J Neural Transm (2014) 121:245–257 DOI 10.1007/s00702-013-1096-8

TRANSLATIONAL NEUROSCIENCES - ORIGINAL ARTICLE

Glutamate release from astrocyte cell-line GL261 via alterations in the intracellular ion environment Kenji Ono • Hiromi Suzuki • Madoka Higa Kaori Tabata • Makoto Sawada

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Received: 29 August 2013 / Accepted: 25 September 2013 / Published online: 8 October 2013 Ó Springer-Verlag Wien 2013

Abstract Astrocytes modify and maintain neural activity and functions via gliotransmitter release such as, glutamate. They also change their properties and functions in response to alterations of ion environment resulting from neurotransmission; however, the direct evidence for whether intracellular ion alteration in astrocytes triggers gliotransmitter release is not indicated. Recent studies have reported that channelrhodopsin-2 (ChR2) is useful for alteration of intracellular ion environment in several types of cells with blue light exposure. Here, we show that ChR2-expressing GL261 (GLChR2) cells, clonal astrocytes, change their properties by photoactivation. Increased intracellular sodium and calcium ion concentrations and an altered membrane potential were observed in GLChR2 cells with blue light exposure. Alterations in the intracellular ion environment caused intracellular acidification and the inhibition of proliferation. In addition, it triggered glutamate release from GLChR2 cells. Glutamate from GLChR2 cells acted on N18 cells, clonal neuronal cells, as both a transmitter and neurotoxin depending on photo-activation. Our results show that the properties of ChR2-expressing astrocytes can be controlled by blue light exposure, and cation influx through photo-activated ChR2 might trigger functional

K. Ono
H. Suzuki (&)
M. Higa
K. Tabata
M. Sawada Department of Brain Function, Division of Stress Adaptation and Protection, Research Institute of Environmental Medicine, Nagoya University, Nagoya, Aichi 464-8601, Japan e-mail: [email protected] Present Address: K. Tabata Department of Chemo-Pharmaceutical Sciences, Graduate School of Pharmaceutical Sciences, Kyushu University, Fukuoka 812-8582, Japan

cation influx via endogenous channels and result in the increase of glutamate release. Further, our results suggest that ChR2-expressing glial cells could become a useful tool in understanding the roles of glial cell activation and neural communication in the regulation of brain functions. Keywords Glutamate release
Astrocytes
Channelrhodopsin-2
Photo-activation Abbreviations ChR2 GLChR2 EYFP DMEM MEM GAPDH DIC RB Carboxy SNARF-1 AM PI BL TTX

Channelrhodopsin-2 ChR2-expressing GL261 Enhanced yellow fluorescent protein Dulbecco’s modified Eagle’s medium Eagle’s minimum essential medium Glyceraldehyde 3 phosphate dehydrogenase Differential interference contrast Recording buffer 5-(and-6)-Carboxy SNARF-1, acetoxymethyl ester, acetate Propidium iodide Blue light Tetrodotoxin

Introduction Many glial cells such as astrocytes, oligodendrocytes, and microglia exist in the brain parenchyma in addition to

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neurons. For a long time, glial cells were considered to be a population of cells that only provided support and protection for neurons in the brain. However, recent accumulating evidence suggests that glial cells actively participate in synaptic transmission and the processing of information (Perea et al. 2009; Achour and Pascual 2010). Astrocytes are the most abundant type of glial cells and are closely associated with blood vessels, neurons, and other glial cells. In addition, astrocytes release gliotransmitter such as glutamate and ATP in response to alteration of ion environment (Hamilton and Attwell 2010). Glutamate release from astrocytes has an aspect of a transmitter between glial cells and also between glial cells and neurons. Most neurons as well as glial cells express multiple types of glutamate receptors such as AMPA receptors, NMDA receptors, and kainate receptors. Moreover, glutamate plays important roles in long-term potentiation, long-term depression, learning, and memory (Aszte´ly and Gustafsson 1996; Debanne et al. 2003; Pe´rez-Otan˜o and Ehlers 2005). On the other hand, glutamate release from astrocytes has another aspect as a neurotoxin. The overstimulation of glutamate receptors causes neuronal damage through excitotoxicity. Excessive glutamate results in the apoptosis of neuronal cells via mitochondrial damage from excessively high intracellular Ca2? concentrations, increases in transcription factors for pro-apoptotic genes, and decreases in transcription factors for anti-apoptotic genes (Manev et al. 1989; Uberti et al. 2000; Leon et al. 2009). Furthermore, excitotoxicity via glutamate receptors is associated with cerebral ischemia, epilepsy, and many neurodegenerative diseases such as Alzheimer’s disease, Parkinsonism, and multiple sclerosis (Werner et al. 2001; Dong et al. 2009; Tymianski 2011; Coulter and Eid 2012). Therefore, astrocytes may control neuronal activity and synaptic transmission through glutamate release (Pascual et al. 2005; Haydon and Carmignoto 2006). Astrocytic proliferation is influenced by glutamate receptor signaling in culture (Kanumilli and Roberts 2006). In addition, there is accumulating evidence that glutamate promotes the proliferation of invasive glial tumors via autocrine or paracrine activation of glutamate receptors (Rzeski et al. 2001; Ishiuchi et al. 2002; Arcella et al. 2005). Proliferation of reactive astrocytes (gliosis) is a common feature of many brain diseases such as epilepsy and neurodegenerative diseases. The gliosis in epilepsy is correlated with an increased concentration and slowed clearance of extracellular glutamate in the hippocampus (Cavus et al. 2005). Reversal of glutamate uptake system and release of additional glutamate occur in parallel with the gliosis in neurodegenerative diseases (Giaume et al. 2007). Astrocytes respond to alterations in extracellular and intracellular ions such as Na?, K? and Ca2? via

