a DNA damage regulator. Understanding how ATM is regulated in the above-mentioned physiological conditions will also help us to identify more promising drug targets and manipulate ATM activity in pathologies related to these stresses.
References
Adamowicz, M., Vermezovic, J., & Fagagna, F. (2016). NOTCH1 Inhibits Activation of ATM by Impairing the Formation of an ATM-FOXO3a-KAT5/Tip60 Complex. Cell Reports,16, 2068–2076.
Ahmad, F., Patrick, S., Sheikh, T., et al. (2017). Telomerase reverse transcriptase (TERT)—enhancer of zeste homolog 2 (EZH2) network regulates lipid metabolism and DNA damage responses in glioblastoma. Journal of Neurochemistry,143, 671–683.
Ali, A., Zhang, J., Bao, S., et al. (2004). Requirement of protein phosphatase 5 in DNA-damage-induced ATM activation. Genes and Development,18, 249–254.
Andrews, F. H., Shinsky, S. A., Shanle, E. K., et al. (2016). The Taf14 YEATS domain is a reader of histone crotonylation. Nature Chemical Biology,12, 396–398.
Aoyama, K., Fukumoto, Y., Ishibashi, K., et al. (2011). Nuclear c-Abl-mediated tyrosine phosphorylation induces chromatin structural changes through histone modifications that include H4K16 hypoacetylation. Experimental Cell Research,317, 2874–2903.
Ayrapetov, M. K., Gursoy-Yuzugullu, O., Xu, C., Xu, Y., & Price, B. D. (2014). DNA double-strand breaks promote methylation of histone H3 on lysine 9 and transient formation of repressive chromatin. Proceedings of the National Academy of Sciences of the United States of America,111, 9169–9174.
Bakkenist, C. J., & Kastan, M. B. (2003). DNA damage activates ATM through intermolecular autophosphorylation and dimer dissociation. Nature,421, 499–506.
Banin, S., Moyal, L., Shieh, S., et al. (1998). Enhanced phosphorylation of p53 by ATM in response to DNA damage. Science,281, 1674–1677.
Bannister, A. J., Zegerman, P., Partridge, J. F., et al. (2001). Selective recognition of methylated lysine 9 on histone H3 by the HP1 chromo domain. Nature,410, 120–124.
Baretic, D., Pollard, H. K., Fisher, D. I., et al. (2017). Structures of closed and open conformations of dimeric human ATM. Science Advances,3, e1700933.
Barlow, C., Hirotsune, S., Paylor, R., et al. (1996). Atm-deficient mice: a paradigm of ataxia telangiectasia. Cell,86, 159–171.
Boulikas, T. (1989). DNA strand breaks alter histone ADP-ribosylation. Proceedings of the National Academy of Sciences of the United States of America,86, 3499–3503.
Bryant, H. E., Schultz, N., Thomas, H. D., et al. (2005). Specific killing of BRCA2-deficient tumours with inhibitors of poly(ADP-ribose) polymerase. Nature,434, 913–917.
Burma, S., Chen, B. P., Murphy, M., Kurimasa, A., & Chen, D. J. (2001). ATM phosphorylates histone H2AX in response to DNA double-strand breaks. Journal of Biological Chemistry,276, 42462–42467.
Canman, C. E., Lim, D. S., Cimprich, K. A., et al. (1998). Activation of the ATM kinase by ionizing radiation and phosphorylation of p53. Science,281, 1677–1679.
Cao, L. L., Shen, C., & Zhu, W. G. (2016a). Histone modifications in DNA damage response. Science China Life Sciences,59, 257–270.
Cao, L. L., Wei, F., Du, Y., et al. (2016b). ATM-mediated KDM2A phosphorylation is required for the DNA damage repair. Oncogene,35, 301–313.
Carney, J. P., Maser, R. S., Olivares, H., et al. (1998). The hMre11/hRad50 protein complex and Nijmegen breakage syndrome: linkage of double-strand break repair to the cellular DNA damage response. Cell,93, 477–486.
Carson, C. T., Schwartz, R. A., Stracker, T. H., Lilley, C. E., Lee, D. V., & Weitzman, M. D. (2003). The Mre11 complex is required for ATM activation and the G2/M checkpoint. EMBO Journal,22, 6610–6620.
Carvalho, S., Vitor, A. C., Sridhara, S. C., et al. (2014). SETD2 is required for DNA double-strand break repair and activation of the p53-mediated checkpoint. Elife,3, e02482.
Celeste, A., Difilippantonio, S., Difilippantonio, M. J., et al. (2003a). H2AX haploinsufficiency modifies genomic stability and tumor susceptibility. Cell,114, 371–383.
Celeste, A., Fernandez-Capetillo, O., Kruhlak, M. J., et al. (2003b). Histone H2AX phosphorylation is dispensable for the initial recognition of DNA breaks. Nature Cell Biology,5, 675–679.
Celeste, A., Petersen, S., Romanienko, P. J., et al. (2002). Genomic instability in mice lacking histone H2AX. Science,296, 922–927.
Chapman, J. R., & Jackson, S. P. (2008). Phospho-dependent interactions between NBS1 and MDC1 mediate chromatin retention of the MRN complex at sites of DNA damage. EMBO Reports,9, 795–801.
