Precision Mapping of Amyloid-β Binding Reveals Perisynaptic Localization and Spatially Restricted Plasticity Deficits

Abstract Secreted amyloid-β (Aβ) peptide forms neurotoxic oligomeric assemblies thought to cause synaptic deficits associated with Alzheimer’s disease (AD). Soluble Aβ oligomers (Aβo) directly bind to neurons with high affinity and block plasticity mechanisms related to learning and memory, trigger loss of excitatory synapses and eventually cause cell death. While Aβo toxicity has been intensely investigated, it remains unclear precisely where Aβo initially binds to the surface of neurons and whether sites of binding relate to synaptic deficits. Here, we used a combination of live cell, super-resolution and ultrastructural imaging techniques to investigate the kinetics, reversibility and nanoscale location of Aβo binding. Surprisingly, Aβo does not bind directly at the synaptic cleft as previously thought but, instead, forms distinct nanoscale clusters encircling the postsynaptic membrane with a significant fraction also binding presynaptic axon terminals. Synaptic plasticity deficits were observed at Aβo-bound synapses but not closely neighboring Aβo-free synapses. Thus, perisynaptic Aβo binding triggers spatially restricted signaling mechanisms to disrupt synaptic function. These data provide new insight into the earliest steps of Aβo pathology and lay the groundwork for future studies evaluating potential surface receptor(s) and local signaling mechanisms responsible for Aβo binding and synapse dysfunction.


Introduction
Amyloid-b (Ab) is widely recognized as a primary neuropathologic agent in Alzheimer's disease (AD). Formed by proteolytic processing of amyloid precursor protein (APP), Ab peptide self-associates to form soluble oligomers and fibrils before eventually depositing into the hallmark plaques associated with AD (Seubert et al., 1992;Shoji et al., 1992;Lambert et al., 1998;Gong et al., 2003;Lesné et al., 2006). While Ab plaques correlate with neuronal dysfunction and cell death, considerable evidence supports a major role for soluble, oligomeric Ab (Abo) in synapse toxicity. For example, synapse loss and memory impairment in AD can occur before widespread plaque formation (Dekosky and Scheff, 1990;Terry et al., 1991). Acute exposure to nano-to picomolar quantities of Abo, either in vivo or in vitro, is sufficient to block neural plasticity within minutes, trigger synapse elimination over days, and eventually cause cell death (Walsh et al., 2002;Shankar et al., 2007Shankar et al., , 2008Wei et al., 2010;Sinnen et al., 2016).
Pioneering studies demonstrated Abo preferentially accumulates at excitatory synapses within minutes following application of synthetic Ab-derived diffusible ligands (ADDLs; Lacor et al., 2004;Koffie et al., 2009;Renner et al., 2010). Consistent with these studies, naturally derived Ab produced over much longer timescales (months/years) also accumulates at synaptic sites in both animal AD models and human AD patients (Games et al., 1995;Gong et al., 2003;Koffie et al., 2009;Pickett et al., 2016). While these observations suggest a direct role of Ab in synapse toxicity, little is known about the earliest steps of Abo binding. Precisely where does Abo engage neurons relative to synaptic connections? Does it bind to presynaptic or postsynaptic compartments? How fast does it associate and dissociate? Are only Abo-bound synapses impaired? Addressing these questions will be important for understanding the mechanisms of Abo toxicity. For example, mapping where Abo binds to neurons with nanometer precision will be imperative for evaluating putative Abo receptors. Thus far, over 20 Ab receptors have been described, each with diverse subcellular localizations, including the presynaptic membrane, the postsynaptic membrane, perisynaptic and nonsynaptic sites, yet whether Abo directly binds at these sites remains unclear. Furthermore, whether Abo binding triggers cell wide synaptic dysfunction or selectively impairs synapses to which it is bound is not known.
Using longitudinal live imaging, we demonstrate acutely applied Abo rapidly forms stable clusters on the neuronal cell surface. In agreement with previous studies, Abo preferentially associates with excitatory synapses. However, super-resolution light microscopy, immunogold electron microscopy (EM), and expansion microscopy (ExM) revealed that Abo does not bind directly at the synaptic cleft, but instead forms stable nanoscale clusters encircling the postsynaptic membrane with a significant fraction also binding the presynaptic axon terminal. Finally, we used two-photon glutamate uncaging at individual synapses to demonstrate plasticity deficits are restricted to Abo-bound spines. Together, these results provide the first quantitative, super-resolution interrogation of the earliest steps of Abo binding, dynamics and local toxicity at synaptic sites. Defining precisely where Abo initially engages the neuronal surface is a key step in understanding how Abo causes synaptic dysfunction and for directing future strategies aimed at preventing Abo-induced pathology.

Cell culture and transfection
All animal procedures were conducted in accordance with the Institutional Animal Care and Use Committee at the University of Colorado, Anschutz Medical Campus. Primary hippocampal cultures were made from postnatal day (P)0 to P1 Sprague Dawley rats as previously described (Sinnen et al., 2016) and maintained in Neurobasal media (Invitrogen) with B27 supplement (Invitrogen) for 15-18 d in vitro (DIV) before experiments. Neurons were typically transfected between DIV15 and DIV17 using Lipofectamine 2000 according to the manufacturer's instructions. Plasmids used in this study include: PSD95 FingR -GFP (Gift from Don Arnold, University of Southern California) and pCAG-mCh, pCAG-GFP and pSyn-tdtomato plasmids (where pCAG is the chicken b-actin promoter; pSyn is human synapsin promoter).

Ab preparation
Soluble Ab1-42 (Anaspec) oligomers were prepared similar to a previously reported method (Klein, 2002). Briefly, Ab was dissolved in 1,1,1,3,3,3-hexafluoro-2-propanol, aliquoted and dried in a chemical fume hood and stored at À80. The day before use, Ab (6 nmol) was dissolved in 4 ml of dimethyl sulfoxide and then 60 ml of PBS was added for a final concentration of 94 mM. The dissolved peptide was incubated at 4°C overnight. Following 12-24 h of incubation, the sample was centrifuged at 14,000 Â g at 4°C. The supernatant was reserved and applied to a size exclusion spin filter (30-kDa cutoff; Millipore, MRCFOR030) and centrifuged for 10 min at 10,000 Â g at room temperature to remove low molecular weight Ab species. The high molecular weight fraction was diluted to a final volume of 600 ml with PBS (working concentration of 10 mm) and stored on ice until use. For experiments using fluorescent Abo, the preparation was conducted as described above with HiLyte647-conjugated Ab (AnaSpec) included at a molar ratio of 1:3, labeled:unlabeled Ab peptide.