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endogenous ion channels, which result in changes in their properties and functions (Butt and Kalsi 2006; Verkhratsky et al. 2012; Kirischuk et al. 2012). In addition, astrocytes often communicate with each other via calcium signaling such as calcium waves (Hassinger et al. 1996; Shigetomi et al. 2010). Intracellular ion elevations also result in glutamate release from astrocytes (Hamilton and Attwell 2010); however, the direct evidence for whether intracellular ion alteration in astrocytes triggers glutamate release is not indicated. Recent studies have reported that channelrhodopsin-2 (ChR2) is a powerful tool for neural circuit analysis (Wang et al. 2007; Arenkiel et al. 2007; Tsai et al. 2009). ChR2 is a seven-transmembrane ion channel from the green algae Chlamydomonas reinhardtii that allows the entry of cations (mostly sodium ions and very low levels of calcium ions) into cells exposed to blue light (Nagel et al. 2003). Furthermore, ChR2 functions in mammals as well as C. elegans and Drosophila without cofactors (Nagel et al. 2005; Zhang et al. 2007b). It is useful for the control of neuronal spiking and synaptic transmission in ChR2-expressing neurons with blue light exposure (Zhang et al. 2007a). Although it is unclear whether ChR2 functions in cells that cannot generate action potentials unlike neurons (Sasaki et al. 2012), if ChR2 on astrocytes can control the intracellular ion environment on blue light exposure, gliotransmitter release from astrocytes may be altered. Here, we demonstrated that ChR2-expressing GL261 (GLChR2) cells, clonal astrocytes, successfully changed their properties of cell proliferation and glutamate release through increases in intracellular cations with blue light exposure. Released glutamate from GLChR2 cells acted on neuronal cells as either a transmitter or neurotoxin depending on photo-activation.

Materials and methods Cell lines GL261 murine astrocyte cell lines, including ChR2-EYFP expressing derivatives were cultured in Dulbecco’s modified Eagle’s medium (DMEM) (Sigma-Aldrich, St Louis, MO, USA) with 10 % fetal bovine serum and penicillin–streptomycin (Life Technologies, Carlsbad, CA, USA) in a 95 % air/5 % CO2-humidified atmosphere. N18, a murine neuronal cell line was cultured in Eagle’s minimum essential medium (MEM) (Sigma-Aldrich, St Louis, MO, USA) with 10 % fetal bovine serum, 1 % non-essential amino acid solution (Sigma-Aldrich, St Louis, MO, USA), 1 mM sodium pyruvate (Sigma-Aldrich, St Louis, MO, USA), and 1 % penicillin–streptomycin (Life Technologies, Carlsbad, CA, USA) in a 95 % air/5 % CO2-humidified atmosphere.

