α₂δ₁ Dihydropyridine Receptor Subunit Is a Critical Element for Excitation-Coupled Calcium Entry but Not for Formation of Tetrads in Skeletal Myotubes

Abstract

It has been shown that small interfering RNA (siRNA) partial knockdown of the α₂δ₁ dihydropyridine receptor subunits cause a significant increase in the rate of activation of the L-type Ca²⁺ current in myotubes but have little or no effect on skeletal excitation-contraction coupling. This study used permanent siRNA knockdown of α₂δ₁ to address two important unaddressed questions. First, does the α₂δ₁ subunit contribute to the size and/or spacing of tetradic particles? Second, is the α₂δ₁ subunit important for excitation-coupled calcium entry? We found that the size and spacing of tetradic particles is unaffected by siRNA knockdown of α₂δ₁, indicating that the visible particle represents the α_1s subunit. Strikingly, >97% knockdown of α₂δ₁ leads to a complete loss of excitation-coupled calcium entry during KCl depolarization and a more rapid decay of Ca²⁺ transients during bouts of repetitive electrical stimulation like those occurring during normal muscle activation in vivo. Thus, we conclude that the α₂δ₁ dihydropyridine receptor subunit is physiologically necessary for sustaining Ca²⁺ transients in response to prolonged depolarization or repeated trains of action potentials.

INTRODUCTION

The dihydropyridine receptor (DHPR) complex in the plasma membrane/transverse tubular system of skeletal muscle is composed of five subunits (α₁, α₂, β, δ, and γ) and serves at least three distinct functions. One of these functions is as a high-voltage-activated, L-type Ca²⁺ channel. As for other voltage-activated Ca²⁺ channels, the structures underlying voltage sensing and ion permeation reside in the α₁ subunit (termed α_1s, CaV1.1 in skeletal muscle). In addition to this function as a calcium channel, the CaV1.1 serves to link depolarization of the plasma membrane to Ca²⁺ release from the type 1 ryanodine receptor (RyR1) in the sarcoplasmic reticulum (SR), which in turn causes contraction. This excitation-contraction (EC) coupling in skeletal muscle does not depend on the L-type Ca²⁺ current and is thought instead to reflect conformational coupling between the CaV1.1 and RyR1. Strong support for such conformational coupling is provided by the discovery of the retrograde interaction between the two molecules (1,2) and freeze fracture electron microscopy. The latter technique reveals that DHPR complexes are arranged in groups of four (termed “tetrads”) such that each tetradic particle, resulting from the fracture of a single complex, is aligned with one of the four identical subunits of an underlying RyR1 (3).

A third function of the DHPR complex is in the recently elucidated process termed “excitation-coupled calcium entry” (ECCE). In ECCE, depolarization causes the entry of extracellular Ca²⁺ via a pathway with pharmacological similarities to the pathway involved in store-operated calcium entry (SOCE) (4,5). However, ECCE differs from SOCE in that it does not require the emptying of intracellular Ca²⁺ stores and it requires the presence of both the CaV1.1 and RyR1. Until our recent discovery (4,5), evidence for depolarization associated Ca²⁺ entry had only been indirectly demonstrated (6–8), but the mechanism for this entry remained elusive. In addition, the fact that muscle twitches (9) could be sustained in Ca²⁺-free external buffer focused attention away from seeking mechanisms for this entry, even though it had been known for a long time that prolonged contractions could not be sustained for long periods of time in the absence of external Ca²⁺ (10,11). Discrimination between depolarization induced SR Ca²⁺ release and sarcolemmal Ca²⁺ entry is technically difficult and therefore has limited investigations of ECCE in adult skeletal muscle fibers. However, recent results from studies using the Mn²⁺ quench technique on single dissociated flexor digitorum brevis fibers have recapitulated the existence of our ECCE results from myotube experiments; ECCE is seen in adult skeletal muscle fibers, and its rate is enhanced by ryanodine pretreatment (12).

An important tool for analyzing how the functions of the DHPR complex depend on its various subunits has been provided by animals bearing null mutations. Clearly important is CaV1.1, which consists of four repeat sequences, each having six transmembrane helices. EC coupling is absent in skeletal muscle of dysgenic mice, which are homozygous null for CaV1.1 (13–19), and there is an absence of large particles in the sarcolemma of these animals when subjected to freeze fracture studies (3). EC coupling is also absent in skeletal muscle of mice or zebrafish null for the cytoplasmic β₁ subunit (16–20). In both mice and zebrafish, the absence of the β₁ subunit results in an impaired trafficking of α_1S to the plasma membrane. Additionally, the work on zebrafish has shown that those α_1S subunits that do reach the plasma membrane lack a well-defined association with RyR1 in β1-null fish (20). Genetic deletion of the γ subunit, which contains four transmembrane helices, does not markedly affect muscle function (13,21).

The α₂ and δ subunits contribute almost half of the total mass of the DHPR and arise by cleavage of a common precursor. The α₂ subunit is positioned on the extracellular side of the plasma membrane and is disulfide bonded to the δ subunit, which has a single transmembrane span (22). Mice null for the α₂δ isoform that is expressed in skeletal muscle (α₂δ₁) appear to have an embryonic or birth lethal phenotype (J. Offord, Pfizer, Groton, CT, personal communication, 2007). Thus small interfering RNA (siRNA) knockdown represents a useful alternative for examining the roles of α₂δ₁ in skeletal muscle. Previously, it was shown that siRNA partial knockdown of α₂δ₁ caused a significant increase in the rate of activation of the L-type Ca²⁺ current in myotubes but had little or no effect on skeletal EC coupling (23). The goal here was to use siRNA knockdown to address two important questions not examined in these previous studies. We chose the myotube system rather than adult fibers because only with myotubes is it possible to study a defined population of cells with the identical amount of gene silencing in each cell and because we previously demonstrated that the two systems recapitulate each other in areas we wished to study.

