We obtained whole-cell current-clamp recordings from CA1 pyramidal neurons in rat hippocampal slices. The effects of the group I mGluR activation on responses to intracellular current injection were examined following bath application of the group I mGluR agonist DHPG (2–4 µM, see Materials and Methods). Step current injections (0.6-s long) that were just above threshold for action potential firing in control elicited increased action potential firing in the presence of DHPG (Figure 1A). In response to longer current injections (4.5 s), DHPG converted a simple pattern of spike-frequency accommodation to a more complex pattern consisting of a high-frequency burst of action potentials, followed by a period of silence, and finally a continuous train exhibiting spike-frequency accommodation. In response to noisy current injections, the number of action potentials was also increased when DHPG was applied (Figure 1B and 1C). DHPG produced a small but statistically significant increase in the input resistance (54.7±3.0 MΩ for control, t = 0 min; 59.7±4.1 MΩ for DHPG, t = 15 min; n = 26, paired t test, p<0.01). This did not noticeably change the subthreshold response to noisy current injection, but enhanced spiking appeared to be attributable to a reduction in the post-spike AHP in the presence of DHPG (Figure 1D).

The effect of DHPG on the post-spike AHP was studied systematically by examining responses to bursts of action potentials evoked by five brief (2 nA, 2 ms) current injections. In normal artificial cerebrospinal fluid (ACSF), a 100 Hz burst of five spikes was followed by an AHP (−3.1±0.2 mV) that reached a peak at 59±5 ms after the last spike. Application of DHPG (or quisqualate, another group I mGluR agonist, Figure S1) eliminated the AHP, resulting in a post-burst ADP (+18.7±0.7 mV) that reached a peak at 34±3 ms after the last spike and decayed to 25% of the peak value in 200±23 ms (n = 20, Figure 1E). We refer to this ADP as a medium ADP to distinguish it from the fast ADP following a single spike in normal ACSF [19]–[23] and a slow DHPG-induced ADP described later in this article. Functionally, the change from post-burst AHP into a post-burst medium ADP made the pyramidal neurons more excitable; current injections that were subthreshold during the post-burst AHP evoked action potential firing during the medium ADP (Figure 1F).

Application of DHPG resulted in a gradual reduction of the AHP and conversion to an ADP. The medium ADP reached its maximum value after several minutes in DHPG and was fully reversible, with a similar time course, upon washout of DHPG (Figure S2); however, the slow onset and reversal of the medium ADP is attributable to the slow speed of the perfusion system, as rapid application of DHPG produced more rapid responses (see below). In all experiments, the amplitude of the post-burst potential (AHP or medium ADP) was quantified at a fixed time, corresponding to the peak of the AHP in normal ACSF (59±5 ms after the last spike). For some analyses, the effect of DHPG was quantified by the change in the post-burst potential (Δ post-burst potential) at this time point in the response (see Figure S3). This measure required comparison of the response in normal ACSF (AHP at t = 0 min) to the response at a fixed time after application of DHPG (e.g., ADP at t = 15 min). Importantly, this measure was not complicated by any slow, drug-independent effects, as the post-burst AHP was stable for typical recording duration in normal ACSF (Figure S3).

