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Original Article

A Whole-Cell and Single-Channel Study

Jacopo Magistretti^a,b and Angel Alonso^a

^a From the Department of Neurology and Neurosurgery, McGill University and Montreal Neurological Institute, Montréal, Québec, H3A 2B4 Canada
^b Dipartimento di Neurofisiologia Sperimentale, Istituto Nazionale Neurologico "Carlo Besta", 20133 Milano, Italy
Department of Neurology and Neurosurgery, Montreal Neurological Institute, McGill University, 3801 University Street, Montréal, Québec, H3A 2B4 Canada.Fax: (514) 398-8106;

mdao{at}musica.mcgill.ca

	ABSTRACT

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The functional and biophysical properties of a sustained, or "persistent," Na⁺ current (I_NaP) responsible for the generation of subthreshold oscillatory activity in entorhinal cortex layer-II principal neurons (the "stellate cells") were investigated with whole-cell, patch-clamp experiments. Both acutely dissociated cells and slices derived from adult rat entorhinal cortex were used. I_NaP , activated by either slow voltage ramps or long-lasting depolarizing pulses, was prominent in both isolated and, especially, in situ neurons. The analysis of the gating properties of the transient Na⁺ current (I_NaT) in the same neurons revealed that the resulting time-independent "window" current (I_NaTW) had both amplitude and voltage dependence not compatible with those of the observed I_NaP , thus implying the existence of an alternative mechanism of persistent Na⁺-current generation. The tetrodotoxin-sensitive Na⁺ currents evoked by slow voltage ramps decreased in amplitude with decreasing ramp slopes, thus suggesting that a time-dependent inactivation was taking place during ramp depolarizations. When ramps were preceded by increasingly positive, long-lasting voltage prepulses, I_NaP was progressively, and eventually completely, inactivated. The V_1/2 of I_NaP steady state inactivation was approximately –49 mV. The time dependence of the development of the inactivation was also studied by varying the duration of the inactivating prepulse: time constants ranging from 6.8 to 2.6 s, depending on the voltage level, were revealed. Moreover, the activation and inactivation properties of I_NaP were such as to generate, within a relatively broad membrane-voltage range, a really persistent window current (I_NaPW). Significantly, I_NaPW was maximal at about the same voltage level at which subthreshold oscillations are expressed by the stellate cells. Indeed, at –50 mV, the I_NaPW was shown to contribute to >80% of the persistent Na⁺ current that sustains the subthreshold oscillations, whereas only the remaining part can be attributed to a classical Hodgkin-Huxley I_NaTW. Finally, the single-channel bases of I_NaP slow inactivation and I_NaPW generation were investigated in cell-attached experiments. Both phenomena were found to be underlain by repetitive, relatively prolonged late channel openings that appeared to undergo inactivation in a nearly irreversible manner at high depolarization levels (–10 mV), but not at more negative potentials (–40 mV).

Key Words: persistent Na⁺ current • window current • stellate cells • oscillations • patch clamp

	INTRODUCTION

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The so-called persistent sodium current (I_NaP)¹ is known to be expressed, together with the classical, transient sodium current (I_NaT), by numerous mammalian neuronal types. Its basic features include persistence during prolonged depolarizations, lower threshold of activation than I_NaT, and low amplitude (the underlying conductance normally representing 0.2–2% of the total sodium conductance; French and Gage 1985; French et al. 1990; for review see Taylor 1993; Crill 1996. After the initial descriptions of the actions of sustained Na⁺ currents on neuronal electroresponsiveness (Hotson et al. 1979; Llinás and Sugimori 1980; Connors et al. 1982), I_NaP was first demonstrated with voltage-clamp studies in neocortical neurons (Stafstrom et al. 1982, Stafstrom et al. 1985), and biophysically characterized in the hippocampus (French and Gage 1985; French et al. 1990). I_NaP has also been observed, directly or indirectly, in an increasing number of neuronal structures including basal ganglia (Chao and Alzheimer 1995; Cepeda et al. 1995), amygdala (Pape and Driesang 1998), thalamus (Jahnsen and Llinás 1984; Parri and Crunelli 1998), hypothalamus (Llinás and Alonso 1992; Uteshev et al. 1995), cerebellum (Jahnsen 1986; D'Angelo et al. 1998; Kay et al. 1998), and peripheral ganglia (Baker and Bostock 1997). Due to its voltage-dependent properties, I_NaP can contribute to important integrative functions such as amplification of excitatory postsynaptic potentials (Deisz et al. 1991; Stuart and Sakmann 1995; Schwindt and Crill 1995; Lipowsky et al. 1996), generation of pacemaker activity (Alonso and Llinás 1989; Amitai 1994; Pennartz et al. 1997; Pape and Driesang 1998), and firing-pattern shaping (Jahnsen and Llinás 1984; Klink and Alonso 1993; Franceschetti et al. 1995; Parri and Crunelli 1998). Since I_NaP can generate membrane bistability and plateau potentials, it has also been implicated in the pathogenesis of some forms of epilepsy (Segal 1994; for review see Ragsdale and Avoli 1998). In addition, the ability of I_NaP to sustain long-lasting Na⁺ influxes, and therefore to steadily increase intracellular Na⁺ concentration, has raised interest as to its possible role in mechanisms of neurodegeneration (Taylor and Meldrum 1995).

The entorhinal cortex (EC) has proven to be a particularly interesting neuronal system for the study of I_NaP functions. In the stellate cells of EC layer II, which give rise to the main cortical projection to the hippocampus (Steward and Scoville 1976), I_NaP has been shown to critically participate in the generation of subthreshold membrane-potential oscillations in the theta-rhythm range (Alonso and Llinás 1989; Klink and Alonso 1993). This particular intrinsic subthreshold activity is considered to be a critical determinant for the generation of the population theta oscillations generated by the EC network (Adey et al. 1957, Adey et al. 1960; Holmes and Adey 1960; Mitchell and Ranck 1980; Alonso and García-Austt 1987a,Alonso and García-Austt 1987b). The theta rhythm has been shown to contribute to synaptic-plasticity processes (Larson and Lynch 1986; Larson et al. 1986; Greenstein et al. 1988; Alonso et al. 1990; Huerta and Lisman 1996; Hölscher et al. 1997), and it is thus believed to play a major role in temporal lobe learning and memory functions (Doyère and Laroche 1992; Buzsáki 1996). On the other hand, since the EC is also known to play a crucial role in temporal lobe epileptogenesis (see references in Dickson and Alonso 1997), it has been hypothesized that the presence of a robust I_NaP in EC neurons may contribute to epileptogenic processes (Klink and Alonso 1993).

For the above considerations, a detailed knowledge of the biophysical properties of I_NaP expressed by EC layer II neurons seems of great interest since it would help to understand how this current specifically influences the physiological behavior of these cells. In this study, we have characterized I_NaP in both acutely isolated and in situ EC layer II neurons. Our results revealed the existence of some interesting biophysical properties of I_NaP that had not been thoroughly investigated yet, including: (a) a slow voltage- and time-dependent inactivation occurring with voltage-dependent time constants in the order of seconds; (b) a full inactivation at sufficiently positive potentials; and (c) a truly persistent, or "window," current arising from the particular activation and steady state inactivation properties of the corresponding conductance. Moreover, we describe the single-channel events that account for all of the above-mentioned macroscopic phenomena.

