All experimental protocols were approved by the University of Otago Animals Ethics Committee and conducted in accordance with New Zealand Animal Welfare Legislation under the ethics approval ET18/15 and AUP-18-136 for cell culture work and DET19/16 for all acute slice work. All experiments conducted in primary hippocampal cultures were prepared from postnatal Sprague-Dawley rat pups (male or female, P0-P1) sourced from a breeding colony maintained at the Hercus Taieri Resource Unit by the University of Otago (Dunedin, New Zealand). The preparation of primary hippocampal cultures followed a modified protocol based on Banker and Goslin (1998) and Kaech and Banker (2006). Hippocampi were dissociated using papain (Sigma) and plated at a low density on glass-bottomed culture dishes (40,000 cells/cm²; Mattek), or 96-well assay plates (67,500 cells/cm²; Corning, #3603) for immunolabelling. Cells were cultured in Neurobasal A medium (Life Technologies #10888-022), supplemented with B27 (Life Technologies, #17504-001) and Glutamax (Life Technologies, #35050-061) at 37°C/5% CO₂ for 21–27 days in vitro (DIV). Control treatments were undertaken at the same time in matched sets of culture dishes (FUNCAT-PLA, immunocytochemistry).

For experiments examining the role of Ca²⁺-permeable receptors the antagonist IEM-1460 was used (100 μM, Abcam, #AB141507). sAPPα production and purification was carried out according to Turner et al. (2007).

Inhibition of Arc synthesis was achieved using Acell^TM siRNA (1 μM, CUGCAGUACAGUGAGGGUA; Dharmacon, #A-080172-15-0020) targeted to the open-reading frame of the Arc gene, as well as the control non-targeting siRNA (1 μM, UGGUUUACAUGUGUCGACUAA; Dharmacon, #D-001910-01-20). All Accell^TM siRNA were prepared in a 1x siRNA reconstitution buffer consisting of (in mM): KCL 300, MgCl₂ 1, HEPES 30 in RNAse-free water (pH 7.3–7.6), and incubated at 37°C with gentle rocking for 70 min, as according to manufacturer’s instructions. Reconstituted siRNA were aliquoted and stored at −20°C until needed. Expression of siRNA in cultured neurons was confirmed by detection of the red non-targeting (NT) control siRNA (1 μM, UGGUUUACAUGUGUCGACUAA; DY-547, Dharmacon, #D-001960-01-05).

For the detection of proteins in situ primary antibodies were used targeting Arc (rabbit polyclonal, 1:1000, synaptic systems, #156003), Biotin (mouse monoclonal, 1:1000, Sigma, #B7653), MAP2 (guinea-pig polyclonal, 1:1000, synaptic systems, #188004; monoclonal, 1:1000, Abcam, #AB11267), Synapsin-1 (mouse monoclonal, 1:1000, synaptic systems, #106011), GluA1 (C-terminal; rabbit polyclonal, 1:500, Abcam, #AB31232), GluA1 (N-terminal; mouse monoclonal, 1:500, #MAB2263), GluA2 (C-terminal; rabbit polyclonal, 1:500, Abcam, #AB1768-I), GluA2 (N-terminal; mouse polyclonal, 1:500, Abcam, #133477), and GluA3 (N-terminal; mouse monoclonal, 1:250, Thermofisher, #32-0400).

Detection of primary antibodies was achieved via addition of the following secondary antibodies: Goat anti-Guinea Pig Alexa Fluor 488 (1:1000, Thermofisher, #A11073), Goat anti-rabbit Alexa Fluor 555 (1:1000, Invitrogen, #A21429), Goat anti-mouse Alexa Fluor 488 (1:500; Thermofisher, #A11001), Donkey anti-mouse PLA^minus probe (1:10, Sigma-Aldrich, #DUO92004), Donkey anti-rabbit PLA^plus probe (1:10, Sigma-Aldrich, #DUO92002), DAPI (1:1000, Thermofisher, #D1306), and Duolink detection reagent Texas Red (1:5, Sigma-Aldrich, #DUO92008).