To determine which group I mGluR subtype was responsible for the DHPG-induced medium ADP, we used the subtype-selective antagonists LY367385 (25 µM, mGluR1 antagonist) and MPEP (10 µM, mGluR5 antagonist). LY367385 blocked the medium ADP (t = 15 min) by 16%, while MPEP blocked it by 73% (Figure 2A), suggesting that the effect of DHPG is mediated primarily by mGluR5 and partially by mGluR1. When applied together, the two drugs blocked the medium ADP by 88% (Figure 2A), suggesting that the effects of mGluR1 and mGluR5 are approximately additive. To determine the subcellular localization of the mGluRs mediating the medium ADP, DHPG was applied locally, by pressure application from a large patch pipette, to either the perisomatic region or the apical dendrites (see Materials and Methods for details). Application of DHPG (500 µM in application pipette) to the apical dendrites had no effect, while perisomatic application produced a medium ADP (Figure 2B). Similarly, when DHPG was present in the bath, application of normal ACSF reduced the medium ADP when applied to the soma, but not when applied to the apical dendrites (Figure 2C). The effects of DHPG were induced within seconds of its application and reversed rapidly when the DHPG application was terminated. Together, these results suggest that perisomatic mGluRs must be activated to elicit an medium ADP in response to somatic action potential firing and that the effect is rapidly induced and reversed. The lack of effect with application to the dendrites does not necessarily imply the absence of group I mGluRs, as backpropagating action potentials could have a different effect from action potentials in the soma (see Discussion).

To determine whether group I mGluR-mediated modulation of the post-burst potential could occur in response to synaptically released glutamate, the post-burst AHP/ADP was compared during control conditions and during high-frequency (50 Hz) activation of Schaffer collaterals. Fast synaptic responses were prevented by blocking glutamate and GABA receptors (see Materials and Methods). Under control conditions, each burst was followed by an AHP. During synaptic stimulation, however, each burst was followed instead by an ADP (Figure 3A). To determine the role of group I mGluR activation in the induction of the post-burst ADP, we applied blockers of mGluR1 and mGluR5 (Figure 3B and 3C). The results of these experiments were compared to a separate group of control experiments occurring over the same time course but without blocker application. In the absence of synaptic stimulation, the post-burst AHP was stable over time, both in control and in the presence of group I mGluR blockers (Figure S4). In the presence of synaptic stimulation, the size of the ADP in the control group gradually increased over the course of the experiment (156%±12% of initial value, n = 5). By contrast, blocking the group I mGluRs resulted in a decrease in the size of the ADP (66%±7% of initial value, n = 5). The magnitude of the ADP at the end of the experiment (60 min) in the presence of group I mGluR blockers was 42% of control, consistent with a substantial contribution of these receptors to induction of the ADP triggered by synaptically released glutamate. The long-term effects of synaptic stimulation on the ADP are interesting but are not considered here. Instead, our focus is on the acute modulation of the AHP/ADP during activation of group I mGluRs.

Bath application of DHPG increased the fast ADP following a single spike, and the size of the medium ADP increased with both the number and the frequency of action potentials, reaching medium ADP values of nearly 15 mV for 5 spikes at 100 Hz (Figure S3). Longer trains (20 or 50 spikes) did not increase the ADP further and in fact resulted in a decrease in the size of the ADP (Figure S5), perhaps due to inactivation of the ADP-producing current or enhanced activation of an AHP-producing current.

The medium ADP following a burst of spikes evoked by a step current injection was blocked by application of tetrodotoxin (TTX; 0.5 µM) to eliminate Na⁺-mediated spikes; however, increasing the magnitude of the current injection to elicit a Ca²⁺ spike [24] restored the medium ADP (Figure 4A and 4B). Under these conditions, the amplitude of the medium ADP increased with the magnitude and duration of the current injection, reaching a maximum of about 10–15 mV for current injections of at least 1.4 nA for 40 ms (Figure 4C and 4D).

The requirement for action potential firing or a Ca²⁺ spike suggests that the medium ADP may require Ca²⁺ entry through voltage-gated calcium channels (VGCCs). Consistent with this idea, we found that the medium ADP was eliminated by switching to a Ca²⁺-free ACSF (Figure 5A) or by bath application of micromolar concentrations of NiCl₂ (Figure 5B–E; IC₅₀ = 23 µM). Nimodipine (10 µM, an L-type calcium channel blocker) did not block the medium ADP (Figure 5B–D). To test whether the medium ADP required elevation of internal Ca²⁺ concentration, we performed experiments with patch-clamp electrodes containing BAPTA (10 mM) and found that this strongly reduced the medium ADP (Figure S6). Blocking Ca²⁺ release from internal stores with cyclopiazonic acid (CPA; 20 µM) also reduced the medium ADP (Figure S6), suggesting that this is another important mechanism for induction of the medium ADP.

