Cardoso et al. Journal of Neuroinflammation (2015) 12:82
DOI 10.1186/s12974-015-0299-3
RESEARCH
JOURNAL OF
NEUROINFLAMMATION
Open Access
Systemic inflammation in early neonatal mice
induces transient and lasting neurodegenerative
effects
Filipa L Cardoso1, Jasmin Herz2, Adelaide Fernandes1,3, João Rocha1, Bruno Sepodes1, Maria A Brito1,3,
Dorian B McGavern2* and Dora Brites1,3*
Abstract
Background: The inflammatory mediator lipopolysaccharide (LPS) has been shown to induce acute gliosis in
neonatal mice. However, the progressive effects on the murine neurodevelopmental program over the week that
follows systemic inflammation are not known. Thus, we investigated the effects of repeated LPS administration in
the first postnatal week in mice, a condition mimicking sepsis in late preterm infants, on the developing central
nervous system (CNS).
Methods: Systemic inflammation was induced by daily intraperitoneal administration (i.p.) of LPS (6 mg/kg) in
newborn mice from postnatal day (PND) 4 to PND6. The effects on neurodevelopment were examined by staining
the white matter and neurons with Luxol Fast Blue and Cresyl Violet, respectively. The inflammatory response was
assessed by quantifying the expression/activity of matrix metalloproteinases (MMP), toll-like receptor (TLR)-4, high mobility
group box (HMGB)-1, and autotaxin (ATX). In addition, B6 CX3CR1gfp/+ mice combined with cryo-immunofluorescence
were used to determine the acute, delayed, and lasting effects on myelination, microglia, and astrocytes.
Results: LPS administration led to acute body and brain weight loss as well as overt structural changes in the brain such
as cerebellar hypoplasia, neuronal loss/shrinkage, and delayed myelination. The impaired myelination was associated with
alterations in the proliferation and differentiation of NG2 progenitor cells early after LPS administration, rather than with
excessive phagocytosis by CNS myeloid cells. In addition to disruptions in brain architecture, a robust inflammatory
response to LPS was observed. Quantification of inflammatory biomarkers revealed decreased expression of ATX with
concurrent increases in HMGB1, TLR-4, and MMP-9 expression levels. Acute astrogliosis (GFAP+ cells) in the brain
parenchyma and at the microvasculature interface together with parenchymal microgliosis (CX3CR1+ cells) were
also observed. These changes preceded the migration/proliferation of CX3CR1+ cells around the vessels at later
time points and the subsequent loss of GFAP+ astrocytes.
Conclusion: Collectively, our study has uncovered a complex innate inflammatory reaction and associated
structural changes in the brains of neonatal mice challenged peripherally with LPS. These findings may explain
some of the neurobehavioral abnormalities that develop following neonatal sepsis.
Keywords: Astrocytes, Microglia, Myelination, Neurons, Autotaxin, HMGB1, Neurodevelopment, LPS
* Correspondence: mcgavernd@ninds.nih.gov; dbrites@ff.ulisboa.pt
2
National Institute of Neurological Disorders and Stroke, National Institutes of
Health, 10 Center Drive, Bethesda, MD 20892-1430, USA
1
Research Institute for Medicines (iMed.ULisboa), Faculdade de Farmácia,
Universidade de Lisboa, Avenida Professor Gama Pinto, 1649-003 Lisbon,
Portugal
Full list of author information is available at the end of the article
© 2015 Cardoso et al.; licensee BioMed Central. This is an Open Access article distributed under the terms of the Creative
Commons Attribution License (http://creativecommons.org/licenses/by/4.0), which permits unrestricted use, distribution, and
reproduction in any medium, provided the original work is properly credited. 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.
Cardoso et al. Journal of Neuroinflammation (2015) 12:82
Background
Sepsis is a common severe inflammatory response to an
infection that may afflict people at any age. Patients have
a high risk of morbid complications and death. Fatality
rates range from 10% to 20% in sepsis, to 20% to 50% in
severe sepsis, and 40% to 80% in septic shock [1]. Neonatal septicemia (first 28 days of life) is the third leading
cause of death in developed and developing countries,
and a gestational age of less than 32 weeks is considered
to be a risk factor for this disease [2-4]. Induction of
lipopolysaccharide (LPS)-induced sepsis during the first
postnatal week in mice [5,6] reproduces many of the
complications observed in late premature human babies
with septicemia [7]. Systemic administration of LPS, an
endotoxin of gram-negative bacteria, is widely used to
induce a neuroinflammatory response associated with
short-term ‘sickness’ behavior [8] in adult [9] and newborn animals [10] as well as during gestation [11]. In
these models, weight loss is a commonly observed sign
of illness [12-14] and is one of the consequences of sepsis
[15,16]. Across species, sepsis survivors frequently experience white matter damage [17], cerebral palsy [18], as well
as cognitive and affective disorders [19]. A single systemic
LPS injection is intended to reproduce the acute systemic
LPS-mediated inflammation [8], whereas a repetitive challenge is used to model a chronic condition [20,21].
Although LPS entrance into the brain is low [22], acute
systemic inflammation is known to induce transient expression of proinflammatory mediators and microglia activation
but only to a mild extent and without neuronal death [23].
However, in contrast with a single LPS application, repeated
systemic challenge of mice was shown to sustain the microglial inflammatory phenotype, trigger the loss of neurons
[24], and produce changes in cerebral vasculature that include upregulation of the major histocompatibility complex
(MHC) class I and II [20].
How peripheral LPS induces its effects on brain is still
unclear, but mechanisms may involve alterations in the
blood-brain barrier (BBB) permeability and function,
stimulation of LPS receptors located outside the BBB
[22,25], or activation of brain microvascular endothelial
cells (BMECs) through the induction of downstream signaling pathways [26]. BMECs physically separate the
brain from the blood, forming the basis of the BBB.
These cells express the toll-like receptor (TLR)-4, whose
activation by LPS leads to the synthesis and release of
pro-inflammatory cytokines [27,28]. In addition, LPS
triggers the release of activated matrix metalloproteinase
(MMP)-9 and MMP-2 as well as granulocyte-macrophage
colony-stimulating factor, which may contribute to the infiltration of monocytes and to BBB breakdown [29,30].
