Involvement of mitochondria in endoplasmic reticulumstress-induced apoptotic cell death pathway triggeredby the prion peptide PrP106–126

Elisabete Ferreiro, Rui Costa, Sueli Marques, Sandra Morais Cardoso, Catarina R. Oliveira andClaudia M. F. Pereira

Center for Neuroscience and Cell Biology, Institute of Biochemistry, Faculty of Medicine, University of Coimbra, Coimbra, Portugal

Prion disorders (PrD) are progressive neurodegenerativediseases characterized by neuronal loss, cognitive dysfunc-tion and dementia. The principal hallmark of these disordersis the spongiform degeneration of the brain because of theextensive neuronal loss, astrogliosis and accumulation ofextracellular deposits of the scrapie isoform of prion protein(PrPSc) resulting from the abnormal conversion of thecellular prion protein (PrPC) (Prusiner 1998). PrPSc isinsoluble in non-denaturating detergents, aggregates easilyand is partially resistant to protease digestion. The PrP106–126,a synthetic peptide homologous to PrPSc, has been used as amodel system to study prion-induced neurodegeneration(Della-Bianca et al. 2001; Perez et al. 2003). PrP106–126 isrich in b-sheet structure, forms aggregates that are detergentinsoluble and proteinase K-resistant and catalyses theaggregation of endogenous PrPC to an amyloidogenic formthat shares several characteristics with PrPSc (Ettaiche et al.2000; Singh et al. 2000).

Endoplasmic reticulum (ER) stress-mediated apoptosis hasbeen shown to be involved in the neuronal death that occursin several neurodegenerative diseases, such as PrD (Nakag-awa et al. 2000; Hetz et al. 2003; Ferreiro et al. 2007).Perturbation of ER Ca2+ homeostasis can lead to the

Received May 29, 2007; revised manuscript received June 14, 2007;accepted September 18, 2007.Address correspondence and reprint requests to Claudia M. F. Pereira,

Institute of Biochemistry, Faculty of Medicine, University of Coimbra,Coimbra 3004-504, Portugal. E-mail: cpereira@cnc.cj.uc.ptAbbreviations used: Ab, amyloid-beta; DTT, dithiothreitol; ER,

endoplasmic reticulum; ETC, electron transport chain; GRP,glucose-regulated protein; mtDNA, mitochondrial DNA; MTT, 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide; PARP, poly(ADP-ribose) polymerase; PBS, phosphate-buffered saline; PrD, priondisorders; PrP, prion peptide; PrPC, cellular prion protein; PrPSc, scrapieisoform of prion protein; RyR, ryanodine receptor; TBS-T, Tris-bufferedsaline with 0.1% Tween 20.

Abstract

Prion disorders are progressive neurodegenerative diseases

characterized by extensive neuronal loss and by the accu-

mulation of the pathogenic form of prion protein, designated

PrPSc. Recently, we have shown that PrP106–126 induces

endoplasmic reticulum (ER) stress, leading to mitochondrial

cytochrome c release, caspase 3 activation and apoptotic

death. In order to further clarify the role of mitochondria in ER

stress-mediated apoptotic pathway triggered by the PrP

peptide, we investigated the effects of PrP106–126 on the

Ntera2 human teratocarcinoma cell line that had been

depleted of their mitochondrial DNA, termed NT2 q0 cells,

characterized by the absence of functional mitochondria, as

well as on the parental NT2 q+ cells. In this study, we show

that PrP106–126 induces ER stress in both cell lines, given that

ER Ca2+ content is low, glucose-regulated protein 78 levels

are increased and caspase 4 is activated. Furthermore, in

parental NT2 q+ cells, PrP106–126-activated caspase 9 and 3,

induced poly (ADP-ribose) polymerase cleavage and

increased the number of apoptotic cells. Dantrolene was

shown to protect NT2 q+ from PrP106–126-induced cell death,

demonstrating the involvement of Ca2+ release through ER

ryanodine receptors. However, in PrP106–126-treated NT2 q0

cells, apoptosis was not able to proceed. These results

demonstrate that functional mitochondria are required for cell

death as a result of ER stress triggered by the PrP peptide,

and further elucidate the molecular mechanisms involved in

the neuronal loss that occurs in prion disorders.

Keywords: apoptosis, Ca2+ homeostasis, endoplasmic reti-

culum, mitochondria, prion disorders, prion peptide.

J. Neurochem. (2008) 104, 766–776.

d JOURNAL OF NEUROCHEMISTRY | 2008 | 104 | 766–776 doi: 10.1111/j.1471-4159.2007.05048.x

766 Journal Compilation � 2007 International Society for Neurochemistry, J. Neurochem. (2008) 104, 766–776� 2007 The Authors

accumulation of unfolded proteins and activate the ER stress-induced apoptotic pathway (Kaufman 1999; Pashen 2001). Inresponse to ER stress, expression of glucose-regulatedprotein (GRP78), an ER chaperone, is induced (Kaufman1999; Rao et al. 2002). Under conditions of irreversible ERstress, caspase 4, the closest homologue of the rodent caspase12 in humans, is activated leading to apoptosis (Hitomi et al.2004).

We have previously shown that, in cortical neurons,PrP106–126 triggers ER stress and the early release of ERCa2+. In addition, we demonstrated that the PrP106–126-induced perturbation of ER Ca2+ homeostasis is responsiblefor the increase in cytosolic Ca2+ concentration, leading tocytochrome c release, caspase 3 activation and apoptosis(Ferreiro et al. 2006). These results suggest themitochondrial involvement in cell death that occurs duringPrP106–126-induced ER stress. To further examine the role ofmitochondria in the ER stress-mediated apoptotic cell deathcaused by the PrP peptide, we analysed the effects ofPrP106–126 on the Ntera2 human teratocarcinoma cell linedepleted of their mitochondrial DNA (mtDNA), termed NT2q0 cells, as well as on the parental NT2 q+ cells. The q0 cellline lack a functional electron transport chain (ETC), as 13protein subunits of the mitochondrial respiratory apparatusare mtDNA encoded (Wallace 1994; Jiang et al. 1999). Wefound that PrP106–126 induces ER stress in both cells lines,given that ER Ca2+ content was depleted, GRP78 levels wereincreased and caspase 4 was activated. In NT2 q+ cells,PrP106–126 increased caspase 9- and 3-like activities, Poly(ADP-ribose) polymerase (PARP) cleavage and also DNAfragmentation. Dantrolene, inhibitor of ER Ca2+ release, wasshown to prevent PrP106–126-induced cell death. However, inPrP106–126-treated NT2 q0 cells, although ER stress isinduced, apoptosis is not able to proceed, as no activationof caspase 9 and 3 nor cleavage of PARP and DNAfragmentation were observed. These results support that afunctional mitochondria is required for PrP-inducedER-mediated apoptosis.

