Next Article in Journal
Localization and Dynamics of the Methionine Sulfoxide Reductases MsrB1 and MsrB2 in Beech Seeds
Next Article in Special Issue
Transcription Factors as the “Blitzkrieg” of Plant Defense: A Pragmatic View of Nitric Oxide’s Role in Gene Regulation
Previous Article in Journal
Crucial Role of FABP3 in αSyn-Induced Reduction of Septal GABAergic Neurons and Cognitive Decline in Mice
Previous Article in Special Issue
New Omics—Derived Perspectives on Retinal Dystrophies: Could Ion Channels-Encoding or Related Genes Act as Modifier of Pathological Phenotype?
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:

Epigenetics of Aging and Aging-Associated Diseases

Dominik Saul
1,2 and
Robyn Laura Kosinsky
Kogod Center on Aging and Division of Endocrinology, Mayo Clinic, 200 First St SW, Rochester, MN 55905, USA
Department of Trauma, Orthopedics and Reconstructive Surgery, Georg-August-University of Goettingen, 37075 Goettingen, Germany
Division of Gastroenterology and Hepatology, Mayo Clinic, 200 First St SW, Rochester, MN 55905, USA
Author to whom correspondence should be addressed.
Int. J. Mol. Sci. 2021, 22(1), 401;
Submission received: 17 November 2020 / Revised: 19 December 2020 / Accepted: 26 December 2020 / Published: 2 January 2021
(This article belongs to the Collection Feature Papers in Molecular Genetics and Genomics)


Aging represents the multifactorial decline in physiological function of every living organism. Over the past decades, several hallmarks of aging have been defined, including epigenetic deregulation. Indeed, multiple epigenetic events were found altered across different species during aging. Epigenetic changes directly contributing to aging and aging-related diseases include the accumulation of histone variants, changes in chromatin accessibility, loss of histones and heterochromatin, aberrant histone modifications, and deregulated expression/activity of miRNAs. As a consequence, cellular processes are affected, which results in the development or progression of several human pathologies, including cancer, diabetes, osteoporosis, and neurodegenerative disorders. In this review, we focus on epigenetic mechanisms underlying aging-related processes in various species and describe how these deregulations contribute to human diseases.

1. Introduction

Aging is a multifactorial biological process of declining physiological functions increasing the susceptibility to aging-related chronic diseases, such as cancer, metabolic, cardiovascular, musculoskeletal, as well as neurodegenerative diseases [1]. Numerous studies have focused on the decipherment of the hallmarks of aging in order to identify potential therapeutic targets to mitigate the aging process. Hallmarks of aging include stem cell exhaustion, altered intercellular communication, senescence, genomic instability, and epigenetic deregulation [2].
Epigenetics refers to reversible heritable mechanisms, which can affect gene expression without underlying changes in DNA sequences, but rather via chromatin modifications. Eukaryotic chromatin is a highly condensed structure containing repeating structural subunits, the nucleosomes. Each nucleosome consists of a histone octamer assembled of two copies of each histone (H2A, H2B, H3, and H4, as well as histone variants, such as macroH2A, H3.3 and H2A.Z), wrapped around by 147 base pairs of DNA [3,4]. Each core histone possesses histone-fold domains serving for the interaction of the histones and N-terminal histone-tails. These tails can be subjected to post-translational modifications, which frequently affect gene expression. These modifications include, for instance, histone acetylation, methylation, phosphorylation and ubiquitination [5].
Epigenetics is a rapidly evolving research field and there is a profound interest in therapies targeting epigenetic as well as aging-related processes. In this review, we focus on aging-associated epigenetic regulatory mechanisms and highlight their implications in aging-related diseases.

2. Epigenetics of Aging and Aging-Related Diseases

2.1. Epigenetic Changes in Aging

2.1.1. Histone and Heterochromatin Loss

The DNA is organized into complex three-dimensional structures; however, for gene transcription, the DNA sequence has to be accessible to key regulators, such as transcription factors and RNA polymerases. Besides chromatin remodeling, which results in the rearrangement of chromatin structures, the global number of histones defines DNA accessibility [6]. In fact, the loss of histones during cellular aging is one of the key observations from simple eukaryotic models, including yeast, to mice and humans. In a micrococcal nuclease-DNA sequencing (MNase-seq) approach detecting protein-unbound DNA regions in young and old Saccharomyces cerevisiae, a nucleosome loss of approximately 50% was detected. As a consequence, global transcription levels were highly upregulated in aged cells [7]. Similarly, aging human fibroblasts grown in vitro showed a replication-associated reduction in histone biosynthesis and quiescent satellite cells displayed decreased histone expression [8,9].
Reduced synthesis of histones together with changes in chromatin structure (see Section 2.1.4 and Section 2.1.5) leads to a global loss of constitutive heterochromatin, one of the earliest models associated with aging. Heterochromatin loss, the transition from highly condensed to tightly packed chromatin structures, during aging has been observed across many species. As a consequence, modified chromatin architecture, the de-repression of silenced genes and global gene expression changes can occur [10].

2.1.2. Histone Variants

Besides the loss of histones, the exchange of canonical histones (H2A, H2B, H3, and H4) with histone variants was observed in aging organisms. These histone variants display distinct primary sequence and properties compared to canonical histones, thereby regulating gene transcription programs. Various aging-related studies evaluating histone variants in murine, primate and human cells implicate a high enrichment of macroH2A (mH2A), H3.3 and H2A.Z. In general, the incorporation of histone variants into the chromatin can be replication-coupled or replication-independent. The replication-coupled process results in a genome-wide incorporation of new nucleosomes into gaps between pre-existing nucleosomes. In contrast, the replication-independent addition of nucleosomes or subunits occurs locally. Thus, during the replication-independent process, histone variants can replace canonical histones, thereby potentially altering gene expression programs [11,12]. The mH2A isoforms are characterized by the presence of a C-terminal 30 kDa non-histone macro domain [13,14], and were shown to facilitate the activation of transcription factors during differentiation processes [13] and the prevention of the reactivation of pluripotency-associated genes [15]. Notably, human fibroblasts undergoing replicative senescence in vitro as well as several tissues isolated from aging mice and primates displayed an enrichment in mH2A levels [16].
Another example is the H3 variant H3.3, which differs from the canonical form by only four amino acids. It was shown to be incorporated only in a replication-independent manner and to be enriched in transcriptionally active chromatin regions. Recent aging studies in mice revealed that H3.3 accumulates in various tissues during aging and that the canonical isoforms have been almost completely replaced by this histone variant by the age of 18 months [17]. Moreover, H3.3 was linked to aging processes in Caenorhabditis elegans. Here, the deletion of H3.3 resulted in profound transcription changes of longevity-associated genes and in decreased survival [18]. Similar results were found when analyzing postmortem human brains where H3.3 levels gradually increased over the first decade of life. In individuals who were 14 to 72 years old, H3.3 amounts remained stable [19].
Another well-characterized histone variant is H2A.Z, which only shares approximately 60% similarity with the canonical H2A form. There are numerous studies elucidating the function of H2A.Z; however, some findings are contradictory. For instance, H2A.Z was associated with transcriptional activation, transcriptional repression, cell cycle control, DNA replication and DNA damage repair [12]. A recent study uncovered a memory-suppressing function of H2A.Z in mice. While H2A.Z accumulated during aging, active learning resulted in H2A.Z reduction and learning-induced gene expression patterns [20].
A variant of H2AX, which is phosphorylated at the C terminal serine-139 by Ataxia-Telangiectasia-Mutated and Ataxia Telangiectasia and Rad3-related (ATM/ATR), appears during the response to double strand breaks (DSBs). Radiation induced phosphorylation of H2AX in short time ranges, referred to as γH2AX, can be used as biological dosimeter. Together with Senescence-Associated β-galactosidase (SA-β-gal) staining, γH2AX is frequently used to detect senescent cells, highlighting DSBs and telomere shortening [21,22,23].

2.1.3. DNA Methylation

Besides histone methylation, DNA can be directly methylated through the covalent linkage of a methyl group to the fifth position of the cytosine ring to generate 5-methylcytosine (5mC). This modification is mainly present in DNA regions rich in cytosine-phospho-guanine (CpG) dinucleotides. While there is extensive evidence that DNA methylation at promoter regions is associated with gene silencing, the decipherment of the function of gene body methylation is still ongoing [24,25,26]. The repression of transcription due to covalent addition of methyl groups onto the DNA can be mediated by interfering with the site-specific binding of transcription factors or by the recruitment of methyl-CpG-binding domain proteins [27,28]. The transfer of this heritable epigenetic mark is mediated by DNA methyltransferases (DNMTs) including DNMT1, DNMT2, DNMT3A, DNMT3B, and DNMT3L. While DNMT1 has a maintenance function, the de novo establishment of DNA methylation is exerted by DNMT3A and DNMT3B alone or in a complex with DNMT3L [29,30].
The conversion of 5mC to the unmodified state is thought to be mediated in an “active”, enzyme-dependent or in a “passive” demethylation process. The family of Ten Eleven Translocation (TET) proteins, TET1, TET2, and TET3, are able to erase DNA methylation in an “active” stepwise process [31,32]. These factors catalyze the oxidation of 5-methylcytosine (5mC) to the intermediates 5-hydroxymethylcytosine (5hmC), 5-formylcytosine (5fC), and 5-carboxylcytosine (5caC). After the recognition of 5fC and 5caC by the Thymine DNA Glycosylase (TDG), the oxidized cytosine base is excised. Finally, this abasic site will be recognized and replaced by an unmodified cytosine residue by Base Excision Repair (BER). During “passive” DNA demethylation, 5-methylcytosine is diluted in a replication-dependent process during cell division [33,34]. Interestingly, it has been demonstrated that a high abundance of 5hmC represses DNMT1 activity by 60-fold, suggesting a role of TET-mediated induction of “passive” demethylation [35].
While methylation-associated control of gene expression pattern is essential for mammalian development and further cellular processes, it was thought to be dispensable in several organisms such as Caenorhabditis elegans and Drosophila melanogaster [36]. Recent studies describe the methylation of exocyclic NH2 groups at the sixth position of the purine ring in adenines (6 mA) in C. elegans, which is thought to be regulated through the DNA demethylase NMAD-1 and the DNA methyltransferase DAMT1 [37]. In addition, there is emerging knowledge on species- and life cycle-dependent 5mC levels among the genomes of the members of genus Drosophila [38].
While it has been known for several decades that DNA methylation can regulate gene expression patterns, biological consequences are still not fully investigated. Generally, it has been described that CpGs at promoter regions display hypermethylation while other CpGs undergo hypomethylation during aging [39]. Remarkably, two large-scale studies significantly contributed to the understanding of the relevance of DNA methylation pattern during aging. The authors identified 353 and 71 CpG sites, respectively [40,41], which were differentially methylated during aging and, therefore, can serve as reliable age predictors in human tissues. In fact, this “epigenetic clock” displays a robust correlation to age (r = 0.96 and r = 0.91, respectively) with minor deviations from the calendar age of analyzed individuals (3.6 and 4.9 years, respectively). In a comparative study evaluating the robustness of biological age predictors (i.e., epigenetic clock, telomere length, composite biomarker predictors, as well as transcriptome-, proteome- and metabolome-based predictors), the epigenetic clock was suggested to be the most reliable readout. However, further confirmatory studies will be needed to additionally evaluate the predictive values of these biological hallmarks of aging [42].

2.1.4. ATP-Dependent Chromatin Remodeling

The chromatin is organized in a highly compact structure and in order to initiate cellular processes such as transcription, DNA replication, and DNA damage repair, it must be remodeled to enable the accessibility for required factors to the DNA. The reorganization of chromatin structures is facilitated by ATP-dependent chromatin remodeling complexes and results in activation or repression of transcription. These remodelers are multi-subunit complexes containing a highly conserved ATPase subunit which belongs to the superfamily II helicase-related proteins [43]. Based on their ATPase-flanking domains, these complexes are categorized into four major subfamilies: switch/sucrose non-fermentable (SWI/SNF), chromodomain helicase DNA-binding (CHD), INO80, and imitation switch (ISWI). They utilize ATP hydrolysis to disrupt interactions between DNA and histones leading, for instance, to nucleosome sliding/repositioning, nucleosome eviction and histone replacement/incorporation [44]. Despite the high complexity and redundance of several remodeling subunits, the understanding of chromatin remodelers and aging has increased over the last years.
Recent data implicate that the two mutually exclusive catalytic ATPase subunits of the SWI/SNF complex, BRM (SMARCA2), or BRG1 (SMARCA4), are involved in telomere maintenance. BRG1 was identified as a negative modulator of the human telomerase reverse transcriptase (hTERT), an enzyme maintaining telomere ends. It was discovered that BRG1 levels are negatively correlated with hTERT in human cervical cancer cells and that BRG1 knockdown promoted hTERT transcription levels [45]. In a later study by these authors, it was shown in human fibroblasts and cervical cancer cells in vitro that BRM is required for the transcription of the telomere-binding proteins TRF1 and TRF2, which are essential to maintain telomere length and structure. This study provided novel insights into the longevity-associated functions of SWI/SNF by maintaining telomere structure and functionality [46]. In C. elegans, SWI/SNF and DAF16/FOXO co-localize at DAF16/FOXO target promoters to induce transcription of genes associated with stress resistance and longevity [47]. Mutations in LET-418, the C. elegans homolog of CHD3/CHD4, increased lifespan and stress resistance and this phenotype depends on DAF16/FOXO activity [48].
Similar to SWI/SNF, INO80 is essential for murine telomere maintenance and its deletion resulted in cellular proliferative defects and activation of p21-dependent cellular senescence [49].
ISW2, the catalytic component of the ISW2 complex, was described as a regulator of aging in S. cerevisiae since its deletion increased yeast replicative lifespan. By de-repressing stress response genes, the deletion of ISW2 contributed to promoting stress resistance pathways [50]. Subsequent research in C. elegans provided evidence that isw1 gene expression is upregulated in response to multiple stressors changing histone levels. Accordingly, deletion of ISW1 resulted in reduced lifespan in the P0 and F1 generation, suggesting that ISW1 regulates longevity in worms [51].

2.1.5. Histone Modifications

N-terminal tails protruding from the four core histones can be subjected to post-translational modifications. Depending on the histone, residue and type of modification (e.g., methylation, acetylation, phosphorylation and ubiquitination), these histone modifications can activate or repress transcription [52]. Importantly, the imbalance of repressing and activating histone modifications can change gene expression programs and, as a consequence, promote aging-associated transcriptome-wide changes.

Histone Methylation

While DNA methylation is highly associated with transcriptional repression, histone methylation can have both activating and repressing effects on gene expression. Previous studies imply that methylation of H3K4, H3K36, and H3K79 promotes transcription and that H3K9, H3K27, and H4K20 negatively affect transcription levels [53].
In C. elegans, the trimethylation of lysine 4 of histone H3 (H3K4me3) was demonstrated to negatively affect longevity. In fact, the loss of the H3K4me3 methyltransferase SET2 as well as WDR5 and ASH2 promoted survival, while the depletion of the H3K4 demethylase RBR2 showed the opposite effect [54]. This finding was supported in a study in Drosophila where the loss of Lid, the ortholog of RBR2, reduced life span in male flies [55]. Notably, 30% of H3K4me3-occupied genes displayed a substantial aging-related deregulation [56]. In agreement with those studies, H3K4me3 levels declined during aging in yeast and the authors described that its loss was linked to the induction of various aging-associated genes [57].
A survival screen in S. cerevisiae revealed that a deficiency in H3K36 methylation decreased life span. Accordingly, deleting the H3K36me2/3 demethylase RPH1 increased H3K36me3 levels and extended survival in yeast [58]. The decline in H3K36me3 occupancy during aging was confirmed in C. elegans and was associated with a transcriptome-wide age-dependent change of gene expression patterns. When inactivating the methyltransferase MET1 in Drosophila, animals displayed lower H3K36me3 levels as well as reduced lifespan [59]. Recently, the importance of H3K36me2 was demonstrated in aging C. elegans. When deleting the H3K36 dimethyltransferase SET18, lifespan was extended in a DAF16-dependent pattern [60]. Despite their distinct functions, these findings support a role of di- and trimethylation of H3K36 in longevity.
The trimethylation on lysine 27 of histone H3 (H3K27me3) is mediated by the Polycomb Repressive Complex 2 (PRC2) and removed by Ubiquitously Transcribed TPR on X 1 (UTX1). H3K27me3 negatively affects gene transcription and was shown, together with UTX1, to be reduced during aging in C. elegans. By depleting the demethylase UTX1 and thereby increasing H3K27me3 levels, the lifespan was increased [61]. Surprisingly, other studies revealed that increased H3K27me3 abundance was detected during aging in other species. For instance, depleting homologs of PRC2 members in flies resulted in increased longevity [62,63]. Moreover, quiescent mouse muscle stem cells and killifish brain tissues displayed an increase in H3K27me3 during aging [8,64].
The relevance of H3K9me3 was implicated in Drosophila when the deletion of KDM4A, a H3K9me3 demethylase, resulted in impaired wing extension and decreased lifespan in male flies [65]. Accordingly, among the factors downregulated after KDM4A loss, was the male sex-determination gene, as well as several genes associated with longevity [65]. In addition, decreased expression of the H3K9me3 methyltransferase SUV39H1 was found during aging of human and murine hematopoietic stem cells [66].
Besides modifications of histone 3, also the trimethylation of lysine 20 on histone 4 (H4K20me3) has been linked to aging. When kidney and liver samples were collected from rats aged 10, 30, 300, and 450 days, a significant enrichment of H4K20me3 was detected in animals older than 30 days. In contrast, the amounts of mono- and dimethylated H3K20 did not change during aging processes [67]. In addition, it was described that senescent primary human IMR90 fibroblasts accumulated H4K20me3 in vitro [68].

