Chd2 interacts with H3.3 to determine myogenic cell fate
2012; Springer Nature; Volume: 31; Issue: 13 Linguagem: Inglês
10.1038/emboj.2012.136
ISSN1460-2075
AutoresAkihito Harada, Seiji Okada, Daijiro Konno, Jun Odawara, Tomohiko Yoshimi, Saori Yoshimura, Hiromi Kumamaru, Hirokazu Saiwai, Toshiaki Tsubota, Hitoshi Kurumizaka, Koichi Akashi, Taro Tachibana, Anthony N. Imbalzano, Yasuyuki Ohkawa,
Tópico(s)Adenosine and Purinergic Signaling
ResumoArticle8 May 2012free access Chd2 interacts with H3.3 to determine myogenic cell fate Akihito Harada Akihito Harada Department of Advanced Medical Initiatives, JST-CREST, Faculty of Medicine, Kyushu University, Fukuoka, Japan Search for more papers by this author Seiji Okada Seiji Okada Department of Advanced Medical Initiatives, JST-CREST, Faculty of Medicine, Kyushu University, Fukuoka, Japan Search for more papers by this author Daijiro Konno Daijiro Konno Laboratory for Cell Asymmetry, Center for Developmental Biology, RIKEN, Kobe, Japan Search for more papers by this author Jun Odawara Jun Odawara Department of Advanced Medical Initiatives, JST-CREST, Faculty of Medicine, Kyushu University, Fukuoka, Japan Search for more papers by this author Tomohiko Yoshimi Tomohiko Yoshimi Department of Bioengineering, Graduate School of Engineering, Osaka City University, Osaka, Japan Search for more papers by this author Saori Yoshimura Saori Yoshimura Department of Bioengineering, Graduate School of Engineering, Osaka City University, Osaka, Japan Search for more papers by this author Hiromi Kumamaru Hiromi Kumamaru Department of Advanced Medical Initiatives, JST-CREST, Faculty of Medicine, Kyushu University, Fukuoka, Japan Search for more papers by this author Hirokazu Saiwai Hirokazu Saiwai Department of Advanced Medical Initiatives, JST-CREST, Faculty of Medicine, Kyushu University, Fukuoka, Japan Search for more papers by this author Toshiaki Tsubota Toshiaki Tsubota Experimental Research Center for Infectious Diseases, Institute for Virus Research, Kyoto University, Kyoto, Japan Search for more papers by this author Hitoshi Kurumizaka Hitoshi Kurumizaka Laboratory of Structural Biology, Graduate School of Advanced Science and Engineering, Waseda University, Tokyo, Japan Search for more papers by this author Koichi Akashi Koichi Akashi Department of Medicine and Biosystemic Science, Faculty of Medicine, Kyushu University, Fukuoka, Japan Search for more papers by this author Taro Tachibana Taro Tachibana Department of Bioengineering, Graduate School of Engineering, Osaka City University, Osaka, Japan Search for more papers by this author Anthony N Imbalzano Anthony N Imbalzano Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA, USA Search for more papers by this author Yasuyuki Ohkawa Corresponding Author Yasuyuki Ohkawa Department of Advanced Medical Initiatives, JST-CREST, Faculty of Medicine, Kyushu University, Fukuoka, Japan Search for more papers by this author Akihito Harada Akihito Harada Department of Advanced Medical Initiatives, JST-CREST, Faculty of Medicine, Kyushu University, Fukuoka, Japan Search for more papers by this author Seiji Okada Seiji Okada Department of Advanced Medical Initiatives, JST-CREST, Faculty of Medicine, Kyushu University, Fukuoka, Japan Search for more papers by this author Daijiro Konno Daijiro Konno Laboratory for Cell Asymmetry, Center for Developmental Biology, RIKEN, Kobe, Japan Search for more papers by this author Jun Odawara Jun Odawara Department of Advanced Medical Initiatives, JST-CREST, Faculty of Medicine, Kyushu University, Fukuoka, Japan Search for more papers by this author Tomohiko Yoshimi Tomohiko Yoshimi Department of Bioengineering, Graduate School of Engineering, Osaka City University, Osaka, Japan Search for more papers by this author Saori Yoshimura Saori Yoshimura Department of Bioengineering, Graduate School of Engineering, Osaka City University, Osaka, Japan Search for more papers by this author Hiromi Kumamaru Hiromi Kumamaru Department of Advanced Medical Initiatives, JST-CREST, Faculty of Medicine, Kyushu University, Fukuoka, Japan Search for more papers by this author Hirokazu Saiwai Hirokazu Saiwai Department of Advanced Medical Initiatives, JST-CREST, Faculty of Medicine, Kyushu