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Published as doi: 10.1096/fj.08-115931.
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(The FASEB Journal. 2009;23:99-106.)
© 2009 FASEB

The deacetylase HDAC4 controls myocyte enhancing factor-2-dependent structural gene expression in response to neural activity

Todd J. Cohen*, Tomasa Barrientos*, Zachary C. Hartman{dagger}, Sean M. Garvey{ddagger}, Gregory A. Cox§ and Tso-Pang Yao*,1

* Department of Pharmacology and Cancer Biology and

{dagger} Department of Surgery, Duke University, Durham, North Carolina, USA;

{ddagger} Robert M. Berne Cardiovascular Research Center, University of Virginia, Charlottesville, Virginia, USA; and

§ The Jackson Laboratory, Bar Harbor, Maine, USA

1 Correspondence: Department of Pharmacology and Cancer Biology, Box 3813, Duke University Medical Center, Durham, NC 27710, USA. E-mail: yao00001{at}mc.duke.edu


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Histone deacetylase 4 (HDAC4) binds and inhibits activation of the critical muscle transcription factor myocyte enhancer factor-2 (MEF2). However, the physiological significance of the HDAC4-MEF2 complex in skeletal muscle has not been established. Here we show that in skeletal muscle, HDAC4 is a critical modulator of MEF2-dependent structural and contractile gene expression in response to neural activity. We present evidence that loss of neural input leads to concomitant nuclear accumulation of HDAC4 and transcriptional reduction of MEF2-regulated gene expression. Cell-based assays show that HDAC4 represses structural gene expression via direct binding to AT-rich MEF2 response elements. Notably, using both surgical denervation and the neuromuscular disease amyotrophic lateral sclerosis (ALS) model, we found that elevated levels of HDAC4 are required for efficient repression of MEF2-dependent structural gene expression, indicating a link between the pathological induction of HDAC4 and subsequent MEF2 target gene suppression. Supporting this supposition, we show that ectopic expression of HDAC4 in muscle fibers is sufficient to induce muscle damage in mice. Our study identifies HDAC4 as an activity-dependent regulator of MEF2 function and suggests that activation of HDAC4 in response to chronically reduced neural activity suppresses MEF2-dependent gene expression and contributes to progressive muscle dysfunction observed in neuromuscular diseases.—Cohen, T. J., Barrientos, T., Hartman, Z. C., Garvey, S. M., Cox, G. A., Yao, T.-P. The deacetylase HDAC4 controls myocyte enhancing factor-2 dependent structural gene expression in response to neural activity.


Key Words: neuromuscular disease • remodeling • pathological induction • muscle integrity


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
MYOCYTE ENHANCER FACTOR-2 (MEF2) family transcription factors promote muscle differentiation through specific binding of AT-rich promoter elements [CTA(A/T)4TAG] and subsequent activation of muscle-specific genes (1) . In addition to its well-characterized role in muscle development, MEF2 is also critical for adult muscle function and regulates both fiber-type specification and muscle integrity (2 , 3) . Genetic studies (2 , 4) in both mice and zebrafish have shown that many genes controlling sarcomeric and contractile function are regulated by MEF2. As a consequence, mutations in MEF2 lead to disorganized and nonfunctional sarcomere structure, further highlighting a critical role for MEF2 in muscle integrity and contractile function (2 , 4) . Despite its crucial function, how MEF2 activity is regulated in vivo in mature skeletal muscle has not been fully characterized. One critical factor that regulates MEF2 activity is neural input. Analysis of MEF2-LacZ transgenic mice showed that prolonged motor nerve stimulation or exercise training can activate MEF2 transcriptional activity (5) . In agreement with this observation, the MEF2-dependent expression of slow myosin light chain (MLC-slow) requires proper innervation (6) . These studies reveal a functional link between neural input and MEF2 activity; however, the mechanism by which neural activity regulates MEF2 function is not well understood.

Cell-based studies (7 , 8) have shown that histone deacetylase 4 (HDAC4) binds and inactivates MEF2. Accordingly, HDAC4 and HDAC5 were thought to act as repressors of muscle differentiation by virtue of their inhibitory effect on MEF2. Interestingly, however, HDAC4 was found to be imported rather than exported from the nucleus on C2C12 myotube differentiation (8) . This finding is not consistent with a simple model that HDAC4 represses muscle differentiation. Rather it suggests that HDAC4 plays an active role in mature skeletal muscle. Supporting this hypothesis, we (9) recently found that HDAC4 is highly induced and accumulates in the nuclei of denervated muscle including neuromuscular diseases such as amyotrophic lateral sclerosis (ALS). These findings led us to propose that HDAC4 is a key effector that controls muscle gene transcription in response to neural activity. Indeed, HDAC4 regulates synaptic gene expression in response to neural activity (9) . However, the functional significance of HDAC4 induction and its pathological implications have not been fully established. Given that HDAC4 activity can be pharmacologically inhibited, elucidating a role for HDAC4 in muscle dysfunction could provide potential therapeutic opportunity for these devastating diseases.

