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1Department of Life Sciences, Graduate School of Arts and Sciences, University of Tokyo, Meguro-ku, 153-8902 Tokyo; 2Molecular Neurobiology Section, Neuroscience Research Institute, National Institute of Advanced Industrial Science and Technology, Tsukuba, 305-8566 Ibaraki; 3Division of Biophysics and Neurobiology, Department of Molecular Physiology, National Institute for Physiological Sciences; and 4Section of Developmental Neurophysiology, Center for Integrative Bioscience, Okazaki National Research Institutes, Myodaiji, Okazaki 444-8585 Aichi, Japan
Submitted 16 January 2004; accepted in final form 30 March 2004
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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; Takahashi and Okamura 1998). However, the physiological meaning of diversity in the temporal expression profiles of ion channels is difficult to be addressed in higher vertebrates due to the complexity of cellular organization and embryogenesis.
Ascidians are popular animals for the study of developmental mechanisms because of the detailed information that can be gained on cell-cleavage patterns, precisely mapped cell lineages, and their kinship to the evolutional origin of chordates as well as simple embryogenesis (Satoh 1994). The ascidian genome was recently sequenced (Dehal et al. 2002), and a database regarding the temporal and spatial profiles of gene expression was constructed (Satou et al. 2002). In a short developmental period, isolated cleavage-arrested ascidian blastomeres express sets of proteins corresponding to their cell fates (Takahashi and Okamura 1998). Although a native ascidian larval cell has a distinct size and morphology compared with a cleavage-arrested cell and spatial arrangements of intracellular organelles may be different between larval muscle and cleavage-arrested muscle cells, there are still many benefits for the stable recording of ion channel currents during the early stages of development. Much information has been accumulated on time-dependent changes in ion channel expression during muscle differentiation (Dallman et al. 1998; Davis et al. 1995; Greaves et al. 1996; Hirano et al. 1984; Nakajo et al. 1999). One important outcome of these studies was that the magnitude of current and types of ion channels correlate well with the pattern of action potentials during development. For example, in muscle-lineage cells of Boltenia villosa, a combination of small-amplitude inward-rectifier K+ current and medium-amplitude voltage-dependent Ca2+ channel (VDCC) currents in a short time window during embryogenesis induces spontaneous Ca2+ spikes, which are required for up-regulation of Ca2+-activated K+ (KCa) channels later in development (Dallman et al. 1998). Furthermore, differentiating muscle cells show different VDCC kinetics, and therefore, distinct firing patterns.
In a previous paper, we showed that cleavage-arrested cells from 16-cell stage Halocynthia roretzi embryos differentiate and express Ca2+-induced Ca2+ release (CICR), which involves ryanodine receptors and L-type–like VDCCs (Nakajo et al. 1999). However, in that study, it was not clearly shown how developmental changes in ion channels correlate with the maturation of firing patterns. Ohmori and Sasaki (1977) previously performed intracellular recordings from tail muscle of immature larvae. However, intracellular recordings from tail muscle of mature larvae have not yet been performed because of their tough tunic that resists penetration by a glass microelectrode. In this study, we examined the developmental changes of ion channels and calcium-induced calcium release in relation to the firing properties of cleavage-arrested ascidian muscle cells using current clamp, voltage clamp, and calcium microfluorometry. We found that the fully mature ascidian muscle-type cell exhibits a characteristic oscillatory membrane potential with the frequency of 15 Hz, which is similar to the tail-beating frequency of swimming larvae. Acquisition of this property depends on the developmental expression of a transient outward current, which is activated by CICR.
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METHODS |
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H. roretzi were used in all experiments. Large, posterior-vegetal blastomeres B5.1 from cleavage-arrested 16-cell embryos were isolated and cultured in artificial seawater that contained 0.5 µg/ml cytochalasin B until recording, as previously described (Nakajo et al. 1999). Untreated sister embryos from the same batch were cultured in parallel at the same temperature as the control so that the developmental stage of the cleavage-arrested cells could be determined. The swimming larvae hatched around 48 h after fertilization at 10°C.
Electrophysiology
Two-electrode voltage clamp.
