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Institute of Physiology, Humboldt University, Tucholskystr. 2, D-10117 Berlin, Germany

	Abstract

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Activation of microglial cells, the resident macrophages of the brain, occurs rapidly following brain injury. De-ramification, i.e. transformation from ramified into amoeboid morphology is one of the earliest manifestations of microglial activation. In the present study, we identified the physiological mechanisms underlying microglial de-ramification induced by lysophosphatidylcholine (LPC). Patch-clamp experiments revealed activation of non-selective cation currents and Ca²⁺-dependent K⁺ currents by extracellular LPC. LPC-activated non-selective cation channels were permeable for monovalent and divalent cations. They were inhibited by Gd³⁺, La³⁺, Zn²⁺ and Grammostola spatulata venom, but were unaffected by diltiazem, LOE908MS, amiloride and DIDS. Ca²⁺ influx through non-selective cation channels caused sustained increases in intracellular Ca²⁺ concentration. These Ca²⁺ increases were sufficient to elicit charybdotoxin-sensitive Ca²⁺-dependent K⁺ currents. However, increased [Ca²⁺]_i was not required for LPC-induced morphological changes. In LPC-stimulated microglial cells, non-selective cation currents caused transient membrane depolarization, which was followed by sustained membrane hyperpolarization induced by Ca²⁺-dependent K⁺ currents. Furthermore, LPC elicited K⁺ efflux by stimulating electroneutral K⁺–Cl^– cotransporters, which were inhibited by furosemide and DIOA. LPC-induced microglial de-ramification was prevented by simultaneous inhibition of non-selective cation channels and K⁺–Cl^– cotransporters, suggesting their functional importance for microglial activation.

(Received 5 January 2004; accepted after revision 10 March 2004; first published online 12 March 2004)
Corresponding author C. Eder: Institute of Physiology, Humboldt University, Tucholskystr. 2, D-10117 Berlin, Germany. Email: claudia.eder{at}charite.de

	Introduction

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Formation of lysophosphatidylcholine (LPC) is enhanced in the pathological central nervous system. Increased concentrations of LPC were detected following ischaemia (Sun & Foudin, 1984; Kinouchi et al. 1990), under inflammatory conditions (Kabarowski et al. 2002) and in epileptic tissue (Yegin et al. 2002). LPC is generated by hydrolysis of phosphatidylcholine (PC) via the action of phospholipase A₂ (PLA₂). Under normal physiological conditions, the majority of LPC formed in the tissue is rapidly metabolized or reacetylated. In pathological situations, overstimulation of PLA₂ results in PC breakdown and subsequent accumulation of LPC in the damaged tissue (Kabarowski et al. 2002; Xu, 2002; Yegin et al. 2002). Increases in PLA₂ activity occur in various brain diseases, such as ischaemia, epilepsy, Alzheimer's disease, schizophrenia and spinal cord injury (Farooqui et al. 1999).

A wide variety of in vitro and in vivo studies has demonstrated that LPC promotes the activation of microglia and other immune cells. LPC-induced activation processes include shape changes (Ousman & David, 2000), stimulation of cell motility (Quinn et al. 1988), expression of surface antigens (Kume et al. 1992), release of proinflammatory cytokines (Huang et al. 1999) and others. Although the capability of LPC to activate immune cells has been well demonstrated, the underlying physiological mechanisms of these activation processes are unknown.

LPC has been found to modulate ion channel activity in various cell preparations. In neurones, LPC causes prolonged membrane hyperpolarization due to the activation of the two-pore domain K⁺ channels TREK-1, TREK-2 and TRAAK (Lesage et al. 2000; Maingret et al. 2000). By shifting the steady-state inactivation curve in depolarizing direction, LPC induced increases of HERG K⁺ currents in HEK cells (Wang et al. 2001). Furthermore, LPC has been shown to evoke sustained intracellular Ca²⁺ increases, which were linked to the activation of either voltage-gated L-type Ca²⁺ channels (Golfman et al. 1999) or non-selective cation channels (Terasawa et al. 2002; Yokoyama et al. 2002). In cardiac myocytes, LPC enhances slow Na⁺ currents (Burnashev et al. 1989; Undrovinas et al. 1992). Besides the up-regulation of certain types of ion channels, LPC has also been demonstrated to inhibit ion channel activity. In coronary smooth muscle cells, LPC causes inhibition of delayed rectifier K⁺ currents by shifting their steady-state activation curve in depolarizing direction (Yeon et al. 2001). LPC-induced reduction of inward rectifier K⁺ currents was detected in cardiac cells by Kiyosue & Arita (1986). Furthermore, inhibitory effects of LPC on fast Na⁺ currents (Burnashev et al. 1989; Sato et al. 1992) and on NMDA-gated currents (Ikeuchi et al. 1995) were previously described. Although numerous studies report LPC-induced effects on electrophysiological properties of various cell types, data on immune cells, including microglia, are not available.

In the present study, we have identified the ion channels and transporters, namely non-selective cation channels, Ca²⁺-dependent K⁺ channels and K⁺–Cl^– cotransporters, which are activated by extracellular LPC in microglia. Furthermore, we provide evidence that non-selective cation channels and K⁺–Cl^– cotransporters are involved in the transformation of microglial cells from ramified into amoeboid morphology.

