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Research ArticleArticle

Cannabinoid Receptors Differentially Modulate Potassium A and D Currents in Hippocampal Neurons in Culture

Jian Mu, Shou-yuan Zhuang, M. Todd Kirby, Robert E. Hampson and Sam A. Deadwyler
Journal of Pharmacology and Experimental Therapeutics November 1999, 291 (2) 893-902;

Abstract

Cannabinoid (CB1) receptor activation produced differential effects on voltage-gated outward potassium currents in whole-cell recordings from cultured (7–15 days) rat hippocampal neurons. Voltage-dependent potassium currents A (IA) and D (ID) were isolated from a composite tetraethylammonium-insensitive current (Icomp) by blockade with either 4-aminopyridine (500 μM) or dendrotoxin (2 μM) and subtraction of the residual IA from Icomp to reveal ID. The time constants of inactivation (τ) of IA and ID as determined in this manner were found to be quite different. The CB1 agonist WIN 55,212-2 produced a 15- to 20-mV positive shift in voltage-dependent inactivation of IA and a simultaneous voltage-independent reduction in the amplitude of ID in the same neurons. The EC50 value for the effect of WIN 55,212-2 on IDamplitude (13.9 nM) was slightly lower than the EC50 value for its effect on IA voltage dependence (20.6 nM). Pretreatment with either the CB1 antagonist SR141716A or pertussis toxin completely blocked the differential effects of WIN 55,212-2 on IA and ID, whereas cellular dialysis with guanosine-5′-O-(3-thio)triphosphate mimicked the action of cannabinoids but blocked the action of simultaneously administered cannabinoid receptor ligands. Finally, the differential effects of cannabinoids on IA and ID were both shown to be mediated via the well documented cannabinoid receptor inhibition of adenylyl cyclase and subsequent modulation of cAMP and protein kinase. These actions are considered in terms of cAMP-mediated phosphorylation of separate IA and ID channels and the contribution of each to composite voltage-gated potassium currents in these cells.

Activation of the cannabinoid receptor in cultured neurons inhibits adenylyl cyclase (Little and Martin, 1989; Bidaut-Russell et al., 1990) and enhances an outward potassium current in cultured hippocampal neurons (Deadwyler et al., 1993, 1995a; Hampson et al., 1995). The mechanism of cannabinoid receptor modulation of this potassium current has been previously described (Deadwyler et al., 1995b). This current has been identified as similar to A current (IA) in other reports on the basis of activation and inactivation time constants (5 and 50 ms, respectively), steady-state inactivation and activation voltage ranges (V1/2 = −70 to −75 and −20 to −15, respectively), refractory period (∼200 ms), insensitivity to tetraethylammonium (TEA; 25 mM), and sensitivity to high concentrations of 4-aminopyridine (4-AP; 5 mM; Storm, 1990). Several Gi/o protein-linked receptor systems, such as γ-aminobutyric acidB (Gage, 1992), serotonin1A (Segal, 1980), and adenosine A1 (Mei et al., 1995), enhance IA by producing a positive shift in steady-state inactivation of the channel, resulting in fewer inactivated channels and hence increased current at membrane potentials between −80 and −50 mV. The cannabinoid receptor agonist WIN 55,212-2 produces a 15- to 25-mV positive shift in IA through a similar second messenger cascade (Hampson et al., 1995; Mu et al., 1996) and can be blocked by SR141716A (Mu et al., 1995), a competitive antagonist of the CB1 cannabinoid receptor (Rinaldi-Carmona et al., 1994).

Initial descriptions of the cannabinoid receptor modulation of IA in hippocampal neurons focused on the cannabinoid produced shift in voltage dependence of steady-state inactivation of IA. However, an additional effect on this current was observed in which the inactivation time constant (τ) was also markedly reduced from 50 to 25 ms after cannabinoid exposure (Mu et al., 1997). This effect was observed only with higher concentrations (>300 μM) of 4-AP. Although voltage-dependent outward potassium currents with both 25 and 50 ms τ values have been previously observed (Storm, 1990; Sheng et al., 1993), it is likely that a composite of two different potassium currents (only one of which was IA) with overlapping voltage dependencies really comprised this TEA-insensitive “control” current reported in those studies.

A likely candidate for the second current was potassium D or delay current (ID) on the basis of the inactivation τ, insensitivity to TEA, and high sensitivity to 4-AP (100–500 μM;Storm, 1990; Wu and Barish, 1992; Locke and Nerbonne, 1997a). ID or ID-like currents have been characterized in several different mammalian neurons, including cultured hippocampal neurons (Wu and Barish, 1992; Luthi et al., 1996). It differs from IA in that ID is steady-state inactivated and activated at approximately 30 mV more positive membrane potentials than IA (Storm, 1990; Wu and Barish, 1992). ID is also slower to activate (τ = 20 ms) and inactivate (τ = 100 ms) with a refractory period of up to 20 s (Storm, 1990). In addition, ID is blocked by low concentrations of 4-AP (<1 mM) and by dendrotoxin (DTX; 2 μM), whereas IA is insensitive to DTX and requires much higher concentrations of 4-AP (5 mM) for blockade.

There have been no prior reports of the sensitivity of ID to cannabinoid receptor modulation in any type of central nervous system neuron. In the present experiments, we therefore investigated two main issues: 1) whether the second current contributing to the composite outward potassium current was ID, and 2) if so, whether that current also was modulated by cannabinoid receptor activation.

Materials and Methods

Cell Culture.

The preparation of hippocampal neurons in culture was similar to that described in several previous reports (Deadwyler et al., 1993, 1995a; Hampson et al., 1995). Hippocampi from fetal (E-18) rats (Zivic-Miller) were incubated with neutral protease (2 U/ml Dispase 1; Boehringer Mannheim Biochemica, Mannheim, Germany) for 40 to 50 min at 37°C. After stopping the enzymatic reaction with 1.0 mM NaEDTA, cells were dissociated by gentle trituration via two flame-polished Pasteur glass pipettes and plated at a density of 3 to 4 × 105 cells/35-mm dish. The plating medium consisted of 59% Dulbecco’s modified Eagle’s medium (1×), 19.5% Ham’s F-12 Nutrient Mixture (1×), 10% FBS, 10% horse serum, and 1% l-glutamine (200 mM) (all from GIBCO BRL, Gaithersburg, MD). Cultures were grown at 37°C in a humidified 5% CO2 incubator. After 48 h, half of the medium was replaced by “feeding” medium, which consisted of 98% neurobasal medium, 2% B-27 supplement, 0.25% l-glutamine (200 mM), 0.1% 2-mercaptoethanol (all purchased from GIBCO BRL), and 25 mM KCl. At 72 h after plating, half the medium was again replaced with feeding medium, and cultures were treated with 0.75 μM cytosine-β-d-arabinofuranoside (Sigma Chemical Co., St. Louis, MO) to prevent proliferation of glia. The culture medium was then changed every 3 days for the remainder of the experiment. Experiments were performed on cultured cells between days 7 and 15.

