Three Kinetically Distinct Ca2+-Independent Depolarization-Activated K+ Currents in Callosal-Projecting Rat Visual Cortical Neurons

Rachel E. Locke and Jeanne M. Nerbonne

Department of Molecular Biology and Pharmacology, Washington University School of Medicine, St. Louis, Missouri 63110

    ABSTRACT
Abstract
Introduction
Methods
Results
Discussion
References

Locke, Rachel E. and Jeanne M. Nerbonne. Three kinetically distinct Ca2+-independent depolarization-activated K+ currents in callosal-projecting rat visual cortical neurons. J. Neurophysiol. 78: 2309-2320, 1997. Whole cell, Ca2+-independent, depolarization-activated K+ currents were characterized in identified callosal-projecting (CP) neurons isolated from postnatal day 7-16 rat primary visual cortex. CP neurons were identified in vitro after in vivo retrograde labeling with fluorescently tagged latex microbeads. During brief (160-ms) depolarizing voltage steps to potentials between -50 and +60 mV, outward K+ currents in these cells activate rapidly and inactivate to varying degrees. Three distinct K+ currents were separated based on differential sensitivity to 4-aminopyridine (4-AP); these are referred to here as IA, ID, and IK, because their properties are similar (but not identical) K+ currents termed IA, ID, and IK in other cells. The current sensitive to high (>= 100 µM) concentrations of 4-AP (IA) activates and inactivates rapidly; the current blocked completely by low (<= 50 µM) 4-AP (ID) activates rapidly and inactivates slowly. A slowly activating, slowly inactivating current (IK) remains in the presence of 5 mM 4-AP. IA, ID, and IK also were separated and characterized in experiments that did not rely on the use of 4-AP. All CP cells express all three K+ current types, although the relative densities of IA, ID, and IK vary among cells. The experiments here also have revealed that IA, ID, and IK display similar voltage dependences of activation and steady state inactivation, whereas the kinetic properties of the currents are distinct. At +30 mV, for example, mean ± SD activation tau s are 0.83 ± 0.24 ms for IA, 1.74 ± 0.49 ms for ID, and 14.7 ± 4.0 ms for IK. Mean ± SD inactivation tau s for IA and ID are 26 ± 7 ms and 569 ± 143 ms, respectively. Inactivation of IK is biexponential with mean ± SD inactivation time constants of 475 ± 232 ms and 3,128 ± 1,328 ms; ~20% of the 4-AP-insensitive current is noninactivating. For all three components, activation is voltage dependent, increasing with increasing depolarization, whereas inactivation is voltage independent. Both IA and IK recover rapidly from steady state inactivation with mean ± SD recovery time constants of 38 ± 7 ms and 79 ± 26 ms, respectively; ID recovers an order of magnitude more slowly (588 ± 274 ms). The properties of IA, ID, and IK in CP neurons are compared with those of similar currents described previously in other mammalian central neurons and, in the accompanying paper, the roles of these conductances in regulating the firing properties of CP neurons are explored.

    INTRODUCTION
Abstract
Introduction
Methods
Results
Discussion
References

A variety of K+ currents with differing kinetic, pharmacological and voltage-dependent properties have been described in neurons in the mammalian CNS (for reviews, see: Rudy 1988; Storm 1990). In addition, in most CNS neurons, multiple types of K+ currents have been shown to be coexpressed. As in other excitable cells (Barry and Nerbonne 1996; Rudy 1988), K+ currents in mammalian CNS neurons appear to be more numerous and more diverse than the other types (i.e., Na+, Ca2+) of currents expressed. This multiplicity and diversity has a physiological significance in that the various K+ currents function to control resting potentials, the threshold for action potential generation, the rate of action potential repolarization, and the rate of repetitive firing. In addition, differences in the types, densities, and/or the detailed properties of the K+ currents expressed are expected to contribute importantly to determining the regular-spiking, fast-spiking, and intrinsically bursting firing patterns in central neurons (Connors and Gutnick 1990; McCormick et al. 1985).

Depolarization-activated, Ca2+-independent K+ currents are expected to influence action potential repolarization and the frequency of repetitive firing. Two types of K+ currents that are likely to subserve these functions have been described in a number of mammalian CNS neurons: a rapidly activating and inactivating current that is blocked by4-aminopyridine and usually is referred to as IA (Beck et al. 1992; Budde et al. 1992; Ficker and Heinemann 1992; Huguenard et al. 1991; Numann et al. 1987; Segal and Barker 1984; Storm 1988; Surmeier et al. 1989; Wu and Barish 1992); and a 4-aminopyridine-insensitive, delayed, outwardly rectifying K+ current, usually referred to as IK (Beck et al. 1992; Budde et al. 1992; Huguenard and Prince 1991; Numann et al. 1987; Rudy 1988; Segal and Barker 1984; Storm 1990). These are broad classifications, however, and the detailed time- and voltage-dependent properties of the various currents termed IA and IK do vary among cells (Rudy 1988; Storm 1990). In addition, in some CNS neurons, an additional, rapidly activating, slowly inactivating Ca2+-independent K+ current has been identified based on sensitivity to micromolar concentrations of 4-aminopyridine and, in some cases, also to the dendrotoxins (Ficker and Heinemann 1992; Hammond and Crepel 1992; McCormick 1991; Stefani et al. 1995; Surmeier et al. 1991; Wu and Barish 1992). These currents have been referred to variably as ID or "slow transient" currents to distinguish them from IA and IK. It is unclear if ID is expressed in all central neurons because, in many cases, this possibility has not been explored directly.

Although there have been many studies characterizing the repolarizing K+ currents in CNS neurons, few have been completed on neocortical neurons (but see Albert and Nerbonne 1995; Foehring and Surmeier 1993; Hamill et al. 1991; Spain et al. 1991). This largely reflects the heterogeneity of cortical neurons and the difficulties associated with reliably identifying specific cell types. In previous studies on neocortical neurons, for example, the cells either were not identified directly (Foehring and Surmeier 1993; Spain et al. 1991) or were identified based on in vitro morphology (Hamill et al. 1991), which, as we have shown previously, is not a reliable method for identifying cortical neurons (Giffin et al. 1991). To circumvent these difficulties, we have exploited in vivo retrograde labeling to permit the in vitro identification of cortical neurons (Giffin et al. 1991; Katz and Iarovici 1990; Katz et al. 1984; Solomon et al. 1993). In the experiments described here, we have characterized the detailed time- and voltage-dependent properties of the Ca2+-independent, depolarization-activated K+ currents in isolated, identified callosal-projecting (CP) neurons from postnatal rat primary visual cortex. CP cells were selected to correspond to the "regular-spiking" phenotype described previously in recordings from cortical neurons in vivo and in the in vitro slice preparation (Connors and Gutnick 1990; McCormick et al. 1985). Here, we show that there are three kinetically and pharmacologically distinct Ca2+-independent, voltage-gated K+ currents expressed in CP cells that we refer to as IA, ID, and IK because their properties are similar (although not identical) to IA, ID, and IK currents described in other mammalian central neurons. In the companion paper (Locke and Nerbonne 1997), we demonstrate that CP neurons are indeed regular spiking, and we explore the roles of IA, ID, and IK in shaping the waveforms of action potentials and in regulating the firing properties of CP neurons.

