© 1997 American Heart Association, Inc.
Articles |
Dystrophin Is Not a Specific Component of the Cardiac Costamere
From the Imperial College School of Medicine at National Heart and Lung Institute (S.S., S.R., N.J.S.), London, England, and the School of Neurosciences (M.J.C.), University of Newcastle upon Tyne (England).
Correspondence to Prof N.J. Severs, Cardiac Medicine, Imperial College School of Medicine at National Heart and Lung Institute, Royal Brompton Hospital, Sydney Street, London SW3 6NP, England. E-mail n.severs{at}ic.ac.uk
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Abstract |
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Abstract Dystrophin is a key component of the subsarcolemmal skeleton of muscle cells, and lack of dystrophin is the direct cause of Duchenne muscular dystrophy. In skeletal muscle, dystrophin is reported to be localized specifically at costameres, transversely oriented riblike subsarcolemmal plaques that mechanically couple the contractile apparatus to the extracellular matrix. Costameres are characteristically rich in vinculin and are prominent in cardiac as well as skeletal muscle. To define the precise spatial relationship between dystrophin in relation to the costamere in cardiac muscle, we applied high-resolution single- and double-immunolabeling techniques, under a range of preparative conditions, with visualization of vinculin (as a costamere marker) and dystrophin by confocal microscopy and by the freeze-fracture cytochemical technique, fracture label. Immunoconfocal visualization revealed dystrophin as a continuous uniform layer at the cytoplasmic surface of the peripheral plasma membrane of the rat cardiac myocyte at both costameric and noncostameric regions. The pattern of labeling was reproducible with three different antibodies and was independent of time and antibody concentration. Platinum/carbon replicas and thin sections of fracture-label specimens permitted high-resolution visualization of the distribution of dystrophin in plane views of the freeze-fractured plasma membrane and in relation to the sarcomeric banding patterns of the underlying myofibrils. These results confirmed no preferential association of dystrophin with costameres or with any region of the sarcomeres of underlying myofibrils in rat cardiac tissue. We conclude that in contrast to skeletal muscle, dystrophin in cardiac muscle is not exclusively a component of the costamere.
Key Words: costamere • dystrophin • vinculin • confocal microscopy • freeze-fracture cytochemistry
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Introduction |
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Dystrophin, the 427-kD protein product of the Duchenne/Becker muscular dystrophy gene, is a major component of the subsarcolemmal skeleton of muscle cells. The subsarcolemmal skeleton acts as a scaffold at the cytoplasmic surface of the plasma membrane, linking the intracellular cytoskeleton to the extracellular matrix. Dystrophin is tightly associated with a series of transmembrane proteins, the sarcoglycan and dystroglycan complexes, which link externally to laminin, a component of the basal lamina.1 2 3 4 5 The interaction of dystrophin with the cytoskeleton within the cell is mediated via binding to F-actin. Lack of dystrophin due to mutations in the dystrophin gene leads to a gradual but remorseless degeneration of skeletal and cardiac muscle with, in the case of Duchenne muscular dystrophy, fatal consequences for the patient. Since the exact function of dystrophin is not yet understood, the precise cellular mechanism initiating myofiber necrosis has yet to be identified. However, because of its position linking the cytoskeleton to the extracellular matrix, the most accepted current hypothesis regarding the role of the dystrophin/glycoprotein complex is that it has a mechanical function, strengthening the plasma membrane during contraction of the muscle.
