Skip Navigation


Human Molecular Genetics Advance Access originally published online on June 26, 2006
Human Molecular Genetics 2006 15(15):2348-2362; doi:10.1093/hmg/ddl160
This Article
Right arrow Abstract Freely available
Right arrow FREE Full Text (PDF) Freely available
Right arrow All Versions of this Article:
15/15/2348    most recent
ddl160v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Add to My Personal Archive
Right arrow Download to citation manager
Right arrow Search for citing articles in:
ISI Web of Science (11)
Right arrowRequest Permissions
Google Scholar
Right arrow Articles by Garvey, S. M.
Right arrow Articles by Hauser, M. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Garvey, S. M.
Right arrow Articles by Hauser, M. A.
Social Bookmarking
Add to CiteULike   Add to Connotea   Add to Del.icio.us  
What's this?

© The Author 2006. Published by Oxford University Press. All rights reserved. For Permissions, please email: journals.permissions@oxfordjournals.org

Transgenic mice expressing the myotilin T57I mutation unite the pathology associated with LGMD1A and MFM

Sean M. Garvey1,2, Sara E. Miller3, Dennis R. Claflin4, John A. Faulkner4,5 and Michael A. Hauser1,2,*

1 Center for Human Genetics, 2 University Program in Genetics and Genomics and 3 Department of Pathology, Duke University Medical Center, Durham, NC 27710, USA and 4 Department of Surgery and 5 Institute of Gerontology, University of Michigan, Ann Arbor, MI 48109-2007, USA

* To whom correspondence should be addressed at: Duke University Medical Center, DUMC Box 3445, Durham, NC 27710, USA. Tel: +1 9196843508; Fax: +1 9196840902; Email: mhauser{at}chg.duhs.duke.edu

Received April 26, 2006; Accepted June 21, 2006


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Myotilin is a muscle-specific Z-disc protein with putative roles in myofibril assembly and structural upkeep of the sarcomere. Several myotilin point mutations have been described in patients with limb-girdle muscular dystrophy type 1A (LGMD1A), myofibrillar myopathy (MFM), spheroid body myopathy (SBM), three similar adult-onset, progressive and autosomal dominant muscular dystrophies. To further investigate myotilin's role in the pathogenesis of these muscle diseases, we have characterized three independent lines of transgenic mice expressing mutant (T57I) myotilin under the control of the human skeletal actin promoter. Similar to LGMD1A and MFM patients, these mice develop progressive myofibrillar pathology that includes Z-disc streaming, excess myofibrillar vacuolization and plaque-like myofibrillar aggregation. These aggregates become progressively larger and more numerous with age. We show that the mutant myotilin protein properly localizes to the Z-disc and also heavily populates the aggregates, along with several other Z-disc associated proteins. Whole muscle physiological analysis reveals that the extensor digitorum longus muscle of transgenic mice exhibits significantly reduced maximum specific isometric force compared with littermate controls. Intriguingly, the soleus and diaphragm muscles are spared of any abnormal myopathology and show no reductions in maximum specific force. These data provide evidence that myotilin mutations promote aggregate-dependent contractile dysfunction. In sum, we have established a promising patho-physiological mouse model that unifies the phenotypes of LGMD1A, MFM and SBM.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
The muscular dystrophies comprise a large number of clinically and molecularly heterogeneous myopathic diseases, characterized by progression in muscle weakness and histological evidence of muscle fiber degeneration and regeneration. A subset, the limb-girdle muscular dystrophies (LGMDs), is characterized by initial or predominant weakness of the shoulder and hip girdle muscles (1). The similarity in clinical presentation of LGMD and other muscle diseases often complicates diagnosis. LGMDs exist in both autosomal dominant forms (type I, LGMD1A-G) and recessive forms (type II, LGMD2A-J). Given the extensive phenotypic heterogeneity of LGMD, sub-types are positionally or molecularly defined; 13 independent gene defects resulting in LGMD have been reported to date (2).

The subject of this article, LGMD1A (MIM 604103 [OMIM] ), is an autosomal dominant muscle disease, with a mean age at onset of 27 years, characterized by initial weakness of proximal girdle muscles (3). Muscular symptoms progress in both severity and distribution, as more distal limb muscles also become affected. Biopsied skeletal muscle shows several signs of myopathy—degeneration of myofibers, fiber splitting, variations in fiber size, fibrosis and centrally located myonuclei (4). Ultrastructural analysis reveals a large number of autophagic vacuoles and extensive Z-disc streaming. Substantial clinical heterogeneity is found within a single large LGMD1A family; approximately half of all patients exhibit dysarthric, nasal speech, and age at onset of patient-reported symptoms varies across more than four decades (5).

LGMD1A is caused by mutation of the myotilin gene (MYOT, TTID). Two independent myotilin missense mutations, T57I and S55F, cause LGMD1A in a large North American family and an Argentinian pedigree, respectively (4,6). These amino acid substitutions occur in the N-terminal domain of myotilin, a domain with no known homology or functional motifs. Myotilin is predominantly expressed in skeletal and cardiac muscle. The 57 kDa, 498 amino acid residue myotilin protein harbors two immunoglobulin-like (IgL) domains at its C-terminus. The N-terminal domain contains a hydrophobic stretch of amino acids embedded within a serine-rich region. Myotilin localizes to the sarcomere Z-disc, the electron-dense structure that contributes to sarcomere assembly, actin filament stabilization, and muscle force transmission (7). At the Z-disc, myotilin interacts with {alpha}-actinin-2 (ACTN2) via residues 80–124 (8). Through one and/or both of its IgL domains, myotilin has also been shown to interact with {gamma}-filamin (FLNC) (9), actin monomers and filaments (10) and calsarcins-1 and -2 (11). The IgL domains of myotilin also mediate homodimerization (10). Salmikangas et al. have shown in vitro that myotilin can bundle actin filaments, either acting alone or cooperatively with ACTN2, and have therefore hypothesized that myotilin plays a significant role in anchoring and stabilizing F-actin (10,12). This hypothesis is also supported by the predominantly sarcomeric pathology observed in patients with myotilin mutations.

Mutations in myotilin have been associated with two other muscular dystrophies: spheroid body myopathy (SBM) and myofibrillar myopathy (MFM). The myotilin S39F mutation causes SBM, an autosomal dominant muscle disease characterized by dense, intrasarcoplasmic bodies (13). In addition to the SBM mutation, several myotilin mutations (S55F, S60C, S60F, S95I) have been identified in a subset of MFM patients (14,15). MFM is a multigenic, autosomal dominant and progressive muscular dystrophy, characterized pathologically by dense patches of SBM-like sarcomeric disarray and aggregation. Genetic studies of MFM have highlighted the importance of additional Z-disc proteins in sarcomere function. Mutation of the myotilin-binding protein, FLNC, causes an MFM (MIM#609524) characterized by intrasarcoplasmic aggregates similar to those seen in myotilin-associated MFM (16). MFM is also caused by mutations in the Z-disc proteins, ZASP (z-band alternatively spliced PDZ-containing protein; MIM#609542) (17), desmin (MIM#601419) (18–20) and the chaperone protein, {alpha}B-crystallin (CRYAB, MIM#608810) (21,22). Despite recent genetic triumphs, LGMD1A, myotilin-related MFM and SBM, collectively termed the myotilinopathies, remain clinically ill-defined muscle disorders with considerable inter- and intra-familial variations.

A thorough molecular dissection of myotilinopathy disease etiology has largely been limited by the availability of human tissue and the lack of an experimental model system. To address these needs, we have created a transgenic mouse model by expressing the human myotilin cDNA bearing the T57I mutation on a wild-type murine myotilin background. These transgenic mice reproduce many of the symptoms and pathology associated with the myotilinopathies—Z-disc streaming, myofibrillar aggregation and muscle weakness. This mouse model not only unifies the diverse phenotypes of the human myotilinopathies, but also promises to be a key resource for understanding myotilin function, unraveling LGMD1A pathogenesis, and investigating possible therapeutics.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Generation of myotilin transgenic mice
We modeled the autosomal dominant disease LGMD1A by co-expressing a mutant human myotilin transgene in the presence of normal levels of endogenous wild-type murine myotilin. The mutant transgene contains the human myotilin cDNA with the LGMD1A T57I point mutation, cloned downstream of the skeletal muscle-specific human skeletal actin (HSA) promoter (Fig. 1). We chose to use the human cDNA because there are sequence differences between murine and human myotilin near the mutation site, and we wanted to avoid the possibility that these differences would alter the effects of the human mutation. Because myotilin antibodies recognizes both human and murine proteins, we incorporated a c-myc epitope tag at the N-terminus of the human transgene product, thus enabling transgene-specific quantification and localization. Transgenes were genotyped by PCR amplification (Fig. 1), and a single site of insertion was verified by Southern blot (data not shown). Transgene expression and transgene-associated pathology were similar in three distinct T57I mutant myotilin lines. We chose to thoroughly characterize line 71, Tg(HSA-MYOT)71Mah, hereafter referred to as TgT57I. We also generated a line of mice expressing a similar transgene containing the wild-type human myotilin cDNA, Tg(HSA-MYOT)12Mah, hereafter referred to as TgWT. These two transgenic constructs differ only by the single base pair that causes the T57I amino acid substitution. Transgenic mice were obtained at the expected Mendelian ratio, and neither TgT57I nor the TgWT mice displayed gross abnormalities or reduced survival. RT-PCR analysis shows that transgene expression is limited to skeletal muscle in both TgT57I (Fig. 1) and TgWT mice (data not shown). Western blotting with a c-myc antibody specific to the transgene product shows that the expected 57 kDa TgT57I protein is appropriately expressed in multiple muscles (Fig. 1). This c-myc antibody also allows a direct comparison of transgene expression levels: TgWT mice express their transgene product at 2- to 3-fold higher levels than do TgT57I mice, as determined by analysis of soluble fractions of quadriceps muscle lysates (Fig. 1). The abundance of TgWT expression is further confirmed using an antibody raised to a peptide that is identical in human and murine myotilin (Fig. 1). Scanning densitometry was used to determine that TgWT levels are 7.7-fold higher than endogenous levels, whereas TgT57I levels are 2.6-fold higher than endogenous levels. Immunostaining of frozen muscle cross-sections shows that transgene expression is uniform across myofibers within a given muscle group and also similar between slow type I and fast type II fibers (Fig. 1).


