Am J Physiol Cell Physiol 290: C539 –C553, 2006.
First published September 7, 2005; doi:10.1152/ajpcell.00592.2004.
A probable role of dihydropyridine receptors in repression
of Ca2⫹ sparks demonstrated in cultured mammalian muscle
Jingsong Zhou,1 Jianxun Yi,1 Leandro Royer,1 Bradley S. Launikonis,1
Adom González,1 Jesús Garcı́a,2 and Eduardo Rı́os1
1
Section of Cellular Signaling, Department of Molecular Biophysics and Physiology, Rush University, Chicago;
and 2Department of Physiology and Biophysics, University of Illinois at Chicago, Chicago, Illinois
Submitted 3 December 2004; accepted in final form 30 August 2005
excitation-contraction coupling; sarcoplasmic reticulum; ryanodine
receptors; Ca2⫹ imaging
contraction is triggered by Ca2⫹ release
from the sarcoplasmic reticulum (SR) through ryanodine receptor (RyR) channels. The state of these channels is determined by the action potential by means of dihydropyridine
receptors (DHPRs), voltage sensors located in the T tubular
membrane. Most current studies of this regulation focus on the
activation or opening of the release channels, which is known
to require an initial action, most probably mechanical, by the
voltage sensors. Ca2⫹ release initiated thus may then be reinforced by Ca2⫹-induced Ca2⫹ release (CICR), a secondary
activation mediated by Ca2⫹ itself (14, 21). In addition to these
positive actions by voltage sensors and Ca2⫹ ions, there is
IN STRIATED MUSCLE,
Address for reprint requests and other correspondence: J. Zhou, Dept. of
Molecular Biophysics and Physiology, Rush Univ., 1750 W. Harrison St.,
Suite 1279 JS, Chicago, IL 60612 (e-mail: [email protected]).
http://www.ajpcell.org
evidence of an inhibitory mechanism that is “dynamic,”
brought to bear only after Ca2⫹ channel opening and Ca2⫹
release (2, 36, 43, 46). This inhibition is thought to be crucial
for maintaining stability and speed of control in a system
whose activation may be highly self sustaining as a consequence of CICR, and appears to be also mediated by Ca2⫹.
Although much attention has been paid to this dynamic inhibition/inactivation mechanism, some evidence has emerged of
a role of DHPRs in inhibition of release channels.
Initial indications of a basal inhibition by DHPRs originated
from comparisons of excitation-contraction (EC) coupling between amphibian and mammalian muscle. The comparisons
led Shirokova et al. (51) and Zhou et al. (67) to conclude, in
agreement with earlier studies (15, 16), that CICR has a limited
or null role in mammalian muscle. An important contribution
of CICR, however, was upheld for frog muscle.
The apparent differences in the contributions of CICR between amphibians and mammals ought to be explained by the
different molecular makeup and supramolecular assembly of
their EC coupling devices. Whereas frog fast-twitch muscles
contain the ␣- and ␤-isoforms, in approximately equal densities (42, 56), most muscles of adult mammals exclusively
express the ␣-homolog, RyR1 (19, 34, 58), whereas some
additionally express the ␤-homolog, RyR3, in minor amounts.
According to Felder and Franzini-Armstrong (17), ␣RyRs
form separate arrays of channels, placed parajunctionally, on
the sides of the triadic junction. Junctional clusters are instead
constituted exclusively by the ␣ (or, in mammals, the RyR1)
isoform. Because only the latter are associated with tetrads of
DHPRs in the T tubule membrane (3), the activation mechanisms of these two sets of channels must be distinct (67).
The parajunctional channels, which lack directly interacting
DHPRs, can be under voltage control only indirectly, presumably by CICR. Conversely, the location of junctional channels
should allow them to be directly controlled by voltage sensors.
In agreement with these functional expectations, studies in 1B5
RyR-deficient myotubes show that RyR1 can be controlled by
membrane voltage, whereas RyR3, if expressed alone, can only
be activated via CICR (18).
The notion of a basal inhibition originated in good measure
from the very different incidence of Ca2⫹ sparks in muscles of
frogs and mammals. Ca2⫹ sparks are discrete local elevations
of Ca2⫹ concentration ([Ca2⫹]), which reflect opening of a
group of RyR channels spontaneously or in response to voltage
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C539
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Zhou, Jingsong, Jianxun Yi, Leandro Royer, Bradley S. Launikonis, Adom González, Jesús Garcı́a, and Eduardo Rı́os. A probable role of dihydropyridine receptors in repression of Ca2⫹ sparks
demonstrated in cultured mammalian muscle. Am J Physiol Cell
Physiol 290: C539 –C553, 2006. First published September 7, 2005;
doi:10.1152/ajpcell.00592.2004.—To activate skeletal muscle contraction, action potentials must be sensed by dihydropyridine receptors (DHPRs) in the T tubule, which signal the Ca2⫹ release channels
or ryanodine receptors (RyRs) in the sarcoplasmic reticulum (SR) to
open. We demonstrate here an inhibitory effect of the T tubule on the
production of sparks of Ca2⫹ release. Murine primary cultures were
confocally imaged for Ca2⫹ detection and T tubule visualization.
After 72 h of differentiation, T tubules extended from the periphery
for less than one-third of the myotube radius. Spontaneous Ca2⫹
sparks were found away from the region of cells where tubules were
found. Immunostaining showed RyR1 and RyR3 isoforms in all areas,
implying inhibition of both isoforms by a T tubule component. To test
for a role of DHPRs in this inhibition, we imaged myotubes from
dysgenic mice (mdg) that lack DHPRs. These exhibited T tubule
development similar to that of normal myotubes, but produced few
sparks, even in regions where tubules were absent. To increase spark
frequency, a high-Ca2⫹ saline with 1 mM caffeine was used. Wildtype cells in this saline plus 50 ␮M nifedipine retained the topographic suppression pattern of sparks, but dysgenic cells in high-Ca2⫹
saline did not. Shifted excitation and emission ratios of indo-1 in the
cytosol or mag-indo-1 in the SR were used to image [Ca2⫹] in these
compartments. Under the conditions of interest, wild-type and mdg
cells had similar levels of free [Ca2⫹] in cytosol and SR. These data
suggest that DHPRs play a critical role in reducing the rate of
spontaneous opening of Ca2⫹ release channels and/or their susceptibility to Ca2⫹-induced activation, thereby suppressing the production
of Ca2⫹ sparks.
C540
REPRESSION OF CA2⫹ SPARKS BY T TUBULES
METHODS
Cultures of skeletal muscle myotubes. Primary cultures were prepared from the skeletal muscle of neonatal mice (postnatal day 0) of
the stock CACNA1SxNIHS-BCfBR (39, 47). Myotubes from wild
type (mdg⫺/⫺) or heterozygotes (mdg⫹/⫺) were undistinguishable.
Dysgenic or mdg myotubes (mdg⫹/⫹) lack DHPRs in the T tubules
(29) and lack EC coupling (59).
Muscle tissue was dissected from the limbs and digested with 2
mg/ml collagenase I (Worthington Biochemical, Lakewood, NJ) in 0
Ca2⫹-0 Mg2⫹ PBS (GIBCO, Carlsbad, CA) at 37°C for 30 min.
AJP-Cell Physiol • VOL
Myoblasts were collected by passing the suspension through a 40␮m-thick nylon filter (Falcon, Bedford, MA), seeded in glass-bottomed microwell dishes (MatTek, Ashland, MA), and maintained in
the growth medium (DMEM, 20% FBS, GIBCO) for 2 days before
they differentiated into myotubes in the differentiation medium
(DMEM, 2.5% horse serum). Images were obtained from cultures that
underwent differentiation for 2– 4 days. Experiments were performed
in compliance with National Institutes of Health guidelines and were
approved by the Institutional Animal Care and Use Committee of
Rush University.
Solutions. The Krebs solution was composed of (in mM) 136 NaCl,
5 KCl, 2.5 CaCl2, 1 MgCl2, 10 HEPES, and 10 glucose, pH 7.0.
High-Ca2⫹ Krebs (K/Ca) was composed of (in mM) 118 NaCl, 5 KCl,
26 CaCl2, 1 MgCl2, 10 HEPES, and 10 glucose, pH 7.0, caffeine
(K/Ca⫹C), with 50 ␮M nifedipine (K/Ca⫹N), or both (K/Ca⫹NC),
were added for different purposes. All solutions contained 1 ␮M
tetrodotoxin to prevent spontaneous activation.
For dye loading, cell cultures were immersed in Krebs solution
with 5 ␮M of either indo-1 AM or mag-indo-1 AM. For membrane
permeabilization, cultures were immersed in an internal solution
containing (in mM) 130 K-glutamate, 10 Trizma maleate, 5 Na2ATP,
10 Tris-PC, 1 EGTA, 5 glucose, 8% dextran, 0.1 CaCl2, 7.23 MgCl2
(free Ca2⫹ ⫽ 50 nM, free Mg2⫹ ⫽ 1 mM), and 0.05 rhod-2 plus
0.002% saponin.
The solutions were prepared and used at 18 –20°C. Chemicals for
these solutions were purchased from Sigma-Aldrich (St. Louis, MO).
Optical measurements and data analysis. For detection of spontaneous Ca2⫹ sparks, cultures were incubated with 5 ␮M fluo-4 AM
(Molecular Probes, Eugene, OR) in Krebs solution for 50 min at 20°C,
then washed and maintained in fresh Krebs solution for an additional
40 min. For double-staining experiments, the cultures were exposed to
10 ␮M 4-{2-[6-(dioctylamino)-2-naphthalenyl]ethenyl}1-(3-sulfopropyl)-pyridinium (di-8-ANEPPS) (Molecular Probes) for ⬍15 min to
minimize its entry into organelles. The dye was then washed away
with Krebs solution.
