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1 Department of Anesthesia and Critical Care, The University of Chicago, Chicago, Illinois 60637; 3 Departments of Neurobiology, Pharmacology and Physiology, The University of Chicago, Chicago, Illinois 60637; 2 Department of Anesthesiology and Pharmacology, Vanderbilt University, Nashville, Tennessee 37232-2520
Submitted 27 June 2003; accepted in final form 25 August 2003
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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). During development, GABAA mediated depolarization has been observed in many regions of the CNS (Ben-Ari 2002). In these cells, the depolarized ECl observed is due to, at least in some cases, a member of the Na+-K+-2Cl– (NKCC) co-transporter family (Alvarez-Leefmans et al. 1988; Rohrbough and Spitzer 1996; Sun and Murali 1999), which driven by Na+ and K+ gradients, acts to accumulate intracellular Cl– (Ben-Ari 2002). Expression of a different co-transporter, KCC2 (K+/Cl–) (Payne et al. 1996), which normally lowers intracellular Cl–, appears to mediate the developmental shift toward inhibition (Ben-Ari 2002; Owens and Kriegstein 2002; Rivera et al. 1999). Embryonic animals appear to have limited expression of KCC2 and consequently exhibit elevated [Cl–]i. As KCC2 expression increases with developmental age, excitatory GABA responses become inhibitory. Interestingly, depolarizing GABAA actions can even be observed in some adult neurons (Alvarez-Leefmans et al. 1988; Staley et al. 1996).
GABA immunoreactive cells are found in the adrenal gland (Oomori et al. 1993). Chromaffin cells possess the enzymes that synthesize and degrade GABA (Fernandez-Ramil et al. 1983; Kataoka et al. 1984). GABA is taken up by chromaffin cells and is released in response to depolarizing stimuli (Kataoka et al. 1984; Oset Gasque et al. 1985). GABA immunoreactive nerve fibers are found throughout the adrenal medulla (Kataoka et al. 1986; Oomori et al. 1993). Chromaffin cells are known to express functional GABAA receptors with properties similar to their neuronal counterparts (Amenta et al. 1988; Bormann and Clapham 1985). Sangiah et al. (1974) first showed that GABA could elicit catecholamine release from isolated perfused adrenal glands, a result that has been repeated (Fujimoto et al. 1987; Kataoka et al. 1984; Kitayama et al. 1984). Even though a variety of studies have demonstrated that GABA depolarizes chromaffin cells, there is still a question of whether the net effect of GABA is excitatory or inhibitory; some studies suggest that GABA acts to inhibit excitation produced by acetylcholine (Ach) or high K+ (Fujimoto et al. 1987; Kataoka et al. 1986), while other studies suggest that GABA facilitates stimulation evoked release (Kitayama et al. 1990c, 1991).
Here we describe the actions of GABA in juvenile chromaffin cells and explore the molecular mechanisms underlying these actions. Exposing these cells to GABA caused a large and rapid elevation in [Ca2+]i, which required Ca2+ influx from the extracellular space. The elevation of [Ca2+]i required the activation of GABAA receptors; the action was mimicked by the selective GABAA agonist muscimol and blocked by the selective GABAA antagonist bicuculline. Exposing chromaffin cells to muscimol elicited significant catecholamine secretion. The extrapolated anion equilibrium potential of approximately –28 mV requires an equivalent intracellular Cl– concentration of approximately 50 mM (although both chloride and bicarbonate contribute to the anion reversal potential, HCO3 effects were minimized by the use of HEPES buffered solutions and because solutions were not bubbled with CO2-containing gases). These results suggest that GABAA receptors are capable of strongly depolarizing chromaffin cells leading to opening of voltage-gated calcium channels. We show that chromaffin cells expressed NKCC1 protein and not KCC2 protein, a result which provides the molecular underpinning for the elevated [Cl–]i levels observed. Consistent with these observations, blockade of NKCC1 with bumetanide significantly shifted ECl, as measured by changes in [Ca2+]i. As discussed above, there is still considerable debate about whether activation of GABAA receptors in chromaffin cells inhibits the excitation produced by the physiological activator of chromaffin cells, ACh (Fujimoto et al. 1987; Kataoka et al. 1986; Kitayama et al. 1990c, 1991). Our results suggest that activation of GABAA receptors in chromaffin cells is excitatory under all conditions.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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Juvenile bovine adrenal glands, from 4- to 5-mo-old animals, were obtained from a local slaughterhouse, and chromaffin cells were prepared by digestion with collagenase followed by density gradient centrifugation as previously described. The cells were plated onto collagen coated coverslips (at a density of 0.15–0.2 x 106 cells/ml for electrophysiology experiments and 0.3–0.4 x 106 cells/ml for [Ca2+]i measurements) and maintained in an incubator at 37°C in an atmosphere of 93% O2-7% CO2 with a relative humidity of 90%. Fibroblasts were effectively suppressed with cytosine-arabinoside (10 µM), leaving relatively pure chromaffin cell cultures. One-half of the culture medium was exchanged every day. This medium consisted of DMEM / F12 (1:1) (Gibco) supplemented with fetal bovine serum (10%), glutamine (2 mM), penicillin/streptomycin (100 unit/ml per 100 µg/ml), cytosine arabinoside (10 µM), and 5-fluorodeoxyuridine (10 µM).
