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Molecular and Cellular Biology, January 2007, p. 518-525, Vol. 27, No. 2
0270-7306/07/$08.00+0     doi:10.1128/MCB.01415-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Regulation of HDAC9 Gene Expression by MEF2 Establishes a Negative-Feedback Loop in the Transcriptional Circuitry of Muscle Differentiation{triangledown}

Michael Haberland, Michael A. Arnold, John McAnally, Dillon Phan, Yuri Kim, and Eric N. Olson*

Department of Molecular Biology, University of Texas Southwestern Medical Center, 6000 Harry Hines Boulevard, Dallas, Texas 75390

Received 1 August 2006/ Returned for modification 29 August 2006/ Accepted 27 October 2006


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Skeletal muscle development is controlled by the myocyte enhancer factor (MEF2) and myogenic basic helix-loop-helix (bHLH) families of transcription factors, which associate and synergistically activate muscle gene expression. Muscle differentiation is further reinforced by positive-feedback loops in which myogenic bHLH proteins activate their own expression and the expression of MEF2, while MEF2 stimulates expression of myogenic bHLH genes and the Mef2c gene. Here we describe a myogenic negative-feedback loop that consists of MEF2 proteins and the transcriptional repressor histone deacetylase 9 (HDAC9). We show that the HDAC9 gene is a direct transcriptional target of MEF2 in vitro and in vivo. HDAC9 can associate with MEF2 proteins and suppress their transcriptional activity. The transcriptional repressor HDAC9 thus forms a negative-feedback loop in the transcriptional circuitry of muscle differentiation.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Muscle differentiation is under the control of two families of transcription factors, the myogenic basic helix-loop-helix (bHLH) proteins and the myocyte enhancer factor 2 (MEF2) family of MADS (MCM1, Agamous, Deficiens, serum response factor) domain-containing proteins (5, 40). Myogenic bHLH proteins recognize a DNA sequence called the E box (CANNTG), which is present in most muscle-specific promoters. Myogenic bHLH proteins dimerize with ubiquitous bHLH proteins known as E proteins and associate and synergistically activate transcription with MEF2 proteins, which bind the A/T-rich DNA consensus YTAWWWWTAR.

The essential role of MEF2 in muscle development was first demonstrated for Drosophila melanogaster, where disruption of the single MEF2 gene D-mef2 results in a block of differentiation of all muscle lineages (8, 24). Consistent with these findings, forced expression of a dominant-negative MEF2 mutant prevents differentiation of mammalian myoblasts in vitro (39).

The existence of four Mef2 genes (Mef2a, Mef2b, Mef2c, and Mef2d) in vertebrates complicates the analysis of their individual functions due to their partial redundancy and overlapping expression patterns (5). Mice with a disrupted Mef2c allele die around embryonic day 9.5 (E9.5) due to improper development of the heart (26) and vascular system (25), whereas Mef2a mutant mice develop mitochondrial insufficiency in cardiomyocytes postnatally and die from sudden cardiac death due to conduction defects (38). MEF2 has also been implicated in the remodeling of the adult heart in response to a variety of stress signals, in control of energy metabolism in striated muscle, and in maintenance of the slow-fiber phenotype of skeletal muscle (5, 34).

An additional layer of complexity in the transcriptional circuitry of muscle differentiation is introduced by multiple feedback mechanisms. Myogenic bHLH proteins activate their own expression as well as the expression of MEF2, while MEF2 stimulates expression of myogenic bHLH protein genes and of the Mef2c gene (12, 13, 15, 16, 48, 49). These auto- and cross-regulatory interactions thereby establish a mutually reinforcing circuit to induce and maintain the skeletal muscle phenotype (6, 42).

