Human Molecular Genetics, 2007, Vol. 16, No. 22
doi:10.1093/hmg/ddm163
Advance Access published on June 27, 2007
2651–2658
Frataxin is essential for extramitochondrial Fe– S
cluster proteins in mammalian tissues
Alain Martelli1{, Marie Wattenhofer-Donzé1,5{, Stéphane Schmucker1,4, Samuel Bouvet1,
Laurence Reutenauer1,3 and Hélène Puccio1,2,3,4,5*
1
IGBMC (Institut de Génétique et de Biologie Moléculaire et Cellulaire), 1 rue Laurent Fries BP 10142,
Illkirch F-67400, France, 2Inserm, U596, Illkirch F-67400, France, 3CNRS, UMR7104, Illkirch F-67400,
France, 4Université Louis Pasteur, Strasbourg F-67000, France and 5Collège de France,
Chaire de Génétique Humaine, Illkirch F-67400, France
Received May 28, 2007; Revised and accepted June 21, 2007
Friedreich ataxia, the most common recessive ataxia, is caused by the deficiency of the mitochondrial protein
frataxin (Fxn), an iron chaperone involved in the assembly of Fe – S clusters (ISC). In yeast, mitochondria play
a central role for all Fe –S proteins, independently of their subcellular localization. In mammalian cells, this
central role of mitochondria remains controversial as an independent cytosolic ISC assembly machinery has
been suggested. In the present work, we show that three extramitochondrial Fe – S proteins (xanthine oxidoreductase, glutamine phosphoribosylpyrophosphate amidotransferase and Nth1) are affected in Fxn-deleted
mouse tissues. Furthermore, we show that Fxn is strictly localized to the mitochondria, excluding the presence of a cytosolic pool of Fxn in normal adult tissues. Together, these results demonstrate that in mammals, Fxn and mitochondria play a cardinal role in the maturation of extramitochondrial Fe – S proteins.
The Fe – S scaffold protein IscU progressively decreases in Fxn-deleted tissues, further contributing to the
impairment of Fe – S proteins. These results thus provide new cellular pathways that may contribute to molecular mechanisms of the disease.
INTRODUCTION
Friedreich ataxia (FRDA), the most common hereditary
ataxia, is a neurodegenerative disease characterized by
degeneration of the large sensory neurons and spinocerebellar tracts, cardiomyopathy and an increased incidence of diabetes (1). FRDA is caused by a severe deficiency in frataxin
(Fxn) protein, a highly conserved mitochondrial protein, as a
result of a GAA repeat expansion in the first intron (2).
Although the cellular function of Fxn and its yeast
homolog, Yfh1, is still a matter of debate, a role of Fxn as
an iron donor in Fe – S cluster (ISC) biogenesis is widely
accepted. Indeed, Fxn deficiency both in tissues and
cells leads to severe alteration of Fe – S enzymes activities
(3 – 5). Reconstitutional and in vivo studies demonstrated
that yeast Fxn, Yfh1, is required, although not essential,
for ISC biosynthesis (6,7). Yeast Fxn and its bacterial
homolog were shown to form a physical complex with the
central Fe – S cluster scaffold protein, Isu1/IscU, and the
cysteine desulfurase, Nfs1 (8 – 10). Finally, in vitro works
have demonstrated direct iron transfer from Fxn to the
human IscU protein (11).
Biosynthesis and maturation of Fe– S proteins in eukaryotes
is a complex process mediated by a set of highly conserved
components that are located in the mitochondria and cytosol
(12). In yeast, the de novo synthesis of a nascent ISC is
mediated by Isu1/2, the scaffold protein that binds ferrous
iron, and Nfs1, the cysteine desulfurase that provides sulfur
(5). Yfh1 has been suggested to be the iron donor in this
process. In yeast, these three central proteins (Isu1/2, Nfs1
and Yfh1) are found only in mitochondria. Once synthesized
in this compartment, ISCs are incorporated into mitochondrial
apoproteins by a highly regulated chaperone-mediated process
(5,13). For ISC assembly in the cytosol and the nucleus, a
component derived from the de novo ISC synthesis is exported
by the mitochondrial ABC transporter Atm1p (14). The maturation of cytosolic and nuclear Fe– S proteins therefore
requires the close cooperation of the cytosolic assembly
* To whom correspondence should be addressed. Tel: þ33 388653264; Fax: þ33 388653246; Email: [email protected]
{The authors wish it to be known that, in their opinion, the first two authors should be regarded as joint First Authors.
# The Author 2007. Published by Oxford University Press. All rights reserved.
For Permissions, please email: [email protected]
2652
Human Molecular Genetics, 2007, Vol. 16, No. 22
machinery, the mitochondrial de novo synthesis machinery
and the ISC export machinery.