University, Fukuoka, Japan Search for more papers by this author Toshiaki Tsubota Toshiaki Tsubota Experimental Research Center for Infectious Diseases, Institute for Virus Research, Kyoto University, Kyoto, Japan Search for more papers by this author Hitoshi Kurumizaka Hitoshi Kurumizaka Laboratory of Structural Biology, Graduate School of Advanced Science and Engineering, Waseda University, Tokyo, Japan Search for more papers by this author Koichi Akashi Koichi Akashi Department of Medicine and Biosystemic Science, Faculty of Medicine, Kyushu University, Fukuoka, Japan Search for more papers by this author Taro Tachibana Taro Tachibana Department of Bioengineering, Graduate School of Engineering, Osaka City University, Osaka, Japan Search for more papers by this author Anthony N Imbalzano Anthony N Imbalzano Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA, USA Search for more papers by this author Yasuyuki Ohkawa Corresponding Author Yasuyuki Ohkawa Department of Advanced Medical Initiatives, JST-CREST, Faculty of Medicine, Kyushu University, Fukuoka, Japan Search for more papers by this author Author Information Akihito Harada1, Seiji Okada1, Daijiro Konno2, Jun Odawara1, Tomohiko Yoshimi3, Saori Yoshimura3, Hiromi Kumamaru1, Hirokazu Saiwai1, Toshiaki Tsubota4, Hitoshi Kurumizaka5, Koichi Akashi6, Taro Tachibana3, Anthony N Imbalzano7 and Yasuyuki Ohkawa 1 1Department of Advanced Medical Initiatives, JST-CREST, Faculty of Medicine, Kyushu University, Fukuoka, Japan 2Laboratory for Cell Asymmetry, Center for Developmental Biology, RIKEN, Kobe, Japan 3Department of Bioengineering, Graduate School of Engineering, Osaka City University, Osaka, Japan 4Experimental Research Center for Infectious Diseases, Institute for Virus Research, Kyoto University, Kyoto, Japan 5Laboratory of Structural Biology, Graduate School of Advanced Science and Engineering, Waseda University, Tokyo, Japan 6Department of Medicine and Biosystemic Science, Faculty of Medicine, Kyushu University, Fukuoka, Japan 7Department of Cell Biology, University of Massachusetts Medical School, Worcester, MA, USA *Corresponding author. Department of Advanced Medical Initiatives, JST-CREST, Faculty of Medicine, Kyushu University, 3-1-1 Maidashi, Fukuoka 812-8582, Japan. Tel.:+81 92 642 6216; Fax:+81 92 642 6099; E-mail: [email protected] The EMBO Journal (2012)31:2994-3007https://doi.org/10.1038/emboj.2012.136 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Cell differentiation is mediated by lineage-determining transcription factors. We show that chromodomain helicase DNA-binding domain 2 (Chd2), a SNF2 chromatin remodelling enzyme family member, interacts with MyoD and myogenic gene regulatory sequences to specifically mark these loci via deposition of the histone variant H3.3 prior to cell differentiation. Directed and genome-wide analysis of endogenous H3.3 incorporation demonstrates that knockdown of Chd2 prevents H3.3 deposition at differentiation-dependent, but not housekeeping, genes and inhibits myogenic gene activation. The data indicate that MyoD determines cell fate and facilitates differentiation-dependent gene expression through Chd2-dependent deposition of H3.3 at myogenic loci prior to differentiation. Introduction The mechanisms by which a lineage-committed but undifferentiated cell maintains the ability to specifically activate the appropriate differentiation programme upon differentiation signalling is poorly understood. Activation of differentiation-specific genes depends on the binding of lineage-determining transcription factors to specific regulatory regions and on the appropriate regulation of chromatin structure. Hence, the future gene expression pattern of the differentiated cell must be present in the chromatin structure of the undifferentiated cell in the form of some sort of marking of the genome. However, how this marking is established and recognized is not clear. To elucidate the mechanism of this marking of the whole genome, extensive study of chromatin structure in relation to cell differentiation has been undertaken. For example, methylation of specific DNA sequences by DNA methyltransferase activity is required for mouse development (Okano et al, 1999). It has also been reported that maintenance of histone