The study was carried out in accordance with the European Communities Council Directive of September 22nd 2010 (2010 / 63 / EEC) for care of laboratory animals and in accordance with requirements of the animal ethics committee of the local government authority (Landesamt fur Arbeitsschutz, Naturschutz, Umweltschutz und Verbraucherschutz (LANUV), North-Rhine Westphalia). Male Wistar rats were used that were 7–8 weeks old at the time of initial Aβ–treatment. Animals underwent regular physical health checks to verify that they had a continuous clean bill of health throughout the entire study. Animals were group-housed in a temperature- and humidity-monitored vivarium with a constant 12-h light–dark cycle (lights on from 7 a.m. to 7 p.m.) with ad libitum food and water access. For the purpose of in vitro electrophysiology, or immunohistochemistry studies, rats were stereotaxically injected under sodium pentobarbital anesthesia (52 mg/kg, intraperitoneally, i.p.), with Aβ(1–42) oligomers, or scrambled (control) peptide into the lateral cerebral ventricle in a dose of 10 μM, applied in 5 μL, over a 5 min period via a Hamilton syringe (Kalweit et al., 2015). The injection coordinates comprised: 0.5 mm posterior to bregma, 1.6 mm lateral to midline, 5.6 mm depth from skull surface. Further experiments in vitro and immunohistochemistry assessments were conducted 1 week after treatment to assay for early hippocampal changes. Further experiments were conducted 1 month or 4–6 m onths after treatment (Supplementary Figure S1).

The oligomeric Aβ(1–42) peptide was generated as described previously (Bozso et al., 2010; Kalweit et al., 2015). Control peptide comprised a scrambled version of the Aβ(1–42) peptide [referred to here as ScrAβ(1–42)] that contains a permutated amino acid sequence (Bozso et al., 2010; Kalweit et al., 2015). Previous studies confirmed that this scrambled peptide has no significant effect on neuronal function (Südkamp et al., 2021). Before intracerebral treatments, the peptides were incubated in phosphate-buffered saline (PBS) at a pH of 7.4 for 3 h at a concentration of 50 μM and at room temperature to ensure adequate oligomerization of the Aβ(1–42) peptide (Bozso et al., 2010). The mixture was then diluted to the final concentration of 10 μM. This dose was equivalent to 50 pmol, or 225 ng of the respective peptide.

Rats underwent implantation of electrodes into the medial perforant path and the dorsal dentate gyrus, under sodium pentobarbital anesthesia (Nembutal, 52 mg/kg, intraperitoneally (i.p.), Boehringer Ingelheim, Ingelheim, Germany), as described previously (Kalweit et al., 2015). During this procedure, a monopolar recording electrode was implanted in the suprapyramidal granule cell layer of dentate gyrus (3.1 mm posterior to bregma, 1.9 mm lateral to the midline) and a bipolar stimulation electrode was implanted in the medial perforant pathway (6.9 mm posterior to bregma, 4.1 mm lateral to the midline). To enable subsequent peptide treatment, a cannula was implanted into the lateral cerebral ventricle with the following coordinates: 0.5 mm posterior to bregma, 1.6 mm lateral to the midline. For in vitro electrophysiological and immunohistochemistry studies, only a cannula was implanted. Pre- and postsurgical analgesia was conducted using meloxicam (Metacam, 0.2 mg/kg, i.p., Boehringer Ingelheim Vetmedica GmbH, Ingelheim, Germany). Following verification of the integrity of the evoked potentials 1 week after surgery, the first set of experiments were commenced.

