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Original Contribution

Frataxin deficiency in neonatal rat ventricular myocytes targets mitochondria and lipid metabolism Èlia Obis a, Verónica Irazusta b, Daniel Sanchís a, Joaquim Ros a, Jordi Tamarit a,n a b

Departament de Ciències Mèdiques Bàsiques, IRB–Lleida, Universitat de Lleida, 25198 Lleida, Spain Instituto de Investigación para la Industria Química, INIQUI–CONICET, Salta, Argentina

art ic l e i nf o

a b s t r a c t

Article history: Received 6 November 2013 Received in revised form 4 April 2014 Accepted 4 April 2014

Friedreich ataxia (FRDA) is a hereditary disease caused by deficient frataxin expression. This mitochondrial protein has been related to iron homeostasis, energy metabolism, and oxidative stress. Patients with FRDA experience neurologic alterations and cardiomyopathy, which is the leading cause of death. The specific effects of frataxin depletion on cardiomyocytes are poorly understood because no appropriate cardiac cellular model is available to researchers. To address this research need, we present a model based on primary cultures of neonatal rat ventricular myocytes (NRVMs) and short-hairpin RNA interference. Using this approach, frataxin was reduced down to 5 to 30% of control protein levels after 7 days of transduction. At this stage the activity and amount of the iron–sulfur protein aconitase, in vitro activities of several OXPHOS components, levels of iron-regulated mRNAs, and the ATP/ADP ratio were comparable to controls. However, NRVMs exhibited markers of oxidative stress and a disorganized mitochondrial network with enlarged mitochondria. Lipids, the main energy source of heart cells, also underwent a clear metabolic change, indicated by the increased presence of lipid droplets and induction of medium-chain acyl-CoA dehydrogenase. These results indicate that mitochondria and lipid metabolism are primary targets of frataxin deficiency in NRVMs. Therefore, they contribute to the understanding of cardiac-specific mechanisms occurring in FRDA and give clues for the design of cardiac-specific treatment strategies for FRDA. & 2014 Elsevier Inc. All rights reserved.

Keywords: Friedreich ataxia Mitochondria Iron Lipid metabolism Oxidative stress Free radicals

Friedreich ataxia (FRDA)1 is an inherited human disease caused by decreased expression of the mitochondrial protein frataxin. The most common mutation consists of increased GAA triplet repeats in the first intron of the nuclear frataxin gene resulting in decreased protein expression, to less than 30% of normal values [1]. Patients with FRDA present symptoms including progressive gait and limb ataxia, hypertrophic cardiomyopathy, and diabetes mellitus [2]. Frataxin is a highly conserved protein related to processes such as iron–sulfur cluster biosynthesis [3], heme biosynthesis [4], iron storage [5] or sensing [5,6], electron transfer to flavoproteins [7], or holo-aconitase reconstitution [8]. Among these, the “iron–sulfur hypothesis” has received major support. Initial studies suggested frataxin as the iron donor in this process. However, recent work considers frataxin a modulator of the biogenesis rate [9,10], which would agree with the evidence that frataxin is not essential for iron–sulfur biogenesis [11]. Finally, it should be mentioned that frataxin deficiency is associated with increased sensitivity to oxidative stress in most models [12].

Abbreviations: FRDA, Friedreich ataxia; FFA, free fatty acid; tBHP, tert-butylhydroperoxide; IRP, iron regulatory protein; qPCR, quantitative real-time PCR n Corresponding author. E-mail address: [email protected] (J. Tamarit).

Most FRDA patients have evidence of cardiac dysfunction [13]. Indeed, cardiomyopathy is the leading cause of death in these patients [14]. Frataxin is highly expressed in tissues with a high metabolic rate [2]. Alterations of cardiac function may be caused by dysfunction of heart myocytes, which are rich in mitochondria and present high oxidative metabolism. Advances in understanding the cardiac phenotype of FRDA have been obtained from analysis of human hearts and conditional knockout mice with decreased frataxin levels in cardiac tissues. In hearts from FRDA patients, decreased activity of iron–sulfur-containing proteins and increased iron accumulation have been reported [15]. Recent observations indicate that such iron accumulation is highly localized and not restricted to the mitochondria [16]. Conditional cardiac and skeletal muscle mice (known as the MCK model) reproduced some characteristic features of the disease, including hypertrophic cardiomyopathy [17]. It also displayed some features that resembled those found in patients’ hearts, such as decreased activities of iron–sulfur enzymes, mitochondrial iron accumulation, and alterations in mitochondrial number, distribution, and morphology [18]. Despite the progress obtained with human hearts and mouse models, the specific effects of frataxin depletion on heart myocytes are poorly understood because no cardiac cellular model is available to analyze the consequences of frataxin depletion in

http://dx.doi.org/10.1016/j.freeradbiomed.2014.04.016 0891-5849/& 2014 Elsevier Inc. All rights reserved.

Please cite this article as: Obis, È; et al. Frataxin deficiency in neonatal rat ventricular myocytes targets mitochondria and lipid metabolism. Free Radic. Biol. Med. (2014), http://dx.doi.org/10.1016/j.freeradbiomed.2014.04.016i

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these cells. Thus, we have developed a cardiac model of frataxin deficiency to investigate cardiac-specific mechanisms occurring in FRDA. In our model, primary cultures of rat neonatal ventricular myocytes (NRVMs) are transduced with lentivirus containing short-hairpin RNA (shRNA) against frataxin mRNA. Lack of frataxin triggered oxidative stress, mitochondrial disarrangements, and an alteration in lipid metabolism that could be the origin of the altered cardiac status of most FRDA patients. This model also may be useful to analyze the effects of potential therapeutic agents on frataxin-depleted cardiac cells.

Materials and methods

vector SHC002) served as a control. A green fluorescent protein (GFP)-containing vector was used to optimize transduction efficiency. Lentiviral particles were produced by cotransfection of HEK293T cells using the plasmids pMD2.G and psPAX2 (Addgene) and polyethylene as described previously [21]. After 3 days, lentiviruses were collected from the culture medium, centrifuged for 5 min at 1000 rpm, filtered through a 0.45-μm filter (Millipore), concentrated by centrifugation through 100,000 MWCO Vivaspin 20 columns (Sartorius), and stored at  80 1C. Lentiviruses were titered using the Quicktiter Lentivirus ELISA kit (Cell Biolabs), which detects the amount of p24 protein. For NRVM transduction, 5.5 ng of p24 per 1000 cells was added to the medium 4 h after plating, and 20 h later, the culture medium was replaced with fresh medium. This protocol provided a transduction efficiency greater than 70%, according to the GFP control vector.