    METHODS
Abstract
Introduction
Methods
Results
Discussion
References

Isolation and identification of callosal-projecting neurons

Whole cell voltage-clamp recordings were obtained from identified CP visual cortical neurons isolated from postnatal day 7-16 (P7-P16) Long Evans rat pups. Methods for identifying and isolating CP neurons have been previously described (Giffin et al. 1991; Solomon et al. 1993). Briefly, CP neurons were labeled in vivo by pressure injection of fluorescein tagged latex beads (Katz and Iarovici 1990) into the right visual cortex on postnatal day 4, 5, or 6. On postnatal day 7-16, animals were anesthetized with 5% halothane, and the left visual cortex was removed. Cells were dissociated, plated on a glial monolayer (Raff et al. 1979), and maintained in a 95% O2-5% CO2 37°C incubator. CP neurons were identified in vitro before electrophysiological recordings under epifluorescence illumination by the presence of fluorescently labeled beads in the cell soma.

Electrophysiological recordings

Whole cell patch-clamp recordings were obtained at room temperature (23-25°C) from identified, CP neurons within 30 h of cell dissociation. Data were collected using a Dagan 8900 patch-clamp amplifier, and experimental parameters were controlled by an IBM-compatible 486 computer with TL-1 interface using PClamp (Axon Instruments). Neurons without extensive processes were selected for recording (see below). All voltage-clamp data were collected in the presence of 1 µM tetrodotoxin (TTX) and 100 µM Cd2+ to block voltage-gated Na+ and Ca2+ currents, respectively. Electrodes were fabricated from soda lime glass (Kimble No. 73811) on a two-stage puller and the shanks (within ~50 µm of the tips) were coated with silicone elastomer (Sylgard; Dow Corning). Pipette resistances were 1.2-2.5 MOmega after fire polishing. Series resistances, estimated from the decays of the uncompensated capacitative transients, were 2.5-5 MOmega and were compensated electronically by 80-90%. Because peak current amplitudes were <8 nA, voltage errors arising from the uncompensated series resistance were always <8 mV and were not corrected. Depolarization-activated K+ currents were evoked routinely during voltage steps to potentials between -50 and +60 mV from a holding potential (HP) of -70 mV. Other voltage-clamp protocols are described in the text. Data were sampled at various rates (1-70 kHz) depending on the particular experimental paradigm, and current signals were filtered at 5 kHz before digitization and storage.

Solutions

The standard bath solution contained (in mM) 140 NaCl, 2.5 CaCl2, 4 KCl, 2 MgCl2, 5 glucose, 10 N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES), 0.1 CdCl2, and 1 µM TTX (pH 7.4; 305 mOsm). Because millimolar concentrations of Cd2+ have been shown in some cells to alter the time- or voltage-dependent properties of K+ currents (Mayer and Sugiyama 1988), some experiments also were completed with the organic Ca2+ channel blocker nifedipine used in place of Cd2+ to block currents through voltage-gated Ca2+ channels. No differences in the time- or voltage-dependent properties of the K+ currents were observed. The pipette solution contained (in mM) 135 KCl, 10 ethylene glycol-bis(beta -aminoethyl ether)-N,N,N',N'-tetraacetic acid, 10 HEPES, 5 glucose, 3 MgATP, and 0.5 NaGTP (pH 7.4; 305 mOsm). Stock solutions of 4-aminopyridine (4-AP) or dendrotoxin (DTX) were diluted into the standard bath solution immediately before experiments. Tetraethylammonium chloride (TEACl) was substituted for NaCl in the bath solution on an equimolar basis. All chemicals, except DTX, were obtained from Sigma. DTX was obtained from Alamone Laboratories, Israel. Drugs were applied using a perfusion pipette, which consisted of four pieces of PE10 tubing encased in a small glass tube. During superfusion, the pipette was placed within 200 µm of the cell, and the bath volume was kept constant using a small glass tube, attached to a large reservoir, that controlled the solution level by capillary action.

Data analyses

Data were compiled and analyzed using Clampfit (Axon Instruments), Excel (Microsoft), CSS Statistica (Stat Soft), and Nfit (Island Products). All data are means ± SD. The plateau current was defined as the current remaining 150 ms after the onset of voltage steps, and the peak current was defined as the maximum value the current attained during the 160-ms pulse. For each cell, the spatial control of membrane voltage was assessed by directly analyzing the decay of the capacitative currents. Only cells with capacitative transients well described by single exponential functions were analyzed further. Leak currents were always <100 pA at -70 mV and were not corrected. The mean ± SD input resistanceof analyzed CP cells was 1.7 ± 1.1 GOmega (n = 70), and the mean ±SD whole cell membrane capacitance of these cells was 14.4 ± 6.6 pF (n = 70). Equations used in fitting the data are presented in the text. Statistical significance was examined using analysis of variance (ANOVA) and Student's t-test; P values are presented in the text.

    RESULTS
Abstract
Introduction
Methods
Results
Discussion
References

Depolarization-activated currents in callosal-projecting neurons

In the experiments here, CP rat visual cortical neurons were labeled in vivo by pressure injection of fluorescein-tagged latex beads (Katz and Iarovici 1990) into the right visual cortex on postnatal day 4, 5, or 6. After isolation at postnatal day 7-16, CP neurons were identified in vitro under epifluorescence illumination before electrophysiological recordings. With voltage-gated Ca2+ and Na+ currents blocked, whole cell, depolarization-activated outward currents were recorded routinely from isolated, identified P7-P16 CP neurons (n = 70) during 160-ms voltage steps to potentials between -50 and +60 mV from an HP of -70 mV (Fig. 1). The absolute rates of rise and the amplitudes of the currents increase with increasing depolarization; the largest and most rapidly activating current in each panel in Fig. 1 was evoked at +60 mV. No depolarization-activated currents were evoked when the K+ in the recording pipette was replaced by Cs+ (n = 4). The currents recorded and analyzed here, therefore, are interpreted as reflecting only the activation of Ca2+-independent, depolarization-activated K+ channels. Interestingly, the waveforms of the currents evoked by small depolarizations inactivate more rapidly and to a larger extent than the currents evoked by large depolarizations. In addition, although the currents in all cells activate rapidly, the relative amplitudes of the peak and plateau currents vary markedly among cells (Fig. 1). For the cell in Fig. 1A, for example, the amplitude of the peak current is nearly twice that of the plateau current; the peak-to-plateau current ratios are ~1.5 and 1 for the cells in Fig. 1, B and C, respectively. Similar variability was observed among cells isolated from animals at postnatal ages (P7-P12),as well as in cells maintained for various times in vitro(3-30 h).


