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Volume 17, Number 18,
Issue of September 15, 1997
pp. 6918-6928
Copyright ©1997 Society for Neuroscience
Regulation of Mouse Skeletal Muscle L-Type Ca2+
Channel by Activation of the Insulin-Like Growth Factor-1 Receptor
Osvaldo Delbono1, 2,
Muthukrishnan Renganathan2, and
María Laura Messi 1
Departments of 1 Physiology and Pharmacology and
2 Internal Medicine (Gerontology), The Bowman Gray School of
Medicine of Wake Forest University, Winston-Salem, North Carolina 27157
 ABSTRACT
- INTRODUCTION
- MATERIALS AND METHODS
- RESULTS
- DISCUSSION
- FOOTNOTES
- REFERENCES
ABSTRACT 
We investigated the modulation of the skeletal muscle L-type
Ca2+ channel/dihydropyridine receptor in response to
insulin-like growth factor-1 receptor (IGF-1R) activation in single
extensor digitorum longus muscle fibers from adult C57BL/6 mice. The
L-type Ca2+ channel activity in its dual role as a
voltage sensor and a selective Ca2+-conducting pore
was recorded in voltage-clamp conditions. Peak Ca2+
current amplitude consistently increased after exposure to 20 ng/ml
IGF-1 (EC50 = 5.6 ± 1.8 nM). Peak IGF-1
effect on current amplitude at  20 mV was 210 ± 18% of the
control. Ca2+ current potentiation resulted from a
shift in 13 mV of the Ca2+ current-voltage
relationship toward more negative potentials. The IGF-1-induced
facilitation of the Ca2+ current was not associated
with an effect on charge movement amplitude and/or voltage
distribution. These phenomena suggest that the L-type
Ca2+ channel structures involved in voltage sensing
are not involved in the response to the growth factor. The modulatory
effect of IGF-1 on L-type Ca2+ channel was blocked
by tyrosine kinase and PKC inhibitors, but not by a cAMP-dependent
protein kinase inhibitor. IGF-1-dependent phosphorylation of the L-type
Ca2+ channel  1 subunit was demonstrated by
incorporation of [ -32P]ATP to monolayers of adult
fast-twitch skeletal muscles. IGF-1 induced phosphorylation of a
protein at the 165 kDa band, corresponding to the L-type
Ca2+ channel 1 subunit. These results
show that the activation of the IGF-1R facilitates skeletal muscle
L-type Ca2+ channel activity via a PKC-dependent
phosphorylation mechanism.
Key words:
insulin-like growth factor;
calcium channel;
skeletal
muscle;
muscle fiber;
phosphorylation
INTRODUCTION
IGF-1 is a peptide structurally
related to proinsulin and has a primary role in promoting the
differentiation and growth of skeletal muscle, effects that occur in a
relatively slow time scale (delayed effects) (Florini et al., 1996 ).
Manipulation of IGF-1 expression in vivo and in
vitro provides fundamental clues about its action on skeletal
muscle growth and differentiation (DeVol et al., 1990; Vandenburgh et
al., 1991; Coleman et al., 1995). In addition to the delayed effects on
cellular trophism, it has been shown that IGF-1 stimulates
Ca2+ influx in clonal pituitary and neuroblastoma
cell lines (Kleppisch et al., 1992; Selinfreund and Blair, 1994).
Although IGF-1 also exerts potent trophic and developmental effects on
skeletal muscle (Cohick and Clemmons, 1993), modulatory effects of
IGF-1 on skeletal muscle Ca2+ channels have not been
studied.
Because of the role of Ca2+ ions in mediating and/or
triggering short- and long-lasting cellular responses (Berridge, 1993), it is relevant to identify the signaling pathway linking trophic factor
receptor activation and voltage-gated Ca2+ channel
function. In skeletal muscle the L-type Ca2+
channel, a dihydropyridine-sensitive subtype, serves in its dual role
as a voltage sensor and a pore-conducting Ca2+ ion
pathway. Both functions reside in the 1 subunit,
resulting from the expression of the 1S gene (Tanabe et
al., 1987). The L-type Ca2+ channel, as a voltage
sensor, releases Ca2+ from the sarcoplasmic
reticulum as a result of interaction with the ryanodine receptor (RYR1)
(Meissner, 1995; Delbono and Meissner, 1996). As a pore-conducting
pathway for Ca2+ ions, the L-type channel may
participate in the activation of long-lasting intracellular signaling
cascades, relevant for muscle fiber differentiation and trophism at
different stages of ontogenic development. Phosphorylation potentiates
Ca2+ influx through voltage-gated
Ca2+ channels (Sculptoreanu et al., 1993).
Intracellular Ca2+ elevations have been involved in
sustained kinase activation and signaling to the nucleus with
consequent modulation of gene expression (Nishizuka, 1995). The force
of muscle contraction can be modified according to the duration and
frequency of stimulation. During single twitches, contraction is not
dependent on extracellular Ca2+, and the L-type
Ca2+ channels may function only as voltage sensors.
However, prolonged or repetitive contractions are dependent on
extracellular Ca2+ and are sensitive to L-type
Ca2+ channel antagonists (Kotsias et al., 1986;
Dulhunty et al., 1988; Sculptoreanu et al., 1993).
Ca2+ entry through this channel is thought to
replenish intracellular Ca2+ (Oz et al., 1991).
Also, a phosphorylation-dependent potentiation of this current
increased contractile force (Schmid et al., 1985; Arreola et al., 1987;
Huerta et al., 1991).
In the present work we determined whether the skeletal muscle L-type
Ca2+ channel dihydropyridine receptor is a potential
target for the IGF-1R, based on the observation that tyrosine
kinase-linked receptors promote phosphorylation of diverse cellular
proteins and that the L-type Ca2+ channel undergoes
phosphorylation at defined consensus sequences (see below). In this
paper we report the novel finding that the L-type
Ca2+ channel can be phosphorylated in adult living
skeletal muscle by Ca2+-independent protein kinase C
(PKC) isoforms on IGF-1R activation.
MATERIALS AND METHODS
Single skeletal muscle fiber isolation. Single
extensor digitorum longus (EDL) muscle fibers from 14-month-old C57BL/6
mice were used. Mice were obtained from the National Center for
Toxicological Research (Jefferson, AK). Animals were housed in a
pathogen-free area at The Bowman Gray School of Medicine (BGSM;
Winston-Salem, NC) until the day of experimentation. Animal handling
and procedures followed an approved protocol by the Animal Care and Use
Committee of BGSM. Muscles were dissected after the mice were
decapitated. Anesthesia was not provided because the effects of
anesthetics on the specific endpoints of our research are not
characterized. In addition, procedures were optimized to minimize
animal pain or discomfort. Electrophysiological recordings were
performed within 5 hr of muscle dissection. Single fibers were
dissected in "dissecting solution" and transferred to the recording
chamber containing "mounting solution." Fibers with a small radius
(30-40 µm), representing 61 ± 5% (n = 10 EDL
muscles) of the muscle fiber population, were selected to reduce
nonuniformities in the voltage clamp of the T tubule membrane.
