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Short-Term Regulation of Excitation-Contraction Coupling by the ß_1a Subunit in Adult Mouse Skeletal Muscle

María C. García ^*, Elba Carrillo ^*, José M. Galindo ^*, Ascensión Hernández ^*, Julio A. Copello , Michael Fill  and Jorge A. Sánchez ^*

^* Departmento de Farmacología, Centro de Investigación y de Estudios Avanzados del I.P.N., Mexico, D.F. 07360, Mexico; Department of Pharmacology, Southern Illinois University, Springfield, Illinois; and Department of Physiology, Loyola University Chicago, Maywood, Illinois

Correspondence: Address reprint requests to Dr. Jorge A. Sánchez, Dept. of Pharmacology, Cinvestav Apartado Postal 14-740, Mexico, D.F. 07360. Tel.: 52-55-5061-3301; Fax: 52-55-5577-7090; E-mail: jsanchez{at}cinvestav.mx.

	ABSTRACT

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
The ß_1a subunit of the skeletal muscle voltage-gated Ca²⁺ channel plays a fundamental role in the targeting of the channel to the tubular system as well as in channel function. To determine whether this cytosolic auxiliary subunit is also a regulatory protein of Ca²⁺ release from the sarcoplasmic reticulum in vivo, we pressure-injected the ß_1a subunit into intact adult mouse muscle fibers and recorded, with Fluo-3 AM, the intracellular Ca²⁺ signal induced by the action potential. We found that the ß_1a subunit significantly increased, within minutes, the amplitude of Ca²⁺ release without major changes in its time course. ß_1a subunits with the carboxy-terminus region deleted did not show an effect on Ca²⁺ release. The possibility that potentiation of Ca²⁺ release is due to a direct interaction between the ß_1a subunit and the ryanodine receptor was ruled out by bilayer experiments of RyR1 single-channel currents and also by Ca²⁺ flux experiments. Our data suggest that the ß_1a subunit is capable of regulating E-C coupling in the short term and that the integrity of the carboxy-terminus region is essential for its modulatory effect.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Dihydropyridine receptors (DHP) play an essential role as the voltage sensors that link excitation with contraction in skeletal muscle (for reviews, see Ríos and Pizarro and others 1–4). In addition to their role as voltage sensors, these receptors are also permeant to Ca²⁺, generating a very slowly activated L-type Ca²⁺ current ((5) for a review, see Melzer et al. (4)). DHP receptors are composed of the main subunit _1s, and the auxiliary subunits, ₂-, ß₁, and -subunits. The _1s subunit (now referred to as the Ca_V1.1 channel (6)) contains the pore (7), the voltage sensor of excitation-contraction coupling, and the DHP binding sites (for reviews, see Ríos and Pizarro and others (1,8–10)).

The ß_1a subunit is the main isoform among the ß₁ subunits present in skeletal muscle (11) and its role has been explored by inactivating the ß₁ gene using gene targeting techniques. This approach has revealed that this subunit plays an essential role in the assembly of DHP receptors in their correct position. ß-null cells have undetectable levels of the _1s subunit in the cell membrane and show greatly reduced L-type Ca²⁺ currents and charge movement (12,13), an effect that can be reversed by the introduction of the cDNA of the ß_1a subunit (14). However, much less is known about the potential role of this auxiliary subunit as a modulatory protein of E-C coupling, regulating Ca²⁺ release in the short term. In this regard, we have recently described that the addition of the ß_1a subunit to a cell-free preparation enhances the amplitude of L-type current within minutes, consistent with a short-term modulatory role on Ca_V1.1 channels (15). The aim of this study was to examine whether the ß_1a subunit also has a short-term effect on Ca²⁺ release induced by action potentials in adult skeletal muscle fibers.

Preliminary results have been published (16).

	MATERIALS AND METHODS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Fiber preparation
BALB/c mice (aged 9 weeks) were used. These were killed by cervical dislocation, after which the flexor digitorum brevis muscles (FDB) of the hindlimbs were isolated and incubated at 34°C for 60 min in a Ca²⁺-Mg²⁺-free Tyrode solution plus 10% fetal calf serum (Gibco-Invitrogen, Carlsbad, CA) and collagenase (0.5 mg/ml, type lV, Sigma, St. Louis, MO). The muscles were then rinsed and dissociated by gently triturating the enzyme-treated muscles through a fire-polished Pasteur pipette with collagenase-free Tyrode solution containing (mM): NaCl, 146; KCl, 5; CaCl₂, 2; MgCl₂, 1; glucose, 11; HEPES, 10; at pH = 7.4. The experiments were performed according to the guidelines of the local animal care committee.

