Glutamate Receptor-Mediated Toxicity in Optic Nerve Oligodendrocytes
Astroglial plasticity and glutamate function in a chronic mouse model of Parkinson's disease
Transcript of Astroglial plasticity and glutamate function in a chronic mouse model of Parkinson's disease
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Experimental Neurology
Astroglial plasticity and glutamate function in a chronic mouse model of
Parkinson’s disease
Adrian G. Dervana, Charles K. Meshulb,c, Mitchell Bealesa, Gethin J. McBeand, Cindy Mooreb,c,
Susan Totterdelle, Ann K. Snydera, Gloria E. Mereditha,*
aDepartment of Cellular and Molecular Pharmacology, Rosalind Franklin University of Medicine and Science, The Chicago Medical School, North Chicago,
IL 60064, USAbResearch Services, V.A. Medical Center, Portland, OR 97239, USA
cDepartment of Behavioral Neuroscience, Oregon Health and Science University, Portland, OR 97239, USAdDepartment of Biochemisty, Conway Institute Biomolecular & Biomedical Research, University College Dublin, Dublin 4, Ireland
eDepartment of Pharmacology, University of Oxford, Oxford OX1 3QT, UK
Received 20 May 2004; revised 25 June 2004; accepted 8 July 2004
Abstract
Astrocytes play a major role in maintaining low levels of synaptically released glutamate, and in many neurodegenerative diseases,
astrocytes become reactive and lose their ability to regulate glutamate levels, through a malfunction of the glial glutamate transporter-1.
However, in Parkinson’s disease, there are few data on these glial cells or their regulation of glutamate transport although glutamate
cytotoxicity has been blamed for the morphological and functional decline of striatal neurons. In the present study, we use a chronic mouse
model of Parkinson’s disease to investigate astrocytes and their relationship to glutamate, its extracellular level, synaptic localization, and
transport. C57/bl mice were treated chronically with 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine and probenecid (MPTP/p). From 4 to 8
weeks after treatment, these mice show a significant loss of dopaminergic terminals in the striatum and a significant increase in the size and
number of GFAP-immunopositive astrocytes. However, no change in extracellular glutamate, its synaptic localization, or transport kinetics
was detected. Nevertheless, the density of transporters per astrocyte is significantly reduced in the MPTP/p-treated mice when compared to
controls. These results support reactive gliosis as a means of striatal compensation for dopamine loss. The reduction in transporter
complement on individual cells, however, suggests that astrocytic function may be compromised. Although reactive astrocytes are important
for maintaining homeostasis, changes in their ability to regulate glutamate and its associated synaptic functions could be important for the
progressive nature of the pathophysiology associated with Parkinson’s disease.
D 2004 Elsevier Inc. All rights reserved.
Keywords: Glial fibrillary acidic protein; MPTP; Astrocytosis; GLT-1; Stereology
Introduction
Astrocytes, which can be detected by glial fibrillary
acidic protein (GFAP) immunoreactivity (Eng and Lee,
1995), are critical for maintaining the homeostatic extrac-
0014-4886/$ - see front matter D 2004 Elsevier Inc. All rights reserved.
doi:10.1016/j.expneurol.2004.07.004
* Corresponding author. Department of Cellular and Molecular
Pharmacology, The Chicago Medical School, Rosalind Franklin University
of Medicine and Science, 3333 Green Bay Road, North Chicago, IL 60064.
Fax: +1 847 578 3268.
E-mail address: [email protected] (G.E. Meredith).
ellular environment. They regulate glutamate uptake and
play an important role in synaptogenesis and synaptic
plasticity (Araque et al., 1998; Bowers and Kalivas, 2003;
Slezak and Pfrieger, 2003; Ullian et al., 2001). Pathological
processes in the brain, such as those seen in Alzheimer’s and
Huntington’s disease and in amyotrophic lateral sclerosis,
are associated with a significant astroglial reaction (Eddles-
ton and Mucke, 1993), altered glutamate uptake, and
changes in the expression of glutamate transporters (Beh-
rens et al., 2002; Lin et al., 1998; Rothstein et al., 1992,
1995; Scott et al., 2002). In Parkinson’s disease (PD),
190 (2004) 145–156
A.G. Dervan et al. / Experimental Neurology 190 (2004) 145–156146
however, astrocytosis has not been reported for postmortem
striatum (Forno et al., 1992), and nothing is known of
glutamate transport kinetics.
Under normal conditions, striatal neurons are protected
by an efficient glutamate transport system in neighboring
astrocytes, primarily mediated by the glial glutamate trans-
porter (GLT-1). However, under conditions of oxidative
stress such as occurs in PD and experimental animals treated
with 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP),
the functionality of this system might be changed (cf.
Greene and Greenamyre, 1995). When astrocytic cultures
are treated with MPP+, glutamate transporter function is
significantly impaired (Di Monte et al., 1999) and exper-
imental studies in vivo have proposed that striatal neuron
damage arising after dopamine (DA) depleting lesions is
due to increased excitatory activity (Dervan et al., 2003a;
Ingham et al., 1989; McNeill et al., 1998; Meredith et al.,
1995). Certainly, extracellular glutamate is elevated follow-
ing striatal DA loss with 6-hydroxydopamine (6-OHDA) or
acutely administered MPTP (Araki et al., 2001; Meshul et
al., 1999, 2002; Robinson et al., 2003; Sheng et al., 1993).
MPP+, when administered directly into the striatum,
increases the release of glutamate (Carboni et al., 1990),
and glutamatergic receptor blockade seemingly ameliorates
the neurotoxic effects of MPTP (Brouillet and Beal, 1993;
Turski et al., 1991). Nevertheless, less severe treatments
with MPTP, i.e., subacute administration, does not increase
levels of striatal glutamate (Robinson et al., 2003). More-
over, the striatal astroglial reaction to 6-OHDA is unstable
over time, rising, then falling, and rising yet again several
weeks post-lesion (Tripanichkul, 2000). When considered
together, these data indicate that astrocytes and their
regulation of glutamate after DA loss are imperfectly
understood.
Treatments with MPTP administered acutely or 6-OHDA,
either in vivo or in vitro, bear little resemblance to the
progressive nature of PD. Therefore, we questioned whether
a chronic model that is produced by combining MPTP with
probenecid (Lau et al., 1990) to enhance and prolong DA
loss (Petroske et al., 2001) would be more appropriate to
study astrocyte function, glutamate and its uptake. Some of
these results were presented previously (Dervan et al.,
2003b).
Materials and methods
Animals and treatment
One hundred fifty, adult, C57BL/6 male mice (Charles
River Laboratories, Wilmington, MA) at 8 weeks of age and
weighing 20–30 g were housed 2 to the cage. All protocols
were conducted in accordance with the National Institutes of
Health Publications No. 80–23 and were approved by the
Institutional Animal Care and Use Committees at the
Chicago Medical School and Portland VA Medical Center.
Two groups of mice (n = 50 per group) were treated
with 10 doses (2� week; 3.5 days apart) either of MPTP
hydrochloride (25 mg/kg in saline, s.c., Sigma-Aldrich,
St. Louis, MO) and probenecid (250 mg/kg in dimethyl
sulfoxide, i.p., Sigma-Aldrich) (MPTP/p), or probenecid
alone (controls). Previous studies have confirmed that
probenecid administration does not differ from saline
treatment in relation to DA cell number and DA function
(Lau et al., 1990; Petroske et al., 2001). Nevertheless,
results from probenecid-treated mice were compared to
saline-treated controls (n = 24) in some experiments.
