G protein activation by uncaging of GTP-γ-S in the leech giant glial cell

A large variety of cellular responses in neurons and glial cells is mediated by metabotropic membrane receptors linked to G proteins(Gilman, 1987; Brauner-Osborne et al., 2007). One of the most prevalent pathways initiated by activation of G proteins is the activation of phospholipase C (PLC), which leads to the formation of inositol-trisphosphate (IP₃) and the release of Ca²⁺from intracellular stores (Berridge et al.,2000). As electrically inexcitable cells, glial cells use this pathway in response to many kinds of stimuli, including neurotransmitters,neuromodulators, growth factors and cytokines(Lohr and Deitmer, 2006; Fiacco and McCarthy, 2006). In the nervous system, glial cells often respond to active neurons releasing neurotransmitters, which activate metabotropic receptors in the glial cells. Recent evidence suggests that glial cells may themselves release transmitters such as glutamate and ATP, mediated by cytosolic Ca²⁺ transients(Araque et al., 2001; Pascual et al., 2005). Due to the presence of several G proteins, which initiate or suppress different signalling cascades (for a review, see Luttrell, 2006), we tried to elucidate the dominant pathways initiated by global activation of G protein.

We have previously shown in the leech giant glial cell in situthat stimulation of the Leydig neuron elicits a membrane hyperpolarization due to an increased K⁺ conductance in the giant glial cell(Britz et al., 2002). We were able to show that Leydig neurons release the peptide myomodulin, which activates metabotropic membrane receptors in the glial cell membrane and results in the activation of G protein-coupled adenylyl cyclase, leading to an increase in K⁺ conductance and a membrane hyperpolarization(Britz et al., 2004). A similar signalling cascade to that of Leydig neuron stimulation and myomodulin application is initiated by 5-hydroxytryptamine (5-HT)(Britz et al., 2005), a major neurotransmitter and neuromodulator in the leech central nervous system(Lent et al., 1991). The action of both ligands, myomodulin and 5-HT, was suppressed after blocking G proteins by injecting GDP-β-S into the glial cell. These studies raised the question of whether the negative membrane potential in this and other glial cells may in part be attributable to neuronal activity by activating G protein-coupled glial membrane receptors. Since this K⁺ conductance was not affected by inhibition of protein kinase A (PKA), we suggested that it is mediated by ion channels directly gated by cyclic nucleotides(Britz et al., 2004; Britz et al., 2005).

As 5-HT also evokes cytosolic Ca²⁺ transients in this glial cell(Lohr and Deitmer, 1999),another signalling cascade can be triggered by the same ligand, presumably due to the expression of different types of 5-HT receptors in the leech central nervous system. Although the K⁺ conductance induced by 5-HT was not dependent on external Ca²⁺(Britz et al., 2005), the question remained of whether conductance changes evoked by agonists of metabotropic transmitters and modulators, which are mediated by G protein activation, are principally independent of changes in cytosolic Ca²⁺, even if cytosolic Ca²⁺ rises at the same time.

In the present study, we activated G proteins in the leech giant glial directly by injecting caged GTP-γ-S, which was released by brief UV illumination. We assume that this method would not discriminate between different G proteins in the giant glial cell and would give us the effects of`global' G protein activation. This should also give us an idea about the most dominant effects mediated by G protein activation in this glial cell. Our results show that glial cells readily hyperpolarize due to an increase of the membrane K⁺ conductance and evoke cytosolic Ca²⁺ and Na⁺ transients, suggesting that the glial membrane potential is under metabotropic control of neuronal activity.

The experiments were performed on exposed giant glial cells in the neuropil of isolated ganglia of the leech Hirudo medicinalis L. For recording with intracellular conventional and ion-selective microelectrodes, and for confocal imaging in the giant glial cell, the ganglionic capsule and some overlying neuronal cell bodies of the ganglia were mechanically removed on the ventral side after prior enzymatic treatment with collagenase/dispase (2 mg ml^–1 for up to 30 min) and fine micro-dissection. The steps of dissection and the preparation have been described before(Britz et al., 2002). Isolated ganglia were stored in leech saline at 4°C, before being transferred into the experimental chamber.

