Control of Action Potential-Driven Calcium Influx in GT1 Neurons by the Activation Status of Sodium and Calcium Channels
Fredrick Van Goor,
Lazar Z. Krsmanovic,
Kevin J. Catt and
Stanko S. Stojilkovic
Endocrinology and Reproduction Research Branch National
Institute of Child Health and Human Development National Institutes
of Health Bethesda Maryland 20892
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ABSTRACT
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An analysis of the relationship between electrical
membrane activity and Ca2+ influx in
differentiated GnRH-secreting (GT1) neurons revealed that most cells
exhibited spontaneous, extracellular
Ca2+-dependent action potentials (APs). Spiking
was initiated by a slow pacemaker depolarization from a baseline
potential between -75 and -50 mV, and AP frequency increased with
membrane depolarization. More hyperpolarized cells fired sharp APs with
limited capacity to promote Ca2+ influx,
whereas more depolarized cells fired broad APs with enhanced capacity
for Ca2+ influx. Characterization of the inward
currents in GT1 cells revealed the presence of
tetrodotoxin-sensitive Na+,
Ni2+-sensitive T-type
Ca2+, and dihydropyridine-sensitive L-type
Ca2+ components. The availability of
Na+ and T-type Ca2+
channels was dependent on the baseline potential, which determined the
activation/inactivation status of these channels. Whereas all three
channels were involved in the generation of sharp APs, L-type channels
were solely responsible for the spike depolarization in cells
exhibiting broad APs. Activation of GnRH receptors led to biphasic
changes in cytosolic Ca2+ concentration
([Ca2+]i), with an
early, extracellular Ca2+-independent peak and
a sustained, extracellular Ca2+-dependent
phase. During the peak
[Ca2+]i response,
electrical activity was abolished due to transient hyperpolarization.
This was followed by sustained depolarization of cells and resumption
of firing of increased frequency with a shift from sharp to broad APs.
The GnRH-induced change in firing pattern accounted for about 50% of
the elevated Ca2+ influx, the remainder being
independent of spiking. Basal
[Ca2+]i was also
dependent on Ca2+ influx through AP-driven and
voltage-insensitive pathways. Thus, in both resting and
agonist-stimulated GT1 cells, membrane depolarization limits the
participation of Na+ and T-type channels in
firing, but facilitates AP-driven Ca2+ influx.
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INTRODUCTION
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The mammalian hypothalamus contains about 1000 GnRH neurons that
are diffusely distributed within the mediobasal region, but
nevertheless operate in a highly synchronized manner to release pulses
of GnRH into the hypothalamo-hypophyseal portal vessels. This GnRH
neuronal network has been termed the GnRH pulse generator (1). The
propensity of GnRH neurons for pulsatile neuropeptide secretion is
common among mammalian species and is critical for the episodic release
of gonadotropins from the pituitary gland into the systemic circulation
(2). In the monkey, the mediobasal hypothalamus exhibits periodic
electrical activity that is highly synchronized with pulsatile LH
release into the circulation. These observations are consistent with
the hypothesis that simultaneous discharge of GnRH from nerve terminals
in the median eminence is a consequence of action potential (AP) firing
by the GnRH pulse generator (1).
Significant progress in elucidating the mechanism of basal GnRH
secretion has been made by studies on GnRH-secreting immortalized
neurons (GT1 cells) and cultured embryonic GnRH neurons (3). Several
in vitro experiments have shown that changes in cytosolic
Ca2+ concentration ([Ca2+]i)
determine the secretory pattern of GnRH (4), suggesting that
Ca2+ plays a central role in the signal transduction
processes that lead to exocytosis. Furthermore, GnRH secretion from
perifused GT1 and hypothalamic cells is reduced by L-type
Ca2+ channel inhibitors and augmented by activation of
voltage-gated Ca2+ channels (VGCC) (5). GT1 cells express a
variety of plasma-membrane channels, including tetrodotoxin
(TTX)-sensitive Na+ channels, transient and sustained
Ca2+ channels, inward rectifier K+ channels,
and several types of outward K+ channels (6). Embryonic
GnRH neurons also express a comparable set of plasma membrane channels
(7). In addition, single embryonic GnRH neurons and GT1 cells fire
spontaneous action potentials (APs) and exhibit fluctuations in
[Ca2+]i (8, 9). GT1 cells also express
several gap junction proteins (10) that probably permit electrical
coupling between these cells and transmit their synchronized
intercellular Ca2+ waves (11).
To date, the mechanism of AP generation in these cells and the channels
involved in spontaneous firing, as well as their relevance to the
control of [Ca2+]i, have not been completely
characterized. For example, the importance of TTX-sensitive
Na+ channels in AP generation and their relevance to
voltage-gated Ca2+ influx have not been clarified. Although
several adenylyl cyclase and phospholipase C-coupled plasma membrane
receptors are expressed in GnRH neurons (12, 13, 14, 15, 16, 17, 18), their effects on
spontaneous electrical activity have not been addressed. Also, the
mechanism for repletion of the endoplasmic reticulum
Ca2+ pool after activation of intracellular
Ca2+ release through inositol
1,4,5-trisphosphate-controlled Ca2+ channels has not been
determined in these cells. This problem has been studied in lactotrophs
(19) and other excitable cells, but the mechanism by which
voltage-gated Ca2+ influx and inositol
1,4,5-trisphosphate-induced Ca2+ signaling are integrated
has not been clarified. The present study was performed to analyze the
spontaneous spiking activity of isolated GT1 neurons and its importance
in the control of [Ca2+]i, and to
characterize the nature and role of the plasma membrane
Ca2+ oscillator in these cells. The major focus of these
investigations was on inward Na+ and Ca2+
currents and their roles in electrical activity in spontaneously active
cells and those stimulated by Ca2+-mobilizing agonists.
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RESULTS
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Patterns of AP Firing in Spontaneously Active GT1 Cells
The electrical membrane properties of GT17 neurons (hereafter
described as GT1 cells) were monitored using perforated-patch
recording techniques in the current-clamp mode. To eliminate the
influences of electrical and synaptic coupling between cells, only
isolated bipolar neurons with rounded perikarya and small neurites were
chosen. Potential autocrine and paracrine effects due to endogenous
neuropeptide and neurotransmitter release were minimized by constant
bath perifusion. Under these recording conditions, 95% (n = 237)
of the GT1 neurons exhibited self-sustained AP firing, a characteristic
feature of single-cell oscillators (Fig. 1
). The majority (n = 211) of these
cells exhibited either regular or irregular spiking activity, whereas
14 cells showed bursts of four to eight APs separated by quiescent
periods of 1020 sec. The pattern of bursting activity in these cells
was similar to that observed by others (8, 9). The remaining cells
(n = 12) did not exhibit spontaneous AP spiking, but transient
hyperpolarizing current injections elicited one to two rebound APs (not
shown). Since GT1 cells secrete GnRH and express GnRH receptors (14),
electrical activity was also analyzed in cells perifused with a GnRH
antagonist. Application of 1 µM
[D-pGly1,D-Phe2,D-Trp3,6]GnRH,
which completely inhibits agonist binding to GT1 cells (14), did not
affect the firing pattern (not shown).

