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• W1520391040 abstract "Non-technical summary Ca2+ increases in astrocytes have been suggested to trigger the release of neuroactive compounds called gliotransmitters. To study the mechanism of gliotransmitter release, it is desirable to have a method able to selectively and reproducibly evoke Ca2+ increases in astrocytes. Here, we show that the optogenetic tool the light-gated glutamate receptor 6 (LiGluR) reproducibly evokes Ca2+ rises in cultured astrocytes. Combining LiGluR photoactivation and evanescent-field Ca2+ imaging, we explored the cellular mechanism of gliotransmitter release from astrocytes in an all-optical manner. The results suggest that high Ca2+ in astrocytes triggers the release of glutamate via anion channels rather than vesicular exocytosis. Abstract Increases in astrocyte Ca2+ have been suggested to evoke gliotransmitter release, however, the mechanism of release, the identity of such transmitter(s), and even whether and when such release occurs, are controversial, largely due to the lack of a method for selective and reproducible stimulation of electrically silent astrocytes. Here we show that photoactivation of the light-gated Ca2+-permeable ionotropic GluR6 glutamate receptor (LiGluR), and to a lesser extent the new Ca2+-translocating channelrhodopsin CatCh, evokes more reliable Ca2+ elevation than the mutant channelrhodopsin 2, ChR2(H134R) in cultured cortical astrocytes. We used evanescent-field excitation for near-membrane Ca2+ imaging, and epifluorescence to activate and inactivate LiGluR. By alternating activation and inactivation light pulses, the LiGluR-evoked Ca2+ rises could be graded in amplitude and duration. The optical stimulation of LiGluR-expressing astrocytes evoked probabilistic glutamate-mediated signalling to adjacent LiGluR-non-expressing astrocytes. This astrocyte-to-astrocyte signalling was insensitive to the inactivation of vesicular release, hemichannels and glutamate-transporters, and sensitive to anion channel blockers. Our results show that LiGluR is a powerful tool to selectively and reproducibly activate astrocytes. Electrically non-excitable astrocytes respond to neurotransmitters with Ca2+ elevations (Agulhon et al. 2008) that have been suggested to trigger the release of neuroactive gliotransmitters like glutamate and ATP, and to mediate communication between glia and between glia and neurons (Perea & Araque, 2007; Agulhon et al. 2008; Fiacco et al. 2009). Gliotransmission has emerged in recent years as an additional layer of cellular communication that could contribute to information processing by the brain (Halassa & Haydon, 2010). However, the Ca2+ signals required for gliotransmitter release, the release mechanism, and indeed the very existence of such release are highly controversial (Fiacco et al. 2009; Agulhon et al. 2010; Hamilton & Attwell, 2010). One obstacle to a resolution of these questions is that astrocytes share many ligand-gated receptors with neurons (Fiacco et al. 2009). To activate astrocytes specifically, a transgenic mouse has been generated which selectively expresses a foreign G protein coupled receptor (GPCR) in astrocytes. Strikingly, in these mice, triggering specifically Ca2+ elevations in astrocytes with a ligand had an effect neither on synaptic transmission and plasticity nor on neuronal excitability (Fiacco et al. 2007; Agulhon et al. 2010). This contrasts with other results showing Ca2+-dependent gliotransmitter release (Perea & Araque, 2007; Andersson & Hanse, 2010; Gomez-Gonzalo et al. 2010; Halassa & Haydon, 2010). The reasons for these contradictory results are unclear (Agulhon et al. 2010; Halassa & Haydon, 2010). The coupling between Ca2+ signals and gliotransmitter release is still debated. Astrocytic Ca2+ signals rely on plasma membrane receptors and channels, and on intracellular stores (Parpura et al. 2011) and near-membrane spontaneous and ATP-evoked Ca2+ signals depend on different Ca2+ sources (Shigetomi et al. 2010). This diversity of Ca2+ signals could explain why different Gq GPCRs are not equally competent to triggering glutamate release (Shigetomi et al. 2008). A better understanding of Ca2+-dependant gliotransmitter release will require the development