Journal of Applied Physiology
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

J Appl Physiol 98: 1554-1561, 2005. First published October 29, 2004; doi:10.1152/japplphysiol.01070.2004
8750-7587/05 $8.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF) Free
Right arrow All Versions of this Article:
98/4/1554    most recent
01070.2004v1
Right arrow Submit a response
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Web of Science (3)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Bellott, A. C.
Right arrow Articles by Burkholder, T. J.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Bellott, A. C.
Right arrow Articles by Burkholder, T. J.

HIGHLIGHTED TOPICS
Biomechanics and Mechanotransduction in Cells and Tissues

Reduction of caveolin-3 expression does not inhibit stretch-induced phosphorylation of ERK2 in skeletal muscle myotubes

Anne Claire Bellott,1,3 Kalpit C. Patel,2,3 and Thomas J. Burkholder2,3

1School of Mechanical Engineering, 2School of Applied Physiology, and 3Interdisciplinary Bioengineering Program, Georgia Institute of Technology, Atlanta, Georgia

Submitted 27 September 2004 ; accepted in final form 22 October 2004


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Mechanotransduction is critical to the maintenance and growth of skeletal muscle, but the mechanism by which cellular deformations are converted to biochemical signals remains unclear. Among the earliest and most ubiquitous responses to mechanical stimulation is the phosphorylation and activation of mitogen-activated protein kinases, in particular ERK2. Caveolin-3 (CAV-3) binds ERK2 and its upstream activators in inactive states on the caveolae of resting muscle. Caveolae are deformed by stretch, and it was hypothesized that this deformation might disrupt the CAV-3-dependent inhibition of ERK2 to affect stretch-induced activation. Stretch-induced phosphorylation of ERK2 in myotubes was both amplitude and velocity dependent, consistent with a viscoelastic mechanism, such as deformation of caveolae. Chemical disruption of caveolae by cholesterol depletion increased ERK2 activation by up to 176%. Small interfering RNA oligomers were then used to knock down expression of CAV-3 in cultured myotubes before mechanical stimulation, with the expectation that reducing CAV-3 expression would eliminate the stretch-induced activation of ERK2. Knockdown reduced CAV-3 protein content by 55% but did not significantly alter the stretch-induced increase in ERK2 phosphorylation, suggesting that CAV-3 is not an essential element of the mechanotransduction pathway, although the limited extent of knockdown limits the strength of this conclusion.

mechanotransduction; caveolae; mitogen-activated protein kinase

THE IMPORTANCE OF MECHANICAL signaling to the normal maintenance of muscle function has been extensively studied (1, 7, 17, 24). Mechanical signals drive postnatal muscle growth (63) and lead to dramatic alterations in adult muscle structure (19, 62), whereas removal of mechanical stimulation by disuse or age leads to muscle atrophy (24, 26), and diseases that disrupt the mechanotransduction machinery can have devastating consequences (39, 55). Mechanical stimulation induces production and release of growth factors (59, 66), activation of signaling cascades (30, 35) leading to increased protein synthesis (2), and activation of quiescent satellite cells (57). Several mechanisms for this mechanotransduction have been suggested, including integrins (7, 29), membrane disruption (10, 31), and caveolae (22).

Caveolae are rounded, smooth invaginations of the plasma membrane 50–100 nm in diameter (34, 37). A subset of lipid rafts, caveolae are microdomains within the plasma membrane that have a high concentration of sphingolipids and cholesterol (25, 48, 49). They have been shown to open during stretch (11, 41), possibly to shield the cells from damage (43, 67). In addition, many signaling molecules colocalize with caveolae, including PKC{alpha}, Src, MEK, ERK, H-Ras, G-proteins, and eNOS (14, 25, 46, 53). Caveolae depend on caveolin for their characteristic structure (45, 47).

There are three known forms of the mammalian caveolin protein, caveolin-1, -2, and -3, of which caveolin-3 (CAV-3) is specific to striated muscle (56, 61). Expression of the CAV-3 gene is induced during myoblast differentiation (54), and the protein localizes at the sarcolemma, where it forms a complex with dystrophin (16, 54). Of particular interest, CAV-3 has also been shown to bind many of the caveolae-associated signaling molecules in their inactive form (40), and many of these molecules are activated by stretch.

Of note, disruption of CAV-3 has been shown to also result in the hyperactivation of the ERK2 cascade (64). The traditional ERK cascade, which is entirely associated with and inhibited by caveolin (12), consists of three sequentially activated kinases: Raf-1 or B-Raf, which phosphorylates MEK1/2, which phosphorylates ERK1/2 (8, 12). ERK is quickly and strongly phosphorylated after stretch (30). After activation, ERK can regulate cellular proliferation and control the cell cycle (20, 50). Previous work with smooth muscle has shown that caveolins keep ERK inactive in static cells (21). In cardiac myocytes, cyclic stretch leads to activation and dissociation of RhoA and Rac1 from caveolae, which facilitates nuclear translocation of ERK (22).

Intact caveolin filaments associate with and inhibit signaling molecules involved in muscle growth and hypertrophy (12), but stretch deforms those caveolae (11) and may disrupt the carefully organized caveolin structure (21). This disruption may eliminate the inhibitory influence of the caveolin scaffolding domain on signaling molecules, resulting in the activation of those signaling molecules (60, 64). In this model, the mechanical properties of caveolae and the biological activity of CAV-3 play central roles in the transduction of mechanical signals.

