APP fragment controls both ionotropic and
non-ionotropic signaling of NMDA receptors

Jade Dunot, Sebastien Moreno, Carine Gandin, Paula A Pousinha, Mascia Amici, Julien Dupuis, Margarita Anisimova, Alex Winschel, Magalie Uriot, Samuel J Petshow, et al.

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To cite this version:

Jade Dunot, Sebastien Moreno, Carine Gandin, Paula A Pousinha, Mascia Amici, et al. APP fragment controls both ionotropic and non-ionotropic signaling of NMDA receptors. Neuron, 2024, 112 (16), pp.2708-2720.e9. 10.1016/j.neuron.2024.05.027. hal-04742865

HAL Id: hal-04742865

https://hal.science/hal-04742865v1

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# APP fragment controls both ionotropic and non-ionotropic signaling of NMDA receptors

## Highlights

* AETA hampers NMDA receptor ionotropic activity by competing with its co-agonists

* AETA modifies NMDA receptor conformation and enhances ion-flux-independent signaling

* AETA level is raised by increased neuronal activity

* Deletion of AETA modifies NMDA receptor signaling and associated synapse plasticity

## Authors

Jade Dunot, Sebastien Moreno, Carine Gandin, ..., Laurent Groc, Michael Willem, Hélène Marie

## Correspondence

mwillem@med.uni-muenchen.de (M.W.),
marie@ipmc.cnrs.fr (H.M.)

## In brief

Dunot et al. identify AETA, a bioactive cleavage product of amyloid-β precursor protein, as a novel endogenous NMDA receptor regulator. AETA competes with receptor co-agonists, thereby modulating ionotropic and non-ionotropic receptor activities. Chronic AETA loss impairs synaptic plasticity and memory. These findings highlight a new mechanism regulating brain information processing.
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Report

# APP fragment controls both ionotropic and non-ionotropic signaling of NMDA receptors

Jade Dunot,¹,²,¹¹ Sebastien Moreno,¹,¹¹ Carine Gandin,¹ Paula A. Pousinha,¹ Mascia Amici,³ Julien Dupuis,⁴

Margarita Anisimova,⁵ Alex Winschel,⁶ Magalie Uriot,⁴ Samuel J. Petshow,⁵ Maria Mensch,¹ Ingrid Bethus,¹

Camilla Giudici,⁷ Heike Hampel,⁸ Benedikt Wefers,⁷,⁹ Wolfgang Wurst,²,⁷,⁹ Ronald Naumann,¹⁰ Michael C. Ashby,³

Bodo Laube,⁶ Karen Zito,⁵ Jack R. Mellor,³ Laurent Groc,⁴ Michael Willem,⁸,* and Hélène Marie¹,¹²,*

¹Université Côte d'Azur, CNRS, INSERM, Institut de Pharmacologie Moléculaire et Cellulaire, 06560 Valbonne, France

²Munich Cluster for Systems Neurology (SyNergy), 81377 Munich, Germany

³Centre for Synaptic Plasticity, School of Physiology, Pharmacology and Neuroscience, University of Bristol, Bristol BS8 1TD, UK

⁴Université de Bordeaux, CNRS, Interdisciplinary Institute for Neuroscience, 33076 Bordeaux Cedex, France

⁵Center for Neuroscience, University of California, Davis, Davis, CA 95618, USA

⁶Department of Biology, Neurophysiology und Neurosensory Systems, TU Darmstadt, 64287 Darmstadt, Germany

⁷German Center for Neurodegenerative Diseases (DZNE) Munich, 81377 Munich, Germany

⁸Biomedical Center (BMC), Division of Metabolic Biochemistry, Faculty of Medicine, Ludwig-Maximilians-Universität München, 81377 Munich, Germany

⁹Institute of Developmental Genetics, Helmholtz Zentrum München, 85764 Neuherberg, Germany

¹⁰Max Planck Institute of Molecular Cell Biology and Genetics, 01307 Dresden, Germany

¹¹These authors contributed equally

¹²Lead contact

*Correspondence: mwillem@med.uni-muenchen.de (M.W.), marie@ipmc.cnrs.fr (H.M.)

https://doi.org/10.1016/j.neuron.2024.05.027

## SUMMARY

NMDA receptors (NMDARs) are ionotropic receptors crucial for brain information processing. Yet, evidence also supports an ion-flux-independent signaling mode mediating synaptic long-term depression (LTD) and spine shrinkage. Here, we identify AETA ($A_η$), an amyloid-$β$ precursor protein (APP) cleavage product, as an NMDAR modulator with the unique dual regulatory capacity to impact both signaling modes. AETA inhibits ionotropic NMDAR activity by competing with the co-agonist and induces an intracellular conformational modification of GluN1 subunits. This favors non-ionotropic NMDAR signaling leading to enhanced LTD and favors spine shrinkage. Endogenously, AETA production is increased by in vivo chemogenetically induced neuronal activity. Genetic deletion of AETA production alters NMDAR transmission and prevents LTD, phenotypes rescued by acute exogenous AETA application. This genetic deletion also impairs contextual fear memory. Our findings demonstrate AETA-dependent NMDAR activation (ADNA), characterizing AETA as a unique type of endogenous NMDAR modulator that exerts bidirectional control over NMDAR signaling and associated information processing.

## INTRODUCTION

The *N*-methyl-D-aspartate receptor (NMDAR) is a critical glutamate-gated ion channel that plays a fundamental role in various brain functions, particularly in regulating synaptic strength through processes like long-term potentiation (LTP) and long-term depression (LTD), which are essential for memory formation.¹,² Co-agonist binding of glycine or D-serine, in addition to glutamate, is required for the ionotropic activity of NMDARs.³ While the ion channel function is well established,³,⁴ an unconventional ion-flux-independent signaling mode of NMDARs promoting synapse depression and spine shrinkage has been described,¹,⁵⁻¹³ although this evidence has been challenged.¹⁴ Notably, this unconventional NMDAR function is characterized by glutamate binding in the absence of co-agonist binding.⁹,¹²

How this mode of NMDAR activity is controlled endogenously remains unknown. Here, we identified AETA ($A_η$) as a molecular key that controls both ionotropic and ion-flux-independent NMDAR signaling. AETA is a recently discovered peptide derived from amyloid-$β$ precursor protein (APP) cleavage, with potential bioactivity at excitatory synapses.¹⁵,¹⁶ Following the shedding of the APP ectodomain by $η$-secretase and subsequent cleavage by $α$- or $β$-secretases, long and short forms of AETA are generated ($A_η-α$ and $A_η-β$, respectively) (Figure 1A) and subsequently secreted into the interstitial space.¹⁶ Our findings demonstrate that AETA competes with the co-agonists of NMDARs and modifies the conformation of the GluN1 intracellular tails. AETA holds a unique capacity to impact both NMDAR functions, as it inhibits NMDAR ionotropic activity and associated calcium entry while favoring ion-flux-independent
A η-Secretase Pathway

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synapse weakening and spine shrinkage. Furthermore, we pro-
vide evidence that AETA production is activity-dependent and
essential for proper NMDAR transmission, LTD, and memory
processing. Thus, our study unveils a novel endogenous modu-
lator of NMDARs, shedding light on its crucial role in brain
signaling.

RESULTS

AETA inhibits NMDAR ionotropic activity

Upon the discovery of AETA and its ability to reduce hippocam-
pal LTP,¹⁵,¹⁶ a process of synaptic potentiation that depends on
NMDARs for induction and AMPA receptors (AMPARs) for
expression, we first asked if AETA could directly modulate the
activity of these glutamatergic receptors. We expressed recom-
binant forms of NMDARs (specifically GluN1/GluN2A or GluN1/
GluN2B subtypes) or AMPARs (GluA1/GluA2 or GluA2/GluA3
subtypes) in Xenopus oocytes. We investigated the acute effects
of AETA (100 nM) or a control peptide (CtrlP) (100 nM) on the cur-
rent amplitudes of AMPARs and NMDARs. While AMPAR cur-
rents remained unaltered in the presence of AETA, there was a
significant inhibition of NMDAR currents (Figure 1B). To further
characterize this effect, we conducted a dose-response curve
to determine the impact of varying concentrations of AETA on
NMDAR currents. The results revealed a dose-dependent reduc-
tion in NMDAR current, allowing us to calculate the half-maximal
inhibitory concentration (IC50) values for both receptor subtypes
(1.62 nM for GluN1/N2A; 15.45 nM for GluN1/N2B) (Figure S1A).
These findings confirm the direct modulatory action of AETA on
NMDARs. Next, we evaluated the influence of AETA on native
NMDAR in cultured neurons and in hippocampal slices. We pre-

viously reported bioactivity of AETA within a low nanomolar con-
centration range (5–10 nM).¹⁵ To ensure the specificity of our
findings and avoid any potential nonspecific effects, we con-
ducted all subsequent analyses on native receptors using a con-
centration of 10 nM AETA. Notably, this concentration also in-
hibited recombinant GluN1/N2B NMDARs in oocytes (62.98 ±
8.28%; normalized to averaged baseline; data not shown). In
cultured neurons, application of NMDA puffs resulted in currents
that remained unchanged after a 10-min incubation with CtrlP
(Figure S2A). By contrast, the presence of AETA led to a progres-
sive decrease in current amplitude, which eventually reached a
plateau at 55% of the original amplitude (Figure S2A). Similarly,
in hippocampal slices from young adult mice, native synaptic
NMDAR currents recorded at the CA3–CA1 synapse (comprising
approximately 80% GluN1/N2A and 20% GluN1/N2B sub-
types¹⁷) were reduced by AETA reaching a plateau at 65% of
the original amplitude, while no change was observed with CtrlP
(Figures 1C and S2B). Using pharmacological blockage of
GluN2A- or GluN2B-containing NMDARs with TCN-201 or ifen-
prodil, respectively, we observed that AETA’s effect was stron-
ger on native GluN2A-containing than on GluN2B-containing
NMDARs (Figure S2C). Conversely, native synaptic AMPAR cur-
rents measured in hippocampal slices remained unaffected by
AETA (Figure S3A). In addition, spontaneous excitatory postsyn-
aptic currents (sEPSCs) mediated by NMDARs were also
impacted by AETA but not by CtrlP, leading to a decrease in
NMDAR sEPSC frequency (Figures S4 and S5). Again, sEPSCs
mediated by AMPARs showed no significant alterations
(Figures S3B–E and S5). The paired-pulse ratio (PPR), which
serves as a measure of presynaptic short-term plasticity, re-
mained unaltered (Figure S3F), providing evidence against any

**Figure 1. AETA inhibits NMDA receptor ionotropic activity**

(A) Depicted is the η-secretase-dependent processing of APP. Shedding occurs within the luminal ectodomain, liberating sAPP-η, while the membrane-bound C-terminal fragment (CTF-η) is further processed alternatively by either α- or β-secretases releasing longer or shorter forms of AETA, respectively (Aη-α and Aη-β).

(B) (Top) Representative current traces quantified in oocytes expressing recombinant AMPARs containing GluA1/GluA2 (left) or NMDARs containing GluN1/GluN2B (right) before (black) and after (red) application of AETA (100 nM). NMDAR recordings were made in the presence of 100 µM L-glutamate/1 µM glycine. AMPAR recordings were made in the presence of L-glutamate (300 µM) and cyclothiazide (CTZ) (100 µM). (Bottom) Graph represents % peak current (normalized to averaged baseline) of AMPARs (either GluA1/GluA2 or GluA2/GluA3) and NMDARs (either GluN1/GluN2A or GluN1/GluN2B) after application of either AETA (100 nM) or control peptide (CtrlP) (100 nM). *n* = between 8 and 11 independent oocytes per condition.

(C) (Top) Representative traces of NMDAR currents recorded at CA3-CA1 synapses of adult mouse slices before (baseline, black) and after application of CtrlP (10 nM, gray) or AETA (10 nM, red). (Bottom) Bar graph of AETA or CtrlP effect on NMDAR EPSC calculated 25-30 min post application (full time course shown in Figure S2B). *n* = number of patched neurons per condition.

(D) Two-photon image of CA1 hippocampal neuron filled with Alexa 594 (red) and Fluo-5F (green). Boxed area is enlarged on right with uncaging spot indicated in yellow and line scan indicated in white. Dual channel line scan images with response to five uncaging events (50 Hz, yellow dots) shown below.

(E) Line scan profiles across spine heads (scale bars: 100 ms/ΔF/2 a.u.) and simultaneous electrophysiological recordings of uEPSP (scale bars: 50 ms/0.5 mV) in response to five uncaging events before (black) and after D-APV (100 µM, blue), AETA (10 nM, red), or CtrlP (10 nM, gray).

(F) Peak spine calcium transients are depressed after 10 min of 10 nM AETA application (n = 6 neurons) but not after CtrlP application (n = 5).

(G and H) Effect of AETA (100 nM) on NMDAR current in oocytes expressing GluN1/GluN2B in the presence of different concentrations of glutamate (1; 10 or 100 µM) (G) or glycine (0,1; 1 or 10 µM) (H). *n* = oocytes per condition.

(I) Dose-response activation curves for glycine in absence or presence of AETA (100 nM) measured in oocytes expressing either GluN1/GluN2A (left) or GluN1/
GluN2B (right).

(J) Dose-response curve of AETA's effect (10 nM) on native NMDAR current at CA3-CA1 synapses in mouse hippocampal slices measured by patch clamp in the presence of different concentrations of D-serine (0, 1, 3, 10, 30, and 100 µM) in recording solution. *n* = neurons patched per condition.

(K) (Left) Time course of AETA's effect (10 nM) on NMDAR currents at CA3-CA1 synapses in mouse hippocampal slices measured during different washout protocols. Protocol 1, 20 min AETA (10 nM) followed by washout with aCSF; protocol 2, 20 min AETA (10 nM) followed by addition of D-serine (100 µM) but in continuous presence of AETA; protocol 3, 20 min AETA (10 nM) followed by substitution with D-serine (100 µM). (Right) Time course data plotted as a bar graph showing % NMDAR current measured during last 5 min for each protocol (normalized to averaged baseline current of that protocol).

Error bars represent standard error of the mean (s.e.m.); \* *p* < 0.05; \*\* *p* < 0.01; \*\*\* *p* < 0.001; \*\*\*\* *p* < 0.0001. Statistics: One-way ANOVA followed by Tukey's multiple comparisons test (B, G, H, J, and K); Mann-Whitney test (C); Paired Student's t test (F). See supplemental statistics (Data S1) for full statistics. See also Figures S1–S5 for additional experiments related to Figure 1.
A

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B

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C

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CtrlP

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AETA

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Clusters

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Cells

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D

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E

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F

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G

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H

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CtrlP

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I

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J

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presynaptic effect of AETA on release probability. Collectively, the data, obtained from three distinct analysis systems, consistently support the conclusion that AETA acts directly and acutely at the postsynaptic site, specifically inhibiting NMDAR transmission while not affecting AMPAR transmission.

