From the Laboratoire de Biologie du Stress Oxydant
(LBSO), Faculté de Pharmacie, Domaine de La Merci, 38706 La
Tronche-Grenoble cedex 9, France, the ¶ Unité de Virologie
Immunologie Moléculaires, Institut National de la Recherche
Agronomique (INRA), 78350 Jouy-en Josas, France, the
Institut de
Génétique Humaine, CNRS U.P.R. 1142, 141, rue de la
Cardonille, 34396 Montpellier Cedex 5, France, and the
** Laboratoire des Lésions des Acides Nucléiques,
UMR CNRS/CEA/UJF, 5046, Avenue des Martyrs, 38000 Grenoble, France
Received for publication, November 20, 2002
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ABSTRACT |
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The N-terminal region of the prion protein
PrPC contains a series of octapeptide repeats. This
region has been implicated in the binding of divalent metal ions,
particularly copper. PrPC has been suggested to be involved
in copper transport and metabolism and in cell defense mechanisms
against oxidative insult, possibly through the regulation of the
intracellular CuZn superoxide dismutase activity (CuZn-SOD) or a
SOD-like activity of PrPC itself. However, up to now
the link between PrPC expression and copper metabolism or
SOD activity has still to be formally established; particularly because
conflicting results have been obtained in vivo. In this
study, we report a link between PrPC, copper binding, and
resistance to oxidative stress. Radioactive copper (64Cu)
was used at a physiological concentration to demonstrate that binding
of copper to the outer plasma cell membrane is related to the level of
PrPC expression in a cell line expressing a
doxycycline-inducible murine PrPC gene. Cellular PIPLC
pretreatment indicated that PrPC was not involved in copper
delivery at physiological concentrations. We also demonstrated that
murine PrPC expression increases several antioxidant enzyme
activities and glutathione levels. Prion protein may be a stress
sensor sensitive to copper and able to initiate, following copper
binding, a signal transduction process acting on the antioxidant
systems to improve cell defenses.
Prion diseases form a group of fatal neurodegenerative disorders
including Creutzfeldt-Jakob diseases, Gerstmann-Sträussler Syndrome, Kuru and Fatal Familial Insomnia in humans, and scrapie and
bovine spongiform encephalopathy in animals (1). All these disorders
are characterized by the accumulation of an abnormally folded isoform
of the cellular prion protein
PrPC,1 denoted
PrPSc, which represents the major component of infectious
prion diseases (2). The formation of PrPSc from
PrPC is accompanied by profound changes in structure and
biochemical properties. PrPC rich in Human PrPC has 253 amino acids and is mainly expressed on
neurons (5, 6). In its N-terminal region, a repeated sequence of five
octapeptides can be found, which was shown to bind copper and zinc
(7-9). The protein may have some superoxide dismutase-like activity
and therefore a possible protective function against oxidative stress
(10). Wild-type mouse brains have a significantly higher level of
membrane-associated copper than PrPC-deficient mice.
Treatment with phosphatidylinositol phospholipase C (PIPLC)
specifically reduced the copper content from wild type mice but had no
effect on the copper content of PrPC knockout mice (8).
However, these results have not been confirmed (11). Incorporation of
radioactive-labeled copper into CuZn-SOD was found to be proportional
to the level of PrP expression (12). Pattison and Jebbett (13) noticed
more than 30 years ago the similarity between prion histopathology to
the histopathology induced by a copper chelator, cuprizone. The
incidence of chronic Wasting disease (CWD), a sporadic prion disease of
deer and elk, was observed to be higher in regions where the soil had a
low copper content (14). Therefore, prion diseases may be related to an
alteration of copper transport and a loss of copper-enzyme activities.
In a previous work, we demonstrated that neuronal cells infected with
prion strains resulted in an alteration of the molecular mechanism
promoting cellular resistance to ROS (15). The same alteration of
antioxidant enzymes was shown in infected animals (16, 17). In the
present study, we used a transfected transgenic cell line with a
doxycycline-inducible murine PrPC gene to investigate the
involvement of PrPC in copper metabolism and in the
resistance mechanism to toxic stress.
Cell Culture and Construction--
Murine PrPC was
cloned in the pTRE plasmid (Clontech), and the
resulting plasmid was transfected by the LipofectAMINE method (Invitrogen) into rabbit kidney epithelial cells (RK13) (18, 19).
Stable transfectants were selected in the presence of puromycin (1 µg/ml), and one (A74) was amplified for further study. RK13 and A74
cells were grown at 37 °C in a 5% CO2-enriched
atmosphere in minimal essential medium supplemented with 10%
heat-inactivated fetal calf serum and were usually split at one-fourth
dilution each week.
Immunofluorescence and Western Blot
Analysis--
Immunofluorescence analysis on living A74 cells was
performed at 4 °C, with rabbit polyclonal antibody P45-66, raised
against synthetic peptide encompassing mouse PrPC (MoPrP)
residues 45-66. Fluorescein-conjugated IgG was used as second antibody.
For Western blot analysis, confluent cells were washed twice with cold
phosphate-buffered saline, calcium- and magnesium-free, and lysed for
30 min at 4 °C in Triton-deoxycholate lysis buffer (1× buffer is
150 mM NaCl, 0.5% Triton X-100, 0.5% sodium deoxycholate, and 50 mM Tris-HCl, pH 7.5) plus protease inhibitors. After
1 min of centrifugation at 10,000 × g, the supernatant
was collected, and its protein concentration was measured by the BCA
assay (Pierce). The equivalent of 20 µg of total protein in SDS
loading buffer was subjected to 12% SDS-PAGE electrophoresis followed
by electroblotting on polyvinylidene difluoride in Tris-glycine buffer
containing 20% methanol. The membrane was blocked with 5% nonfat dry
milk in TBST (0.1% Tween 20, 100 mM NaCl, 10 mM Tris-HCL, pH 7.8) for 1 h at room temperature, and
MoPrP was detected by immunoblotting with P45-66 antibody as
previously described (20). After adding the second antibody
(horseradish peroxidase-coupled rabbit IgG), immunoreactive proteins
were detected with ECL Western blot system. Quantification was achieved
by densitometric scanning.