The non-hydrolyzable G protein activator GTP-γ-S, dissolved in H₂O, was injected into both giant glial cells as inactive `caged'compound, S-(DMNPE-caged) {guanosine 5′-O-(3-thiotriphosphate),P^3(S)-[1-(4,5-dimethoxy-2-nitrophenyl)ethyl]ester, triammonium salt} (Molecular Probes, Eugene, OR, USA) by constant or pulsed (2 s at 0.25 Hz) iontophoretic currents of –5 nA for 10 min via a microelectrode containing 10 mmol l^–1 caged GTP-γ-S. After the injection, 30 min was allowed for diffusion of GTP-γ-S within the giant glial cell before the caged compound was uncaged. Photolysis of caged GTP-γ-S was carried out by illumination using a mercury arc lamp(model USH-102D; Ushio Electric, Tsukuba, Japan) at 355 nm. The light was focussed onto the ganglion and carefully adjusted to allow full photolysis of the caged compound without affecting neuronal and glial properties (e.g. membrane potential changes) when applied alone. The UV photolysis of caged GTP-γ-S results in the release of the active, non-hydrolyzable, GTP analogue in the glial cells (Dolphin et al., 1988).

Caged IP₃ (NPE-caged Ins-1,4,5-P₃) (Molecular Probes), dissolved in H₂O, was injected into the glial cells, and photolysis was induced in a similar way to caged GTP-γ-S.

For injection of GDP-β-S, which blocks G protein-mediated responses,or, as a control, GDP, a micropipette was filled with either 10 mmol l^–1 GDP-β-S or 10 mmol l^–1 GDP(Sigma-Aldrich Chemie, Taufkirchen, Germany), dissolved in 0.5 mol l^–1 K-acetate. In order to minimize leakage of the negatively charged drugs from the pipette, a positive backing current was delivered by clamping the cells at a holding potential 1–3 mV positive to the resting membrane potential. Injection was then obtained with sustained or repetitive hyperpolarizing voltage pulses of 3 mV for 8 min.

Electrophysiological recordings were performed with intracellular microelectrodes filled with 2 mol l^–1 K-acetate/20 mmol l^–1 KCl or 3 mol l^–1 KCl for membrane potential measurements. In two-electrode voltage-clamp experiments, the microelectrodes were filled with 3 mol l^–1 KCl for potential recording and 2 mol l^–1 K-citrate, adjusted to pH 7.0 with HCl, for current injection (resistance 1–3×10⁷Ω). For current and voltage-clamp recordings, Axoclamp 2A and/or 2B amplifiers (Axon Instruments, Union City, CA, USA) were used. Membrane currents were recorded by the built-in current measurement circuit of the headstages. The experimental bath was grounded via a chloride-covered silver wire in agar dissolved in normal saline. For further details, see Deitmer and Schneider (Deitmer and Schneider, 1995). The membrane input resistance was measured by injecting hyperpolarizing current pulses (10–20 nA for 1–2 s).

A Leydig neuron was stimulated with single square current pulses (2–4 nA, 10–40 ms) via an intracellular sharp microelectrode,controlled by digitimers (D4030 and DS2; Digitimer Limited, Welwyn Garden City, UK), to induce firing frequencies of 10 Hz (for details, see Britz et al., 2002; Britz et al., 2004).

For measurements of the intracellular Na⁺ activity and the membrane potential, single-barrelled Na⁺-selective and voltage-sensing reference microelectrodes were used. The ion-selective electrode was silanized with a drop of 5% tri-N-butylchlorosilane in 99.9% pure carbon tetrachloride, backfilled into the tip. The micropipette was baked for 4.5 min at 450°C on a hotplate. For Na⁺-selective microelectrodes, a drop of Na⁺-cocktail [mixture of Na⁺-ionophore (Fluka 71739), organic solvent (Fluka 73732) and a lipophilic salt (Fluka 72018) at a mass percent ratio of 10:89.5:0.5,respectively] was backfilled into the silanized barrel of the electrode, which was then filled up with 0.1 mol l^–1 NaCl/10 mmol l^–1 3-(N-morpholino) propanesulphonic acid (MOPS),pH 7.0. The reference electrode was filled with 3 mol l^–1KCl. The ion-selective and the reference electrodes were connected by chloride-covered silver wires to the headstages of an electrometer amplifier. The electrodes were calibrated in leech salines with different Na⁺concentrations (85 mmol l^–1, 15 mmol l^–1, 10 mmol l^–1, 8.5 mmol l^–1, 5 mmol l^–1) prior to and at the end of every experiment. In calibrating solutions with reduced Na⁺ concentration,Na⁺ was replaced by equimolar amounts of N-methyl-d-glucamine (NMDG, Fluka). Only electrodes with a response greater than 50 mV for a 10-fold change in the Na⁺concentration were used for the experiments.

