From the Department of Pharmacology and Physiology, University of Rochester School of Medicine and Dentistry, Rochester, New York 14642
Received for publication, October 23, 2000
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ABSTRACT |
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Skeletal muscle obtained from mice that lack the
type 1 ryanodine receptor (RyR-1), termed dyspedic mice, exhibit
a 2-fold reduction in the number of dihydropyridine binding sites
(DHPRs) compared with skeletal muscle obtained from wild-type mice
(Buck, E. D., Nguyen, H. T., Pessah, I. N., and Allen,
P. D. (1997) J. Biol. Chem. 272, 7360-7367 and
Fleig, A., Takeshima, H., and Penner, R. (1996) J. Physiol. (Lond.) 496, 339-345). To probe the role of
RyR-1 in influencing L-type Ca2+ channel (L-channel)
expression, we have monitored functional L-channel expression in the
sarcolemma using the whole-cell patch clamp technique in normal,
dyspedic, and RyR-1-expressing dyspedic myotubes. Our results indicate
that dyspedic myotubes exhibit a 45% reduction in maximum
immobilization-resistant charge movement (Qmax) and a 90%
reduction in peak Ca2+ current density. Calcium current
density was significantly increased in dyspedic myotubes 3 days after
injection of cDNA encoding either wild-type RyR-1 or E4032A, a
mutant RyR-1 that is unable to restore robust voltage-activated release
of Ca2+ from the sarcoplasmic reticulum (SR) following
expression in dyspedic myotubes (O'Brien, J. J., Allen, P. D., Beam,
K., and Chen, S. R. W. (1999) Biophys. J. 76, A302
(abstr.)). The increase in L-current density 3 days after expression of
either RyR-1 or E4032A occurred in the absence of a change in
Qmax. However, Qmax was increased 85% 6 days
after injection of dyspedic myotubes with cDNA encoding the
wild-type RyR-1 but not E4032A. Because normal and dyspedic myotubes
exhibited a similar density of T-type Ca2+ current
(T-current), the presence of RyR-1 does not appear to cause a general
overall increase in protein synthesis. Thus, long-term expression of L-channels in skeletal myotubes is promoted by
Ca2+ released through RyRs occurring either spontaneously
or during excitation-contraction coupling.
The skeletal muscle dihydropyridine receptor
(DHPR)1 functions both as a
slowly activating L-type Ca2+ channel (L-channel) and as a
voltage sensor that controls the activity of the type 1 ryanodine
receptor (RyR-1) present in the sarcoplasmic reticulum (SR). During
excitation-contraction (EC) coupling, sarcolemmal depolarization
(e.g. an action potential) induces voltage-driven
conformational changes in the DHPR, which can be measured
electrophysiologically as nonlinear capacitative currents, termed
intramembrane charge movements or gating currents. These charge
movements are thought to mechanically activate RyR-1 proteins during EC
coupling and thus lead to a massive release of SR calcium (orthograde
signal of EC coupling; see Ref. 1 for review).
Analysis of skeletal myotubes derived from RyR-1-knockout (dyspedic)
mice has revealed that in addition to the orthograde signal of EC
coupling (signal transmitted from the DHPR to the RyR-1), there is also
a retrograde signal whereby RyR-1 promotes the calcium conducting
activity of the skeletal L-channel (2, 3). This conclusion was inferred
from the observation that despite a significant surface density of
DHPRs, dyspedic myotubes exhibit a marked (~90%) reduction in
L-current. Moreover, short-term (2-4 days) expression of RyR-1 in
dyspedic myotubes considerably enhances L-current density in the
absence of a change in intramembrane charge movement (2, 3). These
observations indicate that RyR-1 promotes the L-channel activity of the
skeletal muscle DHPR in a manner that is independent of L-channel
expression (retrograde signal of EC coupling).
Dyspedic muscle exhibits a 25-50% reduction in total DHP binding (4,
5). Accordingly, we have reported that dyspedic myotubes possess a
significant reduction in maximal intramembrane charge movement compared
with normal myotubes (3). This apparent reduction in the number of
functional DHPRs in the sarcolemma cannot completely account for the
~90% decrease in L-current density found in dyspedic myotubes. In
fact, dyspedic myotubes exhibit a nearly 5-fold reduction in the
current-to-charge and conductance-to-charge (Gmax/Qmax) ratios, compared with both normal
and RyR-1-expressing dyspedic myotubes (3). These observations support
the conclusion of Nakai et al. (2) that RyR-1 promotes the
Ca2+ conducting activity of the skeletal muscle L-channel.