Recording Methods.

The procedure for whole-cell recording was similar to that reported previously (Deadwyler et al., 1993). Briefly, patch electrodes were prepared from 1.5-mm o.d./1.1-mm i.d. borosilicate glass capillaries to produce 1- to 2-μm (2–5 MΩ) tip openings. Electrodes were filled by suction and backfilling with a standard intracellular solution of 140 mM KCl, 11 mM EGTA, 1 mM CaCl2, 2 mM MgCl2, 2 mM ATP, 200 μM GTP, and 20 mM HEPES buffer (Sigma Chemical Co.). Hippocampal cells in primary culture (7–15 days) were washed and constantly perfused with extracellular medium consisting of 140 mM NaCl, 5 mM KCl, 2.5 mM CaCl2, 2 mM MgCl2, 10 mM glucose, and 20 mM HEPES, with 1 μM tetrodotoxin (TTX; Sigma Chemical Co.) added to block voltage-gated sodium channels. Slowly activating/noninactivating potassium currents were blocked by 35 mM TEA, leaving only the transient IA and ID (Storm, 1990; Wu and Barish, 1992). Cultured cells were heated (37°C) and perfused with oxygenated (95% O2/5% CO2) bathing medium throughout the experiment. The osmolality of the bath was 320 ± 10 mOsm, and the pipette solution was 280 ± 10 mOsm. Osmolality was adjusted to prevent cell shrinkage or swelling resulting from dialysis with the recording pipette solution. Cells were covered to a depth of 2 ml with bathing medium and placed in a Leiden microincubator (Medical Systems, Inc., Greenvale, NY) on a Nikon Diaphot inverted microscope. Positive pressure was applied to the recording pipette as it was lowered into the medium and approached the cell membrane. Constant negative pressure was applied to form the seal. A sharp pulse of negative pressure opened the cell membrane for whole-cell recording (Hamill et al., 1981).

Voltage-clamp recordings and command voltage steps were performed with an AxoPatch 1D amplifier and TL-1-LabMaster controller (Axon Instruments, Burlingame, CA) connected to a PC-compatible computer. Whole-cell records were acquired and stored on magnetic disk using pClamp CLAMPEX software (Axon Instruments). Pipette tip junction potentials were continuously monitored and compensated as necessary at the amplifier before breakage of the seal. Access resistance was typically 2 to 5 MΩ, and series resistance compensation was not usually necessary because of low resistance of the pipette. Leakage correction and capacitance compensation (typically 10–30 pF) used dialed-in compensation adjustment at the amplifier, as well as a P/−4 subtraction procedure within the acquisition program. Cells were voltage-clamped and held near resting membrane potential (usually −50 mV). An indication that the cells were adequately space-clamped was that sodium currents in non-TTX-treated cells were not delayed relative to the onset of depolarizing voltage steps, and peak IA in TTX-treated cells did not vary over a range of 5.0 ms with any of the protocols used. A standard steady-state inactivation protocol using a single depolarizing pulse (+50 mV) preceded by series of hyperpolarizing prepulses (−120 to −50 mV) was used to elicit the IA. Activation of IA and ID used a protocol consisting of a multiple depolarizing pulses (−30 to +40 mV) preceded by a single prepulse step of either −120 or −40 mV that either enhanced or inactivated IA, respectively. Because the activation and inactivation voltages for IAand ID overlapped (>−60 mV activation, <−70 mV inactivation for IA, >−70 mV activation, <−40 mV inactivation for ID), it was possible to record IA in the absence of ID by using the IAinactivation protocol and pharmacologically blocking ID with 4-AP or DTX. IDcould be recorded in the absence of IA by using a −40-mV, 50-ms prepulse that steady-state inactivated most IA channels (Storm, 1990; Wu and Barish, 1992). The measurement of current amplitudes and time constants was made with the pClamp software.

Drug Preparation.

The cannabinoid drug was applied via a pressure pipette (10- to 50-μm tip opening) controlled by a solenoid valve (Picospritzer II; General Valve Co., Fairfield, NJ) modified to eject a steady stream of drug-containing media over the surface of the cell. WIN 55,212-2 (Sterling Drug Co., Malvern, PA) was prepared daily from a 10 mM stock solution in ethanol, diluted with extracellular bathing medium, and the ethanol evaporated under a constant stream of nitrogen (Deadwyler et al., 1993). Equivalent bath concentrations corresponding to the pressure pipette concentrations of WIN 55,212-2 are reported in the text. The drug solution was titrated to the same osmolality as the extracellular bathing medium. Due to the lipophilicity of the drug, a 30-s ejection via the application pipette was followed by a washout period of at least 2 min. Previous studies demonstrated that the effects of pressure pipette applications of WIN55,212-2 were rapid (∼10-s onset) and were fully reversed after 2-min perfusion of bathing medium after the application; therefore, all current traces were obtained during this 30-s application period (Deadwyler et al., 1993). No effect was observed on whole-cell currents with the application of vehicle-only solution via the same procedure. Controls consisted of vehicle-only applications to separate neurons, as well as pre- and post-drug measurements of IA and ID within the same WIN 55,212-2-treated neurons. DTX (Sigma Chemical Co.) used to selectively block ID (Storm, 1990; Wu and Barish, 1992) was dissolved directly in bathing medium to a concentration of 2 μM. Controls consisted of recordings in normal medium before perfusion with DTX-containing medium. In experiments designed to inhibit cannabinoid receptor coupling to G proteins, cultured neurons (6–15 days) were pretreated with pertussis toxin (PTX; islet-activating protein; Sigma Chemical Co.; Deadwyler et al., 1993). PTX (10 μg/ml) was added to the culture medium 18 h before recording and to the pipette solution for dialyzation into the cell during recording. Control cells were from the same batch of culture plates with no PTX added. To irreversibly activate Gi/o proteins, 600 μM guanosine-5′-O-(3-thio)triphosphate (GTPγS), a nonhydrolyzable GTP analog, was added to the pipette solution. Controls were conducted with neurons in the same culture plate not exposed to GTPγS. All results were from cells exposed to only one of the above conditions unless otherwise specified. The water-soluble cAMP analog 8-bromo-cAMP (8-br-cAMP; 10 μM) was applied to the cells via the bathing medium. The protein kinase A inhibitors IP-20 (“Walsh” inhibitory peptide, Sigma Chemical Co.) and Rp-cAMPS [(Rp)-diastereomer of cAMP; Sigma Chemical Co.] were dialyzed into the cell through the recording solution. The cannabinoid receptor (CB1-specific) antagonist SR141716A (provided by Sanofi Reserche, Montpelier, France) was prepared as a 1 mM stock solution in ethanol and diluted in bathing medium to 300 to 500 nM, and the ethanol was removed by evaporation under nitrogen.