BMECs are part of the neurovascular unit that also
includes the basement membrane, astrocytic endfeet, pericytes, microglia, neurons, and even oligodendrocytes [31].
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The direct response of these cell types to inflammatory
stimuli or the released pro-inflammatory signals may
exacerbate the damaging effects on BMECs [26,32].
Sensitive time windows for LPS-induced alterations in
neurodevelopment result from the fact that neuronal migration, gliogenesis, and myelinogenesis occur at a late
gestational age and predominate in the first 2 weeks of
postnatal life [33]. Microvessel ensheathment by astrocyte endfeet takes place during the first postnatal week
meaning that some barrier properties should be acquired
after birth, at least in rodents [34]. In addition, LPS triggers astrocytic production of pro-inflammatory cytokines,
particularly in immature cells [35], directly influencing
neurodegeneration [36]. LPS also induces brain-resident
immune cells like microglia to release pro-inflammatory
cytokines and other inflammatory mediators such as the
high-mobility group box 1 (HMGB1) [37] and autotaxin
(ATX) [38] that mediate changes in neuronal network activity and apoptosis [39]. Though microglia progenitors
colonize the mouse brain early in embryogenesis, the
main transition from amoeboid into a ramified shape
occurs during the second week after birth, along with
increased microglial numbers and the maturation of
neurons [28,40]. The morphological features of microglia
and their colonization of the mouse brain are similar in
humans [41].
A robust inflammatory response to LPS is mounted
during the acute phase (first hours), which to some degree
is counterbalanced by an anti-inflammatory response during the later stages [42]. Despite induction of an antiinflammatory program, studies have shown that repeated
injection of LPS (prolonged sepsis) can potentiate proinflammatory cytokine levels in the brain [43]. It is therefore
important to understand the pathogenesis of sepsis and its
sequelae when proinflammatory cytokine levels are sustained in the developing neonatal brain for days in order
to develop novel ways to improve survival and preventing
adverse outcomes. Indeed, most of the studies have been
performed in young or aged mice [20,44-46] instead of the
neonatal period [47]. Thus, we set out to investigate the
effects of repetitive LPS injections during the first postnatal week on murine development. We found that sustained
systemic inflammation interferes with central nervous system (CNS) maturation by causing neuronal atrophy, a
delay in myelination, and acute reactive gliosis.
Materials and methods
Animals
Pregnant CD1 wild-type (WT) mice at embryonic day
(E)16 were purchased from Harlan Ibérica (Spain) laboratories and gave birth in the animal facility of the Faculdade
de Farmácia, Universidade de Lisboa. To better explore
microglia activation, we used heterozygous C57BL/6 (B6)
CX3CR1gfp/+ mice. These mice were generated by crossing
Cardoso et al. Journal of Neuroinflammation (2015) 12:82
B6 WT with B6 CX3CR1gfp/gfp mice that were purchased
from The Jackson Laboratory (Bar Harbor, ME, USA) and
maintained in a closed breeding facility at The National
Institutes of Health (NIH), Bethesda. The insertion of the
green fluorescent protein (GFP) allows the tracking of
CX3CR1+ cells, which is important to visualize microglia
dynamic changes [48,49]. Homozygous B6 CX3CR1gfp/gfp
mice are CX3CR1 deficient and do not respond to fractalkine. On the other hand, both B6 WT and CX3CR1gfp/+
mice respond similarly to LPS [50].
Mice were housed with a 12-h light/dark cycle and were
provided with ad libitum access to a standard laboratory
chow diet and drinking water. This study was carried out in
strict accordance with the recommendations of European
Convention for the Protection of Vertebrate Animals Used
for Experimental and other Scientific Purposes (Council
Directive 86/609/EEC), as well as with those in the Guide
for the Care and Use of Laboratory Animals at the NIH.
The animal study protocol was approved by the NINDS
Animal Care and Use Committee (Assurance Number:
A4149-01). All experimental procedures were performed
under anesthesia, conducted in a manner to minimize
animal suffering, and all efforts were made to use the
minimum number of animals.
Drug administration
The day of birth was defined as PND1. For each strain,
offspring of both genders were randomly divided into
two groups and were treated from PND4 to PND6 with
daily i.p. injections of either endotoxin-free saline [control (W/O LPS); n ≥ 4 per analysis] or of LPS [6 mg/kg,
Escherichia coli 055:B5; Calbiochem (Merck, Darmstadt,
Page 3 of 18
Germany); n ≥ 4 per analysis] to induce systemic inflammation [51,52]. CD1 WT mice were sacrificed 1 day
after the final LPS administration (LPS1) and at LPS9 to
evaluate acute and lasting effects, respectively. B6
CX3CR1gfp/+ mice were sacrificed at LPS1/3/5/6/7/9 not
only to determine the acute and lasting effects but also
the delayed effects. Injection and sampling regimens are
depicted in Figure 1.
Tissue process
For paraffin-histological analysis, CD1 WT mice were
perfused via the ascending aorta with 4% paraformaldehyde (PFA) in PBS. Brains were post-fixed in the indicated
fixative for at least 24 h. Brain tissue was processed for
paraffin and cut into 6-μm sagittal sections. For gelatin
zymography and Western blot analysis, the same animals were perfused via the ascending aorta with phosphate buffer saline solution, pH 7.4 (PBS). Brains were
quickly removed, snap-frozen, and cryopreserved at −80°C
for at least 24 h. Protein extracts were obtained by lysing
the brain tissue with radioimmunoprecipitation assay
(RIPA) buffer (Tris Buffer 1 M pH 8.0, EDTA 0.5 M
pH 8.0, NaCl 5 M, 10% NP-40, 50% glycerol, 10% SDS)
[53]. For cryo-histological analysis, B6 CX3CR1gfp/+ animals were perfused with 4% PFA in PBS. Brains were
post-fixed in the same fixative for 24 h, followed by 15%
and 30% sucrose solutions, each for at least 16 h. Brain tissue was embedded in TFMTM Tissue Freezing Medium
(TBS® Triangle Biomedical Sciences, Durham, NC, USA),
frozen at −80°C and cut with a cryomicrotome into 25-μm
sagittal sections.
