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1 Laboratory of Cellular Biochemistry, Department of Animal Resource Sciences and Veterinary Medical Sciences, The University of Tokyo, Tokyo 113-8657, Japan
2 Laboratory of Mouse Model, Experimental Research Center for Infectious Diseases, Institute for Virus Research, Kyoto University, Kyoto 606-8507, Japan
3 Department of Molecular Genetics and Medicine, Albert Einstein College of Medicine, Bronx, NY 10461, USA
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Abstract |
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Introduction |
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Regions with hypermethylated CpGs are found in condensed chromatin, whereas the hypomethylated regions are associated with relaxed chromatin (Bird 2002). Chromatin structure is also associated with the modification of histone amino-terminal tails. Methylation at lysines 9 and 27 of histone H3 (H3-K9, -K27) is often found at heterochromatic regions and transcriptionally-inactive loci in the genome in a wide variety of organisms (Sims et al. 2003). Most known histone methyltransferases (HMTs) have a SET domain, and over 300 SET domain proteins have been found (Sims et al. 2003). Several mammalian HMTs mediating H3-K9 and -K27 methylation have been identified. SUV39H/Suv39h, ESET (officially SETDB1/Setdb1), GLP (officially EHMT1/Ehmt1) and G9a (officially EHMT2/Ehmt2) catalyze H3-K9 methylation. HMTs catalyzing H3-K27 methylation include EZH2/Ezh2, GLP and G9a. Mice deficient for these HMTs show embryonic lethality, except for Suv39h, which predominantly localizes at centromeric heterochromatin (Aagaard et al. 1999).
G9a catalyzes mono and dimethylation of both H3-K9 and H3-K27 in vitro (Tachibana et al. 2001, 2002), although H3-K27 methylation activity in vivo has not been demonstrated. As G9a is localized in euchromatin (Tachibana et al. 2002), it is presumed to influence epigenetic regulation in these gene-containing regions. So far, however, only a few genes have been identified as targets of G9a: Mage-a2 (Tachibana et al. 2002) and Snrpn (Xin et al. 2003) in mouse.
We hypothesized that the formation of cell-type/tissue-dependent DNA methylation profiles is influenced by histone tail modifications. A model system using G9a deficient cells allows this hypothesis to be tested. A causal relationship would be supported by G9a deficiency causing a combination of decreased DNA methylation as well as reduced H3-K9 and/or H3-K27 methylation levels at specific loci. Restriction landmark genomic scanning (RLGS) using Not I, a DNA methylation sensitive restriction enzyme, enabled us to analyze the DNA methylation status of approximately 2000 CpG-rich loci in wild-type, G9a–/– and G9a–/– ES cells with a G9a transgene (G9a–/–Tg). We found multiple specific G9a target loci distributed within the euchromatic compartment of the genome, at which G9a deficiency was associated with concordant decreases in DNA and histone tail methylation. Thus, G9a activity contributes to the establishment of the DNA methylation profile of mammalian cells.
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Results |
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While the RLGS profile of wild-type ES cells contained about 1300 spots that were distinguishable from each other, the profile of G9a–/– ES cells had new 32 spots in addition to the 1300 spots observed in wild-type cells (Fig. 1). The emergence of these new RLGS spots indicated that G9a deficiency caused DNA demethylation at multiple loci, at which DNA is normally hypermethylated in wild-type ES cells.
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Identification of G9a target loci by Vi-RLGS and methylation-sensitive quantitative real-time PCR
Candidates for each of the 32 loci affected by G9a deficiency were selected from the virtual-image RLGS (Vi-RLGS) database (Matsuyama et al. 2003; Hattori et al. 2004a), which shows all possible RLGS spots (Supplementary Fig. S1). We tested the methylation status of the candidate loci using methylation-sensitive quantitative real-time PCR and succeeded to identify ten Not I sites that represent G9a targets (Table 1A). The four hypermethylated and two hypomethylated control loci that were randomly selected from the Vi-RLGS negative set did not show differences in DNA methylation levels between wild-type and G9a–/– ES cells (Table 1B).