modifications in the respective promoters of the HNF-1, HNF-4 and albumin genes through the cell cycle in hepatocytes facilitates expression of these genes (Kouskouti and Talianidis, 2005). Moreover, examination of histone acetylation levels in embryonic stem (ES) cells indicates hyperacetylation of histones H3 and H4 in the undifferentiated state (Meshorer et al, 2006). In fact, gene expression patterns are marked from an early stage for the maintenance of differentiation. Recently, characteristic histone variants have been identified that mark the active and the inactive state (Hake et al, 2006). For example, H3.3 has been found to be enriched with active H3K4me2/3, H3K9Ac and H3K14Ac marks and to be predominantly incorporated in the regulatory regions of transcriptionally active genes (Wirbelauer et al, 2005). In contrast, H3.2 is enriched with repressive H3K27me2/3 and H3K9me2 marks (Hake et al, 2006; Garcia et al, 2007). Therefore, exchange of histone variants is involved in the appropriate switching on and off of genes. In mouse ES cells, H3.3 is found at many developmental regulatory genes that are ‘bivalent genes’, marked with transcription-repressing H3K27me and transcription-activating H3K4me3 (Goldberg et al, 2010). In addition, over-expression of H3.3 results in maintenance of the transcriptionally active pattern of gene expression in specific tissue (Ng and Gurdon, 2008). These findings show that replacement of the histone variant (such as H3.3) contributes to the determination of selective gene expression, likely before histone modification (Hake and Allis, 2006). Induction of transcriptional factors is also a method of controlling gene expression. For example, the myogenic transcription factor MyoD induces myogenic differentiation and can even promote reprogramming from a non-muscle cell to a muscle cell (Davis et al, 1987). A similar phenomenon is observed by introduction of specific lineage-determining regulators such as PPARγ2, or the four transcription factors that regulate formation of the induced pluripotent stem cell (Tontonoz et al, 1994; Takahashi and Yamanaka, 2006). Therefore, these transcription factors are required for reprogramming and alteration of the genomic state. It is known that these transcription factors regulate chromatin structure; to date, histone modification and chromatin remodelling have been identified as resultant changes, but a relationship between transcription factors and the type of histone variant (such as incorporation of H3.3) has not been recorded. MyoD is expressed in committed but undifferentiated cells, but how MyoD identifies genes for activation during differentiation is unknown. We hypothesized that a chromatin modifying or remodelling enzyme was likely involved. We identified chromodomain helicase DNA-binding domain 2 (Chd2), a member of the SNF2 family of chromatin remodelling enzymes, as a MyoD-interacting protein that facilitates cell fate determination via marking of myogenic genes by incorporation of the variant histone H3.3. Results Chd2 interacts with MyoD It was previously shown that MyoD associates with Brg1, an enzyme of the mammalian SWI/SNF class of ATP-dependent chromatin remodelers, in differentiating muscle cells (Simone et al, 2004; de la Serna et al, 2005). We theorized that chromatin remodelling enzymes also might interact with MyoD in undifferentiated cells. Using monoclonal antibodies we had generated against Brg1, Brm, Chd1, and Chd2 (Ohkawa et al, 2009; Okada et al, 2009; Harada et al, 2010b; Yoshimura et al, 2010), we identified Chd2, but not Brg1 or Brm, as a MyoD co-immunoprecipitation (co-IP) product in C2C12 myoblasts (Figure 1A; Supplementary Figure S1A). The closely related protein Chd1 (60% homology) was not co-immunoprecipitated with MyoD. Reciprocal co-IP confirmed that Chd2 interacted with MyoD (Figure 1A). Additionally a proximity ligation assay (PLA) was used to demonstrate interaction between Chd2 and MyoD. Interactions between Chd2 and MyoD were observed in both myoblasts and differentiated cells (Figure 1B). As a control, we examined interactions between MyoD and Brg1, which as expected, were greatly enhanced in differentiated cells (Figure 1C). Immunocytochemistry revealed that a subset of Chd2 and