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Epigenetic mechanisms and genome stability
Clinical Epigenetics volume 2, pages299–314(2011)
Epigenetic marks are well recognized as heritable chemical modifications of DNA and chromatin that induce chromatin structural changes thereby affecting gene activity. A lesser-known phenomenon is the pervasive effects these marks have on genomic integrity. Remarkably, epigenetic marks and the enzymes that establish them are involved in multiple aspects of maintaining genetic content. These aspects include preserving nucleotide sequences such as repetitive elements, preventing DNA damage, functioning in DNA repair mechanisms and chromatin restoration, and defining chromosomal organization through effects on structural elements such as the centromere. This review discusses these functional aspects of epigenetic marks and their effects on human health and disease.
Epigenetics is broadly defined as the heritable, regulatory elements of a genome exclusive of its primary DNA sequence. Methylation of DNA and post-translational modifications of the core histone proteins are the most well understood epigenetic marks. Together, they organize DNA into chromatin and ensure proper gene regulation. Additionally, epigenetic regulation of eukaryotic genomes has been increasingly shown to impart stability on the DNA sequence and to aid in maintenance of genomic integrity. Epigenetic aberrations causing genetic instability are at the root of developmental abnormalities such as ICF (immunodeficiency, centromere instability, and facial anomalies) syndrome, which is characterized by immune defects, heterochromatin instability, and mental retardation, and disease states such as cancer. This review discusses the fundamentals of epigenetics and presents its many roles in maintaining genome stability.
DNA methylation as an epigenetic mark
To regulate genetic information, a cell's immense quantity of DNA must be systematically packaged and organized, a function provided by chromatin structure. Chromatin can be distinguished as heterochromatin, characterized as a highly condensed and transcriptionally inert state, or as euchromatin, a loosely packaged DNA arrangement easily accessible by transcription factors and the transcription machinery. These states are largely determined by covalent modifications of the histone proteins contained within chromatin and by DNA methylation.
DNA methylation is a covalent modification of DNA that, in vertebrates, predominantly occurs at the carbon-5 position of cytosine nucleotides followed by a guanine (CpG). Patterns of DNA methylation are strongly associated with transcriptional repression and are localized to domains of heterochromatin (Miller et al. 1974). On a global level, in differentiated somatic cells, CpG dinucleotides are methylated at about 80%, whereas CpG islands, which contain longer stretches of CpG-containing sequence and are typically associated with gene promoters, often remain unmethylated, presumably to permit gene activation (Bird 1999, 1986; Gardiner-Garden and Frommer 1987; Jaenisch and Bird 2003).
The cell's DNA methylation “signature” provides explicit instructions for directing functions such as maintaining pluripotency or promoting tissue-specific gene expression and controlling growth and proliferation. Establishment of proper DNA methylation patterns and maintenance of those patterns through mitotic division is critical for cellular function. During mammalian development soon after fertilization, gene expression profiles are “reset” by a wave of widespread DNA demethylation, especially of the paternal genome (Jaenisch and Bird 2003; Mayer et al. 2000; Oswald et al. 2000). Subsequently, genome-wide de novo methylation commences following implantation of the zygote. The DNA methyltransferases (DNMT) DNMT1, DNMT3A, DNMT3B, and DNMT3L actively contribute to and determine these DNA methylation patterns in mammalian cells by differential means. DNMT1 has a strong bias for hemi-methylated DNA and localizes to DNA replication foci; these features are consistent with DNMT1 playing a major role in maintenance of methylation in dividing cells (Bestor 1992; Leonhardt et al. 1992). DNMT3A and DNMT3B, on the other hand, operate equally well on unmethylated or hemi-methylated DNA substrates and thus are classified as possessing de novo methylation activity (Liang et al. 2002; Okano et al. 1999; Yokochi and Robertson 2002). Their activity is enhanced by DNMT3L, a catalytically inactive protein with sequence similarity to DNMT3A and DNMT3B (Chedin et al. 2002; Hata et al. 2002; Jia et al. 2007; Margot et al. 2003; Suetake et al. 2004). Both de novo and maintenance methylation are required for development, since deficiency for any of the DNMTs affects viability (Okano et al. 1999). DNA methylation marks are interpreted by methyl-CpG-binding proteins including MBD1, MBD2, MBD3, and MeCP2, which generally act to silence gene expression and recruit histone deacetylases (HDACs), which are required for chromatin condensation (Bowen et al. 2004; Boyes and Bird 1991; Jaenisch and Bird 2003; Lewis et al. 1992; Meehan et al. 1989; Nan et al. 1997). Thus, DNA methylation and the proteins associated with writing (DNMTs) and reading (MBDs) this mark provide a robust mechanism for regulation of transcriptional activity, but more importantly, the stability of DNA methylation as a covalent modification and the replicable nature of DNA methylation marks facilitate the inheritance of epigenetic information during important processes such as X chromosome inactivation, imprinting, silencing of repetitive elements, transposable elements, and pericentromeric repeats, and developmental gene regulation (Gopalakrishnan et al. 2008; Li 2002; Slotkin and Martienssen 2007).
The key question regarding DNA methylation is what factors or stimuli determine how methylation patterns are established. Studies in Arabidopsis show that positioning of nucleosomes along DNA is directly related to the patterning of DNA methylation: nucleosome-bound DNA is enriched for methylation compared to nucleosome-flanking sequences (Chodavarapu et al. 2010). Likewise, the histones comprising the nucleosome core bear post-translational modifications involved in directing DNA methylation patterns, chromatin restructuring, and transcriptional effects (Hu et al. 2009; Ooi et al. 2009, 2007; Zhang et al. 2010).
Histone modifications in chromatin function
The basic repeating unit of chromatin is the nucleosome, containing an octamer of four core histones, H2A, H2B, H3, and H4, around which double-stranded DNA is coiled. Each histone consists mostly of a globular domain except for an N-terminal tail, which protrudes from the nucleosome where it is accessible for binding to DNA, for protein interactions, and for covalent attachment of small molecules. Covalent modifications of amino acid residues in histones direct DNA methylation, chromatin conformation, and gene transcription by promoting or restricting the recruitment of gene regulatory proteins (Jenuwein and Allis 2001). The most well understood histone modifications include methylation, phosphorylation, acetylation, and ubiquitination, and chromatin regulation is conferred by the combination of these modifications on different residues and, in the case of histone methylation, by the number of moieties attached to a specific residue (Kouzarides 2007). Methylation of histone H3 lysine 9 (H3K9), H3K27, H4K20, and H3 arginine 2 (R2) are just a few of the marks determining heterochromatin formation and transcriptional repression (Kirmizis et al. 2007; Kouzarides 2007). Euchromatin is characterized by methylation of H3K4 and acetylation of H3 and H4 histone tails. For histone methylation, the number of attached methyl groups also influences its role as a regulatory mark. Lysines and arginines are mono-, di-, or trimethylated. Di- and trimethylation of H3K4 (H3K4me2 and H3K4me3), for example, is strongly associated with regions of transcriptional permissiveness, and hypomethylation of this residue is correlated with loss of H3K9 acetylation (Bernstein et al. 2002; Santos-Rosa et al. 2002; Schneider et al. 2004). While H3K9me3, H3K27me3, and H4K20me3 are all heterochromatic marks, mono-methylation of these same residues correlates with transcriptional permissiveness. Additionally, some histone marks show specificity for certain gene regions. Transcriptionally active genes are characterized by H3K4me2 within their coding regions, while histone acetylation and H3K4me3 are enriched in promoter regions, and H3K4me1 occurs in enhancer elements (Bernstein et al. 2002; Heintzman et al. 2007; Liang et al. 2004; Schneider et al. 2004). H3R2me2, which is mutually exclusive with H3K4me3, is found within transcriptionally inactive genes and in the 3′ region of transcribed genes (Kirmizis et al. 2007).
A collection of histone-modifying enzymes including lysine methyltransferases, arginine methyltransferases, serine-threonine kinases, and acetyltransferases is involved in writing each mark. These enzymes prove to be highly specific in recognizing a single amino acid substrate, and multiple enzymes provide functional redundancy for writing each mark and/or the ability of particular marks to respond to many different stimuli (Kouzarides 2007). For instance, the SET domain methyltransferases SUV39H1, SUV39H2, G9, and SETDB1, among others, show strict specificity for methylating H3K9. Likewise, mixed lineage leukemia enzymes 1–5 specifically recognize and methylate only H3K4 residues (Kouzarides 2007).
The combinations of these modifications at specific loci are highly dynamic and capable of rapid change depending on a cell's response to cell–cell signaling and environmental cues. For instance, pluripotent cells possess a state of “bivalency” in which H3K27 methylation, a mark of heterochromatin, overlaps with loci containing euchromatic H3K4 methylation (Azuara et al. 2006; Bernstein et al. 2006; Ku et al. 2008). These opposing marks occupy genes encoding many developmentally important transcription factors that were presumably “poised” for either activation or dense repression by stabilization of one of these marks upon differentiation. Thus, some have questioned whether histone modifications should be referred to as epigenetic marks under a strict classical definition of a stable element, faithfully inherited throughout many cell generations (Kouzarides 2007). Recent evidence has shown that at least some histone marks are indeed preserved and replicated during cell division. The repressive mark H3K27me3 is maintained during DNA replication by binding of the polycomb repressive complex 2 to H3K27me3 within promoters of transcriptionally repressed genes (Hansen et al. 2008). Identifying additional mechanisms that allow for the reproduction of histone modifications within the chromatin of newly synthesized DNA will be important for understanding how gene expression programs are controlled. Regardless of their heritability, these modifications provide important information crucial for genomic stability and proper DNA methylation; so, for the purpose of this review, we continue to refer to them as epigenetic marks.
As previously mentioned, histone modifications are important in establishing DNA methylation marks. Repressive histone marks may be established independently of DNA methylation and are capable, in some cases, of inducing de novo DNA methylation. DNMT3L recognizes loci lacking unmethylated H3K4, and an interaction between DNMT3L and the most N-terminal residues of histone H3 is required for de novo DNA methylation (Hu et al. 2009; Ooi et al. 2007). Furthermore, DNMT3A and DNMT3B interact with SUV39H1 and EZH2, which are H3K9 and H3K27 histone methyltransferases, respectively, as well as residues 1–19 in the N-terminal tail of histone H3 (Lehnertz et al. 2003; Vire et al. 2006; Zhang et al. 2010). Conversely, DNA methylation may also direct establishment of repressive histone marks. In ICF cells deficient for DNMT3B function, loci affected by loss of DNA methylation also demonstrated a loss of repressive H3K27 methylation marks and a coordinated gain of H3K9 acetylation and H3K4 trimethylation (Jin et al. 2008).
Epigenetics and genomic stability
Originally, DNA methylation may have been evolutionarily acquired to defend against genomic disruption by parasitic insertional element translocations in eukaryotic genomes, an idea that has become widely supported (Robertson and Wolffe 2000; Slotkin and Martienssen 2007; Yoder et al. 1997). Subsequently, functionality for DNA methylation in other modes of genomic stability may have been co-opted (Colot and Rossignol 1999). In the following sections, we focus on the evidence that epigenetic mechanisms help to preserve genomic content.
