Increases in Intracellular Calcium Triggered by Channelrhodopsin-2 Potentiate the Response of Metabotropic Glutamate Receptor mGluR7

Abstract

The metabotropic glutamate receptor 7a (mGluR7a), a heptahelical Gα_i/o-coupled protein, has been shown to be important for presynaptic feedback inhibition at central synapses and certain forms of long term potentiation and long term depression. The intracellular C terminus of mGluR7a interacts with calmodulin in a Ca²⁺-dependent manner, and calmodulin antagonists have been found to abolish presynaptic inhibition of glutamate release in neurons and mGluR7a-induced activation of G-protein-activated inwardly rectifying K⁺ channel (GIRK) channels in HEK293 cells. Here, we characterized the Ca²⁺ dependence of mGluR7a signaling in Xenopus oocytes by using channelrhodopsin-2 (ChR2), a Ca²⁺-permeable, light-activated ion channel for triggering Ca²⁺ influx, and a GIRK3.1/3.2 concatemer to monitor mGluR7a responses. Application of the agonist (S)-2-amino-4-phosphonobutanoic acid (l-AP4) (1–100 μm) caused a dose-dependent inward current in high K⁺ solutions due to activation of GIRK channels by G-protein βγ subunits released from mGluR7a. Elevation of intracellular free Ca²⁺ by light stimulation of ChR2 markedly increased the amplitude of l-AP4 responses, and this effect was attenuated by the calcium chelator 1,2-bis(2-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid tetrakis (acetoxymethyl ester). l-AP4 responses were potentiated by submembranous [Ca²⁺] levels within physiological ranges and with a threshold close to resting [Ca²⁺]_i values, as determined by recording the endogenous Xenopus Ca²⁺-activated chloride conductance. Together, these results show that l-AP4-dependent mGluR7a signaling is potentiated by physiological levels of [Ca²⁺]_i, consistent with a model in which presynaptic mGluR7a acts as a coincidence detector of Ca²⁺ influx and glutamate release.

Metabotropic glutamate receptors (mGluRs)4 are G-protein-coupled receptors that play a key role in synaptic transmission. Eight mGluRs have been identified and grouped into three classes (1). mGluR7 is a Gα_i/o protein-linked member of the group III mGluRs that is widely expressed at both glutamatergic and non-glutamatergic synapses and has a crucial role in some forms of hippocampal plasticity (2, 3). In addition, mGluR7^–/– mice develop spontaneous seizures (4). Therefore, mGluR7 is a current target for the development of novel antiepileptic drugs.

mGluR7 has been localized to presynaptic terminals, where it inhibits neurotransmitter release via a G-protein-dependent pathway (5). In cortical presynaptic terminals, mGluR7 colocalizes primarily with and inhibits N-type Ca²⁺ channels. However, in cerebellar granule cells, mGluR7 inhibits P/Q-type Ca²⁺ channels via a phospholipase C-dependent pathway (6). mGluR7 may act as a presynaptic feedback inhibitor; elevated synaptic glutamate activates mGluR7, thereby decreasing presynaptic Ca²⁺ influx and reducing glutamate release (7).

Biochemical studies indicate that mGluR7 signaling is regulated by a variety of intracellular proteins. Specifically, the cytoplasmic C-terminal tail of the predominant splice variant mGluR7a has been shown to bind proteins whose properties are directly or indirectly influenced by changes in intracellular Ca²⁺ levels, such as calmodulin (CaM) (8), protein kinase C (9), and protein interacting with protein kinase C 1 (PICK-1) (10, 11). CaM binds to mGluR7a (and other group III mGluRs) in a Ca²⁺-dependent manner, and CaM inhibitors have been shown to prevent presynaptic inhibition at glutamatergic autapses and to disrupt mGluR7a signaling in Xenopus oocytes (8). Moreover, inhibition of P/Q voltage-gated calcium channels by l-AP4 is blocked by BAPTA, suggesting that Ca²⁺ is required for mGluR7-mediated presynaptic inhibition in cerebellar granule cells (6). These data support the view that presynaptic inhibition by mGluR7 depends on [Ca²⁺]_i, likely through regulation of complex protein interactions in the presynaptic nerve terminal (reviews in Refs. 12 and 13).

Here, we tested the dependence of mGluR7a activation on submembranous Ca²⁺ by using the G-protein-activated inwardly rectifying K⁺ channel (GIRK; Kir3.1/3.2) in a heterologous expression system in which we could temporally control Ca²⁺ influx with channelrhodopsin-2 (ChR2), a light-activated cation channel (14), monitor submembranous Ca²⁺ levels with the endogenous Ca²⁺-activated chloride conductance (15), and study mGluR7a in relative isolation. mGluR7a, the Gβγ effector GIRK, and ChR2 were co-expressed in Xenopus oocytes, and membrane currents were measured under two-electrode voltage clamp. Our results show that mGluR7a signaling is potentiated with increasing [Ca²⁺]_i.

