Department of Pharmacology, University of Virginia School of Medicine, Charlottesville, Virginia 22908
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ABSTRACT |
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Independently, plasma K+ and ANG II stimulate aldosterone secretion from adrenal glomerulosa (AG) cells, but together they synergistically control production. We studied mechanisms to mediate this synergy using bovine AG cells studied under physiological conditions (in 1.25 mM Ca2+ at 37°C). Increasing K+ from 2 to 5 mM caused a potentiation of ANG II-induced aldosterone secretion and a substantial membrane depolarization (~21 mV). ANG II inhibited a K+-selective conductance in both 2 and 5 mM K+ but caused only a slight depolarization because, under both conditions, membrane potential was close to the reversal potential of the ANG II-induced current. ANG II activated calcium/calmodulin-dependent protein kinase II (CaMKII) equivalently in 2 and 5 mM K+. However, CaMKII activation caused a hyperpolarizing shift in the activation of T-type Ca2+ channels, such that substantially more current was elicited at membrane potentials established by 5 mM K+. We propose that synergy in aldosterone secretion results from K+-induced depolarization and ANG II-induced modulation of T-type channel activation, such that together they promote enhanced steady-state Ca2+ flux.
low-voltage-activated calcium channels; calcium/calmodulin-dependent protein kinase II; membrane potential; potassium channels
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INTRODUCTION |
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EXTRACELLULAR K+ and ANG II are powerful independent regulators of aldosterone secretion from the adrenal zona glomerulosa (17). However, the potency of these agonists in the physiological control of aldosterone secretion depends on their combined activities and their synergy. For example, when circulating ANG II is low, K+ is a less potent stimulator of aldosterone production. This is observed in anephric patients (42), patients with hyporeninemic hypoaldosteronism, whose aldosterone output is low despite an elevated level of serum K+ (50), and patients with primary aldosteronism, who are less responsive to changes in serum K+ (21). Conversely, the potency of ANG II as a stimulator of aldosterone secretion depends on the prevailing concentration of extracellular K+. Thus the hypokalemia associated with Bartter's syndrome or the misuse of diuretics is accompanied by levels of plasma aldosterone that are inappropriately low for the observed high levels of plasma renin activity (52). In contrast, K+ supplementation given to healthy human subjects on a low-Na+ diet can effect a sustained increase in aldosterone output, despite an accompanying reduction in plasma renin activity (26). K+, as a potentiator of ANG II-elicited aldosterone secretion, has been amply documented in normal humans, dogs, and sheep, in which K+ infusions that result in only small elevations in plasma K+ elicit large and proportional increases in aldosterone (6, 7, 14, 15, 18, 41). In vitro, the combination of physiological concentrations of K+ and ANG II also evokes a greater aldosterone secretory response from isolated glomerulosa cells than either stimulus alone, indicating that the synergistic control by ANG II and K+ in the control of aldosterone production is mediated at the level of the glomerulosa cell (20, 43). K+ does not potentiate ANG II-stimulated aldosterone secretion by increasing the affinity or the number of ANG II receptors (20). However, potentiation may be mediated by an elevation in intracellular Ca2+ (43). In bovine glomerulosa cells, ANG II induces a sustained increase in cytosolic Ca2+ that occurs during prolonged stimulation. This elevation in cytosolic Ca2+ is dependent on extracellular Ca2+ and rises with the concentration of extracellular K+. Although these findings suggest that coordinated effects on Ca2+ influx may account for the observed synergistic effect on aldosterone secretion, the mechanism underlying K+-induced potentiation of ANG II-stimulated aldosterone secretion remains undefined.
We show that the membrane potential of acutely dispersed AG cells in physiological K+ solutions follows closely the K+ equilibrium potential; the membrane potential depolarizes strongly with slight elevations in K+, but changes only modestly with ANG II. We show further that calcium/calmodulin (Ca2+/CaM)-dependent protein kinase II (CaMKII) is activated by ANG II and that this CaMKII activation induces a hyperpolarizing shift in the V1/2 (i.e., the potential corresponding to half-maximal activation) of activation of the low-voltage-activated T-type Ca2+ channel, the major voltage-dependent Ca2+ entry pathway in AG cells. These results suggest a mechanism by which Ca2+ entry can be enhanced jointly by ANG II and K+, via the modulation of CaMKII and membrane potential, and which may underlie the synergistic control exerted by these stimuli in their control of aldosterone secretion.