Effects of epigenetic marks on microsatellite repeat stability
DNA methylation and DNA methyltransferases have been implicated in affecting stability of microsatellites within the genome for some time. Microsatellite loci are repetitive sequences typically consisting of one to four nucleotide repeats, and they are particularly susceptible to length change mutations. Microsatellite repeat instability (MSI) presents a notable hazard in light of its involvement in diseases such as Huntington's, myotonic dystrophy, and cancer among others (see below) and is often associated with defects in the mismatch repair (MMR) machinery, a system that specifically recognizes and repairs errors in nucleotide base pairing.
Repetitive regions are subject to expansion or contraction when single-stranded DNA becomes exposed, allowing repetitive sequences to form secondary structures (i.e., DNA hairpins), particularly during replication, transcription, and DNA repair (Pearson et al. 2005). These secondary structures may then become incorrectly resolved through MMR or cause slippage of DNA polymerases along repetitive sequence during replication. For a history of MSI discovery and characterization, see Laghi et al. (2008).
Epigenetic mechanisms appear to protect against MSI both directly and indirectly (Table 1). Disruption of DNA methylation or the DNMTs destabilizes repeats through a direct local chromatin remodeling effect, such as transcriptional de-repression, or possibly another unknown mechanism. Alternately, DNA methylation affects repeat stability secondarily by producing global changes in gene expression that alter expression of proteins responsible for repeat stabilization (i.e., MMR genes). Here, we discuss the evidence for these epigenetic effects on MSI and their potential mechanisms.
Several groups have shown that MSI results from disruption of DNMT1 or of DNA methylation patterns. Most notably, DNMT1 mutants were isolated from an insertional mutagenesis screen for ES cell clones exhibiting defects in MMR (Guo et al. 2004). To measure MSI in these mutants, a “slippage” assay was employed in which an insertional construct containing repetitive sequence was monitored for gain or loss of nucleotides. DNMT1 mutants showed a higher rate of MSI compared to wild-type cells (Guo et al. 2004). Since expression of MMR components was unaltered, the consequences of DNMT1 deficiency appeared to be direct rather than an indirect consequence of genomic hypomethylation or global changes in gene activity (Guo et al. 2004). In several independent studies, similar roles for DNMT1 were also reported (Gorbunova et al. 2004; Kim et al. 2004; Wang and James Shen 2004). In these reports, DNMT1 deficiency resulted in a significant increase in MSI for both endogenous microsatellite loci (Wang and James Shen 2004) and transgenic slippage reporter constructs (Kim et al. 2004). In a genetic selection assay for trinucleotide repeat contractions, the DNA methyltransferase inhibiting drugs 5-aza-2′-deoxycytidine (5-azadC) and hydralazine further corroborated these findings—treatment with 5-azadC produced a 1,000-fold increase in the rate of MSI (Gorbunova et al. 2004). Although this selection assay was biased for the identification of contraction mutants, additional experiments utilizing human fibroblasts derived from myotonic dystrophy patients also showed large expansions of repeat tracts in response to 5-azadC treatment. These differing results hint at an underlying complexity in the involvement of DNMT1 in MSI.
One suggestion for the influence of DNMT1 on MSI is that microsatellite methylation provides a mechanism for length stabilization by subsequent transcriptional repression of genes containing or proximal to microsatellites with methylated CpG repeats (Table 1). Transcription near or through microsatellites exposes single-stranded DNA to secondary structure formation that may be erroneously resolved through MMR (Lin et al. 2006; Lin and Wilson 2007). DNA methylation may stabilize these sequences by preventing them from being transcribed. In each of the above-mentioned analyses for MSI, non-CpG-containing repeat sequences were examined, thus DNA hypomethylation of the repetitive sequence itself is likely not the cause of MSI (Gorbunova et al. 2004; Kim et al. 2004; Wang et al. 2004). Furthermore, in one study, methylation levels of flanking DNA, which may also assist in stabilizing repeat sequences, did not differ between wild type and DNMT1 mutant cells (Kim et al. 2004). In other analyses, however, CpG content was implicated in MSI. For CpG-containing human repeat sequences transfected into primate cells, DNA methylation stabilized CGG repeats during replication in instances where DNA was premethylated prior to transfection (Nichol Edamura et al. 2005). CpG methylation of these microsatellites significantly decreased the frequency of length change events and the magnitude of those changes (Nichol Edamura et al. 2005). CpG methylation of regions adjacent to microsatellites also contributed to repeat stabilization (Brock et al. 1999; Nichol and Pearson 2002). In one analysis which examined ten independent CAG-repeat-containing (non-CpG-containing) endogenous loci, differences in instability among the loci were noted (Brock et al. 1999). These differences were independent of microsatellite length but were correlated with flanking CpG content (Brock et al. 1999). Thus, in some cases, CpG content of microsatellites or neighboring cis-sequence does appear to play a role in stabilizing repeat length.
DNMT1 also influences transcriptional repression and MSI through chromatin remodeling (Table 1). Following DNA replication, DNMT1 mediates transcriptional repression and chromatin condensation through the histone deacetylases HDAC1 and HDAC2 (Fuks et al. 2000; Robertson et al. 2000; Rountree et al. 2000). Enhanced trinucleotide repeat slippage in DNMT1 mutants was correlated with an increase in H3 acetylation at the repeat, a mark of transcriptional permissiveness (Kim et al. 2004), and in a Drosophila model of repeat instability; reduced activity of the histone acetyltransferase CREB-binding protein (CBP) was responsible for CAG repeat expansion (Jung and Bonini 2007). Histone acetylation, however, cannot be entirely responsible for MSI as the HDAC inhibitor sodium butyrate produced only minimal destabilization of a trinucleotide repeat sequence (Gorbunova et al. 2004). Furthermore, contrary to arguments that transcriptional repression of microsatellite regions protects against MSI, overexpression of the HDAC SIRT1, which functions in gene silencing through chromatin remodeling, was linked to CpG island hypermethylation and high MSI in cancer cells (Nosho et al. 2009). This latter observation may allude to an indirect mechanism of global gene hypermethylation and gene silencing on MSI (as discussed below), and highlights the complexity of the links between epigenetic marks and MSI.
In addition to promoting microsatellite stability through transcriptional repression, another means by which DNMT1 promotes repeat stability is through a direct relationship between DNA methylation and MMR (Table 1). DNA methylation has been suggested to provide a mechanism for strand recognition following DNA replication—hemimethylated DNA generated by DNA replication may facilitate the MMR machinery in distinguishing the nascent strand from the template strand (Hare and Taylor 1985). The methyl CpG-binding protein MBD4/MED1 may provide the functional link between MMR and strand recognition. MBD4 co-localizes with foci of methylated DNA and was identified in a yeast 2-hybrid screen as interacting with the MMR component MLH1 (Bellacosa et al. 1999; Hendrich and Bird 1998). Although the mechanistic relationship between MBD4 and the MMR machinery remains undefined, MBD4 contains glycosylase repair activities with a preference for hemi-methylated CpG/CpT mismatches, thus providing a direct functional link between DNA methylation and MMR (Hendrich et al. 1999).
Yet another potential functional link between DNMT1 and MMR was revealed when DNMT1 was shown to interact with proliferating cell nuclear antigen (PCNA), the DNA polymerase processivity factor (Chuang et al. 1997). Loss of PCNA causes MSI, likely due to its function in DNA repair (Baida et al. 2003). PCNA promotes both MMR and nucleotide-excision repair through its interaction with the MMR factors MLH1 and MSH2, and PCNA rapidly recruits DNMT1 to sites of DNA repair through a direct interaction with DNMT1 (Mortusewicz et al. 2005; Nichols and Sancar 1992; Umar et al. 1996). Although the physical mechanism by which DNMT1 and these MMR proteins facilitate microsatellite stability is not known, the association between these components and MSI is quite strong.
Microsatellite instability and human disease
Epigenetics, MMR, and cancer
An alternate, indirect mechanism in which genomic destabilization occurs is through DNA methylation-induced global changes in gene expression that alter transcriptional regulation of proteins responsible for repeat stabilization (Table 1), an effect that has been observed in cancer. In general, the genome of tumor cells is hypomethylated, particularly in regions of repetitive DNA. Widespread hypomethylation contributes to a hyper-mutation state in cancer, which may generate oncogenic mutations or loss of heterozygosity of tumor suppressor genes. Additionally, tumor cells also demonstrate sites of local hypermethylation, frequently at CpG islands that would, under normal circumstances, be hypomethylated (Robertson and Wolffe 2000). Site-specific CpG island hypermethylation affects genomic stability when it targets promoters of MMR genes. Along these lines, MSI is a common pathway to tumor development, and many MSI tumors contain mutations in MMR genes or epigenetic repression of MMR genes (Laghi et al. 2008).
An underlying methylation defect has been described in a subset of colorectal cancers in which promoter CpG islands are hypermethylated at high frequency, a condition termed the CpG island methylator phenotype (CIMP; Issa 2004; Toyota et al. 1999). One class of CIMP tumors displays MSI and concurrent inactivation of MLH1 by promoter hypermethylation (Issa 2004). The first evidence of CIMP came from the finding that colorectal tumors lacking MLH1 expression displayed MLH1 promoter hypermethylation but no MLH1 mutations (Kane et al. 1997), and subsequent reports have characterized MSI tumors as having a strong association with promoter hypermethylation compared to non-MSI tumors (Ahuja et al. 1997; Veigl et al. 1998). Moreover, experiments introducing a transgenic reporter gene into cancer cell lines revealed that expression of the reporter was associated with genetic instability: upon introduction of the transgene, cell lines deficient in MMR showed strong repression of the reporter gene compared to MMR proficient lines (Lengauer et al. 1997). The contrasting behavior of the transgene was linked to cell line-specific methylation differences that were negated by 5-azadC treatment, rescuing transgene expression (Lengauer et al. 1997). These reports are consistent with the idea that CIMP is associated with increased DNMT1 expression or enhanced DNA methyltransferase activity (Teodoridis et al. 2008). Additional evidence from experiments in mice has further highlighted the importance of DNA methyltransferase activity in CIMP tumors: mice deficient for Mlh1 expression due to Mlh1 promoter hypermethylation were less susceptible to intestinal tumor formation when Dnmt1 function was also reduced (Trinh et al. 2002). Interestingly, although Mlh1/Dnmt1-deficient mice were less susceptible to intestinal tumorigenesis, they were more susceptible to lymphoid tumor development, a result linked to hypomethylation-induced genomic instability (Trinh et al. 2002). In Dnmt1-deficient mice, a similar phenotype was observed including genomic hypomethylation, increased lymphomagenesis, and marked genomic instability, particularly trisomy of chromosome 15 (Gaudet et al. 2003). Thus, DNA methylation is a double-edged sword in maintaining proper growth controls in that too much methylation enhances tumor-associated repression of repair genes, but too little methylation enhances genomic instability in repetitive regions that require a heterochromatic state for their stability. Obtaining a better understanding of the mechanisms of maintaining the proper balance and distribution of methyl marks in normal tissues will be critical for understanding how disruption of this balance contributes to cancer.