The transcriptional activity of MEF2 factors is influenced by a variety of intracellular signaling pathways and by interaction with an array of coactivators and corepressors. Class II histone deacetylases (HDACs) have recently emerged as central regulators of MEF2 activity and the downstream programs of gene expression they control (11, 47, 50, 53). The four class II HDACs, HDAC4, -5, -7, and -9, interact with the MADS domain of MEF2 through an N-terminal peptide sequence and prevent activation of MEF2 target genes by establishing corepressor complexes (11, 27, 28, 30, 32). Signaling from G protein-coupled receptors leads to the phosphorylation of class II HDACs via several calcium-dependent protein kinases, resulting in dissociation from MEF2 and nuclear export (2, 9, 29, 41). The histone acetyltransferase (HAT) p300/CBP docks on the same domain of MEF2 as class II HDACs, thereby providing a binary switch-type mechanism in which MEF2 target genes can be activated or repressed by the mutually exclusive interaction of MEF2 with p300/CBP or class II HDACs, respectively (27, 32).

The MEF2 family of transcription factors, especially MEF2C and MEF2D, are highly enriched in skeletal and cardiac muscle as well as in neuronal tissues (5, 17). A subset of the class II HDACs (HDAC5 and HDAC9) show similar expression patterns with HDAC9 showing the highest expression in adult skeletal and cardiac muscle (10, 52). As MEF2 and class II HDACs interact in an intricate regulatory network via protein-protein interactions (9, 30, 31, 33), these overlapping expression patterns prompted us to investigate if this interaction could also be observed to occur at the transcriptional level.

Here we describe a transcriptional negative-feedback mechanism at the center of the myogenic gene program. We identified the transcriptional repressor HDAC9 as being directly controlled by MEF2 in vitro and in vivo. HDAC9 associates with MEF2 proteins and suppresses their transcriptional activity. These findings reveal a molecular basis for robustness of the muscle phenotype and fine-tuning of muscle gene expression through the regulated expression and activity of HDAC9, a key negative regulator of the muscle gene program.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Plasmids. Promoter fragments were generated by PCR amplification of genomic DNA from tail biopsy specimens of C57BL/6 mice. A 921-bp promoter fragment (corresponding to chromosome 12 positions 35,114,099 to 35,115,019 in the UCSC genome browser, February 2006 assembly [21]) containing 23 bp of the HDAC9 5' untranslated region was used in these studies. Promoter mutants were generated through PCR mutagenesis. The MEF2 site was mutated (underlined residues) from the wild-type sequence ACCTATTTTTGGCT to the mutant sequence ACCTACCGGTGGCTC. PCR products were cloned into PCR2.1Topo (Invitrogen) and then subcloned into pGL4.10 luciferase reporter constructs (Promega) and the hsp68-lacZ vector (15, 43). Expression constructs for MEF2, HDAC9/MITR, HDAC9{Delta}MEF, MyoD, and E12 have been described previously (28, 35, 43, 51, 52). All plasmids were verified by sequencing.

Generation of transgenic and knockout mice. Reporter constructs containing the 921-bp HDAC9 promoter fused to hsp68-lacZ were digested with SalI to remove the vector backbone, gel purified, and injected into fertilized eggs from B6C3F1 female mice. Injected eggs were implanted into pseudopregnant ICR mice as previously described (43). Embryos were collected and stained for ß-galactosidase activity as described previously (37). Mef2c-null animals have been described previously (25, 26). Mef2c conditional knockout mice were generated by flanking exon 2 of the coding region with loxP sites (to be described in detail elsewhere) and crossing with myogenin-Cre deleter mice (23). Mef2d knockout mice were generated by replacing exon 2 with a neomycin resistance cassette (to be described in detail elsewhere). Correct targeting and germ line transmission were confirmed for all animals by Southern blotting, genomic sequencing, and reverse transcriptase PCR (RT-PCR). For the analysis of HDAC9 expression, hind limb muscle was isolated from 4-week-old animals on a mixed C57BL/6/SV129 background. All animal experiments were conducted according to procedures approved by the Institutional Animal Care and Use Committee at the University of Texas Southwestern Medical Center.