EXPERIMENTAL PROCEDURES

Chemicals—l-AP4 was obtained from Calbiochem (Darmstadt, Germany), and niflumic acid (NFA) was obtained from Cayman Chemical (Ann Arbor, MI). Stock l-AP4 (100 mm) was dissolved in 100 mm NaOH and stored at –20 °C. l-AP4 was diluted 10³-10⁵-fold for experiments (working concentration 1–100 μm). NFA was dissolved in DMSO (300 mm stock), kept frozen at –20 °C, and diluted 1000-fold for a working concentration of 300 μm. The K_i for NFA block of Ca-activated Cl channels in oocytes is 10 μm. BAPTA-AM (Sigma product number A1076) was dissolved in DMSO, stored under argon at a stock concentration of 100–300 mm, and diluted at least 1000-fold for experiments.

Expression Constructs and RNA Synthesis—The mGluR7a cDNA encoding the short isoform of mGluR7 (Ref. 16); (gift from Dr. S. Nakanishi, University of Kyoto, Japan) and the GIRK (Kir3.1/3.2) construct (gift from Dr. Andreas Karschin, University of Würzburg, Germany) have been described previously (8). The ChR2 construct used was a C-terminally truncated version (amino acids 2–315) that lacks most of the C-terminal cytoplasmic domain (14) and contains a single point mutation (H134R), which results in larger currents but does not change ion selectivity (17). In vitro transcription was performed using commercial kits (Stratagene, Agilent Technologies, Santa Clara, CA, or Ambion, Applied Biosystems, Darmstadt, Germany). Synthetic RNAs were stored at –80 °C in aliquots at a concentration of 1 μg/μl.

Oocyte Expression and Electrophysiological Recordings—Stage V–VI oocytes were removed from anesthetized Xenopus laevis and enzymatically treated to remove follicular cells. The oocytes were incubated at 18 –19 °C in oocyte saline (ND-96, in mm: 96 NaCl, 2 KCl, 1 CaCl₂, 1 MgCl₂, 5 HEPES, pH 7.4) and injected with in vitro transcribed RNA as follows. For recordings from cells injected with mGluR7a and GIRK RNA, 1 μl (25 ng) of mGluR7a RNA was mixed with 1 μl of GIRK RNA (∼50 pg). Light-activable ChR2 has retinal bound (14). Hence, oocytes were injected on day 1 with a 1:1 mixture of GIRK and mGluR7a RNAs (50 nl/oocyte) and 48 h later with ChR2 RNA (20 ng) followed by incubation in oocyte solution containing trans-retinal (1 μm). Oocytes were used for recordings from day 3 to day 7 after the initial injection using two-electrode voltage clamp recording. The volume of the recording chamber was 15 μl, and this volume was exchanged with 0.5–1 ml of perfusion solution in ∼1s.K⁺ was substituted for Na⁺ in solutions in which K⁺ was elevated. Increases in Ca²⁺ were achieved by substituting for Na⁺. Electrodes had resistances of 0.4-2 megaohms and were filled with 3 m KCl in 0.2% (w/v) agar. Data were acquired with an Axon 2B amplifier, Digidata 1200 A/D converter, and pCLAMP 9 software (MDS Inc., Toronto, Canada).

Photoactivation of ChR2—ChR2 was activated in electrode-impaled oocytes with a mercury arc lamp and a 450 ± 25-nm band filter (5 milliwatt/mm²) as described (14). Neutral density filters with a computer-activated shutter were used to control light intensity.

RESULTS

Control of Submembranous Ca²⁺ with ChR2—ChR2 is a light-activated eukaryotic cation channel that is permeable to Ca²⁺ (14). We utilized ChR2 to introduce Ca²⁺ into the submembranous compartment of Xenopus oocytes to mimic Ca²⁺ influx through voltage-gated calcium channels into the presynaptic nerve terminal. We took advantage of the well characterized, endogenous Ca²⁺-activated chloride current (15, 18) to monitor free Ca²⁺ concentrations beneath the oocyte plasma membrane. Fig. 1A shows the voltage clamp protocol and resulting currents in the presence and absence of extracellular Ca²⁺. ChR2 was activated by illuminating the oocyte immediately before recording, and the oocyte remained illuminated during the record. In the absence of extracellular Ca²⁺, light-induced currents at +40 mV were small and stable. In the presence of 2 mm extracellular Ca²⁺, a large Cl^– current (I_Cl(Ca)) was evident during the second pulse to +40 mV (note also the Cl^– current during the pulse to –140 mV). Thus, Ca²⁺ entry via the ChR2 channel activates I_Cl(Ca).