	Methods

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Cell culture

Microglia were obtained from brain cell cultures of newborn NMRI mice, supplied by Charles River (Sulzfeld, Germany). Mixed brain cell cultures were prepared as previously described (Eder et al. 1997). Brain cortices were enzymatically dissociated (15 min at 37°C with 0.125% trypsin, type XI, Sigma, Germany) and a single-cell suspension was achieved by repeated triturations. Cells were seeded into tissue culture flasks at a density of 2–4 x 10⁶ (10 ml)^–1 in Dulbecco's modified Eagle's medium (DMEM) (Gibco; Germany) supplemented with 10% heat-inactivated fetal calf serum (FCS) (Gibco; Germany) and 30% supernatant of L-929 fibroblasts as a source of macrophage colony-stimulating factor (M-CSF). After at least 10 days of incubation, microglia were harvested by shaking the cultures (30 min, 300 r.p.m) to detach weakly adherent microglial cells from the astrocytic monolayer. Isolated microglia were seeded on glass coverslips in 24 well Costar plates (3 x 10⁴ (0.5 ml)^–1). In order to induce ramification, microglial cells were cultured in astrocyte-conditioned medium (ACM) as previously described (Eder et al. 1999). Experiments were performed on microglial cells, which had been exposed to ACM for at least 3 days. The immortalized mouse microglial cell line BV-2 (kindly provided by Dr E. Blasi, Perugia, Italy) was used in some experiments. BV-2 microglial cells were cultured permanently in DMEM supplemented with 10% FCS and 2 mML-glutamine. BV-2 cells were split twice a week, and were plated on glass coverslips at a density of 3 x 10³ (0.5 ml)^–1 for subsequent patch-clamp experiments. BV-2 cells on coverslips were cultured in ACM for at least 3 days. In all experiments performed, no differences were found between cultured primary microglial cells and BV-2 microglial cells regarding their responses to LPC. Therefore, data of all cells were pooled and analysed together.

Electrophysiological recordings

Whole-cell membrane currents were measured using an EPC-9 patch clamp amplifier (HEKA, Lambrecht/Pfalz, Germany). The amplifier was interfaced to an IBM computer for pulse application and data recording. Series resistance compensation was routinely used to reduce the effective series resistance by about 70%. Patch electrodes of 2–4 M were fabricated on a two-stage puller (Narishige PP-83, Tokyo, Japan) from borosilicate glass (outer diameter 1.5 mm and inner diameter 1 mm; Hilgenberg, Malsfeld, Germany). Intracellular solution I₁ contained (mM): KCl, 120; CaCl₂, 1; MgCl₂, 2; Hepes, 10; EGTA, 11 (pH 7.3). Although no ATP or GTP is included, experiments using perforated-patch recording (Figs 6 and 7) showed identical behaviour with physiological levels of ATP and GTP. Intracellular solution I₂ contained (mM): CsCH₃SO₃, 125; MgCl₂, 2; Hepes, 10; BAPTA, 1 (pH 7.3). In some experiments, CsCH₃SO₃ was substituted by an equimolar concentration of NMGCl. The low Ca²⁺-buffered intracellular solution I₃ contained (mM): KCl, 120; MgCl₂, 2; Hepes, 10; BAPTA, 0.1 (pH 7.3). For perforated-patch-clamp experiments, the tips of the pipettes were filled with the following solution (mM): K₂SO₄, 120; KCl, 16; MgSO₄, 5; Hepes, 10 (pH 7.3). A stock solution of nystatin in DMSO (25 mg ml^–1) was prepared daily and subsequently diluted in the pipette solution to a final concentration of 150 µg ml^–1. After sonication, this solution was used for backfilling the pipettes as previously described (Horn & Marty, 1988). The extracellular solution E₁ contained (mM): NaCl, 130; KCl, 5; CaCl₂, 2; MgCl₂, 1; Hepes, 10; D-glucose, 8 (pH 7.4). In some experiments, 130 mM NaCl were substituted by 70 mM NaCl and 60 mMD-mannitol (extracellular solution E₂). In some cases, CaCl₂ was omitted from the extracellular solution, while 1 mM EGTA was added and the concentration of MgCl₂ was increased to 4 mM (Ca²⁺-free extracellular solution E₃). In some other cases, Cl^– was substituted with gluconate (Cl^–-free extracellular solution E₄). The extracellular solution E₅ contained (mM): NaCl, 140; MgCl₂, 2; Hepes, 10 (pH 7.4). All recordings were done at room temperature (20–23°C). Whole-cell currents were filtered at 3 kHz and stored on computer disk for subsequent analyses. Analyses were performed on IBM computers with the Pulse/PulseFit programme (HEKA, Lambrecht/Pfalz, Germany). Data were corrected for liquid junction potentials. Data are presented as mean values ± standard error of the mean (S.E.M.). The numbers of experiments (n) are indicated.

View larger version (19K): [in this window] [in a new window]   Figure 6.  Activation of Ca2+-dependent K+ currents by LPC BV-2 microglial cells were held at –60 mV. Voltage ramps were applied from –120 to +30 mV for a duration of 300 ms every 20 s. A–B, measurements were performed either in the whole-cell configuration using low Ca2+-buffered intracellular solution I3 (A) or in the perforated-patch-clamp configuration (B). Currents were recorded before (control) and during application of 15 µM LPC using Ca2+-containing extracellular solution E1. C, whole-cell current recordings before (control) and during application of 15 µM LPC using low Ca2+-buffered intracellular solution I3 and Ca2+-free extracellular solution E3. D, currents evoked by 15 µM LPC were recorded in the absence (LPC) and in the presence of 50 nM CTX. Whole-cell recordings were performed using low Ca2+-buffered intracellular solution I3 and Ca2+-containing extracellular solution E1.

View larger version (27K): [in this window] [in a new window]   Figure 7.  LPC-induced changes in membrane potential of BV-2 microglial cells A and B, current clamp recordings were performed using either the whole-cell patch-clamp technique with low Ca2+-buffered intracellular solution I3 (A) or the perforated-patch-clamp technique (B). Following application of 15 µM LPC, cells depolarized transiently, and membrane hyperpolarization occurred subsequently. C, LPC-induced membrane depolarization was inhibited reversibly by Gd3+ and La3+. Recordings were performed using high Ca2+-buffered intracellular solution I1 to prevent activation of Ca2+-dependent K+ channels. D, LPC-induced membrane hyperpolarization was abolished by CTX. Recordings were performed using the perforated-patch-clamp technique. At t= 0 s, the cell was exposed for 220 s to LPC. A–D, cells were superfused with Ca2+-containing extracellular solution E1.

