Metabotropic Acetylcholine and Glutamate Receptors Mediate PI(4,5)P₂ Depletion and Oscillations in Hippocampal CA1 Pyramidal Neurons in situ

Abstract

The sensitivity of many ion channels to phosphatidylinositol-4,5-bisphosphate (PIP₂) levels in the cell membrane suggests that PIP₂ fluctuations are important and general signals modulating neuronal excitability. Yet the PIP₂ dynamics of central neurons in their native environment remained largely unexplored. Here, we examined the behavior of PIP₂ concentrations in response to activation of Gq-coupled neurotransmitter receptors in rat CA1 hippocampal neurons in situ in acute brain slices. Confocal microscopy of the PIP₂-selective molecular sensors tubby_CT-GFP and PLCδ1-PH-GFP showed that pharmacological activation of muscarinic acetylcholine (mAChR) or group I metabotropic glutamate (mGluRI) receptors induces transient depletion of PIP₂ in the soma as well as in the dendritic tree. The observed PIP₂ dynamics were receptor-specific, with mAChR activation inducing stronger PIP₂ depletion than mGluRI, whereas agonists of other Gα_q-coupled receptors expressed in CA1 neurons did not induce measureable PIP₂ depletion. Furthermore, the data show for the first time neuronal receptor-induced oscillations of membrane PIP₂ concentrations. Oscillatory behavior indicated that neurons can rapidly restore PIP₂ levels during persistent activation of Gq and PLC. Electrophysiological responses to receptor activation resembled PIP₂ dynamics in terms of time course and receptor specificity. Our findings support a physiological function of PIP₂ in regulating electrical activity.

Introduction

Phosphatidylinositol-4,5-bisphosphate (PIP₂) directly controls many cellular functions, including membrane and cytoskeletal dynamics and the activity of membrane proteins^1–4. These regulatory effects of PIP₂ are mediated by modulation of activity or of membrane association of PIP₂-interacting proteins. In particular, many ion channels are highly sensitive to manipulation of PIP₂ levels^5,6.

In cell culture models activation of Gα_q-coupled receptors can deplete the PIP₂ content of the plasma membrane by activating phospholipase Cβ (PLCβ)^1–3,7. Such PIP₂ concentration changes were also observed in primary cultures of purkinje⁸ and hippocampal^9–12 neurons. A series of thorough analyses of PIP₂ signaling in response to muscarinic receptor activity in isolated sympathetic ganglion neurons (SGC)^13–18, established that in these neurons activation of some Gq-coupled receptors leads to transient depletion of PIP₂, which in turn inhibits Kv7.2/3-mediated M-currents. Hence, PIP₂ depletion downstream of receptor-mediated pathways may be a ubiquitous principle controlling neuronal activity by modulating ion channels^19,20. Thus it seems likely that the well-known increase of excitability by modulatory neurotransmitters, e.g. in hippocampal pyramidal neurons^21,22, is mediated by deactivation of PIP₂ -dependent channels²³. There is also some evidence for PIP₂-dependent regulation of Kir and K2P channels in striatal and thalamic neurons, respectively^24,25. However, channel modulation may as well be mediated by other cellular signals downstream of Gq activity. Thus Kv7 channels are also inhibited by intracellular Ca²⁺ elevation²⁶ and inhibition of Gq-sensitive TASK channels is mediated by DAG²⁷. Both, Ca²⁺ and DAG signals may occur without a substantial drop in PIP₂ downstream of PLCβ^28–30.

Another issue is the method used to assess PIP₂ dynamics. The standard approach is live-cell fluorescence microscopy using genetically encoded sensors built upon PIP₂-binding domains fused to fluorescent proteins³. The most popular sensor domain also used by the studies cited above is the pleckstrin homology domain from PLCδ1 (PLCδ1-PH)^31,32. However, interpretation of the observations is complicated by the IP₃ affinity of PLCδ1-PH, compromising its suitability as a sensor of PIP₂ following PLC activation^33,34. More recently, the PIP₂-specific tubby_CT domain enabled unequivocal measurement of Gα_q-induced PIP₂ depletion in cultured sympathetic and hippocampal neurons^15,33. While these findings derived from isolated neurons are consistent with substantial PIP₂ concentration changes, they might still not reflect physiological conditions. Studies in cardiac myocytes showed that PIP₂ content can considerably differ between isolated cells and cells in situ³⁵. Besides differences in the extent of PIP₂ depletion, differences in time course may also have relevance for signaling via this pathway. Thus, knowledge of PIP₂ dynamics in native neurons is required for understanding their physiological significance.

To address these issues, we characterized the PIP₂ concentration behavior induced by activation of Gα_q/PLC-coupled transmitter receptors in hippocampal CA1 pyramidal neurons in situ in acute brain slices. These neurons receive modulatory cholinergic input from the septohippocampal pathway³⁶, which is mediated postsynaptically by Gq-coupled M1/M3 receptors and results in transient changes of excitability^37–39. We find that mAChR and mGluRI receptors induce robust PIP₂ depletion in soma and main apical dendrites. Strikingly, both receptors induced PIP₂ oscillations. Moreover, PIP₂ depletion was receptor and neuron type-specific. Correlation with changes in electrophysiological activity supports an instructive signaling role of these neuronal PIP₂ dynamics.

Results

Muscarinic receptors mediate PIP₂ dynamics in CA1 neurons in situ

We began our investigation of PIP₂ dynamics in acute brain slices by examining the response of hippocampal CA1 pyramidal neurons to activation of their muscarinic ACh receptors. In order to measure Gα_q induced PIP₂ dynamics in situ, we expressed genetically encoded PIP₂ sensors by stereotaxic injection of lentiviral expression vectors into the hippocampi of juvenile (P21) rats (see Methods). Two different GFP-fused sensor domains were used, tubby_CT-GFP^15,40,41 and PLCd1-PH-GFP^31,32. Both sensors work as ‘translocation sensors’, i.e. their degree of membrane association is a direct measure for PIP₂ concentration and its temporal dynamics³.

Fluorescence of neurons in acute slices from rats (P26-32) infected with the vector encoding tubby_CT-GFP indicated successful expression in CA1 pyramidal neurons. As shown in Fig. 1a GFP fluorescence was primarily localized to the plasma membrane of the soma and the dendritic tree.

