Vitamin E blocks early events induced by

1-methyl-4-phenylpyridinium (MPP+) in cerebellar granule cells

Rosa A. Gonzalez-Polo,* German Soler,* Alberto Alvarez,� Isabel Fabregat� and Jose M. Fuentes§

*Departamento de Bioquımica y Biologıa Molecular y Genetica, Facultad de Veterinaria, Universidad de Extremadura, Caceres, Spain

�Centro de Citometrıa de Flujo y Microscopıa Confocal. Universidad Complutense de Madrid, Madrid, Spain

�Departamento de Bioquımica y Biologıa Molecular, Facultad de Farmacia (Centro Mixto CSIC/UCM), Universidad Complutense de

Madrid, Madrid, Spain

§Departamento de Bioquımica y Biologıa Molecular y Genetica, EU Enfermerıa y TO, Universidad de Extremadura, Caceres, Spain

Abstract

Exposure of cerebellar granule cells (CGCs) to 1-methyl-4-

phenylpyridinium (MPP+) results in apoptotic cell death, which

is markedly attenuated by co-treatment of CGCs with the

radical scavenger vitamin E. Analysis of free radical produc-

tion and mitochondrial transmembrane potential (DYm), using

specific fluorescent probes, showed that MPP+ mediates early

radical oxygen species (ROS) production without a loss of

DYm. Exposure to MPP+ also produces an early increase in

Bad dephosphorylation and translocation of Bax to the mito-

chondria. These events are accompanied by cytochrome c

release from mitochondria to cytosol, which is followed by

caspase 3 activation. Exposure of the neurons to vitamin E

maintains Bad phosphorylation and attenuates Bax translo-

cation, inhibiting cytochrome c release and caspase activa-

tion. MPP+-mediated cytochrome c release is also prevented

by allopurinol, suggesting the participation of xanthine oxidase

in the process. Our results indicate that free radicals play an

active role in the MPP+-induced early events that culminate

with cell death.

Keywords: apoptosis, cerebellar granule cells, cytochrome c,

1-methyl-4-phenylpyridinium (MPP+), vitamin E.

J. Neurochem. (2003) 84, 305–315.

1-Methyl-4-phenylpyridinium (MPP+) is a well-known neu-

rotoxin (Langston et al. 1983;Blum et al. 1993),which causes

apoptotic cell death at concentrations ranging from 40 to

300 lM (Du et al. 1997). MPP+ incorporation into the cells isproduced essentially through the dopamine (Chiba et al. 1985;

Gonzalez-Polo et al. 2001) or the cation amino acid

(Gonzalez-Polo et al. 2001) transporters. The principal target

ofMPP+ is themitochondria, where it inhibits Complex I in the

mitochondrial respiratory chain, with the consequent cessation

of oxidative phosphorylation (Tipton and Singer 1993).

Several studies have shown the involvement of radical

oxygen species (ROS) in MPP+-induced neurotoxicity

(Castagnoli et al. 1985; Rossetti et al. 1988; Kitamura et al.

1998). The importance of MPP+-mediated ROS production is

demonstrated in vivo by the fact that transgenic mice that

overexpress copper/zinc superoxide dismutase are signifi-

cantly more resistant to MPP+-induced toxicity than wild-

type mice (Przedborski et al. 1992). In vitro MPP+-induced

cell death is abolished by co-treatment with radical scav-

engers, such as ascorbic acid (Akaneya et al. 1995),

a-tocopherol (Odunze et al. 1990) and others (Akaneya

et al. 1995) in several cell types. Free radical production in

MPP+-exposure has been classically associated with inhibi-

tion of electron transport in mitochondria (Nicklas et al.

1985; Mizuno et al. 1989). However, in other models of

programmed cell death cytosolic free radicals have been

shown to play active roles in the generation of apoptosis,

NADPH-oxidase (Tammariello et al. 2000) or xanthine

oxidase (Atlante et al. 2001) being involved in its

Received June 4, 2002; revised manuscript received October 2, 2002;

accepted October 8, 2002.

Address correspondence and reprint requests to Jose M. Fuentes,

Departamento de Bioquımica y Biologıa Molecular, EU Enfermerıa y

TO Universidad de Extremadura, Avenida de la Universidad s/n 10071,

Caceres, Spain. E-mail: jfuentes@unex.es

Abbreviations used: CGC, cerebellar granule cells; CMXRos, chlo-

romethyl-X-rosamine; DFCH, 2¢-7¢-dichlorodihydrofluorescein; DMEM,Dulbecco’s modified Eagle’s medium; HE, hydroethidine; MCB, mon-

ochlorobimane; MPP+, 1-methyl-4-phenylpyridinium; MTT, 3-(4,5-

dimehylthiazol-2-yl)-2,5-diphenyltetrazolium bromide; PBS, phosphate-

buffered saline; PI, propidium iodide; PVDF, polyvinylidene difluoride;

ROS, radical oxygen species; SDS, sodium dodecyl sulphate; DYm:mitochondrial transmembrane potential.

Journal of Neurochemistry, 2003, 84, 305–315

� 2003 International Society for Neurochemistry, Journal of Neurochemistry, 84, 305–315 305

production. Interestingly, MPP+ has been described as a

substrate for xanthine oxidase and it may lead to ROS

formation (Klaidman et al. 1993). In any case, free radical

production has normally been measured over a relatively

long period (about 1 h after beginning MPP+ or other toxic

treatment). Recently, a new and exciting line of evidence has

emerged reporting, in glutamate-exposed and potassium-

deprived cerebellar granule cells (CGCs), early (before

30 min apoptotic injury) free radicals production, being

these free radicals responsible for cytochrome c release

(Atlante et al. 2000; Valencia and Moran 2001), as a cell

defense system against oxidative stress (Atlante et al. 1999,

2000). However, taking into account that cytochrome c

release has been related to apoptosis in several apoptotic

injuries, including MPP+-treatment (Du et al. 1997; Dodel

et al. 1998; Leist et al. 1998), this release could also activate

the apoptotic machinery, such as caspase activation, DNA

fragmentation and cell death.

The release of cytochrome c is associated with the

translocation to the mitochondria of certain bcl-2-family

proteins, such as Bax or Bad (Desagher and Martinou 2000).

Bax may play a central role in mediating mitochondria-

dependent apoptosis in neurons (Putcha et al. 1999).

Following a death signal, Bax can be translocated from

cytosol to mitochondria, which is rapidly followed by

cytochrome c release. The Bax effect can be prevented by

the presence of Bcl-2 or Bcl-xL proteins in the mitochondria

(Gross et al. 1999). Bad is another member of the pro-

apoptotic Bcl-2 proteins, which is dephosphorylated during

the apoptosis process. Phosphorylated Bad is normally

sequestered in the cytosol through its binding to 14-3-3

protein. Dephosphorylation of Bad by phosphatases of the

PP1 or PP2 families promotes its translocation to mitochon-

dria, where it binds to Bcl-xL, inhibiting its death-repressor

activity (Zha et al. 1996; Desagher and Martinou 2000).

In this paper we describe how vitamin E, a biological anti-

oxidant that is involved in neuroprotection from several

insults, blocks all the early events observed in CGCs after

MPP+-exposure. We propose for the first time that free

radicals, produced through the xanthine oxidase system, play

an active role in the MPP+-induced early events that

culminate with cell death.

