The FASEB Journal express article 10.1096/fj.00-0439fje. Published online March 12, 2001.
Astrocytes of the mouse neocortex express functional Nmethyl-D-aspartate receptors Carola G. Schipke+, Carsten Ohlemeyer+, Marina Matyash, Christiane Nolte, Helmut Kettenmann, and Frank Kirchhoff* Max Delbrück Center for Molecular Medicine, Cellular Neurosciences, Robert-Rössle-Straße 10, D-13092 Berlin-Buch, Germany * Max Planck Institute for Experimental Medicine, Neurogenetics, Hermann-Rein-Str. 3, D37075 Göttingen, Germany Corresponding author: Frank Kirchhoff, Max Planck Institute for Experimental Medicine, Neurogenetics, Hermann-Rein-Str. 3, D-37075 Göttingen, Germany. E-mail:
[email protected] + Both authors contributed equally. ABSTRACT In the brain, N-methyl-D-aspartate (NMDA)-type glutamate receptors are important elements for the manifestation of memory as well as mediators of neurotoxicity, and they are thought to be exclusive to neurons. To test for the expression of functional NMDA receptors on astrocytes, we generated transgenic mice in which glial fibrillary acidic protein (GFAP)-positive astrocytes are labeled by a green fluorescent protein and tested their responses to NMDA in acute cortical slices by patch-clamp recording and Ca2+ imaging. The NMDA-evoked currents reversed at 0 mV; could be blocked by MK-801; persisted in the absence of synaptic transmission; were sensitive to Mg2+; and were accompanied by focal Ca2+ elevation, indicating the presence of functional NMDA receptors. Furthermore, we detected mRNAs for NMDA receptor subunits in freshly isolated astrocytes purified by fluorescence-activated cell sorting. We conclude that processes of cortical astrocytes enwrapping synaptic regions express high densities of NMDA receptors that could be involved in neurone-glia signaling. Key words: NMDA receptor • astrocyte • glia • GFAP promoter • enhanced green fluorescent protein • patch-clamp recording • calcium imaging • mouse •ÜFRUWH[
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strocytes are in intimate contact with neurons in gray matter, particularly at synapses. Several recent studies indicate that there is an intense cross-talk between these two cell types. The astrocytes express the prerequisite to participate in neuronal communication (i.e., receptors) for a variety of neurotransmitters and –hormones (1). Within neurons the information is propagated by electrical activity, but the astrocytic form of intracellular signal propagation is dominated by Ca2+ signaling (2). These increases in cytosolic calcium can affect neuronal pathways (3). The first indications for this astrocyte-neuronal cross-talk came from cell
cultures (4), whereas recent studies demonstrate that such signal exchanges can also occur in situ. Studies in the isolated retina, for instance, illustrate that astrocytic calcium signals lead to modulation of light-induced excitation of ganglion cells (5). Glutamate receptors seem to play a key role in this neuron-glia interaction, as has been shown for the hippocampus (6). Numerous glutamate-binding molecules, such as glutamate transporters, the non-N-methyl-DDVSDUWDWH.-amino-3-hydroxy-5-methylisoxazole-4-propionic acid (NMDA/AMPA) / kainatetype glutamate receptors, and metabotropic glutamate receptors, have been identified on astrocytes. For neuron-glia signaling via transmitter receptors, the non-NMDA receptors are the prominent communication channels, whereas there is little evidence for the involvement of NMDA receptors in astrocytes. Studies in cultured astrocytes have long demonstrated that these cells are unresponsive to NMDA but are prominently activated by non-NMDA agonists (7). Despite unanimous agreement that cultured astrocytes are devoid of functional NMDA receptors, there is some evidence that NMDA can elicit signals in astrocytes in situ. Bergmann glial cells respond to NMDA with an inward current, but this response does not reflect features of NMDA receptors because the response is Mg2+-insensitive (8). In addition, the Bergmann glia NMDA response was frequently inhibited by tetrodotoxin, which suggests that the NMDA action is mediated by indirect mechanisms (9). Similar Mg2+-insensitive responses were detected in early postnatal astrocytes from spinal cord slices (10). The astrocytes, however, are only responsive until 2 weeks after birth, whereas most astrocytes in the young adult animals did not respond. Although we lack functional data on astrocytic NMDA receptors, several reports indicate the presence of NMDA receptor subunits (NR) in glial membranes. NR1 immunoreactivity can be found in astrocytic processes close to dendrites and axons in the nucleus accumbens (11), the amygdala (12), and the cat visual cortex (13, 14). Some researchers could not detect NR1 labeling in glial cells of the cortex (15), whereas others described labeling for NR1 and NR2A/B subunits confined to distal processes with features of astrocytes (16). There is also some evidence