Department of Biochemistry and Molecular Biology, University of Maryland School of Medicine, Baltimore, Maryland 21201
Submitted 21 February 2003 ; accepted in final form 22 April 2003
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ABSTRACT |
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calcium; ryanodine receptor; dihydropyridine receptor; muscle development
Differences in the expression profile as well as the structural arrangement of excitation-contraction (EC) coupling proteins between embryonic and adult mammalian skeletal muscle may contribute to the rare detection of Ca2+ sparks in the adult muscle. One protein that is differentially expressed during development of mammalian skeletal muscle is the RyR. In adult skeletal muscle the type 1 isoform (RyR1) is predominantly expressed (21, 31), whereas in the embryonic tissue RyR3 is coexpressed with RyR1 at high levels. The expression level of RyR3 declines to very low levels in adult muscle (16) during the postnatal period. Whereas RyR1 is characterized by its ability to functionally couple to dihydropyridine receptor (DHPR), giving skeletal muscle a voltage-dependent sarcoplasmic reticulum (SR) Ca2+ release mechanism independent of Ca2+ flux across the sarcolemma (28), RyR3 does not couple to the DHPR, instead being activated by voltage-independent mechanisms (14, 27), presumably by Ca2+-induced Ca2+ release (CICR). Given the developmental expression of RyR3, it is tempting to speculate that the presence of RyR3 contributes to the relative abundance of Ca2+ sparks in embryonic but not in adult mammalian skeletal muscle.
The occurrence of Ca2+ sparks in embryonic mammalian skeletal muscle may also be attributed to an altered EC coupling mechanism in the embryonic muscle. In adult skeletal muscle, EC coupling occurs independently of external Ca2+ (1), whereas the twitch tension in neonatal mouse and rat EDL muscles was found to decrease with lowered extracellular Ca2+ concentration ([Ca2+]o) (11, 25). Other investigators have observed a Ca2+ current-dependent component of contraction in developing cultured myotubes (8). Embryonic and neonatal mammalian skeletal muscles thus appear to posses a cardiac type of EC coupling in which influx of extracellular Ca2+ initiates RyR Ca2+ release by CICR due to a local elevation of Ca2+ in the vicinity of the RyR. This raises the possibility that extracellular Ca2+ might also modulate the production of Ca2+ sparks in embryonic fibers.
The presence of Ca2+ sparks in embryonic but not intact adult mammalian skeletal muscle appears to be a complex issue. To examine the production of Ca2+ sparks during muscle development, we have performed a detailed and objective examination of the frequency of occurrence and spatial-temporal properties of skeletal muscle Ca2+ release events over the course of muscle development (i.e., late embryonic through postnatal) in mouse diaphragm and extensor digitorum longus (EDL) skeletal muscle. We have shown that the frequency of occurrence of spontaneous Ca2+ sparks declines dramatically in isolated mouse skeletal muscle fibers after only 1 wk postbirth. The decreasing frequency is not associated with a decline in the relative expression of RyR3 compared with RyR1; in fact, RyR3 shows little decline through postnatal day 14. Thus differential expression of RyR isoforms does not appear to play a central role in Ca2+ spark production in developing mammalian skeletal muscle or in the decline of spark occurrence during early postnatal development.
We also explored possible mechanisms by which RyRs are activated to produce Ca2+ sparks in embryonic skeletal muscle. We have shown that the frequency of occurrence of Ca2+ sparks in embryonic diaphragm is influenced by the [Ca2+]o and specifically by Ca2+ influx via L-type Ca2+ channels. A possible interpretation of our findings is that Ca2+ sparks in embryonic muscle are triggered by local elevation of [Ca2+] due to Ca2+ influx via L-type Ca2+ channels. A decrease in Ca2+ influx via L-type channels during early postnatal development, possibly due to formation of molecular coupling between the DHPRs and RyR1 (30) or to some other developmental change resulting in decreased opening of DHPRs in resting fibers, might account for the observed decrease in spontaneous event occurrence after birth.
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MATERIALS AND METHODS |
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Embryonic and postnatal CD-1 mice (Charles River) at gestation day 18 (E18) and postnatal days 1, 7, and 14 (P1, P7, and P14) were killed by CO2 asphyxiation followed by rapid decapitation. Diaphragm and EDL muscles were dissected and enzymatically dissociated with 2 mg/ml collagenase (type 1; Sigma) dissolved in medium (MEM with Earle's salts; GIBCO) containing 10% fetal bovine serum (Biofluids) and 100 µM gentamicin (Sigma) at 37°C for 2.53.5 h. Fiber bundles were then transferred to serum-supplemented medium without collagenase, teased apart, and separated by gentle trituration. Intact single fibers were plated on extracellular matrix (Sigma)-coated coverslips attached across a 15-mm hole in the bottom of 35-mm petri dishes (MatTek) in serum-supplemented medium. The isolated diaphragm and EDL fibers were incubated overnight in a 5% CO2 incubator at 37° in serum-supplemented medium to allow retention of only viable fibers.