(1)

where E_rev is the measured reversal potential, P is the permeability, R is the gas constant, T is the absolute temperature, and F is the Faraday constant.

The anion permeability of the channel was very small. Therefore, it was ignored in subsequent calculations of cation permeabilities. Next, the reversal potential of the current was obtained under various ionic conditions and the relative permeability of Na⁺, Cs⁺, K⁺ and NMG⁺ for the current was calculated in the absence of Ca²⁺ using the following equation (Hille, 2001):

(2)

where C is the test permeant cation.

The permeability of Ca²⁺ relative to Cs⁺ was calculated from the reversal potential using the following equation:

(3)

Ca²⁺ imaging

Microglial cells were loaded with 3 µM fura-2 acetoxymethylester (fura-2 AM, Molecular Probes, Eugene, OR, USA) in extracellular solution for 30 min at room temperature (20–23°C). After washing, coverslips were mounted in a chamber on an inverted Olympus IX 50 microscope (Olympus Optical Co. GmbH, Hamburg, Germany) equipped with a water immersion objective 40x UApo/340 (Olympus). The fluorescence imaging system consisted of a monochromator, a charge-coupled device (CCD) camera and the Windows NT-based image processing software (Till Photonics, München, Germany). Microglial cells were exposed to alternating 340 ± 5 nm and 380 ± 5 nm wavelengths of UV light and emission light was passed through a 400 nm dichroic mirror and a 420 nm long pass emission filter (both Olympus) prior to acquisition by the CCD camera. Images were collected every 20 s. In some experiments images were collected every 5 s. The ratio of the two background-corrected fluorescence intensities was converted to the intracellular Ca²⁺ concentration ([Ca²⁺]_i) of a single cell according to the equation of Grynkiewicz et al. (1985):

(4)

where K_d= 224 nM is the dissociation constant of fura and R is the measured ratio. R_min (0.375) and R_max (3.930) were determined using 10 µM ionomycin in extracellular solution containing zero Ca²⁺ (using 2 mM EGTA) or 10 mM Ca²⁺, respectively, and ß (5.66) was calculated as the ratio of the fluorescence intensities at 380 nm in 0 and 10 mM Ca²⁺. The extracellular solution E₁ was used for Ca²⁺ imaging experiments. Some experiments were performed using Ca²⁺-free extracellular solution E₃.

Morphometric quantification

For statistical evaluation of cell morphology, microglial cells were examined by light microscopy using an inverted Olympus IX 50 microscope (Olympus Optical Co. GmbH) equipped with a water immersion objective 40x UApo/340 (Olympus, Hamburg, Germany) and a CCD camera system (Till Photonics). For each group microglial cells from at least six different coverslips of at least three different cultures were analysed using the programme Image Tool 2.0 (UTHSCSA, San Antonio, TX, USA). Exclusively microglial cells were analysed, which exhibited ramified morphology at the beginning of experiments. Ramified microglia were defined as cells with distinct processes at least greater than one cell diameter in length (Korotzer & Cotman, 1992). Primary cultured microglial cells and BV-2 microglial cells showed a similar degree of ramification before application of LPC under the various experimental conditions. Both cell types were used in experiments studying effects of LPC, LPC + Gd³⁺ and LPC + Gd³⁺+ DIOA on microglial morphology, while the remaining experiments were performed on BV-2 microglial cells exclusively. Under the various experimental conditions, microglial morphology was unchanged 30 min after exposure to inhibitors in the absence of LPC. For quantification, a ramification index (= form factor) was calculated using the following formula (Wilms et al. 1997):

(5)

where RI is the ramification index. Ramification indices for each cell population are presented as mean values ±S.E.M. Amoeboid cells have RI values close to 1, whereas ramified cells show RI values that approach 0. During the process of data analyses experimenters were kept unaware of the experimental conditions and were not informed about any working hypotheses.

Determination of extracellular K⁺ concentration

Microglial cells were seeded in Petri dishes at a density of 5 x 10⁵ and were cultured in ACM for 3 days. Immediately before experiments, cells were washed 2 times with extracellular solution E₁. Then cells were treated with 1 ml extracellular solution containing 15 µM LPC in the absence of any inhibitor, in the presence of 100 µM Gd³⁺, or in the presence of both, 100 µM Gd³⁺ and 100 µM DIOA. After 30 min, supernatants were collected and the concentration of K⁺ was determined using an ABL 725 radiometer with a K⁺ sensitive electrode (Radiometer A/S, Bronshoj, Denmark). Data were obtained from three individual microglia cultures in three different Petri dishes per experimental condition.

Drug application

L--lysophosphatidylcholine (LPC), palmitoyl (C16: 0) was dissolved in water at a concentration of 10 mM and stored in aliquots at –20°C. Prior to experiments, aliquots were diluted in bath solution to a final concentration of 15 µM. Preparation and dilution steps were followed by 10 min sonication. In Ca²⁺ imaging experiments, drug application was achieved by complete exchange of the bath solution. In patch-clamp experiments, cells were superfused continuously with LPC-free control solution or with solutions containing LPC with and without channel inhibitors. These solutions were applied using a four-barrel microperfusion pipette, positioned at a distance of about 30–50 µm from the recorded cell to permit a rapid exchange of solutions. The flow rate was adjusted by hydrostatic pressure. The following inhibitors of ion channels and ion transporters were tested: GdCl₃ (10–100 µM); LaCl₃ (10–100 µM); ZnCl₂ (100–500 µM); amiloride (1 mM), diltiazem (100 µM); 4,4'-diisothiocyanatostilbene-2,2'-disulphonic acid (DIDS; 200 µM); 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB; 200 µM); R(+)-[(2-n-butyl-6,7-dichloro-2-cyclopentyl-2,3-dihydro-1-oxo-1H-inden-5-yl), oxy] acetic acid (DIOA; 100 µM); furosemide (1 mM); bumetanide (100 µM); Grammostola rosea brown morph (=G. spatulata) venom (1: 2000; Spider Pharm, Yarnell, USA); LOE908MS (50 µM; kind gift of Boehringer Ingelheim, Biberach, Germany); charybdotoxin (CTX; 50 nM; Latoxan, Rosana, France). If not stated otherwise drugs were obtained from Sigma, Germany.