Three specific predictions made by this model were tested. First, deformation of caveolae requires viscous flow of the lipid membrane into a new configuration and elastic deformation of the protein coat, leading to the hypothesis that stretch-induced ERK2 phosphorylation should reflect a viscoelastic response and be influenced by both stretch amplitude and velocity. Varying stretch conditions revealed that stretch-induced phosphorylation of ERK2 increased with both amplitude and velocity, consistent with a mechanism dependent on membrane flow. Second, the model predicts that ERK2 is maintained in a nonphosphorylated state by intact caveolae and CAV-3, leading to the hypothesis that chemical disruption of caveolae should increase ERK2 phosphorylation in nonstretched myotubes. Destabilization of the caveolae structure by cholesterol depletion resulted in changes consistent with inhibition of ERK2 phosphorylation by intact caveolae. Third, the model predicts that stretch-induced ERK2 phosphorylation is mediated by mechanical disruption of constitutive CAV-3 inhibition, leading to the hypothesis that elimination of CAV-3 should increase ERK2 phosphorylation in nonstretched myotubes, but stretch of those myotubes should result in no additional increase in ERK2 phosphorylation. Knockdown of CAV-3 expression by small-interference RNA (siRNA) increased resting ERK2 phosphorylation in myotubes grown on elastic substrates, but stretch induced still further phosphorylation, suggesting that caveolin is not required for phosphorylation of ERK2 after stretch.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Cell culture.   C2C12 myoblasts (ATCC, Manassas, VA) were maintained in DMEM supplemented with 10% FBS and antibiotics. Cells were seeded at 0.5 x 104 cells/cm2 onto Matrigel-coated membranes. Cultures were switched to DMEM supplemented with 10% horse serum and antibiotics to induce differentiation and subjected to 25% stretch to induce alignment.

Fusion of C2C12 cultures is relatively limited, and additional experiments were performed using primary culture. Myoblasts were isolated by enzymatic digestion of hindlimb musculature of neonatal CFW mice (Swiss-Webster, Charles River Labs) (42), using procedures reviewed and approved by Georgia Institute of Technology's Institutional Animal Care and Use Committee. Two- to 5-day-old mice were killed by CO2 asphyxiation, decapitated, and immersed in 70% ethanol. Hindlimb musculature was separated from the bone and skin and minced with razor blades in dissociation solution (PBS containing 10 mM CaCl2, 1.5 U/ml collagenase, 2.4 U/ml dispase). The tissue slurry was incubated for 30 min at 37°C, diluted in growth medium [Ham's F10 nutrient solution (F10) supplemented with 20% FBS, 2.5 ng/ml basic FGF (bFGF), 100 IU/ml penicillin, and 100 µg/ml streptomycin], and filtered through a 100-µm mesh, and the cells were pelleted for 5 min at 350 g. Cells were plated in the growth medium on collagen-coated dishes and identified as passage 0. Cultures were enriched for myoblasts through 10–12 successive passages by selective trypsinization and differential adhesion (42), and myogenic potential was determined by desmin staining. In the following experiments, no cells older than passage 19 were used.

Cells were seeded at 104 cells/cm2 on Matrigel-coated silicone membranes held in stretcher cassettes (4). Cells were allowed to proliferate to roughly 70% confluence. The cultures were then switched to differentiation media (DMEM supplemented with 2% horse serum and antibiotics) and stretched by 25% of seeding the length to promote unidirectional fusion of the myotubes (4). Stretches were performed using a custom, stepper motor-driven device controlled by Labview software. Cells were allowed to differentiate into myotubes for 2–3 days without media change.

Mechanical stimulation.   Cultures were preincubated in CMG (PBS with 0.01% MgCl2, 0.015% CaCl2, and 0.07% glucose) before stretch to minimize serum-stimulated ERK2 activation. C2C12 myotubes, uniformly aligned near the axis of stretch, were subjected to cyclic uniaxial stretch for 15 min by using a range of velocities between 4 and 40%/s and a range of amplitudes from 10 to 30%. The stretch profile was sinusoidal and symmetric about the resting length of the membrane, and reported amplitudes are peak to peak. CFW myotubes were subjected to stretch for 10 min of 30% (±15%) amplitude. After stretch, the myotubes were washed twice in ice-cold PBS and harvested in 100–150 µl of nondenaturing lysis buffer [1% Triton X-100, 50 mM Tris, 250 mM NaCl, 25 mM EDTA, 0.1% protease inhibitors (Sigma), 100 mM NaF, and 4 µg/ml NaVO3]. Cells were lysed for 30 min on ice and cleared by centrifugation at 5,000 g for 5 min. Soluble protein concentration was determined by bicinchoninic acid assay (BCA, Pierce). All samples were stored at –20°C before analysis.

Cholesterol depletion.   To disrupt the caveolae structure, some cultures were incubated for 1 h in CMG plus 0.1% or 0.5% methyl-{beta}-cyclodextrin (MCD) before stretch. To validate cell and ERK2 viability, static cultures were subjected to cholesterol depletion by MCD, followed by addition of 2.5 ng/ml bFGF to assess maximal ERK2 stimulation levels. Treatment with bFGF for 0, 5, 15, 25, and 40 min after the initial 60-min preincubation encompassed the time course of a normal stretch experiment.

CAV-3 knockdown.   Three validated siRNA sequences (Ambion) were evaluated for knockdown efficacy. RNA oligos were transfected into differentiated myotubes using a commercially available cationic lipid reagent (Oligofectamine, Invitrogen), according to the manufacturer's instructions. Briefly, siRNA was diluted to 18.3 µg/ml in serum-free DMEM, and Oligofectamine was diluted 1:5 in serum-free DMEM. The diluted lipid reagent was then added 2:9 to the siRNA mixture. The cells were rinsed once with serum-free DMEM and bathed in serum-free DMEM. Lipid:RNA complexes were applied to the cells, and an equivalent amount of lipid reagent was applied to control cultures. Cultures were incubated with the transfection mixture for 6 h, rocked every 2 h, and then supplemented with 125 µl/well of DMEM plus 6% horse serum. Parallel cultures were transfected with fluorescein-tagged nonsilencing RNA oligomers (Qiagen) and imaged under epifluorescent illumination to determine transfection efficiency. Cultures were maintained for an additional 2 days before mechanical stimulation. Pilot experiments indicated that that the three oligomers resulted in equivalent knockdown (91.2 ± 8.9%, 91.0 ± 5.0%, and 86.5 ± 6.6%), and the "cav3-1" oligomer (5'-GGACAUUGUGAAGGUAGAUtt-3') was used for all further experiments. Parallel control cultures were subjected to sham knockdown either with nonsilencing RNA oligomers (Qiagen) or with Oligofectamine alone. No differences were noted between the two sham treatments, which were then pooled for statistical analysis.