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METHODS |
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Cell isolation. Neonatal bovine AG cells (2-7 days old) were prepared by collagenase digestion, as described previously (19, 33, 34). Thin layers of zona glomerulosa were prepared from adrenal cortex and collected in a Ca2+-free Krebs-Ringer bicarbonate (KRB) that contained (in mM) 120 NaCl, 25 NaHCO3, 3.5 KCl, 1.2 MgSO4, 1.2 NaH2PO4, 1.25 CaCl2, 0.1% dextrose, and 0.2% BSA, equilibrated with 95% air-5% CO2 (pH 7.4). The slices were digested for 10 min at 37°C in KRB containing 0.625 mM CaCl2, 0.1% BSA, and 0.35 U/ml collagenase (Worthington, Freehold, NJ). The cells were dispersed mechanically, filtered through a 20-µm nylon mesh filter (Tetko, Elmsford, NY), collected by centrifugation, and resuspended in KRB containing 1.25 mM CaCl2 and 0.2% BSA. This procedure was repeated once. Second digestion cells were collected and further purified on a two-step (30%/56%) discontinuous Percoll gradient (Pharmacia, Piscataway, NJ) and resuspended in standard KRB buffer. Cells were plated to glass coverslips in media (1:1, DMEM:F12) containing 100 U/ml penicillin and 100 µg/ml streptomycin. All patch-clamp studies were performed within 24 h of isolation.
Recording. The methods used for
electrophysiological experiments have been described previously (33,
34). Briefly, patch electrodes with resistances of 2-4 M were
fabricated from capillaries of 1724 (Garner, Claremeon, CA) or 0010 (World Precision Instruments, Sarasota, FL) glass. AG cells adherent to
noncoated glass coverslips were transferred to a 500-µl
microincubator recording chamber (Medical Systems, Greenvale, NY),
which was continuously perfused by gravity at a rate of 0.5 ml/min.
Typical cells, having diameters of 12-18 µm, were voltage
clamped at 33-37°C using the whole cell configuration of the
patch-clamp technique. Current was sampled at 12.5, 4, or 1 kHz and
filtered with an eight-pole low-pass Bessel filter (Frequency Devices,
Haverhill, MA) set at a cutoff frequency (
3 dB) of 2.5, 2, or
0.25 kHz, respectively. Pulse generation and data acquisition were
performed using a ZEOS 486 computer with pCLAMP 6.0 software and an
Axolab interface (Axon Instruments, Foster City, CA). We recorded
resting membrane potential (Em) in current
clamp and corroborated in voltage clamp by determining the zero-current
potential from current-voltage (I-V)
relationships. Under whole cell voltage clamp, macroscopic current was
elicited by applying voltage ramp commands to
140 mV from a
holding potential of
50 mV (0.095 V/s), delivered at intervals
of 10 s. For each cell, two to five episodes were averaged for control
and experimental groups. In preliminary experiments, membrane
potentials determined from hyperpolarizing and depolarizing ramp
protocols were equivalent [
97.9 ± 3.8 vs.
92.1 ± 3.5; n = 8, P = not significant (NS), according to
paired 2-tailed Student's
t-test], and the hyperpolarizing voltage-ramp protocol provided current-voltage curves that were similar
to steady-state I-V curves constructed
from voltage-step commands (100 ms). Under whole cell voltage clamp, we
elicited Ca2+ currents by applying
voltage-step commands from a holding potential of
95 mV. Tail
currents were elicited on repolarization to
80 mV
(Vr) after a
10.4-ms test pulse from
75 to
5 mV, at intervals of 6 s.
Linear leak and capacitative transient currents were subtracted digitally using scaled hyperpolarizing steps of 1/4 amplitude. Liquid
junction potentials were measured as described (5), and command
potentials were corrected for the liquid junction potential associated
with each set of bath and pipette solutions. With 1.25 mM
Ca2+ as the charge carrier,
Ca2+-channel tail currents were
fitted by one exponential plus a constant using the Levenberg-Marquardt
nonlinear curve-fitting algorithm. We blanked 250 µs of the 4-ms
fitting region to eliminate any possible contamination with L-type
Ca2+-channel currents. The
reported tail current values corresponded to amplitudes obtained from
the exponential function at the end of the blanking period. Tail
currents at Vr
were monoexponential, decaying with a time constant of 0.95 ± 0.09 ms, and were eliminated by prepulse to
40 mV. We determined the
voltage dependence of activation of T-type
Ca2+ channels by plotting the
relative amplitude of the slowly deactivating component of the tail
current vs. the test potential. For each cell, the data set (15 test
potentials in 5-mV increments, from
75 to
5 mV; 10.4 ms)
was fit by a two-state Boltzmann distribution given by the equation
I/Imax = {1 + exp[(V1/2
Vt)/k])
1},
where k is the slope factor
(mV/e-fold change),
Vt is the test potential, and
Imax is the
maximal elicitable current.
V1/2 values for
all cells within a group were averaged. The dependence of channel
inactivation on voltage was determined in response to depolarization by
holding at various potentials (
100 to
35 mV; 5-mV
increments) for 6 s to effect a steady-state level of inactivation and
by application of brief depolarizing test pulses (+15 mV; 8 ms) to open
available channels, followed by repolarization to
80 mV. The
relative amplitude (h) of the slowly
deactivating component of the tail current vs. the test potential was
fitted to the equation h = I/Imax = {(1 + exp[(Vp
V1/2)/k])
1},
where Vp is the
prepulse potential and all remaining parameters are as defined above.
