Contribution of L-type Ca2+ channels to evoked transmitter release in cultured Xenopus nerve-muscle synapses

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Contribution of L-type Ca²⁺ channels to evoked transmitter release in cultured Xenopus nerve-muscle synapses

Olav Sand *, Bo-Ming Chen and Alan D. Grinnell

Department of Physiology, Jerry Lewis Neuromuscular Research Center, University of California Los Angeles School of Medicine, Los Angeles, CA 90095, USA and * Department of Biology, University of Oslo, PO Box 1051 Blindern, N-0316 Oslo, Norway

MS 12232 Resubmitted 23 January 2001; accepted after revision 24 May 2001

ABSTRACT

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Abstract
Introduction
Methods
Results
Discussion
References

Simultaneous pre- and postsynaptic patch recordings were obtained from the varicosity synapses formed by Xenopus motoneurons on muscle cells in embryonic cultures, in order to elucidate the contribution of N- and L-type Ca²⁺ channels to the varicosity Ca²⁺ current (I_Ca) and evoked transmitter release.
Although N-type channels are predominant in the varicosities and generally thought to be responsible for all evoked release, in most synapses a fraction of I_Ca and release could be reversibly blocked by the L-type channel antagonist nifedipine, and enhanced by the agonist Bay K8644. Up to 50 % (mean, 21 %) of the I_Ca evoked by a voltage clamp waveform mimicking a normal presynaptic action potential (APWF) is composed of L-type current.
Surprisingly, the nifedipine-sensitive (L) channels activated more rapidly (time-constant, 0.46 ms at +30 mV) than the nifedipine-insensitive (N) channels (time constant, 1.42 ms). Thus the L-type current would play a disproportionate role in the I_Ca linked to a normal action potential.
The relationship between I_Ca and release was the same for nifedipine-sensitive and -resistant components. The N- and L-components of I_Ca are thus equally potent in evoking release. This may represent an immature stage before N-type channels become predominant.
Replacing Ca²⁺ in the medium with Ba²⁺ strongly enhanced the L-type component, suggesting that L-type channels may be inactivated at Ca²⁺ levels close to those at rest.
We speculate that populations of L-type channels in different parts of the neuron may be recruited or inactivated by fluctuations of the cytosolic Ca²⁺ concentration within the physiological range.

INTRODUCTION

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Abstract
Introduction
Methods
Results
Discussion
References

In the large preponderance of nerve terminals studied to date, Ca²⁺ influx through P/Q- or N-type channels triggers neurotransmitter release (Meir et al. 1999). However, in some terminals the release is mediated by more than one Ca²⁺ channel type (Luebke et al. 1993; Regehr & Mintz, 1994; Reuter, 1995; Wheeler et al. 1995; Wu et al. 1999). L-type channels have been shown to be involved in comparatively few nerve terminals, including goldfish retinal bipolar cell synapses (Heidelberger & Matthews, 1992; Mennerick & Matthews, 1998), murine inner hair cells (Platzer et al. 2000), and some peptide-secreting terminals (Rane & Holz, 1987; Loudes et al. 1988).

In mature motor nerve terminals innervating skeletal muscle, evoked release is mediated by Ca²⁺ predominantly through P/Q-type channels in mammals and through N-type channels in lower vertebrates (see Uchitel, 1997). However, in developing neuromuscular junctions L-type Ca²⁺ channels may be involved in different aspects of presynaptic function (Gray et al. 1992; Sugiura & Ko, 1997; Siri & Uchitel, 1999). At the developing frog neuromuscular synapse, the frequency of spontaneous, miniature postsynaptic currents is increased by ATP, elevated [K⁺]_o and phorbol esters (Fu & Huang, 1994). This potentiation is dependent on Ca²⁺ influx through L-type channels, since it is reduced by nifedipine and enhanced by Bay K8644. It was suggested that L-type Ca²⁺ channels, via spontaneous transmitter release, serve an important trophic function in synaptic maturation. However, recordings from the terminals to demonstrate the presence of L-type channels were not performed, and a possible contribution of such channels to evoked transmitter release was not examined.

Co-cultures of embryonic Xenopus laevis spinal neurons and skeletal muscle cells have previously been used to study presynaptic Na⁺ (Kidokoro & Sand, 1989), Ca²⁺ (Hulsizer et al. 1991; Meriney et al. 1991; Yazejian et al. 1997) and Ca²⁺-activated K⁺ currents (Yazejian et al. 1997), as well as the coupling between Ca²⁺ current and transmitter release (Yazejian et al. 1997). Surprisingly, there is still uncertainty regarding the possible existence of functional presynaptic L-channels in this preparation. Meriney et al. (1991) found no effect of Bay K8644 on Ca²⁺ currents in the varicosities, whereas the Ca²⁺ currents in the soma were potentiated. However, Barish (1991) found nifedipine to have no effect on somatic Ca²⁺ currents. Yazejian et al. (1997) reported that the N-channel blocker $omega$ -conotoxin ( $omega$ -CgTX) blocked all transmitter release and almost 85 % of the presynaptic Ca²⁺ current. The possible existence of L-type Ca²⁺ channels in the varicosities was suggested by the observation that the dihydropyridine nimodipine (2 µM) blocked 18 % of the I_Ca. This dihydropyridine-sensitive population of channels was apparently also blocked by $omega$ -CgTX, however, since the addition of nimodipine after application of $omega$ -CgTX caused no further block. The effect of the nimodipine-sensitive I_Ca on release was not examined in that study (Yazejian et al. 1997). That $omega$ -CgTX might block both N- and L-type channels is not unprecedented, having been shown in several other neuronal preparations (McCleskey et al. 1987; Plummer et al. 1989; Wang et al. 1992; Pearson et al. 1995).

