Background
Parkinson’s disease (PD) is the most common neurodegenerative movement
disorder. It is an age-dependent disease characterized by a severe depletion of
dopamine (DA)-producing neurons in the substantia nigra pars compacta (SNpc) that
project to the striatum. The gradual loss of nigrostriatal pathway function results
in slowly progressing clinical symptoms including tremor, rigidity and slowness of
movement. Despite intensive research, the causes of PD remain unknown. Over the last
decade, a vast number of published studies have indicated that inflammation-derived
oxidative stress and cytokine-dependent neurotoxicity are likely to contribute to
nigrostriatal pathway degeneration [
1‐
5], the pathological hallmark of PD. Specifically, post-mortem analyses of
brains from PD patients confirmed the presence of inflammatory mediators in the area
of the SNpc where maximal destruction of melanin-containing DA-producing neurons
occurs [
6‐
11]. Signs of inflammation included activated microglia and accumulation of
cytokines (including TNF, IL-1β, IL-6, and IFNγ), some of which exert
neurotoxic effects on DA neurons [
5]. In addition to extrinsic factors, SNpc dopaminergic neurons may be
uniquely vulnerable to neuroinflammatory insults that enhance cellular oxidative
stress. For example, the higher sensitivity of nigral DA neurons to injury induced
by neuroinflammatory mediators may be secondary to a reduction of endogenous
anti-oxidant capacity (such as glutathione depletion). Pharmacologically, chronic
infusion of various anti-inflammatory compounds (including COX-2-selective NSAIDs or
soluble TNF-selective inhibitors) rescues nigral DA neurons from progressive
degeneration and death [
12‐
14]. These findings raise the interesting possibility that environmental
triggers may initiate cytokine-driven neuroinflammation and may contribute to the
development of PD in humans.
Monogenic forms of PD have been linked to loss-of-function mutations in a number of
genes, giving rise to autosomal recessive parkinsonism [
15], including mutations in
parkin, which encodes an E3 ligase, and
in
DJ-1, which encodes a putative redox sensor that associates with
chaperones [
16] and translocates to mitochondria during conditions of oxidative stress [
17‐
22]. In addition to its proposed role as a redox sensor [
23], DJ-1 may also have important functions as an RNA binding protein [
24]. Although
DJ-1−/− mice have been reported to be
hypersensitive to the neurotoxin 1-methyl-4-phenyl-1,2,3,6-tetrahydropyrindine
(MPTP) [
25], and to display abnormalities in dopaminergic function when exposed to
the herbicide paraquat [
26], these mice do not develop nigrostriatal degeneration in the absence of
stresses [
17,
25,
27‐
29]. Moreover,
DJ-1−/− dopaminergic neurons or
siRNA-mediated knockdown of DJ-1 mRNA in primary embryonic midbrain DA neurons
resulted in increased sensitivity to toxins that induce oxidative stress [
30]. DJ-1 is also abundantly expressed in non-neuronal cells and the
bacterial endotoxin lipopolysaccharide (LPS) induces a robust increase in DJ-1
expression in inflammatory cells such as peritoneal macrophages [
31]. LPS exposure causes astrocytes derived from
DJ-1−/−
mice to generate ten times more nitric oxide than astrocytes derived from wild-type
mice [
32]. These studies suggest that DJ-1 loss-of-function mutations affect both
neuronal and non-neuronal cell types and could result in enhanced microglial
activation upon neuroinflammatory insults. Therefore, the purpose of our study was
to investigate whether loss of DJ-1 protein increases the vulnerability for
inflammation-induced nigrostriatal degeneration
in vivo. To this end, we
investigated the extent to which repeated intranasal administration of soluble tumor
necrosis factor (inTNF) or repeated intraperitoneal (i.p) injections of low-dose LPS
might induce neuroinflammation, locomotor deficits, enhance oxidative stress and/or
elicit nigral DA neuron loss in
DJ-1−/− mice compared to
wild-type mice.
