Channel-mediated astrocytic glutamate modulates hippocampal synaptic plasticity by activating postsynaptic NMDA receptors
- Hyungju Park†1,
- Kyung-Seok Han†1, 2,
- Jinsoo Seo†3,
- Jaekwang Lee†1,
- Shashank M Dravid4,
- Junsung Woo1, 2,
- Heejung Chun1,
- Sukhee Cho3,
- Jin Young Bae5,
- Heeyoung An1, 6,
- Woohyun Koh1, 2,
- Bo-Eun Yoon1, 7,
- Rolando Berlinguer-Palmini8,
- Guido Mannaioni9,
- Stephen F Traynelis10,
- Yong Chul Bae5,
- Se-Young Choi3Email author and
- C Justin Lee1, 2, 6Email author
© Park et al.; licensee BioMed Central. 2015
Received: 19 December 2014
Accepted: 15 January 2015
Published: 3 February 2015
Activation of G protein coupled receptor (GPCR) in astrocytes leads to Ca2+-dependent glutamate release via Bestrophin 1 (Best1) channel. Whether receptor-mediated glutamate release from astrocytes can regulate synaptic plasticity remains to be fully understood.
We show here that Best1-mediated astrocytic glutamate activates the synaptic N-methyl-D-aspartate receptor (NMDAR) and modulates NMDAR-dependent synaptic plasticity. Our data show that activation of the protease-activated receptor 1 (PAR1) in hippocampal CA1 astrocytes elevates the glutamate concentration at Schaffer collateral-CA1 (SC-CA1) synapses, resulting in activation of GluN2A-containing NMDARs and NMDAR-dependent potentiation of synaptic responses. Furthermore, the threshold for inducing NMDAR-dependent long-term potentiation (LTP) is lowered when astrocytic glutamate release accompanied LTP induction, suggesting that astrocytic glutamate is significant in modulating synaptic plasticity.
Our results provide direct evidence for the physiological importance of channel-mediated astrocytic glutamate in modulating neural circuit functions.
KeywordsAstrocytes Bestrophin 1 Ca2+-activated anion channel Synaptic plasticity Glutamate NMDA receptor LTP PAR1
Growing evidence has supported the idea that astrocytes are actively involved in modulating synaptic strength by affecting neuronal properties [1-3]. At glutamatergic synapses, astrocytically released glutamate has been suggested to play a crucial role in mediating neuronal-glial circuits. Astrocytes not only clear presynaptically released glutamates during synaptic transmission, but can also release glutamate via diverse pathways such as soluble NSF attachment protein receptor (SNARE)-dependent exocytosis [4-6] and the glutamate permeable anion channel [7-11], in response to increased intracellular Ca2+ concentration by activation of G-protein coupled receptors expressed at the astrocytic membrane. In turn, this Ca2+-dependent glutamate release from astrocytes can be sensed by presynaptic or postsynaptic glutamate receptors such as the metabotropic glutamate receptor (mGluR) [6,12] or NMDAR [13,14], both of which are known to modify presynaptic and postsynaptic activities, or synaptic plasticity.
However, the role of astrocytes in synaptic function is still in question, because recent studies have given contradictory reports of the involvement of receptor-mediated Ca2+ signals in astrocytic glutamate release [15-18]. In order to pinpoint the exact role of astrocytic glutamate in synaptic functions, we thus have searched for an effective and reliable tool for triggering glutamate release from astrocytes. Mounting evidence supports the ability of PAR1 to trigger the Ca2+-dependent signaling pathways crucial for astrocytic glutamate release. Activated by endogenous agonist (thrombin, plasmin) or TFLLR-NH2 peptide agonist (TFLLR) [19,20], PAR1 can elevate astrocytic intracellular Ca2+ levels via downstream pathways associated with Ca2+ release from internal stores [19,20]. In addition, PAR1 activation was shown to be effective for triggering Ca2+-dependent astrocytic glutamate release when compared with the activation by other GPCRs [14,21]. Furthermore, due to selective and functional expression of PAR1 in astrocytes in the hippocampal CA1 area [20,22], PAR1 signaling has the additional advantage of inducing astrocyte-specific Ca2+-dependent signaling in the hippocampal CA1 area without affecting pre- or postsynaptic neurons. Therefore, PAR1 appears to be a useful tool for selective induction of Ca2+-dependent glutamate release from astrocytes in vitro and in vivo [10,11,14,20,23-26].
