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Toxicology 290 (2011) 230–240

Contents lists available at SciVerse ScienceDirect

Toxicology

journa l homepage: www.e lsev ier .com/ locate / tox ico l

echanism of Alternariol monomethyl ether-induced mitochondrial apoptosis inuman colon carcinoma cells

atma Bensassi a,b, Cindy Gallernec, Ossama Sharaf el deinc, Mohamed Rabeh Hajlaouib,assen Bachaa,∗, Christophe Lemairec,d

Laboratory for Research on Biologically Compatible Compounds, Faculty of Dentistry, Rue Avicenne, 5019 Monastir, TunisiaLaboratory of Plant Protection, The National Institute for Agricultural Research, INRA Tunisia, Rue Hedi Karray, 2049 Ariana, TunisiaINSERM U769, Signalisation et Physiopathologie Cardiaque, Faculté de Pharmacie, Châtenay-Malabry, FranceUniversité de Versailles St. Quentin, Versailles 78035, France

r t i c l e i n f o

rticle history:eceived 10 August 2011eceived in revised form8 September 2011ccepted 30 September 2011vailable online 6 October 2011

eywords:ME

a b s t r a c t

Alternariol monomethyl ether (AME) is a major mycotoxin produced by fungi of the genus Alternariaand a common contaminant of food products such as fruits and cereals worldwide. AME can cause seri-ous health problems for animals as well as for humans. In this study, human colon carcinoma cells(HCT116) were used to explore the mechanisms of cell death induced by AME. Exposure of HCT116cells to AME resulted in significant cytotoxicity manifested by a loss in cell viability mainly mediatedby activation of apoptotic process. AME activated the mitochondrial apoptotic pathway evidenced bythe opening of the mitochondrial permeability transition pore (PTP), loss of the mitochondrial trans-membrane potential (��m) downstream generation of O2

•−, cytochrome c release and caspase 9 and

poptosisitochondria

TP�m

3 activation. Experiments conducted on isolated organelles indicated that AME does not directly targetmitochondria to induce PTP-dependent permeabilization of mitochondrial membranes. Moreover, no dif-ference was observed in Bax-KO cells in comparison to parental cells, suggesting that the pro-apoptoticprotein Bax is not involved in AME-induced mitochondrial apoptosis. Our findings demonstrate for thefirst time that AME induces cell death in human colon carcinoma cells by activating the mitochondrialpathway of apoptosis.

. Introduction

A great number of Alternaria species are of a major concernecause of their toxigenic properties. They can lead, under suit-ble conditions, to the production of a range of metabolites, some ofhich are powerful mycotoxins (Woody and Chu, 1992) with muta-

enic (Schrader et al., 2001) and teratogenic properties (Griffin andhu, 1983) and have been linked to certain forms of cancer (Viscontind Sibilia, 1994; Scott, 2001).

Literature on the occurrence of Alternaria mycotoxins showshat a wide range of Alternaria metabolites have been describedlthough relatively few have been fully characterized. Among theajor Alternaria mycotoxins, the Alternariol monomethyl ether,

lso known as AME, has been shown to occur naturally (Panigrahi,997). AME is a member of the dibenzo-�-pyrone group of

ycotoxins which include Alternariol (AOH) and Altenuene (ALT)

Bottalico and Logrieco, 1998). The natural occurrence of AME haseen reported in several cereal grains all over the world such as

∗ Corresponding author. Tel.: +216 73 42 55 50; fax: +216 73 42 55 50.E-mail address: hassen.bacha@fmdm.rnu.tn (H. Bacha).

300-483X/$ – see front matter © 2011 Elsevier Ireland Ltd. All rights reserved.oi:10.1016/j.tox.2011.09.087

© 2011 Elsevier Ireland Ltd. All rights reserved.

wheat, barley and sorghum (Li and Yoshizawa, 2000; Medina et al.,2006; Webley et al., 1997).

There are some reports on the mutagenicity and carcinogenicityof AME and their relevance to the etiology of human oesophagealcancer (An et al., 1989; Liu et al., 1992; Davis and Stack, 1994;Yekeler et al., 2001; Schrader et al., 2006). AME might cause cellmutagenicity and transformation. Indeed, it was demonstrated tocombine with the DNA isolated from human foetal oesophagealepithelium, activate oncogens and promote proliferation of humanfoetal oesophageal epithelium in vitro. Furthermore, AME appearsto exhibit a pronounced genotoxicity in most assays (Marko, 2007;Pfeiffer et al., 2007). This toxin showed also cytotoxic activity tobacterial and mammalian cells and fetotoxicity and teratogenicityto mice and hamsters (Scott and Stoltz, 1980; Visconti and Sibilia,1994).

At the cellular level, the presence of mycotoxins induces numer-ous biochemical mechanisms including temporary modificationsin gene expressions. There is a widespread interest in the cellu-

lar mechanisms used by organisms to deal with the disruption inhomeostasis. Apoptosis plays a critical role in the normal devel-opment and maintenance of homeostasis in organisms and alsoin toxicants-induced cell death (Meier et al., 2000; Dunn and Weis,

cology

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F. Bensassi et al. / Toxi

009). It is a genetically directed process of cell self-destruction thats marked by a series of morphological and biochemical changes,ncluding chromatin condensation, fragmentation of nuclear DNAnd caspase activation.