Chen, Y., & Zhu, W. G. (2016). Biological function and regulation of histone and non-histone lysine methylation in response to DNA damage. Acta Biochimica et Biophysica Sinica (Shanghai),48, 603–616.
Chou, D. M., Adamson, B., Dephoure, N. E., et al. (2010). A chromatin localization screen reveals poly (ADP ribose)-regulated recruitment of the repressive polycomb and NuRD complexes to sites of DNA damage. Proceedings of the National Academy of Sciences of the United States of America,107, 18475–18480.
Cook, P. J., Ju, B. G., Telese, F., Wang, X., Glass, C. K., & Rosenfeld, M. G. (2009). Tyrosine dephosphorylation of H2AX modulates apoptosis and survival decisions. Nature,458, 591–596.
Daniel, J. A., Pellegrini, M., Lee, J. H., Paull, T. T., Feigenbaum, L., & Nussenzweig, A. (2008). Multiple autophosphorylation sites are dispensable for murine ATM activation in vivo. Journal of Cell Biology,183, 777–783.
Das, C., Lucia, M. S., Hansen, K. C., & Tyler, J. K. (2009). CBP/p300-mediated acetylation of histone H3 on lysine 56. Nature,459, 113–117.
de Murcia, J. M., Niedergang, C., Trucco, C., et al. (1997). Requirement of poly(ADP-ribose) polymerase in recovery from DNA damage in mice and in cells. Proceedings of the National Academy of Sciences of the United States of America,94, 7303–7307.
Di Micco, R., Sulli, G., Dobreva, M., et al. (2011). Interplay between oncogene-induced DNA damage response and heterochromatin in senescence and cancer. Nature Cell Biology,13, 292–302.
Difilippantonio, S., Celeste, A., Fernandez-Capetillo, O., et al. (2005). Role of Nbs1 in the activation of the Atm kinase revealed in humanized mouse models. Nature Cell Biology,7, 675–685.
Dupre, A., Boyer-Chatenet, L., & Gautier, J. (2006). Two-step activation of ATM by DNA and the Mre11-Rad50-Nbs1 complex. Nature Structural and Molecular Biology,13, 451–457.
Dupre, A., Boyer-Chatenet, L., Sattler, R. M., et al. (2008). A forward chemical genetic screen reveals an inhibitor of the Mre11-Rad50-Nbs1 complex. Nature Chemical Biology,4, 119–125.
Elson, A., Wang, Y., Daugherty, C. J., et al. (1996). Pleiotropic defects in ataxia-telangiectasia protein-deficient mice. Proceedings of the National Academy of Sciences of the United States of America,93, 13084–13089.
Falck, J., Coates, J., & Jackson, S. P. (2005). Conserved modes of recruitment of ATM, ATR and DNA-PKcs to sites of DNA damage. Nature,434, 605–611.
Fernandez-Capetillo, O., Lee, A., Nussenzweig, M., & Nussenzweig, A. (2004). H2AX: the histone guardian of the genome. DNA Repair (Amst),3, 959–967.
Fnu, S., Williamson, E. A., De Haro, L. P., et al. (2011). Methylation of histone H3 lysine 36 enhances DNA repair by nonhomologous end-joining. Proceedings of the National Academy of Sciences of the United States of America,108, 540–545.
Fontana P, Bonfiglio JJ, Palazzo L, Bartlett E, Matic I, Ahel I. Serine ADP-ribosylation reversal by the hydrolase ARH3. Elife 2017; 6.
Fukuto, A., Ikura, M., Ikura, T., et al. (2018). SUMO modification system facilitates the exchange of histone variant H2A.Z-2 at DNA damage sites. Nucleus-Phila,9, 87–94.
Ge, Z., Nair, D., Guan, X., Rastogi, N., Freitas, M. A., & Parthun, M. R. (2013). Sites of acetylation on newly synthesized histone H4 are required for chromatin assembly and DNA damage response signaling. Molecular and Cellular Biology,33, 3286–3298.
Giannattasio, M., Lazzaro, F., Plevani, P., & Muzi-Falconi, M. (2005). The DNA damage checkpoint response requires histone H2B ubiquitination by Rad6-Bre1 and H3 methylation by Dot1. Journal of Biological Chemistry,280, 9879–9886.
Gibson, S. L., Bindra, R. S., & Glazer, P. M. (2005). Hypoxia-induced phosphorylation of Chk2 in an ataxia telangiectasia mutated-dependent manner. Cancer Research,65, 10734–10741.
Goodarzi, A. A., Jonnalagadda, J. C., Douglas, P., et al. (2004). Autophosphorylation of ataxia-telangiectasia mutated is regulated by protein phosphatase 2A. EMBO Journal,23, 4451–4461.
Gravells, P., Grant, E., Smith, K. M., James, D. I., & Bryant, H. E. (2017). Specific killing of DNA damage-response deficient cells with inhibitors of poly(ADP-ribose) glycohydrolase. DNA Repair (Amst),52, 81–91.
Gravells, P., Neale, J., Grant, E., et al. (2018). Radiosensitization with an inhibitor of poly(ADP-ribose) glycohydrolase: A comparison with the PARP1/2/3 inhibitor olaparib. DNA Repair (Amst),61, 25–36.
