1 Department of Molecular Sciences and 2 Department of Physiology, University of Tennessee Health Sciences Center, Memphis, Tennessee 38163
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ABSTRACT |
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The dystrophin-glycoprotein complex (DGC)
is a sarcolemmal complex whose defects cause muscular dystrophies. The
normal function of this complex is not clear. We have proposed that
this is a signal transduction complex, signaling normal interactions
with matrix laminin, and that the response is normal growth and
homeostasis. If so, the complex and its signaling should be altered in
other physiological states such as atrophy. The amount of some of the DGC proteins, including dystrophin, -dystroglycan, and
-sarcoglycan, is reduced significantly in rat skeletal muscle
atrophy induced by tenotomy. Furthermore, H-Ras, RhoA, and Cdc42
decrease in expression levels and activities in muscle atrophy. When
the small GTPases were assayed after laminin or
-dystroglycan
depletion, H-Ras, Rac1, and Cdc42 activities were reduced, suggesting a
physical linkage between the DGC and the GTPases. Dominant-negative
Cdc42, introduced with a retroviral vector, resulted in fibers that
appeared atrophic. These data support a putative role for the DGC in
transduction of mechanical signals in muscle.
syntrophin; sarcoglycan; dystroglycan; Cdc42; retrovirus; tenotomy
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INTRODUCTION |
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INTEGRINS AND
DYSTROGLYCAN are coexpressed in skeletal muscle and play critical
roles during skeletal muscle differentiation and development (36,
43). Dystroglycan links the extracellular matrix with the actin
cytoskeleton, and it exists as a noncovalently linked complex of -
and
-subunits (10, 20, 31, 61). In skeletal muscle
cells, dystroglycan forms a tight complex with dystrophin and
dystrophin-associated proteins, including
-,
-,
-, and
-sarcoglycans, and syntrophins (10, 32, 39, 46, 47, 53,
61). The integrity of the complex is essential for the viability
of muscle cells because disruption of the complex due to a defect in
dystrophin or any one of the sarcoglycans has been reported to cause
various forms of inherited muscular dystrophy (10, 39, 45, 47,
53, 61).
The dystrophin-glycoprotein complex (DGC) was unknown before 1990, and
its physiological role is not currently known. We first suggested that
the DGC protein complex was a signal transduction complex in 1992 (41). Since then, calmodulin kinase II (40), stress-activated protein kinase (SAPK) 3 (28), nitric
oxide synthetase (1, 24), voltage-gated Na+
channels (23), phosphatidylinositol 4,5-bisphosphate
(15), growth factor receptor-bound protein 2 (grb2)
(48), and other cell signaling components have all
been localized to this complex. The interaction of the -dystroglycan
cytoplasmic region with the SH3 domain of grb2, an
adaptor protein involved in signal transduction and
cytoskeletal organization, has been reported (64).
Cavaldesi et al. (13) found that grb2 mediates the
interaction of
-dystroglycan from brain synaptosomes with focal
adhesion kinase p125FAK, a nonreceptor tyrosine kinase that
also participates in the intracellular signaling pathways triggered by
integrins. There is now a general agreement that the complex probably
serves a role in cell signaling.
Dystroglycan binds to the muscle laminin merosin (29), which resembles fibronectin binding to initiate integrin signaling. There is a great similarity between what is known of the signaling proteins associated with the DGC and that of the integrin signaling, as pointed out by Yoshida et al. (65). Thus signaling cellular attachment to laminin might be the long-sought function of the DGC complex. A vast array of signaling molecules and cascades has been connected to integrin signaling, including p125FAK, protein kinase C, mitogen-activated protein kinase, phosphatidylinositol 3-kinase, Ras, and Rho (16). Ras and Rho proteins are low-molecular-weight GTPases. Ras proteins are central to the control of cellular growth and division. Ras binds to and is regulated by grb2-associated Sos (57). The Rho family of GTPases, which include RhoA, Rac1, and Cdc42, are critical for skeletal muscle differentiation and can regulate the expression of MyoD and myogenin (12, 58). An early effect of integrin-type signaling is the induction of the GTP-bound form of small GTPases, and one of the end results is the eventual NH2-terminal phosphorylation of c-Jun by c-Jun NH2-terminal kinase (JNK) (33). Phosphorylation of c-Jun prevents apoptosis and promotes homeostasis and hypertrophy.
We formulated a hypothesis that, just as integrins bind fibronectin to initiate signaling, laminin binding to the DGC complex initiates signaling; i.e., the DGC complex is a laminin receptor. By linking the cytoskeleton to the matrix, the DGC complex could sense the mechanical forces (stretching) of the sarcolemma that accompany contraction. Mechanical stretch, even in the absence of nerve activity, has long been recognized as a powerful modulator of gene expression in skeletal muscle. Muscle that is stretched exhibits a hypertrophic response, whereas an atrophic response is observed in slackened muscle (6, 25, 30, 55, 60). The initiating signal transduction mechanism for stretch has yet to be identified. In our model, the DGC complex is this stretch receptor. This stretch signaling is envisioned to promote cell growth and prevent apoptosis much as does integrin signaling. If this model is correct, atrophy is expected to cause profound changes in the DGC complex. Unused receptors may be upregulated or downregulated, but they are seldom unchanged by disuse. To investigate this model, we used the tenotomy model of gastrocnemius atrophy (30).
