Research ArticleNeuroscience

Hippocampal mGluR1-dependent long-term potentiation requires NAADP-mediated acidic store Ca2+ signaling

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Science Signaling  27 Nov 2018:
Vol. 11, Issue 558, eaat9093
DOI: 10.1126/scisignal.aat9093

Acidic Ca2+ stores in synaptic plasticity

Neurotransmitter signaling and neuronal Ca2+ fluxes are critical to learning and memory. Foster et al. found connections between the glutamate receptor mGluR1 and so-called “acidic Ca2+ stores” (Ca2+ stored in acidic organelles) in mouse hippocampal neurons (see the Focus by Patel and Brailoiu). Activation of mGluR1 induced the production of the molecule NAADP, which triggered a cascade of organellar ion channel–mediated Ca2+ release from relatively acidic endosomes and lysosomes to the endoplasmic reticulum. The resulting increase in intracellular Ca2+ depolarized the neurons by paradoxically inhibiting Ca2+-activated SK-type K+ channels, possibly through Ca2+-dependent activation of a phosphatase. These findings identify potential therapeutic targets for patients with neuronal disorders associated with mGluR1 and lysosomal dysfunction.


Acidic organelles, such as endosomes and lysosomes, store Ca2+ that is released in response to intracellular increases in the second messenger nicotinic acid adenine dinucleotide phosphate (NAADP). In neurons, NAADP and Ca2+ signaling contribute to synaptic plasticity, a process of activity-dependent long-term potentiation (LTP) [or, alternatively, long-term depression (LTD)] of synaptic strength and neuronal transmission that is critical for neuronal function and memory formation. We explored the function of and mechanisms regulating acidic Ca2+ store signaling in murine hippocampal neurons. We found that metabotropic glutamate receptor 1 (mGluR1) was coupled to NAADP signaling that elicited Ca2+ release from acidic stores. In turn, this released Ca2+-mediated mGluR1-dependent LTP by transiently inhibiting SK-type K+ channels, possibly through the activation of protein phosphatase 2A. Genetically removing two-pore channels (TPCs), which are endolysosomal-specific ion channels, switched the polarity of plasticity from LTP to LTD, indicating the importance of specific receptor store coupling and providing mechanistic insight into how mGluR1 can produce both synaptic potentiation and synaptic depression.


Acidic storage vesicles, such as lysosomes and endosomes, have traditionally been viewed as compartments for degradation and recycling of cellular metabolites. However, they have also been recognized as having additional signaling roles, including in intracellular Ca2+ signaling, as through their release of Ca2+ in response to the second messenger nicotinic acid adenine dinucleotide phosphate (NAADP) (16). At present, the functions of NAADP signaling in the central nervous system (CNS) are incompletely understood. NAADP evokes Ca2+ release from brain microsomes (7), and NAADP-binding sites are present throughout the brain (8), suggesting that NAADP is an important signaling molecule in the CNS. Thus far, several key areas in which NAADP signaling plays a role have been identified: (i) in differentiation and growth of (911) and augmentation of neurite outgrowth of cortical neurons (12); (ii) in autophagy in astrocytes in which NAADP promotes autophagosome formation, which can be reduced in the presence of a dominant-negative two-pore channel 2 (TPC2), a lysosomal two-pore channel isoform targeted by NAADP (13) or with the NAADP antagonist Ned-19 (14); (iii) and in regulating neuronal excitability, neurotransmitter release, and synaptic plasticity. The application of membrane-permeable NAADP (NAADP-AM) reportedly causes membrane depolarization in medullary neurons (15), and application of NAADP enhances neurotransmitter release in Aplysia californica (16) and at neuromuscular junctions in frogs (17). Work from our laboratory has shown that acidic store Ca2+ signaling enhances neurotransmitter release (18) and is essential for dendritic spine growth associated with late-phase long-term potentiation (LTP) (19) in hippocampal CA1 pyramidal neurons from rodents.

The activation of various metabotropic receptors reportedly induces NAADP synthesis in non-neuronal cell types (2024), and the application of glutamate, which can activate metabotropic glutamate receptors (mGluRs), also reportedly stimulates NAADP synthesis in both neurons (25) and astrocytes (14). However, the specific glutamate receptor subtype(s) responsible for NAADP synthesis has thus far remained elusive. Here, we examined NAADP-mediated changes in neuronal excitability and found that these NAADP-mediated effects are driven by mGluR1 activation. Critically, this pathway led us to identify a unique role for NAADP in the genesis of mGluR1-mediated synaptic potentiation.


NAADP-AM causes membrane depolarization in pyramidal neurons of the hippocampus

NAADP is a charged molecule, making it necessary to bond it to an acetoxymethyl (AM) ester group for it to be membrane permeable (26). To explore the effect of NAADP on membrane excitability, we introduced this engineered molecule (NAADP-AM) extracellularly, via a glass pipette, onto pyramidal neurons in CA3 and CA1 regions of hippocampal slice cultures. We chose cells that were localized to the surface of the tissue to improve the delivery of NAADP-AM.

Extracellular application of NAADP-AM (pipette concentration of 300 μM) produced transient membrane depolarizations (denoted by ΔVM; Fig. 1, B to D) in both CA3 and CA1 pyramidal neurons of the hippocampus. The effect was highly consistent because the application of NAADP-AM at 30-s intervals produced reproducible membrane depolarization. Application of vehicle alone or NAADP alone, in other words the “no AM” group, failed to produce depolarization (Fig. 1, B and C).

Fig. 1 NAADP causes membrane depolarization in pyramidal neurons of the hippocampus in a manner dependent on acidic store signaling, intracellular Ca2+, and RyR.

