Department of Medicine, Baylor College of Medicine, Houston, Texas 77030
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ABSTRACT |
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-Sarcoglycan (ASG) is a transmembrane
protein of the dystrophin-associated complex, and absence of ASG causes
limb-girdle muscular dystrophy. We hypothesize that disruption of the
sarcoglycan complex may alter muscle extensibility and disrupt the
coupling between passive transverse and axial contractile elements in
the diaphragm. We determined the length-tension relationships of the diaphragm of young ASG-deficient mice and their controls during uniaxial and biaxial loading. We also determined the isometric contractile properties of the diaphragm muscles from mutant and normal
mice in the absence and presence of passive transverse stress. We found
that the diaphragm muscles of the null mutants for the protein ASG show
1) significant decrease in muscle extensibility in the
directions of the muscle fibers and transverse to fibers, 2)
significant reductions in force-generating capacity, and 3) significant reductions in coupling between longitudinal and transverse properties. Thus these findings suggest that the sarcoglycan complex serves a mechanical function in the diaphragm by contributing to muscle
passive stiffness and to the modulation of the contractile properties
of the muscle.
diaphragm mechanics; force transmission; mechanics of breathing; respiratory muscle mechanics; transmembrane proteins
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INTRODUCTION |
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LIMB-GIRDLE MUSCULAR
DYSTROPHY (LGMD) is a result of deficiencies in the sarcoglycan
complex and is a disorder of skeletal muscles. -Sarcoglycan (ASG) is
a transmembrane protein situated along the length of fibers of skeletal
and cardiac muscles. ASG is one of at least five glycoproteins that are
essential to the function of the sarcoglycan complex, and its
deficiency results in the downregulation of other glycoproteins.
These sarcoglycans, along with the syntrophins and
- and
-dystroglycans, comprise the dystrophin-glycoprotein complex.
Mutations in the ASG gene cause LGMD type 2D (15), an
autosomal recessive disorder. ASG-deficient mice develop progressive muscular dystrophy and, in contrast to other animal models for muscular
dystrophy, show ongoing muscle necrosis with age, a characteristic of
the human disease. Molecular analysis of the ASG-deficient mice
demonstrated that absence of ASG resulted in less pronounced sarcoglycan complex, sarcospan, and a disruption of the
-dystroglycan association with membrane recessive disorder
(10, 17). ASG is a structural protein that could be a
load-bearing element in the plane of the cell membrane. ASG could be
similar to the extracellular protein, merosin, in that it can transmit
forces, mostly in shear, between the cytoskeleton and the collagen
matrix fibers. Therefore, most probably, force transmission pathways in
both the longitudinal and transverse direction of the myofibers are
disrupted in an ASG-null mouse diaphragm. We wondered whether force
production and transmission are altered in skeletal muscles when the
sarcoglycan complex is disrupted.
In this study, we hypothesize that disruption of the sarcoglycan complex may alter muscle extensibility and disrupt the coupling between passive transverse elements and axial contractile elements in the diaphragm. To test this hypothesis, we measured passive length tension curves in the directions of the fibers and transverse to the fibers of the muscles of the diaphragm and biceps femoris. Furthermore, we measured the isometric contractile properties of the diaphragm in the absence and presence of passive transverse fiber stress.
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METHODS |
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Animals and tissue preparation. The experimental protocols for this study utilized forty-three 129/SvJ ASG-null and normal wild-type mice weighing 16-24 g. The mice were anesthetized with an intravenous injection of pentobarbital (0.5-0.7 ml/kg). Either the diaphragmatic muscle or biceps femoris muscle was excised and immediately immersed into a muscle bath containing a modified Krebs-Ringer solution (in mM: 137 NaCl, 5 KCl, 1NaH2PO4, 24 NaHCO3, 2 CaCl2, 1 MgSO4, pH 7.4) bubbled with 95% O2-5% CO2 (7). The solution was maintained at a temperature of 25°C throughout the muscle preparation and experimental phase. We excised either left hemidiaphragms or left biceps femoris. The muscle of the diaphragm included the origin on the central tendon and insertion on the rib cage. The biceps femoris included the entire muscle fibers from origin to insertions on the muscle tendonous junctions. Four silk suture position markers (7-0 or 8-0 Surgilene) were sutured on the surface of the appropriate muscle. All markers were placed in the central region of the muscle to minimize the contribution of the mechanical effects of muscle attachments. The markers were placed in a square configuration ~1 mm apart from each other. Two pairs of marker were aligned in the direction along the fibers and two transverse to the fibers.
