Human Molecular Genetics, 2001, Vol. 10, No. 18 1935-1944
© 2001 Oxford University Press
Frataxin deficiency enhances apoptosis in cells differentiating into neuroectoderm
1Department of Medicine, Centre Hospitalier de l’Université de Montréal, Hôpital Notre-Dame, Montréal, Québec, H2L 4M1, Canada, 2UnIGENe, Instituto de Biologia Molecular e Celular, 4150-180 Porto, Portugal
Received April 30, 2001; Revised and Accepted July 5, 2001.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Deficiency of the mitochondrial matrix protein frataxin causes Friedreich ataxia. Frataxin function is believed to be related to mitochondrial iron metabolism and free radical production. In Friedreich ataxia, loss of dorsal root ganglia neurons occurs early in life, suggesting a developmental process. In addition, frataxin knockout mice die during embryonic life, further suggesting that frataxin is necessary for normal development. In this study we examine the role of frataxin in neuronal differentiation by using the P19 embryonic carcinoma cell line as a model system. We produced stably transfected clones with antisense or sense frataxin constructs. During retinoic acid-induced neurogenesis of frataxin-deficient cells there was a striking rise in cell death, while cell division remained unaffected. However, frataxin deficiency does not affect cell survival in cells induced to differentiate into cardiomyocytes. Frataxin deficiency enhances apoptosis of retinoic acid-stimulated cells, and the number of neuronal-like cells expressing MAP2 was dramatically reduced in these clones. In addition, we found that antisense clones induced to differentiate into neuroectoderm with retinoic acid have increased production of reactive oxygen species, and that only cells non-committed to the neuronal lineages could be rescued by the addition of the antioxidant N-acetyl-cysteine (NAC). However, NAC treatment had no effect in increasing the number of terminally differentiated neuronal-like cells in frataxin-deficient clones. Our results suggest that frataxin deficiency may render cells susceptible to apoptosis after exposure to appropriate stimuli.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Friedreich ataxia (FRDA) is an autosomal recessive degenerative disease affecting the nervous system and the heart (1,2). Its neuropathology is characterized by atrophy of the sensory pathways, with early loss of large neurons in the dorsal root ganglia (DRG), sensory axonal neuropathy and degeneration of the posterior columns of the spinal cord (2,3). Onset is usually in childhood or adolescence, but may be delayed to middle or later years of life (4–6).
A considerable breakthrough towards the understanding of the molecular pathogenesis of FRDA was achieved by the discovery of its causative gene (7). The encoded protein, named frataxin, is localized in mitochondria (8–11). Frataxin is highly conserved in evolution, with homologs in essentially all eukaryotes and some prokaryotes (7,8). Yeast cells with a disrupted frataxin homolog gene (YFH1) accumulate 10-fold more iron in mitochondria than wild-type, lose mitochondrial DNA, and become unable to carry out oxidative phosphorylation (9,12). Loss of respiratory competence requires the presence of iron and occurs more rapidly as iron concentration in the medium is increased (13). The mechanism of mitochondrial iron accumulation is unknown, but Yfh1p has been shown to induce a flux of non-heme iron out of mitochondria (13). Iron in mitochondria amplifies the toxicity of reactive oxygen species (ROS) leaking from the respiratory chain. The free hydroxyl radical (OH·), in particular, may be produced by Fenton chemistry and causes lipid peroxidation, protein and nucleic acid damage. Occurrence of the Fenton reaction in 
YFH1 yeast cells is suggested by their enhanced sensitivity to H2O2 (9). Many lines of evidence indicate that frataxin function is conserved in humans (14), suggesting that the mechanism by which cellular damage might occur in FRDA patients involves mitochondrial dysfunction triggered by ROS-mediated damage (15,16).
That frataxin plays an important role during early development is exemplified by the observation that frataxin knockout mice die in utero shortly after implantation at embryonic day (E)6.5 (17).
