Alcohol Tolerance in Large-Conductance, Calcium-Activated Potassium Channels of CNS Terminals Is Intrinsic and Includes Two Components: Decreased Ethanol Potentiation and Decreased Channel Density

Explant preparation and culture

Male Sprague Dawley adult rats (3–5 weeks old) were obtained from Taconic Farms (Germantown, NY). All procedures were performed in accordance with the National Institutes of Health guidelines on the Care and Use of Animals and study protocol approved by the University of Massachusetts Animal Care and Use Committee. After decapitation, the brain and pituitary with intact pituitary stalk were removed very rapidly from the skull and chilled in oxygenated dissection buffer (in mm): 120 NaCl, 5 KCl, 1.0 MgCl₂, 1.9 CaCl₂, 2.0 EGTA, 10 HEPES, 5.0 NaHCO₃, 15.0 glucose, 0.5 glutamine (Glutamax; Invitrogen, Gaithersburg, MD), and 100 U/ml to 0.1 mg/ml penicillin–streptomycin (Invitrogen), pH 7.2, 310 mOsm. To increase cell survival, dissection buffer contained: (1) low levels of free calcium (3 μm), (2) B27 supplement with antioxidants (1:50; Invitrogen), and (3) 1 mm nitro-arginine and kynurenic acid. Under a dissection microscope, the anterior pituitary was removed, and a hexagonal block of tissue was carved out (by cutting rostral to the optic chiasm, parallel to the optic tract, lateral to the median eminence and caudal to the stalk, and undercutting at a depth of 1–2 mm). Next, tissue was trimmed to the final thickness of 550–650 μm to enable nutrients and oxygen penetration despite the lack of vascular perfusion (Fujii et al., 1982; Nicholson and Hounsgaard, 1983). Explants were cultured individually in a humidified incubator, at 34°C, in 5% CO₂, on Millicell CM inserts (Millipore, Bedford, MA), placed in Petri dishes (60 mm; Corning, Corning, NY). They were cultured in Neurobasal A medium (310 mOsm) with B27, glutamine, penicillin–streptomycin (as in dissection buffer), and 10% fetal calf serum (FCS; Sigma, St. Louis, MO) in the air-medium interface (Stoppini et al., 1991; Brewer, 1997; House et al., 1998; Xiang et al., 2000). Medium was changed on the day after dissection to ethanol-free medium (naive group) or medium containing 20 mm ethanol (chronic group). The control (uncultured) group was immediately fixed after isolation.

Ethanol treatment

A constant ethanol concentration was maintained by placing an open dish with 20 mm ethanol near the dish containing the ethanol-treated explant, in a closed, but not sealed container, which allowed exchange of gases. Ethanol levels measured in both the media and the tissue, using an NAD-ADH kit (Sigma) according to manufacturer's instructions, indicated a constant level of 20 mm ethanol at all exposure times tested (0.5, 1, 3, 6 and 24 hr). For exposures <30 min, terminals were dispersed first and then exposed to ethanol during electrophysiological recording.

Viability assay

Immunostaining. Explants were fixed in 4% paraformaldehyde (PFA). After immersion in 20% sucrose, explants were embedded in 6% gelatin–egg yolk mixture in cardboard molds, and exposed to concentrated formaldehyde vapors for at least 3 d (4°C). After hardening, 60 μm sections were cut on a freezing microtome, and sections were placed in Net-well carriers. The floating sections were permeabilized and blocked and incubated alternately with the primary monoclonal mouse antibodies: G38 (against AVP-neurophysin) or G41 (anti-OXT–neurophysin) (1:300; a gift from H. Gainer, Laboratory of Neurochemistry, National Institute on Neurological Disorders and Stroke, National Institutes of Health, Bethesda, MD) (Ben Barak et al., 1985; Whitnall et al., 1985). Next, sections were incubated with Alexa 488-tagged, anti-mouse secondary antibody (Molecular Probes, Eugene, OR), counterstained with Hoechst dye (1:5000), and mounted on microslides (SuperFrost Plus; VWR Scientific, West Chester, PA) in Prolong Antifade medium (Molecular Probes). Alternatively, sections were incubated with biotinylated secondary antibody, processed using Vectastain Elite ABC kit and Peroxidase Substrate kit (Vector Laboratories, Burlingame, CA), counterstained with Hematoxylin QS (Vector Laboratories), dehydrated with ascending alcohols, cleared with xylene, and mounted in VectaMount (Vector Laboratories).