In mammalian cells, the central role of mitochondria in ISC
biogenesis for extramitochondrial proteins remains controversial. Indeed, the existence of an independent cytosolic machinery was proposed after the identification of cytosolic isoforms
of both IscU (IscU1– p15) and Nfs1, which were shown to
promote de novo ISC formation in vitro (15). The possibility
that de novo ISC biogenesis also occurs in cytosol is further
reinforced by the recent identification of a pool of cytosolic
Fxn in cultured cells (16 – 18). However, the recent silencing
of the mitochondrial Nfs1 in murine macrophages or HeLa
cells revealed that the protein was necessary not only for mitochondrial Fe– S proteins, but also for cytosolic Fe – S proteins
such as xanthine oxidoreductase (XOR) or iron regulatory
protein-1 (IRP-1) (19,20). Furthermore, deletion of ABCb7,
the mammalian Atm1p homolog, in mouse liver leads to a
decrease in XOR activity, suggesting that the mitochondrial
ISC export machinery is also necessary for the assembly of
cytosolic Fe– S protein in mammalian tissues (21). Together,
these results suggest that a sulfur moiety necessary for extramitochondrial ISC biosynthesis is most likely provided by the
mitochondrial Nfs1 and transported to the cytosol via Abcb7.
However, the source of iron for Fe– S protein assembly in the
cytosol is unknown.
We have previously shown, using conditional mouse model
of FRDA, that Fxn deletion in cardiac tissue (MCK model)
leads to a drastic impairment of mitochondrial Fe– S
enzymes which precedes any detectable pathological and
functional changes including the mitochondrial iron accumulation (4). Furthermore, the cytosolic aconitase/IRP-1
balance was affected, suggesting that the mitochondrial ISC
machinery is essential for cytosolic Fe– S proteins (22).
IRP-1 activation was also observed in HeLa cells that were
depleted for Fxn by RNAi (23). However, the intrinsic link
between IRP-1 and iron metabolism prevents a clear interpretation. Here, we show that Fxn deletion in mammalian tissues
affects three different extramitochondrial enzymes that are
dependent on ISC biosynthesis for their activity or maturation,
but that are not otherwise linked to iron metabolism. Furthermore, we show the absence of a pool of cytosolic Fxn, providing evidence that the mitochondrial protein is necessary for
extramitochondrial ISC biosynthesis, thus further supporting
the cardinal role of mammalian mitochondria for ISC proteins.
RESULTS
Frataxin is necessary for cytosolic ISC protein integrity
Xanthine oxidoreductase, a cytosolic enzyme containing two
[2Fe – 2S] clusters involved in electron transfer, catalyzes the
degradation of xanthine and hypoxanthine to yield uric acid
(24). XOR activity was initially measured in Fxn-deleted
cardiac tissues of the MCK mouse model, and only a late
reduction of XOR activity was observed (data not shown).
However, as heart samples showed low level of XOR activity
(,2 mmol/min per microgram) and no detectable protein (data
not shown), the heart tissue is probably not appropriate to
assess the effect of Fxn deletion on XOR activity. XOR
activity was therefore measured in liver samples deleted for
Figure 1. Xanthine oxidoreductase (XOR) activity in liver samples from the
ALB mice. (A) XOR activity, expressed as percentage of the control, was
measured in control (n 3) and mutant (n 3) mice. (B) mRNA levels of
XOR were determined by quantitative RT–PCR in control (n ¼ 3) and
mutant (n ¼ 3) mice; representative western blot analysis with an anti-XOR
antibody was carried out on the liver extracts used for the activity measurements. P , 0.005.
Fxn (ALB mutant; Supplementary Material, Fig. S1). A significant and drastic decrease in XOR activity was observed
in the liver of mutant mice compared with control, both at 2
and 4 weeks (Fig. 1A). XOR mRNA level was not transcriptionally altered in mutant mice (Fig. 1B). Although a slight
decrease in XOR protein level was seen in mutant mice
(Fig. 1B), it is not sufficient to account for the strong decrease
in activity. Isocitrate dehydrogenase, a mitochondrial Fe –S
independent enzyme, showed no significant change in activity
(data not shown).