In order to study the role of frataxin during cell differentiation and development, we turned to an embryonic carcinoma (EC) cell model system, the P19 mouse EC cells, which can be induced to differentiate into a variety of cell types (18,19). P19 cells resemble those of the inner mass of the blastocyst, and their differentiation is believed to closely mimic critical events in early embryogenesis. Under appropriate culture conditions, P19 cells display the ability to differentiate into derivatives of three germ layers; endoderm, mesoderm and ectoderm. Treatment of aggregated P19 cells with retinoic acid (RA) effectively induces the development of neurons, astroglia and microglia, cell types normally derived from the neuroectoderm (18). Aggregates of P19 cells exposed to dimethyl sulfoxide (DMSO) differentiate into endodermal and mesodermal derivatives, including cardiac and skeletal muscle (19). In this study we evaluated how frataxin levels change during differentiation and characterized the effect on morphology, growth and differentiation of stable transfectants containing either a frataxin sense or antisense vector.
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RESULTS |
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Frataxin expression during neuronal and cardiomyocyte differentiation
P19 mouse EC cells are induced to differentiate into neuronal precursors upon treatment with RA and aggregation on bacterial-grade Petri dishes for 4 days, followed by dissociation and 4 days further growth on tissue culture plates (19). Untreated P19 cells grow densely packed and display a characteristic cuboidal morphology, evident in phase contrast microscopy. Treatment with 1 µM RA results in a dramatic change in morphology and differentiation to a neuron-like phenotype with neurite-like morphology. Neuronal differentiated P19 cells are prominently stained by an antibody directed against the microtubule associated protein-2 (MAP2).
We investigated frataxin expression in P19 cells during differentiation at the protein level by immunoblotting. Frataxin levels were found to increase 2–3-fold over levels in untreated cells upon aggregation in the presence of RA between days 2 and 4, followed by a decrease after dissociation and re-growth on tissue culture plates (Fig. 2A). By comparison, ß-tubulin levels remained unchanged. The development of neurons was confirmed here by the appearance of the neuron-specific marker MAP2.
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We next examined frataxin expression during cardiomyocyte and endodermal differentiation. To induce muscle and cardiomyocyte differentiation, P19 cultures were grown as aggregates in bacterial dishes in the presence of 1% DMSO and then plated in cell culture-treated dishes on day 4 without DMSO. Rhythmically beating regions were detectable by microscopic examination from day 6. By immunoblot, frataxin levels were observed to dramatically increase from day 2 (Fig. 2B), in a very similar way as when cells were differentiated primarily into endodermal cells by aggregation in the absence of inducer (Fig. 2C). An actin antibody (clone 5C5) and the TROMA-1 antibody were used to follow the synthesis of cardiac actin and cytokeratin 8 intermediate filament (EndoA) during cardiomyocyte and endodermal differentiation, respectively (Fig. 2B and C).
To further explore whether the increased frataxin levels during P19 cell differentiation were related to an increase in the number of mitochondria, we examined the levels of other mitochondrial proteins, such as cytochrome-c and COX IV. The variations in the levels of these proteins and frataxin were very similar when cells were exposed to DMSO (Fig. 2B), as well as when cells were differentiated in the neuronal lineage (Fig. 2A). Hence, frataxin induction during P19 differentiation may eventually be explained by an increase in mitochondrial number.
P19 clones stably transfected with antisense or sense frataxin
To investigate the role of frataxin in RA- and DMSO-induced differentiation of P19 cells, expression of frataxin was suppressed via constitutive expression of antisense RNA, or overexpressed in cells transfected with sense RNA. The vector employed, the pCI-neo, takes advantage of the cytomegalovirus (CMV) promoter which has been shown to express antisense RNA and to suppress targeted protein expression in other teratocarcinoma cells (20). We obtained four ‘empty-vector’ transfected control clones, which had similar frataxin levels to wild-type, non-transfected cells, and were therefore pooled together and designated NP, three antisense harboring clones (clones AS7, AS11 and AS12) and two frataxin overexpressing clones (OV1 and OV8). Frataxin expression was monitored by immunoblot (Fig. 3A) and RNAse protection assay (data not shown). Basal levels of frataxin RNA and protein were significantly reduced in the antisense clones, while in the sense clones they were increased compared with control, empty-vector transfected clones.
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Frataxin deficiency diminishes the number of multipotent, neuronal precursors and terminally differentiated neuronal cells, but does not affect cell division of precursors cells
To begin to monitor the effect of frataxin deficiency on neuronal differentiation, we first examined cell viability either by trypan blue exclusion or by vital dye reduction (Fig. 3B). Expression of antisense frataxin resulted in a 2–5-fold decrease in total cell numbers after RA induction, dissociation and re-growth on tissue culture plates. In contrast, frataxin-overexpressing clones showed a 2–5-fold increase in cell number compared with clones transfected with empty pCI-neo.