Figure 1601
View larger version (26K):
[in this window]
[in a new window]
  		Figure 1. Transgene design and expression. (A) The human myotilin cDNA was cloned downstream of the skeletal muscle-specific HSA promoter and the VP1 splice donor and acceptor sites. The human myotilin cDNA includes the full-length coding region (gray box) and 5'- and 3'-UTRs (white boxes). A c-myc epitope tag (red line) was appended to the myotilin N-terminus. Both wild-type (TgWT) and T57I point mutant (TgT57I) myotilin transgenic mice were generated. (B) The transgene was genotyped by PCR using a forward HSA primer and a reverse myotilin 5'-UTR primer. +/Tg, transgene-positive heterozygote; +/+, control littermate. (C) Reverse transcription–PCR of total RNA from brain (B), heart (H), kidney (K), liver (L), spinal cord (SC) and skeletal muscle (SM) shows that the TgT57I transcript is observed only in skeletal muscle. (D) Western blot survey of several soluble fractions of muscle lysates from 3-month-old TgT57I mice. The myotilin C-term peptide antibody reacts to both murine and transgenic human myotilin peptides; the c-myc antibody is specific to the transgene products. Ab, abdomenal wall muscle; Bi, biceps; Di, diaphragm; Ed, extensor digitorum longus; Ga, gastrocnemius; Qu, quadriceps; So, soleus; Ti, tibialis. (E) Soluble fractions of quadriceps lysates from 3-month-old transgenic mice were analyzed by western blot to show relative expression levels. The c-myc antibody shows that the TgWT product is expressed at 2.3-fold higher levels than the TgT57I product. The myotilin antibody shows that the TgWT levels are 7.7-fold higher than endogenous myotilin levels. TgT57I levels are 2.6-fold higher than endogenous levels, as determined by optical scanning densitometry on diluted samples. (F and G) A Cy3-conjugated c-myc antibody was used to specifically immunolocalize transgene products in soleus muscle cross-sections. TgWT (F) staining is stronger than TgT57I (G) staining. (H and I) Double-labeling of TgT57I quadriceps muscle transverse sections with a slow isoform-specific myosin antibody (H) and the c-myc antibody (I) shows that the transgene is expressed in both slow type I and fast type II fibers.

 
There is no overt toxicity associated with overexpression of the human N-terminally epitope-tagged myotilin transgene product (TgWT) in mouse skeletal muscle, even at 2 years of age. Although we do observe age-related pathology common to transgene-negative littermate controls (i.e. tubular aggregates and rimmed vacuoles), minimal transgene-specific pathology is also observed. Histology of 18-month TgWT triceps, quadriceps, extensor digitorum longus (EDL) and soleus muscle is comparable with 12-month control muscle (Fig. 2). We observe small eosinophilic aggregates beneath the sarcolemma in quadriceps and triceps muscles of older TgWT mice as result of significant myotilin overexpression. Because the TgWT protein is expressed at several-fold higher levels than the TgT57I product, any pathology in TgT57I mice must be caused solely by the T57I point mutation.


Figure 1602
View larger version (126K):
[in this window]
[in a new window]
  		Figure 2. TgT57I mice develop myofibrillar aggregates. (A and B) Light microscopic evaluation of TgT57I muscle. (A) Masson's trichrome-stained cross-section of a 12-month-old TgT57I quadriceps myofiber. The sarcoplasmic aggregate is dense and amorphous and stains dark red or bluish green. (B) The amorphous structure of the aggregate and the presence of associated vacuoles are further revealed in this toluidine blue-stained 0.5 µm epoxy cross-section. (C–N) Masson's trichrome-stained cross-sections of quadriceps, triceps, EDL and soleus muscles from TgT57I (12-month), TgWT (18-month) and control littermate (+/+, 12-month) mice. (C, F, I) Myofibrillar aggregates (arrows) are prevalent in TgT57I quadriceps, triceps and EDL muscles. Adipose infiltration (asterisks) and an increase in the number of tubular aggregates (white circles) are also common in older TgT57I muscle. (L) TgT57I soleus muscle exhibits no myopathology. (D, G, J, M) TgWT muscle does not develop myofibrillar aggregates. (E, H, K, N) Cross-sections of wild-type muscle. Scale bar: 10 µm (A and B); 50 µm (C–N).

 
TgT57I mice develop progressive myofibrillar pathology
Expression of T57I mutant myotilin results in a myofibrillar muscle pathology remarkably similar to that seen in the human myotilinopathies. TgT57I mice develop dense and irregular myofibrillar aggregates (Fig. 2). A survey of muscles from a 12-month-old TgT57I mouse shows that the myofibrillar aggregates are prevalent in two proximal muscle groups, the quadriceps and triceps, and also in the EDL muscle (Fig. 2). Fibrosis, adipose infiltration and increased tubular aggregation also occur in older TgT57I muscle (Fig. 2).

Muscle pathology in the TgT57I mouse is progressive, with both the size and number of aggregates increasing with age. At 2 weeks of age, aggregates are small focal points, which subsequently expand up to 40 µm in diameter, often occupying the entire cross-sectional areas of myofibers in older mice (Fig. 3). The number of affected fibers also increases with age: survey of at least 800 myofibers from the quadriceps shows that the proportion containing aggregates increases from 11.7% at 6 months to 17.8% at 12 months. During the same time period, the number of affected fibers in the triceps muscle increases from 11.6% to 18.2%.


Figure 1603
View larger version (68K):
[in this window]
[in a new window]
  		Figure 3. Myopathology is progressive in TgT57I mice. (A) Masson's trichrome-stained cross-section of quadriceps muscle from a 2-week-old TgT57I mouse. Myofibrillar aggregates are small and focal (arrows). (B) Cross-section of quadriceps muscle from a 12-month-old TgT57I mouse. Larger aggregates (arrows) populate many of the myofibers. Pathology also includes a tract of fibrosis (white asterisk) and adipose infiltration (black asterisk). Scale bar: 25 µm.

 
Different muscle groups show different levels of myopathic involvement (Table 1). Quadriceps, triceps and hamstring muscles exhibit a higher proportion of myofibers with aggregates than EDL, gastrocnemius, abdominal wall and tibialis muscles. Interestingly, the diaphragm, soleus, biceps and ulnar muscles show no evidence of myofibrillar aggregation. This selectivity of muscle group involvement is also seen in human patients: magnetic resonance imaging of myotilin-related MFM patients has revealed selective muscle wasting in the leg (15). The three most severely affected muscles in TgT57I mice are upper forelimb and hindlimb muscles—precisely the same muscle groups that show initial weakness in LGMD1A patients.


View this table:
[in this window]
[in a new window]
  		Table 1. Myofibrillar aggregation is muscle-group dependent in TgT57I mice

 
Ultrastructural analysis of TgT57I muscle
TgT57I muscle recapitulates the ultrastructural defects observed in human myotilinopathy patients. Broadening of isolated Z-discs is observed in both single LGMD1A sarcomeres and in TgT57I muscle (Fig. 4). Disrupted Z-discs may subsequently merge into streaming bodies involving multiple Z-discs. Z-disc streaming in TgT57I mice is indistinguishable from the pathology seen in human LGMD1A patients (Fig. 4). TgT57I muscle also develops striking patches of sarcomeric deterioration and aggregation. These myofibrillar aggregates are irregular, amorphous and devoid of any peripheral or internal membranous material. Myofibrillar aggregates are often associated with autophagic vesicles (Fig. 4), a vacuolar pathology that is prevalent in LGMD1A muscle. Myofibrillar aggregates are also sometimes associated with tubular aggregates, a non-specific pathology observed in older inbred male mice (23). These tubular aggregates are more prevalent in TgT57I mice than in male littermate controls (data not shown). Tubular aggregates have been reported to be associated with the intrasarcoplasmic aggregates in SBM patient muscle (24). Centrally located nuclei are observed in TgT57I thin sections, but not regularly. Sarcolemmal, mitochondrial and nuclear structures all appear normal by electron microscopy. TgWT muscle exhibits normal sarcomeric ultrastructure; however, small aggregates can be found beneath the sarcolemma in older mice (Fig. 4). These sub-sarcolemmal aggregates are commonly <5 µm in diameter and contain embedded, often fragmented, vesicles. The appearance of these aggregates contrasts sharply with that of larger myofibrillar aggregates in TgT57I mice that are devoid of any membranous deposits. Ultrastructural analysis of TgT57I muscle further highlights the largely myofibrillar pathology that is common to the human myotilinopathies.


