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Human Molecular Genetics Advance Access originally published online on October 3, 2005
Human Molecular Genetics 2005 14(22):3397-3405; doi:10.1093/hmg/ddi367
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© The Author 2005. Published by Oxford University Press. All rights reserved.
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RNAi-mediated suppression of the mitochondrial iron chaperone, frataxin, in Drosophila

Peter R. Anderson1, Kim Kirby1, Arthur J. Hilliker2 and John P. Phillips1,*

1Department of Molecular and Cellular Biology, University of Guelph, Guelph, Ontario, Canada N1G 2W1 and 2Department of Biology, York University, Toronto, Ontario, Canada M3J 1P3

* To whom correspondence should be addressed. Tel: +1 5198244120 ext. 52796; Fax: +1 5198372075; Email: jphillip{at}uoguelph.ca

Received August 4, 2005; Accepted September 27, 2005


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
The mitochondrial iron chaperone, frataxin, plays a critical role in cellular iron homeostasis and the synthesis and regeneration of Fe–S centers. Genetic insufficiency for frataxin is associated with Friedreich's Ataxia in humans and confers loss of function of Fe-containing proteins including components of the respiratory chain and mitochondrial and cytosolic aconitases. Here, we report the use of RNA-interference (RNAi) to suppress frataxin in the multicellular eukaryote, Drosophila. Phenotypically, suppression of the Drosophila frataxin homologue (dfh) confers distinct phenotypes in larvae and adults, leading to giant long-lived larvae and to conditional short-lived adults. Deficiency of the DFH protein results in diminished activities of numerous heme- and iron–sulfur-containing enzymes, loss of intracellular iron homeostasis and increased susceptibility to iron toxicity. In parallel with the differential larval and adult phenotypes, our results indicate that dfh silencing differentially dysregulates ferritin expression in adults but not in larvae. Moreover, silencing of dfh in the peripheral nervous system, a specific focus of Friedreich's pathology, permits normal larval development but imposes a marked reduction in adult lifespan. In contrast, dfh silencing in motorneurons has no deleterious effect in either larvae or adults. Finally, overexpression of Sod1, Sod2 or Cat does not suppress the failure of DFH-deficient animals to successfully complete eclosion, suggesting a minimal role of oxidative stress in this phenotype. The robust developmental, biochemical and tissue-specific phenotypes conferred by DFH deficiency in Drosophila provide a platform for identifying genetic, nutritional and environmental factors, which ameliorate the symptoms arising from frataxin deficiency.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Friedreich's Ataxia (FRDA) is a neurodegenerative disorder that arises from a deficit of the mitochondrial iron chaperone, frataxin (1Go–3Go). Frataxin deficiency is associated with the expansion of the GAA triplet repeat in intron I of the human FRDA gene (4Go). The progressive loss of coordination of limb movements, dysarthria, nystagmus, scoliosis, and diabetes characteristic of FRDA usually manifest before adolescence. The peripheral nervous system (PNS) and the heart are among the most severely affected tissues, and more than half of those afflicted eventually succumb to cardiac related complications (4Go–6Go).

Current understanding of the molecular origins of FRDA pathogenesis and disease progression has been aided by the investigations of yeast and animal models of the disorder. Characterization of yeast mutants lacking the yeast frataxin homolog (Yfh) (7Go) has revealed important roles for frataxin in mitochondrial iron storage (7Go–15Go), regulation of intracellular iron levels (7Go,8Go,10Go,16Go), biogenesis of iron–sulfur (Fe–S) clusters (16Go–20Go) and heme (14Go,21Go), reactivation of the labile Fe–S cluster of mitochondrial aconitase (22Go) and in the prevention of deleterious free radical production (12Go,15Go,23Go). Murine models with tissue-specific frataxin deficiency recapitulate important biochemical and whole organism phenotypes that typify FRDA (24Go) including cardiac hypertrophy, large sensory neuron dysfunction and marked reduction of the activities of several Fe–S cluster enzymes (aconitase and complexes I–III of the respiratory chain). These model systems have given valuable insight into the biochemistry of frataxin and its role in the molecular underpinnings of FRDA.

Unfortunately, there is no cure for this debilitating disease and important questions, such as the origin of the wide variation in the manifestation of FRDA symptoms (5Go) and the contribution of oxidative stress to FRDA pathology, still remain. Development of a Drosophila model of FRDA has the potential to accelerate progress in identifying genetic, nutritional and environmental factors that ameliorate the phenotypic outcomes associated with frataxin deficiency. The identification of a gene encoding a Drosophila frataxin homolog (dfh) (25Go) set the stage for the development of such a model. Here, we describe the use of RNA-interference (RNAi) methodology to impose widespread and tissue-specific downregulation of dfh in Drosophila, and we report the molecular and phenotypic consequences of dfh deficiency in this model system.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
DFH levels vary widely throughout development
DFH is predicted from genomic sequencing to consist of 190 amino acids including an N-terminal mitochondrial transit peptide (25Go). Using a polyclonal antibody directed against DFH peptides (see Materials and Methods), we determined that DFH is of low abundance overall in Drosophila, reaching its highest level in late embryos, diminishing in larvae and puparia, then increasing in pharate adults to reach modest levels in young adults (Fig. 1). Because high levels of DFH are detected in early embryos, we used males to assay DFH levels in adults to avoid the possible contribution of embryo-derived DFH in females. The double band detected in embryo and adult extracts (Figs 1 and 2) may correspond to processed and unprocessed forms of DFH, in agreement with findings in mammals and yeast (26Go,27Go). The two bands correspond in size to those predicted for the larger unprocessed precursor (~21 kDa) and the smaller mature form (~15 kDa) (25Go). Failure to detect the smaller form in flies overexpressing dfh may arise from the reduced amount of extract loaded (1/10) to compensate for an overly heavy DFH signal. Overexpression of yfh in yeast was reported to yield a very high ratio of processed to unprocessed YFH (22Go). Moreover, expression of DfhIR transgenes suppresses both forms, confirming their identity as DFH (Fig. 2).



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  		Figure 1. Steady state levels of DFH vary widely throughout development. Extracts were electrophoresed through polyacrylamide gels, transferred to nitrocellulose and probed with anti-DFH antibody. Seventy-five microgram of total protein was applied to each lane except dfh/+ which contained only 7.5 µg protein to reduce the total signal. dfh/+, adult males over-expressing DFH (UAS-Dfh/+; daG32Gal4/+); Embryos, 0–2, 2–6 and 6–24 h; Larvae, first, second and third instar; LP, light pupae; PA, pharate adults; adult males, 1–2 days old.