Other human disease associations with MSI
Trinucleotide repeat expansion is a hallmark of more than 20 human diseases including fragile X syndrome, Huntington's disease, and myotonic dystrophy (Mirkin 2007). These diseases are characterized by an “anticipation” phenomenon in which microsatellite expansion and disease severity worsen with each generation (Mirkin 2007). DNA methylation defects may directly contribute to the etiology of some repeat expansion diseases, because the developmental stages in which MSI occurs coincide with two key periods during which epigenetic reprogramming occurs (Gorbunova et al. 2004; Pearson 2003; Reik et al. 2001; Yoon et al. 2003). Moreover, DNA methylation was linked to stabilization of the CGG repeats whose expansion causes fragile X syndrome (Wohrle et al. 1998).
A key question is how do repeat expansions cause disease? One mechanism is through disruption of gene function: microsatellite expansions residing in exon or intron sequences disrupt the normal function of the gene product. Repeat expansions also produce local epigenetic changes. These expansions disrupt local chromatin structure and alter the expression of adjacent genes, an effect that has been extensively studied in myotonic dystrophy patients (Gatchel and Zoghbi 2005; Mirkin 2007). Myotonic dystrophy is associated with expansion of a CAG repeat within the 3′UTR of the DMPK gene. Repeat length is inversely correlated with expression of the flanking genes DMPK and SIX5/DMAHP, both of which have been implicated in the pathology of myotonic dystrophy (Klesert et al. 2000, 1997; Thornton et al. 1997). Analysis of chromatin from skeletal muscle of individuals with myotonic dystrophy showed that the region surrounding the CAG expansion was protected from DNase digestion (Otten and Tapscott 1995), presumably due to a change in chromatin structure resulting in chromatin condensation (Otten and Tapscott 1995). The normal CAG locus was also shown to contain a region of H3K9 methylated heterochromatin embedded in a euchromatic H3K4 methylated region (Cho et al. 2005). The expanded repeat allele, in contrast, was associated with spreading of heterochromatin (and H3K9 methylation) into gene containing regions. Heterochromatin boundaries are normally limited by binding of the zinc-finger insulator protein CTCF; CTCF binding, however, is lost at the CAG boundaries in myotonic dystrophy (Cho et al. 2005; Filippova et al. 2001). Methylation of CTCF DNA binding sites prevents CTCF binding, which may, in turn, permit the spread of heterochromatin into surrounding euchromatic regions (Filippova et al. 2001).
Epigenetic modifications and DNA damage
In addition to repeat stabilization, DNA methylation and chromatin structure strongly influence other forms of genomic instability such as mutations and chromosomal rearrangements. For example, hypomethylation in Dnmt1-deficient murine ES cells produced a striking increase in gene mutations, deletions, and chromosomal deficiencies (Chen et al. 1998). Likewise, knockout of DNMT1 and DNMT3B in a human colon cancer cell line caused large-scale, randomly occurring chromosomal translocations and aneuploidy (Karpf and Matsui 2005).
DNA methylation and homologous recombination
DNA methylation is well known to protect against unlicensed homologous recombination (HR), which is the basis for many mutational events. Following generation of a DNA double-strand break (DSB), HR is employed to form a synapse between homologous chromosomes, a circumstance that can generate gene conversions, insertions or deletions, or more extensive chromosomal loss. Transcription enhances HR frequency by exposing single-stranded DNA and allowing it to invade its homologous sequence, while DNA methylation-mediated transcriptional silencing inhibits HR (Dominguez-Bendala and McWhir 2004; Ikeda and Matsumoto 1979; Maloisel and Rossignol 1998). Several reports support the idea that DNMT1 represses HR. Dnmt1 deficiency in mice resulted in elevated loss of heterozygosity due to mitotic recombination (Eden et al. 2003), Dnmt1 deficiency in mouse ES cells enhanced gene targeting by HR (Dominguez-Bendala and McWhir 2004), and embryonic carcinoma cells carrying integrated transgenic reporter constructs displayed increased recombination among transgenic sequences in response to depletion of DNA methylation by 5-azadC (McBurney et al. 2001). In a complementary study, hypermethylation of CpG islands within extrachromosomal fragments reduced the occurrence of deletions within and dimerizations between fragments, presumably due to reduced HR (Rizwana and Hahn 1999). Finally, DNMT1 and DNMT3 activity at repetitive pericentromeric and centromeric loci effectively limits mitotic recombination frequency and stabilizes centromeric repeats against length changes (Jaco et al. 2008).
Role of chromatin and DNA methyltransferases in DNA damage repair
In addition to the role of DNA methylation in prevention of HR, DNMT1 unexpectedly performs a role in DSB repair that appears to be independent of its maintenance methylation activity. DNMT1 has been shown to either assist in DSB detection or operate in the repair process itself. Induction of DSBs by irradiation or by 5-azadC shows an accumulation of DNMT1, its binding partners PCNA and MBD4, and the repair-associated factor MLH1 at foci of DSBs, and DNMT1 deficiency reduces the recruitment of DNA repair proteins to these foci (Mortusewicz et al. 2005; Palii et al. 2008; Ruzov et al. 2009). The well-established PCNA-DNMT1 interaction has recently been shown to be nonessential for DNMT1's maintenance methylation function (Schermelleh et al. 2007; Spada et al. 2007), and DNMT1 associates with chromatin independently of replication (Easwaran et al. 2004), suggesting a role for the PCNA-DNMT1 interaction separate from maintenance methylation. PCNA acts in multiple DNA repair processes including MMR and nucleotide excision repair, and thus DNMT1 may facilitate this function (Mortusewicz et al. 2005; Nichols and Sancar 1992; Umar et al. 1996). In summary, DNMT1 functions in the DNA damage response, perhaps through a methylation-independent role, highlighting an important and novel function for DNMT1 that deserves further examination.
DSB repair must operate in the context of complex higher-order chromatin structure and orchestrate restoration of chromatin and epigenetic marks at the site of DNA damage. Chromatin remodeling, like repair of the DNA itself, is critical for maintaining genomic integrity. Decondensation of chromatin in response to DNA damage may be required to loosen or open the chromatin structure and allow access for repair machinery (Ziv et al. 2006). Consistent with this notion, in Saccharomyces cerevisiae, HATs, including Gcn5 and Esa1, are actively recruited to sites of DSBs presumably to assist in promoting open chromatin (Groth et al. 2007; Tamburini and Tyler 2005; van Attikum and Gasser 2005). This activity is only transient, and HDACs rapidly remove acetylation marks likely to prevent aberrant transcriptional activity in the newly repaired region (Tamburini and Tyler 2005). Chromatin remodeling following DNA damage is also mediated by the DNA damage checkpoint kinase CHK1. CHK1 is normally associated with chromatin in undamaged cells and specifically phosphorylates histone H3 at threonine 11 (H3T11) promoting acetylation of H3K9 and continued maintenance of cell cycle regulatory genes in a transcriptionally active state. Upon DNA damage, CHK1 dissociates from chromatin, and levels of phosphorylated H3T11 and acetylated H3K9 decline (Shimada et al. 2008; Smits et al. 2006). This deacetylation may function to repress transcription at the site of damage, providing protection for the decondensed chromatin against further vulnerability to mutations during the repair process.
Another critically important histone modification associated with DNA damage is phosphorylation of H2A, particularly at serine 129 and serine 1 (van Attikum and Gasser 2005). Recruitment of phosphorylated H2A.X (γH2A.X), a variant of histone H2A, to sites of DSBs is a critical step in the repair process (Celeste et al. 2003, 2002). γH2A.X-containing nucleosomes assemble along the regions bordering DSBs, as observed by immunofluorescence microscopy, and aid in the recruitment and/or maintenance of DNA break repair factors and histone modifiers at the site of damage (Rogakou et al. 2000, 1998; van Attikum and Gasser 2005). Notably, recruitment of γH2A.X containing histones to sites of DSBs is also DNMT1 dependent (Palii et al. 2008). In addition, in both fertilized zygotes and cloned embryos, the occurrence of γH2A.X-populated DSBs increased in a temporal pattern that coincided remarkably well with the period of active, DNA replication-independent demethylation during zygotic pronuclear development (Wossidlo et al. 2010). Although the mechanism underlying this connection is unclear, the association between these two events and the requirement for DNMT1 in the recruitment of γH2A.X is consistent with a functional involvement of DNA methylation in DNA repair. It will be of great interest to determine the role of DNMT1 at sites of DNA damage and whether it is methylation dependent, as this may provide important information on how DNA methylation inhibitors exert their antitumor effects.
Chromatin modifications and DNA methylation in centromere function
Epigenetic determinants of centromere identity
A functional centromere is vital for guarding against chromosomal instability during cell division by maintaining proper DNA segregation, averting chromosomal rearrangements, and preventing changes in ploidy. Cytologically, the centromere appears as a chromosomal constriction, but it actually consists of a molecularly unique and complex configuration of chromatin. Centromeric chromatin forms a densely compact structure that provides attachment sites for kinetochore proteins and spindle microtubules. The flanking pericentromeric chromatin recruits a high concentration of cohesin molecules providing the site for sister chromatid attachment (Allshire and Karpen 2008).
With every cell generation, each chromosome's centromere is habitually re-generated at the same location. Providing a single site for kinetochore attachment is necessary to ensure proper chromosomal segregation and stability, and numerous lines of evidence have established that centromere location/identity is determined epigenetically (Allshire and Karpen 2008; Amor and Choo 2002; Henikoff et al. 2001; Karpen and Allshire 1997; Sullivan et al. 2001). For example, cross species comparison shows that centromeric DNA sequences vary widely suggesting the lack of a single centromere recognition sequence. Likewise, human α-satellite repeat sequence within centromeric DNA is insufficient to generate a functional centromere, since chromosomes that contain two regions of centromeric α-satellite DNA, still possess only one functional kinetochore attachment region. Additionally, in the absence of a functional centromere, a neocentromere, which is a functional ectopic centromere, may form at alternate non-centromeric genetic loci that are devoid of α-satellite repeats (Amor and Choo 2002; Choo 2000). Having epigenetic centromeric determinants as opposed to sequence-specific determinants is advantageous because it provides a reliable mechanism for transmitting genetic material in the event of mutational damage to the centromere locus. Epigenetic determination also allows for adaptive karyotypic evolution and may provide an important mechanistic means for speciation (Henikoff et al. 2001). Thus, the epigenetically determined centromere provides a versatile system for consistent and faithful transmission of chromosomal material and for accommodation of karyotype alterations.