of new specific tools for a reliable time-locked control of Ca2+ signals, which need to be associated with imaging techniques for the simultaneous monitoring of Ca2+ signals in electrically silent astrocytes. In an attempt to develop such methods, we set out to control astrocytic Ca2+ rises using the light-gated Ca2+-permeable ionotropic glutamate receptor 6 (LiGluR), the monovalent cationic channel channelrhodopsin 2 (ChR2), and the new Ca2+-translocating channelrhodopsin (CatCh). These channels have been used for a remote non-invasive activation of neurons with high spatio-temporal resolution (Szobota et al. 2007; Gradinaru et al. 2009; Wyart et al. 2009), and ChR2 has been used to activate astrocyte-dependent neuronal responses in vivo (Gradinaru et al. 2009; Gourine et al. 2010). Using cultured cortical astrocytes, we first show that LiGluR and, to a lesser extent, CatCh, provide a more reliable means for triggering reproducible intracellular Ca2+ elevations than the enhanced ChR2 mutant (ChR2(H134R)). We then combined LiGluR activation with evanescent-field excitation Ca2+ imaging to study the responses of LiGluR non-expressing (LiGluR(−)) astrocytes to the photoactivation of their LiGluR-expressing (LiGluR(+)) neighbours. We find that light-evoked Ca2+ elevation in LiGluR(+) astrocytes triggers delayed short-lasting small-amplitude Ca2+ transients in adjacent LiGluR(−) astrocytes, and that this LiGluR-evoked astrocyte-to-astrocyte communication involves a glutamate-permeable anion channel. Our results demonstrate the utility of LiGluR-based optogenetic approaches for studying communication between electrically silent cells. All experiments followed the European Union and institutional guidelines for the care and use of laboratory animals (Council directive 86/609EEC). Cortical astrocytes were prepared from P0–1 (P0 being the day of birth) NMRI mice as previously described (Li et al. 2009). The neocortex was dissected and mechanically dissociated. Cells were plated and maintained on poly-ornithine-coated coverslips (no. 1, BK-7, 25 mm diameter, Menzel-Gläser GmbH) for 1 week to reach confluence. Then, 0.15 mm dibutyryl cAMP (Sigma) was added to induce astrocyte differentiation into a more stellate morphology and form non-confluent islands of several astrocytes. All cultures were kept at 37°C in a humidified 5% CO2 atmosphere. Primary astrocyte cultures were maintained in supplemented Dulbecco's modified Eagle's medium (DMEM, Invitrogen) with 5% fetal bovine serum (FBS), penicillin (5 U ml−1) and streptomycin (5 μg ml−1). The recordings were made during the following week. For neuronal culture, cortical neurons and astrocytes were isolated from embryonic mice (E16). Cells were seeded on poly-l-lysine-treated coverslips and maintained in a medium containing minimum essential medium (MEM) with 5% FBS, 0.3% high glucose MEM, serum extender (1/1000) and penicillin/streptomycin (1/500). Cells were then maintained in half of this serum and half of medium of primary astrocytes containing growth factors for neurons. Cell media and supplements were from Invitrogen (Cergy Pontoise, France). Recordings were made at room temperature (RT) in solution containing (in mm): 140 NaCl, 5.5 KCl, 1.8 CaCl2, 1 MgCl2, 20 glucose, 10 Hepes (pH 7.3, adjusted with NaOH). Ca2+-free extracellular solutions contained nominally zero Ca2+ and 5 mm EGTA. Cell media, supplements, Ca2+ indicators (Fluo-4 AM, Xrhod-1 AM), and dyes (FM4–64, calcein AM, acridine orange, the mitochondrial marker pyridinium, 4-(2-(4-(dimethylamino)phenyl)ethenyl)-1-methyl iodide (DASPMI)) were purchased from Invitrogen (Cergy Pontoise, France), P2 receptor antagonists (pyridoxal-phosphate-6-azophenyl-2′,4′-disulfonate (PPADS), suramin, 2′-deoxy-N6-methyladenosine 3′,5′-bisphosphate (MRS2179)), mGluR receptor antagonists (α-methyl-4-carboxyphenylglycine (MCPG), 2-methyl-6-(phenylethynyl)pyridine (MPEP)), glutamate transporter inhibitor (dl-threo-β-benzyloxyaspartic acid (TBOA)) and thapsigargin were from Tocris (Bristol, UK), bafilomycin A1 from Calbiochem (Merck, Lyon, France) and all other compounds from Sigma-Aldrich. For fluorescence immunostaining experiments, we used the following antibodies: rabbit polyclonal glutamate antibody (1:200, ab9440, Abcam