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      The blood–brain barrier as an endocrine tissue

      Nature Reviews Endocrinology

      Springer Science and Business Media LLC

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          Pericyte loss influences Alzheimer-like neurodegeneration in mice

          Pericytes are vascular mural cells embedded within the basement membrane of blood microvessels, a site of the blood–brain barrier in vivo 1. Pericytes are uniquely positioned within the neurovascular unit between endothelial cells of brain capillaries, astrocytes and neurons2. Recent studies have shown that pericytes regulate the key neurovascular functions including blood–brain barrier formation and maintenance, vascular stability and angioarchitecture, regulation of capillary blood flow, and clearance of toxic cellular byproducts necessary for normal functioning of the central nervous system3 4 5 6 7. Studies using adult viable pericyte-deficient mice have shown that pericyte loss leads to brain vascular damage by two parallel pathways. The first is blood–brain barrier breakdown associated with brain accumulation of serum proteins and several potentially toxic blood-derived products. The second is the reduction in brain microcirculation causing diminished brain capillary perfusion and tissue hypoxia, ultimately leading to secondary neuronal degenerative changes3 5 7 8. Neurovascular dysfunction8 9 10, microvascular reductions2 8 9 10 and pericyte degeneration and loss8 11 12 13 have been demonstrated in Alzheimer’s disease (AD), a neurodegenerative disorder associated with abnormal elevation of amyloid β-peptide (Aβ)14 15 16, tau pathology17 18 and neuronal loss14 15 16 17 18. Whether pericyte loss can influence the natural course of AD-like neurodegeneration and contribute to disease pathogenesis and accumulation of AD pathology remains, however, unknown. To address this question, we crossed transgenic mice overexpressing the Swedish mutation of human Aβ-precursor protein (APP sw/0 )19 with pericyte-deficient platelet-derived growth factor receptor-β (Pdgfrβ +/− ) mice4 5. APP sw/0 mice develop Aβ elevation, amyloid plaques and correlative memory deficits but do not have significant tau pathology or neuronal loss19 20. Pdgfrβ +/− mice exhibit a moderate but age-dependent progressive loss of brain pericytes because of PDGFRβ deficiency in pericytes that disrupts normal endothelial-derived platelet-derived growth factor B (PDGF-BB) signal transduction to PDGFRβ, regulating pericyte proliferation, migration and recruitment to the vascular wall4 5. Here we show that pericyte loss in APP sw/0 mice elevates brain Aβ levels and accelerates amyloid angiopathy and cerebral β-amyloidosis by diminishing clearance of soluble Aβ from the brain interstitial fluid prior to Aβ deposition. We further show that pericyte deficiency leads to the development of tau pathology and an early neuronal loss that is normally absent in APP sw/0 mice, resulting in accelerated cognitive decline. Thus, pericyte loss has an effect on multiple steps of AD-like neurodegeneration pathogenic cascade in APP sw/0 mice suggesting that pericytes may represent a novel therapeutic target to modify disease progression in AD. Results Pericyte loss accelerates Aβ pathology in APP sw/0 mice APP sw/0 ; Pdgfrβ +/− mice exhibited an accelerated age-dependent loss of pericytes compared with either control APP sw/0 ; Pdgfrβ +/+ mice or Pdgfrβ +/− mice beginning at 1 month of age and reaching within 9 months a significant 55% loss compared with 17 and 26% pericyte losses found in APP sw/0 ; Pdgfrβ +/+ and Pdgfrβ +/− littermates, respectively (Fig. 1a,b). Compared with APP sw/0 ; Pdgfrβ +/+ mice, pericyte-deficient APP sw/0 ; Pdgfrβ +/− littermates developed robust Aβ pathology within 9 months of age including a twofold increase in total human Aβ40 and Aβ42 levels (Fig. 2a,b) and an accelerated development of cerebral amyloid angiopathy and parenchymal amyloid deposition, as illustrated in the cortex by multiphoton imaging of X04-positive amyloid21, indicating fivefold and threefold elevations in cerebral amyloid angiopathy and parenchymal amyloid load, respectively (Fig. 2c–e). Aβ load determined by immunostaining was also increased by five- to sixfold in the cortex and hippocampus of APP sw/0 ; Pdgfrβ +/− compared with APP sw/0 ; Pdgfrβ +/+ mice, respectively (Fig. 2f–h). Surprisingly, loss of pericytes increased by 12- to 15-fold cortical and hippocampal murine endogenous Aβ40 and Aβ42 levels in APP sw/0 ; Pdgfrβ +/− mice normally absent in APP sw/0 mice (Fig. 3a,b). At 9 months of age, murine Aβ contributed to 13 and 30% of total Aβ40 and Aβ42 levels found in brains of APP sw/0 ; Pdgfrβ +/− mice, respectively, compared with <1% in APP sw/0 ; Pdgfrβ +/+ controls (Fig. 3c,d), and co-localized with human Aβ in brain lesions (Fig. 3e). Pathological recruitment of endogenous Aβ has not been shown before in transgenic models of AD. Whether endogenous Aβ has a role in the progression of cerebral β-amyloidosis as reported previously for human Aβ22 23 and/or endogenous prion protein in prion disease24 remains to be determined. Nevertheless, these data indicate that pericytes have important functions in regulating human as well as murine Aβ metabolism. To determine whether loss of the Pdgfrβ allele can affect Aβ accumulation without the interaction with the APP sw/0 gene, we analysed Aβ levels in 9-month-old Pdgfrβ +/− and Pdgfrβ +/+ mice. In both Pdgfrβ +/− and Pdgfrβ +/+ mice, human Aβ was undetectable as expected (Fig. 2a,b) but murine brain Aβ40 and Aβ42 levels were comparable (Fig. 3a,b), suggesting that pericyte-deficient mice do not show a detectable accumulation of Aβ in the absence of the APP gene at an age when age-matched APP sw/0 ; Pdgfrβ +/− mice develop significant increases in both human and endogenous murine Aβ levels compared with their littermate APP sw/0 ; Pdgfrβ +/+ controls (Figs 2a,b and 3a,b). Lack of Aβ elevation in 9-month-old Pdgfrβ +/− mice