Protein kinase C modulates NMDA receptor trafficking and gating

© 2001 Nature Publishing Group http://neurosci.nature.com

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Protein kinase C modulates NMDA receptor trafficking and gating Jian-yu Lan1, Vytenis A. Skeberdis1, Teresa Jover1, Sonja Y. Grooms1, Ying Lin1, Ricardo C. Araneda1,2, Xin Zheng1, Michael V. L. Bennett1 and R. Suzanne Zukin1 1Department of Neuroscience, Albert Einstein College of Medicine, 1300 Morris Park Avenue, Bronx, New York 10461, USA 2Present address: Department of Biological Sciences, Columbia University, New York, New York 10027, USA

Correspondence should be addressed to ([email protected])

Regulation of neuronal N-methyl-D-aspartate receptors (NMDARs) by protein kinases is critical in synaptic transmission. However, the molecular mechanisms underlying protein kinase C (PKC) potentiation of NMDARs are uncertain. Here we demonstrate that PKC increases NMDA channel opening rate and delivers new NMDA channels to the plasma membrane through regulated exocytosis. PKC induced a rapid delivery of functional NMDARs to the cell surface and increased surface NR1 immunofluorescence in Xenopus oocytes expressing NMDARs. PKC potentiation was inhibited by botulinum neurotoxin A and a dominant negative mutant of soluble NSF-associated protein (SNAP-25), suggesting that receptor trafficking occurs via SNARE-dependent exocytosis. In neurons, PKC induced a rapid delivery of functional NMDARs, assessed by electrophysiology, and an increase in NMDAR clusters on the surface of dendrites and dendritic spines, as indicated by immunofluorescence. Thus, PKC regulates NMDAR channel gating and trafficking in recombinant systems and in neurons, mechanisms that may be relevant to synaptic plasticity.

Protein kinase C (PKC) is implicated in NMDAR-dependent longterm pontentiation (LTP) and may contribute to enhanced synaptic efficacy at hippocampal CA1 synapses1–7. Micro-injection of active PKC into the postsynaptic cell enhances synaptic transmission3, whereas block of PKC activity by intracellular delivery of PKC inhibitors blocks induction of LTP6. Moreover, mice with targeted deletion of the PKCγ gene exhibit greatly diminished LTP7. Activation of PKC increases NMDA-elicited currents and channel open probability in neurons, and potentiates NMDA currents in Xenopus oocytes expressing recombinant NMDARs8–11. Activation of a number of G-protein-coupled receptors, including phosphoinositol-coupled metabotropic glutamate receptors12,13, µ opioid receptors14 and muscarinic receptors15, potentiates NMDA receptors via activation of PKC (but see ref. 16). The molecular mechanisms underlying PKC potentiation of NMDARs are as yet uncertain. Target sites of kinase-induced phosphorylation have been identified on the NR1, NR2A and NR2B subunits17,18. However, NMDARs assembled from mutant subunits lacking all known sites of PKC phosphorylation show marked PKC potentiation11. Thus, potentiation is likely to occur through phosphorylation of receptor-associated protein(s) involved in signaling and/or trafficking. PKC modulates channel activity not only by changes in intrinsic channel properties, but also by regulation of receptor/channel trafficking. PKC promotes insertion of Ca2+ channels into the plasma membrane of Aplysia bag cell neurons19 and causes internalization of GABAA and GABAC receptors expressed in HEK-293 cells and Xenopus oocytes20,21. Here we examined the hypothesis that PKC potentiation of NMDARs expressed in Xenopus oocytes and in hippocampal neurons occurs at least in part by rapid recruitment by means of exocytosis of vesicle-asso382

ciated channel molecules to the cell surface. Xenopus oocytes expressing recombinant receptors provide geometric simplicity and express a homogenous population of receptors, presumably in the absence of PSD-95, that inhibit PKC potentiation22. Moreover, the molecular machinery for protein trafficking is highly conserved from yeast to mammals23. Hippocampal neurons in culture provide a physiological milieu in which to examine mechanisms of kinase-regulated receptor gating and trafficking. Experiments involving patch-clamp and whole-cell recording, western blot analysis of cell surface proteins, and immunofluorescence reveal that PKC increases NMDA channel opening rate and delivers new NMDA channels to the plasma membrane of oocytes and neurons by regulated exocytosis, mechanisms that may be relevant to synaptic plasticity.

RESULTS TPA increases channel number times open probability To examine effects of PKC activation on NMDA channel gating and trafficking, we recorded channel activity in outside-out patches excised from oocytes expressing NR1-4b/NR2A receptors before and after treatment with 12-O-tetradecanoyl phorbol-13-acetate (TPA), a PKC activator. NR1-4b has the shortest C-terminal tail, and NR1-4b/NR2A and -B have the highest cell surface expression24 and highest degree of PKC potentiation11. In control patches, NMDA (10 µM) activated channels with a single channel conductance (58 ± 1 pS at –60 mV), which did not vary with voltage (Erev ≈ 0, n = 5; Fig. 1a and d). NMDA channel activity in patches excised after application of TPA (100 nM, 10 min) was markedly potentiated (Fig. 1b). TPA increased the number of active channels times channel open probability, npo, by ∼6.7-fold from 0.023 ± 0.004 before TPA (n = 5) nature neuroscience • volume 4 no 4 • april 2001