Gel shift assays. Oligonucleotides corresponding to the conserved MEF2-binding site in the HDAC9 promoter, a mutated site, and a bona fide MEF2 site from the MCK enhancer (45) were synthesized (Integrated DNA Technologies), annealed, radiolabeled with [32P]dCTP using the Klenow fragment, and purified with G50 columns (Roche). The following sequences were used: for the HDAC9 wild type, TGCTTGACCTATTTTTGGCTCAGGC; for the HDAC9 mutant (mutated residues are underlined), TGCTTGACCTACCGGTGGCTCAGGC; and for the muscle creatine kinase (MCK) MEF2 site, GATCGCTCTAAAAATAACCCTGTCG. Cell extracts were isolated from 293T cells that were transfected with either a MEF2C expression vector or a control plasmid. Reaction conditions for gel mobility shift assays were as previously described (43). DNA-protein complexes were resolved on 5% polyacrylamide native gels and visualized by phosphorimaging.

RNA analysis. Total RNA was isolated from cells or tissues with TRIzol (Invitrogen). For semiquantitative PCR analysis, a OneStep RT-PCR (Invitrogen) was used. Quantitative analysis was performed by real-time PCR using TaqMan OneStep chemistry on an ABI PRISM 7000 sequence detection system (Applied Biosystems). Predesigned intron-spanning primers were purchased from Applied Biosystems. The relative quantities of mRNA were determined using the comparative cycle threshold method as described previously (14).

Cell culture, transfection, and luciferase assays. C2C12 cells (7) were cultured in Dulbecco's modified Eagle's medium with 10% fetal bovine serum and penicillin-streptomycin. For myogenic differentiation, cells were shifted to medium containing 2% horse serum and penicillin-streptomycin. 293T cells were maintained in Dulbecco's modified Eagle's medium with 10% fetal bovine serum and penicillin-streptomycin. Transfections were performed using Gene Jammer (Stratagene) according to the manufacturer's instructions. To control for transfection efficiency, a ß-galactosidase expression plasmid was cotransfected. Forty-eight hours after transfection, cells were lysed and assayed for luciferase expression by use of a luciferase assay kit (Promega). Promoter activities are expressed as relative luminescence units normalized for ß-galactosidase expression in the cell extracts.

ChIP assay. The protocol supplied by Upstate was used for chromatin immunoprecipitation (ChIP) using salmon sperm-blocked protein A agarose (Upstate) and the recommended buffers. Precipitated DNA was used for PCR assays with primers either spanning the MEF2 site in the HDAC9 promoter or spanning a fragment of similar size in the GAPDH (glyceraldehyde-3-phosphate dehydrogenase) promoter. For quantitative ChIP, the PCR was performed using SYBR green on an ABI PRISM 7000 sequence detection system. Levels of enrichment (n-fold) between specific antibody and unspecific antibody were calculated using the comparative cycle threshold method. MEF2 antibody (C-21) was purchased from Santa Cruz. This antibody recognizes MEF2A as well as MEF2C and MEF2D. Primer sequences are available on request.

Statistical analysis. Results are expressed as means ± standard deviations. Differences between groups were tested for statistical significance using the unpaired two-tailed Student t test with Welch correction. P values of less than 0.05 were considered significant.

Nucleotide sequence accession number. The GenBank accession number for the HDAC9 promoter fragment used in this study is EF026095.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
The HDAC9 promoter contains an evolutionarily conserved MEF2 site. MITR, a splice variant of HDAC9, has been shown to act as a signal-responsive inhibitor of myogenesis via its repressive effect on MEF2 (52). To further explore the role of MITR/HDAC9 in muscle differentiation, we analyzed the promoter of the HDAC9 gene for evolutionarily conserved regulatory sequences and noted a highly conserved A/T-rich element that resembled the consensus sequence for MEF2 binding (Fig. 1A). Gel mobility shift assays with this sequence and extracts from 293T cells transfected with a MEF2C expression plasmid showed that MEF2 could bind this site in vitro (Fig. 1B). Binding of MEF2C to the putative MEF2 element from the HDAC9 promoter was abolished in the presence of an excess of the unlabeled sequence or the bona fide MEF2 site from the MCK enhancer as a competitor (20), whereas a mutant sequence failed to compete for MEF2C binding. MEF2A and MEF2D were also able to bind this site (data not shown).


