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Journal of Physiology (2001), 536.2, pp. 397-407
© Copyright 2001 The Physiological Society
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ABSTRACT |
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INTRODUCTION |
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Denervation induces profound changes in the organization of skeletal muscle, reflecting orchestrated alterations in gene expression, including those encoding ion channels. Expression of different ion channels may be increased or decreased, including for example expression of different acetylcholine receptor isoforms as well as redistribution of the receptors, changes in Na+ channel isoforms, decreased expression of Cl- and inward rectifier K+ channels, and increased expression of a small-conductance Ca2+-activated K+ (SK) channel (Hartzell & Fambrough, 1972; Rogart & Regan, 1985; Mishina et al. 1986; Heathcote, 1989; Brenner et al. 1990; Gonoi & Hasegawa, 1991; Lupa & Caldwell, 1994; Pribnow et al. 1999). The altered ion channel expression in denervation results in a decrease in the resting membrane potential and an increase in membrane excitability (Tower, 1939; Albuquerque & Thesleff, 1968; Robbins, 1977).
Skeletal muscle hyperexcitability may result from many different imbalances. In two cases, denervation and the inherited disorder myotonic muscular dystrophy, SK channel expression is implicated in the genesis of the hyperexcitability. Electromyographic recordings have shown that the selective SK channel inhibitor, apamin, suppresses the hyperexcitability in rat denervated skeletal muscle (Vergara et al. 1993) and the myotonic runs in the thenar muscle of patients with myotonic muscular dystrophy (Behrens et al. 1994). These results suggest that SK channels play an important role in the hyperexcitability of skeletal muscle. This finding presents an apparent paradox: how might expression of a hyperpolarizing K+ channel, which normally suppresses excitability, result in hyperexcitability?
SK channels were first recorded from cultured skeletal muscle myotubes and are selectively blocked by the bee venom peptide toxin, apamin (Blatz & Magleby, 1986). Several reports have indicated that there is increased expression of SK channels in denervated muscle and myotonic dystrophy (Barrett et al. 1981; Schmid-Antomarchi et al. 1985; Blatz & Magleby, 1987; Kimura et al. 2000). We have previously shown that mRNA encoding SK3 is induced upon denervation in skeletal muscle (Pribnow et al. 1999). SK3 protein levels, as measured by 125I-apamin binding, were also increased after denervation but with about a 24 h delay compared with the increase in mRNA levels (Pribnow et al. 1999).
SK channels are probably tetrameric assemblies of subunits that share the transmembrane topology of voltage-gated K+ channels, having six membrane-spanning domains with the N- and C-termini residing within the cell (Köhler et al. 1996). However, SK channels are not voltage activated, being gated solely by intracellular Ca2+ ions. Ca2+ gating of SK channels is accomplished through a constitutive association of calmodulin with the C-terminal portion of the channel (Ishii et al. 1997; Xia et al. 1998; Keen et al. 1999). SK channels are activated by submicromolar concentrations of Ca2+ with kinetics that are Ca2+ dependent. The time constant for activation during a jump in cytoplasmic Ca2+ from subnanomolar concentrations to 10 µM is ~5 ms and the deactivation time constant on return to subnanomolar concentrations of Ca2+ is ~50 ms (Hirschberg et al. 1998; Xia et al. 1998).
Excitation-contraction (E-C) coupling in skeletal muscle requires physical coupling between the Ca2+ channel on the T-tubular membrane and ryanodine receptors (RyRs) on the sarcoplasmic reticulum (SR) membrane. Membrane depolarization in skeletal muscle causes a movement of the L-type Ca2+ channel voltage sensor, which in turn activates the RyRs (for reviews see Schneider, 1994; Rios & Stern, 1997). The opening of RyRs results in a rapid (~1 ms) release of Ca2+ from the SR and a transient increase in myoplasmic [Ca2+] to about 10 µM that activates actin/myosin, resulting in filament shortening and muscle contraction. The elevated levels of myoplasmic Ca2+ are buffered by parvalbumin, limiting the duration of the contraction, and Ca2+-ATPases located in the SR subsequently return Ca2+ to the internal stores. Unlike cardiac muscle, neither extracellular Ca2+ nor Ca2+ influx through voltage-gated Ca2+ channels is required (Armstrong et al. 1972). Influx of Ca2+ through L-type Ca2+ channels in skeletal muscle is much slower (time constants, ~30-40 ms) than the release of Ca2+ from the SR or the time delay before muscle contraction (Baylor et al. 1983; Garcia et al. 1992).
