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In Alzheimer’s disease (AD), amyloid-β (Aβ) deposits are frequently surrounded by activated microglia but the precise role of these cells in disease progression remains unclear. The chemokine receptor CX3CR1 is selectively expressed in microglia and is thought to modulate their activity. To study the specific effects of microglia activation on amyloid pathology in vivo, we crossbred mice lacking CX3CR1 with the Alzheimer’s mouse model CRND8. Surprisingly, we found that CX3CR1 deficient mice had lower brain levels of Aβ40 and Aβ42 and reduced amyloid deposits. Quantification of Aβ within microglia and time-lapse two photon microscopy in live mice revealed that these cells were highly effective at the uptake of protofibrillar amyloid but were incapable of phagocytosis of fibrillar congophilic Aβ. CX3CR1 deletion was associated with increased phagocytic ability which led to greater amyloid content within microglial phagolysosomes. Furthermore, CX3CR1 deficient mice had an increased number of microglia around individual plaques due to higher proliferative rates, which likely contributed to an overall greater phagocytic capacity. CX3CR1 deletion did not affect the degree of neuronal or synaptic damage around plaques despite increased microglia density. Our results demonstrate that microglia can regulate brain Aβ levels and plaque deposition via selective protofibrillar Aβ phagocytosis. Modulation of microglia activity and proliferation by CX3CR1 signaling may represent a therapeutic strategy for AD.
Alzheimer’s disease (AD) is characterized by the presence of extracellular amyloid β peptide (Aβ) deposits surrounded by activated glia and dystrophic neurites. Microglia are the resident immune cells of the brain and can become activated by a variety of changes in its structural and biochemical integrity (Hanisch and Kettenmann, 2007). Microglia activation and clustering around amyloid deposits tends to occur in early stages of the disease process. Microglia have been shown to be capable of phagocytosis of amyloid fibrils (2) and deficits in microglia accumulations can accelerate amyloid deposition (El Khoury et al., 2007). However, the role of microglia in AD has not been studied in great detail using in vivo models and it remains unclear to what extent microglia are capable of removing amyloid from pre-existing fibrillar congophilic plaques (Stalder et al., 2001) or of preventing the deposition of smaller protofibrillar Aβ aggregates (Grathwohl et al., 2009). Thus, it is not clear how or to what extent microglia control brain amyloid levels or deposition.
To address these questions we investigated mice lacking the chemokine receptor CX3CR1 (Jung et al., 2000). In the brain, this receptor is exclusively expressed in microglia and selectively modulates microglial activity in response to its ligand the chemokine fractalkine (Harrison et al., 1998). Thus, CX3CR1 could potentially play a role in modulating microglia function in AD. CX3CR1−/− mice were crossbred with transgenic mice (CRND8) harboring the human amyloid precursor protein gene with the Indiana and Swedish mutations (Chishti et al., 2001). To determine the role of CX3CR1 in microglia phagocytic activity and regulation of amyloid levels, we implemented new in vivo methods for tracking the interactions between microglia and both fibrillar and non-fibrillar amyloid material using longitudinal imaging with high resolution confocal as well as transcranial two photon microscopy.
Previous in vivo results range from showing that microglia are effective at phagocytosis of fibrillar amyloid (Frautschy et al., 1991; Bacskai et al., 2002; Bolmont et al., 2008) to suggesting that microglia play no role in the control of amyloid deposition (Grathwohl et al., 2009). By contrast, in our study, microglia were incapable of phagocytosis of congophilic fibrillar amyloid from plaques but were very effective at the uptake of oligomeric and protofibrillar Aβ. This selective uptake ability was critical for the regulation of total brain amyloid levels and depositon. CX3CR1 deletion increased microglia proliferation and numbers specifically around plaques, which coupled with their increased phagocytic ability, resulted in decreased brain amyloid levels and deposition.
Conflicting evidence in the literature suggests either neurotoxic or neuroprotective effects of CX3CR1 deletion in various disease models (Cardona et al., 2006; Fuhrmann et al., 2010). However, we found no difference in either neuronal loss or synaptic injury around plaques. This occurred despite significant differences in the density of plaque associated microglia which in their activated status are thought to have neurotoxic potential (Block et al., 2007). Thus, our data demonstrate that microglia play an important role in selective phagocytosis of oligomeric and protofibrillar Aβ but not of preexisting congophilic amyloid plaques. This function may be critical in controlling Aβ deposition as well as levels of potentially neurotoxic Aβ oligomers (Lambert et al., 1998). Thus enhancing microglia phagocytosis and proliferation by blocking CX3CR1 signaling could constitute a therapeutic strategy for AD.
