1 Department of Human Nutrition, Foods and Exercise, Virginia Polytechnic Institute and State University, Blacksburg, Virginia 24061-0430; 2 Department of Physiology, University of Texas Southwestern Medical Center at Dallas, Dallas, Texas 75235-9040
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ABSTRACT |
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Loss of the dystrophin-glycoprotein
complex from muscle sarcolemma in Duchenne's muscular dystrophy (DMD)
renders the membrane susceptible to mechanical injury, leaky to
Ca2+, and disrupts signaling, but the precise mechanism(s)
leading to the onset of DMD remain unclear. To assess the role of
mechanical injury in the onset of DMD, extensor digitorum longus (EDL)
muscles from C57 (control), mdx, and
mdx-utrophin-deficient [mdx:utrn(/
); dystrophic] pups aged 9-12 days were subjected to an acute
stretch-injury or no-stretch protocol in vitro. Before the stretches,
isometric stress was attenuated for mdx:utrn(
/
) compared
with control muscles at all stimulation frequencies (P < 0.05). During the stretches, EDL muscles for each genotype
demonstrated similar mean stiffness values. After the stretches,
isometric stress during a tetanus was decreased significantly for both
mdx and mdx:utrn(
/
) muscles compared with
control muscles (P < 0.05). Membrane injury assessed
by uptake of procion orange dye was greater for dystrophic compared
with control EDL (P < 0.05), but, within each
genotype, the percentage of total cells taking up dye was not different for the no-stretch vs. stretch condition. These data suggest that the
sarcolemma of maturing dystrophic EDL muscles are resistant to acute
mechanical injury.
Duchenne's Muscular Dystrophy; mice; membrane damage
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INTRODUCTION |
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DUCHENNE'S MUSCULAR DYSTROPHY (DMD) is an X-linked lethal muscle-wasting disease caused by deletions that disrupt the open reading frame of the gene that codes for dystrophin, an integral membrane protein (4, 19). Dystrophin is normally expressed in skeletal, smooth, and cardiac muscle and in brain (19), but its absence in DMD is particularly conspicuous in skeletal and cardiac muscle. DMD is clinically apparent between ages 2 and 3 yr and demonstrates relentless progression that yields muscle weakness and wasting, contractures of the ankles and hips, scoliosis, and premature death, usually from respiratory or cardiac deficiency by age 20 yr (19). Although the genetic reason for this severe disease is known, the mechanism(s) by which the progressive muscle wasting is initiated are not yet clearly defined. One proposed possibility is that the absence of dystrophin leads to a more injury-susceptible sarcolemma (23, 24, 32).
The muscle isoform of dystrophin (427 kDa; see Ref. 4) is localized to the sarcolemma as an integral protein in the dystrophin-glycoprotein complex (DGC; see Ref. 33). The DGC along with additional proteins form rib-like lattices on the cytoplasmic face of the sarcolemma known as costameres that help stabilize the cytoskeleton to the extracellular matrix (ECM; see Refs. 4 and 29). Costameres act as mechanical couplers to distribute contractile forces generated in the sarcomere laterally through the sarcolemma to the basal lamina (8) and thereby facilitate uniform sarcomere length between fibers of active and nonactive motor units (29). Dystrophin is also found in abundance in the myotendinous junction and is thought to facilitate the transmission of forces from the muscle fibers to the tendon (36). Thus dystrophin is considered a key structural element in the muscle fiber.
The absence of expressed dystrophin in DMD can lead to the complete
loss of the DGC (22) and a disrupted costameric lattice (38). Loss of the DGC is considered to render the membrane
less stiff and more readily damaged by mechanical stress
(23). Thus sarcolemmal structural integrity in DMD could
be compromised in early development because of the absence of
dystrophin and continue to degrade with time (19), thereby
representing a major determinant of the onset of the dystrophic
process. Fast fibers are considered most susceptible to damage because
of the greater forces they generate (21). Increased
membrane damage assessed by uptake of a fluorescent dye after
stretch-induced injury has been reported for the fast-twitch extensor
digitorum longus (EDL) muscles from adult mdx
(24) and mdx:utrophin knockout
[mdx:utrn(/
); see Ref. 9] mice compared
with control mice. Although both of these studies demonstrated that the
absence of dystrophin yielded a sarcolemma that was more susceptible to
injury in older dystrophic mice, the age of the animals precluded
drawing conclusions about the role of dystrophin in the onset of muscle
injury. Thus, we considered that, if the absence of dystrophin results
in structurally weak membranes, and this is the primary cause for the
onset of DMD, then a stretch-injury protocol in fast-twitch EDL muscles of young dystrophic mice should induce significant membrane damage. Furthermore, we considered that if utrophin compensated for the absence
of dystrophin to stabilize muscle membranes of mdx mice (14, 39), then mice lacking both dystrophin and utrophin
(11) might demonstrate greater injury.
In this study, we tested the hypothesis that the sarcolemmal membranes
of dystrophic EDL muscles obtained from young mice [mdx and
mdx:utrn(/
) genotypes] are more susceptible to injury after a stretch-injury protocol compared with 1) unstretched
dystrophic EDL muscles and 2) stretched or unstretched
control EDL muscles. We demonstrate that EDL muscles from both
dystrophic models exhibit only modestly greater membrane damage
compared with control muscles. We also report that the stretch-injury
protocol did not induce greater membrane damage in either the control
or dystrophic EDL muscles of young pups aged 9-12 days.
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METHODS |
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Our objective in this report was to determine if the sarcolemma
of EDL muscles from young dystrophic mouse pups were susceptible to
damage after a stretch-injury protocol and thereby could contribute to
the onset of DMD. EDL muscles obtained from control, mdx,
and mdx:utrn(/
) pups aged 9-12 days were subjected
to a stretch-injury protocol and muscle membrane damage assessed with
the fluorescent dye procion orange. In addition, the presence of
centralized nuclei to indicate the presence of regenerating fibers and
the distribution of myosin heavy chain (MyHC) isoforms in the EDL
muscles for each of the genotypes was also determined. The Virginia
Tech Animal Care Committee approved all procedures used in this study.
