Koizumi et al. Journal of Neuroinflammation
https://doi.org/10.1186/s12974-019-1467-7(2019) 16:79

RESEARCH

Open AccessTransiently proliferating perivascular
microglia harbor M1 type and precede
cerebrovascular changes in a chronic
hypertension model
Takashi Koizumi1, Katsutoshi Taguchi2, Ikuko Mizuta1, Hiroe Toba3, Makoto Ohigashi3, Okihiro Onishi4,
Kazuya Ikoma4, Seiji Miyata5, Tetsuo Nakata3, Masaki Tanaka2, Sébastien Foulquier6, Harry W. M. Steinbusch7 andToshiki Mizuno1*

Abstract
Background: Microglia play crucial roles in the maintenance of brain homeostasis. Activated microglia show abiphasic influence, promoting beneficial repair and causing harmful damage via M2 and M1 microglia, respectively.
It is well-known that microglia are initially activated to the M2 state and subsequently switch to the M1 state, called
M2-to-M1 class switching in acute ischemic models. However, the activation process of microglia in chronic andsporadic hypertension remains poorly understood. We aimed to clarify the process using a chronic hypertension
model, the deoxycorticosterone acetate (DOCA)-salt-treated Wistar rats.
Methods: After unilateral nephrectomy, the rats were randomly divided into DOCA-salt, placebo, and control
groups. DOCA-salt rats received a weekly subcutaneous injection of DOCA (40 mg/kg) and were continuously
provided with 1% NaCl in drinking water. Placebo rats received a weekly subcutaneous injection of vehicle andwere provided with tap water. Control rats received no administration of DOCA or NaCl. To investigate thetemporal expression profiles of M1- and M2-specific markers for microglia, the animals were subjected to theimmunohistochemical and biochemical studies after 2, 3, or 4 weeks DOCA-salt treatment.
Results: Hypertension occurred after 2 weeks of DOCA and salt administration, when round-shaped microglia withslightly shortened processes were observed juxtaposed to the vessels, although the histopathological findings werenormal. After 3 weeks of DOCA and salt administration, M1-state perivascular and parenchyma microglia
significantly increased, when local histopathological findings began to be observed but cerebrovascular destruction
did not occur. On the other hand, M2-state microglia were never observed around the vessels at this period.
Interestingly, prior to M1 activation, about 55% of perivascular microglia transiently expressed Ki-67, one of the cellproliferation markers.
(Continued on next page)* Correspondence: mizuno@koto.kpu-m.ac.jp
1
Department of Neurology, Graduate School of Medical Science, Kyoto
Prefectural University of Medicine, 465 Kajii-cho Kamigyo-ku, Kyoto 602-8566,
Japan
Full list of author information is available at the end of the article© The Author(s). 2019 Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0
International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, andreproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to
the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver(http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.

Koizumi et al. Journal of Neuroinflammation(2019) 16:79

Page 2 of 13(Continued from previous page)

Conclusions: We concluded that the resting perivascular microglia directly switched to the pro-inflammatory M1state via a transient proliferative state in DOCA-salt rats. Our results suggest that the activation machinery of
microglia in chronic hypertension differs from acute ischemic models. Proliferative microglia are possible initial key
players in the development of hypertension-induced cerebral vessel damage. Fine-tuning of microglia proliferation
and activation could constitute an innovative therapeutic strategy to prevent its development.
Keywords: Neuroinflammation, Cerebral small vessel disease, Chronic hypertension, Perivascular microglia,
Proliferation

Background
Microglia are the resident immune cells in the brain
and play pivotal roles in environmental surveillance
to maintain brain homeostasis. Inflammation or cellular damage can stimulate microglia to increase the activity of immune functions [1]. In vivo two-photon
microscopy studies showed that activated microglia
rapidly migrate to and accumulate at sites of pathological lesions, such as ischemic lesions [2] or newly
formed amyloid-β plaques [3].
The activated microglia show a biphasic influence,
promoting beneficial repair and causing harmful damage. Those responsible for the former are sometimes referred to as anti-inflammatory M2 microglia and thelatter as pro-inflammatory M1 microglia [4]. These different types of activated microglia can be distinguishedbased on the expression of specific markers. These activated microglia are involved in various neurological disorders [1, 5] and also influence the function andintegrity of the blood-brain barrier (BBB) [6, 7]. In thepresent study, we focused on the interaction between
microglia dynamics and cerebrovascular disease.
According to previous studies, microglia are activated in
the acute phase of ischemic stroke as shown in animalmodels of transient middle cerebral artery occlusion(tMCAO) [8–10]. In the tMCAO model, numbers of
M2-state microglia rapidly increase around vessels in the
penumbra after infarction, and, in a few days, M1-state
microglia dominantly increase. This process is called“M2-to-M1 phenotype-switching” or “shift in the M2-toM1 phenotypes” [11, 12]. In addition to microglia activation, various molecules are involved in the acute phase ofischemic strokes, such as free radicals [13], damage-associated molecular patterns (DAMPs) [14], and T cells [15].
DAMPs include heat shock protein, high-mobility groupbox 1 (HMGB1), and peroxiredoxin [14]. For example,
HMGB1 is produced by ischemic neuronal cells about 2–4
h after an ischemic event and peaks at around 6 h. HMGB1
affects vascular endothelial cells and induces BBB destruction, as well as microglia activation [16].
In contrast, in cerebral small vessel disease (CSVD), only
a limited number of reports referred to M1 and M2 microglia phenotyping and their molecular mechanisms. Chronic

hypertension being the major risk factor of CSVD [17, 18],
we focused on a hypertensive CSVD model.
The aim of this study is to clarify microglia involvement in CSVD caused by chronic hypertension using deoxycorticosterone acetate (DOCA)-salt-treated Wistar
rat (DOCA-salt rat) as a model mimicking sporadic andchronic hypertension.