MyoD, both of which are exclusively nuclear, were co-localized prior to cell differentiation (Supplementary Figure S1B). A cross-correlation analysis (vanSteensel et al, 1996) of confocal images of Chd2 and MyoD provided further support for co-localization in myoblasts as well as in differentiated C2C12 cells, suggesting that the MyoD–Chd2 interaction persists during differentiation (Supplementary Figure S1B). The specificity of the MyoD–Chd2 association was further assessed by examining co-localization between MyoD and Chd1. Whereas 35–40% of the MyoD co-localized with Chd2 in both myoblast and myotube nuclei, only 7–15% of the MyoD co-localized with Chd1 (Supplementary Figure S1C). Chd2 protein levels were not significantly different in myoblasts and in differentiated cells (Figure 1D). Figure 1.Chd2 interacts with MyoD. (A) Reciprocal IPs were performed from C2C12 myoblast extracts using MyoD- and Chd2-specific antibodies or IgG as a control. (B) PLAs indicating interaction of MyoD and Chd2 in both proliferating myoblasts and differentiated cells, in contrast to (C) the differentiation-specific interactions of MyoD and Brg1. Quantification represents the mean of three independent experiments, each of which analysed at least three separate fields±s.d. Scale bars=12.5 μm. (D) Western blot analysis of Chd2 levels in C2C12 cells under growth (G) or differentiation (D) conditions. Download figure Download PowerPoint Chd2 binds to myogenic gene promoters Chromatin IP (ChIP) was used to analyse whether Chd2 is localized at differentiation-dependent myogenic genes. Because Chd2 was identified as a MyoD-interacting protein, we focused on regulatory sequences containing E-boxes. Chd2 interacted with the promoters of numerous myogenic gene loci in undifferentiated as well as differentiated C2C12 cells but not with housekeeping genes such as Gapdh or the inactive Igh enhancer (Figure 2A; Supplementary Figure S1D). To examine whether Chd2 recruitment was dependent on MyoD, we performed ChIP assays in NIH3T3 cells directed to undergo myogenesis by ectopic expression of MyoD (Davis et al, 1987). We observed MyoD-dependent binding of Chd2 specifically at myogenic gene promoters but not at housekeeping or silent gene promoters (Figure 2B). Coincident binding of MyoD at these same myogenic sequences was confirmed (Supplementary Figure S1E). Western blot analysis showed that the expression of MyoD in these cells did not alter Chd2 levels (Figure 2C). In addition, MyoD levels in these cells were not over-expressed relative to MyoD expression in C2C12 cells (Supplementary Figure S1F). Figure 2.Chd2 interacts with MyoD and myogenic gene regulatory sequences. (A) ChIP assays for Chd2 binding at differentiation-dependent and skeletal muscle-specific (Acta1, Myl3, Myog, Cdkn1a, Ank1, Dmd), housekeeping (Gapdh, Ef1alpha), and silent (IgH enhancer, Pdx1, Neurod6) gene promoters were performed in C2C12 cells under growth and differentiated conditions. Relative recruitment was defined as the ratio of amplification of the PCR product relative to 1% of input genomic DNA. Values obtained from Acta1 at the growth stage were defined as 1 and all other values were expressed relative to that value. Each value was standardized by the amplification efficiency of each primer pair. Quantification represents the mean of three independent experiments±s.d. (B) Ectopic expression of MyoD induces Chd2 recruitment onto the promoter regions of myogenic genes. ChIP assays were performed as in (A) in fibroblast cells expressing MyoD or empty vector that were subjected to the differentiation protocol. (C) Western blot analysis for MyoD and Chd2 expression in MyoD-infected fibroblasts. H3 levels were monitored as a control. (D) siRNA-mediated MyoD knockdown inhibits Chd2 recruitment onto the promoter regions of myogenic genes. ChIP assays for Chd2 binding were performed as described in (A) in C2C12 myoblasts treated with either control siRNA or MyoD siRNA. (E) siRNA-mediated knockdown of the endogenous MyoD protein in C2C12 cells. A western blot analysis of C2C12 cells treated with either control siRNA or MyoD siRNA using antibodies against MyoD, Chd2, and H3 is shown. (F) Relative expression of skeletal muscle marker genes was reduced in C2C12 