In this study we characterized SK currents recorded using the whole-cell patch clamp configuration from single fast-twitch skeletal fibres acutely dissociated from innervated and denervated flexor digitorum brevis (FDB) muscle. Ca2+ released from the SR during short pulses of up to 80 ms primarily activated SK channels. In current clamp mode, the presence of SK channels decreased the action potential threshold by ~8 mV and apamin reversed this decrease. The results are consistent with a model in which SK channel activity in the T-tubules causes a local increase in K+ concentration, resulting in an increase in the T-tubular resting membrane potential that decreases the apparent action potential threshold.
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METHODS |
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Denervation
Animal care and handling were in accordance with Animal Care and Use Committee of OHSU guidelines. C57/Bl-6 mice (8-10 days old) were anaesthetized with a 2.5 % isoflourane-97.5 % oxygen mix. A section of the sciatic nerve (1-2 mm) was excised through a small (3-5 mm) incision over the hip. The incision was sutured with a single stitch and formulated cyanoacrylate (Surgibond, Vetus Animal Health) was applied to the wound. Animals were killed for fibre isolation 4-10 days after denervation by deep halothane anaesthesia and cervical dislocation.
Isolation of FDB fibres
FDB muscle, isolated from the hindfoot, was pinned out in a 35 mm dish coated with Sylgard (Dow Corning, MI, USA). The muscles were incubated in Dulbecco's modified Eagle's medium (DMEM) containing 10 % fetal calf serum, 100 µg ml-1 penicillin-streptomycin and 0.05 % collagenase 1A (Sigma, St Louis, MO, USA) for 1.5-2 h in an incubator at 5 % CO2 and 37 °C. Single muscle fibres were dislodged by gentle tituration in DMEM and were studied within 6 h.
Electrophysiology
An aliquot of the muscle fibres was plated in a recording chamber perfused with Ca2+-free Tyrode solution containing (mM): 140 NaCl, 5 KCl, 2.8 MgCl2, 10 dextrose, 10 Hepes, pH 7.35. The whole-cell recording configuration was established with 2-4 M
electrodes filled with (mM): 145 potassium aspartate, 3.5 NaCl, 1 MgCl2, 6.5 NaOH, 0.05 EGTA, 10 Hepes, 5 MgATP, pH 7.1. Following whole-cell formation, the external bath solution was switched to Tyrode solution containing 1.8 mM Ca2+. The majority of fibres contracted in response to membrane depolarization in the absence of external Ca2+, consistent with skeletal muscle E-C coupling, and only these fibres were studied. All experiments were performed at room temperature.
Fibre length and width were measured using an ocular micrometer with 100 µm divisions. Fibres were observed with a 
40 objective yielding an optical resolution of 2.5 µm between divisions. Half the distance between divisions was easily resolved giving a measured error of ±1.25 µm. Sarcomere spacing, determined from the number of sarcomeres over 10 divisions, was routinely 2-2.5 µm. Only fibres with a sarcomere spacing greater than 2 µm were considered relaxed and used in these experiments.
Whole-cell recordings were performed using an EPC9 (HEKA Elektronik) or Axopatch 200B (Axon Instruments) amplifier and Pulse software (HEKA Elektronik) interfaced to a Macintosh Power PC computer. In voltage clamp mode, series resistance and whole-cell capacitance were electronically compensated; 50-80 % of the series resistance was electronically compensated. Constant current clamp mode was used for action potential measurements. The resting membrane potential was set to -70 mV. Series resistance, measured as an instantaneous jump in membrane potential to an applied current pulse, was nullified by applying series resistance compensation with the percentage compensation set at 100 (Axopatch 200B).
In experiments investigating reversal potential, equimolar KCl was substituted for NaCl. The pipette solution for testing the effects of high internal EGTA comprised (mM): 120 potassium aspartate, 10 NaCl, 1 MgCl2, 0.7 CaCl2, 20 EGTA, 10 Hepes, 5 MgATP, pH adjusted to 7.1 with KOH; the free Ca2+ concentration was ~10 nM.
