Department of Orthopedics, University of Maryland School of Medicine, Baltimore, Maryland 21201
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ABSTRACT |
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In a series of experiments,
cultured myotubes were exposed to passive stretch or pharmacological
agents that block contractile activation. Under these experimental
conditions, the formation of Z lines and A bands (morphological
structures, resulting from the specific structural alignment of
sarcomeric proteins, necessary for contraction) was assessed by
immunofluorescence. The addition of an antagonist of the voltage-gated
Na+ channels [tetrodotoxin (TTX)] for 2 days in
developing rat myotube cultures led to a nearly total absence of Z
lines and A bands. When contractile activation was allowed to resume
for 2 days, the Z lines and A bands reappeared in a significant way.
The appearance of Z lines or A bands could not be inhibited nor
facilitated by the application of a uniaxial passive stretch.
Electrical stimulation of the cultures increased sarcomere assembly
significantly. Antagonists of L-type Ca2+ channels
(verapamil, nifedipine) combined with electrical stimulation led to the
absence of Z lines and A bands to the same degree as the TTX treatment.
Western blot analysis did not show a major change in the amount of
sarcomeric -actinin nor a shift in myosin heavy chain phenotype as a
result of a 2-day passive stretch or TTX treatment. Results of
experiments suggest that temporal Ca2+ transients play an
important factor in the assembly and maintenance of sarcomeric
structures during muscle fiber development.
muscle; development; Z lines; A bands
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INTRODUCTION |
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MUSCLE FIBERS CHANGE THEIR myosin heavy chain (MHC) phenotype in response to a number of external perturbations, such as increased or decreased usage (5), electrical stimulation (37), denervation (44), anabolic steroid combined with exercise (36), or thyroid hormone (30). Although in vivo experiments can explore long-term changes, they make it difficult to dissect the individual contributions of mechanical loading or neural recruitment on muscle fibers. A cell culture approach allows the manipulation of one variable at a time, and the use of a developmental model may assist in the elucidation of mechanisms, which can then be applied in adult muscle fibers.
Contractile activation has an effect on a number of molecules, for
example, acetylcholinesterase (45), L-type
Ca2+ channels (18), and glucose transporter 4 (25), and it sets off a series of second messenger
cascades (43). The role of mechanical factors in cellular
development is not well established, but it is receiving more attention
(27, 47). The independent contribution of mechanical
factors and neural factors in myofibrillogenesis remains to be
established. In cultured myosatellite cells, long-term electrical
stimulation induces a slower contractile phenotype (53)
and neural input is a requirement for specific stages of myogenesis
(15). The molecular mechanisms responsible for the phenotypic changes as well as the mechanisms for the subsequent sarcomere assembly remain to be resolved. Sarcomere assembly is likely
to entail several mechanisms, such as that illustrated by the
organization of -actinin in the formation of premyofibrils (13) or by the self-assembly of myosin into thick
filaments before they are inserted into sarcomeres (3,
26).
To study the separate contributions of mechanical stretch and contractile activation on sarcomere assembly, I used a mammalian cell culture model coupled with the controlled application of passive stretch or electrical stimulation and evaluated myofibrillogenesis. My findings suggest that contractile activation, and not mechanical stretch, is important for myofibrillogenesis. Moreover, low-frequency electrical stimulation and its subsequent increased intracellular flux of Na+ and especially Ca+ transients are required for the assembly and maintenance of the Z lines and A bands.
