pTau pathology in the retina of TAU58 mice: association with ganglion cell degeneration and implications on seeding and propagation of pTau from human brain lysates

Abstract

The accumulation of abnormal phosphorylated Tau protein (pTau) in neurons of the brain is a pathological hallmark of Alzheimer’s disease (AD). PTau pathology also occurs in the retina of AD cases. Accordingly, questions arise whether retinal pTau can act as a potential seed for inducing cerebral pTau pathology and whether retinal pTau pathology causes degeneration of retinal neurons. To address these questions, we (1) characterized pTau pathology in the retina of TAU58 mice, (2) determined the impact of pTau pathology on retinal ganglion cell density, and (3) used this mouse model to test whether brain lysates from AD and/or non-AD control cases induce seeding in the retina and/or propagation into the brain. TAU58 mice developed retinal pTau pathology at 6 months of age, increasing in severity and extent with age. TAU58 mice showed reduced retinal ganglion cell density compared to wild-type mice, which declined with age and pTau pathology progression. Brain lysates from non-AD Braak neurofibrillary tangle (NFT) stage I controls increased retinal pTau pathology after subretinal injection compared to phosphate-buffered saline (PBS) but did not accelerate pTau pathology in the brain. In contrast, subretinally injected AD brain lysates accelerated pTau pathology in the retina and the contralateral superior colliculus. Subretinal injection of AD brain lysates, but not of non-AD brain, induced in this context a neuroinflammatory response in the retina and in the contralateral primary visual cortex. These results lead to the following conclusions: (1) Brain lysates from AD and non-AD sources can accelerate tauopathy within the retina. (2) The anterograde propagation of pTau pathology from the retina to the brain can be triggered by subretinal injections of AD brain lysates. (3) Such subretinal injections also provoke a neuroinflammatory response in both the retina and the visual cortex. (4) The accumulation of retinal pTau is associated with the degeneration of the involved ganglion cells, indicating that retinal tauopathy might contribute to vision impairment in the elderly and underscore the retina’s potential role in spreading tau pathology to the brain.

Introduction

The retina has been considered to represent a window to the brain [1, 2]. Its accessibility, combined with advancements in ocular imaging techniques, makes retinal imaging a potentially highly promising biomarker for neurodegenerative diseases. For example, ocular imaging methods such as optical coherence tomography (OCT) and fundus photography offer non-invasive, detailed views of retinal structures, potentially enabling early detection of neurodegenerative processes [3,4,5]. Pathological conditions such as Alzheimer’s disease (AD), other tauopathies, transactive response DNA-binding protein (TDP)-43 proteinopathies, multiple sclerosis, and Parkinson’s disease often manifest in specific retinal changes that can be seen by pathological analysis [6,7,8,9,10,11,12,13,14]. Therefore, it is essential to learn how the retina is involved in the respective neurodegenerative diseases such as AD to correctly interpret biomarker findings.

AD is the most prevalent neurological disorder leading to dementia [15]. The neuropathological hallmarks of AD include the aggregation of phosphorylated Tau protein (pTau) leading to the formation of neurofibrillary tangles (NFTs) and the extracellular deposition of amyloid-β (Aβ) peptides in the brain [16,17,18,19]. pTau pathology, however, is not restricted to AD. Other tauopathies also develop an accumulation of pTau in the brain [20]. The retina has been shown to exhibit pTau pathology as well, in AD but also in other tauopathies and even in non-AD control cases [11, 21,22,23]. Primary age-related tauopathy (PART) describes AD-like NFTs and neuropil threads in brains lacking Aβ plaques [24]. Whether PART is the earliest manifestation of AD tauopathy [25, 26] or an independent tauopathy [27, 28] is not conclusively answered today, although morphological, molecular, and cryo-EM patterns argue in favor of a strong link to AD pTau pathology [25, 29, 30]. To better understand the role of retinal pTau pathology in the context of cerebral tauopathies, it is important to know whether retinal pTau accumulation can cause neurodegeneration in the retina and can propagate into the brain. The first evidence for pTau-driven neurodegeneration was reported in Tau transgenic mice by functional alterations observed in retinal ganglion cells [31].

The distribution pattern of pTau pathology in the human brain and its hierarchical sequence involving different brain regions [18, 32] led to the hypothesis that pTau pathology propagates anterogradely along axonal routes [18, 33]. Evidence from animal experiments in Tau transgenic and wild-type mice as well as in neuronal cell culture systems suggests propagation of pTau pathology in an anterograde manner [34,35,36,37,38]. Whether such a propagation from the retina to the brain is possible, is not yet clear. Intravitreal injections of pTau and α-synuclein seeds did not induce retinal seeding or propagation into the brain of Tau transgenic and wild-type mice [39, 40]. However, uptake of injected pTau in ganglion cells was reported as well as pTau expression in mice with a Thy1 promoter-driven transgenic Tau expression [39].

Thus, to investigate pTau toxicity and propagation in the retina, an animal model for tauopathies is needed. Several mouse models producing pTau have been generated based on the transgenic expression of human mutant 4-repeat Tau carrying either the P301S or the P301L mutation [36, 41, 42]. The C57BL/6J.Tg(Thy1TAU)58/2 (TAU58) mouse model expresses human 4-repeat Tau with the P301S mutation driven by the neuron-specific Thy1 promoter and develops a predictable pattern of pTau pathology [41] that can be accelerated by additional Aβ pathology [43]. Accordingly, the tauopathy in the TAU58 mice appears to have similar properties as in AD and may be, therefore, well suited for studying retinal affection and its contribution to cerebral pTau pathology. Considering the uptake of injected pTau in retinal ganglion cells and the expression of pTau in retinal ganglion cells of transgenic mice [39], TAU58 mice appear to be a promising model to clarify the anterograde propagation of pTau from the retina to the brain although not modelling perfectly the retinal tauopathy observed in the humans [11, 23, 44, 45]. However, it is a very good model to investigate whether pTau is principally capable of inducing neurodegeneration and/or spreading towards the brain.

Here, we used TAU58 mice to clarify whether retinal pTau pathology can induce neurodegeneration in the retina and whether seeding and propagation of retinal pathology can be induced by AD and/or non-AD/PART brain lysates. To overcome the inner limiting membrane, which is a barrier to retinal uptake of intravitreally injected brain lysates [39], we performed subretinal injections. Our results indicate that pTau pathology can cause degeneration of retinal ganglion cells and that AD brain lysates can induce seeding and propagation of pTau pathology from the retina to the brain.

Materials and methods

Animal models

A total of 63 female TAU58 heterozygous transgenic mice were enrolled in the study: 30 underwent subretinal injections, while 33 remained non-injected, across various age groups (6 months, n = 9; 9 months, n = 10; 12 months, n = 10; 15 months, n = 4). Furthermore, 14 wild-type (WT) littermates (6 months, n = 7; 12 months, n = 6) were included for comparative analysis. TAU58 transgenic mice were specifically chosen for this study because they express human 4-repeat Tau with the P301S mutation, driven by the Thy1 promoter [41]. Notably, TAU58 mice begin developing cortical and brainstem pTau lesions at an early stage, as early as 2 months of age, with a progressive increase in quantity and distribution across various brain regions with advancing age [41]. The transgenic expression of pTau ensures the presence of sufficient pTau in the retina to induce/accelerate its aggregation by seeds [41, 46]. Previous studies have further confirmed the expression of pTau in the retinas of mice carrying the P301S mutation [47, 48].

Female mice were used for these experiments to reduce sex-related variations in the results leading to a reduction in the number of animals in accordance with the 3R principle [49]. Previous studies have not reported mechanistic differences between male and female mice regarding pTau seeding and propagation [34, 35, 50, 51], supporting our rationale for restricting the study to female mice. Studies showed that female TAU58 mice have a milder pTau pathology profile, offering distinct advantages in controlled seeding effects within the superior colliculus (SC), lateral geniculate nucleus (LGN), and the primary visual cortex (V1) [41, 43].

TAU58 heterozygous mice were crossbred with C57BL/6J WT mice. Retinas were examined histopathologically, and any mice exhibiting visible ocular abnormalities were excluded from the study.

Genotyping was performed as reported [41]. Mice were maintained on a 12-hour light/dark cycle, with access to food and water ad libitum. All animal care and experimental procedures were conducted in accordance with the ethical standards set by the KU Leuven Ethical Committee (P169/2020) and in compliance with Belgian and European law.

