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From the Departments of Molecular Pharmacology and Medicine,* Albert Einstein College of Medicine, Bronx, New York; Muscular and Neurodegenerative Disease Unit, University of Genova and G. Gaslini Pediatric Institute, Genova, Italy; Department of Cancer Biology, Kimmel Cancer Center, Thomas Jefferson University, Philadelphia, Pennsylvania; and H. Houston Merritt Clinical Research Center for Muscular Dystrophy and Department of Neurology, Columbia University, College of Physicians and Surgeons, New York, New York
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Abstract |
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The mechanisms underlying the formation of TAs and their functional significance in skeletal muscle remains unknown. As these structures stain positive with the nicotinamide-adenine dinucleotide (NADH)-tetrazolium reductase reaction, they were initially believed to originate from mitochondria or to consist of mitochondrial aggregates. However, work by several groups has now shown that tubular aggregates arise from the terminal cisternae or longitudinal components of the sarcoplasmic reticulum.13 Interestingly, a recent study has again advocated that a mitochondrial component may be involved in the formation of tubular aggregates,14 clearly illustrating that the pathophysiology of TA formation remains controversial. In addition, several reports have demonstrated that tubular aggregate formation is often found associated with the abnormal proliferation and accumulation of mitochondria in myopathic skeletal muscle fibers.
Unfortunately, the study of the molecular basis of TA formation has been hampered by the lack of a genetic mouse model, demonstrating TA formation as a primary defect. The natural occurrence of TAs in the skeletal muscle was detected in several different mouse strains, including the senescence-accelerated mouse,15 a dystrophic mouse strain,16 and the MLR+/+ substrain mouse.17 In addition, Agbulut et al have reported that TA formation occurs in all inbred mouse strains they examined, in an age- and sex-dependent manner.18 Tubular aggregates were observed only in male mice, with their incidence increasing with age.
In this article, we describe the presence of tubular aggregates and mitochondrial proliferation/aggregation in the skeletal muscle fibers of a mouse model that harbors the genetic ablation of the caveolin (Cav)-2 gene. Our results suggest that loss of Cav-2 function may represent a novel underlying cause for tubular aggregate formation.
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Materials and Methods |
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Antibodies and their sources were as follows. Anti-Cav-1 polyclonal antibody (pAb) (N-20), anti-mitochondrial-specific creatine kinase (sMtCK) pAb (C-18), anti-MCK pAb (N-13), and anti-dihydropyridine receptor (DHPR) pAb (N-19) were from Santa Cruz Biotechnology (Santa Cruz, CA). Anti-Cav-2 monoclonal antibody (mAb) (clone 65) and anti-Cav-3 mAb (clone 26) were generous gifts of Roberto Campos-Gonzalez, BD-Pharmingen (San Diego, CA). Anti-GRP-78 pAb, anti-sarcoplasmic and endoplasmic reticulum calcium ATPase-1 (SERCA-1) mAb, and anti-SERCA-2 mAb were from Affinity Bioreagents (Golden, CO). Anti-calsequestrin mAb and anti-M-cadherin mAb were from BD-Pharmingen. Anti-actin mAb was from Sigma (St. Louis, MO). Anti-heat shock protein 60-kd (HSP-60) pAb was from Stressgen (Ann Arbor, MI). Anti-dicarboxylate carrier (DIC) pAb was the gift of Philipp E. Scherer (Albert Einstein College of Medicine, Bronx, NY).
Animal Studies
All animals were housed and maintained in a pathogen-free environment/barrier facility at the Institute for Animal Studies at the Albert Einstein College of Medicine and at Thomas Jefferson University, under the National Institutes of Health (NIH) guidelines. Cav-1-, Cav-2-, and Cav-3-deficient mice were generated, as we previously described,19-21 and were in the C57Bl/6 genetic background. INK4a(–/–) mice (in the C57Bl/6 background) were used for the generation of the myoblasts and were the generous gift of Dr. Ron DePinho (Harvard Medical School, Boston, MA).
Myoblast Cell Line Generation
One-month-old male INK4a(–/–) mice were sacrificed, and the skeletal muscle was carefully removed under sterile conditions, cleaned of surrounding tissue, and washed in Hanks’ balanced salt solution (HBSS) supplemented with 10 mmol/L 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES). Tissue samples were minced and placed in a 10-ml solution of HBSS-HEPES containing 2 mg/ml collagenase type I (Worthington Biochemical, Freehold, NJ) and agitated for 60 minutes at 
175 rpm at 37°C. Then, the samples were gently centrifuged, and the supernatant was removed by aspiration. Cell pellets were washed twice by centrifugation and plated on 10-cm plastic dishes (Corning, Acton, MA) in Dulbecco’s modified Eagle’s medium containing glutamine, antibiotics (penicillin and streptomycin), and 10% fetal calf serum. The cells were allowed to attach for 24 hours, after which time they were passed and replated. Two days after confluence, the cells were noted to begin differentiating into myotubes. After 4 days in conditioned media, the myotubes began spontaneously contracting.