	MATERIALS AND METHODS

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Slice and Cell Preparation
Young-adult Long-Evans rats (P25–P35) were killed by decapitation. The brain was quickly removed under hypothermic conditions, blocked on the stage of a vibratome (Pelco), and submerged in an ice-cold cutting solution containing (mmol/liter): 115 NaCl, 5 KCl, 4 MgCl₂, 1 CaCl₂, 20 PIPES, and 25 D-glucose, pH 7.4 with NaOH, bubbled with pure O₂. Osmotic pressure () of the latter solution, as measured with a Micro Osmette 5004 osmometer, was typically 320 mOsm. Horizontal slices of the retrohippocampal region were cut at 350–400 µm. For in situ recordings, slices were stored at room temperature in the above solution until use. For recordings on isolated neurons, the layer II of medial entorhinal cortex was dissected from each slice (White et al. 1993). Neurons were acutely isolated from the tissue fragments thus obtained following an enzymatic and mechanical dissociation procedure described elsewhere (Magistretti and de Curtis 1998).

Whole-Cell Recordings on Slices
The recording chamber was mounted on the stage of an upright microscope (see below). Slices were laid onto the bottom of the chamber and perfused with an extracellular solution containing (mmol/liter): 34 NaCl, 26 NaHCO₃, 80 tetraethylammonium (TEA)-Cl, 5 KCl, 3 CsCl, 2 CaCl₂, 3 MgCl₂, 2 BaCl₂, 2 CoCl₂, 0.4 CdCl₂, 4 4-aminopyridine (4-AP), 10 glucose, pH 7.4 when bubbled with 95% O₂, 5% CO₂ ( 320 mOsm). The association of Co²⁺ and Cd²⁺ was found to depress residual inward rectification insensitive to the application of tetrodotoxin (TTx; Sigma-Aldrich Canada Ltd.) more effectively than Cd²⁺ alone in slices (but not in freshly dissociated neurons; see below). Preliminary experiments were performed in in situ neurons in the absence of extracellular Co²⁺ and Ba²⁺ and in the presence of 0.2 rather than 0.4 mM Cd²⁺. Both average peak amplitude and current density of ramp (50 mV/s)-evoked, TTx-subtracted I_NaPs recorded under the former and latter ionic conditions (–179.3 ± 120.7 vs. –195.8 ± 100.0 pA, respectively, and –17.7 ± 29.1 vs. –12.9 ± 10.6 pA/pF, respectively, n = 54 and 8, respectively) were not significantly different (P = 0.68 and 0.63, respectively). Hence, possible enhancing effects of nonphysiological extracellular divalent cations on I_NaP amplitude (Cummins et al. 1998) did not significantly affect measures in our experimental conditions. Patch pipettes were fabricated from thick-wall borosilicate glass capillaries by means of a P-97 horizontal puller (Sutter Instruments Co.). The intrapipette solution contained (mmol/liter): 110 CsF, 10 HEPES-Na, 11 EGTA, 2 MgCl₂, pH 7.25 with CsOH ( adjusted to 290 mOsm with mannitol). When filled with the above solution, the patch pipettes had a resistance of 3–5 M. Slices were observed with an Axioskop microscope (Carl Zeiss, Inc.) equipped with a 40x water-immersion objective lens and differential-contrast optics. A near-infrared charge-coupled device camera (XC-75; Sony Corp.) was also connected to the microscope, and used to improve cell visualization for identification of neuron types and during the approaching and patching procedures. With this equipment, the principal cells of EC layer II were easily distinguished based on their somato-dendritic shape (stellate cells: Ramón y Cajal 1902), size, and position (Klink and Alonso 1997). Patch pipettes were brought in close proximity to the selected neurons while manually applying positive pressure inside the pipette. Tight seals (>100 G) and the whole-cell configuration were obtained by suction (Hamill et al. 1981). Series resistance (R_s) was always compensated by 55% with the amplifier's built-in compensation section. R_s, as estimated off-line from the peak amplitude of averaged capacitive transients evoked by –5-mV voltage square pulses (with the low-pass filter set at 10 kHz), was on average 8.5 ± 2.1 M (n = 54). Cell capacitance was evaluated online by canceling the fast component of whole-cell capacitive transients evoked by –10-mV voltage steps with the amplifier compensation section, and reading out the corresponding value. Voltage-clamp recordings were performed at room temperature (22°C) using an Axopatch 1D amplifier (Axon Instruments). The general holding potential was –80 mV.

Whole-Cell Recordings on Isolated Neurons
The recording chamber was mounted on the stage of an inverted microscope (see below). After seeding into the chamber, dissociated cells were perfused with a standard HEPES buffer containing (mmol/liter): 140 NaCl, 5 KCl, 10 HEPES (free acid), 2 CaCl₂, 2 MgCl₂, 25 glucose, pH 7.4 with NaOH, bubbled with pure O₂ ( 320 mOsm). After wash-out of cell debris, cell perfusion was switched to a solution suitable for Na⁺-current isolation containing (mmol/liter): 100 NaCl, 40 TEA-Cl, 10 HEPES (free acid), 2 CaCl₂, 3 MgCl₂, 0.2 CdCl₂, 5 4-AP, 25 glucose, pH 7.4 with NaOH, bubbled with pure O₂ ( 318 mOsm). The intrapipette solution was the same as described in the previous paragraph. Cells were observed at 400x with an Axiovert 100 microscope (Carl Zeiss, Inc.). After tight-seal formation (>100 G) and the establishment of the whole-cell configuration, series resistance was on average 12.0 ± 4.5 M (n = 38), and was always compensated by 70%. The remaining procedures and experimental conditions were the same as described in the previous paragraph.

Single-Channel Recordings
Single-channel, cell-attached experiments were performed in acutely isolated neurons. After seeding into the recording chamber, cells where initially perfused with the same solution as described in the previous paragraph. The pipette solution contained (mmol/liter): 130 NaCl, 35 TEA-Cl, 10 HEPES-Na, 2 CaCl₂, 2 MgCl₂, 5 4-AP, pH 7.4 with HCl ( 338 mOsm). Single-channel patch pipettes had resistances ranging from 10 to 35 M when filled with the above solution, and were always coated with Sylgard^® (Dow Corning Corp.) from the shoulder to a point as close as possible to the tip so as to minimize stray pipette capacitance. After obtaining the cell-attached configuration, the extracellular perfusion was switched to a high-potassium solution containing: 140 K-acetate, 5 NaCl, 10 HEPES (free acid), 4 MgCl₂, 0.2 CdCl₂, 25 glucose, pH 7.4 with KOH ( 320 mOsm) so as to hold the neuron resting membrane potential at or near 0 mV. Recordings were performed at room temperature with an Axopatch 200B amplifier (Axon Instruments). Capacitive transients and linear current leakage were minimized online by acting on the respective built-in compensation sections of the amplifier. Long-duration (20-s) depolarizing voltage steps were delivered one every 40 s from a holding potential of –80 or –100 mV.