Human autopsy cases

Human brain autopsy brain tissue (temporal cortex Brodmann area 21/22) form two patients was used in this study to prepare seeds with negligible and high pTau levels: 1 non-AD control with PART (Braak stage I) and 1 AD patient (Braak Stage VI) (Tab. S1). The PART case showed Braak stage I. Accordingly, the temporal cortex of Brodmann areas 21/22 of the PART case did not exhibit microscopic pTau pathology. Given that nearly all individuals over the age of 40 years develop at least initial pTau lesions in the brain, this non-AD case with an initial PART stage is considered representative of the age-matched general population [18]. Therefore, it was used as a “non-disease” control in this study. The brains were collected at UZ Leuven (Belgium) and the laboratory of Neuropathology of Ulm University (ethical approval: 54/08 (Ulm); S-52971 (Leuven)) and used for this study under approval by the ethical committee from UZ Leuven (S-64492) and in accordance with the Belgian law. Neuropathological characterization of all cases was performed by D.R. Thal.

The phases of Aβ deposition within the medial temporal lobe (referred to as AβMTL phases) were evaluated according to recommended criteria [52, 53], serving as a valid approximation for Aβ accumulation throughout the entire brain [16]. The progression of NFTs across the brain was determined using Braak NFT stages, utilizing sections immunostained with anti-pTau^S202/pT205 (AT8, Invitrogen Thermo Fisher Scientific, 1/1000) [32]. Specifically, Braak NFT stages were assessed in accordance with the methodology established by Braak et al. [18].

The frequency of pTau-positive neuritic plaques was evaluated following the guidelines set by the Consortium to Establish a Registry for Alzheimer’s Disease (CERAD) [54]. Furthermore, the degree of AD pathology was determined according to the National Institute of Aging-Alzheimer Association (NIA-AA) [52]. This determination took into consideration the Aβ-phases, the Braak NFT stage, and the CERAD score pertaining to neuritic plaque pathology. The distribution of TDP-43 pathology was determined according to the recommendations for the neuropathological assessment of limbic predominant age-related TDP-43 encephalopathy neuropathological changes (LATE-NC) [55].

Protein extraction and BCA assay

To extract insoluble material containing Tau protein, fresh-frozen brain tissue from the temporal cortex underwent a series of centrifugation steps using progressively stronger detergents, following a previously established protocol [56]. Initially, the brain tissue was homogenized in a buffer consisting of high salt and Triton X (HS-TX) and then subjected to ultracentrifugation at 121,656 × g for 30 min at 4 °C. The resulting pellet was subsequently resuspended in HS-TX solution containing 20% sucrose, followed by another round of ultracentrifugation at 121,656 × g for 30 min at 4 °C. Next, the resulting pellet was resuspended in HS-TX solution containing 2% sarkosyl and subjected to ultracentrifugation at 121,656 × g for 30 min at room temperature. Afterwards, the pellet was resuspended in phosphate-buffered saline (PBS) and underwent two rounds of ultracentrifugation at 121,656 × g for 30 min each, at room temperature. This step was performed to eliminate the detergents from the homogenates. The final supernatant was resuspended in PBS, briefly sonicated, and then stored at -80 °C for further use. The overall protein content was measured utilizing the Pierce^® BCA Protein Assay Kit (ThermoFisher Scientific) according to the manufacturer’s instructions.

ELISA

To determine the concentration of pTau in patient-derived brain homogenates, an ELISA assay was conducted (pTau181 human ELISA Kit, ThermoFisher Scientific, #KHO0631), following the manufacturer’s instructions. Duplicates were run for all samples, and final concentrations were calculated by interpolating within the polynomial range of the standard curve for each assay. The resulting interpolated concentrations and the specific amounts of injected pTau per site are presented in Table S1.

Tau biosensor cell line

To confirm the seeding potential of the sarkosyl-insoluble fraction derived from the AD and non-AD/PART cases, we used the Tau repeat domain (RD) P301S FRET Biosensor HEK-293 cell line (CRL-3275, ATCC- LGC Standards, Molsheim Cedex, France), which stably expresses constructs where the RD of Tau-P301S is fused to CFP and YFP. Incubation with Tau seeds nucleates the aggregation of these Tau reporter proteins, thereby producing a FRET signal [57]. For seeding experiments, the cells were cultured in DMEM medium, supplemented with 10% FBS, 1 mM sodium pyruvate, and non-essential amino acids (Gibco, Thermofisher Scientific, Waltham, MA, USA), under an atmosphere of 5% CO2 at 37 °C. Cells were plated at 5.000 cells/well in poly-L-Lysine-coated 384-well PhenoPlates (PerkinElmer, Mechelen, Belgium). After 16 h, cells were incubated with brain extracts using Lipofectamine 3000 (Thermofisher Scientific, Waltham, MA, USA) according to the manufacturer’s protocol. Before the incubation, the samples were sonicated for 15 min (30 s on, 30 s off at 10 A) (Bioruptor Pico, Diagenode, Seraing, Belgium). Each sample was mixed with Lipofectamine 3000 reagent and added to a mixture of Opti-MEM medium (Gibco, Thermofisher Scientific, Waltham, MA, USA) with Lipofectamine 3000. After a 15 min incubation at room temperature, 4µL of mixture was added per well in a total volume of 40µL. After 48 h, cell medium was replaced with 40µL 4% formaldehyde for 5 min, then washed three times with PBS. Nuclear staining was performed with DAPI (Thermofisher Scientific, Waltham, MA, USA). Three individual plate preparations were performed per sample as independent experiments (n = 3). High-content imaging was performed at the VIB Imaging Core (Leuven, Belgium) (Operetta, PerkinElmer, Mechelen, Belgium). Segmentation analysis of 15 fields in 6 planes at a 40x magnification was performed using the Columbus Plus digital platform (PerkinElmer, Mechelen, Belgium).

Subretinal injections

Two-month-old female TAU58 mice (Tab. S2) were anesthetized by intraperitoneal (i.p.) injection of a mixture of ketamine (Nimatek, 100 mg/mL) and medetomidine (Domitor, 1 mg/mL) in saline. In addition, eye drops with local anesthesia (oxybuprocaine 0.4%, Unicaïne, Théa Pharma) were applied on the injected eye, followed by pupil dilation eye drops (tropicamide 0.5%, Tropicol, Théa Pharma). Carbomer hydrogel (carbomer 0.2%, Vidisic, Bauch & Lomb) was applied on the non-injected eye to prevent drying of the cornea. The subretinal injection was performed under direct visualization of the fundus. A trans-vitreal injection of 1 µL of the sarkosyl insoluble fraction of the human brain lysates into the subretinal space of the lateral part of the right eye was performed using a Hamilton syringe (65RN, 600, Fisher 7633-01) equipped with a beveled 34G needle (6x, GA 33, 51 mm, Fisher 7762-06). The injections were performed with standardized lysate volume representing the sarkosyl insoluble fraction of approx. 0,5 mg brain tissue/µl. The pTau concentration in the lysates was determined by ELISA as described. Following the injection, antibiotic treatment (Tobrex, tobramycinum 3 mg/g, Alcon) was applied to the injected eye. The animals were sacrificed 16 weeks later at 6 months of age by decapitation under terminal anesthesia. Eyes and brains were removed for pathological and immunohistochemical analysis and fixed in 4% phosphate-buffered paraformaldehyde.

Immunohistochemistry

The mouse brains and eyes were fixed in 4% paraformaldehyde for three days and then embedded in paraffin. Paraffin-embedded tissue sections were cut at 5 μm thickness and subsequently deparaffinized. Heat-induced epitope retrieval was carried out using Envision™ Flex Target Retrieval Solution Low pH (Dako, K8005) for 10 min at 97 °C. Endogenous mouse peroxidase was blocked with a Peroxidase-Blocking Reagent (Dako) for 5 min to prevent nonspecific reactions. Mouse anti-pTau^S202/T205 primary antibody (AT8, Invitrogen Thermo Fisher Scientific, 1/500) conjugated to biotin was applied and incubated overnight. Immunohistochemical labeling was conducted using the ABC method, as previously described [58]. 3,3’-diaminobenzidine was utilized as a chromogen, and counterstaining was performed with hematoxylin using a Leica Autostainer (Leica, Wetzlar, Germany).