Guo, Z., Kozlov, S., Lavin, M. F., Person, M. D., & Paull, T. T. (2010). ATM activation by oxidative stress. Science,330, 517–521.
Guo, C. Y., Mizzen, C., Wang, Y., & Larner, J. M. (2000). Histone H1 and H3 dephosphorylation are differentially regulated by radiation-induced signal transduction pathways. Cancer Research,60, 5667–5672.
Guo, C. Y., Wang, Y., Brautigan, D. L., & Larner, J. M. (1999). Histone H1 dephosphorylation is mediated through a radiation-induced signal transduction pathway dependent on ATM. Journal of Biological Chemistry,274, 18715–18720.
Gupta, A., Sharma, G. G., Young, C. S., et al. (2005). Involvement of human MOF in ATM function. Molecular and Cellular Biology,25, 5292–5305.
Haince, J. F., Kozlov, S., Dawson, V. L., et al. (2007). Ataxia telangiectasia mutated (ATM) signaling network is modulated by a novel poly(ADP-ribose)-dependent pathway in the early response to DNA-damaging agents. Journal of Biological Chemistry,282, 16441–16453.
Hayakawa, K., Hirosawa, M., Tani, R., Yoneda, C., Tanaka, S., & Shiota, K. (2017). H2A O-GlcNAcylation at serine 40 functions genomic protection in association with acetylated H2AZ or gammaH2AX. Epigenetics Chromatin,10, 51.
Hendriks, I. A., Treffers, L. W., Verlaan-de Vries, M., Olsen, J. V., & Vertegaal, A. C. (2015). SUMO-2 orchestrates chromatin modifiers in response to DNA damage. Cell Reports,10, 1778–1791.
Holt, M., & Muir, T. (2015). Application of the protein semisynthesis strategy to the generation of modified chromatin. Annual Review of Biochemistry,84, 265–290.
Hong, Z., Jiang, J., Lan, L., et al. (2008). A polycomb group protein, PHF1, is involved in the response to DNA double-strand breaks in human cell. Nucleic Acids Research,36, 2939–2947.
Hottiger, M. O. (2015). Nuclear ADP-Ribosylation and its role in chromatin plasticity, cell differentiation, and epigenetics. Annual Review of Biochemistry,84, 227–263.
Huang, C., Cheng, J., Bawa-Khalfe, T., Yao, X., Chin, Y. E., & Yeh, E. T. H. (2016). SUMOylated ORC2 recruits a histone demethylase to regulate centromeric histone modification and genomic stability. Cell Reports,15, 147–157.
Huen, M. S., Grant, R., Manke, I., et al. (2007). RNF8 transduces the DNA-damage signal via histone ubiquitylation and checkpoint protein assembly. Cell,131, 901–914.
Huyen, Y., Zgheib, O., Ditullio, R. A., Jr., et al. (2004). Methylated lysine 79 of histone H3 targets 53BP1 to DNA double-strand breaks. Nature,432, 406–411.
Ikura, M., Furuya, K., Matsuda, S., et al. (2015). Acetylation of histone H2AX at Lys 5 by the TIP60 histone acetyltransferase complex is essential for the dynamic Binding of NBS1 to damaged chromatin. Molecular and Cellular Biology,35, 4147–4157.
Ikura, T., Tashiro, S., Kakino, A., et al. (2007). DNA damage-dependent acetylation and ubiquitination of H2AX enhances chromatin dynamics. Molecular and Cellular Biology,27, 7028–7040.
Jackson, S. P., & Bartek, J. (2009). The DNA-damage response in human biology and disease. Nature,461, 1071–1078.
Jacobs, S. A., & Khorasanizadeh, S. (2002). Structure of HP1 chromodomain bound to a lysine 9-methylated histone H3 tail. Science,295, 2080–2083.
Jazayeri, A., Balestrini, A., Garner, E., Haber, J. E., & Costanzo, V. (2008). Mre11-Rad50-Nbs1-dependent processing of DNA breaks generates oligonucleotides that stimulate ATM activity. EMBO Journal,27, 1953–1962.
Jha, D. K., & Strahl, B. D. (2014). An RNA polymerase II-coupled function for histone H3K36 methylation in checkpoint activation and DSB repair. Nat Commun,5, 3965.
Jing, Y., Liu, Z., Tian, G., Bao, X., Ishibashi, T., & Li, X. D. (2018). Site-specific installation of succinyl lysine analog into histones reveals the effect of H2BK34 succinylation on nucleosome dynamics. Cell Chem Biol,25(166–174), e167.
Johansen, K. M., & Johansen, J. (2006). Regulation of chromatin structure by histone H3S10 phosphorylation. Chromosome Research,14, 393–404.
Jungmichel, S., Rosenthal, F., Altmeyer, M., Lukas, J., Hottiger, M. O., & Nielsen, M. L. (2013). Proteome-wide identification of poly(ADP-Ribosyl)ation targets in different genotoxic stress responses. Molecular Cell,52, 272–285.
Kaidi, A., & Jackson, S. P. (2013). KAT5 tyrosine phosphorylation couples chromatin sensing to ATM signalling. Nature,498, 70–74.
Kanu, N., & Behrens, A. (2007). ATMIN defines an NBS1-independent pathway of ATM signalling. EMBO Journal,26, 2933–2941.
