DNA methylation and chromatin modifications

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DNA methylation and chromatin modifications

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Zahra Sepehri, Tasnim H. Beacon, Fadumo D.S. Osman, Sanzida Jahan, James R. Davie Department of Biochemistry and Medical Genetics, University of Manitoba, Winnipeg, MB, Canada

1. Epigenetics and chromatin organization Epigenetic refers to a variety of processes that have heritable effects on gene expression programs without changes in DNA sequence [1,2]. Key players in epigenetic control are DNA methylation and histone modifications which, in concert with transcription factors, chromatin remodeling complexes, nuclear architecture and non-coding RNAs, define the chromatin structure of a gene and its transcriptional activity. Cellular differentiation is initiated and maintained by epigenetic mechanisms. Although epigenetic marks are established early during development and differentiation, adaptations occur throughout life in response to intrinsic (e.g. oncogene expression) and environmental stimuli (e.g. diet) and may lead to late life disease (e.g. cancer). Thus, the life of an individual is not only defined by his/her genome, but also by his/her numerous epigenomes, with different epigenomes being generated through development, not only during fetal development but also during the plastic phase of early childhood, and existing in different cell types. Moreover, epigenomes react to environmental influence including maternal care, diet, exposure to toxins and xenobiotics. Epigenetic responses to environmental stimuli may have long-term consequences, even affecting future generations. The task ahead of us, to decipher all normal epigenomes and dysfunctional epigenomes leading to the vast array of diseases and cancers is colossal. The basic repeating structural unit in chromatin is the nucleosome. The nucleosome consists of a histone octamer, arranged as an (H3eH4)2 tetramer and two H2A-H2B dimers, around which DNA is wrapped. The H1 histones bind to the DNA, the linker DNA that join the nucleosomes together [3]. The core histones (H2A, H2B, H3, H4) have a similar structure with a basic N terminal domain, a globular domain organized by the histone fold, and a C terminal tail. The N terminal tails emanate from the nucleosome in all directions and interact with linker DNA, nearby nucleosomes or with other proteins [4]. The histones undergo numerous reversible post-translational modifications (PTMs), including acetylation, methylation and phosphorylation [5]. Histone PTMs are added and removed from specific sites by a variety of enzymes [6]. Some histone PTMs (active marks) are associated with transcribed chromatin regions, while others (repressive marks) are present in silent regions. Histone acetylation and H3K4me3 are active gene marks, whereas H3K27me3 is a repressive mark. Histone PTMs function to alter chromatin structure and/or provide a “code” for recruitment or occlusion of Nutritional Epigenomics. https://doi.org/10.1016/B978-0-12-816843-1.00002-3 Copyright © 2019 Elsevier Inc. All rights reserved.

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nonhistone chromosomal proteins to chromatin. The nucleosomal DNA may also be methylated (five methyl cytosine) [7]. Thus, the nucleosome serves multiple roles in regulating gene expression, a structural role and a role as a signaling module [8,9]. Adding to this complexity, most histones have variants, products of different genes [10]. Vertebrate histone variants are classified as replication-independent and replication-dependent. For example, H3.1 and H3.2 are replication-dependent, while H3.3 is replication-independent. Being synthesized throughout most of the cell cycle, the H3.3 variant is available to re-build nucleosomes displaced by transcription factors and the transcription machinery [11]. H3.3 is enriched in PTMs that are associated with transcriptional activity (for example, S28ph, K9ac, K14ac, K4me3) [12e14]. Mutations in H3.3 are associated with several cancers (e.g. pediatric glioblastoma) [15]. The structure and composition of a nucleosome depend to a certain extent on the DNA sequence associated with the core histones. A nucleosome with a DNA sequence that has binding sites for pioneer transcription factors and located in a tissue-specific enhancer may have an accessible configuration [16]. A pioneer transcription factor such as FoxA1 can bind to its binding site in a nucleosome context and in a repressive heterochromatin environment [17]. Nucleosomes associated with non-methylated CpG islands may be rich in H3K4me3. SETD1A/B, which catalyzes trimethylation of H3K4, has among its subunits, CXXC1 (also known as CFP1), a CpG island binding protein [18]. A nucleosome that has H3K4me3 attracts multiple lysine acetyltransferases (KATs) like HBO1 and SAGA which contain subunits that bind to K4me3. Thus, a nucleosome with H3K4me3 will have much different acetylation dynamics than a nucleosome next to it that does not have this modification [18e20]. Nucleosome structure is also radically changed during the process of transcription, which generates atypical nucleosomes known as U-shaped nucleosomes [21,22]. The open, U-shaped nucleosome will retain this altered configuration as long as its acetylation status remains high [23].

1.1 Three-dimensional structure of chromatin The organization of the genome plays an important role in the regulation of gene expression (Fig. 2.1). With the advancement of high-resolution techniques such as 3C and 3C based strategies (e.g. 3C, 4C, 5C, ChIA-PET and Hi-C analysis) [24], the 3D structure of chromatin has become an important model to address questions such as how the conformation of this structure controls interactions of regulatory elements, transcriptional activities and what factors/key players are involved in the maintenance of such conformation. High-resolution mapping of nucleosomes has allowed us to study misregulation of nucleosome positioning at the basic level of compaction of chromatin [25]. Chromatin is further organized into functional territories based on their state of condensation into euchromatin and heterochromatin. A higher level of compaction is required to fit the DNA and interacting proteins within the nucleus. As research advances to understand the underlying mechanism of how chromatin organization impacts gene regulation, major efforts have been made to understand the organization of these epigenetic compartments, map their interaction and determine their spatial arrangement [26]. Chromatin has different levels of spatial organization ranging from the nucleosome to domains. Current literature subcategorizes these spatial scales into three levels: large, intermediate and small. Compartmentalization and chromatin interactions are confirmed mainly by microscopic observations and by the Chromosome Conformation Capture (3C) technique. Microscopy is used to detect fluorescent probes used to track the spatial location of whole chromosomes and the relative positions of locus of interest, while 3C methods quantifies the interaction of two distal segments of the DNA