The first question posed was, is the α₂δ₁ DHPR subunit important for ECCE (4,5)? The second question was, does the α₂δ₁ subunit contribute to the size and/or spacing of tetradic particles? With respect to the latter question, we found that the size and spacing of tetradic particles is unaffected by siRNA knockdown of α₂δ₁, which may indicate that the α₂δ₁ protein does not affect the fracturing process and that the visible particle represents the α_1s subunit. Strikingly, in response to the first question we found that knockdown of α₂δ₁ leads to a nearly complete loss of physiologically stimulated ECCE, which results in a rapid decrement in the size of myoplasmic Ca²⁺ transients during bouts of repetitive stimulation like those occurring during normal muscle activation in vivo.

MATERIALS AND METHODS

Cell culture and siRNA transduction

Primary murine myoblasts were cultured on collagen-coated 10 cm plates (Costar, Cambridge, MA) for propagation and on Matrigel (BD Bioscience, San Jose, CA) coated plates (96-well plates (Opticlear Costar 3614) for imaging, and 10 cm dishes for membrane preparations) in Ham's F10 20% fetal bovine serum, 5 ng/ml basic fibroblast growth factor, 100 μg/ml streptomycin sulfate, 100 units/ml penicillin-G in 5% CO₂. Myoblasts were transduced three times with one of two different pSiREN retrovirus constructs (Fig. 1 A) containing α₂δ₁ complementary sequences connected by a central hairpin loop SIR1: 5′… G TGT GAG CGG ATT GAC TCT … 3′, and SIR2: 5′… G GCA CGC TTT GTG GTG ACA …3′ (Fig. 1 A) and selected for puromycin resistance conferred by the vector. Resistant cells were then plated at clonal density and 20 individual clones screened for α₂δ₁ messenger RNA (mRNA) expression by TaqMan real time polymerase chain reaction (PCR) analysis to select for cells expressing <2% of control. A previously created TRPC3 siRNA knockdown cell line was used as a control for possible off-site nonspecific effects of siRNA (24). For Ca²⁺ imaging, electrophysiology, freeze fracture, or biochemistry transduced cells were allowed to reach 60%–70% confluence and were induced to differentiate into myotubes for 5 days by changing the growth medium to Dulbecco's modified Eagle's medium containing 2% heat-inactivated horse serum, 100 μg/ml streptomycin sulfate, and 100 μg/ml penicillin-G in 20% CO₂.

FIGURE 1.

siRNA vector construction and effects of α₂δ₁ siRNA knockdown on expression of α₂δ₁ protein in primary myoblasts and myotubes. (A) Sequences of siR1 and siR2 and vector map of retroviral construct used to transduce primary muscle cells. (B) Western blot showing expression of α₂δ₁ in pools of primary myoblasts after transduction three times with retrovirus expressing constructs for siR1 and siR2. (Note siR2 did not significantly lower α₂δ₁ expression.) (C) Western blots showing expression of α₂δ₁ in myotubes derived from 27 individual myoblast clones that had been transduced three times with retrovirus expressing siR1. α-tubulin (lower band) used as the loading control. All clones with the exception of C12 showed negligible expression compared to control. At the right extracts from siR1 pooled myoblasts and control myoblasts are shown as controls.

Membrane preparation and immunoblotting

Crude membrane preparations were made from myotubes after 5 days in differentiation media. Myotubes were harvested in harvest buffer (137 mM NaCl, 3 mM KCl, 8 mM Na₂HPO₄, 1.5 mM KH₂PO₄, and 0.6 mM EDTA, pH 7.2) from 10–15 10 cm plates and centrifuged for 10 min at 250 × g. The pellet was resuspended in buffer consisting of 250 mM sucrose, 10 mM HEPES, pH 7.4, supplemented with 1 mM EDTA, 10 μg/ml leupeptin, 0.7 μg/ml pepstatin A, 5 μg/ml aprotinin, and 0.1 mM phenylmethanesulfonyl fluoride and then homogenized using a Polytron cell disrupter (Brinkmann Instruments, Westbury, NY). The whole cell homogenates were centrifuged for 20 min at 1500 × g max, and the supernatants were collected and recentrifuged for 60 min at 100,000 × g max at 4°C. The membrane pellets were finally resuspended in 250 mM sucrose, 20 mM HEPES, pH 7.4, frozen in liquid N₂, and stored at −80°C. Western blot analysis was done on a heavy SR fraction from control and α₂δ₁ DHPR knockdown myotubes using a method published previously (25). The membrane fractions were solubilized in sodium dodecyl sulfate and size fractionated by polyacrylamide gel electrophoresis. The polyacrylamide gels were then blotted onto polyvinylidene fluoride membranes and, after blocking, hybridized with the appropriate primary antibody. They were then probed with a horseradish peroxidase secondary antibody and level of expression determined with chemilumenescence. The samples were probed for RyR1, CaV1.1, α₂δ₁, skeletal calsequestrin (CSQ), FK506-binding protein 12 kDa (FKBP12), SR/ER Ca²⁺-ATPase (SERCA1), and skeletal triadin (25). Anti-β-tubulin (26) was used as a loading control.

Quantitative PCR

TaqMan Real Time PCR analysis was performed on total RNA extracted from two 10 cm plates of differentiated myotubes using RNeasy Kit (Qiagen, Valencia, CA). α₂δ₁ mRNA levels relative to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) mRNA levels were determined by quantitative reverse transcription-polymerase chain reaction using predeveloped commercial TaqMan Gene Expression Assays from Applied Biosystems (Assay ID: Mm00486607_m1; Foster City, CA) according to the manufacturer's instructions in an ABI PRISM 7700 Sequence Detection System for 50 cycles. α₂δ₁/GAPDH mRNA ratios were obtained from the equation 2^−ΔCT, where ΔCT is the difference in threshold cycles between α₂δ₁ and GAPDH.

Immunohistochemistry analysis

Cells were fixed in cold methanol (27,28). After blocking, fixed cells were hybridized with an anti-α₂δ₁ primary mouse monoclonal antibody (20A, Hybridoma Studies Bank, University of Iowa) and then hybridized with a goat anti-mouse Cy3 labeled secondary antibody and observed with a 100× oil immersion lens on an Olympus IX70 microscope (Olympus, Tokyo, Japan).