Methods
Animals

Adult male Wistar rats (150–180 g) were purchasedfrom SHIMIZU Laboratory Supplies Co, Ltd. (Kyoto,
Japan). Protocols were approved by Animal Care and
Use Committees of Kyoto Prefectural University of
Medicine and Kyoto Pharmaceutical University. We
studied Wistar rats fed standard chow and water ad libitum. Care and use of rodents met the standards set by
the National Institutes of Health for experimental animals. The rats were housed under specific pathogen-free
conditions and fed standard laboratory chow and water
ad libitum before entering the study. They were maintained on a 12-h light/day cycle at 20–22 °C and 40–50%
humidity.
Preparation of the chronic hypertension model

Rats were anesthetized by combination anesthesia administered i.p. with 0.375 mg/kg of medetomidine, 2.0
mg/kg of midazolam, and 2.5 mg/kg of butorphanol andunderwent unilateral nephrectomy. After a recovery
period of 7 days, the rats were randomly divided into a
deoxycorticosterone acetate (DOCA)-salt group, placebogroup, and control (Fig. 1a). DOCA-salt group rats received a weekly subcutaneous injection of DOCA
(40mg/kg body weight (Nacalai Tesque, Kyoto, Japan))suspended in carboxymethylcellulose and were provided
with 1% NaCl in drinking water for 2, 3, or 4 weeks(DOCA2W, DOCA3W, and DOCA4W, respectively, n= 3 for each). Placebo group rats received a weekly subcutaneous injection of vehicle and were provided withtap water for 2, 3, or 4 weeks (n = 3 for each period).
Control rats (n = 3) received no administration of DOCA
or NaCl. The rat was placed in a restraint cage in a
warm (38 °C) condition for approximately 2–3 min, thena(2019) 16:79

Page 3 of 13b

Hemi-nephrectomy

One week recovery
periodControl
0W

DOCA4W

DOCA3W

DOCA2W

Control (0W )cPlacebo
2W
3W
4W

DOCA
2W
3W
4W

Systolic blood pressure (mmHg)

Koizumi et al. Journal of Neuroinflammation

240

DOCA
placebo

*

*

200
160
120

0
Control(0W)

Cortex

2W

3W

4W

White matter

1

5

9

13

2

6

10

14

3

7

11

15

4

8

12

16

hematoxylin and eosin*

Klüver-Barrera

Fig. 1 Progression of hypertension and histological damage in deoxycorticosterone acetate (DOCA)-salt rats. a Experimental grouping forhistological analysis of the model animals. b Systolic blood pressure in DOCA2W, DOCA3W, and DOCA4W was compared with that in the control.
Values are expressed as the means ± SEM (n = 3 in each group, *p < 0.05). c Hematoxylin and eosin or Klüver-Barrera staining of brain tissues. In
DOCA3W, focal vascular remodeling including the perivascular space enlargements (open arrowheads) and the formation of vacuoles in thewhite matter (closed arrowheads) appeared. In DOCA4W, cerebral hemorrhage (dotted area) and myelinoclasis lesions were found. Scale bars
500 μm (1–4), 20 μm (5–12), and 40 μm (13–16)

systolic blood pressure and heart rate in a conscious
state were measured by the tail-cuff method (BP-98A-L,
Softron, Tokyo, Japan). The values were measured
three times for each rat and the average value wascalculated. After resting overnight under a lightshield, the rats underwent magnetic resonanceimaging (MRI) under anesthesia with the inhalationof isoflurane. Immediately after MRI, the rats were

perfused transcardially with 4% paraformaldehyde inphosphate buffer, and their brains were removed. For
immuno- or pathological staining, brains were
post-fixed in the same fixative overnight at 4 °C andfurther cryoprotected sequentially in 5, 10, 15, and
25% sucrose. Brains embedded in optimal cutting
temperature compounds were stored at − 20 °C until
examination. Frozen sections of a brain were cut into

Koizumi et al. Journal of Neuroinflammation(2019) 16:79

Page 4 of 13

20-μm-thick slices with a cryostat (CM1850, Leica,
Germany).
For biochemical analysis, fresh brain tissues rapidly
frozen in liquid nitrogen were prepared from the control, DOCA2W, and DOCA3W (n = 4, in each group)
rats under anesthesia.
Evaluation items