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FIG. 1. Waveforms of Ca2+-independent, depolarization-activated K+ currents in callosal-projecting (CP) visual cortical neurons vary among cells. Whole cell K+ currents displayed in this and in subsequent figures were evoked as described in METHODS, unless stated otherwise. Currents plotted in A-C were recorded from 3 different CP cells isolated at postnatal day 9 ~10 h after plating; the cells in A and C were in the same preparation.

Separation of K+ current components

The variability in peak-to-plateau current ratios, as well as the differences in the waveforms of the currents evoked by small and large depolarizations (Fig. 1), suggested that the total Ca2+-independent, depolarization-activated K+ currents recorded in CP visual cortical neurons reflected the simultaneous activation of two (or more) K+ channel types and that these are distributed differentially among CP cells. In some cells, rapidly activating, rapidly inactivating currents (termed IA) have been isolated by taking advantage of the sensitivity of IA to holding potential. Preliminary experiments on CP cells, however, revealed that neither the amplitudes nor the waveforms of the currents are affected by changes in HP between -90 and -70 mV. The waveforms of the outward currents evoked from HPs of -30 and -70 mV are also not significantly different, although the amplitudes of the currents recorded from an HP of -30 mV are reduced substantially (not shown). Similar results were obtained in experiments conducted on 34 cells, suggesting either that CP neurons do not express multiple types of K+ channels or that the voltage dependences of the different K+ channels in these cells are not significantly different. To distinguish between these possibilities, subsequent experiments focused on examining the pharmacological sensitivities of the K+ currents in CP neurons to the K+ channel blocker 4-AP primarily because different types of K+ currents in other cells have been shown to display differential sensitivities to 4-AP (Rudy 1988; Storm 1990).

In these experiments, the effects of varying concentrations (50 µM to 10 mM) of 4-AP on the K+ currents in CP neurons were examined. Control K+ currents were recorded before superfusion of 4-AP-containing bath solutions, and when the effect of 4-AP had reached a steady state, outward currents were recorded again. To obtain the current(s) blocked by different concentrations of 4-AP, records obtained in the presence of (each concentration of) 4-AP were subtracted digitally from the controls (recorded in the absence of 4-AP). Typical examples of the subtracted current waveforms blocked by different concentrations of 4-AP are shown in Fig. 2. As is evident, the current blocked by 50 µM 4-AP (Fig. 2A1) activates rapidly and decays only slightly during the course of 160-ms voltage steps. Increasing the concentration of 4-AP resulted in marked changes in the waveforms of the subtracted current records (Fig. 2A, 2-4). Specifically, the amplitudes of the currents at early times after the onset of depolarizing voltage steps in these subtracted records are increased (relative to the 50-µM subtracted records) as a function of the concentration of 4-AP (Fig. 2A). In contrast, there appears to be little or no change in the amplitudes of the plateau currents in the subtracted records as a function of 4-AP concentration (Fig. 2A). To quantify the effects of varying concentrations of 4-AP, peak and plateau current amplitudes were measured at 3 and 150 ms, respectively, and the percent suppression of the peak and plateau currents (relative to the control peak and plateau current amplitudes in the same cell) was calculated. The percent suppression of the peak and plateau currents were determined in many cells, and mean normalized data are presented in Fig. 2B. As is evident, concentrations of 4-AP >50 µM block increasing amounts of the peak, but not of the plateau, currents. Maximal block of the plateau currents is achieved at 50 µM 4-AP, whereas maximal block of the peak currents is observed at 5 mM (Fig. 2B).


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FIG. 2. Increasing concentrations of 4-aminopyridine (4-AP) block increasing amounts of the peak, but not of the plateau, current. Currents were recorded in control bath solution and in the presence of varying concentrations of 4-AP. Current records displayed in A were obtained by digital subtraction of the current waveforms recorded in 4-AP from the control records in the same cell. Peak and plateau current amplitudes in individual cells were measured at 3 and 150 ms, respectively, after the onset of depolarizing voltages to +30 mV, and the percentages of the peak and plateau currents blocked by 4-AP were determined. A: representative waveforms of the currents blocked by 50 µM (1), 100 µM (2), 500 µM (3), and 5 mM 4-AP (4). Note that the records in 1 and 4 are from the same cell; the records in 2 and 3 are from 2 other cells. B: mean ± SD percentages of the peak and plateau currents blocked by increasing concentrations of 4-AP. n values refer to the number of cells studied.

To isolate the currents blocked by the high concentrations of 4-AP, currents recorded in the presence of 5 mM 4-AP (Fig. 3C) were subtracted from those recorded in 50 µM4-AP (Fig. 3B). Comparison of the waveforms of these currents (Fig. 3E) and the 50-µM 4-AP-sensitive currents (Fig. 3D) reveals that, although both currents activate rapidly, the decay phases of the currents are markedly different. The 50-µM-sensitive currents (Fig. 3D) inactivate slowly and resemble currents referred to as ID, or slow transient currents, in other preparations (Ficker and Heinemann 1992; Storm 1988; Wu and Barish 1992). The currents blocked by 5 mM (but not by 50 µM) 4-AP, in contrast, inactivate rapidly (Fig. 3E) and are similar to IA currents in other cells (for reviews, see Rudy 1988; Storm 1990). The currents remaining in the presence of 5 mM 4-AP (Fig. 3C) are kinetically distinct from both of the 4-AP-sensitive currents (Fig. 3, D and E). Specifically, the 5-mM 4-AP-insensitive currents activate slowly and do not inactivate during160-ms depolarizations at all test potentials, much like delayed rectifier, or IK, currents described in other cell types (for reviews, see Rudy 1988; Storm 1990). Taken together, these data suggest the presence of at least three Ca2+-independent K+ currents in isolated, identified P7-P16 CP neurons. We refer to these currents as IA, ID, and IK primarily because the properties of the currents in CP cells are qualitatively similar to those of currents referred to by these names in other cells (see DISCUSSION).


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FIG. 3. Separation of 2 distinct 4-AP-sensitive current components from the total Ca2+-independent, outward currents. After control currents (A) were recorded, the cell was 1st exposed to 50 µM and then to 5 mM 4-AP. Currents recorded in the presence of 50 µM and 5 mM 4-AP are displayed in B and C, respectively. Currents blocked by 50 µM 4-AP (D) were obtained by subtraction of records in B from those in A. The 5 mM, but not 50 µM, 4-AP-sensitive currents (E) were obtained by subtraction of records in C from those in B. Note that the currents blocked by low concentrations of 4-AP inactivate much more slowly than the currents sensitive only to the high concentration of 4-AP. Similar results were obtained on 13 cells.