 


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  		Figure 2. DfhIR mediates the suppression of DFH through dfh transcript degradation. (A) DfhIR promotes degradation of dfh mRNA. Total RNA (10 µg) isolated from wandering third instar larvae of different genotypes was electrophoresed, transferred to nylon membranes and hybridized with digoxigenin-labeled probes for dfh and the ubiquitously expressed marker, Rp49. Genotypes are +/+;daG32Gal4/+ (lane1), +/+;UDIR1/daG32Gal4 (lane 2), UDIR2/+;daG32Gal4/+ (lane 3). (B) Extracts from adult males were electrophoresed, transferred to nitrocellulose membranes and probed for DFH as described in Figure 1. Nitrocellulose membranes were probed with anti-Actin antibodies as a loading control. Approximate molecular weights are indicated.

 
RNAi suppresses DFH in larvae and adults
The single-copy dfh gene contains a single intron and comprises ~1 kb of the gene-rich 8C/D region of the Drosophila melanogaster X-chromosome (FlyBase: fh). No mutations of dfh have been reported, although several expansive X-chromosome deletions that encompass the dfh locus (and several other genes) are available. To investigate the biological consequences of DFH deficiency, we developed an inverted-repeat, GAL4 regulated transgene (UAS-DfhIR; see Materials and Methods for details). This approach circumvents the lack of available genomic mutants, allows us to mimic FRDA by reducing rather than completely ablating DFH and avoids the possibility of early developmental lethality (28Go). To obtain widespread dfh silencing, transgenic lines carrying UAS-DfhIR transgenes were crossed to the daG32GAL4 driver line. This driver expresses Gal4 widely throughout development and in most, if not all, tissues under the control of regulatory sequences from the class I helix-loop-helix transcription factor, daughterless (da) (29Go,30Go). Expression of DfhIR results in the degradation of dfh transcripts (Fig. 2A) and DFH protein is reduced to undetectable or near undetectable levels in both adults (Fig. 2B) and larvae (data not shown).

Dfh silencing in Drosophila produces giant, long-lived larvae and moribund, short-lived adults
Dfh suppression causes profound and differential effects in larvae and adults. Dfh-suppressed larvae exhibit retarded development and reduced viability (Fig. 3A). By the time, DfhIR larvae are just beginning to initiate pupariation, control strains have completed metamorphosis and eclosed as adults. DfhIR larvae subsequently endure an extended larval phase, during which they continue to forage and grow. This extended larval phase can last as long as 45 days, allowing some individuals become a full 2-fold larger (Fig. 3B).



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  		Figure 3. Dfh suppression retards larval development. (A) First instar larvae hatching within a 3 h window were transferred into vials and scored for pupariation initially twice daily and later once daily at 25°C. The vertical bar indicates the point when all control flies had eclosed. G32: daG32Gal4 driver. (B) Photographs of control and DfhIR wandering third instar larvae. Genotypes are +/+;daG32Gal4/+, and +/+;UDIR1/daG32Gal4. Genotypes are +/+;daG32Gal4/+ (A), +/+;UDIR1/daG32Gal4 (B), UDIR2/+;daG32Gal4/+.

 
DfhIR larvae from UDIR1 and UDIR2 lines described herein are not capable of becoming viable adults at 25°C, though lines exhibiting reduced levels of DFH diminution were occasionally able to produced occasional (<0.5%) adult flies. The failure of DfhIR larvae to become adults at 25°C can be partially abrogated by culture at 18°C beginning at pupariation. Under this condition, 1–2% of DfhIR pupae develop into adults. For reasons not understood, female escapers are five times more prevalent than males (the control strains show the expected 1:1 ratio). The survival of these escaping DfhIR adults is short but biphasic (Fig. 4). By day 3, survival of DfhIR females is reduced to 40–60%, whereas it takes 47 days for the control strain adults to reach 50% survival. Following the initial high-mortality phase during which 70–90% of the population is lost, the remaining cohort of survivors persists with relatively low mortality for up to 40 more days. Experiments performed with fewer numbers of male flies yielded nearly identical survival curves.



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  		Figure 4. Dfh suppression causes early mortality in young adults. Survival at 25°C of 200 adult females of each genotype on standard cornmeal food (20 flies/vial) was followed with enumeration and transfer of survivors to fresh vials every 2–3 days. G32: daG32Gal4 driver.

 
Mitochondrial Fe–S enzymes are impaired by Dfh silencing in both larvae and adults
Silencing of dfh in Drosophila larvae leads to reduced activities of [4Fe–4S]-containing mitochondrial aconitase and of respiratory complexes II, III and IV by 40–60% (Fig. 5). Enzymes like aconitase with a solvent exposed iron–sulfur (Fe–S) cluster are vulnerable to reversible inactivation by superoxide (31Go) and show reduced activities in SOD-deficient flies (32Go). Therefore, we verified that the reduced activities arising from DFH depletion do not result from loss of SOD1 and SOD2 activities (Fig. 6D). Furthermore, mitochondrial aconitase activity lost by DFH depletion could not be restored by in vitro reactivation of its Fe–S center (data not shown) as is the case in SOD2 deficiency (33Go). Overall, these results suggest that DFH is required for Fe–S recycling in Drosophila. We were unable to detect complex I activity in larval mitochondria. However, complex I activity was readily detected in adults and showed a marked reduction in specific activity (37±4%, P<0.001) in response to DFH depletion. Overall, the DfhIR-mediated loss of mitochondrial aconitase and respiratory complex II, III and IV activities in larvae are mirrored in DfhIR adults (data not shown). To determine whether the effects of DFH depletion are confined to iron-dependent enzyme function, we assayed the activity of the mitochondrial iron-independent enzyme, citrate synthase. As shown in Figure 5, the activity of this iron-independent enzyme is reduced by 10–20%. Because the level of mitochondrial aconitase protein is unaffected by DFH depletion (Fig. 5), suggesting that specific protein import is not compromised, these data suggest that DFH depletion exerts a general deleterious effect on mitochondria that extends beyond its specific role in Fe–S center biogenesis.



