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The Journal of Neuroscience, March 1, 1999, 19(5):1675-1690
Somatic Recording of GABAergic Autoreceptor Current in Cerebellar
Stellate and Basket Cells
Christophe
Pouzat and
Alain
Marty
Arbeitsgruppe Zelluläre Neurobiologie, Max-Planck-Institut
für Biophysikalische Chemie, 37077 Göttingen, Germany
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ABSTRACT |
Patch-clamp recordings were performed from stellate and basket
cells in rat cerebellar slices. Under somatic voltage clamp, short
depolarizing pulses were applied to elicit action potentials in the
axon. After the action potential, a bicuculline- and
Cd2+-sensitive current transient was observed. A
similar response was obtained when eliciting axonal firing by
extracellular stimulation. With an isotonic internal
Cl solution, the peak amplitude of this current
varied linearly with the holding potential, yielding an extrapolated
reversal potential of  20 to 0 mV. Unlike synaptic or autaptic
GABAergic currents obtained in the same preparation, the current
transient had a slow rise-time and a low variability between trials.
This current was blocked when 10 mM BAPTA was included in
the recording solution. In some experiments, the current transient
elicited axonal action potentials. The current transient was reliably
observed in animals aged 12-15 d, with a mean amplitude of 82 pA at
70 mV, but was small and rare in the age group 29-49 d. Numerical simulations could account for all properties of the current transient by assuming that an action potential activates a distributed GABAergic conductance in the axon. The actual conductance is probably restricted to release sites, with an estimated mean presynaptic current response of 10 pA per site (70 mV, age 12-15 d). We conclude that in
developing rats, stellate and basket cell axons have a high density of
GABAergic autoreceptors and that a sizable fraction of the
corresponding current can be measured from the soma.
Key words:
cerebellum; basket cell; stellate cell; GABA; IPSCs; axon; autoreceptors; autapses
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INTRODUCTION |
The axonal membrane of brain neurons
is responsible for the generation and propagation of action potentials
as well as for the consequent depolarization of nerve terminals and
associated Ca2+ entry. In addition, axon terminals
and adjacent areas contain receptors for neurotransmitters, called
presynaptic receptors (for review, see McGehee and Role, 1996 ; Langer,
1997; Wu and Saggau, 1997). Presynaptic receptors can be activated by
the release of neurotransmitter coming from the same neurons, in which
case they act as autoreceptors, by stimulation of axo-axonic synapses, or by spillover of neurotransmitter released from nearby neurites belonging to other neurons. Activation of presynaptic receptors in turn
can potently modify the strength of neurotransmitter release.
Among the many subtypes of presynaptic receptors that are believed to
exist, GABAA receptors are perhaps the most extensively studied. Activation of presynaptic GABAA receptors has been
proposed as the cause of presynaptic inhibition first at the crayfish
neuromuscular junction (Dudel and Kuffler, 1961), and since then in a
number of preparations, notably in the mammalian brain (for review, see McGehee and Role, 1996). The mode of action of presynaptic
GABAA receptors has been best studied in two preparations
where the size of the presynaptic structure is large enough to allow
intracellular recording: the sensory axons in the mammalian spinal cord
(for review, see Rudomin, 1990) and mammalian peptidergic pituitary terminals (Zhang and Jackson, 1993). In both cases, presynaptic inhibition is associated with a depolarization of the presynaptic element. In most brain neurons, however, the size of axons and terminals is very small. Although successful presynaptic recordings have recently been obtained in certain brain preparations [calyx of
Held: Forsythe (1994); cerebellar basket cells: Southan and Robertson
(1998)], such recordings remain difficult, so that alternative approaches to the study of brain autoreceptors would be desirable.
In the present work, we present such an approach, which developed from
the chance observation of an unconventional current response when
recording from cerebellar stellate and basket cells under voltage
clamp. If action potentials are elicited in the axon by applying short
depolarizing pulses in the soma, two distinct bicuculline-sensitive
current signals are obtained during repolarization. One of these
signals corresponds to the activation of somatodendritic autaptic
currents, as reported earlier (Pouzat and Marty, 1998). The second
component is described in the present study. As we shall see, it
corresponds to the activation of GABAergic autoreceptors in the axon of
the recorded cell.
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MATERIALS AND METHODS |
Preparation. Rats were decapitated under deep
anaesthesia (metofane; Mallinckrodt Veterinary, Bray, Ireland), and
parasagittal cerebellar slices were prepared as described (Llano and
Gerschenfeld, 1993). Slices were used 1-5 h after preparation. Two
groups of rats (12- to 15-d-old, and 29- to 49-d-old) were used in the
present study. The data base used to determine the proportion of cells displaying autoreceptor currents or/and autaptic currents overlapped with that used in Pouzat and Marty (1998).
Compositions of intracellular and extracellular recording
solutions. The intracellular recording solution contained (in
mM): 150 KCl, 4.6 MgCl2, 0.1 CaCl2, 1 EGTA, 10 KHEPES, 0.4 NaGTP, and 4 NaATP.
The external solution contained (in mM): 125 NaCl, 2.5 KCl,
2 CaCl2, 1 MgCl2, 1.25 NaH2PO4, 26 NaHCO3,
and 10 glucose. Bicuculline methochloride (10 µM; Tocris
Cookson, Bristol, UK) was used in some experiments to block
GABAA receptors.
Recording from stellate and basket cells. The procedures
described by Llano and Gerschenfeld (1993) were used to identify interneurons in the molecular layer and to record from them. Series resistance values during recording ranged from 10 to 35 M and were
partially compensated (50-75%). Currents were recorded with a sample
interval of 50 or 100 µsec and were filtered at one-fifth of the
sampling frequency. To subtract leak and capacitive currents, a series
of hyperpolarizing voltage pulses were applied from the resting
potential with a fraction of the amplitude of the test depolarizing
pulse, and the resulting currents were added to the response to the
depolarizing pulse.
All experiments were performed at room temperature (20-23°C).
Morphology. Neurobiotin (2 mg/ml) was included to the
pipette solution in some experiments. Slices were then processed after the end of the recordings as explained (Pouzat and Hestrin, 1997). In
other experiments, Lucifer yellow (saturating concentration) was
included in the pipette solution; cells were then examined under
epifluorescence during the experiment, and results were stored using a
CCD camera. Unlike neurobiotin staining, Lucifer yellow staining was
always successful. However, the morphology that was obtained with
Lucifer yellow (e.g., see Fig. 9) was less precise than that obtained
after processing neurobiotin-filled cells (e.g., see Fig. 8).
Numerical methods for simulation of an active cable. We
assume here a uniform cable of length l and radius
a (both in centimeters), voltage-clamped at the origin, with
a sealed end at x = l. This cable has voltage-dependent
Na+ and K+ conductances, as well
as a synaptic conductance, which is either localized or uniformly
distributed. The electric parameters of the cable are
Ri, the cytoplasmic resistivity (in
k·cm); Rm, the resistance of a unitary
surface of membrane (in k·cm2);
Cm, the capacitance of a unitary surface of
membrane (in µF/cm2);
GK, the density of K+
conductance (in mS/cm2); GNa,
the density of Na+ conductance (in
mS/cm2); Gs, the density of
synaptic conductance (in mS/cm2).
EK, ENa,
Es, and EL (all in mV)
are, respectively, the reversal potential of the K+
current, the Na+ current, the synaptic current, and
the leak current. The voltage dependence of the K+
and Na+ currents are modeled following Hodgkin and
Huxley (1952). V(x, t) (in mV) is the membrane potential.
To have a consistent set of units, times are in milliseconds.
With the assumptions outlined above the cable equation takes the form
(Hodgkin and Huxley, 1952; Rall, 1977):
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(1)
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with:
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(2)
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(3)
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(4)
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Here, n, m, h,  n,
 n, m,
m, and h, h
are defined as in Hodgkin and Huxley (1952). The boundary conditions
are:
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(5)
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where Vc(t) is the command
potential applied in the soma, and:
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(6)
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The initial conditions are:
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(7)
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where v(x) is the potential profile at time 0 (usually, v(x) = 70).
