Amyloid-β oligomers increase the binding and internalization of tau oligomers in human synapses

Shrinath Kadamangudi; Michela Marcatti; Wen-Ru Zhang; Anna Fracassi; Rakez Kayed; Agenor Limon; Giulio Taglialatela

doi:10.1007/s00401-024-02839-2

. 2024 Dec 17;149(1):2. doi: 10.1007/s00401-024-02839-2

Amyloid-β oligomers increase the binding and internalization of tau oligomers in human synapses

Shrinath Kadamangudi ¹, Michela Marcatti ¹, Wen-Ru Zhang ¹, Anna Fracassi ¹, Rakez Kayed ¹, Agenor Limon ¹, Giulio Taglialatela ^1,^✉

PMCID: PMC11652575 PMID: 39688618

Abstract

In Alzheimer’s disease (AD), the propagation and spreading of CNS tau pathology closely correlates with cognitive decline, positioning tau as an attractive therapeutic target. Amyloid beta (Aβ) has been strongly implicated in driving tau spread, whereas primary tauopathies such as primary age-related tauopathy (PART)—which lack Aβ pathology—exhibit limited tau spread and minimal-to-no cognitive decline. Emerging evidence converges on a trans-synaptic mechanism of tau spread, facilitated by the transfer of misfolded tau aggregates (e.g. soluble oligomers). However, it is unclear whether Aβ oligomers modulate the binding and internalization of tau oligomers in human synapses. Our translationally focused paradigms utilize post-mortem brain specimens from Control, PART, and AD patients. Synaptosomes isolated from the temporal cortex of all three groups were incubated with preformed recombinant tauO (rtauO), ± preformed recombinant AβO (rAβO), and oligomer binding/internalization was quantified via flow cytometry following proteinase K (PK) digestion of surface-bound oligomers. TauO-synapse interactions were visualized using EM immunogold. Brain-derived tau oligomers (BDTO) from AD and PART PBS-soluble hippocampal fractions were co-immunoprecipitated and analyzed via mass spectrometry to compare synaptic tauO interactomes in primary and secondary tauopathies, thereby inferring the role of Aβ. AD synaptosomes, enriched in endogenous Aβ pathology, exhibited increased rtauO internalization compared to PART synaptosomes. This observation was mirrored in Control synaptosomes, where recombinant rAβO significantly increased rtauO binding and internalization. PK pre-treatment abolished this effect, implicating synaptic membrane proteins in AβO-mediated tauO internalization. While both PART and AD BDTO were broadly enriched in synaptic proteins, AD BDTO exhibited differential enrichment of endocytic proteins across pre- and post-synaptic compartments, whereas PART BDTO showed no significant synaptic enrichment. This study demonstrates that Aβ oligomers enhance tau oligomer binding and drive its internalization through synaptic membrane proteins. These findings offer novel mechanistic insights underlying pathological tau spreading directly within human synapses and emphasize the therapeutic potential of targeting Aβ-tau interactions.

Supplementary Information

The online version contains supplementary material available at 10.1007/s00401-024-02839-2.

Keywords: Trans-synaptic tau spreading, Tau oligomers, Aβ oligomers

Introduction

Alzheimer’s disease (AD) is the most common and devastating progressive form of neurodegenerative dementia, characterized by the pathological accumulation of amyloid-β (Aβ) plaques and neurofibrillary tau tangles (NFT) within the CNS [39, 110, 126]. Although Aβ pathology is implicated in AD pathogenesis, extensive neuropathological and radiological investigations have found that cognitive decline correlates best with NFT burden [58, 103, 107, 127]. Composed primarily of the microtubule-associated protein tau, NFTs originate in medial temporal lobe (MTL) structures (i.e., hippocampus and entorhinal cortex) and progressively infiltrate functionally connected neocortical regions of the temporal, parietal, and frontal lobes [17, 19, 20, 28, 145]. Beyond the mere formation and accumulation of tau pathology within MTL structures, amnestic changes result from this spatiotemporal progression (spreading) of pathological tau [77, 103, 143]; this is best evidenced by contrasting the pathology of late-stage AD to primary age-related tauopathy (PART). Neuropathological evaluations of PART report pathological tau (hyperphosphorylated tau and NFT) accumulation restricted to the hippocampus and entorhinal cortex, avoiding neocortical infiltration (as seen in late-stage AD) [35, 37, 64]. PART pathology is almost universally detectable among elderly individuals (> 75 years) and is thought to represent typified age-related neuropathological changes [36, 37, 87, 151, 155]; more importantly, patients with PART exhibit little-to-no amnestic changes that extend beyond age-related cognitive decline [10, 11, 27, 138]. Mitigating the regional spread of pathological tau within the canonical ‘Braak staging’ framework, as observed in PART, may thus serve as a rational and clinically effective therapeutic strategy to combat AD.

PART is further contrasted from AD by the absence of Aβ plaque pathology [36, 37, 64]. This is of particular significance after multiple, independent lines of evidence, both in humans [1, 28, 82, 93, 144] and animal models [61, 112], support the notion that Aβ promotes pathological tau propagation and regional spreading within the CNS. Of note, isolated evaluation of Aβ burden demonstrates minimal association with specific domains of cognitive function, further highlighting the importance of targeting Aβ and tau’s “toxic pas de deux” [14, 23, 58, 61, 127]. In summary, while Aβ plays a central role in AD pathogenesis, it appears that the interaction with tau, particularly in facilitating its spread, drives cognitive decline [28, 55, 139]. However, the precise molecular mechanisms underlying this interaction remain unresolved.

Recent studies offer strong evidence for tau spreading through a trans-synaptic, ‘prion-like’ mechanism, driven by proteopathic ‘seeds’ [5, 29, 101, 136, 145]. Further research suggests that soluble forms of amyloid oligomers are among the seeds with the highest potential for propagation [48, 50, 79, 79, 80, 128]. Notably, oligomeric forms of Aβ (AβO) and tau (tauO) have been consistently associated with synaptic dysfunction underlying dementia in AD, where disease-specific tauO conformers have been further shown to drive clinical presentation [48, 57, 79, 114]. In this paradigm, proteopathic tauO seeds are transferred between diseased and healthy neurons, resulting in templated aggregation of endogenous tau at the synaptic terminal and subsequent propagation of pathological tau beyond the somato-dendritic compartment [49, 60, 62, 85, 114]. Conventionally regarded as an intracellular protein, several secretion pathways have been implicated in the release of misfolded tau [22]. While a fraction of extracellular tau has been observed in exosomes or ectosomes, the majority (~ 90%) exists in a vesicle-free form [22, 70]. At the synaptic interface, tau secretion has been observed under physiological conditions [111], with evidence of truncated tau fragments released from AD cortical synapses [69]. Interestingly, Aβ-induced alterations in synaptic activity have been suggested to increase tau release into the extracellular space [93, 112].

Building on this framework, our recent findings suggest that AβO broadly facilitates the engagement of tauO within murine and human synaptosomes [89]. However, the precise role of AβO in selectively modulating tauO synaptic binding and internalization remains unclear. We, thus, hypothesize that AβO promotes tauO binding and internalization in human synapses—a critical process underpinning pathological tau spreading. To test this, we established two key objectives: (1) to assess tauO binding and internalization in synaptosomes with pre-existing amyloid pathology (AD vs. PART), and (2) to investigate how AβO dynamically inform tauO engagement in healthy Control synaptosomes. To address these objectives, we developed an experimental platform using human synaptosomes derived from post-mortem brain specimens, enabling the study of oligomer dynamics in a system directly relevant to human neurodegenerative pathology. Moreover, this platform is uniquely suited to assess molecular mechanisms directly in human tauopathies, namely AD and PART, providing crucial insights into how Aβ pathology may drive selective vulnerability in primary and secondary tauopathies.

Materials and methods

Clinical and neuropathological assessment of donor brain tissue

Donor subjects underwent comprehensive annual neurological and neuropsychological evaluations, including the Clinical Dementia Rating (CDR), administered by experienced clinicians. Neuropathological evaluations were performed by expert neuropathologists who assessed the presence of Aβ plaques and neurofibrillary tangles using the standardized Consortium to Establish a Registry for Alzheimer’s Disease (CERAD) criteria and Braak staging. Control subjects (CDR = 0) received regular neurological assessments and did not meet clinical or pathological criteria for AD. AD subjects were diagnosed through clinical consensus, met the NINDS-Alzheimer’s Disease and Related Disorders Association criteria, had a CDR > 1.0, and were confirmed neuropathologically at autopsy (plaques = 1–2; Braak stage = 5–6). PART cases, defined by mild-to-moderate neurofibrillary degeneration in the hippocampus without accompanying Aβ plaques [35, 37, 64], were identified through neuropathological assessments.

All samples were de-identified prior to arrival at the University of Texas Medical Branch (UTMB), eliminating the need for additional IRB approval under CFR §46.101(a)(1). Detailed clinical (diagnosis, sex, age at death, last mini-mental status exam [MMSE] score before death) and neuropathological (postmortem interval [PMI], brain region, Braak stage) data for the cases used in this study are summarized in Supp. Table 1.

Standardized isolation and preparation of synaptosomes

Synaptosomes were prepared from individual fresh frozen superior medial temporal gyri (SMTG) autopsy brain specimens obtained from Control (n = 8; 4M, 4F) subjects, as well as PART (n = 7; 2M, 5F), and AD (n = 8; 5M, 3F) patients. The synaptosomes, or isolated nerve terminals, were prepared using a published protocol [44, 45, 89, 90, 121]. We have standardized this protocol in our lab to ensure consistent quality and yield across different preparations. To preserve tissue microstructure and cytoarchitecture, synaptosomes were only isolated from tissues with a PMI of 4 h or less. Briefly, 20 mg of snap frozen brain tissue were homogenized in 400 µL of SynPER lysis buffer (cat# 87793; ThermoFisher Scientific) supplemented with 1% protease and phosphatase inhibitor cocktails. Homogenates were centrifuged at 1200×g for 10 min at 4 °C to clear debris. The resulting supernatants, containing synaptosomes, were further centrifuged at 15,000×g for 20 min at 4 °C to pellet the synaptosomes.

The synaptosome pellets were resuspended in 100 µL of HEPES-buffered Krebs-like solution (HBK; 143.3 mM NaCl, 4.75 mM KCl, 1.2 mM MgSO₄·7H₂O, 1.2 mM CaCl₂, 20.1 mM HEPES, 0.1 mM NaH₂PO₄, and 10.3 mM D-glucose, adjusted to pH 7.4). Notably, HBK was used within 1 week of preparation to preserve its chemical stability and minimize contamination risks. To prevent synaptosome aggregation, 0.5% Pluronic F-68 non-ionic surfactant (cat# 24040-032; ThermoFisher Scientific) was added. Synaptosome concentrations were quantified using the Guava EasyCyte 8 flow cytometer (EMD Millipore) by calculating the total number of events within the pre-established synaptosome size gate (Supp. Figure 6). Concentrations were adjusted to a final density of 1 million synaptosomes per microliter.