Whole-cell measurements of Ca²⁺ currents

Ca²⁺ currents were measured (29,30) using whole-cell patch-clamping with a Warner PC501 patch amplifier (Hamden, CT). Cells were patched with pipettes that were pulled from borosilicate glass capillaries to a resistance of 1.7–3.0 MΩ. Pipettes were filled with internal solution containing (in mM) 140 CsAsp, 5 MgCl₂, 10 Cs₂ EGTA, and 10 HEPES, with the pH adjusted to 7.4 with CsOH. After a seal was formed, the bath was perfused with an additional ∼10 ml of the external solution containing (in mM) 145 tetraethylammonium (TEA), 10 CaCl₂, 0.003 tetrodotoxin, and 10 HEPES, pH 7.4, with CsOH. Control currents, elicited by 20–40 ms voltage steps from −80 mV to −110 mV, were used to determine linear capacitance (cells >800 pF were excluded from further analysis). The voltage clamp command sequence was to sequentially step from: a holding potential of −80 mV, to −20 mV (200 ms), to −50 mV (75 ms), to the test potential (20 ms), to −50 mV (125 ms), and back to −80 mV. Test potentials ranged in 10 mV increments from voltages from −20 to +60 mV/+80 mV. Test currents were corrected for linear components of leak and capacitive currents by digital scaling and subtraction of the average of 10 control currents, which were elicited with the pulse protocol just described, with a 200 ms test potential of −80 mV.

Peak current density-voltage relationships of individual cells were fitted with the equation

where I is the peak current density at test potential V, G_max the maximum Ca²⁺ conductance, V_rev the reversal potential, V_1/2 the potential for half-maximal activation of conductance, and k the slope factor. Time constants of current activation were determined by fitting individual current records with a single (α₂δ₁ knockdown) or double (normal) exponential function.

All data are presented as mean ± SE.

Freeze fracture and measurements of intratetrad distance

The cells were washed twice in phosphate buffer saline at 37°C, fixed in 3.5% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.2, stored in fixative for 1–4 weeks, and briefly infiltrated in 30% glycerol. A small piece of the coverslip was mounted with the cells facing a droplet of 30% glycerol, 20% polyvinyl alcohol on a gold holder, and then frozen in liquid nitrogen-cooled propane. The coverslip was flipped off to produce a fracture that followed the cell surface originally facing it. The fractured surfaces were shadowed with platinum at 25° while rotating and then replicated with carbon in a Balzers freeze fracture machine (model BFA 400; Balzers Spa, Milan, Italy). Replicas were photographed in a Philips 410 Electron Microscope 410 (Philips Electron Optics, Mahwah, NJ). The specimen holder was positioned in the eccentric position to minimize variations in electron optical magnification. Parallel culture dishes were immunolabeled (see Immunohistochemistry analysis above).

Measurements of intratetrad distance

The distances between the pale center of each black ring of platinum (representing a DHPR complex) and the centers of its nearest neighbors in a tetrad were measured using the NIH Image software on micrographs taken at a magnification of 33,900 and scanned at a resolution of 1200 dpi. Measurements on α₂δ₁ knockdown cells were done on clones C12 and C14. To minimize variations due to distortion of the proteins during fracturing, measurements were taken only from portions of tetrads in which the angle subtending the lines connecting three adjacent particles was 90° or close to it. Incomplete tetrads with one particle missing as well as complete four-particle tetrads were used for the measurements. Blind data collection was insured by randomly mixing images from control and knockdown cells.

Myotube Ca²⁺ imaging

Stock concentrations of caffeine solutions were prepared in imaging buffer (125 mM NaCl, 5 mM KCl, 2 mM CaCl₂, 1.2 mM MgSO₄, 6 mM glucose, and 25 mM HEPES, pH 7.4). In KCl-containing solutions the concentration of NaCl was adjusted as necessary to maintain the total ionic strength (KCl + NaCl) equal to 130 mM.

Differentiated myotubes were loaded with 5 μM Fluo-4 AM (Molecular Probes, Eugene, OR) at 37°C for 20 min in imaging buffer as described previously (31). The cells were then washed three times with imaging buffer and transferred to a Nikon TE2000 microscope (Tokyo, Japan). Fluo-4 was excited at 494 nm, and fluorescence emission was measured at 516 nm using a Nikon 40× FluorApo oil 1.3 numerical aperture (na) objective. Data were collected with an intensified 12-bit digital intensified charge-coupled device (ICCD; Stanford Photonics, Stanford, CA) from regions consisting of 10–20 individual cells and analyzed using QED software (QED, Pittsburgh, PA). Test agents were delivered using the Multivalve Perfusion System (Automate Scientific, Oakland, CA). A dose response curve for a single agent was performed by adding an ascending concentration of that agent in any given well to compare the response to any given agent of the cells expressing the α₂δ₁ siRNA to that of cells expressing wild-type-type (wt) α₂δ₁ protein.

To compare different experiments, individual response amplitudes were normalized to the maximum fluorescence obtained in the same cell depolarized with 60 mM KCl. To test for differences in Ca²⁺ stores, a group of both wt and α₂δ₁ knockdown cells were exposed to 200 nM thapsigargin (TG) in the presence of nominal Ca²⁺-free media and the Ca²⁺ release response recorded. To test that a deficiency in Ca²⁺ entry during depolarization might be responsible for the observed responses of α₂δ₁ knockdown cells to depolarization, cells were pretreated with nominal Ca²⁺-free buffer for 1 min before depolarization in the same nominal Ca²⁺-free buffer with 60 mM KCl. Where indicated, nonlinear regression with sigmoidal dose-response analysis was performed using Prism version 4.0. (GraphPad Software, San Diego, CA). Data are presented as mean ± SE.

Additional experiments were performed to measure cell responses to electrical pulse trains. Myotubes were loaded with Fluo-4 as described above and fluorescence emission (516 nm) acquired with an IC-300 ICCD camera (Photon Technology International, Lawrenceville, NJ). Data were captured, digitized, and stored on computer using ImageMaster software (Photon Technology International). Bipolar electrical field stimuli were applied using two platinum electrodes fixed to opposite sides of the well and connected to an AMPI Master 8 stimulator set at 3 V, 25 ms pulse duration, over a range of frequencies (0.05–20 Hz). In these experiments, data were acquired at 200 ms intervals. In separate experiments, a fatigue stimulus protocol was applied. Electrical pulses (7 V, 1 ms duration) were applied at 100 Hz for 500 ms interspersed with 500 ms rest for a total duration of 5 min. The frequency-transient amplitude relationship was studied using 3 s pulse trains in a range of 1–120 Hz.