cells treated with MyoD siRNA. The levels of the indicated mRNAs were analysed by Q-PCR. The values in the differentiated cells expressing control siRNA were set to 1. Data represent the average of three independent experiments±s.d. (G) Chd2 and MyoD co-recruitment at the Ckm but not the Gapdh promoter is shown by re-ChIP. Re-ChIP experiments sequentially used antibodies against Chd2 and MyoD, as indicated. Relative recruitment was defined as the ratio of amplification of the PCR product relative to 1% of input genomic DNA. Values obtained from Ckm at the growth stage with 1st IP were defined as 1 and all other values were expressed relative to that value. Each value was standardized by the amplification efficiency of each primer pair. Quantification represents the mean of three independent experiments±s.d. Download figure Download PowerPoint To further demonstrate that Chd2 recruitment is MyoD-dependent, we reduced the expression of MyoD in C2C12 cells by siRNA treatment and observed that Chd2 binding to myogenic genes did not occur (Figure 2D). Western blot analysis confirmed that MyoD protein levels were reduced by the siRNA treatment and that Chd2 protein levels were not affected (Figure 2E). As expected, siRNA-mediated reduction of MyoD also compromised differentiation-dependent myogenic gene activation (Figure 2F). We then performed re-ChIP assays (Ohkawa et al, 2006). In C2C12 myoblasts maintained in growth media, Chd2 was simultaneously present with MyoD on the Ckm promoter but not on the Gapdh locus (Figure 2G). In differentiated C2C12 cells, Chd2 and MyoD were both present at the Ckm locus, but to a somewhat lesser extent than in myoblasts (Figure 2G). Collectively, these data strongly suggest that Chd2 is targeted to the Ckm promoter via MyoD and are consistent with results demonstrating widespread MyoD binding to myogenic genes in undifferentiated myoblasts (Cao et al, 2010). Chd2 promotes myogenic gene expression To explore the requirement for Chd2 in myogenesis, we suppressed Chd2 expression by stably introducing two microRNAs (miRNA) that target Chd2 (Chd2miR3139 and Chd2miR5111) in C2C12 cells. We used cells stably transfected with lacZ-targeted miRNA (Chd2WT) as a control. To indirectly monitor miRNA expression, enhanced green fluorescent protein—nuclear localization signal (EGFP–NLS) was expressed co-cistronically with the miRNA (Figure 3A). Analysis of myogenic gene expression in these cells indicated that myosin heavy chain expression, which is a late myogenic marker, was completely suppressed in both Chd2miR3139 and Chd2miR5111-expressing cells (n=70 and 63 GFP-positive cells, respectively; Figure 3A). The expression of myogenin, which is a marker of the early phase of myogenesis, was decreased in both cell lines to 6% (Chd2miR3139, n=109 GFP-positive cells) and 13% of wild-type (WT) (Chd2miR5111, n=76 GFP-positive cells; Figure 3A). Moreover, myotube formation was not observed in either miRNA-expressing cell line upon differentiation. Compared with controls, the mRNA expression levels of every differentiation-dependent myogenic gene tested was significantly repressed in Chd2miR3139 and Chd2miR5111-expressing cells (Figure 3B). In contrast, expression of the housekeeping gene, Gapdh, was unaffected (Figure 3B). Figure 3.Chd2 is required for skeletal muscle differentiation. (A) The expression of myogenin (Myog) and myosin heavy chain (MHC) was not induced in C2C12 cells when Chd2 expression was suppressed by miRNAs targeting Chd2. To indirectly monitor miRNA expression, EGFP–NLS (EGFP fused with a nuclear transport signal) was expressed co-cistronically with miRNA. Scale bars=5 μm. (B) The transcription of skeletal muscle marker genes was suppressed in C2C12 cells expressing miRNA targeting Chd2. mRNA levels were analysed by Q-PCR; data represent the average of three independent experiments±s.d. Gapdh levels are shown as a control. (C) Western blot evaluating Chd2 protein knockdown and the expression of MyoD and other indicated proteins. Download figure Download PowerPoint Control experiments determined that Chd2 transcript levels were not affected in cells expressing the Chd2-targeting miRNAs (Supplementary Figure S2A), but Chd2 protein expression was repressed (Figure 3C). This