Fluorescence staining was conducted using overnight incubation with the following primary antibodies: polyclonal guinea pig anti-NeuN (neuronal nuclei, Synaptic Systems, 1/200), rabbit anti-RBPMS (RNA-binding protein with multiple splicing, PhosphoSolutions, 1/250), goat anti-Iba1 (ionized calcium-binding adapter molecule 1, Abcam, 1/300), guinea pig anti-GFAP (glial fibrillary acidic protein, Synaptic Systems, 1/400) and biotin-conjugated mouse anti-pTau^S202/T205 antibody (AT8, Invitrogen Thermo Fisher Scientific, 1/500). Subsequently, a 90-minute incubation was performed with the secondary antibodies: goat-anti rabbit (conjugated with Cy2 fluorochrome), donkey-anti goat (conjugated with Cy2 fluorochrome), goat-anti guinea pig (conjugated with Cy5 fluorochrome), and streptavidin (conjugated with Cy3 fluorochrome). Hoechst 33,342 stain (Thermo Fisher Scientific) was used to visualize nuclei. Slides were mounted using a Glycergel mounting medium (Dako). Microscopic images of the eyes were captured using a DCC 290 microscope paired with a 20 × or 40 × objective with a DFC7000 T camera (Leica). Mouse brain slides were scanned using the Zeiss Axio Scan Z.1 (Zeiss) at a 10× magnification (0.65 μm/pixel).

Positive and negative staining controls were performed.

Gallyas silver staining and quantification of fibrillar tau pathology

The silver-impregnation method of Gallyas-Braak was used to visualize filamentous Tau pathology on paraffin-embedded, 5 μm coronal sections, as previously described [59, 60]. Both positive and negative staining controls were included to ensure staining specificity.

For the quantification of fibrillar Tau, Gallyas staining was performed on both the subretinally injected (right) eyes and the non-injected (left) eyes across the examined groups (PBS, non-AD/PART, AD). Five retinal sections were analyzed: one central section containing the optic nerve, with each of the remaining sections spaced 150 μm apart, dorsally and ventrally. Only sporadic fibrillar material was seen in the retina. To simplify the interpretation the examination of the changes was performed on the entire retina cross-sections without separation into the peripheral and central parts. In the optic projection areas (SC, LGN, V1), four sections per brain region, spaced 250 μm apart, were analyzed to cover the entire regions of the SC, LGN, and V1, separately for the respective ipsi- and contra-lateral hemispheres. The presence/absence of fibrillar aggregates was presented as binary data for clarity and ease of interpretation (Tab. S3). Image analysis was performed in a blinded manner.

Quantification of neuronal density and pTau pathology

After immunolabeling with anti-NeuN and anti-pTau^S202/T205 antibodies, brain sections were scanned (Zeiss Axio Scan Z.1) and images were imported into QuPath image analysis software [61]. Three sections per brain, spaced 250 μm apart, were analyzed to cover the layers of the LGN, SC, and V1. Neurons were selected from the NeuN channel, using a defined threshold for precise visualization, and cell validation was performed through colocalization with Hoechst 33,342 nuclear staining. Neuronal density (NeuN-positive neurons/mm²) and the ratios of pTau-positive (NeuN-positive, pTau^S202/T205-positive) neurons relative to the total number of neurons (NeuN-positive) were quantified. Specific layers of the LGN, SC, and V1 were manually outlined (Fig. S1), referencing the Allen Reference Atlas – Mouse Brain (atlas.brain-map.org).

For mouse retinas, three sections were analyzed, including the central section with the optic nerve, and two sections extending 150 μm dorsally and ventrally, respectively (Fig. S2). QuPath was used for manual quantification of RBPMS-positive retinal ganglion cells and the pTau^S202/T205 immuno-positive area across the retinal ganglion cell layer. Per section, the number of RBPMS-positive cells was quantified in four regions of interest, comprising both the central and peripheral retina (Fig. S2a). The rationale for using the RBPMS marker was to distinguish the retinal ganglion cells from amacrine and glial cells [62, 63]. The presence of pTau was measured by assessing the total length of the pTau^S202/T205 positive area versus the total length of the retina, and the results were expressed as a percentage (Fig. S2b).

The evaluation of pTau threads in the optic nerve of mice receiving subretinal injections was conducted on one central sagittal section of the eye, on which the optic nerve was visible. The presence or absence of pTau alterations in each optic nerve (left/right) was evaluated via immunostaining with the anti-pTau^S202/T205 antibody, and presented as binary data for clarity and simplicity of interpretation.

Image analysis was performed in a blinded manner.

Quantification of the microglia and astrocytes

A morphometric quantitative analysis was performed to measure the percentage of GFAP-positive area, and percentage of Iba1-positive area, in the right (subretinally injected) eyes across the groups (PBS, non-AD/PART, AD) and the corresponding contralateral brain regions (SC, LGN, and V1) using specific antibodies to detect astrocytes (GFAP) and microglia cells (Iba1) following immunofluorescence staining. Digital images of the peripheral and central retinal regions, as well as two images per brain region (SC, LGN, and V1), covering the entirety of each region, were captured using a Leica DM2000 LED microscope equipped with a Leica DFC 700T camera at ×10 magnification and processed using LAS V4.8 software.

For each mouse, four images were taken from the peripheral retina (defined as 100 μm from the ora serrata) and four from the central retina (defined as 100 μm from the optic nerve). Additionally, two images were taken from each contralateral brain region (SC, LGN, and V1), analyzing the whole region for each.

The percentage of GFAP- and Iba1-positive areas were measured using ImageJ software (NIH, Bethesda, USA). For the retina, measurements were taken over a 1 mm length in both peripheral and central regions, encompassing all retinal layers except the photoreceptor layer. For the brain regions (SC, LGN, V1), the entire region imaged and analyzed at the same magnification to ensure consistency. The presence of astrocytes and microglia was quantified as the percentage of the GFAP- or Iba1-positive area relative to the total ROI area. A brightness threshold was applied to detect GFAP-positive astrocytes and Iba1-positive microglia, with the consistent threshold settings in ImageJ ranging between 64 and 255 for GFAP and 90–255 for Iba1. All samples were stained simultaneously on the same bench to ensure uniformity, and background noise was carefully eliminated to include only GFAP- and Iba1-positive areas in the analysis. Image analysis was performed in a blinded manner.

Statistical analysis

Statistical analyses were performed using Prism v.9 (GraphPad) and IBM-SPSS Statistics 25 (SPSS, Chicago, IL, USA). Linear regression investigated the association between pTau-positive area percentage and age in non-injected TAU58 mice across different ages (Tab. S4). Two-way ANOVA with Tukey’s test compared RBPMS-positive ganglion cells between 6-month-old and 12-month-old TAU58 mice and age-matched WT littermates, separately for peripheral and central retina (Tab. S5). Linear regression analyzed the association between RBPMS-positive ganglion cells and age, separately for peripheral and central retina (Tab. S6). Linear regression further examined the association between RBPMS-positive ganglion cells and pTau-positive area percentage, with age as a controlled variable, separately for the peripheral and central retina (Tab. S7). One-way ANOVA with Tukey’s test assessed differences in pTau seeding potential among brain homogenates (Tab. S8). One-way ANOVA with Dunnett’s test compared pTau-positive area percentage after subretinal injection with brain lysates, with separate analyses for left and right eyes, using PBS groups as a control (Tab. S9). An unpaired t-test compared pTau-positive area and RBPMS-positive neurons between PBS-injected and non-injected eyes (Tab. S10). Multinomial logistic regression evaluated the association between fibrillar Tau presence in the right injected eye and the respective injected brain lysates, using the PBS group as the reference category (Tab. S11). Multinomial logistic regression evaluated the association between pTau threads presence in the optic nerve and injected brain lysates, separately for the left non-injected eye and the right injected eye, using the PBS group as the reference category (Tab. S12). A one-way ANOVA with Dunnett’s test compared RBPMS-positive neurons after subretinal injections with brain lysates, with separate analyses for the peripheral and central regions of the left eye and the peripheral and central regions of the right eye, using PBS groups as a control (Tab. S13). A one-way ANOVA with Dunnett’s test compared the percentage of GFAP and Iba1-positive areas in the central and peripheral retina of injected eyes, using PBS as control (Tab. S14). Linear regression examined the association between the percentage of the pTau-positive area in the right (injected) eye and the percentages of GFAP-positive and Iba1-positive areas, separately for the peripheral and central retina (Tab. S15). Kruskal-Wallis test (non-parametric one-way ANOVA) with Dunn’s test assessed pTau pathology percentage in brain regions (SC, LGN, V1) after subretinal injection with brain lysates, with separate analyses for each brain layer, using PBS groups as a control (Tab. S16). A one-way ANOVA with Dunnett’s test examined NeuN-positive neuron count in brain regions (SC, LGN, V1) after subretinal injection with brain lysates, with separate analyses for each brain layer, using PBS groups as a control (Tab. S17). Finally, a one-way ANOVA with Dunnett’s test compared the percentage of GFAP and Iba1-positive areas in contralateral optic areas (SC, LGN, V1), between injected lysates, with separate analyses for each brain, using PBS as control (Tab. S18). Differences were considered statistically significant for two-sided p-values < 0.05 (*p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001).

