Next Article in Journal
The Vacuolar H+ ATPase α3 Subunit Negatively Regulates Migration and Invasion of Human Pancreatic Ductal Adenocarcinoma Cells
Next Article in Special Issue
3,4-Dihydroxybenzalacetone (DBL) Prevents Aging-Induced Myocardial Changes in Senescence-Accelerated Mouse-Prone 8 (SAMP8) Mice
Previous Article in Journal
Dynamic Genome Editing Using In Vivo Synthesized Donor ssDNA in Escherichia coli
Previous Article in Special Issue
Beyond Tumor Suppression: Senescence in Cancer Stemness and Tumor Dormancy
 
 
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

The Histone Code of Senescence

Department of Medicine, Università degli Studi di Udine. P.le Kolbe 4, 33100 Udine, Italy
*
Author to whom correspondence should be addressed.
Cells 2020, 9(2), 466; https://doi.org/10.3390/cells9020466
Submission received: 2 February 2020 / Revised: 14 February 2020 / Accepted: 17 February 2020 / Published: 18 February 2020

Abstract

:
Senescence is the end point of a complex cellular response that proceeds through a set of highly regulated steps. Initially, the permanent cell-cycle arrest that characterizes senescence is a pro-survival response to irreparable DNA damage. The maintenance of this prolonged condition requires the adaptation of the cells to an unfavorable, demanding and stressful microenvironment. This adaptation is orchestrated through a deep epigenetic resetting. A first wave of epigenetic changes builds a dam on irreparable DNA damage and sustains the pro-survival response and the cell-cycle arrest. Later on, a second wave of epigenetic modifications allows the genomic reorganization to sustain the transcription of pro-inflammatory genes. The balanced epigenetic dynamism of senescent cells influences physiological processes, such as differentiation, embryogenesis and aging, while its alteration leads to cancer, neurodegeneration and premature aging. Here we provide an overview of the most relevant histone modifications, which characterize senescence, aging and the activation of a prolonged DNA damage response.
Keywords:
DNA damage; OIS; RS; SIPS; SAHF; SASP

1. Introduction

Aging is a physiological condition characterized by the functional deficit of tissues and organs due to the accumulation of senescent cells [1]. The key role of senescence in aging is well-established. Clearance of senescent cells in mouse models delays the appearance of age-related tissue and organ disfunctions [2,3]. Senescent cells are characterized by the permanent cell-cycle arrest sustained by the accumulation of cyclin-dependent kinase inhibitors/CDKi), like p16, p21 and p27, as well as by the release of cytokines, chemokines and soluble factors. This modified microenvironment is known as senescence-associated secretory phenotype (SASP) [4]. The senescence state is triggered by different stimuli/stressors. These include the shortening of the telomeres (replicative senescence), the oncogene-induced replication stress, the oncogene-induced senescence (OIS), the accumulation of misfolded protein and/or oxidative stress (stress-induced premature senescence, SIPS) [5].
The impairment of the non-homologous end joining (NHEJ) and homologous recombination (HR) repair mechanisms are common traits of senescent cells [6,7,8,9]. Moreover, a widespread epigenetic resetting characterizes senescent cells and sustains cell-cycle arrest and cellular survival, through the activation of (i) CDKi [10,11], (ii) tumor suppressors [12], and (iii) secretion of chemokines and cytokines, as well as the remodeling of the microenvironment [13].
Macroscopically, senescent cells are characterized by the formation of peculiar areas of heterochromatin, named as SAHF (senescence-associated heterochromatin foci), mainly at E2F loci [14]. However, SAHF do not characterize all senescent cells [15] and are not causally linked to the onset of senescence [16]. Other epigenetic features, like the distension of satellites (senescence-associated distension of satellites, SADS) [17], the re-activation of transposable elements, and of endogenous retroviruses (ERV) [18,19], seem to better qualify different types of senescence. Finally, aging appears to be marked by substantial re-arrangements of the nucleosomes, with the loss of histones H3 and H4 [20,21].
During senescence the epigenome undergoes temporal and sequential modifications that are mandatory to accomplish different cellular adaptations. Initially, this epigenetic resetting is mainly due to the accumulation of irreparable DNA damage. After this first wave of epigenetic modifications, the epigenome is remodeled and fixed in order to sustain the permanent cell-cycle arrest and to modulate the microenvironment.

2. The Epigenome of Replicative Senescence (RS)

The telomeric TTAGGG repeats at chromosome ends protect the genome from degradation and distinguish natural chromosomes ends from double-strand breaks (DSBs) [5,22,23]. Histone and non-histone (Shelterin) proteins sustain the folding of telomeric repeats in high-order chromatin structures that acquire a G-quadruplex shape as a consequence of Hoogsteen base pairing between consecutive guanines [24]. The loss of active telomerase complexes in somatic human cells blocks the lengthening of the telomeric ends. As a consequence, for each successful cell division, telomeres get shorter and cell proliferation is restricted. This phenomenon is defined as replicative senescence (RS) [25]. The accumulation of irreparable DNA damage triggered during RS leads to permanent cell-cycle arrest and is considered among the main driving forces of aging [22].

2.1. Histone Variants

The progressive accumulation of double-strand breaks (DSBs) at the chromosome ends is coupled with a deep epigenetic resetting that can be observed in pre-senescent cells, even distal from telomeres. This epigenetic repertoire builds up an epigenetic clock that dictates the replicative potential of human cells [26]. Late passage IMR90 and WI38 human fibroblasts are characterized by a reduced expression of core histone H3 and H4 [21], of the linker histone H1 [27] and of the histone chaperons ASF1A/B and CAF1-p150/p60 [28]. While the decreased levels of H3 and H4 are due to reduced neosynthesis and increased mRNA degradation [21,29], H1 is post-translationally regulated [27]. Moreover, alternative spliced histone mRNAs belonging to the HIST1 cluster are reported to be accumulated in quiescent and RS-arrested human fibroblasts [30].
The epigenome of RS cells is also characterized by the deposition, at certain genomic loci, of the histone variants H3.3 [31], H2A.J [32] and by the release of genomic DNA from H2A.Z [33,34,35] (Table 1). This redistribution results in chromatin remodeling and promotes the transcription of (i) tumor suppressors [30,31], (ii) inflammatory genes marking the SASP, [32] and iii) the cleavage of H3.3, which mediates the repression of E2F/RB target genes [31]. While in senescence, the HIRA-mediated deposition of H3.3 sustains cell-cycle arrest [31], and in embryonic stem cells ATRX and DAXX recruit H3.3 to repress the transcription of endogenous retroviruses (ERVs) [36].
It is possible that the redistribution of H3.3 between the proliferating and senescent cells, which depends on the detachment from ATRX/DAXX and the complexing to HIRA, is at the basis of the activation of ERVs observed in senescence [37] and aging [38].
The regulated deposition of all these histone variants is necessary [30] and sufficient to sustain cell-cycle arrest [31]. Interestingly, genes encoding these histone variants are frequently mutated in cancer, in confirmation of their tumor-suppressive properties [30,39].

2.2. SAHF an Open Question?

The HIRA chaperone complex (HIRA/UBN1/CABIN1) controls both the deposition of H3.3 [30] and SAHF formation [40]. SAHF accumulation of H3K9me3/H3K27me3/macroH2A blocks on E2F loci characterizes OIS and cells undergoing oncogene-induced replication stress [15]. However, RS is not uniformly characterized by the formation of SAHF [41]. In fact, focused heterochromatinization of E2F loci maintains cell-cycle arrest, also in cells described as SAHF-negative (e.g., BJ and MEFs) [41,42]. SAHF are defined as DAPI-dense nuclear regions characterized by the presence of a central core of condensed chromatin, enriched for H3K9me3 and macroH2A. This core is surrounded by a peripheral ring of H3K27me3 [43,44]. SAHF formation requires p16/INK4 and consists of a deep and focused heterochromatin re-organization [45]. This reorganization is HMGA1/ASF1/HIRA-dependent [40,46] and is triggered by the GSK3β-mediated HIRA re-localization at PML bodies [47]. Even though SAHF dismantling, achieved through HMGA1 [46], ASF1 [40] or GSK3β knockdown [48], allows senescence escape, BJ fibroblasts and Hutchinson–Gilford progeria syndrome (HGPS) cells enter senescence with minimal or no signs of SAHF formation. On the opposite, the SAHF formation in HMEC and MCF10A mammary cells in response to H-RAS/G12V over-expression fails to bring the cells to senescence [15]. Whether SAHF formation is only due to the arising of replication stress and could act as a barrier to DNA double-strand breaks spreading [15], or could mark chromatin regions stitched between remodeled LAD domains [45], needs further investigation. It is possible that a better definition of the SAHF, achieved through the improvement of the resolution of confocal nanoscopy and of ChIP-seq histone mapping, will clarify the debated role played by SAHF in senescence.

2.3. Histone Modifications

A global decrease in H3K9me2/3 and H4K20me but increase in H3K9me1 levels in gene bodies characterize RS in human fibroblasts [49,50]. By contrast, the heterochromatin marker H3K27me3 and the euchromatin marker H3K4me3 are mostly redistributed with respect to proliferating cells (Table 2).
This redistribution correlates well with the expression profile of senescent cells [51,52,53]. A different behavior was reported for the repressive mark H4K20me3 and the activating mark H4K16ac. Although they are enriched in the regulative elements of genes modulated during RS, they do not correlate with gene expression changes observed in senescent cells [30,54]. This apparent paradox could stem from the masking effect imposed by the senescence-specific activation of super-enhancers (marked by H3K27ac/H3K4me1) and the activation of neighbor genes involved in SASP and metabolism [55].
A detailed comparison of H3K4me3 and H3K27me3 levels in proliferating and senescent human fibroblasts evidenced large-scale chromatin modifications during RS [56]. In senescent cells the augmented levels of H3K4me3 and H3K27me3 frequently co-localize in areas defined “mesas” that extend for hundreds of kilobases. Larger domains of RS genome (up to 10Mb) and defined “canyons” are characterized by decreased levels of H3K27me3 [56]. H3K4me3 and H3K27me3 mesas colocalize in LMNB1-associated domains and overlap DNA hypomethylation. Instead, canyons are enriched in gene bodies and enhancers and H3K27me3 loss correlates with the up-regulation of senescent transcriptional programs [56].

2.4. Nuclear Lamins

The loss of LMNB1 typifies all senescence conditions [57] and triggers a deep re-modelling of lamina-associated domains (LADs) [44,56]. LADs re-modelling contributes to re-organizing not only heterochromatin and SAHF [44], but also euchromatin (eLADs) [58]. Moreover, the knockdown of LMNB1 in proliferating cells promotes the premature senescence and gives rise to a H3K4me3/H3K27me3 re-organization similarly to what observed in RS, OIS and HGPS [56]. However, LMNB1 re-expression in RIS senescent cells does not overcome the proliferation of arrest and does not repair nuclear membrane blebbing [59]. It is still unknown if the re-expression of LMNB1 in senescent cells is strong enough to re-establish normal LAD domains or if the co-expression of an epigenetic regulator is needed. Similarly, the ectopic expression of lamina-associated polypeptide 2α restores the proliferation of HGPS cells [60], characterized by the expression of the progerin form of LMNA [61,62]. Unfortunately, the impact of LAP2α expression on LAD domains has not been explored yet. The re-localized LAD domains in RS are characterized also by a general DNA hypomethylation that affects LINE and SINE repetitive elements and pericentromeric satellites [17]. This hypomethylation triggers the general distension of the chromatin associated to these genomic regions (senescence-associated distension of satellites or SADS) and their de-repression [17]. The activation of centromeric and pericentromeric satellite repeats is associated with their exclusion from nucleoli compartments, while all the other nucleoli-associated domains (NADs) are significantly altered in senescent cells [63].
Despite LADs redistribution, 3D chromatin organization of human dermal fibroblasts (HDFs) is only partially altered when proliferating, quiescent and senescent cells are compared. A modest gain of short-range and the loss of long-range intra-chromosome interactions in permanently arrested cells was observed [64]. More evident in quiescent and senescent cells is the switch of topologically associated domains (TADs) from euchromatin areas (Hi-C compartment A) to heterochromatin (Hi-C compartment B) and vice versa. This remodeling reflects the transcriptional status of cell-cycle-associated genes [64]. Most of the heterochromatinization is due to condensin mobilization [65] and PRC2 (EZH2) deposition [66], while MLL1 plays a role in mediating euchromatinization [67,68]. Moreover, RS cells lose the TADs associated to the telomeres [69].