PrPC analysis in culture medium was immunodetected by
Western blot. To release cell surface PrPC, cultures were
treated with PIPLC (0.2 units/ml) in opti-MEM serum-free medium at
37 °C for 2 h. Proteins were precipitated from the PIPLC
incubation medium with at least 4 volumes of methanol at Cellular 64Cu Binding--
Cells (RK13 and A74) were
cultured in 35-mm Petri dishes. Culture medium was replaced by 2 ml of
fresh complete medium containing different concentrations of dox
(0-500 ng/ml) to stimulate murine PrPC expression in A74
cells, and 1.6 µM 64Cu (CIS biointernational,
Gif-sur-Yvette, France; specific activity 20 mCi/mg) to evaluate copper
binding to cells as a function of the level of murine PrPC
expression. Non-transfected RK13 cells were used as control and treated
under the same conditions. Cells were incubated at 37 °C under 5%
CO2. The radioactive medium was removed after 30-40 min,
2, 4, 8, 10, 24, and 26 h. Cells were rinsed twice with 2 ml of
diluted Puck's saline A solution (Invitrogen), and harvested after
addition of 1 ml of 0.25% trypsin. After harvesting, each dish was
rinsed with 1 ml of Puck's saline A solution. The final 2 ml obtained
for each dish were counted for 2 min using a Packard Cobra III, mono
well gamma counter (Packard Instrument Company, Meriden, CT). Protein
content was assayed with the BCA protein assay reagent kit. Data were
analyzed using a "self made" computer half-life calculation
program, to obtain results as µCi of 64Cu incorporated or
retained per mg of protein.
Cellular Copper Determination--
Stimulated (500 ng/ml dox for
24 h) or unstimulated A74 cells were cultured in the presence or
absence of 100 µM CuSO4 for 1 h.
For intracellular copper determination, cells were trypsinized, washed
three times in Ca/Mg-free phosphate-buffered saline, and lysed by three
cycles of freeze-thawing. Lysates (total extract) were then centrifuged
at 13,000 rpm for 10 min to obtain the soluble fraction. Copper
concentration was determined by electrothermal atomic absorption
spectrophotometry (PerkinElmer Life Sciences). Their levels were
normalized to the protein content, measured with a protein assay kit.
Cell Viability Assay--
Cell viability was determined by a
modified 3-(4,5-dimethyl-thiazol-2-yl)-2,5-diphenyl-tetrazolium bromide
(MTT) assay (21). Briefly 3000 cells per well were plated in 96-well
microtiter plates with 100 µl of complete medium. The next day, the
medium was changed, and the cells were challenged for 24 h with
different drugs. The medium was then changed, and the cells were
incubated for an additional 24 h without drugs. For the MTT assay,
10 µl of MTT (5 mg/ml stock in phosphate-buffered saline) were added to each well for 3 h at 37 °C. 100 µl of dimethyl sulfoxide
(Me2SO) were added to dissolve the formazan crystals, and
plates were shaken for 5 min on a plate shaker to ensure adequate
solubilization. The absorbance readings for each well were performed at
570 nm using Multiscan ascent plate reader (Labsystems). The absorbance is proportional to viable cell number, and survival was calculated as
the percentage of the staining values of untreated cultures.
Lipid Peroxidation--
Lipid peroxidation was evaluated using
an assay based on fluorescence of thiobarbituric acid reactants
measured after extraction with n-butyl alcohol (22).
Subconfluent cells were trypsinized in 75-cm2 flasks,
washed three times by 10 ml of isotonic, trace element-free Tris-HCl
buffer (400 mM, pH 7.3), and then lysed in hypotonic Tris-HCl buffer (20 mM) by five freeze-defrost cycles. 750 µl of a mixture of thiobarbituric acid at 8 g/7% perchloric acid (2:1) were added to a 100-µl sample. After agitation the mixture was
placed in a 95 °C water bath for 60 min and then cooled in an ice
bath. The fluorescent compound was extracted by mixing with
n-butyl alcohol for 2 min. After centrifugation the
fluorescence in the n-butyl alchohol phase was determined
with an Aminco-Bowman fluorimeter (PerkinElmer Life Sciences) with
excitation at 532 nm and emission at 553 nm. A blank was run for each
sample. The calibration curve was created with a stock solution of
1,1,3,3-tetraethoxypropane prepared in alcohol. The results were
expressed as TBARS, µmol/g of protein.
Superoxide Dismutase Activity--
For SOD activity,
subconfluent cells in 75-cm2 flasks, were washed three
times and collected in 10 ml of isotonic, trace element-free Tris-HCl
buffer (400 mM, pH 7.3), and lysed in hypotonic Tris-HCl buffer (20 mM) by five freeze-defrost cycles. After 10 min
of centrifugation at 4000 rpm, 4 °C, the lysate was assayed for
metalloenzyme activities and soluble protein content. Total SOD,
Mn-SOD, and CuZn-SOD were determined using the pyrogallol assay
following the procedure described by Marklund and Marklund (23), based on the competition between pyrogallol oxidation by superoxide radicals
and superoxide dismutation by SOD, and spectrophotometrically read at
420 nm. Briefly, 50 µl of the sample were added to 1870 µl of Tris
(50 mM)-DTPA (1 mM)-cacodylic acid buffer, pH
8.3 and to 80 µl of pyrogallol (10 mM) in order to induce
an absorbance change of 0.02 in the absence of SOD. The amount of SOD
inhibiting the reaction rate by 50% in the given assay conditions was
defined as one SOD unit. The specific CuZn-SOD inhibition by KCN (60 µl of KCN, 54 mM) added to 300 µl of lysate allowed the
Mn-SOD determination under the same conditions. Each sample was assayed
twice, and results were expressed as SOD units and normalized to the
cell protein content.
Analysis of Glutathione-dependent Antioxidant
System--
For the determination of total glutathione levels,
confluent cells in 25-cm2 flasks were washed three times in
phosphate-buffered saline and collected in 10 ml of isotonic, trace
element-free Tris-HCl buffer (400 mM, pH 7.3), and lysed in
hypotonic Tris-HCl buffer (20 mM) by five freeze-defrost
cycles. Samples of whole lysate were deproteinized by adding
metaphosphoric acid (6%) (lysate-metaphosphoric 5:1, v/v). After 10 min at 4 °C the solutions were spun at 4000 rpm for 10 °C at
4 °C, and the supernatants were assayed for total glutathione
content according to the Akerboom and Sies method (24).