For experiments examining the effect of siRNA treatment on protein expression, DIV21-27 primary hippocampal neurons were treated with sAPPα (1 nM; 2 h), or culture media only. Following pre-treatment with siRNA (1 μM, 60 min), primary hippocampal cultures were co-treated with either Arc or NT siRNA (1 μM, 2 h), in the presence or absence of sAPPα (1 nM). Following incubation, cells were fixed in 4% paraformaldehyde (pH 7.4; 20 min) in PBS supplemented with 1 mM MgCl₂ and 0.1 mM CaCl₂ (PBS-MC) containing sucrose (PBS-MCS; 155.42 mM), and permeabilized with 0.5% Triton X-100 in PBS (pH 7.4; 15 min; for the detection of cell surface GluA1 using an N-terminal antibody this step was omitted). Cells were then blocked in 4% normal goat serum in PBS (pH 7.4) for 60 min at room temperature (RT). Cells were incubated with primary antibodies of interest (2 h, RT), followed by 3 × 5 min washes (PBS, pH 7.4) and incubation in appropriate secondary antibody (30 min, RT), followed by 3 × 5 min washes (PBS, pH 7.4).

FUNCAT-PLA labeling of newly synthesized proteins was conducted according to a previously published protocol (Dieterich et al., 2007; tom Dieck et al., 2015), adapted for detection of proteins at the cell surface. Cells were incubated in 4 mM L-azidohomoalanine (AHA, Click Chemistry Tools #1066-1000) in the presence or absence of sAPPα or anisomycin. Following incubation, cells were washed (3x quickly) with PBS-MC (pH 7.4), and fixed in PFA in PBS-MCS (pH 7.4). Cell-surface azide-labeled newly synthesized proteins were alkylated with biotin-linked alkyne via a copper-mediated click reaction. Click reaction mixture comprised of 200 μM triazole ligand (Tris ((1-benzyl-1H-1,2,3-triazol-4-yl)methyl) amine; TBTA, Aldrich #678973), 500 μM TCEP (Tris(2-carboxyethyl)phosphine hydro-chloride, Thermo Scientific #PIE-20490), 25 μM Biotin-PEG4-alkyne (Biotin alkyne, Aldrich # B10185) and 200 μM CuSO₄ in PBS pH 7.8 was incubated on cells overnight at RT. For detection of de novo GluA1 and GluA2 proteins cells were permeabilized with 0.5% Triton X-100 in PBS (pH 7.4), and incubated with anti-biotin, anti-GluA1 or anti-GluA2 antibodies diluted in 4% normal goat serum. Donkey anti-mouse PLA^minus, and donkey anti-rabbit PLA^plus probes were applied, followed by ligation and amplification with Duolink detection reagent Texas Red according to the manufacturer’s instructions. Neuronal somata, dendrites and nuclei were visualized by addition of anti-Guinea Pig Alexa Fluor 488 and DAPI, respectively.

The labeling of cell-surface receptor subunit dimers was conducted as per (Lin et al., 2015) and adapted for the detection of AMPARs present at the cell surface. In brief, cells were treated with sAPPα (1 nM) or existing media. Following incubation, cells were washed with PBS-MC (pH 7.4), and fixed in PFA in PBS-MCS (pH 7.4). For experiments investigating GluA1/2-containing AMPARs, using both C-terminal and N-terminal antibodies, cells were incubated in blocking buffer (4% normal goat serum in PBS, 60 min at RT), probed with the N-terminal antibody (90 min, RT), washed (PBS pH 7.4, 3 × 5 min), and subsequently permeabilized with 0.5% Triton X-100 in PBS (15 min, RT). Cells were then probed with the C-terminal antibody in addition to anti-MAP2 for visualization of neuronal structure (90 min, RT) and washed (PBS pH 7.4, 3 × 5 min). For experiments utilizing two N-terminal antibodies, such as those investigating GluA2/3-containing AMPARs, permeabilization was omitted and cells were incubated in blocking buffer (4% normal goat serum in PBS, 60 min at RT) followed by incubation of primary antibodies (90 min, RT). Following this, cells were washed (PBS pH 7.4, 3 × 5 min), permeabilized with 0.5% Triton X-100 in PBS (15 min, RT) and incubated in blocking buffer again (4% normal goat serum in PBS, 60 min at RT) before addition of anti-MAP2 and DAPI for visualization of neuronal structure (90 min, RT). Following addition of antibodies, proximity ligation assay was performed as described above.