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Fig. 1. The PKC activator TPA potentiates NMDA single a b c channel activity. (a, b) Representative traces of NMDAactivated channels recorded in outside-out patches excised from control (a) and TPA-treated (b) oocytes expressing NR1-4b/NR2A receptors. Single-channel currents were elicited by application of NMDA at a low concentration (10 µM with 10 µM glycine) directly to the patch, and recorded at Vh = –60 mV. (c) Quantitation of npo measured in outside-out patches from control and TPA-treated oocytes. TPA induced an increase in npo from 0.023 ± 0.004 d e f in control oocytes (n = 5) to 0.154 ± 0.017 in TPA-treated oocytes (n = 5; p < 0.001). (d) NMDA single-channel current–voltage (I–V) relationship recorded in outside-out patches excised from control (white circles) and TPAtreated (black circles) oocytes. The white circles largely superimpose on the black circles. Single-channel currents were measured from each patch at potentials from –100 to +60 mV, in 20 mV steps. PKC activation did not alter NMDA single-channel conductance (58 ± 1 pS before TPA, i h n = 5, versus 58 ± 1 pS after TPA, n = 5) or reversal poteng tial (Erev ≈ 0 before and after TPA, n = 5 for both). (e, f) Distribution of channel open times in outside-out patches from control (e) and TPA-treated (f) oocytes (representative fits). The distribution of open times was fit by a single exponential consistent with the presence of a single open state. TPA did not alter NMDA channel mean open time (τ = 7.89 ± 1.6 ms in control oocytes, n = 5, versus τ = 7.69 ± 0.24 ms in TPA-treated oocytes, n = 5). (g–i) Representative traces illustrating NMDA-activated l j k channels in outside-out patches excised from oocytes before (g) and after (h) application of TPA to the cell. To elicit maximal channel activity, saturating NMDA (1 mM with 10 µM glycine) was rapidly applied directly to the patch. TPA increased the maximal number of simultaneously open channels per patch, n, measured in outside-out patches excised after application of TPA. (i) The increase in the number of simultaneously active channels was to –5 times that of patches excised before application of TPA (2.2 ± 0.2 for control, n = 15, versus 10.2 ± 1.3 for TPA, n = 15, p < 0.001). Scale bars, 10 pA, 100 ms. (j–l) Representative traces illustrating NMDA-activated channels in cell-attached patches from control (j) and TPA-treated (k) oocytes. Single-channel activity was elicited by inclusion of NMDA (100 µM, 10 µM glycine) in the pipette. Cellattached patches formed before (j, upper trace in k) and after (lower trace in k) TPA application to the same oocyte. TPA application after patch formation increased NMDA channel activity (npo) in the same patch to –1.3 times that before TPA (n = 4, p < 0.01, paired t-test, j, upper trace in k, l). In a new cell-attached patch formed on the same oocyte after TPA application, NMDA channel activity was increased to –5.5 times that in control patches formed before TPA (n = 4, p < 0.001, j, lower trace in k, l).

to 0.154 ± 0.017 after TPA (n = 5; p < 0.001; Fig. 1c). PKC activation did not alter NMDA single-channel conductance or reversal potential, as evidenced by comparison of current–voltage (I–V) relationships (n = 5, Fig. 1e and f). Moreover, TPA did not significantly change the mean duration of openings (Fig. 1e and f). The distribution of open times was fit by a single exponential consistent with the presence of a single open state (Fig. 1e and f; τ = 7.89 ± 1.6 ms before TPA, n = 5, versus τ = 7.69 ± 0.24 ms after TPA, n = 5). Application of TPA to a patch excised from a control oocyte resulted in no detectable potentiation, consistent with a requirement for cytoplasmic elements and/or membrane fusion. In whole-cell measurements using PKC inhibitors, potentiation was blocked by staurosporine and reduced by Ro-32-0432; potentiation was not elicited by the inactive phorbol ester 4α-PDD (data not shown). To estimate the effect of PKC on channel number, we repetitively applied brief pulses of saturating NMDA (1 mM) to patches excised from individual oocytes before and after TPA treatment. In 15 of 15 oocytes, patches excised after TPA exhibited an approximately 5-fold increase in the number of simultaneously open channels per patch at the peak of the response relative to patches nature neuroscience • volume 4 no 4 • april 2001

excised from control oocytes (n = 2.2 ± 0.2 before TPA, n = 15, one patch per cell, versus n = 10.3 ± 1.2 after TPA, n = 15, p < 0.001; Fig. 1g–i). A key to evaluating whether the observed increase in npo involves an increase in n is the channel open probability in control patches; reported po values vary widely from 0.024 measured by whole-cell recording from neurons25 and oocytes (po = 0.04, see below) to 0.3 measured in outside-out patches25,26. Assuming po = 0.3 in control patches, the maximal value measured in excised outside-out patches25, we could not account for the entire increase in npo solely by a change in channel gating. These findings raise the possibility that PKC may promote insertion of new channels and/or unmask silent channels at the cell surface. If PKC potentiation involved delivery of receptors to the cell surface, disruption of exocytosis by formation of a cell-attached patch27 would be expected to reduce the potentiation. Application of TPA after formation of a cell-attached patch increased NMDA channel npo measured in the same patch by ∼1.3-fold (from 0.75 ± 0.07 before TPA, Fig. 1j, to 1.10 ± 0.13 after TPA, n = 4, p < 0.03, Fig. 1k, upper trace and Fig. 1l, left). In contrast, NMDA channel activity recorded in a new cell-attached patch formed on 383



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Fig. 2. PKC increases NMDA channel opening a b c rate and the number of active channels at the oocyte surface. (a) Whole-cell recordings were obtained from Xenopus oocytes expressing NR1-4b/NR2A receptors in Ca2+-free (Ba2+) Ringer’s. TPA (100 nM, 10 min), applied to oocytes via the bath perfusate, potentiated whole-cell currents elicited by application of NMDA (1 µM with 50 µM glycine). Vh = –60 mV. (b) TPA increased the number of functional d e f g h NMDA channels expressed at the cell surface (N). Currents were elicited by application of NMDA (1 mM NMDA with 50 µM glycine) in the continuous presence of the open channel blocker MK-801 (5 µM) from control (left) and TPA-treated (right) oocytes at a holding potential of –60 mV. The NMDA-inward current increased to a peak value, and then decayed exponentially because of MK-801 block of NMDA channels as they opened. The cumulative charge transfer, Q, which is the total current flow during the time interval for complete block by MK-801, was obtained by integration of the current trace over time (area indicated by shading). The larger integrated current in TPA-treated oocytes indicated increased number of functional channels per cell. (c) Agonist-evoked currents in (b) were normalized to the same peak amplitude for comparison of the time course of decay. The more rapid decay of the NMDA current in TPA-treated oocytes indicates increased rate of channel opening, kβ. (d–h) Quantitation of data in (a–c). (d) Potentiation of NMDA whole-cell current, ITPA/Icontrol, was to 4.8 ± 0.6 times control (n = 5). (e) For control and TPA-treated oocytes, N was normalized to the NMDA-elicited whole-cell current, which corrects for any differences in levels of expression in the two groups. Number of functional channels expressed at the surface of TPA-treated oocytes was ∼3.5 times that of control oocytes (p < 0.01; n = 5 for control oocytes; n = 5 for TPAtreated oocytes). (f) Ratio of open probabilities was 1.3. (g) Opening rates, kβ, for control and TPA-treated oocytes. (h) Closing rates, kα, for control and TPA-treated oocytes.