We have re-examined the possible existence of L-type Ca²⁺ channels in the varicosities of embryonic Xenopus laevis spinal neurons co-cultured with muscle cells, and have studied the involvement of these channels in evoked release. We report that the varicosities indeed possess L-type Ca²⁺ channels, that these channels are activated much more rapidly than N-type channels in the same preparation, and that Ca²⁺ current through L-type channels contributes significantly to evoked exocytosis of acetylcholine.

METHODS

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Abstract
Introduction
Methods
Results
Discussion
References

Cell culture

The nerve-muscle co-cultures were prepared from stage 20-22 Xenopus laevis embryos (Nieuwkoop & Faber, 1967) as previously described (Tabti & Poo, 1991; Yazejian et al. 1997). The disaggregated cells were plated on uncoated glass coverslips and incubated for 24-48 h at room temperature (22-24 °C) in a medium composed of either 40 % Iscove's modified Dulbecco's medium or L-15 (Sigma), 50 % normal frog Ringer solution (NFR: 116 mM NaCl, 1 mM NaHCO₃, 2 mM KCl, 1.8 mM CaCl₂, 3 mM D-glucose, 10 mM Hepes, pH 7.3) and 10 % deionized H₂O. The medium was supplemented with 7 µg ml^-1 insulin, 0.3 mg ml^-1 glutamine, 5 ng ml^-1 sodium selenite, 4 µg ml^-1 transferrin, 50 units penicillin-streptomycin (all from Gibco) and 25 ng ml^-1 brain-derived neurotrophic factor (BDNF, PeproTech, London, UK). Within 12 h, the spinal neurons start extending neurites that frequently form varicose regions in contact with the substrate. Occasionally the varicosities contact muscle cells, and in 1-2 day cultures the presynaptic regions are often sufficiently large to be accessed by patch electrodes. The synapses formed on the muscle cells exhibit morphological and physiological properties that parallel closely those of their developing counterparts in vivo (Kullberg et al. 1977).

Electrophysiological recordings

Prior to recording, the culture medium was replaced with either glucose-free NFR containing 1 mM 3,4-diaminopyridine (to reduce outward K⁺ currents) and 300 nM TTX (to abolish inward Na⁺ current), or a corresponding solution where the Ca²⁺ was replaced with 1.8 mM Ba²⁺. The volume of the perfusion chamber was 1.2 ml. Ca²⁺ currents (I_Ca) were recorded from the varicosities using the perforated patch technique. The patch electrodes had a resistance of 4-6 M $Omega$ when filled with the following internal solution: 77 mM CH₃O₃SCs, 38 mM CsCl, 1 mM EGTA, 1 mM 3,4-diaminopyridine (DAP), 5 mM Hepes, pH 7.3, plus 900 µg ml^-1 amphotericin B (Sigma) (Rae et al. 1991). The traditional whole cell patch clamp technique was used to record EPSCs in the myocytes. The whole-cell pipettes were filled with the following solution: 116 mM potassium gluconate, 4 mM NaCl, 1 mM MgCl₂, 10 mM EGTA, 10 mM Hepes, pH 7.3. The electrodes were connected to conventional recording equipment, and recordings were not adjusted for the electrode junction potentials. The muscle cells were clamped at -80 mV during the experiments. The holding potential for the varicosities and neurons was -70 mV unless otherwise stated. All experiments were done at room temperature (22-23 °C).

This study is focused on the I_Ca that may be elicited by presynaptic action potentials. The voltage clamp commands to the varicosities therefore usually had a waveform mimicking a normal action potential, which in the varicosities has an amplitude and duration of about 100 mV and 1.5 ms, respectively (Kidokoro & Sand, 1989). The mock action potential waveform (APWF) was composed of a 0.5 ms, 100 mV step pulse followed by a 1 ms ramp back to the -70 mV holding potential. The 0.5 ms plateau phase was inserted in the APWF in order to make the recordings less sensitive to changes in clamp speed during an experiment. In some experiments the duration of the APWF was increased or the amplitude varied, but the duration of the ramp was always 1 ms. Linear leak and capacitive current subtraction were performed digitally using a P/-4 protocol. Preparations were excluded from analysis if Ca²⁺ tail current decay could not be fitted reasonably well with a single exponential. In most cases, the series resistance of the patched varicosities fell in the range 10-20 M $Omega$ and the capacitance was between 5 and 15 pF. Thus the calculated time constant of voltage change was approximately 120-200 µs. This was confirmed empirically by measuring the time course of the capacitance transient associated with large voltage steps, without capacitance compensation or P/-4 subtraction. With capacitance compensation, the clamp would be even faster. Although we did not record uncompensated capacitive transients before and after each experiment, this was routinely done at intervals both in the present series of experiments and in a parallel study of the effects of Ca²⁺ transients on Ca²⁺-activated potassium current (I_K(Ca)) in the same preparation (Yazejian et al. 2000). In all the cases where the decay of the tail current followed the course of a single exponential, the time constant of the capacitive transient was less than 300 µs. Analog low-pass filtering at 3 kHz was employed for the current signals, which were digitized at 100 kHz and stored on a microcomputer. The pCLAMP 6 suite of programs (Axon Instruments, Foster City, CA, USA) was employed for both the acquisition and most of the data analysis. The significance of the observed effects of nifedipine and Bay K8644 was assessed using Student's t test (P < 0.05).

Drugs

Nifedipine and (-)-Bay K8644 were obtained from Sigma and stored as 6-10 mM stock solutions in DMSO. Before experiments the stocks were diluted 100 times with the bath solution and applied directly to the perfusion chamber to yield final concentrations of 0.8-10 µM for nifedipine and 1-5 µM for Bay K8644. The corresponding DMSO concentrations of less than 0.1 % had no observable effect on I_Ca. All handling and applications of the light-sensitive nifedipine were carried out in subdued light. Following drug application and recordings of possible effects, washout of the chamber was achieved by perfusion (2.5 ml min^-1) with at least 6 ml recording solution, corresponding to 5 times the chamber volume. $omega$ -Conotoxin GVIA ( $omega$ -CgTX) was obtained from Bachem (Torrance, CA, USA).

