Voltage-Gated Ca2+ Conductances in Acutely Isolated Guinea Pig Dorsal Cochlear Nucleus Neurons

Scott C. Molitor1 and Paul B. Manis1,2,3

 1Department of Biomedical Engineering,  2Department of Neuroscience, and  3Department of Otolaryngology---Head and Neck Surgery, The Center for Hearing and Balance, The Johns Hopkins University School of Medicine, Baltimore, Maryland 21205


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
APPENDIX
REFERENCES

Molitor, Scott C. and Paul B. Manis. Voltage-gated Ca2+ conductances in acutely isolated guinea pig dorsal cochlear nucleus neurons. Although it is known that voltage-gated Ca2+ conductances (VGCCs) contribute to the responses of dorsal cochlear nucleus (DCN) neurons, little is known about the properties of VGCCs in the DCN. In this study, the whole cell voltage-clamp technique was used to examine the pharmacology and voltage dependence of VGCCs in unidentified DCN neurons acutely isolated from guinea pig brain stem. The majority of cells responded to depolarization with sustained inward currents that were enhanced when Ca2+ was replaced by Ba2+, were blocked partially by Ni2+ (100 µM), and were blocked almost completely by Cd2+ (50 µM). Experiments using nifedipine (10 µM), omega Aga IVA (100 nM) and omega CTX GVIA (500 nM) demonstrated that a variety of VGCC subtypes contributed to the Ba2+ current in most cells, including the L, N, and P/Q types and antagonist-insensitive R type. Although a large depolarization from rest was required to activate VGCCs in DCN neurons, VGCC activation was rapid at depolarized levels, having time constants <1 ms at 22°C. No fast low-threshold inactivation was observed, and a slow high-threshold inactivation was observed at voltages more positive than -20 mV, indicating that Ba2+ currents were carried by high-voltage activated VGCCs. The VGCC subtypes contributing to the overall Ba2+ current had similar voltage-dependent properties, with the exception of the antagonist-insensitive R-type component, which had a slower activation and a more pronounced inactivation than the other components. These data suggest that a variety of VGCCs is present in DCN neurons, and these conductances generate a rapid Ca2+ influx in response to depolarizing stimuli.


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
APPENDIX
REFERENCES

Postsynaptic Ca2+ plays an important role in many neuronal processes, including the activation of ionic conductances (McManus 1991; Sah 1996) and synaptic plasticity (Linden 1994; Nicoll and Malenka 1995). Voltage-gated Ca2+ conductances (VGCCs) provide a major source of postsynaptic Ca2+ influx in response to synaptic input (Eilers et al. 1995; Jaffe et al. 1994; Miyakawa et al. 1992) and during action potentials (Jaffe et al. 1992; Lev-Ram et al. 1992; Markram et al. 1995). Ca2+ imaging experiments in dorsal cochlear nucleus (DCN) have revealed that action potentials elicit a VGCC-mediated Ca2+ influx in the soma and proximal dendrites of two major populations of DCN neurons (Manis and Molitor 1996; Molitor and Manis 1996). VGCCs also contribute to the electrogenic responses that determine the discharge pattern of neurons during depolarization (Llinás 1988) and may be involved in generating the intrinsic discharge patterns of DCN neurons. A slow depolarization resulting from VGCC activation may be responsible for the complex spiking behavior of cartwheel cells (Agar et al. 1996; Manis et al. 1994; Waller and Godfrey 1994; Zhang and Oertel 1993). In addition, K+ channels activated by a Ca2+ influx through VGCCs may be responsible for the slow afterhyperpolarization observed in pyramidal cells after sustained depolarization (Hirsch and Oertel 1988; Manis 1990). However, no studies have been performed that investigate the biophysical and pharmacological properties of VGCCs that contribute to Ca2+ influx and discharge patterns in DCN neurons.

Mammalian neurons possess a variety of VGCC subtypes that can be distinguished by their voltage dependence and their sensitivity to various pharmacological agents. VGCCs can be classified broadly in terms of the voltage-dependent properties they exhibit: low-voltage activated (LVA), which activate near the resting potential and exhibit a rapid inactivation over this voltage range (Huguenard 1996), and high-voltage activated (HVA), which activate at more depolarized voltages and exhibit a slow inactivation at these depolarized voltages (Bean 1989; Tsien et al. 1988). At the present, at least five HVA subtypes have been found in the CNS: the dihydropyridine-sensitive L type, the omega CTX GVIA-sensitive N type (Fox et al. 1987), the omega Aga IVA and omega CTX MVIIC-sensitive P and Q types (Hillyard et al. 1992; Llinás et al. 1989; Mintz et al. 1992b), and a dihydropyridine- and peptide toxin-insensitive R type, which may be a collection of one or more additional VGCC subtypes (Randall and Tsien 1995; Tottene et al. 1996). The existence of multiple VGCC subtypes within a single neuron could generate differential responses at the synaptic level, having differences in their voltage dependence (Forti et al. 1994; Fox et al. 1987) or in their responses to neurotransmitter modulation (Bean 1989).

The present study addresses two objectives: to survey the VGCC subtypes present in DCN neurons and to estimate the types of stimuli that will generate a Ca2+ influx through VGCCs. To this end, we have investigated the pharmacology and voltage dependence of VGCCs in acutely isolated guinea pig DCN neurons using the whole cell voltage-clamp technique. Our results indicate that various HVA VGCCs are present in DCN neurons, and these conductances are capable of generating a large Ca2+ influx in response to action potentials. The Ca2+ influx and discharge patterns produced by VGCCs may play important roles in the neuronal processing that occurs in the DCN.


    METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
APPENDIX
REFERENCES

Acute cell isolation

Pigmented guinea pigs weighing 150-300 g were anesthetized with pentobarbital (35-40 mg/kg), decapitated, and the brainstem was removed quickly and placed into an oxygenated piperazine-N, N'-bis-(2-ethanesulfonic acid) (PIPES)-buffered dissection solution at 30°C (see composition in Solutions). Using an oscillating tissue slicer, the cochlear nuclei were cut along the strial axis, parallel to the orientation of the parallel fibers, into 400-µm-thick slices. The DCN was isolated from the remainder of the brain stem using Castroviejo scissors. DCN slices were placed in a spinner flask containing oxygenated dissection solution with 0.67 mg/ml of bovine pancreatic trypsin (Sigma, type XI) and 0.5 mg/ml bovine serum albumin (BSA, Sigma, A7638) and were spun slowly (90 rpm) for 30 min. Enzymatically treated slices were thoroughly rinsed in enzyme-free dissection solution with BSA and allowed to incubate for >= 1 h in this solution. Before recording, two or three slices were triturated gently in 0.3-0.5 ml of the dissection/BSA solution with a sequence of three or four fire-polished pipettes having gradually decreasing diameters (2-0.5 mm). Cells were plated on 35-mm culture dishes coated with 10 µg/ml of poly-D-lysine to promote cell adherence (Yavin and Yavin 1974). After waiting 10-15 min for cells to adhere, a continuous flow of a N-(2-hydroxyethyl)piperazine-N'-(2-ethanesulfonic acid) (HEPES)-buffered recording solution (0.5 ml/min) was established to clear away debris and for the exchange of extracellular fluids. Fluid flow was maintained with gravity-fed lines and solution exchange was performed using solenoid valves. Isolated cells were maintained at room temperature (~22°C), and were viable for <= 2-3 h after plating.