Glutamine phosphoribosylpyrophosphate amidotransferase
(GPAT), a key enzyme of the purine biosynthesis, is synthesized as an inactive precursor and requires the incorporation
of a [4Fe – 4S] cluster for the cleavage of the 11 N-terminal
residues propeptide to generate the active enzyme (25). Sitedirected mutagenesis studies of the avian glutamine phosphoribosylpyrophosphate amidotransferase (GPAT) in HeLa cells
demonstrated that prevention of the ISC incorporation leads
to an inactive unprocessed form that accumulates (25). To
assess the maturation of GPAT in Fxn-deleted cardiac tissues,
a polyclonal antibody was generated to detect the processed
and unprocessed forms of murine GPAT (Fig. 2A, m-GPAT
and GPAT –C1F). The signal for the processed form of
GPAT was clearly diminished in mutant mice compared with
control at 5, 7 and 10 weeks (Fig. 2A). The decrease in the
processed GPAT was not due to transcriptional regulation as
real-time polymerase chain reaction (RT– PCR) showed no
difference at 5 and 7 weeks (Fig. 2B). A significant mRNA
decrease was observed in mutant mice at 10 weeks, as previously seen for many other genes (22), which can be attributed
Human Molecular Genetics, 2007, Vol. 16, No. 22
2653
Figure 2. Glutamine phosphoribosylpyrophosphate amidotransferase (GPAT) maturation in cardiac samples from the MCK mice. (A) Western blot using a
GPAT-specific antibody of total heart extracts of control (C) and mutant (M) mice. m-GPAT corresponds to a mature GPAT construct transfected into
COS-1 cells, and GPAT– C1F corresponds to a mutated construct for the expression of the precursor. Estimation of mature GPAT level was done using vinculin
signal as loading control. (B) mRNA levels of GPAT in control (n ¼ 3) and mutant (n ¼ 3) MCK mice, determined by quantitative RT–PCR. P , 0.01.
to a general transcriptional impairment because of the late
disease stage rather than a specific downregulation. Furthermore, a band comigrating with the unprocessed form of
GPAT can already be detected in mutant mice at 5 weeks,
and clearly increases at 7 and 10 weeks, further suggesting
that the maturation step is affected (Fig. 2A). GPAT maturation
was also perturbed in liver of the ALB mouse model with a
strong decrease in the mature form already observed at 2
weeks (Supplementary Material, Fig. S2).
Frataxin is necessary for nuclear ISC protein integrity
To determine whether the absence of Fxn also leads to a deficit
in a nuclear Fe– S protein, we evaluated the activity of Nth1,
the mammalian endonuclease III homolog, in cardiac tissues
deleted for Fxn. Nth1, a [4Fe –4S] cluster protein, is a glycosylase/AP-lyase involved in the base excision repair of oxidized bases such as thymine glycol (Tg) (26). DNA
glycosylase activity was assessed using a synthetic oligonucleotide containing Tg. Nth1 cleaves the strand containing
the lesion via a b-elimination mechanism and its activity
can be distinguished from Neil1, a second glycosylase/
AP-lyase, which cleaves via a b,d-elimination process (27)
(Fig. 3A). A significant decrease in Nth1 activity was observed
in mutant mice at 5 weeks, which was even more pronounced
at 7 and 10 weeks, reaching 50% of control activity
(Fig. 3B). Neil1 activity showed no difference at 5 weeks,
but was increased at 7 and 10 weeks in mutant mice,
suggesting a compensatory effect triggered by the Nth1
impairment. The change in activity was not due to a transcriptional regulation, as RT – PCR showed no difference between
control and mutant mice at 5 and 7 weeks (Fig. 3C). Similar
to other transcripts, a significant decrease in both mRNA in
mutant mice at 10 weeks was observed.
IscU is decreased in Fxn-deleted tissues
We previously showed that IscU mRNA level in the MCK
mutant mice was only decreased at 10 weeks compared with
control (28). However, western blot analysis on total protein
extracts shows a progressive decrease in IscU protein in
Fxn-deficient cardiac samples (Fig. 4A). Interestingly, prohibitin (PHB), a mitochondrial protein, was seen increased, in
agreement with the mitochondrial proliferation as reported
previously in these animals (4). The decrease in IscU was
significant at 5 weeks, reaching 50% of the control level by
7 weeks (Fig. 4B). This decrease was also seen when quantified against tubulin (data not shown). In contrast, Nfs1 was not
downregulated in Fxn-deficient tissues (Fig. 4A and B). IscU
protein level was also decreased at 4 weeks in mutant mice
of the ALB model, while it appeared unchanged at 2 weeks
(Supplementary Material, Fig. S3).
No cytosolic Fxn is detected in adult mouse tissues
In addition to its well-established mitochondrial localization
(29), Fxn has recently been proposed to be cytosolic (17,18).