Molecular biology techniques
Production of the ß_1a subunit in COS-1 cells
We have described the expression and detailed purification procedure of the ß_1a subunit in COS-1 cells elsewhere (15). In brief, COS-1 cells were transiently transfected with a plasmid containing the cDNA of the ß_1a subunit (pSG5-Mß_1a) under the control of the SV-40 promoter. Membrane fraction samples of homogeneized COS-1 cells were loaded onto polyacrylamide gels and stained with Coomassie blue. The gel band that contained the ß_1a subunit was identified by Western blots, cut, and purified by electroelution methods according to Smith (17). After the electroelution procedure, electrodialysis techniques were used to remove all traces of sodium dodecyl sulfate, as described by García et al. (15). The dried protein was stored at –20°C until used.

Bacterial production and purification of the ß_1a subunit
The coding fragment of the ß_1a subunit was liberated from the pSG5 expression vector mentioned above. To this end, pSG5-ß_1a was cleaved with NheI (New England Biolabs, Beverly, MA). NheI sites are present in both ends of the ß_1a fragment. The sticky ends of the fragment were filled with Klenow polymerase and ligated at the SmaI site of the multiple cloning site of the pQE-31 vector (Qiagen, Valencia, CA). This strategy generates an in-frame cloning of the ß_1a protein. Escherichia coli DH5- cells were transformed with this construct and clones with the correct orientation of the insert were selected to achieve transcription of the product with the T5 promoter present in the vector. The deletion of the last 40 amino acids in the carboxy-terminus region of the ß_1a subunit was achieved by cleavage of pSG5-ß_1a plasmid with NotI. The plasmid contains a single recognition site located downstream from the open reading frame. After that, DNA was treated with Bal-31 exonuclease (New England Biolabs) for 15 min. To estimate the size of the deleted products, we used NcoI and HincII to liberate the 3' end of the ß_1a subunit and the products were subsequently sequenced. The fragment containing the truncated ß_1a was liberated from the pSG5 vector by NheI and HincII (New England Biolabs). It was then filled with Klenow polymerase and subcloned in the same way as the nontruncated ß_1a subunit. Both clones were sequenced to confirm the integrity of the open reading frame with Sequenase kit (version 2.0, USB) and specific oligonucleotides for pQE vectors. Deletion of 40 amino acids in the C-terminus region of the ß-subunit is not expected to alter the folding of the protein. In fact, Bogdanov et al. (18) produced chimeras of ß-subunits lacking more amino acids in their C-terminus region than in this study and suggested that no disruption in protein folding takes place. In addition, molecular models of ß-subunits have identified several domains that allow intramolecular interactions, namely, SH3 domain, PDZ domain, and the GK domain (19,20). None of these domains is located in the C-terminus region of the ß-protein and the domain organization of D2 and D4 domains is not altered by a similar C-terminus deletion (21).

E. coli BL21(DE3) (Novagen, San Diego, CA) cells were transformed with the above constructs and used for mass production of the ß_1a subunit protein and its truncated form. The cells were cultured in 100 ml of LB medium in the presence of 100 µg/ml ampicillin and grown at 32°C until an A₆₀₀ of 0.6 was reached. The production of the protein was induced by 1.0 mM isopropyl ß-D-thiogalactopyranoside for 6 h at 32°C. The cells were pelleted and resuspended in 10 ml of NETN buffer (NaCl = 120 mM, Tris-HCl = 20 mM, EDTA = 1 mM, PSMF = 1 mM, and 0.5% NP-40 at pH = 8.0). Cell lysis was achieved with the aid of a French press (Thermo Spectronic, Shelton, CT) at a pressure of 600 psi, applied twice. The lysate was centrifuged at 15,000 x g resulting in a pellet and in a clear supernatant fraction. The ß_1a subunit protein was found in the pellet fraction and was solubilized with 5 M urea. The truncated form of the ß_1a subunit was found in the supernatant. The protein fractions were analyzed by SDS-PAGE and stained with Coomassie blue. The ß_1a subunit was identified by Western blot as described elsewhere (15). The products were obtained by electroelution techniques and the eluted protein products were precipitated with acetone. Thereafter, the samples were dialyzed with water to remove the salt content.