[Ca2+]i measurements
Experiments were conducted 1–5 days after chromaffin cells were prepared. Cells were incubated in HBSS with 2 µM fura-2 AM and 1 mg/ml albumin for 45 min and washed in a fura-2-free solution for 1 h. Fura-2 loading and wash was carried out at 37°C. The coverslip was transferred to an experimental chamber for recording. Backgrounds at 340 and 380 nm were obtained using an area of the coverslip devoid of cells. Data were continuously collected throughout the experiment. On each coverslip, 10–40 chromaffin cells were selected and individually imaged. Image pairs (1 at 340 and 380 nm) were obtained every 2 s by averaging 16 frames at each wavelength. Backgrounds were subtracted from the individual wavelengths, and the 340-nm image was divided by the 380-nm image to provide a ratiometric image. Ratios were converted to free [Ca2+]i by comparing data to fura-2 calibration curves made in vitro by adding fura-2 (50 µM free acid) to solutions that contained known concentrations of calcium (0–2,000 nM). The recording chamber was continually perfused with fresh solution from gravity fed reservoirs.
Electrophysiology
Cells were voltage clamped using an Axopatch 1D amplifier and custom written Axobasic software in the gramicidin perforated whole cell configuration. Electrodes were pulled from microhematocrit capillary tubes (Drummond Scientific, Broomall, PA) and coated with Sylgard (Dow Corning, Midland, MI). After fire polishing, electrodes had resistances of <2 M
. Electrodes tips were briefly dipped in gramicidin-free patch pipette solution to fill the tip; gramicidin (50 µg/ml) containing solution was used to backfill the electrode. After seal formation, gramicidin was slowly inserted into the membrane, and the series resistance was monitored by applying 20-mV steps (from a holding potential of –91 mV) to the cell every 15–20 s. The mean series resistance was 18.4 ± 4.8 M (n = 5). Most of the series resistance was compensated using the compensation circuitry of the Axopatch-1D amplifier. No further correction was applied to the data as the currents were typically small. During experiments, cells were voltage clamped at a holding potential of –91 mV, and the recording bath (total volume 
250–300 µl) was continually washed with fresh extracellular medium at a rate of 3–4 ml/min from gravity-fed reservoirs. Inward GABAA receptor currents were elicited by continual perfusion of a supramaximal dose of muscimol (100 µM). Ramp depolarizations (l00 ms duration from –101 to –51 mV) were applied under control conditions and at the peak of the current elicited by muscimol (see Fig. 6). Currents generated by the ramps were filtered at 2 kHz and sampled every 100 µs. A second computer running Axotape software was used to continually acquire data for 1-min episodes (sampling every 100 or 200 µs) to capture the inward current response to muscimol application.
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Amperometric measurement of catecholamine release
Cells were plated approximately five times more densely for these experiments than for the electrophysiology experiments. The recording bath was designed so that it was small (<1 cm diam), and the volume of the bath was kept at around 300–350 µl. Carbon fiber electrodes (Dagan Instruments) were backfilled with 3 M KCl and held at a potential of +700 mV using a modified Warner PC-501 patch clamp amplifier. Electrodes were positioned within the bathing medium. In this configuration, the electrode detected increases in the local catecholamine concentration due to release from neighboring cells. Amperometric data from the patch clamp was recorded continuously using AxoTape (an 8-pole Bessel filter was set at 300–500 Hz and acquired at 1–5 kHz). The cells were continually washed with HBSS for several minutes prior to application of muscimol or high-K+ containing solutions. Under control conditions (HBSS), stopping the flow of the solution in recording bath for 90–120 s did not produce any change in amperometric current, indicating that the cells were not releasing appreciable amounts of catecholamine under basal conditions. Cells were stimulated by stopping the flow of the bath solution for 15–30 s and directly applying a 50-µl bolus of one of the following: HBSS (negative control), high K+ (50 mM; positive control), or muscimol (100 µM; final bath concentration of approximately 20 µM). After 30–60 s, the flow of HBSS through the bath was resumed to wash away the drug and any released catecholamine. A known concentration of noradrenaline was applied to the electrode in the absence of any cells to calibrate the response.
Solutions
Experiments were carried out in HBSS (Gibco) that contained (in mM) 138 NaCl, 5 KCl, 1.3 CaCl2, 0.3 KH2PO4, 0.8 MgSO4, 0.3 Na2HPO4, 5.6 D-glucose, and 20 HEPES. Nominally Ca2+-free HBSS was prepared by treating Ca2+-free, Mg2+-free, HCO3-free HBSS with Chelex-100 and adding MgCl2 to a final concentration of 1 mM. GABA (Sigma) was prepared as 100 mM stock in H2O. The GABA was diluted to a final concentration of 100 µM with HBSS. Muscimol hydrobromide (Calbiochem) was prepared as a 25-mM stock in H2O. Muscimol was diluted to a final concentration with HBSS. (–)-Bicuculline methiodide (Sigma) was prepared as a 100-mM stock in H2O and diluted to a final concentration of 100 µM with HBSS. Lanthanum chloride (Sigma) was prepared as a 100 mM stock in H2O and diluted to a final concentration of 1 mM with HBSS. Cadmium chloride (Sigma) was prepared as a 100-mM stock in H2O and diluted to a final concentration of 200 µM with HBSS. The solution containing sodium thiocyanate (Sigma) contained (in mM) 140 NaSCN, 1.3 CaCl2, 5 KCl, 1 MgCl2, 5.6 glucose, and 20 HEPES (pH 7.35). The high-K+ solution contained (in mM) 97 NaCl, 50 KCl, 1 MgCl2, 10 glucose, 10 HEPES, and 2 CaCl2 (pH 7.3). Bumetanide was prepared as a 1-M stock in DMSO and diluted to 10 µM with HBSS for pretreatment of the cells. For electrophysiological recordings, the cells were bathed in an extracellular solution that consisted of (in mM) 145 NaCl, 2 KCl, 1 MgCl2, 10 glucose, 10 HEPES, and 2 CaCl2 (pH 7.3; 305–310 mOsm). The patch pipette solution consisted of (in mM) 145 Cs-glutamate, 9.5 NaCl, 0.5 TEA Cl, and 10 HEPES (pH 7.3; 300–305 mOsm). These solutions produced a junction potential of approximately –11 mV, which has been corrected throughout the paper. Gramicidin was prepared at a stock concentration of 10 mg/ml in DMSO daily, and aliquots were diluted in the patch pipette solution and sonicated immediately prior to use (final concentration = 50 µg/ml). For the amperometry experiments, the bath contained HBSS. All experiments were carried out at room temperature (
23°C).