In vivo electrophysiological recordings of local field potentials were obtained from freely behaving rats. For this, an evoked response was triggered by stimulating at low frequency (0.025 Hz) with single biphasic square wave pulses of 0.2 ms duration per half wave, generated by a constant current isolation unit. For each time-point measured during the experiments, five records of evoked responses were averaged. The first 6 time-points recorded at 5 min intervals were used as baseline and all time-points are shown in relation to the average of these 6 points. The evoked potential manifested itself as a positive going field excitatory postsynaptic potential (fEPSP), upon which a negative-going population spike (PS) was imposed. The fEPSP was measured as the slope measured on the first five steepest point of the positive-going potential. The PS was measured as the amplitude obtained on the first negative deflection of the PS. The direction of change of fEPSP and PS was consistently the same (not shown). By means of a stimulus–response determination (evaluation of nine different stimulation intensities from 100 to 900 μA in 100 μA steps) the maximal PS amplitude was found, and during experiments all potentials employed as baseline criteria were evoked at a stimulus intensity that produced 40% of this maximum. Long-term potentiation (LTP) was evoked by means of high-frequency stimulation (HFS, comprised of a stimulus burst of 15 pulses each of 0.2 ms duration applied at 200 Hz and repeated 10 times with a 10 s interval between bursts). For each time-point 5 consecutive evoked responses at 40 s intervals were averaged, and the results were expressed as the mean percentage ± standard error of the mean (S.E.M.) of the average of the first 6 recordings. Recordings were made every 5 min. Until 30 min after HFS and then every 15 min until 4 h had elapsed. The following day an additional 1 h of recordings was obtained. Peptide treatment occured after the initial verification of LTP.

Postmortem analysis of electrode localization was conducted to verify the final location of the electrodes. The brains were carefully removed from the cranium, the tissue was fixed in 4% paraformaldehyde (PFA) in phosphate buffered saline (PBS, pH 7.4) for 5–7 days following by cryoprotection in 30% sucrose. The brain tissue was then cut frontally on a cryostat (Leica CM 3050S) into 30 μm slices. After mounting on gelatin-coated slides, the sections were stained in 0.1% cresyl violet (Sigma-Aldrich Chemie GmbH, Munich, Germany). This staining was conducted in order to visualize the hippocampal structures and identify the electrode tracts. Data from animals with misplaced electrodes were excluded from the study.

Two groups of rats were treated intracerebrally with oligomeric Aβ(1–42), or control peptide (ScrAβ(1–42)), for in vitro electrophysiological studies. The first group was investigated 1 week after injection, and the second after 4–6 months. The range of 4–6 months was chosen for patch clamp experiments because of the increasing difficulty of obtaining effective patch seals on cells from the aging hippocampus (Ting et al., 2014). Animals were deeply anesthetized by inhalation of the anesthetic isoflurane, and then decapitated. The brains were rapidly removed and dissected in ice-cold dissection medium composed of (in mM): NaCl (87), KCl (2.4), MgSO₄ (1.3), CaCl₂ (0.5), NaHCO₃ (26), NaH₂PO₄ (1.25), D-Glucose (2). Transverse hippocampal slices (350 μm thick) were prepared using a vibrating blade microtome (VTS1000, Leica, Germany). After cutting, the slices were incubated in a holding chamber in the dissection medium for 30 min at 35°C. After incubation, the slices were transferred to recording chambers where they were used for patch-clamp recordings. During experiments, the slices were continuously perfused with oxygenated (95% O₂/ 5%CO₂) artificial cerebrospinal fluid (aCSF) of the following composition (in mM): NaCl (125), KCl (3), MgSO₄(1.3), CaCl₂(2.5), NaHCO₃(26), NaH₂PO₄(1.25), D-Glucose (13). The flow rate of aCSF was 1.5–2 mL/min. The temperature in the recording chamber was maintained at 30°C. With the help of an upright BX51WI microscope (Olympus, Japan), whole-cell patch clamp recordings were then performed from visually identified cell bodies of dentate gyrus granule cells using infrared illumination.

Borosilicate glass recording pipettes, used for recordings, were filled with an intracellular solution comprising (in mM): Potassium gluconate (97.5), KCl (32.5), EGTA (5), HEPES(10), MgCl₂(1), Na₂ATP(4) (pH 7.3; 290 mOsm). Recordings were performed in current-clamp mode using an amplifier (EPC10 USB, HEKA Electronic, Germany). The data were subjected to low-pass filtering at 2.9 kHz and digitized at 10 kHz.