Mouse genotypes.
Mice were obtained from our colony at Virginia Tech. Breeder pairs of
control C57BL/6 and mdx mice were originally obtained from
Jackson Laboratories, whereas those for the mdx:utrn(/
) mice were a kind gift from Drs. Mark Grady and Joshua Sanes (Washington University, St. Louis, MO). Because the mdx:utrn(
/
) mice
are not fertile (11), mdx:utrn(
/+) male and
female mice were crossed to generate mice deficient in both dystrophin
and utrophin. Mice genotyping was determined by PCR analysis of DNA
obtained from tail snips.
DNA isolation.
Isolation of DNA followed the methods of Laird et al.
(17). Briefly, mouse tail snips (~0.5 cm) were digested
overnight at 55° C in 1 ml lysis buffer [5 mM EDTA, 0.2% SDS, 200 mM NaCl, 100 mM Tris (pH 8.5), and 0.2 mg/ml proteinase K]. Samples
were then spun at 13,000 revolutions/min for 10 min, the supernatant of
each sample transferred to a clean tube, and the DNA was precipitated by adding an equal volume of 100% ice-cold isopropanol. The strands of
precipitated genomic DNA for each sample were transferred to a clean
tube and resuspended in 10 mM Tris-EDTA (pH 7.4). Samples were stored
at 20°C until subsequent PCR analysis.
PCR analysis.
All PCR reagents, including primers, were obtained from GIBCO-BRL.
Specific forward and reverse primers were used to screen the DNA of
each sample for the dystrophin mutation (i.e., mdx; see Ref.
1) and for the utrophin knockout (Dr. Mark Grady, personal
communication). The mdx and wild-type genotypes for
dystrophin were screened in separate tubes with the following reverse
primers: for mdx (259E), 5'-GTCACTCAGATAGTTGAAGCCATTTAA-3';
and, for wild-type dystrophin (260E),
5'-GTCACTCAGATAGTTGAAGCCATTTAG-3'. Both of these reactions used the
same forward primer (9427) 5'-AACTCATCAAATATGCGTGTTAGTG-3' (1). The screen for the utrophin genotype of each sample
was performed in a single tube using the following primers: forward primer for utrophin knockout (22803),
5'-TGCCAAGTTCTAATTCCATCAGAAGCTG-3'; forward primer for wild-type
utrophin (553),
5'-TGCAGTGTCTCCCAATAAGGTATGAAC-3'; and a common reverse
primer (554), 5'-CTGAGTCAAACAGCTTGGAAGCCTCC-3 (Dr. Mark
Grady, personal communication). PCR reactions were performed on
0.5 µl DNA in a final volume of 25 µl, with final concentrations of
1.5 mM MgCl2, 0.4 mM dNTPs, and 0.025 U/µl platinum
Taq in 1× PCR buffer (GIBCO-BRL). The final concentrations
of the primers were 200 nM. PCR reactions were run on a PTC-150
Minicycler with Hot Bonnet (MJ Research) under the following running
conditions for the utrophin screen: 1) 5 min at 94°C for
initial denaturation of double-stranded DNA, 2) 35 cycles
with three steps per cycle, each for 25 s (50°C, ligation;
72°C, extension; and 94°C, denaturation), and 3) 5 min
at 72°C for final extension. The mdx screen used similar
conditions except the ligation temperature was 48°C. The PCR products
were run on a 3% agarose gel (GIBCO-BRL) containing ethidium bromide
(0.001 mg/ml) in 0.5× Tris-borate-EDTA running buffer (Fisher) at 100 volts for 100 min. Images of the PCR product bands in the gels were
visualized under ultraviolet light (Alpha Innotech Imager) and captured
using Alpha Imager 2000 software. PCR products for both wild-type and
mutated dystrophin yielded a band size of 105 bases; thus,
interpretation of the genotype was based on the reverse primer used in
the reaction. The presence of a band from both screens indicated a
heterozygote (Fig. 1 and Ref.
1). For the utrophin screen, a band of 640 bases indicated wild-type utrophin, and a band of 450 bases indicated the utrophin knockout; the presence of both bands in the same screen indicated a
utrophin heterozygote (Dr. Mark Grady, personal communication and Fig.
1).
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Muscle preparation.
Both EDL muscles were carefully dissected from anesthetized control
(C57bl/6; n = 6), mdx (n = 10), and mdx:utrn(/
) (n = 5) pups aged
9-12 days (2 mg xylazine-20 mg ketamine/100 g body mass ip).
Dissection was performed while viewing the hindlimbs of the pups
through a Reichert Stereo Starzoom stereoscope. EDL muscles were
similarly obtained from additional control, mdx, and
mdx:utrn(
/
) pups aged 9-14 and 20 days for
additional experiments as noted below.
Experimental procedures. One muscle of each pair was maintained at a resting tension of 0.5 g without any electrical stimulation or stretches ("unstretched" muscle) for the same duration required to conduct the contractile measures in the other muscle ("stretched" muscle), including a tension-frequency protocol, five stretches, and the posttwitch and tetanic responses obtained at 15 and 30 min after the last stretch. Resting tension for both muscles was continuously monitored and adjusted as necessary to 0.5 g throughout the experiment. In the stretched condition, after the muscle was mounted to the servomotor arm, it was left quiescent for 10 min and then was subjected to a series of three isometric twitches and tetani (150 Hz) spaced 1 min apart. These were elicited to establish consistent contractile responses before any additional experimental procedures were performed. After an additional 5-min quiescent period, the stretched muscle was subjected to a tension-frequency protocol at electrical stimulation frequencies of 1, 30, 50, 80, 100, and 150 Hz, each for a duration of 900 ms and spaced 1 min apart. After the tension-frequency protocol, and an additional 10 min of rest, muscle length at Lo for both muscles was determined to the nearest 0.1 mm using calipers (Sigma), and then muscles were subjected to the stretch protocol.