The subcellular distribution of dystrophin has been extensively studied in skeletal muscle, where numerous studies have demonstrated its localization at the cytoplasmic surface of the plasma membrane. Initial immunofluorescence studies in skeletal muscle reported a homogeneous distribution beneath the plasma membrane.6 7 8 9 10 However, immunogold localization at the electron microscopic level has suggested a lattice-like organization,11 12 13 14 and a series of more recent immunofluorescence studies, including visualization by confocal microscopy, have reported a regular, nonuniform lattice-like arrangement in which dystrophin is predominantly localized at transversely oriented riblike subsarcolemmal plaques called costameres.15 16 17 18 Costameres, which are found in both skeletal and cardiac muscle, anchor the myofibrils to the plasma membrane,19 20 21 maintain their spatial organization, and serve as sites of mechanical coupling between the contractile apparatus and the extracellular matrix.22 Originally defined by the presence of their high vinculin content, costameres typically contain a range of other proteins, including spectrin, integrins, and desmin.23 24 25
Compared with these investigations of skeletal muscle, fewer studies have investigated dystrophin organization in cardiac muscle, and most have suggested a continuous, uniform distribution at the surface plasma membrane, similar to that described in the earlier investigations of skeletal muscle.9 26 27 The organization of dystrophin specifically in relation to simultaneously identified costameres has not previously been investigated in detail in cardiac muscle. In order to define the precise spatial relationship between dystrophin in relation to the costamere in cardiac muscle, the present study set out to apply high-resolution single- and double-immunolabeling techniques, under a range of preparative conditions, for simultaneous visualization of vinculin and dystrophin by confocal microscopy and freeze-fracture cytochemistry. The results demonstrate that dystrophin in rat cardiac muscle is not uniquely distributed at costameres but is continuously and uniformly distributed at the cytoplasmic surface of the peripheral (ie, nonintercalated disk) plasma membrane.
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Materials and Methods |
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Sources of Tissue
Fifteen male Sprague-Dawley rats (
300 g body wt) were used. The animals were preanesthetized by an intraperitoneal injection of Hypnorm (0.315 mg/mL fentanyl citrate and 10 mg/mL fluanisone, at 0.5 mL/kg body wt) and then anesthetized with intraperitoneal Hypnovel (2.0 mg/kg midazolam hydrochloride) before retrograde perfusion fixation via a catheter in the abdominal aorta. After initial perfusion with heparinized PBS, the hearts were perfused with 2% paraformaldehyde (PBS-buffered, pH 7.4) for 15 minutes. The procedures were conducted according to the Animals (Scientific Procedures) Act, 1986, under license from the Home Office. The fixed heart was removed, and half-ventricle slices were frozen in isopentane cooled with liquid nitrogen for frozen sectioning. For fracture-label electron microscopy, tissue blocks of 3 to 5 mm3 were cryoprotected with 30% PBS-buffered glycerol for 2 hours before mounting and freezing as described below.
Antibodies and Detection Systems
The following three primary antibodies were used for dystrophin labeling: (1) Dy8/6C5, a mouse monoclonal raised against the last 17 amino acids of the COOH terminal domain, (2) P1583, a rabbit polyclonal raised against the same sequence of the COOH terminal domain, and (3) Dy4/6D3, a mouse monoclonal raised against a fusion protein containing a 208–amino acid sequence in the region of exons 26 to 29 (ie, an area near the NH2 terminus of the rod domain; for convenience, referred to here as NH2 terminus antibody). The monoclonals were a gift from Dr Louise Anderson (University of Newcastle upon Tyne); the polyclonal antibody was a gift from Dr Henry Klamut (Ontario Cancer Institute, Toronto, Canada). Western blots confirmed that the antibodies recognized a single band of >400 kD (ie, dystrophin) in cardiac and skeletal muscle (Fig 1
). For vinculin (costamere marker) and 
-actinin, standard commercially available mouse monoclonal antibodies were used (Sigma Chemical Co).
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The secondary antibody/detection systems used for immunoconfocal microscopy were (1) biotinylated goat anti-mouse immunoglobulin and Texas red–streptavidin (Amersham Life Sciences) and (2) goat anti-rabbit FITC (Dako). For fracture-label immunogold electron microscopy, the detection systems were (1) biotinylated goat anti-mouse immunoglobulin used with 10-nm gold–streptavidin complexes (Amersham Life Sciences) and (2) goat anti-rabbit 5-nm gold complexes (British BioCell International).
