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(Circulation. 2000;101:2586.)
© 2000 American Heart Association, Inc.
Clinical Investigation and Reports |
Distinct Patterns of Dystrophin Organization in Myocyte Sarcolemma and Transverse Tubules of Normal and Diseased Human Myocardium
From Cardiac Medicine, The National Heart and Lung Institute, Imperial College School of Medicine (R.R.K., S.S., N.J.S.), London, England, and School of Neurosciences (M.J.C.), University of Newcastle upon Tyne, Tyneside, England.
Correspondence to Dr R.R. Kaprielian, Cardiac Medicine, National Heart and Lung Institute, Imperial College, Sydney Street, London SW3 6NP, England.
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Abstract |
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Background—Genetic mutations of dystrophin and associated glycoproteins underlie cell degeneration in several inherited cardiomyopathies, although the precise physiological role of these proteins remains under discussion. We studied the distribution of dystrophin in relation to the force-transducing vinculin-rich costameres in left ventricular cardiomyocytes from normal and failing human hearts to further elucidate the function of this protein complex.
Methods and Results—Single- and double-label immunoconfocal microscopy and parallel high-resolution immunogold fracture-label electron microscopy were used to localize dystrophin and vinculin in human left ventricular myocytes from normal (n=6) and failing hearts (idiopathic dilated cardiomyopathy, n=7, or ischemic heart disease, n=5). In control cardiomyocytes, dystrophin had a continuous distribution at the peripheral sarcolemma, with concentrated bands corresponding to the vinculin-rich costameres. Intracellular labeling extended along transverse (T) tubule membranes. Fracture-label confirmed this distribution, showing significantly greater label on plasma membrane fractures overlying I-bands (I-band 4.1±0.3 gold particles/µm; A-band 3.3±0.2 gold particles/µm: mean±SE, P=0.02). Hypertrophied myocytes from failing hearts showed maintenance of this surface distribution except in degenerating cells; there was a clear increase in intracellular dystrophin label reflecting T-tubule hypertrophy.
Conclusions—Dystrophin partially colocalizes with costameric vinculin in normal and hypertrophied myocytes, a distribution lost in degenerating cells. This suggests a primarily mechanical role for dystrophin in maintenance of cell membrane integrity in normal and hypertrophied myocytes. The presence of dystrophin in the cardiac T-tubule membrane, in contrast to its known absence in skeletal muscle T-tubules, implies additional roles for dystrophin in membrane domain organization.
Key Words: proteins • myocardium • myocyte • heart failure
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Introduction |
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Genetic mutations of dystrophin and dystrophin-associated glycoproteins (DAG), components of the myocyte membrane cytoskeleton, underlie the degeneration of cardiac and skeletal muscle in several inherited myopathies.1 Dystrophin is a 427-kDa protein with binding affinity to cytoskeletal actin; DAG in turn comprises a trans-membrane complex linked laterally by ß-dystroglycan (formerly known as DAG-43) to
-dystroglycan, a
surface membrane receptor for the extracellular matrix component
laminin,2 suggesting that dystrophin-DAG provides a
structural link between the myocyte cytoskeleton and extracellular
matrix. Study of such protein-binding affinities and patterns of muscle
degeneration in patients with dystrophin/DAG mutations have opened new
perspectives on the pathophysiology of the muscle degeneration in these
diseases and in turn permitted progress toward the identification of