Histone Acetylation

The addition of an acetyl group to the ε-amino group onto a lysine residue of a histone is believed to neutralize the positive charge of the lysine. As a result, the interaction between DNA and histone is weakened, thereby loosening chromatin structures and activating transcription. The addition and removal of acetyl groups is orchestrated by histone acetyltransferases (HATs) and histone deacetylases (HDACs), respectively.
When investigating histone modifications in livers isolated from 6, 15, and 30 month old rats, it was discovered that H3K9ac levels decreased with age [69]. In addition, the function of sirtuin 6 (SIRT6), a H3K9 deacetylase, has been addressed in several studies. In human fibroblasts in vitro, the depletion of SIRT6 resulted in abnormal telomere structures, which were similar to the situation observed in Werner syndrome, a premature aging disorder. The authors postulated that this finding is based on H3K9ac-related changes in chromatin states at telomeres [70]. In addition, SIRT6 appears to severely impact survival by regulating several essential biological functions. For instance, the global deletion of Sirt6 in mice resulted in severe metabolic effects and an overall survival of only four weeks [71]. The observation that Sirt6 deletion resulted in early lethality in mice was supported by another study. Here, it was discovered that inflammatory signaling pathways are targeted by SIRT6. It was shown that SIRT6 represses the expression of Nuclear Factor kappa B (NF-κB) target genes by deacetylating H3K9ac on NF-κB promoters [72]. In addition, lifespan of Sirt6 knockout mice was described to be regulated via the insulin-like growth factor 1 (IGF1) pathway [73]. In agreement with these findings, it was recently demonstrated in a rat model that the overexpression of Sirt6 suppresses senescence as well as apoptosis [74].
After the detection of H3K56ac in yeast [75], it was also discovered in mammals. Interestingly, the mutation of this lysine residue to glutamine (K56Q) or arginine (K56R) was associated with increased levels of spontaneous DNA damage and genotoxic stress. In addition, it was demonstrated that H3K56ac required the presence of the histone chaperone ASF1 and that it is mainly present during the S phase [76]. Besides deacetylating H3K9 as previously mentioned, SIRT6 has been described to deacetylate H3K56ac, thereby controlling DNA damage response and genomic stability [77].
The deacetylase SIRT7 positively regulates HAT1, augmenting the acetylation of H4K12 [78]. The function of SIRT7 lies in preserving the genome integrity and intestinal homeostasis [78]. Interestingly, SIRT7 was described as a regulator of aging since its loss caused aneuploidy and aging phenotypes in mice [78].
In addition, the H4K16 acetylation holds an important role in the regulation of telomere silencing, nucleosome assembly and maintaining chromatin structure [79,80]. The deacetylation of H4K16 is performed by SIRT1, whereas the supplementation of SIRT7 was able to prevent this event. A lower acetylation status was linked to defects in DNA repair and a senescent phenotype in mice [81].
The histone variant H2AX and its phosphorylation on serine 139 (γH2AX) have been tightly connected to aging since they depict an early event in the DNA damage response. Despite appearing in healthy brain areas, the cerebral cortex of senescent mice displayed a strong abundance of γH2AX at double strand breaks and repair sites [82]. In addition, the histone H3 threonine 11 phosphorylation (H3pT11) was described as an indicator for stress and aging in yeast and H3pT11 defective mutants displayed prolonged lifespan [83].

Histone Ubiquitination

Histone ubiquitination constitutes a post-translational modification, the transfer of a small regulatory protein, ubiquitin, to histone core members H2A and H2B. While the H2A ubiquitination mostly inhibits gene expression, H2B ubiquitination enhances transcriptional activity. H2B monoubiquitination is necessary for trimethylation of H3K4 and H3K79 and controls the binding of Cps35 with COMPASS complex, actively regulating transcription [84,85,86]. The age-dependent ubiquitination of H2A was originally found in Drosophila, and verified as evolutionarily conserved in humans [87]. In human glioma cells, the inhibition of the monoubiquitination H2Bub1 induced a cellular senescent phenotype [88].

2.1.6. miRNAs

MicroRNAs (miRNAs), short and noncoding single-stranded RNAs (19-22 nucleotides), have become an emerging aging research interest over the last years. Through sequence-specific binding to their gene targets, miRNAs are able to repress translation or induce mRNA degradation [89,90]. Thereby, they exert a regulatory function in various cellular processes such as proliferation, differentiation and cell death [91]. Nearly 2000 miRNAs have been identified in humans which appear to be involved in the regulation of approximately 60% of all human genes [92,93]. An extensive overview on the individual functions of miRNAs affecting lifespan can be found elsewhere [90].
The most widely used model organism to evaluate the role of miRNAs in aging is C. elegans. One of the first discoveries in this field was that the miRNA lin-4 targeting the transcription factor lin-14 was not only required during the development, but also for aging processes in C. elegans. Lin-4 overexpression or interfering with lin-14 activity extended lifespan in a DAF16/HSF1-dependent manner [94]. These findings were supported by a study in Drosophila where the loss of mir-125 (a homolog of lin-4) reduced lifespan in male flies [95]. However, the function of this miRNA in humans remains unclear.
While there is no to little knowledge regarding miRNA functions in aging across species, a recent study analyzed whole-blood samples from 5000 individuals and identified 127 miRNAs expressed in an age-related pattern (r = 0.70) [96]. In addition, several miRNAs have been implicated during the development or progression of aging-associated diseases as explained in Section 2.2.

2.2. Epigenetic Changes in Aging-Related Diseases

Epigenetic discoveries helped to lay the foundation for a deeper perception of multiple diseases. For instance, epigenetic events can contribute to the “hallmarks of cancer” (i.e., chromatin structure affecting cellular identity or methylation patterns leading to evasion from apoptosis), having led to a revised “hallmark” definition [97,98]. Here, we summarize the latest epigenetic discoveries in a selected range of medical conditions with a focus on certain cancer entities, inflammation, musculoskeletal disorders, neurodegenerative diseases, and nutritional diseases.

2.2.1. Cancer

DNA methylation patterns and miRNAs can influence chromatin state regulation [99,100]. In cancer development, the degenerated cell may unrestrictedly proliferate as a consequence of DNA hypermethylation or deregulation of epigenetic modifiers. Methylation of the promotor region of the tumor suppressor genes, for instance VHL, was associated with angiogenesis, leading to an enhanced supply for the tumor environment. Cell death was shown to be impaired by epigenetic modification of apoptotic or cell cycle key players including CDKN2A, hypermethylation of which leads to a loss-of-function gene in numerous cancers [99,100,101]. Here, we focus on two entities to exemplarily demonstrate age-related shifts and deregulations associated with leukemia and colorectal cancer.


Distinct chromatin states can favor oncogene activation driving cells to the hallmarks of cancer [97,100]. For instance, a mutation or generated fusion-protein in the histone methyltransferase MLL or the histone acetyltransferase p300 can block regulatory regions in leukemia, driving malignant transformation [100,101,102,103,104,105,106,107,108]. The pivotal role of p300 in aging was demonstrated by Sen et al., who used 3000 shRNAs to silence well-known epigenetic proteins (n = 600) in order to identify candidates delaying replicative senescence. Upon treatment with shRNA directed against p300, the replicative lifespan of fibroblasts was substantially increased, accompanied by a reduced occurrence of telomere dysfunction-induced foci (TIFs) [109]. While the downregulation of p300 may delay cellular senescence, the downside of this targeting approach is that it appears to promote cancer. The importance of p300 for leukemogenesis was demonstrated by Cheng et al. The authors used a bone marrow transplantation mouse model for acute leukemia (donors: NHD13 p300flox/flox) in which the deletion of p300 led to reduced survival after acceleration of leukemogenesis. An enhanced self-renewal of hematopoietic stem/progenitor cells, combined with a decreased apoptosis rate and upregulation of cytokine receptor genes in p300-deficient cells drove leukemogenesis, identifying p300 as tumor suppressor [110]. In contrast, in a study where the suppression of p300 has been induced pharmacologically, the clonogenic growth of human leukemic cells was impaired [111]. Thus, these effects of p300 suggest a beneficial effect of reducing p300 activity during aging by enhancing cellular lifespan. However, depending of the biological context, decreasing p300 levels and activity may promote tumor development and progression due to its potential role as a tumor suppressor.
Similarly, a dual function was described for the histone methyltransferase and Polycomb Repressive Complex 2 component EZH2, which has been proposed both as a tumor-suppressor and oncogene [112]. A gain-of-function EZH2 mutation characterized by an aberrant H3K27me3 occupancy blocked B cell development and was found in several lymphomas [112,113], whereas a loss-of-function of EZH2 is common in myelodysplastic syndromes [114]. Comparing human and murine methylation data sets, it was demonstrated that the pattern of overall DNA methylation with age is highly conserved between both species and that CpG islands, which are hypermethylated in an age-dependent manner associate with EZH2 [115].
Leukemogenesis has been found to be linked to further aberrant fusion proteins comprising RUNX1, RARα and CBFB [100,116]. In general, a feature of leukemia is the profound increase in H3K27me3 which can be attributed to GSK126, an inhibitor of EZH2 [113]. Subsequently, EZH2 inhibitors were discussed as therapeutic options in AML patients [117].

Colorectal Cancer

As one of the leading causes of cancer-related deaths worldwide, colorectal cancer can be driven by both genetic and epigenetic alterations within epithelial cells. An overall enhanced DNA methylation pattern has been found not only in inflammatory but also cancerous epithelial tissue [118,119]. In fact, this increase in methylation was associated with higher DNA methyltransferase 1 levels. Guo et al. characterized human colon cancer cell lines (SW480, LoVo, and HT29) as well as publicly available data based on colorectal adenocarcinoma samples to demonstrate a high DNMT1 expression levels, accompanied by DNA hypermethylation. Interestingly, the Wnt signaling pathway was activated in these samples, indicating a positive regulation effect of Wnt signaling members on DNMT1 expression while Wnt signaling inhibition downregulated DNMT1 [120]. The strong association between Wnt signaling and DNMT1 has been further elucidated using mass spectrometry-based proteomics. In colorectal cancer cells, a direct protein-protein interaction between β-catenin, the central effector of canonical Wnt signaling and DNMT1 was identified. DNMT reduction via siRNA reduced β-catenin signaling by decreased methylation at several CpG loci confirming the tight interdependence between the two tumor-promoting factors DNMT1 and β-catenin [121]. Besides epithelial cells, the function of peripheral blood mononuclear cells (PBMCs) was evaluated in colorectal cancer. It was demonstrated in PBMCs from 2453 European blood samples that DMNT1 levels gradually declined with aging until the age of 64 years [122,123,124,125]. Since the risk for colorectal cancer increases with aging, an in-depth analysis of the effect of DNMT1 reduction appears to be necessary. Surprisingly, Yung et al. showed an increased DNA methylation pattern via SssI methylase assays in several tissues along with reduced signs of senescence in heterozygous Dnmt null mice. In contrast, Laird et al. reduced the DNMT1 activity genetically and pharmacologically, resulting in a colonic hypomethylation and reduced number of intestinal neoplasia in ApcMin mice [126,127]. Deductively, the role of DNMT1 in the aging intestine is still ambiguous and needs further in-depth characterization.
Colorectal inflammation, as one of the major risk factors of colorectal cancer, was found to be tightly connected to p16INK4a methylation: Wang et al. detected hypermethylation in the promoter region of p16 in human colonoscopic biopsies of rectal inflammatory mucosa, while Hsieh and colleagues suggested the hypermethylation of the p16INK4a promoter region to occur early during the neoplastic progression of ulcerative colitis in colectomy specimens [122,124]. Consequently, attempts have been made to identify molecular subgroups according to their methylation status in distinct gene sets. Accordingly, a CpG island methylator phenotype (CIMP) was identified for diagnostic and therapeutic stratification [119,120,121,122,123,124,125,126,127,128,129]. Of particular interest is the hypermethylation of the hMLH1 promoter, which was found in half of the patients with microsatellite instability and was associated with drug resistance [130,131]. These findings increase the understanding of epigenetic changes in colorectal carcinoma. The number of disparate critical drivers in colorectal cancer progressively leads to a “multi-gene, multi-drug” therapeutic strategy [132].

2.2.2. Inflammation

It is broadly accepted that inflammation is a common event during aging, referred to as “inflamm-aging” [133]. This process is a underlying condition of several diseases such as sarcopenia, osteoarthritis, and cancer [134]. A hallmark of these processes is an increase in Tumor Necrosis Factor alpha (TNFα) levels. DNA methylation and histone acetylation modify the promoter region of TNFα [135]. The TNFα gene itself does not contain a classical CpG island, however, its promoter and first exon were described to be rich in CpG sequences [135]. Accordingly, methylation on these gene regions has been described to negatively regulate TNFα expression levels [136]. Wang et al. demonstrated in porcine spleens by bisulfite sequencing PCR and qPCR that the TNFα promoter region was increasingly methylated with age, correlating with decreased mRNA expression [137].
Similarly, NF-κB mediates acute, as well as chronic inflammation, and is proposed as one of the key regulators of aging. Via its transcriptional activity, the NF-κB family induces the expression of cytokines and genes associated with apoptosis and senescence as described elsewhere [138]. NF-κB levels can be regulated by various epigenetic mechanisms including the acetylation of histone H3 via the H3 lysine 4 methyltransferase SET7/9 [139] which represents a potential targeting strategy [136]. The link to aging has been validated in the skin where C57BL/6 mice exposed to UVB light for 16 days displayed accelerated aging of the skin via NF-κB activation through the mTORC2 pathway. The post-translational modification of the p65 member of NF-κB at Ser536 enhanced NF-κB activity via increased DNA binding activity. The same phenomenon was detected in physiological aged skin of these mice, demonstrating an accelerated inflammatory status in physiological as pathological (skin) aging [140,141].
Another epigenetic modification, which has been demonstrated to regulate NF-κB activity during inflammation, is the monoubiquitination at lysine 120 of histone H2B (H2Bub1). This monoubiquitination is performed by the RNF20/RNF40 E3 ligase complex and leads to increased chromatin accessibility. This results in eased passage of RNA Polymerase II and highly active transcriptional elongation [142]. Recently, it has been demonstrated that the monoubiquitination of histone H2B regulates NF-κB signaling in intestinal inflammation. However, the function of H2Bub1 in animal models for colitis remains inconclusive [143,144].
In general, the transformation from chronic inflammation to cancer can be promoted via DNA methylation, histone modifications, chromatin remodeling and noncoding RNA regulation, upon which the most important pathways are NF-κB- and STAT3-related [122,145]. The phosphorylation of Tyr705, Ser727, and Ser727 are known to positively activate transcriptional activity of STAT3 [145,146]. One downstream target of this signaling pathway is interleukin-6, which has been shown to be repressed via treatment with the DNMT1 inhibitor 5-azadeoxycytidine (5-AzaC) [122]. MicroRNAs are further key regulators of inflammatory responses and inflamed tissues are characterized by downregulation of TET gene expression due to the upregulation of TET-targeting miRNAs (e.g., MiR20a, MiR26B, MiR29C, Let-7 microRNA) [147,148].
Interestingly, the “epigenetic clock” concept of Horvath, has been demonstrated to be very accurate when methylation levels of CpG sites from white blood cells, the central regulators of immune response, were used [149,150,151]. These methylation levels were even able to predict mortality [152]. Despite a loss of T-cell diversity in old age, an exhausted/senescent CD8+ T cell population increases with age, possibly giving rise to associated diseases [153,154,155,156,157].
Excessive inflammatory response, immunosenescence, and autoimmunity outline the other detrimental side of the inflammatory spectrum. The identification of hypomethylated apoptosis-related genes in naïve CD4+ T cells led to the definition of an evolving autoimmune epigenotype [158,159,160]. Accordingly, in chronic nonbacterial osteomyelitis, a reduced expression of immunoregulatory cytokines (IL-10, IL-19) was centrally involved. The authors demonstrated in monocytes from chronic recurrent multifocal osteomyelitis patients that an altered SP1 activation negatively affected IL10 and IL19 expression. Mechanistically, the reduced phosphorylation of histone 3 serine 10 (H3S10P) and impaired SP1 phosphorylation at the IL10 and IL19 promoter regions impaired IL10 expression. This causes an imbalance towards proinflammatory cytokines (compared to anti-inflammatory IL-10 and IL-19), leading to inflammatory bone loss [161]. Similar reductions of H3S1p levels have been identified in hippocampi of aged mice by Wu et al. [162], which were associated with the inflammation-related decline in spatial learning and memory.

