A role for T-type Ca2+ channels in the synergistic control of aldosterone production by ANG II and K+

Xiao-Liang Chen, Douglas A. Bayliss, Robert J. Fern, and Paula Q. Barrett

Department of Pharmacology, University of Virginia School of Medicine, Charlottesville, Virginia 22908


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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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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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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 MOmega 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 beta -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 [gamma -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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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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Fig. 1.   ANG II and K+ synergize to control aldosterone secretion. Bovine adrenal glomerulosa (AG) cells equilibrated in 3.5 mM K+ Krebs-Ringer bicarbonate (KRB) at 37°C were treated with and without ANG II (between 10 pM and 10 nM) at 2 mM (open circle ) or 5 mM () K+. Aldosterone content of cell medium was determined and secretory rates (pg/min per 106 cells) were calculated during maintained incubation (30-60 min). Values are means ± SE of 4 experiments performed in quadruplicate.

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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Fig. 2.   Elevation of extracelluar K+ from 2 to 5 mM reduces membrane potential and whole cell current. AG cells maintained at 37°C were perfused continuously with Krebs-HEPES Ringer, with extracellular K+ fixed at 2 or 5 mM. A: current-clamp recordings of membrane potential. Values are means ± SE of 4 cells. In 5 mM K+, AG cell membrane potential depolarized approximately +21 mV (from -98 mV to -77 mV). B: voltage-clamp recordings of whole cell current. Membrane current was evoked by voltage-ramp commands to -140 mV from a holding potential of -50 mV. Trace is averaged current from 5 cells. Mean zero-current potential is -98.5 ± 1.7 mV (n = 5) in 2 mM K+ and -77.0 ± 1.5 mV (n = 5) in 5 mM K+.

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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Fig. 3.   Effects of ANG II on whole cell current and membrane potential in 2 mM K. AG cells maintained at 37°C were perfused continuously with Krebs-HEPES Ringer, with extracellular K+ fixed at 2 mM in the absence or 2.5 min after exposure to 10 nM ANG II. A: membrane current was elicited by voltage-ramp commands to -140 mV from a holding potential of -50 mV. Trace is averaged current recorded from 5 cells. ANG II decreased membrane conductance determined ± 5 mV of zero-current potential (P <=  0.05). ANG II-sensitive component of current reversed at -107 ± 4.6 mV, close to equilibrium potential for K+ (EK; -114 mV). B: membrane potential determined in current clamp and from voltage-ramp protocols (zero-current potential). ANG II depolarizes AG cells from -98.5 ± 1.5 mV to -95.6 ± 1.8 mV (n = 11). * P <=  0.05 compared with 2 mM K alone.

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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Fig. 4.   Effects of ANG II on whole cell current and membrane potential in 5 mM K+. AG cells maintained at 37°C were perfused continuously with Krebs-HEPES Ringer, with extracellular K+ fixed at 5 mM, in the absence or 2.5 min after exposure to 10 nM ANG II. A: membrane current was elicited by voltage-ramp commands to -140 mV from a holding potential of -50 mV. Trace is averaged current recorded from 5 cells. ANG II decreased membrane conductance determined ± 5 mV of zero-current potential (P = 0.05). ANG II-sensitive component of current reversed at -87 ± 3.6 mV, close to EK (-90 mV). B: membrane potential determined in current clamp and from voltage-ramp protocols. ANG II depolarizes AG cells from -79.8 ± 1.4 mV to -74.0 ± 2.0 mV (n = 11). * P <=  0.05 compared with 5 mM K alone.

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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Fig. 5.   T-type Ca2+ channel window currents at 37°C, recorded with physiological Ca2+ (1.25 mM) as charge carrier. Slowly deactivating tail currents were elicited at -80 mV after maximal depolarizations (+15 mV, 8 ms) from various holding potentials lasting 6 s (Vp = -100 to -35 mV) or after various depolarizing test pulses (Vt = -75 to -5 mV; 10 ms) from a holding potential of -95 mV. A: sample records from a single cell showing current elicited at representative potentials for each commmand protocol. Inset: tail currents shown on expanded time scale; tau  of deactivation was 0.95 ms at Vr = -80 mV. B: relative amplitude of slowly deactivating component of tail current is plotted vs. Vt or Vp. Data are fit by Boltzmann distributions: I/Imax = {1 + exp[(V1/2 - Vt)/k])-1} for activation, and I/Imax = {1 + exp[(Vp - V1/2)/k])-1} for inactivation, yielding the following parameters. Inactivation: V1/2 = -67.8 ± 0.25 mV, k = 4.5, r = 0.99 (n = 12). Activation: V1/2 = -49.7 ± 0.48 mV, k = 5.0, r = 0.99 (n = 15). Intersection of the 2 relationships defines voltage range for steady-state T-type Ca2+ channel currents.