The requirement for Ca²⁺ entry through VGCCs and the elevation of internal Ca²⁺ concentration is consistent with two different models of the medium ADP. In the first model, the medium ADP is mediated by Ni²⁺-sensitive VGCCs, with their modulation (enhanced activity) by DHPG requiring elevated intracellular Ca²⁺. In the second model, Ca²⁺ entry through VGCCs contributes little to the medium ADP directly but acts as a trigger for a downstream conductance that is modulated by DHPG. For example, Ca²⁺ entry and release from internal stores could activate nonselective cation currents, such as I_CAN, which mediate the medium ADP. Alternatively (or in addition), downregulation of Ca²⁺-activated K⁺ channels by DHPG could unveil the medium ADP. To distinguish between these possibilities, we examined the voltage dependence of the medium ADP. We reasoned that in the first model, where the medium ADP is mediated by VGCCs directly, hyperpolarization should accelerate deactivation of the VGCCs, thus reducing the medium ADP. In the second model, hyperpolarization would not eliminate Ca²⁺ entry during the action potentials, and it would increase the driving force on the cation channels, thus increasing the medium ADP. We found that holding the cell at a hyperpolarized holding potential strongly reduced the amplitude and duration of the medium ADP (Figure 6A and 6B), a finding most consistent with the first model, in which the medium ADP is mediated by VGCCs directly. However, the slowest component of the medium ADP was not reduced by hyperpolarization (Figure 6A), suggesting a contribution of Ca²⁺-activated channels to a slow ADP (see below).

To further examine the voltage dependence of the medium ADP, we delivered short hyperpolarizing current injections (−6 nA, 2 ms) either 8 or 48 ms after the last action potential in a burst (Figure 6C). On its own, the brief current injections produced a hyperpolarization to about −120 mV (see Materials and Methods) and returned to rest in 46±2 ms (n = 12). Hyperpolarization at the early time point reduced the medium ADP amplitude more than expected by simply summing the medium ADP and the short hyperpolarization alone (Figure 6D and 6E), again consistent with the notion that the medium ADP is mediated by a voltage-dependent conductance that can be deactivated by hyperpolarization. This effect was measured at the time when the hyperpolarizing response on its own decayed back to rest (54±2 ms after the last spike). When the hyperpolarizing current step began 48 ms after the last action potential, its effect (again measured when the hyperpolarization on its own decayed to rest; 100±2 ms after the last spike) was not statistically significant (Figure 6D and 6E). This finding suggests that the VGCCs responsible for the medium ADP remain activated for at least 50 ms, but not longer than 100 ms, while the conductance responsible for the slow ADP is not deactivated by hyperpolarization.

The voltage dependence of the medium ADP, combined with its sensitivity to low concentrations of Ni²⁺, suggests that the medium ADP may result from upregulation of Ca_v2.3 R-type calcium channels. However, previous work has indicated that mGluR activation can downregulate K⁺ channels [17],[18],[25]–[28], which could result in inhibition of the AHP and activation of R-type channels, without any actual modulation of the calcium channels by mGluRs. To test this alternative hypothesis, we converted the medium AHP to a medium ADP by injecting a ramp current that followed action potential firing, resulting in an artificial medium ADP in control ACSF, which resembled the medium ADP following application of DHPG. This artificial medium ADP was unaffected by bath application of Ni²⁺, suggesting that the Ni²⁺-sensitive calcium channels are not significantly activated by a post-burst medium ADP in the absence of mGluR activation (Figure 7A and 7B). We also tested the voltage dependence of the medium ADP in the absence of DHPG by varying the initial amplitude of the ramp current. The relationship between the medium ADP amplitude and the current injection was linear (Figure 7C and 7D), suggesting that voltage-dependent conductances do not amplify the medium ADP over this voltage range in the absence of mGluR activation.