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Figure 1. Spontaneous Electrical Activity in GT1 Cells
AC, Typical examples of the range of electrical membrane activity
observed in GT1 cells. D, Expanded time scale of single APs from the
recordings in AC labeled as a, b, and c. In this and the following
figures, perforated-patch recording was employed if not otherwise
specified.
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In spontaneously active neurons, the mean value of the maximum levels
of hyperpolarization observed (hereafter referred to as the baseline
potential), was between -75 and -50 mV (Fig. 1
, panels AC). The
interspike interval was characterized by a slow pacemaker
depolarization to the spike initiation threshold, which was between
-55 and -40 mV, depending on the baseline potential (Fig. 1D
). The
amplitude, duration, and frequency of APs were also dependent on the
baseline potential. In cells with more depolarized baseline potentials,
the frequency and duration of APs were greater, whereas AP amplitude
progressively decreased with increasing depolarization (Figs. 1
and 2A
). A similar relationship between baseline potential and frequency,
duration, and amplitude was observed in response to sequential
depolarizing current injections in a single GT1 cell (Fig. 2B
). Conversely, hyperpolarizing current
injections of -2 to -5 pA reduced AP frequency and duration and
increased the amplitude. Larger hyperpolarizing current injections
abolished AP firing (data not shown).

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Figure 2. Characterization of APs in GT1 Cells
A, Relationship between the baseline potential and frequency, duration,
and amplitude of APs in spontaneously active cells
(r = coefficient of correlation). B, Modulation of
spike frequency, amplitude, and duration of AP in a single cell exposed
to sequential applications of depolarizing current injections. Right
panel, Expanded time scale of APs from the recording shown in
left panel.
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These results indicate that AP properties differ from cell to cell,
depending on the baseline potential reached during AP firing. Moreover,
the pattern of firing in any given cell can be altered by changing the
baseline potential with either depolarizing or hyperpolarizing current
injections. Correlation analysis clearly indicates that the transition
from sharp to broad APs is not an all-or-none event, but rather a
graded, continuous process. The extremes of this continuum are
represented by the neuronal-like (high amplitude, sharp) APs shown in
Fig. 1C
, and the endocrine-like (low amplitude, broadened) APs shown in
Fig. 1A
.
TTX-Sensitive Na+ Channels and Pattern
of AP Firing
Since TTX-sensitive Na+ channels typically participate
in AP generation in neurons, we characterized their involvement in GT1
cells exhibiting different patterns of spontaneous electrical activity
(Fig. 3
). In cells exhibiting a baseline
potential more depolarized than -60 mV and low-amplitude, broad APs,
TTX had no effect on electrical activity (Fig. 3A
). However, in cells
that exhibited a baseline potential around -60 mV, 1 µM
TTX shifted the baseline potential to more depolarized levels, reduced
AP amplitude, and increased AP duration (Fig. 3B
). In cells exhibiting
a baseline potential between -75 and -65 mV and high-amplitude, sharp
APs, TTX transiently abolished spiking. This led to a gradual
depolarization and the transition to low-amplitude, broad AP firing
(Fig. 3C
). The majority of cells exhibited the pattern of firing shown
in Fig. 3B
. These results demonstrate that, when operative,
voltage-gated Na+ channels sharpened the APs in addition to
increasing their amplitude. Thus, the exclusion of the underlying
TTX-sensitive currents effectively transforms AP firing from the
neuronal to the endocrine-like mode.

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Figure 3. Effects of the Voltage-Gated Na+
Channel Blocker, TTX, on Spontaneous Electrical Activity in GT1 Cells
AC, left panels, Representative recordings of the
effects of 1 µM TTX (arrow) on GT1 cells
with varying baseline potentials. The majority of cells exhibited the
pattern of firing shown in panel B. AC, right panels,
Expanded time scale of single APs in the absence (a) or presence (b) of
1 µM TTX.
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To understand how the baseline potential level determines the AP
properties, we examined the electrophysiological characteristics of the
voltage-gated Na+ and Ca2+ currents generating
each AP. To isolate inward Na+ currents, outward currents
were blocked the intrapipette Cs+ and the cells were
bathed with Ca2+-deficient Krebs-Ringer medium containing 5
mM tetraethylammonium (TEA). Under these conditions,
a rapidly activating/inactivating Na+ current was observed
after command potentials more depolarized than -60 mV (Fig. 4
). In cells held at -97 mV, the time to
peak current amplitude between command potentials of -37 mV and 3 mV
was 4.1 ± 0.3 to 2.7 ± 0.1 msec (n = 4), and the
maximum amplitude was around -20 mV (Fig. 4B
). The mean
Na+ current density at the peak of the current-voltage
relation was 235 ± 70 pA/pFarad (pF) After activation, the
Na+ current rapidly inactivated during the 200-msec voltage
steps (Fig. 4A
). The residual (<3%) non-inactivating current may
suggest that an additional Na+ current is present in GT1
neurons, as in fish terminal nerve GnRH neurons (20). However, it may
also be due to imperfect voltage control due to leak subtraction
artifacts. The falling phase of the Na+ current elicited by
command potentials to between -37 mV and 3 mV was fitted with a single
exponential curve, and the calculated time course of inactivation was
between 8 ± 0.6 and 1 ± 0.1 msec (n = 4).

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Figure 4. Voltage-Gated Na+ Current in GT1 Cells
A, Sodium current traces elicited by membrane potential steps to -87,
-67, -47, -37, -27, -7, 13, 33, 53, 73 mV from a holding potential
of -97 mV. B, Current-voltage relation of the voltage-gated
Na+ current in GT1 neurons (mean ± SEM;
n = 4). Inset, Whole-cell voltage-clamp recordings of the effects
of TTX on Na+ current. From a cell hold at -97 mV,
currents were elicited by a 20 msec voltage step to -17 mV. The
average series resistance was 26 ± 2.7 M , with 6080%
compensation.
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The rapidly activating/inactivating Na+ currents in GT1
neurons were sensitive to the sodium channel blocker, TTX (Fig. 4
, inset). Application of 1 µM TTX during voltage
steps to -17 mV (holding potential = -97 mV) reduced the
Na+ current amplitude from -721 ± 143 pA to -7
± 3 pA (P < 0.05; n = 4). Higher concentrations
of TTX had no further effect (data not shown). The inhibitory actions
of TTX on Na+ current amplitude were observed at all
membrane potentials between -57 mV and 83 mV (data not shown).
Therefore, under our recording conditions, a conventional
fast-activating and -inactivating TTX-sensitive Na+ current
is present in GT1 neurons.
Although the rapid activation and current-voltage relation of the
Na+ current suggest that it could generate the sharp
upstroke of the APs in GT1 cells, TTX did not alter AP firing in a
subpopulation of the cells examined. A possible explanation for this is
that a large portion of the Na+ current is not available
for activation in cells with relatively depolarized baseline
potentials. This hypothesis was tested by examining the voltage
dependence of isochronal (steady-state) inactivation of the
Na+ current by using a two-pulse protocol (21). Cells held
at -97 mV were subjected to prepulse potentials ranging from -127 mV
to -7 mV for 100 msec, after which a test pulse to -17 mV was given
(Fig. 5A
). The normalized test current
from five cells was plotted against the prepulse potentials, and the
resulting curve was fitted with a single Boltzmann relation, where
E1/2 and k (see Materials and Methods)
were -68 mV and 9, respectively. This indicated that the amount of
Na+ current available for activation ranges from 6010%,
depending on the baseline potential reached during AP activity.
Moreover, relatively small changes in the level of the baseline
potential can significantly alter the amount of Na+ current
available for activation.