Electron Microscopy
Male mice from the different genotypes were sacrificed, and the gastrocnemius muscle was removed. Tissue samples were fixed with 2% paraformaldehyde and 2.5% glutaraldehyde in 0.1 mol/L cacodylate buffer, postfixed with OsO4, en bloc-stained with uranyl acetate, and embedded in epoxy resin. Ultrathin sections were examined with a JEOL 1200 EX transmission electron microscope (Tokyo, Japan). Cells from the myoblast cell line were processed for electron microscopy in a similar manner as the tissue samples, except that cells were pelleted before embedding and sectioning.
Immunofluorescence
Male mice from the different genotypes were sacrificed, and the gastrocnemius muscle was removed. Tissue samples were fixed in 4% paraformaldehyde overnight, followed by extensive washing in phosphate-buffered saline (PBS) and cryoprotected in 30% sucrose overnight. The samples were then placed in OCT mounting media (Tissue-Tek; Electron Microscopy Sciences, Hatfield, PA) and snap-frozen in liquid nitrogen-cooled isopentane. Five-micron-thick sections were cut and placed on slides, allowed to adhere overnight, and stored at –80°C. Slides were warmed to room temperature, washed with PBS to remove OCT, and incubated with a blocking solution (PBS, 2% goat serum, 1% bovine serum albumin, and 0.1% Triton X-100) for 20 minutes. Then, sections were incubated with primary antibodies for 1 hour at room temperature at the dilution suggested by the manufacturer. The sections were washed in PBS and incubated in the appropriate secondary antibody (Jackson Laboratories) at a dilution of 1:1000 for 30 minutes. After washing, sections were mounted with Slowfade anti-fade reagent (Molecular Probes, Eugene, OR) and examined under an Olympus IX70 inverted microscope (Tokyo, Japan). Images were acquired using a Sensicam QE cooled CCD camera.
Quantification of Tubular Aggregates Size, Muscle Fibers Areas, and Sarcoplasmic Reticulum Volume
Images of cross sections of gastrocnemius muscle were acquired at x20 magnification. The measurement of tubular aggregate size was performed on cross-sections stained with the SERCA-1 antibody. The measurement of muscle fiber area was performed on cross-sections stained with Cav-3 antibody. Electron micrographs were used to measure the volume of the sarcoplasmic reticulum. Images were analyzed using NIH Image J software, and raw data were imported into Microsoft Excel for statistical analysis and graphic representation. Importantly, sampling for tissue sections and electron microscopy was performed in a blinded fashion.
Immunoblot Analysis
Male mice from the different genotypes were sacrificed, and the gastrocnemius muscle was isolated. Skeletal muscle samples were homogenized in lysis buffer [10 mmol/L Tris, pH 7.4, 1% sodium dodecyl sulfate (SDS), and 1 mmol/L orthovanadate] containing protease inhibitors (Roche Diagnostics, Indianapolis, IN) and centrifuged at 13,000 x g for 10 minutes to remove insoluble debris. Protein concentrations were determined using bicinchoninic acid reagent (Pierce, Rockford, IL). Tissue lysates (35 µg) were separated by SDS-polyacrylamide gel electrophoresis (10% acrylamide) and transferred to nitrocellulose. Membranes were subjected to immunoblot analysis as previously described,22 except primary antibody concentrations were as suggested by the company.
Muscle Regeneration
Age-matched 8-month-old male mice from the different genotypes were anesthetized using ketamine and xylazine. The hair from the lower legs was removed. A small horizontal incision was made to expose the lower portion of the gastrocnemius muscle just above the site of tendon insertion. Ten µl of notexin (Sigma) was injected into the length of the right gastrocnemius muscle at a concentration of 50 µg/ml using a 25-µl Hamilton syringe (Reno, NV). The wound was closed with a surgical staple. The left leg was similarly treated, except 10 µl of saline was injected as a control. The site of injection was marked by dipping the tip of the needle into India ink before injection. Three and 7 days after injection, animals were sacrificed, and muscles were harvested, fixed, and processed for cryosectioning. Fifteen-micron-thick sections were prepared from various regions within the muscle and subjected to immunofluorescence analysis.
Mithocondrial Examination
Morphological analysis of skeletal muscle and staining for cytochrome c oxidase (COX), NADH, and succinate dehydrogenase (SDH) activities were performed, as previously described.23 Biochemical analysis of respiratory chain enzyme activities and citrate synthesis in skeletal muscle homogenates was performed as described.24
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Results |
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Cav-3 is highly expressed at the sarcolemma of mature skeletal muscle fibers, whereas Cav-1 and -2 are clearly absent in adult skeletal muscle fibers.6 Nonetheless, Cav-1 and -2 are thought to be expressed in undifferentiated myotube precursor cells (such as satellite cells and myoblasts). As such, it has been hypothesized that Cav-3 functionally replaces Cav-1 and -2 in fully differentiated adult skeletal muscle fibers. In support of this hypothesis, undifferentiated C2C12 cells, a mouse-derived skeletal muscle cell line, co-express Cav-1 and -2 but fail to express Cav-3. Interestingly, upon differentiation of C2C12 from myoblasts to myotubes, Cav-3 expression increases, but Cav-1 and -2 levels remain unchanged, probably because differentiated C2C12 myotubes retain an embryonic phenotype. Thus, it has never been formally demonstrated that Cav-1 and -2 undergo down-regulation during myoblast differentiation.