All experiments conducted on acute tissue were prepared from young adult male Sprague-Dawley rats (42–56 d), as described previously (Mockett et al., 2019). Rats were deeply anesthetized with ketamine (100 mg/kg, i.p.) and decapitated via guillotine. The brains were removed and chilled in ice-cold and oxygenated modified artificial cerebrospinal fluid (aCSF) for which sucrose was substituted for NaCl (composition in mM: sucrose 210, glucose 20, KCl 2.5, NaH₂PO₄ 1.25, NaHCO₃ 26, CaCl₂ 0.5, MgCl₂ 3, pH 7.4 when gassed with 95% O₂-5% CO₂). Hippocampi were dissected and slices (400 μm) cut using a vibroslicer (Leica, VT1000). Slices were transferred to a porous, transparent membrane in an incubation chamber, and maintained at the interface between air and standard aCSF (in mM: NaCl 124, KCl 3.2, NaH₂PO₄ 1.25, NaHCO₃ 26, CaCl₂ 2.5, MgCl₂ 1.3, D-glucose 10, equilibrated with carbogen 95% O₂-5% CO₂; 32°C) for 30 min followed by RT for an additional 90 min. Slices were then transferred to the recording chamber containing recirculating aCSF (95% O₂, 5% CO₂; 32.5°C), superfused continuously at a rate of 2 mL/min. Baseline field excitatory postsynaptic potentials (fEPSPs) were elicited in area CA1 by stimulation of the Schaffer collateral-commissural pathway at 0.017 Hz (diphasic pulses, 0.1 ms half-wave duration) using a Teflon-coated 50 μm tungsten wire monopolar electrode (A-M Systems Inc., Carlsborg, WA) connected to Grass P511 AC amplifiers via high impedance probes (Grass Instruments Company, West Warwick, United States). Signals were amplified (x1000) with half-amplitude filter cut-offs of 0.3 Hz and 3 kHz. Evoked responses were recorded with a glass microelectrode (A-M systems, 1.0 mm x 0.58 mm, 4”; Catalog No. #601000) filled with aCSF (1.9–2.9 MΩ) and placed in stratum radiatum of area CA1 (approximately 300 μm from the stimulating electrode). During periods of baseline recording the stimulation intensity was adjusted to elicit a fEPSP with an initial slope value of 40% of the maximum elicited when delivering 200 μA of current. Non-saturated LTP was induced by applying a mild theta burst-stimulation protocol (TBS; 5 bursts of 5 pulses at 100 Hz delivered at 200 ms intervals) at baseline stimulus intensity as per Mockett et al. (2019). sAPPα (1 nM) and IEM-1460 (100 μM) were bath-applied by switching to an identical preheated and oxygenated aCSF solution that contained the compound of interest. sAPPα was delivered 30 min prior to TBS and IEM-1460 was delivered 20 min prior to and during the sAPPα administration and continued 10 min post-TBS. Control experiments were routinely interleaved randomly between experimental treatments.

Images were acquired using an Olympus IX71 inverted light microscope using a 20x/0.45-N.A objective (LUCPFLN) or 4x/0.13-N.A objective (UPFLN). The images were captured using a Hamamatsu Orca-AG camera (C4742-80-12AG) in 1024 × 1024 pixel 8-bit mode and saved as.tif files.

To quantify the PLA signal a custom-made ImageJ script created by Maximilian Heumüller (Max Planck Institute for Brain Research, Frankfurt) was used (tom Dieck et al., 2015). Following thresholding of the FUNCAT-PLA signal, a ‘MAP2 mask’ was generated capturing the area of the neuron, and the size and intensity of FUNCAT-PLA puncta within the mask were measured and recorded. To analyze the signal within the somatic and dendritic compartments, somata were isolated from each cell as above, and the proximal 50 μm segment all dendrites were straightened using the ‘straighten’ plugin in ImageJ, and analyzed for PLA signal. For the detection of somatic receptor subunit dimers using PLA, analysis included a step to exclude non-specific signal detected within the nucleus using a mask generated by the DAPI channel. For image representation, ImageJ was used to adjust the brightness and contrast equally for all treatment groups. To quantify immunofluorescence, neurons were outlined using ImageJ. An ‘integrated intensity/neuronal area’ value was generated for each cell and somatic compartment, including all dendrites up until intersection with neighboring dendrites. This value was corrected for average background fluorescence by subtracting mean gray values of background fluorescence.