Biaxial mechanical muscle testing. The biaxial tissue testing apparatus was used to apply in-plane uniaxial stress to the tissue along the fiber direction, uniaxial stress applied transverse to the fiber direction, or biaxial stress applied in both directions, along and transverse to the muscle fibers. The description of the biaxial testing apparatus is detailed elsewhere (3). Biaxial loading of a muscle sheet refers to the muscle being subjected to mechanical stresses not only in the direction of the muscle fibers but also in the direction transverse to the fibers.
Measurements of muscle passive mechanical properties. Either costal hemidiaphragms or biceps femoris muscles from eight normal wild-type 129/SvJ mice (weight: 23.6 ± 7.2 g; age: 43.5 ± 15.9 days) and seven 129/SvJ ASG-null mice (weight: 17.8 ± 2.6 g; age: 32.3 ± 2.4 days) were used in these experiments. After the mice were anesthetized, the appropriate muscles were quickly submerged in the oxygenated Krebs-Ringer solution. All mechanical loads were applied in the plane of the muscle sheet. We preconditioned each muscle with five uniaxial lengthening-shortening cycles with the peak tension of 5 g/cm, and quasistatic lengthening-shortening cycles characterized the passive length-tension of the muscle. Muscles were then passively lengthened from unstressed length with a peak tension of about 20 g/cm and then passively shortened until passive force was negligible. The uniaxial stretching maneuver consisted of clamping one end of the muscle sheet at a fixed position and stretching the other end of the muscle. Biaxial maneuvers were only applied to the diaphragm muscle and consisted of passive mechanical stretching of the muscle in the direction transverse to the muscle fibers by either 1 or 2 g and then stretching the sheet axially in the muscle in the fiber direction. Stress was computed as applied force divided by cross-sectional area. Therefore, force was divided by the multiplication of muscle width and unstressed muscle thickness [stress = force/(width × thickness)], where stress is in N/cm2.
Computation of two-dimensional strains.
The strains in the plane of the muscle were computed by the following
procedure. The marked region is divided into triangles with markers
forming the apices. The coordinates of these points in that unstressed
plane of the diaphragm are denoted xi and
yi, (i = 1, 2). The
displacement, ui, from the unstressed state to the deformed state is assumed to be a linear function of position and
is computed as follows
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(1) |
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(2) |
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(3) |
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(4) |
Measurements of contractile properties.
Muscles from seven 129/SvJ wild-type mice (weight: 17.36 ± 4.66 g; age: 30 ± 13 days) and eight ASG-null mice (weight:
17.56 ± 1.66 g; age: 30 ± 11 days) were used. Upon
anesthetizing of the animals, the diaphragm muscle was excised and
placed in the oxygenated muscle bath. The muscle was positioned
horizontally between two stainless steel mesh electrodes, and optimal
length was determined by twitch responses (0.1-ms stimulus duration, super maximal voltage). At optimal length, we tetanically stimulated the muscle at 100 Hz with 90 s of recovery time between
stimulations (super maximal voltage, 0.5-ms pulses, and tetanic train
duration of 500 ms). Tetanic stimulations were repeated during biaxial loading of the muscle sheet. Biaxial loading was achieved by adjusting the muscle to optimal length, and then muscle was stretched in the
direction transverse to the muscle fibers by either 1 or 2 g.
Muscle was then maximally stimulated at 100 Hz, and this sequence of
stimulations during uniaxial and biaxial loading was repeated three
times. Figure 1 demonstrates how the
diaphragm muscle was subjected to a biaxial load before the muscle was
tetanically stimulated. To acquire force frequency curves, a separate
set of experiments using six 129/SvJ normal mice (weight: 20.32 ± 3.72 g; age: 29 ± 3 days) and seven ASG-null mice (weight:
17.59 ± 1.54; age: 30 ± 4 days) was conducted. The
diaphragm muscle was stimulated at 10, 30, 50, 60, and 100 Hz (super
maximal voltage, 0.5-ms pulses, and tetanic train duration of 500 ms)
with 120 s of recovery time between any two consecutive tetanic
stimulations. Two additional force frequency data were acquired during
biaxial loading of the muscle in the presence of either 1 or 2 g
of muscle force applied in the direction transverse to the muscle
fibers. All data for the contractile properties protocols were acquired at 300 Hz using a data acquisition board (model Lab-PC-1200/AI, National Instruments) and LabVIEWsoftware (version 4.0) applied in the
transverse direction to the long axis of the muscle fibers.