To address this issue, we examined the expression levels of caveolin isoforms during skeletal muscle differentiation, using primary cultures of myoblasts derived from INK4a(–/–)-null mice. Importantly, loss of the INK4a cell cycle regulatory proteins (p16 and p19ARF) is sufficient to confer immortalization but not transformation.25
Thus, we used INK4a(–/–)-null skeletal muscle to develop a novel myoblast cell line. We first attempted to evaluate whether this novel myoblast cell line can recapitulate the differentiation process of skeletal muscle in vitro. Under the appropriate conditions, we observe that INK4a(–/–) myoblasts undergo differentiation and spontaneously form myotubes—approximately 2 to 3 days after they reach 100% confluence (Figure 1A)
. Remarkably, when the cell culture dish was gently agitated, many of the myotubes underwent spontaneous contractions or appeared to "beat" (cyclical contractions). We also examined the ultrastructure of these INK4a(–/–) myotubes by transmission electron microscopy. Interestingly, they contain numerous caveolae at the sarcolemma (not shown) and show the development of sarcomeres, with clear Z-lines (Figure 1B)
. Additional evidence for sarcomeric organization within these myotubes was obtained by immunostaining with an antibody that specifically recognizes the sarcomeric myosin heavy chain of vertebrates (mAb MF20). Interestingly, myosin heavy chain exhibited a striated staining pattern in INK4a(–/–) myotubes (Figure 1C)
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. More specifically, during myotube formation, we detected induction of expression of Cav-3, SERCA-1, calsequestrin, 
-actinin, desmin, and -sarcoglycan. In contrast, M-cadherin, a marker of satellite cells, Cav-1, and Cav-2 were all down-regulated during muscle cell differentiation. Finally, the levels of tropomyosin and ß-dystroglycan remained unchanged. These results demonstrate that the novel INK4a(–/–) myoblast cell line represents a valid system for studying the molecular events that occur during skeletal muscle differentiation. In addition, these findings show that skeletal muscle differentiation involves Cav-1 and Cav-2 down-regulation and suggest that the three caveolin isoforms are transiently co-expressed and may interact during myotube differentiation.
Cav-2-Deficient Mice Show Tubular Aggregate Formation in Skeletal Muscle
To assess the physiological role of non-muscle caveolins in skeletal muscle, we next decided to use an in vivo molecular genetic approach. Based on the observations that Cav-1 and Cav-2 expression is elevated in myogenic precursor cells and tightly regulated during muscle differentiation, we would predict that loss of Cav-1 and/or Cav-2 may induce significant disturbances in skeletal muscle.
We first analyzed the effect of a Cav-2 deficiency on skeletal muscle. To this end, we examined by electron microscopy skeletal muscle from 3-month-old Cav-2(–/–)-null mice. It was surprising to observe that Cav-2-deficient skeletal muscle displayed profoundly perturbed skeletal muscle morphology and exhibited tubular aggregate formation. Figure 2
shows that the overall size, shape, and organizational appearance of tubular aggregates can vary widely. Interestingly, when the tubular aggregates occurred in a highly organized fashion, the lumen of the tubules contained an electron-dense core that also appears to be extremely ordered (Figure 2
, inset). However, we could not detect any tubular aggregates in Cav-2-deficient pups (not shown), consistent with the idea that tubular aggregate formation is age-dependent. Importantly, only male mice were affected. In addition, there did not seem to be a relationship between the tubular aggregates and the location of the sarcoplasmic reticulum (SR) in our electron micrographs.
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but not from Cav-3-deficient mice. These results suggest that TA formation is restricted to Cav-1- and Cav-2-null skeletal muscle. In addition, because of the nature of the interaction between Cav-1 and Cav-2, we can attribute this phenotype to Cav-2 loss and not to Cav-1 deficiency. Previous results have demonstrated that in the absence of Cav-1, Cav-2 expression is almost completely abolished. As such, Cav-1-deficient mice are deficient in both Cav-1 and Cav-2.19
However, Cav-2 is not needed for the proper functioning and localization of Cav-1, and the levels of Cav-1 protein are not affected in Cav-2-deficient mice20
(see also Figure 6B
). In addition, we noticed that tubular aggregate formation appears to be restricted to skeletal muscle, since no tubular aggregates were detected in either cardiac or smooth muscle in any of the caveolin knock-out mice examined (data not shown).