This possible pool of cytosolic Fxn could participate in the
maintenance and/or biosynthesis of extramitochondrial Fe –S
proteins. However, fractionation experiments using heart,
liver and cerebellum of control adult mice show a strict mitochondrial localization of Fxn (Fig. 5), even though the Fxn
antibody is capable of detecting ,1 ng of protein (unpublished data). No signal for Fxn that cannot be accounted by
slight mitochondrial contamination was detected in the cytosolic fractions, indicating that a cytosolic pool of Fxn is not
present in the mouse tissues used in the present study. Furthermore, no cytosolic isoform of IscU (IscU1– p15) could
be detected (Fig. 5), suggesting that the extramitochondrial
ISC deficit was directly due to a deficit in the mitochondrial
machinery.
DISCUSSION
The present report provides the first direct evidence in
mammals that Fxn deletion affects the assembly of both cytosolic and nuclear Fe– S proteins, which are not directly
involved in iron regulation. Through different conditional
mouse models, we have shown that Fxn deletion results in a
significant decrease in the specific activities of XOR and
Nth1, two enzymes which required proper ISC assembly for
their activity, as well as incomplete maturation of GPAT, an
2654
Human Molecular Genetics, 2007, Vol. 16, No. 22
Figure 3. Thymine glycol nicking activity of crude nuclear extracts of MCK mice. (A) By-products generated by the nicking activity of Nth1 and Neil1,
observed on a 20% denaturing polyacrylamide gel using extracts of 7-week-old mice. The first lane corresponds to the duplex without any addition of
extract. 1, 2, 3 and 4 indicate the different products of the reaction as illustrated on the scheme. Nth1 product (2) and Neil1 product (3) were quantified independently to determine the activity of each enzyme. (B) Tg nicking activity, expressed as % of control activity, was determined in control (n ¼ 3) and mutant
(n ¼ 3) mice. (C) mRNA levels were determined by quantitative RT– PCR in control (n ¼ 3) and mutant (n ¼ 3) mice. P , 0.05; P , 0.01.
Figure 4. IscU and Nfs1 protein levels in total heart extracts of MCK mice. (A) Representative western blot with control (C) or mutant (M) mice extracts using
anti-IscU, anti-Nfs1 and anti-PHB antibodies. (B) Protein levels were quantified for three mice of each genotype and each age, and expressed as % of control
level using the ratio Iscu:PHB or Nfs1:PHB. P , 0.01.
enzyme which requires the insertion of an ISC for its processing. No change was detectable in the activities of proteins
that do not contain ISC, further pointing to the specificity of
action of Fxn on Fe – S containing enzymes.
Oxidative stress is a known factor of instability of some
ISC, specifically the ones that are exposed to solvent such as
the ISC of aconitases (30) or radical S-adenosyl-L -methionine
enzymes (31). However, we have previously shown the
absence of detectable protein or lipid oxidative damage in
the heart of the conditional cardiac model (22). Therefore,
rather than a destabilization of the specific ISC by reactive
oxygen species (ROS), we suggest that the deficiency is
related to an ISC biosynthesis deficiency as a direct consequence of the absence of Fxn.
Xanthine oxidoreductase activity was drastically decreased
in liver deleted for Fxn (Fig. 1A), whereas its basal activity
in control heart tissues is too close to the detection limit for
reliable measurements. It is worth to note that XOR also
Human Molecular Genetics, 2007, Vol. 16, No. 22
Figure 5. Fractionation of control adult mouse tissues. Mitochondrial (m) and
cytosolic (c) fractions were analyzed by western blot: GAPDH and b-tubuline
are markers of the cytosolic compartment; PHB and Mn-SOD are markers of
the mitochondrial compartment; IscU and frataxin (Fxn).
requires a molybdenum cofactor for its activity. The first step
of molybdenum biosynthesis is performed by MOCS1A, an
Fe –S enzyme (32) that may also be affected by Fxn deletion.
Thus, the drastic decrease in XOR activity could be a cumulative effect of both the reduced molybdenum cofactor biosynthesis as well as its own ISC biosynthesis.
Glutamine phosphoribosylpyrophosphate amidotransferase
maturation process was clearly affected in both heart and
liver, as the level of mature GPAT decreased progressively
(Fig. 2A, Supplementary Material, Fig. S2). Interestingly,
the precursor accumulation does not completely parallel the
decrease in the mature form (Fig. 2A), suggesting that either
the unprocessed form gets degraded, or the precursor is less
soluble as previously suggested for the avian GPAT (25).
The activity of the nuclear DNA repair enzyme Nth1 was
significantly affected in heart tissue, and was brought to
50% of residual activity compared with control mice at 7
and 10 weeks (Fig. 3B). Neil1 activity was shown to increase
significantly at 7 and 10 weeks (Fig. 3B), although the
mRNA level was not modified (Fig. 3C). We exclude the
possibility that Neil1 upregulation is due to a direct competition, as the experimental conditions are in excess substrate.