Ca2+ Imaging
Isolated developing diaphragm and EDL fibers were loaded by exposure to 10 µM fluo 4-AM (Ca2+ indicator dye; Molecular Probes) for 30 min and prepared in Ringer solution (in mM: 135 NaCl, 4 KCl, 1.8 CaCl2, 1 MgCl2, 10 glucose, and 10 HEPES-Na2). Fibers were then bathed in Ringer containing no dye (30 min) to allow dye deesterification. Visual screening was used to prescreen fibers for apparent stress. Only fibers possessing a regular striation pattern and no visible signs of stress (e.g., elevated cytosolic Ca2+, membrane blebs/sprouting) were imaged for this investigation. Local Ca2+ events in diaphragm and EDL fibers were monitored on an inverted microscope (Olympus IX-70 with a x60, 1.3 NA water-immersion objective) coupled to a Bio-Rad MRC-600 laser scanning confocal imaging system, focused near the lower surface of the fiber slightly above the coverslip. An argon ion laser was used to excite fluorescent molecules at 488 nm. Fibers were then imaged in either full-frame (XY) or line-scan (Xt) mode. In XY mode, each 138.2 x 92.2-µm image was acquired in 1 s (2 ms per line). The fiber was oriented roughly parallel to the X scan direction. A series of 30 full-frame images was collected at each of four randomly selected locations on each fiber. In line-scan mode, a 138.2-µm line was placed longitudinally along the fiber and repeatedly scanned at the same location 512 times at 2 ms per line, generating an image of distance (X) vs. time (t). Five sequential line-scan images were captured at each spatial location. When possible, line-scan images were acquired at locations that demonstrated Ca2+ activity during full-frame imaging. In P7 EDL, P14 EDL, and P14 diaphragm fibers, line-scan images were acquired without regard to active sites; rather, the entire diameter of the fiber was scanned at increments of 10 pixels across the fiber.
Effect of Ca2+ Entry on Spark Frequency
Isolated embryonic diaphragm fibers were loaded and imaged as described for the developmental studies, with the following exceptions: each E18 diaphragm fiber was imaged at three separate locations, with 30 or 50 sequential full-frame images (X vs. Y) (768 x 512 pixels; 1 pixel = 0.18 µm) obtained at each of three locations on the fiber while bathed in control Ringer solution. The external solution bathing the fiber was then replaced, and the fiber was imaged again after 10 min in 1.8 mM Ca2+ Ringer (control with physiological concentration of Ca2+), 8 mM Ca2+ Ringer, or 0 Ca2+ Ringer containing 1 mM EGTA (Sigma) at the same fiber locations as imaged in Ringer before the external solution was changed. In other experiments E18 diaphragm fibers were first imaged at three locations in Ringer solution and then imaged again at the same locations of the fiber as imaged in Ringer after 30 min in normal Ringer or Ringer solution containing 5 mM CoCl2, 30 µM nifedipine (Sigma), or 5 µM NiCl2 (Sigma).
Ca2+ Event Identification and Analysis
XY images. Ca2+ sparks were identified as
spatially localized regions of elevated fluorescence using an automatic
detection method (6) modified
for identification of Ca2+ release events in XY
confocal images (33). First,
an average fiber fluorescence image was obtained by summing a sequence of 30
images pixel by pixel and calculating the mean fluorescence at each pixel. The
region of the image corresponding to the fiber was then manually defined as an
area of interest, and the standard deviation (SD) of each fiber pixel was
calculated over the sequence of 30 images. Potential local
Ca2+ release events were identified as contiguous pixels
exhibiting fluorescence 1.5 standard deviations above the mean fiber
fluorescence. Mean fiber fluorescence (F) was then recalculated in each image
pixel by pixel over the 30-image sequence excluding any potential event areas.
The normalized change in fluorescence (
F/F) was calculated pixel by
pixel after three-point smoothing of the F images. Regions of potential local
Ca2+ events were identified in
F/F images as
contiguous regions of pixels having fluorescence values
2 SD above the
mean fluorescence and were selected by the criterion that at least 1 pixel
must exceed 3 SD above the mean. Ca2+ release events
were characterized in the
F/F image by the measured parameters peak
amplitude (peak
F/F) and full area at half-maximal fluorescence (FAHM;
µm2) and by the derived parameters equivalent diameter (EDHM)
and equivalent volume integral at half-maximal fluorescence (VIHM). The
equivalent diameter of Ca2+ events was calculated from a
circle created by fitting pixels from the area at 50% of the peak amplitude as
EDHM = 2(FAHM/
)1/2 (µm). Similarly, the equivalent volume
integral was derived from the area at 50% of the peak amplitude, with the
assumption that the events have a spherical geometry, as a product of the
derived sphere and the mean
F/F at half-maximal fluorescence (mean
F/F): VIHM = (
/6)(EDHM)3(mean
F/F).
Line-scan images. Ca2+ sparks from line-scan
images were also identified as spatially localized regions of elevated
F/F fluorescence by using an automatic spark detection algorithm
(6) with slight modifications
as previously described (32,
33).
Ca2+ sparks were characterized by calculating peak
amplitude (
F/F), spatial width (fit to the Gaussian curve) at 50% of
the peak amplitude (FWHM, µm), duration at 50% of the peak amplitude (FDHM,
ms), and the rise time (RT) between 1090% of the peak amplitude (ms).
Only events adhering to the following criteria were selected: amplitude
F/F values >0.2, FWHM
9 µm, and FDHM
30 ms. Long-duration
events without a poorly defined rising or falling phase of the transient were
categorized by examination of the time course records. The time courses were
measured as the mean
F/F of five adjacent spatial pixels centered at
the peak of the event. Image processing and data analysis for both XY
and line-scan images were performed with custom image analysis routines
written in IDL 5.0 (Boulder, CO).
Statistical Analysis
Statistical significance among variables was determined using the Kruskal-Wallis nonparametric test followed by the Dunn's multiple comparison test (P < 0.05).