The calcium-sensitive fluorescent dye Oregon Green 488 BAPTA-1 (OGB;Molecular Probes) was dissolved in water at a concentration of 5 mmol l^–1 and used to fill a microelectrode. The resistance of the microelectrodes filled with the dye solution was 120–150×10⁶ Ω. The microelectrodes were positioned with a micromanipulator (UM-3FC, U Ltd, Tokyo, Japan), fixed at the stage of an upright microscope (Leica DMRB, Bensheim, Germany). An individual giant glial cell was impaled by microelectrodes with a piezo stepper(P-840.20, Physik Instrumente, Waldbronn, Germany). The dye was injected iontophoretically into the glial cell by a negative current of –3 nA for about 10 min. 15–30 min after the termination of the dye injection, the fluorescence in the cell body and the cell processes reached constant values,indicating that the dye was equally distributed within the cell. OGB was excited by the 488 nm line of a Krypton–Argon laser, and images were taken with a confocal laser scanning microscope (Leica TCS 4D) at a sample rate of 0.25–4.7 Hz. The excitation light and the emission light were separated by a dichroic mirror at 510 nm. The emission light was truncated by a 515–545 nm band-pass filter. Regions of interest were defined in the first of a sequence of images, and the normalized fluorescence changes,Δ F (in %), were measured throughout the image sequence.

The standard physiological leech saline for the experiments contained (in mmol l^–1): NaCl, 85; KCl, 4; CaCl₂, 2;MgCl₂, 1; Hepes(N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid),10; pH adjusted to 7.4 with NaOH. In saline containing 10 mmol l^–1 or 40 mmol l^–1 K⁺, KCl was exchanged for an equivalent amount of NaCl. Cyclopiazonic acid (CPA) (Alexis Biochemicals, Lörrach, Germany) was dissolved in dimethyl sulfoxide(DMSO) as stock solutions and added to saline to make a final concentration of 20 μmol l^–1. The DMSO concentration never exceeded 0.2% in the experimental saline. The leech myomodulin was synthesized according to Wang et al. (Wang et al.,1998) as described before(Britz et al., 2002). All experiments were done at room temperature (22–24°C).

All data are given as means ± standard error of the mean (s.e.m.), N indicates the number of experiments. The glial responses were measured either in the anterior or in the posterior giant glial cell, and the results were pooled, since no differences in the responses of the two glial cells can be observed (Schmidt and Deitmer, 1999; Britz et al.,2002). The statistical significance of the differences between mean values was tested using a standard, or a paired, t-test, if applicable. Differences were indicated in some figures as statistically significant for an error probability of P<0.05, P<0.01 or P<0.001, as indicated by one, two or three asterisks,respectively.

Persistent activation of G proteins was attempted by cell loading with the non-hydrolyzable GTP analogue GTP-γ-S. We injected the GTP-γ-S ionophoretically in caged form into both giant glial cells. When the ganglion was illuminated with moderate UV light (`UV pulse') for 60 s to uncage GTP-γ-S, a rapid membrane hyperpolarization was measured, which did not reverse during the duration of the experiment of up to 1 h(Fig. 1A). UV illumination of ganglia without prior injection of caged GTP-γ-S into the glial cell,had no effect on the glial membrane potential (N=13)(Fig. 1E). The UV illumination as used here had no effect on the resting membrane potential and on the firing frequency and kinetics of spontaneous action potentials of Leydig neurons or N-cells (N=4; not shown).