Finally, two groups of mice (n = 25/group) were treated
acutely with MPTP (4 � 20 mg/kg in saline i.p. every 2
h). Chronically treated groups were sacrificed 4–8 weeks
after treatment and acutely treated groups, 1 week post-
treatment.
Mice used in transporter assays were killed by cervical
stunning and decapitation. All others were deeply
anesthetized with sodium pentobarbital (130 mg/kg) and
treated as follows: (1) After microdialysis studies and for
glutamate immuno-gold investigations, perfusions con-
sisted of 1000 units/ml of heparin in 0.1 M HEPES
buffer (pH 7.3) followed by 2.5% glutaraldehyde/0.5%
paraformaldehyde in 0.1 M HEPES containing 0.1%
picric acid. For immuno-gold studies, brains were post-
fixed overnight. (2) For other light or electron (EM)
microscopy studies, perfusions consisted of 2.5% sucrose
in 0.1 M phosphate-buffered saline (PBS), followed by
3% paraformaldehyde in 0.1 M PB; brains were post-
fixed for 2 h. For light microscopic investigations, brains
were sunk in 20% sucrose overnight. All brains were
blocked and sectioned at 50-Am thickness in the coronal
plane using a vibrating or freezing microtome.
Immunohistochemistry and electron microscopy
Sections through the striatum were collected in series
(1 in 5) in a uniformly random manner. Adjacent series of
sections were immunoreacted with monoclonal mouse
tyrosine hydroxylase antibodies (TH; Immunochemicals,
Stillwater, MN), rabbit anti-GFAP sera (DAKO, Carpen-
teria, CA), or dually immunolabeled for GLT-1 (rabbit
anti-GLT-1A sera were the kind gift of Dr. David Pow,
University of Queensland, Brisbane, Australia) and GFAP.
These GLT-1 antibodies were previously characterized and
their specificity established (Reye et al., 2002). All treated
and control tissues were immunoreacted at the same time
with the same solutions. Sections reserved for TH and
GFAP immunohistochemistry were incubated for 24 h at
48C in anti-TH sera (1:2000) or rabbit anti-GFAP
(1:2500), then in appropriate biotinylated IgG (1:300),
followed by incubation in avidin–biotin–peroxidase
(ABC) complex. Sections were then reacted in 0.05% 3-
3V diaminobenzidine hydrochloride (DAB) with added
0.01% H2O2, mounted onto slides, dehydrated, and
topped with coverslips.
A.G. Dervan et al. / Experimental Neurology 190 (2004) 145–156 147
The third series of sections were incubated in rabbit anti-
GFAP, as above, followed by donkey anti-rabbit IgG
conjugated to CY2 (Jackson ImmunoResearch, West Grove,
PA), 1:500. After extensive rinsing, sections were incubated
in rabbit anti-GLT-1, 1:20,000, overnight at 48C, followedby 2 h in donkey anti-rabbit IgG conjugated to CY3, 1:500.
Sections were mounted onto subbed slides and topped with
coverslips using a premixed mounting medium compatible
with fluorescent labels.
For EM studies of astrocytes (n = 3/group), sections from
the anterior striatum (+1.0 mm anterior; Franklin and
Paxinos, 1997) were treated with 1% osmium tetroxide
(0.1 M PB) for 30 min, dehydrated, and mounted onto slides
in Durcupan resin (ACM, Fluka, UK). Uranyl acetate (1%)
was added to the 70% ethanol step to increase contrast.
Regions of interest, selected with the light microscope, were
glued onto preformed resin blocks. Cut serial ultrathin
sections were mounted on Pioloform-coated, single slot
grids, contrasted with lead citrate and photographed in a
Philips 400 EM. Approximately 30 astrocytes were exam-
ined from each group.
Sections prepared for glutamate immuno-gold labeling
came from mice used for microdialysis. However, these
mice were not challenged with l-DOPA before perfusion,
so a comparison could be made between glutamate
immunolabeling density and basal glutamate. Sections
were incubated at room temperature in 1% osmium
tetroxide/1.5% potassium ferricyanide, rinsed and stained
en block with 0.5% uranyl acetate. Tissue was embedded
in Epon/Spurr resin and ultrathin sections cut at 60–90 nm.
Post-embedding immuno-gold EM was performed accord-
ing to Phend et al. (1992), as modified by Meshul et al.
(1994). The glutamate antibody (non-affinity purified,
rabbit polyclonal; Biogenesis, Brentwood, NH), as pre-
viously characterized by Hepler et al. (1988), was diluted
1:400,000 in 0.05 M Tris-buffered saline, pH 7.6.
Aspartate (1 mM) was added to the glutamate antibody
mixture 24 h before incubation to prevent any cross-
reactivity with aspartate within the tissue. The specificity
of glutamate immunolabeling was previously established
(Meshul et al., 1994). Photographs (10 per animal) were
taken randomly through the neuropil at a final magnifica-
tion of �40,000 using a digital camera (AMT, Boston,
MA). The number of gold particles per nerve terminal
associated with an asymmetrical synaptic contact was
counted and the area of the nerve terminal determined
using Image Pro Plus software (Media Cybernetics, Silver
Springs, MD, Version 3.01). The gold particles contacting
the synaptic vesicles within the nerve terminal were
counted and were considered part of the vesicular or
neurotransmitter pool according to previously established
methods (Meshul et al., 1999; Robinson et al., 2001). The
density of gold particles/Am2 of nerve terminal area was
determined for each animal and the mean density for each
treatment group calculated (mean density + SEM). The
differences between treatment groups were analyzed using
the Student’s t test. The total number of synapses for each
of the treatment groups making an asymmetrical synaptic
contact was as follows: MPTP/p group: 234 synapses,
probenecid group: 228 synapses.
Stereological estimates of astrocyte number and size
Striatal borders were demarcated for the stereological
analysis (see Figs. 1A–C; Franklin and Paxinos, 1997) with
the most posterior border set at the level of the anteriormost
amygdala. After randomly selecting a starting point, nine
sections at equally spaced intervals along the rostrocaudal
extent of the striatum were selected for analysis. The
reference volume for the striatum was estimated according
to Cavalieri principles (Coggeshall, 1992) and the total
number (N) and density (Nv) of astrocytes were estimated
with the optical dissector following fractionator rules (West
and Gundersen, 1990) and a semiautomated system (Stereo-
Investigator, version 4.565, Microbrightfield, Williston,
VT).
Video images of GFAP-immunoreactive astrocytes were
acquired with a 100� oil objective (1.35 NA) on a Nikon
E600 microscope equipped with a CCD camera output to a
high-resolution, computer monitor and a Ludl X–Y–Z
motorized stage (Ludl Electronics Products, Hawthorn,
NY). A counting frame of 75 � 75 Am (Fig. 2) and a
disector height of 8 Am with guard zones of 2 Am were
employed. An astrocyte was counted only if it had a clearly
defined nucleus within the disector area, did not intersect
forbidden lines (Fig. 2) and came into focus as the optical
plane moved through the height of the disector. The
precision of each estimate was expressed as the coefficient
of error (CE; Table 1). The somal volume of each astrocyte
cell body was calculated using the nucleator probe (Moller
et al., 1990).