The generation of CX3CR1 deficient mice and TgCRND8 mice has been previously described. Briefly, the CX3CR1 gene locus underwent targeted deletion and direct replacement by a green fluorescent protein (GFP) reporter gene. CX3CR1+/− mice previously backcrossed to C57BL/6 mice for more than 10 generations were crossbred with TgCRND8 mice to obtain CRND8/CX3CR1−/−, CRND8/CX3CR1+/− and CRND8/CX3CR1+/+ mice. Experiments in which we quantified amyloid plaque density, Aβ concentrations and APP processing/cleavage were done in male mice. All other experiments including quantification of Aβ within microglia and in vivo imaging experiments were done in mixed gender mice but with equal gender distribution on each experimental group. Experimental protocols were approved by the Northwestern University Feinberg School of Medicine Institutional Animal Care and Use Committee.
GFP-labeled microglia and Methoxy-X04 (MX04) labeled plaques were imaged through a thinned skull preparation as previously described (Grutzendler et al., 2002). Briefly, transgenic mice were anesthetized with Ketamine/Xylazine and the skull was exposed with a midline scalp incision. About a 1 mm diameter skull region over the somatosensory cortex was thinned with a high speed drill and a microsurgical blade to a final thickness of ~30μm. The skull was attached to a custom-made steel plate to stabilize the head while imaging. A mode-locked Ti-sapphire laser (Coherent Inc.) was used for two-photon excitation (Prairie technologies) and tuned to 835 nm for dual imaging of GFP and MX04 or 890 nm for GFP alone. Emission wavelengths for MX04 and GFP were separated from each other with a 490 nm dichroic. Images were taken using a water immersion objective (Olympus 40x or 60x, 0.8 and 0.9 N.A. respectively) at z-steps of 1–2 μm and zooms of 1–4x. In most cases images were taken at depths up to ~150 μm below the pial surface. The same regions were relocated at different time points using the unchanged pattern of pial blood vessels as reference as well as registration of X, Y, Z coordinates of each scanning area at the initial time point and relocation of the same coordinates using a precision motorized stage at subsequent time points. The precise identity of the areas is confirmed by the presence of unambiguous microvascular shadows or plaque patterns
A ~500 μm area of the skull was thinned with a high speed micro drill at specific cranial stereotaxic coordinates (3.0mm mediolateral, −8.0mm anterioposterior from Bregma). The tip of a 27g needle was used to make a small cut on the surface of the thinned skull area and into the underlying dura without touching the brain parenchyma. Custom-made polypropylene tubing with a final outer diameter of 90μm was inserted ~300 μm into the subarachnoid space and fixed to the skull with cyanoacrylate glue. The tube was connected to a programmable syringe pump through a Hamilton syringe and 10μl (0.4 μM) of fluorescently labeled Aβ42 monomers (HiLyte Fluor 555-Aβ42, AnaSpec, Inc) freshly dissolved in sterile aCSF immediately prior to injection was infused at a rate of 0.2 μl/min. After infusion, tubing was removed and animal scalp was sutured. Animals were imaged in vivo by TPM or in fixed tissue by confocal microscopy at various time intervals after infusion.
Analysis of microglia phagocytosis of fluor-Aβ42 and presence of Thioflavin-S positive material within microglia was performed using NIH ImageJ software on confocal images of brain sections obtained from Fluor-Aβ infused mice. All images were obtained with standardized parameters of laser intensity, gain and pinhole size. a) fluor-Aβ42 and thioflavin-S content within microglia and astrocytes: microglial or astrocyte cell bodies were identified in z-projections from confocal images. A region of interest (ROI) was drawn around the cell body perimeter as well as a mirror ROI control immediately outside of each microglia or astrocyte. Mean fluorescence intensity values were measured for each cell and control ROI in both red channel for Aβ42 (HiLyte-Fluor-555) and blue channel for Thioflavin-S positive fibrillar material. b) Quantification of Aβ fluorescence outside of microglia: custom made NIH Image J Macros were written with the purpose of masking and subtracting areas where microglia were located. Mean fluorescence intensity values were obtained from the resulting area after subtraction of microglia regions. For the purpose of our study any fluor-Aβ deposit that was not located inside microglia was considered to be in the extracellular space, given our finding of the limited intake of fluor-Aβ by astrocytes (supplementary data). Ultimately however, it is not possible to determine unambiguously if deposits are extracellular given the very small size of this space which is beyond the resolution of light microscopy
Sagital sections of the entire left hemisphere were obtained with a cryostat at 50 μm thickness. 8 sections for each hemisphere were selected and visually matched between animals to represent the same brain regions. Sections from all genotypes were stained with thioflavin-S with a standardize concentration and time. Whole brain sections were imaged with a fluorescence microscope (Zeiss axiovert 200M, objective 20x, 0.75NA) using identical parameters of gain and exposure in the CCD camera. All images were stitched together to generate a single image of the entire brain slice. A custom-made thresholding macro based on NIH-Image J was used to detect all fluorescent amyloid plaques using exactly the same parameters for all sections and genotypes. NIH-Image J particle counting plug-in was used to calculate the total number of amyloid plaques per brain section. A subset of images was counted manually by an unbiased individual to corroborate the accuracy of our automated counting. The use of automated counting allowed us to count a much larger number of plaques representing the majority of the hemisphere than would have been possible with stereological methods.
