ALTERATIONS OF MITOCHONDRIAL BIOGENESIS AND ALTERATIONS OF MITOCHONDRIAL ANTIOXIDANT DEFENSE IN FRIEDREICH’S ATAXIA

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ALTERATIONS OF MITOCHONDRIAL BIOGENESIS AND ALTERATIONS OF MITOCHONDRIAL ANTIOXIDANT

DEFENSE IN FRIEDREICH’S ATAXIA

Submitted for the degree of Doctor in Biomedical and Pharmaceuticals Sciences Daniele MARMOLINO

Jury members:

President: Prof. Magali Waelbroeck Secretary: Prof. Massimo Pandolfo ULB members: Prof. Bernard Dan

Prof. Decio Eizirik

Prof. Jean-Marie Vanderwinden Invited members: Prof. Hélène Puccio

Prof. Jörg Schulz

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ALTERATIONS OF MITOCHONDRIAL BIOGENESIS AND ALTERATIONS OF MITOCHONDRIAL ANTIOXIDANT DEFENSE IN

FRIEDREICH‘S ATAXIA

2 Special thanks

Completing a PhD is truly a marathon event, and I would not have been able to complete this journey without the aid and support of countless people over the past four years.

I would like to thank my supervisor, Prof. Massimo Pandolfo, whose expertise added considerably to my graduate experience. I appreciate his vast knowledge and skill in many areas, and his assistance during this long period, which have on occasion made me "GREEN" with envy.

I would like to thank the other members of the laboratory and all the others colleagues, Satyan, Ajay, Myriam, Chantal, Ana, Giuliana, Neslihan, Isabelle, Marie-Aline, Patrick, Laetitia, Marcelo, David D., Michèle, Pierre, Sabrina, David G., Petra, Perrine, Delphine, Samanta, Souad, Stéphanie, Huy, David O., Shan, Francesco, Luca, for the assistance they provided at all levels of the research project.

I would like to thank Prof. Sergio Cocozza from the University of Naples for taking time out from his busy schedule to serve as my external supporter and collaborator. Thanks to people working in Cocozza‘s laboratory: Dr. Antonella Monticelli, Fabio, Paola and Michele.

Very special thanks goes out to Prof. Mario Manto, without whose motivation and encouragement, I would not have considered to finish my PhD. Mario is the one professor/teacher who truly made the difference in my life. It was under his tutelage that I developed a focus and became interested in all the area of Neuroscience. He provided me with direction, technical support and became more of a mentor and a friend, than a professor. It was through his, persistence, understanding and kindness that I completed my doctorate degree and was encouraged to continue for my carrier. I doubt that I will ever be able to convey my appreciation fully, but I owe him my eternal gratitude.

I must also acknowledge Prof. Serge Shiffman, Prof. Jean-Marie Vanderwinden and Prof. Alban de Kerchove d’Exaerde for their suggestions and helpful discussions during this long period.

A special thank to my love Amélie without whose love and encouragements, I would not have finished this thesis.

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ALTERATIONS OF MITOCHONDRIAL BIOGENESIS AND

ALTERATIONS OF MITOCHONDRIAL ANTIOXIDANT DEFENSE IN FRIEDREICH‘S ATAXIA

Finally, my gratitude goes to my family for the support they provided me through my entire life and in particular, I must acknowledge my mother Laura, father Donato, brother Flavio.

In conclusion, I recognize that this research would not have been possible without the support of the Université Libre de Bruxelles (ULB) and the financial assistance of GoFAR (Italy), Fonds Erasme pour la recherché national (Belgium) and the material support of Takeda Pharmaceuticals (Japan), and express my gratitude to those agencies.

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4 Table of contents

I.

Introduction

Clinical Features and Pathogenesis of Friedreich’s Ataxia p.10

Clinical features p.10

Pathophysiology p.12

Mutations in the FXN Gene Cause FRDA p.16

Structure and Regulation of the FXN Gene p.16

Molecular Mechanism of GAA Triplet Repeat Expansion p.18 Genotype Phenotype Correlation and Point Mutations p.23

Animal and Cellular Models p.26

Frataxin Protein p.28

Frataxin Structure p.28

Cellular Function of Frataxin p.30

Abnormal Iron Homeostasis and Oxidative Stress Defense p.32

Fe-S Cluster Assembly p.32

Iron Homeostasis p.35

Iron Binding Properties p.40

Cellular Antioxidant Defenses p.41

Mitochondrial DNA Alterations p.44

Alterations in Nuclear DNA p.45

Mitochondrial Function p.48

Oxidative Phosphorylation p.49

Mitochondrial DNA: Replication, Transcription and Translation p.49 Regulation of Mitochondrial Biogenesis by PGC-1    p.51 Therapeutics Approaches for the Treatment of FRDA p.56

Antioxidants and Oxidative Phosphorylation p.56

Iron Chelating Strategy p.58

Histone Deacetylase Inhibitors p.60

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Human Recombinant Erythropoietin p.62

Peroxisome Proliferator-Activated Receptor-γ Agonists p.63

Protein Replacement p.64

II. PPAR-gamma agonist Azelaoyl PAF increases frataxin protein and mRNA expression: new implications for the Friedreich's ataxia therapy

Abstract p.67

Introduction p.68

Materials and methods p.69

Cell Culture p.69

Cell Treatments p.69

Western Blot p.69

Real-Time PCR p.70

Statistical Analyses p.70

Results p.70

Discussion p.74

Acknowledgments p.75

References p.76

III. Functional genomic analysis of frataxin deficiency reveals tissue-specific alterations and identifies the PPARgamma pathway as a therapeutic target in Friedreich's ataxia

Abstract p.81

Results p.83

Functional genomic analysis of frataxin deficient mice p.83

FRDA patients are insulin resistant p.89

The PPAR- pathway is deregulated in frataxin deficient mice and

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FRDA patients p.90

PPAR- manipulation affects frataxin levels p.90

Animals and tissue samples p.96

Microarrays p.96

Cell culture p.97

qPCR experiments p.97

Western blot and luciferase assays p.97

Assessment of physical impairment in FRDA patients p.97 Assessment of insulin sensitivity in FRDA patients and healthy controls p.98

Supplementary material p.99

Acknowledgements p.107

Funding p.108

IV. PGC-1alpha down-regulation affects the antioxidant response in Friedreich's ataxia

Abstract p.117

Results p.120

PGC-1a reduction is associated to reduced SOD2 in FRDA fibroblasts and

does not increase after H2O2 incubation p.120

PGC-1a down-regulation by RNAi results in lack of SOD2 response to H2O2 p.121 Effect of PPAR- and AMPK agonists on the antioxidant response in FRDA

fibroblasts p.121

Effect of Pioglitazone in vivo in frataxin-deficient (KIKO) Mice p.127

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Ethics statement p.134

Patients p.134

Cell cultures p.134

Cell Treatment p.135

Animal experiments p.135

Cells transfections p.135

Real-time quantitative PCR p.136

Western blot p.137

Statistical analyses p.137

Supporting Information p.138

Acknowledgments p.139

Author Contributions p.139

V.

