Introduction
Alzheimer’s disease (AD) is so far an incurable, progressive degenerative brain disorder characterized by the presence of extracellular amyloid-beta (Abeta) plaques and intracellular aggregated phosphorylated tau, together with synaptic and neuronal cell loss. Neuroinflammation has been postulated to be a critical factor in the pathogenesis of AD [
16,
17,
19]. The microglial activation process is characterized by remarkable morphological changes (larger cell body with shorter, thicker, and less branched processes) and up-regulation of proinflammatory and/or anti-inflammatory cytokines [
13,
18,
44]. Activated microglia accumulate around Abeta plaques in both transgenic models [
18,
22,
44] and AD patients [
49,
50]. Although the inflammatory response could contribute to Abeta phagocytosis [
11,
20], it has been associated with a neurotoxic detrimental effect mediated by the release of proinflammatory cytokines/chemokines and neurotoxins [
16,
17,
19].
The inflammatory response in the AD brain is probably not exclusively detrimental or beneficial. In fact, recent human genome-wide association studies (GWAS) have identified multiple polymorphisms associated with the microglial immune response in AD [
14,
24]. One such polymorphism, R47H, is located in TREM2 and is associated with an increased risk for late-onset AD [
26]. TREM2 is a lipid sensor that, through its adapter molecule DAP12, supports Abeta-reactive microgliosis and Abeta clearance [
63]. In the absence of TREM2, microglial activation is impaired. In the 5xFAD mouse model, TREM2 deficiency increased Abeta accumulation due to a dysfunctional microglial response and microglia was apoptotic instead of activated [
63]. Thus, a deficient rather than an overactive microglial response could, indeed, be associated with AD development [
54].
The microglial response has been preferentially studied in AD brain areas with relatively high Abeta content or in Abeta-rich transgenic models [
16,
22,
49‐
51]. However, brain regions particularly relevant in AD development, such as the hippocampal formation, present low Abeta accumulation and a high number of phospho-tau-bearing neurons [
4]. Thus, the microglial reaction in the hippocampus could be totally different than that previously described in neocortical areas. In the present work, we evaluated the microglial response in postmortem hippocampal human tissue pathologically diagnosed in Braak II to Braak V–VI stages. Contrary to the marked microglial activation reported in APP-based mice models and in human neocortical AD regions, we demonstrated a prominent degenerative process of the microglia in the dentate gyrus (DG) and CA3 of Braak V–VI samples, probably associated with the accumulation of soluble phospho-tau, as determined by in vitro assays. This degenerative process left most of the affected regions with no immunological coverage and could contribute to AD pathology progression.
Materials and methods
Human samples
Human autopsy specimens from the medial temporal lobe (hippocampal/parahippocampal regions) were obtained from the tissue bank Fundación CIEN (BT-CIEN; Centro de Investigación de Enfermedades Neurologicas; Madrid, Spain) and from the Neurological Tissue Bank of IDIBELL-Hospital of Bellvitge (Barcelona, Spain). The utilization of postmortem human samples was approved by the corresponding biobank ethics committees and by the “Comite de Etica de la Investigacion (CEI), Hospital Virgen del Rocio,” Seville, Spain. All cases were scored for Braak tau pathology. Table
1 summarizes the demographics of the human samples. Only Braak V–VI cases were clinically classified as demented (AD) patients. We considered the age-matched Braak II individuals as controls in our experiments.
Table 1
Human sample information
(a) Unfixed frozen samples |
0 (n = 8) | 62.5 | 37.5 | 49 ± 6 | 8 ± 5 |
II (n = 13) | 61.54 | 38.46 | 78 ± 8.5 | 7 ± 4 |
III–IV (n = 9) | 44.44 | 55.56 | 80 ± 11 | 6 ± 5 |
V–VI (n = 18) | 38.89 | 61.11 | 79 ± 10 | 8 ± 4 |
(b) Fixed samples |
0 (n = 4) | 50 | 50 | 53 ± 16 | 7 ± 5 |
II (n = 7) | 28.6 | 71.4 | 78 ± 8.5 | 6 ± 3 |
III–IV (n = 10) | 50 | 50 | 79 ± 7 | 10 ± 5 |
V–VI (n = 16) | 43.8 | 56.20 | 78 ± 10 | 9 ± 5 |
For morphological studies, 4 % paraformaldehyde fixed samples were sectioned (30 µm thickness) on a freezing microtome. For molecular characterization, unfixed frozen samples were used. The anatomical boundaries of the hippocampal system and parahippocampal gyrus were identified by the anatomical and cytoarchitectonic features in sections stained with cresyl violet using a human stereotaxic brain atlas [
30]. Areas between the bregma coordinates at 17 and 35 mm were analyzed.