Materials and methods

Materials

MPP+ was obtained from Research Biochemicals Inc. (Natick, MA,

USA). Cytosine arabinoside, Dulbecco’s modified Eagle’s medium

(DMEM), fetal calf serum and all the medium and supplements,

3-(4,5-dimehylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT),

poly D-lysine, a-tocopherol (vitamin E), xanthine and xanthine

oxidase, allopurinol and protease inhibitors were obtained from

Sigma (St Louis, MO, USA). Okadaic acid was supplied by Tocris

(London, UK). The fluorescent probes hydroethidine (HE), 2¢-7¢-dichlorodihydrofluorescein diacetate (DFCH-DA) and chloromethyl-

X-rosamine (CMXRos) were from Molecular Probes (Eugene, OR,

USA). Monochlorobimane was supplied by Calbiochem (San

Diego, CA, USA) and caspase 3 substrate (Ac-DEVD-AMC) was

obtained from Pharmingen (San Diego, CA, USA). Anti-Bax

(SC7480) and anti-cytochrome c (SC 7119), anti-actin (SC7210)

and secondary mouse (SC 2005) antibodies were all from Santa

Cruz Biotechnologies (Santa Cruz, CA, USA). Anti-Ser136-phos-

pho-Bad, anti-Ser473-phospho-Akt and cleaved caspase 3 antibod-

ies were obtained from Cell Signalling Technology (Beverly, MA,

USA). The rabbit secondary antibody and electrophoresis reagents

were from Bio-Rad Laboratories (Hercules, CA, USA). ECL Plus

and all the western blot reagents were purchased from Amersham

Biosciences (Buckinghamshire, UK). Other chemicals were of

analytical grade.

Cerebellar granule cell culture

Primary cultures of CGCs were obtained from 7- to 8-day-old

Wistar rats of either sex as described before (Gonzalez-Polo et al.

2001). The animals were housed in a temperature-controlled room

maintained at 12 h light/dark cycles. The standard laboratory animal

food and tap water were available ad libitum for the mothers. The

experimental protocols of this study were approved by the Research

Committee of the University of Extremadura (in accordance with the

National Institutes of Health guidelines) and were designed to

minimize the pain or discomfort of the animals. Briefly, cerebella

dissected free of meninges were chopped into small pieces and

digested with trypsin (2.5 mg/mL, 10 min at 37�C) in a Krebs–Ringer buffer solution, pH 7.4, containing bovine serum albumin

(3 mg/mL). After addition of soybean trypsin inhibitor (0.5 mg/mL)

and DNAse (0.1 mg/mL), the tissue was disrupted by 10 passages

through a fine tip plastic transfer pipette. The resulting cell

suspension was filtered through a 100-lm nylon cloth, centrifuged

and resuspended in DMEM supplemented with 10% fetal calf

serum, 25 mM KCl, 2 mM glutamine, penicillin (50 units/mL) and

streptomycin (50 lg/mL). Then cells were seeded at a density of5 · 105 cells/mL in poly L-lysine-pretreated 24-well or six-well

plates in a humidified atmosphere of 5% CO2 at 37�C. Cytosinearabinoside (10 lM) was added 24 h after plating to arrest thegrowth of non-neuronal cells, mostly astrocytes and microglia.

All the experiments were carried out at 7 days in culture in a

DMEM without fetal calf serum supplemented with 25 mM KCl,

2 mM glutamine, 50 U/mL penicillin, 50 lg/mL streptomycin,

0.1 mg/mL sodium pyruvate, 20 nM progesterone and 5 lg/mLinsulin.

Drug treatment protocol

Cells were exposed to MPP+ (50 lM) and cell viability was

measured 24 h after treatment using the MTT assay (Mosmann

1983). Absorbance at 500 nm in control cultures was used as 100%

viability, typically 0.25 ± 0.005.

DNA fragmentation

After removal of the culture medium, cells grown in six-well plates

were detached with 0.5 mL of 5 mM Tris, 20 mM EDTA pH 7.4 and

transferred to assay tubes. Then, 25 lL Triton X-100 (10% v/v) was

added, and the cells were incubatedwith gentle shaking for 1 h at 4�C.

306 R. A. Gonzalez-Polo et al.

� 2003 International Society for Neurochemistry, Journal of Neurochemistry, 84, 305–315

Nuclei were removed by centrifugation at 15 000 g for 15 min.

The supernatants containing cytosolic DNA were treated with 0.5%

sodium dodecyl sulphate (SDS) and 0.1 mg/mL proteinase K for 3 h

at room temperature. After two phenol–chloroform extractions, the

water-soluble fractions were incubated for 1 h at 37�C with RNAse(15 lg/mL). After two further extractions with phenol–chloroform,DNA was precipitated by adding 0.1 volumes of 2.5 M sodium

acetate, pH 5.2, and three volumes of ice-cold ethanol. DNA pellets

were washed with 70% ethanol, air-dried, and dissolved in 10 mM

Tris, pH 8.0, containing 1 mM EDTA, prior to electrophoresis in

1.5% agarose gels.

Propidium iodide staining

For confocal microscopy visualization of chromatin condensation

and nuclei degradation propidium iodide (PI) was used. After

incubation in the absence or in the presence of 50 lM MPP+, cellswere washed with phosphate-buffered saline (PBS) and incubated

with 0.005% PI in PBS during 10 min. Fluorescence images were

obtained using a MRC-1024 laser confocal microscopy (Bio-Rad,

Hemel Hempstead, UK).

ROS measurement

The oxidation-sensitive fluorescent probes HE and DFCH-DA were

used to analyse the net intracellular generation of ROS by flow

cytometry (Rothe and Valet 1990). CGCs were incubated in the

presence or in the absence of 50 lM MPP+ at different times, cellswere collected in the presence of cold PBS (without calcium and

magnesium salts) with EDTA 1 mM, pH 7.4. Then cells were

centrifuged and diluted in 20 lM HE or 5 lM DFCH-DA in PBS andincubated for 30 min at 37�C. Cells were run in a FACScan flowcytometer (Becton & Dickinson, San Jose, CA, USA) acquiring

10 000 cells per sample.

Analysis of mitochondrial transmembrane potential

The fluorescent probe CMXRos was used to analyse the mito-

chondrial transmembrane potential (DYm) by flow cytometry. CGCswere incubated in the presence or in the absence of 50 lM MPP+.Cells were collected in the presence of cold PBS (without calcium

and magnesium salts) with EDTA 1 mM, pH 7.4. Then cells were

centrifuged and diluted in 0.1 lM CMXRos in PBS for 30 min at

37�C. Cells were run in a FACScan flow cytometer (BD, San Jose,CA) acquiring 10 000 cells per sample.

GSH determination

The glutathion-dependent fluorescent probe monochlorobimane

(MCB) was used to analyse the net intracellular content of GSH

by flow cytometry (Kamencic et al. 2000). CGCs were incubated in

the presence or in the absence of 50 lM MPP+. Cells were collectedin the presence of cold PBS (without calcium and magnesium salts),

with EDTA 1 mM, pH 7.4. Then cells were centrifuged and diluted

in 1 lM MCB in PBS and incubated for 30 min at 37�C. UV laserfrom a LSR flow cytometer (BD) was used to excite MCB, blue

fluorescence was recovered through a 440/40 BP filter and 10 000

cells were acquired per sample.

Caspase 3 activity

Cells were scraped off in PBS, collected by centrifugation and lysed

at 4�C in 5 mM Tris–HCl, pH 8.0, 20 mM EDTA, 0.5% Triton

X-100. Lysates were clarified by centrifugation at 13 000 g for

10 min. The reaction mixture contained 25 lM cellular lysates,

325 lM assay buffer (20 mM HEPES pH 7.5, 10% glycerol, 2 mM

dithiothreitol), and 20 lM caspase 3 substrate (Ac-DEVD-AMC).