To test the validity of the separation of the currents (IA, ID, and IK) based on differential sensitivities to 4-AP described above (Fig. 3), several types of experiments were performed to determine if these current components could be isolated by other methods. The current blocked by high (>= 100 µM) concentrations of 4-AP, which we refer to as IA, inactivates rapidly on membrane depolarization (Fig. 3E), whereas the current component that is blocked completely by 50 µM 4-AP, ID (Fig. 3D), and the 4-AP-insensitive current, IK, inactivate slowly. This finding suggested that it should be possible to isolate IA based on its inactivation kinetics. To explore this possibility, outward K+ currents were recorded during depolarizing voltage steps presented directly from the holding potential of -70 mV (Fig. 4A) and after a 50-ms conditioning prepulse to 0 mV (Fig. 4B). Subtraction of the currents recorded with and without the prepulse revealed rapidly activating and inactivating current waveforms (Fig. 4C) indistinguishable from those obtained on subtraction (Fig. 3E) of currents recorded in the presence of 5 mM (Fig. 3C) from those in 50 µM (Fig. 3B) 4-AP, i.e., IA. Similar results were obtained on eight cells. These results suggest that IA waveforms isolated using 4-AP-sensitive currents reflect the activation of a unique current and not voltage-dependent effects of 4-AP (Thompson 1982).


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FIG. 4. IA also can be isolated using a conditioning prepulse protocol. Currents were recorded during 160-ms voltage steps to potentials of 0, +20, +40, and +60 mV either directly from a holding potential of -70 mV (A) or after a 50-ms prepulse to 0 mV (B); the protocol is illustrated below the records. Current inactivated by the 50-ms prepulse to 0 mV (C) was obtained by subtraction of the records in B from those in A. Similar results were obtained in experiments conducted on 8 other cells.

In several other mammalian neurons, the currents displaying sensitivity to low concentrations (micromolar) of4-AP also are blocked selectively by nanomolar concentrations of the mamba snake toxins, the DTXs (Penner et al. 1986; Stansfeld et al. 1986; Surmeier et al. 1991). To determine the effects of DTX on CP neurons, outward K+ currents were recorded before and after bath application ofalpha -DTX. Although no attempts were made to examine dose-response curves, similar effects were observed at all concentrations of alpha -DTX (20-100 nM) examined (n = 10). A typical example of the effect of alpha -DTX on K+ currents in CP neurons is illustrated in Fig. 5. The alpha -DTX-sensitive currents (Fig. 5C), obtained by subtraction of current recorded in the presence of alpha -DTX (Fig. 5B) from the controls (Fig. 5A), appear indistinguishable from the 50-µM 4-AP-sensitive currents (Fig. 3D). Similar to the findings with 50 µM 4-AP, alpha -DTX blocks a similar percent of the peak (18 ± 9%) and the plateau (17 ± 8%) currents. Thus the pharmacological properties of the current in CP neurons we refer to as ID are similar to those of rapidly activating, slowly inactivating currents described in other cells (Penner et al. 1986; Stansfeld et al. 1986; Surmeier et al. 1991).


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FIG. 5. Rapidly activating, slowly inactivating K+ current (ID) also is blocked by alpha -dendrotoxin (alpha -DTX). Currents were recorded before (A) and after superfusion of 100 nM alpha -DTX (B). DTX-sensitive currents (C), obtained by subtraction of the records in B from those in A, activate rapidly and inactivate slowly, and are similar, therefore, to the currents blocked by 50 µM 4-AP (Fig. 3D). Similar results were obtained on 10 other cells.

As illustrated in Fig. 3C, the currents (IK) remaining in the presence of 5 mM 4-AP are kinetically distinct from the 4-AP-sensitive currents, ID (Fig. 3D) and IA (Fig. 3E). Because IK in other preparations is blocked selectively by millimolar concentrations of TEA (for recent reviews, see Rudy 1988; Storm 1990), the effects of TEA on the Ca2+-independent, depolarization-activated K+ currents in CP neurons also were examined (Fig. 6). The TEA-sensitive currents (Fig. 6C), obtained by subtraction of records in the presence of 30 mM TEA (Fig. 6B) from controls (Fig. 6A), activate slowly and undergo little or no inactivation during 160-ms voltage steps; the waveforms of the TEA-sensitive currents, therefore, appear indistinguishable from the currents remaining in 5 mM 4-AP (Fig. 3C). Similar results were obtained in experiments completed on seven other cells. In addition, currents with identical waveforms (but smaller amplitudes) were obtained in 15 other cells using lower concentrations (3-10 mM) of TEA. Taken together, these experiments have provided independent means of isolating IA (voltage-clamp protocol), ID (DTX sensitivity), and IK (TEA sensitivity), and, in each case, the properties of the currents appear indistinguishable from those separated using 4-AP (see below and DISCUSSION). For all subsequent analyses, therefore, the properties of the currents (IA, ID, and IK) separated using 4-AP were examined. In several cases, the properties of the currents separated using these alternative methods also were analyzed.


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FIG. 6. Tetraethylammonium (TEA) blocks a slowly activating current (IK) in CP neurons. Currents were recorded under control conditions (A) and after superfusion of 30 mM TEA (B). Current blocked by TEA (C) was obtained by subtraction of records in B from those in A. Similar results were obtained on 7 other cells.

It is of interest to note that although IA, ID, and IK were identified readily in all CP cells studied (n = 40), the absolute amplitudes of the individual current components were found to vary markedly among cells. Normalizing current amplitudes for differences in cell sizes revealed similar variability in current densities among cells. IA density, for example, varied between 94 and 489 pA/pF, and the mean IA density was 215 ± 83 pA/pF (n = 40). Similar variations in IK densities were observed; IK density varied over the range of 65 to 465 pA/pF with a mean of 173 ± 94 pA/pF (n = 40). In all cells, the density of ID was substantially less than the density of either IA or IK; ID density ranged from 11 to 127 pA/pF with a mean of 54 ± 31 pA/pF. On average, therefore, the densities of both IA and IK are approximately four times the density of ID. More importantly, these analyses revealed that the relative density of IA, ID, and IK (i.e., the ratio of IA:ID:IK) varies markedly among cells. The variability in the relative densities of IA, ID, and IK likely underlies the differences (Fig. 1) in the waveforms of the total Ca2+-independent outward currents observed among CP neurons (see DISCUSSION).