Given AETA's ability to inhibit NMDAR current, it is expected that it would also reduce NMDAR-dependent calcium entry into dendritic spines, thereby preventing NMDAR ionotropic signaling. To investigate this, we employed two-photon single-spine calcium imaging in conjunction with glutamate uncaging to evoke unitary excitatory postsynaptic potential (uEPSP) recorded at CA3-CA1 synapses in hippocampal slices. By monitoring the activity of individual spines upon glutamate uncaging (Figures 1D and 1E), we confirmed that AETA, but not CtrlP, significantly attenuated NMDAR-dependent calcium entry (Figure 1F) without attenuating EPSP amplitude or changing the resting membrane potential (data not shown). These findings provide further evidence that AETA effectively inhibits NMDAR ionotropic signaling by impeding calcium influx in the presence of glutamate stimulation.

## AETA competes with NMDAR co-agonist to inhibit ionotropic activity

The activation of NMDARs involves complex conformational re-arrangements across extracellular, transmembrane, and cytosolic receptor domains, requiring the binding of glutamate and glycine/D-serine as co-agonists.³,⁴ To investigate the mechanism of AETA's inhibitory activity, we conducted experiments using oocytes to assess its dependence on the concentration of these two ligands. Modulating the levels of glutamate in the recording bath prior to AETA application did not alter its effect (Figure 1G), indicating that glutamate availability did not influence AETA's inhibitory action. However, increasing the levels of glycine significantly diminished the impact of AETA (Figure 1H). These findings suggest that AETA competes with the co-agonist

at the glycine/D-serine binding site. To further validate this hypothesis, we performed dose-response curves for glycine in the absence and presence of AETA on both GluN1/GluN2A and GluN1/GluN2B NMDAR receptor subtypes. AETA shifted these curves to the right and increased the half-maximal effective concentration (EC50) for glycine (Figures 1I, S1B, and S1C), indicating a competitive binding of AETA at the co-agonist site. To confirm the competitive action of AETA on native synaptic NMDARs, we investigated its effect on isolated NMDAR currents of CA3-CA1 synapses in hippocampal slices at varying concentrations of D-serine. We used D-serine instead of glycine when recording NMDAR currents in CA1 neurons because D-serine is the primary co-agonist for native synaptic NMDAR receptors.¹⁸ Increasing the dose of D-serine in the recording bath progressively attenuated the effect of AETA, completely blocking it at 30–100 µM (Figure 1J). We also examined the duration of AETA's effect after the removal of the peptide and its susceptibility to competition by D-serine (Figure 1K). The inhibition of NMDAR current by AETA persisted for at least 20 min after its removal, suggesting that AETA binds to and modifies NMDAR activity in a lasting manner. Moreover, the addition of D-serine (100 µM) to AETA did not relieve the inhibition of NMDARs, indicating that AETA binds to NMDARs with higher affinity than D-serine. Conversely, replacing AETA with D-serine restored NMDAR activity, confirming the competitive nature of AETA's interaction with the co-agonist.

## AETA alters NMDAR conformation

Previous studies showed that agonist and co-agonist binding induce conformational changes in the cytosolic domains of NMDARs, affecting both ionotropic and non-ionotropic signaling pathways.⁷,¹⁹ Given the role of AETA as a potential endogenous regulator of NMDAR co-agonist binding, we investigated whether AETA elicits similar conformational rearrangements. To explore this, we utilized fluorescence lifetime imaging

### Figure 2. AETA modifies NMDAR conformation, enhances LTD via non-ionotropic activity of NMDARs, and promotes spine shrinkage

(A) Schematic principle of intramolecular FLIM-FRET experiment. Hippocampal neurons were transfected either with GluN1-GFP alone (donor only, upper panel) or in combination with GluN1-mCherry (donor + acceptor, lower panel; all constructs co-transfected with GluN2B-flag). Upon excitation of the GFP fluorophore with blue light, the proximity of GluN1-GFP and GluN1-mCherry within a receptor and overlap of GFP emission and mCherry excitation spectra allow resonance energy transfer (black arrow) from the donor fluorophore (GFP) to the acceptor fluorophore (mCherry), causing excitation of the acceptor fluorophore (red arrow) and a subsequent decrease in the fluorescence lifetime of the donor fluorophore (green arrows).

(B) Representative illustrations of GFP lifetime in neuronal fields (upper panels) and GluN1-GFP only (donor only) and GluN1-GFP/GluN1-mCherry (D + A) dendritic spine clusters (lower panels) 10 min after exposure to CtrlP or AETA (10 nM). Graph represents GFP lifetime in GluN1-GFP only (Don. only) and GluN1-GFP/GluN1-mCherry (Don. + Acc.) clusters 10 min after exposure to CtrlP (10 nM; Don. only, n = 738 clusters; Don. + Acc., n = 432 clusters) or AETA (10 nM; Don. only, n = 796 clusters; Don. + Acc., n = 299 clusters).

(C) (Left) FRET efficiency in GluN1-GFP/GluN1-mCherry clusters 10 min after exposure to CtrlP (10 nM; n = 432 clusters) or AETA (10 nM; n = 299 clusters). (Right) FRET efficiency in GluN1-GFP/GluN1-mCherry clusters averaged per cell 10 min after exposure to CtrlP (10 nM; n = 23 cells) or AETA (10 nM; n = 18 cells). (Top) Diagram of movement of GluN1 intracellular tail in control condition (CtrlP) and in the presence of AETA.

(D) Representative traces (left, 1 shows trace pre- and 2 shows trace post-induction) and summary graph (right) of fEPSP slope (% baseline) pre- and post-LTD induction (time 0) in control (Ctrl, aCSF only) or in the presence of AETA (10nM) throughout recording.

(E) Summary of fEPSP magnitude 45–60 min after LTD induction as fEPSP (% baseline) for data shown in (D). n/N = number of slices/mice.

(F–G) same as in (D–E) but in the presence of 100 µM MK801 (3 h pre-incubation and throughout recording).

(H) (Left) Low-magnification image of GFP-labeled CA1 pyramidal neuron from P18-21 GFP-M mice. (Right) High-magnification images of basal dendrites of CA1 pyramidal neurons from GFP-M mice before (time 0) and after (1 and 30 min) high-frequency glutamate uncaging (HFU, yellow cross) at an individual dendritic spine (yellow arrowhead) in the presence of 10 nM CtrlP or AETA.

(I–J) HFU-induced spine growth in the presence of CtrlP (gray filled circles/bar; n/N = 7 cells/6 mice) was converted to spine shrinkage in the presence of AETA (red filled circles/bar; n/N = 7 cells/6 mice). The volume of unstimulated neighboring spines (open circles/bars) was unchanged.

Error bars represent s.e.m.; \* *p* < 0.05; \*\* *p* < 0.01; \*\*\* *p* < 0.001; \*\*\*\* *p* < 0.0001. Statistics: Kruskal-Wallis test followed by Dunn's multiple comparisons test (B); Mann-Whitney test (C); unpaired Student's t test (E and G); two-way ANOVA with Bonferroni's multiple comparisons test (J). See supplemental statistics (Data S1) for full statistics. See also Figures S6 and S7 for additional experiments related to Figure 2.
A

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B

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C

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D

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E

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F

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microscopy (FLIM) in hippocampal neurons expressing recombi-
nant GluN1-NMDAR subunits. Förster resonance energy transfer
(FRET) was measured between C-terminal domains labeled with
green fluorescent protein (GFP) or mCherry fluorophores as an
indicator of intracellular conformational changes (Figures 2A
and 2B).⁷,¹⁹ As expected, the lifetime decreased between the
“donor only” and “donor/acceptor” conditions in the presence
of CtrlP or AETA peptides (Figure 2B, graph). Exposure to CtrlP
for 10 min resulted in fluorescence lifetime values comparable
to those reported in buffer,¹⁹ indicating no significant effect on
conformational changes (Figure 2C). In contrast, AETA exposure
for the same duration induced a substantial increase in FRET ef-
ficiency (Figure 2C). This conformational change is independent
of agonist binding, as AETA equally increases FRET efficiency
in the presence of the NMDAR antagonist APV (50 µM)
(Figures S6A and S6B). These findings demonstrate that, simi-
larly to co-agonists, AETA binding triggers conformational
rearrangements in the C-terminal domains of GluN1, bringing
them into closer proximity in a receptor-activity-independent
manner (Figure 2C, diagram). These rearrangements putatively
modulate associated cytosolic protein complexes and signaling
pathways. There is evidence that activation of the p38 kinase
via its phosphorylation represents a downstream signaling
pathway of NMDARs, notably in the context of ion-flux-indepen-
dent signaling.⁹,¹³,²⁰ In line with these data, we found that AETA
increased p38 phosphorylation in cultured neurons (Figures S6C
and S6D). This suggests that AETA might favor NMDAR ion-flux-
independent signaling.

AETA enhances LTD via non-ionotropic activity of NMDARs and promotes spine shrinkage

The reduced ionotropic signaling of NMDARs, which includes
decreased calcium entry, is consistent with the impairment of
LTP caused by AETA.<sup>15,16</sup> However, it is important to consider
that a decrease in NMDAR-dependent calcium signaling could
also affect LTD, as sustained low levels of NMDAR-dependent
calcium entry have been shown to induce LTD.<sup>2</sup> Alternatively,
the altered conformation of NMDARs caused by AETA and in-
crease in p38 phosphorylation may promote non-ionotropic
NMDAR signaling, which has been reported to also play a critical

role in LTD.<sup>7,9,12,20,21</sup> We thus examined the impact of AETA
on LTD at the CA3-CA1 synapse in hippocampal slices
(Figures 2D and 2E). We found that AETA enhanced the magni-
tude of LTD induced by 900 stimuli at 1 Hz. Additionally, even
with a subthreshold LTD induction protocol of 300 stimuli at
1 Hz that did not induce LTD in control conditions, the presence
of AETA still resulted in the production of LTD (Figures S7A and
S7B). These findings demonstrate that AETA favors LTD, poten-
tially through its influence on NMDAR conformation and non-ion-
otropic signaling pathways rather than solely through alterations
in calcium signaling. Despite some controversy,<sup>8-11,14</sup> evidence
suggests that LTD can be mediated by NMDAR activity without
requiring ion flux, as demonstrated by the occurrence of LTD in
the presence of the ion channel blocker MK801.<sup>9</sup> To validate this,
we replicated the conditions reported previously<sup>9</sup> and observed
LTD at CA3-CA1 synapses in rat hippocampal slices in the pres-
ence of MK801 (Figures S7C and S7D). Next, we examined
whether enhanced LTD mediated by AETA could still occur in
the presence of MK801. In mouse slices, we preincubated the
slices with MK801 for 3 h, confirming that this application of
MK801 completely blocked the ionotropic function of NMDARs
as expected (Figure S7E). Notably, also in mouse slices, we
observed LTD in the presence of MK801 (Figures 2F and 2G).
AETA still enhanced this ion-flux-independent LTD (Figures 2F
and 2G). These data provide evidence that AETA binding
promotes ion-flux-independent NMDAR activity, which medi-
ates LTD.

Synaptic plasticity at excitatory glutamatergic synapses is
closely linked to structural changes in dendritic spines, wherein
synaptic weakening is accompanied by spine shrinkage.<sup>22,23</sup>
Previous studies have provided evidence that glutamate binding
to NMDARs in the absence of ion flux is sufficient to mediate
LTD-induced spine shrinkage,<sup>12,20</sup> and p38 activity is involved
in this phenomenon.<sup>13,20</sup> Moreover, a decrease in co-agonist
levels has been shown to bias spine structural plasticity toward
shrinkage.<sup>24</sup> Given AETA's ability to compete with co-agonist
binding and to increase p38 phosphorylation, we hypothesized
that AETA could promote spine shrinkage. To test this hypothe-
sis, we employed a high-frequency glutamate uncaging (HFU)
protocol typically associated with spine growth at single

<caption>Figure 3. Endogenous increase of AETA by *in vivo* BACE1 inhibition leads to increased LTD, and endogenous AETA production increases with *in vivo* neuronal activity</caption>

(A) Diagram of experimental plan. The BACE1 inhibitor LY2811376 (or saline) was administered by gavage 12 h before sacrificing the mouse. Brains were removed to prepare hippocampal slices for electrophysiology (D–E). The rest of the brains were used for immunoblotting to check for increase in AETA (B–C).

(B) Ponceau staining and immunoblot (M3.2 antibody) of AETA from saline- and LY2811376-treated brains.

(C) Quantification of levels of AETA in saline- and LY2811376-treated brains (normalized to Ponceau). N = number of mice.

(D) Representative traces (left, 1 shows trace pre- and 2 shows trace post-induction; scale bars: 10 ms/0.2 mV) and summary graph (right) of fEPSP slope (% baseline) pre- and post-LTD induction (time 0) at CA3-CA1 synapses in hippocampal slices from saline- and LY2811376-treated mice.

(E) Bar graph of fEPSP magnitude 45–60 min after LTD induction as fEPSP (% baseline) for data shown in (D). n/N = slices/mice.

(F) Diagram of experimental design to activate or inhibit neurons *in vivo* using AAV-hM3Dq-mCherry or AAV-hM4Di-mCherry, respectively, and quantify AETA levels.

(G) (Top) Example of immunoblot showing detection of mCherry, AETA, and β-actin in hM3Dq-transduced tissue after saline or CNO i.p. injection. (Bottom) Quantification of AETA in hM3Dq-SAL and hM3Dq-CNO (normalized to β-actin and mCherry). All quantified blots are provided in Figures S9C and S9D. N = number of mice.

(H) (Top) Example of immunoblot showing detection of mCherry, AETA, and β-actin in hM4Di-transduced tissue after saline or CNO i.p. injection. (Bottom) Quantification of AETA in hM4Di-SAL and hM4Di-CNO (normalized to β-actin and mCherry). All quantified blots are provided in Figures S9E and S9F. N = number of mice.