Antibodies
The following primary antibodies were used: (1) monoclonals, anti-phospho-tau AT100 (pSer212/Thr214) and AT8 (pSer202/Thr205, Thermo Fisher Scientific, New York, USA), anti-total tau (tau46, Cell Signaling Technology, Massachusetts, USA), 4G8 (anti-Abeta 17-24, Biolegend, San Diego, USA), 6E10 (anti-Abeta1-16, Covance, Princeton, USA), 82E1 (anti-Abeta-N-terminal, IBL, Hamburg, Germany), and anti-CD68 (PG-M1, Dako, Glostrup, Denmark); (2) rabbit polyclonals, anti-amyloid fibrils (OC, Merck Millipore, Darmstadt, Germany), anti-CD45 (Abcam, Cambridge, United Kingdom), anti-Iba1 (Wako Pure Chemical Industries, Osaka, Japan), and anti-P2ry12 (Sigma-Aldrich, Saint Louis, USA).
Immunohistochemistry
Sections from control and diseased brains were assayed simultaneously using the same batches of solutions to minimize variability in immunolabeling conditions. After antigen retrieval (80 °C for 20 min in 50 mM citrate buffer, pH 6.0), endogenous peroxidase was inhibited (3 % H2O2/10 % methanol in PBS, pH 7.4 for 20 min) and non-specific staining was avoided using 5 % goat or horse serum (Sigma-Aldrich) in PBS. For single labeling light microscopy, sections were incubated with the primary antibody (24–72 h, at room temperature) followed by the corresponding biotinylated secondary antibody (1:500 dilution, 1 h at room temperature, Vector Laboratories), streptavidin-conjugated horseradish peroxidase (1:2000, 90 min, Sigma-Aldrich), and visualized with 0.05 % 3-3-diaminobenzidine tetrahydrochloride (DAB, Sigma-Aldrich) and 0.01 % hydrogen peroxide in PBS. The specificity of the immune reactions was controlled by omitting the primary antisera. For double or triple immunofluorescence labeling, sections were sequentially incubated with the indicated primary antibodies followed by the corresponding Alexa 488/568/405 secondary antibodies (1:1000 dilution, Invitrogen). AT100-immunolabeled sections were stained with 0.1 % thioflavin-S (Sigma-Aldrich) in 70º ethanol for 10 min. Sections were incubated in an autofluorescence eliminator reagent (Merck Millipore) following the manufacturer’s recommendations and examined under a confocal laser microscope (Leica SP5 II).
Stereological analysis
The number of Iba1-immunopositive microglial cells was stereologically quantified according to the optical fractionator method [
43] using an Olympus BX61 microscope equipped with the NewCAST software package (Olympus, Denmark). Cell counting was performed on every 6th section (with a distance of 180 µm) through the rostrocaudal axis of the hilar region (
n = 5–6 sections/individual). The dentate gyrus boundaries were defined using a 2× objective. Cell number was counted using a 100×/1.35 objective. We used a counting frame of 1722 µm
2 with step lengths of 131.23 µm. The numerical density (cells/mm
3) was estimated as:
\({\text{ND}} = {Q \mathord{\left/ {\vphantom {Q {\left( {\sum {A \times h} } \right)}}} \right. \kern-0pt} {\left( {\sum {A \times h} } \right)}}\), where ‘
Q’ is the number of disector-counted somatic profiles, ‘
\(\sum A\)’ is the area of the counting frame, and ‘
h’ is the section thickness (30 µm).
Image analysis
Microglial loading was defined as the percentage of area stained with anti-Iba1 (total microglia) or anti-P2ry12 (non-activated microglia) in relation to the total area analyzed. Digital images (2 sections/individual) were processed using the Visilog 6.3 image analysis system (Noesis, France). The Iba1- or P2ry12-immunopositive signal within the selected brain region was converted into 8-bit gray scale, and immunostained cells were identified by a threshold level mask. A fixed threshold level (ranging 160–180) was maintained throughout the image analysis of all sections from the same individual brain for uniformity.
