Methylation of cytosines within the CpG dinucleotide by DNA methyltransferases is involved in regulating transcription and chromatin structure, controlling the spread of parasitic elements, maintaining genome stability in the face of vast amounts of repetitive DNA, and X chromosome inactivation. Cellular DNA methylation is highly compartmentalized over the mammalian genome and this compartmentalization is essential for embryonic development. When the complicated mechanisms that control which DNA sequences become methylated go awry, a number of inherited genetic diseases and cancer may result. Much new information has recently come to light regarding how cellular DNA methylation patterns may be established during development and maintained in somatic cells. Emerging evidence indicates that various chromatin states such as histone modifications (acetylation and methylation) and nucleosome positioning (modulated by ATP-dependent chromatin remodeling machines) determine DNA methylation patterning. Additionally, various regulatory factors interacting with the DNA methyltransferases may direct them to specific DNA sequences, regulate their enzymatic activity, and allow their use as transcriptional repressors. Continued studies of the connections between DNA methylation and chromatin structure and the DNA methyltransferase-associated proteins, will likely reveal that many, if not all, epigenetic modifications of the genome are directly connected. Such studies should also yield new insights into treating diseases involving aberrant DNA methylation.
DNA methylation is a complex process whereby one of three DNA methyltransferases (DNMTs) catalyzes the addition of a methyl group from the universal methyl donor S-adenosyl-L-methionine, to the 5-carbon position of cytosine. This modification, occurring predominantly within the CpG dinucleotide, is the most prevalent epigenetic modification of DNA in mammalian genomes. There are currently three known, catalytically active DNMTs, DNMT1, 3a, and 3b and each one appears to play a distinct and critical role in the cell (Bestor, 2000). CpG methylation profoundly influences many processes including transcriptional regulation, genomic stability, chromatin structure modulation, X chromosome inactivation, and the silencing of parasitic DNA elements (Baylin et al., 2001; Jones and Laird, 1999; Robertson, 2001). These diverse processes nevertheless appear to share a common characteristic, that is, they all exert a stabilizing effect which promotes genomic integrity and ensures proper temporal and spatial gene expression during development.
Genomic DNA methylation patterns are not randomly distributed. Rather, discrete regions, including most repetitive and parasitic DNA, are hypermethylated, while other regions, such as CpG-rich regions often associated with the regulatory regions of genes (CpG islands), are hypomethylated (Yoder et al., 1997). Furthermore, DNA methylation patterns change dramatically during embryonic development. Genome wide demethylation after fertilization is followed by waves of de novo methylation upon embryo implantation. Not all sequences in the genome, however, are demethylated upon fertilization and not all sequences become de novo methylated after implantation. These exceptions further emphasize the regional specificity of genomic DNA methylation patterning (Reik et al., 2001). Evidence of the great importance of these methylation patterns can be gleaned by examining the effects of disrupting them in vivo. Engineered disruption of factors governing DNA methylation patterns in mice has revealed their vital role in embryonic development (Li et al., 1992; Okano et al., 1999). Naturally occurring mutations in genes involved in controlling DNA methylation patterns, including one of the DNA methyltransferases, result in ICF, Rett, ATRX and fragile X syndromes (Robertson and Wolffe, 2000). Disruption of normal DNA methylation patterns is one of the most common features of transformed cells and a number of studies have revealed that methylation changes are early events in the tumorigenesis process and contribute directly to transformation. In tumor cells the normal regulation of the DNA methylation machinery is severely disrupted, such that the regional specificity of methylation patterns begins to be reversed, resulting in de novo methylation of CpG islands and hypomethylation of repetitive DNA (Baylin et al., 2001; Jones and Laird, 1999; Robertson, 2001).
The role of DNA methylation in cancer has been reviewed extensively and will not be discussed at length here (Baylin et al., 2001; Jones and Laird, 1999; Robertson, 2001). Rather, I will focus on emerging data that may help to answer one of the most pressing and intriguing questions in the DNA methylation field, namely how cellular DNA methylation patterns are established during mammalian development and then properly maintained in somatic cells. Clues have been contributed by numerous studies in the last few years, which indicate that DNA methylation and chromatin structure, or the ‘tightness’ of packaging of the DNA in nucleosomes and the higher order structures they form, are physically and functionally linked (Bird, 2002). For example, all known catalytically active DNA methyltransferases interact with histone deacetylases and the use of inhibitors of each of these processes has revealed that they work together to repress transcription (Cameron et al., 1999). Studies in plants, mice, and humans, using naturally occurring or engineered mutations in chromatin remodeling machines, have indicated that chromatin structure itself may dictate cellular DNA methylation patterns (Bird, 2001). I will first discuss what is currently known about the components of the cellular DNA methylation machinery, namely the DNA methyltransferases. I will then summarize current knowledge of the interacting protein partners of the DNMTs and why the study of such factors may yield clues to the means by which methylation patterns are established. Lastly I will discuss the connections between DNA methylation and chromatin remodeling and describe a model of how chromatin states may dictate genome-wide DNA methylation patterning.
The mammalian DNA methylation machinery – properties of the DNA methyltransferases (DNMTs)
In this section, I will provide an overview of the enzymes responsible for establishing and maintaining cellular DNA methylation patterns in mammalian cells, namely the (cytosine-5) DNA methyltransferases (Figure 1). Particular emphasis will be placed on the domain structure of these proteins, their tissue-specific patterns of expression, subnuclear localization, alternatively spliced isoforms, and their catalytic activity.
The (cytosine-5) DNA methyltransferase 1, now properly referred to as DNMT1 (Figure 1), was the first enzyme to be isolated as a mammalian DNA methyltransferase, and more importantly, the only one that was identified via biochemical fractionation methodology (Bestor, 2000). All other genes described below were identified by database search. Following the isolation of murine DNMT1 (Bestor et al., 1988), its human homolog was identified in 1992 (Yen et al., 1992), which was mapped to chromosome 19p13.2. Generally, DNMTs are believed to be composed of two parts, a diversified amino terminal region and a relatively conserved carboxy terminal region. DNMT1 has the largest amino terminal region of all the mammalian DNMTs, which has roles in regulating the activity of the carboxy terminal catalytic domain, nuclear localization, zinc binding, and in mediating protein–protein interactions (Figures 1 and 4, Table 1; Bestor, 2000; Robertson, 2001). The carboxy terminal region comprises the catalytic domain that is a common feature of all cytosine DNA methyltransferases.
The biochemical and enzymatic properties of DNMT1 have been studied in considerable detail. DNMT1 has a significant preference for hemimethylated double-stranded DNA relative to unmethylated double-stranded DNA. This unique property is why DNMT1 is commonly referred to as the maintenance methyltransferase (Bacolla et al., 1999; Flynn et al., 1996; Glickman et al., 1997; Pradhan et al., 1997, 1999; Yokochi and Robertson, 2002). Analysis of the subnuclear localization of DNMT1 supports this assignment. During the G1 and G2 phases of the cell cycle, DNMT1 shows a diffuse nucleoplasmic distribution pattern, but associates with sites of DNA synthesis (replication foci) throughout S phase (Leonhardt et al., 1992; Liu et al., 1998). These results led researchers to a classic and still attractive hypothesis. DNMT1, as the primary maintenance methyltransferase, is required to maintain the epigenetic information encoded by genome-wide DNA methylation patterns due to the semiconservative nature of DNA replication. This process results in two hemimethylated daughter chromosomes that must be fully methylated in order for DNA methylation patterns to be properly maintained. In contrast to this simple model, the expression patterns of DNMT1 in certain cell types has been found to be somewhat paradoxical. If DNMT1 expression were strictly linked to DNA replication, then expression of DNMT1 would correlate with proliferative state of the cell. However, Northern blot analysis showed that DNMT1 is highly expressed not only in the placenta and lung, but also in low proliferative tissues such as the heart and brain (Robertson et al., 1999; Yen et al., 1992). This unexpected result may suggest an additional function for DNMT1 in addition to maintenance of methylation at replication foci. Emerging lines of evidence suggest that tissue-specific alternative splicing produces several forms of DNMT1, which result in enzymes designed to carry out distinct roles in DNA methylation metabolism.
An alternative splice variant of human DNMT1 in somatic tissues was reported by two groups and was named DNMT1b (Bonfils et al., 2000; Hsu et al., 1999). This isoform of DNMT1 contains an extra 48 base pairs between exons 4 and 5, generating a protein with an additional 16 amino acids derived from this region (Figure 2; Hsu et al., 1999). Dnmt1b mRNA is ubiquitously expressed in all cell lines tested and the protein possessed enzymatic properties comparable to DNMT1 in vitro (Bonfils et al., 2000). One group described the mRNA expression level of DNMT1b as 40–70% of the level of DNMT1 (Hsu et al., 1999), while another group reported that DNMT1b protein was present at only 2–5% of the level of DNMT1 protein (Bonfils et al., 2000). Thus, the relative abundance of DNMT1b and DNMT1 in somatic tissues remains unclear. The murine DNMT1 gene also undergoes alternative splicing and potentially generates an isoform (murine DNMT1b) similar to human DNMT1b, but would differ by two amino acids (Lin et al., 2000). A biological role for human and murine DNMT1b in somatic tissues and the functional differences between them in vivo remains unknown.
Alternative splicing of sex-specific 5′ exons produces at least two DNMT1 variants at the mRNA level, which have been termed DNMT1o and DNMT1p (Figure 2). A shorter isoform of DNMT1 (DNMT1o), which is specific to oocytes, yields a large amount of enzymatically active protein which accumulates in oocytes during the growth phase (Carlson et al., 1992; Mertineit et al., 1998). In fact, DNMT1o is the sole isoform of DNMT1 not only in the oocyte but also in the preimplantation embryo. It is noteworthy that DNMT1o exhibits an unusual trafficking movement. The majority of DNMT1o is localized in the cytoplasm, and transiently relocates to the nucleus during the eight-cell stage. This observation led to the suggestion that the cytoplasmic-nuclear translocation of DNMT1o at a particular stage of embryogenesis is essential for establishment of normal methylation patterns in imprinted regions (Cardoso and Leonhardt, 1999; Doherty et al., 2002).