Kinetic analyses of IA, ID, and IK

Time constants (tau s) of activation for IA, ID, and IK were determined from single exponential fits to the rising phases of the separated current components. For these analyses, IA and ID were obtained using the subtraction procedures described above and illustrated in Fig. 3, D and E; the current remaining in 5 mM 4-AP (Fig. 3C) was analyzed as IK. For all three components, the rising phases of the currents at each test potential were well described by single exponentials (Fig. 7B). Mean activation time constants for IA, ID, and IK are plotted as a function of test potential in Fig. 7A. As is evident, IA and ID activate rapidly, although at all test potentials, IA activates approximately twofold faster than ID. At +30 mV, for example, mean activation tau s are 0.83 ± 0.24 ms (n = 14) and 1.74 ± 0.49 ms (n = 18) for IA and ID, respectively. The observed differences between IA and ID, activation tau s are statistically significant at all test potentials (P < 0.002 at +60 mV; P < 0.001 at all other voltages). Analyses of the rising phases of IA isolated using the conditioning prepulse protocol (Fig. 4C) and of ID isolated using alpha -DTX (Fig. 5C) yielded results indistinguishable from those obtained for currents isolated using 4-AP. IA activation time constants at +60 mV, for example, are 0.61 ± 0.22 ms (n = 9) using the conditioning prepulse protocol and 0.55 ± 0.16 ms (n = 14) for the currents separated using 4-AP. Activation tau s of the alpha -DTX-sensitive currents (Fig. 5C) and the 50-µM 4-AP-sensitive currents (Fig. 3D) are also not significantly different. At +60 mV, for example, the 50-µM 4-AP- and 10-100 nM alpha -DTX-sensitive currents activate with time constants of 0.93 ± 0.33 ms (n = 18) and 1.29 ± 0.46 ms (n = 11), respectively, consistent with the suggestion that alpha -DTX and 50 µM 4-AP both block ID in CP neurons.


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FIG. 7. Kinetics of activation (A and B) and inactivation (C and D) of IA, ID, and IK are distinct. A and B: time constants (tau ) of activation for IA and ID weredetermined from single exponential fits to the rising phases of the currents in subtracted records, such as those in Fig. 3, D and E, respectively. For IK, activation tau were determined from fits to the rising phases of currents remaining in 5 mM 4-AP in records such as those in Fig. 3C. Typical fits for IA, ID, and IK are illustrated in B. Mean ± SD activation tau s for IA, ID, and IK are plotted as a function of test potential (note that the y-axis scale is different for the IK data). Continuous lines are best fits to the averaged data using tau (V) = a [exp(Vm/b)] c (see text). C: time constants (tau s) of inactivation were determined from exponential fits to the decay phases of currents recorded during 12-s voltage steps to 0, 10, 20, and 30 mV from a holding potential of -70 mV in control bath solution and in the presence of 8 mM 4-AP (see text). Mean ± SD inactivation tau s for IA (n = 7), ID(n = 7), and IK (n = 7) are plotted as a function of test potential. Typical fits are shown in D.

IK activates ~10-fold more slowly than IA or ID at all test potentials; the mean activation time constant for IK at +30 mV is 14.7 ± 4.0 ms (n = 19). Activation tau s of the TEA-sensitive currents (n = 8) also were determined from single exponential fits to the rising phases of subtracted current waveforms such as those in Fig. 6C. These analyses revealed that the time constants of activation of the TEA-sensitive currents and the currents remaining in the presence of 5 mM 4-AP are indistinguishable. The activation time constants of the 4-AP-insensitive currents (Fig. 3C) and the TEA-sensitive currents (Fig. 6C) at +60 mV, for example, are 8.2 ± 2.2 ms (n = 19) and 6.9 ± 2.8 ms (n = 8), respectively. For all three currents, activation time constants are voltage dependent, decreasing with increasing depolarization. The variations in tau  with respect to voltage are well described by single exponential functions of the form
τ(<IT>V</IT>) = <IT>a + b</IT>[exp(−<IT>V</IT><SUB>m</SUB>/<IT>c</IT>)]
where Vm is the test potential and c is a constant that defines the steepness of the voltage dependence. The best fits (continuous lines, Fig. 7A) to the experimental data yielded c = 32.1 (a = 0.38, b = 1.1) for IA; c = 38.4 (a = 0.2, b = 3.4) for ID, and c = 34.8 (a = 3.7, b = 25.5) for IK. Comparison of the values for c indicates that the activation rates of all three currents vary similarly as a function of voltage.

Time constants of inactivation for IA, ID, and IK were determined from exponential fits to the decay phases of current recorded during 12-s voltage steps to 0, 10, 20, and 30 mV in control bath solution and in the presence of 8 mM4-AP (to block IA and ID). The decay phases of the currents blocked by 8 mM 4-AP were well described by the sum of two exponentials (Fig. 7D) with mean time constants of19 ± 7 ms and 569 ± 143 ms (n = 7). Neither time constant varied as a function of voltage (Fig. 7C). The faster time constant is assumed to reflect the inactivation of IA and the slower time constant the inactivation of ID. To test this hypothesis, inactivation time constants for IA were determined from single exponential fits to the decay phases of currents such as those in Figs. 3E and 4C. Inactivation of IA at all test potentials was well described by a single exponential; mean inactivation time constants for IA separated using4-AP (Fig. 3E) and the conditioning prepulse (Fig. 4C) were found to equal 19 ± 5 ms (n = 14) and 17 ± 7 ms (n = 9), respectively. These values are not significantly different from the fast time constant obtained from the double-exponential fits to the 8 mM 4-AP-sensitive currents (19 ± 7 ms); this is consistent with the suggestion (above) that the fast component of the 8 mM 4-AP-sensitive currents reflects inactivation of IA.

The decay phases of the 4-AP-insensitive currents (IK) were well described in all cells and at all test potentials by the sum of two exponentials (Fig. 7D) with mean time constants of 475 ± 232 and 3,128 ± 1,328 ms (n = 7; Fig. 7C). In addition, ~20% of the current remaining in 8 mM 4-AP is noninactivating. Similar to the inactivation time constants for IA and ID, neither component of inactivation of the 8 mM 4-AP-insensitive current varied substantially with voltage (Fig. 7C). The relative amplitudes of the fast and slow components of IK inactivation, however, did vary appreciably among cells, ranging from approx 1 to approx 4. The finding of multiple components of inactivation of the 4-AP-insensitive currents suggests either that the properties of the underlying IK channels are complex or that the 4-AP-insensitive currents comprise distinct conductance pathways (see below and DISCUSSION).

Reversal potentials of IA, ID, and IK

Reversal potentials of IA, ID, and IK, were determined from analyses of tail currents of the individual K+ current components. For these analyses, IA and ID were obtained using the subtraction procedures illustrated in Fig. 3, and the plateau current remaining in the presence of 5 mM4-AP was analyzed as IK. From a holding potential of -70 mV, cells were depolarized to +30 mV for 6 ms (for IA and ID) or 150 ms (for IK) to activate the outward currents and subsequently repolarized to potentials between -30 and -120 mV. Tail current amplitudes at each potential were measured 1.5 ms after the onset of the hyperpolarizing voltage steps and plotted as a function of voltage. Reversal potentials were obtained by linear interpolation. Mean reversal potentials for IA, ID, and IK in 3 mM [K+]o were-73 ± 4 mV (n = 4), -82 ± 6 mV (n = 7), and -68 ± 4 mV (n = 7), respectively. The calculated equilibrium potential (EK) for K+ under the recording conditions employed here is -90 mV, suggesting that the channels underlying IA, ID, and IK are K+ selective. Because the experimentally determined reversal potentials, particularly for IA and IK, are positive to EK, however, the underlying channels also may have finite permeabilities for other ions.