Microglial spatial distribution was determined by the grid analysis [
3]. The dentate gyrus was photographed and processed with the Visilog 6.3 program. Digital images (4×, 2 sections/individual,
n = 5–8 individuals per group) were binarized, and grid analysis was performed by placing a grid (354 × 354 μm/square) of 125,000 μm
2 squares. The hilar region was divided into 30–50 regular squares, covering the total parenchymal space. The microglial spatial distribution was then estimated by measuring the Iba1-positive coverage area in each square of the grid.
Microglial domain was defined as the area (µm
2) covered by a single P2ry12-positive cell. This area was calculated by drawing a polygon connecting the distal end of total branches of the cell (see Fig.
3c). Microglia cells (20–30 cells per section) from Braak II and Braak V–VI sections (2 sections/individual,
n = 5–7 individuals) were randomly selected using stereology-based sampling (NewCAST software from Olympus) and imaged under a 40× objective. The optical disector was set at 7118.3 µm
2 with step lengths of 238.64 µm. Only microglial cells with a visible cell body were photographed. Finally, high-resolution images were processed using the Visilog 6.3 program, and the P2ry12-positive cell domain area was measured.
Transgenic animals
Animal experiments were performed in accordance with the Spanish and the European Union regulations (RD53/2013 and 2010/63/UE) with the approval of the Committees of Animal Research from the Universities of Seville (Spain) and Malaga (Spain). APP/PS1 and Thy-tau22 transgenic animals were used. The APP/PS1mice [
22,
58] over-expressed the human mutant PS1M146L and human APP751 carrying the Swedish (KM670/671NL) and London (V717I) mutations. The Thy-tau22 mice expressed human 4-repeat tau with G272 V and P301S mutations [
48]. Non-transgenic mice (WT) of the same genetic background (C57/BL6) were used as controls. In this work, we used 2-, 6-, 9- and 12-month-old APP/PS1 mice; 2-, 9-, 12- and 16-month-old Thy-tau22; and age-matched WT mice (
n = 5/age and genotype). Mice were killed (sodium pentobarbital, 60 mg/kg) and processed as described [
22,
58]. Fixed brains were serially sectioned (40 µm coronal sections) and assayed for immunohistochemistry. Microglial (Iba1-positive) loading and microglial domain were calculated as described above (7–9 sections/mouse;
n = 4/age/genotype).
Preparation of soluble S1 fractions and sequential protein extraction
Soluble S1 fractions from either mouse models or human samples were prepared as described [
23]. Briefly, human or mouse tissue was homogenized (Dounce homogenizer) in TBS (20 mM Tris–HCl, 140 mM NaCl, pH 7.5) containing protease and phosphatase inhibitors (Roche). Homogenates were ultracentrifuged (4 °C for 60 min) at 100,000×
g (Optima MAX Preparative Ultracentrifuge, Beckman Coulter). Supernatants, S1 fractions, were aliquoted and stored at −80 °C. The pellets (P1) were extracted in RIPA buffer (1 % CHAPS, 1 % Na-deoxycholate, 0.2 % SDS, 140 mM NaCl, 10 mM Tris–HCl, pH 7.4); ultracentrifuged and supernatants, S2 fractions (intracellular particulate proteins), were aliquoted and stored. Pellets (P2) were re-extracted in buffered-SDS (2 % SDS in 20 mM Tris–HCl, pH 7.4, 140 mM NaCl), centrifuged as above and supernatants, S3 (SDS releasable proteins) were stored. Finally, the remaining pellets were extracted in SDS-urea (20 mM Tris–HCl, pH 7.4, 4 % SDS and 8 M urea).
Sarkosyl-insoluble fraction isolation
Sarkosyl-insoluble tau was isolated as described [
29]. Human hippocampi were homogenized in “homogenization buffer” (10 mM Tris, 0.8 M NaCl, 1 mM EGTA, 10 % sucrose, pH 7.4, plus protease, and phosphatase inhibitors). After centrifugation (5,000×
g, 4 °C, 15 min), supernatants were incubated (2.5 h at 37 °C in agitation) with 1 % Sarkosyl (Sigma-Aldrich) and 1 % beta-mercaptoethanol (Sigma-Aldrich) in “homogenization buffer.” Samples were then ultracentrifuged (100,000×
g, 4 °C, 30 min). Supernatants constituted the Sarkosyl soluble fractions, whereas pellets were the Sarkosyl-insoluble fraction. The insoluble fraction was rinsed twice and resuspended in TBS.
