Human Molecular Genetics, 2001, Vol. 10, No. 19 2099-2107
© 2001 Oxford University Press
Frataxin expression rescues mitochondrial dysfunctions in FRDA cells
Department of Molecular Biosciences and 1Department of Nutrition, University of California, Davis, CA 95616, USA and 2Division of Biochemistry and Genetics, Istituto Nazionale Neurologico Carlo Besta, Milan, Italy
Received May 15, 2001; Revised and Accepted July 13, 2001.
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ABSTRACT |
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Friedreich’s ataxia (FRDA) is the result of mutations in the nuclear-encoded frataxin gene, which is expressed in mitochondria. Several lines of evidence have suggested that frataxin is involved in mitochondrial iron homeostasis. We have transfected the frataxin gene into lymphoblasts of FRDA compound heterozygotes (FRDA-CH) with deficient frataxin expression to produce FRDA-CH-t cells in which message and protein are rescued to near-physiological levels. FRDA-CH cells were more sensitive to oxidative stress by challenge with free iron, hydrogen peroxide and the combination, consistent with a Fenton chemical mechanism of pathophysiology, and this sensitivity was rescued to control levels in FRDA-CH-t cells. Iron challenge caused increased mitochondrial iron levels in FRDA-CH cells, and a decreased mitochondrial membrane potential (MMP), both of which were rescued in FRDA-CH-t cells. The rescue of the low MMP, and high mitochondrial iron concentration by frataxin overexpression suggests that these cellular phenotypes are relevant to the central pathophysiological process in FRDA which is aggravated by exposure to free iron. However, even at physiological iron concentrations, FRDA-CH cells had decreased MMP as well as lower activities of aconitase and ICDH (two enzymes supporting MMP), and twice the level of filtrable mitochondrial iron (but no increase in total mitochondrial iron), and the observed phenotypes were either fully or partially rescued in FRDA-CH-t cells. Free iron is known to be toxic. The observation that frataxin deficiency (either directly or indirectly) causes an increase in filtrable mitochondrial iron provides a new hypothesis for the mechanism of cell death in this disease, and could be a target for therapy.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Friedreich’s ataxia (FRDA) is an autosomal recessive neuro- and cardio-degenerative disease characterized by progressive ataxia and cardiomyopathy, with a prevalence of 1 in 50 000 individuals (1,2). FRDA is the result of mutations in frataxin, a 210 amino acid mitochondrial protein (3). Most cases of FRDA are caused by the expansion of a GAA trinucleotide repeat located in the first intron of the frataxin gene (4), resulting in decreased amounts of frataxin mRNA and protein, and there is an inverse correlation between the size of the GAA repeat and the amount of frataxin protein observed in lymphoblasts (5). About 2–5% of FRDA cases are compound heterozygotes (FRDA-CHs), having a GAA expansion at one allele and a point mutation at the other (6). Some FRDA-CH individuals have a very low level of frataxin protein and an earlier onset of disease (4,7,8).
Currently, both the correct physiological function of the frataxin protein, and the process by which its decrease results in pathophysiology are controversial. Deletion of the yeast frataxin homolog, yfh1, results in a 10-fold increase in mitochondrial iron, respiratory deficiency, increased sensitivity to oxidants and mtDNA depletion in some yeast strains, and one inference was that frataxin regulates mitochondrial iron homeostasis, perhaps through modulating mitochondrial iron export (9–11). Increased iron deposits have been observed in the hearts of FRDA patients (12) and deficiencies in activity of the iron-sulfur cluster (ISC)-dependent aconitase and mitochondrial respiratory chain complexes I, II and III have been demonstrated in tissues of FRDA patients (13–15). Although these data support a role for regulation of mitochondrial iron homeostasis by frataxin, there are also data which suggest that either mitochondrial iron homeostasis is differentially regulated, or frataxin-like molecules may function differently, in human and yeast cells. For example, FRDA fibroblasts exposed to supraphysiological concentrations of iron exhibit only a limited increase in mean mitochondrial iron/mg protein (maximum of 50%;16,17) in contrast to the 10-fold increase in mitochondrial iron observed in yeast (9). Furthermore, deletion of the Frda gene in the mouse results in embryonic lethality which may be the result of cardiomyopathy a few days after implantation, but with no evidence of iron accumulation (18). More recently, conditional Frda knockout mice exhibit a neuro- and cardio-degenerative phenotype which precedes the accumulation of iron (19). FRDA-CH individuals contain one allele with a GAA expansion and one with a point mutation. These individuals often have very low levels of frataxin expression and an early age of onset of FRDA symptoms. We have thus characterized multiple cellular, mitochondrial and iron-regulatory differences between FRDA-CHs and control lymphoblasts, and then determined which differences are rescued by frataxin transfection in an attempt to identify pathophysiological defects in FRDA-CH cells.