Conclusions

p.148

VI.

Perspectives

p.152

VII.

Thesis Annex

p.154

Experimental study of sirt1 pathway in neurodegenerative diseases

VIII.

References

p.156

IX.

Curriculum Vitae

p.192

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8 List of abbreviations

ADP Adenosine diphosphate

AICAR 5-Aminoimidazole-4-carboxamide 1-b-D-ribofuranoside ALS Amyotrophic lateral sclerosis

AMPK AMP-protein activated kinase APAF Azelaoyl PAF

ATP adenosine triphosphate BBB Blood brain barrier cDNA Complementary DNA CNS Central nervous system CoQ coenzyme Q

DFP Deferiprone DFX Deferoxiamine DMSO Dimethylsulfoxide DNA Deoxyribonucleic acid DRG Dorsal root ganglia Fe Iron

Fe-S Iron sulphur clusters FRDA Friedreich‟s ataxia FXN Frataxin encoding gene GEO Gene Expression Omnibus HAT Histone acetyl-transferase HDAC Histone deacetylase

HDACi Histone deacetylase inhibitors HP1 Heretochromatin protein 1

huEPO Human recombinant erythropoietin IRE Iron responsive elements

ISC Iron sulphur cluster

KIKO FRDA Knock in/knock out mice mRNA Messenger RNA

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NPC Neural precursor cells

OXPHOS Oxidative phosphorylation PBMC Peripheral blood mononuclear cell PD Parkinson‟s disease

PGC-1α Peroxisome proliferator activated receptors gamma coactivator-1α PPARs Peroxisome proliferator activated receptors

ROS Reactive oxygen species RT-PCR Real Time PCR S Sulphur

SEM Standard error of mean siRNA Small interference RNA

SODs Superoxide dismutases enzymes

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10 I. INTRODUCTION

Clinical Features and Pathogenesis of Friedreich’s Ataxia

Clinical features

Friedreich‘s ataxia (FRDA) was described in 1863 by Nikolaus Friedreich (Fig. 1) (Friedreich, 1863) and accepted as a new disease in 1877 (Friedreich, 1876). The major clinical and pathological features of this hereditary ataxia are: age of onset around puberty, degenerative atrophy of the posterior columns of the spinal cord leading to progressive ataxia, sensory loss and muscle weakness; additional features were scoliosis, foot deformity and cardiac symptoms (Table. 1). In 1890 Ladame published the description of 165 cases (Ladame, 1890). Nevertheless, misdiagnosis have been frequent among FRDA, other hereditary ataxias and Charcot-Marie- Tooth disease (Harding et al., 1981; Salisachs et al., 1982; Panas et al., 2002). In the early 1980s, Harding (Harding et al., 1981) proposed a list of criteria for the diagnosis of FRDA (Ackroyd et al., 1984) and the recessive inheritance of the disease was widely accepted (Webb et al., 1999;

Illarioshkin et al., 2000; Panas et al., 2007).

In 1988, the mutated gene in FRDA was mapped to chromosome 9 by linkage analysis with restriction length polymorphism (RFLP) markers (Chamberlain et al., 1988).

Several research groups allowed the chromosome region at 9q13 to be narrowed down to 150 kb (Fujita et al., 1990; Shaw et al., 1990; Sirugo et al., 1993; Rodius et al., 1994; Montermini et al., 1995) and in 1996 the mutated gene in FRDA (now named FXN gene according to HUGO Gene Nomenclature Committee) and its most common mutation were identified (Campuzano et al., 1996). The majority of FRDA patients (96%) are homozygous for an unstable GAA trinucleotide repeat expansion in the first intron of the FXN gene, but a few patients are heterozygous with other loss-of-function mutations found on the other allele (Campuzano et al., 1996; Cossée et al., 2000). The identification of the FXN gene and of its most frequent mutation provided a valuable tool for the diagnosis of FRDA. Additionally, it demonstrated that the typical and atypical phenotypes of the disease were caused by mutations in the same gene (Chamberlain et al., 1989;

De Michele et al., 1994; Durr et al., 1996; Klockgether et al., 1996).

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Since the beginning, FRDA was suggested to be an inherited metabolic disease, but no specific biochemical deficiency could be detected (Purkiss et al., 1981). However, the mitochondrial nature of the pathology was suspected (Barbeau et al., 1980). The identification of the gene allowed the biochemical defect underlying the disease to be clarified: the GAA repeat expansion causes an abnormal conformation of DNA and a decrease in transcription of the FXN gene, encoding for the protein frataxin (Campuzano et al., 1996). Frataxin localizes to the mitochondria, but its function remains unclear. It has been suggested that frataxin is involved in iron homeostasis (iron-sulfur clusters and heme synthesis), iron strorage and detoxification, respiratory control, and resistance to oxidative stress (Rotig et al., 1997; Babcock et al., 1997;

Muhlenhoff et al., 2002; Lesuisse et al., 2003; Bulteau et al., 2004).

A link to oxidative stress was anticipated due to the similarity of phenotypes between FRDA and ataxia with vitamin E deficiency (AVED) (Gibson et al., 1996). The importance of oxidative damage in FRDA is also supported by some benefical effect of anti-oxidant treatment, e.g. the free-radical scavenger idebenone can in some cases significantly reduce myocardial hypertrophy (Rustin et al. 1999).

Fig. 1 Picture of Dr. Nikolaus Friedreich

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Clinical Features Mean Frequency (%)

Ataxia 98

Areflexia 92.1

Dysarthria 93.6

Extensor plantar responses 83.5

Decreased position and vibratory sense 92.7

Foot Deformity 76.8

Scoliosis 84.3

Abnormalities on ECG 80.3

Abnormalities on cardiac ultrasound 50

Diabetes 8

Nystagmus 32

Hearing loss 20

Table. 1 Clinical features of Friedreich‘s ataxia patients, mean frequency (%).

Pathophysiology

Friedreich‘s ataxia neuropathology shows marked involvement of the spinal cord, peripheral nerves and cerebellum (Pandolfo, 2009). The first site of neurodegeneration is the dorsal root ganglia (DRG), with a loss of large sensory neurons, followed by degeneration of the posterior columns, the cortico-spinal and spino-cerebellar tracts of the spinal cords and the dentate nucleus in the cerebellum (Fig. 2). Patients show axonal neuropathy with a profound reduction in the density of large myelinated fibers (Hughes et al., 1968; Rizzuto et al., 1981; Said et al., 1986). Interestingly, the density of small myelinated fibers remains generally normal (Said et al., 1986) or slightly reduced (Zouari et al., 1998), while the fine unmyelinated fibers remain preserved. Onion-bulb complexes may also be present (Rizzuto et al., 1981; Barreira et al., 1999).