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FIG. 2.1 Organization of Chromatin in the Interphase Nucleus. The chromosome territories in the interphase nucleus are composed of chromatin compartments (A/B). The A-compartments are active and are associated with the active marks and their catalyzing enzymes (KMTs, KDMS, KATs and HDACs). The B compartments are found near the nuclear lamina and contain the repressed mark and associated enzymes (PcG and JmJD). The compartments are further organized into TADs each individually regulating the genes and regulatory elements within the insulating loop. The TAD boundaries are the regions where the CTCF and cohesin complexes bind. Credit: James R. Davie.

based on ligation events that takes place between the chromatin fragments after the steps of fixation and digestion. The proximity relative to one another is then used to determine their genomic location [24]. These organizations are commonly termed A/B compartments, topologically associating domains (TADs), and chromatin loops [27] (Fig. 2.1). Segregating the nuclear organization into different scales has facilitated the study of individual chromatin components correlating their dimensional interaction and overarching function in gene regulation [28]. 3C techniques have been promising in providing insight into the 3D arrangement, while the Hi- C technique appreciates and quantifies the interactions leading to average chromosome conformations. Initially the compartments were defined as closed and open chromatin compartment. The Hi-C mapping was performed at 1 Mb and as low as 100 kb and was validated with 3D-Fish to confirm the proximity [27]. With the advent of higher resolution, the structural domains were classified into 2 major compartments in the interphase stage: A compartment and B compartment (Fig. 2.1). The A compartment houses active chromatin (H3K36me3), while the B compartment is associated with

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Chapter 2 DNA methylation and chromatin modifications

inactive chromatin (H3K27me3). The active state is defined by transcriptional activity, associated histone modifications, and accessible DNA. Active chromatin domains are distinguished by their increased sensitivity to DNase I and association with highly acetylated histones [29e31]. Note that DNase I hypersensitive sites differ from that of DNase I sensitive regions. DNase I hypersensitivity marks regulatory elements in the genome that are typically nucleosome depleted and detected by techniques such as FAIRE and ATAC [29,32e34]. DNase I sensitivity encompasses much broader chromatin regions and correlates with histone acetylation status [35,36]. A/B compartments differ in chromatin composition and histone modifications, which identifies the compartment as active and inactive, respectively. The compartments stay constant in the genome throughout development [37]. However, changes to their associated histone modifications, DNAse I sensitivity or composition can lead to a change in their compartmentalization [38]. A pioneer transcription factor has the potential to convert a B to an A compartment [17]. The A compartment is organized into an inner ring-shaped structure, while the B-compartment is associated with the nuclear lamina and the edges of nucleoli within the nucleus. Single-cell Hi-C studies reveal the presence of these compartments in the only interphase cell. While chromosomes attain different conformations in different cells, the A/B compartments spatially segregate both within chromosomes and globally within nuclei [39,40]. At a more intermediate scale, self-interacting domains are seen which typically range in size from tens of kilobases to a few megabases usually comprising of a small number (e.g., 1e10) of genes. These interacting DNA sequences are commonly termed as ‘Topologically Associating Domains’ (TADs) but also are seen to be defined as sub-TADs, ‘contact domains’ and ‘insulated neighbourhoods’ based on their interaction and regulatory functions [37] (Fig. 2.1). TADs play an important role in domain organization. Experiments involving duplication, deletion or inversion of TADs or TAD boundaries have caused physiological malformations. Therefore, disruption of TADs can have significant effects on gene expression resulting in pathogenic phenotypes, indicating its importance in maintaining genomic architecture [41]. TADs function independently of the compartments [42]. The TAD boundaries interact more frequently with each other compared to the rest of the domain and result in a large fraction of peak (38%) when captured with Hi-C. This peak marks the binding of the CTCF and the cohesin complex [43]. The loop domain formed between CTCF and cohesin binding sites buffers the genes from the influence of enhancers residing in different domains [41,44]. The CTCF motifs of the insulating loop are always oppositely positioned marking the boundaries of the loops [39]. These boundaries are sensitive to DNAse I digestion, which indicates the low density of nucleosomes [45]. Therefore, altering the level of the nucleosome remodeling protein BRG1 was seen to change the density of nucleosomes at the boundary and affect boundary strength as well as CTCF binding [46]. While the loss of cohesin and CTCF was shown to have no effect on the compartmentalization of chromatin and the associated histone marks, it resulted in loss of TADs. TADs are segregated from each other by the formation of an insulating loop, which prevents the enhancer from one TAD to control the transcription of a gene in another TAD leading to insulated neighborhood where the gene and its regulatory elements are unaffected from external influence [47]. One exciting feature of the loop is its dynamic nature. It is observed to be present transiently while in a constant state of forming and collapsing resulting in an unstable structure [48]. One of the successful models describing the insulating loop formation is the ‘loop extrusion model’ where a loop extrusion factor (LEF) would bind to the chromatin and will initiate loop formation with the help of the boundary factors. The loop structure can incorporate additional LEF causing the formation of a secondary loop. The loop