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MATERIALS AND METHODS |
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Muscle cells were isolated from the hindlimbs of neonatal Sprague-Dawley rats. For each individual experiment, the muscles from the hindlimbs of one litter (10 pups) were dissected, freed from connective tissue, minced, pooled in a 60-mm dish, and treated with DNase (10 µg/dish; Sigma Chemical, St. Louis, MO) and collagenase (40 mg/dish; Life Technologies, Gaithersburg, MD) in Wyles solution (137 mM NaCl, 5 mM KCl, 21 mM HEPES, 0.7 mM Na2HPO4, 100 mM glucose, and BSA at 0.1 mg/ml) for 2 h at 37°C. After gentle trituration, the cells were resuspended at a density of 106/ml in sterile DMEM (Life Technologies) containing 10% fetal bovine serum (Life Technologies). To establish the cultures, 0.8 × 106 cells were plated on sterile glass coverslips (VWR, Bridgeport, NJ) or 0.5 × 106 cells were plated on Matrigel (0.15 mg/mm2); on average, each preparation yielded 36 coverslips. The next day, the cultures were supplemented with DMEM containing 10% fetal bovine serum. On day 4, the medium was replaced with DMEM containing 10% fetal bovine serum and cytosine arabinoside (Sigma Chemical) at a final concentration of 20 µM. On day 7, the cultures received DMEM containing 3% horse serum and cytosine arabinoside. The latter was changed every 3 days.
Passive stretch. Cultures were established as above on silicon membranes (Flexercell 2000; Flexercell, McKeesport, PA) coated with Matrigel. The cell culture wells containing the silicon membranes were placed in a computerized, vacuum-operated valve system (Flexercell 2000). A uniaxial passive stretch was applied for 48 h between days 7 and 9 of culture. The passive stretch regimen was adapted from the work presented by other investigators (49). In my experiments, the average passive stretch of the membrane was 5%; the stretch was applied as a train lasting 60 s (1 s on, 2 s off, 20 counts/min), with a rest period of 180 s between trains.
Electrical stimulation. Electrical field stimulation was directly applied to the culture as described by Wehrle et al. (53). Coverslips were first transferred in 3% horse serum in DMEM in a 100-mm sterile petri dish, and single bipolar pulses were applied for 48 h starting on day 7 postplating. A stimulator (S-48; Grass, Warwick, MA) connected to a polarity changer, to eliminate electrolysis, was used with a field pulse with a duration of 2 ms, a frequency of 0.1 Hz, and an amplitude of 400 mV/mm. The two stainless steel electrodes were totally submerged (0.4 cm2) in the medium. These parameters generated an average current in the dish of 1.2-1.4 mA.
Pharmacological agents. Tetrodotoxin (TTX; Sigma Chemical), an inhibitor of voltage-gated Na+ channels, was used at 3 µM (46). Inhibitors of L-type Ca2+ channels (nifedipine and verapamil; RBI, Natick, MA) were used at 10 µM (21). Stretch-activated ion channels were inhibited by using gadolinium (Sigma Chemical) at 10 µM (41), and 2,3-butadione 2-monoxime (BDM) was used to inhibit of myofibrillar ATPase (24). To chelate extracellular or intracellular Ca2+, EGTA (Sigma Chemical) or 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid-acetoxymethyl ester (BAPTA-AM; Calbiochem, San Diego, CA) was used, respectively, at different concentrations. The effect on Z-line and A-band morphology was studied in a dose-response design; in the experiments, control cultures were treated with vehicle only. At these concentrations, toxic effects, such as membrane blebbing or cell detachment, were not observed. Cell viability, determined by trypan blue exclusion, was not affected. The number of myotubes per area was obtained by counting the number of myotubes per area (1.357 mm2). Because myotubes may encompass several optical fields, the stage was moved at least 5 mm to obtain a random sample of different myotubes; only five random images per culture dish were evaluated.
Ca2+ fluorescence imaging.