Isolation and culture of NRVMs Cell viability assays The investigation with experimental animals conformed to the national guidelines for the regulation of the use of experimental laboratory animals from the Generalitat de Catalunya and the Government of Spain (Article 33.a 214/1997) and was evaluated and approved by the Experimental Animal Ethic Committee of the University of Lleida (CEEA). Around 150 neonatal rats were used. P3–P4 Sprague–Dawley rat neonates were sacrificed, following the above guide approved by the CEEA, by decapitation and NRVMs were obtained from the hearts as described previously [19], with some modifications. Briefly, ventricles from newborn rats were minced and cells were isolated by three subsequent digestion steps with 150 U/ml type 2 collagenase (Worthington, USA) and stirring at 37 1C. Noncardiomyocyte cells were separated from the cardiomyocytes by differential preplating. Cardiomyocytes were seeded on 0.2% gelatin-coated culture dishes (Falcon, Becton & Dickinson, USA) at 7.5
104 cells/cm2, giving a confluent monolayer of spontaneously contracting cells after 24 h. To inhibit the proliferation of cardiac fibroblasts, cells were treated with 10 μg/ml mitomycin C (Sigma) for 4 h after the seeding time. The culture medium used was Dulbecco's modified Eagle's medium:M199 3:1, 5 mM glucose, with 8% horse serum and 4% fetal bovine serum, GlutaMAX, and Hepes (all from Gibco). Cardiomyocyte purity was checked by immunofluorescence with a specific cardiac α-actinin antibody (Sigma) and was found to be higher than 90% after 24 h in vitro. When required, fatty acid–albumin solutions were used for free fatty acid (FFA) supplementation of the culture medium. A commercial oleic and linoleic–albumin solution was used to provide these unsaturated fatty acids. Saturated fatty acid– albumin solutions were prepared as described [20], with some modifications. FFA–albumin was dissolved in water at 300 μM concentration and the pH was brought to 9 with sodium hydroxide. An ethanolic solution of the desired fatty acid was prepared, 90 mM for palmitic acid and 45 mM for stearic acid. One volume of this ethanolic solution was added to 50 volumes of the albumin solution with stirring. The required amount of this solution was added to the culture medium to give a final concentration of 60 μM palmitic acid and 30 μM stearic acid. For deferoxamine treatment, the medium was changed and replaced with either control medium or 100 μM deferoxamine-containing medium for the next 16 h. Quantification of FFA in medium was performed with a Free Fatty Acids test (Roche). Production of lentiviral particles The shRNA lentiviral plasmids (pLKO.1-puro) for human/mouse/ rat frataxin were purchased from Sigma. The RefSeq used was NM_008044, which corresponds to mouse frataxin. The clones TRCN0000197534 and TRCN0000178380, named sh34 and sh80 in this study, were designed for mouse frataxin interference in a region 100% identical to rat frataxin. A nontargeted scrambled sequence (the

For the MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) assay, cells were grown in 96-well plates and 7 days after lentivirus transduction, the cells were rinsed and labeled with 0.2 mM MTT for 4 h at 37 1C and 5% CO2 in Hanks’ balanced salt solution (HBSS) with calcium, magnesium, and 5 mM glucose. The precipitates were dissolved in dimethyl sulfoxide and absorbance was read at 540 nm in a BioTek Power Wave XS. Treatment with tert-butylhydroperoxide (tBHP) was conducted before the MTT assay in HBSS buffer without glucose. A Live/Dead viability kit (Invitrogen) was used according to the manufacturer's instructions. Western blot and immunofluorescence Cells were first rinsed in ice-cold phosphate-buffered saline (PBS), pH 7.2, and lysed in 2% SDS, 125 mM Tris, pH 6.8, containing a protease inhibitor cocktail (Roche). Protein concentration was determined by the Qubit assay (Invitrogen). After SDS–polyacrylamide gel electrophoresis, proteins were transferred to Immobilon-P, Immobilon-FL (Millipore), or nitrocellulose membranes. The membranes were probed with the following primary antibodies: frataxin (sc-25820), medium-chain acyl-CoA dehydrogenase (sc365109), very long chain acyl-CoA dehydrogenase (sc-271225), and electron-transferring flavoprotein dehydrogenase (sc-242642) (all from Santa Cruz Biotechnology); electron transfer flavoprotein subunit β (ab104944, Abcam); trifunctional enzyme subunit α (ab54477, Abcam); peroxisome proliferator-activated receptor γ coactivator 1-α (101707, Cayman); anti-Rt/Ms Total OxPhos Complex Kit (458099, Invitrogen); dinitrophenyl (V0401, Dako); mitochondrial aconitase (HPA001097, Sigma); manganese superoxide dismutase (SOD-110, Stressgene); and copper–zinc superoxide dismutase (ab1237, Chemicon). Detection was performed using either peroxidase-conjugated or fluorescent (Cy3- or Cy5conjugated) secondary antibodies. Image acquisition was performed in a ChemiDoc XRS (chemiluminescence) or Versadoc MP (fluorescence), both from Bio-Rad. When required, data were analyzed by Quantity One or Image Lab software (Bio-Rad). For immunofluorescence, cells were fixed in 4% paraformaldehyde for 15 min and permeabilized and blocked with 0.2% Triton X-100, 2% bovine serum albumin, and 2% horse serum containing PBS for 1 h before antibody detection. Secondary antibodies were Alexa Fluor 488 (A11008, Invitrogen) and rhodamine red (715-295-150, Jackson Immunoresearch). Cells were coverslipped with an anti-fading mounting medium (0.1 M Tris–HCl buffer, pH 8.5, 20% glycerol, 10% Mowiol, and 0.1% DABCO). Quantitative real-time PCR For qPCR analysis, total RNA was extracted using the RNeasy kit (Qiagen) according to the manufacturer's instructions. One

Please cite this article as: Obis, È; et al. Frataxin deficiency in neonatal rat ventricular myocytes targets mitochondria and lipid metabolism. Free Radic. Biol. Med. (2014), http://dx.doi.org/10.1016/j.freeradbiomed.2014.04.016i