Analysis.

Time constants (τ) for inactivation of potassium currents were calculated by curve-fitting to the inactivating portion of the outward current trace evoked by a depolarizing voltage step using the CLAMPEX and CLAMPAN patch-clamp acquisition and analysis programs (Axon Instruments). The separation of currents from the composite outward potassium current was also performed using the CLAMPAN program. Measurement of observed changes in voltage dependence of the A and D currents was accomplished by fitting Boltzmann functions to the isolated IA or ID generated by the inactivation and activation protocols (see Deadwyler et al., 1993). Peak current amplitude (Imax) in the inactivation protocol was calculated using a −120-mV hyperpolarizing prepulse, whereas peak activation amplitude was calculated using a depolarizing step to +50 mV. Imax was the same or very near the same for all treatment conditions in a given cell. For each combination of prepulse and step voltage, IA or ID (I/Imax) was converted to relative conductance [G/Gmax; becauseGA = IA/(Vm −EK), with EKdetermined to be approximately −100 mV in cultured hippocampal neurons] and plotted to compare changes in voltage dependence across different drug conditions and different cells (Saint et al., 1990). Boltzmann functions were fitted to mean data points ofG/Gmax and conditioning prepulse voltages (Vpp) via the following formula:G/Gmax = 1/[1 + exp(−(Vpp − V1/2)/k)], where Vpp is the conditioning prepulse potential for inactivation; V1/2 is the voltage at whichG/Gmax = half-maximum conductance, and κ is a slope factor. The coefficients (V1/2 and k) from the above Boltzmann equations were calculated via nonlinear regression (PC-SAS NLIN).

The Boltzmann coefficients were then used to plot a Boltzmann curve over all cells with data points shown as mean ± S.E.G/Gmax. Changes in IA and ID within each protocol were subjected to direct statistical analyses by casting the mean V1/2 values into either two-factor (drug versus voltage step) or three-factor (drug versus voltage step versus concentration) ANOVA designs. Only mean differences in these conductance measures calculated across all cells tested and reaching ap < .01 level of significance (adjusted simple effects contrasts) were considered.

Results

Characterization of Composite Outward Current in Cultured Hippocampal Neurons.

Figure1 (top) shows the nondifferentiated whole-cell current elicited by the voltage step protocol for outward potassium currents in hippocampal cells (Saint et al., 1990; Storm, 1990; Wu and Barish, 1992; Deadwyler et al., 1993). The “composite” current (Icomp) was recorded after the addition of TEA (25 mM) to the external medium, which abolished the noninactivating delayed rectifier, IK, and other noninactivating outward potassium currents elicited by this protocol (Storm, 1990). The residual Icomp displays fast activation time (<5 ms) and an inactivation time constant (“τ”) of 50 ms. The current has a steady-state voltage-dependent activation between −50 and 0 mV and steady-state inactivation between −90 and −50 mV, with a refractory period of approximately 200 ms. Icomp is blocked by high concentrations (>1 mM) of 4-AP (cf. Saint et al., 1990; Wu and Barish, 1992; Deadwyler et al., 1993) and has been extensively characterized in prior studies of cannabinoid receptor modulation (Deadwyler et al., 1993, 1995a,b; Hampson et al., 1995).

Figure 1
Figure 1

Separation and identification of voltage-dependent IA and ID. Total current, whole-cell voltage-gated potassium current elicited by the inactivation protocol shown at the top. The holding potential was −50 mV; prepulses were stepped from −90 to −20 mV in 15-mV increments and then followed by a 375-ms, +50-mV pulse (protocol inset). TTX (0.5 μM) was present in the bath solution for block of voltage-gated sodium current and presynaptic activity. Composite (Icomp), after blocking the noninactivating IK with TEA (25 mM), TEA-insensitive outward potassium current shows time-dependent inactivation. IA, residual inactivating current after the addition of 4-AP (500 μM) or DTX (2 μM) to block ID. The remaining current fits the criteria for IA. ID was calculated by subtraction of the 4-AP- and DTX-insensitive IA from Icomp. Amplitude and inactivation time constant (τ) of subtracted or “recovered” ID are within the same range as determined by direct activation (see Fig. 3). Note differences in inactivation τ and peak current amplitude for IA and ID.

The application of DTX (2 μM) or moderate concentrations of 4-AP (500 μM) reduced the mean τ of Icomp from 50 to 25 ms (Fig. 1, IA). Mean ± S.E. Icomp amplitudes and τ values are given in Table 1 for the indicated number of cells tested. A significant (F1,95 = 12.2,p < .001) reduction in Icomp τ was obtained in all cells exposed to 4-AP or DTX. The remaining current profile of Icomp after 4-AP or DTX exposure meets the characteristics of IA as reported by several investigators (Segal and Barker, 1984; Saint et al., 1990; Deadwyler et al., 1993; Hoffman and Johnston, 1998). The current that was eliminated by 4-AP and DTX was subsequently “recovered” by subtracting the latter IA from the original Icomp curve (Fig. 1, bottom). The recovered current trace has a much slower activation time (∼20 ms) and inactivation τ (100 ms) and meets the characteristics stipulated above for ID (Storm, 1987, 1990; Wu and Barish, 1992; Locke and Nerbonne, 1997b). Thus, Icomp was successfully partitioned into two components: IAand ID.

View this table:
Table 1

Effects of drug treatments on Icomp and ID currents

Effects of Cannabinoids on Icomp.