Intrinsic membrane properties were analyzed using PATCHMASTER acquisition software and AP feature software (MATLAB code developed in Department of Psychology, University of Connecticut, courtesy of Prof. M. Volgushev). Resting membrane potential was determined as the mean value recorded during a continuous period of 10 s. Input resistance was calculated from the slope of the linear fit of the relationship between the change in membrane potential (∆V) and the intensity of the applied current (between −60 pA and + 60 pA) duration 600 ms. To analyze action potential properties, square currents (duration 600 ms) were applied in the range of 5pA to 300pA, using 5pA steps. The amount of current necessary to evoke an action potential from resting membrane potential was determined as a threshold current in pA. The first action potential was subjected to analysis of its active properties, such as the maximum and minimum spike voltage, spike amplitude, action potential width (value measured at the point where the AP had reached 20% of its maximum) and afterhyperpolarisation (AHP) amplitude. The action potential amplitude was measured as voltage difference from the threshold value to spike peak. Firing frequency properties were examined by applying square current pulses (duration 1 s) in the range of 0 pA to 400 pA in 50 pA steps. The firing frequency was analyzed as the number of spikes elicited during applied current (duration 1 s).

Behavioral assessments were conducted in different groups of animals either 1 or 6 months following intracerebral inoculation with oligomeric Aβ(1–42), or control peptide. Behavioral tasks were conducted in an open field arena (80 × 80 × 80 cm). A camera was mounted above the open field to record the experiments for off-line analysis. Exploration of an object was defined as when the rat directly contacted the object with the snout, or with the snout pointed toward the object and was within 2 cm distance of the object.

The rats were habituated to the testing procedure and allowed 10 min of exploration time in the empty open field for the 3 consecutive days preceding the experiment. After habituation, the rats were tested in a spatial recognition task (Figure 1C). In the sample trial, the rat was placed at the center of the open field with two identical objects (blue triangles in Figure 1C). After 5 min the rat was returned to its home box for a pause of 3 h before the test trial. In the test trial, the rat was placed at the center of the open field with the same two objects: one remained at the old location (blue filled triangle in Figure 1C), the other was displaced to a new location (orange filled triangle in Figure 1C). The new and old locations were randomized to avoid any possible preference of the location in the open field. The scorer was blind to the treatment the rats received.

The day after the spatial recognition task, the rats were tested in an object recognition task (Figure 1D). The experimental setup was as described above but used another two sets of novel objects. In the sample trial, the rat was placed at the center of the open field with two identical objects (blue rectangle in Figure 1D). After 5 min the rat was returned to its box for a pause of 3 h before the test trial. In the test trial, the rat was placed at the center of the open field with one old object (blue rectangle in Figure 1D) presented in the sample trial and one new object (orange oval in Figure 1D) and was allowed to explore for 5 min. The objects used as new or old objects were randomized and the location of new object was randomized. The scorer was blind to both the treatment of the rats and whether the object was new or old.

The exploration time of different objects was referred to as a percentage of the sum exploration time. Statistical analysis was conducted between the exploration time of different objects within one group of animals within one experiment.

In a different group of animals, fluorescence in situ hybridization (FISH) was conducted 1 month following intracerebral inoculation with oligomeric Aβ(1–42), or control peptide. Rats were handled by the experimenter and habituated to the recording chamber that measured (40 × 40 × 40 cm) for 3 consecutive days. On the experiment day, in the same chamber, the animals were exposed to two novel objects for 5 min followed by a pause of 3 h duration. In a subsequent exploration period, object A was returned to the chamber in exactly the same position as before, and object B was placed in a new location in the chamber (as shown in Figure 1D and described above). The animals explored for 5 min. During the exploration period, the behavior of the animals was recorded. The brains were rapidly removed 40 min after commencement of the second exploration and then shock-frozen in isopentane at −80°C on dry ice and stored at −80°C until being sectioned 20 μm thick on a Cryostat (Leica CM 3050S). The coronal sections containing hippocampus (3.6–4.0 mm posterior to Bregma) were mounted directly on ‘superfrost plus’ slides (Gerhard Menzel GmbH, Braunschweig, Germany) and stored at −80°C until further processing. We additionally included naïve animals that underwent the same handling and habituation procedure, as was conducted with the experimental groups. These animals did not undergo surgery. On the day of the experiment, they resided in the recording chamber for the same duration as the ‚item-place exploration’ animals until brain extraction, without engaging in an exploration task.

