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MS 9551 Received 26 April 1999; accepted after revision 26 July 1999.
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ABSTRACT |
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(the 'hump' component of charge movement).
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INTRODUCTION |
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In skeletal muscle, Ca2+ is released from the sarcoplasmic reticulum (SR) when a signal is transmitted across the triad as a result of depolarization of the transverse tubular system (T-system). In this process, dihydropyridine (DHP) receptors of the transverse tubules play an essential role as voltage sensors of excitation-contraction (E-C) coupling that produce the charge movement recorded during electrophysiological experiments (for reviews see Rios & Pizarro, 1991; Lamb, 1992; Melzer et al. 1995). In addition to their role as voltage sensors, DHP receptors are also permeable to Ca2+, giving rise to very slowly activated Ca2+ currents (for a review, see Melzer et al. 1995).
In this study, we present a novel preparation for recording the electrical activity of DHP receptors, using spherical vesicles and the whole-cell voltage clamp technique. This preparation has the advantage of allowing control of the composition of the internal medium, as well as the recording of L-type currents, without the retrograde influence of the SR. In addition, recordings of L-type currents can be made without the complications present in intact cells, where complete control of the membrane potential is difficult to achieve due to the presence of the T-system.
We found major changes in the amplitude and time course of the currents, compared with those present in cells. Non-linear capacitive currents with properties of charge 1 were also recorded.
Preliminary results have been published (Camacho & Sanchez, 1997; Camacho et al. 1998).
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METHODS |
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Preparation
Spherical vesicles derived from the plasma membrane of frog skeletal muscle were used. Frogs were killed by decapitation. All experiments were carried out according to the guidelines of the Animal Care Committee in México. The procedure used to produce vesicles by enzymatic treatment was that originally described for single channel experiments by Standen et al. (1984) and later modified for 'whole vesicle' recordings by Camacho et al. (1996). In brief, semitendinosus muscles from Rana montezumae were incubated in 120 mM KCl, buffered at pH 7·2 with Hepes (5 mM), with added collagenase (Sigma, Type IA, 50 units ml-1). This enzymatic treatment does not produce major changes in the single channel behaviour of Na+ or K+ channels present in the vesicles (Standen et al. 1984) and collagenase by itself does not significantly affect the electrophysiological properties of DHP receptors in enzymatically dissociated skeletal muscle fibres (Szentesi et al. 1997). Vesicles formed spontaneously after a period of about 60 min at 20-22°C. Recordings were carried out at 15-17°C.
Solutions
The external solution employed to record Ba2+ currents (IBa) contained (mM): Ba2+, 30; TEA+, 110; and methanesulphonate (CH3SO3-), 170, as anion. The composition of the external solution used to record charge movement was (mM): TEACH3SO3, 110; CaCl2, 10. The pipette solution contained (mM): CsCH3SO3, 125; MgCl2, 2 or 0·1; and EGTA, 1. External and internal (pipette) solutions were buffered with Hepes (5 mM) at pH 7·2 and 7·1, respectively. All chemicals were obtained from either Sigma or Aldrich Chemical Co.