Data Acquisition
Voltage protocols were commanded and current signals were acquired with a Pentium PC interfaced to an Axon TL1 interface, using the Clampex program of the pClamp 6.0.2 software (Axon Instruments). Current signals were filtered online (using the amplifier's built-in low pass filter) and digitized at different frequencies according to the specific experimental aim. Filtering and acquisition frequencies were 5 and 20 kHz, respectively, for I_NaT recordings; 0.1–1 and 0.67–10 kHz (depending on the protocol duration), respectively, for I_NaP recordings; 1 and 2 kHz, respectively, for single-channel recordings. In all of the voltage protocols applied, cell-membrane potential was kept at the holding level for 15 (in whole-cell experiments) or 20 s (in single-channel experiments) between the end of each sweep and the beginning of the subsequent sweep (or of the conditioning prepulse preceding it, when applied). This avoided the development of cumulative voltage-dependent inactivation of I_NaP during consecutive acquisition cycles.

Data Analysis
Whole-cell recordings were analyzed by means of the Clampfit program (Axon Instruments). Offline leak subtraction was performed on I_NaT - (but not I_NaP-) protocol traces. Current density was calculated by dividing the peak current amplitude by cell capacitance, estimated as explained above. Conductance values were calculated from Na⁺-current amplitudes by applying the extended Ohm's law in the form: G_Na = I_Na/(V – V_Na), where V_Na is the nominal (Nernst) Na⁺ reversal potential. Data were fitted with exponential functions, I = (A_i · exp(–t/_i) + C, using Clampfit, or with Boltzmann functions, G = G_max/{1 + exp[(V – V_1/2)/k)]}, using Origin 3.06 (MicroCal Software).

Single-channel recordings were analyzed using Clampfit, Fetchan, and pStat (Axon Instruments). Residual capacitive transients were nullified by offline subtracting fits of average blank traces. Residual leakage currents were carefully measured in every single sweep at trace stretches devoid of any channel openings, and digitally subtracted. Channel dwell times were determined using a standard threshold routine of the Fetchan program. Ensemble-average traces were fitted with single exponential functions, I = A · exp(–t/) + C, using Clampfit. Dwell-time histograms were fitted with double exponential functions, N = A₁ · exp(–t/₁) + A₂ · exp(–t/₂), using pStat.

Average values were expressed as mean ± SD, unless otherwise explicitly stated. Statistical significance was evaluated by means of the two-tail Student's t test for unpaired data.

Modeling I_NaP
For a phenomenological description of I_NaP activation and slow inactivation, a simple Hodgkin-Huxley model was assumed. We applied the basic relationship:

(1)

where

(2)

and m and h are the probabilities of the activating and inactivation particles, respectively, to be in the permissive position. I_NaP activation was assumed to be instantaneous, and m(V) was derived directly from the G_NaP activation curve. h(V) was derived directly from the G_NaP steady state inactivation curve. The transitions of the inactivating particle, h, were modeled according to the following first-order kinetic scheme:

from which it follows:

(3)

where

(4)

and

(5)

Numerical values for the rate constants, and β, were derived from the experimental values of time constants of inactivation and recovery from inactivation (_h) and from the h curve by applying and . After obtaining the analytical functions describing the voltage dependence of the rate constants (see RESULTS), the time course of I_NaPs activated in response to various voltage protocols was numerically reconstructed on the basis of the and , and in its differential form. The simulation programs were compiled using QuickBASIC 4.5 (Microsoft Corp.).

	RESULTS

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General Properties of I_NaP in Acutely Dissociated and In Situ Neurons
In acutely dissociated EC layer II principal neurons, a prominent I_NaP could be evoked by delivering long-lasting depolarizing pulses. Fig. 1 A illustrates current traces from a representative neuron. I_NaP was blocked by 1 µM TTx, and was therefore routinely isolated via TTx subtraction. I_NaP threshold of activation was at about –65 mV, and peak at –30 mV. Average I_NaP absolute amplitude (derived by averaging the data points between 400 and 500 ms from the pulse onset) and current density (calculated as explained in MATERIALS AND METHODS) were –96.5 ± 61.5 pA and –12.0 ± 7.0 pA/pF, respectively, at the peak of the current-voltage (I–V) relationship (n = 5). The average I_NaP I–V relationship showed a linear region from –30 to –5 mV, the linear best fitting of which returned a zero-current level at +63.0 mV (not shown). This value compares favorably with the theoretical Nernst Na⁺ reversal potential calculated for our ionic conditions (V_Na = +61.0 mV). The voltage dependence of the conductance underlying I_NaP (G_NaP, calculated as explained in the MATERIALS AND METHODS) is shown in the average plot of Fig. 1 A, inset. Boltzmann fitting to data points returned a half-activation voltage, V_1/2, of –44.4 mV, and a slope factor, k, of –5.2 mV. The ratio between the peak G_NaP value found in each cell and the maximal value of the conductance underlying the transient Na⁺ current (I_NaT) expressed by the same cell was also calculated, and averaged 0.0187 ± 0.0097 (n = 5).

View larger version (25K): [in this window] [in a new window] [Download PPT slide]   Figure 1 Basic features of INaP in EC layer II principal neurons. (A) INaPs evoked in a representative acutely isolated neuron (cell E6M15) by 500-ms depolarizing square pulses at –65 to –35 mV. All the traces shown are TTx subtracted (control minus 1 µM TTx). The holding potential was –80 mV. Scale bars: 100 pA, 100 ms. (Inset) Average plot of normalized GNaP derived, as explained in MATERIALS AND METHODS, from step protocols (INaP amplitude was measured by averaging the data points between 400 and 500 ms from the pulse onset; n = 5). The best Boltzmann fitting to data points is also shown. Fitting parameters were: V1/2 = –44.4 mV, k = –5.2 mV. (B and C) INaPs evoked in a representative acutely isolated neuron (B; cell A6M15) and a representative in situ neuron (C; cell D7904) by slow voltage ramps (50 mV/s, from –80 to +20 mV; the voltage protocol is schematized above the insets). The currents shown in the main panels have been obtained by digitally subtracting the traces recorded in 1 µM TTx from control traces (both shown in the insets; calibration bars: 100 pA in B, 200 pA in C). (D) Plots of the voltage dependence of activation of INaP in the same two neurons as shown in B and C (D1 and D2, respectively). Na+ conductances (GNaPs) were derived from the early parts of the TTx-subtracted currents by applying the extended Ohm equation (MATERIALS AND METHODS). GNaP plots were normalized for the maximal values and fitted with single Boltzmann functions (empty lines). Fitting parameters were: V1/2 = –47.6 mV, k = –5.5 mV (D1); V1/2 = –50.8 mV, k = –4.5 mV (D2). The perpendicular, dotted lines indicate the half-maximal activation and the corresponding V1/2.