2.2.3. Osteoporosis

Osteoporosis is tightly linked to aging via epigenetic changes in mesenchymal stem cells (MSCs) [163]. Physiologically, the Osterix promoter was shown to entail enriched levels of H3Ac/H3K4me3 and reduced levels of H3K9me3/H3K27me3, inducing the differentiation of MSCs into osteoblasts to mediate skeletal tissue homeostasis [163,164]. Among the key transcription factors for osteogenesis are HOX and RUNX2, both of which are hypermethylated in aged MSCs [165]. Bork and colleagues demonstrated in MSCs isolated from bone marrow aspirates from young and old human donors that long-term cell culture and regular aging result in similar epigenetic profiles. HOXA (2,5,6) and RUNX2, transcription factors involved in osteoblast differentiation, were the most prominent among genes hypermethylated during aging which leading to decreased gene expression and age-related bone loss [166].
To prevent the hypermethylation of RUNX2, the transcriptional activation of the methyltransferase DNMT1 can be inhibited via 5-AzaC treatment. As expected, 5-AzaC leads to a hypomethylation of genomic DNA resulting in increased expression of RUNX2, Osteocalcin (OCN) and Osterix (OSX). The beneficial effect of 5-AzaC was supported by Zhou et al. in cell culture experiments using MSCs. Besides a global reduction in methylation levels, an increase in osteogenic gene expression was detected as demonstrated via enhanced alkaline phosphatase (ALP) activity and, subsequently, aggrandized mineralization [165].
It was shown in human bone marrow stromal cells (BMSCs) that in osteoporosis the number of clonogenic BMSCs was reduced, corresponding to decreased levels of Tet1 and Tet2, factors, which are able to erase DNA methylation. During normal osteogenesis, TET1 and TET2 levels were enhanced with an increased binding to the Osterix promoter [163,167]. Yang et al. detected an osteopenic phenotype and decreased Runx2 expression in Tet1−/−Prx1creTet2fl/fl mice. After the authors demonstrated that the use of siRNAs against TET1 and TET2 led to reduced stem cell properties in bone marrow MSCs (BMMSCs), they found via RNA-seq and qPCR in their mouse model that miRNAs targeting Runx2 gene expression were significantly higher in the knockout mice and a treatment with mimics of these miRNAs increased Runx2 expression in BMMSCs. The authors used chromatin immunoprecipitation (ChIP)-qPCR to demonstrate that TET1 and TET2 directly bind to the CpG island of the P2rX7 promoter, a gene which has been linked to exosome release in earlier studies. After a depletion of Tet1 and Tet2, the subsequent methylation led to impaired self-renewal and differentiation potential on the stem cell level in the bone marrow, thus leading to an osteopenic phenotype [168]. Rising evidence shows a broad involvement of several miRNAs in osteoporosis, such as miR-297a-5p, miR-297b-5p, and miR-297c-5p. These are accumulating intracellularly, inhibiting RUNX2 expression, and thereby promoting an osteoporotic phenotype. This TET/P2rX7/RUNX2 cascade may serve as a target for novel therapeutic approaches [168,169]. Again, the link of Tet1/2 to aging has been made by Gontier et al. in the mouse brain, where a Tet2 reduction was detected in the hippocampi of aged mice, and the application of high-titer lentivirus encoding for Tet2 shRNA in young adult mice caused deficits in short-term and long-term learning [170].
Mechanistically, an imbalance between histone modifications of osteogenic and adipogenic genes was proposed as an underlying mechanism of the development of musculoskeletal diseases. For instance, HDAC3 promotes osteogenesis and inhibits lipogenesis, while EZH2 and HDAC6 show opposite effects [165,171]. In bone marrow aspirates from human adults, it was demonstrated via retroviral-mediated enforced Ezh2 expression in MSCs that the differentiation potential into adipocytes was higher compared to vector control cells, along with reduced Runx2 transcription. Subsequent siRNA-mediated EZH2 depletion led to enhanced RUNX2 expression. Using ChIP-qPCR, the authors demonstrated that enforced EZH2 expression in MSCs resulted in increased H3K27me3 on transcriptional start sites of RUNX2, leading to a suppression of osteogenesis and marking EZH2 as positive regulator of adipogenesis and negative regulator of osteogenesis [171,172,173,174,175]. In another study, age-related bone loss was found linked to an increase in EZH2. In osteoporotic mice, Jing et al. detected via ChIP-qPCR that EZH2 was enriched at promoters of Wnt pathway members in BMSCs and a knockdown of EZH2 decreased H3K27me3 occupancy on these factors enhancing the Runx2 and Osterix expression and subsequently osteogenic differentiation. Accordingly, the authors suggested the H3K27me3 inhibitor DZNep as a potential therapeutic substance for anti-osteoporotic treatment [172].
This translation of epigenetic discoveries into the clinic has already been established for several years. In fact, bisphosphonates (increasing MiR191c-5p and miR-497-5p) and monoclonal antibodies, such as denosumab, regulate—among others—DNA methyltransferases, histone acetylases, deacetalyses, and other key factors associated with detrimental epigenetic alterations [173,174,175].

2.2.4. Neurodegenerative Diseases

Neurodegenerative diseases are tightly linked to the process of aging. Twin-studies demonstrated that accelerated epigenetic age, i.e., a higher methylation level, was linked to the development and progression of neurodegenerative diseases [176,177]. Alzheimer’s disease (AD) and Parkinson’s disease (PD) depict two distinct forms of neurodegenerative diseases that have been found associated with epigenetic modifications [178].

Alzheimer’s Disease

In our aging society, Alzheimer is displays a major health burden and cellular senescence was suggested as a key regulator controlling the shift from physiological aging into neurodegeneration [179]. Interestingly, senescent cells show a decrease in repressive heterochromatin marks, including H3K9me3, H3K27me3, and H4K20me3, leading to altered transcription patterns. In particular, the H4K16ac occupancy was reduced in regulatory regions of genes linked to aging in AD patients. Nativio et al. found in human postmortem brain samples from the temporal lobe via ChIP-seq analysis that H4K16ac levels increased in elderly healthy, but not aged AD patients. In particular, the reduced acetylation in AD patients was prominent on HIC1 (p53-mediated DNA-damage response) and Wnt pathway members (synaptic transmission and plasticity). A protective effect of H4K16ac against neurodegenerative diseases has been suggested and therapeutically explored using a miR-149-5p inhibitor to increase H4K16ac levels in the AD in vitro model 293/APPsw, resulting in reduced β-amyloid formation, potentially attenuating AD progression [180,181,182]. Another gene of interest is the silencing transcription factor REST, promoting cellular cell death in AD and protecting neurons against oxidative stress and β-amyloid toxicity. Notably, it was demonstrated that physiological aged human brains showed increased REST mRNA and protein levels compared to AD patients. While a conditional deletion of Rest in mouse brains (Nestin-Cre:RESTlx/lx) promoted age-related neurodegeneration, the neuroprotective effect of REST was confirmed by transgenic expression of human REST in C. elegans, reducing the sensitivity to oxidative stress and Aβ toxicity [183,184]. Summarizing, REST induction frequently appears in the (beneficial) aging process, presumably protecting the brain, while its decrease (as in AD) was associated with neurodegeneration [181,185]. In addition, Hou and co-workers demonstrated in mice that the loss of REST in AD is connected to abnormalities in miR124 signaling. In fact, Rest overexpression suppressed miR-124 while, consequently, the inhibition of miR-124 decreased tau aggregation and rescued memory deficits in 10-month-old P3001S mutant mice [186]. Moreover, 7mi-RNA was found to be significantly increased in the plasma of AD patients which was also true for hsa-miR-27a-3p in the cerebrospinal fluid, whereas the plasma levels of hsa-let-7d-5p and hsa-let-7-g-5p were discovered as potential biomarkers [187,188].

Parkinson’s Disease

In PD, the methylation by DNA methyltransferase 1 (DNMT1) as regulator of the alpha-synuclein (SNCA) gene is crucial. DNMT1 interacts with α-synuclein and is required for its localization in the cytoplasm, thereby reducing its effect on biophysical properties of the DNA [189,190]. However, as determined in a genome-wide association study in combination with rigid-body dockings simulation, nucleotide polymorphisms in the methylation region of the first intron of SNCA, which is required for the interaction with DNMT1, may influence the susceptibility to PD [191]. While overall a lower methylation rate was observed in diverse brain regions in PD patients, an enrichment of H3K27ac at enhancer regions was found at the SNCA locus in multiple brain regions associated with PD via human genome-scale enhancer identification via ChIP-seq. A possible targeting approach of this hyperacetylation was found by Mittal et al. who used human SK-N-MC neuroblastoma cells to detect compounds to decrease SNCA expression. Β2AR (β2-adrenoreceptor) agonists were found to modulate SNCA transcription through H3K27 deacetylation at its promoter and enhancer sites, promoting dopamine neuron health by reduced SNCA expression. The authors subsequently demonstrated in a nationwide longitudinal analysis that the most commonly used β2AR agonist, salbutamol, was associated with a reduced incidence rate of PD [192,193,194]. Moreover, it has been demonstrated that α-synuclein directly binds histones and inhibits the acetylation of histone H3 via the SIRT2 deacetylase, leading to decreased H3K9 acetylation as found in postmortem Parkinson patients’ primary motor cortex [195,196,197]. The central role of SIRT2 has been demonstrated by Esteves and colleagues in transmitochondrial cybrids, who demonstrated that SIRT2 increases microtubule instability via α-tubulin deacetylation and tau hyperphosphorylation. SIRT2 inhibition (via AK1), on the other side, improved intracellular trafficking [195]. Recently, a part of the SWI/SNF complex, SMARCA4, has been linked to age-related dopaminergic degeneration. Making use of gene co-expression analysis in human brain samples and Drosophila PD models, Sun et al. demonstrated that SMARCA4 was upregulated with aging and a down-regulation of SMARCA4 via siRNAs (drosophila: Brm) in dopaminergic neurons restored life span, indicating possible future diagnostic and therapeutic approaches based on SMRCA4 [198].

2.2.5. Diet, Nutrition, and Type 2 Diabetes

Obesity is pathophysiologically associated with the development of type II diabetes [199,200]. Oxidative stress and inflammation, metabolic impairment and accelerated aging on both the micro- and macrocellular level contribute to the pathogenesis of metabolic diseases [201,202].

Dietary Restriction and Hunger

One of the earliest indications of a regulatory mechanism beyond sole genetics was deducted from the Dutch Hunger Winter families. In this observational study, an early-life (gestation) adverse environmental factor like famine (in the Netherlands and years of World War II 1944–1945) led to decreased methylation levels at distinct CpG islands, for instance on the Insulin-like Growth Factor II (IGF2) gene, influencing long-term metabolic health even six decades later [203,204]. In particular, the risk for obesity and type 2 diabetes was significantly higher when certain CpG sites (cg00574958, cg06500161) exhibited a high methylation status [205,206].
In general, nutritional influences were demonstrated to affect cellular longevity and carcinogenesis by telomerase and CDKN2A (p16) modulation: glucose restriction inhibited cellular senescence via chromatin remodeling and histone acetylation as well as methylation of the CDKN2A promoter, impairing E2F1 binding [205,206]. Consequently, caloric restriction builds a connection between aging and cancer: It has been indicated that caloric restriction increases lifespan via modifying the mammalian Target of Rapamycin (mTOR)-pathway, insulin/IGF-1 like signaling and sirtuins. Li and Tollefsbol investigated the effects of glucose restriction using lung fibroblasts in vitro and found that glucose restriction inhibited cellular senescence by measuring SA-βgal activity. Underlying CDKN2A mRNA did not accumulate in glucose-restricted medium compared to the normal glucose medium which was due to increased H3K9me3 (inactive chromatin marker) occupancy at the CDKN2A promoter. In contrast, active histone marks (H3 acetylation and H3K4me2) were detected to a lower extent at CDKN2A promoter region in glucose-restricted medium. Together, glucose deficiency led to a decrease of age-related histone modifications, subsequently, to a reduced incidence of aging-associated diseases [205,206,207].
On the molecular level, dietary restrictions can be mimicked by polyphenols, a large family of organic compounds naturally contained in fruits, vegetables, and cereals [208]. Interestingly, Sirtuin-1 and the senescence-associated secretory phenotype (SASP) are regulated by polyphenols. With molecular dynamics simulations and fragment-centric topographical binding interface mapping, Hou et al. demonstrated that resveratrol, a natural phenol, stabilized SIRT1/peptide interactions by forming a new binding pocket [209]. Since SIRT1 protein levels were increased after calorie restriction, mimetics such as resveratrol are of great interest in the treatment of aging-related diseases. Additional induction of autophagy and neutralization of free radicals resulted in further modulation of age-related detrimental mechanisms [210,211].

High Fat Diet and Obesity

Since aging and obesity have been demonstrated to be closely linked to each other, demographical changes will lead to an increased incidence of obesity in the future [212]. An increased variability in DNA methylation patterns was determined via observational studies in obesity and related phenotypes, indicating epigenetic dysregulation [213]. Histone modifications display the interface through which diet and stress affect organisms on a subcellular level [214]. Analyzing genome-wide methylation profiles in peripheral blood from obese and lean young humans, distinct CpG sites were associated with obesity and a prediction model was able to project adiposity with 70% confidence. In fact, obesity has been linked to differential methylation patterns and variability in CpG sites in proximity to certain genes, out of which HIF3A, CPT1A, CD38, and PHGDH were described to be most important [202,215].
However, the effects of diet on epigenetic patterns appear to be long-lasting and do not affect just one generation: A high-fat diet in embryonic developmental stages substantially affected later life [216]. The maternal diet during gestational and lactational period resulted in severe long-term effects. Zheng et al. demonstrated that a high-fat diet in maternal rats affected the p16 (Ink4a) protein and mRNA levels in the mammary gland of the offspring. Subsequently, ChIP revealed reduced H4 acetylation at the p16 promoter at the CpG-rich sites and enhanced recruitment of HDAC3 to the p16 promoter regulatory region in the mammary glands of the offspring after a maternal high-fat diet [214,217]. While the effects of a high-fat diet on crucial epigenetic promoter sites affecting Ink4a were investigated in detail, the effects of this diet on p21 (Cip1), another cyclin-dependent kinase inhibitor, appeared to be less crucial. Zhang et al. analyzed the effect of a high-fat diet in the liver of male rats. After obtaining a diet rich in fat for 9 weeks, the senescence marker p16(INK4a) was increased due to a decrease of H3K27me3 in the coding region of p16. Histologically, a hepatic steatosis and fat accumulation were observed. Contrary, the expression and protein level of p21 was decreased after a high-fat diet with no significant chromatin modifications of this gene [214,218]. Intriguingly, consequences of high-fat diets can even be detected after a short time (5 days) overfeeding diet in healthy young men. When evaluating the DNA methylation status in 21 healthy young men’s skeletal muscle biopsies after a diet with 50% extra calories for five days, methylation changes were detected on 6508 genes in the skeletal muscle and could only be partly reversed after 6–8 weeks. This finding indicated a slow reversibility of methylation patterns which, in the long-term, may influence gene expression levels [219].