Electrophysiological methods
The whole-cell configuration of the patch clamp technique was used (Hamill et al. 1981). Pipettes used were as described by Camacho et al. (1996) and were double-pulled from hard glass (KIMAX-51; Kimble Glass, Toledo, OH, USA) using a David Kopf (Tujunga, CA, USA) 700D vertical puller. The tips had resistances of about 6-8 M
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Data collection and pulse protocol
Membrane currents (Im), produced in response to voltage step depolarizations applied from the holding potential (Vh), were measured with an Axopatch amplifier (model 200A, Axon Instruments) and sampled by an IBM-PC/AT-compatible Pentium-based microcomputer. Analog signals were digitized to a resolution of 12 bits through a LabMaster interface (TL-1 DMA interface, Axon Instruments) that also generated the command pulses. Data were analysed with a combination of pCLAMP (version 6.0, Axon Instruments) and in-house software. Im was amplified and filtered with an active 4-pole, low-pass Bessel filter set at a corner frequency of no more than half the sampling frequency. To measure the activation of IBa, command pulses of 500 ms duration and variable amplitude were delivered. The interval between pulses was at least 1 s to avoid changes in channel kinetics by a previous depolarization, as described by Feldmeyer et al. (1990). The pulse sequence was bracketed by five consecutive hyperpolarizing control pulses, -20 mV from Vh that ranged between -80 and -100 mV. The currents generated during the hyperpolarizing pulses were used to calculate the linear membrane capacitance and to subtract the leakage current from the currents generated by the test pulses. In some cases, currents generated by +20 mV pulses applied from a Vh of -100 mV were also used for this purpose. Steady-state inactivation was investigated by delivering 5000 ms prepulses to several potentials followed by 500 ms test pulses to +30 mV. Movement of charge 1 was measured in polarized vesicles by delivering 100 ms pulses to preselected depolarizing potentials. Non-linear currents were obtained off-line after subtraction of the linear components generated by the test pulses. To this end, a control current was used. Integration of currents elicited by negative pulses, relative to Vh, showed that the currents generated by these pulses were mostly linear (see Results). For this reason, a control current was constructed by adding together the currents generated by all the hyperpolarizing pulses, and the sum was subtracted, after appropriate scaling, from the current generated by each one of the test pulses. This procedure produced much lower noise levels than the protocol in which only one of the currents elicited by the hyperpolarizing pulses was used. To maintain the stability of our recordings, we added Ca2+ to the external solutions used in charge movement experiments. However, the presence of Ca2+ tail currents contaminated the records of charge movement during 'off' transients. For this reason, charge movement was measured only during 'on' transients. To determine whether charge 2 was present in the vesicles, pulses ranging from +60 to -170 mV in -10 mV steps were applied from a Vh of 0 mV.
The voltage dependence of activation of charge movement was fitted to the Boltzmann function:
| Q = Qmax/(1 + exp((V - Vm)/k)), | (1) |
where Qmax is the maximum charge, Vm is the membrane potential, V is the potential at which Q = 0·5Qmax and k is a measure of the steepness of the curve.
The voltage dependence of inactivation of Ba2+ currents was fitted to a function of the same form as eqn (1) but with (Vm - V) in the exponential.
A non-linear least-squares algorithm was used to fit numerical formulae to experimental data. Parameter values given in the text and Table 1 are expressed as means ± S.E.M.; n, number of experiments. Student's t test was used to calculate statistical significance, with P < 0·05 being considered significant.
Table 1. Charge movement in vesicles
| n | Qmax (nC µF-1) |
V (mV) |
k (mV) |
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| Charge 1 (control) | 21 | 45·6 ± 5·5 | -58·4 ± 4·3 | 22·3 ± 1·5 |
| Charge 1 (nifedipine) | 11 | 22·9 ± 4·5 * | -43·9 ± 8·6 | 19·9 ± 2·4 |
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RESULTS |
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Ba2+ currents in vesicles
L-type Ca2+ channels are present in the vesicles. Figure 1A illustrates membrane currents obtained during voltage steps to the potentials indicated. For depolarizations up to -10 mV, ionic currents were mostly linear and were cancelled out by the subtraction procedure. With larger depolarizations, inward Ba2+ currents were recorded whose time course became faster with further depolarizations. These non-linear currents were not maintained and were inactivated almost completely during large depolarizing pulses. These currents were preceded by non-linear outward currents or 'on' currents at the beginning of the pulses that correspond to the brief outward deflections shown in Fig. 1A and B. These currents represent the movement of a non-linear charge that it is described below. Immediately after the pulses, large inward non-linear currents were recorded. These will be referred to as 'off' currents. Two components contributed to the 'off' currents: an initial fast large component and a relatively small slow component. The amplitude of the fast component increased with voltage in spite of the fact that inward Ba2+ currents showed a pronounced inactivation during large voltage steps. This was followed by a small component whose magnitude did not change greatly within the same voltage range. As shown in more detail below, two distinct components contributed to these currents: an inward Ba2+ tail current component, arising from deactivation of the channels, and a non-linear capacitive current component.