However, the mechanism(s) underlying the different DHPR expression
levels in normal and dyspedic muscle have yet to be investigated.
Considering the functional effects of reintroduction of RyR-1 in
dyspedic myotubes (i.e. restoration of both retrograde and
orthograde signals of EC coupling), either Ca2+ influx (via
L-channels) and/or SR Ca2+ release (via RyR-1) could play a
critical role in the regulation of sarcolemmal DHPR expression.
Despite the central role that the DHPR plays in skeletal EC
coupling, controversy exists with regard to the mechanisms that control
the expression of L-channels and voltage sensors. In frog skeletal
muscle, long-term blockade of skeletal L-channels increases the number
of sarcolemmal L-channels and voltage sensors, suggesting that
Ca2+ influx (via L-channels) inhibits DHPR expression (6).
However, elevations in extracellular Ca2+ increase
L-current density and intramembrane charge movement in cultured rat
myoballs (7). In the present study, we have investigated the roles of
Ca2+ influx and release on the regulation of DHPR
expression (L-channel activity and charge movement), following
expression in dyspedic myotubes of either wild-type RyR-1 or a mutant
RyR-1 (E4032A) that preferentially restores the retrograde
(i.e. L-current) signal of skeletal muscle EC coupling (8).
Using electrophysiological criteria (Ca2+ current and
charge movement magnitudes) as a measure of functional DHPR activity in
the sarcolemma, our results demonstrate that functional DHPR expression
is promoted by Ca2+ released through RyR-1 proteins and not
via Ca2+ influx through sarcolemmal L-channels.
Preparation of Myotubes--
Myotubes were prepared from primary
culture of normal and dyspedic muscle as described previously (2).
Expression of wild-type RyR-1 and the E4032A point mutation in RyR-1 in
dyspedic myotubes was achieved by nuclear microinjection (9) of the
appropriate expression plasmid (0.5 µg/µl) 6-8 days after initial
plating of myoblasts. The E4032A mutation in RyR-1 was constructed
using a standard two-step site-directed mutagenesis strategy (10). The
entire polymerase chain reaction-modified cDNA portion was ultimately confirmed by sequence analysis. Expressing myotubes were
examined electrophysiologically 2-6 days following nuclear microinjection. In some experiments (Figs. 1, 2, and 5), expressing myotubes were identified by the development of green fluorescence 2-6
days after coinjection with a mixture of either RyR-1 or E4032A cDNAs (0.5 µg/µl) and a cDNA expression plasmid encoding an
enhanced green fluorescence protein (0.1 µg/µl). Coinjection of GFP
cDNA was omitted when cells were to be used for measurements of
intracellular Ca2+ transients (Fig. 4). For these
experiments, expression was established by the presence of either
electrically evoked (8.0 V, 10-30 ms) contractile activity (RyR-1) or
large, slowly activating L-type Ca2+ currents (E4032A).
Measurements of Ionic and Gating Currents--
The whole-cell
variant of the patch clamp technique (11) was used to measure ionic and
gating currents in both normal and dyspedic myotubes, as described
previously (3). Inward L-currents were elicited by 200 ms test pulses
of variable amplitude from a holding potential (HP) of
Immobilization-resistant intramembrane charge movements were measured
following the prepulse protocol and blockade of ionic Ca2+
currents by the addition of 0.5 mM Cd2+ + 0.2 mM La3+ to the extracellular recording solution
(12). The amount of immobilization-resistant charge movement was
estimated by integrating the transient of charge that moved outward
after the onset of the test pulse (Qon) and subsequently
normalized to cell capacitance (nC/µF). The magnitude of the maximum
immobilization-resistant charge movement (Qmax) was
estimated by fitting the Qon data according to Equation 2,
Measurements of Intracellular Ca2+
Transients--
Changes in intracellular Ca2+ were
recorded with Fluo-3 as described previously (13, 10). Briefly, the
salt form of the dye was added to the internal recording solution (see
below). After rupture of the cell membrane and entry into the
whole-cell mode, a waiting period of ~5 min was used to allow the dye