Khoury-Haddad, H., Guttmann-Raviv, N., Ipenberg, I., Huggins, D., Jeyasekharan, A. D., & Ayoub, N. (2014). PARP1-dependent recruitment of KDM4D histone demethylase to DNA damage sites promotes double-strand break repair. Proceedings of the National Academy of Sciences of the United States of America,111, E728–E737.
Kim, Y. C., Gerlitz, G., Furusawa, T., et al. (2009). Activation of ATM depends on chromatin interactions occurring before induction of DNA damage. Nature Cell Biology,11, 92–96.
Koch-Nolte, F., Kernstock, S., Mueller-Dieckmann, C., Weiss, M. S., & Haag, F. (2008). Mammalian ADP-ribosyltransferases and ADP-ribosylhydrolases. Frontiers in Bioscience,13, 6716–6729.
Kolas, N. K., Chapman, J. R., Nakada, S., et al. (2007). Orchestration of the DNA-damage response by the RNF8 ubiquitin ligase. Science,318, 1637–1640.
Kozlov, S. V., Graham, M. E., Jakob, B., et al. (2011). Autophosphorylation and ATM activation: additional sites add to the complexity. Journal of Biological Chemistry,286, 9107–9119.
Kozlov, S. V., Graham, M. E., Peng, C., Chen, P., Robinson, P. J., & Lavin, M. F. (2006). Involvement of novel autophosphorylation sites in ATM activation. EMBO Journal,25, 3504–3514.
Krishnan, V., Chow, M. Z., Wang, Z., et al. (2011). Histone H4 lysine 16 hypoacetylation is associated with defective DNA repair and premature senescence in Zmpste24-deficient mice. Proceedings of the National Academy of Sciences of the United States of America,108, 12325–12330.
Krishnan, N., Jeong, D. G., Jung, S. K., et al. (2009). Dephosphorylation of the C-terminal tyrosyl residue of the DNA damage-related histone H2A.X is mediated by the protein phosphatase eyes absent. Journal of Biological Chemistry,284, 16066–16070.
Kumar, R., Hunt, C. R., Gupta, A., et al. (2011). Purkinje cell-specific males absent on the first (mMof) gene deletion results in an ataxia-telangiectasia-like neurological phenotype and backward walking in mice. Proceedings of the National Academy of Sciences of the United States of America,108, 3636–3641.
Lachner, M., O’Carroll, D., Rea, S., Mechtler, K., & Jenuwein, T. (2001). Methylation of histone H3 lysine 9 creates a binding site for HP1 proteins. Nature,410, 116–120.
Lau, W. C., Li, Y., Liu, Z., Gao, Y., Zhang, Q., & Huen, M. S. (2016). Structure of the human dimeric ATM kinase. Cell Cycle,15, 1117–1124.
Lee, J. H., Choy, M. L., Ngo, L., Foster, S. S., & Marks, P. A. (2010a). Histone deacetylase inhibitor induces DNA damage, which normal but not transformed cells can repair. Proceedings of the National Academy of Sciences of the United States of America,107, 14639–14644.
Lee, J. H., Goodarzi, A. A., Jeggo, P. A., & Paull, T. T. (2010b). 53BP1 promotes ATM activity through direct interactions with the MRN complex. EMBO Journal,29, 574–585.
Lee, C. S., Lee, K., Legube, G., & Haber, J. E. (2014). Dynamics of yeast histone H2A and H2B phosphorylation in response to a double-strand break. Nature Structural and Molecular Biology,21, 103–109.
Lee, J. H., & Paull, T. T. (2004). Direct activation of the ATM protein kinase by the Mre11/Rad50/Nbs1 complex. Science,304, 93–96.
Lee, J. H., & Paull, T. T. (2005). ATM activation by DNA double-strand breaks through the Mre11-Rad50-Nbs1 complex. Science,308, 551–554.
Leidecker, O., Bonfiglio, J. J., Colby, T., et al. (2016). Serine is a new target residue for endogenous ADP-ribosylation on histones. Nature Chemical Biology,12, 998–1000.
Li, T. T., Guan, J. H., Huang, Z. J., Hu, X., & Zheng, X. F. (2014). RNF168-mediated H2A neddylation antagonizes ubiquitylation of H2A and regulates DNA damage repair. Journal of Cell Science,127, 2238–2248.
Li, J., Hart, R. P., Mallimo, E. M., Swerdel, M. R., Kusnecov, A. W., & Herrup, K. (2013). EZH2-mediated H3K27 trimethylation mediates neurodegeneration in ataxia-telangiectasia. Nature Neuroscience,16, 1745–1753.
Li Y, Li Z, Dong L et al. (2018) Histone H1 acetylation at lysine 85 regulates chromatin condensation and genome stability upon DNA damage. Nucleic Acids Res
Li, Z., Li, Y., Tang, M., et al. (2018b). Destabilization of linker histone H1.2 is essential for ATM activation and DNA damage repair. Cell Research ,28, 756–770.
Li, Y., Sabari, B. R., Panchenko, T., et al. (2016a). Molecular coupling of histone crotonylation and active transcription by AF9 YEATS domain. Molecular Cell,62, 181–193.