Assay of SR Store Content
E18 diaphragm fibers were loaded with fura 2-AM for 20 min and then allowed
to equilibrate in normal Ringer containing 20 µM
N-benzyl-p-toluene sulfonamide (BTS; Sigma S949760) for 20
min. All solutions thereafter contained 20 µm BTS to inhibit contraction
(7). Fiber fluorescence
emission ratios R = F380/F358 for excitation at 380 or
358 nm were monitored to assay relative cytoplasmic
[Ca2+]. Fibers were briefly exposed to 20 mM caffeine in
Ringer solution containing 1.8 mM Ca2+ (control) and
then washed with caffeine-free Ringer after the peak of the
Ca2+ transient. When the fiber had recovered completely
from the caffeine-induced Ca2+ transient, the fibers
were then incubated in 0 Ca2+ or 8 mM
Ca2+ Ringer for 10 min and then reexposed to 20 mM
caffeine in 1.8 mM Ca2+ Ringer. The peak amplitudes of
the caffeine-induced R transients were measured and normalized to the
preceding control caffeine response in the same fiber, and the mean of the
normalized responses was calculated.
Crude Membrane Preparation and Western Blot Analysis
Adult frog limb muscles and mouse diaphragm muscles from E18, P1, P7, P14,
and adult (>60 days) mice were rapidly dissected and flash frozen in
2-methylbutane (cooled on dry ice) and stored at 80°C. In each of
three litters of embryos, the tissue from the first two animals was used to
prevent degradative proteolysis of the tissue after asphyxiation. Mouse
diaphragm samples at each time point were pooled because of the small amount
of protein in each sample in E18 and P1 time points. As a control we used frog
hindlimb muscle, which is known to express RyR and RyR
(RyR1/RyR3
homologues) in equal amounts. Frozen tissue (6080 mg) was ground in a
mortar containing liquid nitrogen and then homogenized in 500 µl of
ice-cold homogenization buffer [20 mM HEPES, pH 7.4, 250 mM sucrose, 0.2%
sodium azide, 0.2 mM phenylmethylsulfonyl fluoride (PMSF), 10 µM pepstatin,
10 µM leupeptin, 10 µM aprotinin, 10 µM E-64, and 10 µM antipain]
with a Dounce homogenizer. Crude membrane proteins were extracted by adding
500 µl of extraction buffer (5 mM NaPO3, pH 7.4, 75 mM NaCl, 1%
Triton X-100, 1% deoxycholate, 0.1% SDS, 0.2 mM PMSF, 10 µM pepstatin, 10
µM leupeptin, 10 µM aprotinin, 10 µM E-64, and 10 µM antipain) and
then centrifuging at 10,000 g for 15 min
(18,
35). The supernatant, which
contained the membrane proteins, was stored at 80°C. Protein
concentrations were determined using the Bio-Rad DC protein assay. Membrane
proteins were separated on 4% SDS-PAGE gels as described by Laemmli
(20). Proteins were then
transferred to a polyvinylidene difluoride membrane (Novex) in transfer buffer
containing 12 mM Tris-base, 92 mM glycine, 0.02% SDS, and 20% methanol at 25 V
at 4°C overnight. The membrane was blocked for 3 h in blocking solution
[5% Carnation nonfat dry milk in 150 mM NaCl, 10 mM Tris · HCl, pH 7.4,
0.1% Tween 20 (TBST)] at room temperature and then incubated in monoclonal
anti-RyR antibody (Affinity Bioreagents) diluted 1:5,000 in blocking solution
for 1 h at room temperature. Antigen detection was performed using a
peroxidase-conjugated donkey anti-mouse secondary antibody (Jackson
ImmunoResearch Laboratories) diluted 1:10,000 in blocking solution for 1 h at
room temperature, followed by a chemiluminescent substrate-detection system
(Pierce SuperSignal West Pico chemiluminescent substrate no. 34080).
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RESULTS |
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Figure 1A presents
30 sequential F/F images of a randomly selected region of an E18
diaphragm fiber loaded with fluo 4. These images exemplify the low
Ca2+ event occurrence typically observed in embryonic
skeletal muscle fibers. Surface plots (Fig.
1B) of the fiber displayed in
Fig. 1A show accepted
events (dark arrows) selected by the objective technique utilized in
identification and selection of Ca2+ release events.
Light arrows point to areas of increased fluorescence that do not adhere to
the selection criteria and were not selected as events.
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Our first aim was to objectively quantify the frequency of occurrence of spontaneous Ca2+ events in muscles at various stages of embryonic and postnatal development. Embryonic and postnatal diaphragm and postnatal EDL fibers were loaded with fluo 4, and 30 sequential images were taken on three to four randomly selected regions on each fiber. Ca2+ event frequency was calculated as the mean number of events per image and normalized to the fiber area, which increases during development as a result of splitting of myofibrils and addition of new myofibrils (15). The fiber area of interest increased by 28.6% from E18 to P14 in diaphragm fibers and by 12.4% from P1 to P14 in EDL fibers (data not shown). The mean frequencies were measured in E18, P1, P7, and P14 diaphragm fibers and in P1, P7, and P14 EDL fibers. E18 EDL fibers were not observed because of the difficulty in isolating healthy fibers in which a regular sarcomere pattern was present. Both diaphragm and EDL fibers displayed low levels of spontaneous Ca2+ event activity even from the early stages of development observed. On average, only one event was detected for every five XY images monitored in E18 diaphragm fibers (e.g., Fig. 1A), and the event frequency decreased to much lower levels in postnatal diaphragm and EDL fibers. Normalized to the fiber size, the frequency of events in mouse diaphragm fibers declined 30.2% from E18 to P1, 71.0% from E18 to P7, and 95.4% from E18 to P14 (Fig. 2A). The decline in event frequency during postnatal development was even more dramatic in EDL fibers, which showed a 95.1% decline from P1 to P7 (Fig. 2B). At P14 the event frequency remained at a low level significantly greater than zero.