After 2 h incubation in the dark, enzymatic activity was measured in

a Luminescence Spectrophotometer (Perkin Elmer LS-50) (kexci-tation, 380 nm; kemission, 440 nm). Caspase activity was repre-sented as percentage over untreated cells.

Western blot analysis

To detect cytochrome c and Bax proteins in mitochondria, attached

cells were scraped off in a buffer containing 25 mM Tris–HCl

pH 6.8, 250 mM sucrose, 1 mM EDTA, 0.05% digitonine, 1 mM

dithiothreitol, 0.1 mM phenylmethylsulfonyl fluoride, 1 lg/mLleupeptine, 1 lg/mL pepstatin and 1 lg/mL aprotinin according tothe method previously described (Pique et al. 2000). Samples were

centrifuged at 13 000 g for 3 min at 4�C, pellets containing themitochondrial fraction were denatured in a lysis buffer and

processed as described below. For phospho-Bad, phospho-Akt,

cleaved caspase 3 and total Bax proteins in total cell lysates, cells

were washed with cold PBS at 4�C and lysed in a lysis buffer

containing 50 mM Tris–HCl pH 6.8, glycerol 10%, 2% SDS, 10 mM

dithiothreitol and 0.005% blue brominphenol. In both, mitochon-

drial and total lysates proteins (30–50 lg/condition) were resolvedin 12% SDS gel electrophoresis and transferred to polyvinylidene

difluoride (PVDF) membranes according to the conventional

methods partially modified by Fuentes et al. (2000). Briefly,

proteins were transferred (250 mA for 60 min) to PVDF membranes

using a Mini Trans-Blot Cell apparatus (Bio-Rad). The procedure

for immunodetection, including transfer, blocking of the membrane

(30 min at 37�C) with TTBS (10 mM Tris–HCl pH 7.5, 150 mM

NaCl and 0.2% Tween-20) containing 10% non-fat dried milk and

incubation (60 min at room temperature) with the primary antibody

(diluted 1 : 1000 in TTBS + 5% non-fat dried milk). After washing

(twice for 5 min each time with TTBS), membranes were incubated

(60 min at room temperature) with peroxidase-conjugated secon-

dary antibodies (1 : 5000 in TTBS with 5% non-fat dried milk).

After washing (twice for 5 min and once for 10 min), detection of

bound antibodies was visualized by chemiluminiscence using the

ECL-plus reagent.

Other methods

Protein concentration was measured according Bradford (1976) for

mitochondrial lysates and Lowry et al. (1951) for total cell lysates

using bovine serum albumin as standard in both. All data were

representative of at least three independent neuronal preparations

(with comparable results) each one in triplicate. Statistical analysis

were performed using the Student’s t-test (*p < 0.05).

Results

As previously reported (Du et al. 1997; Gonzalez-Polo et al.

2001) CGCs are sensitive to the toxic properties of MPP+.

Treatment with MPP+ caused a time- and concentration-

dependent increase in cell death and DNA fragmentation. In

our work, CGCs treated with 50 lM MPP+ for 24 h exhibitedmarked cell death (50% approximately, Fig. 1a) and DNA

Vitamin E blocks early cytochrome c release 307

� 2003 International Society for Neurochemistry, Journal of Neurochemistry, 84, 305–315

fragmentation (Fig. 1c), this biochemical change being char-

acteristic of apoptosis. In contrast, untreated control cells

showed noDNA laddering (Fig. 1c). Similarly, apoptotic nuclei

could be detected in MPP+-treated cells following PI staining

and confocal microscopy visualization (data not shown).

We next investigated the effects of a-tocopherol(vitamin E) on the viability of MPP+-treated CGCs. Cultured

CGCs were exposed to MPP+ (50 lM) with and withoutvitamin E for 24 h and subsequently examined for cell

viability. MPP+-induced neurotoxicity was attenuated in the

presence of vitamin E (Fig. 1b). The attenuation of MPP+-

induced neurotoxicity by vitamin E was concentration

dependent (Fig. 1b) with a maximum effect in the range

10–100 lM. Also, biochemical features of apoptosis, such asinternucleosomal DNA fragmentation, were also markedly

attenuated in the presence of vitamin E (Fig. 1c). To exclude

any possible interaction between MPP+ and vitamin E that

could justify the absence of a MPP+ effect, we pre-incubated

cells with vitamin E at different times between 0 and 2 h

before MPP+ addition. Protective effects of vitamin E were

not modified under any of these conditions (data not shown).

As previously described, MPP+ is able to induce ROS

production in several cell cultures (Sinha et al. 1986; Obata

1999), including primary cultures of CGCs. In this study we

could observe (Figs 2a and b) an early ROS production

(30 min after MPP+-exposure) induced byMPP+ and detected

by the oxidation-sensitive fluorescent probes HE and DFCH-

DA. Oxidative stress and the generation of free radicals

induced by MPP+ in CGCs also produced depletion in GSH

intracellular content. As shown in Fig. 2(c) after 30 min of

MPP+ treatment, and coincident with the ROS production, a

significant (around 50%) decrease in glutathione levels was

observed. As expected, both ROS production and GSH

depletion were impaired by vitamin E (results not shown).

Recently Bobba et al. (1999) have proposed an early release

of cytochrome c in glutamate-exposed CGCs, this release from

mitochondria being related to an early ROS production (Atlante

et al. 2000).WhetherMPP+ orMPTP can induce a similar early

release of cytochrome c is still unknown. To address this

question, we assayed the distribution of cytochrome c in CGCs

incubated with 50 lM MPP+. Cytochrome c decreases in themitochondrial fraction in a time- and dose-dependent manner

(Figs 3a and c, respectively) with a maximum after 30 min

exposure to MPP+ (Fig. 3a). Cytochrome c release preceded

caspase 3 activation. A maximum of caspase 3 activation was

evident after 60 min of MPP+ incubation, and by western blot

detection of 17-kDa fragment (active form) (Fig. 3e). As we

expected, both early release of cytochrome c and caspase 3

activationwere completely abolished by vitamin E co-treatment

(Figs 3b and f, respectively).

Because MPP+ was previously described as a substrate for

xanthine oxidase (Klaidman et al. 1993), we next determined

the implication of this enzyme in the early MPP+ induced

apoptosis. The xanthine oxidase inhibitor allopurinol (at a

100

80

60

400

100

80

60

400

0.1 1 10 100

0 6 12 24Time (h)

Via

bilit

y (%

con

trol

)

Via

bilit

y (%

con

trol

)

(a)

(b)

1 2 3 4

(c)

MPP+50 µM

MPP+100 µM

[vit E] µM

Fig. 1 Protection by vitamin E of the apoptosis induced by MPP+ in

primary cultures of cerebellar granule cells. (a) Time-dependence for

MPP+-induced cell death. CGCs were exposed to 50 lM (d) or 100 lM

(s) of MPP+ for 24 h. Cell viability was monitored with the MTT

reduction assay as described in the Materials and methods. Data are

mean ± SEM values of triplicate determination of three independent

experiments. (b) Internucleosomal DNA-fragmentation induced in

CGCs following exposure to MPP+ and its prevention by vitamin

E. CGCs (control, lane 2) were exposed to MPP+ (50 lM, 24 h) in the

absence (lane 3) or in the presence of 100 lM vitamin E (lane 4). DNA

marker is shows in lane 1. (c) Concentration–response relationship for

the protection elicited by vitamin E against MPP+-induced cell death.