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A, current records during voltage steps to the potentials indicated (in mV) after subtraction of linear membrane components. Vh = -80 mV. B, effect of Bay K 8644 (1 µM) on membrane currents recorded in a separate experiment at the same potentials as in A. Note that the amplitude scale in B differs from that in A. Vh = -80 mV. C, relation between peak current and membrane potential from the experiments illustrated in A ( | ||
Figure 1C (open symbols) shows the current-voltage relation from the same experiment. Activation of Ba2+ currents began at about 0 mV. The peak Ba2+ current was largest at a membrane potential of +20 mV; it declined at larger depolarizations and was still inward at +70 mV, indicating a very positive reversal potential. The possibility that IBa flows through DHP-sensitive receptors was investigated. Figure 1B shows inward Ba2+ currents from a vesicle incubated in the presence of the DHP agonist Bay K 8644. Compared with records obtained under control conditions (Fig. 1A), the currents activated at similar potentials and had a similar time course. However, the amplitude of the currents was distinctly larger at all potentials tested. The current-voltage relation of this experiment is shown in Fig. 1C (filled symbols). In 14 experiments, mean peak IBa was -3·0 ± 0·7 µA µF-1 in control solution and -6·8 ± 1·5 µA µF-1 in the presence of Bay K 8644. There was also an increase in the amplitude of the 'off' currents, especially of the fast component at low depolarizations, as shown in Fig. 1B.
L-type currents were not sustained but declined during the depolarizing pulses, indicating that Ca2+ channels inactivate. The inactivation properties of IBa were examined in experiments using a double-pulse protocol, as first described for sodium channels (Hodgkin & Huxley, 1952). A representative experiment is illustrated in Fig. 2. The inset shows superimposed membrane currents (top) generated by the pulse protocol (bottom) and described in Methods. Membrane currents are shown after subtraction of linear membrane components. Consistent with the results illustrated in Fig. 1, Ba2+ currents inactivated during the test pulse. In addition, we found that, as the amplitude of the prepulse increased, the magnitude of IBa during the test pulse decreased progressively to zero, as expected from a voltage-dependent inactivation process. In Fig. 2, the symbols represent relative peak values of IBa. The continuous line is the best fit of the inactivation function (eqn (1)) with the parameters given in the legend. In three experiments, the mean value of the mid-point of steady-state inactivation was -40·2 ± 4·7 mV with a slope of 14·8 ± 5·4 mV.
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The membrane potential was stepped from -80 mV to the conditioning voltage (abscissa) for 5 s. Peak current in response to a test potential to +30 mV was measured and plotted relative to IBa elicited without a prepulse (ordinate). The continuous line is the best fit of a Boltzmann function with V = -30·8 mV and k = 8·0 mV. The inset shows original records from the same experiment (top) and the pulse protocol (bottom). | ||
Facilitation of Ba2+ currents in vesicles
Facilitation of Ca2+ channels of mammalian skeletal muscle has been described previously as an increase in the amplitude of tail currents that depends on the duration of the preceding depolarizing pulse. This potentiation is absent in dyspedic myotubes (prepared from mice homozygous for a disrupted ryanodine receptor gene), and apparently depends on a retrograde signal from the SR to DHP-sensitive channels (Fleig et al. 1996). Although this phenomenon has not been described in frog skeletal muscle, it was interesting to test the effect of different pulse durations on tail currents and to determine whether facilitation was present in the vesicles. Figure 3A shows unsubtracted membrane currents produced by the pulse protocol shown in Fig. 3D. Membrane currents were elicited by depolarizing pulses to +60 mV, whose duration progressively increased. Currents obtained after subtraction of linear current components are shown in Fig. 3B. There was a small and brief outward current at the beginning of each pulse, which was similar to those illustrated in Fig. 1A and B. These currents were followed by an inward Ba2+ current that partially inactivated during the pulses. Immediately after the depolarizing steps, 'off' currents were recorded; the amplitude of the fast component decreased as the duration of the depolarization increased. In 13 experiments, the amplitude of the fast component after a 500 ms pulse decreased to 66 % (± 6 %) of its value after a 50 ms pulse. In contrast, the magnitude of the slow component (indicated by the arrow) developed progressively, and its amplitude increased as the pulse duration increased. An alternative way of showing the presence of this non-linear current component, which only makes use of the test currents illustrated in Fig. 3A without the involvement of leakage currents, is shown in Fig. 3C. The membrane current (which contains linear and non-linear components) shown in Fig. 3A, generated immediately after the depolarization lasting 50 ms, was subtracted from the corresponding currents generated after longer depolarizations, i.e. i - 1 (where i = 2, 3, . . . 