We evaluated time-course changes in the blood pressure(Fig. 1b), clinical features (Table 1), histopathology (Fig. 1cand Additional file 1: Figure S2 and S5c), glial fibrillary
acidic protein (GFAP) immunostaining of astrocyte footprocesses as the basic framework of BBB (Additional file 1:
Figure S1), and magnetic resonance imaging (MRI) (Additional file 1: Figure S5a and S5b) of the rats. Histopathological analysis was performed by hematoxylin and eosin(HE) and Klüver-Barrera (KB) staining. To examine themorphological dynamics of microglia in DOCA-salt rats,
we visualized microglia by ionized calcium-binding
adapter molecule 1 (Iba-1) (Figs. 2, 3, 4, and 5), pro-inflammatory M1-state microglia by CD68 (Fig. 3),
anti-inflammatory M2-state microglia by CD206 (Fig. 4),
and proliferative ability of microglia by Ki-67 (Fig. 5). We
identified the vasculature by phalloidin or 4′, 6-diamino-2-phenylindole (DAPI) staining.
Antibodies

Primary antibodies were used against Iba-1 (mouse monoclonal IgG, 1:500 (Millipore) or rabbit polyclonal IgG,
1:1000 (Wako)), inducible nitric oxide synthase (iNOS)(rabbit polyclonal IgG, 1:200 (Abcam)), CD68 (mousemonoclonal IgG, 1:500 (Bio-Rad)), Arginase-1 (rabbit
polyclonal IgG, 1:2000 (Gene Tex)), CD206 (goatpolyclonal IgG, 1:400 (R &D systems)), Ki-67 (rabbit
polyclonal IgG, 1:2000 (Novocas)), and GFAP (mousemonoclonal IgG, 1:10000 (Millipore)). For detection of the
primary antibodies, Alexa488 or Alexa594-conjugatedsecondary antibodies (anti-rabbit or mouse IgG, 1:2000(Thermo Fisher)) were used. Vessel walls were visualized
by Alexa594-conjugated phalloidin (1:500, (Thermo
Fisher)).

ImmunohistochemistryFree-floating sections were permeabilized withphosphate-buffered saline containing 0.1% Tween 20(PBST) for 30 min at room temperature, and then, antigen retrieval was performed with citrate buffer for 20
min at 75 °C or for 15 min at 95 °C (anti-Ki-67 antibody).
After blocking with 5% normal goat serum diluted in
PBST overnight at 4 °C, the sections were incubated withprimary antibody diluted with PBST for 2 days at 4 °C.
For CD206 staining, donkey serum was used for blocking. Following washing with PBST, sections were incubated with appropriate secondary antibodies with DAPI
for 2 h at room temperature. After PBS washing, the sections were mounted on slides with FluorSave Reagent(Merck Millipore, Darmstadt, Germany). For confocal
observation, images were acquired as Z stacks (10–20 z
sections, 1 μm apart, 1024 × 1024 pixels) using a
Plan-Apochromat 63×/1.40 Oil DIC objective (Carl
Zeiss, Oberkochen, Germany) with an inverted
laser-scanning confocal microscope, LSM510 META
(Carl Zeiss). Image analysis was performed using Zeiss
LSM Image Browser. The image acquisition region including the cortical blood vessels was randomly selected.
Ten images per rat were acquired. We identified the vasculature by phalloidin or DAPI staining. Microglia weredistinguished from macrophages by the Iba-1 expression
level and their morphology. The numbers of cells weremanually counted in each image.
Western blotting

Frozen whole-brain tissue was minced and homogenizedwith a Polytron homogenizer in ice-cold PBS (five times
the brain weight) containing phenylmethylsulfonyl fluoride. The samples were then sonicated. Before Western
blot analysis, protein concentrations were determinedusing the Lowry method. We adjusted the protein concentration to 2 mg/mL and loaded 10 μg of the total protein in each well. Proteins were separated by NuPAGE
Bis-Tris 10% gels (Thermo Fisher) and transferred onto
PVDF membranes (Millipore). The membranes were
blocked with Blocking One (Nacalai) for 1 h, followed by

Table 1 Physical profiles of control, placebo-group, and DOCA-salt group rats
Group

Body weight (g)Systolic blood pressure (mmHg)

Heart rate (/min)

Symptoms

Control

0W

165 ± 4.3

113.0 ± 2.8

427.3 ± 38

None

Placebo2W

261 ± 3.9*

116.3 ± 3.4

414.3 ± 31

None

3W

295 ± 16*125.0 ± 3.7

412.3 ± 33

None

4W

330 ± 11*130.0 ± 7.3

413.0 ± 12

None

DOCA

2W

250.3 ± 8.3*

183.3 ± 24*

416.0 ± 50

None

3W

253.0 ± 30*

204.3 ± 18*407.3 ± 9.5

Hemiparesis, bleeding from the tail vein, and loss of appetite

4W

225.0 ± 14*207.0 ± 30*

450.0 ± 29

Severe inactive state

Rats administrated with DOCA-salt or placebo for 2 weeks (DOCA2W or Placebo2W), 3 weeks (DOCA3W or Placebo3W), and 4 weeks (DOCA4W or Placebo4W)
were analyzed. Values are expressed as means ± SEM (n = 3 in each group). Statistical significance was expressed as *p < 0.05 relative to the control by using t testKoizumi et al. Journal of Neuroinflammation(2019) 16:79

Page 5 of 13aControl (0W)