Live-cell imaging
Live-cell imaging of dissociated neurons was performed at 31°C on an Olympus IX71 equipped with a spinning-disk scan head (Yokogawa). Excitation illumination was delivered from an acousto-optic tunable filter (AOTF) controlled laser launch (Andor).
Images were acquired using a 60Â Plan Apochromat 1.4 numerical aperture objective and collected on a 1024 Â 1024-pixel Andor iXon EM-CCD camera. Data acquisition and analysis were performed with MetaMorph (Molecular Devices), Andor IQ, and ImageJ software.
For live cell Abo binding experiments, z-stacks were acquired every 10 s. Labeled Ab was added to the imaging chamber following a baseline acquisition imaging period. For Abo dissociation experiments, Abo was added to the imaging chamber and allowed to bind for 10 min. The imaging media was then exchanged by washing 3Â with Abo-free media. Synapse-associated Abo was quantified by creating a binary mask based on the postsynaptic density protein 95 (PSD95) signal and then calculating the average, background-subtracted integrated density. Extrasynaptic Abo was quantified within a mask created by subtracting the PSD95 mask from a cell fill mask. Binding kinetics were calculated by fitting plots of the Abo fluorescent signal (F/F 0 ) versus time with a single exponential function.

Fluorescence recovery after photobleaching (FRAP)
Abo was added to coverslips for at least 10 min to allow binding to saturate. Baseline images were acquired once every 15 s for 12 frames. Abo puncta were bleached using galvanometric mirrors (FRAPPA module, Andor Technologies) to steer a diffraction limited excitation spot over the region of interest. Photobleaching was typically conducted using 60% laser power from a fiber-coupled 100-mW 641-nm laser with a dwell time of 1 ms. Following photobleaching, images were acquired at 1 frame/min for 25 min.

Structural LTP/2-photon glutamate uncaging
Structural long-term potentiation (LTP)/two-photon glutamate uncaging two-photon glutamate uncaging and imaging were conducted using a Bruker Optima laser scanning microscope equipped with a Mai-Tai DeepSee laser (Spectra-Physics) for imaging and a Mai-tai laser (Spectra-Physics) for uncaging. Hippocampal neurons transfected with green fluorescent protein (GFP) or tdTomato expressing plasmids were treated with Abo generated with either HiLyte568 or HiLyte488-labeled Ab peptide respectively. Full z-stacks were acquired to identify Abo-bound spines using sequential 920/1040-nm excitation (GFP/HiLyte568) or single 920-nm excitation (tdTom/HiLyte488 Abo). Abo-positive and negative spines were subject to glutamate uncaging in artificial CSF (ACSF) containing 3 mM Ca 21 and lacking Mg 21 . Uncaging power and duration were calibrated so that dendritic spine Ca 21 influx triggered by glutamate uncaging (measured in separate cells expressing GCaMP6) matched Ca 21 influx resulting from spontaneous glutamate release (Sinnen et al., 2016). Spine growth was triggered by MNI-glutamate (2 mM) uncaging at 720 nm with a train of 45 1-ms pulses delivered at 0.5 Hz at a single spot adjacent to the tip of the targeted spine. A mix of Abo-positive and negative spines were selected from each cell. Z-stacks were acquired every 90 s to visualize spine morphology preglutamate and postglutamate uncaging.

Structured illumination microscopy (SIM)
Multichannel SIM images of Ab-treated neurons were acquired with a Nikon N-SIM E SIM using a 100 Â 1.49 NA objective, and reconstructed using Nikon Elements software as described previously with minor modifications (Smith et al., 2014;Crosby et al., 2019). Imaging parameters (laser power, exposure) were optimized for a high signal-to-noise ratio (.8). For each coverslip imaged, the objective correction collar was adjusted automatically and a Fourier transform image was used to confirm optimal correction collar adjustment. Z-stacks (z = 0.2 mm, 13 slices) were reconstructed using Nikon Elements software. For three-dimensional (3D) stack reconstruction, the illumination modulation contrast was set automatically and the high-resolution noise suppression was set to 1, and kept consistent across all images.

SIM analysis
Quantification of Ab density distribution relative to specific proteins of interest was performed using custom analysis software written in MATLAB along with the freely available MATLAB Toolbox DipImage (Delft). Proteins of interest were identified as follows. Each channel of the image was smoothed using a Laplacian or Gaussian filter to enhance punctate objects with a kernel size of two pixels in X and Y and one pixel in Z. An automatic intensity threshold was calculated using the MATLAB multithresh function to identify two threshold levels based on the image intensity histogram. The higher threshold was used to generate a mask for each image. The DipImage label function was then used to identify individual objects from the mask and then a 3D Euclidean distance transform was applied using the MATLAB function bwdistc1.m, (Mishchenko, 2015), resulting in a new distance image in which each voxel of the image represents the 3D distance to the closest masked object. To mask the Ab signal a similar process was used, however, because the Ab puncta were more densely spaced, an additional watershed filter was used to improve segmentation. Watershed lines were computed from the Gaussian filtered (sxy = 1) original image with a connectivity of 1 pixel on a frame-to-frame basis using the DipImage gaussf and watershed functions and then subtracted from the Ab mask. The center of mass positions for each labeled Ab puncta were next identified using the DipImage measure function. The distance image could then be used to identify the shell of voxels within a specified distance from the proteins of interest. The count of Ab puncta center positions within this volume of voxels approximates the Ab density within the specified distance range. Resulting densities were then divided by a normalization term representing the expected density from a uniform Ab distribution, such that values .1 represent an Ab density above uniform. To generate this normalization term, a simulation for each image was performed by randomly distributing the same number of Ab puncta found in the original image within 642 nm (20 Â X-Y pixel size) of the proteins of interest and then calculating the density within each distance range. To prevent any artifact arising from the difference in Z pixel size compared with X-Y pixel size in SIM images, the number of Ab puncta at each z plane was kept the same between the original image and the simulated image.
Direct stochastic optical reconstruction microscopy (dSTORM) imaging and analysis Cells exposed to 500 nM Abo-647 for 10 min or anti-GluA1 for 15 min were fixed and labeled with anti-PSD95 as described above. Secondary antibodies were conjugated to either Alexa Fluor 647 or CF568. Following secondary antibody labeling, cells were postfixed with 4% paraformaldehyde for 15 min. Samples were imaged in a buffer containing 50 mM Cysteamine hydrochloride, 10% glucose, 0.6 mg/ml glucose oxidase from Aspergillus niger, 0.063 mg/ml Catalase from Bovine liver in PBS, pH between 7.5 and 8.0. Imaging was performed on a Zeiss Elyra P.1 TIRF microscope using a Zeiss a Plan Apochromat TIRF 100Â/1.46 NA oil objective and a tube lens providing an extra factor of 1.6Â magnification. Alexa Fluor 647 (or HyLite647) and CF568 dyes were imaged in sequential time-series of ;20,000 frames each. Image size was 256 Â 256 pixels, integration time was 18 ms for both channels. Alexa Fluor 647 or HyLite547 molecules were ground-state depleted and imaged with a 100-mW 642-nm laser at 100% AOTF transmission in ultra-high-power mode (condensed field of illumination), corresponding to ;1.4 W/cm 2 . Emission light passed through a LP655 filter. CF-568 molecules were groundstate depleted and imaged with a 200-mW 561-nm laser at 100% AOTF transmission in ultra-high power mode, corresponding to ;2.5W/cm 2 . Emission light was passed through a BP 570-650 1 LP 750 filter. For each dye, ground-state return was elicited by continuous illumination with a 50-mW 405-nm laser at 0.01-0.1% AOTF transmission. Excitation light was filtered by a 405/488/561/642 filter placed in front of the camera. Images were recorded with an Andor iXon1 897 EMCCD. The camera EM gain was set to 100, which yields an effective conversion of 1 photograph electron into 1.65 digital units. The image pixel size was 100 nm xy.