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We next evaluated whether loss of Cav-2 induces TA formation, secondary to alterations either in caveolae formation or in the expression levels of the muscle-specific caveolin isoform Cav-3. To detect caveolae formation, we examined high-magnification electron micrographs of WT and Cav-2-deficient skeletal muscle. Figure 4A
shows the presence of numerous caveolae at the plasma membrane of skeletal muscle fibers from both WT and Cav-2-deficient mice, suggesting that loss of Cav-2 does not affect caveolae formation. In addition, Figure 4A
shows that a tubular aggregate structure is clearly visible in the micrograph of the Cav-2-deficient skeletal muscle in close proximity to the plasma membrane. To assess whether loss of Cav-2 alters the expression or distribution of Cav-3, in some way resulting in the development of the tubular aggregates, cryosections from WT and Cav-2-deficient skeletal muscle were subjected to immunofluorescence analysis with an antibody directed against Cav-3. Figure 4B
shows that the relative staining intensity and distribution of Cav-3 is unchanged in Cav-2-deficient mice compared with WT controls. In addition, the same sections were double-immunostained with an antibody against SERCA-1, a known marker of tubular aggregates. Figure 4B
demonstrates the presence of numerous tubular aggregates in the skeletal muscle fibers from the Cav-2-deficient mice. However, we detected few, if any, SERCA-1-positive fibers in WT controls (Figure 4B)
. These data confirm that the development of tubular aggregates is due to the loss of Cav-2 in skeletal muscle fibers and does not correlate with impairment of caveolae formation and/or changes in the expression profile of Cav-3.
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The occurrence of TA formation in male WT mice has previously been reported as an age-related phenomenon.18
Consistent with these findings, during the course of our study we were also able to identify the presence of tubular aggregates in WT mice. However, the frequency and the size of individual aggregates appeared strikingly increased in Cav-2-deficient skeletal muscle compared with age-matched WT controls. To quantify these differences, skeletal muscle sections from three different 8-month-old WT and Cav-2-deficient mice were stained with a SERCA-1 antibody to visualize TA formation (Figure 5A)
. SERCA-1 is the best known marker for the detection of TA formation by immunofluorescence. (However, we cannot completely exclude the possibility that we are also detecting expanded SR.)
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. Figure 5B
shows the distribution of the number of TA in WT and Cav-2-deficient skeletal muscle. Of the 300 WT fibers examined, only 17% contained any aggregates, and two-thirds of these had only one or two aggregates per fiber. On the contrary, over 56% of the Cav-2-deficient fibers contained aggregates and over half of those had three or more aggregates per fiber, with 12 individual fibers containing 10 or more individual aggregates.
In addition, Figure 5C
shows that the size of individual aggregates was increased 2.6-fold in Cav-2-deficient fibers compared with WT fibers. Interestingly, at 3 months of age, WT skeletal muscle contained no detectable TAs, whereas Cav-2-deficient skeletal muscle contained numerous tubular aggregates at this age. These data indicate that the onset of tubular aggregate formation is greatly accelerated in Cav-2-deficient mice compared with WT animals.
Expression Levels and Distribution of Sarcoplasmic Reticulum Markers
The pathophysiology of TA formation still remains controversial. Multiple lines of evidence strongly suggest that TAs originate from the SR, whereas independent researchers advocate for additional mitochondrial components.13,14
To gain insights into this issue, we analyzed the distribution of resident markers of the SR in skeletal muscle sections from 8-month-old age-matched WT and Cav-2-deficient mice. We examined the distribution of two membrane-bound enzymes, SERCA-1 and SERCA-2, of an intraluminal calcium-binding protein, calsequestrin, and, finally, of an SR-specific heat-shock protein, GRP78. Figure 6
shows that all four SR-specific markers (SERCA-1, SERCA-2, GRP78, and calsequestrin) localized to tubular aggregates within the muscle fibers of Cav-2-deficient mice, suggestive of a strong SR component in tubular aggregate formation. Interestingly, the localization of calsequestrin is significantly altered in the skeletal muscle of Cav-2-deficient mice compared with WT counterparts. Figure 6
shows that calsequestrin displayed diffuse cytoplasmic staining, consistent with SR localization in WT muscle fibers, whereas it localized to TAs as well as to the plasma membrane in Cav-2-deficient myofibers.
We next asked whether the presence of TAs would change the expression profile of muscle-specific proteins, including SR resident molecules. For this purpose, skeletal muscle samples from 8-month-old age-matched WT, Cav-1-, and Cav-2-deficient mice were subjected to Western blot analysis with antibodies against 1) SR resident proteins SERCA-1, SERCA-2, GRP78, and calsequestrin; 2) the T-tubule-specific marker dihydropyridine receptor-1 (DHPR); 3) a cytoplasmic muscle enzyme, muscle creatine kinase (M-CK); and 4) the caveolin isoforms.