Inc., Cambridge, MA, USA) conjugated to the secondary antibody Alexa 594 goat anti-rabbit IgG (1:500, Invitrogen), mouse glial fibrillary acid protein (GFAP) monoclonal antibody (1:400, MAB360, Chemicon, Temecula, CA, USA) conjugated to the secondary antibody Alexa 488 goat anti mouse IgG (1:500, Invitrogen). Astrocytes were fixed with 1% paraformaldehyde and 0.1% glutaraldehyde (10 min, RT). After permeabilization and blockage of unspecific sites with phosphate buffered saline (PBS) 1×, 0.3% Triton X-100 and 2% bovine serum albumin (BSA) (1 h, RT), astrocytes were probed with respective primary antibody in the same solution overnight at 4°C. After being washed with PBS three times at RT, cells were incubated with secondary antibodies (2 h, RT). After three final washes (PBS, 10 min, RT), cells were mounted with Vectashield onto microscope slides. Immunostained cells were imaged with a Zeiss Axiovert LSM 510 confocal microscope using a ×63, NA 1.4 oil objective. Transfection was performed using lipofectamine 2000 (Invitrogen). Astrocytes were labelled with FM4–64 by incubating cells in dye-containing (6.7 μm) solution for 15 min and rinsing the cells for 30 min prior to imaging. Ca2+ dyes were bulk-loaded into astrocytes as AM-esters in static dye-containing solutions (2 μm, 30 min for Fluo-4; 200 nm, 15 min for Xrhod-1). Another 30 min under continuous perfusion of dye-free solution allowed for the wash-off of extracellular dye and the complete de-esterification of the intracellular dye. During recording, cells were perfused at 0.5 ml min−1 with standard extracellular solution. The glutamate-evoked Ca2+ transients were induced by a brief (1 s) application of glutamate (100 μm) through a dual-channel local perfusion system, one continuously perfusing the control buffer and the other transiently delivering the glutamate-containing solution while the control channel was stopped. The solutions were delivered through plastic tubing (0.8 mm inner diameter, Tygon, Charny, France) to a multi-channel holder (AutoMate Scientific, Berkeley, CA, USA) connected to a small (250 μm ID) silica pipette (WPI, Saratosa, FL, USA). All combinations of excitation wavelengths and dichroic and emission filters used are listed in Supplemental Table 1. A custom-built inverted and upright microscope was used for bright-field, polychromatic epifluorescence imaging and through-the-objective (PlanApo TIRFM, ×60, 1.45 NA, Olympus, Hamburg, Germany) total internal reflection fluorescence microscopy (TIRFM). A Polychrome II (TILL Photonics, Gräfelfing, Germany) provided monochromatic (18 nm FWHM) epifluorescence (EPI) illumination. The 488 and 568 nm lines used for TIRFM were isolated from the beam of an Ar/Kr multi-line laser (CVI Melles Griot, Carlsbad, CA, USA) with an acousto-optical tuneable filter (AA.Opto, Saint Rémy de Chevreuse, France) and directed onto the glass–water interface at a super-critical angle. We estimated the effective penetration depth (1/e2-intensity decay) of the order of 200 nm (Nadrigny et al. 2007). Fluorescence images were further magnified (×2) and projected on an electron multiplying charge-coupled device (EMCCD, QuantEM 512, Princeton Instruments, Trenton, NJ, USA). All devices were controlled by Metamorph (Molecular Devices, Downingtown, PA, USA). The effective pixel size in the sample plane was 133 nm. Time-lapse image stacks were taken at 0.5 Hz with 50–300 ms exposure times unless otherwise indicated. Non-ratiometric Ca2+ indicators (Fluo-4 and Xrhod-1) and TIRFM were used to monitor near-membrane Ca2+ increases throughout this study, unless otherwise indicated. TIRFM confines illumination to the near-membrane, providing the conditions for sensitive detection of local Ca2+ signals. Time-lapse fluorescence changes are plotted as Ca2+-dependent fluorescence measured in 2 μm × 2 μm (15 pixel × 15 pixel) regions of interest (ROIs) normalized to the pre-stimulation baseline (ΔF/F0). The Ca2+ elevations in LiGluR(−) cells were searched for by scanning such ROIs across the LiGluR(−) cell. To avoid a possible interference with light-evoked Ca2+ rises in LiGluR(+) cells, Ca2+ transients in LiGluR(−) cells were searched for in regions more than 2 μm away from the visible interface. Signalling events were identified