Figure 1.

mAChR-mediated PIP₂ depletion in acute brain slices. (a) Representative confocal image of CA1 pyramidal neurons expressing GFP-tagged tubby_CT PIP₂ probes one week after injection of viral vector. (b) Schematic diagram of hippocampal slice and positioning of the application capillary in relation to pyramidal neurons in the CA1 area. (c) tubby_CT-GFP translocation during mAChR activation. Under resting conditions probes are associated with PIP₂ at the membrane. Application of Oxo-M (10 µM) induced probe translocation to the cytosol indicating PIP₂ depletion, followed by reassociation to the membrane indicating PIP₂ recovery. (d) Representative somatic response to Oxo-M. Shown are cytoplasmatic fluorescence relative to fluorescence before stimulation (F/F₀, upper panel) and a kymograph of probe translocation between membrane and cytosol (lower panel). (e) Average response from 28 individual neurons (26 slices; 16 rats). Application bar is set according to mean response latency. Reduced baseline fluorescence level at the end of the experiments in (d,e) results from photobleaching of GFP. (f) Translocation amplitudes upon repeated Oxo-M application (n = 11, 11, 11, 5, 4, and 3 for successive applications). Responses of individual neurons are shown in grey. (g) Confocal image (maximum intensity projection) of a representative CA1 pyramidal neuron expressing PLCδ1-PH-GFP. (h) Average translocation of PLCδ1-PH-GFP upon Oxo-M application (n = 16 neurons, 11 slices, 7 rats). Scale bars in (a,f): 50 µm. Contrast enhancement of 0.4% and 3% was applied to images shown in (c,d), respectively.

Activation of mAChR receptors by application of the specific agonist, oxotremorine-M (Oxo-M), induced massive and reversible translocation of the tubby_CT probes from the membrane to the cytoplasm in >95% of CA1 pyramidal cell somata examined, indicating strong PIP₂ depletion (Fig. 1c). To quantify extent and time-course of probe translocation and hence PIP₂ dynamics we measured fluorescence intensity changes in cytosolic ROIs, as cytosolic signals turned out to be less sensitive to tissue movement than when measuring from the small membrane compartment. Consequently an increase of fluorescence signal corresponds to probe dissociation from the membrane, indicating a loss of PIP₂. Figure 1d shows a representative response to application of Oxo-M for 30 s. The mean tubby_CT translocation reached a peak cytoplasmic amplitude (F/F₀) of 1.73 ± 0.06 (mean ± SEM; n = 28; 26 slices; 16 rats; Fig. 1e). Mean response latency was 8.8 ± 2.5 s and 90% of the peak response (t₉₀) was reached within 19.9 ± 2.4 s. Upon washout of the agonist PIP₂ levels recovered within about 100 s as indicated by re-association of the probe to the membrane. Cytoplasmic fluorescence returned to 10% of peak amplitude (t₁₀, i.e. 90% recovery) in 65.1 ± 7.6 s. To explore the variability of muscarinic PIP₂ dynamics, Oxo-M was applied repetitively with subsequent stimulations separated by a time interval of >10 minutes (Fig. 1f). On average, the degree of PIP₂ depletion exhibited a slight but consistent decline in the course of repetitive stimulation. This is consistent with desensitization of muscarinic signaling previously observed in primary hippocampal neurons¹¹.

Previous experiments with cultured neurons have used another sensor domain, PLCδ1-PH, to examine PIP₂ dynamics^9,17,18. In contrast to the tubby_CT sensor, however, PLCδ1-PH has a significant affinity for IP₃, which is produced whenever PIP₂ is cleaved by PLCβ^{33,34,40,42–44}. Therefore the reliability of PLCδ1-PH as an indicator of PIP₂ dynamics during GqPCR/PLC signaling has remained an unresolved issue^33,34, which provided the rationale for choosing tubby_CT in the present study. Indeed, previous studies showed considerable differences in the behavior of tubby_CT-GFP and PLCδ1-PH-GFP sensors in terms of translocation following PLC activation^15,34,40. Thus, we were interested in comparing both sensors in acute brain slices. When PLCδ1-PH-GFP was expressed in CA1 neurons (Fig. 1g), stimulation of mAChRs resulted in robust translocation of fluorescence in all neurons examined (Fig. 1h), similar to the results obtained with tubby_CT-GFP (PLCδ1-PH-GFP: latency 4.9 ± 1.2 s; t₉₀ 22.3 ± 1.9 s; F/F_0 =1.72 ± 0.07; n = 16, 11 slices, 7 rats). However, the recovery was significantly slower compared to tubby_CT-GFP (t₁₀ = 132.7 ± 20.8 s; t-test p = 0.008), suggesting that responses of the PH sensor are co-determined by IP₃ production.

Receptor-specific PIP₂ depletion in CA1 neurons

Activation of PLCβ and subsequent hydrolysis of PIP₂ to IP₃ and DAG is the main signaling pathway of Gα_q-coupled receptors. Yet it is not known if PIP₂ depletion is generally associated with the activation of Gα_q coupled receptors other than M1/M3 receptors in central neurons. For example, in sympathetic ganglion neurons activity of muscarinic and purinergic receptors results in a depletion of PIP₂, whereas bradykinin receptors generate IP₃-dependent Ca²⁺ signals without substantial changes of the PIP₂ concentration^{17,18,29,45,46}. CA1 pyramidal neurons express various Gα_q-coupled receptors which could potentially induce PIP₂ depletion, including group I metabotropic glutamate receptor⁴⁷, α_1A-adrenoreceptor⁴⁸, bradykinin B₂ receptor⁴⁹, Gq-coupled dopamine_D1-like^50–55, histamine H₁ receptor^56,57, P2Y₁ receptor^58,59, and 5-HT_2A/2C receptors^60–62. We applied specific agonists of group I mGluRs, 5-HT_2A/2C receptors α_1A adrenoreceptors, bradykinin receptors, Gq-coupled dopamine_D1-like receptors, H₁ histamine receptor, or P2Y₁ receptor and monitored PIP₂ concentrations with tubby_CT-GFP. Of these receptors, only mGluRs induced detectable probe translocation indicative for depletion of PIP₂ (Fig. 2a). mGluRI-induced PIP₂ depletion in neuronal somata was consistent across the population of neurons examined (F/F₀ = 1.37 ± 0.05; n = 15; 14 slices from 11 rats; Fig. 2b).