A severe or complete loss of sensory nerve action potentials can be observed electro- physiologically in peripheral nerves, as well as a moderate decreased in nerve conduction velocities (Zouari et al., 1998). The peripheral sensory nerve pathology underlies the loss of deep tendon reflexes in FRDA patients. The cellular events underlying the large myelinated fibers loss

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are not fully understood, but both axonal degeneration and Schwann cells involvement with demyelination have been shown to occur (Lu et al., 2009). Magnetic resonance imaging (MRI) of the cervical spinal cord in FRDA patients shows degeneration of posterior and lateral columns (Mascalchi et al., 1994). Neuronal degeneration in the dorsal columns leads to the loss of position and vibration senses. Loss of myelinated fibers and gliosis is characteristic of these regions of the spinal cord (Hughes et al., 1968; Lamarche et al., 1982; Koeppen et al., 2009). Severe neuronal loss is also observed in the Clarke‘s column, with atrophy in the spino-cerebellar tracts. The Clarke‘s column is a major relay center for unconscious proprioception, from which sensory information is passed to the cerebellar cortex by the spino-cerebellar tracts. The sensory (proprioceptive) component of ataxia in FRDA patients is due to this pathology. Atrophy is also observed in the cortico-spinal motor tracts, which are more severely affected in their distal portions, suggests a ―dying back‖ process (Said et al., 1986). The degeneration of cortico-spinal tracts leads to muscle weakness and extensor plantar responses. In the cerebellum, the dentate nucleus is severely affected but the cortex is spared during the early stages of the disease, until Purkinje cell loss can be observed (Koeppen et al., 2007). Dentate atrophy compromises the cerebellar outflow pathway and underlies the cerebellar component of ataxia in FRDA. Other organs affected in FRDA patients include heart, pancreas and skeleton. The heart is affected in the majority of patients, the most common cardiac lesion being hypertrophic cardiomyopathy in which ventricular and inter-ventricular septum walls are thickened. Iron deposition in the myocardium has also been reported (Sanchez-Casis et al., 1976; Lamarche et al., 1993). Diabetes in FRDA results from a combination of peripheral insulin resistance and insulin deficiency. Islet beta cells are lost without the signs of autoimmune attack associated with type I diabetes (Schoenle et al., 1989).

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Fig. 2 An overview of the main sites of neuronal loss and organ dysfunction in Friedreich‘s Ataxia (FRDA). Large red dots indicate severe neuronal loss. Pathological features are shown for the brain, spinal cord (SC), heart (H) (hypertrophy) and pancreas (P) (diabetes). Adapted with permission from Taroni and Di Donato, 2004.

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Mutations in the FXN Gene Cause FRDA

Structure and Regulation of the FXN Gene

The FXN gene was identified in 1996 (Campuzano et al., 1996). The gene is composed of seven exons (exons 1-4, 5a, 5b and 6) encompassing 85 kb of genomic DNA (Fig. 3). The 5‘ end plus the first exon of the gene encloses an unmethylated CpG island containing several rare restriction sites. The major FXN transcript has a size of 1.3 kb and is composed of five exons, from 1 to 5a (Campuzano et al., 1996). It encodes a 210-amino acid protein called frataxin (isoform A) (Fig. 4). Much less represented alternative transcripts contain exon 5b instead of 5a, in presence or not of the non-coding exon 6 (Campuzano et al. 1996) (isoforms B and B1). Exon 5b is located 40 kb downstream of exon 5a and carries an in-frame stop codon, resulting in a shorter transcript of 171 amino acids (isoform B). This isoform also differs from isoform A for 11 amino acids at the C-terminus (Campuzano et al., 1996). The most conserved domains of frataxin are encoded by exons 4 and 5a. A third transcript was also isolated (Pianese et al., 2002).

It is generated by alternative splicing at intron 4, resulting in an 8 bp insertion between exons 4 and 5a. This splicing introduces a frame shift with the appearance of a new stop codon in exon 5a. The transcript thus encodes a putative 196 amino acid protein (isoform A1) that differs from isoform A after residue 160. This transcript is less abundant than isoform A and is present in brain, cerebellum, spinal cord, heart and skeletal muscle (Pianese et al., 2002). No functional data have been reported regarding the transcript A1 and B.

The promoter region is located in the 1,255-bp region extending 5‘ from the human FXN open reading frame (Greene et al., 2005). There is no TATA sequence in this region, but repetitive elements such as retro-elements (AluJb, AluY and L2) and mammalian-wide interspersed repeats (MIR) are present. The Alu and MIR elements function as enhancer for the promoter (Greene et al., 2005). An E-box element located in the first intron can modulate the activity of the promoter (Greene et al., 2007). No differences have been described between the promoter sequences from unaffected individuals and FRDA patients (Greene et al., 2005). The regulation of the human FXN gene is not completely known. The E-box sequence binds helix- loop-helix family transcription factors and one potential factor for this region is the muscle-

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specific factor Mt (Greene et al., 2007). Some evidence shows that FXN expression is also iron regulated. In fact, the iron chelator deferoxamine significantly reduces frataxin expression (Li et al., 2008). Conversely, addition of ferric ammonium citrate or hemin increases frataxin expression (Sarsero et al., 2003; Li et al., 2008). In promoter-luciferase constructs cell lines, iron seems to act as a transcription regulator of frataxin (Li et al., 2008). Moreover, in mouse, the fxn gene seems to be directly regulated by the transcription factor hypoxia-inducible factor 2α (HIF- 2α, Epas1 gene) (Oktay et al., 2007) and a reduction in frataxin amount (about 50%) in the liver of the Epas^-/- knockout mice was observed. However, no data are yet available about a role of HIF-2α in humans.

Fig. 3 FXN gene structure in normal conditions (9 GAA repeats) and in presence of an expanded GAA trinucleotide repeat (700/1500 GAA repeats) in the first intron of the gene. Ex1: exon 1; Ex2: exon 2; Ex3:

exon 3, Ex4: exon 4: Ex5: exon 5; Ex6 : exon 6.

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Fig. 4 Frataxin alternative splicing. The mature form of the protein is the isoform A (210 aa).

Molecular Mechanism of GAA Triplet Repeat Expansion

In 98% of FRDA patients a GAA trinucleotide repeat expansion has been found in the first intron of the FXN gene (Campuzano et al., 1996). In healthy individuals, shorter GAA repeats are present in the FXN gene. In Europeans, GAA repeats in normal chromosomes are polymorphic with a bimodal distribution of repeat lengths, 83% having 6 to 12 GAA triplets (small normal, SN) and 17% having 13 to 34 triplets (large normal, LN). Half of the normal alleles carry nine GAA triplets (Cossée et al., 1997; Montermini et al., 1997; Justice et al., 2001).

The data was confirmed in African American, African and Syrian populations (Montermini et al., 1997). Less than 1% of LN alleles has more than 34 GAA repeats, which can be interrupted, most commonly by a hexanucleotide repeat (GAGGAA), or be continuous (Cossée et al., 1997;

Montermini et al., 1997). Alleles with interrupted GAA repeats are generally stable, even when their length corresponds to more than 100 GAA repeats (Cossée et al., 1997), but uninterrupted

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runs of 34 GAA repeats or more have been shown to undergo hyperexpansion to several hundred GAA triplets in one generation (Cossée et al., 1997; Montermini et al., 1997; Schols et al., 1997).