to diffuse into the cell interior. A 75 watt xenon bulb and high-speed DeltaRAM illuminator (Photon Technology Incorporated, Monmouth Junction, NJ) were used to excite the dye (480 ± 20 nm) present in a small rectangular region of the voltage-clamped myotube. A
computer-controlled shutter was used to eliminate illumination during
intervals between test pulses. Fluorescence emission was measured using
a dichroic long-pass mirror centered at 505 nm, an emission filter
centered at 535 ± 25 nm, and a photomultiplier detection system
operating in analogue mode (analogue filter set at 0.5 ms) (Photon
Technology Incorporated). The background fluorescence was measured and
canceled by analogous subtraction. Fluorescence traces are expressed as
Recording Solutions--
Ionic and gating currents were recorded
using a pipette solution containing (in mM): Cs-Aspartate
(140), MgCl2 (5), Cs2EGTA (10), and HEPES (10),
pH 7.4. The external solution contained (in mM):
triethylammonium chloride (145), CaCl2 (10),
tetrodotoxin (0.003), and HEPES (10), pH 7.4. For measurements
of intracellular Ca2+ transients, the pipette solution
contained (in mM): Cs-aspartate (145), CsCl (10),
Cs2EGTA (0.1), MgCl2 (1.2), MgATP (5), K5Fluo-3 (0.2), HEPES (10), pH 7.4. The external calcium
current recording solution was supplemented with 0.5 mM
CdCl2 + 0.2 mM LaCl3 for
measurements of intramembrane charge movement.
We have previously shown that compared with normal myotubes, the
maximum immobilization-resistant intramembrane charge movement (Qmax) is ~40% smaller in dyspedic myotubes (3).
Moreover, Qmax remains unchanged even 2-4 days following
reintroduction of RyR-1 into dyspedic myotubes (2, 3). We have tested
whether long-term expression of RyR-1 in dyspedic myotubes restores
Qmax to a value comparable with that of normal myotubes.
Fig. 1 compares immobilization-resistant
charge movements in dyspedic myotubes either 3 or 6 days after nuclear
injection of RyR-1 cDNA (3d-RyR-1 and 6d-RyR-1, respectively). In
agreement with previous reports, 3d-RyR-1 expressing dyspedic myotubes
exhibited Qmax values similar to those of uninjected
dyspedic myotubes (Fig. 1C and Table
I). However, charge movements recorded
from 6d-RyR-1-expressing dyspedic myotubes were significantly larger
than those recorded from either uninjected dyspedic myotubes or
3d-RyR-1-expressing myotubes. On average, Qmax increased
85% 6 days after nuclear microinjection of dyspedic myotubes with
RyR-1 cDNA (Fig. 1 and Table I). Fig. 1C shows the
entire time course of the RyR-1-mediated increase in Qmax.
A progressive increase in Qmax was found each day between 3 and 6 days following RyR-1 cDNA injection (gray bars).
The protracted time course of this effect is consistent with the
initiation of a slow process, such as activation of gene transcription
and subsequent protein synthesis, rather than a simple redistribution
of previously translated but sequestered voltage sensors. By contrast,
Qmax values recorded from uninjected dyspedic myotubes
during a parallel time did not exhibit a significant change in
Qmax (white bars). Thus, the increase in DHPR
expression (Qmax) requires long-term reintroduction of
RyR-1 and therefore does not arise from a general increase in L-channel
expression during prolonged culture. Interestingly, the
6d-RyR-1-expressing dyspedic myotubes exhibit Qmax values that were nearly identical to those obtained from normal myotubes (Table I; see also Ref. 3). Thus, the long-term presence of RyR-1
strongly influences the number of sarcolemmal DHPRs in skeletal myotubes.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
80 mV,
following a prepulse protocol (12, 10) used to inactivate T-type
Ca2+ channels (1 s to
20 mV followed by 50 ms to
50
mV). In some experiments, calcium currents were elicited in the absence
of the prepulse protocol to investigate the T-channels activity (see Fig. 3). Peak L-currents were normalized to cell capacitance (pA/pF), plotted as a function of membrane potential (I-V curves), and fitted
according to Equation 1,
where Vrev is the extrapolated reversal potential of
the L-current, Vm is the membrane potential during the test pulse, Gmax is the maximal L-channel conductance,
VG 1/2 is the voltage for half-activation of
Gmax, and kG is a slope factor.
(Eq. 1)
where Vm, VQ 1/2, and
kQ have their usual meanings with regard to
charge movement.