Figure 2.

Somatic PIP₂ dynamics are receptor specific. (a) Representative experiment showing PIP₂ depletion in a CA1 soma in response to application of mGluRI agonist DHPG (10 µM) as determined by tubby_CT-GFP translocation. (b) Average PIP₂ concentration changes obtained from 15 cells as in (a) (blue). Response to activation of mAChRs (grey) is redrawn from Fig. 1e for comparison (n = 28). (c) Summary of peak PIP₂ depletion (translocation of tubby_CT-GFP) upon application of specific agonists for the Gq-coupled receptors indicated. Agonists applied were Oxo-M (10 µM, n = 28 neurons/26 slices/16 rats), DHPG (10 µM, n = 15/14/11), DOI, 10–20 µM, n = 14/14/6), methoxamine (10–20 µM; n = 14/14/6), bradykinin (10–20 µM, n = 10/10/4), SKF 83959 (10–20 µM, n = 5/5/3), 2-pyridylethylamin (10–50 µM, n = 10/10/4), ADPβS (10 µM; application for 30 to 60 s, n = 13/13/4). Numbers of experiments also indicated above bars. (d) Representative sensor responses measured from the same CA1 neuron during successive application of 5-HT_2A/2C agonist DOI and mAChR agonist Oxo-M. (e) Response to higher concentration (100 µM) of DHPG did not increase tubby_CT-GFP translocation (application for 5 min each; n = 6, 6 slices, 3 rats). Successive responses of individual neurons shown in grey. (f) Average response of PLCδ1-PH-GFP sensor to mGluRI activation (blue, DHPG) and mAChR activation (black; Oxo-M; replotted from Fig. 1h). (g) Peak translocation of PLCδ1-PH-GFP sensor during application of agonists as in (c); Oxo-M (n = 16 neurons, 11 slices, 7 rats), DHPG (n = 15/11/7), DOI (n = 19/13/7), methoxamine (n = 18/12/7), bradykinin (n = 19/13/7), SKF 83959 (n = 16/11/6), 2-pyridylethylamin (n = 15/11/6), ADPβS (n = 19/13/7). (h) Example responses from two different neurons to application of DHPG (10 µM). Note that one response (dark blue) closely matches the average response while the other (light blue) shows pronounced recovery in the presence of the agonist. (i) Responses to repeated DHPG application (n = 11, 11, 8, 3, 3, and 2 for successive applications, respectively; delay between applications ≥ 10 min; individual responses indicated in light blue).

As summarized in Fig. 2c, PIP₂ levels in CA1 neurons were insensitive to activation of any of the other receptors. Importantly, subsequent control application of Oxo-M triggered robust translocation of tubby_CT-GFP in each neuron, indicating proper responsiveness of the cell and appropriate sensitivity of the detection approach (Fig. 2d). Further, the prolonged application of each of the agonists (except muscarinic and glutamatergic) for up to 120 seconds or application of the endogenous ligands serotonin and dopamine did not evoke detectable responses (not shown). Equivalent results were obtained with neurons expressing the alternative PIP₂ sensor domain, PLCδ1-PH-GFP. As shown in Fig. 2f and g, the stimulation of mAChR and mGluRI but none of the other receptors examined induced translocation of PLCδ1-PH-GFP. In summary, results obtained with both sensor domains indicate that α_1A-adrenoreceptor, bradykinin, dopamin_D1-like, histamin-H₁, P2Y₁, and 5-HT_2A/2C do not induce significant PIP₂ depletion in the soma of CA1 pyramidal neurons. Thus, PIP₂ depletion is specific to mAChR and mGluRI, at least in the context of standard experimental conditions. However, response magnitude of mGluR activation was significantly lower compared to muscarinic PIP₂ depletion in a population of cells challenged by both agonists (n = 12, 11 slices, 8 rats; paired t-test p = 0.032). While Oxo-M and DHPG have similar binding affinities for their cognate receptors⁶³, EC50 values for downstream effects such as Ca²⁺ responses are often higher for DHPG than for Oxo-M, raising the possibility that 10 µM of DHPG might not be sufficient to evoke a saturating PIP₂ response. However, increasing the concentration to 100 µM or the duration of agonist application of the glutamatergic agonist (DHPG) did not further increase the response mediated by mGluRs (Fig. 2e) and these responses were significantly smaller than muscarinic responses in the same neurons (p < 0.05, one way ANOVA followed by Tukey post-hoc test, n = 6, 6 slices, 3 rats).

In addition to the smaller responses, mGluRI-induced depletion of PIP₂ also differed in its time course compared to muscarinic stimulation (Fig. 2a,b,f). As measured with tubby_CT-GFP, response latencies (6.81 ± 0.89 s) and rise time (t₉₀ = 12.68 ± 1.15 s) where comparable, but time course of recovery was faster compared to mAChR activation (t₁₀ = 28.38 ± 3.57 s; t-test p = 0.0001) Remarkably, in 6 out of 15 recordings, PIP₂ levels recovered in the continued presence of the agonist DHPG, as illustrated by individual recordings shown in Fig. 2h. Similarly, PIP₂ dynamics as measured with the PLCd1-PH-GFP probe showed a much faster recovery after activation of mGluRI (t₁₀ = 30.44 ± 2.05 s, n = 15, 14 slices, 7 rats) when compared to mAChRs (t₁₀ = 132.74 ± 20.77 s; t-test p = 0.0003; Fig. 2f). As noted with mAChR activation, PIP₂ dynamics induced by mGluRIs showed slight desensitization in response to repeated application of the agonist (Fig. 2i).

Dendritic PIP₂ dynamics

Next, we were interested in the spatial pattern of PIP₂ depletion, in particular with respect to dendritic compartments. Because we probed PIP₂ dynamics with translocation sensors that require microscopic resolution of membrane versus cytoplasm the measurements were confined to dendrites with a diameter of more than 1 µm that were localized close to the slice surface allowing for good optical access. Thus we achieved recordings from the main apical dendrite and its major branches up to 300 µm and basal dendrites to 20 µm distal to the soma. The distance between stratum pyramidale to fissura hippocampi defining the total length of apical dendrites was about 400 µm for our slices.