Methods
Animals
DJ-1−/− mice were generated and characterized as described
previously [
17]. Prior to these studies,
DJ-1−/− mice were
back-crossed onto a C57BL/6 genetic background for over ten generations. Mice
were housed in a pathogen-free, climate controlled facility in the Animal
Resources Center at The University of Texas Southwestern Medical Center at
Dallas and given food and water
ad libitum. All animal studies were
reviewed and approved by the Institutional Animal Care and Use Committee at The
University of Texas Southwestern Medical Center in accordance with the National
Institutes of Health Guide for the Care and Use of Laboratory Animals.
Intranasal tumor necrosis factor administration
Murine soluble TNF (0.5 ng or 5 ng delivered as 5 μL of 0.1 ng/uL or 5
μL of 1 ng/uL) or an equivalent amount of saline vehicle was administered
intranasally via an L-10 Pipetman (Rainin Instrument, Oakland, CA, USA) twice
weekly for the time indicated in each set of experiments. Wild-type or
DJ-1−/− mice were 12 months old at the start of the
study (Additional file
1: Figure S1A) and 15 months
old at the time of sacrifice.
Systemic lipopolysaccharide administration
The regimen of LPS injections was chosen based on our previous work which
demonstrated that this dose and frequency of i.p. LPS triggers a
neuroinflammatory response in the midbrain and elicits nigral DA neuron loss in
parkin−/− mice [
33]. Young adult (6 to 13 week old)
DJ-1−/− mice on
a C57BL/6 background and age-matched wild-type mice were given either 7.5 ×
10
5 EU/kg LPS from
Escherichia coli O111:B4
(Sigma-Aldrich, Saint Louis, MO, USA) or sterile 0.9% sodium chloride vehicle
control (Braun Medical, Inc., Irvine, CA, USA) i.p. injections twice a week for
3 months. A second group of mice (designated as 3-month/3-month wait) was given
systemic LPS or vehicle i.p. injections for 3 months followed by a 3-month wait
period prior to tissue collection, during which no additional i.p. injections
were administered (n = 3 to 6 per group). A third group of mice was given twice
weekly systemic LPS or vehicle injections for 6 months with no wait period
before tissue collection (n = 3 to 7 per group). We would like to note that the
DJ-1−/− versus wild-type mouse studies reported in this
manuscript were performed alongside a cohort of
parkin−/−
mice, the results of which were reported previously [
33] in comparison to the same cohort of wild-type mice used here.
Behavior testing
For all behavioral tests, mice were evaluated at baseline (before i.p. injections
began) and again after 3 or 6 months of treatment.
Open-field
Open-field behavior in a glass container (diameter, 24.5 cm) was recorded for
5 minutes for evaluation of time spent moving and number of rearing events
by an investigator blinded to genotype and treatment history.
Narrow beam walk
A narrow beam (1.1 cm diameter, 80.6 cm testing length) with a home cage at
one end was used. Initial training prior to treatment consisted of three
sessions of three trials per session for 4 consecutive days. Mice received
additional training sessions at 3 months and 6 months after the start of the
treatment regimen consisting of three sessions of three trials per session
on 1 day. Testing was conducted the day after training and consisted of one
session of three trials. The average time to traverse the full length of the
beam was used for data analysis.
Accelerating rotarod
A base speed of 20 rpm with an acceleration of 0.2 rpm/second was used on the
rotarod (Economex 0207–005 M, Columbus Instruments, Columbus, OH,
USA). Mice were trained prior to treatment in three sessions of four trials
each for 4 consecutive days. Mice received additional training sessions at 3
months and 6 months after the start of treatment consisting of three
sessions of four trials per session on 1 day. Testing consisted of one
session of three trials the day after training was completed. Latency to
fall (seconds) was calculated and used for data analysis.