It is possible that the posttranscriptional regulation of Neil1
is to compensate the loss of Nth1 activity, although this
has not previously been reported in the Nth1 knockout
mouse (27).
Furthermore, we showed that no Fxn can be detected by
western blot analysis in the cytosolic fractions of heart, liver
or cerebellum of normal adult mice (Fig. 5), even though
the Fxn antibody used has an extreme sensibility. We thus
exclude the existence of a cytosolic pool of Fxn that may participate directly in cytosolic ISC biosynthesis machinery in
mammalian tissues. These results provide evidence that in
addition to the mitochondrial sulfur moiety provided by
Nfs1, either a mitochondrial iron moiety provided by Fxn,
or a substrate requiring the ISC de novo mitochondrial biosynthesis in which Fxn participates, is required for extramitochondrial ISC protein assembly. Furthermore, we could not detect
the previously described cytosolic IscU – p15 (33), suggesting
that this isoform is not required for the biosynthesis of extramitochondrial ISC in normal conditions. Our results reinforce
2655
the notion that, in mammals, mitochondria play a cardinal role
in the ISC biogenesis.
Our results do not however exclude the existence of a pool
of extramitochondrial Fxn and IscU under certain conditions.
It is interesting to note that overexpression of cytosolic Fxn
was able to rescue FRDA cells against oxidative stress
insults (17), and that the cytosolic form of IscU was shown
to be required for the regeneration of IRP1 after ISC
damage by H2O2 or iron chelator treatment (34). This suggests
that a pool of cytosolic Fxn might exist for the regeneration of
some ISC damaged by ROS.
Interestingly, a progressive downregulation of IscU protein
in Fxn-deficient tissues was observed (Fig. 4), similar to
what has been reported in FRDA patient fibroblasts and lymphoblasts (35). Although IscU downregulation could by itself
be the primary cause of ISC biosynthesis deficiency, the Fe –
S protein impairments in heart and liver samples occur prior
to IscU downregulation. Indeed, in Fxn-deficient liver, XOR
activity is diminished at 2 weeks (Fig. 1A) whereas IscU
protein level is normal (Supplementary Material, Fig. S3).
Moreover, it has previously been shown by RNAi experiments that a threshold of at least two-fold reduction in
IscU must be passed in order to trigger an ISC deficit in
HeLa cells (34). We thus hypothesize that partial ISC biogenesis impairment is a primary effect of Fxn deletion, but
the consequent progressive IscU decrease further contributes,
in a time-dependent manner, to ISC biogenesis deficiency.
The mechanism of the posttranscriptional regulation of
IscU is unknown, but the fact that Fxn has been shown to
form a physical complex with Isu1/IscU (8,9,11) probably
plays an essential role.
Finally, it will be interesting to determine whether some
of the newly identified Fe – S-dependent pathways play a
role in the pathogenesis of FRDA, although primarily considered as a mitochondrial disease. In the context of neurodegenerative disorders, the effect on DNA repair pathways is
of particular interest. Indeed, in addition to Nth1, two
related helicases, XPD and FancJ, involved in the nucleotide
excision repair, and Fanconi anemia repair pathways,
respectively, have recently been identified as Fe – S proteins
(36). This new link between DNA repair, transcription and
Fe – S proteins opens some intriguing possibilities for
FRDA pathology. It will be interesting to determine
whether part of the progressive neurological disease might
be accounted by subtle defects in several DNA repair pathways that would lead to accumulation of DNA damages over
several years. As DNA repair is one of the other common
pathways affected in recessive ataxia (37), this may be the
molecular bridge between this heterogeneous group of
disorders.
In conclusion, we have shown that Fxn is required not only
for the biogenesis of mitochondrial containing ISC protein, but
also for the biogenesis of cytoplasmic and nuclear ISC proteins. The eventual presence of an independent cytosolic ISC
machinery is not sufficient to counteract the mitochondrial
dysfunction triggered by the Fxn deletion, a mitochondrial
protein with no cytosolic form in the tissues. In the context
of FRDA, the identification of ISC-dependent pathways that
may be affected by Fxn deficiency opens new venues to understand the pathology of the disease.
2656
Human Molecular Genetics, 2007, Vol. 16, No. 22
MATERIALS AND METHODS
Animals
All methods employed in this work are in accordance with the
Guide for the Care and Use of Laboratory Animals published
by the US National Institutes of Health (NIH Publications No.
85-23, revised 1996). Animal care and MCK mutant generation were as previously described (4). A liver-specific Fxn
deletion was obtained by crossing DA16 conditional Fxn
allele with a transgenic mouse expressing the CRE recombinase under the control of the albumin promoter (38) (unpublished mouse model).