FIGURE 1.

Activation of I_Cl(Ca) by ChR2. A, control of submembranous [Ca²⁺] by light activation of ChR2 and measurement of I_Cl(Ca) current. The voltage step protocol (top) applies to panels A and B; I_Cl(Ca) was recorded from an oocyte expressing GIRK, mGluR7a, and ChR2. The oocyte was illuminated with blue light (450 nm) during the entire record (3.5 s) at either 0 or 2 mm[Ca²⁺]_o. Ca²⁺ that enters during the voltage step to –140 mV activates I_Cl(Ca) that can be seen as an additional inward current at –140 mV and as the large transient outward current during the second step to +40 mV. B, dependence of chloride current on light intensity. In the presence of 2 mm [Ca²⁺]_o, increasing the intensity of illumination (1, 2, 3, 5, 10, 25, 50, and 100% of maximal intensity) produced correspondingly larger I_Cl(Ca) currents during the second step to +40 mV (peak current indicated by asterisk). C, dependence of peak chloride current upon ChR2 current. I_ChR2 was measured 50 ms after the transient phase of the ChR2 current during the voltage step to –140 mV, and I_Cl(Ca) was measured at the peak of the response during the second step to +40 mV (panel B, asterisk). Data (black squares) were fit with the Hill equation (solid line) of the form V = V_max/(1 + (k/x)ⁿ) with V_max = 7.8 μA, k = 0.46 μA, and n = 2.13. The lower abscissa, open circle data points, dashed line, and right ordinate are data replotted from Haase and Hartung (18), with the Hill equation fit with V_max normalized, k = 500 nm Ca²⁺, and n = 2.3. The dashed horizontal and vertical lines indicate the half-maximal current and K_d, respectively. D, time course of the block of I_Cl(Ca) by BAPTA-AM. Application of BAPTA-AM (100 μm in ND-96) caused a rapid block of the I_Cl(Ca) current (filled circles) while leaving I_ChR2 (filled squares) unaffected. The curve is a first-order exponential fit with τ = 1.4 min. I_Cl(Ca) was reduced to 20% of the original current within 5 min. Lower concentrations of BAPTA-AM took longer to block (for 40 μm, τ = 7 min).

The dependence of I_Cl(Ca) upon activation of ChR2 was determined by varying the light intensity. In the presence of 2 mm [Ca²⁺]_o, peak amplitudes of the Cl^– current were measured during the second step to +40 mV (Fig. 1B, asterisk) at increasing light intensities, which activate an increasing number of ChR2 channels. The relationship between I_Cl(Ca) and ChR2 current is plotted in Fig. 1C. The relationship was fit by the Hill equation (Fig. 1C, solid line), with k = 0.46 μA (value of the ChR2 current at the half-maximal response) and n = 2.13. This relationship is very similar to the relationship between [Ca²⁺] and I_Cl(Ca) as determined by Haase and Hartung (18), who applied Ca²⁺ to the intracellular face of excised inside-out patches of Xenopus oocyte membrane and determined a K_d value of 500 ± 20 nm and a Hill coefficient of n = 2.3 ± 0.12 (the average of their data is superimposed on our data in Fig. 1C). We assume that the K_d for Ca²⁺ determined from excised patches is a constant property of the Ca²⁺-activated chloride channel. Thus, by equating the value of k for the oocyte shown in Fig. 1C with the K_d of I_Cl(Ca) for Ca²⁺ determined by Haase and Hartung (18), we find that I_ChR2 = 0.46 μA corresponded to a free [Ca²⁺]_i of 500 nm. The [Ca²⁺]_i produced by smaller and larger currents was assumed to be proportional to the ChR2 current ([Ca²⁺]_i = (500/k_{I ChR2})* I_ChR2). The calibration shown in Fig. 1, B and C, was performed for each oocyte, and each ChR2 current was then converted to free submembranous [Ca²⁺].

