AMPA Receptor-Mediated Ca2+ Transients in Mouse Olfactory Ensheathing Cells

Introduction

Ca²⁺ signaling in glial cells is involved in various intercellular processes such as the release of gliotransmitters, modulation of synaptic transmission, long-range Ca²⁺ wave propagation, and neurovascular coupling (Haydon, 2001; Carmignoto and Gomez-Gonzalo, 2010). Moreover, intracellular processes such as apoptosis, transcription as well as posttranslational modification are regulated by Ca²⁺ signaling (McConkey and Orrenius, 1997; Flavell and Greenberg, 2008). In glial cells, rises of intracellular Ca²⁺ are predominantly triggered by Ca²⁺ release from internal Ca²⁺ stores induced by metabotropic pathways (Deitmer et al., 1998; Verkhratsky et al., 1998), while Ca²⁺ influx was reported in rare cases in specialized astrocyte-like glial cells such as cerebellar Bergmann glia and retinal Müller glia (Burnashev et al., 1992; Muller et al., 1992; Wakakura and Yamamoto, 1994). However, more recent studies suggest that astroglial cells of different regions of the brain also express ligand-gated ion channels that may trigger Ca²⁺ responses via Ca²⁺ influx from the extracellular space (Schipke et al., 2001; Lalo et al., 2006; Mishra et al., 2016; Droste et al., 2017). In addition, store-operated channels as well as transient receptor potential (TRP) channels contribute to Ca²⁺ signaling in astrocytes (Singaravelu et al., 2006; Rungta et al., 2016; Belkacemi et al., 2017; Rakers et al., 2017; Toth et al., 2019). These studies indicate that Ca²⁺ influx might play a previously underestimated role in glial cell physiology and function. Olfactory ensheathing cells (OECs) represent a specialized population of glial cells, exclusively located in the olfactory nerve layer (ONL) in the olfactory bulb (OB) and the peripheral olfactory mucosa (Su and He, 2010; Lohr et al., 2014). They support growth and guidance of axons of olfactory sensory neurons (OSN) from the olfactory epithelium (OE) into the main OB (Graziadei and Graziadei, 1979; Doucette, 1984; Yang et al., 2015). It is assumed, that OECs generate an environment allowing the lifelong re-growth and re-integration of axons into the cellular network after degeneration by the expression of different neurite growth factors and cell adhesions molecules (Crandall et al., 2000; Woodhall et al., 2001; Cao et al., 2007; Yang et al., 2015). In addition, studies showed that Ca²⁺ signaling in OECs is a critical regulator for neurite outgrowth in OEC/retinal ganglion cell (RGC) co-cultures, suggesting a prominent role for Ca²⁺ signaling in OEC-axon interaction (Hayat et al., 2003). Extrasynaptic release of glutamate and ATP by OSN axons initiates Gq-mediated Ca²⁺ release from internal stores in OECs via mGluR1 and P2Y1 receptors (Rieger et al., 2007; Thyssen et al., 2010). Montague and Greer (1999) investigated the distribution of the ionotropic AMPA receptor subunits, GluA1, GluA2/3 and GluA4 in the ONL and found expression by nerve-associated glial cells. However, whether OECs exhibit functional AMPA receptors, which might conduct membrane currents and mediate Ca²⁺ influx is not known. We performed whole-cell voltage-clamp recordings, showing that kainate application induced AMPA receptor-mediated inward currents in OECs, which are partly mediated by Ca²⁺-permeable AMPA receptors. Kainate also induced Ca²⁺ transients that depended on Ca²⁺ influx via the receptor channel itself, but additionally comprised Ca²⁺ release from internal stores. Electrical stimulation of OSN axons evoked Ca²⁺ transients mediated by metabotropic receptors as well as Ca²⁺-permeable AMPA receptors, suggesting a role for AMPA receptor-mediated Ca²⁺ signaling in axon-OEC communication.

Materials and Methods

Animals and Olfactory Bulb Preparation

Mice of the GLAST-Cre^ERT2 × tdTomato^fl/fl, PLP-Cre^ERT2 × tdTomato^fl/fl (age: p28–60) and NMRI (age: p18–p21; Naval Medical Research Institute) strains (Leone et al., 2003; Mori et al., 2006; Madisen et al., 2010) were kept at the institutional animal facility of the University of Hamburg. These mice expressed Cre recombinase under control of the promoters of the glutamate/aspartate transporter EAAT1 (GLAST; expressed by OECs and astrocytes) and the proteolipid protein (PLP; expressed by OECs and oligodendrocytes), respectively (Droste et al., 2017; Beiersdorfer et al., 2019). Animal rearing and all experimental procedures were performed according to the European Union’s and local animal welfare guidelines (GZ G21305/591-00.33; Behörde für Gesundheit und Verbraucherschutz, Hamburg, Germany). To induce reporter expression in GLAST-Cre^ERT2 × tdTomato^fl/fl and PLP-Cre^ERT2 × tdTomato^fl/fl mice, tamoxifen (Carbolution Chemicals GmbH, St. Ingbert, Germany) was dissolved in 8% ethanol/92% Mygliol^®812 (Sigma Aldrich, Taufkirchen, Germany) and injected intraperitoneally for three consecutive days (starting p21; 100 mg/kg bodyweight). Animals were analyzed 7–12 days after the first injection. OBs were prepared as described previously (Stavermann et al., 2012). Both OBs were removed from the opened head in cool preparation solution (molarities in mM: 83 NaCl, 1 NaH₂PO₄x2H₂O, 26.2 NaHCO₃, 2.5 KCl, 70 sucrose, 20 D-(+)-glucose, and 2.5 MgSO₄ × 7 H₂O). Standard artificial cerebrospinal fluid (ACSF) for experiments and storage of the preparations consisted of (molarities in mM): 120 NaCl, 2.5 KCl, 1 NaH₂PO₄x2H₂O, 26 NaHCO₃, 2.8 D-(+)-glucose, 1 MgCl₂, and 2 CaCl₂). Preparation solution and ACSF were continuously perfused with carbogen (95% O₂ and 5% CO₂) to maintain the pH of 7.4 and to supply oxygen.