The cells were observed by the two-microelectrode voltage clamp method using Axoclamp-2B (Axon Instruments, Foster City, CA). The electrodes were pulled by a P-97 micropipette puller (Sutter Instrument, Novato, CA) from borosilicate glass capillaries GC150TF-10 (Harvard Apparatus, Kent, UK). The composition of the extracellular solution was as follows (in mM): 430 NaCl, 10 KCl, 10 CaCl2, 70 MgCl2, and 5 PIPES (pH 7.0). The 10 mM CaCl2 was replaced with 5 mM MgCl2 and 5 mM MnCl2 for the Ca2+-free solution used in Fig. 3B, and the 100 mM NaCl was replaced with 100 mM tetraethylammonium (TEA) chloride in Fig. 8Ac. For Fig. 7A, the composition of the extracellular solution was as follows (in mM): 100 CaCl2, 200 mM TEACl, 200 mM tetramethylammonium chloride, 10 mM KCl, and 5 PIPES (pH 7.0). Recording and stimulating microelectrodes were filled with 3 M KCl and 5 mM EGTA. Their resistances were 7–12 and 3–8 M
, respectively. The holding potential was set at –70 mV. The data were filtered at 3.3 kHz and acquired at 10 kHz using pCLAMP 6 (Axon Instruments). It was analyzed using IGOR Pro software (WaveMetrics, Lake Oswego, OR).
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were used for recording and current injection. An electrode with a resistance of 3–8 M was used to set the resting membrane potential to –70 mV by applying a steady current. The data were acquired at 1 kHz using pCLAMP 6 and analyzed with IGOR Pro software.
Single channel recording.
Patch electrodes were fabricated by a P-97 micropipette puller from GC150-10 or GC150T-10 borosilicate glass capillaries (Harvard Apparatus), and the pipettes were pulled in four to six steps. The pipette tips were coated with melted sticky wax and fire-polished. Single-channel recordings were performed by the cell-attached patch clamp technique (Hamill et al. 1981) using an Axopatch-200B amplifier (Axon Instruments) controlled by pCLAMP 8 software (Axon Instruments). Single channel currents were sampled at 10 kHz and filtered at 5 kHz with a 4-pole Bessel filter. The holding potential was set at –70 mV. The currents were analyzed with TAC X 4.1.3 software (Bruxton, Seattle, WA). Capacitive transients were subtracted from the current traces, and single channel current amplitudes were determined from amplitude histograms. The pipette resistance before the establishment of the seal ranged from 0.7 to 1.2 M. The composition of the extracellular solution was as follows (in mM): 130 NaCl, 300 KCl, 75 MgCl2, 5 MnCl2, 10 EGTA, and 5 PIPES (pH 7.0). The composition of the pipette solution was as follows (in mM): 435 NaCl, 80 CaCl2, 5 KCl, and 5 PIPES (pH 7.0). All recordings were performed at 10–12°C.
Microfluorometry
Oregon green 488 BAPTA-1 dextran (100 µM; Molecular Probes, Eugene, OR) was mixed with 0.25 M KCl and loaded into the cytosol through the current electrode by air pressure just before the recording. The Oregon green compound was excited at 450–490 nm, and emission signals were detected at 520 nm using a P100 photometry system (Nikon, Tokyo, Japan). Ca2+ signals were recorded using pCLAMP 6.
Drugs
Caffeine (Sigma Chemical, St. Louis, MO) was dissolved in water to make a 100 mM stock solution. Cytochalasin B was dissolved in DMSO to make a 2 mg/ml stock solution. Verruculogen, penitrem A, paxilline, apamin, charybdotoxin, and iberiotoxin were purchased from Alomone Labs (Jerusalem, Israel) and were separately dissolved in water. Each drug was diluted in the recording solution and applied via the bath perfusion.
Analysis of swimming behavior
The swimming behavior of ascidian larvae was studied in a petri dish as previously described (Okada et al. 2002). The movement of larvae was recorded by a high-speed CCD camera system (FASTCAM-Rabbit-mini; PHOTRON, Tokyo, Japan) at 250 frames/s.
Statistical analyses
The data were expressed as the means ± SE, with n indicating the number of samples. A statistical P value was calculated using the Student's paired or unpaired t-test. P value of 0.05 or less was considered to be statistically significant.