Data analysis was performed off-line using PulseFit (HEKA Elektronik) and IGOR software (WaveMetrics, Inc., Lake Oswego, OR, USA). Statistical comparisons were performed with either Student's paired t tests or one-way ANOVAs with Tukey HSD post hoc tests to determine which combinations differed. Post hoc tests were performed only on data sets in which the P value of the ANOVA was less than 0.05. Data sets with P < 0.05 were considered significant. All statistical tests were performed with the Instat program (Graphpad) and IGOR. Data are presented as means ± S.E.M. (n, number of fibres).
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RESULTS |
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FDB muscle fibres were chosen for this study because of their relatively small size for skeletal muscle fibres, which allowed for a better space clamp. The mean length and diameter of fibres isolated from innervated muscles were 377.5 ± 17.6 and 18.3 ± 0.9 µm (n = 8), respectively. The effect of denervation on fibre size was minimal (396.5 ± 7.6 and 15.3 ± 0.4 µm, respectively; n = 72), with the small reduction in fibre diameter being significant (P < 0.05). The capacitance measured from an uncompensated transient component of a hyperpolarizing test pulse for innervated and denervated fibres was 483 ± 19 pF (n = 8) and 391 ± 15 pF (n = 72), respectively. The predicted capacitance, assuming a cylindrical cell and a unit capacitance of 1 µF cm-2, was 269 ± 19 pF (n = 8) and 236 ± 8 pF (n = 72), and the ratio of the measured to the predicted capacitance was 1.82 ± 0.20 and 1.65 ± 0.04, for innervated and denervated fibres, respectively. Although the difference in the capacitance ratio between innervated and denervated fibres was not significant, the trend suggests that denervated fibres have a smaller transverse-tubular (T-tubular) density.
Ca2+-dependent tail currents activated in denervated skeletal muscle
Large outward currents were activated during 500 ms step depolarizing pulses to 40 mV in both innervated and denervated fibres (Fig. 1). The amplitude of the outward current varied between cells and the mean current density was 32.4 ± 2.2 pA pF-1 (n = 8) and 26.5 ± 1.4 pA pF-1 (n = 72) for innervated and denervated fibres, respectively. On repolarization to -40 mV, outward tail currents were observed in both innervated and denervated fibres. In innervated fibres, the tail current deactivated rapidly over a 20 ms time period and was best described by the sum of two exponentials with time constants of 2.5 ± 0.2 and 95.0 ± 25.4 ms (n = 8; Fig. 1A). The slow component of the tail current accounted for 1.5 ± 0.3 % of the total tail current, with an amplitude of 0.24 ± 0.01 nA determined by extrapolation back to the beginning of the tail pulse. The maximum outward tail current measured between 50 and 100 ms following repolarization, relative to the final value at 1.5 s, was 0.13 ± 0.01 nA (n = 8). The tail current in denervated fibres contained a large slowly deactivating outward component (Fig. 1C). This slow component often exhibited a small rising phase that peaked ~70 ms after repolarization. The maximum outward tail current measured in denervated fibres between 50 and 100 ms following repolarization varied and in seven out of 72 fibres was within 1 S.D. of the mean value measured in innervated fibres over the same range (Fig. 1B). The mean amplitude of the maximum outward current measured between 50 and 100 ms following repolarization was 0.78 ± 0.05 nA (n = 72) and 0.13 ± 0.01 nA (n = 8) for denervated and innervated fibres, respectively. The decay of the slow component of the tail current after the peak in denervated fibres was described by a single exponential with a mean time constant of 132.8 ± 7.9 ms (n = 72). For comparison, the time constant of the tail currents measured in innervated fibres over the same time range (250-1500 ms) was 1247 ± 686 ms, and the amplitude was 0.04 ± 0.03 nA (n = 8). Presented as a rate constant, the decay of the tail current in denervated fibres was approximately 4-fold faster than that of innervated fibres (Fig. 1D).