may suggest that under physiological conditions of Aβ production low murine Aβ levels do not pose a significant challenge for Aβ clearance by pericytes, even when the pericyte pool was reduced by 26% (Fig. 1b). However, in 9-month-old human Aβ-overproducing APP sw/0 ; Pdgfrβ +/− mice, the 2–3 orders of magnitude-higher brain Aβ levels likely go above the clearance capability of the considerably diminished pericyte pool (that is, by 55%, Fig. 1b), resulting in substantial human and murine Aβ accumulations. Pericytes control Aβ clearance in APP mice To determine whether pericyte deficiency affects Aβ clearance, brain interstitial fluid (ISF) Aβ levels were monitored by hippocampal in vivo microdialysis25 26 in 3–4-month-old pericyte-deficient APP sw/0 ; Pdgfrβ +/− mice with a 31% loss of pericytes in the hippocampus; age-matched APP sw/0 ; Pdgfrβ +/+ controls have no loss of pericytes at this time point (Fig. 1b). Our data indicate a significant 2.4- and 2.7-fold increase in the steady-state levels of soluble Aβ40 and Aβ42 in brain ISF of APP sw/0 ; Pdgfrβ +/− mice compared with age-matched APP sw/0 ; Pdgfrβ +/+ littermates, respectively (Fig. 4a,b). The half-life of Aβ40 and Aβ42 in brain ISF26 was increased in APP sw/0 ; Pdgfrβ +/− mice compared with APP sw/0 ; Pdgfrβ +/+ controls from 1.3 to 2.2 h, and 1.5 to 2.4 h, respectively (Fig. 4c), suggesting that the increase in ISF Aβ levels was because of diminished Aβ clearance. Importantly, an increase in Aβ ISF levels preceded Aβ and amyloid deposition in APP sw/0 ; Pdgfrβ +/− mice that were absent at 3–4 months of age (Fig. 4d,e) but were begin to accumulate at 6 months of age (Supplementary Fig. S1a,b) and correlated with a more pronounced 39% loss of pericytes (Fig. 1b). In support of our findings that pericyte deficiency leads to diminished Aβ clearance, we show that Aβ production and processing14 are not affected by pericyte degeneration in APP sw/0 ; Pdgfrβ +/− mice, as indicated by unchanged levels of APP, comparable β-secretase activity and unchanged levels of sAPPβ, a soluble form of APP cleaved by β-secretase (Fig. 4f–h). We next show that primary cultured murine brain pericytes6 rapidly clear extracellular Cy3-labelled Aβ40 via low-density lipoprotein receptor-related protein 1 (LRP1), a key Aβ clearance receptor in brain vasculature8 21 27, that is normally expressed in brain pericytes in vivo (Fig. 5a). LRP1-mediated Aβ clearance by cultured pericytes has been demonstrated by administering antibodies to block the function of specific LDL receptors (Fig. 5b,c) and by quantifying the effects of silencing different LDL receptors with specific siRNA-blocking agents6 (Fig. 5d,e). Moreover, adenoviral-mediated re-expression of human LRP1 minigene rescued the ability of pericytes with siRNA-induced LRP1 knockdown to clear Aβ (Fig. 5e). Excessive LRP1-mediated accumulation of Aβ in pericytes over longer periods of time such as 7 days resulted in cell death (Fig. 5f) similar to as reported in human brain pericyte cultures28. These findings not only suggest that Aβ clearance by pericytes is critical for Aβ homeostasis but also show that extreme Aβ accumulation in pericytes leads to cell death. We next show in vivo that Aβ-overproducing APP sw/0 mice have an age-dependent loss of pericytes from 17% at 9 months of age (Fig. 1b) to 35% at 18 months of age as reported by another study29. Thus, with age Aβ progressively depletes the pericyte pool, thereby increasing the Aβ load on the remaining pericytes, possibly establishing a propagating negative spiral that accelerates disease progression. The relative levels of LRP1 expression in brain microvessels and pericytes isolated from APP sw/0 ; Pdgfrβ +/− mice and APP sw/0 ; Pdgfrβ +/+ littermate controls were comparable (Supplementary Fig. S2a,b), suggesting that LRP1 cellular expression is not altered by partial Pdgfrβ gene deletion in pericytes. However, the number of pericytes was significantly reduced by 31% in 9-month-old APP sw/0 ; Pdgfrβ +/− mice compared with age-matched APP sw/0 ; Pdgfrβ +/+ littermate controls (Fig. 1b). Collectively, these data suggest that total amount of LRP in the pericyte pool that is available for clearance of Aβ in APP sw/0 ; Pdgfrβ +/− mice is significantly diminished compared with APP sw/0 ; Pdgfrβ +/+ mice because of the reduced number of pericytes rather than diminished LRP1 cellular expression in Pdgfrβ-deficient pericytes. APP sw/0 ; Pdgfrβ +/− mice did not show changes in brain microvascular expression of other known Aβ transporters such as P-glycoprotein and receptor for advanced age glycation products2 (Supplementary Fig. S2c,d) or changes in the levels of Aβ-degrading enzymes in the brain—that is, insulin-degrading enzyme and neprilysin—compared with APP sw/0 ; Pdgfrβ +/+ controls (Supplementary Fig. S2e,f), thus ruling out these mechanisms as contributory to increased Aβ levels observed in pericyte-deficient APP sw/0 ; Pdgfrβ +/− mice. The levels of PDGFRβ receptor in brain microvessels of APP sw/0 ; Pdgfrβ +/+ and age-matched Pdgfrβ +/+ littermate controls were comparable, suggesting that accumulation of Aβ does not influence the expression of PDGFRβ (Supplementary Fig. S2g,h). On the other hand, there was ~65% reduction in PDGFRβ in brain microvessels of APP sw/0 ; Pdgfrβ +/− mice compared with APP sw/0 ; Pdgfrβ +/+ controls (Supplementary Fig. S2g,h) indicating that after crossing with Pdgfrβ +/− mice APP sw/0 mice develop a severe PDGFRβ deficiency, which contributes to accelerated pericyte loss compared with more moderate losses in Pdgfrβ +/− mice alone or APP sw/0 mice alone (Fig. 1b). APP sw/0 mice develop high plasma Aβ levels, raising a possibility that plasma Aβ may contribute and increase brain Aβ levels by their transport across the blood–brain barrier30. The exact contributions of plasma-derived and brain-derived Aβ to total brain Aβ in APP sw/0 mice remain, however, unknown. Our data in APP sw/0 ; Pdgfrβ +/− mice compared with APP sw/0 ; Pdgfrβ +/+ littermate controls show a