The Crank-Nicolson method (Moore et al., 1975; Hines, 1984; Mascagni,
1989) was used to solve numerically this system of equations. "Staggered time grids" (Hines, 1984; Mascagni, 1989) were used to
compute the last three terms in Equation 1. The algorithm was implemented on Igor (Wavemetrics) and run on a Pentium computer (266 MHz). The integration steps, dx and dt were
chosen smaller than one-tenth of the minimal length constant and of the
minimal time constant, respectively. The minimal length and time
constants are computed when the sum of the Na+,
K+, and synaptic conductances is maximum. To obtain
the value of the minimal constants a guess was made, the simulation was
run, and the minimal constants were computed from the stored values of
the activation parameters (n(x, t), m(x, t), and
h(x, t)). If the guess was larger than the computed value,
the simulation was repeated with the computed value as a new guess.
This procedure was repeated until convergence was reached.
The following parameter values were used for the simulations of Figures
10-12: Ri = 0.1 k·cm,
Rm = 50 k·cm2,
Cm = 1 µF/cm2,
GK = 400 mS/cm2,
GNa = 50 mS/cm2,
EL = 70 mV, EK = 95
mV, ENa = +90 mV. The simulations were always
started from 70 mV, so that the initial conditions were: v(x) = 70 mV.
The functions describing the voltage dependence of the rate constants
and in Equations 2-4 were taken from Traub et al. (1994),
except that the reference potential was taken as the bath potential
instead of the resting potential of the cell (60 mV) as in Traub et
al. (1994), and that it was necessary to shift the activation rate
constant of the K+ conductance
(n) by 5 mV to the left to reduce the window of activation of action potentials by the autoreceptor current (see Fig.
11). The synaptic conductance was described as the product of the peak
conductance with a kinetic factor comprised between 0 and 1:
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(8)
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where gs rises linearly from 0 to 1 and
decays with bi-exponential kinetics. The rise time
( rise) was 1.1 or 1.5 msec, and the decay
kinetics were taken from Llano and Gerschenfeld (1993):
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(9)
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Results of numerical calculations were compared with analytical
solutions for the recorded somatic current and the potential along the
cable in response to three types of perturbations. The first one was a
step change of the voltage command. The second perturbation was a
current injection at one position along the cable. The analytical
solutions for these two situations were taken from Rall and Segev
(1985). The last perturbation was a step increase of a distributed
synaptic conductance. The derivation of the analytical solution in this
last case is presented in the Appendix. In all three cases, numerical
calculations reproduced the predictions of the analytical solutions
(data not shown).
Leakage and capacitive current subtraction was simulated for the
calculations of Figures 11 A and 12 B as during
the real experimental protocol. That is, the response to a voltage
pulse from 70 to 80 mV was computed, and the resulting current
trace was multiplied by 5 before being added to the responses to pulses
from 70 to 20 mV.
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RESULTS |
Characterization of a novel bicuculline-sensitive current in
cerebellar interneurons
A delayed current transient associated with
axonal excitability
If somatic depolarizing pulses of increasing duration were applied
from a holding potential of 70 mV in interneurons, a large amplitude,
quickly decaying inward current was observed past a certain threshold,
reflecting the activation of voltage-dependent Na+
channels (Fig. 1A)
(Llano and Gerschenfeld, 1993). Because the site of initiation of
action potentials in central neurons is located in the axon (for
review, see Stuart et al., 1997), it was assumed that this current
reflects the initiation of an action potential in the axon.
Confirmation of this point will be presented below (see Fig. 9). In
many experiments, this early signal was followed after a short time gap
by a delayed inward current transient having an amplitude of up to
several hundreds of picoamperes (Fig. 1A). The
delayed current transient had much slower onset (10-90% rise-time;
1.9 msec in the example shown) and decay (half-decay time, 41 msec)
kinetics than the early Na+ current, as may be seen
in the bottom panel of Figure 1A. Like the
Na+ early transient, it was an all-or-none event.
The activation thresholds for the two components were identical (Fig.
1B; n = 4). If during the course of one
experiment the threshold of the Na+ current
activation changed as a result of an alteration of the series
resistance, the threshold of the delayed current transient changed in
parallel. After application of tetrodotoxin (TTX, 100 nM), the amplitude of the Na+ current
was first gradually reduced, without any marked associated change in
the delayed current transient. Then, the Na+ current
suddenly failed, and the delayed current transient was simultaneously
abolished (results not shown; n = 4). Taken together, the results show that there is a tight coupling between the delayed current transient and the early Na+ current and
therefore indicate that the delayed current transient is a consequence
of an axonal regenerative depolarization.
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Figure 1.
Voltage pulses elicit two distinct inward current
signals with the same threshold. A, Leak-subtracted current
traces obtained in an interneuron in response to successive voltage
pulses (test potential, 20 mV; pulse frequency, 0.5 Hz) of increasing
duration (0.1-1.0 msec, with 0.1 msec increments). For the three
shortest pulses (0.1-0.3 msec, gray traces) no regenerative
current was elicited. For all other stimuli (0.4-1 msec pulses,
black traces), a regenerative Na+ current
was obtained. For these stimuli, a slow, delayed current transient
(mean amplitude, 118 pA) was also obtained. The bottom panel
shows the same set of traces as the top panel, with
different time and current amplitude scales, to illustrate the delayed
current transient; Na+ currents are truncated.
B, Peak amplitudes of (early) Na+ current
(filled symbols) and of delayed current transient
(open symbols) as a function of pulse duration, from three
consecutive runs like the one illustrated in A. Both
components are all-or-none, and they have the same threshold. The
slight decrease of the Na+ current amplitude for
pulse durations  0.5 msec is attributable to the fact that the peak of
the Na+ conductance occurs after the pulses for
shorter durations and during the pulses for longer durations. In the
latter case, the corresponding Na+ current is
reduced because the driving force is less.
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The delayed current transient is mediated by GABAA
receptors and is blocked by extracellular Cd2+
To study the properties of the delayed current transient further,
the protocol shown in Figure 2 was
adopted. The amplitude and duration of the voltage pulse were adjusted
such that axonal Na+ currents were reliably
activated. However, these parameters were maintained in such a range
that somatic K+ current activation was kept at a
minimum. In this way, voltage-dependent currents returned quickly
toward baseline after the offset of the voltage pulse, allowing the
full time course of the delayed current transient to be resolved. Such
voltage pulses were repeated at a frequency of 0.2-1 Hz. The amplitude
of the delayed current transient ran down during the course of
whole-cell recording (Fig. 2A,B), as will be
discussed below. However, this rundown had a relatively slow time
course, so that it was possible to test the action of potential
blockers by exchanging rapidly the bath.
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Figure 2.
The delayed current transient is abolished by
bicuculline and by Cd2+. A, Two
series of five representative current responses to depolarizing somatic
voltage pulses (test potential, 30 mV; pulse duration, 0.8 msec;
pulse frequency, 0.67 Hz), one obtained shortly after breaking into the
whole-cell recording mode (black traces), and the other
obtained after 7 min of whole-cell recording (gray
traces). B, Decay of the peak amplitude of delayed
inward current transients as a function of time in whole-cell
recording, from the same cell as in A. Six points deviate
strongly from the main decay curve: they correspond to traces that were
contaminated with a spontaneous IPSC. C, In another cell,
responses to depolarizing somatic voltage pulses (test potential, 30
mV; pulse duration, 1 msec; pulse frequency, 0.67 Hz), shortly after
breaking into the whole-cell recording mode (black traces),
and after 7 min of whole-cell recording (gray
traces), in the presence of bicuculline. D, Plot of the
peak amplitude of delayed inward current transients as a function of
time, from the same cell as in C. E, F, Same experiment as
in C and D, except that the current was here
challenged by a solution containing 50 µM
Cd2+.
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We have recently reported that depolarizing somatic voltage pulses
elicit autaptic currents in a fraction of stellate and basket cells
(Pouzat and Marty, 1998). Autaptic currents and delayed current
transients are triggered by the same stimulation protocols and are
superficially similar. However, as will become apparent later, they can
be unambiguously distinguished, and they reflect separate cellular
mechanisms. Autaptic currents may be blocked by bicuculline and are
sensitive to the extracellular Ca2+ concentration
(Pouzat and Marty, 1998). Therefore, the effects of bicuculline and of
the nonselective Ca2+ channel blocker
Cd2+ were next tested on the delayed current
transient. If the bath solution was exchanged to one containing 10 µM bicuculline, a rapid decay of the delayed current
transient was observed (Fig. 2C,D; n = 4). After washout of bicuculline, partial recovery was obtained
(data not shown). These results show that the delayed current transient
is mediated by GABAA receptors.
