Regulation of ion channel localization and

© 2004 Nature Publishing Group http://www.nature.com/natureneuroscience

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Regulation of ion channel localization and phosphorylation by neuronal activity Hiroaki Misonou1,2, Durga P Mohapatra1, Eunice W Park2, Victor Leung3, Dongkai Zhen2, Kaori Misonou1,2, Anne E Anderson3 & James S Trimmer1,2 Voltage-dependent Kv2.1 K+ channels, which mediate delayed rectifier Kv currents (IK), are expressed in large clusters on the somata and dendrites of principal pyramidal neurons, where they regulate neuronal excitability. Here we report activitydependent changes in the localization and biophysical properties of Kv2.1. In the kainate model of continuous seizures in rat, we find a loss of Kv2.1 clustering in pyramidal neurons in vivo. Biochemical analysis of Kv2.1 in the brains of these rats shows a marked dephosphorylation of Kv2.1. In cultured rat hippocampal pyramidal neurons, glutamate stimulation rapidly causes dephosphorylation of Kv2.1, translocation of Kv2.1 from clusters to a more uniform localization, and a shift in the voltagedependent activation of IK. An influx of Ca2+ leading to calcineurin activation is both necessary and sufficient for these effects. Our finding that neuronal activity modifies the phosphorylation state, localization and function of Kv2.1 suggests an important link between excitatory neurotransmission and the intrinsic excitability of pyramidal neurons.

Neurons possess various voltage-dependent ion channels, of which voltage-dependent K+ (Kv) channels are crucial for controlling mem brane electrical excitability. Individual Kv channel subtypes regulate distinct aspects of neuronal function according to their different bio physical properties, highly restricted subcellular localization and sus ceptibility to neuromodulation. Kv channels can be localized to synaptic terminals, axons, somata or dendrites1. The restriction of Kv channels to high-density clusters in these domains not only affects local membrane signaling events but also affords an opportunity for local modulation of channel function. Studies in cortical and hip pocampal pyramidal neurons have shown the importance of active electrical processing of synaptic input by somatodendritic voltagedependent ion channels2,3. IK is important in regulating somatodendritic excitability in hip pocampal and cortical pyramidal neurons4–6, where the Kv2.1 potas sium channel is a principal component of somatodendritic IK (refs. 5,7–10). In these cells, Kv2.1 is found in large clusters (∼1–2 µm in diam eter) on the soma and on the very proximal portion of the apical and basal dendrites11–13. In layer 5 cortical pyramidal neurons, the restricted localization of Kv2.1 protein is consistent with the decrease in density of dendritic IK that occurs with increasing distance from the soma4,6. Notably, many studies of somatodendritic IK and Kv2.1 in pyramidal neurons suggest that they have a function in regulating excitability and Ca2+ influx during periods of repetitive high-frequency firing4–6,14,15, rather than a more classical role in action potential repolarization. Most studies of activity-dependent changes in neuronal function have focused on local and rapid changes in synaptic strength and on global long-term changes caused by altered gene expression.

However, activity-dependent alterations in the intrinsic electrical properties of neurons, especially in the somal and dendritic com partments that are so crucial to the processing of synaptic input, would also affect neuronal function16. Dynamic activity-dependent changes in the localization and biophysical properties of somato dendritic Kv channels, which have an impact on synaptic integra tion and neuronal excitability, represent an attractive mechanism with which to achieve neuronal plasticity. In this study, we tested whether an increase in activity could affect the localization and biophysical properties of pyramidal cell Kv2.1 channels. High-resolution imaging of cortical and hippocampal pyramidal neurons showed marked changes in Kv2.1 localization in a rat model of seizure. Parallel biochemical studies showed a marked dephosphorylation of Kv2.1 protein in the brains of the seizure rats. Combined immunohistochemical, biochemical and electrophysio logical analyses showed a comparable glutamate-induced change in Kv2.1 localization and phosphorylation in cultured hippocampal pyramidal neurons, and a large change in the biophysical properties of IK. These changes, which occurred through receptor-mediated Ca2+-dependent activation of calcineurin, were both rapid and reversible. These results provide compelling evidence for activitydependent changes in the localization and biophysical properties of somatodendritic ion channels in pyramidal neurons. RESULTS Neuronal activity alters localization of Kv2.1 in vivo We evaluated activity-dependent regulation of Kv2.1 in vivo by induc ing continuous motor seizures (class 5 motor seizures consistent with

1Department of Pharmacology, School of Medicine, University of California, Davis, California 95616, USA. 2Department of Biochemistry and Cell Biology, State University of New York at Stony Brook, Stony Brook, New York 11794, USA. 3Cain Foundation Laboratories, Baylor College of Medicine, Houston, Texas 77030, USA. Correspondence should be addressed to H.M. ([email protected]).