Our previous studies have shown that PAR1 activation in hippocampal CA1 astrocytes leads to Ca2+-dependent opening of the glutamate-permeable anion channel, Best1, which mediates Ca2+-dependent astrocytic glutamate release [8,10,11,26]. Not only does the Best1 channel displays a glutamate permeability that is Ca2+-dependent , but it also has a preferential subcellular localization at the microdomains of hippocampal astrocytes located around synaptic terminals . These studies suggest that PAR1-induced Ca2+ elevation at the microdomain directs glutamate release through the Best1 channel, resulting in an increase in glutamate concentration at synaptic clefts. Moreover, Best1-mediated astrocytic glutamate release triggered by PAR1 activation may play a role in modulating in synaptic plasticity, as recent studies show that PAR1-deficient mice display reduced NMDAR-dependent hippocampal LTP and contextual fear memory .
To explore the target and physiological consequences of Best1-mediated glutamate release from astrocytes, we triggered Ca2+-dependent glutamate release from hippocampal CA1 astrocytes by activating PAR1, and examined the effect of astrocytic glutamate on neurotransmission. We demonstrated that synaptic NMDAR is the main target of astrocytic Best1-mediated glutamate, and increased synaptic NMDAR activation leads to NMDAR-dependent potentiation of synaptic transmission. Of equally importance, we also identified an altered NMDAR-dependent synaptic plasticity at hippocampal synapses, when synaptic glutamate was increased by Best1-mediated secretion of glutamate from astrocytes. As well as verifying the functional expression of the mechanism for receptor-mediated glutamate release in astrocytes, our findings provide direct evidence for the involvement of astrocytic anion channel-mediated glutamate release in synaptic modification.
Astrocytes release glutamate via Best1 channel upon PAR1 activation
Of more importance, immunohistochemical analysis by co-staining endogenous PAR-1 and Best1 proteins in the CA1 area, showed that both PAR1 and Best1 are highly co-localized in CA1 astrocytes (PAR1/Best1: 79.8 ± 1.6%, n = 10; Best1/PAR1: 80.5 ± 1.6%, n = 10; Figure 1C). Because the astrocytic Best1 channel is localized at the microdomain of astrocytic processes near the synaptic region (Figure 1D) , and Ca2+-activated Best1 channel showed a significant permeability to glutamate in hippocampal astrocytes , our finding raises a possibility that glutamate release through Best1 channel at astrocytic microdomains could affect synaptic glutamate concentration.
To directly test whether PAR1 activation can induce astrocytic glutamate release through Best1 channel, we monitored extracellular glutamate by using fluorescence resonance energy transfer (FRET)-based glutamate sensor GluSnFR (a glutamate-sensing fluorescent reporter) . This GluSnFR was expressed at the membrane of CA1 astrocytes in hippocampal slices to detect glutamate released from astrocytes. Control experiments showed that astrocytic GluSnFR sensors were able to detect extracellular glutamate in a range from 10−3 to 10−6 M (Figure 1E), similar to that found in cultured astrocytes . We found that bath application of the PAR1 agonist, TFLLR (30 μM) [8,20,26] increases extracellular glutamate level (at the peak; 8.5 ± 1.9 μM, n = 5) around a single CA1 astrocyte, and that this elevation of extracellular glutamate level was significantly reduced in slices of Best1 knockout mice (Best1 KO : Figure 1F,G). In line with previous findings [8,10,11,26], these results indicate that PAR1 activation-triggered astrocytic glutamate release is mediated by Best1 channels, which possibly permeate intracellular glutamate into extracellular synaptic clefts.