Materials and methods

Cell lines and experimental treatmentsNT2 human teratocarcinoma cells (Stratagene, La Jolla, CA, USA)

containing mtDNA (q+) were grown in 75-cm2 tissue flasks in

Optimem Medium (Gibco, Pasley, Scotland, UK) containing 10%

heat-inactivated foetal bovine serum (Gibco), penicillin (100 U/mL)

and streptomycin (100 lg/mL). The mtDNA depleted NT2 rho0 cell

line (q0 cells), obtained after NT2 q+ treatment with ethidium

bromide, was a generous gift from Professor R Swerdlow

(Department of Neurology, University of Virginia, VA, USA). As

described by Swerdlow et al. (1997), cells were grown in 75-cm2

tissue flasks in Optimem Medium containing 10% heat-inactivated

foetal bovine serum, penicillin (50 U/mL) and streptomycin (50 lg/mL), and further supplemented with uridine (150 lg/mL) and

pyruvate (200 lg/mL). Both cell lines were maintained at 37�C in a

humidified incubator under an atmosphere of 95% air and 5% CO2

and were used for experiments after 1 day in vitro. Regular controlmeasurements of mitochondrial cytochrome c oxidase (complex IV)

activity were performed as a guaranty of the absence of ETC activity

in the NT2 q0 cells. NT2 q+ and q0 were treated with synthetic

PrP106–126 (Bachem, Bubendorf, Switzerland) at a concentration of

25 lmol/L for 3, 6 or 24 h, added from a stock solution prepared in

sterile distilled water, at a concentration of 1 mmol/L. Before the

addition of PrP106–126, NT2 q+ cells were pre-incubated for 1 h with

dantrolene (10 lmol/L) (Sigma Chemical Co., St Louis, MO, USA).

Dantrolene per se did not affect any of the parameters evaluated

during this study.

Measurement of ER Ca2+ content

Endoplasmic reticulum Ca2+ content was assessed by single cell

Ca2+ imaging, according to the method described by Nutt et al.(2002), with some modifications. Briefly, cells plated in coverslips

were treated with PrP106–126 (25 lmol/L) for 3 or 24 h. Treated and

control cells were washed two times in Krebs buffer (pH 7.4) (in

mmol/L: 132 NaCl, 4 KCl, 1.4 MgCl2, 6 glucose, 10 HEPES, 10

NaHCO3, 1 CaCl2) and loaded with 5 lmol/L Fura-2 acetoxymethyl

ester (Molecular Probes, Eugene, OR, USA), supplemented with

0.2% pluronic acid (w/v), in Krebs buffer for 40 min at 37�C, in the

dark. Afterwards, cells were washed three times in Ca2+-free Krebs

buffer, supplemented with 50 lmol/L EGTA, and the coverslip was

assembled to the perfusion chamber filled with 500 lL of Ca2+-free

Krebs buffer, in an inverted fluorescence microscope Axiovert 200

(Zeiss, Jena, Germany). Cells were alternately excited at 340 and

380 nm using a Lambda DG4 apparatus (Sutter Instruments

company, Nocato, CA, USA), and emitted fluorescence was

collected with a 40· objective and was driven to a coll SNAP

digital camera (Roper Scientific, Trenton, NJ, USA). After a

baseline was established, cells were stimulated with thapsigargin

(2.5 lmol/L final concentration), in the absence of extracellular

Ca2+, to empty Ca2+ from ER. Acquired values were processed

using the METAFLUOR software (Universal Imaging Corporation,

Buckinghamshire, UK). The peak amplitude of Fura-2 fluorescence

(ratio at 340/380 nm) was used to evaluate ER Ca2+ content.

Caspases activity assayCaspase activity assay was performed as described in Ferreiro et al.(2004). Briefly, untreated or treated NT2 q+/q0 cells were lysed

with a buffer containing (in mmol/L): HEPES-Na 25, MgCl2 2,

EDTA 1, EGTA 1 supplemented with 100 lmol/L phenylmethyl-

sulphonyl fluoride, 2 mmol/L dithiothreitol (DTT) and a protease

inhibitor cocktail (containing 1 lg/mL leupeptin, pepstatin A,

chymostatin and antipain). The cellular suspension was rapidly

frozen/defrosted three times, using liquid N2, and then centrifuged

for 10 min at 20 200 g. The supernatant was collected and assayed

for protein content using the Bio-Rad protein dye assay reagent

(Bio-Rad, Hercules, CA, USA). Aliquots of cell extracts containing

50 or 100 lg of protein were incubated for 2 h at 37�C, in a

reaction buffer containing 25 mmol/L HEPES-Na, 10 mmol/L DTT,

10% sucrose (w/v) and 0.1% CHAPS (w/v) (pH 7.4), with

100 lmol/L Ac-LEHD-pNA (BioSource International, Nivelles,

Belgium), Ac-DEVD-pNA (Calbiochem, Darmastadt, Germany) or

Ac-LEVD-pNA (MBL International Co, Wobum, MA, USA),

� 2007 The AuthorsJournal Compilation � 2007 International Society for Neurochemistry, J. Neurochem. (2008) 104, 766–776

Mitochondria-dependent cell death as a result of ER stress by PrP106–126 | 767

chromogenic substrates for caspase 9, 3 and 4 respectively. Caspases

activity was determined by measuring substrate cleavage at 405 nm

using a microplater reader and results were expressed as the increase

above control absorbance.