Figure 2 shows the effect of exposure to the potent cannabinoid WIN 55,212 (40 nM) on Icomp elicited by the above steady-state inactivation protocol (see Materials and Methods). These effects were compared with those of DTX (2 μM) on Icomp using the same inactivation protocol. In both cases, mean τ was significantly reduced from control Icomp levels (Table 1;F1,95 = 17.75, p < .001). However, an effect not mimicked by DTX was for WIN 55,212-2 to increase the amplitude of Icomp (now IA) at all voltage steps (see intermediate traces shown in Fig. 2, bottom left), resulting in a positive shift in the voltage dependence of steady-state inactivation. This is shown graphically by the Boltzmann curves at the right in Fig. 2. As reported elsewhere (Deadwyler et al., 1993, 1995a;Hampson et al., 1995), both steady-state inactivation (circles) and activation (squares) of Icomp were positively shifted by exposure to WIN 55,212-2 (mean Icomp inactivation V1/2 = −71.3 ± 3.1 mV, mean Icomp + WIN 55,212-2 inactivation V1/2 = −54.3 ± 4.4 mV,F1,95 = 12.57, p < .001; mean Icomp activation V1/2 = −20.1 ± 2.6 mV, mean Icomp + WIN 55,212-2 activation V1/2 = −3.5 ± 2.4 mV,F1,95 = 14.2, p < .001). The positive shift in the Boltzmann curves for Icompproduced by WIN 55,212-2 was blocked by simultaneous exposure to SR141716A, the CB1 receptor antagonist (Table 1). As stated above, this shift in the voltage dependence of Icompsteady-state inactivation did not occur after exposure to DTX (cf. Fig.1; mean Icomp + DTX inactivation V1/2= −72.1 ± 4.8; DTX versus WIN 55,212-2,F1,95 = 0.56, p = .45) or to WIN 55,212-2 + DTX (Fig. 2; mean Icomp + WIN 55,212-2 + DTX inactivation V1/2 = −53.2 ± 2.4, F1,95 = .38, p= .54).

Figure 2
Figure 2

Comparison of the effects of DTX and cannabinoid receptor agonist WIN 55,212-2 on Icomp with cannabinoid enhancement of IA. Left, Icomp elicited with steady-state inactivation protocol under control conditions (top) and after exposure to DTX (2 μM, middle) or WIN 55,212-2 (40 nM, bottom). Both DTX and WIN 55,212-2 produced a similar reductions in inactivation τ values (compare with dashed trace for Icomp), leaving a residual current characteristic in both cases of IA. WIN 55,212-2 also produced an increase in peak amplitudes at intermediate voltage steps, indicating a shift in the voltage dependence of steady-state inactivation of IA. Scale bars: 50 ms, 0.5 nA. Right, Boltzmann curves for steady-state inactivation and activation of IA. IAinactivation (● and ○). Peak IA amplitude (Imax) was measured for −120 mV prepulse with depolarizing step to +50 mV. Hyperpolarizing prepulses were then varied in 15-mV steps (positive), and the ratio of IA to Imaxwas computed. The resulting relative current (IA/Imax) was converted to relative conductance (G/Gmax; see text) and fitted by nonlinear regression to the Boltzmann function:G/Gmax = 1/[1 + exp(−(Vpp − V1/2)/k)]. Symbols indicate mean ± S.E.M. conductance (G/Gmax) across different cells (DTX: n = 8; WIN 55,212-2,n = 12). ○, Boltzmann curve for IA(DTX treated). ●, Boltzmann curve and conductance for WIN 55,212-2-treated IA and reflect a 15-mV positive shift in the steady-state inactivation of IA from the DTX-treated condition. IA activation (▪ and ■). Peak IAamplitude (Imax) was measured for −120-mV prepulse and depolarizing step to +50 mV. Depolarizing prepulse was then varied in 10-mV steps (negative); IA/Imax was computed, converted to conductance, and fitted to the Boltzmann equation:G/Gmax = 1/[1 + exp(+(Vpp − V1/2)/k)].

The Boltzmann curves in Fig. 2 suggest that the changes in voltage dependence of Icomp produced by WIN 55,212-2 can be explained by a change in voltage dependence of IA because they are occur in DTX-containing media. However, the reduction in Icomp τ cannot result from effects on IA because IA inactivation τ is not affected by cannabinoid exposure (Deadwyler et al., 1993). The τ reduction in Icomp can be explained appropriately by selective removal of ID from Icomp, leaving only the τ (25 ms) for IA. Thus, cannabinoids produced two independent changes in Icomp, a positive shift in V1/2of IA, which increased peak amplitude of Icomp and reduced τ suggesting decreased contribution of ID.

Effects of Cannabinoids on ID.

Figure3 shows the mean Boltzmann curves for steady-state inactivation and activation of ID recorded in cultured hippocampal neurons (n = 12). The insets show individual currents developed by the respective steady-state inactivation and activation protocols (see Materials and Methods). Because there is no pharmacological means of blocking IA independent of ID and the inactivation protocol includes voltages within a range that will also affect inactivation of IA, the steady-state inactivation curve for ID (Fig. 3, circles) was constructed by subtracting IA from Icomp. To reconstruct ID,Icomp was recorded at each voltage step in the presence and absence of 4-AP (500 μM) to isolate IA. The latter current (IA) was then subtracted from the untreated Icomp to provide the curve for ID. The V1/2 for ID steady-state inactivation was determined to be −39.9 ± 3.5 as shown in Table 1, 30 mV more positive than IA (Deadwyler et al., 1993). The curve for steady-state activation of ID was constructed with an activation protocol and membrane holding potential at −40 mV (Fig. 3, squares). The V1/2 for steady-state activation of ID was +2.6 ± 2.7 mV (Table 1). Either method (subtraction or direct activation) produced comparable current profiles (insets, Fig. 3) respect to both τ and amplitude of ID. The voltage dependence for both steady-state inactivation and activation were consistent with several previous reports of ID characteristics (Storm, 1987, 1990; Wu and Barish, 1992).

Figure 3
Figure 3

Steady-state inactivation and activation of ID. Peak ID amplitude (Imax) was derived and measured using the same inactivation and activation protocols as in Figs. 1 and 2. IA was pharmacologically separated from Icomp in the steady-state inactivation protocol and then subtracted from Icomp to produce ID. IA (−40-mV prepulse) was eliminated in the activation protocol (−50-mV prepulse, depolarizing step to +50 mV). For each combination of prepulse (inactivation) or step voltage (activation), Boltzmann curves for steady-state inactivation and activation were fitted as in Fig. 2. Points are mean ± S.E. conductance elicited for each voltage (n = 21 cells). Inset traces, isolated ID currents elicited by the inactivation and activation protocols. ×, on the Boltzmann curves, single-cell conductance measurements obtained from the traces shown. Scale bars: 50 ms, 0.5 nA.