In general, the apoptotic process can be subdivided into twoategories, the extrinsic apoptotic pathway initiated by ligandngagement of cell surface receptors such as Fas and TNF receptorsRowinsky, 2005; Fulda and Debatin, 2006), and the intrinsic (mito-hondrial) pathway activated by signals emanating from cellularamage sensors or other types of severe cell stress. This pathway

nvolves the release of pro-apoptotic proteins from mitochondrialntermembrane space that in turn activate caspase proteases. Thentrinsic apoptotic pathway relies on the balance between prond anti-apoptotic members of the Bcl-2 super-family of proteinshich act to regulate the mitochondrial membrane permeabi-

ization (MMP). It is generally accepted that the MMP, a highlyegulated phenomenon, could be mediated by the formation ofax oligomeres or opening of a high-conductance channel namedhe permeability transition pore (PTP) (Leung and Halestrap, 2008;icchelli et al., 2011). Thus, mitochondria play a pivotal role inpoptosis by coordinating caspase activation through the releasef apoptogenic factors such as the cytochrome c (Desagher andartinou, 2000).The cytotoxic effects of different mycotoxins, such as Ochra-

oxin A, Patuline, Fumonisin B1, Zearalenone have been reportedo rely on their ability to trigger an apoptotic process (Dragan et al.,001; Kamp et al., 2005; Wu et al., 2008; Chatti Gazzah et al., 2010).ompared with other mycotoxins, knowledge of the toxicology oflternaria mycotoxins is limited. In particular, the nature of celleath induced in AME-exposed cells has not yet been elucidated.

n this study, we have performed our experiments on HCT116 cells,erived from a human colon carcinoma, as alimentary intoxication

s the main route of exposure to mycotoxins contamination.In this work, we have investigated the cellular and molecu-

ar events of AME-induced cell death in HCT116 cells. This workrovides first evidence that the cytotoxicity of AME is related tohe induction of a p53-dependent activation of the mitochondrialathway of apoptosis, and demonstrates the anticancer property ofME.

. Materials and methods

.1. Chemicals

AME (3,7-dihydroxy-9-methoxy-1-methyl-6H-dibenzo[b,d] pyran-6-one) andonidamine (1-[(2,4-dichlorophenyl)methyl]-1H-in-dazole-3-carboxylic acid)]LND) were purchased from Sigma–Aldrich. Dulbecco’s modified eagle medium-12 (DMEM-F12), foetal bovine serum (FBS), phosphate buffer saline (PBS),rypsin–EDTA, penicillin and streptomycin mixture were from Invitrogen. N-acetylysteine (NAC), nitro blue tetrazolium (NBT) and 5-bromo-4-chloro-3-indolyl phos-hate disodium salt (BCIP) were from Sigma–Aldrich. Mouse monoclonal anti-p53,nti-Caspase 9 and the secondary antibody (phosphatase-conjugated) anti-mousemmunoglobulin were from Invitrogen. For immunofluorescence assay, the anti-ytochrome c antibody (mAb 6H2.B4) was from Pharmingen (BD Biosciences, Sanose, CA). Goat anti-mouse immunoglobulin G (IgG) conjugated with Alexa fluor 350

as from Invitrogen. 2′ ,7′-Dichlorodihydrofluorescein diacetate (DCFH-DA), pro-idium iodide (PI), fluorescein diacetate (FDA), 3,3′-dihexyloxacarbocyanin iododeDiOC6(3)), Hoechst 33348 and chloromethyl-X-rosamine (CMXRos), Calcein-AMC-3100), MitoSOXTM Red reagent and Hanks’ balanced salt solution (HBSS) wereupplied by Invitrogen. The colorimetric CaspACE assay kit (G-7351) was purchasedrom Promega (Madison, USA) and the ECL kit was from Amersham BiosciencesPiscataway, NJ). Z-Val-Ala-DL-Asp-Fluromethylketone (ZVAD-fmk, 100 �M) is aachem product. All other compounds were purchased from Sigma–Aldrich and allhe used chemicals were of analytical grade.

.2. Animals

C57BL/6J female, 6–12 weeks old, mice (Charles River, France) were used. Thenimals were maintained in a room with controlled illumination (lights on 7–21 h),emperature (26–28 ◦C) and humidity (30–70%) and were given free access to regular

ouse diet and water. Animals were fasted overnight before being killed by cere-ral dislocation. All procedures involving animals and their care were conducted in

290 (2011) 230–240 231

conformity with the institutional guidelines and in compliance with national andinternational laws and policies in the Animal Care Facility of the “Institut Fédératif deRecherche-IFR141” of the Paris-Sud University (Council directive # 87-848, October19, 1987, Ministère de l’Agriculture et de la Foret, Service Vétérinaire de la Santé etde la Protection Animale).

2.3. Cell culture and treatment

Human colorectal carcinoma cells HCT116 wild type (Bax+/−) and HCT116-Bax-KO (generously given by B. Vogelstein, Johns Hopkins University School of Medicine,Baltimore, MD) were maintained as monolayer culture in DMEM-F12, supplementedwith 10% FBS, 1% l-glutamine (200 mM), 1% of mixture penicillin (100 IU/ml) andstreptomycin (100 �g/ml), in a humidified incubator at 37 ◦C in an atmosphere of5% CO2 in air.