Figure 1604
View larger version (159K):
[in this window]
[in a new window]
  		Figure 4. Ultrastructural pathology of myotilin transgenic muscle. Electron microscopy was performed on quadriceps muscle dissected from a 12-month-old male +/TgT57I mouse (A–F) and a 12-month-old female +/TgWT mouse (G–I). (A) An electron micrograph of a longitudinal section of TgT57I skeletal muscle shows that Z-discs are often irregular and broadened (bracket), as is also seen in LGMD1A patient muscle (inset). (B) The Z-disc streaming (asterisk) in TgT57I muscle is reminiscent of Z-disc streaming in LGMD1A patient muscle (inset). (C) Expanded regions of sarcomeric dissolution and myofibrillar aggregation are prevalent in TgT57I muscle. (D) The myofibrillar aggregates are irregular and amorphous and expand up to 25–40 µm in both length and diameter. (E) Myofibrillar aggregates are often associated with vacuoles (arrows). (F) Tubular aggregation (TA) is often associated with the transgene-induced myofibrillar aggregates (MA) (G–I). TgWT quadriceps muscle ultrastructure. (G) Longitudinal section of TgWT muscle shows normal sarcomeric structure. (H) Sub-sarcolemmal aggregates with internal vesicles develop in older TgWT muscle. (I) Sarcolemmal ultrastructure is normal in TgWT muscle. Scale bar: 0.5 µm (A); 1.0 µm (B, G, H, I); 5.0 µm (C–F).

 
Mutant myotilin and Z-disc proteins localize to the aggregates
The myofibrillar aggregates in TgT57I muscle harbor several Z-disc proteins, including the mutant transgene product (Fig. 5). Immunostaining for the c-myc epitope tag allows for specific localization of the transgene products. Both the TgWT and TgT57I products localize normally to the Z-disc, but the T57I mutant protein is also found in the myofibrillar aggregates (Fig. 5). Overexpression of wild-type myotilin also drives sub-sarcolemmal accumulation. We investigated the localization of several other Z-disc-associated proteins. Although ACTN2, FLNC and desmin all localize to the Z-disc in muscles from TgT57I, TgWT and control littermates, these proteins also localize heavily to the aggregates in TgT57I muscle. Slight accumulation of FLNC and desmin can be observed in TgWT muscle. Sarcomeric proteins titin and myosin localize to the aggregates as well, however vinculin does not (data not shown). In addition, the aggregates contain ubiquitinated protein (Fig. 5). Similar patterns of ectopic expression have been reported in MFM and SBM muscles (14,15,25,26).


Figure 1605
View larger version (85K):
[in this window]
[in a new window]
  		Figure 5. Immunolocalization of Z-disc proteins in myotilin transgenic muscle. Proteins were immunolocalized in longitudinal sections of quadriceps muscle from 6-month-old +/TgT57I, +/TgWT and control littermate (+/+) mice. (A and B) A Cy3-conjugated c-myc antibody was used to localize the transgene products. (A) TgT57I localizes to the Z-disc and heavily at the myofibrillar aggregates. (B) TgWT localizes to the Z-disc and occasionally accumulates near the sarcolemmal membrane. (C) No background staining is observed in control littermate muscle. (D–F) A myotilin peptide antibody cross-reacts with both the human transgene product and murine myotilin. The myotilin antibody decorates the Z-disc and aggregates in TgT57I mice (D) and the Z-discs of both TgWT and control mice (E and F). ACTN2 (G–I), FLNC (J–L) and desmin (M–O) also localize to the Z-disc in each muscle shown and to the aggregates of TgT57I mice. (P) A mono-ubiquitin antibody stains the aggregates of TgT57I muscle. Scale bars: 10 µm (A–C); 20 µm (D–R).

 
In order to quantitate the level of accumulation of proteins found in the aggregates, we conducted immunoblot analysis of the insoluble fractions of muscle lysates from 12-month-old mice. The aggregates are not represented in the soluble fractions shown in Fig. 1. We specifically examined the soleus, which is spared, and the quadriceps muscles, which are severely affected. Soleus muscle shows minimal accumulation of the TgT57I product, whereas this protein is present at high levels in the quadriceps (Fig. 6). This is most likely a reflection of the difference in aggregate burden between the two muscles. FLNC and desmin levels are increased in the quadriceps, but not in the soleus, whereas ACTN2 levels are unchanged (Fig. 6). These differences reflect the levels of these proteins in the myofibrillar aggregates (Fig. 5).


Figure 1606
View larger version (55K):
[in this window]
[in a new window]
  		Figure 6. Immunoblot analysis of myofibrillar fractions of myotilin transgenic muscle lysates. The insoluble fractions of soleus and quadriceps muscle lysates were prepared from 12-month-old +/TgT57I, +/TgWT and control littermate (+/+) mice. (A) c-myc immunoreactivity shows the relative abundance of myotilin transgene products. TgWT abundance is greater than TgT57I in quadriceps. The myotilin antibody (MYOT) shows 9- and 21-fold greater myotilin abundance in TgT57I and TgWT quadriceps, respectively, compared with the control. Immunoblot analysis shows comparable levels of ACTN2, FLNC and desmin (DES) in the soleus muscles of TgT57I, TgWT and control mice. ACTN2 is also present at similar levels in the quadriceps muscles of transgenic and control mice. FLNC and desmin abundance, however, are higher in myotilin transgenic quadriceps. Desmin levels are significantly elevated in TgT57I quadriceps. (B) Coomassie blue staining of myosin and actin bands confirms comparable loading of samples.

 
Maintenance of sarcolemmal integrity in myotilin transgenic mice
Vital staining with Evans blue dye (EBD) was performed to evaluate plasma membrane permeability in myotilin transgenic mice. EBD is a small molecular weight tracer that binds to serum albumin; the EBD–albumin complex can easily pass into the sarcoplasm of myofibers with a damaged sarcolemma. Macroscopic evaluation of forelimbs and hindlimbs showed little EBD uptake in either 12-month-old TgWT mice and control littermate or 9-month-old TgT57I mice (Fig. 7). EBD uptake results in red fluorescence of myofiber sarcoplasm when viewed under a microscope with a rhodamine filter. Cross-sections of quadriceps muscle from TgT57I, TgWT and littermate control mice show little fluorescence. In contrast, large amounts of internalized EBD is evident by macroscopic and fluorescent evaluations in dystrophin-deficient mdx muscle, a positive control for sarcolemmal damage. Similar patterns of EBD uptake in mdx limbs have been shown previously (27,28). These data indicate that the sub-sarcolemmal mini-aggregates do not disrupt sarcolemmal integrity in TgWT muscle. Moreover, maintenance of sarcolemmal integrity in TgT57I muscle further distinguishes this mouse model from others that have been associated with membrane defects.


Figure 1607
View larger version (62K):
[in this window]
[in a new window]
  		Figure 7. Examination of sarolemmal integrity by vital staining with EBD. Muscle was evaluated 18 h after intraperitoneal injection with EBD. TgWT and control littermate (+/+) mice were 12-months old; TgT57I mice were 9-months old; mdx mice were 2.5-months old. The mdx mouse serves as a postive control for sarcolemmal damage. (A–D) Macroscopic analysis shows minimal EBD uptake in wild-type, TgWT and TgT57I forelimbs. EBD uptake occurs in the triceps of mdx mice. (E–H) Macroscopic analysis of hindlimbs also reveals minimal EBD uptake in control littermate, TgWT and TgT57I mice. EBD uptake is prominent in mdx quadriceps femoris and gastrocnemius. (I–L) EBD vital-stained quadriceps were prepared for cryotomy. Seven micrometer cross-sections were analyzed by fluorescence microscopy with a rhodamine filter. Dye uptake results in bright red emission in muscle cytoplasm. Dye uptake is limited to mdx muscle by fluorescent microscopy.

 
TgT57I mice exhibit contractile dysfunction in selective muscle groups
Significant physiological dysfunction was detected in whole, intact EDL muscles of TgT57I mice. TgT57I EDL muscle mass is reduced by 33% compared with littermate controls (8.8 mg in TgT57I, 13.2 mg in controls, Fig. 8). Similarly, cross-sectional area is reduced by 30% (1.52 mm2 in TgT57I, 2.16 mm2 in controls), and specific maximum force is reduced by 24% (153 kN/mm2 in TgT57I, 200 kN/mm2 in controls, Fig. 8). This reduced specific force and reduced muscle size combine to lower the maximum isometric force generated by the EDL by 46% (230 mN in TgT57I, 426 mN in controls). In striking contrast, overexpression of wild-type human myotilin does not cause such a contractile dysfunction. TgWT EDL and soleus muscles are statistically indistinguishable from littermate controls with respect to mass, mean cross-sectional area and specific maximum force (Fig. 8). Intriguingly, TgT57I soleus mass, cross-sectional area and specific maximum force are comparable with transgene-negative littermate controls (Fig. 8). TgT57I diaphragm also generates specific maximum force similar to controls. Thus, the TgT57I mouse model displays the same variable presentation of different muscle groups that is seen in the human myotilinopathies.