In this study, we identified a subset of muscle contractile and structural genes as transcriptional targets of HDAC4. We show that HDAC4-dependent repression of muscle gene expression occurs in both cultured myotubes and skeletal muscle in vivo. Using surgical denervation and mouse models of ALS, in which HDAC4 levels are dramatically induced, we show that HDAC4 is responsible for the activity-dependent repression of MEF2-regulated structural gene expression, underscoring the pathological relevance of HDAC4 activity in neuromuscular disease. Our results provide evidence that abnormal induction of HDAC4 could contribute to defects in skeletal muscle function and integrity associated with neuromuscular disease.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Cell culture/plasmids/antibodies
C2C12 cells were cultured as described previously (8) . Cells were maintained in 20% FBS/DMEM and differentiated at 90% confluency in 2% horse serum (Hyclone, Urban, UT, USA) for the indicated time points. Human HDAC4 wild-type and 3SA mutant were used throughout this study. HDAC4-3SA corresponds to S246/467/632A mutations, previously shown to retain HDAC4 in the nucleus and cause enhanced repressive activity toward MEF2-dependent transcription (7 , 8) . HDAC4wt and HDAC4-3SA DNA fragments containing 5' ecoRI and 3' SmaI sites were cloned into the RI/SmaI site on the vector green fluorescent protein (GFP) -expressing plasmid, pEGFP-C1 to generate N-terminal GFP-tagged HDAC4wt and HDAC4-3SA mutant. HDAC4 polyclonal antibody (clone 186) was previously described (8) . The following muscle specific antibodies were used: dystrophin (DYS1), β,{gamma}-sarcoglycan, and desmin (Novocastra, Newcastle upon Tyne, UK); monoclonal myogenin antibody (Sigma, St. Louis, MO, USA); monoclonal caveolin-3 antibody (BD Biosciences, San Jose, CA, USA); and myotilin antibody (a kind gift from Mike Hauser, Duke University, Durham, NC, USA; ref. 10 ).

Gene expression analysis and preparation
C2C12 cells stably expressing either control vector or HDAC4 by retroviral-gene transfer were grown to 90% confluency and differentiated in 2% horse serum for 4 days, and RNA was extracted using Tri Reagent (Molecular Reagents Center, Cincinnati, OH, USA). RNA was further DNase treated using DNA-free kit (Ambion, Austin, TX, USA). After purification, the quality of the RNA samples was confirmed and the miroarrays were hybridized at the Duke University Microarray Core. Array data sets were deposited at the National Center for Biotechnology Information’s Gene Omnibus Express in a MIAME-compliant form along with complete details of all procedures as accession GSE and GSE using the platform GPL3223, which corresponds to the Operon Mouse v.3.0 oligo spotted array.

RNA expression/statistical analysis
Expression data from the 8 total-oligo data sets were imported in Genespring 6.2 (Agilent, Santa Clara, CA, USA), where intensity-dependant normalizations were applied and genes were quality filtered by flag calls. To ascertain differentially expressed genes, one-way parametric ANOVA tests were performed using a P value of 0.01. Genes were further filtered by fold change from controls. Statistically significant overrepresentation in Gene Ontology categories of significantly up-regulated or repressed genes was checked using the Database for Annotation, Visualization and Integrated Discovery (http://david.abcc.ncifcrf.gov/) using previously described methods (11) .

Chromatin immunoprecipitation (ChIP) assay
ChIP assay was performed on C2C12 myotubes as described previously (12) . C2C12 myotubes were crosslinked in 1.42% formaldehyde for 15 min at room temperature. Glycine (125 mM) was added to quench formaldehyde. Cells were scraped, centrifuged, and washed twice with cold PBS. Cells were lysed with IP buffer (150 mM NaCl; 50 mM Tris, pH 7.5; 5 mM EDTA; 0.5% Nonidet P-40; and 1% Triton) by being resuspended several times and centrifuged 1 min high speed. Nuclear pellet was washed once with IP buffer, and chromatin was sheared by sonication (30 s on, 30 s on ice, 7 rounds) using Branson Sonifier 100 (Branson, Danbury, CT, USA). Samples were divided into fourths and immunoprecipitated overnight using 2 µg immunoglobulin G (IgG), HDAC4, or MEF2 antibodies (Santa Cruz Biotechnology, Santa Cruz, CA, USA). Beads were washed 5 to 6 times with cold IP buffer and then eluted with elution buffer (1% SDS, 0.1 M NaHCO3) at room temp for 15 min. Crosslinks were reversed by addition of 0.25 M NaCl at 65°C for 4 h. Samples were proteinase K treated at 45°C for 1 h, and DNA was recovered by phenol/chloroform extraction and ethanol precipitation. One microliter of DNA was used in a polymerase chain reaction (PCR) reaction to amplify the {gamma}-sarcoglycan MEF2 binding region. Primers were designed from –647 to –914 spanning the MEF2 binding sites in {gamma}-sarcoglycan promoter: forward, GAGTTGTCACAGTGGACA and reverse, GGCATGAATTAAGATGGAC. The PCR program was used at 30 cycles: 95°C 2 min (1x), 95°C 30 s, 55°C 30 s, 72°C 30 s (30x), and 72°C 5 min (1x).