Mn²⁺ quench and electrical field stimulation

Mn²⁺ influx was measured as previously described (32,33). Briefly, differentiated primary wt or α₂δ₁ knockdown myotubes were loaded with fura-2, the cells were imaged using an ICCD camera with an Olympus 40× oil 1.2 na objective, and data were collected from 3–10 individual cells at five frames per second. The isosbestic wavelength for fura-2 was calibrated at the beginning of each experiment. A final concentration of 500 μM MnCl₂ (in the presence of 1.2 mM external Mg²⁺) was used, and emission was monitored at 510 nm. Cells loaded with fura-2 (5 μM) were stimulated with either local application of K⁺ or by electrical field stimulation as described above. To compare the effects of α₂δ₁ DHPR knockdown on both traditional store depletion induced entry (SOCE) and ECCE, myotubes were pretreated for 10 min with 200 nM TG followed by confirmation that the stores were empty with a challenge with 20 mM caffeine. Mn²⁺ quench experiments were only performed on cells that did not respond to the caffeine challenge (>99%).

RESULTS

Construction of siRNA vectors that efficiently knock down expression of the α₂δ₁ DHPR subunit

We used the following general design guidelines to construct our two siRNA vectors: 1), find 21 nt sequences in the target mRNA that begins with an adenine dinucleotide, 2), select several target sequences with 30%–60% GC content and no stretches of >4 Ts or As at different positions along the length of the gene sequence, and 3), compare the potential target sites to the appropriate genome database to eliminate from consideration any target sequences with more than 16–17 contiguous basepairs of homology to other coding sequences.

Fig. 1 A shows a map of the location of the two selected siRNA sequences in the α₂δ₁ complementary DNA (cDNA). Because myoblasts are difficult to transfect at high efficiency and because of the need to have multiple copies of the vector permanently integrated in the cell to reduce expression of the target mRNA to <2% of control, a pSiREN retrovirus was used to transduce the cells at moiety of infection = 5 on three separate occasions (34,35). Between transductions, cells were selected with puromycin to eliminate any cells that were not transduced with at least one copy of the vector from the cell population. Western blot analysis of pooled cells after the third round of infection with the siRNA retrovirus demonstrated that although both siR1 and 2 reduced the expression of α₂δ₁ in myoblasts, siR1 was much more efficient at this task (Fig. 1 B). Cells from pool 1 were plated at clonal density and the clones expanded. As can be seen in Fig. 1 C, myotubes derived from most of the clones from this pool did not produce sufficient α₂δ₁ protein to be detected by Western blot analysis.

In Fig. 2 A, TaqMan analysis of representative clones demonstrated that transcription of α₂δ₁ mRNA could be reduced to <5% of control in myoblasts and <2% of control in myotubes transduced with siR1. Two myoblast clones, C14 and C21, were chosen for further analysis, and the results of all of the studies done were identical in both. When only one cell line is presented, we present the data from clone C14 as the representative value. Immunohistochemistry (Fig. 2 B) confirms that in wt cells α₂δ₁ DHPR has a punctate pattern of expression (23,36), which is dictated by the positioning of CaV1.1 (36) and RyR1 (27). In α₂δ₁ DHPR siRNA knockdown myotubes, there is no α₂δ₁ DHPR detectable (Fig. 2 B), but the absence of the α₂δ₁ DHPR does not affect the targeting of CaV1.1 or RyR1 (data not shown). Cells transduced with the “control” TRPC3 siRNA, to ascertain possible nonspecific consequences on protein expression, have a wt pattern of α₂δ₁ DHPR expression (data not shown).

FIGURE 2.

Characterization of α₂δ₁ knockdown on α₂δ₁ mRNA levels, α₂δ₁ protein expression and targeting, and expression of muscle triad proteins. (A) Transcription of α₂δ₁ mRNA is significantly knocked down in myoblasts and myotubes of both clones that were selected for further study. (B) Immunohistochemistry with anti-α₂δ₁ antibody demonstrates that in control myotubes, α₂δ₁ DHPR is targeted appropriately with the characteristic punctate appearance near the cell surface, similar to the staining pattern for α_1S DHPR or RyR1 (27). Myotubes from α₂δ₁ knockdown clones 14 and 21 have no visible α₂δ₁ subunit expression. (C) Western blot analysis of myoblasts (left) and myotubes (right) from wt and α₂δ₁ knockdown clones 14 and 21. Note that α₂δ₁ is expressed at high levels in wt myoblasts and myotubes, whereas α_1S DHPR is expressed only in myotubes. The expression of α₂δ₁ protein in knockdown clones 14 and 21 appears to correlate with the α₂δ₁ mRNA levels: traces of protein and transcript were seen in clone 14 myoblasts, but protein and transcript were essentially not expressed in clone 21 myoblasts or clones 14 and 21 myotubes. The lack of expression of α₂δ₁ appears to have no effect on the expression of any other triadic protein probed.

Western blot analysis of heavy SR fractions from control and α₂δ₁ DHPR knockdown myotubes probed for RyR1, α_1S DHPR subunit, CSQ, FKBP12, SERCA1, and skeletal triadin (β-tubulin was used as a loading control) demonstrates that there is no detectable α₂δ₁ DHPR protein in either of the two clones used in this study but that the absence of the α₂δ₁ subunits was not associated with any differences in the expression of any other triadic protein tested in either myoblasts or myotubes (Fig. 2 C). Cells transduced with the “control” TRPC3 siRNA have wt amounts of α₂δ₁ DHPR protein as well as RyR1, CaV1.1, CSQ, FKBP12, SERCA1, and skeletal triadin on Western blot analysis (data not shown), showing that the phenotype of the α₂δ₁ knockdown is not simply the result of a nonspecific “off target” artifact caused by siRNA expression.