Fluorescein-labeled RNA probes were generated using the Ambion MaxiScript Kit (Invitrogen, Carlsberg, United States). Homer1a cDNA plasmid was prepared by Entelechon (Bad Abbach, Germany) and GenScript (GenScript Biotech, New Jersey, United States) using a 1.2 kb Homer 1a transcript according to the sequence of Brakeman et al. (1997). The cDNA probes were prepared from the linearized cDNA using Ambion MaxiScript Kit and premixed RNA labeling nucleotide mix containing the fluorescein-labeled UTP (Invitrogen, Carlsberg, United States). The RNA probes were purified on Mini Quick Spin RNA columns (Roche Diagnostics, Mannheim, Germany) and their yield and integrity were verified using gel electrophoresis.

At the time-point of further processing, one glass slide per animal (including three brain slices each) was chosen and left at room temperature until the slices were defrosted. The glass slide showing slices from the dorsal hippocampus at ~3.8 mm from Bregma was chosen for each individual animal. We then applied the following protocol for the Homer1a mRNA in situ hybridization (Hoang et al., 2018):

After 10 min of fixation in fresh, filtered and ice cold 4% PFA in PBS, the slides were washed for 2 min in 2x saline-sodium citrate buffer (SSC). The slides were then left for 10 min in acetic anhydride solution followed by 5 rinses in 2x SSC. After a final wash in 2x SSC for 5 min, the slides were prehybridized in formamide /4 x SSC solution (1:1) for 10 min at 37°C. The fluorescein-labeled RNA probe was diluted in 1x hybridization buffer (Sigma-Aldrich Chemie GmbH, Munich, Germany) (100 ng/100 μL), heated at 90°C for 5 min and the slides were hybridized for approximately 17 h in a humid chamber at 56°C. To confirm the specificity of the hybridized signal, we additionally conducted a negative control FISH test whereby the fluorescein labeled Homer1a RNA was not added to the brain sections (data not shown). Following the hybridization, the slides were first washed 3 × 5 min using 2 x SSC and the remaining single stranded mRNA was removed using 10 μg/mL RNase A in 2 x SSC for 15 min at 37°C. Afterwards, various washing steps were conducted: 10 min in 2x SSC at 37°C, 10 min in 0.5 x SSC at 56°C, 30 min in 0.5 x SSC at 56°C,10 min in 0.5 x SSC at room temperature (RT), 5 min in 1x SSC at RT twice, and finally 5 min in tris-buffered saline (TBS) thrice at room temperature. After these washing steps, we pretreated the slices with 3% H₂O₂ in TBS for 15 min followed by 3 exposures for 5 min to 1x TBS. In order to prevent unspecific binding of proteins we used 10% n-goat serum in TBS-Tween 20 (Polysorbate, 0.2% Tween, TBS-T) containing 20% Streptavidin (Vector Labs, Burlingame, USA) for 60 min followed by 90 min incubation with anti-fluorescein [1:2000] (Dianova, Hamburg, Germany) in 10% n-goat serum in TBS-T containing 20% Biotin (Vector Labs, Burlingame, USA). Following three washes, each of 5 duration, in TBS the signal was amplified using biotinylated Tyramine (bT. Adams, 1992). The slides were left in TBS containing bT (1%) and H₂O₂ for 20 min and afterwards washed three times for 5 min in TBS. Finally, we visualized the signal using StrepAvidin Cy2 (1:250) (Dianova, Hamburg, Germany) in 10% n-goat serum in TBS-Tween 20 at room temperature for 60 min. The slides were washed again 3 × 5 min in TBS, and quickly washed with distilled water. Afterwards, slides were quickly dipped in 70% ethanol, and stained using 1% Sudan black B (Merck KGaA, Darmstadt, Germany) in 70% ethanol (Oliveira et al., 2010). Slides were air-dried overnight and mounted in DAPI (4′,6-diamidino-2-phenylindole) containing Mounting medium (immunoSelect® Dianova, Hamburg, Germany).