Results

Characterization of pTau changes in the non-injected retina of TAU58 mice

To assess retinal changes associated with pTau in female TAU58 mice, the retinas of non-injected TAU58 mice at different ages (6 months, n = 9; 9 months, n = 10; 12 months, n = 10; 15 months, n = 4), were analyzed using the anti-pTau^S202/T205 antibody. Screening of the TAU58 mice showed pTau-related changes at 6 months of age, persisting thereafter. Specifically, the presence of pTau threads within the retinal nerve fiber layer (RNFL) and inner plexiform layer (IPL), as well as cytoplasmic accumulation of pTau within the ganglion cell layer (GCL) was detected (Fig. 1; a-b, d-e, g-h). Moreover, Gallyas staining further confirmed the presence of fibrillar forms of cytoplasmic pTau in the GCL (Fig. 1; c), whereas the retinal pTau threads remained Gallyas-negative (Fig. 1; f-i). In linear regression analysis, the percentage of retinal pTau showed a significant association with age (Fig. 1; j, Tab. S4; p < 0.0001, β = 0,920). Notably, these pTau-related changes were predominantly observed in the central retina (Fig. 1; o-r) with sporadic pathology in the peripheral retina (Fig. 1; k-n). Additionally, the presence of pTau threads was noted in the optic nerve (Fig. 1; s-v).

Fig. 1

Retinal pTau pathology in the retina non-injected TAU58 mice. Changes in pTau^S202/T205 expression were observed predominantly in the central part of the mouse retina, specifically confined to the inner region. The first cytoplasmic pTau inclusions were observed in ganglion cells of six-month-old female TAU58 mice, affecting the GCL (a, b). These inclusions exhibited a Gallyas-positive fibrillar form (c). Additionally, phosphorylated Tau threads were identified within ganglion cell dendrites in the IPL and in the RNFL (d, e, g, h), showing no reactivity to Gallyas staining (f, i). The linear regression analysis revealed a significant association between pTau pathology (expressed as a percentage of affected retina length) and age (p < 0.0001, β = 0,920) (j). Representative images of retinal pTau changes in the peripheral (k-n) and central (o-r) regions of the retina across different age groups. Panels s-v show representative images of pTau changes in the optic nerve across the same age groups. Magnifications of panels b-c, e-f, and h-i are derived from panels a, d, and g, respectively. (6 months TAU58, n = 9; 9 months TAU58, n = 10; 12 months TAU58, n = 10; 15 months TAU58, n = 4). PRL = photoreceptor layer, ONL = outer nuclear layer; OPL = outer plexiform layer, INL = inner nuclear layer, IPL = inner plexiform layer, GCL = ganglion cell layer, RNFL = retinal nerve fiber layer

The presence of retinal pTau is associated with the loss of ganglion cells in TAU58 mice

To determine whether the presence of retinal pTau pathology correlates with neuronal survival, and to assess the progression of neuronal loss with age, staining with anti-RBPMS antibody was performed to quantify ganglion cells in the retinas of 6-, 9-, 12- and 15-month-old TAU58 mice (6 months: n = 9; 9 months: n = 10; 12 months: n = 10; 15 months: n = 4), alongside age-matched WT littermates (6 months: n = 7; 12 months: n = 6) (Fig. 2; g-r). The analysis demonstrated significant ganglion cell loss in 6-month-old TAU58 mice compared to age-matched WT mice, with a 41.6% cell loss in the peripheral retina and a 25.8% cell loss in the central retina (Fig. 2; a, b, Tab. S5; two-way ANOVA with Tukey’s test; p < 0.0001 and p = 0,0060, respectively), progressing to 51.5% cell loss in the peripheral retina and 42,5% cell loss in the central retina at 12 months of age TAU58 mice compared to age-matched WT mice (Fig. 2; a, b, Tab. S5; one-way ANOVA with Tukey’s test; p < 0.0001 and p = 0.0001, respectively).

Fig. 2

The retinal pTau associates with ganglion cell loss in TAU58 mice. a: A two-way ANOVA revealed a reduced number of RBPMS-positive neurons in the peripheral region of the retinas of 6-month-old and 12-month-old TAU58 mice compared to age-matched WT mice (p < 0.0001 for both). b: Similarly, a two-way ANOVA indicated a reduced number of RBPMS-positive neurons in the central region of the retinas of 6-month-old and 12-month-old TAU58 mice compared to age-matched WT mice (p < 0.0060, p = 0.0001, respectively). c: A linear regression model in the peripheral retina demonstrated an association between age and the number of RBPMS-positive cells in TAU58 mice (p < 0.0001; β=-0.650). d: Likewise, a linear regression model in the central retina showed a negative association between age and the number of RBPMS-positive cells in TAU58 mice (p < 0.0001; β=-0.603). e: In the peripheral retina, a linear regression model controlled for age demonstrated an association between the number of RBPMS-positive cells and the percentage of retinal pTau pathology (p = 0,001; β=-0,698). f: Similarly, in the central retina, a linear regression model controlled for age showed a negative association between the number of RBPMS-positive cells and the percentage of retinal pTau pathology (p = 0,008; β=-0,520). g-l: The upper panel shows RBPMS-positive retinal ganglion cells in the peripheral retina, across different age groups and mouse strains. m-r: The lower panel shows RBPMS-positive ganglion cells in the central retina, across different age groups and mouse strains. Error bars: Min to Max in a, b. Scale bar in g is valid for all panels (g-r). (6 months WT, n = 7; 6 months TAU58, n = 9; 9 months TAU58, n = 10; 12 months WT, n = 6 12 months TAU58, n = 10; 15 months TAU58, n = 4)

Linear regression showed a significant association between age of TAU58 mice and ganglion cell loss in peripheral (Fig. 2; c, Tab. S6; p < 0.0001; β=-0.650) and central (Fig. 2; d, Tab. S6; p < 0.0001; β=-0.603) regions of the retina. Furthermore, after controlling for age, linear regression analysis showed a significant association between the number of RBPMS-positive ganglion cells and the percentage of retinal pTau pathology in the peripheral (Fig. 2; e, Tab. S7; p = 0,001; β=-0,698) and the central (Fig. 2; f, Tab. S7; p = 0,008; β=-0,520) regions of the retina.

Comparing 6- and 12-month-old WT mice, no significant differences in the peripheral and central ganglion cell density were observed (Fig. 2; a, b, Tab. S5; two-way ANOVA with Tukey’s test, p > 0.05).

Patient-derived brain homogenates induce Tau seeding in a biosensor cell line

To confirm the seeding potential of the extracted brain homogenates for subsequent experiments, we performed seeding assays using a previously established tau biosensor cell line [57]. The cell line was incubated with sarkosyl-insoluble homogenates from the temporal cortex of one non-AD/PART and one AD case. The amount of pTau in each lysate was determined by an ELISA assay, revealing a 17-fold higher concentration of pTau in the AD brain lysate compared to the non-AD/PART lysate (Tab. S1). A non-treated cell line served as the negative control. The seeding efficiency of the various extracts was quantified by counting the number of pTau fluorescent positive puncta (spots) per cell, which represent Tau aggregates (Fig. 3; a-c). Cells exposed to AD brain homogenates exhibited significantly higher levels of Tau aggregation compared to the non-AD/PART homogenate-treated cells (Fig. 3; d, Tab. S8; one-way ANOVA with Tuckey’s test; p < 0,0001). This confirms that the AD brain homogenates have a robust seeding potential, ensuring that any negative results in subsequent experiments are not due to the insufficient seeding capacity of the extracts.

Fig. 3

Patient-derived homogenates induce seeding in the pTau biosensor cell lines. a-c: A Tau (P301S) biosensor cell line underwent incubation with sarkosyl-insoluble homogenates from the temporal cortex of 1 non-AD/PART case and 1 AD case; a non-treated cell line served as a control. AD brain homogenates exhibited increased pTau seeding, indicated by an increase in green spots (indicated by white arrows), compared to the non-AD/PART case. d: After 48 h, the number of positive pTau spots/cells was automatically quantified, and the one-way ANOVA with Tukey’s multiple comparisons test revealed a significant difference (p < 0,0001) between the analyzed cases. Error bars: Min to Max in d

Subretinal injection of patient-derived brain homogenates induces seeding of pTau pathology in the mouse retina in TAU58 mice

A total of 30 six-month-old female TAU58 transgenic mice, equally divided into three groups receiving subretinal injections of sarkosyl-insoluble fractions from non-AD/PART or AD brain lysates or PBS as negative control (Fig. 4; a, Tab. S2) into the right eye, were analyzed for seeding and propagation of pTau. The brain lysates used were the same as those used for the seeding experiments in the tau biosensor cell line, with differing pTau concentrations between the sarkosyl insoluble fractions from non-AD, control/PART, and AD cases (Tab. S1). It is important to note that the AD and non-AD/PART lysates represent similar amounts of tissue (approx. 0,5 mg brain tissue / µl). The left eye served as non-injected control. Animals were sacrificed four months later, and the eyes and brains were pathologically examined.