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microgram of total RNA from each sample was converted into cDNA and 50 ng of cDNA used for each individual qPCR. The assays were performed in an iCycler (Bio-Rad) using commercial TaqMan Universal PCR master mix and gene expression assays (Applied Biosystems). Assays used were Fxn (Rn01501404_g1), Actb (Rn00667869_m1), Tfrc (Rn01474701_m1), Pparg (Rn00440945_m1), and Cpt1b (Rn00682395_m1). Actb was used as an internal control. Quantification was completed using iCycler IQ Real-Time Detection System software (version 2.3, Bio-Rad). Relative expression ratios were calculated on the basis of ΔCp values with efficiency correction based on multiple samples [22]. Subcellular labeling and image analysis For mitochondria labeling, cells were loaded with 100 nM MitoTracker Red CMXRos for 3 min and washed with serum-free medium. An Olympus IX71 microscope equipped with epifluorescence optics and a DP70 CCD camera were used to obtain images with a 40
lens. To provide an estimation of the percentage of cells that presented a disarranged mitochondrial appearance, we used the following strategy. Images were coded and blind-counted by two individuals who were asked to classify the observed cells in two groups: those showing the typical organized appearance of cardiac myocytes and those showing a disarranged appearance. In cardiomyocytes, mitochondria are arranged in a very regular manner, distributed like lines of small spots of similar size. Cells were classified as disarranged when their mitochondria did not show such a regular appearance, in terms of both size and spatial distribution. For each condition, at least 75 cells in eight different microscopic fields were evaluated. Number of balloon-like mitochondria was assessed using the “Analyze Particles” tool from ImageJ software. Images were background-subtracted and the following criteria were used to detect particles: size higher than 2 μm2 and circularity 0.5–1. For labeling of lipid droplets, cells were washed with PBS and loaded with 5 μM BODIPY 493/503 (Invitrogen) for 10 min at 37 1C. Nuclear staining with 0.05 μg/ml Hoechst 33258 was done as necessary. An Olympus IX71 microscope equipped with epifluorescence optics and a DP70 CCD camera was used to obtain images with a 40
lens. For mitochondrial membrane potential assessment, tetramethylrhodamine methyl ester (TMRM; Life Technologies) was used. NRVM cultures were labeled with 30 nM TMRM for 1 h as described [23] and treated with 10 μM CCCP to completely depolarize mitochondria and to establish the background levels of fluorescence. Cells were visualized and analyzed using a confocal microscope (FV1000, Olympus) using the oil-immersion 60
magnification objective. Analysis of images was made with Fluoview FV1000 software. ATP content Phosphorylated adenosine nucleotides were quantified by HPLC as described [24], with some modifications. NRVM cultures were rinsed twice with cold PBS and 200 μl of 660 mM HClO4 was added to the culture well. Cells were collected and kept on ice for 10 min and then centrifuged at 14,000 rpm at 4 1C for 10 min. The supernatant was neutralized with 20 μl 2.8 M K3PO4, kept on ice for 10 min and at  80 1C for at least 2 h to promote precipitation of the perchlorate, and then centrifuged again. Supernatants were stored at  80 1C until HPLC assay. Chromatography was performed at a flow rate of 0.2 ml/min on a Waters 625 LC system equipped with a 996 photodiode array detector and a Waters DeltaPak HPI C18 column (2
150 mm, 3-μm particle size). Buffer A contained 25 mM NaH2PO4, 100 mg/liter tetrabutylammonium, pH 5. Organic buffer B was composed of 10% (v/v) acetonitrile in 200 mM NaH2PO4, 100 mg/liter tetrabutylammonium, pH 4.0. A linear gradient from 100% buffer A to 100% buffer B over

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25 min was used. Ten microliters of prepared sample or standard mixture was injected and UV monitored at 260 nm. Peaks were identified by their retention times and quantified based on peak areas compared to standards. Glucose and lactate quantitation For the analysis of glucose consumption and lactate production, NRVMs (375,000 cells) were seeded in 4-cm2 plates and cultured under standard conditions in medium supplemented or not with fatty acids. The medium was replaced daily. At days 4 and 7, glucose and lactate concentrations were quantified in 8-h conditioned medium using commercial kits from BioSystems (for glucose; Barcelona, Spain) and Spinreact (for lactate; Sant Esteve de Bas, Spain). For the calculation of glucose consumption, the remaining glucose concentration was subtracted from the concentration in fresh medium (5 mM). Iron analysis For total iron determination, cells were washed once in PBS with 1 mM EDTA and collected in a Mes buffer, pH 4.5, with 1% SDS and digested in nitric acid (3%). Iron content was determined using bathophenanthroline sulfonate as chelator [25]. Enzyme activities Aconitase activity was measured using a protocol described previously [26], with some modifications. Cells were lysed with 50 mM Tris buffer, pH 7.4, containing 2.5 mM citrate and 0.5% Triton X-100 and protease inhibitors (Roche). Samples were incubated on ice for 5 min and centrifuged at 13,000 g at 4 1C. Activity was measured in 50 mM Tris, pH 7.4, containing 5 mM sodium citrate, 0.2 mM NADP, 0.6 mM MnCl2, and 0.25 U of isocitric dehydrogenase. NADPH formation was measured at 340 nm for 3 min. Citrate synthase was measured in a coupled assay to reduce 5,50 -dithiobis(2-nitrobenzoic acid) [27]. Cells were lysed with the same protocol used for aconitase activity assay and 4 μl of cell extract was used for this activity. Electron transfer activities of respiratory chain complexes were measured as reported [28] from cells lysed using a Teflon pestle in PBS containing 0.05% dodecyl maltoside, 0.005% saponin, and protease inhibitors. Cytochrome c reduction was monitored at 550 nm at 25 1C. Statistical analysis All experiments were performed in at least three completely independent cardiomyocyte preparations, except activities from the mitochondrial electron transport chain (ETC) complexes, which were analyzed in four independent preparations. Values are expressed as means 7 SEM. The data obtained from the independent experiments were used for statistical analysis. Data obtained were compared with control conditions using the Student t test. The p values lower than 0.05 were considered significant.