the subcellular functions of the individual affected proteins. Investigation into the cellular localization of dystrophin may provide further evidence for the precise functional role of this molecule. Dystrophin distribution in skeletal myocytes has been extensively studied with the use of immunolabeling techniques with light and transmission electron microscopy (EM). Early results suggested a continuous subsarcolemmal layer of dystrophin, with absence in the transverse (T) tubules3 4 5 6 ; subsequent studies, however, report dystrophin in a discontinuous cell surface network involving colocalization with the cytoskeletal-membrane–linking protein vinculin at costameres.7 8 9 10 Costameres, which are found in both cardiac and skeletal myocytes, were first defined by the concentration of vinculin in a series of sarcolemmal riblike bands overlying the Z-bands11 and are now known to contain multiple proteins.12 Acting as anchor points between the myofibrils and the plasma membrane, they are implicated in the lateral transduction of contractile force from the myocyte to the extracellular matrix.13 Degeneration of skeletal muscle of patients with dystrophin mutations may occur because disruption of dystrophin at these points of mechanical stress14 results in mechanical fragility of myocyte membranes.15 In comparison to skeletal muscle, relatively few studies have focused on cardiac muscle,6 12 16 17 18 and little information is available on dystrophin distribution in human cardiomyocytes.4 In rat heart, dystrophin is localized as a continuous sheet at the sarcolemma and, unlike skeletal myocytes, is also found along T-tubules.12 17 19 The association of dystrophin with cardiomyocyte T-tubules,12 16 19 which are important in excitation-contraction coupling and do not serve in the transmission of contractile force, suggests that dystrophin may serve diverse roles in myocyte membranes. Indeed, mechanisms other than membrane fragility, such as defects in calcium handling, have also been proposed as a primary cause for myocyte degeneration in inherited muscular dystrophy.20
In the present study, we aimed to further elucidate the function of this protein complex by application of the complementary techniques of immunoconfocal microscopy and immunogold fracture-label EM21 to establish the cellular distribution of dystrophin in relation to force-transducing vinculin in normal and remodeled human cardiac myocytes.
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Methods |
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Tissue Acquisition
Normal human left ventricular tissue (n=6) was obtained from 2 patients with cystic fibrosis undergoing heart-lung transplantation, whose hearts were not subsequently used for domino procedures. Such hearts typically demonstrate isolated right ventricular hypertrophy secondary to pulmonary hypertension. Four further specimens were acquired from clinically normal hearts initially harvested for use as transplantation donor organs but rejected for technical reasons. The cause of death was massive subarachnoid hemorrhage (n=3) or head trauma (n=1).
Human left ventricular samples were also collected from the
explanted hearts of patients undergoing orthotopic heart
transplantation at The Royal Brompton and Harefield NHS Trust
Hospitals. All patients were severely symptomatic (New York
Heart Association grade III/IV), with poor left ventricular
systolic function recorded on
echocardiography, multiple gated acquisition
scanning, and ventriculography. Patients either had normal
coronary arteries and were free of inherited myopathies
(idiopathic dilated cardiomyopathy, DCM (n=7), or
had severe coronary artery disease (ICM, n=5), with a history
of previous myocardial infarction (5 of 5) and/or coronary
artery bypass surgery (3 of 5). Patient demographic and clinical
details are shown in the Table
.
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The pathological tissue was dissected in the operating theater immediately after explantation and was processed for immunoconfocal microscopy and for thin-section and fracture-label EM. The normal hearts had been in cardioplegic solution on ice for up to 3 hours before tissue was available. The left ventricular free wall was sampled from the normal hearts and from patients with DCM. The macroscopically homogenous nature of the left ventricles of these specimens made directed sampling less important. In the ICM hearts, large areas of full and partial thickness scarring were easily discernible. Tissue was taken, as far as possible, from the ventricular free wall distant from macroscopic scarring. If such tissue was not available, then specimens were taken from alternative ventricular segments away from infarcted territories. The study was approved by the local ethics committee; individual patient consent was obtained.