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Figure 5. Isochronal Inactivation and Recovery from
Inactivation of the Na+ Current in GT1 Cells
A, Isochronal inactivation curves for the Na+ current were
generated from prepulse experiments in which the Vm was
stepped to between -127 mV and 3 mV for 100 msec before stepping to a
command potential of -17 mV (holding potential = -97 mV).
Left panel, representative traces of the remaining
currents elicited during the command potential after prepulse
potentials between -127 and 3 mV. Right panel,
isochronal inactivation curve of the Na+ current (mean
± SEM, n = 5). The Na+ current was
normalized to the maximum inward current and fitted with a single
Boltzmann relation. B, Recovery from inactivation. A 100-msec command
potential to -17 mV was given after a prepulse to -17 mV and the
interpulse interval was varied from 10500 msec. Left
panel, Representative current traces elicited during the
command potential at varying interpulse duration. Right
panel, The Na+ current was normalized to the
maximum inward current (mean ± SEM; n = 7) and
plotted against the interpulse interval. The average series resistance
for the recordings in panels A and B were 26 ± 2.7 M , with
6080% compensation. C, Changes in Vm in response to
hyperpolarizing current injections of -10 pA in the absence or
presence of 1 µM TTX. The asterisks
indicate the high-amplitude, rebound AP after transient hyperpolarizing
current injections. The arrow indicates the time of TTX
application, which was present for the remainder of the experiment.
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To determine the time period required to remove Na+ current
inactivation, a 100-msec test pulse to -17 mV was given after a
prepulse to the same amplitude, and the interpulse duration was varied
(holding potential = -97 mV). The normalized test current was
plotted against the interpulse duration, and the time required for
reactivation of the Na+ current from inactivation was
determined (Fig. 5B
). Recovery from inactivation occurred at two
different rates, 50% of the current being recovered in less than 50
msec and the remainder in more than 500 msec (Fig. 5B
). This suggests
that in cells with a sufficiently hyperpolarized baseline potential,
the majority of the Na+ channels will recover from
inactivation during the interspike interval, as in those cells
exhibiting a baseline potential below -60 mV (Fig. 3
, B and C).
Conversely, the inactivation and the time required for its removal
account for the lack of involvement of TTX-sensitive Na+
channels in cells with relatively depolarized baseline potentials and
high AP frequencies, as in cells firing endocrine-like APs (Fig. 3A
).
Accordingly, hyperpolarizing current injections in cells exhibiting
endocrine-like APs generated a single, high-amplitude, sharp AP. In
the presence of 1 µM TTX, hyperpolarizing current
injections did not elicit high-amplitude, sharp APs (Fig. 5C
). Thus,
although transient membrane hyperpolarization could reactivate
Na+ channels in cells exhibiting endocrine-like AP spiking,
return to the depolarized baseline potential and high AP frequency
prevented further Na+ channel participation.
Voltage-Gated Ca2+ Channels and Pattern
of AP Firing
The properties of the VGCC subtypes expressed in GT1 cells were
analyzed in Na+-deficient medium containing TEA and TTX,
and with a pipette solution containing Cs+ and TEA. The
extracellular Ca2+ concentration of 2.6 mM was
the same as that used for the recordings of electrical membrane
activity shown in
Figs. 14


. Under these conditions, command
potentials more depolarized than -60 mV elicited a transient inward
current that inactivated within 200 msec (Fig. 6
, A and B). The inactivation rate of the
transient current elicited during membrane potential
(Vm) steps to between -60 and -40 mV could be
fitted with a single exponential function and was between 110 ± 3
and 19 ± 2 msec (n = 7). These data indicate the existence
of a low voltage-activated (LVA) Ca2+ current in GT1
neurons.

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Figure 6. Electrophysiological Characterization of
Voltage-Gated Ca2+ Currents in GT1 Cells
A, Representative Ca2+ current traces elicited by 200-msec
command potentials to -70, -50, -30, and -10 (holding potential of
-90 mV) using conventional whole-cell recording techniques. B,
Current-voltage relation of the early (open circles,
025 msec) and sustained (filled circles, 190200
msec) Ca2+ currents (mean ± SEM; n =
7). Panels C, D, and E, Isochronal inactivation curves for the LVA and
HVA Ca2+ current. The isochronal inactivation curve for the
LVA Ca2+ current (panel C, open squares) was
generated by giving a 200-msec command potential to -50 mV after 1-sec
prepulse from -120 to -10 mV (holding potential = -90 mV). The
isochronal inactivation curve for the HVA current (panel D,
filled squares) was generated by measuring the sustained
current amplitude during a 200-msec command potential to -10 mV after
a 2-sec prepulse potentials from -120 to -10 mV. The continuous line
for the isochronal inactivation curve of the LVA current is a fitted
Boltzmann relation (E). Currents were normalized to the maximal inward
current elicited during a command potential to -50 or -10 mV, after a
prepulse to -120 mV (mean ± SEM; n = 7). All
calcium current recordings shown in this figure were performed in the
presence of 2.6 mM Ca2+ using regular
whole-cell recording techniques.
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In addition to the LVA Ca2+ current, 200-msec or 2-sec
command potentials more depolarized than -50 mV (holding
potential = -90 mV) elicited a slowly inactivating
Ca2+ current (Fig. 6
, A and B). To determine the
inactivation rate of this high voltage-activated (HVA) Ca2+
current, 2-sec command potentials between -40 and 0 mV were applied,
and the falling phase of the current was fitted with a double or single
exponential curve. At these Vm values, the falling phase of
the current was best described by a double exponential fit. The initial
rates of inactivation were between 20 ± 2 and 36 ± 8 msec,
followed by an inactivation rate between 280 ± 51 msec and
504 ± 116 msec (n = 7). The densities of the early (025
msec) and sustained (190200 msec) Ca2+ currents at the
peaks of their individual current-voltage relations were -9.2 ±
2.4 pA/pF (n = 7) and -3.6 ± 0.9 pA/pF (n = 7),
respectively. These data are consistent with the coexistence of LVA and
HVA Ca2+ currents in GT1 neurons.
The proportions of these Ca2+ currents available for
activation at different Vm values were determined by
analysis of their isochronal inactivation properties in seven cells,
using a two-pulse protocol (21). The isochronal inactivation properties
of the LVA Ca2+ current were examined by applying prepulse
potentials from -120 to -10 mV for 1 sec (holding potential =
-90 mV), followed by a test pulse to -50 mV to activate predominately
LVA currents. The normalized test current was plotted against the
prepulse potentials, and the resulting curve was fitted with a single
Boltzmann relation (Fig. 6
, C and E), where E1/2 and k were
-68 mV and 12, respectively. To examine the isochronal inactivation
properties of the HVA Ca2+ current, the current remaining
at the end of a 200-msec test pulse to -10 mV was plotted against
2-sec prepulse potentials from -120 to -10 mV. The HVA current
exhibited minimal voltage-dependent isochronal inactivation (Fig. 6
, D
and E). These data indicate that, like TTX-sensitive Na+
currents, the proportion of the LVA current available for activation
depends on the baseline potential reached during AP activity. In
contrast, a large proportion of the HVA current is available for
activation, regardless of the baseline potential.
To further characterize the VGCC subtypes present, the effects of
selective antagonists were examined. The T-type channel is a LVA
Ca2+ channel and is sensitive to Ni2+ in the
micromolar concentration range (22). In GT1 neurons, the LVA
Ca2+ current elicited by a 200-msec voltage step to -50 mV
(holding potential = -90 mV) was inhibited by Ni2+ in
a concentration-dependent manner (Fig. 7A
). Application of 100 µM
Ni2+ reduced the LVA current amplitude to less than 10% of
control, whereas up to 20% of HVA current was abolished during
200-msec membrane potential steps to -10 mV. Application of 50
µM and 100 µM Ni2+ reduced the
voltage-gated Na+ current by 12 ± 2% and 14 ±
2% (n = 7), respectively. In cells exhibiting a baseline
potential below -60 mV (four of nine cells), addition of 50
µM (Fig. 7B
) and 100 µM Ni2+
(not shown) reduced the AP frequency but did not abolish spontaneous
spiking. In cells with a baseline potential more depolarized than -60
mV, 50 or 100 µM Ni2+ did not alter spike
frequency. Thus, as in experiments with TTX, the inhibitory effect of
Ni2+ was observed only in spontaneously active cells with
baseline potentials more negative than -60 mV. These results are
consistent with the expression of T-type Ca2+ channels in
GT1 neurons and their participation in pacemaker activity under certain
conditions. As with Na+ channels, the inactivation
properties probably account for the lack of involvement of
Ni2+-sensitive Ca2+ channels in the generation
of APs in cells exhibiting baseline potentials more depolarized than
-60 mV.