We next monitored the appearance of differentiated neurons by using antibodies to neuronal markers, such as MAP2. Stable transformants expressing antisense frataxin RNA to diminish frataxin protein levels were treated with RA to induce neuronal differentiation. After 8 days, cells were fixed and stained with an antibody to MAP2. Many MAP2-positive cells appeared in the control cultures (Fig. 1B and C) and in frataxin overexpressing cultures (Fig. 1D), whereas the number of MAP2-positive cells was dramatically reduced in the frataxin-deficient cultures (Figs 1A and 4A–C). By light microscopy, the size and morphology of aggregates of antisense clones were definitely different from all the other clones, empty vector or frataxin sense transfected, and wild-type P19 cells. The few differentiated MAP2-positive cells in the antisense clones had abnormally shortened neurites (Fig. 1A) and frataxin, readily detectable in mock-transfected cultures by immunocytochemistry, remained undetectable in all morphologically distinct cells from antisense cultures (data not shown). Interestingly, overexpressing clone displayed somewhat longer and well-defined neurites (Fig. 1D). The number of MAP2-positive cells however, was not significantly higher in the frataxin overexpressing cultures compared with the cultures with normal frataxin levels (data not shown).
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Thus, frataxin-deficient cells stimulated with RA had significantly reduced levels not only of cells non-committed to neuronal lineages, but also of cells differentiated into neuron-like, MAP2-positive cells (Fig. 4A–C).
Next, we determined whether the decrease in MAP2-positive cells in frataxin-deficient cell cultures was caused by a decrease in the number of precursor cells. We monitored the appearance of multipotent neuronal precursors (day 3) and unipotent precursors (day 4) by staining RA-treated cells with anti-nestin and anti-Hu antibodies, respectively. The numbers of both nestin-positive and Hu-positive cells in antisense cultures were significantly smaller than in the control cultures (Fig. 5A–D). Thus, frataxin deficiency affects the numbers of both multipotent and unipotent neuronal precursors.
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We next determined whether frataxin deficiency affects the proliferation capacity of multipotent precursor cells, thus causing the significant reduction in precursor cell number. Control and frataxin-deficient cultures were treated with RA, and then [3H]thymidine was added at different times to assess cell proliferation. [3H]thymidine incorporation was similar in the control and frataxin-deficient cultures (data not shown). This finding suggests that frataxin deficiency has no significant effect on cell division of precursor cells.
Frataxin deficiency does not affect cardiomyocyte differentiation
Next we examined the ability of each of the clones to differentiate into functioning cardiomyocytes after DMSO treatment by scoring the percentage of aggregates which formed beating regions. The size and morphology of aggregates of all clones appeared identical to aggregates of wild-type P19 cells. Similarly, differentiated cultures of all clones displayed heterogeneous differentiated morphologies, which are readily distinguishable from the smaller, more regular-shaped undifferentiated P19 cells. The percentage of colonies with any rhythmically contracting regions detectable by light microscopy was similar in all tested clones (Fig. 4D).
Rescue of frataxin-deficient cells non-committed to neuronal lineages by N-acetyl-cysteine (NAC)
Some lines of evidence suggest that the mechanism by which cellular damage might occur in FRDA patients involves mitochondrial dysfunction triggered by ROS-mediated damage (15,16). We tested whether anti-oxidant treatment could rescue the growth of RA-induced frataxin-deficient cells. We found that 2–10 mM NAC could indeed increase the total cell numbers of antisense clones after RA stimulation (Fig. 3C). We also tried to rescue frataxin-deficient cells with ascorbic acid, 
-tocopherol, idebenone and desferioxamine and found that these agents failed to rescue frataxin-deficient cells (data not shown). If anything, desferoxamine, even at low concentrations (20–200 µM), had a dramatic effect in increasing cell death to virtually all cells, in all tested clones, suggesting that RA-stimulated P19 cells are exquisitely sensitive to iron deprivation (data not shown). Iron supplementation, in the form of ferric ammonium sulfate (FAC) to differentiating clones, produced huge aggregates, with no significant alterations in total cell numbers at FAC concentrations up to 100 µM, while higher concentrations increased cell death in all clones, but no significant differences were observed between the frataxin-deficient, frataxin-overexpressing and neo cultures (data not shown). We unsuccessfully attempted to rescue frataxin mutant embryos (frataxin knockout mice) (17) by administration of NAC, as well as the above-mentioned antioxidants and desferoxamine, or even iron supplementation, to pregnant dames.
