Chemico-Biological Interactions 199 (2012) 161–176

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Chemico-Biological Interactions

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EPR spin trapping evaluation of ROS production in human fibroblasts exposedto cerium oxide nanoparticles: Evidence for NADPH oxidase andmitochondrial stimulation

Marcel Culcasi a, Laila Benameur a,b, Anne Mercier a, Céline Lucchesi a, Hidayat Rahmouni a, Alice Asteian a,Gilles Casano a, Alain Botta b, Hervé Kovacic c, Sylvia Pietri a,⇑a CNRS, Aix Marseille Université, Institut de Chimie Radicalaire UMR 7273, Equipe ‘‘Sondes Moléculaires en Biologie et Stress Oxydant’’, Marseille, Franceb Aix Marseille Université, CNRS, EA 1784/FR 3098 ECCOREV, Laboratoire de Biogénotoxicologie et Mutagenèse Environnementale, Marseille, Francec Aix Marseille Université, INSERM UMR 911, CRO2, Marseille, France

a r t i c l e i n f o

Article history:Received 6 February 2012Received in revised form 31 July 2012Accepted 3 August 2012Available online 22 August 2012

Keywords:Free radicalsESR spin trappingCerium oxide nanoparticlesCell signalingFibroblastsCytotoxicity

0009-2797/$ - see front matter � 2012 Elsevier Irelanhttp://dx.doi.org/10.1016/j.cbi.2012.08.007

Abbreviations: ACV, acetovanillone; AFR, ascorbyl fchemiluminescence; DCIP, 2,6-dichlorophenolindophphosphoryl)-5-methyl-1-pyrroline N-oxide; DMEM,medium; DMSO, dimethylsulfoxide; DPI, diphenylenetriaminepentaacetic acid; EPR, electron paramagneticdiaminetetraacetic acid; GPx, glutathione peroxidaseGSSG, glutathione disulfide; KH, Krebs–Henseleit megenase; MDA, malondialdehyde; MTT, 3-(4,5-dimethtetrazolium bromide; nano-CeO2, cerium oxide nanooxide; PBS, phosphate-buffered saline; ROS, reactiveoxide dismutase; TBA, 2-thiobarbituric acid; TEMPO,done-N-oxyl radical.⇑ Corresponding author. Address: Sondes Molécu

Oxydant, CNRS UMR 7273, Institut de Chimie RadicalaCentre scientifique de Saint-Jérôme, Case 522, AvNiemen, F-13397 Marseille, France. Tel.: +33 (0) 49288 758.

E-mail address: sylvia.pietri@univ-amu.fr (S. Pietri

a b s t r a c t

To better understand the antioxidant (enzyme mimetic, free radical scavenger) versus oxidant and cyto-toxic properties of the industrially used cerium oxide nanoparticles (nano-CeO2), we investigated theireffects on reactive oxygen species formation and changes in the antioxidant pool of human dermaland murine 3T3 fibroblasts at doses relevant to chronic inhalation or contact with skin. Electron para-magnetic resonance (EPR) spin trapping with the nitrone DEPMPO showed that pretreatment of the cellswith the nanoparticles dose-dependently triggered the release in the culture medium of superoxide dis-mutase- and catalase-inhibitable DEPMPO/hydroxyl radical adducts (DEPMPO–OH) and ascorbyl radical,a marker of ascorbate depletion. This DEPMPO–OH formation occurred 2 to 24 h following removal of theparticles from the medium and paralleled with an increase of cell lipid peroxidation. These effects ofinternalized nano-CeO2 on spin adduct formation were then investigated at the cellular level by usingspecific NADPH oxidase inhibitors, transfection techniques and a mitochondria-targeted antioxidant.When micromolar doses of nano-CeO2 were used, weak DEPMPO–OH levels but no loss of cell viabilitywere observed, suggesting that cell signaling mechanisms through protein synthesis and membraneNADPH oxidase activation occurred. Incubation of the cells with higher millimolar doses provoked a25–60-fold higher DEPMPO–OH formation together with a decrease in cell viability, early apoptosisinduction and antioxidant depletion. These cytotoxic effects could be due to activation of both the mito-chondrial source and Nox2 and Nox4 dependent NADPH oxidase complex. Regarding possible mecha-nisms of nano-CeO2-induced free radical formation in cells, in vitro EPR and spectrophotometricstudies suggest that, contrary to Fe2+ ions, the Ce3+ redox state at the surface of the particles is probablynot an efficient catalyst of hydroxyl radical formation by a Fenton-like reaction in vivo.

� 2012 Elsevier Ireland Ltd. All rights reserved.

d Ltd. All rights reserved.

ree radical; CAT, catalase; CL,enol; DEPMPO, 5-(diethoxy-Dulbecco’s modified Eagle’siodonium; DTPA, diethylene-

resonance; EDTA, ethylene-; GSH, reduced glutathione;dium; LDH, lactate dehydro-ylthiazol-2-yl)-2,5-diphenyl-particles; PAO, phenylarsineoxygen species; SOD, super-2,2,6,6-tetramethyl-4-piperi-

laires en Biologie et Stressire, Aix-Marseille Université,enue Escadrille Normandie

1 288 579; fax: +33 (0) 491

).

1. Introduction

Engineered materials containing cerium oxide (CeO2) nanopar-ticles, nanoceria (nano-CeO2), are widely used in industry as dieselfuel additives, catalysts, semiconductors, solar cells or oxygen sen-sors [1] and also offer promising perspectives in medical diagnosticor therapy [2]. Perhaps the most remarkable physicochemicalproperty of CeO2 is that its surface contains oxygen vacanciesreleasing electrons which may reduce ceric ions (Ce4+) into cerousions (Ce3+), and in nano-CeO2 both the Ce4+/Ce3+ redox cycle andoxygen exchange are dramatically increased due to its nanoscaledimension [3].

An important consequence of this unique redox chemistry isthat nano-CeO2 is a potential free radical scavenger and antioxi-dant. For this main reason nano-CeO2 has been tested in a number

162 M. Culcasi et al. / Chemico-Biological Interactions 199 (2012) 161–176

of biological systems where formation of reactive oxygen species(ROS), including free radicals, is suspected to occur. Thus nano-CeO2 has been found protective against cellular damage inducedby toxicants [4,5], radiation [6] or in pathological situations suchas cardiac [7] or brain [8] ischemia/reperfusion. As a mechanisticexplanation for these observations physicochemical assays, includ-ing electron paramagnetic resonance (EPR), have proposed thatnano-CeO2 can alternatively behave like superoxide dismutase(SOD) [9,10] or catalase [11].

Paradoxically many papers about the toxicity of nano-CeO2 inrelation with ROS production have been published. Hence whennano-CeO2 was added to the culture medium of a variety of normalcells [12–14] or administered to healthy animals [15–17] a numberof adverse effects were reported such as cytotoxicity, genotoxicity,lipid and protein oxidative damage, antioxidant depletion, lifespanreduction, systemic inflammation or cardiovascular alterations. Be-cause cytotoxicity linked to oxidative stress was observed uponin vivo administration of CeCl3 to rats [18,19] it was investigatedwhether cerous ions could efficiently trigger ROS formationin vitro. Based on the EPR spin trapping technique two studies[19,20] proposed that Ce3+ can convert hydrogen peroxide (H2O2)into the highly-reactive hydroxyl radical (HO�) by a Fenton-likemechanism (Eq. (1)), thereby setting the conditions for oxidativestress to occur:

Ce3þ þH2O2 þHþ ! Ce4þ þHO� þH2O ð1Þ

Besides this potential mechanism of nano-CeO2 toxicity, apH-dependent but H2O2-independent oxidase-like activity of theparticles has been recently demonstrated in vitro for a series of or-ganic substrates or in low pH cell organelles [21] and tumoralmicroacidic environment [22] upon nanoceria internalization. Thisprooxidant activity of nano-CeO2 has been used to design targetedanticancer agents [21,22]. Another possible oxidative mechanismnot involving the transient formation of HO� may involve the inter-action of the Ce3+ redox state of CeO2 with H2O2 to convertadsorbed organics into hydroperoxides [23].

In most of the mechanisms cited above, the Ce4+/Ce3+ redox cy-cle will trigger oxidative stress only if cellular sources of H2O2 and/or its biological precursor superoxide (O2

��) would become acti-vated. In this regard adding nano-CeO2 to several biological sys-tems activates oxidative stress genes and apoptosis [13] orstimulates the release of inflammatory cytokines and chemokines[17].

At the molecular level knowledge is still lacking concerning thenature and cellular sources of the species produced in response tonano-CeO2 interactions and the relationship between their poten-tial ROS-generating capability, their impact on cell signaling andthe development of pathophysiological outcomes [24]. Using awide array of spectral and biochemical techniques, and a cell trans-fection approach, we explored herein some chemical and biochem-ical facets of cell/nano-CeO2 concentration-dependent interactionand the possible cellular H2O2 generating sources [25] involved(i.e., the mitochondrial respiratory chain and NADPH oxidase) intwo models of fibroblasts.

2. Materials and methods

2.1. Chemicals, reagents and analytical techniques

Solvents, starting materials and reagents, including diethylene-triaminepentaacetic acid (DTPA), ethylenediaminetetraacetic acid(EDTA), H2O2, 1,1,3,3-tetramethoxypropane (TMP), 2-thiobarbitu-ric acid (TBA), 2,6-dichlorophenolindophenol (DCIP), 2-deoxy-d-ri-bose (deoxyribose), 2(N-morpholino) ethanesulfonic acid (MES),reduced glutathione (GSH), glutathione peroxidase (GPx), dimeth-

ylsulfoxide (DMSO), acetovanillone (apocynin, ACV), phenylarsineoxide (PAO), diphenyleneiodonium (DPI), 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), Rhodamine 123,Hoechst 33342, 7-amino-actinomycin (7-AAD+), 2,2,6,6-tetra-methyl-4-piperidone-N-oxyl radical (TEMPO), catalase (CAT) andxanthine oxidase from bovine liver, and SOD from bovine erythro-cytes, were from Sigma–Aldrich Chimie (Saint Quentin Fallavier,France). Annexin V-APC was from BD Biosciences, France. The spintrap 5-(diethoxyphosphoryl)-5-methyl-1-pyrroline N-oxide (DEP-MPO) was synthesized and purified according to [26,27]. For cellculture, Dulbecco’s modified Eagle’s medium (DMEM), HEPES buf-fer, phosphate-buffered saline (PBS), trypsin–EDTA, Ham’s F12medium and fetal calf serum (FCS) were obtained from Life Tech-nologies Inc. (Gaithersburg, MD, USA). Doubly distilled deionizedwater was used throughout.

Analytical grade Ce salts and derivatives used in this study(from Sigma–Aldrich) were Ce(III): CeCl3, Ce2(SO4)3 and Ce2(CO3)3,and Ce(IV): Ce(SO4)2, (NH4)2Ce(NO3)6 and CeO2. Nano-CeO2, theformula of which being CeO2(HNO3)0.5(H2O), were provided byRhodia Chemicals (France) and are ellipsoidal monocrystallites ofcerianite with a mean diameter of 7 nm and a specific surface areaof 400 m2/g [14]. Stock solutions of CeO2 particles were preparedby vortexing for 10 min suspensions of the compound (1–3 mg/ml) in either DMEM (cell incubations) or water (test-tube controls)prior to use.