2.5. The CpG Methylation Clock

In general, DNA methylation during aging progressively involve both hypomethylation and hypermethylation events. Importantly, the methylation status of a limited number of well-defined CpG islands associate well with human aging [26]. The quantification of the methylation rate of these CpG island allows a good prediction of human biological age [26,70,71,72]. According to these estimations, the methylome of men ages 4% faster than women [70]. Moreover, the methylation clock is accelerated in patients affected by neurodegenerative diseases [73,74,75], chronic stress and insomnia [76], as well as in two premature aging disorders like Werner’s syndromes and Hutchinson–Gilford Progeria syndrome [76,77], while it is subverted in cancer [78,79].
Aging is characterized by enhancer hypomethylation [41] and this correlates with the loss-of-function of stem cells [80]. Similar changes in the methylation prolife also characterize RS [81], and global hypomethylation characterizes both genomic and mitochondrial DNA [82]. CpG hypermethylated regions are associated with H3K27me3, H3K4me3 and H3K4me1, whereas hypomethylation is observed in the constitutive heterochromatin and lamina-associated domains (LADs) [83].

3. The Epigenome of Oncogene-Induced Senescence (OIS)

The expression of a certain number of oncogenes (K-RAS and H-RAS, BRAFV600E, myrAKT1, STAT5, nuclear HDAC4, N1ICD, ERBB2 and β-catenin) [84,85,86,87,88,89,90,91,92] or ablation of tumor suppressors (PTEN, APC and AXIN) in normal cells triggers a permanent and premature cell-cycle arrest defined as OIS [93,94]. OIS can occur also in the presence of ectopic TERT expression [6]. The most studied model of OIS is the RAS-induced senescence (RIS), obtained by over-expressing oncogenic mutants of RAS in fibroblasts, melanocytes and retinal pigment epithelial cells. The consistency of the RIS model has been validated in mice, after conditional induction of the monoallelic expression of K-RAS/G12V. Mice develop lung adenomas characterized by the accumulation of senescent cells [95]. Similarly, premature senescence blocks the spreading of oncogenic lesions in BRAF/V600E expressing melanocytic nevi [89]. Curiously, RAS fails to induce senescence in immortalized human mammary epithelial cells (HMECs), even after the ectopically expression of p16/INKa [96,97]. This failure has been associated to defects in the TGFβ signaling pathway [97].
It is generally accepted that the premature cell-cycle arrest characterizing OIS is elicited by the accumulation of irreparable DNA damage. In fact, the abrogation of ATM signaling allows senescence escape [6]. It is important to note that the DDR dependence was observed for RAS but its involvement in the case of other oncogenes, such as NOTCH and AKT1, needs further validations [90,98]. As explained above, RIS is characterized by the accumulation of SAHF [14]. However, the oncogenic activation of AKT1 and NOTCH or the knockdown of PTEN trigger OIS without SAHF formation [85,86,98]. In the case of NOTCH, this is due to the repression of HMGA1 [90]. Differently, in the case of AKT1 it was hypothesized a mechanism operating through GSK3β inhibition. In fact GSK3β controls the phosphorylation-dependent HIRA sub-compartmentalization [99]. In this respect, a similar defect in HIRA signaling prevents SAHF formation in senescent murine embryonic and skin fibroblasts expressing RAS, but not to the accumulation of nuclear macroH2A.1 [42]. The deposition of the histone variant macroH2A.1 is not only required for SAHF formation. It also sustains a chromatin re-organization that allows the focused histone acetylation and the expression of the typical SASP cytokines and chemokines [100]. Moreover, the knock-down of macroH2A.1 in H-RAS/V12 IMR90-expressing cells not only blunts SASP, but also decreases the phosphorylation of γH2AX [100].
A common property of OIS cells is the loss of LMNB1 [57]. This causes a deep re-organization of LAD domains [44], similarly to what was observed during RS [56]. As a consequence, OIS is characterized by the re-localization and re-organization of LAD-associated heterochromatin domains [43,101]. This re-organization is sculptured in the distribution of H3K4me3 and H3K27me3 “mesas” and H3K27me3 “canyons”, as described above for RS cells [56]. Interestingly, the expansion of H3K27me3 “canyons” achieved though EZH2 repression sustains OIS by promoting the expression of cytokines [56] and of CDKi [66,102,103].
Similarly to RS, the histone variant H3.3 [31,104] and its Cathepsin L-mediated cleavage product H3.3cs1 [104] are deposited in RIS cells in the regulative elements of RB/E2F target allowing the permanent removal of H3K4me3 [31] and the increase in H3K9me3 levels [14]. Contemporary, the removal of H2A.Z and the de-methylation of H3K27me3 at tumor suppressor loci sustains cell-cycle arrest [33,66]. The inflammatory response is achieved through the deposition of the histone variant H2A.J [32] and the binding of HMGB2, which excludes these loci from SAHF [41]. HMGB2 also allows the H3K4 trimethylation mediated by the methyltransferase MLL1 [53]. Moreover, SASP loci are localized in newly formed super-enhancers that require the binding of BRD4 to promote their transcription [105,106].
Finally, OIS in human fibroblasts is characterized by the spatial rearrangement of pre-existing heterochromatin that give rise to SAHF [45]. Differently, HGPS cells are refractory to SAHF formation [45,107,108] and to heterochromatin focusing [45], probably because the huge alterations in the lamina compartment of these cells prevents the proper heterochromatin 3D-structure organization [109]. A similar displacement of H3K9me3 from LADs is observed in OIS, but it is followed by an increase in local interactions between H3K9me3 domains and sharp heterochromatinization [45]. Curiously and differently from RS and aging, OIS cells do not display any changes in CpG island methylation levels [110].

Different Types of OIS?

While studies are increasingly describing RIS epigenetics, detailed data about the epigenetic modifications in other types of OIS are unavailable. Additional data are desirable since increasing evidences highlight the key roles played by epigenetic regulators in maintaining OIS and counteracting oncogenic transformation [84,111,112,113,114,115,116]. For example, in melanomas the activation of H3K9me3 demethylases (LSD1 and JMJD2C) selectively de-represses E2F target genes, allowing senescence escape and tumorigenesis [117]. This result reinforces the idea that a better investigation of the epigenetic mechanism that sustains the early step of tumorigenesis is desperately needed.

4. The Epigenome of Stress Induced Premature Senescence (SIPS)

SIPS is characterized by the accumulation of ROS (reactive oxygen species), due to mitochondrial disfunctions, ER stress or the exogenous administration of oxidative compounds [118,119,120]. Even though SIPS onset is independent from telomere attrition, ROS accumulation can cause telomere disfunction and fusion in primary cells, thus sustaining cell-cycle arrest [120,121]. Accordingly, murine embryonic fibroblasts (MEFs) cultured in normoxia undergo premature senescence due to the accumulation of ROS-induced DNA damage, while the same cells cultured in hypoxia tend to indefinitely proliferate in virtue of the long telomeres [122].
When human cells are exposed to oxidative stress in vitro, they undergo stochastic transcriptional changes which resemble aged tissue. Recently, it has been demonstrated that oxidative stress contributes not only to aging but also to age-related diseases [123]. Moreover, the accumulation of 8-oxo-7,8-dihydro-2′-deoxyguanosine (8-oxodG) has been observed in the liver of aged mice [124]. Global histone methylations of H3K4, K27 and K9 are increased when BEAS-2B cells are exposed to H2O2, and preincubation with ascorbate reverse these changes [125]. These evidences are confirmed also in other contexts [126]. The general heterochromatinization observed after H2O2 treatment is achieved in two steps. Firstly, by reducing acetylation (H3K9ac, H4K8ac, H4K16ac) [125,127] in a HDAC-dependent manner [127,128]. Secondly, by recruiting histone methyltransferases (HMTs) and inducing H3K27me3, H3K9me3 and H4K20me3 [127,129]. Chromatin condensation is an attempt to preserve the DNA from genotoxicity [130].
The downstream pathways that lead to cell-cycle arrest in cells exposed to oxidative stress imply the up-regulation of CDKi, the DDR response and SASP production, similarly to cells undergoing RS [131,132]. In addition, mitochondrial dysfunctions in IMR90 human fibroblasts lead to the ROS–JNK retrograde signaling pathways, which promote SASP and drive cytoplasmic chromatin fragments (CCFs) [133]. Importantly, the epigenetic homeostasis perturbation, achieved through HMTs or HDACs inhibition [129], is reported to expose cells to endogenous ROS production [134] or to trigger and sustain the senescence induced by the treatment with oxidative compounds [135]. In particular, some ROS generators inhibit PRC2 methyltransferases to allow the focused demethylation and transcriptional activation of CDKi [132]. Similarly, the senescence entry of PAK2 knocked-down MEFs cultured in normoxia is delayed because of the decreased deposition of H3.3 on CDKi loci [136]. At the DNA level, SIPS is characterized by a global DNA hypomethylation that only partially overlaps with the one observed during RS [81].
The altered expression and activation of epigenetic regulators allows cancer cells to escape cell-cycle arrest and to proliferate even in the presence of high levels of oxidative and replication stress, perpetuating the accumulation of DNA damage from generation to generation [137]. In short, histone variants and histone posttranslational modifications taking place during senescence are listed in Table 1 and Table 2.
Table 1. Histone variants that characterize senescence. RS: Replicative senescence; OIS: Oncogene induced senescence; SIPS: Stress induced premature senescence; SASP: Senescence associated secretory phenotype; ↑: Increased expression; ↓: Decreased expression; →: No change; NI: Not investigated.
Table 1. Histone variants that characterize senescence. RS: Replicative senescence; OIS: Oncogene induced senescence; SIPS: Stress induced premature senescence; SASP: Senescence associated secretory phenotype; ↑: Increased expression; ↓: Decreased expression; →: No change; NI: Not investigated.
Histones and Histone VariantsRole during SenescenceChanges during RSChanges during OISChanges during SIPSReferences
H1Chromatin condensation Cells 09 00466 i001 Cells 09 00466 i002NI[27]
H2AChromatin condensationIncreased ratio H2A2/H2A1NINI[15,40]
H3, H4Chromatin condensation Cells 09 00466 i003 Cells 09 00466 i004 Cells 09 00466 i005[21]
H2A.XSensor of DNA damage Cells 09 00466 i006 Cells 09 00466 i007 Cells 09 00466 i008[21]
H2A.JPromote SASP Cells 09 00466 i009 Cells 09 00466 i010 Cells 09 00466 i011[32]
H2A.ZRegulation of CDKi Cells 09 00466 i012 Cells 09 00466 i013NI[33,34]
H3.3, H3.3cs1Gene activation, silencing, chromosome segregation Cells 09 00466 i014 Cells 09 00466 i015 Cells 09 00466 i016[31,36,104]
H3.1Gene activation, silencing, chromosome segregation Cells 09 00466 i017NINI[21]
Table 2. Histone post-translation modifications (PMTs) observed in senescence. RS: Replicative senescence; OIS: oncogene induced senescence; SIPS: Stress induced premature senescence; SASP: Senescence associated secretory phenotype; ↑: Increased expression; ↓: Decreased expression; →: No change; NI: Not investigated.
Table 2. Histone post-translation modifications (PMTs) observed in senescence. RS: Replicative senescence; OIS: oncogene induced senescence; SIPS: Stress induced premature senescence; SASP: Senescence associated secretory phenotype; ↑: Increased expression; ↓: Decreased expression; →: No change; NI: Not investigated.
Histone PMTsEnzymes InvolvedRole during SenescenceChanges during RSChanges during OISChanges during SIPSReferences
H3K9me3KDM4A KDM4BHeterochromatin↓Relocalized in focused area↑Relocalized in focused area Cells 09 00466 i018[43,45,128,129]
H3K27me3KDM6B (JMJD3)Heterochromatin↓Relocalized in focused area↓Relocalized in focused area Cells 09 00466 i019[43,44,45,102,103,127,128,129]
H4K20me3Increased activity of Suv420h2 Heterochromatin gene repression Cells 09 00466 i020 Cells 09 00466 i021 Cells 09 00466 i022[54,127]
H3K4me3KMT2/KDM5D/KDM2BEuchromatin, gene activationRelocalizedRelocalized Cells 09 00466 i023[56,125]
H4K16ac(MOF) Histone acetyltransferaseDNA repair, chromatin compaction Cells 09 00466 i024 Cells 09 00466 i025 Cells 09 00466 i026[125,127]
H3K9acHAT/p300Euchromatin, Cells 09 00466 i027 Cells 09 00466 i028 Cells 09 00466 i029[55,125,127]
H4K8acHAT/p300EuchromatinNINI Cells 09 00466 i030[125,127]