Expression of KCC2 and NKCC1
Expression of KCC2 and NKCC1 in bovine chromaffin cells was determined by immunoblotting of chromaffin cell membranes prepared as described by Williams et al. (1999). Bovine brain membranes were prepared in the same way to ensure that the antibodies recognized the bovine forms of both KCC2 and NKCC1. Membranes proteins (10 µg) were separated by SDS-PAGE and transferred to Immobilon P by standard procedures. For detection of NKCC1, a monoclonal antibody against NKCC1 (T4, Developmental Studies Hybridoma Bank, The University of Iowa) was used at a dilution of 1:20,000, followed by an anti-mouse IgG-horseradish peroxidase-coupled secondary antibody (Jackson ImmunoResearch Laboratories) used at a dilution of 1:150,000. For detection of KCC2, a polyclonal rabbit antibody (Upstate Biotechnology 07-285) was used at a dilution of 1:3,000, followed by an anti-rabbit IgG-horseradish peroxidase–coupled secondary antibody (Jackson ImmunoResearch Laboratories) used at a dilution of 1:100,000. Bound antibody was detected using the ECL Advance System (Amersham).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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Chromaffin cells were studied after 1–5 days in primary culture. A coverslip containing fura-2–loaded cells was transferred to an experimental chamber for each experiment. A field of cells was imaged, and [Ca2+]i measurements from 10–40 cells per coverslip were collected every 2 s. Figure 1A plots the mean [Ca2+]i and SE averaged from 16 cells. Cells were perfused with a Ca2+-containing HBSS solution (see METHODS) and exposed three times to 100 µM GABA as indicated in Fig. 1. The first exposure to GABA elevated [Ca2+]i by about 400 nM (Fig. 1A). After removal of extracellular Ca2+, application of GABA was without effect. When Ca2+-containing HBSS was reintroduced into the bath, subsequent exposure to GABA produced a somewhat smaller, but significant, elevation in [Ca2+]i (Fig. 1A). The mean change in [Ca2+]i elicited by 100 µM GABA in Ca2+-containing HBSS was 441 ± 19 nM (n = 48), while no change in [Ca2+]i was observed when GABA was applied in Ca2+-free solution. These data suggest that GABA elicits an elevation of [Ca2+]i that requires the influx of extracellular Ca2+.
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GABAA receptors mediate the elevation of [Ca2+]i observed
The pharmacology of the GABA effect was explored in greater detail using agonists and antagonists specific for the GABAA receptor. Superfusion of the chromaffin cells with the GABAA agonist muscimol (5 µM) produced an elevation in [Ca2+]i of >400 nM when applied in Ca2+-containing HBSS (Fig. 1B). When the bath solution was subsequently changed to Ca2+-free HBSS, re-application of muscimol was without effect on [Ca2+]i. Re-introduction of the Ca2+-containing solution followed by the application of muscimol produced another large elevation of [Ca2+]i (Fig. 1B). The average change in [Ca2+]i elicited by application of muscimol (5 µM) in Ca2+-containing solution was 361 ± 70 nM (n = 44), while in the Ca2+-free solution, no change in [Ca2+]i was observed. This result suggests that muscimol and GABA elevate [Ca2+]i by the activation of ionotropic GABAA receptors known to exist in bovine chromaffin cells (Bormann and Clapham 1985; Peters et al. 1989).
To further verify the involvement of GABAA receptors, the specific GABAA receptor antagonist bicuculline was applied to chromaffin cells to block the response to muscimol. In the presence of bicuculline, the response to muscimol was completely blocked (Fig. 2A). Partial recovery of the response to muscimol was observed on washout of the bicuculline (data not shown). Muscimol (5 µM) produced an average elevation in [Ca2+]i of 335 ± 26 nM (n = 44) when applied in the absence of bicuculline, but this increase was almost completely blocked, to 11.9 ± 2.4 nM, by bicuculline (Fig. 2B). These data clearly demonstrate that activation of GABAA receptors in chromaffin cells are responsible for large elevations in [Ca2+]i.
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Voltage-dependent Ca2+ channels appear to mediate the Ca2+ influx into chromaffin cells
In chromaffin cells, the primary route for calcium influx into the cells is through voltage-gated calcium channels, although other pathways such as nicotinic ACh receptors or members of the transient receptor potential (TRP) family of ion channels may also allow sufficient Ca2+ into cells to trigger catecholamine release (Obukhov and Nowycky 2002). Depolarization of chromaffin cells by high-K+–based solutions activates voltage-dependent Ca2+ channels, elevates [Ca2+]i, and elicits robust catecholamine secretion. Nonselective Ca2+ channel blockers like La3+ block the Ca2+ influx produced by high-K+ stimulation by blocking voltage-dependent Ca2+ channels. Therefore we have used La3+ to investigate the role of voltage-dependent Ca2+ channels in muscimol-induced increases in [Ca2+]i. As shown in Fig. 3A, Ca2+-influx elicited by high-K+ stimulation of the chromaffin cells was blocked by La3+. La3+ also blocked the elevation of [Ca2+]i in response to muscimol. After washing La3+ out of the bath, increased [Ca2+]i was observed in response to both muscimol and high K+ (Fig. 3A). Mean data from 46 cells show that La3+ completely blocked the [Ca2+]i elevation observed with muscimol stimulation (Fig. 3B). In this set of experiments, muscimol elevated [Ca2+]i by 184 ± 26 nM in the absence of La3+ but by 1.6 ± 0.07 nM in the presence of La3+. (Note that the response to muscimol was decreased in these experiment because muscimol was tested after exposure to La3+ and washout of the La3+ was incomplete.) These results suggest that both high K+ and muscimol elevate [Ca2+]i by pathways that involve activation of voltage-dependent Ca2+ channels. We repeated the experiment with another Ca2+ channel blocker, Cd2+ (200 µM, data not shown). Mean data from 52 cells show that Cd2+ effectively blocked the [Ca2+]i elevation observed with muscimol stimulation. In this set of experiments, muscimol elevated [Ca2+]i by 439.6 ± 33.5 nM in the absence of Cd2+ but by 47.6 ± 4.8 nM in the presence of Cd2+. This supports the hypothesis that muscimol elevates [Ca2+]i by pathways that involve activation of voltage-dependent Ca2+ channels.
