L-type Ca2+ Channels and K+ Channels
Specifically Modulate the Frequency and Amplitude of Spontaneous
Ca2+ Oscillations and Have Distinct Roles in Prolactin
Release in GH3 Cells*
Andrew C.
Charles
§,
Elemer T.
Piros¶,
Chris J.
Evans
, and
Tim G.
Hales**
From the
Department of Neurology, UCLA School of
Medicine, Los Angeles, California 90095, ¶ Department of
Physiology, Cornell University, New York, New York 10021,
Department of Psychiatry, Neuropsychiatric Institute, UCLA
School of Medicine, Los Angeles, California 90095, and ** Department of
Pharmacology, The George Washington University,
Washington, D. C. 20037
 |
ABSTRACT |
GH3 cells showed spontaneous
rhythmic oscillations in intracellular calcium concentration
([Ca2+]i) and spontaneous prolactin release. The
L-type Ca2+ channel inhibitor nimodipine reduced the
frequency of Ca2+ oscillations at lower concentrations
(100nM-1 µM), whereas at higher
concentrations (10 µM), it completely abolished them.
Ca2+ oscillations persisted following exposure to
thapsigargin, indicating that inositol 1,4,5-trisphosphate-sensitive
intracellular Ca2+ stores were not required for spontaneous
activity. The K+ channel inhibitors Ba2+,
Cs+, and tetraethylammonium (TEA) had distinct effects on
different K+ currents, as well as on Ca2+
oscillations and prolactin release. Cs+ inhibited the
inward rectifier K+ current (KIR) and increased
the frequency of Ca2+ oscillations. TEA inhibited outward
K+ currents activated at voltages above -40 mV (grouped
within the category of Ca2+ and voltage-activated currents,
KCa,V) and increased the amplitude of Ca2+
oscillations. Ba2+ inhibited both KIR and
KCa,V and increased both the amplitude and the frequency of
Ca2+ oscillations. Prolactin release was increased by
Ba2+ and Cs+ but not by TEA. These results
indicate that L-type Ca2+ channels and KIR
channels modulate the frequency of Ca2+ oscillations and
prolactin release, whereas TEA-sensitive KCa,V channels
modulate the amplitude of Ca2+ oscillations without
altering prolactin release. Differential regulation of these channels
can produce frequency or amplitude modulation of calcium signaling that
stimulates specific pituitary cell functions.
 |
INTRODUCTION |
There is increasing evidence for distinct roles of different
spatial and temporal patterns of intracellular free Ca2+
concentration ([Ca2+]i) in the regulation of
cellular processes (1). Many cell types exhibit oscillations of
[Ca2+]i that may be differentially modulated to
produce highly specific intracellular signals. For example, the
frequency of Ca2+ oscillations has been shown to regulate
secretion, whereas the amplitude of Ca2+ oscillations has
been shown to regulate gene expression in different cell systems (1,
2). The multifunctional enzyme calmodulin kinase II has been shown to
be capable of decoding different patterns of Ca2+ signaling
into different functional responses. (3).
Endocrine cells have the intrinsic capacity for extensive spontaneous
activity that is independent of stimulation by external factors. In
pituitary cells, this activity is characterized by membrane potential
oscillations, action potentials, and Ca2+ oscillations
(4-7). It is likely that this spontaneous, intrinsic signaling plays a
role in basal hormone release by pituitary cells and other endocrine
cells, although this role has yet to be clearly defined (8-10). In
addition, the individual components of this intrinsic signaling may be
targets of modulation through which diverse signals can induce specific
changes in cellular activity and hormone release.
The rat pituitary growth hormone- and prolactin-secreting
GH3 cell line is a useful and well studied model system for
the study of pituitary cell signaling. GH3 cells express
L-type Ca2+ channels as well as inwardly rectifying and
Ca2+- and voltage-activated K+ channels. The
biophysical and pharmacological properties of these channels have been
extensively characterized in previous studies (11-19). GH3
cells also show spontaneous activity that is generated by the
coordinated action of ion channels, Ca2+ influx,
Ca2+ release from intracellular stores, and other second
messengers including
IP31and cAMP. In
these and other pituitary cells, the relative contributions of each of
these cellular signaling components to overall cellular activity may
vary from cell to cell and under stimulated versus unstimulated conditions (20-22). It is well established that increases in [Ca2+]i directly mediate hormone release in
GH3 cells and other endocrine cell types (10, 23, 24).
Oscillatory patterns of Ca2+ signaling provide the
opportunity for a cell to respond to individual components of a
Ca2+ signal (e.g. base-line
[Ca2+]i, oscillation frequency, oscillation
duration, or oscillation amplitude). The functional response of the
cell may be different if it depends on a "frequency-modulated"
signal versus an "amplitude-modulated" signal (1).
Individual ion channels and second messengers may play specific roles
in generating specific patterns of spontaneous Ca2+
signaling and in turn may generate different patterns of hormone release.