Figure 1
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  		FIG. 1. The HDAC9 promoter contains an evolutionarily conserved MEF2 site. (A) Structures of the HDAC9 promoter and the putative MEF2 site. A sequence comparison among several species is shown. Nonconserved residues are shaded black. The bona fide MEF2-binding consensus is indicated. (B) Binding of the putative MEF2 site by MEF2. Electrophoretic mobility shift assays were performed with a labeled probe corresponding to the MEF2 site from the HDAC9 promoter and extracts from 293T cells expressing either MEF2C- or mock-transfected cells. Different competitors (comp) or an anti-MEF2 antibody (ab) was added to the reaction mixtures. Unlabeled HDAC9 probe and an unlabeled bona fide MEF2 probe from the MCK enhancer were indistinguishable in their abilities to compete with labeled probes to form the MEF2-DNA complex. Mutation (mut) of the core nucleotides abolishes formation of a specific DNA-protein complex (lane 3) as well as competition against wild-type (wt) labeled probe (lane 6). The DNA-protein complex was supershifted in the presence of anti-MEF2 antibody. (C and D) MEF2 binds the HDAC9 promoter in vivo. ChIP assays were performed with chromatin from MEF2C-transfected 293T cells. Chromatin was immunoprecipitated with mouse IgG or an anti-MEF2 antibody, and precipitated genomic DNA was amplified with primers flanking the MEF2 site in the HDAC9 promoter and primers flanking a fragment of the GAPDH promoter. Quantification of the relative enrichment levels was performed by real-time PCR. (E) 293T cells were transfected with MEF2C and MEF2D, and HDAC9 mRNA levels were quantified by real-time quantitative PCR.

 
To determine if MEF2 could bind the HDAC9 promoter in the context of native chromatin, we performed ChIP using primers flanking the MEF2 site. When 293T cells were transfected with a MEF2C expression plasmid and cross-linked protein-DNA complexes were immunoprecipitated with a MEF2 antibody, we found strong enrichment of HDAC9 promoter fragments compared to results from control immunoprecipitations with either untransfected cells or unspecific antibodies (normal immunoglobulin G [IgG]). Amplification of a control sequence in the GAPDH promoter showed no significant enrichment after precipitation with a MEF2 antibody (Fig. 1C and D), confirming the specificity of the ChIP assays.

When 293T cells were transfected with either a MEF2C or a MEF2D expression plasmid, expression of the endogenous HDAC9 gene was increased three- to fivefold as determined by measuring mRNA levels with quantitative RT-PCR (Fig. 1E). These findings suggested that HDAC9 was a transcriptional target of MEF2, which could potentially create a negative-feedback loop for the modulation of myogenesis.

Regulation of HDAC9 and MEF2 during skeletal muscle differentiation. To further explore the potential regulatory relationship between MEF2 and HDAC9, we compared the expression levels of HDAC9 and MEF2C during differentiation of the C2C12 myoblast cell line (6, 7). Differentiation of C2C12 cells in response to serum withdrawal led to a robust increase in MEF2C mRNA expression (Fig. 2A). HDAC9 mRNA levels increased in parallel up to day 3 of differentiation, after which they were reduced to baseline levels (Fig. 2B).


Figure 2
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  		FIG. 2. Expression of HDAC9 and MEF2 in C2C12 cells. (A and B) C2C12 cells were differentiated by culturing in 2% horse serum, and MEF2C and HDAC9 mRNA levels were quantified by real-time quantitative PCR at the indicated time points. (C) ChIP assays were performed with chromatin isolated from undifferentiated and day 3-differentiated C2C12 cells. Chromatin was immunoprecipitated with mouse IgG or an anti-MEF2 antibody, and precipitated genomic DNA was amplified with primers flanking the MEF2 site in the HDAC9 promoter and primers flanking a fragment of the GAPDH promoter. Quantification of the relative enrichment levels was performed by real-time quantitative PCR as described in Materials and Methods. (D) Expression levels of the MEF2 isoforms in C2C12 cells at the indicated time points were measured by real-time quantitative PCR.