Solutions

The dissection solution contained (in mM) 110 NaCl, 5 KCl, 25 glucose, 0.2 CaCl2, 4 MgCl2, and 20 PIPES, pH 7.0 with 5 M NaOH. A Na+/K+ recording solution contained (in mM) 130 NaCl, 5 KCl, 25 glucose, 2.5 CaCl2, 1.3 MgCl2, and 10 HEPES, pH 7.35 with 5 M NaOH. To isolate VGCCs pharmacologically from voltage-gated Na+ and K+ conductances, a Ca2+ recording solution containing (in mM) 120 choline Cl, 20 tetraethylammonium chloride (TEA Cl), 4 4-aminopyridine (4-AP), 25 glucose, 2.5 CaCl2, 1.3 MgCl2, and 10 HEPES, pH 7.35 with 1 M TEA OH, and a Ba2+ recording solution containing (in mM) 115 choline Cl, 20 TEA Cl, 4 4-AP, 25 glucose, 10 BaCl2, 1.3 MgCl2, and 10 HEPES, pH 7.35 with 1 M TEA OH, were used. In some experiments, the choline Cl was replaced with NaCl and 1 µM TTX was added to block Na+ conductances. Currents are referred to as Ca2+ or Ba2+ currents to indicate which recording solution was used. The VGCC antagonists NiCl2 (100 µM), CdCl2 (50 µM), nifedipine (10 µM), omega Aga IVA (100 nM), and omega CTX GVIA (500 nM) were diluted from concentrated stock solutions and added to small aliquots of the Ba2+ recording solution before recording. A Cs+ electrode solution containing (in mM) 130 CsCl, 4 NaCl, 11 EGTA, and 10 HEPES, pH 7.20 with 1 M CsOH and a Tris electrode solution containing (in mM) 90 Tris PO4, 108 Tris base, 20 TEA Cl, 11 EGTA, and 10 sucrose, pH 7.20 with 2 M Tris base were used to provide further block of outward K+ conductances. In addition, 2 mM MgATP, 100 µM leupeptin, 100 µM GTP, 10 mM creatine phosphate, and 100 U/ml creatine phosphokinase were added to the electrode solutions to promote the stability of whole cell recordings and to retard rundown of VGCCs (Horn and Korn 1992). omega Aga IVA was a generous gift of Pfizer, and omega CTX GVIA was obtained from Alomone Labs; the remaining chemicals were obtained from Sigma or Aldrich.

Whole cell recording

Dissociated cells were visualized on an inverted scope (IM-35, Carl Zeiss) with Hoffman modulation contrast optics using ×25, 0.45 N. A. or ×40, 0.65 N. A. objectives. Electrodes were pulled from borosilicate glass capillaries (TW150F-4 glass, World Precision Instruments), fire-polished, and coated with silicone elastomer (Sylgard, Dow Corning) and had a resistance of 3-5 MOmega with either the Cs+ or the Tris electrode solution. Before seal formation, a small amount of enzyme-free electrode solution was sucked into the tip of the pipette to facilitate gigohm seal formation. The whole cell recording configuration (Hamill et al. 1981) was obtained in the Na+/K+ recording solution. Junction potentials for both the Cs+ and Tris electrode solutions were measured to be <2 mV in all recording solutions and were not included in any voltage measures. There was a significant increase in series resistance when using the Tris electrode solution [12.5 ± 1.6 (SE) MOmega in Cs+, n = 18; compared with 24.3 ± 1.2 MOmega in Tris, n = 85]; however, the Tris solution generally produced more stable recordings and provided a better block of K+ currents at depolarized potentials. Recordings in either the Ba2+ or Ca2+ recording solution were obtained if a rapidly inactivating choline- or TTX-sensitive current was observed on depolarization above -45 mV in the Na+/K+ recording solution. Whole cell recordings were obtained using an Axopatch 200 amplifier (Axon Instruments), filtered at 1-50 kHz and digitized at 1-100 kHz with a 12 bit A/D converter (Digidata 1200, Axon Instruments).

Data analysis

Currents elicited over a range of voltages were used to assess the voltage-dependent properties of VGCCs. Averages of four current traces were obtained at a given command step, which were presented sequentially in 5- or 10-mV intervals ranging from -150 to +50 mV. The rate at which command steps were presented was dependent on their duration: 10-ms steps were presented every 250 ms, 100-ms steps were presented every 1 s, and 2- to 4-s steps were presented every 10-15 s. Command steps were presented directly from the holding potential or from a prepulse step; tail currents were elicited by repolarizing steps immediately after the command step. Leakage and residual capacitive currents were eliminated by subtracting scaled versions of the average current generated by a sequence of hyperpolarizing command steps. Current-voltage relationships were constructed by plotting the average or peak current magnitudes against corresponding transmembrane voltage values (see APPENDIX) over specified time windows. Time constants were obtained by performing exponential fits of current traces using a Marquardt-Levenberg minimization algorithm. Exponential fits were limited so that the transmembrane voltage did not deviate >5 mV from the steady-state voltage over the duration of the fit window. Although the same command potentials were used, the transmembrane voltage varied between cells, and a uniform voltage scale was needed to average current magnitudes and time constants across experiments. Therefore the measured currents and time constants were interpolated linearly onto a standardized voltage scale before averaging data across experiments.

VGCC subtype-specific antagonists were used to assess the contribution of VGCC subtypes to the overall current. Currents were elicited with 100-ms steps to -10 mV once every 10 s during exposure to various VGCC antagonists. Despite the inclusion of ATP and an ATP-regenerating system in the pipette solution, a slow rundown of VGCCs was observed that could produce time-dependent errors in estimating the effects of a particular antagonist. To minimize the effects of rundown, an exponential was fit to the time course of the current magnitudes before bath infusion of any antagonists (typically 3-5 min), and the resulting fit was extrapolated over the duration of the experiment (Mintz et al. 1992a). Estimates of antagonist sensitivity were obtained by averaging steady-state current magnitudes over the last minute of antagonist application and normalizing these averages to the extrapolated exponential fit over the same period. This procedure assumes that the rate of rundown was equal among all VGCC subtypes; this is unlikely to be a valid assumption. If one subtype has a faster rundown rate, its contribution would be underestimated and the contribution of the remaining subtypes would be overestimated.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
APPENDIX
REFERENCES

General properties

The vast majority of isolated DCN neurons respond to depolarization with sustained inward currents in the presence of voltage-gated Na+ and K+ channel antagonists. During recording in either the Ca2+ or the Ba2+ solution, depolarizing command steps elicited measurable inward currents in 97 of 103 neurons (Fig. 1A). Peak inward currents averaged -286 ± 35 pA (n = 25) in the Ca2+ recording solution and -550 ± 54 pA (n = 74) in the Ba2+ recording solution. When normalized to membrane capacitance, peak Ca2+ current densities averaged -22.8 ± 3.0 pA/pF and peak Ba2+ current densities averaged -48.7 ± 4.4 pA/pF. Ba2+ currents initially activated between -50 and -40 mV, and the largest inward currents were evoked with command steps between -15 and 0 mV (Fig. 1A). After the activation of inward currents with depolarization, large and rapidly deactivating inward tail currents were observed immediately on repolarization after the command step (arrowhead in Fig. 1A). The magnitude of these tail currents increased monotonically with command step voltage and reached maximal amplitudes with command steps between +20 and +35 mV.



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Fig. 1. Acutely isolated dorsal cochlear nucleus (DCN) neurons respond to depolarization with sustained inward currents in the absence of voltage-gated Na+ and K+ currents. A: averages of 4 current traces elicited by 10-ms command steps ranging from -40 mV (smallest current) to -15 mV (largest current) from a holding potential of -60 mV during recording in the Ba2+ solution. black-triangle-left , large inward tail currents were evoked during repolarization to -55 mV after the command step. Calibration bar: 2 ms, 400 pA. B: calculated transmembrane voltage traces corresponding to the current traces shown in A. ---, transmembrane voltages (Vm); - - -, command potentials (Vc) for comparison. black-triangle-left , transmembrane voltages corresponding to the deactivating tail currents shown in A. Noninteger scale is used for the transmembrane voltages (-39 to -11 mV in 5.6-mV steps) to indicate the magnitude of the difference between command potentials and transmembrane voltages at steady state. Parameters used to calculate the transmembrane voltage for this neuron are: Rs = 11.7 MOmega ; Af = 0.926; tau f = 167.5 µs; tau s = 481.9 µs; Rsc = 10.5 MOmega ; Cmc = 14.1 pF; tau lag = 10 µs; p = 0.80; and c = 0.80. Refer to APPENDIX for more details regarding the calculation of transmembrane voltage. Experiments were performed with Tris electrodes.