We also tested the capacity of NAC to rescue more specifically the number of multipotent, unipotent precursors and terminally differentiated neuron-like, MAP2-positive cells. We found that NAC had no influence in the particular rescue of these cell subtypes (Figs 4C and 5A–D). These data suggest that NAC acts only on cells that are not committed to the neuronal lineage, presumably increasing survival rates in these cells.
Increased apoptosis in frataxin-deficient P19 cells induced to differentiate with RA
Apoptotic cell death of differentiating cells is found widely in the developing fetal brain (21). Similarly, apoptosis is also observed during neuronal differentiation of P19 cells (22,23). We therefore reasoned that low frataxin levels could enhance apoptotic cell death of differentiating cells, leading to a reduction in the number of surviving cells. To test this hypothesis, DNA was isolated from RA-treated cultures and subjected to gel electrophoresis and visualization with ethidium bromide. DNA from antisense clones was found to be fragmented 24–72 h after initial exposure to RA (data not shown). DNA fragmentation, albeit later, was also evident in RA-treated control cultures and frataxin-overexpressing cultures. Quantification of DNA fragmentation by [14C]thymidine incorporation followed by elution clearly shows that all antisense clones tested undergo extensive apoptosis compared with empty-vector transfected clones (Fig. 6A). No quantitative differences in DNA fragmentation were observed between empty-vector transfected clones and frataxin-overexpressing clones (data not shown). Similar results were obtained when DNA fragmentation was detected using the TUNEL assay followed by flow cytometry analysis (data not shown). Significantly, when cells were treated with DMSO to induce cardiomyocyte differentiation, no DNA ladder was evident in parental P19 cells or in any other frataxin sense, antisense and empty-vector transfected clones (data not shown).
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Increased levels of MnSOD in frataxin-deficient clones
The results so far indicate that the extensive cell death occurring in RA-treated frataxin antisense clones is a consequence of increased vulnerability to apoptosis. We therefore examined the expression of genes believed to play pivotal roles in apoptosis. Bcl-2 (20) and MnSOD (24) are believed to play a critical role in protecting neuronal cells from apoptosis, namely by affecting the antioxidant status of cells. We analyzed Bcl-2 and MnSOD protein expression after RA-induction of frataxin-deficient clones. Untreated control and antisense clones have undetectable levels of Bcl-2, which markedly increased after RA exposure (Fig. 6B). The levels of MnSOD proteins were lower in untreated anti-sense cultures (3-fold decrease), and persistently higher (2-fold increase on average between days 1 and 8) in the same clones after RA treatment compared with empty-vector transfected clones. During all the differentiation process, antisense clones have visibly less frataxin levels than control clones (Fig. 6B), a feature also confirmed in DMSO treated cultures (data not shown). These findings suggest that frataxin-deficient cultures are subjected to increased oxidative stress.
Increased ROS production in frataxin-deficient P19 cells induced to differentiate with RA
An important mechanism by which many stimuli induce apoptosis is through generation of ROS (25). To assess intracellular ROS production, we measured oxidation of 2',7'-dichlorofluorescein (DCF) in cell cultures allowed to form floating aggregates with or without RA. RA treatment of antisense clones for 24 h resulted in significant increase in ROS production, while neo-empty vector clones and wild-type clones had rather modest amounts of ROS production compared with aggregated cells in the absence of RA (Fig. 7). In the presence of NAC (5 mM), which can significantly increase total cell numbers of RA-induced antisense clones (Fig. 3C), the ROS production was only modestly inhibited in RA-treated cells. These data suggest that apoptosis susceptibility of frataxin-deficient cells upon RA stimulation may be at least partially mediated by ROS. They also indicate that rescue by NAC of cells non-committed to the neuronal lineages by RA stimulation is not due to a direct decrease in ROS production.