Immunolabeling for Confocal Microscopy
For immunoconfocal microscopy, 10-µm cryosections were cut at -25°C and thaw-mounted onto poly-L-lysine–coated glass slides. They were treated with 0.3% Triton X-100 for 15 minutes to improve permeability to the reagents, followed by 0.5% bovine serum albumin (as blocking agent) at room temperature. The sections were then incubated for single labeling with anti-vinculin antibody (1:50) overnight or with anti-dystrophin antibody used at concentrations of 1:10, 1:50, 1:100, 1:500, and 1:1000 for 30 minutes, 1 hour, 2 hours, 3 hours, 4 hours, or overnight. For double labeling, sections were exposed sequentially to (1) the polyclonal anti-dystrophin, followed by the monoclonal anti-vinculin, or (2) the polyclonal anti-dystrophin (COOH terminus), followed by the monoclonal anti-dystrophin (NH2 terminus) overnight and then by biotinylated anti-mouse/streptavidin–Texas red (1:250) and anti-rabbit–FITC (1:20) for 1 hour each. The following controls were run in parallel: (1) omission of primary antibody, (2) switching of detection systems (eg, using mouse monoclonal followed by anti-rabbit secondary antibody), and (3) reversing the order of primary antibodies in the double-labeling procedure. The sections were mounted with Citifluor mounting medium.
Confocal Laser Scanning Microscopy
The immunolabeled sections were examined by confocal laser scanning microscopy using a Leica TCS 4D equipped with an argon/krypton laser and fitted with the appropriate filter blocks for the detection of fluorescein and Texas red fluorescence. Double-labeled samples were imaged using simultaneous dual-channel scanning. Both single optical sections and projection views from sets of 10 consecutive single optical sections taken at intervals between 0.6 and 1 µm were examined. All specimens were examined within 24 hours of immunolabeling.
Fracture-Label Electron Microscopy
Fracture-label electron microscopy was carried out by following a procedure modified from that described by Pinto da Silva et al.28 A mounting and fracturing technique was developed to increase the incidence of suitably fractured plasma membranes (Fig 2). This involved manual freeze-fracturing of adhesive-mounted tissue in which the long axes of the myocytes were oriented parallel with the plane of the plastic mounts on either side. Samples of the perfusion-fixed glycerinated rat hearts were sandwiched between small squares of Thermanox coverslips using cyanoacrylate adhesive (Perma Bond C, R.S. Components) and rapidly frozen in liquid nitrogen slush (ie, liquid nitrogen cooled to its melting point). The sandwich was fractured by using a blade under liquid nitrogen and allowed to thaw in precooled 2% paraformaldehyde in 30% glycerol for 5 minutes. The thawed specimens were rinsed in 30% glycerol to remove excess fixative and deglycerinated by passage through 1 mmol/L glycylglycine in 30% glycerol for 5 minutes followed by pure 1 mmol/L glycylglycine for a further 5 minutes.
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Primary antibody treatment for single and double labeling of vinculin and dystrophin (both COOH and NH2 termini) was carried out as described for confocal microscopy, followed by the corresponding secondary antibody–gold complex (1:50, 1 hour at room temperature). Gold markers of 10 nm were used with the monoclonal antibodies, and markers of 5 nm were used with the polyclonal antibodies throughout. In double-labeling experiments, each primary antibody treatment was followed by its corresponding secondary detection system (eg, the following sequence: [1] dystrophin COOH-terminal antibody, [2] anti-rabbit/5 nm gold, [3] monoclonal vinculin, and [4] biotinylated anti-mouse/streptavidin–10 nm gold). Separate experiments in which specimens were single-immunogold–labeled for -actinin followed the same detection procedure as used for vinculin. All specimens were rinsed in PBS, postfixed in 2.5% glutaraldehyde for 30 minutes, further rinsed in PBS, and processed for thin sectioning or platinum/carbon replication. Specimens for thin sectioning were postfixed in OsO4, dehydrated through a graded series of ethanols, and embedded in Araldite. Semithin and ultrathin sections were cut at right angles to the fracture plane using a Riechert E ultramicrotome. For replication, the specimens were partially dehydrated (to 70% ethanol), dried, and mounted fracture side up on the stage of a Balzers BAF 400T unit, and platinum-carbon replicas were prepared at ambient temperature.29 The replicas were carefully cleaned in sodium hypochlorite such that the biological material was removed without dislodging the gold. Sections and replicas were examined using a Philips 301 electron microscope.