the same oocyte after PKC activation (Fig. 1k, lower trace and Fig. 1l, right) exhibited ∼5.5-fold potentiation, as observed in patches excised after TPA. Evidently, formation of a gigaohm seal, which can distort the membrane inside the pipette and disrupt exocytosis27, greatly reduced PKC potentiation. PKC increases channel opening rate and number The results reported thus far suggest that PKC increases NMDA channel open probability and/or number of active channels at the cell surface. To analyze independently the effects of PKC on number of functional channels in the membrane, N, and channel open probability, po, we took advantage of the essentially irreversible block of NMDA-elicited currents by the open-channel blocker MK-801 (refs. 25, 26). We recorded NMDA whole-cell currents in the continuous presence of MK-801 (5 µM) in control (Fig. 2b, left) and TPA-treated (Fig. 2b, right) oocytes. To determine N, we calculated cumulative charge transfer, Q, which is the total current flow during the time required for complete block by MK-801. N can be calculated from Q as follows.

N = Q/[γ (V – Erev) tbl ] Here, tbl is the time constant for MK-801 block (tbl = 1/(kbl [MK-801])) and kbl = 2.5 × 107/M/s (ref. 26). For a control oocyte exhibiting an NMDA whole-cell current of 100 nA, the mean number of channels, Ncontrol, was 1.2 ± 0.3 × 106, and the mean channel density was 0.04 µm–2, assuming a surface area for the oocyte of 3 × 107 µm2 (ref. 28). TPA does not detectably alter single channel conductance or mean open time (Fig. 1d–f) and should not affect kbl. For control and TPA-treated oocytes, N was normalized to the initial NMDA-elicited whole-cell current, which corrects for any differences in levels of expression within and 384

between the two groups. Thus, we calculate that PKC activation increased the number of channels per cell by ∼3.5-fold (NTPA/Ncontrol = 3.5; p < 0.005), a value that accounted for a major component if not all of the increase in whole-cell current (I TPA/Icontrol = 4.8 ± 0.6, n = 5). In control and TPA-treated oocytes, po can be calculated from the near steady state current at the peak of the NMDA-elicited whole-cell current, I, the single channel current, i, and N, as follows. po, control = Icontrol/iNcontrol = 0.044 ± 0.006 po, TPA = ITPA/iNTPA = 0.057 ± 0.006 From these equations, the ratio po, TPA/po, control is 1.3. These findings indicate that PKC increased the number of active NMDA channels in the oocyte surface; the small increase in channel open probability was not significant. As an independent measure of the effect of PKC on NMDA channel gating, we analyzed the rate of decay of NMDA-elicited current in the presence of MK-801 (Fig. 2c). Provided that kbl[MK-801] >> kα and kβ (the closing and opening rates, respectively), the decay can be described by a single exponential with the rate constant, k β. Activation of PKC increased k β (from 0.99 ± 0.07/s for control oocytes to 1.90 ± 0.22/s for TPA-treated oocytes, p < 0.01; Fig. 2g). This finding indicates that PKC increases NMDA channel opening rate. From measurements of p o and k β, and the relationship po = kβ/(kα+ kβ), we calculated values of kα = 22 ± 4/s in control oocytes versus 33 ± 8/s in TPA-treated oocytes. This difference was not significant. These values of kα are substantially less than the theoretical value of ∼125/s predicted from the ∼8-ms mean open time of single channels (Fig. 2e and f). The apparent discrepancy nature neuroscience • volume 4 no 4 • april 2001


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Fig. 3. PKC promotes delivery of NMDA channels to the oocyte membrane via exocytosis. Microinjection of the light chain of botulinum toxin type A, BoNT A (50 ng), an enzyme that cleaves SNAP-25 into oocytes 5 h before recording reduced PKC potentiation of NMDA-elicited currents by about 50%. NMDA and glycine as in Fig. 2a. (a) Representative NMDA-elicited whole-cell currents before and after TPA application in an oocyte loaded with BoNT A. (b) BoNT A reduced the degree of PKC potentiation from 7.3 ± 0.9 (n = 5) in control oocytes to 3.9 ± 0.3 in BoNT A-treated oocytes (n = 5, ∗∗p < 0.01). (c) BoNT A reduced the PKC-induced increase in NMDA channel number. The ratio of NTPA/Ncontrol was reduced from 6.3 (n = 4) for control oocytes to 3.0 (n = 5,∗ p < 0.05) for oocytes loaded with BoNT A. N was normalized to the control responses of the control oocytes and of the TPA treated oocytes, which corrects for any differences in levels of expression in the two groups.

arises because the po equation applied to macroscopic current assumes an equilibrium between unliganded and bursting states, and thus will yield a k α value corresponding to termination of bursting rather than of individual openings. Calculated in this manner, kα will be overestimated in that po calculated from N and I is less than the probability that a receptor is liganded.