The glutathione peroxidase (GPX) activity was assayed by the method of
Gunzler et al. (25). GPX was measured in a coupled reaction
with glutathione reductase (GR), using
tert-butylhydroperoxide as substrate. Briefly, 25 µl of
the sample were added to 900 µl of Tris (50 mM), EDTA,
sodium azide buffer, pH 7.6 (azide was included in the assay mixture to
inhibit interference of catalase) and 20 µl of glutathione (0.15 M), 20 µl of glutathione reductase (200 units/ml), 20 µl of NADPH2 (8.4 mM) in order and incubated for 1 min for mixture equilibrium. Then 20 µl of
tert-butylhydroperoxide were added, and the decrease of the
absorbance was monitored at 340 nm for 200 s. The difference in
absorbance per minute was used to calculate the enzyme activity, and
results were expressed as GPX units/g of protein.
GR activity was determined by following the oxidation of NADPH to NADP+
during the reduction of oxidized glutathione (GSSG) (26). The main
reagent was prepared by combining 18 ml of
KH2PO4 (139 mM), EDTA (0.76 mM), pH 7.4, and 2 ml of NADPH2 (2.5 mM). 20 µl of sample were added with 220 µl of the main
reagent, and then 30 µl of GSSG (22 mM) plus 10 µl of
KH2PO4 were added to start the reaction; the
absorbance was followed at 340 nm for 175 s. The difference in
absorbance per min was used to calculate the activity of the enzyme.
The results were expressed as glutathione reductase units/g of protein.
Doxycycline-inducible Expression of Murine PrP in A74
Cells--
We used the tetracycline-inducible (tet-on) system (18, 27)
to achieve regulated high-level expression of the murine
PrPC. After transfection of several cell lines, a strong
inducible expression of murine PrPC was obtained in most of
the clones derived from a rabbit kidney epithelial cell line (RK13).
Data obtained with a representative clone (A74) are presented in this
article. Expression of murine PrPC was related to dox
concentration in the culture medium, detectable at 10 ng/ml dox and
reaching a maximum at 500 ng/ml of dox (Fig. 1). No PrPC was detected in
either unstimulated A74 cells (Fig. 1) or non-transfected RK13 (data
not shown), confirming that expression of endogenous, rabbit PrP was
undetectable in these cells (18). We also studied the induction
kinetics of PrPC expression in A74 cells stimulated with
500 ng/ml dox. Expression of PrPC can be detected 8 h
after induction, and a plateau was obtained at 24 h (Fig.
2). Not all, although up to 32% of
cells, produced PrPC at a high level and expressed it on
the outer face of the plasma membrane (Fig.
3).
64Cu Binding Is Correlated to PrPC
Expression--
No significant difference in copper binding was
observed during the first 4 h following induction of
PrPC expression (Fig.
4A). This apparent lack of
effect might be due to the lack of detectable PrPC 8 h
following addition of dox (Fig. 2). Then, the 64Cu binding
increased proportionately to dox concentration. 26 h following the
addition of 500 ng/ml of dox, copper binding was 2.7-fold higher in
stimulated cells compared with the unstimulated cells. This increase
was due to PrPC expression and not to an effect of dox on
copper binding because dox had no effect in non-transfected RK13
control cells (Fig. 4B).
The copper binding curve obtained with the non-transfected control
cells RK13 was similar to that obtained with unstimulated A74 (Fig. 4,
A and B). This corresponds to the copper uptake
via the classical transport systems (CTR1 and other potential
transporters), while the increase observed in dox-stimulated A74
corresponds to the binding or transport activity of
PrPC.
To study the influence of PrPC cleavage on copper binding,
the cells were treated for 2 h with 0.2 units/ml PIPLC at 37 °C before measuring copper binding. PIPLC pretreatment dramatically decreases 64Cu binding in stimulated A74 cells (500 ng/ml
dox) but had no effect on unstimulated cells (Fig. 4C).
However, there was a small difference in the kinetics of copper binding
between stimulated cells treated with PIPLC and unstimulated cells
presumably because PIPLC might not have cleaved all PrPC
from membranes. PIPLC pretreatment had no effect on stimulated or
unstimulated non-transfected RK13 cell controls (Fig.
4D).
PrPC Binds Copper at the Outer Side of the Cell
Membrane at Physiological Concentrations--
To investigate the
location of bound copper and check for a real entry, cells were rinsed
twice with 2 ml of diluted Puck's saline A solution (Invitrogen)
24 h after PrPC induction with 500 ng/ml of dox and
incubated in a radioactive medium containing 64Cu.
Treatment with PIPLC decreased the amount of PrPC in
stimulated cells (Fig. 5A)
concomitantly with a high decrease in cell-associated radioactivity
(Fig. 5C). Immunoblotting demonstrated that PrPC
was found only in the medium when stimulated cells (500 ng/ml dox) were
treated with PIPLC (Fig. 5B). The release of
PrPC was correlated with a high increase of radioactivity
in the medium (Fig. 5D). These findings indicate that PrP
binds copper at the outer side of the cell membrane and that cleavage
of PrPC liberates copper into the medium. The difference in
copper binding between stimulated and unstimulated cells was abolished
after treatment with PIPLC indicating that no copper had been taken up
by the PrPC. Therefore, PrPC does not transport
copper inside the cell at physiological concentrations. These results
are confirmed by measuring cellular copper content by electrothermal
atomic absorption spectrophotometry. Copper content was increased up to
~2-fold in the total but not the soluble fraction of stimulated cells
(see Table II). Thus, at physiological concentrations PrPC
did not transport copper from the extracellular medium to cytoplasm since there was no difference in 64Cu content between the
cytosolic fractions of stimulated and unstimulated cells.