for the expression of the NMDA receptor mRNA by in-situ hybridization of brain sections, but it seems confined to a very small population of astrocytes (17, 18). The main concern with the immunohistochemical studies, however, is the proper identification of labeled membrane as astrocytic. The classical marker of astrocytes, glial fibrillary acidic protein, is confined to main processes and is not present in the fine appendages that are at the contact site of astrocytes and synapses. Furthermore, the presence of mRNA or even protein does not prove the presence of functional receptors. We have therefore developed a new tool to unequivocally identify astrocytes: a transgenic mouse line in which GFAP-positive cells are labeled by the enhanced green fluorescent protein (19). Through this work we determined whether this population of cells has intrinsic responses to NMDA. Moreover, we were able to use fluorescent-activated cell sorting to purify astrocytes from the brain and to analyze for the presence of NMDA receptor mRNAs. By combining these approaches, we have demonstrated that NMDA elicits intrinsic signals in astrocytes that are mediated by NMDA receptor activation, and that these receptors are confined to distal processes. MATERIALS AND METHODS Generation of transgenic mice
We generated the construct for microinjection of FVB/N oocytes by cloning the 2.2-kB fragment of human GFAP promoter (20–22) 5´ to the enhanced green fluorescent gene enhanced green fluorescent protein (EGFP) (Clontech, Heidelberg, Germany). Functional expression of the transgene was tested in the GFAP-positive human anaplastic glioma cell line U343 MG. Oocytes were injected using conventional microinjection technology by Eurogentec, Seraing, Belgium. Founder lines were mated with nontransgenic FVB/N mice. We will describethe characterization of the GFAP/EGFP transgenic mice elsewhere (19). Already, heterozygous offspring displayed a robust bright EGFP expression in astrocytes of all brain regions visible by conventional fluorescence microscopy. Preparation of brain slices and electrophysiological setup One- to four-week-old mice (either GFAP/EGFP transgenic FVB/N mice or nontransgenic NMRI mice) were decapitated and their brains were dissected and washed, after which the hemispheres were cut into 150- P-thick slices in frontal orientation using a vibratome (FTB, Plano, Marburg, Germany). Slice preparation was performed at 6°C in calcium-free bath solution (see below). Subsequently, slices were stored for at least 30 min in bath solution at room temperature. For electrophysiological recordings, slices were placed in a chamber mounted on the stage of a Zeiss microscope (Axioplan, Zeiss, Oberkochen, Germany) and fixed in the chamber using a U-shaped platinum-wire with a grid of nylon threads (23). The chamber was continuously perfused with carbogen-saturated bath solution, and substances were added by changing the perfusate. Cell somata in the cortex were visible with water immersion optics (Zeiss Achroplan 40×) and could be approached by the patch electrode. The selected cells were located about 10– PEH\RQGWKHVXUIDFHRIWKHVOLFH3RVLWLYHSUHVVXUH was applied to the recording pipette while it was lowered under microscopic control. Thus, cellular debris was blown aside and the tip could be placed onto the surface of a cell soma. Membrane currents were measured with the patch-clamp technique in the whole-cell recording configuration (24). Current signals were amplified (EPC-7 and EPC-9 amplifier, HEKA, Lambrecht, Germany), filtered at 3 kHz, and sampled at 5 kHz by an interface (HEKA) connected to a PC system that also served as a stimulus generator. All patch-clamp data analysis was performed using the WinTida software package (HEKA). The resistance of the patch pipettes was 5-6 M /LPLWDWLRQV RI YROWDJH FODPS FRQWURO LQ FHOOV RI VOLFH SUHSDUDWLRQV DUH discussed in detail elsewhere (25). Solutions and electrodes The standard bath solution contained 134 mM NaCl, 2.5 mM KCl, 2 mM CaCl2, 1.3mM MgCl2, 26 mM NaHCO3, 1.25 mM K2HPO4, 10 mM glucose, and 2 mg/l phenol red as a pH indicator. For calcium recordings the CaCl2 concentration was set to 5 mM. Mg2+-free solution was used when indicated. By gassing the solutions with carbogen (5% CO2/95% oxygen), the pH was adjusted to 7.4–7.5. The pipette solution contained 130 mM KCl, 0.5 mM CaCl2, 3 mM MgCl2, 5 mM EGTA, and 10 mM HEPES. Ca2+ activity of the pipette solution was approximately 11 nM. All experiments were carried out at room temperature (about 22°C). Recording pipettes were fabricated from borosilicate capillaries (Hilgenberg, Malsfeld, Germany). To test for