Reagents

The compounds (2S,3S,4S)-3-(Carboxymethyl)-4-(prop- 1-en-2-yl)pyrrolidine-2-carboxylic acid (kainic acid, agonist of AMPA/kainate receptors), (S)-α-Amino-3-hydroxy-5- methylisoxazole-4-propionic acid, 4-(8-Methyl-9H-1,3-dioxolo [4,5-h][2,3]benzodiazepin-5-yl)-benzenamine hydrochloride (GYKI53655, antagonist of AMPA receptors), (E)-ethyl 1,1a,7,7a-tetrahydro-7-(hydroxyimino) cyclopropa[b]chromene-1a-carboxylate (CPCCOEt, antagonist of mGluR₁), 2-methyl- 6-(phenylethynyl) pyridine (MPEP, antagonist of mGluR₅), 2′-deoxy-N6-methyladenosine 3′,5′-bisphosphate (MRS2179, antagonist of P2Y₁), 4-(2-[7-Amino-2-(2-furyl)[1,2,4]triazolo [2,3-α][1,3,5]triazin-5-ylamino]ethyl) pheno (ZM241385, antagonist of A_2A receptors), (R)-(+)-7-chloro-8-hydroxy-3-methyl-1-phenyl-2,3,4,5-tetrahydro-1H-3-benzazepine (SCH2 3390, antagonist of D1-like dopamine receptors) and carbenoxolone disodium salt (CBX; inhibiting gap junctions) were obtained from Abcam (Cambridge, United Kingdom). The reagents D-2-amino-5-phosphonovaleric acid (D-APV; antagonist of NMDA receptors), N-[3-[[4-[(3-aminopropyl)amino]butyl] amino]propyl]-1-naphthaleneacetamide trihydrochloride (Naspm trihydrochloride antagonist of Ca²⁺-permeable AMPA receptors) and tetrodotoxin (TTX; inhibiting voltage-gated sodium channels) were received from Alomone labs (Jerusalem, Israel). All reagents were stored as stock solutions corresponding to the manufacturer’s instructions and added to ACSF directly before the experiment. Drugs were applied via the perfusion system driven by a peristaltic pump (Reglo MS4/12, Ismatec, Wertheim, Germany).

Electrophysiology and Analysis

For electrophysiological experiments, OB slices (220 μm, horizontal) of GLAST-Cre^ERT2 × tdTomato^fl/fl mice were prepared using a vibratome (Leica VT1200S, Nußloch, Germany). Slices were transferred into a recording chamber and continuously superfused with ACSF via the perfusion system. The experiments were performed at room temperature. Whole-cell voltage-clamp recordings (Multiclamp 700B, Molecular Devices) were performed on OECs, identified by tdTomato expression (excited at 543 nm, emission 553–618 nm), and the distinct localization in the ONL (Au et al., 2002). Recordings were digitized (Digidata 1440A, Molecular Devices) at 10–20 kHz and filtered (Bessel filter 2 kHz). The holding potential was −80 mV, achieved by current injection of −35.14 ± 13.5 pA on average. Patch pipettes had a resistance of 4–7 MΩ when filled with internal solution containing (mM): 105 K-gluconate, 30 KCl, 10 HEPES, 10 phosphocreatine, 4 Mg-ATP, 0.3 Na-GTP, 0.3 EGTA, and pH = 7.43. For visualization of the recorded OEC, ATTO-488 carboxy (20 μM, ATTO-Tech GmbH, Siegen, Germany) was added to the internal solution before the experiments. Agonists were applied via the perfusion system for 30 s. Antagonists were applied via the perfusion system 10 min prior to agonist application. The flow rate of the perfusion system amounted to approximately 2 mL/min. The experimental bath was oval in shape and had a diameter of 2.1 cm, the bath volume amounted to 1 mL. Therefore, agonist and antagonist concentrations continuously increased in the experimental bath, until the final concentration is reached. Vice versa, agonists were slowly washed out, accounting for long lasting effects. Kainate-induced inward currents in OECs were analyzed by measuring amplitude the currents.

Ca²⁺ Imaging and Analysis

For Ca²⁺ imaging experiments, GLAST-Cre^ERT2 × tdTomato^fl/fl and NMRI mice were used. Whole OBs were glued onto coverslips, transferred into a recording chamber and the coverslip secured with a U-shaped platinum wire. For multi-cell bolus loading (Stosiek et al., 2003), a glass pipette with a resistance of ∼3.5 MΩ (when filled with ACSF) was filled with 200 μM Fluo-8 AM (Thermo Fisher Scientific, Darmstadt, Germany) in ACSF, made from a 4 mM stock solution (dissolved in DMSO and 20% pluronic acid). After inserting the injection pipette into the ONL, the Ca²⁺ indicator was pressure-injected with 0.7 bar for 30 s into the tissue (PDES-01 AM, NPI electronic GmbH, Tamm, Germany), followed by an incubation of 20 min. Changes in the cytosolic Ca²⁺ concentration in OECs were detected by the fluorescence of Fluo-8 (excitation: 488 nm; emission: 500–530 nm) using a confocal microscope (eC1, Nikon, Düsseldorf, Germany). Images were acquired at a time rate of one frame every 3 s. A similar perfusion system and flow rate was used as described above. However, the experimental bath for Ca²⁺ imaging experiments was differently shaped (diameter 1 cm, volume 1.9 mL) resulting in differences in the kinetics of wash-in and wash-out of drugs between the electrophysiological and Ca²⁺ imaging experiments. To analyze changes in cytosolic Ca²⁺ in single cell somata, regions of interest (ROIs) were defined using Nikon EZ-C1 3.90 software. OECs were identified by GLAST promoter-driven tdTomato expression in GLAST-Cre^ERT2 × tdTomato^fl/fl mice in most experiments. In wild type mice, OECs were identified by their distinct localization in the ONL. The changes in Ca²⁺ were recorded throughout the experiments as relative changes in Fluo-8 fluorescence (ΔF) with respect to the resting fluorescence, which was normalized to 100%. Quantification of the Ca²⁺ transients was achieved by calculating the amplitude of ΔF.

Statistics

All values are stated as mean values ± standard error of the mean. The number of experiments is given as n = x, where x is the number of analyzed cells. At least 3 animals were analyzed for each set of experiments. Statistical significance was estimated by comparing three means using Friedmann ANOVA and the Wilcoxon post hoc test for paired data, and for comparing two means using the Mann–Whitney U-Test, with the error probability p (^∗ p ≤ 0.05; ^∗∗ p ≤ 0.01; ^∗∗∗ p ≤ 0.001).

Immunohistochemistry

Immunohistochemistry on OBs of PLP-Cre^ERT2 × tdTomato^fl/fl mice (≥P28) was performed as described before (Droste et al., 2017). After dissection, the OBs were kept for 1 h at room temperature (RT) in formaldehyde (4% in PBS, pH 7.4). Afterward, 100 μm thick sagittal slices were prepared with a vibratome (VT1000S, Leica, Nußloch, Germany) and incubated for 1 h in blocking solution (10% normal goat serum (NGS), 0.5% Triton X-100 in PBS) at RT. Subsequently, the slices were incubated for 48 h at 4°C with the following primary antibodies: Guinea pig anti-GluA1 (Alomone labs; 1:200); rabbit anti-GluA2 (Millipore, 1:200); rabbit anti-GluA4 (Millipore; 1:200). To validate the specificity of the GluA antibodies we used cerebellar slices as control, since the distribution of GluA subunits is well documented in this brain area (Supplementary Figure S1). Moreover, the antibodies against GluA1 and GluA4 have been validated in glia-specific GluA1 and GluA4 double knockout mice before (Saab et al., 2012). In our control experiments, the GluA2 antibody only labeled cells known to express GluA2, but not cells that lack GluA2 such as Bergmann glial cells (Burnashev et al., 1992; Muller et al., 1992; Saab et al., 2012), as shown before for the used antibody (Droste et al., 2017). Hence, we consider the used antibodies as efficient and specific. The antibodies were diluted in 1% NGS, 0.05% TritonX-100 in PBS. Slices were incubated in PBS with the following secondary antibodies for 24 h at 4°C: goat anti-rabbit Alexa Fluor 488 (Thermo Fisher Scientific; 1:1000) or goat anti-guinea pig Alexa Fluor 488 (Thermo Fisher Scientific; 1:1000). Moreover, Hoechst 33342 (5 μM; Molecular Probes, Eugene, OR, United States) was added to stain nuclei. Slices were mounted on slides using self-hardening embedding medium (Immu-Mount, Thermo Fisher Scientific). Immunohistological stainings were analyzed using a confocal microscope (Nikon eC1). Confocal images were adjusted for brightness and contrast using ImageJ and Adobe Photoshop CS6.