Mice were injected only once with MX04 2 days before the first imaging time point. TPM imaging was done as described above. Image stacks containing plaques were run through a “despeckle” filter to remove noise, and then a maximal projection through a consistent number of slices was taken using NIH image J software. To achieve accurate plaque area measurements regardless of minor brightness differences across time points, images were thresholded for each time point at the average pixel brightness value between a consistent plaque area and the adjacent background.
Cryosections (50μm thick) underwent sodium citrate antigen retrieval and were immunolabled with the following antibodies: 4G8 (Covance), A11 (Biosource Invitrogen), IBA1 (Wako Chemicals USA, Inc), and LAMP1 (Developmental Studies Hybridoma Bank (Iowa) clone 1D4B) according to manufacturer protocols. Confocal images of plaques selected for their similar size, location and cortical layer were obtained for quantitative imaging using an Olympus FV10i Confocal Microscope (60X objective N.A. 1.2). Image z-stacks were taken with standardized laser intensity, gain and pinhole sizes to allow inter-animal comparisons. LAMP1-positive lysosomes inside IBA1-labeled microglia have distinct fluorescence intensities in comparison to LAMP1-positive structures outside microglia. Using NIH Image J, we applied a fluorescence threshold to the LAMP1 signal and created a binary image that was then combined with the other fluorescent channels in the z-stack to measure 4G8, A11, and Thioflavin S fluorescence intensities within the microglial LAMP1 structures. Measurements were performed on each optical slice of a given z-stack and then averaged.
One day old CX3CR1+/+ and −/− mouse littermates were used for primary cultures of cerebral cortical cells. These cell cultures include abundant microglia, astrocytes and neurons as confirmed by immunocytochemistry. Briefly, following cell dissociation, a mixture of equal number of brain cells from both genotypes was adjusted to 106 cells/ml in DMEM/10% fetal bovine serum (FBS), followed by plating of the cell suspension on a culture flask. Fifty percent of the medium was replaced at days 3, 10 and 17. At day 21, cells reaching 90% confluence were trypsinized and re-seeded to 24-well plates containing poly-lysine coated coverslips at a density of 5×104 cells/ml. The re-seeded cells were rinsed in DMEM containing 10% FBS, pretreated with or without 10μg/ml CX3CR1 neutralizing antibody for 30min and then incubated for 2 hours in the same medium containing either 2μm diameter fluorescent microspheres or 0.05mg/ml fluorescently-conjugated Aβ (HiLyte Fluor 555- Aβ42 AnaSpec, Inc) in DMEM. After washing with DMEM, cells were fixed in 2% paraformaldelhyde/PBS for immunocytochemistry. Fixed cells were rinsed in PBS, blocked in 3% normal goat serum/0.3% triton X-100 for 30min and then incubated with primary antibodies (IBA1, rabbit polyclonal, 1:500 and GFP, mouse monoclonal, 1:1000) overnight. CX3CR1−/− microglia were differentiated from CX3CR1+/+ microglia because they had positive labeling with both IBA1 and GFP antibodies while CX3CR1+/+ microglia were only labeled by IBA1. We also excluded the possibility of other cell types expressing either IBA1 or GFP in control experiments in which we used dual labeling with antibodies specific for astrocytes (GFAP) or neurons (MAP2). In separate culture preparations containing only CX3CR1−/− microglia we determined that 100% of IBA1-positive microglia were GFP positive. Microglia of each genotype represent ~50% of the total microglia population in these mixed cell cultures. Minor differences in the proportion of microglia of each genotype do not affect the final results of the phagocytosis assay given that in a mixture of cells there is equal likelihood for each individual cell regardless of its genotype to be exposed to Fluor 555-Aβ42 or microspheres. For quantification of microglia uptake of microspheres or Aβ, we used a custom-made macro with NIH-Image J software for automated measurement of total fluorescence intensity within individual microglia.
Cryosections (50μm thick) were immunolabled with any of the following antibodies: Anti IBA1 (Wako Chemicals USA, Inc), Synaptophysin (Chemicon NeuN (Chemicon) according to manufacturer protocols. Confocal images of plaques selected for their similar size, location and cortical layer (layers III to VI in somatosensory cortex) were obtained for quantitative imaging using a Zeiss LSM 510 Meta UV microscope (63X objective N.A. 1.4). Images were taken at standardized laser intensity, gain and pinhole sizes to allow inter-animal comparisons. Quantification of microglia and NeuN postitive neuron number was performed using the Image J threshold/analyze particle features. Results were validated by manual counting in a subset of stacks. To control for possible differences in microglia density between CX3CR1 genotypes in the absence of AD pathology, we measured the density of microglia in non-AD/CX3CR1 mutant mice which did not differ between genotypes in the brain region analyzed. We have observed full colocalization of IBA1 and F4/80 (an additional microglia marker) demonstrating that all microglia are IBA1 immunoreactive. Furthermore, IBA1 has a 100% colocalization with GFP-labeled cells in the brain parenchyma of CX3CR1 knock in mice. Thus IBA1 labels all brain microglia but cannot differentiate them from infiltrating macrophages given that macrophages also label with IBA1. Microglia density around plaques was quantified in blindly selected plaques of the same average size to eliminate size as a variable which can lead to differences in microglia number unrelated to the CX3CR1 genotype..