(Eq. 2)
F/F, where F represents the baseline fluorescence immediately prior
to depolarization, and
F represents the fluorescence change
from baseline. Fluorescence amplitudes at the end of the test pulses
were plotted as a function of the membrane potential and fitted
according to Equation 3,
where
(Eq. 3)
F/Fmax is the calculated maximal
fluorescence change, VF 1/2 is the midpoint
potential, and kF is the slope factor.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
Immobilization-resistant charge movement is
increased in dyspedic myotubes following long-term expression of the
skeletal muscle ryanodine receptor (RyR-1). A,
immobilization-resistant charge movements recorded from two
RyR-1-expressing dyspedic myotubes. Dyspedic myotubes were injected
with RyR-1 cDNA either 3 days (3d-RyR-1; empty
squares, left) or 6 days (6d-RyR-1; filled
squares, right) before electrophysiological recordings.
B, voltage dependence of charge movements recorded as in
A. The charge movement occurring during the test pulses
(Qon) was integrated and plotted as a function of the
membrane potential (Vm). Data were obtained from twenty 3-day
and eighteen 6-day RyR-1-expressing dyspedic myotubes. The average
values (± S.E.) for the parameters obtained by fitting each myotube
within a group separately to Eq. 2 are given in Table I (Q-V). The
solid lines were generated using Eq. 2 and the corresponding Q-V
parameters given in Table I. C, time-dependent
effect of RyR-1 on the magnitude of the maximum
immobilization-resistant charge movement (Qmax). The
Qmax values were estimated from dyspedic myotubes that were
previously (3-6 days) injected with RyR-1 cDNA (gray
bars). Qmax did not vary significantly
(p > 0.1) for non-injected dyspedic myotubes
investigated during the same period of time (white bars).
Data were obtained from a minimum of six (dyspedic, 5 days) and a
maximum of 18 (RyR-1, 6 days) myotubes for each condition.
Parameters of fitted I-V and Q-V curves
The ability of DHPRs to function efficiently as Ca2+
permeable L-channels is significantly enhanced by the presence of RyR-1 (retrograde signal of EC coupling; Refs. 2, 3). To investigate whether
or not the additional sarcolemmal DHPRs observed upon prolonged
reintroduction of RyR-1 in dyspedic myotubes are functionally coupled
to RyR-1, we compared L-current magnitudes in 3d-RyR-1- and
6d-RyR-1-expressing dyspedic myotubes. Fig.
2A shows representative L-currents that were obtained from 3d-RyR-1 (left) and
6d-RyR-1 (right) expressing myotubes. As illustrated in Fig.
2, L-current density was significantly greater for 6d-RyR-1-expressing
myotubes compared with 3d-RyR-1-expressing myotubes. On average, (Fig. 2B) peak L-current density increased from 8.5 ± 0.4 pA/pF (3 days; empty squares) to
12.5 ± 1.2 pA/pF (6 days; filled squares). This increase in L-current density
arose primarily from a ~30% increase in the maximal conductance of
L-channels (Gmax) and was not accompanied by significant
alterations in the voltage-dependence of L-channel activation
(VG 1/2 and kG, see also
Table I). Normal myotubes and 6d-RyR-1-expressing dyspedic myotubes
displayed nearly identical peak L-current densities (
12.5 ± 1.2 pA/pF Vs
12.9 ± 1.4 pA/pF) and both Gmax
and Qmax values (Table I; see also Ref. 3).
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To estimate the absolute levels of retrograde EC coupling for 3d-RyR-1- and 6d-RyR-1-expressing dyspedic myotubes, we normalized the L-current amplitudes by their respective Qmax values, to obtain the current-to-charge ratio (3). Following this normalization procedure, current-to-charge ratios were found to be similar for 3d-RyR-1- and 6d-RyR-1-expressing myotubes at every membrane potential (Fig. 2C). In addition, both experimental conditions exhibited nearly identical maximal conductance-to-charge ratios (Gmax/Qmax), which were similar to the corresponding value obtained for normal myotubes (Table I; see also Ref. 3). Consequently, no significant differences were found in either the current-to-charge or conductance-to-charge (Gmax/Qmax) ratios for normal myotubes or either 3d-RyR-1- or 6d-RyR-1-expressing myotubes. These results strongly suggest that a similar proportion of DHPRs interact with ryanodine receptors (as judged by the absolute levels of retrograde coupling) under these three conditions. Thus, the additional sarcolemmal DHPRs observed following long-term RyR-1 reintroduction into dyspedic myotubes represent primarily functional or RyR-1-coupled DHPRs.