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Statistical analysis. Statistical differences between groups were assessed by ANOVA with use of the SAS Procedure "Mixed" Program. The model was a two-factor fixed or random effects model for two groups (ASG-null vs. controls) and two treatments (uniaxial vs. biaxial). A P value of 0.05 was chosen as the acceptable level of significance throughout the analysis of all data.
Thickness measurements. Unstressed muscle thickness measurements were obtained from the excised muscles, a digital image of the muscle surface was generated, and surface area was determined by using Image Tool (version 2.0, http://ddsdx.uthscsa.edu). Excess water was removed from the surface of the tissue with a cotton-tipped swab, and the tissue sample was immediately weighed. Thickness was computed as t = m/Ad, where t is muscle thickness in centimeters, m is muscle mass in grams, A is the surface area of the sample in centimeters squared, and d is the density of the muscle and is equal to 1.06 g/cm3 (5).
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RESULTS |
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Stiffness is increased in the ASG
/
diaphragm.
Data in Fig. 2 show representative
length-tension curves for the ASG
/
and control mice. Both loading
(lengthening) and unloading (shortening) curves are shown. The data
demonstrate that during lengthening, there is a slow and continuous
increase in tension over the range of imposed strains. Both ASG
/
and controls exhibited hysteresis. That is, at the same tension, the muscle exhibited lower mechanical strain on loading than on unloading. It appears that hysteresis is smaller in the ASG
/
muscles compared with normal muscles. Furthermore, the length-tension curve of the
normal mouse diaphragm shifts to the right relative to the length-tension curve in the transverse fiber direction. This suggests that the muscle has greater extensibility in the direction of muscle
fibers than in the transverse plane. The length-tension curves of the
ASG-null diaphragm appear to exhibit similar behavior. Extensibility
ratios (
) for all the ASG-null and normal wild-type mice were
computed at a tension of 5 g/cm. In the direction along the fibers (AF)
of the diaphragm muscle,
is smaller in the ASG-null mice compared
with control mice (ASG
/
AF:
= 1.21 ± 0.07; ASG +/+
AF:
= 1.45 ± 0.14; P < 0.05). In the
direction transverse to muscle fibers (TF),
is smaller in the
ASG-null mice than in the control mice (ASG
/
TF:
= 1.10 ± 0.09; ASG +/+ TF:
= 1.15 ± 0.17;
P < 0.05). At a tension of 5 g/cm, the axial strains
are 24 and 39% for ASG
/
and control mice, respectively, yielding
a compliance ratio of 0.89. A compliance ratio that is less than one
implies that at the same level of applied tension the muscle is less
extensible compared with its counterpart-in this case, between the
muscles of the ASG
/
mice and the control mice. At the same level
of tension, 5 g/cm, the transverse strains for the ASG
/
and normal
wild-type mice are 10 and 15%, respectively, yielding a compliance
ratio of 0.95. These data suggest that muscles lacking ASG are less
extensible and less viscous than in normal mice.
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Loss of coupling between transverse passive stress and contractile
muscle force in the ASG /
mice.
The data in Fig. 5 demonstrate that
biaxial mechanical loading with a transverse force of 1 g
increases muscle tetanic stress only in the normal diaphragm (1 g
biaxial: ASG +/+: 22.04 ± 1.48 N/cm2; ASG +/+:
24.97 ± 1.19 N/cm2; P < 0.05). In
contrast to normal wild-type, differences in tetanic stress in the
ASG-null diaphragm between uniaxial and biaxial loading virtually
vanish regardless of the magnitude of transverse force (1 g biaxial:
ASG
/
16.194 ± 0.78 N/cm2; ASG
/
: 17.12 ± 1.15 N/cm2; 2g biaxial: ASG
/
17.26 ± 1.07 N/cm2). This demonstrates that the effect of transverse
passive stress on the contractile properties in the ASG
/
mice is
negligible, whereas transverse passive stress increases contractile
muscle force.