Error bars represent s.e.m.; \* *p* < 0.05; \*\* *p* < 0.01. Statistics: unpaired Student's t test (C and E); Mann-Whitney test (G and H). See supplemental statistics (Data S1) for full statistics. See also Figures S8 and S9 for additional experiments related to Figure 3.
<figure><img src="image_44.png" /><figcaption>B WT APPdelETA</figcaption></figure>

<figure><img src="image_45.png" /><figcaption>C AETA level</figcaption></figure>

<figure><img src="image_46.png" /><figcaption>D CTF-η level</figcaption></figure>

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dendritic spines on basal dendrites of CA1 hippocampal neurons in acute slices from GFP-M mice, where GFP is selectively expressed in a subset of hippocampal pyramidal neurons (Figure 2H).²⁴ As expected, the HFU protocol led to dendritic spine growth in the presence of the CtrlP, whereas the presence of AETA instead induced spine shrinkage (Figures 2I and 2J). These results show shrinkage under conditions that would normally be expected to induce long-term spine growth, consistent with AETA's promoting non-ionotropic NMDAR signaling.

## Increasing endogenous AETA levels with BACE1 inhibition decreases NMDAR activity and enhances LTD

The experiments described above rely on exogenous application of synthetic AETA. To provide stronger *in vivo* evidence for these findings, we asked if increasing endogenous AETA levels could also lead to alterations in NMDAR activity and synapse plasticity. There is no pharmacological compound that specifically targets AETA production. Yet, we previously reported that BACE1 inhibition leads to an increase in endogenous AETA.¹⁶ We thus inhibited BACE1 *in vivo* with oral gavage of the BACE1 inhibitor LY2811376 (Figure 3A).²⁵ This indeed led to a 2-fold increase in endogenous AETA (Figures 3B and 3C). We prepared hippocampal slices from these mice and kept them in LY2811376 throughout the recordings. We observed that increased AETA correlated with decreased NMDAR sEPSC frequency, but not amplitude (Figure S8), and increased LTD (Figures 3D and 3E), thus reproducing exactly the phenotypes observed with acute application of synthetic AETA.

## AETA production is enhanced by neuronal activity

We here demonstrated that AETA functions as a novel modulator of NMDARs. To further establish its classification as an endogenous neuromodulator, we investigated whether AETA levels can be regulated by neuronal activity, as is the case for known neuromodulators. We employed chemogenetic designer receptor

exclusively activated by designer drugs (DREADD) proteins hM3Dq and hM4Di to manipulate neuronal activity *in vivo*. Neurons in the prefrontal cortex were transduced with an adeno-associated virus (AAV) expressing either hM3Dq or hM4Di along with mCherry (Figures 3F, and S9A, and S9B). After three weeks of expression, mice were injected with either saline (control) or clozapine N-oxide (CNO) to activate or inhibit the transduced neurons (Figure 3F). Thirty minutes after CNO injection, we microdissected the mCherry-labeled region of the prefrontal cortex and quantified AETA levels in these tissues using immunoblotting. Our results revealed a significant 4-fold increase in AETA levels in samples subjected to neuronal activation (Figures 3G, S9C, and S9D). By contrast, inhibition of neuronal activity did not alter AETA levels (Figures 3H, S9E, and S9F). Viral expression did not otherwise alter markers of synapse integrity (Figures S9G and S9H). Therefore, AETA production is specifically linked to an elevation in neuronal activity.

## AETA is necessary for adequate NMDAR ionotropic activity, LTD, and memory processing

To further establish AETA as an activity-dependent regulator of NMDARs, we conducted experiments to determine its necessity for this signaling mechanism. Utilizing CRISPR-Cas9 gene editing, we generated a novel mouse model, termed the APPdelETA mouse, in which the η-secretase cleavage site of the endogenous APP was deleted (Figure 4A). This expectedly resulted in a smaller mutant APP (Figure 4B). Consequently, AETA and its membrane-bound precursor peptide CTF-η were absent in these mice (Figures 4B-D and S10A-C). Recording NMDAR-mediated sEPSCs in CA1 pyramidal neurons of the APPdelETA mice, we observed higher frequency (Figures 4E and 4F) but normal amplitude (Figure S10F) of these currents compared with control littermates. Importantly, we were able to normalize NMDAR sEPSC frequency by the acute exogenous application of AETA (Figures 4E and 4F). By contrast, neither AMPAR

### Figure 4. APPdelETA mice display altered NMDAR transmission, loss of LTD, and reduced memory

(A) Depicted is the prevention of η-secretase-dependent processing of APP due to a 41-amino-acid in-frame deletion (marked in red; CRISPR/Cas9 gene editing) in the APPdelETA mouse model. Due to this deletion, no η-secretase shedding occurs (no sAPP-η), and the membrane-bound C-terminal fragment (CTF-η) and AETA peptides are not produced in homozygous APPdelETA mice.

(B) Example of an immunoblot of hippocampal tissue showing detection of full-length APP (FL-APP), β-actin (loading control), and AETA in WT and APPdelETA hippocampal lysates. Note that FL-APP is smaller in size in APPdelETA tissue due to endogenous deletion.

(C) Quantification of endogenous AETA levels in WT and APPdelETA hippocampi, normalized to β-actin levels. All quantified blots are provided in Figures S10A and S10B. N = number of mice.

(D) Quantification of endogenous CTF-η levels (precursor of AETA) in WT and APPdelETA hippocampi, normalized to β-actin levels. Quantified blot is provided in Figure S10C. N = number of mice.

(E) Representative traces of NMDAR sEPSCs recorded in CA1 pyramidal neurons of slices from WT mice and APPdelETA mice and in APPdelETA mice in the presence of AETA (10 nM) in the recording bath.

(F) NMDAR sEPSC frequency calculated from traces as shown in (E). n/N = neurons/mice.

(G) Traces show ten consecutive synaptic responses recorded at -65 mV and +40 mV evidencing responses (black) and failures (gray). Bar graph shows calculated percentage of silent synapses in WT and APPdelETA neurons. n/N = neurons/mice.

(H) Representative traces (left, 1 shows trace pre- and 2 shows trace post-induction; scale bars: 10 ms/0.2 mV) and summary graph (right) of fEPSP slope (% baseline) pre- and post-LTD induction (time 0) at CA3-CA1 synapses in hippocampal slices of WT and APPdelETA mice without or with supplementation of 10 nM AETA (in recording bath).

(I) Bar graph of fEPSP magnitude 45-60 min after LTD induction as fEPSP (% baseline) for data shown in (H). n/N = slices/mice.

(J) Diagram of contextual fear conditioning behavioral task.

(K) Graph represents % freezing measured during 6 min of training session in the three genotypes. N = number of mice.

(L) Graph represents % freezing measured during 6 min of test session done 24 h after training session.

Error bars represent s.e.m.; \* *p* < 0.05; \*\* *p* < 0.01; \*\*\* *p* < 0.001. Statistics: Mann-Whitney test (C and D); one-way ANOVA followed by uncorrected Fisher's LSD test (F and L); unpaired Student's t test (G); one-way ANOVA followed by Tukey's multiple comparisons test (I); two-way ANOVA (K). See supplemental statistics (Data S1) for full statistics. See also Figures S10-S14 for additional experiments related to Figure 4.
sEPSCs (Figure S10D and S10E) nor the PPR (Figure S10G) showed alterations in these AETA-depleted neurons. Higher NMDAR sEPSC frequency in APPdelETA neurons could be due to an increase in NMDAR-only synapses (called silent synapses). Using a minimal stimulation protocol,²⁶ we quantified these silent synapses in wild-type (WT) and APPdelETA mice. We observed an increase in the percentage of silent synapses in APPdelETA mice (Figures 4G and S11). Yet, this alteration in the number of silent synapses was not correlated to an increase in the number of spines in these neurons as quantified by Golgi-Cox staining (Figure S12). Moreover, LTD could not be induced in hippocampal slices at CA3–CA1 synapses of adult APPdelETA mice, and this deficit was rescued by the acute exogenous application of AETA (Figures 4H and 4I). LTD could not be rescued by a competitive antagonist of the glycine/D-serine site (L689,560) (Figure S13), suggesting that AETA acts differently from these types of pharmacological compounds. By contrast, LTP was normal at these AETA-depleted synapses (Figures S10H and S10I). Finally, to assess the impact of loss of η-secretase-dependent processing of APP on NMDAR-dependent memory processes, we subjected these mice to contextual fear conditioning, a task known to rely on the hippocampus and NMDARs (Figure 4J). Notably, APPdelETA mice exhibited normal behavior during training but reduced freezing behavior 24 h after training compared with control littermates (Figures 4K and 4L). To rescue this deficit, we crossed APPdelETA mice with another mouse line, the AETA-m line, which harbors a transgene expressing a secreted form of human AETA in the brain (Figure S14). Re-expression of AETA normalized the memory phenotype (Figure 4L). Collectively, these data strongly support the notion that AETA is indispensable for proper hippocampal NMDAR-dependent information processing.

DISCUSSION

In this study, we identified AETA as a novel activity-dependent modulator of NMDARs that competes with the co-agonists glycine/D-serine. Our findings provide compelling evidence that AETA is capable of modifying the conformation of NMDARs and possesses a unique dual role as a molecular key. It reduces NMDAR ionotropic activity and associated calcium-dependent signaling, while concurrently promoting ion-flux-independent activity, thereby permitting LTD of synapse strength, and facilitates spine shrinkage. A diagram is shown in Figure S15, where we summarize this new AETA-dependent NMDAR activation mechanism. While numerous molecules have been identified as NMDAR modulators,²⁷,²⁸ to the best of our knowledge, none exhibit the distinct dual property observed with AETA. In fact, until this discovery, no molecule with the property to activate the ion-flux-independent mode of NMDAR signaling had ever been identified. Through our knockout strategy utilizing the APPdelETA mouse model and subsequent rescue experiments via acute application or *in vivo* expression of AETA, we have demonstrated that AETA controls ionic NMDAR function and is both necessary and sufficient for maintaining adequate non-ionotropic NMDAR function that permits LTD at the CA3–CA1 synapse and adequate memory processing. Given that APP, the precursor of AETA, is among the most

abundant proteins in synaptic boutons,²⁹ it occupies an ideal position as a reservoir for AETA, allowing for this activity-dependent neuromodulatory function.

AETA exerts a depressive effect on NMDAR current and calcium influx within spines. This ionotropic activity holds critical importance for synaptic integration and LTP, which in turn have far-reaching implications for dendritic computations,³⁰ sensory perception,³¹ and adaptations to neuronal representations.³² Consequently, the release of AETA as a neuromodulator may have broad impacts on neuronal function, influencing various aspects of behavior. Our findings demonstrate that AETA levels regulate the frequency of NMDAR-mediated sEPSCs without affecting their amplitude. Notably, the modulation of AETA levels does not alter AMPAR sEPSCs or the PPR. These observations suggest that AETA specifically modulates the activity of distinct NMDAR-only clusters (silent synapses), although the precise identity and position of these clusters remains unknown, representing an intriguing area for future investigation. Pharmacological analysis suggests that AETA preferably inhibits NMDARs containing the GluN2A subunit, but future work will be essential to identify its inhibitory activity on the multitude of heteromeric NMDARs.

The importance of the ion-flux-independent activity of NMDAR for LTD remains a subject of debate.⁵,⁸,¹⁰,¹¹,¹⁴ Our findings provide a potential explanation for these conflicting reports, as the observation of non-ionotropic signaling of NMDARs may be dependent on the degree of AETA release and its competitive interaction with glycine/D-serine, which might depend on the experimental preparations. Furthermore, data obtained from the APPdelETA mouse model suggest that the mechanisms underlying LTP and LTD can be further dissociated, with AETA being necessary for the induction of LTD but not essential for LTP induction or expression. However, it should be noted that we previously observed a partial inhibition of LTP upon the application of AETA to control slices.¹⁵,¹⁶ In physiological conditions, it is plausible that an acute increase in AETA levels would result in a subset of NMDARs being driven into an LTD-prone state due to reduced co-agonist binding. As a consequence, these NMDARs become less available to contribute to LTP, thereby reducing the capacity of these synapses to express LTP. Our argument finds strong support in our data on spine shrinkage, where AETA transforms the expected spine growth into spine shrinkage. While we did not directly show that AETA-induced spine shrinkage is independent of ion flux via NMDARs, this AETA effect is identical to what has been observed in the presence of pharmacological antagonists of the glycine/D-serine binding site, 7CK and L698,560,¹²,²⁰ and there is strong evidence that this action is driven by ion-flux-independent NMDAR signaling and mediated by phosphorylation of p38, which is increased by AETA.

Our findings contribute to the growing body of evidence linking APP to NMDAR function, including NMDAR alterations observed during aging.¹⁷,²⁵,³³,³⁴ However, there is still limited information regarding the role of AETA in pathological mechanisms. Given the close association between APP processing and the etiology of Alzheimer's disease (AD),³⁵ there is emerging evidence suggesting that AETA might accumulate in patients with AD.¹⁶,³⁶ With the discovery of this newly identified AETA-driven
mechanism, it becomes crucial to investigate whether it contrib-
utes to NMDAR-dependent synapse dysfunction and spine loss,
both of which are considered early events in AD progres-
sion.³⁷⁻³⁹ Moreover, beyond AD, the endogenous mechanism
of action of AETA on NMDARs identified in our study could be
disrupted in various other neurodegenerative, neurological,
and psychiatric disorders involving this receptor. For instance,
abnormal NMDAR function has been implicated in neurodegen-
erative diseases such as Parkinson's and Huntington's dis-
eases,⁴⁰,⁴¹ traumatic brain injury,⁴² stroke,⁴³ epilepsy,⁴⁴ autism
spectrum disorder and intellectual disability,⁴⁵ anti-NMDA re-
ceptor encephalitis and anti-GluN2 antibodies associated with
systemic lupus erythematosus,⁴⁶ schizophrenia, and depres-
sion.⁴⁷ Our discovery provides the impetus to explore the poten-
tial involvement of AETA-driven regulation of NMDARs in the
pathophysiology of these disorders.

STAR★METHODS

Detailed methods are provided in the online version of this paper and include
the following:

● KEY RESOURCES TABLE

● RESOURCE AVAILABILITY

○ Lead contact

○ Materials availability

○ Data and code availability

● EXPERIMENTAL MODEL AND STUDY PARTICIPANT DETAILS

○ Oocytes

○ Mice and rats

○ Primary neuronal cultures

● **METHOD DETAILS**

○ Peptides

○ Electrophysiology in *Xenopus* oocytes

○ Electrophysiology and p38 activity in neuronal cultures

○ Electrophysiology in hippocampal slices

○ uEPSP and calcium transients in single spines

○ FLIM-FRET of GluN1-GFP/GluN1-mCherry FRET pair

○ Two-photon imaging of spine structural plasticity

○ BACE1 inhibition *in vivo*

○ *In vivo* neuronal activation or inhibition by DREADD

○ Generation of APPdelETA mice

○ Generation of AETA-m mouse line

○ Immunoblotting

○ Golgi-Cox staining for spine density analysis

○ Contextual fear conditioning

○ Statistical analysis

SUPPLEMENTAL INFORMATION

Supplemental information can be found online at https://doi.org/10.1016/j.neuron.2024.05.027.