RESULTS

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Abstract
Introduction
Methods
Results
Discussion
References

Nifedipine and Bay K8466 affect the varicosity Ca²⁺ current

To examine varicosity Ca²⁺ currents like those that would be induced by a normal action potential, we applied the APWF as a command potential to voltage clamped varicosities. The Ca²⁺ current was isolated by replacing K⁺ with Cs⁺ in the recording pipette and adding TTX and DAP to the bath solution. The left panels in Fig. 1 present examples of such recordings. The I_Ca peaked during the falling phase of the APWF, as previously observed (Yazejian et al. 1997). The maximal I_Ca elicited by a normal APWF was highly variable, and ranged from hardly detectable to 750 pA. The mean value was 149 ± 127 pA (S.D., n = 60). There was no obvious correlation between the magnitude of the I_Ca and the size of the varicosity. The range and variability in the I_Ca magnitude also seemed to be similar for varicosities that were not in contact with myocytes and for those attached to myocytes.

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Figure 1. L-type Ca²⁺ current in varicosities of cultured embryonic Xenopus spinal neurons
Inward Ca²⁺ current (I_Ca) was evoked by a depolarizing waveform approximating a normal action potential (see voltage protocol at top). Na⁺ current was eliminated by external application of TTX, and K⁺ current was blocked by external application of diaminopyridine and Cs⁺ in the patch pipette. The holding potential was -70 mV. The upper set of recordings shows reversible inhibition of I_Ca by bath application of 5 µM nifedipine. The lower set of recordings is from a different varicosity, and demonstrates reversible enhancement of I_Ca by 1 µM Bay K8644.

The effect of nifedipine (800 nM to 10 µM) was tested on the APWF-induced I_Ca in 50 varicosities. In 70 % of these, nifedipine clearly reduced the I_Ca. This effect was reversible, as shown in the upper set of recordings in Fig. 1. The average reduction of I_Ca caused by nifedipine was 21 ± 20 % (S.D., n = 50). The variability of the response to nifedipine was striking, with a standard deviation of the inhibition close to the mean value. The largest inhibitory effects we observed were about 50 %. The blocking effects of nifedipine were 80-100 % maximal at 800 nM and showed no further block of the remaining I_Ca at concentrations above 4 µM. Given the large variability in response to nifedipine in different varicosities, in most experiments we used 5 µM nifedipine to be sure to block all nifedipine-sensitive channels. The I_Ca of free varicosities and those in contact with myocytes showed similar sensitivity to nifedipine.

The voltage sensitivity of the nifedipine-sensitive component of I_Ca was studied in standard voltage clamp experiments. Figure 2A presents examples of superimposed recordings of inward currents evoked by step depolarizations from a holding potential of -80 mV. The maximum I_Ca was reduced 41 % by 10 µM nifedipine in this particular cell. The corresponding I-V relationships are displayed in Fig. 2B, together with the I-V plot for the nifedipine-sensitive component of I_Ca. We did not observe a systematic shift of the voltage sensitivity of the nifedipine-sensitive current compared with the control recordings.

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Figure 2. Voltage sensitivity of the inward Ca²⁺ currents in a varicosity of a cultured embryonic Xenopus spinal neuron
I_Ca was isolated as described in the legend to Fig. 1. A, superimposed original recordings of I_Ca evoked by step depolarizations from a holding potential of -80 mV to values between -50 and 50 mV (voltage protocol shown at top). Bath application of 10 µM nifedipine caused a 41 % reduction of the maximum inward current (bottom traces). B, I-V relationships for the peak values of the total, the nifedipine-resistant, and the nifedipine-sensitive Ca²⁺ currents.

The DHP agonist (-)-Bay K8644, which prolongs the openings of L-type Ca²⁺ channels, clearly enhanced the I_Ca elicited by the normal APWF in 13 of 15 varicosities. The potentiation was reversible, as shown in the lower set of recordings in Fig. 1. The observed augmentation was highly variable between individual varicosities, to a degree similar to that observed for the nifedipine-dependent inhibition of I_Ca. The average enhancement of I_Ca caused by 1-5 µM Bay K8644 was 39 ± 30 % (S.D., n = 15). The maximal potentiation observed was 80 %. A 40 % enhancement of the total I_Ca would represent an approximately 200 % increase in the fraction of the I_Ca that is dihydropyridine sensitive. This seems a large enhancement, but much larger effects of Bay K8644 have been reported in other preparations (Lory et al. 1993; Chen et al. 1995).

Time dependence of the nifedipine effect

It has been suggested that the presynaptic action potential is normally too brief to recruit a sufficient number of L-channels to influence evoked release (Lindgren & Moore, 1989; Robitaille & Charlton, 1992). We have therefore compared the effect of nifedipine on the I_Ca evoked by APWFs of 1.5-2.5 ms duration. A sample of current traces from one of these experiments is presented in Fig. 3. The current traces obtained under control conditions and during exposure to 5 µM nifedipine are superimposed. Contrary to our expectations, the nifedipine-sensitive current was almost fully developed with the APWF of 1.5 ms, reaching 87 % of its value evoked by the 2.5 ms APWF. On the other hand, the nifedipine-resistant I_Ca evoked by the 1.5 ms APWF reached only about 46 % of its value with the 2.5 ms APWF. Time to peak of the I_Ca was also slightly delayed in the nifedipine-containing solution.