AME was dissolved in pure DMSO. To obtain the studied concentrations in thecell culture media, the mycotoxin treatment volume was negligible and representedabout 0.025% of the total medium volume. In these conditions, untreated cells andcells receiving this low vehicle volume responded in the same manner. For thisreason, we have chosen untreated cells as control.

2.4. Cell viability assay

The fluorescent probe fluorescein diacetate (FDA) has been used to assess cellviability of HCT116 cells. FDA, a non-fluorescent compound, is able to cross themembrane of living cells. Once inside the cell, esterases will cut the diacetate andfluorescein is released resulting in a measurable fluorescence signal that is directlyrelated to the viability of cells. At 50% confluence, cells were seeded in 24-wellmultidishes and were treated with different concentrations of AME for 24 h. Then,they were incubated for 5 min at 37 ◦C with FDA at 0.2 �g/ml. Finally, cells wereanalyzed by flow cytometry. Living cells are FDA positive, whereas dead cells areFDA negative. IC50 value is defined as the concentration inducing 50% loss of cellviability.

2.5. Cell death determination

To measure necrosis, unpermeabilized HCT116 cells were stained with propid-ium iodide (PI) after exposure to 120 �M AME for 24 h. As positive control, cells weretreated with 1 mM H2O2 during 24 h. Negative control corresponds to untreatedcells. PI (5 �g/ml) was added just before cytometric analysis. This impermeant dyeonly stains the DNA of dead cells with compromised plasma membrane, characteris-tic of late apoptotic/necrotic cells. Consequently, early apoptotic cells are PI negativewhile late apoptotic/necrotic cells are PI positive. During apoptosis, condensationand fragmentation of nuclear DNA occurred. The hypoploid cells (Sub-G1 popula-tion) showing a loss of DNA content can be recorded by flow cytometry. HCT116cells were exposed to AME at 120 �M for 48 h. Negative control corresponds tountreated cells. Cells were harvested and fixed with ice-cold 70% ethanol, storedat −20 ◦C overnight and stained with 50 �g/ml PI in the presence of 250 �g/ml ofRNAse A. For inhibition experiments, cells were pretreated for 2 h with 5 mM NAC.The percentage of cell in Sub-G1 was determined by flow cytometric analysis. A totalof 10,000 cells were analyzed per sample.

2.6. Determination of mitochondrial transmembrane potential (��m) andpermeability transition pore (PTP) opening

To measure the mitochondrial membrane potential, DiOC6(3) or CMXRos wereused. These fluorescent probes specifically accumulate in the mitochondria depend-ing on the ��m. For inhibition experiments, cells were pre-treated for 2 h witheither the caspase inhibitor ZVAD-fmk at 50 �M or with the antioxidant NAC at5 mM. After AME treatment, cells were stained with 100 nM of either DiOC6(3) orCMXRos for 30 min at 37 ◦C and 10,000 cells were analyzed by flow cytometry. PTPopening was assessed as previously described (Deniaud et al., 2008). At 50% conflu-ence, cells were incubated for 15 min at 37 ◦C with 1 �M Calcein-AM and 1 mM CoCl2in Hanks’ balanced salt solution (HBSS) supplemented with 1 mM HEPES, pH 7.3.Before AME treatment, Hanks’ solution was replaced by complete culture medium.

2.7. Effect of AME on purified mitochondria

In order to determine whether mitochondria are direct targets of AME, mito-chondria were isolated from mouse liver by differential centrifugations and purifiedon Percoll gradient according to Jacotot et al. (2001). All assays were performed in96-well plates at 37 ◦C in a spectrofluorimeter (TECAN genios; TECAN, Austria). Forswelling and depolarization measurements, 25 �g of mitochondrial proteins wasdiluted in a hypo-osmotic buffer (10 mM Tris-Mops, pH 7.4, 5 mM succinate, 200 mMsucrose, 1 mM Pi, 10 �M EDTA, 2 �M rotenone) in the presence or in the absence of

various concentrations of AME. The mitochondrial swelling was immediately mea-sured by the decrease in optical density (OD) at 540 nm. The depolarization of themitochondria was measured by the rhodamine 123 (1 �M) fluorescence dequench-ing assay (�exc: 485 nm, �em: 535 nm; Molecular Probes). Calcium, a well knowninducer of both depolarization and swelling, was used at increasing concentrations

2 cology 290 (2011) 230–240

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Fig. 1. Cytotoxic effect of AME on HCT116 cells. Cells were treated with AME atthe indicated concentrations for 24 h. Cell viability was determined using the FDA

32 F. Bensassi et al. / Toxi

from 6 to 50 �M) as positive control and cyclosporin A (CsA, a PTP inhibitor) wassed at 5 �M. Negative control corresponds to untreated mitochondria.

.8. Western blot analysis

At 50% confluence, cells treatment was performed with 120 �M of AME for dif-erent treatment times (6, 16 and 24 h). Then, cells were washed with PBS, harvestednd lysed in lysis buffer (0.5 M HEPES, 0.5% Nonidet-P40, 1 mM PMSF, 1 �g/ml apro-inin, 2 �g/ml leupeptin, pH 7.4). Equal amounts of proteins (30 �g) were separatedy 12% SDS-polyacrylamide gel electrophoresis and subsequently transferred ontoVDF membranes (Millipore, Billerica, MA). Membranes were incubated with therimary antibody, either anti-p53 (1:500 dilution) or anti-caspase 9 (1:500 dilution),vernight at 4 ◦C. After washing, blots were incubated for 1 h with the correspond-ng secondary antibody (1:3000). For caspase 9 cleavage, immunoreactive bands

ere detected by ECL kit (Amersham, Piscataway, NJ). Concerning the expressionf p53, the chromogenic substrates BCIP/NBT were added to stain immunoreactiveroteins. Protein levels were then determined by computer-assisted densitometricnalysis (Densitometer, GS-800, BioRad Quantity One).