Osmotic stress.   Hyperosmotic conditions lead to phosphorylation of the MAP kinase cascade but also to cell shrinkage, which would reduce stress on the cell membrane. The induction of ERK2 phosphorylation resulting from hyperosmotic stress must result from a mechanism other than CAV-3 signaling. To determine whether CAV-3 knockdown adversely altered other signaling pathways, C2C12 cultures were preincubated for 60 min in CMG (304 mosmol/kgH2O), followed by incubation for 15 min in either CMG (304 mosmol/kgH2O), CMG supplemented with an additional 20 mM glucose (324 mosmol/kgH2O), or CMG diluted to 284 mosmol/kgH2O with 0.01% MgCl2, 0.015% CaCl2, and 0.07% glucose to maintain consistent concentrations of Ca2+, Mg2+, and glucose. Cultures were then lysed in the nondenaturing lysis buffer and stored at –20°C before analysis.

SDS page and Western blotting.   All protein samples were separated by SDS-PAGE using 10 or 12% acrylamide gels. For Western blotting, proteins were transferred for 1 h at 1.5 mA/cm2 onto a nitrocellulose membrane by a semidry transfer system. After transfer, the membranes were blocked with 2% BSA in 10 mM Tris, 150 mM NaCl, 0.1% Tween-20, pH 7.5 at room temperature for 1 h or overnight at 4°C. Membranes were then incubated in primary antibody against phosphorylated ERK (1:3,000, Cell Signaling), or CAV-3 (1:2,000, BD), rinsed, and incubated with horseradish peroxidase-conjugated secondary antibody (1:20,000). Membranes were visualized by enhanced chemiluminescence (ECL; Amersham) and exposure to film (Kodak). Membranes probed for phospho-ERK were subsequently stripped and reprobed for ERK2 (1:3,000, Transduction Laboratories). Films were digitized, and optical density was quantitated by use of MATLAB (MathWorks). ERK2 phosphorylation was determined by first normalizing phospho-specific integrated optical density (IOD) to the non-phospho-specific IOD, and then normalizing each sample from a particular film to the mean IOD ratio for the control samples from that film. All ERK2 gels included, as a positive control, a sample of C2C12 myotubes stimulated with 2.5 ng/ml bFGF to maximally phosphorylate ERK2. These samples generally displayed phosphorylation of 10- to 15-fold that of nonstretched controls (data not shown).

For assay of myosin content, gels were fixed (50% MeOH, 10% HOAc) and stained with 0.025% Coomassie blue. Gels were extensively destained and scanned, and relative myosin content was determined by IOD.

Cell morphology.   Fusion index and myotube diameter were measured to evaluate any effect of knockdown on myotube viability or morphology. Briefly, fusion index was measured by counting single nuclei vs. number of nuclei present in multinucleated myotubes. Pictures were taken at x20 magnification. Myotube diameter was measured with Scion Image software.

CAV-3 expression after knockdown was also evaluated by immunocytochemistry. Membranes were rinsed with PBS and fixed with 50% methanol in PBS for 5 min. The membranes were rinsed twice and blocked in 5% BSA in PBS for 1 h. Cultures were incubated with CAV-3 antibody (1:100 Santa Cruz) for 30 min, rinsed, and incubated with fluorescein-tagged secondary antibody (1:100). The membranes were rinsed twice more in PBS for 5 min each and covered in glycerol. Cells were imaged under epifluorescent illumination on a Leica DMLS microscope equipped with a Canon EOS D30 camera.

Statistical analysis.   Results were analyzed by one- or two-way ANOVA using Statview 5.01 (SAS). Fishers paired least-significant difference correction was applied to post hoc t-tests to evaluate specific differences between treatments. The significance threshold was set to P < 0.05, and all data are reported as means ± SD. Where differences between groups are not seen, the distinction is made between groups that are statistically identical, on the basis of post hoc statistical power < 0.10, and groups that are statistically indeterminate (P > 0.05, but power > 0.10).


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
Cell mechanics.   Cyclic stretch of C2C12 myotubes with peak-to-peak amplitudes ranging from 10 to 30% of resting length resulted in a graded increase in ERK2 phosphorylation (Fig. 1A, P < 0.05). Phosphorylation of ERK2 also displayed a graded increase with increasing stretch velocity (Fig. 1B, P < 0.05). These observations indicate that both stretch amplitude and velocity play a role in stretch-induced activation of ERK2, as would be predicted for a viscoelastic process. The phospho-specific antibody recognizes both phospho-ERK2 and phospho-ERK1, but the relative intensity of the two species frequently prevented quantification of the less abundant phospho-ERK1. In cases in which both species could be evaluated, the response of each was similar (data not shown).



View larger version (21K):
[in this window]
[in a new window]
  		Fig. 1. Phosphorylation of ERK2 (pERK2) in C2C12 myotubes after stretch increases with stretch amplitude at 20%/s (A) and velocity at 30% peak to peak (PtP) amplitude (B). *P < 0.05 vs. static; means ± SD; n > 6 per group. pERK1, phosphorylation of ERK1.

 
Further experiments were performed using myotubes derived from CFW mice. Untreated CFW myotubes displayed a muted stretch response relative to the C2C12 myotubes, with 30% cyclic stretch producing 81% increase in ERK2 phosphorylation (Fig. 2). The hypothesis that intact caveolae and CAV-3 filaments are required for suppression of ERK2 activity predicts that disruption of caveolae should increase ERK2 phosphorylation. CFW myotubes were treated with MCD for 60 min to extract cholesterol and disrupt caveolae. Treatment with either 0.1 or 0.5% MCD resulted in increases in resting ERK2 phosphorylation of 176 and 62%, respectively (Fig. 2), and two-way ANOVA confirmed a significant effect of MCD treatment (P < 0.05). MCD-treated CFW myotubes subjected to mechanical stimulation displayed additional increases in phosphorylation of 75–120%, which was similar to increases observed in untreated CFW myotubes, and two-way ANOVA reveals no significant interaction between MCD and stretch (P > 0.5). The lack of difference carries statistical power of 0.14, which fails to support the counterclaim that MCD treatment does not alter ERK2 phosphorylation after stretch. The MCD experiments, therefore, indicate that MCD-induced disruption of caveolae increases ERK2 phosphorylation, but the experiments neither support nor refute a role for caveolae in mechanical signaling.