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  		Figure 5. Dfh suppression leads to reduced activity of Fe–S enzymes in the mitochondrial compartment. Mitochondria were isolated from larvae of the following genotypes: +/+;daG32Gal4/+ (A), +/+;UDIR1/daG32Gal4 (B) and UDIR2/+;daG32Gal4/+ (C). Fe–S activities assayed: aconitase, electron transport complexes II, III and IV; citrate synthase (CS) were assayed as an Fe–S independent control. Values are expressed as a percentage of control (+/+;daG32Gal4/+) specific activity. Data points represent the mean±SD of at least nine independent determinations. Asterisk, double asterisk and triple asterisk indicate statistically significant differences of P<0.05, P<0.01 and P<0.001, respectively. The inset shows an immunoblot of mitochondrial proteins (10 µg) from larvae of the three genotypes probed with antibody raised against Drosophila mitochondrial aconitase.

 


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  		Figure 6. Dfh suppression affects aconitase and FerHCH differentially in larvae and adults. (A) Enzymatic activities of mitochondrial (M-Acon) and cytoplasmic (C-Acon) aconitase following electrophoretic separation on cellulose membrane. Genotypes are +/+;daG32Gal4/+ (lane 1), UDIR2/+;daG32Gal4/+ (lane 2). (B) Immunoblot analysis of mitochondrial aconitase and FerHCH in DfhIR larvae and adults. Protein (10 µg) from third instar larvae or adult males was electrophoresed, transferred to nitrocellulose membranes and probed with antibody raised against Drosophila mitochondrial aconitase and anti-Drosophila FerHCH antibody, respectively. Membranes were re-probed with anti-Actin antibody as a loading control. (C) Northern blot analysis of FerHCH transcripts in DfhIR larvae and adults. (D) SOD1 and SOD2 activities in DfhIR larvae and adults. Extracts from third instar larvae or young adult males were electrophoresed on native polyacrylamide gels, and SOD activity was determined by interference with superoxide-dependent reduction of nitroblue tetrazolium.

 
Cytosolic Fe–S proteins are differentially affected by dfh silencing in larvae and adults
To determine the effects of dfh silencing on Fe-dependent proteins in the cytoplasmic compartment, we determined cytoplasmic aconitase activity and Ferritin Heavy Chain Homolog (FerHCH) protein levels. Iron regulatory protein 1 (IRP1) is a dual-function cytosolic Fe–S protein. The holoenzyme functions as a cytosolic aconitase, whereas the apoprotein serves as an RNA-binding translational regulator. We had earlier established that superoxide-mediated disassembly of the exposed [4Fe–4S]-center in the cytosolic aconitase of Drosophila facilitates its functional transition from an aconitase to IRP1 (32Go). Several proteins involved in the transport, storage or utilization of iron are encoded by mRNAs containing iron response elements (IREs) to which IRP1 can bind. For example, binding of IRP1 to the 5' IRE of ferritin heavy chain mRNA decreases its rate of translation (34Go). To determine the impact of DFH depletion on iron homeostasis in the cytoplasmic compartment, we assayed cytoplasmic aconitase activity and Drosophila FerHCH protein levels and found that both are reduced in DfhIR adults (Fig. 6A and B). By quantitative digitometry, cytoplasmic aconitase and FerHCH in DfhIR adults are reduced to 39 and 36%, respectively, of control values. Interestingly, both cytoplasmic aconitase activity and FerHCH remain relatively unchanged in DfhIR third instar larvae, even though reduction of Fe–S enzyme activities in the mitochondrial compartment is equivalent to those seen in adults (Figs 5 and 6). Furthermore, the relative abundance of FerHCH transcripts in both larvae and adults does not decrease in response to Dfh silencing (Fig. 6C). Thus, it is likely that the loss of FerHCH in DfhIR adults arises from enhanced conversion of c-aconitase to IRP1, which in turn binds to and suppresses the translation of FerHCH transcripts. Why this does not happen in larvae is unknown.

Dfh silencing confers hypersensitivity to iron
The proposed role of DFH as an iron chaperone and storage protein would predict that in its absence, tolerance to elevated dietary iron would be reduced. As shown in Figure 7, DfhIR larvae exhibit striking sensitivity to elevated iron concentration and succumb as first larval instars in standard cornmeal food supplemented with 12 mM FeCl3. At his level of supplemental iron, 78% of control animals successfully form pupae. These data are in accord with the role of DFH as an iron chaperone and storage protein.



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  		Figure 7. Dfh suppression increases susceptibility to iron toxicity. First instar larvae were transferred into vials containing cornmeal food supplemented with iron and scored daily for pupariation at 25°C. Genotypes are +/+;daG32Gal4/+ (A), +/+;UDIR1/daG32Gal4 (B) and UDIR2/+;daG32Gal4/+ (C).

 
Differential neuronal silencing of dfh permits normal larval development but reduces adult lifespan
In humans, frataxin exhibits tissue-specific levels of expression (1Go) and high expression partially correlates with those tissues principally involved in the disease (5Go). Two principal focal tissues include the PNS and the heart. Accordingly, we investigated the consequences of silencing of dfh in the PNS by using the C96-Gal4 driver line (35Go), to drive the expression of DfhIR in the PNS. As a comparison, we also used the D42-Gal4 driver (36Go) to express DfhIR in motorneurons. In striking contrast to the outcome of widespread DfhIR expression conferred by the daG32-Gal4 driver (Fig. 3), DfhIR expression in the motorneurons or in the PNS has no detectable effect on pre-adult development. However, eclosing PNS-DfhIR adults suffer a 40% reduction in adult lifespan (Fig. 8), whereas MN-DfhIR adults are unaffected.



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  		Figure 8. Selective dfh suppression in the PNS spares larvae but leads to early adult mortality. Survival at 25°C of 200 adult males of each genotype on standard cornmeal food (20 flies/vial) was followed with enumeration and transfer of survivors to fresh vials every 2–3 days. C96: PNS Gal4 driver; D42: motorneuron Gal4 driver.