We found robust but receptor-specific PIP₂ depletion in all dendritic compartments examined. Figure 3a shows a representative example illustrating membrane localization of tubby_CT-GFP in a dendrite and its transient redistribution into the dendritic cytosol during pharmacological activation of mAChRs, indicating reversible depletion of PIP₂. With the exception of one apical dendrite (190 µm distance from soma), all dendrites examined (n = 12) responded with the translocation of tubby_CT-GFP. The time course of representative mAChR-induced dendritic PIP₂ dynamics is further shown in Fig. 3d as a kymograph and quantitatively as the change of cytosolic fluorescence intensity. The average peak amplitude derived from dendrites of 12 neurons was F/F₀ = 1.57 ± 0.06 (12 slices, 10 rats). The average latency of the responses was 14.29 ± 5.26 s. We observed a rise time t₉₀ of 17.52 ± 2.67 s and 90% recovery time (t₁₀) was 41.27 ± 7.25 s after the end of the application. As shown in Fig. 3a,e, activation of mGluRI also resulted in translocation of tubby_CT-GFP. However, the degree of PIP₂ depletion was significantly weaker compared to activation of mAChRs (F/F₀ = 1.18 ± 0.02; paired t-test, p = 0.041, n = 3 dendrites in 3 slices from 2 rats. Similar observations were made with PLCδ1-PH-GFP as the PIP₂ probe (Fig. 3c,h). Thus, activation of muscarinic receptors by Oxo-M induced strong translocation (F/F₀ = 1.86 ± 0.07; t₉₀ = 21.86 ± 1.90 s; t₁₀ = 142 ± 30.53 s; n = 7 dendrites, 7 slices, 3 rats), whereas activation of mGluRI receptors by DHPG induced small yet reproducible translocation of PLCdδ1-PH-GFP (F/F₀ = 1.19 ± 0.02; n = 7). Stimulation of various other Gq-coupled receptors did not induce detectable translocation of PLCδ1-PH-GFP (Fig. 3c).

Figure 3.

Dendritic PIP₂ dynamics. (a) Translocation of tubby_CT-GFP in response to Oxo-M application (10 µM) in an apical dendrite 60 µm from the soma. (b) Translocation of tubby_CT-GFP from dendritic membranes induced by mAChR (Oxo-M, 10 µM, n = 12 neurons/12 slices/10 rats) and mGluRI (DHPG, 10 µM, n = 3/3/2) activation, quantified as changes of axial (cytoplasmic) fluorescence changes relative to pre-stimulus fluorescence. (c) Translocation of PLCδ1-PH-GFP from dendritic membranes in response to activation of various Gq-coupled receptors. Agonists used as in Fig. 2 (n = 7 neurons, 7 slices, 3 rats for each condition). (d) Time course of PIP₂ depletion from a dendritic recording 220 µm from the soma, shown as the relative increase of axial fluorescence (upper panel) and corresponding kymograph (lower panel). (e) Representative fluorescence change and kymograph of dendritic tubby_CT-GFP translocation in response to application of DHPG (10 µM; apical dendrite; 20 µm from soma). (f) Average time course of mAChR-induced dendritic PIP₂ dynamics. (g) Amplitudes (black) and time constants (green) of tubby_CT-GFP translocation in response to application of Oxo-M plotted as a function of the distance of the recording location from the neuronal soma. Data from 13 individual neurons. (h) Average time course of dendritic PLCδ1-PH-GFP sensor translocation in response to application of the agonists indicated (n = 7 each). (i) Properties of translocation of PLCδ1-PH-GFP in response to Oxo-M plotted as a function of dendritic recording site relative to soma. Contrast enhancement of 10%, 1% and 3% was applied to images shown in a, d and e, respectively.

We wondered about spatial, basal-to-apical signaling gradients along the dendrites. Figure 3g,i display amplitudes and time constants of PIP₂ depletion as a function of the distance from the soma. We find that neither parameter shows an evident trend along the dendritic distance, suggesting that PIP₂ depletion is mostly homogenous throughout the larger dendritic compartments examined here (Pearson correlation coefficients: tubby_CT-GFP amplitudes, −0.071; tubby_CT-GFP time constants, 0.168;, PLCδ1-PH-GFP amplitudes, −0.209;, PLCδ1-PH-GFP time constants, −0.233).

Taken together, these results indicate that receptor-induced PIP₂ dynamics in the larger dendritic compartments amenable to examination with translocation sensors is similar to the somatic dynamics. Specifically, activation of muscarinic receptors resulted in strong depletion of PIP₂, glutamatergic receptors were considerably less effective, and PIP₂ levels recovered rapidly.

Prolonged receptor activation revealed complexity of PIP₂ dynamics

The occasionally observed early recovery of PIP₂ level during activation of mGluRI prompted us to examine the time course of PIP₂ dynamics during sustained stimulation. Figure 4 shows the resulting PIP₂ dynamics measured with the tubby_CT sensor in neuronal somata during continuous application of the receptor agonists for 5 min. Notably, PIP₂ depletion generally showed a phasic-tonic time course with an initial strong PIP₂ depletion followed by partial recovery in the sustained presence of the agonist as apparent from the average from a larger number of neurons (n = 17 and 16 for mAChR and mGluRI activation, respectively; Fig. 4a). The initial rate of PIP₂ decrease was similar for both receptors (t₉₀ = 39.3 ± 5.3 s and 43.3 ± 8.8 s, respectively), but as previously seen with brief receptor stimulation (Fig. 2), the muscarinic responses had a higher average amplitude compared to glutamatergic responses (F/F₀ = 1.84 ± 0.08 and 1.55 ± 0.07, respectively; paired t-test p = 0.0002, n = 16, 14 slices; 5 rats). PIP₂ recovery after muscarinic receptor activation was more pronounced than for mGluR stimulation such that PIP₂ levels tended towards similar values at the end of receptor stimulation period. For both receptors, recovery of PIP₂ levels after removal of the receptor agonist was slower than observed with brief receptor stimulation (mAChR: t₁₀ 243.2 ± 43.9 s; mGluRI: t₁₀ 167.8 ± 41.5 s; cf. Fig. 2b) and thus depended on the duration of receptor activation.

Figure 4.