Figure 6B
shows that SERCA-2 levels appeared increased in Cav-1- and Cav-2-deficient skeletal muscle. These results are consistent with the immunofluorescence data (not shown), revealing that Cav-2-null fibers express SERCA-2 in type I fibers, as expected, but also in TAs in type II fibers (Supplemental Figure 1
, see http://ajp.amjpathol.org). The fact that the expression levels of other SR proteins is not changed in WT and Cav-2-deficient muscle suggests that the presence of TA does not result in an increase in SR proteins but rather induces the relocation of a certain percentage of the endogenous proteins. In addition, we could not detect changes in the expression levels of markers of the cytoplasm M-CK and of the T-tubule system DHPR. Moreover, Cav-3 expression was unaltered in Cav-1- and Cav-2-deficient muscle. Finally, we demonstrate that loss of Cav-2 does not affect Cav-1 expression in muscle, whereas loss of Cav-1 abrogates Cav-2 expression.
The Sarcoplasmic Reticulum Is Dilated in Cav-2-Deficient Skeletal Muscle
To further explore the nature of TA formation, and to corroborate the idea that TAs originate from abnormal dilatation and proliferation of the SR, we evaluated, by electron microscopy, the status of the SR in skeletal muscle samples from 8-month-old WT and Cav-2 KO mice. Figure 7A
shows representative micrographs to illustrate that the SR tubules appear significantly dilated in Cav-2-deficient skeletal muscle compared with their WT counterparts. Note the substantial increase in tubule number and volume in Cav-2-null skeletal muscle.
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reveals a significant increase in the area occupied by the SR in 3- and 8-month-old Cav-2-deficient skeletal muscle compared with WT controls. More specifically, at 3 months of age, the area occupied by the SR increased more than twice in Cav-2-deficient muscle, with 12.9% of the total muscle area being occupied by SR tubules in Cav-2-null myofibers versus 5.8% in WT controls. A similar pattern was evident at 8 months, with 10.1% of total muscle area occupied by the SR in Cav-2 KO mice and 4.7% in their WT counterparts. Interestingly, at 8 months, Cav-1-deficient mice displayed a phenotype similar to Cav-2-deficient mice, with 9.8% of the total muscle area being occupied by SR tubules in Cav-1-null myofibers. However, Cav-2-deficient pups displayed only a small, not statistically different, increase in the muscle area occupied by the SR. It is interesting to note the correlation between the onset of the tubular aggregate formation and of sarcoplasmic reticulum dilation. In fact, the tubular aggregates could be detected in mice at 3 months, a time when SR dilatation seems to be at its highest, but not in newborns, when SR dilatation is not dramatically different between WT and Cav-2-deficient mice, suggesting that SR enlargement may precede TA formation. Cav-2-Deficient Skeletal Muscle Contains Large Mitochondrial Aggregates
During our detailed examination of electron micrographs, we noticed other interesting changes in Cav-2-deficient skeletal muscle. For example, Figure 8A
reveals large mitochondrial aggregates in 3- and 8-month-old Cav-2 KO mice. These aggregates occurred in two distinct patterns. Panels A and B illustrate examples of an amorphous mass of mitochondria tightly packed together, whereas panels C and D are representative of more organized circular assemblies, containing numerous mitochondria, as well as other vacuolar structures. Mitochondrial aggregates were observed within the muscle fiber and, in many instances, disrupted the organization of the sarcomere itself (see panels A and C).
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shows that the expression levels of two mitochondrial markers, DIC and HSP-60, were increased in Cav-1- and Cav-2-deficient muscle compared with their WT counterparts. Similarly to the tubular aggregates and dilated SR, the mitochondrial aggregates could be detected in 3-month-old Cav-2 KO mice but not in newborn pups (data not shown).
To evaluate further the mitochondrial proliferation of Cav-2-deficient mice, skeletal muscle sections from 8-month-old WT and Cav-2 KO mice were subjected to histochemical examination with mitochondrial-specific enzymatic stains, including COX (Figure 9A)
, NADH, SDH, and trichrome staining (Supplemental Figure 1
, see http://ajp.amjpathol.org). Interestingly, staining with the three mitochondrial enzymes COX, NADH, and SDH yielded similar results. Although WT muscle sections presented few areas of intense staining, denoting mitochondrial aggregates in a limited numbers of muscle fibers, almost all Cav-2-deficient muscle fibers had very large and dense areas of staining, suggesting the presence of large mitochondrial aggregates in all of the myofibers. Consistent with the histochemical analysis, trichrome staining demonstrated an obvious increase in TA and mitochondrial aggregates in Cav-2-deficient mice (Supplemental Figure 1
, see http://ajp.amjpathol.org). Interestingly, all muscle fibers with TAs also contained mitochondrial aggregates, whereas other fibers, which did not contain any TAs, still exhibited mitochondrial aggregates. These results suggest a possible correlation between the onset of these two different phenomenons, tubular and mitochondrial aggregates, since they both represent abnormal growth of organelles within the cell.