Processing
Raw data were processed through a custom pipeline written in MATLAB (MathWorks) made up of a number of modular elements, described below. The Bio-Formats MATLAB toolbox (Linkert et al., 2010) was used to read Zeiss raw data files into MATLAB. Image data were transferred between MATLAB and FIJI using MIJI (http:// bigwww.epfl.ch/sage/soft/mij/). If necessary, raw data were preprocessed with a temporal filter (Hoogendoorn et al., 2014) to remove nonhomogeneous background. The filter radius was set at 51 frames, with a key frame distance of 10 (filter is explicitly calculated only for every 10 frames and interpolated between), the quantile for the filtering was set a 20%. Localization of dye emitters was performed using the ThunderSTORM ImageJ plugin (Ovesný et al., 2014). The camera EM gain was set to 100, which resulted in a photon-to-ADU of 1.65. When the temporal median filter was used, the Offset was set to zero. Image filtering was done with the Wavelet filter setting, with a B-Spline order of three and scale of 2.0. A first pass approximate localization of molecules was achieved with by finding local maximum with a peak intensity threshold of 2.5*std(Wave.F1) and 8-neighborhood connectivity. Weighted least squares fitting of the PSF to achieve subpixel localizations was achieved by use of an integrated Gaussian with a fitting radius of four pixels and an initial s of 1.5. Localizations were filtered based on the attributes of uncertainty (,20 nm) and s (100-200 nm for CF568 and 90-190 nm for Alexa Fluor 647 and Hylite-647). Before each experiment a calibration was calculated to correct for shifts and distortions between the acquired fluorescent channels. Subdiffractive beads, labeled with fluorophores in both channels were imaged. The bead positions were fitted and registered between the fluorescent channels. Registered localizations from multiple bead images were compiled into one data-set. Calibration matrices of the shift in x and y direction between the imaging channels across the full field of view were calculated by either applying a 2D polynomial fit or a localized weighted averaging to the registered bead localizations. In the raw data, the shift and distortion between the imaging channels was up to 100 nm. Applying the calibration to the STORM data yields an RMS error of ,15 nm for the channel misalignment. Drift correction was performed using the redundant cross-correlation method described previously (Wang et al., 2014). The segmentation parameter was set at 500 frames, the bin size used in the cross-correlation was 10 nm, and the error threshold for the recalculation of the drift was five pixels.

dSTORM analysis
Coordinate analysis of our dSTORM data are conceptually similar to methods previously used to classify nanoscale organization at the excitatory synapse (Tang et al., 2016). Synapses for downstream analysis were selected manually from a composite rendered image and ROI coordinates were recorded using a custom ImageJ macro. ROI details were imported into MATLAB using the ReadImageJROI function (https://github.com/DylanMuir/ ReadImageJROI). The postsynaptic density and synaptic Ab localization were segmented using a coordinate-by-coordinate density calculation. Because labeling density could vary greatly, the thresholding parameter was determined from the overall density range of the ROI. Localizations with a local-density in the lower 10% of that range were considered to be outside of the synaptic region/clusters. Boundaries for these regions were delineated using MATLAB's alphaShape function, with an a value of 100. Only regions with an area of 1.5e3 nm 2 or greater were considered for analysis. MATLAB's inShape function was used to determine what percentage of Ab or receptor localizations fell within the PSD boundary. Ab and receptor nanoclusters were defined by a cutoff determined by randomizing the experimental localizations assuming a uniform distribution across the synaptic region. The local density threshold for an experimental coordinate to be considered as part of a nano-cluster was set at the mean local density of the randomized dataset plus 2 SDs. The geometric boundaries of individual nano-clusters were again delineated using the alphaShape function, with an a value of 11. Ab or receptor nanoclusters were classified as overlapping with the PSD if the overlap area had a fraction of 0.23 or greater of the total nanocluster area. The weighted center (mean of coordinates) of each nanocluster was calculated and the center to PSD edge was determined using the nearestNeighbor function. This distance was assigned as zero for PSD overlapping nanoclusters.