Preparation and fluorescent labelling of preformed recombinant tauO

Preformed recombinant tauO (rtauO) were generously supplied by Dr. Rakez Kayed’s laboratory; they were produced and characterized according to established protocols [78, 114, 124]. RtauO were fluorescently labeled with Alexa Fluor™ 488 (rtauO-A488) following a previously published procedure [89, 98, 114]. In brief, 1 mg of Alexa Fluor™ 488 NHS Ester Succinimidyl Ester (Thermo Fisher Scientific, cat# A20000) was dissolved in 0.1 M sodium bicarbonate buffer to yield a 1 mg/mL dye solution, adjusted to pH 8.3. TauO were incubated with the dye solution at a 1:4 (w/w) ratio overnight at 4 °C on an orbital shaker. Unbound dye was removed through centrifugation (30 min at 15,000×g) using 10-kDa Amicon Ultra-0.5 mL centrifugal filter units (EMD Millipore, cat# UFC501024). To ensure thorough purification, labeled rtauO were washed with 1X PBS until the filtrate was visibly clear and were recovered by inverting the filter compartment and centrifuging (2 min at 1000×g). All steps involving rtauO employed low retention plasticware to minimize protein loss. Labeled rtauO concentrations were quantified using the Pierce™ BCA Protein Assay Kit (Thermo Fisher Scientific, cat# 23227) and further validated with the NanoDrop 2000c (Thermo Fisher Scientific, Wilmington, DE, USA) to correct for absorbance shifts introduced by the Alexa fluorophore. Western blot analyses were routinely performed to confirm maintenance of oligomeric conformation. Labeled rtauO were subsequently delivered to synaptosomes for assessment of binding and internalization via flow cytometry.

Preparation of preformed recombinant AβO

Fluorescently conjugated recombinant AβO (rAβO-A647) were prepared following an established protocol [32, 89, 91, 97, 156]. In brief, lyophilized human Aβ1–42 peptide was sourced from the Department of Biophysics and Biochemistry at Harvard University (Cambridge, MA, USA). 0.3 mg of Aβ1–42 and 7 μL of Aβ monomer tagged with HyLite Fluor 647 (cat# AS-64161; AnaSpec Inc., Fremont, CA, USA) were dissolved together in 200 μl of 1,1,1,3,3,3-hexafluoro-2-propanol (HFP) and incubated at RT for 10–20 min. Following incubation, 700 μl of ddH₂O was added to the mixture and stirred magnetically for 48 h at RT in a fume hood, allowing the HFP to evaporate through a vented cap. Resulting rAβO were aliquoted, frozen at − 80 °C, and used within three months. Western blot analysis was routinely utilized to confirm maintenance of oligomeric conformation.

Electron microscopy (EM)

Transmission EM for morphology quality control

The quality and cytoarchitecture of synaptosome preparations were evaluated through transmission electron microscopy (TEM), following a standard protocol established previously in our lab [90]. In brief, pooled synaptosome preparations were concentrated via centrifugation (10,000g for 10 min at 4 °C) to form a visible pellet approximately 50 μL in size. Synaptosome pellets were fixed overnight at 4 °C in a solution of 2.5% formaldehyde and 0.1% glutaraldehyde in 0.05 M cacodylate buffer. Post-fixation, the pellets were rinsed three times with 0.1 M cacodylate buffer and subsequently post-fixed in 1% osmium tetroxide (OsO₄) in 0.1 M cacodylate buffer (pH 7.4). The samples were then en bloc stained with 2% aqueous uranyl acetate in 0.1 M maleate buffer and progressively dehydrated in a graded ethanol series. Dehydrated pellets were infiltrated with increasing concentrations of propylene oxide and Poly/Bed 812 epoxy resin (Polysciences, Warrington, PA, USA), before being embedded in the same resin, which was polymerized at 60 °C overnight. Ultrathin sections were then prepared using a Leica EM UC7 ultramicrotome (Leica Microsystems, Buffalo Grove, IL, USA), stained with 0.4% lead citrate, and examined using a JEM-1400 electron microscope (JEOL USA) at 80 kV. Images were acquired on bottom-mounted Orius SC-200 camera (Gatan, Pleasanton, CA).

EM immunogold staining of ultrathin sections for qualitative validation of synaptic rtauO engagement

EM immunogold labeling was performed for qualitative validation of synaptic rtauO engagement. Synaptosomes, isolated from frozen temporal cortex specimens of C57BL/6 wild-type (WT) mice (n = 4; 2M, 2F; 8–9 months of age; Mus musculus-cat# JAX:000664), were pooled and treated with 2.5 μM rtauO (derived from full-length (2N, 4R) human tau) and thoroughly washed (protocol detailed in Synaptosome binding and internalization assay for Aβ and tau oligomers section). Of note, WT mouse synaptosomes were specifically chosen to minimize primary antibody cross-reactivity (Tau13; BioLegend, cat# 835201) with endogenous human tau and to take advantage of the comparable synaptic binding responses between murine and human synaptosomes, as previously demonstrated by our group [89]. RtauO were detected using the Tau13 total tau antibody, selected for its high specificity for human tau over murine tau [83]. Of note, animal housing conditions adhered to USDA standards, with a 12:12 light/dark cycle and unrestricted access to food and water, and animal care was provided daily by specialists under the supervision of the UTMB vivarium manager. Animals were sacrificed in accordance with protocols approved by UTMB’s Institutional Animal Care and Use Committee, ensuring minimal pain or discomfort.

RtauO-treated mouse synaptosomes were concentrated via centrifugation (10,000 g for 10 min at 4 °C) to form a visible pellet approximately 50 μL in size and were fixed in 2% glutaraldehyde overnight at 4 °C. Briefly, the pellets were stained en bloc with 2% aqueous uranyl acetate, dehydrated in 50% then 75% ethanol and embedded in LR White resin medium grade (Electron microscopy Science EMS, Cat# 14381). Ultrathin sections (70–80 nm) were cut on Leica EM UC7 ultramicrotome and collected onto Formvar-carbon coated nickel grids (EMS, Hatfield, PA). For immunogold labeling, grids were incubated in a wet chamber on drops of blocking buffer BSA-G (0.1% BSA and 0.01 M glycine in 0.01 M PBS) for 15 min. After blocking, grids were incubated with primary antibody (Tau13; BioLegend, cat# 835201), diluted 1:250 in 1% BSA in 0.01 M PBS, for 1 h at room temperature (RT), followed by an overnight incubation at 4 °C in a wet chamber. After washing in blocking buffer for five times, grids were incubated with a gold-conjugated (6 nm) secondary antibody (anti-Mouse IgG-H&L, cat# 25123; Electron Microscopy Sciences), diluted 1:20 in 1% BSA, for 1 h at RT, in the dark, within a moist chamber. After washing five times with BSA-G, then three times with PBS and distilled water, grids were fixed in 2% aqueous glutaraldehyde for 5 min at RT. After fixation, grids were washed three times in ddH₂O for 5 min each. The sections were then stained with 2% aqueous uranyl acetate for 5 min at RT, washed three times in ddH₂O for 5 min each, and air-dried. Finally, the sections were stained with lead citrate for 30 s and examined in a JEM-1400 electron microscope (JEOL USA) at 80 kV. Images were acquired on bottom-mounted Orius SC-200 camera (Gatan, Pleasanton, CA).

Western blot

The synaptosomes isolated from individual AD (n = 8) and PART (n = 7) autopsy specimens were analyzed for Aβ and tau oligomers via conventional western blotting techniques established previously by our research group [12, 89, 114]. Key steps for the detection of each target are outlined below.

For evaluation of tau pathology, the synaptosome samples were prepared in 1X NuPAGE™ LDS Sample Buffer (ThermoFisher, cat# NP0007) and 1X NuPAGE™ Sample Reducing Agent (ThermoFisher, cat# NP0004), followed by denaturation at 75 °C for 10 min. For protein separation, 5 million synaptosomes per well were loaded onto pre-cast NuPAGE 4–12% Bis–Tris SDS-PAGE gels (ThermoFisher, cat# NP0323BOX) and transferred to nitrocellulose membranes (Cytiva, cat# 10600001) using conventional wet-tank transfer at 100 V for 1 h at 4 °C. Membranes were blocked with Intercept^® (TBS) Blocking Buffer (LI-COR Biosciences, cat# 927-60001) for 1 h at RT. Tau detection was carried out by incubating membranes overnight at 4 °C with the primary antibody Tau13 (1:1000, BioLegend, cat# 835201), a well-established total tau antibody selected for its ability to detect a broad range of tau species, including oligomeric assemblies, monomers, and lower-molecular-weight fragments [1–7]. Following primary antibody incubation, membranes were treated with the secondary antibody IRDye^® 800CW anti-mouse (1:10,000, LI-COR Biosciences, cat# 926-32210) for 1 h at RT. Synaptic protein enrichment was confirmed with probes for PSD-95 (1:1000, cat# ab13552; Abcam) and synaptophysin (1:20,000, cat# ab32127, Abcam). Membranes were washed four times (7 min each) in TBS-T (0.1% Tween-20 in 1X TBS) prior to imaging on the Odyssey LI-COR Classic Infrared imaging system (LI-COR Biosciences).

For evaluation of Aβ pathology, the synaptosomes were processed similarly, with samples prepared using 1X Tricine SDS Sample Buffer (ThermoFisher, cat# NP0007) and 1X NuPAGE™ Sample Reducing Agent (ThermoFisher, cat# NP0004), and denatured at 75 °C for 10 min. Five million synaptosomes per well were loaded onto pre-cast Novex 16% Tricine SDS-PAGE gels (ThermoFisher, cat# EC6695BOX). Proteins were transferred to polyvinylidene fluoride (PVDF) membranes (Millipore Sigma, cat# ISEQ00010) via wet-tank transfer (80 V for 45 min at 4 °C). Membranes were blocked in 3% BSA in TBS-T for 1 h at RT. Primary antibody (4G8, 1:1000, BioLegend, cat# 800710) incubation was performed overnight at 4 °C, followed by secondary antibody (HRP-linked IgG anti-mouse, 1:5000, Cell Signaling, cat# 7076S) incubation for 1 h at RT. Post-incubation washes (four, 7 min each in TBS-T) were followed by exposure to regular (Cytiva, Amersham™ ECL Regular, cat# RPN2209), enhanced (ThermoFisher, Pierce™ ECL Plus, cat# 32132), or super-enhanced (ThermoFisher, SuperSignal™ West Atto, cat# A38554) HRP substrates. The membranes were developed on Amersham hyperfilms (Cytiva, cat# 28906842). The substrate selection and exposure times were optimized for the best signal-to-noise ratio.