The effects of α₂δ₁ knockdown on L-type Ca²⁺ current

To determine the effects of α₂δ₁ knockdown on the function of the DHPR complex as Ca²⁺ channel, the whole-cell patch-clamp configuration was used to measure membrane Ca²⁺ currents (Fig. 3, A–C). Comparison of representative records demonstrates that, as reported previously (23), currents from α₂δ₁ knockdown myotubes (Fig. 3 A) activate more rapidly than those from normal myotubes (Fig. 3 B). Activation of currents from normal myotubes was best fitted with the sum of two exponential functions, with a dominant contribution by the slower of these two. The average time constants for these two exponential functions (7.6 ± 0.9 ms and 71.0 ± 3.8 ms; Table 1) were similar to those found previously for normal myotubes (8.5 ± 1.2 ms and 79.5 ± 10.5 ms) by Avila and Dirksen (37). Activation of currents from α₂δ₁ knockdown myotubes could be well fitted with a single exponential function having a time constant (6.4 ± 0.3 ms) similar to that of the faster component for wt myotubes (Table 1). At the potential evoking maximal inward current, α₂δ₁ knockdown currents inactivated on average by 25% ± 4% at the end of a 200-ms test pulse, whereas no inactivation was observed for wt myotubes (Table 1). Peak current-voltage relationships were similar for α₂δ₁ knockdown and normal myotubes, although G_max was somewhat lower for α₂δ₁ knockdown myotubes (Fig. 3 C; Table 1).

FIGURE 3.

Comparison of Ca²⁺ currents in α₂δ₁ knockdown and normal myotubes. Representative currents elicited by 200 ms test pulses to the indicated potentials are shown from α₂δ₁ knockdown (A) and normal (B) myotubes. Scale bars are 2 pA/pF (vertical) and 20 ms (horizontal). (C) Average voltage dependence of peak current densities for α₂δ₁ knockdown (closed symbols, n = 18) and normal myotubes (open symbols, n = 9). Smooth curves represent the Boltzmann equation (see Materials and Methods) with average parameters (Table 1) obtained from best fits of the same equation to current-voltage relationships of individual cells.

TABLE 1.

Characteristics of whole cell Ca²⁺ currents in wt and primary α₂δ₁ knockdown myotubes

	Gmax (pS/pF)	V1/2 (mV)	Vr (mV)	k	Inactivation at 200 ms	τslow (ms)	τfast (ms)	Aslow (pA/pF)	Afast (pA/pF)
Wt (n = 9)	307 ± 24	28.0 ± 1.5	81.1 ± 1.3	5.1 ± 0.1	NA	71.0 ± 3.8	7.6 ± 0.9	9.2 ± 0.8	3.7 ± 1.2
α2δ1 k.d. (n = 18)	248 ± 18*	22.5 ± 1.3	83.1 ± 1.4	6.5 ± 0.3*	(25 ± 4)%	NA	6.4 ± 0.3	NA	9.2 ± 0.8

Current time courses of α₂δ₁ knockdown cells were fitted with a single exponential function, and current time courses of wt myotubes were fitted with a double exponential function.

^*p < 0.05 vs. wt.

α₂δ₁ knockdown does not affect DHPR complex tetrad formation and intratetrad spacing

Clusters of DHPR-complex tetrads are equally visible in control and α₂δ₁ DHPR knockdown cells (Fig. 4). Indeed in this particular set of experiments the clusters in the controls (Fig. 4 A) tended to contain fewer and less complete tetrads than those in the treated cells (Fig. 4 B). Since formation of DHPR-complex tetrads requires an interaction with RyR1, it is clear that α₂δ₁-null DHPR complexes are appropriately targeted to the calcium release units (CRUs) and appropriately positioned relative to RyRs.

FIGURE 4.

Tetrads are unaffected by α₂δ₁ knockdown. (A and B) A dark ringlet of platinum surrounds the freeze fracture particles, each marking the position of the DHPR. Particles and their disposition into tetrads appear identical in the control (left) and α₂δ₁ knockdown cells, indicating that the α₂δ₁ subunit is not part of the DHPR particle seen using this technique. (C) The center-to-center distance between particles constituting tetrads (intertetrad distance) was measured in electron micrographs at a magnification of ∼85,000×. The tetrads to be measured were selected because they all show a minimum of distortion as a result of fracturing. The images from experimental and control cells were mixed during the analysis so that measurements were performed by a “blind” operator. The data are presented in a scattergram in which each dot represents a single measurement and the x axis is the dot number from 1 to 790. There were no differences in intratetrad distance between the two groups (p > 0.7). All myotubes were fixed in glutaraldehyde, cryoprotected in glycerol, freeze fractured, and rotary shadowed at 45°.

The apparent size of the particles, as indicated by the ring of platinum deposited around them, is not different in the two sets of cells (Fig. 4, A and B). Most importantly, the center-to-center distance between adjacent particles in the tetrads, or intratetrad distance, is also not altered by the absence of α₂δ₁. Fig. 4 C shows a scattergram of the measured intratetrad distances, each represented by one dot, blue for controls and orange for treated cells. The x axis of this scattergram is the tetrad number, ranging from 1 to 790. Despite the scatter in the data, due to the distortion that can occur during fracturing, the two average intratetrad distances are the same (19.1 ± 1.9 nm for control and 19.0 ± 1.8 nm for α₂δ₁ DHPR-null cells), and Student's t-test indicates that the two populations are indistinguishable (p = 0.25). This would strongly suggest that 1), the CaV1.1 and not the α₂δ₁ subunit is the protein which is observed as the large intramembrane particle when the sarcolemma is fractured; and 2), the α₂δ₁ subunit, which is proposed to be a largely extracellular protein, plays no role in the appearance of this particle, in tetrad formation, or in the position of the complexes, as indicated by the intratetrad distance.

