1Department of Anesthesia Research, Brigham and Women's Hospital, Harvard Medical School, Boston, Massachusetts 02115; 2Department of Cell and Developmental Biology, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania 19104; 3Centro de Biofísica y Bioquímica, Instituto Venezolano de Investigaciones Científicas, Caracas, Apartado 21827, Venezuela; and 4Laboratory of Cellular Physiology, CeSI, Center for Research on Aging, University G. d'Annunzio School of Medicine, 66023 Chieti, Italy
Submitted 12 May 2003 ; accepted in final form 31 August 2003
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ABSTRACT |
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calcium-induced calcium release; dihydropyridine receptors; excitation-contraction coupling; ryanodine receptors; skeletal muscle
DHPRs and RyRs are the two primary proteins involved in the E-C coupling in skeletal, cardiac, and smooth muscle. However, it has become increasingly evident that the communication between these two proteins in skeletal muscle employs a different mechanism compared with the cardiac and smooth muscle. In cardiac and smooth muscle, activation of the 1DHPR generates a large inward flux of Ca2+ from the extracellular space, which induces the opening of RyRs and a consequent massive release of Ca2+ into the myoplasm. This mechanism has been defined as calcium-induced calcium release, or CICR (6). In skeletal muscle, however, initiation of a contraction can be achieved in the absence of the extracellular Ca2+ (3, 4). In fact, it has been generally accepted that in skeletal muscle
1SDHPR functions predominantly as the voltage sensor (24, 30) that activates the RyRs though a physical interaction, known as the orthograde signaling. Although the role of Ca2+ influx in skeletal muscle E-C coupling is unclear, Ca2+ ions do participate in modulation of the RyR activity through a process similar to that of cardiac CICR (5a, 1719).
In the present work, we report the existence of spontaneous, RyR-mediated Ca2+ transients, as well as spontaneous depolarizations of the plasma membrane in primary myotubes of skeletal origin. The ability of Ca2+ channel blockers to eliminate these events indicates that the repetitive pattern of this activity is dependent upon the influx of Ca2+ through the DHPRs and suggests that Ca2+ current through 1SDHPRs may play a relevant role in developing skeletal muscle cells.
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MATERIALS AND METHODS |
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Immunohistochemistry. The cells were fixed in methanol for a minimum of 20 min at 20°C. To avoid nonspecific detection, cells were blocked for 1 h in PBS supplemented with 1% BSA and 10% goat serum. Cells were incubated at room temperature with the appropriate primary antibody for 2 h and then washed three times for 10 min with PBS/BSA before being incubated for 1 h with secondary antibodies. Code, specificity, working dilution, and the sources of primary antibodies used in single staining experiments are as follows: anti-RyR 34C, 1:30 (2) (Developmental Studies Hybridoma Bank, The University of Iowa); sheep anti-1SDHPR, 1:500, Upstate Biotechnology, Lake Placid, NY. Secondary antibodies were conjugated with either cyanine 3 or cyanine 5 (Jackson ImmunoResearch Laboratories, Lexington, KY). The specimens were viewed on a laser scanning confocal microscope (Zeiss LSM510, Specifics) interfaced with an inverted Zeiss Axiovert microscope.