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Clinicopathologic studies (26–31) and the generation of frataxin knockout mice (17) indicate that frataxin may play an important role during early development. However, further studies on the developmental role of frataxin in knockout mice are hampered by the early embryonic lethality of this animal model. In this study, we examined the role of frataxin in neuronal and cardiomyocyte differentiation by using the P19 EC cell line as a model system. We show that frataxin levels are induced during differentiation into all tree lineages, a feature that can presumably be explained by an increase in number of mitochondria. Stable transfection of undifferentiated P19 cells with a frataxin antisense construct drastically reduced the total number of cells in culture upon RA induction. The drastic reduction in total cell numbers also, but not exclusively, includes terminally differentiated neuronal-like cells, MAP2-positive cells. The loss from antisense cultures of cells committed to the neuronal lineages starts at early stages of RA stimulation, with a reduction in the number of multipotent and unipotent neuronal precursors, resulting from enhanced apoptotic cell death rather than from inhibition of precursors cell division. Furthermore, the surviving MAP2-positive cells in these cultures displayed shortened neurites. Treatment of RA-induced frataxin-deficient clones with the anti-oxidant NAC was able to rescue only cells that were not committed to the neuronal lineage, since no effect on multipotent or unipotent precursors was observed. These observations suggest that frataxin-deficient cells committed to neuronal lineages by RA exposure are exquisitely sensitive to apoptosis, presumably due to increased intracellular ROS production. Significantly, DMSO treatment does not trigger apoptosis in this system, and accordingly frataxin deficiency does not seem to affect cardiomyocyte differentiation. Our in vitro observations show a parallelism with the human disease. In FRDA patients, early loss of large neurons in the DRG seems to occur during development (3,26–28), while cardiomyocyte involvement occurs later in life (29).
Disruption of the gene encoding frataxin in mice results in intrauterine death at E6.5, shortly after implantation, further demonstrating an important role for frataxin during early development (17). What important events happen around that period that might explain the lethality of frataxin deficiency? During early embryogenesis at E4.0, the inner cell mass of the mouse blastocyst consists of a core of embryonic ectoderm cells surrounded by an outer layer of primitive (extra embryonic) endoderm. These layers will subsequently give rise to both visceral endoderm and parietal endoderm. Shortly after blastocyst implantation (E5.0) the solid mass of ectoderm cells is converted by a process known as cavitation into a columnar epithelium surrounding a cavity. By E6.0 this process is completed. Substantial evidence indicates that cavitation is the result of both programmed cell death and selective cell survival, and that the process depends on signals from visceral endoderm (32,33). This scenario will fit with our prediction that frataxin deficiency renders cells more susceptible to apoptotic stimuli. In the complete absence of frataxin, excessive apoptosis occurring at the time of implantation of the embryo ultimately leads to its annihilation. A reduced level of the protein allows survival beyond this stage, as observed in FRDA patients, but enhanced programmed cell death may occur later on in vulnerable cells, as DRG sensory neurons.
What is the mechanism linking a deficit in frataxin to enhanced sensitivity to some apoptosis-inducing stimuli? We provide evidence that frataxin-deficient cells generate more ROS during RA-induced neuronal differentiation. While Bcl-2 expression, an anti-apoptotic protein during P19 neuronal differentiation (20), increased to comparable levels in all clones during differentiation, MnSOD rose to significantly higher levels in frataxin-deficient cultures compared with control cultures. These cells may thus behave differently from the reported yeast model (34), where frataxin deficiency was found to be associated with a lack of induction of both SODs and a lack of anti-oxidant response (34). This difference in MnSOD induction is likely be directly related to the neuronal differentiation process taking place in our cell model, a feature not comparable to the yeast model. MnSOD prevents superoxide accumulation by its conversion to hydrogen peroxide (35) and has an anti-apoptotic effect in neural cells (24). It is increasingly clear that ROS, including those generated in mitochondria, participate in intracellular signaling pathways (25,36). In some situations, particularly during development, a decision between survival and apoptosis may depend on pathways that are affected by ROS levels (33). RA treatment of P19 cells appears to be one example, with increased ROS production significantly tipping the balance towards apoptosis. In the case of the human disease, DRG neurons express the Trk family of neurotrophin tyrosine kinase receptors and undergo programmed cell death in a manner regulated by limiting neurotrophin levels. Neurotrophins inhibit ROS production as an acute effect of their binding to receptor tyrosine kinases, probably through the MAPK pathway (37). It is possible that ROS produced in mitochondria as a consequence of frataxin deficiency diffuse in the cytosol and dampen this effect, rendering cells less responsive to neurotrophins and therefore more vulnerable to programmed cell death.