suggests that the specific miRNAs functioned as translational repressors of Chd2. The GFP expression level remained consistent, suggesting no significant differences in miRNA expression between the cells (Figure 3C). In addition, no significant differences in the expression of MyoD were observed between Chd2WT and miRNA-expressing cells, indicating that Chd2 was not regulating the expression of MyoD (Figure 3C). To confirm that changes in MyoD levels observed during differentiation did not alter Chd2 expression, we ectopically expressed MyoD in the Chd2 miRNA-expressing cells and showed that Chd2 expression (Figure 3C) and differentiation-dependent gene expression (Supplementary Figure S2B) were not rescued. We also determined that cell-cycle progression was not affected by miRNA expression in undifferentiated or differentiated cells as measured by FACS analysis (Supplementary Figure S2C) and western blot analysis of cyclins A and E (Supplementary Figure S2D). These data indicate that Chd2 is not indirectly affecting myogenic gene expression via alteration of cell-cycle arrest. To complement these studies showing a requirement for Chd2 in myogenic differentiation, we reduced Chd2 expression by introducing siRNA molecules that target Chd2. siRNA-treated cells did not form myotubes as demonstrated by MHC staining (Supplementary Figure S3A) and were compromised for differentiation-specific gene expression (Supplementary Figure S3B). Western analysis demonstrated the reduction in Chd2 levels in siRNA-treated cells and no effect on MyoD levels (Supplementary Figure S3C). To further confirm a Chd2-specific function in myogenic gene induction, we rescued the inhibition of Chd2 expression by miRNA via the exogenous introduction of competitive mRNA fragments (Chd2-3011-3283 or Chd2-5004-5177) that were linked to the monomeric Kusabira Orange (mKO1) fluorescent protein containing a nuclear localization signal (Karasawa et al, 2004). In differentiated cells expressing Chd2miR3139, introduction of a competitive mRNA (Chd2-3011-3283) but not an mRNA from a different region of Chd2 (Chd2-5004-5177), restored MHC expression (Figure 4A). Similarly, introduction of the mRNA (Chd2-5004-5177) rescued MHC expression in Chd2miR5111-treated cells whereas introduction of the (Chd2-3011-3283) mRNA did not (Figure 4A). mKO1 expression levels were monitored to confirm equal expression of the competitive mRNAs in the cells (Figure 4A). Furthermore, under conditions where MHC expression was restored, we observed restoration of Chd2 protein levels (Figure 4B) and expression of Myog, Ckm, Des, and Chrna1 at levels comparable to WT (Figure 4C). Figure 4.Myogenic phenotype is rescued by a forced expression of Chd2 partial mRNA in Chd2 knockdown cells. (A) The expression of myosin heavy chain (MHC; blue) was re-induced in C2C12 cells expressing miRNAs targeting Chd2 and a Chd2 partial mRNA that competes each miRNA. To indirectly monitor miRNA and Chd2 partial mRNA expression, EGFP–NLS (EGFP fused with a nuclear transport signal) and mKO1–NLS (mKO1 fused with a nuclear transport signal) were expressed co-cistronically with the miRNA and Chd2 partial mRNA, respectively. Scale bars=5 μm. Q-PCR analysis confirms equivalent expression of GFP and mKO1 in each sample (lower panel). (B) Chd2 expression was rescued in C2C12 cells expressing miRNAs targeting Chd2 and the competing Chd2 partial mRNA. Western blot analysis utilized antibodies against Chd2 and α-tubulin. (C) The transcription of skeletal muscle marker genes was rescued in C2C12 cells expressing miRNAs targeting Chd2 and the competing Chd2 partial mRNA. The levels of the indicated mRNAs were analysed by Q-PCR; data represent the average of three independent experiments±s.d. Download figure Download PowerPoint As a complement to this set of experiments, we attempted to rescue the miRNA-mediated inhibition of Chd2 expression and the inhibition of myogenesis via the exogenous introduction of full-length Chd2 cDNA or a Chd2 deletion mutant (Chd2-chromodomain deletion Δ-281–512 aa). Chromodomains facilitate interaction of proteins with chromatin via interaction with methylated histones (Pray-Grant et al, 2005). The constructs utilized were Flag-tagged and co-expressed the