View larger version (30K):
[in this window]
[in a new window]
  		Fig. 2. pERK2 in CFW myotubes was increased by methyl-{beta}-cyclodextrin (M{beta}CD)-mediated cholesterol depletion, but cholesterol depletion did not reduce stretch-induced phosphorylation. *P < 0.05 vs. nonstretch; means ± SD; n > 8 per group.

 
Caveolin knockdown.   siRNA oligo treatment reduced CAV-3 protein expression by 55 ± 7% in C2C12 myotubes (P < 0.01) but failed to alter either resting or stretch-induced ERK2 phosphorylation (Fig. 3). CAV-3 is induced during myotube fusion, and it was believed that the relatively high proportion of mononucleated cells remaining in C2C12 cultures might mask myotube-specific, CAV-3-dependent responses. Additional experiments were performed in primary culture, which achieve a greater extent of differentiation.



View larger version (29K):
[in this window]
[in a new window]
  		Fig. 3. pERK2 induced by ±15% amplitude and 20%/s velocity stretch in C2C12 myotubes is largely unaffected by siRNA-mediated caveolin-3 (CAV-3) knockdown. *P < 0.05 vs. nonstretch; means ± SD; n = 7–10 per group.

 
When CFW myotubes were subject to siRNA-mediated knockdown of CAV-3, CAV-3 protein levels decreased by 45 ± 20% (P < 0.01) after 48 h. This is distinctly less effective knockdown than the roughly 90% achieved in pilot experiments. Pilot cultures were grown on tissue culture plastic, to which the cells adhere substantially better than the Matrigel-coated silicone membranes used in stretch experiments, and it is believed that detachment of more mature myotubes reduces the net reduction in CAV-3 expression. Immunostaining of knockdown CFW myotubes revealed a dramatic reduction in fluorescence intensity relative to untreated controls (Fig. 4). Because CAV-3 has been implicated in myotube differentiation and fusion, cultures were assayed for markers of differentiation. Myotube diameter was 21 ± 7 µm in untreated CFW myotubes and 21 ± 3 µm in knockdown CFW myotubes, and fusion index was 91% in untreated cultures and 90% in treated cultures. Myosin content did not differ significantly between groups, although knockdown cultures yielded only 80 ± 50% as much myosin as untreated cultures. These observations also provide some assurance that the potential loss of myotubes to adhesion did not systematically alter the cell population.



View larger version (71K):
[in this window]
[in a new window]
  		Fig. 4. CAV-3 knockdown (A) substantially reduced the appearance of CAV-3 by immunofluorescence relative to sham-transfected controls (B).

 
ERK phosphorylation in static CFW myotubes subjected to knockdown was increased by 48 ± 15% (P < 0.01, Fig. 5) over untreated CFW myotubes, as has been seen in other models of CAV-3 disruption (64). Stretch of CFW myotubes subjected to knockdown produced a further 40 ± 25% rise in ERK2 phosphorylation, which was statistically identical to the 81 ± 32% increase seen in stretch of untreated cultures (P > 0.6; power < 0.1). These observations suggest that knockdown of caveolin expression increases resting ERK2 phosphorylation, consistent with a requirement for intact CAV-3 in suppression of ERK2 activity, but fails to block stretch-induced phosphorylation, inconsistent with a requirement for CAV-3 in mechanotransduction.



View larger version (26K):
[in this window]
[in a new window]
  		Fig. 5. ERK2 phosphorylation induced by mechanical stimulation was not substantially reduced by CAV-3 knockdown in primary myotubes. *P < 0.05 relative to nonstretch; n > 8 per group.

 
Osmotic stress.   Exposure of C2C12 myotubes to varying osmotic conditions resulted in small and not significant changes in ERK2 phosphorylation (Fig. 6). The trend of phosphorylation is consistent with previous reports, and it is possible that the muted response reported here results from dilution of the stimulation media with residual preincubation media. Interestingly, CAV-3 knockdown significantly reduced the extent of phosphorylation, independent of osmotic treatment (P < 0.03). This reduction in C2C12 myotubes opposes the 48 ± 15% increase in resting phosphorylation seen in CFW myotubes and the absence of effect seen in C2C12 myotubes grown of silicone membranes. The only apparent difference in growth conditions for the C2C12 cultures that display no effect of knockdown and that display reduced ERK2 phosphorylation is the growth substrate.



View larger version (29K):
[in this window]
[in a new window]
  		Fig. 6. ERK2 phosphorylation induced by changes in extracellular osmolarity: mOsm, change in osmolarity from preincubation media, +20, Hi; 0, Norm; –20, Low (means ± SD; n = 6 per group).

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 
The goal of this project was to evaluate whether mechanically induced phosphorylation of ERK2 is dependent on caveolae and CAV-3. ERK2 phosphorylation is influenced by amplitude and velocity, consistent with a role for deformation of viscoelastic structures like caveolae. Chemical disruption of caveolae with MCD resulted in ERK2 phosphorylation similar to mechanical stretch, although the chemical disruption of caveolae did not significantly alter the stretch response. Similarly, knockdown of CAV-3 expression increased resting ERK2 phosphorylation but failed to eliminate the stretch response. Although these observations indicate the important role of CAV-3 in suppressing ERK2 activation in resting cells, they are inconsistent with a critical role for deformation of caveolae in the transduction of mechanical signals.

The influence of amplitude and velocity on ERK2 activation is important. Previous work has shown that muscle adapts its structure to meet specific functional demands (23, 63) and that, during normal use, muscle frequently generates force near optimal length and velocity (27). This reflects a close correlation between the functional demands on the muscle and its physical structure (5), and it is appealing to think that this correlation might be biologically regulated. To maintain force generation at optimal length and velocity, a sensor of length and velocity must exist. We have previously shown that stretch-induced amino acid incorporation exhibits a velocity dependence (9) and that membrane permeability becomes increasingly velocity dependent as stretch amplitude increases (4); we now add that ERK2 phosphorylation is velocity dependent.

Membrane deformation, which depends on the flow of the fluid membrane, is an attractive candidate for a velocity-dependent mechanism. Caveolae have been shown to deform after stretch (11, 41), and caveolin inhibits an array of classically stretch-activated signaling cascades (36, 51), which makes the proposed model intuitively appealing. The potential role of caveolae and caveolin in mechanotransduction was first indicated by Park and colleagues (38), who demonstrated that cholesterol is critical to the shear-induced activation of ERK and that exposure to shear rapidly increases the surface density of caveolae. Subsequent work has shown that chronic exposure to shear stress leads to upregulation of caveolin-1 and attenuates the shear-induced activation of ERK (3).