10), as indicated by the numbers in Fig. 3A and C. If the amplitude of the fast component decreases as the pulse is lengthened, this subtraction procedure would be expected to give rise to outward deflections. This was indeed the case, as shown in Fig. 3C. These results confirm that the amplitude of the fast component decreased over time. Also, consistent with the results shown in Fig. 3B, the amplitude of the slow component (downward deflections) increased as the duration of the pulse increased. In contrast to the facilitation of the tail current observed in wild-type muscle cells by Fleig et al. (1996), we did not observe a large increase in the amplitude of the tail currents by increasing the duration of the pulses. Instead, the amplitude of the fast component consistently decayed, as mentioned above, and even though the magnitude of the slow inward component increased after a 500 ms pulse to +60 mV (measured as illustrated in Fig. 3C), it represented only 9·6 % (± 3·4 %, n = 13) of the magnitude of the fast component of the 'off' current generated after a 50 ms pulse to the same potential. This indicates that facilitation is almost absent in the vesicles.
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A, records showing unsubtracted currents generated with the pulse protocol shown in D. Numbers indicate the pulse sequence. B, currents in A after subtraction of linear current components. The arrow indicates the presence of a slow inward component. C, subtraction of 'off' currents shown in A. The subtraction procedure is indicated by the numbers below the panel. Note that the amplitude scales of panels A-C are different. D shows the pulse protocol. | ||
Actions of internal Mg2+ on Ba2+ currents
It has been shown previously that a low intracellular [Mg2+] potentiates calcium release in frog muscle (Jacquemond & Schneider, 1992). To investigate the possibility that this effect may be due in part to alterations in the voltage dependence of DHP receptors, we tested the actions of internal Mg2+ on the time course of L-type currents, which is highly dependent on membrane potential. Figure 4 shows the relationship between the time to peak (left ordinate) and voltage. Open symbols represent the mean values (± S.E.M.) from several experiments performed in the presence of 2 mM MgCl2. As the amplitude of the depolarizing pulse increased, the time required to reach the peak value of IBa decreased over the whole voltage range. The time to peak declined progressively reaching a limiting value of 17 ms at large depolarizations. In the presence of low [Mg2+] (0·1 mM), we found that channel activation was shifted by -13 mV, as shown in Fig. 4 (filled symbols). On the other hand, we observed no significant changes in the peak amplitude of IBa in solutions containing low [Mg2+]. In the same experiments, mean peak IBa was 2·72 ± 0·39 µA µF-1 (n = 7).
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The membrane potential of the vesicles was stepped from -80 mV to the membrane potentials indicated (abscissa). The left ordinate corresponds to the experimental values. | ||
Charge movement in vesicles
We found that non-linear capacitive currents could be recorded from the vesicles. Non-linear currents were obtained after subtraction of linear current components using an appropriate control membrane current record. Figure 5A shows non-linear membrane current records obtained at the potentials indicated from a Vh of -100 mV. Non-linear currents were evident after the subtraction procedure, especially for large depolarizing membrane potentials. These currents decayed during the 'on' and 'off' transients, without any indication of different components or 'humps' contributing to the records. Due to the presence of Ca2+ tail currents (see below), we integrated the currents during 'on' transients only. As expected from a capacitive current, the charge saturated at large depolarizations. Figure 5B shows the charge mobilized during the depolarizing pulses as a function of membrane potential. The values of charge during the hyperpolarizing pulses after subtraction of the control current are also illustrated (see Methods). As expected, these values were very close to zero. The symbols represent results from two runs performed in the same vesicle. The continuous line is the best fit of a two-state Boltzmann function to the experimental values. The mean fit parameters from several experiments under control conditions and in the presence of nifedipine (10 µM) are shown in Table 1. Nifedipine reduced the value of Qmax by half.