Published online: 13 June 2004; doi:10.1038/nn1260

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Figure 1 Kainate-induced seizures lead to translocation of Kv2.1 in vivo. Rats were injected intraperitoneally with 15 mg/kg of kainate and monitored to assess behavioral seizures. Brain sections from control and kainate-treated rats were stained with antibody to Kv2.1. The projected images are reconstructed from 35 optical sections (thickness 0.4 µm per section) taken from the cortex and subiculum. Insets are highermagnification views of Kv2.1 staining corresponding to the boxed areas in the main images.

continuous electrical seizures or status epilepticus17) in rats through the intraperi toneal injection of kainate (hereafter these rats are referred to as ‘kainate-treated’). Pyramidal neurons in cortex and subiculum of control (vehicle-injected) rats showed typ ical high-density clusters of Kv2.1 on the somatodendritic membrane (Fig. 1). In brain sections from kainate-treated rats, however, Kv2.1 staining was distributed uniformly over the soma and dendrites of pyramidal neurons in these regions, with little evidence of the characteristic Kv2.1 clustering (Fig. 1). Glutamate changes Kv2.1 clustering in cultured neurons Cultured rat hippocampal pyramidal neurons also express Kv2.1 in large clusters on the plasma membrane of the soma and proximal dendrites18. Stimulation with glutamate (10 µM for 10 min), which induces spontaneous bursting activity19, caused translocation of Kv2.1 from clusters to a more uniform distribution on the membrane (Fig. 2a), similar to that observed in the kainate model in vivo. Glutamate treatment significantly reduced the number of cells expressing Kv2.1 clusters and correspondingly increased the number of cells showing diffuse Kv2.1 staining (Fig. 2b). Glutamate-treated neurons showed small clusters (at 2.5 µM glutamate) or diffuse local ization (at 2.5 and 10 µM glutamate) of Kv2.1 rather than the large clusters seen in control cells (Fig. 3a). To assess further the extent of Kv2.1 clustering, the fluorescence intensity of Kv2.1 staining on a 23-µm line across the soma was

Figure 2 Glutamate-induced lateral translocation of Kv2.1. (a) Cultured neurons were incubated with 10 µM glutamate for 10 min at 37 °C. The cells were then fixed and stained for Kv2.1 (green) and MAP-2 (red). Scale bar, 10 µm. (b) The percentage of control and glutamate-treated cells showing clustered and diffuse Kv2.1 staining was each determined by analyzing images collected from 100 cells from four separate coverslips in two independent experiments. Data are the mean ± s.e.m. Note that cells with large Kv2.1 clusters and substantial intercluster staining are scored as both ‘clustered’ and ‘diffuse’.

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plotted18. Control cells yielded large and regular peaks of fluores cence intensity corresponding to Kv2.1 clusters, whereas cells treated with 2.5 or 10 µM glutamate showed less obvious peaks and higher uniform fluorescence (Fig. 3a). A surface plot analysis of flu orescence intensity yielded similar patterns (Supplementary Fig. 1 online). The average number of Kv2.1 clusters with an area greater than 0.25 µm2 was significantly (P < 0.001, n = 20) decreased on cells treated with 10 µM glutamate (5.8 ± 1.4) versus control cells (25.7 ± 1.8; Fig. 3b); however, the total intensity of fluorescence per cell from anti-Kv2.1 antibody staining did not differ between con trol and glutamate-stimulated neurons (Fig. 3c). There were no obvious changes in the localization of postsynaptic density 95 (PSD-95) and F-actin after treatment with 10 µM glutamate for 10 min (data not shown). Live-cell imaging of enhanced green fluorescent protein (EGFP) tagged Kv2.1 in cultured neurons showed that glutamate (10 µM) stimulation caused a rapid fragmentation of the large EGFP-Kv2.1 clusters into smaller microclusters (Supplementary Fig. 2 online). The finding that the EGFP-Kv2.1 was not converted to a diffuse local ization may be due to an inability to convert the whole population of overexpressed protein to a more diffuse localization. Taken together, these results show an activity-dependent regulation of Kv2.1 localiza tion in this in vitro neuronal culture system comparable to that seen after kainate treatment in vivo. Neuronal activity changes the Kv2.1 phosphorylation state The large (440-amino-acid) serine/threonine-rich carboxyl-terminal cytoplasmic tail of Kv2.1 contains the clustering signal18 and more than 20 candidate phosphorylation sites. This suggests that the rapid, activity-mediated translocation of Kv2.1 in brain and cultured neu rons might be mediated through changes in the phosphorylation state of Kv2.1. On SDS-polyacrylamide gel electrophoresis (SDS-PAGE), the major form of Kv2.1 in whole brain and cultured hippocampal neurons showed substantially higher relative molecular masses (Mr) of ∼125 kDa and ∼115 kDa, respectively, than is predicted from the deduced primary sequence (95.3 kDa), and in vitro phosphatase treat

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Figure 3 Glutamate-induced changes in localization of Kv2.1. (a) Neurons were incubated with 2.5 or 10 µM glutamate for 10 min at 37 °C, fixed and stained for Kv2.1. To quantify cluster formation of Kv2.1 in the neurons, the intensity of Kv2.1 staining was measured over a 23-µm segment on the neuronal cell body at the sites indicated by the white lines in the images. Relative intensity values are presented on an arbitrary scale of 0–100 and are plotted against distance (in µm). Scale bar, 10 µm. (b) Glutamate stimulation decreases the number of large Kv2.1 clusters. Neurons were incubated with or without 10 µM glutamate for 10 min at 37 °C, fixed and stained for Kv2.1. Fluorescence images were transformed to binary images by using a fixed threshold of fluorescence intensity, and spots of signals (clusters) were then counted and measured in area. The number of clusters with an area greater than 0.25 µm2 in control and glutamate-treated neurons (20 cells each) was determined. (c) Total fluorescence intensity integrated over the whole soma was measured and expressed as a percentage of the intensity in control neurons. Data are the mean ± s.e.m. (n = 20 cells for each treatment).