The role of caveolin in transduction of stretch specifically has been previously suggested as well. Kawamura and colleagues (22) demonstrated that stretch leads to the rapid dissociation of RhoA and rac1 from CAV-3 in cardiac myocytes. Dissociation and subsequent actin filament reorganization were found to be necessary for nuclear translocation of activated ERK, and chemical disruption of caveolae was sufficient to block stretch-induced activation of RhoA and rac1. More recently, Kawabe and colleagues (21) demonstrated that stretch leads to a displacement of caveolin protein from the caveolae of smooth muscle, consistent with stretch-induced disruption of the caveolin protein coat.

One of the primary goals of this project was to test whether this association between CAV-3 redistribution and stretch reflects a mechanistic role for CAV-3 in mechanotransduction. It was demonstrated that siRNA-mediated knockdown substantially reduces CAV-3 protein expression, although not to the extent observed in clinically pathological cases, and increases ERK2 phosphorylation, as has been reported (8, 64). This magnitude of knockdown was not sufficient to alter mechanotransduction, although the numerical results suggest a reduction in the magnitude of the stretch response. It is possible that the remaining CAV-3 protein and residual caveolae are sufficient to mediate the response to physiologically relevant stretch amplitudes, which are expected to require opening of only 1–2 caveolae per square micrometer, or only 10–20% of all caveolae (34).

The motivation for surveying a range of stretch conditions was to begin to identify cellular responses with similar mechanical dependency. Although the velocity dependence reported here is similar to that of membrane disruption (4), the amplitude dependence of disruption and ERK2 phosphorylation is not similar. Recent work by Mack and colleagues (28) has suggested a threshold effect of force on focal adhesion remodeling, which is an alternative interpretation consistent with the present stretch amplitude results. Looking at endothelial cells, they report deformations associated with subthreshold force of ~5%, equivalent to a peak-to-peak amplitude of 10% in the present study, and over-threshold deformations of ~13%, equivalent to 26% peak-to-peak. The results Mack and colleagues obtain with cyclic loading could also be interpreted as consistent with the present results. They observe a reduction in translocation of focal adhesions subjected to cyclic loading with increasing frequency over the frequency range used in the present study and suggest that this reflects preferential loading of the elastic component of a viscoelastic response unit. This suggests that similar mechanical events trigger ERK2 phosphorylation and focal adhesion remodeling.

Expression of CAV-3 is important to the growth and maintenance of skeletal muscle. Either disruption or overexpression of CAV-3 results in a dystrophic phenotype (13, 14), so balanced expression of the protein appears to be necessary. The underlying mechanisms of CAV-3-associated pathologies are not entirely clear. Disruption of CAV-3 has been variously associated with exaggerated growth and fusion (60), diminished differentiation and fusion (15), and exaggerated apoptosis (52). If intact CAV-3 were a negative control on growth, then disruption of CAV-3 would be expected to stimulate growth, whereas runaway growth might be expected to activate antagonist mechanisms.

Appropriate expression of CAV-3 does appear to be important for normal muscle development. Disruptions in the CAV-3 gene underlie the family of disorders recently identified as "caveolinopathies," which include limb-girdle muscular dystrophy 1C (LGMD-1C), rippling muscle disease, distal myopathy, and hyperCKemia (65). These diseases are characterized by a loss of CAV-3 expression at the sarcolemma and present a rage of phenotypes that vary in age of onset, severity, and survivability (65). LGMD-1C is characterized by muscle weakness, hypertrophy, and myopathic changes (32, 33), whereas hyperCKemia shows elevated levels of creatine kinase but lacks muscle weakness or observable changes in the muscle histology (6). Targeted disruption of CAV-3 leads to myopathic changes similar to LGMD-1C (13, 18). Overexpression of CAV-3 can also have adverse effects in skeletal muscle. Transgenic overexpression of CAV-3 leads to a dystrophic phenotype (14), and both mdx mice and Duchenne muscular dystrophy humans show a compensatory increase in CAV-3 expression in their muscles and increased density of caveolae (44, 58).

The principal observations from the present study support the involvement of a length- and velocity-dependent mechanism in mechanotransduction, which may be mimicked superficially by disruption of caveolae but suggest that CAV-3 is not essential to that mechanism. The conceptual model remains extremely appealing, and, given the limited reduction of CAV-3 expression obtained by siRNA-mediated knockdown, we do not believe that the results definitely disprove the hypothesis. An experimental model that reduces CAV-3 expression to 10% of normal or caveolae density to fewer than 2/µm2 is required, but knockout and mutant models suffer from alterations in differentiation and fusion that may make them difficult to compare with wild types.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
ACKNOWLEDGMENTS
 REFERENCES
 
This work was supported by a grant from the National Institute of Arthritis and Musculoskeletal and Skin Diseases (AR-48664).


   ACKNOWLEDGMENTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
REFERENCES
 
The authors are grateful for the insightful comments of Dr. Hanjoong Jo and Dr. Alfred Merrill.


   FOOTNOTES
 

Address for reprint requests and other correspondence: T. J. Burkholder, School of Applied Physiology, Georgia Institute of Technology, 281 Ferst Dr., Atlanta, GA 30332-0356 (E-mail: thomas.burkholder{at}ap.gatech.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.