It is not known whether sAPPα induces the rapid synthesis of GluA1-containing AMPARs or whether these de novo molecules are trafficked to the cell surface and thus contribute to plasticity. Thus, to examine specifically the cell-surface population of de novo GluA1, we utilized FUNCAT-PLA under detergent-free conditions. We found that sAPPα treatment (1 nM; 30 min) significantly increased the cell-surface expression of de novo GluA1 in both the soma (2.01 ± 0.33 mean ± SEM, p = 0.0039; Figure 1A) and dendrites (3.97 ± 0.32, p = 0.0009; Figure 1B) of cultured hippocampal neurons. Interestingly, following 2 h of sAPPα treatment, there was no detectable change in de novo somatic (0.83 ± 0.15, p = 0.265; Figure 1A) or dendritic (0.86 ± 0.29, p = 0.673; Figure 1B) GluA1. Indeed, cell-surface expression of GluA1 at 2 h was significantly decreased relative to 30 min of sAPPα treatment on both the soma (p = 0.0058) and dendrites (p = 0.0005). Interestingly, in parallel experiments using immunocytochemistry with non-permeabilized conditions to allow detection of both extant and newly made cell surface GluA1, we were able to detect increases in GluA1 in the dendrites. We observed a modest increase by 30 min (2.37 ± 0.37, p = 0.0787), which reached statistical significance by 2 h (3.00 ± 0.44, p = 0.0002) and remained significantly elevated at 4 h (2.50 ± 0.39, p = 0.0204). We found no changes in the soma at any time point (30 min: 1.09 ± 0.12, p = 0.8998; 2 h: 1.066 ± 0.82, p = 0.2694, 4 h: 1.043 ± 0.15, p = 0.3191; see Supplementary Figure 1). Together, these results suggest that sAPPα induces the rapid de novo synthesis of GluA1 subunits, which are trafficked to the cell surface by 30 min. These newly made subunits are later internalized within 2 h of treatment, yet levels of extant GluA1-containing receptors remain elevated for up to 4 h.

As the GluA1 subunit is a constituent of either GluA1/2 or GluA1-homomeric receptors, to determine the likely composition of de novo AMPARs formed following sAPPα treatment, we next examined the temporal expression of de novo GluA2 at the cell surface, in both the somatic and dendritic compartments. Here, we found that de novo cell-surface GluA2 remained unaffected at the soma by either 30 min (0.86 ± 0.28, p = 0.654) or 2 h (1.32 ± 0.21, p = 0.148; Figure 2A) sAPPα treatments. Conversely, there was an early decrease of de novo cell-surface GluA2 at the dendritic cell surface (30 min: 0.54 ± 0.17, p = 0.009), which returned to control levels within 2 h (1.92 ± 1.08, p = 0.395; Figure 2B). From this, we can infer that while sAPPα upregulates the expression of de novo GluA1, it inhibits the synthesis or otherwise restricts expression of de novo GluA2 at the cell surface. Together, these results indicate that sAPPα acts both to upregulate cell surface de novo GluA1-containing homomeric AMPARs and downregulate de novo GluA2-containing AMPARs.

To further explore the localization of de novo GluA1 subunits, we determined the proportion of dendritic GluA1-containing AMPARs, identified by FUNCAT-PLA, associated with the presynaptic marker synapsin-1 (Figures 3A,B). Synaptic overlap of GluA1 puncta was determined using Mander’s overlap coefficient (MOC, Figure 3C). We found no significant difference between control and sAPPα-treated (1 nM, 30 min) conditions (control MOC: 0.35 ± 0.03 mean ± SEM, sAPPα: 0.31 ± 0.02; p = 0.7038; Figure 3C), indicating that 30 min sAPPα treatment did not increase the proportion of de novo synaptic GluA1. We next determined the proportion of synaptic, extrasynaptic and non-synaptic cell surface de novo GluA1 puncta relative to proximity to synapsin-1-positive synapses. We defined synaptic, extrasynaptic, and non-synaptic PLA puncta as 0–2, 2–4, and > 4 μm from the closest synapsin-1 center of mass, ‘. This analysis showed a shift in the frequency of de novo GluA1 puncta present at synapses following sAPPα treatment, increasing the proportion of GluA1 puncta 2–4 μm proximal to the synapse (Figure 3D). In line with this, we found a significant increase in the number of de novo GluA1 puncta at extrasynaptic sites (control: 1.23 ± 0.09 puncta; sAPPα: 1.92 ± 0.14; p = 0.0003). No change was detected at synaptic (control: 1.28 ± 0.10; sAPPα: 1.34 ± 0.12; p = 0.9872), or non-synaptic (control: 1.52 ± 0.09; sAPPα: 1.45 ± 0.11; p = 0.9655) sites (Figure 3E). These results suggest that sAPPα rapidly enhances the extrasynaptic pool of de novo GluA1-containing AMPARs within the dendrites.