Figure 1 Schematic representation of the early induction of systemic inflammation. Offspring of both genders were randomly divided into two
groups and treated with three intraperitoneal injections of either saline solution or lipopolysaccharide (LPS) at days 4, 5, and 6 after birth to
induce systemic inflammation. CD1 wild-type mice were sacrificed at 1 and 9 days after LPS injections, and C57BL/6 (B6) CX3CR1gfp/+ mice were
sacrificed at days 1/3/5/6/7/9 following LPS administration.
Cardoso et al. Journal of Neuroinflammation (2015) 12:82
Staining for Luxol Fast Blue and Cresyl Violet
Paraffin sections were stained with Luxol Fast Blue
(VWR, Radnor, PA, USA) for oligodendrocyte myelin
followed by with Cresyl Violet (Sigma, St. Louis, MO,
USA) for neuronal Nissl bodies’ assessment. Sections
were rehydrated in xylene for 20 min and decreasing
concentrations of ethyl alcohol for 10 min each. Sections
were then incubated with a 0.1% Luxol Fast Blue solution in 70% ethyl alcohol overnight at 56°C. After rinsing
with 70% ethyl alcohol followed by distilled water to remove excess stain, the tissue was differentiated in a 0.5%
lithium carbonate solution. Slides were then rinsed in
distilled water followed by 2% acetic acid solution for
5 min. Next, tissue was counterstained with a 1% Cresyl
Violet solution for 10 min. After rinsing in distilled
water, sections were differentiated in 37.5% acetic acid
and rinsed in distilled water. Slides were mounted with
Fluoromount-G (Southern Biotech, Birmingham, AL,
USA) and visualized using a DFC 490 camera (Leica,
Wetzlar, Germany) adapted to an Axioskop bright field
microscope (Zeiss, Oberkochen, Germany). Width of
cerebellar layers, number of neurons per field, and the
area of their soma were analyzed using ImageJ software
(NIH, Bethesda, MD, USA). Intensity of staining of the
external germinal layer at LPS1 and of the white matter
layer at LPS9 was analyzed using the same software and
was expressed per square micrometers.
Gelatin zymography
Determination of MMP-9 and MMP-2 was evaluated as
previously described [29] with minor alterations. In
short, 40 μg of protein from tissue extracts were analyzed by SDS-PAGE zymography in 0.1% gelatin/10%
acrylamide gels under non-reducing conditions. After
electrophoresis, gels were washed for 1 h with 2.5% Triton X-100 (in 50 mM Tris pH 7.4, 5 mM CaCl2, 1 mM
ZnCl2) to remove SDS and renature the MMPs species
in the gel. Gels were then incubated in developing buffer
(50 mM Tris pH 7.4, 5 mM CaCl2, 1 mM ZnCl2) for
72 h at 37°C to induce gelatin lysis. For enzyme activity
analysis, gels were stained with 0.5% Coomassie Brilliant
Blue R-250 (Bio-Rad, Hercules, CA, USA) for 3 h at
room temperature (RT) and distained in 30% ethanol/
10% acetic acid/H2O. Gelatinase activity, detected as a
white band on a blue background, was quantified by
computerized image analysis using Quantity One 1-D
Analysis Software (Bio-Rad, Hercules, CA, USA).
Western blot
Western blot analysis was performed as previously described [53] with minor alterations. Briefly, 100 μg of
protein from tissue extracts were separated on a 12%
SDS-PAGE gel. Following electrophoretic transfer onto a
nitrocellulose membrane and blocking with 5% milk
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solution, the blots were incubated with primary antibody
overnight at 4°C [rabbit anti-TLR4 (Santa Cruz, Dallas,
TX, USA, #sc-10741; 1:500), mouse anti-HMGB1 (BioLegend, San Diego, CA, USA, #651402; 1:500), rabbit
anti-ATX (Millipore, Billerica, MA, USA, #ABT28; 1:500),
or mouse anti-β-actin (Sigma, USA, #A5441; 1:5,000)] and
with horseradish peroxidase-labeled secondary antibody
[anti-mouse or anti-rabbit (Santa Cruz, USA, #sc-2005
and #sc-2004, respectively; 1:5,000)] for 1 h at RT. Protein
bands were detected by LumiGLO® (Cell Signaling,
Danvers, MA, USA) and visualized by chemiluminescence with ChemiDoc (Bio-Rad, Hercules, CA, USA).
Expression was quantified by computerized image analysis
using Quantity One 1-D Analysis Software (Bio-Rad,
Hercules, CA, USA).
Cryo-immunofluorescence
Frozen sections were used to analyze the expression of
the fractalkine receptor, CX3CR1, as well as NG2+ glia,
GFAP, myelin basic protein (MBP), and cluster of differentiation (CD)31 in B6 CX3CR1gfp/+ mice. Sections were
fixed for 15 min with 1% PFA in PBS. Sections were
then treated with an Avidin/Biotin Blocking Kit (Vector
Laboratories, Burlingame, CA, USA, #SP-2001) per the
manufacturer’s instructions followed by 20-min treatment with Background Buster (INNOVEX Biosciences,
Richmond, CA, USA, #NB306). Tissue was incubated 1 h
at RT with primary antibodies: rabbit anti-NG2 (Millipore,
USA, #AB5320; 1:100), rabbit anti-GFAP (DAKO, Glostrup,
Denmark #Z0334; 1:200), rat anti-MBP (Millipore, Billerica,
MA, USA, #MAB386; 1:100), or Armenian hamster
anti-CD31, clone 2H8 (Chemicon, Temecula, CA, USA,
#MAB1398Z; 1:200). Following the incubation with primary antibodies, sections were washed and incubated for
1 h at RT with secondary antibodies (all from Jackson
ImmunoResearch Laboratories, West Grove, PA, USA)
donkey Alexa Fluor 647 anti-rabbit IgG (H&L) (#711-605152; 1:200), donkey Alexa Fluor 647 F(ab) anti-rat IgG
(H&L) (#712-606-150; 1:200), or goat Rhod-X antiArmenian hamster IgG (H&L) (#127-295-160; 1:200).