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eventually extends to reach the TAD boundaries and are temporarily stabilized [49]. The loop will then disrupt shortly following the dissociation of the LEF and boundary factors. This model is useful to predict the role of CTCF at the TAD boundaries while setting the stage to hypothesize models to understand the underlying mechanism of their function in maintaining the conformation of the 3D structure of the chromatin. To understand how the large and intermediate level structures complement each other, experiments have been designed to explore how altering one structure affected the other. While the loss of CTCF and cohesin-associated factors (Rad21 and Nipbl) led to the disruption of the TADs and loops, deletion of WAPL (cohesin antagonist) had the opposite effect. Meanwhile loss of CTCF and cohesin-associated factors had no effect on the A/B compartment structures, indicating TADs and A/B compartments to be independent structures [50]. Another important factor was the contribution of these structures in different species. Experiments have shown that mammals show strong CTCF/cohesin loop anchors at TAD boundaries [50], whereas in Drosophila CTCF occupancy at the TAD boundaries are comparatively less and are not usually in inverted orientation [28]. These structures can be concluded as species-dependent for the factors involved in domain formation. Although the variation in abundance of CTCF and cohesin has led to different theories in their role in gene regulation, different theories have been speculated on their function leading up to the 3D organization of the chromatin. On a small-scale chromatin organization, variations to the 3C methods have been applied to detect enhancer-promoter interaction, which functions to regulate transcription. To map the interactions of the regulatory region, techniques such as OCEAN-C have been a helpful tool to identify hubs of open chromatin regions (HOCIs) and their regulatory functions. The cells are fixed, digested, ligated, and sonicated to desired fragment sizes. Following the phenol-chloroform extraction, the biotinylated DNA is analyzed for sequencing. As the technique combines FAIRE-seq and Hi-C, it eliminates the limitations of the techniques by improving the background and only captures the peaks representing open chromatin interaction. Important HOCI interactions include promoter-enhancer, promoterpromoter, and enhancer-enhancer interactions. These interactions are key to study the changes in open chromatin conformations as well as the regulation of the transcription of genes localized in proximal and distal domains [51]. Overall, studying chromatin in different scales has greatly advanced our understanding of the 3D architecture of chromatin by defining compartments, TADs and loops. Studying the interactions of the regulatory elements provides a topological basis to correlate open chromatin conformations to transcriptional regulation [52]. Knowledge of CTCF and cohesin factors along with the individual chromatin structures have laid the foundation of the mammalian 3D genome to explore conformational changes and identify novel epigenetic regulators mediating gene expression.

2. DNA methylation DNA methylation is a crucial epigenetic modification found in the genome of various organisms. It is involved in development, differentiation, tissue-specific gene expression and cellular function. Moreover, it plays a fundamental role in epigenetic reprogramming, X-chromosome inactivation and genomic imprinting. Recently, an interesting discovery of the role of DNA methylation in individual CpG sites as an epigenetic age-predictor in both human and mice was made [53,54]. Despite the availability of such tools for predicting biological age, which is quite different from chronological age, a majority of people are not willing to examine CpG DNA methylation for age prediction.

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Chapter 2 DNA methylation and chromatin modifications

However, this information is clinically relevant as a potential biomarker and in determining better therapeutic options for individuals according to their biological age. In mammals, methylation involves the addition of a methyl group at position five of the cytosine ring in the CpG dinucleotides by DNA methyltransferase enzymes (DNMTs) [55,56]. There are two classes of methyltransferases: the de novo methyltransferase (DNMT3a and DNMT3b) and the maintenance methyltransferase (DNMT1). The de novo methyltransferases establish the methylation mark onto the unmethylated CpG region whereas DNMT1 maintains the methylation through addition of methyl groups to hemi-methylated regions during replication. DNMT3l is a DNMT-related protein which binds H3 at lysine 4 (H3K4) and helps in guiding DNMT3a and DNMT3b to their target DNA region [57,58]. In addition to the enzymatic activity of DNMTs, these enzymes can contribute to gene inactivation through modifying the chromatin by binding to both histone deacetylase (HDAC) and lysine methyltransferases (SUV39h1/2 and G9; enzymes which methylate H3K9) [59]. Five-methylcytosine (5-mC) is predominantly found at CpG-rich sites but there are exceptions [60]. For example, human neurons and embryonic stem cells show intragenic methylation in a non-CpG context [61]. CpG islands, located usually at the 5’ end of genes, are generally unmethylated, and their methylation is more often restricted to genes which require stabilization for long-term silenced state such as imprinted genes and genes exclusively expressed in germ cells rather than somatic cells. The function of methylation varies with different genomic contexts [62,63] (Fig. 2.2). For example, methylation in the gene bodies supports gene expression and does not prevent transcriptional elongation. Recently, intragenic DNA methylation in vertebrates was shown to prevent cryptic transcription initiation within the coding region of expressed genes [64]. The intragenic methylation is catalyzed by DNMT3b, which is recruited to the gene body by H3K36me3 (catalyzed by SetD2) (Fig. 2.2). This mechanism involving crosstalk between SetD2, H3K36me3, DNMT3b and 5-mC is mediated by elongating RNA polymerase II [64]. By contrast, DNA methylation of promoter regions and transcription start sites blocks initiation and has been linked to gene silencing, either directly through blocking the binding of transcription factors [65] or indirectly through the involvement of methyl-CpG-binding proteins, which in turn recruit transcriptional repression and chromatin remodeling complexes [66] (Fig. 2.2). The methyl CpG-binding protein (MBP) family is responsible for identifying methylation including MBD1, MBD2, MBD4, MeCP2 and Kaiso. All the MBPs contain a methyl-CpG binding domain (MBD) that specifically recognizes methylated DNA [67] except for Kaiso, which depends on its zinc-finger (ZF) domain in the C terminus for methyl- CpG recognition [68]. Moreover, these proteins can function as methylation-dependent transcriptional repressors. Interestingly, not all transcription-factor binding is blocked by methylation as some factors like SP1 can bind strongly to methyl-CpG sequences and mediate gene expression through the passive removal (replication and enzyme independent demethylation) of 5-mC [69,70]. In the context of repetitive DNA sequences, methylation plays a repressive role by silencing transposable elements [71]. Similar effects are observed in methylated regulatory elements where the enhancer’s activity is reduced [72] and the binding of CTCFs is blocked [73]. However, for some enhancers, cytosine methylation is involved in maintaining an active enhancer state [74]. Also to note, several transcription factors, like SP1, bind to methylated DNA [75]. Overall, as the interpretation of 5-mC changes in a specific genomic and cellular context from another, its impact has gone beyond gene expression control to include genome stability and splicing regulation [61].