Prolonged treatment of sAPPα (1 nM, 2 h) enhances the de novo transcription and translation of the immediate early gene Arc, a protein intrinsically linked to glutamate receptor expression at the cell surface (Livingstone et al., 2019). Treatment of primary hippocampal cultures with an siRNA (1 μM, pre-treatment: 60 min, treatment: 2 h) targeting Arc mRNA or a non-targeting (NT) siRNA with no known homology to rat or human genes had no significant effect on basal Arc protein expression in either the soma (Arc siRNA: 0.78 ± 0.06, p ≥ 0.99; NT siRNA: 1.11 ± 0.09, p = 0.0509; Figure 4A) or the dendrites (Arc siRNA: 1.26 ± 0.12, p = 0.8774; NT siRNA: 1.28 ± 0.08, p = 0.7561; Figure 4B). As previously shown (Livingstone et al., 2019), sAPPα treatment (1 nM, 2 h) enhanced both somatic (2.05 ± SEM, p = 0.0306; Figure 4A) and dendritic (2.33 ± 0.29, p = 0.0010; Figure 4B) Arc protein expression. Co-treatment with sAPPα and the NT siRNA (1 μM, pre-treatment: 60 min, co-treatment: 2 h) also enhanced Arc expression (soma: 1.59 ± 0.23, p = 0.0198; dendrites: 1.98 ± 0.18, p = 0.0022), and no significant difference was detected between sAPPα-treated and sAPPα + NT siRNA in either the soma or dendrites (p ≥ 0.99). Thus, NT siRNA does not affect the expression of Arc protein. By contrast, co-treatment with sAPPα and the Arc siRNA significantly reduced Arc protein expression in both the soma (0.72 ± 0.27, p = 0.0100) and dendrites (1.16 ± 0.68, p = 0.0066) relative to sAPPα treatment alone, and was not significantly different from control in either compartment (p ≥ 0.99). Together, these results indicate that Arc siRNA effectively inhibits sAPPα-induced de novo Arc synthesis in our primary hippocampal cultures.

As de novo GluA1 levels at the cell surface did not persist following prolonged (2 h) treatment of sAPPα, we next employed siRNA knockdown of de novo Arc protein to assess whether inhibition of Arc protein synthesis affects de novo GluA1 cell surface levels. As expected, sAPPα treatment for 2 h resulted in no change in either somatic (0.73 ± 0.16, p = 0.118) or dendritic (0.85 ± 0.85, p = 0.374) expression of de novo cell surface GluA1, relative to control conditions (Figures 4A,B). However, in the presence of Arc siRNA (preincubation: 1 μM, 60 min, followed by co-incubation with sAPPα: 1 nM, 2 h), we observed a significant increase in the expression of de novo GluA1 at both the somatic (1.55 ± 0.36, p = 0.0043; Figure 5A) and dendritic (4.14 ± 0.65, p ≤ 0.0001; Figure 5B) cell surface, relative to sAPPα-only conditions. Finally, treatment of cultures with the NT siRNA alone had no observable effect on the levels of de novo cell surface GluA1 at the soma (0.94 ± 0.15, p = 0.7247, n = 23; Supplementary Figure 2A), or dendrites (1.08 ± 0.21, p = 0.7050, n = 89; Supplementary Figure 2B) relative to control, nor relative to NT + sAPPα treatment in the soma (0.69 ± 0.12, p = 0.2790; n = 18; Supplementary Figure 2A) or dendrites (0.95 ± 0.20, p = 0.2077, n = 98; Supplementary Figure 2B), indicating that the observed effects are due to the specific to the addition of Arc siRNA and subsequent knockdown of Arc protein expression and not as a result of non-specific siRNA-mediated effects. Therefore, these results show that following the rapid expression of de novo GluA1-containing AMPARs at the cell surface, these AMPARs are subsequently endocytosed in an Arc-dependent manner.

In response to synaptic activity (Bagal et al., 2005; Kopec et al., 2006; Guire et al., 2008; Tanaka and Hirano, 2012; Park et al., 2016; Pandya et al., 2018), behavioral learning (Whitlock et al., 2006; Fachim et al., 2016), and neuromodulators (Leonoudakis et al., 2008; Jourdi and Kabbaj, 2013), cell surface or synaptic accumulation of GluA2-containing AMPARs has been shown to occur, typically following the incorporation of GluA1-containing homomers (Shi et al., 1999; Bellone and Lüscher, 2005, 2006; Guire et al., 2008; Hong et al., 2013; Park et al., 2016).