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represents the mitochondrial activity normalized to citrate synthase activity levels. Interestingly, mitochondrial enzymatic activity levels were normal in Cav-2-deficient muscle, suggesting that Cav-2-deficient mitochondria have the ability to function within normal parameters. Cav-2-Deficient Skeletal Muscle Displays an Increased Number of Satellite Cells
We show here that Cav-2 deficiency profoundly perturbs the morphology of skeletal muscle, by inducing the formation of tubular aggregates and mitochondrial proliferation/aggregation. However, because Cav-2 is not normally expressed in mature skeletal muscle, such muscular phenotypes were very surprising and raised questions regarding the underlying molecular mechanisms. As such, we decided to evaluate the status of skeletal muscle precursor cells, or satellite cells, which, given their undifferentiated status, still express the non-muscle caveolins Cav-1 and Cav-2. For this purpose, we subjected WT and Cav-2-null skeletal muscle sections to immunofluorescence analysis with an antibody against a satellite cell marker M-cadherin. Figure 10A
reveals that Cav-2-deficient myofibers exhibit a significant increase in M-cadherin staining compared with their WT counterparts.
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. We reasoned that increased M-cadherin expression levels could result from abnormal proliferation of M-cadherin expressing cells—satellite cells—in Cav-2-deficient skeletal muscle. Because Cav-2 has been considered an "accessory" protein for Cav-1, this idea is quite novel. There is a single report linking Cav-2 loss of function to increased proliferation: Cav-2-deficient mice were shown to display lung abnormalities, with thickened alveolar septa, with the hyperproliferation of endothelial cells.20
Nevertheless, the role of Cav-2 in the regulation of cell proliferation and growth remains undefined.
To assess directly whether loss of Cav-2 induces the abnormal proliferation of satellite cells, we performed a double labeling experiment with antibodies against M-cadherin and Cav-1, another marker of satellite cells, on skeletal muscle sections from WT and Cav-2 KO mice. Then, we monitored the number of satellite cells by counting the numbers of M-cadherin- or Cav-1-positive cells. Figure 10B
shows a clear increase in the number of M-cadherin- and Cav-1-positive cells in Cav-2-deficient muscle. In addition, note the close co-localization of Cav-1 and M-cadherin within the skeletal muscle. We then proceeded to quantify the number of satellite cells in WT and Cav-2 KO skeletal muscle sections. Interestingly, Figure 10C
shows that Cav-2-deficient skeletal muscle displays an increase of 37 and 40% in Cav-1-positive cells and in M-cadherin-positive cells, respectively. These results clearly suggest that loss of Cav-2 disrupts anti-proliferative signals within satellite cells and induces their abnormal proliferation.
Such significant increases in the number of satel-lite cells could lead to gross abnormalities of the skeletal muscle itself, for example an increase in muscle fiber size. To test this idea, we measured the cross-sectional area of over 600 WT and Cav-2-deficient muscle fibers. Figure 10D
reveals that the cross-sectional area of the Cav-2-deficient muscle fibers was significantly enlarged compared with WT fibers (311 versus 249 µm2).
Cav-2-Deficient Skeletal Muscle Displays a Delay in the Regeneration Process
As Cav-2-deficient skeletal muscle shows satellite cell abnormalities (increased cell number and increased M-cadherin expression), we next examined the capacity of Cav-2-deficient satellite cells to undergo differentiation in vivo by studying their regenerative response to a myotoxic substance. The gastrocnemius muscles from age-matched 8-month-old WT and Cav-2-null male mice were injected with a neurotoxic drug, notexin, and harvested either 3 or 7 days after injury. To evaluate their regeneration status, we monitored M-cadherin expression. Figure 11
(top panels) shows that, 3 days after injury, many M-cadherin-positive cells were present both in WT and in Cav-2-deficient skeletal muscle, suggesting that satellite cells are similarly proliferating and that the regenerative process has started in a similar fashion, both in WT and in Cav-2-deficient mice. However, 7 days after injury, the regenerative process was far advanced in WT skeletal muscle compared with their Cav-2-deficient counterparts. Figure 11
(bottom panels) shows the presence of multiple individual M-cadherin-positive fibers in WT sections, suggesting that WT satellite cells have the ability to fuse and generate mature muscle fibers. On the contrary, Cav-2-deficient skeletal muscle still displayed features of damage, with M-cadherin staining being less well defined and few, if any, individual muscle fibers detected. Taken together, these results suggest that Cav-2 ablation induces the abnormal proliferation of satellite cells, which have an impaired ability to undergo proper regeneration after injury.
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Discussion |
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TopAbstractMaterials and MethodsResultsDiscussionReferences |
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The principal structural proteins of caveolae membranes belong to the caveolin gene family, which consists of three members, caveolin-1, -2, and -3.36-39 Caveolin-1 (Cav-1) and caveolin-3 (Cav-3) share a relatively high degree of identity, whereas caveolin-2 (Cav-2) is the most divergent member of the family. Interestingly, despite the homology between Cav-1 and Cav-3 genes, their expression profiles are vastly different. Endothelial cells, adipocytes, fibroblasts, type I pneumocytes, and epithelial cells highly co-express Cav-1 and -2, whereas Cav-3 is restricted to muscle cell types. Interestingly, smooth muscle cells are the only cell type in which all three proteins are co-expressed, possibly because of its hybrid fibroblast/muscle-like nature. In contrast, Cav-2 expression is directly dependent on Cav-1 expression. Cav-2 requires the presence of Cav-1 for proper membrane targeting and stabilization, such that in the absence of Cav-1, Cav-2 is retained in the endoplasmic reticulum (ER)/Golgi complex and undergoes degradation through a proteaosomal pathway.19,40 However, Cav-1 expression and function do not rely on Cav-2 expression.