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RESULTS |
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H. roretzi larvae hatch and start swimming 48 h after fertilization (Nakajo et al. 1999). At this stage, larvae show sporadic single twitches in one direction, which corresponds to the "tail flick" previously described for other ascidians (Bone 1989). Larvae rarely show symmetrical swimming; their swimming is clumsy and lasts for only a few seconds. Within 24 h after hatching, the larvae swim quickly and smoothly. The swimming duration also becomes longer, up to several tens of seconds. The tail beating frequencies of swimming tadpole larvae in other ascidian species are between 10 and 20 Hz (Bone 1989; Dallman et al. 2000). Since the beating frequency of H. roretzi larvae has not been characterized, we recorded the swimming motion of larvae with a high-speed video camera at 250 frames/s and compared the tail beating frequency between the two developmental stages (Fig. 1A). The larvae swam with mean frequencies of 10.1 ± 0.4 at 48 h and 14.5 ± 0.3 Hz at 69 h (n = 10 at each developmental stage; P < 0.001 between 48 and 69 h; Fig. 1B). Thus the tail beating frequency became significantly faster after hatching.
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It is notable that the tail-beating frequency of larvae 69 h after fertilization (see Fig. 1) was close to the oscillation frequency of the membrane potential recorded from mature cleavage-arrested muscle cells (Figs. 1B and 2C). This led us to speculate that the intrinsic firing property of muscle cells may correspond to the tail-beating frequency of swimming larvae.
Transient outward K+ currents (Ito) develop in a maturation-dependent fashion in the ascidian muscle cell
Previous studies showed that ascidian muscle cells expressed VDCCs and at least two types of outward K+ channels during development (Davis et al. 1995; Greaves et al. 1996; Hirano et al. 1984; Shidara and Okamura 1991). To define the ionic currents that account for the distinct firing properties between the two developmental stages (just after hatching and 1 day later), we recorded the ionic currents expressed in the ascidian muscle cell in voltage-clamp mode and compared them between the two stages. At 48 h, inward currents and delayed outward currents were observed (Fig. 3A, 48 h). At 72 h, both currents became larger, and transient outward currents (Ito) appeared (Fig. 3A, 72 h). Ito was seen at membrane potentials between –40 and –20 mV in a 10 mM external Ca2+ solution. At the initial stage of the experiment, we assumed that Ito was a fast inactivating A-type K+ current. However, changing the holding potential did not significantly affect the current amplitude of Ito, thereby refuting this possibility (data not shown). We found that not only the inward current but also Ito disappeared when the external bath solution was replaced with Ca2+-free solution (Fig. 3B). This observation suggested that inward current was calcium influx and was required for the activation of Ito.
To examine the ionic basis of Ito, we changed the external K+ concentration and measured the reversal potentials (Fig. 4A). The reversal potential changed from –65 mV in 10 mM to –42 mV in 40 mM K+ solutions (Fig. 4B). According to the Nernst equation, the reversal potential change should be 34 mV when the concentration of extracellular K+ ion is fourfold greater. Our recorded value was smaller than the expected value for the reversal potential change of K+. Similar deviation of reversal potential of K+ channel from Nernst equation was observed in the previous study (Shidara and Okamura 1991). The accumulation of potassium ions just outside of the plasma membrane during depolarization has been proposed as the mechanism for such observation as in other marine invertebrates (Kukita 1988; Shidara and Okamura 1991). Alternatively, disagreement in these values may be due to the contamination of leak current. We concluded that this transient outward current mainly resulted from the outward K+ flux.
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As shown in Fig. 3B, Ito requires ICa. Since Ca2+ influx evokes Ca2+-induced Ca2+ release from internal stores in the ascidian muscle cell (Nakajo et al. 1999), we tested whether activation of Ito depends on CICR by examining the sensitivity of Ito toward caffeine. Caffeine constitutively opens ryanodine receptors. We therefore applied 10 mM caffeine and waited for >5 min before starting the subsequent recording. Under this condition, the intracellular Ca2+ store is depleted, and no calcium release is evoked by depolarization in the ascidian muscle cell as previously observed in the same preparation (Nakajo et al. 1999). Caffeine (10 mM) eliminated Ito (Fig. 6A). The application of caffeine also slightly affected the amplitude and kinetics of the inward current. However, this was probably due to calcium-release dependent inactivation of the VDCC seen in the ascidian muscle cell (Nakajo et al. 1999). Despite the potential inaccuracy of the subtraction strategy due to sensitivity of the calcium current kinetics to CICR, the subtracted traces represent the caffeine-sensitive component of the current, which was transient and outward (Fig. 6A). These results indicate that the activation of Ito requires Ca2+ release from internal Ca2+ stores.