ment of extracts prepared from either brain or cultured neuron shifted the Mr of Kv2.1 to ∼100 kDa (Fig. 4a,c). Stimulating neurons with glutamate (10 µM for 10 min) also led to a marked shift in the Mr of the major ∼125-kDa band of Kv2.1 to ∼100 kDa (Fig. 4a). In addition, in vitro phosphatase treatment of the sam ple prepared from glutamate-treated neurons yielded no further elec trophoretic shift in the ∼100-kDa band, and Kv2.1 in both control and glutamate-treated samples electrophoretically comigrated after in vitro phosphatase digestion. Taken together, these data show that the differences in electrophoretic mobility are due to changes in the phos phorylation state of Kv2.1 (Fig. 4a). A significant increase in the dephosphorylated (lower Mr) form of Kv2.1 was also evident in membrane fractions prepared from cor tices, hippocampi and whole brains (Fig. 4b,c) of rats subjected to kainate-induced status epilepticus. The Mr of two other highly phosphorylated Kv channels, Kv4.2 (Fig. 4c) and Kv1.2 (data not shown), was not affected by either kainate-induced status epilepti

cus in vivo (Fig. 4c) or glutamate treatment of cultured neurons (data not shown). Glutamate does not alter surface levels of Kv2.1 We next examined whether the changes in Kv2.1 localization were due to internalization or lateral translocation by incubating control and glutamate-treated neurons with the membrane-impermeant biotiny lation reagent sulfo-NHS-SS-biotin to biotinylate surface proteins. Biotinylated proteins were specifically precipitated with avidinagarose, and the amounts of Kv2.1, GluR1 and β-tubulin in biotiny lated and nonbiotinylated fractions were assessed by immunoblotting. Almost all β-tubulin remained in the nonbiotinylated fraction in samples prepared from both control and glutamate-treated neurons, showing that the overall integrity of the cell membrane was not altered by glutamate treatment (Fig. 4d). Glutamate treatment did not significantly alter the amount of biotinylated Kv2.1 and GluR1 on the neuronal surface (Fig. 4d). In addition, both the ∼125-kDa and

Figure 4 Activity-dependent changes in Kv2.1 phosphorylation in vivo and in vitro. (a) Neurons were incubated with or without 10 µM glutamate (Glu) for 10 min at 37 °C. The cell lysates were incubated with or without 1 U/ml of lambda protein phosphatase (λPP) for 3 h at 30 °C. Proteins were separated by SDS-PAGE and analyzed for Kv2.1 by immunoblotting. Numbers on the left indicate the mobility of molecular weight standards in kDa. (b) Crude membranes were prepared from whole brain or dissected brain regions of control (C) and kainate-treated (K) rats, and analyzed by immunoblotting for Kv2.1. As a control, a fraction of crude whole-brain membranes from control rats was incubated for 2 h with 0.1 U/ml of alkaline phosphatase (AP). Bands of total and phosphorylated (Mr ≈ 125 kDa) Kv2.1 were quantified. Numbers indicate the amount of phosphorylated Kv2.1 as a percentage of total Kv2.1 and are the means from two independent experiments (n = 3 samples each). (c) Crude membranes prepared from the whole brains of rats were incubated without (Mock) or with alkaline phosphatase and analyzed simultaneously for Kv2.1 and Kv4.2. (d) The level of surface Kv2.1 was determined by surface biotinylation assay. Neurons were incubated first with glutamate and then with sulfo-NHS-SS-biotin. Biotinylated proteins were analyzed by immunoblotting for Kv2.1, GluR1 and β-tubulin. Note that four times more eluent from avidin beads (Avidin ppt.) was loaded on the gel as compared with total lysate (Total) and supernatant (Sup.). Avidin bead precipitate without surface biotinylation was used as a negative control (NC).


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Figure 5 Ca2+ influx is essential for glutamate-induced dephosphorylation of Kv2.1. (a) Dose dependence and time course of glutamate-induced dephosphorylation of Kv2.1. Neurons were incubated with 0, 1, 2.5, 5 or 10 µM glutamate for 10 min, or with 10 µM glutamate for the indicated durations. Numbers on the left indicate the mobility of molecular weight standards in kDa. (b) Recovery of Kv2.1 phosphorylation. Neurons were incubated with 10 µM glutamate for 10 min, washed, and then incubated for 30, 60 or 120 min in normal media. The graph shows phosphorylated Kv2.1 (Mr ≈ 125 kDa, arrow) as a percentage of the control value. Data are the mean ± s.e.m. from three independent experiments. (c) Effects of glutamate receptor antagonists. Neurons were incubated with 100 µM AP-5 or 10 µM CNQX for 10 min, and then with 10 µM glutamate for 10 min. Control lanes represent Kv2.1 in crude rat brain membranes without (RBM) or with alkaline phosphatase treatment (AP). (d,e) Role of Ca2+ influx. Neurons were incubated either with 10 µM glutamate (Glu) in the presence or absence of 5 mM EGTA for 10 min at 37 °C (d), or with 200 µM CdCl2 or 10 µM nitrendipine for 10 min, and then with 10 µM glutamate for 10 min (e). (f) Effect of depolarization. Neurons were stimulated with 50 mM KCl for 1, 5 or 10 min.