Prior to blocking, membranes were stained with Revert™ 700 Total Protein Stain (LI-COR Biosciences, cat# 926-11011) for normalization. Protein abundance was quantified using ImageJ FIJI software (NIH, Bethesda, MD, USA), and all values were normalized to the total protein signal for each blot.

Preparations of preformed rAβO and rtauO were also routinely assessed by western blotting to verify structural conformation. For these preparations, the reducing agent and heating steps were omitted to maintain oligomeric structure. Similarly, brain-derived tau oligomers were evaluated using total tau antibodies Tau5 (1:1000, cat# 806401, BioLegend) and EPR2396 (1:1000, cat# ab109392, Abcam) under non-reducing, non-denaturing conditions.

Synaptosome binding and internalization assay for Aβ and tau oligomers

Synaptic oligomer binding and internalization assays implemented in this study were adapted directly from our recent published work [89]; all experimental conditions were carefully maintained to ensure the accurate extension of prior findings. To minimize sample loss, low-retention pipette tips and microcentrifuge tubes were employed throughout all experiments. All procedures were conducted on ice, with centrifugation and wash steps performed at 10,000 g for 10 min at 4 °C, unless otherwise specified. We have previously demonstrated that the magnitude of engagement for both rAβO and rtauO remains consistent across individual human synaptosome preparations [89]. Therefore, the synaptosome preparations were pooled to increase sample yield and enhance experimental feasibility.

Each binding and internalization assay used 2 million pooled synaptosomes from various pathological conditions and followed four key steps (Supp. Figure 1, left column): (1) incubation with rtauO-A488 and/or rAβO-A647, (2) treatment with proteinase K (PK; cat# 70663-4; EMD Millipore) to distinguish surface-bound from internalized oligomers, (3) washes, and (4) flow cytometry analysis. All dilutions, treatments, and washes were performed using freshly prepared HBK buffer.

Role of AβO in tauO binding and internalization in PART vs. AD synaptosomes

The first set of experiments utilized pooled SMTG synaptosomes from PART (n = 7) and AD (n = 8) patients. Both PART and AD synaptosomes were treated with rtauO-A488 (2.5 µM). The synaptosomes without oligomers were used as gating controls for flow cytometric evaluation. To differentiate surface-bound from internalized oligomers, each sample was divided equally by volume, with one set treated with PK (final concentration: 1 mg/mL) to estimate internalized oligomers, while the other set remained untreated to estimate total oligomer engagement. Both sets were incubated for 30 min at 37 °C. Following oligomer and PK treatment, synaptosomes were washed three times through serial centrifugation (10,000 g for 10 min at 4 °C) and gentle pipetting in 50 μL of HBK buffer. The final wash step involved re-suspension in 50 μL of HBK for flow cytometry analysis, either immediately or stored at 4 °C for up to 24 h. Prior to flow cytometric measurement, 200 μL of DPBS was added and samples were gently vortexed.

PK digestion of synaptic membrane proteins

The role of synaptic membrane proteins in mediating the Aβ-tau interaction was assessed through PK pre-treatment (PK_pre) experiments (Supp. Figure 1, right column). For these experiments, the synaptosomes were incubated with 1 mg/mL of PK for 30 min at 37 °C prior to being challenged with labeled rAβO and rtauO. Following PK pre-treatment, the samples were cooled in an ice-slush for 2 min, washed twice with ice-cold HBK-PK neutralization buffer (HBK buffer with 1 × protease inhibitor cocktail), and subsequently treated with oligomers as described above.

Role of AβO in tauO binding and internalization in control synaptosomes

The second set of experiments utilized pooled synaptosomes from Control subjects (n = 8) who lack prior Aβ and tau pathology. The control synaptosomes were incubated with rtauO-A488 (2.5 μM) alongside rAβO-A647 at concentrations of 0, 1, 2.5, 5, and 10 μM 1 h at RT. The synaptosomes without oligomers were used as gating controls for flow cytometry evaluation. Wash steps, differentiation between surface-bound and internalized rtauO, as well as the role of synaptic membrane proteins in this process, were performed and assessed as described above.

Ancillary experiments

In addition to the primary experiments, four supplementary flow cytometric studies were conducted using pooled Control SMTG synaptosomes (n = 8). First, the specificity of rAβO’s influence on synaptic rtauO engagement was tested by substituting rAβO-A647 with varying concentrations of recombinant rtauO-A647. Second, synaptic tauO engagement was assessed across a lower range of rtauO concentrations (0–1 μM) in the presence and absence of rAβO. Third, a targeted experiment to evaluate the role of low-density lipoprotein receptor-related protein 1 (LRP1) on the synaptic binding and internalization of rtauO was performed by delivering varying concentrations of receptor-associated protein (RAP; cat# 553506-M, Millipore Sigma) [0, 0.5, and 5.0 µM], a high-affinity inhibitor of LRP1, on to synaptosomes treated with either rtauO (2.5 µM) alone or rtauO (2.5 µM) + rAβO (5.0 µM). 5.0 μM rAβO was selected for the latter two experiments as this was the lowest tested concentration at which significantly increased synaptic rtauO binding and internalization was observed. Fourth, a time-course analysis of rtauO binding and internalization was conducted by treating synaptosomes with 2.5 µM rtauO-A488, with aliquots collected at multiple incubation intervals (15 min, 1 h, 4 h, and 24 h at RT). Each aliquot was subsequently treated with PK, washed, and analyzed via flow cytometry.

Calcein AM assay for assessment of synaptosome membrane integrity

Calcein AM labeling was employed to assess the membrane integrity of pooled SMTG synaptosome preparations from PART (n = 7) and AD (n = 8) post-mortem brain specimens. The synaptosomes were incubated with 100 nM Calcein AM (ThermoFisher, cat# C3100MP) at RT for 30 min, allowing the dye to passively diffuse into the synaptosomes. Within intact synaptosomes, intracellular esterases cleave the Calcein AM, yielding a fluorescent calcein signal indicative of preserved membrane integrity. For experimental validation, a subset of synaptosomes were pre-treated with 2% Tween-20 for effective membrane permeabilization. Calcein-positive events were quantified via flow cytometry, following the same protocol as used for binding and internalization assays.

Synaptosome immunophenotyping

Synaptosome immunophenotyping was performed to evaluate the efficacy of PK-mediated cleavage/digestion of synaptic membrane proteins. Immunophenotyping for flow cytometry detection was performed using pooled Control synaptosomes. Low-retention pipette tips and microcentrifuge tubes minimized sample loss, and all procedures were conducted on ice with centrifugation and wash steps performed at 10,000g for 5 min at 4 °C unless specified otherwise.

The following buffers were prepared: HBK (recipe outlined in Standardized isolation and preparation of synaptosomes section); Sucrose/EDTA/Tris (SET) buffer (320 mM Sucrose, 1 mM EDTA, 5 mM Tris base, pH 7.4); CellTrace neutralization buffer (1% Bovine Serum Albumin (BSA) in HBK); SET-PK neutralization buffer (SET buffer with 1× protease inhibitor cocktail); fixation buffer (4% Paraformaldehyde (PFA) in PBS); blocking-only buffer (5% BSA in SET); blocking + permeabilization buffer (5% BSA with 0.1% Tween-20 in SET); and antibody diluent (0.5% BSA with 0.1% Tween-20 in SET).

The synaptosome aliquots were rapidly thawed at 37 °C for no more than 2 min to prevent aggregation. For CellTrace labeling, 1 M synaptosomes were treated with 5.0 μM CellTrace in 100 μL of HBK buffer (final synaptosome concentration: 10,000/μL) and incubated for 20 min at 37 °C, protected from light. Following incubation, CellTrace neutralization buffer was added at 5× the original staining volume to remove excess dye. The synaptosomes were centrifuged, resuspended in SET buffer, and allowed to rest for 10 min at RT for acetate hydrolysis.

If samples were designated for PK_pre treatment, synaptosomes were pelleted, resuspended in PK (100 μL), and incubated for 30 min at 37 °C. After treatment, samples were cooled in an ice-slush for 2 min and washed twice with ice-cold SET-PK neutralization buffer. Samples were then resuspended in 100 μL of SET buffer. Samples not undergoing PK pre-treatment proceeded directly to fixation.

For fixation, 100 μL of fixation buffer was added to yield a 200 μL solution of 2% PFA, and synaptosomes were fixed for 15 min at RT. Post-fixation, synaptosomes were washed with 100 μL blocking-only buffer to neutralize free PFA. The pellet was then resuspended in 100 μL of blocking + permeabilization buffer and incubated for 30 min at RT, protected from light. During this incubation period, primary and secondary antibody solutions were prepared in antibody diluent at their respective working concentrations.

To assess the selective vulnerability of synaptic membrane proteins versus cytosolic proteins to PK cleavage, protein density was examined for two membrane proteins, GluA1 [1:400 or 2.5 μg/mL] (cat# ABN241, EMD Millipore) and NR2B [1:200 or 2.75 μg/mL] (cat# 21920-1-AP, ThermoFisher Scientific), as well as two cytosolic proteins, PSD95 directly conjugated to Alexa750 [1:200 or 8.65 μg/mL] (cat# NB300-556AF750, Novus Biologicals) and GAD1 directly conjugated to phycoerythrin (PE) [1:100 or 3.3 μg/mL] (cat# NBP2-79937PE, Novus Biologicals). After blocking and permeabilization, samples were incubated with the primary antibody solution (100 μL) for 30 min at room temperature, protected from light. Following primary antibody incubation, samples were washed twice with SET buffer and then incubated with the secondary antibody solution (100 μL) for 30 min at room temperature, also protected from light. Detection of GluA1 and NR2B was performed using an anti-rabbit IgG secondary antibody conjugated to Alexa647 [1:10,000] (cat# ab150079, Abcam). Finally, samples were washed twice with SET buffer, resuspended in SET buffer, and either analyzed immediately via flow cytometry or stored at 4 °C for up to 24 h. Prior to flow cytometric analysis, 200 μL of DPBS was added, and samples were gently vortexed.

Flow cytometric (FC) evaluation of synaptosomes

Instrumentation

Synaptic oligomer binding and internalization was evaluated by the Guava EasyCyte 8 flow cytometer with GuavaSoft InCyte v2.7 software (EMD Millipore, Burlington, MA). The Blue and Red lasers in the Guava EasyCyte 8 were configured at 488 nm with a power of 150 mW and 642 nm at 100 mW respectively. Each laser is equipped with dedicated detectors. Synaptosomes were measured through side scatter (SSC) triggering (Blue laser detector; bandpass filter: 488/16). TauO-Alexa488 associated synaptosomes were detected in the Green channel (Blue laser detector; bandpass filter: 525/30), while tauO-Alexa647 associated synaptosomes were detected in the Red2 channel (Red laser detector; bandpass filter: 661/15). The gains for SSC, Green, and Red2 channels were set to “High”. To minimize particle coincidence or swarm effects, the synaptosomes were acquired at a flow rate of 0.5 µL/s using the “Low” setting. A total of 10,000 events were acquired for each sample. Due to minimal spectral overlap between Alexa488 and Alexa647, no compensation was applied. Data acquisition was performed over a 5-decade acquisition range.