Unbiased stereological determination of magnocellular neuron viability. An unbiased stereological technique (West et al., 1991) was used to estimate the total number of AVP and OXT neurons in the supraoptic nucleus (SON) and its retrochiasmatic part (SONr). All stereological analyses were performed using Stereo Investigator 4.33 (MicroBrightField, Colchester, VT) and a Leica DMRE microscope (Deerfield, IL). We used 18 (n = 6 for each group) supraoptic nuclei in the analysis. All sections were alternately stained against AVP or OXT (starting randomly with either AVP or OXT antibodies), and AVP- or OXT-positive magnocellular cell bodies were counted. Healthy cells had large cell bodies (∼40 μm diameter) filled with hormone, with long processes and a large nonpyknotic nucleus. Magnocellular neurons with shrunken cytoplasm and/or pyknotic nuclei were considered apoptotic (Vutskits et al., 1998). Apoptotic cells constituted <3% of neurons in each group and were not included in the final cell number. Estimates of the total number of neurons were generated by the Stereo Investigator using a fractionator correction factor of 7.42. Differences between left and right nuclei were negligible. Sampling was performed to maintain a coefficient of error <10% (Gundersen and Jensen, 1987; Gundersen et al., 1999).

NH terminal preparation

Neurohypophysial terminals were homogenized as previously described (Cazalis et al., 1987) in a solution containing 270 mm sucrose, 10 mm HEPES, and 1 mm EGTA, pH 7.3. The homogenate was then plated onto a poly-d-lysine-coated 35 mm polystyrene dish or a coverslip, and after attachment, terminals underwent electrophysiological measurements or immunocytochemistry.

Electrophysiology

Dissociated terminals were identified by their spherical shape, size (2–8 μm in diameter), lack of nuclei, and golden color under Hoffmann–phase contrast optics. First they were bathed in ethanol-free low-calcium (3 μm) Locke's solution (in mm: 142 NaCl, 2.0 KCl, 2.0 CaCl₂, 2.0 MgCl₂, 13.0 glucose, and 15.0 HEPES, pH 7.3, 305 mOsm), which subsequently was changed into ethanol-free normal calcium Locke's solution (2.2 mm CaCl₂, no EGTA). Average time before ethanol challenge was 60 min.

Whole-cell recordings

Electrodes were pulled from thin-walled borosilicate glass (Drummond Scientific, Broomall, PA) on a vertical puller (David Kopf Instruments, Tujunga, CA) and fire-polished (Narashige, Tokyo, Japan) to give a final resistance of 5–10 MΩ (2–4 MΩ for perforated patch) when filled with pipette solution (in mm): 120.0 potassium gluconate, 15.0 HEPES, 2.2 CaCl₂, 1.0 MgCl₂, 4.0 EGTA, and 4.0 HEDTA, pH 7.35, 305 mOsm (∼4.0 μm free Ca²⁺). For perforated patch technique, 30 μm amphotericin B was added to the pipette solution (120.0 potassium gluconate, 15.0 KCl, 10 HEPES, 2.2 CaCl₂, and 1.0 MgCl₂, pH 7.3, 295 mOsm). Currents were recorded using an Axopatch 200B (Axon Instruments, Foster City, CA) or a Dagan amplifier at a sampling rate of 1–5 kHz and leak-subtracted on line. Series resistance never exceeded 15 MΩ and was compensated. Data were acquired and analyzed with pClamp6 (Axon Instruments).

Single-channel recordings

Unitary current recordings were obtained from excised, inside-out patches using standard patch-clamp techniques. Currents were recorded in symmetrical 135 mm K⁺ using a patch-clamp amplifier (Axopatch 200; Axon Instruments) at a sampling rate of 5 kHz and filtered at 1 kHz using an eight pole Bessel filter. Data were acquired and analyzed using pClamp 6.02 software.

Superfusion and drug application

Terminals in whole-cell configuration or patches excised from the membrane were exposed to 5 μm [Ca²⁺]_free control solution flowing from a micropipette (1 mm diameter; WPI, Sarasota, FL), and the potential across the membrane was set to produce a relatively low open probability (NP_o ∼0.2). BK channel activity was recorded for 30 sec, and then patches were superfused with ethanol and washed. Because chronic ethanol exposure reduces BK potentiation by subsequent acute ethanol challenge (Knott et al., 2002), we used a high (100 mm) ethanol concentration for acute challenge, to ensure a measurable ethanol effect. When studying gating properties of the BK channel, the intracellular surface of patches was exposed to solutions with a final [Ca²⁺]_free of 1, 5, and 10 μm. The calculation of [Ca²⁺]_free was computed using a custom-written program, based on calculations by Fabiato (1988). At each potential, channel activity was recorded for 30 sec.