The activation protocol was therefore used to determine the effects of WIN 55,212-2 (10 and 40 nM) and DTX (2 μM) on ID. The traces in Fig.4 show the effects of WIN 55,212-2 (10 and 40 nM) on ID recorded in TEA. The effects of DTX (2 μM) are shown below for comparison. Exposure to WIN 55,212-2 (40 nM) reduced the peak ID amplitude by 60% (Table 1, n = 12 cells, mean IDamplitude = 0.64 ± 0.09, mean ID + WIN 55,212-2 amplitude = 0.25 ± 0.03,F1,95 = 16.53, p < .001) relative to the TEA control amplitude. The addition of DTX either alone or in combination with WIN 55,212-2 also produced a 70% reduction in ID amplitude relative to control (mean ID + DTX amplitude = 0.22 ± 0.05 nA, mean ID + WIN 55,212-2 + DTX amplitude = 0.21 ± 0.03; all F1,95 > 7.01,p < .01). The reduction of IDamplitude by WIN 55,212-2 was blocked by the cannabinoid receptor antagonist SR141716A (Table 1).

Figure 4
Figure 4

Cannabinoid effects on ID. Left, WIN 55,212-2 effects on ID activation (steady-state activation protocol). TEA, activation of TEA-insensitive ID in normal media with 25 mM TEA added. Steady-state inactivation of IAis effected by a step to −40 mV before depolarizing steps to 0 to +50 mV (inset, ID activation protocol). WIN 55,212-2, exposure to 10 and 40 nM WIN 55,212-2 (in TEA) produces a dose-dependent decrease in ID amplitude with no change in τ. DTX, exposure to DTX (2 μM) produced a slightly greater reduction in ID. Scale bar: 50 ms, 0.5 nA. Right, Boltzmann curves for steady-state inactivation and activation of ID. Exposure to WIN 55–212-2 (solid lines) produced an apparent shift in voltage dependence of inactivation and activation compared with control (dashed line); however, the shifts were accompanied by symmetric decreases in slope for activation and inactivation (see text).

The reduction of ID amplitude by WIN 55,212-2 was not accompanied by a shift in voltage dependence of steady-state inactivation of ID. This is suggested by the consistency of the relative amplitudes at intermediate voltage steps in the traces in Fig. 4 (left), which show a decrease in the ID amplitude for each trace. This voltage-independent decrease in amplitude is reflected by the altered slope of the Boltzmann curves in Fig. 4 after exposure to WIN 55,212-2 (10 nM, filled symbols; 40 nM, open symbols). The change in slope was responsible for the apparent shift in V1/2 for the same Boltzmann curves and therefore did not reflect a change in voltage dependence of ID. Figure 4 shows that the effect of cannabinoids on ID peak amplitude was also dose dependent but not as effective as DTX (see below).

Relative Potency and Efficacy of Cannabinoids on IA and ID.

Figure 5shows the concentration-effect (2.5–40 nM) curves for the WIN 55,212-2 modulation of ID amplitude (filled circles) and shift in V1/2 for steady-state inactivation of IA(open squares). A comparison of the EC50 values for each effect revealed that the cannabinoid reduction in mean IDamplitude occurred at lower concentrations (13.9 ± 1.6 nM) than the mean V1/2 shift in IA voltage dependence (20.6 ± 1.9 nM; T14 = 2.69,p < .01). Thus, ID was 34% more sensitive to the influence of cannabinoids than IA, a further indication that cannabinoid alteration of Icomp was produced by the above demonstrated independent effects on IA and ID.

Figure 5
Figure 5

Concentration-dependent effects of WIN 55,212-2 on IA and ID (range, 2.5–100 nM). Dose-effect curves for WIN 55,212-2 effect on reduction of ID amplitude (●). Half-maximal effect (EC50) was 13.9 nM, and the maximal effect was obtained at 70 nM (not shown). The dashed line shows the effects of the same concentrations of WIN 55,212-2 on the shift in voltage dependence (V1/2) for steady-state inactivation of IA (■). The EC50 value for cannabinoid effects on IA was 20.6 nM, and the maximal effect was obtained at 80 nM (at least 4 cells/point; range, 4–21).

Cannabinoid Effects on IA and ID Are Mediated by Gi/o Proteins, cAMP, and Protein Kinase A (PKA).

It has been firmly established that cannabinoid receptor inhibition of adenylyl cyclase is mediated through PTX-sensitive G proteins (Howlett et al., 1986; Bidaut-Russell et al., 1990; Pacheco et al., 1991). Consistent with this effect of cannabinoid receptor activation, previous reports demonstrated that the effects of cannabinoids on IA were G protein (Deadwyler et al., 1993) and cAMP dependent (Deadwyler et al., 1995a). In those studies, increased levels of cAMP (8-br-cAMP), PKA, protein kinase inhibitors (IP-20 and Rp-cAMPS), and WIN 55,212-2 were all used (Deadwyler et al., 1993, 1995a; Hampson et al., 1995). The results of similar treatments with respect to effects on IA were replicated in the present study and are summarized in Table 1 and Fig.6, A and B. In general, manipulations that increased cAMP levels produced a negative shift in the voltage dependence of IA but not the slope of the steady-state activation and inactivation curves (Deadwyler et al., 1995a; Hampson et al., 1995). Treatments that decreased cAMP levels or blocked PKA produced a positive shift in IA activation and inactivation with no change in slope of the Boltzmann curves (Fig. 6, A and B). Because the effects on PKA presumably altered phosphorylation of membrane channel proteins, the catalytic subunit of PKA (Cat.S.; Table1) and the phosphatase inhibitor okadaic acid (OA; Table 1) were also tested. Both Cat.S. and OA induced a negative shift in IAvoltage dependence that was similar to increased cAMP levels (8-br-cAMP). Table 1 shows that the effects of WIN 55,212-2 on IA and Icomp were similar in direction and magnitude to decreased cAMP levels or PKA inhibition (especially IP-20) and opposite in effect to increased cAMP levels and enhanced PKA-dependent phosphorylation by other agents. These results indicate that the differential shifts in IA voltage dependence were regulated by the phosphorylation status of the IA channel protein. Furthermore, because exposure to 8-br-cAMP, Cat.S., or OA blocked cannabinoid effects on IA (Table 1) and IP-20 or Rp-cAMPS plus WIN 55,212-2 were not additive, it is therefore likely that the same cAMP/PKA cascade was involved for all of the above influences on IA.