The size of the glial membrane hyperpolarization following the uncaging of GTP-γ-S was dependent on the resting potential of the cell; the less negative the resting potential, the larger was the glial response to uncaging of GTP-γ-S (N=22) (Fig. 1B). The regression line (coefficient of correlation r=–0.943) indicated a reversal potential of the glial response of –75 mV, which is close to the K⁺ equilibrium potential(Britz et al., 2005). A second UV pulse (N=11), or a UV pulse on non-injected cells (N=13),had no effect on the glial membrane potential (open symbols in Fig. 1B).

We also measured the input resistance of the glial membrane before and after UV illumination of caged GTP-γ-S-injected and non-injected glial cells. The input resistance, recorded by repeated negative currents injected into the glial cell, decreased after uncaging GTP-γ-S(Fig. 1C), on average by 36%from 315±32 kΩ to 200±28 kΩ (N=7; P<0.01) (Fig. 1D). By contrast, non-injected glial cells showed no change in their input resistance before (369±46 kΩ) and after (370±48 kΩ)UV illumination (N=6) (Fig. 1E,F). These results indicate that uncaging GTP-γ-S induces a persistent membrane hyperpolarization and an increase in membrane conductance of the glial membrane.

Iontophoretic injection of caged GTP-γ-S alone (without uncaging) had no effect on the glial cell, and Leydig interneuron stimulation (10 Hz, 1 min)(Fig. 2A), myomodulin(MMHir) (3 μmol l^–1)(Fig. 2B) and 5-HT (20 μmol l^–1; not shown) still evoked a membrane hyperpolarization in the glial cell, as described previously(Britz et al., 2002; Britz et al., 2004). After uncaging of GTP-γ-S and the subsequent membrane hyperpolarization,neither MMHir nor Leydig neuron stimulation had any additional effect on the glial membrane potential (Fig. 2 A,B). Repeated UV illumination had only a small or no effect on the glial membrane potential once the glial cell membrane had hyperpolarized. In glial cells not injected, Leydig neuron stimulation elicited a membrane hyperpolarization, but UV pulses had no effect (Inset, Fig. 2A).

The K⁺ permeability was evaluated by the membrane depolarisation induced by raising the extracellular K⁺ concentration from 4 to 10 mmol l^–1 for 1–2 min (see Fig. 2A–C). The amplitude and the rate of rise of this depolarization give an indication of the relative K⁺ permeability (dV/dtindicative of the induced membrane current). The glial membrane depolarization to 10 mmol l^–1 K⁺ depended on the resting membrane potential but was considerably larger in glial cells, which had been hyperpolarized following the uncaging of GTP-γ-S(Fig. 2D). The rate of rise of the glial membrane depolarisation in 10 mmol l^–1K⁺ (dV/dt), indicative of the current flow upon raising the K⁺ concentration, was almost doubled, rising from 15.3±2.5 mV min^–1 to 28.5±5.3 mV min^–1 (N=9–12; P<0.05)(Fig. 2E) after uncaging GTP-γ-S. These results show that the membrane hyperpolarization induced by uncaging GTP-γ-S was accompanied by a rise in the relative K⁺ conductance of the glial cell membrane, indicating that G protein activation increases the K⁺ permeability of the glial cell membrane, as had been shown to be induced by Leydig neuron stimulation,myomodulin and 5-HT (Britz et al.,2004; Britz et al.,2005).

The increase in K⁺ conductance induced by the uncaging of GTP-γ-S was not due to the hyperpolarization per se, since the glial cell current–voltage relationship is nearly linear over the potential range of –50 to –100 mV (not shown here)(Munsch and Deitmer, 1994). The glial responses to myomodulin and Leydig neuron stimulation and to 5-HT could be blocked by the injection of the non-hydrolyzable G protein inhibitor GDP-β-S (Britz et al.,2004; Britz et al.,2005). We measured the membrane current and membrane resistance following G protein blockade by injecting the G protein inhibitor GDP-β-S into the glial cell. In voltage-clamped glial cells, GDP-β-S, but not GDP, led to a net inward current (Fig. 3A). The membrane conductance of the glial membrane decreased after injection of GDP-β-S by 20%, from 4.9±0.4 μS to 3.9±0.3 μS (Fig. 3B,C). Injection of the hydrolyzable analogue GDP had no effect on membrane potential and membrane resistance of the giant glial cell(Fig. 3D,E).