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Ito is synchronized with fluctuations in intracellular calcium
If Ito is dependent on CICR, Ito should be synchronized with local intracellular calcium transiently released from Ca2+ stores. Before confirming this prediction, we first recorded the intracellular calcium concentration of mature muscle cell under voltage-clamp with Oregon green BAPTA-1. To clearly observe calcium fluctuation, we increased the extracellular calcium concentration to 100 mM (see METHODS). Under this condition, we observed two Ca2+ increase phases: fast and slow Ca2+ rises (Fig. 7A, control). When the intracellular Ca2+ store was depleted by 10 mM caffeine, the fast Ca2+ rise disappeared (Fig. 7A, 10 mM caffeine). This suggests that the caffeine-sensitive fast Ca2+ rise is caused by CICR, and the slow Ca2+ rise is due to Ca2+ influx via VDCCs.
We then tried simultaneous recordings of two-electrode voltage-clamp and intracellular calcium fluorometry by Oregon green BAPTA-1 in a normal (10 mM Ca2+) extracellular solution. In representative traces shown in Fig. 7B, it can be seen that Ito is synchronized with the initial fast rise in intracellular calcium concentration, which is probably due to CICR. A second rise seen at –40 and –30 mV may be the subsequent Ca2+ oscillation and may not be located just underneath the plasma membrane.
Pharmacological properties of Ito
In the presence of 100 mM TEA, Ito is abolished (Nakajo et al. 1999). We therefore further examined the effects of other K+ channel blockers. Partial inhibition of Ito was observed with the application of 100 nM verruculogen, penitrem A, and paxilline (Fig. 8, A and B). These compounds are tremorgenic fungal toxins that inhibit BK channels (Knaus et al. 1994). However, other common large conductance Ca2+-activated K+ (BK) channel blockers (100 nM charybdotoxin and 50 nM iberiotoxin) and a small conductance Ca2+-activated K+ (SK) channel blocker (100 nM apamin) were not effective (data not shown). The fungal toxins mildly (
40%) inhibited Ito at –30 mV (Fig. 8, A and B; n = 7, 4, and 2 for verruculogen, penitrem A, and paxilline, respectively). Remaining Ito was completely blocked by 100 mM TEA (Fig. 8Ac). Subtracted traces before and after penitrem A showed its transient kinetics and voltage dependency (Fig. 8A, a – b). Although Ito might be composed of several kinds of ionic currents, the K+ current sensitive to tremorgenic fungal toxins is a significant contributor to Ito.
These inhibitors also affected the membrane oscillation (Fig. 8C). Initial depolarization by current injection often exhibited an overshooting action potential in the presence of verruculogen (Fig. 8C). Verruculogen significantly reduced the oscillation frequency (Fig. 8D, n = 3). However, these effects were not as dramatic as those of caffeine (Fig. 6, C and D).
Single channel activities of the caffeine-sensitive Ca2+-activated K+ channel in the ascidian muscle cell
The correlation between Ito and intracellular calcium dynamics suggests that this putative Ca2+-activated K+ (KCa) current is activated by local CICR. To verify this, we tried to record single channel currents activated by CICR. Under conditions where a VDCC and KCa channel were present in the same patched membrane with ryanodine receptors (calcium release channels) located just underneath the plasma membrane, we expected that an influx in Ca2+ through the VDCC within the patch would induce CICR and thereby activate the nearby KCa channel. The single channel activity of the KCa current was recorded by a cell-attached patch with high Ca2+ (80 mM) present exclusively inside the patch pipette and no Ca2+ in the bathing solution. Ca2+ influx should only occur across the patched membrane such that it induces local Ca2+ release. To set the transmembrane potential near zero, a high K+ concentration (300 mM K+; see METHODS) was used in the bath solution. Under this condition, we identified two types of channels that carried the outward current; one showed a small conductance of 5 pS (unpublished observations), and the other showed a conductance of 60 pS (Fig. 9A). The zero current potential, which was estimated by simple linear extrapolation, was close to the reversal potential of K+ (data not shown). The voltage dependency of the open probability and first opening latency suggested that the 5-pS channel might be a delayed rectifier K+ channel. Additionally, the 5-pS channel did not show any caffeine sensitivity (data not shown). Therefore we decided not to study this channel further, and instead, focused our investigation on the 60-pS channel.