Surgical procedures and in vivo microdialysis
Eight weeks after the last MPTP/p (n = 7) or probenecid
(n = 5) injection, animals were anesthetized (0.1 ml/kg of
2.5% ketamine, 1% xylazine, and 0.5% acepromazine in
normal saline) and placed in a stereotaxic apparatus
(Cartesian Research, Inc., Oregon). A small hole was drilled
in the skull, and the dura was punctured at the following
coordinates from Bregma (Franklin and Paxinos, 1997):
anterior: +1.2 mm; lateral: +1.8 mm. A stainless-steel guide
cannula (8-mm long, 21-gauge; Small Parts, Miami Lakes,
FL) was lowered 0.8 mm from the surface of the skull to the
level of the corpus callosum. The guide cannula was held in
a fixed position by one stainless steel screw attached to the
skull and encompassed by cranioplastic (Plastics One, Inc,
Roanoke, VA).
The dialysis probes, prepared as described by Robinson
and Wishaw (1988) with modifications (Meshul et al.,
1999), were 210 Am in diameter and 2 mm in length. One
day before use, the efficiency of transmitter recovery by the
Fig. 1. (A–C) Light photomicrographs of sections from rostral (A), middle (B), and caudal (C) levels of the striatum in probenecid-treated animals. The lines
drawn in A, B, and C show the boundaries used to delineate the striatum for stereological investigation. (D) In probenecid-treated controls, GFAP-positive
astrocytes are sparsely distributed (arrows) at all levels. (E) In MPTP/p-treated animals, intense GFAP immunostaining is visible (arrows) and corresponds to an
increase in the number of astrocytes and their processes (see Table 1). cc, corpus callosum; Nacc, nucleus accumbens; ac, anterior commissure; gp, globus
pallidus. Scale bars A–C = 500 Am; D and E = 100 Am.
A.G. Dervan et al. / Experimental Neurology 190 (2004) 145–156148
probe was determined by collecting three 10-min samples
(perfusion flow rate of 2 Al/min) after placing the probe in a
solution of glutamate (200 pg/Al) in artificial cerebrospinal
fluid (aCSF, pH 7.4).
Following the collection of recovery samples and the
day before the actual dialysis procedure, the probe was
lowered into the guide cannula with the entire length of
the dialysis probe in the striatum. The probe was secured
to the guide cannula with epoxy. The artificial aCSF
flowed through the probe overnight at a rate of 0.5 Al/min. The following morning, the pump speed was
increased to 2 Al/min for 1 h and then four samples
were collected every 15 min to determine the basal level
of extracellular glutamate. We have previously verified
that changes in extracellular striatal glutamate are depend-
ent on the presence of calcium within the aCSF (Meshul
et al., 2002). Both the probenecid and MPTP/p-treated
groups were then challenged with 15 mg/kg l-DOPA/12.5
mg/kg benserazide (Sigma) and an additional six 15-min
samples were collected. After the experiment, Vibratome
sections (100 Am) were stained with thionin and the site
of the probe placement within the striatum verified
histologically. Probe placement extended 2 mm along
the lateral quadrant of the striatum. If the placement was
not correct (i.e., outside the striatum), the data from that
animal were discarded. The four baseline data points and
the six l-DOPA-challenged data points were separately
averaged at each time point and a grand mean determined
for either the baseline or l-DOPA-challenged samples.
Values of extracellular striatal glutamate are expressed as
the mean F SEM in pmol/Al. The mean probe recovery
was 10.4 F 1.2%. Data were analyzed using a one-way
ANOVA and significant main effects were further
characterized using Peritz’ f test for comparison of
multiple means.
HPLC detection of dialysate glutamate levels
Glutamate concentration in each dialysate sample was
determined using a Hewlett Packard HPLC 1090 interfaced
with a Hewlett Packard 1046A Programmable Fluorescence
Detector. Dialysates were derivatized with o-phthalaldehyde
and chromatographed according to a modification of the
method of Schuster (1988), as previously reported (Meshul
et al., 1999, 2002; Robinson et al., 2003). Assay sensitivity
was in the subpicomole range.
Glutamate transport assays
To establish whether glutamate transporter function was
impaired, we decapitated mice either 3 days after acute
MPTP or saline treatment or 5 weeks after MPTP/p,
Fig. 2. Light micrograph illustrating the stereological counting frame.
Pictured are two astrocytes from a probenecid-control striatum. Only cells
that lay within the volume (75 � 75 � 8 Am) of the frame and/or touched
the green lines were counted and those that crossed the red lines were
excluded from analysis. Estimates of cell size were generated using the
nucleator probe (white lines). For each cell, eight isotropic lines converged
on the nucleus and intersected the somal boundary. Scale bar = 10 Am.
A.G. Dervan et al. / Experimental Neurology 190 (2004) 145–156 149
probenecid, or saline treatment. Pooled striata (n = 5 per
group) were homogenized in ice-cold medium of 0.32 M
sucrose in 50 mM Tris buffered to pH 7.5. The homogenate
was centrifuged 5 min at 1200 � g at 48C (Sorvall
centrifuge, SS-34 rotor). The supernatant was removed
and centrifuged for 12 min at 17,000 � g at 48C. The pelletwas washed in ice-cold gradient medium, collected by
Table 1
Detailed results from the striatum showing total volume, astrocyte number, densi
Probenecid
controls
Chronic
MPTP/P
V (mm3) CE Astrocytes
counted Q�Disec
P0381 5.681 0.07 258 502
P0382 5.636 0.06 275 500
P0384 5.024 0.05 172 450
P0385 5.829 0.08 214 517
Mean F SE 5.543 F 0.17 0.06 230 492
MP0354 5.103 0.09 572 523
MP0356 5.832 0.07 489 453
MP03266 5.926 0.07 598 527
MP03258 5.430 0.07 547 469
MP03267 6.009 0.06 705 514
Mean F SE 5.660 F 0.18a 0.07 582 497
a P = 0.651 (unpaired Student’s t-test); no significant difference compared to cob P = b0.001 (unpaired Student’s t-test); significantly greater than controls.c P = b0.001 (unpaired Student’s t-test); significantly greater than controls.d P = b0.05 (unpaired Student’s t-test); significantly greater than controls.
centrifugation, and resuspended in a final volume of 750 Alof homogenization medium.
High-affinity Na+-dependent transport was assayed at
substrate concentrations between 4 � 10�6 and 40 �10�6 M essentially as described previously (De Souza et
al., 1999). Briefly, synaptosomes were added to duplicate
microcentrifuge tubes containing Krebs solution, pH 7.4.
Following protein determination, the synaptosomes were
incubated at 378C with 15 nCi of d-[2,3-3H]-aspartic acid
(New England Nuclear, Downers Grove, IL). The
reaction was stopped with 200 Al of ice cold 1 mM d-
aspartate, followed by microcentrifugation at 15,700 � g
at 48C. The pellets were washed twice without resus-
pending using sodium-free Krebs medium, microcentri-
fuged as described and dissolved by overnight incubation
in 200 Al of 2% sodium dodecyl sulfate and the next day
suspended in 5 ml of Ecoscint A (National Diagnostics,
Atlanta, GA). The quantity of radioactivity was deter-
mined by liquid scintillation spectroscopy and the rate of
transport was plotted as a function of substrate concen-
tration, using nonlinear regression analysis.