The BDFACSymphony A5 SE flow cytometer (BD Biosciences, San Jose, CA) was selected for synaptosome immunophenotyping due to its capability to detect a broad range of fluorophores within a single sample. This expanded optical configuration facilitates the simultaneous detection and quantification of multiple fluorophores, which is essential for accurate synaptic protein profiling. The instrument's lasers were configured as follows: 355 nm (UV) at 60 mW, 405 nm (Violet) at 200 mW, 488 nm (Blue) at 150 mW, 561 nm (Yellow) at 150 mW, and 640 nm (Red) at 140 mW, each equipped with a dedicated set of detectors. Synaptosome acquisition was carried out using instrument software (BDFACS Diva Software 9.0, BD Biosciences) with dual thresholding via SSC (SSC detector; bandpass filter: 488/10) and fluorescence (UV446 detector; bandpass filter: 446.5/67). The CellTrace Blue Cell Proliferation Kit (cat# C34568, ThermoFisher Scientific) facilitated fluorescence triggering, providing a consistent signal in structurally intact synaptosomes. The detector settings for the fluorophores used were: Alexa488 (B537 detector; bandpass filter: 537/32), Alexa647 (R675 detector; bandpass filter: 675/20), Alexa750 (R780 detector; bandpass filter: 780/60), and PE (YG585; bandpass filter: 585/30). Compensation was performed using BDFACS Diva software in accordance with standard guidelines [59]. Single-color controls were established using compensation beads (cat# 01-2222-42, ThermoFisher Scientific), and the compensation matrix was calculated for each sample. Compensation was applied at the start of every flow cytometry run.

Daily calibration of the Guava EasyCyte 8 flow cytometer involves running the Guava® easyCheck™ Kit (cat# SKU 4500-0025, Cytek), which is designed to assess both particle counting accuracy and fluorescence detection using standardized fluorescent bead reagents. To ensure consistent forward scatter (FSC) positioning across experiments, bead standards were run daily to calibrate the size gates (Supp. Figure 5B). For smaller particles, the ApogeeMix "Micro" (cat# 1493, Apogee Flow Systems) was used, comprising a mixture of non-fluorescent silica and fluorescent polystyrene beads, with sizes ranging from 80 nm to 1.3 µm. For larger particles, non-fluorescent polystyrene bead standards ranging from 2.0 to 7.56 µm (Spherotech Inc.) were employed. The positions of these bead populations on FSC versus SSC plots were carefully maintained within predefined gates stored in the QC workspace, with only minor adjustments made as necessary.

Acquisition parameters for small particle measurement

Flow cytometry provides significant advantages for the study of submicron particles such as synaptosomes, though technical challenges, including instrument sensitivity and ensuring linear range of detection, must be carefully managed to ensure data reliability [106]. Instrument sensitivity for both the Guava EasyCyte 8 and BD FACSymphony was assessed using standardized beads (ApogeeMix “Micro” and Spherotech polystyrene (PS) beads) (Supp. Figure 6B). This calibration informed the establishment of a size gate (0.5–3.3 µm) based on the estimated size of synaptosomes [59, 89, 134]. Size gating was performed using density plots of FSC versus SSC, followed by hierarchical gating of fluorescent channels on bivariate density plots of SSC versus the corresponding fluorophore (e.g., Alexa488, Alexa647) (Supp. Figure 5C-D). Although the typical size of single synaptosomes ranges from 0.5 to 1.5 µm [56, 59, 134], we included signals from larger particles, such as synaptosome doublets and triplets, to enhance signal detection, as isolating single synaptosomes was not critical for assessing synaptic binding and internalization of oligomers. On the Guava EasyCyte 8, the SSC gain was adjusted to place 500-nm PS beads just above the center on the SSC axis, ensuring detection above the SSC noise threshold while keeping 800-nm PS and 1300-nm silica beads within scale (Supp. Figure 5A-B).

Optimization of the linear range of detection was specifically focused on the Guava EasyCyte 8, the primary instrument used for our binding and internalization assays. Serial dilutions of ApogeeMix “Micro” beads, which have known proportions of constituent beads, were analyzed to identify the dilution that provided (1) the best discrimination of individual bead populations, as well as (2) the most accurate and precise proportion of fluorescent beads ( $f l u o r e s c e n t / (f l u o r e s c e n t + n o n f l u o r e s c e n t)$ ) within the designated size gate. Additionally, we prepared pooled Control synaptosome samples at varying concentrations and performed serial reading dilutions in HBK buffer to assess particle event rates (particles/s) within our synaptosome size gate. Linear regression analysis was utilized to determine the linear range of detection, which informed the optimal conditions for minimizing coincidence or ‘swarm’ detection. Lastly, we implemented additional measures to minimize coincidence detection and particle aggregation, as recommended by recent studies [59, 148]. These included reducing spin times, maintaining optimal sample dilution, and utilizing non-ionic buffers when feasible.

FC data analysis

In this study, we aimed to assess the density of synaptic oligomer engagement within individual synaptosomes across various biological contexts. To achieve this, median fluorescence intensity (MFI) was selected as the preferred flow cytometric parameter. MFI effectively captures the entire spectrum of fluorescence signals, facilitating the precise detection of changes in oligomer engagement. This approach minimizes artifacts arising from population heterogeneity, ensuring that even subtle shifts in binding density are accurately quantified—a critical factor when analyzing small particles such as synaptosomes [149]. All flow cytometry data analysis, including gating, quantification, and generation of density plots and histograms, was performed using FlowJo v10 (BD Life Sciences, Ashland, OR). Key metrics such as the number of events and channel statistics (e.g., MFI) were exported via the “Batch Export” function for further statistical analysis.

The independent samples were analyzed in technical duplicate or triplicate within a given experimental day, with experiments repeated across multiple days to ensure consistency. Data interpretation varied across different experimental paradigms, requiring distinct normalization strategies. However, within each paradigm, all data were normalized in the same manner. Specific normalization methods and statistical analyses for each paradigm are outlined in corresponding results sections and figure captions.

Isolation of brain-derived tau oligomers

Preparation of PBS-soluble brain homogenates

Brain-derived tau oligomers (BDTO) were prepared, following an established protocol [49, 114, 124], from fresh frozen hippocampal autopsy brain specimens of AD patients (n = 4; 2M, 2F) and PART patients (n = 4; 1M, 3F). Although less critical for AD pathology, isolating BDTOs from the hippocampus aligns with the primary site of tau aggregation in PART [35, 37, 64] and ensured sufficient yield for subsequent analyses. Low retention plasticware, including tips and tubes, was used to minimize sample loss. All steps for BDTO isolation were performed on ice or at 4 °C unless otherwise specified.

Fresh frozen hippocampal specimen (75 mg) from each patient or individual was gently homogenized in a Dounce homogenizer (10–15 gentle strokes until no visible tissue chunks) in ice-cold Dulbecco’s phosphate-buffered saline (DPBS; 2.68 mmol KCl, 1.47 mmol KH₂PO₄, 136.89 mmol NaCl, 8.10 mmol Na₂HPO₄ (anhydrous); Corning, cat. 21-031-CV), supplemented with protease (Roche, cat. 11836153001) and phosphatase inhibitors (Roche, cat. 490683700), at a 1:5 ratio (w/v) of brain:DPBS + protease + phosphatase buffer (henceforward referred to as PBS lysis buffer). Brain homogenates were transferred to a 1.5 mL microcentrifuge tube and centrifuged at 9168×g for 10 min at 4 °C. The supernatant (i.e., PBS-soluble homogenate) was carefully removed and either immediately used for subsequent co-immunoprecipitation or stored at − 80 °C.

Co-immunoprecipitation

BDTO, alongside their interacting proteins, were isolated via co-immunoprecipitation (co-IP) using the Pierce™ Co-Immunoprecipitation Kit (ThermoFisher, cat. 26149). The primary modification was substitution the of IP lysis/wash buffer with 1X PBS lysis buffer to minimize protein complex disruptions by detergents, thereby preserving the integrity of the isolated interactomes. Briefly, spin columns were prepared by coupling 75.0 µg of T18 (toxic tau conformation 18) antibody with the AminoLink Plus Coupling Resin, as directed by the manufacturer’s protocol. Of note, T18 is an in-house antibody developed by Dr. Rakez Kayed’s group, initially characterized by LoCasio et al. (2020) for targeting disease-relevant tau oligomers [86]. Since its development, T18 has been effectively used to detect and immunopurify pathogenic tau aggregates from human autopsy specimens and transgenic AD models [113, 115]. Furthermore, T18 is part of a standardized antibody panel designed specifically for tau oligomer preparation from brain lysates, underscoring its specificity and reliability in detecting misfolded tau species [124]. Spin columns for isotype controls were prepared in parallel using rabbit IgG (Millipore; cat. 12-370). Prior to resin quenching, the antibody coupling efficiency for each column was verified by measuring IgG A280 (Nanodrop 2000c; ThermoFisher Scientific, Wilmington, DE, USA).

The resin-antibody mixtures were equilibrated with PBS lysis buffer and incubated with equal amounts of individual brain homogenates overnight at 4 °C. Following incubation, the resin-antibody-homogenate mixtures were centrifuged, and the flowthroughs were collected and stored at -80⁰C for subsequent analyses. Spin columns were then washed three times with PBS to reduce non-specific interactors. BDTO were eluted twice in 50 µL of elution buffer and immediately neutralized with 1.0 M Tris (pH 8.0), as per protocol recommendations. Post-elution, BDTO samples were placed on ice for 30 min to promote meta-stability of BDTO conformers. Resin-antibody mixtures were regenerated by washing twice and stored in diluted coupling buffer at 4⁰C.

Liquid-chromatography with tandem mass spectrometry (LC–MS/MS)

Sample digestion

The samples were prepared for LC–MS/MS analysis as outlined previously [6]. Beads were rinsed twice with 50 μL of 50 mM TEAB, pH 7.1. Proteins were eluted by adding 50 μL of 5% SDS, 50 mM TEAB, pH 7.1 and incubating at 37 °C for 30 min. The sample was reduced in a solution of 10 mM TCEP (cat# 77720, ThermoFisher Scientific) and incubated at 65 °C for 30 min. The sample was then cooled to RT, alkylated with 1 μL of 500 mM iodoacetamide, and allowed to react for 30 min at RT in the dark. 2.7 μL of 12% phosphoric acid was added to the 54.7 μL protein solution. 165uL of binding buffer (90% Methanol, 100 mM TEAB pH 8.5) was then added to the solution. The resulting solution was added to S-Trap spin column (protifi.com) and passed through the column using a bench top centrifuge (60 s spin at 1000g). The spin column washed with 150uL of binding buffer, centrifuged, and repeated twice. 30 μL of 20 ng/μL trypsin was added to the protein mixture in 50 mM TEAB pH 8.5 and incubated at 37 °C overnight. Peptides were eluted twice with 75 μL of 50% acetonitrile, 0.1% formic acid. Aliquots of 20 μL of eluted peptides were quantified using the Quantitative Fluorometric Peptide Assay (cat# 23275, Thermo Fisher Scientific). Peptides are dried in a speed vac, suspended in 1.67% acetonitrile, 0.08% formic acid, 0.83% acetic acid, 97.42% water, and placed in an autosampler vial.