While previous research examining acute sAPPα treatments (Mockett et al., 2019) found no evidence for an increase in de novo GluA2 synthesis in response to sAPPα, this does not exclude the possibility that sAPPα additionally promotes the trafficking of pre-existing GluA2-containing AMPARs to the cell surface. Therefore, we extended our analysis to examine cell surface populations of GluA1- and GluA2-containing (GluA1/2) AMPARs within somatic and dendritic compartments. Here, we utilized PLA to detect the coincident proximity of GluA1 and GluA2 AMPAR complexes at the cell surface. We found no significant increase in GluA1/2-containing AMPARs at the cell surface following a 30 min sAPPα treatment, in either the soma (1.04 ± 0.11, p = 0.709; Figure 6A) or dendrites (1.05 ± 0.09, p = 0.54; Figure 6B) of cultured primary hippocampal neurons. However, the cell surface expression of GluA1/2 AMPARs significantly increased in both the soma (2.15 ± 0.30, p = 0.0013; Figure 6A), and dendrites (2.28 ± 0.14, p ≤ 0.0001; Figure 6B) by 2 h. These results indicate that GluA1/2 AMPARs are expressed at the cell surface following prolonged sAPPα treatment.

We further sought to determine the synaptic localization of cell surface GluA1/2 AMPARs (Figures 7A,B). Synaptic overlap of GluA1/2 puncta was determined using MOC (Figure 7C). Here, we found a significant increase in MOC in sAPPα-treated (1 nM, 2 h) conditions (MOC: 0.295 ± 0.02, p = 0.0467, Figure 7C), relative to control (control: 0.231 ± 0.02; Figure 7C), indicating that sAPPα treatment increased the proportion of GluA1/2 AMPARs at synaptic sites. To validate this finding, we determined the proximity of synaptic, extrasynaptic, and non-synaptic cell surface GluA1/2 puncta relative to the proximity of synapsin-1-positive synapses, as above (Figure 7). Supporting the observed increase in MOC, we observed a shift in the frequency ofnd remained compara GluA1/2 AMPARs present at synapses following sAPPα treatment, increasing the proportion of GluA1 puncta 0–1 μm proximal to the synapse (Figure 7D). Expanding on this, we found a significant increase in the number of GluA1/2 at synaptic sites (control: 1.35 ± 0.018 puncta; sAPPα: 2.51 ± 0.27, p = 0.0029), however, no change was detected at extrasynaptic (control: 2.41 ± 0.24; sAPPα: 2.44 ± 0.27, p = 0.9997), or non-synaptic (2.18 ± 0.26; sAPPα: 1.85 ± 0.21, p = 0.7089) sites (Figure 7E). These results show that sAPPα enhances the synaptic pool of GluA1/2 AMPARs within the dendrites of primary hippocampal neurons, potentially replacing de novo GluA1 homomers.

GluA2- and GluA3-containing (GluA2/3) AMPARs comprise the second largest subtype of hippocampal AMPARs. The presence of GluA2/3 AMPARs is thought to denote synaptic maturity, with expression increasing throughout development and acting to replace GluA1/2 AMPARs at the synapse via constituent recycling (Zhu et al., 2000; Shinohara and Hirase, 2009). Interestingly, previous research has shown that GluA3-containing AMPARs are regulated by in vivo LTP (Williams et al., 2007), and in vitro LTD (Holman et al., 2007), and growth factor treatment (Narisawa-Saito et al., 1999). Curiously, GluA3-containing AMPARs do not appear to directly regulate the expression of LTP or context fear memory formation (Meng et al., 2003; Humeau et al., 2007), but their removal from the synapse may be an essential step required for LTD (Holman et al., 2007). Regardless, their synaptic expression is considered essential for basal synaptic transmission (Meng et al., 2003).

Using PLA to label cell surface GluA2/3-containing AMPARs, we found that sAPPα (1 nM, 30 min) significantly decreased the cell surface expression of GluA2/3-containing AMPARs within the dendrites of cultured neurons (0.63 ± 0.12, p = 0.0033; Figure 8B), which remained decreased at 2 h (0.57 ± 0.11, p = 0.0002; Figure 8B). Interestingly, somatic levels of GluA2/3 AMPARs remained unaffected following both 30 min (0.72 ± 0.23, p = 0.255; Figure 8A), and 2 h (1.19 ± 0.32, p = 0.557) treatments. These results may indicate that GluA3-containing AMPARs are removed from the dendritic cell surface to permit the insertion of GluA1 homomers or GluA1/2- AMPARs (Shi et al., 2001). Alternatively, the removal of GluA3-containing AMPARs may reflect homeostatic processes, maintaining synaptic activity within a physiological range (Rial Verde et al., 2006; Diering et al., 2014; Tan et al., 2015).