We show here for the first time that proteins of DGC complex decrease
in muscle atrophy and that the GTP-bound, activated state of small
GTPases like H-Ras, RhoA, and Cdc42 that are downstream (of the DGC
complex and integrin systems) also decrease in muscle atrophy. H-Ras,
Rac1, and Cdc42 are physically associated with laminin receptor and
with the complex containing -dystroglycan. Furthermore,
dominant-negative Cdc42 (constitutively inactive as the GDP-bound form)
transfected into the muscle of rats brought about changes in muscle
fibers that resemble muscle atrophy. These results are all consistent
with the DGC participating in the transduction of mechanical signals
that allow the muscle to adapt to muscle activity by either atrophy or hypertrophy.
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METHODS |
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Muscle preparation. Sprague-Dawley rats (200-300 g) were used for all experiments. Animals were housed in light-and temperature-controlled quarters where they received food and water ad libitum. Animals were anesthetized with isofluorane for surgery and tissue removal. The Animal Care and Use Committee of the University of Tennessee Health Science Center approved all procedures. Muscle atrophy was induced by tenotomy of the Achilles tendon, and in some cases the plantar tendon, in one of the limbs. An identical incision was made in the contralateral limb, and the Achilles tendon was isolated, but not severed, to serve as a sham-operated control. The gastrocnemius muscle was used for the experiments involving measurement of DGC complex proteins and small GTPase activity. Contralateral muscles served as a control.
For measurement of DGC complex proteins and GTPases, tenotomy was performed on the left limb in four rats and on the right limb in four rats (which served as duplicates). Ten days after surgery, rats were checked for weight gain and then anesthetized. The gastrocnemius muscles were removed from both the limbs, weighed, clamp frozen in liquid nitrogen, and stored atSkeletal muscle membrane preparation for DGC proteins. Total muscle membranes were prepared by using the procedure of Ohlendieck et al. (49). The frozen muscle was powdered under liquid nitrogen with the use of a BioPulverizer (Biospec Products) and homogenized in 7 vols of homogenization buffer (20 mM sodium pyrophosphate, 20 mM sodium phosphate, 1 mM MgCl2, 0.3 M sucrose, and 0.5 mM EDTA, pH 7.0) in the presence of a protease inhibitor cocktail (1 µg/ml pepstatin A, 10 µg/ml leupeptin, 10 µg/ml aprotinin, 1 mM benzamidine, and 0.1 mM phenylmethylsulfonyl fluoride) to minimize protein degradation. The homogenate was centrifuged at 13,000 g for 15 min at 4°C. The supernatant was centrifuged for 30 min at 30,600 g at 4°C to pellet total muscle membranes.
To examine dystrophin, syntrophin,Muscle membrane preparation for GTPases. For following the GTPases, the powdered muscle sample was divided into three equal portions (by weight). Two portions were homogenized in 7 vols of the same homogenization buffer containing the protease inhibitor cocktail. The third portion was homogenized in homogenization buffer with 10 mM MgCl2 and protease inhibitors. The total muscle membranes were then isolated as described earlier.
The membrane preparation of the first two portions were suspended in 50 mM Tris, 0.3 M NaCl, and 0.5 mM EDTA (with protease inhibitors) for inducing GTP- and GDP-bound forms. The third portion was suspended in 50 mM Tris, 0.3 M NaCl, 10 mM MgCl2, and 0.5 mM EDTA (with protease inhibitors) for measuring the endogenous amount of GTP-bound GTPases. The protein concentration of the three different samples was balanced by absorption at 280 nm as described earlier. Triton X-100 (25%) was added to all three samples to a final concentration of 1%. The samples were solubilized by mixing at 4°C for 30 min. Guanosine 5'-O-(3-thiotriphosphate) (1 mM; for GTP-loaded samples) and GDP (for GDP-loaded samples) were added to the first and second portions, respectively, and mixed for 5 min. This was followed by the addition of 10 mM MgCl2 to each sample and mixing for another 15 min. All three samples were centrifuged in microfuge tubes at 14,000 g for 15 min to remove the insoluble material. The final solubilized preparations were used for GTPase trapping.Preparation of PAK1-, PKN-, and Raf-Sepharose. Glutathione S-transferase (GST) fusions of the p21-binding domains p21-activated kinase 1 (PAK1), protein kinase N (PKN), and Raf were generated in pGEX-2T vector (37). These proteins were purified to homogeneity on glutathione agarose columns (54). The protein concentration was determined by Bradford's method (8) with bovine serum albumin (BSA) as the standard. The purified GST-fusion proteins (5 mg of protein, 2.5 mg/ml) were covalently coupled to 1 g of cyanogen bromide-activated Sepharose (Sigma) by following procedures recommended by the manufacturer (Pharmacia). The supports were then washed with the coupling buffer (0.1 M NaHCO3, pH 8.3, and 0.5 M NaCl) and blocked for 24 h with 0.1 M Tris · HCl and 0.5 M NaCl, pH 8.0. The amount of the protein coupled (0.7-0.8 mg fusion protein/ml Sepharose beads) was determined by the difference between the absorption at 280 nm of added protein and that recovered from the washes. For controls, activated Sepharose was used, to which either no protein or GST was coupled.