Cells were imaged with a confocal scanner equipped with a ⫻40
water-immersion objective (model MRC-1000, Zeiss; numerical aperture 1.2). Dual images were obtained with excitation light of 488 nm
and simultaneous recording at ⬁ ⫽ 530 nm (⫾25-nm bandwidth) for
detection of cytosolic Ca2⫹ events, and ⬁ ⬎588 nm for detection of
structures stained by di-8-ANEPPS. Ca2⫹ events were detected automatically in xt or xy images and normalized as described by Brum et
al. (4). The detection program carries out automatic determination of
spark parameters: amplitude (peak F/F0), full width above half magnitude (FWHM), full duration above half magnitude (FDHM), and
rise time (from 10% to full peak at the spatial center of the spark) (66).
The location of sparks in myotubes relative to the surface membrane was represented by a number that quantified it relative to the
cell radius, with 0 corresponding to the membrane and 1 to the cell
center. The fractional radius of the myotube invaded by T tubules (fT)
was determined by comparison of T-tubule-free and occupied areas
(dashed line in Fig. 1B) with midcell xy images of doubly stained
myotubes.
SEER imaging of myotubes. Two applications were implemented:
determination of [Ca2⫹]cyto in intact cells loaded with indo-1 and
imaging of [Ca2⫹]SR in membrane-permeabilized cells with magindo-1 inside organelles.
SEER (31) required the simultaneous acquisition of two confocal
fluorescence images (named F11 and F22) produced by alternating line
by line (“line interleaving”) two excitation lights (351 and 364 nm)
and two fluorescence emission ranges (390 – 440 nm and 465–535
nm). A third image, F33, was triply interleaved when rhod-2 was
present in the perfusion medium, for the purpose of demonstrating
permeabilization of the plasmalemma. F33 was excited at 543 nm and
acquired at 550 – 630 nm. Imaging was done with the confocal system
(model TCS SP2; Leica Microsystems, Exton, PA), using a ⫻63
water-immersion objective, numerical aperture 1.2. Examples of im-
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or other stimuli (6, 61). The activation of multiple channels in
Ca2⫹ sparks is believed to involve Ca2⫹ as mediator. This
consensus derives largely from evidence collected in frog muscle, including that sparks are promoted by increases in resting
cytosolic [Ca2⫹] ([Ca2⫹]cyto) (28), by increases in SR Ca2⫹
([Ca2⫹]SR), and by caffeine (24), which increases RyR sensitivity to activation by Ca2⫹ (15, 35).
In contrast with amphibians, sparks are rarely observed (10)
or not found at all (66) in intact adult skeletal muscle cells of
mammals, and are not elicited in adult cells by voltage-clamp
depolarization (11, 52). Spark frequency, however, becomes
much greater under other conditions, including membrane
permeabilization by saponin or its removal by “peeling” (27,
66) and interference with mitochondrial function (26). Therefore, sparks are possible in the mammal, but appear to be
suppressed. Comparisons of spark frequency in 1B5 dyspedic
cells expressing alternatively either isoform demonstrated that
the suppression affects largely isoform 1 (62).
Suppression of a related sort was revealed by Murayama and
Ogawa (37, 38) affecting bovine RyR1 and amphibian ␣-isoforms. Because ␤ was not affected, the stabilization may
explain the greater contribution of CICR in the amphibian. The
authors provided evidence pointing at FKBP12 as a possible
effector of this inhibition in mammalian muscle.
The inhibition has an intriguing functional feature. Using
primary mammalian myotubes, Shirokova et al. (53) showed
dual spatially segregated manifestations of Ca2⫹ release.
Whereas some regions of the myotube featured strict control of
Ca2⫹ release by membrane voltage (of the “skeletal” type, i.e.,
not dependent on Ca2⫹ entry), other regions did not respond to
voltage, but produced Ca2⫹ sparks essentially unrelated to the
depolarizing pulses. These two forms of Ca2⫹ release appear to
be mutually exclusive (regions that respond to voltage do not
produce sparks), which suggests that something in the devices
underpinning voltage control, perhaps the DHPR itself, suppresses spark generation. Consistent with these ideas, Lee et al.
(33) provided evidence of an inhibitory allosteric effect in 1B5
myotubes. A putative molecular locus underpinning the suppression is indicated by the existence of a malignant hyperthermia-inducing mutation in the III-IV loop of the DHPR in
humans (65).
Motivated by these varied observations, the present study set
out to test whether T tubules play a role in the suppression of
sparks. We also explored, using myotubes lacking DHPRs,
whether the voltage sensors themselves are needed for the
inhibition. Because the production of sparks in mdg muscle
required the use of special nonphysiological conditions, we
used the shifted excitation and emission ratio (SEER) imaging
technique (31) to compare [Ca2⫹]cyto and intrastore [Ca2⫹] in
both types of cells under those conditions.
REPRESSION OF CA2⫹ SPARKS BY T TUBULES
C541
Fig. 1. Images of a dually stained myotube. Repeated confocal
images of fluorescence of fluo-4:Ca were obtained simultaneously with fluorescence of image showing 4-{2-[6-(dioctylamino)-2-naphthalenyl]ethenyl}1-(3-sulfopropyl)-pyridinium (di-8-ANEPPS). A: a spark, normalized to an average of
6 images without Ca2⫹ events. B: average of 6 images, contrast-enhanced fluorescence of di-8-ANEPPS, showing developing T tubules. Note that sparks usually occur in central areas,
free of developing T tubules. The dashed line, demarcating the
area free of T tubules, was drawn by sight and used as
described in METHODS to calculate a numerical indicator of
radial development of T tubules (fT).
关Ca 2⫹兴 ⫽ KD␤
R ⫺ Rmin
Rmax ⫺ R
(1)
where Rmin and Rmax are minimum and maximum ratio values, KD
is the dissociation constant of the dye, and ␤ is the ratio of F22 at
[Ca2⫹] ⫽ 0 to F22 at saturating [Ca2⫹]. Parameter values are given by
Launikonis et al. (31). Equation 1 and the equation published by
Grynkiewicz et al. (25) differ only in the definition of ␤.
Measurement of [Ca2⫹]cyto. For the measurement of [Ca2⫹]cyto
with indo-1, the culture dishes were loaded and then imaged intact. As
with any procedure on intact cells loaded with a dye in AM form,
there will be some interference by the dye that accumulates inside
organelles. A correction for this error, developed in the APPENDIX,
leads to an equation isomorphic with 1
关Ca 2⫹兴 ⫽ Keff␤
R ⫺ R0
Rmax ⫺ R
(2)
where Keff and R0 are effective values of KD and Rmin. Their
expressions in terms of the other parameters are given in the APPENDIX.
Calibration experiments to determine the parameters of Eq. 2 were
carried out in cultures loaded with indo-1, so that the organelles would
cause similar interference as in the intact cells. After incubation with
indo-1 AM, the loaded cells were membrane permeabilized and SEER
imaged in solutions with known buffered [Ca2⫹] and 50 ␮M indo-1.
Keff ␤ was determined by fitting Eq. 2 to the data points (R, [Ca2⫹]),
where R was the ratio averaged in those areas that satisfy a criterion
for good staining described above. The parameter values were R0 ⫽
0.51, Rmax ⫽ 3.94, Keff␤ ⫽ 1,620 nM, and ␤ ⫽ 4.11, implying that
Keff was 394 nM.
AJP-Cell Physiol • VOL
Statistics. Line graphs, such as Fig. 6F, plot averages of SEER ratio
values over all well-stained pixels of a cell or cells within an image.
The standard errors of such means are uniformly smaller than the
symbol size. When a bar is plotted, the symbols represent means over
averages of pieces of a cell or image. In those cases, the bar covers
means ⫾ 2 SE. Such piecewise approach was necessary to exclude
well-stained debris, the inclusion of which would have biased the
average. Significance of differences indicated in vertical bar graphs is
determined by two-tailed t-tests.
Immunofluorescence staining. Cultures were fixed for 20 min at
⫺20°C with methanol-acetone (1:1) precooled at ⫺20°C. Fixed cells
were incubated with blocking solution overnight at 4°C or 1 h at room
temperature. The blocking solution was PBS (GIBCO) containing
10% normal goat serum, 1% BSA, and 0.1% Triton X-100 (SigmaAldrich). The cells were incubated with the primary antibodies,
anti-RyR1 (1:1,000) or anti-RyR3 (1:2,000), diluted in the blocking
solution for 1 h at room temperature or overnight at 4°C, then
incubated with secondary antibodies, Cy2-conjugated goat anti-rabbit
(Jackson ImmunoResearch Laboratories, West Grove, PA), in the
blocking solution for 45 min at room temperature. Control experiments were done by replacing primary antibody with 1:1,000 normal
rabbit serum (Sigma-Aldrich) in blocking solution. The primary
antibodies, a gift from Dr. V. Sorrentino (University of Siena, Italy),
have been shown not to cross-react with each other (22).
RESULTS
A pattern of location of Ca2⫹ sparks in developing muscle.
A first set of experiments used developing myotubes, in which
T tubules were incomplete and largely occupied the peripheral
region, to explore whether there was a topographical correspondence between T tubule structures and the occurrence of
spontaneous sparks.
In myotubes produced in primary cultures from normal
newborn mice, Ca2⫹ sparks were observed consistently at a
low frequency. A total of 79 sparks were detected and characterized in xy images from 17 intact cells immersed in Krebs
solution. Sparks occurred more frequently in myotubes of
larger diameter, such as the one shown in Fig. 1. The culture
was dually stained with fluo-4 AM and the membrane-soluble
dye di-8-ANEPPS, and images were acquired simultaneously
at two emission bands. In Fig. 1A, the fluorescence of fluo-4 in
one image is normalized to the average of six other images
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ages F11(x,y) and F22(x,y) of cells loaded with indo-1 AM are
provided in Fig. 6, A and B. Images of a permeabilized cell loaded
with mag-indo-1AM are shown in Figs. 9, A and B, and 10B. F33 is
shown in Figs. 9 and 10. All other images are of the xy ratio [R(x,y)],
defined as F11(x,y)/F22(x,y), shown in areas considered to be well
stained (in other areas color was set to gray). The well-stained areas
were those with dye concentration comprised between the mean and
the mean plus three times the standard deviation of the pixel-by-pixel
distribution of dye concentration within the image. Dye concentration
was calculated from a linear combination of F11 and F22 (31).