Quantitative Analysis of Dystrophin Immunogold Labeling
To examine the distribution of plasma membrane dystrophin in relation to the Z/I-band and A-band regions of the underlying contractile apparatus, 15 to 20 thin-section micrographs of plasma membrane P-face fractures (magnification, x36 000) from each of four separate fracture-label runs of specimens labeled with P1583 and Dy4/6D3 anti-dystrophin antibodies were analyzed. The number of gold particles per unit length of the plasma membrane overlying the Z/I-band region and the A-band region was determined using VIDS III image analysis software (Analytical Measuring Systems). Statistical significance was assessed using the nonparametric Mann-Whitney U test.
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Immunoconfocal Localization
Confocal microscopy of single-labeled sections consistently revealed characteristic patterns of vinculin and dystrophin localization (Figs 3
and 4). Vinculin was distributed in a prominent punctate pattern around the outer circumference of the cells, in the classical positions of the costameres (Fig 3A and 3B). This pattern of vinculin distribution was so sharply defined that it was readily visible in low-magnification survey views (Fig 3A). Vinculin was particularly conspicuous at the transversely oriented portions of the intercalated disks, corresponding to the positions of the fascia adherens junctions but absent from the longitudinal segments of intercalated disk membrane. In higher magnification views (Fig 3B), less prominent striations of vinculin immunoreactivity, in register with the costameres, were visible. This immunolabeling extended as finger-like projections deep into the cell, as confirmed by serial optical sectioning, and was identified as being associated with transverse tubules.
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Dystrophin, by contrast, appeared uniformly distributed over the cell surface; a punctate pattern was never observed (Fig 4
). The dystrophin label was intense and continuous, apart from gaps at the end-on abutments between the cells corresponding to fascia adherens junctions of the intercalated disk membrane. The continuous pattern was consistent irrespective of the antibody concentration or period of incubation and was confirmed in three dimensions by taking serial optical sections through the tissue slice. As with vinculin, higher magnification views disclosed less pronounced labeling within the cell in the form of discontinuous striations or regular punctate patterns (Fig 4B). This signal was weaker than that found in the corresponding position for vinculin and was demonstrated by serial optical sectioning to be organized as finger-like projections within the cell. That this dystrophin labeling was transverse tubular rather than Z-band–associated was further confirmed by its being quite distinct from the immunolabeling pattern for -actinin (not illustrated).
The spatial relationship between the distributions of vinculin and dystrophin was dramatically apparent when the two components were simultaneously visualized by dual-channel imaging of double-label preparations, as illustrated in Fig 5. In this composite figure, the immunolabeling patterns for vinculin and dystrophin are presented separately and simultaneously in longitudinal and transversely sectioned myocytes. As with the corresponding single-labeling experiments, vinculin reveals a clear punctate distribution at the cell surface, whereas dystrophin shows a continuous pattern (Fig 5A through 5C), apparent in both longitudinal and transversely sectioned cells. Simultaneous viewing clearly demonstrated that dystrophin does not localize specifically to the vinculin-rich costameres but has a widespread distribution close to the cell surface, present both at the vinculin-rich punctae and in the intervening regions of membrane. The interior labeling for dystrophin associated with transverse tubules was found to coincide with that of vinculin, although the signal for the latter was the more intense.
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Immunogold Fracture-Label Electron Microscopy
Thin-section examination of fracture-labeled specimens confirmed that vinculin is specifically localized at the plasma membrane overlying the Z disks of the superficial myofibrils, in a position corresponding to the costamere (Fig 6A). No vinculin was detectable in the intervening membrane areas. The label was apparent only on those fractured fragments of tissue containing the P half of the plasma membrane (ie, the half-membrane leaflet attached to the protoplasm32 ). E halves (ie, half-membrane leaflets attached to the extracellular space) were unlabeled. Dystrophin, by contrast, was uniformly distributed at the level of the plasma membrane, with no preferential association with any region of the sarcomere (Fig 6B). This pattern of distribution was confirmed with three different anti-dystrophin antibodies used independently and simultaneously for double labeling (Fig 6B). In both cases, dystrophin labeling was predominantly but not exclusively associated with the plasma membrane P half (gold label sixfold more abundant on the P half than the E half). Cross-fractured myofibrils (ie, within the cell, below the level of the plasma membrane) revealed no detectable dystrophin at the Z/I-band region or any other part of the myofibril (Fig 6C and 6D). Proteins such as -actinin, known to be present at the Z disk, were readily detectable with the same approach (Fig 7).