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Figure 7. Pharmacological Characterization of Voltage-Gated
Ca2+ Currents in GT1 Cells
A, Concentration-dependent effects of Ni2+, a blocker of
T-type LVA Ca2+ channels, on isolated Ca2+
currents elicited by a 200-msec command potential to -40 mV (holding
potential = -90 mV). Calcium current recordings were performed in
the presence of 2.6 mM Ca2+ using regular
whole-cell patch-clamp techniques. B, Effect of Ni2+ on the
pattern of spontaneous spiking. C, Effect of nifedipine, a blocker of
L-type HVA Ca2+ channels, on isolated Ba2+
currents. Voltage-clamp recording of isolated Ba2+ currents
elicited by a 200-msec command potential to -10 mV (holding potential
-90 mV) before and during the application of 1 µM
nifedipine. D, Current-clamp recording of electrical membrane activity
during the application of nifedipine (bar) and 5 min
after its removal (right panel). Recordings shown in
panels B, C, and D were done under perforated patch-clamp conditions.
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HVA Ca2+ channels that are sensitive to the
1,4-dihydropyridine inhibitor, nifedipine, are frequently associated
with Ca2+ entry during AP activity (22). The nifedipine
sensitivity of the HVA currents in GT1 neurons was determined by
measurement of the sustained Ba2+ current amplitude and
electrical membrane activity in cells bathed in
Ca2+-containing medium. As shown in Fig. 7
, C and D,
application of 1 µM nifedipine reversibly reduced the
amplitude of the sustained current elicited by a 200 msec voltage step
to -10 mV (holding potential = -90 mV), from -97 ± 20 pA
to -13 ± 6 pA (P < 0.05; n = 4). This
inhibition was observed during all command potentials more depolarized
than -50 mV. Nifedipine did not affect voltage-gated Na+
currents elicited by voltage steps to -17 mV (holding potential =
-97 mV; control = 1.22 ± 0.20 nA vs. 1
µM nifedipine = 1.21 ± 0.18 nA;
P > 0.05; n = 4) or the LVA Ca2+
currents elicited by voltage steps to -50 mV (holding potential of
-90 mV; control = -122 ± 21 pA vs. 1
µM nifedipine = -122 ± 27 pA;
P > 0.05; n = 5). Spontaneous spiking activity
was reversibly abolished by 1 µM nifedipine in
TTX-sensitive and -insensitive cells (Fig. 7D
).
Application of 10 µM Bay K 8644, an L-type
Ca2+ channel agonist, during a 200-msec voltage pulse to
-10 mV (holding potential = -90 mV) increased the sustained
Ba2+ current amplitude from -102 ± 46 pA to
-290 ± 69 pA (Fig. 8A
;
P < 0.05; n = 4). This increase was observed only
during command potentials between -50 mV and -10 mV, resulting in a
shift in the peak of the current-voltage relation to more
hyperpolarized Vm; however, the activation threshold of the
Ba2+ current was not altered (Fig. 8B
). Application of 10
µM Bay K 8644 increased the amplitude and duration of AP
spiking in GT1 neurons, consistent with its stimulatory actions on
Ba2+ current amplitude (Fig. 8
, C and D). Stronger membrane
after-hyperpolarizations were also observed (Fig. 8C
), suggesting that
the threshold [Ca2+]i for activation of
Ca2+-controlled K+ currents is reached. The
increase in spike amplitude was abolished by 1 µM TTX or
50 µM Ni2+ (data not shown), again confirming
that strong after-hyperpolarizations are sufficient to relieve a
proportion of the voltage-gated Na+ and T-type
Ca2+ currents from inactivation. In addition to these
expected effects of Bay K 8644 on the firing pattern, its facilitation
of Ca2+ influx was associated with an increase in the AP
frequency (Fig. 8
, C and D). Bay K 8644-induced increases in spike
frequency were also observed in cells bathed in TTX- or
Ni2+-containing medium (data not shown). These findings
demonstrate that L-type Ca2+ current is the major spike
depolarization current in spontaneously active GT1 neurons. They also
further indicate that in cells with relatively depolarized baseline
potentials, in which the TTX-sensitive and T-type channels are under
inactivation, L-type Ca2+ channels contribute to both
pacemaking and spike depolar-izations.

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Figure 8. Bay K 8644 Sensitivity of Spontaneous Spiking and
Ba2+ Currents in GT1 Cells
A, Voltage-clamp recording of isolated Ba2+ currents
elicited by a 200-msec command potential to -10 mV (holding
potential = -90 mV) in the absence or presence of 10
µM Bay K 8644. B, Current-voltage relation of the
Ba2+ current shown in panel A. C, Current-clamp recording
of electrical membrane activity before, during (bar),
and after 10 µM Bay K 8644 application to cells bathed in
Ca2+-containing medium (2.6 µM). D, Expanded
time scale of the recording in panel A before (solid
line) and during (dotted line) Bay K 8644
application.
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Plasma-Membrane Ca2+ Oscillator
In spontaneously active GT1 cells firing regular or irregular APs,
each spike generated a discrete, transient increase in
[Ca2+]i, producing an oscillatory-like
pattern of [Ca2+]i signaling (Fig. 9A
). The capacity of each AP to drive
Ca2+ depended primarily on its duration, as the amplitude
of the [Ca2+]i transients were larger in
cells exhibiting broad APs than in those exhibiting sharp,
high-amplitude APs (Fig. 9
, C and D). In the small percentage of GT1
cells exhibiting bursting activity, the intermittent trains of AP
spiking and the associated [Ca2+]i transients
combined to generate larger amplitude [Ca2+]i
fluctuations (Fig. 9B
). These results indicate that the properties of
each AP, as well as the overall pattern of AP spiking, can influence
Ca2+ signaling in GT1 cells.