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RESULTS |
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Transfection of P131 and P585 results in increases of frataxin message and protein to near-physiological levels
To engineer cells to express the full-length frataxin protein at physiological levels, the complete coding sequence of human frataxin tagged with the HA epitope was cloned into the pcDNA3.1 vector and was transfected into two FRDA-CH lymphoblast lines (P131, P585). Transfections of control cells C333 and C621 to make C333-t and C621-t employed pcDNA3.1 with no insert. Two transfected cell populations with stable, high level expression of frataxin (P131-t, P585-t) were selected by growth in geneticin; controls transfected with empty vector were similarly selected (C333-t, C621-t). An increased level of steady-state frataxin RNA expression was observed in P131-t and P585-t relative to P131 and P585, by both semiquantitative and quantitative anchored-RT–PCR (Fig. 1A). The expression of the HA1-tagged frataxin was confirmed by dot blot using the anti-HA1 antibody (Fig. 1D). Untransfected P131 and P585 had 47 and 34% of control mRNA levels, respectively. Transfection resulted in an mRNA level of 147% (P131-t) and 85% (P585-t) of average control values by quantitative PCR on the LightcyclerTM, a 3.1- and 2.5-fold increase, respectively, compared with the untransfected cells (Fig. 1A and B). Untransfected P131 and P585 had 34 and 15% of control frataxin protein, respectively. Transfection rescued protein to near-physiological levels in both transfected lines by western blot densitometry (Fig. 1C).
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FRDA-CH cells are sensitive to oxidative stress, and transfection restores resistance
Earlier experiments with fibroblasts from FRDA homozygous for (GAA)n expansions have indicated increased sensitivity to free iron and hydrogen peroxide exposure (16,20); thus the sensitivity of FRDA-CH lymphoblastoid cells to oxidative stress was tested. An increased sensitivity of mutant cells to 1 and 2.5 mM iron chloride (FeCl3) and 1 mM hydrogen peroxide, and the combination of 0.5 mM iron and 1 mM hydrogen peroxide, was observed in FRDA-CH cells; frataxin-transfected cells essentially exhibited control sensitivity (Fig. 2A and B).
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Iron challenge causes increased mitochondrial iron in mutant cells, but not in control or mutant transfected cells. One possible explanation of the decreased viability of FRDA-CH cells compared with controls is that iron differentially enters the mitochondria of FRDA-CH cells to damage some mitochondrial function. Also, yeasts in which the frataxin homolog yfh1 is deleted have a large increase in mitochondrial iron (9). In contrast, fibroblasts from patients with two GAA expansions have a much more modest increase in mean mitochondrial iron (i.e. an apparent maximum of 50%), and this is under conditions of iron challenge (i.e. exposure to supraphysiological concentrations of free iron) (16,17). To address this we studied mitochondrial iron levels in cells exposed to 0.5 mM FeCl3, which does not cause large amounts of cell death. We observed that when grown in media with high concentration of FeCl3 (0.5 mM) for 16 h, mitochondrial iron levels were significantly higher in P131 and P585 mutants, 741 ± 94 (2 x SEM) nmol/mg of protein, than in controls 425 ± 92 (2 x SEM) nmol/mg of protein (Fig. 3). In contrast, in P131-t and P585-t cells challenged with free iron the mean mitochondrial iron level was 409 ± 54 (2 x SEM) nmol/mg of protein which is indistinguishable from the control values. In normally distributed data 2 x SEM = 95% confidence interval (CI). Thus challenge with supraphysiological levels of free iron does lead to excess iron in FRDA-CH mitochondria but not controls, and FRDA-CH-t mitochondria are rescued for this endpoint.