significant increase in the half-life of soluble Aβ in brain ISF after inhibition of Aβ production, suggesting that reduced Aβ clearance26 27 mediates Aβ accumulation in pericyte-deficient APP mice. To validate our findings in an APP model with low plasma Aβ levels, we have performed a limited number of experiments in Pdgfrβ +/− mice crossed with transgenic Dutch/Iowa mice (APP swDI ) expressing low levels of human APP under control of Thy 1.2 neuronal promoter harbouring Swedish mutation and the Dutch and Iowa vasculotropic Aβ mutations27 31. APP swDI mice express human APP exclusively in neurons and have extremely low plasma Aβ levels (<30 pM)31, thereby ruling out a possibility of any significant contribution of plasma Aβ transport to the elevated brain Aβ levels. Notably, APP swDI mice exhibit earlier onset and more robust Aβ pathology than APP sw/0 mice because the Dutch/Iowa Aβ mutant peptides produced by these mice are poorly cleared from the brain and at the blood–brain barrier compared with their respective wild-type Aβ40 and Aβ42 isoforms produced by APP sw/0 mice, as we reported27. After 5 months, pericyte-deficient APP swDI ; Pdgfrβ +/− mice had approximately seven- to eightfold greater Aβ load in the cortex and hippocampus compared with APP swDI ; Pdgfrβ +/+ littermate controls (Fig. 6a,b). These data suggest that pericyte loss worsens Aβ clearance in APP mice regardless of whether plasma Aβ levels are high as in APP sw/0 mice30 or extremely low as in APP swDI mice27 31, and that the effect of pericytes on Aβ clearance from the brain rather than on plasma Aβ has a major role in accelerating Aβ pathology in APP mice. Pericyte loss triggers tau pathology in APP sw/0 mice Next, we studied whether pericyte loss can influence the development of tau pathology and neurodegenerative changes in APP sw/0 mice. Our immunocytochemical analysis in APP sw/0 ; Pdgfrβ +/− mice shows that pericyte loss leads to a significant tau hyperphosphorylation in cortical and hippocampal neurons (Fig. 7a–d), appearance of caspase-cleaved tau in neurons that has been shown to facilitate nucleation-dependent tau filament formation32 and conformational changes in tau as shown with the early pathological tau marker MC132 (Fig. 7e,f) also confirmed using ELISA for insoluble tau (not shown). Notably, changes in tau pathology were not observed in age-matched control APP sw/0 ; Pdgfrβ +/+ mice (Fig. 7a–e) or Pdgfrβ +/− mice (Supplementary Fig. S3a,b) and/or older APP sw/0 ; Pdgfrβ +/+ mice (Supplementary Fig. S3c,d) with brain Aβ40 and Aβ42 levels comparable to those found in younger 9-month-old pericyte-deficient APP sw/0 ; Pdgfrβ +/− mice. These data suggest that Aβ alone and/or moderate pericyte deficiency alone causing vascular damage and blood–brain barrier breakdown5 are not sufficient to trigger early tau pathology, which we show requires a combined action of the two hits as illustrated in APP sw/0 ; Pdgfrβ +/− mice. Pericyte loss leads to early neuronal loss in APP sw/0 mice Importantly, pericyte deficiency led to progressive neuronal degenerative changes as evidenced by ~50% reductions in the neurite density and 23–25% loss of neurons in the cortex and hippocampus in 9-month-old APP sw/0 ; Pdgfrβ +/− mice compared with their respective age-matched APP sw/0 ; Pdgfrβ +/+ littermate controls (Fig. 8a–c). In the absence of the APP transgene, pericyte deficiency leads, however, to only modest 8% loss of neurons in Pdgfrβ +/− mice as reported previously5, whereas APP sw/0 mice did not show neuronal loss at the same age (Fig. 8a,c). Consistent with significant neurodegenerative changes, 9-month-old pericyte-deficient APP sw/0 ; Pdgfrβ +/− mice performed poorly on several hippocampal-dependent behavioural tests including burrowing, nest construction and novel object location compared with their age-matched APP sw/0 ; Pdgfrβ +/+ littermate controls or Pdgfrβ +/− mice of the corresponding age (Fig. 8d–f). At an early 1 month of age, a modest 17% pericyte loss in APP sw/0 ; Pdgfrβ +/− mice compared with no loss in APP sw/0 ; Pdgfrβ +/+ controls (Fig. 1b) did not affect Aβ levels, tau hyperphoshorylation, neurite density and/or number of neurons (Supplementary Fig. S4a–f). Measurements of sensorimotor cortex activity in response to hindlimb stimulation determined by in vivo voltage-sensitive dye (VSD) imaging in 1- to 2-month-old APP sw/0 ; Pdgfrβ +/− mice (Supplementary Fig. S4g–i) indicated no changes in cortical activity as shown by comparable time-lapse imaging profiles of the spreading of depolarization and no changes in the peak of VSD amplitude and/or time-to-peak of the depolarization wave. Consistent with these findings, no behavioural changes were noticed in 1- to 2-month-old APP sw/0 ; Pdgfrβ +/− mice (Supplementary Fig. S4j,k). Vascular damage in pericyte-deficient APP sw/0 mice An early and progressive blood–brain barrier breakdown and microvascular reductions have been described in pericyte-deficient mice3 4 5 and APP sw/0 mice8 33 as well as in AD individuals8 10 11 12 13 and have also been shown at an early stage in 1-month-old pericyte-deficient APP sw/0 ; Pdgfrβ +/− mice compared with APP sw/0 ; Pdgfrβ +/+ control mice, as indicated by a eightfold increase in vascular leakage of immunoglobulin G (Fig. 9a,b) and 29% decrease in the total capillary length (Fig. 9c). At this early stage, APP sw/0 ; Pdgfrβ +/− mice also developed a twofold greater accumulation of immunoglobulin G (IgG) and 12% greater microvascular reductions compared with Pdgfrβ +/− mice, respectively (Fig. 9b,c). This initial vascular damage did not affect, however, neuronal function (Supplementary Fig. S4) that is consistent with previous reports5 6. At a later stage, the degree of blood–brain barrier breakdown as indicated by the magnitude of IgG extravascular deposition and reductions in the capillary length became more pronounced and remained significantly greater in APP sw/0 ; Pdgfrβ +/− mice compared to either APP sw/0 ; Pdgfrβ +/+ mice or Pdgfrβ +/− mice alone (Fig. 