Statistics

The statistical significance of differences between experimental groups was evaluated by Student's t test and by the Mann–Whitney U test using the SPSS programme. Data were considered to be statistically significant with P < 0.05.

	Results

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LPC-induced changes in microglial morphology

Application of LPC rapidly activated microglial cells as monitored by changes in cell morphology (Fig. 1A–C). Within the first minutes after exposure to 15 µM LPC, cell processes became thinner and were retracted partially, while cell bodies appeared to increase slightly in size (Fig. 1B). The majority of microglial cells retracted their processes completely, i.e. transformed from their ramified into amoeboid morphology, within 30 min of LPC stimulation (Fig. 1C). Changes in microglial morphology were quantified by determination of the ramification index. As demonstrated in Fig. 1D, the ramification index of microglial cells was increased significantly (P < 0.001) 30 min after treatment with 15 µM LPC, indicating a less pronounced ramification. LPC induced a similar extent of de-ramification in primary cultured microglial cells and in BV-2 microglial cells. In control experiments, we did not observe any significant changes in cell morphology and ramification index of untreated microglia held for 30 min in extracellular solution (Fig. 1D). At LPC concentrations of less than 15 µM, microglial shape changes occurred more slowly than at 15 µM LPC. In the presence of 5–10 µM LPC, the first signs of microglial de-ramification were visible about 30 min after the beginning of LPC application, while no complete transformation of microglial cells from ramified into amoeboid morphology was observed within 60 min of LPC stimulation.

View larger version (37K): [in this window] [in a new window]   Figure 1.  Effects of LPC on microglial morphology A–C, primary cultured microglial cells before (A), and 10 min (B) and 30 min (C) following exposure to 15 µM LPC. A, untreated resting microglia exhibited ramified morphology. C, microglial cells stimulated for 30 min with LPC exhibited amoeboid morphology. D, ramification index (mean ±S.E.M.) of microglia exposed for 30 min to extracellular solution in the absence (n= 85) or presence (n= 435) of 15 µM LPC. Open bars (control), mean ramification index of cells determined at the beginning of the measurements. Filled bars, mean ramification index of the same cells held for 30 min under control conditions or stimulated with 15 µM LPC.

In order to identify the physiological mechanisms underlying LPC-induced shape changes of microglial cells, patch-clamp experiments were performed. To study LPC-induced changes in microglial physiology over a time course similar to that of LPC-induced changes in microglial morphology, 15 µM LPC was used in further experiments. Figure 2A demonstrates whole-cell recordings of a microglial cell recorded with KCl-containing intracellular solution I₁ and extracellular solution E₁ (NaCl based, Ca²⁺ containing) under control conditions and following exposure to 15 µM LPC. Before application of LPC, inward and outward rectifier K⁺ currents were detected. These two types of microglial K⁺ currents have been characterized in detail elsewhere (for review, see Eder, 1998). Following extracellular application of LPC, an additional ion current activated. At –60 mV, amplitudes of the whole-cell currents increased from –14 ± 3 to –434 ± 77 pA (n= 5) in primary microglial cells, and from –7 ± 4 to –416 ± 84 pA (n= 9) in BV-2 microglial cells. Amplitudes were determined from ramp currents, because LPC-induced currents exhibited neither time-dependent activation nor inactivation (see below, Fig. 3). Furthermore, in the presence of LPC, the current reversal potential shifted in a depolarizing direction from –64.1 ± 4.2 to –4.4 ± 2.7 mV (n= 5) in primary microglial cells and from –45.7 ± 7.3 to –4.0 ± 2.0 mV (n= 8) in BV-2 microglial cells. Because neither the amplitude nor reversal potential of LPC-induced currents differed significantly (P > 0.8 in each case) between primary microglial cells and BV-2 microglial cells, data from both preparations were pooled in subsequent analyses. Figure 2B illustrates an isolated LPC-induced current, i.e. the difference current of the two currents shown in Fig. 2A.

View larger version (20K): [in this window] [in a new window]   Figure 2.  Activation of LPC-induced membrane current in BV-2 microglia Cells were held at –60 mV, and voltage ramps were applied from –90 to +60 mV for a duration of 300 ms every 10 s. Measurements were performed using K+-containing intra- and extracellular solutions (I1 and E1). A, current recordings before (control) and during application of 15 µM LPC (LPC). B, LPC-induced current isolated by subtracting LPC-control from A. C, examples of currents recorded before (1), 160 s (2) and 340 s (3) following the onset of LPC application and 180 s after washout (4) of LPC. D, amplitudes of the current shown in C were measured at –60 mV and plotted as a function of time. Current amplitudes were determined in intervals of 10 s.

View larger version (19K): [in this window] [in a new window]   Figure 3.  Kinetic properties of LPC-induced non-selective cation currents A, currents were evoked by 200 ms voltage pulses to potentials between –90 and +60 mV in 10 mV increments. Holding potential was 0 mV. Non-selective cation currents of the BV-2 microglial cell were measured using K+-free intra- and extracellular solutions. B, amplitudes of the current shown in A were plotted as a function of membrane potential. Before LPC treatment, this cell had an input resistance of 4.2 G.