Results Frataxin silencing in rat NRVMs NRVM cultures were transduced with lentiviral particles containing vectors encoding two different short-hairpin RNA sequences against frataxin (named sh34 and sh80). A scrambled (SCR; random interference) sequence was used as a control. This sequence does not present homology to any known gene of the rat genome. Four and seven days after infection, interference was

Please cite this article as: Obis, È; et al. Frataxin deficiency in neonatal rat ventricular myocytes targets mitochondria and lipid metabolism. Free Radic. Biol. Med. (2014), http://dx.doi.org/10.1016/j.freeradbiomed.2014.04.016i

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verified by qPCR and Western blot. A marked decrease in frataxin mRNA content was observed 4 days postinfection in cells transduced with sh34 and sh80 shRNA (Fig. 1A). In all cases, mRNA content was below 10% of that found in cells transduced with SCR shRNA. Similar results were observed at 7 days postinfection. Western blot (Fig. 1B and C) indicated a marked decrease in protein content, with residual frataxin protein levels in cells transduced with sh34 and sh80 below 30% of the SCR level. This decrease was less marked at 4 days, probably because of slow turnover of the protein. Thus, we opted to perform all subsequent analyses in 7-day-transduced cultures because the remaining levels of frataxin found in those cells were similar to those in FRDA patients. Cell viability was evaluated using a Live/Dead test based on propidium iodide and calcein staining. Low frataxin levels did not exert a marked effect on cell viability, which was only marginally and nonsignificantly affected in 7-day-transduced cultures (data not shown).

Mitochondrial morphology, distribution, and membrane potential Mitochondrial morphology was evaluated in our model at 7 days posttransduction using MitoTracker Red CMXRos. In SCR cells, mitochondria were evenly arranged, showing a typical fibrous-like appearance (Fig. 2A). Marked alterations in mitochondrial size and distribution could be observed in the sh34- and sh80-silenced cells. In these cells mitochondria were disarranged, and enlarged balloon-like mitochondria were concentrated around the nucleus. These alterations are reminiscent of those in cardiomyocytes from the conditional mouse model and in human heart [13]. The percentage of cells presenting a disarranged mitochondrial appearance and the number of balloon-like mitochondria were calculated as described under Materials and methods. As shown in Fig. 2B and C, the sh34 and sh80 cultures presented a higher presence of cells with a disarranged mitochondrial appearance and increased presence of balloon-like mitochondria than SCR cultures. To analyze mitochondrial function in more detail, NRVMs were stained with TMRM and analyzed by confocal microscopy. TMRM is a cationic fluorescent dye that accumulates inside mitochondria according to the membrane potential. The intensity of the TMRM fluorescence was determined in individual cells before and after CCCP addition. CCCP was used to dissipate transmembrane potential and provide a measurement of background fluorescence. As shown in Fig. 2D, no significant differences in the intensity of TMRM fluorescence could be observed between SCR and silenced NRVMs. TMRM staining confirmed the presence of balloon-like mitochondria in frataxin-deficient cells (Fig. 2E). Mitochondrial respiratory complex activity The results in the previous section indicate that loss of frataxin promotes marked disarrangements in mitochondria distribution. Therefore, we decided to analyze the content and activity of several members of the mitochondrial electron transport chain to determine the effects of frataxin loss on this process. We performed Western blot against five representative subunits from components of the mitochondrial respiratory complexes: Nd6 (complex I), Sdhb (complex II), Uqcrc (complex III), Mtco1 (complex IV), and Atp5a1 (complex V). Fig. 3A shows that no significant changes were found in any of these polypeptides in frataxindeficient cells. We also used coupled assays of complex I þ III and II þ III activities (Fig. 3B) to analyze respiratory complex activity in total cell extracts. None of these assays revealed significant differences between SCR and sh34 or sh80 cells. To further analyze the energy status of frataxin-silenced cells, we measured ATP/ADP and AMP/ATP ratios in total cell lysates by an HPLC-based method (see Materials and methods). Some models of FRDA have decreased ATP levels [29]. However, in our model there were no significant differences between silenced or scrambled NRVMs in the ATP/ADP or AMP/ATP ratios (Fig. 3C). This indicates the capacity of frataxin-deprived NRVMs to sustain their production of ATP, at least in the early stages of depletion. These results are consistent with the presence of high-potential mitochondria (as indicated by TMRM probe) and normal respiratory chain complex activity.

Fig. 1. Frataxin depletion in rat NRVMs. NRVMs were transduced with lentivirus vectors containing sh34, sh80, or scrambled shRNA. (A) Relative frataxin mRNA expression measured by qPCR, 4 and 7 days after transduction. Actin expression was used as an internal control to normalize expression levels. (B) Protein extracts from 4- and 7-day-transduced cultures were probed with anti-frataxin antibody by Western blot analysis. In each gel lane, 30 μg of protein from whole-cell extracts was loaded. Protein load was verified by post-Western Ponceau staining. (C) Histogram represents the quantification of the relative expression of frataxin measured by Western blot. Both protein and mRNA data are represented as means 7 SEM from three independent cultures. In each experiment, protein content or mRNA level in scrambled control cells was used as a 100% reference value.

Aconitase activity Frataxin has been associated with iron–sulfur protein biogenesis [10]. Therefore, we analyzed the activity of the iron–sulfur-containing protein aconitase as an indicator of enzyme status. Two aconitases are present in mammalian cells. Aconitase 1, located in the cytoplasm, also functions as an iron sensor known as iron regulatory protein 1 (IRP1). Aconitase 2 (encoded by Aco2) is located

Please cite this article as: Obis, È; et al. Frataxin deficiency in neonatal rat ventricular myocytes targets mitochondria and lipid metabolism. Free Radic. Biol. Med. (2014), http://dx.doi.org/10.1016/j.freeradbiomed.2014.04.016i

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Fig. 2. Mitochondrial labeling and mitochondrial membrane potential assessment. (A) Representative microscopic images of cultured NRVMs stained with MitoTracker Red CMXRos. Images were acquired 7 days after transduction with lentivirus vectors containing sh34, sh80, or scrambled shRNA. A disturbed mitochondrial pattern and increased presence of enlarged mitochondria can be observed in frataxin-depleted cells. Insets are magnified images to show enlarged mitochondria in more detail. (B) Percentage of NRVMs presenting an altered mitochondrial network. For each condition, at least 75 cells in 8 different microscopic fields were evaluated. Data are presented as means 7 SEM from three independent experiments. (C) Average number of balloon-like mitochondria. For each condition, at least 50 cells in 6 different microscopic fields were evaluated using ImageJ software. (D) Histograms show the difference in fluorescence intensity of TMRM-stained cells before and after CCCP addition. Data represent means 7 SEM. A minimum of 40 cells from 10 different microscopic fields were used for each condition. (E) Representative microscopic images of cultured NRVMs stained with TMRM, before and after CCCP addition. TMRM staining indicates that no major changes in mitochondrial membrane potential occurred after frataxin depletion.