(A) Diagram showing the experimental configuration to record membrane potential of CA1 or CA3 pyramidal neurons in hippocampal slices while NAADP-AM was applied locally. (B) Example voltage traces recorded while applying NAADP-AM, NAADP, or vehicle. Arrowheads indicate the start of delivery, and gray bar indicates the total time of application. (C) Transient membrane depolarization (ΔVM) upon application of NAADP-AM (300 μM; n = 12 cells), NAADP (300 μM; n = 5), or vehicle (n = 6). Data are means ± SEM. (D) Mean ΔVM upon application of NAADP-AM (300 μM; n = 11 cells) alone or (left to right) in combination with a desensitizing concentration of NAADP (1 mM) inside the internal solution of the patch pipette (n = 6) after preincubation with the NAADP antagonist Ned-19 (100 μM, 40 min; n = 6) and preincubation with the vacuolar H+-ATPase inhibitor bafilomycin (4 μM, 40 min; n = 5), and with BAPTA (15 mM) inside the internal solution of the patch pipette (n = 5) after preincubation with ryanodine (30 μM, 40 min; n = 4). Significance was assessed with Kruskal-Wallis and post hoc Dunn’s tests. Data are means ± SEM. n = single cells. ***P < 0.005 by Kruskal-Wallis and post hoc Dunn’s tests.

To then examine the mechanism by which NAADP-AM produced the membrane depolarization, we performed a series of pharmacological manipulations in combination with the extracellular application of NAADP-AM. A characteristic feature of NAADP signaling in mammals is that at high concentrations, NAADP itself can desensitize the NAADP receptor (27). This feature allows for a specific pharmacological inhibition of NAADP signaling. Desensitization of the NAADP receptor with 1 mM NAADP in the patch pipette abolished responses to NAADP-AM (Fig. 1D), as did pharmacological antagonism of the NAADP receptor with Ned-19 (Fig. 1D) (28). These data show that NAADP-mediated depolarization depends on canonical mechanisms of NAADP signaling.

We next examined whether an NAADP-mediated increase in intracellular Ca2+ concentration was required for the depolarization we had observed. By introducing the fast Ca2+ chelator BAPTA [1,2-bis (o-aminophenoxy) ethane-N,N,N′,N′-tetraacetic acid] via the patch pipette (15 mM), we clamped intracellular Ca2+, and depolarization by NAADP-AM was inhibited (Fig. 1D). Whereas it was now clear that depolarization was dependent both on Ca2+ and NAADP, the intracellular source of the Ca2+ required confirmation. NAADP reportedly causes Ca2+ release from acidic stores (29, 30). Bafilomycin A1 is reported to abrogate endolysosomal Ca2+ release by inhibiting vacuolar H+–adenosine triphosphatases (ATPases) and thereby preventing Ca2+ loading into the store (31, 32). Preincubation with bafilomycin A1 abolished depolarization by NAADP-AM (Fig. 1D), suggesting that NAADP-mediated membrane depolarization requires Ca2+ release from an acidic store.

Amplification of acidic store Ca2+ signals by Ca2+-induced Ca2+ release (CICR) from ryanodine receptors (RyRs) has been reported in neurons of the medulla (15); therefore, we examined whether this signaling motif was conserved in pyramidal neurons. Ryanodine inhibits RyRs in the hippocampus at micromolar concentrations (33). Bath application of ryanodine at 30 μM inhibited NAADP-mediated membrane depolarization (Fig. 1D), suggesting that the CICR amplification motif is conserved.

Ca2+-mobilizing second messengers are not generally associated with driving membrane depolarization; thus, we explored whether NAADP is unique in its ability to do so. We dialyzed the known Ca2+-mobilizing second messengers [NAADP, inositol 1,4,5-trisphosphate (IP3), and cyclic adenosine diphosphate ribose (cADPR)] into CA1 pyramidal neurons via a patch pipette and measured membrane potential. We found that only when NAADP was included in the pipette was membrane depolarization observed (Fig. 2, A and B). Critically, the magnitude of depolarization generated in response to dialysis with NAADP produced a “bell-shaped” distribution, a hallmark of NAADP signaling where supramaximal concentrations of NAADP cause inhibition of the associated response (27, 34, 35). The median effective concentration was determined to be 12.5 nM, and the median inhibitory concentration was 7.07 μM. The Hill coefficients were +0.527 for stimulation and −0.927 for inhibition. Furthermore, the maximal ΔVM achieved was at the concentration of 100 nM NAADP, which is within the range previously reported for maximum Ca2+ mobilization from an acidic store (4, 36). Neither IP3 nor cADPR, at any concentration, produced significant depolarization (Fig. 2B). These results suggest that NAADP is unique in its capacity to produce a membrane depolarization among Ca2+-mobilizing second messenger family.

Fig. 2 NAADP is unique among second messengers in its ability to depolarize hippocampal pyramidal neurons.

(A) Example voltage traces for dialysis of CA1 pyramidal neurons patched with internal solutions containing various concentrations of the Ca2+-mobilizing second messengers NAADP, IP3, and cADPR. Changes to membrane potential were recorded over time as the second messengers dialyzed into the patched cell. (B) Transient membrane depolarization (ΔVM) of the cells described in (A) in response to increasing concentrations of the Ca2+-mobilizing second messengers. Data are means ± SEM. n = single cells, indicated above each column; n > 4 for all concentrations of second messengers. ***P < 0.005 and *P <0.05 by Kruskal-Wallis and post hoc Dunn’s tests.