For data analysis, we focused on changes in Homer1a expression in the soma of dentate gyrus granule cells, given that this is the region from which electrophysiological recordings were obtained in both the in vitro patch clamp and the in vivo synaptic plasticity experiments. We looked for Homer1a mRNA expression within the nuclei of the granule cells by obtaining z-stacks at a 63x magnification using a Zeiss ApoTome (Figure 1E). Three consecutive slices of each animal were used for the analysis, whereby we analyzed one hemisphere of each slice and calculated the mean of these three slices. Using Fiji software (Schindelin et al., 2012) the complete DAPI stained nuclei that were not cut on the edges either in the x, y or z direction, were marked. Afterwards, they were checked for Homer1a mRNA expression that peak in the nuclei of the granule neurons and the percentages of the Homer1a mRNA positive cells were calculated per total counted neurons for each subregion of each rat. The designation “positive nuclei” was given to cells that contained intense intranuclear-foci of Homer1a mRNA fluorescent signals (Figure 1G, green dots). Nuclei that did not contain any intranuclear-foci representing a fluorescent signal of Homer1a mRNA were counted as negative. The total number of dentate gyrus granule cells analyzed for each brain slide of each animal ranged from 80 to 100 in the upper (suprapyramidal) blade and, from 70 to 90 in the lower (infrapyramidal) blade. Then the averaged values from both blades of the dentate gyrus were calculated for each animal. Final results are presented as average mean of percentages ± SEM for each group. The analysis was conducted without the experimenter knowing the identity of the different animal groups.

For the purpose of immunohistochemical analysis, another two groups of rats were treated intracerebrally with oligomeric Aβ(1–42) (n = 16), or control peptide (n = 16) and sacrificed either 1 week or 6 months following this procedure. Glia analysis was performed in a smaller group of animals (N = 16) as a pilot experiment. After transcardial perfusion with 4% paraformaldehyde (PFA), brains were carefully removed and further fixated in 4% PFA overnight, followed by cryoprotection in sucrose solution (30%) at 4°C. Immunostaining was performed on free-floating horizontal 30 μm sections. We used Anti-Aβ1-16 primary antibodies (1:400, mouse monoclonal AB, SIG-39320, Covance Inc., United States) to detect Aβ plaque deposition. As a positive control we tested the brains of 9 month old transgenic mice that express a mutant amyloid precursor protein (tgAPP) that is known to result in significant plaque expression in the brains of the animals at this age (Sasaguri et al., 2017).

Anti-glial fibrillary acidic protein (Anti-GFAP) primary antibodies (AB, 1:2000, polyclonal rabbit AB, Z0334, DakoCytomation Denmark A/S, Denmark) were used to assess differences in astroglia cell population (Yang and Wang, 2015). Anti-ionized calcium-binding adapter molecule 1 (Anti-Iba1) primary antibodies (1:500, polyclonal rabbit AB, #019–19,741, Wako Pure Chemical Industries, Japan) were used to investigate possible differences in microglia cell populations (Ito et al., 1998) and Anti-GluN1 primary antibodies (1:200, monoclonal mouse AB, 556308, BD Biosciences Pharmingen, United States) were used to determine N-methyl-D-aspartate receptor (NMDAR) expression. Sections from control- and Aβ-treated animals underwent masked randomization and were processed together in order to avoid variations in the processing of data sets.

Sections were treated with 0.3% H₂O₂ in PBS for 20 min in order to deactivate endogenous peroxidases. To reduce unspecific background staining, endogenous biotin and electrostatic loading proteins were blocked by 10% normal serum, 20% Avidin (Avidin/Biotin Blocking Kit SP-2001, Vector Labs, Burlingame, United States) in PBS containing 0.2% Triton X-100 (Tx, Sigma-Aldrich Chemie GmbH, Munich, Germany). Sections were then incubated overnight (12 to 20 h) in a solution containing 1% normal serum, 20% Biotin (Avidin/Biotin Blocking Kit SP-2001, Vector Labs, Burlingame, USA) and the respective primary antibody (AB) in PBS-Tx. Sections were then incubated in PBS-Tx solution containing 1% normal serum and the corresponding secondary AB (biotinylated goat-anti-rabbit BA-1000 or biotinylated horse-anti-mouse BA-2001, Vector Labs, Burlingame, United States) at a dilution of 1:500 for 90 min. To detect the antibody binding an ABC elite kit (Vectastain® Elite ABC Kit PK-6100, Vector Labs, Burlingame, USA) was applied at a dilution of 1:1000 for 90 min. The peroxidase binding was visualized by incubation in 0.05% 3,3’-Diaminobenzidine-solution (DAB, Sigma-Aldrich) with 0.01% H₂O₂ for 10 min.