CD31 staining was amplified with donkey Rhod-X F(ab)
anti-goat IgG (H&L) (#705-296-147; 1:200) for 1 h at
RT. All working stocks of primary and secondary reagents were diluted in PBS containing 2% fetal bovine
serum (FBS) + 0.5% Triton X-100. Nuclei were counterstained with DAPI dye, and sections were mounted
with IMMU-MOUNT (Thermo-Scientific, Waltham,
MA, USA, #9990402). Between incubations, sections were
washed three times with PBS. Apoptosis was detected in
frozen sections with the ApopTag® Red In Situ Apoptosis
Detection Kit (Chemicon, Temecula, CA, USA, #S7165),
which specifically detects DNA cleavage and chromatin
condensation associated with apoptosis, in accordance with
the manufacturer’s instructions. Images were captured from
Cardoso et al. Journal of Neuroinflammation (2015) 12:82
stained frozen sections using an Olympus FV1200 confocal
microscope equipped with 20× and 40× objectives. Images
were collected using sequential scanning with the 405-,
488-, 559-, and 635-nm laser lines to produce four
color overlays. Cerebellar area was measured in tiles of
DAPI-counterstained brain sections using ImageJ (NIH,
Besthesda, MD, USA). Area fraction and colocalization
of the staining per field of each protein was quantified
by computerized image analysis using ImageJ (NIH,
Besthesda, MD, USA).
Sholl analysis of CX3CR1+ cells
To quantify morphological changes of the CX3CR1+
cells, consecutive Z-stack images were converted to a
maximum intensity projection image by ImageJ software
(NIH Besthesda, MD, USA). Using the Sholl analysis
plugin, concentric circles were created centered on the
soma, beginning at 5.5-μm radii and increasing 2 μm
Page 5 of 18
with every circle. We determined the number of intersections made by microglia branching processes with
each successive increasing circle, the maximum number
of intersections for the cell (Nm), as well as the critical
value at which Nm occurred and the maximum length
at which a branch intersection was observed [54].
Statistical analysis
Results are expressed as means ± SEM from, at least,
four independent animals in each treatment group. Significant differences between groups were determined
by the two-tailed t-test performed on the basis of equal
and unequal variance as appropriate. Comparison of
more than two groups was done by ANOVA using
GraphPad Prism® 5.0 (GraphPad Software, San Diego,
CA, USA). Statistical significance was considered when
P values were lower than 0.05.
Figure 2 Early lipopolysaccharide (LPS) administration causes transient body and brain weight loss and decreases the cerebellar area. (A) Body
and brain of CD1 wild-type mice were weighed at 1 and 9 days post-LPS administration. (B) Body weight of C57BL/6 (B6) CX3CR1gfp/+ mice was
assessed at days 1/3/5/7/9 after LPS injection. (C) Cerebellar area was measured in tiled confocal images of brain sections from B6 CX3CR1gfp/+
mice at days 1/3/5/7/9 post-LPS (representative images with DAPI in (D). Results are mean ± SEM from at least four animals. **P < 0.01 vs. without
(W/O) LPS.
Cardoso et al. Journal of Neuroinflammation (2015) 12:82
Figure 3 (See legend on next page.)
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Cardoso et al. Journal of Neuroinflammation (2015) 12:82
Page 7 of 18
(See figure on previous page.)
Figure 3 Lipopolysaccharide (LPS) administration triggers transient shrinkage of cerebellar layers, acute neuronal loss, and sustained atrophy.
Paraffin sections from CD1 wild-type mice at 1 and 9 days post-LPS administration were stained with Luxol Fast Blue (myelin, blue) followed by
Cresyl Violet (Nissl bodies, purple). (A) Representative images are shown for each condition in the pons, hippocampus, and cerebellum. (B) The
widths of each cerebellar layer [external germinal layer (EGL), molecular layer (ML), Purkinje layer, internal granular layer (IGL), white matter layer
(WML)] were measured. (C,D) Intensity of Cresyl Violet staining in the EGL and of Luxol Fast Blue in WML were quantified per square micrometer
at 1 and 9 days after LPS administration, respectively. (E,F) The number of neurons per field and the area of neuronal cell body (soma) was quantified
throughout the pons, in the CA3 hippocampal region, and in the cerebellar Purkinje layer (PL). All determinations were done using ImageJ software
(NIH, USA). Results are mean ± SEM from at least five animals. *P < 0.05 vs. without (W/O) LPS.
Results
LPS administration in the early neonatal period triggers
acute weight loss and cerebellar hypoplasia
Administration of LPS is known to induce a sickness behavior in adult mice, including weight loss [55,56]. To assess the impact of LPS exposure in the first postnatal week,
we initially assessed body and brain weight oscillations,
which were acutely decreased by LPS (Figure 2A). Body
weight loss was sustained up to LPS7 (P < 0.01) but eventually recovered by LPS9 (Figure 2B). We further explored
acute brain weight loss by measuring the sagittal cerebellar
area. This revealed a disruption in its development evidenced by an approximately twofold reduction at LPS5 that
was still evident at LPS7 (P < 0.01). No difference from the
control group was observed at LPS9 (Figure 2C,D).
Induction of neuronal atrophy in newborn mice by systemic
inflammation is more marked at LPS1 than at LPS9
Given the observed decrease in the cerebellar area, we
next explored the width of each layer: the external germinal layer (with proliferating neuroepithelial cells), the
molecular layer (containing the axons of granule cells
and dendrites of Purkinje cells), the Purkinje neuronal
layer, the granular layer (with small neurons called
granule cells), and the white matter layer (with myelin
fibers) (representative images of cerebellum are shown
in Figure 3A). There was a significant shrinkage of the
neuron-containing layers at 24 h after the last LPS injection (P < 0.05 for the Purkinje layer; P < 0.01 for the
external germinal layer and granular layer) but not at
LPS9 (Figure 3B). In agreement, the density of neurons
based on the intensity of staining per square micrometer of cells in the external germinal layer was also
markedly reduced at LPS1 (approximately twofold, P <
0.05) (Figure 3C). Lastly, a negative impact on the density
of Luxol Fast Blue-labeled myelin fibers in the cerebellum
was evident at LPS9 (Figure 3D, P < 0.05).