Submembranous Ca²⁺ can be sequestered with Ca²⁺ chelators without altering Ca²⁺ flux through ChR2. Fig. 1D demonstrates that extracellular application of the fast, membrane-permeable Ca²⁺ chelator BAPTA-AM largely inhibited I_Cl(Ca) within 5–8 min (69.4 ± 5.7%; average ± S.E., n = 10) and had no effect on ChR2 current (Fig. 1D, squares). The failure to achieve complete block of I_Cl(Ca) raises the possibility that some ChR2 and chloride channels are spatially apposed or colocalize in macromolecular complexes. This block of I_Cl(Ca) corroborates that I_Cl(Ca) reflects [Ca²⁺]_i and that BAPTA-AM treatment does not affect ChR2 function and therefore transmembrane Ca²⁺ flux.

To characterize submembranous Ca²⁺ kinetics, we designed protocols to monitor the time course of ChR2-induced changes in [Ca²⁺]_i. In oocytes, the increase in submembranous [Ca²⁺] depends on the rates of both Ca²⁺ entry through ChR2 and Ca²⁺ buffering and removal. Fig. 2A shows the protocol and currents recorded for increasing durations of illumination. The peak Cl^– current is plotted versus light duration for a representative cell in Fig. 2B. The current increased with a time constant of 630 ± 125 ms (average ± S.E., n = 5). Thus, in 1–2 s, the intracellular Ca²⁺ concentration reaches a steady state. To determine the time required for [Ca²⁺]_i to return to the resting level, the light was turned off during the –140-mV step, and the second step to +40 mV was delayed in 100-ms increments (see the protocol illustrated in Fig. 2C, top). I_Cl(Ca) rapidly declined (peak current during the second step to +40 mV, Fig. 2C) following the termination of illumination. The time course for recovery is shown in Fig. 2D. The time constant for the return to resting [Ca²⁺]_i levels was 181 ± 14 ms (n = 4). It is possible that the kinetics of changes in I_Cl(Ca) are limited by the rates of activation and deactivation of the chloride channel. Evidence from excised patches of oocyte membrane where [Ca²⁺] was increased or decreased rapidly (within a few milliseconds or less) at the cytoplasmic surface have shown that, at the concentrations relevant for our experiments (∼0.5 μm), the time constant of I_Cl(Ca) activation is between 50 (15) and 500 ms (18); the latter value is close to the activation time constant we measured (Fig. 2C). Likewise, the deactivation time constants varied from 30 (15) to 100–200 ms (18). Again, the latter values are close to those measured here for changes in I_Cl(Ca) (Fig. 2D). Thus, our ability to measure changes in [Ca²⁺] could be limited by the inherent kinetics of the chloride channel, and changes in submembranous [Ca²⁺] may be faster than the time courses shown in Fig. 2, B and D. We conclude that ChR2 channel activation allows us to increase submembranous [Ca²⁺] in less than 1–2 s to known free Ca²⁺ levels, and once the light is turned off, [Ca²⁺]_i returns to basal levels in much less than a second. Thus, the temporal profile of submembranous [Ca²⁺] can be tightly controlled through illumination.

FIGURE 2.

Time course of light-induced changes in I_Cl(Ca). A, longer duration of ChR2 activation increases chloride peak currents during the second step to +40 mV. An arrow indicates onset of illumination, and filled dots indicate when illumination ceased and the membrane voltage was stepped to +40 mV. Illumination duration increased in increments of 100 ms. B, time course of the increase in chloride current (from records in panel A). Peak currents (during second step to +40 mV) were plotted versus illumination time. Data were fit by a single exponential with τ = 486 ms. C, delayed voltage step for I_Cl(Ca) shows that submembranous Ca²⁺ is cleared quickly after ChR2 inactivation. Illumination started at the arrow and ceased for all traces at the filled circle. The second step to +40 mV was delayed in increasing steps of 100 ms after the termination of illumination. D, decay of chloride current after cessation of illumination. Peak currents during the second step to +40 mV are plotted versus time after the light was turned off (from records in panel C). Data were fit by a single exponential with τ = 183 ms.