Li, L., Shi, L., Yang, S., et al. (2016b). SIRT7 is a histone desuccinylase that functionally links to chromatin compaction and genome stability. Nature Communications,7, 12235.
Li, L., & Wang, Y. (2017). Cross-talk between the H3K36me3 and H4K16ac histone epigenetic marks in DNA double-strand break repair. Journal of Biological Chemistry,292, 11951–11959.
Li, Z., & Zhu, W. G. (2014). Targeting histone deacetylases for cancer therapy: from molecular mechanisms to clinical implications. International Journal of Biological Sciences,10, 757–770.
Liokatis, S., Stutzer, A., Elsasser, S. J., et al. (2012). Phosphorylation of histone H3 Ser10 establishes a hierarchy for subsequent intramolecular modification events. Nature Structural and Molecular Biology,19, 819–823.
Lorkovic, Z. J., Park, C., Goiser, M., et al. (2017). Compartmentalization of DNA damage response between heterochromatin and euchromatin is mediated by distinct H2A histone variants. Current Biology,27, 1192–1199.
Lou, Z., Minter-Dykhouse, K., Franco, S., et al. (2006). MDC1 maintains genomic stability by participating in the amplification of ATM-dependent DNA damage signals. Molecular Cell,21, 187–200.
Ma, T., Chen, Y. B., Zhang, F., Yang, C. Y., Wang, S. M., & Yu, X. C. (2013). RNF111-Dependent neddylation activates DNA damage-induced ubiquitination. Molecular Cell,49, 897–907.
Mailand, N., Bekker-Jensen, S., Faustrup, H., et al. (2007). RNF8 ubiquitylates histones at DNA double-strand breaks and promotes assembly of repair proteins. Cell,131, 887–900.
Matsuoka, S., Huang, M., & Elledge, S. J. (1998). Linkage of ATM to cell cycle regulation by the Chk2 protein kinase. Science,282, 1893–1897.
Matsuoka, S., Rotman, G., Ogawa, A., Shiloh, Y., Tamai, K., & Elledge, S. J. (2000). Ataxia telangiectasia-mutated phosphorylates Chk2 in vivo and in vitro. Proceedings of the National Academy of Sciences of the United States of America,97, 10389–10394.
Mattiroli, F., Vissers, J. H., van Dijk, W. J., et al. (2012). RNF168 ubiquitinates K13-15 on H2A/H2AX to drive DNA damage signaling. Cell,150, 1182–1195.
Melander, F., Bekker-Jensen, S., Falck, J., Bartek, J., Mailand, N., & Lukas, J. (2008). Phosphorylation of SDT repeats in the MDC1N terminus triggers retention of NBS1 at the DNA damage-modified chromatin. Journal of Cell Biology,181, 213–226.
Messner, S., & Hottiger, M. O. (2011). Histone ADP-ribosylation in DNA repair, replication and transcription. Trends in Cell Biology,21, 534–542.
Meyer, B., Fabbrizi, M. R., Raj, S., Zobel, C. L., Hallahan, D. E., & Sharma, G. G. (2016). Histone H3 Lysine 9 Acetylation Obstructs ATM Activation and Promotes Ionizing Radiation Sensitivity in Normal Stem Cells. Stem Cell Reports,7, 1013–1022.
Miller, K. M., Tjeertes, J. V., Coates, J., et al. (2010). Human HDAC1 and HDAC2 function in the DNA-damage response to promote DNA nonhomologous end-joining. Nature Structural and Molecular Biology,17, 1144–1151.
Mortusewicz, O., Fouquerel, E., Ame, J. C., Leonhardt, H., & Schreiber, V. (2011). PARG is recruited to DNA damage sites through poly(ADP-ribose)- and PCNA-dependent mechanisms. Nucleic Acids Research,39, 5045–5056.
Moyal, L., Lerenthal, Y., Gana-Weisz, M., et al. (2011). Requirement of ATM-dependent monoubiquitylation of histone H2B for timely repair of DNA double-strand breaks. Molecular Cell,41, 529–542.
Murga, M., Jaco, I., Fan, Y., et al. (2007). Global chromatin compaction limits the strength of the DNA damage response. Journal of Cell Biology,178, 1101–1108.
Musselman, C. A., Avvakumov, N., Watanabe, R., et al. (2012). Molecular basis for H3K36me3 recognition by the Tudor domain of PHF1. Nature Structural and Molecular Biology,19, 1266–1272.
Musselman, C. A., Gibson, M. D., Hartwick, E. W., et al. (2013). Binding of PHF1 Tudor to H3K36me3 enhances nucleosome accessibility. Nat Commun,4, 2969.
Namdar, M., Perez, G., Ngo, L., & Marks, P. A. (2010). Selective inhibition of histone deacetylase 6 (HDAC6) induces DNA damage and sensitizes transformed cells to anticancer agents. Proceedings of the National Academy of Sciences of the United States of America,107, 20003–20008.
Nielsen, P. R., Nietlispach, D., Mott, H. R., et al. (2002). Structure of the HP1 chromodomain bound to histone H3 methylated at lysine 9. Nature,416, 103–107.
Nielsen, S. J., Schneider, R., Bauer, U. M., et al. (2001). Rb targets histone H3 methylation and HP1 to promoters. Nature,412, 561–565.