Best1-mediated astrocytic glutamate enhances basal synaptic transmission
Our data showed that application of TFLLR to slices expressing Best1-shRNA globally in the CA1 region (mock-treated hGFAP-CreERT2 mice that were injected with pSicoR-Best1-shRNA) did not produce a TFLLR-induced increase in amplitude of eEPSPs (% baseline, 103.9 ± 15.7, n = 7 slices; Figure 3D,E), as observed from slices of Best1 KO mice (Figure 2C,D). However, when pSicoR-Best1-shRNA was injected into tamoxifen-treated hGFAP-CreERT2 mice (to recover Best1 expression specifically in astrocytes), TFLLR treatment was sufficient for inducing potentiated eEPSP responses (% baseline, 202.6 ± 45.0, n = 6 slices; Figure 3D,E). By contrast, an attempt to rescue Best1 expression in CA1 neurons, by injecting pSicoR-Best1-shRNA into CaMKIIα-Cre mice that express the Cre recombinase in CA1 pyramidal neurons , did not produce eEPSP potentiation upon TFLLR application (% baseline, 96.6 ± 16.9, n = 7 slices; Figure 3D,E). Together, these data indicate that the astrocyte-specific Best1 channel is required for enhanced basal synaptic transmission induced by astrocytic glutamate secretion upon PAR1 activation, implying a role of Best1-mediated glutamate in modulating synaptic activity.
Best1-mediated astrocytic glutamate elevates synaptic glutamate
We next explored how astrocytic glutamate may enhance synaptic activity. Given the preferential distribution of astrocytic Best1 channels at microdomains that wrap around excitatory synaptic structures , the glutamate concentration at synaptic clefts may be directly affected by astrocytic glutamate release.
We also showed that the increased τdecay of CTZ-induced AMPAR-EPSCs by TFLLR application was mediated by selective activation of astrocytic PAR1, because TFLLR application in the presence of CTZ was unable to induce an increase in τdecay of AMPAR-EPSCs measured from hippocampal slices of PAR1 knockout mice (PAR1 KO) (% increase in τdecay, wild type: 52.6 ± 11.6, n = 5 slices; PAR1 KO: 6.9 ± 7.9, n = 5 slices; Figure 4C,D) or in mice expressing astrocyte-specific Par1-shRNA (% increase in τdecay, scrambled shRNA: 68.1 ± 11.8, n = 5 slices; Par1-shRNA: 11.2 ± 28.5, n = 5 slices; Figure 4E-G). These results indicate that the prolonged activation of synaptic AMPARs is caused by an elevated synaptic glutamate level that results from astrocytic glutamate upon PAR1 activation. To examine whether Best1 channels are required for the TFLLR-induced increase in synaptic glutamate concentration, we compared the increased τdecay of AMPAR-EPSCs by PAR1 activation in Best1 KO hippocampal slices with that in wild type slices. Our data showed that TFLLR application fails to induce an increase in τdecay of AMPAR-EPSCs from hippocampal slices of Best1 KO mice as shown in slices of wild type mice (% increase in τdecay, wild type: 48.4 ± 10.9, n = 10 slices; Best1 KO: 15.6 ± 4.6, n = 7 slices; Figure 4H,I). Taken together, these results indicate that astrocytic glutamate released via Best1 channel targets synaptic clefts, causing an elevated synaptic glutamate concentration when combined with presynaptically released glutamate.
By contrast, we found no significant effect of TFLLR application on amplitudes and τdecay of normal AMPAR-EPSCs in the absence of CTZ (% amplitude of AMPAR-EPSCs after TFLLR application = 102.0 ± 7.6, n = 8 slices; τdecay after TFLLR application = 11.3 ± 1.0 ms, n = 9 slices; Figure 4J,K). In addition, presynaptic release properties were not affected by astrocyte-mediated increased synaptic glutamate, because PAR1 activation did not cause any significant alteration in the paired-pulse facilitation (PPF) of AMPAR-EPSCs (% potentiation; 20 ms IPI, control: 97.3 ± 10.8, TFLLR: 105.2 ± 10.6, n = 4 slices; 50 ms IPI, control: 79.7 ± 11.1, TFLLR: 85.1 ± 11.1, n = 5 slices; Figure 4L,M). These results suggest that elevated synaptic glutamate level by astrocytic glutamate affect neither postsynaptic AMPARs nor presynaptic glutamate receptors that can regulate the release probability. In support of this view, our previous study demonstrated unaltered frequencies and amplitudes of AMPAR-mediated miniature EPSCs (mEPSCs) upon PAR1 activation .