Figure 1608
View larger version (26K):
[in this window]
[in a new window]
  		Figure 8. Physiological analysis of TgWT and TgT57I muscles. Whole EDL, soleus (SOL) and diaphragm (DI) muscles were dissected from 8–10-month-old male TgWT and littermate control mice [+/+ (I)] and from 6–8-month-old male TgT57I and littermate control mice [+/+ (II)]. (A) TgT57I EDL muscle's mass is reduced compared with TgWT and wild-type muscle (P=0.001). Soleus muscles from TgWT and TgT57I mice show no differences in mass. (B) The cross-sectional area of TgT57I EDL muscles is also smaller (P=0.0005). (C) The maximum force generated by whole EDL muscles from TgT57I mice is lower than TgWT and littermate controls (P=0.001). The maximum force of TgWT and TgT57I soleus muscles are similar to controls. (D) The specific maximum force of EDL muscles from TgT57I mice is also decreased (P=0.044). (E) Muscles were subject to lengthening contractions (2 x 30%) to determine force deficit, a function of contraction-induced injury. TgT57I EDL (P=0.007) and diaphragm (P=0.0006) show reduced force deficit values (i.e. resistance to injury), following lengthening contractions. Asterisks denote statistically significant differences (P<0.05) between transgenic data sets and the respective littermate control data sets. P-values were determined by two-tailed t-test for two samples, assuming equal variance.

 
EDL and diaphragm muscles from TgT57I mice are less vulnerable to contraction-induced injury than those from littermate controls. Susceptibility to injury is measured by stimulating a muscle to contract while simultaneously stretching it. Maximum isometric force generation before and after the stretch is used to calculate a force deficit—muscles susceptible to damage will show a large force deficit. After a lengthening contraction protocol, EDL and diaphragm of TgT57I show less of a force deficit than muscles from littermate controls (Fig. 8). Lengthening contraction-induced force deficit values for TgWT EDL and soleus are statistically indistinguishable from littermate controls.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
We have generated a transgenic mouse model that successfully recapitulates the pathological features of human myotilinopathies. The myotilinopathies, including LGMD1A, MFM and SBM, comprise an autosomal dominant class of muscle diseases, all caused by missense mutations in exon 2 of the myotilin gene. Despite the proximity and similarity of these mutations, the reported clinical features of LGMD1A, MFM and SBM are somewhat divergent. LGMD1A patients typically present with proximal muscle weakness, whereas distal muscle weakness is common in myotilin-related MFM (14). The defining features of MFM and SBM are dense sarcoplasmic bodies, yet this pathology has not yet been reported in LGMD1A. However, the transgenic mouse model we have presented here unites these disparate phenotypes. The TgT57I mouse displays expansive myofibrillar aggregation very similar to the hallmark pathologies observed in MFM and SBM. At the same time, these mice display abundant autophagic vesicles and Z-disc streaming that are characteristic of LGMD1A patients. This strongly suggests that the T57I myotilin mutation can induce all of the skeletal muscle pathologies observed in the human myotilinopathies and raises the possibility that variations in the presentation of human patients are the result of modifier loci.

The myotilin T57I mutation promotes myofibrillar aggregation
The myofibrillar aggregates and Z-disc streaming found in our TgT57I mouse model reflect dramatic disruptions in muscle structure—it is surprising that such profound changes can arise from a seemingly conservative amino acid replacement in the myotilin protein. We propose two potential mechanisms: first, mutant myotilin could act directly to nucleate protein aggregates, and second, mutant myotilin could act indirectly by disrupting actin tethering at the Z-disc.

In the direct model, myotilin mutations could nucleate myofibrillar aggregation by promoting heightened intermolecular associations. Myotilin binds promiscuously, homodimerizing as well as interacting with multiple muscle proteins. Many of these interactions are mediated by its two C-terminal IgL domains. Proteins containing tandem or serially arrayed IgL domains have previously been shown to be aggregate-prone (29). Myotilin mutations could promote oligomeric self-association, leading to pathogenic aggregation. This kind of single-molecule-dependent nucleation and aggregation model is supported by in vitro cross-linking experiments in FLNC-related MFM (16). The FLNC W2710X mutation causes a truncation at the C-terminal IgL domain 24, a domain that mediates dimerization of the wild-type protein. The truncation may promote misfolding and increase intra-molecular association of FLNC IgL domains, as could also be the case for myotilin mutations. Myotilin mutations could also promote inter-molecular aggregation with other Z-disc proteins that harbor similar IgL domains, including palladin, myopalladin and titin (30). The abundance of IgL domain-containing proteins at the Z-disc may well mediate rapid association of proteins to promote Z-disc assembly. In the TgT57I mouse, it does not appear that Z-disc assembly is adversely affected, as aggregates are not observed in younger mice. Direct nucleation mechanisms such as these imply that the missense substitution in the N-terminal domain of myotilin is able to alter the binding properties of the C-terminal IgL domains.

Alternatively, myotilin mutations could indirectly lead to myofibrillar aggregation by destabilizing actin tethering at the Z-disc. The myofiber Z-disc houses a dense network of proteins that anchor actin filaments, thus facilitating the production and transmission of actin–myosin-initiated force. The chief F-actin cross-linking protein of the Z-disc is {alpha}-actinin, which interacts with myotilin (8) and a number of other Z-disc proteins, including nebulin (31), the {alpha}-actinin-associated LIM-protein (32), the calsarcins (33), titin (34,35), ZASP (36) and myopalladin (30). Myotilin also associates directly with F-actin, thereby bundling and stabilizing actin filaments. Through possible combinatorial interactions with additional actin and alpha-actinin binding proteins that localize to the Z-disc (i.e. ACTN2, FLNC, calsarcin-2), myotilin may be even more intimately involved with actin stabilization and Z-disc function. Clearly, myotilin mutations could easily interfere with the total actin-tethering capacity of the Z-disc. This indirect mechanism of aggregate formation is strongly supported by the pathology in TgT57I muscle. The spectrum of Z-disc pathology, from discrete broadening to streaming to myofibrillar aggregation, suggests that initial Z-disc destabilization leads to the stochastic increase in protein aggregation. In vitro work in COS-7 cells suggests that the T57I mutation has no effect on actin binding or bundling, nor does it induce formation of aggregates (12). The dramatic effect of this T57I mutation in live, contracting murine muscle suggests that additional functional testing of mutant myotilin is required.

TgT57I muscle pathophysiology is muscle group dependent
In the TgT57I mouse, muscle groups with differences in fiber composition displayed significant variation in pathological involvement. The skeletal muscles of adult wild-type mice are composed of three different fiber types, one slow type I fiber type and two fast type II fiber types, IIA and IIB, also termed slow-oxidative, fast oxidative-glycolytic and fast glycolytic fibers (37). The three fiber types have unique physiological characteristics and genomic expression profiles (37,38). The EDL muscle of the mouse is composed completely of fast fibers with ~87% fast type IIB fibers (39). Gastrocnemius also contains predominantly type IIB fibers (79%) (39). Adult murine diaphragm muscle largely contains type IIA fibers, totaling ~78% (39). In contrast, the soleus muscle is composed of approximately equal percentages of slow type I and fast type IIA fibers (39). These ratios are not significantly altered in the TgT57I mouse (data not shown). The EDL and gastrocnemius muscles of the TgT57I mouse are heavily populated with myofibrillar aggregates, and the EDL displays physiological deficits. The soleus and diaphragm muscles, however, are completely spared of any pathology or physiological deficits. The fast type IIB fibers prevalent in the affected EDL and gastrocnemius muscles have few mitochondria, and consequently fatigue more rapidly than either of the other two fiber types. In addition, fast type II fibers have thinner Z-discs than slow type I fibers (40). Type II fibers have the highest velocity of shortening and are the most powerful fibers per unit mass (41–43), but the difference in specific forces developed by type IIB fibers is not sufficient to account for the increased pathological involvement. Despite this, the predominance of fast type IIB fibers in the EDL muscle may be responsible for the severe contractile dysfunction observed as a result of myotilin mutation.

The phenotypic differences between TgT57I muscles could be explained by fiber type- and muscle-specific gene expression profiles. Hierarchical clustering analysis of gene expression patterns can be used to easily distinguish EDL and soleus muscles of the mouse (44). The presence or absence of a specific protein in fast type IIB fibers in muscles of TgT57I mice may increase their risk to damage. For example, {alpha}-actinin-3 (ACTN3), a structural protein of the Z-disc, is expressed exclusively in fast type IIB fibers (45).

A major difference between fast type II and slow type I fibers and muscles is their susceptibility to contraction-induced injury. The fast type II fibers, and muscles largely composed of these fibers, display a three- to four-fold greater force deficit for a given strain or for a given amount of work done to stretch a fiber (46,47). Surprisingly, the EDL and diaphragm muscles in the TgT57I mice do not display the high force deficits expected. Similar observations of low values for maximum force and for force deficits have been made in fatigued muscles and in the muscles of desmin-deficient mice (48,49). This resistance to contraction-induced injury may be secondary to the reduced force-generating capacity of the EDL muscle. This interpretation is supported further by the data on the soleus muscles of TgT57I mice compared with littermate controls, which did not differ in either force-generating capacity or susceptibility to contraction-induced injury. The magnitude of the force deficit correlates highly with the work required to stretch the muscle, which is the product of the displacement and the average force developed by the muscle during the strain (47,50,51). Consequently, the EDL muscle of the TgT57I mouse may have a very low force deficit because it is damaged and only able to generate reduced force. The same explanation may hold true for the reduced force deficit in the diaphragm, although the decrease in specific force for this muscle does not reach the level of statistical significance.