5. The Epigenome at DSBs and during DDR: Early Epigenetic Events in the Senescent Response

Double-strand DNA breaks cause massive epigenetic changes. Immediately, at the damaged sites, the DNA unwraps from the histones and the chromatin undergoes a de-structuration, thus losing compactness [138]. This response is achieved mainly through the chaperone-mediated nucleosomes disassembly [139]. Functionally, it allows the recruitment of proteins involved in DDR. Structurally it causes significant topological alterations in the DNA fiber. These modifications are limited by the subsequent heterochromatinization upstream and downstream from the damage site, which restrains relaxation [140].
The phosphorylation by PIKK kinases (ATM, ATR and DNA-PK) of ser 139 of H2AX (γH2AX) is the widespread histone modification that extends for megabases around a DSBs. γH2AX acts as a platform for the recruitment of MDC1 and 53BP1. This recruitment is sustained by the accumulation at the damaged site of DDR RNAs [141]. The general relaxation of the chromatin is achieved through different mechanisms, which include (i) the RNF8/Ubc13/HUWE1 ubiquitylation of Histone H1 [142,143], (ii) the RNF168 mediated K63-ubiquitylation of H2A/B and H2AX [144], and (iii) the PARylation-dependent, proteasomal degradation of H1.2 [145].
In addition to these huge chromatin remodeling, local histone modifications surrounding the DSBs take place in cells undergoing NHEJ or HR, thus sculpturing an epigenetic pattern specific for each of the two repair mechanisms [138]. The activation of NHEJ is supported by a general chromatin expansion around the DSBs, by the deposition of the histone variants H3.3 [146] and H2AZ [147] and by the monoubiquitylation of H2BK120. This monoubiquitylation promotes the appearance of H3K4me3 and the binding of the SWI/SNF remodeling complex. Moreover, the TIE2-mediated phosphorylation of H4Y51 [148], as well as the RNF20/40-mediated H2BK120ub [149] and the RNF168-mediated H2AK15ub [144] act as platforms to recruit DDR proteins, like ABL1 [148], Ku70/80 [149] and 53BP1 [150,151]. The latter modification and the successful recruitment of 53BP1, which controls end re-sectioning, dictate the choice of the NHEJ pathway in spite of HR [152,153]. H4K20me2 is another anti-resection modification that reinforce 53BP1 binding to the chromatin [154]. However, the co-presence of H2AK15Ub and H4K20me2 in the proximity of the DSBs buffers NHEJ processing by recruiting the HAT Tip60/KAT5, which triggers H2AK15ac and 53BP1 displacement. An activity that favors the establishing of HR [155]. 53BP1 displacement seems to occur during S/G2 transition, as in G1 cells 53BP1 stably occupying the HR sites [156]. The general loss of nucleosome occupancy achieved in proximity to DSBs is counterbalanced by the distal accumulation of H3K36me3 [156]. This PTM can be used as a scaffold for the binding of HDACs, thus ensuring the transcriptional silencing of genomic loci affected by DSBs [157,158].
A different epigenetic environment is associated to the activation of the HR pathway [158]. In this context, macroH2A is found to be more abundant than H2AZ, the acetylation of H2BK120 overcomes ubiquitylation in a SAGA complex-dependent manner [159] and γH2AX is more spread [156]. As explained above, Tip60/KAT5 acetylates H2AK15, thus displacing 53BP1 [155]. Tip60/KAT5 also maintains the acetylation of H4K16, which is required for keeping an open chromatin status [160] at the damaged site. An opposite epigenetic mark, H3K9me2, is required for the BARD1/HP1γ mediated retention of BRCA1 [161] and to allow the loading of the pro-resection factor CtIP [162]. In active genes affected by DSBs, the loss in H3K79me2 and of H4 acetylation levels [156], as well as the recruitment of macroH2A [163] and of repressive protein complexes (HP1,KAP1,SUV39H1,PRDM2,HDACs), seems to compensate for the general loss of histones that characterizes the damaged sites [164]. The spatio-temporal regulation of chromatin remodeling at DSBs is achieved and sustained also by the deposition of other histone variants, like the HIRA-assisted H3.3 and the CAF-1-mediated H3.3 [23].
In summary, from the literature emerges a bleeding/vasoconstriction model in which the chromatin expands and becomes flexible at the damaged sites to accommodate the repair complexes, but heterochromatinization and the creation of a transcriptional repressive environment is required later on to allow the repair before restarting the transcription. The successful prediction of DSBs by looking at the histone positioning and PTMs [165], the emerging roles played by epigenetic regulators in DDR as well as the impact of epigenetic drugs on DDR [166] confirm that the epigenetic status of the chromatin not only identifies sites of genome fragility, but it is also causally linked to the successful repair [23].

6. An Anti-Apoptotic Response Sustains the Survival of Damaged Cells Exposed to Irreparable DNA Damage

Senescent cells display an increased resistance to pro-apoptotic stimuli that is achieved through the up-regulation of the pro-survival gene Bcl2 and Bcl-XL and the down-regulation of the pro-apoptotic genes Bid and Bax [127]. The TP53-dependent up-regulation of the CDKi p21 plays a key role in sustaining the survival of damaged cells [167], while DNMT3a and HDAC1 are recruited on the p21 promoter to switch off its expression during apoptosis [168]. The increase in p21 promotes the cell-cycle arrest allowing the activation of DDR; cellular proliferation is subsequently permanently locked through the activation of INK4 CDKi (mainly p16 and p15) [169]. Many of the anti-apoptotic functions of p21 are achieved through the modulation of TFs (p300, STAT, JUN and TP53) [170] and are sustained by an epigenetic reprogramming [127]. In senescent cells, high levels of H4K20me3 and low levels of H4K16ac keep down the transcription of pro-apoptotic genes, while the increased acetylation of H4K16 characterizes pro-survival loci [127]. The heterochromatinization that surrounds and borders DSBs enhances these pro-survival responses and any relaxation of these structures, obtained for example through HDAC inhibition, triggers apoptosis [6]. Finally, the mTORC1 and PGC-1β dependent metabolic reprogramming [171] observed in senescent cells and in different models of aging [5] ensures energy supply.
Since the epigenetic regulators require cofactors that are produced in a large majority in mitochondria [172,173], any mitochondrial dysfunction can affect both the transcriptional and the metabolic fitness of the cells and lead to senescence even in absence of DNA damage (MiDAS, mitochondrial dysfunction-associated senescence) [171].
Severe and acute stresses induce cell death, while prolonged and mild insults lead to cellular senescence and survival, probably because the cells have the time and a not completely compromised ability to mount the epigenetic, transcriptional and metabolic responses that characterize them [174].

7. Final Considerations: The Link between Senescence, Aging, Epigenetics and DDR

The accumulation of DSBs is a general hallmark of senescence and aging [6]. The main endogenous sources of DSBs are telomere attrition and replicative stress. Replication stress is commonly observed in RS, OIS and aging. In all these conditions cells slow down DNA synthesis and replication fork progression. However, the reduced replication fork speed activates dormant origin to preserve replication timing during replication stress [175]. This adaptive response allows the maintenance of an unaltered replication timing also in cells entering senescence [176]. On the other side it exposes common fragile sites (CFSs), which are genomic loci more prone to breakage after DNA polymerase inhibition, and the accumulation of genomic alterations. CFS alterations are typically observed in pre-neoplastic lesions [177]. Similarly, cells exposed to genotoxic agents (e.g., chemotherapeutic agents, pollutants and toxins) or to oxidative stress activate the DDR that frequently leads to cell-cycle arrest.
Whatever the origin, the accumulation of irreparable DNA damage gives rise to a univocal response characterized by the activation of tumor suppressors and CDKi and by the release of pro-inflammatory cytokines [5].
Global histone loss, as well as the focused deposition of histone variants (H2AX, H2AZ, H2AJ, H3.3 and macroH2A) and the redistribution of H3K4me3, H3K27me3 and H3K36me3 characterize both DDR, DSBs and senescence. The chromatin remodeling observed in different senescence models seems to represent a temporal and spatial evolution of what is observed after a short-time treatment of the cells with DNA damaging agents. Although it is clear that an altered epigenome can expose cells to DSBs [134] and that epigenetic regulators control the fate of damaged cells [15,55,66,115,178,179,180], investigations on the epigenetic inheritance in daughter cells coming from DNA-damaged cells are still in their infancy [177].
Cancer cells appear as forgetful cells that have lost the epigenetic memory of a healthy genome. Aging seems to be predisposed to this memory loss. One of the major challenges of the future regarding the treatment of aging and cancer, will be the identification of the framework of epigenetic changes that can restore this memory.

Author Contributions

H.P., writing—original draft preparation; E.D.G., writing—original draft preparation; C.B., conceptualization, writing—original draft preparation, writing—review and editing, visualization, supervision, project administration, and funding acquisition. All authors have read and agreed to the published version of the manuscript.