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A, macroscopic membrane currents in vesicles. The records show non-linear currents during voltage steps to the potentials indicated (in mV) from Vh = -100 mV. B, non-linear charge movement as a function of voltage. The continuous line is the best fit of a Boltzmann function with Qmax = 37·2 nC µF-1, V = -65·2 mV and k = 16·4 mV. | ||
To record non-linear currents, we also measured the area under the unsubtracted current records as an alternative means of showing the presence of non-linear charge movement. This gives an estimate of the total charge transferred by the pulses. First, pulses from +60 to -180 mV in -10 mV steps were applied to a depolarized vesicle (Vh = 0 mV). Then, the holding potential was set to -100 mV and a 3 min interval was allowed before another sequence of pulses that spanned the same voltage range starting from -180 mV was delivered. Figure 6A shows the relation between the total charge (ordinate) and the membrane potential (abscissa) after the vesicle was held at -100 mV. Open symbols represent 'on' charge and filled symbols the charge under 'off' transients. The straight line is the best fit of a linear function to the data points between -180 and -100 mV extrapolated to the whole voltage range. For large depolarizations, the data points lay above the straight line indicating the presence of an excessive charge in polarized vesicles. 'Off' charge was distinctly larger than 'on' charge suggesting the presence of Ca2+ tail currents. This was confirmed after block of L-type Ca2+ currents with nifedipine (10 µM) in polarized vesicles held at -100 mV. Under these conditions, the ratio of 'on' and 'off' charges at +50 mV was 0·96 ± 0·05 (n = 7), indicating equality of charge. This indicates that the 'off' currents, present after nifedipine blockade, are capacitive in nature. A clear separation in the values of 'on' and 'off' charges in the control solution began with depolarizations to -30 mV. In contrast, activation of IBa during step depolarizations became evident at about 0 mV (Fig. 1). The contamination of 'off' charge by Ca2+ tail currents at lower potentials may be due to the difference in the ionic composition of the solutions. The presence of high concentrations of divalent cations would in fact shift the voltage dependence of channel activation towards more positive potentials, due to surface charge effects (Hille, 1992).
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Symbols represent integration of 'on' ( | ||
Values for non-linear charge movement in polarized vesicles were obtained as the difference between the total 'on' charge at each membrane potential and the linear charge calculated by extrapolation of the straight line. These values were fitted to a Boltzmann function. Table 1 shows the mean values of the Boltzmann parameters fitted to non-linear charge movement obtained in several experiments.
Evidence for non-linear currents when the vesicle was depolarized is shown in Fig. 6B, which shows the relation between total charge and membrane potential from the same experiment as that in Fig. 6A. Vh was maintained at 0 mV. An excessive 'on' charge appeared in the hyperpolarizing range of membrane potentials. A straight line was fitted to the data points obtained with positive potentials and extrapolated to negative values. The 'on' transients associated with hyperpolarizing pulses were not blocked by nifedipine (data not shown). On the other hand, 'off' charge values at negative potentials lay very close to the straight line indicating that 'off' transients are linear within the whole voltage range. Similar results were obtained in another eight similar experiments. Examples of original records from this experiment are shown in the insets to Fig. 6A and B.
The slopes of the straight lines in Fig. 6A and B correspond to the vesicle capacitance under polarized and depolarized conditions, respectively. There was an increase in the membrane capacitance when the vesicles were depolarized. The mean ratio of the slopes in depolarized and repolarized vesicles was 1·23 ± 0·03 (n = 8).
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DISCUSSION |
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This study describes the properties of ionic currents and charge movement recorded from L-type Ca2+ channels in vesicles derived from frog skeletal muscle. These vesicles are free of internal particles or cytoplasmic organelles and lack invaginations (Camacho et al. 1996). These features are most convenient for studying the activity of DHP-sensitive Ca2+ channels under better voltage clamp conditions than would be possible to achieve in skeletal muscle fibres, where voltage control of the T-system is complicated.