In the perforated patch configuration, BK current was isolated by adding 4-aminopyridine to the bath (7 mm final concentration) to block the fast inactivating (I_A) potassium current (Thorn et al., 1991; Knott et al., 2002). Baseline current was first established by stepping from the holding potential of ∼90 mV to +40 mV, for 500 msec, with a15 sec recovery interval between steps. During ethanol application, BK current was determined by averaging a set of four traces obtained within each minute. Ethanol potentiation of the BK current was expressed as a percentage above the baseline level.

Data analysis

NP_o was determined in the presence of 10 μm Ca²⁺ and was used as an index of channel activity. NP_o values were calculated from all-points amplitude histograms by fitting the histogram with a sum of Gaussian functions using a Levenberg–Marquardt algorithm. NP_o data as a function of voltage were fitted with a Boltzmann function of the type: P_o = (1 + exp – K(V – V_0.5))^–1, where K is the logarithmic potential sensitivity and V_0.5 the potential at which P_o is half-maximal. When the NP_o–voltage relationship is fitted by a Boltzmann curve, a plot of lnNP_o versus voltage is linear at low values of P_o (Dopico et al., 1996, 1998). In this plot, the reciprocal of the slope is the potential needed to produce an e-fold change in NP_o, which is routinely used to measure the voltage dependence of BK channel gating.

The relationship between [Ca²⁺]_i and the channel steady-state P_o is usually a sigmoid function that can be fitted according to a Hill equation: P_o = P_max[Ca²⁺]_i^nH/(K + [Ca²⁺]_i^nH)where P_o is the steady-state open probability, P_max is the maximum open probability, n^H is the slope factor (or Hill coefficient), and the nth root of K gives an estimate of the midpoint of the activation curve and the sensitivity of the channel to [Ca²⁺]_i. The slope factor provides a minimum estimate of the number of Ca²⁺ ions involved in maximally activating the channel (McManus, 1991). Thus, the required [Ca²⁺]_i to achieve the midpoint of activation (P_0.5) at 0 mV, usually designated [Ca²⁺]_0.5, is a widely used parameter to measure the calcium sensitivity of BK channels.

Duration of open times was measured with half-amplitude threshold analysis. Mean open time was calculated based on an equation: T_o = NP_oT/X_o for multiple channel recordings [N = number of channels; P_o = open probability; T = time of observation (in milliseconds); X_o = number of openings]. A maximum-likelihood minimization routine was used to fit curves to the distribution of open times. Determination of the minimum number of terms for adequate fit was established using a standard F statistic table (significance level, p < 0.01). The unitary conductance (γ) was taken as the slope of the unitary current amplitude–voltage relationship. Values for unitary current were obtained from the Gaussian fit of all-points amplitude histograms by measuring the distance between the modes corresponding to the closed state and the first opening level. For all experiments, voltages given correspond to the potential at the intracellular side of the membrane.

Immunocytochemistry, confocal analysis, and quantification of BK expression

Terminals were isolated as described above, fixed in 2% PFA, and double-labeled using mouse antibodies against AVP (or OXT) and rabbit antibodies against BK channel α subunit (1:200; Affinity Bioreagents, Golden, CO) and secondary antibodies (anti-mouse Alexa 594, anti-rabbit Alexa 488, respectively; Molecular Probes). Coverslips were then mounted to microslides (SuperFrost Plus; VWR Scientific) using Prolong Antifade medium (Molecular Probes). Laser-scanning fluorescence and differential interference contrast images were acquired simultaneously using a Leica laser-scanning microscope. The laser power was set to avoid photobleaching. Intensity of the signal assessed before and after capturing of images showed no decrease or saturation. AVP and OXT terminals of similar size (average: 3.31 ± 0.36 μm) were scanned using six to eight Z-sections (average: 420 ± 30.0 nm/step). Using Leica software (version 2.00; Microsystems, Heidelberg, Germany), two to four middle sections of each terminal were selected for the analysis. Stacks of images were saved as TIFF files and analyzed off-line. The number of BK clusters, their size, and the intensity profiles were determined. The intensity of specific BK staining in the terminals was calculated by subtracting average background intensity (<1% of the specific staining), obtained from three nonfluorescent areas. The quantitative analysis was performed blind using a Photoshop-based (Adobe Systems, San Jose, CA) protocol. Each section of the terminal was divided into a membrane region and an interior region, using the assumption that the membrane-associated fluorescent signal has a width of 0.45 μm (Chiu et al., 2002). Boundaries of regions were determined using the DIC image of the terminal overlapped onto the BK expression image. In control experiments using antibodies preincubated with the antigenic peptides, we observed no specific staining.

