Dihydropyridine-induced Ca2+ release from ryanodine-sensitive Ca2+ pools in human skeletal muscle cells (2024)

1. Dihydropyridines (DHPs) are widely used antihypertensive drugs and inhibit excitation-contraction (E–C) coupling in vascular smooth muscle and in myocardial cells by antagonizing L-type Ca²⁺ channels (DHP receptors). However, contradictory reports exist about the interaction of the DHP with the skeletal muscle isoform of the DHP receptor and E–C coupling in skeletal muscle cells.

2. Using the intracellular fluorescent Ca²⁺ indicator fura-2, an increase in [Ca²⁺]_i was observed after extracellular application of nifedipine to cultured human skeletal muscle cells. The rise in [Ca²⁺]_i was dose dependent with a calculated EC₅₀ of 614 ± 96 nm nifedipine and a maximum increment in [Ca²⁺]_i of 80 ± 3.2 nm. Similar values were obtained with nitrendipine.

3. This effect of DHPs was restricted to differentiated skeletal muscle cells and was not seen in non-differentiated cells or in PC12 cells. In spite of the observed increase in [Ca²⁺]_i, whole-cell patch clamp experiments revealed that 10 μm nifedipine abolished inward Ba²⁺ currents through L-type Ca²⁺ channels completely.

4. Similar to nifedipine, (±)Bay K 8644, an agonist of the L-type Ca²⁺ channel, also increased [Ca²⁺]_i. This effect could not be enhanced by further addition of nifedipine, suggesting that both DHPs act via a common signalling pathway.

5. Based on the specific mechanism of the skeletal muscle E-C coupling, we propose the stabilization of a conformational state of the DHP receptor by DHPs, which is sufficient to activate the ryanodine receptor.

Materials

Sera and media for cell culture were obtained from PAA (Linz, Austria), trypsin, EDTA, glutamine, penicillin, streptomycin, gentamicin and amphotericin B were obtained from Gibco, xestospongin C from Calbiochem, aprotinin from Bayer AG (Wuppertal, Germany), Pefabloc from Boehringer Mannheim (Mannheim, Germany) and nifedipine from Sigma Chemical Co. and Calbiochem. (±)Bay K 8644 was a generous gift from Bayer (Austria). All other chemicals were from Sigma Chemical Co.

Cell culture

The study conformed to the code of ethics of the World Medical Association (Declaration of Helsinki) and was approved by the Ethics Committee of the General Hospital Vienna. After informed and written consent was obtained from the patients, waste material of muscle biopsies (200-900 mg) was obtained from individuals undergoing the in vitro contracture test to verify susceptibility for malignant hyperthermia (MH). For the present study cells from 13 individuals who were found to be MH non-susceptible were used. Human skeletal muscle cell culture was performed according to the method of Brinkmeier et al. (1993). In brief, muscle tissue was freed from fat and connective tissue, cut into small pieces and digested in three 20 min steps, first with trypsin (0.05 %) and EDTA (0.02 %) and then twice with collagenase (Type IA, 200 U ml⁻¹ in Hanks’ balanced salt solution). The cell suspension was centrifuged at 250 g and resuspended in wash medium (Ham's F12 supplemented with 20 % horse serum). The resuspended material was filtered through a 70 μm nylon mesh and finally seeded in growth medium into 50 ml cell culture flasks for proliferation. Growth (GM) and differentiation (DM) media were prepared according to Baroffio et al. (1993). GM contained Ham's F12 supplemented with 15 % fetal calf serum, 10 ng ml⁻¹ epidermal growth factor, 200 ng ml⁻¹ insulin, 400 ng ml⁻¹ dexamethasone, 0.5 mg ml⁻¹ fetuin, 0.5 mg ml⁻¹ BSA, 7 mM glucose, 4 mM L-glutamine, 200 U ml⁻¹ penicillin, 200 μg ml⁻¹ streptomycin and 2.5 μg ml⁻¹ amphotericin B. DM contained Dulbecco's modified Eagle's medium supplemented with 5 % horse serum, 4 mM L-glutamine, 100 ng ml⁻¹ insulin and 0.1 μg ml⁻¹ gentamicin.

Cells were incubated at 37°C under 5 % CO₂ until almost confluent, and then reseeded on 25 mm glass coverslips coated with fibronectin. Adherent cells were exposed to DM to promote fusion of myogenic satellite cells to myotubes.