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Figure 9. AP-Driven Ca2+ Influx in GT1 Cells
AC, Simultaneous measurements of spontaneous firing of APs and
[Ca2+]i. A, AP-driven Ca2+
transients in a cell exhibiting relatively regular spiking behavior. B,
High-amplitude [Ca2+]i fluctuations in a cell
exhibiting bursting spiking behavior. C, Expanded time scale of APs and
their associated [Ca2+]i transients in cells
exhibiting narrow (left panel) and broad (right
panel) spiking. D, Relationship between AP duration and the
maximum amplitude of the [Ca2+]i transients
(r = coefficient of correlation; n = 53).
|
|
Inhibition of spiking by hyperpolarizing current injections, or by
addition of the nonselective Ca2+ channel blocker,
Cd2+ (50 µM), or 1 µM
nifedipine, abolished the [Ca2+]i transients
and decreased global [Ca2+]i. Figure 10A
illustrates the effects of
nifedipine on spontaneous electrical activity and
[Ca2+]i in a patch-clamped and indo-1-loaded
cell. Perifusion of cells with Ca2+-deficient medium was
also associated with abolition of spiking. However, unlike nifedipine
and Cd2+ treatments, depletion of extracellular
Ca2+ hyperpolarized the membrane (upper panel)
and further reduced [Ca2+]i after addition of
nifedipine (bottom panel) or Cd2+ (not shown).
Similar effects of nifedipine and addition of
Ca2+-deficient medium were observed in unpatched and
fura-2-loaded cells (Fig. 10B
). Conversely, AP broadening by the
application of 10 µM Bay K 8644 increased the amplitude
of the [Ca2+]i transients and increased total
[Ca2+]i (Fig. 10C
). Thus, voltage-gated
Ca2+ entry through L-type Ca2+ channels
contributes to AP generation in GT1 cells and drives changes in
[Ca2+]i, consistent with the operation of a
plasma membrane Ca2+ oscillator in these cells. Moreover,
an increase in AP duration generates larger amplitude
[Ca2+]i fluctuations, despite the decrease in
AP amplitude. Finally, another Ca2+-conducting current
participates in the control of baseline potential and
[Ca2+]i.

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Figure 10. Effects of Nifedipine,
Ca2+-Deficient Medium, and Bay K 8644 on Spontaneous
Electrical Activity and [Ca2+]i in GT1 Cells
A, Simultaneous measurement of Vm and
[Ca2+]i during the application of 1
µM nifedipine followed by removal of extracellular
Ca2+ (-Ca2+) in indo-1-loaded cells. B,
Effects of 1 µM nifedipine and extracellular
Ca2+ removal on [Ca2+]i in
fura-2-loaded and unpatched GT1 cells. The trace represents the mean
ratio (F340/F380) from 44 separate cells. The
dashed line in panels A and B illustrates the
[Ca2+]i in the presence of nifedipine. C,
Simultaneous measurements of APs and [Ca2+]i
in the absence (upper panel) or presence (lower
panel) of 10 µM Bay K 8644. Electrical membrane
activity and [Ca2+]i were recorded in the
presence of normal Krebs-Ringers medium using the perforated
patch-clamp recording technique.
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GnRH-Induced Shift in the Pattern of AP Firing
Activation of Ca2+-mobilizing GnRH receptors in GT1
cells generated biphasic changes in Vm and
[Ca2+]i. GnRH initially induced a spike
increase in [Ca2+]i that transiently
hyperpolarized the plasma membrane (Fig. 11A
). This was followed by sustained
depolarization with a change in baseline potential from -63.4 ±
1.8 mV to -54.3 ± 2.2 mV (P < 0.001; n =
15) and a concomitant increase in spike frequency, which coincided with
a sustained elevation in [Ca2+]i (Fig. 11A
).
In addition to cell depolarization and an increase in spike frequency,
GnRH caused an increase in AP duration and a decrease in AP amplitude
(Fig. 11B
). The increase in AP duration was accompanied by an increase
in the amplitude of the AP-driven [Ca2+]i
transients (Fig. 11B
). These effects were similar to those observed
after current-induced membrane depolarization (Fig. 2B
). This is
consistent with the hypothesis that GnRH-induced depolarization
increases the number of TTX-sensitive Na+ channels and
T-type Ca2+ channels under inactivation. This, in turn,
should reduce their participation in AP generation, shifting the AP
firing pattern from neuronal-like to endocrine-like and thereby
increasing the capacity of the plasma-membrane Ca2+
oscillator to drive Ca2+.

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Figure 11. GnRH-Induced Modulation of Spontaneous Electrical
Activity in GT1 Cells
A Left panel, Simultaneous measurements of
Vm and [Ca2+]i in response to
application of 100 nM GnRH. Right panel,
Increase in spike frequency (mean ± SEM; n = 15)
after the application of 100 nM GnRH. B, left
panel, Expanded time-scale of single APs and associated
[Ca2+]i transients from the recording in
panel A labeled a and b. Right panel, Effects of 100
nM GnRH on spike amplitude and duration (mean ±
SEM; n = 15). Asterisks indicates
significant differences compared with control values:
P < 0.05, Students
t-test.
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To test this hypothesis, the actions of VGCC antagonists and
extracellular Ca2+ depletion during GnRH stimulation were
examined. To avoid the possible impact of patch clamping on
GnRH-induced Ca2+ influx, and to monitor changes in
[Ca2+]i from several cells simultaneously,
Ca2+ imaging experiments using fura-2 AM-loaded GT1 cells
were performed. As in patch-clamped and indo-1-loaded cells,
application of 100 nM GnRH stimulated a biphasic increase
in [Ca2+]i, composed of a transient spike
phase and a sustained plateau phase (Fig. 12A
). In the presence of 1
µM nifedipine, the GnRH-induced spike phase was not
affected but the [Ca2+]i plateau was reduced
by about 50% (Fig. 12
, A and B). Similar results were observed during
nonselective blockade of VGCC with Cd2+ (Fig. 12B
).
Application of 1 µM nifedipine during sustained GnRH
stimulation also reduced the [Ca2+]i plateau
phase by about 50% (Fig. 12C
). Simultaneous measurement of
Vm and [Ca2+]i further indicated
that the GnRH-induced [Ca2+]i plateau was
mediated in part by voltage-gated Ca2+ entry (Fig. 12E
).
Conversely, addition of 100 nM GnRH in the presence of
Ca2+-depleted medium generated a monophasic increase in
[Ca2+]i (Fig. 12A
), since the
[Ca2+]i plateau phase was completely
abolished (Fig. 12B
). Also, unlike blockade of voltage-gated
Ca2+ entry, inhibition of voltage-gated Na+
channels by 1 µM TTX did not alter the GnRH-induced
[Ca2+]i spike or plateau phase (Fig. 12D
).
Thus, an increase in the capacity of the plasma-membrane
Ca2+ oscillator to drive Ca2+, due to increased
AP frequency and duration, contributes to the GnRH-induced
[Ca2+]i plateau phase. However, a
VGCC-independent Ca2+ influx pathway also contributes to
this phase of the GnRH-induced Ca2+ response.