A larger form of DNMT1 mRNA was detected exclusively in pachytene spermatocytes (Mertineit et al., 1998), and thus, the transcript was named DNMT1p (Figure 2; Doherty et al., 2002). It was originally concluded that DNMT1p was not translated, possibly due to the inhibitory effect of multiple short upstream open reading frames (Mertineit et al., 1998). However, the patterns of transcription and translation of this isoform remain controversial. In 2000, it was reported that an alternative DNMT1 transcript in skeletal muscle, specifically differentiated myotubes, was identical to DNMT1p (Aguirre-Arteta et al., 2000). It was also shown that this muscle-specific isoform was translatable in both transfected cells and in differentiated muscle cells. As with DNMT1o, DNMT1p may play an important role during oogenesis, gametogenesis, or myogenesis.
In 1998, attempts to identify novel putative de novo DNA methyltransferases in mammalian cells resulted in the identification of the DNMT2 gene (Figure 1), which shares homology with the pmt1 gene of fission yeast (Okano et al., 1998b; Yoder and Bestor, 1998). Although the human DNMT2 gene was mapped to chromosome 10p12-10p14 (Yoder and Bestor, 1998), the human genome project revealed that its actual location is 10p15.1. DNMT2 mRNA is ubiquitously expressed at very low levels (Okano et al., 1998b; Yoder and Bestor, 1998). The characteristic motifs found in all other active DNA methyltransferases, including the conserved proline-cysteine dipeptide at the active site, are well conserved in both the human and murine DNMT2 genes. Interestingly, DNMT2 may be the most highly conserved of all DNMTs across species since, unlike DNMT1 and DNMT3, DNMT2-like proteins have been found in yeast, plants, and the fruit fly Drosophila (Hung et al., 1999; Lyko et al., 2000; The Arabidopsis Genome Initiative, 2000; Wilkinson et al., 1995). At this time, however, no methyl-transfer activity of the DNMT2 gene product has been reported using in vitro assays (Dong et al., 2001; Okano et al., 1998b; Yoder and Bestor, 1998), although the DNMT2 protein does form stable DNA-protein complexes in vitro (Dong et al., 2001). Elucidation of the exact function of this protein in vivo will be of great importance. DNMT2 could, for example, be the catalytic subunit of a DNA methylase complex that is inactive when expressed without one or more of its associated factors. Alternatively it may possess a more complex DNA recognition sequence beyond the CpG dinucleotide.
Low level, stable de novo methylation activity could be observed in embryonic stem (ES) cells lacking DNMT1, supporting the notion that enzymes other than DNMT1 contribute to de novo methylation in vivo (Lei et al., 1996). DNMT3a and DNMT3b (Figure 1), two genes sharing significant homology to DNMT1, were identified by EST database searches in 1998 using the conserved methyltransferase motifs as bait (Okano et al., 1998a). The human DNMT3a gene was mapped to chromosome 2p23 (Robertson et al., 1999) and shows 96% amino acid identity to its murine counterpart (Xie et al., 1999). This is significantly higher than the identity between human and murine DNMT1 (78%), and suggests a critical function for this enzyme preserved throughout evolution. The carboxy terminal portion of DNMT3a includes the catalytic motifs highly conserved in all cytosine DNA methyltransferases. DNMT3a has been shown to be enzymatically active both in vitro and in vivo by several groups, although these reports differ somewhat in the exact substrate preference of DNMT3a (Aoki et al., 2001; Gowher and Jeltsch, 2001; Hsieh, 1999; Lyko et al., 1999; Okano et al., 1998a; Yokochi and Robertson, 2002).
Proteins of the DNMT3 family commonly have a cysteine-rich domain in the amino terminal region, which is referred to as the PHD (plant homeodomain) region or ATRX-like domain (Figure 1) because of its homology with the PHD region of the ATRX gene. ATRX is a member of the SNF2/SWI2 family of ATP-dependent chromatin remodeling enzymes. This similarity suggests that DNMT3 proteins may be associated with structural changes in chromatin via protein–protein interactions at the amino terminal region. DNMT3a transcripts were ubiquitously expressed in adult tissues, most tumor cell lines, early embryos, and embryonic stem (ES) cells (Okano et al., 1998a; Robertson et al., 1999; Xie et al., 1999). Northern blot analysis showed that DNMT3a activity is particularly high at embryonic day 10.5 (Okano et al., 1998a). Unlike DNMT3b, the level of DNMT3a mRNA was less affected when cells were arrested in the G0/G1 phase of the cell cycle, suggesting that DNMT3a expression is not regulated in a cell cycle-dependent manner like DNMT1 (Robertson et al., 2000b). In contrast to DNMT1, DNMT3a was found at discrete nuclear foci throughout the cell cycle that were not associated with DNA synthesis. During late S phase, however, when heterochromatic regions of the genome are known to be replicated, some of the DNMT3a-enriched foci appeared to overlap with replication foci (Bachman et al., 2001).
DNMT3b and its multiple isoforms
The human DNMT3b gene was mapped to chromosome 20q11.2 (Robertson et al., 1999; Xie et al., 1999) and is 85% identical to murine DNMT3b (Figure 1). The catalytic domain, located at the carboxy terminus, is well conserved between DNMT3a and DNMT3b (more than 80% identity), whereas their amino terminal regions are poorly conserved (less than 30%). This further underscores the notion that the variety in function of each of the DNMTs is most likely due to their diverse N-terminal regions, while the highly conserved C-terminal regions, which are common among all mammalian DNMTs, have similar functions. DNMT3b, like DNMT3a, was shown to be an active DNA methylstransferase in vivo and in vitro (Aoki et al., 2001; Hsieh, 1999; Okano et al., 1998a; Qiu et al., 2002).
Compared to DNMT3a, the expression levels of DNMT3b are very low in most tissues. The testis, however, expressed high levels of DNMT3b, suggesting a crucial function for DNMT3b in spermatogenesis (Okano et al., 1998a; Robertson et al., 1999; Xie et al., 1999). DNMT3b shows a diffuse distribution pattern throughout the nucleus in NIH3T3 cells, while targeting to pericentromeric heterochromatin was observed in undifferentiated ES cells (Bachman et al., 2001). As will be discussed later in this review in the context of ICF syndrome, DNMT3b appears to be specialized for the establishment and/or maintenance of DNA methylation of the minor satellite repeats (satellites 2 and 3 on human chromosomes 1, 9, and 16) and its co-localization with pericentromeric heterochromatin in consistent with this function.
In contrast to DNMT3a, there are several isoforms (five for human and eight for mouse) of DNMT3b that result from alternative splicing. Three major isoforms, namely DNMT3b1, 3b2, and 3b3, have been identified (Figure 3) (Okano et al., 1998a), and were shown to be expressed in a tissue-specific manner (Robertson et al., 1999). DNMT3b1 is the longest form and is usually regarded as the typical gene product of DNMT3b. All other splice variants encode smaller proteins. DNMT3b2 also demonstrated methyltransferase activity in vitro, however DNMT3b3, an even shorter isoform lacking 63 amino acids within the central region of the catalytic domain, did not (Aoki et al., 2001; Okano et al., 1998a; Qiu et al., 2002). Several other splice variants, DNMT3b4 and DNMT3b5 (Figure 3) were identified and are expressed predominantly in the testis (Robertson et al., 1999). Interestingly, these forms of DNMT3b lack several of the conserved catalytic motifs and possess additional novel sequences resulting from frameshifts after alternative splicing and are therefore unlikely to be catalytically active. These splice variants may possess the critical functions encoded by the N-terminal region of DNMT3b, including protein–protein interaction sites, but lack catalytic activity. They may therefore act to inhibit some function of DNMT3b1-2, like de novo methylation of certain sequences, by interacting with the same set of targeting proteins.
The human DNMT3L (DNMT3-like, Figure 1) gene located on human chromosome 21q22.3 was originally isolated by database analysis of the genome sequence in 2000 (Aapola et al., 2000). The identification of the murine DNMT3L gene (Aapola et al., 2001), which is 61% identical to its human counterpart, soon followed. Similarity between DNMT3L and DNMT3a and DNMT3b is restricted mainly to the cysteine-rich region encompassing the PHD/ATRX-like region in the amino terminus of DNMT3L. DNMT3L lacks the critical catalytic motifs commonly seen in all other DNA methyltransferases, including the ‘FGG’ sequence in conserved motif I, the ‘PC’ catalytic active site in motif IV, and the ‘ENV’ sequence in motif VI believed to be involved in cofactor binding. Thus, the gene product is almost certainly catalytically inactive, although this has yet to be tested directly. DNMT3L is highly expressed in testis and mouse embryos (Aapola et al., 2001), and is likely involved in gametogenesis for the process of establishing genomic imprints (Bourc'his et al., 2001).
Insights into functions of the DNA methylation machinery derived from knockout models
The recent history of DNA methylation research has been punctuated by several elegant studies using murine knockout models to elucidate the role of DNA methylation in general, and of the functions of the individual DNMTs in development, transcriptional regulation, chromatin structure maintenance and genomic imprinting. Comparison of the transgenic knockout mice with the corresponding knockout ES cells provides additional data on the role of each of the DNMTs in a pluripotent cell versus a more differentiated cell type. This section of the review will summarize the effects of inactivating each of the DNMTs at the individual cell and the whole organism levels.
Murine ES cells lines homozygous for a DNMT1 knockout were obtained by targeted disruption (Li et al., 1992). The mutant ES cells were viable and showed normal morphology. Only residual DNA methyltransferase activity was observed in lysates derived from these homozygous mutant cells. DNA from homozygous mutant cells was found to have a 5-methylcytosine content of roughly 30% of the DNA from wild type cells, in which about 60% of all CCGG sites were methylated (that is, a 70% decrease in 5-methylcytosine content in the CCGG sequence) (Li et al., 1992). Interestingly, targeted disruption of the DNMT1 gene in a human colorectal carcinoma cell line yielded somewhat different results (Rhee et al., 2000). DNA from the DNMT1 deficient HCT116 cell line had a 5-methylcytosine content of about 80% of the DNA from wild type cells, in which approximately 4% of all cytosines were methylated (that is, a 20% decrease in total genomic 5-methylcytosine content). A direct comparison of the results between these two experimental systems is somewhat difficult due to the different methods used for measuring the amount of DNA methylation in the original wild type cells. Regardless of whether this discrepancy arises from different experimental methods or from the radically different cell types used in each study, these results suggest that methyltransferases other than DNMT1 contribute significantly to the homeostasis of DNA methylation patterns. Transgenic mice heterozygous for the DNMT1 knockout were indistinguishable from wild type littermates. Homozygous mutation of DNMT1 was embryonic lethal and mutant embryos failed to develop beyond mid-gestation, strongly suggesting that DNA methylation is essential for mammalian development. DNMT1 mutants demonstrated significant hypomethylation of several genes, which affected a variety of epigenetic events including genomic imprinting (Li et al., 1993), X chromosome inactivation (Beard et al., 1995), and the suppression of transcription from parasitic elements (Walsh et al., 1998).