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The mean values of spatial and mass parameters for the population of
identified sparks in both diaphragm and EDL fibers exhibited generally
decreasing trends as the fibers become more developed. In diaphragm fibers,
the peak amplitude (F/F) decreased signifi-cantly from 1.24 ±
0.02 for E18 diaphragm fibers to 0.65 ± 0.05 for P14 fibers (P
< 0.05) (Fig. 3A).
In EDL fibers, peak amplitude did not change significantly during postnatal
development (Fig. 3B).
However, there was a significant decrease in the equivalent diameter, which
dropped from 1.56 ± 0.04 to 1.10 ± 0.05 µm
(Fig. 3D). In
contrast, the decrease in equivalent diameter with postnatal age for the
events in diaphragm fibers was not significant
(Fig. 3C). The VIHM
decreased significantly for both diaphragm and EDL fibers, from 3.79 ±
0.22 to 0.80 ± 0.25 µm3 ·
F/F in E18 to P14
diaphragm fibers (Fig.
3E) and from 2.17 ± 0.20 to 0.66 ± 0.12
µm3 ·
F/F in P1 to P14 EDL fibers
(Fig. 3F). Taken
together, these results suggest either that in the more developed fibers less
Ca2+ was released during a spontaneous release event or
that increases in endogenous Ca2+ buffering or some
other changes during development cause the spark parameters to decrease.
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To evaluate the effect of the Ca2+ event selection
criterion on the developmental change in spark frequency and properties,
images were reanalyzed using a less stringent criterion of at least 1 pixel
having F 2.5 SD above the mean. The resulting frequency of selected events
increased in relative proportion at all stages of development. The same events
identified using the
3 SD criterion were also identified when the
2.5
SD criterion was used. Moreover, use of the
2.5 SD criterion identified
many additional events that displayed smaller areas and lower amplitudes that
were more difficult to discern from background noise (data not shown).
However, a similar relative declining trend in event frequency with
development as shown in Fig. 2
was still seen.
Lack of Correlation Between Ca2+ Event Occurrence and RyR3 Expression
Expression of RyR3 has been detected in embryonic mammalian skeletal muscle
but occurs at diminished levels in adult mammalian diaphragm and is
nonexistent in adult mammalian fast twitch skeletal muscle
(16). RyR1 () and RyR3
(
) are known to exhibit different mobilities on SDS-PAGE
(23), so to evaluate a
possible relationship between the occurrence of Ca2+
events and the expression of RyR3, we performed Western blot analysis on
pooled mouse diaphragm samples at E18, P1, P7, P14, and adult (>60 days).
As a control we used frog hindlimb muscle, which is known to express
RyR
and RyR
in approximately equal amounts
(23)
(Fig. 4, A and
B, lane 6). Densitometry was used to quantify
expression of RyR1 (
) and RyR3 (
). Because the amounts of total
RyR protein may vary compared with the total membrane proteins isolated at
each stage of development, we chose to take the ratio of RyR3 to RyR1, which
also helped to eliminate discrepancies in gel loading. In agreement with
previous reports (16), our
results show that RyR3/RyR1 ratios exhibit only a very slight decrease from
E18 to P14 (Fig. 4, A and
B, lanes 14) but a marked reduction in
the adult (>60 days) diaphragm (lane 5). Compared with the
frequency of Ca2+ event occurrence, which declines very
rapidly during postnatal development and is already at very low levels by P14,
the RyR3/RyR1 ratio remains relatively high in P14 diaphragm fibers,
suggesting that the occurrence of Ca2+ sparks does not
correlate to the expression of RyR3.
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Ca2+ Release Events in Line-Scan Images
In the series of 30 images in Fig.
1A, three of the four Ca2+ events
are shown to have occurred at the same location in this fiber. Within the
entire population of fibers examined, the repetitive nature of
Ca2+ spark occurrence was examined by evaluating events
that recurred within 3 µm of each other in two or more different images in
the same image series. With this analysis it was determined that 29% of
all event locations had reoccurring Ca2+ release events.
Thus events often occur repetitively at the same locations.
The repetitive occurrence of Ca2+ release events at
particular spatial locations was used advantageously in placement of the line
location during detection in the line-scan mode of the confocal microscope.
Spontaneous Ca2+ release events were identified in
line-scan images, and their spatial and temporal properties were characterized
using a computer algorithm. Figure
5 presents F/F image strips and corresponding fluorescence
time courses of Ca2+ sparks recorded in line-scan mode.
For those events satisfying our predefined spatial and temporal selection
criteria (see MATERIALS AND METHODS), the mean property values were
similar for both E18 diaphragm and P1 EDL fibers
(Table 1). Because of the low
frequency of Ca2+ event occurrence at later
developmental stages, many line-scan images were monitored in an attempt to
capture enough events for characterization. However, the numbers of events
characterized in line-scan mode were limited because of the rare occurrence of
events in postnatal skeletal muscle fibers and the challenge of finding and
placing a line through an event before it occurs. The numbers of events
acquired were too few for comparisons of event properties as a function of
developmental stage.