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Four patches contained single K+ channels, evidenced by an absence of overlapping channel activity. We therefore determined the probabilities of channel opening in these patches. The open probability of a single 60-pS K+ channel was plotted against the membrane potential before and after the application of 10 mM caffeine to each patch (Fig. 9B), and probability was found to be variable for each patch. For example, one patch showed its highest open probability at a voltage of +110 mV (Fig. 9Ba), whereas another showed a low open probability except at a voltage of +10 mV (Fig. 9Bc). Such variations could reflect different basal levels of [Ca2+]i underneath the patched membrane. Although detailed profiles of the open probabilities were variable from patch to patch, the open probability, which was diminished by caffeine, was relatively high at voltages around 0 mV for every case. Thus these results suggest that the 60-pS channel is activated by CICR.
We also quantified the first opening latency of this channel in the presence and absence of CICR. The first opening latency of the KCa channel becomes shorter with increasing intracellular calcium concentration (Ikemoto et al. 1989). We thus expected to see a shorter first opening latency for the K+ channel at a membrane potential of 0 mV in the presence of CICR. As expected, the first opening latency was smaller at –10 to +30 mV, where open probability was larger (Fig. 9B). In contrast, the blockade of CICR should prolong the first opening latency. The first opening latency was elongated at membrane potentials around 0 mV when 10 mM caffeine was applied (Fig. 9C). These findings further support the hypothesis that activation of the 60-pS K+ channel is CICR-dependent.
Timing of Ito activation during action potentials
To understand the temporal properties of Ito activation in the oscillatory pattern, we performed membrane potential waveform clamp (Dallman et al. 2000). The oscillatory waveform of the membrane potential was first obtained under current-clamp mode. This waveform was used as the voltage command for the same cell under voltage-clamp mode (Fig. 10Ad). The current obtained through this technique was composed of ionic, capacitive, and leak currents. To isolate Ito, we plotted the subtracted trace (Fig. 10Ac), which is the difference between the current traces before (Fig. 10Aa) and during (Fig. 10Ab) the application of 1 µM verruculogen. A trace of the caffeine-sensitive component was also obtained in a similar manner and superimposed on the verruculogen-sensitive component and the action potential waveform command (Fig. 10B). The amplitude of the caffeine-sensitive component was greater than that of the verruculogen-sensitive component because verruculogen partially block Ito, while caffeine completely inhibits Ito (see Figs. 6 and 8). Despite the difference in amplitudes, the phases of activation and inactivation in the caffeine- and verruculogen-sensitive components were nearly identical. Also, a phase shift existed between the subtracted traces and voltage command waveform, which indicated that the subtracted component was not derived from a linear leakage. Ito activated just prior to the peak of oscillation and subsequently hyperpolarized the membrane potential. Ito peaked maximally 15 ms after its activation. The decay of Ito was slow, and it took about 45 ms to reach the minimum current amplitude (Fig. 10B). The voltage command waveform depolarized the membrane potential before Ito returned to its minimum value.
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Comparison with other systems that exhibit coupling between KCa channels and VDCCs
Ca2+ release-activated K+ currents are found in neurons (Akita and Kuba 2000; Merriam et al. 1999; Mitra and Slaughter 2002a,2002b) and smooth muscle cells (Benham and Bolton 1986; Bolton and Imaizumi 1996). These transient outward currents are composed of random, spontaneous currents, called the spontaneous miniature outward currents (SMOCs) and spontaneous transient outward currents (STOCs). Mitra and Slaughter (2002a) reported that retinal amacrine cells of the aquatic tiger salamander exhibit Ito and STOCs that are sensitive to caffeine and iberiotoxin. A major difference between the amacrine cell and ascidian muscle cell is that STOCs in the amacrine cell are observed at more hyperpolarized membrane potentials (–40 to –60 mV). The activity of the KCa channel at subthreshold levels suppresses STOCs, and as a result, Ito becomes more prominent (Mitra and Slaughter 2002a). The wide-field amacrine cell, one of many amacrine cell types, exhibits intrinsic oscillatory membrane potentials with some features similar to those of the ascidian muscle cell (Solessio et al. 2002; Vigh et al. 2003). The membrane potential oscillation of the wide-field amacrine cell is mediated by a feed-back loop between voltage-gated Ca2+ and KCa currents as in the ascidian muscle cell. However, the frequency in the wide-field amacrine cell is higher. This may be due to the feed-back loop between voltage-gated Ca2+ and KCa currents in the wide-field amacrine cell is a direct coupling rather than indirect coupling via Ca2+ release as occurs in the ascidian muscle cell.