Glutamate release from astrocyte cell-line GL261

Plasmids and electroporation pcDNA3.1/hChR2-EYFP plasmids were a generous gift from Dr. Karl Deisseroth (Stanford University). GL261 cells (1 9 106 cells/400 ll) were mixed with 10 lg of plasmids in a 4-mm gap cuvette. The cuvette was set in an ECM830 electroporator (BTX Instrument Division Harvard Apparatus, Inc., Holliston, MA, USA) and electroporation was performed under the following conditions (choose mode: LV mode, set voltage: 170 V, set pulse length: 70 ms, set number of pulses: 1). Transfected cells were cultured in the medium with 400 lg/ml of G418 for the selection of mixed clones expressing ChR2-EYFP for 7 days. Several kinds of single clones were picked up from mixed clones after limiting dilutions and a single clone, 1F12, was used in this study. RNA extraction and RT-PCR Total RNA was extracted from cells using the RNeasy Mini kit and RNase-free DNase set (QIAGEN, Hilden, Germany) according to the manufacturer’s instructions. RNA (1 lg) was reverse transcribed at 37 °C for 90 min in a mixture containing 100 U of recombinant M-MLV reverse transcriptase, 0.1 lg DNA random hexamers, 40 U RNase inhibitor, and 1.4 mM dNTPs in a final volume of 50 ll. cDNA was amplified with Taq DNA polymerase (Takara, Tokyo, Japan), using primer pairs specific to ChR2 (sense primer, TAA TCC TGT GGT GGT GAA CG; antisense primer, TCA GCA GCA AAA TGC TGA AT) and EYFP (sense primer, GGG CAC AAG CTG GAG TAC A; antisense primer, GGG GGT GTT CTG CTG GTA) for 35 cycles (94 °C for 1 min, 55 °C for 1 min, and 72 °C for 2 min) and glyceraldehyde 3 phosphate dehydrogenase, GAPDH (sense primer, TGC ACC ACC AAC TGC TTA G; antisense primer, GAT GCA GGG ATG ATG TTC) for 30 cycles, respectively. PCR products were resolved by electrophoresis on 2 % agarose gels stained with ethidium bromide, and were then photographed using Light Capture (ATTO Corporation, Tokyo, Japan). Time-lapse fluorescent imaging Time-lapse series of the cells were taken at 37 °C using a microscope (Axiovert 40; Carl Zeiss, Oberkochen, Germany) equipped with a CCD camera and a humidified CO2 chamber, and operated by Axiovision software. Differential interference contrast (DIC) and fluorescent images were taken and the motion of the cells and fluorescent intensity were analyzed. The CoroNa Red fluorescent probe (Life Technologies, Carlsbad, CA, USA) was used for the visualization of the Na? influx. CoroNa Red, which is a sodium indicator and increases red fluorescence in a concentration-

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dependent manner, was loaded onto GL261 cells and GLChR2 cells at 1 lM in recording buffer (RB) containing 137 mM sodium chloride, 4.2 mM sodium carbonate, 0.34 mM disodium hydrogenphosphate, 5.4 mM potassium chloride, 0.44 mM potassium dihydrogenphosphate, 0.81 mM magnesium sulfate, 1.26 mM calcium chloride, 5.55 mM D-glucose, and 20 mM HEPES (pH 7.4) for 30 min. After removing over-loaded CoroNa Red, timelapse series of the cells were taken in phenol red-free medium for 10 min. Blue light was applied for 200 ms every 30 s. For the evaluation of depolarization, DiBAC4(5) (Life Technologies, Carlsbad, CA, USA), which is a slow membrane potential indicator that increases and accumulates intracellular red fluorescence accompanying depolarization, was loaded onto cells at 1 lM in RB for 20 min. After removing over-loaded DiBAC4(5), time-lapse series of the cells were taken in phenol red-free medium for 30 min. Blue light was applied for 200 ms every 5 min. 5-(and-6)-carboxy SNARF1, acetoxymethyl ester, acetate (carboxy SNARF-1 AM; Life Technologies, Carlsbad, CA, USA) was used for the visualization of intracellular pH, as SNARF-labeled cells display red fluorescence at an alkaline pH that disappears when the pH becomes acidic. Carboxy SNARF-1 AM was loaded onto cells at 1 lM in RB for 20 min. After removing over-loaded carboxy SNARF-1 AM, time-lapse series of the cells were taken in phenol red-free medium for 20 min. Blue light was applied for 50 ms every 30 s. The Rhod-3 Imaging Kit (Life Technologies, Carlsbad, CA, USA) was used for the visualization of the Ca2? influx. Rhod-3AM, which is a calcium indicator that increases red fluorescence in a concentrationdependent manner, was loaded onto GL261 cells and GLChR2 cells at 10 lM in RB for 30 min. After removing over-loaded Rhod-3 AM, cells were incubated in RB containing 2.5 mM probenecid. Time-lapse series of the cells were taken in RB for 10 min. Blue light was applied for 500 ms every 15 s. Fluorescent intensity from the time-lapse series was measured using Photoshop software (Adobe Systems Inc., San Jose, CA, USA). In some cases, time-lapse series of the cells were taken in the presence or absence of 100 lM EDTA, 1 lM TTX, 10 lM verapamil, or 1 lM riluzole. They were also taken in sodium ion-free buffer containing 137 mM choline chloride, 5 mM potassium chloride, 2 mM calcium chloride, 1 mM magnesium sulfate, and 10 mM HEPES (pH 7.4). WST assay Cell proliferation was determined by analyzing the conversion of WST-1 (light red) to its formazan derivate (dark red) using the WST-1 Cell Counting Kit (Dojindo Laboratories, Kumamoto, Japan). Each cell such as N18 cells, GL261 cells, and GLChR2 cells was plated in 96-well culture plates at a density of 5 9 103 cells/100 ll culture