as ΔF/F0 increases larger than three times the standard deviation (SD) of the baseline. Traces were corrected for photobleaching as follows: individual reference traces were separately obtained by imaging dye-loaded cells without photoactivation, from which the mean decay time constant was obtained from a mono-exponential fit and then applied to correct the Ca2+ traces in photostimulation experiments. Ca2+ signals in astrocytes co-labelled with LiGluR–green fluorescent protein (GFP) and FM4–64 were imaged with Xrhod-1 following the triple-colour detection scheme as described previously (Li et al. 2008). Fluo-4 AM was used for Ca2+ imaging of cells labelled with LiGluR–red fluorescent protein (RFP) and calcein. Ca2+ imaging was performed ∼5–10 min after pharmacological treatment of astrocytes unless otherwise stated. For the experiments done in zero Ca2+, cells were kept in Ca2+-free buffer for less than 5 min to avoid an interference with internal Ca2+ stores (Li et al. 2008). Finally, Ca2+ elevations were recorded only from cultured astrocytes showing no spontaneous Ca2+ elevations. The engineering of LiGluR and synthesis of the tethered photoswitch maleimide-azobenzene-glutamate (MAG) have been previously described (Volgraf et al. 2006; Gorostiza et al. 2007; Numano et al. 2009). Cultured astrocytes were transfected with LiGluR-GFP, LiGluR-RFP, or cotransfected with LiGluR-RFP and AsRed under the control of the cytomegalovirus (CMV) promoter, using lipofectamine 2000 following the standard protocol. Cells were maintained in culture for 1–2 days after transfection to allow the proper targeting of LiGluR to the cell surface. Stock MAG solutions of 5–10 mm were prepared in anhydrous dimethyl sulfoxide (≥99.9%, Sigma) and stored at −20°C. To efficiently conjugate the photoswitch to LiGluR, MAG was pre-illuminated with 385 nm light (0.3 mW mm-2) for 1 min (Gorostiza et al. 2007) before dilution in standard imaging solution to a final concentration of 10 μm. Transfected cells were then incubated in MAG-containing solution for 30 min in the dark followed by another 30 min washing with control buffer. During incubation, 5 μm concanavalin A (Sigma) was added to the medium to block LiGluR desensitisation (Numano et al. 2009). To switch on LiGluR, a 385 nm LED source (Thorlabs, Maisons Lafitte, France) was focused through an air objective (×10, NA 0.25, Olympus) to excite the cells in a 2 mm wide field, and the light pulse duration was 50–200 ms, unless otherwise stated. Where indicated, 488 nm light from the monochromator was used to EPI-illuminate the target cell and switch off LiGluR (39.1 mW mm-2). Astrocytes and neurons were transfected with ChR2(H134R)-GFP using the same protocol as used for LiGluR, and examined 1–2 days after transfection. CatCh–yellow fluorescent protein (YFP) plasmid was provided by Dr Ernst Bamberg (MPI für Biophysik, Frankfurt, Germany). The red Ca2+ dye Xrhod-1 (200 nm, 15 min) was used for Ca2+ imaging of cells expressing ChR2 or CatCh. ChR2 and CatCh were photoactivated by 458 nm light (27.3 mW mm-2 and 15.1 mW mm-2, respectively) generated by the monochromator and delivered through the inverted epifluorescence pathway. For all light-gated channels, Ca2+ imaging was temporally ceased during photoactivation and deactivation. All data are expressed as mean ± standard deviation (SD), and Student's t test was used for testing the significance of P values. Non-normally distributed data were compared using their median ± absolute deviation and non-parametric tests (Kolmogorov–Smirnov, KS). All statistical operations used Matlab (The MathWorks, Natick, MA, USA). *P < 0.05, **P < 0.01, ***P < 0.001. LiGluR is a light-gated channel consisting of a synthetic cysteine-reactive photoisomerizable agonist MAG covalently attached to a cysteine-substituted ionotropic glutamate receptor GluR6 (Volgraf et al. 2006; Numano et al. 2009). To examine the ability of LiGluR to control astrocytic Ca2+ increase, mouse cortical astrocytes in culture were transfected with LiGluR-RFP to which MAG was covalently attached. Short pulses of 385 nm light (0.3 mW mm-2, 50 ms) evoked robust and reliable Ca2+ rises monitored with the green-fluorescent Ca2+ indicator Fluo-4 (ΔF/F0= 29.9 ± 4.0%, n= 23 trials from 7 cells; Fig. 