Voltage dependences of activation and steady state inactivation of IA, ID, and IK

The voltage dependences of activation of IA, ID, and IK were determined from analyses of current records such as those in Fig. 3. IA and ID amplitudes were measured at the peak of subtracted current records such as those in Fig. 3, E (IA) and D (ID), and IK amplitudes were measured at the plateau (150 ms) of the current remaining in 5 mM 4-AP (Fig. 3C). IA, ID, and IK conductances at each test potential were calculated in each cell using the experimentally determined reversal potentials and subsequently normalized to their respective conductance values at +60 mV (measured in the same cell). Data for each cell were fitted to a Boltzmann relation of the form
<IT>G = G</IT><SUB>max</SUB>/[1 + exp{(<IT>V − V</IT><SUB>1/2</SUB>)/<IT>k</IT>}]
where G is the conductance at each potential, V; Gmax is the maximal conductance; V1/2 is the voltage of half-maximal activation; and k is the slope factor. The V1/2 and k values from individual cells then were averaged. Mean normalized conductances for IA (n = 14), ID (n = 18), and IK (n = 29) are plotted as a function of test potential in Fig. 8A; the solid lines reflect the best Boltzmann fits to the averaged data. MeanV1/2 (and k) values for IA, ID, and IK are 6 ± 6 mV (k =16 ± 1 mV), 12 ± 8 mV (k = 11 ± 1 mV), and 12 ± 5 mV (k = 14 ± 2 mV), respectively. The activation threshold of IA (approximately -50 mV) is slightly more hyperpolarized than that of either ID or IK; both ID and IK begin to activate around -40 mV. In addition, although all three currents activate over similar voltage ranges, the V1/2 of IA (6 ± 6 mV) is significantly more hyperpolarized than the V1/2 values of either ID (12 ± 8 mV; P < 0.05) or IK (12 ± 4 mV; P < 0.002).


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FIG. 8. IA, ID, and IK display similar voltage dependences of activation (A) and steady state inactivation (B). To examine the voltage dependences of current activation, IA, ID, and IK conductances at each test potential in individual cells were calculated from records such as those in Fig. 3,C-E, using the experimentally determined reversal potentials. Conductances at each test potential were normalized to their respective conductance values at +60 mV (in the same cell). Mean normalized conductances are plotted as a function of test potential and the Boltzmann fits to the averaged data are shown (------). B: to examine the voltage dependences of steady state inactivation, IA, ID, and IK amplitudes, evoked at +30 mV after 15- to 20-s conditioning prepulses to potentials between -90 and +20 mV were measured; inset: protocol is illustrated. Using the experimentally determined reversal potentials, conductances then were calculated and normalized to their respective conductance values after the -90 mV conditioning prepulse. Mean normalized IA (n = 9), ID (n = 5), and IK (n = 5) conductances are plotted as a function of conditioning potential, and the Boltzmann fits to the averaged data are shown (------).

The voltage dependences of steady state inactivation of IA, ID, and IK were examined during voltage steps to +30 mV presented after 15- to 20-s conditioning prepulses to potentials between -90 and +10 mV (protocol shown in Fig. 8B, inset). Protocols were repeated under control conditions and in the presence of 4-AP or DTX. Subtracted current records (such as those described above and illustrated in Figs. 3 and 5) were obtained to isolate IA and ID. Current amplitudes recorded during the +30-mV steps were measured at the peak of subtracted records for IA (n = 9) and ID (n = 5) and at the plateau (150 ms) of the current remaining in 5 mM 4-AP for IK (n = 5). IA, ID, and IK conductances then were calculated for each conditioning potential and normalized to their respective conductance values after the conditioning step to -90 mV. Mean normalized IA, ID, and IK conductances are plotted with respect to conditioning potential in Fig. 8B; the continuous lines represent the best Boltzmann fits to the averaged data. Mean V1/2 (and k) values for IA, ID, and IK are -38 ± 3 mV (k = -8 ± 2 mV), -21 ± 4 mV (k = -9 ± 3 mV), and -26 ± 2 mV (k = -13 ± 4 mV), respectively. All three currents are sensitive to steady state inactivation over similar voltage ranges, although the V1/2 of IA (-38 ± 3 mV) is significantly (P < 0.005) more hyperpolarized than the V1/2 values of either ID or IK. In addition, these experiments revealed that ~20% of the plateau current (IK) does not inactivate. The finding that steady state inactivation of the remaining4-AP-insensitive current (i.e., the inactivating current) is well described by a single Boltzmann, suggests that the two inactivation time constants (475 and 3,128 ms; Fig. 8C) reflect inactivation of the same population of IK channels.

Recovery from steady state inactivation of IA, ID, and IK

To examine the time dependences of recovery from steady state inactivation of IA, ID, and IK, cells first were depolarized to 0 mV for 12 s to inactivate the currents (longer depolarizations did not lead to further inactivation), subsequently hyperpolarized to -70 mV for varying times ranging from 0 to 5,000 ms to allow recovery, and, finally, stepped to +30 mV for 160 ms to activate the currents; the protocol is displayed below the current records in Fig. 9A. Peak (3 ms) and plateau (150 ms) control current amplitudes after each recovery time were measured, normalized to their respective current amplitudes after the 5-s recovery, and plotted as a function of recovery time (Fig. 9B). The normalized recovery data for both the peak and plateau currents were well fitted by the sum of two exponentials (Fig. 9B, solid lines). For this cell, recovery taus for the peak current were 36 and 738 ms; recovery time constants for the plateau current were 73 and 618 ms. Similar results were obtained in eight cells; mean recovery time constants were 38 ± 7 and 670 ± 202 ms for the peak current and 79 ± 26 and 734 ± 203 ms for the plateau current.


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FIG. 9. Rates of recovery of IA, ID, and IK are distinct. After inactivating the currents by a 12-s prepulse to 0 mV, the cell was hyperpolarized to -70 mV for times ranging from 0 to 5,000 ms before a test depolarization to +30 mV. Typical current waveforms recorded during the +30 mV depolarization after variable (0-5,000 ms) recovery periods are displayed in A; protocol is beneath the records. Note that the current recorded after a 10 ms recovery time at -70 mV (bullet ) activates and inactivates rapidly (see text). B: amplitudes of the peak and plateau currents evoked at +30 mV after each recovery period were measured, normalized to their respective (peak and plateau; black-square) amplitudes after a 5,000-ms recovery period and plotted as a function of time. Recovery of the peak and plateau currents was best described by the sum of 2 exponentials (------). Inset: initial phase of recovery of both the peak and the plateau currents is shown on an expanded time scale. Similar results were obtained on 7 cells.