Ion selectivity and kinetics of the LPC-induced membrane current

Almost identical currents were activated by LPC with K⁺-containing (Fig. 2) and K⁺-free (Figs 3 and 4) intracellular solutions, regardless of whether potassium ions were substituted by either NMG⁺ or Cs⁺. The reversal potential of LPC-induced currents measured with NMGCl in the pipette was –3.2 ± 0.6 mV (n= 5), while currents reversed at –4.2 ± 0.7 mV (n= 20) using Cs⁺-based intracellular solution I₂. These data suggest that LPC-activated ion channels were not highly selective for K⁺. Furthermore, they also suggest that LPC had little effect on pre-existent K⁺ currents, or that any such effect was outweighed by the appearance of a new conductance. LPC-induced currents were also detected after lowering the intracellular Cl^– concentration from 126 mM (Fig. 2) to 4 mM (Figs 3 and 4) (n= 26), or by equimolar substitution of extracellular Cl^– with gluconate (n= 4, data not shown), suggesting that LPC-induced currents were not carried exclusively by chloride ions. This finding is supported by the observation that extracellular application of the chloride channel inhibitors DIDS (200 µM; n= 7; data not shown) or NPPB (200 µM; n= 2; data not shown) did not have any effect on LPC-induced currents. Using CsCH₃SO₃-based intracellular solution I₂, neither the mean amplitude determined at –60 mV (–484 ± 106 pA; n= 20) nor the mean reversal potential (–4.2 ± 0.7 mV; n= 20) of LPC-induced currents differed significantly from the corresponding mean values of LPC-induced currents measured with KCl-based intracellular solution I₁ (see above).

View larger version (26K): [in this window] [in a new window]   Figure 4.  Pharmacological properties of LPC-induced non-selective cation currents Currents of BV-2 microglial cells evoked by 15 µM LPC were recorded in the absence (LPC) and in the presence of 100 µM Gd3+ (Gd3+; A), 100 µM La3+ (La3+; B), 500 µM Zn2+ (Zn2+; C), Grammostola spatulata venom, diluted 1: 2000 (G.s.; D), 100 µM diltiazem (dilt.; E), or 1 mM amiloride (amil.; F).

In order to study isolated non-selective cation currents in more detail, we used CsCH₃SO₃-based intracellular solution I₂ and Ca²⁺-free extracellular solution E₅ in further experiments. Cells were held at a holding potential of 0 mV, i.e. close to the reversal potential of the LPC-induced currents, in order to prevent continuous ion flux through LPC-activated channels and subsequent cell swelling.

To evaluate time-dependent gating of the current, hyperpolarizing and depolarizing voltage steps were applied to the cells. As shown for the example in Fig. 3, LPC-induced currents exhibited neither time-dependent activation nor inactivation at potentials between –90 and +50 mV. At +60 mV, slight current decay was detected in 3 out of 19 cells, while the remaining 16 cells did not show any time dependence at all test potentials.

Pharmacological properties of the LPC-induced non-selective cation current

Lanthanides have been reported to be effective inhibitors of non-selective cation currents in a variety of cell preparations (Sachs & Morris, 1998). In microglia, amplitude of LPC-induced currents was reduced by 61.8 ± 21.7% (n= 6) and by 91.2 ± 6.9% (n= 11) in the presence of 10 and 50 µM Gd³⁺, respectively. The currents were abolished by 100 µM Gd³⁺ in primary cultured microglia (n= 5) and in BV-2 microglial cells (n= 14) as demonstrated in Fig. 4A. La³⁺ was slightly more effective than Gd³⁺ in blocking LPC-induced cation currents. Complete current inhibition was detected at 50 µM (n= 13) and 100 µM La³⁺(n= 10) (Fig. 4B), while currents were reduced by 83.9 ± 9.5% (n= 6) in the presence of 10 µM La³⁺. LPC-induced currents were also inhibited by the divalent cation Zn²⁺, although microglial non-selective cation channels exhibited a lower sensitivity to Zn²⁺ than to Gd³⁺ and La³⁺. LPC-induced currents were reduced by 58.1 ± 12.3% (n= 4) in the presence of 500 µM Zn²⁺ (Fig. 4C), while they were not significantly affected by 100 µM Zn²⁺ (n= 4; data not shown). In some cell preparations, G. spatulata spider venom (e.g. Hu & Sachs, 1996; Suchyna et al. 2000), LOE908 (e.g. Krautwurst et al. 1993), diltiazem (e.g. Hashimoto et al. 2000) or amiloride (e.g. Lane et al. 1991; Duranton et al. 2002) have been shown to inhibit non-selective cation channels. In our study on microglia, G. spatulata spider venom induced partial inhibition of LPC-induced currents when applied at high concentration, i.e. at a dilution of 1: 2000 (Fig. 4D). On average, G. spatulata spider venom reduced LPC-induced currents by 39.2 ± 15.3% (n= 7). Microglial non-selective cation currents were unaffected by 50 µM LOE908MS (n= 7; data not shown), 100 µM diltiazem (n= 7; Fig. 4E) or 1 mM amiloride (n= 7; Fig. 4F).

LPC-induced changes in intracellular Ca²⁺ concentration and activation of Ca²⁺-dependent K⁺ currents

Bath application of 15 µM LPC elicited sustained increases in the intracellular Ca²⁺ concentration ([Ca²⁺]_i) as shown in Fig. 5A. The time course of LPC-induced [Ca²⁺]_i increases was identical to that of LPC-induced activation of non-selective cation currents. The onset of the Ca²⁺ signals was delayed by about 2 min (128 ± 3 s; n= 168). During LPC application, Ca²⁺ signals were detected in all cells investigated. Changes in [Ca²⁺]_i were exclusively caused by Ca²⁺ influx from the extracellular solution. Ca²⁺ signals were not detected following bath application of 15 µM LPC (n= 143) in experiments with Ca²⁺-free extracellular solution E₃ (Fig. 5B). Furthermore, LPC-induced [Ca²⁺]_i increases were abolished in the presence of 100 µM La³⁺(n= 53) or 100 µM Gd³⁺(n= 230) (Fig. 5C), whereas 1 µM Gd³⁺ did not affect LPC-induced Ca²⁺ signals (n= 62; data not shown).