Similar interactions between these currents are also found in hair cells and the synaptic terminals of neurons. In the hair cell of lower vertebrates, interplay between VDCCs and KCa channels determines the resonant frequency of the membrane potential (Art and Fettiplace 1987; Hudspeth and Lewis 1988). In presynaptic terminals, KCa channels that co-localize with VDCCs are activated following microdomain Ca2+ concentration changes near the VDCC pore (Robitaille et al. 1993; Yazejian et al. 2000). In these cases, coupling between VDCC and KCa channels is fast and is not mediated by the CICR. Direct coupling between KCa channels and VDCC is weak in the ascidian muscle cell because Ito is completely suppressed by caffeine. The possibility that the distance between KCa channels and VDCCs is greater in the ascidian muscle cell than in the vertebrate hair cell may explain this phenomenon. Alternatively, the Ca2+ sensitivity of KCa channels may be lower in the ascidian muscle cell.
Functional implications of coupling between KCa channels and CICR
Compared with direct coupling between the KCa channel and VDCC, several physiological advantages of coupling between KCa channels and CICR may exist. First, coupling between KCa channels and CICR is advantageous for activities that require slow signal transduction. In systems that require rapid signal transduction, the mechanical auditory system, for instance, the membrane potential must follow the stimuli of the auditory frequency, which is as large as several hundred Hertz in turtle hair cells (Fettiplace and Fuchs 1999). For such high-speed oscillation, direct coupling between KCa and calcium channels is essential. On the other hand, in a system where CICR mediates the coupling between KCa channel and calcium channels, CICR provides a time delay before the KCa channels open. Deactivation of the KCa channel is also slower because it takes more time to extrude the amplified intracellular Ca2+ signal. These delays suit activities that involve relatively slow signal transduction such as larval locomotion.
The second advantage of coupling between the KCa channel and CICR is that a high local intracellular calcium concentration resulting from CICR will shift the threshold potential of the KCa channel. The threshold of CICR shifts to more negative potentials compared with the I-V relation of VDCCs in the ascidian muscle cell (unpublished observations). This shift activates KCa channels at a more negative membrane potential close to the threshold of VDCC activation and subsequently induces two changes in the action potentials. First, the shift in the threshold decreases the amplitude of overshooting of an action potential. This is consistent with our finding that the action potential overshoots when caffeine eliminates the KCa channel by depleting the Ca2+ store. Second, the low KCa channel activation threshold decreases cell input resistance, and therefore a large depolarizing stimulus is required to cause changes in the membrane potential. A larger depolarizing current is required to fire action potentials in a mature muscle cell that expresses a KCa current compared with an immature muscle cell that does not express one (see Fig. 2). The activity of the KCa channel at threshold of the membrane potential may suppress the spontaneous firing of larval muscle cells and thus restrict muscle contraction to an event of synchronized synaptic transmitter release, which occurs during organized symmetrical swimming. Interestingly, episodes of asymmetrical single twitches, possibly activated by the spontaneous firing of muscle cells, are frequently observed just after hatching, but they are silenced as development proceeds (Bone 1989).
Identity of the CICR-activated K+ current
Based on the following results, we conclude that Ito flowed through CICR-activated K+ channels. First, the removal of Ca2+ from the external solution or bath application of caffeine completely abolished Ito (Figs. 3B and 6A). Second, the kinetics of Ito were similar to those of the intracellular Ca2+ concentration (Fig. 7). Finally, for the cell-attached patch recording, where Ca2+ was only present in the patch pipette, 60-pS conductance K+ channels exhibited a high probability of opening (Po = approximately 0.4) at +10 mV with a low threshold of between –50 and –30 mV (Fig. 9, A and B). K+ channel activity decreased remarkably when the Ca2+ store was depleted by caffeine.