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Evidence of a depolarized anion equilibrium potential in chromaffin cells
For GABA to depolarize a cell sufficiently to activate voltage-dependent Ca2+ channels, it appears that intracellular chloride levels must be high. To test this hypothesis, we altered the Cl– reversal potential by replacing most of the extracellular Cl– with SCN–, an anion that is more permeant in GABAA receptors and other chloride channels than Cl– itself (Bormann et al. 1987; Currie et al. 1995). Replacing Cl– with SCN– anions typically results in a negative shift in the anion equilibrium potential. If the shift in the anion reversal potential is large enough, it would preclude activation of voltage-dependent Ca2+ channels in response to muscimol. Indeed, when cells were exposed to 5 µM muscimol in the presence of SCN– (140 mM), only a small elevation of [Ca2+]i was observed (Fig. 4A). The mean response of chromaffin cells to muscimol in a solution containing SCN– was an elevation in [Ca2+]i of 38.2 ± 6.8 nM, but an elevation in [Ca2+]i of 377.9 ± 25.8 nM (n = 51) was observed when applied in HBSS (Fig. 4B). These data support the idea that the chloride equilibrium potential of the cells is such that application of muscimol produces a depolarization that is strong enough to activate voltage-dependent Ca2+ channels.
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To further explore this hypothesis, the response to muscimol in chromaffin cells was studied using perforated whole cell patch-clamp conditions. Anion impermeant gramicidin channels were used as the perforating agent to keep intracellular Cl– intact. Figure 5A shows that muscimol elicited an inward current that decayed with time in cells voltage clamped at –101 mV. As the current peaked, a voltage ramp from –101 to –51 mV (after correction for junction potential; inset) was applied to the cell. Ramp currents obtained in the presence and absence of muscimol are plotted in Fig. 5B. Note that without muscimol present, virtually no current was observed in the voltage range tested. In this range, neither Na+ nor Ca2+ channels are active and K+ channels were blocked with Cs+ in the pipette. The difference between the "control" current and the current observed with "muscimol" in the bath represents the GABAA receptor current.
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Catecholamine secretion in response to muscimol
The robust elevations of [Ca2+]i observed on exposing cells to either GABA or muscimol suggest that activation of GABAA receptors may be able to elicit catecholamine secretion in chromaffin cells. To test this hypothesis, amperometry was used to measure catecholamine release. An amperometric electrode was positioned inside a small bath, adjacent to a group of chromaffin cells, and was used to oxidize and thereby detect catecholamines released into the bath. Catecholamine release was measured after each of three different solutions were introduced into the bath: 50 mM K+ (high K+; top), control HBSS solution (middle), or muscimol in HBSS (Fig. 6A). Experiments were performed in a static bath because rapid perfusion of fresh solution through the recording chamber rapidly dilutes and washes away secreted catecholamines. With the flow stopped, catecholamines released by the cells accumulate rapidly in the bath. Figure 6 shows that under control conditions (HBSS), there was no increase in bath catecholamine concentration. With the flow of solution through the bath stopped, both KCl and muscimol caused release of catecholamines that accumulated in the bath and led to an increase in the amperometric current recorded by the carbon fiber electrode. Resuming the flow of solution rapidly washed the released catecholamines out of the bath causing the amperometric current to return to baseline. Mean data from five experiments are shown in Fig. 6B and illustrate that muscimol, although not as effective as KCl, produces significant release of catecholamines from chromaffin cells. Thus in addition to elevating [Ca2+]i, activation of GABAA receptors also promotes the release of catecholamines in chromaffin cells.
Expression of KCC2 and NKCC1 co-transporters
In many embryonic neurons, GABA is an excitatory neurotransmitter, due to the high levels of intracellular Cl– found in these cells. After birth, the Cl– reversal potential changes to more negative potentials, in many cases due to expression of the K+-Cl– co-transporter KCC2. Although the bovine chromaffin cells used in this study were from juvenile and not embryonic animals, the elevated [Cl–]i suggests that these cells do not express the KCC2 transporter. To determine whether KCC2 is expressed in chromaffin cells, membrane proteins from the chromaffin cells and from bovine brain were separated by SDS-PAGE, transferred to polyvinylidene fluoride membranes, and probed with a polyclonal antibody to KCC2. A protein of the appropriate size for KCC2 (approximately 140 kDa) was observed in brain membranes but not in membranes from chromaffin cells (Fig. 7A). This indicates that KCC2 is not expressed in bovine chromaffin cells.
In many cases, expression of the NKCC1 co-transporter plays a role in the high Cl– levels observed. To determine whether NKCC1 is expressed in chromaffin cells, immunoblotting was carried out as described for KCC2. Proteins of the appropriate size for NKCC1 (approximately 180 kDa) were detected in both chromaffin cell membranes and bovine brain membranes. (Fig. 7A). These results indicate that NKCC1 is expressed in bovine chromaffin cells.