neurotransmitter-evoked responses, transmitters as well as their agonists and antagonists were applied to the bath solution in the following concentrations (in µM): NMDA 100, kainate 100, Ltrans-pyrollidine-2,4-dicarboxylic acid (PDC) 50, 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX) 20, Cd2+ 100, (+)MK-801 20, tetrodotoxin (TTX) 1-5, dihydrokainate 200, and D,L-threohydroxyaspartate 20. Labeling of cells for morphological analysis The morphology of green fluorescent astrocytes was visualized by confocal laser scanning microscopy (CLSM, Sarastro 2000, Molecular Dynamics, Sunnyvale, Calif.); the scanning unit was mounted on an upright microscope (Axioskope, Zeiss Jena, Germany) equipped with water immersion optics (40×, 0.75 aperture). Glial cells were fluorescent due to either GFAP-driven EGFP expression in the case of transgenic animals or intracellular dialysis with 0.1% Lucifer Yellow-CH lithium salt (LY, Fluka, Seelze, Germany) (added to the pipette solution). During recording, cells were filled with LY by dialyzing the cytoplasm with the patch pipette solution. At the end of the recording, large hyperpolarizing (LY) current injections were applied to enhance cell loading. For further histochemical analysis, slices were fixed in phosphate buffer (PB, 100 mM sodium phosphate pH 7.4) containing 4% paraformaldehyde for 2 h up to overnight at 5°C. The slices were washed in PB for 1 h and mounted with Mowiol (Hoechst, Frankfurt, Germany). CLSM images were stored and processed on an Indigo workstation using the program Imagespace. Calcium recordings For Ca2+ recordings in GFAP/EGFP transgenic and nontransgenic mice, Calcium orange (200 µM) or Fluo-4 (100 µM) were added to calcium/EGTA free pipette solution, respectively. After 10 min of dialysis, cells were sufficiently filled with the indicator dye. Agonist-evoked Ca2+ transients were recorded by CLSM. Before, during, and after agonist application, images of 256 × 256 pixels were recorded every 4–6 s, then stored and processed on an Indigo workstation using the program Imagespace. Fluo-4 and Calcium orange were excited at 488 or at 514 nm, and emission was detected with 510-nm or 600-nm long pass filters, respectively. Ca2+ concentration changes are shown as fluorescent intensity ratios F/F0. The resting fluorescence value F0 was determined at the beginning of each experiment. Immunohistochemistry Mice were deeply anesthetized with sodium pentobarbital (100 mg/kg body weight; Sanofi, Paris, France), and perfused intracardially with a solution of 4% paraformaldehyde in PB. Brains were dissected and postfixed for 2 h at 4°C. After two washes in PB, brains were incubated overnight in 30% sucrose in PB. Brains were quickly frozen in isopentane cooled by dry ice. Cryosections (20 µm thick) were mounted on gelatin-coated slides and allowed to dry for at least 30 min at room temperature. Sections were permeabilized with 0.1% Triton X-100 (TX100) in PB for 20 min and incubated in blocking buffer (0.5% BSA, 1% horse serum, 4% normal goat serum, 0.01% TX100 in PB) for 1 h at room temperature. Polyclonal rabbit anti-GFAP antibodies (Boehringer, Mannheim, Germany) were diluted 1:1000 (in PB/1% BSA/1% horse serum) and incubated with the sections overnight at 4°C. Primary antibodies were detected by
application of Cy3-conjugated goat antirabbit IgG (Jackson/Dianova, Hamburg, Germany). Secondary antibodies were used at a dilution of 1:250 and incubated for at least 2 h at room temperature. After three washes, sections were mounted with Mowiol and inspected in the confocal microscope. The specificity of immunoreactivity was controlled by incubation of tissue sections in dilution buffer instead of primary antibodies. In these control experiments, the immunocytochemical reactions were always negative. Fluorescence-activated cell sorting (FACS) Brains were removed from 2-week-old transgenic GFAP/EGFP and wild-type mice. Meninges were removed with fine forceps. Tissues were minced into small pieces and incubated for 30–45 min at 37°C in Hank’s balanced salt solution (HBSS) containing papain (26) (Worthington, Freehold, N.J.). Subsequently, tissue pieces were triturated gently in Dulbecco’s modified Eagle medium (DMEM; Life Technologies, Grand Island, N.Y,) containing 10% fetal calf serum (FCS; Life Technologies) in the presence of DNAse (0.5 mg/ml; Sigma, Diesenhofen, Germany). Cells – were centrifuged at 800 min 1 in a tabletop centrifuge (Eppendorf centrifuge 5034) for 10 min and resuspended in PBS. For flow cytometry, cells were resuspended in DMEM/FCS (0.5–1 × 6 10 FHOOV O DQG VRUWHG XVLQJ D )$&6FDQ IORZ F\WRPHWHU LQ FRPELQDWLRQ ZLWK WKH CELLQuest software (Becton Dickinson, Bedford, Mass.). Dead cells were excluded from 4 analysis by electronic gating. In a typical experiment, approximately 1 × 10 live EGFP-positive cells per animal could be isolated. RT-PCR analysis of FACS-isolated astrocytes Total RNA obtained from FACS-sorted astrocytes or from total brain of 3- to 4- week-old transgenic and nontransgenic mice was prepared using the Trizol method (LifeTechnologies, Eggenstein, Germany) (27). cDNAs were prepared using Superscript RT™ (LifeTechnologies) and amplified in a Thermocycler 9600 (Applied Biosystems, Weiterstadt, Germany) by hot-start PCR (5 min, 94°C) in 35–40 cycles (30 s, 94°C