Pressure injection techniques
To introduce the ß_1a subunit into muscle fibers, we used an intracellular micropipette to which pressure was applied. The micropipettes were pulled in a Brown-Flaming horizontal puller (Sutter Instruments, San Francisco, CA) from Kwik-fill glass capillaries (WPI, New Haven, CT) and had an average resistance of 25 M when filled with 3 M KCl. They were mounted in a plastic holder and driven by a hydraulic micromanipulator (Narishige MO-150, Tokyo, Japan). Pipettes were filled with a solution containing (mM): KCl, 140; MgCl_2, 1; EGTA, 1; and HEPES, 10, at pH = 7.1. Fibers were impaled at a point located 125 µm from the position where light was detected with the photodiode. The ß_1a subunit, purified either from COS-1 cells or from bacteria, was used at a concentration of 0.35 µg/µl. Most experiments were performed with ß_1a subunits purified from COS-1 cells. Experiments that were performed with ß_1a subunits from bacteria are clearly indicated in the text. Control experiments involved pressure injection of a heat-inactivated ß_1a subunit added at the same concentration or pressure injection of the micropipette saline solution. In both cases, these control injections gave similar results and are considered together in the Results section. Two trains of pulses separated by a 30-s interval were delivered. Each train consisted of five consecutive 150 kPa pressure pulses, each lasting 600 ms. We estimate that this procedure allowed the injection of 40–50 pl of solution. Action potentials were elicited by passing rectangular current pulses between two platinum plate electrodes placed symmetrically on either side of the muscle chamber.

The microinjected ß_1a subunit is expected to diffuse away from the point of injection along the axis of the muscle fiber. To estimate the diffusion of the ß_1a subunit in our experiments, we used Fick's second law (Eq. 1). This equation was used by Papadopoulos et al. (22) to account for the axial spread of microinjected proteins in muscle. In Eq. 1, D is the diffusion coefficient and C is the concentration of the injected protein in the sarcoplasm that changes as a function of time (t) and as a function of the distance from the point of injection (x). Equation 1 describes the proportionality between the change of C along the diffusion pathway C/x and the change in concentration with time C/T. This equation assumes a one-dimensional diffusion process with an infinite extension of the diffusion path.

(1)

The diffusion coefficient (D) in Eq. 1 was measured experimentally in the myoplasm of muscle fibers by Papadopoulos et al. (22) for proteins of different masses. In our estimations we used D = 6.2 x 10^–8 cm² s^–1. This value corresponds to the diffusion coefficient measured for hemoglobin (22), a protein that has a similar mass (64.5 kDa) to that of the ß_1a subunit (55 kDa (10)). The analytical solution of Eq. 1 can be achieved using a Dirac delta function applied at t = 0. The solution is Eq. 2 where A is an amplitude factor.

(2)

Optical techniques
We used Fluo-3 AM (1–10 µM) (Molecular Probes, Eugene, OR) to monitor the levels of intracellular Ca²⁺. This dye undergoes large fluorescence changes upon Ca²⁺ binding, it has a large dynamic range, low compartmentalization (23), and it has been used extensively in muscle (24–26). Fibers were mounted in a chamber placed on the stage of an Optiphot microscope (Nikon, Tokyo, Japan). The fluorescence emitted by a preselected region of a stained muscle fiber, illuminated episcopically with monochromatic light at a wavelength of 485 nm, was filtered with a high-pass barrier filter (cut-on wavelength 535 nm), and detected with a low noise photodiode connected in a photovoltaic configuration. The basal fluorescence (F) from the same region of the muscle fiber was recorded continuously on video tape. Its mean value during 300 ms before electrical stimulation, was used to scale Ca²⁺ signals as F/F. This procedure minimizes the possible effects of changes in the concentration of the dye on fluorescence signals and it has been used extensively by others (24,27). No attempts were made to calculate the actual myoplasmic Ca²⁺ concentration. To prevent mechanical artifacts, intact single fibers were suspended in 0.35% agar gel, following a procedure similar to that described in (28) except that we used a lower agar concentration and that our experiments were performed at a lower temperature (20–22°C).

Ca²⁺ leak measurements
Skeletal muscle heavy sarcoplasmic reticulum (SR) microsomes were obtained following procedures by Saito et al. (29). The net rate of Ca²⁺ leak from these microsomes was measured at room temperature (20–24°C) with a spectrophotometer (Cory 50, Varian, Palo Alto, CA) using the Ca²⁺-sensitive dye antipyrylazo III (APIII). Changes in Ca²⁺ concentration over time were measured as the absorbance difference between 710 and 790 nm as described in (30). Briefly, microsomes (50 µg protein) were incubated with 1 ml of assay solution containing 100 mM potassium phosphate, 0.2 mM APIII, 4 mM MgCl₂, and 2 mM ATP (pH 7). Microsomes were actively preloaded with Ca²⁺ (via the SR Ca²⁺ ATPase) by adding 3 aliquots of 40 nM CaCl₂ while stirring. The microsomes were then incubated for 5 min with standard buffer (control) or buffer containing ruthenium red (5 µM), normal ß_1a subunit, or the truncated ß_1a subunit. After this incubation period, 25 µM cyclopiazonic acid (CPZ) was added to inhibit the SR Ca²⁺ ATPase and then Ca²⁺ leak from the microsomes was monitored. When caffeine (2.5 mM) was applied, it was applied simultaneously with CPZ. Ca²⁺ leak rate in each experimental condition was measured from the slope of the APIII Ca²⁺ signal.