Abeta and Tau quantification by sandwich ELISA
Total soluble Abeta or tau in S1 fractions was determined using a commercial sandwich ELISA (human Abeta x-40, Invitrogen; Abeta x-42, DRG; human Tau, Invitrogen) following the manufacturer’s recommendations. For each assay, 25 μg of protein from the pooled-soluble fractions were used. The ELISA experiments were repeated four times in independent experiments using triplicate replicas.
Total RNA and protein extraction
Total RNA and proteins were extracted using TriPure Isolation Reagent (Roche) [
22]. RNA integrity (RIN) was determined by RNA Nano 6000 (Agilent). Although no differences between Braak groups were observed, the RIN was lower in human samples compared with transgenic models (RIN: 4.95 ± 1.4 or 8.5 ± 0.5 for human and mouse samples, respectively). RNA was quantified using NanoDrop 2000 spectrophotometer (Thermo Fischer). Proteins were quantified using Lowry’s method.
Retrotranscription and quantitative real-time RT-PCR
Retrotranscription (RT) (4 μg of total RNA) was performed with the High-Capacity cDNA Archive Kit (Applied Biosystems). For real-time qPCR, 40 ng of cDNA were mixed with 2× Taqman Universal Master Mix (Applied Biosystems) and 20× Taqman Gene Expression assay probes (Applied Biosystems, supplemental Table 1). Quantitative PCR reactions (qPCR) were done using an ABI Prism 7900HT (Applied Biosystems). The cDNA levels were determined using GAPDH and beta-actin. We observed a highly significant linear correlation between the cycle threshold (Ct) of both genes (beta-actin vs GAPDH, r = 0.912, F(1,46) = 157.73, p < 0.0001). Thus, normalization using either beta-actin or GAPDH produced identical results. Routinely, we used GAPDH as housekeeper. Results were expressed using the comparative double-delta Ct method (2-ΔΔCt). ΔCt values represent GAPDH normalized expression levels. ΔΔCt was calculated using Braak 0 for human samples or 9-month-old WT mice for transgenic models.
Western blots
Western blots were performed as previously described [
22,
23]. Abeta peptides were detected after 16 % SDS-Tris-Tricine-PAGE using PVDF membranes (Immobilon-P Transfer Membrane, Millipore) and a mixture of 6E10 (1/6000) and 82E1 (1/5000) antibodies. For Tau (Tau46, AT8 or AT100, 1/1000), proteins were loaded on 4–20 % SDS-Tris–Glycine-PAGE (Bio-Rad) and transferred to nitrocellulose (Optitran, GE Healthcare Life Sciences). Dot blots using OC were performed as previously described [
22].
Cell cultures
BV2 microglial cells were grown (37 °C and 5 % CO2) in RPMI 1640, 2 mM glutamine, 10 % (v/v) fetal bovine serum, plus penicillin/streptomycin (all from Biowest). SH-SY5Y neuronal cells (SH-control cells) were grown in DMEM-F12, 2 mM glutamine, 10 % fetal bovine serum, penicillin/streptomycin, and 1 % non-essential amino acids (Biowest). SH-SY5Y-Tau (SH-tau cells, see [
8]) stably transfected neuronal cells (expressing 3R human tau) were selected using G418 (0.2 µg/ml, Biowest). For co-culture experiments, BV2 cells were plated (15.000 cells/cm
2, 12 h) and SH-SY5Y cells were added at a ratio of 1/2.5 (SH-SY5Y/BV2 cells). Cells were co-cultured for 12 h in RPMI 1640. After co-culture, the cells were detached by trypsin and analyzed by flow cytometry. Apoptosis of SH-SY5Y cells (either control or tau-expressing cells) was induced using 1 µg/ml staurosporine B (Sigma-Aldrich) for 2 h [
25]. This short treatment produced 72 ± 5.7 % of apoptotic SH-SY5Y cells. The absence of any direct effect of staurosporine B on BV2 cells was analyzed by omitting the SH-cells. Primary murine microglial or astroglial cultures were performed as previously described [
22].