Effect of PrPC Expression on Transition Metal
Toxicity--
As induction of PrPC expression did not
increase the incorporation of physiological levels of copper, we
decided to investigate whether PrPC-dependent
bound copper could play a role in the protection of cells toward copper
toxicity. Since it was suggested that manganese can compete with copper
for the binding sites (28), the resistance to this metal was also
tested. The data presented in Fig. 6,
A and B clearly show that cells overexpressing
PrPC (1000 dox) withstood higher copper (but not manganese
doses, Fig. 6B) than unstimulated (0 dox) cells or
stimulated RK13 cells. This resistance to copper was more related to
stimulated cells, with an LC50 of 540 and 341 µM
CuSO4 when they were compared with unstimulated cells.
Therefore PrP increases cellular resistance to copper but not to
manganese toxicity.
PrPC Overexpression Increases Resistance to Oxidative
Stress and Antioxidant Enzyme Activities--
Because it has been
suggested that one of the physiological functions of PrPC
could be in the protection of cells toward an oxidative stress (10), we
investigated both the resistance to an oxidative stress and the
activities or levels of the main antioxidant systems in cells
overexpressing PrPC. To study the relationship between
prion protein expression and resistance to oxidative stress, MTT
assays were performed following 3-morpholinosydnonimine
(SIN-1) treatments, which generates different free radicals:
O
We also evaluated the involvement of PrPC expression in the
cellular defense against oxidative stress by measuring different antioxidant activities such as SOD, GPX, GR, and glutathione levels. Induction of PrP increases total SOD (~21%), CuZn-SOD (~27%), GR
(~64%) activities, and GSH levels (~78%), while the activities of
GPX and mitochondrial Mn-SOD remain unchanged (see Table
I). Interestingly, Western blot detection
of CuZn-SOD in A74 cells indicates that the total level of this protein
was unchanged in stimulated and unstimulated cells (Fig.
8). This may reflect an increase in
CuZn-SOD activity of stimulated cells resulting from increased copper
incorporation into SOD or from SOD-like activity of PrP-copper
complexes. Finally to detect if PrP expression decreases oxidative
damage, lipid peroxidation was evaluated by measuring the formation of
TBARS as a stress biomarker. The basal level of oxidative damage
was significantly lower in stimulated cells as compared with
unstimulated and control cells. These data indicate that
PrPC expression increases resistance to basal as well as
induced oxidative stress by increasing cellular defenses.
In this study, we used a cellular model derived from a
heterologous epithelial cell line (RK13) in which the expression of murine PrPC was regulatable in a dose-dependent
manner by a doxycycline treatment. Actually most epithelial cell lines
we have tested, unlike, RK13, do express
PrP.2 The RK13 cells were
chosen because they express no detectable levels of endogenous PrP. The
risk that endogenous PrP could interfere with the function of
transfected PrP is therefore reduced. This may be a reason why we
succeeded in a previous study to infect RK13 cells transfected with
ovine PrP (18). It was then logical to generate a clone of cells
overexpressing murine PrP, which is used for cell biology and
transmission studies. RK13 cells are the only available cell lines
allowing a PrP expression from zero to high levels. In the present work
we used radioactive copper (64Cu) to study the effect of
PrPC expression on copper binding and uptake.
There is an increasing amount of data supporting a functional role for
PrPC in copper metabolism. First the N-terminal half of
PrPC contains five or six highly conserved octapeptide
tandem motifs of the general form PHGGGWGQ, which are capable of
binding copper ions with micromolar affinity (7, 8). Indeed,
PrPC isolated from hamster brain can bind a copper affinity
column (29). Second, copper content of membrane-enriched brain extract from PrP PrPC is normally attached to the cell membrane via a
phosphatidylinositol anchor (34). It has been shown that the enzyme PIPLC releases PrPC from the cells into media (35). Our
ex vivo experiments confirm the copper binding activity of
the PrPC protein, because we established a correlation
between copper binding and PrPC expression. This finding
was further confirmed when PrPC was cleaved with PIPLC
prior to 64Cu labeling. However, our work does not support
that PrPC could be involved in the copper transport across
the membrane, as suggested by studies reporting
histidine-dependent uptake of 67Cu proportional
to PrPC expression in cerebellar cells derived from three
lines of mice expressing various amounts of PrPC (36).
However, Pauly and Harris (37) have reported that copper stimulates
endocytosis of both mouse PrP and chicken PrP on the cell surface of
N2a mouse neuroblastoma cells via clathrin-coated pits. They suggested
that PrPC could serve as a recycling receptor for the
uptake of copper ions from the extracellular milieu. Also, it has been
shown that 100 µM copper resulted in the rapid
endocytosis of biotinylated murin PrPC expressed in human
neuroblastoma SH-SY5Y cells (38). In these two studies the minimum
concentration of CuSO4 required to produce an observable
increase in PrPC internalization was ~100
µM, which is 15-fold greater than the estimated
Kd for binding to synthetic PrP peptides and recombinant PrP (7, 9, 39). In our work we used very low levels of
copper, which we believe renders our results much closer to
physiological conditions. In any case when high concentrations of
copper (100 µM) are used in our cultures the
intracellular copper level increased up to 2-fold in both total and
soluble fractions in stimulated cells (see Table
II). So, only under high copper
concentrations PrPC expression increases copper uptake in
the cell. However, our results clearly demonstrated that at
physiological concentrations of copper, murine PrPC binds
copper at the outer side of the cell membrane but also indicates that
PrPC does not function as a copper transporter from the
extracellular medium to the cytoplasm. This supports the hypothesis
that PrPC may rather be an extracellular copper sensor
(33).
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-helical regions is
converted into a molecule with highly
-sheeted structures and
partial resistance to proteolytic digestion (2, 3). The conversion of
PrPC into PrPSc remains enigmatic. Biosynthesis
of PrPC is necessary for PrPSc formation, as
mice lacking PrPC are resistant to scrapie infection (4).
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
20 °C,
collected by centrifugation, and immunoblotted with P45
66 antibody
using ECL visualization.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
PrPC induction is
doxycycline-dependent. Dox was added at different
concentrations (0, 10, 25, 50, 100, 500, and 1000 ng/ml) to medium for
24 h, and PrPC expression was determined in A74 cells
by Western blot. The equivalent of 20 µg of protein (as determined
with the BCA protein assay kit) were loaded to a 12% polyacrylamide
gel, transferred onto polyvinylidene difluoride membrane, and
PrPC was detected with antibody P45-66 raised against the
N terminus of the protein. PrPC expression reaches a
maximum at 500 ng/ml of dox; after this concentration we have a
plateau. Specific murine PrPC bands were quantified by
densitometry and plotted as a percentage of maximum signal of
PrPC expression in A74 cells. Molecular mass markers are
indicated on the left in kDa.