PKC promotes exocytosis of NMDARs To distinguish between insertion of new channels and unmasking of silent channels, we did two additional experiments. First, we examined the effects of loading oocytes with the light chains of type A botulinum neurotoxin (BoNT A), which inactivates SNAP-25, and prevents SNAP-25-dependent exocytosis27. Treatment of oocytes with BoNT A reduced the degree of TPA potentiation of NMDA-elicited currents by about 50% (Fig. 3a and b). BoNT A also reduced the TPA-induced increase in NMDA channel number (Fig. 3c). In contrast, loading the light chains of BoNT B, which inactivates the v-SNARE synaptobrevin27, did not affect potentiation (Fig. 3b). No significant change in the resting potential, input resistance or basal NMDA response was caused by BoNT A or B (data not shown). No effects on TPA potentiation were observed when the vehicle (DTT, 10 mM) was injected. Presumably, toxin treatment attenuated the number of new channels inserted by TPA. Second, because the usual target of BoNT A is SNAP-25, we examined whether PKC potentiation could be inhibited by a dominant-negative mutant of SNAP-25. SNAP-25(∆20), a truncation mutant lacking the C-terminal 20 amino acids, corresponds to yeast sec9-∆17, a dominant negative mutant of the yeast SNAP-25 homolog27. The SNAP-25 mutant was expressed in oocytes by injection of cRNA 48 hours after injection of NMDAR cRNAs. PKC potentiation was measured in oocytes about 20 hours after injection of wild-type SNAP-25 or SNAP-25(∆20). TPA induced about a sevenfold potentiation of NMDA currents in control oocytes (n = 3; Fig. 4a and d) and in oocytes expressing full-length SNAP-25 (n = 4; Fig. 4b and d), but only a 2.49 ± 1.45-fold potentiation in oocytes expressing SNAP-25(∆20) (n = 5, p < 0.01; Fig. 4c and d). Expression of the dominant negative mutant of SNAP-25 did not affect the NMDA-elicited current or endogenous Ca2+-activated Cl– currents27. These findings indicate that

NMDA channels inserted at the cell surface by TPA are delivered by SNAP-25-mediated form of SNARE-dependent exocytosis. The results reported thus far do not exclude a contribution from reduction in internalization. As an estimate of the rate of constitutive exocytosis in the oocyte, we measured re-appearance of functional channels at the cell surface after quasi-irreversible block by MK-801. MK-801 (1 µM) completely blocked the NMDA-elicited current (Fig. 5b, first response). A test application of NMDA at 10 min elicited a small, transient response, indicative of a small number of newly inserted channels that were opened by agonist and then blocked by residual MK-801 (Fig. 5b, second response). The small size of the test response is indicative of a relatively slow rate of constitutive exocytosis and is consistent with the half-time of –40 min reported for recovery of the acetylcholine response after irreversible block of nicotinic acetylcholine receptors expressed in Xenopus oocytes29. Because under steady-state conditions the rate of internalization equals the rate of exocytosis, and the rate of exocytosis is slow (Fig. 5b), inhibition of internalization (or of silencing) could account for, at most, a very small component of PKC potentiation. As a further test of PKC-induced recruitment of new channels to the cell surface, we blocked NMDA currents with MK-801 and then applied TPA in the presence of MK-801 for 10 min (Fig. 5c). Following wash-out of TPA and MK-801, a test application of NMDA elicited a much larger response, ∼16 times that observed in control oocytes at 10 min after MK-801 block without TPA (p < 0.001, Fig. 5d) and ∼6 times that of the control response (p < 0.01; Fig. 5d). If PKC potentiation were due entirely to insertion of a new channel molecules (Fig. 2), and constitutive insertion were negligible, the amplitude of the test NMDA current after TPA application in the presence of MK-801 b (ITPA, MK-801) would equal the amplitude of the test NMDA current after TPA alone (ITPA) minus the amplitude of the initial control current (Iconc trol). ITPA, MK-801 did not differ significantly from the predicted value (Fig. 5d). These findings demonstrate

d Fig. 4. A dominant negative mutant of SNAP-25 reduces PKC potentiation of NMDARs. (a) PKC potentiation of NMDA-elicited currents in oocytes expressing NR1-4b/NR2A receptors as in Fig. 2a. (b) Co-expression of wild-type SNAP-25 with NMDARs had no effect on PKC potentiation. (c) Co-expression of SNAP-25(∆20), a dominant negative mutant of SNAP-25, with NMDARs markedly reduced PKC potentiation. (d) Quantitation of the effects of wild-type and mutant SNAP-25 on PKC potentiation. nature neuroscience • volume 4 no 4 • april 2001

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Fig. 5. PKC-regulated exocytosis is fast relative to constitutive exocytosis of NMDARs. Whole-cell recordings were obtained from Xenopus oocytes expressing NR1-4b/NR2A receptors as in Fig. 2a. The open-channel blocker MK-801 (1 µM) was used to estimate the rate of delivery of functional channels to the cell surface in control and TPA-treated oocytes. NMDA, 300 µM; glycine, 10 µM. (a) In control oocytes, TPA (100 nM, 10 min) induced a –6-fold potentiation of NMDA-elicited currents. (b) MK-801 applied during the steady-state response to NMDA completely blocked the NMDA current. Ten minutes after block and wash-out of NMDA, MK-801 was removed; a test application of NMDA then elicited a very small response (–10% of control), due to either recovery of a small number of channels from block or insertion of new channels. (c) Experimental procedure as described in (b), except that oocytes were incubated with TPA and MK-801 for 10 min. A test application of NMDA elicited a much greater response, –16 times that after MK-801 block in the absence of TPA (compare responses to the second NMDA application in Fig. 5b and c), an increase too great to be accounted for by a 6-fold potentiation of unblocked NMDA channels (Fig. 5a). The test responses were transient, which is ascribable to block by residual MK-801. (d) Quantitation of (a–c); for (b) and (c), test responses were normalized to initial currents. Bar 1, ITPA/Icontrol; bar 2, ITPA, MK-801/IMK-801; bar 3, ITPA, MK-801/Icontrol. ITPA is the NMDA-elicited current after TPA; ITPA, MK-801 is the current after block of the control response by MK-801 and potentiation by TPA; IMK-801 is the current after block of the control response by MK801 followed by 10-min recovery.