In summary, our results suggest that frataxin is essential for the development of the neural lineages and that reduced expression of frataxin increases susceptibility to apoptosis of differentiating neuroectodermal cells. These results raise the possibility that the early lethality of frataxin knockout mice and large DRG neuronal loss in FRDA may be due, at least in part, to higher sensitivity to apoptosis of frataxin-deficient embryonic cells. A general point suggested by our observations is that the deleterious effects of increased ROS production resulting from frataxin deficiency should not only be viewed in terms of direct damage to cellular components, but also as a consequence of interference with signaling pathways that are critical for the survival of specific cell types.
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MATERIALS AND METHODS |
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Cell culture and differentiation
The P19 clone was obtained from the American Type Culture Collection (Rockville, MD). Cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS) in a 5% CO2 humidified chamber. To promote cellular differentiation, cells were cultured as aggregates in bacterial Petri dishes in media supplemented with 1 µM RA. Cells (1 x 105/ml culture medium) were plated into bacterial Petri dishes (day 0) and incubated with RA for 2 days. After 2 days, floating aggregates were collected and washed with DMEM and re-plated onto bacterial grade Petri dishes and incubated with RA for an additional 2 days. The aggregates were harvested, dissociated by treatment with phosphate-buffered saline (PBS) containing 1 mM EDTA, and then transferred to tissue culture plates and cultured in medium lacking RA for 4 days.
For quantification of cardiac differentiation, the ‘hanging drops’ method was used (38). Briefly, 20 µl drops of cells were pipetted onto the roof of a bacterial dish, which was then inverted over PBS (to maintain humidity). On day 4, the drops were collected, the medium was replaced, and cells were replated on tissue culture plates in fresh medium lacking DMSO. On day 8, the percentage of aggregates per dish containing a region of any size which was beating were scored. Triplicate samples were counted in each experiment, and in three separate experiments.
Plasmids and stable transfection
Plasmids harboring the antisense sequence from nucleotides 347 to 1 (pMP257) or sense sequence from nucleotides 1 to 1033 (pMP254) derived from the cDNA sequence of mouse frataxin were engineered into the pCI-neo mammalian expression vector (Promega Corporation) using standard recombinant DNA techniques. The P19 cells were transfected with these plasmids using LipofectAMINETM (Gibco BRL, Life Technologies, Inc.) following the manufacturer’s protocol. Positive transfectants harboring the neo resistance gene were selected and cloned in the presence of G418 (400 mg/ml, Gibco BRL). Stable transfectants were maintained in media containing 200 mg/ml G418. The ability of the antisense RNA to suppress the expression of frataxin in the stable transfectants was determined by immunoblotting with antibody specific for frataxin and by RNase protection assay.
Immunocytochemistry
After RA treatment and culture as aggregates, the cells were dissociated and re-plated onto cover glasses (Fisher Scientific). Cover glasses were rinsed with PBS prior to fixation with 4% paraformaldehyde for 15 min. After fixation, the cover glasses were rinsed with PBS and cells were permeabilized with a solution containing 0.1% TritonX-100 in 0.1% sodium citrate for 2 min on ice. As a neuronal marker, a mononuclear antibody to MAP2 (Boehringer Mannheim) was used. Anti-frataxin monoclonal antibody 1G2 was kindly provided by Dr M.Koenig [Institut de Genetique et de Biologie Moleculaire et Cellulaire (IGBMC), Strasbourg]. The fixed and permeabilized cells were incubated with primary antibodies for 1 h at 37°C and then washed three times with PBS. Biotinylated secondary antibody was added and cover glasses were incubated for 45 min at 37°C. The Vectastain ABC kit was used to detect the biotinylated antibodies (Vector Laboratories). For Hu and nestin staining, the aggregates were harvested at days 3 and 4, respectively, dissociated by treatment with collagenase/dispase (Boehringer Mannheim), counted, and 104 cells were then fixed with acid ethanol (95% ethanol/5% acetic acid) for 30 min at room temperature. The fixed cells were washed with PBS, incubated with a mAb to Hu (Molecular Probes) or to nestin (1:20, Developmental Studies Hybridoma Bank, Rat-401). After overnight incubation at 4°C, the samples were further washed and incubated with anti-mouse Igs conjugated to fluorescein isothiocyanate (FITC) or to rhodamine (Boehringer Mannheim). Stained preparations were resuspended in the same volume and examined under epifluorescence microscopy.
