GluD2 is an ionotropic glutamate receptor (iGluR) family member 1 almost exclusively expressed by cerebellar Purkinje cells 2 . Its absence or mutation causes ataxia, deficits in motor learning 3 , and cognitive disorders in rodents 4 5 6 7 . Its N‐ and C‐terminal domains respectively control parallel fiber (PF) synapse development/maintenance and long‐term depression (LTD) 3 7 8 9 10 . Evidence that the channel‐pore of GluD2 is functional exists 1 10 11 , but this point is still debated since no gating ligand has ever been identified 12 . Interestingly, the subtype 1 metabotropic glutamate receptor (mGlu1) associates with GluD2 and transient receptor potential cation channel TRPC3 in Purkinje cells 13 . Moreover, some mGlu1‐activated conductances share properties with GluD2 14 . The occurrence of a functional crosstalk between mGlu1 and iGluRs 15 16 17 and the report of a mGlu1‐mediated slow current carried by TRPCs 18 19 20 prompted us to investigate if a similar functional coupling occurs between mGlu1 and GluD2 receptors. Here we show that mGlu1 indeed triggers the opening of the GluD2 channel.

Representative PF‐slow ESPCs (C) and DHPG currents (D) from WT (left) and HO‐Nancy (right) Purkinje cells. Histograms: averaged peak amplitude of PF‐slow ESPCs (C) or DHPG currents (D) from all the cells (± s.e.m.).

Mice from the Hotfoot (HO) family bear deletions in the GluD2 encoding gene Grid2 . The HO‐Nancy GluD2 proteins lack their channel and ligand‐binding domain (LBD) ( 28 6 , Fig 3 A), providing a model to test mGlu1 currents in the absence of GluD2 pore. Immunohistochemistry in WT mice confirmed that GluD2s locate in thin dendrites of Purkinje cells ( 7 , Fig 3 B). In HO‐Nancy, although a fraction of GluD2 proteins remained trapped at the level of Purkinje cells somata (Fig 3 B), their dendrites, including proximal ones, were also labeled, supporting the previous observation that HO‐Nancy GluD2s reach the membrane 8 . An anti‐GluD1/2 antibody gave similar results ( supplementary Fig S5 ). Then, we quantified the mGluR1‐mediated PF‐slow EPSC in HO‐Nancy Purkinje cells using calibrated PFs stimulation to compare with WT (see supplementary Fig S4B–C ). Half of the HO‐Nancy cells display no PF‐slow EPSC ( n = 19/37, Fig 3 C, left trace on right panel, versus n = 4/72 in WT). In the 18 others, a small slow EPSC was recorded (71 ± 18 pA), which was blocked by AIDA (Fig 3 C, right trace on right panel). In the HO‐4j mice, extrasynaptic mGlu1 currents were shown to increase 13 . To record both synaptic and extrasynaptic mGlu1‐currents, we used bath applications of DHPG (50 μM, 30 s) in the presence of TTX, NBQX, D‐APV and bicuculline (resp. 1, 10, 50, 20 μM). The DHPG current was detectable in all the HO‐Nancy Purkinje cells but was smaller than in WT cells, (respectively 234 ± 60 pA, n = 13 versus 773 pA ± 136 pA, n = 9, P < 0.01; Fig 3 D). Thus, in Ho‐Nancy Purkinje cells, mGlu1 currents are smaller than in WT cells, the reduction being more pronounced at synapses than outside. Importantly, this does not result from a reduced number of mGlu1s at the membrane since their amount does not change in the absence of GluD2 3 13 . Together, these data suggest that GluD2 channels participate to the mGlu1‐activated currents in Purkinje cells, and support our previous conclusion that GluD2 gating is triggered by mGlu1s.