Human centromeres are characterized by several thousand kilobases of a 171-base pair AT-rich repeat referred to as α-satellite DNA (Amor and Choo 2002). Satellite DNA sequences typically do not possess similarity among chromosomes or across species, but their DNA-binding proteins are frequently conserved. One such important conserved centromeric chromatin protein is CENP-A, a centromere-specific core histone that substitutes for histone H3 in the nucleosome core (Palmer et al. 1987). Centromeric chromatin is composed of alternating blocks of H3 and CENP-A-containing nucleosomes (Blower et al. 2002). The structural significance of this CENP-A distribution is not known, but it may impart directionality onto the centromere structure that is necessary for kinetochore attachment and for bi-directional, anaphase migration of sister chromatids (Allshire and Karpen 2008). Importantly, CENP-A is necessary and sufficient for recruitment of kinetochore proteins for mitosis (Palmer et al. 1991; Palmer et al. 1987). CENP-A deficiency disrupts kinetochore assembly and progression of mitosis, whereas CENP-A overexpression induces ectopic spreading of centromeric chromatin to non-centromeric locations and subsequent redistribution of kinetochore proteins (Allshire and Karpen 2008; Sullivan et al. 2001). Interestingly, CENP-A is recruited to DSBs, and cell survival following generation of DSBs is proportional to CENP-A activity (Zeitlin et al. 2009). Thus, CENP-A is multifaceted in its involvement in genomic maintenance and stability. H2A.Z, another histone variant replacing H2A in non-CENP-A-containing centromeric and pericentromeric nucleosomes, is also required for centromere structure and sister chromatid adhesion prior to chromosomal segregation (Greaves et al. 2007). The interplay between CENP-A and H2A.Z histone variants along with their unique arrangements of histone modifications (see below) appear to be key determinants in the organization and function of the centromere.
Histone modifications in centromere function and chromosomal condensation
Centromeric and pericentromeric chromatin contain unique and largely non-overlapping configurations of histone marks that are critical for centromere function (Fig. 1). Interestingly, despite the highly condensed nature of centromeric chromatin, its array of histone modifications is quite distinct from constitutive heterochromatin. Centromeric heterochromatin contains a characteristic combination of both euchromatic and heterochromatic marks (Lam et al. 2006; Sullivan and Karpen 2004). For example, condensed transcriptionally inert heterochromatin is defined by H3K9 di- and tri-methylation, but immunofluorescence studies showed that centromeric chromatin, despite being highly condensed, does not contain these marks (Sullivan and Karpen 2004). Surprisingly, CENP-A containing centromeric chromatin contains H3K4me2, a mark associated with euchromatin and transcriptional permissiveness (Sullivan and Karpen 2004). However, other marks which confer an “open” euchromatic and transcriptionally active signature, such as H3K4me3 and histone acetylation, are absent from centromeres (Sullivan and Karpen 2004). Pericentromeric heterochromatin, on the other hand, more closely resembles canonical heterochromatin. It is hypoacetylated and enriched for both H3K9me2 and H3K9me3; however, H3K9me3 domains are located distally from the centromere, a configuration that may function to establish centromeric boundaries (Sullivan and Karpen 2004). Consistent with a functional role for H3K9 trimethylation in pericentromeric heterochromatin, the H3K9 methylases, SUV39H1 and SUV39H2, are critical for mitotic progression. SUV39H1 and SUV39H2 accumulate at centromeric domains during mitosis, and their loss, or the loss of H3K9 methylation, in pericentromeric regions is detrimental to chromosome segregation (McManus et al. 2006; Melcher et al. 2000; Peters et al. 2003, 2001; Rice et al. 2003). Also within pericentromeric domains, two additional histone methylases, SUV4-20H1 and SUV4-20H2, were shown to recognize H4K20 substrates for methylation, a modification necessary for chromosomal condensation and stability during mitosis (Oda et al. 2009; Schotta et al. 2004). SUV39H1/2 and SUV4-20H1/2 enzymes are also associated with recruitment and stabilization of heterochromatic protein 1 (HP1) to pericentromeric chromatin to mediate gene silencing and chromosomal condensation (Bannister et al. 2001; Lachner et al. 2001; Maison et al. 2002; Schotta et al. 2004; Souza et al. 2009). Suv39H1/2 activity is also important for CENP-A assembly into centromeric chromatin, since the fission yeast homolog of SUV39, Clr4, along with HP1 and other heterochromatin associated factors, is necessary for loading CENP-A nucleosomes into centromeric regions (Folco et al. 2008). These studies emphasize how critical the distinctive patterns of centromeric and pericentromeric histone marks and their associated methylases are for the proper recruitment and stabilization of centromeric proteins. Global changes in histone methylation are also critical for whole chromosomal condensation and centromere function during mitosis. Inhibition of overall histone methyltransferase activity just prior to prophase causes defects in chromosome alignment and segregation due to expansions and loosening of centromeric chromatin, unstable microtubule associations with kinetochores, and improper kinetochore protein localization (Heit et al. 2009).
Phosphorylation of serine and threonine residues, particularly on histones H1 and H3, is another modification critical for chromosomal condensation during cell division. Both histone H1 and H3 become phosphorylated upon entering mitosis and are dephosphorylated upon exiting mitosis (Gurley et al. 1978; Nowak and Corces 2004). The accumulation pattern of H3S10 phosphorylation is consistent with its requirement during mitosis given that it is initiated within pericentromeric regions during late G2 phase and is later removed during anaphase (Hendzel et al. 1997; Van Hooser et al. 1998; Wei et al. 1998, 1999). Absence of H3S10 phosphorylation yields incomplete chromosome condensation and defects in chromosomal segregation (Hsu et al. 2000; Wei et al. 1998, 1999). CENP-AS7 is phosphorylated similarly to H3S10, and the residues surrounding CENP-AS7 share similarity with those surrounding H3S10, suggesting functional conservation (Zeitlin et al. 2001a). Aurora kinases are responsible for H3S10 and CENP-AS7 phosphorylation, and inhibition of their activity results in loss of pericentromeric H3S10P (Kunitoku et al. 2003; Monier et al. 2007). Aurora kinase phosphorylation of H3S10 recruits condensin to centromeric and pericentromeric regions to promote chromosomal condensation and rigidity (Giet and Glover 2001; Hsu et al. 2000; Ono et al. 2004; Zeitlin et al. 2001b). One of the critical factors recruited by H3S10P is HP1α. Binding of the heterochromatic factor HP1α at centromeric and pericentromeric marks throughout the cell cycle may be affected by the size of the H3S10P expression domain (Monier et al. 2007).
Phosphorylation of H2AS1 and H4S1 is also closely associated with mitosis and is similarly timed with the presence of H3S10P (Xu et al. 2009). Likewise, H3T3 shows patterns of mitosis-specific phosphorylation with particularly strong accumulation in centromeric regions during chromatin condensation (Dai et al. 2006, 2005; Eswaran et al. 2009; Polioudaki et al. 2004). H3T3 phosphorylation is generated by the kinase Haspin in regions devoid of H3K4 methylation, and its deficiency causes reduced sister chromatid cohesion and defects in metaphase chromosomal misalignment (Dai et al. 2006, 2005; Eswaran et al. 2009).
Another important chromatin configuration required for chromosomal condensation is the removal of histone acetylation via the HDACs (David et al. 2003). Acetylation is largely a mark of euchromatin and transcriptional permissiveness, and aberrant acetylation during mitosis reduces chromosome condensation and impairs sister chromatid separation (Cimini et al. 2003; Jeppesen et al. 1992). Inhibition of HDACs also causes loss of transcriptional repression in centromeric regions, chromosomal segregation defects, and chromosomal loss in fission yeast (Ekwall et al. 1997). Importantly, in fission yeast, the chromosomal condensation and segregation defects caused by HDAC inhibition persist even when HDAC activity is restored, suggesting the presence of an inherited mechanism for replicating mitotic deacetylation activity (Ekwall et al. 1997). In mammalian centromeres, histone deacetylase activity relies upon DNA methylation since DNMT1 is required for maintenance of histone deacetylation in pericentromeric domains (Xin et al. 2004).
DNA methylation in centromere function and kinetochore assembly
Centromeric and pericentromeric DNA is heavily methylated at CpG dinucleotides, a status that persists throughout the cell cycle and is important for centromere function (Monier et al. 2007; Wong et al. 2006). The requirement for this centromere-specific methylation is quite apparent in diseases characterized by chromosomal segregation defects, such as some cancers and ICF syndrome. These diseases, which demonstrate large-scale gain or loss of chromosomes, are often associated with loss of DNA methylation at centromeric and/or pericentromeric loci (Gisselsson et al. 2005; Hansen et al. 1999; Lengauer et al. 1997; Wong et al. 2001). For example, a subset of hepatocellular carcinomas has been defined by particular chromosome 1 rearrangements stemming from a pericentromeric breakpoint, which was highly correlated with hypomethylation of pericentromeric satellite-2 CpG dinucleotides (Wong et al. 2001). The requirement for DNA methylation is also apparent in patients with ICF syndrome. ICF syndrome patients exhibit hypomethylation of heterochromatin, particularly of pericentromeric satellite-2 repeats within chromosomes 1, 9, and 16, resulting in decondensation of pericentromeric chromatin (Gisselsson et al. 2005; Hansen et al. 1999; Miniou et al. 1994, 1997). DNMT3B's PWWP domain is essential for methylation of these repeats, and significantly, the majority of ICF syndrome patients have mutations in DNMT3B that at least partially disrupt its methylation activity (Chen et al. 2004; Hansen et al. 1999; Xu et al. 1999).
The effects of DNA methylation and DNMT activity in pericentromeric and centromeric heterochromatin formation appear to be mediated through histone-modifying enzymes, heterochromatin factors, and centromeric proteins. Depletion of DNMT1 prevents accumulation of H3S10P foci, perhaps by affecting the subnuclear localization or the binding affinity of Aurora kinases to the centromere (Monier et al. 2007). DNMT3B also facilitates centromeric heterochromatin formation and chromosomal condensation through an association with condensin complexes (Geiman et al. 2004). DNMT3A and DNMT3B are recruited by H3K9me3-containing nucleosomes and SUV39H1 to pericentromeric domains, where they also interact with the heterochromatin factors HP1β and HP1α (Fuks et al. 2003; Geiman et al. 2004; Lehnertz et al. 2003).
DNA methylation and centromere function are further linked by the recent finding that DNMT3B interacts with the constitutive centromere protein CENP-C. CENP-C is an essential kinetochore protein that binds centromeric α-satellite DNA and is required for kinetochore assembly during mitosis (Kalitsis et al. 1998; Kwon et al. 2007; Politi et al. 2002). DNMT3B and CENP-C interact and become co-enriched at centromeric foci, reaching peak enrichment during metaphase (Gopalakrishnan et al. 2009). Importantly, this co-enrichment requires the mutual activity of both factors, since siRNA knockdown of either DNMT3B or CENP-C reduces the other's binding to centromeric and pericentromeric regions. This enrichment also facilitates DNMT3B's enzymatic activity, since CENP-C knockdown resulted in significant reduction in DNA methylation at both centromeric α-satellite DNA and pericentromeric satellite-2 sequence. Both CENP-C and DNMT3B are required for HP1α recruitment to centromeric and pericentromeric regions, and their loss affects chromosomal segregation and mitotic progression. Based on these findings, it is possible that loss of centromeric CENP-C and HP1α recruitment in DNMT3B-deficient ICF patients contributes to the marked centromeric instability that is a hallmark of this disorder (Gopalakrishnan et al. 2009).