Antibodies
The following antibodies were used in the study. Dystrophin: (1)
mouse monoclonal antibody (Dr Louise Anderson, University of Newcastle
on Tyne) to the last 17 amino acids of the -COOH
terminal domain of the dystrophin molecule (Dy8/6C5) was diluted 1:1000
for immunofluorescence and used undiluted for
fracture-label; (2) rabbit polyclonal antibody (Dr Henry Klamut,
Ontario Cancer Institute, Toronto, Canada) to the same sequence
(P1583) was diluted 1:100 for double immunolabeling. Vinculin antibody
was obtained from Sigma Immunochemicals (clone No. VIN-11 to 5) and
diluted 1:50. ß-Dystroglycan: mouse monoclonal antibody (43 Dag1/8D5)
produced against the last 16 amino acids of the C-terminal domain (1:50
dilution).22 The secondary antibodies
(immunofluorescence) were (1) goat anti-mouse Cy3
at 1:500 dilution, (2) goat anti-rabbit Cy3 at 1:500 dilution, and (3)
goat anti-mouse FITC at 1:25 dilution (all Chemicon International). For
fracture-label immunogold labeling, biotinylated goat anti-mouse
immunoglobulin (1:50) with 10-nm gold-streptavidin complexes (Amersham
Life Sciences) was used to label dystrophin, and goat anti-mouse 15-nm
gold complexes (British BioCell International) at 1:50 dilution were
used to label vinculin. All antibody dilutions and washes were
performed with 0.5% BSA in PBS containing 0.05% sodium azide. All
incubations were performed at room temperature.
Immunoconfocal Fluorescence Microscopy
Small blocks of tissue were rapidly frozen in liquid
nitrogen–cooled isopentane. Ten-micrometer-thick cryosections were
either initially fixed in freshly prepared 2%
paraformaldehyde for 5 minutes and subsequently blocked
in 0.5% BSA/PBS for 45 minutes or thawed directly in the blocking
solution. Primary antibody was applied (dystrophin 180 minutes;
vinculin and ß-dystroglycan overnight), followed by the appropriate
secondary antibody for 1 hour. Negative controls were performed by
omission of the primary antibody. Dystrophin/vinculin double labeling
was performed on unfixed cryopreserved tissue with a procedure modified
from that above. Sections were incubated with rabbit polyclonal
anti-dystrophin antibody (P1583) overnight, washed, and treated with
mouse monoclonal anti-vinculin antibody for 5 hours. A mixed solution
of the secondary antibodies was applied for 60 minutes before mounting
as described above. Specificity of the labeling was confirmed with
appropriate controls. Sections were examined by confocal microscopy
with a Leica TCS 4D equipped with an argon/krypton laser and
appropriate filter blocks for either single- or dual-channel scanning.
All images were recorded within 24 hours.
Standard Thin-Section EM
Tissue fixed in 2.5% glutaraldehyde in
0.1 mol/L sodium cacodylate buffer, pH 7.3, postfixed for 2 hours in
cacodylate-buffered 2% osmium tetroxide, was processed for standard
thin-section EM. Ultrathin sections were stained with uranyl acetate
and lead citrate before examination.
Fracture-Label EM
A modification of the fracture-label
technique,21 which facilitates plasma membrane fracturing
in cardiac muscle, was used.19 Tissue was divided into
2-mm blocks, fixed for 5 or 15 minutes in freshly made 2%
paraformaldehyde in PBS, incubated overnight in 30%
glycerol/PBS for cryoprotection, and processed as described by
Stevenson et al.19
Quantitative Analysis of Dystrophin Immunogold
Label
The distribution of dystrophin gold label per unit length
of plasma membrane was analyzed in relation to the underlying
sarcomeres. Thin-section images of clearly identified fractures of the
surface membrane (P-half) showing well-preserved underlying sarcomeres
in longitudinal section were recorded at a final magnification of
x35 880. Because of the variable contractile state of the muscle,
individual sarcomere lengths were measured and each sarcomere was then
divided into a central region half of the length of the full sarcomere
(A-band) flanked by 2 equal lengths of membrane (I-band). The number of
gold particles per unit length for each defined A-band and I-band was
determined with the use of VIDS III image analysis software
(Analytical Measuring Systems). Data were expressed as mean±SD for
both groups and compared with the use of the Student’s t
test. Analysis was performed along the whole length of cleanly
fractured membrane to include lengths of high and low labeling.