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G9a targets are located in euchromatic gene loci distributed genome-wide
The relationships of the G9a target Not I sites and their surrounding genes are shown in Fig. 2B. Each of the Not I sites is located at a gene-containing region that has one (#110, 188, 289, 291) to 12 (#284) genes within the flanking 100 kbp. These targets do not concentrate in any particular genomic region. Eight out of the ten loci reside in CpG islands (#188) or CpG-rich regions (#110, 258, 282, 284, 287, 289 and 291; Table 1A). Although spots #284, 289 and 290 loci are in regions relatively nearer to centromeres or telomeres, none is within 2 Mbp of these chromosomal regions as judged by their positions in the May 2005 assembly of the mouse genome at the UCSC Genome Browser <http://genome.ucsc.edu/>. These data indicated that the ten G9a target loci are distributed genome-wide at gene loci within euchromatin.
H3-K9 and/or H3-K27 demethylation at the loci where DNA was demethylated by G9a deficiency
At genuine G9a target loci, G9a deficiency should impair local histone H3-K9 methylation. We investigated the H3-K9 methylation status of wild-type and G9a–/– ES cells at the ten identified target loci by using the chromatin immunoprecipitation (ChIP) assay (Fig. 2C). The Mage-a2 gene locus was a positive control known G9a target locus (Tachibana et al. 2002). In wild-type ES cells, H3-K9 dimethylation was observed at the nine loci (#110, 188, 258, 282, 287, 289, 290, 291 and 377), while a weaker signal was detected at the #284 locus. H3-K9 dimethylation levels showed a tendency to decline at all loci except the #284 locus in G9a–/– ES cells. At all ten loci, H3-K27 dimethylation was also detected in wild-type ES cells (Fig. 2C). A decrease of H3-K27 methylation levels was observed at all loci except spots #287 and 289, suggesting that G9a also catalyzes H3-K27 dimethylation in vivo.
The levels of dimethylation at the H3-K9 and H3-K27 residues were not always concordant with each other at a given locus. At the locus corresponding to spot #284, G9a deficiency caused a reduction of the methylation level of H3-K27 but not H3-K9, while the opposite was observed at the loci corresponding to spots #287 and 289 (Fig. 2C). However, the levels of H3-K9 and/or H3-K27 dimethylation were reduced at all of the ten loci in the G9a–/– cells with various ranges among the loci.
DNA demethylation in G9a–/– cells extends beyond a nucleosome unit
A subset of loci (#110, 291, 282 and 284) was further analyzed by the bisulphite sequencing method (Fig. 3). At the #110 locus, all 16 CpGs in the 468 bp region were hypermethylated (78%–90%) in wild-type ES cells, while the levels were reduced to 53%–63% in the G9a–/– cells (Fig. 3A(a)). Analysis of the #291 locus also revealed that all 14 CpGs in 442 bp were markedly demethylated in G9a–/– ES cells (Fig. 3A(b)). Even at the #284 locus, where the H3-K9 methylation status was less responsive to the G9a deficiency (Fig. 2A), the 30 CpGs in the 559 bp around Not I site were strongly hypomethylated in G9a–/– cells (Fig. 3A(c)). More extensive DNA demethylation was also observed at the 5'-upstream region of the first exon of Tm7sf2 gene (#284, region II), which is 2.5 kbp distant from the Not I site (region I; Fig. 3A(c)). Thus, alteration of DNA methylation by G9a deficiency extends beyond a nucleosome unit (
200 bp) flanking these target loci.