More recently, conditional knockouts of DNMT1 have been generated to examine its role in the development and maintenance of specific tissues, particularly the central nervous system. In one study, the cre/lox system was used to delete the DNMT1 locus from primary fibroblasts in culture (Jackson-Grusby et al., 2001). The resulting DNMT1-null cells displayed extensive genomic hypomethylation and uniform cell death by apoptosis within 6 days of DNMT1-deletion. The connection between reduced DNA methylation and apoptosis was further underscored by showing that interfering with p53 function, a key mediator of apoptosis, prolonged the life span of the DNMT1-null cells and reduced the level of apoptosis. Microarray expression studies revealed that 4–10% of all genes were upregulated and 1–2% were down-regulated when DNMT1 was deleted and that these genes were involved in a multitude of cellular functions (Jackson-Grusby et al., 2001). Conditional deletion of DNMT1 in neural precursor cells early in development resulted in genome-wide demethylation and embryonic lethality, although brain structure appeared normal. Mice delivered by caesarian section exhibited aberrant breathing and their lungs failed to inflate, indicating potential defects in respiratory rhythmogenesis or neurotransmission. Deletion of DNMT1 at this early developmental stage in a small fraction of neural precursor cells resulted in a marked selection against the demethylated cells such that the mice developed normally and were devoid of demethylated, DNMT1-deficient cells (Fan et al., 2001). Deletion of DNMT1 from post-mitotic neurons later in development was compatible with both normal development and normal DNA methylation patterns. Thus, these studies strongly suggest that DNA methylation is critical for normal mammalian development during embryogenesis and may play a particularly important role in brain development and function as has been suggested previously (Fan et al., 2001; Robertson, 2001; Tucker, 2001).
Since DNMT1o demonstrated a unique nuclear-cytoplasmic trafficking movement during oogenesis and preimplantation development, a functional role for DNMTo in genomic imprinting in the oocyte was proposed (Carlson et al., 1992; Mertineit et al., 1998). Although males homozygous for a deletion of the maternal-specific exon were normal and fertile, homozygous females were infertile. That is, most heterozygous fetuses of homozygous females died during the last third of gestation, indicating that deletion of DNMT1o causes a pure-maternal phenotype during oogenesis (Howell et al., 2001). DNA methylation at certain imprinted loci, but not the whole genome, was lost in the heterozygous embryos. Therefore it appears that DNMT1o is required to maintain methylation patterns at specific imprinted loci during the fourth embryonic S phase.
Murine ES cells with a homozygous disruption of the DNMT2 gene appeared to be normal, suggesting that DNMT2 is not essential for cell proliferation, growth, and differentiation. No significant change in the methylation status of both genomic DNA and newly integrated provirus DNA was observed in the DNMT2−/− ES cells. These results indicated that DNMT2 is required for neither maintenance nor de novo methylation in vivo (Okano et al., 1998b).
ES cell lines homozygous for a DNMT3a knockout (DNMT3a−/−) retained their undifferentiated morphology and showed normal de novo methylation activity on newly integrated proviral DNA following retroviral infection (Okano et al., 1999). DNMT3a heterozygous mice were normal and fertile. Although DNMT3a−/− homozygous mice appeared to be normal at birth, undergrowth of such mutant mice by 18 days was obvious and all of the animals died by 4 weeks of age.
A homozygous knockout of the DNMT3b gene in ES cells yielded a phenotype similar to that of DNMT3a disruption. DNMT3b−/− ES cell lines and embryos exhibited comparable degrees of demethylation to those of DNMT3a−/− homozygous ES cells and embryos and a comparable ability to de novo methylate retroviral sequences (Okano et al., 1999). However, an analysis of early embryos revealed clear evidence that DNMT3a and DNMT3b have independent functions. Unlike DNMT3a−/− mice, no viable DNMT3b−/− mice were recovered at birth. Interestingly, the minor satellite repeats in the pericentromeric region were significantly hypomethylated in DNMT3b−/− but not in DNMT3a−/− cells, indicating that the minor satellite repeats are specific targets of DNMT3b. As will be discussed later, naturally occurring mutations in the human DNMT3b gene give rise to ICF syndrome, which is also characterized by hypomethylation of pericentromeric repeats (Hansen et al., 1999; Okano et al., 1999; Xu et al., 1999).
DNMT3a−/−, DNMT3b−/− double mutant ES cells were also generated and were viable. However, de novo methylation of newly integrated retroviral sequences was no longer observed, suggesting that the gene products of DNMT3a and DNMT3b have overlapping functions in ES cells with regard to de novo methylation of parasitic elements. Embryonic defects in the double mutant mice were more severe than those of DNMT3a−/− single mutant mice. Global methylation levels in ES cell lines and E9.5 day embryos were dramatically reduced in DNMT3a−/−, DNMT3b−/− knockout mutants compared to the single mutants (Okano et al., 1999), supporting the notion that DNMT3a and DNMT3b have at least partially overlapping functions in the establishment of cellular DNA methylation patterns during development.
Targeted disruption of DNMT3L gene was recently reported (Bourc'his et al., 2001). Homozygous animals of both sexes were viable but sterile. Bisulfite genomic sequencing revealed that a defect in DNMT3L resulted in the disruption of maternal methylation imprints in homozygous oocytes, while genome-wide methylation patterns appeared to be normal, indicating that the DNMT3L protein contributes to establishment of genomic imprints during oogenesis. As was discussed in the previous section, DNMT3L is almost certainly not a functional DNA methyltransferase therefore the observation that its disruption results in any loss of DNA methylation indicates that DNMT3L may target other functional DNA methyltransferases, such as DNMT3a and DNMT3b, to specific loci in the genome via direct or indirect protein–protein interaction.
Chromatin-associated proteins that interact with DNMTs
In vitro studies of the DNMTs have shown that they exhibit little sequence preference beyond the CpG dinucleotide, while the knockout studies emphasize the non-random nature of DNA methylation and the distinctive roles for the individual DNMTs. A prime mediator of these distinctive roles is most likely the complement of proteins that interact with the DNMTs at specific stages of development and differentiation, and within the environment of chromatin. A number of proteins have now been identified which interact with one or more of the DNMTs (Figure 4, Table 1; Robertson, 2001). This section of the review will focus specifically on chromatin-related proteins known to associate with the DNMTs and discuss the functional consequences of these interactions.
The retinoblastoma protein, Rb
The retinoblastoma protein, Rb, is a protein with intimate links to transcriptional regulation in the context of chromatin. Robertson et al. (2000a) initially identified an interaction between Rb, E2F1, and DNMT1 via biochemical fractionation, and this result was recently confirmed by others (Figure 4; Pradhan and Kim, 2002). The amino terminal region of DNMT1 could interact with the A/B pocket region of Rb (Robertson et al., 2000a), and also appears to be capable of interacting with the B/C pocket (Pradhan and Kim, 2002). Rb and DNMT1 cooperate to repress transcription from E2F-responsive promoters in vivo although this repression did not depend on the catalytic activity of DNMT1 (Robertson et al., 2000a). A recent report demonstrated that Rb binding to DNMT1 inhibited the ability of DNMT1 to bind to DNA in vitro and exerted a strong negative effect on DNMT1 catalytic activity. Overexpression of Rb in cells resulted in a significant reduction in total genomic 5-methylcytosine levels (Pradhan and Kim, 2002). Rb, like DNMT1, also co-localizes with DNA replication foci, specifically early S phase perinucleolar replication foci (Kennedy et al., 2000). The role of Rb at these sites remains unclear.
Hypophosphorylated Rb interacts with E2F family members and inhibits their transactivation function. As cells prepare to divide, Rb is phosphorylated and dissociates from E2F, allowing it to recruit co-activators and stimulate transcription of genes involved in cell cycle progression (Dyson, 1998). Interestingly, Rb has been shown to repress transcription at E2F-responsive promoters by recruitment of both HDACs and histone methyltransferases (HMTs) and subsequent binding of the methylated lysine binding proteins of the heterochromatin protein (HP) 1 family (Brehm et al., 1998; Luo et al., 1998; Nielsen et al., 2001). As will be discussed in the next section, DNMT1 can also interact directly with HDAC1 and HDAC2 (Robertson et al., 2000a; Rountree et al., 2000) and methylated cytosine itself serves as a recognition site for another class of repressors of the methyl-CpG binding protein (MBD) family. The MBDs have also been shown to recruit HDACs to methylated DNA and repress transcription (Bird and Wolffe, 1999).
What are the functional consequences of the interaction between Rb and DNMT1? We have previously proposed a model wherein DNMT1 binding to Rb at E2F-containing promoters may be a mechanism to sequester DNMT1 from the genome and prevent promiscuous DNA methylation events (Robertson, 2001). This is supported by recent findings that the catalytic activity of DNMT1 is inhibited by binding to Rb (Pradhan and Kim, 2002). This study also speculated that DNMT1 bound to Rb would not be capable of binding to PCNA, another DNMT1-associated protein thought to recruit DNMT1 to replication foci. Therefore this model may require revision to take into account these new observations. The exact nature of the complexes formed between DNMT1, Rb, and PCNA in silent chromatin versus replication foci will require extensive additional study however we propose a model (Figure 5) for how the nature of these interactions may change during the cell cycle and how they may effect the catalytic activity of DNMT1. It is likely that temporal changes in the complement of DNMT1-associated proteins, particularly Rb and HDAC2, which associate with early and late replication foci, respectively, will be critical modulators of DNMT1 activity at this site.