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Replicas of fracture-labeled specimens provided en face views of dystrophin distribution at the cell surface (Fig 8). In favorable views, the positions of the costameres were discernible as transverse elevations at the surface, resulting from "sinking" of surrounding areas during specimen drying. The lack of any preferential association of dystrophin with the costameres was confirmed in these preparations (Fig 8), as was the absence of label in cross-fractured specimens in which the myofibrils had been exposed (Fig 9).
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Fracture-labeled tissue processed for double-labeling confirmed that vinculin and dystrophin have quite distinctive distributions (Fig 10). As in the single-labeling experiments, dystrophin labeling was observed along the entire lengths of plasma membrane profiles, whereas vinculin labeling was confined to clusters at the Z disk.
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Quantitative analysis of specimens labeled with the two anti-dystrophin antibodies (P1583 and Dy4/6D3) revealed no significant differences in the extent of labeling over A-band versus Z/I-band plasma membrane regions (A band, median 7.86 gold particles/µm; Z/I band, median 7.14 gold particles/µm; P>.05; n=83).
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Discussion |
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In the present investigation, we have applied simultaneous dual-channel scanning immunoconfocal microscopy and complementary double-immunogold electron microscopy to investigate the spatial relationship between dystrophin and costameres in cardiac muscle. For localization at the electron microscopic level, we elected to use the technique of fracture label,28 one of a range of methods in freeze-fracture cytochemistry.30 In fracture label, cytochemical labeling is performed immediately after samples have been freeze-fractured and thawed. Because membranes are split along their hydrophobic interior when fractured at low temperature, the label has unrestricted access to the entire face-on aspects of plasma membranes of cells within the tissue sample. A further feature of the technique, of particular relevance to the present study, is that both integral membrane components and their associated peripheral proteins on the cytoplasmic side of the membrane are rendered accessible for labeling. This happens because upon exposure to aqueous media at the thawing stage, the fractured half-membrane leaflets become reorganized into a discontinuous bilayer, thereby exposing underlying cytoplasmic or extracellular components.31 Fracture label is thus particularly well suited to the investigation of proteins (such as dystrophin and vinculin) that are closely associated with the plasma membrane. Our observation that in fracture-label the dystrophin antibodies labeled the plasma membrane P half rather than the E half indicates that in cardiac muscle, both the carboxy- and amino-terminal domains of dystrophin are closely associated with the protoplasmic side of the membrane, as reported in an earlier fracture-label study in skeletal muscle (in which the carboxy-terminal domain was localized to the P half33 ) and as widely depicted in current models of the dystrophin-glycoprotein complex.3 4 34
The key question we sought to address was whether dystrophin in cardiac muscle is specifically associated with costameres, as reported in skeletal muscle, or whether some other distinctive arrangement characterizes cardiac muscle. Costameres were originally defined in both skeletal and cardiac muscle by the presence of their high vinculin content.19 20 21 22 35 The intense punctate immunofluorescent labeling of vinculin we observed at the plasma membrane in the present study is fully consistent with these earlier observations. Immunogold fracture label confirmed that these patches of vinculin are localized in the characteristic position of the costamere, at the level of the plasma membrane overlying the Z bands of the superficial myofibrils. Other features of vinculin distribution observed in the present study, ie, the presence of high concentrations of vinculin at the fascia adherens junctions of the intercalated disk and vinculin associated with the transverse tubular system penetrating into the cell, accord with the established literature.20 36 37 These comparisons confirm that immunolabeling of vinculin, under the conditions applied in the present study, provided a reliable means for the identification of costameres.