Electrophysiological methods
Dissociated FDB muscle fibers were used in voltage clamp experiments. To minimize mechanical artifacts during measurements of membrane currents, fibers were not embedded in agar as in optical experiments. Instead, movement artifacts were greatly suppressed by a previous incubation for 3 h in the cell permeant calcium buffer BAPTA AM (10 mM) (Molecular Probes).

The whole-cell patch-clamp technique was used to record Ca_V1.1 currents (31). Pipettes (1–1.2 M) were double-pulled from hard glass (KIMAX-51; Kimble Glass, Toledo, OH) and were filled with 5 µl of the internal solution. Ca_V1.1 currents were measured 10 min after achieving the whole cell configuration. The ß_1a subunit was tested by adding 0.30–0.35 µg µl^–1 of the ß_1a subunit to the pipette solution. Currents were recorded with an Axopatch 200A (Axon Instruments, Foster City, CA) amplifier; 60–80% of the series resistance was electronically compensated.

To measure charge movement, command pulses of 60-ms duration and variable amplitude were delivered. The pulse sequence was bracketed by 16 consecutive hyperpolarizing control pulses, –20 mV from the holding potential (E_h) that was set at –100 mV. The currents generated during these pulses were used to subtract linear membrane components, to calculate the linear membrane capacitance, and to measure the leakage current during the experiment.

The voltage dependence of activation of nonlinear charge movement was fitted to the Boltzmann function:

(3)

where Q_max is the maximum value of charge, V_m is the membrane potential, V is the potential where Q = 0.5 Q_max, and k is a measure of the steepness of the curve.

To measure Ca²⁺ currents (I_Ca) the same pulse protocol was used except that the duration of the pulses was 750 ms and E_h = –80 mV. The interval between pulses was 4 s. The peak Ca²⁺ current values were fitted to Eq. 4, which is similar to that used by Wang et al. (32) to describe the current-voltage relationship of L-type Ca²⁺ currents in muscle fibers.

(4)

In Eq. 4, G_max is the maximum conductance and V_rev is the reversal potential. The other parameters have the same meaning as in Eq. 3.

Single channel measurements of skeletal RyR1 channels were recorded by fusing SR microsomes into artificial planar lipid bilayers as previously described by Copello et al. (33). Briefly, planar bilayers were formed by painting a mixture (5:4:1) of phosphatydilethanolamine, phosphatydilserine, and phosphatydilcholine (50 mg/ml decane) across a 100-µm hole separating two 1-ml solutions. One solution (trans) contained 250 mM HEPES/50 mM Ca(OH)₂ (pH 7.4) and was clamped to 0 mV using an Axopatch 200B patch-clamp amplifier (Axon Instruments). The other solution (cis) was held at virtual ground and contained 1 mM CsCl, 250 mM HEPES/Tris (pH 7.4), and 1 mM CaCl₂. The SR microsomes (5 µg protein) were added to the cis solution while stirring. In this situation, the RyR1 channels incorporate with their cytosolic surface facing the cis solution. Channels were identified by their high conductance, Ca²⁺ selectivity, and gating characteristics. After RyR1 incorporation, the cis solution was perfused with 30 vol of 250 mM HEPES/Tris (pH 7.4) and the free Ca²⁺ level was adjusted to 2 µM (1.4 mM CaCl₂, 1 mM BAPTA, and 1 mM DiBromoBAPTA). The cis solution also contained 7 mM ATP and 5.5 mM Mg²⁺. Single-channel recordings were made before and after addition of ß_1a subunit (1–4 µM; stirring for 2 min) to the cis solution. Recordings were filtered at 1000 Hz and digitized at 20 KHz with a Digidata 1360 acquisition system (Axon Instruments). Data were analyzed using pClamp9 (Axon Instruments). Open probability (Po) was determined from at least 8 min of recording.

Electrophysiological and optical experiments were carried out at room temperature (20–22°C).