The effect of TTX concentration on the Ca2+ transient in
the myotube preparation was assessed in fluo 4-loaded myotubes grown on
glass for 7 days, similar to studies using disassociated myofibers (33, 46). Myotube cultures were loaded for 15-30 min
with 10 µM fluo 4-AM (Molecular Probes, Eugene, OR) in Krebs-Ringer solution (in mM: 135 NaCl, 4.0 KCl, 1.8 CaCl2, 1.0 MgCl2, 10.0 glucose, and 10.0 HEPES, pH 7.4) . After the
loading protocol, the cells were allowed to rest in Krebs-Ringer
without the fluorescent dye. Myotube cultures were then incubated in
either control Ringer or Ringer with TTX (1 nM to 10 µM). Myotubes
were selected randomly and monitored for functional Ca2+
release at 37°C in a custom-built air-jacketed chamber on an inverted
microscope (Olympus IX-70 ×60-1.4 numerical aperture oil or
×60-1.3 numerical aperture water objective) coupled to a Bio-Rad
MRC-600 laser scanning confocal system (488-nm excitation) used in
xy-mode (1-s acquisition time; 2 ms/line, 768 pixels/line). Representative mature myotubes were selected by phase-contrast microscopy; on average, 10 myotubes per 35-mm dish were analyzed, and
data were collected from the central part of the myotube. After a
baseline collection (3-4 images), a voltage test pulse (2 ms at
400 mV/mm) was delivered ~100 ms into the subsequent image generating
a brief fluorescent transient. Pretest pulse baseline images were
summed and averaged to generate an average fluorescence image
(F0), which was then subtracted from each of the subsequent
images in the series to create a change in fluorescence images (F F0). The mean peak F
F0 (>10% of
peak F
F0) during the test pulse was determined by
manually selecting pixel regions of the Ca2+ transient. The
average pixel intensity of the region of interest was quantified by
image analysis software (IDL 5.0, Boulder, CO).
Antibodies.
Antibodies against sarcomeric -actinin (clone EA-53) were obtained
from Sigma Chemical. Antibodies against the following rat MHCs were
used: fast MHC (clone MY 32, which cross-reacts with neonatal MHC,
Sigma), types IIa and IIb MHC (clones SC-71 and BF-F3, respectively;
American Type Culture Collection, Manassas, VA), and
developmental/neonatal and slow MHC (clones RNMy2/9D2 and WB-MHCs,
respectively; Novacastra, Newcastle-upon-Tyne, UK).
Immunolabeling.
Cultures that were maintained under control conditions or were
experimentally manipulated by mechanical stretch, electrical stimulation, or addition of drugs were fixed for 10 min with 2% paraformaldehyde in PBS (pH 7.2). After permeabilization with 0.5%
Triton X-100 in PBS, the samples were incubated with BSA-PBS (1 mg/ml) for 30 min. The primary antibodies were applied at a concentration of 5 µg/ml for 1 h. The samples were washed three times in BSA-PBS and incubated for 1 h with a
fluorescein-conjugated secondary antibody (goat anti-mouse IgG; Jackson
Immunochemicals, West Grove, PA) diluted at 1:100. After three washes
in PBS-BSA, coverslips were mounted in 90% glycerol and 10% 1 M
Tris · HCl, pH 8.0, supplemented with 1 mg/ml
p-phenylenediamine and observed through a PlanNeofluar
×63-1.40 numerical aperture oil-immersion objective on a Zeiss IM
35 microscope (Oberkochen, Germany). In double-labeling experiments, A
bands were visualized with monoclonal antibodies against MHC (clone
MY-32), then rabbit anti-mouse Fab fragments (Jackson Immunochemicals),
and then tetramethylrhodamine-conjugated goat anti-rabbit IgG (Jackson
Immunochemicals). The Z lines were visualized with the monoclonal
antibody (clone EA-53), which was then labeled by a
fluorescein-conjugated goat anti-mouse IgG. Digital images were
obtained with a Zeiss 410 confocal laser-scanning microscope (Carl
Zeiss) without further processing. The slides were randomized, and the
integrity and presence of Z lines (Fig. 1E) or A bands (Fig.
1F) were evaluated in eight random fields in each slide.
Approximately 100 myotubes were scored per slide. Myotubes labeled with
anti--actinin continuous Z lines (e.g., Fig. 1E) were
counted as positive cells. Partial Z lines and interrupted Z lines were
counted as negative (e.g., Figs. 1B and 5E).
Similarly, myotubes labeled for MHC that did not show spatially
segregated A bands (e.g., Fig. 1D) were counted as not
having A bands in fully mature myofibrils. The data were presented as a
percentage of total cells containing Z lines or A bands. ANOVA and
appropriate post hoc tests (Tukey's, Scheffé's) were used to
analyze the data. The data are expressed as mean ± SE.