 
Supplemental expression of Sod1, Sod2 or Cat does not rescue the eclosion block conferred by DfhIR
Interference with iron homeostasis raises the risk of damaging iron-catalyzed oxidative damage. We tested the possibility that such oxidative damage contributes to the deleterious effects of DFH deficiency. We generated compound transgenic strains over-expressing Sod1, Sod2 or Cat in concert with dfh suppression and assayed the impact on the capacity of DfhIR animals to escape pre-adult mortality. Crosses generating flies hemizygous for the daG32Gal4 driver and UDIR1 or UDIR2, and either UAS-Sod1, UAS-Sod2 or UAS-Cat, yielded no adult progeny, whereas control crosses generating flies hemizygous for the daG32Gal4 driver and Sod or Cat transgenes in the absence of UDIR1 or UDIR2 yielded an average of 210±80 adult flies with no significant difference observed between the various control crosses (data not shown). Flies hemizygous for both the daG32Gal4 driver and UAS-Sod1, UAS-Sod2 or UAS-Cat exhibited increased enzymatic activities of 203±10, 254±19 and 496±50% of control flies (+/+; daG32Gal4/+), respectively. Because the enzymatic reactive oxygen scavenging system is not compromised in DfhIR larvae (Fig. 6), and because overexpression of Sod1, Sod2 or Cat does not improve the capacity of DfhIR larvae to develop into viable adults, we conclude that reactive oxygen is not an important contributor to the deliterious phenotype displayed by DFH-deficient larvae.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
Systemic suppression of dfh in Drosophila elicits the primary biochemical signature of FRDA, namely, marked impairment of the activities of iron cofactor-dependent enzymes and loss of intracellular iron homeostasis. Comparatively, the Drosophila response is similar to that seen in yfh-null yeast (16Go–18Go,37Go) and in Frataxin-deficient mouse models of FRDA (24Go). Our data on Drosophila, an insect invertebrate, are thus consistent with the functionally conserved role of DFH as an iron chaperone in the synthesis and homeostatic maintenance of Fe–S clusters (14Go,19Go–21Go).

There are no known mutations of the dfh gene. Therefore, we developed a dfh inverted-repeat transgene to bypass the lack of traditional genetic mutants and to reduce, rather than completely eliminate, DFH. In this way, we sought to (a) more accurately mimic the underpinning genetic origin of FRDA, which arises from diminution rather than complete loss of frataxin and (b) to avoid the potential early embryonic lethality reported in frataxin-null mice (28Go). We had confidence in the utility and the accuracy of inverted-repeat RNAi transgene methodology, because we had earlier used this approach to successfully suppress the expression of two other genes, Sod1 and Sod2. RNAi targeted to the Sod1 gene produced an accurate phenotype of pre-existing Sod1-null mutants (32Go). In the second instance, RNAi-mediated suppression of Sod2 (33Go) generates a spectrum of phenotypes which accurately predicted the phenotype of Sod2-null mutants that were subsequently produced by Duttaroy et al. (38Go).

The phenotypes arising from broadly expressed DfhIR transgenes include larvae with reduced viability, protracted development and failure to initiate and complete metamorphosis. We note with interest that prolonged larval development leading to high larval mortality is a feature of Drosophila mutants defective in mitochondrial DNA maintenance (39Go–41Go). This prompts the testable hypothesis that DFH depletion leads to increased mitochondrial DNA damage. Such damage would add to the burden of mitochondrial dysfunction initiated by DFH depletion. Such mitochondrial DNA damage could partially account for the loss of mitochondrial enzyme activity observed in DfhIR mitochondria (Figs 5 and 6); however, the loss of mitochondrial aconitase activity (a nuclear encoded protein imported into the mitochondria) would suggest that if this is the case, it is secondary to a loss of capacity for Fe–S center biogenesis.

We previously showed that conversion of cytoplasmic aconitase to IRP1 in Drosophila can be enhanced by the ablation of SOD1 which increases the availability of superoxide (32Go). Here, we show that this conversion leads to the depression of FerHCH as would be predicted by enhanced binding of IRP1 to the 5' IRE of the FerHCH transcript (42Go). Similar observations have been made in HeLa cells treated with RNAi to deplete frataxin (43Go). Interestingly, no such changes in cytoplasmic aconitase activity or in coupled FerHCH changes are manifested in larvae in response to dfh depletion (Fig. 6). Drosophila FerHCH is known to generate multiple transcripts that vary in relative abundance between larvae and adults, and the balance of these transcripts in specific tissues can shift towards those lacking an IRE in response to iron availability (44Go). In addition as shown here, larvae and adults exhibit markedly different ratios of mitochondrial to cytosolic aconitase activity (Fig. 6). This suggests that the differential phenotypic responses of larvae and adults to DFH depletion reflect distinctly different requirements and utilization strategies for iron. This larval/adult dichotomy exemplified by the Drosophila model of FRDA could provide a unique opportunity for investigating aspects of iron metabolism that may be relevant to FRDA. It should be noted that the half-life of DFH has yet to be determined and as such we cannot rule out the possibility that a small amount of maternal DFH persists throughout the larval phase. Although if this is the case, we have been unable to detect this protein using our immunoblotting assay.

A small percentage of DfhIR larvae can form viable adults if incubated at 18°C as pupae. These adults exhibit high-initial mortality with the majority dying within 3–4 days (Fig. 4). However, following this initial phase of high mortality, the remaining cohort of survivors persists with low mortality for up to 40 more days. If this late-phase survival plateau represents a true adaptive response of adults to DFH depletion, it could have important therapeutic implications for FRDA. Although we cannot, at this point, rule out the possibility that these lingering individuals arise from variation in the extent of RNAi-mediated knock between individual animals, it is intriguing to note that a similar phenomenon is observed in yfh-null yeast strains that exhibit variability and instability of the phenotypes associated with Frataxin deficiency (21Go).

Because one of the principal foci of FRDA is the PNS (45Go), we expressed DfhIR in the PNS of Drosophila. This led to two important outcomes. First, it allows larvae to bypass the deleterious effects conferred by ubiquitous dfh suppression. Secondly, the adults arising from these larvae exhibit a markedly reduced lifespan (Fig. 8). Thus, the PNS of adults is sensitive to dfh suppression. Moreover, the vulnerability of the adult PNS is not apparently shared by the motorneuron component of the central nervous system, because DfhIR expression in motorneurons has no apparent affect. The D42-Gal4 driver is known to promote robust expression of UAS-linked transgenes and has been used previously in this laboratory to rescue the lifespan of Sod1-null mutants and to extend lifespan of Sod1+ flies by ectopic expression of Sod1 in the motorneurons (36Go). It is therefore unlikely that the lack of effect of DfhIR expression in motorneurons could be attributed to weak expression of the D42-GAL4 activator in that tissue. Considering that degeneration of the PNS is a principal feature of FRDA (45Go) and that motorneurons appear unaffected (46Go), our results suggest that the vulnerability of the PNS to Frataxin depletion is conserved across a wide spectrum of animal taxa and provide further validation for the Drosophila model of FRDA.