PC12 cells (a generous gift of Dr Seemann, Knoll AG, Ludwigshafen, Germany) were cultured at 37°C, 5 % CO₂, in RPMI 1640 medium supplemented with 10 % horse serum, 5 % fetal calf serum, 4 mM L-glutamine, 200 U ml⁻¹ penicillin, 200 μg ml⁻¹ streptomycin and 2.5 μg ml⁻¹ amphotericin B, in poly-D-lysine-coated tissue culture flasks. For measurement, PC12 cells were plated on collagen-coated 25 mm glass coverslips. Nerve growth factor (200 ng ml⁻¹) was added to the medium and cells were incubated for a further 7 days to allow differentiation.

Determination of Ca²⁺ concentration

For fluorescence measurements cells were incubated with fura-2 AM. The loading buffer was Tyrode solution (mM: NaCl, 137; glucose, 5.6; KCl, 5.4; NaHCO₃, 2.2; MgCl₂, 1.1; NaH₂PO₄, 0.4; Hepes-Na, 10; CaCl₂, 1.8; pH 7.4) supplemented with 10 μM fura-2 AM and 0.025 % pluronic acid. Cells were incubated for 35–45 min in loading buffer at 37°C. Unloaded dye was washed out and coverslips were placed into a thermostatically controlled (26°C) chamber of a Nikon fluorescence microscope at ×400 magnification. Fluorescence intensity was monitored at an emission wavelength of 510 nm by altering excitation wavelengths between 340 and 380 nm using a rotating filterwheel (Applied Imaging, Newcastle upon Tyne, UK).

The sample interval was 40 ms for the determination of rate constants of depolarization-induced Ca²⁺ release with HK solution (Tyrode solution with 60 mM KCl and 80 mM NaCl) and 400 ms for nifedipine-induced Ca²⁺ release. No average was used in these cases. For all other measurements of [Ca²⁺]_i, four successive signals were averaged to reduce noise and the sampling interval varied between 1.6 and 5 s. Stored images were analysed using the QC 700 software package from Applied Imaging. For determination of [Ca²⁺]_i, regions of interests were defined covering the whole visible area of a cell. The light intensity values from these specified areas were integrated and exported into the SigmaPlot program (SPSS Inc., Erkrath, Germany). Background subtraction, ratioing and calculation of [Ca²⁺] were done off-line. Resting [Ca²⁺]_i was defined by the average of the first ten data points before a specified intervention. The magnitude of a [Ca²⁺]_i transient was defined by the peak value reached in response to a particular intervention.

Calibration of fura-2 fluorescence signals to calculate [Ca²⁺]_i values was done according to a procedure described by using the equation of Grynkiewicz et al. (1985). R_max (fluorescence ratio in the presence of saturating Ca²⁺), R_min (fluorescence ratio in the absence of Ca²⁺) and β (ratio of fluorescence values for Ca²⁺-bound/Ca²⁺-free indicator at 380 nm) were determined with 5 μM of the pentapotassium salt of fura-2 in solutions mimicking the intracellular milieu. The K_D of Ca²⁺ for fura-2 was assumed to be 224 nM; R_min was found to be 0.50, R_max 4.56 and β 4.81.

Electrophysiology

Whole-cell patch experiments were performed using a HEKA EPC-9 patch clamp amplifier with the pulse software package (HEKA Elektronik, Lambrecht, Germany). Series resistance compensation and leak subtraction were not used. Patch pipettes were pulled from PG10150 glass capillaries (WPI, Berlin, Germany) to a resistance of 1.8-2.5 MΩ. The intracellular solution contained (mM): CsCl, 130; EGTA-Na, 11; MgCl₂, 2; CaCl₂, 1; Hepes-Na, 10; ATP-Na, 4.2; pH 7.2. To measure currents through Ca²⁺ channels Ba²⁺ was used as a charge carrier; the extracellular solution contained (mM): CsCl, 110; BaCl₂, 25; Hepes-Cs, 10; CaCl₂, 2; pH 7.4. Cells were clamped to a holding potential of −110 mV. A cumulative dose-response curve for nifedipine was determined by single step depolarizations from −110 to 0 mV 20 s after application of the respective concentration of the substance. For assessment of current-voltage relationships step depolarizations to potentials between −60 and +50 mV were applied for 1000 ms with repolarization periods of 6 s. A prepulse to −25 mV for 30 ms followed by a 2 ms repolarization to −80 mV was used to inactivate occasionally occurring fast components of the current (Sipos et al. 1997). Activation parameters (half-maximal activation potential (V_0.5), slope factor (k)) were obtained by a non-linear fitting procedure of current-voltage relationships according to .