View larger version (26K): [in this window] [in a new window]   Figure 5.  Effects of LPC on intracellular Ca2+ concentration Measurements of [Ca2+]i were performed in fura-2 AM-loaded BV-2 microglial cells. A, 15 µM LPC caused sustained increases in [Ca2+]i. B, lack of Ca2+ elevation in Ca2+-free extracellular solution. C, in the presence of 100 µM Gd3+, LPC-induced Ca2+ increases were inhibited. The right panel of the figure represents mean intracellular Ca2+ concentrations of the individual cells (n= 46 cells in A; n= 16 cells in B; n= 23 cells in C), which are shown at the left side of the figure.

LPC-induced changes in membrane potential

Next, we determined LPC-induced changes in membrane potential. Using high Ca²⁺-buffered intracellular solution I₁ (KCl based), application of LPC elicited membrane depolarization. Using low Ca²⁺-buffered intracellular solution I₃ (KCl based), a more complex pattern of membrane potential changes occurred in response to LPC stimulation. As demonstrated in Fig. 7A, LPC caused first a transient membrane depolarization, which was followed by sustained membrane hyperpolarization. Similar membrane potential changes were observed in perforated-patch-clamp recordings, as shown for the example in Fig. 7B. The onset of the LPC-induced membrane depolarization was delayed on average by 75 ± 15 s (n= 9) in whole-cell recordings using low Ca²⁺-buffered intracellular solution I₃ and by 86 ± 39 s (n= 3) in perforated-patch-clamp recordings. Both values did not differ significantly from each other and from the delay in the onset of the LPC-induced cation current activation and Ca²⁺ increases (see above). In whole-cell recordings with low Ca²⁺-buffered intracellular solution I₃ we determined a mean resting membrane potential of –45.6 ± 7 mV (n= 4) before application of LPC and mean values of –17 ± 5.7 and –70.9 ± 1.9 mV (n= 4) for LPC-induced membrane depolarization and hyperpolarization, respectively. In perforated-patch-clamp measurements, cells had a mean resting membrane potential of –40.3 ± 10.7 mV (n= 3), depolarized to a mean value of –11.2 ± 2.5 mV and hyperpolarized to –68 ± 2.2 mV (n= 3). Mean values of resting membrane potential, LPC-induced depolarization and LPC-induced hyperpolarization did not differ significantly between whole-cell and perforated-patch recordings. LPC-induced depolarization could be reversed by 100 µM Gd³⁺(n= 4) or 100 µM La³⁺(n= 4) (Fig. 7C), while membrane hyperpolarization was abolished by 50 nM CTX (n= 4) (Fig. 7D).

Non-selective cation channels but not Ca²⁺-dependent K⁺ channels are important for LPC-induced morphological changes in microglia

To clarify whether LPC-induced de-ramification was due to increases in [Ca²⁺]_i following Ca²⁺ influx through non-selective cation channels, Ca²⁺ was omitted from the extracellular solution in some experiments. LPC caused almost identical changes in microglial morphology in measurements with Ca²⁺-free and Ca²⁺-containing extracellular solutions. The ramification index of microglial cells treated with LPC in Ca²⁺-free external solution was slightly larger than that of cells in Ca²⁺-containing solution (Fig. 8A). These data implied that changes in microglial morphology were not Ca²⁺-dependent.

View larger version (27K): [in this window] [in a new window]   Figure 8.  Involvement of non-selective cation channels and K+–Cl– cotransporters in LPC-induced microglial de-ramification A, partial inhibition of LPC-induced microglial de-ramification by blockers of either non-selective cation channels or K+–Cl– cotransporters. Ramification index (mean ±S.E.M.) of untreated microglia (n= 85) and of microglia exposed for 30 min to LPC in Ca2+-containing (n= 435) or Ca2+-free (n= 150) solution, or in the presence of 100 µM Gd3+(n= 258), 100 µM La3+(n= 176), 1 mM furosemide (n= 116) or 100 µM DIOA (n= 118). B, complete inhibition of LPC-induced microglial de-ramification by combined application of blockers of non-selective cation channels and blockers of K+–Cl– cotransporters. Ramification index (mean ±S.E.M.) of untreated microglia (n= 85) and of microglia exposed for 30 min to LPC and 100 µM Gd3+ in the absence (n= 258) or in the presence of 1 mM furosemide (n= 148), 100 µM DIOA (n= 166), 100 µM bumetanide (n= 244), or in Cl–-free extracellular solution (n= 181). C, example of primary cultured microglial cells exposed to 100 µM Gd3+ and 100 µM DIOA before (C1) and 30 min after (C2) application of 15 µM LPC.

In further experiments, we investigated whether inhibition of non-selective cation channels influences LPC-induced shape changes. As demonstrated in Fig. 8A, the ramification index of LPC-treated microglial cells was significantly (P < 0.001) smaller in the presence of 100 µM Gd³⁺ than in the absence of Gd³⁺, suggesting that the activity of non-selective cation channels is necessary for LPC-induced morphological changes. Partial inhibition of LPC-induced de-ramification by Gd³⁺ was observed in both primary cultured and BV-2 microglial cells. Almost identical observations were made when non-selective cation currents were abolished by 100 µM La³⁺ (Fig. 8A).