We next searched for changes in neuronal density and
morphology by examining brain regions such as the
pons (mediator of cerebellar input and output) [57], the
CA3 subregion of the hippocampus (responsible for
short-term memory) [58], and the Purkinje cell layer that
provides the output of all motor coordination in the
cerebellar cortex [59] (representative images are showed
in Figure 3A). The number of neurons was acutely decreased at LPS1 (P < 0.05), particularly in the pons where
it was still evident at LPS9 (P < 0.05). The neuronal loss
in both the hippocampus and cerebellum seen acutely at
LPS1 was restored by LPS9 (Figure 3E). On the other
hand, the marked soma reduction observed in neurons at
LPS1 persisted until LPS9 in all three regions (Figure 3F,
P < 0.01).
LPS administration leads to a reduced myelination
Given the significant impact on the myelin layer and the
amount of neuronal damage evident in the pons and
cerebellum, we further analyzed the effects of LPS on
myelination in these two brain regions at days LPS1,
LPS3, LPS5, LPS7, and LPS9 (representative images in
Figure 4A). Reduced levels of MBP per unit area were
observed at all time points, but the decrease was particularly evident at LPS5 (minimum values) and LPS9 in both
brain regions when compared to controls (Figure 4B,
P < 0.01). Considering the significant reduction of MBP at
LPS5, we decided to investigate the contribution of
CX3CR1+ microglia to this abnormality. We hypothesized
that their phagocytotic activity might be associated with the
delayed myelination. However, as shown in Figure 4C, this
was not the case, as microglia did not contain cytoplasmic
myelin signal at LPS1 or LPS3. To determine whether
apoptosis of oligodendrocyte precursor cells (OPCs) could
contribute to the decreased myelination, we stained tissues
at LPS1 with ApopTag and anti-NG2 antibodies (expressed
by OPCs) (Figure 4D). Increased apoptosis was observed in
the cerebellum (P < 0.05) (particularly in the external germinal layer) as well as in the pons; however, no overlap of
NG2+ labeling with ApopTag was noticed. Moreover, we
observed an increased number of OPCs in the cerebellum
(fourfold) and pons (1.4-fold) (Figure 4E, P < 0.05). These
data suggest that there may be an increased proliferation of
OPCs, or, alternatively, a delay in their maturation to myelinating cells resulting from neuroinflammation.
Early neonatal LPS administration acutely decreases ATX
levels while increasing other inflammatory biomarkers
We next explored the neuroinflammatory reaction to
systemic LPS injection by quantifying the expression of
inflammatory biomarkers in whole brain tissue. The
Cardoso et al. Journal of Neuroinflammation (2015) 12:82
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Figure 4 Lipopolysaccharide (LPS) administration disrupts myelination in the cerebellum and pons of neonatal mice. Brain cryosections from
C57BL/6 CX3CR1gfp/+ mice at days 1/3/5/7/9 post-LPS administration were immunolabeled for myelin (myelin basic protein, MBP, red). Representative
confocal images of the cerebellum and pons are displayed in (A), where the overlapping of MBP and microglial marker CX3CR1 (green) is seen in yellow.
(B) Area fraction per field of MBP staining was quantified per region using ImageJ software (NIH, USA). (C) To assess microglial phagocytosis,
confocal images at days LPS1 and LPS3 were amplified (overlapping of MBP and CX3CR1 is visible in yellow). Sections of animals at LPS1 were
immunolabeled for apoptosis (ApopTag, red) and oligodendrocytes precursor cells (NG2 cells, green). (D) Cerebellar layers are identified
[external germinal layer (EGL), molecular layer (ML), Purkinje layer, internal granular layer (IGL)]. (E) Area fraction per field of ApopTag and
of NG2 positive stainings were determined using ImageJ software (NIH, USA). *P < 0.05 and **P < 0.01 vs. without (W/O) LPS.
biomarkers MMP-9, MMP-2, TLR4, HMGB1, and ATX
are known to be expressed by astrocytes, microglia, and/
or BMECs during inflammatory conditions. We therefore
evaluated if they were affected by the LPS administration
and, if so, whether the effects were lasting (Figure 5). With
the exception of MMP-2, all biomarkers were acutely
modified by LPS administration. MMP-9 activity was
modestly increased at LPS1 (P < 0.05), whereas TLR4 and
HMGB1 expression levels were more elevated (P < 0.05
and P < 0.01, respectively). ATX expression, on the other
hand, was significantly decreased (P < 0.01). The results indicate that the pro-inflammatory response observed 24 h
after LPS administration was not sustained, as all biomarkers were restored to control levels at LPS9.
Neonatal inflammation decreases the microglia transition
from an amoeboid to ramified morphology
The alterations in inflammatory biomarkers suggested that
microglia might be involved in the response to peripheral
LPS challenge. Therefore, we decided to examine microglial
morphology following neonatal LPS administration in the
pons - the brain region most affected by this endotoxin.
Analysis of parenchymal CX3CR1gfp/+ cells in control mice
revealed that microglia gradually changed morphology from
Cardoso et al. Journal of Neuroinflammation (2015) 12:82
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Figure 5 Lipopolysaccharide (LPS) challenge promotes differential expression of inflammatory biomarkers in the brain. Whole brain lysates of CD1
wild-type mice at 1 day post-lipopolysaccharide (LPS) administration were used to determine the activities of metalloproteinase(MMP)-9 and
MMP-2 by gelatin zymography, alongside with the expression of toll-like receptor (TLR)-4, high-mobility group box 1 (HMGB1) and autotaxin
(ATX) by western blot. Results are expressed as fold-change from animals without (W/O) LPS, and are mean ± SEM from at least four animals in
each group. *P < 0.05 and **P < 0.01 vs. W/O LPS.
an amoeboid phenotype at the end of the first week of life
to a more ramified morphology by the second week
(Figure 6A). This is supported by our quantitative data
showing that the maximum number of microglia
process interactions (process maximum, Nm) and the
distance from the soma where these interactions occurred (critical value) increased over time in control
mice (Figure 6B,C; P < 0.05 for Nm and P < 0.01 for
critical value). However, these parameters were significantly decreased by LPS treatment. In fact, at LPS1 and
LPS3, CX3CR1+ cells were still amoeboid. By LPS5,
microglia showed elongated soma and few secondary
processes, which progressed to further enlargement of
the soma and a reduced number of short processes at
LPS7 and LPS9 (Figure 6A). Quantitatively, at LPS9,
the maximum radius at which a branch intersection
occurred (maximum branch length) was significantly
reduced (25%, P < 0.01). However, no alterations in the
number of branches that originated from microglia
soma (number of primary branches) or in the cell
branching density (Schoenen ramification index) were
noticed (Figure 6B,C).