GTPase protein trapping. Raf, PKN, or PAK1 (50 µg) coupled to Sepharose beads (3 tubes each of Raf and PKN, 6 tubes of PAK1) were washed three times with 50 mM Tris, 0.3 M NaCl, 10 mM MgCl2, and 1% Triton X-100. The three different solubilized samples (GTP loaded, GDP loaded, and endogenous GTP bound) from the membrane preparations were added to the beads (to Raf-Sepharose for H-Ras, to PKN-Sepharose for RhoA, and to 2 sets of PAK1-Sepharose for Rac1 and Cdc42). They were mixed for 45 min to 1 h at 4°C. The supernatant was removed by centrifugation in microfuge tubes for 2 min. Each resin was washed three times with 50 mM Tris, 0.3 M NaCl, 10 mM MgCl2, and 1% Triton X-100 and then boiled in 50 µl of SDS-polyacrylamide gel electrophoresis sample buffer for 5-10 min at 95°C. Proteins were fractionated on a 12% SDS-polyacrylamide gel. Prestained SDS-PAGE molecular weight markers were obtained from Bio-Rad (Low-range markers).
Antibodies to DGC proteins and GTPases.
Monoclonal antibodies against dystrophin (NCL-DYS2), -dystroglycan
(NCL-
-DG), and
-sarcoglycan (NCL-
-SARC) were from Novacastra. The antibody against recombinant
-syntrophin was produced in rabbit
and purified by affinity chromatography on syntrophin A-Sepharose. The
monoclonal antibodies against H-Ras and RhoA were from Santa Cruz
Biotechnology, the monoclonal antibody against Rac1 was from Upstate
Biotechnology, and the monoclonal antibody against Cdc42 was from
Transduction Laboratories. Goat anti-mouse IgG-horseradish peroxidase
(HRP) and anti-rabbit IgG-HRP were from Santa Cruz Biotechnology.
Immunoblot analyses.
Transfer of proteins to nitrocellulose was performed according to
Towbin et al. (62) followed by immunoblot staining with antibodies. After blocking in TBST (20 mM Tris · HCl,
pH 7.5, 0.5 M NaCl, 0.2% Tween 20) containing 10 mg/ml BSA overnight, blots were washed three times (for 5 min each) with TBST containing 1 mg/ml BSA and incubated for 4 h at room temperature with 1:100 diluted antibodies for dystrophin, -dystroglycan, or
-sarcoglycan and 1:200 diluted antibody for syntrophin. For measuring GTPases, the
immunoblots were washed and incubated with 1:500 diluted antibody for
H-Ras, 1:50 diluted antibody for RhoA, 1:1,000 diluted antibody for
Rac1, or 1:200 diluted antibody for Cdc42. After the nitrocellulose membranes were washed three times (for 5 min each) in TBST with 1 mg/ml
BSA, immunoblots were incubated for 1 h with 1:2,000 diluted, HRP-conjugated goat anti-mouse IgG for all the proteins except syntrophin. For syntrophin, 1:2,000 diluted HRP-conjugated goat anti-rabbit IgG was used. Finally, immunoblots were again washed three
times with TBST with 1 mg/ml BSA and left in 1 M Tris · HCl, pH
8.5, until developed. The blots were developed using the enhanced
chemiluminescence method with luminol and coumaric acid. Solution
A (15 ml of 0.4 mM coumaric acid, 2.5 mM luminol, and 0.1 M
Tris · HCl, pH 8.5), and solution B (7.2 µl of
30% H2O2 in 15 ml of 0.1 M Tris · HCl,
pH 8.5) were mixed in the dark room. The membranes were soaked in this
mixture for 3-5 min and dried by blotting with Whatman filter
paper. The blots were then exposed for various time intervals to X-ray
films (Kodak) and developed with an automatic developer. Densitometric
scanning of different exposures of the same blot was carried out on a
computing densitometer (AlphaImager 2000, Alpha Innotech) and averaged.
Small GTPase trapping after laminin and -dystroglycan
depletion.