The deviation of the transmembrane voltage from the desired command potential complicates the interpretation of voltage-clamp data. During Ba2+ current activation, the transmembrane voltage time course is determined by the flow of current across the uncompensated series resistance. Current flow into the pipette due to VGCC activation (Fig. 1A) produces a voltage drop across the finite series resistance, resulting in transmembrane voltages (Fig. 1B, ---) that are positive relative to the command voltage (Fig. 1B, - - -). Although the difference between the command and transmembrane voltage is usually a few millivolts once steady state is attained (0.5-4 mV in Fig. 1B), the delay to steady state can be >= 1 ms, depending on the membrane properties and the magnitude of the Ba2+ currents. The voltage errors during Ba2+ current deactivation are more pronounced (arrowhead in Fig. 1B; see also Figs. 5B and 6B), partially due to the large magnitude of the tail currents elicited during repolarization. The rapid time course of these tail currents augments these voltage errors: the voltage errors due to current flow across the uncompensated series resistance are superimposed on the errors due to the filtering of the repolarizing step by the uncompensated membrane capacitance. Because the dependence of activating and deactivating Ba2+ current magnitudes on the transmembrane voltage is a critical measure of VGCC function, a method for estimating the transmembrane voltage was developed to more accurately assess the biophysical properties of VGCCs (see APPENDIX).

Divalent ion pharmacology

The divalent ion pharmacology of the Ca2+ and Ba2+ currents recorded from isolated DCN neurons suggests that these inward currents were generated by VGCCs. Peak inward currents recorded with Ca2+ (2.5 mM) as the charge carrier were 73.5 ± 3.5% (n = 6) smaller than the corresponding peak inward currents recorded when Ba2+ (10 mM) was the charge carrier (Fig. 2A). Adding Ni2+ (100 µM) to the Ba2+ solution reduced the peak inward current by 55.6 ± 6.5% (n = 6), whereas Cd2+ (50 µM) reduced the peak Ba2+ current by 98.4 ± 0.4% (n = 4). Although Ca2+ is the primary charge carrier through VGCCs in vivo, Ba2+ was used in the majority of these experiments. The substitution of Ba2+ for Ca2+ increased current magnitudes, allowing for a more accurate characterization of VGCC properties in most cells. In addition, Ba2+ blocked residual outward currents, providing a more complete isolation of currents carried by VGCCs. However, with the exception of the outward currents in the Ca2+ solution at depolarized voltages, the steady-state voltage dependence of the inward currents was not altered when Ba2+ replaced Ca2+ as the primary charge carrier (Fig. 2B). Therefore the remaining data were obtained in the Ba2+ recording solution to assess the pharmacological and voltage-dependent properties of VGCCs.



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Fig. 2. Sustained inward currents recorded from isolated DCN neurons exhibit responses to divalent ions that are typical of voltage-gated Ca2+ conductances (VGCCs). A: average magnitude of normalized Ba2+ currents (100%), Ca2+ currents (26.5 ± 3.5%, n = 7), or Ba2+ currents with either 100 µM Ni2+ (44.4 ± 6.5%, n = 6) or 50 µM Cd2+ (1.6 ± 0.4%, n = 4) added. Current magnitudes were obtained by averaging currents over the last 10 ms of a 100-ms command step to -10 or 0 mV, normalizing to Ba2+ current magnitude obtained in the same cell, and averaging across 4-7 cells. Inset: averages of 4 currents elicited by a 100-ms command step to 0 mV from a holding potential of -70 mV in the presence of the Ca2+ recording solution, the Ba2+ recording solution, or the Ba2+ recording solution with either 100 µM NiCl2 or 50 µM CdCl2 added. B: steady-state voltage dependence of Ca2+ () and Ba2+ (black-triangle) currents. Averages of 4 current traces elicited by 100-ms command steps ranging from -100 to +50 mV from a holding potential of -70 mV were obtained in the Ca2+ and Ba2+ recording solutions. Steady-state current-voltage relationships were calculated by averaging the current and transmembrane voltage over the last 10 ms of the command step, normalizing the measured currents to the magnitude of the largest steady-state Ba2+ current obtained, and averaging across 6 cells. Although the same command steps were used in each experiment, the resulting transmembrane voltage varied between cells. Therefore the normalized current magnitudes were interpolated linearly onto a standardized voltage base before averaging across cells. Experiments were performed using Cs+ electrodes.

VGCC subtypes

The Ba2+ currents recorded from acutely isolated DCN neurons were carried by a variety of VGCC subtypes. The VGCC subtype-specific antagonists nifedipine (10 µM), omega Aga IVA (100 nM), and omega CTX GVIA (500 nM) were used to assay for the presence of known VGCC subtypes. The L-type antagonist nifedipine produced a consistent reduction in Ba2+ current magnitude in all cells tested (Fig. 3, A-D). The P/Q-type antagonist omega Aga IVA blocked a significant portion of VGCCs in some neurons (Fig. 3, A and C) but had little or no effect in others (Fig. 3, B and D). Similarly, the N-type antagonist omega CTX GVIA blocked a substantial fraction of the Ba2+ current in some cells (Fig. 3, B and C) and had little or no effect in others (Fig. 3, A and D). In most cells, a portion of the Ba2+ current was resistant to nifedipine, omega Aga IVA, and omega CTX GVIA but could be blocked by 50 µM Cd2+ (Fig. 3, A-D). Thus the Ba2+ currents in most isolated DCN neurons were carried by three or more known HVA VGCC subtypes.



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Fig. 3. Ba2+ currents recorded from acutely isolated DCN neurons are carried by various VGCC subtypes. Currents were elicited by a 100-ms command step to -10 mV from a holding potential of -70 mV once every 10 s; data points represent the average current over the last 50 ms of the command step. Bars indicate addition of one or more of the following VGCC subtype-specific antagonists to the Ba2+ recording solution: 100 nM omega Aga IVA, 500 nM omega CTX GVIA, 10 µM nifedipine, or 50 µM CdCl2. - - -, exponential fits of the control period extrapolated over the experiment duration to provide an estimate of rundown. Small letters indicate where averages of 6 current traces were obtained to calculate difference currents shown in each inset. A-D: current magnitude time course from four different cells during exposure to VGCC antagonists. Insets: difference between averaged current traces obtained before and during exposure to an antagonist to estimate the portion of the overall Ba2+ current blocked by that antagonist. Experiments were performed using Tris electrodes.

With the exception of the component insensitive to nifedipine, omega Aga IVA, and omega CTX GVIA, VGCC subtypes contributing to the overall Ba2+ current appeared to possess similar biophysical properties. Current traces were averaged before and during exposure to a particular antagonist and were subtracted to estimate the portion of the Ba2+ current blocked by that antagonist (Fig. 3, A-D, insets). Although the rundown of VGCCs could confound these results, similarities were observed between difference traces from cells in which rundown was not prominent (Figs. 3, A and C, insets). Measures of activation rates and inactivation extent were obtained from difference currents to quantify the biophysical properties of the individual VGCC subtypes. The average activation time constants were 1.4 ± 0.2 ms for the omega Aga IVA-sensitive component (n = 10), 1.6 ± 0.2 ms for the omega CTX GVIA-sensitive component (n = 10), and 1.5 ± 0.2 ms for the nifedipine-sensitive component (n = 10). The antagonist-insensitive component had an average activation time constant of 2.7 ± 0.4 ms (n = 10) and was significantly slower than the other three components (unpaired Student's t-test, P < 0.02 for all comparisons). The magnitude of inactivation was 6.8 ± 2.0% for the omega Aga IVA-sensitive component (n = 11), 6.0 ± 1.5% for the omega CTX GVIA-sensitive component (n = 12), and 0.1 ± 0.1% for the nifedipine-sensitive component (n = 13). The antagonist-insensitive component exhibited 16.5 ± 2.9% inactivation (n = 13), which was significantly larger than the other three components (unpaired Student's t-test, P < 0.02 for all comparisons). The nifedipine-sensitive component also had significantly less inactivation than the omega Aga IVA- and omega CTX GVIA-sensitive components (unpaired Student's t-test, P < 0.001 for both comparisons).

The order in which antagonists were applied was varied to minimize the effects of rundown in determining the relative contributions of VGCC subtypes to the overall Ba2+ current. Two different antagonist application sequences were developed under the assumption that the effects of nifedipine, but not those of omega Aga IVA or omega CTX GVIA, were reversible over the time course of these experiments. The first sequence consisted of applying nifedipine after the peptide toxins omega Aga IVA and omega CTX GVIA had been applied (Fig. 3A); the second sequence consisted of applying the peptide toxins during a continuous application of nifedipine (Fig. 3, B-D). No statistically significant differences could be detected in the relative proportions of VGCC subtypes observed in cells in which nifedipine was presented before and during presentation of the peptide toxins (Fig. 4A) when compared with cells in which nifedipine was presented after omega Aga IVA and omega CTX GVIA had been applied (Fig. 4B). Across all experiments, nifedipine blocked 32.5 ± 3.6% (n = 13) of the Ba2+ current, omega Aga IVA blocked 15.5 ± 4.2% (n = 11), and omega CTX GVIA blocked 23.2 ± 6.3% (n = 12). Some of the Ba2+ current was insensitive to nifedipine, omega Aga IVA, and omega CTX GVIA but still could be blocked by Cd2+ (23.3 ± 4.1%, n = 11), whereas a small fraction of the overall Ba2+ current was not blocked by any antagonists, including Cd2+ (5.5 ± 1.0%, n = 12).