Normal myotubes express two main types of voltage-dependent
calcium channels in the sarcolemma: L-channels and T-type calcium channels (T-channels). L-channels and T-channels exhibit distinct biophysical and pharmacological properties including thresholds of
activation, rates of channel activation and inactivation, unitary conductance, and sensitivity to dihydropyridines (14, 15). To
investigate whether the presence of RyR-1 also regulates T-channel expression, we compared T-currents recorded from normal and dyspedic myotubes (Fig. 3). T-channel activity was
dissected by recording calcium currents in the absence (total
ICa = T-current + L-current; Fig. 3, A-a,
B-a, C-a) and presence of a conditioning prepulse (12) designed to inactivate T-channels (L-current only; Fig. 3,
A-b, B-b, C-b). T-currents were then
revealed following offline subtraction of L-currents from the total
calcium current (Fig. 3, A-c, B-c,
C-c).
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Fig. 3C shows average current to voltage relationships of total ICa (a), L-current (b), and T-current (c). I-V curves were obtained from 17 normal (filled circles) and 22 dyspedic myotubes (empty circles). In normal myotubes, total ICa exhibits a prominent shoulder that divides the I-V curve at ~0 mV into the following two components: 1) a low-threshold component (more prominent at negative membrane potentials) and 2) a high threshold component (most clearly resolved at positive potentials). Application of the conditioning prepulse selectively eliminates the low-threshold component, without significantly affecting the high-threshold component (b, L-current). The low-threshold, rapidly inactivating component was then dissected by subtracting b from a (c, T-current). As reported previously (2, 4, 3), dyspedic myotubes exhibit very modest L-currents compared with those recorded from normal myotubes (Fig. 3, C-b). This arises from a 5-fold reduction in macroscopic channels conductance and an ~45% reduction in the number of L-channels in the sarcolemma. Interestingly, average T-channel current density was similar for both normal and dyspedic myotubes (Fig. 3, C-c). Apparently, only the number of sarcolemmal L-channels and not the number of T-channels are significantly influenced by the presence or absence of RyR-1. Thus, the RyR-1-mediated increase in L-channel expression does not represent a general overall increase in ion channel biosynthesis.
Introduction of RyR-1 into dyspedic myotubes restores the following two important Ca2+ homeostatic mechanisms: 1) an SR Ca2+ release pathway or skeletal-type EC coupling and 2) a Ca2+ influx pathway that is manifested as a ~10-fold increase in L-type current magnitude (2, 3). Because elevations in intracellular calcium concentration ([Ca2+]i) in response to activation of voltage- or ligand-gated Ca2+ channels are known to influence a variety of biochemical process in excitable cells (16), we investigated whether the RyR-1-mediated increase in DHPR expression requires restoration of the Ca2+ release pathway, Ca2+ influx pathway, or both.
Substitution of an alanine residue for a highly conserved glutamate
residue in either RyR3 (E3885A) or RyR-1 (E4032A) dramatically reduces
Ca2+ activation of the resulting SR Ca2+
release channel (17, 18). In addition, O'Brien et al. (8) found that expression of E4032A in dyspedic myotubes preferentially restores skeletal L-current activity (i.e. retrograde
coupling), but not robust, voltage-activated SR Ca2+
release (i.e. orthograde coupling). We exploited the ability of E4032A to preferentially restore the retrograde coupling to determine the relative importance of the Ca2+ influx and
release pathways on the RyR-1-mediated increase in DHPR expression.
Fig. 4 illustrates that expression of the
E4032A mutant in dyspedic myotubes fully restored the Ca2+
influx pathway (L-current or retrograde coupling) but not the Ca2+ release pathway (orthograde coupling). Fig.
4A shows L-currents (lower traces) and
intracellular calcium transients (upper traces) recorded
simultaneously from a normal myotube (left), a 3-day RyR-1-expressing dyspedic myotube (middle), and a 3-day
E4032A-expressing dyspedic myotube (right). The 3d-RyR-1-
and the 3d-E4032A-expressing dyspedic myotubes exhibited similar
L-current densities, which are ~45% smaller than L-currents recorded
from normal myotubes (Fig. 4A). As exemplified by the
representative experiments shown in Fig. 4A, normal and
3d-RyR-1-expressing dyspedic myotubes displayed similar robust
voltage-activated Ca2+ transients, whereas
3d-E4032A-expressing dyspedic myotubes exhibited only very modest
voltage-activated Ca2+ transients. On average, the presence
of the E4032A mutation caused an ~7-fold reduction in maximal SR
Ca2+ release compared with wild-type RyR-1 (Fig.