Stretch-injury protocol. The stretch-injury protocol described by Petrof et al. (24) was used. The muscle was stimulated at 80 Hz for 700 ms. During the first 500 ms, contraction was isometric at Lo, whereas during the final 200 ms the muscle was stretched at 0.5 Lo/s to yield a total displacement of 0.1 Lo (e.g., an eccentric contraction). This stretch was repeated five times, once every 4 min. At 15 and 30 min after the final stretch, isometric twitch and tetanic (150 Hz) responses from the stretch muscle were elicited. After the final tetanic contraction, the muscles were incubated in a 0.2% procion orange/PSS solution (Reactive Orange 14, mol wt 631; lot number 46H0459; Sigma-Aldrich) for 60 min to assess membrane damage. Muscles were then washed two times, each time for 5 min in fresh PSS and lightly blotted; tendons were dissected free, and the mass to the nearest 0.1 mg was determined on an A-200D electronic analytical balance (Denver Instrument). Muscles were then mounted for sectioning as described below. Note that muscles were incubated in the dye after all contractile measures were obtained. This was done because, in preliminary studies, both control and dystrophic EDL muscles obtained from young mice incubated in the dye demonstrated attenuated force responses (data not shown). We found that the dye batch was critical, because force responses in the presence or absence of the procion orange dye originally used by Petrof et al. (24; Sigma-Aldrich Lot no. 89F0671, a kind gift from Lee Sweeney, University of Pennsylvania) were not different. However, the supply of this dye was limited.
Assessment of membrane damage from handling dystrophic EDL
muscles.
The potential effect on membrane damage resulting from handling between
excision of the muscle and mounting it to the servomotor followed by
isometric stimulation was assessed in paired EDL muscles excised from
anesthetized mdx pups aged 11 days (n = 3 pairs; 30°C). In addition, EDL muscles obtained from
mdx:utrn(/
) pups aged 20 days (n = 3 pairs; 30°C) were also assessed. This age was selected with the idea
that the dystrophic process was in progress, and therefore membrane
damage arising from handling the muscles might be more evident. After
the muscles were excised from each animal, one muscle of each pair
floated free in a beaker of 0.2% procion orange-PSS ("not mounted"
condition), whereas the other muscle was mounted in the contractile
apparatus at a resting tension of 0.5-1.0 g ("mounted"
condition), and then, after determination of Lo,
was subjected to a tension-frequency but not a stretch protocol. This
muscle was then immersed in 0.2% procion orange-PSS. Both muscles were
incubated in the dye for 30-60 min and then washed two times with
PSS and mounted for sectioning as described below.
Muscle sectioning, fiber uptake of procion orange dye, fiber
morphology, and fiber typing.
Muscles incubated in the procion orange dye from the stretch or
no-stretch conditions were mounted on cork in a mixture of gum
tragacanth (Sigma-Aldrich) and Histoprep (Fisher) and frozen in
isopentane cooled in liquid nitrogen. EDL muscles obtained from a
separate group of control, mdx, and
mdx:utrophin(/
) pups aged 9-14 days (mean ± SE; 11.0 ± 0.4) were similarly mounted and frozen to determine
general muscle morphology, including the number of centralized nucleii.
All samples were stored at
80°C until further analysis. Cross
sections were obtained from the midbelly of each muscle sample on a
cryostat (Microm HM 505 N) at
21°C and transferred to glass slides.
Procion orange uptake. EDL muscle sections (10 µm) were viewed under a fluorescent microscope (Nikon Eclipse E400) at ×25 magnification through a Nikon B-2A filter, with the excitation wavelength range set at 450-490 nm and the emission range set at 505-520 nm. Color images were captured on 800 Speed film (Fuji) with a Nikon 35-mm camera (Nikon N6006) fitted to the microscope. To count damaged cells, a plastic sheet was placed over the photographic image, and a dot was marked over each fluorescent cell. Any fluorescent cells noted within a two-cell depth from the border of the muscle and extending radially in the cross section were not included in the overall damaged cell count, since these could have resulted from dissection. When all damaged cells were counted, the total number of cells in the cross section was determined in a similar manner. The extent of damage was determined from the ratio of cells taking up dye to the total number of cells in the section and was expressed as a percentage.
Hematoxylin and eosin staining. Muscle cross sections (12 µm) were fixed for 10 min in 10% formalin in PBS and then stained with hematoxylin and eosin (H&E) as described previously (15). Sections were viewed with a Nikon Eclipse E400 microscope, and a single region (entire field of view with a ×20 objective lens) from the middle portion of each muscle was randomly selected and digitized using an image analysis system (Scion) driven by NIH Image analysis software. All of the fibers within the digitized region were counted, and the proportion of fibers with clearly discernable central nuclei (i.e., cytoplasm completely surrounding the nucleus) was determined for each muscle. The average number of fibers analyzed per muscle was 143.
MyHC isoforms.
The proportional content of MyHC isoforms was determined using a
modified method of Talmadge and Roy (34). Briefly, frozen EDL muscles obtained from control, mdx, and
mdx:utrn(/
) pups aged 9-14 days were thawed on ice
in microfuge tubes, homogenized in 200 µl sample buffer
(16) using a micropestle, and heated to 60°C for 10 min.
Each sample (40 µl) was subjected to SDS-PAGE using 8% separating
and 4% stacking gels (34). Samples were run on a 20-cm
vertical slab gel unit (CBS Scientific) for 40 h at a constant
current of 4.2 mA. Gels were stained with Coomassie blue, destained,
and scanned using an image analysis system (Alpha Imager 2000). Band
densities for each of six MyHC isoforms were expressed as a percentage
of the total MyHC band density.
