Soluble Oligomers of Amyloid-β Peptide Disrupt Membrane Trafficking of α-Amino-3-hydroxy-5-methylisoxazole-4-propionic Acid Receptor Contributing to Early Synapse Dysfunction^*

Primary Neuronal Culture

Neurons were cultured from E14.5–15.5 C57BL/6 wild-type mouse embryo forebrains or embryos of heterozygous APP_Sw,Ind x non-transgenic crossings. Cells were enzymatically and mechanically disrupted in the presence of trypsin and DNase I before plating in poly-d-lysine (100 μg/ml)-coated 24-well plates, 35- to 60-mm dishes, or on coverslips. Cells were seeded at a density of 5 × 10⁴ cells/cm² in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum (FBS, heat inactivated), 50 units/ml penicillin, 50 μg/ml streptomycin, 2 mm glutamine, and 30 mm glucose. Three hours after seeding, medium was replaced with serum-free Neurobasal medium supplemented with 2% B27 (Invitrogen), 50 units/ml penicillin, 50 μg/ml streptomycin, 2 mm glutamine, 30 mm glucose, which yielded nearly pure neuronal cultures (21). Culture medium was partially replaced every 3–4 days with fresh Neurobasal supplemented with B27. Cell cultures were kept at 37 °C in a humidified incubator with 5%CO₂/95%air, and neurons were used for experiments after 12–14 days in vitro.

Aβ Oligomer Preparation

Synthetic Aβ_1–42 (Bachem, United Kingdom) was dissolved in hexafluor-2-propanol and kept at −80 °C after evaporation of hexafluor-2-propanol. Aβ oligomers (oAβs) were prepared freshly by dissolving the peptide film with DMSO and cold F-12 medium without phenol red to yield a 100 μm stock as previously described (22). Samples were incubated at 4 °C for 24 h and centrifuged at 14,000 × g for 10 min at 4 °C. The supernatant containing a mixture of oAβ was biochemically analyzed by SDS-PAGE and electron microscopy. For negative staining analysis, 5 μl of the sample was placed on copper grids covered with carbon and counterstained with 2% uranyl acetate, using the droplet technique, and examined in a JEOL JEM-2011 transmission electron microscope.

Cell Stimulation and Lysate Preparation

Cultures were first incubated in ACSF for 30 min at room temperature (in mm): 125 NaCl, 2.5 KCl, 1 MgCl₂, 2 CaCl₂, 33 d-glucose, and 25 HEPES (pH 7.3), followed by stimulation with 5 μm oAβ in ACSF (no MgCl₂). After 10 min of stimulation, neurons were replaced in regular ACSF and then subjected to different procedures at indicated time points. For total lysate preparation, Cultures were then washed once with ice-cold PBS and scraped in cold 1% Nonidet P-40 homogenization buffer (in mm: 20 Tris, pH 7.5, 150 NaCl, 5 EDTA, 1 PMSF, 1 Na₂VO₄, 1 × Sigma protease inhibitor and phosphatase inhibitor cocktails) to obtain cell lysates. Mouse hippocampi were lysed in 0.2 ml of cold lysis buffer (50 mm Tris-HCl, pH 7.4, 150 mm NaCl, 2 mm EDTA, 0.5% Triton X-100, 1% Nonidet P-40, 0.1% SDS, 1 mm Na₃VO₄, 50 mm NaF, and 1 mm PMSF) supplemented with protease and phosphatase inhibitors. Tissue was homogenized during 20 s using a pellet pestle (Sigma) and kept 1 h at 4 °C. Lysates were centrifuged at 12,000 × g for 10 min at 4 °C, and the protein in the supernatant was quantified by a DC protein assay kit based on the Bradford method (Bio-Rad Laboratories, Inc.).