Besides all other aforementioned conditions, diabetes has been previously associated with the involvement of epigenetics. In fact, epigenetic deregulation has frequently been observed in the β-cells of the pancreatic islands which showed an impaired capacity to secrete insulin in diabetes patients [201]. Using pancreatic islets from human donors, it was demonstrated that, along with DNA methylation changes at their CpG sites, a high-glucose stimulation altered expression patterns of a subset of genes including CHRNA5, GLRA1, and PDX1. Generally, diabetic islets-associated genes showed an overall trend towards increased DNA methylation and a decreased expression of key genes linked with impaired insulin secretion. Pathogenetically, the DNA methylation status (especially CpG sites annotated to the transcription factor PDX1) can be directly elevated via high glucose and glycated hemoglobin (HbA1c) while differentially methylated regions in pancreatic islets (PDX1, TCF7L2, ADCY5) promote islet dysfunction [220,221]. Another underlying mechanism, supporting the “inflamm-aging” theory, has been unveiled by Sandovici et al. The authors analyzed pancreatic islets from young and old rats and found differential expression of genes regulating islet cell function, type 2 diabetes genes and inflammatory processes, while age-associated transcriptional differences were negatively correlated with promoter DNA methylation at these gene loci [222].
The multifactorial and complex polygenicity of diabetes mellitus suggests that the effect size of single genes and CpG sites is low. Among the genes that show decreased DNA methylation and increased gene expression rates in diabetic patients, CDKN1A (p21), PDE7B, and SEPT19 are of predominant interest. It was discovered in pancreatic islands from diabetic and non-diabetic donors that CDKN1A was differentially expressed in diabetic islands. In fact, the impaired glucose-stimulated insulin secretion was associated with an increased expression and reduced DNA methylation pattern of CDKN1A. Via transfection of clonal β-cells with mock-methylated or methylated constructs, the promoter methylation of CDKN1A (p21) has been demonstrated to directly negative affect the transcriptional activity [223]. As a cyclin-dependent kinase inhibitor, CDKN2A (p16) regulates the cell-cycle progression to G1 and an overexpression led to decreased cell proliferation, an “aged” phenotype, influencing both islet and non-islet mechanisms driving the diabetic phenotype [223,224,225].
The majority of association studies has shown multiple gene loci for epigenetic regulation in these central mediators of type II diabetes, β-cells. Chen and colleagues characterized Ezh2fl/fl mice and Cdkn2a−/− mice to reveal that an increased Ink4a and Arf expression in β-cells was linked to a reduced proliferative capacity. While Ezh2 levels declined throughout aging, INK4A levels increased. ChIP analysis uncovered that H3K27me3 occupancy regulating Ink4a and Ezh2 was declining with age, while H3K4me3 and histone acetylation at the Ink4a locus ascended in older mice. The authors concluded from their study that EZH2-dependent histone methylation and repression of the Ink4a/Arf locus are required for β-cell expansion [223,226]. In a further study, the methylome of β cells was analyzed pancreatic islets from young and old mice using whole genome shotgun bisulfite sequencing (WGSBS). Overall, higher methylation rates (especially in CpGs with low methylation levels in youth), accompanied by a decline in replicative capacity, increased promoter methylation and decreased expression of cell cycle regulators were detected in “healthy” old β-cells. Intriguingly, this observation was associated with a functional improvement in aged murine and human islets [223,227].
In addition, a high relevance of immunoregulatory mechanisms has been detected in type I diabetes. Miao et al. compared blood cells from type I diabetes patients to healthy controls and performed ChIP to identify diabetic methylation profiles. Variations in H3K9ac levels were found at upstream regions of human leukocyte antigen (HLA)-DRB1 and HLA-DQB1 in monocytes from diabetic patients. An increase in H3K9ac levels at the same promoter regions of HLA-DQB1, which was hyperacetylated in type I diabetes patients was found after Interferon gamma (IFNγ) and TNFα stimulation. This suggests that methylation changes in the human leukocyte antigen (HLA) region could precede the development of type I diabetes [228,229]. HLA-DQB1 is among the most relevant HLAs for genetic predisposition since it was identified to be highly probable to contain methylation-variable positions, possibly indicating an early etiological state in the development of type I diabetes [230].
Upon further epigenetic regulatory elements in diabetes, micro-RNAs, such as miR-15a and miR-29b, were found to be downregulated in type 2 diabetes, whereas miR-27a and miR-320a were upregulated and might open the possibility for new diagnostic markers [187,231,232,233].

3. Conclusions and Perspectives

This review provides a comprehensive overview of the key role of epigenetic mechanisms in controlling aging as well as the development of aging-associated pathologies. Numerous studies focusing on aging-related molecular mechanisms using cell-based systems, experimental animal models, as well as primary tissues have profoundly contributed to the current knowledge on this topic.
It has been shown that distinct epigenetic mechanisms are enriched in aging organisms including the accumulation of histone variants as well as the loss of histones and heterochromatin (Figure 1). In addition, histones show aberrant post-translational modifications leading to the imbalance of activating and repressing modifications. Moreover, remodeling complexes modulate chromatin accessibility and there is an aberrant expression/activity of miRNAs. Together, these epigenetic deregulations contribute to aging-associated changes in gene transcription and, as a consequence, translation as well as the stabilization or degradation of molecular factors. While mechanisms underlying aging-related pathologies remain to be elucidated in detail, various studies demonstrate an epigenetic component. In fact, the aforementioned epigenetic modifications were shown to play essential roles in diseases including inflammation, cancer, osteoporosis, neurodegenerative diseases, and diabetes.
While the precise mechanisms and connections between several epigenetic changes and human pathologies are still poorly understood, state-of-the-art next generation sequencing methods will allow researchers to address remaining questions. For instance, chromatin accessibility using Assay for Transposase-Accessible Chromatin using sequencing (ATAC-seq) can be coupled to ChIP-seq as well as gene expression studies (mRNA-seq) using bulk mRNA or even analyzing single cells (scRNA-seq). In addition, advances in molecular biology and cell culture approaches (for instance Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas9) will be beneficial in clarifying aging-processes across species.
An improved understanding of epigenetic mechanisms affecting longevity will be deciding crucial step towards the identification of new potential therapeutic targets. In fact, epigenetic drugs are of particular interest to the clinic due to their reversible and transient effect.
A limitation of manifold epigenetic studies, however, are the variations among single cells (both on an individual and tissue level), which occur with an even higher frequency in aged organisms [234,235]. This biologically relevant heterogeneity might be further investigated, understood and potentially deconstructed with the help of new technological approaches like single-cell genomics [236,237,238].
Together, characterizing molecular changes in different species during aging using state-of-the-art techniques will provide key insights into the relevance of epigenetics of aging and aging-associated diseases.

Author Contributions

All authors contributed crucially to the manuscript. Conceptualization: D.S. and R.L.K. Methodology: D.S. and R.L.K. Analysis of the articles: D.S. and R.L.K. Writing—original draft preparation D.S. and R.L.K. Writing—review and editing: D.S. and R.L.K. Supervision D.S. and R.L.K. Project administration: D.S. and R.L.K. Funding acquisition: D.S. and R.L.K. All authors have read and agreed to the published version of the manuscript.


D.S. was funded by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation)—413501650. R.L.K. was funded by the German Cancer Aid (Mildred Scheel postdoctoral fellowship).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Data sharing not applicable.

Conflicts of Interest

The authors declare no conflict of interest.