To further test the hypothesis that upregulation of R-type calcium channels is responsible for the DHPG-induced medium ADP, we performed experiments on Ca_v2.3 knockout mice [29]. The medium ADP induced by DHPG was significantly smaller in the knockout mice, compared to wild-type controls (Figure 8). Because the Ca_v2.3 knockout mice are a hybrid of C57BL/6J (black) and 129S1/SvImJ (brown) mice, pups had different coat colors (black, dark brown, light brown; see Materials and Methods). We analyzed the results from different-colored mice separately and found no differences between these groups. Furthermore, we performed control experiments using black and brown mice and found similar DHPG-induced medium ADP amplitude in each of the wild-type strains (Figure S7).

We also used voltage-clamp recording to measure isolated R-type calcium currents in CA1 pyramidal neurons (see Materials and Methods). The resulting currents were upregulated by DHPG when activated by large but not small depolarizing steps (Figure 9A and 9B), consistent with modulation of R-type (high voltage activated) but not T-type (low voltage activated) calcium currents. Finally, to explore the ability of mGluR5 activation to regulate current mediated by calcium channels, we co-expressed, in Xenopus oocytes, mGluR5 along with either Ca_v2.3 α₁ (plus α₂δ₁ and β₃) or Ca_v3.2 α₁ (see Materials and Methods; [30]). We found that DHPG application upregulated barium currents in oocytes expressing mGluR5 and Ca_v2.3 but not mGluR5 and Ca_v3.2, suggesting subunit selective modulation of R-type, but not T-type, calcium channels by mGluR5 activation with DHPG (Figure 9C and 9D).

The observation that the slow ADP was not eliminated by hyperpolarization suggests that a different class of channels may contribute to the slow ADP. We therefore examined this component of the ADP pharmacologically. It was not blocked by NiCl₂ (Figure S8A and S8B), consistent with the idea that it is not mediated by the VGCCs responsible for the medium ADP. The slow ADP was also unaffected by nimodipine (Figure S8A and S8B).

We performed a battery of pharmacological experiments to explore the signal transduction mechanisms responsible for the post-burst medium ADP and slow ADP. These experiments (Figure S8C and S8D) required either intracellular drug application or pre-incubation of the drug in the bath. Like the medium ADP, the slow ADP was blocked by group I mGluR antagonists, especially the mGluR5 blocker MPEP. The slow ADP was also reduced by intracellular BAPTA, but it was not blocked by drugs that interfere with Ca²⁺ release from intracellular stores (CPA, ruthenium red, or heparin). All of these drugs inhibited the medium ADP, suggesting the Ca²⁺ release is required for the medium ADP, but not the slow ADP. The medium ADP was blocked by intracellular GDP-β-S, which interferes with G-protein coupled signaling, or by the PLC inhibitor U73122 (but not the inactive analog U73343). None of these drugs blocked the slow ADP, however, suggesting further that distinct signaling mechanisms mediate the DHPG-induced medium ADP and slow ADP.

We examined the activity dependence of the DHPG-induced post-burst medium ADP by delivering pairs of bursts at intervals of 0.1 to 20 s. Three-spike bursts were used in order to limit the size of the medium ADP so that either facilitation or inhibition could be observed. At intervals up to 200 ms, the second burst evoked a medium ADP almost twice the size of the first; at intervals of 1–5 s, the second burst was reduced by about 25% (Figure 10). The data were well fit by a model consisting of two processes: a facilitation process with a decay time constant of 0.25 s and an inhibition process with a decay time constant of 10 s (Figure S9 and Text S1). In the model, inhibition affected the fraction of the current available to be activated and facilitation affected the probability of activation (by a burst) of the available current. A key feature of the model was that any portion of the current could be inhibited, regardless of whether or not it was activated. This model predicted that facilitation is not expected, but inhibition persists, for a third burst delivered following two bursts (Figure S9). We conducted this experiment and the results were consistent with the predictions of the model (Figure 10). By contrast, a model in which inactivation was limited to the activated channels could not explain the results of the three-pulse experiment (Figure S9 and Text S1).