Figure 6
Figure 6

cAMP/PKA modulation of IA and ID steady-state inactivation and activation. A, Boltzmann curves (see Fig. 2) for steady-state inactivation of IA(see Protocol inset) calculated for currents recorded in 8-br-cAMP (10 μM), WIN 55,212-2 (WIN 55,212-2; 40 nM), or DTX (2 μM) in the bathing solution or after dialyzing the cell, via the recording pipette solution, with the PKA inhibitory peptide IP-20 (6 μM) or partial PKA inhibitor Rp-cAMPS (2 μM). Symbols reflect mean ± S.E. conductance across all cells for each prepulse voltage (see Table 1 for V1/2 and n values). B, activation of IA. Treatment conditions were the same as above for Inactivation. Symbols reflect mean ± S.E. conductance across cells for each depolarizing step voltage shown in Protocol inset. Inset, current traces show change in peak IA (activation) after exposure to IP-20 and 8-br-cAMP and under control conditions. C, ID inactivation. Boltzmann curves for steady-state inactivation of ID (by subtraction of IA from Icomp) were calculated by subtraction of IAfrom Icomp (Fig. 2, see also Table 1 for V1/2 and n values). D, Boltzmann curves for activation of ID. Treatment conditions and symbols are the same as in B. Inset, current traces elicited by the IDsteady-state activation protocol for 8-br-cAMP, IP-20, and control. Scale bar: 50 ms, 0.5 nA

Table 1 also indicates similar tests of G protein and cAMP involvement conducted on activation and inactivation of ID. The reduction by WIN 55,212-2 of ID amplitude was blocked by PTX (mean PTX amplitude = 0.60 ± 0.05 nA; mean PTX + WIN 55,212-2 amplitude = 0.59 ± 0.08 nA;F1,95 = 0.19, p = .65), but importantly, PTX did not alter the ability of DTX to reduce ID amplitude (mean PTX + DTX amplitude = 0.24 ± 0.04 nA, F1,95 = 6.47;p < .01, Table 1). Cells (n = 8) dialyzed with the G protein activator GTPγS (600 μM) also exhibited reduced ID amplitude (mean GTPγS amplitude = 0.30 ± 0.08; F1,95 = 6.11,p < .01) while blocking the effect of WIN 55,212-2 (mean GTPγS + WIN 55,212-2 amplitude = 0.31 ± 0.07 nA,F1,95 = 0.18, p = .67, Table 1).

Because cannabinoid receptor effects on ID were shown to be dependent on linkage to Gi/oproteins, further tests of dependence on the cAMP/PKA cascade were performed. Figure 6D (inset) shows that exposure to 8-br-cAMP (10 μM) significantly increased ID amplitude by 25% (mean 8-br-cAMP amplitude = 0.81 ± 0.08 nA, mean control amplitude = 0.64 ± 0.09 nA,F1,95 = 6.29, p < .01, Table 1), whereas IP-20, markedly reduced IDamplitude by 63% (mean IP-20 amplitude = 0.24 ± 0.09 nA,F1,95 = 6.51, p < .01). Boltzmann curves for steady-state inactivation and activation of ID are shown in Fig. 6, C and D, for the same treatment conditions shown in Fig. 6, A and B. Several differences are immediately apparent. First, the effects of 8-br-cAMP on ID were not equivalent for steady-state inactivation versus activation of ID. The shift in voltage dependence in ID produced by 8-br-cAMP was unchanged for steady-state inactivation of IDrelative to control conditions (Fig. 6C) but shifted significantly negative for steady-state activation of ID (Fig.6D, squares). Second, PKA reduction (Rp-cAMPS) produced a small but significant negative shift in steady-state inactivation of ID (Fig. 6C). In all other instances, a change in slope of the Boltzmann curve for either steady-state inactivation and/or activation of ID was obtained (Fig. 6, C and D).

The change in slope in the ID Boltzmann curves in Fig. 6, C and D, indicates a change in the kinetics of the ID channel. One possible source of that change is a shift in sensitivity of voltage dependence “outside” the range of the protocol used, which would have reduced maximum current (Imax; see Materials and Methods) capable of being evoked by the protocol. If so, different Imax values would be obtained depending on the range of voltages used in the protocols. Alternatively, the change in slope of the Boltzmann curves may have resulted from total inactivation (lack of conductance) of a subpopulation of IDchannels (Hille, 1992), also resulting in a decrease in Imax. In the latter case, Imax would be relatively unaffected by changes in voltage range. Figure 7 shows the maximum (i.e., peak) ID amplitude recorded using the same activation protocol at a higher voltage (+80 mV depolarization) but preceded by one of three different levels of hyperpolarizing prepulses (−120, −80, or −40 mV) to maximally activate ID. The three conditions provided a greater range of depolarization to assess whether maximum IDamplitudes were differentially altered by the indicated cAMP/PKA treatments. The bar graph in Fig. 7 shows no significant difference in current amplitude recorded as a function of the three protocols; this indicates that the reduction in ID, and hence the change in slope of the Boltzmann curves in Fig. 6, C and D, produced by cAMP/PKA inhibitors, did not result from a shift in voltage dependence but rather a decrease in the maximum current that could be evoked regardless of voltage. Thus, in contrast to effects on IA, manipulations of the cAMP/PKA cascade and consequent phosphorylation status of channel proteins altered the conductance or availability of ID channels and not the voltage dependence of those channels.

Figure 7
Figure 7

Maximum ID activation. ID was recorded using three combinations of prepulse (holding) voltage (−120, −80, and −40 mV) with a +80-mV depolarizing voltage in the protocol indicated. Voltages were selected to exceed the range used to elicit maximum currents in Fig. 6, C and D, to test whether the slope changes resulted from a shift in voltage dependence or decreased maximal conductance. Peak current was not significantly different for each protocol (Control) but was greatly reduced after exposure to PKA inhibitors, WIN 55,212-2 and DTX. Rp-cAMPS, thio-cAMP stereoisomer, WIN 55,212-2, cannabinoid agonist WIN 55,212-2; and IP-20, PKA inhibitory peptide (20 amino acid).

Contributions of IA and ID to Icomp.