DOCA 2W

DOCA 3W

Iba-1 / Phalloidin

Iba-1 / Phalloidin

Iba-1 / Phalloidinb
DOCA 2W
Iba-1

c

Perivascular microglia
per vessel (number)

CD163 / Iba-1

**

4

CD163

**

3
2
1
0

Control
(0W)

DOCA
2W

DOCA
3W

Fig. 2 Dynamic morphological changes of microglia in DOCA-salt rats. a Microglia and the cerebral vasculature were visualized by Iba-1 (green)and phalloidin (actin, red), respectively, in the control (left column), DOCA2W (middle column), and DOCA3W (right column). The white dottedsquare in the upper row was magnified in the lower row. b Perivascular macrophages (CD163-positive cells, green) showed a flattened shapeand low expression of Iba-1 (open arrowheads). In contrast, microglia had fine processes and showed high expression of Iba-1 (closed
arrowheads). c Perivascular microglia increased in number after DOCA and salt treatments. Significance is expressed as **p < 0.01 using TukeyKramer test. Scale bars 50 μm

overnight incubation at 4 °C with primary antibodies in theblocking solution. Primary antibodies for iNOS (mousemonoclonal IgG (Pharmingen)), Arginase-1 (rabbit polyclonal IgG (Gene Tex)), and α-Tubulin (mouse monoclonal(Novus Biologicals)) were used. After 1-hour incubation ina horseradish peroxidase-conjugated secondary antibody,immunoreactive bands were visualized by enhanced chemiluminescence (ECL prime, GE Healthcare) and ImageQuant LAS4000 mini (GE Healthcare). Intensities of the
bands of interest were quantified using Image J software.

Koizumi et al. Journal of Neuroinflammation(2019) 16:79

Page 6 of 13aControl (0W)

DOCA 2W

DOCA 3W

CD68 / Iba-1 / DAPI

CD68 / Iba-1 / DAPI

CD68 / Iba-1 / DAPI

CD68 / Iba-1

CD68 / Iba-1

CD68 / Iba-1

c
**

100

#

*

80

DOCA-salt

Control
(0W)
1

60

2

3

2W
4

1

2

3

3W
4

1

2

3

4

iNOS
40
α-Tubulin

20
0
Control
(0W)

2W

3W

4W

DOCA-saltdiNOS / α-Tubulin(normalized to the control )CD68- positive microglia
in total microglia (%)

b

5
4

*

3
2
1
0

Control 2W 3W(0W)
DOCA-salt

Fig. 3 M1 switching of microglia in DOCA3W-rats. a Microglia and macrophages were visualized by Iba-1 (red) and CD68 (M1 marker, green),
respectively, in the control (left column), DOCA2W (middle column), and DOCA3W (right column). The white dotted square in the upper row ismagnified in the lower row. CD68-positive microglia were observed in DOCA3W (closed arrowheads) but not in DOCA2W (open arrowheads). In
the white square, only Iba-1 staining was noted. Through the periods, CD68-positive perivascular macrophages existed (arrows). b CD68-positive
microglia increased in number in DOCA3W. c, d The expression level of iNOS protein (M1 microglia marker) was increased after DOCA and salt
treatment. Significant differences are expressed as *p < 0.05 and **p < 0.01 relative to the control rats and #P < 0.05 relative to DOCA2W using
Tukey-Kramer test. The values represent the means ± SEM. Scale bars 50 μm

MRI

Isoflurane-anesthetized rats underwent MRI in a prone
position. The head was kept in a fixed position during
the scanning. The breathing rate was monitored
throughout the experiment. MRI was performed using a

7.04 Tesla (Agilent Technologies, Palo Alto, CA, USA).
T2-weighted contrast images were obtained using thefollowing parameters: echo time = 50 ms, repetition time= 2000 ms, field of view = 2.5 × 2.5 cm2, matrix = 512 ×
512, and slice thickness = 1 mm. To select the imaging

Koizumi et al. Journal of Neuroinflammation(2019) 16:79

Page 7 of 13a
DOCA 2W

DOCA 3W

CD206 / Iba-1 /DAPI

CD206 / Iba-1 / DAPI

CD206 / Iba-1 / DAPI

CD206 / Iba-1

CD206 /
Iba-1

CD206 / Iba-1

bc

DOCA-salt
Control(0W)
1 2 3 4

2W
1

2

3

3W
4

1

2

Arginase-1
α-Tubulin

3

4

Arginase-1 / α-Tubulin(normalized to the control )Control (0W)2
1
0Control 2W
3W(0W)
DOCA-salt

Fig. 4 Absence of M2 switching in DOCA-salt rats. a Microglia and macrophages were visualized by Iba-1 (red) and CD206 (M2 marker, green) in
the control (left column), DOCA2W (middle column), and DOCA3W (right column). The white dotted square in the upper row is magnified in thelower row. CD206-positive microglia were not observed in DOCA-salt rats (open arrowheads). In the white square, only Iba-1 staining was noted.
CD206-positive perivascular macrophages were present in all periods (arrows). b, c No significant increase in the expression level of arginase-1
protein was observed after DOCA and salt treatment. Scale bars 50 μm

position, proton density-weighted images were obtainedusing the following parameters: echo time = 11 ms, repetition time = 2000 ms, field of view = 2.5 × 2.5 cm2,
matrix = 512 × 512, and slice thickness = 1 mm.
Hematoxylin and eosin (HE) or Klüver-Barrera (KB)
staining