Prolonged receptor activation reveals PIP₂ oscillations. (a) Average time course of somatic PIP₂ dynamics during extended application of mAChR and mGluRI agonists Oxo-M (n = 17 neurons/15 slices/5 rats) and DHPG (n = 16/14/5) as measured by translocation of tubby_CT-GFP. (b) Distribution of distinct temporal patterns of tubby_CT-GFP sensor translocation within the population of neurons challenged by sustained application (5 min) of agonists. Responses were categorized as constant, desensitizing (partial recovery), transient (full recovery) and oscillating (≥2 peaks). (c) Distinct temporal behavior of PIP₂ dynamics of an individual pyramidal neuron to successive applications of Oxo-M and DHPG. Lower panel shows a kymograph illustrating the response to Oxo-M. (d) Oscillatory PIP₂ dynamics in a CA1 neuron in response to prolonged activation of mACh receptors. Note the pronounced repetitive phases of rapid PIP₂ depletion and replenishment during stimulation. (e) Oscillatory PIP₂ dynamics of a neuron in response to continuous activation of mGluRs. Contrast enhancement of 0.4%, 1% and 5% was applied to kymographic images shown in c, d and e, respectively.

While demonstrating partial desensitization of PIP₂ responses as a common pattern, the prolonged stimulation also revealed considerable variability and complexity in the time course of PIP₂ dynamics (Fig. 4b). Most strikingly, PIP₂ depletion was often multiphasic or oscillatory. Examples for such complex PIP₂ dynamics are shown in Fig. 4c–e. In these cells, PIP₂ levels returned to baseline despite sustained presence of the agonist, and moreover, multiple depletion events occurred in rapid succession (Fig. 4d,e). Altogether, 8 out of 17 neurons showed oscillatory PIP₂ concentration dynamics with Oxo-M and two out of 16 during application of DHPG. These observations suggest that PIP₂ dynamics may be subject to complex temporal regulation and indicate potent PIP₂ resynthesis capability of CA1 neurons during receptor activation.

Modulation of electrical behavior by receptor stimulation

The well-described effects of muscarinic activity on the electrical properties of CA1 neurons – including inhibition of M-currents – may (at least partially) be mediated by PIP₂ concentration dynamics. We thus were interested in differential effects on excitability of the various Gq/PLC-coupled receptors examined for their coupling to PIP₂ dynamics.

To this end, we performed patch clamp experiments in current clamp mode in acute brain slices prepared from rats at P14 to P21. Current step protocols were used to assess membrane potential, input resistance, spiking behavior, and afterpolarisation (Fig. 5a,b). Experiments were performed in the presence of inhibitors of GABA_A/B and ionotropic glutamate receptors (see Methods) to exclude effects resulting from network activity.

Figure 5.

Electrophysiological responses to activation of Gq-coupled receptors. (a,b) Current clamp protocol (lower panel) and representative recordings of corresponding membrane potential (V_m) responses of CA1 pyramidal neurons before (light traces) and during (dark traces) activation of mAChRs or mGluRIs by 10 µM Oxo-M or DHPG, respectively. (c) Changes of resting membrane potential (ΔV_m) displayed as the difference before and upon application of various GqPCR activators. Agonists applied were Oxo-M (10 µM; n = 10 neurons/10 slices from 7 rats; data points with plateau potentials were excluded for this analysis) for AChRs, DHPG (10 µM; n = 9/8/5) for mGluRIs, DOI (20 µM; n = 8/8/7) for 5-HT_2A/C, methoxamine (20 µM; n = 8/8/7) for α₁-adrenergic receptors, bradykinin (20 µM; n = 7/7/6), SKF 83959 (20 µM; n = 7/7/6) for D1-like dopamine receptors, 2-pyridylethylamin (20 µM; n = 8/8/7) for H₁ histamine receptors, ADPβS (10 µM; n = 15/15/11) for P2Y₁-R. Asterisks indicate significance of difference to control application of ACSF (n = 10/10/7) with p < 0.05 (*), 0.01 (**) and 0.001 (***; one-way ANOVA followed by Dunnett multiple comparison test). (d) Difference in number of action potentials (AP) triggered during 600 ms depolarizing current step. Numbers of experiments as in (c). (e) Changes in afterpolarisation where a negative Δ indicates an AHP_m increase and positive Δ indicates an AHP reduction or afterdepolarisation. Numbers of experiments as in (c). (f) Mean time courses of V_m modulation during application of ACSF as a control (grey), Oxo-M (black), DHPG (blue), and DOI (purple). Vm was measured at the first 500 ms of each trace in the absence of spiking; see (a). (g) Example membrane voltage oscillation during mAChR activation in a P27 slice.

Overall, we found pronounced changes in electrical behavior following the activation of muscarinic mAChR receptors and mGluRI but little effects of other Gq-coupled receptors. Consistent with previous findings^64–67 agonists of both mAChR (n = 11, 11 slices, 10 rats) and mGluRI (DHPG 10 µM, n = 9, 8 slices, 5 rats) induced depolarization of the resting membrane potential and an increase in firing frequency during depolarization (number of action potentials, NAP; Fig. 5a–d,f). In CA1 cells, a train of action potentials is usually followed by an afterhyperpolarization (AHP)^37,68,69. Application of either Oxo-M or DHPG resulted in the disappearance of the AHP and the appearance of an afterdepolarisation (ADP; Fig. 5a,b,e). All of these receptor-induced changes are consistent with the deactivation of potassium conductances such as M currents^37,68,70,71. In some neurons, activation of mAChR receptors induced sustained depolarization (plateau potentials) subsequent to the 600 ms current step, as also described previously⁷².

As shown in Fig. 5c–f, stimulation of other Gq-coupled receptors, including 5-HT_2A/C receptors, α₁ adrenergic receptors, bradykinin receptors, D₁-like dopamine receptors, H₁ histamine receptors, and purinergic P2Y₁ receptors had no significant effects on either of these electrophysiological characteristics. Noteworthy, the mAChR and mGluRI induced depolarization did not always persist during the agonist application, as occasionally initial transient hyperpolarisation and oscillations of the membrane potential was observed during mAChR activation. During muscarinic stimulation, 9 of 11 neurons showed substantial recovery from depolarization in the presence of Oxo-M; of those, 6 neurons showed complete repolarization or even hyperpolarisation before washout. In the presence of DHPG, 3 neurons showed a substantial recovery from initial depolarization. Oscillatory behavior was observed in neurons from younger animals (P14-21), but also in slices age-matched to the PIP₂ imaging experiments, as shown in an exemplary recording (Fig. 5g) obtained from a neuron in a P27 slice.