1H, 13C and 31P NMR spectra were recorded on a Bruker AVL 300spectrometer. Chemical shifts d are reported as ppm relative toexternal 85% H3PO4 (31P) or tetramethylsilane as internal standard(other nuclei). High resolution mass spectrometry–electron sprayionization (HRMS–ESI) analysis was performed using a QStar Eliteinstrument (AB Sciex, USA).

2.2. Modified procedure for mitoPBN synthesis

[4-[4-[[(1,1-Dimethylethyl)-oxidoimino]methyl]phenoxy]butyl]triphenylphosphonium bromide (mitoPBN; 3) was synthesized inthree steps using a modification (Scheme 1) of the procedurereported in [28]. The first step involved synthesis of 4-(4-iodobut-oxy) benzaldehyde (1) from 4-hydroxybenzaldehyde and 1,4-diiodobutane in the presence of K2CO3 by adapting a protocoldescribed for x-dibromoalkanes [29]. Compound (1), which wasrecovered with a high purity, was then converted into the triphen-ylphosphonium iodide (2), which ultimately led to mitoPBN (3) byreductive condensation with 2-methyl-2-nitropropane. These twolatter synthetic steps used the conditions described in [30]. Theexperimental detail and the characterization of the intermediates,including NMR and HRMS–ESI analysis, is given in the appendix.

2.3. Cell culture procedures and treatment with nano-CeO2

2.3.1. Experiments on normal human fibroblastsPrimary cultures of human fibroblasts were isolated by the out-

growth method using infant foreskins obtained after circumcision[14]. The dermis was cut into small pieces of 0.5–1 mm3 understerile conditions. The tissue pieces were seeded in culture dishesand incubated in DMEM supplemented with 10% FCS, 2 mM l-glu-tamine, 1 mM sodium pyruvate, 100 U/ml penicillin and 100 lg/mlstreptomycin. Fibroblasts were cultured at 37 �C in a humidifiedatmosphere of 5% CO2 in air in antibiotic-free DMEM and isolatedcells were generally obtained after 2 weeks where the culturemedium was changed every 2 days.

Isolated fibroblasts (105 cells/ml) were plated in 25 cm2 flasksin a humidified atmosphere of 5% CO2 in air and exposed at 37 �Cfor 2 or 24 h to freshly prepared dilutions of nano-CeO2 stocksolution (i.e., 6 � 10�5; 6 � 10�4 or 6 � 10�1 g/l, corresponding to

Scheme 1. Modified synthesis of mitoPBN (3).

M. Culcasi et al. / Chemico-Biological Interactions 199 (2012) 161–176 163

a range of 0.22 lM–2.2 mM). The supernatants were timely col-lected, placed in cryotubes and either quickly frozen in liquidnitrogen for delayed assessment of their ascorbyl free radical(AFR)–DMSO content [31] or stored at �80 �C for later extracellularlactate dehydrogenase (LDH) assay (see below). The remainingadherent cells were trypsinized and PBS-resuspended (giving�50,000 cells/ml) for lipid peroxidation and protein measurement(see below).

To investigate the mechanisms involved in the short- (2 h) orlong- (24 h) term effects of nano-CeO2 on cells the same parame-ters as described above were measured in additional fibroblaststhat were (i) first pretreated with the NADPH oxidase inhibitorsPAO (0.1 mM), DPI (5 lM) or ACV (0.3 mM) for 30 min, 1, or 4 h,respectively, and, after removal of the incubation medium, reincu-bated with 6 � 10�5–6 � 10�1 g/l nano-CeO2, or (ii) coincubatedfor up to 24 h with nano-CeO2 and either DEPMPO (0.1 mM), CAT(2 mU/ml) or a mixture of SOD (5 U/ml) and CAT (2 mU/ml), or(iii) first exposed to 6 � 10�1 g/l nano-CeO2 for 23 h before coincu-bating with 10 lM DPI for 1 h, or (iv) pretreated with mitochon-dria-targeted mitoPBN (20 lM) for 1 h followed by removal ofcell medium and incubation for 24 h with 6 � 10�1 g/l nano-CeO2

in DMEM, a concentration reported to affect the metabolic mito-chondrial activity [14].

Other more specific indices of cell viability were determined onfibroblasts coincubated up to 48 h with graded nano-CeO2 doses(10�5�1 g/l, corresponding to concentrations ranging 0.036 lM–3.36 mM), i.e., Trypan blue exclusion and MTT assays, the latterbeing an indicator of mitochondrial dehydrogenase activity. Theprotocol for the MTT assay was adapted from [22]: following incu-bation of the cells with nano-CeO2, MTT (0.5 mg/ml) was added tothe medium, incubation was extended for 1 h and the conversionof MTT to a purple formazan precipitate was monitored at 570 nm.

Nano-CeO2-induced mitochondrial toxicity was assessed by theretention of Rhodamine 123, a membrane-permeable fluorescentcationic dye. Cells were first incubated for 24 h with the particlesas described in the viability tests, then Rhodamine 123 (10 lM)and Hoechst 333412 (10 lg/ml) were successively added to themedium and incubation was extended for 15 min. After beingwashed twice with calcium- and magnesium-containing PBS, cellswere observed under an oil immersion 40�/1.4 plan apochromatobjective on a confocal Leica SP5 microscope (Leica Microsystems,Nanterre, France).

The effect of nano-CeO2 on cell apoptosis was determined usingquantification of Annexin V and 7-AAD staining, a combinationallowing the differentiation among viable cells (Annexin V nega-tive, 7-AAD negative), early apoptotic cells (Annexin V positive,7-AAD negative) and late apoptotic or necrotic cells (Annexin V po-sitive, 7-AAD positive). Following a 48-h exposure with nano-CeO2

(10�5�1 g/l) cells were harvested and stained for 20 min with An-nexin V and 7-AAD in binding buffer containing 0.1 M HEPES (pH7.4), 1.4 M NaCl and 25 mM CaCl2 at room temperature in the dark.

At the end of incubation, cells were resuspended in binding bufferand analyzed by flow cytometry on a FACSCalibur flow cytometer(BD Biosciences) according to [32].

2.3.2. Experiments on transfected human fibroblastsPlasmids for p47phox wild-type, p47-M1 (dominant negative

mutant) and p47-M8 (constitutively active mutant) were kindlyprovided by Dr. L.S. Terada (University of Texas, USA). p47-M1plasmid (W193R mutation) has been shown to lack binding func-tion of the first p47 SH3 domain, a function critical to assemblyof the oxidase [33]. p47-M8 plasmid (S303, 304, 328D mutation)mimics phosphorylation of three critical serines and appears to in-duce unmasking of the tandem SH3 regions needed for oxidaseassembly and activation [33]. Human dermal fibroblasts were har-vested in single cell suspension by treatment with trypsin–EDTA,and subsequently transfected by amaxa nucleofector according tothe manufacturer’s protocol. Transfected cells were seeded in sixwell plates at a density of 20,000 cells/cm2 and incubated for24 h with 6 � 10�1 g/l nano-CeO2 in DMEM. Following incubationthe supernatants were collected and stored at �80 �C for laterLDH assay (see below). Transfection controls for p47-M1 andp47-M8 plasmids were done using nucleofection without plasmid(control WP cells) and transfection with p47phox wild-type. Con-trol WP cells incubated for 24 h with H2O2 (0.5 and 500 lM) wereused as positive controls.

2.3.3. Preparation of samples of nano-CeO2 exposed human fibroblastsfor spin trapping analysis

Following exposure to nano-CeO2, the culture medium of eachflask was removed and stored for LDH assay. Cells were washedthree times with PBS and cells were reincubated for 15 min at37 �C in 5 ml of DTPA (0.3 mM)- and DEPMPO (20 mM)-containingPBS in order to reach 105 cells/ml. The supernatant was then trans-fered into cryotubes and immediately frozen in liquid nitrogenprior to EPR analysis. This procedure was applied to prevent anyphysiologically unrelevant, nano-CeO2-catalyzed formation ofDEPMPO spin adducts (see results and discussion).

2.3.4. Preparation of whole cell lysates and western blottingThe adherent cells from flasks assayed by spin trapping as de-

scribed above were scraped and lysed in bromophenol blue-freeLaemmli buffer supplemented with complete protease inhibitorcocktail (Roche Applied Science, France) and centrifuged for10 min at 10,000g. Supernatant protein concentration was deter-mined using the bicinchoninic acid assay-reducing agent compati-ble (Pierce, France) following the manufacturer’s instructions.Equal amounts of protein from lysates were resolved by 12%SDS–PAGE and transferred to Hybond ECL nitrocellulose mem-branes (Amersham Pharmacia Biotech, France). Membranes wereblocked in low-fat milk and incubated with primary antibodiesovernight at 4 �C. As primary antibodies, 0.5 lg/ml of mouse

164 M. Culcasi et al. / Chemico-Biological Interactions 199 (2012) 161–176

monoclonal anti-actin (Sigma–Aldrich, clone AC15), 1 lg/ml ofrabbit polyclonal anti-Nox4 (SantaCruz Biotechnology, sc-30141)and a 1/500 dilution of rabbit polyclonal anti-p47-phox (UpstateBiotechnology, 07–001) were used. Membranes were washed andsubsequently incubated for 1 h with an anti-mouse or anti-rabbitHRP-conjugate secondary antibody. Proteins were quantified usingan enhanced chemiluminescence detection kit (Amersham Phar-macia Biotech, France).

2.3.5. Experiments on murine 3T3 fibroblasts3T3 NIH fibroblasts (ATCC–LGC Promochem, Molsheim, France)

were maintained in culture at 37 �C in a 5% CO2-humidified atmo-sphere in DMEM containing 1% glucose and supplemented with10% FCS, 100 U/ml penicillin and 100 mg/ml streptomycin. Cellswere plated in 24-well dishes and medium was replenished every2–3 days until confluency which was checked by microscopicobservation.

For cell exposure 10-ll aliquots of nano-CeO2 stock solutionwere transferred into each culture well of confluent cells previ-ously filled with phenol red-free DMEM containing 1% glucose, inorder to reach a final volume of 0.5 ml/well and final nano-CeO2

concentrations of 6 � 10�4 or 6 � 10�1 g/l. Cell cultures were thenincubated for 24 h at 37 �C in a 5% CO2-humidified atmosphere,culture medium was sampled for extracellular LDH assay andscraped cells were stored at �80 �C until antioxidant and lipid per-oxidation assays. Additional inhibitions with PAO, SOD, CAT ormitoPBN were performed as described above for human fibro-blasts. In all these experiments the controls represented cells beingincubated in DMEM alone for the same period.

2.4. Biochemical assays

Nano-CeO2-treated scraped 3T3 fibroblasts were successivelythawed and homogenized (�105 cells/0.5 ml) on ice in cold MESbuffer (0.04 M; pH 6.5) containing 1 mM EDTA and centrifugedfor 15–20 min (10,000g) at 3 �C. The concentration of total gluta-thione was measured in the supernatants by the enzymatic recy-cling method involving 5,50-dithio-bis-2-(nitrobenzoate) whichreacts with the sulfhydryl group of GSH using a MP96 microplatereader (SAFAS, Monaco) equipped with a 405–414 nm filter. Gluta-thione disulfide (GSSG) was determined after derivatization of GSHwith 2-vinylpyridine [34]. The concentration of GSH was calculatedfrom the difference between total glutathione and GSSG. Data areexpressed in nmol/106 cells and are the means of at least 10 exper-iments made in duplicate.

Ascorbate content in cell homogenates was determined by anassay involving reduction of DCIP dye by ascorbic acid [35]. Dataare expressed in nmol/106 cells and are the means of at least 10independent experiments made in triplicate.