9d–f). Significant microvascular reductions in APP sw/0 ; Pdgfrβ +/− mice can reduce the surface area for transvascular and perivascular Aβ clearance, which may additionally contribute to reduced Aβ elimination from the brain. Our findings are consistent with a concept that pericyte degeneration causing accelerated vascular damage from one end, and accelerated Aβ accumulation from the other, creates a double hit in brains of pericyte-deficient APP sw/0 ; Pdgfrβ +/− mice that leads to severe tau pathology (Fig. 7a–f), neuronal loss (Fig. 8a,c) and cognitive changes (Fig. 8d–f). Discussion Collectively, our findings suggest that accelerated pericyte loss in APP sw/0 mice because of aberrant PDGFRβ signalling in pericytes influences several steps within the AD pathogenic cascade by generating multiple negative downstream spiral effects, as illustrated in Fig. 10. From one end, pericyte loss diminishes early in the disease process clearance of soluble Aβ accelerating accumulation and deposition of Aβ in the brain, which in turn self-amplifies Aβ-induced pericyte loss in a negative spiral. From the other end, loss of pericytes disrupts cerebrovascular integrity and leads to microvascular reductions amplifying vascular damage. Together, these two hits act in parallel to lead to the development of a complete spectrum of AD pathology including, in addition to accelerated Aβ accumulation, the appearance of tau pathology, neuronal degeneration and loss. Our data suggest that neither Aβ on its own nor a partial Pdgfrβ genetic deletion alone causing a moderate loss of pericytes4 5 7 can trigger tau pathology and a substantive neuronal loss as seen at an early disease stage in double transgenic APP sw/0 ; Pdgfrβ +/− mice. We and others have previously reported that Pdgfrβ +/− mice do not express detectable levels of PDGFRβ in neurons and/or other cell types of the neurovascular unit including astrocytes and endothelial cells4 5 34 35. In addition, it is well established that loss of PDGF-BB/PDGFRβ signalling because of partial and/or global Pdgfb and/or Pdgfrβ gene deletion or malfunction results in vascular phenotype in the central nervous system caused by loss of pericytes, not neuronal loss3 4 5 35 36. These data suggest that increased Aβ burden, tau pathology and neuronal loss that are observed in double transgenic APP sw/0 ; Pdgfrβ +/− mice cannot be attributed to a downstream effect of PDGFRβ signalling in neurons and/or astrocytes that is independent of vascular pericytes. The present data support the two-hit vascular hypothesis of AD8, suggesting that vascular damage (in the present study a deficiency in vascular pericytes) and Aβ act in parallel to initiate and/or accelerate a chronic neurodegenerative disorder. Interestingly, intracellular accumulation of Aβ alone at the levels that exceed Aβ clearance capability of pericytes leads to pericyte cell death both in vitro and in vivo, which might be particularly relevant to degeneration and loss of pericytes in sporadic AD8 11 12 13. In summary, we show that brain pericytes control an AD-like neurodegenerative process in APP sw/0 mice and therefore may represent a novel therapeutic target to modify disease progression in AD. Future studies should explore whether pericyte rescue by re-expression of the Pdgfrβ gene will slow down Alzheimer-like neurodegeneration cascade in APP sw/0 ; Pdgfrβ +/− mice with accelerated pericyte loss. Although our data show that accumulation of Aβ in APP sw/0 ; Pdgfrβ +/+ mice does not reduce PDGFRβ levels in brain microvessels compared with control Pdgfrβ +/+ mice, future studies should determine whether Aβ accumulation in pericytes in APP sw/0 mice and AD individuals can lead to functional changes in PDGF-BB signal transduction to PDGFRβ in pericytes that might trigger pericyte loss without affecting PDGFRβ levels. Search for molecular cues that cause loss of pericytes in AD models and AD may ultimately lead to the discovery of new therapeutics to control pericyte loss and consequently slow down the pathogenic neurodegeneration cascade in AD. Methods Animals Mice were housed in plastic cages on a 12-h light cycle with ad libitum access to water and a standard laboratory diet. All procedures were approved by the Institutional Animal Care and Use Committee at the University of Southern California and the University of Rochester with National Institutes of Health guidelines. Pdgfrβ +/− was generated as previously described5 37. APP sw/0 mice expressing human APP transgene with the K670M/N671L (Swedish) double mutation under control of the hamster prion promoter19 were crossed with Pdgfrβ +/− mice to generate pericyte-deficient APP sw/0; Pdgfrβ +/− mice and their corresponding littermate controls. In a limited number of experiments, Pdgfrβ +/− mice were crossed with transgenic Dutch/Iowa mice (APP swDI ) expressing low levels of human APP under control of Thy 1.2 neuronal promoter harbouring Swedish mutation and the Dutch and Iowa vasculotropic Aβ mutations27 31. To minimize confounding effects of background heterogeneity, all experiments were performed using age-matched littermates. All the animals were included in the study. In all experiments, we used animals of both sexes. Pdgfrβ +/+, Pdgfrβ +/−, APP sw/0 ; Pdgfrβ +/+ and APP sw/0 ; Pdgfrβ +/− mice were 1, 3, 6 and 9 months old. APP swDI ; Pdgfrβ +/+ and APP swDI ; Pdgfrβ +/− mice were 5 months old. All animals were randomized for their genotype information. All experiments were blinded; the operators responsible for experimental procedure and data analysis were blinded and unaware of group allocation throughout the experiments. Multiphoton imaging One day before imaging animals received an intraperitoneal injection of 10 mg kg−1 methoxy-X04 (Neuroptix, Acton, MA, USA). The following day the mice were anesthetized using initially 5% isoflurane, and then within 15–30 s mice were placed on a heating pad (37 °C) and maintained under anesthesia using a face mask with a continuous delivery of air containing 