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 CaMKIIalpha 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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Fig. 6.   ANG II stimulates CaMKII activity equivalently in 2 and 5 mM K+. Ca2+-independent CaMKII phosphotransferase activity (a reporter activity for activation of kinase in situ) was measured in samples prepared from AG cells treated for 30 s with 2 or 5 mM K+, in absence and presence of ANG II (from 100 pM to 10 nM). Ca2+-independent activity is expressed as percentage of total (Ca2+-dependent) CaMKII activity and normalized to the basal value obtained in 2 mM K+. Values are means ± SE of 4-10 experiments performed in triplicate. * P <=  0.05 vs. 2 mM K+ control.

However, if CaMKII activity regulates the physiological V1/2 of activation of the T-type Ca2+ channel, then during cell stimulation with ANG II, cell depolarization by 5 mM K+ could elicit a greater increase in channel open probability. Figure 7A shows that the voltage range of activation of the T-type Ca2+ channel can be controlled by the activation state of CaMKII. In the presence of 5 mM ATP and 0.2 µM pipette CaM, increasing free Ca2+ from 23 nM to 1.2 µM evokes commensurate hyperpolarizing shifts in the V1/2 of activation, from -47.9 ± 1.8 mV (n = 5) at 27 nM Ca2+ to -49.3 ± 1.5 mV (n = 3) at 250 nM, -53.1 ± 1.4 mV (n = 5) at 800 nM, and -58.4 ± 1.2 mV (n = 5) at 1.2 µM Ca2+ (27 nM vs. 1.2 µM, P = 0.05). A comparison of the two Boltzmann functions that fitted the data at 27 nM and 1.2 µM Ca2+ over the range of membrane potentials established by physiological K+ solutions (Fig. 7B) indicates that, during ANG II stimulation (maximal CaMKII activation), 5 mM K+ depolarization should increase channel open probability more than 100-fold (from 0.02 to 2.5%). This 100-fold increase compares with an approximate 30-fold increase (from 0.01 to 0.3%) in channel open probability effected by 5 mM K+ in the absence of ANG II. Importantly, in the presence of ANG II, CaMKII-activated channels are eight times more likely to open in 5 mM K+ (Vm = -74 mV) than nonmodulated channels (Vm = -79 mV). Thus increased Ca2+ entry through T-type Ca2+ channels could account for the ability of K+ to enhance the physiological secretory potential of ANG II. The failure of 5 mM K+ to stimulate aldosterone secretion elicited by supraphysiological concentrations of ANG II likely reflects additional inhibitory effects of high concentrations of ANG II on aldosterone secretion previously reported (24).


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Fig. 7.   Modulation of T-type Ca2+-channel current by intracellular Ca2+ concentration. A: with 0.2 µM CaM, 5 mM ATP, and 1 mM GTP in pipette solution, free Ca2+ was fixed at 27 nM, 250 nM, 800 nM, or 1.2 µM using 11 mM 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA). Ca2+ shifted the V1/2 of activation to more negative potentials: open circle , V1/2 = -47.9 ± 1.8 mV (n = 5, 27 nM Ca2+); black-down-triangle , -49.3 ± 1.5 mV (n = 3, 250 nM Ca2+); black-diamond , -53.1 ± 1.4 mV (n = 5, 800 nM Ca2+); black-triangle, -58.4 ± 1.2 mV (n = 5, 1.2 µM Ca2+). P <=  0.05: 27 nM vs. 800 nM or 1.2 µM Ca2+; 250 vs. 1.2 µM Ca2+. B: data from A plotted on expanded scale marking expected fractional current for nonmodulated and CaMKII-activated channels at membrane potentials established by physiological K+ solutions.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

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 minfinity (probability that a Ca2+ channel is activated) × hinfinity (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.


    ACKNOWLEDGEMENTS

We are grateful for the conceptual input of Dr. Eugene Barrett during development of this project.


    FOOTNOTES

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.


    REFERENCES
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ABSTRACT
INTRODUCTION
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RESULTS
DISCUSSION
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