We induced this slow mGlu1‐mediated EPSC with 8 pulses of 100 Hz PF‐stimulation (Fig 2 A), in the presence of GABA‐A, AMPA and NMDA receptor inhibitors (bicuculline, NBQX, D‐APV, resp. 20, 10, 50 μM). Its intensity was 281 ± 26 pA ( n = 46, Fig 2 B), and it was blocked by the mGluR1/5 antagonist AIDA (150 μM, supplementary Fig S4A ). As expected 18 19 20 , it was reduced by 35 ± 9% by the TRPC3s inhibitor Pyr3 (10 μM, n = 8, P < 0.01). The GluD2 blocker NASPM (100 μM) further decreased it to 78.3 ± 3% of the control (Fig 2 B). D‐serine (10 mM) also reduced this slow current by 58.5 ± 4.3% when considering its NASPM‐sensitive fraction ( n = 7, Fig 2 C). Thus, GluD2 channels likely contribute to the mGlu1‐dependent slow EPSC in Purkinje cells. If so, the dominant‐negative GluD2V617R mutant should reduce it. To test this, we transduced cerebella of WT mice with recombinant Sindbis virus carrying the sequences of GluD2V617R and the marker green fluorescent protein (GFP) (Fig 2 D). In all the Purkinje cells expressing GluD2V617R tested, the PF‐slow EPSC was reduced (87 ± 36 pA, n = 8, Fig 2 E) as compared to cells transfected with GFP only (265 ± 23 pA, n = 5; P < 0.01, Fig 2 E) or to those not transfected (281 ± 26 pA; n = 46; P < 0.01). In the GFP alone condition, the averaged trace displayed slower kinetics, two of the five cells recorded here having slow mGlu1 currents. However, this could not be attributed to the presence of GFP, but simply reflected a cell‐to‐cell variability that we observed in all the experiments of the study (see another example in supplementary Fig S4A ), whatever the genotype or manipulation. In our hands, a quarter of the Purkinje cells had such slow kinetics, however we can provide any explanation for this.

In cerebellar Purkinje cells, repetitive stimulations of PFs induce a mGlu1‐mediated slow excitatory post‐synaptic current (PF‐slow EPSC) carried by several conductances, notably TRPC3s 18 19 20 . As mGlu1s, TRPC3s and GluD2s associate and regulate the mGlu1‐dependent current at PF‐Purkinje cell synapses 13 26 27 , we tested whether GluD2 pore contributes to the mGlu1‐dependent PF‐slow EPSC.

To verify that it is carried by GluD2 but not by other conductances, we tested the mGlu1‐mediated slow current in cells transfected with a dominant‐negative GluD2 subunit. Introducing an arginine near the Q/R editing site of GluD2 disrupts the channel pore and turns it into a dominant‐negative subunit as for AMPA/KA‐Rs 12 24 . We generated the GluD2V617R subunits by replacing a valine at position 617 by an arginine in the GluD2 amino‐acid sequence (Fig 1 D). DHPG elicited no current in cells transfected with mGlu1 and GluD2V617R (Fig 1 D, n = 7). Thus, this current requires functional GluD2 channels. In cells transfected with mGlu1, wild‐type (WT) GluD2 and GluD2V617R, the DHPG current decreased as the quantity of GluD2V617R plasmids co‐transfected increases (Fig 1 E and F). It was almost abolished when WT and dominant‐negative GluD2s were transfected in equal proportions (Fig 1 E and F). Thus, GluD2 channels themselves carry the slow current triggered by mGlu1s. This supports the view that they are functional 5 25 , and designates mGlu1 as their physiological activator.

To examine the coupling between mGlu1 and GluD2 receptors, we first used HEK293 cells and the mGlu1alpha variant that we activated with 3,5‐dihydroxyphenylglycine (DHPG, 100 μM). Membrane expression of the various constructs was verified ( supplementary Figs S1–S3 ). In cells transfected with GluD2 or mGlu1 separately, DHPG did not induce any current (Fig 1 A). Conversely, in cells co‐transfected with mGlu1 and GluD2 receptors, DHPG induced a slow inward current (81 ± 12 pA; n = 27, Fig 1 A). This current relied on GluD2 as DHPG elicited no current in cells co‐transfected with mGlu1s and NMDA receptors (Fig 1 A). In cells co‐expressing mGlu1 and GluD2, the current‐voltage (I–V) relationship of the DHPG‐induced current was similar to that of the constitutively opened Lurcher‐ or chimeric‐GluD2s channels, with a characteristic inward rectification around + 20 mV 11 21 (Fig 1 B, n = 7). Two inhibitors of GluD2, the calcium‐permeable AMPA receptor (AMPA‐R) blocker NASPM (100 μM) and D‐serine (10 mM) 1 11 22 23 , both reduced the slow DHPG‐induced current (Fig 1 C), by 95 ± 8% ( n = 7, P < 0.01) and by 64 ± 15% ( n = 8, P < 0.02). On the contrary, the TRPC3 inhibitor Pyr3 did not change it ( n = 5, P = 0.31, Fig 1 C). These data indicate that mGlu1 triggers GluD2 opening, resulting in a slow current with biophysical and pharmacological features typical of GluD2 currents.