Luciferase assay
Luciferase assay was performed as described previously (8) . Briefly, C2C12 cells were plated at 20,000 cells/well in a 24-well plate. At 50% confluency, cells were transfected with Fugene reagent (200 ng reporter, 50 ng MEF2D, 100 ng HDAC4, 100 ng β-gal, 3 µl Fugene, and 98 µl serum-free medium per well). Cells were transfected for 36–48 h and lysed with luciferase lysis buffer (50% glycerol, 2.5% Triton X-100, 3 g Tris base, and 0.695 g CDTA per 200 ml, pH 7.8) for 20 min at room temperature. Ten microliters of cell lysate was used with 200 µl luciferase assay buffer [15 mM MgSO4, 15 mM K2HPO4, 4 mM EGTA, 1 mM dithiothreitol (DTT), and 1 mM ATP], and samples were read on Lumat LB 9507 luminometer (Berthold, Bad Wildbad, Germany). Twenty microliters of lysate was also used in β-gal assay [10 mg ortho- nitrophenyl-β-galactoside (ONPG) and 17 µl β-mercaptoethanol per 5 ml Z-buffer] to determine transfection efficiency. β-Gal results were read on a plate reader at a 405 nm wavelength. All samples were analyzed in triplicate. {gamma}-Sarcoglycan wild-type reporter was kindly provided by Satoru Noguchi (National Institute of Neuroscience, Tokyo, Japan; ref. 13 ). Site-directed mutagenesis was performed (Stratagene, La Jolla, CA, USA) to delete the AT-rich elements at –858 and –800 upstream of {gamma}-sarcoglycan transcription start generating {Delta}AT1, {Delta}AT2, and {Delta}AT1/{Delta}AT2 double mutant constructs. The primers used for the mutagenesis were as follows (5'->3'): {Delta}AT1: GTTGAAACTGCCCTGGGGCCTGCAGCTGTTCC and GGAACAGCTGCAGGCCCCAGGGCAGTTTCAAC; and {Delta}AT2: GAAAGAGTTCTAAGTGCCAAGATTAGTTGAAACTGC and GCAGTTTCAACTAATCTTGGCACTTAGAACTCTTTC. Proper deletion of MEF2 elements was confirmed with the sequencing primer CCATCAGTCTGGAAGGTCAAGG. Myotilin reporters were kindly provided by Olli Carpen (University of Helsinki, Helsinki, Finland; ref. 14 ).

Mouse procedures
Mice were handled and electroporated as described previously (9) . The 6-wk-old C57/BL mice (Jackson Laboratory) were anesthetized with a ketamine/xylazine mixture (25 mg/ml ketamine and 1 mg/ml xylazine in 0.9% NaCl; 100 µl/mouse). Hair was removed from area surrounding muscle. For overexpression electroporation analysis, 25 µg GFP, HDAC4-GFP, or HDAC4-3SA-GFP was used for injection, with 5 µg of GFP-alone plasmid included in HDAC4 electroporated muscle to allow positive identification of electroporated fibers. Stealth siRNA duplexed oligos were from Invitrogen (Carlsbad, CA, USA). HDAC4 siRNA sequences were as follows: mouse HDAC4: 5'-CACCGGAACCUGAACCACUGCAUUU-3'. DNA and siRNA were directly injected into tibialis anterior muscles using cemented MicroSyringe (VWR, West Chester, PA, USA). ECM 830 electroporator (BTX, Hawthorne, NY, USA) was used for all electroporations. Tweezertrodes (model 520; BTX) were coated with transmission gel and were placed outside the skin around the muscle belly and pulsed 5 times at 50 V, 60 ms duration, with a 200 ms interval time. Mice were allowed to recover in their cages. All mice were housed at the Duke University mouse facilities in accordance with the Institutional Animal Care and Use Committee.

Western blot analysis
Western blotting was carried out as described previously (9) . For cell culture, C2C12 myotubes were harvested at indicated time points and lysed in 150 mM NETN buffer (0.5% Nonidet P-40; 20 mM Tris, pH 8; 1 mM EDTA; and 150 mM NaCl) for 30 min at 4°C and then centrifuged at high speed for 15 min. Protein concentrations were determined using Bradford assay. 50 µg total protein lysate was used per well. For mouse muscles, tibialis anterior muscles were isolated and frozen in liquid nitrogen. Muscles were dounce homogenized on ice using a 1 ml glass-on-glass homogenizers and grinded in RIPA buffer (0.05 M NaCl and 0.02 M Tris, pH 7.6, and 1 mM EDTA, supplemented with leupeptin, aprotinin, PMSF, NaF, sodium orthovanadate, and 1 mM DTT). Lysates were incubated with 1x detergent buffer (1% Nonidet P-40, 0.5% sodium deoxycholate, and 0.1% SDS) for 10 min rocking at 4°C and centrifuged at 14,000 rpm for 10 min to pellet debris. Protein was quantitated using Bradford assay (Bio-Rad, Hercules, CA, USA). Fifty micorgrams of protein lysate was used for Western blot analysis using the antibodies described above.

RNA analysis
For RNA analysis, GFP-positive fibers were isolated using a dissecting microscope. Muscle tissue was dounce homogenized in 1 ml Tri reagent (MRC, Cincinnati, OH, USA) and incubated at 4°C overnight. Two hundred microliters of chloroform was added, and after vigorous shaking, samples were centrifuged at 12,000 rpm for 10 min. The top aqueous layer was added to 500 µl isopropanol, mixed well, and centrifuged at 12,000 rpm for 10 min. RNA pellet was washed with 70% ethanol, air-dried, and resuspended in diethylpyrocarbonate (DEPC) water for reverse transcriptase-PCR (RT-PCR) analysis. For RT-PCR analysis, total RNA was DNase treated using DNA-free kit (Ambion), and 1 µg RNA was used for cDNA synthesis reaction using Iscript RT kit (Bio-Rad). Samples were diluted 1:50, and 5 µl cDNA was used per RT-PCR reaction. Real-time quantitative PCR was performed using iQ syber green supermix on the iCycler iQ detection system (Bio-Rad). Sample volume was 20 µl/reaction. Efficiency and specificity of primers were confirmed by standard PCR and DNA electrophoresis.

RT-PCR Program on iCycler was 95°C 15 s, 60°C 30 s, and 72°C 30 s. All real-time PCR values were normalized to actin as indicated. All experiments were confirmed with n = 3 animals. In vivo overexpression studies are normalized to GFP control fibers, which is set to 1 along the y axis. Therefore, all data points reflect a fold difference compared with GFP control fibers.