Funding

This study was supported by PRIN 2017JL8SRX “Class IIa HDACs as therapeutic targets in human diseases: new roles and new selective inhibitors” and Interreg Italia-Osterreich rITAT1054 EPIC to C.B.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Van Deursen, J.M. The role of senescent cells in ageing. Nature 2014, 509, 439–446. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  2. Baker, D.J.; Wijshake, T.; Tchkonia, T.; LeBrasseur, N.K.; Childs, B.G.; van de Sluis, B.; Kirkland, J.L.; van Deursen, J.M. Clearance of p16Ink4a-positive senescent cells delays ageing-associated disorders. Nature 2011, 479, 232–236. [Google Scholar] [CrossRef] [PubMed]
  3. Baker, D.J.; Childs, B.G.; Durik, M.; Wijers, M.E.; Sieben, C.J.; Zhong, J.A.; Saltness, R.; Jeganathan, K.B.; Verzosa, G.C.; Pezeshki, A.; et al. Naturally occurring p16Ink4a-positive cells shorten healthy lifespan. Nature 2016, 530, 184–189. [Google Scholar] [CrossRef] [Green Version]
  4. He, S.; Sharpless, N.E. Senescence in Health and Disease. Cell 2017, 169, 1000–1011. [Google Scholar] [CrossRef] [PubMed]
  5. Kuilman, T.; Michaloglou, C.; Mooi, W.J.; Peeper, D.S. The essence of senescence. Genes Dev. 2010, 24, 2463–2479. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  6. Di Micco, R.; Fumagalli, M.; Cicalese, A.; Piccinin, S.; Gasparini, P.; Luise, C.; Schurra, C.; Garre’, M.; Giovanni Nuciforo, P.; Bensimon, A.; et al. Oncogene-induced senescence is a DNA damage response triggered by DNA hyper-replication. Nature 2006, 444, 638–642. [Google Scholar] [CrossRef]
  7. D’Adda Di Fagagna, F. Living on a break: Cellular senescence as a DNA-damage response. Nat. Rev. Cancer 2008, 8, 512–522. [Google Scholar] [CrossRef]
  8. Seluanov, A.; Mittelman, D.; Pereira-Smith, O.M.; Wilson, J.H.; Gorbunova, V. DNA end joining be comes less efficient and more error-prone during cellular senescence. Proc. Natl. Acad. Sci. USA 2004, 101, 7624–7629. [Google Scholar] [CrossRef] [Green Version]
  9. Pal, S.; Postnikoff, S.D.; Chavez, M.; Tyler, J.K. Impaired cohesion and homologous recombination during replicative aging in budding yeast. Sci. Adv. 2018, 4. [Google Scholar] [CrossRef] [Green Version]
  10. Zhang, W.; Hu, D.; Ji, W.; Yang, L.; Yang, J.; Yuan, J.; Xuan, A.; Zou, F.; Zhuang, Z. Histone modifications contribute to cellular replicative and hydrogen peroxide-induced premature senescence in human embryonic lung fibroblasts. Free Radic. Res. 2014, 48, 550–559. [Google Scholar] [CrossRef]
  11. Bracken, A.P.; Kleine-kohlbrecher, D.; Dietrich, N.; Pasini, D.; Gargiulo, G.; Beekman, C.; Minucci, S.; Porse, B.T.; Marine, J.; Hansen, K.H.; et al. The Polycomb group proteins bind throughout the. Genes Dev. 2007, 21, 525–530. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  12. Kaneda, A.; Fujita, T.; Anai, M.; Yamamoto, S.; Nagae, G.; Morikawa, M.; Tsuji, S.; Oshima, M.; Miyazono, K.; Aburatani, H. Activation of Bmp2-Smad1 Signal and Its Regulation by Coordinated Alteration of H3K27 Trimethylation in Ras-Induced Senescence. PLoS Genet. 2011, 7, e1002359. [Google Scholar] [CrossRef] [PubMed]
  13. Coppé, J.P.; Patil, C.K.; Rodier, F.; Krtolica, A.; Beauséjour, C.M.; Parrinello, S.; Hodgson, J.G.; Chin, K.; Desprez, P.Y.; Campisi, J. A human-like senescence-associated secretory phenotype is conserved in mouse cells dependent on physiological oxygen. PLoS ONE 2010, 5, e9188. [Google Scholar] [CrossRef] [PubMed]
  14. Narita, M.; Nũnez, S.; Heard, E.; Narita, M.; Lin, A.W.; Hearn, S.A.; Spector, D.L.; Hannon, G.J.; Lowe, S.W. Rb-mediated heterochromatin formation and silencing of E2F target genes during cellular senescence. Cell 2003, 113, 703–716. [Google Scholar] [CrossRef] [Green Version]
  15. Di Micco, R.; Sulli, G.; Dobreva, M.; Liontos, M.; Botrugno, O.A.; Gargiulo, G.; Dal Zuffo, R.; Matti, V.; D’Ario, G.; Montani, E.; et al. Interplay between oncogene-induced DNA damage response and heterochromatin in senescence and cancer. Nat. Cell Biol. 2011, 13, 292–302. [Google Scholar] [CrossRef] [Green Version]
  16. Kosar, M.; Bartkova, J.; Hubackova, S.; Hodny, Z.; Lukas, J.; Bartek, J. Senescence-associated heterochromatin foci are dispensable for cellular senescence, occur in a cell type- And insult-dependent manner, and follow expression of p16ink4a. Cell Cycle 2011, 10, 457–468. [Google Scholar] [CrossRef] [Green Version]
  17. Cruickshanks, H.A.; McBryan, T.; Nelson, D.M.; VanderKraats, N.D.; Shah, P.P.; van Tuyn, J.; Singh Rai, T.; Brock, C.; Donahue, G.; Dunican, D.S.; et al. Senescent cells harbour features of the cancer epigenome. Nat. Cell Biol. 2013, 15, 1495–1506. [Google Scholar] [CrossRef]
  18. De Cecco, M.; Criscione, S.W.; Peckham, E.J.; Hillenmeyer, S.; Hamm, E.A.; Manivannan, J.; Peterson, A.L.; Kreiling, J.A.; Neretti, N.; Sedivy, J.M. Genomes of replicatively senescent cells undergo global epigenetic changes leading to gene silencing and activation of transposable elements. Aging Cell 2013, 12, 247–256. [Google Scholar] [CrossRef]
  19. Chiappinelli, K.B.; Strissel, P.L.; Desrichard, A.; Li, H.; Henke, C.; Akman, B.; Hein, A.; Rote, N.S.; Cope, L.M.; Snyder, A.; et al. Inhibiting DNA Methylation Causes an Interferon Response in Cancer via dsRNA Including Endogenous Retroviruses. Cell 2015, 162, 974–986. [Google Scholar] [CrossRef] [Green Version]
  20. Feser, J.; Truong, D.; Das, C.; Carson, J.J.; Kieft, J.; Harkness, T.; Tyler, J.K. Elevated Histone Expression Promotes Life Span Extension. Mol. Cell 2010, 39, 724–735. [Google Scholar] [CrossRef] [Green Version]
  21. O’Sullivan, R.J.; Kubicek, S.; Schreiber, S.L.; Karlseder, J. Reduced histone biosynthesis and chromatin changes arising from a damage signal at telomeres. Nat. Struct. Mol. Biol. 2010, 17, 1218–1225. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  22. Günes, C.; Rudolph, K.L. The Role of Telomeres in Stem Cells and Cancer. Cell 2013, 152, 390–393. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  23. Dabin, J.; Fortuny, A.; Polo, S.E. Europe PMC Funders Group Epigenome maintenance in response to DNA damage. 2017, 62, 712–727.
  24. Moye, A.L.; Porter, K.C.; Cohen, S.B.; Phan, T.; Zyner, K.G.; Sasaki, N.; Lovrecz, G.O.; Beck, J.L.; Bryan, T.M. Telomeric G-quadruplexes are a substrate and site of localization for human telomerase. Nat. Commun. 2015, 6, 7643. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  25. Ishikawa, F. Portrait of replication stress viewed from telomeres. Cancer Sci. 2013, 104, 790–794. [Google Scholar] [CrossRef] [Green Version]
  26. Wagner, W. The link between epigenetic clocks for aging and senescence. Front. Genet. 2019, 10, 1–6. [Google Scholar] [CrossRef] [Green Version]
  27. Funayama, R.; Saito, M.; Tanobe, H.; Ishikawa, F. Loss of linker histone H1 in cellular senescence. J. Cell Biol. 2006, 175, 869–880. [Google Scholar] [CrossRef] [Green Version]
  28. Volk, A.; Crispino, J.D. The role of the chromatin assembly complex (CAF-1) and its p60 subunit (CHAF1b) in homeostasis and disease. Biochim. Biophys. Acta - Gene Regul. Mech. 2015, 1849, 979–986. [Google Scholar] [CrossRef] [Green Version]
  29. Kaygun, H.; Marzluff, W.F. Translation Termination Is Involved in Histone mRNA Degradation when DNA Replication Is Inhibited. Mol. Cell. Biol. 2005, 25, 6879–6888. [Google Scholar] [CrossRef] [Green Version]
  30. Rai, T.S.; Cole, J.J.; Nelson, D.M.; Dikovskaya, D.; Faller, W.J.; Vizioli, M.G.; Hewitt, R.N.; Anannya, O.; McBryan, T.; Manoharan, I.; et al. HIRA orchestrates a dynamic chromatin landscape in senescence and is required for suppression of neoplasia. Genes Dev. 2014, 28, 2712–2725. [Google Scholar] [CrossRef] [Green Version]
  31. Duarte, L.F.; Young, A.R.J.; Wang, Z.; Wu, H.-A.; Panda, T.; Kou, Y.; Kapoor, A.; Hasson, D.; Mills, N.R.; Ma’ayan, A.; et al. Histone H3.3 and its proteolytically processed form drive a cellular senescence programme. Nat. Commun. 2014, 5, 5210. [Google Scholar] [CrossRef] [Green Version]
  32. Contrepois, K.; Coudereau, C.; Benayoun, B.A.; Schuler, N.; Roux, P.-F.; Bischof, O.; Courbeyrette, R.; Carvalho, C.; Thuret, J.-Y.; Ma, Z.; et al. Histone variant H2A.J accumulates in senescent cells and promotes inflammatory gene expression. Nat. Commun. 2017, 8, 14995. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  33. Gévry, N.; Ho, M.C.; Laflamme, L.; Livingston, D.M.; Gaudreau, L. p21 transcription is regulated by differential localization of histone H2A.Z. Genes Dev. 2007, 21, 1869–1881. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  34. Svotelis, A.; Gévry, N.; Gaudreau, L. Regulation of gene expression and cellular proliferation by histone H2A.Z. Biochem. Cell Biol. 2009, 87, 179–188. [Google Scholar] [CrossRef]
  35. Lazorthes, S.; Vallot, C.; Briois, S.; Aguirrebengoa, M.; Thuret, J.Y.; Laurent, G.S.; Rougeulle, C.; Kapranov, P.; Mann, C.; Trouche, D.; et al. A vlincRNA participates in senescence maintenance by relieving H2AZ-mediated repression at the INK4 locus. Nat. Commun. 2015, 6. [Google Scholar] [CrossRef] [PubMed]
  36. Elsässer, S.J.; Noh, K.-M.; Diaz, N.; Allis, D.; Banaszynski, L.A. Histone H3.3 is required for endogenous retroviral element silencing in embryonic stem cells. Nature 2015, 522, 240–244. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  37. Colombo, A.R.; Elias, H.K.; Ramsingh, G. Senescence induction universally activates transposable element expression. Cell Cycle 2018, 17, 1846–1857. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  38. Cardelli, M. The epigenetic alterations of endogenous retroelements in aging. Mech. Ageing Dev. 2018, 174, 30–46. [Google Scholar] [CrossRef]
  39. Rivera-Casas, C.; Gonzalez-Romero, R.; Cheema, M.S.; Ausió, J.; Eirín-López, J.M. The characterization of macroH2A beyond vertebrates supports an ancestral origin and conserved role for histone variants in chromatin. Epigenetics 2016, 11, 415–425. [Google Scholar] [CrossRef]