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Fig. 4. Summary of Ba2+ current responses to the VGCC subtype-specific antagonists omega Aga IVA (100 nM), omega CTX GVIA (500 nM), and nifedipine (10 µM). Relative contribution of each VGCC subtype to the overall Ba2+ current was estimated by the reduction in Ba2+ current during the bath infusion of a particular antagonist. Relative contribution of VGCC subtypes resistant to the 3 subtype-specific antagonists used in these experiments was estimated by the reduction of Ba2+ current during exposure to CdCl2 (50 µM), after the other antagonists had been applied. Pharmacological presentations were designed under the assumption that the effects of nifedipine, but not those of omega Aga IVA or omega CTX GVIA, were reversible over the time course of these experiments. Ba2+ current magnitudes (data points in Fig. 3, A-D) were averaged over the last minute of antagonist application and normalized to the extrapolated control period fit (Fig. 3, A-D, - - -) to account for the rundown of VGCCs over the duration of these experiments. A: relative contributions of VGCC subtypes sensitive to nifedipine (37.3 ± 3.4%, n = 8), omega Aga IVA (11.7 ± 4.0%, n = 6) and omega CTX GVIA (18.5 ± 6.0%, n = 7) from experiments in which nifedipine was presented before and during applications of omega Aga IVA and omega CTX GVIA (e.g., Fig. 3, B-D). Most of the Ba2+ current resistant to the subtype-specific antagonists was blocked by Cd2+ (25.6 ± 3.1%, n = 6), whereas a small fraction of the overall Ba2+ current was resistant to all applied antagonists (6.0 ± 1.0%, n = 7). B: relative contributions of VGCC subtypes sensitive to omega Aga IVA (20.1 ± 7.9%, n = 5), omega CTX GVIA (29.7 ± 12.8%, n = 5) and nifedipine (24.9 ± 6.8%, n = 5) from experiments in which nifedipine was presented after application of omega Aga IVA and omega CTX GVIA (e.g., Fig. 3A). Again most of the Ba2+ current resistant to the subtype-specific antagonists was blocked by Cd2+ (20.5 ± 8.7%, n = 5), and a small fraction of the overall Ba2+ current was resistant to all applied antagonists (4.8 ± 1.9%, n = 5). Experiments were performed using Tris electrodes.

Voltage dependence of Ba2+ currents

Open-channel Ba2+ current-voltage relationships were constructed by measuring the magnitudes of tail currents elicited on return to various repolarizing voltage steps after the activation of VGCCs with a command step to +50 mV. Under the conditions of these experiments, there are two problems with the interpretation of these data. First, the transmembrane voltage (Fig. 5B, ---) does not follow the repolarizing step potential (Fig. 5B, ···) over the duration of the tail current. Large differences between the repolarizing step potential and the estimated transmembrane voltage could be observed at the tail current peak (estimated transmembrane voltages at the tail current peak are indicated in Fig. 5B, - - -). To minimize errors due to the difference between repolarizing and transmembrane voltages, tail current magnitudes (Itail in Fig. 5A) were plotted as a function of the estimated transmembrane voltage at the tail current peak (Vtail in Fig. 5B). Second, tail current peaks are delayed relative to the onset of the repolarizing step (peak times indicated in Fig. 5A, - - -). A significant number of channels could close during the delay to the tail current peak, resulting in an underestimation of the open-channel current (Taylor 1988). Unfortunately, accurate exponential fits of tail current decay could not be obtained to correct for the underestimation of open-channel current magnitudes. The decay of larger tail currents did not follow an exponential time course, presumably due to a changing transmembrane voltage throughout the duration of the tail current (Fig. 5B). Despite the likely underestimation of open-channel currents, large inward open-channel Ba2+ currents were observed at hyperpolarized voltages (Fig. 5C). Open-channel Ba2+ current magnitudes decreased monotonically with voltage; an inward rectification at depolarized voltages prevented the reversal to an outward current.



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Fig. 5. Voltage dependence of open-channel Ba2+ currents. A: averages of 4 Ba2+ tail currents elicited by repolarizing steps ranging from -110 mV (largest tail current) to -20 mV (smallest tail current) after a 5-ms command step to +50 mV. Residual capacitive transients at the offset of the command step have been blanked for clarity. - - - , times of tail current peaks; the magnitudes of these peaks (Itail) were used to construct the open-channel Ba2+ current-voltage relationship shown in C. Calibration bar: 0.3 ms, 2 nA. B: transmembrane voltage traces corresponding to the current traces shown in A. ---, transmembrane voltages; ···, command potentials for comparison; - - -, indicates times of the tail current peaks shown in A; transmembrane voltage at the tail current peaks (Vtail) were used to construct the open-channel Ba2+ current-voltage relationship shown in C. Parameters used to calculate the transmembrane voltage for this neuron are: Rs = 13.0 MOmega ; Af = 0.924; tau f = 200.2 µs; tau s = 665.9 µs; Rsc = 12.5 MOmega ; Cmc = 11.2 pF; tau lag = 10 µs; p = 0.70; and c = 0.70. C: voltage dependence of open-channel Ba2+ currents. Averages of 4 tail current traces were elicited by repolarizing potentials ranging from -150 to +45 mV after the activation of VGCCs with a 5-ms command step to +50 mV. Open-channel current-voltage relationships were constructed by plotting the magnitudes of tail currents elicited during the repolarizing step (Itail in A) as a function of the transmembrane voltage at the tail current peak (Vtail in B). Open-channel currents were normalized to their magnitude at 0 mV and averaged across 6 cells. Although the same repolarizing voltage steps were used in each experiment, the transmembrane voltage at tail current peaks varied between cells. Therefore tail current magnitudes were interpolated linearly onto a standardized voltage base before averaging across cells. Experiments were performed with Tris electrodes.

Similar to the open-channel Ba2+ current, the voltage dependence of steady-state VGCC open probability could be estimated by measuring the voltage dependence of tail current magnitudes. Tail currents were elicited at a repolarizing potential of -80 or -55 mV after the activation of VGCCs with command steps ranging from -100 to +50 mV. Steady-state open probability then was estimated by plotting tail current magnitudes (Itail in Fig. 6A) as a function of the steady-state transmembrane voltage (Vss in Fig. 6B). However, the magnitudes of these tail currents are also a function of the transmembrane voltage at each tail current peak (Vtail in Fig. 6B). Therefore the transmembrane voltage at the tail current peak Vtail must be held constant to obtain an accurate estimate of the dependence of the open probability on the steady-state voltage Vss. Because of variations in the transmembrane voltage at each tail current peak (Fig. 6B), an alternative method for estimating the steady-state open probability was required.