RT-PCR primer sequences were as follows: G-actin: forward, ACCCAGGCATTGCTGACAGGATGC and reverse, CCATCTAGAAGCATTTGCGGTGGACG; G-SARC: forward TCAGAAGGGGAGGTCACAGGCAG and reverse, TTGGGGCATCCATGCTTAGACTCC; B-SARC: forward, GCTGAGGTTCAAGCAAGTG and reverse, CTTCGCACAATTGCACGC; A-SARC: forward, CAGCCTACAATCGAGACAG and reverse, GTCCCTCAATAGGAAGAGGGA; and dystrophin: forward, GTTCATTGATGGAGACGGAA and reverse, GCACTTCAGCTTCTTCATCT. Additional structural primer sequences were generously provided at http://muscle.cribi.unipd.it/microarrays/atrophy/RT-primers.htm or are available on request.

Immunofluorescence and microscopy
Mouse muscle tissues were excised and placed in trays containing optimal cutting temperature (OCT) compound (VWR). Sections were prepared using a cryostat at 4°C. Sections were kept at –80°C until use. For immunostaining, sections were fixed for 10 min in 4% paraformaldehyde at room temperature. After being washed in PBS 3 x 5 min, sections were blocked in 5% goat serum for 2 h and then incubated overnight with primary antibodies. Primary antibody dilutions were as follows: HDAC4 (clone 186) 1:200 and dystrophin (Novocastra) 1:20. Sections were washed in PBS 3 x 5 min, blocked 90 min in goat serum, and incubated in secondary antibody for 2 h as indicated. Secondary antibodies used were from Jackson ImmunoResearch (West Grove, PA, USA). Nuclei were identified using Hoechst dye. Sections were immersed in mounting medium (fluoromount G, Southern Biotech, Birmingham, AL, USA) for visualization. All images shown were taken with a Zeiss Axioskop compound microscope (Carl Zeiss, Oberkochen, Germany). GFP alone transfected fibers were visualized using fluorescent GFP channel, while HDAC4wt and HDAC4-3SA-GFP fibers were visualized using HDAC4 antibodies, which provided increased sensitivity for detecting overexpressed HDAC4. For muscle integrity analysis, 25 µg GFP, HDAC4-GFP, or HDAC4-3SA-GFP plasmids was electroporatred into tibialis anterior muscles for 7 days. Five micrograms of GFP marker was added to HDAC4wt and 3SA electroporated muscle to allow unambiguous visualization of HDAC4-expressing fibers, thus yielding a diffuse GFP staining pattern for HDAC4wt- and HDAC4-3SA-expressing fibers. Evans blue dye (EBD; Sigma) was intraperitoneally injected (1% solution) 24 h before muscle harvest. Muscles were sectioned and subsequently analyzed by fluorescent microscopy to identify GFP- and EBD-positive fibers.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
HDAC4 represses MEF2-regulated structural and contractile gene expression in cultured myotubes
We previously showed that HDAC4 accumulates in nuclei of denervated muscle, suggesting a regulatory role for HDAC4 in skeletal muscle. Prior studies (7 , 8) have indicated a role for HDAC4 in the repression of MEF2 target gene transcription; however, the identification of these genes in vivo has not been clearly established. With the use of C2C12 muscle cells, HDAC4 potently repressed a synthetic MEF2-driven luciferase reporter (Fig. 1A ). To identify downstream transcriptional targets of HDAC4, microarray analysis was employed using C2C12 myotubes stably overexpressing HDAC4. We identified a subset of muscle structural and contractile genes that was significantly repressed by HDAC4 (Fig. 1B ). On closer examination, many of these gene promoters are reported to contain MEF2 binding sites (Fig. 1B , underlined genes), including myomesin-2, myozenin-1 (FATZ), myotilin, several dystrophin-complex members, and the MEF-2 gene itself. To confirm these findings, the protein levels of several structural genes were evaluated in differentiated myotubes by Western blot analysis. As shown in Fig. 1C , consistent with the array results, the proteins levels of many structural genes were dramatically decreased in HDAC4-expressing myotubes. The repression caused by HDAC4 overexpression was not due to defects in the differentiation program per se, as the induction of myogenin and myotube fusion were not affected (Fig. 1C ; ref. 8 ). In addition, not all muscle genes were regulated by HDAC4, as expression of desmin was unaffected (Fig. 1C ). These results indicate that HDAC4 represses a specific subset of structural genes in cultured muscle myotubes.


Figure 1
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  		Figure 1. HDAC4 represses MEF2-regulated contractile gene expression in cultured muscle myotubes. A) C2C12 myoblasts were transfected with a synthetic 3X-MEF2-element driven luciferase reporter in the absence or presence of MEF2D and/or HDAC4 as indicated. Cells were analyzed for luciferase expression 48 h after transfection. All samples were analyzed in triplicate. B) Microarray analysis was performed on control and HDAC4-expressing C2C12 myotubes. Genes were identified through comparison to controls using a parametric 1-way ANOVA with P = 0.01. Analysis of repressed genes using DAVID revealed a highly significant (P=2.3x10E-5) functional cluster of genes with known sarcomere functions. The genes underlined have been previously shown to contain MEF2 binding sites in their upstream promoters. C) control and HDAC4 stably expressing C2C12 myoblasts were differentiated and harvested at indicated time points for Western blot analysis using antibodies against indicated structural and contractile proteins.