  40. Zhang, R.; Poustovoitov, M.V.; Ye, X.; Santos, H.A.; Chen, W.; Daganzo, S.M.; Erzberger, J.P.; Serebriiskii, I.G.; Canutescu, A.A.; Dunbrack, R.L.; et al. Formation of macroH2A-containing senescence-associated heterochromatin foci and senescence driven by ASF1a and HIRA. Dev. Cell 2005, 8, 19–30. [Google Scholar] [CrossRef] [Green Version]
  41. Cole, J.J.; Robertson, N.A.; Rather, M.I.; Thomson, J.P.; McBryan, T.; Sproul, D.; Wang, T.; Brock, C.; Clark, W.; Ideker, T.; et al. Diverse interventions that extend mouse lifespan suppress shared age-associated epigenetic changes at critical gene regulatory regions. Genome Biol. 2017, 18, 58. [Google Scholar] [CrossRef] [Green Version]
  42. Kennedy, A.L.; McBryan, T.; Enders, G.H.; Johnson, F.B.; Zhang, R.; Adams, P.D. Senescent mouse cells fail to overtly regulate the HIRA histone chaperone and do not form robust Senescence Associated Heterochromatin Foci. Cell Div. 2010, 5, 1–11. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  43. Chandra, T.; Kirschner, K.; Thuret, J.; Pope, B.D.; Ryba, T.; Newman, S.; Ahmed, K.; Samarajiwa, S.A.; Salama, R.; Carroll, T.; et al. Independence of Repressive Histone Marks and Chromatin Compaction during Senescent Heterochromatic Layer Formation. Mol. Cell 2012, 47, 203–214. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  44. Sadaie, M.; Salama, R.; Carroll, T.; Tomimatsu, K.; Chandra, T.; Young, A.R.J.; Narita, M.; Pérez-Mancera, P.A.; Bennett, D.C.; Chong, H.; et al. Redistribution of the Lamin B1 genomic binding profile affects rearrangement of heterochromatic doma.pdf. Genes Dev. 2013, 27, 1800–1808. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  45. Chandra, T.; Narita, M. High-order chromatin structure and the epigenome in SAHFs. Nucleus 2013, 4, 23–28. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  46. Narita, M. Cellular senescence and chromatin organisation. Br. J. Cancer 2007, 96, 686–691. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  47. Ye, X.; Zerlanko, B.; Zhang, R.; Somaiah, N.; Lipinski, M.; Salomoni, P.; Adams, P.D. Definition of pRB- and TP53-Dependent and -Independent Steps in HIRA/ASF1a-Mediated Formation of Senescence-Associated Heterochromatin Foci. Mol. Cell. Biol. 2007, 27, 2452–2465. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  48. Liu, S.; Fang, X.; Hall, H.; Yu, S.; Smith, D.; Lu, Z.; Fang, D.; Liu, J.; Stephens, L.C.; Woodgett, J.R.; et al. Homozygous deletion of glycogen synthase kinase 3β bypasses senescence allowing Ras transformation of primary murine fibroblasts. Proc. Natl. Acad. Sci. USA 2008, 105, 5248–5253. [Google Scholar] [CrossRef] [Green Version]
  49. Jørgensen, S.; Schotta, G.; Sørensen, C.S. Histone H4 Lysine 20 methylation: Key player in epigenetic regulation of genomic integrity. Nucleic Acids Res. 2013, 41, 2797–2806. [Google Scholar] [CrossRef]
  50. Sidler, C.; Kovalchuk, O.; Kovalchuk, I. Epigenetic Regulation of Cellular Senescence and Aging. Front. Genet. 2017, 8. [Google Scholar] [CrossRef] [Green Version]
  51. Zhang, W.; Li, J.; Suzuki, K.; Qu, J.; Wang, P.; Zhou, J.; Liu, X.; Ren, R.; Xu, X.; Ocampo, A.; et al. A Werner syndrome stem cell model unveils heterochromatin alterations as a driver of human aging. Science 2015, 348, 1160–1163. [Google Scholar] [CrossRef] [Green Version]
  52. Benayoun, B.A.; Pollina, E.A.; Brunet, A. Epigenetic regulation of ageing: Linking environmental inputs to genomic stability. Nat. Rev. Mol. Cell Biol. 2015, 16, 593–610. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  53. Capell, B.C.; Drake, A.M.; Zhu, J.; Shah, P.P.; Dou, Z.; Dorsey, J.; Simola, D.F.; Donahue, G.; Sammons, M.; Rai, T.S.; et al. Mll1 is essential for the senescenceassociated secretory phenotype. Genes Dev. 2016, 30, 321–336. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  54. Nelson, D.M.; Jaber-Hijazi, F.; Cole, J.J.; Robertson, N.A.; Pawlikowski, J.S.; Norris, K.T.; Criscione, S.W.; Pchelintsev, N.A.; Piscitello, D.; Stong, N.; et al. Mapping H4K20me3 onto the chromatin landscape of senescent cells indicates a function in control of cell senescence and tumor suppression through preservation of genetic and epigenetic stability. Genome Biol. 2016, 17, 1–20. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  55. Sen, P.; Lan, Y.; Li, C.Y.; Sidoli, S.; Donahue, G.; Dou, Z.; Frederick, B.; Chen, Q.; Luense, L.J.; Garcia, B.A.; et al. Histone Acetyltransferase p300 Induces De Novo Super-Enhancers to Drive Cellular Senescence. Mol. Cell 2019, 73, 684–698.e8. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  56. Shah, P.P.; Donahue, G.; Otte, G.L.; Capell, B.C.; Nelson, D.M.; Cao, K.; Aggarwala, V.; Cruickshanks, H.A.; Rai, T.S.; McBryan, T.; et al. Lamin B1 depletion in senescent cells triggers large-scale changes in gene expression and the chromatin landscape. Genes Dev. 2013, 27, 1787–1799. [Google Scholar] [CrossRef] [Green Version]
  57. Freund, A.; Laberge, R.-M.; Demaria, M.; Campisi, J. Lamin B1 loss is a senescence-associated biomarker. Mol. Biol. Cell 2012, 23, 2066–2075. [Google Scholar] [CrossRef]
  58. Pascual-Reguant, L.; Blanco, E.; Galan, S.; Le Dily, F.; Cuartero, Y.; Serra-Bardenys, G.; Di Carlo, V.; Iturbide, A.; Cebrià-Costa, J.P.; Nonell, L.; et al. Lamin B1 mapping reveals the existence of dynamic and functional euchromatin lamin B1 domains. Nat. Commun. 2018, 9, 3420. [Google Scholar] [CrossRef]
  59. Dou, Z.; Xu, C.; Donahue, G.; Shimi, T.; Pan, J.A.; Zhu, J.; Ivanov, A.; Capell, B.C.; Drake, A.M.; Shah, P.P.; et al. Autophagy mediates degradation of nuclear lamina. Nature 2015, 527, 105–109. [Google Scholar] [CrossRef]
  60. Vidak, S.; Kubben, N.; Dechat, T.; Foisner, R. Proliferation of progeria cells is enhanced by lamina-associated polypeptide 2α (LAP2α) through expression of extracellular matrix proteins. Genes Dev. 2015, 29, 2022–2036. [Google Scholar] [CrossRef] [Green Version]
  61. Kubben, N.; Adriaens, M.; Meuleman, W.; Voncken, J.W.; Van Steensel, B.; Misteli, T. Mapping of lamin A- and progerin-interacting genome regions. Chromosoma 2012, 121, 447–464. [Google Scholar] [CrossRef] [Green Version]
  62. Kudlow, B.A.; Kennedy, B.K.; Monnat, R.J. Werner and Hutchinson-Gilford progeria syndromes: Mechanistic basis of human progeroid diseases. Nat. Rev. Mol. Cell Biol. 2007, 8, 394–404. [Google Scholar] [CrossRef] [PubMed]
  63. Dillinger, S.; Straub, T.; Nemeth, A. Nucleolus association of chromosomal domains is largely maintained in cellular senescence despite massive nuclear reorganisation. PLoS ONE 2017, 12, e0178821. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  64. Criscione, S.W.; Teo, Y.V.; Neretti, N. The Chromatin Landscape of Cellular Senescence. Trends Genet. 2016, 32, 751–761. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  65. Iwasaki, O.; Tanizawa, H.; Kim, K.-D.; Kossenkov, A.; Nacarelli, T.; Tashiro, S.; Majumdar, S.; Showe, L.C.; Zhang, R.; Noma, K. Involvement of condensin in cellular senescence through gene regulation and compartmental reorganization. Nat. Commun. 2019, 10, 5688. [Google Scholar] [CrossRef] [Green Version]
  66. Ito, T.; Teo, Y.V.; Evans, S.A.; Neretti, N.; Sedivy, J.M. Regulation of Cellular Senescence by Polycomb Chromatin Modifiers through Distinct DNA Damage- and Histone Methylation-Dependent Pathways. Cell Rep. 2018, 22, 3480–3492. [Google Scholar] [CrossRef] [Green Version]
  67. Katada, S.; Sassone-Corsi, P. The histone methyltransferase MLL1 permits the oscillation of circadian gene expression. Nat. Struct. Mol. Biol. 2010, 17, 1414–1421. [Google Scholar] [CrossRef]
  68. Wang, Y.; Hou, N.; Cheng, X.; Zhang, J.; Tan, X.; Zhang, C.; Tang, Y.; Teng, Y.; Yang, X. Ezh2 Acts as a Tumor Suppressor in Kras-driven Lung Adenocarcinoma. Int. J. Biol. Sci. 2017, 13, 652–659. [Google Scholar] [CrossRef] [Green Version]
  69. Robin, J.D.; Ludlow, A.T.; Batten, K.; Magdinier, F.; Stadler, G.; Wagner, K.R.; Shay, J.W.; Wright, W.E. Telomere position effect: Regulation of gene expression with progressive telomere shortening over long distances. Genes Dev. 2014, 28, 2464–2476. [Google Scholar] [CrossRef] [Green Version]
  70. Hannum, G.; Guinney, J.; Zhao, L.; Zhang, L.; Hughes, G.; Sadda, S.; Klotzle, B.; Bibikova, M.; Fan, J.-B.; Gao, Y.; et al. Genome-wide Methylation Profiles Reveal Quantitative Views of Human Aging Rates. Mol. Cell 2013, 49, 359–367. [Google Scholar] [CrossRef] [Green Version]
  71. Petkovich, D.A.; Podolskiy, D.I.; Lobanov, A.V.; Lee, S.-G.; Miller, R.A.; Gladyshev, V.N. Using DNA Methylation Profiling to Evaluate Biological Age and Longevity Interventions. Cell Metab. 2017, 25, 954–960.e6. [Google Scholar] [CrossRef] [Green Version]
  72. Wang, T.; Tsui, B.; Kreisberg, J.F.; Robertson, N.A.; Gross, A.M.; Yu, M.K.; Carter, H.; Brown-Borg, H.M.; Adams, P.D.; Ideker, T. Epigenetic aging signatures in mice livers are slowed by dwarfism, calorie restriction and rapamycin treatment. Genome Biol. 2017, 18, 1–11. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  73. Levine, M.E.; Lu, A.T.; Bennett, D.A.; Horvath, S. Epigenetic age of the pre-frontal cortex is associated with neuritic plaques, amyloid load, and Alzheimer’s disease related cognitive functioning. Aging (Albany. NY). 2015, 7, 1198–1211. [Google Scholar] [CrossRef] [PubMed]
  74. Horvath, S.; Gurven, M.; Levine, M.E.; Trumble, B.C.; Kaplan, H.; Allayee, H.; Ritz, B.R.; Chen, B.; Lu, A.T.; Rickabaugh, T.M.; et al. An epigenetic clock analysis of race/ethnicity, sex, and coronary heart disease. Genome Biol. 2016, 17, 171. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  75. Raina, R.; Sen, D. Can crosstalk between DOR and PARP reduce oxidative stress mediated neurodegeneration? Neurochem. Int. 2018, 112, 206–218. [Google Scholar] [CrossRef] [PubMed]