The gastrocnemius muscle (1 g) from normal rats was removed and frozen,
and total muscle membranes were prepared as described earlier. The
membrane pellet was suspended in 1.5 ml of immunoprecipitation buffer
(1% Triton X-100, 50 mM Tris · HCl, pH 7.5, 150 mM NaCl, 10 mM
MgCl2, 0.5 mM EDTA, 10 mM NaF, and 0.4 mM
Na3VO4, with protease inhibitors as described
earlier) and mixed for 30 min at 4°C. The suspension was centrifuged
in a microfuge, and the solubilized protein in the supernatant was
divided into three equal portions. Samples (60 µl each) of the
immunoprecipitation buffer and an antibody against laminin (Santa Cruz
Biotechnology) or an antibody against
-dystroglycan (a kind gift of
Dr. Tamara Petrucci) were added to the three portions, respectively,
and mixed for 2 h at 4°C. As a control, samples were also
prepared that were depleted of the
Na+-K+-ATPase
1-subunit with a
monoclonal antibody against the protein (
6F antibody deposited at
the Developmental Studies Hybridoma Bank, University of Iowa, by Dr.
D. M. Fambrough). The antibody-treated samples and the control
sample were mixed with 100 µl of protein-A Sepharose for 1 h at
4°C. The beads were centrifuged, and the supernatant was removed for
the GTPase-trapping assay described above. In this case, only the
endogenous GTP-bound forms were assayed for H-Ras, RhoA, Rac1, and
Cdc42. Protein A-Sepharose was washed three times with
immunoprecipitation buffer, and bound proteins were examined by electrophoresis.
Immunocytochemistry.
Glass slides were coated with 1% gelatin preserved with 0.02% sodium
azide. Serial sections (10 µm) of frozen tissues were collected on
the coated slides. The sections were allowed to air dry before being
fixed in 4% paraformaldehyde in PBS for 10 min at room temperature.
The slides were washed twice with PBS. The polyclonal antibody against
syntrophin and the monoclonal antibodies against -dystroglycan,
H-Ras, or Cdc42 were diluted 1:10 in PBS with 3% BSA and applied
directly to the sections; the slides were incubated for 30 min at room
temperature in a humidified chamber and washed in three changes of PBS
(5 min each). The labeled secondary antibody [anti-mouse or
anti-rabbit IgG, fluorescein isothiocyanate (FITC), or
tetramethylrhodamine isothiocyanate (TRITC) conjugate] at 1:100
dilution in PBS with 3% BSA was applied, and the slides were incubated
for another 30 min. The slides were washed with three changes of PBS,
and the sections were mounted with glycerol mounting medium.
Plasmid injection into muscle.
For plasmid injection into muscle, a 1-cm lateral incision in the lower
hindlimb was made to expose the soleus muscle (59). Plasmid (25 µg) in 10 µl of saline was injected in small aliquots throughout the muscle. The wound was closed, and the animals were allowed to recover for 1 wk. Plasmid constructs were on the pMX-green fluorescent protein (GFP) internal ribosome entry site retroviral backbone, containing either wild-type Cdc42 (67),
dominant-negative Cdc42 (T17N) (38), fast-cycling active
Cdc42 (F28L) (37), or stuffer DNA (LacZ). This plasmid
allows expression of bicistronic mRNA of GFP and small GTPases and
permits direct visualization of the small GTPases expressing
fluorescent tissues. After 1 wk of recovery, the animals were infused
intravenously with 10 ml of 4% paraformaldehyde in PBS and allowed to
fix for 10 min. The soleus muscles were removed and fixed in ice-cold
4% paraformaldehyde in PBS for an additional 10 min. The muscles were
cryoprotected overnight in 20% sucrose in PBS at 4°C before being
frozen in OCT mounting medium at 20°C.
Fluorescence microscopy of plasmid-injected fibers. Fibers that had incorporated the plasmid constructs and were expressing Cdc42 were identified by their GFP fluorescence. Cross sections (16 µm thick) of the soleus muscle were fixed to gelatin-coated slides and mounted with glycerol mounting medium. Fluorescence was identified by using a FITC or TRITC filter combination. Image fields containing fluorescing fibers and surrounding nonfluorescing fibers were collected. Fibers were defined and the fiber area calculated using NIH Image.
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RESULTS |
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The gastrocnemius muscle of the limb that underwent tenotomy showed a 20 ± 3% reduction in muscle mass (10 days postsurgery) compared with the contralateral gastrocnemius muscle. To examine the status of dystrophin and dystrophin-associated proteins in atrophied muscle, we isolated heavy microsomes from muscle homogenates. Heavy microsomes exhibited a highly reproducible protein composition and have been used as the source for purifying the DGC (19).
Immunoblots of heavy microsomes from normal and atrophied
gastrocnemius were examined for the relative abundance of
dystrophin, -dystroglycan,
-sarcoglycan, and
-syntrophin.