The disruption of iron homeostasis resulting from dfh suppression might be expected to lead to the generation of deleterious reactive oxygen species via iron-catalyzed Fenton chemistry. However, transgenic augmentation of the primary reactive oxygen metabolizing enzymes SOD1, SOD2 or CAT provides no measurable improvement in viability. Because ectopic overexpression of these enzymes has been shown to effectively suppress the deleterious effects of endogenous and applied oxidative stress (47Go–49Go), this strongly suggests that oxidative stress in larvae or pupae is not a major contributor to the eclosion block arising from DFH depletion. Further experiments will be required to determine whether ectopic overexpression of these enzymes can rescue the shortened lifespan conferred by ubiquitous or PNS-targeted DFH depletion in adults. The possible contribution of thiol-based antioxidants like thioredoxin and glutathione also needs to be determined to affirm this conclusion.

There are currently no cures for FRDA. The wide variation in the symptoms and age of onset (5Go) suggests that factors other than the degree of triplet repeat expansion can influence disease progression. The robust phenotypes arising from DFH deficiency in Drosophila set the stage for exploiting the genetics of this model organism to identify genetic, nutritional and environmental factors that can alleviate these phenotypes and which could lead in turn to the development of novel treatment strategies for reducing the burden of FRDA.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
Drosophila stocks and culture methods
Drosophila stocks were maintained at 25°C on standard cornmeal agar medium unless otherwise stated. CO2 anesthesia was used, allowing at least 5 h of recovery at room temperature before commencement of experimentation (50Go). The daG32Gal4, the D42-Gal4 and C96-Gal4 drivers were originally obtained from Boulianne (University of Toronto, Toronto). The daG32Gal4 driver expresses GAL4 widely throughout development and in most, if not all tissues under the control of regulatory sequences from the class I helix-loop-helix transcription factor, da (29Go,30Go). The D42-Gal4 driver expresses GAL4 broadly during embryogenesis but becomes restricted to motorneurons and interneurons within the larval nervous system, and with the exception of a few unidentified neurons within the central brain, it is restricted to motorneurons in the adult nervous system (36Go). The C96-Gal4 driver expresses GAL4 strongly in neurons of the adult peripheral nervous system with weak expression observed in the larval salivary gland and in the margin of the wing imaginal disc in larvae (35Go) (unpublished data).

Construction of UAS-transgenes
For overexpression studies, D. melanogaster cDNAs for Sod1 and dfh were isolated from a Canton S 2-week-old male and female cDNA library (Stratagene, La Jolla, CA, USA) and inserted into pBluescriptSK (±) (Stratagene) before subcloning into the pP[UAST] transformation vector (51Go) to generate UAS-Sod1 and UAS-Dfh transgenes. The UAS-Cat transgene was generated similarly by inserting the Cat cDNA (a kind gift from B. Orr) into p[PUAST]. For RNAi-mediated silencing of dfh, inverted repeats containing the first 391 nucleotides of the dfh coding region were subcloned into pP[UAST] to generate the UAS-DfhIR transgene.

Generation of transformants
Transgenic flies were generated by standard embryo injection protocols (52Go). The UAS-Sod1, UAS-Cat, UAS-Dfh and UAS-DfhIR transgenes were independently co-injected with the p{pi}25.7wc helper plasmid into w1 recipient embryos. G0 transformants were identified by outcrossing to w1 and balanced over FM7A, SM5 and TM3 balancer chromosomes (53Go). Transgenic lines were tested using the daG32Gal4 driver. UAS-Sod1, UAS-Cat and UAS-Dfh lines exhibiting high expression were selected for use in these studies. DfhIR transgenic lines were likewise examined for dfh silencing. UAS-DfhIR1 (UDIR1) and UAS-DfhIR2 (UDIR2) exhibited robust dfh silencing and were selected for use in these studies. To avoid potential recessive effects associated with P-vector insertions, all of the experiments reported here were conducted with flies hemizygous for UAS-transgenes and/or GAL4 drivers.

Antibody design and purification
Rabbit polyclonal antibodies were raised against the peptides QIETESTLDGATYERVCSDT and CLQIGWFKAGSALNRMKELAQ corresponding to amino acids 61–80 of the predicted amino acid sequence of DFH (25Go) and the C-terminal 21 amino acids of Drosophila mitochondrial aconitase (FlyBase: mitochondrial aconitase), respectively. In both cases, the peptides were conjugated to a keyhole limpet hemocyanin carrier protein prior to injection into rabbits. Antibodies were affinity purified by the manufacturer (Bethyl).

Transcript analysis by northern blotting
Total RNA was extracted from 25 wandering third instar larvae by using TRIzol reagent (Invitrogen). After electrophoretic separation of denatured RNA (10 µg per lane) through 1.5% formaldehyde agarose gels, the RNA was transferred and UV-crosslinked to a nylon membrane (Roche Diagnostics). Dfh, FerHCH and Rp49 (control) RNA were detected by using digoxigenin-labeled DNA probes made with a digoxigenin non-radioactive DNA-labeling kit (Roche Diagnostics).

Immunoblotting
Preparation of Drosophila samples for immunoblotting was performed as described previously (33Go). For this and all other procedures, protein levels were determined with the Bio-Rad protein assay. Samples were separated on 4% stacking, 15% separating SDS polyacrylamide gels. Resolved proteins were electroblotted to HybondC Extra nitrocellulose membranes (Amersham, Piscataway, NJ, USA). Rabbit polyclonal antibodies raised against DFH and Drosophila mitochondrial aconitase peptides and rabbit polyclonal antibodies against the Drosophila FerHCH (a gift from F. Missirlis) were used in combination with goat anti-rabbit IgG horseradish peroxidase conjugate (Stressgen, San Diego, CA, USA) and detected with Amersham Pharmacia ECL immunoblotting detection reagents. Mouse anti-Actin monoclonal IgM antibodies (developed by J.J.-C. Lin and obtained from the Developmental Studies Hybridoma Bank, University of Iowa) were used in combination with anti-mouse IgM alkaline phosphatase conjugate and detected according to the manufacturer's instructions (Sigma, St Louis, MO, USA). Pre-stained protein ladders (Invitrogen) were run in parallel with samples to estimate protein molecular weight.

Assay of SOD and CAT activities
SOD activity was assayed spectrophotometrically using the 6-hydroxydopamine autoxidation method as described (36Go,54Go,55Go) or by the in-gel activity assay (32Go). CAT activity was assayed spectrophotometrically as described (56Go).

Preparation of crude mitochondrial extracts
Mitochondria were prepared using a procedure adapted from that described by Van den Bergh (57Go). The final enriched mitochondrial pellet was resuspended in extraction buffer and sonicated for 15 s.