Ramp protocols were also used in experiments performed in in situ neurons. In this situation, TTx subtraction always returned prominent I_NaPs in isolation, whose I–V relationship closely resembled that of I_NaPs in acutely dissociated neurons (Fig. 1 C). I_NaP amplitude, measured at the peak of the I–V relationship, was significantly higher in in situ than in isolated neurons (–179.3 ± 120.7 pA, n = 54, vs. –65.6 ± 37.9 pA, n = 38; P < 5 x 10^–7), whereas the current density did not significantly differ in the two situations (–17.7 ± 29.1 pA/pF, n = 54, vs. –16.5 ± 8.6 pA/pF, n = 38; P = 0.8). These findings strongly suggest that the channels responsible for I_NaP are located not only on the soma, but also on neuronal processes severed by the dissociation procedure. Activation curves of I_NaPs recorded in both in situ and isolated neurons were also constructed. Conductance values were derived from I_NaPs by applying the extended Ohm's law (see MATERIALS AND METHODS), and the resulting activation curves were fitted with single Boltzmann functions (Fig. 1 D). Average half-activation potentials and slope factors were very similar in in situ neurons (V_1/2 = –51.3 ± 3.9 mV, k = –4.0 ± 0.7 mV, n = 39) and isolated neurons (V_1/2 = –48.7 ± 4.7 mV, k = –4.4 ± 0.9 mV, n = 19). These values compare favorably with those obtained from step protocols (see above), and are also in good agreement with the activation parameters previously reported for I_NaPs expressed in other neuronal systems (French et al. 1990; Brown et al. 1994; Baker and Bostock 1997).

Given the effectiveness of TTx subtraction in isolating I_NaPs in both isolated and in situ neurons, this procedure was routinely used in our study. All of the data presented from this point on are from TTx-subtracted currents.

I_NaP Is Not a Window Current Generated by Transient Na⁺ Channels
It is well known that a noninactivating, "window" current (I_NaTW) can be generated by the gating properties of fast, transient Na⁺ channels (Hodgkin and Huxley 1952). To address the issue of whether the I_NaP expressed by EC layer II neurons can be accounted for by a classical window conductance, we analyzed the voltage-dependence properties of I_NaT in acutely dissociated neurons. Fig. 2 A shows Na⁺-current traces recorded in a representative neuron in response to an activation–inactivation pulse protocol. Peak-current amplitudes were measured and used for deriving conductance values (G_NaT); normalized activation and steady state inactivation plots were then constructed (Fig. 2 B), and fitted with Boltzmann functions. The optimal approximation to data points returned by single Boltzmann functions suggested the existence of functionally homogeneous, transient Na⁺ channels in the neuronal preparation under examination. In eight cells, average V_1/2 and k were –32.5 ± 6.5 mV and –3.6 ± 0.9 mV, respectively, for the activation function, and –59.8 ± 5.2 mV and 4.5 ± 0.9 mV, respectively, for the steady state inactivation function. These values are similar to those reported in a number of other studies on neuronal I_NaT voltage dependence (e.g., Sah et al. 1988; Huguenard et al. 1988; Cummins et al. 1994). The product of the activation and steady state inactivation functions was then calculated in individual cells to derive the theoretically predicted voltage dependence of the window conductance (G_NaTW) arising from transient Na⁺ channels (Fig. 2 B, dotted line). Fig. 2 C illustrates the I_NaP evoked by a standard ramp protocol, and isolated via TTx subtraction, in the same cell as in A and B. The conductance underlying I_NaP (G_NaP) was calculated and compared with the predicted G_NaTW (Fig. 2 D); an evident discrepancy in the voltage dependence of the two conductances could be observed at potentials positive to about –30 mV, where G_NaTW rapidly fell towards zero, whereas G_NaP maintained relatively high values, though it also showed a characteristic decline from its maximum (see below). The same discrepancy between G_NaTW and G_NaP was observed in four other acutely dissociated neurons, in which both well-clamped I_NaT s and sizable I_NaPs could be recorded. In a broader cell population, the amplitudes of the reconstructed G_NaTW and of G_NaP were measured both at the peak [G_(max)] and at a voltage point positive by 20 mV to that of the peak [G₍₊₂₀₎]. The average ratios G_NaTW(+20)/G_NaTW(max) and G_NaP(+20)/G_NaP(max) were 0.076 ± 0.057 (n = 8) and 0.715 ± 0.102 (n = 12), respectively (P < 5 x 10^–12). In addition, in those neurons in which both I_NaT and I_NaP were quantified, the size of the predicted I_NaTW was much smaller than that of the observed I_NaP (see Table ). These data clearly indicate that by far most of the I_NaP expressed by EC layer II neurons is not a classical I_NaTW, similar to what has been previously observed in hippocampal neurons (French et al. 1990).

View larger version (19K): [in this window] [in a new window] [Download PPT slide]   Figure 2 The INaP expressed by EC layer II principal neurons is largely not accounted for by a classical Hodgkin-Huxley window current. (A) Voltage protocols used to study the voltage dependence of activation and steady state inactivation of the fast Na+ current (INaT) (A1), and leak-subtracted currents recorded in an exemplary acutely isolated cell (A2; cell L7719). The duration of the inactivating prepulse was 120 ms. (Inset) The current recorded at –30 mV is expanded in amplitude to highlight the presence of a persistent current (calibration bars: 20 pA, 25 ms). (B) Plots of the voltage dependence of activation () and steady state inactivation (•) of INaT in the same cell as in A. Na+ conductances (GNaT s) were derived from peak INaT amplitudes by applying the extended Ohm equation (MATERIALS AND METHODS). GNaT plots were normalized for the maximal values and fitted with single Boltzmann functions (continuous lines). Fitting parameters were: V1/2 = –58.2 mV, k = 5.0 mV (steady state inactivation); V1/2 = –29.9 mV, k = –3.9 mV (activation). The predicted window conductance, GNaTW (RESULTS), is also shown (dotted line: note the different amplitude scale, shown on the right axis). (C) Currents evoked by a slow voltage ramp (50 mV/s) in the same cell as in A and B, before and after the application of 1 µM TTx (inset; calibration bar: 20 pA), and TTx-subtracted INaP (main panel). (D) The voltage dependence of the conductance (GNaP) underlying the INaP shown in C is compared with that of the GNaTW reconstructed in the same cell. Both conductances have been normalized to the maximal values.

View larger version (13K): [in this window] [in a new window] [Download PPT slide]   Figure 3 INaP looks different, depending on the slope of the ramp applied to elicit it. (A) Voltage ramps exploring the –80/+20-mV voltage range with different depolarization speeds (100, 66.7, 50, 25, 12.5, and 6.25 mV/s) were tested (A1), and the TTx-sensitive currents recorded in an exemplary in situ neuron (cell B7902) are depicted in A2, all over the same time scale. (B) Average, normalized INaP peak amplitude as a function of the inverse of ramp slope (n = 12; same ramp protocols as illustrated in A1). The plot could be fitted by a double exponential function [I= A1 · exp(–1/s) + A2 · exp(–2/s) + C, where s is the ramp slope and 1 are slope constants] with A1 = 0.295, 1 = 0.317 V/s, A2 = 0.382, 2 = 20.2 mV/s (continuous line).