Results

Distribution of GluA Subunits in the ONL

AMPA receptors constitute of four subunits (GluA1-GluA4), which form complex heteromeric cation channels, activated by glutamate or selective receptor agonists such as AMPA and kainate (Steinhauser and Gallo, 1996). The composition of the subunits is especially critical to determine the Ca²⁺ permeability of the channel. Thus, GluA2-lacking AMPA receptors show a high permeability for Ca²⁺ whereas GluA2-containing AMPA receptors are impermeable for Ca²⁺ (Hollmann et al., 1991). The OB is divided into several clearly distinguishable layers (Figures 1A,B) and immunoreactivity against GluA1, GluA2/3, and GluA4 demonstrated a wide distribution of these subunits in all layers, while mRNA expression analyses showed low amounts of GluA3 in the OB (Montague and Greer, 1999; Horning et al., 2004). We aimed to analyze the distribution of GluA1, GluA2, and GluA4 specifically in the ONL by immunohistochemistry using PLP-Cre^ERT2 × tdTomato^fl/fl mice. The plp gene encodes for the PLP, expressed by oligodendrocytes, and its slice variant DM20, expressed by oligodendrocytes and OECs, enabling the specific identification of oligodendrocytes and OECs by Cre-recombinase-driven tdTomato expression (Griffiths et al., 1995; Dickinson et al., 1997). OECs are present exclusively in the ONL, whereas oligodendrocytes are located in the glomerular layer (GL) and deeper layers but not in the ONL (Figures 1C,D; Doucette, 1990, 1991). Therefore, PLP-Cre^ERT2-driven tdTomato expression in the ONL is assumed to be restricted to OECs (Figures 1C,D; Beiersdorfer et al., 2019; Piantanida et al., 2019). GluA1 immunoreactivity was widely distributed in the entire ONL, colocalizing intensely with tdTomato-expressing OECs (Figures 1E,F). Homogenous GluA2 immunoreactivity was detected in the ONL, with scattered colocalization with tdTomato-positive OECs (Figures 1G,H). In the GL, GluA2-positive juxtaglomerular cells were present (Figure 1G, arrow; Droste et al., 2017). Additionally, GluA4 immunoreactivity was found in the ONL, however, it sparsely colocalized with tdTomato-positive OEC somata (Figures 1I,J). At higher magnification, GluA4 immunoreactivity was found in close approximation of tdTomato-expressing OECs, likely indicating GluA4 located in cell processes of OECs (Figure 1J, see arrowheads). The results suggest that OECs express GluA1 and GluA2, while GluA4 appears not to be present in OEC somata but presumably in cell processes.

FIGURE 1

Figure 1. Distribution of AMPA receptor subunits in the olfactory nerve layer (ONL). (A) Laminar organization of the olfactory bulb. Nuclei were stained with Hoechst (5 μM). (B) The ONL, glomerular layer (GL), external plexiform layer (EPL) and granule cell layer (GCL) are clearly distinguishable. Scale bar: 200 μm. (C) Distribution of PLP-Cre-dependent tdTomato expression in the ONL and GL. Olfactory ensheathing cells (OECs) are located in the ONL, whereas tdTomato-expressing oligodendrocytes are located in the GL. Nuclei are stained with Dapi (5 μM; blue). The assumed border between the ONL and GL is indicated by the dotted line. Glomeruli are marked by asterisks. Scale bar: 150 μm. (D) Magnified view from (C). OECs are located in the ONL (arrowhead) and oligodendrocytes (arrow) in the GL visualized by PLP-Cre-dependent tdTomato expression (red). Scale bar: 50 μm. (E) Distribution of GluA1 immunoreactivity (green) in the ONL of PLP-Cre^ERT2 × tdTomato^fl/fl mice (red). Nuclei are stained with Dapi (blue). Glomeruli are marked by asterisks. Scale bar: 50 μm. (F) Magnified view from (E). GluA1 immunostaining (green) colocalize with tdTomato-expressing OECs (red) in the ONL (arrowheads). Scale bar: 25 μm. (G) Distribution of GluA2 immunoreactivity (green) in the ONL of PLP-Cre^ERT2 × tdTomato^fl/fl mice (red). GluA2 is widely distributed in the ONL, as well as in the GL. Nuclei are stained with Dapi (blue). Glomeruli are marked by asterisks. Scale bar: 50 μm. (H) Magnified view from (G). GluA2 immunoreactivity colocalizes with tdTomato-expressing OECs (red) in the ONL (arrowheads). Scale bar: 25 μm. (I) Distribution of GluA4 immunoreactivity (green) in the ONL of PLP-Cre^ERT2 × tdTomato^fl/fl mice (red). Nuclei are stained with Dapi (blue). Glomeruli are marked by asterisks. Scale bar: 50 μm. (J) Magnified view from (I). GluA4 immunoreactivity (green) is sparsely colocalized with tdTomato-positive OEC somata (red), but is localized adjacent to OEC somata (arrowheads). Scale bar: 25 μm.

Kainate Induces AMPA Receptor-Mediated Inward Currents in OECs

Functional AMPA/kainate receptors have been investigated in macroglial cells, namely astrocytes and oligodendrocyte precursor cells in gray and white matter as well as microglial cells (Burnashev et al., 1992; Muller et al., 1992; Jabs et al., 1994; Seifert and Steinhauser, 1995; Noda et al., 2000; Zonouzi et al., 2011; Hoft et al., 2014; Droste et al., 2017). Based on our immunohistochemical data, we suggested that OECs in the ONL might also express functional AMPA receptors. Therefore, we performed electrophysiological recordings in OECs in acute OB slices of GLAST-Cre^ERT2 × tdTomato^fl/fl mice. OECs were identified by GLAST promoter-driven tdTomato expression in the ONL (Figure 2A). Additionally, a voltage-step protocol was applied to characterize the I/V relationship of the recorded cell. All OECs analyzed showed a characteristic linear I/V relationship (Figures 2B,C; Rela et al., 2010; Beiersdorfer et al., 2019). We used kainate to agonize AMPA/kainate receptors, since kainate induced a non-desensitizing AMPA receptor-mediated inward current in acutely isolated hippocampal glial cells (Seifert and Steinhauser, 1995). Kainate application (100 μM, 30 s) induced a prominent slowly rising and long lasting inward current with a mean amplitude of 231.5 ± 58.9 pA (at −80 mV holding potential) in OECs (Figure 2D). To specifically isolate kainate-induced responses in OECs, all experiments were performed in the presence of the gap junction inhibitor carbenoxolone (CBX, 100 μM) to avoid intercellular panglial communication to juxtaglomerular astrocytes that respond to kainate and might interfere with recordings in OECs (Droste et al., 2017; Beiersdorfer et al., 2019). Additionally, neuronal activity was suppressed by inhibiting voltage-gated sodium channels by tetrodotoxin (TTX, 1 μM). Application of GYKI53655 (100 μM), a selective AMPA receptor antagonist, significantly reduced kainate-induced inward currents in OECs by 88.8 ± 3.2% of the control (n = 5; p = 0.022), suggesting that the kainate-induced response is mainly mediated by AMPA receptors in OECs (Figures 2D,E). To further characterize the AMPA receptor subtype involved in kainate-induced inward currents, GluA2-lacking Ca²⁺-permeable AMPA receptors were inhibited by the selective antagonist, NASPM (100 μM) (Koike et al., 1997). NASPM reduced kainate-mediated inward currents in OECs significantly by 63.9 ± 2.7% of the control (n = 6; p = 0.019), suggesting that OECs exhibit functional Ca²⁺-permeable AMPA receptors (Figures 2F,G).