If NKCC1 is involved in maintaining the high [Cl–]i necessary for the depolarizing effects of muscimol, inhibition of NKCC1 should reduce the ability of muscimol to raise [Ca2+]i. Therefore we tested the ability of bumetanide, an inhibitor of NKCC1 to inhibit the effects of muscimol. One group of cells was pre-exposed to bumetanide (10 µM) for 3 h, while a second group was not. Muscimol was then applied to both groups of cells. [Ca2+]i was elevated by 299.3 ± 20.1 nM (n = 67) in control cells treated with muscimol but was elevated by only 169.5 ± 19.7 (n = 75; P < 0.01) in cells pre-exposed to bumetanide (Fig. 7B). Note that the baseline [Ca2+]i, prior to muscimol exposure was not different in either case (data not shown). Three-hour exposures to DMSO, the carrier for bumetanide, did not affect responses to muscimol {change in [Ca2+]i to muscimol plus DMSO was 239.31 ± 35.60 (n = 9), while control change in response to muscimol alone was 252.99 ± 37.46 (n = 12)}. In addition, high-K+ depolarizations produced virtually identical Ca2+ transients in both groups of cells, indicating that the voltage-dependent Ca2+ channels and intracellular Ca2+ buffering machinery were unaffected by bumetanide.
Co-activation of ACh and GABAA receptors
ACh released by presynaptic splanchnic nerve terminals is thought to be the physiological activator of adrenal chromaffin cells. In some cells, activation of GABAA receptors is depolarizing but is still inhibitory (Zhang and Jackson 1993). In chromaffin cells, there is considerable controversy whether GABA can inhibit responses to high K+ or ACh, even though GABA itself is excitatory (Fujimoto et al. 1987; Kataoka et al. 1986; Kitayama et al. 1990c, 1991). To address this issue, we examined elevations in [Ca2+]i in response to ACh, in the presence and absence of muscimol, to determine whether activation of GABAA receptors would inhibit the response to ACh. As shown in Fig. 8, responses to ACh (100 µM; a nonsaturating concentration) were virtually identical in the presence or absence of muscimol. The order of presentation of ACh or ACh plus muscimol did not affect the responses (see Fig. 8, A and B). In addition, applying muscimol 2 min prior to ACh application did not alter the response (data not shown). In this set of experiments, ACh elevated [Ca2+]i by 771 ± 11 nM in the absence of muscimol and by 748 ± 8 nM in the presence of muscimol (n = 59).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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; Kataoka et al. 1984); GABA is taken up by chromaffin cells and released in response to depolarizing stimuli (Kataoka et al. 1984; Oset Gasque et al. 1985, 1990); GABA immunoreactive nerve fibers are found throughout the adrenal medulla (Kataoka et al. 1986; Oomori et al. 1993) and bundles of GABA immunoreactive nerve fibers run into the medulla with varicosities often in close contact with chromaffin cells (Kataoka et al. 1986; Oomori et al. 1993); chromaffin cells express GABAA receptors (Amenta et al. 1988; Bormann and Clapham 1985; Peters et al. 1989); and immunoblots indicate the presence of 
1, 4, 
1–3, and 
2 subunits (Parramon et al. 1995a). Functionally it has been shown that GABA can elicit catecholamine release from isolated perfused adrenal glands (Fujimoto et al. 1987; Gonzalez et al. 1992; Kataoka et al. 1984; Kitayama et al. 1984; Sangiah et al. 1974). Consistent with earlier studies (Doroshenko 1989; Kitayama et al. 1990a; Gonzalez et al. 1992), our results show that exposing chromaffin cells to GABA caused a large and rapid elevation in [Ca2+]i. Removing extracellular Ca2+ or blockade of Ca2+ influx with La3+ or Cd2+ abolished the action, suggesting that influx through voltage-gated calcium channels was required for the elevation of [Ca2+]i. While it is possible that release of Ca2+ from intracellular stores plays a role in the responses observed here, perhaps via Ca2+-dependent Ca2+ release, such release does not appear to initiate the measured response. The effects of GABA were mimicked by muscimol and blocked by bicuculline and therefore appears to be mediated by activation of GABAA receptors.
Ionotropic GABAA receptors are usually associated with inhibition, but during development, GABAA mediated depolarization has been observed in many regions of the CNS (Ben-Ari 2002; Chen et al. 1996; Eilers et al. 2001; Luhmann and Prince 1991; Mueller et al. 1984; Owens et al. 1996; Yuste and Katz 1991). The action of GABAA receptors in neurons is determined by the transmembrane Cl– gradient and the resultant ECl (Staley et al. 1996). The depolarized ECl observed is due to, at least in some cases, a member of the NKCC cotransporter family (Alvarez-Leefmans et al. 1988; Rohrbough and Spitzer 1996; Sun and Murali 1999), which driven by Na+ and K+ gradients, acts to accumulate intracellular Cl– (Ben-Ari 2002). Expression of another co-transporter that normally lowers intracellular Cl–, KCC2 (Payne et al. 1996), appears to mediate the developmental shift toward inhibition (Ben-Ari 2002; Owens and Kriegstein 2002; Rivera et al. 1999). Embryonic animals have limited expression of KCC2 and consequently exhibit elevated [Cl–]i. As KCC2 expression increases with developmental age, excitatory GABA responses become inhibitory, although the developmental stage at which KCC2 expression increases varies somewhat in different neurons (Mikawa et al. 2002). The juvenile animals used in our study are 4- to 6-mo-old and are quite mature. They should be similar to adult animals in terms of the switch in neuronal KCC2. Interestingly, depolarizing GABAA responses have also been observed in some adult neurons. Dorsal root ganglion neurons exhibit depolarizing GABA responses as ECl is 50 mV more depolarized than the cellular resting potential (Alvarez-Leefmans et al. 1988; Staley et al. 1996; Sung et al. 2000). In the retina, KCC2 is expressed in neurons where GABA is inhibitory, whereas NKCC is expressed in neurons where GABA is excitatory (Vardi et al. 2000).