In this study, we have used the combination of patch clamp
measurements, fluorescence imaging of intracellular Ca2+
concentration, and a sensitive enzyme-linked immunosorbent assay (ELISA) for prolactin to study the ionic mechanisms controlling hormone
release from GH3 cells. We have investigated the role of
specific patterns of spontaneous Ca2+ signaling in
prolactin release from GH3 cells by using Ca2+
and K+ channel antagonists to modulate the patterns of
Ca2+ signaling.
 |
EXPERIMENTAL PROCEDURES |
Cell Culture--
GH3 cells, obtained from American
Type Culture Collection, Manassas, VA (CCL 82.1), were maintained in
Dulbecco's modified Eagle's medium supplemented with 10% (v/v) fetal
calf serum, penicillin (0.05 IU/ml), and streptomycin (50 µg/ml) and
incubated in a humid atmosphere of 5% CO2, 95%
O2 at 37 °C. Cells were harvested once a week by
treatment with a phosphate-buffered saline containing EDTA (1 mM) and reseeded at 20% original density, either into 6-well plates for prolactin release assays, 35-mm diameter culture dishes for electrophysiological studies, or
poly-D-lysine-coated coverslips for
Ca2+-imaging studies. The incubation medium was changed
every 2-3 days.
Electrophysiological Recordings--
Single cells were
voltage-clamped, and voltage-activated K+ channel activity
was recorded from whole GH3 cells using a List EPC-7
patch-clamp amplifier. For the recording of KCa,V channel activity, cells were superfused with a solution containing 140 mM NaCl, 2.8 mM KCl, 2 mM
MgCl2, 1 mM CaCl2, 10 mM HEPES, 6 mM glucose, 5 × 10
4 mM tetrodotoxin (pH 7.2 with NaOH). The
recording electrode contained 120 mM KCl, 1 mM
EGTA, 1 mM MgCl2, 3 mM Mg-ATP, 10 mM HEPES (pH 7.2 with KOH) (all from Sigma). Currents were
activated by step depolarizations of membrane potential from a holding
potential of
80 mV for 100 ms every 10 s. Capacitance
compensations were achieved using the patch-clamp amplifier. Residual
artifacts and leakage currents were nulled using a P/4 subtraction.
Whole-cell KIR current recordings were performed using
extracellular solutions containing 140 mM KCl, 4 mM MgCl2, 1 mM CaCl2, 10 mM HEPES, 7 mM glucose, 5 × 10
4 mM tetrodotoxin (pH 7.2 with KOH). The
solution in the recording electrode contained 140 mM KCl,
10 mM EGTA, 2 mM MgCl2, 10 mM HEPES, 3 mM Mg-ATP (pH 7.2 with KOH).
Currents were evoked by hyperpolarizing from a
40 mV holding
potential (duration 1.5 s, frequency 0.03 Hz). No leak subtraction
was employed in these experiments.
Measurement of
[Ca2+]i--
[Ca2+]i was
measured using a fluorescence imaging system that has previously been
described in detail (25). Briefly, cells on
poly(D-lysine)-coated glass coverslips were loaded with fura2 by incubation in 5 µM fura2-AM for 40 min. Cells
were then washed and maintained in normal medium for 30 min before
experimentation. Coverslips were excited with a mercury lamp through
340- and 380-nm band-pass filters, and fluorescence at 510 nm was
recorded through a 10× or 20× objective with a silicon intensified
target camera to an optical memory disc recorder. Images were then
digitized and subjected to background subtraction and shading
correction, after which [Ca2+]i was calculated on
a pixel-by-pixel basis, as described previously, by a frame grabber and
image analysis board (Data Translation). Data acquisition and analysis
software were written by Dr. Michael Sanderson. Tracings in all figures
are based upon fluorescence of a 4 × 4 pixel area located within
each cell body.
Experiments were carried out in Hanks' balanced salt solution with 10 mM HEPES buffer, pH 7.4 (HBSS/HEPES) at 22 °C. Agents were applied by perfusion of coverslips with at least 2 ml of HBSS/HEPES containing the particular agent.
ELISA--
A competitive ELISA has been developed for measuring
prolactin secreted by GH3 cells. The assay utilizes an
antibody (raised in rabbit) against rat prolactin. Both antisera
(prolactinS-9) and standards (prolactin RP-3) were kindly provided by
the National Institute of Diabetes and Digestive and Kidney Diseases
(NIDDK). For the collection of samples, GH3 cells were
seeded into 6-well tissue culture plates 2 days before experiments
(0.5-0.7 million cells/well). Release experiments were conducted at
37 °C in a humidified incubator with 5% CO2.
Immediately before each experiment, cells were washed gently with media
(Dulbecco's modified Eagle's medium with 20 mM HEPES and
0.1% bovine serum albumin, pH 7.4, with NaOH). After washing, aliquots
of media (1 ml) were added to each well for 0.5-h time points such that
release could be monitored before, during, and after exposure to drugs.
The amount of prolactin (ng/ml/106 cells) released in
0.5 h in the presence of drugs was expressed as a percentage of
release from the same cells during 0.5 h under control conditions
before exposure to drugs. After incubation with the cells, each media
aliquot was centrifuged at 1800 rpm at 4 °C then stored at
20 °C or assayed directly by ELISA for prolactin. For the
competitive prolactin ELISA, 96-well Nunc-Immuno Maxisorp Plates from
Life Sciences, Denver, CO were used. Each well was coated with
prolactin by incubation of 100 µl of 0.1 M
NaHCO3, pH 9.5, containing 1 ng of prolactin for 20-24 h
at 4 °C. Before assay, prolactin-coated plates were washed with
assay buffer containing 0.5 M NaCl, 20 mM
NaH2PO4, 0.05% Tween 20, 0.5% bovine serum
albumin, pH to 7.4, then incubated with assay buffer for 30 min at room
temperature to remove prolactin binding weakly to the plate. After
further washing with assay buffer, undiluted samples (100 µl) or
standards (0.02-40 ng) dissolved in 100 µl of media were added to
the wells, followed by the addition of 50 µl of prolactin antibody at
a dilution of 1:40,000. After incubation for 2 h at room
temperature, bound antibody was detected using peroxidase-conjugated
anti-rabbit antibody (Vector, Burlingame, Ca) with tetramethylbenzidine
(Life Technologies, Inc.) as substrate. The peroxidase reaction was
terminated by 1 N H2SO4, and
absorbance was measured by a microplate reader (Molecular Devices) at
450 nm. To determine the amount of prolactin present in the samples, a
standard curve was generated. The percent of maximum absorbance (corresponding to no prolactin added) was plotted against known amounts
of prolactin (0.02-40 ng). All samples were assayed in quadruplicates
from three separate determinations.