BV2 treatment with S1 soluble fractions
BV2 cells (15.000 cells/cm2) were serum deprived for 12 h and then treated (0.1 µg soluble protein/µl of culture medium) for 12 h with soluble S1 fractions (from Braak 0 to V–VI or transgenic mice models). The effect on the cell survival was assessed by flow cytometry.
Flow cytometry analysis of cell viability and apoptosis
The viability and apoptosis of BV2 cells were assessed using the apoptosis detection kit Annexin V-FITC (Immunostep) following the manufacturer’s specifications and were analyzed using a FACSCanto II flow cytometer (BD Services, San Jose, CA, USA). The pro-apoptotic effect of SH-SY5Y cells on BV2 cultures was evaluated by double staining the cells with CD45-PE (BV2) and Annexin V-FITC (apoptosis). Control experiments using single cell cultures (either viable or apoptotic BV2 or SH-SY5Y) were conducted to determine the discrimination settings (CD45-positive (BV2) and -negative (SH-SY5Y) cells). With this protocol, 86 ± 2.3 % of viable BV2 cells were discriminated from SH-SY5Y cells. Control experiments also demonstrated that a fraction (16 ± 4.3 %) of apoptotic BV2 cells displayed a reduction in the expression of CD45 and cannot be distinguished from SH-SY5Y cells.
BV2 and SH-SY5Y phagocytosis assay (TAMRA assay)
Phagocytosis was assessed by loading SH-SY5Y or SH-SY5YTau cells with 5 µM TAMRA-ester (5-(and-6)-carboxytetramethylrhodamine succinimidyl ester, Sigma-Aldrich) [
12]. TAMRA-loaded SH-cells were then treated (2 h) with staurosporine (1 µg/ml of culture medium; apoptotic cells) or PBS (control cells) and co-cultured with BV2 microglia (1/2.5 SH- vs BV2). Phagocytic BV2 cells were identified by CD45-PE and TAMRA fluorescence. For the inhibition of phagocytosis, cytochalasin B (5 μg/ml) was added to culture 30 min prior the experiment and maintained throughout the assay.
Statistical analysis
Normality of data was assessed using the Kolmogorov–Smirnov test. Normally distributed data were expressed as the mean ± SD. Non-normally distributed data were represented using box-plots (Sigmaplot) or scatter-plots with the median and interquartile range (GraphPad). For normally distributed data, mean values were compared using ANOVA followed by Tukey’s test (more than two groups) or two-tailed t test (for two group comparisons). The data which were not normally distributed were compared by the Mann–Whitney U test (for two groups comparisons) or Kruskal–Wallis tests (more than two groups) followed by Dunn’s post-hoc test. The significance was set at 95 % of confidence. Linear correlations were analyzed using the Kendall tau-beta test. In all the cases, IBM SPSS (v23) Statistics software was used.
Discussion
The role of the inflammatory response in Alzheimer’s disease is far from elucidated. In this study, we evaluated the microglial response in the hippocampus of postmortem human samples classified from Braak 0 to Braak VI stages. Contrary to the expected results based on APP-transgenic models [
16,
22,
33], an attenuated rather than massive microglial activation was observed in Braak V–VI samples (AD cases). More importantly, the present results also demonstrate the existence of a microglial degenerative process in the Braak V–VI hippocampus. This degenerative process was more prominent in the hilar region of the dentate gyrus and the CA3 region of the hippocampus proper. The microglial pathology is characterized by the following: (1) the presence of degenerative microglial cells, which exhibit shortened and less branched processes, cytoplasmic fragmentation (cytorrhexis) and spheroids formation (see also [
54,
57]); (2) a reduction (at least in 4 of 9 Braak V–VI cases tested) in the microglial numerical density; and (3) a dramatic decrease in the area of surveillance of individual microglial cells, described here as the microglial domain. These pathological modifications produce a prominent decrease in the parenchymal area covered by microglia in these particular hippocampal subfields (DG and CA3), leaving most of the parenchymal space with no immune coverage, including Abeta plaques and vascular Abeta depositions.
It could be argued that this microglial degenerative process reflects the advanced pathological status of demented Braak V–VI individuals. If this is the case, we speculate that non-demented cases with lower pathology (Abeta and/or tau accumulation), such as Braak II or Braak III–IV samples, could better reflect the early microglial response [
47]. However, microglial cells from Braak II individuals presented a highly ramified morphology with a regular spatial distribution, covered the total parenchymal space, and displayed absolutely no signs of activation or degeneration. Furthermore, neither molecular nor morphological evaluation of the Braak III–IV samples indicated the existence of microglial activation outside of that restricted to Abeta plaques or vascular deposits. Concerning microglial degeneration, Braak III–IV samples were highly heterogeneous. Approximately 50 % of the tested Braak III–IV population displayed similar morphological and molecular parameters to that of the Braak II population, whereas the other 50 % better resembled the Braak V–VI cases (see Figs.