Lipid peroxidation was assessed in human or 3T3 fibroblasts bymeasuring MDA–TBA levels as described in [36]. One milliliter ofthe cell homogenate supernatant, treated with 0.5 ml acetic acid(20%) and 0.5 ml TBA (10 g/l), was incubated for 20 min at 95 �C,cooled down at room temperature and assayed at 532 nm. A cali-bration curve was obtained from standard MDA–TBA samples pre-pared by using 0.25 ml aliquots of 0.1–10 mM TMP solutions.Protein determination was performed in two randomly selectedflasks or wells [36]. Data are representative of 6–12 independentexperiments.

Release of cytoplasmic LDH was determined by assaying LDHactivity in the supernatant of human or 3T3 cells using a commer-cial detection kit (Roche Diagnostic, Mannheim, Germany) anddata (in (UI/l/mg protein) are representative of 6–12 independentexperiments. To estimate the total LDH content a control measure-ment was performed for each set of experiments, by treating cellstreated with 1% Triton X-100 to induce both total LDH release in

the supernatant and 100% loss of viability. The percentage of LDHreleased was calculated with respect to the total amount whichis the sum of the enzymatic activity in the lysate and in the culturemedium.

The determination of intracellular SOD, CAT and GPx activitieswas performed at room temperature in samples prepared from105 scraped 3T3 cells homogenized in 0.5 ml of appropriate cell ly-sis buffers according to reported procedures [37]. Color develop-ment was determined at 440, 540 and 340 nm for SOD, CAT andGPx, respectively. Data are expressed in UI/106 cells according tostandard calibration curves obtained from commercial enzymesand are representative of 6–12 independent experiments. Oxidizedcarbonylated proteins were determined at 450 nm with the OxySe-lectTM protein carbonyl ELISA detection kit (Cell Biolabs, Inc., USA)using a 96-wells microplate reader and bovine serum albumin asthe standard. Data are expressed in nmol/mg cell lysate accordingto standard calibration curves. The viability of 3T3 cells incubatedwith nano-CeO2 in the range 6 � 10�5–6 � 10�1 g/l was measuredby the MTT assay as previously described [22].

2.5. EPR spectroscopy

2.5.1. Cell culturesIn the culture medium of cells undergoing oxidative stress a

moderate DEPMPO concentration of 10–20 mM was consideredappropriate to allow EPR detection of spin adducts from ROS[38]. In contrast at a concentration too low to give significant spinadduct formation (i.e., 0.1 mM) the nitrone was rather expected toplay a protective role against antioxidant depletion [27]. Thusthawed samples containing 20 mM DEPMPO were directly intro-duced into a 10-mm quartz flat cell for EPR examination while thatcontaining 0.1 mM of the nitrone were first added to DMSO (1:1 v/v) and then introduced into calibrated 50-ll glass capillary tubes(Hirschmann Laborgeräte, Eberstadt, Germany) for AFR-DMSOmeasurement.

The EPR spectra were recorded at room temperature with a Bru-ker ESP 300 spectrometer (Karlsruhe, Germany) operating at9.8 GHz with a 100-kHz modulation frequency and a microwavepower of 10 mW. Other parameters for DEPMPO spin trapping(AFR–DMSO) were: modulation amplitude, 0.497 (0.883) G; recei-ver gain, 2 � 105 (5 � 105); time constant, 40.96 (81.92) ms; fieldresolution, 4096 (2048) points; sweep width, 140 (45) G; sweeprate, 3.34 (1.07) G/s; number of averaged scans, 2 (4). Spectralacquisition was initiated 1 min after complete thawing of the sam-ple (DEPMPO spin trapping) or 2 min after addition of DMSO. EPRsignals were quantitated by double integration of the simulatedspectra using the Winsim program [39] and radical concentrationswere estimated by calibration with either TEMPO (for spin ad-ducts) or freshly prepared AFR–DMSO (from sodium ascorbate)in PBS. Data represent means of 6–12 independent experimentsfor each tested experimental condition.

2.5.2. Test-tube controlsExperiments aimed at forming or inhibiting typical DEPMPO

spin adducts were carried out in 20 mM KH2PO4 buffer (pH 7) con-taining 1 mM DTPA (termed as ‘KH–DTPA’ throughout). The initialfree radical generator was either the H2O2 (1 mM)–FeSO4 (1 mM)Fenton system or a similar reagent where 1–10 mM Ce compoundwas used instead of Fe2+. Both systems were allowed to react with22–50 mM DEPMPO alone or in combination with 20 mU/ml SODor excess competitors such as DMSO or EtOH (15%, v/v). In anotherset of experiments Ce salts or nano-CeO2 (1–10 mM) were com-pared to Fe3+ (2 mM, as FeCl3) as catalysts of the nucleophilic syn-thesis of the DEPMPO/OH adduct (DEPMPO–OH) from 50 mMaqueous DEPMPO. EPR signals were recorded with a flat cell usingtypical conditions as described above. Except the microwave

M. Culcasi et al. / Chemico-Biological Interactions 199 (2012) 161–176 165

power, all other settings were adapted depending on the sampleand are noted in each figure legend.

2.6. Estimation of the interaction of Ce compounds with superoxideand hydrogen peroxide in vitro

Superoxide radicals were produced from xanthine (50 lM)-xan-thine oxidase (10 mU/ml) in EDTA (0.1 mM)-supplemented 0.2 Msodium phosphate buffer, pH 7.5 and their formation was moni-tored for 10 min by lucigenin (5 lM) chemiluminescence (CL) asdescribed in [40]. This system was used to (i) evaluate the O2

��

scavenging effect of a series of Ce derivatives (0.005–1 mM) ascompared to SOD, and (ii) to check whether their pre-incubationup to 24 h with SOD (5–20 mU/ml) at 37 �C in 0.2 M KH2PO4, pH7.5 will modify enzymatic activity. In control experiments the po-tential of tested Ce derivatives to inhibit xanthine oxidase wasevaluated spectrophotometrically by monitoring uric acid forma-tion from xanthine.

The deoxyribose assay [41] was used as another method to as-sess if HO� radicals are formed in the reaction of Ce derivatives withH2O2. Briefly, reaction tubes contained 1 mM H2O2, 1.1 mM EDTAand either 0.2 mM FeSO4 or various Ce compounds (0.2–1 mM)in 20 mM KH2PO4 (final volume, 1 ml), pH 7.4. Control samplescontained 5–15 mM of the HO� radical scavenger mannitol orKNO3, as a model for nitrate which enters the nano-CeO2 composi-tion (see above). The mixtures were incubated at 37 �C for 1 h andthe MDA–TBA assay was carried out as described above. To checkfor any Ce-chelating effect of the buffer, the experiments were re-peated in HEPES and all measurements were compared againstblanks where water was used instead of H2O2 or Ce compound.Data are the means of 6–10 experiments/concentration.

2.7. Statistical analysis

All values are expressed as the means ± SEM unless otherwisenoted. Statistical analysis was carried out using Student’s t testor one-way ANOVA followed by a a posteriori Newman–Keuls orDuncan’s tests. Intergroup differences were considered significantat P < 0.05.

3. Results and discussion

3.1. Formation of DEPMPO hydroxyl radical adducts from aqueousnano-CeO2 by a mechanism independent from free radical generation

In biological spin trapping four major pathways can lead to theformation of HO� spin adducts of five-membered ring pyrroline N-oxides such as DEPMPO, namely (1) direct scavenging of the radi-cal, (2) reduction of the O2

�� spin adduct by enzymes such as GPx,(3) nucleophilic addition of water catalyzed by metal ions such asFe3+ (Forrester–Hepburn mechanism) [26,42], and (4) hydrolysis ofthe cation radical resulting from single electron oxidation of thenitrone (inverted spin trapping) [43], the two latter mechanismsnot requiring the presence of HO� radicals in the system.

To evaluate whether Ce compounds could act as catalysts fornucleophilic addition DEPMPO (50 mM) was incubated with10 mM nano-CeO2 in deionized water (final pH �3). A strong EPRsignal characteristic of DEPMPO–OH was readily detected(Fig. 1A) which was simulated as a cis (trans) mixture of diastere-oisomers (relative to the diethoxyphosphoryl group): 46% (54%)with parameters aN = 14.05 (14.05) G; aP = 47.29 (47.29) G andaHb = 14.24 (12.73) G, see structure in Fig. 1) [44]. When the aque-ous DEPMPO and nano-CeO2 were mixed in the presence of excessDMSO, DEPMPO–OH was still the only detected species (Fig. 1B)instead of the expected 12-line signal of DEPMPO/methyl radical

adduct which would have formed if HO� radicals had first attackedthe competitor [36]. The signal of Fig. 1A was dose-dependentlyinhibited by addition of the chelator DTPA (0.5–5 mM) in 20 mMKH2PO4 (final pH, 7.1; Fig. 1C). Consistent with a nucleophilic addi-tion mechanism [45], mixing DEPMPO and nano-CeO2 in puremethanol afforded DEPMPO–OMe (recorded as a mixture of trans(cis) diastereoisomers: 76% (24%); aN = 13.03 (12.86) G; aP = 40.16(46.91) G; aHb = 7.48 (7.29) G; aHc = 1.73 G (not resolved))(Fig. 1D). Since these data strongly suggest that DEPMPO–OH seenin Fig. 1A can form by a HO�–unrelated mechanism involving ionscontained in nano-CeO2 that can be inactivated by chelation, wefirst incubated aqueous DEPMPO (50 mM) with 10 mM of any ofthe Ce(III) salts CeCl3, Ce2(CO3)3 or Ce2(SO4)3 for up to 30 min. Un-der these conditions no EPR signal was observed as illustrated byFig. 1E for cerous carbonate. When the same experiment was per-formed with 1–10 mM cerium sulfate, a Ce4+ salt, a strong signalwas detected (Fig. 1F), assigned to DEPMPOX (aN = 7.14 G;aP = 38.69 G; aHb = 3.69 G (2H)), an oxidized form of DEPMPO[46]. However Fig. 1G shows that using aqueous CeO2 suspensionsyielded weak DEPMPO–OH signals (about 80-fold lower than withnano-CeO2) that were inhibited in the presence of a chelated phos-phate buffer (Fig. 1H). This suggests that unlike free Ce3+/Ce4+ ions,covalently bound Ce(IV) in CeO2 behaves as a weak catalyst in anucleophilic-type reaction with DEPMPO and that this propertyis strongly enhanced in the nano-structured material.

The mechanism not related to true spin trapping by which DEP-MPO–OH forms in the presence of CeO2 or nano-CeO2 in aqueousmedium remains to be elucidated. In this regard inverted spin trap-ping, which primarily requires single electron oxidation of DEP-MPO, appears unlikely due to the high oxidation potential of thisnitrone (E(DEPMPO�+/DEPMPO) = 2.24 V versus NHE in acetonitrile[47]). On the other hand water is a very poor nucleophile and bind-ing of a metal ion to the negatively charged oxygen of the nitronylfunction is required for the Forrester–Hepburn reaction to occursubstantially [42]. In line with Fig. 1C and H, the recent finding thatphosphate anions can link to cerium and modify the Ce3+/Ce4+ re-dox cycle at the surface of nano-CeO2 [48] suggests an indirectinvolvement of these DTPA-chelatable ions despite they do notparticipate directly in the observed build up of DEPMPO–OH.