Electrophysiology
Field recordings were performed as previously described using two-to three-week-old C57BL/6 mice (Freund et al., 2016). Mice were killed and brains rapidly removed and immersed in ice-cold sucrose-containing cutting buffer (2 mM KCl, 12 mM MgCl 2 , 0.2 mM CaCl 2 , 1.3 mM NaH 2 PO 4 , 10 mM D-glucose, 220 mM sucrose, 26 mM NaHCO 3 , 1.77 mM sodium ascorbate, and 2 mM N-acetylcysteine). Coronal slices containing hippocampus (400-mm thickness) were prepared using a McIlwain tissue chopper/ slicer and recovered at 27°C for .60 min in ACSF (84.3 mM NaCl, 3 mM KCl, 1.8 mM CaCl 2 , 1.3 mM NaH 2 PO 4 , 4.7 mM MgSO 4 , 26 mM NaHCO 3 , 10 mM glucose, 70.4 mM sucrose, 1.2 mM sodium ascorbate, and 0.65 mM N-acetylcysteine). After recovery, a single slice was transferred to a recording chamber and superfused with ACSF at a flow rate of 2-3 ml/ min at 31°C. The ACSF contained the following: 124 mM NaCl, 3.5 mM KCl, 1.3 mM MgCl 2 , 2.5 mM CaCl 2 , 1.3 mM NaH 2 PO 4 , 10 mM D-glucose, and mM 26 NaHCO 3 . Field recordings were made with a glass micropipette filled with ACSF placed in CA1 stratum radiatum ;200-300 mm from the cell body layer. Synaptic field EPSPs (fEPSPs) were evoked with bipolar tungsten electrodes placed in the Schaffer collateral axon pathway. For each slice, an input-output curve was generated by increasing the stimulus voltage and recording the synaptic response until either a maximum was reached or evidence of a population spike was observed on the fEPSP response. The control stimulus intensity was set to 40% to 50% of the maximum synaptic response, and a baseline recording was obtained delivering one test pulse every 20 s for 20 min. To elicit LTP, we delivered two trains of 100-Hz stimuli lasting 1 s each, with an intertrain interval of 5 min. This protocol reliably produced LTP that persisted for .45 min. We recorded the maximum amplitude of the fEPSPs as well as their initial slopes, measured between 10% and 40% from the point of negative deflection.

EM and immunogold labeling
For EM and immunogold labeling of neuronal cultures, we used osmium-free processing as previously described (Phend et al., 1995). Briefly, samples were fixed with 2.5% glutaraldehyde in 0.1 M phosphate buffer, and sequentially treated with 1% tannic acid (EM Sciences), 1% uranyl acetate, 1% PPD, 0.2% iridium tetrabromide, and then were dehydrated and embedded for sectioning. Following ultrathin microtome sectioning, samples were stained for antibodies against Ab using previously described postembed gold methods (Aoki, 2003). Briefly, grids, were blocked in tris-buffered saline (0.9% sodium chloride) containing 0.1% Triton X-100 (TBST; pH 7.6) and incubated in primary antibody diluted in TBST at room temperature overnight. The following day, grids were washed with TBST and then blocked with TBST, pH 8.2. Grids were incubated for 1 h in donkey anti-mouse secondary conjugated to 10 nM gold (EMS, 25825). Grids were washed, with TBST, H 2 O, postfixed with 1% glutaraldehyde, and counterstained with Reynold's lead citrate.

ExM
Samples were prepared according to the ExM protocol outlined in Zhang et al. (2020). Live DIV16 hippocampal cultures were incubated with rabbit anti-GluA1 for 10 min before fixation (Kennedy et al., 2010;Hiester et al., 2017) Abo (500 nM) was added to live cells for 15 min at 37°C. Cells were fixed for 15 min using 4% paraformaldehyde (Electron Microscopy Sciences) and washed with PBS. Cells were blocked for 30 min in PBS containing 5% bovine serum albumin (Sigma) and stained with mouse anti-Ab (6E10, Biolegend, 803014). Cells were then permeabilized with 0.2% Triton X-100 (Fisher Scientific) and incubated with guinea pig anti-bassoon (Synaptic Systems, 141-004). Following incubation with secondary antibodies (secondary antibodies were generated in goat and include anti-rabbit Alexa Fluor 488; anti-rabbit Alexa Fluor 568; anti-guinea pig Alexa Fluor 488; anti-guinea pig Dylight 550; anti-mouse Abberior Star 635) cells were exposed to a second round of fixation with 3% PFA/0.1% glutaraldehyde (Electron Microscopy Sciences) and sequentially washed in 1Â PBS and distilled water. Acroloyl-X SE (AcX; Invitrogen) diluted at 1:100 in PBS was added to each well and cells were refrigerated overnight. On day 2 of the protocol, the AcX solution was removed and coverslips were washed twice in 1Â PBS for 5 min (samples were placed on ice at the start of the second wash). Gelation solution was prepared by combining chilled reagents: Stock X (Zhang et al., 2020), TEMED (Fisher Bioreagents), and ammonium persulfate (APS; Sigma) at a volumetric ratio of 98:1:1. After removing PBS, 500 ml of gelation solution was added to each well for 5 min on ice. Coverslips were then placed cell-side down atop a gelation chamber constructed using a glass microscope slide and cover glass (Zhang et al., 2020); 60 ml of gelation solution was quickly added underneath each coverslip, and the chambers incubated at 37°for 1 h in a humidified chamber. Gelled samples were submerged in 10-ml digestion buffer (Zhang et al., 2020) containing proteinase K in a 100 Â 20-mm Petri dish (Corning) on an orbital shaker at 60 rpm at room temperature overnight. On day 3 of the protocol, digestion buffer was removed and gels were incubated with 10-ml distilled water on an orbital shaker at 60 RPM for 10 min to allow for gel expansion. This was repeated two additional times. The expansion process was then repeated a third time, for 20 min. Following the final expansion step, gels were cut and plated on 35 Â 10-mm glass-bottom plates (Ted Pella) coated with 0.1% poly-Llysine (Sigma). A total of 2 ml of distilled water was then added to each plate and samples imaged on a spinning disk confocal microscope.

ExM analysis
Ab distribution along the presynaptic to postsynaptic axis was calculated using custom analysis software in MATLAB. Presynaptic and postsynaptic proteins and Ab puncta objects were identified, labeled and the center of mass position was calculated as described in the SIM analysis section in Materials and Methods. Synapses were filtered based on the following criteria: (1) they contained both a presynaptic and postsynaptic object, (2) there was an Ab puncta within 20 pixels in X-Y and three pixels in the Z dimension, and then further selected manually based on the criterion that the synaptic alignment relative to the imaging plane resulted in good separation between presynaptic and postsynaptic objects. Regions were selected in ImageJ and imported into MATLAB using the function ReadImageJROI.m Dylan Muir 2014. For each synapse, the center position for all Ab puncta with 20 pixels in X-Y and three pixels in Z was mapped onto a cylindrical coordinate system having a longitudinal axis defined by the vector between the postsynaptic and presynaptic center of mass positions with an origin at the midway point.