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medium and cultured in the presence or absence of 10 lM NBQX with or without blue light exposure for 24 or 72 h. Each plate was exposed to blue light for 1 min every 5 min until 72 h. At the end of the experiments, cells were incubated with 10 ll of the WST-1 reagent for 4 h at 37 °C in 5 % CO2. The absorbance at 450 and 620 nm (as a reference) was measured using a micro-plate reader (GE Healthcare, Chalfont St. Giles, United Kingdom). FACS analysis Fluorescence in transfected cells and apoptosis in the coculture were detected using a FACSCalibur cell sorter (BD Bioscience, San Jose, CA, USA) equipped with two lasers (a 488 nm argon laser and 635 nm diode laser) and CellQuest software (BD Bioscience, San Jose, CA, USA). Each cell was plated in a 60-mm Cell Culture Dish at a density of 2.0 9 105 cells and was cultivated in the presence or absence of glutamate or NBQX with or without blue light exposure for 72 h. Each cell was exposed in the culture medium to blue light for 1 min every 5 min until 72 h using the Safe Imager Blue-Light Transilluminator (Life Technologies, Carlsbad, CA, USA). The EYFP fluorescence of GLChR2 cells was analyzed without staining. Cell apoptosis was detected by staining with propidium iodide (PI) and Alexa Fluor 647 conjugated Annexin V (Life Technologies, Carlsbad, CA, USA).

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Results GL261 cells transfected with the ChR2-EYFP gene (GLChR2 cells) To confirm the gene transfer of ChR2-EYFP after electroporation, ChR2 and EYFP mRNA expression were analyzed by RT-PCR (Fig. 1a). Expression of ChR2 and EYFP mRNA was clearly found in GLChR2 cells. To clarify the expression of the ChR2-EYFP fusion protein, GLChR2 cells were then analyzed using a flow cytometer (Fig. 1b). EYFPspecific fluorescence was apparently detected in GLChR2 cells. In addition, ChR2-expressing cells were identified by observing EYFP fluorescence on a fluorescent microscope (Fig. 1c). As the fluorescence was found on the cell surface, GLChR2 cells were confirmed to express the ChR2-EYFP fusion protein on the surface of the plasma membrane. Na? and Ca2? influx and the membrane potential of GLChR2 cells by blue light exposure To clarify whether ChR2 functioned on GL261 cells with blue light exposure, intracellular Na? concentration was

Measurement of extracellular glutamate GLChR2 cells were plated in a 60-mm Cell Culture Dish at a density of 2.0 9 105 cells and were cultivated in the presence or absence of 100 lM EDTA, 1 lM TTX, 10 lM verapamil, 1 lM riluzole, or 1 lM ionomycin with or without blue light exposure for 72 h. They were exposed in the culture medium to blue light for 1 min every 5 min until 72 h using the Safe Imager Blue-Light Transilluminator (Life Technologies, Carlsbad, CA, USA). The culture medium was collected on Day 1 and Day 3. For the measurement of glutamate concentration, YAMASA L-Glutamate Assay Kit II (Yamasa Corporation, Tokyo, Japan) was used. Each culture medium was mixed with the enzyme reagent according to the manufacturer’s instructions, the absorbance at 595 nm was measured, and the concentration of glutamate was calculated from a standard curve. Statistical analysis Statistical analysis was performed using a two-tailed t test or one-way ANOVA and post hoc tests. Differences were considered significant when the p value was less than 0.05.

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Fig. 1 GL261 cells (GLChR2 cells) transfected with the ChR2-EYFP gene. The gene transfer and expression of ChR2 and EYFP mRNA was confirmed by RT-PCR (a). The expression of the ChR2-EYFP fusion protein in GLChR2 cells was detected by FACS (b). The filled histogram indicates GL261 cells and the solid line indicates GLChR2 cells. Photographs of GLChR2 cells (DIC and ChR2-EYFP) were taken using a fluorescent microscope (c). Scale bar 50 lm. DIC differential interference contrast

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monitored by a CoroNa Red fluorescent probe, which is one of the Na? indicators and increases red fluorescence in a concentration-dependent manner of intracellular Na?. Although intracellular fluorescence did not increase in GL261 cells with blue light exposure, it significantly increased in GLChR2 cells (Fig. 2a, b). Increase of CoroNa Red fluorescence in photo-activated GLChR2

cells was not observed in Na?-free buffer (Fig. 3a), indicating extracellular Na? influx. Intracellular Ca2? concentration was then monitored by a Rhod-3 fluorescent probe, which is one of the Ca2? indicators. Intracellular fluorescence apparently increased in GLChR2 cells with blue light (Fig. 2c, d). Accompanied with increase of intracellular sodium and calcium ions, the

Fig. 2 Na? and Ca2? influx and the membrane potential of GLChR2 cells by blue light exposure. Na? influx in GLChR2 cells labeled with CoroNa Red, by blue light (BL) exposure was observed using timelapse imaging. a Photographs showed fluorescent images of GL261 cells and GLChR2 cells at 0, 2.5, 5, and 7.5 min with BL exposure. Scale bar 25 lm. The intracellular red fluorescence of CoroNa Red was measured and the results were summarized in a graph (b). Values represent the mean ± SD. *p \ 0.05, **p \ 0.01, ***p \ 0.001 vs GL261 cells at the same time point. Ca2? influx in GLChR2 cells labeled with Rhod-3, by BL exposure was observed using time-lapse imaging. c Photographs showed fluorescent images of GLChR2 cells at 0–10 min without BL and at 10–20 min with BL. Scale bar 50 lm.