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Discussion |
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The present data showed DNA demethylation in G9a–/– ES cells, indicating that G9a contributes to maintaining DNA methylation. In addition, G9a also functions to facilitate de novo methylation of DNA because a G9a transgene reconstituted the normal DNA methylation status in G9a–/– ES cells. Since G9a does not have the catalytic domain of Dnmts, G9a's effect on DNA methylation must be indirect, possibly through H3-K9 and/or -K27 methylation. The histone H3 tail methylated at K9 and K27 interacts with CHROMOMETHYLASE3 (CMT3), a DNA methyltransferase which induces cytosine methylation at CpNpG sequences in Arabidopsis (Lindroth et al. 2004). This interaction requires the chromodomain of CMT3, whereas a mammalian functional Dnmt with a chromodomain has not been reported. In mammals, methylated H3-K9 binds to the chromodomain of HP1 (Lachner et al. 2001), which interacts in turn with Dnmts (Fuks et al. 2003). Although G9a is excluded from HP1-dense heterochromatic regions (Tachibana et al. 2001), HP1 also functions at euchromatic genes such as cyclin E (Nielsen et al. 2001), which has a CpG island and is expressed in a tissue-specific pattern in adult mouse (Geng et al. 2001). At the tissue-specific beta-Globin and Pgk-2 genes, Lsh, a member of the SNF2 chromatin remodelling family, is required for DNA methylation (Dennis et al. 2001). The multiple G9a target loci identified in this study will provide clues to reveal how G9a directs DNA methylation with various epigenetic factors including uncharacterized proteins.
In vitro studies have proved that G9a has catalytic activities for both H3-K9 and -K27 dimethylation (Tachibana et al. 2001). The present study demonstrated that G9a induces H3-K27 dimethylation in vivo. Until now, it has been believed that G9a does not participate in H3-K27 methylation in vivo because G9a deficiency did not reduce overall level of H3-K27 dimethylation detected by immunocytochemical and Western blot analyses (Peters et al. 2003). In addition, G9a induced H3-K9 methylation but not H3-K27 methylation at the Xist locus (Rougeulle et al. 2004). Thus, no report has documented in vivo target loci for G9a-induced H3-K27 methylation. On the other hand, it has been well characterized that G9a induces H3-K9 dimethylation at euchromatic regions in vivo (Tachibana et al. 2002; Peters et al. 2003; Xin et al. 2003). We showed that not only H3-K9 but also H3-K27 methylation level was reduced in G9a–/– cells at multiple genomic loci. It is clear that G9a is responsible for H3-K9 as well as H3-K27 methylation in vivo.
Repetitive sequences have been characterized as hot spots where HMTs control DNA methylation as well as histone methylation. Transposons are the target of H3-K9 methyltransferases Dim-5 in Neurospora crassa (Tamaru & Selker 2001) and KRYPTONITE in Arabidopsis (Jackson et al. 2002) because the DNA methylation status at these regions was impaired when these enzymes were mutated. The DNA methylation status of satellite repeats in mouse pericentromeric region was altered in Suv39h deficient cells (Lehnertz et al. 2003). Murine G9a deficient cells showed decrease in overall levels of H3-K9 methylation detected by immunohistochemistry (Tachibana et al. 2002), which would mostly detect modification status in bulk interspersed repetitive sequences in the genome. Our analysis does not exclude the possibility that G9a could contribute to DNA methylation on these non-CpG-rich repetitive sequences, because RLGS tests almost exclusively CpG-rich gene loci. Recently, Xin et al. (2003) found that G9a is required for DNA methylation at an imprinted gene, Snrpn, which is located in euchromatin. Taken together with the present study, the experimental evidence indicates that G9a functions to regulate DNA methylation status at mammalian euchromatic gene loci in association with regulation of methylation of H3-K9 and/or -K27.
As described above, HMTs are capable of directing DNA methylation. Significantly, the DNA methylation levels of the G9a-deficient ES cells were almost half of the wild-type levels. In addition, residual methylation of H3-K9 and -K27 was detected at the G9a target loci in G9a–/– ES cells. The partial demethylation of DNA may be due to compensation for G9a deficiency by other HMTs such as GLP (Tachibana et al. 2005,), ESET (Schultz et al. 2002) and Ezh2 (Cao et al. 2002) that help to maintain the residual levels of H3-K9 and/or -K27 methylation that we observed. This possible compensation by other HMTs may explain the relatively small change in H3-K9/K27 dimethylation at the G9a target loci in G9a–/– ES cells, although future study will be needed.