Western blottingFor the preparation of total cell extracts, cells which were either

treated or untreated with the PrP106–126 peptide were scraped in

200 lL of ice-cold lysis buffer containing (in mmol/L): HEPES-Na

25, MgCl2 2, EDTA 1, EGTA 1, supplemented with 100 lmol/L

phenylmethylsulphonyl fluoride, 2 mmol/L DTT and protease

inhibitor cocktail (containing 1 lg/mL leupeptin, pepstatin A,

chymostatin and antipain). Cell lysates were frozen/defrosted three

times in liquid N2 and were centrifuged at 140 g, during 2 min, to

remove nuclei and large debris. Protein concentration in the

supernatant was measured using the Bio-Rad protein dye assay

reagent. Samples were denaturated at 95�C for 3 min in a 6x

concentrated sample buffer (mmol/L): Tris 500, DTT 600, 10.3%

sodium dodecyl sulphate (w/v), 30% glycerol (v/v) and 0.012%

(w/v) bromophenol blue. Equal amounts (60 lg) of each protein

sample were separated by electrophoresis on a 10% sodium dodecyl

sulphate–polyacrylamide gel electrophoresis and electroblotted onto

polyvinylidene difluoride membranes (Amersham Pharmacia Bio-

tech, Buckinghamshire, UK). The identification of proteins of

interest was facilitated by the usage of a pre-stained precision

protein standard (Bio-Rad) which was run simultaneously. After

the proteins were electrophoretically transferred, the membranes

were incubated for 1 h at 25�C in Tris-buffer (mmol/L): NaCl 150,

Tris–HCl 25 (pH 7.6) with 0.1% Tween 20 (v/v) (TBS-T) containing

5% (w/v) non-fat dry milk to eliminate non-specific binding and

were next incubated overnight at 4�C in TBS-T containing 1% (w/v)

non-fat dry milk with a mouse monoclonal antibody against PrP

(1 : 250 dilution) (DakoCytomation, Glostrup, Denmark), a rabbit

polyclonal primary antibody against PARP (1 : 1000 dilution) (Cell

Signaling, Danvers, MA, USA), a mouse monoclonal primary

antibody against GRP78 (1 : 250 dilution) (BD Pharmigen, San

Diego, CA, USA), a rabbit polyclonal primary antibody against Bax

(1 : 1000 dilution) (Cell Signaling) or with a mouse monoclonal

primary antibody against Bcl-2 (1 : 500 dilution) (Santa Cruz

Biotechnology, Santa Cruz, CA, USA). The membranes were

washed several times and then incubated in TBS-T with 1% non-fat

dry milk for 2 h at 25�C, with the appropriate alkaline phosphatase-

conjugated anti-rabbit or anti-mouse secondary antibody at a

dilution of 1 : 25 000 or 1 : 20 000 respectively (Amersham

Pharmacia Biotech). Immunoreactive bands were detected after

incubation of membranes with ECF reagent (Amersham Pharmacia

Biotech) for 5–10 min, on a Bio-Rad Versa Doc 3000 Imaging

System (Bio-Rad).

Cell viability analysisThe reduction of 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazoli-

um bromide (MTT) (Sigma Chemical Co.) to formazan was

determined according to the method described by Mosmann

(1983), slightly modified. In brief, NT2 q+/q0 cells were washed

in sodium medium (in mmol/L): NaCl 132, KCl 4, NaH2PO4 1.2,

MgCl2 1.4, glucose 6, HEPES-Na 10 and CaCl2 1, pH 7.4 and were

incubated in MTT (0.5 mg/mL)-containing sodium medium for 2 h

at 37�C. After this period, the blue formazan crystals were dissolved

in an equal volume of 0.04 mol/L HCl in isopropanol and quantified

by measuring the absorbance at 570 nm. MTT reduction was

expressed as the percentage (%) of control cells absorbance.

TUNEL assayDNA fragmentation was analysed by TUNEL staining, which was

performed using an in situ Cell Death Detection Kit, Fluorescein

(Roche Applied Science, Mannheim, Germany) according to the

manufacturer’s directions. Control or PrP106–126-treated cells were

washed two times in phosphate-buffered saline (PBS) buffer (pH

7.4) and were fixed with 4% (w/v) p-formaldehyde for 30 min at

25�C. After washing twice in PBS buffer, the slides were immersed

in 0.1% Triton X-100 (v/v), supplemented with 0.1% (w/v) sodium

citrate, in PBS, for 2 min on ice, to permeabilize the cells. Slides

were further washed three times and incubated in a mixture of the

enzymatic solution with the label solution, for 1 h at 37�C, in the

dark. Finally, slides were washed again three times with PBS and

mounted with DakoCytomation Fluorescent solution (DakoCyto-

mation, Carpinteria, CA, USA) on a microscope slide for visual-

ization in a Axiovert Microscope 200 (Zeiss, Thornwood, NY,

USA). Dnase pre-treated cells were used as a positive control. The

photographs were taken at a magnification of 400·. All experiments

were performed in duplicate and total number and positive-TUNEL

cells were scored in three microscope fields for each coverslip.

Statistical analysisResults are expressed as mean ± SEM of the number of experiments

indicated in the figure captions. Statistical significance was

performed using ANOVA, followed by Dunnett’s post hoc tests for

multiple comparisons or by the unpaired two-tailed Student’s t-test.A p < 0.05 value was considered statistically significant.

Results

PrP106–126 induces ER stress both in parental NT2 q+ andmtDNA-depleted q0 cellsTo initially evaluate whether ER stress induced by PrP106–126is affected by the presence of a functional mitochondria, weused mtDNA depleted NT2 q0 cells and the parental cellline, NT2 q+ cells. To achieve this purpose, ER Ca2+

homeostasis and ER stress markers, namely GRP78 andcaspase 4, which are affected under ER stress conditions(Rao et al. 2002; Hitomi et al. 2004), were investigated, inboth NT2 q+ and q0 cells upon PrP106–126 treatment, ERCa2+ content was analysed by single cell Ca2+ imaging,GRP78 levels were measured by western blotting andcaspase 4 activity was determined by measuring the degra-dation of a chromogenic substrate.

In cells incubated with PrP106–126 (25 lmol/L) for 3 or24 h and loaded with the Ca2+ sensitive fluorescent dye Fura-2 acetoxymethyl ester, ER Ca2+ content was evaluatedindirectly by promoting the release of Ca2+ from ER storeswith thapsigargin, an inhibitor of the ER Ca2+ ATPase, in theabsence of external Ca2+. Representative traces of Fura-2

Journal Compilation � 2007 International Society for Neurochemistry, J. Neurochem. (2008) 104, 766–776� 2007 The Authors

768 | E. Ferreiro et al.

fluorescence ratio at 340 and 380 nm are presented inFig. 1a–d. In the parental NT2 q+ cells, basal values of Fura-2 fluorescence, measured in Ca2+-free medium before theaddition of thapsigargin, were shown to be higher in cells

treated with PrP106–126 for 24 h then in controls (Fig. 1b),whereas at 3 h of incubation no clear difference wasobserved between the two conditions (Fig. 1a). In NT2 q0cells, both at 3 and 24 h, basal values of Fura-2 fluorescence

Thapsigargin

Control

Thapsigargin

NT2 ρ0 cells

NT2 ρ+ cells

NT2 ρ0 cells

NT2 ρ+ cells

Rat

io F

340/

F38

0

0.6

0.65

0.7

0.75

0.8

0.85

Rat

io F

340/

F38

0

0.6

0.65

0.7

0.75

0.8

0.85

Rat

io F

340/

F38

0

0.6

0.65

0.7

0.75

0.8

0.85

Rat

io F

340/

F38

0

0.6

0.65

0.7

0.75

0.8

0.85

0 6 9 123

Time (min)0 6 9 123

Time (min)

0 6 9 123

Time (min)

0 6 9 123

Time (min)

PrP106–126

Control

PrP106–126

Control

PrP106–126

Control

PrP106–126

Control

PrP106–126

***

***

24 h 24 h

0.00

0.25

0.50

0.75

1.00

1.25

******

*

ER

Ca2+

con

tent

(a.

u.)