α₂δ₁ is not essential for EC coupling but is important for maintaining Ca²⁺ transients during sustained depolarization

α₂δ₁ knockdown cells, in which the mRNA levels were reduced to <2% of control, produced Ca²⁺ transients in response to caffeine which were not significantly different from wt in terms of the peak amplitude (0.75 ± 0.02 F_max for wt and 0.76 ± 0.3 F_max for α₂δ₁ knockdown), rate of decay (data not shown), and concentration for half-maximal activation (EC₅₀ = 4.6 mM in α₂δ₁ knockdown vs. 4.6 mM for wt). Moreover, α₂δ₁ knockdown had no significant effect on the amplitude and rate of rise of the Ca²⁺ transient after depolarization with 60 mM KCl (Fig. 5). However, the rate of decay of the Ca²⁺ transient during maintained depolarization was significantly more rapid in α₂δ₁ knockdown myotubes than in wt. The half-decay time measured as the time (x) axis width of the transient at half-maximum tension fell from 24.7 ± 1.5 s in wt to 5.9 ± 0.5 s in α₂δ₁ knockdown myotubes (Fig. 5). With this exception the responses from myotubes permanently transduced with the control siRNA were indistinguishable from those seen in controls, demonstrating that the cause of the decrease in the duration of the Ca²⁺ transient and by inference all the other findings were not due to nonspecific “off target” siRNA effects.

FIGURE 5.

Compared to wt myotubes, α₂δ₁ knockdown myotubes have abbreviated Ca²⁺ transients during KCl depolarization. Representative, raw Fluo-4 signals during a 30-s 60-mM KCl depolarization of wt (left) and α₂δ₁ knockdown (right) myotubes. The bar graphs in the bottom panel show that the half-decay time for the Fluo-4 transient in response to a 30-s 60-mM KCl depolarization is significantly shorter in α₂δ₁ knockdown myotubes than in wt myotubes. Data are shown as mean ± SE. **p < 0.01; n = 130 myotubes for wt and n = 240 for α₂δ₁ knockdown.

Importantly, the difference in the rate of decay of the Ca²⁺ transient during KCl depolarization between α₂δ₁ knockdown and wt myotubes could be abolished if extracellular Ca²⁺ was reduced to 7 μM and extracellular Mg²⁺ increased to 3 mM to maintain the concentration of external divalent cations. Under these conditions, the rate of decay of the Ca²⁺ transient in both groups of myotubes was similar to that of α₂δ₁ knockdown myotubes in Ca²⁺-replete medium (mean ½ decay time of ∼5.9 s) (Fig. 6). This strongly suggests that a loss of external Ca²⁺ entry during the 30 s depolarizing stimulus contributes to the reduced duration of the Ca²⁺ transient in the α₂δ₁ knockdown myotubes.

FIGURE 6.

α₂δ₁ knockdown and wt myotubes have Ca²⁺ transients of similar duration in low Ca²⁺ buffer. Representative average raw Fluo-4 signals during 30-s 60-mM KCl depolarization of n = 65 wt (left) and n = 72 α₂δ₁ knockdown (right) myotubes when the depolarization is done in a buffer containing ∼7 μM Ca²⁺ and 3 mM Mg²⁺. The summary below shows the average time to ½ amplitude ± mean ± SE. There is no significant difference between the two genotypes under these conditions. p > 0.8.

The relationship between stimulation frequency and Ca²⁺ transient amplitude was studied with pulse trains ranging from 1 to 120 Hz (3 ms duration) and was not found to differ between wt and α₂δ₁ knockdown myotubes at any these frequencies (data not shown). Fig. 7 A shows that the amplitudes of Ca²⁺ transients produced by short electrical stimuli are not significantly different between wt and α₂δ₁ knockdown myotubes. However, unlike wt myotubes, α₂δ₁ knockdown myotubes failed to maintain the peak amplitude of their Ca²⁺ transients for a prolonged, full 120 s, pulse train when the stimulus frequency was increased above 1 Hz. The rate of decay was directly related to the stimulus frequency (Fig. 7 A). Cells were also tested for the rate of fall in their peak Ca²⁺ transient in response to a stimulus protocol similar to those used on adult skeletal muscle cells to produce fatigue. Fig. 7 B shows that the amplitude of the Ca²⁺ transient was unaffected during a long (5 min) pulse train composed of bursts lasting 500 ms interspersed by a 500 ms rest period as described in Materials and Methods in wt cells. By contrast, α₂δ₁ knockdown cells subjected to the same stimulus protocol had a steady decline in the amplitude of their Ca²⁺ transient. Thus we conclude that the α₂δ₁ DHPR subunit is physiologically necessary for sustaining Ca²⁺ transients in response to prolonged depolarization or repeated trains of action potentials.

FIGURE 7.

α₂δ₁ knockdown myotubes fail to maintain their Ca²⁺ transient amplitude with electrical pulse trains >1 Hz. (A) Cells were stimulated by sequential bipolar electrical pulse trains lasting 120 s at frequencies ranging from 0.17 to 20 Hz with a 1 min rest period between each frequency tested. The rate of decline of the peak Ca²⁺ transient amplitude was proportional to the stimulus frequency. These data are representative of measurements acquired from n = 3 wt and n = 8 α₂δ₁ knockdown myotubes using the indicated stimulus sequence protocol. (B) Ca²⁺ transient amplitudes were resistant to rundown in wt, but not α₂δ₁ knockdown, myotubes stimulated with electrical pulse trains similar to those commonly used to elicit fatigue in adult fibers, as described in Materials and Methods. The data are representative of n = 20 wt and n = 36 knockdown cells.

α₂δ₁ knockdown inhibits ECCE

We next tested the hypothesis that the mechanism for the more rapid decay of Ca²⁺ transient amplitude in α₂δ₁ knockdown myotubes was that α₂δ₁ is needed to maintain ECCE, SOCE, or both. To test the effects of α₂δ₁ knockdown on SOCE, we depleted SR Ca²⁺ stores completely with TG while exposing the cells to nominally Ca²⁺-free external medium. After store depletion was confirmed with the absence of any Ca²⁺ response to exposure to 20 mM caffeine, addition of 2 mM Ca²⁺ to the external medium clearly showed the presence of SOCE (as measured by the Ca²⁺ transient that accompanied the return to Ca²⁺-replete medium) (Fig. 8). Using this depletion protocol, the same rate and amplitude of SOCE was measured in both wt and α₂δ₁ knockdown myotubes (Fig. 9 A, upper traces). In a separate set of experiments, Ca²⁺-depleted myotubes were perfused with an external medium containing 40 mM K⁺ and 2 mM Ca²⁺ (Fig. 9 A, lower traces). K⁺ depolarization elicited rapid Ca²⁺ entry in wt cells via ECCE, whereas α₂δ₁ knockdown myotubes failed to show any detectable Ca²⁺ entry while the cells were depolarized. When K⁺ was removed from the Ca²⁺-replete external medium, a rapid Ca²⁺ entry was observed in the α₂δ₁ knockdown myotubes similar to their previous response with no depolarization (Fig. 9 A, lower traces). Thus the abolition of ECCE during KCl depolarization in the α₂δ₁ knockdown myotubes allowed the direct demonstration that SOCE is completely inhibited by sarcolemmal depolarization.