Assay of aconitase activity and reactivation
Aconitase activity in crude mitochondrial extracts was assayed spectrophotometrically by determining the conversion of isocitrate to cis-aconitate at 240 nm (58Go). Alternatively, mitochondrial and cytoplasmic aconitase activities in whole-animal extracts were assayed after electrophoretic separation (33Go). In vitro reactivation of mitochondrial aconitase prior to electrophoretic separation was carried out as described previously (33Go).

Assay of mitochondrial respiratory complex activities
The enzymatic activities of mitochondrial electron transport complexes I, III and IV and citrate synthase were assayed in crude mitochondrial extracts as described (59Go). Complex II activity was assayed in crude mitochondrial extracts by using the dichlorophenolindophenol method as described (60Go).

Eclosion measurements
Virgin w1; X/X;daG32/daG32 (X being either UAS-Sod1 or UAS-Sod2 or UAS-Cat) females were crossed to either w1; +/+;UDIR1/UDIR1 or w;UDIR2/UDIR2;+/+ or w1;+/+; +/+ males and allowed to lay eggs for 4 days in cornmeal vials; eclosing adult progeny were scored.

Pupariation
First instar larvae hatching within a 3 h window were transferred into vials (30 larvae per vial) and scored for pupariation initially twice daily and later once daily at 25°C as described (61Go).

Larval iron toxicity
First instar larvae hatching within a 3 h window were transferred into vials (30 larvae per vial) containing cornmeal medium fortified with increasing concentrations of FeCl3 and scored for pupariation initially twice daily and later once daily at 25°C as described (61Go).

Adult lifespan determination
Survival of 200 adult males or females on standard cornmeal medium (20 flies per vial) was followed at 25°C with transfer of survivors to fresh vials every 2–3 days.

Larval photography
Wandering third instar larvae were collected, washed twice briefly in distilled water and photographed using a Leica model MZFLIII stereoscope (Northvale, NJ, USA), Qimaging Retiga 1300R digital camera system (Burnaby, BC, USA) and Improvision Openlab 4.0.2 software.

Digitometry of western immunoblots and aconitase strip assays
Digitometry was carried out using NIH Imaging V1.63 software.

Statistical analysis
Results are expressed as mean±SD. The Student's unpaired t-test was used to determine the significance of differences between means. P<0.05 is considered significant.


   ACKNOWLEDGEMENTS
 
We thank Shawna Wicks and Dr Fanis Missirlis for the critical reading of the manuscript and Reynald Tremblay and Dr Joesph Colasanti for assistance with the collection of digital images. This work was supported by a grant from the National Ataxia Foundation (to J.P.P.) and by grants from the Canadian Institutes of Health Research (CIHR) and the Natural Sciences and Engineering Research Council of Canada (NSERC) (to J.P.P and A.J.H.). P.R.A. is supported by Graduate Scholarships from CIHR and NSERC. Funding to pay the Open Access publication charges for this article was provided by the National Ataxia Foundation.

Conflict of Interest statement. None declared.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 

  1. Campuzano, V., Montermini, L., Lutz, Y., Cova, L., Hindelang, C., Jiralerspong, S., Trottier, Y., Kish, S.J., Faucheux, B., Trouillas, P. et al. (1997) Frataxin is reduced in Friedreich ataxia patients and is associated with mitochondrial membranes. Hum. Mol. Genet., 6, 1771–1780.[Abstract/Free Full Text]

  2. Koutnikova, H., Campuzano, V., Foury, F., Dolle, P., Cazzalini, O. and Koenig, M. (1997) Studies of human, mouse and yeast homologues indicate a mitochondrial function for frataxin. Nat. Genet., 16, 345–351.[CrossRef][ISI][Medline]

  3. Tan, G., Chen, L.S., Lonnerdal, B., Gellera, C., Taroni, F.A. and Cortopassi, G.A. (2001) Frataxin expression rescues mitochondrial dysfunctions in FRDA cells. Hum. Mol. Genet., 10, 2099–2107.[Abstract/Free Full Text]

  4. Patel, P.I. and Isaya, G. (2001) Friedreich ataxia: from GAA triplet-repeat expansion to frataxin deficiency. Am. J. Hum. Genet., 69, 15–24.[CrossRef][ISI][Medline]

  5. Delatycki, M.B., Williamson, R. and Forrest, S.M. (2000) Friedreich ataxia: an overview. J. Med. Genet., 37, 1–8.[Abstract/Free Full Text]

  6. Alper, G. and Narayanan, V. (2003) Friedreich's ataxia. Pediatr. Neurol., 28, 335–341.[CrossRef][ISI][Medline]

  7. Babcock, M., de Silva, D., Oaks, R., Davis-Kaplan, S., Jiralerspong, S., Montermini, L., Pandolfo, M. and Kaplan, J. (1997) Regulation of mitochondrial iron accumulation by Yfh1p, a putative homolog of frataxin. Science, 276, 1709–1712.[Abstract/Free Full Text]

  8. Radisky, D.C., Babcock, M.C. and Kaplan, J. (1999) The yeast frataxin homologue mediates mitochondrial iron efflux. Evidence for a mitochondrial iron cycle. J. Biol. Chem., 274, 4497–4499.[Abstract/Free Full Text]

  9. Adamec, J., Rusnak, F., Owen, W.G., Naylor, S., Benson, L.M., Gacy, A.M. and Isaya, G. (2000) Iron-dependent self-assembly of recombinant yeast frataxin: implications for Friedreich ataxia. Am. J. Hum. Genet., 67, 549–562.[CrossRef][ISI][Medline]

  10. Chen, O.P. and Kaplan, J. (2001) YFH1-mediated iron homeostatis is independent of mitochondrial respiration. FEBS Lett., 509, 131–134.[CrossRef][ISI][Medline]

  11. Gakh, O., Adamec, J., Gacy, A.M., Twesten, R.D., Owen, W.G. and Isaya, G. (2002) Physical evidence that yeast frataxin is an iron storage protein. Biochemistry, 41, 6798–6804.[CrossRef][Medline]

  12. Park, S., Gakh, O., Mooney, S.M. and Isaya, G. (2002) The ferroxidase activity of yeast frataxin. J. Biol. Chem., 277, 38589–38595.[Abstract/Free Full Text]

  13. Nichol, H., Gakh, O., O'Neill, H.A., Pickering, I.J., Isaya, G. and George, G.N. (2003) Structure of frataxin iron cores: an X-ray absorption spectroscopic study. Biochemistry, 42, 5971–5976.[CrossRef][Medline]