Fluorescence measurements. Intracellular Ca2+ imaging was performed as described previously (22, 26). Briefly, the differentiation media were removed and cells were washed twice with imaging buffer (IB) containing 125 mM NaCl, 5 mM KCl, 1.2 mM MgSO4, 6 mM glucose, 25 mM HEPES, 0.05% BSA, 2 mM CaCl2, pH 7.4 (for those conditions where depolarization of the myotubes was required, IB contained 50 mM NaCl and 80 mM KCl). Cells were then loaded for 30 min with Ca2+ indicator dye (fluo 4-AM, 10 µM) and washed several times with IB to terminate further loading. Whole cell fluorescence changes were detected using PTI delta-RAM as the light source with a 12-bit digital intensified charge-coupled device (Stanford Photonics) interfaced with an inverted microscope equipped with an Olympus Uapo/340 x40 oil immersion objective. Changes in intracellular Ca2+ were characterized as changes in fluo 4 fluorescence intensity. All experiments were conducted at room temperature (22°C). Solution exchange within each well was achieved via pressure controlled perfusion system (Automate Scientific, Berkley, CA). The perfusion inlet was positioned close to the cells to allow a very efficient and rapid change of solution. Detected changes in fluorescence from the regions of interest within each cell were analyzed using QED imaging software (QED Software, Pittsburgh, PA). The resulting fluorescence changes were corrected for the background fluorescence within individual cells by dividing the value of the fluorescence intensity at each measured interval by the mean fluorescence intensity of a 30-s quiescent period within that cell to give the F/F0 values.
Microelectrode preparation and membrane potential recording. Microelectrodes used in the recording of the membrane potential were prepared from thin-walled 1.5/1.12 mm internal diameter borosilicate glass capillaries with internal filaments (WPI-TW150-4). Before pulling, the capillaries were washed with 1 M HCl and distilled water and dried at 150°C for 3 h. The clean glass capillaries were drawn into microelectrodes by using a Flaming Brown puller model P-87 (Sutter Instruments, San Francisco, CA). The microelectrodes were backfilled with filtered 3 M KCl immediately before use and had a tip resistance ranging from 10 to 15 M. The bath reference electrode was an Ag-AgCl pellet.
Single myotubes were carefully impaled with the aid of an inverted compound microscope (Axiovert 10) fitted with a x10 eyepiece and a x40 dry objective. The potential from the 3 M KCl barrel (Vm) was recorded with a WPI high-impedance amplifier F-223A (Sarasota, FL). The Vm potential was filtered at 510 KHz to improve the signal to noise ratio and was stored for further analysis. All membrane potential recordings were carried out at 22°C.
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RESULTS |
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To determine whether the spontaneous activity was generated by Ca2+ release through the RyRs, the myotubes were incubated with 0.5 mM ryanodine for 30 min at 37°C. Ryanodine is a plant alkaloid that binds specifically to the open state of the RyRs and at concentrations above 10 µM locks the channel in a conformation that does not allow for the release of Ca2+ (35). As demonstrated in Fig. 1C, application of the ryanodine resulted in complete elimination of all detectable Ca2+ release activity, suggesting that the observed oscillations in Ca2+ occurred as a direct result of the opening of the RyRs.
Expression of key E-C coupling proteins and formation of Ca2+ release units. It has been previously reported that immunolabeling of either RyR1 or 1SDHPRs in skeletal muscle myotubes results in a characteristic punctate pattern localized at the periphery of the cell. This pattern indicates clustering of the RyRs and DHPRs and corresponds to the formation of junctions, or calcium release units (CRUs), between SR and exterior membranes in developing myotubes (10, 25, 27). Colocalization of
1SDHPRs and RyR1s is also an indication of correct assembly of skeletal CRUs (10, 27). As demonstrated in Fig. 2, A and B, wild-type myotubes exhibit a punctate pattern of fluorescence when immunolabeled with either anti-RyR1 or anti-
1SDHPRs antibodies, respectively. The two proteins are not only clustered in bright foci but are also colocalized as demonstrated in Fig. 2C, indicating the formation of functional CRUs.