For synaptophysin quantification, multi-channel 8 bit images were obtained in which plaques were labeled with Thioflavin S in one channel and synaptophysin in a second channel. Using a custom ImageJ macro, the resultant stacks were analyzed slice by slice to determine the fluorescence of synaptophysin as a function of distance from the plaque edge. The plaque edge was determined by thresholding the Thioflavin S channel at a constant predetermined value of 115. This edge was selected and then enlarged by 2 μm (the selection grows in all directions out from its center for 2 μm) to form an even starting point. Then, 10 donut-shaped selection bands of 3 μm width were created at 3 μm intervals, covering a 30 μm circular area out from the evened plaque edge. This essentially creates concentric circular bands of selection, which were then used to measure average fluorescence intensity in the synaptophysin channel, with the first band nearly adjacent to the plaque (all pixels 2–5 μm from the thresholded plaque edge), and the last band about 30 um away from the plaque edge (all pixels 32–35 μm from the thresholded plaque edge).
a) For cell body motility, in vivo TPM images of both microglia and plaques were taken over 4–7 days. For long-term monitoring, a subset of mice was imaged at 1 and 4 month intervals. Microglia cell bodies adjacent to the plaque were counted for each time point and microglia stability was calculated as the percent of all adjacent cell bodies in the first time point which remained in the same location relative to the plaque at the later time point. b) For cell process dynamics, image stacks of microglia processes at two time points 1 hour apart were aligned using ImageJ plugins, maximally projected, matched for brightness, and then thresholded. Each time point was subtracted from the other in order to determine regions which were “lost” (apparent in the first image but not the second) or “gained” (apparent in the second image but not the first). The areas of regions of gains or losses above an arbitrary cutoff of 3.2 μm2 were then measured and counted per image stack.
Mice received daily injections of bromodeoxyuridine (BrdU) (50 mg/kg, i.p.) for 14 days. Brains were removed following 4% PFA perfusion and cryosectioned at 50μm thickness. BrdU was detected after denaturing with 5N HCl followed by incubation with rat anti-BrdU antibody (1:400, Serotec, Oxford, UK) and rabbit anti-IBA1 antibody for microglia detection. BrdU and IBA1 were detected with AlexaFluor 555/488 conjugated secondary antibodies and imaged with confocal microscopy. BrdU-labeled microglia were quantified from confocal image stacks in areas in the vicinity (25 μm) and distant from plaques.
Aβ42 and Aβ40 were measured by commercially available ELISA kits (Biosource). Brain hemispheres of CX3CR1+/+, CRND8/CX3CR1+/+ and CRND8/CX3CR1−/− mice were homogenized in cold 5 M guanidine HCl/50mM Tris HCl. ELISA assays were performed according to manufacturer’s protocol.
Brain tissue lysates were prepared in ice-cold TEEN-Tx (TEEN buffer (50 mM Tris-HCl, pH 7.4, 1 mM EDTA, 1 mM EGTA and 150 mM NaCl) containing 1% Triton X-100) by sonication. This was followed by centrifugation, protein quantification, addition of buffer, and boiling for 5 minutes. Proteins were separated using 15% tris-tricine polyacrylamide gel and incubated with APP C-terminal-specific rabbit anti-APP-CT20 antibody (1:10000; Calbiochem) and mouse anti-α-tubulin (1:5,000; Upstate). Densitometric quantification of band intensity was performed using NIH ImageJ software.
Mouse presenilin-1 and beta-site APP-cleaving enzyme 1(BACE1) mRNA expression were detected by RT-PCR. Briefly, total RNA was extracted and cDNA was obtained by Reverse Transcription according to manufacturer’s instruction. The following pairs of primers were used: 5′-AGC ATG ACA GGC AGA GAC TTG ACA-3′ and 5′-AAC GTA GTC CAC GGC GAC ATT GTA-3′ for presenilin 1, 5′-ATC TAC ACG CCC TTC ACG GAG G-3′ and 5′-TGG GCA GTT TCC ACC AGC ATC-3′ for presenilin 2, 5′-AGG GCT ACT ATG TGG AGA TGA CCG TA-3′ and 5′-TCC CAC TGT CCA CAA TGC TCT TGT-3′ for BACE1, 5′-AGA TGA GAA TGC CAG TCG CTC CTT-3′ and 5′-TGC ACA GTT GAG GTT CCC GAC TAA-3′ for BACE2, and 5′-AAC TTT GGC ATT GTG GAA GGG CTC-3′ and 5′-ACC CTG TTG CTG TAG CCG TAT TCA-3′ for GAPDH (internal control).