In summary, we find that pronounced changes in membrane potential and firing rates paralleled neuronal PIP₂ depletion in terms of effect size, time course and receptor specificity. Thus, depolarization, increased spike rates and PIP₂ dynamics were largely restricted to the activation of mAChR and mGluRI receptors.

Neuron type-specific PIP₂ dynamics: dentate gyrus granule cells

To extend our observations on PIP₂ dynamics to additional neuronal cell types, we measured PIP₂ dynamics following activation of the same receptors (mAChR and mGluRI) in dentate gyrus granule neurons (Fig. 6a). Granule neurons express both mAChR and mGluRI receptors at the soma^47,73,74. However, detectable depletion of PIP₂ upon stimulation with Oxo-M was observed in only three independent experiments (n = 24; Fig. 6b,c). None of the six neurons stimulated with DHPG (10 µM, n = 2) or Glutamate (100 µM, n = 4) showed any detectable sensor translocation (Fig. 6d). Thus, the induction of PIP₂ dynamics may be highly specific between different types of neurons and this specificity seems to be dependent on mechanisms other than the expression of Gq/PLC-coupled transmitter receptors.

Figure 6.

Dentate gyrus granule neurons largely lack receptor-induced PIP₂ depletion. (a) Confocal image of gyrus dentatus granule neurons in an acute hippocampal slice, expressing tubby_CT-GFP after sterotactic injection of lentiviral vector. (b) Confocal images demonstrating a minor degree of probe translocation (1% contrast enhanced; first neuron from left in (a)). (c) Only 3 out of n = 24 neurons showed mAChR-induced PIP₂ depletion, as indicated by sensor translocation (3 individual neurons from separate animals). Black trace shows mean fluorescence time course of 21 non-responsive neurons. Time 0 indicates start of the application shown by bars. (d) Application of the mGluRI agonist DHPG (n = 2 neurons, both of which were responsive to Oxo-M) or glutamate (n = 4) did not induce PIP₂ depletion.

Discussion

Direct observation of PIP₂ dynamics in central neurons in situ

While there is good evidence for PIP₂ depletion in response to activation of Gq-coupled receptors for some types of neurons studied in the cell culture dish, surprisingly little is known about the prevalence and spatiotemporal properties of PIP₂ dynamics in central neurons under physiological conditions. PIP₂ levels and their dynamic regulation may be largely different in vivo, as embedding in the native environment and full differentiation of neurons may impact on relevant factors such as expression and spatial subcellular organization of receptors and downstream components of the signaling cascade and the enzymes that resynthesize PIP₂. Therefore, information on PIP₂ concentration behavior in intact tissue preparations such as brain slices is required. Previous studies with organotypic slices were consistent with PIP₂ depletion in-situ triggered by synaptic release of glutamate onto cerebellar Purkinje neurons⁷⁵ or by muscarinic agonist application in cortical pyramidal cells⁷⁶. However, both studies were not fully conclusive with respect to PIP₂ signaling because PLCδ1-PH was used as a sensor domain, which has a similar affinity for PIP₂ and IP₃ and may report the production of IP₃ rather than depletion of PIP₂^44,77. In fact, Okubo et al.⁷⁵ interpreted probe translocation in terms of IP₃ production rather than depletion of PIP₂.

Here, by using tubby_CT, as an alternative PIP₂ sensor insensitive to IP₃^33,34,40, our present data now show unequivocally that muscarinic and metabotropic glutamate receptors indeed trigger PIP₂ dynamics in a prototypic central neuron in situ. Of note, in cultured cell lines tubby_CT previously failed to respond or only weakly translocated upon PLC-mediated PIP₂ depletion^34,40, which initially was attributed to a higher affinity for PIP₂ compared to PLCδ1-PH. However, quantitative titration of PIP₂ in living cells showed that its affinity to PIP₂ is actually lower, which should make it a useful PIP₂ sensor⁴¹. In fact, our current results demonstrate that tubby_CT readily translocates in a neuronal cellular environment. The difference in behavior between experimental conditions is not yet understood, but may suggest cell type-specific segregation of PIP₂ into distinct pools selectively accessible to the different PIP₂-binding domains⁷⁸. In any case, our findings show that in the native neuronal system tubby_CT is a much better reporter of PIP₂ dynamics than might have been anticipated²⁰. Thus, using tubby_CT-GFP allowed us to systematically assay PIP₂ dynamics without confounding effects of IP₃ dynamics.

Spatiotemporal properties of PIP₂ dynamics

We found that in the larger compartments accessible to measuring sensor translocation, receptor-induced PIP₂ depletion appeared largely homogenous without evidence for substantial subcellular differences. This suggested that induced PLC activity is similar in somatic and dendritic compartments. This finding was not entirely expected, because while M1 receptors show high densities throughout soma and dendrites⁷³, the most prominent mGluRI receptor of CA1 neurons, mGluR₅, has a relatively low density in soma compared to dendrites⁴⁷. Since glutamatergic PIP₂ depletion in dendrites was not stronger than in the somatic compartment, the PIP₂ depletion pattern does not appear to correlate closely with receptor distribution. It is worth noting that the moderate degree of PIP₂ sensor translocation in dendrites did not result from the distinct dendritic geometry, because the smaller volume-to-membrane area ratio in the dendrites should rather result in a stronger relative increase of sensor fluorescence when sensors dissociate from the membrane. Also, muscarinic stimulation elicited larger dendritic responses than glutamatergic stimulation, showing that PIP₂ sensor response was not saturated by mGluR stimulation. Of note, a similar observation was made by Nakamura et al.⁷⁹ for Ca²⁺ dynamics in CA1 pyramidal cells: activation of mGluRI, mAChR and 5-HT₂R elicited comparable Ca²⁺ waves despite different receptor distribution. Thus, these neurons may possess mechanisms to globalize Gq signaling including PIP₂ depletion.

To our knowledge our results for the first time demonstrate oscillations of the PIP₂ concentration in a neuron. While IP₃, DAG and Ca²⁺ are known to undergo oscillatory concentration dynamics in neurons⁸⁰, previous observations on PIP₂ dynamics in primary dissociated neurons^16,18,33 seemed to indicate that PIP₂ concentrations essentially remained depleted during prolonged receptor activity. Oscillatory translocation of the PLCδ1-PH sensor domain observed occasionally has been understood as dynamics of the IP₃ signal picked up by the PH domain^23,44,81. More recently, careful observations also including the specific tubby_CT sensor showed bona-fide oscillations of PIP₂ in mast cells^82,83. Our observations suggest that such dynamics may be a more general phenomenon with implications for neuronal biology.