Decreased astrocytosis occurs after LPS5 and is inversely
associated with microgliosis
To further evaluate the inflammatory reaction to LPS,
we assessed glial responses (astrocytes and microglia, including their interaction with the vasculature) in the
pons (Figure 7A). There was an acute reactive astrogliosis
at LPS1 that extended until LPS3 in the parenchyma and
to LPS5 around the microvessels (P < 0.05). By LPS7 and
LPS9, the GFAP+ staining per unit in both locations was
approximately half that of controls (Figure 7B, P < 0.05 in
parenchyma and P < 0.01 at vasculature). The closest
GFAP+-labeled area fraction to controls was at LPS5,
when astrocytes showed long thin processes as seen in
Figure 7A. Examination of microgliosis revealed that the
area occupied by CX3CR1+ cells was markedly increased
in association with the microvessels at LPS1 (P < 0.01) and
LPS3 (P < 0.05) (Figure 7C). A transient decrease was observed in the pons parenchyma at LPS5, when microglia
showed a dystrophic morphology (Figure 6A), followed by
a remarkable increase at LPS7 and LPS9 (P < 0.05), but
not around the microvessels (Figure 7C). Collectively,
these results suggest that the kinetics of astrocytosis and
microgliosis are inversely correlated in the parenchyma
from LPS1 to LPS9.
Increased density of CX3CR1+ cells around the vessels
precedes the loss of GFAP+ cells in pons
Our data pointed to a transition from astrocyte gain to
loss between LPS5 and LPS7. In order to better understand this transition, we evaluated the glial-vasculature
response at an intermediate time point - LPS6 (Figure 8A).
At this time point, GFAP+ staining per unit in the parenchyma was similar to control and still higher around
the vasculature (Figure 8B, P < 0.01) similar to LPS5
(Figure 7B). On the other hand, the parenchymal area
occupied by CX3CR1+ cells at LPS6 had already increased to control levels (P < 0.05 compared to LPS5).
Surprisingly, the area of microvasculature covered by
these cells was significantly increased at this time point
(Figure 8C, P < 0.01 compared to respective control and
to LPS5), which was not visible either at LPS5 or at
LPS7 (Figure 7C). These data suggest that CX3CR1+
cells migrate towards the microvasculature prior to the
loss of perivascular GFAP+ cells. Lastly, we performed
ApopTag staining at LPS6 to determine if the significant loss of GFAP+ staining at LPS7 was due to apoptosis. No difference in apoptotic cells was observed
between the control and LPS-treated mice at this time
point, nor did the ApopTag staining co-localize with
microglia or astrocytes (Figure 8D,E). Thus, the reduction in GFAP+ staining at LPS7 likely results from a
mechanism other than cell death.
Cardoso et al. Journal of Neuroinflammation (2015) 12:82
Figure 6 (See legend on next page.)
Page 10 of 18
Cardoso et al. Journal of Neuroinflammation (2015) 12:82
Page 11 of 18
(See figure on previous page.)
Figure 6 Morphological changes in microglia following lipopolysaccharide (LPS) administration. (A) Representative confocal images of CX3CR1+
cell morphology from brain cryosections of C57BL/6 CX3CR1gfp/+ mice at days 1/3/5/7/9 post-lipopolysaccharide (LPS) administration are shown.
(B,C) Using ImageJ software (NIH, USA), we performed Scholl analysis (>40 cells per animal) at days 1 and 9 post-LPS administration. Results are
mean ± SEM from at least four animals. *P < 0.05 and **P < 0.01 vs. without (W/O) LPS; §P < 0.05 and §§P < 0.01 vs. day 1 post-LPS.
Discussion
Here, we demonstrate that repeated peripheral LPS administration in the first week of life, a condition that
mimics septicemia in the premature human infants
[7,60,61], alters normal mouse brain development over
the following week. This leads to a delayed recovery of
the LPS-induced neuronal atrophy as well as myelination deficits. These alterations reproduce the neuronal
dysfunction, white-matter damage, and cerebral palsy
associated with human perinatal brain injury following
sepsis [62-66]. We also provide evidence that neonatal
exposure to LPS causes a robust pro-inflammatory reaction in the CNS characterized by astrocyte and
microglia activation followed by astrocyte loss. These
structural and inflammatory changes may explain some
of the CNS abnormalities observed in humans after
neonatal sepsis [67,68].
Our study is novel in that it evaluates the effects of
repetitive LPS injections (6 mg/kg of body weight) in
mice from PND4 to PND6. Most previous studies were
performed in adult or aged mice [20,44-69] and only
one isolated administration of LPS was used [5,70] and
at lower doses [71,72]. Lower LPS concentrations do
not always elicit a septic response, and concentrations
up to 25 mg/kg may be necessary depending on the mouse
model [73]. In our study, we injected a high dose of LPS
daily to induce a sustained septic state. This approach has
been used by other investigators but not during the perinatal period [74,75]. Body weight, myelination, neuronal
density, inflammatory biomarkers, and glial cell reactivity
were evaluated for a week after the last injection to establish the impact of peripheral LPS challenge on the
developing neonatal brain.
An acute loss in body weight following LPS administration is usually indicative of a sickness behavior in newborns
[56], and a three injection regimen with 3 mg/kg in 6to 8-week-old CD-1 mice resulted in significant weight
loss, although the animals survived [75]. Loss of body
weight is one of the consequences of sepsis [15], even in
humans [16], and is a sign of illness in animal models
[12-14]. We observed in our model that body weight decreased immediately following LPS injection. In a study by
Du et al. [76], the effects of LPS in newborn mice were
also studied, although the exposure period was longer
(PND3 to PND11) and less LPS was administered per day
(0.3 mg/kg). The animals examined at PND12 in this study
were shown to have recovered their body weight. In our
model, we observed a decrease in weight over the 7 days
following the last LPS injection. Body weight returned to
normal on day 9. Our findings are in line with a study
showing weight loss in PND5 rats 24 h after LPS injection
(2 mg/kg) [77].