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FIG. 2.2 The Role of DNA Methylation in Gene Expression. (A) The CpG islands (CGI) within the promoter region and transcriptional start site (TSS) are normally unmethylated in active genes. The DNA methyltransferases (DNMT) are recruited by H3K36me3 to the gene body. Intragenic methylation (me) is positively correlated with active transcription and does not block the elongation process, but spurious transcription initiation is inhibited. (B) In inactive genes, methylation of promotors inhibits transcription initiation. The addition of methyl group is coupled with repressive chromatin modifications by DNMT-linked lysine methyltransferase (KMT) and histone deacetylase (HDAC). Additionally, methyl-binding domain (MBD) proteins recognize methylated regions and recruit chromatin remodeling corepressor complexes. Credit: James R. Davie.

2.1 DNA demethylation There are different pathways used in different tissues for the removal of 5-mC [76]. In mammals, 5-mC is passively demethylated during replication and cell division, while active demethylation occurs in cycling and non-replicating cells like neurons. In the oxidation pathway, 5-mC is oxidized by TET enzymes to 5-hmC which can be further oxidized to 5-formlycytosine (5-fC) and 5-carboxylcytosine (5-caC). 5-fC and 5-caC are cleaved by thymine DNA glycosylase (TDG) resulting in the involvement of the base excision repair (BER) machinery. On the other hand, the deamination pathway involves first the deamination of 5-mC by the activation-induced cytidine deaminase (AID)/apolipoprotein B mRNA editing enzyme, catalytic polypeptide (APOBEC) family producing a T:G mismatch identified by TDG which in turn creates an abasic site to be repaired by the BER machinery [76].

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In 2009, the first member of the ten-eleven translocation family of methylcytosine dioxygenases (TET1) was discovered to be responsible for 5-mC oxidation and demethylation to 5-hydroxymethylcytosine (5-hmC) [77]. The TET family of enzymes is characterized as iron- and oxoglutarate-dependent enzymes with their respective binding domains. In contrast to DNMT1, the CXXC zinc-finger DNA-binding domain of TET1 and TET3 has the ability to bind both methylated and hydroxymethylated DNA regions [78]. Five-hmC levels in adult tissues are moderately low ranging between >0.1 % and w0.7% with the highest level in the tissues of the central nervous system [79]. Five-hmC does more than its role as a demethylation intermediate, and TET enzymes are more than their catalytic function. Hydroxymethylation is enriched in euchromatic regions within the gene bodies, TSS and upstream of TSS. Gene expression can be controlled either negatively or positively by 5-hmC and TETs. The presence of 5-hmC in gene bodies prevents the binding of the DNA methylation machinery and MBPs allowing for the activation of genes [80e82]. However, TET enzymes can play another role in silencing genes by their interaction with SIN3A co-repressor complex [83]. Furthermore, the presence of 5-hmC in the case of bivalent genes (transcriptionally poised genes) facilitates the recruitment and binding of polycomb repressive complex 2 (PRC2), which catalyzes the trimethylation of H3 at lysine 27, thus maintaining the repressed state of these genes [84].

2.2 Mitochondrial DNA methylation Unlike nuclear DNA, mitochondrial DNA (mtDNA) has a 16,569-base-pair circular structure and lacks histones. It consists of a heavy (H) strand and a light (L) strand, encoding 37 genes: 13 oxidative phosphorylation-related protein-encoding genes, 22 transfer RNAs and 2 ribosomal RNAs [85]. MtDNA is believed to be modified by both 5-mC and 5-hmC, however, the presence and function of such epigenetic modifications are still highly debated. In 2011, a mitochondrially targeted isoform of DNMT1 (mtDNMT1) was discovered and accounts for 1e2% of total DNMT1 transcripts. Hypoxiaresponsive transcription factors such as peroxisome proliferator-activated receptor gamma coactivator 1 alpha (PGC1a) and nuclear respiratory factor 1 (NRF1) are responsible for regulating the mtDNMT1 expression [86]. Moreover, other DNMTs and TET enzymes were identified to be present in the mitochondria in a tissue-type-dependent manner. For example, the localization of DNMT3A in excitable tissues such as skeletal and heart muscles [87]. DNA methylation and hydroxymethylation patterns are observed across the mitochondrial genome, particularly their main location in the D-loop region where all the major promoters are located. Interestingly, more non-CpG compared to CpG is methylated in the D-loop region [88]. To date, the findings supporting the functional and regulatory roles of mtDNA methylation are based on associations without confirmatory evidence on its mechanism of action. The changes in mitochondrial gene transcription as a result of mtDNMT1 upregulation are not random, but rather genespecific leading to the activation of ND1 and repression of ND6 [86]. Transcription of mtDNA is dependent on mainly three factors: mitochondrial transcription factor B2 (TFB2M), mitochondrial transcription factor A (TFAM) in addition to mitochondrial RNA polymerase (POLRMT). Methylation might influence the ability of TFAM to bind the promoter regions directly or by the recruitment of proteins that add post-transitional modifications to TFAM [89]. Further work is required to fully understand and determine the detailed mechanism as well as to identify the players regulating mitochondrial DNA methylation.