The threshold length for instability and expansion seems to be between 26 and 44 uninterrupted GAA repeats (Sharma et al., 2002; Pollard et al., 2004). Alleles carrying more than 44 uninterrupted repeats have been reported to be already associated with FRDA symptoms (Sharma et al., 2004), but GAA expansion in most FRDA patient contain 500-1000 repeats, up to 1700 (Campuzano et al., 1996; Durr et al., 1996; Filla et al., 1996; Epplen et al., 1997; Montermini et al., 1997; Sharma et al., 2004).

Analysis of intergenerational variability shows that paternal transmission is most often accompanied by a contraction of the repeats while maternal transmission may result in expansion or contraction (Monros et al., 1997; Pianese et al., 1997; Schols et al., 1997; De Michele et al., 1998; Delatycki et al., 1998). Reversion of the expansion to a normal length allele is very rare (De Michele et al., 1998; Bidichandani et al., 1999; Sharma et al., 2002).

Expanded GAA triplet repeats also show extensive instability in cultured cells, in peripheral blood leukocytes, in the central nervous system, in the DRG, the spinal cord and the heart (Bidichandani et al., 1999; Sharma et al., 2002; De Biase et al., 2007; Montermini et al., 1997). This leads to somatic mosaicism. DRG are highly sensitive to frataxin deficiency and their degeneration is among the primary causes of neurological problems in FRDA patients (Simon et al., 2004). In DRG somatic instability starts after early embryonic development and continues after birth throughout life, resulting in progressive, age-dependent accumulation of larger GAA triplet repeat expansions (De Biase et al., 2007). Thus, DRG somatic instability may contribute to disease progression. This was not seen in other regions of the central nervous system.

In vitro, GAA expanded repeats can adopt unusual structures (presumably a triplex) for sequences containing 79 and 100 GAA repeats, but not for those containing 45 repeats (Bidichandani et al., 1998). In a study of long tracts of GAA·TTC (150 and 270 repeats), a novel DNA structure was described (sticky DNA) which results from intra-molecular association of triplexes (Sakamoto et al., 1999; Le Proust et al., 2000; Heidenfelder et al., 2003).

Accordingly, the transcription silencing caused by pathological expansions has been attributed to the formation of these non-B DNA structures, either intramolocular triplexes and sticky DNA, or persistent RNA·DNA hybrids (Wells et al., 2008). The proposed molecular mechanism

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underlying the inhibition of transcription by sticky DNA involves the sequestration of RNA polymerase by its direct binding to the complex DNA structure (Sakamoto et al., 2001).

Transcription of a GAA·TTC template (88 repeats) using T7 DNA polymerase showed that the polymerase paused at the distal end of the repeat (Grabczyk et al., 2000) and that this was tightly linked to RNA·DNA hybrid formation (Grabczyk et al., 2007) (Fig. 5). These in vitro studies demonstrated that RNA polymerase is arrested by triplex structures, preventing transcription elongation.

Epigenetic studies in the FXN promoter and intron regions flanking the GAA repeat expansion revealed marks of condensed heterochromatin, indicating that an epigenetic mechanism may be ultimately responsible of FXN silencing in vivo. These marks include increased methylation of specific CpG sites in FRDA patients‘ lymphoblasts (Greene et al., 2007), peripheral blood (Castaldo et al., 2008), brain and heart tissues (Al-Mahdawi et al., 2008) as well as reduction of histone H3 and H4 acetylation levels and increased histone H3 lysine 9 (H3K9) trimethylation (Greene et al., 2007; Herman et al., 2007; Al-Mahdawi et al., 2008).

Histone hypoacetylation was not observed in the promoter region (Herman et al. 2007) (Fig. 6).

The GAA·TTC duplex and triplex structures can reduce the efficiency of nucleosome assembly (Ruan et al., 2008). Thus, the non-B structures adopted by long tracts of GAA repeats may generate heterochromatin-dependent and -independent gene silencing.

DNA replication, recombination and repair are also affected by triplex and sticky DNA structures (Wells et al., 2003). In Escherichia coli and Saccharomyces cerevisiae the presence of a GAA repeat led to attenuation of replication, to the occurrence of small slippage events and large contractions (Heidefelder et al., 2003; Krasilnikova et al., 2004; Pollard et al., 2004). The somatic instability observed in post-mitotic neurons suggest that other mechanisms than replication, such as transcription and post-replicative DNA repair could be responsible for the triplet repeat expansions observed in FRDA patients.

Recent studies on a model for the study of GAA repeat expansion in human cells show that transcription through the repeat tracts is a major contributor for expansions (Ditch et al., 2009).

Other uninterrupted GAA repeat sequences up to (GAA)44 have been found in the human and mouse genomes, but only FXN GAA repeats of the same size show a high mutation load, suggesting that somatic instability is locus-specific (Rindler et al., 2006).

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Fig. 5 GAA repeat-mediated gene silencing in FRDA. Blue, GAA strand; yellow, TTC strand; violet, Pol II: mRNA polymerase; orange, DNA_RNA hybrid; green, transcribed RNA. Distal pause is the site where the Pol II is blocked. Normal frataxin transcription. In the presence of a normal GAA repeat (GAA <35, blue strand), Pol II normally transcribes frataxin (RNA in green). DNA triplex formation in FRDA. In the presence of a pathological (GAA)n repeat expansion (GAA >61 to 1,700 repetitions, blue strand with red dots), the (GAA)n generates a rotation of the DNA helix with subsequent formation of an abnormal third strand (DNA triplex). Transcriptional block in FRDA. Once the triplex is formed, a block of the transcription occurs (distal pause), generating an interruption in the starting frataxin transcription (smaller green mRNA). DNA_RNA hybrid formation in FRDA. With the transcriptional block at the distal pause, the nascent mRNA transcript (smaller green mRNA) can anneal to the free single-strand of TCC template (in yellow) with the subsequent formation of an RNA_DNA hybrid (orange/yellow double helix) that displaces the much less stable triplex structure. Adapted with permission from Marmolino and Acquaviva, 2009.

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Fig. 6 A model for heterochromatin formation in FRDA. In the absence of a pathological GAA repeat (GAA <35 repetitions) in the first intron of the FXN gene, the switching between accessible (left side) and inaccessible (right side) chromatin is regulated by two classes of enzymes, histone acetyl transferases (HATs in orange; transfer of acetyl group (Ac in grey) on histone residues (green)) and histone deacetylases (HDACs in grey; deacetylated histone residues). In FRDA, the pathological (GAA)n repeat expansion (GAA >61 to 1,700 repetitions) enhances the formation of a non B-DNA structure and may induce an increase in the recruitment of HDACs that remove the acetylated groups. Moreover, in this condition, histone methyltransferase enzymes (HMTransferase in violet) methylate histone residues (Me in violet) generating a much more inaccessible form of chromatin. This switch to a hypoacetylated and hypermethylated form of chromatin is called heterochromatin and is associated with gene silencing.