Solutions
The external solution employed to record I_Ca contained (mM): 10 Ca²⁺, 140 TEACH₃SO₃ (tetraethylamonium methanesulphonate), and 2 MgCl₂. The pipette solution contained (mM): 140 Cs-aspartate, 5 MgCl₂, and 10 EGTA. The composition of the external solution used to record charge movement was similar, except that the concentration of external Ca²⁺ was reduced to 1 mM. The presence of Ca²⁺ was required to maintain the stability of our recordings because fibers do not generally tolerate the absence of Ca²⁺ well. We did not use Ca²⁺ channel blockers because they alter the voltage dependence of charge movement and may have deleterious effects on leakage currents (34). Due to the contamination of "off" charge by tail Ca²⁺ currents, we restricted our measurements to "on" charge only. Contamination of "on" charge by I_Ca is likely to be minor because charge moves at more negative potentials than I_Ca and the time course of activation of I_Ca is very slow compared to that of "on" charge (32). In fact, it has been shown that integration of "on" charge provides an accurate measure of charge movement in Ca²⁺-contaning solutions, even without Ca²⁺ channel blockers (35).

Extracellular and intracellular solutions were buffered with HEPES (10 mM) at pH 7.2 and 7.1, respectively. Chemicals were obtained from either Sigma Chemical or Aldrich Chemical (St. Louis, MO).

The fitting of numerical formulas to experimental data employed a nonlinear, least squares algorithm. Parameter values given in the text are expressed as mean ± SE. Student's t-test was used at the level p < 0.05 to calculate statistical significance of the data.

	RESULTS

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Actions of the ß_1a subunit on intracellular Ca²⁺ signals
Fig. 1 A shows Ca²⁺ transients from a control experiment recorded from a single FDB skeletal muscle fiber. The recorded region was located 125 µm from the point of injection. Illustrated are Ca²⁺ signals recorded before and after the injection of a control solution (at t = 0). The change in fluorescence of the dye is expressed as a F/F ratio. The transient increase in Fluo-3 fluorescence induced by an action potential rapidly reaches a peak and decays quickly initially and then, more slowly to prestimulus levels. The fiber was stimulated extracellularly at the times indicated (in minutes) by the numbers above each trace. The Ca²⁺ transients were quite similar before and after pressure injection indicating that the injection, by itself, had only minor effects on Ca²⁺ release. A very different result was observed when the ß_1a subunit was injected. The ß_1a subunit had remarkable effects on Ca²⁺ signals generated by action potentials (Fig. 1 B). The amplitude of the Ca²⁺ transients increased and this potentiation was evident a few minutes after the ß_1a subunit was pressure injected. The potentiation reached a peak 20 min after the injection and then slowly declined to preinjection values. The time course of ß_1a subunit action is summarized in Fig. 2. To compare data from different experiments, the ratio between the peak amplitude of the Ca²⁺ signal, at any given time, relative to its mean value before pressure injection, was computed for every experiment. Their average values are shown in Fig. 2. Each symbol represents mean values (± SE). Open symbols represent results from fibers that were pressure injected with a control solution. Solid circles represent results from fibers that were pressure injected with the ß_1a subunit. The smooth curve is the solution of Eq. 2 with the parameters indicated in the legend. It describes rather well the time course of the effect of the ß_1a subunit on peak F/F signals. The most significant effect of the ß_1a subunit was to increase the peak amplitude of the Ca²⁺ transient. There were only minor changes in their time course that were not statistically significant. Before pressure injection of the ß_1a subunit, the half-time and decay-time constant of the Ca²⁺ transients averaged 42.8 ± 4.9 (n = 13) ms and 40.9 ± 4.5 (n = 13) ms, respectively. After the injection of the ß_1a subunit, these parameters were 46.1 ± 4.8 (n = 13) ms and 38.6 ± 4.3 (n = 13) ms, respectively.

View larger version (16K): [in this window] [in a new window]   FIGURE 1  The action of the ß1a subunit upon Ca signals. (A) The records show Fluo-3 Ca2+ signals associated with action potentials from a muscle fiber that was pressure injected with a control solution. (B) Ca2+ signals from a separate fiber to which the ß1a subunit was pressure injected. The numbers below each trace indicate time in minutes computed since microinjection was done.

View larger version (15K): [in this window] [in a new window]   FIGURE 2  The time course of action of the ß1a subunit. Circles represent mean values (± SE) of peak Ca2+ signals as a function of time. Values were normalized for every experiment as: Peak j / Peak a, where j is every individual peak Ca2+ signal value and a is the average peak value of signals recorded before pressure injection; () represents values of normalized records obtained from pressure-injected fibers with the control solution; (•) represents values from pressure-injected fibers with the ß1a subunit. The arrow indicates time of microinjection. Unless otherwise indicated by the number in parentheses, each symbol represents the mean value of at least five to seven separate experiments. The smooth curve is the graph of Eq. 2 at x = 125 µm and D = 6.2 x 10–8 cm2 s–1.