Statistical analysis was performed using two-tailed Student’s t-test or linear correlation analysis. P< 0.05 was considered significant.
To examine the role of microglia in the process of brain amyloid buildup, we crossbred CRND8 mice with mice lacking the CX3CR1 receptor (CX3CR1−/−). At 2 months of age, both CRND8/CX3CR1−/− mice and their wild-type littermates (CRND8/CX3CR1+/+) demonstrated low levels of total brain Aβ40 and Aβ42 peptides (Figures 1A, B). At 5 months, brain Aβ40 and Aβ42 concentrations had risen markedly but were significantly lower in CRND8/CX3CR1−/− mice as compared with CRND8/CX3CR1+/+ littermates (Figures 1A, B). The effect of CX3CR1 deficiency on Aβ deposition was further examined by quantifying the density of Thioflavin-S (TS) labeled fibrillar amyloid deposits. We found that CRND8/CX3CR1−/− mice had a significantly reduced number of amyloid plaques at both 3 and 5 months of age (Figure 1C, D). The disparity in brain Aβ levels and amyloid plaque number between CX3CR1 genotypes were likely due to reduced Aβ accumulation rather than decreased Aβ production given that there was no significant difference in amyloid precursor protein (APP) concentrations (Figure 1E), mRNA expression of APP processing enzymes (BACE1 and 2; PS1 and 2) (Figure 1F) or levels of APP cleavage products (Figure 1G).
Given that the expression of CX3CR1 in the brain is microglia specific, the reduction in Aβ levels and plaque density in CRND8/CX3CR1−/− mice suggests a critical role of microglia and CX3CR1 in controlling Aβ accumulation.
Although microglia appear to be capable of soluble (Mandrekar et al., 2009) and fibrillar (Frautschy et al., 1992) Aβ uptake, their phagocytic properties in vivo are not well characterized. To determine the mechanisms by which microglia control the accumulation of Aβ in the brain, we examined their phagocytic capabilities in living mice by time-lapse transcranial two-photon microscopy (TPM). We first labeled plaques by intraperitoneal injection of the fluorescent amyloid-binding dye methoxy-XO4 (Klunk et al., 2002) 48 hours prior to the first imaging session and without relabeling on subsequent sessions. We reasoned that if microglia phagocytosis of fibrillar amyloid were a robust phenomenon in vivo, the size and shape of the initial fluorescent plaque label would change significantly overtime as portions of amyloid deposits are removed by microglia. However, time-lapse TPM images over intervals of up to 1 month showed virtually no change in the size or shape of the original fluorescent plaque label (Figure 2A, B). In addition, we labeled fixed brain slices with the fluorescent fibrillar amyloid binding dye Thioflavin-S (TS) and with IBA1 antibody to visualize fibrillar amyloid contained within microglia using high resolution confocal microscopy. Quantification of TS fluorescence within IBA1-immunolabeled cells revealed absence of any intracellular TS labeling (Figure 2C, D) regardless of the CX3CR1 genotype (Figure 2D).
To further establish whether fibrillar amyloid within plaques is stable or is constantly being removed, we developed a method for subarachnoid infusion of fluorescently tagged Aβ peptides. We obtained time-lapse images with two-photon microscopy while Aβ42 (HiLyte)-555 was being infused. During imaging, we observed very rapid binding of Aβ42 (within minutes) to preexisting amyloid plaques (Figure 3A) suggesting a very high affinity of soluble Aβ42 for fibrillar amyloid within plaques. However, non-plaque areas immediately adjacent demonstrated very low fluorescence levels (Figure 3A) suggesting that unbound Aβ42 was rapidly cleared possibly through the blood brain barrier (DeMattos et al., 2002). Once bound to plaques, we found by time-lapse imaging that Aβ42 was highly stable over many weeks like we observed with methoxy-XO4. This was formally quantified from fixed tissue confocal images of amyloid plaques obtained from animals infused with Aβ42 (HiLyte)-555 and sacrificed either 1 or 90 days post-infusion. We reasoned that if there were any significant phagocytosis of plaque-associated Aβ, we would observe a reduction in the average Aβ42 (HiLyte)-555 fluorescence intensity and/or plaque diameter over time. However, we found complete stability of the label at least over this 90 day interval (Figure 3B, C).
Thus, the long-term persistence of methoxy-XO4 and Aβ42 (HiLyte)-555 plaque labeling coupled with the absence of TS label inside microglia strongly suggests that little phagocytosis of fibrillar amyloid takes place in vivo.