  76. Horvath, S. DNA methylation age of human tissues and cell types. Genome Biol. 2013, 14, R115. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  77. Heyn, H.; Moran, S.; Hernando-Herraez, I.; Sayols, S.; Gomez, A.; Sandoval, J.; Monk, D.; Hata, K.; Marques-Bonet, T.; Wang, L.; et al. DNA methylation contributes to natural human variation. Genome Res. 2013, 23, 1363–1372. [Google Scholar] [CrossRef] [Green Version]
  78. Baylin, S.B.; Jones, P.A. A decade of exploring the cancer epigenome—Biological and translational implications. Nat. Rev. Cancer 2011, 11, 726–734. [Google Scholar] [CrossRef]
  79. Mays-Hoopes, L.L. DNA methylation in senescence, aging and cancer. Oncoscience 2019, 44, 291. [Google Scholar]
  80. Neri, F.; Rapelli, S.; Krepelova, A.; Incarnato, D.; Parlato, C.; Basile, G.; Maldotti, M.; Anselmi, F.; Oliviero, S. Intragenic DNA methylation prevents spurious transcription initiation. Nature 2017, 543, 72–77. [Google Scholar] [CrossRef]
  81. Zhang, W.; Ji, W.; Yang, J.; Yang, L.; Chen, W.; Zhuang, Z. Comparison of global DNA methylation profiles in replicative versus premature senescence. Life Sci. 2008, 83, 475–480. [Google Scholar] [CrossRef]
  82. Yu, D.; Du, Z.; Pian, L.; Li, T.; Wen, X.; Li, W.; Kim, S.J.; Xiao, J.; Cohen, P.; Cui, J.; et al. Mitochondrial DNA hypomethylation is a biomarker associated with induced senescence in human fetal heart mesenchymal stem cells. Stem Cells Int. 2017, 2017, 14–16. [Google Scholar] [CrossRef] [PubMed]
  83. Hänzelmann, S.; Beier, F.; Gusmao, E.G.; Koch, C.M.; Hummel, S.; Charapitsa, I.; Joussen, S.; Benes, V.; Brümmendorf, T.H.; Reid, G.; et al. Replicative senescence is associated with nuclear reorganization and with dna methylation at specific transcription factor binding sites. Clin. Epigenetics 2015, 7, 1–19. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  84. Paluvai, H.; Di Giorgio, E.; Brancolini, C. Unscheduled HDAC4 repressive activity in human fibroblasts triggers TTP53-dependent senescence and favors cell transformation. Mol. Oncol. 2018, 12, 2165–2181. [Google Scholar] [CrossRef] [PubMed]
  85. Astle, M.V.; Hannan, K.M.; Ng, P.Y.; Lee, R.S.; George, A.J.; Hsu, A.K.; Haupt, Y.; Hannan, R.D.; Pearson, R.B. AKT induces senescence in human cells via mTORC1 and TP53 in the absence of DNA damage: Implications for targeting mTOR during malignancy. Oncogene 2012, 31, 1949–1962. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  86. Teo, Y.V.; Rattanavirotkul, N.; Olova, N.; Salzano, A.; Quintanilla, A.; Tarrats, N.; Kiourtis, C.; Müller, M.; Green, A.R.; Adams, P.D.; et al. NOTCH Signaling Mediates Secondary Senescence. Cell Rep. 2019, 27, 997–1007.e5. [Google Scholar] [CrossRef] [Green Version]
  87. Mallette, F.A.; Moiseeva, O.; Calabrese, V.; Mao, B.; Gaumont-Leclerc, M.F.; Ferbeyre, G. Transcriptome analysis and tumor suppressor requirements of STAT5-induced senescence. Ann. N. Y. Acad. Sci. 2010, 1197, 142–151. [Google Scholar] [CrossRef]
  88. Serrano, M.; Lin, A.W.; McCurrach, M.E.; Beach, D.; Lowe, S.W. Oncogenic ras Provokes Premature Cell Senescence Associated with Accumulation of TP53 and p16INK4a. Cell 1997, 88, 593–602. [Google Scholar] [CrossRef] [Green Version]
  89. Michaloglou, C.; Vredeveld, L.C.W.; Soengas, M.S.; Denoyelle, C.; Kuilman, T.; Van Der Horst, C.M.A.M.; Majoor, D.M.; Shay, J.W.; Mooi, W.J.; Peeper, D.S. BRAFE600-associated senescence-like cell cycle arrest of human naevi. Nature 2005, 436, 720–724. [Google Scholar] [CrossRef]
  90. Parry, A.J.; Hoare, M.; Bihary, D.; Hänsel-Hertsch, R.; Smith, S.; Tomimatsu, K.; Mannion, E.; Smith, A.; D’Santos, P.; Russell, I.A.; et al. NOTCH-mediated non-cell autonomous regulation of chromatin structure during senescence. Nat. Commun. 2018, 9, 1–15. [Google Scholar] [CrossRef] [Green Version]
  91. Trost, T.M.; Lausch, E.U.; Fees, S.A.; Schmitt, S.; Enklaar, T.; Reutzel, D.; Brixel, L.R.; Schmidtke, P.; Maringer, M.; Schiffer, I.B.; et al. Premature senescence is a primary fail-safe mechanism of ERBB2-driven tumorigenesis in breast carcinoma cells. Cancer Res. 2005, 65, 840–849. [Google Scholar]
  92. Gu, Z.; Tan, W.; Feng, G.; Meng, Y.; Shen, B.; Liu, H.; Cheng, C. Wnt/β-catenin signaling mediates the senescence of bone marrow-mesenchymal stem cells from systemic lupus erythematosus patients through the TP53/p21 pathway. Mol. Cell. Biochem. 2014, 387, 27–37. [Google Scholar] [CrossRef]
  93. Lee, J.J.; Kim, B.C.; Park, M.J.; Lee, Y.S.; Kim, Y.N.; Lee, B.L.; Lee, J.S. PTEN status switches cell fate between premature senescence and apoptosis in glioma exposed to ionizing radiation. Cell Death Differ. 2011, 18, 666–677. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  94. Verschuren, E.W.; Ban, K.H.; Masek, M.A.; Lehman, N.L.; Jackson, P.K. Loss of Emi1-Dependent Anaphase-Promoting Complex/Cyclosome Inhibition Deregulates E2F Target Expression and Elicits DNA Damage-Induced Senescence. Mol. Cell. Biol. 2007, 27, 7955–7965. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  95. Collado, M.; Gil, J.; Efeyan, A.; Guerra, C.; Schuhmacher, A.J.; Barradas, M.; Benguría, A.; Zaballos, A.; Flores, J.M.; Barbacid, M.; et al. Tumour biology: Senescence in premalignant tumours. Nature 2005, 436, 642. [Google Scholar] [CrossRef] [PubMed]
  96. Olsen, C.L.; Gardie, B.; Yaswen, P.; Stampfer, M.R. Raf-1-induced growth arrest in human mammary epithelial cells is p16-independent and is overcome in immortal cells during conversion. Oncogene 2002, 21, 6328–6339. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  97. Cipriano, R.; Kan, C.E.; Graham, J.; Danielpour, D.; Stampfer, M.; Jackson, M.W. TGF-β signaling engages an ATM-CHK2-TP53-independent RAS-induced senescence and prevents malignant transformation in human mammary epithelial cells. Proc. Natl. Acad. Sci. USA 2011, 108, 8668–8673. [Google Scholar] [CrossRef] [Green Version]
  98. Kennedy, A.L.; Morton, J.P.; Manoharan, I.; Nelson, D.M.; Jamieson, N.B.; Pawlikowski, J.S.; McBryan, T.; Doyle, B.; McKay, C.; Oien, K.A.; et al. Activation of the PIK3CA/AKT Pathway Suppresses Senescence Induced by an Activated RAS Oncogene to Promote Tumorigenesis. Mol. Cell 2011, 42, 36–49. [Google Scholar] [CrossRef] [Green Version]
  99. Ye, X.; Zerlanko, B.; Kennedy, A.; Banumathy, G.; Zhang, R.; Adams, P.D. Downregulation of Wnt Signaling Is a Trigger for Formation of Facultative Heterochromatin and Onset of Cell Senescence in Primary Human Cells. Mol. Cell 2007, 27, 183–196. [Google Scholar] [CrossRef] [Green Version]
  100. Chen, H.; Ruiz, P.D.; McKimpson, W.M.; Novikov, L.; Kitsis, R.N.; Gamble, M.J. MacroH2A1 and ATM Play Opposing Roles in Paracrine Senescence and the Senescence-Associated Secretory Phenotype. Mol. Cell 2015, 59, 719–731. [Google Scholar] [CrossRef] [Green Version]
  101. Lenain, C.; De Graaf, C.A.; Pagie, L.; Visser, N.L.; De Haas, M.; De Vries, S.S.; Peric-Hupkes, D.; Van Steensel, B.; Peeper, D.S. Massive reshaping of genome-nuclear lamina interactions during oncogene-induced senescence. Genome Res. 2017, 27, 1634–1644. [Google Scholar] [CrossRef]
  102. Agger, K.; Cloos, P.A.C.; Rudkjaer, L.; Williams, K.; Andersen, G.; Christensen, J.; Helin, K. The H3K27me3 demethylase JMJD3 contributes to the activation of the INK4A-ARF locus in response to oncogene- and stress-induced senescence. Genes Dev. 2009, 23, 1171–1176. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  103. Barradas, M.; Anderton, E.; Acosta, J.C.; De Li, S.; Banito, A.; Rodriguez-Niedenführ, M.; Maertens, G.; Banck, M.; Zhou, M.M.; Walsh, M.J.; et al. Histone demethylase JMJD3 contributes to epigenetic control of INK4a/ARF by oncogenic RAS. Genes Dev. 2009, 23, 1177–1182. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  104. Corpet, A.; Olbrich, T.; Gwerder, M.; Fink, D.; Stucki, M. Dynamics of histone H3.3 deposition in proliferating and senescent cells reveals a DAXX-dependent targeting to PML-NBs important for pericentromeric heterochromatin organization. Cell Cycle 2014, 13, 249–267. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  105. Tasdemir, N.; Banito, A.; Roe, J.-S.; Alonso-Curbelo, D.; Camiolo, M.; Tschaharganeh, D.F.; Huang, C.; Aksoy, O.; Bolden, J.E.; Chen, C.-C.; et al. BRD4 Connects Enhancer Remodeling to Senescence Immune Surveillance. Cancer Discov. 2016, 6, 612–629. [Google Scholar] [CrossRef] [Green Version]
  106. Watanabe, S.; Kawamoto, S.; Ohtani, N.; Hara, E. Impact of senescence-associated secretory phenotype and its potential as a therapeutic target for senescence-associated diseases. Cancer Sci. 2017, 108, 563–569. [Google Scholar] [CrossRef] [Green Version]
  107. Swanson, E.C.; Manning, B.; Zhang, H.; Lawrence, J.B. Higher-order unfolding of satellite heterochromatin is a consistent and early event in cell senescence. J. Cell Biol. 2013, 203, 929–942. [Google Scholar] [CrossRef]
  108. Shumaker, D.K.; Dechat, T.; Kohlmaier, A.; Adam, S.A.; Bozovsky, M.R.; Erdos, M.R.; Eriksson, M.; Goldman, A.E.; Khuon, S.; Collins, F.S.; et al. Mutant nuclear lamin A leads to progressive alterations of epigenetic control in premature aging. Proc. Natl. Acad. Sci. USA 2006, 103, 8703–8708. [Google Scholar] [CrossRef] [Green Version]
  109. McCord, R.P.; Nazario-Toole, A.; Zhang, H.; Chines, P.S.; Zhan, Y.; Erdos, M.R.; Collins, F.S.; Dekker, J.; Cao, K. Correlated alterations in genome organization, histone methylation, and DNA-lamin A/C interactions in Hutchinson-Gilford progeria syndrome. Genome Res. 2013, 23, 260–269. [Google Scholar] [CrossRef] [Green Version]
  110. Xie, W.; Kagiampakis, I.; Pan, L.; Zhang, Y.W.; Murphy, L.; Tao, Y.; Kong, X.; Kang, B.; Xia, L.; Carvalho, F.L.F.; et al. DNA Methylation Patterns Separate Senescence from Transformation Potential and Indicate Cancer Risk. Cancer Cell 2018, 33, 309–321.e5. [Google Scholar] [CrossRef] [Green Version]