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Spontaneous depolarizations in myotubes. As demonstrated in Fig. 1, the observed spontaneous Ca2+ oscillations generally appeared at the same frequency and amplitude within each given cell. The uniformity of the release suggests that this activity might be controlled in part by changes in the plasma membrane potential. We have observed that in addition to the cells exhibiting stable resting membrane potential, a subset of cells (30%) exhibited spontaneous fluctuations in the membrane potential. Figure 3, A and B, shows membrane potential recordings from a myotube that does not exhibit spontaneous fluctuations in the membrane potential, and from one that does, respectively. Consistent with previous reports (13, 15, 34), the average value of the resting membrane potential was 62 ± 0.44 mV (n = 48). The recorded oscillations in the membrane potentials did not appear to resemble action potentials, because the magnitude of the depolarization, 26 ± 1 mV (n = 146), was insufficient to reach the action potential threshold. However, on the basis of previous reports, these depolarizations should be sufficient to activate Ca2+ influx through the L-type Ca2+ channels (32). The percentage of cells tested in each culture exhibiting spontaneous membrane depolarizations correlated with the percentage of cells that exhibited spontaneous intracellular Ca2+ oscillations as described in Fig. 1. Analogous to the changes in the intracellular Ca2+, the spontaneous depolarizations occurred without any stimulation of the cells.
Role of DHPR and Ca2+ influx in spontaneous oscillations. Because membrane depolarizations activate the voltage sensors in the surface membrane/TT, we sought to determine whether DHPR is directly involved in eliciting these events. To do so, we applied 5 µM nifedipine to the extracellular bathing solution. Nifedipine is a DHP-specific antagonist, which promotes the inactivation of the channel (21, 29) and has a blocking effect on the Ca2+ current through the TT membrane (21). As demonstrated in Fig. 4, application of nifedipine abolished the spontaneous RyR-mediated Ca2+ release. However, nifedipine could not abolish the KCl-induced Ca2+ transients, which could still be elicited in these cells. Further evidence for the involvement of the DHPRs in the spontaneous activity was obtained from experiments conducted with muscular dysgenesis myotubes (mdg), which do not express any functional DHPRs (16, 23). This phenotype renders DHPRs in these cells unable to conduct Ca2+ (1) or to participate in E-C coupling (31). None of the tested mdg myotubes exhibited any type of spontaneous activity (Fig. 5). To determine whether these cells expressed functional RyR Ca2+ release channels, mdg cells were challenged with 40 mM caffeine. All tested cells produced a robust Ca2+ transient in response to the caffeine, indicating a sufficient expression of the RyRs and the viable status of the cells. From these results, it could be inferred that although Ca2+ oscillations occurred without any external stimuli, they were under control of the DHPRs.
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Ca2+ influx through DHPR. To confirm that the abolition of spontaneous activity by nifedipine was based on its inhibition of Ca2+ influx and not due to a conformational inactivation of the DHPR, we tested the effects of Cd2+ and La3+, which have been previously described as potent blockers of the 1SDHPR channel pore (12, 31). As shown in Fig. 6, application of Cd2+/La3+ completely blocked the spontaneous Ca2+ oscillations and did so in every cell that exhibited this phenomenon. Washing the cells with the Cd2+/La3+-free solution could reverse the Cd2+/La3+ effects. In most cases, the spontaneous activity returned in those cells that exhibited this activity before the application of the Cd2+/La3+. The presence of Cd2+/La3+ did not interfere with the large Ca 2+ transient elicited by depolarization, showing that similarly to nifedipine, Cd2+/La3+ blocked spontaneous activity but not the coupling between
1SDHPRs and RyR1.
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DISCUSSION |
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The spontaneous Ca2+ transients were observed in a subpopulation of differentiated myotubes and depending on the relative level of cellular differentiation, constituted 30 to 50% of the entire cell culture. The difference in differentiation rate is due to a number of factors including distribution of myoblasts on the culture plate and regional differences in the concentrations of excreted growth factors. These global oscillations in intracellular Ca2+, although exhibiting variable frequency and amplitude between the cells, were not initiated by any type of applied stimulus and were always significantly smaller then the Ca2+ transients detected upon cellular depolarization with KCl. Similar spontaneous Ca2+ transients have been previously reported in the cultured human (33) and chicken myotubes (9) and C2C12 mouse cell line (9), as well as embryonic Xenopus myocytes (8). However, one of the major differences from previous reports is that the events reported here appear to require the influx of extracellular Ca2+, seemingly through the DHPRs.