Hippocampal slices were prepared from male Wistar rats 25–45 d old or mice (C57BL/6J or 129S1/SvImJ or Ca_v2.3 knockout) 21–35 d old. Voltage-clamp experiments were done on slices prepared from younger rats (13–18 d old) in order to reduce space-clamp problems. Knockout mice were derived from 129S1/SvImJ (brown mouse) embryonic stem-cell injections in C57BL/6J (black) mice [29]. The founder mice were bred to C57BL/6J females; therefore, offspring of the knockout mice had either light brown, dark brown, or black coat color.

Following anesthesia with halothane or isoflurane, animals were perfused through the heart with ice-cold ACSF (see below). The brain was removed rapidly and mounted in a near-horizontal plane for preparation of 300 µm hippocampal slices using a vibratome. Slices were prepared in either ice-cold ACSF or sucrose-based solution, then transferred to a chamber containing oxygenated ACSF (no sucrose) at approximately 35°C for half an hour. The slice chamber was then maintained at room temperature and slices were removed individually for electrophysiological recordings.

Whole-cell current-clamp recordings were obtained at 33±2°C. Patch-clamp electrodes were pulled from 2.0 mm outer diameter borosilicate glass and filled with a K-gluconate-based intracellular solution (see below). Electrode resistance was 3–6 MΩ in the bath and series resistance was 5–20 MΩ during the recordings. Current-clamp recordings were obtained with Dagan BVC-700 amplifiers, using appropriate bridge balance and electrode-capacitance compensation. Voltage-clamp recordings with appropriate capacitance and series resistance compensation were performed at room temperature (23–25°C) and monitored with an Axopatch 200B amplifier (Molecular Devices, Union City, CA). Data acquisition and analysis was performed using custom software written for Igor Pro. Statistical tests included the paired or unpaired t test and analysis of variance (one-way ANOVA or repeated measures one-way ANOVA) with Tukey's post hoc comparisons. All statistical analyses were performed using Prism 4 software and in most cases detailed results are provided in the figure legends.

For the hyperpolarizations shown in Figure 6, the −6 nA, 2 ms current steps produced hyperpolarizations that briefly (1–8 ms) exceeded the −10 V limit of the analog-to-digital converter. The peak hyperpolarization was therefore estimated by extrapolating double-exponential fits of the response to the peak time predicted from linear fits of the rising phase. The estimated peaks were −119±3 mV for the early steps (n = 12) and −117±2 mV for the late steps (n = 11). The small clipping effect had an insignificant effect on the subtraction procedure used (see Results), because the effect is measured when the hyperpolarization had decayed back to rest, more than 40 ms after the clipping ended.

To test whether synaptic activation can induce the post-burst ADP, 5 brief action potentials were somatically injected either with or without synaptic stimulation and the responses were monitored once every 5 min with 1 min delay between two conditions. In both the MPEP/LY and control groups, experiments were performed in the presence of blockers of ionotropic glutamate receptors (30 µM CNQX and 50 µM D-AP5) and GABA receptors (2 µM SR95531 and 1 µM CGP55845). Bipolar borosilicate theta glass stimulation electrodes (Sutter Instruments) filled with ACSF were used in conjunction with Dagan BSI-950 biphasic stimulus isolator. Stimulating electrodes were placed in proximal stratum radiatum and at least 100 µm away from the recorded cell and toward CA3. Stimulus intensity was set to produce a 5–11 mV ADP during synaptic stimulation.