1A). Light-evoked Ca2+ responses were neither observed in the absence of LiGluR-RFP (−0.1 ± 1.5%, n= 5 cells), nor in cells expressing LiGluR without MAG (0.3 ± 2.0%, n= 5 cells), nor in cells lacking LiGluR and MAG (ΔF/F0= 0.1 ± 1.6%, n= 5 cells) (Supplemental Fig. 1A). The LiGluR-evoked Ca2+ rises recorded in the thin processes of LiGluR(+) astrocytes were synchronized and their amplitude was maintained along the process as expected for a direct activation of LiGluR targeted to the process membrane (Fig. 1B). Finally, the amplitude of LiGluR-elicited Ca2+ signals correlated with the duration of illumination (Supplemental Fig. 1B). LiGluR evokes precisely timed and shaped Ca2+ rises in astrocytes A, dual-colour TIRF image of a cultured cortical astrocyte transfected with LiGluR-RFP conjugated with the photoswitch MAG and loaded with the green-fluorescent Ca2+ indicator Fluo-4 (left). Dashed line shows the contour of the cell. The pseudocolour kymograph and green trace illustrate reproducible Ca2+ rises evoked by 385 nm light pulses (violet arrows, 0.3 mW mm-2, 50 ms). B, LiGluR photoactivation evoked repetitive and synchronized Ca2+ rises in astrocytic soma (ROI-1) and processes (ROI-2 and ROI-3). C, temporal shaping of astrocytic Ca2+ rises by switching LiGluR on and off with alternate 385 nm (violet arrows) and 488 nm (blue arrows, 39.1 mW mm-2, 200 ms) EPI light pulses. D–F, astrocytic Ca2+ elevations induced by LiGluR-GFP photoactivation and monitored with the red-fluorescent Ca2+ indicator Xrhod-1. The Ca2+ signals were shaped by switching off LiGluR with variable delay using 488 nm EPI light from the monochromator. Bars, 10 μm. Besides being activated by UV light, LiGluR can be switched off with 488 nm blue light (Szobota et al. 2007). Because the spectral band for deactivating LiGluR overlaps with the 488 nm excitation wavelength of Fluo-4, epifluorescence Ca2+ imaging quickly terminates LiGluR-evoked responses (Supplemental Fig. 1C and D). To minimize imaging-light induced LiGluR deactivation, and to shape the Ca2+ increases with a pulse of EPI-illumination deactivation, we confined Ca2+ imaging to the near-membrane cellular compartment, using total internal reflection fluorescence microscopy (TIRFM) (Supplemental Fig. 1E). In these conditions, alternating pulses of 385 nm and 488 nm light evoked reproducible Ca2+ transients (Fig. 1C). To minimize LiGluR deactivation during Ca2+ imaging, we also used a LiGluR-GFP construct with the red-fluorescent Ca2+ indicator Xrhod-1 AM. This combination allowed us to monitor LiGluR-evoked Ca2+ responses (ΔF/F0= 31.3 ± 6.0%, n= 12 trials from 5 cells), and to shape the Ca2+ rise by altering the interval between the activating and deactivating light pulses (Fig. 1D–F). Together, these results indicate that photoactivation of LiGluR provides a robust, graded and reliable control of Ca2+ signalling in astrocytes. Previous reports indicate that ChR2 can activate astrocytes (Gradinaru et al. 2009; Gourine et al. 2010); therefore we decided to compare the ability of ChR2 and LiGluR to elicit Ca2+ elevations in astrocytes. We first confirmed our ability to activate ChR2-expressing neurons in culture by showing that the repetitive photoactivation of ChR2(H134R) (Nagel et al. 2005) with 458 nm light pulses (27.3 mW mm-2, 500 ms) evoked Ca2+ transients (Fig. 2A) in 9 of 14 cortical neurons expressing ChR2(H134R)-GFP. Light-evoked Ca2+ transients were absent in control neurons without ChR2(H134R)-GFP (Fig. 2B), and were inhibited by cobalt, a non-selective blocker of voltage-gated Ca2+ channels (VGCCs) (Fig. 2C), indicating that neuronal Ca2+ rise is mainly due to a secondary entry via VGCCs. Ca2+ signalling in astrocytes evoked by photoactivation of the mutant channelrhodopsin ChR2(H134R) and the Ca2+-permeable channelrhodopsin CatCh A, in a neuron expressing ChR2(H134R)-GFP and loaded with the red Ca2+ indicator Xrhod-1, 458 nm EPI pulses (27.3 mW mm−2, 500 ms) evoked reproducible Ca2+ rises (ΔF/F0= 46.0 ± 13.2%, n= 9 cells). B, absence of light-evoked Ca2+ increases in neurons not expressing ChR2 (ΔF/F0= 1.3 ± 6%, n= 4 cells). C, neuronal ChR2-evoked Ca2+ rises were inhibited by the non-selective voltage-gated Ca2+ channel (VGCC) blocker, cobalt (Co, 1 mm, ΔF/F0= 