  111. Braig, M.; Lee, S.; Loddenkemper, C.; Rudolph, C.; Peters, A.H.F.M.; Schlegelberger, B.; Stein, H.; Dörken, B.; Jenuwein, T.; Schmitt, C.A. Oncogene-induced senescence as an initial barrier in lymphoma development. Nature 2005, 436, 660–665. [Google Scholar] [CrossRef]
  112. Di Giorgio, E.; Franforte, E.; Cefalù, S.; Rossi, S.; Dei Tos, A.P.; Brenca, M.; Polano, M.; Maestro, R.; Paluvai, H.; Picco, R.; et al. The co-existence of transcriptional activator and transcriptional repressor MEF2 complexes influences tumor aggressiveness. PLoS Genet. 2017, 13, e1006752. [Google Scholar] [CrossRef] [PubMed]
  113. Wilting, R.H.; Dannenberg, J.H. Epigenetic mechanisms in tumorigenesis, tumor cell heterogeneity and drug resistance. Drug Resist. Updat. 2012, 15, 21–38. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  114. Di Giorgio, E.; Paluvai, H.; Picco, R.; Brancolini, C. Genetic programs driving oncogenic transformation: Lessons from in vitro models. Int. J. Mol. Sci. 2019, 20. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  115. Di Giorgio, E.; Dalla, E.; Franforte, E.; Paluvai, H.; Minisini, M.; Trevisanut, M.; Picco, R.; Brancolini, C. Different class IIa HDACs repressive complexes regulate specific epigenetic responses related to cell survival in leiomyosarcoma cells. Nucleic Acids Res. 2020, 48, 646–664. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  116. Cutano, V.; Di Giorgio, E.; Minisini, M.; Picco, R.; Dalla, E.; Brancolini, C. HDAC7-mediated control of tumour microenvironment maintains proliferative and stemness competence of human mammary epithelial cells. Mol. Oncol. 2019, 13, 1651–1668. [Google Scholar] [CrossRef] [Green Version]
  117. Yu, Y.; Schleich, K.; Yue, B.; Ji, S.; Lohneis, P.; Kemper, K.; Silvis, M.R.; Qutob, N.; van Rooijen, E.; Werner-Klein, M.; et al. Targeting the Senescence-Overriding Cooperative Activity of Structurally Unrelated H3K9 Demethylases in Melanoma. Cancer Cell 2018, 33, 322–336.e8. [Google Scholar] [CrossRef] [Green Version]
  118. Frippiat, C.; Chen, Q.M.; Zdanov, S.; Magalhaes, J.P.; Remacle, J.; Toussaint, O. Subcytotoxic H2O2 Stress Triggers a Release of Transforming Growth Factor-β1, Which Induces Biomarkers of Cellular Senescence of Human Diploid Fibroblasts. J. Biol. Chem. 2001, 276, 2531–2537. [Google Scholar] [CrossRef] [Green Version]
  119. Duan, J.; Duan, J.; Zhang, Z.; Tong, T. Irreversible cellular senescence induced by prolonged exposure to H 2O2 involves DNA-damage-and-repair genes and telomere shortening. Int. J. Biochem. Cell Biol. 2005, 37, 1407–1420. [Google Scholar] [CrossRef]
  120. Coluzzi, E.; Leone, S.; Sgura, A. Oxidative Stress Induces Telomere Dysfunction and Senescence by Replication Fork Arrest. Cells 2019, 8, 19. [Google Scholar] [CrossRef] [Green Version]
  121. Fumagalli, M.; Rossiello, F.; Mondello, C.; D’Adda Di Fagagna, F. Stable cellular senescence is associated with persistent DDR activation. PLoS ONE 2014, 9, 44–46. [Google Scholar] [CrossRef]
  122. Parrinello, S.; Samper, E.; Krtolica, A.; Goldstein, J.; Melov, S.; Campisi, J. Oxygen sensitivity severely limits the replicative lifespan of murine fibroblasts. Nat. Cell Biol. 2003, 5, 741–747. [Google Scholar] [CrossRef] [PubMed]
  123. Liu, Z.; Zhou, T.; Ziegler, A.C.; Dimitrion, P.; Zuo, L. Oxidative Stress in Neurodegenerative Diseases: From Molecular Mechanisms to Clinical Applications. Oxid. Med. Cell. Longev. 2017, 2017, 2525967. [Google Scholar] [CrossRef] [PubMed]
  124. Mikkelsen, L.; Bialkowski, K.; Risom, L.; Løhr, M.; Loft, S.; Møller, P. Aging and defense against generation of 8-oxo-7,8-dihydro-2′-deoxyguanosine in DNA. Free Radic. Biol. Med. 2009, 47, 608–615. [Google Scholar] [CrossRef] [PubMed]
  125. Niu, Y.; DesMarais, T.L.; Tong, Z.; Yao, Y.; Costa, M. Oxidative stress alters global histone modification and DNA methylation. Free Radic. Biol. Med. 2015, 82, 22–28. [Google Scholar] [CrossRef] [Green Version]
  126. Nishida, N.; Kudo, M.; Nagasaka, T.; Ikai, I.; Goel, A. Characteristic patterns of altered DNA methylation predict emergence of human hepatocellular carcinoma. Hepatology 2012, 56, 994–1003. [Google Scholar] [CrossRef]
  127. Sanders, Y.Y.; Liu, H.; Zhang, X.; Hecker, L.; Bernard, K.; Desai, L.; Liu, G.; Thannickal, V.J. Histone modifications in senescence-associated resistance to apoptosis by oxidative stress. Redox Biol. 2013, 1, 8–16. [Google Scholar] [CrossRef] [Green Version]
  128. Chen, F.; Li, X.; Aquadro, E.; Haigh, S.; Zhou, J.; Stepp, D.W.; Weintraub, N.L.; Barman, S.A.; Fulton, D.J.R. Inhibition of histone deacetylase reduces transcription of NADPH oxidases and ROS production and ameliorates pulmonary arterial hypertension. Free Radic. Biol. Med. 2016, 99, 167–178. [Google Scholar] [CrossRef] [Green Version]
  129. Bosch-Presegué, L.; Raurell-Vila, H.; Marazuela-Duque, A.; Kane-Goldsmith, N.; Valle, A.; Oliver, J.; Serrano, L.; Vaquero, A. Stabilization of Suv39H1 by SirT1 Is Part of Oxidative Stress Response and Ensures Genome Protection. Mol. Cell 2011, 42, 210–223. [Google Scholar] [CrossRef]
  130. Raurell-Vila, H.; Bosch-Presegue, L.; Gonzalez, J.; Kane-Goldsmith, N.; Casal, C.; Brown, J.P.; Marazuela-Duque, A.; Singh, P.B.; Serrano, L.; Vaquero, A. An HP1 isoform-specific feedback mechanism regulates Suv39h1 activity under stress conditions. Epigenetics 2017, 12, 166–175. [Google Scholar] [CrossRef]
  131. Bielak-Zmijewska, A.; Mosieniak, G.; Sikora, E. Is DNA damage indispensable for stress-induced senescence? Mech. Ageing Dev. 2018, 170, 13–21. [Google Scholar] [CrossRef]
  132. Ryu, Y.S.; Kang, K.A.; Piao, M.J.; Ahn, M.J.; Yi, J.M.; Bossis, G.; Hyun, Y.M.; Park, C.O.; Hyun, J.W. Particulate matter-induced senescence of skin keratinocytes involves oxidative stress-dependent epigenetic modifications. Exp. Mol. Med. 2019, 51. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  133. Vizioli, M.G.; Liu, T.; Miller, K.N.; Robertson, N.A.; Gilroy, K.; Lagnado, A.B.; Perez-garcia, A.; Kiourtis, C.; Dasgupta, N.; Lei, X.; et al. Mitochondria-to-nucleus retrograde signaling drives formation of cytoplasmic chromatin and inflammation in senescence. Genes Dev. 2020, 1–18. [Google Scholar] [CrossRef] [PubMed]
  134. Itahana, K.; Zou, Y.; Itahana, Y.; Martinez, J.-L.; Beausejour, C.; Jacobs, J.J.L.; Van Lohuizen, M.; Band, V.; Campisi, J.; Dimri, G.P. Control of the replicative life span of human fibroblasts by p16 and the polycomb protein Bmi-1. Mol. Cell. Biol. 2003, 23, 389–401. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  135. Zheng, H.; Huang, Q.; Huang, S.; Yang, X.; Zhu, T.; Wang, W.; Wang, H.; He, S.; Ji, L.; Wang, Y.; et al. Senescence inducer shikonin ROS-dependently suppressed lung cancer progression. Front. Pharmacol. 2018, 9, 1–15. [Google Scholar] [CrossRef]
  136. Lee, J.S.; Mo, Y.; Gan, H.; Burgess, R.J.; Baker, D.J.; van Deursen, J.M.; Zhang, Z. Pak2 kinase promotes cellular senescence and organismal aging. Proc. Natl. Acad. Sci. USA 2019, 116, 13311–13319. [Google Scholar] [CrossRef] [Green Version]
  137. Cheng, Y.; He, C.; Wang, M.; Ma, X.; Mo, F.; Yang, S.; Han, J.; Wei, X. Targeting epigenetic regulators for cancer therapy: Mechanisms and advances in clinical trials. Signal Transduct. Target. Ther. 2019, 4, 62. [Google Scholar] [CrossRef] [Green Version]
  138. Dantuma, N.P.; Attikum, H. Spatiotemporal regulation of posttranslational modifications in the DNA damage response. EMBO J. 2016, 35, 6–23. [Google Scholar] [CrossRef] [Green Version]
  139. Goldstein, M.; Derheimer, F.A.; Tait-Mulder, J.; Kastan, M.B. Nucleolin mediates nucleosome disruption critical for DNA double-strand break repair. Proc. Natl. Acad. Sci. USA 2013, 110, 16874–16879. [Google Scholar] [CrossRef] [Green Version]
  140. Hinde, E.; Kong, X.; Yokomori, K.; Gratton, E. Chromatin dynamics during DNA repair revealed by pair correlation analysis of molecular flow in the nucleus. Biophys. J. 2014, 107, 55–65. [Google Scholar] [CrossRef] [Green Version]
  141. Francia, S.; Michelini, F.; Saxena, A.; Tang, D.; de Hoon, M.; Anelli, V.; Mione, M.; Carninci, P.; d’Adda di Fagagna, F. Site-specific DICER and DROSHA RNA products control the DNA-damage response. Nature 2012, 488, 231–235. [Google Scholar] [CrossRef]
  142. Thorslund, T.; Ripplinger, A.; Hoffmann, S.; Wild, T.; Uckelmann, M.; Villumsen, B.; Narita, T.; Sixma, T.K.; Choudhary, C.; Bekker-Jensen, S.; et al. Histone H1 couples initiation and amplification of ubiquitin signalling after DNA damage. Nature 2015, 527, 389–393. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  143. Mandemaker, I.K.; Van Cuijk, L.; Janssens, R.C.; Lans, H.; Bezstarosti, K.; Hoeijmakers, J.H.; Demmers, J.A.; Vermeulen, W.; Marteijn, J.A. DNA damage-induced histone H1 ubiquitylation is mediated by HUWE1 and stimulates the RNF8-RNF168 pathway. Sci. Rep. 2017, 7, 1–11. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  144. Mattiroli, F.; Vissers, J.H.A.; Van Dijk, W.J.; Ikpa, P.; Citterio, E.; Vermeulen, W.; Marteijn, J.A.; Sixma, T.K. RNF168 ubiquitinates K13-15 on H2A/H2AX to drive DNA damage signaling. Cell 2012, 150, 1182–1195. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  145. Li, Y.; Li, Z.; Dong, L.; Tang, M.; Zhang, P.; Zhang, C.; Cao, Z.; Zhu, Q.; Chen, Y.; Wang, H.; et al. Histone H1 acetylation at lysine 85 regulates chromatin condensation and genome stability upon DNA damage. Nucleic Acids Res. 2018, 46, 7716–7730. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  146. Luijsterburg, M.S.; de Krijger, I.; Wiegant, W.W.; Shah, R.G.; Smeenk, G.; de Groot, A.J.L.; Pines, A.; Vertegaal, A.C.O.; Jacobs, J.J.L.; Shah, G.M.; et al. PARP1 Links CHD2-Mediated Chromatin Expansion and H3.3 Deposition to DNA Repair by Non-homologous End-Joining. Mol. Cell 2016, 61, 547–562. [Google Scholar] [CrossRef] [Green Version]