The response to lengthening contractions illustrates that myotilin mutations give rise to a dramatically different myopathy than mutations in the dystrophin-associated glycoprotein (DAG) complex. The muscles of mdx mice display a significant reduction in specific force (52); however, they are highly sensitive to contraction-induced injury (53). With the absence of dystrophin and the DAG complex in the mdx mouse, the large force deficits of the EDL and soleus muscles arise from disruptions of the sarcolemma and of mechanical properties of the cytoskeleton (47). Myotilin mutations do not interfere with the link between the DAG complex and the contractile elements. Consequently, TgT57I muscles are not exposed to this acute injury response to lengthening contractions. Rather, the myotilin mutations directly impair the development of force. Further investigations of the contractility and susceptibility to contraction-induced damage in the TgT57I mouse model are necessary for a more complete understanding of the pathophysiology of mutations in Z-disc proteins. In particular, physiological measurements of single fast type IIB fibers may shed additional light on their role in the myotilinopathies.

TgT57I mouse complements current MFM models
Mutations in multiple genes other than myotilin have been shown to cause MFM—mouse models incorporating several of these genes have been reported. Adult desmin knock-out mice show signs of muscle degeneration, Z-disc streaming and myofibril misalignment, but myofibrillar aggregation does not occur (54,55). In contrast to normal TgT57I diaphragm and soleus muscles, desmin-null diaphragm exhibits increased degeneration, and desmin-null soleus muscle generates 90% less maximum isometric force (54). These pathophysiological differences can be explained both by desmin's additional role at myofiber costameres, structures that connect Z-discs to the sarcolemma, and by the approach of targeted inactivation of desmin, rather than transgenic expression. Indeed, transgenic expression of the MFM desmin allele, D7-des (deletion of seven amino acids, 173–179), causes intrasarcoplasmic aggregation in cardiac myocytes. Cardiac-restricted expression by the {alpha}-myosin heavy chain promoter prevents the opportunity to study skeletal myopathy in this model (56).

Targeted inactivation of another MFM gene, {alpha}B-crystallin (CRYAB), causes non-specific histological features of muscular dystrophy and small patches of amorphous granules in older mice, but no significant myofilbrillar aggregation (57). In the same way as the transgenic desmin allele, cardiac-restricted transgenic expression of the CRYAB R120G MFM mutation causes eosinophilic aggregation in cardiac myocytes (58). Targeted inactivation of cypher, the murine ortholog of ZASP, causes perinatal lethality, right and left ventricular dilation and disorganization of skeletal muscle Z-discs, and no aggregation is observed (59). The desmin, CRYAB and ZASP mouse mutants have proven very successful in the study of cardiac failure associated with MFM. The transgenic myotilin T57I mouse, however, best captures the skeletal muscle phenotype of MFM. The desmin/CRYAB and myotilin transgenic mouse models complement each other well in the effort to identify the causes and functional impacts of myofibrillar aggregation in cardiac and skeletal muscles.

Therapeutic potential of myotilin transgenic mouse models
Analysis of our TgWT mouse demonstrates that in vivo expression of high levels of wild-type myotilin protein results in small sub-sarcolemmal aggregates and slight accumulation of FLNC and desmin in TgWT mice (Figs 5 and 6). However, these minor abnormalities do not induce any changes in sarcolemmal integrity (Fig. 7) or physiological function of the muscle (Fig. 8). The lack of toxicity of high levels of myotilin protein raises the possibility that myotilin overexpression could be used therapeutically to treat muscle diseases caused by sarcomeric dysfunction, especially those characterized by destabilization of F-actin tethering. A preliminary test of this hypothesis is currently underway by crossing the TgWT and TgT57I lines. The TgWT mouse serves as a promising genetic resource for testing the therapeutic effects of myotilin overexpression in other established mouse models.

The TgT57I mouse also provides a useful system to test disease therapeutics such as RNA-directed approaches to correct the mutant myotilin transcript or the oral administration of anti-aggregation drugs. Doxycycline and trehalose have proven effective in reducing nuclear aggregation and muscle weakness in a mouse model of oculopharyngeal muscular dystrophy (60,61), and rapamycin has also been shown to have potent anti-aggregation activity in vivo (62).

We have generated an excellent murine model that unites the pathological and physiological phenotypes of the human myotilinopathies. This mouse model recapitulates many of the features of human patients, including the selective pattern of muscle group weakness. Further characterization of the TgT57I mouse promises to reveal the mechanistic roots and impacts of myofibrillar aggregation in muscle disease and allows for the development and testing of therapeutic approaches to all the myotilinopathies.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
Generation and characterization of myotilin transgenic mice
The wild-type myotilin cDNA was isolated from a pTriplEx human skeletal muscle cDNA library (BD Clontech) and cloned into a modified pBluescript vector (Stratagene) containing an enhanced HSA promoter cassette and two SV40 polyadenylation signals. This HSA promoter fragment, –2139 to +239, has previously been shown to drive skeletal muscle-specific expression (63), as well as the enhanced HSA cassette (HSA-VP1) containing the 400 bp SV40 VP1 intron (64–66). The myotilin cDNA includes the 1530 bp coding region, 281 bp of 5'-UTR and 485 bp of 3'-UTR. Recombinant PCR was used to incorporate the c-myc epitope tag and the T57I point mutation; sub-cloning was facilitated by EcoRI and SphI double digestion of recombinant products and subsequent ligation into the wild-type clone. The 5.9 kb linear transgenic constructs were released by KpnI/NaeI digestion, purified with gel extraction columns (QiaGen), sterilized by ethanol precipitation and microinjected into B6SJLF2 one-cell embryos at the Duke University Transgenic Mouse Facility. Transgene-positive founder mice (F0, also B6SJLF2) were subsequently backcrossed to C57BL/6. Data reported in this manuscript were gathered from progeny that were backcrossed at least five times to C57BL/6 [N5(B6SJLF2XB6)], with the exception of F2N3 animals used to assess TgWT and control littermate muscle physiology. Because the SJL strain develops myopathy owing to a null mutation in the dysferlin gene (67), homozygosity of the wild-type C57BL/6 allele was tested and confirmed in all transgenic F2N1 breeders. Myotilin transgene genotyping was done by PCR amplification of a 394 bp fragment from tail clip DNA with a forward HSA-VP1 primer (5'-GCTCCTGTTAATTGGTATAAC-3') and a reverse myotilin 5'-UTR primer (5'-AACCCACTATTGAAGGGAAG-3'). C57BL/6 breeders were purchased from Harlan. Male C57BL/10ScSn-Dmdmdx/J mice were purchased from The Jackson Laboratory. The mice were bred under standard conditions in the Genome Sciences Research Building II Animal Facility at Duke University.

Analysis of myotilin transgene expression
Total RNA was prepared from dissected and homogenized brain, kidney, heart, liver, spinal cord and quadriceps tissues (SV Total RNA Isolation System, Promega). For RT–PCR, 1 µg of total RNA was reverse transcribed using random hexamers (Promega). A transgene-specific 481 bp product was amplified with a forward HSA exon 1 primer (5'-GAGTAGCAGTTGTAGCTACC-3') and a reverse myotilin exon 2 primer (5'-AGGAGGCTGCAATCTGGAGC-3'). Muscle tissue for downstream western blotting was snap-frozen in liquid nitrogen, ground in a mortar and pestle and resuspended (200 µl/100 mg tissue) in 1% SDS, supplemented with a mammalian protease inhibitor cocktail (Sigma). For crude soluble fraction isolation, lysates were vortexed for 30 s, boiled for 5 min, vortexed for 30 s, boiled for 60 s and then spun at 12 000g for 60 s. The supernatant is the soluble fraction. To obtain the insoluble fraction, muscle lysates were spun at 12 000g for 20 min. The pellet was resuspended in 1x SDS buffer (62.5 mM Tris pH 8.8, 6% SDS, 50 mM DTT), vortexed for 1 min, boiled for 5 min, vortexed for 1 min and then further resuspended with an electric homogenizer. Prior to electrophoresis, aliquots of lysates were mixed 1:1 with 2x Laemmli buffer, boiled for 1 min, spun for 60 s and loaded onto denaturing 4–15% polyacrylamide Tris–HCl gels (Bio-Rad). Proteins were separated at 15 mA for 3 h and mobilized by tank transfer (40 V for 4 h) to a 0.45 µm nitrocellulose membrane (Bio-Rad). Membranes were blocked with 5% non-fat dry milk + 1% donkey serum + 0.1% Tween 20 in TBS and stained with primary antibody for 60 min at room temperature in blocking buffer. Membranes were washed 1x15 min, then 3x5 min each in TBS + 0.1% Tween 20 (TBST), stained with horseradish peroxidase-conjugated goat anti-rabbit secondary antibody (1:10 000, Jackson Immunoresearch) for 45 min at room temperature in blocking buffer and washed 1x15 min and then 3x5 min each in TBST. Immunodetection was carried out with the enhanced chemiluminescence (ECL) Plus Western Blotting Detection System (Amersham Biosciences). Primary antibodies for western blot included myotilin peptide antibody (1:4000, rabbit polyclonal, generated by Bethyl Laboratories, using the peptide corresponding to myotilin residues 473–488, CVKQAFNPEGEFQRLAAQ); anti-cmyc (1:10 000, rabbit polyclonal, RDI Division of Fitzgerald Industries); anti-ACTN (sarcomeric), clone EA-53 (1:4000, Sigma); rabbit anti-FLNC-A1 (1:1500, L. Kunkel) (68); anti-desmin, clone DE-R-11 (1:400, Novocastra). Optical scanning densitometry was performed with Image-Pro Plus software (MediaCybernetics).