2,
4). We do not know what determines this heterogeneity. However, the existence of a highly significant correlation between two different microglial markers, P2ry12 and Iba1, in both the DG and CA3 subfields from samples of different Braak pathology (from Braak II to Braak V–VI individuals) strongly suggests that the microglial degeneration in Braak V–VI patients, at these particular hippocampal areas, could be part of a continuum along with the progression of the AD.
As mentioned above, microglial degeneration was particularly evident in the hilus of the dentate gyrus and in the CA3 subfield. Interestingly, these two hippocampal areas also displayed the highest microglial loading compared with other hippocampal subfields in Braak II individuals. Although we do not know what determines this microglial regionalization, it is obvious that the higher microglial load in Braak II samples and the microglial degeneration in AD patients seem to be associated with the DG-CA3 connection by the mossy fibers. In this sense, recent studies have suggested the existence of hippocampal hyperexcitability in both AD patients [
2] and transgenic models [
40,
60]. This hyperactive status is particularly evident at the level of DG to CA3 connection. Although we cannot conclude that a hyperactive status of the mossy fibers is involved in the microglial degeneration observed in AD patients based on the present data, this idea could be a potential explanation, and further investigation is clearly needed.
The implication of microglial cells in the development of neurodegenerative disorders is generally accepted [
16,
19,
59]. However, microglial dysfunction in AD has been primarily associated with over-activation and cytotoxicity of these cells [
19,
59]. Microglial activation has been undoubtedly observed in cerebral regions with the early and abundant extracellular Abeta deposits [
15,
36,
49,
50]. In fact, microgliosis has been associated with the disease duration and the neurodegenerative progression of AD [
49‐
51]. Furthermore, in the APP-based transgenic models, microglial activation may drive the AD pathology. In this sense, increases in microglial reactivity [
32] or, in contrast, pharmacological depletion of microglial cells [
32,
36,
53,
63] clearly enhanced or reduced Alzheimer’s disease pathology, respectively, with consequent changes in tau phosphorylation, synaptic strength, and neuronal cell lost. Therefore, the data presented here seem to contradict those previously reported. The most parsimonious explanation to reconcile these, apparently, contradictory observations is the different Abeta and/or tau content between different brain regions in AD patients and/or animal models. Most of these observations are based on Abeta producing mice, similar to our APP/PS1 model, or in Abeta-rich cortical areas from AD patients. However, the hippocampus of AD patients exhibits low and late Abeta pathology, whereas phospho-tau accumulates starting in the early stages of the disease [
4]. Thus, the preferential accumulation of phospho-tau over Abeta plaques could induce a totally different microglial response.
The pathological consequence(s) of a deficient immunological protection due to the microglial degeneration observed in AD patients are unknown. Microglial cells are implicated in the maintenance of synaptic integrity [
62] and, in fact, could promote learning-dependent synaptic formation [
41]. In addition, microglia is involved in Abeta phagocytosis [
27,
64,
65], senile plaque compaction, and limitation of Abeta toxicity [
7,
66]. Furthermore, microglia play a relevant role in removing damaged neurons and neuronal components, such as aberrant synaptic terminals or demyelinated axons. In this sense, deficiencies in key genes for microglial survival and/or proliferation (such as CSF1R or TREM2) are associated with rare hereditary neurodegenerative diseases, such as adult-onset leukoencephalopathy with axonal spheroids or Nasu–Hakola disease (respectively) [
6,
38,
39]. In both diseases, the microglial response and, more relevant, the microglial survival seem to be compromised. Moreover, the rs75932628 polymorphism results in an R47H missense mutation in TREM2 and increases the risk for late-onset AD [
14,
26,
37,
65]. Furthermore, TREM2 knock-out models display dystrophic microglial cells similar to that described in this work [
42], deficiencies in microglial survival, and aggravation in AD pathology [
63,
65]. Although the presence of any of these rare TREM2 variants in our limited AD cohort is highly improbable, these studies together with our present data strongly suggest that microglial pathology, with the consequent deficient immunoprotection in relatively large areas of the hippocampus, such as the dentate gyrus and CA3, might, indeed, contribute to the progression of AD pathology and cognitive impairment.