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DISCUSSION |
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In this study, we found that muscle extensibility is decreased in skeletal muscles from the ASG-null mice. We also found that muscle force-generating capacity is depressed in the diaphragm muscle of these mice. Furthermore, we found that passive transverse stress has no effect on the contractile properties of the diaphragm in ASG-null mice, whereas muscle force is increased in the presence of transverse stress in the normal mouse.
The mouse model used in our study was developed by Liu and Engvall
(17) and essentially follows the autosomal
recessive LGMD exhibited by humans with a primary sarcoglycan gene
defect. Analogous to this mouse model, Duclos et al.
(10) developed an ASG-null mutant mouse that
exhibited matching trends in complex formation and localization of the
sarcoglycan complex. Both models of the null mutant ASG exhibited
histopathological features of autosomal LGMD about 1 wk after birth
with ongoing necrosis until the age of 9 mo. Histological and
immunofluorescence analysis of skeletal muscles in 2-mo-old mice
demonstrated the established sarcoglycan complex in the control mice
and the degeneration of the -sarcoglycan in the ASG-null mice. This
deficiency is also noticed in the
- and
-sarcoglycans, leading to
a disruption in the sarcoglycan complex (17).
The passive and maximally stimulated uniaxial length-tension curves of the diaphragm muscle have been measured (11, 12, 16, 18-22). The length-tension relationship of a homogeneous elastic material is stiffer when subjected to a biaxial than when subjected to a uniaxial load. This is an added complexity in relating the length-tension relationship measured in isolated diaphragm preparations to in vivo measurements. Limited data are available on the diaphragm muscle properties under biaxial loading (4, 5, 13), and the relationship between stress in the transverse direction and the length-tension properties of the muscle in the direction of the fibers is unknown. We measured the longitudinal and transverse strains in the excised (4) and intact (5) diaphragms. Data from these studies have shown that the diaphragm muscle stiffness properties are anisotropic, with a greater stiffness in the transverse fiber direction than that in the fiber direction. If the diaphragm were extensible in the direction transverse to muscle fibers, then during active contraction, as transdiaphragmatic pressure and stress in both directions increase, the diaphragm would expand in the transverse direction as it contracts along the muscle fibers direction (6).
We recently investigated the mechanical role of desmin (3), a cytoskeletal protein in muscles. We demonstrated that desmin integrates the three-dimensional properties of skeletal muscles by coupling the longitudinal and transverse properties of the diaphragm. In our current study, we demonstrated that transverse loads increase muscle maximal contractile force production in the diaphragm, indicating the presence of structures that couple longitudinal and transverse properties in the diaphragmatic muscle. Our data suggest that desmin may not be the only structural protein that integrates the longitudinal and transverse elements in skeletal muscles. The sarcoglycan complex may be another structural element that could transmit muscle force between the longitudinal and transverse elements. ASG is one of the many components of the dystrophin-associated complex, which could function at least in part to transmit forces generated by the sarcomeric proteins across the cell membrane. This group of membrane-associated structural proteins is found to be highly concentrated at the ends of muscle fibers at the muscle tendonous junction where contractile muscle forces and passive forces would be transmitted across the cell membrane (3).
In contrast to desmin-null mice, which show more pronounced muscle extensibility in the transverse plane to the fibers, skeletal muscles from the ASG-null mice are stiffer in the axial, as well as in the transverse, direction to the muscle fibers compared with muscles from age-matched normal wild-type mice. Our findings are in agreement with those published by Duclos et al. (10) on hindlimb muscles. Although data were not presented, the investigators (10) stated that their data on passive stretch of the extensor digitorum longus and soleus (EDL) muscles demonstrated an increase in the resistance to passive stretch in the ASG-null muscles compared with controls.
Reduced muscle compliance has generally attributed to collagen accumulation (2, 14, 23). However, there are other studies that have demonstrated no correlation between the proportion of collagen and passive stiffness in striated muscles. In particular, in the rat soleus muscle, the increased collagen content during ageing is not associated with changes in muscle stiffness (1). Furthermore, diaphragm stiffness is decreased in the cardiomyopathic Syrian hamster, whereas the surface area of collagen was increased in those animals (8). The ASG-null mice develop muscle necrosis on day 7 after birth, and according to the data by Duclos et al. (10), these mice show ongoing muscle necrosis with increasing age. Our experiments were conduced on mice that are about 30 days old. Therefore, we cannot rule out the possibility that the sarcoglycan complex is a load-bearing element in skeletal muscles despite the increased stiffness in the ASG-null mice.