FIGURE 2

Figure 2. Kainate induces AMPA receptor-mediated inward currents in olfactory ensheathing cells (OECs). (A) Distribution of OECs in the olfactory nerve layer (ONL), identified by GLAST-Cre^ERT2-driven tdTomato expression. Scale bar: 50 μm. (B) Voltage-step protocol applied to an OEC, recorded in whole cell voltage clamp mode. (C) All recorded OECs showed a linear current-voltage relationship. (D) The application of kainate (100 μM) induced an inward current in OECs, that was reduced by the selective AMPA receptor inhibitor GYKI 53655 (100 μM). All experiments were performed in the presence of TTX (1 μM) and CBX (150 μM). (E) Averaged amplitudes and statistical analysis of kainate-induced inward currents in OECs. (F) NASPM (100 μM), a selective inhibitor of Ca²⁺-permeable AMPA receptors, reduced kainate-induced inward currents significantly. (G) Averaged amplitudes and statistical analysis of kainate-induced inward currents in OECs. ^∗p ≤ 0.05.

Kainate-Mediated Ca²⁺ Transients in OECs Depend on Extracellular Ca²⁺

Kainate-induced Ca²⁺ transients in Bergmann glial cells of the cerebellum result from Ca²⁺ influx directly through the AMPA receptor channel (Burnashev et al., 1992; Muller et al., 1992). However, AMPA/kainate receptors may also activate metabotropic pathways (Wang et al., 1997; Rozas et al., 2003; Takago et al., 2005). We performed confocal Ca²⁺ imaging experiments to investigate the source of Ca²⁺ that fuels Ca²⁺ transients in OECs in response to kainate application. The Ca²⁺ indicator Fluo-8 AM was pressure injected into OB in-toto preparations of GLAST-Cre^ERT2 × tdTomato^fl/fl mice (Figures 3A,B). Experiments were performed in the presence of CBX and TTX to suppress indirect effects. Kainate application induced Ca²⁺ transients in OECs with a mean amplitude of 107.8 ± 5.3% ΔF. GYKI 53655 (100 μM), an AMPA receptor antagonist, greatly reduced kainate-evoked Ca²⁺ transients in OECs by 84.0 ± 6.1% of the control (n = 27; p = 5.93 × 10^–6), showing that kainate-evoked Ca²⁺ transients are mediated by AMPA receptors (Figures 3C,D). Removal of extracellular Ca²⁺ reduced kainate-induced Ca²⁺ transients by 91.9 ± 2.3% (n = 43; p = 1.16 × 10^–8). Kainate-evoked Ca²⁺ responses recovered after readdition of extracellular Ca²⁺, the average amplitude reached 122.0 ± 18.3% of the control (p = 0.16) (Figures 3E,F). To estimate the impact of intracellular Ca²⁺ stores on kainate-induced Ca²⁺ transients in OECs, we applied cyclopiazonic acid (CPA, 20 μM) to inhibit Ca²⁺ uptake into Ca²⁺ stores via SERCA (sarcoplasmic/endoplasmic reticulum calcium ATPase). Kainate-induced Ca²⁺ transients were significantly reduced by 57.6 ± 4.2% upon Ca²⁺ store depletion (n = 70; p = 8.18 × 10^–8) (Figures 3G,H). Our results suggest that Ca²⁺ influx via AMPA receptor channels is essential to trigger Ca²⁺ transients in OECs, whereas Ca²⁺ release from internal stores is a secondary effect as a result of the kainate-evoked Ca²⁺ influx. To verify that the AMPA-evoked membrane current and associated Ca²⁺ influx was not elicited by a preceding Ca²⁺ increase, we evoked a Ca²⁺ transient by application of ADP and recorded membrane currents. ADP induced a large Ca²⁺ increase but failed to elicit membrane currents, demonstrating that Ca²⁺-dependent membrane conductances are negligible in OECs (Supplementary Figure S2).

FIGURE 3

Figure 3. Kainate-induced Ca²⁺ transients in olfactory ensheathing cells (OECs) depend on extracellular Ca²⁺ and intracellular Ca²⁺ stores. (A) Schematic illustration of the experimental set up of Ca²⁺ imaging experiments. Olfactory bulb (OB) in-toto preparations of GLAST-Cre^ERT2 × tdTomato^fl/fl mice were fixed and placed under the confocal microscope. Fluo-8 was pressure injected into the tissue. The focal plane as seen in panel (B) is indicated by the dotted line. (B) Confocal image of tdTomato-expressing OECs in an OB in-toto preparation. Arrowheads highlight OEC somata located in the ONL. One glomerulus is indicated by an asterisk. Scale bar: 100 μm. (C) Kainate-induced Ca²⁺ transients in OECs in the presence of TTX (1 μM) and CBX (150 μM) were reduced by GYKI 53655 (100 μM). (D) GYKI 53655 significantly reduced kainate-mediated Ca²⁺ transients. (E) Withdrawal of extracellular Ca²⁺ abolished kainate-induced Ca²⁺ transients in OECs completely. Restitution of extracellular Ca²⁺ caused a recovery of kainate-induced Ca²⁺ transients in OECs. (F) Averaged amplitudes and statistical analysis of kainate-mediated Ca²⁺ transients. (G) Upon intracellular Ca²⁺ store depletion by CPA (20 μM) kainate-induced Ca²⁺ transients were significantly reduced. (H) Averaged amplitudes and statistical analysis of the effect of CPA on kainate-mediated Ca²⁺ transients. ^∗∗∗p ≤ 0.001; ^∗p ≤ 0.05. n.s.: not significant.