HDAC1 and HDAC2
Several studies have now shown that DNMT1, DNMT3a, and DNMT3b associate with HDAC1 and HDAC2 in vitro and in vivo (Figure 4 and Table 1; Bachman et al., 2001; Fuks et al., 2000, 2001; Robertson et al., 2000a). Histone deacetylases can remove acetyl groups from the amino terminal core histone tails, which are critical modulators of chromatin structure, leading to the assembly of tightly packed chromatin and rendering the sequence inaccessible to the transcription machinery (Jenuwein and Allis, 2001). DNMT1-mediated transcriptional repression has been shown to be comprised of both HDAC-dependent and HDAC-independent components. The HDAC-dependency can be demonstrated using the HDAC inhibitor trichostatin A (TSA), which relieves a substantial amount of the DNMT1-mediated repression (Fuks et al., 2000; Robertson et al., 2000a). DNMT1 was shown to bind HDAC1 via a transcriptional repression region other than the HRX-homology domain (Figure 4; Fuks et al., 2000). A direct interaction between the amino terminal half of DNMT1 and HDAC2 has also been demonstrated (Figure 4; Rountree et al., 2000). At present, there is no evidence to believe that there are major functional differences between HDAC1 and HDAC2 since they are highly similar proteins (85% identical at the amino acid level in humans). Thus, the interaction of DNMT1 with HDAC1 or HDAC2 likely has the same functional consequence. The TSA-insensitive component of DNMT1 repression may be mediated by its interaction with DNMT1-associated protein (DMAP) 1 (Rountree et al., 2000). DMAP1 binds to the extreme amino terminus of DNMT1, a region missing in certain germ-cell specific DNMT1 splice variants (Figures 2 and 4). DMAP1 interacts with another potent transcriptional repressor TSG101, although the existence of a DNMT1, DMAP1, and TSG101 ternary complex has not been reported. DMAP1 co-localizes with DNMT1 at replication foci throughout S phase (Figure 5) and may affect catalytic activity or targeting of DNMT1.
The DNMT3s also interact with HDAC1 through their ATRX-homology/PHD regions (Bachman et al., 2001; Fuks et al., 2001), which are not present in DNMT1 (Figure 4 and Table 1). DNMT3a and DNMT3b also exhibited a TSA-insensitive transcriptional repression component, however the protein–protein interactions mediating this repression are unknown (Bachman et al., 2001). Therefore, it is likely that the repression capability of DNMT1 and DNMT3s are the result of distinct protein–protein interactions, although histone deacetylase activity appears to be involved in both.
What is the functional consequence of the HDAC-DNMT interactions? This extremely important question remains unanswered. The potential importance is further underscored by the now universal nature of the association. All known catalytically active DNMTs have been shown to interact with HDACs (Table 1). One scenario that has been proposed previously relates to the temporal association of DNMT1 and HDAC2 at replication foci (Figure 5). Heterochromatic, hypermethylated sequences enriched in hypoacetylated histones are replicated in late S phase. The recruitment of HDAC2 to replication foci by DNMT1 may help to coordinate remethylation of the newly synthesized DNA and deacetylation of newly deposited histones in late S phase (Rountree et al., 2000). It is possible that DNMT recruitment to, and methylation of, a specific genomic region then brings in HDACs to deacetylate core histones in the newly methylated region and allow for chromatin compaction and transcriptional silencing. This implies that DNA methylation is the primary event in transcriptional silencing and chromatin modifications follow. Evidence gathered from experimental systems in which gene silencing occurs in a regulated manner, such as X chromosome inactivation in females and host genome defense-mediated silencing and de novo methylation of retroviral DNA, suggests that other mechanisms may also exist. In X chromosome inactivation, transcriptional silencing and histone modifications (both deacetylation and methylation) occur well before de novo DNA methylation (Csankovszki et al., 2001; Heard et al., 2001). Furthermore, transcriptional silencing of newly introduced retroviral sequences occurs before de novo methylation and can occur in the complete absence of the de novo DNA methyltransferases (Pannell et al., 2000), which have been shown to be responsible for methylation of this class of sequence (Okano et al., 1999). Therefore an alternative scenario is one in which HDACs are recruited to a region destined to undergo long-term transcriptional silencing via interaction with other proteins. Once the region is deacetylated and possibly with the help of other chromatin remodeling factors, the DNMTs gain access to the DNA or are attracted to a particular chromatin structural feature, the region is methylated, and stable, heritable, long-term gene silencing and chromatin compaction is achieved (Figures 5 and 6). Alternatively, both models may operate and depend on the transcriptional and replicative state of the cell.
As has been mentioned previously, there is a great need to better understand how cellular DNA methylation patterns are targeted to, or restricted from, certain regions of the genome. A fascinating recent study, directly relevant to the targeting issue, showed that DNMT1 and DNMT3a can interact with the leukemia-promoting PML–RAR fusion protein (Figure 4 and Table 1; di Croce et al., 2002). Acute promyelocytic leukemia (APL) is caused by a reciprocal translocation of the retinoic acid receptor α (RARα) gene on chromosome 17 to one of several other chromosomes. To generate the PML–RAR fusion, the most common in APL, the RARα gene is fused to the promyelocytic leukemia gene (PML) on chromosome 15. PML is a critical component of discrete nuclear structures referred to as PML-oncogenic domains (PODs, ND10 and nuclear bodies). PODs, which are composed of a number of proteins, including Sp100, Sp140, SUMO-1 and Daxx, may be involved in transcriptional activation, DNA replication, and apoptosis (Li et al., 2000; Muller et al., 2001; Zhong et al., 2000a,b). The PML–RAR fusion protein, which retains the DNA and ligand binding domains of RARα, disrupts the PODs, but they can be restored by treatment of cells with retinoic acid (RA). RA treatment and POD reorganization correlates with differentiation of the APL cells, indicating that PODs may have a critical role in differentiation of promyelocytes (di Croce et al., 2002; Zhong et al., 2000a).
di Croce et al. (2002) showed that PML–RAR could repress a model RA target gene, RARβ2, and that the repression coincided with de novo methylation of the 5′ end of the endogenous RARβ2 promoter CpG island. Co-immunoprecipitation studies revealed an interaction between PML–RAR and DNMT1 and DNMT3a, and immunofluorescence studies showed co-localization of PML–RAR and DNMT1 and Dnmt3a when over expressed. It appeared that DNMT3a could interact with regions on both the PML and RAR portions of the fusion protein. Treatments of cells with the DNA methylation inhibitor 5-aza-2′-deoxycytidine (5-aza-dC) or the HDAC inhibitor TSA, revealed that the PML–RAR-mediated transcriptional repression and differentiation blockage was due to the temporally distinct recruitment of both histone deacetylases (at early time points) and DNA methyltransferases (at later time points) (di Croce et al., 2002). The functional significance of the interaction between DNMT1 and DNMT3a and PML in normal cells is unclear, although it is possible that the DNMTs may carry out some critical function in the PODs. Interestingly, another PML-associated POD protein and transcriptional repressor, Daxx, has been shown to associate with DNMT1 and it remains possible that Daxx bridges the interaction between PML–RAR and DNMT1 (Li et al., 2000; Michaelson et al., 1999; Zhong et al., 2000b). Perhaps most significantly are the implications of this work in cancer since there have been no prior studies showing how regional hypermethylation events, common to so many tumor cells, might occur.
DNMT3a and DNMT3b were recently shown to interact directly with a protein called RP58 via the ATRX-like domain (Figure 4 and Table 1; Fuks et al., 2001). This is also the region of DNMT3a and DNMT3b that interacts with HDAC1 (and most likely HDAC2). RP58 is a sequence-specific zinc finger DNA binding protein and transcriptional repressor associated with heterochromatin (Aoki et al., 1998; Meng et al., 2000). It contains a POZ domain and several Krüppel-type zinc fingers commonly seen in other transcriptional repressors (Ryan et al., 1999). The repression activity of RP58 was enhanced by co-expression of DNMT3a and the cooperative effect did not require the catalytic activity of DNMT3a (Fuks et al., 2001). This suggests that DNMT3a acts as a structural component, rather than an active enzyme, in this repression pathway. Nuclear localization studies using a DNMT3a fragment lacking the catalytic domain also support this notion. The isolated amino terminal regulatory domain of DNMT3a co-localized with heterochromatin-associated proteins like HP1α as well as methyl-CpG binding proteins like MeCP2 (Bachman et al., 2001). Thus, the co-localization of DNMT3a with these types of proteins suggests that it may be an important component of densely methylated, pericentromeric heterochromatin. Although the ability of DNMT3a and RP58 to cooperate in transcriptional repression was not dependent on the catalytic activity of DNMT3a, it still remains possible that one function of RP58 could be to target DNMT3a to specific DNA sequences that are destined to become de novo methylated. Post-translational modifications, other protein cofactors expressed in a tissue-specific manner (RP58 is highly expressed in the brain for example (Meng et al., 2000)), missing from the cell culture system used to first characterize this interaction, could influence the function of the RP58 – DNMT3a complex.
MBD2 and MBD3
A physical interaction between DNMT1 and methyl-CpG binding proteins was also reported (Figure 4 and Table 1). DNMT1 was co-immunoprecipitated with MBD2 and MBD3 as one potential complex (Tatematsu et al., 2000). MBD2 and MBD3 have recently been reported to be components of the large macromolecular MeCP1 repressor complex, which is capable of preferentially binding, remodeling, and deacetylating methylated DNA-containing nucleosomes in vitro (Feng and Zhang, 2001). MBD2 and MBD3 co-localized with DNMT1 at replication foci in late S phase. Furthermore, the MBD2–MBD3 complex demonstrated binding affinity for both hemimethylated and fully methylated DNA and repressed transcription in a TSA-sensitive manner (Tatematsu et al., 2000). Thus, these results suggest the possibility of an ultimate ‘all-in-one’ type transcriptional repression complex, which contains a DNA modification module (DNMT), a methylcytosine reocognition subunit (methyl-CpG binding protein), and a histone modifying subunit (HDAC). This complex could have roles in directing DNMT1 to hemimethylated sequences following DNA replication, silencing of genes during S phase, or deacetylation of newly deposited histones in a manner akin to the previously described DNMT-DMAP1-HDAC2 interactions.