∼100-kDa forms of Kv2.1 were found in the biotinylated fraction, indicating that glutamate may induce lateral translocation of Kv2.1 on the somatic membrane without affecting its surface expression. Reversible modulation of Kv2.1 The effect of glutamate stimulation on the phosphorylation state of Kv2.1 in cultured neurons was dependent on both dose and time (Fig. 5a). The effects of glutamate were notably rapid, with marked changes in Kv2.1 phosphorylation observed within 5 min of glutamate addition. The glutamate effects were reversible, in that fully phospho rylated Kv2.1 (∼125 kDa) recovered to control levels after glutamate washout (34.7% after glutamate versus 77.2% after washout; Fig. 5b). The effects of glutamate on Kv2.1 clustering were also reversible. In glutamate-treated cultures, 88.6 ± 4.0% of cells showed diffuse Kv2.1 staining, as compared with only 12.8 ± 2.0% of cells in control cul tures. Removal of the glutamate stimulus led to a gradual recovery of clustering, such that the fraction of cells showing diffuse Kv2.1 stain ing decreased to 44.8 ± 8.6% after 1 h of recovery and to 29.7 ± 7.7% after 2 h of recovery. These results show that the glutamate effects on Kv2.1 phosphory lation and clustering are both rapid and reversible, thereby indicating the specific regulation of Kv2.1 phosphorylation as opposed to gen eral effects on altered metabolic state or cell viability. Glutamate acts through ionotropic glutamate receptors To address the involvement of ionotropic NMDA and AMPA-kainate glutamate receptors in the induction of Kv2.1 dephosphorylation, we examined effects of antagonists of these receptors on glutamateinduced dephosphorylation of Kv2.1. Cells were pretreated with 100 µM D-2-amino-5-phosphonopentanoate (AP-5) or 10 µM 6-cyano 7-nitroquinoxaline-2,3-dione (CNQX) for 10 min and then incu bated with 10 µM glutamate. Both AP-5 and CNQX resulted in partial blockade of glutamate-induced dephosphorylation of Kv2.1 (Fig. 5c), and they completely blocked the effect of glutamate when applied

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altogether (data not shown). Using tetrodotoxin (TTX) to block elec trical activity affected neither the glutamate effects nor any of the antagonist effects (data not shown). Taken together, these results suggest that the effect of glutamate on Kv2.1 requires activation of both AMPA-kainate and NMDA recep tors, as previously observed for glutamate-induced changes in phos phorylation of GluR1 (ref. 20). Glutamate effects on Kv2.1 require Ca2+ influx We began to dissect pharmacologically the glutamate response by first removing Ca2+ from the extracellular medium. Removal of Ca2+ completely blocked the effects of glutamate on Kv2.1 dephosphoryla tion, suggesting that Ca2+ influx is necessary for glutamate-induced dephosphorylation of Kv2.1 (Fig. 5d), although the constitutive phos phorylation state of Kv2.1 was not altered. Inclusion of CdCl2 (200 µM) in the extracellular medium also blocked glutamateinduced dephosphorylation of Kv2.1, consistent with a requirement for Ca2+ influx in the signaling pathway (Fig. 5e). However, a specific L-type voltage-gated Ca2+ channel inhibitor (nitrendipine) had no effect on glutamate-induced dephosphorylation of Kv2.1 (Fig. 5e). To determine whether Ca2+ influx alone was sufficient to induce Kv2.1 dephosphorylation, we examined the effects of membrane depo larization in the absence of glutamate stimulation. Depolarization by KCl in the presence of 1 µM TTX was sufficient to induce dephospho rylation of Kv2.1 (Fig. 5f). Treatment with the Ca2+ ionophore A23187 in the presence of TTX also induced dephosphorylation of Kv2.1 (Fig. 6d). Taken together, these results suggest that Ca2+ influx through a non-L-type Ca2+ channel permeation pathway is necessary and suffi cient for glutamate-induced dephosphorylation of Kv2.1. Phosphatases inducing Kv2.1 dephosphorylation To identify the protein phosphatases that induce dephosphorylation of Kv2.1 after glutamate-induced Ca2+ influx, we treated neurons with 0.1 or 1 µM okadaic acid, which potently inhibits the protein