4B). Interestingly, E4032A-expressing dyspedic myotubes
exhibited a slight (~20 mV), but significant (p < 0.01) depolarizing shift in VF 1/2 compared with
those obtained from either normal myotubes or RyR-1-expressing dyspedic
myotubes. This observation is consistent with the notion that the
E4032A mutation stabilizes a SR Ca2+ release channel closed
state(s), thus causing the requirement of stronger depolarizations to
activate the release channel. Nevertheless, Ca2+
transients recorded from normal myotubes, as well as dyspedic myotubes
expressing either RyR-1 or E4032A, each exhibited a sigmoidal voltage-dependence demonstrating the presence of a skeletal-type (as
opposed to a Ca2+ influx-dependent) EC coupling
mechanism (Fig. 4C).
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As shown in Fig. 4, the E4032A mutation in RyR-1 provides a powerful
tool for dissecting the putative roles of calcium influx and calcium
release on functional DHPR expression. To test if the E4032A mutant
mimics the ability of wild-type RyR-1 to increase DHPR functional
expression, we compared L-current densities (Fig. 5, A and B) and
immobilization-resistant charge movements (Fig. 5, C and
D) for 3-day E4032A- and 6-day E4032A-expressing dyspedic myotubes. Remarkably, no significant differences were found in the peak
L-current density, Gmax, or Qmax values (Fig.
5, B and D; Table I). In addition, the
voltage-dependence of charge movements and L-current activation were
similar for 3-day E4032A- and 6-day E4032A-expressing dyspedic myotubes
(Fig. 5 and Table I). Thus, despite the restoration of large L-currents
(i.e. retrograde coupling), the E4032A mutant failed to
increase the number of functional L-channels over a time interval
during which wild-type RyR-1 expression caused a ~2-fold increase in
DHPR expression. These data indicate that restoration of the
Ca2+ release pathway, rather than the Ca2+
influx pathway, is required for the ability of RyR-1 to increase functional DHPR expression in skeletal myotubes.
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DISCUSSION |
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In this study, we have demonstrated that long-term expression of RyR-1 in dyspedic myotubes increases functional DHPR expression in the sarcolemma (i.e. increased both peak L-current and maximal intramembrane charge movement). The influence of RyR-1 appears to be restricted to L-channel expression because T-current density was similar for both normal and dyspedic muscle. Interestingly, long-term expression of E4032A, a mutant RyR-1 that preferentially affects the orthograde signal of E-C coupling (i.e. fully restores L-channel activity but not SR Ca2+ release) failed to increase functional DHPR expression. These results suggests that Ca2+ release through SR Ca2+ release channels (but not Ca2+ influx through L-channels) influences a signaling pathway in myotubes that promotes the expression of functional L-type Ca2+ channels.
There is increasing evidence to suggest that the expression level of
skeletal muscle DHPRs is subject to changes in intracellular Ca2+. In amphibian (frog) skeletal muscle, long-term
blockade of skeletal L-channels increases DHP binding and charge
movement, consistent with the notion that Ca2+ influx
inhibits DHPR expression in frog skeletal muscle (6). However, opposing
observations have been made using mammalian skeletal muscle
preparations. Specifically, elevations in extracellular Ca2+ (7) increase L-type tail current density and
intramembrane charge movement in mammalian (rat) cultured myotubes. Our
results support the observations of Renganathan et al. (7),
but appear to be in contrast to the postulated role of Ca2+
influx in frog skeletal muscle (6). However, numerous differences between DHPR function in amphibian and mammalian skeletal muscle have
been identified (e.g. molecular determinants of slow
activation, regulation by cAMP-dependent protein kinase A,
presence of Q, prepulse-induced kinetic acceleration; see O'Connell
and Dirksen, Ref. 19 and Melzer et al., Ref. 1 for review).
Thus, it is conceivable that the signaling pathways that modulate
functional DHPR surface expression represent yet another fundamental
difference between mammalian and amphibian skeletal muscle.