Because Cav-2 has been considered an "accessory" protein that functions in conjunction with Cav-1, the physiological role of Cav-2 remains elusive. The Cav-2 protein sequence contains few of the conserved regions that are believed to be involved in the known functions of Cav-1; ie, membrane attachment,41 formation of caveolae,42 and compartmentalization and inhibition of signaling molecules.43 Currently, the only known physiological roles of Cav-2 are hetero-oligomerization with Cav-144 and pulmonary dysfunction observed on Cav-2 deletion in mice.20
Phenotypic Characterization of Cav-2 Knockout Skeletal Muscle
In the present study, we describe new unexpected phenotypes in the skeletal muscle of Cav-2-deficient mice. The presence of mitochondrial aggregates and of tubular aggregates in skeletal muscle fibers of male Cav-2 mice at a young age (as early as 3 months) is surprising given that Cav-2 is not expressed in adult skeletal muscle. However, both Cav-1 and Cav-2 are expressed in myoblasts, before terminal differentiation, and in myotube precursor cells, or satellite cells.
The formation of tubular aggregates is not dependent on Cav-3 expression or the formation of caveolae in skeletal muscle, since both seem normal in the Cav-2-deficient mice. In addition, our work clearly demonstrates that tubular aggregates originate mainly from the sarcoplasmic reticulum. This is supported by the fact that TAs in Cav-2-deficient mice contain known SR resident proteins, such as SERCA-1, SERCA-2, GRP-78, and calsequestrin. Interestingly, we also show that loss of Cav-2 induces the mis-localization of calsequestrin from the SR to the plasma membrane.
Early during myofiber development, peripheral coupling between the SR and the plasma membrane occurs. For example, the DHPR is transiently expressed on the plasma membrane as well as on SR membranes.45,46
Because Cav-2 is expressed at the plasma membrane in myogenic precursor cells, it is possible that loss of Cav-2 affects the development or maturation of muscle fibers, resulting in the altered localization and expression of SR resident proteins. On the other hand, Cav-2 may be required for the proper localization of certain proteins, such as calsequestrin, and in the absence of Cav-2, calsequestrin is mislocalized to the plasma membrane. For example, in muscular dysgenesis mice, the 1 subunit of the DHP receptor is lacking, resulting in mistargeting of the 2 subunit to the plasma membrane instead of the triad junction.47
Further work is needed to address directly the relationship between Cav-2 expression and calsequestrin localization.
In all previous studies, including the current study, the appearance of TAs occurs in an age-dependent manner. Although Agbulut et al have shown that many inbred mouse strains examined develop TAs, they do not become abundant until 10 months of age.18
The senescence-accelerated SAMP8 mouse and the MRL+/+ mouse develop TAs as early as 6 months.15, 17
However, we show here that Cav-2-deficient mice develop TAs as early as 3 months of age in conjunction with severely dilated SR tubules. As such, Cav-2 KO mice constitute the first well-defined genetic model to study the pathogenesis of tubular aggregate formation.
Tubular Aggregates and Calcium Homeostasis
Several lines of evidence suggest that a defect in Ca2+ homeostasis is responsible for the formation of TAs: 1) the SR functions in intracellular Ca2+ homeostasis; 2) TAs have been shown to have calcium-loading capabilities;13 3) several human skeletal muscle conditions, known to exhibit TA formation, including periodic paralysis and myalgia/cramps syndrome, display perturbed intrasarcoplasmic Ca2+ homeostasis;48, 49 and 4) treatment with Dantrolene, a muscle-specific relaxant that diminishes Ca2+ release from the SR, has been shown to provide beneficial effects in patients with a variety of neuromuscular disorders known to contain TAs.50 Additional indications for a correlation between TA formation and Ca2+ homeostasis come from the observations that there is a pronounced difference in Ca2+ uptake between type I and type II muscle fibers, that type II fibers have a higher volume percentage of SR (see review51 ), and that TA formation is only seen in type II fibers. In addition, we show that TA formation correlates with enlargement of the SR in Cav-2-deficient mice. Taken together, these findings support the idea that TAs may represent an adaptive mechanism to compensate for increased intracellular levels of calcium and to avoid muscle fibers hypercontraction and necrosis. In response to the increase in intracellular levels of Ca2+, the terminal cisternae of the SR would undergo hypertrophy with a subsequent rearrangement into TAs in an attempt to act as a calcium sink and prevent irreversible contraction of the fiber or cell death.