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In E18 diaphragm fibers as well as P1, P7, and P14 EDL fibers, some events were detected in line-scan images that could not be characterized with our predefined spatial and temporal parameters. Time courses of Ca2+ events that possessed a FDHM >30 ms (4.3% of all events fulfilling the other selection criteria; Fig. 5B) were compared with accepted Ca2+ spark time courses that exhibited property values within the accepted range (Fig. 5A). Temporal traces of longer duration events displayed a variety of fluorescence time courses (Fig. 5B). Some time courses displayed a repetitive firing pattern, whereas others showed a sustained fluorescence, reminiscent of long-duration Ca2+ sparks seen previously (19, 32), often preceded by a graded fluorescence increase and/or followed by a graded decrease in fluorescence.
Extracellular Ca2+ Modulates Ca2+ Event Frequency
Ca2+ entry via L-type Ca2+ channels is reported to be important for EC coupling in embryonic muscle (11, 25). We therefore investigated the possible role of Ca2+ influx in activation of Ca2+ sparks in E18 diaphragm fibers. We first examined the effects of [Ca2+]o on the occurrence of Ca2+ sparks. For these studies, each enzymatically isolated E18 diaphragm fiber was first imaged at three locations in XY mode while bathed in 1.8 mM Ca2+ Ringer solution (control). The same fiber was then imaged again at the same locations after 10 min of additional exposure to control Ringer (1.8 mM Ca2+), 0 Ca2+ Ringer containing 1 mM EGTA, or 8 mM Ca2+ Ringer, and the Ca2+ event frequency was determined while the fiber was bathed in the test solution (0, 1.8, or 8 mM Ca2+ Ringer). The mean event frequency of all fibers in a given solution was normalized to the mean event frequency of the same fibers while bathed in the initial control Ringer solution. The resulting mean relative Ca2+ spark rates of fibers bathed in 0, 1.8, or 8 mM Ca2+ Ringer during the test period were, respectively, 0.61 ± 0.09, 1.11 ± 0.15, and 1.90 ± 0.21 of the rate of events from the same fibers in the initial 1.8 mM Ca2+ Ringer (control) (Fig. 6A). The increase of the mean Ca2+ spark rate in fibers bathed in high concentrations of external Ca2+ and the decrease of event frequency in fibers bathed in 0 Ca2+ clearly demonstrate that Ca2+ spark production is strongly influenced by extracellular Ca2+, possibly due to changes in Ca2+ influx. However, it has been previously shown that Ca2+-free solution causes slight depolarization of neonatal fibers (11), which could also increase spark frequency. Our experiments argue against this possibility because application of a nonspecific Ca2+ channel blocker, CoCl2 (5 mM), also suppressed spark frequency (Fig. 7), an effect independent of fiber depolarization (11). Furthermore, although surface charge effects might partially contribute to the observed effect of 0 mM Ca2+ Ringer solution, changes in surface charge would be opposite with cobalt addition.
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Possible alterations in SR store Ca2+ content were assayed in a separate group of fibers by using fura 2 (see MATERIALS AND METHODS). Figure 8A presents fura 2 ratio signals from a fiber in response to application of 20 mM caffeine in control (1.8 mM Ca2+) Ringer solution and from the same fiber after 10 min of to exposure to 0 mM Ca2+ Ringer. The similarity of the Ca2+ transients indicates minimal alteration in store content after 10 min in 0 Ca2+ solution. Average peak amplitudes of test caffeine responses after 0 mM (n = 2) or 8 mM (n = 3) Ca2+ Ringer, each normalized to control response in the same fiber, were similar (Fig. 8B). This finding indicates minimal external Ca2+-dependent change in store content. Thus the observed effect of Ca2+ on Ca2+ spark frequency is not likely to be due to changes in store content.
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An influx of extracellular Ca2+ could conceivably
increase the frequency of Ca2+ sparks by increasing the
cytosolic [Ca2+] in the vicinity of the RyRs, thereby
increasing the probability that a RyR Ca2+ release
channel would open by a CICR mechanism. Ca2+ influx
could initiate the opening of RyR Ca2+ release channels
via CICR either by raising the average level of cytosolic
Ca2+ or by producing local increases of
Ca2+ in the vicinity of the RyRs. To determine whether
average cytosolic [Ca2+] was altered when
Ca2+ spark frequency was found to vary with
[Ca2+]o, relative levels of cytosolic
Ca2+ were monitored using the fiber fluorescence within
the same fiber images, as used for spark detection (above). Fiber fluorescence
was measured in the fluorescence (F) images after exclusion of areas
encompassing potential events (F/F
1.5 SD). The mean fluorescence
of each region of each fiber bathed in 0, 1.8, or 8 mM
Ca2+ Ringer was normalized to the mean fiber
fluorescence of the same region of the same fiber while bathed in the initial
normal Ringer. The relative fiber fluorescence exhibited no significant change
with regard to [Ca2+]o
(Fig. 6B), suggesting
that the mean cytosolic [Ca2+] does not play a
signifi-cant role in stimulating Ca2+ spark production.
However, even though there was no significant change in global cytosolic
[Ca2+], it is still possible that
[Ca2+] in the microdomain of the triad junction varied
with the applied [Ca2+]o without any observed
changes in average fiber fluorescence.