Rapid bath exchange with a solution containing 50 µM
Cd2+ similarly led to an abolition of the delayed
current transient (Fig. 2E,F; n = 4), indicating that Ca2+ entry through
voltage-gated Ca2+ channels was involved. Thus
autaptic currents and delayed current transients share the same
sensitivity to depolarization-induced Ca2+ entry.
The delayed current transient depends linearly on potential
To obtain information on the I-V characteristics of
the delayed current transient, the repolarization potential was altered (Fig. 3A). Over a range
from 110 to 60 mV, the dependence of the peak amplitude of the
transient on repolarization potential was linear (Fig.
3B). From the intersect of the linear fit of this
I-V curve with the voltage axis, values of reversal
potentials comprised between 18 and 0 mV (n = 4;
values corrected for the junction potential arising between the pipette
and the bath solutions) were inferred. The results indicate that the
delayed current transient reflects a conductance increase with a
depolarized reversal potential and are consistent with the notion that
the transient involves the activation of
Cl-selective GABAergic channels. However, because
the pipette solution was loaded with a high Cl
solution, the expected Cl reversal potential
assuming full equilibration is very close to 0 mV, whereas negative
values were usually found for the extrapolated reversal potential.
Furthermore, data obtained with more depolarized repolarization
potentials indicated an upward bend of the I-V curve,
leading to even more negative extrapolated reversal potential values
(results not shown). This deviation from linearity, as well as the
discrepancy between the expected value of ECl
and the value obtained by extrapolation of the I-V curve,
can be explained by the model proposed later in this study (see Fig.
12).
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Figure 3.
The delayed current transient depends linearly on
potential. A, In this experiment the repolarization
potential was changed from 110 to 60 mV. Depolarizing pulse was
20 mV (0.8 msec duration; repetition rate, 0.5 Hz). Traces
were leak-subtracted. B, Plot of the peak amplitude of the
delayed current transient (mean ± SEM) as a function of the
repolarization potential, from four consecutive series such as the one
shown in A. The regression line yields an extrapolated
reversal potential of 13 mV.
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The delayed current transient runs down during
whole-cell recording
The variation of the amplitude of the delayed current transient as
a function of time in whole-cell recording followed several phases
(Figs. 2B,
4C). Directly after the onset
of whole-cell recording, there was a short period of increase with a
modest and variable amplitude ("run up"), which lasted for up to 3 min. We presume that this increase at least in part reflects the
equilibration of the Cl-rich pipette solution with
the cell compartment (the axon, see below) where the GABA receptors are
located. The transient then reached a maximum, and irreversibly
decreased thereafter ("run down"; Fig. 2B). When
examined over a longer period, the run down appeared as biphasic (Fig.
4C). The main part of the current decreased rather abruptly
after 10-30 min of whole-cell recording, leaving a small amplitude
component of up to 20 pA, which was almost totally insensitive to run
down. This component, like the main component, was abolished by
bicuculline (results not shown).
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Figure 4.
The delayed current transient fades during
prolonged whole-cell recording. A, Mean delayed current
transient at three distinct periods of whole-cell recording, showing a
marked decay during the initial 20 min period, followed by a
stabilization at a low level (gray traces).
B, Superimposed normalized traces at 1 and 20 min of
whole-cell recording, showing that the reduction of amplitude with time
is accompanied by a lengthening of the time to peak and a decrease of
the time of decay. C, Kinetics of washout, from the same
data (stimulation frequency, 0.25 Hz; series resistance during
recording, 14 M). D, E, Relation of time to peak
(D, measured from the onset of the voltage pulse) and half
decay time (E) on peak current amplitude. Data are from four
cells as the one illustrated in A and B, and from
a fifth cell in which the current was inhibited by addition of
bicuculline. Each data point corresponds to averages of ~10-20
traces with homogeneous amplitudes. Results for individual cells are
indicated by specific symbols and have been fitted by individual
regression lines. Note that the slopes of the regression lines are all
negative in D and positive in E.
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Changes in kinetics associated with a reduction of the delayed
current transient
A characteristic kinetic change occurred during washout, as became
apparent after scaling up the reduced current (Fig.
4B). As the peak amplitude was reduced, the rise time
was increased and the decay time was decreased. Summary data from five
cells including that illustrated in Figure 4B are
presented in Figure 4D,E. Four of
the data sets represent washout experiments, and the fifth plots the
results of a cell where the current was reduced by addition of
bicuculline. In each case the regression line through the data
indicates a positive correlation between amplitude and decay time, and
a negative one between amplitude and time to peak. Similar trends were
obtained in other experiments with either bicuculline or
Cd2+ (results not shown). Thus the correlations
apparent in Figure 4D,E apply regardless of
whether the delayed current transient was reduced as a result of
rundown or after addition of bicuculline or
Cd2+.
The delayed current transient is blocked by dialyzing the cell with
10 mM BAPTA
When an intracellular solution including 10 mM BAPTA
was used instead of the usual 1 mM EGTA, the relation of
the delayed current transient amplitude on whole-cell recording time
changed drastically (Fig. 5). There was
no run up, and the maximum current amplitude was less than in control
conditions. The kinetics of rundown was monophasic and markedly
accelerated with respect to control experiments. These results show
that the delayed current transient is sensitive to the
Ca2+ homeostasis of the recorded cell.
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Figure 5.
The delayed current transient is blocked by
intracellular BAPTA. A, Mean delayed current transient (6 consecutive traces each; stimulation frequency, 0.25 Hz) recorded
shortly after break-in (labeled 30 s) and after
approximately 8 min of whole-cell recording. The pipette solution
contained 10 mM BAPTA instead of the usual 1 mM
EGTA. B, Time course of decay of the peak amplitude; series
resistance during recording, 24 M. C, D, Summary results
for six control cells and six BAPTA-dialyzed cells, showing values
(mean ± SEM) for the maximum peak delayed current transient
(obtained during the first minutes of whole-cell recording) and for the
time to half decay of this current. Average series resistance values
were 24 ± 5 M for the control cells and 36 ± 6 M for
the BAPTA-dialyzed cells.
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The rise-time and decay of the delayed current transient are longer
than those of IPSCs
Spontaneous IPSCs recorded in stellate or basket cells have sharp
rising phases, with a mean time to peak of 1.1 msec at room temperature
(Llano and Gerschenfeld, 1993). In each cell where measurements of the
delayed current transient were performed, the slow rise of the delayed
current transient contrasted strikingly with the fast rise-times of the
spontaneous IPSCs (Fig.
6A). In addition, decay
times were systematically slower for the delayed current transient than
for IPSCs (Fig. 6A3). We considered the possibility
that the conductances accompanying the axonal action potential might
decrease the quality of the voltage clamp, leading to a distortion of
the delayed current transient. This seemed unlikely because autaptic
currents, when observed, have fast rise-times (Pouzat and Marty, 1998),
although they are triggered by the same stimulations as delayed current
transients, and furthermore because spontaneous IPSCs with fast
rise-times are occasionally superimposed with the delayed current
transient (data not shown). Nevertheless, the possibility of a
distortion through the stimulated action potential was examined in a
series of experiments in which extracellularly evoked IPSCs and delayed
current transients were elicited in the same cell with variable
intervals. When the stimuli were timed such that the extracellularly
evoked IPSCs occurred near the onset of the delayed current transients,
the IPSCs retained their fast rise-times (Fig. 6B). These
results suggest that the slow kinetics of the delayed current transient
is not simply derived from a deterioration of the voltage clamp
after the axonal action potential.
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Figure 6.
The delayed current transient has slow onset and
decay kinetics. A1, A2, Examples of superimposed traces of
delayed current transient (A1) and of spontaneous IPSCs
taken in the same recording (A2). A3, Comparison between
scaled averages of the autoreceptor current (continuous
line, average of 142 sweeps) and of the spontaneous IPSCs
(dotted line, average of 150 IPSCs; peak amplitude before
scaling: 210 pA). B1, Examples of extracellularly evoked
IPSCs, from another cell. B2, Here extracellular
stimulations (at the time indicated by the star) were
combined with somatic depolarizations. B3, Comparison of
single evoked IPSCs obtained without (black trace) or with
(gray trace) somatic depolarization. Rise-times are
rapid in both cases. B4, A further series of evoked IPSCs,
obtained with another location of the stimulation pipette. Here the
evoked IPSCs are seen to superimpose on slowly rising delayed current
transients.