 |
RESULTS |
The majority of GH3 cells (approximately 70%,
n > 1500 cells in 50 experiments) showed spontaneous
oscillations in [Ca2+]i. The pattern of these
oscillations in [Ca2+]i varied considerably from
cell to cell. Ca2+ oscillations had a periodicity ranging
from 3-30 s and a peak amplitude ranging from 40-300 nM
in different cells. Some cells showed Ca2+ oscillations
with a relatively consistent frequency, amplitude, and shape
(e.g. Fig. 1, Cell
#2), whereas other cells showed a more random pattern of
oscillations (e.g. Fig. 1, Cell #28). Increasing the temperature of the bath solution from room temperature to 37 °C
increased the frequency of spontaneous Ca2+ oscillations
but did not result in any qualitative changes in spontaneous
Ca2+ oscillations or in their response to the conditions
described below (data not shown).

View larger version (44K):
[in this window]
[in a new window]
|
Fig. 1.
The effect of nimodipine on spontaneous
Ca2+ oscillations. The left panel is a
raster plot of [Ca2+]i versus time in
a microscopic field of 30 different GH3 cells. Each
row represents an individual cell, and
[Ca2+]i is represented by changes in
gray-scale as shown on the bar at the upper left.
The right panel shows line tracings of
[Ca2+]i in an exemplary individual cell
(top trace) as well as the average
[Ca2+]i for the entire field of cells. Cells in
the field show heterogeneous patterns of spontaneous Ca2+
oscillations, with a wide variation in frequency and amplitude (peak
minus base). Bath application of 1 µM nimodipine results
in a reduction in the frequency of Ca2+ oscillations in
some cells (e.g. Cell #2) and abolishes
Ca2+ oscillations in other cells. Bath application of 10 µM nimodipine abolishes Ca2+ oscillations in
all cells.
|
|
Ca2+ oscillations were completely abolished in all cells
during perfusion with medium containing no added Ca2+, and
they were restored immediately upon replacement of Ca2+
(n = 120 cells in three experiments, data not shown).
We have previously shown that GH3 cells show predominantly
L-type Ca2+ currents that are inhibited by nimodipine (19).
Nimodipine inhibited spontaneous Ca2+ oscillations in a
concentration-dependent fashion (Fig. 1). Nimodipine (1 µM) reduced the frequency of Ca2+
oscillations in some cells and abolished them in other cells. Interestingly, although 1 µM nimodipine reduced the
frequency of Ca2+ oscillations in some cells, in most of
these cells it did not significantly affect their amplitude. Nimodipine
(10 µM) abolished Ca2+ oscillations in all
cells (n = 100 cells in three experiments). Nimodipine
(10 µM) also reduced spontaneous prolactin release by
32.9 ± 9.1% (see Fig. 9). The persistence of prolactin release despite the abolition of Ca2+ oscillations shows that a
significant portion of the spontaneous hormone release does not require
spontaneous Ca2+ signaling.
Exposure to 1 µM thapsigargin resulted in an increase in
base-line [Ca2+]i and a slight increase in the
frequency of Ca2+ oscillations in most cells
(n = 120 cells in four experiments). Base-line
[Ca2+]i returned to previous levels after 3-5
min in thapsigargin. After exposure to thapsigargin, Ca2+
oscillations continued to occur in 69% of cells showing spontaneous oscillations (59/87 cells in three experiments), although in some cells
their amplitude and frequency were reduced (Fig.
2). The phospholipase C inhibitor U73122
had no significant effect on Ca2+ oscillations, suggesting
that the active formation of IP3 is not required for
spontaneous Ca2+ oscillations (n = 100 cells in three experiments, data not shown). The activity of both
thapsigargin and U73122 were verified by the observation that each
agent completely inhibited the rapid peak increase in
[Ca2+]i induced by thyrotropin-releasing hormone
(data not shown). These results show that the activity of phospholipase C and the release of thapsigargin-sensitive intracellular
Ca2+ are not required for spontaneous Ca2+
oscillations in most cells but may contribute to modulation of their frequency and amplitude.

View larger version (36K):
[in this window]
[in a new window]
|
Fig. 2.
The effect of thapsigargin on spontaneous
Ca2+ oscillations. The left panel is a
raster plot of [Ca2+]i versus time in
a field of 30 different GH3 cells showing the effect of
bath application of 1 µM thapsigargin after 90 s of
spontaneous activity. Most cells show a transient increase in base-line
[Ca2+]i with a superimposed increased frequency
of Ca2+ oscillations, but some cells instead show a
decrease or no change in base-line [Ca2+]i and
Ca2+ oscillation frequency. After at least 15 min of
exposure to thapsigargin, most cells continue to show spontaneous
Ca2+ oscillations. The right panel shows
line tracings of [Ca2+]i in a typical
cell (top trace) as well as the average
[Ca2+]i for the entire field.
|
|
The effect of nimodipine suggests that Ca2+ oscillations
are generated by spontaneous depolarizations with resultant influx of
Ca2+ through voltage-gated channels. Consistent with this
hypothesis, spontaneous Ca2+ oscillations were highly
sensitive to depolarization induced by increasing extracellular
[K+]. Increasing extracellular [K+] by as
little as 3 mM resulted in a increase in base-line
[Ca2+]i as well as an increase in the frequency
of Ca2+ oscillations (n = 100 cells in
three experiments, not shown).