The results from neurons suggest that the excitatory responses of chromaffin cells to GABA could be due to depolarization because of elevated [Cl–]i. Consistent with this hypothesis, we showed that replacing most of the extracellular Cl– with SCN– (which should shift ECl in the hyperpolarizing direction) reduced the [Ca2+]i response to muscimol. Using the gramicidin perforated patch-clamp technique (gramicidin channels are impermeable to Cl and so maintain the endogenous [Cl–]i), we extrapolated an anion equilibrium potential of approximately –28 mV, which requires an equivalent intracellular anion concentration of approximately 50 mM. We showed that the chromaffin cells express NKCC1 protein but not KCC2 protein, a result that provides the molecular underpinning for the elevated [Cl–]i levels. Furthermore, pharmacological blockade of the NKCC1 co-transporter with bumetanide significantly inhibited the response to muscimol, most likely by altering ECl. Although it has been reported that a different NKCC1 inhibitor furosemide can inhibit GABAA receptors at concentrations >1 mM (Nicoll 1978), we expect that bumetanide would be quite selective at the low concentration used in the current study. The inhibition by bumetanide, while significant, was incomplete probably because the dissipation of the intracellular anionic gradient was slow. We have no reason to doubt that bumetanide completely blocked NKCC1. Longer exposures to bumetanide gave progressively larger effects but very long exposures were technically difficult to do. So we compromised by using a 3-h exposure where we observed an incomplete response.
The role of GABA in the adrenal gland is still not well understood. There is even some debate as to whether GABA is truly excitatory in adrenal chromaffin cells. Several studies have suggested that GABA acts to inhibit excitation produced by other neurotransmitters (Fujimoto et al. 1987; Kataoka et al. 1986) and that activation of GABAA receptors at first lead to action potential generation, followed by the arrest of action potentials (Busik et al. 1996). However, Kitayama et al. (1990c, 1991) showed that GABA facilitated catecholamine release elicited by ACh or high K+. Earlier measurements in chromaffin cells placed ECl at –51.4 mV (Gonzalez et al. 1992), suggesting that GABA will depolarize chromaffin cells only weakly if at all. Our data suggest that GABA is excitatory under all conditions. Responses to ACh were not diminished by co-activation of GABAA receptors. Furthermore, we provide evidence that ECl is much more depolarized than previously appreciated. Please note that any rectification of the channels would alter our estimated reversal potential but that it would be quite depolarized under all conditions.
It should also be noted that GABA responses are more complex than simply altering membrane potential. Activation of GABAA receptors induced c-Fos immunoreactivity and increased brain-derived neurotrophic factor (BDNF) mRNA expression in embryonic hippocampal neurons but failed to have any effect in older neurons (Berninger et al. 1995). GABA stimulates DNA synthesis (Haydar et al. 2000; LoTurco et al. 1995), and interestingly, appears to regulate KCC2 expression (Ganguly et al. 2001). It is possible that GABA may mediate other effects in chromaffin cells, but this remains to be determined.
Several studies also implicate GABAB receptors in the actions of GABA on catecholamine secretion from chromaffin cells(Castro et al. 1989; Oset-Gasque et al. 1993; Parramon et al. 1995b). Our own studies were not designed to investigate GABAB-mediated responses per se, but they suggest that GABAB receptors were not involved in the actions of GABA described in this paper. Activation of GABAB receptors mobilizes Ca2+ from intracellular stores. Because it takes the intracellular stores in chromaffin cells quite a while to rundown in Ca2+-free solutions, we would expect a Ca2+-transient when GABA was applied in Ca2+-free solution if GABAB receptors were present. Because we saw no response, we concluded that most likely there were no GABAB receptors present in the cells. In mammals, GABAC receptors are only expressed in the retina and the subcortical visual system (Boller and Schmidt 2003) and are therefore unlikely to play a role in the responses reported here.
Overall, the physiological roles of GABA in the adrenal gland are likely to be complex and may involve actions on gene expression, control of membrane potential, calcium influx, and catecholamine release. Our data clearly support an exclusively excitatory role for GABA that is mediated through the selective expression of NKCC1 but not KCC2. It also suggests that clinically relevant drugs that interact with GABAA receptors including anesthetics and other neuroactive agents like benzodiazepines and barbiturates may have stimulatory effects on catecholamine release (Joyce et al. 1982; Kitayama et al. 1989). Further studies will be needed to determine if this is the case and to elucidate other functional consequences of the elevated [Cl–] found in chromaffin cells.
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Address for reprint requests and other correspondence: Z. Xie, The Univ. of Chicago, Dept. of Anesthesia and Critical Care, 5841 S. Maryland, MC 4028, Chicago, IL 60637 (E-mail: jxie{at}airway.bsd.uchicago.edu).
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONDISCLOSURESREFERENCES |
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Amenta F, Collier WL, Erdo SL, Giuliani S, Maggi CA, and Meli A. GABAA receptor sites modulating catecholamine secretion in the rat adrenal gland: evidence from 3H-muscimol autoradiography and in vivo functional studies. Pharmacology 37: 394–402, 1988.[Web of Science][Medline]
Ben-Ari Y. Excitatory actions of gaba during development: the nature of the nurture. Nat Rev Neurosci 3: 728–739, 2002.[Web of Science][Medline]
Berninger B, Marty S, Zafra F, da Penha Berzaghi M, Thoenen H, and Lindholm D. GABAergic stimulation switches from enhancing to repressing BDNF expression in rat hippocampal neurons during maturation in vitro. Development 121: 2327–2335, 1995.[Abstract]
Boller M and Schmidt M. GABAC receptors in the rat superior colliculus and pretectum participate in synaptic neurotransmission. J Neurophysiol 89: 2035–2045, 2003.