Potentiation of mGluR7a Agonist Responses by Changes in [Ca²⁺]—Having established a system in which submembranous [Ca²⁺]can be temporally controlled and accurately measured, we next characterized the Ca²⁺ dependence of mGluR7a signaling. Oocytes co-expressing mGluR7a, GIRK 3.1/3.2, and ChR2 were voltage-clamped, and in each oocyte, the relationship between I_ChR2 and I_Cl(Ca) was examined as described above. Subsequently, the Ca²⁺-activated chloride channel was blocked with NFA (300 μm). Following calibration of each oocyte and NFA treatment, the group III-specific agonist l-AP4 was applied. Repeated applications of the same concentration of l-AP4 during increasing levels of illumination, and therefore increasing levels of submembranous [Ca²⁺], were employed to determine changes in mGluR7a function in the presence of Ca²⁺, as described in more detail below. Fig. 3A illustrates a typical experiment. Oocytes were bathed in a solution containing 80 mm K⁺ (high potassium) to increase the flux of K⁺ through the inwardly rectifying GIRK channel. Once l-AP4-independent GIRK channel currents were stable, l-AP4 was applied in high potassium. This resulted in a characteristic inward current, indicative of l-AP4-dependent Gβγ activation of GIRK (19). Currents were recorded with [l-AP4] (100 μm) and all other conditions constant while varying illumination intensity (and thus submembranous [Ca²⁺]). As expected from Figs. 1 and 2, increasing light intensity resulted in larger ChR2 currents, as indicated by the short latency, fast-desensitizing currents immediately following illumination with a smaller maintained ChR2 current (Fig. 3A). l-AP4 was applied during the plateau phase of the ChR2 current. Most notably, as ChR2 current increased, the responses to l-AP4 were progressively potentiated in comparison with the l-AP4 currents obtained in the absence of illumination. Control experiments showed that l-AP4 responses were blocked by Ba²⁺, which inhibits GIRK currents, and that l-AP4 had no effect on ChR2 current (data not shown).

FIGURE 3.

Potentiation of mGluR7a agonist responses upon ChR2 activation. A, continuous recording from an oocyte co-expressing GIRK, mGluR7a, and ChR2. l-AP4 (100 μm) was applied during the periods indicated by bars. Arrows indicate the onset of illumination, and filled circles indicate the time at which the light was turned off. The relative light intensity (percentage of maximum) is indicated above each arrow. Increasing light intensities produced correspondingly larger ChR2 currents, with large initial transients followed by smaller plateau currents. The inset shows the transient and plateau ChR2 currents and the l-AP4 response at an expanded scale (scale bars are 100 nA and 5 s). The same concentration of l-AP4 resulted in larger responses with increasing intensity of illumination. Transients seen in the initial application of l-AP4 were not consistently observed. B, dependence of the l-AP4 response on [Ca²⁺]_i. Data are presented for five oocytes. [Ca²⁺]_i concentrations were calculated from calibrations done individually for each oocyte (as shown in Fig. 1C). Data were fit by with a binding isotherm (I_L-AP4 = B_Max × [Ca²⁺]_i/(K_d + [Ca²⁺]_i); see “Results”). Data for the cell in panel A are plotted as filled circles. Values for B_Max (in nA) and K_d (in nm) were 571 and 741 (▴); 330 and 233 (▾); 460 and 1205 (•); 205 and 71 (▪); and 99 and 417 (♦). C, inhibition by BAPTA-AM of light-induced mGluR7a potentiation. l-AP4 was applied to Xenopus oocytes expressing mGluR7a, GIRK, and ChR2. Responses to l-AP4 were compared in the absence and presence of ChR2 activation with blue light (left panel). Cells were then incubated with BAPTA-AM (100 μm) for 8–12 min, which is sufficient time for a maximal block of I_Cl(Ca), and l-AP4 responses were recorded again with and without light (right panel). Thin lines, no light; thick lines, ChR2 activation by light. The thin and thick traces were superimposed at the beginning of the l-AP4 application. The no light response to l-AP4 application tended to increase in the presence of BAPTA-AM, but this was not statistically significant (average increase = 24 ± 18%, S.E., n = 5, p = 0.12). D, as the inset shown in the lower right of panel A, but repeated illumination with the same light intensity in the presence of variable extracellular [Ca²⁺] (from 0.1 to 5 mm with the [Ca²⁺] shown adjacent to each response). l-AP4 responses are superimposed at the beginning of agonist application (note that the responses are small because l-AP4 was 1 μm, rather than 100 μm in panel A, and the driving force for K⁺ influx was smaller due to differences in the membrane potential, see below). E, the peak l-AP4 response (maximal response for 0.1 mm, and peaks indicated in panel D by asterisks below the 0.3–5 mm traces) are plotted (with others not illustrated in panel D) along with post-BAPTA-AM responses of the same cell (filled circles). Cells were voltage-clamped at –60 mV (panels C–E) or –100 mV (panels A and B).

Results similar to those shown in Fig. 3A were observed in seven out of eight oocytes (one oocyte had no increase in agonist-induced GIRK current). The peak l-AP4-induced current amplitude data for five oocytes are plotted in Fig. 3B as a function of [Ca²⁺]_i. All responses increased with increasing illumination, and these increases began at very low [Ca²⁺]_i (at or below ∼100 nm). The relationship between the agonist-induced currents and [Ca²⁺]_i could be fitted by binding isotherms, with a mean equilibrium dissociation constant of K_d = 533 ± 201 nm and a mean maximal current of 333 ± 84 nA (means ± S.E.). However, there was considerable variability between individual oocytes. This variability in the sensitivity of the response to [Ca²⁺]_i is likely to reflect differences in the expression ratios of GIRK to mGluR7a between oocytes.