Endogenous Glutamate Release Induces AMPA Receptor-Mediated Ca²⁺ Transients in OECs

Electrical stimulation of OSN axons evokes ectopic release of glutamate and ATP, which initiates mGluR1 and P2Y1 receptor-mediated Ca²⁺ rises in OECs (Berkowicz et al., 1994; Rieger et al., 2007; Thyssen et al., 2010). Here, we showed that OECs express functional Ca²⁺-permeable AMPA receptors which may also contribute to Ca²⁺ transients in response to neuronal activity. We endogenously released glutamate upon electrical stimulation of OSN axons (20 Hz, 2 s) and recorded Ca²⁺ transients in Fluo-8-loaded OECs in wild type animals. Electrical stimulation of OSN axons induced Ca²⁺ transients in OECs with an amplitude of 51.8 ± 5.0% ΔF (n = 46). By eliminating purinergic (MRS2179), dopaminergic (SCH23390) and mGluR1-mediated Ca²⁺ transients (CPCCOEt) as well as possible indirect NMDA-mediated neuronal responses (D-APV), the mean amplitude of stimulation-induced Ca²⁺ transients in OECs were significantly reduced to 61.0 ± 9.3% of the control (p = 3.68 × 10^–4). Additional inhibition of Ca²⁺-permeable AMPA receptors by the selective antagonist NASPM further reduced the mean amplitude to 30.6 ± 5.8 (p = 1.36 × 10^–5) of the control. NBQX almost completely abolished stimulation-induced Ca²⁺ transients in OECs and the amplitude further decreased significantly to 13.01 ± 3.3% of the control (p = 0.002). Receptor inhibition was reversible and stimulation-induced Ca²⁺ transients in OECs recovered after wash out (Figure 4). The results indicate that endogenous glutamate release triggers AMPA receptor-mediated Ca²⁺ responses in OECs, suggesting a possible role for AMPA receptors in axon-OEC communication.

FIGURE 4

Figure 4. Electrical stimulation of OSN axons evokes AMPA-mediated Ca²⁺ transients in olfactory ensheathing cells (OECs).(A) Electrical stimulation of OSN axons evoked Ca²⁺ transients in OECs in control conditions (ACSF), and in the presence of purinergic (MRS2179, 100 μM; ZM241385; 0.5 μM), dopaminergic (SCH23390; 5 μM) an glutamatergic (CPCCOEt, 100 μM; D-APV, 100 μM) receptor antagonists. The additional application of NASPM (100 μM) further reduced Ca²⁺ transients in OECs. NBQX (20 μM) abolished stimulation-induced Ca²⁺ transients. (B) Averaged amplitudes of stimulation-evoked Ca²⁺ transients in OECs. ^∗∗∗p < 0.0001; ^∗∗p < 0.01; ^∗p < 0.05.

Discussion

In the present study, we investigated the role of AMPA receptors in OEC physiology and axon-OEC communication. Kainate induced inward currents and Ca²⁺ transients in OECs that were partially mediated by the GluA2-lacking Ca²⁺-permeable AMPA receptor subtype. Endogenous glutamate release initiated AMPA receptor-mediated Ca²⁺ transients in OECs, indicating the relevance of AMPA receptors for axon-OEC communication.

We showed clear GluA1 and GluA2 immunoreactivity in the ONL colocalized with tdTomato-expressing OECs, while GluA4 immunoreactivity rarely colocalized with tdTomato in OECs. However, in the present study tdTomato was predominantly localized in OEC somata, limiting the detection of OEC processes, in which GluA4 might be enriched, as it has been demonstrated in a study by Montague and Greer (1999), showing strong GluA4 immunoreactivity in cell processes of presumed olfactory nerve-associated glial cells. Therefore, the results of our and other studies suggest the presence of at least three of four GluA subunits in OECs.

While the presence of GluA2 in OECs argues against Ca²⁺ permeability of the AMPA receptors, our physiological data implies the involvement of Ca²⁺ influx via Ca²⁺-permeable AMPA receptors. AMPA receptor-dependent Ca²⁺ influx could either be conducted by Ca²⁺-permeable AMPA receptors or by voltage-gated Ca²⁺ channels activated by AMPA receptor-mediated depolarization (Porter and McCarthy, 1995). However, OECs do not generate voltage-dependent Ca²⁺ influx, e.g., via voltage-gated Ca²⁺ channels, ruling out that a depolarization by AMPA receptors is causative for the kainate-evoked Ca²⁺ signals (Thyssen et al., 2010). In addition, both kainate-evoked membrane currents and Ca²⁺ transients were largely reduced by NASPM, a selective blocker of Ca²⁺-permeable AMPA receptors (Koike et al., 1997). Hence, taking into account that on the one hand side OECs show immunoreactivity to the GluA2 subunit and on the other hand side the current and Ca²⁺ responses are NASPM-sensitive, the results suggest that OECs express both Ca²⁺-permeable as well as Ca²⁺-impermeable AMPA receptors. This is confirmed by the NASPM-insensitive but GYKI-sensitive fraction of kainate-evoked inward currents. In addition to Ca²⁺ influx via Ca²⁺-permeable AMPA receptors, Ca²⁺ release from internal stores contributes to AMPA receptor-mediated Ca²⁺ transients. OECs generate prominent Ca²⁺-induced Ca²⁺ release (CICR) and even a small Ca²⁺ increase generates a fast and large Ca²⁺ transient by CICR (Stavermann et al., 2012). Thus, Ca²⁺ influx through AMPA receptors upon kainate application could induce CICR that boosts the Ca²⁺ increase and accounts for the fast rising kinetics of the Ca²⁺ signal as compared to the rising phase of the kainate-evoked membrane current. We conclude that Ca²⁺ influx is the initial Ca²⁺ signal that triggers subsequent internal Ca²⁺ release such as CICR, as shown by the entire suppression of the Ca²⁺ signal by external Ca²⁺ withdrawal but only partial reduction upon Ca²⁺ store depletion.

Ca²⁺ signaling in OECs may serve different functions. In other glial cells such as astrocytes, increases in the cytosolic Ca²⁺ concentration has been reported to evoke release of so-called gliotransmitters that affect nearby neurons and synapses, but also release of vasoactive substances such as prostaglandins and arachidonic acid (Bazargani and Attwell, 2016; Guerra-Gomes et al., 2017). OECs enwrap blood vessels in the ONL (Herrera et al., 2005) and respond to neurotransmitters extrasynaptically released from axons of OSN with Ca²⁺ signals (Rieger et al., 2007; Thyssen et al., 2010; this study). Increases in intracellular Ca²⁺ in OECs results in constriction of adjacent capillaries (Thyssen et al., 2010). Hence, Ca²⁺ signals evoked by glutamate release from axons in the ONL and subsequent activation of Ca²⁺-permeable AMPA receptors may lead to vasoresponses that adjust blood flow to the metabolic demand upon action potential firing, a mechanism termed neurovascular coupling.

In summary, our results show that release of glutamate from axons of OSN results in Ca²⁺ transients in OECs that are partially mediated by AMPA receptors. AMPA receptor-mediated Ca²⁺ transients were due to Ca²⁺ influx, most likely through Ca²⁺-permeable AMPA receptors themselves, and subsequent Ca²⁺ release from internal Ca²⁺ stores.

Data Availability Statement

The datasets generated for this study are available on request to the corresponding authors.

Ethics Statement

The animal study was reviewed and approved by GZ G21305/591-00.33; Behörde für Gesundheit und Verbraucherschutz, Hamburg, Germany.

Author Contributions

AB and CL designed the experiments and wrote the manuscript. AB performed the experiments and analyzed the data.

Funding

This work was supported by the Deutsche Forschungsgemeinschaft (LO779/10 and SFB 1328 TP-A07).