Normal ACSF had the following composition (mM): 125 NaCl, 2.5 KCl, 25 NaHCO₃, 1.25 NaH₂PO₄, 1 MgCl₂, 2 CaCl₂, 25 Dextrose (Fisher Scientific; Sigma). In some cases slices were prepared in a modified ACSF in which 125 mM NaCl was replaced with 75 mM NaCl and 75 mM sucrose. In many experiments, drugs were added to the bath (see below). The bath perfusion rate was 2–3 ml/min.

The intracellular recording solution had the following composition (mM): 115 K-gluconate, 20 KCl, 10 Na₂phosphocreatine, 10 HEPES, 2 MgATP, 0.3 NaGTP, 0.1% Biocytin (Fisher Scientific; Sigma). In some experiments, drugs were added to the intracellular solution (BAPTA, GDP-β-S, U73122, U73343, ruthenium red, and heparin; see below); for BAPTA-containing internal solution, the K-gluconate concentration was reduced to 100 mM. The K-gluconate based internal solution was used because the properties of CA1 pyramidal neurons are more stable with this solution than with K-Methylsulfate based solutions [56].

For voltage-clamp experiments in slices, patch electrodes (3–6 MΩ in bath) were filled with intracellular solution containing the following (in mM): 110 Cs-gluconate, 25 TEA-Cl, 10 HEPES, 2 EGTA, 4 Mg-ATP, and 0.5 Na-GTP, 5 Na₂-phosphocreatine, pH 7.3 with CsOH. R- and T-type calcium currents were isolated pharmacologically by preincubating the slices in a mixture containing ω-conotoxin MVIIC (2 µM), ω-conotoxin-GVIA (2 µM), and ω-agatoxin IVA (0.2 µM) to block N-, P-, and Q-type Ca²⁺ currents and cytochrome c (0.1 mg/ml to block nonspecific toxin binding) for>1 h at room temperature. Nifedipine (20 µM) and TTX (1 µM) were bath applied to block L-type Ca²⁺ currents and Na⁺ currents, respectively. Recordings were performed in modified ACSF solution containing the following (in mM): 125 NaCl, 25 NaHCO₃, 2.5 KCl, 1.0 MgCl₂, 2.0 CaCl₂, 1.25 NaH₂PO₄, 25 NaHCO₃, and 10 dextrose, 2 CsCl, 5 4-AP, 10 TEA-Cl, pH 7.4.

The following drugs were obtained from Tocris: (S)-3,5-Dihydroxyphenylglycine (DHPG), (S)-(+)-a-Amino-4-carboxy-2-methylbenzeneacetic acid (LY367385), 2-Methyl-6-(phenylethynyl)pyridine hydrochloride (MPEP), Ammoniated ruthenium oxychloride (Ruthenium Red), 1,4-Dihydro-2,6-dimethyl-4-(3-nitrophenyl)-3,5-pyridine dicarboxylic acid 2-methyloxyethyl 1-methylethyl ester (Nimodipine), (6aR,11aS,11bR)-rel-10-Acetyl-2,6,6a,7,11a,11b-hexahydr o-7,7-dimethyl-9H-pyrrolo[1′,2′:2,3]isoindolo[4,5,6-cd] indol-9-one (CPA), D-(−)-2-Amino-5-phosphonopentanoic acid (D-AP5), 6-Cyano-7-nitroquinoxaline-2,3-dione disodium (CNQX disodium salt), (2S)-3-[[(1S)-1-(3,4-Dichlorophenyl)ethyl]amino-2-hydro xypropyl](phenylmethyl)phosphinic acid hydrochloride (CGP 55845 hydrochloride), and Octahydro-12-(hydroxymethyl)-2-imino-5,9∶7,10a-dimethan o-10aH-[1],[3]dioxocino[6,5-d]pyrimidine-4,7,10,11,12-pen tol citrate (Tetrodotoxin citrate).