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3. Histone modifications and their distribution in the genome Histones (H1, H2A, H2B, H3, H4) are basic proteins subject to a multitude of PTMs, which are mostly reversible [5,90]. Histone PTMs can affect almost all genomic events such as transcription, replication, recombination, DNA repair, kinetochore, chromatin remodeling and centromere formation [91]. Together with histone modifying enzymes and proteins that recognize specific histone PTMs, which are categorized as “Reader,” “Writer,” “Eraser,” “Effector” and “Presenter”, histone PTMs can regulate the transcriptional state [92,93]. Changes in histone PTMs can alter the active chromatin state into an inactive state and vice versa. Histone crosstalk is defined as a combination of histone PTMs that can code for transcriptional activation or repression in a context-dependent manner [94,95]. Histone PTM crosstalk can occur either cis or trans, involving events on the same histone tail or nearby histone tail within the same or neighboring nucleosome [96]. For example, it was demonstrated that serine 10 phosphorylation on H3 enhances the GCN5 mediated acetylation of H3 at lysine 14 [97]. Histone crosstalk can be initiated by preventing the nearby histone PTM. Indeed, H3 asymmetric di-methylation (H3R2me2a) was shown to prevent the MLL mediated formation of di- and tri-methylation of H3 lysine 4 (H3K4me3/H3K4me2) [98]. Interestingly, the presence of H3K4me3 prevents protein arginine methyltransferase (PRMT) 6-mediated H3R2me2a [98]. The advancement in the technology with tools such as chromatin immunoprecipitation (ChIP) and ChIP-sequencing (ChIP-seq) enables one to determine the crosstalk between different writers and readers or effector molecules. Lysine acetylation (all histones), lysine and arginine methylation (H3, H4, and H2B), serine and threonine phosphorylation (all histones), lysine ubiquitination (H2A, H2B), ADP- ribosylation at glutamine (H1) and sumoylation are some of the well-known histone PTMs [5]. Several of the histone marks are exclusively associated with active chromatin state (H3K4me3, H3K27ac), while others are with the inactive chromatin state (H3K27me3) [96].

3.1 Histone acetylation Histone acetylation was first reported by Vincent Allfrey and his group in 1964. Allfrey’s group described the dynamic and rapid histone acetylation using nuclei isolated from calf thymus [99]. Following on this study, they later reported that histone acetylation occurs on ε-amino lysine residue, and they also identified histone deacetylase (HDAC) activity in the nuclei. In 1978, both Dr. Davie and Dr. Allfrey reported for the very first time that n- butyrate acts as an HDAC inhibitor [100,101]. Hyperacetylation of H3 and H4 and to a lesser extent H2A and H2B were observed upon sodium butyrate treatment in the cell line investigated [100]. DNA sequences associated with the hyperacetylated histones showed increased DNase I sensitivity in HeLa and chicken erythrocyte cells [101]. The first report of a direct link between histone acetylation and transcriptionally active chromatin came from the study by Dr. Crane Robinson’s group using chicken erythrocytes [102]. In this study, the ChIP assay was used for the first time to demonstrate that acetylated histones are associated with transcriptionally active DNA sequences [102]. The relationship between histone acetylation and transcription became established after the discovery of KATs were co-activators [103]. KATs are categorized into four different groups; GCN5, MYST (SAS/MOZ), P300/CBP and SRC/p160 nuclear receptor coactivator family [104]. Acetylation of histone and non-histone proteins is catalyzed by KATs [105]. Dynamic and reversible histone acetylation is catalyzed by KATs and HDACs. The rate of histone acetylation can vary across the genomic regions with some regions having a faster rate of dynamic acetylation while some have slower or none [104].

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Histone acetylation can prevent H1-mediated salt insolubility, facilitating solubility of the region at physiological salt concentration [35,106,107].

3.2 Histone lysine methylation Lysine and arginine located in the histone N-terminal tails are methylated by lysine methyltransferases (KMTs) or PRMTs [5]. Mono-, di- or tri-methylation of lysine and mono- or di-methylation of arginine can be distinguished as active or repressive chromatin marks [108,109]. H3K4me1 along with H3K27ac typically marks active enhancers. H3K27me3 is a strong repressive mark catalyzed by the polycomb complex [110]. With our current knowledge of the histone PTMs associated with regulatory elements such active enhancers, poised enhancers, upstream promoters and silenced genes, we can now read histone PTM tracks and have a good idea of the function of the associated DNA sequence [111]. H3K4me3 is an active mark found in the upstream promoter region and strongly positioned after the first exon of transcribed genes [112]. The mitotic inheritance (memory) of the transcriptional state of genes is dependent upon H3K4me3 [113]. Bryan Turner’s group noted that the H3K4me3 regions on mitotic chromosomes coincided with CpG islands, which are found at the promoter and 50 regions of genes [18,114]. H3K4me3 has several roles in the regulation of gene expression. H3K4me3 located in the 50 coding region of expressed genes interacts with the proteins involved in pre-mRNA splicing (reviewed in Ref. [18]). Although most genes have H3K4me3 limited to the promoter region and the 50 region of the gene body, a small subset of genes have broad H3K4me3 regions (called H3K4me3 buffer domains). The function of H3K4me3 buffer domain is to ensure transcriptional consistency of the gene, that is, the same transcription rate in each cell [113,115]. Of note, these genes with the H3K4me3 buffer domains have a role in cell identity [115]. The top 5% broadest H3K4me3 regions in a given cell type were most enriched for genes that had important functions for that specific cell type (for example, genes involved in muscle function in skeletal muscle or stem cell regulators in embryonic stem cells) [115]. In normal but not cancer cells, tumor suppressor genes have broad H3K4me3 peaks [116]. Importantly genes coding for transcription factors, which were critical for cell identity or cell fate, have H3K4me3 buffer domains in the relevant cell type/tissue. Through the identification of genes with the H3K4me3 buffer domains, Anne Brunet and colleagues found genes coding for transcription factors and non-coding RNA that were novel regulators of neural progenitor cells [115]. Knocking down the genes with the broadest H3K4me3 domains resulted in decreased neural progenitor cell proliferation and decreased neurogenesis. Lysine and arginine methylation of histones can serve either as a binding site or occlude the binding of other modifiers to the site and thereby play a crucial role in histone PTM mediated signaling events. Due to the existing signaling event, aberrant binding of the modifying enzymes can lead to diseased state as observed for several cancers [117e119]. EZ, SET1, SET2, SMYD, SUV39, SUV4-20, RIZ are among the major family of lysine methyltransferases [120]. S-adenosyl methionine (SAM) serves as methyl donor and co-factor for both KMTs and PRMTs [121].