Measurements of resting membrane potential were performed in Tyrode solution using a pipette solution comprising (mM): KCl, 135; EGTA-Na, 11; MgCl₂, 2; CaCl₂, 1; Hepes-Na, 10; ATP-Na, 4.2; pH 7.2.

Experiments were done within a period of 3–6 days after changing the culture medium from GM to DM. For experiments we used multinucleated cells which were either contractile upon stimulation with HK or showed a rise in [Ca²⁺]_i when rinsed with Ca²⁺-free depolarization solution. According to the existing model of skeletal muscle E-C coupling, such cells were assumed to possess mature skeletal muscle-type DHP and ryanodine receptors and were regarded to be differentiated muscle cells.

Application of substances to the cells was performed by a superfusion system with a 7-channel perfusion pipette (List Electronic, Darmstadt, Germany), driven by a valvebank (TSE, Bad Homburg, Germany) with solution exchange times of less than 500 ms. Cells were constantly rinsed during measurements. Electrical stimulation was mimicked by depolarization with HK solution. Ca²⁺-free assay conditions were obtained by including 0.5 mM EGTA in solutions without any added Ca²⁺. Prior to the application of substances in the absence of Ca²⁺, cells were rinsed with Ca²⁺-free solution for a few seconds.

[³H]Ryanodine binding

Heavy sarcoplasmic reticulum (HSR) membranes and microsomal fractions (100000 g) from human skeletal muscle cells were prepared by differential centrifugation according to Wyskovsky et al. (1990) and subjected to high-affinity [³H]ryanodine binding. Binding experiments were carried out as described previously (Klinger et al. 1999). Briefly, HSR membranes or microsomal fractions from human skeletal muscle cells (50 μg) were incubated in 50 μl buffer containing 20 mM Hepes (pH 7.4), 200 mM KCl, 20 nM [³H]ryanodine, 100 nM aprotinin, 1 μM leupeptin, 100 μM Pefabloc, and a free Ca²⁺ concentration of 0.6 μM, which was adjusted with 1 mM EGTA and 0.9 mM CaCl₂. Nifedipine was added in the concentration range 0.01-30 μM and the corresponding concentrations of the solvent DMSO were used as controls. The incubation was carried out at 30°C for 40 min. The reactions were terminated by filtration over glass-fibre filters (pre-soaked in 1 % polyethylenamine) using a Skatron vacuum filtration device. Non-specific binding was determined in the presence of 1000-fold excess of unlabelled ryanodine, which had been added to the incubation mixture prior to addition of the labelled ligand. Non-specific binding was not affected by different assay conditions. The experiments were carried out in duplicate and repeated with two different HSR preparations and microsomal fractions.

All experiments were repeated at least three times. Data are given as means ±s.e.m.; n, number of cells analysed. Statistical analysis was performed using Student's t test.

Dihydropyridines elevate [Ca²⁺]_i in differentiated muscle cells

Two to 3 days after changing the medium from growth medium (GM) to differentiation medium (DM) cultured satellite cells started to fuse and formed multinucleated myotubes. Three to 6 days after initiation of differentiation the resting [Ca²⁺]_i in myotubes was 72 ± 1.7 nM (n = 163). Stimulation with HK solution led to a rapid rise in [Ca²⁺]_i (Fig. 1A). This effect could not be observed in cells which did not show the typical appearance of a differentiated muscle cell. The average maximal [Ca²⁺]_i in differentiated myotubes upon stimulation with HK was found to be 594 ± 48 nM (n = 49) in the presence of extracellular Ca²⁺. Stimulation of the cells in the absence of extracellular Ca²⁺ increased the [Ca²⁺]_i to only 442 ± 19 nM (n = 101). In addition to the smaller amplitude, a different shape of the Ca²⁺ transients was also observed under these conditions. When extracellular Ca²⁺ was omitted the Ca²⁺ transient was sharp. An accelerated inactivation (Brum et al. 1988; Schnier et al. 1993) is likely to be the reason for such a difference in shape.