  14. Park, S., Gakh, O., O'Neill, H.A., Mangravita, A., Nichol, H., Ferreira, G.C. and Isaya, G. (2003) Yeast frataxin sequentially chaperones and stores iron by coupling protein assembly with iron oxidation. J. Biol. Chem., 278, 31340–31351.[Abstract/Free Full Text]

  15. O'Neill, H.A., Gakh, O., Park, S., Cui, J., Mooney, S.M., Sampson, M., Ferreira, G.C. and Isaya, G. (2005) Assembly of human frataxin is a mechanism for detoxifying redox-active iron. Biochemistry, 44, 537–545.[CrossRef][Medline]

  16. Chen, O.S., Hemenway, S. and Kaplan, J. (2002) Genetic analysis of iron citrate toxicity in yeast: implications for mammalian iron homeostasis. Proc. Natl Acad. Sci. USA, 99, 16922–16927.[Abstract/Free Full Text]

  17. Foury, F. (1999) Low iron concentration and aconitase deficiency in a yeast frataxin homologue deficient strain. FEBS Lett., 456, 281–284.[CrossRef][ISI][Medline]

  18. Muhlenhoff, U., Richhardt, N., Ristow, M., Kispal, G. and Lill, R. (2002) The yeast frataxin homolog Yfh1p plays a specific role in the maturation of cellular Fe/S proteins. Hum. Mol. Genet., 11, 2025–2036.[Abstract/Free Full Text]

  19. Muhlenhoff, U., Richhardt, N., Gerber, J. and Lill, R. (2002) Characterization of iron–sulfur protein assembly in isolated mitochondria. A requirement for ATP, NADH, and reduced iron. J. Biol. Chem., 277, 29810–29816.[Abstract/Free Full Text]

  20. Muhlenhoff, U., Gerber, J., Richhardt, N. and Lill, R. (2003) Components involved in assembly and dislocation of iron–sulfur clusters on the scaffold protein Isu1p. EMBO J., 22, 4815–4825.[CrossRef][ISI][Medline]

  21. Lesuisse, E., Santos, R., Matzanke, B.F., Knight, S.A., Camadro, J.M. and Dancis, A. (2003) Iron use for haeme synthesis is under control of the yeast frataxin homologue (Yfh1). Hum. Mol. Genet., 12, 879–889.[Abstract/Free Full Text]

  22. Bulteau, A.L., O'Neill, H.A., Kennedy, M.C., Ikeda-Saito, M., Isaya, G. and Szweda, L.I. (2004) Frataxin acts as an iron chaperone protein to modulate mitochondrial aconitase activity. Science, 305, 242–245.[Abstract/Free Full Text]

  23. Wilson, R.B. and Roof, D.M. (1997) Respiratory deficiency due to loss of mitochondrial DNA in yeast lacking the frataxin homologue. Nat. Genet., 16, 352–357.[CrossRef][ISI][Medline]

  24. Puccio, H., Simon, D., Cossee, M., Criqui-Filipe, P., Tiziano, F., Melki, J., Hindelang, C., Matyas, R., Rustin, P. and Koenig, M. (2001) Mouse models for Friedreich ataxia exhibit cardiomyopathy, sensory nerve defect and Fe–S enzyme deficiency followed by intramitochondrial iron deposits. Nat. Genet., 27, 181–186.[CrossRef][ISI][Medline]

  25. Canizares, J., Blanca, J.M., Navarro, J.A., Monros, E., Palau, F. and Molto, M.D. (2000) dfh is a Drosophila homolog of the Friedreich's ataxia disease gene. Gene, 256, 35–42.[CrossRef][ISI][Medline]

  26. Cavadini, P., Adamec, J., Taroni, F., Gakh, O. and Isaya, G. (2000) Two-step processing of human frataxin by mitochondrial processing peptidase. Precursor and intermediate forms are cleaved at different rates. J. Biol. Chem., 275, 41469–41475.[Abstract/Free Full Text]

  27. Gordon, D.M., Kogan, M., Knight, S.A., Dancis, A., Pain, D., Gordon, B.N. and Kogan, K.L. (2001) Distinct roles for two N-terminal cleaved domains in mitochondrial import of the yeast frataxin homolog, Yfh1p. Hum. Mol. Genet., 10, 259–269.[Abstract/Free Full Text]

  28. Cossee, M., Puccio, H., Gansmuller, A., Koutnikova, H., Dierich, A., LeMeur, M., Fischbeck, K., Dolle, P. and Koenig, M. (2000) Inactivation of the Friedreich ataxia mouse gene leads to early embryonic lethality without iron accumulation. Hum. Mol. Genet., 9, 1219–1226.[Abstract/Free Full Text]

  29. Wodarz, A., Hinz, U., Engelbert, M. and Knust, E. (1995) Expression of crumbs confers apical character on plasma membrane domains of ectodermal epithelia of Drosophila. Cell, 82, 67–76.[CrossRef][ISI][Medline]

  30. Smith, J.E. and Cronmiller, C. (2001) The Drosophila daughterless gene autoregulates and is controlled by both positive and negative cis regulation. Development, 128, 4705–4714.[Abstract/Free Full Text]

  31. Gardner, P.R. and Fridovich, I. (1991) Superoxide sensitivity of the Escherichia coli Aconitase. J. Biol. Chem., 266, 19 328–19 333.

  32. Missirlis, F., Hu, J., Kirby, K., Hilliker, A.J., Rouault, T.A. and Phillips, J.P. (2003) Compartment-specific protection of iron–sulfur proteins by superoxide dismutase. J. Biol. Chem., 278, 47 365–47 369.

  33. Kirby, K., Hu, J., Hilliker, A.J. and Phillips, J.P. (2002) RNA interference-mediated silencing of Sod2 in Drosophila leads to early adult-onset mortality and elevated endogenous oxidative stress. Proc. Natl Acad. Sci. USA., 99, 16162–16167.[Abstract/Free Full Text]

  34. Eisenstein, R.S. and Ross, K.L. (2003) Novel roles for iron regulatory proteins in the adaptive response to iron deficiency. J. Nutr., 133, 1510S–1516S.[Abstract/Free Full Text]

  35. Gustafson, K. and Boulianne, G.L. (1996) Distinct expression patterns detected within individual tissues by the GAL4 enhancer trap technique. Genome, 39, 174–182.[Medline]

  36. Parkes, T.L., Elia, A.J., Dickinson, D., Hilliker, A.J., Phillips, J.P. and Boulianne, G.L. (1998) Extension of Drosophila lifespan by overexpression of human SOD1 in motorneurons. Nat. Genet., 19, 171–174.[CrossRef][ISI][Medline]