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 ACKNOWLEDGMENTS
 REFERENCES
 

  1. Aronson D, Violan MA, Dufresne SD, Zangen D, Fielding RA, and Goodyear LJ. Exercise stimulates the mitogen-activated protein kinase pathway in human skeletal muscle. J Clin Invest 99: 1251–1257, 1997.[Web of Science][Medline]
  2. Baar K and Esser K. Phosphorylation of p70S6k correlates with increased skeletal muscle mass following resistance exercise. Am J Physiol Cell Physiol 276: C120–C127, 1999.[Abstract/Free Full Text]
  3. Boyd NL, Park H, Yi H, Boo YC, Sorescu GP, Sykes M, and Jo H. Chronic shear induces caveolae formation and alters ERK and Akt responses in endothelial cells. Am J Physiol Heart Circ Physiol 285: H1113–H1122, 2003.[Abstract/Free Full Text]
  4. Burkholder TJ. Permeability of C2C12 myotube membranes is influenced by stretch velocity. Biochem Biophys Res Commun 305: 266–270, 2003.[CrossRef][Web of Science][Medline]
  5. Burkholder TJ, Fingado B, Baron S, and Lieber RL. Relationship between muscle fiber types and sizes and muscle architectural properties in the mouse hindlimb. J Morphol 221: 177–190, 1994.[CrossRef][Web of Science][Medline]
  6. Carbone I, Bruno C, Sotgia F, Bado M, Broda P, Masetti E, Panella A, Zara F, Bricarelli FD, Cordone G, Lisanti MP, and Minetti C. Mutation in the CAV3 gene causes partial caveolin-3 deficiency and hyperCKemia. Neurology 54: 1373–1376, 2000.[Abstract/Free Full Text]
  7. Carson JA and Wei L. Integrin signaling's potential for mediating gene expression in hypertrophying skeletal muscle. J Appl Physiol 88: 337–343, 2000.[Abstract/Free Full Text]
  8. Chang F, Steelman LS, Lee JT, Shelton JG, Navolanic PM, Blalock WL, Franklin RA, and McCubrey JA. Signal transduction mediated by the Ras/Raf/MEK/ERK pathway from cytokine receptors to transcription factors: potential targeting for therapeutic intervention. Leukemia 17: 1263–1293, 2003.[CrossRef][Web of Science][Medline]
  9. Clark CB, Burkholder TJ, and Frangos JA. Uniaxial strain system to investigate strain rate regulation in vitro. Rev Sci Instrum 272: 2415–2422, 2001.[CrossRef]
  10. Clarke MS, Khakee R, and McNeil PL. Loss of cytoplasmic basic fibroblast growth factor from physiologically wounded myofibers of normal and dystrophic muscle. J Cell Sci 106: 121–133, 1993.[Abstract]
  11. Dulhunty AF and Franzini-Armstrong C. The relative contributions of the folds and caveolae to the surface membrane of frog skeletal muscle fibres at different sarcomere lengths. J Physiol 250: 513–539, 1975.[Abstract/Free Full Text]
  12. Engelman JA, Chu C, Lin A, Jo H, Ikezu T, Okamoto T, Kohtz DS, and Lisanti MP. Caveolin-mediated regulation of signaling along the p42/44 MAP kinase cascade in vivo. A role for the caveolin-scaffolding domain. FEBS Lett 428: 205–211, 1998.[CrossRef][Web of Science][Medline]
  13. Galbiati F, Engelman JA, Volonte D, Zhang XL, Minetti C, Li M, Hou H Jr, Kneitz B, Edelmann W, and Lisanti MP. Caveolin-3 null mice show a loss of caveolae, changes in the microdomain distribution of the dystrophin-glycoprotein complex, and t-tubule abnormalities. J Biol Chem 276: 21425–21433, 2001.[Abstract/Free Full Text]
  14. Galbiati F, Volonte D, Chu JB, Li M, Fine SW, Fu M, Bermudez J, Pedemonte M, Weidenheim KM, Pestell RG, Minetti C, and Lisanti MP. Transgenic overexpression of caveolin-3 in skeletal muscle fibers induces a Duchenne-like muscular dystrophy phenotype. Proc Natl Acad Sci USA 97: 9689–9694, 2000.[Abstract/Free Full Text]
  15. Galbiati F, Volonte D, Engelman JA, Scherer PE, and Lisanti MP. Targeted down-regulation of caveolin-3 is sufficient to inhibit myotube formation in differentiating C2C12 myoblasts. Transient activation of p38 mitogen-activated protein kinase is required for induction of caveolin-3 expression and subsequent myotube formation. J Biol Chem 274: 30315–30321, 1999.[Abstract/Free Full Text]
  16. Galbiati F, Volonte D, Minetti C, Chu JB, and Lisanti MP. Phenotypic behavior of caveolin-3 mutations that cause autosomal dominant limb girdle muscular dystrophy (LGMD-1C). Retention of LGMD-1C caveolin-3 mutants within the golgi complex. J Biol Chem 274: 25632–25641, 1999.[Abstract/Free Full Text]
  17. Goldspink G, Williams P, and Simpson H. Gene expression in response to muscle stretch. Clin Orthop 403, Suppl: S146–S152, 2002.
  18. Hagiwara Y, Sasaoka T, Araishi K, Imamura M, Yorifuji H, Nonaka I, Ozawa E, and Kikuchi T. Caveolin-3 deficiency causes muscle degeneration in mice. Hum Mol Genet 9: 3047–3054, 2000.[Abstract/Free Full Text]
  19. Huet de la Tour E, Tardieu C, Tabary JC, and Tabary C. Decrease of muscle extensibility and reduction of sarcomere number in soleus muscle following a local injection of tetanus toxin. J Neurol Sci 40: 123–131, 1979.[CrossRef][Web of Science][Medline]
  20. Jones NC, Fedorov YV, Rosenthal RS, and Olwin BB. ERK1/2 is required for myoblast proliferation but is dispensable for muscle gene expression and cell fusion. J Cell Physiol 186: 104–115, 2001.[CrossRef][Web of Science][Medline]
  21. Kawabe J, Okumura S, Lee MC, Sadoshima J, and Ishikawa Y. Translocation of caveolin regulates stretch-induced ERK activity in vascular smooth muscle cells. Am J Physiol Heart Circ Physiol 286: H1845–H1852, 2004.[Abstract/Free Full Text]
  22. Kawamura S, Miyamoto S, and Brown JH. Initiation and transduction of stretch-induced RhoA and Rac1 activation through caveolae: cytoskeletal regulation of ERK translocation. J Biol Chem 278: 31111–31117, 2003.[Abstract/Free Full Text]