Immunoblotting

Primary antibodies were: anti-phospho-Ser-831-GluA1, anti-phospho-Ser-845-GluA1, anti-GluA1 (1:1,000, Santa Cruz Biotechnology, Santa Cruz, CA); anti-phospho-Ser-845-GluA1, anti-GluA1, and anti-GluA2 (1:1,000, Millipore); anti-phospho-Ser-880-GluA2 (1:1,000, Abcam); anti-Calcineurin A and B (1:1,000, BD Bioscience); anti-GAPDH (1:40,000, Ambion Inc.); and anti-β-amyloid (1:1,000, 6E10, Signet). Samples were separated on 7.5 or 10% SDS-PAGE and transferred onto Hybond-C Extra, nitrocellulose membranes (Amersham Biosciences). Blots were blocked at room temperature for 1 h with 10% dry milk, 0.1% BSA (fraction V), pH 7.4, in PBS and incubated at 4 °C overnight with primary antibody in PBS 0.1% BSA, pH 7.4. After washing, blots were then incubated with horseradish peroxidase-conjugated secondary antibodies diluted in blocking buffer and developed using the ECL^TM Western blotting Detection Reagents (Amersham Biosciences). Semi-quantitative analysis of immunoblots was performed by densitometry using ImageJ (National Institutes of Health, Bethesda, MD), and protein levels were corrected for corresponding loading control.

Surface Biotinylation

After oAβ stimulations, cultured neurons were transferred to ice-cold PBS-Ca²⁺-Mg²⁺ buffer (pH 7.4, 1 mm CaCl₂, 0.1 mm MgCl₂), followed by biotinylation in 1 mg/ml biotin (EZ-Link Sulfo-NHS-SS-Biotin, Pierce) for 30 min with slow agitation. Free biotin was quenched by 3× wash in cold PBS-Ca²⁺-Mg²⁺ + glycine (0.1 m). Cell cultures were immediately scraped in cold 1% Triton X-100 homogenization buffer (in mm: 50 NaCl, 10 EDTA, 10 EGTA, 1 Na₃VO₄, 50 NaF, 25 NaPP_i, 1β-glycerophosphate, 1 PMSF, 1 × protease inhibitor mixture, 1 × phosphatase inhibitor mixture, and 50 HEPES, pH 7.5). Solubilization was performed in 1% Triton X-100 to avoid solubilization postsynaptic densities (23) and is, therefore, selective for extrasynaptic AMPARs. Homogenates from cultures were centrifuged at 10,000 × g for 20 min to pellet insoluble fraction. 75 μl of the supernatant was mixed and heated with 25 μl of 4× SDS sample buffer to determine total fraction of GluA1 (surface plus internal). Biotinylated surface proteins in the remaining supernatant (∼225 μl) were pulled down with 40 μl of 50% avidin-agarose beads (ImmunoPure Immobilized Avidin, Pierce) overnight at 4 °C. The beads were pelleted, and 75 μl of the supernatant (internal fraction) was mixed and heated with 25 μl of 4× SDS sample buffer. The beads were then rinsed three times with 1% Triton X-100 homogenization buffer and heated in 100 μl of 2× SDS sample buffer (surface fraction). Equal volumes of the total, internal, and biotinylated fractions were subjected to 10% SDS-PAGE, probed for total GluA1, and normalized to GAPDH.

Immunoprecipitation

Neurons were washed in ice-cold PBS and immediately scraped in cold 1% Nonidet P-40 homogenization buffer (400 μl/2 × 60-mm plate). Homogenates from cultures were centrifuged at 10,000 × g for 10 min to pellet insoluble fraction. 75 μl of the supernatant was mixed and heated with 25 μl of 4× SDS sample buffer (total homogenate). The remaining supernatants (∼300 μl) were immunoprecipitated with 1 μg of anti-GluA1 (Millipore) overnight at 4 °C. Incubations continued for 1 h at 4 °C in the presence of 40 μl of 50% slurry protein-G-Sepharose beads (Amersham Biosciences). The beads were pelleted, and 75 μl of the supernatant (unbound fraction) was mixed and heated with 25 μl of 4× SDS sample buffer. The beads were then rinsed three times with 1% Nonidet P-40 homogenization buffer and heated in 100 μl of 2× SDS sample buffer (immunoprecipitation fraction). Equal volumes of each fraction were detected by immunoblotting.