3 h 3 h

Thapsigargin

Thapsigargin

NT2 ρ+ cells NT2 ρ0 cells

***

(a) (b)

(c)

(e)

(d)

Fig. 1 PrP106–126 induces the decrease of ER Ca2+ content in mtDNA-

depleted q0 cells, as well as in the parental NT2 q+ and cells. The

fluorescence ratio, at 340/380 nm, of the Ca2+ sensitive probe Fura-2

acetoxymethyl ester was monitored during 12 min, in the absence of

external Ca2+, in untreated or PrP106–126-treated NT2 q+ and q0 cells.

After 2 min, 2.5 lmol/L thapsigargin was added to deplete ER Ca2+

levels. Representative F340/F380 traces of NT2 q+ treated with

PrP106–126 for 3 h (a) or 24 h (b) and NT2 q0 treated with PrP106–126 for

3 h (c) or for 24 h (d) and corresponding controls are presented. The

peak amplitude of Fura-2 fluorescence was used to evaluate ER Ca2+

content and data from at least three different experiments were

expressed as the normalization relative to NT2 q+ control values (e).

Data were expressed as the mean ± SEM. *p < 0.05; ***p < 0.001

significantly different when compared with the respective (q+ or q0)

untreated cells.

� 2007 The AuthorsJournal Compilation � 2007 International Society for Neurochemistry, J. Neurochem. (2008) 104, 766–776

Mitochondria-dependent cell death as a result of ER stress by PrP106–126 | 769

were higher in PrP106–126-treated cells when compared withthe control. These data demonstrate that intracellular Ca2+

levels, associated with release from internal stores, areincreased in NT2 q+ cells 24 h after PrP106–126 addition,while in NT2 q0 this rapid increment in cytosolic Ca2+ levelsis observed after 3 h incubation. Furthermore, we alsoobserved that basal values of Fura-2 fluorescence, in theabsence of PrP106–126 treatment, were higher in NT2 q0 cellsthan in NT2 q+ cells (Fig. 1b and d). These results suggestthat the absence of functional mitochondria per se, compro-mises the cellular Ca2+ buffering capacity.

Endoplasmic reticulum Ca2+ content was analysed by thecalculation of the peak over baseline, obtained uponthapsigargin treatment, and the results normalized to ERCa2+ content of NT2 q+ control values. In this cell line,PrP106–126 (25 lmol/L) induced a significant decrease in theER Ca2+ content: 85.2% decrease at 3 h (p < 0.001) and76.5% decrease at 24 h (p < 0.001) (Fig. 1e). In NT2 q0cells, PrP106–126 also induced a significant decrease in theER Ca2+ content: 90.6% decrease at 3 h (p < 0.001) and73.6% decrease at 24 h (p < 0.001) (Fig. 1e). NT2 q+ cellsare able to slightly recover ER Ca2+ content after theincubation with PrP106–126 for 24 h, when compared withdata at 6 h of incubation. Differences in ER Ca2+ contentbetween NT2 q+ and q0 cells exposed to PrP106–126 werenot statistically different, demonstrating that PrP106–126-induced ER Ca2+ release is not affected in the absence offunctional mitochondria. In addition, it was possible toobserve that, in the absence of PrP106–126-treatment, ERCa2+ content in NT2 q0 cells was by about 20% lower incomparison with that determined in the parental cells(Fig. 1e). These results are in accordance with the datapresented in Fig. 1b and d.

Treatment of NT2 q+ with PrP106–126 (25 lmol/L) induceda twofold increase in GRP78 levels at 6 h (p < 0.05) whencompared with untreated cells, which persisted at 24 h ofincubation. Similarly, in NT2 q0 cells, the same concentra-tion of PrP106–126 also induced an increase in GRP78 levelsat 6 h (p < 0.05) and at 24 h of incubation (p < 0.05)(Fig. 2), with no significant differences observed betweenNT2 q+ and q0 cells upon PrP106–126 treatment.

Furthermore, after 6 h incubation with PrP106–126(25 lmol/L) an almost twofold increase in caspase 4 activitywas observed both in NT2 q+ cells (p < 0.001) and in NT2q0 cells (p < 0.001). Similar results were obtained 24 h afterthe addition of PrP106–126 to NT2 q+ cells (p < 0.001) or inNT2 q0 cells (p < 0.05) (Fig. 3). No significant differencesin caspase 4 activation were observed between PrP106–126-treated NT2 q+ and q0 cells, both at 6 or 24 h.

Therefore, from the data obtained in the present study, wecan state that the synthetic PrP peptide PrP106–126 induces ERstress both in the presence and in the absence of functionalmitochondria, activating ER pathways that can potentiallylead to apoptotic cell death.

PrP106–126 induces apoptosis in parental NT2 q+ but not inmtDNA depleted q0 cellsIn order to investigate whether PrP106–126 activates apoptoticcell death as a result of ER stress in the absence of functionalmitochondria, cellular viability and apoptotic features wereevaluated in mtDNA-depleted q0 cells, and also in theparental q+ cell line, upon exposure to PrP106–126. In controland PrP106–126-treated cells the activities of caspase 9 and 3,an initiator and an effector caspase, respectively, involved inapoptosis (Kilic et al. 2002), was determined by measuringthe degradation of the chromogenic substrates Ac-LEHD-pNA and Ac-DEVD-pNA. The levels of a PARP fragment

Control

PrP106–126

6 h

PrP106–126

24 h

* **

*

GRP78 78 kDa

0

50

100

150

200

250

GR

P78

leve

ls (%

of

cont

rol)

NT2 ρ+ cells NT2 ρ0 cells

Fig. 2 PrP106–126 increases the levels of the ER chaperone GRP78 in

mtDNA-depleted q0 cells and in q+ cells. Total cell extracts were ob-

tained from NT2 q+ or q0 cells treated with PrP106–126 (25 lmol/L) for 6

or 24 h and the levels of GRP78 were determined by western blotting,

using a specific antibody. A representative immunoblot is depicted.