in mitochondria. The relative contributions of both enzymes to total cellular aconitase activity depend on the type of cell and growth conditions [30]. We measured total cellular aconitase in whole-cell lysates of SCR-, sh34-, and sh80-transduced NRVMs by a spectrophotometric method. This approach provided accurate and rapid quantification of labile enzymatic activity. Of note, in heart cells the

mitochondrial isoenzyme accounts for nearly 98% of total cellular aconitase activity [31]. Citrate synthase activity was monitored as a control, as this mitochondrial enzyme does not contain an iron–sulfur cofactor. Finally, the content of Aco2 was measured by Western blot and by immunofluorescence. Fig. 4A and B show that no significant

Please cite this article as: Obis, È; et al. Frataxin deficiency in neonatal rat ventricular myocytes targets mitochondria and lipid metabolism. Free Radic. Biol. Med. (2014), http://dx.doi.org/10.1016/j.freeradbiomed.2014.04.016i

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Fig. 3. Analysis of components of the OXPHOS system. (A) Protein extracts from 7-day-transduced cultures were probed with the indicated antibodies by Western blot analysis. In each gel lane, 15 μg of protein from whole-cell extracts was loaded. Protein load was verified by post-Western Coomassie Brilliant Blue (CBB) staining. Histogram represents the quantification of the relative expression of the indicated proteins measured by Western blot. Protein content in scrambled control cells was used as a 100% reference value. (B) Activity from the mitochondrial ETC complexes measured in whole-cell extracts as reduction of cytochrome c using either NADH (complexes I þ II) or succinate (complexes II þ III) as substrate. Units represent micromoles of cytochrome c reduced per minute. (C) AMP/ATP and ATP/ADP ratios were measured by HPLC as described under Materials and methods. From (A) to (C), data are presented as means 7 SEM from three independent cultures.

changes were observed in the ratio of aconitase to citrate synthase activities, nor in the content of Aco2 protein in frataxin-silenced cells compared to SCR cells. Immunofluorescence using Aco2 antibodies (Fig. 4D) confirmed the Western blot results and the altered mitochondrial pattern observed with MitoTracker and TMRM dyes. Markers of iron status In several models, loss of frataxin activates the cellular response to low iron conditions. In mammalian cells this may occur through the activation of IRP1 or IRP2 [32]. Activation of these proteins leads to upregulation of iron-uptake systems and

downregulation of some iron-containing proteins. Mitochondrial aconitase (Aco2) is an iron-responsive protein regulated by IRPs with decreased expression under iron-limiting conditions. The presence of normal Aco2 content in frataxin-silenced NRVMs (Fig. 4B) suggested that IRPs had not been activated in those cells. To confirm this point in our model, other markers of iron status were evaluated in SCR, sh34, and sh80 cells. Transcript levels from a protein involved in cellular iron uptake, transferrin receptor 1 (TFR1), were evaluated by qPCR. Total cellular iron content was evaluated colorimetrically after wet acid digestion of cells. No significant changes were observed in any of these parameters between silenced and SCR cells (Fig. 4E and F), suggesting that iron metabolism was not disturbed in silenced NRVMs under the

Please cite this article as: Obis, È; et al. Frataxin deficiency in neonatal rat ventricular myocytes targets mitochondria and lipid metabolism. Free Radic. Biol. Med. (2014), http://dx.doi.org/10.1016/j.freeradbiomed.2014.04.016i

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Fig. 4. Aconitase activity and markers of iron status. (A) Aconitase (ACO2) and citrate synthase (CS) activities were measured in protein extracts from 4- and 7-daytransduced cultures with the indicated shRNAs. Citrate synthase activity was used as a marker of mitochondrial content. Data are presented as means 7 SEM from three independent cultures. (B) Mitochondrial aconitase (Aco2) content analyzed by Western blot in whole-cell protein extracts from 4- and 7-day-transduced cultures. (C) Aconitase content measured by Western blot in control cells treated with deferoxamine (DFO) for 16 h. In both (B) and (C), 15 μg of protein was loaded in each gel lane. Protein load was verified by post-Western CBB staining. (D) Immunofluorescence assay was performed in 7-day-transduced NRVMs with mitochondrial aconitase antibodies. Hoechst staining was used to visualize nuclei. (E) Total iron content per 35-cm2 culture dish well was measured by a photometric method in 7-day-transduced cultures as described under Materials and methods. (F) Relative mRNA expression of transferrin receptor 1 (TFR1) in sh34- and sh80-inhibited cultures, compared to scrambled control cultures. The relative change in TFR1 mRNA expression triggered by 16 h DFO treatment in control cells is also shown. Data are presented as means 7 SEM from three independent cultures. Actin expression was used as an internal control to normalize expression levels. In each experiment, mRNA levels under control conditions were used as the reference value. nnp o 0.01; highly significant difference compared to control conditions.

conditions analyzed. To confirm that the IRP pathway could be activated in NRVMs by iron deprivation, control NRVMs were treated with the iron chelator deferoxamine. Aco2 content and

TFR1 expression were analyzed. We observed a decrease in Aco2 content and increased expression of the TFR1 mRNA, confirming that NRVMs can activate the IRP pathway (Fig. 4C and F).