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Iron challenge results in lower mitochondrial membrane potential (MMP) in FRDA-CH cells as compared with controls, and membrane potential is rescued in transfected cells. Since challenge with free iron led to decreased viability and a higher level of mitochondrial iron in the FRDA-CH cells, it suggested that mitochondrial function might be compromised in these cells. One measure of mitochondrial bioenergetic function is the MMP. Therefore MMP was studied after exposure to 0.5 mM FeCl3 for 16 h (a concentration that does not cause excessive cell death in FRDA-CH cells), using the membrane potential probe, TMRM, on a flow cytometer (Fig. 4A and B). Iron challenge decreased the MMP more in the FRDA cells than in controls, and this phenotype was rescued in the frataxin-transfected cells. Thus a 0.5 mM micromolar free iron challenge induces a bioenergetic defect in CH lymphoblasts, which is rescued by frataxin transfection.
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MMP in FRDA-CH lymphoblasts is compromised even at physiological iron concentration. In previous studies (16,17), which have focused on FRDA cells from individuals with two GAA expansions, MMP has not been investigated. However, other mitochondrial bioenergetic parameters have been shown to be deficient in muscle of FRDA patients (14,15). Our measurements of MMP demonstrated a defect in FRDA-CH cells and rescue of this phenotype in transfected cells (Fig. 4A and B). When MMP was studied in FRDA-CH cells cultured in media without iron loading, it was found to be significantly lower as compared with controls (Fig. 4B). Also in this case, frataxin transfection rescued the phenotype (CH = 74.8 ± 6.4%; CTL = 100 ± 4.4%; CH-t = 94.0 ± 4.0%). The data therefore suggest that FRDA-CH cells have an intrinsic bioenergetic defect which is evident even at physiological iron levels, and is largely dependent on frataxin function as demonstrated by rescue of the phenotype following frataxin transfection. Frataxin expression partially rescues the decrease of aconitase and isocitrate dehydrogenase activities in FRDA-CH mutant lymphoblasts. The MMP is supported by the enzymes of mitochondrial oxidative phosphorylation, including the tricarboxylic acid (TCA) cycle and the electron transport chain. One TCA enzyme, aconitase, has been observed to be deficient in activity in FRDA patients and frataxin-deficient mice (13,15,19). We assayed aconitase activity in extracts of P131 and P585, and observed a significant decrease of total aconitase activity versus CTL cells C333-t and C621-t. On the other hand, there was a corresponding significant increase of mean aconitase activity in both P131-t and P585-t (Fig. 5A). In contrast to our other measurements which were very similar between the two mutant cell lines, there was significantly less aconitase activity in P585 compared with P131, which may result from a greater severity of the genotype of P585, (GAA)760/Y118 stop, versus the P131 genotype, (GAA)600/W173G. A similar enzyme activity profile was observed for another TCA cycle enzyme, isocitrate dehydrogenase, and all differences were significant at P < 0.05 or less (Fig. 5B). Thus frataxin overexpression significantly stimulates the activity of at least two TCA enzymes, which are significantly deficient in FRDA-CH cells. At physiological iron concentrations there is no significant iron excess in FRDA-CH lymphoblasts. To examine the possible causes of mitochondrial bioenergetic dysfunction and decreased enzyme activity at physiological iron concentrations (i.e. in the micromolar range), mitochondrial iron concentration was measured using the isotope 59Fe. Cells were loaded with 5 µM 59FeCl3 in the culture medium for a period of 16 h and the uptake of radiolabeled iron into CTL, FRDA-CH, and FRDA-CH-t mitochondria was compared. Although there was a small increase in the mean iron concentration in FRDA-CH versus control mitochondria, the difference was not statistically significant (918.3 ± 146.2 versus 773.6 ± 134.4 pmol/mg protein, P > 0.05, Fig. 6A).