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Figure 1. Depolarization induced activation of an outward tail current in denervated FDB muscle fibres A and C, representative current traces evoked by 500 ms depolarization to 40 mV from a holding potential of -50 mV in innervated (A) and denervated (C) fibres. Repolarization to a tail potential of -40 mV activated a rapidly decaying outward current in innervated fibres that was fitted with the sum of two exponentials with time constants of 1.6 and 55.9 ms (A, inset; vertical lines indicate 50-100 ms). The fast component accounted for 98 % of the current. The slow decaying outward current in denervated fibres (measured from 250-1500 ms of the tail pulse) was fitted with a single exponential with a time constant of 124 ms (C, inset; vertical lines indicate 50-100 ms). Bottom panel in C: voltage protocol. B, scattergram of maximum outward tail current measured between 50 and 100 ms following repolarization (vertical lines in inset of A and B) in innervated and denervated fibres. The horizontal bar represents the mean in each category and the dashed line represents the mean plus 1 S.D. for innervated fibres. D, bar graph of rate constant calculated as the reciprocal of the time constant determined from fits of a single exponential to the decay of the outward current over the time range 250-1500 ms. Data are means ± S.E.M. | ||
Activation of the slow component of the tail current in denervated fibres required internal Ca2+ (Fig. 2). Dialysis of denervated fibres with 20 mM EGTA in the patch pipette solution yielded tail currents with kinetics and amplitudes similar to those recorded in innervated fibres (compare Fig. 1A and Fig. 2B). Dialysis of EGTA was rapid and completely suppressed the depolarization-induced contractions within 2 min. The amplitude of the tail current measured upon establishment of the whole-cell recording configuration decreased rapidly and reached a steady state within 2 min, generally within the time course of adjustment of the series resistance compensation. The mean maximum tail current measured between 50 and 100 ms following repolarization after 5 min of dialysis with 20 mM EGTA in the patch pipette solution was 0.10 ± 0.04 nA (n = 6; Fig. 2C). The mean tail current recorded with the standard internal solution on the same day was 0.82 ± 0.18 nA (n = 8).
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Figure 2. Suppression of tail current with intracellular dialysis of EGTA in denervated fibres A, representative current trace from a denervated fibre dialysed with the control minimal EGTA-containing internal solution (0.05 mM). Inset, amplification of outward tail current; vertical lines indicate 50-100 ms. The initial 15 ms of the tail current was deleted. B, representative current trace from another denervated fibre dialysed with 20 mM EGTA in the internal pipette solution. Inset, amplification of outward tail current; vertical lines indicate 50-100 ms. The initial 15 ms of the tail current corresponding to capacitance and delayed rectifier K+ current was deleted. C, bar graph of mean maximum outward tail current measured 50-100 ms following repolarization in denervated fibres dialysed with the control (Control Int, n = 8) and 20 mM EGTA-containing (n = 6) internal solution. Data are means ± S.E.M. | ||
Tail currents are K+ selective and apamin sensitive
The ion selectivity of the tail currents was investigated by increasing the concentration of external K+ from 5 to 50 mM. Currents were activated by a 20 ms depolarizing command to 40 mV from a holding potential of -50 mV, and tail currents were measured over a range of tail potentials from -80 to -10 mV preceded by a gap potential to -50 mV for 20 ms to deactivate the delayed outward rectifier and Ca2+ currents (Fig. 3A). The slopes of deactivation measured from 0.14 to 0.2 s (Fig. 3A, vertical lines) were plotted as a function of the tail potential for each concentration of external K+ and the reversal potential calculated from the voltage intercept of a line fitted to the data points by linear regression. Increasing the external K+ concentration increased the reversal potential (Fig. 3B). Mean reversal potentials changed from -72 ± 2.9 mV in 5 mM K+ to -31.7 ± 2.3 mV in 50 mM K+ (n = 4, Fig. 3C). The mean values were plotted as a function of the K+ concentration and fitted with a logarithmic regression, yielding a slope of 40.7 mV per decade (Fig. 3C). Although these results are consistent with a K+-selective ion channel, the slope was slightly less than that predicted for a purely K+-selective channel (dashed line, Fig. 3C). The measured reversal potential deviated from the predicted value primarily in low (5 mM) K+.