Data from at least three different experiments were analysed by

densitometry (graphs) and were expressed as the mean ± SEM.

*p < 0.05 significantly different when compared with the respective

(q+ and q0) untreated cells.

Control

PrP106–126

***

**** ***

0.0

0.5

1.0

1.5

2.0

2.5

6 h 6 h24 h 24 h

Ac-

LE

VD

-pN

A c

leav

age

(a.u

.)NT2 ρ+ cells NT2 ρ0 cells

Fig. 3 PrP106–126 activates caspase 4 in mtDNA-depleted cells and in

parental cells. NT2 q+ and q0 cells were incubated for 6 or 24 h with

PrP106–126 peptide (25 lmol/L). Protein extracts obtained from both

untreated and treated cells were used to evaluate caspase 4-like

activity, by measuring at 405 nm, the cleavage of the substrate

Ac-LEVD-pNA, as described in Materials and methods section. The

results, expressed as the increase above control values, are the

mean ± SEM of values corresponding at least to three experiments,

each value being the mean of duplicate assays. *p < 0.05;

***p < 0.001 significantly different when compared with the respective

(q+ or q0) untreated cells.

Journal Compilation � 2007 International Society for Neurochemistry, J. Neurochem. (2008) 104, 766–776� 2007 The Authors

770 | E. Ferreiro et al.

that results from caspase 3 cleavage (Nicholson et al. 1995)was analysed by western blotting and DNA fragmentationwas evaluated by the TUNEL assay. Furthermore, cellviability was measured by the MTT colorimetric assay. Toconfirm that ER stress is involved in PrP-induced cell deathin NT2 q+ cells, dantrolene, which inhibits the release ofCa2+ from ER through ryanodine receptors (RyR), was addedbefore PrP106–126 treatment.

In NT2 q+ cells, a significant increase in caspase 9- and 3-like activities was observed after 6 or 24 h PrP106–126addition (p < 0.001) (Figs 4a and 5a). Furthermore, whenNT2 q+ cells were treated with PrP106–126 for 24 h, in thepresence of dantrolene, a significant decrease in caspase 9-and 3-like activities was observed, demonstrating that RyR-mediated ER Ca2+ release is involved in caspase activationtriggered by PrP peptide (Figs 4b and 5b). In contrast to theresults obtained with the NT2 q+ cells, PrP106–126 peptidewas not able to induce caspase 9 and 3 activation in NT2 q0cells (Figs 4a and 5a). Caspase 9- and 3-like activities inPrP106–126-treated q+ and q0 cells were significantly different(p < 0.001) both at 6 and 24 h. PrP106–126 also increased the

***

Control

NT2 ρ+ cells NT2 ρ0 cells

6 h 6 h24 h 24 h

0

1

2

3***

#

###

0

1

2

3***

##

Control

PrP106–126

PrP106–126

PrP106–126

+ dantrolene

Ac-

LE

HD

-pN

A c

leav

age

(a.u

.)

Ac-

LE

HD

-pN

A c

leav

age

(a.u

.)

(a)

(b)

Fig. 4 PrP106–126 activates caspase 9 in parental NT2 q+ cells but not

in NT2 q0 cells and ER Ca2+ release is involved in PrP106–126-induced

caspase 9 activation. (a) NT2 q+ and q0 cells were incubated for 6 or

24 h with PrP106–126 peptide (25 lmol/L) and caspase 9-like activity

was determined. (b) NT2 q+ cells were treated for 24 h with PrP106–126

(25 lmol/L), in the absence or in the presence of dantrolene (10 lmol/

L). The activity of caspase 9 was determined measuring the cleavage

of the chromogenic substrate Ac-LEHD-pNA at 405 nm, as described

in Materials and methods section. The results, expressed as the in-

crease above control values, are the mean ± SEM of values corre-

sponding at least to three experiments, each value being the mean of

duplicate assays. ***p < 0.001 significantly different when compared

with untreated NT2 q+ cells and #p < 0.05, ##p < 0.01 and###p < 0.001 significantly different when compared with PrP106–126-

treated NT2 q+ cells.

Control

PrP106–126

6 h

PrP106–126

24 h

0

1

2

***

***

##

Cleaved PARP

NT2 ρ+ cells NT2 ρ0 cells

89 kDa

Cle

aved

PA

RP

(a.u

.)

Fig. 6 PARP cleavage occurs in NT2 q+ after incubation with PrP106–

126 but is not observed in treated q0 cells. Total cell extracts were

obtained from NT2 q+ and q0 cells treated with PrP106–126 (25 lmol/L)

for 6 or 24 h and from respective controls. The levels of a PARP

fragment (89 kDa) resulting from caspase 3 cleavage were deter-

mined by western blotting. Data from at least three different experi-

ments were analysed by densitometry and represented as graphics

and were expressed as the mean ± SEM. ***p < 0.001 significantly

different when compared with untreated NT2 q+ cells. #p < 0.05 sig-

nificantly different when compared with PrP106–126-treated NT2 q+

cells.

Control

***

***

0.0

0.5

1.0

1.5

2.0

2.5

######

NT2 ρ+ cells NT2 ρ0 cells

6 h 6 h24 h 24 h

***

#

0.0

0.5

1.0

1.5

2.0

2.5

Control

Ac-

DE

VD

-pN

A c

leav

age

(a.u

.)

Ac-

DE

VD

-pN

A c

leav

age

(a.u

.)

PrP106–126

PrP106–126

PrP106–126

+ dantrolene

(a)

(b)

Fig. 5 Activation of caspase 3 is induced by PrP106–126 in parental

NT2 q+ cells but not in NT2 q0 cells and dantrolene prevents caspase

3 activation induced by PrP106–126. (a) NT2 q+ and q0 cells treated for

6 and 24 h with PrP106–126 (25 lmol/L). (b) NT2 q+ cells were incu-

bated for 24 h with PrP106–126 (25 lmol/L) in the absence or in the

presence of dantrolene (10 lmol/L). Caspase 3-like activity was

determined in protein extracts by measuring the cleavage of the

chromogenic substrate Ac-DEVD-pNA at 405 nm, as described in

Materials and methods section. The results, expressed as the

increase above control values, are the mean ± SEM of values corre-

sponding at least to three experiments, each value being the mean of

duplicate assays. ***p < 0.001 significantly different when compared

with untreated NT2 q+ cells and #p < 0.05 and ###p < 0.001 signifi-

cantly different when compared with PrP106–126-treated NT2 q+ cells.