Cells were treated with the indicated vitamin E concentrations at the

same time as 50 lM MPP+. The solid bar represents untreated cells

and the open bar represents cells treated with 50 lM MPP+ for 30 min.

Cell viability was measured after 24 h of MPP+ treatment. Data are

mean ± SEM values of three experiments in triplicate.

308 R. A. Gonzalez-Polo et al.

� 2003 International Society for Neurochemistry, Journal of Neurochemistry, 84, 305–315

concentration of 100 lM) was assessed by its ability to inhibitapoptotic events in MPP+. This concentration of allopurinol

impaired the mitochondrial cytochrome c release observed at

30 min with 50 lM MPP+ (Fig. 4a). Interestingly, artificial

ROS generation with the xanthine plus xanthine oxidase

system also stimulated early (30 min) cytochrome c release in

CGCs, as detected by western blot analysis (Fig. 4b). These

results are in agreement with those shown in Fig. 4(a) after

MPP+-exposure. In both assays, allopurinol largely prevented

cytochrome c release, indicating the participation of xanthine

oxidase in the early MPP+-induced cytochrome c release.

Cytochrome c release can be produced through the

mitochondrial permeability transition pore, which is fre-

quently coincident with loss of DYm (Petit et al. 1998;

Kroemer and Reed 2000). As MPP+-induced mitochondrial

permeability transition and loss of DYm have been describedin several cell models (Cassarino et al. 1999; Chalmers-

Redman et al. 1999), we decided to analyse whether MPP+

induced early changes in DYm in CGCs, by using the

fluorescent probe CMXRos in a flow cytometry assay. As

shown in Fig. 5, MPP+ had no effect on DYm at 30 min ofMPP+-exposure and it only induced loss of DYm at 6 h (datanot shown). This result demonstrates that MPP+-induced

early cytochrome c release precedes loss of DYm and that novariation in mitochondrial permeability transition is neces-

sary for MPP+-induced early cytochrome c release.

To identify the possible factors responsible for cytochrome

c release in CGCs exposed to MPP+, we prepared a

mitochondria-purified fraction and total cell lysates from

cells incubated with and without 50 lM MPP+. As Bad andBax are Bcl2-related proteins involved in the regulation of

cytochrome c release from mitochondria in several models of

apoptosis (von Harsdorf et al. 1999; Desagher and Martinou

2000), we determined their translocation to the mitochondria

in MPP+-induced cell death. Untreated cells showed unde-

tectable levels of Bax protein in mitochondrial fraction.

However, this protein translocated from cytosol to the

mitochondria (with a maximum over 30 min, data not

shown) after MPP+ exposure. The exact mechanism by

which MPP+ contributes to Bax translocation to mitochon-

dria is as yet unknown. Since this early translocation of Bax

was parallel with cytochrome c release, we next decided to

assay whether Bax translocation was inhibited by vitamin E.

Figure 6(a) shows that co-incubation with 100 lM vitamin Epartially blocked early MPP+-induced Bax translocation.

Total levels of Bax protein in the cell were not altered at the

same times (data not shown). Figure 6(b and c) shows the

changes in the patterns of Bad phosphorylation in MPP+-

exposed CGCs. A strong signal for phosphorylated Bad

protein was detected in lysates from control cells. In contrast,

phosphorylated Bad protein was barely detectable after

30 min of incubation with 50 lM MPP+. Exposure of the

neurons to 100 lM vitamin E (Fig. 6b) or 100 lM allopur-inol (Fig. 6a) maintained Bad phosphorylation even in the

(c)

(b)

(a)

*

*

*

Fig. 2 MPP+ effect on intracellular ROS content and GSH levels in

primary cultures of cerebellar granule cells. (a) and (b) CGCs in the

absence or presence of MPP+ (30 min) were incubated for a further

30 min with 20 lM or 5 lM, respectively, of the oxidation-sensitive

fluorescent probes hydroethidine (DHE) or 2¢-7¢-dichlorodihydrofluo-

rescein (DFCH), measuring the fluorescence intensity by flow cytom-

etry, as described in the Materials and methods. In both, the solid bar

shows untreated cells and the open bar shows the cells treated with

50 lM MPP+ for 30 min. Data are expressed as a percentage of DHE

or DFCH fluorescence with respect to control value and are

mean ± SEM values of three independent experiments. *Significantly

different (p < 0.05) from untreated cells. (c) Effect of MPP+ on GSH

levels in primary cultures of cerebellar granule cells. Cells were treated

(solid bar) or untreated (open bar) with 50 lM MPP+. After 30 min cells

were incubated with 1 lM of monochlorobimane (MCB), measuring the

fluorescence intensity by flow cytometry as described in the Materials

and methods. Data are expressed as a percentage of DHE or MCB

fluorescence with respect to control value and are mean ± SEM values

of three independent experiments. *Significantly different (p < 0.05)

from untreated cells.

Vitamin E blocks early cytochrome c release 309

� 2003 International Society for Neurochemistry, Journal of Neurochemistry, 84, 305–315

presence of MPP+. Total levels of Bad protein in the cell

were not altered at the same times (data not shown). Bad

protein is probably dephosphorylated by protein phospha-

tases as calcineurin, PP1a and others (Desagher and

Martinou, 2000). To evaluate whether participation of protein

phosphatases in the early events implicated MPP+-induced

apoptosis, we assayed the broad-range protein phosphatase

inhibitor okadaic acid. Figure 7(a and b) shows that 1 lMokadaic acid inhibited both Bad dephosphorylation and

cytochrome c release induced by 50 lM MPP+. These data

suggest a protein phosphatase participation in early apoptosis

induction in CGCs by MPP+. Interestingly phospho-Akt

levels (enzyme that phosphorylates Bad) are not modified by

MPP+ exposure (Fig. 7c). Because phospho-Akt is the active

form (Mora et al. 1999) and the medium conditions (high

potassium and insulin) favour this status, the decrease in

phospho-Bad protein shows that an early protein phosphatase

activation is implicated in its dephosphorylation, without

involvement of the PI3-K/Akt pathway.

Discussion

Apoptosis is thought to play an important role in the neuronal

loss in many neurological disorders, including Parkinson’s

disease. Cells undergoing apoptotic death exhibit several

morphological characteristics, such as chromatin condensa-

tion, nuclear fragmentation or apoptotic cell body formation.

Numerous studies suggest that MPP+ is able to induce

apoptosis in vitro in several cell types (Hartley et al. 1994;

Itano and Nomura 1995; Chalmers-Redman et al. 1999;

Yoshinaga et al. 2000) including CGCs (Dipasquale et al.

1991; Gonzalez-Polo et al. 1901; Du et al. 1997), some of

Fig. 3 Vitamin E inhibition of cytochrome c release and caspase 3

activation in primary cultures of cerebellar granule cells. CGC cultures

were exposed to: (a) 50 lM MPP+ at the times indicated, (b) 50 lM

MPP+ or 50 lM MPP+ plus 100 lM vitamin E for 30 min or (c)

increasing concentrations of MPP+ (5–200 lM) for 30 min. In both,

mitochondrial proteins (30–50 lg protein/lane) were size-fractionated

by SDS–PAGE, transferred onto PVDF and probed with cytochrome c

antibody as described in the Materials and methods. Actin content was

analysed as control Blots are representative of at least three inde-

pendent experiments. (d) Caspase 3 activity. CGCs cultures were

exposed to 50 lM MPP+ plus 100 lM vitamin E for 60 min after cells

were lysed and caspase 3 was assayed as described in the Materials

and methods. Data are expressed as a percentage of caspase 3

activity with respect to control value and are mean ± SEM values of

three independent experiments. *Significantly different (p < 0.05) from

untreated cells. (e) Proteolytic activation of caspase 3 and (f) its

prevention by 100 lM vitamin E. Cells were size-fractionated by SDS–

PAGE, transferred onto PVDF and probed with cleaved (active)

caspase 3 (17 kDa) antibody as described in the Materials and

methods. Actin content was analysed as control. Blots are represen-

tative of at least three independent experiments.