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The delayed current transient is developmentally regulated
It was reported recently that activation of AMPA receptors
increases the rate of miniature IPSCs in mouse cerebellar stellate and
basket cells, indicating the presence of axonal ionotropic glutamate
receptors in this preparation (Bureau and Mulle, 1998). This effect
wanes during development, suggesting a downregulation of presynaptic
AMPA receptors. In view of these results, we compared the properties of
the GABAergic delayed current transient during two age periods: 12-15
d and 29-49 d. Results presented so far correspond to the first of
these periods. For this age group, the delayed current transient was
consistently observed (92% of the recordings, n = 41;
estimated threshold for detection: 10 pA). When observed, the delayed
current transient had a mean amplitude of 82 ± 17 pA. By
contrast, delayed current transients in the older age group were
observed much more sparingly, with an incidence rate of 14%
(n = 35). When observed, they were of smaller
amplitude, with a mean amplitude of only 15 ± 3 pA (Fig.
7B1).
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Figure 7.
The delayed current transient is associated with
enhanced axonal excitability. A1, Superimposed exemplar
traces of delayed current transient; regenerative responses are present
in three traces out of six. No such responses were seen in the presence
of bicuculline (thick line; average trace). A2,
Three further traces from the same experiment, shown with a less
expanded vertical scale. Two of the traces contained large amplitude
regenerative responses. B1, B2, Similar results obtained
from an experiment with an older animal (27-d-old instead of 15-d-old
in A1, A2). In this case the amplitude of the delayed
current transient is small (~30 pA), but regenerative responses are
nevertheless observed before application of bicuculline.
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Autapses were observed in the same series of experiments. As reported
earlier (Pouzat and Marty, 1998), autapses are rather frequent in the
group 26-49 d, with an incidence rate of 26%. In two cells from this
group (6% of the cases), autaptic currents and autoreceptor currents
were superimposed. At age 12-15 d, autapses occurred in only 5% of
the recordings (2/42 cells). In these two cases the probability of
failure of the autaptic currents was very low, as expected from a
previous study on the evolution of the probability of release at the
interneuron-Purkinje cell synapse (Pouzat and Hestrin, 1997). Because
of the high activity of autaptic currents, it could not be decided in
these two cells whether an autoreceptor current was also present.
Active responses associated with the delayed current transient
In some experiments, fast inward current signals were superimposed
on the early part of the decay phase of the delayed current transient
(Fig. 7). These signals occurred often with two different amplitudes,
one of the order of a few tens of picoamperes, and the other on the
order of 1 nA (Fig. 7 A1,A2). The time course and
amplitude of the larger signals were similar to those of the Na+ current transients directly associated to the
voltage pulse, suggesting that the delayed current transient induced
voltage-clamp escape in the axon. The smaller events appeared earlier
than the larger ones (Fig. 7A2,B2), and occasionally
a transition was observed from the smaller to the larger level (Fig.
7B2, arrow). Although in most recordings each
delayed current transient produced maximally a single spike, in one
experiment up to three spikes were obtained, with regular interspike
intervals of ~10 msec (data not shown).
Such regenerative signals were found in both of the age groups studied.
However, they were more prominent in the older age group, where they
occurred in 60% of the recordings displaying delayed current
transients (3/5 cells), compared with a proportion of 8% in the
younger age group (3/36 cells). Furthermore, in accordance with the
general difference in peak amplitudes between the two groups, spikes
were associated with delayed current transients of a few tens of
picoamperes in the older age group, as opposed to 100 pA or more in the
younger age group (compare Fig. 7, A1 and B1;
note vertical scale difference). In all recordings where delayed
current transients elicited spikes, spontaneous IPSCs (occurring at
ordinary synapses) with amplitudes similar to or larger than that of
the delayed current transient were present; none of these IPSCs
elicited active currents. These observations show that the active
currents are not caused by voltage escape resulting from current flow
across the series resistance to the cell, and that they are
specifically associated with the delayed current transient.
After application of bicuculline, the secondary axonal spikes were
abolished in parallel with the delayed current transients (Fig.
7A1,B1).
All of these observations indicate that the delayed current transient
elicits voltage-clamp escape in the axon and that this occurs more
readily in the older than in the younger age group.
Results with neurobiotin-filled cells
In a series of experiments, neurobiotin was included in the
pipette solution, and the slices were processed with the hope of
recovering the morphology of the cells. However, the proportion of
interneurons that gave good stainings was low (<10%), whereas parallel staining of Purkinje cells was almost always successful. Success rates were particularly low if the intracellular solution was
rich in Cl ions, as required for the measurement
of autaptic currents and delayed current transients. As a result, only
three cells yielded electrophysiological recordings together with
satisfactory staining. One of these cells had an autaptic current, and
the two others had only an autoreceptor current. Two of the cells,
including the cell with autaptic currents, had regions of close
apposition between axonal and dendritic domains, with potential
autaptic sites. The third cell is illustrated in Figure
8. There is clearly no autaptic site in
this cell, and yet a substantial delayed current transient was
measured. This demonstrates that the delayed transient current does not
require an autapse in the recorded cell. In addition, the axonal and
dendritic domains are largely distinct in the cell of Figure 8; this
also applied to other cases in which large delayed transient currents
were recorded (Fig. 9). One branch of the
axon does appear to overlap with the dendritic tree in Figure
8A, but close inspection reveals that this branch was
above the plane of the dendrites and remained at a distance of 2 µm
of the nearest dendrite. At such distances dilution by diffusion from a
point source is expected to reduce the local GABA concentration below the threshold for activation of GABAA receptors (Clements,
1996; Barbour and Häusser, 1997). This evidence suggests that the
extent of overlap between axonal and dendritic domains of interneurons is quite limited and that one cannot account for the current on the
basis of spillover from the axonal to the dendritic region.
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Figure 8.
The delayed current transient does not require
close contact or extensive overlap between the axonal and dendritic
domains. Top, Camera lucida drawing of a basket cell
(14-d-old animal) that was filled with neurobiotin during recording and
processed for histology. The short thick neurites radiating from the
soma are dendrites. The axon leaves the soma on the right and takes an
almost straight course, sending several collaterals along the way. It
is prolonged beyond the boundary of the drawing. The Purkinje cell
layer is roughly horizontal and is located at the level of the tips of
the axon collaterals at the lower boundary of the cell.
Middle, Composite photograph from the same cell. In the
lower scale photograph the dendrites are in focus; the axon collateral
that appears to overlap with the dendritic field in the drawing of the
top panel is then not in focus. Insert, Blowup of
the somatodendritic region with the axon collateral in focus. The
shortest distance between axonal and dendritic domains in this cell was
2 µm. Bottom, Average responses to depolarizing pulses
from the same cell, with and without bicuculline, showing the presence
of a normal size, delayed current transient.
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Figure 9.
The delayed current transient can be obtained by
extracellular stimulation. A, Morphology of the recorded
interneuron (a basket cell) with Lucifer yellow staining. The Purkinje
cell layer (PCL) corresponds roughly to the lower border of
the field. The location of the stimulation pipette was alternated
between a position 160 µm left of the soma of the recorded cell
(Site 1) and another position 160 µm right of the soma of
the recorded cell (Site 2; data not shown). B,
Current traces obtained for different stimulation conditions. Series of
pulses of 0.2 msec in duration and of various amplitudes were delivered
with a frequency of 0.25 Hz at each site. Between each series of
pulses, responses to somatic depolarizations were recorded.
B1, A 600 µA stimulation at Site 1 failed to elicit any
response. Only the stimulation artifact is obtained. B2,
Increasing the stimulation intensity to 700 µA resulted in a large
Na+ current (which just saturated the recording
amplifier at 2 nA; data not shown) and a 90 pA delayed current
transient. B3, Response to somatic stimulation.
Na+ current amplitude = 1.75 nA (data not
shown). Note the similarity of the amplitude and time course of the
delayed current transient with that obtained by direct axon
stimulation. B4, Lack of response at Site 2, with a
stimulation intensity largely beyond the threshold at Site 1.