Monolayer of skeletal muscle preparation for phosphorylation
studies. White (fast-twitch) leg muscles were dissected and rinsed in a modified Ringer's solution (see Solutions and Materials) at room
temperature. Whole muscles were transferred to a dissecting solution
(described below), and multiple monolayers of skeletal muscle were
dissected under a stereoscope. Connective tissue was removed carefully
without damaging the cells. Skeletal muscle fiber monolayers were
sectioned into smaller fragments of ~500 µM length to
facilitate the exchange of intracellular constituents. Short muscle
monolayers were stored in a phosphate-free solution similar to the
intracellular solution used for Ca2+ current
recording (see below). The following protease inhibitors were added to
the phosphate-free solution and solubilization buffer: phenylmethylsulfonyl fluoride (0.2 mM), aprotinin (100 nM), leupeptin (1 µM), pepstatin (1 µM), and diisopropyl fluorophosphate (1 mM).
L-type Ca2+ channel 1 subunit
phosphorylation. Phosphorylation studies were performed, using
conditions optimized in previous studies (O'Callahan et al., 1988;
Chang et al., 1991). Muscle monolayers were preincubated for 20 min at
room temperature in phosphate-free solution and for another 15 min in
the same solution to which 50 µCi of [-32P]ATP
(specific activity of 10 Ci/mmol) was added. The phosphorylation reaction was initiated by adding IGF-1 to a final concentration of 20 ng/ml for 20 min and was stopped with two volumes of stop buffer
containing (in mM) 50 HEPES-NaOH, pH 7.4, 50 NaF, and 20 EDTA plus protease inhibitors. The radiolabeled muscle fibers were
washed three times with phosphate-free and low Ca2+
buffer (see below) and were homogenized in a Thomas Tissue grinder (Hand homogenizer) 3431-E20 (A. H. Thomas, Philadelphia, PA) with 20 hand strokes. Protein concentration of the homogenate was estimated by Coomassie protein assay, using bovine serum albumin as the protein
calibration standard. The homogenate was mixed with an equal volume of
2× sample buffer containing 62.5 mM Tris-HCl, 2% SDS, 100 mM DTT, 10% glycerol, and 0.02% bromophenol blue. The
sample prepared for sodium dodecyl sulfate (SDS) gel electrophoresis was boiled for 5 min at 95°C, cooled for 5 min, and centrifuged in a
microfuge at 5000 rpm for 3 min. Gradient electrophoresis was performed
in a 5-15% SDS polyacrylamide gel, stained with Coomassie blue, and
destained to visualize the proteins. From a 200 mg pool of skeletal
muscle monolayers, 0.8-1.6 µg of protein was obtained. In all of the
experiments, 100 µg protein/lane was loaded. SDS-PAGE 42,699-200,000
(Bio-Rad, Richmond, CA) was used as a high molecular range standard.
Protein molecular weight calculation was performed by logarithmic
extrapolation. The gel was incubated with 10% glycerol for 2 hr and
dried at 55°C for 2 hr under vacuum. Gels were subjected to
autoradiography, using Kodak X-Omat film and Cronex enhancing screens
for 3 d. For quantitation of protein phosphorylation in films, the
average density of pixels across the band width was integrated over the
band height with an optical scanner. For this analysis a Pdi image
system, together with Quantity One 2.6 software (New York, NY), was
used. The magnitude of protein phosphorylation was expressed as the
integral of the optical density (O.D. × mm2).
L-type Ca2+ channel 1 subunit antibody
coupling to Sepharose 4B and immunoprecipitation. For antibody
coupling to Sepharose 4B, 100 mg of freeze-dried cyanogen
bromide-activated Sepharose 4B powder was suspended in 5 ml of ice-cold
1 mM HCl for 15 min and then washed four times with 5 ml of
ice-cold 1 mM HCl. The final aliquot of 1 mM
HCl was aspirated from the beads, and then the beads were transferred
immediately to mouse monoclonal antibody specific for L-type
Ca2+ channel 1 subunit (Chemicon,
Temecula, CA). A total of 100 ml of monoclonal antibody was mixed
previously with 4× coupling buffer containing 0.1 M
NaHCO3, pH 8.3, and 0.5 M NaCl to
facilitate the coupling of antibody to the beads and to yield a volume
of 0.5 ml of beads. The beads in antibody solution were mixed in an
end-over-end mixer overnight at 4°C. The beads were centrifuged at
low speed, transferred to a buffer with blocking agent (0.2 M glycine, pH 8.0), and mixed for 16 hr at 4°C to block
the remaining active groups. To remove the excess uncoupled ligand, we
washed the adsorbent alternately with high pH coupling buffer and a low pH acetate buffer solution five times. The low pH buffer solution contained 0.1 M acetate and 0.5 M NaCl, pH 4.0. Antibody/Sepharose 4B beads were stored at 4°C in PBS buffer in the
presence of 1 mM sodium azide. Muscle fibers were
phosphorylated as described before but homogenized in 25 mM
NaKPO4 (20 mM Na2HPO4
plus 5 mM KH2PO4), 20 mM NaF, and 0.1% digitonin to prevent dephosphorylation during immunoprecipitation (Gutierrez et al., 1991). The
antibody/Sepharose 4B beads were washed five times with incubation
buffer consisting of (in mM) 25 NaKPO4,
20 NaF, and 200 NaCl with 0.1% digitonin and 1 mg/ml bovine serum
albumin; then the beads were incubated with muscle homogenate (2 mg of
protein equivalent of muscle homogenate was added to 200 ml of
Sepharose 4B beads) at 4°C overnight. The beads were washed three
times with incubation buffer, and the immunoprecipitated proteins were
eluted by boiling the beads in SDS sample buffer before analysis by SDS
gel electrophoresis, immunoblot, and autoradiography. The extent of
immunoprecipitation was estimated from an analysis of the supernatant
in SDS-PAGE.
Muscle fiber voltage-clamp, ionic current, and charge movement
recording. EDL fibers were voltage-clamped, using the double Vaseline gap technique at a holding potential
(Vh) of 90 mV, as described previously
(Delbono, 1992). This technique has been used for adult skeletal muscle
voltage clamp in the present work because it was demonstrated that the
double Vaseline gap resulted in an improvement in the frequency
response, compensation for external series resistance, and compensation
for the complex impedance of the current-passing pathway, as compared
with other potentiometric methods (Hille and Campbell, 1976). Since
these initial reports, the double Vaseline gap technique has been used
for charge movement and ICa recordings in mature
amphibian and mammalian fibers (Hui and Chandler, 1990; Delbono, 1992).
For voltage-clamp recordings a muscle fiber was mounted in the
recording chamber. Two Vaseline strands separated a central pool,
equilibrated with the "external solution," from lateral
compartments, equilibrated with the "internal solution." Glass
bridges containing agar equilibrated in 1 M-KCl provided
electrical connections between each compartment, and separate wells
were filled with 3 M KCl and fit with Ag-AgCl pellets. Command pulses referred to ground were applied at the central pool. The
current was injected into one of the end pools via a variable gain
feedback amplifier. The negative input of the amplifier was connected
to the other end pool. Passive muscle membrane properties were measured
according to Irving et al. (1987). Temperature was maintained at 22°C
throughout the experiments and monitored with a thermistor probe
positioned close to the fiber in the middle pool.
For charge movement recording, 1 µM nifedipine was
preferred over divalent cations such as Co2+ and/or
Cd2+. We found the use of nifedipine less
deleterious to the fiber than the use of Co2+ and/or
Cd2+ at the concentrations required to block the
ionic current completely (2 and 1 mM). We selected a
procedure to block the ionic current with less deleterious effect on
the integral of the charge movement and on the passive membrane
properties. Nifedipine induced a decrease in the integral of the charge
movement recorded 1 hr after complete blockade of the ionic
conductance, which was 15 ± 4.2% of control (n = 7). The decrease induced by the combination of Co2+
and Cd2+ was 31 ± 2.4% (n = 7). Co2+ plus Cd2+ also promoted
a higher increase in the holding current than nifedipine. The range of
increase was 20-35 nA (n = 7) and 5-20 nA
(n = 8), respectively. Membrane current during a
voltage pulse (P) initially was corrected by analog subtraction of
linear components. The remaining linear components were subtracted
digitally by automated scaling of control pulses, which were
1/4 of P (Bezanilla, 1986; Delbono et al., 1991).