Statistics

Unless otherwise indicated, statistical significance between the various treatment groups was analyzed using Student's t test (Statistica, version 5.5; StatSoft, Tulsa, OK). All data are expressed as mean ± SEM, and p values <0.05 were considered to be statistically significant.

HNS explants contain viable AVP and OXT magnocellular neurons with intact axons

Because of the nonplanar route of axons from the SON to the NH (Silverman and Zimmerman, 1983; Armstrong, 1995), it is not possible to prepare an intact HNS explant using a straight slice technique. Thus, it was necessary to “sculpt” the tissue manually with microtools, using caution to avoid axotomy. The resulting HNS explants were laid on Millipore CM inserts, in Petri dishes in the interface between medium and air. An image of an actual HNS explant is shown in Figure 1A. Elements of the optic system [optic nerves (ON), optic chiasm (OC), optic tract (OT)] are apparent and were used as landmarks in assessing the position of the supraoptic nuclei (marked as SON and SONr for the retrochiasmatic part of the SON), where cell bodies of the neurons of interest are located (Fig. 1B). To ensure that the axonal tract was intact in the HNS explant, we performed immunofluorescent staining using antibodies against AVP and OXT. A semi-sagittal section of the AVP-stained explant in Figure 1C confirms that the axonal path from cell bodies to terminals was well preserved in the explant. Similar results were obtained for OXT-containing neurons (data not shown).

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Figure 1.

HNS explant contains viable AVP and OXT neurons of the intact SON–NH. A, Bright-field top view of an HNS explant. B, Diagram of HNS explant showing position of magnocellular neuron compartments: cell bodies, axons, and terminals. C, Immunofluorescent image (above) and sagittal diagram (below) of the SON–NH tract (AVP-positive, similar OXT-positive results not shown) indicate that axonal projections from SON magnocellular neurons to the neurohypophysis were preserved, and the SON–NH tract is the only structure present in the HNS explant. Note the winding path of axons from the SON, above the optic tract, below the sole of the third ventricle, through the median eminence to the neurohypophysis. D, Magnocellular neurons in SON were counted to estimate the viability of the HNS explants. Representative immunofluorescent grayscale images (top) of AVP neuron in the SON from explants cultured in 20 mm ethanol for 24 hr (similar data for OXT and fresh and naive explants not shown). Note the large cell body with long processes filled with hormone (left image) and large, nonapoptotic nucleus with diffuse Hoechst staining (right image, arrow).Other neurons are not visible because of the irout-of-focus location. The exact number of neurons in SON was estimated using stereological techniques (see Material and Methods) and is shown in the graph (bottom). Cap on each bar represents SD. ME, Median eminence; MM, medial mammilliary nucleus; PVN, paraventricular nucleus; 3V, third ventricle. Scale bars: A, 1.5 mm; C, 430 μm; D, 20 μm.

Chronic, low-dose ethanol treatment does not influence magnocellular neuron viability