Calcium Imaging

Primary neurons grown onto poly-lysine-coated coverslips for 12 days were loaded with Fura-2/AM (4 μm, Molecular Probes, Invitrogen) for 1 h at room temperature. Coverslips were washed with ACSF buffer and mounted in a static chamber at room temperature on an inverted Nikon TE2000U microscope. Cells were excited alternatively at 340 and 380 nm using a monochromator (Cairn Research Ltd.), and emission light was collected at 510 nm every 4 or 20 s. Images were acquired by using a 12-bit CCD ERG ORCA Hamamatsu camera and processed with Metafluor (Universal Imaging). When appropriated, cells were treated with oAβ (5 μm) in ACSF (no MgCl₂). N > 50 cells were analyzed in each experiment (three independent experiments were performed). Data were analyzed with Excel (Microsoft, Seattle, WA) and Prism (GraphPad, San Diego, CA) software.

Calcineurin Activity

Calcineurin activity was determined with the calcineurin cellular activity assay kit (Calbiochem). Briefly, neurons were homogenized in lysis buffer (25 mm Tris-HCl, pH 7.5, 0.5 mm dithiothreitol, 50 μm EDTA, 50 μm EGTA, and 0.2% Nonidet P-40). Free phosphate was eliminated using a desalting column, and an equal amount of protein was incubated with the calcineurin substrate RII phosphopeptide (1.64 mg/ml) for 30 min at 30 °C. The reaction was stopped by adding 100 μl of GREEN TM reagent, and fluorescence was measured at 620 nm using a microtiter plate reader.

APP Transgenic Mice and Morris Water-maze Test

APP_Sw,Ind transgenic mice (line J9, C57BL/6 background) expressing mutant human APP695 isoform harboring the FAD-linked Swedish (K670N/M671L) and Indiana (V717F) mutations under the neuronal PDGFβ promoter have been previously described (24). Mice were age-matched littermate males obtained by crossing heterozygous APP_Sw,Ind x non-transgenic (WT) crossings. The Morris water-maze test was performed as previously described (25, 26). Experimenters of the behavioral tests were blind to the genotypes of the mice. Animal procedures were performed in accordance with institutional and national guidelines following approval by the Animal Care and Ethical Committee (CEEAH) of the Universitat Autònoma de Barcelona.

Statistical Analysis

Statistical analysis of the biochemical experiments was performed using one-way analysis of variance and the Newman-Keuls multiple comparison post hoc test. The behavioral data were analyzed using two-way analysis of variance with repeated measures and the Scheffé test for post hoc comparisons. Data were shown as the mean ± S.E. Differences with p < 0.05 were considered significant.

oAβ Induce Ser-845-GluA1 Dephosphorylation and a Decrease in Surface Expression of GluA1

We first evaluated the effects of oAβ on critical phosphorylation sites of AMPAR subunits that are important for cell surface receptor expression and regulation of synaptic plasticity (8). Primary neuronal cultures were treated with freshly prepared oligomers of Aβ (oAβ 5 μm, supplemental Fig. S1) prepared as described under “Experimental Procedures,” and phosphorylation levels of Ser-831 and Ser-845 in GluA1 and Ser-880 in GluA2 were analyzed by immunoblotting. We found that oAβ significantly reduced phosphorylation levels (∼35%) of GluA1 Ser-845 in a time-dependent manner but had no effect on GluA1 Ser-831 or GluA2 Ser-880. There were no significant changes in the total amount of GluA1 and GluA2 in the cells treated with oAβ (Fig. 1, A–C).