V1/2 values for
all cells within a group were averaged. We evaluated statistical
differences using paired or nonpaired (where appropriate) two-tailed
Student's t-test. We assumed
significance when P
0.05.
Solutions. For current clamp recordings, we used an external (bath) HEPES-based Ringer solution that contained (in mM) 140 NaCl, 2-5 KCl, 1.25 CaCl2, 2 MgCl2, 10 glucose, and 10 HEPES-NaOH, pH 7.3 (with NaOH). The internal (pipette) solution contained (in mM) 17.5 KCl, 122.5 potassium gluconate, 9 NaCl, 1 MgCl2, 5 Mg-ATP, 1 Mg-GTP, 0.2 EGTA, and 10 HEPES-KOH, pH 7.2 (with KOH). For the recording of Ca2+ currents, the external (bath) solution contained (in mM) 134 tetraethylammonium chloride (TEA-Cl), 1.25 CaCl2, 2 MgCl2, 10 HEPES-Cs, 5 dextrose, and 32 sucrose, pH 7.3 (with CsOH). The internal (pipette) solution contained 95 mM CsCl, 1 mM tetrabutylammonium chloride (TBA-Cl), 1 mM MgCl2, 5 mM Mg-ATP, 1.0 mM Mg-GTP, 20 mM HEPES/Cs, 11 mM 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA), 0.2 µM calmodulin, and CaCl2 (see below), pH 7.2 (with CsOH). All solutions were filtered through 0.22-µm nitrocellulose filters (Micron Separations, Westboro, MA). Free Ca2+ in the pipette solution (Ca2+i) was calculated using the ligand-binding program EQCAL (Biosoft, Ferguson, MO), with the following Kd values: BAPTA, 190 nM (for 0.15 M ionic strength) (23); and calmodulin, 33, 0.7, 25, and 0.4 µM (28). With BAPTA (11 mM) and calmodulin (0.2 µM) in the pipette solution, added CaCl2 (in mM) fixed the Ca2+i at 27 nM (0.9), 240 nM (4.9), 800 nM (8.0), and 1.2 µM (8.8).
Preparation of samples for CaMKII phosphotransferase
assays. After successive 30-min equilibrations at room
temperature and 37°C, AG cells were resuspended at 5 × 106 cells/ml of KRB containing 3.5 mM K+ and preincubated for an
additional 30 min at 37°C. Incubations with agonist were initiated
by the addition of an equal volume of KRB, to give 2.0 or 5.0 mM
K+ (isosmotic replacement) and
0.1-10 nM ANG II, and continued for 30 s at 37°C. At the end
of treatment, cells were briefly (5 s) pelleted by centrifugation,
resuspended in ice-cold lysis buffer (at 5 × 106 cells/ml), and immediately
sonicated in an ice bath (2 × 5 s; 140 W with an XL2020
sonicator). The lysis buffer contained 50 mM HEPES, pH 7.5, 50 mM
-glycerophosphate, 4 mM EGTA, 4 mM EDTA, 0.1 µM microcystin LR
(Calbiochem, San Diego, CA), 1 mM benzamidine, 25 µg/ml leupeptin, 25 µg/ml aprotinin, and 0.1% Triton X-100. The resulting sonicate was
held on ice for immediate use or frozen in liquid
N2 and stored at
80°C.
The protein concentration of the whole cell extract was 0.4-0.9
mg/ml, as determined by the method of Pierce using BSA as a standard
(51).
CaMKII phosphotransferase assay.
CaMKII activity was measured by modification of a previously described
protocol (19). In the assay for
Ca2+/CaM-dependent (or total)
CaMKII activity, the standard kinase buffer contained 50 mM HEPES, 0.5 mM EGTA, 0.1 mg/ml BSA, 10 magnesium acetate, 25 µM
[-32P]ATP (3-6
Ci/mmol), 10 µM autocamtide-2, 1.81 mM
CaCl2, and 1.4 µM CaM, in a
final reaction volume of 50 µl. We calculated the free
Ca2+ concentration in this buffer
to be 100 µM using the ligand-binding program EQ-CAL (Biosoft,
Ferguson, MO). To assay for
Ca2+/CaM-independent (or
autonomous) CaMKII activity, we omitted
CaCl2 and CaM.
The kinase reaction was initiated by the addition of 10 µl of AG cell sonicate (2.5-5 µg of protein) and continued for 40 min at 4°C. The reaction was terminated by the addition of trichloroacetic acid to 5%, followed by centrifugation at 14,000 g for 1 min at 4°C to remove large, precipitated proteins. We spotted 50 µl of the resulting supernatant onto a strip of P81 phosphocellulose paper (Whatman, Hillsboro, OR). P81 strips were washed in distilled H2O, dehydrated in 100% ethanol, and air dried; then radioactivity was quantified by liquid scintillation counting. We determined 32P incorporation into autocamtide-2 by subtracting background counts (from assays in the absence of substrate) from total counts. CaMKII activity measured in this manner was linear with respect to time and protein concentration. Under these assay conditions, CaMKII activity in the presence of 100 µM free Ca2+ and 1.4 µM CaM was 62 ± 4 pmol Pi incorporated per minute per milligram of protein (n = 10). Ca2+/CaM-independent (autonomous) CaMKII activity was calculated as a percentage of Ca2+/CaM-dependent CaMKII activity.