Importantly, DNA methylation within the centromere is not entirely ubiquitous and unrestrained, but it instead consists of specific non-methylated sites within extensively methylated regions. Non-methylated regions are particularly important for the kinetochore protein CENP-B. CENP-B, which interacts with both CENP-A and CENP-C, binds a specific 17-base pair sequence within α-satellite DNA and provides an important mechanism for centromere identity (Ando et al. 2002; Masumoto et al. 1989). Loss of CENP-B does not affect kinetochore assembly or function (Choo 2001; Hudson et al. 1998; Kapoor et al. 1998; Perez-Castro et al. 1998), but it instead functions to effectively limit multi-centromere formation and promote heterochromatin formation via SUV39H1 recruitment and DNA methylation (Okada et al. 2007). The centromeric pattern of methylation consisting of unmethylated CENP-B binding sites intermingled with extensively methylated regions is important for these functions since aberrant DNA methylation or 5-azadC treatment disrupts binding of CENP-B to these loci and alters its distribution (Miniou et al. 1997; Mitchell et al. 1996; Okada et al. 2007; Tanaka et al. 2005).
Interestingly, centromeric and pericentromeric DNA methylation is distinctive in that its function is not simply to promote widespread transcriptional silencing. Despite the dense DNA methylation in these regions, these genetic loci show signs of transcriptional activity (Lehnertz et al. 2003). Analysis of DNA methylation of neocentromeres showed that most CpGs are hypermethylated, except for small domains of hypomethylation corresponding to transcriptionally permissive sites (Wong et al. 2006). Furthermore, centromeres on human artificial chromosomes showed loss of function when transcription within the centromere was altered (Nakano et al. 2008). Either strong transcriptional repression or broad transcriptional activation caused loss of CENP-A, -B, and -C binding, loss of kinetochore assembly, and chromosomal missegregation (Nakano et al. 2008). Thus, studies using artificial chromosomes and neocentromeres suggest that a delicate balance between transcriptional activity and repression is required to maintain centromere functionality.
In summary, DNA methyltransferases affect chromosomal stability through DNA methylation, chromatin remodeling, and recruitment of heterochromatin specific factors. Genetic loss-of-function experiments involving the DNA methyltransferases have only just begun to elucidate their pervasive effects on genomic stability and the crosstalk between DNA methylation and the histone code. Our understanding of how DNA methylation and the histone code affect each other to differentially establish functional, region-specific chromatin structure is still in its early stages, but it is increasingly evident that these epigenetic marks are critical for genomic stability and disruption of these mechanisms contributes to a growing number of human malignancies.
Ahuja N, Mohan AL, Li Q, Stolker JM, Herman JG, Hamilton SR, Baylin SB, Issa JP (1997) Association between CpG island methylation and microsatellite instability in colorectal cancer. Cancer Res 57:3370–3374
Allshire RC, Karpen GH (2008) Epigenetic regulation of centromeric chromatin: old dogs, new tricks? Nat Rev Genet 9:923–937
Amor DJ, Choo KH (2002) Neocentromeres: role in human disease, evolution, and centromere study. Am J Hum Genet 71:695–714
Ando S, Yang H, Nozaki N, Okazaki T, Yoda K (2002) CENP-A, -B, and -C chromatin complex that contains the I-type alpha-satellite array constitutes the prekinetochore in HeLa cells. Mol Cell Biol 22:2229–2241
Azuara V, Perry P, Sauer S, Spivakov M, Jorgensen HF, John RM, Gouti M, Casanova M, Warnes G, Merkenschlager M et al (2006) Chromatin signatures of pluripotent cell lines. Nat Cell Biol 8:532–538
Baida A, López A, Marcos R, Velázquez A (2003) Germline mutations at microsatellite loci in homozygous and heterozygous mutants for mismatch repair and PCNA genes in Drosophila. DNA Repair (Amst) 2:827–833
Bannister AJ, Zegerman P, Partridge JF, Miska EA, Thomas JO, Allshire RC, Kouzarides T (2001) Selective recognition of methylated lysine 9 on histone H3 by the HP1 chromo domain. Nature 410:120–124
Bellacosa A, Cicchillitti L, Schepis F, Riccio A, Yeung AT, Matsumoto Y, Golemis EA, Genuardi M, Neri G (1999) MED1, a novel human methyl-CpG-binding endonuclease, interacts with DNA mismatch repair protein MLH1. Proc Natl Acad Sci USA 96:3969–3974
Bernstein BE, Humphrey EL, Erlich RL, Schneider R, Bouman P, Liu JS, Kouzarides T, Schreiber SL (2002) Methylation of histone H3 Lys 4 in coding regions of active genes. Proc Natl Acad Sci USA 99:8695–8700
Bernstein BE, Mikkelsen TS, Xie X, Kamal M, Huebert DJ, Cuff J, Fry B, Meissner A, Wernig M, Plath K et al (2006) A bivalent chromatin structure marks key developmental genes in embryonic stem cells. Cell 125:315–326
Bestor TH (1992) Activation of mammalian DNA methyltransferase by cleavage of a Zn binding regulatory domain. EMBO J 11:2611–2617
Bird AP (1986) CpG-rich islands and the function of DNA methylation. Nature 321:209–213
Bird A (1999) DNA methylation de novo. Science 286:2287–2288
Blower MD, Sullivan BA, Karpen GH (2002) Conserved organization of centromeric chromatin in flies and humans. Dev Cell 2:319–330
Bowen NJ, Palmer MB, Wade PA (2004) Chromosomal regulation by MeCP2: structural and enzymatic considerations. Cell Mol Life Sci 61:2163–2167
Boyes J, Bird A (1991) DNA methylation inhibits transcription indirectly via a methyl-CpG binding protein. Cell 64:1123–1134
Brock GJ, Anderson NH, Monckton DG (1999) Cis-acting modifiers of expanded CAG/CTG triplet repeat expandability: associations with flanking GC content and proximity to CpG islands. Hum Mol Genet 8:1061–1067
Celeste A, Petersen S, Romanienko PJ, Fernandez-Capetillo O, Chen HT, Sedelnikova OA, Reina-San-Martin B, Coppola V, Meffre E, Difilippantonio MJ et al (2002) Genomic instability in mice lacking histone H2AX. Science 296:922–927
Celeste A, Difilippantonio S, Difilippantonio MJ, Fernandez-Capetillo O, Pilch DR, Sedelnikova OA, Eckhaus M, Ried T, Bonner WM, Nussenzweig A (2003) H2AX haploinsufficiency modifies genomic stability and tumor susceptibility. Cell 114:371–383
Chedin F, Lieber MR, Hsieh CL (2002) The DNA methyltransferase-like protein DNMT3L stimulates de novo methylation by Dnmt3a. Proc Natl Acad Sci USA 99:16916–16921
Chen RZ, Pettersson U, Beard C, Jackson-Grusby L, Jaenisch R (1998) DNA hypomethylation leads to elevated mutation rates. Nature 395:89–93
Chen T, Tsujimoto N, Li E (2004) The PWWP domain of Dnmt3a and Dnmt3b is required for directing DNA methylation to the major satellite repeats at pericentric heterochromatin. Mol Cell Biol 24:9048–9058
Cho DH, Thienes CP, Mahoney SE, Analau E, Filippova GN, Tapscott SJ (2005) Antisense transcription and heterochromatin at the DM1 CTG repeats are constrained by CTCF. Mol Cell 20:483–489
Chodavarapu RK, Feng S, Bernatavichute YV, Chen PY, Stroud H, Yu Y, Hetzel JA, Kuo F, Kim J, Cokus SJ et al (2010) Relationship between nucleosome positioning and DNA methylation. Nature 466:388–392
Choo KH (2000) Centromerization. Trends Cell Biol 10:182–188
Choo KH (2001) Domain organization at the centromere and neocentromere. Dev Cell 1:165–177
Chuang LS, Ian HI, Koh TW, Ng HH, Xu G, Li BF (1997) Human DNA-(cytosine-5) methyltransferase-PCNA complex as a target for p21WAF1. Science 277:1996–2000
Cimini D, Mattiuzzo M, Torosantucci L, Degrassi F (2003) Histone hyperacetylation in mitosis prevents sister chromatid separation and produces chromosome segregation defects. Mol Biol Cell 14:3821–3833
Colot V, Rossignol JL (1999) Eukaryotic DNA methylation as an evolutionary device. Bioessays 21:402–411
Dai J, Sultan S, Taylor SS, Higgins JM (2005) The kinase haspin is required for mitotic histone H3 Thr 3 phosphorylation and normal metaphase chromosome alignment. Genes Dev 19:472–488
Dai J, Sullivan BA, Higgins JM (2006) Regulation of mitotic chromosome cohesion by Haspin and Aurora B. Dev Cell 11:741–750
David G, Turner GM, Yao Y, Protopopov A, DePinho RA (2003) mSin3-associated protein, mSds3, is essential for pericentric heterochromatin formation and chromosome segregation in mammalian cells. Genes Dev 17:2396–2405
Dominguez-Bendala J, McWhir J (2004) Enhanced gene targeting frequency in ES cells with low genomic methylation levels. Transgenic Res 13:69–74
Easwaran HP, Schermelleh L, Leonhardt H, Cardoso MC (2004) Replication-independent chromatin loading of Dnmt1 during G2 and M phases. EMBO Rep 5:1181–1186
Eden A, Gaudet F, Waghmare A, Jaenisch R (2003) Chromosomal instability and tumors promoted by DNA hypomethylation. Science 300:455
Ekwall K, Olsson T, Turner BM, Cranston G, Allshire RC (1997) Transient inhibition of histone deacetylation alters the structural and functional imprint at fission yeast centromeres. Cell 91:1021–1032
Eswaran J, Patnaik D, Filippakopoulos P, Wang F, Stein RL, Murray JW, Higgins JM, Knapp S (2009) Structure and functional characterization of the atypical human kinase haspin. Proc Natl Acad Sci USA 106:20198–20203
Filippova GN, Thienes CP, Penn BH, Cho DH, Hu YJ, Moore JM, Klesert TR, Lobanenkov VV, Tapscott SJ (2001) CTCF-binding sites flank CTG/CAG repeats and form a methylation-sensitive insulator at the DM1 locus. Nat Genet 28:335–343
Folco HD, Pidoux AL, Urano T, Allshire RC (2008) Heterochromatin and RNAi are required to establish CENP-A chromatin at centromeres. Science 319:94–97