9.2 ± 4.7%, FWHM = 2.5 ± 1.9 s; vs. control ΔF/F0= 45.4 ± 8.3%, P < 0.01, FWHM = 7.0 ± 2.6 s, P < 0.05, n= 8 trials from 4 cells per condition). D, in astrocytes, short photoactivation (458 nm, 27.3 mW mm-2, 500 ms) of ChR2 did not evoke any near-membrane Ca2+ elevation. E, example Ca2+ responses evoked by longer light pulse (458 nm, 1 s) in ChR2-expressing astrocytes. F, percentage of astrocytes showing light-gated Ca2+ rises, and Ca2+ response amplitude in LiGluR-expressing (60/63 cells) and ChR2-expressing (19/60 cells) astrocytes. The comparison was made using three successive light pulses (385 nm, 100 ms for LiGluR; 458 nm, 1 s for ChR2) applied every 150 s. G, in astrocytes, the photoactivation (1 s, 458 nm) of CatCh evokes fast Ca2+ rises which fade upon repetitive photoactivation. H, CatCh-evoked Ca2+ elevations in astrocytes were absent in 0 mm Ca2+ extracellular solution (0 Ca, solution change was achieved in less than 30 s), and were unaffected by depleting the endoplasmic reticulum Ca2+ store by thapsigargin (TG, 2 μm, 20 min). Recordings were done on the same cells in different conditions. CatCh photoactivation, 2 s 458 nm (CTR, 15 trials, 5 cells; 0 Ca, 6 trials, 3 cells; TG, 18 trials, 6 cells). Bars, 10 μm. We then expressed ChR2(H134R)-GFP in astrocytes. The average ChR2(H134R)-GFP fluorescence intensity was found to be comparable in astrocytes and neurons, suggesting that in our conditions, similar expression levels are achieved in both cell types (Supplemental Fig. 2A). The same light intensity used for neurons, however, evoked small Ca2+ changes in cultured cortical astrocytes expressing ChR2(H134R) (ΔF/F0= 5.1 ± 4.2%, n= 5, Fig. 2D), which were not significantly different from the control responses of ChR2-non-expressing cells (ΔF/F0= 2.2 ± 3.0%, n= 5, P= 0.3). Longer (1 s) 458 nm light pulse occasionally evoked Ca2+ increases in a subset of astrocytes, but responses were variable in duration and amplitude, as imaged with TIRFM (Fig. 2E) and epifluorescence (Supplemental Fig. 2B). In comparison with LiGluR, ChR2(H134R) was significantly less effective in driving astrocytic Ca2+ elevation, in terms of light intensity and duration needed, percentage of responsive cells, and amplitude of light-evoked Ca2+ responses (Fig. 2F). Our inability to trigger reliable Ca2+ rises in astrocyte following ChR2(H134R) photoactivation could be due to a malfunction of this channel when expressed in astrocytes. To investigate this possibility, we compared the performance of ChR2(H134R) in astrocytes and neurons. Since ChR2 is highly permeable to H+ (Nagel et al. 2003), the H+ influx induced by its photoactivation is expected to quench the fluorescence of ChR2(H134R)-GFP (Nadrigny et al. 2006). We observed that ChR2(H134R) photoactivation induced a similar degree of quenching of intracellular ChR2(H134R)-GFP fluorescence in astrocytes and neurons (Supplemental Fig. 2C). This result indicates that ChR2(H134R) is as well-expressed and functional in astrocytes as it is in neurons. Thus, the relative inefficacy of ChR2 to trigger Ca2+ rise in astrocytes might be due to its low Ca2+ permeability. We therefore tested a recently developed Ca2+-permeable channelrhodopsin mutant called CatCh (Kleinlogel et al. 2011). Ca2+ rises could be triggered in astrocytes by optical activation of CatCh, although this required ∼50 times more power and ∼10 times longer illumination (15.1 mW mm-2, 1 s) than with LiGluR and, unlike the reproducible LiGluR-evoked responses, the CatCh-evoked responses faded in amplitude with repeated stimulation (Fig. 2G). Our results indicate that Ca2+-permeable LiGluR and, to a lesser extent, CatCh are more efficient than ChR2 in controlling astrocytic Ca2+ elevation. We next examined the cellular mechanisms of the Ca2+ increase evoked in astrocytes by the photoactivation of CatCh and LiGluR. CatCh is a light-sensitive Ca2+-permeable ChR2 (Kleinlogel et al. 2011), and CatCh-evoked Ca2+ transients were abolished in 0 mm Ca2+ (Fig. 2H). The endoplasmic reticulum (ER) Ca2+ stores contribute to Ca2+ signalling in astrocytes (Agulhon et al. 2008), but do not seem to be involved in CatCh-evoked Ca2+ rises, since the application of thapsigargin (TG, 1 μm), which induced long-lasting cytoplasmic Ca2+ rises