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Figure 3. K+ selectivity of outward tail current in denervated fibres A, family of tail currents measured at tail potentials between -30 and -80 mV (10 mV increments) in 5 mM external K+. From a holding potential of -50 mV the fibre was depolarized to 40 mV for 20 ms, repolarized to -50 mV for 20 ms and subsequently stepped to potentials between -30 and -80 mV (bottom panel). B, the slope of the decaying phase of the tail currents (measured between the vertical lines in A) plotted as a function of voltage for a single fibre bath perfused with either 5 mM ( | ||
Cloned SK channels can be blocked by apamin (Köhler et al. 1996), but not by iberiotoxin, a blocker of the large-conductance Ca2+- and voltage-activated K+ (BK) channel. Application of 100 nM iberiotoxin had no effect on the currents measured in denervated fibres. The amplitude of the tail current in denervated fibres was 1.23 ± 0.05 and 1.21 ± 0.07 nA (n = 4) in control and iberiotoxin-containing bath solution, respectively. In contrast, application of 1 µM apamin nearly completely suppressed the tail current in denervated fibres (Fig. 4). For the representative fibre shown in Fig. 4A, apamin reduced the maximum outward tail current measured between 50 and 100 ms following repolarization from 1.22 to 0.08 nA. Steady-state block was achieved within 15 min of apamin perfusion (Fig. 4A, inset). The block by apamin was reversible but was generally incomplete over the time course of the fibres (30-45 min). The mean reduction in tail current by 1 µM apamin was from 1.38 ± 0.14 to 0.23 ± 0.08 nA (n = 6; P < 0.01; Fig. 4C). Apamin had no effect on currents measured in denervated fibres dialysed with EGTA to eliminate the SK tail current (n = 4; Fig. 4B and C) and the currents measured in innervated fibres were not affected by apamin.
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Figure 4. Apamin sensitivity of outward tail current in denervated fibres A, representative current traces evoked by a 200 ms step from -50 to 40 mV and the resulting tail current measured at -40 mV in control and in 1 µM apamin (ap)-containing solution. Fibre dialysed with control internal solution. Inset, time course of apamin block of tail current measured as the maximum amplitude 50-100 ms following repolarization relative to the final tail current 1.5 s after repolarization. B, representative current traces of a fibre dialysed with an internal solution containing 20 mM EGTA. Inset, time course of tail current. C, bar graph of mean maximum tail current measured between 50 and 200 ms in control bath solution or control bath solution containing 1 µM apamin in fibres dialysed with either control internal solution (Cont Int, n = 6) or EGTA-containing internal solution (EGTA Int, n = 4). Data are means ± S.E.M. | ||
The dependence on internal Ca2+, the K+ selectivity and the apamin sensitivity of tail currents seen only in FDB fibres prepared from denervated skeletal muscle, together with the previous observation of induction of SK3 mRNA in skeletal muscle following denervation (Pribnow et al. 1999), suggests that SK3 channels are responsible for the tail currents.
Time dependence of SK tail currents
Myoplasmic Ca2+ levels can be increased either by Ca2+ released from SR stores or Ca2+ ions entering through voltage-gated Ca2+ channels on the surface membrane. Both of these processes are initiated by voltage but exhibit distinct kinetics. Ca2+ influx through voltage-gated Ca2+ channels is a much slower process, with an activation time constant of ~30 ms at 40 mV, than SR release, which exhibits a delay to half-peak of ~5 ms following T-tubular depolarization (Baylor et al. 1983; Garcia & Schneider, 1993). To examine the source of Ca2+ that activates SK channels in denervated skeletal muscle, the time course of activation of the SK tail current was determined from the envelope of tail currents activated by voltage pulses to 40 mV of varying duration (Fig. 5). A representative family of SK tail currents for pulse durations from 0 to 640 ms in denervated and innervated fibres is shown in Fig. 5A and B, respectively. The maximum outward tail current was measured between 50 and 100 ms following repolarization and the mean maximum tail current from 12 denervated fibres was plotted versus pulse duration (Fig. 5C). The SK tail current in denervated fibres was first detected with pulse durations of > 5 ms and increased rapidly for pulse durations between 5 and 40 ms, then more slowly to a maximum with a pulse duration of 320 ms (Fig. 5C). The envelope of tail currents in innervated fibres revealed small tail currents that reached a steady state within 5 ms (Fig. 5C). The mean tail current measured at a pulse duration of 640 ms in denervated and innervated fibres was 1.25 ± 0.11 nA (n = 12) and 0.07 ± 0.02 nA (n = 8), respectively.