310 R. A. Gonzalez-Polo et al.

� 2003 International Society for Neurochemistry, Journal of Neurochemistry, 84, 305–315

the MPP+ effects being mediated by oxidative stress (Rossetti

et al. 1988; Kitamura et al. 1998). However, detection of

early apoptotic events involved in MPP+-cell death has not

been described yet. In this work we show that CGCs exposed

to MPP+ induce an early (in the 30 min after MPP+-

exposure) production of ROS, detected by the oxidation-

sensitive fluorescent probes HE and DFCH-DA. This

production is accompanied by GSH loss and appears to be

responsible for the apoptotic cell death, with the character-

istic cytochrome c release, caspase 3 activation, DNA

fragmentation and morphological changes, such as con-

densed chromatin. Previous works established a relationship

between MPP+-induced ROS production and inhibition of

mitochondrial complex I, but free radical production in these

models always occurred at later times (Mizuno et al. 1987;

Salinas et al. 2001). More recent works describe earlier ROS

generation in CGCs exposed to glutamate (Atlante et al.

1999) or potassium deprivation (Valencia and Moran 2001),

which appear to be directly involved in cytochrome c release

from mitochondria. Experiments reported in Fig. 1 demon-

strate that MPP+ can produce apoptotic cell death in CGCs

and Figs 2 and 3 also reveal an early cytochrome c release

MPP+ 50 µM

Allopurinol 100 µM

actin

actin

Cyt c mitochondria

Cyt c mitochondria

X/XOD

Allopurinol 100 µM

– + +

– – +

– + +

– – +

(a)

(b)

Fig. 4 Allopurinol blocks cytochrome c release from mitochondria in

both, MPP+-exposed and xanthine/xanthine oxidase system. Cells

were incubated for (a) 30 min with 50 lM MPP+ in the absence or the

presence of 100 lM of allopurinol or (b) 30 min with 10 mU/mL xanthine

oxidase plus 10 lM xanthine. In both mitochondrial proteins (30–50 lg

protein/lane) were size-fractionated by SDS–PAGE, transferred onto

PVDF and probed with cytochrome c antibody as described in the

Materials and methods. Actin content was analysed as control. Blot is

representative of at least three independent experiments.

100

Control

CM

xRos

Flu

ores

cenc

e(%

con

trol

)

MPP+

80

60

40

20

0

Fig. 5 MPP+ does not induce early changes in the mitochondrial

transmembrane potential (DYm) in primary cultures of cerebellar

granule cells. CGCs were incubated for 30 min in the absence (solid

bar) or presence (open bar) of 50 lM MPP+. After 30 min of incubation

with 0.1 lM CMXRos, the intracellular fluorescence intensity was

measured in a FACScan flow cytometer as described in the Materials

and methods. Data are expressed as a percentage of CMXRos

fluorescence with respect to control value and are mean ± SEM values

of three independent experiments.

Bax

MPP+ 50 µMvit E 100 µM

actin

– + +– – +

P-Bad

MPP+ 50 µMvit E 100 µM

actin

– + +– – +

P-Bad

MPP+ 50 µM

Allopurinol 100 µM

actin

– + +– – +

(a)

(b)

(c)

Fig. 6 Vitamin E partially prevents MPP+-induced mitochondrial Bax

translocation and vitamin E and allopurinol prevents Bad dephos-

phorylation in primary cultures of cerebellar granule cells. Cells were

incubated with 50 lM MPP+ in the presence or in the absence of

100 lM vitamin E for 30 min. (a) Mitochondrial proteins were pro-

cessed as described for cytochrome c release and analysed by

electrophoresis and western blot as described in the Materials and

methods using Bax antibody. (b) and (c) Cell lysates (30–50 lg protein

lane) obtained as described in the Materials and methods were sub-

jected to electrophoresis and western blot with an anti-Ser136 phos-

pho-Bad antibody. Actin content was analysed as control. Blots are

representative of at least three independent experiments.

Vitamin E blocks early cytochrome c release 311

� 2003 International Society for Neurochemistry, Journal of Neurochemistry, 84, 305–315

(30 min after MPP+-exposure) and caspase 3 activation

(60 min after MPP+-exposure) with an early oxidative stress

(free radical detection and intracellular GSH depletion).

Vitamin E co-incubation blocks both cell death and cyto-

chrome c release/caspase 3 activation, establishing a strong

relationship between early ROS generation and cytochrome c

translocation. These data have not been previously reported

in any cellular model of MPP+-induced neurotoxicity. The

knowledge that mitochondria are a direct target for MPP+

focused the attention on the cell changes that occurred

simultaneously with this event, all the earlier process being

ignored. However, the implication of mitochondrial free

radical production in MPP+-induced apoptotic cell death is

widely reported (Nicklas et al. 1987; Mizuno et al. 1995).

The fast cytosolic ROS production reported here is possibly a

result of MPP+ interaction with the xanthine oxidase enzyme

(Klaidman et al. 1993), which might lead to the formation of

ROS. Experiments reported in Fig. 4 demonstrate that

cytochrome c release is driven by ROS produced in MPP+-

exposure or by exogenous production via the xanthine/

xanthine oxidase system. This conclusion was supported by

the partial prevention of cytochrome c release with the

specific xanthine oxidase inhibitor allopurinol, which would

block ROS formation by impairing xanthine oxidase activity

in both cases. The MPP+ concentration-dependence and

timing in cytochrome c release shown in Fig. 3b could

suggest that this early translocation is a function of ROS

production in cytosol, probably through the xanthine oxidase

system. Blockage by vitamin E and allopurinol of cyto-

chrome c release demonstrates this fact. The cytosolic MPP+-

induced ROS production by xanthine oxidase would occur

earlier than that produced by mitochondrial impairment and

would be compatible with the events observed in the first

30 min after MPP+ exposure: loss of intracellular GSH and

cytochrome c release from mitochondria, without change in

mitochondrial transmembrane potential. Recent studies have

shown that early translocation of cytochrome c, previous to

mitochondrial collapse, is directly implicated in apoptosis

induction (Granville et al. 2001; Herrera et al. 2001; Li et al.

2001; Suen et al. 2001). In good agreement with these data,

our results clearly show that early cytochrome c release

drives cells to apoptotic cell death, because blockade of this

release by vitamin E increases cell survival and impairs

apoptotic events.

It is important to note that cytochrome c release is closely

connected with the Bcl-2 family proteins. Antiapoptotic

members Bcl-2 and Bcl-XL are, for the most part, mito-

chondrial proteins (Hsu et al. 1997) while other related

proteins, such as Bad or Bax, are cytosolic but translocate to

the mitochondria in response to different apoptotic stimuli

(Desagher et al. 1999; Downward 1999). The mechanisms

that promote Bax activation remain unclear, but glutathione

depletion, induced by an increase in free radical concentra-

tion in the cells, could be involved as has been recently

reported (Jungas et al. 2002). In CGCs exposed to MPP+ we

observed an increase in mitochondrial Bax levels, which is

parallel to free radical production and cytochrome c release.