  23. Koh TJ and Herzog W. Excursion is important in regulating sarcomere number in the growing rabbit tibialis anterior. J Physiol 508: 267–280, 1998.[Abstract/Free Full Text]
  24. Lexell J, Taylor CC, and Sjostrom M. What is the cause of the ageing atrophy? Total number, size and proportion of different fiber types studied in whole vastus lateralis muscle from 15- to 83-year-old men. J Neurol Sci 84: 275–294, 1988.[CrossRef][Web of Science][Medline]
  25. Li S, Couet J, and Lisanti MP. Src tyrosine kinases, G{alpha} subunits, and H-Ras share a common membrane-anchored scaffolding protein, caveolin. Caveolin binding negatively regulates the auto-activation of Src tyrosine kinases. J Biol Chem 271: 29182–29190, 1996.[Abstract/Free Full Text]
  26. Loughna P, Goldspink G, and Goldspink DF. Effect of inactivity and passive stretch on protein turnover in phasic and postural rat muscles. J Appl Physiol 61: 173–179, 1986.[Abstract/Free Full Text]
  27. Lutz GJ and Rome LC. Built for jumping: the design of the frog muscular system. Science 263: 370–372, 1994.[Abstract/Free Full Text]
  28. Mack PJ, Kaazempur-Mofrad MR, Karcher H, Lee RT, and Kamm RD. Force-induced focal adhesion translocation: effects of force amplitude and frequency. Am J Physiol Cell Physiol 287: C954–C962, 2004.[Abstract/Free Full Text]
  29. Maniotis AJ, Chen CS, and Ingber DE. Demonstration of mechanical connections between integrins, cytoskeletal filaments, and nucleoplasm that stabilize nuclear structure. Proc Natl Acad Sci USA 94: 849–854, 1997.[Abstract/Free Full Text]
  30. Martineau LC and Gardiner PF. Insight into skeletal muscle mechanotransduction: MAPK activation is quantitatively related to tension. J Appl Physiol 91: 693–702, 2001.[Abstract/Free Full Text]
  31. McNeil PL and Khakee R. Disruptions of muscle fiber plasma membranes. Role in exercise-induced damage. Am J Pathol 140: 1097–1109, 1992.[Abstract]
  32. Minetti C, Bado M, Broda P, Sotgia F, Bruno C, Galbiati F, Volonte D, Lucania G, Pavan A, Bonilla E, Lisanti MP, and Cordone G. Impairment of caveolae formation and T-system disorganization in human muscular dystrophy with caveolin-3 deficiency. Am J Pathol 160: 265–270, 2002.[Abstract/Free Full Text]
  33. Minetti C, Sotgia F, Bruno C, Scartezzini P, Broda P, Bado M, Masetti E, Mazzocco M, Egeo A, Donati MA, Volonte D, Galbiati F, Cordone G, Bricarelli FD, Lisanti MP, and Zara F. Mutations in the caveolin-3 gene cause autosomal dominant limb-girdle muscular dystrophy. Nat Genet 18: 365–368, 1998.[CrossRef][Web of Science][Medline]
  34. Mobley BA and Eisenberg BR. Sizes of components in frog skeletal muscle measured by methods of stereology. J Gen Physiol 66: 31–45, 1975.[Abstract/Free Full Text]
  35. Nader GA and Esser KA. Intracellular signaling specificity in skeletal muscle in response to different modes of exercise. J Appl Physiol 90: 1936–1942, 2001.[Abstract/Free Full Text]
  36. Okamoto T, Schlegel A, Scherer PE, and Lisanti MP. Caveolins, a family of scaffolding proteins for organizing "preassembled signaling complexes" at the plasma membrane. J Biol Chem 273: 5419–5422, 1998.[Free Full Text]
  37. Palade GE. Fine structure of blood capillaries. J Appl Physiol 24: 1424–1436, 1953.
  38. Park H, Go YM, St John PL, Maland MC, Lisanti MP, Abrahamson DR, and Jo H. Plasma membrane cholesterol is a key molecule in shear stress-dependent activation of extracellular signal-regulated kinase. J Biol Chem 273: 32304–32311, 1998.[Abstract/Free Full Text]
  39. Petrof BJ. Molecular pathophysiology of myofiber injury in deficiencies of the dystrophin-glycoprotein complex. Am J Phys Med Rehabil 81: S162–S174, 2002.[CrossRef][Web of Science][Medline]
  40. Pol A, Calvo M, and Enrich C. Isolated endosomes from quiescent rat liver contain the signal transduction machinery. Differential distribution of activated Raf-1 and Mek in the endocytic compartment. FEBS Lett 441: 34–38, 1998.[CrossRef][Web of Science][Medline]
  41. Prescott L and Brightman MW. The sarcolemma of Aplysia smooth muscle in freeze-fracture preparations. Tissue Cell 8: 248–258, 1976.[Medline]
  42. Rando TA and Blau HM. Primary mouse myoblast purification, characterization, and transplantation for cell-mediated gene therapy. J Cell Biol 125: 1275–1287, 1994.[Abstract/Free Full Text]
  43. Raucher D and Sheetz MP. Characteristics of a membrane reservoir buffering membrane tension. Biophys J 77: 1992–2002, 1999.[Web of Science][Medline]
  44. Repetto S, Bado M, Broda P, Lucania G, Masetti E, Sotgia F, Carbone I, Pavan A, Bonilla E, Cordone G, Lisanti MP, and Minetti C. Increased number of caveolae and caveolin-3 overexpression in Duchenne muscular dystrophy. Biochem Biophys Res Commun 261: 547–550, 1999.[CrossRef][Web of Science][Medline]
  45. Rothberg KG, Heuser JE, Donzell WC, Ying YS, Glenney JR, and Anderson RG. Caveolin, a protein component of caveolae membrane coats. Cell 68: 673–682, 1992.[CrossRef][Web of Science][Medline]
  46. Rybin VO, Xu X, and Steinberg SF. Activated protein kinase C isoforms target to cardiomyocyte caveolae: stimulation of local protein phosphorylation. Circ Res 84: 980–988, 1999.[Abstract/Free Full Text]
  47. Schlegel A, Volonte D, Engelman JA, Galbiati F, Mehta P, Zhang XL, Scherer PE, and Lisanti MP. Crowded little caves: structure and function of caveolae. Cell Signal 10: 457–463, 1998.[CrossRef][Web of Science][Medline]
  48. Schnitzer JE, McIntosh DP, Dvorak AM, Liu J, and Oh P. Separation of caveolae from associated microdomains of GPI-anchored proteins. Science 269: 1435–1439, 1995.[Abstract/Free Full Text]