Finally, several proteins, such as the heterochromatin repressive protein HP1 (HP1 in red), recognize the methylated regions and may further compact chromatin. Adapted by permission from Marmolino and Acquaviva, 2009.

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Genotype Phenotype Correlation and Point Mutations

FRDA patients show a direct correlation between the lengths of GAA repeat expansions and the presence and timing of several features of the disease. An inverse correlation was found between the size of the smaller expansion and both the age of onset and rate of disease progression (time until wheelchair) (Durr et al., 1996; Filla et al., 1996; Montermini et al., 1997;

Schols et al., 1997; Delatycki et al.1999). Cardiomyopathy is also more frequent in patients with large expansions in the smaller allele (Durr et al., 1996; Filla et al., 1996; Montermini et al., 1997; Delatycki et al., 1999). Diabetes does not appear to be associated with either the number of GAA repeats (Durr et al., 1996; Delatycki et al., 1999), but usually develops during the late stages of disease (Filla et al., 1996). Recently, however, insulin sensitivity, regardless of the presence of diabetes or carbohydrate intolerance, was found to be inversely correlated with the number of GAA repeats in the smaller allele (Coppola et al., 2009). Furthermore, severity of dysarthria, skeletal deformities, optic atrophy and hearing loss show direct correlation to GAA expansion size (Filla et al., 1996; Montermini et al., 1997). Additionally, expansion size has been shown to be associated with the severity of sensory neuropathy (Santoro et al., 1999). Loss of large myelinated fibers (>7 μm) is directly correlated to the duration of disease and is inversely correlated to the GAA repeat expansion size in the smaller allele. The methylation of two CpG sites in the genome has been directly correlated to the size of the smaller allele and indirectly correlated to the age of onset in FRDA (Castaldo et al., 2008). Residual levels of frataxin vary according to the expansion and cell type. In peripheral blood leukocytes, frataxin levels in patients range from 5 to 30% of normal (Gellera et al., 2007).

Despite the significant correlation of GAA repeat sizes, particularly the smaller GAA repeat, with age of onset and disease severity, variability among individuals is very high and it is not possible to predict the clinical severity based only on the GAA mutation. Several factors may explain the clinical variability observed among individuals with almost identical number of repeats. One such factor could be mitotic instability causing somatic mosaicism of expansion sizes (Montermini et al., 1997; a et al., 2002). Mitochondrial haplotype may also affect FRDA phenotype, as described for a population from southern Italy (Giacchetti et al., 2004). Other genetic or non-genetic modifiers have still to be identified.

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About 2% of the mutations found in FRDA patients are other loss-of-function mutations (Campuzano et al., 1996; Cossée et al., 1999). So far, 43 mutations in the FXN gene (nonsense, missense, frameshift, splice-site and a deletion) have been described in patients (Fig. 7). All patients carrying these mutations are compound heterozygotes with an expanded GAA repeat on the other allele. The occurrence of these mutations is rare, ~1: million individuals (Delatycki et al., 2000) or 1:2,500 FRDA carriers (Cossée et al., 1999). Deletion of the frataxin-encoding gene in mice causes embryonic lethality (Cossée et al., 2000), suggesting that null mutations in humans may also result in a very severe or lethal phenotype. The effects of genomic mutations on transcript abundance or on the specific protein defect have been poorly studied. 50% of the mutations identified lead to the absence or a truncated form of frataxin. Five of these mutations affect the translation initiation codon, resulting in a translated protein lacking the mitochondrial targeting. Six mutations affect splice-site donors and one mutation a splice acceptor site, leading to aberrant splicing and predicted exon skipping. Studies suggest that aberrantly spliced mRNA may be unstable and rapidly degraded (Gellera et al., 2007). Four nonsense and nine frame shift mutations introduce premature stop codons and nearly all of the predicted proteins lack exons 3- 5. Only one mutation causing a deletion of 2,776 bp was reported (Zuhlke et al., 2004). This mutation is also expected to lead to truncated frataxin. All these mutations are associated with typical FRDA, in some cases with particularly severe phenotypes. A single amino-acid change is predicted for 17 missense mutations. These mutations span across exons 1, 3, 4 and 5, with a cluster of mutations found within exons 4 and 5, which correspond to the C-terminal and most conserved portion of frataxin. The most common mutations are those that affect the ATG translation start codon, G130V and I154F. In particular, evidence of founder events for the M1I (Zuhlke et al., 2004) and G130V (Delatycki et al., 1999) mutations has been documented.

The phenotypic features of patients harboring point mutations are frequently those of typical FRDA, though dysarthria and diabetes are less frequent (Cossée et al., 1999). In general, point mutations that result in complete loss of function of frataxin are associated with a severe phenotype. Conversely, heterozygous patients for the G130V mutation, which causes only a partial loss of function, have a milder disease presentation (Bidichandani et al., 1997; Cossée et al., 1999), with frataxin levels similar to those in healthy carriers (Bidichandani et al., 1997). The I154F mutation, described in Italian families, despite the apparently conservative amino acid

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change results in severe loss of function and patients present with a typical FRDA phenotype (Campuzano et al., 1996; Filla et al., 1996). In all cases, the size of the GAA expansion in the other allele may modulate the effect of the mutation.

The residual functionality of mutated frataxins has been assessed by complementation analysis in fraxain depleted cellular models (yeast and mammalian) and in in vitro reconstructed systems, as detailed below in the section on frataxin function. Point mutations have helped to identify important aminoacid residues for frataxin function.

Fig. 7 FXN gene loss of functions. Upper panel: the FXN gene with a GAA expansion in the first intron.

Lower panel: an illustration of all FXN gene loss of functions so far observed in exons.

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Animal and Cellular Models

Animal and cellular models are essential to study FRDA pathogenesis and to develop and test potential therapeutics. However, there have been major difficulties in generating models that faithfully reproduce the features of the human disease (Puccio, 2009).

Frataxin knock-out mice show early embryonic lethality. Viable mouse models were first obtained through a conditional gene targeting approach, to create floxed exon 4 frataxin alleles.

Muscle and neuron-specific mutants displayed many of the symptoms or pathophysiology of FRDA. A tamoxifen-inducible neuron-specific prion mutant was also produced, generating a progressive neurological phenotype with slow evolution that recreates the neurological features of the human disease. In the effort to create a model closer to the human disease, a (GAA)230

repeat mouse knock-in at the endogenous frataxin gene (frda) called KIKI was generated (Rai et al., 2008). The homozygous mutant only shows a 25% decrease in frataxin expression compared to wild-type. To further decrease frataxin expression, frataxin knock-in-knock-out mice (frda- /230GAA) were further generated, expressing approximately 25-30% of wild-type frataxin levels, a reduction associated with mild, but clinically evident FRDA in humans. These mice manifest no pathology, with only minor motor abnormalities, but changes in gene expression in the CNS, heart, muscle and liver. Two transgenic mouse lines carrying the human FXN were created. They were crossed to frataxin KO mice, to create a mutant only expressing human frataxin mRNA and protein in a homozygous frataxin KO background. The phenotype of the expanded GAA repeat transgenic (Y8 mice) shows decreased aconitase activity, oxidative stress, and develops progressive neurodegenerative and cardiac pathological phenotypes (Al-Mahdawi et al., 2007).