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Results |
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Dystrophin in Normal Human Myocytes
Immunoconfocal microscopy of normal ventricular tissue revealed consistent patterns of dystrophin distribution. In transverse sections, dystrophin was seen as a thin layer of immunofluorescence at the myocyte sarcolemma, with irregular spokelike extensions penetrating toward the center of the cell (Figure 1A
). In longitudinal
sections, and at higher resolution with the x63 lens, the cell surface
labeling was visualized as bright, regularly spaced foci of dystrophin
label superimposed on a less intense continuous background layer of
label (Figure 1B). The intercalated disks were unlabeled. In
optical sections including a tangential surface view (en face) of the
sarcolemma, periodic broad bands of dystrophin were seen in a direction
perpendicular to the long axis of the cell with finer interconnecting
longitudinal strands (Figure 1C). Intracellular dystrophin was
observed as regular cross-striations corresponding to the spokelike
extensions described in transverse sections. This labeling was
identified as T-tubular because of its occurrence at the Z-line level
penetrating into the cell. A similar pattern of T-tubular labeling was
obtained with antibodies against fibronectin, a recognized component of
the basal lamina within cardiac T-tubules (data not shown). Dystrophin
labeling patterns were unaffected by tissue fixation or with the use of
either the monoclonal and polyclonal anti-dystrophin antibodies. Tissue
distribution of ß-dystroglycan mirrored that of dystrophin (Figure 1D). Continuous labeling with a regular punctate pattern of
denser label could be discerned at the myocyte surface membrane;
T-tubules were again clearly labeled with no labeling of the
intercalated disk.
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Vinculin immunolabeling revealed sarcolemmal distribution in the
characteristic costameric pattern of regularly repeating riblike
structures perpendicular to the long axis of the cell, clear T-tubule
labeling, and strong labeling of the intercalated disks (Figure 2). Double-labeled cells viewed
longitudinally at high resolution showed that the clear foci of
vinculin label at the sarcolemmal surface precisely matched the
punctate pattern observed for dystrophin, with the intervening
vinculin-negative membrane revealing lower though clearly detectable
levels of dystrophin (Figure 3). In en
face membrane views, we observed colocalization of vinculin and
dystrophin over the transverse bands encircling the myocyte, again with
fainter dystrophin label occurring as fine longitudinal strands between
the dense bands. The T-tubular label for dystrophin coincided with that
of vinculin.
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Immunogold Fracture-Label EM
Thin-section examination of fracture-labeled tissue was used
to clarify the distribution of both vinculin (Figure 4) and dystrophin (Figure 5) at the ultrastructural level. In
plasma membrane fractures, vinculin labeling was sparse and closely
associated with the Z-band/sarcolemmal junction (Figure 4); in
contrast, dystrophin showed a wider distribution with more frequent
label (Figure 5). There was a visual impression of a higher
concentration of dystrophin label over the I/Z-bands, with gold
particles occurring in clusters in these regions, compared with single
particles in membrane overlying A-bands. There was a much lower level
of label along the surfaces of fractures that entered the cytoplasm
(cross-fractures); in these cases, the gold particles were observed in
association with fractures exposing the T-tubules. Intercalated disks
lacked dystrophin but had prominent vinculin labeling.
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Fracture-label replicas for dystrophin-labeled tissues, which allow
inspection of large expanses of membrane in plan view, showed prominent
label of surface membrane fractures with minimal label of
cross-fractures (Figure 6). Extracellular
(connective tissue) fractures showed negligible label. The positions of
Z-bands of myofibrils subjacent to the surface membrane could be
identified as ridges; dystrophin labeling patterns confirmed the
findings of the thin-section examination, with gold particles found at
higher density in the regions close to Z-bands. Fracture-label controls
with omission of the primary antibody showed very low levels of
background labeling. Quantitative analysis of dystrophin
immunogold distribution (eg, Figure 5
) confirmed a statistically
significant higher density of gold label over the I/Z-band (4.1±3.4
gold particles/µm; standard error=0.26; n=172) than the A-band
(3.3±2.9 gold particles/µm; standard error=0.23; n=154).