Quantitative analysis of GLT and GFAP immunoreactive
profiles
Although high-affinity, Na+-dependent transport assays
give an indication of the number of functional trans-
porters present in synaptosomes prepared from the
striatum, they provide no information about the location
or density of transporters on individual astrocytes.
Therefore, we quantified the density of GLT-1 on
GFAP-immunolabeled cells. To minimize the variability
of immunohistochemical processing, all sections were
processed simultaneously using buffers, antibodies, and
solutions made at the same time. Single images (1024 �1024 pixels) were acquired digitally from an Olympus
Fluoview Series 43 confocal laser-scanning microscope
(CLSM; oil objective 60� and �2 magnification) without
ty, and somal volume
tors, P CE Total number
N (�105)
Astrocyte density
Nv (�103/mm3)
Somal
volume (Am3)
0.06 2.38 4.18 289.0
0.06 2.51 4.45 326.7
0.08 1.54 3.04 301.5
0.06 1.75 2.99 259.2
0.06 2.045 F 0.24 3.67F 0.38 294.1 F 14.0
0.04 4.89 9.59 330.4
0.04 4.24 7.27 319.6
0.04 4.52 7.62 342.6
0.04 4.10 7.55 320.9
0.03 5.21 8.67 343.7
0.04 4.592 F 0.20b 8.14 F 0.43c 331.6 F 5.1d
ntrols.
A.G. Dervan et al. / Experimental Neurology 190 (2004) 145–156150
saturation of pixel intensities and under constant con-
ditions by adjusting the CLSM parameters: pinhole size,
laser power, and detector voltage. Four mice from each
group and a single section through the striatum (mid-
rostral level from Bregma: anterior +1.2 mm; same as in
Fig. 3) for each mouse were selected for study.
Essentially all cells, i.e., approximately 20 per section
(each a merged image of a GFAP-immunoreactive
astrocyte and accompanying GLT-1 immunolabeling)
from the dorsomedial quadrant of the striatum from each
coded section, were captured by an investigator blind to
the animal treatment. The images were imported into the
Image J 1.29 program (NIMH), where the gray level of
each was inverted (no signal = 0 and maximum level =
255, corresponding to eight bits). The gray levels of the
merged images were measured by placing four open
rectangles, each with a diameter of 4 pixels, across the
cell body and proximal astrocytic processes of each cell
(see Results for further detail). Optical density within each
rectangle was recorded and measurements were pooled. The
mean F standard error of the mean (SEM) was established
for each group; group means were compared with a
Student’s t test.
Results
Effect of MPTP/p treatment on striatum
At 3 weeks post-MPTP/p treatment, mice show a
significant behavioral impairment and reduction in DA
neurons and striatal DA function (Dervan et al., 2003a;
Fig. 3. Representative sections of the rostral striatum from probenecid control (A) a
striatum contains numerous TH-immunoreactive fibers. (B) The extent of loss of T
Note the relative sparing in the ventral striatum and along a thin band dorsolater
Meredith et al., 2002; Petroske et al., 2001). In the
present study, mouse striata from MPTP/p-treated mice
that were immunoreacted for TH show a significant
reduction in TH-positive fibers, except for an outer rim
of spared fibers in the dorsolateral striatum and a
relatively spared ventral striatum (compare Fig. 3B with
A). This pattern of TH immunoreactivity is consistent
with that described in our earlier work (Meredith et al.,
2002; Petroske et al., 2001).
Astrocyte distribution, number, and size
In control animals, GFAP immunoreactivity is moderate to
light in the striatum and cortex (Figs. 1A–C); GFAP-
immunopositive cells are typically grouped into small
dislandsT that follow the course of blood vessels. At the
cellular level, each astrocyte in control mice (probenecid- or
saline-treated) has a tightly configured set of fibrous
processes that are initially thick as they emerge from the cell
body but rapidly become fine and wispy distally (Fig. 4A). In
contrast, MPTP/p treatment is associated with an increase in
GFAP-immunoreactive profiles in the striatum (compare Fig.
1E with D), and immunopositive astrocytes display a reactive
morphology with thickened processes (Fig. 4B). The reactive
cells are often located close to the ventricle and along the
corpus callosum. Ultrastructurally, most astrocytes from
MPTP/p-treated, but not control mice, have nuclear inden-
tations and clumped chromatin (Figs. 4C–D).
Stereological estimates showed that the striatum of
MPTP/p-treated animals had significantly more GFAP-
immunoreactive cells than does that of controls. However,
the reference volume of the striatum is not different between
nd MPTP/p (B) treated animals immunoreacted for TH. In controls (A), the
H-immunoreactivity can be seen in the striatum of MPTP/p-treated animals.
ally. Scale bars = 500 Am.
Fig. 5. In vivo microdialysis of the lateral striatum from MPTP/p (n = 7) or
probenecid (n = 5) treated animals, 8 weeks after final administration.
Baseline samples were collected from each group and then both groups
were injected with a single dose of l-DOPA (15 mg/kg, l-DOPA + 12.5
mg/kg, benserazide) and additional samples collected. There was no
difference in the basal extracellular levels of striatal glutamate between the
two groups [Control (baseline) vs. MPTP/p (baseline)]. Following the
injection of l-DOPA, there was a significant increase in the extracellular
levels of glutamate in both groups compared to their respective baseline
levels. Values are means F SEM (pmol/Al). *P b 0.05 and **P b 0.05
compared to control (baseline) or MPTP/p (baseline) groups using Peritz’ f
test for comparison of multiple means.
Fig. 4. Light photomicrographs of GFAP-immunopositive astrocytes of probenecid-treated (A) and MPTP/p-treated (B) animals. Note the increased thickness
of processes on cells after MPTP/p treatment (B). (C) Representative electron micrograph of a striatal astrocyte of a probenecid-treated animal. Note the
electron-lucent cytoplasm and nucleus. In astrocytes from MPTP/p-treated animals (D), the cells are electron-dense and contain an indented nucleus with
clumped chromatin (arrows). Scale bars in A and B = 10 Am and in C and D = 2 Am.
A.G. Dervan et al. / Experimental Neurology 190 (2004) 145–156 151
groups (Table 1). In addition, cell bodies immunopositive
for GFAP are significantly increased in size following
MPTP/p treatment when compared to controls (Table 1).
Extracellular glutamate levels
Eight weeks following administration of MPTP/p or
probenecid (i.e., a total of 13 weeks after the first dose of
MPTP/P), in vivo microdialysis of the lateral striatum
revealed no difference in basal extracellular glutamate
between the two groups (Fig. 5). To determine if the
MPTP/p group is differentially sensitive to the effects of l-
DOPA, used to restore brain levels of DA, both groups
received a single dose of the drug following the collection of
baseline samples. This resulted in a significant increase in
the extracellular levels of striatal glutamate in both groups
compared to baseline (Fig. 5). Although the increase in the
control (probenecid) group (21.4%) is greater than that of
the MPTP/p group (12.3%), this difference is not statisti-
cally significant.