High doses of ethanol (∼100 mm) are known to trigger neuronal apoptosis (Madeira et al., 1993; Ruela et al., 1994), therefore we limited our chronic exposure to clinically relevant (20 mm) concentrations. In exposures of ≥30 min, ethanol penetrated the tissue of the HNS explant and was maintained at a constant 20 mm level throughout the treatment. Viability of magnocellular neurons was unaffected by 48 hr culture with or without 24 hr ethanol treatment (Fig. 1D). Viable AVP and OXT magnocellular neurons have large cell bodies and processes filled with hormone. Using Hoechst dye, we observed viable cells, in which the large, weakly stained nuclei with loosely coiled chromatin suggested the absence of apoptosis (Bonde et al., 2002). Many neurons with these features were observed in the HNS explant. An example of such a cell is shown in Figure 1D. To precisely determine the influence of culturing and ethanol exposure on survival of AVP and OXT magnocellular neurons, we used stereology to assess the number of viable neurons. In freshly isolated tissue (Fig. 1D, graph) the SON contained 3080 ± 42 AVP neurons and a slightly smaller number of OXT neurons (2170 ± 38), in accordance with previous reports (Swaab et al., 1975; Rhodes et al., 1981; Marciano et al., 1989; Harding et al., 1995; House et al., 1998; Burbach et al., 2001). Explants cultured in ethanol-free medium contained 2996 ± 78 AVP cells and 1979 ± 118 OXT cells, whereas explants cultured in 20 mm ethanol contained 3136 ± 76 AVP and 1993 ± 35 OXT-positive cells, respectively. Apoptotic, hormone-positive (AVP or OXT) cells constituted <3% of the total number of SON neurons in fresh explants, as well as in explants cultured in the presence or absence of ethanol. Thus, we did not observe increased cell death during culture or after ethanol exposure.

BK channels retain normal properties in cultured HNS explant

Figure 2A shows typical single-channel recordings from an inside-out patch from a BK channel in a NH terminal from a naive explant. We tested the voltage dependence of this channel by exposing its intracellular face to 5 μm free Ca²⁺ and changing the holding potential. At –60 mV, the BK channel remained mostly in a closed state and showed only brief openings. As more depolarized potentials (–40 and –20 mV) were reached, the time spent in the open state increased (Fig. 2A). We also tested the calcium dependence of BK channels by holding the membrane potential at +30 mV and exposing the cytoplasmic face of the channel to increasing Ca²⁺ concentrations (Fig. 2B). In the presence of 1 μm Ca²⁺, the BK channel showed only sporadic openings. After switching to 5 μm Ca²⁺, channel activity increased, to finally reach near maximum open probability in 10 μm Ca²⁺. Finally, a unitary conductance of 215 pS, a value typical of BK channels, was observed in symmetrical 135 mm K⁺ (Fig. 2C). Thus, BK channel basic properties (Wang et al., 1992; Dopico et al., 1996, 1999a) remained unchanged during HNS explant preparation or culture. We also examined the dependence of ethanol effects on the membrane potential between –10 and +70 mV (Fig. 2D) in whole-cell recordings. We found that 50 mm ethanol potentiated BK currents at all potentials tested (Fig. 2D) and that this effect was voltage-independent (Fig. 2D, inset). Traces in Figure 2D show representative ethanol-mediated potentiation of BK currents recorded from a naive terminal and evoked at different membrane potentials (20 mV steps) before (control) and after 50 mm ethanol application.

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Figure 2.

Voltage and calcium dependence of BK channels in naive NH terminals. A, Representative currents from a single BK channel, recorded in an inside-out patch at different potentials in the presence of 10μm free [Ca²⁺]_i. Channel activity increases as the patch membrane is held at more depolarized potentials. B, Activity of a single BK channel as in A recorded at +30 mV in the presence of various [Ca²⁺]_i. The activity increases as the cytosolic face of the channel is exposed to increasing concentrations of calcium. C and O₁, adjacent to the traces, indicate the closed and the open states, respectively. C, Unitary conductance was determined by fitting the I–V relationship by linear regression.D, Ethanol-mediated potentiation of BK current is voltage-independent. Graph shows the average BK current amplitude (n = 5) as a function of the membrane voltage measured in whole-cell voltage clamp. Each terminal served as its own control. A 50 mm concentration of ethanol increased BK current amplitude at all the potentials tested between –10 and +70 mV in a voltage-independent manner (inset). Representative traces show BK currents recorded from a naive terminal and evoked at different membrane potentials (20 mV steps) before (control) and after 50 mm ethanol application. The fast transient current at the beginning of the voltage pulse is caused by sodium channels (TTX was omitted from the bath solution).

BK channel tolerance to ethanol in terminals is intrinsic

Next, we examined whether 24 hr chronic ethanol treatment of the explant produces changes in BK channel properties similar to those observed in terminals obtained from animals on an ethanol diet (Knott et al., 2002). First, we compared the effect of ethanol on BK channels isolated from fresh terminals to BK channels obtained from explants cultured in ethanol-free medium. Ethanol increased the open probability of BK channels from freshly isolated terminals and did not change channel unitary current amplitude (Fig. 3A, top panels), in agreement with previous findings (Dopico et al., 1996). After 2 d in culture in ethanol-free medium, BK ethanol potentiation was unaltered (Fig. 3A, middle panels). On average, ethanol increased BK channel open probability by more than fivefold (Fig. 3B) (5.26 ± 1.87; n = 7). Interestingly, 24 hr of 20 mm ethanol treatment of the explant produced significantly reduced BK channel potentiation by acute ethanol challenge (1.7 ± 1.1-fold; n = 12) (Fig. 3A, bottom panels, B)(p < 0.001).