HDAC4 represses muscle structural gene expression in vivo in skeletal muscle
To determine whether HDAC4 regulates structural gene expression in skeletal muscle, we electroporated control GFP, HDAC4-GFP, or the constitutively nuclear mutant HDAC4-3SA-GFP into mouse tibialis muscles. The HDAC4-3SA mutant lacks critical serine residues (S246, 467, 632) required for phosphorylation and nuclear export and therefore is predominantly nuclear localized with enhanced repressor activity (7 , 8) . The subcellular localization of wild-type HDAC4 and HDAC4-3SA in vivo in skeletal muscle was determined by immunostaining analysis, which confirmed a pan-cellular distribution of wild-type HDAC4 (Fig. 2A-D ) and a prominent nuclear accumulation of the HDAC4-3SA mutant (Fig. 2E-H ). Total RNAs from transfected muscle were isolated and analyzed by real-time RT-PCR. As shown in Fig. 3A , structural gene expression is moderately reduced in muscles expressing wild type HDAC4-GFP. However, the repressive effect is more prominent with the HDAC4-3SA mutant, consistent with its predominantly nuclear localization (8) . Similar to wild-type HDAC4, muscles electroporated with the deacetylase defective mutant HDAC4-H803A led to repression of several structural genes, suggesting that HDAC4-mediated repression of structural genes is independent of deacetylase activity (see Supplemental Fig. 1 ; also see ref. 15 and references therein). These results demonstrate that HDAC4 can actively repress the expression of structural gene components in skeletal muscle.


Figure 2
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  		Figure 2. Localization of wild-type HDAC4 and HDAC4-3SA in electroporated skeletal muscle. Wild-type HDAC4 (A–D) or HDAC4-3SA (E–H) constructs were electroporated into mouse tibialis muscles for 7 days, and samples were harvested, sectioned, and analyzed by immunohistochemistry using dystrophin antibodies to mark muscle fibers and Hoechst dye to mark nuclei. GFP fluorescence was used to identify HDAC4-positive electroporated myofibers. White arrows indicate nuclear localized, ectopically expressed wild-type HDAC4 and HDAC4-3SA.


Figure 3
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  		Figure 3. HDAC4 represses structural gene expression in skeletal muscle. A) Mouse tibialis muscles were electroporated with control GFP, wild-type HDAC4-GFP, or HDAC4-3SA-GFP constructs for 7 days. GFP-positive fibers were isolated and RT-PCR analysis was performed using structural gene specific primers. Experimental samples were normalized to control GFP samples, and error is reported as SE. B) Tibialis muscles were isolated from control and 150-day-old pathological ALS mice and analyzed similar to A. C) Asymptomatic 135-day-old ALS mice were electroporated with either scrambled control or HDAC4 siRNA and GFP-positive fibers were isolated for RNA analysis using the indicated gene specific primers. D) Mice were electroporated with scrambled control or HDAC4 siRNA for 3 days and subsequently denervated by sciatic nerve dissection for 7 days. Tibialis muscles were analyzed by RT-PCR using primers for the indicated structural genes. SARC, sarcoglycan.

To investigate the physiological significance of HDAC4-mediated repression of MEF2 target genes, we examined skeletal muscle in which HDAC4 levels are elevated. Previous analysis demonstrated that HDAC4 levels are robustly induced in an ALS model of neuromuscular disease (9) . We therefore determined whether MEF2-regulated structural gene expression is repressed in ALS-affected muscles. Similar to ectopic HDAC4 expression, ALS-affected muscles demonstrated significant repression of structural muscle genes (see Fig. 3B ). Importantly, the expression of several of these structural genes can be significantly restored in ALS mice by electroporation of HDAC4 siRNA (see Fig. 3C ; ref. 9 ). Surgically denervated muscles, in which HDAC4 levels are similarly induced (9) , were reported to show a rapid repression of MEF2-regulated structural and contractile genes (16) . The denervation-dependent repression of MEF2-target genes is significantly abrogated in muscles electroporated with HDAC4 siRNA (Fig. 3D ). Together, these results suggest a critical role for nuclear HDAC4 as a global repressor of muscle structural genes in response to reduced neural activity.

HDAC4 represses muscle gene expression through direct binding and inhibition of MEF2 activity
Previous studies (7 , 8) have shown that HDAC4 represses transcription through direct binding and inhibition of MEF2 function. To determine if HDAC4 regulates muscle gene expression via MEF2, we characterized the transcriptional regulation of two candidate genes: {gamma}-sarcoglycan and myotilin. As shown in Fig. 4A , the {gamma}-sarcoglycan promoter contains two putative AT-rich MEF2 response elements within a 50 bp region ~800 bp upstream of the transcriptional start site (13) . Similarly, the myotilin promoter contains multiple MEF2 elements located within a 2 kb region upstream of the myotilin transcriptional start site (14) . To test whether HDAC4 represses structural genes via inhibition of MEF2, we evaluated {gamma}-sarcoglycan and myotilin promoter activity in C2C12 cells in the presence or absence of MEF2 and/or HDAC4 (see Fig. 4B, C ). Consistent with MEF2 being a transcriptional activator, expression of MEF2 markedly activates the wild-type {gamma}-sarcoglycan and myotilin reporter activity. Mutation of either {gamma}-sarcoglycan MEF2 site individually ({Delta}AT1 or {Delta}AT2) led to a slightly impaired activation of {gamma}-sarcoglycan reporter activity; however, mutation of both elements completely abrogated promoter activity (Fig. 4B ; see {Delta}AT1/{Delta}AT2 mutant). Similarly, a myotilin construct lacking critical MEF2 binding elements showed reduced activation in the presence of MEF2 (Fig. 4C ). Notably, HDAC4 represses MEF2-dependent {gamma}-sarcoglycan and myotilin reporter activity but has little effect when the MEF2 binding sites are mutated (Fig. 4B, C , HDAC4 lanes). To further confirm that HDAC4 represses transcription directly, we determined if HDAC4 and MEF2 physically associate with the {gamma}-sarcoglycan promoter by ChIP assay in cultured C2C12 myotubes. As shown in Fig. 4D , ChIP analysis clearly showed that HDAC4 and MEF2 bind a 300 bp genomic region of the {gamma}-sarcoglycan promoter encompassing two AT-rich MEF2 binding sites. These results show that HDAC4 represses gene transcription via binding and inhibition of MEF2 activity.