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Extracellular stimulation of the delayed current transient
It was reasoned that if the delayed current transient was caused
by activation of the axon, it should be obtainable by suitable extracellular stimulation in the axon domain. Interneurons tend to send
their axon in the parasagittal plane, along an axis parallel to the
Purkinje cell layer. However, the direction of the course of the main
axon of any given cell along the Purkinje cell layer cannot be
predicted. After establishing a whole-cell recording on a target cell,
extracellular stimulations were therefore applied in alternation on one
side and on the other. At both locations, the stimulation pipette was
positioned in the proximal part of the molecular layer, where basket
cell axons are normally found, at a distance of ~150 µm from the
soma. It was found that although no response was obtained on one side,
an action potential could be elicited from the other side beyond a well
defined threshold (Fig. 9B1,B2). This was monitored
under somatic voltage clamp as a rapid inward current transient
resembling that obtained with somatic stimulation. After the rapid
inward current, a delayed current transient very similar to that
obtained with somatic stimulation was also obtained (Fig. 9, compare
B2 and B3). This signal decreased during
prolonged whole-cell recording in parallel to the somatically evoked
delayed current transient. When stimulating on the other side of the
soma, neither an action potential nor a delayed current transient was
obtained, even if the stimulation intensity was increased to three
times the threshold value on the responsive side (Fig. 9B4;
n = 3). For these experiments, Lucifer yellow was included in
the pipette solution, and the morphology of the recorded neuron was
determined by epifluorescence after completion of the measurements. The
axon was found on the side where stimulations had been successful (Fig.
9A; n = 3).
These experiments show that direct stimulation of the axon under steady
somatic voltage clamp, like somatic depolarization, elicits the same
sequence of fast Na+ current and delayed current
transient. Under the conditions of Figure 9, somatic voltage clamp
uncouples the dendrites from the axon, effectively preventing
activation of somatodendritic voltage-dependent channels. Therefore,
the Na+ channels activated in Figure 9B2
are axonal rather than somatodendritic. The similarity with the results
obtained when depolarizing the soma confirms that the fast
Na+ current elicited on somatic stimulation reflects
the activation of axonal Na+ channels and that the
delayed current transient requires firing of an action potential in the axon.
Simulations
This section presents simulations of somatic currents elicited in
a voltage-clamped neuron by activation of synaptic conductances located
either in dendrites or in the axon. In performing these simulations,
particular attention was given to the following features of the delayed
current transient: (1) slow onset and decay kinetics compared with
that of spontaneous IPSCs; (2) link between peak amplitude and
kinetics, whereby large events have short rise-times and slow decay
times, whereas small events have long rise-times and short decay times;
(3) negative extrapolated reversal potential; and (4) presence of
spikes on the decay phase in some experiments.
Dendritic receptors
Feature 1 suggests at first that the delayed current transient
could be a filtered version of the autaptic currents reported previously (Pouzat and Marty, 1998) and that they could arise from
axodendritic contacts of the interneuron onto itself. Morphological data (Llano et al., 1997, their Fig. 5; Pouzat and Hestrin, 1997, their
Fig. 1A2, B5) suggest that autaptic contacts are highly localized.
Consequently, simulations were performed with one localized input along
an otherwise uniform cable. The length and radius of the cable were set
at 50 and 0.4 µm, respectively, corresponding to the upper and lower
bounds of values for interneuron dendrites at the age 12-15 d (Llano
et al., 1997, their Fig. 5; Pouzat and Hestrin, 1997, their Fig.
1A2; this work, Figs. 8, 9). Such a cable therefore should
represent maximum dendritic filtering in this preparation. The passive
electrical parameters of the dendrites were as given in Materials and
Methods. No active conductance was included. Synaptic inputs with a
reversal potential at 0 mV were positioned at various locations along
the dendrite. The synaptic conductance had a rise-time of 1.1 msec
(Llano and Gerschenfeld, 1993) and bi-exponential decay kinetics (Eq. 9). Figure 10 shows the results of
such simulations for an input located close to the soma (A)
and for an input located near the tip of the dendrite (B).
Each graph presents several traces corresponding to various peak
conductances. The insets show scaled versions of the largest (continuous line) and smallest (dashed line)
current. In all traces, including those generated at the distal site,
current kinetics are almost identical to those of the underlying
conductance and do not depend significantly on current amplitude. In
particular, no change in rise-time is observed. These results strongly
argue against dendritic filtering as the explanation of the slow
kinetics of the delayed current transient. Nevertheless, we considered the possibility that the cytosol of interneurons may have an unusually low conductivity, thus leading to more severe filtering than predicted in Figure 10A,B. Therefore, additional
simulations were performed assuming larger values of
Ri (0.25 and 0.5 k·cm instead of 0.1 k·cm). With these values of Ri, a
significant attenuation of the peak amplitude was observed for the
largest currents at the distal input and was associated with a slower
decay time course. However, even in these extreme conditions, no
significant increase in rise-time was observed (data not shown).
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Figure 10.
Currents recorded in response to synaptic
conductance changes in passive cables. A, B,
Dendrite-like cable voltage-clamped at the origin with localized
synaptic inputs. The cable is uniform, with a length of 50 µm and a
radius of 0.4 µm. Its electrical parameters are
Ri = 100 ·cm, Rm = 50 k·cm2, Cm = 1 µF/cm2. The synaptic conductance has a rise time
of 1.1 msec and a bi-exponential decay with fast = 9 msec, slow = 40 msec, and a weight of 0.6 for the fast
component. Six different peak conductances are simulated: 0.5, 1.0, 1.5, 2.0, 2.5, and 3 nS. The reversal potential of the synaptic current
is at 0 mV. The origin of the cable is clamped at 70 mV. The
conductance is located 10 µm away from the origin in A and
48 µm away in B. Inset, Scaled largest current
(continuous line) and scaled smallest current (dashed
line); Calibration, 5 msec. C, Axon-like cable
voltage-clamped at the origin with homogeneous synaptic input. The
cable is 200 µm long and has a radius of 0.25 µm. Its electrical
parameters are the same as in A and B. The
synaptic conductance has a rise time of 1.5 msec and the same
bi-exponential decay as above. Six different autoreceptor conductance
densities are simulated: 0.25, 0.64, 0.89, 1.27, 1.91, and 2.55 mS/cm2. As above, the reversal potential of the
synaptic conductance is 0 mV, and the origin is maintained at 70 mV.
As the autoreceptor current density increases, the time to peak
decreases from 3.5 to 2.0 msec, whereas the half decay time increases
from 14.2 to 21.8 msec. Inset, As above, scaled largest
(continuous line) and smallest (dashed line)
currents. Calibration, 5 msec.
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We conclude from these simulations that cable filtering generated at
interneuron dendrites cannot account for the slow rise of the delayed
current transient.
Distributed axonal receptors without active conductances
Having rejected a dendritic site of origin for the delayed current
transient, we performed next a series of simulations on the assumption
that it is generated in the axon. This hypothesis is immediately
compatible with the association of axonal spikes with the delayed
current transient (feature 4). The possibility was considered that the
delayed current transient could be generated at axo-axonic autapses.
However, such autapses have not been observed morphologically in any
preparation. Furthermore, examination of the available data at
interneurons (Llano et al., 1997, their Fig. 5; C. Pouzat, unpublished
results) fails to reveal any apposition between distinct regions of the
axonal tree and thus does not support this possibility. Finally,
axo-axonic autapses would be expected to yield widely fluctuating
currents including failures, whereas the delayed transient current
varied little from trial to trial. Overall, it seemed highly unlikely
that the delayed current transient would be caused by axo-axonic
contacts. We next considered the possibility that this current would
reflect the activation of autoreceptors along the axon, after the
activation of many release sites. We therefore performed simulations of
an axon-like cable with a length of 200 µm, a radius of 0.25 µm, and a uniformly distributed autoreceptor conductance. Passive cable
properties were as described in Materials and Methods. Autoreceptor conductance densities were empirically set at 0.25, 0.64, 0.89, 1.27, 1.91, and 2.55 mS/cm2 to cover the range of
experimentally observed amplitudes of the delayed current transient.
(These numbers correspond to conductances of 0.2, 0.5, 0.7, 1, 1.5, and
2 × 1010 S per site, assuming one site every
5 µm; see below.) The rise-time of the conductance was somewhat
increased with respect to the previous simulation (1.5 msec instead of
1.1 msec) to take into account the jitter expected among release sites
attributable to the propagation of the action potential. (The
propagation time of 0.4 msec was taken in view of the results of Fig.
11B,C.) Decay kinetics were the same as above. A first set of simulations was run
without active conductances (Fig. 10C). Simulated currents have many of the qualitative features of the delayed current
transients. Their kinetics are slower than those of spontaneous IPSCs,
as may be seen in Figure 10 (compare traces in C with those
in A and B) and are within the range of the
observed kinetics of the delayed current transient (Fig.