Histology
Mice were transcardially perfusion-fixed with 4% paraformaldehyde in PBS. Dissected muscle was then fixed at least overnight at room temperature, also in 4% paraformaldehyde in PBS. Tissue was then dehydrated in 70% ethanol and processed via manufacturer's instructions in an automatic processor (Tissue-Tek VIP). Specimens were embedded in paraffin, sectioned at 7 µm and stained with Masson's trichrome stain (69).

Electron microscopy
Mice were transcardially perfusion-fixed with 4% paraformaldehyde and 0.25% glutaraldehyde fixative in HEPES buffer. Whole muscles were dissected, diced into 1–2 mm3 cubes and further fixed with 4% glutaraldehyde in 0.1 M cacodylate buffer for at least 1 h. They were washed in 0.1 M cacodylate buffer containing 7.5% sucrose, 3x20 min each wash. Specimens were then post-fixed in 1% osmium tetroxide in cacodylate buffer, dehydrated through a graded series of ethanol, followed by propylene oxide and embedded in PolyBed 812 epoxy resin (Polysciences). Specimens were baked overnight at 60°C. Semi-thin sections (0.5 µm) were cut and stained with toluidine blue. Ultrathin sections (60–90 nm) were cut with a diamond knife and post-stained with uranyl acetate and lead citrate. Sections were examined in a Philips EM 400 or CM 12 electron microscope.

Immunofluorescence
All immunofluorescence experiments were performed on fresh, frozen muscle sections. Muscle specimens were saturated overnight at 4°C in PBS + 20% sucrose. Tissue was then embedded in optimal cutting temperature (OCT) compound (Tissue-Tek) over liquid nitrogen-cooled isopentane. Frozen sections were cut at 7 µm, washed for 15 min in PBS, blocked for 60 min in 2% bovine serum albumin (BSA), washed 3x10 min each in PBS and incubated for 60 min at room temperature with primary antibody diluted in PBS + 1% BSA + 1% goat serum (Sigma). Sections were washed 3x10 min each with PBS, stained for 60 min at room temperature with the appropriate fluorophore-conjugated secondary antibody diluted in PBS + 1% BSA, washed 3x10 min each in PBS and mounted with ProLong Gold antifade reagent (Invitrogen). The following primary antibodies were used for immunofluorescence: Cy3-conjugated anti-cmyc, clone 9E10 (1:200, Sigma); myotilin peptide antibody (1:800); anti-ACTN (sarcomeric), clone EA-53 (1:500, Sigma); rabbit anti-FLNC-A2 (1:200, L. Kunkel) (68); anti-desmin, clone DE-R-11 (1:50, Novocastra); anti-myosin (FAST), clone MY-32 (1:400, Sigma); anti-myosin (SLOW), clone NOQ7.5.4D (1:400, Sigma) and rabbit anti-ubiquitin (1:80, Sigma). Secondary antibodies included Alexa Fluor 488- and Alexa Fluor 594-conjugated chicken anti-rabbit, goat anti-rabbit and rabbit anti-mouse IgG antibodies (1:1000, Invitrogen).

EBD uptake assay
The mice were administered an intraperitoneal injection of EBD (Sigma) (1 mg/10 g body weight). After 16–18 h, mice were euthanized and skinned. Whole limbs were fixed in 4% paraformaldehyde for 6 h and examined macroscopically for dye uptake. For microscopic analysis, fresh quadriceps muscles were embedded in OCT over liquid nitrogen-cooled isopentane. Seven micrometer sections were cut and examined for dye uptake by fluorescence microscopy, using a rhodamine filter.

Muscle physiology
EDL and soleus muscles were isolated, and the nerves to the muscles were dissected and cut. Ties were placed around the proximal and distal tendons of the muscle, and the muscle was removed. The tissue was placed in buffered mammalian Ringer's solution. The solution was maintained at 25°C and gassed continuously with a mixture of 95% O2 and 5% CO2. One tendon was tied to a servo-motor (Aurora Scientific Inc.) and the other to a force transducer (Kulite model BG-50). The muscle was stimulated directly by an electrical field generated between two large platinum electrodes situated in the bath on either side of the muscle. Square-wave pulses 0.2 ms in duration were amplified to increase current intensity to a sufficient level to produce a maximum response. Muscle length was adjusted for maximum isometric twitch force. The entire diaphragm, including adjacent ribs, was also removed and immersed in an oxygenated bath containing mammalian Ringer's solution at 25°C. Diaphragm strips, 1–2 mm wide, were carefully dissected from the central region of the lateral costal hemi-diaphragm. The connections of the strip of fibers to a small section of a single rib and part of the central tendon were preserved. The rib and the tendon were attached to the force transducer and servo-motor. Diaphragm strips were stimulated directly by an electric field, as described for EDL and soleus. To measure force deficits following lengthening contractions, two stretches of magnitude 0.3 fiber-lengths and velocity 0.5 fiber-lengths/s were initiated from the plateau of an isometric contraction. Lengthening ramps were separated by 10 s and stimulation was terminated at the end of the second lengthening ramp. Force deficit was assessed during a final isometric tetanic contraction 1 min after the second lengthening contraction ended. Further details on these physiological protocols can be found elsewhere (70–73).


   ACKNOWLEDGEMENTS
 
We would like to thank Dr Gregory Cox for many helpful discussions and initial review of the manuscript, as well as the laboratory of Dr Fred Schachat for analysis of myosin isoform expression. FLNC antibodies were a kind gift from Dr Louis Kunkel. This work was supported by grants from the Muscular Dystrophy Association (M.A.H.), Charles E. Culpepper Biomedical Pilot Initiative (M.A.H.) and the NIH/NINDS NS026630 (M.A.H.). S.M.G. was supported by a Ruth L. Kirschstein National Research Service Award pre-doctoral fellowship NIH/NINDS NS047910.

Conflict of Interest statement. The authors report no conflict of interest.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 

  1. Piccolo F., Moore S.A., Mathews K.D., Campbell K.P. (2002) Limb-girdle muscular dystrophies. Adv. Neurol. 88:273–291.[Medline]

  2. Guglieri M., Magri F., Comi G.P. (2005) Molecular etiopathogenesis of limb girdle muscular and congenital muscular dystrophies: boundaries and contiguities. Clin. Chim. Acta 361:54–79.

  3. Gilchrist J.M., Pericak-Vance M.A., Silverman L., Roses A.D. (1988) Clinical and genetic investigation in autosomal dominant limb-girdle muscular dystrophy. Neurology 38:5–8.[Abstract/Free Full Text]

  4. Hauser M.A., Horrigan S.K., Salmikangas P., Torian U.M., Viles K.D., Dancel R., Tim R.W., Taivainen A., Bartoloni L., Gilchrist J.M., et al. (2000) Myotilin is mutated in limb girdle muscular dystrophy 1A. Hum. Mol. Genet. 9:2141–2147.[Abstract/Free Full Text]

  5. Speer M.C., Gilchrist J.M., Stajich J.M., Gaskell P.C., Westbrook C.A., Horrigan S.K., Bartoloni L., Yamaoka L.H., Scott W.K., Pericak-Vance M.A. (1998) Evidence for anticipation in autosomal dominant limb-girdle muscular dystrophy. J. Med. Genet. 35:305–308.[Abstract/Free Full Text]

  6. Hauser M.A., Conde C.B., Kowaljow V., Zeppa G., Taratuto A.L., Torian U.M., Vance J., Pericak-Vance M.A., Speer M.C., Rosa A.L. (2002) Myotilin mutation found in second pedigree with LGMD1A. Am. J. Hum. Genet. 71:1428–1432.[CrossRef][Web of Science][Medline]

  7. Faulkner G., Lanfranchi G., Valle G. (2001) Telethonin and other new proteins of the Z-disc of skeletal muscle. IUBMB Life 51:275–282.[Web of Science][Medline]

  8. Salmikangas P., Mykkänen O.M., Grönholm M., Heiska L., Kere J., Carpén O. (1999) Myotilin, a novel sarcomeric protein with two Ig-like domains, is encoded by a candidate gene for limb-girdle muscular dystrophy. Hum. Mol. Genet. 8:1329–1336.[Abstract/Free Full Text]

  9. van der Ven P.F.M., Wiesner S., Salmikangas P., Auerbach D., Himmel M., Kempa S., Hayess K., Pacholsky D., Taivainen A., Schröder R., et al. (2000) Indications for a novel muscular dystrophy pathway: {gamma}-filamin, the muscle-specific filamin isoform, interacts with myotilin. J. Cell Biol. 151:235–247.[Abstract/Free Full Text]

  10. Salmikangas P., Van Der Ven P.F., Lalowski M., Taivainen A., Zhao F., Suila H., Schroder R., Lappalainen P., Furst D.O., Carpen O. (2003) Myotilin, the limb-girdle muscular dystrophy 1A (LGMD1A) protein, cross-links actin filaments and controls sarcomere assembly. Hum. Mol. Genet. 12:189–203.[Abstract/Free Full Text]

  11. Gontier Y., Taivainen A., Fontao L., Sonnenberg A., van der Flier F.A., Carpen O., Faulkner G., Borradori L. (2005) The Z-disc proteins myotilin and FATZ-1 interact with each other and are connected to the sarcolemma via muscle-specific filamins. J. Cell Sci. 118:3739–3749.[Abstract/Free Full Text]