Functional importance of K⁺–Cl^– cotransporters for LPC-induced changes in microglial morphology

Although 100 µM Gd³⁺ or 100 µM La³⁺ abolished non-selective cation currents, neither Gd³⁺ nor La³⁺ inhibited LPC-induced de-ramification of microglia completely. Therefore, it was investigated whether electroneutral ion transporters were involved additionally in microglial shape changes. As shown in Fig. 8A, inhibition of K⁺–Cl^– cotransporters leads to partial inhibition of microglial de-ramification. Thirty minutes after exposure to LPC, the mean ramification indices of cells treated with either 1 mM furosemide or 100 µM DIOA were significantly smaller than that of cells held in the absence of K⁺–Cl^– cotransporter inhibitors. Figure 8B demonstrates that combined application of 100 µM Gd³⁺ and 1 mM furosemide prevented LPC-induced transformation of microglial morphology from the ramified to the amoeboid form. LPC-induced de-ramification also did not occur in the presence of 100 µM Gd³⁺ and 100 µM DIOA in either primary cultured microglia or BV-2 microglia (Figs 8B and C). The mean ramification index of these cell populations was not significantly different from that of untreated cells (Fig. 8B). In contrast, combined application of 100 µM Gd³⁺ and 100 µM bumetanide failed to prevent microglial de-ramification. LPC-induced shape changes were also not abolished by Gd³⁺ in Cl^– free extracellular solution (Fig. 8B).

To verify the activity of K⁺–Cl^– cotransporters we determined the extracellular K⁺ concentration ([K⁺]_o) of LPC-treated microglia cultures in the absence and presence of DIOA. We determined a mean [K⁺]_o of 7.2 ± 0.3 mM(n= 4) for LPC-stimulated cultures 30 min after exposure of microglia to LPC. In the presence of 100 µM Gd³⁺, [K⁺]_o of LPC-stimulated cell cultures was 6.9 ± 0.2 mM(n= 4). During combined application of Gd³⁺ and DIOA, [K⁺]_o of LPC-stimulated cultures was significantly smaller (P < 0.05) than that of cultures not exposed to DIOA. A mean [K⁺]_o of 6.4 ± 0.1 mM(n= 4) was determined for LPC-stimulated microglial cultures exposed to both 100 µM Gd³⁺ and 100 µM DIOA. These data suggest that K⁺–Cl^– cotransporters contribute to K⁺ efflux from LPC-stimulated microglial cells.

	Discussion

Top Abstract Introduction Methods Results Discussion References

 
In this study we demonstrate for the first time that in microglia non-selective cation channels, Ca²⁺-dependent K⁺ channels and K⁺–Cl^– cotransporters are activated by LPC. Furthermore, we demonstrate the functional importance of non-selective cation channels and K⁺–Cl^– cotransporters for microglial de-ramification.

Physiological concentrations of LPC are in the range of 5–180 µM (Croset et al. 2000; Xu, 2002), varying in different tissues and body fluids. In pathophysiological situations, such as ischaemia, LPC may reach concentrations of up to 200 µM (Akita et al. 1986; Sedlis et al. 1990). Assuming that 90% of LPC is protein bound, it has been suggested that the free concentration of LPC reaches 20 µM in ischaemic tissue (Yu et al. 1998). In microglia, physiological concentrations of LPC evoke membrane depolarization and sustained Ca²⁺ increases by activating non-selective cation channels. Activation of non-selective cation currents in response to LPC has been reported in ventricular myocytes (Magishi et al. 1996;

Hashimoto et al. 2000; Song & Ochi, 2002), arterial smooth muscle cells (Jabr et al. 2000; Terasawa et al. 2002) and endothelial cells (Yokoyama et al. 2002). LPC-activated non-selective cation channels of microglia have similarities with those of other cell preparations. They activated after some delay, were permeable for monovalent and divalent cations and could be inhibited by lanthanides. Diltiazem has been shown to inhibit LPC-induced currents of ventricular myocytes (Hashimoto et al. 2000), but failed to block microglial currents. Grammostola spatulata spider venom had inhibitory effects on LPC-induced currents in microglia similar to those on stretch-activated cation currents in astrocytes and heart cells (Hu & Sachs, 1996; Suchyna et al. 2000). Based on our observations that LPC-induced currents were inhibited by Zn²⁺ but unaffected by amiloride, it can be excluded that microglial channels are related to the amiloride-sensitive ENaC/DEG channel superfamily (Kellenberger & Schild, 2002). LPC-activated cation currents of microglial cells correspond in some of their properties to the currents that are activated by sustained depolarization, ADP-ribose or ßNAD⁺ in peritoneal macrophages (Campo et al. 2003). Moreover, microglial non-selective cation channels resemble closely stretch-activated/mechanosensitive cation channels described in a variety of cell preparations (Sachs & Morris, 1998). Perozo et al. (2002) have recently elucidated the mechanism by which LPC causes activation of cation channels. According to the authors' model, LPC produces local stress due to its insertion into the outer monolayer of the plasma membrane. By sensing this stress, mechanosensitive channels undergo conformational changes and open subsequently (Perozo et al. 2002; Sachs, 2002). Most likely, microglial cation channels are activated by the same mechanism. However, it cannot be excluded that LPC receptor-coupled intracellular signalling pathways modulate channel activity additionally.

We suggest that LPC-induced sustained Ca²⁺ increases are mediated by activation of non-selective cation channels. It can be excluded that Ca²⁺ release-activated Ca²⁺ (CRAC) currents contribute to the LPC-induced Ca²⁺ signal, because Gd³⁺ did not affect LPC-induced Ca²⁺ increases at a concentration of 1 µM, which is sufficient to block CRAC channels completely (Ross & Cahalan, 1995). The LPC-induced Ca²⁺ signal appears to consist of two components. The first transient component of LPC-induced Ca²⁺ signals may reflect swelling-activated Ca²⁺ (SWAC) transients, which have been described first in thymocytes (Ross & Cahalan, 1995), and can also be elicited in microglia by hyposmotic extracellular solutions (T. Schilling, C.Eder, unpublished observations). However, it is also possible that the LPC-induced Ca²⁺ signal decreased to some extent after reaching the initial peak due to Ca²⁺ extrusion mechanisms. Further experiments are required to test whether one of these hypotheses can explain the two components of the LPC-induced Ca²⁺ signal.