Activation of mGluR1 causes NAADP-dependent membrane depolarization

In order that we might begin to understand the importance of the NAADP-mediated depolarization, we sought to identify the mechanism by which NAADP elevations are triggered physiologically. The excitatory neurotransmitter glutamate has previously been shown to generate the synthesis of NAADP in neurons and astrocytes [Pandey et al. (25) and Pereira et al. (14)], although the specific glutamate receptor subtype(s) were not identified. In a variety of other tissue types, NAADP synthesis is reported to occur after the activation of metabotropic receptors (6, 2023), we therefore looked to mGluRs as likely candidates. To examine this, we pharmacologically isolated the mGluRs with antagonists of ionotropic glutamate and γ-aminobutyric acid receptors [50 μM AP5, 10 μM NBQX (2,3-dihydroxy-6-nitro-7-sulfamoylbenzo[f]quinoxaline), 100 μM picrotoxin, and 2 μM CGP 55845] and delivered patterned electrical stimulation (four pulses, 20 Hz), known to activate mGluRs in the hippocampus (37), while measuring the membrane potential from CA1 pyramidal neurons (Fig. 3A). Under these conditions, we observe a depolarization ΔVm of +2.09 ± 0.15 mV (Fig. 3, B and C). To confirm that this was a consequence of mGluR activation, the mGluR antagonist LY341495 (100 μM) was added to the bath at a concentration reported to block all members of the mGluR family (38). This blocked the electrically induced membrane depolarization (Fig. 3, B and C). Now confident that mGluRs are able to produce depolarization, we sought to determine the specific mGluR subtype(s) that mediated the response.

Fig. 3 Activation of mGluR1 in CA1 pyramidal neurons causes a membrane depolarization that depends on NAADP signaling and acidic store Ca2+ signaling.

(A) Diagram showing the experimental configuration to record membrane potential of CA1 pyramidal neurons in hippocampal slices while mGluRs were pharmacologically isolated (50 μM AP5, 10 μM NBQX, 100 μM picrotoxin, and 2 μM CGP 55845) and electrical stimulation was applied to Schaffer collaterals (four pulses, 20 Hz) (n = 11 cells). (B) Typical voltage recordings from single cells upon electrical stimulation with pharmacological isolation of mGluRs or plus antagonism of group II and III mGluRs (100 nM LY341485; n = 6), pan-mGluRs (100 μM LY341485; n = 6), mGluR5 (10 μM MPEP; n = 5), or mGluR1 (300 nM JNJ16259685; n = 5) (top to bottom). The red lines indicate where membrane potentials were compared before and after stimulation. (C and D) Transient membrane depolarization (ΔVM) of CA1 pyramidal neurons after electrical stimulation alone (control; n = 7 cells) or with the presence of (C) the mGluR antagonism described in (B) or (D) pan-mGluR antagonist [100 μM LY341485; n = 6; cells and dataset are independent from those in (C)] with a desensitizing concentration of NAADP inside the internal solution of the patch pipette (1 mM; n = 6) and NAAD inside the internal solution of the patch pipette (1 mM; n = 6), preincubation with the NAADP antagonist Ned-19 (100 μM, 40 min; n = 6), or acute administration of the lysosomal disrupting agent GPN (200 μM; n = 5). Data are means ± SEM. n = single cells. **P < 0.01 and *P < 0.05 by Kruskal-Wallis and post hoc Dunn’s tests.

The mGluR family contains eight members divided into three groups based on their pharmacological and functional profiles: group I (mGluR1 and mGluR5), group II (mGluR2 and mGluR3), and group III (mGluR4, mGluR6, mGluR7, and mGluR8) (39, 40). We systematically isolated members of the mGluR family using a series of mGluR subgroup- and/or subtype-specific pharmacological antagonists. LY341495 acts as an mGluR group II and III antagonist at 100 nM, whereas at 100 μM it acts as a pan-mGluR antagonist (38). The addition of 100 nM LY341495 produced no significant reduction in the depolarization observed upon electrical stimulation), whereas 100 μM LY341495 blocked the membrane depolarization (Fig. 3C). Thus, the depolarization is a group I mGluR-dependent effect. Group I contains two mGluR subtypes, mGluR1 and mGluR5. MPEP [2-methyl-6-(phenylethynyl)pyridine], a selective mGluR5 antagonist (41), did not block the depolarization, whereas JNJ16259685, a selective mGluR1 antagonist (42), abolished the depolarization (Fig. 3C). On the basis of the pharmacological dissection of the response, we conclude that of the mGluR family of receptors, only mGluR1 activation produces membrane depolarization in CA1 pyramidal neurons.

We next sought to examine a link between the mGluR1-mediated membrane depolarization, NAADP signaling, and acidic store Ca2+ release. We examined their relationship by isolating the mGluR response as described above before the introduction of further manipulations. Desensitization of the NAADP receptor with NAADP (1 mM) inside patch pipette abolished mGluR-mediated depolarization, whereas 1 mM nicotinic acid adenosine dinucleotide (NAAD), an inactive metabolite, had no effect (Fig. 3D), suggesting that the action of NAADP is specific. Preincubation with the NAADP receptor antagonist Ned-19 also significantly reduced responses to mGluR depolarization, and disruption of the lysosomes with acute application of glycyl-l-phenylalanine 2-naphthylamide (GPN) abolished mGluR-mediated depolarization (Fig. 3D). GPN prevents acidic store Ca2+ signaling without compromising cell health (19); thus, together, these data suggest that mGluR membrane depolarization is dependent on NAADP and Ca2+ release from acidic stores.

mGluR1-dependent membrane depolarization does not require dendritic IP3 receptors

Our data suggest that NAADP and Ca2+ release from acidic stores drives the membrane depolarization. It therefore seems reasonable that Ca2+ signaling via the acidic store is intimately linked to this process. Several studies report an amplification of acidic store Ca2+ by the endoplasmic reticulum (ER), and we show that the RyRs are essential for NAADP-mediated depolarization (Fig. 1D). However, we are also mindful that group I mGluRs are thought to be involved in IP3 receptor (IP3R)–mediated Ca2+ release (40); therefore, we needed to explore this relationship in greater detail to better understand these Ca2+ signaling events.