Imaging of [Ca2⫹]SR. The technique was essentially that of Launikonis et al. (31). Cells loaded with mag-indo-1 had their plasma
membrane permeabilized, so that the dye remnant inside the cell was
all trapped in organelles, largely the SR. [Ca2⫹]SR is derived from the
ratio through pixel-by-pixel application of the equation
C542
REPRESSION OF CA2⫹ SPARKS BY T TUBULES
Fig. 2. Spatial location of spontaneous sparks of myotubes in Krebs solution.
“Distance” of the spark source from the cell surface along the scanning line is
normalized to cell radius, with 0 corresponding to the membrane and 1 to the
cell center. The bars at top represent the averaged value in 9 cells (0.22 ⫾ 0.06)
of fT, the fractional development of T tubules. The histogram shows the
distribution of 79 sparks detected in xy images from 19 cells.
AJP-Cell Physiol • VOL
(48). Figure 3A shows the image of an mdg cell labeled with
di-8-ANEPPS. After 3– 4 days in differentiation medium, dysgenic myotubes, like their wild-type counterparts, had a partially developed T tubule structure, occupying the peripheral
region to an extent similar to that in normal myotubes (fT ⫽
0.234 ⫾ 0.03, n ⫽ 8).
No spontaneous Ca2⫹ sparks were detected in dysgenic
myotubes in Krebs solution. Caffeine (10 mM) induced a
global release, which demonstrated integrity of the Ca2⫹
source (data not shown). Several probable causes for the
absence of spontaneous sparks were considered. One could be
a low Ca2⫹ load in the SR, which, according to evidence from
cardiac and skeletal muscle, would reduce event frequency (60,
67). It could be also a direct consequence of the absence of
plasmalemmal L-type Ca2⫹ channels, which have a role in the
initiation of Ca2⫹ sparks in embryonic mouse muscle (8), or it
could be a consequence of lower cytosolic Ca2⫹. All these
possibilities were tested by direct measurements, described
below.
To induce spark production, the cultures were exposed to
K/Ca⫹C, a modified Krebs with 26 mM Ca2⫹ and a low dose
of caffeine (1 mM). As shown in Fig. 3B, 315 sparks were
detected and characterized in xy images from 24 dysgenic
myotubes. The most remarkable difference with the normal
cells was that sparks could occur just inside the plasmalemma.
As shown in Fig. 3A, T tubules could be visualized well in
cells stained with di-8-ANEPPS. The spatial location of the
events, represented by the histogram in Fig. 4, bore little or no
relation with the location of T tubules (represented by the
horizontal bar). Therefore, in dysgenic cells, the suppression
effect on the releasing channel opening was lost under the
conditions required to observe sparks. This absence could
reflect a fundamental molecular influence or just a trivial
artifact.
Ca2⫹ overload erases the pattern of spark suppression in
normal cells. The lack of a topographic pattern of spark
generation in mdg myotubes might be simply a consequence of
Ca2⫹ overload (of cytosol or intracellular stores) in the high
extracellular [Ca2⫹] ([Ca2⫹]o) needed to induce spontaneous
sparks. A first control for this possibility was a study of the
spatial distribution of sparks in normal myotubes. The cells
were exposed to K/Ca⫹C, the high-Ca2⫹ Krebs. One hundred
and forty-two sparks were detected in xy images from twentyone cells. As shown by the histogram in Fig. 5A, the pattern of
locations was lost, with sparks appearing equally frequently in
all areas of the cells. According to this result, the loss of the
spatial pattern of spark origination in mdg cells could indeed be
a consequence of Ca2⫹ overload. A more specific control was
introduced, as described next.
Spatial pattern is maintained in high [Ca2⫹] if L-type
channels are blocked. It is reasonable to expect that a normal
myotube in a high-Ca2⫹ extracellular solution will acquire a
higher Ca2⫹ load (in cytosol and stores) than a dysgenic
myotube in the same solution, as L-type channels present in the
normal cell should provide additional paths for entry of Ca2⫹.
An alternative control condition was designed with the aim of
better reproduction of the membrane permeability of dysgenic
myotubes in wild-type cells. Sparks were studied in normal
myotubes in the high-Ca2⫹ solution K/Ca⫹NC, which included nifedipine, a specific L-type channel blocker (41) with
no direct effect on release channels (64). As shown later, in
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from the same region of the specimen. Figure 1B is the
averaged fluorescence of di-8-ANEPPS at longer wavelengths
in the same region. It allows visualization of T tubules.
Like the one shown in Fig. 1, most Ca2⫹ sparks occurred
away from the peripheral region. As shown in Fig. 1B, regions
where sparks were frequent did not have structures stained with
the membrane marker. A numerical location of the events
recorded in these images was defined as the ratio (distance
from center of spark to cell edge)/(cell radius). Figure 2 shows
the histogram of locations of all identified sparks.
The average extent of development of T tubules was measured in images of these and other cells stained with di-8ANEPPS. The extent of occupancy by T tubules, again as a
fraction of cell radius, represented by fT, was calculated as the
ratio of T tubule occupied to total area in images such as that
in Fig. 1B. The average of this ratio in 9 cells, 0.22 ⫾ 0.06, is
indicated by a horizontal bar in Fig. 2. Clearly, sparks occurred
at a very low frequency in the region occupied by T tubules.
When the extent of occupancy by T tubules was calculated as
the square root of the ratio of areas, as would befit cells of
circular perimeter, the radial extent of occupancy by T tubules
was 0.12. Regardless of the approximation used, there was a
close correspondence between the radial extent free of sparks
and that occupied by T tubules, reflecting a pattern of avoidance of T tubules by sparks.
Topographic pattern of spark generation is lost in dysgenic
myotubes. If, as shown, the presence of T tubules suppresses
the generation of Ca2⫹ sparks, a first candidate as molecular
inhibitor ought to be the DHPRs, which interact mechanically
with RyR1 channels in the T-SR junction (40). Skeletal muscle
cells from mice homozygous for the mdg mutation (47), which
lack DHPRs in the T tubules (29), were used to test this
hypothesis. Primary cultures of skeletal muscle taken from
neonatal mdg⫹/⫹ pups produced myotubes that were morphologically similar to those of the wild type, but appeared to have
a somewhat greater diameter, as described by Powell et al.
REPRESSION OF CA2⫹ SPARKS BY T TUBULES
C543
Fig. 3. Structural and functional images of dysgenic myotubes.
A: confocal xy image of fluorescence of a dysgenic mouse
myotube (mdg) stained with di-8-ANEPPS for identification of T
tubules. T tubules had similar development as in the wild-type
cells. B: line scan image of fluorescence of fluo-4 in an mdg
myotube loaded with the Ca2⫹ monitor and immersed in modified Krebs with 26 mM Ca2⫹ and 1 mM caffeine. Note Ca2⫹
sparks originating close to or at the plasmalemma.
Fig. 4. Spatial location of spontaneous sparks of mdg myotubes. Histogram of
location of 315 sparks detected in xy images from 24 mdg cells immersed in
Krebs with high Ca2⫹ and 1 mM caffeine. The bars at top represent fT averaged
in 8 cells (0.23 ⫾ 0.03).
AJP-Cell Physiol • VOL
state of the [Ca2⫹] of interest under the special conditions used
to elicit Ca2⫹ sparks. The study followed the evolution of
[Ca2⫹]cyto and [Ca2⫹]SR as the extracellular solution was
changed from Krebs to the high-[Ca2⫹] K/Ca with or without
nifedipine and/or caffeine.
Evolution of [Ca2⫹]cyto. Figure 6, A–C, shows images F11,
F22, and R (⫽F11/F22) of wild-type myotubes in Krebs solution. Figure 6, D and E, show ratio images in subsequent stages
of work with the same culture dish. The average ratio in Fig.
6C corresponds to a [Ca2⫹]cyto of 152 nM. When the solution
was changed to K/Ca with 50 ␮M nifedipine and 1 mM
caffeine, the ratio did not change (Fig. 6D). After a later
change to K/Ca without nifedipine, the ratio increased substantially, eventually reaching a value corresponding to [Ca2⫹]cyto
of 1.3 ␮M (Fig. 6E). As shown, [Ca2⫹]cyto could then be very
different in different cells. Figure 6F plots the time course of
image-averaged ratio (the right-side axis is labeled with the
corresponding [Ca2⫹] values, calculated using Eq. 2 in METHODS).
Corresponding ratio images for cultures of dysgenic cells are
illustrated in Fig. 7. [Ca2⫹]cyto in Krebs varied near 170 nM.
Upon changing the solution to 26 mM (no caffeine), [Ca2⫹]cyto
remained close to initial values. The addition of 1 mM caffeine
appeared to cause a small increase in some cells. Figure 7D
plots the time course of image-averaged ratio and [Ca2⫹]cyto.
Figure 8 summarizes results with various solutions. Wildtype and dysgenic cells immersed in Krebs had similar
[Ca2⫹]cyto, 152 and 171 nM, respectively. In Krebs with 26
mM Ca2⫹, [Ca2⫹]cyto increased significantly in the wild type,
to 464 nM, but in dysgenic cells it did not (175 nM). The
change in the wild type was essentially suppressed when
nifedipine was added to the high-Ca2⫹ Krebs (166 nM).
As reported there, sparks in high-Ca2⫹ Krebs plus caffeine
occurred at random locations in mdg cells, whereas in normal
myotubes (in the same solution plus nifedipine) events failed to
occur at peripheral locations. As shown in Fig. 8, these conditions led to a somewhat lower [Ca2⫹]cyto in mdg (118 nM vs.