that PKC causes a rapid increase in the number of functional NMDARs in the cell surface. PKC increases surface expression of NMDARs in oocytes We directly measured changes in NR1 subunit surface expression by two approaches. First, we did western blots on cell-surface proteins30. Intact control and TPA-treated oocytes expressing NR1/NR2A receptors were surface labeled with sulfo-NHS-SSbiotin, and biotinylated surface proteins were separated from non-labeled intracellular proteins by reaction with Neutravidin beads. Surface and total cell proteins were treated with DTT (which releases the biotin moiety from surface proteins), subjected to electrophoresis and probed with monoclonal antibody 54.1 directed to the extracellular loop of the NR1 subunit31 (Fig. 6a). Analysis of band densities indicated an increase in surface NR1 expression to 2.8 ± 0.2 times control (p < 0.01; n = 3), with no change in total cell NR1 protein (Fig. 6b). Because the NR1 surface labeling of TPA-treated oocytes lay outside the calibration range, TPA potentiation calculated as the ratio of the two values may be an underestimate. The finding of an increase in surface NR1, but not in total cell NR1, argues against the possibility that the increase in active channels at the cell surface occurs via synthesis of new receptor subunits. Second, we assessed the effect of PKC on receptor surface expression, by immunofluorescence. Control and TPA-treated non-permeabilized oocytes expressing NR1-4b/NR2A receptors were reacted with antibody 54.1, followed by a fluorescein-tagged secondary antibody. A Z-series of cross-sectional images through the oocyte was viewed by confocal microscopy and captured by COMOS (Bio-Rad). Control oocytes expressing NMDARs exhibited clear immunofluorescence, which appeared confined to the external surface in cross-sectional and tangential images (Fig. 7a and c). TPA dramatically increased NR1 surface immunolabeling (Fig. 7b and d). Quantitation of images showed increase in fluorescence to 4.2 ± 0.25 times that in control oocytes (n = 8 control oocytes; n = 8 TPA-treated oocytes; p < 0.001). To address the issue of whether TPA might increase permeability of the oocyte membrane to antibody and thereby increase immunolabeling, we used an antibody directed to an epitope on the intracellular C-terminal domain of the NR1 subunit. Intact oocytes exhibited little or no immunofluorescence with the C-terminal antibody (Fig. 7e and f). In contrast, after permeabilization with Triton-X100, both control and TPA-treated oocytes exhibited intense immunofluorescence (Fig. 7g and h). Moreover, TPA treatment did not detectably alter the leak current of the oocyte 386

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(data not shown), a finding that argues against TPA-induced increase in d membrane permeability. Little or no immunofluorescence was observed for oocytes reacted with FITCtagged secondary antibody in the absence of primary antibody (Fig. 7i) or for water-injected oocytes labeled with the N-terminal antibody (Fig. 7j); these findings indicate specificity of the immunofluorescence labeling. PKC delivers new NMDARs to the surface of neurons To examine PKC-regulated NMDAR trafficking in neurons, we used three experimental approaches. First, we performed wholecell recording on dissociated E18 hippocampal neurons and measured surface expression of functional channels using the quasi-irreversible blocker MK-801. In control neurons, NMDAelicited currents exhibited run-down; the peak response decreased by –20% in 10 min (Fig. 8a). Loading of neurons with PKM (the constitutively active fragment of PKC) potentiated the NMDA peak response by 1.78 ± 0.02-fold (n = 7 for control and for PKM, p < 0.02, Fig. 8b). To determine N in control and PKMloaded neurons, we measured the cumulative charge transfer Q from the total NMDA-elicited current flow during block by MK-801 (40 µM; Fig. 8c). We calculated that delivery of PKM into neurons increased the number of channels per cell by about twofold (NPKM/Ncontrol = 1.9; p < 0.01; Fig. 8e), a value that accounted for virtually all of the increase in whole-cell current; PKM did not significantly alter NMDA channel open probability, po = ∼0.003 or opening rate, kβ = ∼1.4/s (Fig. 8d and e). Second, we examined the effect of PKM on exocytosis of NMDARs in neurons using MK-801. In control neurons, application of MK-801 (40 µM), as in Fig. 5, completely blocked the NMDA response (Fig. 8f, first record). A test application of NMDA at 10 min elicited a very small response (< 5% of the initial, control current), indicative of the relatively slow constitutive exocytosis of NMDA channels at the neuronal plasma membrane (Fig. 8f, second record). We next loaded neurons with PKM and again blocked the NMDA-elicited current with MK-801. A test application of NMDA at 10 minutes in these neurons elicited a markedly greater response (∼3.6 ± 0.4 times the small response observed in the absence of PKM (n = 6; Fig. 8g nature neuroscience • volume 4 no 4 • april 2001


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Fig. 6. Activation of PKC increases NR1 abundance at the cell surface. NR1 surface and total cell expression in control and TPA-treated oocytes, as assessed by western blot analysis of biotinylated surface proteins. (a) Representative western blot of surface protein from oocytes expressing NR1-4b/NR2A receptors probed with anti-NR1 antibody 54.1 (ref. 31). Lanes 3, 6 and 12 indicate micrograms of protein in samples of total cell extract before Neutravidin bead extraction loaded on each lane; ‘surface’ indicates an aliquot of Neutravidin bead-isolated proteins. TPA increased NR1 abundance in samples of surface protein. (b) Density was linearly related to total protein (data normalized to give the same slope for each gel). Surface protein abundance in control (white circles) versus TPA-treated oocytes (black circles). TPA treatment increased surface expression to 2.8 ± 0.2 times control (n = 3, p < 0.01).