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Taken together, these results suggest that glutamate-induced dephosphorylation of Kv2.1 and lateral translocation of Kv2.1 from a clustered to uniform localization are tightly coupled and occur by means of ionotropic glutamate receptor stimulation, leading to Ca2+-dependent activation of calcineurin. Glutamate stimulation alters neuronal IK We have previously shown that alkaline phos phatase–mediated dephosphorylation of re combinant Kv2.1 expressed in transfected cells modifies its voltage-dependent activa Figure 6 Analysis of the signaling pathway involved in glutamate-induced dephosphorylation and tion, with pronounced (>25 mV) hyperpolar translocation of Kv2.1. (a–c) Effects of phosphatase inhibitors. (a) Neurons were incubated with 0.1 µM okadaic acid (OA) or 10 µM cyclosporin A (CsA) for 10 min, and then with 10 µM izing shifts in the conductance-voltage (G-V) glutamate for 10 min. Numbers on the right or left indicate the mobility of molecular weights relationship of macroscopic Kv2.1 currents22. standards in kDa. (b) Neurons were incubated with 0.1 or 1 µM OA for 10 min and then with We therefore examined whether glutamate 10 µM glutamate for 10 min. (c) Neurons were incubated either with 1 µM deltamethrin (DM) for induced dephosphorylation of Kv2.1 altered 10 min and then 10 µM glutamate for 10 min, or with 5 µM A23187 for 10 min at 37 °C in the the G-V relationship of neuronal IK, the bulk presence of 1 µM tetrodotoxin. Control lanes show Kv2.1 in crude rat brain membranes without of which is mediated by Kv2.1 (ref. 8). (RBM) or with alkaline phosphatase treatment (AP). (d–g) Effects of cyclosporin A, AP-5 and CNQX on glutamate-induced translocation of Kv2.1. Neurons were incubated for 10 min with 20 µM The voltage-dependent activation of IK in cyclosporin A or with 100 µM AP-5 and 10 µM CNQX and then were incubated for 10 min with control neurons, and in neurons before glu 5 µM glutamate. Cells were stained for Kv2.1 (green) and MAP-2 (red). Cells treated with vehicle tamate stimulation, could be fit by a single (d), glutamate (e), CsA and glutamate (f), or AP-5, CNQX and glutamate (g) are shown. Arrows Boltzmann distribution with a half-maximal indicate dendrites with ‘beading’. Scale bar, 10 µm. activation membrane potential (V1/2) of +15.5 ± 0.9 mV and was stable throughout the course of a 10-min control experiment phosphatases PP1 and PP2A. Okadaic acid enhanced the phosphory (data not shown). Glutamate stimulation (10 µM for 10 min) led to a lation of Kv2.1 in unstimulated cultures (Fig. 6a), indicating that rapid and large (>20 mV; Fig. 7a) hyperpolarizing shift in the V1/2 of okadaic acid–sensitive phosphatases are involved in constitutive regu neuronal IK to −8.4 ± 0.5 mV (P < 0.001 versus V1/2 before stimula lation of the Kv2.1 phosphorylation state. Glutamate-induced tion). This glutamate-induced shift in the Ik G-V relationship was dephosphorylation of Kv2.1 occurred even in the presence of 1 µM reversible (V1/2 = −0.47 ± 0.9 mV at 1 h and +9.7 ± 0.5 mV at 2 h after okadaic acid and resulted in the appearance of the dephosphorylated washout; Fig. 7a) and inhibited by the inclusion of calcineurin Kv2.1 forms of ∼115 kDa and ∼100 kDa(Fig. 6b). inhibitors in the patch pipette (V1/2 = +10.7 ± 0.4 mV; Fig. 7b). By contrast, cyclosporin A, an inhibitor of PP2B (also known as cal These fundamental changes in the properties of neuronal IK suggest cineurin), inhibited glutamate-induced dephosphorylation of Kv2.1 that glutamate-induced dephosphorylation of Kv2.1 in cultured pyram but had no effect on the constitutive phosphorylation state of Kv2.1 idal neurons leads to marked changes in its function and localization. (Fig. 6a). Deltamethrin, another calcineurin inhibitor, also inhibited glutamate-induced dephosphorylation of Kv2.1 (Fig. 6c). The KCl DISCUSSION and A23187-induced dephosphorylation of Kv2.1 was also inhibited The subcellular localization of Kv2.1 channels in pyramidal cells is one by cyclosporin A (data not shown), suggesting that calcineurin has an of nature’s more notable examples of the restricted localization of a essential role in glutamate-induced, Ca2+-dependent dephosphoryla membrane protein. Pyramidal cells throughout the cortex and hip pocampus express high levels of Kv2.1 protein, which is invariably found tion of Kv2.1 in pyramidal neurons. in large clusters over the soma and proximal dendrites. The precise mechanism of Kv2.1 clustering is unknown, although a specific target Translocation of Kv2.1 is mediated by calcineurin We next investigated the relationship between glutamate-induced ing signal in the C-terminal cytoplasmic tail of Kv2.1 is both necessary dephosphorylation of Kv2.1 and the marked effects of glutamate on and sufficient for Kv2.1 clustering18,23. In the absence of detailed infor Kv2.1 localization. The majority of neurons treated with glutamate mation on the clustering mechanism, insights into how an increase in (80.6 ± 3.5%) showed diffuse Kv2.1 staining, as compared with a neuronal activity alters Kv2.1 localization are not directly apparent. small minority of cells (8.4 ± 2.9%) in control cultures (Figs 6d,e). The idea that these changes in Kv2.1 localization occur as a result of Most glutamate-stimulated neurons (61.7 ± 1.7%) also showed activity-dependent Kv2.1 dephosphorylation is supported by their typical glutamate-stimulated ‘beading’ of dendrites, as assessed by similar activity dependence in vivo (Figs. 1 and 4) and in culture MAP-2 staining21. (Figs. 2 and 4). Moreover, in cultured neurons the changes in Kv2.1 A combination of the ionotropic glutamate receptor antagonists localization and phosphorylation state exhibit similar glutamate dose AP-5 and CNQX significantly reduced the percentage of glutamate- responses (Figs. 3 and 5), kinetics (Fig. 5 and Supplementary Fig. 2 stimulated cells with diffuse Kv2.1 staining (25.8 ± 7.8%) and com online) and pharmacology (Fig. 6). Notably, the Kv2.1 clustering sig pletely eliminated dendritic beading (Fig. 6g). The calcineurin nal is located in a serine- and threonine-rich cytoplasmic tail of Kv2.1, inhibitor cyclosporin A also significantly inhibited glutamate- and the clustering signal itself contains three serine residues that dis induced translocation of Kv2.1 (24.8 ± 4.6% cells showed diffuse rupt clustering when individually mutated to alanine. Numerous examples of phosphorylation-dependent clustering Kv2.1; P < 0.001); however, most cyclosporin A–treated cells (66.4 ± of channels and receptors have been reported. Clustering of the 1.3%) still showed glutamate-induced dendritic beading (Fig. 6f).