Fig. 4

Subretinal injection of the patient-derived homogenates induces pTau seeding in the retina in TAU58 mice. a: Graphical representation of the experiment: Two-month-old female TAU58 mice were injected with PBS, non-AD/PART, and AD human brain homogenates. After 4 months, mice were sacrificed, and immunohistochemistry was conducted to check for the presence of retinal pTau pathology, as well as pTau-positive neurons in the brain b: Significant differences were noted in the percentage of retinal pTau pathology based on the type of seed injected in the right eye, with significant differences observed between PBS vs. non-AD/PART seeds (One-way ANOVA with Dunnett’s test: p = 0.0128), PBS vs. AD seeds (One-way ANOVA with Dunnett’s test: p < 0.0001). c: No significant differences were observed in the percentage of retinal pTau pathology in the non-injected (left) eye. d: In the optic nerve of the injected (right) eye, a multinomial logistic regression model demonstrated an association between the presence of pTau threads (expressed as a percentage of positive cases) and AD brain lysates (p = 0,033; OR = 9,333 [1,193 to 72,991]) e: In the optic nerve of the non-injected (left) eye, a multinomial logistic regression model demonstrated no association between the presence of pTau threads (expressed as a percentage of positive cases) and different types of injected brain lysates. f: No significant differences were found in the number of positive RBPMS cells in the peripheral injected (right) eye. g: Similarly, no significant differences were found in the number of positive RBPMS cells in the peripheral non-injected (left) eye. h: No significant differences were found in the number of positive RBPMS cells in the central injected (right) eye. i: Likewise, no significant differences were found in the number of positive RBPMS cells in the central non-injected (left) eye. Error bars: Min to Max in b, c, f, g, h, i, SEM in d, e (PBS; n = 10, non-AD/PART; n = 10, AD; n = 10 cases)

In comparison to the PBS-injected eyes, the injected eyes with brain lysates exhibited more retinal pTau pathology when injecting non-AD/PART brain lysates, and after AD brain lysate injection (Fig. 4; b, Tab. S9; one-way ANOVA with Dunnett’s test; p = 0,0128 and p < 0.0001, respectively). Conversely, the non-injected left eye did not exhibit significant differences among the examined groups (Fig. 4; c, Tab. S9; one-way ANOVA with Dunnett’s test; p > 0.05). The observed pTau changes in the injected eyes were predominantly seen in the central part of the retina (Fig. S3a-c). No differences in retinal pTau pathology as well as in the number of retinal ganglion cells were noted between non-injected 6-month-old mice and PBS-injected mice, showing no significant effect of subretinal injections on retinal pTau pathology and retinal ganglion cells survival (Fig. S4, Tab. S10, t-test; p > 0.05). To assess the presence of fibrillar Tau in the retina, Gallyas staining was performed, and fibrillar Tau was dichotomously assessed in the retinas and brains of injected mice. Fibrillar Tau was predominantly observed in the injected retinas (Fig. S5b-d, Tab. S3). In the right injected eyes, multinomial logistic regression showed a higher prevalence of fibrillar Tau aggregation with AD brain lysates compared to PBS (Fig. S5a, Tab S11; p = 0.035, OR = 13.5 [1,197 to 152,211]). No significant difference in comparison to PBS injections was observed for non-AD/PART lysates (Fig. S5a, Tab. S11; p > 0,05). In the left non-injected eyes of subretinally injected mice, as well as in the eyes of 6-month-old non-injected mice, no fibrillar Tau aggregation was observed (Tab. S3).

The presence/absence of pTau-positive threads in the optic nerve was analyzed in both non-injected and injected eyes across the examined groups (Fig. S3d-f). In the optic nerve of the injected eyes, a multinomial logistic regression model revealed a significant association between the presence of pTau threads and the injection of AD brain lysates (Fig. 4; d; Tab. S12; p = 0,033, OR = 9,333 [1,193 to 72,991]). Conversely, in the optic nerve of the non-injected (left) eyes, no association was found between the presence of pTau threads and any of the human brain lysates (Fig. 4; e; Tab. S12; p > 0.05).

No significant differences were observed between the types of subretinally injected human brain homogenates, compared to PBS injections, on ganglion cell survival in either the peripheral or central retina in the injected eyes (Fig. 4; f, h, Tab. S13; one-way ANOVA with Dunnett’s test; p > 0.05) or the contralateral, non-injected eyes (Fig. 4; g, i, Tab. S13; one-way ANOVA with Dunnett’s test; p > 0.05).

Astro- and microglial response after subretinal injection of AD, non-AD/PART brain lysates, and PBS

To clarify whether subretinal injections in the lateral part of the eye induce the activation of microglial cells and astrocytes depending on the lysate injected, we stained for astrocytes with an antibody against GFAP to assess the percentage of GFAP-positive area, and for microglial cells with the Iba1 antibody to evaluate the percentage of Iba1-positive area, which detects both residual and activated microglial cells. This analysis was conducted in both central and peripheral regions of the retina. In the central retina (Fig. 5; e-g, Fig. S9a-e, k-o, u-y), the percentage of GFAP-positive and Iba1-positive areas were significantly increased in the AD-brain lysates injected group compared to PBS-injected mice (Fig. 5; a, b; Tab. S14; one-way ANOVA with Dunnett’s test; GFAP: p = 0,0143, Iba1: p = 0,0275). No significant differences were found between the PBS and non-AD/PART brain homogenate-injected groups in the central retina (Fig. 5, a, b; Tab. S14; one-way ANOVA with Dunnett’s test; p > 0.05). In contrast, no significant differences were observed for the percentage of GFAP and Iba1-positive areas in the peripheral retina (Fig. 5; h-j, Fig. S9f-j, p-t, z-d’) across the examined groups (Fig. 5, c, d; Tab. S14; one-way ANOVA with Dunnett’s test; p > 0.05). Additionally, in the linear regression analysis, the percentage of retinal pTau in the injected (right) eye showed a significant association with the percentages of GFAP-positive and Iba1-positive areas in the central retina (GFAP: Table S15; p = 0.020, β = 0.424; Iba1: Table S15; p = 0.045, β = 0.369). In contrast, no significant associations were observed in the peripheral retina (GFAP: Table S15; p > 0.05; Iba1: Table S15; p > 0.05).

Fig. 5

Subretinal injection of AD-derived homogenates accelerates the presence of microglia and astrocytes in the retina of TAU58 mice. a, b; Significant differences were observed in the percentage of the percentage of GFAP and Iba1-positive areas in the central retina (One-way ANOVA with Dunnett’s test: GFAP: p = 0,0143, Iba1: 0,0275). c, d: No significant differences were found in the percentage of GFAP and Iba1-positive areas in the peripheral retina (One-way ANOVA with Dunnett’s test: p > 0,05). Representative merged fluorescence images show astrocytes and microglia in the GCL of the central (e-g) and peripheral (h-j) retinas of 6-month-old female TAU58 mice for AD, non-AD/PART, and PBS-injected groups. The merged images display astrocytes (GFAP) in green, microglia (Iba1) in magenta, pTau (pTau^S202/T205) in red, and nuclei (Hoechst) in blue. Single-channel images for the merged images in e-j are shown in Fig. S9; a-d’. Arrowheads indicate pTau^S202/T205 retinal cytoplasmic inclusions (g, j). Error bars: Min to Max in a, b, c, d. (PBS; n = 10, non-AD/PART; n = 10, AD; n = 10 cases)

Propagation of pTau pathology form the eye to the brain in TAU58 mice

Next, we tested whether subretinal injection with PBS or sarkosyl-insoluble fractions from non-AD/PART or AD brain into the eye initiates the propagation of pTau pathology into the optic projection areas of predominantly the contralateral hemisphere. The analysis covered layers of the SC, LGN, and V1 (Figs. 6 and a and 7 and a, and Fig. 8; a, respectively). Using immunohistochemistry with anti-pTau^S202/T205 antibody, we found only a few pTau neuronal cytoplasmic inclusion in the optic projection areas, predominantly on the contralateral side.