In skeletal muscle, the current consensus from immunofluorescence studies is that dystrophin has a nonuniform distribution at the cytoplasmic surface of the plasma membrane in the form of dense transversely oriented bands at the I/Z-band level (ie, at costameres) linked by finer longitudinally oriented strands, in a pattern that lies in register with -actinin and mirrors that of spectrin and vinculin.15 16 17 Although, in guinea pig muscle, a few small patches of dystrophin label apparently may occur in the absence of vinculin, the two proteins are predominantly colocalized,17 and current models specifically depict dystrophin as a component of the skeletal muscle costamere.34 38
The results of the present study, however, indicate that such models are not universally applicable to cardiac muscle. Instead of having a costameric distribution pattern, our observations indicate that dystrophin appears uniformly distributed at the cytoplasmic surface of the general plasma membrane in rat cardiac muscle. Compared with skeletal muscle, relatively few studies have previously investigated dystrophin organization in cardiac muscle, and none has applied double labeling for vinculin and dystrophin at both the immunoconfocal and immunoelectron microscopic levels to allow the precise spatial localization of dystrophin in relation to the costamere to be defined. Previous reports on cardiac muscle variously describe a continuous or punctate pattern of dystrophin distribution at the surface plasma membrane, and the absence or presence of dystrophin at transverse tubules, although there is general agreement on the lack of dystrophin at the adherens junctions of the intercalated disks.7 9 26 27 39 40 Our present observations on the surface plasma membrane and transverse tubules are in close agreement with those of Frank et al26 and Klietsch et al40 ; although in contrast to the findings of Meng et al,41 we find no evidence for the presence of dystrophin within the Z disks of myofibrils within the cell either by confocal microscopy or fracture-label techniques.
Whereas the demonstration of uniform continuous labeling at the peripheral plasma membrane strongly suggests that dystrophin is ubiquitous at this site, it does not exclude the possibility of local differences in dystrophin abundance. It might be hypothesized, for example, that if dystrophin had a preferential, though nonexclusive, association with costameres, the ability to detect such a relationship would depend critically on preparative conditions. Our demonstration by confocal microscopy that the continuous labeling pattern was demonstrable in rat cardiac myocytes by use of three different antibodies and a wide range of antibody concentration and incubation periods and, in particular, that the continuous labeling was apparent at extremely low antibody concentration and very brief incubations, indicates that, if major local differences exist, they are exceptionally difficult to detect by immunofluorescence. In line with these findings, quantitative analysis of the immunogold results demonstrated no significant difference in the extent of dystrophin labeling in costamere regions versus noncostamere regions of the plasma membrane.
From the point of view of organization of the membrane skeleton, there seems to be no fundamental necessity for dystrophin and vinculin to coexist at the same plasma membrane sites. Immunoconfocal studies on smooth muscle have shown that dystrophin and vinculin are organized in distinct, entirely separate alternating domains.42 Taking this and other published findings together with our present results, the current evidence suggests that the relationship between dystrophin and vinculin may vary in a characteristic and distinctive manner in each muscle type. Whereas in skeletal muscle dystrophin is largely colocalized to the same domains as vinculin, the situation is reversed in smooth muscle, with dystrophin being confined specifically to nonvinculin zones. Cardiac muscle shows yet another distinctive pattern, with dystrophin distributed throughout both the vinculin and nonvinculin domains. Such muscle type–specific features in dystrophin distribution may reflect subtly different roles for dystrophin in myocardium and skeletal muscle that could in turn influence the relative susceptibility of these muscle types to dysfunction in myopathic diseases characterized by deficiencies in dystrophin expression. In Duchenne muscular dystrophy, the loss of dystrophin is as complete in cardiac muscle as it is in skeletal muscle,7 43 but clinically apparent cardiomyopathy, though common, does not normally become evident until relatively late, and in only 10% of cases is death attributable to cardiac failure.44 45 46 Comparable but less severe cardiac abnormalities are apparent in most forms of the clinically milder Becker muscular dystrophy, in which reduced levels or semifunctional forms of dystrophin are expressed.45 46 That dystrophin is ultimately critical to cardiac function, however, is demonstrated by the linkage of mutations in the dystrophin gene to a subset of familial dilated cardiomyopathies that show X-linked inheritance.47 48 Here, mutations specifically affecting dystrophin expression in the heart result in rapidly progressive and fatal heart failure with no or only relatively minor clinical signs of skeletal muscle involvement.47 48 49 50 These findings point to the potential importance of further more detailed investigation of the expression of dystrophin and components of the dystroglycan complex in human cardiomyopathies.
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Acknowledgments |
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This study was supported by grants FS/94044 and PG/93136 from the British Heart Foundation. We thank Steven Coppen for help with the Western blots.
Received September 10, 1996; accepted November 25, 1996.
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References |
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