1.3–1.5% isoflurane. The cranium was firmly secured in a stereotaxic frame. A high-speed dental drill (tip FST 19007-05, Fine Science Tools Inc., Foster City, CA, USA) was used to thin a square cranial window about 2 × 2 mm over the parietal cortex, and 45 degree forceps were used to remove the square piece of skull. Gelfoam (Pharmacia & Upjohn Company, Kalamazoo, MA, USA) was applied immediately to control any cranial or dural bleeding. A sterile 5-mm glass cover slip was then placed on the dura mater and sealed with a 1:1 mixture of bone cement and cyanoacrylate-based glue. Texas-Red-conjugated Dextran (70 kDa; 200 mg kg−1) was injected via tail vein in order to create a fluorescent angiogram. In vivo images were acquired using a Zeiss 5MP multiphoton microscope coupled to a 900-nm mode locked Ti:sapphire laser (Mai Tai, Spectra Physics, Santa Clara, CA, USA)21. Quantification of residual X04 fluorescence was analysed using the NIH Image J software. Confocal microscopy All images were taken with a Zeiss 510 confocal microscopy and analysed using the NIH Image J software5 6 21. Briefly, the number of CD13-positive pericytes and NeuN-positive neurons were analysed using the Image J cell counter tool and expressed per mm2 of total tissue area. The length of capillaries (≤6 μm in diameter) was measured using the Image J ‘Neuro J’ plug-in length analysis tool. The length of capillaries was expressed as mm of lectin-positive vascular profiles per mm3 of brain tissue. To quantify extravascular IgG accumulations, the positive immunofluorescent IgG signals were subjected to threshold-processing and measured using the Image J-integrated density measurement tool. SMI-positive neurofilaments were subjected to threshold-processing and the area of positive neurites was calculated as a percentage of total tissue area. For the quantification of CD13-positive pericyte numbers, capillary length, IgG extravasation, neurite length and NeuN-positive cell numbers six randomly selected fields in the cortex (420 × 420 μm) and four randomly selected fields in the hippocampus (420 × 420 μm) per section from six non-adjacent (~100 μm apart) sections per animal were analysed. At least six animals per group were analysed. Quantification of Aβ load in the cortex and hippocampus was determined by the area occupied by Aβ-positive immunostaining using the NIH Image J software21 38. Tissue immunofluorescent and fluorescent thioflavin-S and lectin staining Mice were anesthetized as described above and transcardially perfused with phosphate buffered saline (PBS) containing 5 U ml−1 heparin. Brains were dissected and embedded into optimal cutting temperature compound (Tissue-Tek, Torrance, CA, USA) on dry ice. Optimal cutting temperature-embedded frozen brain tissue was cryosectioned at a thickness of 14–18 μm and subsequently fixed in ice-cold acetone. Sections were blocked with 5% normal swine serum (Vector Laboratories, Burlingame, CA, USA) for 60 min and incubated in primary antibody diluted in blocking solution overnight at 4 °C. We used the following primary antibodies: rabbit anti-human Aβ (Cell Signaling Technology; no. 8243; 1:200), mouse anti-mouse Aβ (Invitrogen; AMB0062; 1:200), goat anti-mouse PDGFRβ (R&D Systems; AF1042; 1:100), mouse anti-NeuN (Millipore; MAB377; 1:200), mouse anti-neurofilament SMI-311 (Abcam; ab24575; 1:1,000) and goat anti-CD13 (R&D Systems; AF2335; 1:200). Sections were washed in PBS and incubated with the following secondary antibodies: Alexa 488-conjugated donkey anti-rabbit (Invitrogen; A11008; 1:200) to detect human Aβ, Alexa 568-conjugated donkey anti-goat (Invitrogen; A11057; 1:200) to detect mouse neurofilament, PDGFRβ or CD13, Alexa 488-conjugated donkey anti-mouse (Invitrogen; A21202; 1:200) to detect NeuN. To visualize brain microvessels, sections were incubated with Dylight 488-conjugated L. esculentum lectin (Vector Laboratories; DL-1174; 1:100) and coverslipped with fluorescent mounting medium (Dako, Carpinteria, CA, USA). For thioflavin-S staining21 38, frozen brain sections (20 μm) were stained with 0.2% thioflavin-S in PBS for 10 min and washed three times with PBS before imaging. Bright field microscopy analysis Mice were transcardially perfused with 4% paraformaldehyde in 0.1 M PBS. Brains were postfixed in 4% paraformaldehyde overnight at 4 °C and embedded in paraplast. Serial sections were cut at 5 μm using a microtome, mounted on glass slides and rehydrated according to the standard protocols. Mounted slides were pretreated with a citrate buffer, pH 6.0, in a Black & Decker (Hampstead, MD, USA) steamer for 30 min, followed by a 10-min cool down. We used the following primary antibodies: anti-tau pThr231 (Millipore; AB9668; 1:200), anti-phospho-PHF-tau pSer202/Thr205 monoclonal antibody (Thermo Scientific; Clone AT8; MN1020, 1:200), anti-caspase-cleaved tau antibody (Invitrogen; AHB0061; 1:200) and anti-conformation-sensitive tau antibody (Clone MC1, gift of Dr. Peter Davies, Yeshiva University, Bronx, New York, USA, 1:200). Standard 2-day-immunostaining procedures with peroxidase-labelled streptavidin and DAB chromagen were carried out using a Vectastain Elite kit (Vector Laboratories; PK-6100). Hematoxylin counterstaining was used to provide cytological detail. Images were obtained using an inverted microscope (DMI6000B, Leica Microsystems Inc., Buffalo Grove, IL, USA). The number of p-tau (Thr231)-positive neurons and AT8-positive neurons was determined using the Image J software with colour deconvolution plug-in and Cell Counter analysis tools (NIH). Aβ40- and Aβ42-specific ELISA Cortex and hippocampus were dissected and homogenized in ice-cold guanidine buffer (5 M guanidine hydrochloride/50 mM TrisCl, pH 8.0). Aβ40 and Aβ42 levels were determined in samples using human-specific ELISA kits (Invitrogen) according to the manufacturer’s instructions. For human Aβ40 and Aβ42 ELISA, a monoclonal antibody specific against the N-terminus of human Aβ was used as the capturing antibody and a rabbit antibody specific for