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Figure 3.

BK channels exhibit intrinsic tolerance to ethanol after 24 hr. A, Effects of acute ethanol in freshly isolated, naive, and chronic explants. Inside-out terminal patches were exposed to 5 μm free-Ca²⁺ control solution, and the potential across the membrane was set to produce low open probability (NP_o around 0.2), to allow observation of ethanol (100 mm) potentiation. Whereas potentiation is evident in freshly isolated and naive terminals, chronic terminals show lack of ethanol effect on BK activity as shown in representative traces at the left of each panel. All-points amplitude histograms summarizing these effects are shown on the right.The washout histogram is omitted from the graph for clarity. Differences in unitary current amplitude reflect different holding potentials that were –50, –50, and –30 mV for patches from fresh, naive, and chronic conditions, respectively. B, The increase of BK channel open probability by acute ethanol challenge in naive (n = 7) and chronic (n = 12) explants. BK channel activity is expressed as the ratio of BK channel open probability after acute ethanol challenge over its activity before drug application (NP_o EtOH /NP_o Control), where 1 = no potentiation. Chronic ethanol treatment decreases BK channel potentiation by ethanol (p < 0.05) compared with the naive group. C, BK currents in terminals were recorded in whole-cell patch clamp to determine current densities. Membrane potential was held at –70 mV and depolarized for 500 msec from –50 to +90 mV in 20 mV increments. D, Average current densities (±SEM) recorded in naive (n = 5) and chronic (n = 4) terminals were significantly smaller (∼5-fold) after chronic ethanol exposure. Current densities are expressed as picoamperes per 10 μm² (membrane capacitance was used to assess membrane area).

In addition to the decreased potentiation by acute drug challenge, BK current densities obtained in terminals from explants exposed to ethanol for 24 hr were significantly lower than in terminals from naive explants. In terminals from naive explants, depolarizing the membrane to +50 mV (from –70 mV) evoked a strong BK current (196 ± 43 pA). In contrast, after 24 hr ethanol treatment, a similar voltage step elicited only 56 ± 36 pA of BK current (Fig. 3C). When current was expressed as a function of membrane surface area (determined by membrane capacitance), we found a dramatic decrease in current density after chronic ethanol treatment, from 14.2 ± 3 pA/10 μm² (n = 5) in naive explants to 2.71 ± 2 pA/10 μm² (n = 4) in chronic explants (p < 0.05) (Fig. 3D).

Thus, chronic tolerance to ethanol observed in neurohypophysial terminals from cultured explants was intrinsic and did not require the actions of elements originating outside of the HNS.

Time course of tolerance suggests two distinct processes

We next determined the time course, beginning at 30 min, of the two components of tolerance evident at 24 hr. Macroscopic currents in terminals from HNS explants cultured for various periods of time in ethanol were obtained, using whole-cell patch clamp recordings. Two distinct time courses were observed. A rapid reduction in BK ethanol potentiation was evident within 30 min of drug exposure and was sustained for the 24 hr test period, whereas BK current density responded more slowly to the ethanol treatment, showing partial reduction in current density after 6 hr of ethanol exposure, and reaching a near-minimal value after 24 hr (Fig. 4A). The finding that decreased potentiation was already fully present at the 30 min time point led us to examine the development of this form of tolerance during the first few minutes of ethanol exposure. Because short-latency changes might be complicated by a time lag attributable to ethanol penetration into the explant, we altered the protocol, to examine the reduction of ethanol potentiation of BK current during continued exposure to ethanol using the perforated patch clamp technique of an isolated terminal from an ethanol-naive rat. It is evident (Fig. 4B) that potentiation is decreasing within 4 min of drug exposure, and this form of tolerance is fully developed by 12 min of exposure. Introduction of an incremental 50 mm ethanol (final concentration, 100 mm) at the 13 min time point did not result in the reinitiation of potentiation (data not shown). Representative traces are shown in Figure 4C.

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Figure 4.