� 2007 The AuthorsJournal Compilation � 2007 International Society for Neurochemistry, J. Neurochem. (2008) 104, 766–776

Mitochondria-dependent cell death as a result of ER stress by PrP106–126 | 771

levels of a caspase 3-mediated PARP cleavage fragment with89 kDa: an almost twofold increase was observed at 6 h(p < 0.001) and at 24 h, in NT2 q+ cells (p < 0.001)(Fig. 6). However, under the same conditions, PrP106–126did not affect the levels of the PARP cleavage product inNT2 q0 cells (Fig. 6). Differences in the levels of the PARPcleavage product observed between NT2 q+ and q0 cellstreated with PrP106–126 were statistically different (p < 0.05).

In PrP106–126-treated NT2 q+ cells, an approximatelyseven- and tenfold increase in the number of TUNEL-positivecells was observed after 6 and 24 h, respectively, when

compared with control cells (p < 0.05 and < 0.01 respec-tively) (Fig. 7c). Furthermore, when NT2 q+ cells weretreated for 24 h with PrP106–126 in the presence of dantrolene,a significant decrease in the number of TUNEL-positive cellswas observed, demonstrating that ER Ca2+ release throughRyR is involved in PrP-induced cell death (Fig. 7a and d). InPrP106–126-treated NT2 q0 cell line, TUNEL-positive cellswere almost absent (Fig. 7a and d), correlating with the dataobtained before for caspase 9- and 3-like activities and PARPcleavage (Figs 4, 5 and 6). In addition, it is necessary to statethat the number of TUNEL-positive cells in PrP106–126-treatedNT2 q+ cells is low, because of the fact that cells were plated

NT2 ρ+

Control

Control

PrP106–126

NT2 ρ0

MergedTUNELPhase contrast

PrP106–126

PrP106–126

dantrolene

0

1

2

3

4

5

6 ***

#####

Control

Num

ber

of T

UN

EL

-pos

itive

cel

ls(%

of

tota

l num

ber

of c

ells

)

NT2 ρ+ cells

PrP106–126 + dantrolene

PrP106–126

NT2 ρ0 cells

0

25

50

75

100

125

*

###

Control

0

1

2

3

4

5

6

*

**

Control

0

25

50

75

100

125

*

Tot

al n

umbe

r of

cel

ls

###

Control

Num

ber

of T

UN

EL

-pos

itive

cel

ls(%

of

tota

l num

ber

of c

ells

)

NT2 ρ+ cells

PrP106–126 24 h

PrP106–126 6 hPrP106–126

NT2 ρ+ cells NT2 ρ0 cells

Control

(a)

(b)

(d)

(c)

0

25

50

75

100

125

*** Control

PrP106–126

###

NT2 ρ+ cells NT2 ρ0 cells M

TT

red

uctio

n (%

of

cont

rol)

Fig. 8 PrP106–126 peptide affects cell survival in NT2 parental q+ but

not in mtDNA-depleted q0 cells. Twenty-four hours after PrP106–126

(25 lmol/L) addition, cellular viability was evaluated in control or

treated NT2 q+ or q0 cells by measuring the ability to reduce the

tetrazolium salt MTT. The results, expressed as the percentage of the

absorbance determined in control cells, are the mean ± SEM of values

corresponding at least to three experiments, each value being deter-

mined in duplicate. ***p < 0.001 significantly different when compared

with untreated NT2 q+ cells and ###p < 0.001 significantly different

when compared with PrP106–126-treated NT2 q+ cells.

Fig. 7 The number of TUNEL-positive apoptotic cells is increased by

PrP106–126 treatment in the NT2 q+ cell line, but not in q0 cells and

dantrolene protects against PrP-induced apoptosis. TUNEL assay was

performed as described in Materials and methods section. (a) Rep-

resentative phase contrast, fluorescence (TUNEL-positive cells are

green fluorescent) and merged images of untreated and treated NT2

q+ cells with PrP106–126 (25 lmol/L) in the absence or the presence of

dantrolene (10 lmol/L) and untreated and treated NT2 q0 cells with

PrP106–126 (25 lmol/L), for 24 h (400· magnification). (b) Total num-

ber of untreated or PrP106–126-treated NT2 q+/q0 cells. (c) Number of

TUNEL-positive cells of untreated or PrP106–126-treated NT2 q+ for 6

and 24 h. (d) Number of TUNEL-positive cells of untreated and treated

NT2 q+ cells with PrP106–126 (25 lmol/L) in the absence or the pres-

ence of dantrolene (10 lmol/L) and untreated and treated NT2 q0 cells

with PrP106–126 (25 lmol/L), for 24 h. Results correspond to values in

three microscope fields, in duplicate, from at least three independent

experiments. *p < 0.05, **p < 0.01 and ***p < 0.001 significantly dif-

ferent when compared with untreated NT2 q+ cells and ##p < 0.01 and###p < 0.001 significantly different when compared with PrP106–126-

treated NT2 q+ cells.

Journal Compilation � 2007 International Society for Neurochemistry, J. Neurochem. (2008) 104, 766–776� 2007 The Authors

772 | E. Ferreiro et al.

on slides that have no support and that dying cells afterPrP106–126 treatment were detaching from slides and thereforewere impossible to be fixed and counted. Counting the totalnumber of cells present in treated or untreated slides showedthat 30% of NT2 q+ cells detached upon PrP106–126 treatment(p < 0.05). Although the number of total NT2 q0 cells islower than NT2 q+ cells, probably because of different celldivision velocities between the two cell lines, no differenceswere observed between untreated and PrP106–126-treated NT2q0 cells (Fig. 7b).

Finally, after 24 h incubation, PrP106–126 decreased cellviability by about 20% in NT2 q+ (p < 0.001), but had noeffect on NT2 q0 cell survival (Fig. 8). Both inMTTand in theTUNEL assays, significant differences were observed betweenNT2 q+ and q0 cells treated with PrP106–126 (p < 0.001).