To test whether microglia were capable of phagocytosis of soluble or “protofibrillar” Aβ (broadly defined in our study as Aβ deposits that do not co-label with Thioflavin-S), we infused Aβ42(HiLyte)-555 into the brain via the subarachnoid space and obtained brain slices at different time intervals for confocal microscopy. Within hours of infusion, we observed strong binding of Aβ42(HiLyte)-555 to preexisting methoxy-XO4-labeled fibrillar amyloid plaques as seen by in vivo imaging. However, little Aβ42(HiLyte)-555 was observed in areas outside of plaques in the first 12 hours post infusion, likely due to rapid Aβ efflux through the BBB. Beginning at ~24 hours, however, scattered fluorescent Aβ deposits became evident (Figures 4C) possibly due to aggregation of Aβ42(HiLyte)-555 that failed to be cleared through the BBB or due to binding to pre-existing protofibrillar aggregates. Many of these early Aβ deposits outside of microglia were likely to be located in the extracellular space given that they did not colocalize with astrocytes either (Figure 4F and Supplementary Figure 1). However, at this early time point a substantial number of deposits were already observed inside microglia within vesicular compartments consistent with phagolysosomes(Frackowiak et al., 1992) (Figure 4A, C and supplementary movie 1).
Quantification of Aβ42(HiLyte)-555 fluorescence within individual microglia showed a ~3 fold rise 16 days after subarachnoid infusion (Figure 4E). Concurrent with the increase in fluorescence inside microglia, at day 16 there was a large decrease in the number and intensity of fluorescent aggregates outside microglia (Figure 4F) suggesting that these newly formed Aβ deposits were being phagocytosed. Virtually none of these Aβ42(HiLyte)-555 accumulations co-labeled with TS (Figures 4B, D) suggesting that these deposits were composed of material in a protofibrillar Aβ conformation. Together, these data indicate that microglia exhibit an increasing capacity for Aβ phagocytosis as it gradually adopts protofibrillar conformations but fail to phagocytose fibrillar deposits that have adopted a β-pleated sheet conformation.
We then aimed at determining if microglia, in addition to exogenously administered Aβ42, can also uptake endogenously produced amyloid. We obtained high resolution confocal images of brain slices from CRND8 mice stained with A11 and 4G8 antibodies, which detect oligomeric and other protofibrillar conformations respectively (Lesné et al., 2006; Kayed et al., 2007). We found bright clusters of A11 and 4G8-immunoreactive material that did not co-label with TS, mostly inside lysosomal-associated membrane protein (LAMP-1)-immunoreactive vesicles inside microglia (Figures 5A-D and supplementary Figure 2). Thus endogenously produced oligomeric and protofibrillar amyloid but not thioflavin-S positive fibrillar material are efficiently phagocytosed by microglia and may be targeted for degradation given their presence within phagolysosomes.
To explore the possibility that the disparities we observed in brain amyloid accumulation between CX3CR1 genotypes could be explained by differential microglial phagocytic capacity, we quantified A11 and 4G8 immunofluorescence within LAMP-1 labeled phagolysosomes in microglia (Figures 5A–F). We found a significantly increased A11 and 4G8 immunofluorescence in phagolysosomes from CRND8/CX3CR1−/− mice compared to wild-type littermates, suggesting that CX3CR1−/− microglia have an increased capacity to phagocytose endogenously produced amyloid.
To further prove that the increase in protofibrillar Aβ observed within phagolysosomes in vivo in CX3CR1 deficient mice was the result of increased phagocytosis rather than decreased degradation, we performed an in vitro phagocytosis assay. We generated dissociated cortical cell cultures containing a mixture of CX3CR1−/− and CX3CR1+/+ microglia and incubated them for 2 hours with either Aβ42 (HiLyte)-555 or 2 μm fluorescent microspheres. Following incubation we carried out quantitative imaging and compared the average fluorescence intensity within microglia of both genotypes (Figure 6A–D). We found that CX3CR1−/− microglia had a greater capacity to uptake both Aβ peptides and microspheres suggesting an overall enhanced phagocytic activity. Pre incubation with anti-CX3CR1 neutralizing antibody 2 hours prior to the phagocytosis assay led to increased microsphere phagocytosis in CX3CR1+/+ microglia but not those lacking CX3CR1 (Figure 6E) demonstrating rapid and specific modulation of phagocytosis by the reduction of CX3CR1 activity. The fact that microsphere phagocytosis was increased demonstrates that differences in Aβ within microglia are not likely due to intracellular degradation but rather uptake differences. It also indicates that the increased phagocytic capacity is not specific to amyloid but a general property of CX3CR1−/− microglia.
To examine additional cellular mechanisms that might explain differences in Aβ accumulation between CX3CR1 genotypes, we analyzed the spatial relationship between amyloid plaques of similar size and their surrounding microglia. We found a greater average number of microglia within a 25 μm radius of the plaque edge in 5 month-old CRND8/CX3CR1−/− mice than in wild-type littermates (Figure 7A, B). Several mechanisms could explain the observed difference in microglia number such as differential proliferation, migration and/or retention rates.