Figure processing
In some cases, images were expanded 2Â and were interpolated for display only. Volume-rendered images were created using expanded, masked images through the ImageJ volume viewer with a z-aspect of 2. All quantitative analysis was performed on raw image files.

Statistical analysis
Statistical significance for experiments comparing two populations was determined using a two-tailed unpaired Student's t test. When populations were not normally distributed, Mann-Whitney tests were used. In the cases where the two populations represented paired measurements, a paired Student's t test was used. For experiments comparing three or more populations, a One-way ANOVA with Bonferroni multiple comparison test was used. When populations were not normally distributed, Kruskal-Wallis with Dunn's multiple comparisons test were used. Statistical analyses were performed using GraphPad Prism and Microsoft excel. All data are presented as mean 6 SEM unless otherwise stated.

Results
Abo rapidly and stably accumulates on the cell surface at or near excitatory synapses To directly visualize Abo as it binds to neurons, we generated fluorescently labeled Abo using HiLyte647-labeled Ab peptide as previously described (Lacor et al., 2004;Sinnen et al., 2016). We confirmed the labeled peptide formed oligomeric species by western blotting and that it blocked LTP following high frequency stimulation in acute mouse hippocampal slices (Extended Data Fig. 1-1).
We next characterized the rate of Abo binding and its stability at different subcellular locations on excitatory neurons. Here, we performed time lapse confocal imaging as we applied labeled Abo (500 nM) to live dissociated hippocampal neurons expressing mCherry (mCh) to visualize cellular morphology and a GFP-labeled fibronectin intrabody generated with mRNA display (FingR) against PSD95 (PSD95 FingR -GFP) to label excitatory synapses (Fig. 1A). Abo binding was initially detected as diffraction-limited puncta within seconds of Abo application. In most cases, Ab puncta progressively grew in size and intensity but saturated at a maximum plateau value within 5-10 min (Fig. 1B,  C). Once maximum binding was achieved, we observed detectable Abo signal overlapping with 64 6 6.8% of excitatory synapses, in agreement with previous studies (Lacor et al., 2004;Sinnen et al., 2016). Abo binding was not exclusive to synapses; of the total Abo signal, 50 6 6.5% associated with dendritic spines and 50 6 6.5% associated with nonsynaptic sites on the dendritic shaft. Kinetic values for Abo accumulation at PSD95-positive dendritic spines and nonsynaptic sites were estimated by fitting binding curves with a single exponential function (Fig. 1C); t values for Abo association with dendritic spines versus nonsynaptic regions on dendritic shafts were similar; 3.62 6 0.47 and 3.51 6 0.46 min, respectively. However, when we compared maximum Abo fluorescence intensity at PSD95-positive dendritic spines (10 min following Abo addition) with nearby nonsynaptic signal on the dendritic shaft, Abo signal was consistently elevated near PSD95, suggesting a surface receptor(s) that is enriched at but not exclusive to excitatory synapses (Fig. 1C). A possible explanation for this enrichment is that Abo preferentially associates with a structural feature unique to dendritic spines. Previous studies proposed Abo binds directly to spine membranes by recognizing their degree of curvature (Sugiura et al., 2015;Terakawa et al., 2018). Contrary to this hypothesis, we found Abo accumulation at shaft excitatory synapses was not significantly different from spine synapses (Fig. 1D,E). Abo was added at 0 s. Scale bar: 1 mm. C, left, Quantification of Abo association kinetics at PSD95-positive dendritic spines or neighboring PSD95-negative dendritic shafts (n = 10 neurons, 4 independent cultures). Right, Plateau intensity values for Abo binding at PSD951 versus PSD95-(shaft) locations on the same cell (n = 10 neurons; p = 0.0139, paired Student's t test). D, Representative image of Abo localization to spine (arrow) and shaft (arrowhead) PSD95. Scale bar: 1 mm. E, left, Abo binding kinetics at PSD95 puncta on spines or shafts (n = 20 PSD95 puncta from 5 neurons, 3 independent cultures). Right, Plateau intensity values of Abo at PSD95 puncta on the spine and shaft of the same neuron (n = 5 neurons, p = 0.2857, paired Student's t test). ns = not significant. F, Representative image sequence of Abo (teal) bound to a dendritic spine following washout into Abo-free imaging media at t = 0 min. The cell outline is shown as a dashed line, drawn based on a cell fill (data not shown). Scale bar: 1 mm. G, Quantification of Abo intensity following washout. Data are plotted as F/F 0 , with F 0 representing normalized Abo signal immediately before washout (n = 5 neurons). H, Representative image of a dendrite from a hippocampal neuron expressing mCh (dotted line), treated with labeled Abo (cyan, left panel) for 10 min and then an extracellular antibody against Abo to assess Thus, Abo appears to recognize a feature enriched at excitatory synapses regardless of their localization or morphology. We next measured the stability of Abo at synaptic and nonsynaptic sites. Here, we added Abo to hippocampal neurons, allowed binding to saturate for ;10 min and then washed the cells into Abo-free extracellular solution (Fig. 1F). Following washout, Abo clusters remained highly stable at both synaptic and nonsynaptic sites with only 13.2 6 2.8% (synaptic) or 12.7 6 3.3% (nonsynaptic) loss in signal after 15 min (Fig. 1G). We also imaged samples that had been treated with Abo and then fixed to assess the level of photobleaching over the same imaging time window and found photobleaching could account for 2.2 6 1.1% loss in signal. We also tested whether Abo remained on the surface over the time course of this experiment by briefly applying an extracellular Ab antibody either 10 or 45 min following Abo application. In both cases we observed nearly all (92.27 6 0.047% at 45 min) Abo puncta observed directly with HiLyte647-labeled Ab peptide colabeled with the antibody signal. Thus, the observed stability was not the result of internalization into stationary intracellular organelles (Fig. 1H,I).
In a complementary set of experiments, we performed FRAP to investigate surface-bound Abo dynamics. Here, we applied Abo, waited 10 min to allow binding to saturate and then focally photobleached individual Abo puncta at synaptic sites. When we performed FRAP measurements with soluble Abo (500 nM) remaining in the ACSF, we observed limited but significant recovery of bleached surface Abo clusters (synaptic Abo: mobile fraction = 0.25 6 0.02, recovery rate = 0.15 6 0.03 min). Under these conditions, the source of signal recovery could be soluble Abo from the extracellular solution, or laterally diffusing Abo that was already associated with the cell surface (Renner et al., 2010).To distinguish these possibilities, we performed additional FRAP measurements without Abo in the extracellular solution to eliminate the soluble pool. Under these conditions, we observed a ;4-fold decrease in the mobile fraction of bound Abo (0.059 6 0.015) at synaptic sites. Thus, while surface-bound Abo appears stable at steady state, limited exchange with soluble pools can occur (Fig. 1J,K).