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Figure 3. Nifedipine block of I_Ca generated by action potential waveforms (APWFs) of different durations
Sample recordings showing I_Ca evoked by APWFs of 1.5 ms (A) and 2.5 ms (B) durations. The thick continuous traces show control recordings, while the thin traces are recordings from the same varicosity during exposure to 5 µM nifedipine. The lower set of traces reproduce the I_Ca after nifedipine (thin continuous lines) and compare it with the nifedipine-blocked component (dotted traces) obtained by subtraction of responses before and after nifedipine. The recordings were obtained with baseline-to-peak depolarizations of 100 mV. Note that the nifedipine-sensitive component predominated with the 1.5 ms APWF and was near-maximal in amplitude, while the nifedipine-resistant component more than doubled in amplitude with the increase from 1.5 to 2.5 ms duration.

The time course of activation of I_Ca was examined more systematically before and after treatment with the nifedipine, using an abrupt 100 mV step depolarization (-70 to +30 mV, approximating the level of depolarization in an action potential). Figure 4A presents data from one of these experiments. The continuous superimposed traces were recorded in control solution and during exposure to 5 µM nifedipine, respectively. Nifedipine not only reduced the magnitude of the I_Ca, but markedly slowed its time course of development. The difference between these responses (light dotted trace) represents the nifedipine-sensitive component. It clearly was responsible for most of the early portion of the control I_Ca, and had a much faster time course of excitation than the nifedipine-resistant component. Similar recordings were obtained from eleven additional varicosities (Fig. 4B). For a step from -70 to 30 mV the mean activation time constant, $tau$ , for the whole I_Ca was 0.95 ± 0.05 ms, the nifedipine-resistant I_Ca had a $tau$ of 1.42 ± 0.12 ms, and the $tau$ for the nifedipine-sensitive I_Ca was 0.46 ± 0.08 ms (n = 10).

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Figure 4. Activation kinetics of nifedipine-resistant and -sensitive Ca²⁺ currents
A, superimposed traces showing recordings from a voltage-clamped varicosity stepped from -70 to +30 mV before (Control) and after bath application of 5 µM nifedipine. The nifedipine-resistant I_Ca displayed significantly slower activation kinetics than the total current. Subtraction yielded the nifedipine-sensitive I_Ca (Difference). B, the mean time constants of activation ( $tau$ ) of the different components of I_Ca (n = 12).

This difference in activation properties of the two I_Ca components can explain the striking difference in amplitudes of the I_Ca evoked by the 1.5 and 2.5 ms APWFs in Fig. 3. The 1.5 ms APWF, consisting of a 0.5 ms step and a 1 ms declining ramp, would be above the threshold for Ca²⁺ channel activation for only 0.5-0.8 ms, enough to largely activate the L-type component while only beginning to activate the slower N-type component. With prolongation of the step depolarization to 1.5 ms, however, even the N-type component would be largely activated.

Recruitment of dormant L-type Ca²⁺ channels

In previous tests of the effects of nifedipine on the Ca²⁺ currents in the soma of Xenopus embryonic spinal neurons, L-type Ca²⁺ channels were not found (Barish, 1991). This could be due to the absence of such channels in the soma. However, among the high voltage-activated (HVA) Ca²⁺ channels, the L-type channels are most sensitive to Ca²⁺-dependent inactivation (Imredy & Yue, 1994), and the previously reported recordings from the soma (Barish, 1991) were performed in medium with a relatively high Ca²⁺ concentration (10 mM). It is thus possible that dormant L-type Ca²⁺ channels may still be present in the soma, but inactivated by a relatively high level of [Ca²⁺]_i. We have therefore re-examined the effect of nifedipine on the soma I_Ca in our recording solution, which contained 1.8 mM Ca²⁺. Recordings were obtained from 11 somas, and nifedipine clearly reduced I_Ca in six of these. Nifedipine had no obvious effect on the remaining five. The mean reduction of I_Ca by nifedipine was 14 ± 5 % (n = 11). This effect of nifedipine is not significantly less than the observed effect on the varicosities. To further test the possibility that the L-type channels might be completely or partially inactivated at the ambient level of [Ca²⁺]_i, we reduced [Ca²⁺]_i by using Ca²⁺-free saline as the bath solution. Replacing the culture medium with Ca²⁺-free saline causes [Ca²⁺]_i in cultured embryonic Xenopus spinal neurons to stabilize at about 50 % of the normal value within less than 3 min (Stoop & Poo, 1996). To still be able to record current through the Ca²⁺ channels, the Ca²⁺-free bath solution contained 1.8 mM Ba²⁺. Two motoneuron somas were tested under these conditions, and the I_Ca increased by 62 and 64 % over that obtained in 1.8 mM Ca²⁺. The block by nifedipine increased from 16 and 17 % in NFR to 61 and 41 % in the Ba²⁺-containing solution.

To further investigate the possible effects of Ca²⁺ inactivation on the two components of I_Ca, experiments were performed on motoneuron somas using a standard voltage clamp protocol, with depolarizing step potentials from holding potentials of -80 and -40 mV. These holding potentials were selected to discriminate between inward currents exhibiting different degrees of voltage-dependent inactivation. The L-type Ca²⁺ currents characteristically show no voltage-dependent inactivation, in contrast to the N-type currents (Wang et al. 1992). Figure 5 presents data from a soma that was insensitive to nifedipine in the normal 1.8 mM Ca²⁺-containing recording solution. Figure 5A compares the inward Ca²⁺ currents evoked by depolarizing steps to -10 mV from -40 and -80 mV holding potentials. The lower trace shows the difference between these two current traces, which represents the current component showing voltage-dependent inactivation. The I-V relationship for the peak currents elicited at the two holding potentials and the differential current are presented in Fig. 5B. These I-V plots show a negative shift of about 20 mV compared with similar data previously reported by Barish (1991). This shift can be explained by the difference in surface charge in the recording solutions, which contained 10 mM Ca²⁺ in his experiments and 1.8 mM Ca²⁺ in ours.