Tissue harvest
Following the last inTNF administration or i.p. injection and final behavioral
testing, mice in the 3-month, 3-month/3-month wait, and 6-month treatment
cohorts were deeply anesthetized with Euthasol (pentobarbital sodium and
phenytoin sodium) i.p. then intracardially perfused with warm 0.1 M PBS pH 7.4
supplemented with 0.1% glucose and 1 U/mL heparin prior to rapid whole brain
removal. For quantitative real-time PCR (QPCR), brain tissue was microdissected
into four regions on an ice-cold glass Petri dish - olfactory bulb, cerebellum,
ventral midbrain and cortex - then snap-frozen in cryovials in liquid nitrogen
and stored at −80°C until processed for RNA extraction. For
immunohistochemistry, mice in the 3-month, 3-month/3-month wait, and 6-month
treatment cohorts were perfused with warm 0.1M PBS (pH 7.4 supplemented with
0.1% glucose and 1 U/mL heparin) followed by ice cold 4% paraformaldehyde in PBS
(pH 7.4). Brains (in the skull) were post-fixed overnight in 4%
paraformaldehyde. Brains were removed from the skull and then were cryoprotected
for 24 hours in 20% sucrose in 0.1 M PBS pH 7.4, embedded in Neg 50 frozen
section medium (Richard Allen Scientific, Kalamazoo, MI, USA), and frozen in dry
ice-cooled isopentane.
Peritoneal macrophage harvest
Murine peritoneal macrophages were obtained by eliciting an acute peripheral
inflammatory reaction with an i.p. injection of thioglycolate [
34]. Briefly, adult mice were given an i.p. injection of 3%
Brewer’s yeast thioglycolate in normal saline. Three days later, animals
were euthanized and peritoneal exudates were recovered, pelleted and resuspended
in culture media (high-glucose DMEM supplemented with 10% heat-inactivated FBS
from Atlanta Biologicals (Norcross, GA, USA), 1% penicillin/streptomycin, and 1%
L-glutamine (Sigma-Aldrich)). Six hours after the initial plating, cells were
washed twice with PBS without Ca
2+ or Mg
2+ to remove
non-adherent cells and growth medium was replenished to the homogeneous
population of adherent macrophages.
Quantitative real-time polymerase chain reaction
Total RNA was isolated from tissue samples using Tri Reagent® (Molecular
Research Center, Cincinnati, OH, USA), treated with DNAse I (Invitrogen,
Carlsbad, CA, USA), and reverse transcribed to obtain cDNA. QPCR was performed
using SYBR Green Master Mix (ABI) on an Applied Biosystems Prism 7900HT sequence
detection system (Foster City, CA, USA) as described previously [
35]. Primers for each gene (available upon request) were designed using
Primer Express Software (PerkinElmer Life Sciences, Wellesley, MA, USA) and were
validated by analysis of template titration and dissociation curves. Results for
QPCR were normalized to the housekeeping gene cyclophilin B and evaluated by
comparative C
T method (user bulletin No. 2, PerkinElmer Life
Sciences). RNA levels are expressed relative to the wild-type saline-injected
(vehicle) mice.
Immunohistochemistry
Coronal serial sections (30 μm thickness) were cut on a Leica CM 1850
cryostat (Buffalo Grove, IL, USA) and placed on Superfrost/Plus microscope
slides (Fisher Scientific; Pittsburgh, PA, USA). Sections on slides were stored
at −80°C until processed for immunohistochemistry.
Brightfield immunohistochemistry. Sections were stained for tyrosine
hydroxylase (TH) using published protocols [
36,
37]. Sections were permeabilized in 0.3% Triton X-100 in PBS pH 7.4.
Endogenous peroxidases were quenched with 1% H
2O
2 and
non-specific binding was blocked with 5% normal serum (goat or horse,
Equitech-Bio, Inc., Kerrville, TX, USA). Sections were incubated with primary
antibodies against TH (rabbit polyclonal antibody AB152 diluted 1:2000, Chemicon
International, Temecula, CA, USA), or neuronal nuclear antigen (NeuN) (mouse
monoclonal antibody MAB377 diluted 1:1000, Chemicon) overnight at room
temperature followed by biotinylated secondary antibody (goat anti-rabbit or
horse anti-mouse rat absorbed, or goat anti-rat IgG diluted 1:400, Vector
Laboratories, Burlingame, CA, USA) and NeutrAvidin-HRP (diluted 1:5000, Pierce
Biotechnology, Inc., Rockford, IL, USA). The tissue bound peroxidase activity
was developed with 0.024% diaminobenzadine (DAB, Sigma), 0.006%
H
2O
2 in 0.05 M Tris–HCl buffer pH 7.6 for 20
minutes with or without nickel intensification. Tissue sections were dehydrated
in a graded series of ethanols, immersed in xylene, and coverslipped with
Permount (Fisher Scientific).