The DNA methylation profiles of T-DMRs are unique to specific cell types and tissues (Shiota 2004). We previously found that Dnmt3a and 3b regulate DNA methylation at CpG islands including T-DMRs more strongly than Dnmt1, which has a preference for repetitive sequences (Hattori et al. 2004a). Therefore, Dnmt3a and 3b are global regulators of DNA methylation profiles of T-DMRs. In the process of establishment of a DNA methylation profile during events such as differentiation and development, DNA methylation and demethylation occur at specific T-DMRs depending on the cell-types (Ohgane et al. 2002; Shiota et al. 2002). Intriguingly, RLGS analyses on Dnmt1–/– and Dnmt3a–/–3b–/– ES cells showed DNA demethylation at 247 CpG-rich loci (Hattori et al. 2004a). G9a deficiency, in contrast, caused DNA demethylation at 32 loci. This restricted number of the loci affected by G9a suggests that local specific regulation of DNA methylation occurs by participation of G9a. Thus, the formation of cell-type/tissue-dependent DNA methylation profiles is related to other influences including histone tail modifications. The identification of G9a targets provides insights into the question of how these global regulators act locally. G9a is involved in the local regulation of DNA methylation in association with H3-K9 and/or H3-K27 dimethylation at these selected loci.
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Experimental procedures |
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The establishment and characterization of wild-type (TT2 line), G9a–/– and G9a–/–Tg murine ES cell lines were described previously (Tachibana et al. 2002). At least ten generations have already passed since their establishment. These cells were cultivated on a gelatin-coated culture dish in DMEM (Invitrogen, CA) containing 15% fetal calf serum (FCS) and 1000 U/ml of leukemia inhibitory factor (ESGRO, Chemicon, CA).
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RLGS |
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Spot identification by Vi-RLGS
The Vi-RLGS profile was created by Vi-RLGS software package using mouse draft genome sequence (MGCSv3_release3) provided by GENBANK <ftp://ftp.ncbi.nih.gov/genomes/M_musculus/> (Matsuyama et al. 2003; Hattori et al. 2004a). By comparing the Vi-RLGS with RLGS profiles, several candidate sequences for each of the 32 RLGS spots that appeared in G9a–/– ES cells were chosen (Supplementary Fig. S1). These sequences were localized using genome browsers (UCSC <http://genome.ucsc.edu/> and NCBI <http://www.ncbnlm.nih.gov/>) to obtain the surrounding sequence information. The DNA methylation status of the Not I sites in these candidate sequences was analyzed by methylation-sensitive quantitative real-time PCR described below. We also analyzed the negative control loci selected from the Vi-RLGS profile. The CpGPlot program <http://www.ebi.ac.uk/emboss/cpgplot/> indicated regions where (G+C) content are over 0.50 and observed to expected CpG ratio are over 0.60 within ±1 kbp of the Not I sites. These regions were classified as CpG islands or as CpG-rich regions by their length: over 200 bp, and 100–200 bp, respectively.
Methylation-sensitive quantitative real-time PCR
Methylation-sensitive quantitative real-time PCR was performed as previously described (Hattori et al. 2004a). This method utilizes digestion of genomic DNA with Not I, a methylation-sensitive restriction enzyme, prior to quantitative PCR. Briefly, genomic DNA was digested with Pst I and an aliquot was subsequently treated with Not I. Forty nanograms of the DNA treated with or without Not I were subjected to PCR with the primer pair amplifying the genomic fragment containing the Not I site. The amplification was monitored by SYBR green (all loci except spot #258) in a SYBR green PCR Master Mix (Applied Biosystems, Foster City, CA) or Tamra (spot #258) in a TaqMan Universal PCR Master Mix (Applied Biosystems) on an ABI Prism 7000 or a 7900HT Sequence Detection System (Applied Biosystems) or a DNA Engine Opticon 2 system (BioRad, Hercules, CA) according to the manufacturers’ protocols. The DNA methylation level at each Not I site was defined as the proportion of the amount of undigested DNA in the Not I treated solution to that without the Not I treatment, calculated as below: DNA methylation level (%) = 100 x (1 + E)(Ctb–Cta) x k–1, where E is the efficiency of the PCR using the primer pair; Cta and Ctb are Ct (threshold cycle) value in the PCR using DNA with and without the Not I treatment, respectively; k is the proportion of the total amount of Not I treated DNA to untreated DNA. To obtain the k value, real-time PCR was performed using primer pairs of Met, Xist1 and 5-lipo2 that were designed to amplify fragments without the Not I site. For all samples, PCR was performed at least 3 times independently. The primers, probe and PCR efficiencies used in the PCRs are listed in Supplementary Table S1.