Tg nicking activity
Thymine glycol nicking activity was measured as described
previously (39), with minor adaptation. Frozen hearts were
homogenized in buffer A (10 mM HEPES – KOH, pH 7.7,
0.5 mM MgCl2, 10 mM KCl, 1 mM DTT) using a polytron
(Ultra-Turrax T 25 basic) and then centrifuged at 2000g,
48C for 10 min. The pellet was resuspended in buffer B (20
mM HEPES– KOH, pH 7.7, 0.5 mM MgCl2, 420 mM NaCl,
0.2 mM EDTA, 25% glycerol, 1 mM DTT) and gently stirred
at 48C for 15– 20 min to allow an efficient nuclear lysis.
The suspension was centrifuged at 14 000g, 48C for 10 min
and the supernatant (nuclear extract) was dialyzed against
buffer C (40 mM HEPES – KOH, pH 7.7, 50 mM KCl, 2 mM
DTT) during 3 h. After determining the protein concentration
by Bradford assay, nuclear extracts were immediately used for
Tg nicking activity using a 34-mer synthetic oligonucleotide
containing the oxidized base Tg: 50 -GGC TTC ATC GTT
GTC (Tg)CA GAC CTG GTG GAT ACC-30 (kind gift from
Dr Gasparutto, Grenoble). Labeled oligonucleotide containing
Tg was annealed with two equivalents of its non-oxidized
complementary strand. Two microliters of crude nuclear
extract were incubated with 0.1 pmol of labeled DNA
duplex in buffer D (40 mM HEPES –KOH, pH 7.7, 100 mM
KCl, 0.5 mM EDTA, 0.5 mM DTT, 0.2 mg/ml bovine serum
albumin) for 1 h at 378C. The reaction was stopped by the
addition of sodium dodecyl sulfate (SDS; 0.5%) and proteinase K (0.8 mg/ml) and further incubated for 15 min at 558C.
After phenol –chloroform extraction, one volume of loading
buffer (formamide 98%, 10 mM EDTA, xylene cyanol
0.01% and bromophenol blue 0.01%) was added to the
samples. Samples were separated on a 20% acrylamide/
bisacrylamide (19:1) denaturing gel in TBE buffer. Gels
were exposed to a phosphoscreen that was subsequently analyzed using a Typhoon apparatus (GE Healthcare, UK).
Bands corresponding to cleaved and uncleaved products
were quantified using ImageQuant TL software (GE Healthcare, UK). Reported activities correspond to the percentage
of Tg cleavage per microgram total protein per hour.
Xanthine oxidoreductase activity
The method for XOR activity measurement was adapted from
Beckman et al. (40). Frozen heart or liver tissues were homogenized in sucrose buffer, pH 7.4 (0.25 M sucrose, 10 mM
DTT, 0.2 mM PMSF, 0.1 mM EDTA, 50 mM KH2PO4,
pH 7.4) using a polytron (Ultra-Turrax T 25 basic) and
centrifuged at 15 000g, 48C to pellet tissues debris. Clear
supernatants, containing total protein extract, were used for
the measurements of XOR activity on a PTI (Photon Technology International TimeMaster; Bioritech) spectrofluorometer
(excitation, 345 nm; emission, 390 nm) as followed: homogenates were diluted in 1 ml of phosphate buffer, pH 7.4 (50 mM
KH2PO4, pH 7.4, 0.1 mM EDTA) and warmed to 378C in
the cuvette for further baseline drift measurement. Twenty
microliter of 1 mM pterin (Fluka) was then added to measure
the activity of xanthine oxidase. Once a linear rate was
obtained, 20 ml of 1 mM methylene blue (Riedel-de Haën)
was added as an electron acceptor to measure the combined
xanthine oxidase and xanthine dehydrogenase activities. The
reaction was then inhibited by the addition of 20 ml of 3.5
mM allopurinol (Sigma). A 0.5 mM final concentration of isoxanthopterin (Fluka) was added and the immediate fluorescence increase was used as an internal standard for
activity calculation. Activities were calculated as follows:
A ¼ {DF [I]/FI} 0.001 Vc/(Vs P), where A is the
enzyme activity (in mmol/min/mg total protein); DF, the
change in fluorescence per minute; [I], the concentration of
isoxanthopterin added; FI, the immediate increase in the fluorescence produced by the addition of isoxanthopterin; Vc, the
cuvette volume (in mL); Vs, the volume of sample added to
the cuvette (in mL); and P, the protein concentration of the
sample (in mg).