Coordination deficits are also present, together with a progressive decrease in locomotor activity.

These animals also show changes in DNA methylation and histone modifications that indicate epigenetic frataxin gene repression, however levels are close to wild-type levels in many tissues, as the expanded repeat is relatively small (around 200 triplets). The reason for the relatively severe phenotype is because human frataxin only partially complements mouse frataxin.

Drosophila and C. elegans models have been created based on RNAi technology. These models show variable phenotypes according to the timing and severity of frataxin downregulation

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and are of great interest for studies on genetic modifiers (Anderson et al., 2008; Vázquez- Manrique et al., 2006).

Cellular models include cells derived from human FRDA patients, cells derived from animal models, and cells that have been engineered to modify frataxin expression. The available cells from patients are fibroblasts, lymphoblastoid cell lines, and primary lymphocytes. The problem with these cell types is that they do not constitute tissues affected by FRDA. It has so far been impossible to obtain long term cultures of cells entirely devoid of frataxin that can be isolated from conditional KO mouse models, confirming that frataxin is essential for cellular viability in higher organisms. The only way of obtaining partially frataxin deficient cell types that show vulnerability in FRDA has so far been by using RNAi technology. More recently, novel

‗‗humanized‘‘ mouse cell models based on FRDA point mutations were developed (Calmels et al., 2009). These are mouse fibroblast cell lines deleted for endogenous murine frataxin and expressing a human frataxin cDNA carrying pathogenic FRDA missense mutations. These lines reproduce many biochemical phenotypes associated with FRDA, but they have no GAA repeat expansions and are not the cell types primarily targeted buy the disease.

Therefore, new cellular models are needed to pursue investigations into the pathogenesis and physiopathology of FRDA, in particular the basis of cellular specific vulnerability, and to test new treatments.

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Frataxin Protein

Frataxin Structure

Frataxins are small proteins (100-220 amino acids) highly conserved among eukaryotes and some prokaryotes (Gibson et al., 1996). Eukaryotic frataxins are localized to the mitochondrial matrix (Gibson et al., 1996; Campuzano et al., 1997; Koutnikova et al., 1997;

Vàsquez-Manrique et al., 2006; Llorens et al., 2007). Structures were determined in solution by Nuclear Magnetic Resonance (NMR) for the human isoform A (residues 91 to 210) (Musco et al., 2000), the E. coli frataxin homolog CyaY (Nair et al., 2004) and the mature yeast frataxin homologue Yfh1 (He et al., 2004). Crystal structures are also available (Dhe-Paganon et al., 2000).

The frataxin fold is unique and consists of a large, twisted, six-stranded β-antiparallel sheet, flanked by N- and C-terminal α helices (α1 and α2), with no main surface cavity (Fig. 8).

Frataxin presents a patch of negatively charged residues on the helical plane, which could be involved in iron binding (Adinolfi et al., 2002). On the other side, the β sheet surface is mostly uncharged. These surfaces may be responsible for protein-protein interactions. Most of the residues affected by FRDA mutations are in the neutral surface.

Frataxin is translated by cytoplasmic ribosomes (Saint-Georges et al., 2008), then imported into the mitochondria (Koutnikova et al., 1997), where it is proteolytically cleaved in a two-step process to generate the mature protein (Branda et al., 1999; Cavadini et al., 2000). In yeast and humans, frataxin maturation has been shown to depend on the mitochondrial processing peptidase (MPP) (Branda et al., 1999; Cavadini et al., 2000; Gordon et al., 2001). Also the N- terminus of mouse frataxin has been shown to interact with the β-subunit of MPP (Koutnikova et al., 1998). Frataxin contains two MPP cleavage consensus sequences defining a domain I (residues 1- 20) and a domain II (residues 21-51). In Yfh1, deletion of domain II leads to the loss of the mitochondrial targeting signal (Knight et al., 1998; Voisine et al., 2000). In yeast, the Ssc1 Hsp70 family ATPase, constituent of the import motor component of the Translocase of the Inner Mitochondrial membrane (TIM), is crucial for Yfh1 transport across the inner membrane, as

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shown by the observation that ssc1-3 mutants cannot process Yfh1 to the intermediate or mature forms (Voisine et al., 2000).

Recombinant frataxin starting at aminoacid 81 (m81-FXN), where the second cleavage site is located, co-migrates with endogenous frataxin in fibroblasts, lymphoblasts and heart tissue (Condo et al., 2007). It can also rescue aconitase activity deficiency in FRDA patient cells (Condo et al., 2007) and the lethal phenotype in frataxin-deficient murine fibroblasts (Schmucker et al., 2008), supporting the idea that this is the mature, functional form. However, longer intermediate forms can be produced when normal processing is impaired (Condo et al., 2007;

Schmucker et al., 2008), but also by some normal cells. Their functional role remains controversial. Possibly, the two-step cleavage by MPP also has regulatory functions. In humans, frataxin precursor is rapidly cleaved to generate the intermediate form; the second cleavage is slower and limits the overall rate of mature frataxin production in rat liver mitochondria (Cavadini et al., 2000). Probably mitochondrial proteins modulate the processing of the intermediate form.

Fig. 8 Crystal structure of human mature frataxin. From Protein Databank (PDB) accession number 1ekg (Dhe-Paganon et al., 2000).

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Cellular Function of Frataxin

The first data about frataxin function came from the study of frataxin deficient S.

cerevisiae (Δyfh1) cells, which show a severe growth deficit with reduced rate of respiration, loss of mtDNA, high sensitivity to oxidants as hydrogen peroxide and copper, mitochondrial iron accumulation with low cytosolic iron, and hyper-activation of the high affinity iron transport system in the plasma membrane (Fig. 9) (Babcock et al., 1997; Foury et al., 1997; Koutnikova et al., 1997; Wilson et al., 1997). When Yfh1 was reintroduced in the Δyfh1 mutant, iron shifted from mitochondria back to the cytosol (Radisky et al., 1999), suggesting that loss of frataxin prevents iron efflux from mitochondria. Such phenotype indicates that frataxin is important for mitochondrial integrity and function, and that it is involved in mitochondrial iron metabolism.

A role of frataxin in iron homeostasis was subsequently supported by observations in human patients. Iron deposit were observed in the heart of FRDA patients, along with deficiencies of the respiratory chain complexes I, II, and III and of both mitochondrial and cytosolic aconitases (Sanchez-Casis et al., 1976; Lamarche et al., 1993; Rotig et al., 1997). These enzymes all have iron-sulfur clusters (ISCs) in their active sites, which are exquisitely sensitive to reactive oxygen species (ROS). These data thus suggested that the alteration in iron homeostasis caused by frataxin deficiency resulted in increased mitochondrial iron and ROS production by the Fenton reaction, leading to inactivation of ISCs, mtDNA damage and hypersensitivity to oxidative stress (Babcock et al., 1997; Rotig et al., 1997).