Nerve terminal glutamate immuno-gold labeling
In a separate group of MPTP/p- or probenecid-treated
animals, quantitative immuno-gold EM was carried out to
determine the immunolabeling associated with the synaptic
vesicle (i.e., neurotransmitter) pool in endings making an
asymmetrical synaptic contact (Figs. 6A–B). There is no
difference in the density of nerve terminal glutamate
labeling between MPTP/p-treated and control groups (Fig.
Fig. 7. The effect of chronic MPTP/p (A) and acute MPTP (B) treatment on
the kinetics of d-[3H]aspartate transport. Synaptosomes were incubated at
358C for 4 min in Krebs’ medium and the uptake of d-[3H]aspartate at the
specified concentrations was measured. The results from each treatment are
plotted as a mean of five independent experiments, performed in duplicate
and expressed as the mean F SEM.
Fig. 6. Nerve terminal glutamate immuno-gold labeling in the dorsolateral
striatum, 8 weeks after MPTP/p or probenecid (n = 8/group). (A)
Representative electron micrograph from the probenecid (control) group.
Note the number of gold particles (arrowhead) associated with synaptic
vesicles and within three nerve terminals making an asymmetrical synaptic
contact (arrows) with an underlying dendritic spine. (B) Glutamate nerve
terminal immunolabeling from the MPTP/p group: There are four terminals
making an asymmetrical synaptic contact (arrows) with an underlying
dendritic spine. (C) The density of nerve terminal glutamate (gold particles/
Am2) in asymmetrical (excitatory) synapses was determined. There was no
change in the density of gold particles, the number of gold particles, or in
the area of the terminals between the two groups. Values are meansF SEM.
Scale bar = 0.5 Am.
A.G. Dervan et al. / Experimental Neurology 190 (2004) 145–156152
6C). Previous work has shown that 97% of the immuno-
gold labeling is associated with the synaptic vesicle pool
and only 3% with the cytoplasmic pool (Meshul et al.,
1999). This cytoplasmic pool does not change following a
6-OHDA lesion (Meshul et al., 1999). In previous studies,
the mitochondrial (metabolic) pool was analyzed separately
as compared to the vesicular pool. There appears to be no
change in the mitochondrial pool of glutamate after a 6-
OHDA lesion (Meshul and Allen, 2000; Robinson et al.,
2001). In the present study, the gold particles associated
with the mitochondrial pool were not included in the density
measurements of nerve terminal glutamate.
Effect of MPTP treatment on high-affinity, Na+-dependent
transport
We studied the rate of transport of d-[3H] aspartate
into crude synaptosomes from mouse striatum over a
concentration range of 0.4–40 AM and compared high-
affinity transport in chronically (MPTP/p) and acutely
(MPTP) treated mice and their respective probenecid or
saline controls. Fig. 7A shows that Vmax for chronically
MPTP/p-treated mice (2.48 F 0.04 nmol mg protein�1
min�1) does not differ significantly from that of the
probenecid controls (2.36 F 0.12 nmol mg protein�1
min�1; P = 0.616). The values for Km are not
significantly different between groups (7.75 F 1.1 AMfor probenecid vs. 8.69 F 0.43 AM for MPTP/p treated
animals; P = 0.377). Fig. 7B shows that the Vmax for
animals treated acutely with MPTP (2.68 F 0.08 nmol
mg protein�1 min�1) also does not differ significantly
from saline-treated controls (2.71 F 0.07 nmol mg
protein�1 min�1; P = 0.772) and the values for Km also
A.G. Dervan et al. / Experimental Neurology 190 (2004) 145–156 153
did not differ significantly between groups (8.97 F 0.61
AM for saline vs. 9.81 F 0.74 AM for MPTP treated
mice; P = 0.914).
Comparisons between acute and chronic treatments
show no significant difference in Vmax (ANOVA, F =
0.497; P = 0.689) or Km (F = 0.463; P = 0.711) between
groups. Furthermore, there is no difference between the
probenecid and saline control groups (Vmax, P = 0.299;
Km, P = 0.465).
GLT-1 labeling on GFAP-immunolabeled astrocytes
The GLT-1 transporter is the glutamate transporter
commonly found on striatal astrocytes. As our transport
studies showed that the number of functional transporters in
the striata of MPTP/p-treated mice did not differ from that
of controls (Vmax was unchanged), but our stereological
analysis revealed a doubling in the number of astrocytes
after MPTP/p treatment, we hypothesized that the density of
transporters per astrocyte would be lower in the toxin-
treated group compared to controls. To test this possibility,
we employed dual-label confocal microscopy under con-
trolled conditions (Figs. 8A–F). Qualitatively, sections from
MPTP/p-treated mice showed more GFAP-labeled pro-
cesses in each field. GLT-1 immunoreactivity on the cell
soma and proximal processes of GFAP-immunopositive
cells appeared less obvious when compared to controls
(compare Fig. 8C with F). We quantified the density of
GLT-1 immunoreactivity per GFAP-immunopositive astro-
cyte by capturing merged images and averaging the pixel
densities after inversion of each image (see Materials and
methods). After pooling the data for each group and
comparing groups with a Student’s t test, we found that
the mean optical density for the MPTP/p-treated group
(104.6 F 17 SEM) was significantly lower (P b 0.05) than
that in controls (174.6 F 25 SEM). This result can also be
interpreted as a 40% reduction in GLT-1 density per
Fig. 8. Confocal images showing the cellular localization of GFAP (A, D) and GL
month after final treatment. Four animals from the MPTP/p group and four probe
localization on astrocytes from MPTP/p treated animals (F) appears to be less (y
astrocyte after MPTP/p treatment compared to controls
(compare Fig. 8C with F).
Discussion
Many of the classic symptoms of Parkinson’s disease
are thought to arise by elevated glutamate levels follow-
ing the loss of DA (Fornai et al., 1997). Experimental
studies have shown increased striatal neuron activity after
complete 6-OHDA lesions (Schultz and Ungerstedt, 1978)
and elevated glutamate 1 month after acute MPTP
(Robinson et al., 2003). Although glutamatergic pathways
to the striatum could potentially show sustained over-
activity after DA loss (Fornai et al., 1997), there are data
to suggest that the striatum compensates remarkably well
for DA reductions, especially if the loss is progressive
(Zigmond et al., 1990). In the present study, we show for
the first time that extracellular glutamate and its transport
kinetics are not changed 13 weeks after the start of
chronic MPTP/p treatment, although greater than 90% of
the striatal DA function is gone (Petroske et al., 2001).
The question arises as to the mechanism responsible for
such results. Our study indicates that it is due to a
significant increase in reactive astrocytes, which through
their large numbers maintain striatal homeostasis. Never-
theless, each GFAP-immunopositive cell has a reduced
complement of GLT-1 transporters, a change that may
eventually contribute to disease pathophysiology.
Changes in extracellular glutamate could be reflected in
the time course of DA loss. Acute MPTP treatment, which
down-regulates TH levels within 24 h (Mandel et al., 2002),
is associated with significant elevations in extracellular
glutamate after 4 weeks (Robinson et al., 2003). In the
chronic model, DA levels decline progressively (Petroske et
al., 2001), and although earlier time points have not been
studied, striatal glutamate probably rises rapidly after each
T-1 (B, E) in probenecid-treated (A–C) and MPTP/p-treated mice (D–F) 1
necid controls were used in this study. In merged images (C, F), the GLT-1
ellow) compared to the probenecid controls (C). Scale bar = 10 Am.