Nowsheen, S., Aziz, K., Aziz, A., et al. (2018). L3MBTL2 orchestrates ubiquitin signalling by dictating the sequential recruitment of RNF8 and RNF168 after DNA damage. Nature Cell Biology,20, 455–464.
Palazzo L, Leidecker O, Prokhorova E, Dauben H, Matic I, Ahel I. Serine is the major residue for ADP-ribosylation upon DNA damage. Elife 2018; 7.
Pei, H., Zhang, L., Luo, K., et al. (2011). MMSET regulates histone H4K20 methylation and 53BP1 accumulation at DNA damage sites. Nature,470, 124–128.
Pellegrini, M., Celeste, A., Difilippantonio, S., et al. (2006). Autophosphorylation at serine 1987 is dispensable for murine Atm activation in vivo. Nature,443, 222–225.
Peng, J. C., & Karpen, G. H. (2009). Heterochromatic genome stability requires regulators of histone H3 K9 methylation. PLoS Genetics,5, e1000435.
Penicud, K., & Behrens, A. (2014). DMAP1 is an essential regulator of ATM activity and function. Oncogene,33, 525–531.
Pfister, S. X., Ahrabi, S., Zalmas, L. P., et al. (2014). SETD2-dependent histone H3K36 trimethylation is required for homologous recombination repair and genome stability. Cell Reports,7, 2006–2018.
Polo, S. E., & Jackson, S. P. (2011). Dynamics of DNA damage response proteins at DNA breaks: a focus on protein modifications. Genes and Development,25, 409–433.
Ramachandran, S., Haddad, D., Li, C., et al. (2016). The SAGA deubiquitination module promotes DNA repair and class switch recombination through ATM and DNAPK-mediated gammaH2AX formation. Cell Reports,15, 1554–1565.
Robert, T., Vanoli, F., Chiolo, I., et al. (2011). HDACs link the DNA damage response, processing of double-strand breaks and autophagy. Nature,471, 74–79.
Rogakou, E. P., Pilch, D. R., Orr, A. H., Ivanova, V. S., & Bonner, W. M. (1998). DNA double-stranded breaks induce histone H2AX phosphorylation on serine 139. Journal of Biological Chemistry,273, 5858–5868.
Rosenthal, F., & Hottiger, M. O. (2014). Identification of ADP-ribosylated peptides and ADP-ribose acceptor sites. Frontiers in Bioscience (Landmark Ed),19, 1041–1056.
Sabari, B. R., Tang, Z., Huang, H., et al. (2015). Intracellular crotonyl-CoA stimulates transcription through p300-catalyzed histone crotonylation. Molecular Cell,58, 203–215.
Sabari, B. R., Zhang, D., Allis, C. D., & Zhao, Y. (2017). Metabolic regulation of gene expression through histone acylations. Nature Reviews Molecular Cell Biology,18, 90–101.
Sancar, A., Lindsey-Boltz, L. A., Unsal-Kacmaz, K., & Linn, S. (2004). Molecular mechanisms of mammalian DNA repair and the DNA damage checkpoints. Annual Review of Biochemistry,73, 39–85.
Savitsky, K., Bar-Shira, A., Gilad, S., et al. (1995). A single ataxia telangiectasia gene with a product similar to PI-3 kinase. Science,268, 1749–1753.
Sha, M. Q., Zhao, X. L., Li, L., et al. (2016). EZH2 mediates lidamycin-induced cellular senescence through regulating p21 expression in human colon cancer cells. Cell Death Dis,7, e2486.
Sharma, A. K., Bhattacharya, S., Khan, S. A., Khade, B., & Gupta, S. (2015a). Dynamic alteration in H3 serine 10 phosphorylation is G1-phase specific during ionization radiation induced DNA damage response in human cells. Mutation Research,773, 83–91.
Sharma, A. K., Khan, S. A., Sharda, A., Reddy, D. V., & Gupta, S. (2015b). MKP1 phosphatase mediates G1-specific dephosphorylation of H3Serine10P in response to DNA damage. Mutation Research,778, 71–79.
Sharma, G. G., So, S., Gupta, A., et al. (2010). MOF and histone H4 acetylation at lysine 16 are critical for DNA damage response and double-strand break repair. Molecular and Cellular Biology,30, 3582–3595.
Shiloh, Y., & Ziv, Y. (2013). The ATM protein kinase: regulating the cellular response to genotoxic stress, and more. Nature Reviews Molecular Cell Biology,14, 197–210.
Shimada, M., Haruta, M., Niida, H., Sawamoto, K., & Nakanishi, M. (2010). Protein phosphatase 1gamma is responsible for dephosphorylation of histone H3 at Thr 11 after DNA damage. EMBO Reports,11, 883–889.
Shimada, M., Niida, H., Zineldeen, D. H., et al. (2008). Chk1 is a histone H3 threonine 11 kinase that regulates DNA damage-induced transcriptional repression. Cell,132, 221–232.
Shirai, H., Poetsch, A. R., Gunji, A., et al. (2013). PARG dysfunction enhances DNA double strand break formation in S-phase after alkylation DNA damage and augments different cell death pathways. Cell Death Dis,4, e656.