FIGURE 8.

Complete SR Ca depletion protocol for α₂δ₁ knockdown myotubes. Cells were loaded with Fluo-4 as described in Materials and Methods and tested for responses to electrical pulses and K⁺ (40 mM) application in Ca-replete medium ( 2 mM). The extracellular solution was exchanged with one lacking added Ca²⁺ ( free); it elicited spontaneous activity in most cells. TG (200 nM) was then perfused onto the cells in the -free medium until the new baseline was established (typically <10 min) and depletion of SR Ca tested with a 20 mM caffeine challenge. The presence of SOCE was then tested by perfusion of 2 mM Ca²⁺ medium. This protocol completely depleted SR Ca in >95% of all myotubes exhibiting EC coupling. Wt myotubes were indistinguishable from the α₂δ₁ knockdown myotubes shown here.

FIGURE 9.

α₂δ₁ knockdown selectively inhibits ECCE but not SOCE. (A) (Upper traces) α₂δ₁ knockdown had no effect on initial rate and amplitude of SOCE under condition of severe store depletion elicited by TG (200 nM). (Lower traces) ECCE is completely abolished in K⁺-depolarized α₂δ₁ knockdown myotubes. Note that K⁺ depolarization in the presence of external Ca²⁺ reveals the complete inhibition of SOCE even under conditions of severe depletion. (B) Representative traces showing Mn²⁺ quench of fura-2 fluorescence in α₂δ₁ knockdown cells was significantly reduced when a 20 Hz electrical pulse train was applied or completely abolished when the cells were exposed to 60 mM KCl. (C) Representative traces of KCl depolarization induced Mn²⁺ influx pre- and posttreatment with ryanodine. Although there is a brisk Mn²⁺ influx in wt cells when they are exposed to 60 mM KCl, there is no Mn²⁺ influx in α₂δ₁ knockdown cells under normal conditions. However, when the cells are pretreated with high concentrations of ryanodine (500 μM) for 30 min ECCE can be partially rescued (∼40% of wt) by the change in the conformation of RyR1 caused by ryanodine exposure. (D) Even in the presence of ryanodine, the amount of Mn²⁺ entry in α₂δ₁ knockdown cells is significantly reduced compared to wt. N > 20 for each group.

We also examined Mn²⁺ quench of fura-2 fluorescence during electrical (20 Hz trains) and 40 mM K⁺ depolarization. Fig. 9 B shows that the rate of Mn²⁺ entry is significantly reduced (p < 0.0003) in response to either KCl or electrical depolarization in α₂δ₁ knockdown myotubes. The rate of Mn²⁺ entry is reduced by more than twofold in response to electrical stimulation, and it is abolished in response to K⁺ depolarization.

ECCE requires the presence of RyR1 and CaV1.1 and is very dependent on the conformation of RyR1. We have shown that ECCE can be enhanced by two conformational mutations of RyR1 that do not support normal EC coupling (4) or by pretreatment of murine myotubes (4,5) or muscle fibers (12) with 250–500 μM ryanodine. This apparent enhancement of ECCE elicited by changes in the conformation of RyR1 caused by mutations or ryanodine binding is primarily a result of delayed inactivation/deactivation of entry during depolarization and not increased activation. To test the hypothesis that the α₂δ₁ subunit was key to the interaction of the three Ca²⁺ channels participating in ECCE, we pretreated α₂δ₁ knockdown myotubes with either 500 μM ryanodine or solvent (control) for 30 min and performed Mn²⁺ quench experiments similar to those described above. In Fig. 9, C and D, it can be seen that, unlike wt myotubes, 40 mM K⁺ depolarization of α₂δ₁ knockdown myotubes in the absence of ryanodine pretreatment failed to permit Mn²⁺ entry. However α₂δ₁ knockdown myotubes pretreated with ryanodine regained a robust long-lived K⁺-triggered Mn²⁺ entry response, although the initial rate was much (∼60%) smaller than the rate seen in wt (Fig. 9 D). This would indicate that although the α₂δ₁ subunit is required for physiologic ECCE, its absence can be partially overcome when RyR1 is put into a conformation that enhances this depolarization-induced Ca²⁺ entry.

DISCUSSION

α₂δ₁ is not necessary for myotube growth or differentiation

Despite the almost complete absence (<2%) of the α₂δ₁ DHPR subunit, there appears to be no change in the ability of the myoblasts to form myotubes or in the expression levels or targeting of CaV1.1 to the CRU (28,38). This is similar to the report of Obermair et al. (23), who studied transiently siRNA transfected cells with a reduction in α₂δ₁ mRNA to 6%–25% of control. In addition, the absence of the α₂δ₁ DHPR subunit did not affect the expression of several other components of the triad (RyR1, FKBP12, CaV1.1, CSQ, Triadin, SERCA), suggesting that the α₂δ₁ subunit is not important for the expression or normal targeting of these CRU proteins.

Unlike CaV1.1, the α₂δ₁ subunit is expressed in both myoblasts and myotubes, and α₂δ₁-null mice have an embryonic or birth lethal phenotype (J. Offord, Pfizer, Groton, CT, personal communication, 2007), suggesting that the α₂δ₁ may play a role in embryonic development of components other than muscle. Our data show that whatever the early function of α₂δ₁ in myoblasts, it is not necessary for myoblast division, their differentiation into myotubes, or basic EC coupling.

The effects of the absence of the α₂δ₁ subunit on slow voltage-gated Ca²⁺ current

Overall, our results on Ca²⁺ currents in α₂δ₁ knockdown myotubes are in good agreement with those reported previously showing that the fundamental functions of the DHPR complex as Ca²⁺ channel do not depend on the presence of the α₂δ₁ subunit (23). In particular, α₂δ₁ knockdown had little effect on the amplitude or voltage dependence of L-type Ca²⁺ currents but did result in accelerated activation kinetics. Additionally, in response to 200-ms depolarizations, the L-type currents displayed greater inactivation in α₂δ₁ knockdown myotubes, perhaps because inactivation in wt myotubes was obscured by the slower activation kinetics.