ICF syndrome – a disease caused by aberrant DNA methylation and chromatin structure
The identification of numerous interactions between DNA methyltransferases and chromatin associated proteins like histone deacetylases, Rb, and RP58 provides a clear link between DNA methyltransferases and transcriptional regulation and chromatin structure modulation. Is there additional in vivo evidence, particularly in human cells, that these processes are connected? While knockout studies in mouse models have been very revealing, the severity of the phenotype, namely embryonic lethality for DNMT1 and DNMT3b, limits the information that can be gained to embryonic development. Are there models or diseases involving less severe mutations in the DNA methylation machinery that could provide clues? The recent finding of a human genetic disease caused by mutations in a DNA methyltransferase gene, the only such disease known, has provided a wealth of information about the more subtle roles a particular DNMT may play in determining genomic DNA methylation patterns.
ICF syndrome (immunodeficiency, centromere instability, facial anomalies) is a very rare recessive disorder caused by mutations in the DNMT3b gene (Hansen et al., 1999; Okano et al., 1999; Xu et al., 1999). Most ICF patients are compound heterozygotes for their DNMT3b mutations and, with one exception, all of these mutations occur within the carboxy terminal catalytic domain of DNMT3b and likely fully or partially impair catalytic activity (Robertson, 2001; Xu et al., 1999). Phenotypically, ICF syndrome is characterized by a profound immunodeficiency with an absence or severe reduction in at least two immunoglobulin isotypes, variable impairment in cellular immunity, unusual facial features, neurologic and intestinal dysfunction, and delayed developmental milestones (Franceschini et al., 1995; Smeets et al., 1994).
At the cytogenetic level, primary and cultured cells from ICF patients exhibit marked elongation of juxtacentromeric heterochromatin. This elongation occurs most consistently on chromosomes 1 and 16, and to a lesser extent chromosome 9. Abnormalities which have been observed include multiradial chromosomes involving multiple arms (3–12) of one or more of the decondensed chromosomes, whole-arm chromosome deletions or duplications, translocations, centromeric breakage, and in rare cases telomeric associations (Franceschini et al., 1995; Smeets et al., 1994; Tuck-Muller et al., 2000). These observations strongly suggest that defective forms of DNMT3b lead to chromosome instability and large-scale changes in chromatin structure.
Molecular aspects of ICF syndrome
One of the most consistent features of ICF syndrome is juxtacentromeric repeat sequence hypomethylation on chromosomes 1, 9, and 16 (Jeanpierre et al., 1993). Interestingly, these chromosomes contain the largest blocks of classical satellite long tandem repeat arrays (satellite 2 for chromosomes 1 and 16, and satellite 3 for chromosome 9) adjacent to their centromeres (Jeanpierre et al., 1993; Tagarro et al., 1994). These regions are normally hypermethylated in somatic cells and such methylation is likely essential for proper centromere structure and stability (Jeanpierre et al., 1993; Tuck-Muller et al., 2000). A recent study, using bisulfite genomic sequencing, revealed that satellite 2 repeat methylation was reduced from roughly 70% in normal lymphoblasts, to 20% in ICF cells (Hassan et al., 2001). Although there is a drastic loss of methylation from satellite sequences in ICF patients, the overall reduction in cellular 5-methylcytosine levels is relatively small (about 7% in primary ICF brain tissue) further underscoring the idea that mutations in DNMT3b lead to highly selective losses of methylation from the genome (Kondo et al., 2000; Tuck-Muller et al., 2000). Several other regions have been shown to become hypomethylated in ICF patients including, α-satellite and Alu repeats (Miniou et al., 1997; Schuffenhauer et al., 1995), the non-satellite repeats D4Z4 and NBL2 (Kondo et al., 2000), the H19 gene (Schuffenhauer et al., 1995), and several genes (G6PD, SYBL1, AR and PGK1) on the inactive X chromosome of female ICF patients (Hansen et al., 2000). Biallelic expression of several genes and advanced replication timing of the normally late S phase replicating inactive X chromosome have also been noted (Hansen et al., 2000). The marked lack of autosomal genes in the list of affected DNA sequences in ICF syndrome suggests a specialized role for DNMT3b in gene-poor heterochromatin. The transcribed genes (as opposed to repeats of some kind) most consistently affected in ICF syndrome all appear to reside on the inactive X chromosome, whose methylation is likely regulated in a manner distinct from autosomal sequences.
DNA methylation and aberrant chromatin structure in ICF syndrome
In ICF syndrome it appears that loss of methylation from pericentromeric heterochromatin and the inactive X chromosome results in aberrant chromatin structure. Pericentromeric heterochromatin is massively decondensed and promoter regions of genes on the inactive X chromosome showing reactivation demonstrate increased susceptibility to nucleases, indicative of a more open chromatin configuration (Hansen et al., 2000). ICF syndrome also demonstrates that hypomethylation results in aberrant transcription, which may also be directly related to chromatin structure changes since a number of genes whose expression is altered in ICF syndrome do not display DNA methylation changes (Ehrlich et al., 2001). This suggests that DNA methylation is critical for the long-term maintenance of repressive condensed chromatin. Since the regions of the genome which DNMT3b is responsible for methylating are never properly methylated in ICF cells, the subsequent recruitment of other proteins involved in maintaining or reinforcing chromatin compaction, such as the MBDs and their associated HDACs, or the MeCP1 repressor complex, never occurs (Bird, 2002; Feng and Zhang, 2001). Therefore ICF syndrome can be regarded not only as a disease of aberrant DNA methylation patterns, but also of aberrant chromatin structure. The chromatin structural changes may be directly or indirectly related to the ability of DNMT3b to act as a transcriptional repressor via both HDAC-dependent and independent mechanisms. Alternatively, reduced methylation may result in global imbalances in transcription factor binding by allowing transcription factors access to sites that would normally be blocked by closed chromatin configuration, or by excesses of transcriptional repressors, such as the methyl-CpG binding protein-containing complexes, that may gain promiscuous access to important transcriptional control regions. The exact nature of defects in DNA methylation and chromatin structure in ICF cells will require considerable study, however, this disease further emphasizes the connection between DNA methylation, DNA methyltransferases, and chromatin structure.
DNA methylation and chromatin remodeling
We have previously discussed connections between DNA methyltransferases and chromatin in the context of histone tail modifying proteins, namely histone deacetylases. HDACs are associated with all active DNA methyltransferases and likely play an important role in determining which regions of the genome are to be methylated. A significant body of data has accumulated showing that hypermethylated regions are transcriptionally inactive, enriched in hypoacetylated histones, and the chromatin is tightly packed, reducing the access of transcription factors to these regions (Eden et al., 1998). Work over the last few years has provided tantalizing evidence for yet another connection between DNA methylation and chromatin – the possibility that DNA methylation is directly connected to, or targeted by, chromatin remodeling machines. In this last section I will discuss the evidence for this linkage and propose models for how these processes may be coordinated in mammalian cells.
ATP-dependent chromatin remodeling enzymes
Helicases are a large group of proteins, involved in a host of RNA and DNA metabolic processes, which possess a set of seven conserved motifs involved in ATP binding and hydrolysis. The helicase group can be broken down into several families. SNF2 family members are involved in chromatin remodeling (ISWI), transcription (SNF2), DNA repair (ERCC6), and recombination (RAD54) (Table 2). SNF2 (sucrose non-fermenter) was first identified in yeast as a gene essential for transcription of genes involved in sucrose fermentation and mating type switching (Eisen et al., 1995). Since then, a large number of SNF2-like genes have been discovered, most of which have homologs in organisms ranging from yeast to humans. The SNF2 family can be further divided into three subfamilies, the SNF2-like subfamily, the ISWI-like subfamily, and the CHD subfamily, based on the presence of other conserved domains (Table 2; Havas et al., 2001; Varga-Weisz, 2001). None of the SNF2 family members appear to act as helicases, instead they utilize the energy derived from ATP hydrolysis to disrupt histone-DNA interactions and allow for the physical movement, or sliding, of nucleosomes on the DNA. In this way SNF2 proteins can reorganize the chromatin structure of a region, to one more permissive for transcription factor binding and transcriptional activation, or to one with more regularly spaced, tightly packed nucleosomes characteristic of transcriptionally inactive heterochromatin (Vignali et al., 2000; Wolffe and Pruss, 1996). Inherited mutations in SNF2-like genes give rise to a number of human diseases, including Werner, Bloom and Cockayne syndromes, and Xeroderma pigmentosum (Ellis, 1997). In the next section we will review the connections between DNA methylation and chromatin remodeling enzymes of the SNF2 family and finally propose a model for how DNA methylation, histone modification, and chromatin remodeling may act in concert to establish and maintain genomic DNA methylation patterns.
The phenotypic consequences of mutations in the Arabidopsis DDM1 (decrease in DNA methylation 1) gene provided the first evidence that chromatin remodeling may be essential for proper DNA methylation patterning. DDM1 is not a DNA methyltransferase, but rather a member of the SNF2 family (Jeddeloh et al., 1999). DDM1 does not readily fit into one of the three SNF2 subfamilies listed in Table 2 due to its lack of motifs commonly associated with members of each family, but may be distantly related to the ISWI-subfamily. Mutations in DDM1 result in loss of about 70% of the total genomic 5-methylcytosine content, primarily at repetitive elements like satellites and ribosomal DNA (Jeddeloh et al., 1999; Martienssen and Henikoff, 1999). With increasing generations of DDM1 inbreeding, DNA methylation at single copy loci, including imprinted regions, is also lost, suggesting that DDM1 may be involved in maintenance methylation following DNA replication (Jeddeloh et al., 1999; Vielle-Calzada et al., 1999). DDM1 mutant plants exhibited defects in flowering time, floral and leaf morphology, and fertility (Jeddeloh et al., 1999). Similar effects were observed in plants expressing antisense to the Arabidopsis DNMT1-like gene MET1, except that the defects in the MET1 plants tended to occur after fewer generations (Finnegan et al., 1996). Demethylation and activation of transposition from transposable elements has also been observed in DDM1 mutant plants (Miura et al., 2001).