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Figure 5. Recruitment of L-type Ca²⁺ current in a soma in Ca²⁺-free solution containing Ba²⁺
A, the upper superimposed traces show I_Ca evoked by depolarization to -10 mV from holding potentials of -40 and -80 mV in a solution containing 1.8 mM Ca²⁺. B, I-V relations for I_Ca at the two holding potentials and the difference between them, representing the component of I_Ca showing voltage-dependent inactivation. C, corresponding sample recordings in Ca²⁺-free solution containing 1.8 mM Ba²⁺. The inward currents at the two holding potentials were greatly enhanced, while the difference remained virtually unaltered compared with the recordings in normal solution. D, I-V relations for the inward Ba²⁺ current at the two holding potentials and the current difference. E and F, bath application of 5 µM nifedipine blocked mainly the component of the inward current that was resistant to voltage-dependent inactivation.

When this cell was exposed to the Ba²⁺-containing, Ca²⁺-free solution, the inward Ba²⁺ currents were greatly enlarged (Fig. 5C and D) compared with the previously recorded Ca²⁺ currents in the same cell. Augmentation of the current component lacking voltage-dependent inactivation was responsible for nearly all of the increase. The differential current, expressing the component that was inactivated at the shallow holding potential, was virtually unaltered, while the current evoked from the -40 mV holding potential was increased by a factor of 2.7.

The effect of nifedipine in the Ca²⁺-free, Ba²⁺-containing solution (Fig. 5E and F) was in dramatic contrast to the lack of nifedipine-dependent inhibition in the control solution. Application of 5 µM nifedipine reduced the recorded currents at both holding potentials to even below their control values. The current elicited by a voltage step to -10 mV from the -40 mV holding potential was 68 % inhibited, while the differential current was only 34 % reduced. It is likely that a modest rundown of the Ba²⁺ currents during this relatively long experiment was partly responsible for the observed reduction of the inward currents. Similar experiments on five somas yielded results comparable to those shown in Fig. 5. There was an increase of the inward currents in Ba²⁺-containing solution averaging 52 ± 10 % (range, 23-84 %) with the -80 mV holding potential, and this enhancement was predominantly in the component that was still present with a -40 mV holding potential, i.e. the non-voltage-inactivating (L-type) component.

L-type Ca²⁺ current contributes to evoked neurotransmitter release

Simultaneous voltage clamp recordings from presynaptic varicosities and postsynaptic muscle cells made it possible to test directly the effect of nifedipine on evoked transmitter release. The holding potentials for varicosities and myocytes were -70 and -80 mV, respectively, and the varicosities received a series of APWF commands of different peak amplitudes. Each series consisted of 10 repetitions of the same command, at 0.33 Hz. The duration of the APWF was kept constant at the normal value of 1.5 ms, while the amplitude was varied between 60 and 160 mV. Each cell pair was exposed to only a limited number of test series, to avoid depletion of transmitter release. The peak inward current increased with increasing APWF amplitude, and we were thus able to correlate I_Ca with the size of the excitatory postsynaptic current (EPSC).

The left column of Fig. 6A shows combined recordings from a presynaptic varicosity and its postsynaptic muscle cell in normal recording solution, and presents the averaged current traces from four test series with APWF amplitudes between 80 and 140 mV. The upper trace in each pair shows the presynaptic I_Ca, while the lower trace displays the EPSC. Larger APWF amplitudes increased both the peak I_Ca and the peak EPSC (filled symbols in Fig. 6B and C). The right column of Fig. 6A presents corresponding recordings after 2-5 min exposure to 5 µM nifedipine, and it is evident that both the I_Ca and the EPSC were reduced at all APWF amplitudes. Figure 6B and C compare the peak values of the presynaptic I_Ca and the EPSC before and after nifedipine application. The inhibitory effect of nifedipine on both the pre- and postsynaptic currents was significant at all the applied APWF amplitudes. The effect of 5-10 µM nifedipine on evoked transmitter release was tested by combined recordings from 11 varicosity-myocyte pairs. Both the presynaptic I_Ca and the EPSC were clearly inhibited in seven of these cases, whereas nifedipine had no obvious effects in the remaining four.

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Figure 6. Contribution of nifedipine-sensitive Ca²⁺ currents to evoked transmitter release
A, simultaneous voltage clamp recordings of I_Ca in a presynaptic varicosity and the evoked EPSC in the postsynaptic muscle cell. I_Ca was induced by APWFs of 1.5 ms duration and 80-140 mV amplitude. The holding potentials of the varicosity and the muscle cell were -70 and -80 mV, respectively. Each trace presents the average of 10 successive recordings. The amplitudes of the APWFs are indicated on the figure. Bath application of 5 µM nifedipine reduced both I_Ca and the EPSC. B, presynaptic I_Ca as a function of APWF amplitude. The data points in B and C show the mean value of the 10 recordings at each APWF amplitude, and the error bars represent the S.E.M. Nifedipine significantly reduced I_Ca at all APWF amplitudes. C, EPSC as a function of APWF amplitude. Nifedipine significantly reduced the EPSC at all APWF amplitudes. D, the EPSC as a function of the presynaptic I_Ca. The filled symbols represent individual values (not averaged) obtained in normal solution, whereas the open symbols display values recorded after nifedipine application. The corresponding regression lines (continuous: control; dashed: nifedipine) are not significantly different.

Our results show that both N- and L-type Ca²⁺ currents are involved in evoked transmitter release in the developing neuromuscular junctions forming in our embryonic cultures. However, these currents may not be equally potent in evoking release, for instance due to different degrees of co-localization with the release sites. This question can be addressed by correlating the EPSCs to the presynaptic I_Ca before and after nifedipine administration. In the latter situation, a given I_Ca will be composed of relatively more N- and less L-type current than a current of the same magnitude before inhibition of the L-type current by nifedipine. If the N-type current is more potent than the L-type in evoking release, a given presynaptic current amplitude would evoke a larger EPSC after nifedipine inhibition. However, this does not seem to be the case. Figure 6D presents scatter plots relating the peak values of postsynaptic EPSCs and presynaptic Ca²⁺ currents. Each current trace shown in Fig. 6A is the average of 10 individual traces, whereas the scatter plots include each individual pair of peak currents. The regression lines linked to the two sets of data points obtained before and after nifedipine application are not significantly different.

