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