3.2. Evidence against Ce3+– and nano-CeO2-catalyzed hydroxyl radicalformation from hydrogen peroxide by a Fenton-like reaction

A proposed explanation for the toxic properties of nano-CeO2

reported in some biological systems was that, as Fe2+ or Cu+ salts,Ce3+ is capable of splitting H2O2 into harmful HO� species by a Fen-ton reaction [19,20]. Upon mixing 8 mM H2O2, 22 mM DEPMPOand 10 mM nano-CeO2 in KH–DTPA a strong EPR signal was imme-diately observed, consisting of a mixture of DEPMPO–OH (63%) and37% DEPMPO–OOH, the DEPMPO/O2

�� spin adduct (Fig. 2A). In gen-eral the best way to simulate DEPMPO–OOH EPR signals is to con-sider the major trans diastereoisomer is subjected to a dynamicchemical exchange between two rotamers [26,36]. However forthe simulation shown in Fig. 2A we obtained a good fit by assum-ing the trans-DEPMPO–OOH signal is composed of two static com-ponents with equal contribution, having the hyperfine splittingconstants: aN = 13.20 (13.12) G; aP = 50.76 (49.61) G; aHb = 11.60(10.38) G; aHc1 = 1.01 (0.62) G and aHc2 (6 H) = 0.39 (0.29) G. Inagreement with a previous study [36] the weak additional signalseen in Fig. 2A (see the low-field, non-composite doublet in the ex-panded region) was assigned to cis-DEPMPO–OOH (aN = 13.35 G;aP = 40.59 G; aHb = 9.95 G; aHc = 1.35 G; �25% of the total DEP-MPO–OOH signal) and we postulate it may be the non attributedspecies reported in a recent EPR study describing the interactionof H2O2 with CeCl3 [20].

A

D

20 G

simulated

simulated

C

E

F

simulated

× 10

× 10

× 4

DEPMPO-OH

N

(EtO)2(O)P

H3C

H

O

OH

DEPMPO-OMe

N

(EtO)2(O)P

H3C

H

O

OCH3

DEPMPOX

N

(EtO)2(O)P

H3CO

O

B

DEPMPO+nano-CeO2/H2O

+ 22% DMSO

+ 1 mM DTPA/Buffer

DEPMPO+nano-CeO /MeOH2

DEPMPO+Ce /H O3+

2

× 10

DEPMPO+Ce(SO ) /H O24 2

DEPMPO+CeO2/H2O

× 10 + 1 mM DTPA/Buffer

G

H

Fig. 1. EPR spectra following the addition of nano-CeO2 or typical Ce(III) or Ce(IV) salts (10 mM) to DEPMPO solutions. (A) From nano-CeO2 in water. (B) Same as A in thepresence of 22% DMSO. (C) Same as A in KH2PO4 (20 mM)-DTPA (1 mM). (D) From nano-CeO2 in pure methanol. (E) From Ce2(CO3)3 in water. (F) From Ce(SO4)2 in water. (G)From CeO2 in water. (H) Same as G in KH2PO4 (20 mM)-DTPA (1 mM). DEPMPO concentration was 50 mM except in (B) and (F): 22 mM. The simulations correspond to thefollowing adducts and sets of main hyperfine splitting constants: (A): 46% cis- + 54% (trans)-DEPMPO–OH: aN = 14.05 (14.05) G; aP = 47.29 (47.29) G and aHb = 14.24 (12.73) G;(D): 24% cis- + 76% (trans)-DEPMPO-OMe: aN = 12.86 (13.03) G; aP = 46.91 (40.16) G; aHb = 7.29 (7.48) G; (F): DEPMPOX: aN = 7.14 G; aP = 38.69 G; aHb = 3.69 G (2H). Single-scan spectra were acquired at 9.79 GHz 1–3 min after mixing using parameters: resolution, 4096 points/scan; microwave power, 10 mW; modulation frequency, 100 kHz;modulation amplitude, 0.44 G (except for (E): 0.79 G); time constant, 40.96 ms; gain, 2 � 104 (except for (D): 3.2 � 104 and (E): 5 � 104); sweep rate, 3.81 G/s except for (D):1.91 G/s. Spectral intensities scalings are relative to trace A.

166 M. Culcasi et al. / Chemico-Biological Interactions 199 (2012) 161–176

When 1–10 mM of the Ce(IV) compounds Ce(SO4)2 and CeO2

were used instead of nano-CeO2, a slightly weaker EPR signal ap-peared immediately, with a superoxide versus hydroxyl radical ad-duct proportion above 70% (Fig. 2C). However, using 10 mM of theCe(III) derivatives Ce2(SO4)3 or CeCl3 in the system, mixtures ofDEPMPO–OOH (>80%) with DEPMPO–OH only appeared whenincubation was prolonged for more than 25 min (Fig. 2D and E).Furthermore, addition of 20 mU/ml SOD in all [H2O2-DEPMPO–Ce

compound] systems caused no change of the detected EPR signal(Fig. 2F). Because of the strong chelating properties of KH–DTPAused in these experiments, the results at this point suggest thatnano-CeO2 catalyzes DEPMPO adduct formation from H2O2 moreefficiently than its Ce(III) and Ce(IV) components, by a mechanismnot involving formation of O2

�� or nucleophilic synthesis of DEP-MPO–OH. Thus the detection of DEPMPO–OOH in Ce(IV) experi-ments signals here appears consistent with the earlier

A

B

25 G

simulated

G2.5 G

Cerium compounds Iron (II)

× 10

H

C× 10

× 10

× 10

× 10 × 10

D

IEHydrogen peroxide

× 10F

Fig. 2. EPR spectra of DEPMPO radical adducts obtained during incubation of hydrogen peroxide alone or under Fenton conditions using Ce compounds (left part) or Iron(II)(right part) as catalysts. Samples contained 22 mM DEPMPO and 1 mM DTPA in 20 mM phosphate buffer, pH 7.0 and were incubated at room temperature for 1–3 min, exceptwhen indicated. The concentrations of Ce compounds and FeSO4 were 10 and 1 mM, respectively. (A) nano-CeO2 + 8 mM H2O2; (B) same as (A), in the presence of 15% DMSO;(C) Ce(SO4)2 + 8 mM H2O2; (D) CeCl3 + 8 mM H2O2; (E) same as (D), after 1 h incubation; (F) same as (E) in the presence of 20 mU/ml SOD; (G) FeSO4 + 1 mM H2O2; (H) same as(G), in the presence of 15% DMSO; (I) H2O2 (8 mM) alone, after 1 h incubation. Simulations were consistent with (G) DEPMPO–OH and (H) DEPMPO–Me in studies withIron(II), and with DEPMPO–OOH/DEPMPO–OH mixtures with superoxide adduct contribution of (A) 58%; (B) 63%; (C) 72%; (E) 82%; (F) 62%; (I) 77% in other studies.Spectrometer settings were as in the legend of Fig. 1, except receiver gain, 4 � 104 (A–E) and 2 � 104 (F–H), and number of accumulated scans, 6 (E, F, I). The selected regionshows the low-field doublet of cis-DEPMPO–OOH. Spectral intensities scalings are relative to traces (G) and (H).

M. Culcasi et al. / Chemico-Biological Interactions 199 (2012) 161–176 167

mechanism proposed by Czapski et al. [49] for the reduction of Ce4+

to Ce3+ by H2O2 in acidic medium, via intermediate formation ofthe perhydroxyl radical HOO�, the conjugate acid form of O2

��

(Eq. (2) and (3)):

Ce4þ þH2O2 ! Ce3þ þHOO� þHþ ð2Þ

Ce4þ þHOO� ! Ce3þ þ O2 þHþ ð3Þ

To clarify if HO� is nevertheless formed in [nano-CeO2 + H2O2]mixtures we checked for the presence of spin adducts from car-bon-centered radicals by adding excess competitors such as etha-nol or DMSO. When 15% DMSO was added to a [Fe2+

(1 mM) + H2O2 (1 mM)] Fenton reagent in the presence of 22 mMDEPMPO the expected strong DEPMPO–OH signal of Fig. 2G wasimmediately replaced by the signal of the methyl radical adduct(DEPMPO-Me; aN = 15.14 G; aP = 47.66 G; aHb = 22.09 G; Fig. 2H)[36]. However repeating the competition reaction with ethanolor DMSO on 10 mM nano-CeO2 (Fig. 2B) or any of the other tested

Ce compounds did not significantly affect the control [DEPMPO–OOH + DEPMPO–OH] mixture during the first 30 min followingincubation.

As these data, in particular those for Ce(III) compounds, are indisagreement with an initial formation of HO� as proposed byHeckert et al. for CeCl3 [20] (Eq. (1)), we used an alternative meth-od by evaluating the extent to which Ce compounds can cause oxi-dative damage to deoxyribose [41]. Table 1 lists the percentages ofdeoxyribose (1 mM) degradation in phosphate buffer at pH 7.0 fortest compounds and combinations as compared to the value foundwhen the Fenton reagent [FeSO4 (0.2 mM) + H2O2 (1 mM)] was ap-plied for 1 h at 37 �C. In this system, Fe2+-induced formation ofMDA–TBA dose-dependently decreased to the baseline level (i.e.,in the absence of iron or H2O2) by addition of the HO� scavengermannitol in the range 0.2–15 mM, all absorbances being similarwhen HEPES was used instead of KH2PO4 (not shown). Of thetested Ce compounds used instead of Iron(II) in this assay onlynano-CeO2 (up to 1 mM) and, in a lesser extent, Ce(IV) derivativescaused a weak, yet significant concentration-dependent damage todeoxyribose but since this effect was not modified by 15 mM man-

Table 1Effect of hydrogen peroxide and various ions and Ce compounds in the degradation ofdeoxyribose.

Added compound % of Deoxyribose degradation

+H2O2 (1 mM) no H2O2

Fe(II) as FeSO4 (0.2 mM) 100a 0.36 ± 0.01no Fe2+ 0.79 ± 0.01 0.23 ± 0.01Fe(II) as FeSO4 (0.2 mM) + mannitol 0.90 ± 0.01 0.19 ± 0.01Ce(IV) as Ce(SO4)2 (0.2 mM) 1.66 ± 0.01 2.04 ± 0.10Ce(IV) as Ce(SO4)2 (0.2 mM) + mannitol 1.91 ± 0.01Ce(IV) as Ce(SO4)2 (1 mM) 4.09 ± 0.36* 5.46 ± 0.41*

Ce(IV) as Ce(SO4)2 (1 mM) + mannitol 4.14 ± 0.28* 5.54 ± 0.61*

Ce(IV) as CeO2 (0.2 mM) 0.47 ± 0.01Ce(IV) as CeO2 (1 mM) 1.13 ± 0.01Ce(IV) as (NH4)2Ce(NO3)6 (0.2 mM) 1.43 ± 0.16Ce(IV) as (NH4)2Ce(NO3)6 (1 mM) 2.98 ± 0.45Ce(IV) as (NH4)2Ce(NO3)6 (1 mM) + mannitol 2.57 ± 0.56Ce(III) as CeCl3 (0.2 mM) 0.74 ± 0.01Ce(III) as CeCl3 (1 mM) 1.54 ± 0.01 2.54 ± 0.11Ce(III) as CeCl3 (1 mM) + mannitol 1.62 ± 0.01 2.46 ± 0.47nano-CeO2 (0.2 mM) 5.35 ± 0.33* 7.83 ± 0.54*

nano-CeO2 (0.2 mM) + mannitol 5.52 ± 0.16* 6.72 ± 0.61*

nano-CeO2 (1 mM) 11.05 ± 0.36* 12.86 ± 0.72*

nano-CeO2 (1 mM) + mannitol 10.46 ± 0.25* 12.75 ± 0.81*

NO3� as KNO3 (0.2 mM) 0.67 ± 0.01

NO3� as KNO3 (1 mM) 0.55 ± 0.01

a Data are calculated relative to the value for [FeSO4 + H2O2] set at 100% and areexpressed as means ± SD (n = 3). One-way ANOVA followed by Duncan t test.* P < 0.05 versus iron-free buffer and [FeSO4 + H2O2]. The incubation mediumcontained 1 mM deoxyribose in EDTA (1.1 mM)-supplemented phosphate buffer(20 mM), pH 7.4. The concentration of the HO� scavenger mannitol was 15 mM.