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Figure 5. Time-dependent activation of SK tail currents in denervated skeletal muscle Aa, representative family of traces in a denervated fibre for a test pulse to 40 mV for varying pulse durations from 0 to 640 ms from a holding potential of -50 mV. Ab, superimposed tail currents measured on repolarization to -40 mV. The first 15 ms of the tail currents were deleted. Ba and b, representative families of traces for an innervated fibre. C, mean maximum tail current amplitude measured 50-100 ms after repolarization plotted versus pulse duration for denervated ( | ||
Contributions of SR Ca2+ release and Ca2+ entry to SK tail current activation
The envelope of tail currents may reflect contributions from two sources: Ca2+ released from the SR and Ca2+ entering through voltage-gated Ca2+ channels. Therefore, a pharmacological approach was used to discern their relative contribution. SR Ca2+ release was blocked using ryanodine and Ca2+ entry through voltage-gated Ca2+ channels was blocked by external cobalt.
SR release. Application of ryanodine (10 µM) decreased the SK tail current over a 15 min period. Comparison of the envelope of SK tail currents measured 50 ms after repolarization in control solution (Fig. 6A) and in solution containing ryanodine (Fig. 6B) shows that ryanodine reduced the SK tail current at each pulse duration (Fig. 6E). Subsequent addition of cobalt (2 mM) to block Ca2+ entry through voltage-gated Ca2+ channels eliminated most of the remaining tail current, particularly for pulse durations greater than 80 ms (Fig. 6C and E).
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Figure 6. Ryanodine suppression of SK tail currents in denervated skeletal muscle A, envelope of SK tail currents in control solution determined as in Fig. 5. B, envelope of tail currents after 15 min exposure to 10 µM ryanodine; same fibre as in A. C, envelope of tail currents after 5 min exposure to 2 mM Co2+ plus ryanodine; same fibre as in A and B. D, pulse template. E, amplitude of maximum outward tail current measured 50-100 ms after repolarization plotted versus pulse duration for control ( | ||
Ca2+ entry through voltage-gated Ca2+ channels. Application of cobalt (2 mM) partially reduced the SK tail current (Fig. 7). The envelope of tail currents measured in control solution (Fig. 7A) and 5 min after exposure to cobalt (Fig. 7B) shows that cobalt decreased the SK tail currents for pulse durations greater than 80 ms (Fig. 7C).
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Figure 7. Cobalt suppression of SK tail current following pulse durations greater than 80 ms in denervated skeletal muscle A, envelope of SK tail currents in control solution determined as in Fig. 5. B, envelope of tail currents after 5 min exposure to 2 mM Co2+; same fibre as in A. C, pulse template. D, amplitude of maximum outward tail current measured 50-100 ms after repolarization plotted versus pulse duration for control ( | ||
The envelope of tail currents in each fibre was normalized to the maximum tail current in control solution (usually for pulse durations of 320-640 ms) and plotted as a function of pulse duration (Fig. 8). Plotting the envelope of tail currents on an expanded time axis illustrates the reduction of the SK tail current by cobalt only for pulse durations greater than 80 ms, in contrast to ryanodine which decreased the tail current at each pulse duration, including those less than 80 ms (Fig. 8B). Ryanodine reduced the mean normalized tail current measured at 80 and 320 ms to 0.53 ± 0.15 and 0.44 ± 0.08 of the control, respectively. Cobalt had minimal effect on the tail current at 80 ms (0.88 ± 0.18) but reduced the mean tail current at 320 ms to 0.71 ± 0.12 of the control.
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Figure 8. Ca2+ source for activation of SK tail current A, average envelope of SK tail currents normalized to the maximum amplitude in control solution for each fibre; control ( | ||
These results show that either SR Ca2+ release or Ca2+ entry through voltage-gated Ca2+ channels may gate SK channels in denervated skeletal muscle fibres. However, short pulse durations, representative of a single action potential, activate SK tail currents by SR Ca2+ release.