We then investigated the issue of I_NaP inactivation in further detail by analyzing the effects of variable prepulse potentials on ramp-activated I_NaPs. The protocol employed is illustrated in Fig. 4 A1. 50-mV/s voltage ramps were preceded by very-long-lasting (15 s) conditioning prepulses at various voltage levels (V_cond). When V_conds of –90 to –50 mV were used, the ramp started from the same voltage level as V_cond itself, rather than from a fixed, negative voltage level: in this way, the possible occurrence of recovery from inactivation during the initial part of the ramp was avoided. At more positive V_conds, the ramp started from –50 mV, so as to preserve the voltage region of I_NaP peak. Currents recorded with the above protocol in a representative in situ neuron are shown in Fig. 4 A2. I_NaP peak amplitude turned out to markedly depend on the conditioning potential. Average, normalized current traces obtained from seven in situ neurons are depicted in Fig. 4 B1. It can be observed that voltage-dependent steady state inactivation of I_NaP was nearly complete at about –20 mV. The average plot of I_NaP's voltage dependence of inactivation (Fig. 4 B2) could be fitted by a single Boltzmann function, with V_1/2 at about –49 mV and a slope factor, k, of 10 mV. In addition, we also constructed an activation plot from the average I_NaP derived from the same neuron pool, and fitted it with a single Boltzmann function (Fig. 4 B2). Note that, importantly, G_NaP activation and steady state inactivation functions overlapped over a wide voltage range. Due to this phenomenon, a significant window conductance (G_NaPW) is expected to arise from G_NaP. The predicted voltage dependence of G_NaPW is depicted in Fig. 4 B2 (dotted line), and will be compared with relevant experimental data later on in the paper.

View larger version (14K): [in this window] [in a new window] [Download PPT slide]   Figure 4 Voltage dependence of inactivation of INaP. (A) Voltage protocol used for the study of INaP voltage dependence of inactivation (A1), and TTx-subtracted currents recorded in an exemplary in situ neuron (A2; cell D7808). The conditioning prepulses preceding each ramp varied from –90 to –10 mV in 10-mV steps, and their duration was 15 s. Note that here, as well as in all of the voltage-clamp protocols applied in whole-cell experiments, each conditioning prepulse/ramp cycle was preceded by a 15-s period at –80 mV so as to allow recovery from the inactivation developed during the foregoing cycle (MATERIALS AND METHODS). (B) Average currents and steady state inactivation function from seven cells. (B1) Average, normalized currents. The currents evoked in each cell as shown in A were divided by the maximal peak-current value observed in the same cell; the traces were then averaged among cells and further normalized to the maximal peak value thus obtained. Standard deviations are also shown at 5-mV intervals in the traces preceded by conditioning prepulses at –90, –50 and –10 mV. (B2) Average steady state inactivation plot (•). The amplitudes of the currents evoked as shown in A were measured at the ramp potential of –40 mV. The values thus obtained were divided by the maximal value observed in each cell, averaged among cells, and plotted as a function of the conditioning prepulse potential. The inactivation plot was best-fitted by a single Boltzmann function, with V1/2 = –48.8 mV and k = 10.0 mV. An average activation plot was also constructed by deriving the normalized conductance (MATERIALS AND METHODS) from the average, normalized current obtained from the same cells (B1; prepulse potential = –90 mV), and best-fitted with a single Boltzmann function, with V1/2 = –52.6 mV and k = –4.6 mV. Each plot-fitting pair was further normalized to the value of the fitting-function amplitude coefficient. The dotted line represents the product of the two fitting functions, corresponding to the predicted voltage dependence of the resulting window conductance (GNaPW).

View larger version (18K): [in this window] [in a new window] [Download PPT slide]   Figure 5 Time dependence of inactivation of INaP. The voltage protocols applied are shown in B. A shows TTx-sensitive currents evoked with the protocol shown in B in a representative in situ neuron (cell B7808) (prepulse potential = –60 mV). Individual currents have been staggered along the x axis by intervals proportional to the duration of the inactivating (conditioning) prepulse (t, each indicated below the traces). The actual x (voltage) scale is indicated, for the last trace only (top right). (C) Average, normalized amplitudes of INaP as a function of depolarizing prepulse duration (mean ± SEM, n = 6–12). Each point family refers to a single prepulse voltage level. Single-exponential best fittings are also shown (continuous lines), along with time-constant () values.

View larger version (16K): [in this window] [in a new window] [Download PPT slide]   Figure 6 Time dependence of recovery from inactivation of INaP. The voltage protocols applied are shown in B. The duration of the inactivating prepulse at –30 mV was 15 s. (A) TTx-sensitive currents evoked with the protocol (B) in a representative in situ neuron (cell K7805; recovery potential = –80 mV). Individual currents have been staggered along the x axis by intervals proportional to the duration of the recovery (conditioning) prepulse (t, each indicated below the traces). The actual x (voltage) scale is indicated, for the last trace only (top right). (C) Average, normalized amplitudes of INaP as a function of recovery-prepulse duration (means ± SEM, n = 3–8). The recovery potential was –80 (top) or –90 (bottom) mV. Single-exponential best fittings are also shown (continuous lines), along with time-constant () values.

View larger version (14K): [in this window] [in a new window] [Download PPT slide]   Figure 7 INaP inactivation during long-lasting depolarizing voltage steps. (A) Averaged, TTx-subtracted currents (A1) from five in situ neurons as obtained in response to 15-s voltage steps to the test potentials indicated on the left. The voltage protocol applied is shown in A2. Standard deviations for some points, single-exponential best fittings of the decay phases (enhanced lines), and time-constant () values are also shown; the starting point of the fitted part of the traces is indicated by the vertical, dashed line. Scale bars: 50 pA, 2.5 s. (B) The amplitude of the noninactivating ("offset") component of average INaP (•), measured at the different test potentials, is compared with the predicted window current arising from the gating properties of INaP (INaPW, continuous line). The INaPW shown has been derived from the GNaPW depicted in Fig. 4 B2 (dotted line).

Modeling of I_NaP Slow Inactivation
To further clarify the basis of the experimentally observed decline in G_NaP at positive potentials (Fig. 2 D), a theoretical reconstruction of the biophysical properties of I_NaP inactivation was carried out. A simple Hodgkin-Huxley model, considering a single inactivation gate switching between two energy states, was considered in order to give account for the monoexponential time course of I_NaP decay and recovery from inactivation. This reconstruction was merely phenomenological and was given no mechanistic meaning since single-channel data clearly indicated different features of the underlying elementary events (see below). The time constants of I_NaP inactivation and recovery from inactivation and the data on voltage dependence of I_NaP steady state inactivation were processed to derive numerical values for the rate constants of the inactivation-gate transitions, as explained in MATERIALS AND METHODS. Fig. 8 B shows the voltage dependence of the values thus obtained for the two rate constants, and β. The plots were then best fitted with the empirical function, (or β) = (a · V_m + b)/{1 – exp[(V_m + b/a)/k]}, where V_m is the membrane voltage. The numerical values returned by the fittings for the a, b, and k coefficients, in both the and β plots, are indicated in the legend to Fig. 8. The voltage-dependence functions thus obtained for and β were then used to derive the predicted voltage dependence of I_NaP inactivation-gating time constants (see MATERIALS AND METHODS). The concordance between the reconstructed, theoretical function and the real data is shown in Fig. 8 A.