First, we confirmed the requirement of the acidic Ca2+ stores for mGluR1-mediated depolarization by pharmacologically isolating mGluR1s (as previously described) and recording membrane potential while glutamate was bath-applied (Fig. 4, A and B). As expected, acute application of GPN abolished mGluR1-mediated depolarization (Fig. 4C). We next wanted to determine whether the mGluR1-mediated depolarization was also dependent on Ca2+ release from the ER. Therefore, we examined whether RyRs or IP3Rs are essential for mGluR1-mediated membrane depolarization. Acute application of desensitizing concentrations of ryanodine prevented mGluR1-mediated depolarization (Fig. 4C). In contrast, xestospongin C, a potent and selective inhibitor of IP3Rs (43), had no effect on the mGluR1-mediated depolarization (Fig. 4C).

Fig. 4 In CA1 pyramidal neurons, mGluR1-dependent membrane depolarization and Ca2+ release require acidic store signaling and Ca2+ release from the ER via RyRs but not IP3Rs.

(A) Diagram showing the experimental configuration. The membrane potential of CA1 pyramidal neurons in hippocampal slices was recorded while mGluR1 was pharmacologically isolated and extracellular glutamate was applied. (B) Typical voltage recordings recorded upon bath application of glutamate (300 μM, 120 s) or the vehicle. (C) Columns show mean ΔVM of CA1 pyramidal neurons before and after extracellular glutamate application under control conditions (n = 8) and in the presence of the lysosomal disrupting agent GPN (200 μM; n = 6), RyR antagonist ryanodine (40 μM, 15 min; n = 6), IP3R antagonist xestospongin C (2 μM, 15 min; n = 6), “fast” Ca2+ chelator BAPTA (20 μM, 15 min; n = 6), or the “slow” Ca2+ chelator EGTA (20 μM, 15 min; n = 6). (D) Time-series images of CA1 neurons filled with Ca2+ indicator OGB-1 (1 mM) were recorded while mGluR1 was pharmacologically isolated (50 μM AP5, 10 μM NBQX, 100 μM picrotoxin, and 2 μM CGP 55845) and electrical stimulation was applied (four pulses, 20 Hz). Images (from top to bottom) of z stack of the dendritic branch being imaged (green), Ca2+ signal at baseline before stimulation, Ca2+ signal 300 ms after stimulation, and subtraction of Ca2+ at 300 ms from baseline (purple). Scale bar, 0.5 μm. (E) ΔF/F over the imaging time course where mGluR1 was pharmacologically isolated (n = 20 cells) in combination with acute application of LY341495 (100 μM, 10 min; n = 5), preincubation with Ned-19 (100 μM, 1 hour; n = 6), or acute application of ryanodine (20 μM, 10 min; n = 5), xestospongin C (2 μM, 15 min; n = 5), or 2-aminoethoxydiphenyl borate (2-APB) (50 μM, 15 min; n = 5). (F) Columns show mean ΔF/F before and after electrical stimulation for each pharmacological manipulation undertaken. Significance was assessed with Kruskal-Wallis and post hoc Dunn’s tests. Error bars denote SEM. n = single cell. Significant differences indicated by asterisks where ***P < 0.005 and *P < 0.05.

Membrane contact sites between the acidic Ca2+ stores and the ER are reported and suggested to mediate microdomain signaling between these two organelles (44). To assess whether microdomain Ca2+ signaling is likely to occur with mGluR1-mediated membrane depolarization, we compared the action of two Ca2+ chelators: EGTA and BAPTA. EGTA has an “on” rate for binding Ca2+ of 3 to 10 μM−1 s−1, whereas BAPTA is several orders of magnitude faster at 100 to 1000 μM−1 s−1 (45, 46). Consequently, EGTA is thought not to chelate Ca2+ at a rate fast enough to inhibit microdomain Ca2+ signaling, whereas BAPTA can prevent all except elementary Ca2+ signaling events (47, 48). Bath application of membrane-permeable EGTA (EGTA-AM) did not affect mGluR1-mediated cellular depolarization (Fig. 4C), whereas membrane-permeable BAPTA (BAPTA-AM) prevented mGluR1-mediated depolarization (Fig. 4C). These data suggest that mGlurR1-mediated depolarization is dependent on microdomain Ca2+ signaling.

Dendritic mGluR1-dependent Ca2+ transients require NAADP signaling and RyRs but not IP3Rs

To assess more directly the intracellular Ca2+ signaling events initiated by mGluR1 activation, we used confocal microscopy to visualize Ca2+ signals in the dendrites of CA1 pyramidal neurons. Neurons were loaded with the Ca2+-sensitive dye Oregon Green 488 BAPTA-1 (OGB-1), and mGluR1s were pharmacologically isolated as described above.

Electrical stimulation was applied to generate dendritic Ca2+ signals. These Ca2+ signals could be blocked by pharmacologically inhibiting either the NAADP receptor with Ned-19 (Fig. 4, D to F) or all members of the mGluR family with LY341495 (Fig. 4, E and F), again indicating that mGluR1-mediated Ca2+ signals depend on NAADP signaling. We found that ryanodine abolished mGluR1-dependent Ca2+ signals (Fig. 4, E and F), whereas addition of the IP3R antagonists xestospongin C or 2-APB did not reduce mGluR1-mediated Ca2+ signaling (Fig. 4, E and F). Together, these data suggest that mGluR1-mediated Ca2+ signals are dependent on Ca2+ release from both acidic Ca2+ stores and from the ER via RyR but not IP3Rs. As a positive control, we confirmed that xestospongin C inhibited IP3R-mediated Ca2+ release by visualizing Ca2+ signals in CA1 pyramidal neurons patch-clamped with an internal solution containing IP3 under control conditions and after preexposure to xestospongin C (fig. S1, A to C).

mGluR1-dependent depolarization is mediated by the inactivation of small conductance calcium-activated potassium (SK) channels after dephosphorylation

We wished to understand the biophysical basis of the mGluR1-mediated membrane depolarization. A number of studies suggest that group I mGluRs inhibit K+ channels in CA1 pyramidal neurons of the hippocampus (41, 4956), with Tigaret et al. (57) providing evidence that mGluR1 activation inhibits SK channels, an important step for the induction of mGluR1-mediated LTP in the hippocampus.