HE staining was performed to observe the tissue andvascular changes according to the standard procedure.
Briefly, sections were stained with Mayer’s hematoxylin
for 3 min and then washed in running tap water for 10
min. Thereafter, the sections were stained with eosin for
90 s. These sections were subsequently dehydrated andcleared using alcohol and xylene, respectively. The vascular remodeling structure in HE staining was observed
using microscopy (IX73, Olympus, Tokyo, Japan). We

identified vascular remodeling as perivascular enlargement and vessel wall thickening [19]. The image acquisition region was randomly selected so that a cortical
blood vessel was always included in the image. Ten vessels per rat, that is, 30 vessels per group, were acquired.
All images within an experiment were acquired under
the same microscope settings.
KB staining was performed to observe the demyelination according to the standard procedure. Briefly,
sections were stained with Luxol Fast Blue solution in a
56 °C oven overnight and then washed in 95% alcoholand distilled water. Thereafter, the sections were stainedwith lithium carbonate solution for 30 s. Then, sections
continued to undergo differentiation in 70% alcohol
until the gray matter was clear and the white matter
sharply defined. Next, they were counterstained with

Koizumi et al. Journal of Neuroinflammation(2019) 16:79

Page 8 of 13aControl (0W)

DOCA2W

DOCA3W

DOCA4W

DOCA3W

DOCA4W

Ki-67 / Iba-1

b
Control (0W)

DOCA2W

d
100

*

80

Ki-67- positive
perivascular microglia (%)

c

Ki-67- positive microglia
in total microglia (%)

Ki-67 / CD163
††

** ##

60
40
20
0

Control
(0W)

2W

3W

100

*

80

†

*

60
40
20
0

Control

4W(0W)

DOCA-salt

2W

3W

4W

DOCA-saltfeKi-67 / CD68

Percentage of Ki-67 positive
M1-state microglia (%)

DOCA3W

Ki-67 / CD206

††

** ##

100
80
60
40
20
0

Control

(0W)

2W

3W

4W

DOCA-salt

Fig. 5 Transient increase of proliferative microglia before M1 switching. a Microglia were visualized by Iba-1 (red) and proliferative cells by Ki-67(green). Ki-67-positive microglia were increased in DOCA2W and more markedly increased in DOCA3W (closed arrowheads). In DOCA4W, Ki-67positive microglia were markedly decreased. b CD163-positive perivascular macrophages (red, arrows) were not merged with Ki-67 (green, open
white arrowheads) in all stages. c The rate of Ki-67-positive microglia started to increase in DOCA2W. d The rate of Ki-67-positive perivascular
microglia became significantly higher in DOCA2W. e In some microglia of DOCA3W, Ki-67 was co-expressed with CD68 but not with CD206. f
The rate of proliferative M1-state microglia was significantly higher in DOCA3W. Significant differences are expressed as *p < 0.05 and **p < 0.01
relative to control rats, ##p < 0.01 relative to DOCA2W, and †p < 0.05 and ††p < 0.01 relative to DOCA3W using Tukey-Kramer test. The values
represent the means ± SEM. Scale bars 50 μm

cresyl violet acetate. These sections were subsequently
dehydrated and cleared using alcohol and xylene, respectively. The severity of the white matter lesions wasgraded as reported previously [20]. We also confirmed

the presence of a white matter lesion as the formation ofmarked vacuoles or disappearance of myelinated fibers.
We observed images of every slide by microscopy (IX73,
Olympus, Tokyo, Japan).

Koizumi et al. Journal of Neuroinflammation(2019) 16:79

Statistical analysis

All statistical analyses were performed with JMP12 (SASInstitute, Cary, NC, USA). We used Student’s t test,
Dunnett’s test, or Tukey-Kramer test. Error bars represent the means ± SEM in all figures. A p value of < 0.05
was considered significant.

Results
DOCA-salt-mediated hypertension induces abnormalparenchymal and cerebrovascular morphologies

Compared with control, placebo groups remained
normotensive (Fig. 1b) and did not show any abnormal
findings in histology or MR images (Additional file 1:
Figure S2c and S5). DOCA2W showed a marked elevation of the blood pressure in the absence of any clinical,
histopathological and MRI features of CSVD (Fig. 1b, c,
Additional file 1: Figure S5 and Table 1). DOCA3W
demonstrated several changes in addition to the sustained blood pressure elevation. First, DOCA3W showedclinical symptoms including hemiparesis, a decreasedfood intake, and bleeding from the tail vein (Table 1).
Second, vascular remodeling in the cortex and the formation of vacuoles in the white matter were apparent(Fig. 1c). Third, focal high- and low-intensity areas wereseen on T2-weighted on MR images (Additional file 1:
Figure S5a and S5b). In DOCA4W, we observed shrinkage of the astrocyte foot processes (Additional file 1: Figure S1), a marked decrease in movement, myelin
degeneration, and diffuse high-intensity areas on MR
images, in addition to hypertension.
Morphological changes of microglia precede theappearance of histopathological abnormalities