ACKNOWLEDGMENTS

Complementary DNA (cDNA) for expression of recombinant rodent NMDARs
in oocytes were generously provided by Dr. Paoletti (Institut de Biologie de
l'Ecole Normale Supérieure [IBENS], France). cDNA for expression of recom-
binant AMPARs in oocytes were generously provided by Dr. Kristensen (Uni-
versity of Copenhagen, Denmark). We thank Franck Chatelain (IPMC, France)
for technical help with Xenopus oocyte recordings. We thank Maxime Villet
(IPMC, France) for technical help with pictures of virus spread. We thank
Prof. Haass (German Center for Neurodegenerative Diseases Munich;
Biomedical Center, Division of Metabolic Biochemistry, Faculty of Medicine,

Ludwig-Maximilians-Universität München, Germany) for financial support and insightful discussions. This work was supported by the Centre National de la Recherche Scientifique (CNRS); the French Association France Alzheimer (AAP SM 2018 Dossier 1795) to H.M.; the French Fondation Alzheimer (AAP 2015 AETAPHYS) to H.M.; the Agence Nationale de la Recherche (ANR-21-CE16-0032 APPYSYNAPSE) to H.M. and L.G.; the Flag ERA JTC 2019 (MILEDI Project) to S.M. and H.M.; the Investments for the Future LABEX SIG-NALIFE #ANR-11-LABX-0028-01 to M.M.; the Fondation pour la Recherche Médicale (FRM) to L.G.; the European Research Council Synergy grant (ENSEMBLE, #951294) to L.G.; the Institut Neuromod, Université Côte d'Azur, to H.M. and J.R.M.; the MRC (1514380) and EUFP17 Marie Curie Actions (PCIG10-GA-2011-303680) to M.C.A.; the Wellcome Trust (101029/Z/13/ Z) to J.R.M.; the BBSRC (BB/R002177/1) to M. Amici; the NIH (R01 NS062736) to K.Z.; and the Deutsche Forschungsgemeinschaft Walter Benjamin project (468470832) to M. Anisimova. J. Dunot is the recipient of a PhD fellowship from the Ministère de la Recherche, de l'Enseignement Supérieur et de l'Innovation.

AUTHOR CONTRIBUTIONS

J. Dunot, S.M., P.A.P., M.M., and H.M. performed electrophysiological anal-
ysis by field and patch clamp in hippocampal slices. A.W., B.L., C. Gandin,
and H.M. performed analysis on *Xenopus* oocytes. J. Dunot, I.B., and H.M.
performed stereotaxic surgeries. M. Amici, M.C.A., and J.R.M. performed
analysis of EPSP combined with calcium transients in hippocampal slices. J.
Dupuis, M.U., and L.G. performed analysis of NMDAR conformation by
FRET. M. Anisimova, S.J.P., and K.Z. performed analysis of spine structural
plasticity. C. Giudici, M.W., H.H., J. Dunot, and H.M. performed all analysis
by immunoblotting. C. Gandin performed spine density analysis. M.W.,
B.W., R.N., and W.W. created the APPdelETA and AETA-m mouse lines. J. Du-
not performed behavioral analysis. H.M. supervised the full study and wrote
the manuscript with help from other authors.

DECLARATION OF INTERESTS

The authors declare no competing interests.

DECLARATION OF GENERATIVE AI AND AI-ASSISTED
TECHNOLOGIES IN THE WRITING PROCESS

During the preparation of this work, the senior author used ChatGPT in order to
improve readability and language. After using this tool, the author reviewed
and edited the content as needed and takes full responsibility for the content
of the publication.

Received: July 26, 2023

Revised: April 9, 2024

Accepted: May 29, 2024

Published: June 14, 2024

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41. Wang, J., Wang, F., Mai, D., and Qu, S. (2020). Molecular Mechanisms of Glutamate Toxicity in Parkinson's Disease. Front. Neurosci. 14, 585584. https://doi.org/10.3389/fnins.2020.585584.

42. Khormali, M., Heidari, S., Ahmadi, S., Arab Bafrani, M., Baigi, V., and Sharif-Alhoseini, M. (2022). N-methyl-D-aspartate receptor antagonists in improving cognitive deficits following traumatic brain injury: a systematic review. Brain Inj. 36, 1071-1088. https://doi.org/10.1080/02699052.2022.2109749.

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48. Platzer, K., Yuan, H., Schütz, H., Winschel, A., Chen, W., Hu, C., Kusumoto, H., Heyne, H.O., Helbig, K.L., Tang, S., et al. (2017). GRIN2B encephalopathy: novel findings on phenotype, variant clustering, functional consequences and treatment aspects. J. Med. Genet. 54, 460-470. https://doi.org/10.1136/jmedgenet-2016-104509.

49. Mony, L., Zhu, S., Carvalho, S., and Paoletti, P. (2011). Molecular basis of positive allosteric modulation of GluN2B NMDA receptors by polyamines. EMBO J. 30, 3134-3146. https://doi.org/10.1038/emboj.2011.203.

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## STAR★METHODS

### KEY RESOURCES TABLE

<table><thead><tr><th>REAGENT or RESOURCE</th><th>SOURCE</th><th>IDENTIFIER</th></tr></thead><tbody><tr><td colspan="3"><strong>Antibodies</strong></td></tr><tr><td>Anti-rabbit-IgG-HRP conjugated secondary antibody</td><td>Jackson ImmunoResearch</td><td>Cat#111-036-045; RRID: AB_2337943</td></tr><tr><td>Anti-rat-IgG-HRP conjugated secondary antibody</td><td>Jackson ImmunoResearch</td><td>Cat#111-036-062; RRID: AB_2338142</td></tr><tr><td>Anti-mouse-IgG-HRP conjugated secondary antibody</td><td>Jackson ImmunoResearch</td><td>Cat#111-036-146; RRID: AB_2307347</td></tr><tr><td>M3.2</td><td>Biolegend</td><td>Cat#805701; RRID: AB_2564981</td></tr><tr><td>2D8</td><td>Sigma-Aldrich</td><td>Cat#MABN2273</td></tr><tr><td>2E9</td><td>Sigma-Aldrich</td><td>Cat#MABN2295</td></tr><tr><td>22C11</td><td>Merck Milipore</td><td>Cat#MAB348</td></tr><tr><td>Y188</td><td>Abcam</td><td>Cat#ab32136; RRID: AB_2289606</td></tr><tr><td>mCherry</td><td>Takara</td><td>Cat#632496; RRID: AB_10013483</td></tr><tr><td>Synaptophysin</td><td>Sigma-Aldrich</td><td>Cat#S5786</td></tr><tr><td>Synaptobrevin-2</td><td>SYSY</td><td>Cat#104211</td></tr><tr><td>p-38</td><td>Cell Signaling</td><td>Cat#8690; RRID: AB_10999090</td></tr><tr><td>p-p38</td><td>Cell Signaling</td><td>Cat#9216; RRID: AB_331296</td></tr><tr><td>Beta-actin</td><td>Sigma-Aldrich</td><td>Cat#AC-74</td></tr><tr><td colspan="3"><strong>Bacterial and virus strains</strong></td></tr><tr><td>AAV8-hSyn-hM3Dq-mCherry</td><td>Addgene</td><td>Cat#50474</td></tr><tr><td>AAV8-hSyn-hM4Di-mCherry</td><td>Addgene</td><td>Cat#50475</td></tr><tr><td colspan="3"><strong>Chemicals, peptides, and recombinant proteins</strong></td></tr><tr><td>Glycine</td><td>Euromedex</td><td>Cat#26128 6405</td></tr><tr><td>L-Glutamate</td><td>Sigma-Aldrich</td><td>Cat#1446600 USP</td></tr><tr><td>Cyclothiazide</td><td>Biothekne</td><td>Cat#07130-50</td></tr><tr><td>Poly-L-Lysine</td><td>Sigma Aldrich</td><td>Cat#P2636</td></tr><tr><td>Neurobasal medium (Gibco)</td><td>Thermo Fisher Scientific</td><td>Cat#21103049</td></tr><tr><td>B27</td><td>Thermo Fisher Scientific</td><td>Cat#17504044</td></tr><tr><td>GlutaMAX</td><td>Thermo Fisher Scientific</td><td>Cat#35050061</td></tr><tr><td>Penicillin/streptomycin</td><td>Thermo Fisher Scientific</td><td>Cat#15140122</td></tr><tr><td>NaCl</td><td>Sigma Aldrich</td><td>Cat#560 14</td></tr><tr><td>KCl</td><td>Sigma Aldrich</td><td>Cat#P3911</td></tr><tr><td>CaCl2</td><td>Sigma Aldrich</td><td>Cat#C8106</td></tr><tr><td>HEPES</td><td>Sigma Aldrich</td><td>Cat#H3375</td></tr><tr><td>Glucose</td><td>VWR</td><td>Cat#24379294</td></tr><tr><td>Tetrodotoxin</td><td>Alomone Labs</td><td>Cat#T-550</td></tr><tr><td>Cs-gluconate (Cesium Hydroxide)</td><td>Sigma Aldrich</td><td>Cat#232041</td></tr><tr><td>CsCl</td><td>Sigma Aldrich</td><td>Cat#289329</td></tr><tr><td>TEACl</td><td>Sigma Aldrich</td><td>Cat#86614</td></tr><tr><td>EGTA</td><td>Sigma Aldrich</td><td>Cat#E3889</td></tr><tr><td>MgATP</td><td>Sigma Aldrich</td><td>Cat#A9187</td></tr><tr><td>NaGTP</td><td>Sigma Aldrich</td><td>Cat#G8877</td></tr><tr><td>NMDA</td><td>Tocris</td><td>Cat#0114</td></tr><tr><td>Sucrose</td><td>Sigma Aldrich</td><td>Cat#S0389</td></tr></tbody></table>

(Continued on next page)
Continued

<table><thead><tr><th>REAGENT or RESOURCE</th><th>SOURCE</th><th>IDENTIFIER</th></tr></thead><tbody><tr><td>NaH2PO4</td><td>Sigma Aldrich</td><td>Cat#S9638</td></tr><tr><td>MgSO4</td><td>Sigma Aldrich</td><td>Cat#230391</td></tr><tr><td>NaHCO3</td><td>Sigma Aldrich</td><td>Cat#S6014</td></tr><tr><td>Gluconic acid (potassium salt: K-gluconate)</td><td>Sigma Aldrich</td><td>Cat#G1951</td></tr><tr><td>MgCl2</td><td>Sigma Aldrich</td><td>Cat#M2670</td></tr><tr><td>Picrotoxin</td><td>Sigma Aldrich</td><td>Cat#P1675</td></tr><tr><td>DNQX</td><td>Tocris</td><td>Cat#2312</td></tr><tr><td>D-APV</td><td>Tocris</td><td>Cat#0106</td></tr><tr><td>NBQX</td><td>Tocris</td><td>Cat#0373</td></tr><tr><td>Cesium-methanesulfonate (Methanesulfonate)</td><td>Sigma Aldrich</td><td>Cat#64285</td></tr><tr><td>Na2ATP</td><td>Sigma Aldrich</td><td>Cat#A26209</td></tr><tr><td>cAMP</td><td>Sigma Aldrich</td><td>Cat#A9501</td></tr><tr><td>MK801</td><td>Sigma Aldrich</td><td>Cat#M107</td></tr><tr><td>L689,560</td><td>Tocris</td><td>Cat#0742</td></tr><tr><td>Alexa Fluor 594 hydrazide</td><td>Invitrogen</td><td>Cat#A10438</td></tr><tr><td>Fluo-4 pentapotassium salt</td><td>Invitrogen</td><td>Cat#F14200</td></tr><tr><td>Fluo-5F pentapotassium salt</td><td>Invitrogen</td><td>Cat#F14221</td></tr><tr><td>MNI-caged-L-glutamate</td><td>Tocris</td><td>Cat#1490</td></tr><tr><td>Trolox</td><td>Sigma</td><td>Cat#238813</td></tr><tr><td>BrainPhys medium</td><td>STEMCELL Technologies</td><td>Cat#O5790</td></tr><tr><td>Ketamine</td><td>Virbac</td><td>Ketamine 1000</td></tr><tr><td>Xylazine</td><td>Virbac</td><td>Paxman</td></tr><tr><td>LY2811376</td><td>Clinisciences</td><td>Cat#HY-10472</td></tr><tr><td>40% PEG300</td><td>Fluka</td><td>Cat#81160</td></tr><tr><td>5% Tween-80</td><td>Sigma-Aldrich</td><td>Cat#P1754</td></tr><tr><td>CNO</td><td>Enzo</td><td>Cat#BML-NS105-0025</td></tr><tr><td>Acepromazine</td><td>N/A</td><td>Calmivet</td></tr><tr><td>Hyaluronidase</td><td>Sigma-Aldrich</td><td>Cat#H4272</td></tr><tr><td>Diethylamine</td><td>Sigma-Aldrich</td><td>Cat#471216-5mL</td></tr><tr><td>Tris-HCl</td><td>VWR</td><td>Cat#0234-500G</td></tr><tr><td>Na2EDTA</td><td>Sigma-Aldrich</td><td>Cat#E6758-100g</td></tr><tr><td>NP-40 nonidet</td><td>Roche</td><td>Cat#1175459900</td></tr><tr><td>Sodium deoxycholate</td><td>Sigma-Aldrich</td><td>Cat#D5670-25g</td></tr><tr><td>Triton X-100</td><td>Sigma-Aldrich</td><td>Cat#X100</td></tr><tr><td>Protease inhibitors</td><td>Sigma-Aldrich</td><td>Cat#P8340</td></tr><tr><td>Bradford</td><td>Biorad</td><td>Cat#5000205</td></tr><tr><td>Tris-Tricine gels</td><td>Invitrogen</td><td>Cat#EC66252BOX</td></tr><tr><td>I-Block</td><td>Thermo Fisher Scientific</td><td>Cat#T2015</td></tr><tr><td>Tween 20 merck</td><td>Sigma-Aldrich</td><td>Cat#102423676</td></tr><tr><td>Ponceau</td><td>Sigma-Aldrich</td><td>Cat#P71-70</td></tr><tr><td>Chemiluminescence detection reagent ECL</td><td>Perkin Elmer</td><td>Cat#NEL112001EA</td></tr><tr><td>TCN201</td><td>Tocris</td><td>Cat#4154</td></tr><tr><td>DMSO</td><td>Sigma-Aldrich</td><td>Cat#D8418</td></tr><tr><td>Ifenprodil</td><td>Sigma Aldrich</td><td>Cat#I2892</td></tr><tr><td>D-Serine</td><td>Sigma Aldrich</td><td>Cat#S4250</td></tr><tr><td colspan="3">Critical commercial assays</td></tr><tr><td>Human/Rat (Mouse) β-Amyloid (40)</td><td>Wako</td><td>Cat#294-62501</td></tr><tr><td>ELISA Kit</td><td></td><td></td></tr></tbody></table>