Ca2+ Influx from L-type Ca2+ Channels
To determine whether Ca2+ influx via plasmalemma or T-tubule Ca2+ channels was modulating the observed event frequency, we measured the spark rate after blocking specific Ca2+ channels known to contribute to the inward Ca2+ current. The L-type Ca2+ current (Islow) is the primary Ca2+ current measured in adult mammalian skeletal muscle and increases substantially during postnatal development. Thus E18 diaphragm fibers were imaged while in normal Ringer and again after a 30-min exposure to Ringer solution containing 30 µM nifedipine, which is a dihydropyridine that selectively inhibits L-type Ca2+ channels found in the plasmalemma and T-tubules. As a control, embryonic fibers were imaged in Ringer and then imaged again after being bathed in Ringer solution without nifedipine for 30 min. The mean frequency of events in fibers treated with nifedipine dropped to 17.6% compared with the Ringer control after both were normalized to the mean event frequency of those same fibers bathed in Ringer before treatment (Fig. 7A). The significant pronounced decline in Ca2+ spark rate in nifedipine demonstrates that inhibition of the DHPRs drastically reduces the frequency of Ca2+ sparks and indicates that Ca2+ influx via L-type Ca2+ channels can modulate the Ca2+ spark rate in embryonic muscle fibers.
Additionally, embryonic and neonatal mouse skeletal muscles also exhibit a T-type (24) Ca2+ current (Ifast), which disappears by the third week after birth (2). Islow is present in skeletal muscle cells of normal animals but absent from skeletal muscle cells of mdg animals, whereas Ifast is present in developing skeletal muscle cells of both normal and mutant animals, indicating that different channel species give rise to these two currents (3). To determine whether Ca2+ influx via T-type Ca2+ channels has an effect on the Ca2+ spark frequency, we imaged E18 diaphragm fibers in control Ringer solution and then again after a 30-min exposure to 5 µM NiCl2 to block T-type Ca2+ channels (4). Inhibition of T-type Ca2+ channels without affecting L-type Ca2+ channels requires a very low concentration of NiCl2. Berthier et al. (4) reported NiCl2 inhibition of the T-type Ca2+ current with an IC50 of 5.4 µM in embryonic mouse skeletal muscle. Thus in our studies we used 5 µM NiCl2, which should block about half of the T-type Ca2+ channels. Application of 5 µM NiCl2 had no significant affect on the Ca2+ spark frequency (Fig. 7A), suggesting that Ca2+ entering via T-type channels may not play a role or may only play a minor role in contributing to the Ca2+ spark frequency. There were no significant differences in the resting fiber fluorescence (Fig. 7B).
Characterization of Events Influenced by Ca2+ Influx
We also compared the spatial properties of events imaged in XY
mode of fibers bathed in Ringer containing varying concentrations of
Ca2+ or in fibers treated with
Ca2+ channel blockers. The mean properties of peak
amplitude (F/F), EDHM (µm), and VIHM (µm3 ·
F/F) did not change significantly when
[Ca2+]o was altered
(Table 2). With
Ca2+ channel blockers, significant differences were
observed between a limited number of parameter values, but systematic patterns
were not observed (Table 2).
Histograms, normalized to the total number of events, of property
characteristics in each condition showed similar distributions (data not
shown).
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Spontaneous Electrical Activity
We also examined whether the Ca2+ sparks detected in the embryonic diaphragm preparation were produced as a result of spontaneous Na+-dependent action potentials. Treatment with tetrodotoxin (TTX), which blocks Na+ channels, was used to prevent Na+-dependent fiber action potentials. To overcome TTX insensitivity observed in embryonic and neonatal mammalian skeletal muscle (17), we exposed embryonic fibers to high concentrations of TTX (10 µM) (12). Three E18 diaphragm fibers were imaged while in Ringer and were then imaged again immediately after the fiber was exposed to Ringer containing TTX. The mean spark frequency did not change significantly (data not shown), demonstrating that Na+-dependent action potentials are not the cause of Ca2+ sparks in embryonic mammalian skeletal muscle. We also saw no evidence for spontaneous Ca2+ waves in the fibers used for these experiments (data not shown).
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DISCUSSION |
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We next demonstrated that in mouse diaphragm, the ratio of the protein expression levels of the two muscle RyR isoforms, RyR3 to RyR1, does not change appreciably over the developmental period from E18 to P14. This observation confirms the earlier findings of Flucher et al. (16) regarding RyR isoform expression changes during mouse muscle development and establishes that in the diaphragm, the decline in Ca2+ spark frequency from E18 to P14 cannot be attributed to a relative decline in RyR3 protein expression compared with RyR1.
In a third and unanticipated new observation, we have now found that the frequency of occurrence of Ca2+ sparks in embryonic muscle fibers is markedly influenced by Ca2+ influx via L-type Ca2+ channels in the plasmalemma or transverse tubules. Raising or lowering [Ca2+]o causes the frequency of spontaneous Ca2+ sparks in E18 diaphragm to respectively increase or decrease, and inhibition of plasmalemmal or T-tubule Ca2+ channels by either the nonspecific Ca2+ channel blocker CoCl2 (5 mM) or the specific L-type Ca2+ channel blocker nifedipine (30 µM) causes a marked reduction in spark frequency. Thus Ca influx via DHPR (L-type) Ca2+ channels appears to be involved in triggering the opening of the RyR Ca2+ release channel(s) that underlies the spark. The likely mechanism for such an effect in embryonic muscle would be a local elevation of cytosolic [Ca2+] in the immediate vicinity of an open L-type channel and the resulting activation of a RyR Ca2+ release channel by CICR (Fig. 9A). The effect of Ca2+ influx via L-type channels appears to be quite localized, because there were no detectable changes in global cytosolic fluorescence under conditions that caused clear variations in Ca2+ spark frequency. Changes in SR Ca2+ content were not involved in mediating the observed effects of [Ca2+]o on spark frequency.