In the control and placebo groups, resting microglia, morphologically characterized by their fine processes, wereobserved sparsely in the cerebral parenchyma (Fig. 2a, left
column). In DOCA2W, round-shaped microglia withshortened processes were observed juxtaposing vessels in
the cerebral parenchyma (Fig. 2a, middle column), in theabsence of histopathological abnormalities (Fig. 1c).
Thereafter, morphological changes of microglia further
progressed. In DOCA3W, more amoeboid microglia accumulated around structurally altered vessels (Fig. 2a, rightcolumn). In DOCA4W, amoeboid microglia were widespread across the cortex and white matter (Additional file 1:
Figure S3a). The number of microglia juxtaposing vessels
in the cortex was significantly increased in DOCA2W andDOCA3W (Fig. 2c).
We distinguished microglia from perivascular macrophages (PVM) by their Iba-1 intensity and morphology,
based on a previous report that the expression of Iba-1 isweak in macrophages [21]. We confirmed this using amacrophage-specific marker, CD163 [22]. CD163-negative
microglia showed intense Iba-1 immunoreactivity andPage 9 of 13

processes. On the other hand, CD163-positive PVM
showed faint Iba-1 immunoreactivity and a flattened shape(Fig. 2b). Moreover, the distribution of microglia was different from that of the PVM. CD163-positive macrophages were never observed to be accumulated in the
inflammatory lesion (Additional file 1: Figure S3a).
Activated perivascular microglia express a proinflammatory pattern

For characterization of the morphologically activated perivascular microglia, we first studied the expression of CD68
as a pro-inflammatory M1 marker. In the control group,
microglia were CD68-negative (Fig. 3a, left column). Whilemost of the microglia were CD68-negative in DOCA2W(Fig. 3a, middle column), CD68-positive, Iba1-positive
microglia significantly increased around vessel walls andparenchyma in DOCA3W (Fig. 3a, right column). Quantitative analysis showed that the percentage of CD68-positive
microglia increased significantly in DOCA3W comparedwith the control (Fig. 3b). As for the PVM, a sparse distribution pattern of CD68-positive PVMs was similar among
the control, placebo, and DOCA groups (Fig. 3a). Biochemical analysis indicated that the expression level of inducible
nitric oxide synthase (iNOS) as another pro-inflammatory
M1 marker was increased in DOCA3W (Fig. 3c and d).
Next, we studied the expression of CD206 as ananti-inflammatory M2 marker. Perivascular microglia did
not express CD206 in rat brains of any groups (Fig. 4a), except for a few CD206-positive microglia around hemo
rrhage sites (Additional file 1: Figure S3b). As for the PVM,
a sparse distribution pattern of CD206-positive PVMs was
similar among in the control, placebo, and DOCA groups(Fig. 4a). Biochemical analysis also indicated that the expression level of arginase-1, another anti-inflammatory M2marker, was not changed in the rat brains of any groups(Fig. 4b and c). Taken together, direct M1 activation, but
not the M2 state, was identified in our DOCA-salt model.
Activated microglia transiently expressed a cellproliferation marker, Ki-67, prior to M1 switching

The total number of microglia did not change between
the control and placebo groups, whereas it significantly
increased in the DOCA group (Additional file 1: FigureS4a). In the control and placebo groups, no microglia
expressed Ki-67 (Fig. 5a, c and Additional file 1: FigureS4c). The rate of Ki-67-positive microglia significantly
increased to 22% of the total microglia in DOCA2W,
peaked to 54% in DOCA3W, and then decreased to thebaseline level in DOCA4W (Fig. 5a and c). A similar increase in the number of Ki-67-positive microglia wasalso observed (Additional file 1: Figure S4b). The rate of
Ki-67-positive perivascular microglia peaked to 55% of
the total perivascular microglia in DOCA2W and remain
at the same level in DOCA3W (Fig. 5d). Rates of both

Koizumi et al. Journal of Neuroinflammation(2019) 16:79

Page 10 of 13

Ki-67-positive and M1-state microglia were highest in
DOCA3W (Fig. 5e and f). In DOCA4W, proliferative
M1-state microglia markedly decreased (Fig. 5c and d). In
contrast, PVM did not express Ki-67 in any group (Fig. 5b).

Discussion
The dynamics of microglia activation in chronic hypertension model to date have been poorly understood. In thepresent study, we showed microglia undergo dynamic morphological changes in the early stages of chronic hypertension using DOCA-salt-induced hypertension Wistar rats.
At first, proliferative microglia juxtaposed to the cerebral
vessels. Next, they switch to the pro-inflammatory M1state, but not to the anti-inflammatory M2 state (Fig. 6).
On the other hand, dynamic pathological changes of macrophages were not observed in the DOCA-salt rats. In thetMCAO model, the PVM infiltrating from vessels plays
crucial roles in the enhancement of ischemic damage [23,24]. However, this was not observed in our model (Figs. 3aand 4a and Additional file 1: Figure S3a). These results suggest that microglia, rather than PVM, are the key initial
players in the process of cerebral vessel damage in thechronic hypertension model.
Another chronic hypertension model, angiotensin II(AngII)-induced hypertension model mice, showed that
PVMs play important roles in neurovascular regulation

Stage 0Stage 1

Control(0W)