However, the idea that oxidative damage caused by iron accumulation in the mitochondria is responsible for the abnormalities observed in FRDA cells (Foury et al., 1999; Rustin et al., 1999; Chen et al., 2000) was subsequently challenged. In yeast, it was shown that preventing mitochondrial iron accumulation through the use of iron-depleted medium did not fully prevent aconitase deficiency (Foury et al., 1999). When mouse models became available, conditional KO mice showed ISC enzyme deficiencies in the frataxin depleted tissues before iron accumulation could be detectable in mitochondria (Dollé et al., 2000; Puccio et al., 2001). These findings led to the hypothesis that frataxin has a function in the biogenesis of ISCs (Muhlenhoff et al., 2002). At the same time, the observation that mitochondrial iron accumulation is a common feature of impaired ISC synthesis of any cause provided an explanation of why this occurs in frataxin

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deficiency. Data accumulated in the following years that further supported this hypothesis. The exact role of frataxin in ISC biogenesis is, however, still unclear and controversial, as is the possibility that it is involved in additional cellular processes in mitochondria and in the cytosol.

One important controversy concerns the functional role in vivo, if any, of iron-containing oligo- and polymers that frataxins can form in vitro in the presence of excess iron, Yfh1 much better than mammalian frataxins (Adamec et al., 2000). This is further discussed below (Fig. 9).

Fig. 9 The model indicates a possible role of frataxin in iron (Fe)–sulphur (S) cluster (ISC) biogenesis.

ISCs are prosthetic groups that are commonly found in various proteins that participate in oxidation–

reduction reactions and catalysis. The mature form is the (4Fe–4S). Frataxin would interact with IscU and IscS involved in ISC biosynthesis. In the proposed model, frataxin deficiency would result in reduced ISC biosynthesis, intra-mitochondrial iron accumulation, and increased oxidative stress (formation of hydroxyl radicals (OH^-) and reactive oxygen species (ROS)). Thus, the reduction of the mitochondrial ISC- containing enzymes aconitase and respiratory chain complexes I-III and, possibly, of other extra- mitochondrial Fe–S proteins, might be both the direct result of reduced ISC biosynthesis and the indirect effect of oxidant-induced damage of the ISCs. Adapted with permission from Taroni and Di Donato, 2004.

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Abnormal Iron Homeostasis and Oxidative Stress Defenses

Fe-S Cluster Assembly

Multiple functional deficits in proteins containing ISCs systematically occur when frataxin is depleted in eukaryotic cells (Anderson et al., 2005; Busi et al., 2006; Rotig et al., 1997; Wilson et al., 1997; Cossée et al., 2000; Long et al., 2008), though these are not obvious in bacteria (Li et al., 1999; Vivas et al., 2006).

ISCs are ensembles of two or more iron atoms bridged by sulfide centers. A number of metalloproteins (iron sulfur proteins, ISPs) in both prokaryotes and eukaryotes contain ISCs, which are essential for their function, structure and stability (Ye and Rouault, 2010). Most biological ISCs are of the [2Fe-2S] type, consisting in two iron atoms bridged by two sulfides and bound to the protein backbone through four cysteines (two cysteines and two histidines in Rieske proteins), or of the [4Fe-4S] type, which has a cubane structure and is bound to the protein backbone through cysteine residues. ISPs are found in different cell compartments and have diverse functions. In mitochondria, ISPs include, among others, several subunits of respiratory complexes I, II and III and a complex I assembly factor (NUBPL), the Krebs cycle enzyme aconitase, a ferredoxin, ferrochelatase (the enzyme that inserts Fe into protoporphyrin IX to form heme), the molybdenum cofactor synthesis enzyme MOCS1A, the membrane associate protein of unknown function MitoNEET. ISPs are present also in the cytosol and in the nucleus, where they are involved in different biological processes, including control of iron metabolism (IRP1), various metabolic pathways, signaling pathways, and DNA repair. Such a wide distribution and variety of function of ISPs clearly show how their importance cannot be overstated.

Bacteria have several ISC assembly systems, the one encoded by the Isc operon in E. coli being the homolog of the eukaryotic system. It is interesting that the gene encoding bacterial frataxin, CyaY, is not in the Isc operon, nor in any other ISC synthesis operon, suggesting separate regulation. In simple eukaryotes as yeast, ISC biogenesis takes place in the mitochondria. In higher eukaryotes, ISC biogenesis takes place in mitochondria and in the cytosol, but cytosolic ISC synthesis appears to depend on the integrity of the mitochondrial

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assembly system and on the export of a still unknown component from the mitochondria through a carrier known as ABCB7 in mammals (homologus to ATM1 in yeast). Mitochondrial ISC biogenesis in higher eukaryotes is carried out by assembly machinery composed by many factors (Ye and Rouault, 2010). The list is probably not yet complete and the function of all components is not known with certainty. The main ISC biogenesis factors, named according to the human nomenclature, are: the scaffold protein ISCU, where the nascent ISC is assembled; the sulfur donor NFS1, a cysteine desulfurase enzyme (EC 2.8.1.7) which catalyzes the reaction that converts L-cysteine into alanine and a highly reactive persulfide group; the small protein ISD11, which forms a complex with NSF1 and is needed for its function; the ferredoxin FDX1 and the ferredoxin reductase FDXR, which provide reducing equivalents; and the putative alternative scaffold NFU.

Cluster transfer from ISCU to the apoproteins to form holo-ISC proteins requires a further set of factors: the chaperones HSCB and mortalin; the glutaredoxin GLRX5; the GrpE-L1/2 nucleotide exchange factor; and a set of specific assembly factors for certain ISC proteins, including Ind1 for complex I, and ISCA1, ISCA2, IBA57 for SAM-dependent proteins and aconitase.

Many data now support a role of frataxin in mitochondrial ISC biogenesis. The similarity between the phenotype of frataxin deficient yeast and yeast depleted of other ISC biogenesis factors has been the first indirect evidence in this regard. The observation of biochemical and genetic interactions with the highly conserved NFS1/ISCU complex, the two main components central to the ISC assembly machinery (Gerber, Muhlenhoff et al. 2003; Yoon and Cowan 2003;

Ramazzotti, Vanmansart et al. 2004) has further supported this hypothesis. Yet, while these data have received independent confirmation, many questions whose answer would be crucial for our understanding of the frataxin function remain unresolved.

One important point, even preliminary to other functional investigations, is the identification of the functionally relevant form of frataxin and the confirmation of its exclusive subcellular localization in mitochondria (Schmucker and Puccio 2010). Concerning the first point, m81-FXN is clearly the most abundant isoform, retaining all structural features needed to interact with the ISC assembly factors. Controversy concerns m42-FXN, considered by some just a processing intermediate, but by others a functionally relevant isoform because, through

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interactions involving the extra aminoacids at the N-terminus, it is able to oligomerize. Clearly, this is crucial to establish if oligomerization is an accidental, dispensable property only shown by incompletely processed frataxin, or it is an important property for the full functionality of the protein.