We also observed an acute loss in brain weight following
LPS injection. This is consistent with a study demonstrating that intrauterine administration of LPS (125 μg per
dam) in mice at E15 reduces brain weight relative to
controls, even at PND14 [78]. Again, this appears to be
a dose-dependent effect given that intrauterine administration of lower LPS concentrations (80 μg/kg or approximately 3 μg per dam) did not induce brain weight
loss in offspring at PND14 [79]. Nevertheless, this outcome
is relevant to humans, as MRI scans have revealed that neonatal infection can be associated with changes in cerebral
development, including a reduction in cerebral growth
[66,67]. The overall loss of brain weight in our study may
be linked in part to cerebellar hypoplasia. Several cerebellar
regions were reduced in size following LPS administration,
and our results suggest that this decrease might result
from neuronal atrophy or decreased survival. Recent
data demonstrate that LPS has stronger effect on cell
survival, at least in the hippocampus, than on proliferation
during inflammation in the neonatal mouse brain [80]. In
addition, we observed that there was a decreased soma
area and associated neuronal loss in all brain regions
examined. The aforementioned study by Du et al. [76]
also demonstrated a loss of NeuN+ neurons, although
no change was observed in adult animals 24 h after
LPS treatment (1 mg/kg) [81]. Thus, it is likely that the
negative impact of LPS is more pronounced during
neurogenesis and neuronal migration [7] because the
developing brain is particularly sensitive to inflammation
during this developmental period [80,82].
The negative impact of LPS also extended to the
process of myelination when injected into neonatal mice.
We observed that LPS caused a persistent decrease in
MBP levels even at LPS9. Although most studies have
only evaluated this parameter during the prenatal period
[11,83,84], two recent reports documented hypomyelination following injection of low-dose LPS (0.05 mg/kg) at
PND5 [85] and after repeated low-dose LPS (0.3 mg/kg)
from PND3 to PND11 [76], which are consistent with
our findings. In hypomyelinated mice, we did not observe evidence of phagocytic uptake of degraded myelin
by CX3CR1+ myeloid cells but instead we saw an
increased number of NG2+ cells. We postulate based on
these findings that LPS slows or arrests oligodendrocyte
Cardoso et al. Journal of Neuroinflammation (2015) 12:82
Figure 7 (See legend on next page.)
Page 12 of 18
Cardoso et al. Journal of Neuroinflammation (2015) 12:82
Page 13 of 18
(See figure on previous page.)
Figure 7 Early increased vessel coverage by glia is followed by delayed astrocytic loss and parenchymal microgliosis. Brain cryosections of C57BL/6
CX3CR1gfp/+ mice at days 1/3/5/7/9 post-lipopolysaccharide (LPS) administration were immunolabeled for astrocytes (glial fibrillary acidic protein, GFAP,
red) along with vessel marker CD31 (cluster of differentiation 31, gray). Representative confocal images of glia-endothelium interactions in
pons are displayed in (A). (B,C) Area fraction per field of GFAP and CX3CR1 (green) positive staining, as well as their respective colocalization
with the vessels were determined by ImageJ software (NIH, USA). Results are mean ± SEM from at least four animals. *P < 0.05 and **P < 0.01
vs. without (W/O) LPS.
differentiation instead of promoting myelin degradation.
Additional studies are required to prove such theory.
Assessment of inflammatory biomarkers revealed
that LPS triggers an increased expression of TLR4 and
HMGB1, as well as enhanced MMP-9 activity, at LPS1.
MMPs are gelatinases that have the capacity to remodel
the extracellular matrix, promote cellular invasion, and
induce various signaling pathways [86]. We previously
showed that increased release of active MMP-9 and
MMP-2 by BMECs occurred within 24 h of LPS exposure
in vitro [29]. In vivo, we observed an elevated activation of
MMP-9, but not of MMP-2, following LPS administration.
Previous studies have shown that LPS-stimulated pericytes
and microglia can lead to high levels of active MMP-9
[87], which has the potential to disrupt brain homeostasis,
degrade the extracellular matrix, and ultimately weaken
the BBB, giving rise to leakage [88]. Given that MMP-9
can open the BBB [89] and LPS induces its expression
through the TLR4/NF-κB pathway [90], we investigated
whether this receptor was upregulated after the LPS challenge. We observed increased expression of TLR4 at LPS1
but not at LPS9. TLRs are expressed in immune cells,
microglia, as well as in BMECs and respond to microbial
infections. TLR4 is the receptor responsible for the initial
inflammatory response to LPS and is usually elevated
within hours of exposure [91]. Although few studies have
focused on TLR4 expression in newborn mice, a recent
one reported that TLR4 was elevated 24 h after exposure
to 1 mg/kg of LPS in 6-week-old mice [81], what is in
agreement with our data.
Interestingly, it was demonstrated that HMGB1 triggers
MMP-9 upregulation in neurons and astrocytes predominantly via TLR [92,93]. HMGB1 is a nuclear protein and
an alarmin that is secreted by immune cells and endothelial cells, as well as neurons, microglia, and astrocytes, in
response to an inflammatory stimulus [94]. HMGB1 was
acutely elevated in the brain homogenates at LPS1, which
would potentiate TLR4 signaling. Previous studies have
demonstrated that this biomarker is markedly increased
during neonatal inflammation [95,96]. HMGB1 was shown
to contribute to ‘sickness’ behavior and likely causes
decreased food intake [97]. This could explain the body
weight reduction observed in our study. In contrast to
the increase in MMP-9, TLR4, and HMGB1 levels, a
decrease in ATX expression was noticed at LPS1, despite studies showing that its expression is normally
elevated in inflammatory diseases [98]. ATX has been
associated with an anti-inflammatory and defensive
role, at least in microglia [38,99]. Further studies are
required to determine the exact role played by ATX in
the developing neonatal brain following a peripheral
LPS challenge.