The work described above indicates that sAPPα enhances the extrasynaptic expression of de novo GluA1, but not GluA2-containing AMPARs in primary hippocampal neurons (Figures 1, 2). To determine whether these GluA1-containing AMPARs comprise functional homomeric Ca²⁺-permeable receptors, we assessed their involvement in plasticity in acute hippocampal slices, building on Mockett et al. (2019) and Richter et al. (2018) who have previously shown that sAPPα enhances submaximal LTP in area CA1. We first examined the dependence of CP-AMPARs during TBS-induced LTP alone (Figures 9A–C), by utilizing the CP-AMPAR antagonist IEM-1460. Following TBS, continued perfusion of IEM-1460 (100 μM; 10 min) had no significant effect on the induction of LTP (0–10 min post-TBS; control: 52.10 ± 6.19% of baseline, mean ± SD; IEM-1460: 59.66 ± 6.94% of baseline, p = 0.9399; Figures 9C,D), and remained comparable to no-drug control slices following washout (50–60 min post-TBS; control: 26.64 ± 4.33% of baseline; IEM-1460: 27.99 ± 4.12% of baseline, p = 0.9969; Figures 9C,E), supporting past research (Adesnik and Nicoll, 2007; Gray et al., 2007; Asrar et al., 2009). Due to this, we conclude that CP-AMPARs do not play a significant role in the potentiation of synaptic transmission following a mild TBS protocol, and therefore conclusions drawn following treatment of slices with sAPPα may be attributed to mechanisms activated by sAPPα alone.

We next examined whether the enhancement of LTP induction following sAPPα treatment (1 nM, 30 min) was dependent on the function of CP-AMPARs. Here, slices were perfused with IEM-1460 (100 μM, 50 min) before TBS, and 10 min post-TBS stimulation. Slices were also perfused with sAPPα 30 min before TBS in the presence or absence of IEM-1460. As shown in Figure 9, relative to control slices, pre-treatment with sAPPα (1 nM, 30 min) enhanced both the induction (126.2 ± 13.55% of baseline, p ≤ 0.0001; Figures 9C,D) and persistence (77.41 ± 10.66% of baseline, p = 0.0002; Figures 9C,E) of LTP, confirming previously observed enhancements described by Mockett et al. (2019). Co-application of IEM-1460 significantly inhibited the sAPPα-enhanced early potentiation (84.19 ± 11.74% of baseline, p = 0.0334; Figures 9C,D), while wash-out of IEM-1460 resulted in the recovery of potentiation, which was no longer different from sAPPα treatment alone (70.78 ± 15.44% of baseline, p = 0.3518; Figures 9C,E), but was significantly different from control slices (p = 0.0149). These results indicate that at least the initial period of the sAPPα-mediated enhancement of LTP is due to the rapid incorporation of functional CP-AMPARs into the synapse.

Secreted amyloid precursor protein-alpha enhances the synthesis of GluA1 but not GluA2 subunits (Mockett et al., 2019). Although sAPPα alters transcription profiles (Stein et al., 2004; Ryan et al., 2013), it has not been shown to alter levels of AMPAR subunit mRNA within a 2 h treatment window. sAPPα does, however, enhance protein synthesis within isolated hippocampal synapses (Claasen et al., 2009), and it is well established that AMPAR subunits, including GluA1, undergo synthesis within dendrites (Torre and Steward, 1992; Kacharmina et al., 2000; Grooms et al., 2006; Sutton and Schuman, 2006) for direct exocytosis at extrasynaptic sites (Choquet, 2018). Thus, it is likely that the rapid de novo synthesis of GluA1 results from translation of pre-existing mRNA. It is yet to be determined whether sAPPα-induced GluA1 synthesis occurs within dendrites or is delivered to dendrites via lateral diffusion from somatic sites (Borgdorff and Choquet, 2002; Adesnik et al., 2005) or active transport throughout the dendrites (Passafaro et al., 2001; Hangen et al., 2018).

We have shown that de novo GluA1 is trafficked to extrasynaptic sites and is later internalized, but not degraded, as intracellular levels remain elevated (Mockett et al., 2019). As extrasynaptic AMPARs are necessary for the induction of LTP (Oh et al., 2006; Penn et al., 2017) and increasing the proportion of extrasynaptic AMPARs enhances the expression of LTP (Oh et al., 2006), the enhanced GluA1 expression likely provides reservoirs of CP-AMPARs which could underpin the sAPPα-mediated priming of LTP. Indeed, we confirmed that sAPPα does not affect responses to baseline stimulation (Richter et al., 2018; Mockett et al., 2019), nor did IEM-1460 affect baseline responses in sAPPα-treated slices any more than control slices, all supporting the view that the enhancement of cell-surface GluA1 expression by sAPPα is strictly extrasynaptic. However, pre-treatment with sAPPα did enhance LTP induction in a manner that was dependent on CP-AMPARs, likely due to lateral diffusion of the recently inserted extrasynaptic CP-AMPARs.