To label Anti-Aβ1-16, Anti-GFAP and Anti-Iba1, the dilution medium contained 10% normal serum and 20% Avidin (Vector Labs, Burlingame, USA) diluted in PBS containing 0.2% Triton X-100 (Tx, Sigma-Aldrich, United States). To enhance the staining of GluN1, 1% bovine serum albumin (Sigma-Aldrich, United States) in PBS was used as dilution medium, sections were incubated in the primary AB solutions for 5 days at 4°C and the biotinylated tyramine method (as described by Adams, 1992) was applied.

After mounting on gelatin-coated microscope slides and dehydration, sections were cover-slipped using DPX (Sigma-Aldrich, St. Louis, United States). To exclude unspecific binding of secondary AB and the detection system, negative controls were performed in the same staining protocol by omitting the primary AB. No staining could be observed in these negative controls, indicating that the observed staining was specific.

To assess for changes in protein expression, horizontal sections obtained from −4.6 mm and − 7.34 mm relative to bregma were segregated into regions of interest (ROIs). The ROIs comprised the lateral and medial EC (LEC and MEC), the dentate gyrus (DG) with its granule cell layer (GCL), outer and inner molecular layers (oML and iML), the CA1 and CA3 regions and the subiculum (SB).

Statistical analysis was performed using the STATISTICA software version 14.0.1.25, TIBCO Software Inc., Santa Clara, CA, United States. For in situ hybridization data, a one-way analysis of variance (ANOVA) was performed, followed by a Tukey HSD post-hoc test for pairwise comparison (naïve vs. control vs. Aβ). Electrophysiological results were assessed using a multifactorial ANOVA with repeated measures. Behavioral performance and immunohistochemistry results were assessed using the Student’s t-test. The level of significance was set at p < 0.05. All data were shown as mean ± standard error of mean.

One month after intracerebral treatment with oligomeric Aβ (1–42) (n = 7), we assessed the robustness of LTP in the dentate gyrus of freely behaving rats. After measuring basal synaptic transmission for 1 h, high frequency stimulation (HFS) was applied to the perforant path. This protocol induced long-term potentiation that lasted for over 24 h in control animals (Figure 1A).

By contrast, LTP was significantly reduced in Aβ–treated animals compared to controls (n = 7) (Figure 1A): Whereas the initial 15 min of LTP were equivalent in both animal cohorts, deficits became more pronounced from 30 min onwards. Despite the impairments LTP was still present in Aβ-treated animals 24 h after HFS. (ANOVA with repeated measures applied to all time-points recorded after HFS: F (1,12) = 7.33, p < 0.05).

Examination of LTP in the same animals 6 months after treatment revealed that the LTP impairment had become much more pronounced in Aβ-treated animals (Figure 1B). Now both short-term potentiation and LTP were severely impaired compared to responses evoked in control animals. Evoked potentials had returned to pre-HFS levels by 4 h after stimulation. [ANOVA: F(1, 11) = 10.53, p < 0.01].

To test for cognitive impairments, animals engaged in a recognition and a spatial memory task. We observed that 1 month after intracerebral treatment with oligomeric Aβ (1–42), animals (n = 15) successfully recognized the new object (t-test, p < 0.05) with a performance level that was equivalent to controls (t-test, n = 15, p < 0.05) (Figure 1Di). When Aβ–treated animals (n = 13) were faced with cognitively more challenging item-place task, memory deficits were evident: only the control animals (n = 14) successfully noticed that a familiar object had been moved to a new position (t-test, p < 0.05) (Figure 1Ci).

By 6 months after treatment, Aβ–treated animals were impaired in both the item-place task (n = 12) (Figure 1Cii) and the object recognition task (n = 11) (Figure 1Dii). By contrast, control animals exhibited effective memory performance in the item-place task (t-test, n = 10, p < 0.05) (Figure 1Cii) and the object recognition task (t-test, n = 14, p < 0.05) (Figure 1Dii).