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Fig. 6. Voltage dependence of steady-state VGCC open probability. A: averages of 4 Ba2+ current traces elicited by 10-ms command steps ranging from -35 to +10 mV from a holding potential of -60 mV; tail currents were subsequently elicited by a repolarizing step to -80 mV. Residual capacitive transients at the offset of the command step have been blanked for clarity. - - -, times of tail current peaks; magnitudes of these peaks (Itail) were used to construct the steady-state open probability-voltage relationship shown in C. Also shown is the time window over which current magnitudes were averaged (Iss) for the steady-state current-voltage relationship shown in D. Calibration bar: 0.4 ms, 750 pA. B: transmembrane voltage traces corresponding to the current traces shown in A. ---, transmembrane voltages; ··· command potentials for comparison; - - -, times of the tail current peaks shown in A; transmembrane voltage at the tail current peaks (Vtail) were used to calculate the steady-state open probability shown in C. Also shown is the time window over which the transmembrane voltage was averaged (Vss) for the steady-state open probability shown in C and the steady-state current-voltage relationship shown in D. Parameters used to calculate the transmembrane voltage for this neuron are: Rs = 26.4 MOmega ; Af = 0.930; tau f = 167.7 µs; tau s = 645.7 µs; Rsc = 25.0 MOmega ; Cmc = 5.5 pF; tau lag = 20 µs; p = 0.70; and c = 0.70. C: voltage dependence of steady-state VGCC open probability. Averages of 4 tail current traces were elicited by a repolarizing potential of -80 or -55 mV after the activation of VGCCs with 10-ms command steps ranging from -100 to +50 mV. Steady-state open probability was calculated as a ratio of tail current magnitudes (Itail in Figs. 5A and 6A) to account for the variation in the transmembrane voltage at which tail currents peaks were measured (Vtail in B); see text for more details. For each cell, the steady-state open probability Po(Vss) was fit to a modified Boltzmann function
<IT>P</IT><SUB><IT>o</IT></SUB>(<IT>V</IT><SUB><IT>ss</IT></SUB>)<IT>=</IT><IT>A</IT><SUB><IT>min</IT></SUB><IT>+</IT><FR><NU><IT>A</IT><SUB><IT>max</IT></SUB><IT>−</IT><IT>A</IT><SUB><IT>min</IT></SUB></NU><DE><IT>1+exp</IT>[−(<IT>V</IT><SUB><IT>ss</IT></SUB><IT>−</IT><IT>V</IT><SUB><IT>1/2</IT></SUB>)<IT>/</IT><IT>k</IT>]</DE></FR>
where Amin is the minimum, Amax is the maximum, V1/2 is the half-activation voltage, and k is the activation slope. The steady-state open probability subsequently was normalized so that Amin = 0 and Amax = 1, and averaged across 6 cells. ---, best-fit of a modified Boltzmann function with Amin = 0 and Amax = 1 to the steady-state open probability from 6 cells, resulting in a half-activation voltage V1/2 = -12.6 ± 0.3 mV, and an activation slope k = 8.34 ± 0.22 mV. D: voltage dependence of steady-state Ba2+ current magnitudes. Averages of 4 current traces were elicited by 10-ms command steps ranging from -100 to +50 mV from a holding potential of -60 mV. Steady-state current-voltage relationships were constructed by averaging the current (Iss in A) and transmembrane voltage (Vss in B) during the last 1 ms of the command step, normalizing the measured currents to the magnitude of the largest steady-state current obtained, and averaging across 5 cells. ---, voltage dependence of steady-state Ba2+ current magnitudes reconstructed as a product of open-channel Ba2+ current Iopen (Fig. 5C) and steady-state open probability Popen (Fig. 6C), normalized to the magnitude of the largest inward current, and averaged across 5 cells. - - -, mean ± SE of reconstructed steady-state Ba2+ current magnitudes. Because of variations in the steady-state transmembrane voltage between cells, the data in C and D were linearly interpolated onto a standardized voltage base before averaging across cells. Experiments were performed using Tris electrodes.

To correct for variations in the transmembrane voltages at each tail current peak, the steady-state VGCC open probability was estimated as a ratio of tail currents elicited at the same transmembrane voltage. Assuming that the VGCC open probability is maximal at +50 mV, the steady-state open probability Po(Vss) was calculated by comparing the magnitude of a tail current elicited after a step to Vss to the magnitude of a tail elicited after a step to +50 mV
<IT>P</IT><SUB><IT>o</IT></SUB>(<IT>V</IT><SUB><IT>ss</IT></SUB>)<IT>=</IT><FR><NU><IT>I</IT><SUB><IT>tail</IT></SUB>(<IT>V</IT><SUB><IT>ss</IT></SUB><IT>, </IT><IT>V</IT><SUB><IT>tail</IT></SUB>)</NU><DE><IT>I</IT><SUB><IT>tail</IT></SUB>(+<IT>50 mV, </IT><IT>V</IT><SUB><IT>tail</IT></SUB>)</DE></FR>
Itail(Vss, Vtail) is the magnitude of the tail current (Itail in Fig. 6A) elicited by the transition from the steady-state transmembrane voltage (Vss in Fig. 6B) to the transmembrane voltage measured at the tail current peak (Vtail in Fig. 6B). Itail(+50 mV, Vtail), obtained from the open-channel Ba2+ current data in Fig. 5, is the magnitude of a tail current (Itail in Fig. 5A) evoked by the transition from +50 mV to the same transmembrane voltage at the tail current peak (Vtail in Fig. 5B). The resulting steady-state open probability-voltage relationship (Fig. 6C) shows that a large depolarization was required to activate VGCCs: <5% of maximal activation was achieved at -40 mV. However, VGCCs were steeply voltage dependent once activated: 80% of the increase in open probability occurs within a 40-mV range around the half-activation voltage of -13 mV.

The voltage dependence of steady-state Ba2+ current magnitudes was described accurately by a product of the open-channel Ba2+ current and the steady-state open probability. Steady-state current-voltage relationships were constructed by plotting the average current during the last 1 ms of a 10-ms command step (Iss in Fig. 6A) against the average transmembrane voltage over the same period (Vss in Fig. 6B). Measurable inward currents initially were observed at transmembrane voltages above -50 mV, and the largest inward currents were obtained around -8 mV (Fig. 6D, ). Outward currents were sometimes observed around or above +50 mV; however, the lack of intracellular Ba2+ requires that any outward currents be carried by other ions present in the intracellular solution. Steady-state Ba2+ current magnitudes were reconstructed as a product of the steady-state open probability (Fig. 6C) and open-channel Ba2+ current (Fig. 5C) to confirm the accuracy of these estimates (Fig. 6D, ---). The reconstructed current magnitudes compare favorably to the measured values over a wide voltage range. Differences between the measured and reconstructed values at hyperpolarized voltages can be attributed to slightly positive steady-state open probability values and large inward open-channel current magnitudes over this voltage range.

Ba2+ currents activated rapidly over the range of voltages at which these currents could be elicited. The time course of VGCC activation in response to a voltage step will contain multiple components that can be described by a sum of exponentials. However, an imperfect voltage clamp can prevent the accurate measurement of these components: if there is any uncompensated series resistance, changes in current magnitude will produce a corresponding change in transmembrane voltage. Because the time course of VGCC activation is voltage dependent, changes in transmembrane voltage subsequently will alter the current time course. Rather than extending over the entire duration of the 10-ms command step, exponential fits were limited so that the transmembrane voltage at the beginning of the fit did not deviate >5 mV from steady-state values (Fig. 7A, thick lines). This procedure precludes the use of higher-order kinetic models: only the slowest components of VGCC activation could be measured; faster components would have decayed before the beginning of the fit window. The time constants from these exponential fits had a bell-shaped voltage dependence, with the slowest time constants occurring around -20 mV, and the fastest values occurring at the most depolarized voltages (Fig. 7B). It is not possible to correct for the filtering of membrane currents by any remaining uncompensated membrane time constant (Sigworth 1983), which can distort the time course of faster currents (Armstrong and Gilly 1992). It is likely that the time course of VGCC activation is affected by this filtering and would occur more rapidly under an ideal voltage clamp.



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Fig. 7. Voltage dependence of Ba2+ current activation time course. A: exponential fits of activating currents. Thin traces, averages of 4 current traces elicited by 10-ms command steps ranging from -45 mV (smallest current) to -20 mV (largest current) from a holding potential of -60 mV. Thick traces, corresponding exponential fits. Curve fits were limited so that the transmembrane voltage at the beginning of the fit window did not deviate >5 mV from the average transmembrane voltage during the last 1 ms of the command step. Parameters used in the voltage estimation procedure for this neuron are: Rs = 18.2 MOmega ; Af = 0.92; tau f = 274.8 µs; tau s = 634.2 µs; Rsc = 15.8 MOmega ; Cmc = 11.5 pF; tau lag = 20 µs; p = 0.70; and c = 0.70. Calibration bar: 0.75 ms, 500 pA. B: voltage dependence of activation time constants. Time constants were obtained from exponential fits of activating current traces; voltage values are the average transmembrane voltage over the last 1 ms of the command step. Because of variations in the transmembrane voltage between cells, the time constants were linearly interpolated onto a standardized voltage base before averaging across cells. Experiments were performed using Tris electrodes.