SK channels reduce the action potential threshold in denervated muscle
The initiation of action potentials in denervated FDB muscle was examined in constant current mode. A holding current was applied to set the resting membrane potential to -70 mV. Constant current depolarizing pulses (1 s duration) of increasing amplitude were applied to depolarize the membrane potential in ~3 mV increments. The steady-state potential measured at the end of the pulse at which the first action potential was initiated was taken as the action potential threshold. Figure 9 shows voltage records from innervated and denervated FDB muscles in response to increasing depolarizing current pulses. The action potential threshold was reached at -50 mV in the innervated fibre in Fig. 9A. However, the threshold was reduced to -53 mV in the denervated fibre and the delay to action potential initiation was increased (Fig. 9B). Application of apamin (1 µM) to the denervated fibre shown in Fig. 9B increased the threshold to -48 mV, without effect on the delay to action potential onset (Fig. 9C and D). The mean action potential threshold in denervated muscle of -55.6 ± 0.5 mV (n = 21) was significantly less than that measured in fibres obtained from innervated FDB muscle, -47.4 ± 1.2 mV (n = 10, P < 0.01, ANOVA; Fig. 9E). Application of apamin (1 µM) increased the action potential threshold in denervated fibres to -48.2 ± 1.8 mV (n = 6; Fig. 9E), similar to that measured in innervated fibres. Suppression of the SK tail current in denervated fibres by dialysis with 20 mM EGTA in the patch pipette solution, to buffer Ca2+ below 1 nM, yielded on average an action potential threshold of -48.4 ± 0.5 mV (n = 6) similar to that of innervated fibres and denervated fibres exposed to apamin (Fig. 9E). These results show that expression of SK channels in denervated skeletal muscle fibres alters the apparent action potential threshold, consistent with an increase in excitability of the denervated fibres.
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Figure 9. SK channel activity decreases the action potential threshold in denervated skeletal muscle A, action potentials recorded in an innervated fibre in current clamp mode. A holding current was applied to set the resting membrane potential at -70 mV. Superimposed traces are shown for a subthreshold and threshold level of current injection that just initiates an action potential. B, voltage records for subthreshold and threshold current injection in a denervated fibre. C, voltage records from the same fibre as in B perfused with 1 µM apamin. D, overlay of action potentials recorded in innervated fibres (long-dash trace), denervated fibres (continuous trace) and denervated fibres perfused with apamin (short-dash trace). E, bar plot of mean threshold for innervated fibres (n = 10), denervated fibres (n = 21), denervated fibres perfused with 1 µM apamin (n = 6) and denervated fibres dialysed with 20 mM EGTA (n = 6). Data are means ± S.E.M. | ||
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DISCUSSION |
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Skeletal muscle fibres are excitable cells that use Ca2+ to couple action potentials to contractions (E-C coupling). SK channels are Ca2+-gated K+ channels that have been implicated in the genesis of hyperexcitable skeletal muscle resulting from denervation or the inherited myotonic disorder, myotonic dystrophy. In both cases, application of apamin markedly reduces the hyperexcitability (Behrens & Vergara, 1992; Behrens et al. 1994). There is also correlative biochemical evidence for increased expression of SK channels because 125I-apamin binding sites are present at much higher levels than in normal innervated muscle. In addition, SK3 mRNA and protein levels are dramatically increased following denervation in skeletal muscle (Pribnow et al. 1999).
To investigate the roles of SK channels in skeletal muscle, we characterized SK currents in denervated FDB fibres. Using whole-cell voltage clamp recordings, we found that the intracellular Ca2+-dependent tail currents activated by a depolarizing pulse in denervated fibres were K+ selective and inhibited by apamin, consistent with the current being the result of functional SK3 channels. Activation of SK tail currents was rapid and the ryanodine-sensitive, cobalt-insensitive component occurred within the first 80 ms of depolarization, clearly demonstrating that the source of Ca2+ for channel activation during the shorter pulses is derived from SR release (Fig. 8). The increase in global free Ca2+ following a single action potential in frog fast-twitch skeletal muscle was at least 10 µM with a half-width of 10 ms (Maylie et al. 1987). Ca2+ activation of SK channels can be described by first-order rate constants with forward and backward rate constants of 47 µM-1 s-1 and 17 s-1, respectively (Hirschberg et al. 1998). For an average calcium transient of 5 µM lasting 10 ms during a single action potential, ~95 % of the SK channels will be activated. Clearly a single action potential was sufficient to activate the majority of the SK channels.