  49. Schnitzer JE, Oh P, Pinney E, and Allard J. Filipin-sensitive caveolae-mediated transport in endothelium: reduced transcytosis, scavenger endocytosis, and capillary permeability of select macromolecules. J Cell Biol 127: 1217–1232, 1994.[Abstract/Free Full Text]
  50. Seger R and Krebs EG. The MAPK signaling cascade. FASEB J 9: 726–735, 1995.[Abstract]
  51. Smart EJ, Graf GA, McNiven MA, Sessa WC, Engelman JA, Scherer PE, Okamoto T, and Lisanti MP. Caveolins, liquid-ordered domains, and signal transduction. Mol Biol Cell 19: 7289–7304, 1999.
  52. Smythe GM, Eby JC, Disatnik MH, and Rando TA. A caveolin-3 mutant that causes limb girdle muscular dystrophy type 1C disrupts Src localization and activity and induces apoptosis in skeletal myotubes. J Cell Sci 116: 4739–4749, 2003.[Abstract/Free Full Text]
  53. Song KS, Li S, Okamoto T, Quilliam LA, Sargiacomo M, and Lisanti MP. Co-purification and direct interaction of Ras with caveolin, an integral membrane protein of caveolae microdomains. Detergent-free purification of caveolae microdomains. J Biol Chem 271: 9690–9697, 1996.[Abstract/Free Full Text]
  54. Song KS, Scherer PE, Tang Z, Okamoto T, Li S, Chafel M, Chu C, Kohtz DS, and Lisanti MP. Expression of caveolin-3 in skeletal, cardiac, and smooth muscle cells. Caveolin-3 is a component of the sarcolemma and co-fractionates with dystrophin and dystrophin-associated glycoproteins. J Biol Chem 271: 15160–15165, 1996.[Abstract/Free Full Text]
  55. Sunada Y and Campbell KP. Dystrophin-glycoprotein complex: molecular organization and critical roles in skeletal muscle. Curr Opin Neurol 8: 379–384, 1995.[Web of Science][Medline]
  56. Tang Z, Scherer PE, Okamoto T, Song K, Chu C, Kohtz DS, Nishimoto I, Lodish HF, and Lisanti MP. Molecular cloning of caveolin-3, a novel member of the caveolin gene family expressed predominantly in muscle. J Biol Chem 271: 2255–2261, 1996.[Abstract/Free Full Text]
  57. Tatsumi R, Hattori A, Ikeuchi Y, Anderson JE, and Allen RE. Release of hepatocyte growth factor from mechanically stretched skeletal muscle satellite cells and role of pH and nitric oxide. Mol Biol Cell 13: 2909–2918, 2002.[Abstract/Free Full Text]
  58. Vaghy PL, Fang J, Wu W, and Vaghy LP. Increased caveolin-3 levels in mdx mouse muscles. FEBS Lett 431: 125–127, 1998.[CrossRef][Web of Science][Medline]
  59. Vandenburgh HH, Shansky J, Solerssi R, and Chromiak J. Mechanical stimulation of skeletal muscle increases prostaglandin F2 alpha production, cyclooxygenase activity, and cell growth by a pertussis toxin sensitive mechanism. J Cell Physiol 163: 285–294, 1995.[CrossRef][Web of Science][Medline]
  60. Volonte D, Peoples AJ, and Galbiati F. Modulation of myoblast fusion by caveolin-3 in dystrophic skeletal muscle cells: implications for Duchenne muscular dystrophy and limb-girdle muscular dystrophy-1C. Mol Biol Cell 14: 4075–4088, 2003.[Abstract/Free Full Text]
  61. Way M and Parton RG. M-caveolin, a muscle-specific caveolin-related protein. FEBS Lett 378: 108–112, 1996.[CrossRef][Medline]
  62. Williams PE and Goldspink G. Changes in sarcomere length and physiological properties in immobilized muscle. J Anat 127: 459–468, 1978.[Web of Science][Medline]
  63. Williams PE and Goldspink G. The effect of immobilization on the longitudinal growth of striated muscle fibres. J Anat 116: 45–55, 1973.[Web of Science][Medline]
  64. Woodman SE, Park DS, Cohen AW, Cheung MW, Chandra M, Shirani J, Tang B, Jelicks LA, Kitsis RN, Christ GJ, Factor SM, Tanowitz HB, and Lisanti MP. Caveolin-3 knock-out mice develop a progressive cardiomyopathy and show hyperactivation of the p42/44 MAPK cascade. J Biol Chem 277: 38988–38997, 2002.[Abstract/Free Full Text]
  65. Woodman SE, Sotgia F, Galbiati F, Minetti C, and Lisanti MP. Caveolinopathies: mutations in caveolin-3 cause four distinct autosomal dominant muscle diseases. Neurology 62: 538–543, 2004.[Abstract/Free Full Text]
  66. Yang S, Alnaqeeb M, Simpson H, and Goldspink G. Cloning and characterization of an IGF-1 isoform expressed in skeletal muscle subjected to stretch. J Muscle Res Cell Motil 17: 487–495, 1996.[CrossRef][Web of Science][Medline]
  67. Zager RA. Plasma membrane cholesterol: a critical determinant of cellular energetics and tubular resistance to attack. Kidney Int 58: 193–205, 2000.[CrossRef][Web of Science][Medline]



This article has been cited by other articles:


Home page
 Journals of Gerontology Series A: Biological Sciences and Medical SciencesHome page
S. E. Riechman, R. D. Andrews, D. A. MacLean, and S. Sheather
Statins and Dietary and Serum Cholesterol Are Associated With Increased Lean Mass Following Resistance Training
J. Gerontol. A Biol. Sci. Med. Sci., October 1, 2007; 62(10): 1164 - 1171.
[Abstract] [Full Text] [PDF]

Home page
 J. Appl. Physiol.Home page
G. C. Sieck
Commentary
J Appl Physiol, April 1, 2005; 98(4): 1541 - 1541.
[Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF) Free
Right arrow All Versions of this Article:
98/4/1554    most recent
01070.2004v1
Right arrow Submit a response
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Web of Science (3)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Bellott, A. C.
Right arrow Articles by Burkholder, T. J.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Bellott, A. C.
Right arrow Articles by Burkholder, T. J.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2005 by the American Physiological Society.