How does a Cav-2 deficiency relate to calcium signaling and regulation? Several lines of evidence suggest that caveolae constitute a major site of Ca2+ entry. For example, endothelial Ca2+ waves preferentially originate at specific caveolin-rich edges, and molecules that regulate Ca2+ influx into cells, such as the inositol 1,4,5-trisphosphate receptor-like protein, and the plasmalemmal Ca2+-ATPase localize to caveolae.52-54 Although loss of Cav-2 does not affect caveolae formation per se, it may still affect caveolae function in regulating Ca2+ influx into cells. As such, Cav-2-deficient caveolae may provoke alterations in Ca2+ signaling and induce an abnormal accumulation of Ca2+ in the cytosol. In this view, TAs would develop to compensate for the increased intracellular Ca2+ levels. The idea that Cav-2 modulates the dynamics of Cav-1-dependent caveolae is not completely new. For example, Cav-2 phosphorylation on serine 23 and 36 was shown to be necessary for plasma membrane attachment of Cav-1-positive caveolae in prostate cancer cells, suggesting that although not necessary per se for caveolae formation, Cav-2 expression is still needed for specialized caveolar functions.55
Tubular Aggregates and Mitochondrial Dysfunction
Several groups have shown a correlation between mitochondrial dysfunction and the TA formation.14,56 We also show a link between TA formation and mitochondrial abnormalities. As early as 3 months of age, Cav-2-deficient mice develop large mitochondrial aggregates and atypical mitochondrial clusters not seen in age-matched WT controls. Interestingly, TA formation and SR dilatation coincide with the appearance of mitochondrial aggregates, suggesting interdependence between these conditions. Immunoblot analysis of Cav-2-deficient skeletal muscle shows increased levels of DIC and HSP-60 proteins, whereas the levels of other mitochondria-specific markers are unchanged. Mitochondria are extremely sensitive to increased intracellular calcium levels. For example, in Duchenne’s muscular dystrophy and mdx mice, the loss of dystrophin destabilizes the plasma membrane, leading to an increase in intracellular levels of Ca2+57 and subsequent disruption of proper mitochondrial function.58,59
How mitochondrial function and tubular aggregate formation are connected remains unclear. However, the appearance of TAs and mitochondrial aggregates in Cav-2-deficient mice earlier than in any other mouse strain should allow for a detailed examination and identification of the underlying molecular mechanisms. The fact that at 8 months of age all muscle fibers with TA also contain mitochondrial aggregates, whereas other fibers that do not contain any TA still exhibit mitochondrial aggregates, suggests the establishment of a temporal progression between these two phenotypes. As such, we propose that the formation of mitochondrial aggregates may precede the formation of tubular aggregates.
Tubular Aggregates and Skeletal Muscle Regeneration
Skeletal muscle regeneration is a complex, multistep process in which satellite cells are activated, by signals from necrotic muscle cells, to become myoblasts. These myoblasts then replicate and fuse to form myotubes, which then differentiate to become fully formed muscle fibers (see review60 ). We show here that Cav-2 is normally expressed in myogenic precursor muscle cells and that loss of Cav-2 induces abnormally high numbers of satellite cells in skeletal muscle. Our findings are surprising because little is known about the role of Cav-2 in the regulation of cell proliferation. A previous report has shown that Cav-2-null mice display lung abnormalities, with hyperproliferation of lung endothelial cells. That and our current study are the first to suggest that Cav-2 may play a critical role in controlling cell proliferation. Constant satellite cell hyperproliferation and subsequent incomplete differentiation-regeneration cycles may underlie the pathogenesis of tubular and mitochondrial aggregate formation. This hypothesis is corroborated by the fact that tubular and mitochondrial aggregates are absent in newborn mice—in which the defects in differentiation-regeneration cycles have not yet damaged the muscle—and that their numbers increase with aging. In addition, we show here that Cav-2-deficient skeletal muscle displays an obvious delay in regeneration after injury. Interestingly, 3 days after injury, Cav-2-deficient satellite cells have the ability to undergo proliferation but exhibit an inability to form differentiated myofibers. It is very likely that Cav-2-deficient myogenic precursor cells are fully committed into a proliferative pathway and are unable to undergo differentiation, thus impairing their ability to respond to muscle injury and repair skeletal muscle fibers in a proper fashion.
In conclusion, we have identified several novel defects in the skeletal muscle of Cav-2-deficient mice, which seem to be independent of Cav-1 expression levels. These findings open new avenues for further understanding of the physiological functions of Cav-2 in controlling cell proliferation and differentiation in skeletal muscle. In addition, Cav-2-deficient mice will provide an interesting new source for the potential biochemical purification of tubular aggregates.
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Supported by grants from the National Institutes of Health, the Muscular Dystrophy Association, and the American Heart Association (all to M.P.L.).
Supplemental material for this article can be found on http://ajp.amjpathol.org.
Accepted for publication September 20, 2006.
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