Immunoblot analysis of normal muscle with antibodies to dystrophin,
-dystroglycan,
-sarcoglycan, and
-syntrophin demonstrated the
presence of proteins of the expected sizes of 427, 43, 50, and 59 kDa
(Fig. 1), respectively, as previously
described for rabbit skeletal muscle (11, 19, 21). We
observed a reduction in the expression of dystrophin and other DGC
proteins in the atrophied muscle (Fig. 1). Densitometric scanning
revealed an 88 ± 2% reduction for
-dystroglycan and a 90 ± 3% reduction for
-sarcoglycan in atrophied muscle compared with
control muscle membrane. Dystrophin could not be detected in atrophied
muscle. The 59-kDa
-syntrophin did not show a significant change in
the atrophied muscle compared with the control muscle. Typical of
atrophy models and consistent with these data, total protein staining
of heavy microsome gels to verify protein loading revealed changes in
the expression of other proteins, as well. Therefore, it appears that a
specific loss of dystrophin,
-dystroglycan, and
-sarcoglycan
occurred during atrophy (per unit protein), whereas
-syntrophin was
unaffected. Syntrophin also served as a fortuitous internal control in
these experiments.
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Small GTP-binding protein signals were characterized on the basis of their interaction with a GST-fusion protein derived from their corresponding downstream effector targets. When RhoA becomes active, it exchanges bound GDP for GTP and then binds PKN to initiate a downstream event (50). Likewise, Rac1 and Cdc42 activate PAK1 through binding to its p21-binding domain. This sequence, located in the NH2-terminal regulatory part of the protein, contains a highly conserved 14-amino acid Cdc42/Rac-interactive-binding (CRIB) domain (amino acids 74-88) found in many proteins interacting with Rac- or Cdc42-GTP (4). Ras-GTP binds with the Ras-binding domain of Raf with high affinity (18). It is important to maintain the Mg2+ concentration (10 mM MgCl2) in the muscle membrane preparation to trap the endogenously present active GTP-bound form. The role of Mg2+ cofactor in the guanine nucleotide exchange and GTP hydrolysis reactions of Rho family GTP-binding proteins has been discussed by Zhang et al. (66).
The effector fusion proteins used to trap the small GTPases exhibit a
selective affinity for the GTP-bound form of GTPases (42).
We first verified that this specificity for the active conformation of
the GTPases is maintained in the isolated GST-fusion proteins. Purified
GST-fusions were used as a probe in an affinity precipitation assay
with different nucleotide-bound forms of GTPases. There was little or
no interaction with the inactive GDP-bound forms obtained by incubation
in GDP and EDTA. The results obtained for Cdc42 are shown in Fig.
2 and are representative of findings for
the other GTPases (data not shown). We also verified that the GTPases
did not bind nonspecifically to GST beads or to glutathione-Sepharose beads alone (data not shown). Triton X-100 (1%) was used to solubilize the GTPases in the muscle membrane preparations. GTP- and GDP-loaded forms served as positive (total GTPase, activated and trapped) and
negative controls (inactivated and, hence, could not bind to the
protein kinase fusion protein), respectively. These controls allowed us
to determine the percentage of the protein that was present in the
active GTP-bound form in isolated muscle membrane. We found that
94 ± 2% of total H-Ras, 26 ± 5% of total RhoA, 10 ± 4% of total Rac1, and 98 ± 1% of total Cdc42 were present in the active GTP-bound form in normal muscle membrane preparations (Fig.
3). When the total GTPases of normal
muscle were compared with those of the atrophied muscle, there was a
67 ± 4% reduction for H-Ras and a 73 ± 5% reduction for
Cdc42 in atrophied muscle membrane preparation (Fig. 3). RhoA was
either completely lacking or present in low amounts that could not be
detected in atrophied muscle membrane. There was no significant change
in Rac1. The ratio of the active GTP-bound form to the total GTPases in
atrophied muscle was the same as in normal muscle in the case of H-Ras, Rac1, and Cdc42. The ratio for RhoA in atrophied muscle was not determined because RhoA was not detectable in the atrophied muscle.
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If the DGC complex is a laminin receptor involved in small GTPase
signaling, as we propose, then a signaling complex might form
containing laminin, -dystroglycan, and small GTPases. To examine
this possibility, we used an antibody depletion assay. Antibodies
against laminin or
-dystroglycan were used to deplete microsomes
from normal muscle of these proteins, and the remainder was examined
for small GTPase content. The amount of H-Ras, Rac1, and Cdc42 trapped
by their respective effector domain fusion proteins was reduced to
5-10% of the normal level following depletion with either of the
antibodies (Fig. 4). In contrast, RhoA
was considerably less affected by laminin or
-dystroglycan depletion
(Fig. 4). As a control for nonspecific removal of the small GTPases, we immunodepleted the membrane preparations of the
1-subunit of Na+-K+-ATPase. Immunodepletion of
Na+-K+- ATPase did not affect the
amount of the small GTPases remaining in the membrane preparations
(Fig. 4). Immunoblot analysis of the proteins bound to the protein
A-Sepharose beads confirmed that the laminin and
-dystroglycan
depletion effectively immunoprecipitated the targeted proteins (data
not shown).