A.G. Dervan et al. / Experimental Neurology 190 (2004) 145–156154
dose of MPTP but then subsides in parallel with astrocytic
proliferation. Certainly, the striatal, astroglial reaction to 6-
OHDA rises and falls with time after the lesion (Tripanich-
kul, 2000).
The rise in striatal extracellular glutamate following the
single dose of l-DOPA in MPTP/p-treated mice is
consistent with results after acute MPTP treatment (Rob-
inson et al., 2003) and presumably reflects changes in basal
ganglia function (Albin et al., 1989). DA stimulation, either
indirectly via l-DOPA to restore brain levels of DA, or via
DA D-1 or D-2 receptors in the striatum, would lead to a
decrease in the GABAergic output from the basal ganglia to
the motor thalamus, resulting in increased activity of the
corticostriatal pathway.
The striatum appears to adapt to glutamate changes by up-
regulating astrocytic function through an increase in GFAP-
immunoreactive cells. The ability of glia to regulate
glutamate levels is well known, and early elevations in
glutamate could be counteracted by a rapid astrocytic reaction
that fades later in a manner similar to that described in models
of CNS injury (see Ridet et al., 1997). Acute MPTP treatment
does trigger a reactive gliosis, but it is short-lived (Muramatsu
et al., 2003), which may explain the lack of change in
transporter kinetics (present results) at 1 week post treatment,
and the high level of extracellular glutamate 1 month later
(Robinson et al., 2003). Thus, reduced levels of GFAP
immunoreactivity found in postmortem PD striatum (Forno et
al., 1992) could be due to the late stage of the disease and not
to a lack of a glial response, which may have peaked earlier.
Moreover, subacute l-DOPA treatment significantly elevates
GLT-1 mRNA (Lievens et al., 2001), a change that may
enhance glutamate recycling and limit gliosis.
We have measured high-affinity uptake using synaptoso-
mal preparations, which are generally considered to be
neuronal in nature. Nevertheless, there is good evidence that
uptake activities by these particles reflect glial-mediated
transport (see Danbolt, 2001), presumably because glial cell
fragments contaminate the preparation (Nakamura et al.,
1993) and because mice deficient in the GLT-1 transporter
show greatly reduced glutamate uptake from synaptosomes
compared to controls (Tanaka et al., 1997).
Our results show that the kinetics of glutamate are
unchanged after chronic MPTP. Although single application
of MPP+ to astrocytic cultures is accompanied by impaired
glutamate uptake (Di Monte et al., 1999), the lack of change
seen in our study may not be surprising because the animals
were analyzed 5 weeks after an extended exposure to MPTP.
Other in vivo studies have shown that nigrostriatal denerva-
tion has no effect on glutamate transporter mRNA (Lievens et
al., 2001).
As the basis of MPTP toxicity is negotiated via astrocytic
uptake and conversion (Markey et al., 1984), it is unclear
whether MPP+ affects host astrocyte function. When
compared to their mesencephalic counterparts, striatal
astrocytes seem to have a stronger intrinsic facility for
dealing with oxidative stress, especially in the presence of
MPTP (Wong et al., 1999). Striatal astrocytes can increase
levels of superoxide dismutase, which affords them
increased protection by absorbing reactive oxygen species
(Wong et al., 1999). Therefore, it seems unlikely that the
glial changes seen in the present study are strictly due to the
inability of striatal astrocytes to cope with the pyridinium
ion, MPP+, but see Di Monte et al. (1999). Once reactive,
astrocytes show a reduced ability to carry out normal
support functions (Behrens et al., 2002; Ginsberg et al.,
1995; Krum et al., 2002; Masliah et al., 1996; Ridet et al.,
1997). Therefore, their activation could conceivably con-
tribute to progressive brain pathology (Blandini et al., 1996;
Olanow and Tatton, 1999).
Glutamate rapidly diffuses out of the cleft of excitatory
synapses (Clements, 1996) and is quickly cleared by
glutamate transporters, as measured in hippocampal cultures
(Diamond and Jahr, 1997; Tong and Jahr, 1994). Although
reuptake blockers have essentially failed to prolong AMPA
receptor-mediated synaptic currents at some synapses (Hes-
trin et al., 1990; Isaacson and Nicoll, 1993; Sarantis et al.,
1993), glutamate transporters contribute to synaptic efficacy
if AMPA receptor desensitization is blocked (Mennerick
and Zorumski, 1994). Glutamate transporters affect both the
duration and amplitude of excitatory postsynaptic currents
and ultimately influence the induction of synaptic plasticity
(Bowers and Kalivas, 2003; Diamond and Jahr, 1997;
Mennerick and Zorumski, 1994; Turecek and Trussell,
2000). Moreover, any reduction in uptake sites for
glutamate, as seen in the present results, could increase
the vulnerability of striatal neurons to toxic insults (McBean
and Roberts, 1985). Maintaining the normal complement of
glutamate transporters appears to be critical for modulating
synaptic strength, and altered synaptic efficacy, rather than
elevated glutamate, could therefore be a strong contributing
factor to progressive pathogenesis in PD.
Acknowledgments
This research was supported by a USPHS NS41799
(GEM), a Wellcome Trust Biomedical Collaboration Grant
(GEM and ST), and the Department of Veterans Affairs Merit
Review Program (CKM). We thank Dr. David Pow,
University of Queensland, Brisbane, Australia, for his
generous gift of the GLT1 antibodies and Prof. B.L. Roberts
for his comments on the manuscript. We also thank Heather
Milligan and Julie Frederickson for their technical assistance.
References
Albin, R.L., Young, A.B., Penney, J.B., 1989. The functional anatomy of
basal ganglia disorders. Trends Neurosci. 12, 366–375.
Araki, T., Kumagai, T., Tanaka, K., Matsubara, M., Kato, H., Itoyama, Y.,
Imai, Y., 2001. Neuroprotective effect of riluzole in MPTP-treated mice.
Brain Res. 918, 176–181.
Araque, A., Parpura, V., Sanzgirl, R.P., Haydon, P.G., 1998. Glutamate-
A.G. Dervan et al. / Experimental Neurology 190 (2004) 145–156 155
dependent astrocyte modulation of synaptic transmission between
cultured hippocampal neurons. Eur. J. Neurosci. 10, 2129–2142.
Behrens, P.F., Franz, P., Woodman, B., Lindenberg, K.S., Landwehrmeyer,
G.B., 2002. Impaired glutamate transport and glutamate–glutamine
cycling: downstream effects of the Huntington mutation. Brain 125,
1908–1922.
Blandini, F., Porter, R.H., Greenamyre, J.T., 1996. Glutamate and
Parkinson’s disease. Mol. Neurobiol. 12, 73–94.
Bowers, M.S., Kalivas, P.W., 2003. Astroglial plasticity is induced
following withdrawal from repeated cocaine administration. Eur. J.
Neurosci. 17, 1273–1278.
Brouillet, E., Beal, M.F., 1993. NMDA antagonists partially protect against
MPTP induced neurotoxicity in mice. NeuroReport 4, 387–390.
Carboni, S., Melis, F., Pani, L., Hadjiconstantinou, M., Rossetti, Z.L., 1990.