3.3 PRMTs and histone arginine methylation Eleven mammalian PRMTs have been discovered and reported to date. Mono and dimethylation of arginine are catalyzed by three classes of PRMT enzymes. PRMTs catalyze arginine methylation by

3. Histone modifications and their distribution in the genome

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using molecule of S-adenosyl-L-methionine (AdoMet) to form asymmetric (u-NG,NG-dimethylarginine or ADMA) or symmetric (u-NG,N0 G-dimethylarginine or SDMA) or monomethylarginines (MMA). Type I PRMTs, which asymmetrically dimethylate arginine, include PRMT1, 3, 4, 6 and 8, with PRMT1 accounting for most of the type I PRMT activity. PRMT5 and 9 are involved in symmetric dimethylation of arginine and belong to type II, while PRMT7, a type III methyltransferase, contributes to monomethylation of arginine [122]. Therefore, PRMTs have a wide range of substrate specificity. Arginine methylation of proteins and histones catalyzed by PRMTs can be either symmetrical or asymmetrical, and they are categorized based on this chemical feature [123]. Similar to lysine methylation, arginine methylation can contribute to activate or repress the chromatin state in a contextdependent manner [124]. H4R3me2a, an active chromatin mark, is catalyzed by PRMT1, whereas PRMT5 is responsible for the symmetric dimethylation of H4R3 (repressive mark). PRMT6 is the major methyltransferase responsible for the genesis of H3R2me2a (repressive mark) in vivo. H3R2me2s, formed by PRMT5, recruits WDR5, which is a subunit of several co-activator SET1/MLL complexes that produce H3K4me3 (an active mark) [125,126]. Other histone arginine methylations are listed below. H3R17me2a: This arginine modification is catalyzed by PRMT4/CARM1 and is associated with transcriptional activation. This mark is located at the upstream promoter of several genes [119,127,128]. Moreover, it was shown that prior acetylation of H3K18 and K23 promotes H3R17 methylation and activation of estrogen-responsive gene TFF1 [129,130]. H3R26me2a: PRMT4/CARM1 produces H3R26me2a, which is a less studied mark. Similar to H3R17me2a, this mark was also found associated with the upstream promoter region of target genes [131]. H3R2me2a: PRMT6 produces H3R2me2a. This mark has an intriguing feature as it can antagonize H3K4me3 through blocking the binding of WDR5 to the site [132]. This mark is associated with repressive chromatin and found in pericentromeric regions [133]. H3R2me2s: This mark is associated with transcriptionally active chromatin regions and is generated by PRMT5. H3R2me2s recruits WDR5, which is found in complexes such as MLL, SET1A, and SET1B, which catalyze H3K4me3. Therefore, H3R2me2s is located with H3K4me3 [134]. H4R3me2a and H2AR3me2a: PRMT1 is involved in the generation of these two modifications. H4R3me2a was found associated with the upstream promoter regions of several genes [135e138]. H4R3me2a was reported to facilitate the subsequent acetylation of H4 at Lys 8 and 12 and of H3 at Lys 9 and 14 by p300, and therefore H4R3me2a can be considered as an active mark that recruits the KATs, p300 and PCAF [135,139]. Acetylation of H4 Lys 5 results in reduced arginine methylation by PRMT1 but increased activity by PRMT5 [140,141]. Highly acetylated H4 that has H4K5ac is strongly detrimental to PRMT1 mediated H4R3 methylation. In contrast, acetylation of H4 at K16 is a better substrate for PRMT1 than unmodified H4 [141]. This is an interesting observation as H4K16ac is involved in decondensing chromatin [142]. H4R3me2s and H2AR3me2s: PRMT5 generates the repressive marks, H4R3me2s and H2AR3me2s. H4R3me2s was found associated with the upstream promoter region of several silenced genes and with imprinting control regions [143,144]. H3R8me2s: This is a repressive chromatin mark generated by PRMT5 [143]. Prior acetylation was shown to prevent H3R8me2s [145].