The inward Ca2+ current
(ICa)-voltage relationship was fit to
the following equation:
![I<SUB><UP>Ca</UP></SUB>=G<SUB><UP>max</UP></SUB>(V−V<SUB><UP>r</UP></SUB>)/{1+<UP>exp</UP>[<UP>F</UP>(V−V<SUB>1/2</SUB>)/<UP>RT</UP>]},](/content/vol17/issue18/fulltext/6918/img001.gif)
|
(1)
|
where V is the membrane potential,
Vr is the extrapolated reversal potential,
V1/2 is the half-activation potential, F
is the Faraday constant, R is the gas constant, and T is the absolute temperature (296°K).
For the analysis of the voltage dependence of charge movement, the
following equation was used:
![Q<SUB><UP>on</UP></SUB>=Q<SUB><UP>max</UP></SUB>/{1+<UP>exp</UP>[z<SUB><UP>Q</UP></SUB><UP>F</UP>(V<SUB>1/2<UP>Q</UP></SUB>−V<SUB><UP>m</UP></SUB>)/<UP>RT</UP>]},](/content/vol17/issue18/fulltext/6918/img002.gif)
|
(2)
|
where Vm is the membrane potential,
Vr is the extrapolated reversal potential,
Qmax is the maximum charge and
V1/2Q is the charge movement
half-activation potential, F is the Faraday constant, R is the gas
constant, T is the absolute temperature (296°K), and z is
the effective valence.
Cell capacitance was monitored throughout the experiments by applying
brief hyperpolarizing pulses from a holding potential of 90 to 110
mV for 25 msec. Cell capacitance, expressed in F (Faraday), was
calculated from the integral of the current transients and the
amplitude of the voltage step. The specific membrane capacity, expressed in F/cm2, represents the cell capacitance
normalized to the fiber cross-sectional area.
For data acquisition a personal computer was used. D-A and A-D
conversions were done with a Digidata 1200 acquisition board and
interface (Axon Instruments, Foster City, CA). Stimulation protocols
are detailed in Results for each group of experiments. Currents were
sampled at 5-10 kHz according to the stimulation protocol and filtered
at 0.3 of the sampling frequency (3 dB point) with a four-pole
Butterworth low-pass filter (Frequency Devices, Haverhill, MA).
Membrane currents were normalized to membrane capacitance (amperes per
farad).
A dose-reponse curve was obtained by a nonlinear least-squares fit of
a four-parameter function, using a nonlinear curve-fitting procedure
(SigmaPlot, Jansen, San Rafael, CA), where:
![y=(I−F)/(1+([<UP>concentration of factor</UP>]/<UP>EC</UP><SUB>50</SUB>)<SUP><UP>slope</UP></SUP>+F),](/content/vol17/issue18/fulltext/6918/img003.gif)
|
(3)
|
with I being the initial level and F the
final saturating level (Selinfreund and Blair, 1994). Experimental data
are expressed as mean ± SEM.
Solutions and materials. The extracellular solution for
ICa and charge movement recording contained (in
mM): 150 TEA (tetraethylammonium hydroxide)
CH3SO3 (methanesulfonic acid), 2 CaCl2, 2 MgCl2, 5 TEA HEPES,
0.001 tetrodotoxin, 1 9-anthracenecarboxylic acid, and 1 3,4-diaminopyridine. For charge movement recording, 1 µM nifedipine (Sigma, St. Louis, MO) was added from a stock prepared in
ethanol and handled in the dark. The intracellular solution contained
(in mM) 120 Na-glutamate, 15 Na2-EGTA, 3 Mg-ATP, 5 Na2-phosphocreatine, 10 Na-HEPES, and 10 glucose,
pH 7.4, at 300 mOsm. The modified Ringer's solution contained (in
mM) 145 NaCl, 5 KCl, 2.5 CaCl2, 1 MgSO4, 10 Na-HEPES, and 10 glucose. The dissecting
solution contained (in mM) 95 K2SO4, 10 MgCl2, 0.4 CaCl2, and 10 Na-HEPES. The mounting solution
contained (in mM) 150 K-glutamate, 2 MgCl2, 1 K2-EGTA, and 10 K-HEPES. The
phosphate-free solution contained (in mM) 98 K-glutamate,
0.1 K2-EGTA, 0.0082 CaCl2, 5.5 MgCl2, 5.0 glucose, and 5.0 HEPES-KOH. The pH was
adjusted to 7.4 in all of the solutions. Osmolarity was fixed to 300 mOsm. GF-109203X was kindly provided by Dr. Linda McPhail (Department
of Biochemistry, BGSM). Human recombinant IGF-1 was purchased from
Research Biochemicals (Natick, MA). Phorbol-12-myristate, 13-acetate
(PMA), 4-phorbol, genistein, PKC inhibitor peptide 19-36, daidzein,
and herbimycin A were all purchased from Calbiochem (La Jolla, CA).
cAMP-dependent protein kinase A inhibitor 5-24 and fura-2
pentapotassium salt were purchased from Peninsula (Belmont, CA) and
Molecular Probes (Eugene, OR), respectively. The anti-phosphotyrosine
antibody 4G10 was purchased from Upstate Biotechnology (Lake Placid,
NY).
Statistical analysis. Data were analyzed by paired and
unpaired Student's t test, one-way ANOVA with
Student-Newman-Keuls test, as indicated for each group of
experiments.
RESULTS
IGF-1 increases Ca2+ current through L-type
Ca2+ channels
The activity of the dihydropyridine-sensitive
Ca2+ channel was recorded with
Ca2+ as the charge carrier. Ca2+
channels were activated repetitively by 60 msec test depolarizations to
10 or 20 mV from a holding potential
(Vh) of 90 mV. Despite the lower
IGF-1-induced ICa (calcium current) potentiation
at 20 than at 30 mV, repetitive single-pulse stimulation was
performed at 20 mV, because 60 msec pulses at 30 mV evoked smaller
currents in control conditions (see below). Ca2+
channel activity was studied in paralyzed single muscle fibers. Fibers
were immobilized by strong myoplasmic Ca2+ chelation
with 15 mM EGTA added to the internal solution.