168 M. Culcasi et al. / Chemico-Biological Interactions 199 (2012) 161–176

nitol HO� radicals are probably not involved. As a confirmationomitting H2O2 in the samples had no significant effect on MDA–TBA release suggesting deoxyribose degradation may result fromdirect oxidation by ceric ions, which are known to oxidize sugarssuch as glucose [50]. We also checked that this oxidation wasnot induced by NO3

� (a constituent of nano-CeO2) when intro-duced instead of Fe or Ce in the sample (Table 1).

If the two above in vitro approaches suggested that the directinteraction between Ce3+ and H2O2 does not rapidly form HO�,the mechanism by which the mixtures of DEPMPO adducts seenin Fig. 2A–F and reported in [20] are formed with a delay is still un-known. In an attempt to gain insights we found that omitting CeCl3

in the mixture that yielded the spectrum of Fig. 2E afforded a verysimilar background signal after 1 h incubation (Fig. 2I), where stillDEPMPO–OOH was the dominant species (77%) over DEPMPO–OH.Although H2O2 is not prone to significant daylight homolysis it ispossible that low levels of HO� progressively form, which can thenreact with excess hydrogen peroxide to form HOO�, both radicalspecies being trapped by DEPMPO.

3.3. In human fibroblasts micromolar concentrations of nano-CeO2

induced ROS formation at non-toxic levels via NADPH oxidaseactivation

The cell effects of nano-CeO2 in relation to ROS formation[12,13,16] or inhibition [4,5] have been studied by many investiga-tors previously and significant changes in a variety of biochemicalindices of oxidant stress were found to occur over relatively longperiods (>24 h). Since the results of Fig. 1C evidenced weak butdetectable artifactual DEPMPO–OH formation when the nitronewas in contact with nano-CeO2 in buffers, we considered that thebest spin trapping strategy to evaluate ROS release (especiallyH2O2) from cells exposed to the particles would be to avoid pro-longed coincubation with DEPMPO.

We first investigated the effects of nano-CeO2 on human fibro-blasts at several concentrations relevant to common chronic expo-

sure situations, i.e., within the range 10�5–10�1 g/l, whichcorresponds to 0.036 lM–0.36 mM [5,12,13,22,24]. Incubatingthe cells at 37 �C for 24 h with the lowest doses of nano-CeO2

(i.e., 10�5–10�4 g/l) resulted in no significant cytosolic release ofLDH into the medium as compared to control cells incubated inDMEM, which had a LDH activity of 10.7 ± 1.9 U/l/mg protein. ThisLDH release in cell supernatant did not exceed 2% of total cell LDHactivity (i.e., 689 ± 12.4 U/l/mg protein). Cell monolayer stainingwith Trypan blue showed that none of these two lowest doses in-duced cell death after a 24-h incubation with the particles. Increas-ing the nano-CeO2 dose resulted in a decrease of cell viability, asvizualized in Fig. 3A for 10�1 g/l. The same dose-related loss of via-bility was observed using the MTT assay (Fig. 3B). In contrast incu-bating the cells for 24 h with nano-CeO2 induced no significantmitochondrial toxicity even et the highest tested dose, as assessedby Rhodamine 123 staining assay (Fig 3C). These results are com-parable to that of Alili et al. [22] who recently reported no toxicproperties of nano-CeO2 up to 10�1 g/l in human dermal fibro-blasts. Interestingly, application of the highest nano-CeO2 concen-trations for 24 h resulted in a significant increase of total cellularprotein content, reaching 1.71 ± 0.04 mg/ml and 1.53 ± 0.06 mg/ml for 6 � 10�1 g/l and 6 � 10�4 g/l, respectively as compared tothe normal level of 0.76 ± 0.03 mg/ml in control cells (P < 0.05).This protein biosynthesis, which mostly occurred at a non-toxicconcentration of nano-CeO2, paralleled with the increase of the to-tal cell count.

Earlier studies have shown that low doses of CeCl3 stimulatedcell proliferation in cardiac fibroblasts [51] and the synthesis ofcollagen in the rat heart in vivo [18], that could possibly be medi-ated by an early formation of O2

�� capable of promoting growth re-sponses prior to any modifications in cell morphology and viability[51]. On the other hand, pre-incubated nano-CeO2 has been shownto remain internalized within the cytoplasm and vesicles of fibro-blasts even after external particles were completely removed fromculture plates by washing [14]. From these considerations, it couldbe of interest to explore any potential delayed ROS production pro-moted by nano-CeO2 after removal of the medium.

In this context the washing protocol following nano-CeO2 expo-sure, which was considered to comply with our required lack ofprolonged contact between the Ce particles and the spin trap,could also give crucial information on the behavior of particles ascontinuous intracellular ROS generators. Therefore it was adoptedin the next EPR studies where the culture medium of fibroblastsincubated for 2 or 24 h with graded concentrations of nano-CeO2

was then carefully washed with PBS before a 20 mM DEPMPO solu-tion in DTPA (0.3 mM)-supplemented PBS was introduced. DEP-MPO-trappable species still released in the medium at this timewere then allowed to accumulate for 15 min at 37 �C in order toimprove EPR detection. A control sample obtained by mixing6 � 10�4 g/l nano-CeO2 with 20 mM DEPMPO in DMEM for15 min gave no detectable background EPR signal (Fig 4A).

Preliminary cytotoxicity studies showed no significant effect onLDH activity when normal fibroblasts were incubated for 1 h witheither 20 or 40 mM DEPMPO (6.4 ± 0.9 and 9.2 ± 2.3 U/l/mg pro-tein, respectively) compared to DMEM-treated cells (10.7 ± 1.9 U/l/mg protein, <2% of total LDH activity). Incubation of the cellsfor 2 h in plain DMEM gave no EPR signal in the supernatant(Fig. 4B) whereas strong DEPMPO–OH signals were detected when6 � 10�5 g/l nano-CeO2 was applied during the incubation phase(Fig. 4C). The mean DEPMPO–OH concentration significantly in-creased when either the incubation time or nano-CeO2 concentra-tion increased, e.g., it was almost 4-fold higher when shifting from6 � 10�5 g/l to 6 � 10�4 g/l nano-CeO2 and 24 h incubation(Fig. 4F). For both nano-CeO2 concentrations and incubation times,adding 2 mU/ml CAT or [2 mU/ml CAT + 5 U/ml SOD] during incu-bation provoked a significant partial or total inhibition of the DEP-

Fig. 3. Effect of nano-CeO2 on viability and apoptosis of human fibroblasts. Cells were exposed to the particles (10�5�1 g/l, corresponding to the range 0.036 lM�3,66 mM)for 24 h (A�D) or 48 h (E) before processing and analysis. (A) Trypan blue staining. (B) MTT assay. (C) RH-123 fluorescence. (D) Effect of NADPH inhibitors using MTT assay.Cells were pretreated with the NADPH oxidase inhibitors PAO (0.1 mM) or ACV (0.3 mM) for 30 min or 4 h, respectively, before being exposed to 6 � 10�1 g/l nano-CeO2 infresh buffer. DPI was added 1 h before (5 lM) or at the 23rd hour (10 lM; DPI after nano) of nano-CeO2 exposure (E) Annexin V and 7-AAD staining. Symbols represent: h,viable cells (Annexin V- and 7-AAD�); P, early apoptotic cells (Annexin V + and 7-AAD�), and j, late apoptotic cells (Annexin V + and 7-AAD+). Data represent means ± SEM(n = 3–6 independent experiments performed in triplicate). One-way ANOVA followed by Newman–Keuls test: ⁄P < 0.01 versus respective control in DMEM and §P < 0.05 forDPI and PAO pretreated cells versus cells exposed to 6 � 10�1 g/l nano CeO2.

M. Culcasi et al. / Chemico-Biological Interactions 199 (2012) 161–176 169

MPO–OH signal, respectively (Figs. 4D and E). This observationstrongly suggests that cells loaded with nano-CeO2 arecontinuously activated to release H2O2 and O2

��. The intra- versusextracellular site of their interaction with the spin trap yet dependson several factors such as cell penetration of DEPMPO [52] and therelatively low stability of DEPMPO–OOH towards cell reductants or

transition metals [26,52,53]. The partial inhibition by SOD of theEPR signal in Fig. 4D could be explained considering that evenwhen exogenously added, the enzyme can scavenge intracellularlyformed O2

�� [53]. Of interest, formation of DEPMPO–OH adductsconsecutive to extracellular release of H2O2 has been documentedpreviously in murine fibroblasts in situation of oxidant stress [38].

Fig. 4. DEPMPO–OH formation following exposure of human fibroblasts to nano-CeO2 and effect of inhibitors. After removal of Ce particles from the culture medium cellswere reincubated in DEPMPO (20 mM)- and DTPA (0.3 mM)-containing PBS for 15 min at 37 �C. EPR spectra were obtained in the supernatant by signal averaging 2 scansusing the spectrometer settings given in the legend of Fig. 1, except for modulation amplitude, 0.497 G; receiver gain, 2 � 105 and sweep rate, 3.34 G/s. (A) EPR signalfollowing a 15-min contact between 6 � 10�4 g/l nano-CeO2 and 20 mM DEPMPO in DMEM. Other typical EPR signals were recorded after incubation for 2 h with; (B) plainDMEM; (C) 6 � 10�5 g/l nano-CeO2; (D) same as; (C) + 2 mU/ml catalase; (E) same as; (D) + 5 U/ml SOD. (F) Variation of DEPMPO–OH levels with exposure time, nano-CeO2

concentration and inhibitors. As described in Fig. 3, concentrations of PAO and ACV were 0.1 mM and 0.3 mM, respectively and data represent means ± SEM (n = 6–12). One-way ANOVA followed by Newman–Keuls test: ⁄P < 0.05 versus nano-CeO2 with same concentration and incubation time; §P < 0.05 versus 2 h nano-CeO2.

170 M. Culcasi et al. / Chemico-Biological Interactions 199 (2012) 161–176

Of the mechanisms that could lead to the CAT-inhibitable DEP-MPO–OH signal of Fig. 4E a Fenton reaction involving mobilizedtransition metals such as iron, but not Ce3+ (see above) is likely tooccur instead of nucleophilic synthesis because of the presence ofDTPA in the buffer. It has also been shown that human fibroblastscan produce large amounts of H2O2/O2

�� over many hours by stim-ulation of NADPH oxidase located in the outer membrane [54].

Pertinent to this hypothesis, the two NADPH oxidase inhibitors,ACV (0.3 mM) and PAO (0.1 mM), decreased the EPR signal by 60and 85%, respectively (Fig. 4F). In these experimental conditionsPAO and ACV did not significantly affect the cell viability in theMTT assay when preincubated before nano-CeO2 exposure at6 � 10�4 g/l. Thus viabilities for these ACV- or PAO-treated cells were90.3 ± 1.5% and 97.0 ± 1.2%, respectively as compared to 91.3 ± 1.8%in nano-CeO2 treated control cells. Since these inhibitions of ROS pro-duction were not complete, as expected from the optimal low toxicconcentrations used [54,55], NADPH oxidase may not be the uniquecellular source of ROS being activated by nano-CeO2.