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There is increased filtrable iron in CH cells, which is rescued by transfection. Iron exists in both high and low molecular weight forms in cells. The higher molecular weight forms are usually bound to protein or precipitated, whereas lower molecular weight forms are not. Low molecular weight, protein-unbound iron is thought to be more toxic than high molecular weight (i.e. precipitated or protein bound) iron because of the preferential ability of the former to undergo Fenton chemistry. The observation of a small (and statistically non-significant) difference in mean mitochondrial iron concentration at physiological iron levels suggested another experiment, i.e. to evaluate whether the ratio of iron in the protein-unbound form (i.e. with molecular weight > 10 kDa) was different in mutant and transfected cells. Thus cells were exposed to 5 µM 59FeCl3, the mitochondria purified and lysed, and filtrable versus non-filtrable counts separated on Centricon-10 membranes. There was no significant difference in either total or non-filtrable FRDA-CH, FRDA-CH-t and CTL cells. However, the FRDA-CH mitochondria exhibited a higher concentration of filtrable iron in all experiments (98.1 ± 9.2 pmol/mg protein) than CTL cell mitochondria (61.3 ± 13.3 pmol/mg protein), and the difference was significant, P < 0.05. Furthermore, values of filtrable iron were rescued in the frataxin-transfected FRDA-CH-t cells to within control concentrations, 63.0 ± 9.5 pmol/mg protein (Fig. 6C). Expressed in terms of percentages of total soluble counts, the values for unfiltrable iron for CTL, CH and CH-t were 92, 88 and 92%, respectively; and 8, 12 and 9%, respectively, for filtrable counts.
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DISCUSSION |
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Because of the mitochondrial localization of frataxin (21–23), the neuro- and cardio-degeneration observed in FRDA is thought to be the result of a mitochondrial defect. Deficiencies of mitochondrial enzymes and function have been observed in tissues of FRDA patients (13–15). Furthermore, the observation of iron deposits in FRDA heart and brain tissue (12,24), as well as the mitochondrial iron overload in some yeast strains deficient in the frataxin homolog yfh1p (9,11), have suggested a role for iron in the induction of the mitochondrial defect. The target cell types in FRDA, cardiocytes and neurons, are difficult to study in vitro. In contrast, fibroblasts and transformed lymphoblasts are more straightforward to study, but there have been few recent reports of differences between FRDA and control cells fibroblasts and lymphoblasts. We have studied lymphoblasts from FRDA-CH, which usually have an early age of onset and very low frataxin protein expression, and have reconstituted frataxin levels by transfection to near physiological levels by transfection.
FRDA-CH lymphoblasts were more sensitive to oxidative stress, i.e. challenge with free iron, and hydrogen peroxide, and free iron plus hydrogen peroxide, consistent with our earlier results of increased sensitivity in FRDA-GAA fibroblasts (16,20). Transfection with frataxin cDNA reconstituted frataxin to near-physiological levels, and rescued this cellular sensitivity. Since overexpression of frataxin specifically rescued from the excess sensitivity induced by free iron and hydrogen peroxide exposure, these results are consistent with a Fenton chemical mechanism of pathophysiology in these FRDA-CH cells and suggests that cellular conditions which increase free iron or hydrogen peroxide aggravate the pathophysiological process.
To try to determine the cause of the increased sensitivity of FRDA-CH cells to oxidative stress, mitochondrial iron levels and MMP were measured. A 50% increase in mitochondrial iron was observed in the FRDA-CH cells under conditions of iron challenge, consistent with earlier reports using cells homozygous for long (GAA)n expansions (16,17). Thus when challenged with iron, FRDA-CH cells become slightly overloaded with iron just as FRDA-GAA cells do. However, there is no large increase in mitochondrial iron as is the case in yeast yfh1 knockout in some strains (9,11), which may suggest differences in mitochondrial iron homeostasis between yeast and human FRDA-CH cells.
To try to determine what detrimental effect free iron challenge may have on mitochondrial bioenergetic function, MMP was measured. An iron-dependent decrease in MMP was observed in the FRDA-CH cells and MMP was rescued in frataxin transfected cells, implicating frataxin in the rescue of mitochondrial bioenergetic function. The stimulation of MMP by frataxin has been observed previously in different cell system, i.e. cells into which supraphysiological levels of frataxin were transfected (25).