We have also performed similar tests of the effect of 1-5 µM Bay K8644 on evoked release by combined recordings from seven varicosity-myocyte pairs. Bay K8644 had no obvious effects in two of these cases, whereas both the presynaptic I_Ca and the EPSC were clearly enhanced in five of the tested junctions. The results from one of these experiments are shown in Fig. 7. The experimental design corresponded to that described above, except that in this case the amplitude of the APWF was varied between 100 and 160 mV. The rather small EPSCs recorded in this experiment illustrate the large variation in transmitter release and postsynaptic receptor density in developing neuromuscular junctions. Bay K8644 (5 µM) significantly increased the peak presynaptic I_Ca at all the tested APWF amplitudes. The EPSC amplitudes were also significantly enhanced at the APWF amplitudes of 120 and 140 mV. However, Bay K8644 had no significant effects on the EPSCs evoked by the 100 mV and 160 mV APWF. We frequently observed saturation of the EPSC at large presynaptic I_Ca, and both nifedipine and Bay K8644 usually had negligible effect on these saturated EPSCs. At the other end of the scale, where small presynaptic currents often caused highly variable EPSCs with frequent failures, the effects of the DHP drugs were also usually less significant.

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Figure 7. Contribution of Bay K8644-sensitive Ca²⁺ currents to evoked transmitter release
A, simultaneous voltage-clamp recordings of I_Ca in a presynaptic varicosity and the evoked EPSC in the postsynaptic muscle cell. I_Ca was induced by APWFs of 1.5 ms duration and 100-160 mV amplitude. The holding potentials of the varicosity and the muscle cell were -70 and -80 mV, respectively. Each trace presents the average of 10 successive recordings. The amplitudes of the APWFs are indicated on the figure. Bath application of 5 µM Bay K8644 augmented both I_Ca and the EPSC. B, presynaptic I_Ca as a function of APWF amplitude. The data points in B and C show the mean value of the 10 recordings at each APWF amplitude, and the error bars present the S.E.M. Bay K8644 significantly enhanced I_Ca at all APWF amplitudes. C, EPSC as a function of APWF amplitude. Bay K8644 significantly increased the EPSC at the 120 and 140 mV APWF amplitudes. D, the EPSC as a function of the presynaptic I_Ca. The filled symbols represent individual values (not averaged) obtained in normal solution, whereas the open symbols display values recorded after Bay K8644 application. The corresponding regression lines (continuous: control; dashed: Bay K8644) are not significantly different.

The scatter plots of the individual pairs of peak pre- and postsynaptic currents are presented in Fig. 7D. Even in this case the regression lines associated with the peak currents recorded before and after drug application have overlapping 95 % confidence intervals, and are thus not significantly different.

DISCUSSION

Top
Abstract
Introduction
Methods
Results
Discussion
References

Nifedipine inhibits and Bay K8644 potentiates the varicosity I_Ca evoked by action potential-like voltage clamp commands in embryonic spinal motoneurons co-cultured with muscle cells. The varicosities thus possess Ca²⁺ channels with L-type properties. However, the nifedipine-sensitive component of the varicosity I_Ca is also blocked by 1 µM of the N-type channel antagonist $omega$ -CgTX GVIA (Yazejian et al. 1997). Rather than being linked to distinct channel entities, the embryonic spinal neurons may express an immature Ca²⁺ channel type with less specific pharmacological sensitivity than mature channels. We believe, however, that the observed L-type properties are linked to channels separate from the previously described N-type channels in this preparation. This conclusion is based on the findings that nifedipine preferentially inhibits the Ca²⁺ channels lacking voltage-dependent inactivation (Fig. 5), and the nifedipine-sensitive component of the I_Ca exhibits distinctly faster activation kinetics than the remaining component (Fig. 3 and Fig. 4). In other preparations, there is precedent for block of L-type channels by $omega$ -CgTX (McClesky et al. 1987; Plummer et al. 1989; Williams et al. 1992; Pearson et al. 1995). In nerve terminals of rat neurohypophysis the specificity of $omega$ -CgTX is highly concentration dependent, and L-type current is inhibited at concentrations exceeding 300 nM (Wang et al. 1992). It is thus not surprising that the presynaptic L-type current component in Xenopus embryonic cultures should be blocked by 1 µM $omega$ -CgTX (Yazejian et al. 1997). In contrast, we consider it unlikely that nifedipine, at the relatively high concentrations used in our experiments, might have blocked a subset of N-type channels, since the N-type channels in the somas of cultured Xenopus neurons are insensitive to 20 µM nifedipine (Barish, 1991). Furthermore, in our experiments, the approximately 80 % of the varicosity current that was resistant to nifedipine at 0.8-1 µM was also insensitive to concentrations of 5-10 µM.

A possible source of error when studying effects of inhibitors on evoked release is depletion of neurotransmitter, which is more likely to occur in embryonic than in mature junctions (Dennis et al. 1981; Sugiura & Ko, 1997). However, our finding that Bay K8644, a specific DHP agonist, enhanced both I_Ca and evoked release is unambiguous evidence that L-type Ca²⁺ current is involved. Neither the L-type-depleted presynaptic currents after nifedipine application nor the L-type-enhanced currents after Bay K8644 treatment differed significantly from control currents in their ability to evoke transmitter release (Fig. 6 and Fig. 7). At the newly formed synapses in our cultures, therefore, the N- and L-type currents are equally potent in evoking release. This finding is in contrast to the generally accepted situation in mature frog neuromuscular junctions, where the release is thought to be dependent on Ca²⁺ influx through N-type channels (see Uchitel, 1997) co-localized with the release sites (Cohen et al. 1991; Robitaille et al. 1993). In view of the lack of specificity of $omega$ -CgTX GVIA, however, it is possible that L-channels also play a role in mature terminals.