To understand the apparent discrepancy between the lack ofglobal cytotoxicity of the low doses of nano-CeO2 (up to 10�1 g/l)and the fact that they yet trigger ROS generating systems such asmembrane NADPH oxidase, we looked for earlier indices of oxidantstress. Thus experiments were repeated on human fibroblasts inorder to assess the time course of nano-CeO2-induced ascorbate re-lease or lipid peroxidation, and the effects of inhibitors of O2

�� and/or H2O2, or NADPH oxidase. In biological fluids AFR–DMSO EPR sig-nal (giving a doublet with aH = 1.81 G) is a reliable marker of ascor-bate concentration [31]. Fig. 5A shows that ascorbate was mostlyreleased in the cell supernatant at 2 h incubation, at a concentra-tion dependent on the nano-CeO2 dose. Again, the key role ofH2O2 and NADPH oxidase activation in this early cell damagewas supported by the observed inhibition by 2 mU/ml CAT and0.1 mM PAO. The EPR signal was also significantly decreased by

coincubation with 0.1 mM DEPMPO, likely by a mechanism involv-ing direct scavenging of free radicals at the nitronyl site [38]. Par-allel to this ascorbate leakage fibroblasts exposed to Ce particlesshowed increased lipid peroxidation, with a peak of membraneMDA–TBA formation at 24 h incubation of 6 � 10�4 g/l nano-CeO2, and a similar inhibition profile (Fig. 5B).

Together the results presented in this section tend to demon-strate that micromolar concentrations of nano-CeO2 can partiallyactivate NADPH oxidase to release ROS such as H2O2/O2

�� (with apreferential role of Ce(IV) redox state). Rather than sustaining sig-nificant lipid peroxidation and cell death, this activation has thecharacteristics of cell signaling, leading to protein synthesis andcell proliferation. These particular properties may explain whylow nano-CeO2 has been found either detrimental in cardiac stud-ies [18,51] or neuroprotective in in vitro [4] and in vivo [8] exper-iments by increasing cell regeneration and survival. This generalrole in ROS-mediated signaling is shared by Nox1–5 enzymeswhich specifically regulate physiological processes including cellgrowth, apoptosis or differentiation [56]. Thus the next step ofour study was to focus on the specific role of Nox2 and Nox4 iso-forms because (i) they are localized in endosomes, or in the nu-cleus and the endoplasmic reticulum, respectively [56], allsubcellular sites that can be reached by nano-CeO2 agglomeratesas they are uptaken in the cytoplasm [5,13,14,21], and (ii) humanfibroblasts mainly express these two proteins [57].

3.4. Simultaneous stimulation of NADPH oxidase and mitochondrialsources of ROS are involved in the toxicity of millimolar concentrationsof nano-CeO2 in human fibroblasts

To assess the role of Nox2, Nox4 and identify other ROS sourcessustaining the development of toxicity in human fibroblasts weincubated the cells for 24–48 h with higher (above 10�1 g/l) nano-

Fig. 5. Levels of AFR–DMSO and MDA–TBA following short- and long-term exposure of human fibroblasts to non toxic concentrations of nano-CeO2, and effect of inhibitors.(A) EPR signals of ascorbyl free radical (AFR) were obtained at 9.8 GHz from cell supernatant:DMSO (1:1) mixtures by signal averaging 4 scans with instrument parameters:microwave power, 10 mW; modulation frequency, 100 kHz; modulation amplitude, 0.883 G; time constant, 81.92 ms; gain, 5 � 105 and sweep rate, 1.07 G/s. (B) Lipidperoxidation was measured in homogenates from untreated (DMEM) or nano-CeO2-treated cells. Concentrations of inhibitors were as in the legend of Fig. 3, and DEPMPO was0.1 mM. Data represent means ± SEM (n = 6–12). One-way ANOVA followed by Newman–Keuls test: ⁄P < 0.05 versus nano-CeO2 with same concentration and incubationtime;§P < 0.05 versus 2 h nano-CeO2.

M. Culcasi et al. / Chemico-Biological Interactions 199 (2012) 161–176 171

CeO2 concentrations. Cell damage occurred when 6 � 10�1 g/l orabove nano-CeO2 was used, as significant LDH was released extra-cellularly (LDH activity was 137.08 ± 5.1 U/l/mg protein; P < 0.05versus control), corresponding to 22.1 ± 2.6% of control total cellLDH. This cell necrosis was concomitant to a�30–50% loss of viabil-ity, as demonstrated by the MTT assay (Fig. 3B). This finding is inline with the reported 45% impairment of the metabolic mitochon-drial dehydrogenase activity of human fibroblasts exposed to6 � 10�1 g/l nano-CeO2 for 24 h using the WST-1 assay [14]. Inour study despite there was significant cell death (Fig. 3B) anddetachment (not shown) at these high nano-CeO2 concentrations,staining with either Rhodamine 123 or Hoechst 33342 did not showany evidence of significant compromized mitochondrial function orsign of chromatin condensation, respectively (data not shown).Pharmacological NADPH oxidase inhibition by pretreatment with0.1 mM PAO partially reduced the cellular toxicity of 6 � 10�1 g/l

nano-CeO2, while ACV was not efficient to protect cells (Fig. 3D).This protective effect of PAO could be related to the finding that asimilar dose of PAO provided 90% inhibition of H2O2 production inhuman fibroblasts after only 10 min incubation [54]. In our studythe failure of ACV to protect cells might reflect a more toxic and/or pro-oxidant activity of the inhibitor, a feature already reportedin non phagocytic cells [58]. On another hand, preincubation ofDPI, a general flavoprotein inhibitor that almost completely blocksNADPH oxidase activity as compared to PAO or ACV [59], again par-tially preserved the viability of cells exposed to high nano-CeO2

doses (Fig. 3D) showing that NADPH oxidase is likely one but notthe sole of the ROS source involved in cell injury. This effect waseven less marked when DPI was added only at the 23rd hour ofnano-CeO2 treatment.

Taken together, the results suggests that in the presence ofnano-CeO2, mitochondria may also function as a source of signal-

172 M. Culcasi et al. / Chemico-Biological Interactions 199 (2012) 161–176

ing molecules (such as H2O2/O2��) that initiate early mechanisms

such as cell proliferation and apoptosis, leading to mitochondrialdysfunction as described for several other toxic metals [60 and ref-erence therein]. As a confirmation, we found a significant amountof early apoptotic cells upon incubation for 48 h with 6 � 10�1 or1 g/l nano-CeO2 (Fig. 3E).

Using the spin trapping protocol of Fig. 4 the mean DEPMPO–OH levels recovered in the supernatant of normal cells exposedto cytotoxic nano-CeO2 were 25–60 times more elevated thanwhen non toxic concentrations were applied for the same period(Fig. 6A) showing an increased ROS production and suggesting an-other source than NADPH oxidase has been activated. IndeedNADPH oxidases and mitochondrial respiratory chain representtwo main sources of H2O2/O2

�� production in cells. Thus we inves-tigated whether this second source of ROS was implicated by pre-treating the cells for 1 h with 20 lM mitoPBN, a lipophilic,mitochondria-targeted spin trap reported to efficiently protectcells from the consequences of O2

�� formation [28]. Whereas notoxicity was found for mitoPBN at 50 lM in normal untreatedfibroblasts (with a mean LDH release of 12.7 ± 2.4 UI/ mg protein,n = 6) a lower concentration of 20 lM significantly decreased DEP-MPO–OH formation by ca. 60% in treated cells (Fig. 6A). These re-sults unambiguously demonstrate the mitochondrial source ofROS is activated by cytotoxic doses of nano-CeO2.

Fig. 6. Involvement of NADPH oxidase and mitochondria in ROS production from humanfollowing nano-CeO2 exposure in human fibroblasts determined by the cell washing, DEPmethods. Preceding the spin trapping phase, the added inhibitors in control cell medium wof nano-CeO2 exposure) or the specific mitochondrial radical inhibitor mitoPBN (20 lMfibroblasts transfected either with no plasmid (control WP), p47phox wild-type, domrespectively). Inset represents p47phox protein expression and their respective loading coexpressed as percentage of LDH released into the cell medium with respect to totaltransfected human fibroblasts as described in (B). Hydrogen peroxide after 24-h exposexpression in untreated and nano-CeO2 exposed human normal fibroblasts. The right panactin (from 2 independent experiments). Data on ROS production represent means ± SEnano-CeO2 treatment in corresponding control cells. Data on LDH release represent mANOVA followed by Newman–Keuls test: �P < 0.05 versus p47-M8 nano-CeO2 treated ce

Next, we compared the mitochondrial versus NADPH oxidasesources of ROS that become activated by 6 � 10�1 g/l nano-CeO2

by measuring the effect on the DEPMPO–OH signal of DPI. Com-pared to nano-CeO2-treated (24 h) fibroblasts, a 1-h pretreatmentor addition to the particles for the last incubation hour withDPI decreased DEPMPO–OH levels by 80 and 50%, respectively(Fig. 6A). This result implies that overstimulation of NADPH oxi-dase also participates in the observed ROS-induced cytotoxicityof the high nano-CeO2 concentration.

It was reported that DPI can also inhibit mitochondrial ROS pro-duction [59], that is NADPH oxidase inhibition may not account forthe effect on DEPMPO–OH seen in Fig. 6A. Therefore to gain insightinto the involvement of NADPH oxidase in the cytotoxicity of nano-CeO2 we used a molecular approach with transfected cells. SinceNox2-dependent ROS production relies on p47phox phosphoryla-tion, a function critical to assembly and activation of the oxidase[56,57], the effect of the overexpression of wild-type p47phox,dominant negative mutant of p47phox (p47-M1) and constitutiveactive mutant of p47phox (p47-M8) on nano-CeO2 (6 � 10�1 g/l)-induced DEPMPO–OH formation was first evaluated. It has beenshown that P47phox needs to be phosphorylated on different ser-ine residues to bind p22phox and activate NADPH oxidase activity[33]. Also, the P47-M1 mutant inhibits the binding of p47phox top22phox and the P47-M8 mutant mimics serine phosphorylation

fibroblasts ROS exposed to 6 � 10�1 g/l nano-CeO2 for 24 h. (A) DEPMPO–OH levelsMPO (20 mM) spin trapping and EPR recording protocols described in materials andere the NADPH oxidase inhibitor DPI (1 h before (5 lM) or at the 23rd hour (10 lM); 1 h before nano-CeO2 exposure). (B) DEPMPO–OH levels in nano-CeO2-treated

inant negative or constitutive active mutant of p47phox (p47-M1 and p47-M8ntrol (actin) in untreated transfected cells. (C) Lactate dehydrogenase (LDH) release,

LDH, determined after 24-h exposure with or without 6 � 10�1 g/l nano-CeO2 inure at 500 lM in the medium was used as a positive control. (D) Nox4 and actinel represents the densitometric analysis of nano-CeO2 on Nox4 expression relative toM (n = 6–12). One-way ANOVA followed by Newman–Keuls test: ⁄P < 0.05 versuseans ± SEM (n = 4–6 independent experiments performed in triplicate). One-waylls.