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Fig. 2.
Time course of PrPC expression in
A74-stimulated cells. 500 ng/ml of dox was added at time 0, and
cells were harvested and lysed at different times (0, 1, 2, 4, 6, 8, 10, 24, and 48 h). 20 µg of protein were Western blotted and
analyzed with the same antibody P45-65. Expression of PrPC
can be detected 8 h after induction with a plateau at 24 h.
Specific murine PrPC bands were quantified by densitometry
and plotted as a percentage of maximum signal of PrPC
expression in A74 cells. Molecular mass markers are indicated on the
left in kDa.
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Fig. 3.
A74 cells expressed PrPC on the
cell membrane. A, fluorescence microscopy photography
of stimulated (500 ng/ml dox for 24 h) or unstimulated A74 cells.
Fluorescein-conjugated IgG were used as second antibody. B,
phase contrast microscopy for the same cells. 32% of stimulated cells
produced PrPC at the outer side of cell membrane.
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Fig. 4.
Relationship between PrPC
expression and 64Cu binding. A,
64Cu binding increases in proportion to PrPC
expression in A74 cells. 0.1 µg of 64Cu was added to A74
cell culture medium containing different concentrations of dox (0, 25, 100, and 500 ng/ml). Cells were incubated for 0.6, 2, 4, 8, 10, 24, and
26 h in radioactive media and harvested, and 64Cu
binding was measured. Copper binding was correlated to murine
PrPC expression. B, dox treatment did not change
copper binding in non-transfected control cells RK13. 0.1 µg of
64Cu was added to RK13 cell culture medium containing 0 or
500 ng/ml of dox. Cells were incubated for 0.75, 10, and 24 h in
radioactive media, harvested, and 64Cu binding was measured
and normalized to protein cell lysates. C, the experiments
described above were repeated, but cells were treated with 0.2 units/ml
PIPLC before measuring radioactivity in stimulated and unstimulated A74
cells (C) or in stimulated and unstimulated non-transfected
RK13 cells (D). Controls were at times (2.5, 10, 18, and
24 h). Copper binding was quantitated and normalized to the
protein of cell lysates. Data represent the mean of three
experiments ± S.D.
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Fig. 5.
Effect of PIPLC treatment on stimulated and
unstimulated A74 cells. A, after 24 h of
PrPC induction with 500 ng/ml dox in radioactive media,
cells were treated with 0.2 units/ml PIPLC for 2 h at 37 °C,
lysed in lysis buffer and loaded onto a 12% polyacrylamide gel,
transferred onto polyvinylidene difluoride membrane, and
PrPC was detected with antibody P45-66 raised against the N
terminus of the protein. B, proteins in culture were
precipitated with 4 volumes of methanol, and PrPC was
detected with P45-66 antibody. C, 24 h after
PrPC induction with or without 500 ng/ml of dox in a
radioactive medium containing 64Cu, cells were treated with
PIPLC (0.2 units/ml) for 2 h at 37 °C, and radioactivity was
measured. D, radioactivity was measured in the opti-MEM
medium of the same cells and normalized to protein cell lysates and
protein in the media. Results are expressed as the mean of three
experiments ± S.D. of radioactivity (µCi of 64Cu/mg
of protein). *, p < 0.01; #, p < 0.005.
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Fig. 6.
PrPC expression increases
resistance to copper but not to manganese toxicity. Cell lines
were incubated with the indicated concentration of copper
(A) or manganese (B) for 24 h, and viability
was then measured as described under "Experimental Procedures."
Results are expressed as mean percentage ± S.D. of viable cells,
assuming 100% viability for untreated A74 cells. *, p < 0.01; #, p < 0.005.
(46). Stimulated cells (500 ng/ml
dox) presented higher resistance (cell viability, 95%) to this
oxidative stress when compared with unstimulated cells (0 dox) (cell
viability, 59%) or control cells (RK13 0 or 500 dox) (Fig.
7A). In contrast, cells
overexpressing PrPC were surprisingly more susceptible to
hydrogen peroxide than unstimulated cells. Treatment for 3 h with
different concentrations of H2O2 induced a more
severe decrease in viability for doses exceeding 200 µM
(Fig. 7B) in stimulated cells as compared with unstimulated
cells. At 500 µM H2O2-stimulated
cells revealed ~50% lower viability than unstimulated cells. These
data demonstrate that PrPC expression decreases resistance
to peroxide toxicity.
View larger version (34K):
[in a new window]
Fig. 7.
PrPC expression increases
resistance to oxidative stress produced by SIN-1 but not to
H2O2 toxicity. Cell viability was
evaluated by a modified MTT assay as described under "Experimental
Procedures" in stimulated and unstimulated A74 and non-transfected
RK13 cells after 24 h of exposure to 1 mM SIN-1
(A) or after 3 h of exposure to different
concentrations of H2O2 (B). Results
are expressed as mean percentage ± S.D. of survival cell;
survival was calculated as the percentage of the staining values of
untreated cultures. *, p < 0.01; #, p < 0.005.
PrPc and antioxidant enzymes activities
View larger version (37K):
[in a new window]
Fig. 8.
PrPC expression did not change
CuZn-SOD protein level. Cell lysates of stimulated and
unstimulated A74 cells were prepared as in the legend to Fig. 1 and the
total amounts measured by using the BCA protein assay kit. Equal
amounts of proteins were Western blotted with an anti-CuZn-SOD sheep
polyclonal antibody. Molecular mass markers are indicated on the
left in kDa.