Because challenge with free iron caused the three phenotypes (increased cellular toxicity, increased mitochondrial iron concentrations, decreased MMP) which were rescued by frataxin, it is reasonable to infer that these phenotypes are relevant to the pathophysiological process in FRDA. This presumably involves decreased cellular viability which is the result of some deficiency of mitochondrial bioenergetic function or activation of apoptosis which is aggravated by exposure to free iron.
The experiments also demonstrated an intrinsic bioenergetic defect in the CH cells that has not been previously reported, i.e. that these FRDA-CH cells had an intrinsic decrease in MMP, that is, even in the absence of oxidative challenge. The observation of a decreased MMP in unstressed cells, and the rescue of MMP by frataxin transfection, suggested that mitochondrial dysfunction could be studied even in the absence of exposure to oxidative stress.
The deficiency in MMP in FRDA-CH cells also suggested that deficiencies may exist in mitochondrial enzymes that support MMP. It was also observed that FRDA-CH cells had significantly decreased activities of total aconitase and ICDH, and transfection with frataxin significantly activated these activities. Because the TCA cycle and electron transport chain are coupled, and because the electron transport chain supports the membrane potential, a simple explanation for the decreased MMP is decreased mitochondrial enzyme activities, at least those of aconitase and ICDH. This may not be a complete explanation however, as frataxin transfection appeared to completely rescue MMP, but only partly rescues aconitase and ICDH activities. It is also interesting to note that while aconitase requires an ISC for its activity, ICDH does not. Thus if the physiological function of frataxin was at some stage of ISC assembly, then one might not expect ICDH deficiency in FRDA-CH cells, and restoration by frataxin, unless there were effects of inhibition of ISC enzymes on non-ISC enzymes. In fact there is precedent for defects in ISC assembly having negative effects on non-ISC enzymes in mitochondria (26,27).
To better understand what may cause the bioenergetic defect in FRDA-CH mitochondria in the absence of iron challenge, the concentration and condition of iron was measured in the mitochondria. The observation that mitochondrial iron levels in FRDA-CH, frataxin-transfected and normal control cells were similar suggests that mitochondrial iron overload may not be a primary feature of the disease process in FRDA-CH cells. These results are consistent with results from the frataxin knockout mice, in which apparent cardiomyopathy and embryonic lethality occur without iron accumulation (18), and the frataxin conditional knockout mice, in which neuro- and cardio-degeneration precede iron accumulation (19). In contrast, the relatively high ratio of mitochondrial filtrable:non-filtrable iron in CH cells, and its modification following frataxin transfection, suggests that frataxin deficiency (either directly or indirectly) alters the ratio of free to bound iron within mitochondria, i.e. causing an increase in free mitochondrial iron. The levels of free iron are normally very low in the cell, and increases in free iron are known to be toxic.
The observed increase in filtrable iron, and its rescue by frataxin, could have multiple causes, depending on both the correct physiological role of frataxin and the pathogenic mechanism of FRDA. Currently there are at least four non-exclusive hypotheses for frataxin’s correct physiological role, including mitochondrial: (i) iron efflux pump (9); (ii) iron binding/sequestration protein (28,29); (iii) ISC assembly promoter (11,30); and (iv) general stimulator of oxidative phosphorylation (25). If the increase in mitochondrial free iron we observe is the direct result of frataxin deficiency, then this result seems most simply explained by hypotheses (ii) and (iii), because a decreased concentration of an iron sequestration protein, or a decreased activity of insertion of iron into ISCs, could be simply imagined to cause an increase in low molecular weight mitochondrial iron. Our data aren’t easily consistent with hypothesis (i) because we observe no significant change in mitochondrial iron concentration in unchallenged mutant cells, only a shift to a more filtrable state of iron. It is not easy to understand how high or low iron transport would affect the state of iron in the mitochondria. Our membrane potential and enzyme activity data are also consistent with hypothesis (iv), in that as did Ristow et al. (25), we also observe a stimulation of mitochondrial functions (i.e. membrane potential and mitochondrial enzyme activity) in frataxin transfected cells. However, it is difficult for us to imagine how a defect in a stimulator of OXPHOS would cause an increase in free mitochondrial iron, although many studies have shown the converse, i.e. that increasing free iron does cause mitochondrial dysfunction. Nevertheless, there are no published data to our knowledge that exclude the possibility that a decrease in mitochondrial function could cause the increase in free iron observed, and so this remains a formal possibility.