-Dystroglycan was depleted from the membrane
preparations treated with antibodies against laminin or
-dystroglycan, as well as by laminin-Sepharose.
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Immunocytochemistry of the control medial gastrocnemius muscle showed
uniformly arranged intact fibers, whereas the tenotomized muscle fibers
were smaller in size compared with the normal fibers and were pulled
apart from each other in some areas. These data are consistent with
numerous reports from other laboratories. Because H-Ras and Cdc42 were
reduced in muscle atrophy (Fig. 3) and found to be associated with
laminin and -dystroglycan in the depletion assay (Fig. 4), it was of
interest to confirm these observations by immunostaining.
-Dystroglycan, H-Ras, Cdc42, and
-syntrophin were all primarily
localized to the sarcolemma (Fig. 5). The
-dystroglycan fluorescence at the sarcolemma (Fig. 5A)
was reduced in the tenotomized medial gastrocnemius muscle fibers (Fig.
5B). Unlike
-dystroglycan, intracellular nonsarcolemmal immunofluorescence was detectable in addition to the sarcolemmal fluorescence for H-Ras (Fig. 5C) and Cdc42 (Fig.
5E). In tenotomized muscle, approximately one-half of the
fibers lacked H-Ras and Cdc42 staining at the sarcolemma, and the
immunoreactivity was found in aggregates (as punctate fluorescence) in
the remaining fibers. The overall fluorescence was reduced
significantly in tenotomized muscle fibers for both H-Ras and Cdc42
(Fig. 5, D and F, respectively). Labeling with
-syntrophin antibodies indicated no difference between the control
and tenotomized muscle fibers in syntrophin staining (Fig. 5,
G and H). These data are consistent with the
abundance of these proteins in the heavy microsomes isolated from
control and tenotomized muscle (Figs. 1 and 3).
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The four G proteins fall into three distinct groups. One of these,
represented by Rac1, is little affected by atrophy (Fig. 3). In
contrast, RhoA is dramatically affected by atrophy (Fig. 3) but does
not appear to be physically associated with either laminin or
-dystroglycan (Fig. 4). The third group contains H-Ras and Cdc42,
which are both diminished by atrophy and associated with laminin and
-dystroglycan. Like H-Ras, Cdc42 is also found predominantly in the
active form in both control and tenotomized muscle (Fig. 3). This last
group is clearly relevant to muscle atrophy. To further probe whether
Cdc42 activity was a potential signal transduction component affecting
muscle structure, we injected adult rat soleus muscle with retroviral
plasmids coding for forms of Cdc42 with different intrinsic activities.
Skeletal muscle is well known to be able to incorporate and express
from DNA injected directly into the muscle. The soleus muscle was
chosen because of its small mass, its exquisite sensitivity to
mechanical activity for its gene expression, and its surgical
accessibility. We were able to identify muscle fibers expressing the
Cdc42 forms by their coexpression of the GFP that was encoded behind an
internal ribosome entry site in the retroviral Cdc42 construct. Plasmid
vector (Fig. 6E), wild-type
Cdc42 (Fig. 6, A and B), and a fast-cycling
active mutant Cdc42 (data not shown) produce normal fiber appearance. The expression of GFP was in a subsarcolemmal or variegated pattern that was identical to the expression observed previously in muscle injected with retrovirus coding for
-galactosidase
(59). In contrast, the dominant-negative form of Cdc42
(Fig. 6, C and D) caused the muscle to exhibit
fibers that were misshapen and had lost fiber-to-fiber contacts. The
dominant-negative Cdc42-injected muscle also had areas that apparently
had lost muscle fibers (see e.g., Fig. 6C). The pattern of
GFP expression within the fibers did not appear different in the
dominant-negative Cdc42-injected muscles, except that the fluorescence
was now localized to subsarcolemmal regions that were not in cell-cell
contacts in many cases. A comparison of the frequency and distribution
of muscle fiber area in GFP-expressing regions of wild-type and
dominant-negative Cdc42-injected muscle is shown in Fig.
7. It demonstrates the significantly
diminished size of the fibers that received the dominant-negative
mutant (P < 0.001, power > 0.999).
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DISCUSSION |
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Our experiments support the hypothesis that the DGC complex in
skeletal muscle may participate in the transduction of mechanical forces to intracellular signals. Upon tenotomy, the muscle is no longer
subjected to normal stretching and the DGC receptor downregulates (Fig.
1). Dystrophin can no longer be detected 10 days posttenotomy, and
members of the dystroglycan and sarcoglycan complex are also greatly
diminished. -Syntrophin is not changed over this same time period.
Immunocytochemistry for
-dystroglycan and syntrophin further support
these data.
-Dystroglycan staining was reduced significantly, and
syntrophin remained the same (Fig. 5).