The ATRX gene is mutated in a human genetic disease called X-linked, α-thalassemia mental retardation (ATR-X) syndrome (Gibbons et al., 1997). Features of this disease include developmental abnormalities, severe mental retardation, facial dysmorphism, and α-thalassemia. Many of the mutations in the ATRX gene occur within the PHD region, a region highly homologous to the PHD regions of DNMT3a and DNMT3b (Gibbons et al., 1997). Structurally, ATRX belongs to the CHD subfamily (Table 2) of SNF2-like proteins, although it has yet to be purified from cells and shown to possess chromatin remodeling activity (Havas et al., 2001). ATRX localizes to pericentromeric heterochromatin and may act as a transcriptional regulator within a chromatin environment (Berube et al., 2000). Interestingly, it was found that ATR-X patients demonstrate DNA methylation defects in select regions of the genome. The ribosomal DNA repeats (where ATRX has also been shown to localize) were significantly hypomethylated. Conversely, a Y chromosome-specific repeat (DYZ2) was found to be hypermethylated in ATR-X patients (Gibbons et al., 2000). Therefore, unlike the effects of DDM1 mutation, ATRX mutations result in both aberrant losses and gains in DNA methylation in the genome. This result implies that aberrant chromatin structures, which may be the result of an improperly functioning or improperly targeted chromatin remodeling protein, may be able to target DNA methylation to regions that would not normally be methylated. This also supports the notion that chromatin structure changes may precede, and potentially dictate, patterns of DNA methylation.
Lsh (lymphoid-specific helicase, Hells, PASG), the murine homolog of DDM1, was initially identified as a protein highly expressed in lymphoid tissue and thought to be involved in recombination (Jarvis et al., 1996). Since then, its expression has been found to be less restricted, however Lsh expression appears to be tightly correlated with cell proliferation (Geiman and Muegge, 2000; Raabe et al., 2001). Lsh knockout mice were generated and developed relatively normally. Lsh−/− mice were born live but died shortly thereafter, possibly due to renal failure. The Lsh knockout mice show a reduced birth weight, renal lesions, and reduced numbers of lymphoid cells. T cells from Lsh−/− mice demonstrated defects in cell proliferation and high levels of apoptosis (Geiman and Muegge, 2000). Given the homology between Lsh and DDM1 (50% identity in the helicase region), genomic DNA methylation patterns were examined in the Lsh−/− mice. Remarkably, the knockout mice exhibited profound methylation defects (Dennis et al., 2001). An examination of repetitive elements, including the major and minor satellite repeats, intracisternal A-particle retroviral sequences, and Sine B1 repeats, all of which are normally heavily methylated in cells, revealed significant hypomethylation. Several single copy loci were also examined, including the β-globin, Pgk-2 and Pgk-1 genes, and the imprinting control region upstream of the H19 gene, and all revealed losses of methylation relative to normal mice. Total genomic 5-methylcytosine levels were reduced by 50–60% and all tissues appeared to be similarly affected (Dennis et al., 2001). Thus it appeared that most, but not all DNA methylation was affected by the Lsh mutation and unlike ATRX mutation, resulted only in losses of DNA methylation. No aberrant hypermethylation was reported. The Lsh work provides the most compelling evidence that chromatin structure and chromatin remodeling are critical determinants of cellular DNA methylation patterns in mammalian cells.
DNA methylation and chromatin structure – how does it all fit together?
The consistent theme arising from studies on DDM1, ATRX, and Lsh, is that these chromatin remodeling enzymes, most likely acting within large macromolecular remodeling complexes, may remodel chromatin to allow the DNA methyltransferases access to their target sites, or set up a particular chromatin configuration which is recognized by DNA methyltransferases and/or their associated proteins. Thus one could imagine that, (1) the chromatin remodeling enzyme is directly associated with a DNA methylase complex which can both remodel chromatin and methylate DNA or (2), the chromatin remodeling complex may remodel a particular region destined to undergo DNA methylation, depart, then the DNA methylation machinery would follow. The work with DDM1, ATRX, and Lsh firmly establishes an indirect connection between DNA methylation and chromatin remodeling but has not provided any evidence for a direct connection, such as a direct interaction, between a chromatin remodeling enzyme and a DNA methyltransferase (or DNA methyltransferase-associated protein). Given the universal association of HDACs with DNMTs it does not seem unreasonable to speculate that a similar direct association between one or more of the DNMTs and SNF2 family members may soon come to light.
A model which has been proposed previously is that proteins like DDM1 and Lsh may facilitate access of the DNMT to newly replicated DNA following DNA replication (Martienssen and Henikoff, 1999). DDM1 and Lsh may therefore remodel newly deposited histones in such a way as to allow DNMT1 to carry out its maintenance methylation function. The hemimethylated DNA would be converted to fully methylated DNA, and the associated HDAC2 may then deacetylate newly deposited histones and allow for maintenance of a heterochromatic state once the DNA replication machinery has passed (Rountree et al., 2000). The strict correlation between Lsh expression and cell proliferation lends support for this idea (Raabe et al., 2001). This model is highly feasible for methylation in the context of DNA replication (maintenance methylation), however evidence from other cellular processes, particularly related to de novo methylation and methylation remodeling events occurring during embryogenesis, indicates that there may be additional mechanisms. As was mentioned previously, silencing of newly introduced retroviral sequences, and of the X chromosome copy destined for inactivation, occurs well before de novo DNA methylation of those sequences. A similar situation may also be occurring during the rapid genome-wide methylation remodeling events (both demethylation and de novo methylation) that occur during embryogenesis (Reik et al., 2001). Therefore chromatin remodeling and histone modification would seem to set the stage for DNA methylation in some cases. Additional support for this notion comes from recent studies in Neurospora and Arabidopsis, which showed that loss of histone methylation (on histone H3, lysine 9) resulted in a complete or partial loss of genomic DNA methylation, respectively (Tamaru and Selker, 2001; Jackson et al., 2002). Homologous HMTs exist in mammalian cells (Rea et al., 2000), but it remains unclear whether a similar control system exists in mammals, which, unlike Neurospora, absolutely require DNA methylation for proper development.
How might DNA methylation and chromatin remodeling be coupled mechanistically? How are the HDACs also involved in this process? Unfortunately, the in vitro biochemical properties of the DDM1, ATRX and Lsh proteins have not been investigated. The biochemical properties of other SNF2-like chromatin remodeling enzymes, including ones in the ISWI subfamily, have been investigated in considerable detail and we may be able to gain insights into potential mechanisms of Lsh or DDM1 using the properties of these related proteins as a model. In the case of recombinant Drosophila ISWI, or the ISWI-containing complex NURF, it has been demonstrated that the histone H4 tail, which protrudes from the nucleosomal core particle, is essential for ISWI chromatin remodeling activity (Clapier et al., 2001; Corona et al., 2002; Hamiche et al., 2001). The H3 and H4 tails are subject to numerous post-translation modifications including acetylation, methylation and phosphorylation (Jenuwein and Allis, 2001). In particular, it appears that the base of the H4 tail, combined with nucleosome-bound DNA, is the ‘epitope’ recognized by ISWI in chromatin. Interestingly, it was recently shown that acetylation of lysines (K12 and K16) near the base of the H4 tail inhibited the chromatin remodeling ability of ISWI (Clapier et al., 2002). Indirect genetic evidence presented in the previous section indicates that chromatin remodeling is essential for proper DNA methylation patterns. Thus we propose a model (Figure 6) where these three activities, histone deacetylase, ATPase, and DNA methyltransferase, rely on each other. In this model, histone deacetylation would occur first. This may be the point at which transcriptional shutdown of the region would occur. Access of the HDACs themselves to chromatin may require other chromatin remodeling activities (Tong et al., 1998). Following histone deacetylation (and possibly methylation), the chromatin remodeling complex binds and alters the chromatin structure or nucleosome positioning in such a way as to directly facilitate access of the DNMT to the nucleosomal DNA or create a particular ‘epitope’ or signature recognized by a DNMT or DNMT-associated complex. The region is then methylated which locks the chromatin in a silent mode, which would then be further enhanced by the recruitment of methyl-CpG binding proteins and their associated repressive activities (Bird, 2002). There is ample evidence for DNMTs and HDACs as components of one complex. The ATP-dependent chromatin remodeling enzyme may also be part of the complex or act separately. Although speculation at this time, this model is testable using chromatin reconstituted in vitro and recombinant remodeling proteins and will likely be the subject of future studies. Previously proposed models, where DNA methylation occurs first and is followed by histone deacetylation and gene silencing (Baylin et al., 2001), may also operate and are by no means excluded by the model proposed here.
The past decade has seen amazing advances in our understanding of the ways in which DNA methylation contributes to transcriptional regulation and tumorigenesis. Findings from the last few years in particular have revealed the first glimpses of how the myriad epigenetic control mechanisms that exist in mammalian cells, including DNA methylation, histone acetylation, histone methylation, and ATP-dependent nucleosome positioning, may be directly connected. Histone acetylation and methylation patterns may recruit certain chromatin remodeling activities, which in turn may create docking sites for DNA methyltransferase complexes. The CpG methylated region may recruit further chromatin modulatory activities, such as the methyl-CpG binding protein complexes and their associated repressive activities, and finally lock a given region of the genome in a silent state. Emerging evidence strongly suggests that histone modifications set the stage for DNA methylation. Issues that still need to be resolved include the following: (1) which ATP-dependent chromatin remodeling machines are involved in establishing and maintaining DNA methylation patterns; (2) is the association between the ATPases and the DNMTs direct or indirect; (3) does the nature of the interaction change between undifferentiated, differentiated, and transformed cellular states; (4) do each of the three catalytically active DNMTs recognize the same chromatin ‘epitope’ for methylation or do the de novo and maintenance methyltransferases utilize fundamentally different signals, and finally; (5) have we indeed accounted for all of the DNA methylating activities in mammalian cells? Such important questions will likely be answered in the next few years as intensive research identifies and characterizes DNMT-associated proteins, and in vitro DNA methylation and chromatin remodeling systems are established. Such findings will likely be highly relevant to diseases involving aberrant DNA methylation patterns, including cancer, as well as ICF, Rett, and ATRX syndromes, and may provide the understanding to devise completely novel strategies for reversing the defects. Thus, our blurred image of the ‘tangled web’ of DNA methylation, histone modifications, and chromatin remodeling is gradually resolving into that of a precisely patterned, logically woven fabric.