Contractile data. Contractile data were obtained with specialized software (Aurora Scientific) that controlled the isometric and isotonic modes of the servo arms as well as the timing of pulses that triggered the voltage pulse generator to electrically stimulate the muscles. Contractile responses were collected at a sampling frequency of 1,000 Hz and were stored to disk for subsequent analysis to determine contractile properties using specialized analysis software (Aurora Scientific). Force responses were digitally filtered with a dual-pass Butterworth filter with a cut off frequency of 60 Hz. The cross-sectional area for each muscle was determined by dividing the mass of the muscle (g) by the product of its length (Lo, cm) and the density of muscle (1.06 g/cm3; see Ref. 20) and was expressed as square millimeters. Muscle output was expressed as stress (g/mm2) determined by dividing the tension (g) by the muscle cross-sectional area. Twitch contractile parameters [e.g., peak stress, maximum rates of stress development and relaxation, time-to-peak stress (TPS), and half-relaxation time (HRT)], the stress-frequency relation, and the isometric and dynamic stress values resulting from the stretch protocol were also determined. An index of stiffness during each eccentric contraction was determined by dividing the difference in stress produced between the 80-Hz isometric response and the peak stress resulting from the imposed stretch (i.e., change in stress) by the change in length of the muscle during the stretch (0.1 Lo).
Statistical analysis. The differences in contractile properties between the genotypes and during the series of stretches within each genotype were determined by a two-way ANOVA. Contractile properties before and after the stretch protocol were determined similarly. Differences in stress generation at a given time of contraction and dye uptake between the mouse genotypes were analyzed by a one-way ANOVA. The Newman-Keuls multiple range post hoc test was applied to all significant main effects to determine differences between means. Differences were considered significant at P < 0.05. Data are presented as means ± SE.
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RESULTS |
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Genotyping.
Specific primers were used to assay DNA isolated from mouse tail snips
of the mice using PCR to discriminate the mdx and
mdx:utrn(/
) genotypes. When the PCR products were run on
a 3% agarose gel, the various homozygote and heterozygote genotypes
could be readily determined based on the size of the bands, as
described in the legend for Fig. 1.
Morphological data.
The morphological data are reported in Table
1. The mean ages for the three genotypes
were similar as follows: control, 11.0 ± 0.4 days
(n = 6), mdx 10.9 ± 0.3 days
(n = 10), and mdx:utrn(/
) 11.4 ± 1.0 days (n = 5). The mdx pups (8.32 ± 0.02 g; P < 0.05) had a mean body mass
significantly greater than either the control (6.78 ± 0.34 g) or the mdx:utrn(
/
) (7.14 ± 0.60 g) pups.
EDL muscle length was greater for mdx (8.13 ± 0.13 mm;
P < 0.05) compared with control (7.31 ± 0.11 mm)
but was not different from the values for the
mdx:utrn(
/
) (7.90 ± 0.54 mm) pups. The EDL masses
were similar between the control (1.1 ± 0.1 mg) and
mdx:utrn(
/
) (1.0 ± 0.1 mg) pups but were
significantly less than the values for the mdx group
(1.7 ± 0.3 mg; P < 0.05). Both the control
(0.14 ± 0.02 mm2) and mdx:utrn(
/
)
muscle cross-sectional areas (0.13 ± 0.02 mm2) were
similar, but both were significantly less than the mdx values (0.20 ± 0.02 mm2; P < 0.05).
Within each genotype, there were no differences in the number of fibers
per cross section for those EDL muscles in the stretch vs. no-stretch
condition; therefore, these were combined by genotype. There were a
fewer total number of fibers per muscle cross section in the
mdx:utrn(
/
) (666 ± 30 fibers) and mdx
(769 ± 42 fibers) compared with control (932 ± 43 fibers;
P < 0.05) EDL muscles. There were no significant
differences in the percentage of central nucleated fibers (Fig.
2), nor were there significant differences in myosin isoform distribution among the genotypes (Fig.
3).
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Stress frequency relation.
The isometric stress produced by EDL muscles from the
mdx:utrn(/
) pups was less than the values obtained from
the control EDL muscles at each stimulation frequency
(P < 0.05; Fig.
4A). The control and
mdx stresses were similar at each stimulation frequency
except at 50 and 80 Hz (P < 0.05; Fig. 4A).
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Isometric stress at 80 Hz during the stretch protocol.
EDL muscles obtained from pups of each genotype were subjected to five
stretches. This protocol required that one muscle of each pair not be
stretched, whereas the second muscle was stimulated while isometric at
80 Hz for 0.5 s before a stretch of 0.1 Lo imposed at 0.5 Lo/s (24). This
stretch procedure was repeated five times once every 4 min. There were
no differences in the absolute isometric stresses at 80 Hz across the
five stretches within each genotype (Fig. 4B), but there
were differences between the genotypes (P < 0.05),
with stress stimulation of the mdx:utrn(/
) EDL less than
that of the control EDL at each of the five stretches. For example, the
mean stress at 80 Hz for the first stretch for control EDL muscles was
16.39 ± 2.93 compared with 5.45 ± 1.93 g/mm2
for the mdx:utrn(
/
) muscles (P < 0.05).
The mdx EDL isometric responses at 80-Hz stimulation
differed from those of control EDL only for stretches 1 and
2 (e.g., for stretch 2, control = 14.76 ± 2.49 and mdx = 8.59 ± 1.86 g/mm2;
P < 0.05). The mdx:utrn(
/
) values
differed from those for the mdx muscles only for
stretches 4 and 5 (Fig. 4B;
P < 0.05).
Stiffness.
A stiffness value was determined for each stretch response by dividing
the difference between the plateau of the isometric response at 80 Hz
and the peak stress response during the stretch by 0.1 Lo, the imposed length change (Fig.
4C). There were no differences in stiffness within each
genotype across stretches 1-5 or between the genotypes.