denaturation; 30 s, 55–60°C annealing; 30 s + 1 s per cycle, 72°C elongation) and 10 min at 72°C for elongation. Subsequently, the tubes were cooled to 4°C and the products analyzed by agarose gel electrophoresis. Primers for mouse NMDA receptor genes, GFAP, tyrosine hydroxylase, and EGFP were derived from the published sequence delivered to the EMBL database (accession numbers in brackets): mouse GFAP (K01347), sense: 5´- TCC TCC GCC AAG CCA AAC ACG A-3´, antisense: 5´-CCT CAC CAT CCC GCA TCT CCA CA-3´, 438 bp; tyrosine hydroxylase (M69200), sense: 5´-TGT CAG AGC AGC CCG AGG TC-3´, antisense: 5´-CCA AGA GCA GCC CAT CAA AG-3´, 412 bp; EGFP (CV55761), sense: 5´-GCC GCT ACC CCG ACC AC-3´, antisense: 5´-TTC ACC TTG ATG CCG TTC TTC T-3´, 272 bp; NR1 (D10028), sense: 5´-CTG CTG ACA TTC GCC CTG CTT -3´, antisense: 5´-ACG CGC ATC ATC TCA AAC CAG AC -3´, 455 bp; NR2A (D10217), sense: 5´-AGT TTC CAT CGG GTC TCA TTT CA -3´, antisense: 5´-ACT TCG CCT ATC ATT CCA TTC CA -3´, 688 bp; NR2B (D10651), sense: 5´-TCA CCC CCT TTC CGC TTT G 3´, antisense: 5´-GTT CTT CCA TCT CCC CAT CTC CA -3´, 343 bp; NR2C (D10694), sense: 5´-CCT TCT TCG CTG TCA TCT TCC TC -3´, antisense: 5´-TGC TGC TCA TCA CCT CGT TCT TC -3´, 528 bp; NR2D (D12822), sense: 5´-CCT GGG GGA CGA TGA GAT TGA GA 3´, antisense: 5´-GAG CGG GCG GAG ATG GAA AG -3´, 662 bp; rat NR3 (U29873), sense:
5´-TGT CTG CTA TGC CCT TCT GTT TG -3´, antisense: 5´-CTG GTT TTG TCC TTC CTC GTC A -3´, 949 bp. The primer pairs were selected to cross exon borders distinguishing genomic contaminations. Identities of amplimers derived from NMDA receptor messages were confirmed by DNA sequencing carried out in either sense or antisense direction using the Taq DyeDeoxy Terminator Cycle sequencing kit on an automated ABI DNA sequencer (model 373A, Applied Biosystems, Weiterstadt, Germany). RESULTS Isolated astrocytes express NMDA receptor gene products We have generated a transgenic mouse line in which expression of the red-shifted variant of the green fluorescent protein EGFP was controlled by the GFAP promoter to unequivocally label astrocytes. To isolate a pure population of astrocytes from the brain, we prepared single-cell suspensions from cortices of 2-week-old mice and purified astrocytes by fluorescence-activated cell sorting. To test for the presence of mRNAs for NMDA receptors, we sorted those cells displaying a fluorescence that was at least 50 times brighter than cells from nontransgenic animals. Subsequently, cellular RNA was isolated, reverse transcribed, and probed for NMDA receptor gene activity by PCR. We obtained significant amplification signals for NR1, NR2B, and C (Fig. 1). Neuronal contamination of the RNA could be excluded because PCR with primers for the neurone-specific tyrosine hydroxylase did not reveal any amplification product. As a control, we obtained a clear signal for neurone-specific tyrosine hydroxylase with RNA derived from total mouse brain. Acutely isolated astrocytes could yet not be harvested in sufficient amounts to allow the analysis of protein expression by Western blot. NMDA triggers a current response in identified protoplasmic astrocytes To test for the presence of functional NMDA receptors in astrocytes, we made acute slices from the GFAP/EGFP transgenic mice. We studied EGFP-positive cells primarily in the cingulate, retrosplenial, and parietal cortical areas of coronal slices from the forebrain prepared from 1- to 4-week-ROGPLFH6PDOOVRPDWDRIDERXW PVHYHUDOSURFHVVHVZLWKDEXVK\DSSearance, and contacts of endfeet to blood vessels were the morphological features of the green fluorescent cells (Fig. 2A). In addition, GFAP immunostaining labeled the population of EGFP-positive cells (Fig. 2D–F). By comparing EGFP-fluorescence and phase contrast images, individual cells were approached with the patch pipette to establish the whole-cell recording mode. Depolarizing and hyperpolarizing voltage steps (+/– 90 mV in 10-mV steps from the holding potential of –80 or – 70 mV) elicited passive currents with a linear current to voltage (I/V) relationship (Fig. 2B). The membrane potential was –71.8 mV +/– 6.4 mV (n = 133). Thus, the EGFP-positive cells have the characteristic physiological features of astrocytes. NMDA triggered an inward current in most EGFP-positive cells (96 out of 133; Fig. 2C). The NMDA (0.1 mM)-induced current responses ranged from 11 to 814 pA (mean 197 +/– 190 pA). Studying cells with similar morphological features in brain slices of 3- to 4-week-old nontransgenic NMRI mice revealed similar results: 40 out of 54 cells exhibited NMDA