The mechanisms underlying PIP₂ oscillations may include both positive and negative feedback regulation of PIP₂ cleavage by PLC. Such mechanisms have previously been shown to be involved in Ca²⁺ and IP₃ oscillations and include Ca²⁺-dependent activation of PLC^80,81 as a positive feedback. In mast cells, PIP₂ oscillations are probably driven by Ca²⁺ oscillations⁸². Inhibition of Gq signaling by, e.g., PKC, receptor kinases, or RGS molecules^81,84 may contribute to a negative feedback loop controlling PIP₂ degradation. Moreover, our observations reveal an impressive capability of PIP₂ replenishment, as indicated by the rapid and complete recovery of PIP₂ levels in presence of agonists. PIP₂ resynthesis may be increased during GqPCR activation⁸⁵, providing negative feedback to PIP₂ depletion and possibly contributing to observed oscillations. Specifically, PIP₂ replenishment may involve Ca²⁺ and phosphatidic acid-dependent phospholipid exchange at plasma membrane-endoplasmic reticulum (PM-ER) junctions⁸⁶.

Whatever the mechanism underlying the oscillations is, our findings indicate that PIP₂ dynamics may provide neurons with another dimension of effector modulation beyond a simple on/off switch for downstream effectors. Although the consequences of PIP₂ oscillations for electrical neuronal activity remain to be explored, we note that indeed, neurons showed fluctuations of membrane potential and firing frequency during agonist application. It is worth mentioning that mAChR and mGluRI agonists can induce and shift gamma and theta oscillations⁸⁷. In the light of the present data, it is tempting to speculate that PIP₂ oscillations might participate in such frequency modulation.

Neuronal ion channels as effectors of PIP₂ dynamics

Given the known high sensitivity of some ion channels to even a moderate drop in the PIP₂ concentration^5,78, a main potential target of PIP₂ depletion are ion channels and thus electric excitability. Based on studies on isolated neurons, inhibition of Kv7 channels in sympathetic neurons as the direct consequence of PIP₂ depletion is well established^13,17,29. Our data permit the correlation of PIP₂ dynamics and electrophysiology in situ. Activation of mAChR and mGluRI, but not other PLC-coupled receptors known to be present and functional in CA1 neurons induced robust PIP₂ depletion. The same pattern of receptor specificity was observed for modulation membrane potential, firing frequency and afterhyperpolarization, providing at least circumstantial evidence for the causation of channel regulation by PIP₂. Simultaneous recordings of electrical activity and PIP₂ dynamics from the same neuron should be performed in the future to provide more direct evidence.

In conclusion, our data support and generalize the as yet largely hypothetical mechanism of PIP₂ dynamics as a major cellular signal in the control of neuronal activity through regulation of PIP₂-sensitive ion channels such as Kv7. Future studies need to address this issue rigorously by manipulating PIP₂ levels in-situ²⁰. Along those lines a recent study aimed at PIP₂ depletion in hippocampal slice cultures by chemically induced recruitment of a PIP₂ phosphatase⁸⁸. While this approach did not reveal any effects on electrical properties of the neurons, the results appear inconclusive since changes in PIP₂ concentration were not verified.

One of the most intriguing unknowns are the spatiotemporal properties of PIP₂ dynamics during entirely physiological neuronal activity, i.e, during synaptic activity of the modulatory (e.g. cholinergic) and principal (i.e. glutamatergic) inputs into the hippocampal neurons and of the PIP₂ dynamics associated with intrinsic neural (network) activity. Another question is the PIP₂ signaling in the distal smaller dendritic compartments not amenable to analysis by the translocation probes used in this study. In particular, in the immediate postsynaptic compartment, i.e. spines, PIP₂ may have a role in controlling synaptic plasticity^89–91.

Materials and Methods

Virus production and constructs

Lentiviral plasmids pCMVΔR8.9, pVSVG and FUGW were kindly provided by Pavel Osten (MPI for medical research Heidelberg, Germany). The PLCδ1-PH and tubby_CT constructs were provided by Tamás Balla (NIH, Bethesda, USA) and Lawrence Shapiro (Columbia University, USA), respectively. Lentiviral particles were derived by triple transfection of HEK293FT cells with Lipofectamin 2000 (Invitrogen, Darmstadt, Germany). Virus purification from supernatant was achieved by 15 minute centrifugation at 3000 rpm, filtration through a Millex ® HV 0.45 µm filter (Millipore, Darmstadt, Germany) and two successive ultracentrifugation steps (25000 rpm, 1 h 30 min, 4 °C). Pellets were resuspended in TBS-5 buffer (50 mM Tris-HCl, 130 mM NaCl, 10 mM KCl, 5 mM KCl₂) and subjected to a final 30 s centrifugation at 5000 rpm. Aliquots were stored at −80 °C and thawed up to two times.