DISCUSSION and h), indicative of increase in the number of functional channels in the presence of activated PKC. ITPA, MK-801 was smaller than predicted, presumably due to release of a low level of endogenous glutamate, which enabled MK-801 block of a fraction of NMDA channels during the development of potentiation (Fig. 8g). Third, we assessed the effect of TPA (together with the phosphatase inhibitor okadaic acid) on NMDAR surface expression in hippocampal neurons by immunofluorescence. Control (untreated), okadaic acid-treated (100 nM) and TPA/okadaic acid-treated (100 nM of each) intact neurons were reacted with NR1 antibody. A Z-series of optical images through dendrites was acquired by confocal microscopy. Control neurons exhibited light immunofluorescence in cell somata and intense punctate labeling indicative of receptor clusters along the length of their proximal and distal dendrites; the labeling in dendrites appeared confined to the external surface (Fig. 9a, c and e). TPA (with okadaic acid) increased NR1 labeling on the surface of proximal dendrites by 2.12 ± 0.26-fold (p < 0.001; TPA-treated neurons, n = 28; control neurons, n = 18; Fig. 9b, d and f). Okadaic acid enhances TPA potentiation in hippocampal neurons32. Okadaic acid alone did not significantly alter NR1 labeling (n = 21). The number of receptor clusters defined as non-contacting regions of fluorescence was not significantly altered, indicating that average size of receptor clusters was enlarged. Intact neurons labeled under the same conditions for the intracellular protein synapsin exhibited no detectable labeling, whereas when permeabilized, neurons exhibited intense punctate immunofluorescence at presumed sites of synaptic contact (data not shown). These findings indicate that PKC recruits new NMDA channels to the cell surface of neurons, presumably via SNARE-dependent exocytosis.

Fig. 7. TPA increases NR1 surface immunofluorescence. Oocytes expressing NR1-4b/NR2A receptors were incubated in external recording solution in the presence or absence of TPA (100 nM, 10 min) and subjected to immunocytochemistry. Surface fluorescence was expressed as the mean intensity of fluorescence per unit area. Representative cross-sectional and tangential images of oocytes expressing NR1-4b/NR2A receptors from control (a, c) and TPA (b, d) treatment groups. In TPA-treated oocytes (n = 8), fluorescence intensity was 4.2 times that observed in control oocytes (n = 8, p < 0.001). Experiments with an antibody directed against an epitope in the intracellular C-terminal domain of the NR1 subunit and nonpermeabilized oocytes show little or no labeling before or after TPA (e, f). After permeabilization with Triton-X100, immunolabeling with the C-terminal Ab was intense in control (g) and TPA-treated (h) oocytes; fluorescence intensity did not differ significantly in the two groups. Control oocytes labeled by secondary antibody in the absence of primary antibody (i), and water-injected oocytes labeled as in (a–d) showed negligible fluorescence (j). Scale bar, 40 µm. nature neuroscience • volume 4 no 4 • april 2001

The present study demonstrates that PKC activation increases NMDA channel opening rate and recruits new channel molecules to the cell surface by regulated exocytosis. Our previous studies involving truncation and deletion mutants of NMDARs indicate that one or both of these mechanisms occurs indirectly via phosphorylation of receptor-associated protein(s)11. Together, the two studies suggest that PKC-induced insertion of NMDARs involves phosphorylation of a protein involved in receptor trafficking. One potential target of PKC is SNAP-25, which can be directly phosphorylated by PKC in an activity-dependent manner33,34. We show that loading cells with BoNT A, which interferes with exocytosis by cleaving SNAP-25, markedly reduces PKC potentiation of NMDAR responses. Moreover, expression of a dominant negative mutant of SNAP-25 greatly reduces PKC potentiation. A simple scenario is that PKC regulates membrane fusion events of NMDAR-containing vesicles by phosphorylation of SNAP-25. The association of SNAP-25 or its binding partners with NMDARs, however, remains to be established. Specificity of PKC action on NMDARs is indicated by the observation that receptors containing NR2C do not show PKC potentiation35. The physiological relevance of PKC actions is underscored by our observation that PKM recruits new functional channels to the surface of hippocampal neurons.

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Fig. 8. PKC promotes increases in functional NMDA channels in the a b plasma membrane of hippocampal neurons. To estimate the half-time for receptor turnover and the increase in number of functional channels at the cell surface of neurons induced by PKC activation, we performed whole-cell recordings on dissociated E18 hippocampal neurons maintained in culture for 14–21 days. NMDA, 100 µM; glycine, 10 µM; PKM, 2 µM; MK-801, 40 µM. (a) With control solution in the pipette, NMDA-elicited currents declined by 20% over 10 min. (b) Inclusion of PKM in the pipette induced ∼1.7-fold potentiation of the peak current c d e by 10 min. (c) PKM increased N. Currents were elicited by application of NMDA in the continuous presence of the quasi-irreversible blocker MK-801 from control (left) and PKM-loaded (right) neurons at a holding potential of –60 mV. The NMDA-elicited current increased to a peak value, and then decayed exponentially due to MK-801 block of NMDA channels as they opened. The cumulative charge transfer, Q, was measured from the integrated currents. The larger integrated current in PKM-loaded neurons indicated increased N. (d) PKM did not significantly alter the rate of decay of the NMDA current in the presence of f g h MK-801, indicating no detectable change in channel opening rate. (e) Quantitation of data in (a–c). PKM increased the NMDA-elicited whole-cell current and number of functional NMDA channels expressed at the cell surface; channel open probability calculated as for Fig. 3f was not significantly changed. (f) With control solution in the pipette, application of MK-801 with agonist completely blocked the NMDA-elicited current. Ten minutes after block and wash-out of NMDA, MK-801 was washed out; a test application of NMDA elicited a very small response (–5% of control), due to insertion or recovery of a small number of new channels. (g) Experimental procedure as in (f), with PKM in the recording pipette. After wash-out of MK-801, a test application of NMDA elicited a much larger response than the test response in (f), indicating insertion or recovery of new receptors. (h) Quantitation of data in (a), (b), (f) and (g). NMDA-elicited currents after MK-801 were normalized to the initial currents before block. The ratio of NMDA-elicited currents after block by MK-801 with and without PKM was much larger than that predicted by PKC potentiation. These findings indicate that PKC increases the number of active channels in the cell surface.