To examine the extent of microglia proliferation, we administered daily injections of 5-bromo-2-deoxyuridine (BrdU) for 2 weeks in 5 month old mice. We quantified the number of BrdU-labeled microglia located in the immediate vicinity of plaques in mice of both CX3CR1 genotypes. Interestingly, we found that the overall proportion of microglia that were BrdU-labeled was significantly greater in CRND8/CX3CR1−/− mice than in CRND8/CX3CR1+/+ littermates (Figure 7C, D), while non-plaque areas showed substantially lower overall proliferation rates and no significant differences between genotypes (Figure 7D). Thus increased proliferation rates around plaques are likely to explain the greater number of plaque associated microglia in CX3CR1 deficient mice.
To determine if migration/retention also plays a role, we used time-lapse two photon microscopy in live mice to track the dynamics of GFP-labeled microglia around plaques. We found that CX3CR1 deficient microglia display enhanced process dynamism over short intervals (Supplementary Figure 3). Microglia cell bodies however, can remain stable over months (Supplementary figure 4) but no significant difference in stability between mice of different CX3CR1 genotypes was observed (Figures 7E, F). However, technical limitations of in vivo imaging make it unfeasible to quantify microglia cell body dynamics over longer intervals with sufficient precision. Thus we cannot rule out that subtle variations in microglia recruitment/retention could contribute to the observed disparities in microglia number around plaques.
Results of studies in a variety of disease models have yielded conflicting results regarding secondary neurotoxicity in CX3CR1 mutants. For example, mice deficient in CX3CR1 were found to have enhanced neuronal death in models of MPTP toxicity and Amyotrophic Lateral Sclerosis (ALS)(Cardona et al., 2006), while CX3CR1 deficiency was neuroprotective in models of cerebral ischemia (Dénes et al., 2008) and mutant tau/APP associated neurodegeneration(Fuhrmann et al., 2010). Since our results demonstrated that CX3CR1 deficient mice had increased numbers of microglia around amyloid plaques we wondered whether this would lead to increased neuronal injury given that activated microglia can secrete potentially neurotoxic substances (Block et al., 2007). We thus investigated whether CX3CR1 deficiency had any effect on morphological correlates of neurotoxicity as assessed by neuronal and synaptic density in the immediate surroundings of amyloid plaques. Quantification of cells with the neuronal nuclear marker NeuN and of synaptophysin immunoreactive puncta demonstrated no statistically significant difference between wild type and CX3CR1 deficient mice (Figure 8). Thus it appears that CX3CR1 deficiency has neither neuroprotective nor neurotoxic effects around individual plaques at least in this model of Alzheimer’s-like pathology. However, given that CX3CR1 deficient mice have fewer plaques, it is likely that the overall pathological burden is diminished.
Our results provide new insights into the role of microglia in the pathogenesis of Alzheimer’s disease in vivo. We show that deletion of the fractalkine receptor CX3CR1, which in the brain is specifically expressed in microglia, leads to a significant reduction in the total levels of brain Aβ and in the number of amyloid plaques in a mouse model of AD. This demonstrates that microglia play an important role in regulating brain amyloid levels and deposition.
The effect of CX3CR1 deletion on brain amyloid was mediated by a general enhancement of microglia phagocytic ability as demonstrated both in vitro and in vivo. In addition, we also found that with CX3CR1 deficiency, microglia proliferation near plaques was increased leading to a greater density. This may further enhance the global brain capacity for Aβ clearance.
However, in contrast to recent imaging studies (Bolmont et al., 2008) we found that microglia, while highly effective at phagocytosis of oligomeric and protofibrillar Aβ, were incapable of removing fibrillar amyloid from preexisting plaques in vivo. Thus microglia may only be able to remove fibrillar amyloid from plaques during heightened levels of activation such as in injury models (DiCarlo et al., 2001) or after Aβ immunization (Bacskai et al., 2002). Our study also contrasts with recent studies of microglia ablation showing no role for these cells in brain amyloid plaque regulation (Grathwohl et al., 2009). Ablation is likely to induce marked astrocyte activation and could turn these cells into much more effective phagocytes which could compensate for the loss of microglia and thus explain the apparent lack of effect of microglia ablation on Aβ levels.
Although we define microglia uptake of Aβ as “phagocytosis”, the precise mechanism of Aβ uptake is unclear and it has been proposed that microglia can also use fluid phase macropinocytosis to clear Aβ (Mandrekar et al., 2009). However, given that CX3CR1 deletion is able to modulate phagocytosis of polystyrene microspheres, we assume that the disparity in Aβ uptake between genotypes is mostly due to differential phagocytic capacity rather than macropinocytosis. Ongoing microglial phagocytosis around growing fibrillar deposits could reduce levels of oligomeric and protofibrillar Aβ, which may change the equilibrium between monomers, oligomers and fibrils, altering the process of nucleation-dependent polymerization that leads to fibrillar plaque formation (Harper and Lansbury Jr., 1997; Walsh et al., 1999). This not only may reduce the number of amyloid deposits but the overall concentration of oligomeric and protofibrillar Aβ which could have important implications in the progression of AD given the enhanced toxicity of these smaller Aβ conformations (Lambert et al., 1998; Walsh et al., 1999). Thus, it is likely that the main role played by microglia is to ameliorate the formation of new fibrillar plaques rather than to remove preexisting ones, with the most critical effect likely being at the earliest stages of plaque deposition.
In addition to microglia, peripheral macrophages have been shown to be capable of Aβ phagocytosis in AD models (Simard et al., 2006). Although currently it is thought that macrophages may normally not enter the brain in significant numbers in the absence of blood brain barrier disruption(Ajami et al., 2007; Mildner et al., 2007), fractalkine/CX3CR1 signaling may also have chemotactic effects on macrophages(Imai et al., 1997). Thus we cannot rule out that some of the effects we observed are at least partly due to peripheral macrophages entering the brain.
Despite increased clustering of activated microglia in CX3CR1 deficient mice, we did not observe any difference in neuronal or synaptic density around plaques. This suggests that plaque-associated microglia do not exert a significant neuroprotective or neurotoxic “bystander” effect. Microglia have been shown to exacerbate neuronal death mainly in acute and subacute injury models where they are fully activated (Wu et al., 2000; Cardona et al., 2006; Block et al., 2007). However, in slowly progressive neurodegenerative disorders, evidence of neurotoxicity or neuroprotection has been more conflicting. Thus it appears that microglia activation is not an all or none process and its net effect may depend on the particular pathological scenario (Hanisch and Kettenmann, 2007). In the case of CX3CR1, our results indicate that reducing microglia activation or phagocytosis could have deleterious effects in AD. This may at least partly explain why recent attempts at prevention of AD with non-steroidal anti-inflammatory therapy have been unsuccessful or even counterproductive (Martin et al., 2008).
Our findings support the concept that fractalkine through its receptor CX3CR1 (Harrison et al., 1998) acts as a repressor of microglia activation (Cardona et al., 2006). This would explain the increased proliferation and clustering around plaques with CX3CR1 deletion. Although the increased number of microglia around amyloid deposits would likely lead to an overall greater phagocytic capacity, the precise mechanism through which CX3CR1 affects phagocytosis at the single microglia level is less clear. A large number of pattern recognition cell surface receptors in microglia such as the Toll-like family, receptor for advanced glycation end-products (RAGE) and scavenger receptors have been implicated in the recognition of amyloid oligomers and fibrils (Salminen et al., 2009). Signal transduction pathways associated with these receptors are known to modulate phagocytosis (Tahara et al., 2006; Richard et al., 2008) and cytokine production (Reed-Geaghan et al., 2009) after amyloid exposure. Furthermore, crosstalk can occur between chemokine and pattern recognition receptor signaling pathways leading to functional changes in phagocytes (Kishore et al., 2005; Alves-Filho et al., 2009). This receptor transactivation could partly explain the effect of CX3CR1 on phagocytosis. In addition, CX3CR1 deletion has been shown to directly affect cytokine production (Cardona et al., 2006). Thus, CX3CR1 modulation of phagocytosis may involve the indirect effect of some proinflammatory cytokines which can attenuate phagocytosis (Koenigsknecht-Talboo and Landreth, 2005). On the other hand, our experiment showing a rapid increase in phagocytic ability after incubation with anti-CX3CR1 neutralizing antibody suggests that CX3CR1 may also exert direct effects on microglia phagocytosis.
Expression of various chemokines has been shown to be increased in the aging brain (Mo et al., 2003) and plasma levels of fractalkine can be increased in AD patients (Kim et al., 2008). Increased fractalkine signaling in aging could in theory restrain microglia and predispose individuals to amyloid deposition and toxicity. Although our results were obtained in a model where CX3CR1 was developmentally rather than conditionally depleted and should thus be cautiously extrapolated; attempts at treating a variety of conditions with Fractalkine/CX3CR1 agonists or other microglia inhibitors should take into account their potential for exacerbating AD. On the other hand, selective inhibition of fractalkine or CX3CR1 in the brain, which could modulate microglia proliferation and phagocytosis, may represent a novel therapeutic approach for AD.
This project was supported by the National Institute of Health/National Institute on Aging Grant R01AG027855 (JG), Ellison Medical Foundation New Scholar Award in Aging (JG) and Dana Foundation Brain and Immunoimaging Award (JG).
This study was funded by NIH/NIA grant AG027855, and awards from the Ellison Medical Foundation and the Dana Foundation.