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Figure 12. Contribution of Ca2+-Driven APs to
GnRH-Induced Plateau [Ca2+]i Response in GT1
Cells
A, Pattern of GnRH-induced [Ca2+]i response
in Ca2+-containg medium with or without 1 µM
nifedipine or in Ca2+-deficient medium. B, GnRH-induced
change in plateau [Ca2+]i (mean ±
SEM, measured 3 min after GnRH addition) in the absence or
presence of 1 µM nifedipine, 50 µM
CdCl2, or in Ca2+-deficient medium. C, Effects
of sustained addition of 1 µM nifedipine in
GnRH-stimulated cells. D, Lack of effect of TTX on GnRH-induced
[Ca2+]i response. In panels AD, changes in
[Ca2+]i (F340/F380)
were simultaneously monitored in up to 30 individual and unpatched
cells loaded with fura-2, and the mean
[Ca2+]i responses are shown. E, Simultaneous
measurement of Vm and [Ca2+]i in
indo-1-loaded GT1 cells under perforated patch-clamp recording
conditions. The bar and shaded area
indicate the duration of 1 µM nifedipine or
Ca2+-deficient medium application, respectively. GnRH (100
nM) was present from the time of application indicated by
the arrow.
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|
The agonist-induced shift from neuronal- to endocrine-like spiking may
be due to inhibition of TTX-sensitive Na+ channels and
T-type Ca2+ channels and/or augmentation of L-type
Ca2+ channels. To test this, we examined the actions of
GnRH on isolated Na+ and Ca2+ currents, as well
as on Vm in the presence of VGCC antagonists.
Application of 100 nM GnRH did not alter the
Na+ current or the peak or sustained Ca2+
current (Fig. 13
, A and B). Moreover,
the GnRH-induced membrane depolarization was maintained in the presence
of 1 µM nifedipine and 50 µM
Ni2+ (Fig. 13C
). These data indicate that neither
inhibition of Na+ or T-type current, nor direct
augmentation of L-type Ca2+ channels, mediates the actions
of GnRH. This further supports the view that the removal of
Na+ and T-type Ca2+ currents during GnRH action
is due to their inactivation imposed by the GnRH-induced
depolarization of the cell membrane.

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Figure 13. Effects of GnRH on Voltage-Gated Na+
and Ca2+ Currents in GT1 Cells
A, left panel, Superimposed Ca2+ current
traces elicited during a 200-msec voltage-step to -10 mV (holding
potential = -90 mV) in the absence or presence of 100
nM GnRH. Right panel, Ca2+
current amplitude during the peak (025 msec) and sustained (190200
msec) current in the presence or absence of 100 nM GnRH
(mean ± SEM; n = 6). Calcium current recordings
were performed in the presence of 10 mM Ca2+.
B, left panel, Superimposed Na+ current
traces elicited by a 50-msec voltage step to -17 mV in the absence or
presence of 100 nM GnRH. Right panel,
Na+ current amplitude in the presence or absence of 100
nM GnRH (mean ± SEM; n = 6). C,
Application of 100 nM GnRH in the presence of 1
µM nifedipine and 50 µM Ni2+
under current-clamp recording conditions. The dotted
line represents the baseline potential in unstimulated cells.
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|
 |
DISCUSSION
|
---|
Many neuronal and endocrine cells generate APs spontaneously or in
response to agonist stimulation. When such firing is associated with
periodic Ca2+ influx, it is referred to as a plasma
membrane Ca2+ oscillator (23). For example, APs generated
in conjunction with, or solely by, Ca2+ influx through VGCC
transiently drive Ca2+ into the cell to cause discrete
fluctuations in [Ca2+]i. This is an integral
part of the cellular Ca2+ homeostatic pathway in these
cells, and is intimately involved in the control of molecular and
cellular function (24). Earlier studies have indicated that GT1 neurons
express T-type and L-type voltage-gated Ca2+ channels,
voltage-gated Na+ channels, and voltage-dependent and
Ca2+-controlled K+ channels, but do not exhibit
spontaneous AP firing (6). In other reports, bursting AP activity and
high-amplitude [Ca2+]i fluctuations were
observed in isolated and interconnected GT1 cells (9, 11). Embryonic
and terminal nerve GnRH neurons express a comparable set of plasma
membrane channels, but exhibit regular AP firing (7). Like native GnRH
neurons, a majority of the GT1 cells examined in this study exhibited
relatively regular AP firing, whereas a small percentage of the cells
exhibited bursting AP firing comparable to that observed by Charles and
co-workers (8, 9).
The ability of a majority of the GT1 cells examined in the present
study to fire more frequent and relatively regular APs is probably
related to their degree of differentiation. In our experiments,
electrical membrane recordings were acquired from morphologically
differentiated cells that had been cultured in defined medium without
FCS for at least 7 days. In contrast, Bosma (6) and Charles and Hales
(9) employed GT1 cells cultured in serum-containing medium, which
facilitates cell division. This may affect the density of expressed
channels and, consequently, the underlying excitability of the cells.
For example, although both TTX-sensitive Na+ currents and
voltage-gated Ca2+ currents were observed in differentiated
(
Figs. 48