Application of nifedipine to myotubes in concentrations from 30 nM to 30 μM caused a rise in [Ca²⁺]_i in a dose-dependent manner (Fig. 1). The nifedipine dose-response curve could be fitted to the Hill equation giving a maximal calculated amplitude of the nifedipine [Ca²⁺]_i rise of 80 ± 3.2 nM, an EC₅₀ of 614 ± 96 nM nifedipine and a Hill coefficient of 1.12 ± 0.16 (Fig. 1B). The [Ca²⁺]_i increase due to nifedipine was always smaller compared with the response obtained in HK (Fig. 1A) and was not able to induce contractions. The rate constant for the rise time of the nifedipine-induced [Ca²⁺]_i increase was smaller than that for the HK-induced increase (rate constant 0.1-0.2 s⁻¹ for nifedipine compared with a depolarization-induced Ca²⁺ transient of 2.5-5 s⁻¹; the latter was crucially dependent on the rate of rinsing). The nifedipine-dependent [Ca²⁺]_i stayed elevated for as long as nifedipine was applied (1 min in Fig. 1A and 1.5 min in inset to Fig. 6A) when extracellular Ca²⁺ was present. A complete reversal of the effect could be reached within 1–2 min after washout. Subsequent addition of nifedipine again triggered a rise in [Ca²⁺]_i (Fig. 1A). Mononucleated cells such as satellite cells and non-muscle cells did not respond to nifedipine. The possible participation of the solvent in the [Ca²⁺]_i transient was ruled out by the application of 0.1-0.3 % (v/v) DMSO to the myotubes. No increase in [Ca²⁺]_i could be observed (data not shown).

The observation that nifedipine was able to elevate [Ca²⁺]_i raises the question of whether nifedipine was acting as an antagonist of the L-type Ca²⁺ channel in these experiments. The whole-cell patch clamp technique was used to investigate the electrophysiological features of the L-type Ca²⁺ channel. Ba²⁺ currents through L-type Ca²⁺ channels were inhibited by nifedipine in a dose-dependent manner (Fig. 1A (inset) and B) with a calculated IC₅₀ value of 190 ± 6.5 nM and a Hill coefficient of 1.23 ± 0.05.

Nitrendipine, another DHP with antagonistic action on the L-type Ca²⁺ channel, was also able to increase [Ca²⁺]_i in a similar manner. The maximum rise observed was 82 ± 9.5 nM and, therefore, was comparable to that induced by nifedipine. The EC₅₀ value for nitrendipine (1.290 ± 0.64 μM) was about twice that for nifedipine (Fig. 2).

The agonist (±)Bay K 8644 also caused a rise in [Ca²⁺]_i in skeletal muscle cells (Fig. 3). Indeed, 10 μM (±)Bay K 8644 was able to trigger a [Ca²⁺]_i rise (67 ± 8.7 nM, n = 7). Furthermore, the simultaneous application of nifedipine and (±)Bay K 8644 resulted in no further rise in [Ca²⁺]_i. This suggests that nifedipine and (±)Bay K 8644 act via a common mechanism resulting in elevated [Ca²⁺]_i.

It has been demonstrated that phosphorylation of the tetrodotoxin (TTX)-sensitive Na⁺ channel makes it promiscuous with respect to ion selectivity, permitting Ca²⁺ entry (Santana et al. 1998). In order to exclude Ca²⁺ influx due to this so-called ‘slip mode’ conductance of Na⁺ channels, we used 1 μM TTX to block the Na⁺ channels. Na⁺ channel block had no detectable influence on the nifedipine-induced response of [Ca²⁺]_i (data not shown).

To clarify whether nifedipine was able to increase [Ca²⁺]_i in cells other than skeletal muscle cells, we used the well characterized PC12 cells. These cells are known to express both voltage-dependent L-type Ca²⁺ channels (Liu et al. 1996) and intracellular Ca²⁺ release channels, i.e. InsP₃ and ryanodine receptors (Bennett et al. 1998). Nevertheless, a coupling between the DHP receptor and the ryanodine receptor, as in skeletal muscle cells, does not exist. The application of depolarizing solution to these cells elicited a [Ca²⁺]_i transient which was caused solely by an influx of Ca²⁺, as shown by the inability of these cells to react to depolarization under Ca²⁺-free conditions (Fig. 4). Furthermore, nifedipine was not able to stimulate a [Ca²⁺]_i response in PC12 cells but it decreased the amplitude of the Ca²⁺ transient by about 50 % when cells were depolarized by HK (Fig. 4).

Source of Ca²⁺

Three mechanisms responsible for the intracellular [Ca²⁺] rise in skeletal muscle cells must be considered: influx of extracellular Ca²⁺, Ca²⁺ release from intracellular Ca²⁺ pools through ryanodine receptors and Ca²⁺ release from intracellular pools through InsP₃ receptors.

We are grateful to P. Artigas from the Universidad de la Republica, Uruguay, for critical reading and comments on the manuscript. This work was supported by research grants from the Anton Dreher-Gedächtnisstiftung and the OeNB Jubiläumsfonds (6646 and 8266).

Parts of this material were presented at the 43rd Annual Meeting of the Biophysical Society, 13–17 February 1999, Baltimore, MA, USA.

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