Fluorescence immunohistochemistry for Iba1-positive microglia. Brain
sections were stained for microglial markers using a standard immunofluorescence
protocol [
12]. Auto-fluorescence was quenched in 0.2 M glycine in PBS pH 7.4, for 1
hour at room temperature. Sections were then permeabilized in 0.3% Triton X-100
with 1% normal goat serum in 20 mM Tris-buffered saline (TBS) pH 7.4.
Non-specific binding was blocked with species-appropriate 1% normal serum in
TBS. Sections were incubated overnight at 4°C with an antibody specific for
Iba1 (Wako Chemicals, Richmond, VA, USA, 019–19741, diluted 1:10000) and
followed by Alexa-conjugated secondary antibody (Fab) (1:1000 dilution,
Invitrogen) for 4 hr at room temperature. Antibodies were diluted in blocking
buffer with 0.1% Triton X-100. Washes were done in TBS with 0.2% Triton X-100
(TBST). Following secondary antibody incubations, the slides were rinsed briefly
with dH
2O, then counterstained with Hoescht 33258 (diluted 1:20,000,
Invitrogen) for 15 minutes, and coverslipped with aqueous mounting media with
anti-fade (Biomeda Corp, Foster City, CA, USA). Quantification of Iba1-positive
cells was performed on images captured under a 20X objective lens (or 40X for
inset images) on a Nikon 90i fluorescence microscope using thresholding analysis
on Nikon Elements 5 software (Nikon Instruments, Melville, NY, USA). Values
represent the mean ± SEM of Iba1-positive microglia per field from four
fields selected randomly from entorhinal brain sections harvested from two mice
per treatment group.
Stereological analysis
The optical fractionator probe of Stereoinvestigator software (MicroBrightField,
Inc., Williston, VT, USA) was used to obtain an unbiased estimate of TH-positive
and NeuN-positive neurons in the SNpc and ventral tegmental area (VTA) as per
the atlas of the mouse brain by Paxinos and Franklin [
38]. Stereologic parameters were as follows: counting frame, 50 μm
× 50 μm; optical dissector: 20 μm; grid size, 120 μm ×
160 μm. For the population size estimate (number of sections per animal), a
target coefficient of error (Gundersen’s m = 1) of less than 0.10 was
considered acceptable. Neuron counting was performed by two different
investigators blinded to genotype and treatment history.
Striatal tyrosine hydroxylase fiber density and densitometry
Coronal serial sections (30 μm thickness) were cut between Bregma
−1.22 to 1.70 on a Leica CM 1850 cryostat and placed on Superfrost/Plus
microscope slides (Fisher Scientific). Tissue sections were immunostained for TH
and developed using DAB as described above. Images of striatum (caudate putamen)
from 12 tissue sections per animal were taken with a CoolSnap cf digital color
camera mounted on a BX61 microscope (Olympus, Center Valley, PA, USA). Exposure
times were kept constant for all images. TH-positive fiber density was
determined using background corrected integrated optical density measurements
for each section using an Alpha Innotech FluorChem FC2 imaging workstation and
software (Protein Simple, Santa Clara, CA, USA). All sections for each animal
were averaged and group means were used to compare treatment groups.