It is well established that PKA-dependent phosphorylation of GluA1 at Ser-845 increases cell surface expression of AMPARs, whereas NMDA-induced AMPAR dephosphorylation triggers its internalization (12, 27). Because dynamic changes in this phosphorylated site seem to be crucial in the modulation of AMPAR trafficking and synaptic plasticity, we next examined the effect of oAβ on surface GluA1 subunit of AMPAR in primary hippocampal neurons. We found that treatment of cultured neurons with oAβ significantly decreased cell surface GluA1 expression (∼43%) and induced a slight but not significant reduction of GluA2 (Fig. 1, D and E). As a result, the GluA1/GluA2 ratio at the cell surface was significantly decreased (∼40%) after oAβ treatment (Fig. 1F and supplemental Fig. S2).

oAβ Negatively Affects the Interaction between the AMPAR Subunits GluA1 and GluA2

It has been recently reported that ∼80% of synaptic and >95% of somatic extrasynaptic receptors are GluA1/GluA2 heteromers (28). GluA2/GluA3 receptors continuously cycle in and out of synapse, preserving the number of synaptic AMPAR (constitutive pathway), whereas GluA1/GluA2 are added into synapses in an activity-dependent manner during synaptic plasticity (regulated pathway) (29). Because oAβ alters the GluA1/GluA2 ratio of AMPAR across the cell surface, we next tested the possibility that oAβ could be affecting the global subunit composition of AMPARs. For this purpose, we performed immunoprecipitation assays with anti-GluA1 antibodies in neurons treated with oAβ and examined the bound and unbound GluA2. Surprisingly, the levels of GluA2 were decreased in the bounded fraction and increased in the unbounded fraction (∼30%) 60 min after treatment (Fig. 2, A–C). These data revealed that soluble oAβ induced a dissociation of GluA1/GluA2 complexes, which led to a significantly decrease of the GluA2/GluA1 ratio (∼30%) in oAβ-treated neurons (Fig. 2D).

oAβ Induces Calcium Influx into Neurons and Reduces Surface Expression of GluA1 through Ionotropic Glutamate Receptors and Calcineurin Activity

Synapse loss, induced by oAβ in cultured neurons (3), is thought to result from an initial excitotoxicity mediated by oxidative stress and increased [Ca²⁺]_i (30, 31). The increase in [Ca²⁺]_i has been linked to increased NMDAR responsiveness induced by oAβ treatment (32, 33). In support of this idea, our data show that oAβ (5 μm) caused a glutamate receptor-dependent increase in [Ca²⁺]_i (Fig. 3, A and B). Experiments with the membrane-permeable calcium chelator BAPTA-AM (20 μm) and the NMDA and AMPAR antagonists, MK-801 (10 μm) and CNQX (50 μm), revealed that the increase of [Ca²⁺]_i and activation of ionotropic receptors were required for oAβ-mediated AMPAR internalization (Fig. 3, C and D).

When cytosolic calcium reaches critical concentrations, certain LTD-related signaling pathways are activated (34). For instance, NMDAR-dependent LTD in the CA1 region recruits calcineurin (protein phosphatase 2B) (35). Because PKA and calcineurin have been implicated in the regulation of AMPAR trafficking during synaptic plasticity (35–37) and PKA-dependent phosphorylation of GluA1 Ser-845 is reduced by oAβ, we decided to explore the possibility that calcium influx would results in an increase in calcineurin activity. As shown in Fig. 4A, calcineurin activity was significantly increased in the presence of oAβ (∼25% at 30 min). Western blotting analysis revealed unchanged levels of the calcineurin calmodulin-binding catalytic and Ca²⁺-binding subunits in oAβ-treated neurons (Fig. 4B).