) and less-differentiated cells (6), there were significant
differences in the amplitudes of these inward currents. In general, the
amplitudes of Na+ and Ca2+ currents in
differentiated cells were 3- to 4-fold higher than in the
less-differentiated cells. Furthermore, only isolated cells were used
to exclude any effects of electrical or synaptic coupling between cells
that could influence the baseline potentials in connected cells.
Although the presence of TTX-sensitive Na+ channels, as
well as T-type and L-type Ca2+ channels, endows GT1 cells
with the ability to fire spontaneous APs, the relative contribution of
each channel is dependent on the baseline potential, which ultimately
determines the pattern of AP firing. For example, in cells that were
more hyperpolarized than -60 mV, all three currents contributed to AP
firing. Due to their sharp profile, high amplitude, and TTX
sensitivity, these APs were also referred to as neuronal-like. In
contrast, in cells with baseline potentials more depolarized than -60
mV, a large proportion of the Na+ and T-type
Ca2+ currents had undergone inactivation, so that only
L-type Ca2+ channels contributed to AP activity. This
resulted in broad, low-amplitude APs and an increase in firing
frequency. Since many excitable endocrine cell types also exhibit
TTX-insensitive firing (24), this pattern was referred to as
endocrine-like. However, the transition from sharp, neuronal-like to
broad, endocrine-like AP firing is a continuous, graded process that is
determined by the proportion of Na+ and T-type
Ca2+ currents available for activation. Regardless of the
baseline potential, and the resulting composition of the currents
contributing to each AP, transient activation of L-type
Ca2+ channels underlies the basal fluctuations in
[Ca2+]i in all cells. Thus, a plasma membrane
Ca2+ oscillator is operative in cells exhibiting both sharp
and broad APs. The capacity of the plasma membrane Ca2+
oscillator to drive Ca2+ into the cell is dependent on the
baseline potential, which influences both AP frequency and
duration.
In GT1 cells firing sharp APs, depolarization of the baseline potential
by current injection or GnRH increases AP frequency and duration and
facilitates Ca2+ influx. Membrane depolarization brings the
baseline potential closer to the firing threshold, resulting in an
increase in firing frequency and concomitant Ca2+ entry. In
addition, membrane depolarization progressively inactivates
TTX-sensitive Na+ currents and T-type Ca2+
currents, which decreases AP amplitude and increases AP duration.
Similar effects were observed in cells after inhibition of
voltage-gated Na+ currents by TTX. In resting and
GnRH-stimulated GT1 cells, the increase in AP duration increases the
magnitude of AP-driven [Ca2+]i transients,
despite a decrease in AP amplitude. The increase in AP duration caused
by the Ca2+ channel agonist, Bay K 8644, also amplified
AP-driven [Ca2+]i transients. Although there
is a decrease in AP amplitude in cells firing endocrine-like APs,
comparison of AP amplitude and the Ca2+ current/voltage
relation indicate that these APs are able to strongly activate HVA
Ca2+ currents. As in GT1 cells, AP prolongation increased
Ca2+ influx in smooth muscle cells (25). Similarly,
although an increase in AP duration decreases the peak Ca2+
current in rat sympathetic neurons (26) and ventricular myocytes (27),
there is a net increase in Ca2+ influx. In addition,
comparison of high-amplitude, sharp APs and low-amplitude, broad APs in
rat sympathetic neurons indicated that an increase in AP duration
facilitates Ca2+ influx, despite the decrease in AP and
Ca2+ current (26). Therefore, AP duration, as well as AP
amplitude, is critical in determining the net Ca2+ influx
per single spike.
The patterns of firing and associated Ca2+ influx are not
only controlled by the voltage-gated inward currents that were the
primary focus of this study. Voltage-gated K+ channels also
play an important role in the pattern of AP spiking (28). The decrease
in AP amplitude by inactivation of voltage-gated Na+ and
T-type Ca2+ channels could result in a reduction in the net
activation of voltage-gated K+ channels during spike
depolarization. This would reduce the potassium current needed to
repolarize the cells, prolonging the opening of voltage-gated L-type
Ca2+ channels and resulting in AP broadening and enhanced
Ca2+ influx. The increase in AP frequency after membrane
depolarization may also induce AP broadening by frequency-dependent
inactivation of voltage-gated K+ channels (29). The similar
effects of GnRH and depolarizing current injections on AP duration
argue against second messenger-mediated inhibition of K+
channels, as observed in other cell types (30). Thus, voltage-gated
Na+ and T-type Ca2+ channels are critical for
controlling AP duration, presumably by influencing the degree of
voltage-gated K+ channel activation, whereas L-type
Ca2+ currents are the major contributors to spike
depolarization and AP-driven [Ca2+]i
transients in GT1 cells.
In both unstimulated and agonist-stimulated cells, the ability of
spontaneous APs to drive Ca2+ is proportional to the level
of membrane depolarization. The extracellular
Ca2+-dependence and nifedipine, Ni2+,
Cd2+, and TTX insensitivity of the baseline potential, as
well as the partial dependence of basal and GnRH-induced plateau
[Ca2+]i responses on Ca2+ influx
through voltage-insensitive Ca2+ channels, are consistent
with the operation of an additional Ca2+-conducting channel
in GT1 neurons. This current may be similar to the
Ca2+-controlled nonselective cationic currents activated by
Ca2+-mobilizing receptors in other cells (31, 32). It may
also be related to the store-operated Ca2+ currents
observed in nonexcitable cells (33). The lack of effect of GnRH on
isolated Ca2+ or Na+ currents in GT1 cells
further indicates that GnRH-induced membrane depolarization is not due
to direct augmentation of either of these currents. In general,
inhibition of inwardly rectifying K+ channels (6, 35) or
M-like K+ currents (34) may mediate agonist-induced
membrane depolarization, but not the increased Ca2+ influx
and the dependence of baseline potential on extracellular
Ca2+.
In conclusion, the present results demonstrate the physiological
impact of spontaneous and receptor-mediated inactivation of two typical
pacemaker currents carried by TTX-sensitive Na+ channels
and T-type Ca2+ channels. In classical neurons,
TTX-sensitive channels drive AP depolarization. Coexpression of T-
and L-type Ca2+ channels is also frequently observed in
neural and neuroendocrine cells that exhibit spontaneous activity (24),
as well as in other spontaneously firing cell types (36). However,
spontaneous activity frequently occurs at voltages at which T-type
channels are almost completely inactivated. Both Na+ and
T-type Ca2+ channels participate in pacemaking in GT1
neurons, but only in cells with baseline potential between -60 and
-75 mV. Inhibition of these channels by TTX and Ni2+ does
not silence the cells, but promotes Ca2+ influx by
prolonging AP duration. These currents are suppressed during
depolarization of cells to levels that inactivate the majority of the
two channels. Such depolarization is facilitated by activation of
Ca2+-mobilizing GnRH receptors, but also occurs
spontaneously through a mechanism that has not yet been characterized.
In both resting and GnRH-stimulated cells, the depolarizing current
that controls the baseline potential also directly contributes to
Ca2+ influx.
 |
MATERIALS AND METHODS
|
---|
GT1 Cell Culture
All experiments were performed on the GT17 subtype of
immortalized GnRH neurons (3), which were originally provided by Dr.
Richard I. Weiner (University of California, San Francisco, CA). The
cells were grown in 75-ml culture flasks containing culture medium
(DMEM/F-12, 1:1, with L-glutamate, pyridoxine
hydrochloride, 2.5 g/liter sodium bicarbonate, 10% heat-inactivated
FBS, and 100 µg/ml gentamicin; GIBCO, Grand Island, NY). At
confluence, the cells were dispersed by trypsinization (0.05% trypsin)
for 10 min, resuspended in culture medium, and plated (50,000 cells/ml)
in 35-mm tissue culture dishes (Corning, Corning, NY) with or without