Figure 4
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  		Figure 4. HDAC4 directly represses structural gene expression via binding and inhibition of MEF2 activity. A) Promoter sequence of {gamma}-sarcoglycan spanning –812 to –861 nt upstream of transcriptional start. AT-rich MEF2 sites are indicated above, which were mutated singly in or combination for subsequent luciferase assays. B) C2C12 myoblasts were tranfected with luciferase reporter constructs containing wild-type or MEF2 binding site deletion ({Delta}AT1, {Delta}AT2, or {Delta}AT1,2) promoters as indicated. Transfections occurred in the presence of MEF2 and/or HDAC4 as indicated. Luciferase activity was determined 48 h after transfection. C) luciferase analysis was performed using either 2 or 1.3 kb myotilin promoter-driven luciferase constructs (14) . D) ChIP analysis was performed on C2C12 myotubes using either control IgG-, MEF2-, or HDAC4-specific antibodies. Bound DNA was analyzed by standard PCR generating a 300 bp fragment encompassing both MEF2 binding sites in the {gamma}-sarcoglycan promoter. Input lane represents 5% of total starting material.

Nuclear accumulation of HDAC4 causes increased skeletal muscle damage
The global repression of MEF2-target genes by HDAC4 could contribute to muscle defects, as previous studies (2 , 4) have demonstrated that loss of MEF2 leads to impaired structural integrity and defects in sarcomeric organization. To explore this possibility, uptake of EBD dye, an indicator of muscle damage, was examined in muscle fibers expressing either control GFP, wild-type HDAC4, or the nuclear HDAC4-3SA mutant. Mice were injected with EBD 24 h before muscle harvest, and tibialis muscles were subsequently sectioned and analyzed by fluorescent microscopy to identify GFP- and EBD-positive fibers. As shown in Fig. 5 , myofibers expressing wild-type HDAC4 show a moderate uptake of EBD (~8% of the transfected fibers; see Fig. 5E ), while the majority of fibers expressing the constitutively active HDAC4-3SA mutant are positive for EBD (83%, see Fig. 5F ). Importantly, this phenotype is not caused by muscle damage induced by electroporation, as GFP-expressing myofibers are not permeable to EBD (Fig. 5D ). Thus, accumulation of nuclear HDAC4 represses MEF2-dependent structural genes and contributes to defects in muscle integrity.


Figure 5
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  		Figure 5. Nuclear accumulation of HDAC4 causes skeletal muscle damage. Tibialis muscles were electroporated with control GFP (A, D, G), wild-type HDAC4 (B, E, H), or HDAC4-3SA (C, F, I) for 7 days. GFP control marker was coelectroporated at a 1:5 ratio to allow identification of GFP-positive/HDAC4-expressing myofibers; 1% EBD was intraperitoneally injected 24 h before muscle harvest. Tibialis muscles were isolated, sectioned, and analyzed by fluorescent microscopy to identify GFP-positive (A–C) and EBD-positive fibers (D–F), which is represented in merge panels (G–I). The EBD-positive fibers were quantified in GFP- and HDAC4-expressing fibers and represented as percentage of EBD positive fibers out of total electroporated fibers, as indicated below images.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Although HDAC4 has been proposed to regulate muscle function, the physiological targets of HDAC4 in skeletal muscle are largely unknown. Microarray analysis revealed that HDAC4 regulates the expression of a subset of MEF2-dependent contractile and structural genes. Indeed, we show that HDAC4 directly represses MEF2-dependent transcription through an association with AT-rich MEF2 elements. Elevated HDAC4 levels caused by HDAC4 overexpression, surgical denervation, or pathological neuromuscular disease led to a significant repression of MEF2-target genes. Our results establish HDAC4 as a critical in vivo regulator of MEF2 function and provide a novel physiological mechanism by which neural activity regulates muscle gene transcription.

We (9) have previously shown that HDAC4 is robustly induced and accumulates in muscle nuclei in response to denervation. This regulation is in stark contrast to the related deacetylase HDAC9, the levels of which are rapidly reduced after denervation (17) , suggesting a distinct role for HDAC4 in neural activity-dependent signaling. These findings led us to propose that HDAC4 acts as a molecular link between neural activity and the muscle transcriptional machinery. Our current study provides the first evidence that neural activity-regulated HDAC4 controls MEF2 target genes in vivo. The analysis of {gamma}-sarcoglycan and myotilin expression revealed that HDAC4 represses transcription via MEF2-response elements (Fig. 4) . Although not fully characterized, numerous muscle structural genes including dystrophin and {alpha}-sarcoglycan also contain putative MEF2 response elements (18 , 19) . These results indicate a coordinated regulation of a specific subset of muscle genes by the HDAC4-MEF2 transcription complex.