  1. Kennedy, B.K.; Berger, S.L.; Brunet, A.; Campisi, J.; Cuervo, A.M.; Epel, E.S.; Franceschi, C.; Lithgow, G.J.; Morimoto, R.I.; Pessin, J.E.; et al. Geroscience: Linking aging to chronic disease. Cell 2014, 159, 709–713. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  2. López-Otín, C.; Blasco, M.A.; Partridge, L.; Serrano, M.; Kroemer, G. The hallmarks of aging. Cell 2013, 153, 1194–1217. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  3. Luger, K.; Mäder, A.W.; Richmond, R.K.; Sargent, D.F.; Richmond, T.J. Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature 1997, 389, 251–260. [Google Scholar] [CrossRef] [PubMed]
  4. Richmond, T.J.; Davey, C.A. The structure of DNA in the nucleosome core. Nature 2003, 423, 145–150. [Google Scholar] [CrossRef] [PubMed]
  5. Torres, I.O.; Fujimori, D.G. Functional coupling between writers, erasers and readers of histone and DNA methylation. Curr. Opin. Struct. Biol. 2015, 35, 68–75. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  6. Zhang, W.; Qu, J.; Liu, G.-H.; Belmonte, J.C.I. The ageing epigenome and its rejuvenation. Nat. Rev. Mol. Cell Biol. 2020, 21, 137–150. [Google Scholar] [CrossRef] [PubMed]
  7. Hu, Z.; Chen, K.; Xia, Z.; Chavez, M.; Pal, S.; Seol, J.-H.; Chen, C.-C.; Li, W.; Tyler, J.K. Nucleosome loss leads to global transcriptional up-regulation and genomic instability during yeast aging. Genes Dev. 2014, 28, 396–408. [Google Scholar] [CrossRef] [Green Version]
  8. Liu, L.; Cheung, T.H.; Charville, G.W.; Hurgo, B.M.C.; Leavitt, T.; Shih, J.; Brunet, A.; Rando, T.A. Chromatin modifications as determinants of muscle stem cell quiescence and chronological aging. Cell Rep. 2013, 4, 189–204. [Google Scholar] [CrossRef] [Green Version]
  9. 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] [Green Version]
  10. Lee, J.-H.; Kim, E.W.; Croteau, D.L.; Bohr, V.A. Heterochromatin: An epigenetic point of view in aging. Exp. Mol. Med. 2020, 52, 1466–1474. [Google Scholar] [CrossRef]
  11. Marzluff, W.F.; Gongidi, P.; Woods, K.R.; Jin, J.; Maltais, L.J. The human and mouse replication-dependent histone genes. Genomics 2002, 80, 487–498. [Google Scholar] [CrossRef] [PubMed]
  12. Henikoff, S.; Smith, M.M. Histone variants and epigenetics. Cold Spring Harb. Perspect. Biol. 2015, 7, a019364. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  13. Buschbeck, M.; Uribesalgo, I.; Wibowo, I.; Rué, P.; Martin, D.; Gutierrez, A.; Morey, L.; Guigó, R.; López-Schier, H.; Di Croce, L. The histone variant macroH2A is an epigenetic regulator of key developmental genes. Nat. Struct. Mol. Biol. 2009, 16, 1074–1079. [Google Scholar] [CrossRef] [PubMed]
  14. Pehrson, J.R.; Fried, V.A. MacroH2A, a core histone containing a large nonhistone region. Science 1992, 257, 1398–1400. [Google Scholar] [CrossRef] [PubMed]
  15. Gaspar-Maia, A.; Qadeer, Z.A.; Hasson, D.; Ratnakumar, K.; Leu, N.A.; Leroy, G.; Liu, S.; Costanzi, C.; Valle-Garcia, D.; Schaniel, C.; et al. MacroH2A histone variants act as a barrier upon reprogramming towards pluripotency. Nat. Commun. 2013, 4, 1565. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  16. Kreiling, J.A.; Tamamori-Adachi, M.; Sexton, A.N.; Jeyapalan, J.C.; Munoz-Najar, U.; Peterson, A.L.; Manivannan, J.; Rogers, E.S.; Pchelintsev, N.A.; Adams, P.D.; et al. Age-associated increase in heterochromatic marks in murine and primate tissues. Aging Cell 2011, 10, 292–304. [Google Scholar] [CrossRef] [Green Version]
  17. Tvardovskiy, A.; Schwämmle, V.; Kempf, S.J.; Rogowska-Wrzesinska, A.; Jensen, O.N. Accumulation of histone variant H3.3 with age is associated with profound changes in the histone methylation landscape. Nucleic Acids Res. 2017, 45, 9272–9289. [Google Scholar] [CrossRef]
  18. Piazzesi, A.; Papić, D.; Bertan, F.; Salomoni, P.; Nicotera, P.; Bano, D. Replication-Independent Histone Variant H3.3 Controls Animal Lifespan through the Regulation of Pro-longevity Transcriptional Programs. Cell Rep. 2016, 17, 987–996. [Google Scholar] [CrossRef] [Green Version]
  19. Maze, I.; Wenderski, W.; Noh, K.-M.; Bagot, R.C.; Tzavaras, N.; Purushothaman, I.; Elsässer, S.J.; Guo, Y.; Ionete, C.; Hurd, Y.L.; et al. Critical Role of Histone Turnover in Neuronal Transcription and Plasticity. Neuron 2015, 87, 77–94. [Google Scholar] [CrossRef] [Green Version]
  20. Stefanelli, G.; Azam, A.B.; Walters, B.J.; Brimble, M.A.; Gettens, C.P.; Bouchard-Cannon, P.; Cheng, H.-Y.M.; Davidoff, A.M.; Narkaj, K.; Day, J.J.; et al. Learning and Age-Related Changes in Genome-wide H2A.Z Binding in the Mouse Hippocampus. Cell Rep. 2018, 22, 1124–1131. [Google Scholar] [CrossRef] [Green Version]
  21. Biran, A.; Zada, L.; Abou Karam, P.; Vadai, E.; Roitman, L.; Ovadya, Y.; Porat, Z.; Krizhanovsky, V. Quantitative identification of senescent cells in aging and disease. Aging Cell 2017, 16, 661–671. [Google Scholar] [CrossRef] [PubMed]
  22. Pilch, D.R.; Sedelnikova, O.A.; Redon, C.; Celeste, A.; Nussenzweig, A.; Bonner, W.M. Characteristics of gamma-H2AX foci at DNA double-strand breaks sites. Biochem. Cell Biol. 2003, 81, 123–129. [Google Scholar] [CrossRef] [PubMed]
  23. Vidanes, G.M.; Bonilla, C.Y.; Toczyski, D.P. Complicated tails: Histone modifications and the DNA damage response. Cell 2005, 121, 973–976. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  24. Balajee, A.S.; Geard, C.R. Replication protein A and gamma-H2AX foci assembly is triggered by cellular response to DNA double-strand breaks. Exp. Cell Res. 2004, 300, 320–334. [Google Scholar] [CrossRef] [PubMed]
  25. Sharma, A.; Singh, K.; Almasan, A. Histone H2AX phosphorylation: A marker for DNA damage. Methods Mol. Biol. 2012, 920, 613–626. [Google Scholar] [CrossRef] [PubMed]
  26. Jin, B.; Li, Y.; Robertson, K.D. DNA methylation: Superior or subordinate in the epigenetic hierarchy? Genes Cancer 2011, 2, 607–617. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  27. Auclair, G.; Weber, M. Mechanisms of DNA methylation and demethylation in mammals. Biochimie 2012, 94, 2202–2211. [Google Scholar] [CrossRef]
  28. Hyun, J.; Jung, Y. DNA Methylation in Nonalcoholic Fatty Liver Disease. Int. J. Mol. Sci. 2020, 21, 8138. [Google Scholar] [CrossRef]
  29. Lister, R.; Pelizzola, M.; Dowen, R.H.; Hawkins, R.D.; Hon, G.; Tonti-Filippini, J.; Nery, J.R.; Lee, L.; Ye, Z.; Ngo, Q.-M.; et al. Human DNA methylomes at base resolution show widespread epigenomic differences. Nature 2009, 462, 315–322. [Google Scholar] [CrossRef] [Green Version]
  30. Schübeler, D. Function and information content of DNA methylation. Nature 2015, 517, 321–326. [Google Scholar] [CrossRef]
  31. He, Y.-F.; Li, B.-Z.; Li, Z.; Liu, P.; Wang, Y.; Tang, Q.; Ding, J.; Jia, Y.; Chen, Z.; Li, L.; et al. Tet-mediated formation of 5-carboxylcytosine and its excision by TDG in mammalian DNA. Science 2011, 333, 1303–1307. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  32. Ito, S.; Shen, L.; Dai, Q.; Wu, S.C.; Collins, L.B.; Swenberg, J.A.; He, C.; Zhang, Y. Tet proteins can convert 5-methylcytosine to 5-formylcytosine and 5-carboxylcytosine. Science 2011, 333, 1300–1303. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  33. Wu, H.; Zhang, Y. Reversing DNA methylation: Mechanisms, genomics, and biological functions. Cell 2014, 156, 45–68. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  34. Rasmussen, K.D.; Helin, K. Role of TET enzymes in DNA methylation, development, and cancer. Genes Dev. 2016, 30, 733–750. [Google Scholar] [CrossRef]
  35. Hashimoto, H.; Liu, Y.; Upadhyay, A.K.; Chang, Y.; Howerton, S.B.; Vertino, P.M.; Zhang, X.; Cheng, X. Recognition and potential mechanisms for replication and erasure of cytosine hydroxymethylation. Nucleic Acids Res. 2012, 40, 4841–4849. [Google Scholar] [CrossRef]
  36. Weinhouse, C.; Truong, L.; Meyer, J.N.; Allard, P. Caenorhabditis elegans as an emerging model system in environmental epigenetics. Environ. Mol. Mutagen. 2018, 59, 560–575. [Google Scholar] [CrossRef]
  37. Greer, E.L.; Blanco, M.A.; Gu, L.; Sendinc, E.; Liu, J.; Aristizábal-Corrales, D.; Hsu, C.-H.; Aravind, L.; He, C.; Shi, Y. DNA Methylation on N6-Adenine in C. elegans. Cell 2015, 161, 868–878. [Google Scholar] [CrossRef] [Green Version]
  38. Deshmukh, S.; Ponnaluri, V.C.; Dai, N.; Pradhan, S.; Deobagkar, D. Levels of DNA cytosine methylation in the Drosophila genome. PeerJ 2018, 6, e5119. [Google Scholar] [CrossRef] [Green Version]
  39. Day, K.; Waite, L.L.; Thalacker-Mercer, A.; West, A.; Bamman, M.M.; Brooks, J.D.; Myers, R.M.; Absher, D. Differential DNA methylation with age displays both common and dynamic features across human tissues that are influenced by CpG landscape. Genome Biol. 2013, 14, R102. [Google Scholar] [CrossRef] [Green Version]
  40. Horvath, S. DNA methylation age of human tissues and cell types. Genome Biol. 2013, 14, R115. [Google Scholar] [CrossRef] [Green Version]
  41. Horvath, S. Erratum to: DNA methylation age of human tissues and cell types. Genome Biol. 2015, 16, 96. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  42. Jylhävä, J.; Pedersen, N.L.; Hägg, S. Biological Age Predictors. EBioMedicine 2017, 21, 29–36. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  43. Liu, B.; Yip, R.K.; Zhou, Z. Chromatin remodeling, DNA damage repair and aging. Curr. Genomics 2012, 13, 533–547. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  44. Clapier, C.R.; Cairns, B.R. The biology of chromatin remodeling complexes. Annu. Rev. Biochem. 2009, 78, 273–304. [Google Scholar] [CrossRef]
  45. Wu, S.; Ge, Y.; Huang, L.; Liu, H.; Xue, Y.; Zhao, Y. BRG1, the ATPase subunit of SWI/SNF chromatin remodeling complex, interacts with HDAC2 to modulate telomerase expression in human cancer cells. Cell Cycle 2014, 13, 2869–2878. [Google Scholar] [CrossRef] [Green Version]
  46. Wu, S.; Ge, Y.; Li, X.; Yang, Y.; Zhou, H.; Lin, K.; Zhang, Z.; Zhao, Y. BRM-SWI/SNF chromatin remodeling complex enables functional telomeres by promoting co-expression of TRF2 and TRF1. PLoS Genet. 2020, 16, e1008799. [Google Scholar] [CrossRef]
  47. Riedel, C.G.; Dowen, R.H.; Lourenco, G.F.; Kirienko, N.V.; Heimbucher, T.; West, J.A.; Bowman, S.K.; Kingston, R.E.; Dillin, A.; Asara, J.M.; et al. DAF-16 employs the chromatin remodeller SWI/SNF to promote stress resistance and longevity. Nat. Cell Biol. 2013, 15, 491–501. [Google Scholar] [CrossRef]
  48. de Vaux, V.; Pfefferli, C.; Passannante, M.; Belhaj, K.; von Essen, A.; Sprecher, S.G.; Müller, F.; Wicky, C. The Caenorhabditis elegans LET-418/Mi2 plays a conserved role in lifespan regulation. Aging Cell 2013, 12, 1012–1020. [Google Scholar] [CrossRef]
  49. Min, J.-N.; Tian, Y.; Xiao, Y.; Wu, L.; Li, L.; Chang, S. The mINO80 chromatin remodeling complex is required for efficient telomere replication and maintenance of genome stability. Cell Res. 2013, 23, 1396–1413. [Google Scholar] [CrossRef] [Green Version]
  50. Dang, W.; Sutphin, G.L.; Dorsey, J.A.; Otte, G.L.; Cao, K.; Perry, R.M.; Wanat, J.J.; Saviolaki, D.; Murakami, C.J.; Tsuchiyama, S.; et al. Inactivation of yeast Isw2 chromatin remodeling enzyme mimics longevity effect of calorie restriction via induction of genotoxic stress response. Cell Metab. 2014, 19, 952–966. [Google Scholar] [CrossRef] [Green Version]
  51. Matilainen, O.; Sleiman, M.S.B.; Quiros, P.M.; Garcia, S.M.D.A.; Auwerx, J. The chromatin remodeling factor ISW-1 integrates organismal responses against nuclear and mitochondrial stress. Nat. Commun. 2017, 8, 1818. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  52. Iwasaki, W.; Miya, Y.; Horikoshi, N.; Osakabe, A.; Taguchi, H.; Tachiwana, H.; Shibata, T.; Kagawa, W.; Kurumizaka, H. Contribution of histone N-terminal tails to the structure and stability of nucleosomes. FEBS Open Bio 2013, 3, 363–369. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  53. Greer, E.L.; Shi, Y. Histone methylation: A dynamic mark in health, disease and inheritance. Nat. Rev. Genet. 2012, 13, 343–357. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  54. Greer, E.L.; Maures, T.J.; Hauswirth, A.G.; Green, E.M.; Leeman, D.S.; Maro, G.S.; Han, S.; Banko, M.R.; Gozani, O.; Brunet, A. Members of the H3K4 trimethylation complex regulate lifespan in a germline-dependent manner in C. elegans. Nature 2010, 466, 383–387. [Google Scholar] [CrossRef] [Green Version]
  55. Li, L.; Greer, C.; Eisenman, R.N.; Secombe, J. Essential functions of the histone demethylase lid. PLoS Genet. 2010, 6, e1001221. [Google Scholar] [CrossRef] [Green Version]
  56. Pu, M.; Wang, M.; Wang, W.; Velayudhan, S.S.; Lee, S.S. Unique patterns of trimethylation of histone H3 lysine 4 are prone to changes during aging in Caenorhabditis elegans somatic cells. PLoS Genet. 2018, 14, e1007466. [Google Scholar] [CrossRef] [Green Version]
  57. Cruz, C.; Della Rosa, M.; Krueger, C.; Gao, Q.; Horkai, D.; King, M.; Field, L.; Houseley, J. Tri-methylation of histone H3 lysine 4 facilitates gene expression in ageing cells. Elife 2018, 7. [Google Scholar] [CrossRef]
  58. Sen, P.; Dang, W.; Donahue, G.; Dai, J.; Dorsey, J.; Cao, X.; Liu, W.; Cao, K.; Perry, R.; Lee, J.Y.; et al. H3K36 methylation promotes longevity by enhancing transcriptional fidelity. Genes Dev. 2015, 29, 1362–1376. [Google Scholar] [CrossRef] [Green Version]
  59. Pu, M.; Ni, Z.; Wang, M.; Wang, X.; Wood, J.G.; Helfand, S.L.; Yu, H.; Lee, S.S. Trimethylation of Lys36 on H3 restricts gene expression change during aging and impacts life span. Genes Dev. 2015, 29, 718–731. [Google Scholar] [CrossRef] [Green Version]
  60. Su, L.; Li, H.; Huang, C.; Zhao, T.; Zhang, Y.; Ba, X.; Li, Z.; Zhang, Y.; Huang, B.; Lu, J.; et al. Muscle-Specific Histone H3K36 Dimethyltransferase SET-18 Shortens Lifespan of Caenorhabditis elegans by Repressing daf-16a Expression. Cell Rep. 2018, 22, 2716–2729. [Google Scholar] [CrossRef]
  61. Jin, C.; Li, J.; Green, C.D.; Yu, X.; Tang, X.; Han, D.; Xian, B.; Wang, D.; Huang, X.; Cao, X.; et al. Histone demethylase UTX-1 regulates C. elegans life span by targeting the insulin/IGF-1 signaling pathway. Cell Metab. 2011, 14, 161–172. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  62. Kelly, J.T.; Feldman, S.E.; Ross, J.E. Quality review in the Peer Review Organization Program. Qual. Assur. Util. Rev. 1987, 2, 107–110. [Google Scholar] [CrossRef] [PubMed]
  63. Siebold, A.P.; Banerjee, R.; Tie, F.; Kiss, D.L.; Moskowitz, J.; Harte, P.J. Polycomb Repressive Complex 2 and Trithorax modulate Drosophila longevity and stress resistance. Proc. Natl. Acad. Sci. USA 2010, 107, 169–174. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  64. Baumgart, M.; Groth, M.; Priebe, S.; Savino, A.; Testa, G.; Dix, A.; Ripa, R.; Spallotta, F.; Gaetano, C.; Ori, M.; et al. RNA-seq of the aging brain in the short-lived fish N. furzeri—conserved pathways and novel genes associated with neurogenesis. Aging Cell 2014, 13, 965–974. [Google Scholar] [CrossRef]