The cytosolic Ca²⁺, monitored by the use of the Ca²⁺indicator dye Oregon Green BAPTA-1 (OGB) injected into the leech giant glial cell, and with confocal microscopy, increased following uncaging of GTP-γ-S by a UV pulse (Fig. 4A,C). A second UV pulse, or UV illumination to a cell not injected with caged GTP-γ-S, had no effect on the cytosolic Ca²⁺. By contrast, raising the extracellular K⁺concentration from 4 to 40 mmol l^–1, which depolarized the cell to near –30 mV, evoked a Ca²⁺ transient in all cells injected and not injected with GTP-γ-S(Fig. 4A–C). This Ca²⁺ rise had been shown to be due to voltage-dependent Ca²⁺ influx (Lohr and Deitmer,2006; Munsch and Deitmer,1992). The amplitude of the Ca²⁺ transients amounted to about 30% fluorescence change for both stimuli(Fig. 4C). After depletion of the intracellular Ca²⁺ stores with cyclopiazonic acid (CPA; 20μmol l^–1), GTP-γ-S still evoked a membrane hyperpolarization but did not elicit any cytosolic Ca²⁺ change(Fig. 4D,E). This indicates that the Ca²⁺ rise evoked by GTP-γ-S can be attributed to release of Ca²⁺ from intracellular stores. Membrane depolarization by 40 mmol l^–1 K⁺ still induced a rise in cytosolic Ca²⁺, which amounted to 18±2% fluorescence change(N=4) (Fig. 4E). Our experiments suggest that the membrane hyperpolarization following release of GTP-γ-S was independent of the increase in cytosolic Ca²⁺,and presumably due to a Ca²⁺-independent K⁺ conductance increase, as described for the 5-HT-evoked, cAMP-mediated, K⁺conductance in this cell (Britz et al.,2005). We confirmed this conclusion further by the injection of BAPTA into the giant glial cell, which suppressed the Ca²⁺ rise(not shown) but did not abolish the membrane hyperpolarization elicited by uncaging GTP-γ-S (Fig. 4F). The membrane potential was –53.5±1.5 mV before and –51.4±1.6 mV after injection of BAPTA (not significant; N=10); subsequent uncaging of GTP-γ-S hyperpolarized the membrane to –74.3±1.3 mV (P<0.01, N=10).

The release of GTP-γ-S by a UV pulse also affected the cytosolic Na⁺ as measured with ion-selective microelectrodes in the leech giant glial cell. During the membrane hyperpolarization following uncaging of GTP-γ-S, the cytosolic Na⁺ concentration increased from 7.6±1.8 mmol l^–1 to 13.2±1.7 mmol l^–1 (N=8, P<0.01)(Fig. 5A,B). In cells not injected with GTP-γ-S, UV illumination had no effect on the cytosolic Na⁺ (Fig. 5C). Application of 5-HT (20 μmol l^–1), which also hyperpolarizes the cell membrane of the giant glial cell, did not change the cytosolic Na⁺ level, suggesting that it was not the membrane hyperpolarization per se that induced the rise in cytosolic Na⁺ (not shown here).

In order to check if the rise in Na⁺ was linked to the release of Ca²⁺ from intracellular stores, we depleted these stores with CPA prior to releasing GTP-γ-S. The cytosolic Na⁺ did not change after the addition of CPA, nor did it change upon uncaging GTP-γ-S in the presence of CPA (Fig. 5D). As described above, the membrane hyperpolarization evoked by uncaging GTP-γ-S was not affected by CPA. These experiments suggest that the rise of intracellular Na⁺ following uncaging of GTP-γ-S was secondary to the release of Ca²⁺ from intracellular stores,presumably due to enhanced Na⁺/Ca²⁺ exchange activity,and not due to a Na⁺ leak of the cell membrane, which would be expected to increase during the membrane hyperpolarization due to the larger Na⁺ gradient generated.