On longer time scales, a slower component of VGCC activation could be observed in some Ba2+ current traces. This slow activation sometimes was present during the first 100 ms of the 2- to 4-s command steps used to investigate the time course of inactivation (Fig. 8B, black-triangle). In addition, a slowly developing inward current could be elicited by 100-ms command steps (omega CTX-sensitive difference traces in Fig. 3, B and C; nifedipine-sensitive difference traces in Fig. 3, B and D). In most cases, a slowly decaying tail current was associated with the slowly developing inward current. This slow component of VGCC activation and deactivation may correspond to the "mode 2" gating augmented by dihydropyridine agonists during single-channel recordings of L-type VGCCs (Tsien et al. 1986). However, the conditions required to evoke this slow component were not investigated in this study, and it is not clear if this component would be prominent under physiological conditions.



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Fig. 8. Voltage dependence of Ba2+ current inactivation. A: no significant low-threshold inactivation was observed. Averages of 4 current traces elicited by a 100-ms step to 0 mV from a 50-ms prepulse to -100 or -40 mV are superimposed on an average of 4 current traces elicited by a 100-ms step to 0 mV directly from the holding potential of -70 mV. black-triangle, small inward current elicited by the -40 mV prepulse. Calibration bar: 25 ms, 800 pA. B: a slow, high-threshold inactivation was observed. Averages of 4 current traces were elicited by 4-s command steps to -25, -15, and +5 mV from a holding potential of -70 mV. black-triangle, slow component of Ba2+ current activation observed at the beginning of the current traces evoked by command steps to -25 and -15 mV. Calibration bar: 1 s, 300 pA. C: voltage dependence of inactivation time constants. Time constants were obtained from exponential fits over the inactivating phase of currents elicited by 2- to 4-s command steps. Voltage values are the average transmembrane voltage over the duration of the curve fit. Time constants from individual cells are distinguished by different symbols; missing data are due to inactivation amplitudes that were too small to generate accurate time constants. Experiments were performed using both Cs+ and Tris electrodes.

A slow inactivation was observed at depolarized voltages, suggesting that a majority of VGCCs recorded from acutely isolated DCN neurons were of the HVA subtype. Unlike LVA VGCCs, the inward current did not appear to have any significant low-threshold inactivation. No significant changes in the peak current magnitude elicited by a 100-ms command step to 0 mV were observed with 50-ms precommand steps to either -100 or -40 mV from the holding potential of -70 mV (Fig. 8A). Across experiments, 50-ms precommand steps to -100 mV resulted in peak current magnitudes that were 99.6 ± 2.0% (n = 8) of the peak current magnitudes elicited directly from the holding potential of -70 mV; and 50-ms precommand steps to -40 mV resulted in peak current magnitudes that were 92.2 ± 4.2% (n = 8) relative to currents elicited directly from -70 mV. Consistent with the behavior of HVA VGCCs, Ba2+ currents elicited by 2- to 4-s command steps from a holding potential of -70 mV showed a slow inactivation at voltages higher than -25 mV (Fig. 8B). Although this inactivation was faster with more depolarized voltages, inactivation time constants typically remained >1 s at most voltages (Fig. 8C), which is more than three orders of magnitude larger than the activation time constants at similar voltages (Fig. 7B).

It is possible that individual VGCC subtypes exhibited different voltage-dependent properties so that the voltage dependence of the overall Ba2+ current represented the average behavior of these individual subtypes. Comparison of the difference traces obtained during exposure to various antagonists (Fig. 3) suggests that the individual VGCC subtypes shared similar biophysical properties. However, these currents were elicited at a single voltage, and these similarities may not be present over a range of voltages. To compare the biophysical properties of VGCC subtypes over a wide voltage range, Ba2+ currents were elicited by a range of 10-ms command steps before and during the application of 10 µM nifedipine. The nifedipine-sensitive portion of the current was estimated by subtracting the currents obtained during exposure to nifedipine from the currents obtained before nifedipine had been applied. Some differences between nifedipine-sensitive and -insensitive VGCCs were found: nifedipine-insensitive VGCCs had more steeply voltage-dependent open-channel Ba2+ currents (Fig. 9A, ) and activated more rapidly below -15 mV (Fig. 9D, ) when compared with nifedipine-sensitive currents (Fig. 9, A and D, open circle ). However, the steady-state open probability (Fig. 9B) and steady-state current magnitudes (Fig. 9C) were similar over a wide voltage range as were the activation time constants above -10 mV (Fig. 9D). Although the antagonism of VGCCs by some dihydropyridine compounds is modulated by voltage at physiological pH (Sanguinetti and Kass 1984), the saturating concentration of nifedipine used in these experiments should provide a complete block of L-type VGCCs regardless of voltage. Unfortunately, it was not possible to compare voltage dependence of VGCC subtypes in other cells or with additional pharmacological agents; the combination of pharmacological effects and rundown usually resulted in currents that were too small to obtain accurate measures over the voltage range of interest.



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Fig. 9. Voltage-dependent properties of nifedipine-sensitive and insensitive currents. Data in A-D are measures obtained from nifedipine-sensitive (open circle ) and nifedipine-insensitive () currents from an individual cell. Averages of 4 Ba2+ currents were obtained before and during a 5-min exposure to nifedipine (10 µM); nifedipine-sensitive currents were estimated subsequently as the difference between the overall Ba2+ and nifedipine-insensitive currents. Rather than directly subtracting 2 sets of current traces elicited by the same series of command steps, nifedipine-insensitive currents were interpolated linearly onto the transmembrane voltages associated with the overall Ba2+ currents before subtraction. A: voltage dependence of open-channel Ba2+ currents. Averages of 4 tail currents elicited by the transition to repolarizing potentials ranging from -150 to +45 mV after a 5-ms command step to +50 mV were obtained before and during exposure to nifedipine. Open-channel Ba2+ current-voltage relationships were constructed as described in Fig. 5C. B: voltage dependence of steady-state VGCC open probability. Averages of 4 tail currents elicited by the transition to a repolarizing voltage of -80 mV after 10-ms command steps ranging from -100 to +50 mV were obtained before and during exposure to nifedipine. Steady-state open-probability voltage relationships were constructed as described in Fig. 6C. C: voltage dependence of steady-state Ba2+ current magnitudes. Averages of 4 activating currents elicited by 10-ms command steps ranging from -100 to +50 mV from a holding potential of -60 mV were obtained before and during exposure to nifedipine. Steady-state current-voltage relationships were constructed as described in Fig. 6D. D: voltage dependence of the time course of Ba2+ current activation. Exponential fits of the activating currents used in C were performed as described in Fig. 7B. Time constants from the nifedipine-insensitive current data () are missing at more depolarized voltage levels because the current amplitudes over the specified fit windows were too small to generate accurate values. Experiments were performed using Tris electrodes.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
APPENDIX
REFERENCES

The results of this study show that the majority of neurons isolated from guinea pig DCN possess VGCCs. There are many neuronal populations residing within the guinea pig DCN, and our results indicate that a variety of VGCC subtypes are present in these neurons. Given these conditions, it is reasonable to expect VGCCs to exhibit a diverse range of biophysical properties, and the apparent uniformity of VGCC properties in isolated DCN neurons is an intriguing result. In all neurons from which Ba2+ currents were observed, VGCCs had similar steady-state voltage dependence, activated and deactivated rapidly, and exhibited no voltage-dependent inactivation on physiologically relevant time scales. Regardless of subtype, the VGCCs present in isolated DCN neurons are poised to produce a rapid Ca2+ influx in response to a large depolarization.

The use of the isolated cell preparation is advantageous for the characterization of voltage-gated conductances in adult mammalian neurons (Kay and Wong 1986); however, the acute isolation procedure poses some potential problems for the interpretation of the results obtained in the present study. First, the trauma resulting from the enzymatic and mechanical dispersion of neuronal tissue could alter the expression of proteins contributing to the electrophysiologic properties of neurons, resulting in patterns of conductances that would not be observed in recordings from intact preparations. Acutely isolated ventral cochlear nucleus neurons produce responses to depolarizing current injection that are similar to responses obtained from in vitro slice preparations (Manis and Marx 1991), indicating that these neurons do not express different sets of conductances as a result of the acute isolation procedure. Second, the acute isolation procedure may modify the properties of existing membrane proteins, including voltage-gated conductances. Trypsin, the enzyme used in the dissociation procedure, is known to inactivate N-methyl-D-aspartate receptors (Allen et al. 1988). However, the use of trypsin in previous studies did not appear to adversely affect VGCCs (Kay and Wong 1987; Thompson and Wong 1991). In addition, the biophysical and pharmacological properties of VGCCs in the present study are similar to those found in neurons dissociated without the use of trypsin (Regan 1991; Zidanic and Fuchs 1995). Therefore the use of the acute dissociation procedure is not likely to adversely affect VGCCs in the present study.