These results show that in parental NT2 q+ cells,PrP106–126 induces ER stress-mediated apoptosis, whichoccurs after ER Ca2+ depletion, because of ER Ca2+ releasefrom RyR, and activation of ER stress. However, inPrP106–126-treated NT2 q0 cells, in which mtDNA, thatcodifies several proteins of the mitochondrial ETC, has beendepleted, apoptosis is not able to proceed although ER Ca2+

content is decreased and stress is induced. Altogether thepresented results demonstrate that ER stress-mediatedapoptosis induced by the PrP peptide requires the presenceof functional mitochondria.

To elucidate whether the absence of toxicity of PrP106–126in the NT2 q0 cell line is related to the endogenous levels ofPrPC, western blotting was performed. In untreated NT2 q0cells, PrPC levels were significantly higher then in untreatedNT2 q+ cells (Fig. 9). Furthermore, as an attempt to identifya possible mechanism that could contribute to the resistanceof NT2 q0 cells to toxic insults, such as PrP106–126 peptideexposure, the levels of two members of the Bcl-2 family, thepro-apoptotic protein Bax and the anti-apoptotic protein Bcl-2, were evaluated by western blotting. As depicted inFig. 10a, no differences were observed in Bax levels betweenthe two cell lines. However, a significant increase in Bcl-2levels occurred in the NT2 q0 cells, when compared with theNT2 q+ cells (Fig. 10b). As a consequence, a higher Bcl-2/Bax ratio was observed in NT2 q0 cells relatively to theparental cells (Fig. 10c).

Discussion

Recently, we have reported that, in primary cultures ofcortical neurons, PrP106–126 induces ER stress, which isfollowed by a significant increase in the cytosolic Ca2+ levels,cytochrome c release from mitochondria and activation ofseveral caspases, finally leading to apoptotic cell death(Ferreiro et al. 2006). Taken together, these data suggest thatmitochondria are involved in ER stress-mediated apoptoticcell death triggered by PrP106–126. In order to clarify the roleof mitochondria in apoptosis induced by PrP peptide, we usedNtera2 (NT2) human teratocarcinoma cell line that had beendepleted of their mtDNA and therefore are characterized bythe absence of functional mitochondria, termed NT2 q0 cells,and their parental NT2 cells (q+ cell line). mtDNA depletedNT2 q0 cells and the parental NT2 q+ cells had been used inour laboratory to clarify the role of mitochondria in amyloid-beta (Ab) peptide-induced cell death. Results obtained byCardoso et al. (2001, 2002) clearly showed that functionalmitochondria are required for Ab-induced apoptosis. As Aband PrP peptides share several structural and toxic charac-teristics, we decided to use cells lacking functional mito-chondria to investigate the role of this organelle in cell deathas a result of ER stress induced by PrP106–126.

As they lack mtDNA, NT2 q0 cells cannot produce afunctional ETC, because 13 protein subunits of mitochon-

050

100150200250300350400450

** **

)lortnocf o

% (s level

xaB

NT2 ρ+ cells

NT2 ρ0 cells

050

100150200250300350400450

)lortnocfo

%(slevel

2-lcB

050

100150200250300350400450

xaB/2-lc

Boita

R

Bax 27 kDa20 kDa Bcl-2

(a) (b) (c)

Fig. 10 Bcl-2/Bax ratio is increased in the NT2 q0 cell line in com-

parison with the q+ cells. Total cell extracts were obtained from

untreated NT2 q+ or q0 cells and the levels of Bax and Bcl-2 were

determined by western blotting, using specific antibodies. Represen-

tative immunoblots are presented. Data from at least three different

experiments were analysed by densitometry (graphs) and were

expressed as the mean ± SEM. Bcl-2/Bax ratio was calculated and

expressed as the mean ± SEM. **p < 0.01 significantly different when

compared with NT2 q+ untreated cells.

0

100

200

300

400 *

NT2 ρ+ cells

NT2 ρ0 cells

PrPc 33–35 kDa

PrPc le

vels

(%

of

cont

rol)

Fig. 9 PrPc levels are increased upon mtDNA depletion. Total cell

extracts were obtained from untreated NT2 q+ or q0 cells and the

levels of PrPC were determined by western blotting. Representative

immunoblots are presented. Data from at least three different experi-

ments were analysed by densitometry (graphs) and were expressed

as the mean ± SEM. *p < 0.05 significantly different when compared

with NT2 q+ untreated cells.

� 2007 The AuthorsJournal Compilation � 2007 International Society for Neurochemistry, J. Neurochem. (2008) 104, 766–776

Mitochondria-dependent cell death as a result of ER stress by PrP106–126 | 773

drial respiratory apparatus are mtDNA encoded (sevensubunits of NADH-Q reductase, one subunit of cyto-chrome c reductase, three subunits of cytochrome coxidase and two subunits of ATP synthetase) (Wallace1994; Jiang et al. 1999). Two rRNAs and 22tRNAs thatare necessary for the expression of the 13 structuralmtDNA genes are also encoded by mtDNA (Mosmann1983). NT2 q0 cells have no changes in mitochondrialmembrane potential (Dwm), although ETC is non-func-tional, and are still able to undergo apoptosis afterexposure to staurosporine (Jiang et al. 1999). The main-tenance of Dwm can be because of the transport of protonsout of the mitochondrial matrix, caused by the reversal ofATP synthase activity (Nijtmans et al. 1995).

The results presented in this report highlight the role ofmitochondria in ER stress-mediated apoptosis induced by thePrP peptide. The synthetic PrP106–126, which mimics PrPSc

toxicity (Della-Bianca et al. 2001; Perez et al. 2003) wasshown to induce ER stress in NT2 q+ (Ntera2 humanteratocarcinoma cell line) and in q0 cells, which had beendepleted of their mtDNA. In both cell lines, ER stress wasinduced through the disturbance of Ca2+ homeostasis (ERCa2+ levels were depleted, as shown in Fig. 1) and theactivation of the unfolded protein response (Fig. 2 shows theincrease of GRP78 levels), leading to the increase in theactivity of the ER-resident apoptosis-associated caspase 4(Fig. 3). The observed perturbation of Ca+ homeostasis canbe because of the early formation of membranar PrP Ca2+

channels (Lin et al. 1997), a death signal that can beextended to the ER, leading to ER Ca2+ release through theRyR, amplifying cytosolic Ca2+ content. Furthermore, ERstress-mediated apoptosis triggered by the PrP peptide wasobserved in the NT2 q+ cells. In parental NT2 cell line,PrP106–126 was demonstrated to increase caspase 9 and 3activity (Figs 4 and 5), PARP cleavage (Fig. 6) and DNAfragmentation (Fig. 7) and also to reduce cell viability(Fig. 8), by a mechanism involving the release of Ca2+