Differences between data sets were considered to be statistically
significant at P < 0.05.
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Western blotting. Myotube cultures were rinsed three times in PBS medium and harvested at 4°C by scraping in SDS-PAGE sample buffer. After samples were boiled, protein concentrations were determined using amido black (23). Samples were then subjected to SDS-PAGE (29) and transferred to nitrocellulose electrophoretically. Immunolabeling was performed as described above with specific primary antibodies and then goat secondary antibodies conjugated to alkaline phosphatase. The bands were visualized using chemiluminescence (Tropix, Bedford, MA) and radiographic film (Biomax ML; Kodak, Rochester, NY).
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RESULTS |
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I used immunofluorescent labeling with antibodies specific to
-actinin and MHC to assess the presence of Z lines and A bands in
maturing rat myotubes. Figure 1, E and F, shows
the morphological criteria used to determine whether Z lines or A bands
were present. Myotubes gradually attain a sarcomeric pattern as a
function of time in culture. At day 4, most of the
-actinin labeling is present at stress fibers (arrow in Fig.
1A), and, later, a striated pattern can be seen initially
near the sarcolemma (arrow in Fig. 1B). An organized MHC
labeling is first seen as thin myofibrils in the cytoplasm (arrow in
Fig. 1D). Gradually, the majority of the myotubes have
sarcomeric structures.
I used a qualitative method, based on the morphology as depicted in
Fig. 1, E and F, to evaluate the development of
myofibrils in the mammalian muscle cell culture. After cells were
plated, myoblasts proliferated for several days and then fused on
days 4 and 5 to give rise to multinucleated
myotubes. On day 5, some of the myotubes showed spontaneous
twitching, and, consequently, the number of cells containing Z lines
and A bands increased significantly (Fig. 1, H and
I). This progressive development of sarcomeric elements was
complete by day 8, when ~60% of the myotubes displayed Z
lines and A bands (Fig. 1, H and I). Although the
Z lines appeared to develop before fully mature A bands could be
visualized in several cultures, no statistical difference
(P = 0.12) in the rate of development of these
structures was apparent from the results. Similar to the observations
of others (19), I observed that rat muscle cultures grown
on Matrigel showed earlier fusion and maturation, especially in the
development of Z lines. Six days after initial plating on Matrigel,
41 ± 12% of the myotubes contained Z lines compared with 24 ± 5.5% in myotubes grown on glass (P < 0.001, 3 independent experiments). By day 9, 78 ± 5% of the
myotubes grown on Matrigel contained Z lines compared with 52 ± 6% for myotubes grown on glass (P < 0.001). The
appearance of A bands in the myotubes did not show a substrate
dependency. On day 6, 20 ± 19% of the myotubes grown
on glass contained A bands and 16 ± 11% of the cells grown on
Matrigel had fully mature A bands (P = 0.53). The
development of mature myofibrils was dependent on the amount of
spontaneous contractile activation, as TTX was able to block the
development of Z lines in a dose-dependent manner (Fig.
2). The 48-h pharmacological treatment in
these cultures did not alter the number of myotubes per dish. In
control cultures, there were 15.3 ± 5.8 myotubes/mm2;
in TTX-treated cultures, there were 17.9 ± 3.4 myotubes/mm2; and, in the verapamil-treated cultures, there
were 20.3 ± 6.6 myotubes/mm2. No statistical
differences were noted between the number of myotubes per dish.
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Having established reliable conditions for studying the development of
contractile structures in myotubes in vitro, I proceeded first to study
the effects of passive stretch of the myotubes grown on Matrigel. As
reported for chick myotubes (49), after passive mechanical
stretch, the rat myotubes aligned parallel to the longitudinal axis of
the uniaxial deformation (bidirectional arrow in Fig.
3, A and
C). In contrast, the controls showed a more random
orientation of myotubes (Fig. 3B). By applying passive stretch, the number of myotubes containing Z lines was not
significantly affected (71 ± 11%, P = 0.1) (Fig.
4). The application of a 10% stretch led
to the loss of the majority of the myotubes; the effect of other
stretch regimens on the presence of Z lines was not further evaluated.