We chose the biceps femoris because in vivo it experiences mechanical loading only along the length of the muscle fibers, whereas the diaphragm experiences mechanical loads not only along the muscle fibers but also transverse to the muscle fibers. Furthermore, the biceps femoris muscle fibers run along the length of the muscle, whereas muscle fibers in the EDL and soleus muscles are oriented at an angle to the long axis of the muscle. Therefore, it is easier to apply a transverse load to the biceps femoris than to either the EDL or soleus muscles. Furthermore, the biceps femoris is a flat muscle, and, therefore, it is easy to apply mechanical stretch in the plane of the muscle sheet.
The increased muscle stiffness in muscle fibers of the diaphragm during
uniaxial loading is demonstrated in data shown in Fig. 2. The
significant shift to the left of the length tension curves of the
diaphragms of ASG /
mice compared with controls demonstrate reduced
muscle extensibility in the directions of the fibers, as well as in the
direction transverse to the fibers. Similar trends are seen in Fig. 3
where the muscle length-tension relationships during biaxial loading of
the ASG
/
diaphragm demonstrate a very significant increase in
stiffness compared with the controls. The extensibility and muscle
compliance of the diaphragm in the ASG
/
is significantly smaller
during biaxial lengthening than during uniaxial lengthening in the
muscle fiber direction. Differences between passive lengthening and
passive shortening data suggest that the muscles exhibit viscoelastic behavior. Our data suggest that hysteresis is smaller in the ASG
/
compared with controls. This suggests that the ASG complex may
contribute to the viscoelastic properties of the muscles. In
particular, it may serve as an energy-dissipating complex, at least
during passive stretching. The effect of ASG deficiency on muscle
stiffness is not specific to the diaphragm; the biceps femoris muscles
are less extensible in the ASG
/
mice. Furthermore, the effect of
ASG deficiency in altering muscle extensibility and stiffness is more
pronounced in the fiber direction than in the transverse fiber
direction (Fig. 4). These data are consistent with a possible
mechanical role of the sarcoglycan complex in modulating passive
stiffness in skeletal muscles.
Data in Figs. 5 and 6 suggest that the sarcolemma of the diaphragm is not capable of transmitting muscle force in the direction transverse to fibers in the ASG mutant mice. This determines a possible role of ASG complex in transmitting muscle force between the longitudinal and transverse elements. ASG appears to contribute to the coupling of the transverse and longitudinal mechanical properties of the diaphragm muscles through structural elements. These results are similar with those reported by us (3) in which muscle force production was not altered during biaxial loading compared with that during uniaxial loading in desmin-deficient mice.
Our data suggest that the sarcoglycan complex may serve a complex mechanical function in the diaphragm by contributing to muscle stiffness, muscle viscoelasticity, and the modulation of the contractile properties of the muscle.
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ACKNOWLEDGEMENTS |
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We thank Dr. Eva Engvall for providing the ASG /
mouse model,
conducting the immunocytochemical labeling analysis, and providing discussion of the data.
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FOOTNOTES |
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This work was supported by National Heart, Lung, and Blood Institute grant HL-63134.
Address for reprint requests and other correspondence: A. M. Boriek, Baylor College of Medicine, One Baylor Plaza, Dept. of Medicine, Pulmonary Section, Suite 520B, Houston, TX 77030 (E-mail: boriek{at}bcm.tmc.edu).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
10.1152/ajpcell.00326.2002
Received 12 July 2002; accepted in final form 27 November 2002.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Alnaqeeb, MA,
ALZaid NS,
and
Goldspink G.
Connective tissue changes and physical properties of developing and ageing skeletal muscle.
J Anat
139:
677-689,
1984[ISI][Medline].
2.
Borg, TK,
and
Caulfield JB.
Morphology of connective tissue in skeletal muscle.
Tissue Cell
12:
197-207,
1980[ISI][Medline].
3.
Boriek, AM,
Capetanaki YG,
Hwang W,
Officer T,
Badshah M,
Rodarte JR,
and
Tidball JG.
Desmin integrates the three-dimensional mechanical properties of muscles.
Am J Physiol Cell Physiol
280:
C46-C52,
2001
4.