The relative contribution of surface versus tubular membranes to vesicle formation has not yet been established. However, the presence of L-type Ca2+ channels in the vesicles reported here suggests that the T-system does contribute to vesicle formation. In fact, previous work has shown that, in adult skeletal muscle, the vast majority or all DHP receptors are located in the T-system membranes (Nicola Siri et al. 1980; Fosset et al. 1983; Jorgensen et al. 1989; Flucher et al. 1990). The tubular localization of DHP receptors does not imply that all receptors are located in the junctional region and one report suggested that a significant fraction of DHP receptors can be found in discrete subsarcolemmal foci or caveolae (Jorgensen et al. 1989). Other lines of evidence also suggest that vesicles are formed, at least in part, by the T-system. Thus, ATP-sensitive K+ channels are present in T-system membranes (Parent & Coronado, 1989) and similar channels are also found in vesicles (Spruce et al. 1987). Furthermore, a tubular origin for the vesicles is suggested by experiments in which the tubular membrane of frog muscle fibres was labelled selectively with the potentiometric dye RH-795. It was found that the vesicles form at the level of the Z-line and that they take up the tubular fluorescent probe (J. Vergara, personal communication).
L-type Ca2+ channels
Our results show that the vesicles contain functional L-type Ca2+ channels. We base this conclusion on the fact that we recorded inward Ba2+ currents activated by membrane depolarization and potentiated by Bay K 8644, a well-known DHP agonist of these channels (for a review see Melzer et al. 1995). The time course of Ba2+ currents was clearly dependent on membrane potential, activation was faster at large depolarizations and the currents decayed during the pulse suggesting the involvement of a voltage-dependent inactivation process. In frog muscle fibres, L-type currents also decay during long depolarization. However, it has been difficult to establish whether the currents decay by inactivation (Sanchez & Stefani, 1983) or by depletion of divalent cations from the lumen of the T-system (Almers et al. 1981). Since the vesicles lack invaginations, our results favour the view that a voltage-dependent inactivation process is responsible for the decay of the currents. We found that long depolarizing prepulses inactivated IBa during test pulses. In addition, the mid-point of the steady-state inactivation that we report is similar to the value obtained in intact cells, though the slope is greater (Sanchez & Stefani, 1983). Inactivation is probably responsible for the decay in the amplitude of the fast component of the 'off' currents when the duration of the pulse was increased. In fact, it is expected that at least part of the 'off' current represents the closing or deactivation of Ca2+ channels. Therefore, as inactivation proceeds fewer channels are available when the vesicle is repolarized and the magnitude of the jump in the current is consequently smaller. It is likely, however, that not all of the fast component represents a tail current carried by Ba2 +. This is because, even when the channels were largely inactivated, we found that the amplitude of the fast component was quite significant. Also, even after the concentration of divalent cations was reduced, 'off' currents of significant magnitude were recorded. It is possible that part of the fast component is a non-linear capacitive current that represents the movement of the 'off' charge. On the other hand, the origin of the slow component of the 'off' current remains to be identified. Among other criteria, the presence of two tail current components, with different deactivation kinetics, has been used in the past to identify two distinct types of Ca2+ channel in excitable cells (Matteson & Armstrong, 1986); however, we have no other experimental evidence to support the presence of more than one type of Ca2+ channel in the vesicles.