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Development of DHPR-RyR1 Coupling and Cessation of Spontaneous Ca2+ Sparks
Together, our three major findings now suggest the hypothesis that the decline in spark frequency in early postnatal development might be due to either a decline in Ca2+ influx in resting fibers via DHPR L-type channels from E18 to P14 or to a decline in the ability of such Ca2+ influx to initiate opening of RyRs from E18 to P14. Interestingly, the previously studied development of functional and/or structural couplings between the DHPR and RyR1 (26, 27) could provide a molecular mechanism for a developmental decrease in activation of Ca2+ sparks by Ca2+ entry via DHPR L-type channels. Consistent with this hypothesis, it has previously been suggested that sparks in RyR3 knockout myotubes were produced by islands of RyR1 channels, which were not coupled to DHPR voltage sensors (29).
There are several possible mechanisms whereby the development of DHPR-RyR1 coupling could give rise to the observed decrease in the frequency of occurrence of Ca2+ sparks. First, if coupled DHPRs generated much less Ca2+ influx in resting fibers than uncoupled DHPRs (Fig. 9B), the Ca2+ signal for RyR opening by CICR would be decreased by DHPR-RyR1 coupling and fewer sparks would occur.
Alternatively, coupled DHPRs could still carry the same current and exhibit the same opening probabilities and open times in resting fibers as uncoupled DHPRs, but the likelihood that a RyR Ca2+ release channel would be activated by Ca2+ from a nearby DHPR could still be decreased as a result of DHPRRyR1 coupling (not shown in Fig. 9). Decreased RyR activation in response to a given DHPR current after coupling of DHPRs to RyR1 could occur by either or both of two possible mechanisms. In the first mechanism, coupling to a DHPR could directly decrease the Ca2+ sensitivity for CICR activation of a coupled RyR1. In this case, a given Ca2+ current through a coupled DHPR could be unlikely to open its coupled RyR1, whereas the same current through an uncoupled DHPR would be more likely to activate a nearby but uncoupled RyR1 or RyR3. As a second possibility, RyR1s (coupled or uncoupled) could be less susceptible to activation by CICR than RyR3s (5, 9, 14, 22, 32). In this case, the structural reorganization induced by coupling, where RyR1s but not RyR3s are now coupled and thus closest to the DHPR, would restrict RyR3s from closest proximity to coupled DHPRs (13). The RyR3s would then be further from the activating trigger Ca2+ signal and, consequently, would experience lower local [Ca2+] elevation and thus be less likely to open in response to a given Ca2+ influx through coupled than through uncoupled DHPRs.
It is also possible that changes in Ca2+ influx via DHPRs could occur independently of changes in DHPR/RyR coupling (Fig. 9C). For example, Ward and Wareham (34) reported membrane hyperpolarization during postnatal development of mouse EDL fibers (from 42 mV at P4 to 54 mV at P8 and 67 mV at P16), which could give rise to less frequent opening of DHPR Ca2+ channels, resulting in decreased frequency of Ca2+ spark occurrence. These and other hypothetical possibilities require experimental investigation in future studies.
In conclusion, the frequency of spontaneous Ca2+ events is greatest in embryonic mouse skeletal muscle and declines significantly over a 2-wk period postbirth. The Ca2+ sparks in embryonic mouse diaphragm fibers can be modulated by Ca2+ influx via L-type Ca2+ channels.
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DISCLOSURES |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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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.
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REFERENCES |
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2. Beam KG and Knudson CM. Effect of postnatal development on calcium currents and slow charge movement in mammalian skeletal muscle. J Gen Physiol 91: 799815, 1988.[Abstract]
3. Beam KG, Knudson CM, and Powell JA. A lethal mutation in mice eliminates the slow calcium current in skeletal muscle cells. Nature 320: 168170, 1986.[ISI][Medline]
4. Berthier C,
Monteil A, Lory P, and Strube C. 1H mRNA in single
skeletal muscle fibers accounts for T-type calcium current transient
expression during fetal development in mice. J Physiol
539: 681691,
2002.
5. Chen SRW,
Ebisawa K, Li X, and Zhang L. Molecular identification of the ryanodine
receptor Ca2+ sensor. J Biol
Chem 273:
1467514678, 1999.
6. Cheng H, Song
L, Shirokova N, Gonzalez A, Lakatta EG, Rios E, and Stern MD. Amplitude
distribution of calcium sparks in confocal images: theory and studies with an
automatic detection method. Biophys J
76: 606617,
1999.
7. Cheung A, Dantzig JA, Hollingworth S, Baylor SM, Goldman YE, Mitchison TJ, and Straight AF. A small-molecule inhibitor of skeletal muscle myosin II. Nat Cell Biol 4: 8388, 2002.[ISI][Medline]
8. Cognard C, Rivet-Bastide M, Constantin B, and Raymond G. Progressive predominance of "skeletal" versus "cardiac" types of excitation-contraction coupling during in vitro skeletal myogenesis. Pflügers Arch 422: 207209, 1992.[ISI][Medline]
9. Conklin MS,
Ahern CA, Vallejo P, Sorrentino V, Takeshima H, and Coronado R. Comparison
of Ca2+ sparks produced independently by two ryanodine
receptor isoforms (type 1 and type 3). Biophys J
78: 17771785,
2000.
10. Conklin MW,
Barone V, Sorrentino V, and Coronado R. Contribution of ryanodine receptor
type 3 to Ca2+ sparks in embryonic mouse skeletal
muscle. Biophys J 77:
13941403, 1999.