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
/
mice is 10-15-fold lower than in wild type
controls while no significant difference was observed for other metals
(8). Third, neuronal CuZn-SOD from PrP
/
mice showed
decreased activity linked to decreased copper incorporation by the
enzyme (10, 30). Neurons cultured from PrP
/
mice were
also more sensitive to oxidative stress, perhaps because of the
alteration of CuZn-SOD (10). However, the significance of copper
binding on PrPC functions or the role of PrPC
copper metabolism has yet to be clarified. Several hypotheses have been
proposed. Copper could have a role in the conformation of the protein
(31). PrPC could have a role in copper transport across the
cell membrane, and this could follow different processes. Copper needs
to be mobilized from its extracellular ligands (albumin and histidine), and Cu(II) is reduced to Cu(I) by unknown metalloreductases at the
membrane surface prior to its delivery across the plasma membrane by
the high affinity transporter CTR1 (32). These processes are still
unclear. It is also possible that copper may be transported by more
than one transport system, at least in some tissues. The cooperative
copper-binding mode of PrPC within the physiological
concentration range suggests a role in copper transport (33). So, the
contribution of PrPC, if any, in copper uptake by cells
could be a direct transport across the plasma membrane or a binding
step allowing either the mobilization of copper from its ligand or its
reduction prior to its effective transport by other systems.
Determination of cellular copper content under low and high copper
concentrations
Murine PrPC may serve as a copper chelating or buffering agent in the outer side of the cell membrane, and this may serve to protect cells against toxicity of free copper ions or a copper and reactive oxygen species-dependent cleavage of PrP into the octapeptide repeat region. This process may be related to the function of the molecule in the response to oxidative stress and suggests that the binding of copper is important for its processing (40).
The link between copper and PrPC may explain the mechanism
of neurodegeneration in prion diseases because copper and other transition metals play an important role in the neuropathology of
neurodegenerative disorders such as Parkinson's disease (PD), Alzheimer's disease (AD), and Amyotrophic lateral sclerosis (ALS) (41). Copper is an important component of various redox enzymes because
of its ability to readily adopt two ionic states Cu(I) and Cu(II). Free
copper is also a toxic ion, as exemplified by its ability to inactivate
proteins through tyrosine nitration, and both deficiency and excess
lead to disorders such as Menkes syndrome or Wilson's disease (42),
illustrating its physiological importance and duality in the central
nervous system. In the absence of copper chelating agent on the cell
surface, free copper could react with peroxides such as hydrogen
peroxide produced by superoxide dismutation or directly by many enzyme
catabolites such as monoamine oxidase, urate oxidase, glucose oxidase,
D-amino acid oxidase, and others to form the highly
reactive hydroxyl radical (·OH), which can initiate lipid
peroxidation as well as protein oxidation and cause apoptosis. It has
been shown that in the brain, highest concentrations of
PrPC are found at synapses, and copper binding by
PrPC in the synaptic cleft has a significant influence on
synaptic transmission (43). Changes in electrophysiological properties such as long-term potentiation (LTP), circadian rhythm between PrP/
and wild-type mice could be related to a disturbed
copper uptake in PrP
/
mice (43). Stimulated A74 cells
undergo high resistance to copper but not to manganese or cadmium
toxicity when compared with unstimulated or control cells. This
specific protection against copper toxicity may be due to the chelating
or buffering effect of murine PrPC on the cell surface.
Previously PC12, cells selected for resistance to copper toxicity and
oxidative stress showed high levels of PrPC (44). Primary
cerebellar granule culture derived from PrP knockout mice were
significantly more susceptible to H2O2 toxicity
than wild type; this toxicity was related to a significant decrease in
glutathione reductase activity (45) Moreover, increased oxidative damage to proteins and lipids was observed in the brain lysates from
Prnp
/
as compared with wild type mice of the same
genetic background (46, 47). As oxidative stress has been frequently
implicated in neurodegeneration it was very interesting to test the
influence of PrP expression in stimulated A74 on antioxidant enzymes
activities and resistance to oxidative stress. PrPC
induction in stimulated cells increases significantly CuZn-SOD, catalase, glutathione reductase activities, and glutathione levels in
cells. In addition stimulated cells were more resistant to oxidative
stress caused by SIN-1. This active metabolite of the vasodilatatory
drug molsidomine is frequently used as a model for the continuous
release of different free radicals: O
(48). The relationship between PrPC and oxidative stress
arose from results showing an alteration in cellular response to stress
with the decrease in PrPC expression or conversion to the
infectious form. PrP
/
mouse brains have reduced
CuZn-SOD, and cerebellar cells derived from these mice were more
sensitive to oxidative stress (10); increased levels of
PrPC were linked to increased levels of CuZn-SOD activity
because of an increase in copper incorporation (12). In our model, we believe that increased CuZn-SOD activity in stimulated cells is due to
SOD-like activity of PrP-Cu complexes in the outer side of the cell
membrane. Indeed, we detected no change in the protein levels of
CuZn-SOD in stimulated and unstimulated cells (Fig. 8), and we
demonstrated that copper stays at the outer side of the cell membrane.
This antioxidant function in the outer of the cell membrane is very
important, especially in neurons, to detoxify free radicals such as
O
), from reactions between NO· and
O
In conclusion, we have shown that expression of heterologous
PrPC (murine PrPC) in rabbit kidney cells
(RK13) increases copper binding but not uptake, and several antioxidant
enzymes activities confer high resistance to oxidative stress. These
findings are of major importance since oxidative stress is implicated
in several neurodegenerative diseases. We can hypothesize that in prion
diseases the conversion of PrPC to PrPSc
inhibited PrP copper binding. This inhibition could affect, the enzymatic activity of PrP-Cu complexes in the outer of the cell membrane and also, the regulation of the anti-oxidant system, which is
PrP-dependent in neurons. In the future, it will be
important to determine the influence of PrP conversion on copper
binding in a similar model permissive to prion replication (18) and the
influence of trace elements such as copper, zinc, or antioxidant on
prion diseases as a new therapeutic agent to re-equilibrate the
antioxidant deficiencies in these diseases.
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ACKNOWLEDGEMENTS |
---|
We thank David Harris (Washington University, St. Louis) for antibody P45-66, Dr. Josiane Arnaud for trace element measurement by atomic spectrophotometry, and P. Schaeffer for helpful technical assistance.
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FOOTNOTES |
---|
* This work was supported by the European Community QRT-2000-02353.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
§ Supported by le centre evian pour l'eau. To whom correspondence should be addressed: Laboratoire de Biologie du Stress Oxydant (LBSO), Faculté de Pharmacie, Domaine de la Merci, 38706 la Tronche, France. Tel.: 33-4-76-63-74-56; Fax: 33-4-76-63-74-85; E-mail: walid. rachidi{at}ujf-grenoble.fr.