The following drugs were obtained from Sigma: 1,2-Bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid tetrapotassium salt (BAPTA), 1-[6-[((17β)-3-Methoxyestra-1,3,5[10]-trien-17-yl)amino]hexyl]-1H-pyrrole-2,5-dione (U73122), 1-[6-[((17β)-3-Methoxyestra-1,3,5[10]-trien-17-yl)amino]hexyl]-2,5-pyrrolidinedione (Uf73343), Guanosine 5′-[β-thio]diphosphate trilithium salt (GDP-β-S), Heparin sodium salt (from porcine intestinal mucosa average mol wt ∼3,000 kD), Nickel(II) chloride hexahydrate (NiCl₂), 2-(3-Carboxypropyl)-3-amino-6-(4 methoxyphenyl)pyridazinium bromide (SR 95531), Dextrose, K-gluconate, sodium phosphocreatine, HEPES, MgATP, NaGTP, and biocytin.

In most experiments, DHPG was applied by bath perfusion at a concentration of 4 µM. Some batches of DHPG were more potent than others, so in some cases it was necessary to reduce the DHPG concentration to as low as 2 µM in order to prevent additional spiking following the triggered burst of action potentials. Experiments using 2–4 µM DHPG were pooled together for analysis. In most experiments, the membrane potential was held at −65 mV, which required very small current injections (< 50 pA). DHPG application resulted in a depolarization of 3–5 mV, so the holding potential was adjusted to −65 mV with hyperpolarizing holding current. In some experiments (as noted), DHPG (500 µM DHPG) or ACSF was applied to the cell via pressure application from a broken patch pipette. Pressure (10 psi, 0.2 s) was applied via a Dagan PMI-100 pressure micro-injector. Current was injected to the cell within 3 s of pressure application. For experiments with mouse slices, bath application was performed using 10 µM DHPG, yielding an ADP similar to that observed with 4 µM DHPG in rat slices.

The cDNAs for the Ca_v3.2 (accession number AF051946), Ca_v2.3 (L27745), β₃ (M88751), α₂δ₁ (M86621), and mGluR5 (D10891) were subcloned into a high expression vector pGEMHEA, which contains the 5′ and 3′ untranslated regions of the Xenopus β globin gene, linearized, and transcribed into cRNA using T7 RNA polymerase according to the manufacturer's protocol (Ambion, Austin, TX, USA).

Oocytes were obtained from female Xenopus laevis (Nasco, WI, USA) using a standard procedure. Several ovary lobes were surgically removed from mature female Xenopus laevis and torn into small clusters in SOS solution (in mM: 100 NaCl, 2 KCl, 1.8 CaCl₂, 1 MgCl₂, 5 HEPES, 2.5 pyruvic acid, 50 µg/ml gentamicin, pH 7.6). The follicular membranes were removed by digestion in Ca²⁺-free Barth's solution (in mM: 88 NaCl, 1 KCl, 2.4 NaHCO₃, 0.82 MgSO₄, pH 7.4) containing 20 mg/ml collagenase (SERVA, Heidelberg, Germany). Oocytes were injected under a stereo-microscope with 2–20 ng of cRNA using a Drummond Nanoject pipette injector (Parkway, PA, USA) attached to a Narishige micromanipulator (Tokyo, Japan).

Barium currents were measured at room temperature 4 to 8 d after cRNA injection using a two-electrode voltage-clamp amplifier (OC-725C, Warner Instruments, Hamden, CT, USA). Microelectrodes (Warner Instruments, Hamden, CT, USA) were filled with 3 M KCl and their resistances were 0.2–1.0 MΩ. The 10 mM Ba²⁺ bath solution contained (in mM): 10 Ba(OH)₂, 90 NaOH, 1 KOH, 5 HEPES (pH 7.4 with methanesulfonic acid). The currents were sampled at 5 kHz and low pass filtered at 1 kHz using the pClamp system (Digidata 1322A and pClamp 8; Axon instruments, Foster City, CA, USA). Peak currents and exponential fits to currents were analyzed using Clampfit software (Axon instruments, Foster City, CA, USA).