Shogren-Knaak, M., Ishii, H., Sun, J. M., Pazin, M. J., Davie, J. R., & Peterson, C. L. (2006). Histone H4-K16 acetylation controls chromatin structure and protein interactions. Science,311, 844–847.
Shreeram, S., Demidov, O. N., Hee, W. K., et al. (2006). Wip1 phosphatase modulates ATM-dependent signaling pathways. Molecular Cell,23, 757–764.
Singh, N., Basnet, H., Wiltshire, T. D., et al. (2012). Dual recognition of phosphoserine and phosphotyrosine in histone variant H2A.X by DNA damage response protein MCPH1. Proceedings of the National Academy of Sciences of the United States of America,109, 14381–14386.
Sirbu, B. M., & Cortez, D. (2013). DNA damage response: three levels of DNA repair regulation. Cold Spring Harb Perspect Biol,5, a012724.
Spycher, C., Miller, E. S., Townsend, K., et al. (2008). Constitutive phosphorylation of MDC1 physically links the MRE11-RAD50-NBS1 complex to damaged chromatin. Journal of Cell Biology,181, 227–240.
Stewart, G. S., Maser, R. S., Stankovic, T., et al. (1999). The DNA double-strand break repair gene hMRE11 is mutated in individuals with an ataxia-telangiectasia-like disorder. Cell,99, 577–587.
Stucki, M., Clapperton, J. A., Mohammad, D., Yaffe, M. B., Smerdon, S. J., & Jackson, S. P. (2005). MDC1 directly binds phosphorylated histone H2AX to regulate cellular responses to DNA double-strand breaks. Cell,123, 1213–1226.
Stucki, M., & Jackson, S. P. (2006). gammaH2AX and MDC1: anchoring the DNA-damage-response machinery to broken chromosomes. DNA Repair (Amst),5, 534–543.
Sun, Y., Jiang, X., Chen, S., Fernandes, N., & Price, B. D. (2005). A role for the Tip60 histone acetyltransferase in the acetylation and activation of ATM. Proceedings of the National Academy of Sciences of the United States of America,102, 13182–13187.
Sun, Y., Jiang, X., Xu, Y., et al. (2009). Histone H3 methylation links DNA damage detection to activation of the tumour suppressor Tip60. Nature Cell Biology,11, 1376–1382.
Tan, M., Luo, H., Lee, S., et al. (2011). Identification of 67 histone marks and histone lysine crotonylation as a new type of histone modification. Cell,146, 1016–1028.
Tang, M., Li, Z., Zhang, C., et al. (2019). SIRT7-mediated ATM deacetylation is essential for its deactivation and DNA damage repair. Science Advances. https://doi.org/10.1126/sciadv.aav1118.
Thorslund, T., Ripplinger, A., Hoffmann, S., et al. (2015). Histone H1 couples initiation and amplification of ubiquitin signalling after DNA damage. Nature,527, 389–393.
Tian, B., Yang, Q., & Mao, Z. (2009). Phosphorylation of ATM by Cdk5 mediates DNA damage signalling and regulates neuronal death. Nature Cell Biology,11, 211–218.
Tjeertes, J. V., Miller, K. M., & Jackson, S. P. (2009). Screen for DNA-damage-responsive histone modifications identifies H3K9Ac and H3K56Ac in human cells. EMBO Journal,28, 1878–1889.
Toiber, D., Erdel, F., Bouazoune, K., et al. (2013). SIRT6 recruits SNF2H to DNA break sites, preventing genomic instability through chromatin remodeling. Molecular Cell,51, 454–468.
Tripathi, D. N., Chowdhury, R., Trudel, L. J., et al. (2013). Reactive nitrogen species regulate autophagy through ATM-AMPK-TSC2-mediated suppression of mTORC1. Proceedings of the National Academy of Sciences of the United States of America,110, E2950–E2957.
Tsai, W. B., Chung, Y. M., Takahashi, Y., Xu, Z., & Hu, M. C. (2008). Functional interaction between FOXO3a and ATM regulates DNA damage response. Nature Cell Biology,10, 460–467.
Tutton, S., Azzam, G. A., Stong, N., et al. (2016). Subtelomeric p53 binding prevents accumulation of DNA damage at human telomeres. EMBO Journal,35, 193–207.
Uckelmann, M., & Sixma, T. K. (2017). Histone ubiquitination in the DNA damage response. DNA Repair (Amst),56, 92–101.
Uziel, T., Lerenthal, Y., Moyal, L., Andegeko, Y., Mittelman, L., & Shiloh, Y. (2003). Requirement of the MRN complex for ATM activation by DNA damage. EMBO Journal,22, 5612–5621.
Varon, R., Vissinga, C., Platzer, M., et al. (1998). Nibrin, a novel DNA double-strand break repair protein, is mutated in Nijmegen breakage syndrome. Cell,93, 467–476.
Vempati, R. K., Jayani, R. S., Notani, D., Sengupta, A., Galande, S., & Haldar, D. (2010). p300-mediated acetylation of histone H3 lysine 56 functions in DNA damage response in mammals. Journal of Biological Chemistry,285, 28553–28564.
Walker, C., Herranz-Martin, S., Karyka, E., et al. (2017). C9orf72 expansion disrupts ATM-mediated chromosomal break repair. Nature Neuroscience,20, 1225–1235.