11. Dangain J and Neering IR. Effect of low extracellular calcium and ryanodine on muscle contraction of the mouse during postnatal development. Can J Physiol Pharmacol 69: 12941300, 1991.[ISI][Medline]
12. DeDeyne P.
Formation of sarcomeres in developing myotubes: role of mechanical stretch and
contractile activation. Am J Physiol Cell Physiol
279: C1801C1811,
2000.
13. Felder E and
Franzini-Armstrong C. Type 3 ryanodine receptors of skeletal muscle are
segregated in a parajunctional position. Proc Natl Acad Sci
USA 99:
1695700, 2002.
14. Fessenden JD,
Wang Y, Moore RA, Chen SRW, Allen PD, and Pessah IN. Divergent functional
properties of ryanodine receptor types 1 and 3 expressed in a myogenic cell
line. Biophys J 79:
25092525, 2000.
15. Flucher BE. Structural analysis of muscle development: transverse tubules, sarcoplasmic reticulum, and the triad. Dev Biol 154: 245260, 1992.[ISI][Medline]
16. Flucher BE,
Conti A, Takeshima H, and Sorrentino V. Type 3 and type 1 ryanodine
receptors are localized in triads of the same mammalian skeletal muscle
fibers. J Cell Biol 146:
621629, 1999.
17. Harris JB and Marshall MW. Tetrodotoxin-resistant action potentials in newborn rat muscle. Nat New Biol 243: 191192, 1973.[Medline]
18. Kandarian SC,
Peters DG, Taylor JA, and Williams JH. Skeletal muscle overload
upregulates the sarcoplasmic reticulum slow calcium pump gene. Am J
Physiol Cell Physiol 266:
C1190C1197, 1994.
19. Kirsch WG,
Uttenweiler D, and Fink RH. Spark- and ember-like elementary
Ca2+ release events in skinned fibers of adult mammalian
skeletal muscle. J Physiol 537:
379389, 2001.
20. Laemmli UK. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227: 680685, 1970.[ISI][Medline]
21. Marks AR, Tempst P, Hwang KS, Taubman MB, Inui M, Chadwick C, Fleischer S, and Nadal-Ginard B. Molecular cloning and characterization of the ryanodine receptor/junctional channel complex cDNA from skeletal muscle sarcoplasmic reticulum. Proc Natl Acad Sci USA 86: 86838687, 1989.[Abstract]
22. Murayama T, Oba
T, Katayama E, Oyamada H, Oguchi K, Kobayashi M, Otsuka K, and Ogawa Y.
Further characterization of the type 3 ryanodine receptor (RyR3) purified from
rabbit diaphragm. J Biol Chem
274: 1729717308,
1999.
23. Murayama T and Ogawa Y. Purification and characterization of two ryanodine-binding protein isoforms from sarcoplasmic reticulum of bullfrog skeletal muscle. J Biochem (Tokyo) 112: 514522, 1992.[Abstract]
24. Nowycky MC, Fox AP, and Tsien RW. Three types of neuronal calcium channel with different calcium agonist sensitivity. Nature 316: 440443, 1985.[ISI][Medline]
25. Péréon Y, Louboutin JP, and Noireaud J. Contractile responses in rat extensor digitorum longus muscles at different times of postnatal development. J Comp Physiol [B] 163: 203211, 1993.[ISI][Medline]
26. Protasi F,
Franzini-Armstrong C, and Allen PD. Role of ryanodine receptors in the
assembly of calcium release units in skeletal muscle. J Cell
Biol 140:
831842, 1998.
27. Protasi F,
Takekura H, Wang Y, Chen SR, Meissner G, Allen PD, and Franzini-Armstrong
C. RYR1 and RYR3 have different roles in the assembly of calcium release
units of skeletal muscle. Biophys J
79: 24942508,
2000.
28. Schneider MF and Chandler WK. Voltage dependent charge movement of skeletal muscle: a possible step in excitation-contraction coupling. Nature 242: 244246, 1973.[ISI][Medline]
29. Shirokova N,
Garcia J, and Rios E. Local calcium release in mammalian skeletal muscle.
J Physiol 512:
377384, 1998.
30. Takekura H, Flucher BE, and Franzini-Armstrong C. Sequential docking, molecular differentiation, and positioning of T-tubule/SR junctions in developing mouse skeletal muscle. Dev Biol 239: 204214, 2001.[ISI][Medline]
31. Takeshima H, Nishimura S, Matsumoto T, Ishida H, Kangawa K, Minamino N, Matsuo H, Ueda M, Hanaoka M, Hirose T, and Numa S. Primary structure and expression from complementary DNA of skeletal muscle ryanodine receptor. Nature 339: 439445, 1989.[ISI][Medline]
32. Ward CW,
Protasi F, Castillo D, Wang Y, Chen SRW, Pessah IN, Allen PD, and Schneider
MF. Type 1 and type 3 ryanodine receptors generate different
Ca2+ release event activity in both intact and
permeabilized myotubes. Biophys J
81: 32163230,
2001.
33. Ward CW,
Schneider MF, Castillo D, Protasi F, Wang Y, Chen SRW, and Allen PD.
Expression of ryanodine receptor RyR3 produces Ca2+
sparks in dyspedic myotubes. J Physiol
525.1: 91103,
2000.
34. Ward KM and Wareham AC. Changes in membrane potential and potassium and sodium activities during postnatal development of mouse skeletal muscle. Exp Neurol 89: 554568, 1985.[ISI][Medline]
35. Wu K and Lytton
J. Molecular cloning and quantification of sarcoplasmic reticulum
Ca2+-ATPase isoforms in rat muscles. Am J
Physiol Cell Physiol 264:
C333C341, 1993.