The intracellular red fluorescence of Rhod-3 was measured and the results were summarized in a graph (d). Values represent the mean ± SD. *p \ 0.05, **p \ 0.01, ***p \ 0.001 vs GLChR2 cells without BL at the same time point. The membrane potential of GLChR2 cells was visualized by the DiBAC4(5) red fluorescent probe with or without BL exposure using time-lapse imaging. e Photographs showed fluorescent images of GLChR2 cells at 0, 10, 20, and 30 min with or without BL. Scale bar 50 lm. The intracellular red fluorescence of DiBAC4(5) was measured and the results were summarized in a graph (f). Values represent the mean ± SD. **p \ 0.01, ***p \ 0.001 vs GLChR2 cells without BL at the same time point

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Fig. 3 Endogenous ion channels involved in alteration of intracellular ion environment in photo-activated GLChR2 cells. The intracellular red fluorescence of CoroNa Red was measured in the sodium ion-free buffer (a), or in RB in the presence or absence of EDTA (100 lM), TTX (1 lM), verapamil (10 lM) or riluzole (1 lM) (b) at 0–10 min with or without BL and alteration of fluorescence during 10 min was summarized in each graph. The intracellular red

fluorescence of Rhod-3 was measured in the sodium ion-free buffer (c), or in RB in the presence or absence of EDTA, TTX, verapamil or riluzole (d) at 0–10 min with or without BL and alteration of fluorescence during 10 min was summarized in each graph. Values represent the mean ± SD. *p \ 0.05, **p \ 0.01, ***p \ 0.001 vs GLChR2 cells without BL (N). #p \ 0.05, ##p \ 0.01, ###p \ 0.001 vs GLChR2 cells with BL. Sodium-free sodium ion-free

membrane potential of GLChR2 cells was significantly depolarized by blue light exposure monitored by the DiBAC4(5) red fluorescent indicator (Fig. 2e, f). To know whether endogenous ion channels are involved in the alteration of intracellular sodium and calcium under photo-activation, GLChR2 cells were exposed to blue light in the presence or absence of inhibitors of Na? and Ca2? influx such as EDTA, a non-selective chelator; TTX, a kind of sodium channel blocker; verapamil, a voltage-activated calcium channel blocker; and riluzole, a blocker for both sodium channels and calcium channels (Fig. 3). Increase of CoroNa Red fluorescence in photo-activated GLChR2 cells was observed even in the presence of EDTA, TTX, verapamil, or riluzole (Fig. 3b). Since ChR2 is not affected by these inhibitors, Na? influx in photo-activated GLChR2 cells was only via ChR2. On the other hand, Rhod-3 fluorescence in photo-activated GLChR2 cells was decreased to less than baseline in sodium ion-free buffer (Fig. 3c). In addition, it was significantly decreased in the presence of EDTA, TTX, verapamil, or riluzole (Fig. 3d). These results suggested that Na? influx through photoactivated ChR2 triggered Ca2? influx via endogenous channels.

Alterations in the cellular functions of GLChR2 cells by blue light exposure

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We examined whether alterations in the intracellular ionic environment of GLChR2 cells affected intracellular pH. Intracellular pH was monitored by carboxy SNARF-1 AM, which displays red fluorescence at an alkaline pH that disappears as the pH becomes acidic. Intracellular red fluorescence was observed in both GL261 cells and GLChR2 without blue light stimulation. Red fluorescence in GL261 did not change by light exposure, whereas that in GLChR2 decreased by blue light exposure (Fig. 4a, b) indicating that the intracellular pH of GLChR2 cells became acidic with blue light exposure. As it has been shown that intracellular acidification inhibits the proliferative response (Lucas et al. 1988), we compared the WST reduction activity, an indicator of cell number, of GL261 and GLChR2 cells at Day1 and Day3 with and without blue light exposure (Fig. 4c). The WST reduction activity of GL261 cells and GLChR2 cells was similar on Day 1 with or without blue light exposure, whereas that of GLChR2 cells was significantly decreased on Day 3

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Fig. 4 Decrease in intracellular pH resulted in the inhibition of proliferation in photo-activated GLChR2 cells. Intracellular pH in GL261 cells and GLChR2 cells labeled with carboxy SNARF-1 AM, by BL exposure was observed using time-lapse imaging. a Photographs showed fluorescent images of GL261 cells and GLChR2 cells at 0, 5, 10, 15, and 20 min with BL exposure. Scale bar 50 lm. Intracellular red fluorescence of carboxy SNARF-1 AM was measured and the results were summarized in a graph (b). Values represent the mean ± SD. *p \ 0.05, **p \ 0.01, ***p \ 0.001 vs