To assess to what extent the memory deficits were caused by changes in information processing in the hippocampus we examined nuclear immediate early gene expression in the hippocampus following item-place learning (Figures 1E–G). The object interactions (not shown) revealed the same outcome as reported in Figure 1Ci [ANOVA F(1,16) = 15,93,088, p = 0.001; Tukey HSD post-hoc test: control: new vs. old: p = 0.0147; Aβ: new vs. old: p = 0.1762]. The examination of Homer1a expression was conducted in the suprapyramidal and infrapyramidal blades of the dentate gyrus. The average percentage somatic expression of Homer1a mRNA for both dentate gyrus blades was calculated. The low somatic expression level of Homer1a mRNA in the granule cells of naïve animals (no surgery, no exploration) fits very well with reports from other studies (Vazdarjanova et al. 2002; Hoang et al., 2018). Both groups of animals (n = 5 each) that explored the item-place configuration exhibited a significant increase in Homer1a mRNA expression, compared to naïve animals (n = 6) (Figure 1F, one-way ANOVA F(2,13) = 30.533, p = 0.0000, Tukey HSD post-hoc test: control vs. naïve: p = 0.0001, Aβ vs. naïve: p = 0.0015). This result is in line with observations from other studies reporting that novel spatial learning increases somatic Homer1a expression in the hippocampus (Vazdarjanova et al. 2002; Clifton et al. 2017; Hoang et al., 2018, 2021, 2023). Strikingly, one month after Aβ–treatment, animals (n = 5) showed significantly reduced somatic Homer1a expression in the dentate gyrus following the learning task, compared to control animals (n = 5) (Figure 1F, Tukey HSD post-hoc test, control vs. Aβ: p = 0.0271). This result aligns with the abovementioned behavioral data and indicates that oligomeric Aβ(1–42) causes impairments of experience-dependent information encoding 1 month after intracerebral treatment.

Given that we identified deficits in synaptic plasticity and nuclear expression of Homer1a following Aβ(1–42)-treatment, we explored whether physiological properties of dentate gyrus granule cells were altered. Patch clamp recordings from granule cells revealed no differences in firing frequency when we examined responses one week after Aβ(1–42)-treatment (Aβ: N = 8, n = 18; control: N = 6, n = 15, ANOVA: F(1, 248) = 0.283, p = 0.595) (Figure 2A and S2, top chart), but 4–6 months after treatment, firing frequency was significantly reduced in granule cells from animals that had been treated with Aβ compared to ScrAβ(1–42)- treated controls (Figure 2A and S2, bottom chart) [ANOVA: F(1, 208) = 32.38, p < 0.001, Aβ: N = 8, n = 16; control: N = 7, n = 12, Duncan’s test, *, p < 0.05].

Resting membrane potential was significantly more positive in cells from Aβ-treated animals 1 week after treatment (N = 8, n = 18) (Figure 2B, top chart), compared to controls (N = 6, n = 15) [ANOVA, F(1,31) = 9.128, p = 0.005] (top chart), whereas 4–6 months after Aβ–treatment it became significantly more negative (Figure 2B, bottom chart) [ANOVA, F(1,26) = 8.173, p = 0.008] (bottom chart) (Aβ: N = 8, n = 16; Controls: N = 7, n = 12).

The width of the action potential (AP) at 20% of its amplitude was not significantly changed 1 week after Aβ–treatment (Figure 2C, top chart) (ANOVA, F(1,31) = 3.227, p = 0.082) but was significantly increased after 4–6 months (Figure 2C, bottom chart) [ANOVA, F(1,26) = 4.806, p = 0.037] (bottom chart) (Aβ: N = 6, n = 16; Controls: N = 7, n = 12).

Afterhyperpolarization (AHP) depth was significantly reduced in cells from Aβ-treated animals 1 week after treatment (Figure 2D, top chart) [ANOVA, F(1,31) = 5.484, p = 0.025] (Aβ: N = 8, n = 18; Controls: N = 6, n = 15), but not significantly changed 4–6 months after treatment [ANOVA, F(1,26) = 0.184, p = 0.670) (Figure 2D, bottom chart].

The latency from AP peak to AHP minimum was unchanged 1 week after Aβ-treatment (Figure 2E, top chart) [ANOVA, F(1,31) = 2.071, p = 0.160] (top chart), but 4–6 months after treatment (bottom chart), the latency was significantly increased (Figure 2E, bottom chart). [ANOVA, F(1,26) = 11.870, p = 0.0019 (Aβ: N = 6, n = 16; Control: N = 7, n = 12), ANOVA: **, p < 0.01; *, p < 0.05].