The slower time constant for recovery of the peak (670 ± 202 ms) and plateau (734 ± 203 ms) currents are not significantly different (Student's t-test), suggesting that both time constants reflect recovery of the same current component. Because ID is the only one of the three voltage-gated K+ currents in CP cells that contributes to both the peak and plateau currents, it seemed quite likely that the slower time constants of recovery of both the peak and the plateau currents reflect recovery of ID. To test this hypothesis, recovery of ID was examined directly. The protocol outlined above was completed first in control bath solution and subsequently in the presence of 50 µM 4-AP. Analyses of the rates of recovery of the subtracted currents (ID) revealed that recovery was well described by a single exponential in all cells examined. The mean recovery time constant for ID was found to equal 588 ± 274 ms (n = 8), a value not significantly different (ANOVA) from the slower component of recovery of the peak (670 ± 202 ms) and plateau (734 ± 203 ms) components of the control currents. These results confirm the hypothesis (above) that the slow time constants of recovery of the peak and plateau currents reflect recovery of the same population of (ID) channels. Because only ID and IK contribute to the plateau currents, the fast component of recovery of the plateau current (mean tau of 79 ± 26 ms) must reflect recovery of IK. The fast time constant for recovery of the peak (38 ± 7 ms), therefore, reflects IA recovery. The waveforms of the currents recorded after brief recovery times are consistent with this assertion. As illustrated in Fig. 9A, the current recorded after 10 ms recovery (arrow) activates and inactivates rapidly and is indistinguishable from IA isolated using 4-AP or the conditioning prepulse protocol (see Figs. 3E and 4C). Similar results were obtained in experiments completed on six other cells. The mean activation and inactivation time constants for the currents recorded (at +30 mV) after 10-ms recovery at -70 mV were determined to be 0.76 ± 0.20 ms and 21.6 ± 12 ms, respectively. For comparison, the activation and inactivation time constants (at +30 mV) for IA isolated using 4-AP were 0.83 ± 0.24 ms and 19.2 ± 3.8 ms, respectively. These values are not significantly different; this is consistent with the interpretation that the fast component of recovery of the peak current reflects IA.

    DISCUSSION
Abstract
Introduction
Methods
Results
Discussion
References

The results presented here demonstrate the presence of three kinetically and pharmacologically distinct Ca2+-independent, voltage-gated K+ currents in isolated, identified postnatal day 7-16 CP visual cortical neurons. These cells were identified after in vivo retrograde labeling at postnatal day 5 (MATERIALS AND METHODS). Because many more cells in immature, than in mature, rat visual cortex make projections across the corpus callosum (Olavarria and Van Sluyters 1985; J. M. Nerbonne and A. Burkhalter, unpublished observations), some of the cells studied here are not destined to be CP cells in the adult. It is important to emphasize, however, that CP cells were selected for the present studies to represent "regular-spiking" cortical neurons rather than because they project across the callosum. Because neurons that retract their callosal projections during postnatal development do not die or undergo changes in morphology or transmitter phenotype (Olavarria and VanSluyters 1985; J. M. Nerbonne and A. Burkhalter, unpublished observations), both the cells that retain and the cells that retract their callosal projections were expected to display regular-spiking behavior. As demonstrated in the subsequent paper (Locke and Nerbonne 1997), current-clamp recordings have revealed that this hypothesis is correct, i.e., all of the cells labeled in early postnatal animals based on the callosal projection are regular-spiking cortical neurons.

The Ca2+-independent, voltage-gated K+ currents characterized here in CP neurons are referred to as IA, ID, and IK because their properties are similar, although not identical, to depolarization-activated K+ currents termed IA, ID (or slow transient), and IK in other mammalian neurons (Rudy 1988; Storm 1990). We emphasize that the properties of IA, ID, and IK vary in different cell types (see below for further discussion) and that these names were selected to emphasize qualitative similarities. Although IA, ID, and IK were identified readily in all CP cells studied (n = 40), the absolute amplitudes and densities of the individual current components varied markedly among cells. Mean IA, IK, and ID densities determined in the experiments here were 215 ± 83, 173 ± 94, and 54 ± 31 pA/pF, respectively. On average, therefore, the densities of both IA and IK are approximately four times the density of ID. In addition, the relative IA:ID:IK density varies among cells; this variability underlies the observed differences in the waveforms of the total Ca2+-independent outward currents among CP neurons (Fig. 1). It is important to note that, because the experiments here were performed on postnatal (day 7-16) CP neurons, it is certainly possible that the densities (and/or the detailed properties) of IA, ID, and IK in adult CP cells are different. Further experiments will be necessary to explore this possibility directly.

Functionally, IA and ID underlie the peak outward currents in CP cells (Fig. 1); because activation is slow, IK does not contribute appreciably to the peak. IK, however, together with ID, determines the plateau current amplitudes in CP cells (Fig. 1); IA inactivates rapidly and, therefore, does not also contribute to the plateau currents. It is important to note that, because the goal of the experiments here was to define the detailed time- and voltage-dependent properties of the K+ currents expressed in CP neurons, recordings were obtained only from neurons without extensive processes (to maintain adequate spatial control of the membrane voltage), and analyses were completed only on those cells that behaved as a single electrical compartment (i.e., cells withcapacitative transients well described by single exponentials). The currents separated and characterized here, therefore, are assumed to reflect only the activation of voltage-gated K+ channels expressed in CP cell soma. Depolarization-activated K+ channels expressed predominantly or exclusively in dendrites are likely to be overlooked. It is certainly possible, therefore, that novel channel types are expressed in CP cell dendrites or that the relative densities of the K+ channels in dendrites are different from those in cell bodies. Alternative experimental approaches will be needed to explore these possibilities directly.

Comparison of IA, ID, and IK in CP neurons to currents in other mammalian neurons

Rapidly activating and inactivating currents, similar to IA in CP neurons, have been characterized in neurons from rat hippocampus (Ficker and Heinemann 1992; Segal and Barker 1984; Storm 1988), dentate gyrus (Beck et al. 1992), neostriatum (Surmeier et al. 1989), lateral geniculate nucleus (Budde et al. 1992), thalamus (Huguenard et al. 1991), sensorimotor cortex (Foehring and Surmeier 1993; Hamill et al. 1991), and visual cortex (Albert and Nerbonne 1995; Foehring and Surmeier 1993); the guinea pig (Numann et al. 1987) and embryonic mouse (Wu and Barish 1992) hippocampus; and cat sensorimotor cortex (Spain et al. 1991). The time- and voltage-dependent properties and the pharmacological sensitivities of the currents termed IA in these different preparations are similar. In general, for example, IA activates rapidly at all test potentials, and activation is voltage dependent. Inactivation, although also rapid, is voltage independent (but see Beck et al. 1992; Budde et al. 1992; Spain et al. 1991), and IA is blocked by millimolar concentrations of 4-AP (but see Spain et al. 1991). In most cells, the threshold for IA activation lies between -65 and -50 mV, and steady state inactivation of IA is characterized by a V1/2 between -60 and -80 mV. In CP neurons, however, the activation threshold (approximately -50 mV) and the V1/2 values (approximately -40 mV) for steady state inactivation of IA are somewhat more depolarized than the values reported for many other cells. Similar voltage-dependent properties, however, have been described previously for IA in rat cortical (Albert and Nerbonne 1995; Hamill et al. 1991) and neostriatal (Surmeier et al. 1989) neurons. The differences in the voltage-dependent properties of IA in different cell types may reflect molecular heterogeneity in IA channels or cell type-specific posttranslational regulation of IA channel properties. Alternative experimental strategies will be needed to distinguish between these (and other) hypotheses.