the C-terminus of either Aβ40 or Aβ42 was used as the detecting antibody21. For mouse Aβ40-specific sandwich ELISA, the capturing and biotinylated detecting antibodies were monoclonal anti-mouse Aβ raised against amino-acid residues 1–20 (Invitrogen, AMB0062) and rabbit polyclonal anti-Aβ40 biotin conjugate (Invitrogen), respectively38 39. For mouse Aβ42-specific sandwich ELISA, the capturing and detecting antibodies were AMB0062 and rabbit polyclonal anti-Aβ42 biotin conjugate (Invitrogen), respectively38 39. Synthetic mouse Aβ40 and Aβ42 (American Peptide Co., Sunnyvale, CA, USA) were used as standards. The lower detection limits for these ELISA assays are 0.5 pmol g−1 of Aβ40 and Aβ42, as reported38 39. Western blotting All samples were lysed in RIPA buffer (50 mM Tris, pH 8.0, 150 mM NaCl, 0.1% SDS, 1.0% NP-40, 0.5% sodium deoxycholate and Roche Applied Science, Indianapolis, IN, USA, protease inhibitor cocktail). Samples were then subjected to SDS–PAGE gel electrophoresis and transferred to a nitrocellulose membrane. Membranes were blocked with 5% milk, incubated with primary antibody and then incubated with the appropriate horse radish peroxidase-conjugated secondary antibody. Membranes were then treated with Immobilon Western ECL detection buffers (Millipore), exposed to CL-XPosure film (Thermo Scientific) and developed in a X-OMAT 3000 RA film processor (Kodak, Rochester, NY, USA). The following antibodies were used: anti-tau phospho Threonine 231, Millipore), anti-LRP-85 (Abcam), anti-RAGE (Santa Cruz Biotechnology Inc.), anti-neprilysin (R&D Systems), anti-IDE (Millipore, AB9210), anti-APP (Millipore) and horse radish peroxidase-conjugated goat anti-mouse IgG (Bio-Rad). Full scans of western blots are provided in Supplementary Fig. S5. Measurement of sAPP-β levels and β-secretase activity sAPP-β levels in the brain of APP sw/0 Pdgfrβ +/+ and APP sw/0 Pdgfrβ +/− mice were measured using the ELISA kit (Covance)39 and β-secretase activity was determined using a β-secretase activity kit (Abcam). In vivo microdialysis and ISF Aβ half-life determination In vivo microdialysis was used to measure soluble Aβ40 and Aβ42 steady-state levels in the hippocampus of awake, freely moving 3- to 4-month-old APP sw/0 Pdgfrβ +/+ and APP sw/0 Pdgfrβ +/− mice. Under isoflurane anesthetic, an intracerebral guide cannula MRB-5 (Bioanalytical Systems, West Lafayette, IN, USA) was stereotaxically implanted into the left hippocampus of the mouse (coordinates: AP −3.1 mm, L +2.4 mm and DV −0.6 mm at a 12° angle). A small head block (Instech laboratories, Plymouth Meeting, PA, USA) that provides tether anchoring to the freely moving system was attached to the skull. The cannula and the head block were cemented into place using dental acrylic. The microdialysis probes had a 2-mm, 38-kDa molecular weight cutoff membrane (Bioanalytical Systems) and were washed with 4% bovine serum albumin -artificial cerebrospinal fluid (aCSF) (Harvard Apparatus, Holliston, MA, USA) before use. After implantation of the guide cannula, the stylet was removed and the microdialysis probe inserted through the guide cannula into the hippocampus. The tethering system connected to a swivel (Instech) and counter balance arm (Instech) allowed unrestricted movement of the animal. The mice were allowed to recover from anesthesia and were housed in the freely moving system with ad libitum access to food and water throughout the experiment. The inlet tract of the microdialysis probe was connected to a PHD 2000 programmable syringe pump (Harvard Apparatus) using FEP tubing (SciPro, Sanborn, NY, USA), and 4% bovine serum albumin-aCSF was perfused continuously at a flow rate of 1 μl min−1. Microdialysates were collected every 60 min into polypropylene tubes in a refrigerated fraction collector (Havard Apparatus). A stable baseline ISF Aβ40 and Aβ42 concentrations were obtained within 4 h followed by an intraperitoneal injection of Compound E (20 mg kg−1, Millipore)25. In order to measure the elimination half-life (t 1/2) of Aβ40 and Aβ42, eight additional 1-h microdialysates were collected. The t 1/2 of Aβ was calculated in the GraphPad Prism 5.0 software using the slope (k′) of the linear regression that included all fractions between drug delivery and when Aβ concentrations plateau (t 1/2=0.693/k, where k=2.303k′)40. The mice were perfused at the end of the experiment and probe-placement was verified. Measurements of soluble human Aβ40 and Aβ42 levels were performed using ELISA as described above. CY3-Aβ40 uptake and internalization by pericytes To isolate brain murine pericytes, isolated microvessel fragments from mouse cortex and hippocampus were digested for 12 h at 37 °C with collagenase A (Roche Applied Science), followed by constant shaking and vigorous pipetting every 3–4 h (ref. 6). The cells were then spun down and washed with PBS and plated in a complete medium containing Dulbecco’s Modified Eagle Medium (DMEM), 10% fetal bovine serum, 1% non-essential amino acids, 1% vitamins and 1% antibiotic/antimycotic on plastic (non-coated) tissue culture plates. After 6–12 h, the non-adherent cells were rinsed away and fresh medium was replaced every 2–3 days. Cultures were confirmed to be morphologically consistent with pericyte cultures and were PDGFRβ-positive, desmin-positive, glial fibrillar acidic protein-negative, aquaporin 4-negative, microtubule-associated protein 2-negative, NeuN-negative, von Willebrand Factor-negative and ionized calcium-binding adapter molecule 1-negative. Primary pericytes were plated into an eight-well chambered coverglass (Nunc, Thermo Scientific) and grown overnight. For Cy3-Aβ40 uptake experiments, pericytes were initially incubated with 1 μM Cy3-Aβ40 in DMEM at 4 °C for 1 h. Unbound Cy3-Aβ40 was removed by several washes with cold DMEM. Cy3-Aβ40 uptake was determined at 37 °C after 30 min incubation with and without 50 μg ml−1 non-immune IgG or specific function-blocking antibodies raised against the extracellular domain of LRP1 (Santa Cruz