  65. Lorbeck, M.T.; Singh, N.; Zervos, A.; Dhatta, M.; Lapchenko, M.; Yang, C.; Elefant, F. The histone demethylase Dmel\Kdm4A controls genes required for life span and male-specific sex determination in Drosophila. Gene 2010, 450, 8–17. [Google Scholar] [CrossRef] [Green Version]
  66. Djeghloul, D.; Kuranda, K.; Kuzniak, I.; Barbieri, D.; Naguibneva, I.; Choisy, C.; Bories, J.-C.; Dosquet, C.; Pla, M.; Vanneaux, V.; et al. Age-Associated Decrease of the Histone Methyltransferase SUV39H1 in HSC Perturbs Heterochromatin and B Lymphoid Differentiation. Stem Cell Reports 2016, 6, 970–984. [Google Scholar] [CrossRef] [Green Version]
  67. Sarg, B.; Koutzamani, E.; Helliger, W.; Rundquist, I.; Lindner, H.H. Postsynthetic trimethylation of histone H4 at lysine 20 in mammalian tissues is associated with aging. J. Biol. Chem. 2002, 277, 39195–39201. [Google Scholar] [CrossRef] [Green Version]
  68. 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, 158. [Google Scholar] [CrossRef] [Green Version]
  69. Kawakami, K.; Nakamura, A.; Ishigami, A.; Goto, S.; Takahashi, R. Age-related difference of site-specific histone modifications in rat liver. Biogerontology 2009, 10, 415–421. [Google Scholar] [CrossRef]
  70. Michishita, E.; McCord, R.A.; Berber, E.; Kioi, M.; Padilla-Nash, H.; Damian, M.; Cheung, P.; Kusumoto, R.; Kawahara, T.L.A.; Barrett, J.C.; et al. SIRT6 is a histone H3 lysine 9 deacetylase that modulates telomeric chromatin. Nature 2008, 452, 492–496. [Google Scholar] [CrossRef]
  71. Mostoslavsky, R.; Chua, K.F.; Lombard, D.B.; Pang, W.W.; Fischer, M.R.; Gellon, L.; Liu, P.; Mostoslavsky, G.; Franco, S.; Murphy, M.M.; et al. Genomic instability and aging-like phenotype in the absence of mammalian SIRT6. Cell 2006, 124, 315–329. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  72. Kawahara, T.L.A.; Michishita, E.; Adler, A.S.; Damian, M.; Berber, E.; Lin, M.; McCord, R.A.; Ongaigui, K.C.L.; Boxer, L.D.; Chang, H.Y.; et al. SIRT6 links histone H3 lysine 9 deacetylation to NF-kappaB-dependent gene expression and organismal life span. Cell 2009, 136, 62–74. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  73. Kanfi, Y.; Naiman, S.; Amir, G.; Peshti, V.; Zinman, G.; Nahum, L.; Bar-Joseph, Z.; Cohen, H.Y. The sirtuin SIRT6 regulates lifespan in male mice. Nature 2012, 483, 218–221. [Google Scholar] [CrossRef] [PubMed]
  74. Chen, J.; Xie, J.-J.; Jin, M.-Y.; Gu, Y.-T.; Wu, C.-C.; Guo, W.-J.; Yan, Y.-Z.; Zhang, Z.-J.; Wang, J.-L.; Zhang, X.-L.; et al. Sirt6 overexpression suppresses senescence and apoptosis of nucleus pulposus cells by inducing autophagy in a model of intervertebral disc degeneration. Cell Death Dis. 2018, 9, 56. [Google Scholar] [CrossRef] [PubMed]
  75. Ozdemir, A.; Spicuglia, S.; Lasonder, E.; Vermeulen, M.; Campsteijn, C.; Stunnenberg, H.G.; Logie, C. Characterization of lysine 56 of histone H3 as an acetylation site in Saccharomyces cerevisiae. J. Biol. Chem. 2005, 280, 25949–25952. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  76. Yuan, J.; Pu, M.; Zhang, Z.; Lou, Z. Histone H3-K56 acetylation is important for genomic stability in mammals. Cell Cycle 2009, 8, 1747–1753. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  77. Kokkonen, P.; Rahnasto-Rilla, M.; Mellini, P.; Jarho, E.; Lahtela-Kakkonen, M.; Kokkola, T. Studying SIRT6 regulation using H3K56 based substrate and small molecules. Eur. J. Pharm. Sci. 2014, 63, 71–76. [Google Scholar] [CrossRef]
  78. Liu, X.; Li, C.; Li, Q.; Chang, H.-C.; Tang, Y.-C. SIRT7 Facilitates CENP-A Nucleosome Assembly and Suppresses Intestinal Tumorigenesis. iScience 2020, 23, 101461. [Google Scholar] [CrossRef]
  79. Krishnan, V.; Liu, B.; Zhou, Z. ‘Relax and Repair’ to restrain aging. Aging (Albany NY) 2011, 3, 943–954. [Google Scholar] [CrossRef]
  80. Xu, H.; Su, T.; Xue, Y. Histone H3 N-terminal acetylation sites especially K14 are important for rDNA silencing and aging. Sci. Rep. 2016, 6, 21900. [Google Scholar] [CrossRef] [Green Version]
  81. Krishnan, V.; Chow, M.Z.Y.; Wang, Z.; Zhang, L.; Liu, B.; Liu, X.; Zhou, Z. Histone H4 lysine 16 hypoacetylation is associated with defective DNA repair and premature senescence in Zmpste24-deficient mice. Proc. Natl. Acad. Sci. USA 2011, 108, 12325–12330. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  82. Barral, S.; Beltramo, R.; Salio, C.; Aimar, P.; Lossi, L.; Merighi, A. Phosphorylation of Histone H2AX in the Mouse Brain from Development to Senescence. Int. J. Mol. Sci. 2014, 15, 1554–1573. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  83. Oh, S.; Suganuma, T.; Gogol, M.M.; Workman, J.L. Histone H3 threonine 11 phosphorylation by Sch9 and CK2 regulates chronological lifespan by controlling the nutritional stress response. Elife 2018, 7. [Google Scholar] [CrossRef] [PubMed]
  84. Maleszewska, M.; Mawer, J.S.P.; Tessarz, P. Histone Modifications in Ageing and Lifespan Regulation. Curr. Mol. Biol. Rep. 2016, 2, 26–35. [Google Scholar] [CrossRef] [Green Version]
  85. Lee, J.-S.; Shukla, A.; Schneider, J.; Swanson, S.K.; Washburn, M.P.; Florens, L.; Bhaumik, S.R.; Shilatifard, A. Histone crosstalk between H2B monoubiquitination and H3 methylation mediated by COMPASS. Cell 2007, 131, 1084–1096. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  86. Cao, J.; Yan, Q. Histone ubiquitination and deubiquitination in transcription, DNA damage response, and cancer. Front. Oncol. 2012, 2, 26. [Google Scholar] [CrossRef] [Green Version]
  87. Yang, L.; Ma, Z.; Wang, H.; Niu, K.; Cao, Y.; Le, S.; Geng, Y.; Yang, B.; Gao, F.; Chen, Z.; et al. Ubiquitylome study identifies increased histone 2A ubiquitylation as an evolutionarily conserved aging biomarker. Nat. Commun. 2019, 10, 2191. [Google Scholar] [CrossRef] [Green Version]
  88. Gao, Z.; Xu, M.S.; Barnett, T.L.; Xu, C.W. Resveratrol induces cellular senescence with attenuated mono-ubiquitination of histone H2B in glioma cells. Biochem. Biophys. Res. Commun. 2011, 407, 271–276. [Google Scholar] [CrossRef]
  89. Bushati, N.; Cohen, S.M. microRNA functions. Annu. Rev. Cell Dev. Biol. 2007, 23, 175–205. [Google Scholar] [CrossRef]
  90. Kinser, H.E.; Pincus, Z. MicroRNAs as modulators of longevity and the aging process. Hum. Genet. 2020, 139, 291–308. [Google Scholar] [CrossRef]
  91. Lai, W.-F.; Lin, M.; Wong, W.-T. Tackling Aging by Using miRNA as a Target and a Tool. Trends Mol. Med. 2019, 25, 673–684. [Google Scholar] [CrossRef] [PubMed]
  92. Kozomara, A.; Griffiths-Jones, S. miRBase: Annotating high confidence microRNAs using deep sequencing data. Nucleic Acids Res. 2014, 42, D68–D73. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  93. Lewis, B.P.; Burge, C.B.; Bartel, D.P. Conserved seed pairing, often flanked by adenosines, indicates that thousands of human genes are microRNA targets. Cell 2005, 120, 15–20. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  94. Boehm, M.; Slack, F. A developmental timing microRNA and its target regulate life span in C. elegans. Science 2005, 310, 1954–1957. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  95. Chawla, G.; Deosthale, P.; Childress, S.; Wu, Y.-C.; Sokol, N.S. A let-7-to-miR-125 MicroRNA Switch Regulates Neuronal Integrity and Lifespan in Drosophila. PLoS Genet. 2016, 12, e1006247. [Google Scholar] [CrossRef] [PubMed]
  96. Huan, T.; Chen, G.; Liu, C.; Bhattacharya, A.; Rong, J.; Chen, B.H.; Seshadri, S.; Tanriverdi, K.; Freedman, J.E.; Larson, M.G.; et al. Age-associated microRNA expression in human peripheral blood is associated with all-cause mortality and age-related traits. Aging Cell 2018, 17. [Google Scholar] [CrossRef]
  97. Hanahan, D.; Weinberg, R.A. The hallmarks of cancer. Cell 2000, 100, 57–70. [Google Scholar] [CrossRef] [Green Version]
  98. Hanahan, D.; Weinberg, R.A. Hallmarks of cancer: The next generation. Cell 2011, 144, 646–674. [Google Scholar] [CrossRef] [Green Version]
  99. Flavahan, W.A.; Gaskell, E.; Bernstein, B.E. Epigenetic plasticity and the hallmarks of cancer. Science 2017, 357, eaal2380. [Google Scholar] [CrossRef] [Green Version]
  100. Nebbioso, A.; Tambaro, F.P.; Dell’Aversana, C.; Altucci, L. Cancer epigenetics: Moving forward. PLoS Genet. 2018, 14, e1007362. [Google Scholar] [CrossRef] [Green Version]
  101. 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] [PubMed]
  102. Cancer Genome Atlas Network. Comprehensive molecular portraits of human breast tumours. Nature 2012, 490, 61–70. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  103. Karsli-Ceppioglu, S.; Dagdemir, A.; Judes, G.; Ngollo, M.; Penault-Llorca, F.; Pajon, A.; Bignon, Y.-J.; Bernard-Gallon, D. Epigenetic mechanisms of breast cancer: An update of the current knowledge. Epigenomics 2014, 6, 651–664. [Google Scholar] [CrossRef] [PubMed]
  104. Yegnasubramanian, S.; Kowalski, J.; Gonzalgo, M.L.; Zahurak, M.; Piantadosi, S.; Walsh, P.C.; Bova, G.S.; de Marzo, A.M.; Isaacs, W.B.; Nelson, W.G. Hypermethylation of CpG islands in primary and metastatic human prostate cancer. Cancer Res. 2004, 64, 1975–1986. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  105. Wu, Y.; Sarkissyan, M.; Vadgama, J.V. Epigenetics in Breast and Prostate Cancer. Methods Mol. Biol. 2015, 1238, 425–466. [Google Scholar] [CrossRef] [Green Version]
  106. Xu, W.; Yang, H.; Liu, Y.; Yang, Y.; Wang, P.; Kim, S.-H.; Ito, S.; Yang, C.; Wang, P.; Xiao, M.-T.; et al. Oncometabolite 2-hydroxyglutarate is a competitive inhibitor of α-ketoglutarate-dependent dioxygenases. Cancer Cell 2011, 19, 17–30. [Google Scholar] [CrossRef] [Green Version]
  107. Su, R.; Dong, L.; Li, C.; Nachtergaele, S.; Wunderlich, M.; Qing, Y.; Deng, X.; Wang, Y.; Weng, X.; Hu, C.; et al. R-2HG Exhibits Anti-tumor Activity by Targeting FTO/m6A/MYC/CEBPA Signaling. Cell 2018, 172, 90–105.e23. [Google Scholar] [CrossRef] [Green Version]
  108. Winters, A.C.; Bernt, K.M. MLL-Rearranged Leukemias-An Update on Science and Clinical Approaches. Front. Pediatr. 2017, 5, 4. [Google Scholar] [CrossRef] [Green Version]
  109. 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] [Green Version]
  110. Cheng, G.; Liu, F.; Asai, T.; Lai, F.; Man, N.; Xu, H.; Chen, S.; Greenblatt, S.; Hamard, P.-J.; Ando, K.; et al. Loss of p300 Accelerates MDS-associated Leukemogenesis. Leukemia 2016, 31, 1382–1390. [Google Scholar] [CrossRef] [Green Version]
  111. Picaud, S.; Fedorov, O.; Thanasopoulou, A.; Leonards, K.; Jones, K.; Meier, J.; Olzscha, H.; Monteiro, O.; Martin, S.; Philpott, M.; et al. Generation of a Selective Small Molecule Inhibitor of the CBP/p300 Bromodomain for Leukemia Therapy. Cancer Res. 2015, 75, 5106–5119. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  112. Comet, I.; Riising, E.M.; Leblanc, B.; Helin, K. Maintaining cell identity: PRC2-mediated regulation of transcription and cancer. Nat. Rev. Cancer 2016, 16, 803–810. [Google Scholar] [CrossRef] [PubMed]
  113. McCabe, M.T.; Ott, H.M.; Ganji, G.; Korenchuk, S.; Thompson, C.; van Aller, G.S.; Liu, Y.; Graves, A.P.; Della Pietra, A.; Diaz, E.; et al. EZH2 inhibition as a therapeutic strategy for lymphoma with EZH2-activating mutations. Nature 2012, 492, 108–112. [Google Scholar] [CrossRef] [PubMed]
  114. Sashida, G.; Iwama, A. Multifaceted role of the polycomb-group gene EZH2 in hematological malignancies. Int. J. Hematol. 2017, 105, 23–30. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  115. Mozhui, K.; Pandey, A.K. Conserved effect of aging on DNA methylation and association with EZH2 polycomb protein in mice and humans. Mech. Ageing Dev. 2017, 162, 27–37. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  116. Singh, A.A.; Mandoli, A.; Prange, K.H.M.; Laakso, M.; Martens, J.H.A. AML associated oncofusion proteins PML-RARA, AML1-ETO and CBFB-MYH11 target RUNX/ETS-factor binding sites to modulate H3ac levels and drive leukemogenesis. Oncotarget 2017, 8, 12855–12865. [Google Scholar] [CrossRef] [PubMed]
  117. Skoda, R.C.; Schwaller, J. Dual roles of EZH2 in acute myeloid leukemia. J. Exp. Med. 2019, 216, 725–727. [Google Scholar] [CrossRef] [Green Version]
  118. Okugawa, Y.; Grady, W.M.; Goel, A. Epigenetic Alterations in Colorectal Cancer: Emerging Biomarkers. Gastroenterology 2015, 149, 1204–1225.e12. [Google Scholar] [CrossRef] [Green Version]
  119. Lao, V.V.; Grady, W.M. Epigenetics and colorectal cancer. Nat. Rev. Gastroenterol. Hepatol. 2011, 8, 686–700. [Google Scholar] [CrossRef]
  120. Guo, Y.; Wang, M.; Jia, X.; Zhu, H.; Zhi, Y.; Yuan, L. Wnt signaling pathway upregulates DNMT1 to trigger NHERF1 promoter hypermethylation in colon cancer. Oncol. Rep. 2018, 40, 1165–1173. [Google Scholar] [CrossRef]
  121. Song, J.; Du, Z.; Ravasz, M.; Dong, B.; Wang, Z.; Ewing, R.M. A Protein Interaction between β-Catenin and Dnmt1 Regulates Wnt Signaling and DNA Methylation in Colorectal Cancer Cells. Mol. Cancer Res. 2015, 13, 969–981. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  122. Yang, Z.-H.; Dang, Y.-Q.; Ji, G. Role of epigenetics in transformation of inflammation into colorectal cancer. World J. Gastroenterol. 2019, 25, 2863–2877. [Google Scholar] [CrossRef] [PubMed]
  123. Foran, E.; Garrity-Park, M.M.; Mureau, C.; Newell, J.; Smyrk, T.C.; Limburg, P.J.; Egan, L.J. Upregulation of DNA methyltransferase-mediated gene silencing, anchorage-independent growth, and migration of colon cancer cells by interleukin-6. Mol. Cancer Res. 2010, 8, 471–481. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  124. Fearon, E.R.; Vogelstein, B. A genetic model for colorectal tumorigenesis. Cell 1990, 61, 759–767. [Google Scholar] [CrossRef]
  125. Ciccarone, F.; Malavolta, M.; Calabrese, R.; Guastafierro, T.; Bacalini, M.G.; Reale, A.; Franceschi, C.; Capri, M.; Hervonen, A.; Hurme, M.; et al. Age-dependent expression of DNMT1 and DNMT3B in PBMCs from a large European population enrolled in the MARK-AGE study. Aging Cell 2016, 15, 755–765. [Google Scholar] [CrossRef]
  126. Yung, R.; Ray, D.; Eisenbraun, J.K.; Deng, C.; Attwood, J.; Eisenbraun, M.D.; Johnson, K.; Miller, R.A.; Hanash, S.; Richardson, B. Unexpected effects of a heterozygous dnmt1 null mutation on age-dependent DNA hypomethylation and autoimmunity. J. Gerontol. A Biol. Sci. Med. Sci. 2001, 56, B268–B276. [Google Scholar] [CrossRef] [Green Version]
  127. Laird, P.W.; Jackson-Grusby, L.; Fazeli, A.; Dickinson, S.L.; Jung, W.E.; Li, E.; Weinberg, R.A.; Jaenisch, R. Suppression of intestinal neoplasia by DNA hypomethylation. Cell 1995, 81, 197–205. [Google Scholar] [CrossRef] [Green Version]
  128. Gerecke, C.; Scholtka, B.; Löwenstein, Y.; Fait, I.; Gottschalk, U.; Rogoll, D.; Melcher, R.; Kleuser, B. Hypermethylation of ITGA4, TFPI2 and VIMENTIN promoters is increased in inflamed colon tissue: Putative risk markers for colitis-associated cancer. J. Cancer Res. Clin. Oncol. 2015, 141, 2097–2107. [Google Scholar] [CrossRef]