SK channels are present in the dendrites of CA1 pyramidal neurons of the hippocampus (58), where their activation produces action potential after-hyperpolarization currents (IAHP). SK channels have also been implicated in regulating dendritic excitability (57, 59, 60).

We sought to determine whether the allosteric inhibitor of SK channels, apamin (61), inhibited mGluR1-mediated depolarization. mGluR1s were pharmacologically isolated (as described above), and membrane potential was recorded from CA1 pyramidal neurons while a patterned of electrical stimulation was delivered. The experiment reveals that synaptic activation of mGluR1 produced membrane depolarization (Fig. 5, A and B), and apamin significantly reduced the amplitude of depolarization (Fig. 5B). GPN was then introduced to determine whether a common signaling pathway was being used. GPN was found to have no further effect on reducing the mGluR1-mediated depolarization (Fig. 5B). These data suggest that inhibition of the SK channels by acidic store Ca2+ is the key intermediate step in producing mGluR1-mediated depolarization.

Fig. 5 In CA1 pyramidal neurons, mGluR1-dependent depolarization occurs through the inactivation of SK channels by possibly PP2A.

(A and B) Representative voltage recordings (A) and mean ΔVM (B) upon electrical stimulation (four pulses, 20 Hz) of CA1 neurons while mGluR1 was pharmacologically isolated, then subsequent addition of apamin (200 nM, 15 min) and, last, GPN (200 μM, 10 min; n = 6 cells). (C and D) Representative voltage recordings (C) and mean Δ VM (D) upon bath application of glutamate (red arrowhead; 300 μM, 120 s) of CA1 neurons while mGluR1 was pharmacologically isolated, then subsequent addition of apamin (200 nM, 15 min) and, last, GPN (200 μM, 10 min; n = 6 cells). (E and F) Representative voltage recordings (E) and mean ΔVM (F) upon electrical stimulation (four pulses, 20 Hz) of CA1 neurons while mGluR1 was pharmacologically isolated in the absence or presence of okadaic acid (100 nM, 15 min, n = 6 cells). Data are means ± SEM, each from n = 6 single cells. *P < 0.05 (relative to mGluR1 isolation-alone condition) by Freidman’s test with post hoc Dunn’s tests (B and D) or a Wilcoxson pair-matched signed-rank test (F). n.s., no significant difference.

To ensure that the addition of apamin and/or GPN had not interfered with presynaptic glutamate release, we repeated this experiment with transient bath application of l-glutamate rather than electrical stimulation. The mGluR1s were again pharmacologically isolated, and neurotransmission was prevented with tetrodotoxin (TTX). Extracellular glutamate caused membrane depolarization (Fig. 5C) and was significantly reduced by the addition of apamin (Fig. 5D). Again, we found that the application of GPN had no further effect on reducing the amount of mGluR1 depolarization after application of apamin (Fig. 5D). Apamin did not affect the resting membrane potential of CA1 neurons of the hippocampus (fig. S2, A and B).

SK channels are subject to modulation. Multiple sites for phosphorylation have been reported (62), with SK channels shown to form macromolecular complexes with protein kinase 2 and protein phosphatase 2A (PP2A) (63, 64). Some members of the PP2A family are Ca2+ sensitive (65), and the presence of Ca2+-binding EF hand domains is noted on regulatory B-type subunits (66). We therefore used the inhibitor of PP2A (okadaic acid) to determine whether SK channel modulation could affect mGluR1-mediated depolarization. We found that acute application of okadaic acid significantly reduced mGluR1-mediated depolarization (Fig. 5, E and F). Therefore, we suggest that activation of mGluR1 and the subsequent Ca2+ signals evoked are key in activating one or more members of the PP2A family.

Last, it has also been suggested that mGluR1-mediated depolarization may occur via activation of a nonselective cation or transient receptor potential (TRP) channel (67, 68). To assess this possibility, we used the broad-spectrum antagonists of nonselective cation channels, flufenamic acid and La3+, to determine whether mGluR1-mediated depolarization could be achieved via these channels. We found that neither had any effect on mGluR1-mediated depolarization (fig. S3, A to C).

mGluR1-dependent plasticity requires inhibition of SK channels by acidic store Ca2+

Several studies suggest that hippocampal LTP can occur after the activation of mGluR1 (57, 6972). We wished to determine whether the signaling pathway that we describe is required for mGluR1-dependent plasticity. We therefore implemented a spike timing–dependent plasticity (STDP) protocol known to produce mGluR1-dependent plasticity (57).

The STDP induction protocol produced LTP of about 150% (Fig. 6A) in CA1 pyramidal neurons. To ensure that the protocol induced mGluR1-dependent LTP, we repeated the experiment in the presence of an mGluR1-specific antagonist (JNJ16259685), which blocked the LTP (Fig. 6B).

Fig. 6 In CA1 pyramidal neurons, mGluR1-dependent synaptic plasticity requires inhibition of SK channels via NAADP signaling.