To measure charge movement, voltage clamp experiments were carried out in FDB muscle fibers. We chose this preparation because of the small fiber size that favors diffusion of the ß_1a subunit and provides better space clamp than large fibers. The mean fiber length and diameter were 565 ± 26 µm (n = 12) and 29 ± 2 µm (n = 12), respectively. The mean capacitance was 899 ± 72 pF (n = 12). The expected capacitance, assuming a value of 1 µF cm^–2, was 508 ± 26 pF (n = 12).

Fig. 3 A illustrates the relationship between the amount of mobilized charge and membrane potential from control experiments. Each symbol represents average values ± SE. The smooth curve is the best fit of Eq. 3 with the parameters indicated in the legend. The Boltzmann parameters in Fig. 3 A are similar to those found by Collet et al. (36) in FDB muscle fibers. The inset shows nonlinear currents from a representative experiment at the potential indicated (in mV) with the numbers beside each trace. Fig. 3 B summarizes results from experiments in which the pipette contained the ß_1a subunit. The maximum charge values and the voltage dependence of charge movement were very similar to those of the control experiments. This indicates that, in the short term, the ß_1a subunit has no effect on charge movement.

View larger version (14K): [in this window] [in a new window]   FIGURE 3  The action of the ß1a subunit on charge movement. (A) () Represents mean values (± SE) of charge movement as a function of membrane potential from control experiments (n = 16). The smooth curve is the best fit of Eq. 3 to the data points with Qmax = 31.3 nC/µF, V = –36.6 mV, and k = 15.0 mV. (B) (•) Represents mean values (± SE) of charge movement as a function of membrane potential in the presence of the ß1a subunit (n = 16). Eh = –100 mV. The smooth curve is the Boltzmann fit with Qmax = 30.2 nC/µF, V = –36.0 mV, and k = 13.9 mV. The insets in panels A and B show records of nonlinear currents from representative experiments. The amplitude and timescale in panel A also applies to panel B.

View larger version (20K): [in this window] [in a new window]   FIGURE 4  The action of the ß1a subunit on L-type currents. (A) The records show nonlinear currents from a control experiment at the potentials indicated with the numbers (in mV) between traces. (B) Records of membrane currents at the same potentials from a separate experiment performed in the presence of the ß1a subunit. Currents were normalized per unit capacitance. (C) The current-voltage relationship of Ca2+ currents. Symbols represent mean values (± SE) of peak Ca2+ currents from control experiments () (n = 21) and from experiments performed in the presence of the ß1a subunit (•) (n = 16). Eh = –80 mV. The smooth curve is the best fit of Eq. 4 to the data points with Gmax = 122.5 µS/µF, V = 3.4 mV, k = 4.2 mV, and Vrev = 70.2 mV () and Gmax = 163.6 µS/µF, V = 7.0 mV, k = 3.1 mV, and Vrev = 81.7 mV (•). (D) The voltage dependence of the normalized conductance. Mean current density values from panel C were divided by Gmax x (Vm – Vrev). Smooth curves are best fits of a Boltzmann function with V = 3.4 mV and k = 4.1 mV () and V = 7.0 mV and k = 3.0 mV (•).

Role of the carboxy-terminus on potentiation of Ca²⁺ signals
The carboxy-terminus region of the ß_1a subunit is essential for the restoration of E-C coupling in ß-null myotubes to levels similar to those of control cells (37). Therefore, the role of this region in the potentiation of Ca²⁺ transients induced by action potentials was examined. Fig. 5 A shows representative Ca²⁺ signals from a fiber that was pressure injected with the ß_1a subunit purified from bacteria. Consistent with the data illustrated in Fig. 1 B, there was a gradual increase in the amplitude of the signals that reached a peak value 16 min after injection. After that, there was a slow return to control values. In contrast, when a truncated ß_1a subunit (lacking 40 amino acids in the carboxy-terminus region) was pressure-injected, the amplitude of the Ca²⁺ signals remained unchanged. Similar results were observed in 15 other experiments. The peak F/F values averaged 4.1 ± 0.2 (n = 16) before pressure injection and 3.9 ± 0.2 (n = 16), 20 min after the truncated form of the ß_1a subunit was pressure injected. Likewise, electrophysiological experiments revealed that the truncated form of the ß_1a subunit had no effect on L-type currents or on charge movement.