(Continued on next page)
Continued

<table><thead><tr><th>REAGENT or RESOURCE</th><th>SOURCE</th><th>IDENTIFIER</th></tr></thead><tbody><tr><td>Golgi-Cox FD rapid GolgiStain kit, FD Neurotechnologies, USA</td><td>FD Neurotechnologies</td><td>Golgi stain</td></tr><tr><td colspan="3">Experimental models: Organisms/strains</td></tr><tr><td>APPdelETA mice</td><td>DZNE Munich -B6J- App&lt;em1Bwef&gt; (+em3)</td><td>private MGI: 7281307 (under registration at Jackson Laboratories)</td></tr><tr><td>AETA-m mice</td><td>LMU Munich -Tg(Thy1-APP)A2haa</td><td>private MGI: 7281720 (under registration at Jackson Laboratories)</td></tr><tr><td>C57BL/6</td><td>Charles River</td><td>RRID: IMSR_JAX:000664</td></tr></tbody></table>

Oligonucleotides

<table><tr><td>APPintron11-12for</td><td>Eurogentec</td><td>AAGCTCTGACTTTCCTTAAGGTGC</td></tr><tr><td>APPintron12-13rev</td><td>Eurogentec</td><td>TAGGAGTGGTATCCCTGCGGGT</td></tr><tr><td>AETA</td><td>Peptide Specialty Laboratories (PSL GmbH), Munich, Germany</td><td>MISEPRISYGNDALMPSLTETKTTVE LLPVNGEFLDDLQPWHSFGADSPVANTE NEVEPVDARPAADRGLTTRPGSG LTNIKTEEISEVKMDAEEFRHDSGYEVHHQK</td></tr><tr><td>CtrlP</td><td>Peptide Specialty Laboratories (PSL GmbH), Munich, Germany</td><td>KQHHVEYGSDHRFEADMKVESIEETKINTL GSGPRTTLGRDAAPRADVPEVENETNA PVSDAGFSHWPQLDLSFEGNVPLLE VTTKTETLSPMLADNGYSIRPESIM</td></tr><tr><td>crRNA APP-Ex12dn</td><td>IDT</td><td>CGCTCTCATGCCTTCGCTGA</td></tr><tr><td>crRNA APP-Ex12up</td><td>IDT</td><td>AACTACTCCGACGATGTCT</td></tr></table>

Recombinant DNA

<table><tr><td>Human GluN1-1a</td><td>Dr Laube, TU Darmstadt, Germany</td><td>Platzer et al.<sup>48</sup></td></tr><tr><td>Human GluN2A</td><td>Dr Laube, TU Darmstadt, Germany</td><td>Platzer et al.<sup>48</sup></td></tr><tr><td>Human GluN2B</td><td>Dr Laube, TU Darmstadt, Germany</td><td>Platzer et al.<sup>48</sup></td></tr><tr><td>Rat GluN1-1a</td><td>Dr Paoletti, IBENS, France</td><td>Mony et al.<sup>49</sup></td></tr><tr><td>Rat GluN2A</td><td>Dr Paoletti, IBENS, France</td><td>Mony et al.<sup>49</sup></td></tr><tr><td>Rat GluN2B</td><td>Dr Paoletti, IBENS, France</td><td>Mony et al.<sup>49</sup></td></tr><tr><td>Rat GluA1</td><td>Dr Kristensen, University of Copenhagen, Denmark</td><td>Stenum-Berg et al.<sup>50</sup></td></tr><tr><td>Rat GluA2<sub>R</sub></td><td>Dr Kristensen, University of Copenhagen, Denmark</td><td>Stenum-Berg et al.<sup>50</sup></td></tr><tr><td>Rat GluA3</td><td>Dr Kristensen, University of Copenhagen, Denmark</td><td>Stenum-Berg et al.<sup>50</sup></td></tr><tr><td>GluN1-GFP</td><td>Dr De Koninck, CERVO Brain Research Center, Canada</td><td>Doré et al.<sup>51</sup></td></tr><tr><td>GluN1-mCherry</td><td>Dr De Koninck, CERVO Brain Research Center, Canada</td><td>Doré et al.<sup>51</sup></td></tr><tr><td>Flag-GluN2B</td><td>Dr Groc, IINS, France</td><td>Ferreira et al.<sup>19</sup></td></tr></table>

Software and algorithms

<table><tr><td>GraphPad Prism 9</td><td>Dotmatics</td><td>Academic module</td></tr><tr><td>Packwin; freezing module</td><td>Panlab</td><td>N/A</td></tr><tr><td>Clampfit 10.6</td><td>Axon Instruments</td><td>N/A</td></tr><tr><td>Clampex</td><td>Molecular Devices</td><td>N/A</td></tr><tr><td>Multiclamp</td><td>Molecular Devices</td><td>N/A</td></tr></table>

RESOURCE AVAILABILITY

Lead contact

Further information and requests for resources and reagents should be directed to and will be fulfilled by the lead contact, Hélène Marie (marie@ipmc.cnrs.fr).
<figure><img src="image_58.png" /></figure>

## Materials availability

The new APPdelETA and AETA-m mouse lines have been preregistered to Jackson Laboratories for unrestricted availability upon publication.

## Data and code availability

* All data reported in this paper will be shared by the [lead contact](#) upon request.

* This paper does not report original code.

* Any additional information required to reanalyze the data reported in this paper is available from the [lead contact](#) upon request.

## EXPERIMENTAL MODEL AND STUDY PARTICIPANT DETAILS

### Oocytes

Oocyte harvest was performed in accordance with the European directives 2010/63/EU on the Protection of Animals used for Scientific Purposes in the framework of project authorization APAFIS#14012-2018030214144339 as delivered by the competent French authorities and by the German local animal care and use committee (II25.3-19c20/15, RP Darmstadt, Germany).

### Mice and rats

All experiments and protocols on mice and rats were performed in accordance with the European Communities' Council Directive 2010/63/EU. Protocols used in this study were approved by the committee for the Care and Use of Laboratory Animals and governments of the relevant countries as detailed in the relevant sections in the methods details below. The different mouse and rat species used are reported in the methods details where relevant. In all animal houses the rodents had *ad libitum* access to water (tap water) and standard chow and were maintained under constant environmental conditions (12:12 h light/dark cycle, 23 ± 2°C and humidity of 55%). They were housed in groups of 5-6 animals by sex in standard cages (mouse: 542 cm²; rat: 2065 cm²) in animal houses under specific pathogen free (SPF) status. For *ex vivo* experiments, sex used is reported in each section whenever relevant. None of the mice used in the reported experiments were used in other previous non-reported *in vivo* procedures. For the behavioral experiment (Figures 4I-4K), only adult males (2-3 months old) were considered and the result cannot be generalized to both sexes for this experiment. AETA-m mice were backcrossed regularly (at least 8 times prior to experiments presented in this study) on C57BL6J background (Charles River) to obtain WT and transgenic mutants. APPdelETA homozygous and WT littermates were generated by crossing heterozygous male and female mice from different litters with intermittent backcrossing on C57BL6J background (Charles River) to obtain new heterozygous mice for the crossings. Littermates of the same sex were randomly assigned to experimental groups. Generation of double mutants (APPdelETA/AETA-m mice) was obtained in two steps. We first crossed homozygous male homozygous APPdelETA with female transgenic AETA-m mice. Upon obtaining heterozygous/transgenic females from this crossing, these females were bred with heterozygous APPdelETA males to obtain WT, homozygous and homozygous/transgenic genotypes from same litters for the behavioral experiment.

### Primary neuronal cultures

E15 RjOr1:Swiss and E18 Sprague-Dawley rat embryos were used for neuronal cultures. Sex of embryos was not considered and both male and females embryos were used. Details of culture conditions are provided in relevant sections below.

## METHOD DETAILS

### Peptides

Synthetic AETA and control peptide (CtrlP, representing the reverse sequence of AETA) were obtained from Peptide Specialty Laboratories (PSL GmbH; Heidelberg, Germany) and consisted of the following sequences:

AETA sequence (108 amino acids):

MISEPRISYGNDALMPSLTETKTTVELLPVNGEFSLDDLQPWHSFGADSVPANTENEVEPVDARPAADRGLTTRPGSGLTNIKTEEISEVKM
DAEFRHDSGYEVHHQK

CtrlP sequence (108 amino acids):

KQHHVEYGSDHRFEADMKVESIEETKINTLGSGPRTTLGRDAAPRADVPEVENETNAPVSDAGFSHWPQLDDLSFEGNVPLLEVTTKTETL
SPMLADNGYSIRPESIM

The peptides were dissolved in dimethyl sulfoxide (DMSO) at 100 µM and placed at -80°C for long term storage. For recordings in *Xenopus* oocytes, these aliquots were further dissolved to 100 nM in oocyte recording solution on day of experiment. For all other experiments, the peptides were further diluted to 10 µM in artificial cerebrospinal fluid (aCSF) and placed at -80°C for long term storage. On day of experiment, aliquots were diluted to 10 nM in aCSF.

### Electrophysiology in *Xenopus* oocytes

Human GluN1-1a (named GluN1 herein), GluN2A and GluN2B subunits of NMDAR (Laube lab)⁴⁸ or pcDNA3-based plasmids for rodent GluN1-1a, GluN2A and GluN2B subunits of NMDAR (Marie lab)⁴⁹ or pXOOF-based plasmids for rat GluA1, GluA2R and GluA3⁵⁰
were used for expression in *Xenopus* oocytes. All constructs were linearized with appropriate enzyme and transcribed into cRNA (mCAP mRNA Capping Kit, Ambion, or AmpliCap-Max T7 High Yield Message Maker KitA, TEBU). Oocytes from female *Xenopus laevis* were prepared as described previously.⁵²,⁵³ *Xenopus laevis* oocytes were injected with 30 ng cRNAs in a volume of 50 nL of a 1:1 ratio of GluN1 per GluN2A or GluN2B, or 30 ng a 2:1 ratio of GluA2R per GluA1 or GluA3. 1–2 days after injection, two-electrode voltage clamp (TEVC) recording was performed at a holding potential of –60 mV or –70 mV at room temperature. The two microelectrodes were filled with 3M KCl and had a resistance of 0.5–1 MOhm. Currents were acquired at 200 Hz with a Geneclamp 500B amplifier, a Digidata 1322A digitizer and Clampex 9.2 software (Molecular Devices, USA). Recombinant NMDAR-mediated currents were induced in Mg²⁺-free frog Ringer solution by co-application of L-glutamate (100 µM) and glycine (1 µM), unless otherwise indicated in figure caption. Recombinant AMPAR-mediated currents were induced by application of L-glutamate (300 µM) and cyclothiazide (CTZ, 100 µM). For determining dose-response relations, superfusion was switched to desired concentrations of peptides/drugs in Ringer solution. Glutamate/glycine, dissolved in bath solution, was applied either alone for 10 s (Laube lab) or 30 s (Marie lab) or after 15 s (Laube lab) or 1 min (Marie lab) pre-application of the appropriate peptide also dissolved in bath solution. Currents were measured with Clampfit 9.2 software (Molecular Devices, USA) and results were analyzed using GraphPad Prism version 9 (GraphPad, USA). 100% current value represented current obtained upon glutamate application prior to peptide application. Current value obtained after peptide application was normalized to this control current value for each oocyte recording. The dose-response curves of glutamate-induced peak currents were normalized to the maximal current value (Imax) obtained without AETA (Figure S1A) or with 100 µM glycine (Figure 1b). IC50 (half-maximal inhibitory concentration) and EC50 (half-maximal effective concentration) were calculated from fitting dose-response curves to a sigmoidal dose-response curve using the Hill equation. All experiments were from at least three batches of oocytes.

## Electrophysiology and p38 activity in neuronal cultures

Primary hippocampal neurons were isolated from 15-day-old mouse embryo brain (E15) from RjOrl:Swiss pregnant female. Cells were mechanically dissociated and plated on 12-mm glass coverslips pre-treated with poly-L-Lysine (0.1 mg mL⁻¹; Sigma) and maintained at 37°C, 5% CO₂ in Neurobasal medium (Gibco) supplemented with 2% B27 (Gibco), 2 mM GlutaMAX (Gibco) and 1% penicillin/streptomycin (10 000UI, 10 000 µg/mL; Gibco) for 10–14 days prior to use.

To test p38 activity, we incubated neurons (200,000 cells/sample) with AETA (10 nM) for 20 min. We collected control neurons and AETA-exposed neurons, lysed them in RIPA (see below) and processed the lysate for immunoblotting (see below).

Neurons were recorded in whole-cell patch-clamp configuration at –65 mV at room temperature (RT) in extracellular solution containing in mM: NaCl 150, KCl 2.5, CaCl₂ 2.0, HEPES 10, glucose 10 and glycine 0.01, pH 7.4 supplemented with Tetrodotoxin (100 nM) and NBQX (1µM). Recording pipettes (5–6 MΩ) were filled with Cs-gluconate solution (in mM): 117.5 Cs-gluconate, 15.5 CsCl, 10 TEACl, 8 NaCl, 10 HEPES, 0.25 EGTA, 4 MgATP and 0.3 NaGTP (pH 7.3; osmolarity 290–300 mOsm).

NMDA currents were evoked 5 times by 10 s application of 30 µM NMDA separated by 40 s washout (extracellular solution). After 10 min of peptide incubation (10 nM CtrlP or AETA in bath solution), neurons were stimulated again with another 5 pulses of NMDA application. Local application of NMDA was delivered by an automated perfusion system (ValveLink 8.2, Science Products) with a 350 µm tip that was set close to the cell of interest. For analysis, mean peak amplitude measured for the 5 stimulations before incubation was set as 100% current and a ratio amplitude was established on this mean for each evoked current. Two factor analysis of variance (two-way ANOVA) followed by Sidak's post-hoc was performed to analyze significance among the conditions.