Plasmids
Full length GPAT was PCR amplified using EST clone
IMAGp998H2314089Q1 (F: 50 -CGGCGGGAATTCCCGAG
GCGCCATGGAGC-30 ; R: 50 -CGGCGGCTCGAGCTACCA
TTCCAGCTCCACAG-30 ) and cloned into pcDNA3.1 (Invitrogen) to obtain pcDNA3 – GPAT. A truncated GPAT corresponding to the mature form was generated by PCR
amplification (F: 50 - CGGCGGGAATTCCCACCATGTGTG
GTGTGTTTGGGTGCATC-30 ; R: 50 -CGGCGGCTCGAG
CTACCATTCCAGCTCCACAG-30 ) and subcloned into
pcDNA3.1 to obtain pcDNA – m-GPAT. pcDNA3 – GPAT –
C1F, a noncleavable GPAT precursor bearing a mutation at
Cys-12, was generated by directed mutagenesis (F: 50 -GATC
CGCGAGGAATTTGGTGTGTTTGGGT-30 ; R: 50 -ACCCAA
ACACACCAAATTCCTCGCGGATC-30 ). The mature form
of the mouse mitochondrial IscU was amplified by nested
PCR (F: 50 -CCGCCGGAATTCCACAAGAAGGTTGTGG-30 ;
R:
50 -CCGCCGCTCGAGTCACTGCTTCTCTGGCTC-30 )
using mouse heart cDNA and cloned into pGEX4T1 to give
pGEX – IscU. All constructs were verified by DNA
sequencing.
Recombinant protein expression
Recombinant GPAT proteins were generated by transient
transfection of COS-1 cells using Fugene Reagent 6 (Roche
Diagnostic). After 24 h, cells were lysed and protein extracts
were used as size markers. GST – IscU protein was obtained
from bacterial expression of pGEX –IscU in XL1 Blue
cells and purified on glutathione sepharose 4B column according to the manufacturer’s recommendations (Amersham
Biosciences).
Human Molecular Genetics, 2007, Vol. 16, No. 22
Antibodies
A polyclonal antibody against mouse GPAT (R2374) was
raised against peptide PH228 (CLTGQYPVELEW). Affinity
purification was performed using a peptide-coupled sulfo-link
gel column according to the manufacturer’s protocol (Pierce).
A polyclonal antibody against mouse IscU (R2386) was raised
against peptide PH234 (CADYKLKQESKKEEPEKQ). Affinity purification was performed using a GST – IscU sulfo-link
gel coupled column (Pierce).
Protein electrophoresis and western blots
Total extracts were obtained by homogenizing frozen tissues
in 0.15 M Tris– HCl, pH 6.8, 25% glycerin, and 5% SDS as
described previously (4). Two different protein electrophoresis
systems were used [SDS – Tris/glycine –polyacrylamide gel
electrophoresis (PAGE) and SDS – Tricine – PAGE] and
western blot were carried out as previously described (4).
Antibodies were diluted as follows: anti-Fxn (R1270) and antivinculine (VIN-11-5, Sigma) at 1/1000; anti-GPAT (R2374)
and anti-IscU (R2386) at 1/2000; anti-Nfs1, anti-XOR
(Ab6194, Abcam), anti-b tubuline (MAB3408, Chemicon
International), anti-PHB (RB-292-P, Neomarkers Inc.) and
anti-Mn SOD (SOD-110, StressGen) at 1/5000; antiglyceraldehyde 3-phosphate dehydrogenase (GAPDH;
MAB374, Chemicon International) at 1/50 000. HRP-coupled
secondary antibodies were diluted at 1/5000.
Subcellular fractionation
The method was adapted from Gray and Whittaker (41).
Briefly, mouse tissues were homogenized in sucrose –
HEPES buffer (0.32 M sucrose, 4 mM HEPES, pH 7.4) in
the presence of a protease inhibitor cocktail using a glass
dounce. Homogenates were filtered on Sefar Nitrex 03-70/33
(Merck) and centrifuged at 1000g, 48C for 10 min. The supernatant was centrifuged at 10 000g, 48C for 10 min. The pellet,
corresponding to the mitochondrial fraction, was washed once
with the sucrose – HEPES buffer. The supernatant was further
centrifuged at 100 000g, 48C for 60 min to get the soluble
cytosolic fraction.
mRNA level determination
Total RNA from tissues was obtained using Trizol according
to the manufacturer’s protocol (Invitrogen). cDNA was generated using the Superscript II kit (Invitrogen) according to the
manufacturer’s protocol with oligo dT. Quantitative RT –PCR
was achieved using LightCycler (Roche Biosciences) and the
following primers: Nth1, forward 50 -CAAGATGGCACACT
TGGCTATG-30 , reverse 50 -GTCCGTTGACCTCACTCCAC
AG-30 ; Neil1, forward 50 -CCTGCTGGAACTGTGTCAC
TTG-30 , reverse 50 -CAGCTGTGTCTCCTGTGACTTC-30 ;
XOR, forward 50 -CCATTGAGTTCAGAGTATCCCTG-30 ,
reverse 50 -TTCTGGTGTTCCAGTGGCACACAG-30 ; GPAT,
reverse
forward
50 -ATCCGTGCTTCATGGGAATA-30 ,
0
5 -TTCTGGTGTTCCAGTGGCACACAG-30 ; hypoxanthineguanine phosphoribosyltransferase (HPRT), forward 50 -GTA
2657
ATGATCAGTCAACGGGGGAC-30 , reverse 50 -CCAGCAA
GCTTGCAACCTTAACCA-30 . HPRT was used as control.