.9. Measurement of reactive oxygen species (ROS) production

The intracellular amounts of ROS was measured using the fluorogenic freelyermeable tracer DCFH-DA which is deacylated by intracellular esterases to the non-uorescent compound DCFH and oxidized to the fluorescent compound DCF throughhe action of peroxides in the presence of ROS such as hydroxyl radical and especiallyydrogen peroxide (Le Bel et al., 1992; Gomes et al., 2005). At 50% confluence, cellsere treated with AME at 120 �M for 24 h. A positive control was treated withmM H2O2 during 24 h. Negative control corresponds to untreated cells. Cells wereentrifuged, washed with PBS and incubated with DCFH-DA at 10 �M in HBSS for0 min at 37 ◦C. DCF fluorescence was immediately analyzed by flow cytometry.o measure the relative levels of mitochondrial superoxide anion, O2

•− , the probeitoSOXTM was used. Once in the mitochondria, MitoSOXTM reagent is oxidized by

uperoxide anion and exhibits red fluorescence detected by flow cytometry. At 50%onfluence, cells were treated with AME at 120 �M for 24 h. A positive control wasreated with 1 mM H2O2 during 24 h. Negative control corresponds to untreatedells. Then, the cells were centrifuged, washed with PBS and incubated with 2 �Mf MitoSOXTM for 10 min at 37 ◦C. Mitosox fluorescence was immediately analyzedy flow cytometry.

.10. Caspase 3 activity assay

Caspase 3 activity was measured using a commercially available kit, accordingo the manufacturer’s instructions. At 50% confluence, HCT116 cells were culturedn the absence or the presence of AME at 120 �M for 6, 16 and 24 h at 37 ◦C. Cells

ere harvested and centrifuged at 5000 × g at 4 ◦C for 10 min. The cell pellet wase-suspended in lysis buffer and kept on ice bath for 30 min and then centrifugedt 5000 × g for 20 min, and supernatant was used for caspase-3 assay. 50 �g totalrotein of each supernatant was incubated along with acetylated tetrapeptide (Ac-EVD) substrate labeled with the chromophore p-nitroaniline (pNA) in the presencef caspase 3 buffer in a 96-well microplate in triplicate. In the presence of activeaspase 3, cleavage and release of pNA from the substrate occurs. Free pNA producesyellow colour detected by a spectrophotometer at 405 nm. A standard curve was

ealized in order to determine the correspondence between absorbance and pNAoncentration. Results were expressed as caspase 3 specific activity (pmol pNA/h/�grotein) calculated according to the manufacturer.

.11. Immunocytochemistry analysis

For the determination of cytochrome c localization, HCT116 cells were seedednto glass coverslips in complete medium. At 50% confluence, cells were incubatedith 120 �M of AME. After 16 h of treatment, cells were fixed in 3.7% paraformalde-yde for 10 min at 37 ◦C and permeabilized for 3 min at −20 ◦C with cold acetone.fter two washes with PBS, cells were saturated with PBS/BSA (3%) for 30 min. Pri-ary antibody was added for 1 h, and then secondary antibody was added for 30 min.uclei were stained by the addition of 1 �g/ml Hoechst 33348 for the last 5 min.ells fluorescence was examined under a DMR LEICA microscope equipped with anlympus DP70 photo camera (Rueil-Malmaison, France).

.12. Flow cytometric analysis

Cells were analyzed with Cell Lab Quanta MPL flow cytometer (Beckman-oulter, Paris, France). Data were analyzed using Cell Lab Quanta Analysis software.

.13. Statistical analysis

Each experiment was done independently three times. Values are presented asean ± SD. Statistical differences between control and treated groups for all experi-ents were determined by Student’s t-test. Differences were considered significant

t p < 0.05.

assay and expressed as percentages of viable cells (FDA+). Data are expressed asthe mean ± SD of three independent experiments. Values are significantly different(p < 0.05) from control.

3. Results

3.1. AME induces cell death

Cytotoxic effect of AME on HCT116 cells after 24 h of incuba-tion was measured by FDA assay. Results showed a dose-dependentinhibition of cell viability at concentrations ranging from 10 to200 �M of AME (Fig. 1). The IC50 value determined after 24 h ofcell treatment from the viability curve was about 120 �M. Thisconcentration was used in all further experiments.