The application of the 5% passive stretch resulted in well-developed
focal adhesion-like complexes near the substrate-attached membrane (see
arrow in Fig. 3E). These complexes contained dartlike -actinin-labeled structures associated with stress fibers
(phalloidin staining is not shown), similar to structures reviewed in
Ref. 7. The application of more vigorous passive stretch
protocols resulted in the detachment of most of the myotubes. The data
suggest that passive stretch as it was applied here does not affect Z lines. Similar to the Z lines, the presence of A bands was not influenced by the application of passive stretch (not shown).
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I used TTX, an inhibitor of voltage-gated Na+ channels, to
block the spontaneous contractile activation of the myotubes in developing cultures from days 7 to 9 postplating.
A dose response for TTX was established (Fig. 2), and, in all the
experiments described herein, we used 3 µM. The lack of contractile
activation resulted in myotubes lacking Z lines, and the sarcomeric
-actinin labeling was found in stresslike fibers (arrow in Fig.
3H) and in a punctate pattern throughout the sarcoplasm
(arrowhead in Fig. 3H). The application of passive stretch
did not reverse the absence of Z lines. In the TTX-treated cells, under
both stretched and nonstretched conditions, the number of myotubes with
Z lines decreased to 2 ± 3% and 3 ± 3%, respectively
(Fig. 4). Similar changes occurred in the organization of MHC in that
the TTX treatment resulted in the nearly complete absence of
recognizable A bands (data not shown). An experiment in which
spontaneously twitching myotubes were treated with gadolinium (10 µM,
48 h) showed no significant alteration in the number of
A-band-containing myotubes (64 ± 7%, n = 3).
However, a 24-h treatment of 8-day-old cultures with BDM (5 µM),
which inhibits myosin-actin interaction, led to a decrease in the
number of Z-line-containing myotubes compared with control cultures
(39.6 ± 23 and 78.5 ± 7.7%, respectively; n = 3, P = 0.05, power = 0.50),
and a similar effect was noticed in the number of A-band-containing
myotubes (39.6 ± 23 and 76 ± 1.5%, respectively;
n = 3, P = 0.025, power = 0.71).
Passively stretching the TTX-treated myotubes had no effect on either Z lines or A bands. These results suggest that contractile activation and
perhaps the internal mechanical strains are a major factor in
controlling myofibrillogenesis.
The effect of TTX was at least partially reversible. In these
experiments, the removal of TTX on day 9 after a 4-day
treatment and the resultant reallowance of spontaneous contractile
activation for a 2-day recovery period resulted in the appearance of Z
lines. Under these culture conditions, 39.3 ± 7.5% (see Fig.
6A) of the myotubes contained Z lines and 35 ± 6.5%
of the cells contained A bands (see Fig. 6B). The removal of
TTX (recovery) resulted in Z lines and A bands having a morphology
similar to that of the control cultures (Fig.
5, C and D). In
11-day control cultures, the number of cells containing Z lines was
57.3 ± 4.3% (Fig. 6A) and the number containing A bands was 60.3 ± 4.4% (Fig.
6B), which was similar to the data acquired previously when
the myotubes were grown on glass (Fig. 1).
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To test whether contractile activation is required for the maintenance of Z lines or A bands, TTX was added at different time points during development. Myotubes were treated with TTX during sarcomeric assembly (days 5-11) or when the assembly process was already accomplished for most of the myotubes (days 9-11). As mentioned above, the addition of TTX from days 5 to 11 resulted in a loss of sarcomeric structures. The number of cells containing Z lines was 0.75 ± 0.1% (Fig. 6A), and the number of cells having recognizable A bands was 1.1 ± 0.5% (Fig. 6B). To determine whether the Z lines and A bands were susceptible to disassembly, TTX was added at a later stage (days 9-11), at which time most of the myotubes had assembled these structures. The TTX treatment resulted in the nearly complete loss of assembled Z lines and A bands (Fig. 5, E and F). Under these conditions, 0.83 ± 0.5% of the cells contained Z lines (Fig. 6A) and 2.3 ± 1 of the cells contained A bands (Fig. 6B). These experiments suggest that electrical activity of the sarcolemma and contractile activation, by either a direct or an indirect mechanism, are required for the development and maintenance of normal sarcomeric structures.