4D,E). The simulations display correlations between amplitude and rise-time on the one hand, and between amplitude and decay time on the other, which match those observed experimentally (compare Fig. 10C, inset with Fig. 4B).
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Figure 11.
Axon-like cable with homogeneous autoreceptor
current input. Simulations of a cable 200 µm long with a radius of
0.25 µm. The parameters describing the passive electrical properties
of the cable are as in Figure 10. The cable has a uniform
Na+ conductance density (50 mS/cm2 with a reversal potential at +90 mV) and a
uniform K+ conductance density (400 mS/cm2 with a reversal potential at 95 mV). The
kinetic parameters of these conductances are given in Materials and
Methods. Results of simulations are displayed with different uniform
autoreceptor conductance densities. The kinetic parameters and reversal
potential of the autoreceptor conductance are the same as in Figure 10.
A, Currents elicited by a short voltage pulse (duration, 0.5 msec; amplitude, 50 mV) at the origin of the cable. Traces are
"leak-subtracted" (see Materials and Methods). The autoreceptor
conductance is activated 1 msec after the beginning of the pulse.
Autoreceptor densities of 0.64 mS/cm2 (dashed
trace), 1.27 mS/cm2 (continuous
trace with the two secondary spikes), 2.55 mS/cm2 (gray trace), and 5.1 mS/cm2 (continuous trace without
secondary spikes) are represented. B, C, Voltage time
courses at four different locations along the cable with two different
autoreceptor conductance densities: 1.27 mS/cm2 for
B and 5.1 mS/cm2 for C. Here
only 10 msec are shown. The first spike is the one evoked by the
voltage pulse at the origin. The second, antidromic spike in
B corresponds to the first of the two secondary spikes in
A. D, Peak-scaled traces corresponding to autoreceptor
conductance density values of 0.64 mS/cm2
(dashed line) and 5.1 mS/cm2
(continuous line). Times to peak are, respectively, 8.4 and
3.6 msec, and half-decay times are 25 and 28 msec. Calibration, 5 msec.
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To obtain insight into the factors that determine the characteristics
of the delayed current transient, an analytical solution was derived
for a simplified but closely related situation: a semi-infinite cable
with an instantaneously rising distributed conductance (Appendix A).
The analytical solution shows that for large synaptic conductance
densities, the peak current grows like the square root of the synaptic
conductance density (Eq. A.20) and furthermore that the rise-time is
proportional to the inverse of the square of the peak current amplitude
(Eq. A.22).
Other simulations were performed with different cable lengths ranging
from 100 to 600 µm (data not shown). In general, the results were
almost totally independent of the cable length over this range, except
that with the shortest cable length (100 µm), peak amplitudes for
small synaptic current densities were somewhat reduced, and decay
kinetics were accelerated. This acceleration was the same for all
autoreceptor conductance densities, so that the correlation between
amplitude and kinetics was left unchanged. Consistent with these
simulations, an analytical solution for a finite cable submitted to a
step increase in distributed conductance predicts that the amplitude of
the recorded current saturates as a function of cable length (Appendix
B). These results indicate that somatic recordings gather information
from the proximal end of the cable only, essentially from the first 100 µm. Therefore, attaching axon collaterals to the main axon should
have no major consequence on the predicted currents. Other simulations
were performed changing the values of Ri (range,
0.1-0.25 k·cm) and Rm (range, 50-100
k·cm2): the qualitative features apparent in
Figure 10C were maintained throughout this range of parameters.
Distributed axonal receptors with active conductances:
secondary spikes
In the next series of simulations, voltage-dependent
Na+ and K+ conductances were
included. Because quantitative information on axonal voltage-dependent
conductances in this preparation is almost totally lacking, the choice
of the parameters describing these conductances was largely arbitrary.
We chose a set of parameters derived from simulations performed by
Traub et al. (1994) for the axonal membrane of hippocampal pyramidal
cells, as detailed in Materials and Methods. The chief goal of these
simulations was to see whether the features apparent in Figure
10C would be conserved with a more realistic set of axonal
conductances, and also whether features 3 and 4 of the delayed current
transient could be duplicated with reasonable parameters for
Na+ and K+ conductances. Again,
the cable length was set at 200 µm and its radius at 0.25 µm. The
responses of such a cable to a voltage pulse having a duration of 0.5 msec and an amplitude of 50 mV were computed. The synaptic conductance
was triggered 1 msec after the beginning of the voltage pulse, and its
kinetic parameters were the same as in Figure 10C.
Figure 11A shows the computed currents for a series
of autoreceptor conductance densities. It is immediately apparent that the model accounts for the presence of spikes on the decay phase of the
current (feature 4). The presence of spikes was critically dependent on
the properties of voltage-dependent conductances as well as on the
autoreceptor conductance density. The interplay between
voltage-dependent and GABAergic conductances may be appreciated by
examination of the time course of the voltage at different locations
along the cable (here 50, 100, 150, and 199 µm from the soma; Fig.
11B,C). Figure 11B illustrates such voltage
profiles for an autoreceptor conductance density value (1.27 mS/cm2) within the critical range where secondary
spikes occur during the decay of the delayed current transient. The
secondary spike, which is triggered by the autoreceptor conductance, is
generated at the end of the cable and is therefore an antidromic spike. Figure 11C shows similar voltage profiles for a higher
density of axonal GABAergic channels. Comparison between B
and C reveals the reasons why some synaptic conductance
densities give rise to secondary spikes whereas others do not. If the
density is too large, it maintains the membrane potential too
depolarized to allow Na+ conductance to recover from
inactivation, as exemplified in Figure 11C. If it is too
small, the depolarizing autoreceptor current is not strong enough to
bring the membrane to threshold. In Figure 11, only one autoreceptor
conductance density (1.27 mS/cm2) gave rise to
spikes, whereas smaller and larger densities did not. Increasing the
Na+ conductance density (to 100 or 200 mS/cm2) and/or reducing the K+
conductance density (to 200 mS/cm2) increased the
range of synaptic conductance densities that led to spike generation
(down to 0.75 and up to 2 mS/cm2). Slowing down the
kinetics of the voltage-dependent conductances (by a factor of 2) had
the same consequence (data not shown). Changing the length of the cable
(range, 100-600 µm) did not substantially alter the results (data
not shown).
It should be noted that in the simulations, secondary spikes were not
assumed to generate any autoreceptor conductance increase. In reality,
however, a secondary spike would be followed by its own autoreceptor current.
Distributed axonal receptors with active conductances: amplitude
and kinetics of the delayed current transient
Figure 11D shows a peak scaling of two of the
currents of Figure 11A. The continuous line is from
the autoreceptor conductance density of 5.1 mS/cm2,
and the dashed line is from the autoreceptor conductance density of
0.64 mS/cm2. It may be seen that the higher peak
amplitude gives rise to faster onset and slower decay. Thus the model
reproduces the correlation between peak amplitude and kinetics that is
observed experimentally (Fig. 4).
Figure 12 illustrates the effects of
active conductances on the measured delayed current transient. In
Figure 12A, currents corresponding to a conductance
of 0.64 mS/cm2 are compared without (dashed
line) and with (thick continuous line) active
conductances. Inclusion of voltage-dependent channels results in a
reduction of the peak amplitude of the delayed current transient and in
a lengthening of its rise-time.
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Figure 12.
Effects of active conductances on the kinetics
and peak amplitude of the delayed current transients. A,
Superimposed traces representing simulations of the delayed current
transient for a 200 µm long homogeneous cable containing an
autoreceptor conductance density of 0.64 mS/cm2,
without (dashed trace, from Fig. 10C) and with
(thin continuous trace, from Fig. 11A)
voltage-dependent conductances. The thick trace represents
the subtraction of the currents obtained with and without
autoreceptors, in the presence of voltage-dependent conductances.
B, Peak delayed current transient as a function of
autoreceptor conductance density, as calculated with the analytical
model of a semi-infinite passive cable (dashed line), with
the numerical simulation of a passive cable (triangles, from
Fig. 10C) and with the numerical simulation of an active
cable (circles, from Fig. 11A). C, Simulations of
the experiments performed to estimate the reversal potential of the
delayed current transient. Top part, schematic
representation of the voltage command applied at the origin of the
cable. Bottom part, current traces obtained with an
autoreceptor conductance density of 5.1 mS/cm2
(these traces are leak-subtracted). D, Plot of the peak
delayed current transient versus repolarization potential. The
extrapolated reversal potential is 33 mV.