3.2. Determination of the mode of cell death induced by AME

Cell death is known to occur mainly by two alternative modes:apoptosis, a programmed form of cell death, and necrosis, anunordered and accidental form of cellular dying. Apoptotic cellsdisplay specific morphological and biochemical features that dis-tinguish them from living and necrotic cells. They characteristicallyshow nuclear collapse and chromatin fragmentation which can bedetected by flow cytometric DNA content analysis. Indeed, apop-totic cells with fragmented DNA appear as cells with a hypoploidDNA content and are represented as a hypoploid Sub-G1 peak(Fig. 2A). HCT116 cells were treated with AME at 120 �M for 48 hand the percentage of apoptotic cells was quantified (Fig. 2B).The Sub-G1 population significantly increased from 9.59 ± 0.96%in control (untreated cells) to 58.15 ± 2.3 in AME-treated cells. Thehallmark of the necrotic process is the ultimate breakdown of theplasma membrane, due to the early loss of membrane integrity,leading to the release of cytoplasmic content into the extracellu-lar fluid. Therefore, necrosis was evaluated by labeling cells withthe membrane impermeant fluorochrome PI, which is excludedfrom viable cells but stained necrotic cells. As compared to a pos-itive control treated with 1 mM H2O2, in which the percentage ofnecrotic cells (PI+) increased by 5-fold compared with control, AMEdid not significantly induced necrosis (Fig. 2C). These results clearlydemonstrate that AME triggers an apoptotic process in HCT116 cellsrather than a necrotic death.

3.3. AME causes loss of the mitochondrial transmembranepotential (��m) and PTP opening

In order to assess the involvement of the intrinsic pathway inAME-induced apoptosis, we investigated the effect of this myco-toxin on MMP. Cells were treated with 120 �M of AME for 6, 16,24 and 48 h, and the ��m was evaluated using the membrane

potential-sensitive probe CMXRos. Cells showing a dissipation of��m present a lower CMXRos fluorescence (CMXRos-). As shownin Fig. 3A, AME induced a significant time-dependent loss of ��m.The ��m dissipation was significant after 16 h, the percentages

F. Bensassi et al. / Toxicology 290 (2011) 230–240 233

Fig. 2. AME induced nuclear events of apoptosis. HCT116 cells were treated for 48 h with 120 �M AME, fixed and DNA content was analyzed by flow cytometry. The percentageo ell cyto C). H2

t contr

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f hypoploid cells with fragmented DNA is presented on control and AME-treated cn histograms (B). The percentage of necrotic cells was detected after PI staining (hree independent experiments. (*) Values are significantly different (p < 0.05) from

f CMXRos- cells reaching about 3-fold and 4.5-fold of control val-es after 24 h and 48 h of mycotoxin exposure, respectively. Theitochondrial permeability transition is caused by the opening

f the permeability transition pore (PTP) which provokes matrixwelling and outer membrane rupture. To examine the role of PTP,e used the calcein/cobalt assay (Petronilli et al., 1999; Poncet

t al., 2003). Our results showed that AME induced the openingf PTP in 57 ± 4.1% of HCT116 cells after 24 h of mycotoxin’s expo-ure (Fig. 3B). Therefore, AME induces the mitochondrial pathwayf apoptosis at least by activating the opening of PTP.

.4. AME does not directly target mitochondrial PTP

A variety of different apoptosis inducers are able to act directlyn mitochondria and induce PTP opening, dissipation of ��m,

ograms (A). The percentage of cells in Sub-G1 of different experiments is presentedO2 (1 mM) was used as a positive control. Data are expressed as the mean ± SD ofol.

matrix swelling and other mitochondrial alterations. With respectto AME-induced PTP opening observed in cellulo, we evaluatedthe ability of this mycotoxin to directly target mitochondria. Thus,increasing concentrations of AME, ranging from 5 to 120 �M, wereadded to purified mitochondria. Calcium (6–50 �M) was used as apositive control and CsA was added to prevent calcium-induced PTPopening. Kinetics of the loss of ��m (increase in rhodamine 123fluorescence, Fig. 4A) and mitochondrial matrix swelling (decreasein OD, Fig. 4B) were concomitantly recorded. Our results indicatedthat AME induced neither depolarization (Fig. 4A(b)) nor swelling(Fig. 4B(d)) of mitochondria, in contrast to calcium, which induced

both a dose-dependent depolarization (Fig. 4A(a)) and swelling(Fig. 4A(c)) totally inhibited by CsA. Thus, these results indicatethat AME does not directly target mitochondria to induce PTP-dependent MMP.

234 F. Bensassi et al. / Toxicology

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Fig. 3. Effect of AME on mitochondrial transmembrane potential (��m) and PTPopening in HCT116 cells. Loss of ��m was assessed by flow cytometry after cellstreatment with AME (120 �M) at indicated times, and then stained with CMXRos (A).Opening of PTP was measured by the calcein/cobalt assay in response to AME treat-mtc

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ent (120 �M, 24 h) by flow cytometry (3B). Data are expressed as the mean ± SD ofhree independent experiments. (*) Values are significantly different (p < 0.05) fromontrol.

.5. AME induces MMP and cytochrome c release fromitochondria

MMP and release of cytochrome c from mitochondria are con-idered as key initial steps in the proteolytic cascade activationf the apoptotic process. To determine if the observed openingf PTP leads to MMP, we examined the intracellular localiza-ion of cytochrome c by immufluorescence in response to AME.

e observed that in control cells, presenting intact nuclei, theytochrome c fluorescence was punctuated and characteristic ofitochondrial localization (Fig. 5). After treatment with AME,

poptotic cells with condensed or fragmented nuclei showed aiffuse staining pattern for cytochrome c, indicating MMP andelease of this apoptogenic protein from mitochondria to the cyto-ol.