View larger version (25K): [in this window] [in a new window]   FIGURE 5  The involvement of the carboxy-terminus region of the ß1a subunit on Ca2+ signal potentiation. (A) The records show Fluo-3 Ca2+ signals associated with action potentials from a muscle fiber that was pressure injected with the full-length ß1a subunit purified from bacteria. (B) Ca2+ signals from a separate fiber to which a truncated form of the ß1a subunit, lacking 40 amino acids in the carboxy-terminus region, was pressure injected. The numbers below each trace indicate time in minutes computed since microinjection was carried out. The dotted lines indicate resting fluorescence and peak F/F before pressure injection.

Action of the ß_1a subunit on SR Ca²⁺ release channels
The possibility that the ß_1a subunit protein is acting directly on SR Ca²⁺ release channels was tested by performing SR Ca²⁺ leak and bilayer experiments. Fig. 6 A shows the action of the ß_1a subunit on RyR-mediated Ca²⁺ leak from a population of heavy SR membrane microsomes. The SR microsome population was actively Ca²⁺ loaded using the ATP driven SERCA pump. The pump was then blocked with CPZ and the rate of Ca²⁺ leak from the microsomes measured in five different experimental conditions (control, caffeine, ruthenium red, ß_1a, and truncated ß_1a). Open diamonds and squares show leak rates in the presence of ruthenium red and caffeine, respectively. Ruthenium red slowed leak rate whereas caffeine accelerated the leak. The average leak rate (nM Ca mg protein^–1min^–1) was 17.7 ± 2.4 (n = 6) in ruthenium red and 489 ± 55 (n = 6) in caffeine. Open and solid circles show leak rates in the presence of the ß_1a subunit and truncated ß_1a subunit, respectively. The average leak rate was 64.9 ± 7.7 (n = 6) with the ß_1a subunit added and 62.8 ± 8.8 (n = 6) when the truncated ß_1a subunit was added. The leak rates with the normal and truncated subunits were virtually identical and overlap control leak data (i.e., leak after CPZ application without another added reagent; data not shown). This result suggests that the ß_1a subunit does not activate RyR-mediated Ca²⁺ release from SR microsomes.

View larger version (55K): [in this window] [in a new window]   FIGURE 6  The action of the ß1a subunit on Ca2+ leak from SR microsomes and on single RyR1 channel function. (A) The Ca2+ leak rate of Ca2+ loaded skeletal muscle SR microsomes (50 µg) was recorded after SERCA pump blockade by 25 µM cyclopiazonic acid (CPZ). Leak was measured in different experimental conditions (, 2.5 mM caffeine; , 5 µM ruthenium red; •, ß1a subunit; , truncated ß1a subunit). (B) Single RyR1 channel activity in planar lipid bilayers was monitored in the absence (control) and presence of the ß1a subunit. The charge carrier was Ca2+ and holding potential was 0 mV. The solution in the cytosolic chamber had 2 µM free Ca2+, 5.5 mM total Mg2+, and 7.0 mM total ATP.

	DISCUSSION

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
Role of the ß-subunit on Ca²⁺ release
The major finding of this study is that the acute application of ß_1a subunits increases the amount of Ca²⁺ that is released by an action potential. Although the underlying mechanism of the potentiation of Ca²⁺ signals by ß_1a subunits is unknown, the fact that no changes in charge movement were observed suggests that the ß_1a subunit improves the efficiency of the coupling mechanism, where the same number of voltage sensors would open more Ca²⁺ release channels. Alternatively, the ß_1a subunit may improve the effectiveness of the Ca²⁺-induced Ca²⁺ release mechanism that is known to operate in skeletal muscle, although Ca²⁺-induced Ca²⁺ release is a minor component in mammalian skeletal muscle (38) and no direct action of the ß_1a subunit on the RyR1 channel was detected in our studies.

An alternative explanation would involve binding of the ß_1a subunit to intracellular factors that regulate Ca²⁺ release. The SR Ca release process is regulated by numerous factors in cells. The presence of some of these factors (e.g., sorcin) may chronically inhibit the SR Ca²⁺ release process. If these chronically inhibitory factors also interact with the ß_1a subunit, the application of excess ß_1a subunits could alter the way this factor regulates release. The addition of excess ß_1a protein could hypothetically result in larger Ca²⁺ release simply by interacting with this factor. Based on the current absence of data supporting this alternative interpretation, we believe an increase in coupling efficiency is more likely the underlying mechanism involved.

From a structural point of view, potentiation of Ca²⁺ signals could be explained in two ways. The first possibility is that not all _1s subunits in the transverse tubular system (TTS) membranes are associated with ß_1a subunits. In this scenario, the exogenously added ß_1a subunit would bind to free _1s subunits to exert their action. However, this is very unlikely because the ß_1a subunit is essential for the targeting of the channel (12) and thus all correctly targeted _1s subunits should have associated ß_1a subunits. As discussed by Dolphin (39), however, it is possible that the affinity of _1s subunits for ß_1a subunits is reduced once the channel has reached the TTS membranes. The second possibility is that there are multiple binding sites between _1s and ß_1a subunits (40). In this scenario, additional ß_1a subunits would bind to a single _1s subunit modulating the interaction between the _1s subunit and the RyR1 channel. In either case, the ß_1a subunit would modulate the signal transduction process involved in EC coupling.