Nevertheless, it will be interesting to determine whether SR
Ca2+ release also influences functional DHPR expression in
frog skeletal muscle.
The precise molecular mechanism(s) by which SR Ca2+ release
increases the number of functional sarcolemmal DHPRs is currently unclear. Conceivably, alterations in myoplasmic Ca2+
homeostasis could alter pathways that control DHPR
1-subunit (or a DHPR regulatory protein such as the
-subunit) gene transcription (16), message stabilization (20),
and/or degradation. Global elevations in resting intracellular
Ca2+ levels have been reported to increase L-type
Ca2+ channel expression in both cardiac (21) and skeletal
muscle cells (7). However, Ca2+-mediated alterations in
gene transcription may not necessarily require global changes in
resting intracellular Ca2+ levels. Under certain
conditions, activity-dependent gene transcription has been
demonstrated to be activated by privileged local Ca2+
signaling mechanisms. For example, in hippocampal neurons,
Ca2+ influx through neuronal L-type Ca2+
channels activates the nuclear translocation of specific
Ca2+-dependent transcription factors, such as
CREB (22, 23) and NF-ATc4 (24). A similar privileged role for
Ca2+ influx through L-type Ca2+ channels in
CREB activation has recently been observed in vascular smooth muscle
(25). This elegant study demonstrated that depolarization-induced increases in Ca2+ influx through
voltage-dependent Ca2+ channels promotes CREB
phosphorylation (P-CREB) and results in increased c-fos
mRNA levels in intact mouse cerebral arteries. Interestingly,
Ca2+ release through RyRs in cerebral arteries reduced
P-CREB and c-fos mRNA levels, presumably by causing
membrane hyperpolarization following Ca2+ spark-mediated
activation of Ca2+-sensitive K+ channels (25,
26). Thus, at least one mechanism for activity-dependent control of gene expression in excitable cells involves the privileged ability of Ca2+ influx through voltage-gated
Ca2+ channels to fine-tune protein expression during a
process referred to as excitation-transcription coupling (27).
Our results demonstrate that long-term expression of the skeletal
muscle RyR-1 appears to modulate functional DHPR expression through a
novel form of excitation-transcription coupling. Because SR
Ca2+ release is an essential component of Ca2+
signaling in skeletal muscle cells, RyR-1 activity could serve a
central role in regulating protein expression in skeletal muscle. Insulin-like growth factor-1 induces skeletal muscle hypertrophy through activation of the Ca2+-dependent
transcription factor NF-ATc1 in myotubes (28, 29) and promotes
L-channel gene expression in rat skeletal muscle (30). Our data provide
strong evidence for a novel pathway that involves the privileged
ability of Ca2+ released through RyR-1s, rather than influx
through L-type Ca2+ channels, to strongly influence the
degree of functional DHPR expression in cultured myotubes. Thus, it
will be important for future studies to determine the precise
downstream molecular components (e.g. NF-ATc1, CREB) that
are activated by Ca2+ released through RyR-1s and if this
pathway is recruited by growth factors such as insulin-like growth
factor-1. Independent of the precise molecular mechanism, our data
demonstrate the requirement for robust SR Ca2+ release in
the up-regulation of the number of functional DHPRs and provides an
explanation for the reduction in DHPR levels observed in both dyspedic
muscle homogenates (4, 5) and cultured dyspedic myotubes (3).
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ACKNOWLEDGEMENTS |
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We thank Drs. Kurt G. Beam and Paul D. Allen for providing us access to the dyspedic mice used in this study as well as for their advice and continued support.
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FOOTNOTES |
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* This work was supported by National Institutes of Health Grant AR44657 (to R. T. D.) and a Neuromuscular Disease research grant (to R. T. D.).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.
Recipient of Consejo Nacional de Ciencia y Technologia
(CONACYT) Postdoctoral Fellowship 990236.
§ To whom correspondence should be addressed. Tel.: 716-275-4824; Fax: 716- 273-2652; E-mail: Robert_Dirksen@URMC.rochester.edu.
Published, JBC Papers in Press, January 22, 2001, DOI 10.1074/jbc.M009685200
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ABBREVIATIONS |
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The abbreviations used are: DHPR, dihydropyridine receptor; L-type Ca2+ channel, L-channel; EC, excitation-contraction; SR, sarcoplasmic reticulum; RyRs, ryanodine receptors; CREB, cAMP-response element-binding protein.
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