Super-resolution microscopy reveals Abo binds predominantly at perisynaptic sites
The nanoscale distribution of Abo surface binding remains poorly characterized. We first used SIM (Gustafsson, 2005;Crosby et al., 2019), which has ;2-fold higher resolution compared with confocal microscopy as well as improved resolution in z, to localize Abo (500 nM, applied 10 min before fixation) relative to the excitatory postsynaptic proteins GluA1 and PSD95, detected by antibody staining (Fig. 2A). Surprisingly, our SIM images revealed that most Abo surface clusters do not actually overlap with PSD95 or GluA1 signal as previously reported in studies using wide field or confocal microscopy (Lacor et al., 2004;Renner et al., 2010). Instead, most Abo signal appeared immediately adjacent to the PSD (Fig. 2B). In contrast, the synaptic receptor GluA1 appeared highly colocalized with PSD95, confirming the observed perisynaptic localization of Abo was not because of optical or SIM reconstruction artifacts (Fig. 2B). Using our SIM dataset, we also wanted to quantify the degree to which total surfacebound Abo is enriched near excitatory synapses. We developed an unbiased analysis routine that calculates the number of segmented Abo clusters encountered at different voxel distances from a segmented synaptic marker protein ( Fig.  2C; for details, see Materials and Methods). The number of Abo clusters at each distance is then divided by a randomly simulated data set with the same labeling density. Thus, if Abo binding is random with respect to the protein of interest, our analysis reports a value of one. While there was little direct overlap between Abo and excitatory synaptic proteins, we did observe a high degree of enrichment of total Abo signal within 200 nm of excitatory synapses. As a control, we found no significant enrichment of Abo at inhibitory synapses labeled with gephyrin, confirming specificity for excitatory synaptic connections ( Fig. 2D; Lacor et al., 2004;Renner et al., 2010). Consistent with perisynaptic or extrasynaptic localization of bound Abo, our analyses yielded a much greater degree of enrichment of the synaptic protein GluA1 near PSD95 (Fig. 2E).
To more precisely characterize the nanoscale organization of Abo binding with respect to synapses, we used dSTORM, which has 3-to 4-fold greater resolution than SIM (Heilemann et al., 2008). Here, we resolved Abo signal into discrete nanoscale clusters immediately adjacent to, but generally nonoverlapping with the postsynaptic density, labeled by immunostaining PSD95 (Fig. 3A). On average, Abo-bound synapses were associated with 8.04 6 1.36 Abo clusters, with individual clusters having an average area of 4548 6 600 nm 2 . To quantify the degree of Abo overlap with the PSD, we employed a density-based clustering algorithm to define the PSD (based on a threshold density of PSD95 localizations) and then calculated the fraction of individual Abo localizations that fell within the segmented PSD region (Fig. 3A). For comparison to a known synaptic protein, we also continued surface localization (green). Scale bar: 1 mm. I, Fraction of Abo puncta (averaged per cell) labeled with an extracellular antibody (i.e., localized to the cell surface) 10 and 45 min following Abo application (n = 5 neurons per group, two independent cultures). ns = not significant, Student's t test. J, Representative time series for FRAP experiments. Shown is a single Abo-bound spine. The Abo signal was photobleached and signal recovery was monitored over time. Arrowheads indicate the location of photobleaching and signal recovery. Scale bar: 1 mm. K, Kinetics and extent of Abo recovery following photobleaching in the continued presence (no wash, orange) or absence (wash, red) of Abo in the extracellular solution (wash: n = 23 spines, from 6 neurons and 2 independent cultures; no wash: n = 12 spines from 4 neurons and 2 independent cultures). Quantification of the mobile Abo fraction is shown to the right under each condition (***p = 0.0007, Student's t test).

Figure 2.
Super-resolution localization of Abo relative to excitatory and inhibitory synapses. A, Representative SIM images of Abo (teal) with excitatory synaptic proteins PSD95 (yellow), GluA1 (red), and the inhibitory synaptic protein Gephyrin (green). Bottom panels show control, GluA1 (red) with PSD95 (yellow). Scale bars:1 mm. B, Expanded regions showing individual synapses from panel A. Scale bars: 500 nm. The graphs to the right of each example plot pixel intensities for each channel along a line drawn diagonally through representative synapses. 3D volume renderings of masked and segmented synapses are shown to the right. C, Approach for quantifying the spatial relationship between synaptic proteins and Abo. i, Representative SIM image showing PSD95 (green) and Abo (cyan). The outline of the cell (dashed line) was drawn using the signal from an mCh cell fill (data not shown). ii, The Abo signal (cyan) is masked and binarized (blue). Right, PSD95 (green) is masked and binarized (red). iii, Magnified red box from ii. The number of Abo puncta are counted at increasing concentric voxel distances around masked synaptic marker. D, The number of Abo puncta (quantified as described in panel B) at different distances from either PSD95 (orange; n = 48 neurons) or gephyrin (green; n = 18 neurons), normalized to randomly localized simulated data (black; average of seven independent simulations). A value of 1 indicates no spatial relationship, .1 a positively correlated spatial relationship, and ,1 a negatively correlated spatial relationship. E, Abo is enriched near the excitatory PSD. Plotted is the average number density of segmented Abo puncta 0-64 nm from PSD95, GluA1, or gephyrin (PSD95 n = 48 neurons, GluA1 n = 16 neurons, gephyrin n = 18 neurons). The average number density of the synaptic protein GluA1 relative to PSD95 is plotted for comparison (n = 7 neurons); ****p , 0.0001, one-way ANOVA. ns = not significant. performed this analysis with GluA1 (Fig. 3B). While 77 6 1.9% of GluA1 localizations were observed within the PSD boundary, only 41 6 4.2% of Abo localizations fell within the PSD (Fig. 3C). We also performed a separate analysis where we segmented discrete, spine-localized Abo clusters and quantified their percentage overlap with the segmented PSD. Only 12.9% of spine Abo clusters fully overlapped with the PSD compared with 50.9% for GluA1. Conversely, we observed that 61.1% of spine Abo clusters were completely excluded from the PSD compared with only 28.7% for GluA1 (Fig. 3D). Combined, our super-resolution imaging data reveal the majority of spine-localized Abo does not actually bind directly at the synaptic cleft, but localizes to perisynaptic sites immediately adjacent to the PSD on the dendritic spine.