In our conditions Bax translocation is blocked by vitamin E,

linking the Bax translocation to the free radical generation.

This result disagrees with that recently observed by Hart-

mann et al. (2001). In our study Bax is detected in

mitochondrial extract 30 min after incubation with 50 lMMPP+ while in Hartmann’s work Bax localization was

measured at 12 h with a significantly lower MPP+ concen-

tration (1–3 lM). The different time of exposure and MPP+

concentration can explain this disagreement. On the other

hand Bax-induced cytochrome c release is often associated

with changes in DYm (Lotharius et al. 1999; Antonson andMartinou 2000). Loss in DYm is also observed in severalMPP+-induced apoptosis systems (Lambert and Bond 1989;

Cassarino et al. 1999; Chalmers-Redman et al. 1999).

P-Bad

MPP+ 50 µM

Okadaic acid 1 µM

actin

– + +– – +

Cyt c mitochondria

MPP+ 50 µM

Okadaic acid 1 µM

actin

– + +– – +

P-Akt 60kDa

MPP+ 50 µM

actin

– +

(a)

(b)

(c)

Fig. 7 Okadaic acid blocks both MPP+-induced Bad dephosphoryla-

tion and cytochrome c release in primary cultures of cerebellar granule

cells. CGCs were exposed to 50 lM MPP+ for 30 min with or without

preincubation with 1 lM okadaic acid. (a) Cell lysates (30–50 lg pro-

tein/lane) were separated on polyacrylamide gels and western blot

analysis (as described in the Materials and methods) was performed

with an anti-Ser136 phospho-Bad antibody to detect the phosphory-

lation state of Bad. (b) Cytochrome c was detected in mitochondrial

extracts (30–50 lg protein/lane) by western blot as described in Fig. 3

and the Materials and methods using cytochrome c antibody.

(c) Phosphorylation state of Akt. Cells were also incubated for 30 min

in the presence or absence of 50 lM MPP+. After this time cell lysates

were extracted and Akt activity was analysed by electrophoresis

(30–50 lg protein/lane) and western blot using antibodies (specific for

Ser473) against the active form (P-Akt). Actin content was analysed as

control Blots are representative of at least three independent experi-

ments.

312 R. A. Gonzalez-Polo et al.

� 2003 International Society for Neurochemistry, Journal of Neurochemistry, 84, 305–315

However, we observed no changes in DYm coincident withBax translocation and cytochrome release. Our results are in

agreement with other recent studies, which implicated early

events in the development of apoptotic cell death (Annis

et al. 2001; Suen et al. 2001). Bax could form a channel in

mitochondria, allowing cytochrome c release, without mito-

chondrial damage as has been reported (Antonsson et al.

1997).

Another pro-apoptotic Bcl-2-related protein is Bad. In the

absence of apoptotic stimuli Bad is phosphorylated and

sequestered in the cytosol by binding to 14-3-3 protein (Zha

et al. 1996). During apoptosis Bad is dephosphorylated and

promotes cell death by binding to bcl-XL (Zha et al. 1996).

Bad is phosphorylated preferably, but not only, by Akt

(Dudek et al. 1997) and dephosphorylated by PP1 and PP2

phosphatases (Desagher and Martinou, 2000). In our work,

Bad is strongly dephosphorylated at the same time as Bax

translocation to the mitochondria. Bad can bind Bcl-XL and

this fact would contribute to the inhibition of the death-

repressor activity of Bcl-XL, facilitating the Bax-induced

cytochrome c release. Pre-incubation of CGCs with okadaic

acid, a broad-range protein phosphatase inhibitor partially

inhibits both early Bad dephosphorylation and cytochrome c

release. This result clearly indicates that phosphatases are

activated in the 30 min after MPP+ exposure and that this

activation would be involved in cytochrome c release.

Because Bad dephosphorylation is also inhibited by vitamin

E and allopurinol, we could hypothesize that phosphatase

activities are switched on by the ROS produced in the first

minutes after MPP+ exposure. Vitamin E also partially

inhibits Bax translocation to mitochondria, indicating that the

impairment of early MPP+-induced free radical production

eliminates all the apoptotic events initiated by MPP+.

In conclusion, our results (summarized in Fig. 8) demon-

strate that MPP+ produces early Bax translocation, Bad

dephosphorylation, cytochrome c release and caspase acti-

vation, concluding with cell death. All events are blocked by

vitamin E, suggesting that the early events implicated in

MPP+-induced apoptosis in CGCs are related to oxidative

stress probably as a result of activation of the xanthine

oxidase system. These results together point towards the

existence of some early events (30 min) that play an essential

Fig. 8 Summary of the proposed mechanism for the early apoptotic

events induced by MPP+ in primary cultures of cerebellar granule cells.

MPP+ production of ROS could be mediated by xanthine oxidase. The

oxidative stress would contribute to Bad dephosphorylation (by protein

phosphatase activation) and Bax translocation to the mitochondria,

which would mediate cytochrome c release (without loss of DYm),

caspase 3 activation and cell death. Vitamin E is able to block all the

observed events, including cell death.

Vitamin E blocks early cytochrome c release 313

� 2003 International Society for Neurochemistry, Journal of Neurochemistry, 84, 305–315

role in MPP+-induced apoptosis. Further work will be

necessary to understand completely the molecular mechan-

ism by which oxidative stress provokes Bax translocation

and protein phosphatase activation and all the events that

conclude with apoptotic cell death.

Acknowledgements

This work was supported, in part, by grants 2PR01A079, to GS

(from Junta de Extremadura, Spain), 01/51 to JMF (from Junta de

Extremadura, Spain), FIS010797 to IF (from Ministerio de Sanidad

y Consumo, Spain) and CAM-08.1/0078/2000 to IF (from Comu-

nidad de Madrid, Spain). RAGP was supported by a Spanish

Ministerio de Educacion, Cultura y Deporte predoctoral fellowship.

The authors also thanks to J. C. Alonso and J. M. Moran for their

inestimable helpful technical assistance.

References

Akaneya Y., Takahashi M. and Hatanaka H. (1995) Involvement of free

radicals in MPP+ neurotoxicity against rat dopaminergic neurons

in culture. Neurosci. Lett. 193, 53–56.

Annis M. G., Zamzami N., Zhu W., Penn L. Z., Kroemer G. and Leber

B. and Andrews D. W. (2001) Endoplasmic reticulum localized

Bcl-2 prevents apoptosis when redistribution of cytochrome c is a

late event. Oncogene 20, 1939–1952.

Antonsson B. and Martinou J. C. (2000) The Bcl-2 protein family. Exp.

Cell. Res. 256, 50–57.

Antonsson B., Conti F., Ciavatta A., Montessuit S., Lewis S., Martinou

I., Bernasconi L., Bernard A., Mermod J. J., Mazzei G., Maundrell

K., Gambale F. and Sadoul R. and. Martinou J. C. (1997) Inhibition

of Bax channel-forming activity by Bcl-2. Science 277, 370–372.

Atlante A., Calissano P., Bobba A., Azzariti A., Marra E. and Passarella

S. (2000) Cytochrome c is released from mitochondria in a reactive

oxygen species (ROS)-dependent fashion and can operate as a

ROS scavenger and as a respiratory substrate in cerebellar neurons

undergoing excitotoxic death. J. Biol. Chem. 275, 37159–37166.

Atlante A., Calissano P., Bobba A., Giannattasio S., Marra E. and

Passarella S. (2001) Glutamate neurotoxicity, oxidative stress and

mitochondria. FEBS Lett. 497(1), 1–5.

Atlante A., Gagliavoli S., Marra E., Calissano P. and Passavella S.