Published, JBC Papers in Press, December 23, 2002, DOI 10.1074/jbc.M211830200
2 F. Archer and H. Laude, unpublished data.
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ABBREVIATIONS |
---|
The abbreviations used are: PrPC, cellular isoform of prion protein; SOD, superoxide dismutase; dox, doxycycline; PIPLC, phosphoinositol phospholipase C; ROS, reactive oxygen species; GPX, glutathione peroxidase; GR, glutathione reductase; MDA, malondialdehyde acid; SIN-1, 3-morpholinosydnonimine; PrPSc, scrapie isoform of prion protein; MTT, (4,5-dimethyl-thiazol-2-yl)-2,5-diphenyl-tetrazolium bromide; TBARS, thiobarbituric acid reactants.
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REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Parchi, P., and Gambetti, P. (1995) Curr. Opin. Neurol. 8, 286-293[Medline] [Order article via Infotrieve] |
2. |
Prusiner, S. B.
(1998)
Proc. Natl. Acad. Sci. U. S. A.
95,
13363-13383 |
3. | Horiuchi, M., and Caughey, B. (1999) Structure Fold Des. 7, R231-40[Medline] [Order article via Infotrieve] |
4. | Bueler, H., Aguzzi, A., Sailer, A., Greiner, R. A., Autenried, P., Aguet, M., and Weissmann, C. (1993) Cell 73, 1339-1347[Medline] [Order article via Infotrieve] |
5. | Kretzschmar, H. A., Stowring, L. E., Westaway, D., Stubblebine, W. H., Prusiner, S. B., and Dearmond, S. J. (1986) Dna 5, 315-324[Medline] [Order article via Infotrieve] |
6. | Kretzschmar, H. A., Prusiner, S. B., Stowring, L. E., and DeArmond, S. J. (1986) Am. J. Pathol. 122, 1-5[Abstract] |
7. | Hornshaw, M. P., McDermott, J. R., and Candy, J. M. (1995) Biochem. Biophys. Res. Commun. 207, 621-629[CrossRef][Medline] [Order article via Infotrieve] |
8. | Brown, D. R., Qin, K., Herms, J. W., Madlung, A., Manson, J., Strome, R., Fraser, P. E., Kruck, T., von Bohlen, A., Schulz-Schaeffer, W., Giese, A., Westaway, D., and Kretzschmar, H. (1997) Nature 390, 684-687[CrossRef][Medline] [Order article via Infotrieve] |
9. | Stockel, J., Safar, J., Wallace, A. C., Cohen, F. E., and Prusiner, S. B. (1998) Biochemistry 37, 7185-7193[CrossRef][Medline] [Order article via Infotrieve] |
10. | Brown, D. R., Schulz-Schaeffer, W. J., Schmidt, B., and Kretzschmar, H. A. (1997) Exp. Neurol. 146, 104-112[CrossRef][Medline] [Order article via Infotrieve] |
11. |
Waggoner, D. J.,
Drisaldi, B.,
Bartnikas, T. B.,
Casareno, R. L.,
Prohaska, J. R.,
Gitlin, J. D.,
and Harris, D. A.
(2000)
J. Biol. Chem.
275,
7455-7458 |
12. | Brown, D. R., and Besinger, A. (1998) Biochem. J. 334, 423-429[Medline] [Order article via Infotrieve] |
13. | Pattison, I. H., and Jebbett, J. N. (1971) Nature 230, 115-117[Medline] [Order article via Infotrieve] |
14. | Purdey, M. (2000) Med. Hypotheses 54, 278-306[CrossRef][Medline] [Order article via Infotrieve] |
15. |
Milhavet, O.,
McMahon, H. E.,
Rachidi, W.,
Nishida, N.,
Katamine, S.,
Mange, A.,
Arlotto, M.,
Casanova, D.,
Riondel, J.,
Favier, A.,
and Lehmann, S.
(2000)
Proc. Natl. Acad. Sci. U. S. A.
97,
13937-13942 |
16. | Lee, D. W., Sohn, H. O., Lim, H. B., Lee, Y. G., Kim, Y. S., Carp, R. I., and Wisniewski, H. M. (1999) Free Radic. Res. 30, 499-507[Medline] [Order article via Infotrieve] |
17. | Choi, S. I., Ju, W. K., Choi, E. K., Kim, J., Lea, H. Z., Carp, R. I., Wisniewski, H. M., and Kim, Y. S. (1998) Acta Neuropathol. (Berl.) 96, 279-286[CrossRef][Medline] [Order article via Infotrieve] |
18. |
Vilette, D.,
Andreoletti, O.,
Archer, F.,
Madelaine, M. F.,
Vilotte, J. L.,
Lehmann, S.,
and Laude, H.
(2001)
Proc. Natl. Acad. Sci. U. S. A.
98,
4055-4059 |
19. | Christofinis, G. J., and Beale, A. J. (1968) J. Pathol. Bacteriol. 95, 377-381[Medline] [Order article via Infotrieve] |
20. |
Lehmann, S.,
and Harris, D. A.
(1995)
J. Biol. Chem.
270,
24589-24597 |
21. | Hansen, M. B., Nielsen, S. E., and Berg, K. (1989) J. Immunol. Methods 119, 203-210[CrossRef][Medline] [Order article via Infotrieve] |
22. |
Richard, M. J.,
Portal, B.,
Meo, J.,
Coudray, C.,
Hadjian, A.,
and Favier, A.
(1992)
Clin. Chem.
38,
704-709 |
23. | Marklund, S., and Marklund, G. (1974) Eur. J. Biochem. 47, 469-474[Medline] [Order article via Infotrieve] |
24. | Akerboom, T. P., and Sies, H. (1981) Methods Enzymol. 77, 373-382[Medline] [Order article via Infotrieve] |
25. | Gunzler, W. A., Kremers, H., and Flohe, L. (1974) Z. Klin. Chem. Klin. Biochem. 12, 444-448[Medline] [Order article via Infotrieve] |
26. | Spooner, R. J., Delides, A., and Goldberg, D. M. (1981) Biochem. Med. 26, 238-248 |
27. | Gossen, M., and Bujard, H. (1992) Proc. Natl. Acad. Sci. U. S. A. 89, 5547-5551[Abstract] |
28. |
Brown, D. R.,
Hafiz, F.,
Glasssmith, L. L.,
Wong, B. S.,
Jones, I. M.,
Clive, C.,
and Haswell, S. J.