Abo binds both presynaptic and postsynaptic membranes
Abo is reported to rapidly disrupt both presynaptic and postsynaptic function, yet the relative extent of direct Abo association with axonal terminals and dendritic spines remains largely uncharacterized. To address this, we first performed postembedding immunogold EM. We treated hippocampal cultures for 10-45 min with Abo (500 nM) or an equal volume of PBS (negative control) before fixation. Following fixation, embedding, and cutting, we labeled sections with an Abo antibody and a gold-conjugated secondary antibody. We imaged samples by scanning EM and quantified the number of gold particles per linear micrometer of presynaptic or postsynaptic membrane (Fig. 4A). Given the estimated size of the primary and secondary antibodies (;10 nm each), we only included gold particles that were within 20 nm of the plasma membrane. Importantly, we observed little background signal in control (no added Abo) samples (Fig. 4B). In samples treated with Abo, 32.5% and 67.5% of the total synaptic signal was observed at the presynaptic and postsynaptic membrane respectively (Fig. 4C). Of the total postsynaptic signal, ;75% was nonoverlapping with the PSD, consistent with our dSTORM and SIM data demonstrating perisynaptic binding (Figs. 3, 4D).
Given the sparse labeling and extensive fixation/processing steps required for immunogold-EM, we took an . Abo binds at both presynaptic and postsynaptic sites. A, Representative postembedding immunogold electron micrograph of a synapse exposed to 500 nM Abo and labeled with an Ab antibody and gold-conjugated secondary. The green box (magnified in inset) highlights immunogold signal. Gold particles within 20 nm of the cell membrane were considered plasma membrane-associated based on the size of the primary and secondary labeling antibodies. Scale bar: 200 nm. B, Quantification of total membrane-associated Abo (measured as total number of gold particles per linear micron of plasma membrane) from samples treated with PBS alone or PBS with 500 nM Abo. (PBS: n = 60 spines; Ab: n = 66 spines; p , 0.0001, Mann-Whitney test). C, Quantification of Abo label on the presynaptic or postsynaptic membrane (p = 0.0005, Mann-Whitney test). D, Percentage of the total dendritic spine gold particles that localized directly at the PSD or perisynaptic regions within 200 nm of the PSD (n = 27 gold particles). E, Representative image of a dendritic segment processed for ExM, labeled for presynaptic bassoon (green), postsynaptic GluA1 (red), and Abo (cyan). The lower panels show two representative synapses (labeled 1 and 2) from the larger image and their respective 3D volume renderings. F, top, Schematic of the analysis used to quantify presynaptic and postsynaptic Abo signal. The synaptic axis is defined by a line drawn between the centers of mass of masked bassoon (presynaptic marker) and GluA1 (postsynaptic marker) signals. A vector is generated from the middle of the pre/post axis to the center of mass of the segmented Abo, with the component vectors representing the radial distance from the synapse center (r) and the distance along the pre/post axis (z). Bottom, Quantification of Abo signal along the pre/post and radial axes. Negative and positive values indicate postsynaptic and presynaptic localization respectively. The number of Abo puncta at different distances along the pre/post axis are summed and plotted in the histogram to the right (n = 89 synapses from 26 neurons from 3 independent cultures). independent fluorescence-based approach to quantify presynaptic and postsynaptic Abo. We used ExM, which allows acquisition of three or more fluorescent labels (an advantage over dSTORM) with higher spatial resolution compared with SIM. We labeled live hippocampal neurons with Abo and then fixed and stained for bassoon (presynaptic), GluA1 (postsynaptic), and Ab and expanded the preparation according to (Zhang et al., 2020;Fig. 4E). Using cellular nuclei as a reference, we estimate our samples were expanded 4-fold. We quantified Abo signal in 3D with respect to a plane perpendicular to an axis connecting the center of mass of the segmented presynaptic and postsynaptic signals ( Fig. 4F; for details, see Materials and Methods). We found that 40.8% and 59.2% of Abo signal associated with the presynaptic and postsynaptic label respectively, in agreement with our immunogold labeling. Once more, Abo did not directly overlap with synaptic markers (Fig. 4E,F). Combined, these experiments pinpoint Abo binding to sites immediately adjacent to the synaptic cleft with significant amounts of Abo directly binding both presynaptic and postsynaptic compartments.

Plasticity is disrupted specifically at Abo-bound spines
While most synapses associate with Abo at the concentration used here (500 nM), a small fraction of spines do not appear to bind Abo. This allowed us to test whether Abo selectively impairs synapses to which it is bound (Fig.  5A). One of the hallmark synaptic pathologies of Abo is LTP impairment. LTP is associated with structural enlargement of dendritic spines (sLTP) and can be locally triggered at targeted synapses using focal two-photon glutamate uncaging (Matsuzaki et al., 2004). We used an established uncaging protocol (45 pulses, 0.5 Hz) to induce sLTP at individual Abo-bound and Abo-free spines . In control cells that were not treated with Abo, our glutamate uncaging protocol triggered robust and persistent spine growth (Fig. 5B,C). Abo-bound spines from cells treated with Abo for 30-45 min before sLTP induction initially exhibited a comparable degree of Figure 5. Abo-mediated plasticity impairment is locally restricted near sites of surface binding. A, left, Representative image of a dendritic segment from a neuron transfected with tdTomato (red) and treated with Abo-488 (teal). Asterisks designate the location of MNIglutamate uncaging. The closed arrowhead shows a spine with bound Abo (Ab1), and the open arrowhead shows a neighboring spine lacking Abo (Ab-). Right, Time course of Abo-spine (top) and Abo1 spine (bottom) from the same dendritic segment before the uncaging stimulus and up to 24 min following the stimulus. Scale bars: 5 mm (left panel) and 1 mm (right panels). B, Quantification of spine size (based on the cell fill intensity) before and after MNI-glutamate uncaging for control spines not treated with Abo (blue, n = 14 spines, N = 5 neurons, 3 independent cultures), adjacent spines that were not stimulated (green, n = 12 spines, N = 5 neurons, 3 independent cultures), Ab1 spines from cultures treated with 500 nM Abo for at least 25 min (maroon, n = 16 spines, N = 8 neurons, 3 independent cultures), and neighboring Abo-lacking spines (black, n = 14 spines, 8 neurons, 3 independent cultures). C, Average increase in spine cell fill signal during the final 3 min of imaging compared with baseline for control (n = 14 spines, N = 5 neurons, 3 independent cultures), Abo1 (n = 16 spines, N = 8 neurons, 3 independent cultures), and Abo-(n = 14 spines, 8 neurons, 3 independent cultures; *p 0.05, Student's t test). ns = not significant. D, Average F/F 0 over the final 3 min of imaging compared with baseline at Abo-bound spines and neighboring Abo-free spines on the same neurons (eight neurons, three independent cultures, *p = 0.0116, paired t test). growth as controls, but returned to baseline levels several minutes following plasticity induction. Surprisingly, neighboring Abo-free spines exhibited persistent growth that was indistinguishable from control spines. In nearly every case, Abo-free spines exhibited increased persistent growth compared with nearby Abo-bound spines on the same cell (Fig. 5D). Since the extent of spine growth depends on initial size, we compared the average baseline sizes (calculated as spine area from 2D-projected images) of Abo-bound and unbound spines targeted for sLTP. There was no significant difference between the two populations (Abo-bound, 1.76 6 0.25 mm 2 ; Abo-free 1.69 6 0.19 mm 2 ). Thus, Abo disrupts plasticity in a spatially restricted manner, presumably through local signaling mechanisms constrained near sites of Abo surface engagement.