In this study, we observed an ∼6-fold potentiation of NMDA currents by TPA in oocytes and an ∼2-fold potentiation by PKM in neurons. Potentiation of neuronal NMDARs by PKC varies from ∼1.2-fold in embryonic hippocampal neurons8,10 to more than 2-fold in postnatal (P15 to P22) hippocampal neurons32. Moreover, activation of mGluR1 potentiates NMDA currents –4-fold in postnatal rat striatal neurons, an effect which is likely to be PKC-mediated36. The higher potentiation observed in striatal neurons may be due to the fact that, unlike hippocampal neurons, striatal neurons express high levels of NR1-4a and 4b splice variants, which exhibit highest PKC potentiation in oocytes11. In this study, PKC potentiation was about 50% inhibited by BoNT A (Fig. 3). This inhibition was relatively specific, as surface expression of endogenous Ca2+-activated Cl– channels, endogenous voltage-gated Ca2+ current and transfected epithelial Na+ channels and NMDARs in Xenopus oocytes are not reduced (data not shown; see also ref. 27). Moreover, the time course and efficacy of the inhibitory action on PKC potentiation by BoNT A are similar to that in a number of other systems. The maximum inhibition by BoNT A of Isoc current in oocytes27 of insulin-stimulated glucose uptake in adipocytes37 and of neurotransmission at Aplysia synapses38 is in the range of 40% to 50%, similar to the maximal reduction of NMDAR potentiation observed in our experiments. The incomplete block by BoNT A in these systems could be for any of a number of reasons: BoNT A might not cleave the endogenous SNAP-25 in these cell types efficiently, other trafficking proteins might ‘substitute’ for SNAP-25 when it is inactivated, and/or the cleaved SNAP-25 is still partially active. Our findings suggest a mechanism whereby PKC could regulate cell surface expression of NMDARs at the postsynaptic membrane, thereby modulating neuronal excitability. Turnover studies of NMDARs in cultured cerebellar granule cells indicate that there is a large intracellular pool of NR1 subunits (∼60% 388

total NR1) in contrast to NR2A and NR2B subunits, which are largely found in surface membrane. These observations suggest that in neurons, unassembled NR1 subunits are present in large numbers in dendritic shafts and spines, and that assembly of NR1 with NR2 subunits occurs immediately before insertion into the postsynaptic membrane39. Our observations of PKCinduced regulation of NMDAR trafficking in oocytes and in hippocampal neurons are consistent with the observation that the molecular machinery for secretion is conserved from yeast to mammals23. We demonstrate rapid insertion of NMDARs in the cell membrane of oocytes and neurons, presumably via a SNARE-dependent mechanism. However, rapid trafficking of AMPARs between the plasma membrane and an intracellular compartment under conditions of normal synaptic transmission40–42 and during synaptic plasticity43 is well established. Regulation of protein trafficking by kinases, and, by implication, phosphatases, is likely to be a universal mechanism critical for maintenance of the precise and highly ordered organization of the postsynaptic membrane. Kinase and phosphatase signaling complexes are strategically localized together with ionotropic glutamate receptors at PSDs by means of receptor-associated proteins. The PKC family of serine-threonine kinases and the src family of tyrosine kinases potentiate NMDAR activity9,10,32; the corresponding phosphatases depress NMDAR activity44,45. Our finding that PKC increases NMDA channel opening rate is consistent with observations that PKC increases NMDA channel open probability in neurons8 and in HEK-293 cells expressing recombinant NMDARs10. Our finding of PKC-induced delivery of new channel molecules to the plasma membrane via exocytosis demonstrates regulation of NMDAR trafficking by PKC in oocytes expressing recombinant NMDARs and in neurons. There is evidence that membrane fusion events contribute to NMDA receptor-dependent LTP46, but the relative contribution of NMDA nature neuroscience • volume 4 no 4 • april 2001



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and AMPA receptors is controversial4. Moreover, persistent activation of PKC is required during the maintenance phase of LTP4. Although activity-dependent targeting of NMDARs to postsynaptic sites occurs on the order of many hours or days47, experience-dependent insertion of new NMDARs into the membrane in the visual cortex can be achieved within one to two hours and may underlie rapid modification in synaptic strength48. Findings from the present study suggest that PKC may phosphorylate one or more proteins involved in receptor recycling and targeting, thereby increasing NMDAR density at synapses. In this manner, insertion and retrieval of NMDA receptors to and from the plasma membrane may be common and powerful mechanisms for regulating excitatory synaptic transmission. Given that PKC and NMDA receptors are widely expressed throughout the central nervous system, regulation by PKC of NMDA channel activity provides a potentially important way to modulate efficacy of synaptic transmission, long-term potentiation (LTP), and long-term depression (LTD) and alter the LTP/LTD modification threshold48.

METHODS Expression constructs. We cloned rat NR1-4b (NR1100) cDNA; mouse ε1 (corresponding to rat NR2A) cDNA was a gift of M. Mishina (Tokyo, Japan). SNAP-25 and SNAP-25 (∆20) cDNAs were gifts of the HHMI Investigator R. Y. Tsien (Howard Hughes Medical Institute, San Diego, California). Capped mRNAs were synthesized as run-off transcripts from linearized plasmid cDNAs with T3 or T7 polymerase (Ambion mMessage mMachine Transcription Kit, Austin, Texas)11. Culture and recording of hippocampal neurons. Dissociated cell cultures of hippocampus were prepared from rat embryos day 18 (E18) as described by Banker and Goslin49. Briefly, dissociated neurons were attached to poly-L-lysine coated coverslips and co-cultured with astrocytes in defined media supplemented with N2. Neurons were used after 14–21 days in culture. Electrophysiological recording from hippocampal neurons was done on the stage of an inverted phase-contrast microscope at room temperature. Pipette resistance ranged from 2 to 4 MΩ with an internal solution consisting of 140 mM Cs methyl SO4, 11 mM EGTA, 1 mM CaCl2, 2 mM MgCl2, 10 mM HEPES, 2 mM TEA, 4 mM K2ATP and 0.0001 mM leupeptin (pH 7.3). Cells were bathed in an nature neuroscience • volume 4 no 4 • april 2001