.6. AME-induced apoptosis is dependent upon caspase activation

It is well known that redistribution of cytochrome c to cyto-ol initiates the activation of the proteolytic cascade of caspases,

class of cysteine proteases that ultimately lead to apoptosisBoatright and Salvesen, 2003). To elucidate the role of caspasesn AME-induced apoptosis, whole cell lysates were prepared fromCT116 cells stimulated with AME for different treatment times

6, 16 and 24 h) and the activation of the upstream mitochon-rial initiator caspase 9 was analyzed by western blot. As shown

n Fig. 6A, the level of the cleaved active form of caspase 9ncreased with AME treatment in a time-dependent manner. Inddition, the activity of the downstream executioner caspase 3as recorded by spectrometry. Caspase 3 activity significantly

ncreased according to the incubation time with AME as comparedo control. Indeed, this activity increased from 4.97 ± 0.5 pmolNA/h/�g of protein after 6 h of AME exposure to 10.3 ± 1.2 pmolNA/h/�g of protein after 24 h (Fig. 6B). The implication of

290 (2011) 230–240

caspases was further tested by pre-treating cells for 2 h with thegeneral caspase inhibitor ZVAD-fmk before 24 h treatment withthe mycotoxin. The ��m was then measured by flow cytome-try. Fig. 6C showed that ZVAD-fmk pre-treatment decreased thepercentage of cell with low ��m (CMXRos-) (49 ± 3.2% in AME-treated cells vs. 21.9 ± 1.7% in the presence of ZVAD-fmk) andtherefore protected cells from mitochondrial apoptosis. Our datashowed that AME triggers a caspase-dependent apoptotic pro-cess.

3.7. AME induces superoxide anion generation

We wondered whether AME-cytotoxicity implicated reactiveoxygen species (ROS) generation. The production of cellular ROS(mainly H2O2) was analyzed using the ROS sensitive DCFH-DAprobe. Cellular ROS generation did not exceed 1.3 folds of con-trol in the presence of AME (120 �M) as compared to the positivecontrol cells treated with H2O2 (1 mM), in which an importantincrease in DCF fluorescence was observed (about 10-fold com-pared to untreated control). Thus, AME treatment does not inducea significant H2O2 generation (Fig. 7A). Mitochondrial superox-ide anion levels were also detected by staining cells with theMitoSOX probe. Mitochondrial O2

•− level increased from 8.4 ± 0.7%in control to 36.2 ± 2.8% in the presence of AME (Fig. 7B). Thisobservation indicated that ROS produced in response to AMEin HCT116 cells appears to be mainly mitochondrial superoxideanion.

Many reports have showed that ROS generation may be eithera cause or a consequence of the PTP opening and mitochondrialalterations in apoptosis (Skulachev, 1996; Le Bras et al., 2005).To determine whether mitochondrial ROS production are earlyor late events of the mitochondrial alterations, cells were pre-treated with 5 mM of the antioxidant N-acetylcysteine (NAC) andboth the ��m and nuclear fragmentation were measured follow-ing AME treatment. Concerning the ��m dissipation (CMXRos-cells), no significant difference was observed in cells incubatedwith AME alone or in the presence of NAC (Fig. 7C). By contrast,NAC partially inhibited AME-induced DNA fragmentation (Fig. 7D).Consequently, mitochondrial ROS appears to be generated down-stream of mitochondrial alterations as a consequence of the MMPtriggered by AME.

3.8. AME increases p53 protein level

In response to a variety of stresses such as DNA damage, onco-gene activations or xenobiotic exposure, the tumor suppressor p53can induce cell cycle arrest and/or apoptosis (Vousden and Lu, 2002;Wang et al., 2007). We investigated whether AME triggers a p53-dependent or -independent pathway in HCT116 cells. As shownin Fig. 8(A and B), AME treatment increased p53 protein level inHCT116 cells in a time-dependent manner with a maximal levelbeing reached after 16 h of exposure. This result suggests that p53protein is required in AME-mediated apoptosis in the studied coloncarcinoma cell line.

3.9. Bax is not involved in the regulation of AME-induced MMP

Among the numerous genes known to be transcriptionally acti-vated by p53, the pro-apoptotic protein Bax plays a crucial rolein inducing MMP either by forming oligomeres in the outer mito-chondrial membrane or by activating the PTP opening. To assess theinvolvement of Bax in the MMP induced by AME, we took advan-

tage of human colon carcinoma HCT116 cells deficient for Bax (HCTBax-KO). Thus, Parental (HCT116) and cells lacking Bax (HCT Bax-KO) were treated for 24 h with AME. As shown in Fig. 9, neitherthe dissipation of ��m (Fig. 9A) nor the opening of PTP (Fig. 9B)

F. Bensassi et al. / Toxicology 290 (2011) 230–240 235

Fig. 4. Effects of AME on mitochondrial membrane depolarization and swelling. Measurements were performed in the absence (Co) or in the presence of indicated concen-t ence oi depo� e dec

wctwbtMa

rations of AME. Calcium was used as a positive control in the absence or in the presn hypo-osmotic buffer containing 1 �M rhodamine 123. Mitochondrial membraneem = 535 nm) for 20 min. (B) Mitochondrial swelling was measured by recording th

as modified by the deficiency of Bax, comparatively to positiveontrol cells treated with LND (250 �M), an antineoplastic drughat induces apoptosis via activation of the mitochondrial path-ay (Ravagnan et al., 1999), which were greatly protected against

oth the loss of ��m and the PTP opening. These results indicatehat the pro-apoptotic protein Bax is not involved in AME-induced

MP and is not required for the opening of PTP and subsequentpoptosis.

f 5 �M CsA (an inhibitor of PTP). (A) Isolated mitochondria were incubated at 37 ◦Clarization was analyzed by measuring the increase of fluorescence (�exc = 485 nm,

rease of OD at 540 nm for 20 min. Experiments were done in triplicate.