 
ChIP assays using an antibody that recognizes MEF2A, -C, and -D (clone C-21; Santa Cruz) with primers spanning the MEF2 site in the proximal promoter of the HDAC9 gene revealed that MEF2 proteins were bound to this promoter region at day 3 of differentiation, whereas only minimal binding could be detected in undifferentiated cells (Fig. 2C). Quantitative RT-PCR revealed the expression of MEF2A, MEF2C, and MEF2D at day 3 of differentiation (Fig. 2D).

Activation of the HDAC9 promoter by MEF2. We next tested if MEF2 could activate the HDAC9 promoter. A 921-bp fragment containing 23 bp of the 5' untranslated region was cloned upstream of a promoterless luciferase reporter construct and used in cotransfection experiments. When 293T cells were transfected with this construct and increasing amounts of a MEF2D expression plasmid, we observed robust activation of the reporter (Fig. 3A). A similar induction was observed when MEF2C was used in cotransfection assays (data not shown). As myogenic bHLH proteins often show synergistic activation with proteins of the MEF2 family, we also cotransfected increasing amounts of MyoD either alone or with MEF2D. However, we did not observe synergistic activation of the HDAC9 promoter with MEF2D and MyoD. Indeed, when increasing amounts of MyoD were cotransfected with MEF2D, we saw less activation than when MEF2D was transfected alone, an effect that might be attributed to squelching. Mutation of the four central nucleotides (underlined) in the MEF2 site (wild-type sequence ACCTATTTTTGGCT to mutant sequence ACCTACCGGTGGCTC), which was sufficient to abolish MEF2 binding (Fig. 1B), severely blunted the ability of MEF2D to activate the promoter (Fig. 3B).


Figure 3
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  		FIG. 3. Activation of the HDAC9 promoter by MEF2. (A) Responsiveness of the HDAC9 promoter to MEF2. 293T cells were transfected with a pGL4-Luc reporter containing the region from –898 bp to +23 bp of the HDAC9 gene and increasing amounts of MEF2D and MyoD as indicated. (B) The MEF2 responsiveness of the HDAC9 promoter is dependent on the MEF2 site. The pGL4-Luc reporter containing the wild-type (wt) HDAC9 promoter was transfected in 293T cells with increasing amounts of MEF2D as indicated. Mutation of the MEF2 site abolishes the responsiveness of the HDAC9 reporter construct to MEF2D. (C) Inhibition of MEF2 transactivation by HDAC9. Activation of the HDAC9 pGL4-Luc reporter construct in 293T cells is inhibited by cotransfection of HDAC9 but not by cotransfection of an HDAC9 mutant lacking the MEF2-binding domain (HDAC9{Delta}MEF).

 
We next tested if HDAC9 could prevent activation of its own promoter by MEF2. Indeed, coexpression of HDAC9 with MEF2D completely repressed the HDAC9 promoter (Fig. 3C). In contrast, an HDAC9 mutant protein lacking the MEF2 interaction domain (HDAC9{Delta}MEF) was defective in repressing the HDAC9 promoter (Fig. 3C). This HDAC9 mutant lacks the MEF2-binding domain between amino acids 135 and 152 but still can recruit corepressor complexes such as CtBP (35, 51, 52).

MEF2 is necessary to drive HDAC9 expression in vivo. To determine if the MEF2 site in the HDAC9 promoter was necessary for the expression of HDAC9 in vivo, we fused the 921-bp proximal promoter to the hsp68 basal promoter and analyzed transgene expression in F0 mouse embryos. As shown in Fig. 4, the HDAC9-lacZ transgene was expressed in the embryonic heart, dorsal root ganglia, and neural tube at E11.5. At E12.5, the HDAC9 promoter directed expression in heart and skeletal muscle, as well as dorsal root ganglia, neural tube, and retina. Mutation of the MEF2-binding site completely abolished transgene expression in the heart and markedly reduced expression in neuronal tissues (Fig. 4).