Furthermore, we provide evidence that a Ca²⁺-dependent K⁺ current is activated secondary to LPC-induced Ca²⁺ entry through non-selective cation channels. We have shown previously that Ca²⁺-dependent K⁺ currents of murine microglia are voltage independent, sensitive to inhibitors of intermediate conductance (IK) channels, such as charybdotoxin and clotrimazole, but insensitive to an inhibitor of small conductance (SK) channels, apamin, and to an inhibitor of large conductance (BK) channels, paxilline. It was also demonstrated that the channel is encoded by the gene IKCa1 (Eder et al. 1997; Schilling et al. 2002, 2004). The Ca²⁺-dependent K⁺ current evoked by LPC was identical to the IKCa1 Ca²⁺-dependent K⁺ current described before in murine microglia. Similar to IKCa1 currents, LPC-evoked Ca²⁺-dependent K⁺ currents exhibited voltage-independent activation behaviour, reversed at potentials close to the K⁺ equilibrium potential and were inhibited by charybdotoxin. K⁺ current activation did not occur by using high Ca²⁺-buffered intracellular solution or Ca²⁺-free extracellular solution, suggesting that increases in the intracellular concentration of Ca²⁺ were responsible for current activation. Importantly, LPC-activated Ca²⁺-dependent K⁺ currents caused membrane hyperpolarization, which occurred secondary to the initial depolarization that was evoked by non-selective cation currents. We propose that the membrane hyperpolarization that results from Ca²⁺-activated K⁺ channels increases the driving force for Na⁺ and Ca²⁺ entry through non-selective cation channels. Thus, Ca²⁺-dependent K⁺ channels help maintain substantial cation influx.

In our experiments, microglial cells transformed almost completely from a ramified to an amoeboid morphology following stimulation with LPC. Intriguingly, microglial de-ramification occurred independently of intracellular and extracellular Ca²⁺. This observation appears to be in contrast to some reports linking cytoskeletal reorganization to increases in intracellular Ca²⁺ (e.g. Onuma & Hui, 1988; Li et al. 2001). LPC-induced morphological changes of microglial cells were partially inhibited in the presence of Gd³⁺, La³⁺, furosemide or DIOA, suggesting involvement of non-selective cation channels and K⁺–Cl^– cotransporters in microglial de-ramification. Microglial shape changes were prevented by combined application of Gd³⁺ and either furosemide or DIOA. This observation led us to conclude that simultaneous activity of both non-selective cation channels and K⁺–Cl^– cotransporters is required for microglial de-ramification. At a concentration of 100 µM, DIOA specifically inhibits K⁺–Cl^– cotransporters without affecting Na⁺–K⁺–2Cl^– cotransporters (Garay et al. 1988). Although furosemide is capable of inhibiting both K⁺–Cl^– cotransporters and Na⁺–K⁺–2Cl^– cotransporters (Lauf & Adragna, 2000; Russell, 2000), our data argue against a contribution of Na⁺–K⁺–2Cl^– cotransporters. Specific inhibition of Na⁺–K⁺–2Cl^– cotransporters either by using Cl^–-free extracellular solution or by application of bumetanide failed to inhibit LPC-induced morphological changes. Further experiments are required to clarify whether K⁺–Cl^– cotransporters are activated by LPC directly or secondarily to other LPC-induced processes. It has been demonstrated for other cell preparations that K⁺–Cl^– cotransporters can be activated by cell swelling, intracellular acidification, reactive oxygen radicals, or intracellular signalling processes, such as protein phosphorylation (Lauf & Adragna, 2000).

In contrast to non-selective cation channels and K⁺–Cl^– cotransporters, Ca²⁺-dependent K⁺ channels appear to play only a minor role in regulating microglial shape changes. In experiments at which Ca²⁺-dependent K⁺ channel activation was prevented using Ca²⁺-free extracellular solution, LPC-induced microglial de-ramification was not inhibited. In contrast, under these conditions de-ramification was slightly more pronounced than in experiments using Ca²⁺-containing solutions. It is possible that Ca²⁺-dependent K⁺ channels control the amount of LPC-induced cell body swelling by counteracting the activity of non-selective cation channels. Most likely, osmolarity and subsequent swelling of the cell body is reduced to some extent due to K⁺ efflux through Ca²⁺-dependent K⁺ channels. Loss of up to 20% of the cell volume was attributed to the activity of IKCa1 Ca²⁺-dependent K⁺ channels in other cell preparations (Schneider et al. 2000).

We hypothesize that rearrangement of the cytoskeleton following exposure of microglia to LPC is accompanied by changes in osmolarity. To explain the physiological mechanisms underlying LPC-induced microglial de-ramification, we propose the following model (Fig. 9): Extracellular application of LPC leads to simultaneous activation of non-selective cation channels and K⁺–Cl^– cotransporters. We suggest that the expression of cation channels is higher in cell bodies than in cell processes, while K⁺–Cl^– cotransporters are predominantly expressed in cell processes. Cation influx through non-selective cation channels increases the osmolarity inside the cell. The subsequent osmotic entry of water results in swelling of cell bodies. In contrast, KCl efflux due to activity of K⁺–Cl^– cotransporters reduces osmolarity in the processes. Subsequently, water leaves cell processes causing their shrinkage and retraction. These two opposing mechanisms occur simultaneously and lead finally to complete retraction of cell extensions and increased size of cell bodies, i.e. the cells exhibit amoeboid morphology (Fig. 9).

View larger version (17K): [in this window] [in a new window]   Figure 9.  Proposed model of physiological mechanisms of microglial de-ramification induced by LPC (see Discussion)

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