Irrespective of the relationship between the correct role of frataxin (which is still controversial) and the increase in mitochondrial free iron we observed, which could be a direct or indirect consequence of frataxin function, there are multiple hypotheses for the proximal causes of neurodegeneration and cardiodegeneration observed in FRDA. One major hypothesis is decreased mitochondrial function in mutant cardiocytes and neurons that is the result of either: (i) increased mitochondrial superoxide generation (13); (ii) increased free iron (this work); (iii) starvation for ISC (11,30); or (iv) depletion of a mitochondrial OXPHOS activator (25). A second major hypothesis is activation of the apoptotic program in mutant cells (16), which could be the result of direct mitochondrial damage mediated by superoxide or free iron. Each of these makes different predictions regarding the proximal toxic event which causes the neuro- and cardio-degeneration in FRDA. Explicit testing of the hypotheses for both primary frataxin function, and the proximal cause of cell death in FRDA cells could provide a rational basis for design of pharmacotherapy.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Vectors
The pcDNA3.1 vector was purchased from Invitrogen (Carlsbad, CA). Polyclonal antibody against frataxin was obtained from the Laboratory of Cellular Pathology, Istituto Nazionale Neurologico Carlo Besta (Milan, Italy). Anti-HA1 monoclonal antibody was purchased from Boehringer Mannheim (Indianapolis, IN). Secondary antibodies and chemiluminescent western detection kit were purchased from Bio-Rad (Hercules, CA). Escherichia coli-competent cells, RT–PCR system, DMRIE-C, cytokines and PHA were purchased from Life Technologies (Gaithersburg, MD). SV total RNA isolation system and Miniprep DNA purification system were purchased from Promega (Madison, WI). Most biochemical reagents were from Sigma-Aldrich (St Louis, MO). Fluorescent dyes were obtained from Molecular Probes (Eugene, OR).
Cell lines and mutation detection
Peripheral blood lymphocytes from two FRDA patients with severe clinical phenotypes and from two normal controls were transformed with EB virus and grown in the RPMI-1640 medium (Life Technologies) supplemented with 15% fetal bovine serum, 50 mM sodium pyruvate, 100 U/ml penicillin and 100 µg/ml streptomycin in a humidified atmosphere containing 5% CO2 at 37°C. Cells were grown at an approximate density of 5 x 105 cells/ml, with daily supplementation of fresh medium, and complete change of medium every 7 days. Two FRDA patients (P131 and P585) referred to the Istituto Nazionale Neurologico ‘C. Besta’ (Milan, Italy) were identified to be compound heterozygous for a (GAA)n expansion and a mutation in the frataxin gene by long-range PCR analysis of the (GAA)n tract and direct sequence of the five frataxin exons, respectively (C.Gellera and F.Taroni, manuscript in preparation). The genotype of P131 was (GAA)600/W173G while the genotype of P585 was (GAA)760/Y118stop. The oligonucleotide primers used for sequencing and long-range PCR were as described previously (31).
Plasmid constructs and gene transfection
Plasmid pcDFRDAHA1 was derived from the parental vector pcDNA3.1 and encodes the 210 amino acid human frataxin precursor tagged at the C-terminal end with the HA1 epitope. The transcriptional unit was under the control of the CMV immediate-early promoter. The plasmid also encodes the geneticin resistance gene for selection of transfectants. The inserted sequence was confirmed by DNA sequencing. Plasmid DNA was prepared using a DNA miniprep commercial kit (Promega, WI). DNA quality was determined by restriction endonuclease digestion and quantified by UV spectrophotometry. Four lymphoblast lines were prepared by growing in fresh medium for 16 h, and transiently transfected with 2 µg/ml pcDFRDAHA1 expression vector or pcDNA3.1 empty vector, or 1 µg/ml of the reporter gene plasmid pCMV.sport-ßgal using DMRIE-C (Life-Tech, CA) according to the manufacturer’s protocol for suspension cells. Each transfection was performed in triplicate in 6-well plates with 2 µg of plasmid DNA, 6 µl of DMRIE-C and 2 x 106 cells mixed in 1.2 ml/well of OPTI-MEM low-serum medium. Five hours after transfection, fresh culture medium was added. Twenty-four hours after transfection, cells were stained with X-gal to determine transfection efficiency and selected with 400 µg/ml geneticin for 12 days. Frataxin gene expression was examined by semiquantitative and quantitative RT–PCR and anchored-RT–PCR, western blot and dot blot as described below. Cell lines expressing high frataxin levels were selected for assays, and aliquots of cells were frozen for experiments. Frataxin mRNA expression levels were periodically examined by quantitative RT–PCR on the lightcycler.