Discussion

Here, we demonstrate that mGlu1s activation triggers a current carried by GluD2 channels, showing for the first time that WT GluD2 have an ionotropic nature and that their gating can be, at least indirectly, triggered by glutamate.

An important consequence of this coupling is that GluD2s as well as TRPCs contribute to the mGlu1‐activated currents in Purkinje cells 2930. This unexpected finding may explain why several previous studies disagreed on the nature of mGlu1 currents, or more recently, on the effects of TRPCs inhibitors on them 14313233. The respective contribution of TRPC1/3 and GluD2 may depend on the experimental conditions and/or on the splicing variants of mGlu1, as these latter vary among cerebellar regions 34. These conditions remain to be clarified.

The presence of the dominant‐negative or the HO‐Nancy GluD2s could have changed the number of mGlu1s or TRPC3 1335 thereby explaining the decrease of the mGlu1 current. This is very unlikely. GluD2s do not seem to behave as scaffold or auxiliary proteins 313. Moreover, the existence of a GluD2‐dependent mGlu1 current in HEK293 cells that is inhibited by D‐serine, NASPM and GluD2V617R but not Pyr3 shows that the mGlu1 current flows through GluD2s, and not through some other interacting channel.

Even if the coupling between mGlu1 and GluD2 is reminiscent of other metabotropic‐ionotropic receptor crosstalks 1536, it appears unique in that mGlu1 triggers, rather than modulates, the gating of GluD2. However, none of our data suggest that it involves a direct activation of GluD2 by mGlu1. Intermediate steps cannot be excluded. Remarkably, the GluD2s gating mechanism supports the view that their LBD works differently from that of other iGluRs 112522. It also makes glutamate their indirect activator, which eventually brings these orphans back to their original family of ionotropic glutamate receptors. As such, GluD2s have some permeation, regulation and trafficking properties of AMPA/KA‐R, but they also require mGlu1 for their activation. Thus, GluD2 currents display chimeric properties that derive from both mGlu1 and AMPA/KA‐Rs.

Some of our results seem to contradict previous ones showing no reduction of mGlu1 currents and a redistribution of mGlu1s and TRPC3s in the absence of GluD2 13. However, these studies have been made in HO‐4j mice, where truncated GluD2s are retained in the endoplasmic reticulum, which is not the case for HO‐Nancy. Thus, the two models are different.

Some studies question the ionotropic nature of GluD2, based on the fact that PF LTD and the establishment of normal climbing fiber connection do not to depend on GluD2 channels 1237. However, this is not enough to refute the existence of GluD2 currents. Such currents could be necessary for other aspects of cerebellar physiology. As GluD2s are much less permeable to calcium than TRPC3 2138, the GluD2/TRPC3 ratio could set the permeability of the mGlu1 conductance to calcium which could, for example, determine the polarity (LTP/LTD) of synaptic plasticity.

In contrast to conventional fast AMPA‐Rs, the mGlu1/GluD2 duo converts high frequency input into a slow current. This duo is expressed in the vast majority of PF synapses 27 whereas more than 80% of these synapses are silent 39, and thus have no or few AMPA‐Rs. This suggests that most PF synapses act as a highpass filter and that Purkinje cells relay high versus low frequency PF inputs with very different dynamics.

The other delta family member GluD1 has 60% sequence homology with GluD2 40 and is similarly endowed with a functional channel pore domain 2541. We suggest that GluD1 might also be activated by metabotropic receptors, which would provide synapses with differential dynamical response to low and high frequency inputs.

Finally, the metabotropic‐dependent gating of GluDelta receptors enriches the computational repertoire of synapses. It makes these former orphans prodigal children of the glutamate receptor family.