Fuks F, Burgers WA, Brehm A, Hughes-Davies L, Kouzarides T (2000) DNA methyltransferase Dnmt1 associates with histone deacetylase activity. Nat Genet 24:88–91
Fuks F, Hurd PJ, Deplus R, Kouzarides T (2003) The DNA methyltransferases associate with HP1 and the SUV39H1 histone methyltransferase. Nucleic Acids Res 31:2305–2312
Gardiner-Garden M, Frommer M (1987) CpG islands in vertebrate genomes. J Mol Biol 196:261–282
Gatchel JR, Zoghbi HY (2005) Diseases of unstable repeat expansion: mechanisms and common principles. Nat Rev Genet 6:743–755
Gaudet F, Hodgson JG, Eden A, Jackson-Grusby L, Dausman J, Gray JW, Leonhardt H, Jaenisch R (2003) Induction of tumors in mice by genomic hypomethylation. Science 300:489–492
Geiman TM, Sankpal UT, Robertson AK, Chen Y, Mazumdar M, Heale JT, Schmiesing JA, Kim W, Yokomori K, Zhao Y et al (2004) Isolation and characterization of a novel DNA methyltransferase complex linking DNMT3B with components of the mitotic chromosome condensation machinery. Nucleic Acids Res 32:2716–2729
Giet R, Glover DM (2001) Drosophila aurora B kinase is required for histone H3 phosphorylation and condensin recruitment during chromosome condensation and to organize the central spindle during cytokinesis. J Cell Biol 152:669–682
Gisselsson D, Shao C, Tuck-Muller CM, Sogorovic S, Palsson E, Smeets D, Ehrlich M (2005) Interphase chromosomal abnormalities and mitotic missegregation of hypomethylated sequences in ICF syndrome cells. Chromosoma 114:118–126
Gopalakrishnan S, Van Emburgh BO, Robertson KD (2008) DNA methylation in development and human disease. Mutat Res, Fundam Mol Mech Mutagen 647:30–38
Gopalakrishnan S, Sullivan BA, Trazzi S, Della Valle G, Robertson KD (2009) DNMT3B interacts with constitutive centromere protein CENP-C to modulate DNA methylation and the histone code at centromeric regions. Hum Mol Genet 18:3178–3193
Gorbunova V, Seluanov A, Mittelman D, Wilson JH (2004) Genome-wide demethylation destabilizes CTG.CAG trinucleotide repeats in mammalian cells. Hum Mol Genet 13:2979–2989
Greaves IK, Rangasamy D, Ridgway P, Tremethick DJ (2007) H2A.Z contributes to the unique 3D structure of the centromere. Proc Natl Acad Sci USA 104:525–530
Groth A, Rocha W, Verreault A, Almouzni G (2007) Chromatin challenges during DNA replication and repair. Cell 128:721–733
Guo G, Wang W, Bradley A (2004) Mismatch repair genes identified using genetic screens in Blm-deficient embryonic stem cells. Nature 429:891–895
Gurley LR, D'Anna JA, Barham SS, Deaven LL, Tobey RA (1978) Histone phosphorylation and chromatin structure during mitosis in Chinese hamster cells. Eur J Biochem 84:1–15
Hansen RS, Wijmenga C, Luo P, Stanek AM, Canfield TK, Weemaes CM, Gartler SM (1999) The DNMT3B DNA methyltransferase gene is mutated in the ICF immunodeficiency syndrome. Proc Natl Acad Sci USA 96:14412–14417
Hansen KH, Bracken AP, Pasini D, Dietrich N, Gehani SS, Monrad A, Rappsilber J, Lerdrup M, Helin K (2008) A model for transmission of the H3K27me3 epigenetic mark. Nat Cell Biol 10:1291–1300
Hare JT, Taylor JH (1985) One role for DNA methylation in vertebrate cells is strand discrimination in mismatch repair. Proc Natl Acad Sci USA 82:7350–7354
Hata K, Okano M, Lei H, Li E (2002) Dnmt3L cooperates with the Dnmt3 family of de novo DNA methyltransferases to establish maternal imprints in mice. Development 129:1983–1993
Heintzman ND, Stuart RK, Hon G, Fu Y, Ching CW, Hawkins RD, Barrera LO, Van Calcar S, Qu C, Ching KA et al (2007) Distinct and predictive chromatin signatures of transcriptional promoters and enhancers in the human genome. Nat Genet 39:311–318
Heit R, Rattner JB, Chan GK, Hendzel MJ (2009) G2 histone methylation is required for the proper segregation of chromosomes. J Cell Sci 122:2957–2968
Hendrich B, Bird A (1998) Identification and characterization of a family of mammalian methyl-CpG binding proteins. Mol Cell Biol 18:6538–6547
Hendrich B, Hardeland U, Ng HH, Jiricny J, Bird A (1999) The thymine glycosylase MBD4 can bind to the product of deamination at methylated CpG sites. Nature 401:301–304
Hendzel MJ, Wei Y, Mancini MA, Van Hooser A, Ranalli T, Brinkley BR, Bazett-Jones DP, Allis CD (1997) Mitosis-specific phosphorylation of histone H3 initiates primarily within pericentromeric heterochromatin during G2 and spreads in an ordered fashion coincident with mitotic chromosome condensation. Chromosoma 106:348–360
Henikoff S, Ahmad K, Malik HS (2001) The centromere paradox: stable inheritance with rapidly evolving DNA. Science 293:1098–1102
Hsu JY, Sun ZW, Li X, Reuben M, Tatchell K, Bishop DK, Grushcow JM, Brame CJ, Caldwell JA, Hunt DF et al (2000) Mitotic phosphorylation of histone H3 is governed by Ipl1/aurora kinase and Glc7/PP1 phosphatase in budding yeast and nematodes. Cell 102:279–291
Hu J-L, Zhou BO, Zhang R-R, Zhang K-L, Zhou J-Q, Xu G-L (2009) The N-terminus of histone H3 is required for de novo DNA methylation in chromatin. Proc Natl Acad Sci USA 106:22187–22192
Hudson DF, Fowler KJ, Earle E, Saffery R, Kalitsis P, Trowell H, Hill J, Wreford NG, de Kretser DM, Cancilla MR et al (1998) Centromere protein B null mice are mitotically and meiotically normal but have lower body and testis weights. J Cell Biol 141:309–319
Ikeda H, Matsumoto T (1979) Transcription promotes recA-independent recombination mediated by DNA-dependent RNA polymerase in Escherichia coli. Proc Natl Acad Sci USA 76:4571–4575
Issa JP (2004) CpG island methylator phenotype in cancer. Nat Rev Cancer 4:988–993
Jaco I, Canela A, Vera E, Blasco MA (2008) Centromere mitotic recombination in mammalian cells. J Cell Biol 181:885–892
Jaenisch R, Bird A (2003) Epigenetic regulation of gene expression: how the genome integrates intrinsic and environmental signals. Nat Genet 33(Suppl):245–254
Jenuwein T, Allis CD (2001) Translating the histone code. Science 293:1074–1080
Jeppesen P, Mitchell A, Turner B, Perry P (1992) Antibodies to defined histone epitopes reveal variations in chromatin conformation and underacetylation of centric heterochromatin in human metaphase chromosomes. Chromosoma 101:322–332
Jia D, Jurkowska RZ, Zhang X, Jeltsch A, Cheng X (2007) Structure of Dnmt3a bound to Dnmt3L suggests a model for de novo DNA methylation. Nature 449:248–251
Jin B, Tao Q, Peng J, Soo HM, Wu W, Ying J, Fields CR, Delmas AL, Liu X, Qiu J et al (2008) DNA methyltransferase 3B (DNMT3B) mutations in ICF syndrome lead to altered epigenetic modifications and aberrant expression of genes regulating development, neurogenesis and immune function. Hum Mol Genet 17:690–709
Jung J, Bonini N (2007) CREB-binding protein modulates repeat instability in a Drosophila model for polyQ disease. Science 315:1857–1859
Kalitsis P, Fowler KJ, Earle E, Hill J, Choo KH (1998) Targeted disruption of mouse centromere protein C gene leads to mitotic disarray and early embryo death. Proc Natl Acad Sci USA 95:1136–1141
Kane MF, Loda M, Gaida GM, Lipman J, Mishra R, Goldman H, Jessup JM, Kolodner R (1997) Methylation of the hMLH1 promoter correlates with lack of expression of hMLH1 in sporadic colon tumors and mismatch repair-defective human tumor cell lines. Cancer Res 57:808–811
Kapoor M, de Oca M, Luna R, Liu G, Lozano G, Cummings C, Mancini M, Ouspenski I, Brinkley BR, May GS (1998) The cenpB gene is not essential in mice. Chromosoma 107:570–576
Karpen GH, Allshire RC (1997) The case for epigenetic effects on centromere identity and function. Trends Genet 13:489–496
Karpf AR, Matsui S (2005) Genetic disruption of cytosine DNA methyltransferase enzymes induces chromosomal instability in human cancer cells. Cancer Res 65:8635–8639
Kim M, Trinh BN, Long TI, Oghamian S, Laird PW (2004) Dnmt1 deficiency leads to enhanced microsatellite instability in mouse embryonic stem cells. Nucleic Acids Res 32:5742–5749
Kirmizis A, Santos-Rosa H, Penkett CJ, Singer MA, Vermeulen M, Mann M, Bahler J, Green RD, Kouzarides T (2007) Arginine methylation at histone H3R2 controls deposition of H3K4 trimethylation. Nature 449:928–932
Klesert TR, Otten AD, Bird TD, Tapscott SJ (1997) Trinucleotide repeat expansion at the myotonic dystrophy locus reduces expression of DMAHP. Nat Genet 16:402–406
Klesert TR, Cho DH, Clark JI, Maylie J, Adelman J, Snider L, Yuen EC, Soriano P, Tapscott SJ (2000) Mice deficient in Six5 develop cataracts: implications for myotonic dystrophy. Nat Genet 25:105–109
Kouzarides T (2007) Chromatin modifications and their function. Cell 128:693–705
Ku M, Koche RP, Rheinbay E, Mendenhall EM, Endoh M, Mikkelsen TS, Presser A, Nusbaum C, Xie X, Chi AS et al (2008) Genomewide analysis of PRC1 and PRC2 occupancy identifies two classes of bivalent domains. PLoS Genet 4:e1000242
Kunitoku N, Sasayama T, Marumoto T, Zhang D, Honda S, Kobayashi O, Hatakeyama K, Ushio Y, Saya H, Hirota T (2003) CENP-A phosphorylation by Aurora-A in prophase is required for enrichment of Aurora-B at inner centromeres and for kinetochore function. Dev Cell 5:853–864
Kwon MS, Hori T, Okada M, Fukagawa T (2007) CENP-C is involved in chromosome segregation, mitotic checkpoint function, and kinetochore assembly. Mol Biol Cell 18:2155–2168
Lachner M, O'Carroll D, Rea S, Mechtler K, Jenuwein T (2001) Methylation of histone H3 lysine 9 creates a binding site for HP1 proteins. Nature 410:116–120
Laghi L, Bianchi P, Malesci A (2008) Differences and evolution of the methods for the assessment of microsatellite instability. Oncogene 27:6313–6321
Lam AL, Boivin CD, Bonney CF, Rudd MK, Sullivan BA (2006) Human centromeric chromatin is a dynamic chromosomal domain that can spread over noncentromeric DNA. Proc Natl Acad Sci USA 103:4186–4191