GH3 cells display multiple types of K+
currents, including an inward rectifying (KIR) current and
outward currents mediated by Ca2+ and voltage-activated
channels (15-17, 26). For the purposes of this study, the different
outward K+ currents recorded were grouped into a single
category of Ca2+ and voltage-activated (KCa,V)
currents. KCa,V channel activity was recorded using the
whole-cell patch-clamp configuration by depolarizing GH3
cells held at
80 mV to between
50 and 60 mV in 10-mV increments
(Fig. 3). Using the specified internal
and external solutions (see "Experimental Procedures"), outward
K+ currents were observed with a threshold of activation of
approximately
40 mV. Different recording solutions (see
"Experimental Procedures") and hyperpolarizing steps from a
40 mV
holding potential to between
50 and
120 mV (10 mV decrements) were
used to specifically activate currents mediated by KIR
channels. KIR channel activity recorded at more depolarized
potentials than
40 mV was overwhelmed by currents through
KCa,V channels. KIR and KCa,V
currents were inhibited selectively by different agents (Fig.
4). TEA (1 mM) inhibited outward KCa,V currents but not KIR currents.
The outward K+ currents had both transient and sustained
components; the inhibition of outward K+ current by TEA was
most marked at the end of the voltage step, indicating a relatively
selective block of the sustained current component. Ba2+ (1 mM) inhibited both KIR and KCa,V
(with no obvious specificity for the transient or sustained
components), whereas Cs+ inhibited only KIR
currents (Fig. 4).

View larger version (22K):
[in this window]
[in a new window]
|
Fig. 3.
Whole-cell K+ currents recorded
from GH3 cells. A, outward K+
currents through KCa,V channels were recorded using the
whole-cell patch-clamp technique. Currents were evoked by depolarizing
from a 80 mV holding potential to voltages between 50 to 60 mV (10 mV increments) for 100 ms. Traces represent averaged
currents recorded from 6 cells. B, relationship between the
peak K+ current amplitude and test pulse are shown
graphically. Data were obtained from the same cells as in A.
Vertical error bars, when bigger than symbols, represent
±S.E. C, KIR currents were activated by
hyperpolarizing pulses (duration 1.5 s) to voltages between 50
to 120 mV (10 mV decrements) from a 40 mV holding potential.
Equimolar KCl (120 mM) solutions were used to record
KIR channel activity (see "Experimental Procedures").
Tracings represent currents averaged from 5 cells.
D, the graph shows the current-voltage
relationship of KIR channel activity recorded from the same
cells as in C. Both peak ( ) and sustained ( ) current
amplitude is illustrated. Vertical error bars represent ± S.E.
|
|

View larger version (26K):
[in this window]
[in a new window]
|
Fig. 4.
The effect of K+ channel
antagonists on KCa,V and KIR channels in
GH3 cells. A, control and Ba2+
(1 mM)-inhibited KCa,V currents.
Ba2+-inhibited peak and sustained current components
indicating blockade of both a transient A current and longer lasting
currents, consistent with delayed rectifying and
Ca2+-activated K+ currents. B, TEA
(10 mM) predominantly inhibited the sustained current
component. In both A and B, superimposed
traces are the averages of two currents activated by depolarizing
from -80 to 0 mV in the presence and absence of the inhibitors
indicated. C, the graph shows the effects of TEA,
Ba2+, and Cs+ (all 1 mM) on
KCa,V current amplitude. Current amplitudes were averaged
over 5 ms at the end of each depolarizing (-80 to 0 mV) voltage step.
Bars represent current amplitudes in the presence of the
inhibitors expressed as a percentage of the control current amplitudes.
Currents through KCa,V channels were inhibited by TEA
(n = 5, p < 0.005) and
Ba2+ (n = 4, p < 0.005)
but were not modulated by Cs+ (n = 4, p < 0.005). D, Ba2+ (1 mM)- and (E) Cs+ (1 mM)-inhibited KIR currents activated by
hyperpolarizing cells from -40 to -90 mV with equal K+
concentrations in the electrode and bath solutions (see "Experimental
Procedures"). Superimposed traces are the averages of two
currents recorded in the presence and absence of the inhibitors
indicated. F, inward rectifying KIR currents
were not sensitive to TEA (n = 6 cells) but were
inhibited by the application of Ba2+ (n = 6, p < 0.05) and Cs+ (n = 4, p < 0.005). p values were determined by
a paired sample t test. Current measurements were made at
the end of the hyperpolarizing step (as described in
C).
|
|
Cs+, Ba2+, and TEA also had distinct effects on
Ca2+ oscillations. TEA (1 mM) induced a marked
increase in the amplitude of Ca2+ oscillations, a small
increase in their frequency, and had no significant effect on base-line
[Ca2+]i (Fig. 5;
see Fig. 8). By contrast, Ba2+ (1 mM) induced a
marked increase in base-line [Ca2+]i as well as a
significant increase in the frequency of Ca2+ oscillations
and a small but significant increase in the amplitude of
Ca2+ oscillations (Fig. 6;
see Fig. 8). Cs+ caused a significant increase in the
frequency of Ca2+ oscillations as well as a slight increase
in base-line [Ca2+]i but had no significant
effect on the amplitude of Ca2+ oscillations (Fig.