(2000)
EMBO J.
19,
1180-1186 |
29. |
Pan, K. M.,
Stahl, N.,
and Prusiner, S. B.
(1992)
Protein Sci.
1,
1343-1352 |
30. | Brown, D. R., Wong, B. S., Hafiz, F., Clive, C., Haswell, S. J., and Jones, I. M. (1999) Biochem. J. 344, 1-5[CrossRef][Medline] [Order article via Infotrieve] |
31. | Wadsworth, J. D., Hill, A. F., Joiner, S., Jackson, G. S., Clarke, A. R., and Collinge, J. (1999) Nat. Cell Biol. 1, 55-59[CrossRef][Medline] [Order article via Infotrieve] |
32. |
Lee, J.,
Pena, M. M.,
Nose, Y.,
and Thiele, D. J.
(2002)
J. Biol. Chem.
277,
4380-4387 |
33. | Kramer, M. L., Kratzin, H. D., Schmidt, B., Romer, A., Windl, O., Liemann, S., Hornemann, S., and Kretzschmar, H. (2001) J. Biol. Chem. 27, 27 |
34. | Stahl, N., Borchelt, D. R., Hsiao, K., and Prusiner, S. B. (1987) Cell 51, 229-240[Medline] [Order article via Infotrieve] |
35. | Stahl, N., Borchelt, D. R., and Prusiner, S. B. (1990) Biochemistry 29, 5405-5412[Medline] [Order article via Infotrieve] |
36. | Brown, D. R. (1999) J. Neurosci. Res. 58, 717-725[CrossRef][Medline] [Order article via Infotrieve] |
37. |
Pauly, P. C.,
and Harris, D. A.
(1998)
J. Biol. Chem.
273,
33107-33110 |
38. | Perera, W. S., and Hooper, N. M. (2001) Curr. Biol. 11, 519-523[CrossRef][Medline] [Order article via Infotrieve] |
39. | Hornshaw, M. P., McDermott, J. R., Candy, J. M., and Lakey, J. H. (1995) Biochem. Biophys. Res. Commun. 214, 993-999[CrossRef][Medline] [Order article via Infotrieve] |
40. |
McMahon, H. E.,
Mange, A.,
Nishida, N.,
Creminon, C.,
Casanova, D.,
and Lehmann, S.
(2001)
J. Biol. Chem.
276,
2286-2291 |
41. | Sayre, L. M., Perry, G., and Smith, M. A. (1999) Curr. Opin. Chem. Biol. 3, 220-225[CrossRef][Medline] [Order article via Infotrieve] |
42. | Mercer, J. F. (2001) Trends Mol. Med. 7, 64-69[CrossRef][Medline] [Order article via Infotrieve] |
43. | Kretzschmar, H. A., Tings, T., Madlung, A., Giese, A., and Herms, J. (2000) Arch. Virol. Suppl. 16, 239-249[Medline] [Order article via Infotrieve] |
44. | Brown, D. R., Schmidt, B., and Kretzschmar, H. A. (1997) Int. J. Dev. Neurosci. 15, 961-972[CrossRef][Medline] [Order article via Infotrieve] |
45. |
White, A. R.,
Collins, S. J.,
Maher, F.,
Jobling, M. F.,
Stewart, L. R.,
Thyer, J. M.,
Beyreuther, K.,
Masters, C. L.,
and Cappai, R.
(1999)
Am. J. Pathol.
155,
1723-1730 |
46. | Wong, B. S., Liu, T., Li, R., Pan, T., Petersen, R. B., Smith, M. A., Gambetti, P., Perry, G., Manson, J. C., Brown, D. R., and Sy, M. S. (2001) J. Neurochem. 76, 565-572[CrossRef][Medline] [Order article via Infotrieve] |
47. | bio Klamt, F., Dal-Pizzol, F., Conte da Frota, M. L., Walz, R., Andrades, M. E., da Silva, E. G., Brentani, R. R., n Izquierdo, I., and Fonseca Moreira, J. C. (2001) Free Radic. Biol. Med. 30, 1137-1144[CrossRef][Medline] [Order article via Infotrieve] |
48. |
Gergel, D.,
Misik, V.,
Ondrias, K.,
and Cederbaum, A. I.
(1995)
J. Biol. Chem.
270,
20922-20929 |
49. |
Radi, R.,
Beckman, J. S.,
Bush, K. M.,
and Freeman, B. A.
(1991)
J. Biol. Chem.
266,
4244-4250 |
50. | Radi, R., Beckman, J. S., Bush, K. M., and Freeman, B. A. (1991) Arch Biochem. Biophys. 288, 481-487[Medline] [Order article via Infotrieve] |
51. | Beckman, J. S., Ischiropoulos, H., Zhu, L., van der Woerd, M., Smith, C., Chen, J., Harrison, J., Martin, J. C., and Tsai, M. (1992) Arch. Biochem. Biophys. 298, 438-445[Medline] [Order article via Infotrieve] |
52. |
Brito, C.,
Naviliat, M.,
Tiscornia, A. C.,
Vuillier, F.,
Gualco, G.,
Dighiero, G.,
Radi, R.,
and Cayota, A. M.
(1999)
J. Immunol.
162,
3356-3366 |
53. |
Mouillet-Richard, S.,
Ermonval, M.,
Chebassier, C.,
Laplanche, J. L.,
Lehmann, S.,
Launay, J. M.,
and Kellermann, O.
(2000)
Science
289,
1925-1928 |
54. | Wong, B. S., Brown, D. R., Pan, T., Whiteman, M., Liu, T., Bu, X., Li, R., Gambetti, P., Olesik, J., Rubenstein, R., and Sy, M. S. (2001) J. Neurochem. 79, 689-698[CrossRef][Medline] [Order article via Infotrieve] |