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ARTICLES GluR2 AMPA receptor subtype at synapses is regulated by phosphorylation of a spe cific serine residue on the GluR2 cytoplas mic tail24. Phosphorylation of this serine by protein kinase C interferes with GluR2 binding to the PDZ domain–containing clustering protein GRIP and leads to GluR2 binding to PICK1, resulting in receptor internalization and long-term depression25. Kir2.3 K+ channels26, β-adrenergic recep tors27, cystic fibrosis transmembrane con ductance regulator (CFTR) channels28 and acid-sensing ion channels29 also show phosFigure 7 Neuronal activity alters the properties of I current in neurons. (a) Inset, representative I phorylation-sensitive binding to PDZ currents in a cultured hippocampal neuron recordedK under whole-cell patch clamp. Scale bars, 25Kms domain–containing proteins, although (horizontal); 2 nA (vertical). The membrane potential was held at −80 mV and depolarized to +50 mV phosphorylation-dependent changes in the in 10-mV increments for 200 ms. A 30-ms pre-pulse of −10 mV was given before each test localization of these proteins in wild-type depolarization. The main plot shows the G-V relationship of peak IK currents recorded from neurons cells have not been reported. In contrast to with (filled circles) or without (open circles) glutamate stimulation, and neurons incubated for 2 h after these established examples, Kv2.1 does not glutamate washout (squares). Data are the mean ± s.e.m. (n = 5). (b) IK currents were recorded from neurons before (open circles) and after (filled circles) glutamate stimulation, and after stimulation in contain a consensus PDZ domain–binding the presence of cyclosporin A and cyclophilin A (1 µM each) in the internal solution (squares). The motif, and phosphorylation seems to be calcineurin inhibitors themselves did not affect I currents during the course of experiments. K positively correlated with clustering. Thus, Cyclophilin A was included to compensate for loss of the endogenous molecule through dialysis. any phosphorylation state–specific interac tion of Kv2.1 with clustering proteins would be predicted to occur through a mechanism distinct from those of brane Kv2.1 clusters in pyramidal neurons lie over subsurface cisthe established examples. ternae13intracellular endoplasmic reticulum–derived memWe found that marked changes in the phosphorylation state and branes rich in inositol triphosphate and ryanodine receptors. These localization of Kv2.1 in pyramidal cells were coupled to a significant specialized sites, where intracellular Ca2+ stores come into close modulation of the biophysical properties of neuronal IK. The voltage- apposition with the plasma membrane, represent a specialized neu dependent activation of Kv2.1 is regulated by the cytoplasmic C ter- ronal signaling domain that may also contain increased amounts of minus30 via altered phosphorylation of Kv2.1 (ref. 22). The hyper- voltage-dependent Ca2+ channels32. polarizing shifts in macroscopic Kv2.1 currents on treatment with Calcineurin binds to both inositol triphosphate and ryanodine alkaline phosphatase22 and in neuronal IK on stimulation with gluta- receptors33, suggesting a possible mechanism for an indirect Kv2.1 mate (our current results) are of similar magnitude (∼25 mV) and are calcineurin association. Although the specific role of Kv2.1 at these consistent with the biochemically observed dephosphorylation of sites is not yet clear, antisense knockdown of Kv2.1 expression in hip Kv2.1. Such large shifts in activation would be expected to affect the pocampal neurons has profound effects on dendritic intracellular active electrical properties of pyramidal cell somata and proximal Ca2+ transients5. The activity-dependent translocation of Kv2.1 dendrites, especially during periods of high-frequency firing when observed here would lead to a selective uncoupling of Kv2.1 from Kv2.1 and/or IK has a crucial role5. these domains, which could affect the local driving force for transOur data indicate that the effects of glutamate on Kv2.1 localization membrane Ca2+ fluxes in the restricted space where the plasma mem in cultured pyramidal neurons may be mediated by calcineurin- brane and subsurface cisterns come into close apposition and could dependent dephosphorylation. An increase in intracellular Ca2+, prevent Ca2+ overload and excitotoxicity. Kv2.1 channels are also either through activation of ionotropic glutamate receptors or implicated in oxidant- and staurosporine-induced apoptotic signalthrough membrane depolarization by KCl or ionophore, is sufficient ing cascades in mammalian neurons10; thus, activity-dependent to induce dephosphorylation and translocation of Kv2.1. Blocking dephosphorylation of Kv2.1 could affect neuronal survival. Ca2+ influx with Ca2+-free medium or CdCl2 also inhibits the glutaThere are many examples of neuromodulation of neuronal ion mate-induced effects. An increase in intracellular Ca2+ in pyramidal channel function34, whereas few examples of dynamic changes in the cell dendrites has diverse effects on protein phosphorylation, and not localization of voltage-dependent ion channels have been reported. It all intracellular Ca2+ may act in the same manner31. Several dendritic is clear that dynamic activity-dependent changes in the localization protein kinases are activated directly or indirectly by an increase in and clustering of synaptic AMPA receptors underlie many of the intracellular Ca2+, which serves as a major mechanism for both short- changes associated with both long-term potentiation and long-term depression in hippocampal pyramidal neurons35. In addition, and long-term activity-dependent changes in neuronal function. Protein phosphatases such as calcineurin, PP1 and PP2A can be increases in intracellular Ca2+ lead to the physical synapse-to-nucleus also activated directly or indirectly by increased intracellular Ca 2+, translocation of signaling molecules that regulate diverse aspects of such that potentially competing but spatially segregated kinase and neuronal function36. Our data provide a unique example of activityphosphatase reactions can be simultaneously mobilized by an dependent changes in the localization and function of a somatoden increase in neuronal activity. Our results show that an increase in dritic Kv channel in pyramidal neurons. In addition, they offer a intracellular Ca2+ results in rapid and efficient calcineurin- previously unknown mechanism for cellular plasticity, in which an dependent dephosphorylation of Kv2.1, suggesting that Kv2.1 clus- increase in intracellular Ca2+ leads to changes in the input-output ters may be localized at or near membrane-associated pools of relationships of pyramidal neurons through phosphorylation calcineurin. Electron microscopic analyses show that plasma mem- dependent translocation of ion channels.