poly-L-lysine-coated (0.01%) coverslips. After incubation
for 48 h, the culture medium was replaced with medium containing
B-27 serum-free supplement (GIBCO) to induce morphological
differentiation of the cells. All experiments were performed 3 to 5
days after serum removal.
Electrophysiological Recordings
Ionic currents were measured using the whole-cell, gigaohm-seal
(37) or perforated-patch recording technique (38). All current-clamp
recordings of Vm were carried out using the
perforated-patch recording technique. Current- and voltage-clamp
recordings were performed at room temperature using an Axopatch 200 B
patch-clamp amplifier (Axon Instruments, Foster City, CA) and were
low-pass filtered at 2 kHz. For perforated patch recordings, patch
pipette tips (35 M
) were briefly immersed in amphotericin B-free
solution and then backfilled with amphotericin B (240
µg/ml)-containing solution. Before seal formation, liquid junction
potentials were canceled. An average series resistance of 19 ± 1
M
(n = 237) was reached 10 min after the formation of a gigaohm
seal (seal resistance > 5 G
) and remained stable for up to
1 h. When necessary, series resistance compensation was optimized
and current records were corrected for linear leakage and capacitance
using a P/-N procedure (21). Pulse generation, data
acquisition, and analysis were done with an PC equipped with a Digidata
1200 A/D interface in conjunction with pCLAMP 7 (Axon Instruments). All
recordings of Vm were digitized at 2 kHz using the software
package AxoScope 1.1 (Axon Instruments). In some cases, the
current-voltage relations were fit with a single Boltzmann relation:
I/Imax = Imax + exp[(E -
E1/2)/k]; where Imax is the maximum inward
current, E is the command potential, E1/2 is the
Vm at which there is 50% of the maximal current, and k is
the slope factor. Exponential fits were performed using clampfit. For
the determination of the AP properties in individual cells, the mean
baseline potential, as well as AP duration (half-amplitude) and
amplitude (threshold to peak) were determined from at least five
individual cells under control or experimental conditions. Action
potential frequency was determined during a 1- to 2-min period. For the
determination of the early (025 ms) and sustained (190200 ms)
Ca2+ current amplitude, the peak current amplitude during
the respective time periods was determined. All values in the text are
reported as mean ± SEM. Differences between groups
were considered to be significant when P < 0.05 using
paired t-test or ANOVA, followed by Fishers least
significant difference test.
Simultaneous Measurement of
[Ca2+]i and
Vm
GT1 neurons were incubated for 30 min at 37 C in phenol red-free
medium 199 containing Hanks salts, 20 mM sodium
bicarbonate, 20 mM HEPES, and 0.5 µM indo-1
AM (Molecular Probes, Eugene, OR). The coverslips with cells were then
washed twice with modified Krebs-Ringers solution containing (in
millimolar concentrations): 120 NaCl, 4.7 KCl, 2.6
CaCl2, 2 MgCl2, 0.7 MgSO4, 10
HEPES, 10 glucose (pH adjusted to 7.4 with NaOH) and mounted on the
stage of an inverted epifluorescence microscope (Nikon, Melville, NY).
A photon counter system (Nikon) was used to simultaneously measure the
intensity of light emitted at 405 nm and at 480 nm after excitation at
340 nm. Background intensity at each emission wavelength was corrected.
Perforated patch recording techniques (see above) were used to monitor
Vm. The data were digitized at 4 kHz using a PC equipped
with the pCLAMP 7-software package in conjunction with a Digidata 1200
A/D converter (Axon Instruments. The [Ca2+]i
was calibrated in vivo according to Kao (39). Briefly,
Rmin was determined by exposing the cells to 10
µM Br-A23187 in the presence of Krebs-Ringers solution
with 2 mM EGTA and 0 Ca2+ for 60 min; 15
mM Ca2+ was then added to determine
Rmax. The values used for Rmin,
Rmax, Sf,480/Sb,480, and
dissociation constant (Kd) were 0.472, 3.634, 3.187, and
230 nM, respectively.
Measurement of
[Ca2+]i in Unpatched
Cells
To determine the Ca2+ signaling patterns in cells
not under patch-clamp recording conditions, single-cell fura-2
Ca2+ imaging techniques were used. GT1 neurons were
incubated for 30 min at 37 C in phenol red-free medium 199 containing
Hanks salts, 20 mM sodium bicarbonate, 20 mM
HEPES, and 0.5 µM fura-2 AM (Molecular Probes). The cells
on coverslips were subsequently washed with Krebs-Ringer solution and
mounted on the stage of an Axiovert 135 microscope (Carl Zeiss,
Oberkochen, Germany) attached to the Attofluor Digital Fluorescence
Microscopy System (Atto Instruments, Rockville, MD). Cells were
examined under a 40x oil immersion objective during exposure to
alternating 340- and 380-nm light beams, and the intensity of light
emission at 520 nm was measured. The ratio of light intensities,
F340/F380, which reflects changes in
[Ca2+]i, was simultaneously measured in
several single cells.
Chemicals and Solutions
Stock solutions of TTX citrate (Research Biochemicals
International, Natick, MA) and GnRH (Peptides International,
Louisville, KY) were prepared in double-distilled, deionized water.
Stock solutions of nifedipine and S(-)-Bay K 8644 (Research
Biochemicals International) were dissolved in dimethylsulfoxide and
ethanol, respectively. The maximum final concentrations of
dimethylsulfoxide and ethanol were 0.1% and 0.01%, respectively,
neither of which altered electrical membrane activity or ionic
currents. For recordings of electrical membrane activity and total
inward and outward currents, the extracellular medium contained
modified Krebs-Ringer salts and the pipette solution contained (in
millimolar concentration): 70 KCl, 70 K-aspartate, 1 MgCl2,
and 10 HEPES (pH adjusted to 7.2 with KOH). In some experiments,
Krebs-Ringer salts without CaCl2 was used and is referred
to as Ca2+-deficient medium since it may contain residual
amounts of Ca2+ in the absence of added EGTA. To record
isolated Na+ currents, the extracellular medium contained
Krebs-Ringers solution without CaCl2 and with 20
mM TEA, 5 mM 4-AP, and 50 µM
CdCl2, and the pipette contained [in millimolar
concentration (mM) ]: 70 CsCl, 70
Cs-methanesulfonate, 2 MgCl2, and 10 HEPES (pH adjusted to
7.2 with CsOH). To record isolated Ca2+ currents using
conventional whole-cell recording techniques, the extracellular medium
contained (in mM): 100 tetramethylammonium-Cl, 20
TEA, 2.6 or 10 CaCl2, 1 MgCl2, 1
µM TTX and 10 HEPES (pH adjusted to 7.4 with
tetramethylammonium-OH). The pipette solution contained (in
mM): 120 CsCl, 20 TEA-Cl, 4 MgCl2, 10 EGTA, 9
glucose, 20 HEPES, and 0.3 Tris-GTP, 4 Mg-ATP, 14
creatine-PO4, and 50 U/ml creatine phosphokinase (pH
adjusted to 7.2 with Tris base). To record isolated Ba2+
currents using perforated-patch techniques, the extracellular solution
contained (mM): 120 TEA-Cl, 30 BaCl2, 2
MgCl2, 10 glucose, and 10 HEPES (pH adjusted to 7.2 with
TEA-OH), and the pipette solution was the same as that used for
isolated Na+ current recordings. All reported
Vm values under total current and isolated Na+
current recording conditions were corrected for a liquid junction
potential between the pipette and bath solution of +10 mV and +7 mV,
respectively, calculated using the JPCalc program (Axon
Instruments, Ref. 40). The junction potentials for isolated
Ca2+ and Ba2+ current recordings were less than
3 mV and were not corrected for. The bath contained less than 500 µl
of saline and was continuously perfused at a rate of 2 ml/min using a
gravity-driven superfusion system. The outflow was placed near the
cell, resulting in complete solution exchange around the cell within 2
sec. A solid Ag/AgCl reference electrode was connected to the bath via
a 3 M KCl agar bridge.
 |
FOOTNOTES
|
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Address requests for reprints to: Dr. Stanko Stojilkovic, National Institute of Child Health and Human Development, Endocrinology and Reproduction Research Branch, Building 49, Room 6A-36, 49 Convent Drive, Bethesda, Maryland 20892-4510. E-mail:
stankos{at}helix.nih.gov
Received for publication August 31, 1998.
Revision received December 14, 1998.
Accepted for publication December 16, 1998.
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