(A) A causal STDP protocol was used to induce mGluR1-dependent LTP, in which one causal presynaptic stimulation is paired with two backpropagating action potentials (bAPs) (100 Hz) at a 10-ms interval. The induction protocol is delivered, where t = 0, indicated by the black triangles. Example excitatory postsynaptic potential (EPSP) traces before (black) and after (red) STDP induction are shown at the top right of each graph. Scale bar, 5 mV by 50 ms. This STDP protocol produces LTP lasting at least 30 min (n = 7 cells). (B) LTP in the STDP protocol described in (A) with mGluR1-specific antagonism with JNJ16259685 (300 nM; n = 5 cells). (C) LTP as described in (A) upon prevention of NAADP/acidic store Ca2+ signaling with a desensitizing concentration of NAADP (5 mM; n = 5 cells). (D) Magnitude of LTP upon induction of STDP in the presence of SK channel antagonist apamin (200 nM; n = 7 cells). (E) LTP as described in (A) in the presence of apamin and JNJ16259685 (300 nM; n = 6 cells). (F) Mean change in synaptic strength at 25 to 30 min, expressed as a percentage of the baseline. Data are means ± SEM. n = single cell. *P < 0.05 by Kruskal-Wallis and post hoc Dunn’s tests.

Next, we selectively abolished NAADP signaling with a desensitizing concentration of NAADP (73, 74) in the patch pipette. We found that this manipulation prevented LTP (Fig. 6C), indicating that mGluR1-dependent LTP requires NAADP-mediated acidic store Ca2+ signaling.

Because the inhibition of SK channels via mGluR1 is thought to be required for mGluR1-mediated LTP, pharmacological inactivation of SK channels with apamin should have no impact on the LTP; SK channels should be inhibited by mGluR1 activation, thereby occluding the apamin’s action. We found that apamin alone had no effect on the magnitude of LTP compared to control experiments (Fig. 6D). Critically, we also found that antagonism of mGluR1s with JNJ16259685, while simultaneously inhibiting SK channels with apamin, rescues the ability of the STDP protocol to produce mGluR1-dependent plasticity (Fig. 6E). These findings (summarized in Fig. 6F) indicate that activation of mGluR1 can produce LTP in the hippocampus and that LTP is achieved via the modulation of SK channel activity.

TPCs are essential for mGluR1-dependent membrane depolarization and LTP

We have shown that NAADP signaling is required for mGluR1-mediated depolarization and LTP. Here, we add to the pharmacological evidence by genetically manipulating the signaling pathway. Two-pore channels (TPC1 and TPC2) are localized to acidic Ca2+ stores in mammalian cells and have been shown to be essential for NAADP-mediated Ca2+ release (13, 35). First, we wanted to determine whether TPCs were required for NAADP-mediated events in our experimental preparation. Therefore, we dialyzed NAADP into CA1 pyramidal neurons via a patch pipette at the maximally effective concentration (determined in the data shown in Fig. 2B) while recording membrane potential. NAADP-mediated depolarization was reduced in neurons from Tpcn1−/− mice and abolished in neurons from Tpcn2−/− mice (fig. S4, A and B).

Next, we explored whether either TPC1 or TPC2 was required for mGluR1-mediated depolarization and mGluR1-dependent LTP. To achieve this, we first assessed whether mGluR1-dependent depolarization could be produced in CA1 neurons of wild-type (WT) mice. We pharmacologically isolated mGluR1s (as described above) and delivered electrical stimulation, confirming that mGluR1 activation produced depolarization (Fig. 7, A and B). In contrast, depolarization was not observed in CA1 pyramidal neurons of either Tpcn1−/− or Tpcn2−/− animals.

Fig. 7 In CA1 pyramidal neurons, TPCs are required for mGluR1-mediated membrane depolarization and mGluR1-dependent LTP.

(A) Representative average (five traces) voltage recordings from CA1 neurons in hippocampal slice preparations from WT, Tpcn1−/−, and Tpcn2−/− mice (n = 6 mice each). Recordings were obtained upon pharmacological isolation of mGluR1 (50 μM AP5, 10 μM NBQX, 100 nM LY341495, 10 μM MPEP, 100 μM picrotoxin, and 2 μM CGP 55845) and electrical stimulation (four pulses, 20 Hz) of afferent fibers in stratum radiatum. Solid red lines indicate where membrane potentials were compared before and after stimulation. (B) Columns show mean ΔVM of CA1 pyramidal neurons before and after electrical stimulation described in (A). (C) A causal STDP protocol was used to induce mGluR1-dependent LTP after a baseline of EPSPs were recorded for 5 min (indicated by marker at 0 min) in WT (n = 4), Tpcn1−/− (n = 5), and Tpcn2−/− (n = 7) animals. One casual presynaptic stimulation is paired with two bAPs (100 Hz) at a 10-ms interval. (D) Mean change in synaptic strength at 25 to 30 min shown/described in (C), expressed as a percentage of the baseline (red dashed line). Data are means ± SEM. n = single cell. **P < 0.01 and *P < 0.05 by Kruskal-Wallis and post hoc Dunn’s tests.

Fig. 8 Proposed model for mGluR1-dependent plasticity.

(A) Model of SK channel activation, wherein (i) synaptic glutamate activates GluA (AMPA) receptors to produce (ii) membrane depolarization and (iii) Ca2+ entry via VDCCs. This causes (iv) activation of SK channels and local hyperpolarization, resulting in inhibition of GluNs (NMDAs) by reinstating Mg2+ block, thereby reducing Ca2+ entry through the GluNs and reducing the probability of LTP induction. Where synaptic activity is sufficiently strong, the mGluR1 receptors are recruited. (B) The proposed model for SK channel inhibition mediated by mGluR1 signaling; GluA/VDCC regulation of SK channels is also present but not shown. (i) Glutamate activates mGluR1 receptors and causes (ii) NAADP synthesis, which results in (iii) acidic store Ca2+ release, which is amplified through activation of RyRs in the ER. This somehow inactivates SK channels (iv), which in turn prevents local hyperpolarization and (v) allows greater Ca2+ entry through the GluN receptors, which facilitates the induction of LTP.