Figure 4
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  		FIG. 4. MEF2-dependent transcriptional control of HDAC9 in mouse embryos. (Upper panel) Transgenic mice were generated with the HDAC9-lacZ transgene, as described in Materials and Methods. Robust expression is seen in the heart (h), the dorsal root ganglia (drg), the neural tube (nt), and the brain at E11.5 in F0 transgenic embryos. Mutation of the MEF2 site abolishes expression in the heart and reduces expression in the dorsal root ganglia and the neural tube and brain. At E11.5, 10 of 15 transgenic embryos harboring the wild-type transgene showed lacZ staining similar to that shown, whereas 0 of 9 embryos harboring the transgene with the mutant MEF2 sites showed lacZ expression outside neural tissue. (Lower panel) E12.5 F0 embryos show expression in skeletal muscle (skm) of the body wall, limbs, and face as well as in heart and neuronal tissues. Mutation of the MEF2 site abolishes expression in muscle tissue. At this stage, 9 of 12 transgenic embryos harboring the wild-type transgene showed lacZ staining similar to that shown, whereas 0 of 15 embryos harboring the transgene with the mutant MEF2 sites showed lacZ expression in heart and muscle.

 
Down-regulation of HDAC9 in MEF2 mutant mice. To further test our hypothesis that HDAC9 expression is under the control of MEF2 transcription factors, we analyzed HDAC9 expression in skeletal muscle from MEF2 mutant mice. Mouse embryos homozygous for a Mef2c deletion die around E9 from severe cardiac abnormalities (26). We therefore analyzed mice with conditional deletion of Mef2c in skeletal muscle (to be described in detail elsewhere). This was achieved by breeding mice harboring a conditional Mef2c allele in which exon 2, encoding the MADS and MEF2 domains, was flanked by loxP sites with transgenic mice expressing Cre recombinase under control of the myogenin promoter and Mef2c enhancer (23). MEF2C mRNA in the skeletal muscle of these mice was significantly reduced (Fig. 5A and B). Consistent with the apparent requirement of MEF2 for HDAC9 expression, HDAC9 transcripts were dramatically reduced in skeletal muscle of these mice (Fig. 5C). We also generated Mef2d-null mice (to be described in detail elsewhere) and analyzed HDAC9 expression levels in skeletal muscle. These mice showed no detectable MEF2D transcript in their skeletal muscle (Fig. 5D and E), and HDAC9 mRNA expression was markedly reduced (Fig. 5F).


Figure 5
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  		FIG. 5. Down-regulation of HDAC9 in MEF2 mutants. (A) Skeletal muscle-specific deletion of MEF2C. Conditional MEF2C mice carrying a floxed MEF2C allele were crossed with myogenin-Cre deleter mice. Semiquantitative PCR shows robust reduction of MEF2C mRNA in hind limb muscle. (B) Quantification of the MEF2 isoforms in hind limb skeletal muscle of MEF2Cflox/KO, myogenin-Cre mutant mice by real-time quantitative PCR. (C) Quantification of HDAC9 mRNA in hind limb skeletal muscle of MEF2Cflox/KO, myogenin-Cre mutant mice by real-time quantitative PCR. **, P < 0.005. (D) Knockout of MEF2D. Semiquantitative PCR of hind limb mRNA of mice harboring a homozygous targeted deletion of the MEF2D gene shows the absence of MEF2D mRNA. (E) Quantification of the MEF2 isoforms in hind limb skeletal muscle of MEF2D-null mutant mice by real-time quantitative PCR. (F) Quantification of HDAC9 mRNA in MEF2D-null hind limb skeletal muscle by real-time quantitative PCR. *, P < 0.05. wt, wild type; mut, mutant.