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Figure 12B compares the peak delayed current
transient values obtained from the analytical solution of the infinite
passive cable (dashed line, from Eq. A.17), from the passive cable
simulation (triangles, from Fig. 10C), and from
the active cable simulation (circles, from Fig.
11A). At any density of autoreceptor currents, addition of voltage-dependent conductances results in a strong reduction of the peak amplitude of the delayed current transient.
Finally, Figure 12C,D is a simulation of the
experiments used to evaluate the reversal potential of the delayed
current transient. The reduction of the apparent peak amplitude of the
delayed current transient increases with depolarization because of the
voltage dependence of the K+ conductance, leading to
an apparent negative reversal potential (Fig. 12D).
The I-V curve nevertheless remains approximately linear in
the voltage range used in the tail experiments (110 to 60 mV; Fig.
12D). The deviation is actually larger in Figure
12D than experimentally observed, probably because
the K+ conductance assumed in the simulation is
larger than in reality.
An estimate of the contribution of each individual release site to
the total autoreceptor current
The simulations presented in Figure 12A
(circles) indicate that a somatically recorded delayed
current transient of 80 pA (the average value at age 12-15 d)
corresponds to an axonal conductance of 1.4 mS/cm2.
This is not an accurate estimate because it depends on specific assumptions concerning the voltage-dependent K+
conductances in the axon, which are poorly characterized. Nevertheless, this estimate is rather consistent when changes of the
K+ current characteristics are performed within the
frame of parameters compatible with the known properties of interneuron
currents (size of somatically measured K+ currents,
kinetic properties of the delayed current transient, etc.), so that it
provides a useful indication of the order of magnitude of the
autoreceptor current.
Because of the steep dependence of the concentration on distance during
diffusion from a point source, and of the relatively low affinity of
GABAA receptors for GABA, the transmitter that is released
at one varicosity is unlikely to activate GABAA receptors that are much beyond the limits of the synaptic cleft (Clements, 1996;
Barbour and Häusser, 1997). For this reason, the currents that
are measured most likely reflect the activation of receptors that
belong to the presynaptic membrane of each varicosity. Assuming an axon
radius of 0.25 µm and a spacing between varicosities of 5 µm
(Pouzat and Kondo, 1996), the above current density translates to a
mean current value (at 70 mV) of ~10 pA per varicosity.
Because the simulation performed in Figures 10-12 assumes an axon 200 µm in length, the same current density corresponds to a total maximum
current (assuming a driving force of 70 mV, and a homogeneous
distribution of one varicosity every 5 µm of axon length) of 400 pA,
five times larger than the value of the delayed current transient. This
estimate needs to be corrected upward to account for the presence of
side branches in the axonal tree (Bishop, 1993; Pouzat and Kondo,
1996), leading to a corrected value on the order of 1-2 nA.
It is instructive to compare the value of 10 pA per varicosity with an
estimate of the postsynaptic current obtained under symmetrical
Cl conditions. The average quantal size obtained
at single-site interneuron-interneuron synapses is 61 pA, and the
corresponding release probability is 0.28 [Kondo and Marty (1998); the
latter value is derived from their Table 1]. Thus one release site
gives on average a postsynaptic current of 17 pA per action potential, a value only slightly larger than the estimate for the presynaptic autoreceptor current. These estimates suggest that the number of
presynaptic receptors may be only slightly smaller than that of
postsynaptic receptors in the present preparation.
Although the estimated density of 1.4 mS/cm2 is much
smaller than the values assumed for axonal Na+ and
K+ conductances, it may have a significant effect on
axonal excitability. In pituitary gland nerve terminals, the density of
presynaptic GABAA conductance was estimated (under
symmetrical Cl conditions) at only 0.38 mS/cm2, yet this density was shown to significantly
inhibit action potential firing by interfering with
Na+ current inactivation (Zhang and Jackson, 1995a;
Jackson and Zhang, 1995).
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DISCUSSION |
The delayed current transient: a Ca2+-dependent
GABAergic current with unconventional properties
The delayed current transient is blocked by bicuculline,
indicating that it is mediated by GABAA receptors. It is
blocked by extracellular Cd2+, indicating that it
depends on voltage-dependent Ca2+ entry. On the
basis of these criteria, the delayed current transient is similar to
the many examples of GABAergic IPSCs that have been described in CNS
preparations. However, in other respects the delayed current transient
has quite unusual properties. It is blocked by perfusion of the
recorded cell with BAPTA, suggesting that the release of GABA takes
place in the very cell in which currents are recorded. It also has
remarkable kinetic properties: it displays a slow rise-time, suggesting
attenuation through cable filtering, and a unique correlation between
peak amplitude and the kinetics of both onset and decay. Finally, its
fluctuations from one trial to the next are much smaller than for
ordinary GABAergic IPSCs of comparable mean amplitude. We will next
examine various possible interpretations of the delayed current
transient in the light of these observations.
The delayed current transient is not an autaptic current
We demonstrated recently the presence of functional autapses in
cerebellar stellate and basket cells by using essentially the same
experimental protocol as that of the present study (Pouzat and Marty,
1998). Like the delayed current transient, autaptic currents are
triggered by an axonal spike, they are blocked by bicuculline, and they
depend on voltage-dependent Ca2+ entry (Pouzat and
Marty, 1998). However, autaptic currents in interneurons have sharply
rising onsets that contrast with the slow rise of the delayed current
transient. Furthermore, autaptic currents fluctuate widely from one
trial to the next. Quantal sizes ranging from 21 to 178 pA (symmetrical
Cl conditions) have been estimated from these
fluctuations, and failure rates average 80%. In all these respects,
autaptic currents resemble ordinary interneuron-interneuron IPSCs, but
differ markedly from the delayed current transients described here. A
further difference concerns developmental regulation: whereas the rate of occurrence of autaptic currents grows during development, the delayed current transient is much less prominent in adult animals than
at 2 weeks.
This series of differences between delayed current transient and
autaptic currents suggests that the underlying cellular mechanisms are distinct.
The delayed current transient is not mediated by electrical
junctions among interneurons
It is conceivable that delayed current transients could reflect
the detection of IPSCs elicited in neurons that would be postsynaptic to the recorded cell if presynaptic and postsynaptic neurons were linked by electrical junctions. In previous publications, we have investigated the possible presence of electrical junctions among interneurons. Although there is some evidence for the presence of
electrical junctions, these junctions are rare and when present are
associated with a small conductance [on the order of 1 nS at most; see
Vincent and Marty (1996); Pouzat and Marty (1998)]. The rare
occurrence of electrical junctions, compared with the very high rate of
occurrence of delayed current transients, make it unlikely that one
could be the source of the other. Second, the conductance of electrical
junctions, when present, is much too low to ensure action potential
transfer from one cell to the next [as discussed in Pouzat and Marty
(1998)]. Third, if delayed current transients were caused by the
detection of IPSCs elicited in neighboring postsynaptic interneurons
through gap junctions, the high variability and high failure rate of
interneuron-interneuron IPSCs (Kondo and Marty, 1998) would be
expected to yield a corresponding high variation in the delayed current
transients including occasional failures. This contrasts with the low
variability that is experimentally observed for delayed current
transients. Finally, if delayed current transients were generated by
IPSCs in neighboring cells, they should occur randomly because of the
ongoing firing of interneurons. However, no delayed current transient
was ever observed between stimulations. Altogether, the hypothesis of a
participation of electrical junctions in the generation of delayed
current transients can be safely dismissed.
The delayed current transient is attributable to activation of
axonal GABAA receptors
A telltale property of delayed current transients is that they are
often associated with signs of active responses (Fig. 7). This escape
cannot occur in the dendrites because the triggering current is only on
the order of 100 pA. Simulations similar to those illustrated in Figure
10 indicate that injections of such currents in the dendrites elicit
only minor voltage changes ( 6 mV), which are clearly insufficient to
trigger action potentials. Therefore it can be concluded that delayed
current transients elicit voltage escape in the axon. These signals
occur only during the decay phase of the delayed current transients,
and they tend to be larger if they occur later, probably because the
Na+ channels are still recovering from the
inactivation caused by the initial axonal spike. We propose that these
signals are caused by the activation of axonal GABAA
receptors. Because the entire neuron is loaded with the high
Cl solution, activation of axonal receptors
induces local depolarization and may lead to voltage escape.