Another relevant finding of our work is the demonstration of the toxic effect of soluble intra- and/or extracellular phospho-tau on microglial cells by in vitro assays, which strongly supports the hypothesis that phosphorylated tau is the putative toxic agent for microglia in the AD hippocampus. Furthermore, our in vitro experiments are also in concordance with the differential microglial response to the Abeta or phospho-tau pathology observed in transgenic models and with the neuroprotection observed in tau-deficient models [
31,
45]. However, we are also aware that the microglial pathology in the tau model was modest and less obvious than what was observed in the Braak V–VI samples. We do not know why this apparent discrepancy exists, and it clearly indicates that AD pathology is more complex than what is reproduced in mouse models.
We do not know which form(s) of phospho-tau (monomeric vs oligomeric) exerts the toxic effect on microglial cells. However, it is important to emphasize that not all phospho-tau species present in AD samples are toxic for microglial cells, as demonstrated using Sarkosyl-insoluble phospho-tau. It has long been recognized that Sarkosyl-insoluble phospho-tau is part of the intracellular paired helical filaments that constitute the basis for neurofibrillary tangle formation [
28]. Therefore, as proposed by others, tangle formation could represent a cellular protection mechanism that prevents neuronal toxicity [
46] and, as probed in this work, microglial toxicity. The cause of the accumulation of soluble phospho-tau species in AD patients is currently unknown.
Another open question is whether soluble phospho-tau is released by the neurons or, on the contrary, is retained intracellularly. Although there is evidence that supports the neuronal release of tau [
1,
9,
10,
56], our data also demonstrate that soluble intracellular phospho-tau could be toxic for microglia after phagocytosis. The phagocytic capacity of microglial cells is highly induced by apoptotic signals in the affected neurons [
34,
52]. Thus, we postulate that not only the accumulation of intraneuronal soluble phospho-tau but also the induction of apoptosis and the consequent phagocytosis of the phospho-tau affected neurons by the microglia trigger the toxicity in these cells, although the exact intracellular/extracellular cell death mechanism needs to be further investigated. Supporting this suggestion, viable SH-tau cells produced no toxicity, whereas apoptotic tau-expressing SH-cells were highly toxic for BV2 cells. Moreover, phagocytosis of Abeta-expressing N2a-APPswe cells was not toxic for microglial cells. Furthermore, the toxic effect was suppressed by physical (transwell device) and pharmacological (cytochalasin B) inhibition of phagocytosis. Therefore, the toxic effect of soluble phospho-tau in AD cases is likely mediated by the microglial phagocytic capacity for the clearance of apoptotic tau-bearing neurons or, more probably, for the elimination of the neuropil threads. This suggestion is in agreement with a recent work [
21] that demonstrates the involvement of microglia in the phagocytosis of synapses after oligomeric Abeta stimulation and the accumulation of soluble phospho-tau in the presynaptic terminals of demented cases [
55]. Thus, based on these observations, we postulate that the accumulation of soluble phospho-tau forms in aberrant synaptic terminals and/or dystrophic neurites could induce the phagocytic response of the microglial cells, thereby producing toxicity in these cells. This close association between tau-bearing neurons or neuropil threads and severely dystrophic microglial cells has also been reported by Streit and co-workers [
54].
This restricted toxic effect, together with a probably limited regenerative capacity of the microglial cells in AD individuals, could explain the regional pattern of microglial degeneration and also the minor microglial pathology observed in other hippocampal regions, such as CA1 or CA2, with a high accumulation of tangle-bearing neurons.
In summary, our results demonstrate the existence of a significant microglial degenerative process in the hippocampus of AD patients. This degenerative process reduces the parenchymal area covered by microglia and consequently compromises the immune coverage and neuronal survival. Our data also demonstrate that soluble AT8- and/or AT100-positive phospho-tau species, located either extracellular or intracellular after phagocytosis, drive microglial degeneration. The microglial vulnerability in AD pathology provides new insights into the immunological mechanisms underlying this neurodegenerative disease. Finally, our findings highlight the need to improve or develop new animal models, as the current models do not mimic the microglial pathology observed in the hippocampus of AD patients.