Synaptic NMDARs are activated by Best1-mediated astrocytic glutamate
Because both presynaptic glutamate receptor and postsynaptic AMPAR are unaffected by synaptic glutamate elevation (Figure 4J-M), we next tested whether activity of synaptic NMDARs could be enhanced by astrocytic glutamate. We recorded NMDAR-mediated whole-cell currents at SC-CA1 synapses in the presence of low external Mg2+ (5 μM) and tetrodotoxin (1 μM), as previously used for showing that NMDAR-dependent whole-cell currents can be generated from CA1 neurons by Best1-mediated glutamate released upon PAR1 activation .
In order to directly examine whether the synaptic NMDAR activity can be enhanced by astrocytic glutamate, we isolated and measured NMDAR-mediated evoked EPSPs (NMDAR-EPSP: Figure 5C,D) at SC-CA1 synapses in the presence of 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX) and bicuculline. After TFLLR application, the amplitude of evoked NMDAR-EPSPs was gradually increased (% baseline after TFLLR application: 144.2 ± 18.6%, n = 7 slices; Figure 5C,D), consistent with the idea that the activity of synaptic NMDAR is increased by astrocytic synaptic glutamate elevation. When opened synaptic NMDARs during synaptic transmission were fully blocked by bath application with MK-801 (50 μM; Figure 5C), we could not detect any increase in amplitude of NMDAR-EPSP on TFLLR application (% baseline after TFLLR application: 87.1 ± 11.3%, n = 7 slices; Figure 5C,D). Together with evidence for the direct activation of GluN2A-sensitive NMDAR by astrocytic glutamate (Figure 5A,B), these results support the notion that GluN2A-containing synaptic NMDARs are the major target of astrocytic glutamate, and our observation of NMDAR-dependent synaptic strengthening is resulted from increased activation of synaptic NMDARs by astrocytic synaptic glutamate elevation (Figure 2).
Best1-mediated astrocytic glutamate modulates synaptic plasticity
On the other hand, selective activation of astrocytic GPCR, Mas-related G-protein coupled receptor member A1 (MrgA1) by applying its agonist, FMRFamide peptide (FMRFa; ~ 30 min), to hippocampal slices of transgenic mice selectively expressing MrgA1 in astrocytes , was not efficient for altering fEPSP responses (Figure 6F). Moreover, in contrast to PAR1 activation, MrgA1 activation by FMRFa application could not alter the magnitude of LTD compared with control (% baseline, MrgA1: 86.9 ± 5.2, n = 5 slices; MrgA1 + FLRF: 89.5 ± 7.8, n = 4 slices; Figure 6G,H). These data are in agreement with previous reports showing that an increase in astrocytic Ca2+ by MrgA1 activation is not linked to the mechanisms responsible for triggering astrocytic glutamate release [15,16].