Bormann J and Clapham DE. gamma-Aminobutyric acid receptor channels in adrenal chromaffin cells: a patch-clamp study. Proc Natl Acad Sci USA 82: 2168–2172, 1985.
Bormann J, Hamill OP, and Sakmann B. Mechanism of anion permeation through channels gated by glycine and gamma-aminobutyric acid in mouse cultured spinal neurones. J Physiol 385: 243–286, 1987.
Busik J, Nakamura M, Abe Y, Shibuya I, and Kanno T. Effects of GABA on spontaneous [Ca2+]c dynamics and electrical properties of rat adrenal chromaffin cells. Brain Res 739: 97–103, 1996.[Web of Science][Medline]
Castro E, Oset-Gasque MJ, and Gonzalez MP. GABAA and GABAB receptors are functionally active in the regulation of catecholamine secretion by bovine chromaffin cells. J Neurosci Res 23: 290–296, 1989.[Web of Science][Medline]
Chen G, Trombley PQ, and van den Pol AN. Excitatory actions of GABA in developing rat hypothalamic neurones. J Physiol 494: 451–464, 1996.
Currie KP, Wootton JF, and Scott RH. Activation of Ca(2+)-dependent Cl- currents in cultured rat sensory neurones by flash photolysis of DM-nitrophen. J Physiol 482: 291–307, 1995.
Doroshenko PA. gamma-Aminobutyric acid elevates cytosolic Ca in bovine chromaffin cells. Neurosci Lett 104: 83–87, 1989.[Web of Science][Medline]
Eilers J, Plant TD, Marandi N, and Konnerth A. GABA-mediated Ca2+ signalling in developing rat cerebellar Purkinje neurones. J Physiol 536: 429–437, 2001.
Fernandez-Ramil JM, Sanchez-Prieto J, Canadas S, and Gonzalez MP. GABA-T in bovine medulla cells: kinetic properties and comparison with GABA-T from other tissues. Rev Esp Fisiol 39: 299–303, 1983.[Web of Science][Medline]
Fujimoto M, Kataoka Y, Guidotti A, and Hanbauer I. Effect of gamma-aminobutyric acidA receptor agonists and antagonists on the release of enkephalin-containing peptides from dog adrenal gland. J Pharmacol Exp Ther 243: 195–199, 1987.
Ganguly K, Schinder AF, Wong ST, and Poo M. GABA itself promotes the developmental switch of neuronal GABAergic responses from excitation to inhibition. Cell 105: 521–532, 2001.[Web of Science][Medline]
Gonzalez MP, Oset-Gasque MJ, Castro E, Bugeda J, Arce C, and Parramon M. Mechanism through which GABAA receptor modulates catecholamine secretion from bovine chromaffin cells. Neuroscience 47: 487–494, 1992.[Web of Science][Medline]
Haydar TF, Wang F, Schwartz ML, and Rakic P. Differential modulation of proliferation in the neocortical ventricular and subventricular zones. J Neurosci 20: 5764–5774, 2000.
Joyce JT, Roizen MF, Gerson JI, Grobecker H, Eger EI II, and Forbes AR. Induction of anesthesia with halothane increases plasma norepinephrine concentrations. Anesthesiology 56: 286–290, 1982.[Web of Science][Medline]
Kataoka Y, Fujimoto M, Alho H, Guidotti A, Geffard M, Kelly GD, and Hanbauer I. Intrinsic gamma aminobutyric acid receptors modulate the release of catecholamine from canine adrenal gland in situ. J Pharmacol Exp Ther 239: 584–590, 1986.
Kataoka Y, Gutman Y, Guidotti A, Panula P, Wroblewski J, Cosenza-Murphy D, Wu JY, and Costa E. Intrinsic GABAergic system of adrenal chromaffin cells. Proc Natl Acad Sci USA 81: 3218–3222, 1984.
Kitayama S, Morita K, Dohi T, and Tsujimoto A. The nature of the stimulatory action of gamma-aminobutyric acid in the isolated perfused dog adrenals. Naunyn Schmiedebergs Arch Pharmacol 326: 106–110, 1984.[Web of Science][Medline]
Kitayama S, Morita K, Dohi T, and Tsujimoto A. Benzodiazepines facilitate the stimulatory action of gamma-aminobutyric acid (GABA) on basal and veratridine-evoked catecholamine release from cultured bovine adrenal chromaffin cells. Arch Int Pharmacodyn Ther 300: 254–264, 1989.[Web of Science][Medline]
Kitayama S, Morita K, Dohi T, and Tsujimoto A. GABAA receptor-mediated increase of cytosolic Ca2+ in isolated bovine adrenal chromaffin cells. Biochim Biophys Acta 1053: 189–194, 1990a.[Medline]
Kitayama S, Morita K, Dohi T, and Tsujimoto A. Enhancement by GABA of the stimulation-evoked catecholamine release from cultured bovine adrenal chromaffin cells. Naunyn Schmiedebergs Arch Pharmacol 341: 414–418, 1990b.[Web of Science][Medline]
Kitayama S, Morita K, Dohi T, and Tsujimoto A. GABAergic modulation of catecholamine release from cultured bovine adrenal chromaffin cells. Evidence for the involvement of Cl(–)-dependent Ca2+ entry. Naunyn Schmiedebergs Arch Pharmacol 341: 419–424, 1990c.[Web of Science][Medline]
Kitayama S, Nakatsukasa Y, Morita K, Dohi T, and Tsujimoto A. Pharmacological evidence for the possible involvement of repetitive action potentials in facilitation by GABA of catecholamine secretion in bovine adrenal chromaffin cells. Br J Pharmacol 102: 706–710, 1991.[Web of Science]
LoTurco JJ, Owens DF, Heath MJ, Davis MB, and Kriegstein AR. GABA and glutamate depolarize cortical progenitor cells and inhibit DNA synthesis. Neuron 15: 1287–1298, 1995.[Web of Science][Medline]
Luhmann HJ and Prince DA. Postnatal maturation of the GABAergic system in rat neocortex. J Neurophysiol 65: 247–263, 1991.