To evaluate the MHC isoforms and quantitative changes, I used Western
blot analysis. In myotube cultures grown on Matrigel, the 48-h
treatment with passive stretch or TTX did not result in major
differences. I used several antibodies to determine the type of myosin
present in these cultures. Proteins in homogenates from different
experimental groups were separated by SDS-PAGE, transferred to
nitrocellulose membranes, and labeled with antibodies specific for
sarcomeric -actinin, developmental MHC, type IIa MHC, type IIb MHC,
and slow MHC. The results indicate that these cultures contain mostly
developmental (neonatal) MHC and some type IIa (Fig.
7). Separation of the MHC in
high-glycerol-containing SDS gels, according to the work presented by
Caiozzo et al. (8), pointed to the same
conclusions: the predominant type of MHC in these cultures is the
neonatal form, with small amounts of types IIa and IIx present (not
shown). Filters stained with antibodies against type IIb MHC (clone
BF-F3) and slow MHC did not result in a visible signal. We compared the
blots from four independent experiments and could not detect any major
changes in signal intensity (representative blot depicted in Fig. 7).
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Because spontaneous activity (contractile activation) promoted
myofibrillogenesis and blocking inhibited myofibrillogenesis, I
reasoned that exogenous electrical stimulation might accelerate the
development of Z lines and A bands. I therefore exposed the myotube
cultures to electrical stimulation for 2 days at a frequency of 0.1 Hz.
In addition, inhibition with the use of antagonists of the L-type
Ca2+ channels resulted in a decrease in the amount of
myotubes containing Z lines (Fig. 8). As
expected, the electrical stimulation protocol resulted in an increased
number of cells with Z lines (Fig.
9A) and A bands (Fig.
9B) on day 9. In three independent electrical stimulation experiments, the number of Z-line-containing myotubes increased from 50 ± 4% in controls to 73 ± 5%
(P = 0.05), and the number of A-band-containing
myotubes increased from 50 ± 9 to 71 ± 3%
(P = 0.02) (Fig. 9). The addition of TTX in combination with the electrical stimulation led to the nearly complete absence of Z
lines (0.6 ± 0.7%, P < 0.001) and, to a lesser
extent, A bands (10.5 ± 5.5%, P < 0.001) (data
not shown). Thus low-frequency electrical stimulation increased the
number of mature myotubes in the dish, perhaps by synchronizing the
population of developing myotubes.
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I used pharmacological agents to explore L-type Ca2+ channels and their role in the appearance of sarcomeric structures. Blocking the influx of Ca2+ had an effect similar to that of blocking the membrane depolarization by TTX in both the spontaneous and the electrically stimulated cultures. Verapamil and nifedipine (both at 10 µM), L-type Ca2+ channel blockers, led to the absence of the Z lines (Fig. 9A) and A bands (Fig. 9B). In spontaneously twitching cultures, the addition of verapamil reduced the number of Z-line-containing cells to 8.5 ± 3.6% (P < 0.001); the addition of nifedipine had a similar effect (6 ± 3.3%, P < 0.001). The number of A bands in spontaneously contracting myotube cultures was reduced to 3.6 ± 1.9% (P < 0.001) when treated with verapamil and to 8.8 ± 6.5% (P < 0.001) when treated with nifedipine. Electrical stimulation combined with the Ca2+ channel blockers led to similar results. Although the data suggested a lesser effect on the sarcomeric structures by verapamil than by nifedipine, the data were not statistically different from those of the other pharmacologically treated groups. The application of both verapamil and electrical stimulation in the cultures yielded 19 ± 5% Z-line-containing myotubes (Fig. 9A, P < 0.001) and 10.5 ± 4.6% A-band-containing myotubes (Fig. 9B, P < 0.001). The treatments with nifedipine and electrical stimulation led to 2.7 ± 2.5% Z-line-containing myotubes (P < 0.001) and 1 ± 1.7% A-band-containing myotubes (P < 0.001). The data indicate the role of Ca2+ in sarcomeric assembly.