Previous studies (5 , 6) have indicated that, similar to HDAC4, MEF2 function is responsive to neural activity, as endurance training elevates MEF2 activity while surgical denervation represses MEF2 target genes, including MLC2. The results presented in this study suggest that reduced neural activity and nuclear accumulation of HDAC4 are critical for repression of MEF2-target gene expression. Thus, the activity responsiveness of the HDAC4/MEF2 transcription complex could effectively couple neural input to the remodeling of muscle gene transcription. In this regard, note that CaMKII, an NMJ-localized HDAC4 kinase (9) , is thought to act as a calcium sensor in response to changes in neural activity (20) . Therefore, CaMKII could potentially relay changes in neural activity to the HDAC4/MEF2 transcription complex required for muscle remodeling.

The pathophysiological importance of these findings began to emerge when we found that HDAC4 is also robustly induced and accumulates in nuclei of muscle affected by motor neuron disease, such as ALS (9) . ALS mice (SOD1-G93A mutant), in which HDAC4 levels are highly induced, show a reduction in structural components (see Fig. 3 ). In light of our findings that HDAC4 is a potent repressor of structural gene transcription, it is possible that the induction of HDAC4 by chronically reduced neural input, as is the case in ALS-affected muscles, causes reduction in MEF2-regulated structural gene expression, contributing to muscle defects (as shown in Fig. 5 ). Indeed, studies (2 , 4) in both mice and zebrafish have demonstrated an essential role for MEF2 family members in structural integrity and contractile function. This suggests that prolonged nuclear accumulation and "activation" of HDAC4 in muscles affected by neuromusculear diseases such as ALS would phenocopy muscles lacking MEF2, thereby displaying prominent contractile defects. Previous studies (21 , 22) have linked ALS with muscle abnormalities, as ultrastructural analysis of muscle from ALS patients revealed severe disorganization and perturbation of membrane integrity. Therefore, the induction of HDAC4 and subsequent repression of MEF2 target genes could, in fact, be one pathological component associated with ALS and other neuromuscular diseases. If this supposition is true, it would suggest a new therapeutic angle for improving muscle function in neuromuscular disease patients. In this regard, pan-HDAC inhibitors have shown therapeutic potential in the treatment of both ALS and spinal muscular atrophy models (23 24 25) . Inhibition of HDAC4 activity could account, at least in part, for the observed improvement in muscle phenotype seen in these neuromuscular disease models. Although HDAC4 deacetylase activity is not required for inhibition of MEF2 function (Supplemental Fig. 1; ref. 15 ), it remains possible that HDAC4 activity regulates muscle factors other than MEF2.

The characterization of HDAC4 as a critical in vivo regulator of MEF2 target genes and a contributing factor to muscle dysfunction highlights the importance of this neural activity-regulated deacetylase in muscle biology. Elucidating how HDAC4 is regulated by neural activity and whether it affects a broad spectrum of muscle phenotypes would provide critical insight into both physiological and pathological muscle remodeling and pave the way to novel therapeutic opportunities for muscle dysfunction associated with neuromuscular disorders.


   ACKNOWLEDGMENTS
 
We thank Tim Bolger, Xuan Zhao, and Chun Lai for technical assistance. We thank Satoru Noguchi (National Institute of Neuroscience, Tokyo, Japan) and Olli Carpen (University of Helsinki, Helsinki, Finland) for kindly providing mouse {gamma}-sarcoglycan and mouse myotilin reporter constructs and Mike Hauser (Duke University, Durham, NC, USA) for providing myotilin antibody. This work was supported by the Leukemia and Lymphoma Society and grant AR055613 (to T.-P.Y), and the U.S. National Institutes of Health and the ALS Association (to G.A.C.).