VGCC subtypes

Acutely isolated guinea pig DCN neurons possess a variety of HVA VGCC subtypes. The divalent ion pharmacology is more typical of HVA rather than LVA VGCCs: unlike their LVA counterparts, HVA VGCCs are usually more selective for Ba2+ than for Ca2+ and are blocked more potently by Cd2+ than by Ni2+ (Bean 1989; Huguenard 1996). Responses to subtype-specific antagonists also confirmed the presence of HVA VGCCs. All neurons tested contained a significant L-type component; only some neurons contained N- and P/Q-type components, and these appeared in varying amounts. The observation of a consistent nifedipine-sensitive component and variable omega Aga IVA- and omega CTX GVIA-sensitive components could be explained by the differential localization of VGCC subtypes within a single cell. In hippocampal pyramidal cells, L-type VGCCs cluster in the proximal dendrites (Westenbroek et al. 1990); whereas N-type VGCCs are distributed over the entire dendritic arbor (Westenbroek et al. 1992). Because the acute cell isolation procedure produces neurons that only possess a cell body and proximal processes, it is expected that such a distribution of VGCCs would result in a consistent nifedipine-sensitive component, and a reduced, if not erratic, omega CTX GVIA-sensitive component.

Most neurons possess a significant inward current that remains in the presence of nifedipine after exposure to omega Aga IVA and omega CTX GVIA or during a simultaneous exposure to all three antagonists. The majority of this remaining inward current is blocked by 50 µM Cd2+ and may correspond to R-type VGCCs (Ellinor et al. 1993; Zhang et al. 1993). In the present study, the antagonist-insensitive component exhibits a slower activation and a more pronounced inactivation than the L-, N-, and P/Q-type components. This is consistent with the voltage-dependent properties of R-type VGCCs, which exhibit many similarities to LVA VGCCs (Bourinet et al. 1996; Ellinor et al. 1993; Soong et al. 1993). It is possible that this antagonist-insensitive component could consist of LVA VGCCs as well; however, the high sensitivity of residual currents to Cd2+ and the insensitivity of the overall current to a hyperpolarized prepulse voltage suggest that these channels belong to the HVA family. It is also possible that Q-type VGCCs could contribute to this antagonist-insensitive component. Although the concentrations of the antagonists used should provide a complete block of L-, N-, and P-type VGCCs, the concentration of omega Aga IVA used in these experiments was only slightly greater than the IC50 for Q-type channels (Randall and Tsien 1995). In some neurons, it is possible that additional omega Aga IVA would have blocked a larger portion of the antagonist-insensitive current by providing a more complete block of Q-type channels. However, several neurons exhibited little or no response to 100 nM omega Aga IVA, and it is unlikely that additional omega Aga IVA would have provided a further block of the remaining current in these cells.

The results of the study do not provide any information about the distribution of VGCCs throughout the DCN. Various cell populations reside within the guinea pig DCN (Hackney et al. 1990), and the use of the isolated cell preparation does not permit positive morphological identification of cell types. Without morphological identification, there is no way to determine whether the various VGCC subtypes observed in the present study are distributed differentially among the individual DCN cell populations. In addition, the results of this study do not provide any information about the distribution of VGCC subtypes throughout the dendritic arbors of the various cell types. A VGCC-mediated Ca2+ influx into the dendrites of DCN cartwheel and pyramidal cells can be elicited by somatic action potentials (Manis and Molitor 1996; Molitor and Manis 1996); however, these dendritic processes inevitably are destroyed by the cell-isolation procedure. Recordings from hippocampal pyramidal and cerebellar Purkinje neurons have shown that the relative levels of VGCC subtypes differed between the soma and the dendrites of these neurons (Kavalali et al. 1997; Mouginot et al. 1997). Therefore it is possible that the dendrites of DCN neurons possess VGCCs that differ from those characterized in this study.

Voltage-dependent properties

The voltage-dependent properties of VGCCs suggest that these conductances are suited to respond to action potentials. A large depolarization from rest is required to activate VGCCs in isolated DCN neurons: <10% of maximal activation is achieved below -40 mV, whereas >90% of maximal activation is achieved above +10 mV. An action potential would maximally activate VGCCs, whereas a subthreshold synaptic depolarization would result in little or no VGCC activation. In addition, the activation time course is rapid enough at depolarized voltages to reach steady-state levels within the duration of an action potential. The lack of any low-threshold inactivation should prevent previous neuronal activity from altering the response of VGCCs to action potentials; the sustained depolarization required to produce any voltage-dependent inactivation is not likely to occur during periods of normal neuronal activity. However, VGCCs are known to inactivate due to elevated internal Ca2+ (de Leon et al. 1995; Imredy and Yue 1994). It is not clear whether elevated intracellular Ca2+ inactivates VGCCs in DCN neurons; the use of Ba2+ as the charge carrier minimized this effect so that voltage-dependent inactivation could be investigated in isolation.

Once activated, VGCCs are capable of producing a large Ca2+ influx. An asymmetric distribution of Ba2+ in these experiments (10 mM Ba2+ outside, 11 mM EGTA inside) results in large inward currents at hyperpolarized voltages and produces an inward rectification at depolarized voltages, which prevents the reversal to an outward current. A similar asymmetric Ca2+ distribution exists in vivo, and the voltage dependence of the Ca2+ influx should exhibit similar properties under physiological conditions. The voltage dependence of open-channel currents suggests that the bulk of the Ca2+ influx will occur after the action potential peak during the repolarizing phase before VGCCs deactivate in response to hyperpolarized voltage levels (Llinás et al. 1981). However, imaging studies at the parallel fiber-stellate cell synapse in the rat cerebellum indicate that the Ca2+ influx in presynaptic terminals becomes more prominent during the rising phase of parallel fiber-mediated action potentials at physiological temperatures (Sabatini and Regehr 1996). It is possible that VGCCs in DCN neurons behave in a similar fashion at physiological temperatures, resulting in a Ca2+ influx that occurs more rapidly with respect to action potential initiation.

Functional implications

VGCCs may be involved in producing different discharge patterns in DCN neurons. VGCCs are likely to be involved in the evoked responses of cartwheel cells, which are a major population of inhibitory interneurons in the superficial DCN. Cartwheel cells respond to depolarizing current injection with a burst of fast action potentials superimposed on a slow depolarization that has been attributed to VGCCs (Agar et al. 1996; Manis et al. 1994; Zhang and Oertel 1993). In contrast, VGCCs may contribute indirectly to the evoked responses of pyramidal cells, which form the main projection pathway from the DCN to the inferior colliculus. Pyramidal cells are only capable of generating Ca2+-dependent action potentials in the presence of voltage-gated Na+ and K+ conductance antagonists (Hirsch and Oertel 1988), and models incorporating only voltage-gated Na+ and K+ conductances are capable of reproducing many pyramidal cell responses to intracellular current injection (Hewitt and Meddis 1995; Kim et al. 1994). However, a slow afterhyperpolarization observed in pyramidal cells after a sustained discharge of action potentials (Hirsch and Oertel 1988; Manis 1990) may be attributed to Ca2+-activated K+ conductances, which would be activated by the Ca2+ influx through VGCCs. Thus VGCCs may play different roles in determining the patterns of evoked responses across neuronal populations of the DCN.

VGCCs are also capable of regulating neuronal responses through the actions of Ca2+ as an intracellular second messenger. Despite the differential involvement of VGCCs in the generation of evoked responses, somatically evoked action potentials result in a VGCC-mediated Ca2+ influx into the soma and proximal dendrites of both pyramidal and cartwheel cells (Manis and Molitor 1996; Molitor and Manis 1996). Because Ca2+ is an important intracellular second messenger (Tsien and Tsien 1990), an evoked Ca2+ influx through VGCCs could play a role in regulating neuronal responses. Ca2+ can interact with and regulate other signaling pathways in the DCN, such as protein kinase C, which regulates the strength of synaptic transmission at parallel fiber synapses (Francis and Manis 1995; Scott and Manis 1992); and intracellular Ca2+ release through inositol 1,4,5-trisphosphate receptors (Mignery et al. 1989; Ryugo et al. 1995), which may be activated through metabotropic glutamate receptors present at parallel fiber and auditory nerve synapses (Molitor and Manis 1997). A Ca2+ influx through VGCCs could act on many different time scales, from the short-term activation of ionic conductances (such as Ca2+-activated K+ conductances), to the long-term regulation of synaptic responses. Regardless of the contribution of VGCCs to discharge patterns, an influx of Ca2+ through VGCCs may be an important indicator of neuronal activity and may have significant implications for information processing in this nucleus.