Consistent with previous reports [14,15,17,38], activation of astrocytic GPCRs, with the exception of PAR1, was not sufficient to trigger glutamate release from astrocytes (Figures 6F and 7A), supporting the idea that glutamate released upon astrocytic PAR1 activation has a specific modulatory role in synaptic function. The reason that PAR1 is more effective than other GPCRs for inducing glutamate release from astrocytes remains to be determined, but it is possible that activation of MrgA1 or other GPCRs only causes an insufficient Ca2+ levels at microdomains for triggering Ca2+-dependent glutamate release mechanisms . On the other hand, either MrgA1 or other GPCRs may not be associated with the astrocytic molecular mechanisms responsible for glutamate release, whereas PAR1 is linked to multiple channel-mediated glutamate release mechanisms such as G protein-dependent opening of the two-pore K+ channel (TREK-1) and Ca2+-dependent opening of the Best1 channel . Thus, if PAR1 activation occurs during the process of synaptic plasticity, astrocytic glutamate released through channel-mediated mechanisms may play a significant role in modulating hippocampal synaptic plasticity and cognitive functions. In support of this idea, it has been shown that activation of PAR1 by thrombin (a native PAR1 agonist) or TFLLR modulates LTP formation at hippocampal synapses [27,39], and that a reduction in NMDAR-dependent hippocampal LTP and contextual fear memory were observed in PAR1 deficient mice . How astrocytic PAR1 is activated during the process of synaptic plasticity and memory formation is still unclear, but we suggest that astrocytic PAR1 might be activated by endogenous PAR1 agonists that are generated from secreted proteases or their byproducts in an activity-dependent manner. For example, plasmin, which is cleaved from plasminogen by the extracellular tissue plasminogen activator (tPA), can act as a native agonist for PAR1 in the hippocampus , and activity-dependent secretion of tPA from postsynaptic dendrites [40,41] is required for inducing late-phase LTP (L-LTP) at hippocampal synapses [42,43].
Our study has also provided evidence for the involvement of PAR1-triggered astrocytic glutamate in synaptic function by showing that the Best1 channel is required for synaptic NMDAR activation. Ultrastructural analyses showed that Best1 channels are preferentially expressed at the membrane of astrocytic microdomains around synaptic terminals in the hippocampus (Figure 1), proposing a possible influence of Best1-mediated glutamate on the synaptic glutamate level . Consistent with this, we have shown that PAR1 activation prolongs the CTZ-induced AMPAR-EPSC response, an action indicative of increased synaptic glutamate levels, and that this prolonged AMPAR-EPSC response was dependent on Best1 expression (Figure 4). Because the amount of glutamate released through the Best1 channel was much lower (10−5 ~ 10−6 M; Figure 1E-G) than that from presynaptically released glutamate (~10−3 M), a subtle increase in synaptic glutamate levels by PAR1 activation thus had no significant effect on synaptic glutamate receptors, other than NMDARs [2,20,39], as shown by the unchanged presynaptic release probability and AMPAR-mediated postsynaptic activity (Figure 4J-M). Thus, we suggest that enhanced NMDAR-dependent signaling by receptor-mediated astrocytic glutamate release can produce potentiation of AMPAR-mediated synaptic transmission (Figure 2), because AMPAR expression at the postsynaptic membrane could be elevated by activation of NMDAR-dependent signaling [44,45]. However, our study do not exclude the possibility that astrocytic modulation of synaptic NMDAR activity and neurotransmission is mediated by other active substances such as D-serine [46,47], which might be released through Best1 channels in astrocytes. Further studies are required for clarifying the role of astrocytes in modulating neural circuits and behaviors by dissecting the exact functions of Best1 channels in neurotransmission.
In summary, our present study reveals that astrocytes can play a significant role in regulating neural synaptic functions, by releasing glutamate via Ca2+-activated anion channels opened by the activation of receptor-mediated signaling. The models proposed and the tools developed in this study aim to improve our understanding of the physiological role of multiple glutamate release mechanisms, along with their significance in cognitive functions.
All animal protocols were performed in accordance with the institutional guideline of Korea institute of Science and Technology (KIST; Seoul, Korea).
FRET-based glutamate imaging
The amount of released extracellular glutamate was represented by the ratio between the emission intensity of CFP and YFP (CFP/YFP), which was divided by baseline CFP/YFP ratio (relative CFP/YFP ratio). Adenovirus containing pDisplay-GluSnFR  was injected into the hippocampal CA1 region. After one week, FRET imaging in acute hippocampal slices was performed under a microscope (BX50WI; Olympus) equipped with a xenon lamp fitted with a 436/20-excitation filter (D436/20x filter; Chroma). The emission beam was split using a Dual-View (Optical Insights) fitted with a CFP/YFP filter set (OI-05-EX), and recorded with an iXon EMCCD camera (Andor). Imaging Workbench software (INDEC BioSystems) was used for image acquisition and offline image analysis. TFLLR puffs (TFLLR-NH2; Peptron, Korea; 500 μM) were applied using picospritzer-assisted positive pressure (~100 ms).