Role of the C-terminus region
Our data further indicate that the carboxy-terminus region of the ß_1a subunit is essential for potentiation of Ca²⁺ transients. Previous work has established that this region is required for restoration of normal E-C coupling in ß-null cells (37,41) and it is different from the high affinity site in ß (the BID region), a 30-amino-acid N-terminal region of its second conserved domain (42), that interacts with a high-affinity site in the -subunit (the AID region) (43,44). It is thus possible that higher order regulatory complexes are formed where the ß_1a subunit binds to the _1s subunit in different regions. The high-affinity binding site would provide the structural basis for the strong interaction between _1s and ß_1a subunits allowing the trafficking and expression of _1s in TTS. The regulatory effects of the ß_1a subunit on Ca²⁺ release that we report here, might result from the interaction of _1sß_1a complexes with a second ß_1a subunit, possibly mediated by low-affinity sites. This last possibility is fully compatible with these findings, since the time course of the effect of the ß_1a subunit on Ca²⁺ signals could be described by a purely diffusional process. The additional interaction of a ß_1a subunit with the Ca_V1.1 channel would not necessarily lead to the formation of a stable complex as it has been described for the ß-modulation of the _1c channel (45). Also, low-affinity sites in the ₁ subunit of Ca²⁺ channels have been described in regions distant from the AID region (46,44,47,48), and based on the effects of increasing concentrations of ß₃ on _1A subunit, it has been suggested that there are two distinct binding processes for ß-subunits (49). A low-affinity interaction between _1s and ß_1a subunits would be more easily switched "on" and "off" as expected from a regulatory process.

Is the _1s subunit function limited by the ß-subunit?
These results suggest a different role of ß-subunits from the classical action on trafficking of the channels to the plasma membrane. The capability of the ß_1a subunit to potentiate Ca²⁺ transients in the short term, suggests that this auxiliary subunit is a limiting factor regulating the amount of Ca²⁺ that is released by the SR. Therefore, the ₁ subunits of skeletal muscle are not normally saturated with ß, raising the possibility that Ca release may be regulated by ß-subunits under physiological conditions. In this regard, there are several previous observations suggesting that the function of ₁ subunits is limited by the amount of ß-subunits available. Thus, when Ca_V1.1 channels in a cell-free preparation are exposed to additional ß_1a subunits, a potentiation of L-type Ca²⁺ channel currents is observed (15). Also, overexpression of ß-subunits in adult heart cells increases whole-cell, L-type Ca²⁺ currents, and maximal gating charge (48), whereas overexpression of _1c in a transgenic mouse model does not lead to increases in Ca²⁺ channel current density values (50). The observations of Yamaguchi et al. (51) are also consistent with the idea that the concentration of ß-subunits is limiting: the injection of ß₃ subunit protein increases Ca_V1.2 currents in Xenopus oocytes and it has acute effects on the biophysical properties of the channel.

A physiological modulation of Ca release by ß-subunits would not be expected to involve a very high concentration of these proteins. In this regard, we used in these experiments an amount of ß-protein that is distinctly lower (at least five times) than the one used in the experiments of Yamaguchi et al. (51) and Opatowsky et al. (52). A physiological role of the ß-subunits on Ca release in muscle would require the presence of an intracellular pool of ß-proteins that could be shuttled to the membrane where they would bind to ₁ subunits leading to changes in the interaction between ₁ subunits and RyR1. The presence of ß-subunits in skeletal muscle, not associated with ₁ subunits, has indeed been described in tissue homogenates (53).

	ACKNOWLEDGEMENTS

TOP ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS REFERENCES

 
We thank Dr. Roberto Coronado and Dr. Zurisaddai Hernández for relevant discussions and advice, Dr. Patricia Powers for the ß_1a cDNA, and Dr. Julio Vergara for providing the pressure injection system. We thank Rodrigo Sanchez for data analysis. We also thank Oscar Ramírez, Rubén García, Maura Porta, and Dr. Paula Diaz-Sylvester for technical assistance and Ms. Susana Zamudio for secretarial work.

This work was supported by CONACyT grants, 41180-N and 37356-N, and by National Institutes of Health grants R01 HL63903 and HL57832, MDA3699.

Submitted on May 23, 2005; accepted for publication September 13, 2005.

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