  12. von Nandelstadh P., Gronholm M., Moza M., Lamberg A., Savilahti H., Carpen O. (2005) Actin-organising properties of the muscular dystrophy protein myotilin. Exp. Cell Res. 310:131–139.[CrossRef][Web of Science][Medline]

  13. Foroud T., Pankratz N., Batchman A.P., Pauciulo M.W., Vidal R., Miravalle L., Goebel H.H., Cushman L.J., Azzarelli B., Horak H., et al. (2005) A mutation in myotilin causes spheroid body myopathy. Neurology 65:1936–1940.[Abstract/Free Full Text]

  14. Selcen D. and Engel A.G. (2004) Mutations in myotilin cause myofibrillar myopathy. Neurology 62:1363–1371.[Abstract/Free Full Text]

  15. Olive M., Goldfarb L.G., Shatunov A., Fischer D., Ferrer I. (2005) Myotilinopathy: refining the clinical and myopathological phenotype. Brain 128:2315–2326.[Abstract/Free Full Text]

  16. Vorgerd M., van der Ven P.F., Bruchertseifer V., Lowe T., Kley R.A., Schroder R., Lochmuller H., Himmel M., Koehler K., Furst D.O., Huebner A. (2005) A mutation in the dimerization domain of filamin C causes a novel type of autosomal dominant myofibrillar myopathy. Am. J. Hum. Genet. 77:297–304.[CrossRef][Web of Science][Medline]

  17. Selcen D. and Engel A.G. (2005) Mutations in ZASP define a novel form of muscular dystrophy in humans. Ann. Neurol. 57:269–276.[CrossRef][Web of Science][Medline]

  18. Goldfarb L.G., Park K.Y., Cervenakova L., Gorokhova S., Lee H.S., Vasconcelos O., Nagle J.W., Semino-Mora C., Sivakumar K., Dalakas M.C. (1998) Missense mutations in desmin associated with familial cardiac and skeletal myopathy. Nat. Genet. 19:402–403.[CrossRef][Web of Science][Medline]

  19. Paulin D., Huet A., Khanamyrian L., Xue Z. (2004) Desminopathies in muscle disease. J. Pathol. 204:418–427.[CrossRef][Web of Science][Medline]

  20. Munoz-Marmol A.M., Strasser G., Isamat M., Coulombe P.A., Yang Y., Roca X., Vela E., Mate J.L., Coll J., Fernandez-Figueras M.T., et al. (1998) A dysfunctional desmin mutation in a patient with severe generalized myopathy. Proc. Natl Acad. Sci. USA 95:11312–11317.[Abstract/Free Full Text]

  21. Vicart P., Caron A., Guicheney P., Li Z., Prevost M.C., Faure A., Chateau D., Chapon F., Tome F., Dupret J.M., et al. (1998) A missense mutation in the alphaB-crystallin chaperone gene causes a desmin-related myopathy. Nat. Genet. 20:92–95.[CrossRef][Web of Science][Medline]

  22. Selcen D. and Engel A.G. (2003) Myofibrillar myopathy caused by novel dominant negative alpha B-crystallin mutations. Ann. Neurol. 54:804–810.[CrossRef][Web of Science][Medline]

  23. Agbulut O., Destombes J., Thiesson D., Butler-Browne G. (2000) Age-related appearance of tubular aggregates in the skeletal muscle of almost all male inbred mice. Histochem. Cell Biol. 114:477–481.[CrossRef][Web of Science][Medline]

  24. Goebel H.H., Muller J., Gillen H.W., Merritt A.D. (1978) Autosomal dominant ‘spheroid body myopathy’. Muscle Nerve 1:14–26.[CrossRef][Web of Science][Medline]

  25. Selcen D., Ohno K., Engel A.G. (2004) Myofibrillar myopathy: clinical, morphological and genetic studies in 63 patients. Brain 127:439–451.[Abstract/Free Full Text]

  26. Goebel H.H., D'Agostino A.N., Wilson J., Cole G., Foroud T., Koller D., Farlow M., Azzarelli B., Muller J. (1997) Spheroid body myopathy revisited. Muscle Nerve 20:1127–1136.[CrossRef][Web of Science][Medline]

  27. Matsuda R., Nishikawa A., Tanaka H. (1995) Visualization of dystrophic muscle fibers in mdx mouse by vital staining with Evans blue: evidence of apoptosis in dystrophin-deficient muscle. J. Biochem. (Tokyo) 118:959–964.[Abstract/Free Full Text]

  28. Straub V., Rafael J.A., Chamberlain J.S., Campbell K.P. (1997) Animal models for muscular dystrophy show different patterns of sarcolemmal disruption. J. Cell Biol. 139:375–385.[Abstract/Free Full Text]

  29. Wright C.F., Teichmann S.A., Clarke J., Dobson C.M. (2005) The importance of sequence diversity in the aggregation and evolution of proteins. Nature 438:878–881.[CrossRef][Medline]

  30. Bang M.L., Mudry R.E., McElhinny A.S., Trombitas K., Geach A.J., Yamasaki R., Sorimachi H., Granzier H., Gregorio C.C., Labeit S. (2001) Myopalladin, a novel 145-kilodalton sarcomeric protein with multiple roles in Z-disc and I-band protein assemblies. J. Cell Biol. 153:413–427.[Abstract/Free Full Text]

  31. Nave R., Furst D.O., Weber K. (1990) Interaction of alpha-actinin and nebulin in vitro. Support for the existence of a fourth filament system in skeletal muscle. FEBS Lett. 269:163–166.[CrossRef][Web of Science][Medline]

  32. Xia H., Winokur S.T., Kuo W.L., Altherr M.R., Bredt D.S. (1997) Actinin-associated LIM protein: identification of a domain interaction between PDZ and spectrin-like repeat motifs. J. Cell Biol. 139:507–515.[Abstract/Free Full Text]

  33. Faulkner G., Pallavicini A., Comelli A., Salamon M., Bortoletto G., Ievolella C., Trevisan S., Kojic S., Dalla V.F., Laveder P., et al. (2000) FATZ, a filamin-, actinin-, and telethonin-binding protein of the Z- disc of skeletal muscle. J. Biol. Chem. 275:41234–41242.[Abstract/Free Full Text]

  34. Ohtsuka H., Yajima H., Maruyama K., Kimura S. (1997) Binding of the N-terminal 63 kDa portion of connectin/titin to alpha-actinin as revealed by the yeast two-hybrid system. FEBS Lett. 401:65–67.[CrossRef][Web of Science][Medline]

  35. Sorimachi H., Freiburg A., Kolmerer B., Ishiura S., Stier G., Gregorio C.C., Labeit D., Linke W.A., Suzuki K., Labeit S. (1997) Tissue-specific expression and alpha-actinin binding properties of the Z-disc titin: implications for the nature of vertebrate Z-discs. J. Mol. Biol. 270:688–695.[CrossRef][Web of Science][Medline]

  36. Faulkner G., Pallavicini A., Formentin E., Comelli A., Ievolella C., Trevisan S., Bortoletto G., Scannapieco P., Salamon M., Mouly V., et al. (1999) ZASP: a new Z-band alternatively spliced PDZ-motif protein. J. Cell Biol. 146:465–475.[CrossRef][Web of Science][Medline]

  37. Burkholder T.J., Fingado B., Baron S., Lieber R.L. (1994) Relationship between muscle fiber types and sizes and muscle architectural properties in the mouse hindlimb. J. Morphol. 221:177–190.[CrossRef][Web of Science][Medline]

  38. Pette D. and Staron R.S. (2000) Myosin isoforms, muscle fiber types, and transitions. Microsc. Res. Tech. 50:500–509.[CrossRef][Web of Science][Medline]

  39. Agbulut O., Noirez P., Beaumont F., Butler-Browne G. (2003) Myosin heavy chain isoforms in postnatal muscle development of mice. Biol. Cell 95:399–406.[CrossRef][Web of Science][Medline]

  40. Eisenberg B.R. (1986) Quantitative ultrastructure of mammalian skeletal muscle. In Peachey L.D., Adrian R.H., Geiger S.R. (Eds.). Handbook of Physiology (American Physiological Society, Bethesda, MD) pp. pp. 73–112 Section 10.

  41. Bottinelli R., Schiaffino S., Reggiani C. (1991) Force–velocity relations and myosin heavy chain isoform compositions of skinned fibres from rat skeletal muscle. J. Physiol. 437:655–672.[Abstract/Free Full Text]

  42. Galler S., Schmitt T.L., Pette D. (1994) Stretch activation, unloaded shortening velocity, and myosin heavy chain isoforms of rat skeletal muscle fibres. J. Physiol 478:(Pt 3)513–521.[Abstract/Free Full Text]

  43. Larsson L. and Moss R.L. (1993) Maximum velocity of shortening in relation to myosin isoform composition in single fibres from human skeletal muscles. J. Physiol 472:595–614.[Abstract/Free Full Text]

  44. Haslett J.N., Kang P.B., Han M., Kho A.T., Sanoudou D., Volinski J.M., Beggs A.H., Kohane I.S., Kunkel L.M. (2005) The influence of muscle type and dystrophin deficiency on murine expression profiles. Mamm. Genome 16:739–748.[CrossRef][Web of Science][Medline]

  45. Mills M., Yang N., Weinberger R., Vander Woude D.L., Beggs A.H., Easteal S., North K. (2001) Differential expression of the actin-binding proteins, alpha-actinin-2 and -3, in different species: implications for the evolution of functional redundancy. Hum. Mol. Genet. 10:1335–1346.[Abstract/Free Full Text]

  46. Macpherson P.C., Schork M.A., Faulkner J.A. (1996) Contraction-induced injury to single fiber segments from fast and slow muscles of rats by single stretches. Am. J. Physiol. 271:C1438–C1446.