We failed to identify the ion channel species that underlie the CICR-activated K+ current because of its poor sensitivity to various KCa channel blockers. It seems to be related to a family of BK channels, since tremorgenic fungal toxins, which are potent BK channel blockers, mildly inhibited Ito (see Fig. 8). Unlike these toxins, however, the two peptide BK channel blockers iberiotoxin and charybdotoxin failed to suppress Ito. It is possible that Ito is not based on a single population of ion channels but rather on multiple populations. With single channel recording, the putative CICR-activated KCa channel showed a smaller conductance (60.3 pS; Fig. 9A) compared with the conductance of typical vertebrate BK channels. Nevertheless, many KCa channels in the invertebrate nervous system such as molluscan neurons and crayfish muscles are considered to be BK channels based on their voltage dependence and pharmacology despite conductance that ranges around 60–70 pS (Araque and Buño 1999; Crest and Gola 1993; Crest et al. 1992; Gola et al. 1990; Hermann and Erxleben 1987). It is unlikely that the 60-pS conductance channel in our study belongs to a family of intermediate or small conductance KCa channels because a single channel recording showed their voltage dependency. At a higher voltage, the channel showed a shorter opening latency and higher open probability when uncoupled from CICR or VDCCs (see Fig. 9).
Sequenced and coordinated maturation of muscle cell properties
We previously showed that ryanodine receptors and VDCCs appear during the gastrula stage in H. roretzi (Nakajo et al. 1999). The expression of VDCCs occurs during the same stage in B. villosa, another ascidian species (Simoncini et al. 1988). During the gastrula stage, no significant outward K+ channel can be detected, implying that the expression of KCa channels occurs later. Ryanodine receptors and VDCC couple with each other 35 h after fertilization (Nakajo et al. 1999), and the coupling becomes more apparent after 40 h. Our data shows that Ito emerged at between 48 and 72 h, which is later than when the maturation of CICR occurs. Also, the emergence of Ito occurs after the development of an ER-like structure that is formed underneath the plasma membrane (Nakajo et al. 1999). This evidence suggests that KCa channels are recruited in proximity to the ryanodine receptors-VDCC complex after the complex is formed. Such a time sequence of ion channel expression is consistent with a previous report on B. villosa that preceding spontaneous Ca2+ spikes regulate the expression of KCa channels during the maturation of the muscle cell (Dallman et al. 1998). A similar result was obtained regarding the development of mammalian CNS neurons in that the preceding neuronal electrical activities regulate the transcription of a BK channel gene (Muller et al. 1998).
The developmental expression of KCa channels is coordinated with other [Ca2+]i regulatory mechanisms. We have learned that depletion of intracellular Ca2+ stores by caffeine drastically changes the firing pattern (Fig. 6B). Simultaneous recordings of KCa channel activities and [Ca2+]i revealed that the opening of a KCa channel reflects the kinetics of Ca2+ transient evoked by CICR (see Fig. 7B). Thus the intervals between Ca2+ release may determine the interval of KCa channel openings. The rise in [Ca2+]i that is evoked by CICR must be cleared before the next stimulus arrives or else the next Ca2+ release will be weaker. Ca2+ ions are mainly extruded by sarcoplasmic reticulum Ca2+-ATPase pumps in ascidian muscle cells (unpublished observations). In a preliminary experiment, the decay time constant of [Ca2+]i transients that are evoked by a short depolarization was shorter in day 3 cells (48 h after fertilization) than day 2 cells (24 h), which implies that the efficiency of Ca2+ reuptake improves with development.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Address for reprint requests and other correspondence: K. Nakajo, Div. of Biophysics and Neurobiology, Dept. of Molecular Physiology, National Inst. for Physiological Sciences, 38 Nishigonaka, Myodaiji, Okazaki 444-8585 Aichi, Japan (E-mail: knakajo{at}nips.ac.jp).
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REFERENCES |
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78 h old (n = 3). Straight line indicates result of linear extrapolation [y (Hz) =12.5 + 1.0x].
) and 40 mM (
) external K+ solutions are plotted against membrane potential.
,
, cells exhibiting a spiking pattern (n = 9); 
,
, cells exhibiting a oscillatory pattern (n = 13). B: mean current amplitude histograms for Ito and ICa. The same data as in A was used for this plot. Cells exhibiting a spiking pattern (open bars) and oscillatory pattern (filled bars) are compared. ***P < 0.001.