The stiffness values (in
g · mm2 · Lo
1)
for the initial and final stretches for the control EDL were 330.49 ± 17.74 and 371.48 ± 22.82 for control EDL,
250.2 ± 28.2 and 268.9 ± 33.4 for mdx EDL, and
312.0 ± 48.7.4 and 396.5 ± 55.5 for
mdx:utrn(
/
) (Fig. 4C).
Contractile properties before and after the stretch protocol. Isometric twitches and tetani were collected before (initial) and at 15 and 30 min after the conclusion of the stretch-injury protocol (post) to assess the effects of stretch on the subsequent stress-generating capability at low (i.e., twitch)- and high (i.e., tetanus at 150 Hz)-frequency electrical simulation.
Twitch.
For muscles from control animals, mean isometric twitch peak stress was
decreased to 50.4 ± 9.7% (P < 0.05) of the
initial twitch peak stress (4.04 ± 0.65 g/mm2) at 15 min post but was not different from the initial value at 30 min post
(81.7 ± 8.7%; Fig. 5A).
TPS was similar for the initial and 15 and 30 min postcontractions,
with values of 20.8 ± 0.5, 22.0 ± 0.3, and 22.1 ± 1.2 ms, respectively (Fig. 5B). Compared with the initial mean
HRT (16.3 ± 1.7 ms), the mean values were not different at 15 min
(18.8 ± 1.9) or 30 min post (16.5 ± 1.4; Fig.
5C). The maximum rate of stress development
(+dS/dt) was depressed at 15 min post (in
g · mm2 · s
1; 182.5 ± 31.3) compared with both the initial (366.7 ± 64.6) and 30 min
post (306.8 ± 32.6) values. The initial and 30 min post values
were not different. The maximum rate of stress relaxation (
dS/dt) was depressed at 15 min post (
108.2 ± 14.5 g · mm
2 · s
1) compared with
both the initial (
195.3 ± 26.1) and 30 min post (
173.3 ± 17.8; Fig. 5D) values. The initial and 30 min post values were not different.
|
Tetanus.
For control EDL muscles, the initial mean peak tetanic stress of
24.61 ± 2.40 g/mm2 was decreased to 41.9 ± 6.4% of initial values (P < 0.05) at 15 min post and
recovered to 77.1 ± 2.9% (P < 0.05) at 30 min post (Fig. 6). For the mdx EDL
muscles, at 15 min post, the mean peak tetanic stress was only
27.1 ± 2.6% (P < 0.05) of the initial mean
value of 17.66 ± 3.05 g/mm2 and had recovered only to
72.7 ± 10.0% at 30 min (P < 0.05; Fig. 6). For
the mdx:utrn(/
) EDL muscles, the initial mean peak
tetanic stress of 9.41 ± 2.82 g/mm2 was decreased to
34.3 ± 8.9% (P < 0.05) of initial values at 15 min post and showed partial recovery to 53.2 ± 15.8% of initial values at 30 min post (P < 0.05; Fig. 6).
|
Comparisons between the three genotypes.
Initial peak isometric twitch stress generation was similar between
control and mdx EDL muscles but was significantly less for
the mdx:utrn(/
) EDL muscles (P < 0.05).
The initial and 30 min post mdx:utrn(
/
) values for
+dS/dt were significantly attenuated compared with
corresponding control and mdx values (P < 0.05; Fig. 5D). Only the initial mdx:utrn(
/
)
dS/dt value was different from the respective control
value (P < 0.05). TPS for the mdx and
mdx:utrn(
/
) muscles was significantly less compared with
control at 15 and 30 min post (Fig. 5B; P < 0.05), but HRT was similar between the three genotypes at each time
point (Fig. 5C). Comparisons between the contractile and
temporal characteristics of the post and initial twitches within each
genotype (and across all genotypes) revealed a consistent pattern. In
general, at 15 min after the stretch protocol, twitch contractile
properties were attenuated compared with those of the initial twitch
and then demonstrated partial or full recovery toward initial values at
30 min post. The pattern of tetanic response was similar to that of the
twitches, with attenuated stress generation at 15 min post and partial
recovery at 30 min post (Fig. 6). The mean peak stress generated during
the initial tetanus by the mdx:utrn(
/
) EDL muscles was
significantly less than that of control muscles (P < 0.05) but was not different from the mdx initial values. At
15 min post, peak stress generation (in g/mm2) during a
tetanus was depressed for both the mdx and
mdx:utrn(
/
) EDL compared with control EDL values
(P < 0.05). At 30 min post, only the
mdx:utrn(
/
) peak stress was different from the
respective control value (P < 0.05; see legend for
Fig. 6).
Dye uptake as an index of membrane damage.
There was no apparent effect on membrane damage from handling EDL
muscles of mdx:utrn(/
) pups aged 20 days. The percentage of total fibers in a cross section that were dye positive was not
different for the not mounted vs. mounted conditions (n = 3; 14.7 ± 3.8 vs. 9.1 ± 4.4%, respectively; data not
shown). Dye uptake in EDL muscles obtained from mdx mice
aged 11 days were 5.1 ± 0.1 and 4.4 ± 0.6% for the not
mounted and mounted conditions, respectively (data not shown). Both the
not mounted and mounted mean values for these mdx muscles
were less than the mean for the unstretched mdx muscles
(9.2 ± 1.0%; P < 0.05) but were not different
from the mean for the stretched mdx muscles (8.0 ± 1.0%).
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DISCUSSION |
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Major finding. The major finding in this study was that sarcolemmal membranes of EDL muscles obtained from dystrophic pups aged 9-12 days subjected to an acute stretch-injury protocol did not demonstrate greater damage compared with unstretched contralateral muscles, as assessed by uptake of the fluorescent dye procion orange. We hypothesized that, if the absence of dystrophin (and utrophin) rendered the membrane more susceptible to injury as has been shown for muscles of older dystrophic mice (9, 24), and this was a primary mechanism initiating the onset of DMD, then muscles from dystrophic pups at this age would show significant membrane injury when stretched eccentrically in vitro, but this did not occur. These data suggest that the early onset of the dystrophic process may be independent of mechanical perturbation to the sarcolemma.