Levels of striatal DA and its metabolites (DOPAC, HVA and 3-MT) were quantified
by HPLC with electrochemical detection. Mice were euthanized by carbon dioxide
asphyxiation followed by immediate decapitation and dissection of the striatum
on an ice-cold glass Petri dish. The striatum was then weighed and stored at
−80°C. Frozen brain tissue was sonicated in a 49 volume/weight (mg of
tissue) solution of 0.1 M perchloric acid containing 0.2 mM sodium metabisulfite
and centrifuged at 20,000 rpm for 20 minutes at 4°C in a benchtop
centrifuge to clear debris. Cleared supernatant (20 μL) was then injected
onto a C18 HPLC column (ESA MD-150 3 × 150 mm) and separated by isocratic
elution at a flow rate of 0.6 ml/minute using MD-TM mobile phase (ESA Inc.,
Chelmsford, MA, USA). Neurotransmitter monoamines and metabolites were detected
using a BAS electrochemical cell set to a potential of +800 mV and compared to
external standards. Dopamine turnover was calculated as (DOPAC + HVA +
3-MT)/DA.
Statistics
Multiple-way analysis of variance (ANOVA) with significance level α = 0.05
were used as indicated for each set of experimental data. Significant
differences between groups were further evaluated using Tukey’s HSD or
Bonferroni’s post hoc test. Kruskal-Wallis analysis was the
non-parametric statistical test used for testing equality of population medians
of integrated optical density measurements of striatal TH fiber density. Graphs
were generated and statistical analyses performed with the use of GraphPad Prism
5.0 (GraphPad Software, La Jolla, CA, USA).
Discussion
DJ-1 function has been implicated in the regulation of inflammation-induced oxidative
stress
in vitro by reports that LPS can robustly increase DJ-1 expression
in peritoneal macrophages in response to NADPH oxidase-derived reactive oxygen
species [
31]. Moreover, DJ-1 has been shown to be important for mitochondrial function
in astrocytes [
40] and
DJ-1−/− astrocytes overproduce nitric oxide when
stimulated with LPS [
32,
41]. On the basis of these
in vitro studies we expected that the
brains of
DJ-1−/− mice would display exacerbated
neuroinflammatory responses to chronic inflammatory stress and would develop a
nigral degeneration phenotype, given that DA neurons are sensitive to
neuroinflammation. However, we found that the neuroinflammatory responses in
DJ-1−/− mice after repeated i.p. LPS injection were similar
to that of wild-type mice (Figure
6). Chronic i.p. LPS
exposure induced significant increases in midbrain TNF levels in both wild-type and
DJ-1−/− mice; however, the TNF levels were higher in
LPS-treated wild-type mice compared to LPS-treated
DJ-1−/−
mice. It remains possible that
DJ-1−/− mice may have shown
significant loss of nigral dopamine neurons if the LPS dose were increased to cause
midbrain TNF levels to match that of wild-type mice treated with this dose of LPS.
It is worth noting that the microglia burden measured by Iba-1+ cells in the brains
of
DJ-1−/− mice was not different from wild-type mice
(Figure
1), and is consistent with the observation
that the lineage-related myeloid-derived peripheral macrophage population also
displayed inflammatory responses
in vitro that were indistinguishable from
wild-type (Additional file
2: Figure S3). inTNF
administration caused no apparent neuroinflammatory responses in either wild-type or
DJ-1−/− mice. Therefore, the magnitude of the TNF
inflammatory insult may be too low to draw conclusions regarding the effect of DJ-1
mutations on susceptibility to TNF neuroinflammation-mediated neurodegeneration. It
remains possible that we would have observed different responses to TNF in wild-type
and
DJ-1−/− mice if we had used a stronger TNF administration
regimen.
Functionally, the repeated i.p. LPS injection regimen which successfully triggered
nigral degeneration in
parkin−/− mice [
33] did not induce nigral degeneration in
DJ-1−/− mice
(Figure
4), suggesting that loss of DJ-1 function
does not increase vulnerability to inflammation-induced nigral degeneration.
Nevertheless, it remains possible that a higher concentration of inTNF or LPS and/or
a longer regimen (greater than 6 months) and/or longer wait-time (>3 months)
between the delivery of the last set of repeated i.p. LPS injections and the
endpoint of the study may be required to elicit a more robust neuroinflammatory
response and uncover increased vulnerability to inflammation-induced degeneration in
DJ-1−/− mice. Although we observed an intriguing increase
in striatal TH fiber density and striatal DA in
DJ-1−/− mice
treated with LPS for 3 months, we did not observe significant changes in the cohorts
receiving LPS for 6 months or for 3 months with a 3-month wait period. Some caution
should therefore be exercised in interpreting the unexpected increases in striatal
TH fiber density and striatal DA in the 3-month treatment group.