In order to check the effects of a Ca²⁺ increase alone, we injected caged inositol-trisphosphate (IP₃) into the giant glial cell and uncaged it with a moderate UV pulse. This should lead to the release of Ca²⁺ from intracellular stores via activation of IP₃ receptor channels. Monitoring cytosolic Ca²⁺indicated that uncaging of IP₃ evoked a Ca²⁺ rise of similar amplitude as membrane depolarization with 40 mmol l^–1K⁺ (Fig. 6A–C). A fast rise of cytosolic Ca²⁺ was followed by a rapid recovery, and a second UV pulse had no effect on the recovered level of intracellular Ca²⁺. The membrane potential was not affected by uncaging IP₃.

Uncaging of IP₃ also evoked a small but significant rise in cytosolic Na⁺ by 1.4 mmol l^–1 from 8.5±0.5 mmol l^–1 to 9.9±0.6 mmol l^–1(N=7; P<0.05) (Fig. 6D,E). A second UV pulse had only a small or no effect on the cytosolic Na⁺ level. Thus, Ca²⁺ release from intracellular stores evokes an Na⁺ rise, which supports the notion that this Na⁺ is due to an increase of Na⁺/Ca²⁺ exchange. These experiments also indicate that,although uncaging IP₃ elicited a transient rise in cytosolic Ca²⁺ and Na⁺, these ion changes were not sufficient to induce a membrane hyperpolarization.

G protein activation by extracellular signalling molecules is one of the most frequently employed mechanisms to activate cells, including neurons and glial cells in the brain (Gilman,1987; Brauner-Osborne et al.,2007). Our experiments show that activation of G proteins by uncaging of GTP-γ-S mimics the effects of several neurotransmitters/neuromodulators activating metabotropic receptors in the leech central nervous system. Uncaging GTP-γ-S resulted in membrane hyperpolarization, due to a K⁺ conductance increase, and a rise in cytosolic Ca²⁺, due to release from intracellular stores, and cytosolic Na⁺, presumably attributable to increased Na⁺/Ca²⁺ exchange. Our study shows that `global'activation of G proteins in the giant glial cell leads to the activation of a number of signalling cascades, which in turn affects various ion gradients and membrane conductance.

Ligands usually activate metabotropic receptors, which are linked to specific, excitatory or inhibitory G proteins(Luttrell, 2006). However, one or more G proteins can be activated by simultaneous activation of several types of metabotropic receptors (Werry et al., 2003). This may trigger multiple signalling cascades in a given cell, leading to the activation of diverse cell functions. The technique of uncaging caged molecules by UV illumination has become a powerful technique to increase the non-hydrolyzable G protein activator GTP-γ-S in a relatively short time to initiate G protein-mediated cellular activity(Dolphin et al., 1988; Oberhauser et al., 1998). Since we studied effects that were relatively slow in this large cell, we used moderate UV illumination to avoid the side effects of strong UV light. For example, ion channels can be sensitive to UV, and strong illumination may therefore result in a change in neuronal activity(Hof and Fox, 1983; Middendorf et al., 2000). We could observe changes in membrane potential and firing frequency of several neurons in the leech central nervous system when we used strong UV pulses. Therefore, we used moderate UV pulses of longer duration (up to 2 min), which still lead to the uncaging of intracellularly injected caged compounds but do not alter neuronal activity. UV illumination without prior injection of GTP-γ-S also had no effect on the membrane potential of the giant glial cell.