  37. Rotig, A., de Lonlay, P., Chretien, D., Foury, F., Koenig, M., Sidi, D., Munnich, A. and Rustin, P. (1997) Aconitase and mitochondrial iron–sulphur protein deficiency in Friedreich ataxia. Nat. Genet., 17, 215–217.[CrossRef][ISI][Medline]

  38. Duttaroy, A., Paul A., Kundu, M., and Belton, A. (2003) A Sod2 null mutation confers severely reduced adult lifespan in Drosophila. Genetics, 165, 2295–2299.[Abstract/Free Full Text]

  39. Iyengar, B., Roote, J. and Campos, A.R. (1999) The tamas gene, identified as a mutation that disrupts larval behavior in Drosophila melanogaster, codes for the mitochondrial DNA polymerase catalytic subunit (DNApol-gamma125). Genetics, 153, 1809–1824.[Abstract/Free Full Text]

  40. Maier, D., Farr, C.L., Poeck, B., Alahari, A., Vogel, M., Fischer, S., Kaguni, L.S. and Schneuwly, S. (2001) Mitochondrial single-stranded DNA-binding protein is required for mitochondrial DNA replication and development in Drosophila melanogaster. Mol. Biol. Cell, 12, 821–830.[Abstract/Free Full Text]

  41. Iyengar, B., Luo, N., Farr, C.L., Kaguni, L.S. and Campos, A.R. (2002) The accessory subunit of DNA polymerase gamma is essential for mitochondrial DNA maintenance and development in Drosophila melanogaster. Proc. Natl Acad. Sci. USA, 99, 4483–4488.[Abstract/Free Full Text]

  42. Gray, N.K., Pantopoulos, K., Dandekar, T., Ackrell, B.A.C. and Hentze, M.W. (1996) Translational regulation of mammalian and Drosophila citric acid cycle enzymes via iron-responsive elements. Proc. Natl Acad. Sci. USA., 93, 4925–4930.[Abstract/Free Full Text]

  43. Stehling, O., Elsasser, H.P., Bruckel, B., Muhlenhoff, U. and Lill, R. (2004) Iron–sulfur protein maturation in human cells: evidence for a function of frataxin. Hum. Mol. Genet., 13, 3007–3015.[Abstract/Free Full Text]

  44. Georgieva, T., Dunkov, B.C., Harizanova, N., Ralchev, K. and Law, J.H. (1999) Iron availability dramatically alters the distribution of ferritin subunit messages in Drosophila melanogaster. Proc. Natl Acad. Sci. USA, 96, 2716–2721.[Abstract/Free Full Text]

  45. Said, G., Marion, M.H., Selva, J. and Jamet, C. (1986) Hypotrophic and dying-back nerve fibers in Friedreich's ataxia. Neurology, 36, 1292–1299.[Abstract/Free Full Text]

  46. Puccio, H. and Koenig, M. (2000) Recent advances in the molecular pathogenesis of Friedreich ataxia. Hum. Mol. Genet., 9, 887–892.[Abstract/Free Full Text]

  47. Parkes, T.L., Kirby, K., Phillips, J.P. and Hilliker, A.J. (1998) Transgenic analysis of the cSOD-null phenotypic syndrome in Drosophila. Genome, 41, 642–651.[Medline]

  48. Sun, J., Folk, D., Bradley, T.J. and Tower, J. (2002) Induced overexpression of mitochondrial Mn-superoxide dismutase extends the life span of adult Drosophila melanogaster. Genetics, 161, 661–672.[Abstract/Free Full Text]

  49. Orr, W.C. and Sohal, R.S. (1992) The effects of Catalase gene overexpression on life span and resistance to oxidative stress in transgenic Drosophila melanogaster. Arch. Biochem. Biophys., 297, 35–41.[CrossRef][ISI][Medline]

  50. Ashburner, M. (1989) Drosophila A Laboratory Handbook. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.

  51. Brand, A.H. and Perrimon, N. (1993) Targeted gene expression as a means of altering cell fates and generating dominant phenotypes. Development, 118, 401–415.[Abstract]

  52. Rubin, G.M. and Spradling, A.C. (1981) Genetic transformation of Drosophila with transposable element vectors. Science, 218, 348–352.

  53. Lindsley, D.L. and Zimm, G. (1992) The Genome of Drosophila melanogaster. Academic Press, New York, NY.

  54. Heikkila, R.E. and Cabbat, F. (1976) A sensitive assay for superoxide dismutase based on the autoxidation of 6-hydroxydopamine. Anal. Biochem., 75, 356–362.[CrossRef][ISI][Medline]

  55. Duttaroy, A., Parkes, T., Emtage, P., Kirby, K., Boulianne, G.L., Wang, X., Hilliker, A.J. and Phillips, J.P. (1997) The manganese superoxide dismutase gene of Drosophila: structure, expression, and evidence for regulation by MAP kinase. DNA Cell Biol., 16, 391–399.[ISI][Medline]

  56. Griswold, C.M., Matthews, A.L., Bewley, K.E. and Mahaffey, J.W. (1993) Molecular characterization and rescue of acatalasemic mutants of Drosophila melanogaster. Genetics, 134, 781–788.[Abstract]

  57. Van den Bergh, S.G. (1967) Preparation of insect mitochondria. Methods Enzymol., 10, 117–122.[Medline]

  58. Henson, C.P. and Cleland, W.W. (1967) Purification and kinetic studies of beef liver cytoplasmic aconitase. J. Biol. Chem., 242, 3833–3838.[Abstract/Free Full Text]

  59. Trounce, I.A., Kim, Y.L., Jun, A.S. and Wallace, D.C. (1996) Assessment of mitochondrial oxidative phosphorylation in patient muscle biopsies, lymphoblasts, and transmitochondrial cell lines. Methods Enzymol., 264, 484–509.[Medline]

  60. Ackrell, B.A., Kearney, E.B. and Singer, T.P. (1997) Mammalian succinate dehydrogenase. Methods Enzymol., 53, 466–483.

  61. Palladino, M.J., Keegan, L.P., O'Connell, M.A. and Reenan, R.A. (2000) A-to-I pre-mRNA editing in Drosophila is primarily involved in adult nervous system function and integrity. Cell, 102, 437–449.[CrossRef][ISI][Medline]


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P. R. Anderson, K. Kirby, W. C. Orr, A. J. Hilliker, and J. P. Phillips
Hydrogen peroxide scavenging rescues frataxin deficiency in a Drosophila model of Friedreich's ataxia
PNAS, January 15, 2008; 105(2): 611 - 616.
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