Simultaneous intracellular Ca2+ concentration
measurements were performed in a group of experiments, using a 50 µM concentration of the high-affinity
Ca2+ indicator fura-2. The peak myoplasmic
Ca2+ concentration in 15 mM EGTA was
108 ± 11 nM (n = 7). These results did not differ significantly from separate control studies (without EGTA) (117 ± 15 nM, n = 7;
p > 0.05). In these experimental conditions the
preparation was stable and maintained the sarcolemmal electrical properties for >1 hr. The holding current ranged from 5 to 20 nA,
and ICa amplitude and charge movement
(Q) were within 92 ± 2.8% (n = 30) of the initial values 1 hr after the fiber was voltage-clamped in
control experiments (without exposure to IGF-1). Muscle fiber capacitance, calculated from the integral of the current transients and
the amplitude of the voltage step, and the specific membrane capacity
did not change significantly throughout the experiment. Specific
membrane capacity at the beginning of the experiments (switch from
current-clamp to voltage-clamp mode) was 6.8 ± 0.4 F/cm2, whereas at the end its value was 6.5 ± 0.7 F/cm2 (n = 30). IGF-1 was
diluted to a final concentration of 20 ng/ml in the external solution
and was applied to the muscle fiber by completely exchanging the
bathing solution three times in 30 sec. Because the IGF-1R binds IGF-1
with high affinity and IGF-II and insulin with lower affinity (De Pablo
et al., 1990), we tested the effect of insulin and IGF-II on
Ica amplitude. Both insulin and IGF-II failed to
promote Ica potentiation at 20 ng/ml. The ratio
between peak ICa amplitude in the presence and
absence of insulin or IGF-II at 15 min after adding the factor was
0.96 ± 0.11 (n = 5; p > 0.5) and
0.95 ± 0.16 (n = 6; p > 0.5),
respectively. These results are in agreement with previous publications
in which insulin was two orders of magnitude less potent than IGF-1,
and IGF-II was completely ineffective in modulating the electrical properties of GH4C1 cells and cerebellar
granular cells (Selinfreund et al., 1994). The IGF-1 concentration used
in this study was within the range known to promote muscle cell
proliferation and differentiation in vitro (Allen et al.,
1985) and to increase ICa in clonal pituitary
cells (Selinfreund and Blair, 1994). Single depolarizing pulses were
applied to the fiber every 30 sec, and the peak
ICa amplitude during the pulse was computed.
Corrected ICa values to membrane capacitance
were normalized to the maximum amplitude at the end of the pulse and
averaged (Fig. 1A). A
complete solution exchange with the same bathing solution (Fig.
1A) was performed systematically as a control before
IGF-1 application. Peak ICa amplitude
consistently increased within the first 30 sec after exposure to the
growth factor, reaching a maximum effect in 5-7 min and persisting
after its removal (c). Peak IGF-1 effect on
ICa amplitude was 2.1 ± 0.18 times the
control. The IGF-1 effect on peak Ica continued
up to 30 min after its removal from the bathing solution. At that time
the effect was sustained in 1.9 ± 0.25 of the control. Brief
hyperpolarizing pulses elicited by 25 msec pulses from
Vh to 110 mV showed no significant changes in
sarcolemmal linear capacitive transient as an indication of the lack of
alterations in the membrane passive properties throughout the
experiment. The values corresponding to the first and second capacitive
transients are 6.6 and 6.4 F/cm2, respectively.
Figure 1B illustrates the IGF-1-mediated
ICa potentiation within the period indicated in
Figure 1A. A sequence of repetitive pulses before the
drug application (b) was used as the baseline for data
normalization. The initial upper deflection, followed by two downward
deflections, corresponds to charge movement ICa during fiber depolarization and tail current after repolarization, respectively. A clear and significant effect on
ICa amplitude during the pulse and on the tail
current was detected after IGF-1 application (b) and removal
(c). The effect of IGF-1 on ICa
potentiation is dose-dependent. Figure 2
shows the normalized peak Ica-IGF-1 concentration relationship. Experimental points were fit to Equation 3
(see Materials and Methods), giving an EC50 value of
5.6 ± 1.8 nM (n = 6).
Fig. 1.
Time course of IGF-1-induced
ICa potentiation in single skeletal muscle
fibers. A, ICa was elicited
by 60 msec depolarizing test pulses from a
Vh of 90 to 20 mV every 30 sec. A
complete solution exchange with the same bathing solution was used as a control (a). IGF-1 (20 ng/ml) was applied in
b and removed in c. Both
insets illustrate the capacitive transient current in response to brief hyperpolarizations to 110 mV at time 0 and at 15 min after adding IGF-1 to the bathing solution. B,
IGF-1-induced ICa potentiation during the
time indicated in A. IGF-1 application and removal are
depicted by arrows at b and
c, respectively. The interrupted
line indicates the baseline current.
[View Larger Version of this Image (31K GIF file)]
Fig. 2.
IGF-1 induces ICa
potentiation in a dose-dependent manner. ICa
was elicited by 60 msec depolarizing test pulses from a
Vh of 90 to 20 mV every 30 sec.
Increasing concentrations of IGF-1 were added to the muscle fiber. Data
for individual fibers were fit to Equation 3 (see Materials and
Methods). Average values for six complete experiments and fitting to
mean values for each IGF-1 concentration are included in the
plot.
[View Larger Version of this Image (14K GIF file)]
These experiments support the concept that IGF-1R activation triggers
an intracellular and/or membrane signaling cascade having the L-type
Ca2+ channel as a target. It also can be postulated
that the time course of the growth factor effect may result from a
process of phosphorylation and dephosphorylation in which kinase
activation and phosphatase activity result from IGF-1 binding to its
specific tyrosine kinase-linked receptor. The lack of effect on charge movement, together with a potentiation of
ICa, seems to exclude the possibility
that IGF-1 alters the voltage sensor. These findings also suggest that
phosphorylation of Ca2+ channel structures involved
in Ca2+ permeation are a likely coupling mechanism
to the IGF-1R. To test these concepts further, we assessed the role of
protein kinases on L-type Ca2+ channel
voltage-sensing function.
IGF-1 facilitates ICa activation without
affecting the voltage sensor
The voltage distribution for ICa was
studied, using 250 msec pulses, from a Vh = 90
mV, with 10 mV increments, from 80 to +30 mV. The voltage
distribution for ICa was studied in control (before adding IGF-1) and 15 min after starting the incubation in
IGF-1, a time at which a plateau for ICa
potentiation was reached (see Fig. 1A). Figure
3A shows
ICa control traces from 40 mV (threshold for
ICa activation) to 10 mV (saturating potential). The voltage distribution for ICa amplitude is
similar to that described previously for mammalian skeletal muscle
fibers (Delbono, 1992; Delbono et al., 1995). After exposure to IGF-1,
ICa was potentiated significantly at negative
voltages (40 to 10 mV). Figure 3B also shows
ICa potentiation of the tail current amplitude. Tail current amplitudes, measured during the repolarization immediately after voltage-clamp steps, at 20 mV were 4.1 ± 0.51 and
8.3 ± 0.64 µA/µF in control and IGF-1, respectively
(n = 12; p < 0.01). Tail current
amplitudes at 30 mV were 1.3 ± 0.1 and 4.8 ± 0.03 µA/µF in control and IGF-1, respectively (n = 12;
p < 0.01). Experiments in control and in response to
IGF-1 were not significantly different from 0 to 30 mV. Results are
consistent with measurements of tail currents over a voltage range
applied after a constant depolarizing pulse. Tail current amplitudes at
20 mV were 4.7 ± 0.62 and 7.9 ± 0.43 µA/µF in
control and IGF-1, respectively (n = 12; p < 0.01). Tail current amplitudes at 30 mV were
1.7 ± 0.14 and 5.2 ± 0.08 µA/µF in control and
IGF-1, respectively (n = 12; p < 0.01). The voltage dependence of ICa
potentiation is shown in Figure 3C, where control
(filled circles) and IGF-1 (open circles) experiments were plotted (n = 20).
ICa amplitude during the pulse was augmented
twofold at 20 mV and fourfold at 30 mV. At 20 mV, the values for
control and IGF-1 were 1.4 ± 0.03 and 2.9 ± 0.05 µA/µF, respectively, and at 30 mV the values were 0.5 ± 0.06 and 2.1 ± 0.07 µA/µF, respectively
(p < 0.01). Gmax and V1/2 were obtained by fitting the
Ica-voltage relationship to Equation 1 (see
Materials and Methods). Gmax was potentiated
significantly in approximately fourfold, and ICa
potentiation resulted from a shift in 13 mV of the
ICa-voltage relationship toward more negative potentials in response to IGF-1 (Table
1).