Although
DJ-1−/− mice were reported to display normal numbers of
nigral DA neurons in the SNpc, they were shown to have deficits in dopaminergic
function [
17] that are further accentuated when exposed to paraquat [
26] and MPTP [
25], suggesting a protective role for DJ-1 in mitochondrial function and/or
against oxidative stress. In support of this molecular model, DJ-1 has been shown to
translocate to mitochondria [
20] in response to oxidative stress when key cysteine residues become
oxidized [
18], and to also interact with multifunctional regulators of transcription
and RNA metabolism in the nucleus [
42]. In contrast, we found that mice deficient in
DJ-1 displayed
blunted oxidative stress levels in response to LPS treatment relative to wild-type
mice (Figure
7), as measured by expression of Nrf2, HO-1
and iNOS. However, we found that expression of SOD-1 and SOD-2 was increased in
saline-treated
DJ-1−/− mice compared to saline-treated
wild-type mice (Figure
7), which may reflect compensatory
upregulation of certain anti-oxidants in the absence of DJ-1 and may have
contributed to the attenuated expression of oxidative stress genes in response to
LPS. Inflammatory stimuli have been reported to exacerbate the oxidative response of
DJ-1−/− cells, and although we did not observe this in our
study, we only reported responses after chronic LPS treatments (6 months) and did
not have the acute response gene expression after 3 months of LPS injections to draw
conclusions about the kinetics of the oxidative stress response. Additionally, gene
expression was measured in lysates of whole brain sections and it is therefore
likely that the differences we observed in oxidative gene expression in the
DJ-1−/− mouse midbrain is attributed to other cell types in
the brain, such as astrocytes or neurons which also express SOD-1 and SOD-2, for
example. Interestingly, we also observed that saline-treated
DJ-1−/− mice compared to saline-treated wild-type mice have
increased CD45 in the midbrain. Although Iba-1 immunostaining indicated no robust
changes in microglia number or activation status, this increased expression could be
due to increased levels of other lymphocytes or neutrophils which also express CD45 [
43]. Lastly, given that
DJ-1−/− mice have been reported
to have compensatory gene expression in mitochondrial enzymes [
44], we considered and ruled out the possibility that compensatory increases
in Parkin mRNA and protein expression accounted for the lack of vulnerability
against inflammation-induced nigral degeneration.
In summary, we conclude that loss of DJ-1 function, unlike loss of Parkin function,
does not confer increased vulnerability to the neurodegenerating effects of chronic
inflammatory insults induced by LPS. Given the contrasting responses of
parkin−/− and DJ-1−/− mice to repeated
i.p. LPS injections, we conclude that Parkin and DJ-1 are likely to have distinct
and non-overlapping roles in protecting the nigrostriatal pathway against
inflammatory and neurotoxic insults that lead to degeneration.
Competing interests
The authors declare that they have no competing interests.
Authors’ contributions
TAN: experimental design, food intake, dosing, behavior, tissue harvest, tissue
processing, QPCR, HPLC, IHC, stereology, statistical analysis, paper editing. TF-C:
experimental design, dosing, behavior, tissue harvest, tissue processing, IHC,
stereology and striatal density, statistical analysis, paper editing. TNM:
experimental design, dosing, behavior, tissue harvest, tissue processing, IHC,
stereology, fluorescent microscopy, paper editing. KAR: animal colony, dosing,
behavior, tissue harvest, perfusions, cryosectioning, stereology. MM: dosing, tissue
processing for RNA, behavior, HPLC. BC: dosing, tissue harvest. JJH: IHC. IT: IHC,
HPLC. MGT: experimental design, dosing, food monitoring, behavior, stereology,
fluorescence microscopy, paper editing. MSG: experimental design, behavior, food
monitoring, tissue harvests, stereology, striatal dissections, paper editing. All
authors read and approved the final manuscript.