Discussion
Increasing evidence supports a role for secreted, soluble Abo as the primary culprit in AD-associated synapse dysfunction. While many studies have characterized steady state Ab distribution in human postmortem samples and animal models, little is known about the earliest steps of Abo-mediated synapse toxicity, including the spatial and temporal dynamics of its engagement with the neuronal membrane. Here, we used a combination of live and high-resolution imaging modalities to define the rate, stability, nanoscale localization and functional consequences of Abo binding.
To our knowledge, ours is the first to directly visualize Abo longitudinally as it associates with neurons. This analysis revealed that Abo generally nucleates at specific sites and attracts additional Abo with cluster growth saturating within minutes rather than binding as discrete, preformed assemblies. Growth could occur through recruitment of additional Abo from soluble pools and/or by coalescence of laterally diffusing Abo/receptor complexes on the cell surface (Renner et al., 2010). In either case, the rapid surface assembly and synaptic association we observe is consistent with numerous observations that Abo can rapidly (within minutes) affect synaptic function (Shankar et al., 2008;Laurén et al., 2009;Um et al., 2012;Freund et al., 2016;Cook et al., 2019;Gulisano et al., 2019). It is important to note that we observed little Abo internalization over this timescale, consistent with action through a signaling surface receptor(s) rather than direct Abo-mediated interference of intracellular plasticityrelated processes. However, our data do not rule out an intracellular role for Abo in pathologies that manifest over longer timescales, such as synapse elimination or cell death (Takahashi et al., 2002(Takahashi et al., , 2004. Indeed, numerous studies have demonstrated accumulated intracellular pools of Ab at synaptic sites in animal models and human AD samples where Ab is constitutively produced for months or years (Koffie et al., 2009;Pickett et al., 2016).
While we focused on Abo binding at synaptic sites, it should be noted that Abo accumulates at synaptic and nonsynaptic sites with indistinguishable kinetics, suggesting widespread and heterogeneous distribution of Abo receptor(s). The rate of Abo binding was similar at spines and nonsynaptic sites but surface Abo clusters were consistently more intense near excitatory synapses suggesting some degree of receptor enrichment at synaptic sites. While intriguing, the functional significance of synaptic Abo binding has remained unclear. We demonstrate for the first time that Abo-bound spines are more susceptible to plasticity disruption than neighboring Abo-free spines, supporting a model where Abo engages locally restricted signaling mechanisms to impair plasticity. Our experiments were performed over a relatively short timescale, with application of Abo ;30 min before plasticity induction. It remains possible that longer, disease relevant exposure times would lead to more global plasticity disruption. It will also be important to investigate whether other aspects of Abo-mediated pathology, such as synapse loss, occur selectively at sites where Abo initially engages the neuronal surface. In any case, these results emphasize the importance of future experiments unraveling molecular features that shield select synapses from Abo binding and subsequent plasticity deficits.
Our experiments are also the first to map the nanoscale distribution of Abo surface engagement. The imaging techniques used in earlier studies lacked the spatial resolution to precisely map Abo binding sites (Lacor et al., 2004;Koffie et al., 2009;Renner et al., 2010;Um et al., 2012). Surprisingly, we observed very little Abo actually binds directly at the postsynaptic density. Instead, Abo forms nanoscale clusters immediately adjacent to and surrounding the excitatory synaptic cleft. Several reported Abo receptors localize to perisynaptic regions, including a7 nicotinic acetylcholine receptor (a7-nAchR), cellular prion protein (PrP c ), and metabotropic glutamate receptor 5 (mGluR5; Luján et al., 1996;Mironov et al., 2003;Jones and Wonnacott, 2004). Perisynaptic localization also suggests Abo is unlikely to exert its effects through direct binding of synaptic neurotransmitter receptors such as NMDA or AMPA-type glutamate receptors as previously proposed (De Felice et al., 2007;Lacor et al., 2007;Zhao et al., 2010;Texidó et al., 2011). While earlier studies concluded Abo binds primarily to postsynaptic sites on dendritic spines, these experiments primarily relied on diffraction limited imaging techniques. Using multiple approaches, we confirmed binding near the postsynaptic membrane but also observed a substantial fraction of Abo binding to axonal terminals, consistent with rapid Abomediated effects on presynaptic vesicle release probability, glutamate reuptake and structural alterations (Abramov et al., 2009;Huang et al., 2013;He et al., 2019). Whether the same receptor mediates Abo binding to both presynaptic and postsynaptic compartments is unknown, but several reported Abo receptors localize to both sides of the synapse, including a7-nAchR and PrP c (Moya et al., 2000;Fabian-Fine et al., 2001;Barmada et al., 2004;Um et al., 2012Um et al., , 2013. Taken together, our study is the first to interrogate the kinetics, stability, ultrastructural localization and functional consequences of Abo binding. These basic, yet fundamental assessments provide new insight into the earliest steps of Abo toxicity and lay the groundwork for future studies evaluating the relevant receptor(s) responsible for neuronal surface engagement and the local signaling mechanisms leading to synapse dysfunction.