reflecting the leak of Ca2+ from ER stores (ΔF/F0= 71.7 ± 10.4%, n= 3; Fig. 3D), had no effect on the subsequent CatCh-evoked Ca2+ rises (Fig. 2H). These results suggest that CatCh-evoked Ca2+ transients depend only on Ca2+ influx. LiGluR-evoked near-membrane Ca2+ rises in astrocytes depend on Ca2+ influx and uptake into the endoplasmic reticulum A, LiGluR-evoked Ca2+ responses in the absence (control, 12 trials from 4 cells) and in the presence (15 trials, 5 cells) of a competitive GluR6 antagonist NBQX (1 mm). LiGluR-RFP and the green Ca2+ indicator Fluo-4 were used. B, LiGluR-evoked Ca2+ responses in zero Ca2+ (5 trials, 5 cells, left), 1.8 mm Ca2+ (control, 15 trials from 5 cells, middle, black traces), and 5 mm Ca2+ (15 trials, 5 cells; green traces). C, LiGluR activation triggers a rapid near-membrane Ca2+ increase (grey triangles, 0.5 Hz imaging prior to photoactivation. Black triangles, 20 and 100 Hz imaging after 10 ms activation light pulse, purple dots). An intermediate rising step, indicated by a red arrow, was revealed at 100 Hz. Similar results were obtained for six astrocytes. D, the Ca2+-ATPase inhibitor thapsigargin (TG, 1 μm) led to Ca2+ elevation due to Ca2+ release from the internal store. Black arrowhead indicates the beginning of TG application, which was maintained throughout the recording. Each trace depicts the response of a single astrocyte. E, inhibition of Ca2+-ATPase by TG (1 μm, 15 min) enhanced LiGluR-evoked Ca2+ increases (red, 17 trials, 7 cells, vs. control, black, 19 trials, 8 cells. n.s. P > 0.16). LiGluR is derived from the kainate-type glutamate receptor GluR6 (Volgraf et al. 2006), and in line with previous reports (Gorostiza et al. 2007; Numano et al. 2009), the competitive AMPA/kainate receptor antagonist NBQX attenuated the LiGluR-evoked Ca2+ increase (ΔF/F0= 19.3 ± 9.7%, n= 12 trials from 4 cells, vs. control, 43.2 ± 8.5%, n= 15 trials from 5 cells, P < 0.01; Fig. 3A). As predicted by the high Ca2+ permeability of GluR6 (Egebjerg & Heinemann, 1993), the photoactivation of LiGluR in 0 mm Ca2+ failed to trigger detectable Ca2+ rises (ΔF/F0= 0.32 ± 1.4%, n= 5), while the Ca2+ rises evoked in 5 mm extracellular Ca2+ (ΔF/F0= 45.2 ± 14.3%, FWHM = 97.1 ± 43.8 s, n= 15 trials from 5 cells) were larger than the control response evoked in 1.8 mm Ca2+ (ΔF/F0= 21.6 ± 6.7%, P < 0.01; FWHM = 71.3 ± 14.7 s, P < 0.05; n= 15 trials from 5 cells; Fig. 3B). The Ca2+ influx through the ion channel should lead to a rapid rise of the near-membrane Ca2+ concentration detectable with TIRFM (Demuro & Parker, 2005). Sampling LiGluR-evoked Ca2+ rises at 100 Hz, we captured an intermediate ΔF/F0 step one frame after light activation (Fig. 3C, red arrow), suggesting that the Ca2+ rise occurs within 10 ms after LiGluR activation. We then tested a possible involvement of the Ca2+ stores in LiGluR-evoked Ca2+ signalling using thapsigargin (TG, 1 μm, 15 min) to block the ER Ca2+-ATPase and deplete the Ca2+ stores prior to LiGluR activation. Surprisingly, we found that LiGluR-evoked Ca2+ rises were enhanced in TG-treated cells (ΔF/F0= 56.3 ± 16.8%, n= 17 trials from seven cells, vs. control, 23.4 ± 6.2%, n= 19 trials from 8 cells, P < 0.01; Fig. 3E). These results indicate that LiGluR-evoked Ca2+ elevations rely on Ca2+ influx, and that Ca2+ uptake into the ER via the Ca2+-ATPase limits the LiGluR-evoked cytosolic Ca2+ increase. Having established LiGluR as a tool for controlling astrocytic Ca2+ in culture, we investigated if LiGluR-evoked Ca2+ increases can trigger gliotransmitter release. After transfection, only a subset of astrocytes express LiGluR, while the entire population of astrocytes can be loaded with acetoxymethyl ester (AM) derivative of Ca2+ indicators (Fig. 4A). This permitted us to explore astrocyte-to-astrocyte signalling in an all-optical manner, in which light was used to drive Ca2+ elevation in LiGluR expressing (LiGluR(+)) cells as well as to report Ca2+ signals in both the LiGluR(+) and adjacent LiGluR(−) astrocytes. To avoid interference with spontaneous Ca2+ oscillations, recordings were made from astrocytes without spontaneous activity. All optical probing of astrocyte-to-astrocyte communication using LiGluR and TIRF A, AsRed was co-transfected with LiGluR-RFP to i" @default.