Mikawa S, Wang C, Shu F, Wang T, Fukuda A, and Sato K. Developmental changes in KCC1, KCC2 and NKCC1 mRNAs in the rat cerebellum. Brain Res Dev Brain Res 136: 93–100, 2002.[Medline]
Mueller AL, Taube JS, and Schwartzkroin PA. Development of hyperpolarizing inhibitory postsynaptic potentials and hyperpolarizing response to gamma-aminobutyric acid in rabbit hippocampus studied in vitro. J Neurosci 4: 860–867, 1984.[Abstract]
Obukhov AG and Nowycky MC. TRPC4 can be activated by G-protein-coupled receptors and provides sufficient Ca(2+) to trigger exocytosis in neuroendocrine cells. J Biol Chem 277: 16172–16178, 2002.
Oomori Y, Iuchi H, Nakaya K, Tanaka H, Ishikawa K, Satoh Y, and Ono K. Gamma-aminobutyric acid (GABA) immunoreactivity in the mouse adrenal gland. Histochemistry 100: 203–213, 1993.[Web of Science][Medline]
Oset-Gasque MJ, Canadas Correas S, Masso Cordoba JM, and Gonzalez Gonzalez MP. Uptake of GABA by bovine adrenal medulla slices. Naunyn Schmiedebergs Arch Pharmacol 329: 372–375, 1985.[Web of Science][Medline]
Oset-Gasque MJ, Castro E, and Gonzalez MP. Mechanisms of [3H] gamma-aminobutyric acid release by chromaffin cells in primary culture. J Neurosci Res 26: 181–187, 1990.[Web of Science][Medline]
Oset-Gasque MJ, Parramon M, and Gonzalez MP. GABAB receptors modulate catecholamine secretion in chromaffin cells by a mechanism involving cyclic AMP formation. Br J Pharmacol 110: 1586–1592, 1993.[Web of Science][Medline]
Owens DF, Boyce LH, Davis MB, and Kriegstein AR. Excitatory GABA responses in embryonic and neonatal cortical slices demonstrated by gramicidin perforated-patch recordings and calcium imaging. J Neurosci 16: 6414–6423, 1996.
Owens DF and Kriegstein AR. Is there more to GABA than synaptic inhibition? Nat Rev Neurosci 3: 715–727, 2002.[Web of Science][Medline]
Parramon M, Gonzalez MP, Herrero MT, and Oset-Gasque MJ. GABAB receptors increase intracellular calcium concentrations in chromaffin cells through two different pathways: their role in catecholamine secretion. J Neurosci Res 41: 65–72, 1995b.[Web of Science][Medline]
Parramon M, Gonzalez MP, and Oset-Gasque MJ. Pharmacological modulation of adrenal medullary GABAA receptor: consistent with its subunit composition. Br J Pharmacol 116: 1875–1881, 1995a.[Web of Science][Medline]
Payne JA, Stevenson TJ, and Donaldson LF. Molecular characterization of a putative K-Cl cotransporter in rat brain. A neuronal-specific isoform. J Biol Chem 271: 16245–16252, 1996.
Peters JA, Lambert JJ, and Cottrell GA. An electrophysiological investigation of the characteristics and function of GABAA receptors on bovine adrenomedullary chromaffin cells. Pfluegers 415: 95–103, 1989.
Rivera C, Voipio J, Payne JA, Ruusuvuori E, Lahtinen H, Lamsa K, Pirvola U, Saarma M, and Kaila K. The K+/Cl– co-transporter KCC2 renders GABA hyperpolarizing during neuronal maturation. Nature 397: 251–255, 1999.[Medline]
Rohrbough J and Spitzer NC. Regulation of intracellular Cl– levels by Na(+)-dependent Cl– cotransport distinguishes depolarizing from hyperpolarizing GABAA receptor-mediated responses in spinal neurons. J Neurosci 16: 82–91, 1996.
Sangiah S, Borowitz JL, and Yim GK. Actions of GABA, picrotoxin and bicuculline on adrenal medulla. Eur J Pharmacol 27: 130–135, 1974.[Web of Science][Medline]
Staley K, Smith R, Schaack J, Wilcox C, and Jentsch TJ. Alteration of GABAA receptor function following gene transfer of the CLC-2 chloride channel. Neuron 17: 543–551, 1996.[Web of Science][Medline]
Sun D, Murali SG. Na+-K+-2Cl– cotransporter in immature cortical neurons: a role in intracellular Cl-regulation. J Neurophysiol 81: 1939–1948, 1999.
Sung KW, Kirby M, McDonald MP, Lovinger DM, and Delpire E. Abnormal GABAA receptor-mediated currents in dorsal root ganglion neurons isolated from Na-K-2Cl cotransporter null mice. J Neurosci 20: 7531–7538, 2000.
Vardi N, Zhang LL, Payne JA, and Sterling P. Evidence that different cation chloride cotransporters in retinal neurons allow opposite responses to GABA. J Neurosci 20: 7657–7663, 2000.
Yuste R and Katz LC. Control of postsynaptic Ca2+ influx in developing neocortex by excitatory and inhibitory neurotransmitters. Neuron 6: 333–344, 1991.[Web of Science][Medline]
Zhang SJ and Jackson MB. GABA-activated chloride channels in secretory nerve endings. Science 259: 531–534, 1993.
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