monomeric Kusabira Orange 2 (mKO2) fluorescent protein (Karasawa et al, 2004; Sakaue-Sawano et al, 2008). In differentiated cells expressing either Chd2miR3139 or Chd2miR5111, introduction of full-length Chd2 restored MHC expression and myotube formation, whereas expression of the Chd2 mutant lacking the chromodomain did not (Supplementary Figure S4A). Similarly, introduction of the full-length, but not the mutant Chd2, restored expression of Acta1, Myog, Ckm, and Myh4 to levels comparable to those expressed in the control cells (Supplementary Figure S4B). Under conditions where myogenic gene expression and differentiation were restored, we observed restoration of Chd2 protein levels (Supplementary Figure S4C). Chd2 interacts with histone H3.3 Since we observed that Chd2 was present at differentiation-dependent genes in proliferating myoblasts, we wished to address whether Chd2 might facilitate myogenic gene activation prior to the onset of differentiation and myogenic gene expression. CHD1, a related member of the CHD family, incorporates H3.3 into the nucleosome (Konev et al, 2007). H3.3 is a variant of H3 that is incorporated at transcriptionally activated genes (Ahmad and Henikoff, 2002; Jin et al, 2009). To evaluate whether Chd2 interacts with endogenous H3.3, we generated monoclonal antibodies to specifically distinguish H3.3 from H3.1, the major H3 isoform. H3.3 differs from H3.1 at only five residues, only one of which is in the N-terminal tail (position 31). Peptides spanning amino acids 21–39 were used as antigen. The specificity of each antibody was demonstrated by specific recognition of the appropriate recombinant H3.3 or H3.1 protein (Figure 5A). Specificity of the H3.3 antibody was further demonstrated by its specific recognition of modified and unmodified H3.3 and H3.1 peptides (Supplementary Figure S5A). Epitope mapping with recombinant H3.3 and H3.1 proteins revealed that the H3.3 antibody specifically recognized the unique S31 residue present in H3.3 (Supplementary Figure S5B). Since H3.3 is primarily associated with active transcription, we performed immunostaining to gauge the extent to which the H3.3 antibody recognizes euchromatin and heterochromatin, which is marked by intense Hoechst staining. The data indicate that the H3.3 antibody primarily recognizes euchromatic regions of the genome (Supplementary Figure S5C). This result is consistent with previous work showing that exogenous H3.3 is predominantly incorporated into active gene loci (Ahmad and Henikoff, 2002; Jin et al, 2009). Previously, H3.3 has been found to be enriched with active H3K4me2/3 marks, but not H3K9me2 marks (Wirbelauer et al, 2005; Hake et al, 2006; Garcia et al, 2007). Use of the H3.3 antibody in co-localization analyses with either H3K4me3 or H3K9me3 demonstrated significant co-localization with H3K4me3 but not H3K9me3 (Supplementary Figure S5D). Collectively, the data indicate that the H3.3 antibody specifically recognizes endogenous H3.3. We also determined that neither the overall nor the chromatin-associated levels of H3.3 changed as a function of C2C12 cell differentiation (Supplementary Figure S5E). Subsequent experiments showed that Chd2 could co-immunoprecipitate with H3.3, but not H3.1, in undifferentiated cells (Figure 5B). PLA assays also indicated Chd2–H3.3 interactions in myoblasts as well as differentiated cells, whereas the frequency of Chd2–H3.1 interactions was much lower (Figure 5C). Immuno-localization studies indicated Chd2–H3.3 co-localization in both undifferentiated and differentiated cells (Figure 5D). We further analysed the Chd2 and H3.3 interaction by co-IP using Chd2mir3139 cells that exogenously express either Flag-tagged full-length Chd2 or the chromodomain deleted Chd2 mutant. Flag-tagged full-length Chd2 was immunoprecipitated with endogenous H3.3, while the Chd2 mutant was not (Figure 5E). Collectively, these studies suggest a possible link between Chd2 function and H3.3. Figure 5.Chd2 interacts with H3.3 prior to and during skeletal muscle differentiation. (A) H3.3 antibody specifically discriminates between H3.3 and H3.1. Serial dilutions of purified recombinant H3.1 and H3.3 protein were evaluated by immunoblotting u
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