through ER RyR (Figs 4b and 5b; Fig. 7a and d). However,in PrP106–126-treated NT2 q0 cells, apoptosis was not able toproceed, as activation of caspase 9 and 3 (Figs 4a and 5a),PARP cleavage (Fig. 6) and DNA fragmentation were notobserved (Fig. 7a and d), although ER stress was beinginduced. In addition, cell viability was not compromisedupon PrP106–126 addition in NT2 q0 cells (Fig. 8). Toevaluate whether the lack of PrP106–126 toxicity in NT2 q0cells could be a result of altered expression of the endo-genous cellular form of PrP, PrPC levels were measured inboth untreated NT2 q0 and parental NT2 q+ cell lines. PrPC

expression was shown to be increased in NT2 q0 cells whencompared with the parental cell line (Fig. 9). This differencecan be involved in the decreased susceptibility of mtDNA-depleted NT2 q0 cells when compared with the parental cellline and can be related to the protective function of PrPC (Huet al. 2007).

Because of the considerable structural similarity betweenPrPC and Bcl-2 family members, it has been suggested thatPrPC might function as an anti-apoptotic protein (Roucou andLeBlanc 2005). Indeed, it was demonstrated that PrPC

protects human neurons against mitochondrial-mediated celldeath induced by Bax (Bounhar et al. 2001). Finally, thelevels of Bax and Bcl-2 were analysed in untreated NT2 q+and q0 cells, to get better insight into the mechanismsinvolved in the resistance of mtDNA-depleted NT2 q0 cellsagainst PrP106–126 toxicity. We have observed that a fourfoldincrease in Bcl-2 levels occurs in comparison with theparental cell line (Fig. 10b), while Bax levels are unchangedupon depletion of mtDNA (Fig. 10b). These results suggestthat the enhanced expression of the anti-apoptotic proteinBcl-2, a mitochondrial membrane protein that was shown toblock apoptosis (Hockenbery et al. 1990), may be one of themolecular mechanisms involved in the resistance of NT2 q0cells to PrP106–126-induced cell death.

Previous studies have revealed the involvement of ERstress in neuronal death that occurs in PrD. In fact, Hetz et al.

ER

GRP78

ER stress

Mitochondria

?

Caspase-4

Cytochrome c

Caspase-9

Caspase-3

PARP cleavage

NucleusDeath

PrP ?

Ca2+ Ca2+

Fig. 11 Model of PrP peptide-induced cell

death. PrP induces ER stress through the

perturbation of Ca2+ homeostasis and acti-

vation of the unfolded protein response,

subsequently increasing the activity of the

ER-resident caspase 4. Ca2+ released from

ER can enter mitochondria and disturb its

functioning, leading to cytochrome c

release and activation of caspase 9 and 3.

Activated caspase 3 leads to PARP cleav-

age, which translocates to the nucleus,

finally leading to cell death.

Journal Compilation � 2007 International Society for Neurochemistry, J. Neurochem. (2008) 104, 766–776� 2007 The Authors

774 | E. Ferreiro et al.

(2003) refer that caspase 12 activation is detected in cellstreated with PrPSc and that GRP58, a chaperone with proteindisulphide isomerase-like activity, protects cells againstPrPSc toxicity and decreases the rate of caspase 12 activation(Hetz et al. 2005). Increased activity of caspase 4, the humanhomologue of caspase 12 in rodent, has been associated tocaspase 3 activation (Kim et al. 2006). However, theinvolvement of this ER-resident caspase in apoptosis is stillcontroversial. Obeng and Boise (2005) showed that caspase 4activation was not required for the ER stress apoptoticpathway, but pointed to the requirement of caspase 9activation. In fact, we were not able to demonstrate theinvolvement of caspase 4 in PrP-induced cell death, ascaspase 3 activation was not observed in the NT2 q0 cellsdespite caspase 4 is activated. More recently, cells withdamaged ER were shown to be more susceptible to prionreplication and as PrPSc induces ER stress, a vicious cycle isimplicated (Hetz et al. 2007). In addition, it was reported thatER stress conditions impair PrP cotranslational translocationand favour the accumulation of cytosolic species of PrP,which can be potentially toxic (Orsi et al. 2006). These dataconfirm the importance of ER dysfunction in PrD (reviewedin Castilla et al. 2004; Hetz and Soto 2006). The questionthat remained to answer was whether and how ER stresssubsequently leads to apoptotic death in cells exposed to thePrP peptide. Our previous studies in primary neuronalcultures demonstrated that PrP106–126 induces ER stressthrough the release of Ca2+ from ER, leading to mitochon-drial cytochrome c release and activation of caspase-depen-dent apoptotic cell death (Ferreiro et al. 2006). A study by DiSano et al. (2006) showed that ER stress-induced apoptosisis apoptosome dependent. When Apaf-1 deficient cells wereexposed to ER stress inducers, like tunicamycin, thapsigarginor brefeldin A, ER stress was induced, but these cells werenot able to die by apoptosis. More recently, Masud et al.(2007) presented results showing that ER-stress induceddeath depends on mitochondria and on the intrinsic apoptoticpathway. Our results are in accordance with these studies asin PrP106–126-treated NT2 q0 cells, ER stress occurs,although features of apoptosis are absent, implicatingmitochondria in cell death as a result of ER stress inducedby the PrP peptide. In conclusion, we propose that PrPinduces ER stress through the perturbation of Ca2+ homeo-stasis and also through activation of the unfolded proteinresponse pathway. In response to ER stress caspase 4 isactivated. Ca2+ release from ER can enter mitochondria anddisturb mitochondria functioning, leading to cytochrome crelease, caspase 9 and subsequent caspase 3 activation, PARPcleavage, DNA fragmentation and finally cell death(Fig. 11).

Altogether, our results demonstrate that functional mito-chondria are required for ER stress-mediated apoptosisinduced by the PrP106–126 peptide and reinforce the hypoth-esis that both organelles cooperate in PrP-induced apoptosis.

Acknowledgements

The authors greatly thank Professor Russel Swerdlow for the

generous gift of the NT2 q0 cell line and Isabel Nunes (PhD) for the

handling of NT2 q+ and NT2 q0 cell lines. This work was supportedby FCT (Portuguese Research Council) Project no. POCTI/36101/

NSE/2000. The PhD students EF and RC have fellowships from

FCT (Grant no.SFRH/BD/14108/2003 and SFRH/BD/28403/2006

respectively).

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