Fig. 3.
IGF-1 facilitates ICa
activation. ICa was elicited by 250 msec
depolarizations from a Vh of 90 mV, with
10 mV increments, from 80 to +30 mV. ICa
was recorded in a range of membrane potentials (from 40 to +10 mV),
before (A) and after exposure to IGF-1 (15 min; B). Offsets were imposed to the traces to better
observe tail current amplitude. The interrupted line
indicates the baseline current. C, Peak
ICa-V relationship in
control (filled circles) and in IGF-1
(open circles). Experimental points (mean ± SEM) were fit to Equation 1 included in Materials and Methods
(n = 20; p < 0.01 from 40 to
10 mV).
[View Larger Version of this Image (17K GIF file)]
To determine whether IGF-1 alters L-type Ca2+
channel pore-conducting function as well as the voltage sensor, we
performed a series of experiments after Ca2+
conductance was blocked with 1 µM nifedipine. Figure
4 illustrates charge movement traces from
70 to 10 mV in control (A) and after exposure to
IGF-1 for 15 min (B; n = 15). Charge
movement became measurable at 80 to 70 mV and saturated at 10 to
0 mV for both groups. The threshold values for charge movement were
similar in both conditions, 70 ± 3.2 and 72 ± 2.9 mV
(p > 0.5) in control and in IGF-1,
respectively. IGF-1 did not change the voltage distribution significantly for the integral of the charge movement at the beginning of the pulse (charge ON; Fig. 4C). Experimental points were
fit to Equation 2 (see Materials and Methods). The lack of
statistically significant effect on Qmax,
V1/2Q, and zQ
values (Table 1; p > 0.5) supports the concept that
IGF-1 does not alter the voltage sensor.
Fig. 4.
IGF-1 does not alter the voltage sensor. Charge
movement is in response to 120 msec duration pulses at the voltages
indicated on the right. Charge movement traces are shown
in control (A) and after exposure to IGF-1 for 15 min (B). The interrupted line indicates the baseline current. C, Voltage distribution
of the integral of the charge movement at the beginning of the pulse (charge ON) in control (filled circles) and in
IGF-1 (open circles). Data points were fit to Equation 2
detailed in Materials and Methods (n = 15;
p > 0.5).
[View Larger Version of this Image (19K GIF file)]
A potential effect of IGF-1 on the dihydropyridine-sensitive fraction
of charge movement was explored. Ica was blocked
completely with 2 mM Co2+ and 1 mM Cd2+, and 20 ng/ml IGF-1 was added
subsequently. Charge movement parameters were measured 15 min after
adding IGF-1 or the same volume of vehicle to the bathing solution.
Qmax values in the presence and absence of IGF-1
were 8.8 ± 1.1 and 8.3 ± 1.5 pC/nF, respectively (not
statistically significant). V1/2Q values in both
experimental conditions were 44.8 ± 5.1 and 45.5 ± 4.4 mV, respectively (not statistically significant; p > 0.5; n = 10). The lack of effect of IGF-1 on the
dihydropyridine-sensitive fraction of charge movement was confirmed by
using 0.1 mM tetracaine as a ICa
blocker (Lamb, 1986) in a different set of experiments.
Qmax values in the presence and absence of IGF-1
were 9.2 ± 1.4 and 8.9 ± 1.5 pC/nF, respectively (not
statistically significant; p > 0.5).
V1/2Q values in both experimental conditions
were 43.4 ± 4.6 and 44.1 ± 3.7 mV (not statistically
significant; n = 10).
IGF-1 promotes a facilitation of the channel activation that is
manifest in a shift of the Ica-V
relationship toward more negative potentials. This phenomenon is not
associated with changes in the effective valence, suggesting that the
transitions governing the movement of charged particles in the voltage
sensor are not regulated by IGF-1. In summary, IGF-1-mediated
ICa potentiation occurs via a direct effect on
the structures involved in Ca2+ ion permeation,
facilitating the channel activation, and not via an effect on the
voltage sensor.
IGF-1-induced ICa potentiation is mediated
by Ca2+-independent PKC isoforms
Tyrosine kinase activity, which is associated with the IGF-1R,
suggests that the mechanism of interaction with other sarcolemmal proteins such as the L-type Ca2+ channel is via a
phosphorylation mechanism. Phosphorylation sites for several protein
kinases have been identified in skeletal muscle L-type
Ca2+ channels but not for tyrosine kinase. This
prompted us to explore the pathway for IGF-1-induced phosphorylation of
the L-type Ca2+ channel 1 subunit underlying
ICa up-modulation.
Figure 5 shows the lack of IGF-1 effect
on ICa amplitude after muscle fiber incubation
in an intracellular solution containing the tyrosine kinase inhibitor
genistein (100 µM) (open circles) for
15-20 min. Neither genistein nor DMSO (used as a vehicle) had obvious
deleterious effects on the sarcolemmal passive properties or on
ICa amplitude more than a slight expected
run-down (Fig. 5A). In Figure 5 the lack of IGF-1 effect in
the presence of genistein is compared with the profile of the
IGF-1-mediated ICa potentiation shown in Figure
1A (interrupted line). The magnitude of
ICa enhancement in single-pulse experiments is
illustrated in Figure 4B. ICa
traces in control (c) and after exposure to IGF-1
(filled circles) are used as a reference for Figure
5, C and D. C shows a slight decrease in ICa amplitude after exposure to genistein
plus IGF-1 (open circle), as compared with control
(c) (genistein alone). Peak ICa in
control and 15 min after exposure to IGF-1 (in the presence of
genistein) was 3.5 ± 0.11 and 2.9 ± 0.32 µA/µF
(n = 15; p > 0.5), respectively. To
ensure that the functional effect of genistein was not an structural
effect, we tested the inactive analog daidzein. Peak
ICa in control and 15 min after exposure to
IGF-1 (in the presence of 10 µM daidzein) was 3.5 ± 0.27 and 6.3 ± 0.35 µA/µF (n = 8;
p > 0.5), respectively. The involvement of tyrosine
kinase in current potentiation was confirmed by the enzyme inhibitor
herbimycin A. Peak ICa in control and 15 min after exposure to IGF-1 (in the presence of 2 µM
herbimycin A) was 4.1 ± 0.25 and 3.9 ± 0.32 µA/µF
(n = 8; p > 0.5), respectively.
Fig. 5.
Tyrosine kinase and PKC inhibitors preclude
IGF-1-induced potentiation of skeletal muscle
ICa. A, Lack of IGF-1 effect
on ICa amplitude after muscle fiber
incubation in an intracellular solution containing 100 µM
genistein (open circles) or 50 nM GF-109203X (filled diamonds) for 15-20 min before exposure
to IGF-1. The effect of IGF and genistein or GF-109203X are compared
with the effect of IGF-1 alone (dashed line, repeated
from Fig. 1A). B-D, ICa in control (c) and
after exposure to IGF-1 (filled circle), genistein (open circle), or GF-109203X
(filled diamond). The interrupted line indicates the baseline current.
[View Larger Version of this Image (13K GIF file)]
The blockade exerted by genistein and herbimycin A on IGF-1-mediated
ICa potentiation does not necessarily imply that
a direct tyrosine kinase-mediated phosphorylation of the L-type
Ca2+ channel operates in the whole muscle fiber.