View larger version (17K): [in this window] [in a new window] [Download PPT slide]   Figure 8 Modeling of INaP inactivation and of its consequences on ramp protocols. (A) Time constants of INaP inactivation (circles) and recovery from inactivation (triangles) as a function of membrane voltage. Time constants of inactivation (h) determined with both ramp (•) and step () protocols are shown. The continuous line is the theoretical function describing h(V), as obtained from the fitting functions of the voltage dependence of rate constants (see below). Crosses indicate time constants of inactivation of ensemble-average traces obtained in single-channel, cell-attached experiments (see text), here included for comparison. (B) Rate constants of the transitions of the inactivating gate, h, to () and from (β) the permissive position (see RESULTS and MATERIALS AND METHODS for details). Values for (squares) and β (diamonds) have been calculated, as explained in MATERIALS AND METHODS, on the basis of time constants measured from both ramp (filled symbols) and step (empty symbols) protocols. The continuous lines represent the fitting functions (described in RESULTS) for (V) and β(V), with a = –2.88 · 10–3 mV–1 s–1, b = –4.9 · 10–2 s–1, k = 4.63 mV (); a = 6.94 · 10–3 mV–1 s–1, b = 0.447 s–1, k = –2.63 mV (β). (C) Effects of INaP inactivation on the amplitude and shape of the INaPs generated by simulated voltage-ramp protocols with different depolarization slopes. The curves shown correspond to the "ideal" (unperturbed) INaP (Contr.), and to INaPs generated by simulated voltage ramps of 100, 66.7, 50 (dashed line), 25, 12.5, and 6.25 mV/s. For describing the voltage dependence of activation of the conductance underlying INaP (GNaP), the same parameters as obtained from the data of Fig. 4 B2 were used in these simulations. (D) Dependence of INaP peak amplitude on the inverse of voltage-ramp slope in simulated experiments. The data points shown were derived from the same current traces depicted in C. The plot has been fitted with a single exponential function (continuous line), with A = 0.23, = 12.8 mV/s. (E) The voltage dependence of GNaP in a simulated 50-mV/s ramp protocol (dashed line) is compared with that of the ideal (unperturbed) GNaP (continuous line). The initial part (negative to –36 mV) of the same curve has also been best fitted with a single Boltzmann function (dotted line). Fitting parameters were: V1/2 = –53.0 mV, k = –4.5 mV.

Finally, a simulated protocol of steady state inactivation revealed that, during a ramp preceded by a prepulse that fully inactivates I_NaP, a significant recovery from inactivation can occur provided that the ramp starts at sufficiently negative levels and is sufficiently slow. For instance, during a 25-mV/s ramp starting at –80 mV, 17% of the total current can recover from inactivation (not shown). By contrast, voltage protocols similar to those we employed for the study of I_NaP voltage-dependent inactivation (see above) determined no appreciable ramp-dependent recovery from inactivation of I_NaP .

Single-Channel Bases of I_NaP Slow Inactivation and I_NaPW Generation
The single-channel basis of I_NaP in EC layer II neurons has been described elsewhere (Magistretti et al. 1999). In that study, we found that, after membrane step depolarization, I_NaP is generated by early as well as late single Na⁺-channel openings of much more prolonged duration and of significantly higher conductance than the usual, transient Na⁺-channel openings responsible for I_NaT generation. Here, we report how the same persistent Na⁺-channel activity also accounts for some particular biophysical properties described for macroscopic I_NaP , namely I_NaP time-dependent inactivation and I_NaPW generation.

Recordings from cell-attached patches in acutely isolated EC layer II neurons frequently revealed the presence of a persistent Na⁺-dependent channel activity that proved to remain stable even over prolonged periods of time (tens of minutes). When very-long-lasting (20 s) depolarizing steps at –10 mV were commanded from a holding potential of –80 mV, multiple, repetitive single-channel openings were observed that tended to cluster preferentially at the beginning of the test pulse. Typical examples of this channel activity for a series of consecutive 20-s test pulses is shown in Fig. 9 A1. A detail of some prolonged, late channel openings is also provided by Fig. 9 A1, inset. Ensemble averaging of multiple traces was then carried out for each patch (Fig. 9 A2). Due to the very long overall duration of the recording cycle required for every single sweep (40 s), only a limited number of traces could be recorded in each patch (15 on average), what explains the low signal-to-noise ratio of ensemble-average traces. In all cases, however, the averaged currents showed a noticeable trend to decay towards zero, and this decay could be properly fitted by a single exponential function (Fig. 9 A2, blank trace). The time constant of average-current decay was 2.66 ± 0.52 s in six patches at –10 mV, a value that compares favorably with those found in whole-cell protocols on macroscopic I_NaP inactivation at the most positive voltage levels tested (see Fig. 8 A). Ensemble averaging of all of the available sweeps recorded from the same six patches returned a better signal-to-noise ratio (Fig. 9 B). The decay time constant of the resulting average current was 2.46 s, again in good agreement with the data obtained from the analysis of both individual patches and whole-cell recordings.

View larger version (26K): [in this window] [in a new window] [Download PPT slide]   Figure 9 Single-channel basis of INaP slow inactivation. (A) Cell-attached recordings at the test potential of –10 mV in a representative patch from an acutely isolated EC layer II neuron (patch D8708). The patch was held at –80 mV, and 20-s depolarizing pulses were delivered beginning from the time point marked by the arrow. A1 shows 12 sweeps selected from an ensemble of 20 (scale bars: 1 pA, 2 s). (Inset) Detail of long-lasting single-channel openings (scale bars: 1 pA, 200 ms). The shut- (s) and open- (o) channel levels used in dwell-time analysis are marked by the horizontal, dashed lines. A2 illustrates the ensemble average of the 20 sweeps recorded from the same patch as in A1 (scale bars: 0.1 pA, 2 s). The single-exponential best fitting of the average current's decay phase (blank line) and the corresponding time-constant value are also shown; values for A and C (see MATERIALS AND METHODS) were –83.3 and –12.4 fA, respectively. (B) Ensemble average of 86 sweeps recorded at the test potential of –10 mV, with the same experimental protocol as explained above, from six different patches (scale bars: 0.1 pA, 2 s). The single-exponential best fitting of the average current's decay phase (blank line) and the corresponding time-constant value are also shown (values for A and C were –74.4 and –11.7 fA, respectively).

View larger version (9K): [in this window] [in a new window] [Download PPT slide]   Figure 10 Frequency distribution of dwell times of "persistent" channels in the open state in long-lasting (20 s) depolarizing step protocols. (A) Open-time histogram of channel openings recorded at the test potential of –10 mV in the same patch as shown in Fig. 9 A. (Inset) Detail of the histogram's region from 25 ms upwards. A double-exponential best fitting (continuous lines) and the corresponding time-constant values are also shown; the W2/W1 ratio (see RESULTS for details) was 0.081. (B) Open-time histogram of channel openings recorded at the test potential of –40 mV in the same patch as shown in A. A double-exponential best fitting (continuous line) and the corresponding time-constant values are also shown; the W2/W1 ratio was 0.099.