GL261 cells without BL at the same time point. c The proliferation of GL261 cells and GLChR2 cells was investigated on Day 1 and Day 3 with or without BL exposure by the WST assay. Values represent the mean ± SD. ***p \ 0.001 vs GL261 cells without BL. d The apoptosis of GLChR2 cells by BL exposure was investigated by FACS analysis. The ratio of PI and Annexin V expression in GLChR2 cells were measured on Day 3 with BL (BL) and without BL (N). The numbers in each density plot indicate the percentage of PI? and Annexin V? cells

with blue light (Fig. 4c). To confirm whether the decrease in WST reduction activity on Day 3 was due to decrease of proliferative capacity but not to cell death, GLChR2 cells were analyzed by FACS stained with propidium iodide (PI) and Annexin V (Fig. 4d).

There were no differences in the ratio of PI and Annexin V in GLChR2 cells with or without blue light exposure (Annexin V ?PI- GLChR2 cells without blue light exposure; 0.84 ± 0.26 %, with blue light; 1.14 ± 0.32 %, p = 0.276, Annexin V?PI? GLChR2

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Fig. 5 Glutamate release from photo-activated GLChR2 cells. a The concentration of glutamate in the culture medium from GL261 cells and GLChR2 cells was measured on Day 1 and Day 3 with BL (BL) and without BL (N). Values represent the mean ± SD. **p \ 0.01, ***p \ 0.001 vs GL261 cells without BL. b The concentration of glutamate in the culture medium was measured from GLChR2 cells in the presence or absence of EDTA (100 lM), TTX (1 lM), verapamil (10 lM), riluzole (1 lM), or ionomycin (1 lM) on Day 1 with and without BL. Values represent the mean ± SD. *p \ 0.05, **p \ 0.01, ***p \ 0.001 vs GLChR2 cells without BL, ### p \ 0.001 vs GLChR2 cells with BL. c The proliferation of GLChR2 cells was investigated in the presence or absence of NBQX (10 lM) on Day 3 with or without BL exposure by the WST assay. Values represent the mean ± SD. ***p \ 0.001 vs GLChR2 cells without BL

cells without blue light exposure; 0.94 ± 0.25, with blue light; 1.19 ± 0.29, p = 0.321). These results indicated that decreases in cell number (WST reduction

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activity) resulted not from inducible cell death, but from the inhibition of proliferation in response to the blue light.

Glutamate release from astrocyte cell-line GL261

Glutamate release from GLChR2 cells by blue light exposure Astrocytes release gliotransmitters such as glutamate and ATP partly in a calcium-dependent manner for communication between glial cells and also glial cells and neurons (Hamilton and Attwell 2010). We indicated increase of intracellular Na? and Ca2? in GLChR2 by blue light exposure, therefore, we examined whether gliotransmitter release in GLChR2 was induced by blue light; glutamate release was determined by measuring glutamate concentrations of the culture medium (Fig. 5a). The glutamate release from GLChR2 cells without blue light was similar to that from GL261 cells on Day 1 and Day 3 with or without blue light. However, the glutamate release from GLChR2 cells with blue light was 2- to 3-fold higher than that from GL261 cells and GLChR2 cells without blue light on Day 1 and Day 3. To confirm that light-induced glutamate release from GLChR2 cells was calcium dependent, GLChR2 cells were exposed to blue light in the absence or presence of EDTA (100 lM), TTX (1 lM), verapamil (10 lM), riluzole (1 lM), or ionomycin (1 lM) (Fig. 5b). Although the glutamate release was apparently increased by blue light, it returned to baseline levels by EDTA, a non-selective chelator, and verapamil, a voltage-activated calcium channel blocker. In addition, the glutamate release from GLChR2 cells with blue light was slightly decreased by treatment with TTX, a kind of sodium channel blocker. Furthermore, the glutamate release from GLChR2 cells was drastically decreased by treatment with riluzole, which blocks both voltage-dependent sodium channels and voltage-activated calcium channels (Narahashi 2000). On the other hand, ionomycin which raises the intracellular level of Ca2? significantly increased glutamate release from GLChR2 cells, with or without blue light. It is possible that glutamate release results in induction of astrocyte proliferation (Liao and Chen 2001) via several glutamate receptors such as AMPA receptors and kainate receptors expressed on them. Therefore, we examined whether glutamate from GLChR2 cells affected proliferation by autocrine signaling (Fig. 5c). The proliferation of GLChR2 cells with blue light exposure was not significantly altered in the presence of NBQX, a glutamate receptor antagonist on Day 3. Glutamate from photo-activated GLChR2 cells acted on N18 neuronal cells Glutamate is one of gliotransmitters from astrocyte to neurons. To investigate whether glutamate released from photo-activated GLChR2 cells affected neuronal cells, we examined neuron–glia co-culture. Ca2? increase in N18 neuronal cells was measured when GLChR2 cells were