Kinases located downstream in the signaling cascade may mediate IGF-1R
L-type Ca2+ channel 1 subunit interaction.
Because the experiments were performed in conditions of strong
myoplasmic Ca2+ chelation, the possibility of
Ca2+- and calmodulin-dependent kinases and
Ca2+-dependent PKC isoforms phosphorylation was
minimized.
IGF-1-evoked L-type Ca2+ potentiation is
mediated by PKC
We focused the following group of experiments on the elucidation
of a potential role of cAMP-dependent protein kinase and PKC in
inter-receptor signaling. It has been demonstrated that the compound
GF-109203X, used at low concentrations (10-50 nM), acts as
a specific PKC inhibitor. The IC50 is 10 nM for
PKC, 2 µM for cAMP-dependent protein kinase, and >60
µM for tyrosine kinase activity (Toullec et al., 1991).
Therefore, 50 nM GF-109203X was used to investigate a
PKC-mediated L-type Ca2+ channel phosphorylation.
Cells were preincubated with 50 nM GF-109203X for 10-15
min before IGF-1 was applied. Figure 5A shows the lack of
effect of IGF-1 when a muscle fiber was pretreated with the PKC
inhibitor (filled diamonds). Figure 5D
illustrates two pulses from a complete run of repetitive
depolarizations to 20 mV in control (c) and IGF-1-treated
tissues (filled diamond). The magnitude of the
decline in the current amplitude was no more than that expected for
run-down (see above). ICa amplitudes in control
and after exposure to IGF-1, both determined in the presence of
GF-109203X, were 3.9 ± 0.21 and 3.4 ± 0.28 µA/µF
(n = 10), respectively. Control
ICa in the presence of GF-109203X did not differ
significantly from the control in the absence of the compound. The
complete abolition of IGF-mediated ICa
enhancement suggests that Ca2+-independent PKC
isoforms phosphorylate the Ca2+ channel and that the
cAMP-dependent protein kinase is not involved in the inter-receptor
signaling cascade. This point was corroborated by using PKC 19-36, a
specific pseudosubstrate inhibitor of PKC with an IC50 for
PKC of 15 µM (House and Kemp, 1987). A muscle fiber was
incubated in 50 µM PKC(19-36) for 10-15 min before
exposure to IGF-1. ICa amplitudes in control and
IGF-1-treated fibers, both in the presence of the PKC inhibitor
peptide, were 4.1 ± 0.51 and 3.7 ± 0.48 µA/µF
(n = 7), respectively. Control experiments were
performed with the internal solution containing the PKC inhibitor vehicle alone (acetic hydroxide). The ratio between peak
ICa amplitude in the vehicle plus IGF-1 and peak
ICa in the vehicle alone was 1.9 ± 0.28 (n = 5), demonstrating that the vehicle, by itself, did
not inhibit PKC phosphorylation.
PKC activation enhances skeletal muscle
ICa
To determine whether PKC-induced phosphorylation is feasible in
adult whole mouse skeletal muscle fibers, the preparation used in these
studies, we performed a group of experiments that used phorbol esters.
Phorbol esters as PKC agonists have been used widely to promote
PKC-dependent ion channel phosphorylation in other cells (Yang and
Tsien, 1993). Figure 6A
shows the time course of ICa potentiation
induced by 50 nM PMA (filled circles). No
significant changes in the passive sarcolemmal properties were detected
throughout the experiment, as demonstrated by the lack of changes in
the capacitive transients recorded at the beginning and at the end of
the experiment (insets). The time course of the PMA effect
was very similar to that recorded in the presence of IGF-1 (see Fig.
1A). No current potentiation was recorded in the
presence of the vehicle (DMSO; data not shown). As a control, 1 µM 4-phorbol 12,13-didecanoate (4-PDD), an inactive
phorbol ester analog, was used. No ICa
potentiation was detected in the presence of 4-PDD, as shown in
Figure 6A (filled triangles). The
change in ICa amplitude induced by 4-PDD, as
compared with control (without the drug), was 0.96 ± 0.11 (n = 5). The vehicle (DMSO) did not modify calcium
current amplitude (data not shown). The PMA effect on
ICa amplitude decreased with time. At 50 min after the start of the incubation in 50 nM PMA, peak
ICa was 0.98 ± 0.11 (n = 6) of the control. The reduction in current amplitude may reflect PKC
degradation and the requirement for rephosphorylation to maintain a
higher level of channel activity. To determine whether PKC degradation
accounts for the decline in PMA effect on ICa amplitude, we tested IGF-1 in the same preparation. PMA was washed out
several times, and the muscle fiber was incubated in 20 ng/ml IGF-1.
Peak ICa in IGF-1 was 0.97 ± 0.09 (n = 6) of the control (in PMA; p > 0.5).
Fig. 6.
Time course of PKC-dependent phosphorylation of
the L-type Ca2+ channel 1 subunit.
A, PMA-induced ICa
potentiation (filled circles) and lack of effect
of the inactive 4-PDD analog (filled
triangle). Both insets illustrate the linear
capacitive transient current in response to hyperpolarizations to 110
mV at time 0 and at 15 min of exposure to PMA.
ICa in control (B) and
in PMA (C; after 15 min exposure) were elicited by 250 msec depolarizations from a Vh of 90 mV,
with 10 mV increments. The interrupted line indicates the baseline current.
[View Larger Version of this Image (22K GIF file)]
Figure 6, B and C, illustrates the voltage
dependence of PMA-induced ICa potentiation.
Similar to the effect induced by IGF-1, a shift in 15 mV of the
Ica-V relationship was recorded in
the presence of PMA. V1/2 value was 17.3 ± 2.2 and 32.2 ± 4.1 mV in control and PMA, respectively. The
shift in V1/2 was not associated with changes in
the voltage distribution for the charge movement as shown for IGF-1.
V1/2Q in control and PMA was 44.3 ± 2.7 and 43.5 ± 2.9 mV, respectively (n = 10).
Phosphorylation of the L-type Ca2+ channel 1
subunit dependence on IGF-1R activation
Biochemical experiments were performed to ascertain whether
tyrosine kinase- and PKC-dependent phosphorylation underlies the ICa potentiation in response to IGF-1R
activation described above. To this aim, we developed a technique to
induce phosphorylation of the L-type Ca2+ channel
via IGF-1R activation in mature skeletal muscle fibers. The
preservation of functionally viable muscle fibers was a key factor to
explore the IGF-1R-Ca2+ channel signaling. Living
short monolayers of adult skeletal muscle fibers were exposed to an
ATP-depleting solution, loaded with [-32P]ATP, and
subsequently stimulated with IGF-1 (see Materials and Methods). Figure
7 shows a 5-15% SDS polyacrylamide gel
in which the high molecular weight standard (A) and
100 µg of muscle proteins (B) have been loaded.
Figure 7C is a Western blot analysis of muscle proteins with
a monoclonal antibody specific for the L-type Ca2+
channel 1 subunit (see Materials and Methods). A single band at 165 kDa demonstrates the specificity of the antibody against the 1
subunit. Figure 6D shows the lack of the 165 kDa band
in the supernatant after immunoprecipitation of the
Ca2+ channel 1 subunit is induced with the
monoclonal antibody. Figure 7E corresponds to an
autoradiography of the gel that shows protein phosphorylation in the
pellet (immunoprecipitation) 15 min after the addition of 20 ng/ml
IGF-1. These results were repeated in five muscle preparations from
five mice. These studies demonstrate that IGF-1 induces phosphorylation
of the L-type Ca2+ channel 1 subunit.