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Dystrophin in Human Heart Failure
Histological examination of tissue from human
end-stage failing hearts revealed myocyte hypertrophy with
varying degrees of interstitial fibrosis irrespective of
underlying cause. Occasional islands of fibrosis suggesting previous
subsegmental infarction were seen in ICM specimens. Myocyte
hypertrophy has been formally measured in an overlapping
group of patients in a separate study; we showed a predominant increase
in cell length in DCM and predominant increase in cell cross-sectional
area in ICM.23
In longitudinally sectioned tissue from both DCM (Figure 7A) and ICM (Figure 7B),
dystrophin was seen in a continuous surface membrane distribution with
a superimposed regular punctate pattern as described in control
tissues. Intracellular dystrophin distribution seen as linear
structures showed increased tortuosity and size (Figures 7, C
and D) and unlike controls were frequently observed to run in a
longitudinal orientation (Figures 7A and 8A). Thin-section EM confirmed that these
structures represented T-tubules (Figure 8): There
were clear increases in size and number of the T-tubules at EM
examination in both DCM and ICM hearts as compared with normal
controls, with prominence of longitudinally orientated elements (Figure 8, B through D).
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Cardiomyocytes adjacent to larger zones of scarring suggesting previous
localized infarction were studied separately from the noninfarcted
tissue. In contrast to the noninfarcted tissue, certain cells
demonstrated an abnormal distribution of sarcolemmal dystrophin, with
loss of the normal punctate pattern seen in longitudinal section
(Figure 9).
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Discussion |
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Genetic mutations of dystrophin and its associated membrane proteins underlie progressive myocyte degeneration in several inherited cardiomyopathies,1 although the precise role of these proteins in muscle physiology remains under discussion. The novel results of the present study, which help to further our understanding of the function of this protein complex, are that (1) dystrophin is partially colocalized with vinculin in the human cardiac myocyte sarcolemma in a costameric distribution but is also present, though at lower density, in the intervening noncostameric surface membrane; (2) dystrophin and vinculin coexist throughout the T-tubular system in normal cardiac myocytes; (3) the distribution of ß-dystroglycan, a marker for the DAG complex, mirrors that of dystrophin; (4) cardiac cell hypertrophy is associated with maintenance of sarcolemmal dystrophin labeling and increased intracellular dystrophin in parallel with more extensive T-tubular development; and (5) the normal sarcolemmal distribution is lost in degenerating cells in the failing left ventricle. These features, established by single- and double-label immunoconfocal microscopy, were extended and confirmed at high resolution by fracture-label immunogold EM.
The costameric pattern of surface membrane dystrophin label observed
here in human cardiac muscle resembles that reported in skeletal muscle
by confocal microscopy.6 7 8 However, not all studies have
found this pattern either in skeletal or cardiac
muscle.3 4 5 This distribution was difficult to discern on
initial examination of the 10-µm sections with conventional
epifluorescence microscopy but was clearly seen with the
greater resolution of immunoconfocal microscopy. Furthermore, we found
that adjustment of the microscope to provide the thinnest optical
sections of 
0.5-µm thickness optimized the images obtained. These
findings may partly explain the earlier reports of human
cardiomyocytes showing a continuous pattern of sarcolemmal
label. The distribution of dystrophin immunolabeling in the present
study is supported by the consistency of results achieved
with the use of 2 separate antibodies and varying tissue fixation and
extended by the findings of qualitative and quantitative fracture-label
EM. Tissue distribution of ß-dystroglycan as a marker of the DAG
complex was found to be identical to that of dystrophin, again
supporting specificity of labeling in the study.