Animals, stereotactic injection and slice preparation

Wistar rats were obtained from the animal facility of the Philipps University of Marburg (Marburg, Germany) or Charles River (Cologne, Germany) and kept and handled according to German law and institutional guidelines at the Philipps University. All procedures were approved by the Regierungspräsidium Giessen, Germany. Animals were housed with access to ad libitum water and food on a 12-h light/dark cycle. At weaning (postnatal day 21) male and female rats were anesthetized by intraperitoneal injection of a mixture of ketamine (Bela-Pharm, Vechta, Germany) and xylazine (Rompun®, Bayer AG, Leverkusen, Germany) at a dose of 100 and 10 mg per kilogram body weight. Additionally, the mixture included 0.05 mg/kg Atropine (B. Braun, Melsungen, Germany) and 0.1 ml/10 g body weight of a 0.9% NaCl solution for injection (Diaco, Triest, Italy). Under stereotactic control, lentivirus was injected bilaterally using Paxinos and Watson⁹² as a reference. Coordinates were optimized for targeting in juvenile rats by setting the adult references to x = +/−6.125, y = −6.15 and z = −6.2 mm and multiplying by the ratio of the juvenile to atlas (8.7 mm) distance of bregma to lambda. Up to 2.5 μl virus per hemisphere were injected in 500 nl portions going from ventral to dorsal in 0.3–0.35 mm steps during 5–10 minutes. For imaging and electrophysiological experiments, rats were anesthetized with Isoflurane (Baxter, Unterschleißheim, Germany) or Sevoflurane (Sevorane®, Abbott, Wiesbaden, Germany) and sacrificed by decapitation at the ages indicated in results. The head was placed in ice cold sucrose-ACSF (sucrose-artificial cerebrospinal fluid, in mM: 87 NaCl, 25 NaHCO₃, 25 D-glucose, 75 sucrose, 2.5 KCl, 0.5 CaCl₂ and 7 MgCl₂, oxygenated with 95% O₂/5% CO₂) and the hippocampi rapidly removed. 300 μm transversal slices were cut with a vibratome (VT1200, Leica Biosystems, Wetzlar, Germany) and placed into a chamber with 4 °C sucrose-ACSF. After a 35 min recovery period at 35 °C slices were kept at room temperature. For recordings slices were transferred to a submerged chamber and perfused with ACSF (in mM: 125 NaCl, 25 NaHCO₃, 25 D-glucose, 2.5 KCl, 2 CaCl₂ and 1 MgCl₂, oxygenated with 95% O₂/5% CO₂) for at least 20 minutes.

Imaging, electrophysiological recording and data analysis

Confocal imaging was performed with a Zeiss LSM710 (Zeiss, Oberkochen, Germany). The sampling rate for time series experiments was 1.75 s with a pixel size of 0.13 µm. In some cases (especially dendrite measurements) the sampling rate was increased to 1 s. In all cases where the sampling rate slightly deviated the data were resampled to allow averaging across experiments. Overlay with the original was performed to ensure preservation of time scale. Average cytoplasmic fluorescence intensities were determined from regions of interest (ROI) excluding both the plasma membrane (defined as the local intensity max at the cell’s border in the resting cell) and the nucleus. Distance of ROIs to the plasma membrane was >0.5 µm even when slight shifts of the cell’s position occurred during the experiment. ROIs were defined post-hoc using the microscope software ZEN (Versions 2008 and 2009) and obtained average intensities were exported to Igor Pro (Version 6.03 A, Wave Metrics, Portland, OR USA). Traces were background subtracted and normalized to the last time point before beginning of a response (F/F₀ normalized to t₀). Measurements without evident response were corrected for photobleaching according to a biexponential fit to the decaying fluorescence signal and normalized to signal at the onset of agonist application. We found that probe translocation generally prevented the reliable determination of the time course of photobleaching. Therefore most data were not corrected for bleaching which results in apparently lower signals following transient depletion of PIP₂, with bleaching generally being more pronounced for tubby_CT-GFP than for PLCδ1-PH-GFP. Confocal images were further analyzed with ImageJ (National Institutes of Health, USA) to isolate individual images of a time series, create kymographs and set scale bars. Electrophysiological data were recorded with a HEKA EPC10USB amplifier and Patch Master software (Version 2.43 HEKA, Lambrecht, Germany) in current clamp mode. Data were low pass filtered with a 2.9 kHz Bessel filter and digitized at 20 kHz. Borosilicate recording pipettes had a resistance of 3–4 MΩ and were filled with intracellular solution containing (in mM): K-gluconate 135, KCl 20, MgCl₂ 2, Na₂-ATP 2, Na₂-GTP 0.3, HEPES 10 and EGTA 0.1 (adjusted to pH 7.2 with KOH). Series resistance was monitored in voltage clamp mode before and after each current clamp recording, but not corrected for. Measurements with a change in series resistance >40% during the course of the experiment were discarded. Input resistance was assessed by injection of small positive and negative currents steps, followed by a depolarizing current step above action potential threshold to quantify spiking behavior and afterpolarisation (see Fig. 5a,b). Sweep length was 7 seconds. In applications of the P2Y₁ agonist ADPβS a shorter protocol without the positive 20 pA step was used. Medium afterhyperpolarisation (AHP_m) was obtained as the difference of resting V_m and mean V_m at 70 to 120 ms after the depolarizing current step. Changes in AHP value resulting from application of receptor agonists are given as Δafterpolarization such that positive values indicate reduction of AHP or eventually the emergence of an afterdepolarisation. Amplitudes were calculated from averaging at least 10 baseline data points and a minimum of 3 peak points, with avoidance of plateau potentials.

Statistical analysis

Statistical significance was tested in Igor Pro. Randomness, equal variances and normal distribution of the data was tested with Igor’s Runs, Kolmogorow-Smirnow and Jarque-Bera test. In cases where validity of a parametric test was compromised, a Wilcoxon-Mann-Whitney test was performed. Where applicable, groups of two were compared with paired and unpaired Student’s t. Two tailed one-way ANOVA was followed by a Dunett test for comparing multiple groups to a single control or a Tukey test to compare all groups to each other. Unless noted otherwise all values are given ± standard error of the mean.

Chemicals and perfusion system

Oxotremorine-M, DHPG, Bradykinin, SKF83959, DOI, Serotonin and Dopamine were purchased from Tocris and Methoxamine and 2-Pyridylehylamin from Sigma. All other chemicals were from Sigma/Fluka or Merck (Germany). For application of test substances a capillary of 200 to 250 µM inner diameter (TSP200350, BGB Analytik AG, Boeckten, Germany or MicroFil MF28G-5, World Precision Instruments, Berlin, Germany) was placed directly next to the hippocampal recording region. Solution exchange at the tip occurred within 1–2 s. Unless noted otherwise recordings represent first applications of each test substance. To block fast glutamatergic and GABAA/B signaling in electrophysiological recordings, receptor antagonists (4 μM NBQX, 50 μM D-AP5, 50 μM Picrotoxin and 1 μM CGP 55845, all from Tocris) were added both to the bath and local perfusion.

Author Contributions

D.O. and S.H. designed the study. S.H. performed the experiments and analyzed the data. S.H. and D.O. wrote the manuscript. All authors approved of the manuscript.

Data availability

Most data generated or analysed during this study are included in this published article. Additional datasets generated and analysed during the current study are available from the corresponding authors on reasonable request.

Competing Interests

The authors declare no competing interests.