The axonal receptor hypothesis readily accounts for the other unusual
properties of the delayed current transient. Because the axon is thin
and long, and because it contains a high density of voltage-dependent
channels, severe distortions are expected to occur when the axonal
current is monitored in a somatic voltage-clamp recording. This,
according to the simulations of Figures 10-12, accounts for the
difference between the kinetics of the delayed current transient and
that of dendritic IPSCs. Because the axon has many release sites, as
opposed to one or a few release sites in any specific
interneuron-interneuron synapse, relative fluctuations in the total
number of releasing sites are expected to be much less in the first
case than in the second.
Physiological relevance of the autoreceptor current
As in the case of any presynaptic receptor, several functional
roles can be envisaged for the GABAergic autoreceptors of cerebellar interneurons. The first possibility is a participation in axo-axonic synapses, as occurs at the crayfish neuromuscular junction, in the
mammalian spinal cord, and in retinal bipolar cells of fish (for
review, see Rudomin, 1990; McGehee and Role, 1996). In the present case, however, such a mechanism is unlikely, for reasons that
have already been discussed. Alternatively, presynaptic
GABAA receptors could be activated by spillover from nearby
GABAergic terminals, as occurs at peptidergic terminals in the
pituitary (Zhang and Jackson, 1995b). In cerebellar slices, presynaptic GABAergic receptors could be sensitive to the buildup of the bulk concentration of GABA in the molecular layer after sustained activity of interneurons, leading to a negative feedback curtailing excessive inhibitory activity. A final possibility is that presynaptic GABAergic receptors are sensitive to the phasic release of GABA from the cell
that carries the receptors (autoreceptors), as proposed earlier in
hippocampal cultures (Vautrin et al., 1994). The fact that autoreceptor
currents can be measured gives strong support to this possibility. The
currents that are generated in the axon could contribute to regulate
the firing rate. However, it is difficult to predict the exact
consequences of the activation of the autoreceptor current, because
this necessitates a knowledge of the position of the reversal potential
of the GABAergic channels under physiological conditions, which is
presently unknown.
Another open question is whether the axonal and dendritic GABA channels
have the same properties. In fish retinal bipolar cells,
GABAC channels underlie the presynaptic GABA conductance (for review, see McGehee and Role, 1996). In the present work, however, as in the pituitary and the spinal cord of mammals, the presynaptic autoreceptor current is blocked by low concentrations of
bicuculline and thus appears to be carried by GABAA
channels. In the pituitary, presynaptic GABAA channels have
unique single-channel properties: subconductance states are much less
prominent than in somatodendritic channels (Zhang and Jackson, 1995b),
raising the intriguing possibility that axonal and somatodendritic
GABAA currents could be attributable to different
combinations of channel subunits.
| |
FOOTNOTES |
Received Oct. 23, 1998; revised Dec. 17, 1998; accepted Dec. 21, 1998.
This work was supported by a European Community grant (ERB 4061 PL97-0967). We thank L. Forti and I. Llano for comments on this manuscript.
Correspondence should be addressed to Dr. A. Marty, Arbeitsgruppe
Zelluläre Neurobiologie, Max-Planck-Institute für
Biophysikalische Chemie, 37077 Göttingen, Germany.
Dr. Pouzat's present address: Division of Biology 139-74, California
Institute of Technology, 1201E California Boulevard, Pasadena, CA 91125.
| |
APPENDIX A: Current delivered by the amplifier in response to a
step increase of a distributed conductance for a semi-infinite
cable |
We consider here a semi-infinite cable voltage-clamped at its
resting value at the origin. We call V the deviation of the membrane voltage from its resting value, and we assume that at t = 0 the cable is uniformly at rest. At rest, the
cable has a cytoplasmic resistivity Ri, a
membrane resistivity Rm, and a membrane capacitance Cm. At t = 0, a
uniformly distributed conductance density Gs,
with a reversal potential Es, is activated. Once
activated, Gs remains constant.
To simplify calculations, we use dimensionless parameters (Rall, 1977).
Then the cable equation takes the form:
|
(A.1)
|
with the boundary conditions:
|
(A.2)
|
and
|
(A.3)
|
and the initial condition:
|
(A.4)
|
We use the Laplace transform method to solve the problem. Applying
the Laplace operator:
|
(A.5)
|
on both sides of Equation A.1 and using the initial conditions
(Eq. A.4) and the boundary condition (Eq. A.3), we get:
|
(A.6)
|
where:
|
(A.7)
|
Applying the Laplace operator (Eq. A.5) to the boundary conditions
(Eqs. A.2 and A.3) gives:
|
(A.8)
|
and:
|
(A.9)
|
The solution of the inhomogenous ordinary differential equation
(A.6), with the boundary conditions (Eqs. A.8 and A.9), is (Tuckwell; 1988, pp 139-141):
|
(A.10)
|
The inverse transform of Equation A.10 is not easy to find but is
not needed here. Rather, we have to calculate the current delivered by
the amplifier. This is given, according to Equations 3.6 and 3.7 of
Rall (1977), by:
|
(A.11)
|
|
(A.12)
|
where G is the input conductance of the
semi-infinite cable. In view of the identity:
|
(A.13)
|
the problem of calculating ic(t)
amounts to finding the inverse transform of the partial derivative
(with respect to X) of VL at
X = 0. This is, according to Equation A.10:
|
(A.14)
|
Using a table of Laplace transforms (Gaskell, 1970, Eq. 44), we
obtain:
|
(A.15)
|
where:
|
(A.16)
|
Combining with Equation A.11 leads to the solution:
|
(A.17)
|
When T   , ic(T) takes the
value:
|
(A.18)
|
Because the resting conductance of the axon is usually much
smaller than that prevailing at the peak of the delayed current transient, Rm·Gs  1. With this simplification, Equations A.17 and A.18 may be rewritten,
respectively, as:
|
(A.19)
|
and
|
(A.20)
|
The rise-time of the current corresponds to the value of T,
Tr, for which the argument of the erf
function takes a certain value (which may be chosen close to 1 if the
rise-time is defined as the time to peak amplitude). Hence:
|
(A.21)
|
where Ct is a constant. Combining
Equations A.19 and A.21 yields:
|
(A.22)
|
Equation A.22 shows that the rise-time of the delayed current
transient is proportional to the inverse of the square of its peak amplitude.
| |
APPENDIX B: Steady-state solution of the equation of a finite
cable with a uniform autoreceptor conductance density |
We consider a finite cable voltage-clamped at its resting value at
the origin. The electrical parameters of the cable are as defined in
Appendix A. As in Appendix A, we assume that the cable is uniformly at
rest until at time T = 0; a synaptic (or autoreceptor)
conductance is activated and remains constant thereafter. The reduced
length of the cable, L, is related to the actual length,
l, and to the cable length constant,  , by the
equation:
|
(B.1)
|
Finally, a sealed-end boundary condition is used at the end of the
cable. The voltage escape (from the resting value), V, follows the equation:
|
(B.2)
|
with the boundary conditions:
|
(B.3)
|
and
|
(B.4)
|
and the initial condition:
|
(B.5)
|
We aim here at calculating V, the
steady-state solution of Equation B.2. From Equation B.2, we
obtain:
|
(B.6)
|
The solution of the inhomogeneous Equation B.6 with the boundary
conditions (B.3 and B.4) is (Tuckwell, 1988, pp 139-141):
|
(B.7)
|
From Equations B.7 and A.18, one obtains the expression of the
steady-state current delivered by the amplifier:
|
(B.8)
|
which, for Rm·Gs
1, simplifies to:
|
(B.9)
|
In Equation B.9, the length of the cable influences only the
argument of the tanh function. This function rises first proportionally to its argument but then abruptly saturates at 1 as the argument grows
further. Because of the shape of the tanh function, the amplitude of
the current delivered by the amplifier saturates for large values of
the cable length. Let us consider a specific case in which the density
of autoreceptor conductance, Gs, is 1 mS/cm2. Taking our standard values for
Ri (0.1 k·cm), Rm
(50 k·cm2), and a (0.25 µm), we
find that the length constant of the cable is = 791 µm. Applying
Equations B.9 and B.1, it follows that the currents for the cable
lengths of 200 and 400 µm are 116 and 123 pA, respectively (with
Es = +70 mV).
| |
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Top
Abstract
Introduction
Compound via MeSH