To confirm that the potentiation of mGluR7a-stimulated GIRK currents observed upon light illumination is mediated by Ca²⁺, we tested whether BAPTA-AM inhibits light-dependent mGluR7a potentiation (Fig. 3C). BAPTA-AM blocked the ChR2-mediated increase in the mGluR7a response by 65 ± 12% (S.E.; n = 5, p = 0.002). This was similar to the inhibition of I_Cl(Ca) (69%, Fig. 1D) seen under the same conditions. These results are consistent with the ChR2-induced potentiation of mGluR7a currents being primarily mediated by an increase in [Ca²⁺]_i. To further confirm that the increased l-AP4 response seen in panel A is due to Ca²⁺ influx, we repeated the stimulation protocol shown in Fig. 3A with a constant light intensity while varying the extracellular concentration of Ca²⁺. Fig. 3D shows that the peak amplitude of the l-AP4 response increased with increasing [Ca²⁺]_o. The peak l-AP4 response in panel D and additional data are plotted in panel E (Fig. 3). Two additional points are evident from Fig. 3E. First, BAPTA-AM (filled circles) blocked a large portion of the Ca²⁺-dependent l-AP4 response (open circles). Second, at 10 mm [Ca²⁺]_o, the response decreased. The smaller agonist response at the highest [Ca²⁺]_o is likely caused by direct effects of extracellular Ca²⁺ on mGluR7 (see below).

Both modeling and experimental studies suggest that in a restricted space such as the synaptic cleft, extracellular Ca²⁺ can be transiently reduced during periods of high activity, producing short term synaptic depression (20). In addition, mGluRs belong to a superfamily of proteins that bind extracellular Ca²⁺ (21). mGluR1 has been reported to be activated by extracellular Ca²⁺ in the absence of agonist (22), and the agonist responses of the related GABA_B receptor are modulated by extracellular Ca²⁺ (23). Thus, mGluR7a signaling might be regulated not only by changes in [Ca²⁺]_i but also by extracellular Ca²⁺ levels. We therefore examined whether extracellular Ca²⁺ might activate mGluR7a in the absence of l-AP4 and whether mGluR7a currents elicited by l-AP4 are modulated by changes in the extracellular Ca²⁺ concentration. We found that extracellular Ca²⁺ alone was unable to elicit GIRK currents (Fig. 4A, left half, and 4B, filled circles) and that current responses to l-AP4 varied little between 100 μm and 2 mm [Ca²⁺]_o (Fig. 4A, right half, and Fig. 4B, open diamonds). Although extracellular Ca²⁺ modulates the activation of mGluR7a by agonist (l-AP4), this effect will be important only if extracellular Ca²⁺ is reduced to less than 0.1 mm (Fig. 4B), a value well below the changes predicted or measured under physiological conditions at the synapse. Hence, it appears unlikely that physiological changes in extracellular [Ca²⁺]_i influence mGluR7a signaling.

FIGURE 4.

Effects of extracellular Ca²⁺ upon basal and mGluR7a-activated GIRK currents. A, continuous two-electrode voltage clamp recording from an oocyte expressing GIRK and mGluR7a. The oocyte was superfused with normal saline solution (containing 2 mm K⁺) at the beginning of the recording. Replacement of normal saline by high K⁺ bath solution (96 mm K⁺) elicited a rapid inward current due to the basal activity of the GIRK channel. This basal current is invariably observed when mGluRs are expressed in Xenopus oocytes and is thought to be due to agonist-independent stimulation of GIRK by free Gβγ subunits (43). In our oocytes, basal GIRK currents changed very little over a wide range of [Ca²⁺]_o from the low nm range (zero added Ca²⁺ with 1 mm EGTA) up to 10 mm Ca²⁺ (panel B, filled circles). The effect of changing [Ca²⁺]_o on agonist-induced mGluR7a responses was tested in the right half of the panel. The addition of l-AP4 (100 μm) produced an additional inward current, and the amplitude of this l-AP4 current changed as [Ca²⁺]_o was varied from 10 to 0 mm (inward current was maximal at 1 mm Ca²⁺). B, average amplitudes (normalized for each cell to the amplitude at 0.1 mm Ca²⁺) were plotted as a function of [Ca²⁺]_o (open diamonds, l-AP4 response, n = 11; closed circles, basal response, n = 2). The zero Ca²⁺ data were plotted at 5 nm, although the true concentration may have been lower (zero added Ca²⁺ with 1 mm EGTA). Cells were voltage-clamped at –100 mV.