responses in the range of a few hundred pA (mean 234 +/– 182 pA, range 17–718 pA). In addition, astrocytes in nontransgenic FVB/N mice reacted upon NMDA application. We found a large variation in the NMDA current amplitudes in all the mouse strains we studied. MK-801 is a specific noncompetitive open channel blocker of NMDA receptors. Indeed, 1 µM MK-801 almost completely abolished the NMDA-induced current of astrocytes, to 7% as compared with a control (15 pA under MK-801 vs. 211 pA of the control, n = 7, Fig. 3A). This blockade of MK-801 was irreversible, because we were not able to elicit another NMDA response after washout of the blocker for 15–30 min. NMDA responses in astrocytes are influenced by neurones Application of NMDA to brain slices will stimulate all NMDA receptors, those that are profoundly described on neurons and the potential ones in astrocytes. While there is no doubt that the recorded NMDA responses originate from currents across the membrane of astrocytes, we cannot exclude the fact that they are due to indirect effects triggered after activation of neuronal NMDA receptors and stimulated neuronal activity. We therefore used pharmacological tools to block synaptic transmission, action potentials, and glutamate uptake. Cd2+ (100 µM), which is known to block presynaptic Ca2+ channels and thereby inhibit calciumdependent transmitter release, reduced the NMDA-evoked current to 23% of the control value (n=4). Similarly, blocking action potentials by tetrodotoxin (1–5 µM) reduced the response to about 45% as compared with the control (n=2). In the presence of a cocktail of all the blockers (i.e., TTX, PDC, Cd2+, and CNQX), NMDAevoked responses were reduced to about 10–25% as compared with a control, but still present (Fig. 3B). Therefore, we conclude that most of the NMDA-induced current is due to indirect effect via neuronal glutamate release and glial glutamate uptake; but a component was present that would indicate the presence of intrinsic NMDA receptors in astrocytes. Glial NMDA currents display properties of functional NMDA receptors To study NMDA-induced changes in membrane conductance and to determine the reversal potential, we clamped the membrane potentials to a series of depolarizing and hyperpolarizing values (–180 to +70 mV with 25 mV increment, 100 ms per voltage step). This series of voltage steps was repetitively applied every 5.5 s and allowed us to monitor membrane conductance and determine reversal potentials at this frequency (Fig. 4). Unfortunately, it was not possible to block the resting, passive membrane conductance. Neither 20 mM Ba2+ in the bath perfusate, 130 mM CsCl instead of KCl in the patch pipette, nor tetraethylammonium (TEA) significantly affected the resting K+ currents (data not shown). In several recordings the K+ conductances varied, and a reliable reversal potential could not be determined. We therefore selected for cells in which the resting membrane conductance was stable over a period of more than 30 min. The current voltage curve of the NMDA-induced current (at the peak of the response minus control) showed an increase in membrane conductance by 2.9 +/– 1.8 nS (n=9). The reversal potential was more positive than 25 mV (Fig. 4, left). In the presence of Cd2+, CNQX, PDC, and TTX, the conductance increase was significantly smaller (mean 0.39 +/– 0.34 nS, n=6). The
reversal potential, however, was close to 0 mV (–1.3 mV, range –7.5 to +15.6 mV, n=4) (Fig. 4, right), as was expected for responses due to the activation of NMDA receptors. The NMDAinduced current voltage curve was linear (Fig. 4). The effect of the blockers (i.e., shifting the reversal potential from positive values to 0 mV) indicates that the glial NMDA response is due to both activation of glial NMDA receptors and indirect effects, most likely because glutamate transporters are activated after neuronal glutamate release is triggered by NMDA. NMDA responses can be blocked by high Mg2+ Functional NMDA receptor channels display a voltage-dependent Mg2+ block at potentials more negative than –40 mV. Therefore, we investigated the effect of extracellular Mg2+ on intrinsic astrocytic NMDA responses. To isolate the intrinsic astrocytic NMDA response, we performed experiments in the presence of PDC, Cd2+, and CNQX (Fig. 5). Mg2+ concentrations of 4 mM and higher almost abolished NMDA responses as compared with a previous control in Mg 2+-free solution (39.5 pA vs. 431 pA) (n=6; Fig. 5). Under these conditions the Mg2+ block was irreversible and could not be washed out (Fig. 5A). However, this block could be overcome by depolarizing the membrane for 5 s to 0 mV in Mg2+-free solution. After such a depolarization, NMDA elicited responses with an amplitude similar to that of the control prior to high Mg2+ application (Fig. 5B; n=2). NMDA triggers local increases in cytosolic Ca2+ NMDA receptors are Ca2+-permeable, and we therefore tested the effect of NMDA on astrocytic Ca2+ levels. In the first experimental series (Fig. 6A–C), EGFP-positive cells were dialyzed with the red-shifted Ca2+ indicator dye Calcium orange via the recording pipette solution, and thus it was possible to distinguish between the emission of the two dyes in our confocal system. NMDA triggered an increase in the fluorescence signal, indicating an increase of intracellular [Ca2+] (n=3; Fig. 6A–C). We performed the experiments in the absence or