Wang, X., Chu, H., Lv, M., et al. (2016a). Structure of the intact ATM/Tel1 kinase. Nature Communications,7, 11655.
Wang, Q., & Goldstein, M. (2016). Small RNAs recruit chromatin-modifying enzymes MMSET and Tip60 to reconfigure damaged DNA upon double-strand break and facilitate repair. Cancer Research,76, 1904–1915.
Wang, Y., Zhang, N., Zhang, L., et al. (2016b). Autophagy regulates chromatin ubiquitination in DNA damage response through elimination of SQSTM1/p62. Molecular Cell,63, 34–48.
Wang, D., Zhou, J., Liu, X., et al. (2013). Methylation of SUV39H1 by SET7/9 results in heterochromatin relaxation and genome instability. Proceedings of the National Academy of Sciences of the United States of America,110, 5516–5521.
Wang, H., Zhou, W., Zheng, Z., et al. (2012). The HDAC inhibitor depsipeptide transactivates the p53/p21 pathway by inducing DNA damage. DNA Repair (Amst),11, 146–156.
Wang, Z., Zhu, W. G., & Xu, X. (2017). Ubiquitin-like modifications in the DNA damage response. Mutation Research,803–805, 56–75.
Wilson, M. D., Benlekbir, S., Fradet-Turcotte, A., et al. (2016). The structural basis of modified nucleosome recognition by 53BP1. Nature,536, 100–103.
Wu, J., Chen, Y., Lu, L. Y., et al. (2011a). Chfr and RNF8 synergistically regulate ATM activation. Nature Structural and Molecular Biology,18, 761–768.
Wu, Z., Lee, S. T., Qiao, Y., et al. (2011b). Polycomb protein EZH2 regulates cancer cell fate decision in response to DNA damage. Cell Death and Differentiation,18, 1771–1779.
Wu, L., Luo, K., Lou, Z., & Chen, J. (2008). MDC1 regulates intra-S-phase checkpoint by targeting NBS1 to DNA double-strand breaks. Proceedings of the National Academy of Sciences of the United States of America,105, 11200–11205.
Wysocki, R., Javaheri, A., Allard, S., Sha, F., Cote, J., & Kron, S. J. (2005). Role of Dot1-dependent histone H3 methylation in G1 and S phase DNA damage checkpoint functions of Rad9. Molecular and Cellular Biology,25, 8430–8443.
Xiao, A., Li, H., Shechter, D., et al. (2009). WSTF regulates the H2AX DNA damage response via a novel tyrosine kinase activity. Nature,457, 57–62.
Xiong, X., Panchenko, T., Yang, S., et al. (2016). Selective recognition of histone crotonylation by double PHD fingers of MOZ and DPF2. Nature Chemical Biology,12, 1111–1118.
Xu, Y., Ashley, T., Brainerd, E. E., Bronson, R. T., Meyn, M. S., & Baltimore, D. (1996). Targeted disruption of ATM leads to growth retardation, chromosomal fragmentation during meiosis, immune defects, and thymic lymphoma. Genes and Development,10, 2411–2422.
Yang, D. Q., & Kastan, M. B. (2000). Participation of ATM in insulin signalling through phosphorylation of eIF-4E-binding protein 1. Nature Cell Biology,2, 893–898.
Young, L. C., McDonald, D. W., & Hendzel, M. J. (2013). Kdm4b histone demethylase is a DNA damage response protein and confers a survival advantage following gamma-irradiation. Journal of Biological Chemistry,288, 21376–21388.
Zhao, D., Guan, H., Zhao, S., et al. (2016). YEATS2 is a selective histone crotonylation reader. Cell Research,26, 629–632.
Zheng, N., & Shabek, N. (2017). Ubiquitin ligases: structure, function, and regulation. Annual Review of Biochemistry,86, 129–157.
Zhou, Y., Lee, J. H., Jiang, W., Crowe, J. L., Zha, S., & Paull, T. T. (2017). Regulation of the DNA damage response by DNA-PKcs inhibitory phosphorylation of ATM. Molecular Cell,65, 91–104.
Ziv, Y., Bielopolski, D., Galanty, Y., et al. (2006). Chromatin relaxation in response to DNA double-strand breaks is modulated by a novel ATM- and KAP-1 dependent pathway. Nature Cell Biology,8, 870–876.
Acknowledgements
This work was supported by the National Key R&D Program of China (Grant number 2017YFA0503900) and the Shenzhen Municipal Commission of Science and Technology Innovation (Grant number JCYJ20160427104855100 and JCYJ20170818092450901). The authors would like to recognize that there are many other valuable papers that could have been included in this review but space limitations prevented them all from being cited.
Author information
Authors and Affiliations
Corresponding author
Ethics declarations
Conflict of interest
The authors declare no competing financial interests.
Rights and permissions
About this article
Cite this article
Li, Z., Chen, Y., Tang, M. et al. Regulation of DNA damage-induced ATM activation by histone modifications. GENOME INSTAB. DIS. 1, 20–33 (2020). https://doi.org/10.1007/s42764-019-00004-8
Received:
Revised:
Accepted:
Published:
Issue Date:
DOI: https://doi.org/10.1007/s42764-019-00004-8