 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
MEF2 transcription factors are key regulators of muscle development. The activity of MEF2 proteins is tightly regulated by class II HDACs, which interact directly with the MADS domain and inhibit MEF2 activity. The results of the present study demonstrate the HDAC9 gene to be a direct transcriptional target of MEF2 factors, thereby identifying a negative-feedback loop at the center of the transcriptional circuitry that drives myogenic differentiation (Fig. 6).


Figure 6
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  		FIG. 6. A negative-feedback loop for the control of muscle differentiation. Myogenic induction leads to transcriptional activation of myogenic bHLH proteins and MEF2 transcription factors. Myogenic bHLH proteins, such as myogenin (MYOG), activate their own expression as well as the expression of MEF2, while MEF2 stimulates expression of myogenic bHLH proteins and of the Mef2c gene. Myogenic bHLH and MEF2 cooperatively activate downstream genes responsible for muscle differentiation (diff.). MEF2 also activates HDAC9, which forms a negative-feedback loop by suppressing the transcriptional activity of MEF2.

 
MEF2 activates the HDAC9 gene. Entry of myoblasts into the differentiation pathway is accompanied by the up-regulation of MEF2 and HDAC9 expression. Consistent with the conclusion that MEF2 regulates HDAC9 expression at the onset of myogenesis, MEF2 is an effective activator of the HDAC9 promoter in vivo and in vitro and can be detected on the promoter within native chromatin by ChIP assay. Therefore, it is seemingly paradoxical that HDAC9 is regulated differently than most MEF2 target genes, such that its expression is diminished during late stages of muscle development when MEF2 levels exceed a threshold. During muscle differentiation in vitro, for example, HDAC9 expression increases initially but then declines to a basal level, while MEF2 levels continue to increase progressively in parallel with other bona fide MEF2 targets, such as Srpk3 (37). Similarly, HDAC9 is strongly expressed in skeletal muscle during embryogenesis but is down-regulated in skeletal muscle postnatally (52). The unique regulation of HDAC9 is further illustrated by the insensitivity of the promoter to MyoD. Whereas MyoD generally synergizes with MEF2 proteins to activate skeletal muscle genes (5, 37), cotransfection of MyoD did not lead to activation of the HDAC9 promoter, either alone or when cotransfected with MEF2 expression constructs. Presumably, other promoter-binding factors yet to be identified contribute to the distinctive pattern of regulation of HDAC9 and its selective sensitivity to MEF2 during initial phases of muscle development.

Regulation of HDAC9 by MEF2 in vivo. The HDAC9 promoter can direct the expression of a lacZ transgene during mouse embryogenesis in a pattern mirroring that of the endogenous gene (52). Beginning at E11.5, the promoter is active in the heart as well as in the dorsal root ganglia, the neural tube, and the brain. The HDAC9 promoter continues to be expressed in these tissues as well as in skeletal muscle at E12.5. Transcriptional activity of the HDAC9 promoter in skeletal and cardiac muscle is MEF2 dependent, as mutations of the MEF2 site in the promoter abolish transgene expression in muscle tissue. We also observed reduced, but residual, lacZ activity in neural tissues when the MEF2 site in the HDAC9 promoter was mutated, indicating that MEF2 enhances but is not absolutely essential for HDAC9 expression in neurons. It will be of interest to identify the factors that control HDAC9 expression in neural tissues and to determine if they interact with MEF2 to establish a neuronal feedback loop.