Another controversy concerns the presence and functional relevance of a cytosolic pool of frataxin. Some observations suggested that frataxin may also be involved in the assembly of cytosolic and nuclear ISC, at least in some cell types (Acquaviva et al., 2005), others proposed a specific pro-survival function of cytolsolic frataxin (Condo et al., 2006). Data from our laboratory (Jiralerspong, MSc thesis, McGill University and unpublished data), obtained in fibroblasts, lymphoblastoid cell lines and various mouse tissues, go in the direction of an exclusive intramitochondrial localization of mature frataxin in all the explored cell types, with only the unprocessed precursor being present in very low amounts in the cytosol, probably surronding the interactions of frataxin in the mitochondrion and its primary function.

Calorimetric studies on human frataxin have suggested a direct interaction with ISCU (Yoon and Cowan, 2003). Interaction requires iron and a single iron center is sufficient to reach nanomolar affinities (Huang, Dizin et al., 2008). In contrast, prokaryotic frataxin (CyaY) could not be shown to interact with the isolated ISCU homolog (IscU), but rather with the NSF1 homolog IscS (Layer et al., 2006; Adinolfi et al., 2009). In yeast, both the frataxin (Yfh1) and the the NSF1-ISD11 homologs (Nfs1-Isd11) can directly bind to the scaffold Isu1 (the ISCU homolog) in an iron dependent way (Gerber, Muhlenhoff et al. 2003) and the interaction seems to require Yfh1 oligomerization (Li, Gakh et al. 2009). But the most crucial question is why these interactions take place, or, ultimately, what is the primary function of frataxin. This point remains highly controversial. Frataxin has been suggested to act either as a chaperone which delivers iron to the NSF1/ISCU complex (Yoon and Cowan 2003) or as a scavenger (Cavadini, O'Neill et al. 2002) which, through formation of large iron loaded assemblies, prevents iron excesses in mitochondria while still keeping iron biologically available for being delivered to key acceptors. These hypotheses rely on the assumption that frataxin has a functionally relevant ability to bind iron, a point that is further discussed in the next section. More recently, by using an in vitro reconstructed enzymatic ISC synthesis complex composed of IscU and IscS, it was shown that CyaY is able to inhibit ISC formation on the scaffold protein IscU. This occurs without inhibiting

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desulfurase enzymatic activity and suggests a role for frataxin as a gatekeeper of ISC formation (Adinolfi, Iannuzzi et al. 2009). Clearly, these results have to be reconciled with the in vivo finding of decreased ISC biogenesis in frataxin deficiency. In any case, frataxin should play an important but not essential role in this process, since ISC biogenesis can occur in the absence of frataxin, even though at a reduced level (Duby et al., 2002).

Other functions have also been proposed for frataxin. In particular, it has been reported that it can act as an iron chaperone in converting the oxidative damaged (3Fe-4S) cluster into the active (4Fe-4S) cluster of aconitase (Bulteau et al., 2004). Interaction of frataxin with aconitase, in the presence of citrate, would protect the cluster from oxidation, reducing the risk of enzyme inactivation (Bulteau et al., 2004). However, this interaction could not be reproduced in other laboratories (H. Puccio, A. Pastore, personal communications).

Iron Homeostasis

One of the most striking findings in Yfh1 yeast cells was a marked iron accumulation in mitochondria accompanied by a relative depletion in the cytosol (Babcock et al., 1997; Foury et al., 1997; Wilson et al., 1997). The high-affinity iron transport system, whose expression is normally suppressed in iron replete yeast cells, was markedly overxpressed in in Yfh1 as a result of the activation of the iron-sensitive transcription factor Aft1 (Babcock et al., 1997). In yeast, Aft1 senses cytosolic iron levels and activates transcription of the high affinity uptake system when cytosolic iron is low. A severe impairment of iron homeostasis therefore takes place in this model of frataxin deficiency, which behaves as if iron depleted when in fact iron is in excess and trapped inside mitochondria. A simple way of interpreting this abnormality is to assume that the inability of mitochondria to efficiently utilize iron for critical biosynthetic processes, in particular ISC synthesis, on the one hand leads to the participation of this metal in redox chemistries and eventual precipitation within the organelle, and on the other hand generates a signal to the iron- sensing mechanism of the cell that there is a need of iron to synthesisze more ISCs. The nature of the latter signal is still unknown, but it may act through the suppression of iron export out of mitochondria when too few ISCs are made, resulting in low cytosolic iron (Pandolfo, 2002). This model is supported by the observation that all gene knockouts leading to defective ISC synthesis

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in yeast cause the same alteration of iron homeostasis as observed in Yfh1 (Philpott et al., 2008;

Rouault et al., 2008).

The situation in upper eukaryotes is more complex. Iron homeostasis is not directly regulated by an iron-sensitive transcription factor as in yeast, but it mostly takes place at the post- transcriptional level through the binding of regulatory proteins to a specific sequence, the Iron Responsive Element (IRE) in the 5‘ or 3‘ untranslated regions of specific mRNAs (Muckenthaler et al., 2008). Two cytosolic proteins, IRP1 and IRP2, bind IREs when iron is low in their compartment. IRP1 is a cytosolic aconitase containing a cubane (4Fe-4S) cluster when iron levels are high. Its cluster loses one iron atom when iron levels decrease, leading to loss of aconitase activity and stimulation of IRE-binding activity. IRP2 has a similar structure to IRP1, but no enzymatic activity and is degraded through the ubiquitine-proteasome system when iron levels ar high. When iron is low, IRP2 escapes proteolytic degradation and binds to IREs. IRP2 is thought to be the main regulator of iron metabolism in most tissues. IRP binding to a 5‘ IRE prevents mRNA translation, binding to a 3‘ IRE stabilizes the mRNA and enhances translation. Proteins involved in iron uptake, as the transferrin receptor (TfR) have 3‘ IREs in the corresponding mRNAs, proteins involved in iron storage (as ferritin) or utilization have a 5‘ IRE in their mRNAs. If the alteration of iron homeostasis when frataxin is low or absent (as in conditional KO models) in mammalian cells is the same as in yeast, one would thus expect increased IRP activation. Accordingly, there are reports of IRP1 activation in FRDA patients‘ fibroblasts (Li et al., 2008) and in conditional KO mouse models. In addition, the levels of several IRP-regulated proteins have been found to be changed in FRDA patients‘ cells as expected when cytosolic iron is low and IRPs are activated. These include TfR1 and the mitochondrial iron importer mitoferrin-2, which are increased, and the cell membrane iron exporter ferroportin 1, which is decreased (Huang et al., 2009). Furthermore, inactivation of human ISD11, a component of the mitochondrial ISC biogenesis machinery, also results in disruption of iron homeostasis through increased binding activity of IRP1 and increased protein levels of IRP2 (Shi et al., 2009), supporting the concept that a similar alteration of iron homeostasis follows impaired ISC biogenesis in yeast and in upper eukaryotes.

Mitochondrial iron overload has, however, been difficult to prove in cells from FRDA patients and in conditional KO mouse models. No significant iron deposits were seen in the