  129. Requena, D.O.; Garcia-Buitrago, M. Molecular Insights Into Colorectal Carcinoma. Arch. Med. Res. 2020, 51, 839–844. [Google Scholar] [CrossRef]
  130. Fleisher, A.S.; Esteller, M.; Harpaz, N.; Leytin, A.; Rashid, A.; Xu, Y.; Liang, J.; Stine, O.C.; Yin, J.; Zou, T.T.; et al. Microsatellite instability in inflammatory bowel disease-associated neoplastic lesions is associated with hypermethylation and diminished expression of the DNA mismatch repair gene, hMLH1. Cancer Res. 2000, 60, 4864–4868. [Google Scholar]
  131. Arnold, C.N.; Goel, A.; Boland, C.R. Role of hMLH1 promoter hypermethylation in drug resistance to 5-fluorouracil in colorectal cancer cell lines. Int. J. Cancer 2003, 106, 66–73. [Google Scholar] [CrossRef] [PubMed]
  132. Dienstmann, R.; Vermeulen, L.; Guinney, J.; Kopetz, S.; Tejpar, S.; Tabernero, J. Consensus molecular subtypes and the evolution of precision medicine in colorectal cancer. Nat. Rev. Cancer 2017, 17, 79–92. [Google Scholar] [CrossRef] [PubMed]
  133. Franceschi, C.; Bonafè, M.; Valensin, S.; Olivieri, F.; de Luca, M.; Ottaviani, E.; de Benedictis, G. Inflamm-aging. An evolutionary perspective on immunosenescence. Ann. N. Y. Acad. Sci. 2000, 908, 244–254. [Google Scholar] [CrossRef] [PubMed]
  134. Fulop, T.; Larbi, A.; Dupuis, G.; Le Page, A.; Frost, E.H.; Cohen, A.A.; Witkowski, J.M.; Franceschi, C. Immunosenescence and Inflamm-Aging As Two Sides of the Same Coin: Friends or Foes? Front. Immunol. 2017, 8, 1960. [Google Scholar] [CrossRef] [Green Version]
  135. Sullivan, K.E.; Reddy, A.B.M.; Dietzmann, K.; Suriano, A.R.; Kocieda, V.P.; Stewart, M.; Bhatia, M. Epigenetic regulation of tumor necrosis factor alpha. Mol. Cell. Biol. 2007, 27, 5147–5160. [Google Scholar] [CrossRef] [Green Version]
  136. Ray, D.; Yung, R. Immune Senescence, Epigenetics and Autoimmunity. Clin. Immunol. 2018, 196, 59–63. [Google Scholar] [CrossRef]
  137. Wang, H.; Feng, H.; Sun, J.; Zhou, Y.; Zhu, G.; Wu, S.; Bao, W. Age-associated changes in DNA methylation and expression of the TNFα gene in pigs. Genes Genet. Syst. 2018, 93, 191–198. [Google Scholar] [CrossRef] [Green Version]
  138. Tilstra, J.S.; Clauson, C.L.; Niedernhofer, L.J.; Robbins, P.D. NF-κB in Aging and Disease. Aging Dis. 2011, 2, 449–465. [Google Scholar]
  139. Li, Y.; Reddy, M.A.; Miao, F.; Shanmugam, N.; Yee, J.-K.; Hawkins, D.; Ren, B.; Natarajan, R. Role of the histone H3 lysine 4 methyltransferase, SET7/9, in the regulation of NF-kappaB-dependent inflammatory genes. Relevance to diabetes and inflammation. J. Biol. Chem. 2008, 283, 26771–26781. [Google Scholar] [CrossRef] [Green Version]
  140. Wang, Y.; Wang, L.; Wen, X.; Hao, D.; Zhang, N.; He, G.; Jiang, X. NF-κB signaling in skin aging. Mech. Ageing Dev. 2019, 184, 111160. [Google Scholar] [CrossRef]
  141. Choi, Y.J.; Moon, K.M.; Chung, K.W.; Jeong, J.W.; Park, D.; Kim, D.H.; Yu, B.P.; Chung, H.Y. The underlying mechanism of proinflammatory NF-κB activation by the mTORC2/Akt/IKKα pathway during skin aging. Oncotarget 2016, 7, 52685–52694. [Google Scholar] [CrossRef] [PubMed]
  142. Johnsen, S.A. The enigmatic role of H2Bub1 in cancer. FEBS Lett. 2012, 586, 1592–1601. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  143. Tarcic, O.; Pateras, I.S.; Cooks, T.; Shema, E.; Kanterman, J.; Ashkenazi, H.; Boocholez, H.; Hubert, A.; Rotkopf, R.; Baniyash, M.; et al. RNF20 Links Histone H2B Ubiquitylation with Inflammation and Inflammation-Associated Cancer. Cell Rep. 2016, 14, 1462–1476. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  144. Kosinsky, R.L.; Chua, R.L.; Qui, M.; Saul, D.; Mehlich, D.; Ströbel, P.; Schildhaus, H.-U.; Wegwitz, F.; Faubion, W.A.; Johnsen, S.A. Loss of RNF40 Decreases NF-κB Activity in Colorectal Cancer Cells and Reduces Colitis Burden in Mice. J. Crohns. Colitis 2019, 13, 362–373. [Google Scholar] [CrossRef]
  145. Yuan, J.; Zhang, F.; Niu, R. Multiple regulation pathways and pivotal biological functions of STAT3 in cancer. Sci. Rep. 2015, 5, 17663. [Google Scholar] [CrossRef] [Green Version]
  146. Zimmers, T.A.; Fishel, M.L.; Bonetto, A. STAT3 in the Systemic Inflammation of Cancer Cachexia. Semin. Cell Dev. Biol. 2016, 54, 28–41. [Google Scholar] [CrossRef] [Green Version]
  147. Takeshima, H.; Niwa, T.; Yamashita, S.; Takamura-Enya, T.; Iida, N.; Wakabayashi, M.; Nanjo, S.; Abe, M.; Sugiyama, T.; Kim, Y.-J.; et al. TET repression and increased DNMT activity synergistically induce aberrant DNA methylation. J. Clin. Investig. 2020, 130, 5370–5379. [Google Scholar] [CrossRef]
  148. Jiang, S.; Yan, W.; Wang, S.E.; Baltimore, D. Dual mechanisms of posttranscriptional regulation of Tet2 by Let-7 microRNA in macrophages. Proc. Natl. Acad. Sci. USA 2019, 116, 12416–12421. [Google Scholar] [CrossRef] [Green Version]
  149. Horvath, S.; Erhart, W.; Brosch, M.; Ammerpohl, O.; von Schönfels, W.; Ahrens, M.; Heits, N.; Bell, J.T.; Tsai, P.-C.; Spector, T.D.; et al. Obesity accelerates epigenetic aging of human liver. Proc. Natl. Acad. Sci. USA 2014, 111, 15538–15543. [Google Scholar] [CrossRef] [Green Version]
  150. Hanson, B.M.; Tao, X.; Zhan, Y.; Jenkins, T.G.; Morin, S.J.; Scott, R.T.; Seli, E.U. Young women with poor ovarian response exhibit epigenetic age acceleration based on evaluation of white blood cells using a DNA methylation-derived age prediction model. Hum. Reprod. 2020, 35, 2579–2588. [Google Scholar] [CrossRef]
  151. Morin, S.J.; Tao, X.; Marin, D.; Zhan, Y.; Landis, J.; Bedard, J.; Scott, R.T.; Seli, E. DNA methylation-based age prediction and telomere length in white blood cells and cumulus cells of infertile women with normal or poor response to ovarian stimulation. Aging (Albany NY) 2018, 10, 3761–3773. [Google Scholar] [CrossRef] [PubMed]
  152. Marioni, R.E.; Shah, S.; McRae, A.F.; Chen, B.H.; Colicino, E.; Harris, S.E.; Gibson, J.; Henders, A.K.; Redmond, P.; Cox, S.R.; et al. DNA methylation age of blood predicts all-cause mortality in later life. Genome Biol. 2015, 16, 25. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  153. Johnson, P.L.F.; Goronzy, J.J.; Antia, R. A population biological approach to understanding the maintenance and loss of the T-cell repertoire during aging. Immunology 2014, 142, 167–175. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  154. Stevenson, A.J.; McCartney, D.L.; Harris, S.E.; Taylor, A.M.; Redmond, P.; Starr, J.M.; Zhang, Q.; McRae, A.F.; Wray, N.R.; Spires-Jones, T.L.; et al. Trajectories of inflammatory biomarkers over the eighth decade and their associations with immune cell profiles and epigenetic ageing. Clin. Epigenetics 2018, 10, 159. [Google Scholar] [CrossRef] [Green Version]
  155. Quinn, K.M.; Fox, A.; Harland, K.L.; Russ, B.E.; Li, J.; Nguyen, T.H.O.; Loh, L.; Olshanksy, M.; Naeem, H.; Tsyganov, K.; et al. Age-Related Decline in Primary CD8+ T Cell Responses Is Associated with the Development of Senescence in Virtual Memory CD8+ T Cells. Cell Rep. 2018, 23, 3512–3524. [Google Scholar] [CrossRef]
  156. Lau, E.Y.M.; Carroll, E.C.; Callender, L.A.; Hood, G.A.; Berryman, V.; Pattrick, M.; Finer, S.; Hitman, G.A.; Ackland, G.L.; Henson, S.M. Type 2 diabetes is associated with the accumulation of senescent T cells. Clin. Exp. Immunol. 2019, 197, 205–213. [Google Scholar] [CrossRef] [Green Version]
  157. Bektas, A.; Schurman, S.H.; Sen, R.; Ferrucci, L. Human T cell immunosenescence and inflammation in aging. J. Leukoc. Biol. 2017, 102, 977–988. [Google Scholar] [CrossRef] [Green Version]
  158. Surace, A.E.A.; Hedrich, C.M. The Role of Epigenetics in Autoimmune/Inflammatory Disease. Front. Immunol. 2019, 10, 1525. [Google Scholar] [CrossRef] [Green Version]
  159. Mikami, N.; Kawakami, R.; Sakaguchi, S. New Treg cell-based therapies of autoimmune diseases: Towards antigen-specific immune suppression. Curr. Opin. Immunol. 2020, 67, 36–41. [Google Scholar] [CrossRef]
  160. Dozmorov, M.G.; Coit, P.; Maksimowicz-McKinnon, K.; Sawalha, A.H. Age-associated DNA methylation changes in naive CD4+ T cells suggest an evolving autoimmune epigenotype in aging T cells. Epigenomics 2017, 9, 429–445. [Google Scholar] [CrossRef] [Green Version]
  161. Hofmann, S.R.; Kubasch, A.S.; Ioannidis, C.; Rösen-Wolff, A.; Girschick, H.J.; Morbach, H.; Hedrich, C.M. Altered expression of IL-10 family cytokines in monocytes from CRMO patients result in enhanced IL-1β expression and release. Clin. Immunol. 2015, 161, 300–307. [Google Scholar] [CrossRef] [PubMed]
  162. Wu, Z.-X.; Cao, L.; Li, X.-W.; Jiang, W.; Li, X.-Y.; Xu, J.; Wang, F.; Chen, G.-H. Accelerated Deficits of Spatial Learning and Memory Resulting From Prenatal Inflammatory Insult Are Correlated With Abnormal Phosphorylation and Methylation of Histone 3 in CD-1 Mice. Front. Aging Neurosci. 2019, 11. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  163. Cakouros, D.; Gronthos, S. Epigenetic Regulators of Mesenchymal Stem/Stromal Cell Lineage Determination. Curr. Osteoporos. Rep. 2020, 18, 597–605. [Google Scholar] [CrossRef] [PubMed]
  164. Sepulveda, H.; Villagra, A.; Montecino, M. Tet-Mediated DNA Demethylation Is Required for SWI/SNF-Dependent Chromatin Remodeling and Histone-Modifying Activities That Trigger Expression of the Sp7 Osteoblast Master Gene during Mesenchymal Lineage Commitment. Mol. Cell. Biol. 2017, 37, e00177-17. [Google Scholar] [CrossRef] [Green Version]
  165. Wang, R.; Wang, Y.; Zhu, L.; Liu, Y.; Li, W. Epigenetic Regulation in Mesenchymal Stem Cell Aging and Differentiation and Osteoporosis. Stem Cells Int. 2020, 2020, 1–17. [Google Scholar] [CrossRef]
  166. Bork, S.; Pfister, S.; Witt, H.; Horn, P.; Korn, B.; Ho, A.D.; Wagner, W. DNA methylation pattern changes upon long-term culture and aging of human mesenchymal stromal cells. Aging Cell 2010, 9, 54–63. [Google Scholar] [CrossRef] [Green Version]
  167. Cakouros, D.; Hemming, S.; Gronthos, K.; Liu, R.; Zannettino, A.; Shi, S.; Gronthos, S. Specific functions of TET1 and TET2 in regulating mesenchymal cell lineage determination. Epigenetics Chromatin 2019, 12, 3. [Google Scholar] [CrossRef]
  168. Yang, R.; Yu, T.; Kou, X.; Gao, X.; Chen, C.; Liu, D.; Zhou, Y.; Shi, S. Tet1 and Tet2 maintain mesenchymal stem cell homeostasis via demethylation of the P2rX7 promoter. Nat. Commun. 2018, 9, 2143. [Google Scholar] [CrossRef]
  169. Ko, N.-Y.; Chen, L.-R.; Chen, K.-H. The Role of Micro RNA and Long-Non-Coding RNA in Osteoporosis. Int. J. Mol. Sci. 2020, 21, 4886. [Google Scholar] [CrossRef]
  170. Gontier, G.; Iyer, M.; Shea, J.M.; Bieri, G.; Wheatley, E.G.; Ramalho-Santos, M.; Villeda, S.A. Tet2 Rescues Age-Related Regenerative Decline and Enhances Cognitive Function in the Adult Mouse Brain. Cell Rep. 2018, 22, 1974–1981. [Google Scholar] [CrossRef] [Green Version]
  171. Ren, J.; Huang, D.; Li, R.; Wang, W.; Zhou, C. Control of mesenchymal stem cell biology by histone modifications. Cell Biosci. 2020, 10, 1–15. [Google Scholar] [CrossRef] [PubMed]
  172. Jing, H.; Liao, L.; An, Y.; Su, X.; Liu, S.; Shuai, Y.; Zhang, X.; Jin, Y. Suppression of EZH2 Prevents the Shift of Osteoporotic MSC Fate to Adipocyte and Enhances Bone Formation During Osteoporosis. Mol. Ther. 2016, 24, 217–229. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  173. Ma, J.; Lin, X.; Chen, C.; Li, S.; Zhang, S.; Chen, Z.; Li, D.; Zhao, F.; Yang, C.; Yin, C.; et al. Circulating miR-181c-5p and miR-497-5p Are Potential Biomarkers for Prognosis and Diagnosis of Osteoporosis. J. Clin. Endocrinol. Metab. 2020, 105, 1445–1460. [Google Scholar] [CrossRef] [PubMed]
  174. Bone, H.G.; Wagman, R.B.; Brandi, M.L.; Brown, J.P.; Chapurlat, R.; Cummings, S.R.; Czerwiński, E.; Fahrleitner-Pammer, A.; Kendler, D.L.; Lippuner, K.; et al. 10 years of denosumab treatment in postmenopausal women with osteoporosis: Results from the phase 3 randomised FREEDOM trial and open-label extension. Lancet Diabetes Endocrinol. 2017, 5, 513–523. [Google Scholar] [CrossRef]
  175. de Nigris, F.; Ruosi, C.; Colella, G.; Napoli, C. Epigenetic therapies of osteoporosis. Bone 2020, 142, 115680. [Google Scholar] [CrossRef]
  176. Dolinar, A.; Ravnik-Glavač, M.; Glavač, D. Epigenetic mechanisms in amyotrophic lateral sclerosis: A short review. Mech. Ageing Dev. 2018, 174, 103–110. [Google Scholar] [CrossRef]
  177. Ebbert, M.T.W.; Ross, C.A.; Pregent, L.J.; Lank, R.J.; Zhang, C.; Katzman, R.B.; Jansen-West, K.; Song, Y.; Da Rocha, E.L.; Palmucci, C.; et al. Conserved DNA methylation combined with differential frontal cortex and cerebellar expression distinguishes C9orf72-associated and sporadic ALS, and implicates SERPINA1 in disease. Acta Neuropathol. 2017, 134, 715–728. [Google Scholar] [CrossRef]
  178. Tecalco-Cruz, A.C.; Ramírez-Jarquín, J.O.; Alvarez-Sánchez, M.E.; Zepeda-Cervantes, J. Epigenetic basis of Alzheimer disease. World J. Biol. Chem. 2020, 11, 62–75. [Google Scholar] [CrossRef]
  179. Saez-Atienzar, S.; Masliah, E. Cellular senescence and Alzheimer disease: The egg and the chicken scenario. Nat. Rev. Neurosci. 2020, 21, 433–444. [Google Scholar] [CrossRef]
  180. Valenta, T.; Lukas, J.; Doubravska, L.; Fafilek, B.; Korinek, V. HIC1 attenuates Wnt signaling by recruitment of TCF-4 and beta-catenin to the nuclear bodies. EMBO J. 2006, 25, 2326–2337. [Google Scholar] [CrossRef] [Green Version]
  181. Nativio, R.; Donahue, G.; Berson, A.; Lan, Y.; Amlie-Wolf, A.; Tuzer, F.; Toledo, J.B.; Gosai, S.J.; Gregory, B.D.; Torres, C.; et al. Dysregulation of the epigenetic landscape of normal aging in Alzheimer’s disease. Nat. Neurosci. 2018, 21, 497–505. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  182. Chen, W.Y.; Wang, D.H.; Yen, R.C.; Luo, J.; Gu, W.; Baylin, S.B. Tumor suppressor HIC1 directly regulates SIRT1 to modulate p53-dependent DNA-damage responses. Cell 2005, 123, 437–448. [Google Scholar] [CrossRef] [PubMed]
  183. Lu, T.; Aron, L.; Zullo, J.; Pan, Y.; Kim, H.; Chen, Y.; Yang, T.-H.; Kim, H.-M.; Drake, D.; Liu, X.S.; et al. Addendum: REST and stress resistance in ageing and Alzheimer’s disease. Nature 2016, 540, 470. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  184. Lu, T.; Aron, L.; Zullo, J.; Pan, Y.; Kim, H.; Chen, Y.; Yang, T.-H.; Kim, H.-M.; Drake, D.; Liu, X.S.; et al. REST and Stress Resistance in Aging and Alzheimer’s Disease. Nature 2014, 507, 448–454. [Google Scholar] [CrossRef] [Green Version]
  185. Hwang, J.-Y.; Zukin, R.S. REST, a master transcriptional regulator in neurodegenerative disease. Curr. Opin. Neurobiol. 2018, 48, 193–200. [Google Scholar] [CrossRef]
  186. Hou, T.-Y.; Zhou, Y.; Zhu, L.-S.; Wang, X.; Pang, P.; Wang, D.-Q.; Liuyang, Z.-Y.; Man, H.; Lu, Y.; Zhu, L.-Q.; et al. Correcting abnormalities in miR-124/PTPN1 signaling rescues tau pathology in Alzheimer’s disease. J. Neurochem. 2020, 154, 441–457. [Google Scholar] [CrossRef]