Taken together, these data suggest that early, but subtle increases in membrane excitability are succeeded months after treatment by a reduction in neural responsiveness.

In the current study, immunohistochemistry assessments did not detect an increase in the number of Aβ-plaques 6 months after Aβ-treatment (n = 8) compared to control animals (n = 7) (Figure 3A) [t-test: T(13) = 0.242, p = 0.812]. To verify that this result was not due to problems with the antibody used, we assessed for plaque expression in a transgenic mouse model of AD (TgAPP, 9 month old mice) and were able to detect plaques (Figure 3A).

We scrutinized the hippocampus, the lateral and medial entorhinal cortex, and the subiculum and discovered that in 6 month old control animals a small number of plaques are intrinsically present (Figure 3A). Levels in Aβ-treated animals did not differ from levels detected in controls, however.

We then assessed Nissl stained brain slices to examine for cell damage in the hippocampal CA1 region, the dentate gyrus, the subiculum, the lateral entorhinal cortex, or the medial entorhinal cortex (Figures 3B,C). We found no evidence of increased cytoxicity in Aβ-animals (n = 8), 6 months after treatment, compared to controls (n = 7) (Table 1).

A role for microglia in AD progression has been proposed (Edwards, 2019). Here, we used immunohistochemical analysis of Iba1 expression to assess to what extent microglia are activated in the hippocampal CA1 region, the dentate gyrus, the subiculum, the lateral entorhinal cortex and the medial entorhinal cortex at early (1 week) or late (6 month) latencies after Aβ-treatment (Figure 4A). We found that Iba1 expression was increased in the inner molecular layer of the dentate gyrus, the subiculum, the medial entorhinal cortex 1 week after Aβ-treatment (n = 4), compared to controls (n = 4) (Table 2). By contrast, 6 months after treatment Iba1 expression levels were equivalent in Aβ-treated (n = 4) and control animals (n = 4) (Table 3).

To examine to what extent astrocytes might be affected by Aβ-treatment we assessed GFAP levels 1 week and 6 months after treatment (Figures 4B,C) using densitometrical analysis (Tables 4, 5) and an analysis of the percentage of area covered by GFAP positive astrocytes (Tables 6, 7). GFAP is an astroglia cytoskeleton-specific protein which is upregulated in response to reactive astrogliosis (Jurga et al., 2021). We detected reconstruction in the morphology of GFAP-positive cells in Aβ-treated animals, compared to control animals (Figure 4C, photomicrographs). The astrocytes in Aβ-treated animals exhibited a reduction in their complexity (i.e., in arborisation and surface area) compared to control animals. Densitometrical analysis revealed no changes in GFAP expression levels in Aβ-treated (n = 4) and control animals (n = 4) 1 week after treatment (Table 4), whereas the area analysis revealed significant reductions of astrocyte density (Table 5). Six months after treatment, Aβ-treated animals (n = 4) exhibited a significant reduction of GFAP expression in the granule cell, the outer and inner molecular layer of the dentate gyrus and the CA3 region compared to controls (n = 4) in the densitometrical analysis (Table 6), and in the outer molecular layer of the dentate gyrus and the CA3 region in the area analysis (Table 7). Our observations are in line with findings by others, that suggest that a reduction in GFAP positive astrocytes is correlated with changes in astroglial morphology, which occurs during pathological states in the brain (Senitz et al., 1995; Chvátal et al., 2007a,b).

Taken together, these data suggest that an early and transient activation of microglia is followed by a change in morphology of astrocytes following Aβ-treatment.

Changes in NMDAR function have been proposed to contribute to the impairments of LTP seen in various animal models of AD (Rammes et al., 2018; Yang et al., 2018). Here, we investigated the expression of the GluN1 subunit of the NMDAR in the CA1 region, the dentate gyrus, the subiculum, the lateral entorhinal cortex and the medial entorhinal cortex (Figure 5). No changes in receptor expression were detected 1 week after treatment when effects were compared in Aβ-treated (n = 8) and control animals (n = 7) (Table 8). However, 6 months after treatment we detected a significant decrease in GluN1 expression in the granule cell and the inner molecular layers of the dentate gyrus, as well as in the subiculum and the CA1 regions of Aβ-treated animals (n = 8), compared to controls (n = 7) (Table 9).