Lehnertz B, Ueda Y, Derijck AA, Braunschweig U, Perez-Burgos L, Kubicek S, Chen T, Li E, Jenuwein T, Peters AH (2003) Suv39h-mediated histone H3 lysine 9 methylation directs DNA methylation to major satellite repeats at pericentric heterochromatin. Curr Biol 13:1192–1200
Lengauer C, Kinzler KW, Vogelstein B (1997) Genetic instability in colorectal cancers. Nature 386:623–627
Leonhardt H, Page AW, Weier HU, Bestor TH (1992) A targeting sequence directs DNA methyltransferase to sites of DNA replication in mammalian nuclei. Cell 71:865–873
Lewis JD, Meehan RR, Henzel WJ, Maurer-Fogy I, Jeppesen P, Klein F, Bird A (1992) Purification, sequence, and cellular localization of a novel chromosomal protein that binds to methylated DNA. Cell 69:905–914
Li E (2002) Chromatin modification and epigenetic reprogramming in mammalian development. Nat Rev Genet 3:662–673
Liang G, Chan MF, Tomigahara Y, Tsai YC, Gonzales FA, Li E, Laird PW, Jones PA (2002) Cooperativity between DNA methyltransferases in the maintenance methylation of repetitive elements. Mol Cell Biol 22:480–491
Liang G, Lin JCY, Wei V, Yoo C, Cheng JC, Nguyen CT, Weisenberger DJ, Egger G, Takai D, Gonzales FA et al (2004) Distinct localization of histone H3 acetylation and H3-K4 methylation to the transcription start sites in the human genome. Proc Natl Acad Sci USA 101:7357–7362
Lin Y, Wilson JH (2007) Transcription-induced CAG repeat contraction in human cells is mediated in part by transcription-coupled nucleotide excision repair. Mol Cell Biol 27:6209–6217
Lin Y, Dion V, Wilson JH (2006) Transcription promotes contraction of CAG repeat tracts in human cells. Nat Struct Mol Biol 13:179–180
Maison C, Bailly D, Peters AH, Quivy JP, Roche D, Taddei A, Lachner M, Jenuwein T, Almouzni G (2002) Higher-order structure in pericentric heterochromatin involves a distinct pattern of histone modification and an RNA component. Nat Genet 30:329–334
Maloisel L, Rossignol JL (1998) Suppression of crossing-over by DNA methylation in Ascobolus. Genes Dev 12:1381–1389
Margot J, Ehrenhofer-Murray A, Leonhardt H (2003) Interactions within the mammalian DNA methyltransferase family. BMC Mol Biol 4:7
Masumoto H, Masukata H, Muro Y, Nozaki N, Okazaki T (1989) A human centromere antigen (CENP-B) interacts with a short specific sequence in alphoid DNA, a human centromeric satellite. J Cell Biol 109:1963–1973
Mayer W, Niveleau A, Walter J, Fundele R, Haaf T (2000) Demethylation of the zygotic paternal genome. Nature 403:501–502
McBurney MW, Lau S, Jardine K, Yang X, Davies B (2001) Reexpression of a cluster of silenced transgenes is associated with their rearrangement. Genes Chromosom Cancer 32:311–323
McManus KJ, Biron VL, Heit R, Underhill DA, Hendzel MJ (2006) Dynamic changes in histone H3 lysine 9 methylations: identification of a mitosis-specific function for dynamic methylation in chromosome congression and segregation. J Biol Chem 281:8888–8897
Meehan RR, Lewis JD, McKay S, Kleiner EL, Bird AP (1989) Identification of a mammalian protein that binds specifically to DNA containing methylated CpGs. Cell 58:499–507
Melcher M, Schmid M, Aagaard L, Selenko P, Laible G, Jenuwein T (2000) Structure-function analysis of SUV39H1 reveals a dominant role in heterochromatin organization, chromosome segregation, and mitotic progression. Mol Cell Biol 20:3728–3741
Miller OJ, Schnedl W, Allen J, Erlanger BF (1974) 5-Methylcytosine localised in mammalian constitutive heterochromatin. Nature 251:636–637
Miniou P, Jeanpierre M, Blanquet V, Sibella V, Bonneau D, Herbelin C, Fischer A, Niveleau A, Viegas-Pequignot E (1994) Abnormal methylation pattern in constitutive and facultative (X inactive chromosome) heterochromatin of ICF patients. Hum Mol Genet 3:2093–2102
Miniou P, Jeanpierre M, Bourc'his D, Coutinho Barbosa AC, Blanquet V, Viegas-Pequignot E (1997) Alpha-satellite DNA methylation in normal individuals and in ICF patients: heterogeneous methylation of constitutive heterochromatin in adult and fetal tissues. Hum Genet 99:738–745
Mirkin SM (2007) Expandable DNA repeats and human disease. Nature 447:932–940
Mitchell AR, Jeppesen P, Nicol L, Morrison H, Kipling D (1996) Epigenetic control of mammalian centromere protein binding: does DNA methylation have a role? J Cell Sci 109(Pt 9):2199–2206
Monier K, Mouradian S, Sullivan KF (2007) DNA methylation promotes Aurora-B-driven phosphorylation of histone H3 in chromosomal subdomains. J Cell Sci 120:101–114
Mortusewicz O, Schermelleh L, Walter J, Cardoso MC, Leonhardt H (2005) Recruitment of DNA methyltransferase I to DNA repair sites. Proc Natl Acad Sci USA 102:8905–8909
Nakano M, Cardinale S, Noskov VN, Gassmann R, Vagnarelli P, Kandels-Lewis S, Larionov V, Earnshaw WC, Masumoto H (2008) Inactivation of a human kinetochore by specific targeting of chromatin modifiers. Dev Cell 14:507–522
Nan X, Campoy FJ, Bird A (1997) MeCP2 is a transcriptional repressor with abundant binding sites in genomic chromatin. Cell 88:471–481
Nichol Edamura K, Leonard MR, Pearson CE (2005) Role of replication and CpG methylation in fragile X syndrome CGG deletions in primate cells. Am J Hum Genet 76:302–311
Nichol K, Pearson CE (2002) CpG methylation modifies the genetic stability of cloned repeat sequences. Genome Res 12:1246–1256
Nichols AF, Sancar A (1992) Purification of PCNA as a nucleotide excision repair protein. Nucleic Acids Res 20:2441–2446
Nosho K, Shima K, Irahara N, Kure S, Firestein R, Baba Y, Toyoda S, Chen L, Hazra A, Giovannucci EL et al (2009) SIRT1 histone deacetylase expression is associated with microsatellite instability and CpG island methylator phenotype in colorectal cancer. Mod Pathol 22:922–932
Nowak SJ, Corces VG (2004) Phosphorylation of histone H3: a balancing act between chromosome condensation and transcriptional activation. Trends Genet 20:214–220
Oda H, Okamoto I, Murphy N, Chu J, Price SM, Shen MM, Torres-Padilla ME, Heard E, Reinberg D (2009) Monomethylation of histone H4-lysine 20 is involved in chromosome structure and stability and is essential for mouse development. Mol Cell Biol 29:2278–2295
Okada T, Ohzeki J, Nakano M, Yoda K, Brinkley WR, Larionov V, Masumoto H (2007) CENP-B controls centromere formation depending on the chromatin context. Cell 131:1287–1300
Okano M, Bell DW, Haber DA, Li E (1999) DNA methyltransferases Dnmt3a and Dnmt3b are essential for de novo methylation and mammalian development. Cell 99:247–257
Ono T, Fang Y, Spector DL, Hirano T (2004) Spatial and temporal regulation of Condensins I and II in mitotic chromosome assembly in human cells. Mol Biol Cell 15:3296–3308
Ooi SK, Qiu C, Bernstein E, Li K, Jia D, Yang Z, Erdjument-Bromage H, Tempst P, Lin SP, Allis CD et al (2007) DNMT3L connects unmethylated lysine 4 of histone H3 to de novo methylation of DNA. Nature 448:714–717
Ooi SK, O'Donnell AH, Bestor TH (2009) Mammalian cytosine methylation at a glance. J Cell Sci 122:2787–2791
Oswald J, Engemann S, Lane N, Mayer W, Olek A, Fundele R, Dean W, Reik W, Walter J (2000) Active demethylation of the paternal genome in the mouse zygote. Curr Biol 10:475–478
Otten AD, Tapscott SJ (1995) Triplet repeat expansion in myotonic dystrophy alters the adjacent chromatin structure. Proc Natl Acad Sci USA 92:5465–5469
Palii SS, Van Emburgh BO, Sankpal UT, Brown KD, Robertson KD (2008) DNA methylation inhibitor 5-Aza-2′-deoxycytidine induces reversible genome-wide DNA damage that is distinctly influenced by DNA methyltransferases 1 and 3B. Mol Cell Biol 28:752–771
Palmer DK, O'Day K, Wener MH, Andrews BS, Margolis RL (1987) A 17-kD centromere protein (CENP-A) copurifies with nucleosome core particles and with histones. J Cell Biol 104:805–815
Palmer DK, O'Day K, Trong HL, Charbonneau H, Margolis RL (1991) Purification of the centromere-specific protein CENP-A and demonstration that it is a distinctive histone. Proc Natl Acad Sci USA 88:3734–3738
Pearson CE (2003) Slipping while sleeping? Trinucleotide repeat expansions in germ cells. Trends Mol Med 9:490–495
Pearson CE, Nichol Edamura K, Cleary JD (2005) Repeat instability: mechanisms of dynamic mutations. Nat Rev Genet 6:729–742
Perez-Castro AV, Shamanski FL, Meneses JJ, Lovato TL, Vogel KG, Moyzis RK, Pedersen R (1998) Centromeric protein B null mice are viable with no apparent abnormalities. Dev Biol 201:135–143
Peters AH, O'Carroll D, Scherthan H, Mechtler K, Sauer S, Schofer C, Weipoltshammer K, Pagani M, Lachner M, Kohlmaier A et al (2001) Loss of the Suv39h histone methyltransferases impairs mammalian heterochromatin and genome stability. Cell 107:323–337
Peters AH, Kubicek S, Mechtler K, O'Sullivan RJ, Derijck AA, Perez-Burgos L, Kohlmaier A, Opravil S, Tachibana M, Shinkai Y et al (2003) Partitioning and plasticity of repressive histone methylation states in mammalian chromatin. Mol Cell 12:1577–1589
Polioudaki H, Markaki Y, Kourmouli N, Dialynas G, Theodoropoulos PA, Singh PB, Georgatos SD (2004) Mitotic phosphorylation of histone H3 at threonine 3. FEBS Lett 560:39–44
Politi V, Perini G, Trazzi S, Pliss A, Raska I, Earnshaw WC, Della Valle G (2002) CENP-C binds the alpha-satellite DNA in vivo at specific centromere domains. J Cell Sci 115:2317–2327
Reik W, Dean W, Walter J (2001) Epigenetic reprogramming in mammalian development. Science 293:1089–1093
Rice JC, Briggs SD, Ueberheide B, Barber CM, Shabanowitz J, Hunt DF, Shinkai Y, Allis CD (2003) Histone methyltransferases direct different degrees of methylation to define distinct chromatin domains. Mol Cell 12:1591–1598