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exposed to blue light for 10 min (Fig. 6a). Increase of Ca2? concentration in N18 cells was not observed until 8 min, but it increased transiently after 8 min exposure and then decreased. Increase of Ca2? concentration in N18 cells was not observed in the presence of NBQX, suggesting that N18 cells responded to glutamate from photo-activated GLChR2 cells and resulted in increase of intracellular Ca2? concentration. In general, glutamate derived from astrocytes has both bifacial and harmful effects on neurons such as being a transmitter and a neurotoxin, respectively. Then we examined the neurotoxicity of glutamate in N18 cells. The proliferation of N18 cells was inhibited by glutamate in a concentration-dependent manner (Fig. 6b). When N18 cells were stimulated with a high concentration of glutamate (500 lM) for 3 days, apoptosis was induced in N18 cells (Annexin V?PI? cells, 0 lM; 1.41 ± 0.42 %, 500 lM; 20.35 ± 4.07 %, p \ 0.01) (Fig. 6c). We also observed induction of N18 apoptosis in the co-culture with photo-activated GLChR2 cells (Fig. 6d, e). The WST reduction activity of the co-culture was significantly decreased by blue light exposure on Day 3, but NBQX prevented the decrease (Fig. 6d). On FACS analysis N18 cells were easily distinguished from GLChR2 cells by EYFP fluorescence. Apoptotic N18 cells (PI? Annexin V?) were significantly increased in the co-culture with blue light exposure (PI? Annexin V?, 18.41 % ± 1.57) compared to the control (PI? Annexin V?, 4.88 % ± 0.42). NBQX prevented the increase of apoptosis of N18 to the control level (PI? Annexin V?, 5.83 % ± 0.49) (Fig. 6e). Apoptotic GLChR2 cells in the co-culture were not found in the presence or absence of NBQX with or without blue light exposure.

Discussion In this study, we demonstrated that the forced expression of ChR2 in GL261 astrocytes could function as a cation channel regulating the influx of Na? and Ca2? by blue light exposure. Alterations of membrane potentials via cation influx in GL261 astrocytes resulted in the inhibition of cell proliferation (Fig. 4) and promotion of glutamate release (Fig. 5) from these cells in a calcium-dependent manner. Glutamate is known to be one of the gliotransmitter as well as an excitatory neurotransmitter; therefore, it may play important roles in interactions among neurons and glial cells (Jourdain et al. 2007; Perea and Araque 2007). Previous studies reported that glutamate was involved in cognitive functions like learning and memory in the brain (McEntee and Crook 1993; Woo et al. 2012). Further, a recent study demonstrates that ChR2-expressing astrocytes modulate neuronal activity by photo-stimulation using the slice and in vivo system (Sasaki et al. 2012).

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In addition, excitotoxicity due to glutamate was associated with many diseases such as cerebral ischemia (Tymianski 2011), amyotrophic lateral sclerosis (Shaw and

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Ince 1997), epileptic seizure (Cendes et al. 1995), Parkinsonism, and Alzheimer’s disease (Coulter and Eid 2012). In many brain diseases as well as normal conditions,

Glutamate release from astrocyte cell-line GL261 b Fig. 6 Apoptosis of neuronal cells by glutamate release from photo-

activated GLChR2 cells. a The co-culture of N18 cells with GLChR2 cells, which was loaded with Rhod-3, was exposed to BL every 15 s for 10 min. Photographs showed representative serial images (DIC and fluorescence) of the co-culture from 8 min. Arrows indicate that intracellular red fluorescence increased in N18 cells. Scale bar 50 lm. b The proliferation of N18 cells was investigated in the presence of glutamate (0–800 lM) by the WST assay. Values represent the mean ± SD. *p \ 0.05, ***p \ 0.001 vs N18 cells in the absence of glutamate (0 lM). c The apoptosis of N18 cells by glutamate was investigated by FACS analysis. The ratio of PI and Annexin V expression in N18 cells was measured in the absence or presence of 500 lM glutamate on Day 3. The numbers in each density plot indicate the percentage of PI? and Annexin V? cells. d When N18 cells were co-cultivated with GLChR2 cells, the proliferation of N18 cells and GLChR2 cells was investigated in the presence or absence of NBQX (10 lM) on Day 3 with or without BL exposure by the WST assay. Values represent the mean ± SD. ***p \ 0.001 vs the co-culture in the absence of NBQX without BL exposure. e The apoptosis of N18 cells and GLChR2 cells in the co-culture was investigated by FACS analysis. GLChR2 cells were distinguished from N18 cells by EYFP fluorescence. The ratio of PI and Annexin V expression in N18 cells and GLChR2 cells were measured in the presence or absence of NBQX (10 lM) on Day 3 with or without BL exposure. The numbers in each density plot indicate the percentage of PI? and Annexin V? cells