Figure8 shows the net changes in Icomp with different manipulations of cAMP. The solid traces show Icomp recorded from a single neuron under control conditions (i.e., “resting” levels of cAMP) and after exposure to 8-br-cAMP (10 μM) or the cannabinoid agonist WIN 55,212-2 (40 nM). The horizontal lines at the top in Fig. 8 show that increased cAMP levels (8-br-cAMP) reduced the peak amplitude of Icompby 16% relative to control (long vertical arrow); in contrast, cannabinoid exposure increased peak Icomp amplitude by only 4% (short vertical arrow). However, at the same time, increased cAMP levels (8-br-AMP) caused a 27% increase in Icomp τ relative to control, whereas cannabinoids decreased the Icomp τ by 58%, almost twice the change produced by cAMP. The dashed line in Fig. 8 (8-br-cAMP and Cannabinoid) indicates the degree of change in time course of Icomp under both conditions as indicated by the direction of the arrow. Thus, the maximum range in Icomp τ from increased cAMP levels (Fig.8, 8-br-cAMP) to decreased cAMP levels produced by cannabinoid receptor activation (Fig. 8, Cannabinoid) was 68%. The dotted traces in Fig. 8depict IA (8-br-cAMP and Control) and show that Icomp amplitude was reduced in conjunction with IA amplitude after exposure to increased levels of cAMP. Conversely, the dash-dotted traces (Control and Cannabinoid) show the reciprocal change in amplitude and τ of ID as a function of decreased cAMP levels produced by cannabinoid exposure. In each case, it is clear that an increase in cAMP produced an increase in the duration of Icomp but at the expense of a marked reduction in amplitude. Decreasing cAMP levels via cannabinoid receptor activation resulted in a marked decrease in the duration of Icomp, with a relatively insignificant increase in amplitude. This dual modulation of Icomp shown in Fig. 8 is cAMP dependent; however, the differential modulation of IAand ID must occur “downstream” at the level of PKA-dependent phosphorylation, as indicated by the differential effects of IP-20 and Cat.S. on both currents (see Table 1).

Figure 8
Figure 8

Effects of differential modulation of IAand ID by cAMP on Icomp. The three sets of single-cell voltage-clamp current traces show Icomp under control (center), 8-br-cAMP (10 μm, left), and WIN 55,212-2 (40 nM, right) recording conditions. Solid current traces indicate Icomp recorded under each of the three conditions. Dashed current traces at left (8-br-cAMP) and right (WIN 55,212-2 ) are superimposed control Icomp (center) traces. Bold arrows show differential change in τ from control Icomp. Horizontal lines and arrow at top depict change in Icompamplitude (right to left) produced by increasing levels of cAMP. Dotted traces (middle and left) depict decrease in IA as cAMP levels increase (8-br-cAMP). Dash-dotted traces (middle and right) show decrease from maximum ID (8-br-cAMP), through control (middle), to minimum ID after exposure to WIN 55,212-2 (right).

Discussion

The above results clearly demonstrate that a major source of voltage-dependent potassium current (Icomp) in hippocampal cells is selectively modulated by cannabinoid receptor occupancy. IA and ID, which contribute to Icomp in cultured hippocampal neurons, are voltage-dependent outward currents with overlapping activation ranges; both are differentially sensitive to 4-AP, and both are TEA resistant. Because the two currents, IAand ID, inactivate at different τ values (IA ≈ 25 ms; ID ≈ 100 ms), have different voltage dependencies for steady-state inactivation (V1/2: IA ≈ −70 mV; ID ≈ −40 mV), and can be totally differentiated pharmacologically with DTX (2 μM) or low-to-moderate concentrations of 4-AP (500 μM), the cannabinoid receptor modulation of each current was examined independent of the other.

It is clear from the above data that the previously reported effects of cannabinoids (Deadwyler et al., 1993, 1995a; Hampson et al., 1995) on IA in cultured hippocampal neurons can now be identified with respect to the contributions of IA and ID to Icomp (Figs. 2, 6, and 8). Hence the positive shift in voltage dependence of Icomp resulted from cannabinoid-mediated enhancement of IA, whereas the reduction in τ of Icomp was derived from decreased ID amplitude. The addition of the selective channel blocker DTX produced a reduction in the Icomp τ similar to WIN 55,212-2 (Figs. 1 and 2) and blocked any further actions of WIN 55,212-2 (Table 1). Exposure to WIN 55,212-2 also produced a reduction in IDamplitude in both the steady-state inactivation and activation protocols (Fig. 4). DTX, however, did not provoke a change in the voltage dependence of IA (Fig. 2). Further evidence that both the positive shift in IAvoltage dependence and the reduction in IDamplitude were cannabinoid receptor mediated was provided by blockade by the CB1 receptor antagonist SR141716A (Table1). Although a potential alternative interpretation for the dual effects of cannabinoids on Icomp is that the shift in voltage dependence of IA (Fig. 2) could have resulted directly from the elimination or marked reduction of ID (Fig. 4), the removal of a current with a more positive Boltzmann curve (V1/2) than IA (as was the case for ID; see Fig. 6) would have produced a negative (not a positive) shift in the voltage dependence of Icomp (Fig. 2). The predicted negative shift would result only if both currents were activated with the same time constant (cf. Fig. 6). The slower activation time constant for ID (20 ms) requires that the initial peak of Icomp (5-ms duration) consists almost entirely of IA; thus, the positive shift in Icomp voltage dependence was strictly the result of the cannabinoid receptor activation on IA, not ID. Similarly, the reduction in Icomp τ resulted from direct cannabinoid effects on ID amplitude because IA inactivation τ was not altered by cannabinoids (Table 1).

Prior reports have established that cannabinoid receptor modulation of IA voltage dependence can be attributed ultimately to PKA-dependent phosphorylation (Table 1; Hampson et al., 1995; Mu et al., 1996). Several manipulations of the cAMP/PKA cascade, including direct application of Cat.S (Table 1), confirmed that the reduction in ID amplitude produced by cannabinoid receptor activation was consistent with a decrease in PKA dependent phosphorylation (presumably of ID channel proteins) in these cells (Fig. 6, Table 1). Figure 8 further demonstrates that the effect of increased cAMP on ID steady-state inactivation was reciprocal to modulation by the cannabinoid receptor of IA. The fact that the control (untreated) measures of IDamplitude and IA V1/2 were between these two extremes shows that a tonic level of cAMP is normally present and active, allowing both IA and ID to be only partially “expressed” under normal, control conditions (Control trace in Fig. 1). Because there was a change in slope in the Boltzmann curves for ID, it was not possible to determine whether the voltage dependence of either activation or inactivation of ID was altered by cannabinoids and other agents that modulate cAMP levels (Fig. 6). However, there was a cAMP-dependent decrease in ID amplitude, presumably due to total blockade of the channel as occurs with DTX (Storm, 1990; Wu and Barish, 1992). The fact that the EC50 value was lower for cannabinoid reduction in ID amplitude than for shifting the voltage dependence of IA suggests a slight bias toward decreasing the τ of Icomp.