Stress generation is dramatically reduced in
mdx:utrn(/
) EDL muscles.
Stress generation was depressed in mdx:utrn(
/
) compared
with control skeletal muscles obtained from older mice
(9). In mdx:utrn(
/
) compared with control
pups aged 9-12 days, EDL stress generation was also markedly
depressed throughout the stress frequency relation. In contrast, mean
stresses at most stimulation frequencies were similar between
mdx and control EDL muscles. Depressed stress generation in
the mdx:utrn(
/
) muscles could have been the result of
increased numbers of regenerating fibers. The dystrophic process in
skeletal muscles of mdx:utrn(
/
) and mdx mice
occurs at age ~2 and ~3-4 wk, respectively (11,
28). Morphological evidence of dystrophy includes the presence
of central nuclei indicative of regenerating fibers (11).
However, we found that few fibers in EDL obtained from pups of either
dystrophic phenotype exhibited central nuclei based on the H&E stain,
suggesting that fiber regeneration was minimal at this early age. In
the absence of these influences, the severely attenuated stress
generation in the mdx:utrn(
/
) EDL even before stretches
were imposed suggests a potential intrinsic deterioration in the
contractile apparatus. This decrement could be because of alterations
in the distribution of myosin isoforms and/or alterations in myosin function.
Isometric stress at 80 Hz was maintained during the stretch
protocol.
The mean proportion of maximal stress (150 Hz) elicited at 80 Hz during
the stretch protocol was similar between the genotypes [control
66.6%, mdx 65.7%, and mdx:utrn(/
)
68.5%]. Relative stress over the five stretches was unchanged
for control and gradually increased in the mdx EDL but was
depressed ~37% in the mdx:utrn(
/
) EDL muscles.
However, the decrease in the mdx:utrn(
/
) EDL occurred between the second and first stretches and stabilized thereafter. These
data contrast with the ~29% (control) and ~57% (mdx)
reductions in isometric force output at 80 Hz for diaphragm between the
final and initial stretches in mice aged 90-110 days
(24) and the 10, 41, and 76% decreases in tetanic force
observed for EDL muscles of control, mdx, and
mdx:utrn(
/
) EDL muscles, respectively, from mice aged 10 wk subjected to stretches imposed during tetanic contractions
(9). In EDL muscles of adult control and mdx
mice aged 90-125 days subjected to five stretches, we observed
mean decreases from the initial isometric stress at 80 Hz (~80% of maximum tetanic stress) of ~13% (n = 5) and ~22%
(n = 4), respectively [data not shown; note:
mdx:utrn(
/
) mice this age were not available]. These
results indicate that the magnitude of force loss after the stretch
protocol is likely dependent on genotype, age, and muscle type, as well
as the percentage of maximal stress elicited during the stretch
protocol. However, although the relative stress imposed was different
in the studies reported above, it is important to consider the
experimentally imposed mechanical perturbation relative to the stresses
experienced in vivo. Given the limited movement of the pups aged
9-12 days, the acute stretch imposed on the EDL muscles likely
presented a significant challenge.
Stiffness during the stretch protocol was maintained in the
mdx:utrn(/
) EDL.
We considered that an index of stiffness determined from the change in
stress divided by the change in length during the eccentric phase of
the stretch might reveal potential differences in mechanical coupling
between sarcomeres and sarcolemma and between fibers and tendon in the
absence of dystrophin (e.g., mdx EDL; see Refs. 36, 29, and 38) and in the
absence of dystrophin and utrophin [e.g., mdx:utrn(
/
)
EDL]. Surprisingly, stiffness was similar for all genotypes. Stiffness
in the mdx muscles may have been similar to control muscles
because of compensation by utrophin (14, 39) in providing
stability to the DGC and costameres. However, if it is true that
mechanical coupling between sarcomeres, sarcolemma, and tendon are
compromised in the absence of dystrophin alone, then a greater
deficiency would be expected in the absence of both dystrophin and
utrophin. However, stiffness in the mdx:utrn(
/
) EDL
muscles was similar to that of control and mdx muscles.
Membrane damage of dystrophic muscles was not increased by a
stretch-injury protocol.
Membrane injury as assessed by uptake of the fluorescent dye procion
orange after either a no-stretch or stretch condition was ~3.5- to
4.5-fold greater in EDL muscles from both dystrophic mouse models
[mdx and mdx:utrn(/
)] compared with control
muscles (~7-9% vs. ~2%, respectively). Without stretch
(e.g., the mounted and unstretched conditions), the percentage of
dye-positive fibers in young mdx muscles was ~4-9%
compared with ~3% for both adult control (n = 3) and
mdx (n = 3) muscles (data not shown; see
Ref. 24). This small difference suggests that maturing
compared with adult mdx muscles may exhibit slightly greater
and more variable fiber damage in vivo that reflects the early onset of
the dystrophic pathology, and/or they are more sensitive to handling.
Given the small size of the maturing dystrophic muscles we assessed, we cannot discount some fiber damage from handling that possibly masked
effects of the stretch; yet, this appeared to be minor. For example,
within the five mdx:utrn(
/
) paired muscles, four of the
pairs yielded 4.3 ± 0.7% (n = 4) dye-positive
fibers for the unstretched condition, suggesting limited effects of
handling. Overall, the percentage of dye-positive dystrophic fibers for the maturing mdx and mdx:utrn(
/
) EDL muscles
for both the no-stretch and stretch conditions averaged 7-9%. In
contrast, we found the mean percentage of dye-positive fibers in EDL
muscles of adult control and mdx mice subjected to the
stretch protocol were ~6% (control, n = 5) and
~23% (mdx, n = 4) (data not shown),
similar to the mean values of ~4 and ~19%, respectively, reported
previously (24). Thus dye uptake by EDL muscles obtained
from young compared with adult dystrophic mice was approximately
threefold less (e.g., ~7-9 vs. ~23%). This suggests that the
absence of the DGC does not render the membrane acutely susceptible to
mechanical shear stress during early maturation (i.e., >90% of the
total fibers were undamaged by stretch).