(1999) Glutamate neurotoxicity in rat cerebellar granule cells

involve cytochrome c release from mitochondria and mitochondrial

shuttle impairment. J. Neurochem. 73(1), 237–246.

Blum D., Torch T., Lambeng N., Nissou M. F., Benabid A. L., Sadoul R.

and Verna. J. R. (2001) Molecular pathways involved in the neu-

rotoxicity of 6-OHDA, dopamine and MPTP, contribution to the

apoptotic theory in Parkinson’s Disease. Prog. Neurobiol. 65, 135–

172.

Bobba A., Atlante A., Giannattasio S., Sgaramella G., Calissano P. and

Marra E. (1999) Early release and subsequent caspase-mediated

degradation of cytochrome c in apoptotic cerebellar granule cells.

FEBS Lett. 457, 126–130.

Bradford M. (1976) A rapid and sensitive method for the quantitation of

microgram quantities of protein utilizing the principle of protein-

dye binding. Anal. Biochem. 72, 248–254.

Cassarino D. S., Parks J. K., Parker W. D. Jr and Bennett J. P. Jr (1999)

The parkinsonian neurotoxin MPP + opens the mitochondrial

permeability transition pore and releases cytochrome c in isolated

mitochondria via an oxidative mechanism. Biochim. Biophys. Acta

1453, 49–62.

Castagnoli N. Jr, Chiba K. and Trevor K. (1985) Potential bioactivation

pathways for the neurotoxin 1-methyl-4-phenyl-1,2,3,6-tetra-

hydropyridine (MPTP). Life Sci. 36, 225–230.

Chalmers-Redman R. M., Fraser A. D., Carlile W., Pong A. and Tatton

W. G. (1999) Glucose protection from MPP+-induced apoptosis

depends on mitochondrial membrane potential and ATP synthase.

Biochem. Biophys. Res. Commun. 257, 440–447.

Chiba K., Trevor A. and Castaganoli N. Jr (1985) Active uptake of

MPP+, a metabolite of MPTP by brain synaptosomes. Biochem.

Biophys. Res. Commun. 128, 1228–1232.

Desagher S. and Martinou J. C. (2000) Mitochondria as the central

control point of apoptosis. Trends Cell Biol. 10, 369–377.

Desagher S., Osen-Sand A., Nichols A., Eskes R., Montessuit S.,

Lauper S., Maundrell K., Antonsson B. and Martinou J. C. (1999)

Bid-induced conformational change of Bax is responsible for

mitochondrial cytochrome c release during apoptosis. J. Cell Biol.

144, 891–901.

Dipasquale B., Marini A. M. and Youle R. J. (1991) Apoptosis and DNA

degradation induced by 1-methyl-4-phenylpyridinium in neurons.

Biochem. Biophys. Res. Commun. 181, 1442–1448.

Dodel R. C., Du Y., Bales K. R., Ling Z. D., Carvey P. M. and Paul S. M.

(1998) Peptide inhibitors of caspase-3-like proteases attenuate

1-methyl-4-phenylpyridinum-induced toxicity of cultured fetal rat

mesencephalic dopamine neurons. Neuroscience 86, 701–707.

Downward J. (1999) How BAD phosphorylation is good for survival.

Nat. Cell Biol. 1, E33–E35.

Du Y., Dodel R. C., Bales K. R., Jemmerson R., Hamilton-Bird E. and

Paul S. M. (1997) Involvement of a caspase-3-like cysteine

protease in 1-methyl-4-phenylpyridinium-mediated apoptosis

of cultured cerebellar granule neurons. J. Neurochem. 69, 1382–

1388.

Dudek H., Datta S. R., Franke T. F., Birnbaum M. J., Yao R., Cooper

G. M., Segal R. A., Kaplan D. R. and Greenberg M. E. (1997)

Regulation of neuronal survival by the serine-threonine protein

kinase Akt. Science 275, 661–665.

Fuentes J. M., Lompre A. M., Moller J. V., Falson P., le Maire M. (2000)

Clean Western blots of membrane proteins after yeast heterologous

expression following a shortened version of the method of Perini

et al. Anal. Biochem. 285, 276–278.

Gonzalez-Polo R. A., Mora A., Clemente N., Sabio G., Centeno F., Soler

G. and Fuentes J. M. (2001) Mechanisms of MPP+ incorporation

into cerebellar granule cells. Brain Res. Bull. 56, 119–123.

Granville D. J., Cassidy B. A., Ruehlmann D. O., Choy J. C., Brenner

C., Kroemer G., van Breemen C., Margaron P., Hunt D. W. and

McManus B. M. (2001) Mitochondrial release of apoptosis-indu-

cing factor and cytochrome c during smooth muscle cell apoptosis.

Am. J. Pathol. 159, 305–311.

Gross A., McDonnell J. M. and Korsmeyer S. J. (1999) BCL-2 family

members and the mitochondria in apoptosis. Genes Dev. 13, 1899–

1911.

Hartley A., Stone J. M., Heron C., Cooper J. M. and Schapira A. H.

(1994) Complex I inhibitors induce dose-dependent apoptosis in

PC12 cells: relevance to Parkinson’s disease. J. Neurochem. 63,

1987–1990.

Hartmann A., Michel P. P., Troadec J. C., Mouatt-Prigent A., Faucheux

B. A., Ruberg M., Agid Y., Hirsch E. C. (2001) Is Bax a mito-

chondrial mediator in apoptotic death of dopaminergic neurons in

Parkinson’s disease? J. Neurochem. 76, 1785–1793.

Herrera B., Fernandez M., Alvarez A. M., Roncero C., Benito M., Gil J.

and Fabregat I. (2001) Activation of caspases occurs downstream

from radical oxygen species production, Bcl-xL down-regulation,

and early cytochrome C release in apoptosis induced by trans-

forming growth factor beta in rat fetal hepatocytes. Hepatology 34,

548–556.

314 R. A. Gonzalez-Polo et al.

� 2003 International Society for Neurochemistry, Journal of Neurochemistry, 84, 305–315

Hsu Y. T., Wolter K. G. and Youle R. J. (1997) Cytosol-to-membrane

redistribution of Bax and Bcl-X (L) during apoptosis. Proc. Natl

Acad. Sci. USA 94, 3668–3675.

Itano Y. and Nomura Y. (1995) 1-methyl-4-phenyl-pyridinium ion

(MPP+) causes DNA fragmentation and increases the Bcl-2

expression in human neuroblastoma, SH-SY5Y cells, through

different mechanisms. Brain Res. 704, 240–245.

Jungas T., Motta I., Duffieux F., Fanen P., Stoven V. and Ojcius D. M.

(2002) Glutathione levels and Bax activation during apoptosis due

to oxidative stress in cells expressing wildtype and mutant CFTR.

J. Biol. Chem. 277(31), 27912–27918.

Kamemcic H., Lyon A., Paterson P. G. and Juurlink B. H. (2000)

Monoclorobimane fluorometric method to measure tissue gluta-

thione. Anal. Biochem. 286, 35–37.

Kitamura Y., Kosaka T., Kakimura J. L., Matsuoka Y., Kolmo Y., Nomura

Y. and Taniguchi T. (1998) Protective effects of the antiparkinsonian

drugs talipexole and pramipexole against 1-methyl-4-phenylpyridi-

nium-induced apoptotic cell death in human neuroblastoma

SH-Sy5Y cells. Mol. Pharmacol. 54, 1046–1054.

Klaidman L. K., Adams J. D., Leung A. C., Sam Jim S. and Cadenas E.