There are several differences between L-type currents in muscle fibres and those recorded in vesicles. First, the amplitude of Ba2+ currents in the vesicles is smaller than that in cells. Peak values averaged 3 µA µF-1. In contrast, when published data from frog muscle fibres are expressed in the same units (assuming a mean capacitance of 6 µF cm-2), Ca2+ and especially Ba2+ currents have higher values ranging from 10 to 20 µA µF-1 (Almers & Palade, 1981; Sanchez & Stefani, 1983). Several explanations can be put forward to account for the lower current amplitude in the vesicles. First, it is possible that only a fraction of the vesicle membrane is formed by the T-system and, therefore, that the density of DHP receptors would be lower than that in cells. Alternatively, if vesicles derive in part from the surface it is possible that recordings were made from a surface Ca2+ channel with novel properties. Another possible explanation for the small L-type Ca2+ current density in the vesicles is the rundown of Ca2+ currents that has been described in muscle fibres. This process develops with a time course of minutes when fibres are cut (Arreola et al. 1987). It is possible that the complex metabolic machinery that preserves the Ca2+ channel activity in intact cells is altered in the vesicles, leading to a rundown of ionic currents. Finally, another possible explanation could be a missing retrograde signal from the ryanodine receptor to the DHP receptor. It has been shown recently that Ca2+ currents of myotubes prepared from mice homozygous for a disrupted ryanodine receptor gene (dyspedic myotubes) are drastically reduced in size (Nakai et al. 1996). Since the vesicles contain no SR, DHP receptors would lack the retrograde influence of the SR and a reduction in the amplitude of the currents would follow. Our tail current experiments may also be consistent with this possibility. We found only a small increase in the amplitude of the slow 'off' current component when the duration of the pulses was increased. In contrast, it has been shown that long and large depolarizations cause a tail current increase of more than 400 % in mammalian myotubes, an increase that is missing in dyspedic myotubes (Fleig et al. 1996). Nevertheless, it is necessary to point out that, if facilitation is present in frog muscle, other factors may also contribute to its absence in the vesicles. For example, we found that inactivation of L-type currents during large depolarizations is rather pronounced. This would therefore tend to obscure any facilitation.
In addition to differences in current amplitude, the kinetic properties of the Ca2+ channels recorded in the vesicles are clearly different from those previously described in cells. Activation of L-type Ca2+ channels in vesicles is significantly faster. This conclusion was reached when we compared our experimental data with the time-to-peak values obtained using the kinetic model developed by Francini et al. (1996), who described the kinetic properties of L-type Ca2+ channels of frog muscle fibres at 16°C. The continuous line in Fig. 4 shows the relationship, derived from their model, between the time to peak (right ordinate) and membrane potential. The time to peak was about 10 times faster in the vesicles at all membrane potentials. Although the underlying mechanism is still unknown, it is very unlikely that the kinetic differences can be explained by the tubular delay associated with charging of the T-system in muscle fibres. The tubular membrane potential of frog muscle has been measured with potentiometric dyes and it was observed that, when a square voltage pulse is imposed at the surface, it takes the T-system 13 ms to reach 95 % of its final voltage value (Kim & Vergara, 1998). Clearly, this delay makes only a minor contribution to currents lasting seconds, as in the case of L-type currents measured at low temperatures. Another possibility involves changes in channel kinetics produced by the charge carrier used in our experiments. Dirksen & Beam (1995) reported that the rate of activation of L-type Ca2+ channels in mammalian myotubes is accelerated when Ca2+ in the extracellular medium is replaced by a high concentration of Ba2+. However, this effect is far too small to explain our results. For example, using their published kinetic parameters at +50 mV and correcting for temperature differences (Cota et al. 1983), we calculated a time to peak of 110 ms, about 5 times longer than that which we recorded in the vesicles. It is possible that the currents that we recorded are generated by a novel channel that activates with a much faster time course. Alternatively, it is possible that a modulatory influence on channel kinetics, present in the cells, is missing in the vesicles. In this regard, it is interesting to note that though there are no reports on the kinetic properties of Ca2+ channels in dyspedic myotubes, the Ca2+ current records (for example at +10 mV) in the experiments of Nakai et al. (1996) and Fleig et al. (1996) show a distinctly faster activation time course in cells lacking the ryanodine receptor, compared with those recorded in control myotubes.
In our experiments, we found that activation of DHP-sensitive channels was shifted towards more negative potentials by low [Mg2+]. It is possible that a shift in the activation of DHP-sensitive channels may play a role in the potentiation described in cells by Jacquemond & Schneider (1992). The origin of the shift remains to be established, though it seems clear that it cannot be explained by surface charge effects. Thus, decreasing the internal concentration of divalent cations would increase the membrane electric field and would produce shifts in the opposite direction along the voltage axis (Hille, 1992).