4. Discussion

As secondary metabolites, mycotoxins are not essential for fun-gal growth and are produced sporadically under fungal stress.

Unfortunately, due to their toxic properties and their high stabilityto heat treatment, their presence in the food chain is potentiallyhazardous to the health of both humans and animals. Despite themounting concern about the health hazards posed by mycotoxins,

236 F. Bensassi et al. / Toxicology 290 (2011) 230–240

Fig. 5. AME induces nuclear fragmentation and cytochrome c release. Mitochondrial release of cytochrome c as determined by immunofluorescence in response to AME(120 �M) in HCT116 cells. Apoptotic cells were identified by Hoechst staining as bright condensed or fragmented nuclei. White arrows present cells with fragmented nucleiand released cytochrome c. Results are representative of three independent experiments.

Fig. 6. Role of caspases in AME-induced apoptosis. (A) Activation of caspase 9 in AME-treated cells. The levels of cleaved caspase 9 were determined after different treatmenttimes by Western blotting analysis. Tubulin was used as an internal loading control. (B) Activation of caspases 3 in HCT116 cells. Measurement of the activity of caspase 3(DEVD-pNa cleavage) in HCT116 cells exposed to AME (120 �M) at different treatment times. (C) The effect of ZVAD-fmk pre-treatment (2 h at 50 �M) on the loss of ��m wasassessed by flow cytometry. Histograms represent cells with dissipation of ��m (cells CMXRos-). Data are expressed as the mean ± SD of three independent experiments.(*) Values are significantly different (p < 0.05) from control. (�) Values are significantly different from cells treated with AME alone (p < 0.05).

F. Bensassi et al. / Toxicology 290 (2011) 230–240 237

Fig. 7. Effect of AME treatment on ROS generation in HCT116 cells. (A) The per-centage of cells producing H2O2 (cells DCF+) was analyzed by flow cytometry. (B)Superoxide anion (O2

•−) generation in AME treated HCT116 cells. O2•− production

was evaluated by analyzing MitoSOX fluorescence. H2O2 (1 mM) was used as a pos-itive control. Effects of the antioxidant N-acetylcysteine (NAC) pre-treatment onAME-induced ��m loss and nuclear apoptosis. HCT116 cells were pre-treated 2 hwith NAC (5 mM) prior to AME-treatment. (C) Histograms represent cells with ��mloss (cells CMXRos-). (D) Nuclear apoptosis/hypoploidy (Sub-G1) was detected 48 hadV

dcgaicactncb

Fig. 8. Up-regulation of p53 protein in HCT116 cells. p53 protein expression afterdifferent treatment times (6, 16 and 24 h), in response to AME (120 �M) (A) assessed

Golstein and Kroemer, 2007). We also found that AME increasesthe expression of the tumor suppressor protein p53 in HCT116cells (Fig. 8). p53 is activated when DNA repair systems are over-burdened due to too much DNA damage and acts as an upstream

Fig. 9. Determination of the role of the pro-apoptotic protein Bax in the AME-induced mitochondrial permeabilization. (A) Loss of ��m (DiOC-) was analyzedby flow cytometry in HCT116 KO for Bax (HCT116 Bax-KO) after 24 h of treatment

fter treatment with AME. Data are expressed as the mean ± SD of three indepen-ent experiments. (*) Values are significantly different (p < 0.05) from control. (�)alues are significantly different from cells treated with AME alone (p < 0.05).

ata on the toxicities of the mycotoxin AME, relevant for hazardharacterization, are scarce. Currently, there are no statutory oruideline limits set for AME by regulatory authorities. In this study,n attempt has been made to investigate the mechanisms of AME-nduced cell death in vitro. Our results clearly indicated that HCT116ells are sensitive to AME. The choice of toxin concentration tossess AME-induced cell death was made on the basis of the con-entration that reduced cells viability by about 50%. We observed

hat AME reduced HCT116 cells viability in a dose-dependent man-er with an IC50 of 120 �M after 24 h of treatment (Fig. 1). Thisell mortality was triggered by an apoptotic process, characterizedy chromatin condensation and DNA fragmentation. The apoptotic

by western blot analysis and (B) by scanning densitometry. Tubulin was used as aninternal loading control. Data are expressed as the mean ± SD of three independentexperiments. (*) Values are significantly different (p < 0.05) from control.

pattern was further confirmed by the persistence of plasma mem-brane integrity in AME-treated cells (Fig. 2C), contrarily to necrosis,branded by an early loss of integrity of the plasma membrane withresultant swelling of the cell and its organelles (Malhi et al., 2006;

with AME (120 �M). (B) Opening of PTP (Calcein-) in response to 24 h of AME treat-ment (120 �M) as assessed by flow cytometry. LND, 250 �M was used as a positivecontrol. Data are expressed as the mean ± SD of three independent experiments. (*)Values are significantly different (p < 0.05) from control. (�) Values are significantlydifferent from cells treated with LND alone (p < 0.05).