Received for publication June 19, 2008. Accepted for publication August 14, 2008.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Black, B. L., Olson, E. N. (1998) Transcriptional control of muscle development by myocyte enhancer factor-2 (MEF2) proteins. Annu. Rev. Cell Dev. Biol. 14,167-196[CrossRef][Medline]
  2. Potthoff, M. J., Arnold, M. A., McAnally, J., Richardson, J. A., Bassel-Duby, R., Olson, E. N. (2007) Regulation of skeletal muscle sarcomere integrity and postnatal muscle function by Mef2c. Mol. Cell. Biol. 27,8143-8151[Abstract/Free Full Text]
  3. Potthoff, M. J., Wu, H., Arnold, M. A., Shelton, J. M., Backs, J., McAnally, J., Richardson, J. A., Bassel-Duby, R., Olson, E. N. (2007) Histone deacetylase degradation and MEF2 activation promote the formation of slow-twitch myofibers. J. Clin. Invest. 117,2459-2467[CrossRef][Medline]
  4. Hinits, Y., Hughes, S. M. (2007) Mef2s are required for thick filament formation in nascent muscle fibres. Development 134,2511-2519[Abstract/Free Full Text]
  5. Wu, H., Rothermel, B., Kanatous, S., Rosenberg, P., Naya, F. J., Shelton, J. M., Hutcheson, K. A., DiMaio, J. M., Olson, E. N., Bassel-Duby, R., Williams, R. S. (2001) Activation of MEF2 by muscle activity is mediated through a calcineurin-dependent pathway. EMBO J. 20,6414-6423[CrossRef][Medline]
  6. Esser, K., Nelson, T., Lupa-Kimball, V., Blough, E. (1999) The CACC box and myocyte enhancer factor-2 sites within the myosin light chain 2 slow promoter cooperate in regulating nerve-specific transcription in skeletal muscle. J. Biol. Chem. 274,12095-12102[Abstract/Free Full Text]
  7. McKinsey, T. A., Zhang, C. L., Lu, J., Olson, E. N. (2000) Signal-dependent nuclear export of a histone deacetylase regulates muscle differentiation. Nature 408,106-111[CrossRef][Medline]
  8. Zhao, X., Ito, A., Kane, C. D., Liao, T. S., Bolger, T. A., Lemrow, S. M., Means, A. R., Yao, T. P. (2001) The modular nature of histone deacetylase HDAC4 confers phosphorylation-dependent intracellular trafficking. J. Biol. Chem. 276,35042-35048[Abstract/Free Full Text]
  9. Cohen, T. J., Waddell, D. S., Barrientos, T., Lu, Z., Feng, G., Cox, G., Bodine, S. C., Yao, T. P. (2007) The histone deacetylase HDAC4 connects neural activity to muscle transcriptional reprogramming. J. Biol. Chem. 282,33752-33759[Abstract/Free Full Text]
  10. Garvey, S. M., Miller, S. E., Claflin, D. R., Faulkner, J. A., Hauser, M. A. (2006) Transgenic mice expressing the myotilin T57I mutation unite the pathology associated with LGMD1A and MFM. Hum. Mol. Genet. 15,2348-2362[Abstract/Free Full Text]
  11. Dennis, G., Jr, Sherman, B. T., Hosack, D. A., Yang, J., Gao, W., Lane, H. C., Lempicki, R. A. (2003) DAVID: Database for Annotation, Visualization, and Integrated Discovery. Genome Biol. 4,P3[CrossRef][Medline]
  12. Nelson, J. D., Denisenko, O., Bomsztyk, K. (2006) Protocol for the fast chromatin immunoprecipitation (ChIP) method. Nat. Protoc. 1,179-185[CrossRef][Medline]
  13. Wakabayashi-Takai, E., Noguchi, S., Ozawa, E. (2001) Identification of myogenesis-dependent transcriptional enhancers in promoter region of mouse gamma-sarcoglycan gene. Eur. J. Biochem. 268,948-957[Medline]
  14. Mologni, L., Moza, M., Lalowski, M. M., Carpen, O. (2005) Characterization of mouse myotilin and its promoter. Biochem. Biophys. Res. Commun. 329,1001-1009[CrossRef][Medline]
  15. Zhao, X., Sternsdorf, T., Bolger, T. A., Evans, R. M., Yao, T. P. (2005) Regulation of MEF2 by histone deacetylase 4- and SIRT1 deacetylase-mediated lysine modifications. Mol. Cell. Biol. 25,8456-8464[Abstract/Free Full Text]
  16. Raffaello, A., Laveder, P., Romualdi, C., Bean, C., Toniolo, L., Germinario, E., Megighian, A., Danieli-Betto, D., Reggiani, C., Lanfranchi, G. (2006) Denervation in murine fast-twitch muscle: short-term physiological changes and temporal expression profiling. Physiol. Genomics 25,60-74[Abstract/Free Full Text]
  17. Mejat, A., Ramond, F., Bassel-Duby, R., Khochbin, S., Olson, E. N., Schaeffer, L. (2005) Histone deacetylase 9 couples neuronal activity to muscle chromatin acetylation and gene expression. Nat. Neurosci. 8,313-321[CrossRef][Medline]
  18. Delgado-Olguin, P., Recillas-Targa, F., Rosas-Vargas, H., Salamanca, F., Coral-Vazquez, R. M. (2006) Partial characterization of the mouse alpha-sarcoglycan promoter and its responsiveness to MyoD. Biochim. Biophys. Acta 1759,240-246[Medline]
  19. Klamut, H. J., Bosnoyan-Collins, L. O., Worton, R. G., Ray, P. N. (1997) A muscle-specific enhancer within intron 1 of the human dystrophin gene is functionally dependent on single MEF-1/E box and MEF-2/AT-rich sequence motifs. Nucleic Acids Res. 25,1618-1625[Abstract/Free Full Text]
  20. De Koninck, P., Schulman, H. (1998) Sensitivity of CaM kinase II to the frequency of Ca2+ oscillations. Science 279,227-230[Abstract/Free Full Text]
  21. Drachman, D. B., Murphy, S. R., Nigam, M. P., Hills, J. R. (1967) "Myopathic" changes in chronically denervated muscle. Arch. Neurol. 16,14-24[Abstract/Free Full Text]
  22. Fidzianska, A. (1976) Morphological differences between the atrophied small muscle fibres in amyotrophic lateral sclerosis and Werdnig-Hoffmann disease. Acta. Neuropathol. 34,321-327[CrossRef][Medline]
  23. Ryu, H., Smith, K., Camelo, S. I., Carreras, I., Lee, J., Iglesias, A. H., Dangond, F., Cormier, K. A., Cudkowicz, M. E., Brown, R. H., Jr, Ferrante, R. J. (2005) Sodium phenylbutyrate prolongs survival and regulates expression of anti-apoptotic genes in transgenic amyotrophic lateral sclerosis mice. J. Neurochem. 93,1087-1098[CrossRef][Medline]
  24. Brichta, L., Holker, I., Haug, K., Klockgether, T., Wirth, B. (2006) In vivo activation of SMN in spinal muscular atrophy carriers and patients treated with valproate. Ann. Neurol. 59,970-975[CrossRef][Medline]
  25. Minamiyama, M., Katsuno, M., Adachi, H., Waza, M., Sang, C., Kobayashi, Y., Tanaka, F., Doyu, M., Inukai, A., Sobue, G. (2004) Sodium butyrate ameliorates phenotypic expression in a transgenic mouse model of spinal and bulbar muscular atrophy. Hum. Mol. Genet. 13,1183-1192[Abstract/Free Full Text]




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