The non-competitive NMDA-receptor antagonist MK-801 prevents the
massive release of glutamate and aspartate from rat striatum induced by
1-methyl-4-phenylpyridinium (MPP+). Neurosci. Lett. 117, 129–133.
Coggeshall, R.E., 1992. A consideration of neural counting methods.
Trends Neurosci. 15, 9–13.
Clements, J.D., 1996. Transmitter timecourse in the synaptic cleft: its role in
central synaptic function. Trends Neurosci. 19, 163–171.
Danbolt, N.C., 2001. Glutamate uptake. Prog. Neurobiol. 65, 1–105.
Dervan,A.G., Totterdell, S., Lau,Y.-S.,Meredith, G.E., 2003a. Altered striatal
neuronal morphology is associated with astrogliosis in a chronic mouse
model of ParkinsonTs disease. Ann. N. Y. Acad. Sci. 991, 291–294.Dervan, A.G., Beales, M., Frederickson, J., Meshul, C.K., McBean, G.J.,
Snyder, A.K., Meredith, G.E., 2003b. Chronic MPTP/Probenecid
treatment induces a significant gliosis but does not alter striatal
glutamate levels or transport. Soc. Neurosci. Program No 204.13.
Abstract Viewer/Itinerary Planner. Online.
De Souza, I.E., McBean, G.J., Meredith, G.E., 1999. Chronic haloperidol
treatment impairs glutamate transport in the rat striatum. Eur. J.
Pharmacol. 382, 139–142.
Diamond, J.S., Jahr, C.E., 1997. Transporters buffer synaptically released
glutamate on a submillisecond time scale. J. Neurosci. 17, 4672–4687.
Di Monte, D.A., Tokar, I., Langston, J.W., 1999. Impaired glutamate
clearance as a consequence of energy failure caused by MPP(+) in
astrocytic cultures. Toxicol. Appl. Pharmacol. 158, 296–302.
Eddleston, M., Mucke, L., 1993. Molecular profile of reactive astro-
cytes—implications for their role in neurologic disease. Neuroscience
54, 15–36.
Eng, L.F., Lee, Y.L., 1995. Intermediate filaments in astrocytes. In:
Kettenmann, H., Ransom, B.R. (Eds.), Neuroglia. Oxford Univ. Press,
New York, pp. 650–667.
Fornai, F., Vaglini, F., Maggio, R., Bonuccelli, U., Corsini, G.U., 1997.
Species differences in the role of excitatory amino acids in experimental
parkinsonism. Neurosci. Biobehav. Rev. 21, 401–415.
Forno, L.S., DeLanney, L.E., Irwin, I., Di Monte, D., Langston, J.W., 1992.
Astrocytes and Parkinson’s disease. Prog. Brain Res. 94, 429–436.
Franklin, K.B.J., Paxinos, G., 1997. The Mouse Brain in Stereotaxic
Coordinates. Academic Press, San Diego.
Ginsberg, S.D., Martin, L.J., Rothstein, J.D., 1995. Regional deafferenta-
tion down-regulates subtypes of glutamate transporter proteins. J.
Neurochem. 65, 2800–2803.
Greene, J.G., Greenamyre, J.T., 1995. Exacerbation of NMDA, AMPA, and
l-glutamate excitotoxicity by the succinate dehydrogenase inhibitor
malonate. J. Neurochem. 64, 2332–2338.
Hepler, J.R., Toomim, C.S., McCarthy, K.D., Conti, F., Battaglia, G.,
Rustioni, A., Petrusz, P., 1988. Characterization of antisera to glutamate
and aspartate. J. Histochem. Cytochem. 36, 13–22.
Hestrin, S., Sah, P., Nicoll, R.A., 1990. Mechanisms generating the time
course of dual component excitatory synaptic currents recorded in
hippocampal slices. Neuron 5, 247–253.
Ingham, C.A., Hood, S.H., Arbuthnott, G.W., 1989. Spine density on
neostriatal neurons changes with 6-hydroxydopamine lesions and with
age. Brain Res. 503, 334–338.
Isaacson, J.S., Nicoll, R.A., 1993. The uptake inhibitor l-trans-PDC
enhances responses to glutamate but fails to alter the kinetics of
excitatory synaptic currents in the hippocampus. J. Neurophysiol. 70,
2187–2191.
Krum, J.M., Phillips, T.M., Rosenstein, J.M., 2002. Changes in astroglial
GLT-1 expression after neural transplantation or stab wounds. Exp.
Neurol. 174, 137–149.
Lau, Y.-S., Trobough, K.L., Crampton, J.M., Wilson, J.A., 1990. Effects of
probenecid on striatal dopamine depletion in acute and long-term 1-
methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP)-treated mice. Gen.
Pharmacol. 21, 181–187.
Lievens, J.C., Salin, P., Nieoullon, A., Kerkerian-Le Goff, L., 2001.
Nigrostriatal denervation does not affect glutamate transporter mRNA
expression but subsequent levodopa treatment selectively increases
GLT1 mRNA and protein expression in the rat striatum. J. Neurochem.
79, 893–902.
Lin, C.L., Bristol, L.A., Jin, L., Dykes-Hoberg, M., Crawford, T., Clawson,
L., Rothstein, J.D., 1998. Aberrant RNA processing in a neuro-
degenerative disease: the cause for absent EAAT2, a glutamate
transporter, in amyotrophic lateral sclerosis. Neuron 20, 589–602.
Mandel, S., Grunblatt, E., Maor, G., Youdim, M.B., 2002. Early and late
gene changes in MPTP mice model of Parkinson’s disease employing
cDNA microarray. Neurochem. Res. 27, 1231–1243.
Markey, S.P., Johannessen, J.N., Chiueh, C.C., Burns, R.S., Herkenham,
M.A., 1984. Intraneuronal generation of a pyridinium metabolite may
cause drug-induced parkinsonism. Nature 311, 464–467.
Masliah, E., Sisk, A., Mallory, M., Mucke, L., Schenk, D., Games, D.,
1996. Comparison of neurodegenerative pathology in transgenic mice
overexpressing V717F beta-amyloid precursor protein and Alzheimer’s
disease. J. Neurosci. 16, 5795–5811.
McBean, G.J., Roberts, P.J., 1985. Neurotoxicity of l-glutamate and
dl-threo-3-hydroxyaspartate in the rat striatum. J. Neurochem. 44,
247–254.
McNeill, T.H., Brown, S.A., Rafols, J.A., Shoulson, I., 1998. Atrophy of
medium spiny I striatal dendrites in advanced Parkinson’s disease. Brain
Res. 455, 148–152.
Mennerick, S., Zorumski, C.F., 1994. Glial contributions to excitatory
neurotransmission in cultured hippocampal cells. Nature 368, 59–62.
Meredith, G.E., Ypma, P., Zahm, D.S., 1995. Effects of dopamine depletion
on the morphology of medium spiny neurons in the shell and core of the
rat nucleus accumbens. J. Neurosci. 15, 3808–3820.
Meredith, G.E., Totterdell, S., Petroske, E., Santa Cruz, K., Callison Jr.,
R.C., Lau, Y.-S., 2002. Lysosomal malfunction accompanies alpha-
synuclein aggregation in a progressive mouse model of ParkinsonTsdisease. Brain Res. 956, 156–165.