192 nM in the wild type). This difference makes it impossible
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experiments using SEER to monitor [Ca2⫹], these conditions
lead to similar concentrations in the compartments of interest
in mdg and normal myotubes.
The histogram of locations of 361 sparks from 26 normal
myotubes in K/Ca⫹NC is plotted in Fig. 5B. It shows that the
pattern of avoidance of T tubules by sparks, which was lost in
mdg cells, persisted in normal cells under these conditions. The
averaged morphological parameters of 165 events detected in
xt images (see Table 1). The similarity of spark amplitudes in
mdg and wild type suggests that a rough parity of cytosolic and
intra-SR Ca2⫹ prevailed under these conditions, which proved
to be true in direct measurements described next. The result is
consistent with the interpretation that DHPRs are required to
preserve the spatial segregation of T tubules and Ca2⫹ sparks.
Measurements of cytosolic and SR-luminal [Ca2⫹]. Measurements were carried out using SEER (31) to compare the
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REPRESSION OF CA2⫹ SPARKS BY T TUBULES
that the loss of the pattern of spark generation is due to an
increased stimulation by cytosolic Ca2⫹ in the dysgenic cells.
Comparisons of SR luminal [Ca2⫹]. Figure 9 illustrates the
technique. Figure 9, A and B, are images of a cell in a culture
dish loaded with mag-indo-1 AM and then membrane-permeabilized. Figure 9C is the ratio F11/F22, restricted as described
in METHODS to areas of the cell where the dye concentration was
above the cellular mean. This restriction essentially removes
nuclei, as well as debris, from the ratio image. The image in
Table 1. Morphological parameters of sparks detected automatically in xy images
Wild type
mdg
n
Amplitude
FWHM, ␮m
FDHM, ms
Rise Time, ms
165
1,276
0.53⫾0.03
0.41⫾0.01
4.62⫾0.14
4.87⫾0.05
41.5⫾0.76
44.2⫾0.19
29.2⫾0.66
30.2⫾0.19
Values are means ⫾ SE; n, no. of events detected automatically. mdg, dysgenic mouse myotubes. Parameters, including full width and full duration at half
magnitude (FWHM and FDHM), are defined in METHODS. Sparks of mdg myotubes were recorded in Krebs solution with 26 mM Ca2⫹ and 1 mM caffeine. Sparks
of wild-type myotubes were recorded in Krebs solution with 26 mM Ca2⫹, 1 mM caffeine, and 50 ␮M nifedipine.
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Fig. 5. Spatial location of spontaneous sparks in normal myotubes in Krebs
with high Ca2⫹ and 1 mM caffeine. A: without nifedipine sparks originated at
random locations. B: no sparks originated in the region occupied by T tubules
after 50 ␮M nifedipine was administered.
Fig. 9D is of fluorescence F33 of the dye rhod-2, present in the
solution to identify permeabilized myotubes.
In principle, the permeabilization step reduces one’s ability
to follow changes in [Ca2⫹]SR upon changing extracellular
solutions because the SR contents may vary after the plasma
membrane is permeabilized. The approach illustrated in Fig. 10
was designed to evaluate the evolution of [Ca2⫹]SR after
permeabilization. Figure 10A shows fluorescence of rhod-2, in
an mdg culture dish preloaded with mag-indo-1 AM and
immersed in the permeabilizing solution (which contains 50
nM free Ca2⫹). Figure 10B is fluorescence F22 of mag-indo-1,
which is present in cytosol and organelles. A very bright cell in
Fig. 10B corresponds to a dark region of Fig. 10A (dashed
contour), marking a myotube that has not been permeabilized.
The myotube that is highly fluorescent in Fig. 10A has much
lower F22 intensity in Fig. 10B. The cellwide concentration of
mag-indo drops ⬃10 times after permeabilization, consistent
with the presence of much more dye in the cytosol than in
organelles. Figure 10C shows the ratio with a high value in the
permeabilized cell, consistent with the high [Ca2⫹] in the SR,
and a low value in the intact cell, where the dye signal is
largely determined by [Ca2⫹]cyto.
The heterogeneous staining pattern of Fig. 10, A–C, is
characteristic of initial stages of permeabilization. Although
it is not possible to know the exact time, most cells become
permeabilized in 2–3 min. The evolution of [Ca2⫹]SR can
then be followed for tens of minutes after permeabilization,
as illustrated in Fig. 10, D–F. Figure 10D is from a wildtype culture in Krebs, and Fig. 10E is from a different
wild-type dish, exposed to K/Ca for 10 min before permeabilization. The evolution of [Ca2⫹]SR after permeabilization, plotted in Fig. 10F, is slow. Hence, there is time after
permeabilization to obtain several images and derive an
average R that will represent approximately the prevailing
level of [Ca2⫹]SR in the conditions imposed before permeabilization. The plot in red is representative of the substantial increase in [Ca2⫹]SR observed in normal myotubes in
high Ca2⫹ Krebs.
Figure 11 summarizes the measurements of [Ca2⫹]SR. The
ratio measured in wild-type cells immersed in Krebs corresponds to a [Ca2⫹]SR of 82 ␮M. Immersion in K⫹/Ca2⫹ with
nifedipine increased [Ca2⫹]SR significantly, to 144 ␮M. As
expected, immersion in high [Ca2⫹]o without the dihydropyridine caused an even greater increase in [Ca2⫹]SR, to 284 ␮M.
The [Ca2⫹]SR level of mdg cells in Krebs, 206 ␮M, was
significantly greater than that of normal cells in the same
condition. Immersion of mdg cultures in K⫹/Ca2⫹ had an
unexpected result, [Ca2⫹]SR decreased. Serendipitously, this
decrease made the average value in mdg cells identical to that
of wild-type cells in K/Ca⫹N (K/Ca and K/Ca⫹N were the
REPRESSION OF CA2⫹ SPARKS BY T TUBULES
C545
conditions used for comparing location of sparks in mdg and
wild-type cells).
Location of ryanodine receptor isoforms in wild-type and
mdg myotubes The occurrence of fewer sparks in the periphery
of normal myotubes could be due to a paucity of release
channels in this area. Although the adult mammalian skeletal
muscle predominantly expresses RyR1, developing cells express RyR3 at a high level (54). Ca2⫹ sparks could originate
from either isoform (9, 53, 62), although myotubes expressing
RyR3 had a much greater frequency of spontaneous events
than those expressing RyR1 (63). To identify the expression of
these isoforms, we used antibodies against RyR1 or RyR3.
Figure 12, top, shows fluorescence images of myotubes from
normal or mdg cultures immunolabeled with anti-RyR1 or
anti-RyR3 antibodies. Confocal sections ran near the myotube
axis, as evidenced by the presence of nuclei. In normal myotubes, the density of sites reactive to anti-RyR1 antibodies was
greater near the surface in some cases (Fig. 12A) and randomly
distributed in the cytosol in others (Fig. 12B). The RyR3specific reactivity was likewise present nearly everywhere,
although in some images it was denser near nuclei (Fig. 12C).
The distribution pattern of RyR1 and RyR3 in mdg (Fig. 12,
D–F) was not visibly different from that of the wild-type cells,
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including cases, not shown, in which RyR3 was denser in
perinuclear regions. The data suggest that the virtual absence
of events at the periphery of normal myotubes was not due to
a lack of release channels of either isoform.
DISCUSSION
This study demonstrates a straightforward topographic segregation of developing muscle cells into central regions, where
Ca2⫹ sparks occur spontaneously, and peripheral regions, comprising ⬃20% of the radius, where T-tubular structures are
present and Ca2⫹ sparks are largely absent. The study also
found evidence that DHPRs in the T tubules appear to have a
crucial role in this inhibitory effect.
T tubule structures are essential for preventing Ca2⫹ sparks.
Ca2⫹ sparks of skeletal muscle result from the brief concerted
opening of clusters of release channels (23). In mammalian
muscle, few Ca2⫹ sparks were observed in fibers with intact
plasmalemma (11). To induce Ca2⫹ sparks in mammalian
muscle, the T tubule membrane had to be permeabilized by
application of saponin (27, 66). This suggests that the integrity
of the T tubule structure may be necessary to prevent spontaneous opening of release channels. Embryonic myotubes,
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Fig. 6. Imaging cytosolic [Ca2⫹] ([Ca2⫹]cyto) in myotubes. A: fluorescence image F11, excited at 351 nm and emitted in the 390- to 440-nm range. B: fluorescence
image F22, excited at 364 nm and emitted in the 465–535 nm range. C: ratio R ⫽ F11/F22 [the shifted excitation and emission ratio (SEER)]. Culture was immersed
in normal Krebs. D: ratio image, 10 min after changing to high Ca2⫹ Krebs plus nifedipine. Note no appreciable change from ratio in C. E: ratio image after
10 min in high-Ca2⫹ Krebs without nifedipine. Note large increase in the ratio. F: evolution of average ratio, with corresponding [Ca2⫹]cyto in right-side scale.
See METHODS for the definition of vertical bars. Times of acquisition of images C–E are represented by corresponding letters in the graph. Lines at top represent
solution changes: red, high Ca2⫹; green, 50 ␮M nifedipine.
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REPRESSION OF CA2⫹ SPARKS BY T TUBULES
which express Ca2⫹ release channels but have an incompletely
developed T tubular structure, were used to test this hypothesis.
With the use of a double-staining technique, Ca2⫹ sparks and
T tubule structures were imaged simultaneously. First, it was
demonstrated that the regions that gave rise to sparks were
essentially devoid of T tubules. Conversely, the regions that
did not produce sparks correspond to areas of the cell that had
T tubules. It was also shown that the absence of sparks in the
Fig. 8. The effects of extracellular [Ca2⫹] on [Ca2⫹]cyto. Means ⫾ SE over all
images obtained in solutions listed under each bar as follows: K, Krebs; K/Ca,
high Ca2⫹ Krebs; K/Ca⫹N, same plus 50 ␮M nifedipine; K/Ca⫹C, same plus
1 mM caffeine; K/Ca⫹NC, high Ca2⫹ plus nifedipine and caffeine. Individual
values correspond to average R within one image. All images were obtained
from separate fields. The number of fields included is listed inside each bar.