Next, we sought to determine whether the TPCs were important for mGluR1-dependent LTP. We confirmed that it was observed in CA1 pyramidal neurons (Fig. 7, C and D). Upon assessing LTP in knockout mice, we were surprised to find that the STDP protocol that had generated LTP instead induced long-term depression (LTD) in both Tpcn1−/− and Tpcn2−/− animals (Fig. 7, C and D). It would therefore appear that the change in the Ca2+ signaling profile in these neurons resulted in a switch in the polarity of the plasticity for our STDP stimulus regime.


In this work, we show that the metabotropic receptor mGluR1 is specifically linked to the NAADP signaling cascade, a pathway that uniquely mobilizes the release of Ca2+ from acidic organelles such as lysosomes and late endosomes. We found that the activation of mGluR1 produced NAADP-dependent Ca2+ release and also membrane depolarization, both of which requiring Ca2+ amplification via the RyRs. Because we found that cADPR alone was not able to produce depolarization, we suggest that mGluR1/NAADP-mediated Ca2+ release from the acidic stores is the critical first step in this signaling pathway. The pathway is quite specific because we find that mGluR5, the second member of group I mGluR group, fails to trigger NAADP-mediated cellular depolarization. We reveal that this pathway is intimately linked to the induction of mGluR1-mediated LTP. The mechanism by which this occurs appears to be a transient modulation of SK channel function after dephosphorylation by PP2A.

In a previous study, we found that acidic stores were essential for the maintenance of spine growth after LTP induction (19). We now find that in addition to LTP maintenance, acidic stores play a role in the induction of mGluR1-dependent forms of LTP. Whether LTP requires mGluR1, however, depends on the parameters of stimulation, with stronger stimulation protocols capable of bypassing the requirement of mGluRs and, more generally, store Ca2+ release (7577).

Perhaps the least intuitive of our results is the observation that mGluR1 activation modulates SK channel function to produce depolarization of the membrane. Canonically, SK channels are thought to underlie AHPs. In this context, the channels are closed and become active as Ca2+ enters neurons after activity. However, it would appear that the regulation of SK channels by Ca2+ is more complex than once thought, with SK channels differentially regulated by Ca2+ from different sources. Ca2+ that enters via R-type voltage-dependent Ca2+ channels (VDCCs) or N-methyl-d-aspartate receptors (NMDARs) acts to potentiate SK channel opening, causing hyperpolarization. In this context, hyperpolarization helps reinstate the Mg2+ block of NMDARs, reducing the amount of Ca2+ entry and consequently reducing the magnitude of LTP (78, 79). Conversely, we find that Ca2+ release from acidic stores reduces SK channel activity. We do not fully describe the mechanism as to how this occurs; however, we show that PP2A is required to produce mGluR1-mediated depolarization. PP2As form complexes with SK channels and are only active while the channels are in an open state; the result of their activity is to increase the sensitivity of SK channels to Ca2+ (63, 64). Perhaps this increased sensitivity allows detection of acidic store Ca2+ and, in turn, reduce SK channel activity. Further work needs to be undertaken to elucidate this signaling mechanism. In addition, we found that apamin had no effect on resting membrane potential of hippocampal neurons. This is consistent with previous findings (8083) and might be explained by apamin’s allosteric mode of action (61), which perhaps locks SK channels in their current state and thus prevents them from being activated by any source of Ca2+.

Nevertheless, the reduction in SK channel activity is a key step in the induction of metabotropic receptor–dependent forms of LTP (57, 84). In good agreement with Tigaret et al. (57), we confirm that inhibition of SK channels is essential for mGluR1-mediated LTP, but critically we identify the intermediate step and link mGluR1-mediated depolarization to the NAADP signaling pathway (Fig. 8). Underpinning our link to this pathway are the data showing that genetic knockout of either Tpcn1 or Tpcn2 removes both mGluR1-mediated depolarization and mGluR1-mediated LTP. The importance of both TPC1 and TPC2 for mGluR1-mediated LTP may indicate that mGluR1 is interacting with heteromeric complexes of TPC1 and TPC2, an arrangement reported in other cell types (85).

Roles for mGluR1 in both LTD (37, 71, 8688) and LTP (69, 8991) have been reported at CA3-CA1 synapses. Whether LTD or LTP is observed seems to be dependent on the stimulation pattern delivered to the neurons, with stronger stimulation regimes generating LTP and weaker ones LTD (71, 75, 92). mGluR1 can produce both hyperpolarization and depolarization in dopaminergic neurons, with strong stimulation causing depolarization and weak stimulation causing hyperpolarization (93). One clear illustration of the importance of recruiting specific signaling pathways to generate a specific plasticity outcome was seen in our data that showed that when TPCs are genetically removed, our LTP induction protocol induced LTD, not LTP. Collectively, these data suggest that at least two intracellular signaling pathways can be stimulated by mGluR1 activation and that the pathway engaged, and the consequent polarity of plasticity, is dependent on the stimulation pattern.

The importance of understanding the different mGluR1 signaling pathways is clear when interpreting work in vivo. mGluR1 antagonists impair spatial learning (94), and the deletion of the gene encoding mGluR1 reduces LTP in CA1 region of the hippocampus and impairs context-specific associative learning (95) and spatial memory (96). Impaired group I mGluR signaling is also implicated in the pathogenesis of Fragile X syndrome (97, 98) as yet an untreatable disorder; thus, the signaling pathway that we reveal offers new opportunities toward understanding diverse behavioral phenotypes and routes for novel therapeutic intervention.