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3.4 Arginine demethylation Protein arginine deiminase (PAD) family of enzymes catalyze citrullination from the amino acid arginine. To date, PADs, PAD1-4 and PAD6, have been identified [146]. PADs replace the ketamine (¼NH) group of arginine to keto group (¼ O), thereby resulting in no net charge from the positively charged arginine. This change in charge due to citrullination alters the structure and function of the protein as well affect the binding of protein interacting partners [147]. It was reported that both PAD2 and PAD4 could catalyze citrullination on a histone tail, albeit it is PAD4 which is involved in citrullination of monomethyl arginine [148,149]. Symmetric and asymmetric mono and dimethylation of arginine H3/H4 were reported to be catalyzed by Jumonji domain-containing 6 protein (JMJD6) [150]. However, later it was shown that JMJD6 is involved only in the demethylation of mono- and di-methyl H4 arginine residues [151]. Although JMJD6 has been reported as a candidate for demethylation of arginine, there is still a lack of sufficient biochemical evidence for that. Moreover, demethylation of H3R2 has not yet been detected.

4. Metabolism and epigenetics Metabolism is a set of life-saving chemical modifications in the cells [152]. Every cell has a set of responses to changes in its environment. These responses depend on various factors such as cell type and duration of changes, to name a few. Overall, nutrition as an environmental input modulates cell metabolism, and in turn cell metabolism regulates epigenetic processes [153]. More precisely, the dynamic range of physiological concentrations of commensurate intermediates in metabolism will characterize the “kinetics” of crosstalk between metabolism and chromatin [154] (Fig. 2.3).

4.1 Metabolism and chromatin modifications Enzymes that are responsible for histone and DNA modifications need metabolites such as acetyl or methyl groups to do their enzymatic reactions specifically and efficiently. Availability of these metabolites and their localization in the cell play major roles in the activity of the enzymes regulating DNA and histone modifications. For example, efficiency of KATs depends on local subcellular acetylCoA concentration, which is produced during catabolism and anabolism [152,153]. In a nutrient favorable situation, that is, there is a lot of acetyl-CoA precursors entailing glucose, galactose or ethanol, adenosine triphosphate citrate lyase will convert glucose derived citrate into acetyl-CoA in the nucleus [152]. On the other hand, class III HDACs (the sirtuins) deacetylate histones and non-histone proteins dependent on the concentration of available local nicotinamide adenine dinucleotide (NADþ). The Sirtuin family of HDACs (SIRTs) “sense” the positive effects of caloric restriction on physiology of the cell and are a key controller in mitochondrial energy metabolism, senescence of the cell, inflammation and most importantly tumorigenesis [153]. Yet, they are NAD dependent [152]. When fasting happens, available NADþ and activity of SIRT1 are high. In contrast when energy resources are high, NADþ would be converted to NADH and consequently its concentration would decrease. In aging and diabetes, NAD levels significantly decrease [152]. Actually, SIRTs are the only HDACs that are dependent on an endogenous metabolite such as NAD [153]. Although SIRT6 has high affinity to bind to NAD, its dependency to NAD is less. SIRT6 specifically deacetylates pericentric

4. Metabolism and epigenetics

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FIG. 2.3 Interactions Among Environment, Cell Metabolism and Epigenetics. There are dynamic interactions among environmental inputs (such as nutrition, O2), cell metabolism pathways and epigenetic changes. NADþ, involved in redox reactions as electron carrier, is a coenzyme for class III HDACs (sirtuins). Acetyl CoA, which is an oxidizing reagent that conveys carbon atoms in TCA cycle, is a coenzyme for KATs. bOHB, ketone body in starvation and exercise, is an endogenous HDAC inhibitor. SAM, donor of methyl group, is coenzyme for DNMTs and HMTs. Credit: James R. Davie.

heterochromatin H3K18ac [155]. SIRT6 also deacetylates H3K9ac and H3K56ac resulting in repression of transcription of hypoxic inducible factor (HIF)-a driven glycolytic gene [152]. Another interesting possibility about links between nutrition and histone PTMs is that the ketone body b-hydroxybutyrate e product of breakage of fatty acids due to starvation or long time caloric restriction and consequently activation of fatty acid b oxidation - acts as an endogenous HDAC inhibitor [152,153,156,157]. This would lead to increased H3 acetylation (Lys9 and Lys14), and transcription of genes that are controlled with FOXO3a. Other studies confirmed that caloric restriction e mostly low carbohydrate diets-would lead to ketogenesis and this is associated with class I HDACs (yeast, Caenorhabditis elegans, Drosophila) in the extension of life span along with protection against reactive oxygen species (ROS). It seems that this metabolite controlled HDAC is a process that is involved in lots of cell events and not just longevity [153]. SAM is a donor of the methyl group for both DNA and histone methylation. Reduction of threonine, a metabolic precursor for the genesis of SAM, results in reduced levels of H3K4me2 and H3K4me3 but not H3K4me1 [158]. It is important to note that loss of SAM will reduce the activities of the KMTs, PRMTs and DNMTs but unless there is an event removing the histone or DNA modification, the modification will appear unaffected. TET enzymes need substrates and cofactors such as a-ketoglutarate (a-KG), Fe2þ and O2 for stepwise oxidation reactions [152,159]. More interestingly, a-KG, which is normally derived from

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Chapter 2 DNA methylation and chromatin modifications

either glucose or glutamine anabolism [160], plays a key role in the TCA cycle, amino acid synthesis and nitrogen transport. However, in any situation in which high levels of the inhibitor of F2- and a-KG-dependent dioxygenase are present as a result of dysregulative mutations, certain types of cancer would occur [161].