Please cite this article as: Obis, È; et al. Frataxin deficiency in neonatal rat ventricular myocytes targets mitochondria and lipid metabolism. Free Radic. Biol. Med. (2014), http://dx.doi.org/10.1016/j.freeradbiomed.2014.04.016i

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Fig. 5. Markers of oxidative stress and sensitivity to tBHP. (A) Presence of protein carbonyl groups was analyzed in 7-day-transduced cultures by Western blot using α-DNP antibodies. 10 μg of protein was loaded in each gel lane and protein load was verified by post-Western CBB staining. Histograms represent relative carbonyl content in each condition, which was calculated from the chemiluminescent signal of each lane of the Western blot analyzed with Quantity One software (Bio-Rad). Carbonyl levels in scrambled control cultures were used as a reference value. (B) 7-day-transduced cultures were subjected to tBHP treatment at the indicated concentrations, and cell viability was assessed by an MTT assay. Values represent percentage of fluorescence relative to untreated scrambled control cultures. (C) Protein extracts from 7-day-transduced cultures were probed with SOD1 and SOD2 antibodies by Western blot analysis. In each gel lane, 15 μg of protein from whole-cell extracts was loaded. Protein load was verified by post-Western CBB staining. Histograms represent the quantification of the relative expression of the indicated proteins measured by Western blot. Protein content in scrambled control cells was used as a 100% reference value. From (A) to (C), data are presented as means 7 SEM from three independent cultures. np o 0.05 and nn p o 0.01; significant or highly significant difference, respectively, compared to control condition.

Oxidative stress: markers and sensitivity Oxidative stress is a common trait in several models of FRDA [12]. Thus, we evaluated the presence of carbonylated proteins in our model, as well as sensitivity to oxidative stress in frataxindeficient NRVMs. Carbonyls are a common product of the reaction of free radicals with proteins. This reaction can be evaluated by Western blot using antibodies against 2,4-dinitrophenylhydrazone (DNP), a molecule used to label carbonyl groups on proteins [33]. Increased protein carbonyl content has been observed in

frataxin-deficient yeast cells [34], although this was not the case in the conditional frataxin MCK mouse model [35]. In our model, carbonylated proteins after 7 days of transduction were higher in frataxin-deficient cells (Fig. 5A). We also evaluated sensitivity to an oxidant agent such as tBHP. Seven-day-transduced cultures were exposed to 5 and 10 μM tBHP; cellular viability was assessed with MTT assay. A dose-dependent increased sensitivity was observed in frataxin-depleted cells compared to scrambled control cells (Fig. 5B). Finally, we analyzed the content of two antioxidant enzymes, SOD1 and SOD2, which have been found either induced

Please cite this article as: Obis, È; et al. Frataxin deficiency in neonatal rat ventricular myocytes targets mitochondria and lipid metabolism. Free Radic. Biol. Med. (2014), http://dx.doi.org/10.1016/j.freeradbiomed.2014.04.016i

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[36] or repressed [37] in different models of frataxin deficiency. In our model, both enzymes showed a slight but significant increase in protein content (Fig. 5C). Together, these results indicate that frataxin-deficient NRVMs show a mild oxidative stress phenotype, with increased sensitivity toward oxidative stress, slight induction of SOD enzymes, and an increased presence of protein carbonyls. Lipid metabolism—presence of lipid droplets Silenced NRVMs, in contrast to control cells, exhibited small refringent bodies after 7 days in culture. These were positively stained with the neutral dye BODIPY 493/503, confirming that they were lipid droplets (Fig. 6A), and quantified (Fig. 6B). This finding attracted our attention because lipid droplets have been observed in frataxin-deficient Drosophila melanogaster [38] and in hearts from a neuron/cardiac frataxin conditional knockout mouse

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model (the NSE model) [17]. In addition, deregulated lipid metabolism was observed in several pathologies related to mitochondrial dysfunction. As heart relies mostly on fatty acid mitochondrial metabolism to fuel oxidative phosphorylation [39], we tested whether the concentration of this nutrient in the culture medium used was close to physiological concentrations. In fresh culture medium, the FFA concentration was 72 μM, contrasting with the  200 μM concentration typical of rat plasma [40]. To ensure FFA availability, and in an attempt to approach physiological conditions in terms of total concentration and composition of FFA, culture medium was supplemented with a mixture of bovine serum albumin-conjugated FFA. The supplemented medium contained 30 μM each linoleic, oleic, and stearic acid and 60 μM palmitic acid plus FFA provided by the standard medium. It is important to note that control NRVMs were able to consume these lipids in normal as well as supplemented culture medium

Fig. 6. Presence of lipid droplets. (A) Representative microscopic images of 7-day-transduced cultures grown in standard medium loaded with 5 μM BODIPY 493/503 for labeling lipid droplets. (B) Histograms represent both the percentage of cells that present lipid droplets and the average number of lipid droplets per cell in 7-day-transduced cultures grown in nonsupplemented medium. For each condition, at least 100 cells in five different microscopic fields were evaluated. Data are presented as means 7 SEM from three independent experiments. nnp o 0.01; highly significant difference compared to control condition. (C) NRVMs were cultured in standard or FFA-supplemented (FAsup) medium and the concentration of remaining FFA in the medium was measured at time 0 and after 24 h. (D) Representative microscopic images of 7-day-transduced cultures grown in fatty-acid-supplemented medium loaded with 5 μM BODIPY 493/503 for labeling lipid droplets.

Please cite this article as: Obis, È; et al. Frataxin deficiency in neonatal rat ventricular myocytes targets mitochondria and lipid metabolism. Free Radic. Biol. Med. (2014), http://dx.doi.org/10.1016/j.freeradbiomed.2014.04.016i