• W1520391040 created "2016-06-24" @default.
• W1520391040 creator A5018129332 @default.
• W1520391040 creator A5024310714 @default.
• W1520391040 creator A5028468342 @default.
• W1520391040 creator A5081378606 @default.
• W1520391040 creator A5082997007 @default.
• W1520391040 date "2012-02-14" @default.
• W1520391040 modified "2023-10-10" @default.
• W1520391040 title "Optogenetic activation of LiGluR-expressing astrocytes evokes anion channel-mediated glutamate release" @default.
• W1520391040 cites W1533853049 @default.
• W1520391040 cites W1968446819 @default.
• W1520391040 cites W1970987547 @default.
• W1520391040 cites W1973361488 @default.
• W1520391040 cites W1974361256 @default.
• W1520391040 cites W1984122112 @default.
• W1520391040 cites W1984543142 @default.
• W1520391040 cites W1984766476 @default.
• W1520391040 cites W1987436909 @default.
• W1520391040 cites W1995256270 @default.
• W1520391040 cites W2000604662 @default.
• W1520391040 cites W2006395355 @default.
• W1520391040 cites W2011101653 @default.
• W1520391040 cites W2017787995 @default.
• W1520391040 cites W2020138925 @default.
• W1520391040 cites W2021786044 @default.
• W1520391040 cites W2026154459 @default.
• W1520391040 cites W2027270469 @default.
• W1520391040 cites W2027616725 @default.
• W1520391040 cites W2032154466 @default.
• W1520391040 cites W2032159555 @default.
• W1520391040 cites W2035501441 @default.
• W1520391040 cites W2036048932 @default.
• W1520391040 cites W2040109036 @default.
• W1520391040 cites W2044107589 @default.
• W1520391040 cites W2045092834 @default.
• W1520391040 cites W2048105028 @default.
• W1520391040 cites W2050932531 @default.
• W1520391040 cites W2055162787 @default.
• W1520391040 cites W2058085559 @default.
• W1520391040 cites W2063063986 @default.
• W1520391040 cites W2069294819 @default.
• W1520391040 cites W2078102941 @default.
• W1520391040 cites W2079823127 @default.
• W1520391040 cites W2083932518 @default.
• W1520391040 cites W2086512982 @default.
• W1520391040 cites W2086837225 @default.
• W1520391040 cites W2088318073 @default.
• W1520391040 cites W2088944569 @default.
• W1520391040 cites W2089776232 @default.
• W1520391040 cites W2097242525 @default.
• W1520391040 cites W2099640358 @default.
• W1520391040 cites W2102087494 @default.
• W1520391040 cites W2103244356 @default.
• W1520391040 cites W2104417933 @default.
• W1520391040 cites W2111819027 @default.
• W1520391040 cites W2118751104 @default.
• W1520391040 cites W2118788831 @default.
• W1520391040 cites W2124017577 @default.
• W1520391040 cites W2124940469 @default.
• W1520391040 cites W2127183925 @default.
• W1520391040 cites W2127361742 @default.
• W1520391040 cites W2130525167 @default.
• W1520391040 cites W2132949150 @default.
• W1520391040 cites W2134230622 @default.
• W1520391040 cites W2134779286 @default.
• W1520391040 cites W2136095861 @default.
• W1520391040 cites W2140591268 @default.
• W1520391040 cites W2145593101 @default.
• W1520391040 cites W2147376890 @default.
• W1520391040 cites W2147551208 @default.
• W1520391040 cites W2156094132 @default.
• W1520391040 cites W2158256695 @default.
• W1520391040 cites W2159423248 @default.
• W1520391040 cites W2165135104 @default.
• W1520391040 cites W2171799151 @default.
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