Table 2Cell viabilitya and oxidative stress-related parameters in murine 3T3 cells following exposure to nano-CeO2 for 24 h and effect of specific inhibitors.

Inhibitor Viability SOD CAT GPx Protein MDA–TBA GSH Ascorbateactivity activity activity carbonyls

(%) (UI/106 cells) (mUI/106 cells) (nmol/mg) (nmol/10 mg prot) (nmol/106 cells)

none (DMEM) 100 8.7 ± 0.9 36.5 ± 2.6 29.5 ± 2.0 2.98 ± 0.75 8.3 ± 1.8 20.8 ± 0.01 5.35 ± 0.9

[nano-CeO2] = 6 � 10�4 g/lnone 99.1 ± 0.1 10.1 ± 2.8 48.6 ± 3.1* 37.9 ± 2.1* 3.09 ± 0.11 14.1 ± 2.7* 33.7 ± 1.4* 3.93 ± 0.1*

CAT 100.1 ± 0.2 10.8 ± 1.1 41.9 ± 2.2 28.6 ± 3.1 3.02 ± 0.09 10.1 ± 1.9 16.4 ± 1.6§ 6.24 ± 0.7[CAT + SOD] 99.9 ± 0.3 11.8 ± 2.3 37.7 ± 1.2§ 28.1 ± 1.1§ 3.45 ± 0.11 6.9 ± 1.4§ 19.9 ± 1.1§ 6.90 ± 0.2§

PAO 96.1 ± 0.4 12.7 ± 1.8 38.8 ± 1.9 29.2 ± 1.0 2.78 ± 0.07 9.6 ± 2.2 20.6 ± 2.7§ 6.78 ± 0.4§

[nano-CeO2] = 6 � 10�1 g/lnone 64.1 ± 2.4* 4.5 ± 0.5* 18.5 ± 2.4* 16.8 ± 1.7* 4.99 ± 0.67* 27.7 ± 1.5* 9.8 ± 1.1* 2.4 ± 0.9*

CAT 86.1 ± 0.9§ 5.9 ± 0.8 22.9 ± 1.9* 18.9 ± 1.1* 3.90 ± 0.27 14.9 ± 1.2* 13.9 ± 2.0* 2.9 ± 0.8*

[CAT + SOD] 91.1 ± 1.9§ 7.7 ± 1.1§ 27.8 ± 0.9§ 25.2 ± 1.1 2.81 ± 0.09§ 10.3 ± 1.2§ 17.6 ± 0.9*§ 4.8 ± 1.1§

PAO 76.1 ± 1.1 6.8 ± 0.9 23.9 ± 1.2* 18.0 ± 2.1* 3.99 ± 0.24 19.8 ± 1.7* 13.9 ± 1.0* 3.9 ± 0.9*

mitoPBN 89.1 ± 2.1§ 8.9 ± 1.1§ 30.9 ± 1.9§ 25.9 ± 1.7 2.70 ± 0.91§ 9.2 ± 2.9§ 20.3 ± 0.9§ 5.1 ± 0.8§

a Measured by the MTT assay. Data are expressed as means ± SEM (n = 8–14 independent experiments in duplicate/group). The concentrations of added inhibitors were:CAT, 2 mU/ml; SOD, 5 U/ml; PAO, 0.1 mM and mitoPBN, 20 lM. One-way ANOVA followed by Duncan t test.* P < 0.05 versus cells incubated in DMEM.§ P < 0.05 versus nano-CeO2-treated cells at the corresponding dose.

M. Culcasi et al. / Chemico-Biological Interactions 199 (2012) 161–176 173

necessary to activate NADPH oxidase [33]. In line with these dataFig. 6B shows that control transfected cells (control cells withoutplasmid, WP) and wild-type p47phox transfected cell in absenceof nano-CeO2 do not display any significant EPR signal. In nano-CeO2 treated cells a same extent of ROS production was observedin control WP transfected cells and in cells over-expressingwild-type p47phox. ROS production in p47-M1 transfected cellswas decreased by 50%, compared to control transfected cells andwild-type p47phox cells, while it significantly increased withp47-M8, the constitutive active mutant of p47phox (+40% versuswild-type p47phox, P < 0.05). Together these data suggest thatnano-CeO2 exposure induces ROS formation in part through Nox2activation by a mechanism involving p47phox phosphorylationand translocation.

In parallel, LDH activity was monitored in supernatants fromtransfected cells showing that wild-type p47phox expression doesnot increase cell necrosis in absence of nano-CeO2 (Fig 6C) as com-pared to normal fibroblasts. As a positive control, a significant LDHrelease required WP control cells to be treated by 500 lM (Fig 6C),but not 0.5 lM H2O2 for 24 h, in line with the reported propertiesof the oxidant as a necrotic inductor when used in the millimolarrange [61]. In the presence of nano-CeO2 the expression of the con-stitutive active form of p47phox (p47-M8) lead to a significant con-comitant increase in ROS production (Fig 6B) and LDH release (Fig6C) compared to wild-type p47phox expression. In contrast, theexpression of the dominant negative form of p47phox (p47-M1)led to a significant decrease of nano-CeO2 induced ROS productionbut no significant decrease of LDH activity. These results suggestthat the p47-phox dependent NADPH oxidase is not the onlysource mediating nano-CeO2 toxicity in human fibroblasts. A pos-sible involvement of Nox4 might also be partly suggested asnano-CeO2 treatment induced an increase in Nox4 expression(Fig. 6D). In contrast to Nox2, Nox4 does not possess any identifiedregulatory subunit and an increase in Nox4-dependent ROS pro-duction relies exclusively on Nox4 protein induction [56]. To date,there is no particular clue on the pathway leading to the inductionof Nox4 in presence of nano-CeO2.

Finally, we evaluated the impact of Nox2 activation on nano-CeO2-induced protein synthesis. After 24 h incubation with6 � 10�1 g/l nano-CeO2 the total protein content in WT-p47phoxoverexpressing cells increased by 2.2 ± 0.4 times compared to un-treated cells (i.e., 0.59 mg/ml) while it decreased by 1.7 ± 0.3 andraised by 3.8 ± 0.4 times (n = 3) for the p47-M1 and p47-M8 mu-tants, respectively. Treatment with DPI decreased protein synthe-sis by 15 and 30% when administered before (5 lM) or at the

end (10 lM) of nano-CeO2 exposure, respectively, while mitoPBN(20 lM) had no effect (1.70 versus 1.69 mg/ml in nano-CeO2-over-expressing WT-p47phox cells).

Together the results presented in this section suggest that (i)Nox2 activation might participate to nano-CeO2 stimulation of pro-tein synthesis and that this activation is ROS-mediated and (ii) alarge amount of ROS are produced by the mitochondria, a phenom-enon which is likely in relation with the increase in cell mortality,decrease in mitochondrial dehydrogenase activity and early apop-tosis development.

3.5. Impact of nano-CeO2 exposure on the antioxidant and oxidativestress status in murine 3T3 fibroblasts (Table 2)

In a further series of experiments we investigated if the nano-CeO2-induced ROS formation evidenced in the above EPR studies(Figs. 4 and 6) would result in changes in cell viability and an arrayof biomarkers of oxidative stress. Being a more abundant cellsource than primary cultures of human cells, we used murine3T3 fibroblasts which were obtained at 105 cells/ml and incubatedwith 6 � 10�4–6 � 10�1 g/l nano-CeO2 for 24 h. As compared tountreated fibroblasts cell viability (evaluated by the mitochondrialdehydrogenase activity assay) at the end of incubation was not al-tered when the applied dose of nano-CeO2 ranged 6 � 10�4–6 � 10�2 g/l, while it was significantly impaired by 30% with6 � 10�1 g/l of the particles, a pattern paralleled by measurementsof extracellular LDH release (data not shown). In those cells sus-taining a normal viability in the presence of 6 � 10�4 g/l nano-CeO2, ROS formation was nevertheless demonstrated by (i) the sig-nificant variations of the levels of ascorbate (decreasing by 25%)and GSH (increasing by 150%), two antioxidants that function to-gether in the cell, and (ii) a slight but significant increase of lipidperoxidation. However these biochemical events are not detrimen-tal enough as to induce a significant production of protein carbon-yls in cell lysates, an end-point of protein oxidation whichirreversibly accumulate within cells [62]. From the increase ofspecific enzyme activities such CAT and GPx, but not SOD, or theeffects of added antioxidant enzymes or inhibitors, the participa-tion of the NADPH oxidase/H2O2 system in cell damage can be pos-tulated. Since GSSG levels fell under the detection limit when cellswere treated by 6 � 10�4 g/l nano-CeO2, the elevation of GSH levelsreflects mobilizations from GSSG (given that one mole of GSSG willyield two moles of GSH) or the synthesis and/or transport of GSHinto the cells. Together with cell viability, all the above oxidativestress-related biomarkers were markedly depressed when the

Table 3Inhibition of superoxide-induced lucigenin chemiluminescence by Ce compounds.

Added compound % of luminescence

None 100Nano-CeO2 (5 M) 68.8 ± 2.5Nano-CeO2 (10 M) 56.4 ± 1.3Nano-CeO2 (30 M) 27.3 ± 1.1Nano-CeO2 (50 M) 10.6 ± 1.0CeO2 (5 M) 91.3 ± 1.5Ce(SO4)2 (5 M) 54.9 ± 3.5CeCl3 (5 M) 25.6 ± 1.9SOD 15.7 ± 2.1SOD + nano-CeO2 (10 M) 10.4 ± 1.4*

SOD + nano-CeO2 (30 M) 22.4 ± 1.9*

Preincubated [SOD + nano-CeO2 (30 M)] 32.6 ± 2.9*,§

Preincubated SOD 16.9 ± 1.9

Data are calculated relative to the value for (lucigenin + O2��) measured 5 min after

addition of xanthine oxidase and are expressed as means ± SD (n = 6). The incuba-tion medium contained 5 lM lucigenin, 50 lM xanthine in EDTA (0.1 mM)-sup-plemented 0.2 M sodium phosphate buffer, pH 7.5. The reaction was initiated byaddition of xanthine oxidase (10 mU/ml). SOD concentration was 20 mU/mlthroughout and preincubation time before addition of xanthine oxidase was 6 h at37 �C. One-way ANOVA followed by Duncan t test.* P < 0.05 versus SOD.§ P < 0.05 versus preincubated SOD.

174 M. Culcasi et al. / Chemico-Biological Interactions 199 (2012) 161–176

nano-CeO2 dose was increased up to 6 � 10�1 g/l and, if PAO onlyprovided a partial protection, only the mixture of SOD and CAT,or mitoPBN could efficiently restore all these indices. In parallellipid peroxydation and protein carbonyls raised by 3.3 and 1.7times, respectively.

The present data in 3T3 cells provide substantial support for adual mechanistic evidence of oxidative stress associated withexposure to nano-CeO2, with an initial signaling at micromolardoses regulated by thiol-containing compounds. Similar observa-tions have been reported in in vivo studies with Ce compounds[15,63]. In our study the observed inhibitory properties of mitoPBNand the lack of a clear protective effect of PAO confirm that mito-chondria is a major source of ROS in nano-CeO2-induced cytotoxic-ity at millimolar doses. Although PBN and its derivatives are able toquench O2

��, this reaction is not fast enough to compete with dis-mutation (either catalyzed or spontaneous) and therefore the pro-tective effect of mitoPBN in the present study may arise frominhibition of secondary oxidation reactions. Nevertheless, this im-plies that Ce particles could reach the mitochondrial membrane, aproperty that was not clearly established in previous transmissionelectron microscopy studies of different cell types [5,14,21]. Wealso showed that, from the signaling phase, all systems involvedin H2O2 control (i.e., CAT, GPx and GSH) play a primordial role incell response to nano-CeO2 exposure. In this regard the cytotoxicityof high levels of H2O2 in cells, which provokes an irreversible oxi-dation of thiol-containing compounds [25,56,60], can explain thestrong decrease of CAT, GPx and GSH levels observed here. The cru-cial cooperation between GSH and the mitochondrial enzyme glu-tathione peroxidase in the control of H2O2 concentration has beendescribed in details [60].