Nature of the dystrophic process.
The significant depression in stress-generating capability of the
mdx:utrn(/
) EDL independent of membrane damage suggests the presence of an active dystrophic process. This observation could be
interpreted in at least three ways. 1) The absence of dystrophin does not initially render the membrane more susceptible to
injury because the membrane is elastic enough to tolerate and recover
from eccentric contractions; thus, the dystrophic process is dependent
on one or more disrupted signaling pathways. Our data indicate that the
dystrophic process may have been evident in a small percentage of the
total fibers but was not exacerbated by the acute injury protocol.
2) The acute perturbation of the muscle used herein was not
enough to exacerbate the structural weakness of the muscle (i.e., the
muscle was not stressed enough or long enough). Thus an increase in the
percentage of maximal isometric stress before the stretch, the extent
of stretch, the number of stretches, or chronic activity may be
necessary to induce the dystrophic process. These possibilities,
however, must be tempered with the caveat that the imposed challenge(s)
on the muscle reasonably reflects those experienced by pups aged
9-12 days in vivo. 3) Alternatively, the onset of
dystrophy could become evident several hours or even days later after
an acute injury (e.g., muscle injury associated with eccentric work),
but this may also favor a signaling hypothesis to account for the onset of DMD (e.g., triggering of inflammatory and immune responses).
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ACKNOWLEDGEMENTS |
---|
We thank Ludwig Haber for expert technical assistance.
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FOOTNOTES |
---|
This work was supported by a grant from the Muscular Dystrophy Association (to R. W. Grange).
Address for reprint requests and other correspondence: R. W. Grange, Dept. of Human Nutrition, Foods and Exercise, Virginia Polytechnic Institute and State Univ., Blacksburg, VA 24061-0430 (E-mail: rgrange{at}vt.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.
May 29 2002;10.1152/ajpcell.00450.2001
Received 19 September 2001; accepted in final form 22 May 2002.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Amalfitano, A,
and
Chamberlain JS.
The mdx-amplification-resistant mutation system assay, a simple and rapid polymerase chain reaction-based detection of the mdx allele.
Muscle Nerve
12:
1549-1553,
1996.
2.
Burkin, DJ,
and
Kaufman SJ.
The 7
1 integrin in muscle development and disease.
Cell Tissue Res
296:
183-190,
1999[ISI][Medline].
3.
Burkin, DJ,
Wallace GQ,
Nicol KJ,
Kaufman DJ,
and
Kaufman SJ.
Enhanced expression of the 7
1 integrin reduces muscular dystrophy and restores viability in dystrophic mice.
J Cell Biol
152:
1201-1218,
2001.
4.
Campbell, KP.
Three muscular dystrophies: loss of cytoskeleton-extracellular matrix linkage.
Cell
80:
675-679,
1995[ISI][Medline].
5.
Chang, WJ,
Iannacone ST,
Lau KS,
Masters BSS,
McCabe TJ,
McMillan K,
Padre RC,
Spencer MJ,
Tidball JG,
and
Stull JT.
Neuronal nitric oxide synthase and dystrophin-deficient muscular dystrophy.
Proc Natl Acad Sci USA
93:
9142-9147,
1996
6.
Coirault, C,
Lambert F,
Marchand-Adam S,
Attal P,
Chemla D,
and
Lecarpentier Y.
Myosin molecular motor dysfunction in dystrophic mouse diaphragm.
Am J Physiol Cell Physiol
277:
C1170-C1176,
1999
7.
Crosbie, RH.
NO vascular control in Duchenne muscular dystrophy.
Nat Med
7:
27-29,
2001[ISI][Medline].
8.
Danowski, BA,
Imanaka-Yoshida K,
Sanger JM,
and
Sanger JW.
Costameres are sites of force transmission to the substratum in adult rat cardiomyocytes.
J Cell Biol
118:
1411-1420,
1992[Abstract].
9.
Deconinck, N,
Rafael JA,
Beckers-Bleukx G,
Kahn D,
Deconinck AE,
Davies KE,
and
Gillis JM.
Consequences of the combined deficiency in dystrophin and utrophin on the mechanical properties and myosin composition of some limb and respiratory muscles of the mouse.
Neuromuscul Disord
8:
362-370,
1998[ISI][Medline].
10.
Grady, RM,
Grange RW,
Lau KS,
Maimone MM,
Nichol MC,
Stull,
and
Sanes JR.
Role for -dystrobrevin in the pathogenesis of dystrophin-dependent muscular dystrophies.
Nature
1:
215-220,
1999.
11.
Grady, RM,
Teng H,
Nichol MC,
Cunningham JC,
Wilkinson RS,
and
Sanes JR.
Skeletal and cardiac myopathies in mice lacking utrophin and dystrophin: a model for Duchenne muscular dystrophy.
Cell
90:
729-738,
1997[ISI][Medline].
12.
Grange, RW,
Isotani E,
Lau KS,
Kamm KE,
Huang PL,
and
Stull JT.
Nitric oxide contributes to vascular smooth muscle relaxation in contracting fast-twitch skeletal muscles.
Physiol Genomics
5:
35-44,
2001
13.
Hutter, OF.
The membrane hypothesis of duchenne muscular dystrophy: quest for functional evidence.
J Inherit Metab Dis
15:
565-577,
1992[ISI][Medline].
14.
Karpati, G,
Carpenter S,
Morris GE,
Davies KE,
Guerin C,
and
Holland P.
Localization and quantitation of the chromosome 6-encoded dystrophin-related protein in normal and pathological human muscle.