in the presence of Cd2+, CNQX, PDC, and TTX , but always in Mg2+-free bath solution while cells were clamped at –80 mV. Yet Calcium orange yielded only a poor signal amplitude due to its properties, and the recordings showed considerable noise. In a second series of experiments we used Fluo-4, which offers a much better ratio between the Ca2+-bound and the free fluorophore emission (FCa2+bound/ FCa2+-free is 3 and >100 for Calcium orange and Fluo-4, respectively). Because Fluo-4 and EGFP have similar fluorescence spectra, we had to record from astrocytes of nontransgenic FVB/N mice. The better properties of Fluo-4 permitted us to differentiate between responses in selected parts of processes and the soma. In the soma we detected only small responses, but we recorded much larger responses in the distal part of the processes (n=8; Fig. 6 D–E). DISCUSSION The GFAP/EGFP mouse as a unique tool to identify live astrocytes At present, there is no specific surface marker that unequivocally identifies live astrocytes. Previous physiological approaches to study astrocytes in living brain slices either relied on morphological identification (28) or identified astrocytes after recording by filling the cell with Lucifer Yellow, pulling off the recording pipette, fixing the slice, permeabilizing cells and,
finally, immunolabeling for GFAP (29, 30). In our previous studies we intensively used the latter approach, but the success rate was limited: We lost cells at each step, and in many cells (with morphological features of astrocytes) the GFAP immunolabel was very faint or even invisible. For the present study, we generated a GFAP/EGFP transgenic mouse line, which allowed us to unequivocally identify astrocytes. It is well established for the central nervous system that all cells with GFAP promotor activity are astrocytes (20, 31). A similar transgenic mouse line had already been generated using the S65T variant of the green fluorescent protein (32), which is dimmer than the EGFP variant used in this study. So far this type of labeling is unique for live and intact astrocytes, and we used this mouse to sort for a live population of astrocytes and identify astrocytes in living brain slices. The cellular label visualized the specific astrocytic morphological features, such as the endfeet contacting the blood vessels. The membrane currents as studied with the patch clamp technique exhibited the passive features as described for mature astrocytes (28). Astrocytes sense the activation of neuronal NMDA receptors We observed that a large portion of the NMDA current amplitude in astrocytes is blocked in the presence of drugs that interfere with neuronal activity (i.e., TTX blocking voltage-gated sodium channels), synaptic transmission (i.e., Cd2+-inhibiting transmitter release via blocking voltagegated Ca2+ channels and CNQX blocking postsynaptic glutamate receptors of the AMPA-type), and glutamate uptake (i.e., PDC acting on glutamate transporters). Therefore, we conclude that a major component of the astrocytic NMDA response is not directly due to activation of astrocytic NMDA receptors, but rather to an indirect effect involving the activation of neuronal NMDA receptors. Neuronal activity (i.e., firing of action potentials) triggered in response to NMDA receptor activation could result in glial currents by the following mechanisms: First, the increased firing rate leads to an increase in extracellular K+ and thereby triggers an inward K+ current in astrocytes; and second, activity-dependent release of glutamate from neurones could trigger uptake currents in glial cells. The action of astrocytes as a K+ sink or the glutamate uptake current are well described in astrocytes (33–38). The reversal potential of the glutamate uptake current is far in the positive range, which is compatible with our observation in the absence of blockers for uptake and synaptic transmission. Astrocytes express functional NMDA receptors We have isolated the intrinsic NMDA receptor response of the astrocytes by blocking the indirect effects after neuronal NMDA receptor activation. Only under these conditions did we record a conductance increase that reversed at 0 mV. The response is blocked by MK-801, leads to an increase in cytosolic Ca2+, and thus shows similarities to neuronal NMDA receptors (39). A difference in the conventional neuronal or cloned NMDA receptor is the reduced Mg2+ sensitivity and the linear current-voltage relation. However, similar, less voltage-dependent NMDA-evoked responses have been described for immature CA3 pyramidal neurones in early postnatal rats (40). Studies of heteromeric recombinant NMDA receptors revealed a subunit dependent Mg2+ sensitivity (41). Conductances of heteromeric NMDA receptor channels composed of NR1 and NR2C were less reduced at high negative membrane potentials in the presence of Mg2+ than channel complexes containing NR2A or NR2B. Interestingly, we found NR2B and NR2C mRNAs in FACS-sorted astrocytes.