Recordings performed in current clamp mode revealed the importance of SK channel activity for muscle excitability. Current injection initiated an action potential with a time to onset at threshold that was slower in denervated fibres than in innervated fibres. This delay most probably reflects a larger membrane time constant, as a result of the decreased Cl- and inward rectifying K+ conductance that occurs following denervation (Heathcote, 1989; Gonoi & Hasegawa, 1991). In contrast, the threshold for action potential initiation was lower in fibres isolated from denervated muscle than from innervated muscle when stimulated from a similar resting potential. Addition of apamin or intracellular dialysis with EGTA to buffer intracellular Ca2+ to less than 1 nM reversed the decrease in threshold observed in denervated fibres. These results show that SK channels contribute to the hyperexcitability by affecting the apparent action potential threshold in addition to the elevated resting potential that occurs as a result of denervation. Importantly, it also suggests that there is some basal SK channel activity at rest in denervated muscle fibres.
How might the expression of a hyperpolarizing K+ channel paradoxically result in hyperexcitability in denervated muscle? Two observations suggest that SK channels may be localized in the T-tubular network. First, the block by apamin took a relatively long time to develop (~15 min). The T-tubules are long, narrow (20-40 nm) invaginations of the surface membrane that conduct the action potential internally into the muscle fibre to trigger SR Ca2+ release (Franzini-Armstrong et al. 1975; Dulhunty, 1984). The narrowed mouth of the T-tubule may slow the diffusion of apamin into the tubular lumen thereby affecting the time course of block. This is consistent with the finding that the SK currents recorded in cultured myotubes that lack an extensive T-tubular network during the early stages of cell culture were rapidly and completely inhibited by 100 nM apamin (Bond et al. 2000).
Second, the measured reversal potential in 5 mM K+ was less than the reversal potential predicted for a perfectly K+-selective channel. The large outward K+ current during the depolarizing pulse, if equally distributed between the surface and T-tubular membrane, might result in K+ accumulation in the diffusion-limited tubular space. This would cause an underestimation of the reversal potential of the SK tail current, an effect that would be exacerbated at lower external K+ concentrations, such as 5 mM, because the locally accumulated K+ would contribute a larger percentage of the K+ ions. Evidence for K+ accumulation in the T-tubules has been well documented (Almers, 1980). Assuming a narrow T-tubule with a width of 20-40 nm and ignoring K+ reuptake and diffusion, an outward current density of 20 pA pF-1 for 20 ms will result in a K+ accumulation of 4.3-2.2 mM. This will shift the K+ reversal potential from -86 to between -71 and -77 mV, close to the reversal potential measured in 5 mM external K+ (-72.3 mV).
Aberrant expression of SK channels has been implicated in the genesis of the hyperexcitability presented by denervated and myotonic dystrophic skeletal muscle (Behrens & Vergara, 1992; Behrens et al. 1994). In both cases, apamin markedly relieves the hyperexcitability. The unique tubular architecture of skeletal muscle may provide the basis for the apparent paradox that a hyperpolarizing K+ channel may underlie the hyperexcitability in denervated and perhaps myotonic dystrophic skeletal muscle. Localization of SK channels in the T-tubules may mediate an accumulation of tubular K+, as evidenced by the shift in reversal potential seen in 5 mM K+, giving rise to a localized depolarizing condition.
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Acknowledgements
This work was supported by an America Heart Association, Northwest Affiliate Postdoctoral Fellowship Award (T.R.N.), and grants from the Muscular Dystrophy Association, ICAgen Inc., the Human Frontiers of Science Program and the National Institutes of Health (J.P.A. and J.M.).
T. R. Neelands and P. S. Herson contributed equally to this study.
Corresponding author
J. Maylie: Department of Obstetrics and Gynecology, Oregon Health Sciences University, L-458, 3181 S.W. Sam Jackson Park Road, Portland, OR 97201-3098, USA.
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), 15 mM (
) or 50 mM (
) external K+. The reversal potential was determined by interpolation between points on either side of zero for each K+ concentration. C, mean reversal potential (
) and in ryanodine plus Co2+ (