Time course of the development of ethanol tolerance of BK channels. A, HNS explants were cultured on Millipore inserts and exposed to ethanol for 0, 0.5, 1, 6, and 24 hr (n = 4–10). Ethanol tolerance of BK channels in the cultured NH terminals is manifested as both decreased sensitivity to ethanol, evident within 30 min of drug exposure and sustained for the entire 24 hr observation period, and reduced BK current density, evident after 6 hr, and continuing to decrease during the 24 hr observation period. B, The time course of decreased potentiation of BK current during the first few minutes was determined using the perforated patch-clamp technique with an isolated terminal from an ethanol-naive neurohypophysis (n= 2). The graph demonstrates the rapid loss of potentiation in the continued presence of ethanol. C, Representative traces of BK current from specified time points in B.

Single-channel properties of BK remain unchanged after chronic ethanol treatment

The observed decrease in current density could result from either altered individual channel properties or from a downregulation of functional channels (or both). We tested the first possibility by examining gating properties, kinetics, and conductance of terminal BK channels from naive and chronic (24 hr ethanol exposure) explants.

Voltage and calcium activation

Figure 5A shows that the open probability of BK channels obtained from naive (open markers) and chronic (filled markers) terminals, as a function of calcium concentrations and voltage, does not differ. In both groups, for each [Ca²⁺]_i the NP_o–voltage relationship could be well fitted with a sigmoid function (see Materials and Methods). Figure 5B shows the data plotted in a manner that illustrates the lack of effect of chronic treatment on BK channel voltage sensitivity. The reciprocal of the slope of the ln(P_o)–V relationship at low P_o in the presence of 10 μm free-Ca²⁺ indicated that the voltage necessary to produce an e-fold change in open probability was similar in naive (28 ± 1.22 mV) and chronic terminals (29 ± 1.21 mV, see Materials and Methods).

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Figure 5.

Chronic ethanol does not alter BK channel characteristics. A, Normalized NP_o as a function of voltage at three different [Ca²⁺]_i in naive (open symbols) and chronic (filled symbols) BK channels were fitted with a Boltzmann relationship of the type: P_o = (1 + exp – K(V – V_0.5)) ^–1, where K is the logarithmic potential sensitivity, and V_0.5 the potential at which P_o is half-maximal. At each calcium concentration, naive and chronic BK channels are very similar. The P_o–V curve is shifted similarly along the voltage axis to more negative potentials as [Ca²⁺]_i is increased. B, Data plotted in a manner that illustrates the lack of effect of chronic treatment on BK channel voltage sensitivity. The reciprocal of the slope of the ln(P_o)–V relationship at low P_o in the presence of 10μm free-Ca²⁺ indicated that the voltage necessary to produce an e-fold change in open probability was similar in naive (28 ± 1.22 mV) and chronic terminals (29 ± 1.21 mV). C, V_0.5 values, obtained from Boltzmann fits displayed in A (dashed line), are now plotted as a function of [Ca²⁺]_i on a logarithmic scale. Data points are fitted by a linear regression (r = 0.99). The leftward shift inV_0.5 as [Ca²⁺]_i increases is very similar in naive and chronic BK channels. D, The averaged mean open time of BK channels from naive (n = 6) and chronic explants (n = 7) are not different. Mean open time was calculated based on the equation: T_o = NP_oT/Xo for multiple channel recordings [N, Number of channels; P_o, open probability; T, time of observation (in milliseconds), X_o, number of openings]. E, The mean BK channel conductance from naive (n = 12) and chronic (n = 11) terminals obtained by fitting I–V plots (Fig. 2C) are not different.

We compared the calcium sensitivity of BK channels from naive and chronic explants. This is illustrated by the plot of V_0.5 (in millivolts) as a function of free-Ca²⁺, in which V_0.5 decreases as [Ca²⁺]_i increases (Fig. 5C). We observed no difference between the naive and the chronic group. We also calculated the Hill coefficient by fitting the slope of the relationship between NP_o and [Ca²⁺]_i and found it was similar in naive (1.15) and chronic explants (1.07; plot not shown).