J Neuropathol Exp Neurol
52:
119-128,
1993[ISI][Medline].
15.
Kiernan, JA.
Histological and Histochemical Methods: Theory and Practice. New York: Pergamon, 1981.
16.
Laemmli, UK.
Cleavage of structural proteins during the assembly of the head of bacteriophage T4.
Nature
227:
680-685,
1970[ISI][Medline].
17.
Laird, PW,
Zijderveld A,
Linders K,
Rudnick MA,
Jaenisch R,
and
Berns A.
Simplified mammalian DNA isolation procedure (Abstract).
Nucleic Acids Res
19:
4293,
1991[ISI][Medline].
18.
Lau, KS,
Grange RW,
Isotani E,
Sarelius IH,
Kamm KE,
Huang PL,
and
Stull JT.
nNOS and eNOS modulate cGMP formation and vascular responsiveness in contracting fast-twitch skeletal muscle.
Physiol Genomics
2:
21-27,
2000
19.
Mendell, JR,
Sahenk Z,
and
Prior TW.
The childhood muscular dystrophies: diseases sharing a common pathogenesis of membrane instability.
J Child Neurol
10:
150-159,
1995[ISI][Medline].
20.
Mendez, J,
and
Keys A.
Density and composition of mammalian muscle.
Metabolism
9:
184-188,
1960[ISI].
21.
Moens, P,
Baatsen PHWW,
and
Marechal G.
Increased susceptibility of EDL muscles from mdx mice to damage induced by contractions with stretch.
J Muscle Res Cell Motil
14:
446-451,
1993[ISI][Medline].
22.
Ohlendieck, K,
Matsumara K,
Ionasescu VV,
Towbin JA,
Bosch EP,
Weinstein SL,
Sernett SW,
and
Campbell K.
Duchenne muscular dystrophy: deficiency of dystrophin-associated proteins in the sarcolemma.
Neurologia
43:
795-800,
1993.
23.
Pasternak, C,
Wong C,
and
Elson EI.
Mechanical function of dystrophin in muscle cells.
J Cell Biol
128:
355-361,
1995[Abstract].
24.
Petrof, BJ,
Shrager JB,
Stedman HH,
Kelly AM,
and
Sweeney HL.
Dystrophin protects the sarcolemma from stresses developed during muscle contraction.
Proc Natl Acad Sci USA
90:
3710-3714,
1993[Abstract].
25.
Petrof, BJ,
Stedman HH,
Shrager JB,
Eby J,
Sweeney HL,
and
Kelly AM.
Adaptations in myosin heavy chain expression and contractile function in dystrophic mouse diaphragm.
Am J Physiol Cell Physiol
265:
C834-C841,
1993
26.
Phillips, SK,
Bruce SA,
and
Woledge RC.
In mice, the muscle weakness due to age is absent during stretching.
J Physiol (Paris)
437:
63-70,
1991.
27.
Rando TA. The role of nitric oxide in the pathogenesis of muscular
dystrophies: a "Two Hit" hypothesis of the cause of muscle
necrosis. Microsc Res Tech 55: 223-235.
28.
Rezvani, M,
Cafarelli E,
and
Hood DA.
Performance and excitability of mdx mouse muscle at 2, 5 and 13 weeks of age.
J Appl Physiol
78:
961-967,
1995
29.
Rybakova, IN,
Patel JR,
and
Ervasti JM.
The dystrophin complex forms a mechanically strong link between the sarcolemma and costameric actin.
J Cell Biol
150:
1209-1214,
2000
30.
Sander, M,
Chavoshan B,
Harris SA,
Iannaccone ST,
Stull JT,
Thomas GD,
and
Victor RG.
Functional muscle ischemia in neuronal nitric oxide synthase-deficient skeletal muscle of children with Duchenne muscular dystrophy.
Proc Natl Acad Sci USA
97:
13818-13823,
2000
31.
Stamler, JS,
and
Meissner G.
Physiology of nitric oxide in skeletal muscle.
Physiol Rev
81:
209-237,
2001
32.
Stedman, HH,
Sweeney HL,
Shrager JB,
Maguire HC,
Panetierri 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
353:
536-539,
1991[ISI].
33.
Straub, V,
and
Campbell KP.
Muscular dystrophies and the dystrophin-glycoprotein complex.
Curr Opin Neurol
10:
168-175,
1997[ISI][Medline].
34.
Talmadge, RJ,
and
Roy RR.
Electrophoretic separation of rat skeletal muscle myosin heavy-chain isoforms.
J Appl Physiol
75:
2337-2340,
1993[Abstract].
35.
Thomas, GD,
Sander M,
Lau KS,
Huang PL,
Stull JT,
and
Victor RG.
Impaired metabolic modulation of alpha-adrenergic vasoconstriction in dystrophin-deficient skeletal muscle.
Proc Natl Acad Sci USA
95:
15090-15095,
1998
36.
Tidball, JG,
and
Law DJ.
Dystophin is required for normal thin filament-membrane associations at myotendinous junctions.
Am J Pathol
138:
17-21,
1991[Abstract].
37.
Wehling, M,
Spencer MJ,
and
Tidball JG.
A nitric oxide synthase transgene ameliorates muscular dystrophy in mdx mice.
J Cell Biol
155:
123-131,
2001
38.
Williams, MW,
and
Bloch RJ.
Differential distribution of dystrophin and beta-spectrin at the sarcolemma of fast twitch skeletal muscle fibers.
J Muscle Res Cell Motil
20:
383-393,
1999[ISI][Medline].
39.
Wilson, LA,
Cooper BJ,
Dux L,
Dubowitz V,
and
Sewry CA.
Expression of utrophin (dystrophin-related protein) during regeneration and maturation of skeletal muscle in canine X-linked muscular dystrophy.
Neuropathol Appl Neurobiol
20:
359-367,
1994[ISI][Medline].