## Electrophysiology in hippocampal slices

For *ex vivo* electrophysiology recordings, 3–6 weeks old male RjOrl:SWISS mice (Janvier, France), 2–3 months male APPdelETA and WT littermates, or 3–4 weeks old male Sprague-Dawley rats (Janvier, France) were used. Mice were culled by cervical dislocation and hippocampi were dissected and incubated for 5 min in ice-cold oxygenated (95% O₂/5% CO₂) cutting solution (in mM): 234 sucrose, 2.5 KCl, 1.25 NaH₂PO₄, 10 MgSO₄, 0.5 CaCl₂, 26 NaHCO₃, 11 glucose (pH 7.4). Hippocampal slices (250 µm for patch-clamp recordings, 350 µm for field recordings) were cut on a vibratome (Microm HM600V, Thermo Scientific, France). For recovery, slices were then incubated in standard aCSF (in mM): 119 NaCl, 2.5 KCl, 1.25 NaH₂PO₄, 26 NaHCO₃, 1.3 MgSO₄, 2.5 CaCl₂ and 11 D-glucose, oxygenated with 95% O₂ and 5% CO₂, pH 7.4 for 1 h at 37 ± 1°C and then stored at RT until used for recordings. Recordings were done in this standard aCSF (unless otherwise stated) in a recording chamber on an upright microscope with IR-DIC illumination (SliceScope, Scientifica Ltd, UK) using a Multiclamp 700B amplifier (Molecular Devices, San Jose, CA, USA), under the control of pClamp10 software (Molecular Devices, San Jose, CA, USA). Data analysis was executed using Clampfit 10 software (Molecular Devices, San Jose, CA, USA). The Schaffer collateral pathway was stimulated at 0.1 Hz for patch clamp experiments and 0.25 Hz for field experiments (unless otherwise stated) using electrodes (glass pipettes filled with aCSF) placed in the stratum radiatum.

Patch-clamp experiments were performed at 31 ± 1°C. For whole-cell voltage-clamp, recording pipettes (5–6 MΩ) were filled with a solution containing the following: 117.5 mM Cs-gluconate, 15.5 mM CsCl, 10 mM TEACl, 8 mM NaCl, 10 HEPES, 0.25 mM EGTA, 4 mM MgATP and 0.3 NaGTP (pH 7.3; osmolarity 290–300 mOsm). For whole cell current-clamp, the recording pipette solution contained (in mM): 135 gluconic acid (potassium salt: K-gluconate), 5 NaCl, 2 MgCl₂, 10 HEPES, 0.5 EGTA, 2 MgATP and 0.4 NaGTP (pH 7.25; osmolarity 280–290 mOsm). After a tight seal (>1GΩ) on the cell body of the selected neuron was obtained, whole-cell patch clamp configuration was established, and cells were left to stabilize for 2–3 min before recordings began. Holding current and series
resistance were continuously monitored throughout the experiment, and if either of these two parameters varied by more than 20%, the cell was discarded.

NMDAR EPSCs were pharmacologically isolated by adding 50 µM picrotoxin (Sigma-Aldrich, dissolved in DMSO) to block GABAergic transmission and DNQX (10 µM; Sigma-Aldrich, dissolved in DMSO) to block AMPA receptors and recorded at +40mV. The time courses were obtained by normalizing each recording to the average value of all points constituting the first 10 min stable baseline. Graphs of current alteration was measured during the last 10 min of recording and calculated as % change from baseline average (first 10 min).

Spontaneous EPSCs were recorded at -65 mV in presence of picrotoxin (50 µM; Sigma-Aldrich), D-APV (50 µM; Tocris) for AMPAR EPSCs or NBQX (10 µM, Tocris) and Mg²⁺-free aCSF for NMDA EPSCs using the following internal solution (mM): Cesium-methanesulfonate (143), NaCl (5), MgCl₂ (1), EGTA (1), CaCl₂ (0.3), HEPES (10), Na₂ATP (2), NaGTP (0.3) and cAMP (0.2) (pH 7.3 and 290-295 mOsm). To test effect of AETA or CtrlP (10 nM), sEPSC were first recorded in gap-free mode for 5 min (baseline condition), then the peptide was applied for 10 min, and sEPSC were then recorded for another 5 min (peptide condition). For recordings in WT and APPdelETA mice, 5 min were recorded for individual neurons of each genotype. In APPdelETA neurons, after these 5 min of recordings, AETA (10 nM) was bath applied for 10 min, after which another 5 min were recorded. Analysis of sEPSCs was performed on the first and last 5 min to compare frequencies and amplitudes use Clampfit 10. sEPSCs were detected manually by following criteria of peaks with a threshold 2xSD of baseline noise level and a faster rise time than decay time. Analysis was performed blind to experimental condition.

Minimal stimulation experiments were performed as described previously.²⁶ Briefly, EPSCs were recorded in CA1 neurons while stimulating the CA3 Schaffer collaterals in presence of picrotoxin (50 µM). After evoking a small (20-40 pA) EPSC at -65 mV at 0.1 Hz stimulation frequency, stimulation strength was reduced in small increments to the point that failures versus responses could be clearly distinguished visually. Stimulation intensity was then kept constant throughout experiment. 50 sweeps were recorded at -65 mV and 50 sweeps were recorded at +40 mV. Failures and success rates were estimated visually for each voltage by experimenter blind to genotype. Percent silent synapses were calculated as follows: $1 - \ln(F_{-65mV})/\ln(F_{+40mV})$ where F represents failure rate calculated for each voltage.⁵⁴

Field excitatory postsynaptic potentials (fEPSPs) were recorded in the stratum radiatum of the CA1 region (using a glass electrode filled with 1 M NaCl and 10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES), pH 7.4) and the stimuli were delivered to the Schaffer collateral pathway by a monopolar glass electrode filled with aCSF. fEPSP response was set to approximately 30% of the maximal fEPSP response i.e., approx. 0.2-0.3 mV, with stimulation intensity 10 µA ± 5 µA delivered via a stimulation box (ISO-Flex, A.M.P.I. Inc., Israel). Electrodes were placed superficially to maximize exposure to peptides. A stable baseline of 20 min bath application of aCSF under control conditions or with peptide was first obtained before induction for long-term plasticity recordings. The peptide was then also recirculated throughout the 1-h recording after induction. LTP was induced by high frequency stimulation: 2 × 100 Hz/1 s at 20 s interval. Sub-threshold LTD was induced by low frequency stimulation: 300 pulses at 1 Hz. On RjOrl:SWISS mice (3-6 weeks) slices, LTD was induced with low frequency stimulation: 900 pulses of 1 Hz in standard aCSF (see above) in presence of 50 µM picrotoxin (Sigma-Aldrich, dissolved in DMSO). On WT and APPdelETA mice (2-3 months), LTD was induced in a modified aCSF containing in mM: (in mM): 119 NaCl, 2.5 KCl, 1.25 NaH₂PO₄, 26 NaHCO₃, 2 MgSO₄, 4 CaCl₂ and 11 D-glucose, oxygenated with 95% O₂ and 5% CO₂, pH 7.4, without picrotoxin. MK801 (100 µM; Sigma-Aldrich, dissolved in DMSO) was used for some LTD experiments. For all LTD and LTP recordings, only the first third of the fEPSP slope was analyzed to avoid population spike contamination. The time courses were obtained by normalizing each experiment to the average value of all points constituting a 20 min stable baseline before induction. fEPSP magnitude was measured during the last 15 min of recording (45-60 min after induction) and calculated as % change fEPSP slope from baseline average. For analysis of LTD in APPdelETA mice in presence of L689,560, the drug was purchased from Tocris, prepared as a stock solution of 10 mM in DMSO and diluted to 10 µM (as recommended by⁵⁵ to avoid nonspecific effects) in aCSF on day of experiment. For PPRs, an EPSC at -65mV (naive slices from RjOrl:Swiss mice) or a fEPSP of ~50% of the maximum (WT and APPdelETA mice) was obtained and two stimuli were delivered at 100, 200, or 300 ms inter-stimulus interval as indicated in figure. PPR was calculated as fEPSP2slope/fEPSP1slope (10 sweeps average per ISI). Recordings of control and peptide conditions were interleaved within the same day. Recordings of WT and APPdelETA mice were interleaved between days.

## uEPSP and calcium transients in single spines

All procedures were carried out under local institutional guidelines, approved by the University of Bristol Animal Welfare and Ethical Review Board, and in accordance with the UK Animals (Scientific procedures) Act 1986. Transverse hippocampal slices from 4 to 5 week-old male C57/BL6J mice were used for simultaneous 2-photon imaging and current-clamp electrophysiology experiments. Dissection and slicing were performed in ice-cold sucrose based medium (in mM: 205 sucrose, 10 glucose, 26 NaHCO₃, 2.5 KCl, 1.25 NaH₂PO₄, 0.5 CaCl₂, 5 MgSO₄), 400 µm slices were then transferred to aCSF (in mM: 124 NaCl, 3 KCl, 26 NaHCO₃, 1.4 NaH₂PO₄, 10 Glucose, 1.3 MgSO₄, 2.5 CaCl₂, 50 µM picrotoxin) saturated with 95% O₂ and 5% CO₂. Patch pipettes had resistances of 4-6 MΩ and were filled with a potassium-based intracellular solution containing: 150 K⁺ methanesulfonate, 5 KCl, 10 HEPES, 3 MgATP, 0.4 Na₂GTP, with 20 µM Alexa Fluor 594 and 200 µM Fluo-4/5F. Spectra Physics Mai Tai pulsed lasers were tuned to 810 nm for imaging and 730 nm for uncaging. Image acquisition, uncaging and electrophysiological recordings were controlled using PrairieView (Bruker). Laser power at the slice surface was less than 10 mW. Pyramidal cells were visualized using Dodt gradient contrast and
a 60× water immersion objective. Once whole-cell configuration was established, 2-photon excitation was used to visualize dendrites and spines (Alexa Fluo 594). Line scan images were acquired at least 30 min after breaking into the cell to allow for dye diffusion and equilibration. When the aCSF was supplemented with 2.5 mM MNI-caged-L-glutamate and 250 µM Trolox, the perfusion system was switched to a closed system. A series of five brief (1–2 ms) uncaging laser pulses targeted to a single spine was used to evoke EPSPs recorded using a Multiclamp 700B. Laser power, pulse duration and location were finely calibrated to elicit calcium transients only in the target spine as tested by line scans across neighboring spines and dendrites.⁵⁶ Electrophysiological signals were digitized at 20 kHz and low pass filtered at 4kHz. Membrane potentials were not corrected for the junction potential. Line scan series over 400 ms were acquired every 2–3 min. Plot profiles from individual line scan images were collected over the spine using ImageJ and the mean value for each time point quantified. The profile obtained from Fluo-4/5F (green ΔG) was divided by that obtained from Alexa 594 (red R), so ΔF = ΔG/R. ΔF/F values presented were obtained by dividing each ΔF by the 30 ms baseline before uncaging.

## FLIM-FRET of GluN1-GFP/GluN1-mCherry FRET pair

Primary cultures of dissociated hippocampal neurons were prepared from E18 Sprague-Dawley rat embryos, as previously described.⁵⁷ Briefly, cells were plated at a density of 280 × 10³ cells per dish on poly-L-lysine-coated Ø 18 mm glass coverslips kept in Ø 60 mm Petri dishes filled with Neurobasal medium supplemented with B-27 Plus (#A3653401, ThermoFisher Scientific, Waltham, MA, USA), GlutaMAX (#35050061, ThermoFisher Scientific), and 1.5% heat-inactivated horse serum. Cells were kept at 37°C/ 5% CO₂ and were transferred to serum-free medium after 3 days *in vitro* (div), half of which was replaced by fresh medium at 6 div. In order to perform FLIM-FRET measurements, 8–10 div cells were transfected with GluN1-GFP, GluN1-mCherry (gifts from Paul De Koninck),²¹,⁵¹ and Flag-GluN2B (gift from R. Wenthold) at a ratio of 1:3:1 since we previously defined that this ratio provided optimal FRET signal.¹⁹ The various plasmids were expressed in neurons using the calcium-phosphate co-precipitation method.⁵⁸ mCherry-GluN2B was expressed instead of GluN1-mCherry in a subset of cells as a negative control. After transfection, cells were transferred to BrainPhys medium (STEMCELL Technologies, #05790) supplemented with B-27 Plus and maintained at 37°C/5% CO₂ until the day of experimentation (13–15 div). Experiments were carried out at 37°C using an incubator box with an air heater system (Life Imaging Services, Switzerland) installed on an inverted Leica DMI6000B spinning disk microscope (Leica Microsystems, Germany) using the LIFA frequency domain lifetime attachment (Lambert Instruments BV, The Netherlands) and the LI-FLIM software. Cells were imaged with an HCX PL Apo CS 63X NA 1.4 oil-immersion objective using an appropriate filter set. GFP fluorescence was excited using a sinusoidally modulated 3 W/478 nm LED at 36 MHz under wild-field illumination. Emission was collected using an intensified CCD LI2CAM camera (Lambert Instruments BV). Acquisitions were performed after a 10 min incubation with either synthetic AETA or CtrlP peptides (10 nM). The experimenter was blind to the conditions until final analysis. Lifetimes were calibrated using a reference solution of erythrosin B (1 mg/mL) with an average lifetime of 0.086 ns. The lifetime of the sample was determined from the fluorescence phase-shift between the sample and the reference from a set of 12 phase settings using the LI-FLIM software provided by the manufacturer. GFP fluorescence lifetimes were measured in dendritic spine clusters defined by the user based on confocal images of the fluorescence signals of the donor (GluN1-GFP) and acceptor (GluN1-mCherry) fluorophores, blind to the FLIM image. For each neuron, a total of 20–30 dendritic spines clusters were selected, from which lifetime values were extracted using the LI-FLIM software. FRET efficiencies were calculated—either from a random selection of cluster lifetime values or from mean lifetime values per neuron—for each experimental condition using the following equation:

$$FRET efficiency (\%) = \frac{(\tau_D - \tau_{DA}) * 100}{\tau_D}$$

where τD is the GFP fluorescence lifetime when the FRET donor (GluN1-GFP) is expressed alone, and τDA is the GFP fluorescence lifetime when the FRET donor is expressed together with the acceptor (GluN1-mCherry).⁵¹