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because after 24 h the FFA concentration was reduced almost 90% (Fig. 6C). This result is consistent with early observations by other authors, which showed that neonatal rat cardiomyocytes had the capacity to oxidize fatty acids when present in the culture medium. Indeed, fatty acid supplementation resulted in increased expression of several enzymes required for fatty acid oxidation and increased cardiomyocyte fatty acid oxidation capacity [41]. In our model, fatty acid supplementation also promoted an increase in the expression of very long chain acyl-CoA dehydrogenase (VLCAD; shown in Supplementary Fig. 1A). Interestingly, VLCAD also increased with culture time. We also analyzed what effects fatty acid supplementation and culture time had on glucose consumption and lactate production, and we observed that cultures supplemented with fatty acids produced less lactate and showed a tendency toward decreased glucose consumption (Supplementary Fig. 1B). These results suggest that NRVMs decrease their glycolytic flux after several days in culture, a tendency that is more marked by the presence of fatty acids in culture medium. We then analyzed the effects of FA-supplemented medium on SCR and silenced NRVMs. Cells were cultured from day 1 with FFA supplement, and these media were replaced daily. After 6 days (7 days posttransduction), the cells were stained with BODIPY 493/503 dye. Large lipid droplets accumulated in frataxindeficient cells (Fig. 6D), whereas SCR cells scarcely displayed lipid droplet accumulation. Lipid metabolism—peroxisome proliferator-activated receptor (PPAR) pathway To further investigate the origin of lipid droplets in frataxindeficient cells we focused on the PPAR pathway. This pathway regulates many genes involved in lipid metabolism. In addition, one of its components (the PPARγ coactivator 1-α or PGC1α) was found altered in some models of FRDA: in skeletal muscle from a mouse model of Friedreich ataxia and in fibroblasts from FRDA patients it was decreased, whereas in HL-1 cardiomyocytes deficient in frataxin it was found slightly increased [42]. PGC1α is a transcriptional coactivator that positively regulates the expression of diverse mitochondrial, peroxisomal, and ROS-detoxifying genes [43]. PPARs are nuclear receptors that play key roles in the regulation of lipid metabolism [44]. They bind and are activated by fatty acids (FAs) and serve as transcriptional sensors of FAs. Three different PPARs have been described. PPARα is a major inducer of FA oxidation and is highly expressed in tissues with substantial mitochondrial and peroxisomal β-oxidation, such as brown adipose tissue, liver, kidney, and heart. PPARβ (also denominated PPARδ) also contributes to myocardial FA oxidation and is involved in maintaining energy balance and normal cardiac function. Both PPARα and PPARβ are expressed at relatively high levels in the heart where they display a large overlap in target genes. Finally, PPARγ is a major activator of adipocyte differentiation and promotes FFA uptake and lipogenesis in many different cell types. Although PPARγ is expressed at low levels in the heart, induction of PPARγ levels has been observed in hypertrophic cardiomyopathy in mice and humans [45]. Interestingly, cardiac PPARγ-overexpressing mice display increased FFA uptake [46]. Based on this background, we decided to investigate the potential role of the PPAR pathway in promoting lipid accumulation in frataxin-deficient NRVMs. We first analyzed the levels of PGC1α by Western blot. As shown in Fig. 7, we did not observe significant differences in PGC1α content between SCR and silenced NRVMs. We next focused on six proteins required for fatty acid catabolism that are PPARα targets. These were carnitine palmitoyltransferase 1 (CPT1), medium-chain acyl-CoA dehydrogenase (MCAD), VLCAD, the α subunit from the trifunctional enzyme (HADHA), the β subunit from electron transfer flavoprotein (ETFB), and electron

transfer flavoprotein–ubiquinone oxidoreductase (ETFDH). Levels of these proteins were measured by Western blot, except CPT1 expression, which was measured by qPCR. MCAD showed a marked increase after frataxin depletion. CPT1 was increased in sh80, but not in sh34 NRVMs. No significant changes were observed in the levels of VLCAD, HADHA, ETFB, or ETFDH. These events clearly indicate that the PPARα-dependent targets are not downregulated. Indeed a certain activation of this pathway may occur after frataxin depletion, as MCAD and CPT1 were increased. Finally, we measured PPARγ expression by qPCR to analyze if this transcription factor was induced and promoting FFA uptake and lipogenesis in our frataxin-silenced NRVMs. However, we found the opposite: PPARγ levels were lower in silenced NRVMs than in SCR cells (Fig. 7). In summary, these results indicate that lipid accumulation in frataxin-deficient cardiomyocytes is not due to increased FFA uptake promoted by PPARγ or to downregulation of catabolic enzymes (targets of PPARα or PGC1α). Instead, compromised FA catabolism due to mitochondrial defects caused by frataxin depletion may contribute to lipid droplet formation.

Discussion Cardiopathy is the leading cause of death in patients with FRDA, and to date there has been no cellular model to investigate the consequences of frataxin depletion in cardiac cells. We present a new cellular model designed to explore frataxin depletion in NRVMs. We used two different shRNA sequences to silence frataxin, limiting the possibility of off-target effects due to RNA interference. Moreover, the residual levels of frataxin observed in inhibited cultures are close to those observed in FRDA patients. Our results point to marked changes in the morphology of the mitochondrial network and alteration of lipid metabolism in frataxin-deficient NRVMs. Disruption of the mitochondrial network and the presence of enlarged mitochondria in frataxindeficient cells were observed using immunostaining against mitochondrial aconitase and two different probes (MitoTracker and TMRM). The origin of such changes is not clear. In cardiac myocytes, mitochondria represent more than 30% of the cellular mass. They exist primarily as individual entities with tight interactions with the sarcoplasmic reticulum. These interactions are crucial for energy transfer between mitochondria and myofibrils [47] and confer a characteristic spatial organization on the mitochondria [48]. However, mitochondria are dynamic organelles and alterations in the mitochondrial network have been observed in several human diseases [49] and under stress conditions [50]. Therefore, it is conceivable that the presence of enlarged mitochondria in frataxin-deficient cells could be related to a stress situation, as markers of oxidative stress were detected in our model. We can also hypothesize that a disturbed mitochondrial network could compromise the efficiency of energy transfer from mitochondria to myofibrils. Another relevant conclusion is that the cause of mitochondrial alteration cannot be attributed to a loss of proteins requiring iron–sulfur clusters for their activities, because the mitochondrial transport chain complexes and aconitase are not affected. This conclusion is also supported by the presence of high-potential mitochondria (as evidenced by the TMRM probe) and of normal ATP/ADP levels. Another important observation is the increased presence of lipid droplets in frataxin-deficient cells, which indicates impaired lipid metabolism. These lipid deposits are more prominent in fatty-acid-enriched media and therefore may be the consequence of an impaired lipid catabolism. Interestingly, lipid droplets have been observed in frataxin-deficient D. melanogaster [38] and in hearts from a neuron/cardiac frataxin conditional knockout mouse model (the NSE model) [17], indicating that such impairment in

Please cite this article as: Obis, È; et al. Frataxin deficiency in neonatal rat ventricular myocytes targets mitochondria and lipid metabolism. Free Radic. Biol. Med. (2014), http://dx.doi.org/10.1016/j.freeradbiomed.2014.04.016i