  187. Williams, J.; Smith, F.; Kumar, S.; Vijayan, M.; Reddy, P.H. Are microRNAs true sensors of ageing and cellular senescence? Ageing Res. Rev. 2017, 35, 350–363. [Google Scholar] [CrossRef] [Green Version]
  188. Kumar, P.; Dezso, Z.; MacKenzie, C.; Oestreicher, J.; Agoulnik, S.; Byrne, M.; Bernier, F.; Yanagimachi, M.; Aoshima, K.; Oda, Y. Circulating miRNA biomarkers for Alzheimer’s disease. PLoS ONE 2013, 8, e69807. [Google Scholar] [CrossRef]
  189. Labbé, C.; Lorenzo-Betancor, O.; Ross, O.A. Epigenetic regulation in Parkinson’s disease. Acta Neuropathol. 2016, 132, 515–530. [Google Scholar] [CrossRef]
  190. Jiang, K.; Rocha, S.; Westling, A.; Kesarimangalam, S.; Dorfman, K.D.; Wittung-Stafshede, P.; Westerlund, F. Alpha-Synuclein Modulates the Physical Properties of DNA. Chemistry 2018, 24, 15685–15690. [Google Scholar] [CrossRef]
  191. Sharma, A.; Osato, N.; Liu, H.; Asthana, S.; Dakal, T.C.; Ambrosini, G.; Bucher, P.; Schmitt, I.; Wüllner, U. Common genetic variants associated with Parkinson’s disease display widespread signature of epigenetic plasticity. Sci. Rep. 2019, 9, 1–19. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  192. Vermunt, M.W.; Reinink, P.; Korving, J.; de Bruijn, E.; Creyghton, P.M.; Basak, O.; Geeven, G.; Toonen, P.W.; Lansu, N.; Meunier, C.; et al. Large-scale identification of coregulated enhancer networks in the adult human brain. Cell Rep. 2014, 9, 767–779. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  193. Chen, X.; Scholz, S.W. Identification of new α-synuclein regulator by nontraditional drug development pipeline. Mov. Disord. 2018, 33, 402. [Google Scholar] [CrossRef] [PubMed]
  194. Mittal, S.; Bjørnevik, K.; Soon Im, D.; Flierl, A.; Dong, X.; Locascio, J.J.; Abo, K.M.; Long, E.; Jin, M.; Xu, B.; et al. β2-Adrenoreceptor is a Regulator of the α-Synuclein Gene Driving Risk of Parkinson’s Disease. Science 2017, 357, 891–898. [Google Scholar] [CrossRef] [Green Version]
  195. Esteves, A.R.; Palma, A.M.; Gomes, R.; Santos, D.; Silva, D.F.; Cardoso, S.M. Acetylation as a major determinant to microtubule-dependent autophagy: Relevance to Alzheimer’s and Parkinson disease pathology. Biochim. Biophys. Acta Mol. Basis Dis. 2019, 1865, 2008–2023. [Google Scholar] [CrossRef]
  196. Gebremedhin, K.G.; Rademacher, D.J. Histone H3 Acetylation in the Postmortem Parkinson’s Disease Primary Motor Cortex. Neurosci. Lett. 2016, 627, 121–125. [Google Scholar] [CrossRef] [Green Version]
  197. Kontopoulos, E.; Parvin, J.D.; Feany, M.B. Alpha-synuclein acts in the nucleus to inhibit histone acetylation and promote neurotoxicity. Hum. Mol. Genet. 2006, 15, 3012–3023. [Google Scholar] [CrossRef] [Green Version]
  198. Sun, L.; Zhang, J.; Chen, W.; Chen, Y.; Zhang, X.; Yang, M.; Xiao, M.; Ma, F.; Yao, Y.; Ye, M.; et al. Attenuation of epigenetic regulator SMARCA4 and ERK-ETS signaling suppresses aging-related dopaminergic degeneration. Aging Cell 2020, 19, e13210. [Google Scholar] [CrossRef]
  199. Kahn, S.E.; Hull, R.L.; Utzschneider, K.M. Mechanisms linking obesity to insulin resistance and type 2 diabetes. Nature 2006, 444, 840–846. [Google Scholar] [CrossRef]
  200. Desiderio, A.; Spinelli, R.; Ciccarelli, M.; Nigro, C.; Miele, C.; Beguinot, F.; Raciti, G.A. Epigenetics: Spotlight on type 2 diabetes and obesity. J. Endocrinol. Investig. 2016, 39, 1095–1103. [Google Scholar] [CrossRef]
  201. Guzik, T.J.; Cosentino, F. Epigenetics and Immunometabolism in Diabetes and Aging. Antioxid. Redox Signal. 2018, 29, 257–274. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  202. Ling, C.; Rönn, T. Epigenetics in Human Obesity and Type 2 Diabetes. Cell Metab. 2019, 29, 1028–1044. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  203. Heijmans, B.T.; Tobi, E.W.; Stein, A.D.; Putter, H.; Blauw, G.J.; Susser, E.S.; Slagboom, P.E.; Lumey, L.H. Persistent epigenetic differences associated with prenatal exposure to famine in humans. Proc. Natl. Acad. Sci. USA 2008, 105, 17046–17049. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  204. Lumey, L.H.; Stein, A.D.; Kahn, H.S.; van der Pal-de Bruin, K.M.; Blauw, G.J.; Zybert, P.A.; Susser, E.S. Cohort profile: The Dutch Hunger Winter families study. Int. J. Epidemiol. 2007, 36, 1196–1204. [Google Scholar] [CrossRef] [Green Version]
  205. Li, Y.; Tollefsbol, T.O. p16INK4a Suppression by Glucose Restriction Contributes to Human Cellular Lifespan Extension through SIRT1-Mediated Epigenetic and Genetic Mechanisms. PLoS ONE 2011, 6, e17421. [Google Scholar] [CrossRef]
  206. Daniel, M.; Tollefsbol, T.O. Epigenetic linkage of aging, cancer and nutrition. J. Exp. Biol. 2015, 218, 59–70. [Google Scholar] [CrossRef] [Green Version]
  207. Kapahi, P.; Kaeberlein, M.; Hansen, M. Dietary restriction and lifespan: Lessons from invertebrate models. Ageing Res. Rev. 2017, 39, 3–14. [Google Scholar] [CrossRef]
  208. Joseph, S.V.; Edirisinghe, I.; Burton-Freeman, B.M. Fruit Polyphenols: A Review of Anti-inflammatory Effects in Humans. Crit. Rev. Food Sci. Nutr. 2016, 56, 419–444. [Google Scholar] [CrossRef]
  209. Hou, X.; Rooklin, D.; Fang, H.; Zhang, Y. Resveratrol serves as a protein-substrate interaction stabilizer in human SIRT1 activation. Sci. Rep. 2016, 6, 38186. [Google Scholar] [CrossRef]
  210. Li, Y.-R.; Li, S.; Lin, C.-C. Effect of resveratrol and pterostilbene on aging and longevity. Biofactors 2018, 44, 69–82. [Google Scholar] [CrossRef]
  211. Yessenkyzy, A.; Saliev, T.; Zhanaliyeva, M.; Masoud, A.-R.; Umbayev, B.; Sergazy, S.; Krivykh, E.; Gulyayev, A.; Nurgozhin, T. Polyphenols as Caloric-Restriction Mimetics and Autophagy Inducers in Aging Research. Nutrients 2020, 12, 1344. [Google Scholar] [CrossRef] [PubMed]
  212. Almén, M.S.; Nilsson, E.K.; Jacobsson, J.A.; Kalnina, I.; Klovins, J.; Fredriksson, R.; Schiöth, H.B. Genome-wide analysis reveals DNA methylation markers that vary with both age and obesity. Gene 2014, 548, 61–67. [Google Scholar] [CrossRef] [PubMed]
  213. Hannon, E.; Knox, O.; Sugden, K.; Burrage, J.; Wong, C.C.Y.; Belsky, D.W.; Corcoran, D.L.; Arseneault, L.; Moffitt, T.E.; Caspi, A.; et al. Characterizing genetic and environmental influences on variable DNA methylation using monozygotic and dizygotic twins. PLoS Genet. 2018, 14, e1007544. [Google Scholar] [CrossRef] [PubMed]
  214. Molina-Serrano, D.; Kyriakou, D.; Kirmizis, A. Histone Modifications as an Intersection Between Diet and Longevity. Front. Genet. 2019, 10, 192. [Google Scholar] [CrossRef] [PubMed]
  215. Xu, X.; Su, S.; Barnes, V.A.; de Miguel, C.; Pollock, J.; Ownby, D.; Shi, H.; Zhu, H.; Snieder, H.; Wang, X. A genome-wide methylation study on obesity: Differential variability and differential methylation. Epigenetics 2013, 8, 522–533. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  216. Andreas, E.; Reid, M.; Zhang, W.; Moley, K.H. The effect of maternal high-fat/high-sugar diet on offspring oocytes and early embryo development. Mol. Hum. Reprod. 2019, 25, 717–728. [Google Scholar] [CrossRef]
  217. Zheng, S.; Li, Q.; Zhang, Y.; Balluff, Z.; Pan, Y.-X. Histone deacetylase 3 (HDAC3) participates in the transcriptional repression of the p16 (INK4a) gene in mammary gland of the female rat offspring exposed to an early-life high-fat diet. Epigenetics 2012, 7, 183–190. [Google Scholar] [CrossRef] [Green Version]
  218. Zhang, X.; Xu, G.B.; Zhou, D.; Pan, Y.-X. High-fat diet modifies expression of hepatic cellular senescence gene p16(INK4a) through chromatin modifications in adult male rats. Genes Nutr. 2018, 13, 6. [Google Scholar] [CrossRef]
  219. Jacobsen, S.C.; Brøns, C.; Bork-Jensen, J.; Ribel-Madsen, R.; Yang, B.; Lara, E.; Hall, E.; Calvanese, V.; Nilsson, E.; Jørgensen, S.W.; et al. Effects of short-term high-fat overfeeding on genome-wide DNA methylation in the skeletal muscle of healthy young men. Diabetologia 2012, 55, 3341–3349. [Google Scholar] [CrossRef] [Green Version]
  220. Hall, E.; Dekker Nitert, M.; Volkov, P.; Malmgren, S.; Mulder, H.; Bacos, K.; Ling, C. The effects of high glucose exposure on global gene expression and DNA methylation in human pancreatic islets. Mol. Cell. Endocrinol. 2018, 472, 57–67. [Google Scholar] [CrossRef]
  221. Sommese, L.; Benincasa, G.; Lanza, M.; Sorriento, A.; Schiano, C.; Lucchese, R.; Alfano, R.; Nicoletti, G.F.; Napoli, C. Novel epigenetic-sensitive clinical challenges both in type 1 and type 2 diabetes. J. Diabetes Complicat. 2018, 32, 1076–1084. [Google Scholar] [CrossRef] [PubMed]
  222. Sandovici, I.; Hammerle, C.M.; Cooper, W.N.; Smith, N.H.; Tarry-Adkins, J.L.; Dunmore, B.J.; Bauer, J.; Andrews, S.R.; Yeo, G.S.H.; Ozanne, S.E.; et al. Ageing is associated with molecular signatures of inflammation and type 2 diabetes in rat pancreatic islets. Diabetologia 2016, 59, 502–511. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  223. Dayeh, T.; Volkov, P.; Salö, S.; Hall, E.; Nilsson, E.; Olsson, A.H.; Kirkpatrick, C.L.; Wollheim, C.B.; Eliasson, L.; Rönn, T.; et al. Genome-wide DNA methylation analysis of human pancreatic islets from type 2 diabetic and non-diabetic donors identifies candidate genes that influence insulin secretion. PLoS Genet. 2014, 10, e1004160. [Google Scholar] [CrossRef] [PubMed]
  224. Baker, D.J.; Jin, F.; van Deursen, J.M. The yin and yang of the Cdkn2a locus in senescence and aging. Cell Cycle 2008, 7, 2795–2802. [Google Scholar] [CrossRef] [Green Version]
  225. Kong, Y.; Sharma, R.B.; Nwosu, B.U.; Alonso, L.C. Islet biology, the CDKN2A/B locus and type 2 diabetes risk. Diabetologia 2016, 59, 1579–1593. [Google Scholar] [CrossRef]
  226. Chen, H.; Gu, X.; Su, I.; Bottino, R.; Contreras, J.L.; Tarakhovsky, A.; Kim, S.K. Polycomb protein Ezh2 regulates pancreatic beta-cell Ink4a/Arf expression and regeneration in diabetes mellitus. Genes Dev. 2009, 23, 975–985. [Google Scholar] [CrossRef] [Green Version]
  227. Avrahami, D.; Li, C.; Zhang, J.; Schug, J.; Avrahami, R.; Rao, S.; Stadler, M.B.; Burger, L.; Schübeler, D.; Glaser, B.; et al. Aging-Dependent Demethylation of Regulatory Elements Correlates with Chromatin State and Improved β Cell Function. Cell Metab. 2015, 22, 619–632. [Google Scholar] [CrossRef] [Green Version]
  228. Zhang, H.; Pollin, T.I. Epigenetics Variation and Pathogenesis in Diabetes. Curr. Diab. Rep. 2018, 18, 121. [Google Scholar] [CrossRef]
  229. Miao, F.; Chen, Z.; Zhang, L.; Liu, Z.; Wu, X.; Yuan, Y.-C.; Natarajan, R. Profiles of epigenetic histone post-translational modifications at type 1 diabetes susceptible genes. J. Biol. Chem. 2012, 287, 16335–16345. [Google Scholar] [CrossRef] [Green Version]
  230. Rakyan, V.K.; Beyan, H.; Down, T.A.; Hawa, M.I.; Maslau, S.; Aden, D.; Daunay, A.; Busato, F.; Mein, C.A.; Manfras, B.; et al. Identification of type 1 diabetes-associated DNA methylation variable positions that precede disease diagnosis. PLoS Genet. 2011, 7, e1002300. [Google Scholar] [CrossRef]
  231. Vasu, S.; Kumano, K.; Darden, C.M.; Rahman, I.; Lawrence, M.C.; Naziruddin, B. MicroRNA Signatures as Future Biomarkers for Diagnosis of Diabetes States. Cells 2019, 8, 1533. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  232. Taheri, M.; Eghtedarian, R.; Ghafouri-Fard, S.; Omrani, M.D. Non-coding RNAs and type 2 diabetes mellitus. Arch. Physiol. Biochem. 2020, 1–10. [Google Scholar] [CrossRef] [PubMed]
  233. Kamalden, T.A.; Macgregor-Das, A.M.; Kannan, S.M.; Dunkerly-Eyring, B.; Khaliddin, N.; Xu, Z.; Fusco, A.P.; Yazib, S.A.; Chow, R.C.; Duh, E.J.; et al. Exosomal MicroRNA-15a Transfer from the Pancreas Augments Diabetic Complications by Inducing Oxidative Stress. Antioxid. Redox Signal. 2017, 27, 913–930. [Google Scholar] [CrossRef] [PubMed]
  234. Bell, C.G.; Lowe, R.; Adams, P.D.; Baccarelli, A.A.; Beck, S.; Bell, J.T.; Christensen, B.C.; Gladyshev, V.N.; Heijmans, B.T.; Horvath, S.; et al. DNA methylation aging clocks: Challenges and recommendations. Genome Biol. 2019, 20, 249. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  235. Pal, S.; Tyler, J.K. Epigenetics and aging. Sci. Adv. 2016, 2, e1600584. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  236. Povinelli, B.J.; Rodriguez-Meira, A.; Mead, A.J. Single cell analysis of normal and leukemic hematopoiesis. Mol. Aspects Med. 2018, 59, 85–94. [Google Scholar] [CrossRef]
  237. Guo, M.; Peng, Y.; Gao, A.; Du, C.; Herman, J.G. Epigenetic heterogeneity in cancer. Biomark. Res. 2019, 7, 23. [Google Scholar] [CrossRef]
  238. Wu, J.; Tzanakakis, E.S. Deconstructing stem cell population heterogeneity: Single-cell analysis and modeling approaches. Biotechnol. Adv. 2013, 31, 1047–1062. [Google Scholar] [CrossRef] [Green Version]
Figure 1. Epigenetics of aging and aging-related diseases. During aging, various epigenetic alterations occur including accumulation of histone variants, changes in chromatin accessibility mediated by chromatin remodeling complexes, loss of histones and heterochromatin, imbalance of activating/repressing histone modifications and aberrant expression/activity of miRNAs. These deregulations can affect transcription and, subsequently, translation, as well as the stabilization or degradation of molecular components. Consequently, these aberrant epigenetic processes can promote morbidities, which are frequently observed in the elderly populations, including inflammation, cancer, osteoporosis, neurodegenerative diseases, and diabetes.
Figure 1. Epigenetics of aging and aging-related diseases. During aging, various epigenetic alterations occur including accumulation of histone variants, changes in chromatin accessibility mediated by chromatin remodeling complexes, loss of histones and heterochromatin, imbalance of activating/repressing histone modifications and aberrant expression/activity of miRNAs. These deregulations can affect transcription and, subsequently, translation, as well as the stabilization or degradation of molecular components. Consequently, these aberrant epigenetic processes can promote morbidities, which are frequently observed in the elderly populations, including inflammation, cancer, osteoporosis, neurodegenerative diseases, and diabetes.
Ijms 22 00401 g001
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Saul, D.; Kosinsky, R.L. Epigenetics of Aging and Aging-Associated Diseases. Int. J. Mol. Sci. 2021, 22, 401.

AMA Style

Saul D, Kosinsky RL. Epigenetics of Aging and Aging-Associated Diseases. International Journal of Molecular Sciences. 2021; 22(1):401.

Chicago/Turabian Style

Saul, Dominik, and Robyn Laura Kosinsky. 2021. "Epigenetics of Aging and Aging-Associated Diseases" International Journal of Molecular Sciences 22, no. 1: 401.

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