AJP-Cell Physiol • VOL
T tubular regions was not due to a lack of expression of RyR
isoforms, of either sort.
The present work shows that T tubules and sparks occupy
mutually exclusive spaces. As a consequence, the suppression
of sparks must be due to T tubule-associated components, or to
structures that are modified when the T tubule integrity is
affected. At the molecular level, the suppression must be due to
an interaction between release channels and junctional structures of some sort. The conclusion is consistent with the rapid
decrease in frequency of spontaneous Ca2⫹ sparks in mouse
muscle observed in the first two postnatal weeks (8), at a time
when development of T tubules is becoming complete (57). It
is also consistent with the existence of a malignant hyperthermia-inducing mutation in the III-IV loop of the DHPR in
humans (65), which presumably weakens a basal inhibitory
effect on RyR channels.
No T-SR docking is required for production of Ca2⫹ sparks.
According to Takekura et al. (57), ontogenesis of Ca2⫹ release
units requires docking of SR to embryonic plasmalemma or T
tubules, to permit local clustering of release channels and
voltage sensors. In contrast with this view, the present results
demonstrate the presence of functional sources of Ca2⫹, capable of generating sparks in regions of SR not yet docked to the
T tubule network. It follows that clusters of channels capable
of activation form at stages before T-SR docking. Therefore,
docking does not appear to be necessary for development of a
spark-capable cluster. Instead, the effect of linkage with T
tubules appears to be to put such clusters under a strong
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Fig. 7. Imaging [Ca2⫹]cyto in mdg cells. Images are
of SEER ratios. A: cells in normal Krebs. B: 10 min
after being changed to high-Ca2⫹ Krebs (solution
change represented in red; D, top). C: after addition
of 1 mM caffeine (brown line in D). D: evolution of
average ratio. Note that it remains nearly constant
throughout the solution changes. Times of acquisition of images A–C are represented by corresponding letters in the graph.
REPRESSION OF CA2⫹ SPARKS BY T TUBULES
C547
inhibitory control, which suppresses their ability to fire under
normal conditions.
While these “unmoored” regions of SR generate sparks, the
events are profoundly different from sparks observed in adult
muscle (67). Most notably, their spatial width is nearly threefold greater in myotubes, which is only possible if the activation propagates over large, extensive groups of channels.
Again, this is an indication of a susceptibility to activation that
is largely lost on docking with T tubules.
DHPRs are necessary for basal suppression of Ca2⫹ sparks.
DHPRs in the T tubule membrane function as activators, which
open release channels in the SR membrane during an action
potential through a direct interaction. Some studies have also
shown that DHPRs may function as inhibitors for the Ca2⫹
release channels, including the work of Suda (55), indicating
possible involvement of DHPRs in terminating Ca2⫹ release in
rat skeletal myotubes, blockage of Ca2⫹ release from SR
vesicles by a peptide derived from the II-III loop of the DHPR
(13), an effect of DHPR channel blockers on caffeine sensitivity of 1B5 myotubes expressing RyR1 (33) and the demonstration that the R1086H mutation in ␣1S enhances sensitivity
to activation by the voltage sensor and by caffeine (65).
To test for a role of DHPRs in situ we used mdg myotubes,
which form T-SR junctions (20) lacking DHPRs in the T tubule
membrane but are structurally normal otherwise. The location
of Ca2⫹ sparks was random in these cells, indicating that T
tubules without DHPRs no longer prevented spontaneous
opening of the release channels. The DHPR is therefore required for this inhibitory effect.
The test required changes in conditions that made the comparison less clear cut. In dysgenic cells it was necessary to
increase [Ca2⫹]o radically and apply 1 mM caffeine to even
observe sparks. Complicating matters, when wild-type cells
AJP-Cell Physiol • VOL
were observed in the high-[Ca2⫹] solution with caffeine, an
even higher frequency of sparks was induced, together with the
loss of the characteristic pattern of locations.
In an attempt to compare wild-type and mdg cells under
similar conditions of load, the normal myotubes were imaged
in high Ca2⫹, the addition of 50 ␮M nifedipine to block Ca2⫹
entry through L-type Ca2⫹ channels. The similarity in morphological parameters of Ca2⫹ sparks recorded in mdg and in
normal cells under these conditions was an indication that
rough parity in cytosolic and intra-SR Ca2⫹ levels had been
reached. The normal myotubes in higher [Ca2⫹]o still maintained the suppression pattern, consistent with a necessary role
of DHPRs in the basal inhibition of release channels.
We initially thought that the reason for the absence of sparks
in mdg cells could be a lower [Ca2⫹]cyto caused by decreased
Ca2⫹ influx due to the absence of DHPRs (the L-type Ca2⫹
channels in the plasmalemma), a lower load of Ca2⫹ in the SR
secondary to the diminished influx, or both. Indeed, when
[Ca2⫹]o was increased, the dysgenic cells generated sparks at a
higher rate, comparable with the wild-type cells in Krebs.
However, direct measurements of [Ca2⫹]cyto and [Ca2⫹]SR
did not support this explanation because [Ca2⫹]cyto was nearly
identical in wild-type and mdg cells immersed in Krebs solution. In any case, the final conditions used for comparing
patterns of spark location led to similar [Ca2⫹]cyto and
[Ca2⫹]SR in both types of cells. The [Ca2⫹] measurements
therefore support an inhibitory role of the DHPR, as they show
that the difference in patterns may not be attributed to differences in relevant [Ca2⫹] under the conditions that made the
comparison possible.
The suppression effect in embryonic cells is conserved in
adults. As first described by Shirokova et al. (53) the topographic segregation between T tubules and Ca2⫹ sparks has a
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Fig. 9. Imaging sarcoplasmic reticulum (SR) luminal
[Ca2⫹]. A and B: portions of images F11 and F22, with
excitation and emission wavelengths as described for Fig. 6,
in a culture loaded with mag-indo-1 and membrane-permeabilized with saponin. C: ratio F11/F22 in areas where dye
concentration was above the mean (other areas are colored
gray). D: fluorescence F33 of rhod-2, present in the saponin
solution to monitor permeabilization.
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REPRESSION OF CA2⫹ SPARKS BY T TUBULES
two-way functional correlate, which consists of an absence of
voltage-operated release in the regions where sparks are observed, plus a pattern of voltage-operated release (where it
exists) that is devoid of sparks. The failure of voltage sensors
in the mammal to elicit sparks was first shown in adult muscle
(52). The picture was recently completed with the finding that
Fig. 11. The effects of extracellular [Ca2⫹] on [Ca2⫹]SR. Means ⫾ SE over all
mag-indo-1 SEER images obtained in the same condition. The number of fields
included in the averages is listed. Note the near identity of levels attained in
wild-type cells in K⫹/Ca2⫹ solution with nifedipine, and in mdg cells in K/Ca.
AJP-Cell Physiol • VOL
the elementary events produced by depolarization in the rat are
“embers,” resulting from the opening of single RyR channels
(11).
Thus it appears that DHPRs and RyRs engage in a peculiar
functional relationship. It includes a basal inhibition of sparks,
plus a dynamic, voltage-dependent activation of opening accompanied by continued inhibition of sparks even when channels open. This peculiar feature is also present in myotubes, as
shown by Shirokova et al. (53). Therefore, the DHPR-RyR
interaction at work in the embryonic cells produces fully the
“adult” effect.
The fact that the inhibitory effect of the DHPR on sparks is
observed in both adult and immature mammalian muscle cells
has intriguing implications. We showed above that the inhibition of sparks associated with T-tubular structures (and presumably the DHPRs) occurs even in regions where RyR3
isoforms are present. The inhibitory effect, which could be
attributed to the mechanical interaction between DHPRs and
RyR1, somehow is capable of inhibiting the other isoform,
which is believed not to be under any sort of mechanical
interaction with voltage sensors (49). That the effect of Ttubular structures on RyR1 channels also prevents RyR3 channels from causing sparks is difficult to explain, as RyR3 are
capable of producing sparks when expressed alone in 1B5 cells
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Fig. 10. The evolution of SR luminal [Ca2⫹]. A: image F33 of rhod-2 fluorescence, 1 min after being immersed in an mdg culture dish in permeabilizing solution.
B: fluorescence F22 of mag-indo-1. Note bright cell in B, a myotube not yet permeabilized, seen as a dark region in A (dashed contour). C: ratio of images F11
(not shown) and F22. Note lower ratio in the cell that is not permeabilized. D and E: ratio images in wild-type culture dishes, permeabilized after 10 min in Krebs
(D) or K/Ca (E). F: evolution of average ratio (and [Ca2⫹]SR) vs. time after immersion in permeabilizing solution of the culture in D (black symbols) or E (red).
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REPRESSION OF CA2⫹ SPARKS BY T TUBULES
(63) or in RyR1-null mouse embryonic muscle cells (9). It
suggests an intricate interaction, such that the isoform 3 channels cannot engage in coordinated opening unless the RyR1
channels are also enabled.
DHPRs are not sufficient for basal suppression of Ca2⫹
sparks. Although the present studies point at an important role
of the DHPRs in the suppression of sparks, they also demonstrate that the suppressive effect can be overcome. This occurs
when normal myotubes are exposed to a high [Ca2⫹]o. Chun et
al. (8) demonstrated an association between “spontaneous”
Ca2⫹ sparks in myotubes and Ca2⫹ current through L-type
Ca2⫹ channels. The increased unitary current that presumably
crosses these channels in conditions of high [Ca2⫹]o appears to
be sufficient to overcome the basal inhibitory effect.