Lack of the α₂δ₁ subunit reveals the position of CaV1.1 in DHPR complex tetrads

Freeze fracture reveals the position of intramembranous proteins, but it cannot be reliably used to define the size and shape of the proteins that are fractured in the frozen state. For example, the very large ryanodine receptor of the SR produces a barely visible shallow hump when fractured (39). Nevertheless, we are surprised to find that even though the α₂δ₁ is a very large subunit, the DHPR complex particles are neither absent nor grossly distorted in its absence. The importance of this result lies in the fact that we can, albeit at a low level of resolution, predict the position of the α_1s DHPR mass relative to the four RyR subunits (40). The channel would be located somewhat peripherally relative to the RyR outline in such a manner that some portion of it could extend over the side of RyR1.

α₂δ₁ is not essential for EC coupling, but its absence alters properties of the Ca²⁺ transient

In agreement with Obermair et al. (1), we find that the α₂δ₁ DHPR subunit is not needed for skeletal-type EC coupling since there was no difference between α₂δ₁-deficient and wt myotubes in either the magnitude or the activation kinetics of the Ca²⁺ transient elicited by electrical or KCl depolarization in nominally Ca²⁺-free buffer (7 μM). However, during sustained depolarization in Ca²⁺-containing (2 mM) medium, the duration of Ca²⁺ transients in α₂δ₁ knockdown myotubes was reduced ∼5-fold compared to wt myotubes, despite similar sized SR Ca²⁺ stores, as indicated by caffeine responses. Two observations support the idea that this reduced duration in α₂δ₁ knockdown cells is a consequence of a loss of Ca²⁺ entry via ECCE. First, measurements of Mn²⁺ quench indicate that ECCE is greatly reduced in α₂δ₁ knockdown myotubes. Second, the duration of Ca²⁺ transients in wt myotubes is decreased fivefold when the external medium is made nominally Ca²⁺-free (Mg²⁺ increased to 3 mM), whereas this manipulation has no effect on the duration of the Ca²⁺ transient in α₂δ₁ knockdown cells. Although these results point to the importance of Ca²⁺ entry via ECCE for maintaining Ca²⁺ transients during prolonged depolarization, two limitations should be mentioned. First, removal of external Ca²⁺ would eliminate all sources of Ca²⁺ entry (not just that via ECCE). Second, removal of external Ca²⁺ might be expected to cause the voltage sensor for EC coupling to inactivate more rapidly (41,42). However, if either of these represented major mechanisms whereby Ca²⁺ removal caused a reduced duration of the Ca²⁺ transient in wt myotubes, it would not explain why Ca²⁺ removal had essentially no effect on α₂δ₁ knockdown cells.

α₂δ₁ is essential for physiologically engaging ECCE but not SOCE

In cells treated with TG, there was no difference in the kinetics or magnitude of the refilling response between α₂δ₁ DHPR knockdown and wt myotubes, showing that the absence of the α₂δ₁ subunit has no detectable effect on SOCE. Interestingly, when another set of myotubes from the same myoblast clones were depolarized with 40 mM KCl concurrently with the addition of 2 mM Ca²⁺ in the external medium, there was a striking difference between the two cell types. As expected (5), wt cells had a rapid Ca²⁺ transient with an even faster rise time than was seen with the addition of Ca²⁺ alone (5,43–45). α₂δ₁-deficient cells, on the other hand, showed no Ca²⁺ entry response at all during the KCl depolarization but did show a rate of cation entry similar to the rate of the SOCE transient when the KCl was removed. This result suggests that the mechanism behind the rapid decline in the Ca²⁺ transient during a 30 s KCl exposure or trains of 20 Hz electrical stimuli in α₂δ₁ knockdown myotubes was either a marked reduction or a complete absence of ECCE. In addition these data directly demonstrate and confirm the suggestions by others that SOCE is significantly inhibited by depolarization (46–48).

The results of our studies using Mn²⁺ quench of fura-2 fluorescence clearly confirm that that α₂δ₁ is an essential regulator of physiologic ECCE in skeletal myotubes and the degree of the dysfunction depends on 1), the nature of the depolarizing trigger (electrical trains versus [K⁺]), and, importantly, 2), the conformation of RyR1. From our results, it appears that if the RyR1 Ca²⁺ release channel is in or is placed into a conformational state different from the state found at rest in the cell (e.g., with the conformational state induced by certain RyR1 mutations or ryanodine), the effect of the absence of α₂δ₁ on ECCE can be partially overcome. This observation makes it unlikely that the absence of α₂δ₁ has either negatively influenced the expression or targeting of the ECCE channel to the surface membrane. Presumably the reason that ECCE is not completely abolished in α₂δ₁-deficient myotubes during electrical stimulation is that at a fairly low frequency of stimulation the voltage sensor has the opportunity to reset itself between individual stimuli. It is possible that if the stimulus frequency were raised higher than 20 Hz the reduction in the rate of Mn²⁺ entry would be more profound. This would suggest that the physiological engagement of ECCE may be most significant under conditions of extreme activity and that changes in the efficiency of Ca²⁺ entry during depolarization may be a factor in the processes leading to muscle fatigue.

In conclusion, the evidence gained from these studies confirms the previously defined role of the α₂δ₁ DHPR subunit as the particle needed to stabilize the CaV1.1 channel in the slowly activating state. Further it provides conclusive evidence that the α₂δ₁ subunit is not the particle that forms tetrads in freeze fracture studies of DHPRs in the surface membrane. Most importantly, it demonstrates that in addition to its previously defined role, the α₂δ₁ subunit also plays an important role in the three-way interaction of the DHPR complex, RyR1, and ECCE channels during normal EC coupling. However, unlike the CaV1.1, the role of the α₂δ₁ subunit in coordinating ECCE is important but not essential, as the absence of α₂δ₁ can be partially overcome by repriming the voltage sensor and almost completely overcome when the RyR1 assumes a conformation that enhances ECCE.