-Syntrophin is also known to
form complexes with utrophin at neuromuscular junctions that are devoid
of dystrophin (2); the
-syntrophin in these other
complexes may be more abundant or exhibit a compensatory increase with
tenotomy such that an effect on DGC syntrophin is masked. Furthermore,
the role of syntrophin in binding Na+ channels suggests
that there may be other complexes rich in syntrophin (23),
as do the observations that the syntrophins are often abundant in
tissues other than muscle or brain that express very little dystrophin
or utrophin (e.g., testes) (34). All the activities involving DGC complex proteins appear to be related in some way to cell
signaling, making it likely that they form a signal transduction complex. That it is altered in atrophy suggests that it may be important for regulating normal muscle growth and homeostasis.
Dystrophin is located in the sarcolemma, the myotendinous junctions,
and costameres (52). Myotendinous junctions are extensive and highly specialized subsarcolemmal adherens junctions, and costameres provide myofibril-to-sarcolemma attachment sites. We extracted and detected dystrophin from whole gastrocnemius muscle sarcolemmal membrane preparations (heavy microsomes), the source for
all purifications of the DGC. This may explain, in part, why we report
decreased dystrophin in the tenotomized gastrocnemius muscle, whereas
other investigators have reported increased dystrophin in denervated
muscle (5, 7, 51). In addition to contrasting atrophy
models with respect to the integrity of the neuromuscular junction, the
reports mentioned above also used cruder protein extracts from muscle
(only one of which was gastrocnemius). Whereas this earlier report used
a crude membrane fraction from the superficial portion of the
gastrocnemius muscle dissected free of adhering connective tissue
(7), we used isolated sarcolemma-derived microsomes from
the whole gastrocnemius muscle for our experiments. In our membranes
enriched in the DGC, we observed that dystrophin, -dystroglycan, and
-sarcoglycan are all decreased during the atrophy. We conclude that
the DGC is downregulated under these conditions. Nevertheless, the
proposed role of the DGC as a mechanical receptor is not excluded by
either an upregulation (5, 7, 51) or downregulation (this
report) that may occur in different animal models.
Concurrent with the downregulation of the DGC complex, we also observed changes in the small GTPases associated with the DGC. Small GTPases have been found to be important for integrin signaling and, by analogy, may be important in DGC signaling. The effects on Cdc42 and Ras appear quite similar (Fig. 3) in that almost all of the GTPase present is active and the amount present is greatly diminished by atrophy. The immunocytochemistry results for H-Ras and Cdc42 (Fig. 5) support these results from the in vitro assays using muscle membrane preparations. RhoA is the most dramatically affected of all the small GTPases, becoming undetectable in atrophied muscle (Fig. 3). Although RhoA was not present in detectable levels in our protein extractions from atrophied muscle, this does not mean that it was completely absent. Rac1 is seemingly unaffected by atrophy. Paradoxically, this may suggest that it is particularly important to atrophy and its recovery. Muscles recover from atrophy when stretching and contractile activities are resumed, so the atrophied muscle must contain within it the components necessary for recovery. Because Rac1 is available and only modestly activated in normal and atrophied muscle, it has a potentially large range of response to changes in the muscle.
The results of the laminin and -dystroglycan depletion assay show
that there is a physical link between the DGC and the small GTPases.
The association of H-Ras, Rac1, and Cdc42 with laminin and
-dystroglycan (Fig. 4) and the localization of
-dystroglycan, H-Ras, and Cdc42 at the sarcolemma (Fig. 5) are notable in this regard.
In support of this suggestion, we also found that the DGC proteins are
directly associated with the small GTPase Rac1 through grb2 and Sos
(48). Laminins are major components of the basement
membrane. Cells bind directly to laminins via a subset of integrins and
other non-integrin-type receptors, such as dystroglycans. Reduction of
the small GTPases H-Ras, Rac1, and Cdc42 due to laminin depletion
implies that these GTPases may be signal transduction elements
reflecting mechanical attachment between the cell cytoskeleton and the
extracellular matrix. The similar magnitude of reduction in
these GTPases due to
-dystroglycan depletion shows the involvement of DGC in GTPases signaling.
The retroviral transformation data (Fig. 6) complement the GTPase trapping data (Fig. 3). The fast-cycling active mutant of Cdc42 does not alter fiber morphology, similar to the vector and wild-type controls. Because of its rapid cycling property and the >10-fold excess of GTP over GDP in cells, this protein functions as though it were constitutively active. Because Fig. 3 shows that virtually all of the Cdc42 present in muscle is active, the fast-cycling active mutant form is in the same high-activation state as the native protein, and this may be why it had no effect. However, when constitutively inactive (dominant negative) Cdc42 is introduced, it clearly alters the muscle fiber morphology (Figs. 6 and 7). Muscle fiber diameter decreases in atrophy, so in that sense, the dominant-negative phenotype is similar to atrophy. We also performed analogous experiments using retroviral constructs containing variants of RhoA. All were without any discernable effect (data not shown). Thus we conclude that Cdc42 has the special property of affecting muscle fiber morphology, a property not shared with RhoA.