A second instance, in which the inhibitory effect is relieved
to a degree, is the chemical permeabilization of the plasmalemma by brief exposure to the mild detergent saponin. The
interaction between voltage sensors and release channels may
be altered upon exposure to saponin (32). Alternatively, Shirokova and coworkers (26) proposed that an alteration of
mitochondrial function, occurring progressively after saponin
treatment, is what determines the production of sparks and
overcomes the inhibition by T tubule structures.
Work on mechanically skinned skeletal muscle fibers supports the alternative. After the plasmalemma is peeled, T
tubules reseal, restore their transmembrane potential, and
maintain voltage-dependent skeletal type EC coupling (30).
This implies that in peeled fibers DHPRs must be in a normal
functional and structural interaction with Ca2⫹ release channels. As shown by Kirsch et al. (27) and confirmed with rat
muscle in our laboratory (B. S. Launikonis and E. Rı́os,
unpublished observations), spontaneous sparks occur in these
fibers, albeit at a lower rate.
AJP-Cell Physiol • VOL
Therefore, the presence of the DHPRs is one of the conditions necessary to prevent the occurrence of Ca2⫹ sparks in the
mammal. It should be noted that the functional state of the
DHPR is immaterial for this effect, as the suppression persists
in depolarized fibers (hence with voltage-inactivated DHPRs;
Refs. 11 and 53) or as shown here, in the presence of a high
dose of nifedipine, which drives the DHPRs to the same
inactivated state.
The studies of Murayama and Ogawa (37, 38) identified an
inhibition, described as stabilization against CICR activity,
affecting specifically ␣RyRs in the frog and RyR1 in the
mammal. This stabilization was not correlated in any detectable degree with a T-SR interaction. The same is true of the
inhibition reported by Zhou et al. (67) in muscle of both
mammals and amphibians, which they ascribed tentatively to
interactions with SR-luminal structures, perhaps calsequestrin.
While these effects, and interactions with FKBP12, calsequestrin or other molecules, may be important determinants of the
function in situ, they cannot explain the topographic segregation of T tubules and spark generation. Several layers of
control are likely to be operating in the native system, reminiscent of those described elsewhere within supramolecular
arrays of proteins (12), which endow the participating molecules with properties that can substantially diverge from, and
are more subtly controlled than, those observed for the same
molecules in isolation.
In conclusion, there are now many evidences for an inhibitory effect of T tubule-associated structures on Ca2⫹ release
channels, which leads to an effective suppression of Ca2⫹
sparks in situ. The evidence indicates that the RyR1 isoform
(or ␣) is the primary recipient of the inhibition, but the RyR3
isoform is also affected. While DHPRs are needed, they are not
sufficient for the inhibitory effect. Many perturbations of the
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Fig. 12. Immunolocalization of ryanodine receptor (RyR) isoforms in myotubes. A–C: fluorescence of fluorescein conjugates of wild-type primary myotubes labeled with the anti-RyR1 (A
and B) or anti-RyR3 antibody. D–F: corresponding images of dysgenic myotubes. Images reveal
a clustered distribution of both RyR1 and RyR3
in the entire cytosolic region of the cells.
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AJP-Cell Physiol • VOL
In adult cells, two types of structures were described, with
clearly different SEER ratios: areas of high ratio, corresponding to SR terminal cisternae, and longitudinal structures of low
ratio, identified as mitochondria (31). As shown, for example,
in Figs. 9 and 10, there is no comparable pattern in myotubes.
It is clear that nuclei do not load well with dye. But perinuclear
areas sometimes exhibit a different ratio (Figs. 9C and 10C).
Perhaps relative development of mitochondria and SR change
among different cultures. Given this variability, we have not
attempted functional and structural separation of the main
organelles in these cells.
The resting [Ca2⫹] in stores was 82 ␮M in normal myotubes. This is substantially lower than in adult frog muscle,
where the same technique yields an average of 470 ␮M for
permeabilized fibers equilibrated in a solution with 100 nM
[Ca2⫹]cyto (31). The measurement in myotubes may have been
distorted by a contribution from mitochondria. In addition, the
concentration of intra-organellar dye was two- to fourfold
lower in myotubes than in adult cells, which may have biased
the ratio toward lower values. In conclusion, the present
measurements, which indicate a major difference in SR load
between embryonic mouse and adult frog cells, may have
underestimated [Ca2⫹]SR to some extent.
The measurement, however, should provide a good comparison tool. Thus, it demonstrated an increase in organellar
[Ca2⫹], partially prevented by nifedipine, upon exposing normal myotubes to elevated [Ca2⫹]o. This result is consistent
with the observation of higher [Ca2⫹]cyto under the same
conditions (Fig. 8). The fact that the increase in [Ca2⫹]cyto took
many minutes, even in [Ca2⫹]o as high as 26 mM, is consistent
with the observation that Ca2⫹ load in stores of embryonic
muscle fibers did not change measurably after 10 min of
exposure to 8 mM [Ca2⫹]o (8).
[Ca2⫹]SR was expected to be close to normal in mdg cells, as
many structure-function studies have demonstrated (and relied
upon) the integrity of the Ca2⫹ store in these cells. Surprisingly, intra-organellar [Ca2⫹] was significantly greater in the
dysgenic cells, by about twofold. The present study is not the
first to imply greater load in dysgenic cells; Weiss et al. (65)
found that the maximal cytosolic Ca2⫹ transient in response to
caffeine was greater in uninjected than in ␣-expressing dysgenic cells, by nearly 40%. As we have shown, dysgenic cells
do not produce spontaneous Ca2⫹ sparks in Krebs, but normal
myotubes do. The difference in [Ca2⫹]SR, could be in part due
to the greater leak constituted by large and frequent release
events in normal cells at rest.
This explanation may help justify the paradoxical effect of
exposure to high [Ca2⫹]o in dysgenic cells: a decrease in
[Ca2⫹]SR. As shown previously, elevated [Ca2⫹]o does not
significantly increase [Ca2⫹]cyto in these cells, perhaps because
their only pathways of Ca2⫹ entry may be SOC or ECCE
channels (7, 44), with currents that saturate at relatively low
[Ca2⫹]o (7). High [Ca2⫹]o causes sparks to appear, providing a
leak that may shift the steady [Ca2⫹]SR to a lower value.
However, these explanations are incomplete, as they do not
provide a reason for the absence of Ca2⫹ sparks in mdg
myotubes at rest, or their appearance in high [Ca2⫹]o.
The conditions found appropriate for comparing the patterns
of spark generation in normal and dysgenic myotubes included
high [Ca2⫹]o, 1 mM caffeine, and, in normal myotubes, 50 ␮M
nifedipine. Under those conditions, free [Ca2⫹]cyto was 192 nM
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supramolecular arrangement (saponin, mechanical peeling, and
mitochondrial alteration) or an increase in the inward current
through L-type channels are capable of overcoming the functional inhibition.
Measuring [Ca2⫹]cyto with SEER. By increasing the dynamic range of ratiometric measurements, SEER makes it
possible to image [Ca2⫹] inside organelles, as initially demonstrated using mag-indo-1 (31). Here we used the higher-affinity
dye indo-1 to monitor [Ca2⫹] in the cytosol. Like other measurements with dyes loaded into the cytosol in AM form, the
present one is compromised by entry of dye into organelles. In
the APPENDIX, we introduce an approach to correct such errors,
which results in Eq. 2 relating [Ca2⫹] to the measured ratio.
Equation 2 is isomorphic with the conventional Eq. 1, but has
an effective dissociation constant, Keff, smaller than KD, and an
effective minimum ratio, R0, greater than Rmin. This correction
is liable to introduce errors, as it assumes the presence of a
constant fraction of the dye in the organelles. However, the
APPENDIX shows that the magnitude of the correction is
proportional to the combined fractional volume of SR and
mitochondria in myotubes, which is probably ⬍10% of the
total (C. Franzini-Armstrong, unpublished observations).
Therefore, even if the dye load and volume of organelles
change from cell to cell, the errors in Eq. 2 will probably
stay small.
The normal myotubes studied here had on average a
[Ca2⫹]cyto of 152 nM. This value is in good agreement with
results obtained in normal myotubes by Shimahara et al. (50)
using fura-2, and those measured by Pérez et al. (45) in 1B5
myotubes expressing RyR1 in a careful study with Ca2⫹sensitive microelectrodes, but greater than values of Avila et al.
(1) using a null-point calibration approach in normal myotubes
with indo-1. Considering the many sources of error, the agreement among all these approaches appears reasonable.
[Ca2⫹]cyto measured in mdg myotubes was on average 172
nM, which is very close to the value in wild-type cells. Similar
concentrations in these two types of cells were also reported by
Shimahara et al. (50). While we know of no other comparisons
between normal and dysgenic myotubes, Weiss et al. (65)
reported that uninjected and DHPR ␣1s-expressing mdg cells
had the same resting indo-1 fluorescence ratios. In all, different
approaches lead to the conclusion that lack of the L-type Ca2⫹
channel/voltage sensor per se does not determine a major
change in resting cytosolic [Ca2⫹].
Cytosolic [Ca2⫹] increased significantly in normal myotubes
exposed to high [Ca2⫹]o. The increase was due in large part to
flux through a DHP-sensitive pathway, as it was nearly abolished by 50 ␮M nifedipine. In dysgenic cultures however, the
exposure to high [Ca2⫹]o did not lead to higher [Ca2⫹]cyto. One
may conclude from these observations that exposure to a high
[Ca2⫹]o increases [Ca2⫹]cyto through an increase in basal entry
of Ca2⫹ through L-type Ca2⫹ channels open at rest.
Imaging [Ca2⫹] in stores of normal and dysgenic myotubes.
The present study includes for the first time confocal ratiometric images of a [Ca2⫹]-sensitive dye in stores of myotubes.