To ascertain whether the TTX treatment during the electrical
stimulation protocol led to a lack of Ca2+ fluxes, an
experiment was undertaken to visualize the intracellular amount of
Ca2+ in these developing myotubes (Fig.
10). The data show that, under the TTX
concentrations and electrical stimulation conditions used in the
experiments, Ca2+ flow was significantly inhibited. In an
additional set of experiments, a Ca2+ transient was
visualized at a TTX concentration of 10 µM by increasing the voltage
in the stimulation protocol to 800 mV/mm (not shown). To
determine whether the internal Ca2+ pools contributed to
sarcomeric assembly, I used a pharmacological approach. The addition of
EGTA to the medium led to a dose-dependent decrease in the number of
Z-line-containing myotubes (Fig.
11A), whereas the chelation
of internal Ca2+ by using BAPTA-AM at different
concentrations did not affect Z line formation (Fig. 11B).
To reverse the effect of reduced external Ca2+ entry by
verapamil, the release of internal Ca2+ stores was induced
by using nanomolar amounts of ryanodine (Fig. 11B); such an
approach did not lead to an increase in Z-line-containing myotubes.
Immunofluorescent labeling with antibodies against the ryanodine
receptor indicated a punctate, suggesting an unorganized internal membrane system (data not shown).
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DISCUSSION |
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In vivo models of muscle development and muscle plasticity may not be able to determine completely the independent contributions of the mechanical environment (i.e., the stresses and strains generated within the tissue) or of the electrical activity (dictated by the neural recruitment of the motoneurons). The independent effects of passive stretch (an external strain) and contractile activation (inducing internal strains) on muscle development may be different. In an effort to study these effects, I used a mammalian cell culture model of myofibrillogenesis. By combining pharmacological agents with either passive stretch or electrical stimulation, I was able to assess the effects of these treatments on the development of sarcomeric structures. A passive, external stretch, as was applied here, had no apparent effect. In contrast, electrical activity, by either spontaneous contractile activation or induced electrical stimulation, is not only required for the assembly of Z lines and A bands but is also necessary for the maintenance of these structures.
Role of passive stretch in myofibrillogenesis.
The in vitro application of passive stretch is used to evaluate the
synthesis of prostaglandins in fibroblasts (1), and the
deformation of the substrate in this application can be modeled mathematically (20). The application of a passive stretch
is used in chick cultures to assess -actin gene transcription
(9) or to evaluate morphological changes
(49). I applied passive stretch to the cultured rat
myotubes by using the approach based on the work of Vandenburgh and
Karlisch (49). Using this method, I could (by adding
pharmacological agents such as antagonists for voltage-gated
Na+ or Ca2+ channels) inhibit the spontaneous
contractile activation, an approach that was used earlier (4,
46). I could thereby independently test the effects of passive
stretch or contractile activation on the development of sarcomeres in
mammalian skeletal muscle cells. Using morphological criteria similar
to those used by others (40), I could not show that a
cyclic passive stretch regimen (averaging 5%) facilitated or inhibited
the formation of Z lines or A bands. In vitro, passive stretch may
actually have a series of catabolic effects in muscle, such as a
decreased level of transcription (9), induction of
membrane damage (12), and the generation of
phospholipase-derived products (11, 50, 51). It is
possible that passive stretch may have a role during the myoblast stage rather than at the myotube stage, similar to what is shown in alveolar
cells (42) and cardiac fibroblasts (34). In
cultured, mononuclear cells, passive stretch gives rise to increased
cell proliferation (42) and activation of cell
proliferation-related kinases, such as extracellular signal-related
kinase and c-Jun NH2-terminal kinase (34).