Aapola U, Lyle R, Krohn K, Antonarakis SE, Peterson P . 2001 Cytogenet. Cell Genet. 92: 122–126
Aapola U, Shibuya K, Scott HS, Ollila J, Vihinen M, Heino M, Shintani A, Kawasaki K, Minoshima S, Krohn K, Antonarakis SE, Shimizu N, Kudoh J, Peterson P . 2000 Genomics 65: 293–298
Aguirre-Arteta AM, Grunewald I, Cardoso MC, Leonhardt H . 2000 Cell Growth Differ. 11: 551–559
Aoki A, Suetake I, Miyagawa J, Fujio T, Chijiwa T, Sasaki H, Tajima S . 2001 Nucleic Acids Res. 29: 3506–3512
Aoki K, Meng G, Suzuki K, Takashi T, Kameoka Y, Nakahara K, Ishida R, Kasai M . 1998 J. Biol. Chem. 273: 26698–26704
Bachman KE, Rountree MR, Baylin SB . 2001 J. Biol. Chem. 276: 32282–32287
Bacolla A, Pradhan S, Roberts RJ, Wells RD . 1999 J. Biol. Chem. 274: 33011–33019
Baylin SB, Esteller M, Rountree MR, Bachman KE, Schuebel K, Herman JG . 2001 Hum. Mol. Genet. 10: 687–692
Beard C, Li E, Jaenisch R . 1995 Genes. Dev. 9: 2325–2334
Berube NG, Smeenk CA, Picketts DJ . 2000 Hum. Mol. Genet. 9: 539–547
Bestor T, Laudano A, Mattaliano R, Ingram V . 1988 J. Mol. Biol. 203: 971–983
Bestor TH . 2000 Hum. Mol. Genet. 9: 2395–2402
Bird A . 2001 Science 294: 2113–2115
Bird A . 2002 Genes. Dev. 16: 6–21
Bird AP, Wolffe AP . 1999 Cell 99: 451–454
Bonfils C, Beaulieu N, Chan E, Cotton-Montpetit J, MacLeod AR . 2000 J. Biol. Chem. 275: 10754–10760
Bourc'his D, Xu G-L, Lin C-S, Bollman B, Bestor TH . 2001 Science 294: 2536–2539
Brehm A, Langst G, Kehle J, Clapier CR, Imhof A, Eberharter A, Muller J, Becker PB . 2000 EMBO J. 19: 4332–4341
Brehm AB, Miska EA, McCance DJ, Reid JL, Bannister AJ, Kouzarides T . 1998 Nature 391: 597–601
Cameron EE, Bachman KE, Myohanen S, Herman JG, Baylin SB . 1999 Nature Genet. 21: 103–107
Cardoso MC, Leonhardt H . 1999 J. Cell Biol. 147: 25–32
Carlson LL, Page AW, Bestor TH . 1992 Genes. Dev. 6: 2536–2541
Clapier CR, Langst G, Corona DFV, Becker PB, Nightingale KP . 2001 Mol. Cell. Biol. 21: 875–883
Clapier CR, Nightingale KP, Becker PB . 2002 Nucleic Acids Res. 30: 649–655
Corona DFV, Clapier CR, Becker PB, Tamkun JW . 2002 EMBO Rep. 3: 242–247
Csankovszki G, Nagy A, Jaenisch R . 2001 J. Cell Biol. 153: 773–783
Dennis K, Fan T, Geiman T, Yan Q, Muegge K . 2001 Genes. Dev. 15: 2940–2944
Deuring R, Fanti L, Armstrong JA, Sarte M, Papoulas O, Prestel M, Daubresse G, Verardo M, Moseley SL, Berloco M, Tsukiyama T, Wu C, Pimpinelli S, Tamkun JW . 2000 Cell 5: 355–365
Di Croce L, Raker AA, Corsaro M, Fazi F, Fanelli M, Faretta M, Fuks F, Coco FL, Kouzarides T, Nervi C, Minucci S, Pelicci PG . 2002 Science 295: 1079–1082
Doherty AS, Bartolomei MS, Schultz RM . 2002 Dev. Biol. 242: 255–266
Dong A, Yoder JA, Zhang X, Zhou L, Bestor TH, Cheng X . 2001 Nucleic Acids Res. 29: 439–448
Dyson N . 1998 Genes. Dev. 12: 2245–2262
Eden S, Hashimshony T, Keshet I, Cedar H, Thorne AW . 1998 Nature 394: 842
Ehrlich M, Buchanan KL, Tsein F, Jiang G, Sun B, Uicker W, Weemaes CMR, Smeets D, Sperling K, Belohradsky BH, Tommerup N, Misek DE, Rouillard J-M, Kuick R, Hanash SM . 2001 Hum. Mol. Genet. 10: 2917–2931
Eisen JA, Sweder KS, Hanawalt PC . 1995 Nucleic Acids Res. 23: 2715–2723
Ellis NA . 1997 Curr. Opin. Genet. Dev. 7: 354–363
Fan G, Beard C, Chen RZ, Csankovszki G, Sun Y, Siniaia M, Biniszkiewicz D, Bates B, Lee PP, Kuhn R, Trumpp A, Poon C-S, Wilson CB, Jaenisch R . 2001 J. Neurosci. 21: 788–797
Feng Q, Zhang Y . 2001 Genes. Dev. 15: 827–832
Finnegan EJ, Peacock WJ, Dennis ES . 1996 Proc. Natl. Acad. Sci. USA 93: 8449–8454
Flynn J, Glickman JF, Reich NO . 1996 Biochemistry 35: 7308–7315
Franceschini P, Martino S, Ciocchini M, Ciuti E, Vardeu MP, Guala A, Signorile F, Camerano P, Franceschini D, Tovo PA . 1995 Eur. J. Pediatr. 154: 840–846
Fuks F, Bergers WA, Brehm A, Hughes-Davies L, Kouzarides T . 2000 Nature Genet. 24: 88–91
Fuks F, Burgers WA, Godin N, Kasai M, Kouzarides T . 2001 EMBO J. 20: 2536–2544
Geiman TM, Muegge K . 2000 Proc. Natl. Acad. Sci. USA 97: 4772–4777
Gibbons RJ, Bachoo S, Picketts DJ, Aftimos S, Asenbauer B, Bergoffen J, Berry SA, Dahl N, Fryer A, Keppler K, Kurosawa K, Levin ML, Masuno M, Neri G, Pierpont ME, Slaney SF, Higgs DR . 1997 Nature Genet 17: 146–148
Gibbons RJ, McDowell TL, Raman S, O'Rourke DM, Garrick D, Ayyub H, Higgs DR . 2000 Nature Genet. 24: 368–371
Glickman JF, Pavlovich JG, Reich NO . 1997 J. Biol. Chem. 272: 17851–17857
Gowher H, Jeltsch A . 2001 J. Mol. Biol. 309: 1201–1208
Hamiche A, Kang J-G, Dennis C, Xiao H, Wu C . 2001 Proc. Natl. Acad. Sci. USA 98: 14316–14321
Hansen RS, Stoger R, Wijmenga C, Stanek AM, Canfield TK, Luo P, Matarazzo MR, D'Esposito M, Feil R, Gimelli G, Weemaes CMR, Laird CD, Gartler SM . 2000 Hum. Mol. Genet. 9: 2575–2587
Hansen RS, Wijmenga C, Luo P, Stanek AM, Canfield TK, Weemaes CMR, Gartler SM . 1999 Proc. Natl. Acad. Sci. USA 96: 14412–14417
Hassan KMA, Norwood T, Gimelli G, Gartler SM, Hansen RS . 2001 Hum. Mol. Genet. 109: 452–462
Havas K, Whitehouse I, Owen-Hughes T . 2001 Cell. Mol. Life Sci. 58: 673–682
Heard E, Rougeulle C, Arnaud D, Avner P, Allis CD, Spector DL . 2001 Cell 107: 727–738
Howell CY, Bestor TH, Ding F, Latham KE, Mertineit C, Trasler JM, Trasler JM, Chaillet JR . 2001 Cell 104: 829–838
Hsieh C-L . 1999 Mol. Cell. Biol. 19: 8211–8218
Hsu D-W, Lin M-J, Lee T-L, Wen S-C, Chen X, Shen C-KJ . 1999 Proc. Natl. Acad. Sci. USA 96: 9751–9756
Hung M-S, Karthikeyan N, Huang B, Koo H-C, Kiger J, Shen C-KJ . 1999 Proc. Natl. Acad. Sci. USA 96: 11940–11945
Ito T, Levenstein ME, Fyodorov DV, Kutach AK, Kobayashi R, Kadonaga JT . 1999 Genes. Dev. 13: 1529–1539
Jackson JP, Lindroth AM, Cao X, Jacobsen SE . 2002 Nature advance online publication 17 March
Jackson-Grusby L, Beard C, Possemato R, Tudor M, Fambrough D, Csankovzki G, Dausman J, Lee P, Wilson C, Lander E, Jaenisch R . 2001 Nature Genet. 27: 31–39
Jarvis CD, Geiman T, Vila-Storm MP, Osipovich O, Akella U, Candeias S, Nathan I, Durum SK, Muegge K . 1996 Gene 169: 203–207
Jeanpierre M, Turleau C, Aurias A, Prieur M, Ledeist F, Fischer A, Viegas-Pequignot E . 1993 Hum. Mol. Genet. 2: 731–735
Jeddeloh JA, Stokes TL, Richards EJ . 1999 Nature Genet. 22: 94–97
Jenuwein T, Allis CD . 2001 Science 293: 1074–1080
Jones PA, Laird PW . 1999 Nature Genet. 21: 163–166
Kal AJ, Mahmoudi T, Zak NB, Verrijzer CP . 2000 Genes Dev. 14: 1058–1071
Kennedy BK, Barbie DA, Classon M, Dyson N, Harlow E . 2000 Genes Dev. 14: 2855–2868
Kondo T, Bobek MP, Kuick R, Lamb B, Zhu X, Narayan A, Bourc'his D, Viegas-Pequignot E, Ehrlich M, Hanash SM . 2000 Hum. Mol. Genet. 9: 597–604