The observed elevation in extrasynaptic de novo GluA1 occurs alongside a reduction in cell-surface de novo GluA2 and extant dendritic GluA2/3 while no cell-wide reduction in levels was evident. As GluA2 mRNA is also expressed in dendrites, local downregulation of GluA2 synthesis may be controlled locally via the RNA interacting protein, CPEB3 (Savtchouk et al., 2016) or FMRP (Muddashetty et al., 2007), or via microRNA (miRNA). Indeed, we have previously shown that sAPPα transiently upregulates miR-30 (Ryan et al., 2013), a miRNA known to regulate the expression of GluA2 (Song et al., 2019). Alternatively, association with Protein Interacting with C Kinase-1 (PICK1) may restrict GluA2 trafficking (Terashima et al., 2004; Jaafari et al., 2012) or AP2-GluA2 binding may also promote internalization (Hanley, 2018). Furthermore, the NO-cGMP-PKG and MAPK pathways have been found to be involved in de-clustering of GluA2/3-containing AMPARs (Endo and Launey, 2003), an event which is typically associated with subsequent internalization of receptors (Matsuda et al., 2000). As synaptic protein synthesis, expression of Arc protein, and GluA1 trafficking are dependent on the activity of PKG and MAPK (Claasen et al., 2009; Livingstone et al., 2019; Mockett et al., 2019), sAPPα’s induction of these pathways may also contribute to GluA2/3 receptor internalization. Further, rapid downregulation of GluA2-containing calcium impermeable AMPARs (CI-AMPARs) may facilitate GluA1 cell-surface expression via increasing the available “slots” within the PSD (MacDougall and Fine, 2013) to be filled by CP-AMPARs (Bellone and Lüscher, 2005, 2006; Hong et al., 2013). Indeed, in vitro live-cell imaging shows simultaneous downregulation of cell-surface GluA2/3 and upregulation of GluA1-containing AMPARs following LTP induction (Tanaka and Hirano, 2012).

Interestingly, we also observed a delayed increase in the proportion of GluA1/2 AMPARs following sAPPα treatment, at both the somatic and dendritic cell-surface primarily at synaptic sites. These receptors were likely derived from pre-existing pools as they were not tagged in the FUNCAT-PLA assays. Previous work has shown that chronic overexpression of APP in vitro upregulates cell-surface GluA2 expression (Lee et al., 2010), indicating that APP, and likely sAPPα, may regulate the persistence of synaptic plasticity through the long-term expression of GluA2-containing AMPARs. As their increase occurs at a timepoint at which both de novo GluA1 and extant GluA2/3 are internalized, this would result in a switch to synapses expressing GluA1/2 CI-AMPARs (Shi et al., 2001; Bellone and Lüscher, 2005; Sutton et al., 2006; Mameli et al., 2007), as observed during reconsolidation of memories (MacDougall and Fine, 2013). As we have shown that sAPPα enhances Arc expression and that internalization of de novo GluA1 is Arc dependent, Arc expression likely facilitates the addition of synaptic GluA1/2-containing AMPARs (McCormack et al., 2006; Hong et al., 2013). Indeed, previously knockdown of Arc has been shown to increase cell-surface GluA1- but not GluA2-containing AMPARs (Shepherd et al., 2006), while overexpression of Arc protein significantly decreases CP-AMPAR–associated rectification in neurons (DaSilva et al., 2016), a mechanism which may permit addition of CI-AMPARs. Recently, increased Arc expression has been shown to protect neurons against CP-AMPAR–mediated oxyhemoglobin excitotoxicity (Chen et al., 2020), via endocytosis of these AMPARs. While reduced phosphorylation of GluA1_S845 may also contribute to internalization (Man et al., 2007; Lussier et al., 2015), our observations support a role for Arc-dependent endocytosis in the regulation of de novo CP-AMPARs.

Of note, sAPPα has been previously shown to enhance LTP and plasticity-related protein synthesis via a mechanism involving α7nAChRs (Richter et al., 2018; Livingstone et al., 2019) in that antagonism of α7nAChRs during the priming phase, but not during the induction of LTP, impairs the expression of sAPPα-enhanced LTP (Richter et al., 2018). Our current work suggests that this occurs by promoting the synthesis and trafficking of CP-AMPARs. Indeed, activation of postsynaptic α7nAChRs by acute nicotine treatment promotes the rapid trafficking of CP-AMPARs (Lozada et al., 2012; Halff et al., 2014; Tang et al., 2015), while chronic α7nAChR activation enhances the cell-surface expression of GluA1/2 AMPARs (Duan et al., 2015).