Fig. 6

The pTau pathology in the SC of 6-month-old TAU58 mice brain after subretinal injection of brain lysates. a: Schematic representation of a coronal section showing the delineated layers of SC, with superficial layers marked in purple and intermediate and deeper layers (DLSC) marked in blue. The Kruskal-Wallis test, followed by Dunn’s test, was conducted within the layers of the SC in the left (b, d, f, h, j, l, n) and right (c, e, g, i, k, m, o) hemispheres. b: Significant differences in the number of pTau-positive neurons were observed in the zonal layer of the SC between the PBS and AD brain homogenate injected groups (p = 0.0198). j: Similarly, significant differences were noted in the number of pTau-positive neurons in the intermediate white layer of the SC between the PBS and AD brain homogenate injected groups (p = 0.0198). No significant differences were found in the number of pTau-positive neurons between the seed injection groups for panels c, d, e, f, g, h, i, k, l, m, n and o. Panels p-r show fluorescence staining images of the contralateral (left) superior colliculus with zonal layers marked in white after subretinal injection into the right eye of 6-month-old female TAU58 mice for AD, non-AD/PART, and PBS-injected groups. The arrowhead indicates pTau pathology. Panel s: Magnified region from (a), showing neurons positive for pTau (white) in the zonal layer following the injection of AD brain homogenates. The merged image on the left displays neurons (NeuN) in red and nuclei (Hoechst) in blue, with arrowheads indicating pTau^S202/T205 cytoplasmic inclusions. The schematic figure in a was made with the help of the Allen Institute for Brain Science. (2011), pp. http://mouse.brain-map.org. Error bars: Min to Max. SC = superior colliculus, zo = zonal layer, sg = superficial gray layer, op = optic layer, ig = intermediate gray layer, iw = intermediate white layer, dg = deep gray layer, dw = deep white layer

Fig. 7

The pTau pathology in the LGN of 6-month-old TAU58 mice brain after subretinal injection of brain lysates. a: Schematic representation of a coronal section showing the delineated LGN (purple), which is part of the thalamus (blue). The Kruskal-Wallis test, followed by Dunn’s tests, was performed within the layers of the LGN for the left (b) and right (c) hemispheres. The analysis revealed no significant difference in the number of pTau-positive neurons between the seed injection groups. The schematic figure in a was made with the help of the Allen Institute for Brain Science. (2011), pp. http://mouse.brain-map.org. Error bars: Min to Max. LGN = lateral geniculate nucleus

Fig. 8

The pTau pathology in the V1 of 6-month-old TAU58 mice brain after subretinal injection of brain lysates. a: Schematic representation of a coronal section showing the delineated layers of V1. The Kruskal-Wallis test, followed by Dunn’s tests, was conducted within the layers of the V1 in the left (b, d, f, h, j, l) and right (c, e, g, i, k, m) hemispheres. h: Significant differences were observed in the number of pTau-positive neurons in the Internal Pyramidal Layer of the V1 between the PBS and AD brain homogenate injected groups (p = 0.0318). No significant difference in the number of pTau-positive neurons was found between the seed injection groups for panels a, b, c, d, e, f, h, i, j, k, and l. The schematic figure in a was made with the help of the Allen Institute for Brain Science. (2011), pp. http://mouse.brain-map.org. Error bars: Min to Max.V1 = primary visual area, VISp1 = Molecular Layer, VISp2/3 = External Granular and External Pyramidal Layer, VISp4 = Granular Layer, VISp5 = Internal Pyramidal Layer, VISp6a = Multiform Layer (sublayer), VISp6b = Multiform Layer (sublayer), LGN = lateral geniculate nucleus

Quantification of pTau-positive neurons in the SC revealed significant differences in the contralateral hemisphere, specifically in the zonal layer (SCzo) and the intermediate white layer (SCiw). Compared to the PBS-injected group, propagation of pTau pathology occurred only after subretinal injections of AD brain lysates with a high concentration of pTau (Fig. 6; b, j; Tab. S16; Kruskal-Wallis with Dunn’s test; SCzo: PBS vs. AD; p = 0,0198, SCiw: PBS vs. AD; p = 0,0198). Importantly, only 4 out of 10 AD brain lysate-injected mice exhibited propagation of pTau pathology into the contralateral zonal layer of the superior colliculus SCzo (Fig. 6; p-s), but none of the PBS and non-AD control brain lysate injected animals had pTau aggregates in this region. Other contralateral layers of the SC showed an elevated number of pTau-positive neurons; however, the differences between analyzed groups were not significant (Fig. 6; d, f, h, l, n; Tab. S16; Kruskal-Wallis with Dunn’s test; p > 0.05). Additionally, the ipsilateral layers of the SC did not exhibit differences among PBS, non-AD, and AD brain lysate-injected groups (Fig. 6; c, e, g, i, k, m, o; Tab. S16; Kruskal-Wallis with Dunn’s test; p > 0.05).

No differences were found in the percentage of pTau-positive neurons in the LGN (Fig. 7; b, c; Tab. S16; Kruskal-Wallis with Dunn’s test; p > 0.05).

Among all layers of the contralateral V1, only the internal pyramidal layer (V1Sp5) showed a higher percentage of pTau-positive neurons after subretinal injection with AD brain homogenates compared to PBS-injected mice (Fig. 8; h; Tab. S16; Kruskal-Wallis with Dunn’s test; p = 0,0318). The other layers of the contralateral V1 exhibited elevated numbers of pTau-positive neurons; however, the differences between analyzed groups were not significant (Fig. 8; b, d, f, j, l; Tab. S16; Kruskal-Wallis with Dunn’s test; p > 0.05). The ipsilateral layers of V1 also did not show differences (Fig. 8; c, e, g, i, k, m; Tab. S16; Kruskal-Wallis with Dunn’s test; p > 0.05).

No differences in the density of NeuN-positive neurons were observed between the groups after subretinal injection of PBS or sarkosyl-insoluble fractions from the respective brain lysates within the layers of the SC, LGN, and V1 (Figs. S6, S7, and S8, respectively; Tab. S17; One-way ANOVA with Dunnett’s test; p > 0.05).

Induction of neuroinflammatory response in the superior colliculus and primary visual cortex after subretinal injection of AD brain lysates

To clarify whether propagation of pTau pathology and/or injection of AD brain lysates subretinally have an impact on the neuroinflammatory response, we analyzed the brains for signs of neuroinflammation as indicated by increased astroglial and/or microglial activation. In the SC, the percentage of GFAP-positive area was elevated after subretinal injection of the AD-brain lysates compared to PBS-injected mice (Fig. 9; a; Tab. S18; One-way ANOVA with Dunnett’s test; p = 0,0426) (Fig. 9: c-h, Fig. S10a-d’). However, no differences in the percentage of Iba1-positive area were observed between the AD and PBS groups in the SC (Fig. 9; b; Tab. S18; One-way ANOVA with Dunnett’s test; p > 0.05). In the LGN, no significant differences were observed for either the percentage of GFAP (Fig. 10; a; Tab. S18; One-way ANOVA with Dunnett’s test; p > 0.05) or Iba1-positive areas (Fig. 10; b; Tab. S18; One-way ANOVA with Dunnett’s test; p > 0.05) between the groups (Fig. 10: c-h, Fig. S11a-d’). In the V1, both the percentage of GFAP (Fig. 11; a; Tab. S18; One-way ANOVA with Dunnett’s test; p = 0,0130) and Iba1-positive areas (Fig. 11; b; Tab. S18; One-way ANOVA with Dunnett’s test; p = 0,0035) were increased in the AD-injected group compared to PBS-injected mice, reflecting a pronounced glial response in this region (Fig. 11: c-h, Fig S12a-d’). No differences were observed in the percentage of GFAP and Iba1-positive area between the non-AD/PART and PBS groups in these brain regions.