Lei H, Oh SP, Okano M, Juttermann R, Goss KA, Jaenisch R, Li E . 1996 Development 122: 3195–3205
Leonhardt H, Page AW, Weier H, Bestor TH . 1992 Cell 71: 865–873
LeRoy G, Loyola A, Lane WS, Reinberg D . 2000 J. Biol. Chem. 275: 14787–14790
Li E, Beard C, Jaenisch R . 1993 Nature 366: 362–365
Li E, Bestor TH, Jaenisch R . 1992 Cell 69: 915–926
Li H, Leo C, Zhu J, Wu X, O'Neil J, Park E-J, Chen JD . 2000 Mol. Cell. Biol. 20: 1784–1796
Lin M-J, Lee T-L, Hsu D-W, Shen C-KJ . 2000 FEBS Lett. 469: 101–104
Liu Y, Oakeley EJ, Sun L, Jost J-P . 1998 Nucleic Acids Res. 26: 1038–1045
Luo RX, Postigo AA, Dean DC . 1998 Cell 92: 463–473
Lyko F, Ramsahoye BH, Kashevsky H, Tudor M, Mastrangelo M-A, Orr-Weaver TL, Jaenisch R . 1999 Nature Genet. 23: 363–366
Lyko F, Whittaker AJ, Orr-Weaver TL, Jaenisch R . 2000 Mech. Dev. 95: 215–217
Martienssen R, Henikoff S . 1999 Nature Genet. 22: 6–7
Meng G, Inazawa J, Ishida R, Tokura K, Nakahara K, Aoki K, Kasai M . 2000 Gene 242: 59–64
Mertineit C, Yoder JA, Taketo T, Laird DW, Trasler JM, Bestor TH . 1998 Development 125: 889–897
Michaelson JS, Bader D, Kuo F, Kozak C, Leder P . 1999 Genes. Dev. 13: 1918–1923
Miniou P, Jeanpierre M, Bourc'his D, Barbosa ACC, Blanquet V, Viegas-Pequignot E . 1997 Hum. Genet. 99: 738–745
Miura A, Yonebayashi S, Watanabe K, Toyama T, Shimada H, Kakutani T . 2001 Nature 411: 212–214
Muller S, Hoege C, Pyrowolakis G, Jentsch S . 2001 Nature Rev. Mol. Cell Biol. 2: 202–210
Nielsen SJ, Schneider R, Bauer U-M, Bannister AJ, Morrison A, O'Carroll D, Firestein R, Cleary M, Jenuwein T, Herrera RE, Kouzarides T . 2001 Nature 412: 561–565
Okano M, Bell DW, Haber DA, Li E . 1999 Cell 99: 247–257
Okano M, Xie S, Li E . 1998a Nature Genet. 19: 219–220
Okano M, Xie S, Li E . 1998b Nucleic Acids Res. 26: 2536–2540
Pannell D, Osborne CS, Yao S, Sukonnik T, Pasceri P, Karaiskakis A, Okano M, Li E, Lipshitz HD, Ellis J . 2000 EMBO J. 19: 5884–5894
Poot RA, Dellaire G, Hulsmann BB, Grimaldi MA, Corona DFV, Becker PB, Bickmore WA, Varga-Weisz PD . 2000 EMBO J. 19: 3377–3387
Pradhan S, Bacolla A, Wells RD, Roberts RJ . 1999 J. Biol. Chem. 274: 33002–33010
Pradhan S, Kim G-D . 2002 EMBO J. 21: 779–788
Pradhan S, Talbot D, Sha M, Benner J, Hornstra L, Li E, Jaenisch R, Roberts RJ . 1997 Nucleic Acids Res. 25: 4666–4673
Qiu C, Sawada K, Zhang X, Cheng X . 2002 Nature Struct. Biol. 9: 217–224
Raabe EH, Abdurrahman L, Behbehani G, Areci RJ . 2001 Dev. Dyn. 221: 92–105
Rea S, Eisenhaber F, O'Caroll D, Strahl BD, Sun Z-W, Schmid M, Opravil S, Mechtler K, Ponting CP, Allis CD, Jenuwein T . 2000 Nature 406: 593–599
Reik W, Dean W, Walter J . 2001 Science 293: 1089–1093
Rhee I, Jair K-W, Yen R-WC, Lengauer C, Herman JG, Kinzler KW, Vogelstein B, Baylin SB, Schuebel KE . 2000 Nature 404: 1003–1007
Robertson KD . 2001 Oncogene 20: 3139–3155
Robertson KD, Ait-Si-Ali S, Yokochi T, Wade PA, Jones PL, Wolffe AP . 2000a Nature Genet. 25: 338–342
Robertson KD, Keyomarsi K, Gonzales FA, Velicescu M, Jones PA . 2000b Nucleic Acids Res. 28: 2108–2113
Robertson KD, Uzvolgyi E, Liang G, Talmadge C, Sumegi J, Gonzales FA, Jones PA . 1999 Nucleic Acids Res. 27: 2291–2298
Robertson KD, Wolffe AP . 2000 Nature Rev. Genet. 1: 11–19
Rountree MR, Bachman KE, Baylin SB . 2000 Nature Genet. 25: 269–277
Ryan RF, Schultz DC, Ayyanathan K, Singh PB, Friedman JR, Fredericks WJ, Rasucher FR . 1999 Mol. Cell. Biol. 19: 4366–4378
Schuffenhauer S, Bartsch O, Stumm M, Buchholz T, Petropoulou T, Kraft S, Belohradsky B, Hinkel GK, Meitinger T, Wegner R-D . 1995 Hum. Genet. 96: 562–571
Smeets DFCM, Moog U, Weemaes CMR, Vaes-Peeters G, Merkx GFM, Niehof JP, Hamers G . 1994 Hum. Genet. 94: 240–246
Tagarro I, Fernandez-Peralta AM, Gonzales-Aguilera JJ . 1994 Hum. Genet. 93: 383–388
Tamaru H, Selker EU . 2001 Nature 414: 277–283
Tatematsu K-i, Yamazaki T, Ishikawa F . 2000 Genes Cells 5: 677–688
The Arabidopsis Genome Initiative. 2000 Nature 408: 796–815
Tong JK, Hassig CA, Schnitzler GR, Kingston RE, Schreiber SL . 1998 Nature 395: 917–921
Tran HG, Steger DJ, Iyer VR, Johnson AD . 2000 EMBO J. 19: 2323–2331
Tucker KL . 2001 Neuron 30: 649–652
Tuck-Muller CM, Narayan A, Tsien F, Smeets DFCM, Sawyer J, Fiala ES, Sohn OS, Ehrlich M . 2000 Cytogenet. Cell Genet. 89: 121–128
Varga-Weisz P . 2001 Oncogene 20: 3076–3085
Vielle-Calzada J-P, Thomas J, Spillane C, Coluccio A, Hoeppner MA, Grossniklaus U . 1999 Genes Dev. 13: 2971–2982
Vignali M, Hassan AH, Neely KE, Workman JL . 2000 Mol. Cell. Biol. 20: 1899–1910
Wade PA, Gegonne A, Jones PL, Ballestar E, Aubry F, Wolffe AP . 1999 Nature Genet. 23: 62–66
Walsh CP, Chaillet JR, Bestor TH . 1998 Nature Genet. 20: 116–117
Wang W, Cote J, Xue Y, Zhou S, Khavari PA, Biggar SR, Muchardt C, Kalpana GV, Goff SP, Yaniv M, Workman JL, Crabtree GR . 1996 EMBO J. 15: 5370–5382
Whitehouse I, Flaus A, Cairns BR, White MF, Workman JL, Owen-Hughes T . 1999 Nature 400: 784–787
Wilkinson CRM, Bartlett R, Nurse P, Bird AP . 1995 Nucleic Acids Res. 23: 1995
Wolffe AP, Pruss D . 1996 Cell 84: 817–819
Xie S, Wang Z, Okano M, Nogami M, Li Y, He W-W, Okumura K, Li E . 1999 Gene 236: 87–95
Xu G-L, Bestor TH, Bourc'his D, Hsieh C-L, Tommerup N, Bugge M, Hulten M, Qu Russo JJ, Viegas-Pequignot E . 1999 Nature 402: 187–191
Yen R-WC, Vertino PM, Nelkin BD, Yu JJ, El-Deiry W, Cumaraswamy A, Lennon GG, Trask BJ, Celano P, Baylin SB . 1992 Nucleic Acids Res. 20: 2287–2291
Yoder JA, Bestor TH . 1998 Hum. Mol. Genet. 7: 279–284
Yoder JA, Walsh CP, Bestor TH . 1997 Trends Genet. 13: 335–340
Yokochi T, Robertson KD . 2002 J. Biol. Chem. 277: 11735–11745
Zhong S, Salomoni P, Pandolfi PP . 2000a Nature Cell Biol. 2: E85–E90
Zhong S, Salomoni P, Ronchetti S, Guo A, Ruggero D, Pandolfi PP . 2000b J. Exp. Med. 191: 631–639
KD Robertson is a Cancer Scholar supported by the National Cancer Institute (NIH grant CA84535-01). I thank Andrea Kahler Robertson for critical reading of the manuscript. This article is dedicated to my former mentor and friend, Alan Wolffe, who died tragically last year.
About this article
Cite this article
Robertson, K. DNA methylation and chromatin – unraveling the tangled web. Oncogene 21, 5361–5379 (2002). https://doi.org/10.1038/sj.onc.1205609
- DNA methyltransferase
- DNA methylation
- histone deacetylase
- histone methylase
- transcriptional repression
Deleterious point mutations in T‐cell acute lymphoblastic leukemia: Mechanistic insights into leukemogenesis
International Journal of Cancer (2021)
Expert Review of Clinical Immunology (2021)
Virus Research (2021)
Bioactive Materials (2021)