View larger version (30K): [in this window] [in a new window] [Download PPT slide]   Figure 11 Single-channel basis of INaPW generation. (A) Cell-attached recordings at the test potential of –40 mV in a representative patch from an acutely isolated EC layer II neuron (patch D8617). The patch was held at –80 mV, and 20-s depolarizing pulses were delivered beginning from the time point marked by the arrow. The traces shown are 12 consecutive sweeps (scale bars: 2 pA, 2 s). (Inset) Time expansion of some single-channel openings (scale bars: 2 pA, 200 ms). The shut- (s) and open- (o) channel levels used in dwell-time analysis are remarked by the horizontal dashed lines. (B) Ensemble average of 33 sweeps recorded at the test potential of –40 mV, with the same experimental protocol as explained above, from three different patches (scale bars: 0.1 pA, 2 s). The single-exponential best fitting of the average current's decay phase (blank line) and the corresponding time-constant value are also shown; values for A and C (see MATERIALS AND METHODS) were –51.7 and –17.3 fA, respectively.

The observation that the channel mean open times are far exceeded by the time constants of ensemble-average current inactivation clearly indicates that the latter do not reflect average channel-opening lifetimes in a simple model considering two open states each undergoing one single closure process (with rate constant b) towards an absorbing state, like:

with the index i being either 1 or 2, and with a_i >> b_i. Rather, alternative models considering late first openings and/or late reopenings must be taken into account. In an extreme situation, late channel openings such those of Fig. 9 A1 might all be first openings of as many distinct channels inactivating towards an absorbing state, in a model qualitatively similar to that depicted in Scheme I, but in which the kinetics of the macroscopic inactivation process would rather reflect the rate constants of the closed-to-open reactions, a_i. This interpretation, which resembles the classical Aldrich-Corey-Stevens model of the kinetics of transient Na⁺ channels (Aldrich et al. 1983), seems very unlikely since, in patches containing most likely only one "persistent" channel, delayed first openings occurred very infrequently, whereas late reopenings were often observed (Magistretti et al. 1999). An alternative possibility is that the channel, once open, can reach multiple nonconductive states, one of which is virtually absorbing at positive potentials, thus causing a true channel inactivation. Under this assumption, a channel behavior such as that in Fig. 9 A1 could be accounted for by repetitive reopenings of a small number of persistent channels. The most economical kinetic scheme of such a behavior is seen in Fig. 2.

View larger version (6K): [in this window] [in a new window] [Download PPT slide]   Scheme S2

	DISCUSSION

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
The present study provides a biophysical characterization of the I_NaP expressed by rat EC layer II principal neurons. The major issues we addressed deal with the mechanism of generation of I_NaP , the possible influence of the voltage protocols employed for the study of I_NaP biophysical properties, and the real degree of I_NaP persistence over time and specific voltage windows.

In general, the hypotheses on generation of persistent Na⁺ currents consider two main possibilities: (a) I_NaP simply derives from an incomplete steady inactivation of transient Na⁺ channels over a narrow voltage window, due to the partial superimposition of activation and steady state inactivation voltage-dependence curves of the corresponding conductance; (b) I_NaP is the result of channel openings that do not functionally behave according to the properties of transient Na⁺ channels, but derive from a rare and atypical gating modality of the same channels (Alzheimer et al. 1993b), possibly under modulatory control by G proteins (Ma et al. 1997) or other factors, or, alternatively, from a different channel species. To discriminate between these two possibilities, it is necessary to accurately determine the properties of the window current generated by transient Na⁺ channels and compare them with those of I_NaP present in the same cell preparation. The study of acutely dissociated EC neurons allowed us to perform careful measurements on I_NaT, and therefore to make reliable predictions on the resulting I_NaTW. The comparison between I_NaTW and I_NaP was done both on a statistical basis and, in some cases, in the same cells. Our data on the voltage dependence and amplitude of these currents indicate that I_NaTW, in contrast with what was reported in a previous study (Fan et al. 1994), cannot be a major source of the I_NaP expressed by EC principal neurons. An alternative, specialized mechanism must be implied in the generation of the large I_NaPs found in these neurons. An exciting possibility, already suggested elsewhere on the basis of whole-cell data (French et al. 1990), is that I_NaP is generated by a specialized Na⁺ channel at least biophysically distinct from those generating the fast transient Na⁺ current (Masukawa et al. 1991). Indeed, single-channel, patch-clamp experiments in EC layer II neuron somata indicate that while I_NaT is generated by a typical 15-pS channel with fast activation and inactivation kinetics, I_NaP is due to an 20-pS channel activity with a 10-mV lower threshold of activation, and a sustained, high open probability during prolonged depolarizations (Magistretti et al. 1999). The issue regarding the molecular diversity of Na⁺ channels (see Ragsdale and Avoli 1998, for recent review) expressed by EC layer II neurons is beyond the scope of the present study and will be discussed elsewhere.

Another interesting issue raised by our data relates to the definition itself of I_NaP. Due to their practicality, ramp protocols are the most widely used means for eliciting and isolating I_NaP. However, the employment of such protocols presupposes that they can adequately reproduce the voltage dependence of I_NaP without significantly recruiting any other kinetically different (i.e., faster-decaying) Na⁺-current component. This is often implicitly assumed rather than demonstrated. Moreover, the notion of "slow voltage ramp" suitable for I_NaP activation varies considerably among different experimental works (Alzheimer et al. 1993a; Uteshev et al. 1995; Cepeda et al. 1995; Pennartz et al. 1997; Parri and Crunelli 1998; Cummins et al. 1998). The experiments we carried out by running voltage ramps of variable slopes clearly indicate that the amplitude of the ensuing I_NaP strictly depends on the depolarization rate applied. Our data show how, when ramps of progressively decreasing slopes (from 100 to 6.25 mV/s) are commanded, I_NaP amplitude decreases in a roughly biexponential fashion. The existence of a faster exponential component may be due to the presence of Na⁺-current components kinetically intermediate between classical, "fast" Na⁺ currents and the persistent Na⁺ current. These intermediate kinetic components were easily observed in our step-protocol recordings (see Fig. 1 A and 7 A1), and their properties in EC layer II stellate cells will be described in detail elsewhere. The measurements on I_NaPs evoked with "slow" voltage ramps may therefore be contaminated by the superimposition of such current components, unless the commanded ramp is slow enough.

Moreover, our data show that, importantly, the process of I_NaP slow inactivation is a potential source of distortions in the measurements of I_NaP biophysical parameters (amplitude, voltage dependence of activation, reversal) when this current is elicited by voltage ramps. All these considerations point to the importance of accurately choosing the voltage protocol applied for the study of I_NaP in each specific experimental situation. We propose that some previously reported, atypical biophysical features of I_NaP , particularly regarding its nonsigmoidal voltage dependence of activation (Brown et al. 1994), are the result of the interaction between multiple Na⁺-current slow decay components and the ramp protocols employed. In our study, the ramp protocols routinely used were chosen so as to minimize both the possible contribution of intermediate-kinetics Na⁺ current components and the effects of voltage- and time-dependent inactivation of I_NaP.