Production of shRNA containing lentivirus and delivery into mouse hippocampus
Scrambled shRNA, mBest1-shRNA, and Par1-shRNA were inserted into pSicoR lentiviral vector (provided by Dr. T. Jacks through Addgene Inc.;  as previously described . For shRNA expression in hippocampal CA1 region, lentivirus (produced by Macrogen, Korea) was introduced into the hippocampal CA1 region by the stereotaxic surgery method . hGFAP-CreERT2 transgenic mice were provided by Dr. Ken McCarthy. CaMKIIα-Cre transgenic mice were purchased from The Jackson Laboratory. hGFAP-CreERT2 mice were used at the age of 7 weeks for tamoxifen or sunflower oil injection (intraperitoneal injection, once per day for 5 days). The lentivirus carrying shRNA was injected 1 day after the fifth day injection. CaMKIIα-Cre mice were used at 8 weeks of age for virus injection and were used at around 9 weeks for electrophysiological recordings. Only male mice were used throughout the study. Injected mice were sacrificed for electrophysiological recordings at 8 ~ 9 weeks of age.
Slice preparation and electrophysiology
Horizontal or transverse mouse brain slices (300 ~ 400 μm) containing hippocampus were acutely prepared as previously described . Prepared slices were left to recover for at least 1 hour before recording, in oxygenated (95% O2 and 5% CO2) artificial cerebrospinal fluid (ACSF; in mM, 130 NaCl, 24 NaHCO3, 3.5 KCl, 1.25 NaH2PO4, 1 CaCl2, 3 MgCl2 and 10 glucose, pH 7.4; room temperature). The standard ACSF recording solution was composed of (mM): 130 NaCl, 24 NaHCO3, 3.5 KCl, 1.25 NaH2PO4, 1.5 CaCl2, 1.5 MgCl2 and 10 glucose saturated with 95% O2 and 5% CO2, at pH 7.4. To block the effect of neuronal spontaneous activity on astrocytes, TTX (0.5 μM; Tocris Bioscience) was added into the ACSF. Experiments with a holding current of more than −100 pA or in which there was a change in input resistance >30% of the control were rejected. Recordings were obtained using Axopatch 200A (Axon Instruments) and were filtered at 2 kHz. mEPSC recordings were digitized at 10 kHz and analyzed using pCLAMP 9 software (Axon Instruments) and Mini Analysis Program software (Synaptosoft) as previously described . Whole-cell recordings from CA1 neuron, mEPSC recordings, and eEPSP recordings were carried out as previously described . For making Zn-included ACSF solution, 250 nM ZnCl2 was used in 10 mM Tricine with the relation [Zinc]free = [Zinc]applied/200 as previously described [34,35]. CA1 fEPSPs were evoked by Schaffer collateral using a bipolar electrode and quantified as the initial slope of fEPSP as previously described . To examine the effect of TFLLR on synaptic plasticity at different stimulation frequencies, slices were perfused with TFLLR (30 μM) ~15 min before stimulation. Electrical stimulations were given as 1 Hz (900 sec), 10 Hz (90 sec), and theta-burst stimulation (consisting of four trains containing ten bursts (each with four pulses at 100 Hz) of stimuli delivered every 200 msec). For 40 Hz + TFLLR-induced LTP induction experiments, TFLLR (30 μM) was applied ~8 min before 40 Hz stimulation and washed out 2 min after stimulation. APV (Tocris Bioscience; 50 μM), niflumic acid (Sigma; 100 μM) was co-treated with TFLLR.