Western blot analysis
Cell lysis was achieved by sonication, followed by centrifugation at 5000 g for 10 min. The supernatant was collected and protein determined with the Protein Assay Kit (Bio-Rad). Protein (50 µg for each sample) was resolved on a 12% SDS-polyacrylamide gel and transferred onto nitrocellulose membrane. The following primary antibodies were employed: anti-frataxin monoclonal antibody 1G2, kindly provided by Dr M.Koenig; anti-ß-tubulin and rabbit anti-bcl-2 (Ab-2) (Calbiochem); rabbit anti-MnSOD (Research Diagnostics, Inc.); and anti-cytochrome-c (PharMigen). For the detection of the immunocomplexes formed, the secondary antibody peroxidase-conjugated goat anti-mouse IgG (Jackson Immunoresearch Laboratories, Inc.) or goat anti-rabbit IgG (Calbiochem) was used. Staining intensity was developed with the chemiluminescence system (RENAISSANCETM, NEN Life Science Products).
Measurement of ROS generation
ROS was assessed by the dye DCF-1 diacetate (DCF-DA; Molecular Probes, Inc.). DCF-DA is a non-fluorescent cell-permeant compound. Once inside the cell, it is cleaved by endogenous esterases and thereby trapped within the cell. The de-esterified product becomes fluorescent compound DCF on oxidation by ROS. Cells were loaded with 100 µM DCF for 30 min at 37°C, washed in PBS, and the fluorescence intensity measured flow cytometry analysis (FACS) in a Coulter Epics Elite (Coulter).
Apoptosis assay
Apoptotic cells were determined by three different methods. Microscopic observation after Hoechst 33342 staining (Molecular Probes) was employed to detect apoptotic morphology. The TUNEL assay for the detection of DNA fragmentation by FACS was used according to the manufacturer’s instructions (In Situ Cell Death detection kit, fluorescein, Boehringer Mannheim). DNA fragmentation was quantified after [14C]thymidine incorporation followed by cell lysis and elution. Radioactivity in the eluates and in the filters was determined in a scintillation counter.
Cell viability
Cell viability was assessed either by trypan blue exclusion or by vital dye reduction. 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) (Sigma) was used for the quantification of living metabolically active cells (39). Mitochondrial dehydrogenases metabolize MTT to blue formazan dye that can be detected spectrophotometrically. Cells (2.0–2.5 x 104) were plated onto 24-well plates and incubated in culture medium for 48 h. The medium was then removed and replaced by DMEM without phenol red or FBS and MTT solution (2 mg/ml phosphate buffer) for 4 h. After removing the DMEM and MTT solutions the remaining formazan blue crystals were dissolved in a 0.04 N HCl-isopropyl alcohol mixture. The absorbance of the reaction solution at 570 nm was recorded. The absorbance at 630 nm was used as a reference. The net A570–A630 was taken as the index of cell viability.
Proliferation assays
A total of 2 x 104 cells/well were cultured in triplicate in 96-well round-bottomed plates with or without 1 µM RA. Incorporation of [3H]thymidine was measured after 24, 48 and 72 h in culture. During the last 14 h of culture, cells were pulsed with 1 µCi of [3H]thymidine and cells were harvested on GF/A filters. Radioactivity was measured using a scintillation counter.
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ACKNOWLEDGEMENTS |
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We thank Laura Montermini, Patricia Maciel, Nathalie Droin and Xiaochun Wan for technical tips. mAbs to nestin and Troma-1 were obtained from the Developmental Studies Hybridoma Bank, maintained by the University of Iowa, Department of Biological Science. This work was supported by grants from the National Institute of Neurological Diseases and Stroke (NINDS, grant no. NS34192), the Medical Research Council of Canada (MRC, grant no. FRN14689) and the Muscular Dystrophy Association (MDA), USA. M.P. is supported by an MRC scientist award. M.M.S. is supported by a scholarship from Fundação para a Ciência e a Tecnologia – PRAXIS XXI (BPD/18833/98).
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FOOTNOTES |
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+ To whom correspondence should be addressed at: Service de Neurologie, Hôpital Erasme, Route de Lennik 808, 1070 Bruxelles, Belgium. Tel: +32 2 555 33 46; Fax: +32 2 555 39 42; Email: massimo.pandolfo@ulb.ac.bePresent address:Keiichi Ohshima, Growth Factor Division, National Cancer Center Research Institute, Tokyo 104-0045, Japan
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