astrocytes become a source of extracellular glutamate (Hamilton and Attwell 2010), hence it is important to understand the roles of glutamate released from astrocytes. In this study, we demonstrated that glutamate release from astrocyte was controlled by photo-activated ChR2. As it was controlled by blue light exposure, ChR2-expressing astrocytes might become a useful tool for clarifying interaction and reaction via glutamate between neurons and astrocytes or between other glial cells and astrocytes in vitro and in vivo. Our results showed that glutamate from GLChR2 cells displayed bifacial aspects. Glutamate released in a short and transient term, for example, for 10 min in our experiment, promoted neuronal activation; we demonstrated intracellular Ca2? increase in N18 neuronal cells co-cultured with photo-activated GLChR2 (Fig. 6a). On the other hand, glutamate released in a long-lasting time, for example, for 3 days in our experiment, induced neuronal damage. Both effects were mediated via glutamate receptors because NBQX prevented them (Fig. 6d, e). As the amount of glutamate released from photo-activated GLChR2 was dependent on length of light exposure (Fig. 5a), GLChR2 might be controlled in its characteristics for either neuronal activity regulation or neuronal apoptosis by time of exposure. Therefore, this study suggested that ChR2-expressing astrocytes could become a useful tool to understand the roles of glial cell activation and neural communication in the regulation of brain functions. Deprivation of Na? but not Ca2? from culture media prevented the increase of intracellular Na? concentration in photo-activated GLChR2 cells (Fig. 3), indicating that the

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increase was due to Na? influx from extracellular environment; whereas, either deprivation of Na? or Ca2? prevented the increase of intracellular Ca2? concentration in photo-activated GLChR2 cells suggesting that the increase of Ca2? was due to Ca2? influx from extracellular environment triggered by Na? influx. We also demonstrated that treatment of photo-activated GLChR2 cells with TTX, verapamil, or riluzole prevented Ca2? influx, but not Na? influx (Fig. 3c, d). As TTX and verapamil block endogenous sodium and calcium channels, respectively (Zhang and Oertner 2007), without any blocking effect on ChR2, endogenous sodium and calcium channels should be associated with Ca2? influx triggered by ChR2 activation. A previous study reported that the increase in intracellular Ca2? induced glutamate release from astrocytes (Parpura et al. 2011). Since glutamate release from photoactivated GLChR2 cells correlated with concentrations of intracellular Ca2? (Fig. 5b) and ionomycin, a calcium ionophore, promoted glutamate release without photoactivation, these results indicate that the increase in Ca2? influx triggered glutamate release from GLChR2 cells. Furthermore, riluzole fully inhibited glutamate release from photo-activated GLChR2 cells. Riluzole acts as not only a blocker of both sodium and calcium ion channels and prevents glutamate release (Wang et al. 2004; Lamanauskas and Nistri 2008), but also as an activator glutamate transporters and enhancer of glutamate uptake (Frizzo et al. 2004). In this study, riluzole seemed to block endogenous Na? and Ca2? channels, which was involved in the increase in Ca2? influx by photo-activated ChR2 and moreover remove glutamate through activated glutamate transporter. Our findings suggested that Na? influx through photo-activated ChR2 should trigger functional Ca2? influx via endogenous channels and result in increases in glutamate release. We also indicated a possibility that alteration of intracellular ion environment might control proliferation of astrocytic cells. The proliferative capacity of GLChR2 cells was significantly decreased by blue light exposure for 3 days (Fig. 4c) in a parallel with intracellular acidification (Fig. 4a, b), but not in parental GL261 cells. It has been shown that intracellular acidification inhibited the proliferative response (Lucas et al. 1988). Thus, it may be possible that astrocytic proliferation, especially gliosis and glioma, can be controlled by light exposure with intracellular acidification via ChR2. This may provide a new technique to manipulate ectopic gliosis or malignant glioma. In conclusion, we demonstrated that the properties of ChR2-expressing astrocytes were controlled by blue light exposure, and cation influx through photo-activated ChR2 could trigger functional Ca2? influx via endogenous channels and resulted in both inhibition of proliferation of

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astrocytic cells and increase of glutamate release. Further, released glutamate from astrocytes by photo-activated ChR2 acted on neuronal cells as both a transmitter and neurotoxin. Our results suggest that ChR2-expressing glial cells could become a useful tool in understanding the roles of glial cell activation and neural communication in the regulation of brain functions. Acknowledgments We thank Dr. Karl Deisseroth (Stanford University) for providing expression vectors. This study was supported by the Industrial Technology Research Grant Program from the New Energy and Industrial Technology Development Organization (NEDO) of Japan, the Hori Information Science Promotion Foundation, and the Research Foundation for Opto-Science and Technology. Conflict of interest

The authors declare no conflict of interest.

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