Conflict of Interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Acknowledgments

We thank A. C. Rakete and M. Fink for technical assistance. We also thank A. Scheller and F. Kirchhoff for providing transgenic animals.

Supplementary Material

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fncel.2019.00451/full#supplementary-material

FIGURE S1 | Control stainings of the cerebellum to validate the specificity of anti-GluA antibodies. GluA1 was detected in Purkinje neurons (arrowhead) and Bergmann glial cells (arrow), GluA2 in Purkinje neurons and GluA4 in Bergmann glial cells, in line with published data. Scale bars: 50 μm.

FIGURE S2 | ADP evokes Ca²⁺ transients but not membrane currents. (A) The application of kainate (100 μM) induced an inward current in OECs, whereas the application of ADP (100 μM) did not. (B) Average current amplitudes of kainate and ADP-induced currents in OECs. (C) Application of kainate and ADP induced Ca²⁺ transients in OECs. (D) Average amplitudes of kainate- and ADP-evoked Ca²⁺ transients.

References

Au, W. W., Treloar, H. B., and Greer, C. A. (2002). Sublaminar organization of the mouse olfactory bulb nerve layer. J. Comp. Neurol. 446, 68–80. doi: 10.1002/cne.10182

PubMed Abstract | CrossRef Full Text | Google Scholar

Bazargani, N., and Attwell, D. (2016). Astrocyte calcium signaling: the third wave. Nat. Neurosci. 19, 182–189. doi: 10.1038/nn.4201

PubMed Abstract | CrossRef Full Text | Google Scholar

Beiersdorfer, A., Scheller, A., Kirchhoff, F., and Lohr, C. (2019). Panglial gap junctions between astrocytes and olfactory ensheathing cells mediate transmission of Ca(2+) transients and neurovascular coupling. Glia 67, 1385–1400. doi: 10.1002/glia.23613

PubMed Abstract | CrossRef Full Text | Google Scholar

Belkacemi, T., Niermann, A., Hofmann, L., Wissenbach, U., Birnbaumer, L., Leidinger, P., et al. (2017). TRPC1- and TRPC3-dependent Ca(2+) signaling in mouse cortical astrocytes affects injury-evoked astrogliosis in vivo. Glia 65, 1535–1549. doi: 10.1002/glia.23180

PubMed Abstract | CrossRef Full Text | Google Scholar

Berkowicz, D. A., Trombley, P. Q., and Shepherd, G. M. (1994). Evidence for glutamate as the olfactory receptor cell neurotransmitter. J. Neurophysiol. 71, 2557–2561. doi: 10.1152/jn.1994.71.6.2557

PubMed Abstract | CrossRef Full Text | Google Scholar

Burnashev, N., Khodorova, A., Jonas, P., Helm, P. J., Wisden, W., Monyer, H., et al. (1992). Calcium-permeable AMPA-kainate receptors in fusiform cerebellar glial cells. Science 256, 1566–1570. doi: 10.1126/science.1317970

PubMed Abstract | CrossRef Full Text | Google Scholar

Cao, L., Zhu, Y. L., Su, Z., Lv, B., Huang, Z., Mu, L., et al. (2007). Olfactory ensheathing cells promote migration of Schwann cells by secreted nerve growth factor. Glia 55, 897–904. doi: 10.1002/glia.20511

PubMed Abstract | CrossRef Full Text | Google Scholar

Carmignoto, G., and Gomez-Gonzalo, M. (2010). The contribution of astrocyte signalling to neurovascular coupling. Brain Res. Rev. 63, 138–148. doi: 10.1016/j.brainresrev.2009.11.007

PubMed Abstract | CrossRef Full Text | Google Scholar

Crandall, J. E., Dibble, C., Butler, D., Pays, L., Ahmad, N., Kostek, C., et al. (2000). Patterning of olfactory sensory connections is mediated by extracellular matrix proteins in the nerve layer of the olfactory bulb. J. Neurobiol. 45, 195–206. doi: 10.1002/1097-4695(200012)45:4<195::aid-neu1>3.0.co;2-y

PubMed Abstract | CrossRef Full Text | Google Scholar

Deitmer, J. W., Verkhratsky, A. J., and Lohr, C. (1998). Calcium signalling in glial cells. Cell Calcium 24, 405–416. doi: 10.1016/s0143-4160(98)90063-x

PubMed Abstract | CrossRef Full Text | Google Scholar

Dickinson, P. J., Griffiths, I. R., Barrie, J. M., Kyriakides, E., Pollock, G. F., and Barnett, S. C. (1997). Expression of the dm-20 isoform of the plp gene in olfactory nerve ensheathing cells: evidence from developmental studies. J. Neurocytol. 26, 181–189. doi: 10.1023/a:1018584013739

PubMed Abstract | CrossRef Full Text | Google Scholar

Doucette, J. R. (1984). The glial cells in the nerve fiber layer of the rat olfactory bulb. Anat. Rec. 210, 385–391. doi: 10.1002/ar.1092100214

PubMed Abstract | CrossRef Full Text | Google Scholar

Doucette, R. (1990). Glial influences on axonal growth in the primary olfactory system. Glia 3, 433–449. doi: 10.1002/glia.440030602

PubMed Abstract | CrossRef Full Text | Google Scholar

Doucette, R. (1991). PNS-CNS transitional zone of the first cranial nerve. J. Comp. Neurol. 312, 451–466. doi: 10.1002/cne.903120311

PubMed Abstract | CrossRef Full Text | Google Scholar

Droste, D., Seifert, G., Seddar, L., Jadtke, O., Steinhauser, C., and Lohr, C. (2017). Ca(2+)-permeable AMPA receptors in mouse olfactory bulb astrocytes. Sci. Rep. 7:44817. doi: 10.1038/srep44817

PubMed Abstract | CrossRef Full Text | Google Scholar

Flavell, S. W., and Greenberg, M. E. (2008). Signaling mechanisms linking neuronal activity to gene expression and plasticity of the nervous system. Annu. Rev. Neurosci. 31, 563–590. doi: 10.1146/annurev.neuro.31.060407.125631

CrossRef Full Text | Google Scholar

Graziadei, G. A., and Graziadei, P. P. (1979). Neurogenesis and neuron regeneration in the olfactory system of mammals. II. Degeneration and reconstitution of the olfactory sensory neurons after axotomy. J. Neurocytol. 8, 197–213. doi: 10.1007/bf01175561

PubMed Abstract | CrossRef Full Text | Google Scholar

Griffiths, I. R., Dickinson, P., and Montague, P. (1995). Expression of the proteolipid protein gene in glial cells of the post-natal peripheral nervous system of rodents. Neuropathol. Appl. Neurobiol. 21, 97–110. doi: 10.1111/j.1365-2990.1995.tb01035.x

PubMed Abstract | CrossRef Full Text | Google Scholar

Guerra-Gomes, S., Sousa, N., Pinto, L., and Oliveira, J. F. (2017). Functional roles of astrocyte calcium elevations: from synapses to behavior. Front. Cell Neurosci. 11:427. doi: 10.3389/fncel.2017.00427