7 and Fig.
8). Both Ba2+ and TEA
significantly increased average [Ca2+]i, whereas
Cs+ induced only a small increase in average
[Ca2+]i.

View larger version (50K):
[in this window]
[in a new window]
|
Fig. 5.
The effect of TEA on spontaneous
Ca2+ oscillations. The left panel is a
raster plot of [Ca2+]i (gray-scale)
versus time (x axis) in a microscopic field of 30 GH3 cells. Each row in the plot
represents an individual cell. The right panel shows
line tracings of [Ca2+]i
versus time for a representative cell (top trace)
as well as the average [Ca2+]i for the entire
field. In most cells, bath application of TEA evokes a marked increase
in the amplitude of Ca2+ oscillations (peak minus base-line
[Ca2+]i for each oscillation) and a slight
increase in base-line [Ca2+]i but does not
significantly alter the frequency of Ca2+ oscillations. The
average [Ca2+]i for the entire field is
significantly increased compared with base line.
|
|

View larger version (45K):
[in this window]
[in a new window]
|
Fig. 6.
The effect of Ba2+ on spontaneous
Ca2+ oscillations. The left panel shows
[Ca2+]i versus time in a field of 30 cells; the right panel shows line tracings of
[Ca2+]i versus time in a
representative cell (top) and the average
[Ca2+]i (bottom) for the entire field.
Bath application of 1 mM Ba2+ evokes a marked
increase in a base-line [Ca2+]i as well as an
increase in the frequency and amplitude of Ca2+
oscillations.
|
|

View larger version (40K):
[in this window]
[in a new window]
|
Fig. 7.
The effect of Cs+ on spontaneous
Ca2+ oscillations. The raster plot and line
tracings show the effect of bath application of Cs+ on
a field of GH3 cells. Cs+ evokes an increase in
the frequency of Ca2+ oscillations as well as a slight
increase in base-line [Ca2+]i but does not
significantly affect the amplitude of Ca2+ oscillations.
There is only a slight increase in the average
[Ca2+]i for the entire field.
|
|

View larger version (37K):
[in this window]
[in a new window]
|
Fig. 8.
The effects of K+ channel
antagonists on characteristics of Ca2+ oscillations.
Bars represent average percentage change in each of the
parameters from control (100%), determined on an individual cell basis
following bath application of 1 mM TEA (n = 60 cells from 3 experiments), Ba2+ (1 mM,
n = 70 cells from 3 experiments), or Cs+ (1 mM n = 60 cells from 3 experiments). TEA
and Cs+, both, evoked a slight but significant
(p < 0.005) increase in base-line
[Ca2+]i, whereas Ba2+ evoked a large
increase in base-line [Ca2+]i (p < 0.001) (left, average control base-line
[Ca2+]i values for TEA, Ba2+, and
Cs+ were 63 ± 28, 63 ± 32, and 67 ± 33 nM ± S.D., respectively). TEA evoked a large increase in
Ca2+ oscillation amplitude (p < 0.001),
Ba2+ evoked a slight but significant (p < 0.05) increase in Ca2+ oscillation amplitude, and
Cs+ did not have a significant effect on Ca2+
oscillation amplitude (middle, average control
Ca2+ oscillation amplitude values for TEA,
Ba2+, and Cs+ were 121 ± 48, 131 ± 52, and 158 ± 50 nM, respectively). Both
Ba2+ and Cs+ evoked a significant increase in
Ca2+ oscillation frequency (p < 0.005),
whereas TEA did not have a significant effect (right,
average control Ca2+ oscillation frequency values for TEA,
Ba2+, and Cs+ were 0.17 ± .06, 0.16 ± .04, and 0.14 ± .05 Hz, respectively). p values
were determined by a paired-sample t test.
|
|
These K+ channel antagonists also had distinct effects on
prolactin release (Fig. 9). TEA had no
significant effect on prolactin release. By contrast, Ba2+
evoked a large increase in prolactin release, whereas Cs+
evoked a small but significant increase in prolactin release.

View larger version (81K):
[in this window]
[in a new window]
|
Fig. 9.
The effects of K+ and
Ca2+ channel antagonists on prolactin release.
Histogram bars represent percent change in prolactin release
compared with control release measured immediately before antagonist
application (100%). Each value is the average of at least four
experiments. Control values for prolactin secretion in 0.5 h from
cells before their exposure to TEA, Ba2+, and
Cs+ were 2.6 ± 1.2, 1.7 ± 0.3, and 2.2 ± 0.1 ng/ml/106 cells, respectively. Both Ba2+ (1 mM) and Cs+ (1 mM) caused a
significant increase in spontaneous prolactin release
(p < 0.05), whereas nimodipine (10 µM)
significantly inhibited release (p < 0.05). TEA did
not have a significant effect on prolactin release. Error
bars represent S.E..
|
|
 |
DISCUSSION |
GH3 cells show spontaneous activity characterized by
rhythmic Ca2+ oscillations and prolactin release.