Aldosterone radioimmunoassay. After
overnight storage at 4°C (in a 95% air-5%
CO2 evacuated container), AG cells
were equilibrated and preincubated at 37°C. Incubations (0.5 × 106 cells/ml of KRB) with
agonists were initiated on the addition of an equal volume of KRB to
give 2 or 5 mM K+ with and without
ANG II (from 10 pM to 10 nM). After 0.5 or 1 h incubation, cells were
placed on ice to prevent further steroidogenesis and centrifuged at
1,000 g for 10 min at 4°C. Medium
was removed from the pelleted cells and stored at 20°C.
Aldosterone content in the samples was measured by radioimmunoassay
(Diagnostic Products, Los Angeles, CA) using a specific antibody.
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RESULTS |
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Potentiation of ANG II-stimulated aldosterone
secretion by extracellular
K+. Figure
1 shows that 5 mM extracellular
K+ increases the secretory
response to physiological concentrations of ANG II. We measured
aldosterone secretion at 0.5 and 1 h of incubation at the extremes of
plasma K+ that are encountered
physiologically. In Fig. 1, the sustained rate of aldosterone secretion
(0.5-1 h of stimulation) was calculated at 2 and 5 mM
extracellular K+. At 2 mM
K+, ANG II elicited a
dose-dependent increase in aldosterone secretion, stimulating
production ~13-fold at 10 nM ANG II, with an
ED50 of 1 nM. In the absence of
ANG II, raising extracellular K+
from 2 to 5 mM increased secretion fourfold. Importantly, in 5 mM
K+, the secretory response to ANG
II was markedly potentiated, especially at the lower doses. Thus 30 pM
ANG II, which elicited <20% of a maximal response in 2 mM
K+, increased aldosterone
secretion to >80% when extracellular
K+ was 5 mM. The secretory
response elicited by supraphysiological doses of ANG II (10 nM) was not
potentiated in 5 mM K+. Thus
ambient K+ modulates the secretory
potential of ANG II, but only at physiological concentrations. These
data mimic the amplification elicited by in vivo
K+-infusion studies (6, 15, 18)
and confirm in vitro cell studies using canine and adult bovine AG
cells (20, 43).
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Effect of
K+ and ANG II on
membrane potential. The membrane potential of the AG
cell is largely determined by the
K+ equilibrium potential (44).
Figure 2 shows that the membrane potential
is strongly influenced by extracellular
K+. To maintain the putative
hyperpolarizing influence of the Na-K-ATPase pump on membrane
potential, and thus establish a membrane potential relevant to
secretory activity in vivo, we performed our recordings at
33-37°C. At 2 mM extracellular
K+, the resting membrane potential
(Em) of the AG
cell was 97.5 ± 1.8 mV (n = 4). Under current-clamp recording conditions, bath perfusion with 5 mM
K+ caused a large maintained
depolarization (~21 mV) associated with a decreased input resistance
(data not shown), with a prompt repolarization to resting values on
reperfusion with 2 mM K+ (Fig.
2A). All cells studied responded to
5 mM K+, with a depolarization to
77.0 ± 1.5 mV (n = 4); no cells showed evidence of spontaneous action potentials at
either 2 or 5 mM K+. Voltage-ramp
protocols were used to corroborate the measurements of membrane
potential made under current clamp. As illustrated in Fig.
2B, macroscopic current demonstrated a
nonlinear dependence on voltage that was weakly outwardly rectifying
over the voltage range of
140 to
50 mV, not unlike the
I-V curves recorded from acutely
dissociated rat AG cells that had been maintained in culture for <24
h. Increasing extracellular K+
from 2 to 5 mM increased the overall slope conductance of the I-V relationship and shifted the zero
current potential
(Em) toward more positive voltages. The zero current potential was
97.5 ± 1.7 mV (n = 5) in 2 mM
K+ and
78.0 ± 1.4 mV
(n = 5) in 5 mM
K+; these values for
Em in 2 and 5 mM
K+ are indistinguishable from
those made under current-clamp recording conditions (
98 and
77 mV). Moreover, these values are near the predicted
equilibrium potential for K+
[EK =
114 mV (2 mM K+);
90 mV (5 mM K+)]
indicating that, at 37°C, the membrane potential of the AG cell is
determined largely by a
K+-selective conductance(s).