The reduction of H-Ras, Rac1, and Cdc42 signals in response to laminin
or -dystroglycan depletion (Fig. 4) strongly suggests that they are
physically linked to one another in a signaling complex. However, this
alone does not establish a cause-and-effect relationship. Thus much
remains to be done to establish their functional links. Nonetheless,
the studies presented arose from a hypothesis that predicts such a link
and would be hard to explain by any other existing model for the
function of the DGC complex. Because muscle is a complex tissue, there
are many possible transduction mechanisms that may influence muscle
structure and function. Members of the Rho family of GTPases have
previously been implicated in the sarcomere assembly: Rac1 activity
inhibits normal assembly during myotube differentiation
(22), and RhoA activity initiates reorganization during
cardiac hypertrophy (14). Our studies show the possible
involvement of H-Ras and Cdc42 GTPases signaling in muscle atrophy. Of
these two, we have shown that the dominant-negative Cdc42 results in
abnormal muscle cells. It is important to determine the role of H-Ras
by studying the effect of dominant-negative H-Ras on muscle fibers,
which we plan for future work. Furthermore, we have recently shown that
grb2 binds
-syntrophin (48), and it is already known
that grb2 also binds to
-dystroglycan (64), suggesting
that signaling through the Ras family small GTPase proteins is likely
to be relevant to the DGC complex.
The signal transduction mechanisms for stretch in striated muscle are not well understood. Several studies have demonstrated heterotrimeric G protein activation (17, 27, 63) or focal adhesion kinase (FAK) activation (26). However, there is little information linking skeletal muscle stretch to modulation of small G protein activity. Nonetheless, results from smooth and cardiac muscle support the model presented here. For example, the elastin-laminin receptor functions as the stretch receptor in vascular smooth muscle, and a peptide derived from elastin can inhibit it. This stretch signaling involves grb2 recruitment, and cRas-dominant-negative transfection prevents signaling whereas an inhibitor of stretch-activated cation channels does not (56), showing that grb2/small GTPase signaling is important to stretch signaling in smooth muscle. In bladder smooth muscle, stretch activates SAPK2 and JNK (44), a frequent consequence of grb2/small GTPase signaling. In cardiomyocytes, stretch activation of another mitogen-activated protein kinase, extracellular signal-regulated kinase 2 (ERK2), requires an intact Rac1/RhoA pathway. Dominant-negative constructs of Rac1 and RhoA prevent activation of ERK2 by stretch, as does inhibition of RhoA (3). The data presented here are consistent with these reports of modulation of small G proteins by stretch. Because the DGC complex is also present in cardiac and smooth muscle, it may be a common upstream transducer of mechanical stretch.
The model proposed here of DGC being involved in mechanotransduction
mechanisms regulating muscle size has thus survived our initial
examination. This hypothesis predicts that the DGC complex, involved in
many devastating muscular dystrophies, is also involved in muscle
atrophy (a normal process of muscle remodeling). We now have
experimental data to support this prediction. Our prediction is also
supported by Brown et al. (9), who showed that the treatment of cultured myotubes with a monoclonal antibody that blocks
-dystroglycan binding to laminin leads to the induction of a
dystrophic phenotype in vitro. The phenotype is characterized by
reduced myotube size, myofibril disorganization, loss of contractile activity, and reduced spontaneous clustering of acetylcholine receptors; this phenotype is reversed by addition of exogenous laminin
2. They concluded that
-dystroglycan may be part of a signaling
pathway for the maturation and maintenance of skeletal myofibers.
The hypothesis presented here, if it survives subsequent testing, could represent an important advance. Detailed knowledge of this signaling may provide insights into the molecular pathology of the various inherited muscular dystrophies, and identify valuable pharmacological targets and new therapeutic strategies. A consequence of the model is that these therapies may also be effective against muscle atrophy in the convalescing and elderly.
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ACKNOWLEDGEMENTS |
---|
We acknowledge the support of the Muscular Dystrophy Association (H. W. Jarrett), National Institute of General Medical Sciences Grant GM-60523 (Y. Zheng), and the American Diabetes Association and American Heart Association (D. B. Thomason).
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FOOTNOTES |
---|
Present address of P. S. Chockalingam: Wyeth-Genetics Institute, Musculoskeletal Sciences, 200 Cambridge Park Dr., Cambridge, MA 02140.
Address for reprint requests and other correspondence: H. W. Jarrett, Dept. of Molecular Sciences, Univ. of Tennessee Health Sciences Center, Memphis, TN 38163 (E-mail: hjarrett{at}utmem.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
April 3, 2002;10.1152/ajpcell.00529.2001
Received 6 November 2001; accepted in final form 27 March 2002.
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