DISCUSSION

In this study, we provide direct evidence showing that elevated [Ca²⁺]_i modulates agonist activation of mGluR7a. Our results are based on the use of the light-activated cation channel ChR2 (14) for inducing Ca²⁺ influx and of the endogenous calcium-activated chloride channel of Xenopus oocytes for estimating free intracellular Ca²⁺ concentrations. The magnitudes of I_Cl(Ca) allowed us to determine free [Ca²⁺]_i at the inner face of the membrane, where regulation of mGluR7a activity occurs. Light activation of ChR2 provided rapid, reproducible, and readily reversible changes in submembranous [Ca²⁺]. Importantly, increases in submembranous [Ca²⁺] potentiated the mGluR7a response to the selective agonist l-AP4, as monitored by activation of co-expressed GIRK, and this potentiation could be detected at concentrations close to resting levels of [Ca²⁺]_i. A scheme summarizing the experimental system used and our interpretation of the results obtained are presented in Fig. 5. Accordingly, elevation of [Ca²⁺]_i by light activation of ChR2 results in Ca²⁺ binding to CaM or other Ca²⁺-sensitive proteins that interact with the C-terminal tail of mGluR7a in the presence of Ca²⁺. This would displace prebound Gβγ from mGluR7a and thereby trigger K⁺ influx via Gβγ binding to GIRK. Although not indicated in the figure, it is possible that Ca²⁺ and Ca²⁺-binding proteins act directly on GIRK. This alternative explanation seems unlikely because we have found no reports describing potentiation of GIRK currents by increases of intracellular Ca²⁺. Conversely, there is mixed evidence for a role of Ca²⁺ in G-protein-coupled receptor-mediated inhibition of GIRK currents, with some laboratories finding enhanced or accelerated inhibition and others finding no effect of Ca²⁺ (24–26). We observed no increase in deactivation or inhibition by Ca²⁺. The scheme in Fig. 5 agrees well with our previous model of presynaptic mGluR regulation by Ca²⁺-CaM (8, 12), which was based on the finding that Ca²⁺-CaM and Gβγ compete for overlapping binding sites in the C-terminal tail of mGluR7a that are conserved in other presynaptic group III mGluRs (27).

FIGURE 5.

Mechanism proposed for potentiation of mGluR7a agonist responses in Xenopus oocytes. Heterologously expressed proteins are underlined (GIRK, mGluR7a, and ChR2). The endogenous Ca²⁺-activated chloride channel and Ca²⁺-sensitive proteins (for example, calmodulin) are included, and Gβγ subunits are depicted as linked to the plasma membrane. mGluR7a exists as a dimer but is pictured as a monomer for simplicity. Blue light (450 nm) activates ChR2, producing a cation current that results in an influx of Ca²⁺. This activates the endogenous Ca²⁺-activated chloride channel. The current through this channel (I_Cl(Ca)) enables monitoring and calibration of submembranous [Ca²⁺] as a function of light intensity. Concurrently, Ca²⁺ activates Ca²⁺-sensitive proteins, which interact with mGluR7a and thus potentiate agonist-dependent Gβγ signaling. Gβγ signaling is monitored by GIRK activation and influx of K⁺.

What are the implications and predictions of our findings for synaptic physiology? Since mGluR7a is localized in presynaptic terminals of both excitatory and inhibitory synapses and known to inhibit presynaptic Ca²⁺ channels (6, 28), its sensitivity to [Ca²⁺]_i will provide a gain control for negative feedback inhibition of neurotransmitter release. Active synapses with elevated presynaptic [Ca²⁺]_i will be more potently inhibited upon mGluR7a activation. Such a regulation by Ca²⁺ implies that inhibition by mGluR7a will be enhanced if the receptor is positioned close to Ca²⁺ channels. Furthermore, the [Ca²⁺]_i required to modulate mGluR7a should lie in a range that is similar to the [Ca²⁺]_i, which triggers synaptic vesicle exocytosis. In the literature, the [Ca²⁺]_i values reported to initiate neurotransmitter release vary widely (reviewed in Ref. 29). In some excitable cells, exocytosis requires such a high Ca²⁺ concentration (∼100 μm) that the fusion machinery is assumed to be within 20 nm of the Ca²⁺ channels. For other neurons, lower values are reported, and the [Ca²⁺]_i necessary for exocytosis appears to reflect a more global concentration estimate within the terminal (neuromuscular junction, 2–4 μm (30); calyx of Held, ∼10 μm (31)). Thus, the range of free [Ca²⁺]_i over which mGluR7a modulation was observed here is well within or even below that initiating neurotransmitter release from presynaptic terminals. In conclusion, our results are consistent with the concept that presynaptic group III mGluRs act as activity-dependent regulators of neurotransmitter release.