Rizwana R, Hahn PJ (1999) CpG methylation reduces genomic instability. J Cell Sci 112(Pt 24):4513–4519
Robertson KD, Wolffe AP (2000) DNA methylation in health and disease. Nat Rev Genet 1:11–19
Robertson KD, Ait-Si-Ali S, Yokochi T, Wade PA, Jones PL, Wolffe AP (2000) DNMT1 forms a complex with Rb, E2F1 and HDAC1 and represses transcription from E2F-responsive promoters. Nat Genet 25:338–342
Rogakou EP, Pilch DR, Orr AH, Ivanova VS, Bonner WM (1998) DNA double-stranded breaks induce histone H2AX phosphorylation on serine 139. J Biol Chem 273:5858–5868
Rogakou EP, Nieves-Neira W, Boon C, Pommier Y, Bonner WM (2000) Initiation of DNA fragmentation during apoptosis induces phosphorylation of H2AX histone at serine 139. J Biol Chem 275:9390–9395
Rountree MR, Bachman KE, Baylin SB (2000) DNMT1 binds HDAC2 and a new co-repressor, DMAP1, to form a complex at replication foci. Nat Genet 25:269–277
Ruzov A, Shorning B, Mortusewicz O, Dunican DS, Leonhardt H, Meehan RR (2009) MBD4 and MLH1 are required for apoptotic induction in xDNMT1-depleted embryos. Development 136:2277–2286
Santos-Rosa H, Schneider R, Bannister AJ, Sherriff J, Bernstein BE, Emre NC, Schreiber SL, Mellor J, Kouzarides T (2002) Active genes are tri-methylated at K4 of histone H3. Nature 419:407–411
Schermelleh L, Haemmer A, Spada F, Rosing N, Meilinger D, Rothbauer U, Cardoso MC, Leonhardt H (2007) Dynamics of Dnmt1 interaction with the replication machinery and its role in postreplicative maintenance of DNA methylation. Nucleic Acids Res 35:4301–4312
Schneider R, Bannister AJ, Myers FA, Thorne AW, Crane-Robinson C, Kouzarides T (2004) Histone H3 lysine 4 methylation patterns in higher eukaryotic genes. Nat Cell Biol 6:73–77
Schotta G, Lachner M, Sarma K, Ebert A, Sengupta R, Reuter G, Reinberg D, Jenuwein T (2004) A silencing pathway to induce H3-K9 and H4-K20 trimethylation at constitutive heterochromatin. Genes Dev 18:1251–1262
Shimada M, Niida H, Zineldeen DH, Tagami H, Tanaka M, Saito H, Nakanishi M (2008) Chk1 is a histone H3 threonine 11 kinase that regulates DNA damage-induced transcriptional repression. Cell 132:221–232
Slotkin RK, Martienssen R (2007) Transposable elements and the epigenetic regulation of the genome. Nat Rev Genet 8:272–285
Smits VA, Reaper PM, Jackson SP (2006) Rapid PIKK-dependent release of Chk1 from chromatin promotes the DNA-damage checkpoint response. Curr Biol 16:150–159
Souza PP, Volkel P, Trinel D, Vandamme J, Rosnoblet C, Heliot L, Angrand PO (2009) The histone methyltransferase SUV420H2 and heterochromatin proteins HP1 interact but show different dynamic behaviours. BMC Cell Biol 10:41
Spada F, Haemmer A, Kuch D, Rothbauer U, Schermelleh L, Kremmer E, Carell T, Langst G, Leonhardt H (2007) DNMT1 but not its interaction with the replication machinery is required for maintenance of DNA methylation in human cells. J Cell Biol 176:565–571
Suetake I, Shinozaki F, Miyagawa J, Takeshima H, Tajima S (2004) DNMT3L stimulates the DNA methylation activity of Dnmt3a and Dnmt3b through a direct interaction. J Biol Chem 279:27816–27823
Sullivan BA, Karpen GH (2004) Centromeric chromatin exhibits a histone modification pattern that is distinct from both euchromatin and heterochromatin. Nat Struct Mol Biol 11:1076–1083
Sullivan BA, Blower MD, Karpen GH (2001) Determining centromere identity: cyclical stories and forking paths. Nat Rev Genet 2:584–596
Tamburini BA, Tyler JK (2005) Localized histone acetylation and deacetylation triggered by the homologous recombination pathway of double-strand DNA repair. Mol Cell Biol 25:4903–4913
Tanaka Y, Kurumizaka H, Yokoyama S (2005) CpG methylation of the CENP-B box reduces human CENP-B binding. FEBS J 272:282–289
Teodoridis JM, Hardie C, Brown R (2008) CpG island methylator phenotype (CIMP) in cancer: causes and implications. Cancer Lett 268:177–186
Thornton CA, Wymer JP, Simmons Z, McClain C, Moxley RT 3rd (1997) Expansion of the myotonic dystrophy CTG repeat reduces expression of the flanking DMAHP gene. Nat Genet 16:407–409
Toyota M, Ahuja N, Ohe-Toyota M, Herman JG, Baylin SB, Issa JP (1999) CpG island methylator phenotype in colorectal cancer. Proc Natl Acad Sci USA 96:8681–8686
Trinh BN, Long TI, Nickel AE, Shibata D, Laird PW (2002) DNA methyltransferase deficiency modifies cancer susceptibility in mice lacking DNA mismatch repair. Mol Cell Biol 22:2906–2917
Umar A, Buermeyer AB, Simon JA, Thomas DC, Clark AB, Liskay RM, Kunkel TA (1996) Requirement for PCNA in DNA mismatch repair at a step preceding DNA resynthesis. Cell 87:65–73
van Attikum H, Gasser SM (2005) The histone code at DNA breaks: a guide to repair? Nat Rev Mol Cell Biol 6:757–765
Van Hooser A, Goodrich DW, Allis CD, Brinkley BR, Mancini MA (1998) Histone H3 phosphorylation is required for the initiation, but not maintenance, of mammalian chromosome condensation. J Cell Sci 111(Pt 23):3497–3506
Veigl ML, Kasturi L, Olechnowicz J, Ma AH, Lutterbaugh JD, Periyasamy S, Li GM, Drummond J, Modrich PL, Sedwick WD et al (1998) Biallelic inactivation of hMLH1 by epigenetic gene silencing, a novel mechanism causing human MSI cancers. Proc Natl Acad Sci USA 95:8698–8702
Vire E, Brenner C, Deplus R, Blanchon L, Fraga M, Didelot C, Morey L, Van Eynde A, Bernard D, Vanderwinden JM et al (2006) The polycomb group protein EZH2 directly controls DNA methylation. Nature 439:871–874
Wang KY, James Shen CK (2004) DNA methyltransferase Dnmt1 and mismatch repair. Oncogene 23:7898–7902
Wang H, Wang L, Erdjument-Bromage H, Vidal M, Tempst P, Jones RS, Zhang Y (2004) Role of histone H2A ubiquitination in polycomb silencing. Nature 431:873–878
Wei Y, Mizzen CA, Cook RG, Gorovsky MA, Allis CD (1998) Phosphorylation of histone H3 at serine 10 is correlated with chromosome condensation during mitosis and meiosis in Tetrahymena. Proc Natl Acad Sci USA 95:7480–7484
Wei Y, Yu L, Bowen J, Gorovsky MA, Allis CD (1999) Phosphorylation of histone H3 is required for proper chromosome condensation and segregation. Cell 97:99–109
Wohrle D, Salat U, Glaser D, Mucke J, Meisel-Stosiek M, Schindler D, Vogel W, Steinbach P (1998) Unusual mutations in high functioning fragile X males: apparent instability of expanded unmethylated CGG repeats. J Med Genet 35:103–111
Wong N, Lam WC, Lai PB, Pang E, Lau WY, Johnson PJ (2001) Hypomethylation of chromosome 1 heterochromatin DNA correlates with q-arm copy gain in human hepatocellular carcinoma. Am J Pathol 159:465–471
Wong NC, Wong LH, Quach JM, Canham P, Craig JM, Song JZ, Clark SJ, Choo KH (2006) Permissive transcriptional activity at the centromere through pockets of DNA hypomethylation. PLoS Genet 2:e17
Wossidlo M, Arand J, Sebastiano V, Lepikhov K, Boiani M, Reinhardt R, Scholer H, Walter J (2010) Dynamic link of DNA demethylation, DNA strand breaks and repair in mouse zygotes. EMBO J 29:1877–1888
Xin H, Yoon HG, Singh PB, Wong J, Qin J (2004) Components of a pathway maintaining histone modification and heterochromatin protein 1 binding at the pericentric heterochromatin in mammalian cells. J Biol Chem 279:9539–9546
Xu GL, Bestor TH, Bourc'his D, Hsieh CL, Tommerup N, Bugge M, Hulten M, Qu X, Russo JJ, Viegas-Pequignot E (1999) Chromosome instability and immunodeficiency syndrome caused by mutations in a DNA methyltransferase gene. Nature 402:187–191
Xu D, Bai J, Duan Q, Costa M, Dai W (2009) Covalent modifications of histones during mitosis and meiosis. Cell Cycle 8:3688–3694
Yoder JA, Walsh CP, Bestor TH (1997) Cytosine methylation and the ecology of intragenomic parasites. Trends Genet 13:335–340
Yokochi T, Robertson KD (2002) Preferential methylation of unmethylated DNA by mammalian de novo DNA methyltransferase Dnmt3a. J Biol Chem 277:11735–11745
Yoon SR, Dubeau L, de Young M, Wexler NS, Arnheim N (2003) Huntington disease expansion mutations in humans can occur before meiosis is completed. Proc Natl Acad Sci USA 100:8834–8838
Zeitlin SG, Barber CM, Allis CD, Sullivan KF (2001a) Differential regulation of CENP-A and histone H3 phosphorylation in G2/M. J Cell Sci 114:653–661
Zeitlin SG, Shelby RD, Sullivan KF (2001b) CENP-A is phosphorylated by Aurora B kinase and plays an unexpected role in completion of cytokinesis. J Cell Biol 155:1147–1157
Zeitlin SG, Baker NM, Chapados BR, Soutoglou E, Wang JY, Berns MW, Cleveland DW (2009) Double-strand DNA breaks recruit the centromeric histone CENP-A. Proc Natl Acad Sci USA 106:15762–15767
Zhang Y, Jurkowska R, Soeroes S, Rajavelu A, Dhayalan A, Bock I, Rathert P, Brandt O, Reinhardt R, Fischle W et al (2010) Chromatin methylation activity of Dnmt3a and Dnmt3a/3 L is guided by interaction of the ADD domain with the histone H3 tail. Nucleic Acids Res 38:4246–4253
Ziv Y, Bielopolski D, Galanty Y, Lukas C, Taya Y, Schultz DC, Lukas J, Bekker-Jensen S, Bartek J, Shiloh Y (2006) Chromatin relaxation in response to DNA double-strand breaks is modulated by a novel ATM- and KAP-1 dependent pathway. Nat Cell Biol 8:870–876
Work in the Robertson laboratory is supported by funds from the National Institutes of Health and the Georgia Cancer Coalition. KDR is a Georgia Cancer Coalition Distinguished Cancer Scholar.
Conflict of interest
The authors have no conflicts to declare.
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Putiri, E.L., Robertson, K.D. Epigenetic mechanisms and genome stability. Clin Epigenet 2, 299–314 (2011) doi:10.1007/s13148-010-0017-z
- DNA methylation
- Histone modifications
- Microsatellite instability (MSI)
- DNA repair