Together these results provided evidences indicating that oAβ increased calcium influx and calcineurin activity in an ionotropic glutamate receptor-dependent manner. To establish the relationship between these data and the decrease in phosphorylation levels of GluA1 Ser-845 and cell surface GluA1, we performed experiments in the presence of FK-506, a calcineurin inhibitor (10 μm). We observed that FK-506 was able to prevent the decrease in the levels of phosphorylated Ser-845-GluA1 and cell surface expression of GluA1 induced by oAβ (Fig. 4, C–E).

oAβ Blocks the Extrasynaptic Delivery of AMPAR Mediated by Chemical Potentiation

Although the effect of Aβ on LTD has been less examined than its effects on LTP, there is compelling evidence that high amounts of Aβ actually induce a “chemical” LTD (cLTP) (17, 38). The most notable physiological disruption of synaptic function by synthetic and natural Aβ is the inhibition of LTP, and this effect seems to be caused specifically by oligomeric forms of Aβ (6, 39). Because the mechanisms by which Aβ oligomers inhibit LTP are unclear, we focused our attention in the possible effect of oAβ on inhibition of LTP. To induce cLTP, we used a chemical stimulation protocol of forskolin plus rolipram (F/R) that results in prolonged NMDAR-dependent LTP (40) and increases phosphorylation of GluA1 at Ser-845 and cell surface GluA1 levels (12) (supplemental Fig. S2). Treatment of neuronal cultures with F/R (50 μm/0.1 μm) for 10 min increased basal GluA1 Ser-845 phosphorylation at 30 min (∼70%, Fig. 5A), which correlated with a significant increase in the levels of cell surface expression of GluA1 but not GluA2 (Fig. 5B and supplemental Fig. S3). Bath application of oAβ to cultured neurons prior to F/R treatment partially blocked F/R-mediated Ser-845 phosphorylation and completely blocked surface delivery of GluA1. These results support the hypothesis that oAβ is affecting the bidirectional process of LTP and LTD through the regulation of GluA1 phosphorylation at Ser-845.

Naturally Secreted Aβ Reduces Surface AMPA Receptors

To compare the effects on AMPAR of synthetic oAβ and naturally secreted Aβ, we established primary neurons from a β-amyloid precursor protein (APP) transgenic mouse (APP_Sw,Ind) that develops age-dependent amyloid pathology and memory deficits (24, 25). Neurons from APP_Sw,Ind embryos expressed human APP (∼2-fold) and released soluble Aβ peptides without causing gross morphological synaptic changes (26). Using the biotinylation assay, we found that surface expression of GluA1 was reduced in 12 days in vitro cultured neurons from APP_Sw,Ind mice compared with neurons from wild-type littermates, although total GluA1 levels were not affected (Fig. 6). Surface expression of GluA1 in neurons from APP_Sw,Ind was ∼60% of that observed in wild-type neurons. Treatment of APP_Sw,Ind neurons with the γ-secretase inhibitor DAPT for 72 h restored surface GluA1 to that of control neurons.

To further investigate potential mechanisms underlying synaptic dysfunction between APP_Sw,Ind and wild-type mice, we analyzed the levels of phosphorylated levels of GluA1 Ser-845 in GluA1 in hippocampus from 6-month-old wild-type and APP_Sw,Ind mice, when initial hippocampal Aβ accumulation and spatial memory deficits are detected (24, 25). We examined spatial learning and memory in the Morris water maze, a hippocampal-dependent spatial memory task. After the second trial day, APP_Sw,Ind mice required significantly longer latencies to locate the platform during training (Fig. 7A; two-way analysis of variance; latencies: genotype effect, F₍₁₎ = 10.71; day effect, F₍₄₎ = 24.38; p < 0.0001). Interestingly, levels of phosphorylated GluA1 Ser-845 were increased after the second day of training in the hippocampus of control mice, whereas that increased was not observed in APP_Sw,Ind mice. In addition, total hippocampal GluA1 levels were decreased (45%) in APP_Sw,Ind mice after 2 days of training (Fig. 7, B and C). These results revealed a relationship between initial accumulation of Aβ, memory deficits, and deregulation of AMPARs.