238 F. Bensassi et al. / Toxicology 290 (2011) 230–240

y of ap

ieNa2Awaa

iite(dpPcipdrbaosPcBtiot

Fig. 10. Signaling pathwa

nducer of apoptosis by activating the expression of genes-encodingither MMP-inducing proteins of the Bcl-2 family, such as Bax, Bid,oxa and PUMA, or other apoptotic and cell cycle regulators (Orennd Prives, 1996; Zhivotovsky and Kroemer, 2004; Bouleau et al.,007). Thus, our results involving p53 up-regulation suggest thatME could induce DNA damage in HCT116 cells and are consistentith other reports demonstrating that AME-mediated cell death is

ssociated with induction of DNA strand breaks in cultured humannd animal cells (Pfeiffer et al., 2007; Fehr et al., 2009).

In another set of experiments, we have demonstrated that thentrinsic/mitochondrial apoptotic pathway contributes to the AME-nduced apoptosis. Indeed, we demonstrated that AME provokeshe permeabilization of mitochondrial membranes. MMP is consid-red to be a central event and the point-of-no-return in apoptosisGalluzzi et al., 2006). To promote permeabilization of mitochon-rial membranes, different models were proposed, most notablyroteins of the Bcl-2 family (Bax, Bak, Bid) and/or the opening ofTP (Kroemer et al., 2007). The composition of PTP, a multiproteinomplex spanning the inner and outer mitochondrial membranes,s still a matter of debate. However, its opening results in dissi-ation of the mitochondrial inner membrane potential (��m),isruption of the outer mitochondrial membrane and subsequentelease of proteins such as the cytochrome c from the intermem-rane space. In this study, we demonstrated that AME inducespoptosis via a PTP-mediated MMP (Fig. 3B). Nevertheless, basedn the experiments conducted on isolated mitochondria, we clearlyhowed that AME does not directly act on mitochondria to induceTP opening. Moreover, we observed that AME triggers a mito-hondrial apoptotic process to a similar extent in wild-type andax-deficient colon carcinoma cells. These results strongly suggest

hat the MMP induced by AME is mainly due to the PTP open-ng in a Bax-independent manner. It could not be excluded thatther pro-apoptotic proteins such as Bak or Bid may substitutehe function of Bax in our system to favor PTP-induced MMP in

optosis induced by AME.

response to AME. Further investigations are needed to test thishypothesis.

The toxicities of different mycotoxins such as T-2 toxin, Pat-uline, Zearalenone and Aflatoxin B1 have been associated to thegeneration of oxidative stress (El Golli et al., 2006; Liu et al., 2007;Chatti Gazzah et al., 2010; El-Nekeety et al., 2011). In our model, anegligible increase in the production of H2O2 was observed in AME-treated cells (Fig. 7A), whereas the generation of mitochondrialO2

•− was clearly stimulated by AME (Fig. 7B). This O2•− produc-

tion occurs primarily on the matrix side of the inner mitochondrialmembrane (Orrenius et al., 2007). In addition, the antioxidant NACfailed to prevent the dissipation of ��m (Fig. 7C), but decreasedthe DNA fragmentation induced by AME (Fig. 7D). This suggests thatROS production occurs downstream as results of PTP opening andmitochondrial alterations.

Following MMP, the release into the cytosol of cytochrome cleads to its interaction with the adaptor molecule Apaf-1, result-ing in the recruitment and activation of the mitochondrial initiatorpro-caspase 9 in the presence of dATP (Martinou et al., 2000; Green,2005). The activated caspase 9, in turn, further leads to the acti-vation of downstream executioner caspases such as caspase 3 (Liet al., 1997; Antonsson, 2001). Caspase 3 is believed to be respon-sible for the cleavage of various proteins leading to biochemicaland morphological features characteristic of apoptosis (Earnshawet al., 1999; Tang et al., 2009). In HCT116 cells, the initiator caspase9 was activated by AME treatment, and triggered the activationof the executioner caspase 3. In the presence of the broad-rangeinhibitor of caspases ZVAD-fmk, the mycotoxin-induced MMP wasreduced, suggesting either that a caspase activated upstream ofmitochondria could favor MMP or that ZVAD-fmk inhibits a cas-

pase 3-mediated feedback amplification loop that amplifies MMPand mitochondrial alterations, as proposed previously (Chen et al.,2000). Altogether, these data demonstrated that HCT116 cells,treated by AME underwent a caspase-dependent apoptotic process.

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Taken together, our findings suggest a possible underlyingolecular mechanism for AME-induced cell death in vitro, and

mphasize the anticancer property of this mycotoxin in humanolon carcinoma cells. This mycotoxin provokes DNA damage thatnduces a p53-dependent PTP-mediated activation of the mito-hondrial pathway of apoptosis (Fig. 10). To the best of ournowledge, this is the first in vitro study which describes that thelternaria mycotoxin AME induces a mitochondrial apoptotic path-ay in human colon cells, and highlights the anticancer property

f AME against colon carcinoma.

onflict of interest statement

Authors assure that there are no conflicts of interest:

cknowledgments

This study was supported by “Le Ministère Tunisien de’Enseignement Supérieur, de la Recherche Scientifique et de laechnologie”. We thank Dr. Mbarek Chetoui (Faculty of Sciences,unisia) for his help and Dr. Salwa Bacha (University of La Manouba,unisia) for the correction of the English manuscript.

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