  47. Consolino C.M. and Brooks S.V. (2004) Susceptibility to sarcomere injury induced by single stretches of maximally activated muscles of mdx mice. J. Appl. Physiol. 96:633–638.[Abstract/Free Full Text]

  48. Sam M., Shah S., Friden J., Milner D.J., Capetanaki Y., Lieber R.L. (2000) Desmin knockout muscles generate lower stress and are less vulnerable to injury compared with wild-type muscles. Am. J. Physiol. Cell Physiol. 279:C1116–C1122.[Abstract/Free Full Text]

  49. McCully K.K. and Faulkner J.A. (1986) Characteristics of lengthening contractions associated with injury to skeletal muscle fibers. J. Appl. Physiol. 61:293–299.[Abstract/Free Full Text]

  50. Brooks S.V., Zerba E., Faulkner J.A. (1995) Injury to muscle fibers after single stretches of passive and maximally stimulated muscles in mice. J. Physiol. 488:459–469.[Abstract/Free Full Text]

  51. Brooks S.V. and Faulkner J.A. (1996) The magnitude of the initial injury induced by stretches of maximally activated muscle fibers of mice and rats increases in old age. J. Physiol. 497:573–580.[Abstract/Free Full Text]

  52. Lynch G.S., Hinkle R.T., Chamberlain J.S., Brooks S.V., Faulkner J.A. (2001) Force and power output of fast and slow skeletal muscles from mdx mice 6–28 months old. J. Physiol. 535:591–600.[Abstract/Free Full Text]

  53. Brooks S.V. (1998) Rapid recovery following contraction-induced injury to in situ skeletal muscle in mdx mice. J. Muscle Res. Cell Motil. 19:179–187.[CrossRef][Web of Science][Medline]

  54. Li Z., Mericskay M., Agbulut O., Butler-Browne G., Carlsson L., Thornell L.E., Babinet C., Paulin D. (1997) Desmin is essential for the tensile strength and integrity of myofibrils but not for myogenic commitment, differentiation, and fusion of skeletal muscle. J. Cell Biol. 139:129–144.[Abstract/Free Full Text]

  55. Milner D.J., Weitzer G., Tran D., Bradley A., Capetanaki Y. (1996) Disruption of muscle architecture and myocardial degeneration in mice lacking desmin. J. Cell Biol. 134:1255–1270.[Abstract/Free Full Text]

  56. Wang X., Osinska H., Dorn G.W., Nieman M., Lorenz J.N., Gerdes A.M., Witt S., Kimball T., Gulick J., Robbins J. (2001) Mouse model of desmin-related cardiomyopathy. Circulation 103:2402–2407.[Abstract/Free Full Text]

  57. Brady J.P., Garland D.L., Green D.E., Tamm E.R., Giblin F.J., Wawrousek E.F. (2001) AlphaB-crystallin in lens development and muscle integrity: a gene knockout approach. Invest. Ophth. Vis. Sci. 42:2924–2934.[Abstract/Free Full Text]

  58. Wang X., Osinska H., Klevitsky R., Gerdes A.M., Nieman M., Lorenz J., Hewett T., Robbins J. (2001) Expression of R120G-alphaB-crystallin causes aberrant desmin and alphaB-crystallin aggregation and cardiomyopathy in mice. Circ. Res. 89:84–91.[Abstract/Free Full Text]

  59. Zhou Q., Chu P.H., Huang C., Cheng C.F., Martone M.E., Knoll G., Shelton G.D., Evans S., Chen J. (2001) Ablation of Cypher, a PDZ-LIM domain Z-line protein, causes a severe form of congenital myopathy. J. Cell Biol. 155:605–612.[Abstract/Free Full Text]

  60. Davies J.E., Wang L., Garcia-Oroz L., Cook L.J., Vacher C., O'Donovan D.G., Rubinsztein D.C. (2005) Doxycycline attenuates and delays toxicity of the oculopharyngeal muscular dystrophy mutation in transgenic mice. Nat. Med. 11:672–677.[CrossRef][Web of Science][Medline]

  61. Davies J.E., Sarkar S., Rubinsztein D.C. (2006) Trehalose reduces aggregate formation and delays pathology in a transgenic mouse model of oculopharyngeal muscular dystrophy. Hum. Mol. Genet. 15:23–31.[Abstract/Free Full Text]

  62. Berger Z., Ravikumar B., Menzies F.M., Oroz L.G., Underwood B.R., Pangalos M.N., Schmitt I., Wullner U., Evert B.O., O'Kane C.J., Rubinsztein D.C. (2006) Rapamycin alleviates toxicity of different aggregate-prone proteins. Hum. Mol. Genet. 15:433–442.[Abstract/Free Full Text]

  63. Brennan K.J. and Hardeman E.C. (1993) Quantitative analysis of the human alpha-skeletal actin gene in transgenic mice. J. Biol. Chem. 268:719–725.[Abstract/Free Full Text]

  64. Crawford G.E., Faulkner J.A., Crosbie R.H., Campbell K.P., Froehner S.C., Chamberlain J.S. (2000) Assembly of the dystrophin-associated protein complex does not require the dystrophin COOH-terminal domain. J. Cell Biol. 150:1399–1410.[Abstract/Free Full Text]

  65. Wehling M., Spencer M.J., Tidball J.G. (2001) A nitric oxide synthase transgene ameliorates muscular dystrophy in mdx mice. J. Cell Biol. 155:123–131.[Abstract/Free Full Text]

  66. Spencer M.J., Guyon J.R., Sorimachi H., Potts A., Richard I., Herasse M., Chamberlain J., Dalkilic I., Kunkel L.M., Beckmann J.S. (2002) Stable expression of calpain 3 from a muscle transgene in vivo: immature muscle in transgenic mice suggests a role for calpain 3 in muscle maturation. Proc. Natl Acad. Sci. USA 99:8874–8879.[Abstract/Free Full Text]

  67. Bittner R.E., Anderson L.V., Burkhardt E., Bashir R., Vafiadaki E., Ivanova S., Raffelsberger T., Maerk I., Hoger H., Jung M., et al. (1999) Dysferlin deletion in SJL mice (SJL-Dysf) defines a natural model for limb girdle muscular dystrophy 2B [letter]. Nat. Genet. 23:141–142.[CrossRef][Web of Science][Medline]

  68. Thompson T.G., Chan Y.-M., Hack A.A., Brosius M., Rajala M., Lidov H.G.W., McNally E.M., Watkins S., Kunkel L.M. (2000) Filamin 2 (FLN2): a muscle-specific sarcoglycan interacting protein. J. Cell Biol. 148:115–126.[CrossRef][Web of Science][Medline]

  69. Sheehan D. and Harpchak B. (1980) Theory and Practice of Histotechnology 2nd edn. (C.V. Mosby Co, St Louis).

  70. Brooks S.V. and Faulkner J.A. (1988) Contractile properties of skeletal muscles from young, adult and aged mice. J. Physiol. 404:71–82.[Abstract/Free Full Text]

  71. Brooks S.V. and Faulkner J.A. (1990) Contraction-induced injury: recovery of skeletal muscles in young and old mice. Am. J. Physiol. 258:C436–C442.

  72. Brooks S.V. and Faulkner J.A. (1991) Maximum and sustained power of extensor digitorum longus muscles from young, adult, and old mice. J. Gerontol. 46:B28–B33.[Abstract]

  73. Faulkner J.A., Brooks S.V., Dennis R.G. (1998) In Morgan J. and Yarmush M.L. (Eds.). Methods in Molecular Medicine (Humana Press Inc, Totowa, NJ) pp. pp. 155–172.


Add to CiteULike CiteULike   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us    What's this?


This article has been cited by other articles:


Home page
 FASEB J.Home page
T. J. Cohen, T. Barrientos, Z. C. Hartman, S. M. Garvey, G. A. Cox, and T.-P. Yao
The deacetylase HDAC4 controls myocyte enhancing factor-2-dependent structural gene expression in response to neural activity
FASEB J, January 1, 2009; 23(1): 99 - 106.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
T. J. Cohen, D. S. Waddell, T. Barrientos, Z. Lu, G. Feng, G. A. Cox, S. C. Bodine, and T.-P. Yao
The Histone Deacetylase HDAC4 Connects Neural Activity to Muscle Transcriptional Reprogramming
J. Biol. Chem., November 16, 2007; 282(46): 33752 - 33759.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow FREE Full Text (PDF) Freely available
Right arrow All Versions of this Article:
15/15/2348    most recent
ddl160v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Add to My Personal Archive
Right arrow Download to citation manager
Right arrow Search for citing articles in:
ISI Web of Science (11)
Right arrowRequest Permissions
Google Scholar
Right arrow Articles by Garvey, S. M.
Right arrow Articles by Hauser, M. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Garvey, S. M.
Right arrow Articles by Hauser, M. A.
Social Bookmarking
Add to CiteULike   Add to Connotea   Add to Del.icio.us  
What's this?