    APPENDIX
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
APPENDIX
REFERENCES

The study of voltage-dependent conductances requires the accurate control of transmembrane voltage. In many circumstances, the single-electrode, whole cell voltage-clamp technique can be used to provide this control. However, the experimental conditions in the present study resulted in deviations of the transmembrane voltage from the presented command potential. This discrepancy arises from two sources: the voltage drop due to the flow of ionic current across a finite series resistance and the filtering of the command waveform by the membrane capacitance and series resistance. Therefore accurate estimates of the transmembrane voltage are needed to assess the voltage dependence of the Ba2+ currents presented in this study.

A method for calculating the transmembrane voltage can be derived from a simple circuit used to model the isolated neurons in these experiments. The cell is modeled as a membrane resistance Rm in parallel with a membrane capacitance Cm and a current source Ix(t); an additional resistance Rs is added in series to represent the access between the pipette tip and cell interior. Under the conditions of these experiments, it is assumed that Rm >> Rs, which effectively eliminates Rm and reduces the model circuit to Rs in series with Cm and Ix(t). The solution to the differential equation governing this reduced circuit can be expressed in the Laplace domain as a function of complex frequency (s)
<IT>V</IT><SUB><IT>m</IT></SUB>(<IT>s</IT>)<IT>=</IT><FR><NU><IT>V</IT><SUB><IT>c</IT></SUB>(<IT>s</IT>)<IT>−</IT><IT>R</IT><SUB><IT>s</IT></SUB><IT>I</IT><SUB><IT>x</IT></SUB>(<IT>s</IT>)</NU><DE><IT>1+</IT><IT>s</IT><IT>&tgr;</IT><SUB><IT>m</IT></SUB></DE></FR> (A1)
where Vm(s) is the transmembrane voltage, Vc(s) is the command potential, Ix(s) is the sum of all ionic currents flowing across the membrane, and tau m is the membrane time constant, which is a product of Rs and Cm. The physical interpretation of Eq. A1 is that Vm(t) can be obtained by filtering the difference Vc(t) - Rs·Ix(t) with a single-pole low-pass filter having a -3 dB cutoff frequency of 1/2pi tau m. In practice, the transmembrane voltage is calculated by convolving Vc(t- Rs·Ix(t) with the impulse response of the membrane filter, which is a decaying exponential with amplitude 1/tau m and time constant tau m.

The properties of the membrane filter must be known to calculate the transmembrane voltage. The values of Rs and tau m can be obtained from currents elicited by command steps that do not activate ionic currents. In the absence of any on-line amplifier compensation or off-line capacitive current subtraction, the response of the model circuit to a hyperpolarizing command step of magnitude Delta V will be a decaying exponential with amplitude Delta V/Rs and time constant tau m. For the majority of neurons used in the present study, the capacitive currents evoked by hyperpolarizing command steps were more accurately described by a sum of two exponentials
<IT>I</IT><SUB><IT>c</IT></SUB>(<IT>t</IT>)<IT>=</IT><FR><NU><IT>&Dgr;</IT><IT>V</IT></NU><DE><IT>R</IT><SUB><IT>s</IT></SUB></DE></FR> [<IT>A</IT><SUB><IT>f</IT></SUB><IT> exp</IT>(−<IT>t</IT><IT>/&tgr;</IT><SUB><IT>f</IT></SUB>)<IT>+</IT>(<IT>1−</IT><IT>A</IT><SUB><IT>f</IT></SUB>)<IT> exp</IT>(−<IT>t</IT><IT>/&tgr;</IT><SUB><IT>s</IT></SUB>)] (A2)
where 0 <=  Af <=  1 and tau f < tau s. For these neurons, the impulse response of the membrane filter is also a sum of two exponentials, the first with amplitude Af/tau f and time constant tau f and the second with amplitude (1 - Af)/tau s and time constant tau s. A biexponential capacitive current suggests that the isolated neurons in these experiments did not have a uniform transmembrane voltage and were more accurately modeled as two isopotential compartments joined in series with a finite resistance. However, typical values for the amplitudes (Af > 0.9) and time constants (tau s congruent  3 · tau f) of capacitive currents suggest that in most cells this second compartment possessed only a small fraction of the total membrane capacitance and, hence, a small fraction of the total membrane surface. Therefore it is reasonable to assume that the majority of VGCCs contributing to the overall Ba2+ current in the present study reside in one compartment and are responding to a uniform transmembrane voltage.

The true command potential also must be known to calculate the transmembrane voltage. Many voltage-clamp amplifiers provide compensation circuitry that use estimates of Rs and Cm to modify the command potential so that the transmembrane voltage will more faithfully replicate the desired command potential. The Axopatch 200 amplifier used in these experiments provides two separate compensation pathways by which the command potential is modified: prediction and correction compensation. Prediction compensation reduces low-pass filtering of the membrane by boosting the high-frequency components of the command potential
<IT>V</IT><IT><SUP>*</SUP></IT><SUB><IT>c</IT></SUB>(<IT>s</IT>)<IT>=</IT><IT>V</IT><SUB><IT>c</IT></SUB>(<IT>s</IT>) <FR><NU><IT>1+</IT><IT>s</IT><IT>&tgr;</IT><SUB><IT>p</IT></SUB></NU><DE><IT>1+</IT><IT>s</IT><IT>&tgr;</IT><SUB><IT>p</IT></SUB>(<IT>1−</IT><IT>p</IT>)</DE></FR> (A3)
where tau p is the prediction compensation time constant, which is a product of the series resistance Rsc and membrane capacitance Cmc amplifier settings; and p is the fraction of prediction compensation (%/100) amplifier setting. If the amplifier settings are such that tau p congruent  tau m, the prediction compensation circuitry effectively reduces the membrane time constant from tau m to tau p (1 - p). Correction compensation reduces the voltage drop due to the flow of ionic current across a finite series resistance by adding a scaled and filtered version of this current to the command potential
<IT>V</IT><IT><SUP>*</SUP></IT><SUB><IT>c</IT></SUB>(<IT>s</IT>)<IT>=</IT><IT>V</IT><SUB><IT>c</IT></SUB>(<IT>s</IT>)<IT>+</IT><IT>cR</IT><SUB><IT>sc</IT></SUB> <FR><NU><IT>I</IT><SUB><IT>x</IT></SUB>(<IT>s</IT>)</NU><DE><IT>1+</IT><IT>s</IT><IT>&tgr;</IT><SUB><IT>lag</IT></SUB></DE></FR> (A4)
where tau lag is the lag filter time constant and c is the fraction of correction compensation (%/100) amplifier setting. If Ix(t) is not significantly attenuated by the lag filter, the correction compensation circuitry effectively reduces the series resistance from Rs to Rs-c·Rsc. The modifications to the command voltage by the prediction and correction compensation superimpose, so that the modified command potential V*c(t) is obtained by filtering the original command potential Vc(t) with the prediction compensation circuitry (Eq. A3), and adding the lag-filtered product of c·Rsc·Ix(t) (Eq. A4). The difference between the modified command potential V*c(t) and the product Rs·Ix(t) then can be attenuated by the membrane filter to obtain the transmembrane voltage Vm(t) (Eq. A1).


    ACKNOWLEDGMENTS

We thank Dr. Nicholas Saccomano of Pfizer for the generous gift of omega Aga IVA.

This work was supported by National Institute of Deafness and Other Communication Disorders Grants RO1 DC-00425 and K04 DC-00048 to P. B. Manis, a National Science Foundation predoctoral fellowship to S. C. Molitor, and a grant from the W. M. Keck Foundation.


    FOOTNOTES

Address for reprint requests: S. C. Molitor, The Johns Hopkins University School of Medicine, 420 Ross Research Bldg., 720 Rutland Ave., Baltimore, MD 21205.

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received 10 June 1998; accepted in final form 30 October 1998.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
APPENDIX
REFERENCES

0022-3077/99 $5.00 Copyright © 1999 The American Physiological Society