There are several lines of evidence suggesting that this type of activity is not simply random Ca2+ release from the SR. The fact that the release was uniform throughout the cell and that the oscillations appeared at fixed intervals and with fairly constant amplitudes suggests that there are specific cellular factors that govern the initiation of each oscillation. Results in Figs. 4 and 6 show that by blocking Ca2+ influx into the cells either by nonspecific cation channel blockers, such as Cd2+ and La3+, or more selectively by nifedipine, these oscillations could be completely inhibited. The fact that application of either of these blockers does not eliminate the depolarization-elicited Ca2+ transients, that is, the functional components of the skeletal E-C coupling were still preserved, suggests that the only cause for the change is the abolition of the DHPR Ca2+ current. And the fact that intracellular Ca2+ oscillations reappear upon the washout of the blockers that restrict the flow of Ca2+ suggests that the Ca2+ influx is a necessary component of the initiation of this type of activity.
We also demonstrate that a population of myotubes exhibits spontaneous depolarizations of the plasma membrane. It should be pointed out that the observed depolarizations did not resemble typical action potentials. Because the subthreshold depolarizations were not of sufficient magnitude (35 mV) to reach the threshold of initiation of an action potential, the observed depolarizations could therefore not possibly elicit action potentials. However, these depolarizations must be large enough to activate L-type Ca2+ channels to a level sufficient to generate a significant Ca2+ influx. Although the membrane potential recordings were performed independently of the intracellular Ca2+ measurements, we believe that because the frequency of occurrence of these two observations was similar in both preparations, the two phenomena are related to the same process. If the spontaneous depolarizations precede the activation of Ca2+ release from the SR, then it is possible that they activate the voltage sensors in the TT and, subsequently, the DHPR Ca2+ channels.
It has been generally accepted that skeletal muscle, unlike cardiac or smooth muscle, does not require the influx of extracellular Ca2+ to achieve contraction. One of the reasons it is believed that the influx of Ca2+ is simply a vestigial process is because the kinetics of activation are too slow and the magnitude of the current is simply too small for Ca2+ to diffuse rapidly from the DHPR Ca2+ channel and overcome the Mg2+ inhibition of the RyR. To achieve Ca2+ influx-induced Ca2+ release in skeletal muscle, the skeletal DHPRs would have to behave similar to those of cardiac type with respect to the magnitude and the kinetics of the Ca2+ influx. This condition could be potentially achieved in the skeletal muscle if the cells were stimulated by repetitive depolarization (7, 11), analogous to those exhibited in Fig. 3. It has been previously reported that repetitive depolarizations of skeletal muscle fibers at short intervals, as infrequently as 1.7 Hz, result in acceleration, as well as in the potentiation, of the Ca2+ currents (7, 11). Additionally, it has now been suggested that the magnitude of Ca2+ influx through the skeletal DHPR could be sufficient to induce CICR (14). Thus it is conceivable that the initiation of Ca2+ release within each oscillation could be triggered by the Ca2+ entry through the 1s of DHPRs (5).
In summary, skeletal myotubes exhibit spontaneous oscillations in intracellular Ca2+, as well as spontaneous depolarizations of the plasma membrane. Pharmacological data indicate that initiation of Ca2+ oscillations in developing skeletal muscle cells is dependent on the Ca2+ influx through the 1s-subunit of the DHPR.
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ACKNOWLEDGMENTS |
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GRANTS
This work was supported by National Institute of Arthritis and Musculoskeletal and Skin Diseases (NIAMS) Grant P01 AR-44650 (to P. D. Allen), Muscular Dystrophy Association Grant MDA 2688 (to P. D. Allen and F. Protasi), and NIAMS Grant AR-49160-2 (to A. Shtifman).
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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