Unlike the images of indo-1 in the cytosol, those of mag-indo-1
in organelles reveal structural detail. As the ratiometric measurement is independent, to a good degree, of local dye
concentration and volume of the stained organelle, the local
variations in the ratio reflect changes in organellar [Ca2⫹].
REPRESSION OF CA2⫹ SPARKS BY T TUBULES
in the wild-type cells and 118 nM in the dysgenic cells (Fig. 8).
In similar solutions, organellar [Ca2⫹] was 144 ␮M in both
types of cells (Fig. 11). Given these results, the characteristic
spatial location of Ca2⫹ sparks in the wild type is not likely to
be due to differences in modulation by Ca2⫹. It may reflect
instead a direct inhibitory effect of the DHPR, missing in
dysgenic cells.
APPENDIX
共1 ⫺ v兲共f11,Ca关Ca2⫹兴 ⫹ f11KD兲
F 11 ⫽ vf11,Ca ⫹
关Ca2⫹兴 ⫹ KD
(3)
where f11 and f11,Ca are, respectively, the fluorescence per unit
concentration of free and Ca2⫹-bound dye. An analogous expression
applies for F22. Solving for [Ca2⫹] the definition equation R ⫽
F11/F22, it follows that
关Ca2⫹兴 ⫽ KD␤共1 ⫺ v兲
共␤共1 ⫺ v兲 ⫹ v兲R ⫺ ␤共1 ⫺ v兲Rmin ⫺ vRmax
共␤共1 ⫺ v兲 ⫹ v兲共Rmax ⫺ R兲
(4)
where ␤ is f22/f22,Ca, Rmax is f11.Ca/f22,Ca, and Rmin is f11/f22.
This is the equation of Grynkiewycz et al. (25) (Eq. 1 in METHODS),
modified to correct for the interference of a dye in organelles. It can
be shown that Eqs. 4 and 1 are isomorphic, by defining
K eff ⫽ KD共1 ⫺ v兲
(5)
␤共1 ⫺ v兲Rmin ⫹ vRmax
␤共1 ⫺ v兲 ⫹ v
(6)
and
R0 ⫽
Equation 2, repeated here, is obtained substituting these definitions in
Eq. 4:
关Ca 2⫹兴 ⫽ Keff␤
R ⫺ R0
Rmax ⫺ R
(7)
The interference from dye in organelles therefore results in a change
in KD to a value smaller than KD, and an effective minimum ratio, to
a value greater than Rmin [R0 is an average of Rmin and Rmax with
weights ␤ (1 ⫺ v) and v]. The modified parameters, determined in an
on-cell calibration, gave Keff ⫽ 394 nM and R0 ⫽ 0.51.
The limitations of this correction should be obvious: it assumes the
presence of a constant fraction of the dye in the organelles, and it
assumes, perhaps more safely given the high affinity of indo-1, that
[Ca2⫹] there is sufficient to saturate the dye. While the first assumption is unlikely to hold as more than a rough approximation, the
correction is small. The combined volume of SR and mitochondria in
myotubes is probably ⬍10% of the total (C. Franzini-Armstrong,
AJP-Cell Physiol • VOL
unpublished observations). If v is 0.1, then Keff will be 0.9 KD and R0
will be very close to Rmin (⬃30 times closer to Rmin than Rmax,
according to Eq. 6). Therefore, even if the dye load and volume of
organelles change from cell to cell, the errors in Eq. 7 (see Eq. 2) will
probably remain low.
ACKNOWLEDGMENTS
We are grateful to V. Sorrentino (University of Siena) for the generous gift
of antibodies against RyR1 and RyR3, to Y. Chu (Rush University) for critical
help with immunostaining, to Clara Franzini-Armstrong (University of Pennsylvania) for unpublished information on the structure of mouse myotubes, and
to G. Brum (Univ. de la República, Montevideo), T. Shannon (Rush University) and L. Csernoch (Medical University of Debrecen) for valuable suggestions on this work.
Present address for A. González: Fundación Instituto de Estudios Avanzados, Caracas, Venezuela.
GRANTS
This study was funded by National Institute of Arthritis and Musculoskeletal and Skin Diseases Grants AR-049184 and AR-032808 (to E. Rı́os). At the
time this study was performed, J. Zhou was a Merrick Scholar of Rush
University and B. S. Launikonis was a C. J. Martin Fellow of the National
Health and Medical Research Council of Australia.
REFERENCES
1. Avila G, O’Brien JJ, and Dirksen RT. Excitation-contraction uncoupling by a human central core disease mutation in the ryanodine receptor.
Proc Natl Acad Sci USA 98: 4215– 4220, 2001.
2. Baylor SM, Chandler WK, and Marshall MW. Sarcoplasmic reticulum
calcium release in frog skeletal muscle fibres estimated from Arsenazo III
calcium transients. J Physiol 344: 625– 666, 1983.
3. Block BA, Imagawa T, Campbell KP, and Franzini-Armstrong C.
Structural evidence for direct interaction between the molecular components of the transverse tubule/sarcoplasmic reticulum junction in skeletal
muscle. J Cell Biol 107: 2587–2600, 1988.
4. Brum G, González A, Rengifo J, Shirokova N, and Rı́os E. Fast
imaging in two dimensions resolves extensive sources of Ca2⫹ sparks in
frog skeletal muscle. J Physiol 528: 419 – 433, 2000.
5. Chaudhari N. A single nucleotide deletion in the skeletal muscle-specific
calcium channel transcript of muscular dysgenesis (mdg) mice. J Biol
Chem 267: 25636 –25639, 1992.
6. Cheng H, Lederer WJ, and Cannell MB. Calcium sparks: elementary
events underlying excitation-contraction coupling in heart muscle. Science
262: 740 –744, 1993.
7. Cherednichenko G, Hurne AM, Fessenden JD, Lee EH, Allen PD,
Beam KG, and Pessah IN. Conformational activation of Ca2⫹ entry by
depolarization of skeletal myotubes. Proc Natl Acad Sci USA 101: 15793–
15798, 2004.
8. Chun LG, Ward CW, and Schneider MF. Ca2⫹ sparks are initiated by
Ca2⫹ entry in embryonic mouse skeletal muscle and decrease in frequency
postnatally. Am J Physiol Cell Physiol 285: C686 –C697, 2003.
9. Conklin MW, Ahern CA, Vallejo P, Sorrentino V, Takeshima H, and
Coronado R. Comparison of Ca2⫹ sparks produced independently by two
ryanodine receptor isoforms (type 1 or type 3). Biophys J 78: 1777–1785,
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: 1394 –1403, 1999.
11. Csernoch L, Zhou J, Stern MD, Brum G, and Rı́os E. The elementary
events of Ca2⫹ release elicited by membrane depolarization in mammalian
muscle. J Physiol 557: 43–58, 2004.
12. Davare MA, Avdonin V, Hall DD, Peden EM, Burette A, Weinberg
RJ, Horne MC, Hoshi T, and Hell JW. A ␤2 adrenergic receptor
signaling complex assembled with the Ca2⫹ channel Cav1.2. Science 293:
98 –101, 2001.
13. El-Hayek R, Antoniu B, Wang J, Hamilton SL, and Ikemoto N.
Identification of calcium release-triggering and blocking regions of the
II-III loop of the skeletal muscle dihydropyridine receptor. J Biol Chem
270: 22116 –22118, 1995.
14. Endo M, Tanaka M, and Ogawa Y. Calcium induced release of calcium
from the sarcoplasmic reticulum of skinned skeletal muscle fibres. Nature
228: 34 –36, 1970.
290 • FEBRUARY 2006 •
www.ajpcell.org
Downloaded from http://ajpcell.physiology.org/ by 10.220.32.247 on April 12, 2017
Ratiometric measurements of [Ca2⫹]cyto corrected for dye inside
organelles. Like other ratiometric measurements with dyes loaded
into cells in AM form, the SEER measurement introduced in the text
is affected by errors due to entry of dye into organelles. The following
is an approach to correct for such errors.
Organelles occupy a certain fraction of the cell volume, and load
dye, free or bound, at concentrations that may or may not be equal to
that in the cytosol. Let v represent the fraction of the dye load present
in organelles (equal to the fractional volume where the organellar dye
would partition, if its concentration was the same in organelles and
cytosol). Assuming that luminal [Ca2⫹] is sufficient to saturate the
dye, fluorescence F11 will satisfy
C551
C552
REPRESSION OF CA2⫹ SPARKS BY T TUBULES
AJP-Cell Physiol • VOL
38. Murayama T and Ogawa Y. RyR1 exhibits lower gain of CICR activity
than RyR3 in the SR: evidence for selective stabilization of RyR1 channel.
Am J Physiol Cell Physiol 287: C36 –C45, 2004.
39. Nabhani T, Zhu X, Simeoni I, Sorrentino V, Valdivia HH, and Garcia
J. Imperatoxin a enhances Ca2⫹ release in developing skeletal muscle
containing ryanodine receptor type 3. Biophys J 82: 1319 –1328, 2002.
40. Nakai J, Dirksen RT, Nguyen HT, Pessah IN, Beam KG, and Allen
PD. Enhanced dihydropyridine receptor channel activity in the presence of
ryanodine receptor. Nature 380: 72–75, 1996.
41. Neuhaus R, Rosenthal R, and Lüttgau HC. The effects of dihydropyridine derivatives on force and Ca2⫹ current in frog skeletal muscle fibres.
J Physiol 427: 187–209, 1990.
42. Ogawa Y, Kurebayashi N, and Murayama T. Ryanodine receptor
isoforms in excitation-contraction coupling. Adv Biophys 36: 27– 64, 1999.
43. Pape PC, Jong DS, and Chandler WK. Calcium release and its voltage
dependence in frog cut muscle fibers equilibrated with 20 mM EGTA.
J Gen Physiol 106: 259 –336, 1995.
44. Parekh AB and Putney JW. Store-operated calcium channels. Physiol
Rev 85:757– 810, 2005.