Chronic ethanol causes declustering and redistribution of BK channels

The finding that single-channel properties were unaltered after the development of tolerance suggested that the decrease in current density might result from a decrease in BK channel expression in the terminal membrane. To address this question, we measured expression of BK channel protein in terminals, using a specific antibody directed against the main (pore-forming) α subunit of the channel. Although both naive and chronic explants contained BK channels in both the membrane and interior of the terminals, their distribution was different (Fig. 6A). Quantitative analysis revealed that chronic ethanol did not significantly change the total BK expression in the terminals, but shifted the expression of BK from the membrane to the interior (Fig. 6B, total). In naive explants, membrane-associated and internal BK channels constituted ∼20 and 80%, respectively, of the total BK expressed in these terminals. After chronic ethanol treatment, many more BK channels were found in the interior (94%), whereas membrane-expressed BK channels comprised only 6% of the total BK. Thus, chronic ethanol treatment caused a significant (p < 0.05) decrease in the proportion of BK channels localized in the membrane of neurohypophysial terminals and a significant (p < 0.05) increase in the proportion of BK channel present in the interior (Fig. 6B, bottom panels).

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Figure 6.

Chronic ethanol changes distribution of BK channels in neuronal terminals. A, Representative immunofluorescent confocal images (in a grayscale) of naive and chronic HNS terminals stained for BK pore-forming α subunit. Neurohypophysial terminals were homogenized and dropped onto a poly-l-lysine-coated Petri dish, fixed in paraformaldehyde, and double-labeled against AVP–OXT and BK channel α subunit. Laser-scanning fluorescence and DIC images were acquired simultaneously using a Leica laser-scanning microscope. Boundaries of the terminals were determined using DIC image, and each section of the terminal was divided into a membrane and an interior regions based on a membrane-associated fluorescence signal width of 0.45 μm (Chiu et al., 2002). Scale bar, 700 nm. B, Quantitative analysis in naive (n = 12) and chronic (n = 14) terminals showed that total BK expression in NH terminals is not significantly changed after chronic ethanol treatment (total). In contrast, membrane-associated BK expression decreases (membrane), whereas the amount of BK expressed in the interior increases (interior) after chronic ethanol treatment. *p < 0.05.

In addition to BK channel redistribution, we also found that chronic ethanol disturbed the expression pattern of BK channels in the terminal membrane. In terminals obtained from naive explants, BK channels were present mainly in clusters, whereas in the membrane of the terminals from chronic explants, clusters were rarely seen, and BK expression was more diffuse. Quantitative analysis showedthat in naive terminals, BK clustersaveraged 6.5 per 100 μm² of terminal membrane (Fig. 7A), whereas in terminals isolated from chronic explants, clusters were observed in less than half of the terminals (43%), and when present, averaged only 1 per 100 μm² (Fig. 7A) (p < 0.003). In the remaining 57% of chronic terminals, BK channels (although present in the membrane) were not clustered. Thus, chronic ethanol caused declustering of BK channels in the membrane of neuronal terminals.

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Figure 7.

Chronic ethanol causes declustering of BK channels and reduces density of BK channels in clusters. Confocal images from naive and chronic terminals were used in this study. The number of BK clusters, their size, and the intensity profiles were determined using Leica quantification software (version 2.00; Microsystems). A, Quantification of the number of clusters in naive (n = 12) and chronic (n = 14) terminals shows a significant decrease after chronic ethanol treatment. B, Representative images of a naive and a chronic cluster showing axes used to determine three-dimensional density of BK in clusters. C, Intensity profiles of BK clusters in naive and chronic terminals measured along X, Y, and Z axes as shown in B. D, Quantification of BK density in naive (n = 40) and chronic (n = 8) clusters indicates that chronic ethanol causes an approximately twofold decrease in the BK density per cluster. (*p < 0.05).

Chronic ethanol decreases BK channel density in clusters

Next, we determined whether BK channel density in clusters was altered after chronic ethanol treatment. Using confocal sections (in which clusters appeared oval-shaped), the dimensions of each cluster along the X, Y, and Z axes (Fig. 7B), and the intensity of BK channel expression in each section were determined. The dimensions of clusters in naive terminals (n = 40) (X, Y, and Z measurements) were 367 ± 82, 261 ± 56, and 843 ± 175 nm, respectively, and did not differ significantly from the corresponding values obtained from ethanol-treated terminals (n = 8) (397 ± 55, 234 ± 52, and 834 ± 38 nm). However, although the size of clusters remained the same, the intensity of BK signal in each cluster was decreased along all axes after chronic ethanol treatment (Fig. 7C). Figure 7D illustrates that 24 hr chronic ethanol treatment of HNS explants caused a significant (p < 0.05) decrease in BK channel density in clusters in neuronal terminals.