Quantitative RT–PCR analysis
Total RNA was prepared using the SV total RNA extraction kit (Life Technologies), and RNA concentration was determined by UV spectrophotometry. Reverse transcription of 1 µg RNA was performed using an RT–PCR kit (Life-Tech), and reactions were performed in 20 µl volume. Two microlitres of cDNA from 20x dilution of RT reaction was used for PCR. Plasmid DNA of pcDFRDAHA1 and ß-actin cDNA from RT–PCR were prepared and purified, and DNA concentration was determined with an UV spectrophotometer, and the corresponding copy number was calculated. Serial dilution of 10 ng:0.1 pg plasmid DNA and ß-actin cDNA were prepared in ddH2O for quantitative PCR standard curve. The primers of PCR are as follows:
Frataxin (323 bp) forward, 5'-ACT AGC AGA GGA AAC GCT GG-3'; reverse, 5'-GGA ATA GGC CAA GGA AGA CA-3'; FRDAHA1 (318 bp) forward, 5'-GAA GAC CTT GCA GAC AAG CC-3'; reverse, 5'-TGG GAC GTC ATA TGG ATA AGC-3'; ß-actin (553 bp) forward, 5'-AGA AAA TCT GGC ACC ACA CC-3'; and reverse, 5'-AGG AAG GAA GGC TGG AAG AG-3'. All PCR reaction mixtures contained 1x PCR buffer (200 mmol/l Tris–HCl pH 8.4, 500 mmol/l KCl), 2.5 mmol/l MgCl2, 0.8 mmol/l dNTP (Life Technologies), 0.2 µmol/l each primer, and 1 U of Platinum Taq DNA polymerase (Life Technologies), Syber green, and 1.5 g/l bovine serum albumin (BSA; Sigma). Reactions on the LightCycler (Roche Diagnostics, Indianapolis, IN) were performed in 20 µl volumes. The standard temperature profile for PCR included an initial denaturation for 1 min at 95°C, followed by 30 cycles of denaturation at 95°C for 0 s, annealing at 56°C for 5 s, and an extension with fluorescence monitoring at 68°C for 15 s. The final frataxin copies were nomalized by ß-actin cDNA copies.
Western blot analysis
Cells were lysed in a buffer containing 20 mM HEPES pH 7.4, 100 mM NaCl, 0.1% deoxycholic acid, 1% NP-40, 1 mM EDTA, 1 mM EGTA, 10% glycerol, 1 mM PMSF, 10 µg/ml aprotinin, 10 µg/ml leupeptin and 50 mM sodium fluoride at 4°C for 30 min. Insoluble material was removed by centrifugation, and equal amounts of lysates (20 µg) were used for immunoblotting. Protein concentration was estimated using the Bradford protein assay system (Bio-Rad). Samples were resolved on a 15% SDS–polyacrylamide gel and then transferred to a PVDF membrane (Millipore, Bedford, MA) by electroblotting. After blocking with 5% non-fat dry milk, the blot was incubated with anti-frataxin polyclonal antibody (0.5 µg/ml, total serum protein) and was developed with AP-conjugated secondary antibodies using a chemiluminescent substrate. A densitometer was used for quantitation of signals on films. Dot blots were performed for detection of HA1 epitope tag of pcDFRDAHA1 in transfected cells. 0.5 µg protein was dotted on the PVDF membrane, and western blot procedure was conducted.