4.2 Hypoxia-induced epigenetic changes Hypoxia is defined as inadequate available oxygen (O2) for the demands of target tissues. Demethylation enzymes such as LSD1, JHDMs, JMJDs, JARIDs, UTX, and TETs need oxygen as a substrate and/or cofactor [154]. When tissue metabolism exceeds metabolic supplies, adaptation and maladaptation processes will start. These processes occur to prevent cell death. In the presence of enough O2, cells oxidize glucose to provide adenosine triphosphate (ATP) as the source of energy. In hypoxia, glucose is converted to lactate. This switch from oxidative to glycolytic metabolism is mediated by hypoxiainducible factor 1 (HIF-1) [162]. Hypoxia might occur as chronic, intermittent, and acute. Adaptive responses to chronic hypoxia are due to binding of HIF-1 to a hypoxia-response element (HRE) located near the target gene. This HIF-1-dependent gene regulation is inherently sensitive to cytosine methylation by DNA methyltransferases. The changes in DNA methylation alter gene expression patterns, in some cases irreversibly and last long after hypoxia is resolved [163]. Living in low barometric pressure at high altitude such as in the mountains is one of the best examples of chronic exposure to hypoxia [164]. Preeclampsia is a pathologic situation of chronic hypoxia and changes in gene expression due to disruption in DNA methylation [165]. In the tumor context, hypoxia is responsible for about half of DNA hypermethylation in solid tumors [166,167] and triggers initiating tumor occurrence in cells through epigenetic changes [168,169]. In cancer cells, there is often a lower expression of TET. However, the low oxygen levels in hypoxia also reduce the activity of the TET enzymes. Together, the low protein levels and reduced activity of the TET enzymes may alter DNA methylation and expression of some genes [167,168,170e173]. Sub-acute intermittent hypoxic exposure may result in reversible epigenetic adaptations, while chronic hypoxia leads to irreversible epigenetic adaptations [164]. One example of long-term intermittent hypoxia is obstructive sleep apnea (OSA). OSA is a common clinical problem that affects one in 5 (approximately 20%) healthy individuals and 40%e70% of obese people [174] and even children [175]. Long-term chronic intermediate hypoxia increased DNA methylation and down-regulation of anti-oxidant genes, leading to persistent cardiorespiratory abnormalities [176,177]. The expression of the anti-oxidant genes was restored after decitabine treatment. This treatment also normalized cardiorespiratory function [176]. There are fewer studies about short-term acute nonlethal hypoxia situation. A study on hypoxia induced epigenetic reprogramming in hippocampal neurons showed that acute hypoxia causes upregulation in the genes more than downregulation. In addition, promoters and CpG islands remained hypomethylated several days after hypoxic exposure. It supports the idea that whole genome methylation reprogramming is correlated with gene expression days after sub-lethal hypoxic stress [178].

5. Concluding remarks There is still much to learn about the plethora of histone PTMs and how these modifications crosstalk with each other and with DNA modifications. Several of the histone PTMs play a role in altering

Acknowledgments

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chromatin structure (such as H4K16ac); however, many do not. The latter serve important roles in attracting or repelling the binding of non-histone proteins and/or chromatin modifying enzymes (the readers). There is much to be worked out in order to understand how the readers recognize the histone and DNA modifications and what the roles of the reader once bound are. Metabolism plays a major role in regulating histone and DNA modifications involved in epigenetic processes [179]. Metabolism is impacted in many disease states particularly in cancer [180,181]. The mitochondria dynamic network “communicates” with the nucleus in the regulation of metabolism and epigenetic processes; there is much to be learned about this relationship and how it is altered in disease. Each cell may have up to 10,000 mitochondria and each mitochondrion may have up to ten circular mitochondrial DNAs. How do mitochondria numbers and network play into the dynamic web of events involving mitochondria, nucleus, metabolism and epigenetic processes? How does exercise fit into this picture? Clearly, the external (diet, environment in which we live) and internal environments (metabolic enzymes, hypoxia) will impact epigenetic programming and could potentially have long term, transgenerational consequences. As we will all find out one day, our metabolism changes as we age. NAD levels drop [182] and as a consequence, epigenetic processes do not perform with the same efficiency as they did when we were younger. With the decline of NAD, acetylation of histone and non-histone proteins may increase due to reduced activity of the sirtuins. The increased acetylation of histone and non-histone proteins will have consequences on their activities and cellular levels. In cases where the acetylated sites are also ubiquitinated, the acetylation event may prevent turnover of the protein by the proteasome [183]. For example, SIRT1 deacetylates acetylated HDAC1 [184,185]. Acetylated HDAC1 is not active and trans-represses the activity of HDAC2. Acetylation of p53 increases the stability and activity of this tumor suppressor by suppressing ubiquitination. HDAC1 and SIRT1 deacetylate acetylated p53, regulating its activity and levels. Loss of SIRT1 results in the accumulation of acetylated HDAC1 and acetylated p53 [184]. The genomic levels of the replication-independent histone variants, such as H3.3 increase with age and with the elevated levels of H3.3 comes an increase in the PTMs associated with this variant [186]. With old age, DNA methylation of the genome declines [187]. Neurological disorders, which escalate with age, are a consequence of deregulation in the epigenetic processes. As many players in these epigenetic processes are druggable, refinement of our drug arsenal may counter neural decline during normal aging [188]. As our understanding of the mechanisms regulating metabolism and epigenetic processes matures, we will realize the importance of our lifestyle choices such as diet and regular exercise in healthy living and healthy aging.

Acknowledgments This work is supported by an Environments, Genes and Chronic Disease Canadian Institutes for Health Research Team Grant (144626), a Natural Sciences and Engineering Research Council of Canada Grant (RGPIN-201705927), and a CancerCare Manitoba Foundation Grant (761020234). Ms Beacon and Dr. Sepehri were funded by the Graduate Enhancement of Tri- Council Stipends (GETS) through the University of Manitoba. Ms Osman was funded by CancerCare Manitoba/Children’s Hospital Research Institute of Manitoba Master’s Studentship. Dr. Sepehri thanks the Zabol University of Medical Sciences for the opportunity to study abroad in Dr. Davie’s research group.

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