Fig. 9. PKC activation increases NR1 surface immunofluorescence in hippocampal neurons. Representative images of longitudinal sections through the center of proximal dendrites of untreated neurons (a, c, e) and neurons treated with TPA (100 nM) in the presence of okadaic acid (100 nM) for 10 min (b, d, f). Cells treated with TPA and okadaic acid show markedly increased surface immunofluorescence relative to that of control cells or cells treated with okadaic acid alone. Scale bar, 4 µm (g) Quantitation of surface NR1 surface immunofluorescence at 10 min in response to control, okadaic acid (100 nM) and TPA (100 nM) in the presence of okadaic acid (100 nM). **p < 0.005 compared with control or okadaic acid alone.

external solution containing 140 mM NaCl, 1.3 mM CaCl2, 5 mM KCl, 25 mM HEPES, 33 mM glucose and 0.0005 mM tetrodotoxin (pH 7.4). Test solutions containing NMDA (100 µM with 10 µM glycine) were delivered to the patch from a multibarrel array fed by gravity. Expression of recombinant NMDA receptors in Xenopus oocytes. Selected Stage V and VI oocytes from adult female Xenopus laevis (Xenopus I) were injected with a mixture of in vitro transcribed mRNAs (–20 ng mRNA/cell; NR1:NR2, 1:2). Whole-cell currents were recorded from oocytes (two to six days after injection) as described11. Single-channel currents were recorded from outside-out patches excised from devitellinized oocytes. Single-channel current amplitudes were determined from means of Gaussian fits to all-point amplitude histograms. We calculated npo as the total channel open time divided by recording time. Open time durations were calculated from single channel openings above baseline, but presumably include events from two or more channels in the same patch. All data are presented as mean ± s.e.m. for 4–15 experiments done with different oocytes. Statistical significance was assessed by the Student’s t-test (SigmaPlot 3.0, Chicago, Illinois). Western blot analysis of surface proteins. Xenopus oocytes expressing NR1-4b/NR2A receptors were washed twice and surface proteins were biotinylated with the membrane-impermeable reagent sulfosuccinimidyl 2-(biotinamido) ethyl-1,3´dithiopropionate (sulfo-NHS-SS-biotin; Pierce, Rockford, Illinois)30. Cell extracts were prepared50. To isolate biotinylated surface proteins from non-surface proteins, cell extracts were incubated with Neutravidin-linked beads (Pierce) for 2 h at 4°C, centrifuged and washed. Bound proteins were eluted from beads by incubation with SDS-PAGE gel loading buffer containing DTT (which releases the biotin moiety from labeled proteins), and subjected to gel electrophoresis. Cell surface immunolabeling of oocytes and neurons. Xenopus oocytes expressing NR1-4b/NR2A receptors and hippocampal neurons were labeled with monoclonal antibody 54.1 directed to the extracellular loop of the NR1 subunit31. Oocytes were screened for NMDA currents in the range of 100 to 300 nA and TPA potentiation of ∼8-fold. Oocytes were incubated in 20 ml external recording solution in the absence or presence of TPA (100 nM, 10 min), devitellinized, and fixed in 4% paraformaldehyde/2% sucrose (1 h at RT). Oocytes were then incubated with NR1 Ab (10 µg/ml; 4° C overnight) and reacted with biotinylated horse anti-mouse IgG (Vector Laboratories, Burlingame, California; 10 µg/ml; 1.5 h at RT) followed by FITC-conjugated avidin (Vector Laboratories; 10 µg/ml; 1.5 h at RT). Oocytes were rinsed, placed in a chamber 1.5 mm deep with a coverslip forming the bottom (MatTek Corp.), and covered with ProLong mounting medium (Molecular Probes, Eugene, Oregon) to reduce fluorescence quenching. Cross-sectional and tangential images of oocytes were viewed by a Bio-Rad MRC 600 Kr/Ar laser scanning confocal microscope and acquired using COMOS software (Hercules, California). FITC fluorescence was measured (excitation wavelength, 488 nm; emission, 515 nm). A 40× objective lens (numerical aperture 1.3) was used; laser intensity, photomultiplier gain and pinhole aperture were kept constant. Dissociated hippocampal neurons isolated from E18 rat embryos were maintained on glass coverslips for 14–21 days as described above. Batches of neurons were screened for NMDA currents in the range of 100 to 300 pA and PKC potentiation of 1.3- to 2.0-fold. Neurons were incubated in external recording solution in the absence or presence of okadaic acid (100 nM) or TPA (100 nM) with okadaic acid (100 nM) for 10 min at 35°C and subjected to immunocytochemistry. Immediately 389



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after incubation, control (untreated), okadaic acid-treated and TPA-treated neurons were fixed in paraformaldehyde (4%), blocked with horse serum (10%), and incubated with NR1 Ab (5 µg/ml;1 h at RT), followed by biotinylated secondary antibody and FITC-conjugated avidin as above. Images were viewed by a confocal microscope and acquired as above. For quantitation of immunofluorescence intensity, a Z-series of optical images through proximal dendrites of control (n = 18), okadaic acid-treated (n = 21) and TPA/okadaic-acid treated (n = 28) neurons was acquired with a Bio-Rad MRC 600 Kr/Ar laser scanning confocal microscope and processed using IP Lab Spectrum software (Scanalytics, Fairfax, Virginia) running on a Macintosh G3. Exposure times were kept constant and below gray-scale saturation to permit a linear response to light intensity. We analyzed regions of interest (ROIs) taken from longitudinal images selected from the center of the Z series for 2–6 proximal neurites per neuron. The ROI was defined as the entire cross-sectional area of the dendrite between the soma and a point–20 µm distal to the soma. The fraction of pixels exhibiting fluorescence signal and the number of receptor clusters, defined as contiguous regions of fluorescence, were measured using a thresholding algorithm.

ACKNOWLEDGEMENTS The authors thank G. Bassell and D. Faber for their helpful comments on the manuscript and A.P. Wang and M. Martinez for technical support. We acknowledge the Analytical Imaging Facility of the Albert Einstein College of Medicine (M. Cammer, Director). This work was supported by NIH grants NS 20752 and NS 31282 (to R.S.Z.) and NS 07512 (to M.V.L.B.). M.V.L.B. is the Sylvia and Robert S. Olnick Professor of Neuroscience.

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