NanoLC MS/MS analysis

Peptide mixtures were analyzed by nanoflow liquid chromatography-tandem mass spectrometry (nanoLC-MS/MS) using a nano-LC chromatography system (UltiMate 3000 RSLCnano, Dionex), coupled on-line to a Thermo Orbitrap Fusion mass spectrometer (ThermoFisher Scientific, San Jose, CA) through a nanospray ion source. A trap and elute method was used. The trap column was a C18 PepMap100 (100 μm × 2 cm, 5 μm particle size) from Thermo Scientific. The analytical column was an Acclaim PepMap 100 (75 μm × 25 cm) from Thermo Scientific. After equilibrating the column in 98% solvent A (0.1% formic acid in water) and 2% solvent B (0.1% formic acid in acetonitrile (ACN)), the samples were injected onto the trap column and subsequently eluted (300 nL/min) by gradient elution onto the C18 column as follows: isocratic at 2% B, 0–5 min; 2–4% B, 5–6 min; 4–25% B, 6–59 min; 25–44% B, 59–64 min; 44–90% B, 64–66 min; isocratic at 90% B for 1 min, 90–5% B, for 1 min; isocratic at 5% B for 1 min (increase flow to 600 nL/min); 5–90% B from 69 to 71 min; isocratic at 90% B for 2 min; 90–2% B, for 1 min; isocratic at 2% B, 74–86 min; isocratic at 2% B for 1 min (reduce flow to 450 nL/min); isocratic at 2% B for 2 min (reduce flow to 300 nL/min); and isocratic at 2% B till 90 min.

All LC–MS/MS data were acquired using XCalibur, version 4.4 (ThermoFisher Scientific, San Jose, CA) in positive ion mode using a data-dependent acquisition (DDA) method with a 3 s cycle time. The survey scans (m/z 375–2000) are acquired in the Orbitrap at 120,000 resolution (at m/z = 400) in profile mode, with a maximum injection time of 50 ms and an AGC target of 400,000 ions. The S-lens RF level is set to 60. Isolation was performed in the quadrupole with a 1.6 Da isolation window, and HCD MS/MS acquisition as performed in centroid mode with detection in the Ion Trap, with the following settings: parent threshold = 25,000; normalized collision energy = 32%; maximum injection time 35 ms; AGC target 20,000 ions. Monoisotopic precursor selection (MIPS) and charge state filtering were on, with charge states 2–10 included. Dynamic exclusion was used to remove selected precursor ions, with a ± 10 ppm mass tolerance, for 30 s after acquisition of one MS/MS spectrum.

Database searching

Tandem mass spectra were extracted and charge state deconvoluted by Proteome Discoverer (version 2.5.0.402, ThermoFisher Scientific, San Jose, CA). Deisotoping was not performed. All MS/MS spectra were searched against a Uniprot database of reviewed human sequences (obtained June 11th, 2019) and common laboratory contaminants (cRAP, thegpm.org) using Sequest. Searches were performed with a parent ion tolerance of 10 ppm and a fragment ion tolerance of 0.6 Da. Trypsin was specified as the enzyme, allowing for two missed cleavages. Fixed modification of carbamidomethyl (C) and variable modifications of oxidation (M) as well as protein terminal acetylation, methionine loss, and acetylation with methionine loss were specified in Sequest. Protein and peptide FDR weres set to 1% and protein identification required 2 peptides per protein.

Synaptic enrichment analyses

To interpret the synaptic interactome of AD and PART BDTO, we utilized Synaptic Gene Ontology (SynGO; v1.2), a web-based repository and -omics tool that provides detailed, experimentally validated annotations of synaptic genes, including their associated processes and subcellular locations within the synapse [75]. Prior to enrichment analysis, contaminant proteins, reverse sequences, and proteins identified “only by site” were filtered out. In addition, proteins identified by a single peptide and not identified or quantified consistently in the same condition were removed. Two levels of enrichment analyses were performed. First, all proteins from the AD and PART BDTO interactomes were individually submitted to SynGO for synaptic enrichment. Second, proteins were categorized based on their presence in AD-specific, PART-specific, or shared BDTO interactomes using Venny 2.1 [108] and subsequently analyzed for further synaptic enrichment using SynGO. For all enrichment analyses, evidence filtering was set to ‘Stringent’ to ensure maximum reliability of annotations. This level of stringency includes data derived only from neuronal biological systems, experimentally validated techniques that excluded overexpression or pharmacological interventions, and analyses that did not rely on annotator inference.

Statistical analysis

Prior to hypothesis testing, all data were assessed for normality through quantitative (Shapiro–Wilk or Kolmogorov–Smirnov tests) and qualitative (histograms, QQ plots) methods. Outliers were identified using the ROUT method (Q = 1%) [100]. For comparisons between two independent groups (e.g., PART vs. AD), either a two-tailed independent samples t-test (for parametric data) or a Wilcoxon signed-rank test (for non-parametric data) was employed (α = 0.05). Linear regression was used to assess correlations between variables.

For comparisons across multiple groups, ANOVA or Kruskal–Wallis tests were utilized under parametric or non-parametric conditions, respectively. Post-hoc analyses were conducted with Dunnett’s (parametric) or Dunn’s (non-parametric) tests. TauO measurements (i.e., total engagement, binding, and internalization) across treatment groups (e.g., AβO or RAP) were normalized to the Control, with all baseline values scaled to 1. Such normalization minimized inter-run variance and allowed treatment effects to be expressed as fold changes relative to the Control. In addition to the Kruskal–Wallis test, a one-sample t-test was employed to compare the sample means of each treatment group to a hypothetical value of 1 (representing the Control). Although both tests yielded comparable results, Kruskal–Wallis statistics were reported due to the test’s flexibility with respect to underlying data distributions and its correction for multiple comparisons, thus reducing the risk of false positives [76].

All statistical analyses and graphical representations were performed using GraphPad Prism 10.0 (GraphPad Software, La Jolla, California, USA). Figures were prepared with BioRender (BioRender.com) and Adobe Illustrator (Adobe Inc., San Jose, CA, USA).

Results

Synaptic Aβ pathology is associated with increased tauO internalization in AD versus PART synaptosomes

To ensure scientific rigor, key aspects of our experimental paradigms (illustrated in Fig. 1 and Supp. Figure 1) were systematically optimized and validated. Briefly, this included quality control assessments of our recombinant oligomer preparations (Supp. Figure 2), confirmation of synaptosome structural integrity through TEM imaging (Supp. Figure 3), visualization of synaptic rtauO engagement via EM immunogold (Supp. Figure 4), optimization of flow cytometry gating (Supp. Figure 5) and acquisition (Supp Fig. 6) and parameters for small particle assessment (i.e., synaptosomes), validation of the PK-digestion assay for evaluating oligomer internalization (Supp. Figure 7), and titration of synaptic rtauO binding and internalization as a function of incubation time (Supp. Figure 8). The details of these validation steps are outlined in the Supplementary Information section.

Fig. 6 — Synaptic protein interactome of PART and AD brain-derived tau oligomers (BDTO). BDTO were co-immunoprecipitated (antibody: T18) from the PBS-soluble fractions of PART and AD human hippocampal autopsy tissue. Pooled PART (n = 4) and AD (n = 4) BDTO samples were subjected to proteomic analysis via LC–MS/MS and subsequent Synaptic GO (SynGO) enrichment analysis with evidence filters set to maximum stringency. SynGO results are displayed as sunburst plots, where color-coded segments denote significantly enriched synaptic compartments or pathways. a Enrichment of synaptic compartments associated with PART (top panel) and AD (bottom panel) BDTO. b Venn diagram showing the number of shared and unique proteins between PART and AD BDTO. Unique proteins for each condition were further analyzed via SynGO enrichment. Individual proteins comprising enriched terms are listed below the respective sunburst plots

Before evaluating synaptic rtauO binding and internalization in the context of pre-existing amyloid pathology, we first assessed the levels of endogenous Aβ and tauO in AD versus PART synaptosomes. Synaptosomes were isolated from the SMTG of individual PART (n = 7) and AD (n = 8) cases. Notably, Aβ was consistently detected in all AD synaptosomes, while no Aβ signal was observed in any of the PART samples (Fig. 2a). In addition, the tauO levels were significantly higher in AD synaptosomes compared to PART (t(13) = 2.55, p = 0.02) (Fig. 2d, f). To determine whether Aβ was associated with tauO pathology in AD synaptosomes, a linear regression analysis was performed, which revealed a positive correlation between Aβ and tauO, though this relationship did not reach statistical significance (slope = 2.33, R² = 0.48, F(1, 5) = 4.57, p = 0.09) (Fig. 2d). Enrichment of synaptic proteins in PART and AD synaptosomes was confirmed by the detection of PSD-95 (Supp. Figure 9a) and synaptophysin (Supp. Figure 9b), both of which showed comparable expression across conditions. Additionally, pooled synaptosomes from both conditions exhibited trends in Aβ and tau pathology consistent with those observed in individual synaptosome preparations (Supp. Figure 10).

Fig. 2 — Western blot analysis of endogenous Aβ and tau pathology in SMTG synaptosomes from PART and AD cases. Synaptosomes isolated from the SMTG of PART (n = 7) and AD (n = 8) post-mortem brain specimens, used in binding and internalization assays, were evaluated for endogenous Aβ and tau pathology. Each lane represents an individual patient sample, with identifiers indicated above each lane. Pertinent experimental details are provided on the right side of each blot. a and b Aβ detection with corresponding total protein staining for normalization, performed using the Tricine gel system. d and e tauO detection with corresponding total protein staining for normalization, performed using the Bis–Tris gel system. Red dashed boxes highlight quantified regions for each probe. c Linear regression analysis of Aβ and tauO levels in AD synaptosomes. f Quantification of tauO in PART versus AD synaptosomes. Statistical significance was assessed using an independent samples t-test (α = 0.05), with *p < 0.05 indicating significance

Next, we compared the binding and internalization of rtauO in PART versus AD synaptosomes. Total rtauO engagement did not differ significantly between PART and AD synaptosomes (Fig. 3a). However, surface-bound rtauO was significantly higher in PART synaptosomes compared to AD, where no synaptic surface binding was observed (t(8) = 5.65, p < 0.0001) (Fig. 3b). In contrast, internalized rtauO was markedly higher in AD synaptosomes as compared to PART (t(8) = 5.41, p = 0.0006) (Fig. 3c).