## Two-photon imaging of spine structural plasticity

Acute hippocampal slices were prepared from P18-P21 GFP-M mice⁵⁹ of both sexes, as described.¹³ All experimental protocols were approved by the University of California Davis Institutional Animal Care and Use Committee. GFP-expressing CA1 pyramidal neurons were imaged using a custom two-photon microscope.⁶⁰ For each neuron, image stacks (512 × 512 pixels; 0.02 µm per pixel; 1-µm z-steps) were collected from a tertiary basal dendrite at 5 min intervals at 30°C in recirculating artificial cerebral spinal fluid (aCSF; in mM: 127 NaCl, 25 NaHCO₃, 1.2 NaH₂PO₄, 2.5 KCl, 25 D-glucose, aerated with 95%O₂/5%CO₂, ~310 mOsm, pH 7.2) with 2 mM Ca²⁺, 0.1 mM Mg²⁺, and 1 µM TTX. Slices were pre-incubated for 20 min with CtrlP or AETA. Images are maximum projections of image stacks after applying a median filter (2 × 2) to raw image data. Estimated spine volume was measured from background-subtracted green fluorescence using the integrated pixel intensity of a boxed region surrounding the spine head, as described.⁶⁰ High-frequency uncaging (HFU) consisted of 60 pulses (720 nm; 2 ms duration, ~12 mW at the sample) at 2 Hz delivered in ACSF containing (in mM): 2 Ca²⁺, 0.1 Mg²⁺, 0.001 TTX, and 2.5 MNI-glutamate. The beam was parked at a point 0.5–1 µm from the spine at the position farthest from the dendrite. Cells for each condition were obtained from 6 independent hippocampal acute slices preparations of both sexes. Data collection and analysis was done blind to the experimental condition. All statistics were calculated across cells. two-way ANOVA with Bonferroni's multiple comparisons was used.
## BACE1 inhibition in vivo

To increase endogenous AETA levels, 6 weeks old male RjOrl:SWISS mice (Janvier, France) were administered the BACE1 inhibitor LY2811376 (100 mg/kg) by oral gavage.²⁵ LY2811376 was obtained from Medchem Express (Sweden) and prepared in 10% DMSO, 40% PEG300, 5% Tween-80, and 45% saline. After slicing and recovery (1h at 37°C), slices were incubated in LY2811376 (5 µM) throughout experiment. NMDAR sEPSC and LTD recordings were performed as described above.

## In vivo neuronal activation or inhibition by DREADD

The procedure was performed essentially as described previously⁶¹ in accordance with the recommendations of the European Commission (2010/63/EU) for care and use of laboratory animals and approved by the French National Ethical Committee (#16459-2018061116303066). AAV8-hSyn-hM3Dq-mCherry (#50474) and AAV8-hSyn-hM4Di-mCherry (#50475) were purchased from Addgene and injected in pre-frontal cortex by stereotaxic surgery in 8 week-old male C57/BL6J mice (Janvier Labs, France). Stereotaxic injections were performed using a stereotaxic frame (Kopf Instruments). General anesthesia was achieved using a mix of ketamine (150 mg/kg) and xylazine (10 mg/kg). Viruses were injected bilaterally at a rate of 100 nL/min for a final volume of 500 nL per site. Stereotaxic coordinates were (in mm): antero-posterior (AP): +1,7; Mediolateral (ML): +/- 1,5; dorsoventral (DV): -1,5; based on the Paxinos atlas of the adult mouse brain. Coordinates were taken from bregma for AP and ML coordinates, and from skull at the site of injection for DV. Mice were given a 3-week-recovery period to allow sufficient viral expression. CNO (1 mg/kg, Sigma-Aldrich, France) or saline (0.9% NaCl, 10 mL/kg) was administered by intraperitoneal injection 30 min before microdissection of transduced tissue. Brains were then removed, mCherry-positive tissues were visualized with a DFP-1 duel fluorescence protein flashlight (NIGHTSEA, USA) and dissected out to be snap-frozen in liquid nitrogen for immunoblotting (see immunoblotting section). A subgroup of slices with transduced virus were mounted and visualized under a Vectra 3 (PerkinElmer) microscope to assess virus spread.

## Generation of APPdelETA mice

APPdelETA mice were generated by CRISPR/Cas9-assisted gene editing in zygotes as described previously.⁶² Briefly, pronuclear stage zygotes were obtained by mating C57BL/6J males with superovulated C57BL/6J females (Charles River, Germany). Embryos were then microinjected into the male pronucleus with an injection mix containing APP-specific CRISPR/Cas9 ribonucleoprotein (RNP) complexes. RNPs consisted of 50 ng/µL Cas9 protein (IDT, Coralville, USA), 1 µM crRNA APP-Ex12up (protospacer GAACTACTCCGACGATGTCT; IDT), 1 µM crRNA APP-Ex12dn (protospacer CGCTCTCATGCCTTCGCTGA; IDT), and 1 µM tracrRNA (IDT, Coralville, USA). After microinjection, zygotes were cultured in KSOM medium until transferred into pseudo-pregnant CD-1 foster animals. A mutant founder carrying a 123 bp deletion was crossed to a C57BL/6J animal to establish the stable APPdelETA line. All mice were handled according to institutional guidelines approved by the animal welfare and use committee of the government of Upper Bavaria and housed in standard cages in a specific pathogen-free facility on a 12-h light/dark cycle with *ad libitum* access to food and water. To identify putative off target sites of the APP-specific crRNAs, the CRISPOR online tool⁶³ was used. TOP12 predicted sites (combined CFD and MIT score) were chosen for off-target analysis. For analysis, genomic DNA of wildtype and heterozygous mutant F1 APPdelETA mice was isolated and predicted loci were PCR amplified with primers flanking the putative cut sites and subsequently Sanger sequenced using the PCR primers. No off-target events were detected. Genotyping of the line is performed with this set of primers in a standard PCR reaction from tail biopsies: APPintron11-12for (AAGCTCTGACTTTTCCTTAAGGTGC) and APPintron12-13rev (TAGGAGTGGTATCCCTGCGGGT). The PCR products are cleaved with the restriction enzyme Bcl1 (New England Biolabs, Germany) with no cleavage for the KI/KI (510 bp) and two bands for the WT allele resulting products of 318 bp and 192 bp. To quantify AETA and CTF-1 levels, hippocampi of APPdelETA mice and WT littermates were dissected out, snap frozen in liquid nitrogen and kept in -80°C freezer until processed for immunoblotting.

## Generation of AETA-m mouse line

AETA-overexpressing mice (Thy-1.2-Aη-α; AETA-m) were generated using C57Bl6/NCrI Donor mice mated by superovulation (46 h between pregnant mare serum gonadotropin (PMSG) and human chorionic gonadotropin (HCG)) at the midpoint of the dark period (12 h/12 h, 6 a.m. to 6 p.m. light circle). After positive plug detection in the morning, the cumulus complexes were isolated and zygotes removed with a treatment of hyaluronidase (Sigma-Aldrich; final concentration of 0.1% (801 U/ml)). Transgenic mice were backcrossed on C57Bl6/J. The linearized Thy1.2 cassette coding for a secreted version of human AETA DNA plasmid was injected into the male pronucleus of fertilized zygotes by using a motor driven manipulator-based microinjection stage. About 2 h after injections, the surviving embryos were transferred into Crl:CD1(ICR) pseudopregnant recipient female mice (ca. 20 embryos per recipient). The recipient mice were mated with sterile males (vasectomized) Crl:CD1(ICR). By detection of a copulation plug in the morning of the transfer day, pseudopregnant mice can be used for the unilateral surgical embryo transfer procedure into the oviduct. Anesthesia was induced by intraperitoneal (i.p.) injection with a mix of Ketamin/Xylazin and Acepromazine. Mice were generated under the license 24-9168.11-9/2012-5. All animal experiments were performed in accordance with the European Communities Council Directive (86/609/EEC), and were approved by the local ethics committee (Government of Saxony, Germany). Immunoblotting of rodent AETA levels and human AETA levels was performed as described below. Levels of Aβ₁-40 was quantified using the Wako Human/Rat (Mouse) β-Amyloid (40) ELISA Kit (Catalog Number: 294-62501) as per manufacturer's instructions.
## Immunoblotting

The brain homogenates for the analysis of APP processing products (APP-FL, CTF-η, AETA) were essentially prepared as described previously.¹⁶ In brief, DEA lysates (0.2% Diethylamine in 50 mM NaCl, pH 10) and RIPA lysates (20 mM Tris-HCl pH 7.4, 150 mM NaCl, 1 mM Na₂EDTA, 1% NP-40, 0.5% sodium deoxycholate, 0.05% Triton X-100) with protease inhibitors (Sigma-Aldrich, P8340) were prepared from brain samples using the Precellys system (Bertin) for homogenisation followed by ultracentrifugation. For the DEA and RIPA samples we used the Bradford (Biorad) to measure the protein concentration, which was adjusted equally to all samples before Western blot analysis. For detection by Western blotting, proteins were separated on 8–12% Tris-Glycine gels or alternatively on Tris-Tricine (10–20%, Thermo Fisher Scientific) gels, transferred to nitrocellulose membranes (0.1 µm, GE Healthcare) which were boiled for 5 min in PBS and subsequently incubated with the blocking solution containing 0.2% I-Block (Thermo Fisher Scientific) and 0.1% Tween 20 (Merck) in PBS for 1 h, followed by overnight incubation with 2 µg/mL antibody in the blocking solution. Antibody detection was performed using the corresponding anti-rat/mouse/rabbit-IgG-HRP conjugated secondary antibody (Thermo Fisher Scientific) and chemiluminescence detection reagent ECL (Thermo Fisher Scientific). Antibodies used for immunoblotting are M3.2 (Biolegend, #805701; mouse IgG) for detection of mouse AETA, antibodies 2D8 and 2E9 for human AETA,¹⁶ 22C11 (Merck, #MAB348; mouse IgG) for APP-FL, Y188 (abcam, #32136; rabbit IgG) for CTF-η, a polyclonal serum for mCherry (Takara, # 632496, rabbit IgG), Synaptophysin (Sigma-Aldrich, S5786, mouse IgG) and Synaptobrevin-2 (SYSY, #104211, mouse IgG), p-38 (#8690, Cell Signaling, rabbit IgG), p-p38 (#9216, Cell Signaling, mouse IgG). For the loading control, when necessary, we used an antibody specific to β-actin (Sigma-Aldrich, #AC-74). AETA was quantified in DEA fraction and CTF-η was quantified in RIPA fraction. For analysis of APPdelETA and WT mouse tissue, peptide levels were normalized to β-actin or ponceau (as stated in results description), and normalized to WT average for each immunoblot for statistical analysis (Mann-Whitney test) using GraphPrism 8. For DREADD experiment, AETA levels and mCherry levels were normalized to β-actin. AETA levels were further normalized to mCherry levels. Outliers in values obtained for AETA/mCherry ratio (1 hM3D-SAL and 1 hM4D-CNO) were not considered for statistical analysis (see Figure S4). Ratios were further normalized to average Saline of each experiment to pool all experiments together and perform statistical analysis (Mann-Whitney test).

## Golgi-Cox staining for spine density analysis

Brains of WT and APPdelETA mice were collected and processed for Golgi-Cox impregnation. Brains were removed and impregnated in a Golgi-Cox solution (1% potassium dichromate, 1% mercuric chloride, 0.8% potassium chromate) for 3 weeks at room temperature according to manufacturer instruction (FD rapid GolgiStain kit, FD Neurotechnologies, USA). Brains were sectioned coronally (80 µm) using a vibratome and stained and mounted according to protocol. Images were acquired under white light on a DMD108 Leica microscope (×60 magnification). Spine density (number of spines per 1 µm length) in selected segments (minimum of 30 per mouse) of secondary dendrites of CA1 pyramidal neurons in stratum radiatum was estimated using ImageJ and manual count.

## Contextual fear conditioning

Behavioral analysis was performed in accordance with the European directives 2010/63/EU on the Protection of Animals used for Scientific Purposes in the framework of project authorizations APAFIS#6856-2016091610462338 and APAFIS#37493-2022050311352580 as delivered by the competent French authorities. All animals were daily handled for 2 min during one week. To measure electric shock-induced freezing, each mouse was placed individually in a soundproof test chamber containing a floor made of a grid with 27 stainless-steel rods (diameter 4 mm) spaced 1 cm apart and connected to a generator to allow shock delivery (Shocker LE 100-26 Panlab Harvard Apparatus Bioseb). Mice were left to freely explore the apparatus for 3 min and then received three consecutive electrical foot shocks (intensity: 0.7 mA, duration: 2 s) with a 1 min interval between each shock. Mice remained in the test chamber 1 min after the last foot shock. Activity levels of mice were recorded through a high-precision sensor plate placed beneath the floor grid (Load cell coupler LE 111 Panlab Harvard Apparatus Bioseb) to assess the variations of weight induced by the movements of the mice. To test for aversive memory formation, mice were re-exposed 24 h after to the same context for 6 min without any shock. Freezing was defined as total lack of movement aside from breathing for a cumulative duration of at least 2s. The freezing behavior was scored using high-precision sensor plate.

## Statistical analysis

Results are shown as mean ± s.e.m. Numbers and their correspondence are given in each figure. Statistical analysis was performed with GraphPrism software. Statistical analyses are described in brief in figure legends and are presented in detail in the supplemental statistics (Data S1). Statistical significance was set at *p* < 0.05; **p** < 0.05; ***p** < 0.01; ****p** < 0.001.
Supplemental information

APP fragment controls both ionotropic
and non-ionotropic signaling of NMDA receptors

Jade Dunot, Sebastien Moreno, Carine Gandin, Paula A. Pousinha, Mascia Amici, Julien Dupuis, Margarita Anisimova, Alex Winschel, Magalie Uriot, Samuel J. Petshow, Maria Mensch, Ingrid Bethus, Camilla Giudici, Heike Hampel, Benedikt Wefers, Wolfgang Wurst, Ronald Naumann, Michael C. Ashby, Bodo Laube, Karen Zito, Jack R. Mellor, Laurent Groc, Michael Willem, and Hélène Marie
**Supplemental information**

APP fragment controls both ionotropic and non-ionotropic signaling of NMDA
receptors.