ChIP assay
The ChIP assay was performed according to our previous report (Hattori et al. 2004b). Harvested ES cells were incubated in DMEM containing 15% FCS and 1% formaldehyde at 25 °C for 20 min to cross-link chromatin. The cells were sonicated to shear their DNA to lengths between 200 and 1000 bp. Aliquots of sonicated cell lysates (1 x 106 cells) were incubated with antibodies against dimethylH3-K9 (Cat. # 07–212, Lot. # 27563, Upstate, NY) and dimethylH3-K27 (Cat. # 07–452, Lot. # 24461, Upstate) at 4 °C for 16 h to immunoprecipitate chromatin. For a negative control, IgG from non-immunized rabbit (Normal rabbit IgG, Cat. # 12-370, Upstate) was used. Recovered DNA was analyzed by PCR with 33 cycles. Intensities of PCR bands stained with ethidium bromide were measured using NIH image 1.61 software <http://rsb.info.nih.gov/nih-image/>. Relative intensity of a PCR band amplified from the immunoprecipitated chromatin that was normalized with the input DNA was calculated as follows: Relative intensity = ([IP]—[IgG]) / [Input], where [IP], [IgG] and [Input] are intensities of PCR bands from 0.25% of input DNA ([Input]) immunoprecipitated DNA with anti-dimethylH3-K9 or K27 antibody ([IP]) and negative control IgG ([IgG]). Primers used in the PCRs are listed in Supplementary Table S1.
Sodium bisulphite genomic sequencing
This was performed as previously described (Hattori et al. 2004a). Briefly, genomic DNA denatured by NaOH was incubated with sodium metabisulphite (pH 5.0) at 55 °C for 16 h in the dark. The modified DNA was purified, and then the bisulphite reaction was completed with NaOH. The DNA solution was neutralised by addition of ammonium acetate (pH 7.0). The recovered DNA was amplified by PCR with primers listed in Supplementary Table S1. Each amplified fragment was cloned and sequenced.
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Acknowledgements |
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Footnotes |
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* Correspondence: E-mail: ashiota{at}mail.ecc.u-tokyo.ac.jp
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References |
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Received: 30 May 2006
Accepted: 20 September 2006
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  S. Yagi, K. Hirabayashi, S. Sato, W. Li, Y. Takahashi, T. Hirakawa, G. Wu, N. Hattori, N. Hattori, J. Ohgane, et al. DNA methylation profile of tissue-dependent and differentially methylated regions (T-DMRs) in mouse promoter regions demonstrating tissue-specific gene expression Genome Res., December 1, 2008; 18(12): 1969 - 1978. [Abstract] [Full Text] [PDF]
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A. Smallwood, P.-O. Esteve, S. Pradhan, and M. Carey Functional cooperation between HP1 and DNMT1 mediates gene silencing Genes & Dev., May 15, 2007; 21(10): 1169 - 1178. [Abstract] [Full Text] [PDF]
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 N. Hattori, Y. Imao, K. Nishino, N. Hattori, J. Ohgane, S. Yagi, S. Tanaka, and K. Shiota Epigenetic regulation of Nanog gene in embryonic stem and trophoblast stem cells Genes Cells, March 1, 2007; 12(3): 387 - 396. [Abstract] [Full Text] [PDF]
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