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ARTICLES METHODS Materials. All chemicals, unless stated otherwise, were purchased from Sigma. Lamda phosphatase was obtained from NEB. Okadaic acid, cyclosporin A, deltamethrin and nitrendipine were purchased from Calbiochem.


Cell culture. Hippocampal cultures were prepared on coverslips as described37,38. For neurons on plastic dishes, we changed the media to serumfree astrocyte-conditoned media. Cells that had been cultured for 10–18 d were used in the experiments. Live cell imaging. A EGFP-Kv2.1 complementary DNA37 was used to transfect neurons at 8 d in culture using Lipofectamine2000 (Invitrogen) as described18. After 3 d, EGFP-Kv2.1 was visualized under epifluorescence illumination by using a 24-bit color digital camera installed on an Axiovert 200M microscope (Zeiss) with a 63×, 1.3 NA lens and a CO2 incubation stage coupled to Axiovision software (Zeiss). Biochemical analysis of neuronal proteins. Neurons were washed twice with Hank’s balanced salt solution and incubated with drugs as indicated in the fig ure legends. For surface biotinylation, cells were incubated with 2.5 mg/ml of sulfo-NHS-SS-biotin (Pierce) for 1 h at 4 °C. The cells were then washed twice with ice-cold Locke’s solution (154 mM NaCl, 5.6 mM KCl, 2.3 mM CaCl2, 1 mM MgCl2, 5 mM glucose and 5 mM HEPES, pH 7.4), collected and cen trifuged at 12,000g for 30 min at 4 °C. The pellets were extracted by adding sample buffer (2% SDS, 1 mM EDTA, 10% glycerol, 0.001% bromophenol blue, 5% β-mercaptoethanol and 62.5 mM Tris-HCl, pH 6.8) and sonicating briefly. We concentrated biotinylated proteins on avidin-agarose beads and eluted them by adding sample buffer. Rat brain membranes were prepared and treated with purified phosphatases as described12,39. Proteins were separated on 7.5% SDS-PAGE gels, transferred to nitrocellu lose membrane and immunoblotted with mouse monoclonal antibody K89/41 to Kv2.1 (ref. 37), a mouse monoclonal antibody to Kv4.2 (ref. 40), or a rabbit polyclonal antibody to GluR1 (Upstate Biotechnology). The blot was then incubated with horseradish peroxidase (HRP)-conjugated secondary antibod ies (ICN), followed by enhanced chemiluminescence reagent (Perkin Elmer). We visualized immunoreactive bands by exposing the blot to X-ray films. Immunoreactive bands were quantified after scanning by National Institutes of Health (NIH) Image software (ImageJ) with a gel-plotting macro program. Quantification was done with samples from at least three different cultures. We compared the mean intensity of signals statistically by Student’s t-test. Immunofluorescence staining of neurons. Neurons were incubated with drugs as indicated in the figure legends, and then fixed and stained with anti Kv2.1 rabbit polyclonal8 and anti–MAP-2 (Sigma) mouse monoclonal anti bodies and Alexa-conjugated isotype-specific secondary antibodies as described19. Fluorescent images were taken with a 24-bit color digital camera installed on a Axiscop 2 microscope with a 100×, 1.4 NA lens coupled to Axiovision software (Zeiss). Images were transferred to PhotoShop software (Adobe) as JPEG files. Quantitative analyses of Kv2.1 clustering were done on the raw images by Image software. In the line scan analyses, background fluorescence has been subtracted from each value, and data are shown as a percentage of the maxi mum values in each plot. To count the number of large clusters, we converted images to binary images by using a fixed threshold of fluorescence intensity. The clusters were then counted and measured in the binary images by NIH Image. Surface plot analyses were done by NIH Image with a macro program. Preparation of seizure model. All animal use procedures were done in strict accordance with the Guide for the Care and Use of Laboratory Animals described by the National Institutes of Health. Rats were injected with 15 mg per kg (body weight) of kainate dissolved in saline or the vehicle. The progres sion of seizure was assessed by recording the behavioral seizure stage after injection as described17: class 1, mouth and facial movements; class 2, head nodding; class 3, forelimb clonus; class 4, rearing; class 5, rearing and falling. A full behavioral seizure, with loss of postural control, was considered as a class 5 motor seizure. Rats showing class 5 seizure were subjected to biochemical and immunohistochemical analysis.