During the review of this paper, Thaler et al. (2001) reported a substantial L-type Ca²⁺ current, coupled to release, in the same preparation. These authors found that the L-type channel blockers nitrendipine and nimodipine (1 µM) blocked 28 and 17 % of I_Ca, respectively. The P/Q-channel blocker $omega$ -agatoxin ( $omega$ -Aga IVA) had no effect, while $omega$ -CgTX GVIA blocked 91 % of the current. Thus, consistent with our present findings and those of Yazejian et al. (1997), $omega$ -CgTX GVIA blocks some or all of the L-type channels as well as N-type. $omega$ -CgTX MVIIC (10 µM) proved to be a more specific N-type channel blocker (Thaler et al. 2001).

The presence of several types of Ca²⁺ channels in a nerve terminal may enhance synaptic plasticity, and it is therefore not surprising that multiple types of Ca²⁺ channels exist also in neuromuscular synapses. Atchison (1989) and Pancrazio et al. (1989) reported that although L-type Ca²⁺ channels may be present at adult neuromuscular junctions in rat and mouse, respectively, these channels play no role in transmitter release under physiological conditions. In the neuromuscular junction of adult frog, Arenson & Gill (1996) have shown that nimodipine had no effect on either spontaneous or evoked transmitter release. However, application of the phosphatase inhibitor okadaic acid increased the frequency of spontaneous release, and this enhancement was blocked by nimodipine. Their results suggest that a population of normally silent L-type Ca²⁺ channels in the presynaptic terminals may be recruited by phosphorylation. Unfortunately, the effect of okadaic acid on evoked release was not examined.

Although we cannot rule out the possibility that the presence of L-type channels in embryonic Xenopus motoneurons is an artifact of the culture conditions, it seems more likely that their expression in the cells we have studied is another in a growing list of cases in which there are changes in Ca²⁺ channel expression and function during development. In developing rat neuromuscular junctions, for example, pharmacological manipulations indicate that N-type channels play a significant role in release early in development, but not in mature animals, where P/Q-type channels are predominant (Siri & Uchitel, 1999). L-type channels are prominent and coupled to release in regenerating rat motor terminals (Katz et al. 1996), or terminals recovering from botulinum toxin poisoning (Santafe et al. 2000), but not in normal mature terminals. In chick ciliary ganglion neurons, there is a developmental shift from DHP-sensitive to DHP-insensitive Ca²⁺ channels (Gray et al. 1992), and Iwasaki et al. (2000) have shown that N-type channels contribute to thalamic and cerebellar IPSCs during the post-natal period, but have little effect after day P13. In embryonic rat motor nerve terminals and regenerating junctions in adult rat and frog (Rana pipiens), Sugiura & Ko (1997) reported that L-type channel blockers enhance evoked but not spontaneous release, while no effect was found on mature neuromuscular junctions. Considering these examples of the presence of functional L-channels in developing and regenerating neurons in several species, we find it likely that L-channels do indeed play a prominent role in the developing terminals in Xenopus.

In the Xenopus culture system, moreover, nifedipine-sensitive Ca²⁺ channels are well coupled to evoked release. The suggestion that the activation kinetics of the L-type channels may be too slow to allow a substantial Ca²⁺ influx during the brief action potential (Lindgren & Moore, 1989; Robitaille & Charlton, 1992) is evidently not the case in our embryonic preparation, where the nifedipine-sensitive current displayed much faster activation than the remaining current component (Fig. 3 and Fig. 4). Indeed, because their activation kinetics are faster than that of the nifedipine-insensitive component, the L-type channels can contribute a disproportionate fraction of the I_Ca generated by a brief action potential-like waveform (Fig. 3), and hence play a disproportionate role in triggering release. In an independent series of experiments, we have observed that large conductance K_Ca channels are coupled to both nifedipine-sensitive and -resistant Ca²⁺ currents, and that most of the early Ca²⁺-activated K⁺ current (I_K(Ca)) is blocked by nifedipine in a way that reflects the quantitative difference in kinetics of the L- and N-type currents we have described in the present report (A. D. Grinnell & B.-M. Chen, unpublished data). There is a precedent for L-type channels with comparably fast activation times. In the synaptic terminals of goldfish retinal bipolar neurons, Mennerick & Matthews (1998) described an L-type current with an activation time constant of 0.62 ms at -10 mV. An even faster L-type channel, with an activation time constant of approximately 0.31 ms, has recently been described in murine inner hair cells (Platzer et al. 2000). These are both preparations in which channels with fast kinetics may be necessary for rapid neurotransmission. Together with the channels we describe in Xenopus, these instances suggest that a category of L-channels with fast kinetics may be relatively wide-spread and play an important role in synaptic function.

The relationship between I_Ca and EPSC amplitude for both L- and N-type current components (Fig. 6 and Fig. 7) showed a slope of only about 2, less than the relationship often cited for dependence of release on [Ca²⁺]_o of about 3-4 (Dodge & Rahamimoff, 1967). Direct correlation between I_Ca and release has been achieved in only a few synapses. In the squid giant synapse, the relationship has a maximum slope of about 4 (Augustine & Charlton, 1986), and measurements in the calyform synapses of the medial nucleus of the trapezoid body have yielded values of 4 (Borst & Sakmann, 1996) and 2.7 (Wu et al. 1999). Correlating calculated release rates with [Ca²⁺]_i derived from photorelease from DM-nitrophen yields exponent values of 4.2-4.4 (Schneggenburger & Neher, 2000; Bollmann et al. 2000). In vertebrate preparations, exponents of 3-4 typically are observed only when [Ca²⁺]_o is quite low or I_Ca is small, and decrease at higher levels of Ca²⁺. The relationships we observed in the present experiments are consistent with those observed in the Xenopus nerve-muscle synapses in earlier reports (Yazajian et al. 1997, 2000).