Given the increased expression of inflammatory biomarkers during neonatal sepsis, we decided to evaluate
glial reactivity and their interactions with the brain microvasculature. Both astrocytes and microglia became reactive
and were associated with increased coverage of blood
vessels early after LPS administration. This response
waned over time, with astrocytes eventually showing
reduced vascular interactions. A sustained glial response
to LPS was recently described in the hippocampus and
brainstem of adult mice following induction of sepsis
[100]. In addition, it has been shown that microglia and
astrocytes proliferate in response to LPS [81,101] and E.
coli [102]. Our results are in agreement with the findings
of Gómez-Nicola et al. [103], who showed that LPS administration alone can induce reactivity of both microglia
and astrocytes. It is conceivable that the neuronal atrophy
and delayed myelination observed in our study is in fact
linked to the LPS-induced glial response. Astrocytes and
microglia are essential for the formation, trimming, and
function of developing synapses [104,105], as well as for
CNS myelination by promoting OPC migration, proliferation, and differentiation [106]. Studies have reported that
the loss or dysfunction of astrocytes can lead to demyelination or inhibited oligodendrocyte maturation [107,108].
Others have also shown that disruption of microgliamediated synaptic pruning contributes to neurodevelopmental disorders [109] and produces long-lasting defects
in oligodendrocyte maturation and myelination [110,111].
The divergent early and late astrocytic responses to
LPS administration may be linked to the concept of tertiary
brain damage [112], especially given the role that astrocytes
play in brain homeostasis and BBB maintenance. Sherwin
et al. [113] showed that LPS binds to microglia and astrocytes during LPS-induced neonatal neuroinflammation,
and the response was particularly intense on astrocytes surrounding blood vessels. LPS was shown to increase BBB
resistance by inducing a protective response in endothelial
cells and astrocytes [114]. Therefore, the initial increased
astrocytic coverage of microvessels here reported is in line
with a protective neuroinflammatory reaction. With time,
Cardoso et al. Journal of Neuroinflammation (2015) 12:82
Page 14 of 18
Figure 8 Astrocytic loss in neonatal inflammation is preceded by proliferation/migration of CX3CR1+ cells in the pons. Brain cryosections of
C57BL/6 CX3CR1gfp/+ mice at day 6 post-lipopolysaccharide (LPS) administration were immunolabeled for astrocytes (glial fibrillary acidic protein,
GFAP, red) and brain microvascular endothelial cells (cluster of differentiation, CD31, gray). Representative confocal images of each condition in the
pons are depicted in (A). (B,C) Area fraction per field of GFAP and CX3CR1 (green) positive staining, along with their respective colocalization with
vessels, were determined using ImageJ software (NIH, USA). Representative confocal images of sections immunolabeled for astrocytes (GFAP, red) and
for ApopTag (gray) are depicted in (D). Quantification of ApopTag+ staining area fraction, using the abovementioned software, is shown in (E). Results
are mean ± SEM from at least four animals. **P < 0.01 vs. without (W/O) LPS; §P < 0.05 and §§P < 0.01 vs. day 5 post-LPS.
the upregulation of the chemoattractant CX3CL1 by astrocytes in response to inflammatory mediators [115]
may attract microglia, which is supported by the
increased CX3CR1+ staining and re-distribution we observed. Interestingly, the reduced association of GFAP+
cells with microvessels over time coincided with the
Cardoso et al. Journal of Neuroinflammation (2015) 12:82
morphological transformation of microglia into a bushy,
activated state. This is consistent with studies showing
that reduced astrocytosis may follow a concomitant increase in microgliosis after a neuroinflammatory stimulus
[116,117].
Conclusion
Our data demonstrate that systemic inflammation profoundly alters several anatomical and inflammatory aspects
of the brain when experienced during the early neonatal
period. Cerebellar hypoplasia and neuronal atrophy coincided with reduced myelination, delayed differentiation of
microglia from an amoeboid into the ramified state, and
enhanced glial coverage of cerebral blood vessels. This was
followed by decreased astrocytosis and increased microgliosis. Our results not only expand upon previous literature
but are also the first to document the ongoing progression
of neuroinflammatory changes that occur in the week that
follows repeated LPS administration. Future studies on the
neurodevelopmental outcome and glial response to a second LPS stimulus in young adult mice will help elucidate
tertiary mechanisms of damage and whether or not the
brain becomes sensitized to further injury. These studies
should hopefully aid in the development of therapies that
promote the functional recovery of microglia and astrocytes
in the inflamed brain following neonatal sepsis.
Abbreviations
ATX: autotaxin; BBB: blood-brain barrier; BMECs: brain microvascular
endothelial cells; CD31: cluster of differentiation 31; CNS: central nervous system;
CX3CR1: fractalkine receptor; GFAP: glial fibrillary acid protein; GFP: green
fluorescent protein; HMGB1: high-mobility group box 1; LPS: lipopolysaccharide;
MBP: myelin basic protein; MMP: matrix metalloproteinase; PND: postnatal day;
TLR4: toll-like receptor 4..
Competing interests
The authors declare that they have no competing interests.
Authors’ contributions
DB conceived the study and assisted in the experimental design together
with MAB and DBM. JR and BS helped in the establishment of the neonatal
inflammatory mouse model. FLC conducted the LPS administrations and
performed the experiments. JH and AF assisted in the experimental studies.
FLC, DBM, and DB analyzed the data and drafted the manuscript. All authors
read and approved the final manuscript.
Acknowledgements
This study was supported by the National Institutes of Health (NIH)
intramural program (to DBM), by FEDER (COMPETE Program), and by
National Portuguese funds (Fundação para a Ciência e a Tecnologia - FCT)
through the projects PTDC/SAU-FAR/118787/2010 (to DB) and PEst-OE/SAU/
UI4013/2011-2013 (to iMed.ULisboa). FLC is a recipient of a PhD fellowship
(SFRH/BD/62959/2009) from FCT, a fellowship (2014/CON3/CAN30) from
FLAD and a short-term fellowship (ASTF 31-2014) from EMBO. The funding
organizations had no role in study design, data collection and analysis, decision
to publish, or preparation of the manuscript.
Author details
1
Research Institute for Medicines (iMed.ULisboa), Faculdade de Farmácia,
Universidade de Lisboa, Avenida Professor Gama Pinto, 1649-003 Lisbon,
Portugal. 2National Institute of Neurological Disorders and Stroke, National
Institutes of Health, 10 Center Drive, Bethesda, MD 20892-1430, USA.
3
Department of Biochemistry and Human Biology, Faculdade de Farmácia,
Page 15 of 18
Universidade de Lisboa, Avenida Professor Gama Pinto, 1649-003 Lisbon,
Portugal.
Received: 2 December 2014 Accepted: 10 April 2015
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