For immunoblotting, brains were kept whole or dissected into hippocampi and cortices, and a crude membrane fraction was prepared12. For immunohis tochemical analyses, sagittal brain sections were prepared from rats fixed by perfusion of 4% paraformaldehyde. The sections were blocked with 5% goat serum and stained with rabbit antibody to Kv2.1 (ref. 8) and an Alexa-conju gated secondary antibody (Molecular Probes). Fluorescent images were taken as 35 optical sections (0.4 µm thickness) with a 24-bit color digital camera installed on a Axiovert 200M microscope with a 63×, 1.3 NA lens and an ApoTome coupled to Axiovision software (Zeiss). Images were reconstituted as a projected image on the software. Electrophysiology. Current recordings were made with the whole-cell patchclamp configuration. Electrodes (0.8–2 MΩ) were pulled from borosilicate glass tubing, heat-polished and filled with pipette solution (140 mM KCl, 5 mM EGTA, 2 mM MgCl2, 10 mM glucose and 10 mM HEPES, pH 7.3). Currents were recorded with an EPC-10 patch-clamp amplifier (HEKA Elektronik), sampled at 10 kHz and filtered at 2 kHz by a digital Bessel filter. All currents were capacitance- and leak-subtracted by a standard P/n procedure. The external solution contained 0.1 µM tetrodotoxin, 140 mM NaCl, 5 mM KCl, 2 mM CaCl2, 2 mM MgCl2, 10 mM glucose and 10 mM HEPES, pH 7.3. For the recovery experiments, we stimulated cells by bath application of 10 µM glu tamate for 10 min. Currents were recorded from cells incubated with or without glutamate stimulation, and from cells incubated for 2 h after glutamate washout. For the calcineurin inhibition experiments, cyclosporin A and cyclophilin A (1 µM each) were included in the pipette solution, and glutamate (10 µM) was applied to neurons for 10 min with a polytetrafluorethylene glass multiple-bar rel perfusion system. Currents were recorded from a cell before and after gluta mate stimulation. All recordings were done at room temperature (23–25 °C). The membrane potential was held at −80 mV and depolarized for 200 ms to +50 mV in 10-mV increments. Before the start of the test depolarization, a sin gle pre-pulse to −10 mV was given for 30 ms. The peak current amplitude (I) at each test potential was converted into conductance (G) using the equation G = I/(V − EK). The Nernst potassium ion equilibrium potential EK was calculated as −84 mV. The normalized conductance G were plotted against the test poten tial (V) and fitted to a single Boltzmann equation G = Gmax/(1 + exp[–(V – V1/2)/k]), where Gmax is the maximum conductance, V1/2 is the test potential at which the channel has half-maximal conductance, and k is the parameter that represents the slope of the activation curve. We used an unpaired Student’s ttest to evaluate the significance of changes in mean values. Note: Supplementary information is available on the Nature Neuroscience website. ACKNOWLEDGMENTS We thank A. Illausky for technical assistance; G. Mandel for allowing use of equipment; P. Brehm for chemical reagents; L. Taylor for assistance in live-cell imaging; and A.C. Bonham, J. Engebrecht, M.N. Rasband and K.J. Rhodes for critically reading the manuscript. This work was supported by the National Institute of Neurological Disorders and Stroke (NIH/NINDS; grants NS42225 to J.S.T. and NS39943 to A.E.A.). COMPETING INTERESTS STATEMENT The authors declare that they have no competing financial interests. Received 5 April; accepted 4 May 2004

Published online at http://www.nature.com/natureneuroscience/

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