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Inhibition of a resting K+
conductance by ANG II has been hypothesized to mediate aldosterone
secretion by causing a membrane depolarization and the consequent
opening of voltage-dependent Ca2+
channels (31, 38, 46). To date, five different types of K+ currents have been identified
in zona glomerulosa cells (transient A current, delayed rectifier,
inward rectifier, Ca2+-activated
current, and leak current) with the relative expression of each
dependent on species (rat vs. bovine vs. human) and culturing conditions (8, 27, 38, 39, 44, 53). In cultures of both rat and bovine
AG cells, ANG II inhibits inward rectifier and delayed rectifier
K+ channels (27, 30, 38), whereas
in freshly dispersed rat AG cells, leak channels are the target of ANG
II action (30). However, the functional importance of
K+ channel inhibition (i.e., the
degree of depolarization induced by a change in
K+ conductance) will depend
critically on the net driving force on
K+ ions (i.e., the difference
between the Em of
the cell and
EK) and the
conductance of other ions relative to
K+. Therefore, we examined
1) whether ANG II induces a change
in K+ conductance that results in
a membrane depolarization and 2) whether the depolarization is influenced by different physiological K+ solutions. As depicted in Fig.
3A, 10 nM
ANG II reduced whole cell conductance at all potentials examined when
applied in a bath containing 2 mM
K+. To quantify the
effect of ANG II on resting membrane conductance, we fitted the
I-V relationship by linear regression
(±5.0 mV of the zero-current potential under control conditions).
ANG II decreased the resting membrane conductance by 38.6 ± 7.3%
(n = 6, P 0.02). The intersection of the
I-V relationships recorded with and
without ANG II represents the reversal potential of the ANG
II-sensitive current. In 2 mM K+,
the ANG II-modulated current reversed at
107 ± 4.6 mV
(n = 4), a value that is very close to
the Em of the
cell (
98 mV). Thus, given such a small driving force for
K+ movement
(Erev
Em =
9 mV,
where Erev is the
reversal potential), the ANG II-induced reduction in conductance would
be expected to have minimal effects on membrane potential. This
prediction was substantiated in current-clamp experiments, in which we
found that membrane potential depolarized only slightly after exposure to ANG II (n = 4). Data pooled from
voltage-ramp and current-clamp measurements (Fig.
3B) indicate that ANG II, after 2.5 min, depolarized the cells only 3-5 mV (from
98.5 ± 1.6 mV to
95.6 ± 1.8 mV; n = 11, P
0.01). This effect was stable
with time, as values obtained after 5 min of exposure were identical
(96.8 ± 4.2 mV; n = 5). Therefore,
at 37°C in 2 mM K+, the
membrane potential of the AG cell was close enough to
Erev that the ANG
II-induced change in conductance resulted in only a small depolarizing
net current.
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In 5 mM K+ (Fig.
4A), ANG
II also induced a decrease in conductance over the entire voltage
range, reducing the resting membrane conductance by 44 ± 7.0%
(n = 5, P 0.05). The
Erev of the ANG II-modulated current in 5 mM K+
was
87 ± 3.6 mV (n = 4), a
value that is ~20 mV depolarized to
Erev in 2 mM
K+ (
107 mV); this value is
very close to EK
in 5 mM K+ (
87 mV), but
also close to Em
(
78 mV) measured in 5 mM
K+. A
K+-induced shift in the
Erev of the ANG
II-modulated component of current is consistent with modulation of a
K+ conductance, although the shift
was somewhat less than that predicted by the Nernst equation for an
exclusively K+-selective
conductance (~24 mV). Figure 4 also shows that, in 5 mM
K+, the ANG II-induced inhibition
of this K+ conductance resulted in
a small depolarizing current that was sufficient to depolarize the cell
~6 mV (from
79.8 ± 1.4 mV to
74.0 ± 2.0 mV,
n = 11, P
0.05) 2.5 min after ANG II
exposure. This depolarizing effect of ANG II was maximal at 2.5 min, as values obtained after 5 min of ANG II exposure were identical (
75.5 ± 2.4 mV; n = 5). In
addition, the inclusion of calmodulin to the pipette solution to
promote the activation of CaM- or CaMKII-dependent conductances did not
augment this depolarization (
75.9 ± 1.9 mV;
n = 9). Thus, because membrane
potential remains close to the
Erev of the ANG
II-modulated conductance in 5 mM
K+, modulation of this
K+-selective conductance by ANG II
remains only weakly depolarizing. As is evident in our
I-V plots (Fig.
4A), current injection to move the
membrane potential of the AG cell away from the
Erev of the ANG
II-sensitive current would be predicted to increase the driving force
for K+ movement and amplify the
depolarizing effect of ANG II. This prediction finds support in
previously published studies using rat AG cells, in which the magnitude
of the depolarization induced by ANG II was shown to be dependent on
membrane voltage (45). Nevertheless, these data refute the frequently
quoted hypothesis that a major component of the mechanism of action of
ANG II in the physiological control of steroidogenesis is the opening
of voltage-dependent Ca2+ channels
via ANG II-induced membrane depolarization (38, 46, 52).