Fig. 7.
IGF-1-dependent phosphorylation of the
skeletal muscle L-type Ca2+ channel 1 subunit.
A, High molecular weight standard (42,699-200,000). B, SDS polyacrylamide gradient gel (5-15%)
electrophoresis of 100 µg of skeletal muscle proteins.
C, Western blot analysis of the L-type
Ca2+ channel 1 subunit, using a
monoclonal antibody. D, Electrophoresis of the muscle
proteins supernatant after immunoprecipitation of the L-type
Ca2+ channel 1 subunit.
E, Autoradiography of the pellet after IGF-1-induced phosphorylation and immunoprecipitation of the L-type
Ca2+ channel 1 subunit.
[View Larger Version of this Image (48K GIF file)]
Figure 8A shows the
time course of the L-type Ca2+ channel 1 subunit
phosphorylation. At time 0, a light band can be detected that
represents a phosphorylation independent of the activation of the
IGF-1R cascade. At 5 min of exposure to 20 ng/ml IGF-1, the signal
increased and reached a peak at 15 min. At 30 min the signal slightly
decreased. To quantitate the time course and magnitude of the L-type
Ca2+ channel phosphorylation, we scanned
autoradiographs, and the signals were expressed as the integral of the
optical density corresponding to each band (see Materials and Methods).
The magnitudes of the channel phosphorylation (in optical density, O.D. × mm2) were 0.31 ± 0.02, 0.63 ± 0.05, 2.33 ± 0.16, and 2.31 ± 0.21 at 0, 5, 15, and 30 min of
exposure to IGF-1, respectively. Differences in the optical density
were significant when zero, 5, and 15-30 min points were compared
(paired t test, n = 6; p < 0.005; one-way ANOVA, p = 0.005). However, differences
between signals at 15 and 30 min were not statistically significant
(ANOVA, p > 0.5). The lack of statistically
significant differences at 15 and 30 min was confirmed by a
Student-Newman-Keuls test. The ratio of the Ca2+
channel phosphorylation at the peak of IGF-1 activity (15-30 min) over
control (time 0) was 7.5 ± 0.52 (n = 5). These
results are in close agreement with the time course of the
ICa potentiation described above (see Fig.
1A).
Fig. 8.
L-type Ca2+ channel 1
subunit phosphorylation dependence on IGF-1R activation. Optical
scanning of autoradiographs depicting phosphorylation of the 165 kDa
band corresponding to the L-type Ca2+ channel
1 subunit. Phosphorylation was induced with 20 ng/ml IGF-1 on monolayers of adult fast-twitch skeletal muscles fibers incubated in 50 µM [-32P]ATP (specific
activity of 10 Ci/mmol) and electrophoresed on SDS polyacrylamide gels.
A, Time course of the L-type Ca2+
channel 1 subunit phosphorylation in control (time 0, before adding IGF-1) and 5, 15, and 30 min after adding 20 ng/ml IGF-1 to the bathing solution. B, Phosphorylation of the
L-type Ca2+ channel 1 subunit after
15 min incubation in 20 ng/ml IGF-1 and inhibition of IGF-1-dependent
channel phosphorylation by preincubation in 100 µM
genistein for 15 min. C, IGF-1-dependent phosphorylation of the L-type Ca2+ channel 1 subunit,
inhibition of channel phosphorylation by preincubation in 50 µM protein kinase C inhibitor peptide 19-36 for 15 min,
and channel phosphorylation after incubation in PMA for 15 min.
[View Larger Version of this Image (62K GIF file)]
To explore further the involvement of tyrosine kinase in L-type
Ca2+ channel 1 subunit phosphorylation, we
preincubated muscle fibers in 100 µM genistein for 15-20
min. Figure 8B shows an almost complete inhibition of
the IGF-1-dependent Ca2+ channel phosphorylation by
genistein. The ratio of the IGF-1-induced phosphorylation over
IGF-1-induced phosphorylation in cells preincubated in genistein was
4.4 ± 0.22 (n = 5). These results provide support for the concept that the activation of tyrosine kinase phosphorylation cascade is required to enhance ICa through the
dihydropyridine-sensitive Ca2+ channel.
The involvement of Ca2+-independent PKC isoform(s)
has been supported by a complete suppression of the IGF-1-dependent
ICa potentiation by PKC inhibitors (see Fig. 5).
Figure 8C shows a strong inhibition of the 165 kDa band
phosphorylation by preincubation in the PKC inhibitor peptide 19-36. The ratio of the IGF-1-induced phosphorylation over IGF-1-induced
phosphorylation in cells preincubated in the PKC inhibitor peptide
19-36 was 4.3 ± 0.31 (n = 5). The phosphorylation remaining after the PKC inhibitor has been used may correspond to
channel phosphorylation through IGF-1-independent pathways. Figure
8C shows the effect of the PKC activator PMA on L-type Ca2+ channel 1 subunit phosphorylation. The ratio
of the PMA-induced phosphorylation over IGF-1-induced phosphorylation
in cells preincubated in the PKC inhibitor peptide 19-36 was 5.1 ± 0.41 (n = 5). These experiments also demonstrate
that the molecule phosphorylated in response to IGF-1R activation
corresponds to that phosphorylated by the phorbol ester PMA. An
additional series of experiments did not support a role for
cAMP-dependent protein kinase in IGF-1 signaling. The ratio of the
phosphorylation in control (before adding IGF-1) over IGF-1-induced
phosphorylation in cells preincubated with the PKC inhibitor peptide
19-36 was 0.96 ± 0.05 (n = 6). The cAMP-dependent
protein kinase A inhibitor 5-24 (IC50, 0.8 µM; Scott et al., 1985) did not inhibit further the
effect of the PKC inhibitor peptide 19-36 on IGF-1-induced
phosphorylation. The ratio of the IGF-1-induced phosphorylation in PKC
inhibitor over phosphorylation in PKC plus 10 µM PKA
inhibitor was 0.89 ± 0.18. Moreover, the magnitude of inhibition
of the IGF-1-mediated Ca2+ channel phosphorylation
with the cAMP-dependent protein kinase A inhibitor 5-24 was not
statistically significant. The ratio of the IGF-1-dependent
phosphorylation over phosphorylation in cells preincubated in 10 µM PKA inhibitor was 0.95 ± 0.12 (n = 8).
In summary, the interaction between the L-type Ca2+
channel and the IGF-1R is not mediated primarily by tyrosine kinase.
Other protein kinases such as Ca2+-independent PKC
isoforms are activated secondarily, promoting phosphorylation of the
skeletal muscle L-type Ca2+ channel 1
subunit.
DISCUSSION
In this work we found that IGF-1R modulates the skeletal muscle
L-type Ca2+ channel 1 subunit via a
phosphorylation mechanism involving a tyrosine kinase and
Ca2+-independent PKC isoform(s). The
dihydropyridine-sensitive L-type Ca2+ channel is the
only high-voltage-activated Ca2+ channel expressed
in adult mammalian muscle. Thus, the IGF-1-mediated ICa potentiation recorded in single skeletal
muscle corresponds to the enhancement of multiple units of the same
Ca2+ channel class (dihydropyridine-sensitive,
L-type). IGF-1 induced a 13 mV shift in the
Ica-V relationship toward more
negative potentials. Potentiation and changes in the voltage dependence
of ICa were not associated with alterations in
the maximum charge movement and voltage distribution. A plausible
mechanism that may account for these results is that L-type
Ca2+ channel 1 subunit
phosphorylation facilitates relatively uncharged transitions leading to
the opening of the channel that occur after the movement of the
voltage-sensing particles.