PubMed Abstract | CrossRef Full Text | Google Scholar

Hayat, S., Wigley, C. B., and Robbins, J. (2003). Intracellular calcium handling in rat olfactory ensheathing cells and its role in axonal regeneration. Mol. Cell. Neurosci. 22, 259–270. doi: 10.1016/s1044-7431(03)00051-4

PubMed Abstract | CrossRef Full Text | Google Scholar

Haydon, P. G. (2001). GLIA: listening and talking to the synapse. Nat. Rev. Neurosci. 2, 185–193. doi: 10.1038/35058528

PubMed Abstract | CrossRef Full Text | Google Scholar

Herrera, L. P., Casas, C. E., Bates, M. L., and Guest, J. D. (2005). Ultrastructural study of the primary olfactory pathway in Macaca fascicularis. J. Comp. Neurol. 488, 427–441. doi: 10.1002/cne.20588

PubMed Abstract | CrossRef Full Text | Google Scholar

Hoft, S., Griemsmann, S., Seifert, G., and Steinhauser, C. (2014). Heterogeneity in expression of functional ionotropic glutamate and GABA receptors in astrocytes across brain regions: insights from the thalamus. Philos. Trans. R. Soc. Lond. B Biol. Sci. 369, 20130602. doi: 10.1098/rstb.2013.0602

PubMed Abstract | CrossRef Full Text | Google Scholar

Hollmann, M., Hartley, M., and Heinemann, S. (1991). Ca²⁺ permeability of KA-AMPA–gated glutamate receptor channels depends on subunit composition. Science 252, 851–853. doi: 10.1126/science.1709304

PubMed Abstract | CrossRef Full Text | Google Scholar

Horning, M. S., Kwon, B., Blakemore, L. J., Spencer, C. M., Goltz, M., Houpt, T. A., et al. (2004). Alpha-amino-3-hydroxy-5-methyl-4-isoxazolepropionate receptor subunit expression in rat olfactory bulb. Neurosci. Lett. 372, 230–234. doi: 10.1016/j.neulet.2004.09.044

PubMed Abstract | CrossRef Full Text | Google Scholar

Jabs, R., Kirchhoff, F., Kettenmann, H., and Steinhauser, C. (1994). Kainate activates Ca(2+)-permeable glutamate receptors and blocks voltage-gated K+ currents in glial cells of mouse hippocampal slices. Pflugers. Arch. 426, 310–319. doi: 10.1007/bf00374787

PubMed Abstract | CrossRef Full Text | Google Scholar

Koike, M., Iino, M., and Ozawa, S. (1997). Blocking effect of 1-naphthyl acetyl spermine on Ca(2+)-permeable AMPA receptors in cultured rat hippocampal neurons. Neurosci. Res. 29, 27–36. doi: 10.1016/s0168-0102(97)00067-9

PubMed Abstract | CrossRef Full Text | Google Scholar

Lalo, U., Pankratov, Y., Kirchhoff, F., North, R. A., and Verkhratsky, A. (2006). NMDA receptors mediate neuron-to-glia signaling in mouse cortical astrocytes. J. Neurosci. 26, 2673–2683. doi: 10.1523/JNEUROSCI.4689-05.2006

PubMed Abstract | CrossRef Full Text | Google Scholar

Leone, D. P., Genoud, S., Atanasoski, S., Grausenburger, R., Berger, P., Metzger, D., et al. (2003). Tamoxifen-inducible glia-specific Cre mice for somatic mutagenesis in oligodendrocytes and Schwann cells. Mol. Cell. Neurosci. 22, 430–440. doi: 10.1016/s1044-7431(03)00029-0

PubMed Abstract | CrossRef Full Text | Google Scholar

Lohr, C., Grosche, A., Reichenbach, A., and Hirnet, D. (2014). Purinergic neuron-glia interactions in sensory systems. Pflugers Arch. 466, 1859–1872. doi: 10.1007/s00424-014-1510-6

PubMed Abstract | CrossRef Full Text | Google Scholar

Madisen, L., Zwingman, T. A., Sunkin, S. M., Oh, S. W., Zariwala, H. A., Gu, H., et al. (2010). A robust and high-throughput Cre reporting and characterization system for the whole mouse brain. Nat. Neurosci. 13, 133–140. doi: 10.1038/nn.2467

PubMed Abstract | CrossRef Full Text | Google Scholar

McConkey, D. J., and Orrenius, S. (1997). The role of calcium in the regulation of apoptosis. Biochem. Biophys. Res. Commun. 239, 357–366. doi: 10.1006/bbrc.1997.7409

PubMed Abstract | CrossRef Full Text | Google Scholar

Mishra, A., Reynolds, J. P., Chen, Y., Gourine, A. V., Rusakov, D. A., and Attwell, D. (2016). Astrocytes mediate neurovascular signaling to capillary pericytes but not to arterioles. Nat. Neurosci. 19, 1619–1627. doi: 10.1038/nn.4428

PubMed Abstract | CrossRef Full Text | Google Scholar

Montague, A. A., and Greer, C. A. (1999). Differential distribution of ionotropic glutamate receptor subunits in the rat olfactory bulb. J. Comp. Neurol. 405, 233–246. doi: 10.1002/(sici)1096-9861(19990308)405:2<233:aid-cne7<3.0.co;2-a

PubMed Abstract | CrossRef Full Text | Google Scholar

Mori, T., Tanaka, K., Buffo, A., Wurst, W., Kuhn, R., and Gotz, M. (2006). Inducible gene deletion in astroglia and radial glia–a valuable tool for functional and lineage analysis. Glia 54, 21–34. doi: 10.1002/glia.20350

PubMed Abstract | CrossRef Full Text | Google Scholar

Muller, T., Moller, T., Berger, T., Schnitzer, J., and Kettenmann, H. (1992). Calcium entry through kainate receptors and resulting potassium-channel blockade in Bergmann glial cells. Science 256, 1563–1566. doi: 10.1126/science.1317969

PubMed Abstract | CrossRef Full Text | Google Scholar

Noda, M., Nakanishi, H., Nabekura, J., and Akaike, N. (2000). AMPA-kainate subtypes of glutamate receptor in rat cerebral microglia. J. Neurosci. 20, 251–258. doi: 10.1523/jneurosci.20-01-00251.2000

PubMed Abstract | CrossRef Full Text | Google Scholar

Piantanida, A. P., Acosta, L. E., Brocardo, L., Capurro, C., Greer, C. A., and Rela, L. (2019). Selective Cre-mediated gene deletion identifies connexin 43 as the main connexin channel supporting olfactory ensheathing cell networks. J. Comp. Neurol. 527, 1278–1289. doi: 10.1002/cne.24628

PubMed Abstract | CrossRef Full Text | Google Scholar

Porter, J. T., and McCarthy, K. D. (1995). GFAP-positive hippocampal astrocytes in situ respond to glutamatergic neuroligands with increases in [Ca2+]i. Glia 13, 101–112. doi: 10.1002/glia.440130204

PubMed Abstract | CrossRef Full Text | Google Scholar

Rakers, C., Schmid, M., and Petzold, G. C. (2017). TRPV4 channels contribute to calcium transients in astrocytes and neurons during peri-infarct depolarizations in a stroke model. Glia 65, 1550–1561. doi: 10.1002/glia.23183