The down-regulation of HDAC9 expression in mice lacking MEF2C and MEF2D provides strong support for the conclusion that HDAC9 is a direct target gene of MEF2 factors in skeletal muscle. Given that the transcriptional activity of the HDAC9 promoter was not completely abolished in MEF2C and MEF2D mutant mice, it is likely that the four MEF2 isoforms exhibit partial functional redundancy on this promoter. In this regard, MEF2C is up-regulated upon deletion of MEF2D (Fig. 5E), which may account for the decreased reduction of HDAC9 mRNA in MEF2D mutant mice. On the other hand, functional redundancy between class II HDACs might explain why HDAC9 mutant mice do not display an overt muscle phenotype. HDAC9-null mice are viable but display hypersensitivity to MEF2 in the heart and skeletal muscle postnatally (35, 50). In skeletal muscle, HDAC9 appears to repress activity-dependent genes after birth, such that HDAC9-null mice display heightened expression of such genes following denervation. Similarly, following cardiac stress, stress-responsive genes are superactivated in the hearts of HDAC9 mutant mice (10, 50). As HDAC4 and HDAC5 are highly expressed in skeletal muscle, they could take over the function of HDAC9 in the skeletal muscle of HDAC9 mutant animals.

A negative-feedback loop within the transcriptional network for myogenesis. Transcriptional networks commonly contain positive- and negative-feedback loops, which provide robustness and fine-tuning to gene programs (1, 3, 4, 18, 19, 22, 36, 44). Muscle differentiation represents a classic example of the importance of such regulatory loops (5). Myogenic bHLH proteins, such as MyoD, not only activate downstream muscle structural genes but also act on their own promoters to amplify and maintain their own expression. MyoD additionally activates the Mef2c gene; MEF2C then feeds back to up-regulate its own promoter as well as the promoters of some of the myogenic bHLH genes (12, 15, 16, 42, 46, 48). Our results reveal an additional layer of regulatory control within the transcriptional circuit for myogenesis, in which MEF2C activates the HDAC9 gene. HDAC9 can, in turn, associate with MEF2 proteins and repress their transcriptional activity, dampening HDAC9 transcription (Fig. 6). The integration of this negative-feedback loop into the myogenic transcriptional circuitry would be expected to precisely titrate the expression of MEF2 and its corepressor and to stabilize the muscle differentiation program.

Transcriptional repression by class II HDACs is thought to be mediated by their association with DNA-bound MEF2 and the consequent modulation of the chromatin structure of MEF2-dependent promoters (27, 28, 52). This interaction is additionally under the control of promyogenic signals that can disrupt MEF2-HDAC interactions by phosphorylating class II HDACs and thereby promote their nuclear export during muscle differentiation (30). Indeed, expression of a mutated, signal-resistant HDAC9 leads to a complete block of myogenesis (52). These findings suggest that the MEF2-HDAC loop is under the control of a myogenic "rheostat" that modulates muscle differentiation in response to extracellular cues.

Implications for other tissues. In addition to their roles in muscle development, MEF2 and class II HDACs have been implicated in diverse processes of tissue growth and remodeling, including cardiac growth, chondrocyte hypertrophy and bone development, maintenance of vascular integrity, and proliferation and apoptosis of immune cells (10, 11, 35, 41, 47, 50). It will be of interest to determine whether regulatory circuits similar to the MEF2-HDAC9 feedback loop described in the present study are operative in these settings and whether such feedback loops can be manipulated to regulate these cellular processes.


   ACKNOWLEDGMENTS
 
We thank J. Brown for editorial assistance.

M.H. was supported by a grant from the Deutsche Forschungsgemeinschaft (HA 3335/2-1), and work in the laboratory of E.N.O. was supported by grants from the National Institutes of Health, The D. W. Reynolds Clinical Cardiovascular Research Center, and the Robert A. Welch Foundation.


   FOOTNOTES
 
* Corresponding author. Mailing address: Department of Molecular Biology, University of Texas Southwestern Medical Center, 6000 Harry Hines Boulevard, Dallas, TX 75390. Phone: (214) 648-1187. Fax: (214) 648-1196. E-mail: eric.olson{at}utsouthwestern.edu. Back

{triangledown} Published ahead of print on 13 November 2006. Back


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
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Molecular and Cellular Biology, January 2007, p. 518-525, Vol. 27, No. 2
0270-7306/07/$08.00+0     doi:10.1128/MCB.01415-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.



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