Adult mice were deeply anesthetized with 2% avertin and perfused with 0.1 M PBS followed by 4% paraformaldehyde. Brains were postfixed in 4% paraformaldehyde at 4°C for 24 hr and 30% sucrose at 4°C for 48 hr. Brains were then cut into 30 μm coronal cryosections. Sections were blocked in 0.1 M PBS containing 0.3% Triton X-100 (Sigma) and 2% Donkey Serum (GeneTex) for 30 min at room temperature. Primary antibody was then applied at the appropriate dilution and incubated overnight at 4°C. Sections were then washed three times in 0.1 M PBS and incubated in secondary antibody for 2 h. After three rinses in 0.1 M PBS and DAPI staining at 1:1000 (Pierce), the sections were mounted on polysine microscopic glass slides (Thermo Scientific). Images were acquired using a Nikon A1R confocal microscope.
Animals were deeply anesthetized with sodium pentobarbital (80 mg/kg, i.p.) and perfused transcardially with heparinized normal saline (10 ml for mouse and 100 ml for rat), followed by a freshly prepared mixture (50 ml for mouse and 500 ml for rat) of 4% paraformaldehyde and 0.01% glutaraldehyde in 0.1 M phosphate buffer (PB), pH 7.4. Hippocampus was removed and postfixed in the same fixative for 2 hours at 4°C. Sagittal sections (60 μm) were cut with a vibratome and cryoprotected in 30% sucrose in PB overnight at 4°C. Sections were frozen on dry ice for 20 minutes, thawed in phosphate-buffered saline (PBS; 0.01 M, pH 7.2) to enhance penetration. They were pretreated with 1% sodium borohydride for 30 minutes to quench glutaraldehyde and then blocked with 3% H2O2 for 10 minutes to suppress endogenous peroxidases, and with 10% Normal Donkey Serum (NDS, Jackson ImmunoResearch, West Grove, PA) for 30 minutes to mask secondary antibody binding sites. For double immunostaining of GFP and Best1, sections of hippocampus pretreated as above were incubated overnight in a mixture of mouse anti-GFP (1:400, MAB3580, Millipore, Temecula, CA) and rabbit anti-Best1 (1:200) antibodies. After rinsing in PBS, sections were incubated with a mixture of biotinylated donkey anti-mouse (1:200, Jackson ImmunoResearch) and 1 nm gold-conjugated donkey anti-rabbit (1:50, EMS, Hatfield, PA) antibodies for 2–3 hours. The sections were postfixed with 1% glutaraldehyde in PB for 10 minutes, rinsed in PB several times, incubated for 4 minutes with HQ silver enhancement solution (Nanoprobes, Yaphank, NY), and rinsed in 0.1 M sodium acetate and PB. Serially cut thin sections were collected on Formvar-coated single slot nickel grids and stained with uranyl acetate and lead citrate. Grids were examined on a Hitachi H-7500 electron microscope (Hitachi, Tokyo, Japan) at 80 kV accelerating voltage. Images were captured with Digital Montage software driving a MultiScan cooled CCD camera (ES1000W, Gatan, Pleasanton, CA) attached to the microscope and saved as TIFF files.
All data were shown as mean ± standard error of the mean (s.e.m.). Statistical analyses were performed by using Sigma Plot software (ver. 10.0; Systat Software Inc., San Jose, CA). Detailed information of the statistical tests was indicated in the figure legends.
We thank Dr. T. Jacks for providing us with pSicoR lentiviral vector through Addgene Inc., Dr. R. Tsien for the pDisplay-GluSnFR vector, and Dr. K. McCarthy for hGFAP-CreERT2 transgenic mice. HP was supported by the Korea Institute of Science and Technology (KIST; Star-Postdoc. Fellowship). GM and RPL were supported by the Ente Cassa di Risparmio di Firenze. JS, S-HC, and S-Y C were supported by the Korea Research Foundation (KRF; grant 2014050477). SFT was supported by NIH (grant NS039419). CJL was supported by NIH (grant NS43875), KRF (grant KRF-2005-070-C00096), KIST institutional program (Project No. 2E25210), and Brain Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Science, ICT & Future Planning (NRF-2012M3C7A1055412).
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