DOCA2W[25]. Administration of AngII reaches the perivascularspace and acts on AngII type 1 receptors on PVMs,
which results in the activation of NADPH oxidase 2
and reactive oxygen species production. These oxidative stresses lead to neurovascular dysfunction within 2
weeks of AngII administration. However, in human,blood-AngII levels are usually normal in benign anduncomplicated essential hypertension [26]. DOCA-salt
rat is characterized by low renin-AngII levels [27, 28],
and BBB permeability is sustained in the early stage (atleast 3 weeks after DOCA-salt administration) of hypertension in DOCA-salt rat [27, 29]. The present study
aimed to evaluate the impact of the progression ofchronic hypertension induced experimentally, in order
to exclude from our study the potential influence ofgenetic factors on microglia density and/or phenotypes
around cerebral vessels. We have therefore selected the
Wistar DOCA-salt hypertension rat model instead ofthe more popular spontaneous hypertension rat (SHR)or stroke-prone spontaneous hypertension rat (SHR-SP)
that carry strong differential genetic backgrounds [30,31]. We, therefore, consider that DOCA-salt-induced
hypertension Wistar rat model is suitable for the studyof chronic hypertension in cerebrovascular diseases.
Microglia activation with a transient proliferative stateand pro-inflammatory M1 state was observed earlier

Stage 2

Stage 3Hypertension

DOCA3W
DOCA4W

Microglia
Ki-67 (+)
CD206 (-)
CD68 (-)

Ki-67 (+)
CD206 (-)
CD68 (+)

Ki-67 (-)
CD206 (-)
CD68 (+)

Perivascular
microglia

Perivascularmacrophage
Ki-67 (-)
CD206 (+)
CD68 (+)

Perivascular spaceenlargement

Fig. 6 Summary of the dynamics of microglia in the process of the chronic hypertensive cerebrovascular disorderKoizumi et al. Journal of Neuroinflammation(2019) 16:79

than cerebral vessel damage in our model. Regarding abnormal findings of the cerebral vasculature, perivascularspace enlargement was observed in DOCA3W, and apparent BBB breakdown identified by astrocyte footshrinkage was noted in DOCA4W (Additional file 1: Figure S1). These results are consistent with a previous
study indicating that BBB permeability remained unchanged in DOCA3W mice using the Evans Blue extravasation method [27].
Regarding microglia dynamics, our findings differed fromthose of previous reports using acute cerebrovascular disease models, such as tMCAO. In the tMCAO model,
microglia were activated to an anti-inflammatory M2 state
followed by a transition to a pro-inflammatory M1 state[11]. The variation in microglia dynamics between chronic
hypertension and acute ischemic models may be due to thedifference of microglia-activating factors. In the tMCAO
model, various factors were produced in the ischemic brain
several hours or days after reperfusion. Interferon regulatory factor 4, known as an M2-switching factor, was rapidly
upregulated within 2 h after ischemic insult [32]. HMGB1,
one of the DAMPs, was produced several hours after ischemic events, and HMGB1 was able to activate microglia[16]. ATP or excessive glutamate was immediately released
from necrotizing neuronal cells and could activate microglia [33]. In a chronic hypertension model, other than the
DOCA-model, a previous study using partial renal artery
occlusion model showed that microglia were activated to
an inflammatory state within 5 weeks after the operation[34]. In this model, chronic hypertension increased the expression of adhesion molecules such as JAM-1, ICAM-1,
and VCAM-1 on the cerebral endothelium and this led todeposition of platelets. Deposited platelets produced
CD40L, which mediated the activation of pro-inflammatory
microglia and activated NFκB and mitogen-activated protein kinase signaling in microglia.
A previous report showed that extracellular signal-regulated kinase (ERK)-activated microglia acquired a proliferative ability and produced mainly pro-inflammatory
cytokines, which cause synaptic and neuronal losses in thebrain and result in lethal neurodegenerative disease in adultmice [35]. This suggests that microglia proliferate prior toactivation of the M1 state. We, therefore, analyzed the expression of Ki-67, as a cell proliferation marker, in microglia
by applying immunostaining to our model. In our model,
proliferative microglia had a close relationship with
M1-state activation. Interestingly, such relationship was recently demonstrated using transgenic mice harboring somatic BRAF mutation, p.V600E. Activation of the MEK-ERK
pathway induces microglia proliferation and is associatedwith an upregulation of pro-inflammatory cytokines fromthe proliferative microglia [35]. The rate of Ki-67-positive
microglia significantly increased in DOCA2W, peaked in
DOCA3W, and then decreased to the baseline inPage 11 of 13