Charge movement
We found that vesicles held at negative potentials have non-linear capacitive currents activated during depolarizing steps. These currents show some similarities to the movement of charge 1 recorded in muscle fibres (Chandler et al. 1976). As in cells, the charge associated with the currents saturates when large voltage steps are applied. In addition, nifedipine blocks a substantial amount of charge movement in the vesicles, as it does in muscle fibres (for a review see Rios & Pizarro, 1991). Some differences were also evident. The values of Qmax that we recorded are higher than those reported in cells, which range between 22 and 38 nC µF-1 when Ca2+ is present in the extracellular solution (for a review see Rios & Pizarro, 1991). It is conceivable that the slower depolarization of the T-system when a voltage step is imposed at the surface (Adrian & Peachey, 1973; Heiny & Vergara, 1982) may result in slow current components that have remained undetected. In agreement with this possibility, Kim & Vergara (1998) have recently shown an increase of 37 % in Qmax in their supercharging experiments. The most significant difference between the records of movement of charge 1 in the vesicles and those previously described in muscle fibres is the absence of 'humps' that may be identified with Q. This charge component is closely associated with E-C coupling and may originate from a distinct population of voltage sensors. Alternatively, it may be a consequence of Ca2+ release, an idea proposed by Garcia et al. (1991). Our experiments do not provide direct evidence that distinguishes between these two possibilities, but, if Q is a consequence of Ca2+ release, then the absence of Q in the vesicles, where no Ca2+ release is present, would be readily explained.
In addition to DHP receptors, gating of other voltage-dependent channels may contribute in principle to charge movement in the vesicles. Gating of Na+ channels has a much faster time course, therefore a significant role in charge movement is unlikely. On the other hand, gating of K+ channels in frog skeletal muscle has some similarities with charge movement. However, as reviewed by Huang (1993), the density of K+ channels in muscle is far too low to account for all of the charge movement and since the K+ conductance in vesicles is even lower (Camacho et al. 1996), the contribution of K+ channel gating to charge movement is likely to be less significant in this preparation than in muscle fibres.
In depolarized skeletal muscle fibres, non-linear currents are recorded during hyperpolarizing voltage steps (Adrian et al. 1976). A portion of these currents is generated by a Ca2+-dependent conductance (Brum & Rios, 1987). The remaining currents correspond to non-linear charge movement (charge 2). This charge has been proposed to represent the voltage sensor of E-C coupling in the inactivated state (Brum & Rios, 1987). The voltage dependence of charge 2 has been analysed by the fitting of a Boltzmann function, yielding values for the mid-point of activation that range between -74 and -108 mV and values for the slope of between 19 and 39 mV (Rios & Pizarro, 1991). We also recorded non-linear currents in depolarized vesicles, albeit only during 'on' transients. These currents are similar to those described by Brum & Rios (1987) who proposed that they are carried by Ca2+, since they are eliminated in Ca2+-free solutions. Unfortunately, we could not confirm whether they are indeed Ca2+ dependent, since recordings in the vesicles were unstable in the absence of Ca2+. The nature of the underlying conductance remains to be identified, though its insensitivity to nifedipine suggests that DHP receptors are not involved.
We found no evidence of non-linear charge movement in depolarized vesicles. Indeed, we observed that the relation between the charge under 'off' transients and voltage was linear along the whole voltage range. This may indicate that charge 2 is not present in the vesicles or alternatively that its voltage dependence is extremely flat. Thus, if the movement of charge 2 does not reach saturation within the voltage range that we explored, then integration of transients in depolarized vesicles may appear linear, but may in fact contain some contribution of charge 2. In agreement with this possibility, we consistently observed that the membrane capacitance (measured as in Fig. 6) was larger in depolarized than in polarized vesicles.
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The authors are grateful to Dr F. Francini who provided the rate constants and advice on the use of his kinetic model, to Dr W. Melzer for critically reading the manuscript and to Dr J. Vergara for sharing his unpublished results. We thank A. Aldana for developing analysis programs and Ms S. Zamudio for excellent secretarial work. J. Camacho was supported by a fellowship from CONACyT.
Corresponding author
J. A. Sánchez: Department of Pharmacology, Cinvestav, Apartado Postal 14-740, México D.F. 07300, México.
Email: jsanchez{at}mail.cinvestav.mx
Author's present address
J. Camacho: Max-Planck Institut für experimentelle Medizin, MBNS, Hermann-Rein-Strasse 3, D-37075 Göttingen, Germany.
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