Ca2+ channels, K+ channels, Ca2+
influx, and release of intracellular Ca2+ play distinct
roles in this spontaneous activity. A number of previous studies have
demonstrated a primary role for release of Ca2+ from
intracellular stores in the spontaneous and stimulated activity of
pituitary cells (20, 23, 27-30). Our results show that spontaneous Ca2+ oscillations in GH3 cells are completely
dependent on influx of Ca2+ and in the majority of cells,
do not require release of intracellular Ca2+ from
thapsigargin-sensitive stores. The efficacy of thapsigargin in
depleting Ca2+ stores was verified by the complete
inhibition of the peak increase in [Ca2+]i in
response to thyrotropin-releasing hormone, as reported previously by
Nelson and Hinkle (31). The frequency and amplitude of Ca2+
oscillations were diminished in some cells in response to thapsigargin, suggesting that intracellular Ca2+ stores may play a
modulatory rather than a primary role in Ca2+ oscillations.
This modulatory role may involve Ca2+-induced
Ca2+ release through interaction of Ca2+ with
the IP3 receptor (28, 30, 32). Thapsigargin did abolish spontaneous Ca2+ oscillations in a significant proportion
of cells (approximately 30%), suggesting that there is a subset of
cells whose spontaneous Ca2+ signaling does require
intracellular Ca2+ release from thapsigargin-sensitive
stores. A similar dependence on IP3-sensitive intracellular
Ca2+ stores in a subset of GH3 cells is
suggested by Varney et al. (33), who found that chronic
treatment with Li2+, which reduces basal levels of
IP3, decreased the number of cells showing spontaneous
Ca2+ oscillations.
The effects of nimodipine indicate that spontaneous Ca2+
oscillations are generated by influx of Ca2+ through L-type
channels that have previously been shown to be inhibited by nimodipine
in GH3 cells (19). At lower concentrations, nimodipine
often reduced the frequency of Ca2+ oscillations without
significantly altering their amplitude. This observation suggests that
the frequency of Ca2+ oscillations is dependent upon the
activation state of Ca2+ channels, whereas the base-to-peak
amplitude of spontaneous Ca2+ oscillations is an
all-or-none phenomenon that is not regulated by the state of
Ca2+ channels under unstimulated conditions. The central
role of Ca2+ channels in the overall activity of the cell
makes them a logical target for inhibitory or excitatory modulation by
external ligands. The observations that somatostatin receptors and
expressed opioid receptors in GH3 cells modulate
Ca2+ channels is consistent with their role as a target for
receptor-mediated modulation (19, 34).
Nimodipine inhibited basal prolactin release by approximately 30%,
indicating that a significant proportion of basal prolactin release is
stimulated by spontaneous Ca2+ signaling. However, the
persistence of prolactin release in the absence of any detectable
Ca2+ signaling shows that a high proportion of basal
prolactin release is independent of spontaneous Ca2+
oscillations. This finding is consistent with the report of
Masumoto et al.(10), who found that basal
exocytosis continued in the absence of increases in
[Ca2+]i in pituitary gonadotrophs. It is likely
that this represents an unregulated pathway of secretion that has been
reported in GH3 cells (35) and other cell types.
Different K+ channel antagonists have distinct effects on
Ca2+ signaling in GH3 cells. TEA evoked a
dramatic increase in the amplitude of Ca2+ oscillations;
this effect is likely because of prolongation of the action potential
and subsequent increase in the action potential-induced influx of
Ca2+. This is consistent with the relatively selective
effect TEA has on sustained outward K+ currents recorded
from GH3 cells, with little effect on the A current (15).
Interestingly, the large increase in the amplitude of Ca2+
oscillations was not associated with a large increase in base-line [Ca2+]i, showing that base-line
[Ca2+]i is regulated by a mechanism other than
TEA-sensitive outward K+ currents. Both Ba2+
and Cs+ increased the frequency of Ca2+
oscillations, whereas TEA did not. These results indicate that the
inward rectifier K+ channel plays a primary role in setting
the frequency of spontaneous Ca2+ oscillations. They are
consistent with the results of Barros et al. (36, 37), who
report a primary role for the inward rectifier in the regulation of
GH3 cell excitability. Base-line [Ca2+]i was increased dramatically by
Ba2+, and to a lesser extent, by Cs+ and TEA.
These results suggest that Ba2+ modulates base-line
[Ca2+]i by a mechanism that is distinct from the
inward rectifier, and as discussed above, distinct from TEA-sensitive
outward K+ currents.
Prolactin release was also differentially modulated by different
K+ channel antagonists. Despite the fact that TEA caused a
marked increase in the amplitude of Ca2+ oscillations and a
significant increase in average [Ca2+]i, it did
not increase prolactin release. By contrast, Cs+, which
increased the frequency of Ca2+ oscillations but caused a
much smaller increase in average [Ca2+]i than
TEA, did cause a significant increase in prolactin release. These
findings are consistent with the stimulation of hormone release by an
increase in the frequency of spontaneous Ca2+ oscillations
but not in their amplitude. The marked effect of Ba2+
suggests that the combination of increased base-line
[Ca2+]i and increased Ca2+
oscillation frequency can cause a much greater increase in hormone release than increased Ca2+ oscillation frequency alone.