Boriek, AM,
Kelly NG,
Rodarte JR,
and
Wilson TA.
Biaxial constitutive relations for the passive canine diaphragm.
J Appl Physiol
89:
2187-2190,
2000
5.
Boriek, AM,
Rodarte JR,
and
Reid MB.
Shape and tension distribution of the passive rat diaphragm.
Am J Physiol Regul Integr Comp Physiol
280:
R33-R41,
2001
6.
Boriek, AM,
Wilson TA,
and
Rodarte JR.
Displacements and strains in the costal diaphragm of the dog.
J Appl Physiol
76:
223-229,
1994
7.
Campbell, KP,
and
Kahl SD.
Association of dystrophin and an integral membrane glycoprotein.
Nature
338:
259-262,
1989[ISI][Medline].
8.
Coirault, C,
Samuel JL,
Chemla D,
Pourny JC,
Lambert F,
Marotte F,
and
Lecarpentier Y.
Diaphragm Compliance is increased in the cardiomyopathic Syrian hamster.
J Appl Physiol
85:
1762-1769,
1998
10.
Duclos, F,
Straub V,
Moore SA,
Venzke DP,
Hrstka RF,
Crosbie RH,
Durbeej M,
Lebakken CS,
Ettinger AJ,
van der Meulen J,
Holt KH,
Lim LE,
Sanes JR,
Davidson BL,
Faulkner JA,
Williamson R,
and
Campbell K.
Progressive muscular dystrophy in -sarcoglycan-deficient mice.
J Cell Biol
142:
1461-1471,
1998
11.
Farkas, GA,
and
Rochester DF.
Functional characteristics of canine costal and crural diaphragm.
J Appl Physiol
65:
2253-2260,
1988
12.
Faulkner, JA,
Maxwell LC,
Ruff GL,
and
White TP.
The diaphragm as a muscle contractile properties.
Am Rev Respir Dis
119:
89-92,
1979[ISI][Medline].
13.
Gates, F,
McCammond D,
Zingg W,
and
Kunov H.
In vivo stiffness properties of the canine diaphragm muscle.
Med Biol Eng Comput
18:
625-632,
1980[ISI][Medline].
14.
Gosselin, LE,
Martinez DA,
Vailas AC,
and
Sieck GC.
Passive length-force properties of senescent diaphragm: relationship with collagen characteristics.
J Appl Physiol
76:
2680-2685,
1994
15.
Hack, AA,
Groh ME,
and
McNally EM.
Sarcoglycans in muscular dystrophy.
Microsc Res Tech
48:
167-180,
2000[ISI][Medline].
16.
Kim, JA,
Walter SD,
Danon J,
Machnach W,
and
Sharp JT.
Mechanics of the canine diaphragm.
J Appl Physiol
41:
369-382,
1976
17.
Liu, LA,
and
Engvall E.
Sarcoglycan isoforms in skeletal muscles.
J Biol Chem
274:
38171-38176,
1999
18.
McCully, KK,
and
Faulkner JA.
Length-tension relationships of mammalian diaphragm muscles.
J Appl Physiol
54:
1681-1686,
1983
19.
Reid, MB,
Feldman HA,
and
Miller MJ.
Isometric contractile properties of diaphragm strips from alcoholic rats.
J Appl Physiol
63:
1156-1164,
1987
20.
Rochester, DF,
and
Farkas GA.
Performance of respiratory muscles in situ.
In: The Thorax, Part B: Applied Physiology, edited by Roussos C.., 1995, p. 1127-1159.
21.
Sant'Ambrogio, G,
and
Saito F.
Contractile properties of the diaphragm in some mammals.
Respir Physiol
70:
349-357,
1970.
22.
Singh, YN,
and
Dryden WF.
Isometric contractile properties and caffeine sensitivity of the diaphragm, EDL and soleus in the mouse.
Clin Exp Pharmacol Physiol
16:
581-589,
1989[ISI][Medline].
23.
Stedman, HH,
Sweeney HL,
Shrager JB,
Maguire HC,
Panattieri RA,
Petrof B,
Narusawa M,
Leferovich JM,
Sladky JT,
and
Kelly AM.
The mdx mouse diaphragm reproduces the degenerative changes of Duchenne muscular dystrophy.
Nature
352:
536-539,
1991[ISI][Medline].