Fig. 3 — rtauO internalization is increased in AD synaptosomes, as compared to PART, and is modulated by synaptic membrane proteins. The binding and internalization of rtauO was assessed by flow cytometry in human synaptosomes isolated from the SMTG of PART and AD cases. a Total engagement, b surface-bound, and c internalized rtauO were measured and normalized to the maximum total engagement MFI within each group. d–f To evaluate the role of synaptic membrane proteins, the same parameters were assessed in synaptosomes pre-treated with proteinase K (PK_pre). Representative flow cytometry histograms display fluorescence intensity distributions for total engagement and internalized tauO. Biological replicates, n_PART = 7, and n_AD = 8; independent experiments, n = 5. Statistical analysis was performed using an independent samples t-test (α = 0.05). Significance levels are indicated as follows: ***p < 0.001

To elucidate the role of synaptic membrane proteins in the binding and internalization of rtauO, PART and AD synaptosomes were pre-treated with proteinase K (PK_pre, 1 mg/mL) prior to oligomer exposure. Consistent with our findings in untreated synaptosomes, total rtauO engagement did not differ significantly between PART and AD PK_pre synaptosomes (Fig. 3d). However, in contrast to untreated samples, PK_pre treatment eliminated differences in both rtauO surface binding (t(8) = 1.91, p = 0.09) and internalization (U = 6.50, p = 0.25) between PART and AD synaptosomes (Fig. 3e and f). Representative flow cytometry histograms further illustrate the density of total engaged and internalized rtauO in PART and AD synaptosomes under these conditions (Fig. 3d–f).

AD neuropathology has been associated with synaptic membrane damage [16, 67, 73, 95, 142], potentially confounding interpretations of AβO’s role in synaptic tauO internalization. To evaluate membrane integrity, the set of same pooled SMTG synaptosomes (as used in Fig. 3) from PART and AD cases were treated with Calcein AM, a membrane-permeable dye that fluoresces when retained within structurally intact cellular compartments. Both PART and AD synaptosomes exhibit high and comparable levels of Calcein positivity, with 86.3% in PART and 88.9% in AD (Supp. Figure 13, middle panels). Treatment with 2% Tween-20, significantly reduced Calcein positivity to 30.5% in PART and 34.1% in AD synaptosomes, validating the assay’s sensitivity to membrane integrity. These findings demonstrate that both PART and AD synaptosomes exhibit comparable levels of synaptic membrane integrity, with a high proportion of intact synaptosomes in both conditions.

AβO modulates synaptic binding and internalization of tauO in control synaptosomes

To further investigate the influence of rAβO on synaptic rtauO binding and internalization, we exposed standardized synaptosomes isolated from the SMTG of healthy Control brain specimens to rtauO (2.5 µM) in the presence of varying concentrations of rAβO (0–10 µM). To minimize variance due to potential confounding factors (e.g. detector sensitivity, variance in rtauO fluorophore conjugation efficiency, synaptosome aggregation, etc.) data from multiple experiments were pooled. Prior to pooling, data within each paradigm were normalized to the MFI of rtauO in synaptosomes treated exclusively with rtauO, with this baseline value set to 1. Such normalization ensures consistency in comparative analyses where subsequent values reflect fold changes relative to rtauO alone.

Our results demonstrate that rAβO significantly enhances synaptic rtauO total engagement, binding, and internalization in a concentration-dependent manner (tot. eng.: (H(4) = 23.97, p < 0.0001); binding: (H(4) = 23.99, p < 0.0001); internalization: (H(4) = 12.39, p = 0.0062) (Fig. 4). RtauO total engagement increased approximately fivefold at 5.0 µM rAβO (p = 0.0014) and eightfold at 10 µM rAβO (p < 0.0001) compared to untreated samples—0 µM rAβO (Fig. 4a). Additionally, when tested across a lower concentration range (0.05–1.0 µM), rtauO total engagement was also notably increased in the presence of 5.0 µM rAβO (Supp. Figure 8). Both surface-bound and internalized rtauO levels were elevated, with surface-bound rtauO increasing approximately tenfold at 5.0 µMr AβO (p = 0.0013) and 13-fold at 10 µM rAβO (p < 0.0001) (Fig. 4b) and rtauO internalization increasing approximately 1.5-fold at both 5 µM (p = 0.0086) and 10 µM rAβO (p = 0.00142) (Fig. 4c). Representative flow cytometry dot plots illustrate the fluorescence intensity of total engaged and internalized rtauO under varying rAβO concentrations, corroborating quantified outcomes (Fig. 4d).

Fig. 4 — rAβO increase the binding and internalization of rtauO in human synaptosomes. The effect of rAβO on the binding and internalization of rtauO in standardized human synaptosomes, isolated from the SMTG of Control cases, was assessed by flow cytometry. a Total engagement of rtauO (2.5 µM) as a function of increasing rAβO concentrations (0–10.0 µM) is shown. b Surface-bound rtauO and c internalized rtauO per synapse were measured under the same conditions. Biological replicates, n = 8; independent experiments, n = 8. Statistical analysis was performed using Kruskal–Wallis (α = 0.05) with Dunn’s post-hoc test. Significance levels are indicated as follows: ****p < 0.0001, **p < 0.01, *p < 0.05. d Representative flow cytometry dot plots displaying the fluorescence intensity of total engaged and internalized rtauO in the presence of varying rAβO concentrations

Supplementary experiments were conducted to investigate rtauO’s propensity for self-engagement and its interaction dynamics with other protein oligomers at the synaptic interface. These experiments revealed that rtauO does not modulate its own synaptic engagement across a range of concentrations (Supp. Figure 11). These findings suggest a specific enhancing effect of AβO on tauO synaptic engagement.

Role of synaptic membrane proteins in tauO binding and internalization

To examine the role of synaptic membrane proteins in differential rtauO engagement in the presence of rAβO, the same parameters were examined in synaptosomes pre-treated with proteinase K (PK_pre). Despite PK_pre treatment of synaptosomes, rAβO continued to increase total rtauO engagement (H(4) = 19.75, p = 0.0002); post-hoc analyses revealed increases of approximately twofold at 5 µM rAβO (p = 0.04) and threefold at 10 µM rAβO (p < 0.0001) compared to untreated samples—0 µM rAβO (Fig. 5a). Surface-bound rtauO exhibited a similar trend (H(4) = 16.93, p = 0.0007), with an approximately six-fold increase at 5 µM rAβO (p = 0.03) and 12-fold increase at 10 µM rAβO (p = 0.001) compared to untreated samples (Fig. 5b). Conversely, rtauO internalization was not significantly modulated by rAβO across any of the tested concentrations (H(4) = 4.64, p = 0.20) (Fig. 5c).

Fig. 5 — Synaptic membrane proteins modulate the binding and internalization of rtauO in human synaptosomes. The influence of synaptic surface proteins on the binding and internalization of rtauO (2.5 µM) was assessed via flow cytometry in Control human SMTG synaptosomes pre-treated with PK (PK_pre, 1 mg/mL). a Total engagement, b surface bound, and c internalized rtauO, were measured as a function of rAβO (0–10.0 µM). Data are expressed as fold change from baseline (0 µM rAβO). Biological replicates, n = 8; independent experiments, n = 4. Statistical analysis was performed using Kruskal–Wallis (α = 0.05) with Dunn’s post-hoc test. Significance levels are indicated as follows: ****p < 0.0001, ***p < 0.001, **p < 0.01, *p < 0.05. d Representative flow cytometry dot plots display the fluorescence intensity of total engaged and internalized rtauO in the presence of varying rAβO concentrations

Among the many proteins that regulate synaptic membrane trafficking and transportation, LDL receptor-related protein 1 (LRP1) has emerged as a potential mediator of Aβ-tau synergy [23, 152]. Notably, LRP1 is well-established for its role in Aβ production and clearance [47, 68, 68], with emerging evidence revealing its importance in tau spreading [116] and engagement of high molecular weight (oligomeric) tau species [33, 34]. To investigate the role of LRP1 in modulating AβO-driven tauO synaptic binding and internalization, we conducted focused experiments using standardized SMTG Control synaptosomes (n = 8). We first confirmed the presence of full-length LRP1 in human synaptosomes via LC–MS/MS analysis (Supp. Figure 9a). To elucidate LRP1’s functional role in synaptic tauO dynamics, we examined rtauO binding and internalization in the presence of receptor-associated protein (RAP), a high-affinity LRP1 inhibitor [103, 119, 120]. RAP treatment significantly reduced both the binding and internalization of rtauO (binding: H(3) = 7.69, p = 0.01; 0.5 µM RAP, p = 0.04; 5.0 µM RAP, p = 0.03) (internalization/uptake: H(3) = 8.26, p = 0.003; 5.0 µM RAP, p = 0.01) (Supp. Figure 14, b-1 and b-2). However, RAP's inhibitory effects were negated by introduction of rAβO (Supp. Figure 14, b-3 and b-4), suggesting that AβO may modulate the interaction between LRP1 and tauO at the synaptic interface.

Synaptic interactome of PART and AD brain-derived tau oligomers (BDTO)

To identify candidate proteins involved in modulating the Aβ-tau interplay, BDTO and their interacting proteins, were isolated from post-mortem hippocampal specimens of PART (A−, T+) and AD (A+, T+) cases (n = 4 in each condition). BDTO were co-immunoprecipitated from PBS-soluble hippocampal lysates using T18, a conformation-specific antibody targeting toxic tau oligomers [86, 113, 115, 124], and subsequently analyzed by LC–MS/MS. By comparing the tau oligomer interactome in the presence and absence of Aβ pathology, we aimed to investigate whether and how Aβ influences the protein network of native tau oligomers (i.e. BDTO) within the soluble brain parenchyma, particularly at the synaptic interface. BDTO were isolated specifically from the hippocampus, where tau pathology predominantly localizes in PART cases [35, 37, 64]. Additionally, isolating BDTO from the PBS-soluble total homogenate fraction offered two advantages: (1) prioritizing “soluble” tau oligomers, widely regarded as the most toxic forms [48, 57, 79, 114], and (2) enabling an unbiased interactome analysis by avoiding forced detection of synaptic proteins (when restricted to the synaptosomal fraction).