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Fig. 7. PPAR pathway. Protein extracts from 7-day-transduced cultures were probed with the indicated antibodies by Western blot analysis. In each gel lane, 15 μg of protein from whole-cell extracts was loaded. Protein load was verified by post-Western CBB staining. Histograms represent the quantification of the relative expression of the indicated proteins measured by Western blot. Protein content in scrambled control cells was used as a 100% reference value. PPARγ and CPT1 gene expression was assessed by qPCR. Actin expression was used as an internal control to normalize expression levels. In each experiment, the mRNA level under control conditions was used as the reference value. Data are presented as means 7 SEM from three independent cultures. np o 0.05 and nnp o 0.01; significant or highly significant difference compared to control condition.

lipid metabolism can also occur in vivo. Although the metabolism of the neonatal heart differs from that of the adult heart, the relevance of lipid metabolism in NRVMs is not negligible at all. In most mammals, expression of several enzymes required for lipid metabolism increases soon after birth, and fatty acid oxidation becomes the dominant oxidative substrate during the first week of age [51]. In rat heart, this may occur very early, as several acyl-CoA dehydrogenases reach 30 to 50% of adult levels at day 2 and remain at this level during the suckling period [52]. As the NRVMs used in this work were obtained at day 3 or 4, they had already experienced such increase in fatty acid oxidative capacity. In fact, several authors have demonstrated that NRVM cultures have the capacity to consume FA when present in culture medium [41,53], and our results confirm these previous observations. In addition, it is worth mentioning that alterations in lipid metabolism have been associated with mitochondrial dysfunction in other cardiopathies [54]. Another recent report indicates that mice heterozygous for Opa1, a protein involved in mitochondrial fusion, display altered lipid metabolism, enlarged mitochondria, and cardiac hypertrophy, whereas mitochondrial respiratory chain activities are maintained [55]. As these features resemble those described in our results, we can hypothesize that deficiency in

lipid metabolism in frataxin-deficient NRVMs can be a consequence of the altered mitochondrial network. Thus, a link between frataxin deficiency, mitochondrial dynamics, lipid metabolism, and cardiac dysfunction can be envisaged and merits further investigation. An additional point to consider is the involvement of the PPAR pathway in the metabolic alterations observed. PGC1α, which is a coactivator of PPARs, was decreased in skeletal muscle from a mouse model of Friedreich ataxia and in fibroblasts from FRDA patients. In contrast, it was slightly increased in HL-1 cardiomyocytes deficient for frataxin [42]. Our results are consistent with this last observation, as no significant changes in PGC1α content were detected by Western blot in frataxin-silenced NRVMs, but increased expression of some genes involved in lipid catabolism such as MCAD and CPT1 could be observed. As these genes are PPARα targets, its induction could be indicating a mild activation of this pathway, probably due to altered lipid metabolism, as PPARα can be activated by intracellular lipids [56]. In this context, upregulation of PPARα target genes MCAD and CPT1 (but not PPARα itself) was associated with intramitochondrial lipid accumulation, contractile dysfunction, and heart failure [57]. On the other hand, decreased expression of PPARγ could be a

Please cite this article as: Obis, È; et al. Frataxin deficiency in neonatal rat ventricular myocytes targets mitochondria and lipid metabolism. Free Radic. Biol. Med. (2014), http://dx.doi.org/10.1016/j.freeradbiomed.2014.04.016i

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compensatory mechanism to avoid an increase in FA uptake, as both PPARα and PPARγ promote FA uptake. Thus, alterations in the PPAR pathways in cardiac frataxin-deficient cells are most probably a consequence of altered lipid metabolism rather than its cause. Nevertheless, such alterations could result in further complications as mice with cardiac-specific knockout of PPARγ display cardiac hypertrophy and increased expression of NF-κB [44]. Interestingly, PPARγ agonists have already been proposed as therapeutic agents in FRDA, as the PPARγ agonist Azelaoyl PAF is able to increase frataxin levels in FRDA cells [58]. The decreased expression of this transcription factor in frataxin-deficient NRVMs could further support the use of such agents in FRDA patients. However, we should be cautious about this point, as PPARγ agonists could have an undesired effect such as promoting an increase in FA uptake without restoring mitochondrial alterations. Some features described in other models of the disease, or even in samples from FRDA patients, were not observed in our model. These include decreased ATP levels, lower aconitase activity, or altered mitochondrial complex function [15,29]. The amounts of the antioxidant enzymes SOD1 and SOD2 were increased in our model, consistent with previous observations in yeast [36]. However, these enzymes were decreased in other models of frataxin deficiency [37]. The reason for such discrepancies could be the stage at which those biochemical features were analyzed. For instance, in the MCK model, SOD2 protein content is slightly increased in frataxin-deficient heart at week 5, but decreased at week 10 [35]. In addition, loss of aconitase or complex II activities would be a late event promoted by oxidative stress or IRP activation. In agreement with this notion, the late activation of similar pathways leading to iron–sulfur protein deficiency has been reported in frataxin-deficient yeasts [59,60]. Another point for discussion is iron metabolism. In contrast to many models of the disease, total iron levels and expression of ironrelated genes were not affected in our model. Our results are consistent with those obtained in conditional mice and patient hearts, which indicate a mild or late effect on iron metabolism. Patient hearts do have iron deposits, but they are very localized and do not affect total heart iron levels [16]. In the MCK conditional mice, iron deposits and changes in the expression of iron metabolism genes are observed after 10 weeks of age [17,35]. Thus, our model provides useful data about the early effects of frataxin deficiency on cardiomyocytes, whereas many studies performed on human tissue or mouse models may detect later consequences of frataxin deficiency. In summary, the present research indicates that frataxin deficiency in NRVMs leads to mitochondrial dysfunction and oxidative stress, which would affect lipid metabolism. In the adult heart, these alterations could compromise the heart contractile function and the survival of patients with FRDA. Moreover, this cell model can be used to evaluate the effects of potential therapeutic interventions on cardiac cells.

Acknowledgments This work was supported by grants to J.T. (La Marató de TV3 2010) and to J.R. (BFU2010-19193, CSD2007-00020 Consolider Ingenio 2010 from the Ministerio de Economia y Competitividad (Spain), and SGR2009-00196 from the Generalitat de Catalunya). We thank Roser Pané for technical assistance. We also thank Elaine Lilly, Ph.D., for English language review.

Appendix A. Supplementary material Supplementary data associated with this article can be found in the online version at http://dx.doi.org/10.1016/j.freeradbiomed. 2014.04.016.

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