Role of contractile activation and electrical stimulation in sarcomeric formation. By using pharmacological agents, I could inhibit the ion flow during spontaneous contractile activation or electrical stimulation. Although adult skeletal muscle responds to TTX at nanomolar levels, in denervated and developing muscle a different voltage gated Na+ channel (SkM2) is expressed that is sensitive to TTX in the micromolar range (55). In culture, both genes are expressed (55), necessitating the use of TTX in the micromolar range (Fig. 2). Blocking the extracellular Na+ movement with or without a passive stretch regimen led to the absence of Z lines and A bands, suggesting that membrane depolarization is more important for sarcomeric assembly than is passive stretch. Direct electrical stimulation of the cultures combined with the pharmacological manipulations pointed more immediately to this conclusion. Inhibition of either the voltage-dependent Na+ channels or the L-type Ca2+ channels in the mammalian myotube culture resulted in the failure to assemble and maintain Z lines and A bands.
Experiments in which whole muscle is exposed to altered loading (5, 14, 48) convincingly showed altered gene expression, which could be the result of either a modified mechanical environment or an altered neural recruitment. To complement these experiments, additional structural studies are necessary to understand sarcomere assembly, an issue of particular importance for muscle, since its structural characteristics are related to its functional properties. My experiments suggest that, in developing myotubes, the role of contractile activation on myofibrillogenesis is more important than the influence of passive mechanical factors. Parallel experiments emphasize the importance of neural activity on the MHC gene expression by applying electrical stimulation to cultured muscle cells (16) or by using nerve-muscle cocultures (15). How depolarization and the inward Ca2+ flow may facilitate Z-line assembly are not yet known, but these processes may preferentially affectSynthesis and assembly of sarcomeric structures.
An important aspect of muscle development is the orchestrated interplay
between synthesis, degradation, and assembly of contractile elements. I
did not detect major changes in the level of MHC and -actinin
protein in two different experimental conditions, but the experiments
were relatively short. Changes in muscle gene expression and altered
phenotype as a result of altered loading or electrical stimulation are
seen only after several days to weeks (5, 37). Increased
turnover of myofibrillar proteins is shown in chick muscle culture
grown for 6 or more days in high-K+-containing medium (12 mM) or in normal medium supplemented with TTX (4).
Similarly, electrical stimulation of cultured muscle satellite cells
induces changes at the MHC mRNA level only after several weeks
(16, 35, 53).
Role of Ca2+ in myofibrillogenesis.
Altered neural activity and the altered electrical pulse pattern might
lead directly to gene transcription (6), as shown for the
MHC (35). The pulse pattern might lead to temporally and
spatially defined increases in intracellular Ca2+ and thus
gene expression. A Ca2+-dependent gene transcription is
proposed in MHC fiber-type transformation (10, 28), but
change in muscle phenotype as it occurs during development and in adult
life would also have to entail a well-controlled protein degradation
and sarcomeric assembly process. The data presented herein indicate a
role for temporal, extracellular Ca2+ fluxes in sarcomere
assembly and maintenance. It is interesting to note that calpain, a
Ca2+-dependent protease, is able to remove Z lines from
muscle fibrils without causing the degradation of -actinin and
without affecting its binding capability to actin (22,
31). Treatments of muscle cell cultures with phorbol esters
result first in the disassembly of Z lines and subsequently in the
disassembly of A bands (32). Thus Ca2+ and
related second messenger pathways might activate both selective gene
transcription and sarcomeric assembly.
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ACKNOWLEDGEMENTS |
---|
I appreciate the help of Dr. Chris Ward during the Ca2+ imaging experiments, and I am grateful for the suggestions from Dr. Susan D. Kraner regarding the SkM2 Na+ channel. I also thank Dr. Martin Schneider and especially Dr. Robert Bloch for comments and helpful suggestions, Rick Meyer for technical assistance, and Dori Kelly for editorial assistance.
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FOOTNOTES |
---|
This work was possible through the support of the National Center for Medical Rehabilitation Research.
Address for reprint requests and other correspondence: P. G. De Deyne, PhD, MPT, Division of Orthopedics, MSTF, Rm. 400, 10 South Pine St., Baltimore, MD 21201 (E-mail: pdedeyne{at}smail.umaryland.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.
Received 28 January 2000; accepted in final form 7 July 2000.
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