Meshul, C.K., Allen, C., 2000. Haloperidol reverses the changes in striatal
glutamatergic immunolabeling following a 6-OHDA lesion. Synapse
36, 129–142.
Meshul, C.K., Stallbaumer, R.K., Taylor, B., Janowsky, A., 1994.
Haloperidol-induced synaptic changes in striatum are associated with
glutamate synapses. Brain Res. 648, 181–195.
Meshul, C.K., Emre, N., Nakamura, C.M., Allen, C., Donohue, M.K.,
1999. Time-dependent changes in striatal glutamate synapses following
a 6-OHDA lesion. Neuroscience 88, 1–16.
Meshul, C.K., Kamel, D., Moore, C., Kay, T.S., Krentz, L., 2002. Nicotine
alters striatal glutamate function and decreases the apomorphine-
induced contralateral rotations in 6-OHDA lesioned rats. Exp. Neurol.
175, 257–274.
Mfller, A., Strange, P., Gundersen, H.J., 1990. Efficient estimation of cell
volume and number using the nucleator and the dissector. J. Microsc.
159, 61–71.
Muramatsu, Y., Kurosaki, R., Watanabe, H., Michimata, M., Matsubara,
M., Imai, Y., Araki, T., 2003. Cerebral alterations in a MPTP-mouse
model of Parkinson’s disease—an immunocytochemical study. J.
Neural Transm. 110, 1129–1144.
Nakamura, Y., Iga, K., Shibata, T., Shudo, M., Kataoka, K., 1993. Glial
plasmalemmal vesicles: a subcellular fraction from rat hippocampal
homogenate distinct from synaptosomes. Glia 9, 48–56.
A.G. Dervan et al. / Experimental Neurology 190 (2004) 145–156156
Olanow, C.W., Tatton, W.G., 1999. Etiology and pathogenesis of
Parkinson’s disease. Annu. Rev. Neurosci. 22, 123–144.
Petroske, E., Meredith, G.E., Callen, S., Totterdell, S., Lau, Y.-S., 2001.
Mouse model of Parkinsonism: a comparison between subacute MPTP
and chronic MPTP/probenecid treatment. Neuroscience 106, 589–601.
Phend, K.D., Weinberg, R.J., Rustioni, A., 1992. Techniques to optimize
post-embedding single and double staining for amino acid neuro-
transmitters. J. Histochem. Cytochem. 40, 1011–1020.
Reye, P., Sullivan, R., Fletcher, E.L., Pow, D.V., 2002. Distribution of two
splice variants of the glutamate transporter GLT1 in the retinas of
humans, monkeys, rabbits, rats, cats, and chickens. J. Comp. Neurol.
445, 1–12.
Ridet, J.L., Malhotra, S.K., Privat, A., Gage, F.H., 1997. Reactive
astrocytes: cellular and molecular cues to biological function. Trends
Neurosci. 20, 570–577.
Robinson, T.E., Wishaw, I.Q., 1988. Normalization of extracellular
dopamine in striatum following recovery from a partial unilateral 6-
OHDA lesion of the substantia nigra: a microdialysis study in freely
moving rats. Brain Res. 450, 209–224.
Robinson, S., Krentz, L., Moore, C., Meshul, C.K., 2001. Blockade of
NMDA receptors by MK-801 reverses the changes in striatal glutamate
immunolabeling in 6-OHDA-lesioned rats. Synapse 42, 54–61.
Robinson, S., Freeman, P., Moore, C., Touchon, J.C., Krentz, L., Meshul,
C.K., 2003. Acute and subchronic MPTP administration differentially
affects striatal glutamate synaptic function. Exp. Neurol. 180, 73–86.
Rothstein, J.D., Martin, L.J., Kuncl, R.W., 1992. Decreased glutamate
transport by the brain and spinal cord in amyotrophic lateral sclerosis.
N. Engl. J. Med. 326, 1464–1468.
Rothstein, J.D., Van Kammen, M., Levey, A.I., Martin, L.J., Kuncl, R.W.,
1995. Selective loss of glial glutamate transporter GLT-1 in amyo-
trophic lateral sclerosis. Ann. Neurol. 38, 73–84.
Sarantis, M., Ballerini, L., Miller, B., Silver, R.A., Edwards, M., Attwell,
D., 1993. Glutamate uptake from the synaptic cleft does not shape the
decay of the non-NMDA component of the synaptic current. Neuron 11,
541–549.
Schultz, W., Ungerstedt, U., 1978. Short-term increase and long-term
reversion of striatal cell activity after degeneration of the nigrostriatal
dopamine system. Exp. Brain Res. 33, 159–171.
Schuster, R., 1988. Determination of amino acids in biological, pharma-
ceutical, plant and food samples by automated precolumn derivatization
and high-performance liquid chromatography. J. Chromatogr. 431,
271–284.
Scott, H.L., Pow, D.V., Tannenberg, A.E., Dodd, P.R., 2002. Aberrant
expression of the glutamate transporter excitatory amino acid trans-
porter 1 (EAAT1) in Alzheimer’s disease. J. Neurosci. 22, RC206.
Sheng, J.G., Shirabe, S., Nishiyama, N., Schwartz, J.P., 1993.
Alterations in striatal glial fibrillary acidic protein expression in
response to 6-hydroxydopamine-induced denervation. Exp. Brain Res.
95, 450–456.
Slezak, M., Pfrieger, F.W., 2003. New roles for astrocytes: regulation of
CNS synaptogenesis. Trends Neurosci. 26, 531–535.
Tanaka, K., Watase, K., Manabe, T., Yamada, K., Watanabe, M., Takahashi,
K., Iwama, H., Nishikawa, T., Ichihara, N., Kikuchi, T., Okuyama, S.,
Kawashima, N., Hori, S., Takimoto, M., Wada, K., 1997. Epilepsy and
exacerbation of brain injury in mice lacking the glutamate transporter
GLT-1. Science 276, 1699–1702.
Tong, G., Jahr, C.E., 1994. Block of glutamate transporters potentiates
postsynaptic excitation. Neuron 13, 1195–1203.
Tripanichkul, W., 2000. Associations between glia and sprouting of
dopaminergic axons. PhD thesis. Monash University, Australia.
Turecek, R., Trussell, L.O., 2000. Control of synaptic depression by
glutamate transporters. J. Neurosci. 20, 2054–2063.
Turski, L., Bressler, K., Rettig, K.-J., Lfschmann, P.-A., Wachtel, H., 1991.
Protection of substantia nigra from MPP+ neurotoxicity by N-methyl-d-
aspartate antagonists. Nature 349, 414–418.
Ullian, E.M., Sapperstein, S.K., Christopherson, K.S., Barres, B.A., 2001.
Control of synapse number by glia. Science 291, 657–661.
West, M.J., Gundersen, H.J., 1990. Unbiased stereological estimation of the
number of neurons in the human hippocampus. J. Comp. Neurol. 296,
1–22.
Wong, S.S., Li, R.H., Stadlin, A., 1999. Oxidative stress induced by MPTP
and MPP+: selective vulnerability of cultured mouse astrocytes. Brain
Res. 836, 237–244.
Zigmond, M.J., Abercrombie, E., Berger, T.W., Grace, A.A., Stricker, E.M.,
1990. Compensations after lesions of central dopaminergic neurons:
some clinical and basic implications. Trends Neurosci. 13, 290–296.