The interaction between IGF-1R and L-type Ca2+
channel in skeletal muscle and native neurons, cells in which IGF-1
exerts potent trophic and developmental effects, has not been explored.
A tyrosine kinase-dependent phosphorylation of the L-type
Ca2+ channel in response to IGF-1 stimulation has
been reported in clonal pituitary and neuronal cells (Kleppisch et al.,
1992; Selinfreund and Blair, 1994). However, the lack of consensus
sequence in the L-type Ca2+ channel for tyrosine
kinase phosphorylation suggested to us that IGF-1R exerts a modulatory
effect on the Ca2+ channel 1 subunit through an
indirect pathway, as demonstrated for skeletal muscle in the present
study.
Other trophic factors such as insulin and epidermal growth factor
induce a long-term potentiation of Ca2+ channels in
clonal pituitary cells (Meza et al., 1994). Despite the fact that the
mechanism for this long-term effect is not clear, the increase in
ICa density suggests alterations in gene
expression or changes in L-type Ca2+ turnover. The
long-term modulation clearly differs from the rapid regulation exerted
by IGF-1R activation on the L-type Ca2+ channel
described in this work. Unfortunately, IGF-1-dependent long-term
modulation of Ca2+ channels cannot be explored in
mature skeletal muscle, because cells in culture undergo a process of
dedifferentiation.
Phosphorylation sites for PKC, cAMP-dependent, and
Ca2+-calmodulin-dependent protein kinase in the
L-type Ca2+ channel 1 and  subunits have been identified (Jahn et al., 1988; O'Callahan et al.,
1988; Chang et al., 1991; Rotman et al., 1995), but not for tyrosine
kinase. In this work we found no modulatory effect on L-type
Ca2+ channels when PKC was inhibited, supporting the
concept that the channel is not modulated directly by tyrosine kinase.
Although it is not known if the channel is phosphorylated on ser/thr or tyr residues, it is likely that autophosphorylated IGF-1R activates a
second messenger such as PKC via phospholipase C- (Nishibe et al.,
1990; Nishizuka, 1995), and PKC phosphorylates the L-type Ca2+ channel. In summary, our results differ from
previous studies in which Ca2+ channels are
modulated in terms of seconds to a few minutes and require a sequential
activation of two protein kinases.
It has been demonstrated that purified and reconstituted
Ca2+ channels are activated by phosphorylation of
their subunits (Flockerzi et al., 1986; Hymel et al., 1988; Nunoki et
al., 1989). Although Ca2+ channels can be
phosphorylated in colchicine-treated myotubes (Sculptoreanu et al.,
1993) or in membranes extracted from mature muscle, in the present
study channel phosphorylation has been induced by activation of the
tyrosine kinase-linked IGF-1 receptor in vivo and in adult
fast-twitch muscle, before tissue grinding.
Cultured myotubes from embryonic rat skeletal muscles exhibit a
voltage- and frequency-dependent potentiation of L-type
Ca2+ currents that is caused by a left shift in the
voltage dependence of channel activation toward more negative
potentials and that requires cyclic AMP-dependent protein kinase
phosphorylation (Sculptoreanu et al., 1993). These investigations
reported that phosphorylation induced a potentiation of the L-type
Ca2+ current at negative voltages, which is similar
to the results of our studies. However, some differences between both
works have to be emphasized: (1) we used freshly dissociated muscle
fibers from 14-month-old rats instead of colchicine-treated cultured myotubes, (2) we induced current potentiation by IGF-1R activation instead of using high rates of sarcolemmal depolarization, (3) the
frequency of fiber stimulation used in the present work (one pulse
every 30 sec) did not induce current potentiation (as shown in Fig.
1A, before exposure to IGF-1) and did not enhance
Ca2+ current in myotubes, and (4) the use of a
pseudosubstrate inhibitor of PKC did not occlude the
stimulation-dependent ICa potentiation in
myotubes in contrast to a complete inhibition of the current potentiation in adult muscle fibers. In summary, PKC- and
cAMP-dependent protein kinase participate as second messengers in cell
signaling, depending on the specific trigger. An example of this is the
L-type Ca2+ channel phosphorylation by
cAMP-dependent protein kinase in response to high-frequency sarcolemmal
stimulation or by PKC in response to IGF-1R activation.
Although the effect of ICa enhancement on
sarcolemmal excitation-sarcoplasmic reticulum Ca2+
release-contraction coupling has been excluded during single twitches,
it is likely that during prolonged or repetitive depolarizations current enhancement has a role (for review, see Melzer et al., 1995).
This theory is based on a series of reports supporting a
phosphorylation-dependent potentiation of contractile force and on the
detrimental effect of L-type Ca2+ channel
antagonists during prolonged muscle activation (see above). Also, it is
likely that the increase in Ca2+ influx promoted by
IGF-1R activation is related to sarcolemmal-nuclear signaling and
ultimately to gene expression.
The development of a technique to study the incorporation of
radiolabeled ATP into the L-type Ca2+ channel 1
subunit in living adult muscle fibers provides important evidence for a
role of tyrosine kinase and PKC on Ca2+ channel
phosphorylation. Two Ca2+-dependent (PKC- and
PKC-) and two Ca2+-independent (PKC- and
PKC- ) PKC isoforms have been identified in rat skeletal muscles
(Yamada et al., 1995). In our recording conditions only
Ca2+-independent kinases potentially are involved in
this signaling cascade, because high Ca2+ buffer
concentrations completely prevent increases in myoplasmic Ca2+ concentration. Therefore, the participation of
the Ca2+-dependent PKC isoforms (PKC- and
PKC-) cannot be ruled out in physiological conditions. To elucidate
the role of Ca2+-dependent PKC isoforms in ion
channel phosphorylation, experiments in stretched fibers and free
Ca2+ movement are required. It has been demonstrated
that insulin increases the membrane-associated immunoreactive
Ca2+-independent PKC isoforms in rat muscles (Yamada
et al., 1995). Because PKC- expression was found to be restricted to
the sarcolemmal fraction in the neuromuscular junction (Hilgenberg and
Miles, 1995), it is unlikely that this PKC isoform phosphorylates the L-type Ca2+ channel, which is widely expressed at
the T tubule membrane. Thus, it is likely that the PKC- isoform is
involved in Ca2+ channel 1 subunit
phosphorylation on IGF-1R activation. Although the L-type
Ca2+ channel 1 subunit is phosphorylated by PKC,
phosphorylation by secondarily activated kinases (i.e., tyrosine
kinase) cannot be ruled out completely.
In summary, this work demonstrates that, in addition to long-term
effects of IGF-1 on skeletal muscle, a direct modulation of the L-type
Ca2+ channel operates in matured tissue in a shorter
time period via a phosphorylation mechanism involving tyrosine kinase
and PKC.
FOOTNOTES
Received March 27, 1997; revised June 30, 1997; accepted July 7, 1997.
This work was supported by National Institutes of Health/National
Institute on Aging Grants K01 AG00692, R29AG13934, P60-AG 10484, and
T-32 AG00182 and indirectly from the Muscular Dystrophy Association. We
are very grateful to Dr. William E. Sonntag for helpful comments on
this work.
Correspondence should be addressed to Dr. Osvaldo Delbono, Department
of Internal Medicine, Gerontology, The Bowman Gray School of Medicine,
Medical Center Boulevard, Winston-Salem, NC 27157.
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