Biotechnology; sc-16166), LRP2 (a generous gift from Dr Scott Argraves, Medical University of South Carolina), very low-density lipoprotein receptor (R&D Systems; AF2258) and low-density lipoprotein receptor (R&D Systems; AF2255)6. Cy3-Aβ40 (1 μM) internalization and lysosomal colocalization in pericytes were determined 48 h after siRNA silencing of Lrp1, Lrp2, Vldlr, Ldlr, Apoer2 or control siRNA (siCtrl)6. Adenoviral-mediated re-expression of the LRP1 minigene (Ad.mLRP1) or GFP was performed in pericytes with silenced Lrp1. LysoTracker green DND-26 (Invitrogen) was added at 100 nM and incubated for 30 min at the end of the experiment. Cells were then fixed with 4% paraformaldehyde (PFA), washed with PBS, briefly incubated with 0.1% Triton X-100 for 20 s and then stained with Alexa Fluor phalloidin conjugates, F-actin (Invitrogen) and Hoechst 33342 (Invitrogen). Slides were scanned using Zeiss 510 confocal microscope (Carl Zeiss MicroImaging Inc., Thornwood, NY, USA). The Cy3-Aβ40 relative intensity and its lysosomal colocalization in pericytes were measured with the NIH Image J software. Cell death in Aβ-treated pericyte cultures Brain murine pericytes were plated into an eight-well chambered coverglass (Nunc, Thermo Scientific). Cells were cultured for 7 days in the presence and absence of 5 μM Aβ40. Medium with and without 5 μM Aβ40 was replaced after 3 days. Cell viability was quantified using a fluorescent Live/Dead Viability/Cytotoxicity kit according to the manufacturer’s instruction (Invitrogen). In some experiments, cells were treated with anti-LRP1 antibody or siLrp1 for 7 days, as described above. Images were obtained using an inverted microscope (DMI6000B, Leica Microsystems Inc., Buffalo Grove, IL, USA). Data were analysed with the NIH ImageJ counter tool. Behavioural testing For the novel object recognition test41 42, mice were acclimatized to a 25-cm3 cubic box for 10 min, and then exposed at three-time-point trials to two objects affixed to the floor, equidistant to the two nearest walls. Mice were placed in a corner equidistant to both objects, minimizing spatial memory confounds. Starting position for mice was rotated, and objects counterbalanced throughout the test. All trials were videotaped for 5 min. Baseline trial was performed 24 h after acclimatization with two identical objects. A choice trial was performed 90 min after baseline, replacing one familiar object with a novel object and keeping one baseline object constant. Total duration of exploratory approaches to familiar or novel objects was measured and was defined as sniffing or touching an object with the snout at a critical distance of <1 cm from object. The novelty exploration index was calculated as time spent exploring the novel object over total time exploring both objects. To assess burrowing behaviour, mice were individually placed in rat cages equipped with a burrow made from a 200-mm long and 70-mm-diameter tube of polyvinyl chloride (PVC) plastic43. One end of the tube was closed by a PVC cap. The open end of the tube was raised ~30 mm by drilling in two supporting screws44. The burrow was filled with 200 g of mouse food pellets, and the mice were allowed to burrow for 3 h. The weight of the remaining food pellets inside the burrow was determined to obtain a measurement of the amount burrowed. For the younger age group (1-month old), a PVC tube of 50 mm in diameter and 250 g of mouse food were used instead. All other procedures were performed identically to the older mouse groups. To assess nest construction behaviour, mice were individually placed in their home cages with a preweighed nestlet ~1 h before the dark phase. The nests were assessed the next morning and given a score of 1–5 based on the nest construction score41. Any unused nestlet was weighed to determine the percentage of nestlet used. Voltage-sensitive dye imaging of cortical activity For VSD imaging6, a cranial window was placed over the somatosensory cortex using the same procedure as for Multiphoton imaging above. After removing the dura, without causing any cranial bleeding, RH-1692 VSD (Optical Imaging), dissolved in aCSF was applied to the exposed cortex. The brain was washed with aCSF for 5 min, covered with low-melt agarose dissolved in aCSF (~1.3%), sealed with a coverslip and the skull was secured to a custom-built microscopy frame. Images were collected using a Pixelfly CCD camera coupled to the CamWare 3.0 software. RH-1692 was excited using a 627-nm LED light source and imaged using a custom-built Olympus 2 × epifluorescent microscope. Images were collected for 500 ms before and after a mechanical deflection of the hindlimb 5 ms in duration. The responses were averaged from 10–20 trials per animal. Stimulation trial signals were divided by baseline signal profiles collected in the absence of stimulation. The signal intensity was quantified by placing a circular region area of interest over the hindlimb region using the NIH Image software. The change in fluorescent intensity (ΔF/F0) was calculated as a percent change by dividing the signal intensity change after stimulation (ΔF) by the average intensity taken before stimulation (F0)6. The time-lapse ΔF/F0 profiles of VSD signal responses were plotted using ImageJ. The peak amplitude and time-to-peak in fluorescent VSD signal in the hindlimb somatosensory cortex after stimulation were taken from readings of generated VSD time-lapse profiles6. Laser doppler flowmetry Cerebral blood flow responses to vibrissal stimulation in anesthetized 3- to 4-month-old APP sw/0 Pdgfrβ +/+ and APP sw/0 Pdgfrβ +/− mice (750 mg kg−1 urethane and 50 mg kg−1 chloralose) were determined with laser Doppler flowmetry5 21. The tip of the laser Doppler probe (Transonic Systems Inc., Ithaca, NY, USA) was stereotaxically placed 0.5 mm above the dura of the cranial window. The right vibrissae was cut to about 5 mm and stimulated by gentle stroking at 3–4 Hz for 1 min with a cotton-tip applicator. The percentage increase in cerebral blood flow (CBF) because of vibrissal stimulation was obtained from the baseline CBF and stable maximum plateau and was averaged for the three trials. Arterial blood pressure was monitored continuously during the experiment. pH and blood gases were monitored before and after CBF recordings. No significant changes in these physiological parameters were found between different genotypes and ages. Statistical analysis Sample sizes were calculated using nQUERY assuming a two-sided alpha-level of 0.05, 80% power and homogeneous variances for the two samples to be compared, with the means and common s.d. for different parameters predicted from published data and our previous studies. All animals were included in the study. F-test was conducted to ensure that the data meet the assumptions of the tests. The variance was similar between the groups that are statistically compared. Data were analysed using Student’s t-test or using multifactorial analysis of variance followed by Tukey’s post-hoc tests. A P-value <0.05 was considered statistically significant in all studies. Author contributions A.P.S. performed biochemical analyses and in vivo microdialysis experiments. R.D.B. performed in vivo multiphoton and behavioural experiments. Z.Z. performed immunohistochemistry experiments. Q.M. performed tau immunohistochemistry and in vitro pericyte culture experiments. E.A.W. performed experiments. A.R. performed in vivo microdialysis experiments. B.V.Z. designed experiments, analysed data and wrote the paper. Additional information How to cite this article: Sagare, A. P. et al. Pericyte loss influences Alzheimer-like neurodegeneration in mice. Nat. Commun. 4:2932 doi: 10.1038/ncomms3932 (2013). Supplementary Material Supplementary Information Supplementary Figures S1-S5
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            Permeability studies on in vitro blood-brain barrier models: physiology, pathology, and pharmacology.

            (1) The specifically regulated restrictive permeability barrier to cells and molecules is the most important feature of the blood-brain barrier (BBB). The aim of this review was to summarize permeability data obtained on in vitro BBB models by measurement of transendothelial electrical resistance and by calculation of permeability coefficients for paracellular or transendothelial tracers. (2) Results from primary cultures of cerebral microvascular endothelial cells or immortalized cell lines from bovine, human, porcine, and rodent origin are presented. Effects of coculture with astroglia, neurons, mesenchymal cells, blood cells, and conditioned media, as well as physiological influence of serum components, hormones, growth factors, lipids, and lipoproteins on the barrier function are discussed. (3) BBB permeability results gained on in vitro models of pathological conditions including hypoxia and reoxygenation, neurodegenerative diseases, or bacterial and viral infections have been reviewed. Effects of cytokines, vasoactive mediators, and other pathogenic factors on barrier integrity are also detailed. (4) Pharmacological treatments modulating intracellular cyclic nucleotide or calcium levels, and activity of protein kinases, protein tyrosine phosphatases, phospholipases, cyclooxygenases, or lipoxygenases able to change BBB integrity are outlined. Barrier regulation by drugs involved in the metabolism of nitric oxide and reactive oxygen species, as well as influence of miscellaneous treatments are also listed and evaluated. (5) Though recent advances resulted in development of improved in vitro BBB model systems to investigate disease modeling, drug screening, and testing vectors targeting the brain, there is a need for checking validity of permeability models and cautious interpretation of data.
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              Two defects contribute to hypothalamic leptin resistance in mice with diet-induced obesity.

              Obesity in humans and in rodents is usually associated with high circulating leptin levels and leptin resistance. To examine the molecular basis for leptin resistance, we determined the ability of leptin to induce hypothalamic STAT3 (signal transducer and activator of transcription) signaling in C57BL/6J mice fed either low-fat or high-fat diets. In mice fed the low-fat diet, leptin activated STAT3 signaling when administered via the intraperitoneal (ip) or the intracerebroventricular (icv) route, with the half-maximal dose being 30-fold less when given by the icv route. The high-fat diet increased body-weight gain and plasma leptin levels. After 4 weeks on the diet, hypothalamic STAT3 signaling after ip leptin administration was equivalent in both diet groups. In contrast, peripherally administered leptin was completely unable to activate hypothalamic STAT3 signaling, as measured by gel shift assay after 15 weeks of high-fat diet. Despite the absence of detectable signaling after peripheral leptin at 15 weeks, the mice fed the high-fat diet retained the capacity to respond to icv leptin, although the magnitude of STAT3 activation was substantially reduced. These results suggest that leptin resistance induced by a high-fat diet evolves during the course of the diet and has at least two independent causes: an apparent defect in access to sites of action in the hypothalamus that markedly limits the ability of peripheral leptin to activate hypothalamic STAT signaling, and an intracellular signaling defect in leptin-responsive hypothalamic neurons that lies upstream of STAT3 activation.
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                Author and article information

                Journal
                Nature Reviews Endocrinology
                Nat Rev Endocrinol
                Springer Science and Business Media LLC
                1759-5029
                1759-5037
                May 24 2019
                Article
                10.1038/s41574-019-0213-7
                © 2019

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