Co-immunoprecipitated proteins were further analyzed via SynGO enrichment, allowing for the generation of disease-specific synaptic BDTO interactomes. SynGO results were visualized as sunburst plots, highlighting enriched synaptic sub-compartments (pre- vs. post-synaptic) and associated pathways (Fig. 6). BDTO from PART cases (Fig. 6a, top panel) exhibited overall enrichment in synaptic proteins (− log10 Q-value ≥ 6). Specifically, PART BDTO demonstrated enrichment in both pre-synaptic and post-synaptic proteins (− log10 Q-value ≥ 6 and ≥ 5, respectively), with a preferential enrichment towards the pre-synaptic compartment. Within the pre-synapse, PART BDTO were notably enriched in proteins associated with synaptic vesicle cycling (− log10 Q-value ≥ 4), whereas the post-synaptic compartment was enriched in cytoskeletal, membrane, and cytosolic proteins (− log10 Q-value ≥ 2). In contrast, BDTO from AD cases (Fig. 6a, bottom panel) exhibited greater overall enrichment of synaptic proteins (− log10 Q-value ≥ 8) as compared to PART BDTO (− log10 Q-value ≥ 6). AD BDTO also showed enrichment in both pre-synaptic and post-synaptic proteins, but unlike PART BDTO, the gradient of enrichment was more evenly distributed between the two compartments (− log10 Q-value ≥ 8 for both). Within the pre-synapse, AD BDTO were enriched in proteins associated with the endocytic zone, cytoskeleton, and synaptic vesicle (− log10 Q-value ≥ 4, 3, and ≥ 2, respectively), while the post-synaptic compartment was primarily enriched in cytoskeletal proteins (− log10 Q-value ≥ 5).

A total of 442 proteins were identified across both AD and PART BDTO interactomes; 140 were common to both conditions, while 77 and 225 proteins were unique to PART and AD, respectively (Fig. 6b). Further SynGO enrichment analysis of these unique proteins revealed no significant associations in PART BDTO. Conversely, proteins uniquely associated with AD BDTO exhibited enrichment in both pre-synaptic and post-synaptic compartments (− log10 Q-value ≥ 5 and ≥ 3, respectively), with preferential post-synaptic enrichment. In AD BDTO, the synaptic enrichment profile was more diverse, with significant involvement of endocytic zone (AP2M1, CLTC, CTBP1, PACSIN1) and cytosolic (GDA, SH3GL2, STXBP1, SYT1) proteins in the pre-synaptic compartment (log10 Q-value ≥ 3 and ≥ 2, respectively), as well as cytoskeletal (ABI2, ACTN1, ACTN4, NEFL, SPTBN2) and endocytic zone (AP2A1, CLTC, RAB5A) proteins within the post-synaptic compartment (− log10 Q-value ≥ 4 and ≥ 2, respectively).

In sum, the enrichment analysis of proteins within PART and AD BDTO interactomes reveals a shift in tau’s synaptic distribution: from a predominantly pre-synaptic focus in PART to a more balanced presence across both pre- and post-synaptic compartments in AD. This shift suggests a role for Aβ in expanding tau’s synaptic reach, potentially facilitating trans-synaptic tau oligomer spreading. Furthermore, the enrichment of distinct proteins in AD vs. PART BDTO interactomes is suggestive of the notion that Aβ pathology in AD may drive tau oligomer internalization through endocytic pathways at the post-synaptic interface, potentially accelerating the propagation of tau pathology.

Discussion

Growing evidence underscores the critical role of Aβ in driving pathological tau propagation and facilitating its regional spread within the CNS. In this study, we provide the first direct evidence in human synapses that Aβ oligomers significantly enhance tau oligomer binding and internalization. Our work not only reinforces established distinctions between PART and AD, but also reveals, for the first time, the complete absence of pathological Aβ in PART synapses, compared to AD. The presence of endogenous Aβ in AD synaptosomes appears to promote tau internalization, further supporting a direct role for Aβ in modulating tau pathology. PK_pre experiments support the hypothesis that AβO enhance tauO uptake via synaptic membrane proteins. By isolating BDTO from both PART and AD cases, we identified candidate proteins and potential pathways that may mediate the Aβ-tauO interplay at the post-synaptic interface. These findings offer new insights into the molecular mechanisms that govern pathological tau spreading in AD, shedding light on how Aβ and tau oligomers synergistically contribute to synaptic dysfunction.

This study leverages rigorously validated human synaptic preparations, providing clinically meaningful insights into AD pathogenesis and enhancing translational relevance beyond animal models. Scientific rigor was ensured through systematic validation, including quality control of recombinant oligomer preparations, TEM imaging to confirm synaptosome integrity, and EM immunogold labeling to visualize synaptic rtauO engagement. Flow cytometry parameters were optimized for small-particle synaptosome analysis, while PK-digestion assays and incubation titration experiments reliably assessed rtauO binding and internalization. Together, these approaches establish a robust framework for investigating tau oligomer dynamics and Aβ’s modulatory effects at the synaptic interface in AD.

To address our first objective, we assessed the extent of pre-existing Aβ and tau pathology in PART and AD synaptosomes. Given that PART is operationally defined as a primary tauopathy [37], the absence of Aβ pathology in PART synaptosomes was expected. However, the complete absence of Aβ in all PART cases and its consistent presence in all AD cases is particularly striking. In AD synaptosomes, Aβ was resolved as a prominent band within the 8.5–12 kDa range, a finding consistent with previous reports that emphasize the prevalence of low molecular weight/low-n oligomeric Aβ species—primarily dimers and trimers—detected within synaptosome-enriched P2 fractions from AD neocortical autopsy specimens using SDS-PAGE [13, 43, 133]. Additionally, numerous studies have isolated naturally occurring low-n Aβ oligomers, also resolving primarily as dimers and/or trimers via SDS-PAGE, which have been shown to drive impairments in synaptic plasticity and memory function [30, 74, 94, 129, 140].

Notably, AD synapses exhibited significantly higher endogenous tauO burden compared to PART synapses, prompting us to investigate whether Aβ levels were correlated with tauO pathology. While not statistically significant, a positive association between Aβ and tauO levels was evident in AD synaptosomes. Consistent with prior studies showing AβO precede hyperphosphorylated tau in human AD synaptosomes [13, 43, 88], our findings suggest AβO may also inform synaptic tauO accumulation.

Emerging evidence within the trans-synaptic spreading framework indicates that misfolded tau can be secreted by neurons [22], with additional studies suggesting that Aβ-induced alterations in synaptic activity may enhance tau release into the extracellular space [93, 112]. Furthermore, we recently demonstrated that preformed rAβO promote the general engagement of rtauO in human frontal cortex synaptosomes from non-demented Control subjects [89]. Based on these findings, we first hypothesized that pre-existing Aβ pathology would also modulate the binding and internalization of rtauO in human AD synapses compared to PART (lacking Aβ). We further hypothesized that preformed rAβO would modulate the binding and internalization of rtauO in Control synaptosomes. Consistent with our prior observations [89], we found that rAβO significantly increased the total engagement of rtauO, as well as its surface binding and internalization in Control SMTG synaptosomes. Interestingly, rtauO did not facilitate its own engagement, suggesting a specific modulatory role of AβO in tauO dynamics. Although numerous studies have demonstrated tau’s capacity for self-propagation [49, 51, 79, 101], the observation that tauO does not facilitate its own synaptic engagement is not entirely surprising. Most primary tauopathies—such as PART [36], frontotemporal dementia [25], argyrophilic grain disease [18], and corticobasal degeneration [40]—are characterized by localized tau accumulation and regionally confined pathology. This distinction likely underscores the subtle difference between the ability of pathological tau to “propagate” within specific regions and its capacity to “spread” across broader neural networks.

While we observed no significant differences in the overall engagement of rtauO between PART and AD synaptosomes, we found a marked increase in rtauO internalization in AD synaptosomes, with little-to-no detectable surface-bound rtauO. However, in Control synaptosomes exposed to rAβO—a scenario likely reflecting the early phase of AD disease progression—rtauO internalization was accompanied by an increase in surface-bound rtauO. This suggests that AD synaptosomes may be primed to favor tau oligomer internalization. One potential explanation is the formation of pre-existing annular ion pores by AβO in AD synaptosome membranes, facilitating tauO entry [72, 81, 125]. Such pores have been hypothesized to destabilize membrane integrity [16, 24, 67] and promote oligomer uptake [15, 81, 125], thus warranting further investigation as a potential mechanistic link between Aβ and tau interactions in AD. However, the high and comparable levels of Calcein positivity between AD and PART synaptosomes indicate that, while AβO may alter membrane properties, synaptic membranes in AD remain largely intact, minimizing the likelihood of pervasive membrane destabilization.

Our findings indicate that Aβ oligomers promote tau oligomer internalization in human synapses through pathways reliant on synaptic membrane proteins, as evidenced by the complete abolition of rtauO internalization in PK_pre treated AD synaptosomes and Control synaptosomes exposed to rAβO + rtauO. To explore the role of synaptic proteins in the Aβ-tau interplay, BDTO were co-immunoprecipitated from hippocampal tissues of PART (A⁻, T⁺) and AD (A⁺, T⁺) cases, analyzed via LC–MS/MS, and interpreted using SynGO enrichment to explore how Aβ informs the protein network of native tau oligomers (i.e. BDTO) within the soluble brain parenchyma. Several noteworthy observations emerged from these analyses. First, AD BDTO exhibited greater enrichment of synaptic proteins compared to PART BDTO, consistent with the broader trends observed in this study. Interestingly, PART BDTO showed preferential enrichment toward the pre-synaptic compartment, whereas AD BDTO displayed a more evenly distributed interactome across both pre- and post-synaptic compartments. This is supported by prior observations in iPSC models of human primary tauopathies, such as FTD and PSP, which reveal a strong association between tau and pre-synaptic proteins [141], pre-synaptic vesicles [154], and network-level deficiencies in pre-synaptic signaling [66]. In contrast, studies of AD human brain tissue, along with cellular and animal models, indicate a more global synaptopathy that affects both pre- and post-synaptic compartments, as highlighted in several independent investigations and reviews [23, 31, 54, 65, 92, 118, 135]. This shift suggests that the presence of Aβ may fundamentally alter the reach of synaptic tau pathology, extending from a primarily pre-synaptic focus [53] to one that encompasses both synaptic regions, possibly indicative of an Aβ-facilitated trans-synaptic tauO spreading as observed in AD.

More intriguing results emerged when analyzing the unique proteins found in AD BDTO, after excluding those present in PART. These findings revealed a specific enrichment of post-synaptic proteins, suggesting that Aβ may recruit tauO to the post-synaptic compartment. Notably, proteins related to canonical endocytic process [41, 117], namely AP2M1 (adaptor related protein complex 2 subunit mu 1), CLTC (clathrin heavy chain), CTBP1 (C-terminal binding protein), and PACSIN1 (protein kinase C and casein kinase substrate in neurons 1) were observed in the pre-synapse, while AP2A1 (adaptor related protein complex 2 subunit alpha 1), CLTC, and RAB5A (Ras-related protein Rab-5A) were observed in the post-synapse. Cross-referencing these findings with NeuroPro, a comprehensive database of proteomic changes in human AD brain tissue [4], revealed that AP2A1, CLTC, and PACSIN1 have been found in association with both neurofibrillary tangle and amyloid plaque pathology, while AP2M1, CTBP1, and RAB5A were exclusively linked to plaque pathology. The concordance of these observations with previous reports in human AD pathology strengthens the reliability of our findings, while also offering novel evidence for the involvement of these proteins in native BDTO. The presence of CLTC is particularly compelling, as recent studies implicate clathrin-mediated endocytosis in the internalization of both Aβ [47, 63, 130] and tau oligomers [34, 46, 114].