The increase in the membrane K⁺ conductance and the subsequent hyperpolarization following uncaging of GTP-γ-S was likely to be attributable to cAMP produced by the activation of adenylyl cyclase. This had been studied in detail for the neurotransmitters 5-HT and myomodulin, which both evoked a GDP-β-S-sensitive increase in the membrane K⁺conductance of the leech giant glial cell, and which were blocked by inhibition of adenylyl cyclase and mimicked by membrane-permeant cAMP analogues (Britz et al., 2004; Britz et al., 2005). It can be concluded that `global' activation of G proteins by uncaging of GTP-γ-S activates the same signalling pathways leading to the conductance increase as initiated by 5-HT and myomodulin and following Leydig neuron stimulation. Since inhibition of PKA did not inhibit this effect, the K⁺conductance was suggested to be due to activation of cyclic nucleotide-gated channels in the glial membrane (Britz et al., 2004; Hirth and Deitmer,2006). The K⁺ conductance elicited by 5-HT was also shown to be independent of Ca²⁺(Britz et al., 2005), which is supported by our present findings that the membrane hyperpolarization induced by uncaging of GTP-γ-S was neither suppressed by injection of the fast Ca²⁺ chelator BAPTA nor by prior depletion of intracellular Ca²⁺ stores by CPA.

The other signalling pathway activated by uncaging GTP-γ-S in the giant glial cell resulted in the release of Ca²⁺ from intracellular stores mediated by phospholipase C (PLC) and the production of IP₃(Berridge et al., 2003). We could show that uncaging of IP₃ elicited a similar transient Ca²⁺ rise as uncaging of GTP-γ-S. The rise of cytosolic Na⁺, however, was apparently secondary to the cytosolic Ca²⁺ increase, since it was abolished after the intracellular Ca²⁺ stores were depleted by CPA, suggesting increased Na⁺/Ca²⁺ exchange as the source for the elevated cytosolic Na⁺. Na⁺/Ca²⁺ exchange is one of the main processes to regulate cytosolic Ca²⁺ and is activated by increased intracellular Ca²⁺ concentrations(Annunziato et al., 2004; DiPolo and Beaugé,2006). In the leech giant glial cell, we could show that both plasmalemmal Ca²⁺-ATPase and Na⁺/Ca²⁺exchange contribute to the recovery of Ca²⁺ from a cytosolic rise(Nett and Deitmer, 1998a). In rat cerebellar astrocytes, Na⁺/Ca²⁺ exchange was shown to be responsible for maintaining a low resting cytosolic Ca²⁺(DiPolo and Beaugé,2006). We hypothesize that a transient intracellular Na⁺ rise may generally follow Ca²⁺ release from stores and global cytosolic Ca²⁺ signalling in glial cells and other cell types, as initiated by activation of metabotropic receptors. A rise of cytosolic Na⁺ would be expected to reduce membrane transport processes driven by the Na⁺ gradient across the glial cell membrane, such as the excitatory amino acid transporter EAAT, which is instrumental for the clearance of synaptically released glutamate. Hence,cytosolic Ca²⁺ rises in glial cells may modulate glutamatergic transmission by decreasing Na⁺-dependent glutamate uptake(Marcaggi and Attwell,2004).

Uncaging IP₃ elicited cytosolic Ca²⁺ and Na⁺ transients, as uncaging of GTP-γ-S did, but no membrane hyperpolarization, confirming the above conclusion that the G protein-mediated increase in K⁺ conductance and membrane hyperpolarization is not Ca²⁺- or Na⁺-dependent. In the leech giant glial cell,an increased K⁺ conductance can be elicited by removal of external Na⁺, which is Ca²⁺ independent and presumably attributable to a decreased cytosolic Na⁺ concentration(Nett and Deitmer, 1998b). The experiments also support the notion that it was the release of Ca²⁺from intracellular stores that caused the rise in intracellular Na⁺, and not G protein activation by GTP-γ-S per se. It can be concluded that the cAMP-mediated K⁺ conductance in the leech giant glial cell, elicited either by neurotransmitters or, as in the present study, by G protein activation, is neither Ca²⁺- nor Na⁺-dependent. The rise in cytosolic Na⁺ may itself affect Na⁺-dependent transport processes, in particular glial glutamate uptake, which is greatly dependent on the Na⁺ gradient across the membrane. This would leave more glutamate for a longer while in synaptic areas and thus may be a glial feedback response to the synaptic activity between neurons.

We thank Christian Lohr for his comments on an earlier version of this manuscript and Hans-Peter Schneider for technical assistance during some experiments. We gratefully acknowledge the financial support by the Deutsche Forschungsgemeinschaft (De 231/18-1, 18-2).