The results described here demonstrate that, at non-saturating agonist concentrations, elevations in [Ca²⁺]_i result in increased mGluR7a responses. An unusual feature of mGluR7 is its reportedly low affinity for glutamate (K_d ∼1 mm; (2, 32)). Therefore, it has been suggested that mGluR7 is only activated during intense synaptic activity or under extreme circumstances, such as epileptic seizures or ischemia. Indeed, l-AP4 has been found to be neuroprotective under conditions of excitotoxicity, e.g. in the presence of elevated extracellular glutamate (33). The enhanced survival of cultured cerebellar neurons observed in the presence of this agonist does not involve large increases in [Ca²⁺]_i and appears to depend on mGluR7 (34). Additional studies also have implicated mGluR7, working by inhibition of glutamate release, in neuroprotection. The increased l-AP4 response seen here at elevated [Ca²⁺]_i (Fig. 3) predicts that concentrations of glutamate well below the K_d would be effective if intracellular Ca²⁺ levels were raised. Thus, mGluR7 may be important under physiological conditions, rather than solely providing neuroprotection in situations involving excessive stimulation.

The results reported here provide a new perspective for interpreting the phenotypes of mGluR knock-out mice. Synaptic facilitation depends on the [Ca²⁺]_i level in the presynaptic terminal. Potentiation of mGluR7a by increased [Ca²⁺]_i, as reported here, should enhance feedback inhibition of Ca²⁺ channels and thus accelerate recovery from facilitation. Consistent with this prediction, mGluR7-deficient mice show a delayed recovery from synaptic facilitation (4). A train of action potentials can increase presynaptic [Ca²⁺]_i, and under these conditions, mGluR7 responses should be enhanced and their speed increased. Thus, mGluR7 could act as a frequency-dependent filter for synaptic transmission by more effectively reducing release at high frequencies. The magnitude of this effect may be modest since there is evidence at the calyx of Held that presynaptic mGluRs contribute only about 10% to synaptic depression (35). However, even a 10% change may be sufficient to produce susceptibility to epilepsy, and indeed, mGluR7 knock-out mice are prone to epileptic seizures, whereas mice deficient in mGluRs 1, 2, 4, 5, 6, or 8 do not show such a phenotype (4).

Increases in [Ca²⁺]_i can result not only from the opening of voltage-gated Ca²⁺ channels but also from the activation of intracellular Ca²⁺ stores. Thus, Ca²⁺ release from intracellular stores might provide a molecular link between different mGluR subtypes because activation of group I mGluRs results in the production of inositol 1,4,5-trisphosphate, and hence, the release of stored Ca²⁺ (36). Since both group I and III receptors have been shown to be presynaptic at the Schaffer collateral-CA1 synapse in the hippocampus (37), release of intracellular Ca²⁺ could create another pathway for synergy between mGluRs, as postulated for some aspects of long term potentiation and long term depression (38).

Long term depression is the most obvious form of synaptic plasticity in which mGluR7 might be involved since mGluR7 inhibits Ca²⁺ channels and synaptic transmission. Indeed, at glutamatergic synapses formed by hippocampal mossy fibers on stratum lucidum interneurons, high frequency stimulation has been shown to induce presynaptic long term depression that is mimicked by l-AP4 at concentrations sufficient to activate mGluR7 and is blocked by the group III-specific mGluR antagonist (RS)-α-methylserine-O-phosphate (3). However, mGluR7 is also present at inhibitory nerve terminals, and inhibition of such terminals should produce excitation due to a reduced release of GABA. Consistent with such a mechanism, activation of group III mGluRs has been found to facilitate glutamate release in some layers of the cortex (39). Also, glutamate spillover resulting from mossy fiber stimulation in cerebellar glomeruli has been shown to suppress GABA release via presynaptic mGluR activation (40). All these results are consistent with an important role of group III mGluRs in bidirectional synaptic modifiability. Furthermore, several lines of evidence indicate that the magnitude of the rise in presynaptic [Ca²⁺]_i following stimulation is a major factor in determining whether a synapse shows potentiation or depression (41, 42). It therefore is tempting to speculate that the sensitivity of mGluRs to [Ca²⁺]_i described here for mGluR7a might play an important role in the selection of long term potentiation versus long term depression.