We recognize that a non-crosslinked co-IP of native BDTO likely captures only strong, irreversible interactors, requiring consideration of additional mechanisms beyond the interactome data. In this context, LDL receptor-related protein 1 (LRP1) emerges as a potentially pivotal mediator of the Aβ-tau synergy observed here. Compelling evidence has demonstrated that LRP1 interacts with Aβ, APP, and APOE4, influencing both the production and clearance of Aβ [47, 68, 68]. Moreover, recent studies highlight LRP1’s pivotal role in tau endocytosis and regional spread [116], with further evidence suggesting that LRP1 can bind high molecular weight/oligomeric tau [33, 34]. The fact that LRP1 employs clathrin-mediated mechanisms for ligand internalization [21, 34, 131], strengthens the premise for our investigation of LRP1’s role within this study, particularly in light of our observed enrichment of endocytic proteins associated with AD BDTO. In our focused investigation, we confirmed the presence of full-length LRP1 in human synaptosomes—a significant finding given that while LRP1 is well-characterized in murine synapses [102] and has been identified in human neurons [26, 38, 68] and astrocytes [84, 119, 153], this is, to our knowledge, the first report of full-length LRP1 in human synapses. Consistent with previous observations that RAP-mediated inhibition of LRP1 reduces tau binding [34, 116], we found that RAP treatment significantly diminished both rtauO binding and internalization in human synapses. Notably, the addition of rAβO appeared to abolish RAP’s inhibitory effects, suggesting two possible interpretations: (1) Aβ oligomers may modulate the interaction between LRP1 and tau oligomers, potentially by inducing a conformational change in LRP1 affecting its affinity for RAP, or (2) Aβ oligomers may promote tau oligomer internalization via LRP1-independent pathways. Given the significant enrichment of endocytic proteins within the AD BDTO interactome, we favor the former interpretation. Nonetheless, these findings highlight the need for a more comprehensive investigation into LRP1’s mechanistic role in mediating and/or modulating the interplay between Aβ and tau oligomers in human synapses.

The overall findings of this study should be considered within the context of certain key limitations. Our focus on the interactions between Aβ and tau oligomers was guided by strong evidence identifying these species as the most pathogenic and synaptotoxic conformations of the respective proteins [9, 30, 50, 105, 146]. However, the effects of other aggregation states, such as Aβ monomers or fibrils, on tau oligomer dynamics, as well as whether tau monomers can be internalized by synaptosomes or require an oligomeric conformation, warrant further investigation. The use of post-mortem tissue limits the bioenergetic potential of fully functional synapses, restricting our ability to investigate energy-intensive pathways. Nevertheless, synaptosomes retain a degree of energy-dependent functionality [7, 71, 104], with further evidence indicating that even in ATP-depleted cells, ligands can still be efficiently sequestered into deeply invaginated pits, rendering them inaccessible to extracellular agents [120, 122, 123]. While the use of human synaptosomes enhances the translational relevance of our study, the inherent heterogeneity of synaptic sub-types (i.e. excitatory vs. inhibitory, pre- vs. post-synaptic, etc.) complicates the interpretation of our data within the context of selective neuronal vulnerability. Furthermore, the experimental approach implemented in this study does not capture the full spectrum of mechanisms by which toxic oligomers spread. For instance, while most extracellular tau appears to be non-vesiculated [22, 70, 96], the potential involvement of vesiculated oligomers in tau spread remains an important avenue for exploration. Moreover, studies indicate that glial cells, including microglia [3, 52, 109, 147, 150] and astrocytes [42, 99, 132]—both independently and in concert [2, 8, 137]—play pivotal roles in the progression of tau pathology.

Looking ahead, our findings underscore the need for a deeper understanding of the molecular interplay between Aβ and tau oligomers at the synaptic interface. A logical next step would involve a more comprehensive screening of the proteins mediating this interaction, expanding upon and reinforcing our current results. Transitioning from synaptosomes to cell-based models—such as those utilizing mutagenesis paradigms or microfluidic chambers—could offer greater insight into the cellular mechanisms driving AβO-tauO dynamics. Lastly, validation through in vivo models will be essential to enhance our understanding of the spatiotemporal dynamics of AβO-driven tau pathology in human disease.

Conclusion

Our findings provide novel insights into the synaptic interplay between pathological Aβ and tau, emphasizing the role of Aβ oligomers in driving the binding and internalization of tau oligomers in human synapses. This study builds on our prior understanding of human tauopathies, offering potential mechanisms that may explain the disparity in clinical outcomes between PART and AD. The identification of synaptic proteins, particularly those involved in endocytosis, reveal critical pathways through which Aβ and tau oligomers may synergistically accelerate AD progression. The outcomes of this study highlight the need to further investigate such mechanisms, aimed at disrupting the synergy between pathological Aβ and tau oligomers in human synapses.

Supplementary Information

Below is the link to the electronic supplementary material.

Supplementary file1 (DOCX 6089 KB)^{(5.9MB, docx)}

Acknowledgements

This work was supported by the UTMB Kempner Fellowship awarded to SK, as well as NIH/NIA grants F30AG085974 to SK, 1R01AG073133 to GT, and R01AG072883 to GT and AL. We extend our sincere gratitude to the donors and families that make these studies possible. We thank Dr. Peter Nelson from the University of Kentucky for dissection and neuropathological evaluation of the post-mortem autopsy specimens utilized in this study. We also thank Drs. Nemil Bhatt and Md Anzarul Haque (members of RK lab group) for providing preformed and tested recombinant tauO; Meredith Weglarz and the UTMB Flow Cytometry Core Facility for their training and assistance with the flow cytometry FC synaptosome immunophenotyping experiments, as well as general FC data analysis; Dr. Lee Palmer, Dr. William Russell, and the UTMB Mass Spectrometry Core Facility, for their assistance with acquisition and interpretation of mass spectrometry data; Dr. Vsevolod Popov and the UTMB Electron Microscopy for their training and assistance with acquisition of electron microscopy data.

Author contributions

SK was involved in project conceptualization, data collection and analysis, as well as drafting, revision, and approval of the manuscript. MM assisted with preparation of AβO, as well as establishment of and training in key experimental methods. WZ assisted with acquisition of Western blot data. AF assisted with protocol optimization and acquisition of EM images. RK provided preformed recombinant tauO used for all binding and internalization experiments. AL assisted with procurement and banking of Control, AD, and PART human brain tissue. GT contributed to project conceptualization, data analysis and interpretation, as well as drafting, revision, and approval of the manuscript. All authors reviewed the manuscript, provided revisions if necessary, and agreed to publication.

Funding

This work was supported by the UTMB Kempner Fellowship awarded to SK, as well as NIH/NIA grants F30AG085974 to SK, 1R01AG073133 to GT, and R01AG072883 to GT and AL.

Availability of data and materials

Raw proteomic and flow cytometry data will be made available through the AD Knowledge Portal or provided by the authors upon reasonable request.

Declarations

Conflict of interest

The authors declare that they have no financial or non-financial competing interests to disclose.

Ethical approval and consent to participate

Postmortem frozen brain tissues used in this study were obtained through established material transfer agreements with the Alzheimer’s Disease Research Center (ADRC) at Sanders-Brown Center on Aging, University of Kentucky, and the Layton Aging and Alzheimer’s Disease Center (ADC) at Oregon Health and Science University. All donors were enrolled in ongoing brain aging studies, with informed consent obtained prior to participation, and all protocols adhered to Institutional Review Board (IRB) guidelines at each institution. All donor subject samples were de-identified before being provided to the University of Texas Medical Branch (UTMB); therefore, no approval was required from the UTMB IRB per CFR §46.101(a). Animals (mice) used procure EM immunogold images were housed in adherence with USDA standards and sacrificed in compliance with UTMB’s Institutional Animal Care and Use Committee (IACUC), as well as UTMB IRB, approved protocols.

Footnotes

Publisher's Note

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplementary file1 (DOCX 6089 KB)^{(5.9MB, docx)}

Data Availability Statement

Raw proteomic and flow cytometry data will be made available through the AD Knowledge Portal or provided by the authors upon reasonable request.

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PERMALINK

Amyloid-β oligomers increase the binding and internalization of tau oligomers in human synapses

Shrinath Kadamangudi

Michela Marcatti

Wen-Ru Zhang

Anna Fracassi

Rakez Kayed

Agenor Limon

Giulio Taglialatela

Abstract

Supplementary Information

Introduction

Materials and methods

Clinical and neuropathological assessment of donor brain tissue

Standardized isolation and preparation of synaptosomes

Preparation and fluorescent labelling of preformed recombinant tauO

Preparation of preformed recombinant AβO

Electron microscopy (EM)

Transmission EM for morphology quality control

EM immunogold staining of ultrathin sections for qualitative validation of synaptic rtauO engagement

Western blot

Synaptosome binding and internalization assay for Aβ and tau oligomers

Role of AβO in tauO binding and internalization in PART vs. AD synaptosomes

PK digestion of synaptic membrane proteins

Role of AβO in tauO binding and internalization in control synaptosomes

Ancillary experiments

Calcein AM assay for assessment of synaptosome membrane integrity

Synaptosome immunophenotyping

Flow cytometric (FC) evaluation of synaptosomes

Instrumentation

Acquisition parameters for small particle measurement

FC data analysis

Isolation of brain-derived tau oligomers

Preparation of PBS-soluble brain homogenates

Co-immunoprecipitation

Liquid-chromatography with tandem mass spectrometry (LC–MS/MS)

Sample digestion

NanoLC MS/MS analysis

Database searching

Synaptic enrichment analyses

Statistical analysis

Results

Synaptic Aβ pathology is associated with increased tauO internalization in AD versus PART synaptosomes

Fig. 1.

Fig. 6.

Fig. 2.

Fig. 3.

AβO modulates synaptic binding and internalization of tauO in control synaptosomes

Fig. 4.

Role of synaptic membrane proteins in tauO binding and internalization

Fig. 5.

Synaptic interactome of PART and AD brain-derived tau oligomers (BDTO)

Discussion

Conclusion

Supplementary Information

Acknowledgements

Author contributions

Funding

Availability of data and materials

Declarations

Conflict of interest

Ethical approval and consent to participate

Footnotes

References

Associated Data

Supplementary Materials

Data Availability Statement

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