Simultaneous measurements of membrane capacitance and
[Ca2+]i by multiple investigators have provided
strong evidence that each oscillatory increase in
[Ca2+]i is capable of evoking secretion of
hormone (24, 38-42). Our results suggest that increasing the peak
[Ca2+]i associated with each spontaneous
Ca2+ oscillation does not increase hormone release; this
may be because there is a maximum number of vesicles released per
Ca2+ oscillation once the peak
[Ca2+]i has reached a certain threshold or
because buffering of Ca2+ prevents this increased peak
[Ca2+]i from being sensed by the secretory
apparatus. By contrast, the increase in hormone release associated with
an increase in Ca2+ oscillation frequency suggests that
modulation of Ca2+ oscillation frequency does represent a
mechanism for stimulation of hormone release. Increasing base-line
[Ca2+]i, in addition to increasing
Ca2+ oscillation frequency, results in a further increase
in prolactin release. Possible explanations for the specific effects of
base-line [Ca2+]i and oscillation frequency on
secretion include direct effects on the exocytotic trigger mechanism
(43) or the recruitment of additional pools of vesicles with different
Ca2+ sensitivities (44, 45).
Ca2+ channels, K+ channels, Ca2+
influx, and intracellular Ca2+ release play specific roles
in the generation of spontaneous activity in GH3 cells. In
turn, specific characteristics of spontaneous Ca2+
signaling, namely base-line [Ca2+]i and the
frequency of Ca2+ oscillations, are correlated with changes
in hormone release. Our studies identify L-type Ca2+
channels and the inward rectifier K+ channel as key
components of the cellular signaling machinery whose modulation results
in the specific changes in the patterns of spontaneous cellular
activity that regulate changes in basal prolactin release.
TEA-sensitive outward K+ currents may affect other cellular
processes through amplitude-modulated Ca2+ signaling.
 |
FOOTNOTES |
*
This work was supported by National Institutes of Health
(NINDS) Grants R29 NS32283 and P01-NS 02808 (to A. C. C.) and
National Institute on Drug Abuse Grants DA05010 (to T. G. H. and
C. J. E.) and DA05627 (to E. T. P.).The costs of publication of this article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement" in accordance with 18 U.S.C. Section
1734 solely to indicate this fact.
§
To whom correspondence should be addressed: UCLA Dept. of
Neurology, 710 Westwood Plaza, Los Angeles, CA 90095. Tel.:
310-794-1870; Fax: 310-794-1871; E-mail: acharles{at}ucla.edu.
 |
ABBREVIATIONS |
The abbreviations used are:
IP3, inositol 1,4,5-trisphosphate;
ELISA, enzyme-linked immunosorbent assay;
TEA, tetraethylammonium.
 |
REFERENCES |
-
Berridge, M. J.
(1997)
Nature
386,
759-760[CrossRef][Medline]
[Order article via Infotrieve]
-
Dolmetsch, R. E.,
Lewis, R. S.,
Goodnow, C. C.,
and Healy, J. I.
(1997)
Nature
386,
855-858[CrossRef][Medline]
[Order article via Infotrieve]
-
De Koninck, P.,
and Schulman, H.
(1998)
Science
279,
227-230[Abstract/Free Full Text]
-
Schlegel, W.,
Winiger, B. P.,
Mollard, P.,
Vacher, P.,
Wuarin, F.,
Zahnd, G. R.,
Wollheim, C. B.,
and Dufy, B.
(1987)
Nature
329,
719-721[CrossRef][Medline]
[Order article via Infotrieve]
-
Wagner, K. A.,
Yacono, P. W.,
Golan, D. E.,
and Tashjian, A. H., Jr.
(1993)
Biochem. J.
292,
175-182[Medline]
[Order article via Infotrieve]
-
Scherubl, H.,
Hescheler, J.,
Bychkov, R.,
Cuber, J. C.,
John, M.,
Riecken, E. O.,
and Wiedenmann, B.
(1994)
Ann. N. Y. Acad. Sci.
733,
335-339[Abstract]
-
Li, Y. X.,
Rinzel, J.,
Vergara, L.,
and Stojilkovic, S. S.
(1995)
Biophys. J.
69,
785-795[Abstract]
-
Malgaroli, A.,
and Meldolesi, J.
(1991)
FEBS Lett.
283,
169-172[CrossRef][Medline]
[Order article via Infotrieve]
-
Miki, H.,
Maercklein, P. B.,
and Fitzpatrick, L. A.
(1995)
Endocrinology
136,
2954-2959[Abstract]
-
Masumoto, N.,
Tasaka, K.,
Mizuki, J.,
Fukami, K.,
Ikebuchi, Y.,
and Miyake, A.
(1995)
Cell Calcium
18,
223-231[Medline]
[Order article via Infotrieve]
-
Shangold, G. A.,
Kongsamut, S.,
and Miller, R. J.
(1985)
Life Sci.
36,
2209-2215[Medline]
[Order article via Infotrieve]
-
Matteson, D. R.,
and Armstrong, C. M.
(1986)
J. Gen. Physiol.
87,
161-182[Abstract]
-
Simasko, S. M.,
Weiland, G. A.,
and Oswald, R. E.
(1988)
Am. J. Physiol.
254,
E328-E336[Abstract/Free Full Text]
-
Rogawski, M. A.
(1989)
Mol. Pharmacol.
35,
458-468[Abstract]
-
Oxford, G. S.,
and Wagoner, P. K.
(1989)
J. Physiol. (Lond)
410,
587-612[Abstract]
-
Dubinsky, J. M.,
and Oxford, G. S.
(1985)
Proc. Natl. Acad. Sci. U. S. A.
82,
4282-4286[Abstract]
-
Simasko, S. M.
(1991)
Am. J. Physiol.