Fracture label is particularly useful for the study of sarcolemma-associated proteins such as dystrophin because the fracturing process exposes hydrophobic internal membrane faces (membrane fractures) and crosses through cells, exposing the underlying cytoplasm (cross-fractures).21 The small absolute differences and high standard deviation in dystrophin label density observed in this study reflect the tendency for fractured hydrophobic surfaces to randomly reassociate into bilayer segments after exposure to aqueous media at the thawing stage.21 Where such bilayer segments occur, access of antibodies to any underlying epitopes are blocked, leaving the areas devoid of label.
Stevenson et al19 recently showed that dystrophin is not confined to costameres in rat cardiac muscle but rather has a continuous and uniform distribution at the cell surface. Our present finding of an increased concentration of dystrophin and ß-dystroglycan at costameres with a lower density over the A-band in human cardiac muscle suggests the existence of species differences in cardiac dystrophin distribution between large and small animals. Previous work has shown that skeletal muscle from the mdx mouse model24 -which, like the muscle of patients with Duchenne muscular dystrophy, lacks dystrophin-sustains less severe degenerative changes.25 26 The higher stress to which human muscle is exposed because of the larger body mass may therefore be important for myocyte damage in Duchenne muscular dystrophy.8 In a similar way the human heart, being larger and generating a higher a higher wall shear stress according to Laplace’s law, may display clustering of dystrophin over the costameric regions of the membrane (the points of highest shear force) as an adaptation to provide the necessary increased mechanical strength. The functional importance of such membrane localization to cell survival is underlined by preservation of the sarcolemmal pattern in hypertrophied myocytes subjected to higher mechanical stress in human heart failure, whereas other cytoskeletal proteins are known to show disrupted patterns.27
The differences in sarcolemmal colocalization of dystrophin with vinculin in our study and in previous reports in both striated19 and smooth muscle cells28 suggest that the function of dystrophin is not entirely associated with the transmission of force, as has been proposed for costameric proteins.13 Furthermore, a nonmechanical function of dystrophin is suggested by its association with the cardiac T-tubule, a structure not directly affected by membrane distortion during contraction.29 The importance of this association is underlined in the current study by preservation of submembrane dystrophin in the extended T-tubules of hypertrophied cells from failing hearts. T-tubular dystrophin previously has been postulated to be important in maintenance of tubular patency during myocyte contraction.12 However, the absence of dystrophin in the skeletal muscle counterpart (distinguished morphologically by a lack of a basement membrane and the presence of triadic junctional complexes) and the consequent differing mechanisms triggering contraction in these muscle cell types suggest an alternative hypothesis of selective association of dystrophin with distinct functional membrane domains. A role for dystrophin in membrane domain organization is also supported by the existence in skeletal muscle of sarcolemmal and T-tubular membrane subcompartments separable by density-gradient centrifugation with variable contents of dystrophin30 and the clustering of dystrophin in the troughs of the muscle end-plate at the neuromuscular junction.31 Accumulating evidence also implicates dystrophin in regulatory interactions with other membrane proteins.22 32
Our results suggest that distinct muscle type and species differences exist for the distribution of the dystrophin membrane cytoskeleton. In human myocardium, dystrophin is distributed at the sarcolemma, with concentration at the lateral costameres suggesting a mechanical role in surface membrane support. The localization of dystrophin in the cardiac T-tubule suggests a cellular function besides that of mechanical support and transmission of force. Further study of dystrophin distribution and molecular interactions is necessary to precisely define the function of this system in human cardiomyocytes in health and disease.
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Acknowledgments |
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We thank the British Heart Foundation for support of this work (grant FS/94073). Dr Cullen is supported by The Wellcome Trust. We thank Professor Sir Magdi Yacoub and Mr Asghar Khaghani at Harefield Hospital for the generous provision of human specimens.
Received May 27, 1999; revision received December 13, 1999; accepted December 22, 1999.
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