It is generally believed that HO� plays an important role in H2O2-induced cell injury leading to the oxidation of membrane lipids andproteins and that the molecular mechanism requires the presenceof catalytic metal ions. From our discussion above and literaturedata [23] HOO� is the most likely intermediate formed whennano-CeO2 interferes with H2O2 within the cell (Eq. (1)). ThenHO�-induced toxicity, if any, will not proceed via Ce3+ catalysisbut probably by ‘conventional’ Fenton chemistry, e.g., with muchmore catalytically active iron or copper ions released upon mem-brane disruption. Furthermore a nonradical, H2O2-independentoxidase-like activity of Ce compounds, including nanoparticleshas been described for many biologically-occurring organic sub-stances [19,21,23,50].

3.6. Antagonist properties of cerium compounds towards superoxideand superoxide dismutase (Table 3)

In a final series of experiments we first investigated whetherthe protection of SOD levels of 3T3 cells seen in Table 2 could berelevant to O2

�� scavenging properties of nano-CeO2 in micromolarconcentrations. Therefore CL determinations were performed usinglucigenin (5 lM) as a chemiluminogenic probe for O2

�� generatedby the xanthine-xanthine oxidase reaction in the absence or pres-ence of Ce compounds (5–50 lM). Incubation of lucigenin alonewith Ce compounds (up to 50 lM) for up to 15 min yielded no lightemission whereas O2

��-induced lucigenin CL at 5 min was concen-tration-dependently inhibited by nano-CeO2 suggesting the parti-cles are good O2

�� scavengers. Lucigenin CL inhibition data ofindividual Ce compounds at 5 lM indicated the Ce3+ componentof nano-CeO2 (tested as CeCl3) to be a far better radical scavengercompared to Ce4+ (as Ce(SO4)2) or CeO2. In this assay addition ofSOD (20 mU/ml) inhibited 85% of lucigenin CL and uric acid absor-bance was not modified by Ce compounds showing none of theminhibit xanthine oxidase (data not shown).

Second we tested the possibility that depressed SOD levels seenin cells treated with a millimolar concentration of nano-CeO2 (Ta-

ble 2) may partially reflect a nonradical mechanism between theparticles and the enzyme. While a mixture of SOD (20 mU/ml)and nano-CeO2 (10 lM) acted synergistically on lucigenin CL ascompared to each component alone, this inhibitory effect droppedmarkedly when nano-CeO2 was present above 30 lM. This declinein lucigenin CL inhibition was even more pronounced when theparticles and the enzyme were coincubated for 6 h at 37 �C beforeO2�� generation was initiated, a result not due to an intrinsic loss of

SOD activity during incubation.By showing the prominent role of Ce3+ versus Ce4+ constituents

in the O2�� scavenging effect of nano-CeO2 in vitro, our data are

consistent with other studies on nanoceria having variable Ce3+/Ce4+ surface ratios [9,10]. Previous investigators [9] have proposedthat this active redox [Ce4+/Ce3+] couple in nano-CeO2 possess a po-tent SOD-like activity as described by Eqs. (4) and (5):

Ce3þ þ O��2 þ 2Hþ ! H2O2 þ Ce4þ ð4Þ

Ce4þ þ O��2 ! O2 þ Ce3þ ð5Þ

Besides its direct reaction with O2�� (Eq. (5)) it has also been

postulated that Ce4+ can be reduced into Ce3+ in the presence ofH2O2 according to a catalase-mimetic mechanism [11].

The results of Tables 2 and 3 support the reported significance ofO2�� scavenging in the protection by nano-CeO2 of cells under differ-

ent oxidative stress [4,6,8] and pH [22] conditions. Moreover therehave been several cell studies describing an upregulation of SODactivity consecutive to nano-CeO2 treatment at low doses [6,22].On the other hand the results of SOD preincubations with higher,but still micromolar concentrations of nano-CeO2 (Table 3) stronglysuggest, however, that their accumulation within the cell may resultin an overall loss of antioxidant defenses because the intrinsic O2

��

scavenging potential of the particles will not compensate the de-crease of SOD activity. This assumption of a detrimental effect ofnano-CeO2 overload in cells is in line with studies on bovine eryth-rocytes showing that CeCl3 can bind the active site of SOD, and while[Ce3+] <15 lM stimulated SOD activity, it was markedly decreasedwhen Ce3+ was added up to 25 lM in the system [64].

4. Conclusions

The present study imparts new information on the apparentcontradictory results of the literature where nanoceria is consid-

M. Culcasi et al. / Chemico-Biological Interactions 199 (2012) 161–176 175

ered either a protective antioxidant and free radical scavenger, orpro-oxidant and cytotoxic. In this regard our work on human andmurine fibroblasts identifies two key factors: (i) the possibility thatparticles can accumulate within a biological system followingchronic exposure (a case environmentally relevant to the wide-spread use of nanoceria), and (ii) the dual signaling/damaging roleof the H2O2/O2

�� couple. At micromolar doses (<300 lM), nano-CeO2 activates Nox2 (and possibly Nox4)-dependent ROS produc-tion at the membrane, interacts with signaling pathways leadingto protein synthesis activation and increases H2O2- and GSH-re-lated enzyme activity. However these pro-oxidant properties oflow nano-CeO2 do not lead to any significant cytotoxicity despitecell antioxidant status is altered. At millimolar doses of nano-CeO2 both the NADPH oxidase and the mitochondrial sources ofROS becomes highly activated leading to many known aspectsof cellular oxidative stress such as antioxidant depletion and lossof viability. In that case mitochondria is supposed to commandalternative signaling pathways leading to early apoptosis. In thiscascade of damaging events where formation of HO� is prone to oc-cur, Ce3+ ions are not likely to play a significant catalytic role, e.g.,compared to iron. This can be considered as a great advantage forbiomedical applications when non-toxic, micromolar doses of nan-oceria are used.

Conflict of Interest Statement

The authors declare that there are no conflicts of interest.

Acknowledgments

This work was funded by grants from the CNRS and the AgenceNationale de la Recherche (INTOX, ANR-08-CES-002 and ROS SIG-NAL, ANR-10-BLAN-003) at SMBSO-ICR UMR CNRS 7273. LB wassupported by a doctoral grant from Post-Grenelle funding via theAntiopes network (Impecnano project) and thanks J. Rose and M.Auffan at CEREGE (UMR 6635 CNRS, Aix-en-Provence, France) fortheir kind help. We also thank P. Bernasconi, G. Gosset, A.S. Saba-tier at DIPTA (Aix-en-Provence, France) and C. Questaigne at IN-SERM UMR 911 (Marseille, France) for their kind technicalassistance.

Appendix A. Details of syntheses and characterization

A.1. Synthesis of mitoPBN (3) (Scheme 1)

A.1.1. 4-(4-Iodobutoxy)benzaldehyde (1)To a cold solution of 4-hydroxybenzaldehyde (2 g, 16.39 mmol)

in acetonitrile (25 ml) was added solid K2CO3 (3.85 g, 27.86 mmol).After stirring for 10 min in an ice-water bath, 1,4-diiodobutane(25 g, 80.91 mmol) was added dropwise over 30 min, the reactionmixture was allowed to increase to room temperature over 1 h andwas then stirred at room temperature for 12 h. After removal of theprecipitate, the organic layer was concentrated under vacuum andthe crude product was purified by flash column chromatographyon silica gel (CH2Cl2) to give 4-(4-iodobutoxy)benzaldehyde (1)as a yellow oil (4.0 g, 13.10 mmol, 80%). 1H NMR (300 MHz, CDCl3):1.82–2.03 (m, 4 H, O–CH2-CH2; I–CH2-CH2), 3.21 (t, 2 H, J = 6.8 Hz,I–CH2), 4.01 (t, 2 H, J = 6.0 Hz, O–CH2), 6.93 (dd, 2 H, J = 2.00, 8.9 Hz,o-H, Ar–O–C), 7.76 (dd, 2 H, J = 2.00, 8.9 Hz, m-H, Ar–O–C), 9.82 (s,1 H, CHO); 13C NMR (75 MHz, CDCl3): 6.13 (I–CH2), 29.77 (I–CH2-CH2; O–CH2-CH2), 66.85 (O–CH2), 114.52 (o-C, Ar–O–C), 129.68(p-C, Ar–O–C), 131.76 (m-C, Ar–O–C), 163.65 (Ar–O–C), 190.53(CHO). Anal. HRMS–ESI: Calcd for C11H13IO2: 303.9960. Found303.9963.

A.1.2. [4-(4-Formylphenoxy)butyl]triphenylphosphonium iodide (2)A reaction mixture containing (1) (3 g, 9.86 mmol) and tri-

phenyl phosphine (4.4 g, 16.79 mmol) was refluxed in toluene(40 ml) for 12 h in the dark. After cooling down at room tempera-ture, the mixture evaporated in vacuo, the residue was dissolved ina minimal amount of dichloromethane and this solution was addeddropwise to excess ether (100 ml) under vigorous stirring. Theresultant yellow residue was allowed to settle for 1 h and the sol-vent was decanted. The resulting precipitate was again submittedthree times to the above precipitation process and the final residuewas dried in vacuo for 2 h to afford (2) as a pale yellow solid(5.13 g, 92%). 1H NMR (300 MHz, CDCl3): 1.87 (m, 2 H, P+–CH2-CH2), 2.25 (2H, quin, J = 6.4 Hz, O–CH2-CH2), 3.85 (m, 2 H, P+–CH2), 4.22 (t, 2 H, J = 5.7 Hz, OCH2), 6.95 (d, 2 H, J = 8.7 Hz, o-H,Ar–O-C), 7.6–7.9 (m, 17 H, ArH), 9.86 (s, 1 H, CHO); 31P NMR(121.5 MHz, CDCl3): 25.24 ppm; 13C NMR (75 MHz, CDCl3): 19.34(d, J = 4.4 Hz, P+–CH2-CH2), 22.70 (d, J = 50 Hz, P+–CH2), 29.22 (d,J = 16.00 Hz, P+–CH2-CH2-CH2), 67.12 (O–CH2), 115.01 (o-C, Ar–O–C), 118.60 (d, J = 95.7 Hz, �P+Ph3 ipso), 130.05 (p-C, Ar–O–C),130.62 (d, J = 12.6 Hz, �P+Ph3 meta), 132.13 (m-C, Ar–O–C),133.77 (d, J = 9.9 Hz, �P+Ph3 ortho), 135.23 (d, J = 2.7 Hz, �P+Ph3

para), 163.88 (Ar–O–C), 190.94 (CHO).

A.1.3. [4-[4-[[(1,1-Dimethylethyl)oxidoimino]methyl]phenoxy]butyl]-triphenylphosphonium bromide (mitoPBN; 3)

MitoPBN (3) was obtained (0.95 g; 46%) by reducing (2) (2.0 g,3.5 mmol) with Zn powder in dry ethanol, following the experi-mental procedure reported in [28].

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