The functional data on NMDA receptor expression are complemented by several observations that both mRNA and NMDA receptors can be detected in glial cells, particularly astrocytes. NR2C mRNA was detected in small cells, which the authors considered to be ”most likely glial cells” (42). NR2B mRNA was detected in Bergmann glial cells by in-situ hybridization (43). Using immunoelectron microscopy, several authors reported labeling of glial processes with antibodies against NR1 (11, 12). NMDA receptors are confined to astroglial processes The NMDA-triggered Ca2+ signals in our recordings were confined to peripheral parts of processes and were not observed in the soma. This resolution was only achieved by applying confocal microscopy and can explain why we had not detected NMDA-triggered Ca2+ signals in Bergmann glial cells in a previous study using conventional imaging techniques (8). These functional restrictions to distal parts of astrocytic processes suggest that astrocytic NMDA receptors are confined to these regions. Indeed, this view is supported by immunohistochemical localization of different NMDA receptor subunits at the ultrastructural level. As reported in several studies, the protein is enriched in areas where astroglial membranes enwrap synaptic regions (11–14, 16, 17, 44–48). ACKNOWLEDGMENTS This work was supported by grants from the Federal Ministry for Education and Research of Germany (BMBF), the German Research Foundation (SFB 505), and the Volkswagen foundation. The authors wish to thank Sybille Just for excellent technical assistance. The many valuable discussions with Dr. Christian Steinhäuser, Bonn, and Dr. Alexej Verkhratsky, Manchester, are gratefully acknowledged. REFERENCES 1. Verkhratsky, A., Orkand, R. K., and Kettenmann, H. (1998) Glial calcium: homeostasis and signaling function. Physiol. Rev. 78, 99–141 2. Verkhratsky, A., and Kettenmann, H. (1996) Calcium signalling in glial cells. Trends. Neurosci. 19, 346–352 3. Araque, A., Parpura, V., Sanzgiri, R. P., and Haydon, P. G. (1999) Tripartite synapses: glia, the unacknowledged partner. Trends. Neurosci. 22, 208–215 4. Parpura, V., Basarsky, T. A., Liu, F., Jeftinija, K., Jeftinija, S., and Haydon, P. G. (1994) Glutamate-mediated astrocyte-neuron signalling. Nature 369, 744–747 5. Newman, E. A., and Zahs, K. R. (1998) Modulation of neuronal activity by glial cells in the retina. J. Neurosci. 18, 4022–4028
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Fig. 1
Figure 1. Freshly isolated astrocytes express NMDA receptor mRNAs. RT-PCR analysis of NMDA receptor gene activity in cortical astrocytes acutely isolated by FACS sorting from GFAP/EGFP transgenic mice. In FACS-sorted astrocytes, GFAP mRNA and EGFP mRNA could be amplified via RT-PCR. However, the neurone-specific tyrosine hydroxylase message (TH) could not be detected. Significant levels of amplification products were found for NR1, NR2B, and C,, but not for NR2A, 2D, and 3. RNA isolated from the cortex of transgenic GFAP/EGFP mice was used as the positive control. mRNAs encoding EGFP, TH, and all NMDA receptor subunits could be amplified.