Fig. 9

Presence of microglia and astrocytes in the SC of 6-month-old TAU58 mice brain after subretinal injection of brain lysates. a: Significant differences were observed in the percentage of GFAP-positive area in the contralateral SC between PBS- and AD-brain lysate-injected mice. However, no significant difference was found between PBS- and non-AD/PART-injected mice (One-way ANOVA with Dunnett’s test, PBS vs AD: p = 0,0426, PBS vs non-AD/PART: p = 0,6175), b: No significant differences were observed in the percentage of the percentage of Iba1-positive area between the examined groups (One-way ANOVA with Dunnett’s test: p > 0,05). Representative merged fluorescence images of the contralateral SC (c-e) and their higher magnifications (f-h) show astrocytes (GFAP) in green, microglia (Iba1) in magenta, pTau (pTau^S202/T205) in red, and nuclei (Hoechst) in blue. The white-outlined rectangular area in panel h highlights pTau^S202/T205 immunoreactivity localized in proximity to reactive microglia and astrocytes. Single-channel images for the merged images in c-h are shown in Fig. S10; a-d’. Magnifications of panels f, g, and h are derived from panels c, d, and e, respectively. Error bars: Min to Max in a, b. Scale bar in c is valid for panels c-e and scale bar in f is valid for panels f-h. (PBS; n = 10, non-AD/PART; n = 10, AD; n = 10 cases)

Fig. 10

Presence of microglia and astrocytes in the LGN of 6-month-old TAU58 mice brain after subretinal injection of brain lysates. One-way ANOVA with Dunnett’s test was performed on the contralateral LGN to the percentage of GFAP- (a) and Iba1-positive areas (b). The analysis showed no significant differences in the percentage of GFAP and Iba1-positive areas between PBS-injected and human-derived homogenate-injected groups (One-way ANOVA with Dunnett’s test: p > 0,05). Representative merged fluorescence images of the contralateral LGN (c-e) and their higher magnifications (f-h) depict astrocytes (GFAP) in green, microglia (Iba1) in magenta, pTau (pTau^S202/T205) in red, and nuclei (Hoechst) in blue. Single-channel images for the merged images in c-h are shown in Fig. S11a-d’. Magnifications of panels f, g, and h are derived from panels c, d, and e, respectively. Error bars: Min to Max in a, b. Scale bar in c is valid for panels c-e and scale bar in f is valid for panels f-h. (PBS; n = 10, non-AD/PART; n = 10, AD; n = 10 cases)

Fig. 11

Presence of microglia and astrocytes in the V1 of 6-month-old TAU58 mice brain after subretinal injection of brain lysates. a: Significant differences were observed in the percentage of the percentage of GFAP -positive area in the contralateral V1 between PBS- and AD-brain lysate-injected mice, while no significant difference was found between PBS- and non-AD/PART-injected mice (One-way ANOVA with Dunnett’s test, PBS vs AD: p = 0,0130, PBS vs non-AD/PART: p = 0,4174), b: Significant differences were also observed in the percentage of the percentage of Iba1-positive area in the contralateral V1 between PBS- and AD-brain lysate-injected mice, with no significant difference between PBS- and non-AD/PART-injected mice (One-way ANOVA with Dunnett’s test, PBS vs AD: p = 0,0035, PBS vs non-AD/PART: p = 0,6704). Representative merged fluorescence images of the contralateral V1 (c-e) and their higher magnifications (f-h) show astrocytes (GFAP) in green, microglia (Iba1) in magenta, pTau (pTau^S202/T205) in red, and nuclei (Hoechst) in blue. Single-channel images for the merged images in c-h are shown in Fig. S12a-d’. Magnifications of panels f, g, and h are derived from panels c, d, and e, respectively. Error bars: Min to Max in a, b. Scale bar in c applies to panels c-e, and scale bar in f applies to panels f-h. (PBS; n = 10, non-AD/PART; n = 10, AD; n = 10 cases)

Discussion

This study revealed several novel findings: (1) Subretinal injection of temporal cortex lysates from non-AD control/PART (Braak NFT stage I) and AD (Braak NFT stage VI) brains, accelerates the development of pTau pathology in retinal ganglion cells in TAU58 mice. Fibrillar pTau pathology was in this context only induced by AD brain lysates. (2) Additional accelerated pTau pathology was observed in the optic nerve and the contralateral brain hemisphere of the injected eye, particularly affecting the SC and V1 regions, in mice receiving the sarkosyl-insoluble fraction of AD-brain lysates. This effect was absent when injecting brain lysates from a non-AD control case with Braak NFT stage I or PBS. (3) In TAU58 mice, phosphorylation and aggregation of pTau increase with age in the retina and are associated with retinal ganglion cell loss. (4) Subretinal injection of AD brain lysates, but not of non-AD brain, induced a neuroinflammatory response in the form of astroglial and microglial activation in the retina and in the contralateral primary visual cortex. In the superior colliculus only astroglial activation was seen.

The first finding of this study is that AD brain and non-AD control/PART lysates can induce pTau seeding in the retina of a Tau overexpression mouse model. However, only AD brain lysates led to an increase of argyrophilic, fibrillar Tau. Previous research suggested that retinal ganglion cells can internalize human pTau after intravitreal injection [39] but seeding of aggregated forms of pTau in the neuropil was not observed, possibly due to a limited amount of pTau seeds and the intravitreal injection method as used in the respective study [39], which restricts the amount of pTau seeds reaching the retina due to diffusion barriers. Our study, employing subretinal injection, overcomes the inner limiting membrane barrier and, by doing so, delivers higher doses of aggregated pTau seeds to the outer part of the retina. Notably, the observed induction of pTau pathology in the retina following subretinal injections was induced not only by AD brain lysates (Braak NFT stage VI) with high pTau concentration but also by non-AD control brain lysates of a Braak NFT stage I case with very low levels of pTau as occurring in “normal brain” whereas induction of fibrillar AD-Tau was restricted to the AD brain lysates. Accordingly, our findings challenge the view that only high concentrations of pTau, as found in AD, can induce pre-tangle stages of retinal tauopathy. This suggests that even nearly unaffected brain tissue without local detection of pTau pathology, such as neuropathologically unaffected brain regions of PART, is sufficient to exaggerate retinal Tau pathology when applied by subretinal injection. This aligns with previous findings from in-vitro seeding assays, showing that seeding can be induced even by Braak NFT stage I cases [64]. In this context, it is important to mention that both PART and AD pTau share the same cryo-electron microscopic signature [29]. Therefore, these results provide novel insights into the modification of Tau pathology manifestations, probably modified by increased dosage or the presence of fibrillar Tau. That the induction of fibril formation was restricted to the AD brain lysate points towards preformed Tau fibrils as critical players in NFT induction. A limitation of our study is that the initial research question did not include whether pTau interacting proteins as reported for phosphorylated TDP-43 or cellular prion protein in AD brain [43, 65, 66] play a role in the local exaggeration of pTau pathology and/or fibril formation. Further research is therefore required to address this question.

Our second finding expands the current knowledge by showing that anterograde propagation of pTau pathology requires higher levels of pTau aggregates than those typically found in the sarkosyl-insoluble fraction of the temporal cortex lysate of a Braak NFT stage I case. Moreover, the induction of fibrillar pTau in retinal ganglion cells by AD brain lysates appears to be a prerequisite for proper propagation. Notably, a PART case exhibiting a more advanced Braak NFT stage (stage IV) has been earlier reported to induce propagation [50] highlighting the pivotal role of pTau aggregate concentration in this process. Additionally, non-AD/PART brain lysates with lower pTau concentrations exhibited lower in-vitro seeding activity compared to AD brain lysates and failed to induce (1) fibrillar pTau in retinal ganglion cells and (2) detectable propagation to the brain. These results align with earlier studies demonstrating anterograde pTau pathology propagation in various brain regions and in vitro [34, 35, 50, 51], and extend these findings by demonstrating that (a) the retina can also serve as a site for pTau seeding and as an origin for anterograde propagation and (b) the local induction of neuronal, argyrophilic, fibrillar Tau aggregates appears to be a prerequisite for the propagation of pTau pathology. Furthermore, the concentration/fibrillar Tau-dependent differences in propagation-induction provide insights into the gradual progression of neurodegeneration observed in human tauopathies, likely attributable to pTau pathology levels and pathology accelerators such as Aβ [42, 43, 67], which was absent in the non-AD control case used in our study but present in the AD brain.

In this context, it needs to be mentioned that, to our knowledge, this is the first demonstration of the propagation of pTau pathology from the retina to the brain in an in vivo model [39]. Although the induction of pTau pathology in the contralateral SCzo as shown here is a subtle finding only observed in a subset of the AD brain lysate-receiving TAU58 mice, none of the PBS or non-AD control brain lysate-receiving animals exhibited pTau aggregates in the SCzo. The relatively mild pTau pathology in SCzo is explained by anatomically receiving only a few projections from retinal ganglion cells [68]. Accordingly, it is likely that pTau pathology propagation took place only in those mice receiving AD brain lysates, with this spread being associated with astro- and microglial activation. This finding suggests that the retina can potentially act as an origin for anterograde pTau pathology spreading, given that there are sufficient aggregates in the retina. Inflammation involving astrocytes and microglia might be associated with this propagation [69, 70]. According to our results, and those of de Fisenne et al. [39], low doses of pTau may not be enough to start spreading of a tauopathy from the retina into the brain. For retinal pTau pathology observed in the human eye, this would support the hypothesis that the retinal tauopathy observed in AD and non-AD controls is presumably not sufficient to induce propagation to the brain, as the pTau levels in these retinas were lower than those in AD as well as in early Braak NFT stage III PART brain [23].