Enzyme assays
To stabilize the ISC of aconitase, total cell extracts were prepared by disrupting the cells resuspended in HDGC buffer (20 mM HEPES pH 7.4, 1 mM DTT, 10% glycerol and 2 mM sodium citrate, 0.5 µg/ml leupetin, 0.7 µg/ml pepstatin and 0.2 mM PMSF) using a microtip sonic oscillator (Fischer Scientific, Pittsburg, PA). Isocitrate dehydrogenase activity was measured by modifying the method of a Sigma kinetic kit. L-Isocitrate was mixed with NADP, protein extracts in the presence of excess Mn2+. The conversion of NADP to NADPH was spectrophotometrically monitored over time at 340 nm at 30°C. Aconitase activity was determined by coupled reaction of aconitase and isocitrate dehydrogenase (32). The change of absorbance by NADPH at 340 nm was followed at 30°C. Protein concentration was assayed by the Bradford method using BSA (fraction V) as the standard.
Cell viability assay
Cell viability was determined as described (16). Briefly, 5 x 105 cells/ml were grown in RPMI 1640 medium 16 h prior to use. Cells were plated in 12-well plates at 5 x 105 cells in 0.2 mM uridine medium, and multiple different stresses were carried out. Viable and dead cells were counted by trypan blue exclusion assay. Six independent experiments were repeated, and Student’s t-test was carried out to determine the significance.
Determinations of total and filtrable iron at 0.5 mM and 5 µM iron
Cells were grown in fresh growth media in presence or absence of 0.5 mM FeCl3 for 16 h, mitochondria were isolated from lymphoblasts as described previously (32). Iron concentration was determined by Bradford assay. Mitochondrial iron was measured by flame atomic absorption spectrophotometry (16). Five independent experiments were carried out. For low-iron measurements, cells were incubated with 5 µM 59FeCl3 for 16 h, and then mitochondria were isolated as above, and then lysed with 100 µl lysis buffer (0.5% Triton X-100, 50 mM NaCl, 10 mM Tris–HCl pH 7.4). Protein concentration was determined, and the solution was filtered through Centricon-10 membranes (Amicon) by centrifugation at 14 000 g for 50 min. The supernatant (non-filtrable) and flow-through (filtrable) radioactivity were counted using Gamma 5000 counter. These experiments were repeated five times.
Measurement of MMP
MMP was measured in digitonin-treated cells with TMRM by FACScan flow cytometry as described previously (33). Briefly, cells were grown in fresh media with or without 5 µM or 500 µM FeCl3 for 16 h, then washed three times with cold PBS, and resuspended in KCl medium (80 mM KCl/10 mM Tris–HCl/3 mM MgCl2/1 mM EDTA/5 mM KH2PO4 pH 7.4) containing 10 mM succinate and 1 µM rotenone. Cells (1 x 106) were incubated with 10 ng digitonin in 1 ml media for 5 min on ice, washed two times and resuspended in 1 ml KCl buffer. Cells were incubated with 100 nM TMRM at room temperature for 10 min, and washed two times, and resuspended in 2 ml medium. Aliquots of cells were incubated in 10 nM DNP for 5 min before measurement of fluorescence. Cytofluorimetric analysis was performed on the FACScan flow cytometer (Becton-Dickinson, San Jose, CA) equipped with a 488 nm argon laser. TMRM signal was analyzed in the FL2 channel, equipped a bandpass filter at 580 ± 30 nm; the photo-multiplier value of the detector was 631 V. Data were acquired on a logarithmic scale. Arithmetic mean values of the fluorescence signal in arbitrary units were determined for each sample for subsequent graphical representation. All experiments were repeated three times.
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ACKNOWLEDGEMENTS |
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This work is supported by USPHS grants AG11967, AG16719, EY12245, a pilot project supported by P30ES05707 to G.A.C. and a Telethon-Italia grant (E.514) to F.T.
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FOOTNOTES |
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+ To whom correspondence should be addressed. Tel: +1 530 754 9665; Fax: +1 530 754 9342; Email: gacortopassi@ucdavis.edu
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REFERENCES |
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