Mechanism of Cannabinoid Modulation of IA and ID.

The above results provide evidence that a reciprocal relationship exists between IA and ID with respect to cannabinoid receptor modulation of levels of cAMP. Modulation of voltage-gated potassium channel currents (Icomp) can play a major role in altering the temporal and amplitude characteristics of action potentials in hippocampal neurons (Segal et al., 1984). Such reciprocity is likely explained by differences in the subtypes of K+ channels producing IA and ID. If the two currents were produced by different subtypes (i.e., shaker-type Kv1 versusshal-type Kv4 channels or even Kv4.2 versus Kv4.3 channels), then the common factor of cannabinoid receptor-mediated inhibition of adenylyl cyclase, PKA, and subsequent protein phosphorylation could be translated into opposite actions on IA and ID. The IA channel described in these studies fits the description of the Kv4.2 or Kv4.3 subtypes, which have been shown to be present in hippocampal pyramidal cells and interneurons (Villarroel and Schwarz, 1996; Serodio and Rudy, 1998). Although the profile of IA is also satisfied by Kv1.4, this can be ruled out on the basis of a lack of effect of H2O2 on either IA or Icomp (Mu et al., 1997). Different inactivation mechanisms have been demonstrated for different types of potassium channels (Levitan, 1994). When phosphorylated, the Kv4.2- and Kv4.3-type homomultimers have low conductances that change drastically toward maximum under conditions of dephosphorylation (Aiello et al., 1995). Thus, for this putative IA channel, cannabinoid receptor-mediated decreases in PKA phosphorylation would lead to a positive shift in the inactivation voltage of the IAchannel (Deadwyler et al., 1993; Hampson et al., 1995).

The same cannabinoid-induced decrease in cAMP and consequent decrease in PKA dependent phosphorylation were also associated with a decreased amplitude of ID (Figs. 4, 6, and 7). Decreased phosphorylation in Kv3.3- or Kv3.4-type K+channels (possible candidates for ID) results in a decrease in channel conductance (Massengill et al., 1997). Because amplitude reduction in ID may occur without a shift in voltage dependence, the phosphorylation site on the ID channel may not be associated with an inactivation or activation process as it is in IA(Fig. 6). Thus, in the ID-type channel, cAMP-dependent reduction in current amplitude may reflect a more complex interaction, involving multiple phosphorylation sites (Fadool and Levitan, 1998). Such a mechanism could be responsible for the altered slope in the Boltzmann curves in Fig. 6, showing that peak conductance of ID at all voltage levels was markedly reduced by decreases in cAMP and/or PKA inhibition. Hence, the effects of cannabinoid drugs on IA and ID are coupled through the cannabinoid receptor/cAMP cascade. Receptor occupancy thus appears to have opposite functional consequences, presumably due to different conductance states of the two channels with respect to PKA-dependent phosphorylation (Levitan, 1994).

The reciprocal nature of cannabinoid receptor-meditated effects on IA and ID provides insight into prior reports that did not entirely distinguish among Icomp, IA, and ID in central nervous system neurons (Brew and Forsythe, 1995; Zhang and McBain, 1995; Keros and McBain, 1997). IA and ID were distinguished on the basis of changes in synaptic and action potential profiles as a function of sensitivity to low versus high concentrations of 4-AP (Storm, 1987, 1990; Hamon et al., 1995; Inokuchi et al., 1997). Low concentrations of 4-AP (30–500 μM) enhanced excitatory postsynaptic potentials (Southan and Owen, 1997), increased action potential duration (Storm, 1987, 1987), and decreased interspike intervals (Brew and Forsythe, 1995; Locke and Nerbonne, 1997a), presumably as the result of a selective blockade of ID (Fig. 8, Cannabinoid, ID). High concentrations of 4-AP were shown to enhance these effects as would be predicted by eliminating the residual Icomp shown in Fig. 8 (Cannabinoid, solid line), which is mostly IA. In contrast, cannabinoids reduce ID amplitude while simultaneously producing a positive voltage shift in inactivation and activation of IA (Figs. 2, 6, and 8), thereby counteracting the spike-broadening effects (Storm, 1987; Locke and Nerbonne, 1997a) without influencing the decrease in interspike interval produced by blockade of ID (Locke and Nerbonne, 1997a). Therefore, the positive voltage shift in IAproduced by cannabinoid receptor inhibition of adenylyl cyclase and subsequent blockade of the cAMP/PKA cascade would increase the number of action potentials resulting from the decrease in Icomp τ (Fig. 8, Cannabinoid, ID) while limiting calcium influx during the action potential (Storm, 1987; Locke and Nerbonne, 1997a). These results suggest the role of cannabinoids is to “fine-tune” the activity and excitability of neurons through the modulation of voltage-dependent potassium channels.

Footnotes

  • Send reprint requests to: Dr. Sam A. Deadwyler, Dept. of Physiology and Pharmacology, Wake Forest University School of Medicine, Winston-Salem, NC 27157-1983. E-mail:sdeadwyl{at}wfubmc.edu

  • ↵1 This work was supported by National Institute on Drug Abuse Grants DA03502, DA07625, and DA00119 to S.A.D.

  • Abbreviations:
    IA
    voltage-sensitive potassium A current
    ID
    voltage-sensitive potassium D current
    Icomp
    composite tetraethylammonium-insensitive voltage-sensitive potassium current
    GTPγS
    guanosine-5′-O-(3-thio)triphosphate
    PTX
    pertussis toxin
    PKA
    protein kinase A
    TTX
    tetrodotoxin
    8-br-cAMP
    8-bromo-cAMP
    TEA
    tetraethylammonium
    Rp-cAMPS
    (Rp)-diastereomer of cAMP
    Cat.S.
    catalytic subunit of PKA
    OA
    okadaic acid
    4-AP
    4-aminopyridine
    DTX
    dendrotoxin
    V1/2
    half-inactivation voltage of steady-state inactivation (of IA and ID)
    τ
    time constant of inactivation of voltage-sensitive potassium current
    • Received April 22, 1999.
    • Accepted July 21, 1999.

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Cannabinoid Receptors Differentially Modulate Potassium A and D Currents in Hippocampal Neurons in Culture

Jian Mu, Shou-yuan Zhuang, M. Todd Kirby, Robert E. Hampson and Sam A. Deadwyler
Journal of Pharmacology and Experimental Therapeutics November 1, 1999, 291 (2) 893-902;
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