DOCA4W (Fig. 5a and c). On the other hand, the rate of
Ki-67-positive perivascular microglia peaked in DOCA2W
and DOCA3W (Fig. 5d) and similarly decreased to thebaseline in DOCA4W. These results indicate that resting
microglia were preferentially activated around the vessels,
and those perivascular microglia acquired the ability to proliferate earlier than other microglia apart from the vessels.
Our results suggest that specific cytokines released fromthe vessels induced perivascular microglia to enter a proliferative state. Further studies are required to clarify the molecular signaling involved in this phenomenon.
According to our immunohistochemical study, we
concluded that perivascular microglia were from residential microglia. We distinguished microglia from macrophages by Iba-1 intensity and cell morphology. In thepresent study, we did not employ flow cytometry,lineage-specific markers, or other reporter methods.
Flow cytometry succeeded to detect infiltration of macrophages in the ischemic brain by using tMCAO mice inwhich ischemic changes were observed globally inre-perfusion area [15]. However, in DOCA-salt rats,
localization of vascular damages was sparse, and hence,
we performed immunohistochemical analysis rather thanflow cytometry analysis of brain homogenate. According
to a previous study, TMEM119 antigen is expressed specifically in residential microglia in the brain but not inmacrophages [36]. Unfortunately, we failed to detect
TMEM119 in rat brain using antibody raised againstmouse TMEM119 [37]. This may be due to the difference of amino acid sequence in the epitope region between mouse and rat. Finally, reporter method will be a
powerful strategy and may be possible if the DOCA-salt
model is prepared by transgenic mice expressing reporter gene such as EGFP under TMEM promoter. We
like to leave this strategy for our future plan to explore
the process of microglia activation under hypertension.

Conclusions
The present study demonstrates that perivascular microglia proliferate transiently and subsequently underwent
direct M1 switching, prior to cerebral vessel destruction.
Our findings raise the intriguing possibility of a link between perivascular microglia activation and initiation of
cerebrovascular diseases induced by chronic hypertension, and that both anti-hypertensive therapy and thefine-tuning of microglia proliferation might generate asynergistic effect.
Additional file
Additional file 1: Figure S1. Morphological changes of astrocyte footprocesses making up the small vessel wall in DOCA-salt rats. Figure S2.
Quantitative analysis of vessel wall thickening and perivascular space enlargement. Figure S3. Distribution of perivascular macrophages andKoizumi et al. Journal of Neuroinflammation(2019) 16:79

microglia in DOCA4W, and presence of CD206-positive M2-state microglia
around a site of hemorrhage. Figure S4. Quantitative analysis of microglia. Figure S5. Sequential MRI analysis of rat brains. (PPTX 8857 kb)

Abbreviations
AngII: Angiotensin II; BBB: Blood-brain barrier; CSVD: Cerebral small vessel
disease; DAMPs: Damage-associated molecular patterns; DAPI: 4′, 6-Diamino2-phenylindole; DOCA: Deoxycorticosterone acetate; ERK: Extracellular signalregulated kinase; GFAP: Glial fibrillary acidic protein; HE: Hematoxylin andeosin; HMGB1: High-mobility group box 1; Iba-1: Ionized calcium-binding
adapter molecule 1; iNOS: Inducible nitric oxide synthase; KB: Klüver Barrera;
MRI: Magnetic resonance imaging; PBST: Phosphate-buffered saline
containing 0.1% Tween 20; PVMs: Perivascular macrophages;tMCAO: Transient middle cerebral artery occlusion
Acknowledgements
We thank Hiromi Yasuike (Kyoto Prefectural University of Medicine) for
technical support.
Funding
This work was supported by the construction and application of a database
for CADASIL, a hereditary small-vessel disease, by the Japan Agency for Medical Research and Development (AMED), and by a grant-in-aid for Researchon Intractable Disease “Research group on medical infrastructure for adultonset leukoencephalopathy” from the Japanese Ministry of Health, Labour,and Welfare, Japan.
Availability of data and materials
The datasets used and/or analyzed during the current study are available
from the corresponding author on reasonable request.
Authors’ contributions
TK, KT, TN, MT, and MT designed research; TK, KT, HT, MO, OO, and KI
performed research; SM contributed new reagents/analysis tools; TK and KT
analyzed the data; TK, KT, IM, SF, HS, and TM wrote the paper. All authors
read and approved the final manuscript.
Ethics approval and consent to participate
Protocols were approved by Animal Care and Use Committees of Kyoto
Prefectural University of Medicine and Kyoto Pharmaceutical University. Careand use of rodents met the standards set by the National Institutes of Healthfor experimental animals.
Consent for publication
Not applicable.
Competing interests
The authors declare that they have no competing interests.

Publisher’s NoteSpringer Nature remains neutral with regard to jurisdictional claims inpublished maps and institutional affiliations.
Author details
1
Department of Neurology, Graduate School of Medical Science, Kyoto
Prefectural University of Medicine, 465 Kajii-cho Kamigyo-ku, Kyoto 602-8566,
Japan. 2Department of Anatomy and Neurobiology, Graduate School of
Medical Science, Kyoto Prefectural University of Medicine, Kyoto, Japan.
3
Department of Clinical Pharmacology, Division of Pathological Sciences,
Kyoto Pharmaceutical University, Kyoto, Japan. 4Department of Orthopaedics,
Graduate School of Medical Science, Kyoto Prefectural University of Medicine,
Kyoto, Japan. 5Department of Applied Biology, Kyoto Institute of Technology,
Kyoto, Japan. 6Department of Pharmacology and Toxicology, School for
Mental Health and Neuroscience, Maastricht University, Maastricht, theNetherlands. 7Department of Neuroscience, School for Mental Health and
Neuroscience, Maastricht University Medical Center +, Maastricht, The
Netherlands.

Page 12 of 13

Received: 19 July 2018 Accepted: 26 March 2019

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