Fig. 2
Figure 2. Morphological and electrophysiological features of EGFP-positive astrocytes in the slice. (A) Confocal laser-scanning microscopical image of an astrocyte from a cortical slice expressing the green fluorescent protein under the control of the GFAP promoter. Astrocytes under study were characterized by their numerous, highly branched processes. One endfeet contacted a blood vessel (upper right) while other processes ended within the neuropil enwrapping synaptic regions or neuronal somata. (B) Whole-cell currents of the cell shown in A. The cell is characterized by a symmetrical pattern of non-inactivating outward and inward currents elicited by depolarizing and hyperpolarizing voltage steps (from – 170 to +10 mV). The membrane was clamped at –80 mV. (C) NMDA was applied (100 µM) to the same cell in the presence of 50 µM PDC and 50 µM CNQX.(D–F) Green fluorescent cells in the mouse forebrain with the morphological and electrophysiological properties of protoplasmic astrocytes express the intermediate filament protein GFAP. (D) Micrograph of green fluorecent cells expressing EGFP in a brain slice; (E) the same area, in which GFAP expression is visualized by specific polyclonal rabbit antibovine GFAP antibodies; (F) overlay of the two fluorescence images. The horizontal bars represent 15 µm in A and 30 µm in D–F.
Fig. 3
Figure 3. Pharmacological properties of astroglial NMDA responses. (A) Block of the NMDA response by MK-801. NMDA (10–4 M) elicited an inward current (left trace). In the presence of MK-801 (10–5 M), the response to NMDA was abolished (middle trace) and did not recover after washout (right trace). The interval between the applications was 20 min. Time of application is indicated by the bars. (B) In the presence of tetrodotoxin (to block voltage-gated Na+ currents mediating action potentials), PDC (to block glutamate transporters), CNQX (to block AMPA receptor currents), and Cd2+ (to inhibit Ca2+-dependent synaptic glutamate release), NMDA (10–4 M) evoked responses in astrocytes were reduced to 10% to 25 % (middle trace) as compared with the control before (left trace) and after the application of the blockers (right trace).
Fig. 4
Figure 4. Reversal potential of NMDA responses in the presence and absence of blockers. A series of depolarizing and hyperpolarizing voltage steps (+25, 50, 75, 100, 125, and 150 mV positive and –25, –50, –75, and –100 mV to the holding potential of –80 mV) was repetitively applied every 3 s, and NMDA (100 µM) was applied as indicated in the upper traces. The recording on the left was obtained in the absence (control) and the recording on the right in the presence, of the blockers Cd2+, CNQX, PDC, and TTX from the same astrocyte. The inset on the right shows the NMDA-induced inward current at the holding potential of –80 mV and is a blow-up of the main trace. To eliminate the display of the voltage steps, we selected a low sampling frequency (1.8 Hz) collecting the data points between the voltage jumps (sampling frequency of the original trace, 500 Hz). In the middle, a series of voltage jumps are displayed with a better time resolution prior to the NMDA () and at the peak of the NMDA (inward current at –80 mV, ). The triangles at the top and the middle traces correspond. The current (I) to voltage (U) curve of the NMDA-triggered response was obtained by subtracting current responses at the peak (of the response) from those recorded before NMDA application. The reversal potential in control solution was 25 mV; in the presence of the blockers 2 mV.
Fig. 5
Figure 5. Mg2+ block of the NMDA response can be released by depolarization. (A) Block of the NMDA response by high Mg2+. NMDA (10–4 M) elicited an inward current (left trace) in the control solution containing nominally free Mg2+. In the presence of 10-mM Mg2+ (10–5 M), the response to NMDA was abolished (middle trace) and did not recover after washout (right trace). The interval between the applications was 20 min. The time of application is indicated by the bars. The blockers Cd2+, CNQX, and PDC were present before, during, and after the NMDA application, as indicated by the gray bars. (B) In a similar experiment as described in A, the cell was depolarized for 5 s to 0 mV after applying NMDA in the presence of 4 mM Mg2+. Note that the subsequent NMDA application (right trace) triggered a response with an amplitude similar to the one in the control (left trace).
Fig. 6
Figure 6. NMDA triggered an increase in [Ca2+]i. (A) The confocal laser-scanning micrograph displays the EGFP fluorescence of an astrocyte in a cortical slice obtained from a GFAP/EGFP transgenic mouse. The square indicates the sectors shown in B displaying the fluorescence of the Ca2+ indicator calcium orange prior to (upper micrograph) and during NMDA application (lower micrograph). The sector in B outlines the area for measuring the fluorescence change (F/Fo) in calcium orange as displayed in the upper trace in C. The lower trace in C is the corresponding current trace, which was recorded simultaneously. NMDA (100 µM) was applied as indicated by bars. (D) Micrograph of Fluo-4 fluorescence: A cell with the morphological features of an astrocyte was dialyzed with Fluo-4 via the patch pipette from a nontransgenic FVB/N mouse. Note the recording pipette approaching the cell from top. Fluorescence changes (F/Fo) were analyzed in the areas denoted by the squares and are displayed in the top three traces in E. The lower trace is the simultaneously recorded current response. NMDA (100 µM) and CNQX, TTX, Cd2+ were applied as indicated by bars.