The inactivation properties of what, according to our operative definition, can be considered as I_NaP have been characterized in detail in our study. Our experiments indicate that, in our preparation, the steady state inactivation of the conductance underlying I_NaP (G_NaP) (a) has a voltage dependence that extends over a wide voltage window, and (b) reaches a nearly complete level at –20 to –10 mV. The former of these features, together with the relative position of the G_NaP activation curve along the voltage axis, is expected to give rise to a major, time-independent, "window" current over a limited voltage range. This window current (I_NaPW), clearly different from that arising from the voltage-dependence properties of the transient Na⁺ conductance (I_NaTW), could also be directly demonstrated both as nonzero baselines at the beginning of ramp protocols on voltage dependence of inactivation (see Fig. 4), and as offsets, or pedestals, in exponentially decaying I_NaPs elicited by long-lasting voltage steps (Fig. 7). The peak of the observed I_NaPW occurred at voltage levels very close to those at which I_NaP-dependent, theta-like subthreshold membrane-potential oscillations are generated by EC stellate cells (Alonso and Llinás 1989; Alonso and Klink 1993; Klink and Alonso 1993; van der Linden and Lopes da Silva 1998). Our data indicate that the contribution of I_NaPW to the total persistent Na⁺ current over the voltage range of subthreshold-oscillation generation must be substantial, since the peak I_NaPW amplitude was estimated to exceed that of the predicted I_NaTW by more than four times. Since the subthreshold oscillations generated by the stellate cells can last indefinitely, our observations also imply that only a fraction of the total I_NaP, namely I_NaPW itself, is sufficient for sustaining them.

I_NaP slow inactivation and recovery from inactivation were found to occur with voltage-dependent time constants in the order of a few seconds. We worked out an analytical reconstruction of the kinetics of these processes which, if introduced into suitable neuronal models, should allow us to make predictions on the effects of slow voltage-dependent inactivation on I_NaP modulation of membrane-voltage events. We demonstrated that I_NaP inactivation can considerably affect the apparent current maximal amplitude during the delivery of slow depolarizing ramps. Therefore, it is conceivable that the recruitment of I_NaP and its impact onto membrane-voltage dynamics can be significantly influenced by the speed of membrane depolarization. For instance, the ability of I_NaP to bring the membrane potential towards threshold for action-potential firing may be expected to be higher in response to a step depolarizing current injection than to slower or sustained depolarizations. This is consistent with the role of I_NaP, demonstrated in various central nervous system neurons (Jahnsen and Llinás 1984; Franceschetti et al. 1995; Parri and Crunelli 1998) including EC stellate cells (Alonso and Klink 1993; Klink and Alonso 1993) in promoting transient low-threshold spikes and sustaining phasic firing in response to fast depolarizations. On the other hand, I_NaP inactivation, which we found to be eventually complete above a physiologically interesting range of membrane potentials, may have an important role in limiting detrimental Na⁺ influx during pathological conditions such as seizures and ischaemia (see also Taylor and Meldrum 1995).

The question can then be raised whether the biophysical properties we describe here for the I_NaP expressed by EC stellate cells represent a general feature of this current in various neuronal populations. Slow time-dependent inactivation of I_NaP has been previously reported in neocortical pyramidal neurons (Fleidervish and Gutnick 1996), although in that case the voltage dependence of the process was not investigated in detail, whereas a nondecaying "offset" I_NaP component corresponding to 30% of the total was found in experiments on time dependence of inactivation even after inactivating prepulses at +20 mV. This is in contrast with our finding of a virtually complete inactivation at approximately –20/–10 mV. The discrepancy may imply the existence of interesting functional differences among I_NaPs expressed by different neuronal populations, which encourages further studies in other cell systems; alternatively, it may be the consequence of the inactivation protocol employed in Fleidervish and Gutnick 1996, in which the depolarizing ramp after the inactivating prepulse started from a fixed, negative voltage level (–80 mV): this may allow some degree of recovery from inactivation during the early phase of the ramp, as also suggested by the output of our modeling study (RESULTS).

Finally, the present study provides an insight into the fine mechanisms underlying the complex biophysical features displayed by I_NaP. Our previous work has already clarified the nature and elementary properties of the single-channel events responsible for I_NaP generation in EC principal neurons (Magistretti et al. 1999). The data we report here clearly point to the importance of late Na⁺-channel (re)openings, many times longer in duration than those generating classical, transient I_NaT in central neurons, for determining the behavior of slow kinetic components of macroscopic I_NaP. Late Na⁺-channel (re)openings have already been identified as a possible source of I_NaP in neocortical neurons (Alzheimer et al. 1993b) and ventricular myocytes (Ju et al. 1994), but those we observed were comparatively much longer-lived. The slow inactivation of whole-cell I_NaP we characterized was paralleled by that of ensemble-average traces from single-channel, cell-attached recordings. In turn, the latter appeared to be the consequence of the slow delivery of channels to an inactivated state. This can be the consequence of either late first openings or, much more probably, low-rate transitions from open states that are more likely to switch to a close, re-recruitable state, or vice versa. The inactivated state was nearly absorbing at –10 mV, but not at –40 mV, where the existence of a major window current generated by the voltage-dependent properties of G_NaP had been predicted and demonstrated (see above). Again, whole-cell I_NaPW had its single-channel correlate in late, repetitive openings able to generate measurable net inward currents after as long as 20 s of membrane depolarization.

In conclusion, the I_NaP expressed by EC layer II principal neurons is a prominent current operating in a subthreshold range of membrane potentials, most of which is generated by a process independent of the classical gating behavior of the transient Na⁺ conductance. It also displays complex and previously nonrecognized biophysical characteristics that appear to be tailored to the specific role this current is known to play in the generation of the sub- and near-threshold membrane-potential events typical of the same neurons. The concept of "persistent Na⁺ current" may turn out to be susceptible to some critical revision also in other experimental situations, with reference to both the existence of multiple and heterogeneous functional components, and the expression of kinetic and voltage-dependence properties that might influence their impact onto neuronal function in previously unforeseen ways.

Portions of this work were previously published in abstract form (Magistretti, J., and A. Alonso. 1998. NYAS Meetings. Abstr. No. PI–29; Alonso, A., and J. Magistretti. 1998. Soc. Neurosci. Abstr. 24:2035).

	ACKNOWLEDGMENTS

 
J. Magistretti thanks the Human Frontier Science Program Organization (HFSPO), the Istituto Nazionale Neurologico "Carlo Besta," and Dr. Marco de Curtis for support.

This work has been funded by grants from the Medical Research Council of Canada, HFSPO, and the North Atlantic Treaty Organization to A. Alonso.

Submitted: 2 June 1999
Revised: 4 August 1999
Accepted: 9 August 1999

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