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T-type
Ca2+ channel
currents.
It has been hypothesized that steady-state current, through
low-voltage-activated, T-type Ca2+
channels, contributes to the Ca2+
signal that is critical to sustaining stimulated aldosterone secretion
in AG cells (3, 48). Yet, to date, experiments on T-type currents have
been performed under nonphysiological conditions designed to optimize
the amplitude and kinetic isolation of these currents. Therefore,
because of charge screening and/or binding under conditions of high
extracellular Ca2+ and recordings
made at room temperature, the reported voltage range over which these
window currents are elicited was, at best, 20 mV more depolarized than
values for Em
previously recorded for AG cells (3, 12, 39, 48). Here, we recorded
Ca2+ currents at ~37°C,
using 1.25 mM Ca2+ as the charge
carrier, to determine if T-type
Ca2+ channels activate over the
range of membrane potentials established by physiological
K+ solutions. Because inward
Ca2+ currents are small with
reduced charge carrier and because the open-channel
I-V relationship for T-type
Ca2+ channels is highly nonlinear,
we used tail currents, rather than peak inward currents, to quantify
the voltage dependence of activation and inactivation of T-type
Ca2+ channels (25,
37). Tail currents were elicited upon repolarization to
80 mV
(Vr).
Slowly deactivating tail current amplitudes evoked on repolarization
from a holding potential of
100 mV by test depolarizations from
75 to
5 mV defined a voltage dependence of activation
that is well fit by a Boltzmann distribution (Fig. 5). Under these recording conditions, the
V1/2 of
activation of this current was
49.7 ± 0.48 mV
(n = 15). This half-activation potential is ~30 mV more hyperpolarized than values we reported previously for AG cells recorded at room temperature with a 20 mM
Ca2+ bath solution (34).
Ca2+-induced gating shifts have
been explained by changes in the surface potential near channel
proteins (29). Nonetheless, despite the more hyperpolarized value for
the V1/2 of
activation, channel open probability was not >0.25% at membrane
voltages negative to
80 mV. Figure 5 also depicts the voltage
dependence of inactivation constructed from slowly deactivating tail
current amplitudes evoked on repolarization by a test depolarization to
+15 mV from varying holding potentials. Under these recording
conditions, T-type Ca2+ channel
current inactivated with a
V1/2 of
67.8 ± 0.25 mV (n = 12), a
value that compares favorably to values previously reported by our
laboratory (
58 mV, 22°C) (3). Because the voltage
range over which steady-state window current is available (
75 to
40 mV) is still considerably more depolarized than the range of
membrane voltages measured in 2 and 5 mM
K+ solutions (
98 to
74 mV), with the foot of the activation curve defining the
extent of overlap, small changes in the
V1/2 of
activation could greatly alter the magnitude of
Ca2+ channel flux through T-type
Ca2+ channels during AG cell
stimulation.
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CaMKII activation and modulation of T-type
Ca2+
channel current.
Our laboratory has previously reported that low-voltage-activated,
T-type Ca2+ channels are regulated
by CaMKII (34) and/or the heterotrimeric G protein,
Gi (33). Each of these mechanisms
causes an approximate 10-mV hyperpolarizing shift in the
V1/2 of
activation without affecting the
V1/2 for
inactivation. Although at present, the manner in which these mechanisms
interact remains undefined, cell stimulation with
K+ (10 mM) or ANG II (10 nM)
increases CaMKII activity in the AG cell (19). To determine if
physiological K+ solutions
increase the activation state of CaMKII in intact AG cells during
stimulation with ANG II, we examined cell sonicates for
Ca2+-independent CaMKII activity.
Activation of CaMKII in situ results in autophosphorylation of
Thr286, rendering the kinase
partially active when assayed in the absence of activating
Ca2+. Autocamtide-2
(KKALRRQETVDAL), a synthetic peptide modeled on the sequence (RQETV)
containing the autonomy site
(Thr286) in the CaMKII
autoregulatory domain (22), was used as a specific substrate for CaMKII
(22) in the kinase assay. In 2 mM
K+, AG cell
Ca2+-independent CaMII activity
was 7.04 ± 2.8% of total
(Ca2+ dependent) CaMKII activity.
Cell stimulation with ANG II dose dependently (0.1 to 10 nM) increased
CaMKII activity from 125.6 ± 3.6 to 159.0 ± 15% of control
values (P
0.05, n = 9). This level of activation
compares well with the maximal activity of the
Ca2+-independent form of CaMKII
(227%) that can be generated by in vitro incubation with ATP and
Ca2+. Cell stimulation with 5 mM
K+ neither enhances CaMKII
activation (2 vs. 5 mM K+; NS) nor
augments the activation evoked by ANG II (Fig.