Hippocampal slice preparation

All animal work was carried out in accordance with the Animals (Scientific Procedures) Act 1986 (UK) and under the project and personal licenses approved by the Home Office (UK). Slice cultures of the hippocampus were prepared from male Wistar rats (P6 to P8) or WT, Tpcn1−/−, or Tpcn2−/− mice (35). The hippocampi were isolated in ice-cold Earle’s balanced salt solution with added 21 mM Hepes and 27.8 mM D-glucose (pH adjusted to 7.2 to 7.4 with NaOH) and cut into slices of 350-μm thickness with a McIlwain tissue chopper. Slices were placed into Millicell CM culture plate inserts (polytetrafluoroethylene filter; pore size, 0.4 μm; diameter, 12 mm) in a six-well Millicell culture plate (both supplied by Merck Millipore) with 1 ml of culture medium and stored at 34.5°C at 5% CO2. Culture medium was composed of 78.8% minimum essential medium with GlutaMAX (Gibco), 20% heat-inactivated horse serum, 1% B27 with added 1 mM CaCl2, 30 mM Hepes, 26 mM D-glucose, 5.8 mM NaHCO3, and 2 mM MgSO4. Culture media were renewed every 3 to 4 days.

During experiments, slices [10 to 14 days in vitro (DIV)] were perfused (1 to 2 ml/min) with heated (32° to 34°C) artificial cerebrospinal fluid, which is composed of 145 mM NaCl, 2.5 mM KCl, 1.2 mM KH2PO4, 16.0 mM NaHCO3, 11.0 mM glucose, 3.0 mM CaCl2, and 2.0 mM MgCl2 aerated with 95% O2 and 5% CO2.


Whole-cell patch clamp recordings were performed on pyramidal neurons from either CA3 or CA1 neurons of hippocampal slice cultures. To minimize intracellular dialysis, high-resistance patch electrodes (16 to 20 megohms) were used, where mGluR1 was isolated pharmacologically and all other experiments were undertaken with lower-resistance electrodes (5 to 8 megohms). Voltage signals were detected using an Axoclamp 2B (Axon Instruments/Molecular Devices) amplifier, signals were digitized using a Digidata 1440A and then recorded digitally with WinWCP V4.7.9 (Strathclyde Electrophysiology Software), and 50-Hz noise was eliminated with a Hum Bug (Quest Scientific). The internal solution contained (135 mM K-gluconate, 10 mM KCl, 10 mM Hepes, 2 mM MgCl2, 2 mM Na2–adenosine triphosphate, and 0.4 mM Na3–guanosine triphosphate; pH 7.2 to 7.4). Pharmacological agents/second messengers added to the internal solution were conjugated to K+ salts, and when present, the K-gluconate concentration was reduced by equal molarity to maintain osmolarity. Electrical stimulation was applied using a tungsten stimulating electrode with an isolated constant current stimulator (Digitimer Ltd.). To produce mGluR1-mediated depolarization, the stimulation pattern used consisted of four pulses at 20 Hz. The mGluR1 LTP protocol consisted of causally pairing one presynaptic stimulus (to produce a subthreshold EPSP) with two bAPs (100 Hz) elicited in the postsynaptic neuron via current injection at a 10-ms interval. This paired induction protocol was repeated 300 times at 5 Hz and delivered within 5 min of whole-cell breakthrough to prevent dialysis of factors required for mGluR1-dependent LTP.

Ca2+ imaging

Pyramidal neurons in the CA1 region of hippocampal slice cultures, 10 to 14 DIV, were filled with OGB-1 via a patch clamp in whole-cell configuration for 1 min. The patch electrode’s internal solution consisted of 135 mM K-gluconate, 10 mM KCl, 10 mM Hepes, 1 mM MgCl2, 1 mM OGB-1, and 10 mM QX314. Apical dendrites were imaged using confocal laser scanning microscopy and a 488-nm argon laser, whereas an isolated constant current stimulator was used to deliver electrical stimulation (four pulses, 20 Hz) presynaptic neurons.


NAADP-AM and NAADP were synthesized in-house (26, 99). Other drugs were purchased from the following suppliers: Abcam (LY341495, CGP 55845, D-AP5, Ned-19, JNJ16259685, MPEP, QX314, and TTX), Sigma-Aldrich (ryanodine, bafilomycin A1, picrotoxin, NBQX, NAAD and BAPTA, BAPTA-AM, and EGTA-AM), Santa Cruz Biotechnology (GPN), Thermo Fisher Scientific (OBG-1), and Tocris (U73122 and U73343).


Fig. S1. Xestospongin C inhibits somatic IP3-mediated Ca2+ release in CA1 pyramidal neurons in the hippocampus.

Fig. S2. Apamin does not affect resting membrane potential of CA1 pyramidal neurons.

Fig. S3. mGluR1-mediated depolarization unlikely to occur via TRP channel activation.

Fig. S4. NAADP-mediated membrane depolarization in CA1 pyramidal neurons requires Tpc1 and Tpc2.


Acknowledgments: We thank G. Churchill and C. Garnham for synthesizing and providing us the NAADP-AM. Funding: This work was funded by the Wellcome Trust Senior Investigator Enhancement Grant, “A messenger role for NAADP in the central nervous system” to A.G., by an Alison Brading Scholarship from Lady Margaret Hall, Oxford to W.J.F., and by a grant from the BBSRC to N.J.E. Author contributions: W.J.F. and H.B.C.T. performed and analyzed the experiments. W.J.F., Z.P., and A.F.J. designed the experiments. W.J.F. and N.J.E. wrote the manuscript. A.G. and N.J.E. supervised the project. Competing interests: The authors declare that they have no competing interests. Data and materials availability: All data needed to evaluate the conclusions in the paper are present in the paper or the Supplementary Materials.
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