PubMed Abstract | CrossRef Full Text | Google Scholar

Rela, L., Bordey, A., and Greer, C. A. (2010). Olfactory ensheathing cell membrane properties are shaped by connectivity. Glia 58, 665–678. doi: 10.1002/glia.20953

PubMed Abstract | CrossRef Full Text | Google Scholar

Rieger, A., Deitmer, J. W., and Lohr, C. (2007). Axon-glia communication evokes calcium signaling in olfactory ensheathing cells of the developing olfactory bulb. Glia 55, 352–359. doi: 10.1002/glia.20460

PubMed Abstract | CrossRef Full Text | Google Scholar

Rozas, J. L., Paternain, A. V., and Lerma, J. (2003). Noncanonical signaling by ionotropic kainate receptors. Neuron 39, 543–553. doi: 10.1016/s0896-6273(03)00436-7

PubMed Abstract | CrossRef Full Text | Google Scholar

Rungta, R. L., Bernier, L. P., Dissing-Olesen, L., Groten, C. J., LeDue, J. M., Ko, R., et al. (2016). Ca(2+) transients in astrocyte fine processes occur via Ca(2+) influx in the adult mouse hippocampus. Glia 64, 2093–2103. doi: 10.1002/glia.23042

PubMed Abstract | CrossRef Full Text | Google Scholar

Saab, A. S., Neumeyer, A., Jahn, H. M., Cupido, A., Simek, A. A., Boele, H. J., et al. (2012). Bergmann glial AMPA receptors are required for fine motor coordination. Science 337, 749–753. doi: 10.1126/science.1221140

PubMed Abstract | CrossRef Full Text | Google Scholar

Schipke, C. G., Ohlemeyer, C., Matyash, M., Nolte, C., Kettenmann, H., and Kirchhoff, F. (2001). Astrocytes of the mouse neocortex express functional N-methyl-D-aspartate receptors. FASEB J. 15, 1270–1272. doi: 10.1096/fj.00-0439fje

PubMed Abstract | CrossRef Full Text | Google Scholar

Seifert, G., and Steinhauser, C. (1995). Glial cells in the mouse hippocampus express AMPA receptors with an intermediate Ca²⁺ permeability. Eur. J. Neurosci. 7, 1872–1881. doi: 10.1111/j.1460-9568.1995.tb00708.x

PubMed Abstract | CrossRef Full Text | Google Scholar

Singaravelu, K., Lohr, C., and Deitmer, J. W. (2006). Regulation of store-operated calcium entry by calcium-independent phospholipase A2 in rat cerebellar astrocytes. J. Neurosci. 26, 9579–9592. doi: 10.1523/JNEUROSCI.2604-06.2006

PubMed Abstract | CrossRef Full Text | Google Scholar

Stavermann, M., Buddrus, K., St John, J. A., Ekberg, J. A., Nilius, B., Deitmer, J. W., et al. (2012). Temperature-dependent calcium-induced calcium release via InsP3 receptors in mouse olfactory ensheathing glial cells. Cell Calcium 52, 113–123. doi: 10.1016/j.ceca.2012.04.017

PubMed Abstract | CrossRef Full Text | Google Scholar

Steinhauser, C., and Gallo, V. (1996). News on glutamate receptors in glial cells. Trends Neurosci. 19, 339–345. doi: 10.1016/0166-2236(96)10043-6

PubMed Abstract | CrossRef Full Text | Google Scholar

Stosiek, C., Garaschuk, O., Holthoff, K., and Konnerth, A. (2003). In vivo two-photon calcium imaging of neuronal networks. Proc. Natl. Acad. Sci. U.S.A. 100, 7319–7324. doi: 10.1073/pnas.1232232100

PubMed Abstract | CrossRef Full Text | Google Scholar

Su, Z., and He, C. (2010). Olfactory ensheathing cells: biology in neural development and regeneration. Prog. Neurobiol. 92, 517–532. doi: 10.1016/j.pneurobio.2010.08.008

PubMed Abstract | CrossRef Full Text | Google Scholar

Takago, H., Nakamura, Y., and Takahashi, T. (2005). G protein-dependent presynaptic inhibition mediated by AMPA receptors at the calyx of Held. Proc. Natl. Acad. Sci. U.S.A. 102, 7368–7373. doi: 10.1073/pnas.0408514102

PubMed Abstract | CrossRef Full Text | Google Scholar

Thyssen, A., Hirnet, D., Wolburg, H., Schmalzing, G., Deitmer, J. W., and Lohr, C. (2010). Ectopic vesicular neurotransmitter release along sensory axons mediates neurovascular coupling via glial calcium signaling. Proc. Natl. Acad. Sci. U.S.A. 107, 15258–15263. doi: 10.1073/pnas.1003501107

PubMed Abstract | CrossRef Full Text | Google Scholar

Toth, A. B., Hori, K., Novakovic, M. M., Bernstein, N. G., Lambot, L., and Prakriya, M. (2019). CRAC channels regulate astrocyte Ca(2+) signaling and gliotransmitter release to modulate hippocampal GABAergic transmission. Sci. Signal. 12:eaaw5450. doi: 10.1126/scisignal.aaw5450

PubMed Abstract | CrossRef Full Text | Google Scholar

Verkhratsky, A., Orkand, R. K., and Kettenmann, H. (1998). Glial calcium: homeostasis and signaling function. Physiol. Rev. 78, 99–141. doi: 10.1152/physrev.1998.78.1.99

PubMed Abstract | CrossRef Full Text | Google Scholar

Wakakura, M., and Yamamoto, N. (1994). Cytosolic calcium transient increase through the AMPA/kainate receptor in cultured Muller cells. Vision Res. 34, 1105–1109. doi: 10.1016/0042-6989(94)90293-3

PubMed Abstract | CrossRef Full Text | Google Scholar

Wang, Y., Small, D. L., Stanimirovic, D. B., Morley, P., and Durkin, J. P. (1997). AMPA receptor-mediated regulation of a Gi-protein in cortical neurons. Nature 389, 502–504. doi: 10.1038/39062

PubMed Abstract | CrossRef Full Text | Google Scholar

Woodhall, E., West, A. K., and Chuah, M. I. (2001). Cultured olfactory ensheathing cells express nerve growth factor, brain-derived neurotrophic factor, glia cell line-derived neurotrophic factor and their receptors. Brain Res. Mol. Brain Res. 88, 203–213. doi: 10.1016/s0169-328x(01)00044-4

PubMed Abstract | CrossRef Full Text | Google Scholar

Yang, H., He, B. R., and Hao, D. J. (2015). Biological roles of olfactory ensheathing cells in facilitating neural regeneration: a systematic review. Mol. Neurobiol. 51, 168–179. doi: 10.1007/s12035-014-8664-2

PubMed Abstract | CrossRef Full Text | Google Scholar

Zonouzi, M., Renzi, M., Farrant, M., and Cull-Candy, S. G. (2011). Bidirectional plasticity of calcium-permeable AMPA receptors in oligodendrocyte lineage cells. Nat. Neurosci. 14, 1430–1438. doi: 10.1038/nn.2942

PubMed Abstract | CrossRef Full Text | Google Scholar