Jade Dunot, Sebastien Moreno, Carine Gandin, Paula A. Pousinha, Mascia Amici, Julien Dupuis, Margarita Anisimova, Alex Winschel, Magalie Uriot, Samuel J Petshow, Maria Mensch, Ingrid Bethus, Camilla Giudici, Heike Hampel, Benedikt Wefers, Wolfgang Wurst, Ronald Naumann, Michael C. Ashby, Bodo Laube, Karen Zito, Jack R. Mellor, Laurent Groc, Michael Willem and Hélène Marie

Figures S1-S15
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<figure><img src="image_62.png" /><figcaption>Figure S1 (related to Figure 1) AETA inhibits recombinant GluN1/GluN2A and GluN1/GluN2B NMDARs in oocytes by competing with glycine.</figcaption></figure>

(A) AETA inhibition curves for GluN1/GluN2A and GluN1/GluN2B, from which the respective IC₅₀ of AETA inhibition were calculated. Data represent mean ± s.e.m. n = 6 independent oocytes per condition. (B-C) pEC₅₀ values and EC₅₀ were calculated from dose-response activation curves for glycine (Figure 1I) for GluN1/GluN2A (B) and GluN1/GluN2B (C). n= oocytes per condition. See supplemental statistics (*Data S1*) for full statistics.
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<figure><img src="image_65.png" /><figcaption>Figure S2 (related to Figure 1) AETA inhibits native NMDAR current in cultured hippocampal neurons and adult CA1 pyramidal neurons exhibiting a stronger inhibition for GluN2A containing NMDARs.</figcaption></figure>

(A) (Top) Representative traces of NMDAR current elicited in cultured neurons in response to 5 applications of NMDA (30 µM/10 s) before (baseline) and 10 minutes after incubation with 10nM of CtrlP or AETA; scale bars: 10 s/100 pA. (Bottom) Graph represents time course of % peak current (normalized to baseline average of the 5 pulses) of recordings as shown in top. n= 6 neurons per condition. (B) Time course of % NMDAR current (normalized to averaged baseline) recorded at CA3-CA1 synapse of adult mouse slices upon application of 10 nM CtrP or AETA (Data quantified and shown in Figure 1C). (C) (Left, Right) Representative traces and (middle) Bar graph of % NMDAR current (normalized to averaged baseline) recorded at CA3-CA1 synapse of adult mouse slices before (baseline) and after application of AETA (10 nM) or (30 nM) in aCSF or aCSF containing GluN2A current inhibitor TCN 201 (10 µM) or aCSF containing GluN2B current inhibitor ifenprodil (5 µM). 10 nM AETA still had an effect in presence of ifenprodil (blockage of GluN2B). While 10 nM AETA did not have any effect in presence of TCN 201 (blockage of GluN2A), 30 nM AETA still partially inhibited the TCN 201-insensitive current. See supplemental statistics (*Data S1*) for full statistics.
<figure><img src="image_66.png" /><figcaption>Figure S3 (related to Figure 1) - AETA does not impact native synaptic AMPAR currents nor short term pre-synaptic plasticity.</figcaption></figure>

(A) Bar graph of evoked AMPAR current peak amplitude at CA3-CA1 synapse in mouse hippocampal slices 25-30 min post application of either AETA (10 nM) or CtrlP (10 nM). (B) Representative traces of AMPAR sEPSCs before and after application of CtrlP (top) or AETA (bottom) (10 nM). (C) Representative recorded trace of AMPAR sEPSCs (stars) that disappear upon application of AMPAR blocker NBQX. (D) AMPAR sEPSC frequency and (E) amplitude calculated before (baseline, BL) and after bath application of CtrlP (top) or AETA (bottom) (10nM). (F) Paired-pulse ratio (100 ms inter-stimulus interval) measured before (baseline, BL) and after (10 min) application of AETA (50 nM) at CA3-CA1 synapse in mouse hippocampal slices. (n/N= neuron/mice). See supplemental statistics (*Data S1*) for full statistics.
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Figure S4 (related to Figure 1) – AETA modifies spontaneous NMDAR EPSCs frequency but not amplitude.

(A) Representative traces of NMDAR sEPSC before and after application of CtrlP (top) or AETA (bottom) (10 nM). (B) Representative recorded trace of NMDAR sEPSCs (stars) that disappear upon application of NMDAR blocker D-APV. Note also the shift in tonic current that represents blockage of tonically activated NMDARs. (C) NMDAR sEPSC frequency and (D) Amplitude calculated before (baseline, BL) and after (10 min) bath application of AETA or CtrlP (10 nM). See supplemental statistics (*Data S1*) for full statistics.
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<figure><img src="image_73.png" /><figcaption>Figure S5 (related to Figure 1) : Kinetics of AMPAR and NMDAR EPSCs.</figcaption></figure>

(A) representative trace of NMDAR EPSCs with zoom windows on small and large EPSCs. (B);

representative trace of AMPAR EPSCs with zoom windows on small and large EPSCs. (C):

Representative averaged (20 traces) AMPAR and NMDAR EPSCs with their kinetic values.
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<figure><img src="image_77.png" /><figcaption>Figure S6 (related to Figure 2) : AETA still modifies NMDAR conformation in presence of NMDAR competitive antagonist APV and increases p38 phosphorylation.</figcaption></figure>

(A) Graph represents GFP lifetime in GluN1-GFP only (Don. only) and GluN1-GFP/GluN1-mCherry (Don. + Acc.) clusters 10 min after exposure to Ctrl (Tyrode) (Don. Only, n = 62 clusters; Don. + Acc., n = 69 clusters) or APV (50µM; Don. Only, n = 74 clusters; Don. + Acc., n = 65 clusters) or AETA (10 nM; Don. Only, n = 62 clusters; Don. + Acc., n = 69 clusters) (B) Graph represents GFP lifetime in GluN1-GFP and GluN1-GFP/GluN1-mCherry clusters 10 min after exposure to Ctrl (Tyrode; Don. only n = 62; Don. + Acc. n = 79) or APV (50µM; Don. only n = 79; Don. + Acc. n = 65) then 10 min after exposure to AETA (10 nM). Bottom, Illustration of movement of GluN1 intracellular tail in basal condition (Ctrl or APV) and in presence of AETA. (C) Example immunoblotting of phosphorylated p38 (p-p38) and p38 (same gel) of neuron culture lysates in control (Ctrl) or after 20 min of incubation with AETA (10 nM). The analysis was performed in duplicate. Full blots are provided in additional supplementary material (Data S2). (D) Ratio of p-p38 on total p38 obtained from immunoblots shown in (C). n= number of independent samples. Statistics: (A) paired t-test and Mann Whitney; (B) paired t-test and Wilcoxon test; (D) Mann Whitney test (D). See supplemental statistics (Data S1) for full statistics.
<figure><img src="image_78.png" /><figcaption>Figure S7 (related to Figure 2) – LTD is induced by sub-threshold protocol in presence of AETA and MK801-independent LTD is observed at CA3-CA1 synapse in rat hippocampal slices.</figcaption></figure>

(A) Summary graph (right) of fEPSP slope (% baseline) pre- and post-subLTD induction (time 0) in control (Ctrl, aCSF only) or in presence of AETA (10 nM) throughout recording. (B) Summary of fEPSP magnitude 45-60 min after subLTD induction as fEPSP (% baseline) for data shown in (A). (C) Summary graph of fEPSP slope (% baseline) pre- and post-LTD induction (time 0) in control (Ctrl, aCSF only) or after 3h of pre-incubation with MK801 (100 µM, also present in recording bath) recorded at CA3-CA1 synapse in rat hippocampal slices. (D) Summary of fEPSP magnitude 45-60 min after LTD induction as fEPSP (% baseline) for data shown in (C). (E) Representative trace of EPSC recorded in control (black), after NBQX (red) and APV (orange) without (left) or after 3 hours of preincubation of slice in MK801. The NMDAR-dependent current, normally observed under NBQX (red trace on left recording) is completely absent after MK801 preincubation (red trace on right recording). τ weight value is also shown for the two black traces showing loss of slow current after MK801 incubation. Statistics: Two-tailed unpaired Student's t-test (B and D). See supplemental statistics (Data S1) for full statistics.
<figure><img src="image_79.png" /><figcaption>Figure S8 (related to Figure 3) : Endogenous increase of AETA by *in vivo* BACE-1 inhibition leads to decreased NMDAR EPSC frequency.</figcaption></figure>

(A) Representative traces of NMDAR sEPSCs recorded in CA1 pyramidal of hippocampal slices from saline and LY2811376 treated mice. (B) NMDAR sEPSC frequency and (C) Amplitude calculated from traces as shown in (A). n/N= neurons/mice. See supplemental statistics (*Data S1*) for full statistics.
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<figure><img src="image_87.png" /><figcaption>Figure S9 (related to Figure 3) – *In vivo* viral expression of hM3Dq-mCherry and hM4Di-mCherry and blots for quantification of AETA and synaptic proteins in hM3Dq and hM4Di transduced tissues.</figcaption></figure>

(A) Representative image of AAV-hM3Dq-mCherry transduction in the prefrontal cortex (PFC). (B) Representative image of AAV-hM4Di-mCherry transduction in the PFC. (C-D) 18 mice were transduced with AAV-hM3Dq-mCherry and were injected i.p. with saline (9 mice) or CNO (9 mice). 30 min post injection, mCherry fluorescent tissues (both PFC injection area per mouse) were collected and processed for immunoblotting for AETA, mCherry and β-actin. 4 control PFC samples were also processed and blotted to check for specificity of mCherry antibody. (E-F) Same experiment as in (C-D) but for 20 mice transduced with AAV-hM4Di-mCherry (10 saline and 10 CNO). (G-H) overall health of infected tissue for each viral transduction was confirmed by reblotting some of the membranes with two synaptic markers: synaptophysin and synaptobrevin-2. * denotes two samples that were excluded from statistical analysis because AETA/mCherry ratios were detected as outliers in GraphPrism. Full blots are provided in additional supplementary material (Data S2).
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Figure S10 (related to Figure 4) - Blots used for quantification of AETA and CTF-η in WT and APPdelETA hippocampi and measures of AMPAR sEPSC, NMDAR sEPSC amplitude, paired pulse ratios (PPR) and LTP in APPdelETA mice.

(A-B) Two immunoblots for analysis of AETA and β-actin (loading control) levels in DEA samples. AETA level was normalized to β-actin level and then normalized to average WT level for each immunoblot.

(C) Immunoblot for analysis of CTF-η and β-actin (loading control) levels in RIPA samples. CTF-η level was normalized to β-actin level and then normalized to average WT level. Full blots for (A-C) are provided in additional supplementary material (Data S2). (D) Representative traces of AMPAR sEPSC recorded in WT (top) or APPdelETA (bottom) CA1 pyramidal neurons. (E) AMPAR sEPSC frequency (top graph) and amplitude (bottom graph calculated from traces as shown in (D)). (F) NMDAR sEPSC amplitude is normal in APPdelETA mice and not modulated by application of synthetic AETA (10 nM).

(G) Example traces and PPR measured at three interstimulus intervals (ISI; 100, 200 and 200 ms) at CA3-CA1 synapse in hippocampal slices of WT and APPdelETA mice. (H) Example traces and summary graph of fEPSP slope (% baseline) pre- and post-LTP induction (2x 100Hz/1s at 20 s inter-stimulus interval; time 0) at CA3-CA1 synapse in hippocampal slices of WT and APPdelETA mice. (I) Bar graph of fEPSP magnitude 45-60 min after LTP induction as fEPSP (% baseline) for data shown in (H). n/N= neurons/mice for E-F; n/N= slices/mice for G-I. See supplemental statistics (*Data S1*) for full statistics.
<figure><img src="image_99.png" /><figcaption>Figure S11 (related to Figure 4) : Absence of η-secretase processing of APP leads to an increase in the number of NMDAR-only containing synapses (silent synapses) in CA1 pyramidal neurons.</figcaption></figure>

(A) Example of a minimal stimulation experiment showing decrease in failures when recording EPSCs at +40 mV versus -65 mV because of activation of silent synapses at +40 mV. (B) Average percent success rate when recording at -65 mV versus +40 mV in WT (n/N = 19/3; left panel) and APPdelETA (n/N (neurons/mice) = 14/3; right panel) neurons. See supplemental statistics (Data S1) for full statistics.
<figure><img src="image_100.png" /><figcaption>Figure S12 (related to Figure 4) : Absence of η-secretase processing of APP does not perturb the number of spines in the stratum radiatum of CA1 pyramidal neurons.</figcaption></figure>

(A) Example images of segments of dendrites stained by Golgi-Cox and used for analysis of spine density. (B) The number of spines per micron was counted in segments from dendrites present in the stratum radiatum of CA1 pyramidal neurons from WT and APPdelETA mice (170 and 174 segments, respectively). (C) Same data as in (B) but averaged per mouse (4 mice per genotype). See supplemental statistics (Data S1) for full statistics.
<figure><img src="image_101.png" /><figcaption>**Figure S13 (related to Figure 4):** L689,560 (L689), a potent antagonist of the glycine/D-serine site of NMDARs, does not rescue LTD in APPdelETA mice. (A) Representative traces (left, 1 shows trace pre, 2 shows trace post-induction; scale bars: 10 ms / 0,2 mV) and summary graph (right) of fEPSP slope (% baseline) pre- and post-LTD induction (900 pulses at 1 Hz, time 0) at CA3-CA1 synapse in hippocampal slices of WT and APPdelETA mice without or with supplementation of 10 µM L689 (in recording bath). (B) Bar graph of fEPSP magnitude 45-60 min after LTD induction as fEPSP (% baseline) for data shown in (A). n/N= slices/mice. See supplemental statistics (*Data S1*) for full statistics.</figcaption></figure>
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Figure S14 (related to Figure 4) - characterization of AETA-m mouse overexpressing secreted human AETA in the brain. (A) Diagram of production of AETA-m line. (B) Example of immunoblot to detect endogenous AETA (rodent (rod); M3.2 antibody) and recombinant secreted human (hu) AETA (long AETA-alpha form) (2D8 and 2E9 antibodies) and β-actin in brains. Full blots are provided in additional supplementary material (Data S2). (C) Quantification of endogenous rodent AETA in brain by immunoblotting. (D) Quantification of Aβ40 in brain by ELISA. See supplemental statistics (Data S1) for full statistics.
<figure><img src="image_106.png" /><figcaption>**Figure S15 – Summary diagram of AETA-dependent NMDAR activation (ADNA) mechanism.** AETA, which is produced by increased neuronal activity, competes with D-serine/Glycine to inhibit NMDAR ionotropic activity. AETA binding leads to a conformational change of NMDARs. AETA also increases p38 phosphorylation. AETA-dependent modulation of NMDARs induces long-term depression (LTD), a NMDAR function that is independent of ion flux. AETA also favors spine shrinkage, in conditions identical to spine shrinkage driven by ion flux-independent and p38-dependent NMDAR signaling. Diagram created by Biorender.com.</figcaption></figure>