Slow transient currents, which are distinct from IA and IK and often are referred to as ID, have been identified in neurons from rat prefrontal cortex (Hammond and Crepel 1992), neostriatum (Surmeier et al. 1991), hippocampus (Ficker and Heinemann 1992), visual cortex (Albert and Nerbonne 1995; Foehring and Surmeier 1993), globus pallidus (Stefani et al. 1995) and dorsal root ganglia (DRG) (Stansfeld et al. 1986) and from rat sensorimotor cortex (Foehring and Surmeier 1993), guinea pig lateral geniculate nucleus (McCormick 1991) and DRG (Penner et al. 1986), and mouse hippocampus (Wu and Barish 1992). In general, slow transient currents activate fast on membrane depolarization, inactivate slowly and are blocked by micromolar concentrations of 4-AP and often by DTX (Foehring and Surmeier 1993; Penner et al. 1986; Stansfeld et al. 1986; Surmeier et al. 1991). Inactivation is voltage independent, and best described by the sum of two exponentials with time constants of 100-500 ms and 2-7 s (Ficker and Heinemann 1992; McCormick 1991; Wu and Barish 1992). The V1/2 of steady state inactivation of slow transient currents usually occurs between -95 and -60 mV; ID channel availability, therefore, can be influenced by changes in the membrane potential around rest. In the experiments here, however, ID in P7-P16 CP neurons is sensitive to steady state inactivation only at potentials positive to approximately -60 mV. It is certainly possible that the voltage-dependent properties of ID vary during postnatal development and that the V1/2 for steady state inactivation of ID is more hyperpolarized in adult CP neurons. Further experiments will be necessary to test this hypothesis directly.

In some preparations, slow transient currents begin to activate between -75 and -60 mV, i.e., at potentials hyperpolarized to the action potential threshold (McCormick 1991; Storm 1988; Surmeier et al. 1991) and, as described originally in hippocampal neurons (Storm 1988), the activation of ID in these cells controls the "delay" to firing in response to threshold current injections. In other preparations, however, slow transient currents begin to activate at more depolarized potentials: ID in neonatal rat SCP neurons (Albert and Nerbonne 1995) and embryonic mouse hippocampal neurons (Wu and Barish 1992), Itslow in embryonic rat hippocampal neurons (Ficker and Heinemann 1992), Ikm in rat LGN neurons (Budde et al. 1992) for example, begin to activate around -35 mV, similar to ID in CP neurons (threshold approximately -40 mV; Fig. 8A). The depolarized activation thresholds of these currents suggest that they may contribute less to controlling delay to firing an action potential in response to depolarizing inputs than IA. One factor underlying the differences in activation thresholds in different studies may be the age of the animals from which the cells were obtained: currents with depolarized activation thresholds were recorded from embryonic or neonatal neurons, whereas the currents with hyperpolarized activation thresholds were recorded from adult neurons. It is possible, therefore, that the activation threshold of ID shifts toward more hyperpolarized potentials during development and that the functional role of ID in adult CP (and other) neurons includes controlling the delay to firing in response to threshold depolarizing inputs. Further experiments aimed at examining the properties and densities of ID in CP (and other) neurons isolated at later stages of development will be necessary to test this hypothesis directly.

The slowly activating, TEA-sensitive K+ current, which we have termed IK, in CP neurons closely resembles delayed rectifier type-currents described in neurons from rat thalamus (Huguenard and Prince 1991), sensorimotor cortex (Foehring and Surmeier 1993; Hamill et al. 1991), dentate gyrus (Beck et al. 1992), lateral geniculate nucleus (LGN) (Budde et al. 1992), hippocampus (Segal and Barker 1984), globus pallidus (Stefani et al. 1995), and visual cortex (Albert and Nerbonne 1995; Foehring and Surmeier 1993) and from guinea pig hippocampus (Numann et al. 1987). As in CP neurons, the threshold of IK activation in these cells is between -45 and -35 mV, i.e., positive to that of IA described in the same cells. Activation (tau ~10-30 ms at 0 mV) and inactivation (tau more than ~1 s) of IK are slow, and, in some preparations, inactivation is best described by two exponentials (Beck et al. 1992; Huguenard and Prince 1991) or is incomplete even during prolonged depolarizing voltage steps (Beck et al. 1992; Foehring and Surmeier 1993; Hamill et al. 1991; Huguenard and Prince 1991; Numann et al. 1987). In CP neurons, inactivation of IK is also slow and biexponential (taus = 475 and 3,128 ms) and, in addition, ~20% of IK is noninactivating. The finding that the voltage dependences of the inactivating components of the 4-AP-insensitive currents are the same (Fig. 9B) suggests that the biexponential kinetics reflects the gating properties of the underlying IK channels. In many preparations, the rates of recovery of IK from steady state inactivation are substantially slower than for IA or ID (in the same cells) and generally proceed during a time course of 100-500 ms (but see Hamill et al. 1991; Huguenard et al. 1991). In CP neurons, however, IK recovers from inactivation somewhat more rapidly (79 ± 26 ms at -70 mV) and is similar therefore to IK in rat LGN neurons (116 ms at -90 mV) (Budde et al. 1992). The voltage dependence of steady state inactivation of IK varies widely among cell types with V1/2 values ranging from -87 mV in guinea pig hippocampal cells (Numann et al. 1987) to -49 mV in rat LGN neurons (Budde et al. 1992). Additional experiments will be necessary to determine whether the differences in the time- and voltage-dependent properties of the delayed rectifier currents in various preparations reflect the expression of distinct types of IK channels or cell type-specific regulation of the properties of a single type of IK channel.

    ACKNOWLEDGEMENTS

  We thank J. Doyle, R. J. Martinez, and J. M. Coates for expert technical assistance with the in vivo retrograde labeling and with the preparation and the maintenance of cortical glial cultures. In addition, we thank Drs. Andreas Burkhalter, Joel Solomon, and Jennifer L. Albert for many helpful discussions.

  Finally, we acknowledge the financial support provided by the National Institute of Neurological Disorders and Stroke Grant NS-30676.

    FOOTNOTES

  Address for reprint requests: J. M. Nerbonne, Dept. of Molecular Biology and Pharmacology, Washington University School of Medicine, 660 South Euclid Ave., Box 8103, St. Louis, MO 63110.

  Received 3 September 1996; accepted in final form 30 June 1997.

    REFERENCES
Abstract
Introduction
Methods
Results
Discussion
References

0022-3077/97 $5.00 Copyright ©1997 The American Physiological Society