  147. Xu, Y.; Ayrapetov, M.K.; Xu, C.; Gursoy-Yuzugullu, O.; Hu, Y.; Price, B.D. Histone H2A.Z Controls a Critical Chromatin Remodeling Step Required for DNA Double-Strand Break Repair. Mol. Cell 2012, 48, 723–733. [Google Scholar] [CrossRef] [Green Version]
  148. Hossain, M.B.; Shifat, R.; Johnson, D.G.; Bedford, M.T.; Gabrusiewicz, K.R.; Cortes-Santiago, N.; Luo, X.; Lu, Z.; Ezhilarasan, R.; Sulman, E.P.; et al. TIE2-mediated tyrosine phosphorylation of H4 regulates DNA damage response by recruiting ABL1. Sci. Adv. 2016, 2, e1501290. [Google Scholar] [CrossRef] [Green Version]
  149. Moyal, L.; Lerenthal, Y.; Gana-Weisz, M.; Mass, G.; So, S.; Wang, S.Y.; Eppink, B.; Chung, Y.M.; Shalev, G.; Shema, E.; et al. Requirement of ATM-Dependent Monoubiquitylation of Histone H2B for Timely Repair of DNA Double-Strand Breaks. Mol. Cell 2011, 41, 529–542. [Google Scholar] [CrossRef]
  150. Fradet-Turcotte, A.; Canny, M.D.; Escribano-Díaz, C.; Orthwein, A.; Leung, C.C.Y.; Huang, H.; Landry, M.-C.; Kitevski-LeBlanc, J.; Noordermeer, S.M.; Sicheri, F.; et al. 53BP1 is a reader of the DNA-damage-induced H2A Lys 15 ubiquitin mark. Nature 2013, 499, 50–54. [Google Scholar] [CrossRef] [Green Version]
  151. Wilson, M.D.; Benlekbir, S.; Fradet-Turcotte, A.; Sherker, A.; Julien, J.P.; McEwan, A.; Noordermeer, S.M.; Sicheri, F.; Rubinstein, J.L.; Durocher, D. The structural basis of modified nucleosome recognition by 53BP1. Nature 2016, 536, 100–103. [Google Scholar] [CrossRef]
  152. Chapman, J.R.; Taylor, M.R.G.; Boulton, S.J. Playing the End Game: DNA Double-Strand Break Repair Pathway Choice. Mol. Cell 2012, 47, 497–510. [Google Scholar] [CrossRef] [Green Version]
  153. Uckelmann, M.; Sixma, T.K. Histone ubiquitination in the DNA damage response. DNA Repair (Amst). 2017, 56, 92–101. [Google Scholar] [CrossRef] [PubMed]
  154. Botuyan, M.V.; Lee, J.; Ward, I.M.; Kim, J.-E.; Thompson, J.R.; Chen, J.; Mer, G. Structural Basis for the Methylation State-Specific Recognition of Histone H4-K20 by 53BP1 and Crb2 in DNA Repair. Cell 2006, 127, 1361–1373. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  155. Tang, J.; Cho, N.W.; Cui, G.; Manion, E.M.; Shanbhag, N.M.; Botuyan, M.V.; Mer, G.; Greenberg, R.A. Acetylation limits 53BP1 association with damaged chromatin to promote homologous recombination. Nat. Struct. Mol. Biol. 2013, 20, 317–325. [Google Scholar] [CrossRef] [PubMed]
  156. Clouaire, T.; Rocher, V.; Lashgari, A.; Arnould, C.; Aguirrebengoa, M.; Biernacka, A.; Skrzypczak, M.; Aymard, F.; Fongang, B.; Dojer, N.; et al. Comprehensive Mapping of Histone Modifications at DNA Double-Strand Breaks Deciphers Repair Pathway Chromatin Signatures. Mol. Cell 2018, 72, 250–262.e6. [Google Scholar] [CrossRef] [Green Version]
  157. Carrozza, M.J.; Li, B.; Florens, L.; Suganuma, T.; Swanson, S.K.; Lee, K.K.; Shia, W.J.; Anderson, S.; Yates, J.; Washburn, M.P.; et al. Histone H3 methylation by Set2 directs deacetylation of coding regions by Rpd3S to suppress spurious intragenic transcription. Cell 2005, 123, 581–592. [Google Scholar] [CrossRef] [Green Version]
  158. Clouaire, T.; Legube, G. A Snapshot on the Cis Chromatin Response to DNA Double-Strand Breaks. Trends Genet. 2019, 35, 330–345. [Google Scholar] [CrossRef] [Green Version]
  159. Morgan, M.T.; Haj-Yahya, M.; Ringel, A.E.; Bandi, P.; Brik, A.; Wolberger, C. Structural basis for histone H2B deubiquitination by the SAGA DUB module. Science 2016, 351, 725–728. [Google Scholar] [CrossRef] [Green Version]
  160. Horikoshi, N.; Sharma, D.; Leonard, F.; Pandita, R.K.; Charaka, V.K.; Hambarde, S.; Horikoshi, N.T.; Gaur Khaitan, P.; Chakraborty, S.; Cote, J.; et al. Pre-existing H4K16ac levels in euchromatin drive DNA repair by homologous recombination in S-phase. Commun. Biol. 2019, 2, 1–12. [Google Scholar] [CrossRef]
  161. Wu, W.; Nishikawa, H.; Fukuda, T.; Vittal, V.; Asano, M.; Miyoshi, Y.; Klevit, R.E.; Ohta, T. Interaction of BARD1 and HP1 Is Required for BRCA1 Retention at Sites of DNA Damage. Cancer Res. 2015, 75, 1311–1321. [Google Scholar] [CrossRef] [Green Version]
  162. Khurana, S.; Kruhlak, M.J.; Kim, J.; Tran, A.D.; Liu, J.; Nyswaner, K.; Shi, L.; Jailwala, P.; Sung, M.-H.; Hakim, O.; et al. A Macrohistone Variant Links Dynamic Chromatin Compaction to BRCA1-Dependent Genome Maintenance. Cell Rep. 2014, 8, 1049–1062. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  163. Burgess, R.C.; Burman, B.; Kruhlak, M.J.; Misteli, T. Activation of DNA Damage Response Signaling by Condensed Chromatin. Cell Rep. 2014, 9, 1703–1717. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  164. Hauer, M.H.; Seeber, A.; Singh, V.; Thierry, R.; Sack, R.; Amitai, A.; Kryzhanovska, M.; Eglinger, J.; Holcman, D.; Owen-Hughes, T.; et al. Histone degradation in response to DNA damage enhances chromatin dynamics and recombination rates. Nat. Struct. Mol. Biol. 2017, 24, 99–107. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  165. Mourad, R.; Ginalski, K.; Legube, G.; Cuvier, O. Predicting double-strand DNA breaks using epigenome marks or DNA at kilobase resolution. Genome Biol. 2018, 19, 1–14. [Google Scholar] [CrossRef] [Green Version]
  166. Krumm, A.; Barckhausen, C.; Kucuk, P.; Tomaszowski, K.H.; Loquai, C.; Fahrer, J.; Kramer, O.H.; Kaina, B.; Roos, W.P. Enhanced histone deacetylase activity in malignant melanoma provokes RAD51 and FANCD2-triggered drug resistance. Cancer Res. 2016, 76, 3067–3077. [Google Scholar] [CrossRef] [Green Version]
  167. Childs, B.G.; Baker, D.J.; Kirkland, J.L.; Campisi, J.; van Deursen, J.M. Senescence and apoptosis: Dueling or complementary cell fates? EMBO Rep. 2014, 15, 1139–1153. [Google Scholar] [CrossRef] [Green Version]
  168. Zhang, Y.; Gao, Y.; Zhang, G.; Huang, S.; Dong, Z.; Kong, C.; Su, D.; Du, J.; Zhu, S.; Liang, Q.; et al. DNMT3a plays a role in switches between doxorubicin-induced senescence and apoptosis of colorectal cancer cells. Int. J. Cancer 2011, 128, 551–561. [Google Scholar] [CrossRef]
  169. Takeuchi, S.; Takahashi, A.; Motoi, N.; Yoshimoto, S.; Tajima, T.; Yamakoshi, K.; Hirao, A.; Yanagi, S.; Fukami, K.; Ishikawa, Y.; et al. Intrinsic cooperation between p16INK4aand p21 Waf1/Cip1in the onset of cellular senescence and tumor suppression in vivo. Cancer Res. 2010, 70, 9381–9390. [Google Scholar] [CrossRef] [Green Version]
  170. Karimian, A.; Ahmadi, Y.; Yousefi, B. Multiple functions of p21 in cell cycle, apoptosis and transcriptional regulation after DNA damage. DNA Repair (Amst). 2016, 42, 63–71. [Google Scholar] [CrossRef]
  171. Correia-Melo, C.; Marques, F.D.; Anderson, R.; Hewitt, G.; Hewitt, R.; Cole, J.; Carroll, B.M.; Miwa, S.; Birch, J.; Merz, A.; et al. Mitochondria are required for pro-ageing features of the senescent phenotype. EMBO J. 2016, 35, 724–742. [Google Scholar] [CrossRef]
  172. Ren, R.; Ocampo, A.; Liu, G.-H.; Izpisua Belmonte, J.C. Regulation of Stem Cell Aging by Metabolism and Epigenetics. Cell Metab. 2017, 26, 460–474. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  173. Ciotti, S.; Iuliano, L.; Cefalù, S.; Comelli, M.; Mavelli, I.; Di Giorgio, E.; Brancolini, C. GSK3β is a key regulator of the ROS-dependent necrotic death induced by the quinone DMNQ. Cell Death Dis. 2020, 11, 2. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  174. Rivera-Mulia, J.C.; Schwerer, H.; Besnard, E.; Desprat, R.; Trevilla-Garcia, C.; Sima, J.; Bensadoun, P.; Zouaoui, A.; Gilbert, D.M.; Lemaitre, J.M. Cellular senescence induces replication stress with almost no affect on DNA replication timing. Cell Cycle 2018, 17, 1667–1681. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  175. Rivera-mulia, J.C.; Dimond, A.; Vera, D.; Trevilla-garcia, C.; Sasaki, T.; Dupont, C.; Gribnau, J.; Fraser, P.; Gilbert, D.M.; Gilbert, D.M.; et al. Allele-specific control of replication timing and genome organization during development DNA replication timing, hybrid mouse ES cells, Hi-C., SNPs in vitro. bioRxiv 2018, 800–811. [Google Scholar]
  176. Tsantoulis, P.K.; Kotsinas, A.; Sfikakis, P.P.; Evangelou, K.; Sideridou, M.; Levy, B.; Mo, L.; Kittas, C.; Wu, X.R.; Papavassiliou, A.G.; et al. Oncogene-induced replication stress preferentially targets common fragile sites in preneoplastic lesions. A genome-wide study. Oncogene 2008, 27, 3256–3264. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  177. Thorsson, V.; Gibbs, D.L.; Brown, S.D.; Wolf, D.; Bortone, D.S.; Ou Yang, T.-H.; Porta-Pardo, E.; Gao, G.F.; Plaisier, C.L.; Eddy, J.A.; et al. The Immune Landscape of Cancer. Immunity 2018, 48, 812–830.e14. [Google Scholar] [CrossRef] [Green Version]
  178. Arora, M.; Moser, J.; Phadke, H.; Basha, A.A.; Spencer, S.L. Endogenous Replication Stress in Mother Cells Leads to Quiescence of Daughter Cells. Cell Rep. 2017, 19, 1351–1364. [Google Scholar] [CrossRef] [Green Version]
  179. Di Giorgio, E.; Hancock, W.W.; Brancolini, C. MEF2 and the tumorigenic process, hic sunt leones. Biochim Biophys Acta Rev Cancer 2018, 1870, 261–273. [Google Scholar] [CrossRef]
  180. Peruzzo, P.; Comelli, M.; Di Giorgio, E.; Franforte, E.; Mavelli, I.; Brancolini, C. Transformation by different oncogenes relies on specific metabolic adaptations. Cell Cycle 2016, 15, 2656–2668. [Google Scholar] [CrossRef] [Green Version]

Share and Cite

MDPI and ACS Style

Paluvai, H.; Di Giorgio, E.; Brancolini, C. The Histone Code of Senescence. Cells 2020, 9, 466. https://doi.org/10.3390/cells9020466

AMA Style

Paluvai H, Di Giorgio E, Brancolini C. The Histone Code of Senescence. Cells. 2020; 9(2):466. https://doi.org/10.3390/cells9020466

Chicago/Turabian Style

Paluvai, Harikrishnareddy, Eros Di Giorgio, and Claudio Brancolini. 2020. "The Histone Code of Senescence" Cells 9, no. 2: 466. https://doi.org/10.3390/cells9020466

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop