Vascular β-amyloid and early astrocyte alterations impair cerebrovascular function and cerebral metabolism in transgenic arcAβ mice
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Cerebrovascular lesions related to congophilic amyloid angiopathy (CAA) often accompany deposition of β-amyloid (Aβ) in Alzheimer’s disease (AD), leading to disturbed cerebral blood flow and cognitive dysfunction, posing the question how cerebrovascular pathology contributes to the pathology of AD. To address this question, we characterised the morphology, biochemistry and functionality of brain blood vessels in transgenic arctic β-amyloid (arcAβ) mice expressing human amyloid precursor protein (APP) with both the familial AD-causing Swedish and Arctic mutations; these mice are characterised by strong CAA pathology. Mice were analysed at early, mid and late-stage pathology. Expression of the glucose transporter GLUT1 at the blood–brain barrier (BBB) was significantly decreased and paralleled by impaired in vivo blood-to-brain glucose transport and reduced cerebral lactate release during neuronal activation from mid-stage pathology onwards. Reductions in astrocytic GLUT1 and lactate transporters, as well as retraction of astrocyte endfeet and swelling consistent with neurovascular uncoupling, preceded wide-spread β-amyloid plaque pathology. We show that CAA at later disease stages is accompanied by severe morphological alterations of brain blood vessels including stenoses, BBB leakages and the loss of vascular smooth muscle cells (SMCs). Together, our data establish that cerebrovascular and astrocytic pathology are paralleled by impaired cerebral metabolism in arcAβ mice, and that astrocyte alterations occur already at premature stages of pathology, suggesting that astrocyte dysfunction can contribute to early behavioural and cognitive impairments seen in these mice.
KeywordsAlzheimer’s disease Congophilic amyloid angiopathy Cerebral glucose metabolism Astrocytes
Vascular accumulation of β-amyloid (Aβ) results in congophilic amyloid angiopathy (CAA), a vascular lesion diagnosed as part of β-amyloid pathology in up to 90% of post-mortem brains with a neuropathological diagnosis of Alzheimer’s disease (AD) . CAA is associated with a plethora of abnormalities and malfunctioning of the cerebrovasculature [6, 15, 40, 49, 52, 64] including ruptures of the vessel walls causing microhaemorrhages and lesions of the blood–brain barrier (BBB), resulting in inflammation, vascular oedema and uncontrolled influx of peripheral blood components into the brain parenchyma . As a result of CAA, vascular basement membranes (VBM) thicken and smooth muscle cells (SMC) degenerate, jointly leading to impaired cerebral blood flow (CBF) [15, 18, 40]. Physiological functions of the cerebral vasculature are compromised early, possibly even preceding the onset of AD pathology including the onset of CAA, and raising the possibility of an early involvement of vascular dysfunction in the pathophysiology of the disease with disturbances in angiogenesis [11, 54]. Cerebral hypoperfusion and hypometabolism occur in patients with mild cognitive impairment (MCI), and were also observed in transgenic mouse models of AD in which they precede the onset of β-amyloid plaque pathology [5, 10]. While vascular and metabolic disorders including atherosclerosis and diabetes can strongly impair cerebral circulation , neurotoxic soluble Aβ species can also diminish performance of the cerebral vasculature [16, 18], presumably already at early and pre-pathological stages of β-amyloid pathology. As a consequence, Aβ-related disruption of brain blood flow could lead to neuronal impairments by compromising a steady vascular supply of essential metabolic precursors such as oxygen, amino acids and glucose [4, 24, 65], as well as by decreasing cerebral lymphatic drainage of metabolic products [58, 59]. Astrocytes play a pivotal role in regulating these functions by mediating signalling and nutrient exchange between neurons and vascular endothelium [1, 23] via astrocyte endfeet that contain numerous nutrient transporters including the glucose transporter GLUT1 and the monocarboxylate transporter 1 (MCT1), in addition to many receptors essential for signal transduction [1, 41]. Interactions of astrocyte endfeet with the VBM are mediated via the dystroglycan protein complex that binds to laminin and neuronal-expressed neurexin [2, 56, 63]. Astrocyte endfeet can retract and swell during pathological conditions including neuroinflammation in multiple sclerosis and in experimental autoimmune encephalomyelitis . Because both astrogliosis surrounding β-amyloid plaques and neuroinflammation occur in AD, alterations in astrocyte endfeet structure and function are likely to be present [13, 63, 64]; such changes consistent with loss of contact between astrocyte endfeet and the cerebrovasculature were indeed observed in a transgenic mouse model of AD . To further understand the role of the cerebrovasculature and astrocytes as well as their influence on cerebral metabolism in β-amyloid pathology, we characterised cerebral blood vessels and astrocytes in transgenic arcAβ mice expressing both the Swedish and Arctic APP mutations. This mouse model exhibits severe vascular AD pathology with significant electrophysiological and behavioural deficits preceding β-amyloid deposition [28, 35]. The Swedish double mutation (K670N and M671L) together with the arctic mutation (E693G) leads to an over-production of aggregation-prone Aβ42 and accumulation of Aβ at brain blood vessels [28, 35]. Similar as was described for other mouse models of AD [22, 42, 55, 62], our studies identified severe morphological and biochemical abnormalities of CAA-affected vessels including stenoses, leakages, loss of vascular smooth muscle cells, and decreased GLUT1 expression. We also observed a unique astrocytic pathology commencing at the early disease stages. These included reduced GLUT1 and MCT1 expression, decreased astrocytic release of lactate in vivo and ex vivo, increased laminin secretion and retraction and swelling of astrocyte endfeet surrounding diffuse and vascular Aβ depositions, signs of neurovascular uncoupling and BBB disruption. In vivo microdialysis revealed severe reductions in brain glucose concentrations at baseline during neuronal stimulation and impaired blood-to-brain glucose uptake. These findings establish an astrocytic involvement in the early stages of the cerebrovascular and metabolic pathology in this transgenic mouse model of AD.
Materials and methods
ArcAβ mice were produced as described previously by Knobloch and colleagues . Non-transgenic littermates were used as controls when specified. All animals were housed 3–5 per cage and had access to food and water ad libitum under a 12-h light/dark cycle. All in vivo experiments conducted complied with Swiss federal and cantonal law and were performed with the approval of the Swiss Veterinary Council, approval number 47/2008, 3781.
Classification of disease pathology
Transgenic arcAβ mice were grouped according to the disease pathology in the following groups: early-stage pathology: 6 months old, mid-stage pathology: 9–13 months old and late-stage disease pathology: 16–22 months old. Disease stage pathology was based on overall cerebral Aβ plaque burden including CAA and cognitive performance according to Knobloch et al. .
Perfusion of mice and collection of brain material
Mice were deeply anaesthetised using a cocktail of 1.25% ketamine and 0.25% xylazine. Mice used for histochemistry purposes were transcardially perfused with ice-cold PBS (10 mM Na2HPO4, 2.5 mM NaH2PO4, 150 mM NaCl, 3 mM KCl) followed by ice-cold paraformaldehyde (PFA) 4.0% in phosphate buffer 0.15 M (Na2HPO4 + NaH2PO4 in double-distilled water), pH 7.4. Brains were incubated in PFA 4.0% in PBS for 4 h at 4°C followed by incubation in sucrose 30% in PBS for 24–36 h. Mice serving biochemistry purposes were transcardially perfused with ice-cold PBS only. Hippocampi were dissected out and cortices, hippocampi and cerebella were separately snap-frozen in liquid nitrogen and stored at −80°C.
Vascular corrosion casts and scanning electron microscopy
All procedures were carried out as described previously . In brief, anaesthetised mice (100 mg/kg pentobarbital) were first perfused with artificial cerebrospinal fluid (aCSF) containing heparin followed by a perfusion with 4% PFA in PBS. Subsequently, mice were perfused with the casting resin PU4ii (vasQtec, Switzerland). After a curing period of 7 days, soft tissue was macerated in 7.5% potassium hydroxide at 50°C for 24–72 h and the bones decalcified in 5% formic acid at 50°C for 24–72 h. Vascular corrosion casts were washed with water, dried by lyophilisation and sputter-coated with gold particles for scanning electron microscopy (SEM) purposes.
Extraction of membrane brain proteins
Working on ice, brain materials (hippocampi and cortices) were homogenised separately in homogenisation buffer [Tris-HCl 10 mM, pH 7.4, EDTA 1.0 mM, sucrose 250 mM and protease inhibitors (Complete®, Roche, Switzerland)] with 30 strokes using a glass Dounce homogeniser with Teflon pestle (Wheaton, USA). Homogenates were centrifuged at 900×g and 4°C for 10 min. The formed supernatants were subsequently centrifuged at 110,000×g and 4°C for 75 min. Supernatants were discarded and the pellets dissolved in solubilisation buffer [Tris-HCl 10 mM, pH 7.4, EDTA 1.0 mM, Triton X-100 0.50%, sodium deoxycholate 0.50% and protease inhibitors (Complete®, Roche, Switzerland)] by continuous rotation at 4°C for 1 h. The dissolved pellets were centrifuged at 14,000×g and 4°C for 10 min. Supernatants were collected and protein concentrations measured using the Pierce® BCA Protein Assay Kit (Thermo Scientific, Rockford, IL, USA).
Equal amounts of total protein or equal volumes of cell culture media were subjected to separation on 10–20% Tricine gels (Invitrogen, Basel, Switzerland), blotted on nitrocellulose membranes Immobilon-P PVDF membranes (0.45 μm, Millipore, Switzerland). The immunoblot was then incubated with primary antibodies followed by incubation with HRP-tagged secondary antibodies. Detection was performed using chemiluminescence visualised using ECL WB reagents (Amersham Pharmacia, GE, Germany) or SuperSignal West Dura Extended Duration reagents (Pierce, Rockford, IL, USA) on BIOMAX films (GE, Germany).
Paraformaldehyde-fixed and cryoprotected brains were cut into 30 μm thick slices at ~−80°C using a microtome (Leica Jung HN40) and kept at −20°C in an anti-freeze solution (phosphate buffer 0.50 M in MilliQ water:ethyleneglycol:glycerol = 1.3:1:1) until staining was performed. All immunohistochemical stainings were executed using the free-floating method. Washing steps were carried out between all incubations using washing buffer (TBS pH 7.4 containing 0.2% Triton X-100) at RT. Antigen retrieval was performed when required using the proteinase K antigen retrieval method [incubation of sections in proteinase K solution (proteinase K 20 μg/ml in Tris base 50 mM + EDTA 1.0 mM, pH 8.0)] at 37°C for 7 min. Slices were blocked for 1 h at RT using blocking buffer (5.0% goat serum 5.0% donkey serum in washing buffer). Blocked slices were incubated overnight at 4°C with slight agitation in primary antibody incubation buffer (2.5% goat serum and 2.5% donkey serum in washing buffer) containing the primary antibody/antibodies. Subsequently, secondary antibody incubations were carried out for 2 h at RT. Slices were washed in washing buffer, mounted on chrom-gelatin-coated microscopy slides (SuperFrost Plus, Menzel, Braunschweig, Germany) and glass-covered using Hydromount® (National Diagnostics, Hull, UK).
Fluorescent immunohistochemical images were acquired on a Leica DM4000B microscope using an Olympus DP71 colour digital camera and newCAST software (Visiopharm, Copenhagen, Denmark). Image analysis was carried out with ImageJ software (NIH, USA).
High-resolution imaging was performed using a TCS/SP2 Leica confocal laser scanning microscope (Leica, Wetzlar, Germany) with 63× objective (water, NA: 1.2) where mentioned in the figure legends. All confocal images are maximal intensity projections of z stacks composed of multiple images.
Trypan Blue BBB leakage experiments
Mice received an intraperitoneal injection of 200 μl of a 0.4% Trypan Blue solution in 0.85% saline (Gibco, Switzerland). Thirty minutes after Trypan Blue administration, mice were perfused and their brains processed for histological analysis as described above. Trypan Blue was visualised using immunofluorescence with excitation and emission wavelengths at 642 and 660 nm, respectively. Protocol adapted from Persson et al. .
Prussian Blue and Thioflavin S stain for detection of haemorrhages and CAA
Haemorrhages were visualised using the Prussian Blue stain method on free-floating brain sections. Free-floating brain sections were incubated in a mixture of equal volumes of 10% potassium ferrocyanide (K4Fe(CN)6 trihydrate) in d.d. H2O and 20% hydrochloric acid (HCl) in d.d. H2O for 30 min. Brain sections were subsequently washed with d.d. H2O, counterstained with nuclear fast red solution for 10 min and washed with d.d. H2O. To co-stain for CAA/fibrillar Aβ, sections were subsequently incubated in a 0.25% aqueous solution of potassium permanganate for 20 min, washed, incubated in an aqueous solution of 2% potassium disulphite and 1% oxalic acid for 2 min; washed, incubated in 0.25% acetic acid for 5 s; washed, incubated in a 50% ethanol solution of 0.0125% Thioflavin S; washed and mounted as described above. All chemicals were purchased from Sigma-Aldrich, Switzerland. Prussian Blue was detected using bright field microscopy; ThioS using fluorescence microscopy with excitation and emission wavelengths at 490 and 514 nm, respectively.
Ex vivo astrocyte cultures
Astrocytes were cultured from early and mid-stage pathology TG and arcAβ NTG mice. For the isolation of adult mouse brain astrocytes, mice were deeply anaesthetised using a cocktail of 1.25% ketamine and 0.25% xylazine. Working on ice, meninges and cerebellum + brain stem were carefully removed, the remaining brain tissue minced into small pieces and homogenised in culture medium (DMEM/F12 containing l-glutamine 2 mM, FCS 10%, PenStrep 1.0% (all from Invitrogen, Switzerland)) using a glass homogeniser with Teflon pestle (Wheaton, USA). Brain homogenates were cultured in poly-l-lysine-coated 75 cm2 cell culture flasks (TPP, Switzerland) at 37°C, 5.0% CO2 and 100% humidity. After the cells had reached 80–90% confluence (about 4–6 weeks), the oligodendrocytes layer was detached and removed by shaking of the culture flasks and the medium aspirated. Trypsin/EDTA buffer (Invitrogen, Switzerland) was added to the remaining astrocytes and left for 2 min at 37°C. Culture medium was added and the astrocytes in the medium centrifuged at 1,250 rpm for 10 min. Pellets were homogenised in a fresh culture medium and astrocytes were seeded in poly-l-lysine-coated 6-well culture plates (TPP, Switzerland), 1.0 × 106 cells in 2 ml culture medium per well or 10 cm culture Petri dishes (2.0 × 106 cells in 10 ml culture medium per dish). Cells were grown to 80–90% confluence and split or frozen and stored in liquid nitrogen at −196°C.
Surgery for microdialysis studies was performed in mid-stage pathology arcAβ mice which were deeply anaesthetised by an intraperitoneal injection (100 μl/10 g body weight) of a cocktail consisting of fentanyl 50 μl/ml, midazolam 5.0 mg/ml and medetomidin 1 mg/ml in 0.9% sterile saline. To prevent drying of the cornea, an eye cream (Viscotears®, carbomerum 980, 2.0 mg) was applied onto the eyes. Body temperature was controlled throughout all the surgical procedures by means of a rectal probe connected to a whole body heating pad set at 37°C. Mice were placed in a stereotactic frame and the skull was exposed. A small hole was drilled to allow the implantation of a microdialysis guide cannula (CMA/7, Aurora Borealis Control B.V. Schoonebeek, The Netherlands) in the hippocampus [relative to bregma (in mm): AP: −3.1; L: +3.0; V: −2.0]. Two support screws were placed to hold the guide, which was glued to the skull with dental acrylic cement (Pattern Resin, GC Europe, Switzerland). A subcutaneous injection of the analgesic Metacam (1 μg/g body weight) was administered directly after surgery followed by an antidote (intraperitoneal injection of 100 μl/10 g body weight of a mixture consisting of naloxon 0.4 mg/ml, flumazenil 0.1 mg/ml and atipamezol 5.0 mg/ml in 0.9% sterile saline) to antagonise the anaesthetic.
Eighteen hours prior to the experiment, the microdialysis probe (CMA/7, cut-off 6 kDa, membrane length 1.0 mm, Aurora Borealis Control B.V., Schoonebeek, the Netherlands) was inserted into the guide cannula. The inlet of the microdialysis probe was connected by fluorinated ethylene propylene tubing (FEP tubing, CMA, Sweden) to a syringe pump (Beehive, Bas Technicol, Congleton, UK). The probe was perfused with artificial extracellular fluid (aCSF, composition in mM: NaCl 145; KCl 2.7; CaCl2 1.2; MgCl2 1.0 in a 2 mM phosphate buffer, pH 7.4) at a flow rate of 1 μl/min. The outlet was connected to another piece of FEP tubing with sample vial attached. Eighteen hours after insertion and start of perfusion of the probe, the actual experiment was started (between 08:00 and 09:00 a.m.). To assess the baseline glucose differences between TG and NTG arcAβ mice, 20 samples with 30-min intervals were collected. Each sample vial was put on dry ice directly after the sampling time was over and exchanged for a new, empty one. For glucose uptake measurements, mice were fasted for 12 h prior to the experiment. After collection of three 30-min interval microdialysis samples (for baseline glucose measurements), mice were given an intraperitoneal injection of 250 mg glucose in PBS/kg body weight during continuous microdialysis sampling. Twelve 5-min interval microdialysis samples were collected, followed by four 30-min samples. Fifteen min before the sampling time of the last 30-min interval sample had ended, aCSF perfusate was changed to aCSF containing potassium chloride (KCl) 120 mM using a liquid switch (CMA, Sweden). These 15 min were calculated based on the dead volume of the FEP tubing and microdialysis probe in- and outlet. Two 5-min samples were collected after which an intraperitoneal injection of 250 mg glucose in PBS/kg body weight was given during continuous microdialysis sampling. Twelve more 5-min interval samples were taken followed by two 30-min collections. Each sample vial was put on dry ice immediately after its sampling time was over and exchanged for a new, empty one.
Glucose and lactate assays
Glucose was measured using the Amplex® Red Glucose/Glucose Oxidase Assay Kit from Invitrogen, Switzerland. Microdialysis samples were diluted 1:5 and blood samples 1:20, cell culture media were used undiluted; processing of samples and assay was performed according to the manufacturer’s directions. Lactate assays were performed with an in-house made lactate assay: cell culture media samples were used undiluted whereas microdialysis samples were diluted 1:10 in reaction buffer (KH2PO4 10 mM, pH 6.6, with flavin adenine dinucleotide 0.01 mM) and 50 μl of these samples and 50 μl of lactate calibration standards (l-lactate, Sigma-Aldrich, Switzerland) were mixed with 50 μl lactate assay reaction mix [5 ml lactate reaction mix contained 30 μl Amplex® Red 10 mM (Invitrogen, Switzerland), 80 μl horseradish peroxidase 10 U/ml (Invitrogen, Switzerland), 30 μl lactate oxidase 100 U/ml (Sigma-Aldrich, Switzerland) in 4.86 ml reaction buffer] followed by a 20 min incubation at 37°C. Fluorescence was read using a fluorescence reader (SpectraMax Gemini XS, Bucher Biotec AG, Switzerland). Excitation and emission were set at 540 and 590 nm, respectively.
Primary antibodies used for immunohistochemistry and western blotting: glucose transporter 1 (GLUT1, Alpha Diagnostic, GT11-A; GLUT1, Abcam, ab40084), glucose transporter 3 (GLUT3, Alpha Diagnostic, GT31-A), endothelial cells (CD31/PECAM1, BD Pharmingen, 553370), APP/Aβ (6E10, Covance/Signet, SIG-39300), Aβ (in-house produced antibody recognising fibrillar Aβ), laminin (Sigma-Aldrich, L9393), β actin (Abcam, ab6276), lactate transporter 1/monocarboxylate transporter 1 (MCT1, Alpha Diagnostic, MCT-13A), glial fibrillary acidic protein (GFAP, Advanced Immunochemical Inc., 031223), alpha smooth muscle actin (Sigma-Aldrich, A2547), β dystroglycan (Abcam, ab49515).
Decreased endothelial and astrocytic GLUT1 expression in mid-stage arcAβ mice
Reduced brain baseline glucose concentrations and impaired blood-to-brain glucose uptake in mid-stage arcAβ mice
Impaired lactate release upon neuronal stimulation and reduced expression of the astrocytic lactate transporter MCT1 in mid-stage arcAβ mice
Early-stage astrogliosis precedes wide-spread Aβ plaque burden and is paralleled by increased vascular basement laminin staining and increased soluble laminin secretion by TG arcAβ astrocytes ex vivo
Aβ induces astrocyte endfeet retraction leading to neurovascular uncoupling, starting at early-stage pathology
Loss of astrocytic β dystroglycan expression in late-stage TG arcAβ mice is consistent with decreased interactions between astrocyte endfeet and vasculature and is paralleled by cerebral extravasation of blood-derived IgG
β-amyloid-related pathology of cerebral arteries and arterioles in late-stage arcAβ mice
Cerebral hypoperfusion and small cerebral vessel leakages without significant haemorrhaging in late-stage arcAβ mice
Unlike TBA extravasations, cerebrovascular haemorrhages as detected by Prussian Blue staining were rarely seen but likewise associated with CAA (Fig. 9b, asterisk and 9c) with brain areas presenting abundant CAA, e.g. (pre-) frontal cortex, mainly affected. However, the extent of haemorrhagic lesions did not correlate with the severity degree of CAA. Furthermore, casting resin protrusions were not chronically paralleled by haemorrhages (Fig. 9d, arrows).
CAA-dependent abnormalities of vascular basement membranes and degeneration of smooth muscle cells in late-stage arcAβ mice
Vascular pathology is a critical and more and more recognised component of AD, impairing blood-to-brain influx of nutrients essential for cerebral functioning and brain-to-blood efflux of cerebral waste products leading to cognitive deterioration [10, 11, 64, 65]. By characterising the transgenic arcAβ mouse model of AD—which presents a strong vascular pathology—at early, mid and late disease stages we investigated what processes could possibly instigate the vascular and metabolic impairments early on in the disease and which mechanisms aggravate the vascular pathology at later stages in this AD mouse model. Decreased endothelial GLUT1 expression has been found in transgenic mouse models of AD and in human AD patients [15, 22, 26, 47]. We observed lowered GLUT1 protein expression in TG animals starting at mid-stage disease pathology when CAA formation is apparent in these mice and diffuse Aβ plaques are becoming wide-spread. As GLUT1 is the sole glucose transporter expressed by brain endothelial cells and peripheral supply is the principal source of glucose for the brain , it can be appreciated that a decrease in GLUT1 expression will have significant outcomes on cerebral glucose uptake. Indeed, cerebral imaging data from human subjects show a clear correlation between GLUT1 expression and blood-to-brain glucose uptake . Our findings of decreased brain glucose influx and concentration as measured by in vivo microdialysis in TG arcAβ mice, are in line with these data. The majority of glucose entering the brain via endothelial GLUT1 is subsequently taken up by astrocytes via astrocytic GLUT1 . Astrocyte pathology with altered expression of astrocyte-specific proteins essential for cerebral functioning and metabolism is recognised as a significant factor in AD and subject to intensive research [3, 48]. To understand if a decreased astrocytic GLUT1 expression could also be an underlying factor in the reduced glucose uptake in our microdialysis studies, we cultured astrocytes from the same cohort of arcAβ mice ex vivo. We found that its expression was severely affected, starting at mid-stage disease pathology. Astrocytic glucose uptake is a vital step in the cerebral energy apparatus because astrocytes convert glucose into lactate which is used by neurons during neuronal stimulation [14, 29, 38, 43, 44]. Abnormalities in this essential astrocytic contribution to neurometabolic coupling have detectable outcomes on cerebral performance and are known to occur in AD [3, 50]. The impaired lactate release we observed upon in vivo neuronal stimulation during microdialysis in TG arcAβ mice, was confirmed by a decreased astrocytic MCT1 expression and lactate release deficiency ex vivo. The reciprocal character of the expression profiles of MCT1 and GLUT1 as shown by Maurer and colleagues  makes plausible that the decreased astrocytic MCT1 protein levels could be related to lower astrocytic GLUT1. Astrocytic GLUT1 and MCT1 are mainly expressed on astrocyte endfeet which are crucial for neurovascular coupling [1, 3, 65]. Impaired neurovascular coupling is common in neurodegenerative disorders including AD and AD mouse models with astrocyte endfeet retraction and swelling compromising the cross-talk between astrocytes, blood vessels and neurons and decreased cerebral metabolism as histological and functional hallmarks [3, 37, 61]. We detected signs of neurovascular uncoupling represented by astrocyte endfeet retraction and swelling, possibly also (partly) explaining our findings on impaired metabolism. We observed this starting at an early pathological stage of the disease, at which extracellular Aβ plaque burden is small in the arcAβ mouse. Only astrocytes surrounding diffuse Aβ plaques and vascular Aβ deposits showed endfeet abnormalities even at the late-stage pathology, suggesting that Aβ instigates the loss of neurovascular coupling rather than the process of ageing or the mere presence of the APP transgene in these mice. The mechanism by which Aβ prevents contact between endfeet could be the formation of a physical barrier between blood vessels and astrocytes. Another likely mechanism underlying this impaired astrocyte-endothelium interaction could be the loss of expression of the astrocyte endfeet—VBM linker β dystroglycan as we observed in our TG arcAβ mice [2, 53]. It is unclear whether Aβ toxicity is involved in the attenuated β dystroglycan expression or that the changes in the VBM we describe here and were described by others  are culprit. Interestingly, however, the degree of β dystroglycan loss was paralleled by the presence of diffuse, parenchymal Aβ plaques and leakage of endogenous IgG into the brain. Astrocyte endfeet form an essential part of the BBB  and the question is if detachment of astrocyte endfeet from the vasculature leading to disruption of the BBB and hence extravasation of serum proteins into the CNS or that entry of serum proteins into the brain is responsible for this aspect of neurovascular uncoupling. Brains of multiple sclerosis patients and its in vivo counterpart experimental autoimmune encephalomyelitis show that neuroinflammation and extravasation of components of the peripheral immune system induce astrocyte endfeet abnormalities including retraction and swelling [2, 63]. It is well established that CNS entry of components of the peripheral immune system is part of the AD pathology . Taking into account our findings on the pathological cerebral extravasation of the BBB viability marker Trypan Blue and vessel wall instability as shown by leakage of vascular casting resin in the TG arcAβ mice, a role for components of the peripheral immune system in (further) triggering pathological astrocyte responses including endfeet retraction could indeed be possible [1, 13, 65]. It is clear that with astrocyte endfeet abnormalities already present at the early-stage pathology, impaired BBB functioning is a premature process in the arcAβ mouse. An astrocytic involvement at premature time points in the pathology was further indicated by astrogliosis with paralleled increase in vascular laminin and astrocytic laminin secretion ex vivo in the early-stage mice. Increase in laminin VBM in these relatively young mice with little Aβ plaque burden seems to be contradictory with the unique CAA-related increase in laminin we found in mid-stage and late-stage mice. It is interesting, however, that mainly the incoming arteries from leptomeningeal origin showed this increased astrocyte activation in the early-stage TG mice with concomitant increase in VBM laminin. It is exactly these vessels which are the first to be affected by CAA at later disease stages in the arcAβ mouse and other transgenic AD mouse models [58, 59]. The induction of signal transduction cascades within the endothelium can trigger astrocytic release of endothelium-protective molecules . This protective reaction could be elicited by soluble Aβ species which are known to induce astrocyte activation and to be toxic to endothelial cells and SMCs [18, 51, 56, 60]. Furthermore, laminin expression is known to be up-regulated by damage to the vasculature [4, 65]. Soluble Aβ species mainly as toxic protofibrils are abundantly present in arcAβ mice from 2 months of age onwards [31, 35] and could be directed to the vasculature by the forces of the cerebrovascular drainage pathways in which especially the cerebral arteries are involved [9, 20, 58], providing a possible explanation for the unique location of these reactive astrocytes around blood vessels. Seen in this light, laminin is known to interact with Aβ, inhibiting its fibrillisation and diluting its neurotoxicity [27, 36], giving foundation to the idea that astrocytic laminin release as we detected could be a protective mechanism to both strengthen the vessel wall by increasing VBM thickness and an attempt to reduce Aβ-induced neuro- and vascular toxicity by interfering with (further) Aβ fibrillisation. Although we found astrocytes to secrete laminin as was observed by others as well [8, 45], it cannot be excluded that endothelial cells themselves are involved too as these cells can secrete laminin . Although possibly being an astrocytic protection attempt, an increase in laminin secretion paralleled by VBM thickening leads to vessel wall rigidity and hence reduced CBF, especially the leptomeningeal vessel wall thickening we observed in early-stage TG arcAβ mice which directly affects downstream perfusion of the arterioles and capillaries [23, 30]. Cerebral hypoperfusion—and with that reduced oxygen and glucose supply and hindered cerebral drainage of cerebral waste products—starting at such early age could be at the base of the accumulation of Aβ and the severe CAA-related vascular pathology with vascular SMC loss and BBB impairments as we observed later on at mid- and late-stage pathology in the arcAβ mice.
Together, our data underscore the role of the impaired brain vascular system accompanying β-amyloid pathology [4, 7, 17, 59, 64] and they establish that astrocytes are already involved early on in the disease in arcAβ mice, with further age-related progression in severity. Further, our data suggest that impaired functions of astrocytes related to cerebral energy deficits may contribute to decreased neuronal and vascular functions followed later by pathological stages of β-amyloid pathology in arcAβ mice.
We thank Drs. Antonella Chadha-Santuccione and Zoë Goodger for careful reading of the manuscript and Daniel Schuppli for breeding and caretaking of the arcAβ mice. This work was funded by the Swiss National Science Foundation SNSF, grant number 3200B0-112616/1, and the National Centre of Competence in Research NCCR “Neural Plasticity and Repair”, project 2.
Conflict of interest
The authors declare that they have no conflict of interest.
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