6). Because CaMKII activation is not
modulated by extracellular K+, our
data raise the possibility that mobilization of intracellular stores
may provide the Ca2+ responsible
for CaMKII activation during ANG II stimulation. Nonetheless, our data
show that K+ effects on the
activity of CaMKII cannot account for the potentiating effect of
K+ on ANG II-elicited aldosterone
secretion.
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DISCUSSION |
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We applied whole cell patch-clamp recording techniques to provide the first measurements of the membrane potential of bovine AG cells at 37°C in physiological K+ solutions. We measured whole cell currents under similar recording conditions and Ca2+ currents through T-type Ca2+ channels using physiological levels of Ca2+ as the charge carrier to assess the possible importance of these currents to the synergy exhibited by K+ and ANG II in controlling aldosterone secretion.
The resting membrane potential of the AG cell was
K+ dominated; it was close to
EK (14 mV more
depolarized) and shifted +21 mV with a change in extracellular
K+ from 2 to 5 mM (+24 mV expected
for a perfectly K+-selective
conductance). The value calculated based on our data for
Em at 4 mM
K+ (84 mV) is comparable to
values of
78 mV and
85 mV reported previously for acutely
dissociated bovine and rat AG cells, respectively, but was
significantly more hyperpolarized than potentials recorded from
cultured AG cells (
50 mV at 5.4 mM
K+).
Although numerous reports have identified members of all of the major classes of K+ channels in AG cells, the contribution of individual K+ channels to the resting membrane conductance appears to vary among species and among preparations (8, 27, 30, 38, 39, 53). In acutely dispersed rat glomerulosa cells, weakly voltage-dependent K+ leak channels appear to be the primary determinant of resting membrane (30, 44). Inhibition of current, through this leak channel, by Cs causes a large membrane depolarization, and presumably the opening of voltage-dependent Ca2+ channels that leads to the stimulation of aldosterone production (31). Nonetheless, ANG II, which inhibited this conductance more modestly, did not lower membrane potential (27) but stimulated aldosterone secretion (31). Unlike freshly dispersed AG cells, a charybdotoxin-sensitive, Ca2+-dependent maxi-K+ channel has been suggested to play a pivotal role in controlling membrane potential in cultured rat glomerulosa cells (38). The expression of this conductance is dominant in primary cultures of rat AG cells after 48 h of culture (32, 38). Inhibition of this conductance by ANG II effects a large 23-mV membrane depolarization, an increase in cytosolic Ca2+, and a putative increase in aldosterone secretion (38). Similar differences exist between freshly dissociated and cultured preparations of bovine AG cells.
Cultured preparations exhibit a prominent voltage-dependent transient
outward macroscopic current, with little or no evidence of inward
current at membrane voltages negative to 50 mV (8). In acutely
dispersed preparations, macroscopic inward current is measurable (44),
but the observed low density of inward rectifiers that can be inhibited
by ANG II (27, 53) cannot account for this inward current. We show, in
acutely dispersed bovine AG cells that maintain membrane potentials
close to EK in
physiological K+ solutions, a
macroscopic current that is weakly outwardly rectifying with an
I-V relationship that is strikingly
similar to that previously reported for acutely dissociated AG cells
(30, 44). Inhibition of the resting membrane conductance by ANG II
defines an ANG II-sensitive component of current that has a reversal
potential close to
EK and that
shifts as the Nernst equilibrium potential when extracellular K+ is changed. The component of
current that is active at the
Em of the AG cell
resembles current that is conducted by weakly voltage-dependent K+ leak channels (30). As in rat
AG cells, in our acutely dispersed preparation of bovine AG cells, we
found no evidence of an inwardly rectifying macroscopic current.
We demonstrate that inhibition of this resting
K+ conductance by ANG II does not
result in a large change in membrane potential. This is to be expected
because the depolarizing effect of a reduction in a
K+-selective conductance is
proportional to the driving force on K+. We show here that the
difference between the
Em and the
Erev of the ANG
II-sensitive component of current in acutely dissociated bovine AG
cells is 9 mV in physiological
K+ solutions. This small driving
force contrasts with that found in cultured preparations that maintain
a membrane potential that is well displaced from
EK and, thus, in
which a similar change in conductance by ANG II would cause a much
larger membrane depolarization. Because the membrane potential of the
AG cell is K+ dominated in the
presence of ANG II, extracellular
K+ will be the primary determinant
of membrane potential and
Ca2+-channel open probability.