(1993) Redox cycling of MPP+: Evidence for a new mechanism

involving hydride transfer with xanthine oxidase, aldehyde dehy-

drogenase, and lipoamide dehydrogenase. Free Radic. Biodiv.

Med. 15, 169–179.

Kroemer G. and Reed J. C. (2000) Mitochondrial control of cell death.

Nat. Med. 6, 513–519.

Lambert C. E. and Bondy S. C. (1989) Effects of MPTP, MPP+ and

paraquat on mitochondrial potential and oxidative stress. Life Sci.

44, 1277–1284.

Langston J. W., Ballard P., Tetrud J. W. and Irwin I. (1983) Chronic

Parkinsonism in humans due to a product of meperidine-analog

synthesis. Science 219, 979–980.

Leist M., Volbracht C., Fava E. and Nicotera P. (1998) 1-Methyl-4-

phenylpyridinium induces autocrine excitotoxicity, protease acti-

vation, and neuronal apoptosis. Mol. Pharmacol. 54, 789–801.

Li P., He Q. P., Ouyang Y. B., Liu C. L., Hu B. R. and Siesjo B. K.

(2001) Early release of cytochrome C and activation of caspase-3

in hyperglycemic rats subjected to transient forebrain ischemia.

Brain Res. 896, 69–76.

Lotharius J., Dugan L. L. and O’Malley K. L. (1999) Distinct mech-

anisms underlie neurotoxin-mediated cell death in cultured dop-

aminergic neurons. J. Neurosci. 19, 1284–1293.

Lowry O. H., Rosebrough N. J., Farr A. L. and Randall R. J. (1951)

Protein measurement with the Folin phenol reagent. J. Biol. Chem.

193, 265–271.

Mizuno Y., Ohta S., Tanaka M., Takamiya S., Suzuki K., Sato T., Oya

H., Ozawa T. and Kagawa K. (1989) Deficiencies in complex I

subunits of the respiratory chain in Parkinson’s Disease. Biochem.

Biophys. Res. Commun. 163, 1450–1455.

Mizuno Y., Saitoh T. and Sone N. (1987b) Inhibition of mitochondrial

NADH-ubiquinone oxidoreductase activity by 1-methyl-4-phe-

nylpyridinium ion. Biochem. Biophys. Res. Commun. 143, 294–

299.

Mora A., Gonzalez-Polo R. A., Fuentes J. M., Soler G. and Centeno F.

(1999) Different mechanisms of protection against apoptosis by

valproate and Li+. Eur J. Biochem. 266, 886–891.

Mosmann T. (1983) Rapid colorimetric assay for cellular growth and

survival: application to proliferation and cytotoxicity assays.

J. Immunol. Meth. 65, 55–63.

Nicklas W. J., Vyas I. and Heikkila R. E. (1985) Inhibition of NADH-

linked oxidation in brain mitochondria by 1-methyl-4-phenyl-

1,2,5,6-tetrahydropyridine. Life Sci. 36, 2503–2508.

Nicklas W. J., Youngster S. K., Kindt M. V. and Heikkila R. E. (1987)

MPTP, MPP+ and mitochondrial function. Life Sci. 40(8), 721–

729.

Obata T. (1999) Reserpine prevents hydroxyl radical formation by MPP+

in rat striatum. Brain Res. 828, 68–73.

Odunze I. N., Klaidman L. K. and Adams J. D. Jr (1990) MPTP toxicity

in the mouse brain and vitamin E. Neurosci. Lett. 108, 346–349.

Putcha G. V., Deshmukh M. and Johnson E. M. Jr. (1999) BAX trans-

location is a critical event in neuronal apoptosis: regulation by

neuroprotectants, BCL-2, and caspases. J. Neurosci. 19(17), 7476–

7485.

Petit P. X., Goubern M., Diolez P., Susin S. A., Zamzami N. and

Kroemer G. (1998) Disruption of the outer mitochondrial

membrane as a result of large amplitude swelling: the impact of

irreversible permeability transition. FEBS Lett. 426, 111–116.

Pique M., Barragan M., Dalmau M., Bellosillo B., Pons G. and Gil J.

(2000) Aspirin induces apoptosis through mitochondrial cyto-

chrome c release. FEBS Lett. 480, 193–196.

Przedborski S., Kostic V., Jackson-Lewis V., Naini A. B., Simonetti S.,

Fahn S., Carlson E., Epstein C. J. and Cadet J. (1992) Transgenic

mice with increased Cu-Zn-superoxide dismutase activity are

resistant to N-methyl-4-phenyl-1,2,3,6-tetrahydropyridine-induced

neurotoxicity. J. Neurosci. 12, 1658–1667.

Rossetti Z., Sotgiu A., Sharp D. E., Hadgiconstantinou M. and Neff

N. H. (1988) 1-methyl-4-phenyl-1,2,5,6-tetrahydropyridine (MPTP)

and free radicals in vitro. Biochem. Pharmacol. 37, 4573–4574.

Rothe G. and Valet G. (1990) Flow cytometric analysis of respiratory

burst activity in phagocytes with hydroethidine and 2¢,7¢-dichlo-rofluorescin. J. Leukoc. Biol. 47, 440–448.

Salinas M., Martın D., Alvarez A. and Cuadrado A. (2001) Akt1/PKBaprotects PC12 cells against the Parkinsonism-inducing neurotoxin

1-methyl-4-phenylpyridinium and reduces the levels of oxygen-

free radicals. Mol. Cell. Neurosci. 17, 67–77.

Sinha B. K., Singh Y. and Krishna G. (1986) Formation of su-

peroxide and hydroxyl radicals from 1-methyl-4-phenylpyridi-

nium ion (MPP+): reductive activation by NADPH cytochrome

P-450 reductase. Biochem. Biophys. Res. Commun. 135, 583–

588.

Suen Y. K., Fung K. P., Lee C. Y. and Kong S. K. (2001) Gliotoxin

induces apoptosis in cultured macrophages via production of

reactive oxygen species and cytochrome c release without mito-

chondrial depolarization. Free Radic. Res. 35, 1–10.

Tammariello S. P., Quinn M. T. and Estus S. (2000) NADPH oxidase

contributes directly to oxidative stress and apoptosis in nerve growth

factor-deprived sympathetic neurons. J. Neurosci. 20, 123–132.

Tipton K. F. and Singer T. P. (1993) Advances in our understanding of

the mechanisms of the neurotoxicity of MPTP and related com-

pounds. J. Neurochem. 61, 1191–1206.

Valencia A. and Moran J. (2001) Role of oxidative stress in the apoptotic

cell death of cultured cerebellar granule neurons. J. Neurosci. Res.

64, 284–297.

von Harsdorf R., Li P. F. and Dietz R. (1999) Signaling pathways in

reactive oxygen species-induced cardiomyocyte apoptosis. Circu-

lation 99, 2934–2941.

Yoshinaga N., Murayama T. and Nomura Y. (2000) Apoptosis induction

by a dopaminergic neurotoxin, 1-methyl-4-phenylpyridinium ion

(MPP+), and inhibition by epidermal growth factor in GH3 cells.

Biochem. Pharmacol. 60, 111–120.

Zha J., Harada H., Yang E., Jockel J. and Korsmeyer S. J. (1996) Serine

phosphorylation of death agonist BAD in response to survival

factor results in binding to 14-3-3 not BCL-X (L). Cell 87, 619–

628.

Vitamin E blocks early cytochrome c release 315

� 2003 International Society for Neurochemistry, Journal of Neurochemistry, 84, 305–315