261,
E66-E75[Abstract/Free Full Text]
-
Barros, F.,
Delgado, L. M.,
del Camino, D.,
and de la Pena, P.
(1992)
Pfluegers Arch. Eur. J. Physiol.
422,
31-39[Medline]
[Order article via Infotrieve]
-
Piros, E. T.,
Prather, P. L.,
Loh, H. H.,
Law, P. Y.,
Evans, C. J.,
and Hales, T. G.
(1995)
Mol. Pharmacol.
47,
1041-1049[Abstract]
-
Kukuljan, M.,
Rojas, E.,
Catt, K. J.,
and Stojilkovic, S. S.
(1994)
J. Biol. Chem.
269,
4860-4865[Abstract/Free Full Text]
-
Gollasch, M.,
Haller, H.,
Schultz, G.,
and Hescheler, J.
(1991)
Proc. Natl. Acad. Sci. U. S. A.
88,
10262-10266[Abstract]
-
Bauer, C. K.,
Davison, I.,
Kubasov, I.,
Schwarz, J. R.,
and Mason, W. T.
(1994)
Pfluegers Arch. Eur. J. Physiol.
428,
17-25[Medline]
[Order article via Infotrieve]
-
Tse, F. W.,
Tse, A.,
Hille, B.,
Horstmann, H.,
and Almers, W.
(1997)
Neuron
18,
121-132[Medline]
[Order article via Infotrieve]
-
Tse, A.,
Tse, F. W.,
Almers, W.,
and Hille, B.
(1993)
Science
260,
82-84[Medline]
[Order article via Infotrieve]
-
Charles, A. C.,
Merrill, J. E.,
Dirksen, E. R.,
and Sanderson, M. J.
(1991)
Neuron
6,
983-992[Medline]
[Order article via Infotrieve]
-
Ritchie, A. K.
(1987)
J. Physiol. (Lond.)
385,
611-625[Abstract]
-
Kukuljan, M.,
Vergara, L.,
and Stojilkovic, S. S.
(1997)
Biophys. J.
72,
698-707[Abstract]
-
Li, Y. X.,
Keizer, J.,
Stojilkovic, S. S.,
and Rinzel, J.
(1995)
Am. J. Physiol.
269,
C1079-C1092[Abstract/Free Full Text]
-
Stojilkovic, S. S.,
Tomic, M.,
Kukuljan, M.,
and Catt, K. J.
(1994)
Mol. Pharmacol.
45,
1013-1021[Abstract]
-
Tse, F. W.,
Tse, A.,
and Hille, B.
(1994)
Proc. Natl. Acad. Sci. U. S. A.
91,
9750-9754[Abstract/Free Full Text]
-
Nelson, E. J.,
and Hinkle, P. M.
(1994)
Endocrinology
135,
1084-1092[Abstract]
-
Tse, A.,
Tse, F. W.,
and Hille, B.
(1994)
J. Physiol. (Lond.)
477,
511-525[Abstract]
-
Varney, M. A.,
Galione, A.,
and Watson, S. P.
(1994)
Br. J. Pharmacol.
112,
390-395[Abstract]
-
Piros, E. T.,
Prather, P. L.,
Law, P. Y.,
Evans, C. J.,
and Hales, T. G.
(1996)
Mol. Pharmacol.
50,
947-956[Abstract]
-
Varro, A.,
Nemeth, J.,
Dickinson, C. J.,
Yamada, T.,
and Dockray, G. J.
(1996)
Biochim. Biophys. Acta
1313,
101-105[Medline]
[Order article via Infotrieve]
-
Barros, F.,
Villalobos, C.,
Garcia-Sancho, J.,
del Camino, D.,
and de la Pena, P.
(1994)
Pfluegers Arch. Eur. J. Physiol.
426,
221-230[Medline]
[Order article via Infotrieve]
-
Barros, F.,
del Camino, D.,
Pardo, L. A.,
and de la Pena, P.
(1996)
Pfluegers Arch. Eur. J. Physiol.
431,
443-451[Medline]
[Order article via Infotrieve]
-
Thomas, P.,
and Waring, D. W.
(1997)
J. Physiol. (Lond.)
504,
705-719[Abstract]
-
Maruyama, Y.,
Inooka, G.,
Li, Y. X.,
Miyashita, Y.,
and Kasai, H.
(1993)
EMBO J.
12,
3017-3022[Abstract]
-
Engisch, K. L.,
Chernevskaya, N. I.,
and Nowycky, M. C.
(1997)
J. Neurosci.
17,
9010-9025[Abstract/Free Full Text]
-
Chow, R. H.,
Klingauf, J.,
Heinemann, C.,
Zucker, R. S.,
and Neher, E.
(1996)
Neuron
16,
369-376[Medline]
[Order article via Infotrieve]
-
Mansvelder, H. D.,
and Kits, K. S.
(1998)
J. Neurosci.
18,
81-92[Abstract/Free Full Text]
-
Heinemann, C.,
von Ruden, L.,
Chow, R. H.,
and Neher, E.
(1993)
Pfluegers Arch. Eur. J. Physiol.
424,
105-112[Medline]
[Order article via Infotrieve]
-
Horrigan, F. T.,
and Bookman, R. J.
(1994)
Neuron
13,
1119-1129[Medline]
[Order article via Infotrieve]
-
Giovannucci, D. R.,
and Stuenkel, E. L.
(1997)
J. Physiol. (Lond.)
498,
735-751[Abstract]
Copyright © 1999 by The American Society for Biochemistry and Molecular Biology, Inc.