The finding that brain lysates from a non-AD control case with Braak NFT stage I, i.e., early stage of PART, accelerate local pTau pathology in the retina but are not sufficient for the induction of spreading into the brain as seen for AD brain lysates, broadens our understanding about the pathogenesis of AD/PART. This result demonstrates that brain lysates from cases that do not meet the criteria for AD pathology [52] have already enough pTau seeds to accelerate a local tauopathy despite the low amounts of pTau compared to those cases in advanced Braak stages with symptomatic AD [71]. Given that the tauopathy in AD and PART shows a nearly identical pattern regarding the maturation of the aggregates [30] and cryo-electron microscopic structure [29], our finding that early-stage PART brain lysates already accelerate pTau pathology strongly argues in favor of PART being considered as a precursor of AD. Once present, Aβ aggregates may accelerate pTau spreading, as previously shown in a mouse model for AD pathology [43]. A similar potential to induce seeding has also been shown for brain lysates from cases with initial Aβ deposits, when used to seed Aβ plaques in APP transgenic mouse brains [72], supporting the idea that AD neuropathological changes already have a disease-driving potential in very early disease stages.

The third important finding of this study reveals a strong association between pTau aggregation in the retina and the reduction of retinal ganglion cell density in TAU58 mice. This is in line with the role of pTau in neuronal degeneration in the brain. First, pTau accumulates in neurons before configurational changes leading to the presence of paperclip formation of Tau. Finally, fibrillar, argyrophilic pTau aggregates are formed, representing NFTs. Later on, NFT-bearing neurons die and leave ghost tangles behind [17, 30, 73]. The presence of argyrophilic NFTs in the TAU58 mouse retinal ganglion cells argues in favor of a similar process in retinal ganglion cells as in cerebral neurons. Although the TAU58 mouse model can illustrate the main effects of pTau on retinal integrity, it does not perfectly represent PReT, i.e., human primary retinal tauopathy. This is a limitation for translating the findings in the mouse model to the human retina. However, TAU58 mice provide valuable insights into the general principles of retinal pTau and its potential role in retinal degeneration. Importantly, we demonstrate that Tau accumulation in the retina is associated with significant ganglion cell loss. This may explain the functional impact of pTau on visual acuity, as reported in previous studies [48]. Our findings are consistent with the known neurotoxic effects of Tau in the brain, and we extend these observations to the retina [69, 74]. The loss of retinal ganglion cells parallels the loss of neurons in tauopathies, and this supports the hypothesis that retinal Tau pathology could contribute to vision impairment seen in the elderly [23]. An argument against the contribution of pTau aggregates to the degeneration of retinal ganglion cells is the finding that subretinal injections of AD brain lysates in TAU58 mice increased pTau pathology but did not cause an accelerated degeneration of retinal ganglion cells. Given that TAU58 mice already produce retinal pTau pathology, it cannot be excluded that the addition of a few pTau aggregates with the subretinal injections is not sufficient for an obvious additional effect on pTau toxicity, since seeds do not significantly increase the concentration of a given protein rather than accelerating aggregation and fibril formation by secondary nucleation [75]. Therefore, it is in our opinion more likely that retinal tauopathy can lead to neurodegeneration as seen in TAU58 mice compared to wild-type littermates. Accordingly, a similar role of pTau pathology in the human retina on neuronal integrity can be considered and may have a functional impact, particularly in glaucoma, a leading cause of blindness, where Tau phosphorylation is associated with disease progression [76, 77]. Similar associations have been observed in Lewy body disease, where retinal α-synuclein inclusions impact retinal degeneration and visual function [10, 78]. It remains unclear which type of pTau pathology (microscopically seen NFTs or small fibrils or oligomers in intra- or extracellular fluid compartments) primarily contributes to neuronal degeneration. Regardless of which forms of pTau are the ultimate executors of pTau toxicity, our results of ganglion cell loss in the retina of TAU58 mice may provide a possible mechanism of pTau toxicity that could contribute to the worsening of vision in individuals with a retinal tauopathy [23].

Our fourth finding demonstrates that injection of AD brain lysates but not of non-AD/PART lysates induces astrocytic and microglial response in the retina as well as in connected brain regions. The induction of neuroinflammation in connected brain regions is documented in SC and V1 with increased micro- and/or astroglial activation. Considering the organization of the mouse visual system and its connectivity, approximately 85% of visual information from retinal ganglion cells projects directly to the SC before further transmission to V1 [79, 80]. Based on our results, there are two possible explanations for the induction of astro- and/or microglial activation after injection of AD brain lysates: (1) the induced Tau pathology in retinal ganglion cells and in the contralateral superior colliculus and visual cortex triggers this neuroinflammatory response or (2) the damage induced in the retina by injecting the lysates leads to reactive changes in retina and its transmission that induce the respective alterations. Arguments for the first explanation are: (a) Tau propagation can be observed in the superior colliculus and V1, and (b) injection of non-AD brain lysates did not induce neuroinflammatory response although inducing a higher pTau expression in retinal ganglion cells. Arguments favoring the second explanation are: (a) glial activation in the superior colliculus is not seen in the zonal layer where pTau occurs, and (b) not all animals with a neuroinflammatory response show significant pTau propagation into the brain. In light of earlier findings that NFTs are associated with astro- and microglial activation in AD [81] and in mouse models of Tau pathology [46, 69], as well as evidence that inflammasome activation supports the development of Tau pathology [82], it is plausible that the first explanation, where Tau propagation is responsible for the reported neuroinflammatory response, is more likely. However, it is also possible that the microglial response itself can accelerate the development of pTau pathology [69, 82]. A third potential explanation could involve a combination of microglial activation and Tau seed propagation.

In light of the ongoing challenge in diagnosing AD during its early stages, there is a critical need for affordable, non-invasive, and easily accessible biomarkers. With recent advancements in retinal imaging, such as ocular imaging techniques like OCT and hyperspectral imaging, the retina emerges as a promising target for AD diagnosis [4, 22, 83,84,85,86]. Our findings on retinal Tau pathology, specifically in relation to (a) neuronal functionality of the retina and (b) AD-related Tau pathology and its association with neuroinflammatory responses, shed light on the potential of detecting distinct patterns of Tau changes and neuroinflammatory activation in the retina as diagnostic markers for AD neuropathologic changes (ADNC).

Limitations of this study are: (1) The expression of the promoter driving the transgene influences the spatial distribution of the observed pTau pathology. This leads to different distributions of pTau pathology in TAU58 mouse retina and human retinas with PReT. Accordingly, we can conclude only on general mechanisms of retinal pTau pathology whereas direct translation to PReT is not possible. Especially for retinal imaging studies targeting pTau pathology, transgenic mice with promoter-driven Tau expression should not be used, as they may lead to results with very limited translational potential. Other models, such as target replacement mice [39] or animals with spontaneous retinal tauopathy might be better suited. (2) The projections of RGCs to the SC and LGN differ between humans and mice, mainly because of differences in the organization and function of the visual pathways [87, 88], which may affect the translation of findings from mouse models to human data. Therefore, transgenic mouse models may not fully reflect all aspects of human retinal tauopathy. (3) Another limitation of our study is the absence of biochemical analysis to evaluate the levels of insoluble proteins, particularly insoluble Tau. Although this was not possible due to the lack of frozen retina samples, similar biochemical studies in mouse models that share the same Thy1 promoter and P301S mutation as the TAU58 model have reported lower levels of pTau in the retina compared to the brain, which is consistent with our findings [39, 89].

Conclusions

Our results demonstrate that pTau pathology can be seeded in the retina and spread to the brain given the presence of sufficient amounts of seeds as found in AD but not in non-AD control brain lysates. This process including the propagation of pTau pathology into the brain is accompanied with the activation of microglia and astrocytes highlighting a potential role of neuroinflammation in the pathology propagation process in AD. In this context, it is important to note that even low levels of pTau aggregates in Braak NFT stage I temporal neocortex (considered as non-AD control tissue) can already accelerate local retinal pTau pathology. The general potential of early-stage PART, which is often considered as non-AD control tissue, to accelerate local pTau accumulation compared to PBS, without further spreading, points to a novel view on initial disease stages of AD/ PART in the pathogenesis of AD by their disease-accelerating potential. Moreover, our study demonstrates age-dependent, progressive neurodegeneration of retinal ganglion cells in TAU58 mice, with increasing severity of retinal tauopathy. This supports the hypothesis that retinal tauopathy can lead to vision impairment and may play a significant role in the decline of visual performance in the elderly.