The high sensitivity to Ca²⁺-dependent inactivation may render a large fraction of the L-type channels inactivated at normal levels of [Ca²⁺]_i, as suggested by the enhancement of the inward current in Ca²⁺-free solution containing Ba²⁺ (Fig. 5), a condition in which resting [Ca²⁺]_i is much reduced (Stoop & Poo, 1996). The augmentation was too large to be easily accounted for by increased permeation of Ba²⁺ through individual Ca²⁺ channels (Church & Stanley, 1996), and was restricted primarily to the current component lacking voltage-dependent inactivation. We suggest, therefore, that the enhanced inward current in Ca²⁺-free solution containing Ba²⁺ may be due partially to the unmasking of previously inactivated L-type Ca²⁺ channels. A similar preferential enhancement of the L-type current component by replacing Ca²⁺ with Ba²⁺ in the recording saline has previously been observed in nerve terminals of the rat neurohypophysis (Wang et al. 1992).

Although the evidence in favour of our tentative interpretation of the data presented in Fig. 5 is not conclusive, additional evidence for [Ca²⁺]_i regulation of L-channel recruitment comes from the work of Urbano & Uchitel (1999), who recorded the perineurial Ca²⁺ current outside mouse motor nerve terminals. In normal medium nifedipine had no effect on this current. However, nitrendipine partially blocked the perineurial Ca²⁺ currents when preparations were pre-incubated with BAPTA-AM. These nerve terminals thus possess dormant L-type Ca²⁺ channels that may be unmasked by introducing the Ca²⁺ buffer BAPTA into the cytosol. Evoked transmitter release from the terminals was not affected by nitrendipine, excluding involvement of the unmasked L-type Ca²⁺ channels in this process (Urbano & Uchitel, 1999). L-type Ca²⁺ channels in intact smooth muscle cells are 50 % inactivated at a [Ca²⁺]_i of 260 nM (Schumann et al. 1997). Thus, not only phosphorylation (Arenson & Gill, 1996), but also physiological variations of the level of [Ca²⁺]_i may be a factor in the modulation of L-type Ca²⁺ channel activity.

It is feasible that there exist regional differences of [Ca²⁺]_i within the neuron, and such differences may determine the degree of Ca²⁺-dependent inactivation of the L-channels. For example, growth cones of embryonic Xenopus motoneurons have a significantly lower level of [Ca²⁺]_i than the soma, but the level increases after the initial contact with a muscle cell (Dai & Peng, 1993). Moreover, Ca²⁺ mediates the turning behaviour of growth cones of cultured Xenopus neurons in an extracellular gradient of the diffusable guidance factor, netrin-1 (Hong et al. 2000). The netrin-1-induced turning response depends on Ca²⁺ influx through L-type Ca²⁺ channels as well as Ca²⁺-induced Ca²⁺ release. The direction of growth cone turning depends on the magnitude of regional differences in [Ca²⁺]_i across the growth cone and within the same neuron (Hong et al. 2000; Zheng, 2000). Dynamic changes of [Ca²⁺]_i in developing neurons might thus control several factors, including recruitment and inactivation of L-type Ca²⁺ channels.

In conclusion, embryonic Xenopus spinal motoneurons in co-culture with muscle cells possess L-type Ca²⁺ channels. A normal presynaptic action potential with a duration of about 1.5 ms will activate the L-type channels to a much greater extent than N-type channels in the same cells. The N- and L-components of I_Ca are equally potent in evoking transmitter release. This may represent an immature stage before L-type channels become less closely co-localized with the release sites or disappear altogether. We speculate that populations of L-type channels in different parts of the neuron may be recruited or inactivated by fluctuations of the cytosolic Ca²⁺ concentration within the physiological range. This would be a novel mechanism for modulation of Ca²⁺ currents in nerve terminals.

REFERENCES

Top
Abstract
Introduction
Methods
Results
Discussion
References

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Acknowledgements

We are grateful to Li Qiuhe for preparation of the cell cultures and to Dr X.-P. Sun for helpful suggestions during the course of the work. This work was supported by grants from NSF and NIH.

Corresponding author

A. D. Grinnell: Department of Physiology, Jerry Lewis Neuromuscular Research Center, University of California Los Angeles School of Medicine, Los Angeles, CA 90095, USA.

Email: adg{at}ucla.edu

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ARENSON, M. S. & GILL, D. S. (1996). Differential effects of an L-type Ca²⁺ channel antagonist on activity- and phosphorylation-enhanced release of acetylcholine at the neuromuscular junction of the frog in vitro. European Journal of Neuroscience 8, 437-445	[Medline]
ATCHISON, W. D. (1989). Dihydropyridine-sensitive and -insensitive components of acetylcholine release from rat motor nerve terminals. Journal of Pharmacology and Experimental Therapeutics 251, 672-678	[Abstract]
AUGUSTINE, G. J. & CHARLTON, M. P. (1986). Calcium dependence of presynaptic calcium current and post-synaptic response at the squid giant synapse. Journal of Physiology 381, 619-640	[Abstract]
BARISH, M. E. (1991). Voltage-gated calcium currents in cultured embryonic Xenopus spinal neurones. Journal of Physiology 444, 523-543	[Abstract]
BOLLMANN, J. H., SAKMANN, B. & BORST, J. G. G. (2000). Calcium sensitivity of glutamate release in a calyx-type terminal. Science 289, 953-957
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