Despite numerous reports characterizing the gating properties of T-type
Ca2+ channels in AG cells (12, 16,
35, 36, 49), the values for the
V1/2 of
activation and inactivation that have been previously defined are not
useful for predicting channel open probability under physiological
conditions because the open state probability for voltage-dependent
Ca2+ channels depends on the type
and concentration of charge carrier used (11, 13). In these studies, we
recorded at 37°C and used a physiological concentration of charge
carrier (1.25 mM Ca2+). The
Boltzmann function describing the voltage dependence of activation of
the slowly deactivating Ca2+
current had a
V1/2 of
activation of 44.7 mV, a value that is at least 15 mV more
hyperpolarized than most previously reported values (
30 mV to
15 mV). Combining these data with the measurements of membrane
potential, we can predict the probability that a channel will be open
in the steady state using the multiplicative product of
m
(probability
that a Ca2+ channel is activated) × h
(probability that a Ca2+ channel
is not inactivated). For example, we can predict that at
98.6 mV
(the Em measured
in 2 mM K+), the probability
that a channel will be open in the steady state is vanishingly small
(0.02%), the product of 0.0002 × 0.999. Even given the large
surface-to-volume ratio in AG cells and assuming that the
Ca2+ influx necessary to maintain
secretion can occur over several minutes, our data argue that the
stimulation of aldosterone secretion by ANG II in 2 mM
K+ does not depend on T-type
Ca2+-channel activity (i.e., the
membrane potential never approaches the voltage range over which the
channel is open). An alternative mechanism that might account for the
extracellular Ca2+ dependence of
ANG II-stimulated aldosterone secretion in 2 mM K+ is the capacitative entry
channel. ANG II stimulates the formation of inositol
1,4,5-trisphosphate in AG cells, and the depletion of
1,4,5-trisphosphate-sensitive Ca2+
stores activates a capacitative
Ca2+ influx (9, 47). Recent
studies suggest that secretion stimulated by high concentrations of ANG
II exhibits a stronger dependence on
Ca2+ influx mediated by the
capacitative Ca2+ entry channel
(9). The requirement for higher concentrations of ANG II to elicit
equivalent secretory responses in 2 mM
K+, and the strong electrical
driving force that maximizes Ca2+
flux through this channel, is consistent with an important role for
capacitative Ca2+ entry in
sustaining ANG II-stimulated aldosterone secretion in 2 mM
K+. Our results further show that
in 5 mM K+ (
79 mV), the
probability that a T-type Ca2+
channel will be open in the steady state will increase to 0.3% (0.003 × 0.999), concomitant with a twofold increase in aldosterone secretion. Simultaneous activation of CaMKII by ANG II shifts the
voltage range of activation of T-type
Ca2+ channels and induces a
further slight depolarization to
74 mV, such that the T-channel
open probability would be expected to increase to ~2.2% (0.0253 × 0.871) in 5 mM K+ and ANG
II. This expected increase in Ca2+
entry through T-type Ca2+ channels
effected by K+ depolarization
contrasts with a predicted decrease in
Ca2+ entry through capacitative
entry pathways. In our studies, 100 pM ANG II stimulated cellular
CaMKII activity only weakly, yet potentiated secretion
robustly. This discrepancy in dose effects is not
surprising because the diffusion of
Ca2+ within cells is very limited,
and suggests that the pool of CaMKII responsible for regulating T-type
Ca2+ channel activity is located
near the site of Ca2+ elevation.
In conclusion, our results suggest that increases in extracellular K+ move the membrane potential of the AG cell to a value where Ca2+ channels are active in the steady state. This, together with an ANG II-induced shift in the voltage range of activation and a slight change in membrane voltage, would promote even more Ca2+ channel activity in the steady state and thus could account for the synergy between K+ and ANG II in the control of Ca2+ influx. The importance of this regulated flux to aldosterone secretion is supported by previous reports that CaM antagonism (1, 2, 54) and CaMKII inhibition (40) attenuate ANG II-stimulated aldosterone secretion without effecting basal aldosterone production or production from 22(R)-hydroxycholesterol. However, because CaM antagonism in permeabilized bovine glomerulosa cells also prevents Ca2+ stimulation of aldosterone production, sites downstream of Ca2+ influx that are critical to stimulated secretion must also be targets of CaM activation (10). Nonetheless, the importance of voltage-dependent Ca2+ channel activity to secretion stimulated by 100 pM ANG II in 5 mM K is supported by the dramatic reduction of stimulated secretion effected by either a cationic (Ni2+, <100 µM) or organic "T-selective" Ca2+-channel inhibitor (mibefradil, 1.2 µM IC50) (4). Thus the potentiating effect of K+ on ANG II-stimulated aldosterone production likely mirrors the synergy with which ANG II and K+ regulate the activity of low-voltage-activated, T-type Ca2+ channels.
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ACKNOWLEDGEMENTS |
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We are grateful for the conceptual input of Dr. Eugene Barrett during development of this project.
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FOOTNOTES |
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This work was supported by National Heart, Lung, and Blood Institute Grant HL-36977 (to P. Q. Barrett) and National Institute of Neurological Disorders and Stroke Grant NS-33583 (to D. A. Bayliss).
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. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: P. Q. Barrett, Dept. of Pharmacology, Univ. of Virginia School of Medicine, 1300 Jefferson Park Ave., Charlottesville, VA 22908 (E-mail: pqb4b{at}virginia.edu).
Received 13 July 1998; accepted in final form 7 January 1999.
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