45. Pérez CF, López JR, and Allen PD. Expression levels of RyR1 and
RyR3 control resting free Ca2⫹ in skeletal muscle. Am J Physiol Cell
Physiol 288: C640 –C649, 2005.
46. Pizarro G and Rı́os E. How source content determines intracellular Ca2⫹
release kinetics. Simultaneous measurement of [Ca2⫹] transients and [H⫹]
displacement in skeletal muscle. J Gen Physiol 124: 239 –258, 2004.
47. Powell JA and Fambrough DM. Electrical properties of normal and
dysgenic mouse skeletal muscle in culture. J Cell Physiol 82: 21–38, 1973.
48. Powell JA, Petherbridge L, and Flücher BE. Formation of triads
without the dihydropyridine receptor alpha subunits in cell lines from
dysgenic skeletal muscle. J Cell Biol 134: 375–387, 1996.
49. 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:
2494 –2508, 2000.
50. Shimahara T, Bournaud R, Inoue I, and Strube C. Charge movement
and Ca2⫹ release in normal and dysgenic foetal myotubes. J Physiol 86:
117–121, 1992.
51. Shirokova N, Garcı́a J, Pizarro G, and Rı́os E. Ca2⫹ release from the
sarcoplasmic reticulum compared in amphibian and mammalian skeletal
muscle. J Gen Physiol 107: 1–18, 1996.
52. Shirokova N, Garcı́a J, and Rı́os E. Local calcium release in mammalian
skeletal muscle. J Physiol 512: 377–384, 1998.
53. Shirokova N, Shirokov R, Rossi D, González A, Kirsch WG, Garcı́a J,
Sorrentino V, and Rı́os E. Spatially segregated control of Ca2⫹ release in
developing skeletal muscle of mice. J Physiol 521: 483– 495, 1999.
54. Sorrentino V and Reggiani C. Expression of the ryanodine receptor type
3 in skeletal muscle. A new partner in excitation-contraction coupling?
Trends Cardiovasc Med 9: 54 – 61, 1999.
55. Suda N. Involvement of dihydropyridine receptors in terminating Ca2⫹
release in rat skeletal myotubes. J Physiol 486: 105–112, 1995.
56. Sutko JL and Airey JA. Ryanodine receptor Ca2⫹ release channels: does
diversity in form equal diversity in function? Physiol Rev 76: 1027–1071,
1996.
57. Takekura H, Flücher BE, and Franzini-Armstrong C. Sequential
docking, molecular differentiation, and positioning of T tubule/SR junctions in developing mouse skeletal muscle. Dev Biol 239: 204 –214, 2001.
58. 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: 439 – 445, 1989.
59. Tanabe T, Beam KG, Powell JA, and Numa S. Restoration of excitation-contraction coupling and slow calcium current in dysgenic muscle by
dihydropyridine receptor complementary DNA. Nature 336: 134 –149,
1988.
60. Terentyev D, Viatchenko-Karpinski S, Valdivia HH, Escobar AL, and
Györke S. Luminal Ca controls termination and refractory behaviour of
Ca-induced Ca release in cardiac myocytes. Circ Res 91: 414 – 420, 2002.
61. Tsugorka A, Rı́os E, and Blatter LA. Imaging elementary events of
calcium release in skeletal muscle cells. Science 269: 1723–1726, 1995.
62. Ward CW, Protasi F, Castillo D, Wang Y, Chen SR, 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: 3216 –3230, 2001.
290 • FEBRUARY 2006 •
www.ajpcell.org
Downloaded from http://ajpcell.physiology.org/ by 10.220.32.247 on April 12, 2017
15. Endo M. Conditions required for calcium-induced release of calcium from
the sacoplasmic reticulum. Proc Jpn Acad 51: 467– 472, 1975.
16. Endo M. Calcium release from the sarcoplasmic reticulum. Physiol Rev
57: 71–108, 1977.
17. 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: 1695–1700, 2002.
18. Fessenden JD, Wang Y, Moore RA, Chen SR, Allen PD, and Pessah
IN. Divergent functional properties of ryanodine receptor types 1 and 3
expressed in a myogenic cell line. Biophys J 79: 2509 –2525, 2000.
19. Flücher 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: 621– 630, 1999.
20. Flücher BE, Phillips JL, Powell JA, Andrews SB, and Daniels MP.
Coordinated development of myofibrils, sarcoplasmic reticulum and transverse tubules in normal and dysgenic mouse skeletal muscle, in vivo and
in vitro. Dev Biol 150: 266 –280, 1992.
21. Ford LE and Podolsky RJ. Regenerative calcium release within muscle
cells. Science 167: 58 –59, 1970.
22. Giannini G, Conti A, Mammarella S, Scrobogna M, Sorrentino V. The
ryanodine receptor/calcium channel genes are widely and differentially
expressed in murine brain and peripheral tissues. J Cell Biol 128: 893–904,
1995.
23. González A, Kirsch WG, Shirokova N, Pizarro G, Brum G, Pessah IN,
Stern MD, Cheng H, and Rı́os E. Involvement of multiple intracellular
release channels in calcium sparks of skeletal muscle. Proc Natl Acad Sci
USA 97: 4380 – 4385, 2000.
24. González A, Kirsch WG, Shirokova N, Pizarro G, Stern MD, and Rı́os
E. The spark and its ember: separately gated local components of Ca2⫹
release in skeletal muscle. J Gen Physiol 115: 139 –158, 2000b.
25. Grynkiewicz G, Poenie M, and Tsien RY. A new generation of Ca2⫹
indicators with greatly improved fluorescence properties. J Biol Chem
260: 3440 –3450, 1985.
26. Isaeva EV, Shkry VM, and Shirokova N. Mitochondrial redox state and
Ca2⫹ sparks in permeabilized mammalian skeletal muscle. J Physiol 565:
855– 872, 2005.
27. Kirsch WG, Uttenweiler D, and Fink RH. Spark- and ember-like
elementary Ca2⫹ release events in skinned fibres of adult mammalian
skeletal muscle. J Physiol 537: 379 –389, 2001.
28. Klein MG, Cheng H, Santana LF, Jiang YH, Lederer WJ, and
Schneider MF. Two mechanisms of quantized calcium release in skeletal
muscle. Nature 379: 455– 458, 1996.
29. Knudson CM, Chaudhari N, Sharp AH, Powell JA, Beam KG, and
Campbell KP. Specific absence of the ␣1 subunit of the dihydropyridine
receptor in mice with muscular dysgenesis. J Biol Chem 264: 1345–1348,
1989.
30. Lamb GD and Stephenson DG. Effects of intracellular pH and [Mg2⫹]
on excitation-contraction coupling in skeletal muscle fibres of the rat.
J Physiol 478: 331–339, 1994.
31. Launikonis BS, Zhou J, Royer L, Shannon TR, Brum G, and Rı́os E.
Confocal imaging of [Ca2⫹] in cellular organelles by SEER, shifted
excitation and emission ratioing of fluorescence. J Physiol 567: 523–543,
2005.
32. Launikonis BS and Stephenson DG. Effect of saponin treatment on the
sarcoplasmic reticulum of rat, cane toad and crustacean (yabby) skeletal
muscle. J Physiol 504: 425– 437, 1997.
33. Lee EH, Lopez JR, Li J, Protasi F, Pessah IN, Kim do H, and Allen
PD. Conformational coupling of DHPR and RyR1 in skeletal myotubes is
influenced by long-range allosterism: evidence for a negative regulatory
module. Am J Physiol Cell Physiol 286: C179 –C189, 2004.
34. Marks AR, Tempst P, Hwang KS, Taubman MB, Inui M, Chadwick
C, Fleisher 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:
8683– 8687, 1989.
35. Meissner G. Ryanodine receptor/Ca2⫹ release channels and their regulation by endogenous effectors. Annu Rev Physiol 56: 485–508, 1994.
36. Melzer W, Rı́os E, and Schneider MF. Time course of calcium release
and removal in skeletal muscle fibers. Biophys J 45: 637– 641, 1984.
37. Murayama T and Ogawa Y. Selectively suppressed Ca2⫹-induced Ca2⫹
release activity of ␣-ryanodine receptor (␣-RyR) in frog skeletal muscle
sarcoplasmic reticulum: potential distinct modes in Ca2⫹ release between
␣- and ␤-RyR. J Biol Chem 276: 2953–2960, 2001.
REPRESSION OF CA2⫹ SPARKS BY T TUBULES
63. Ward CW, Schneider MF, Castillo D, Protasi F, Wang Y, Chen SR,
and Allen PD. Expression of ryanodine receptor RyR3 produces Ca2⫹
sparks in dyspedic myotubes. J Physiol 525: 91–103, 2000.
64. Weigl LG, Hohenegger M, and Kress HG. Dihydropyridine-induced
Ca2⫹ release from ryanodine-sensitive Ca2⫹ pools in human skeletal
muscle cells. J Physiol 525: 461– 469, 2000.
65. Weiss RG, O’Connell KM, Flücher BE, Allen PD, Grabner M, and
Dirksen RT. Functional analysis of the R1086H malignant hyperthermia
mutation in the DHPR reveals an unexpected influence of the III-IV loop
C553
on skeletal muscle EC coupling. Am J Physiol Cell Physiol 287: C1094 –
C1102, 2004.
66. Zhou J, Brum G, González A, Launikonis BS, Stern MD, and Rı́os E.
Ca2⫹ sparks and embers of mammalian muscle. Properties of the sources.
J Gen Physiol 122: 95–114, 2003.
67. Zhou J, Launikonis BS, Rı́os E, and Brum G. Regulation of Ca2⫹
sparks by Ca2⫹ and Mg2⫹ in mammalian and amphibian muscle. An RyR
isoform-specific role in excitation-contraction coupling? J Gen Physiol
124: 409 – 428, 2004.
Downloaded from http://ajpcell.physiology.org/ by 10.220.32.247 on April 12, 2017
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