SUPPLEMENTARY MATERIAL
Supplementary Material is available at HMG Online.
ACKNOWLEDGEMENTS
We thank Michel Koenig for fruitful discussions, Didier Gasparutto (CEA-Grenoble) for the kind gift of synthetic oligonucleotides, and Cécile Bouton for the kind gift of the Nfs1
antibody. We also thank Françoise Hoegy and Isabelle
Schalk (ESBS, Illkirch) for help in the fluorometric measurements. This work was funded by the French National Agency
for Research (ANR-05-MRAR-013-01), the French Ministry
for Research (ACI-JC5375) and the French Medical Research
Foundation ‘Equipe FRM 2005’ (DEQ2005-1205774). M.W.
is ATER Collège de France.
Conflict of Interest statement: None declared.
REFERENCES
1. Harding, A.E. (1981) Friedreich’s ataxia: a clinical and genetic study of
90 families with an analysis of early diagnostic criteria and intrafamilial
clustering of clinical features. Brain, 104, 589– 620.
2. Campuzano, V., Montermini, L., Molto, M.D., Pianese, L., Cossee, M.,
Cavalcanti, F., Monros, E., Rodius, F., Duclos, F., Monticelli, A. et al.
(1996) Friedreich’s ataxia: autosomal recessive disease caused by an
intronic GAA triplet repeat expansion. Science, 271, 1423–1427.
3. 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.
4. 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.
5. 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.
6. Duby, G., Foury, F., Ramazzotti, A., Herrmann, J. and Lutz, T. (2002)
A non-essential function for yeast frataxin in iron-sulfur cluster assembly.
Hum. Mol. Genet., 11, 2635–2643.
7. 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.
8. Gerber, J., Muhlenhoff, U. and Lill, R. (2003) An interaction between
frataxin and Isu1/Nfs1 that is crucial for Fe/S cluster synthesis on Isu1.
EMBO Rep., 4, 906 –911.
9. Ramazzotti, A., Vanmansart, V. and Foury, F. (2004) Mitochondrial
functional interactions between frataxin and Isu1p, the iron –sulfur cluster
scaffold protein, in Saccharomyces cerevisiae. FEBS Lett, 557, 215–220.
10. Layer, G., Ollagnier-de Choudens, S., Sanakis, Y. and Fontecave, M.
(2006) Iron–sulfur cluster biosynthesis: characterization of Escherichia
coli CYaY as an iron donor for the assembly of [2Fe –2S] clusters in the
scaffold IscU. J. Biol. Chem., 281, 16256– 16263.
11. Yoon, T. and Cowan, J.A. (2003) Iron– sulfur cluster biosynthesis.
Characterization of frataxin as an iron donor for assembly of [2Fe–2S]
clusters in ISU-type proteins. J. Am. Chem. Soc., 125, 6078– 6084.
12. Lill, R., Dutkiewicz, R., Elsasser, H.P., Hausmann, A., Netz, D.J.,
Pierik, A.J., Stehling, O., Urzica, E. and Muhlenhoff, U. (2006)
Mechanisms of iron-sulfur protein maturation in mitochondria, cytosol
and nucleus of eukaryotes. Biochim. Biophys. Acta, 1763, 652–667.
2658
Human Molecular Genetics, 2007, Vol. 16, No. 22
13. Lutz, T., Westermann, B., Neupert, W. and Herrmann, J.M. (2001) The
mitochondrial proteins Ssq1 and Jac1 are required for the assembly of iron
sulfur clusters in mitochondria. J Mol Biol, 307, 815 –825.
14. Kispal, G., Csere, P., Prohl, C. and Lill, R. (1999) The mitochondrial
proteins Atm1p and Nfs1p are essential for biogenesis of cytosolic Fe/S
proteins. EMBO J., 18, 3981–3989.
15. Li, K., Tong, W.H., Hughes, R.M. and Rouault, T.A. (2006) Roles of the
mammalian cytosolic cysteine desulfurase, ISCS, and scaffold protein,
ISCU, in iron– sulfur cluster assembly. J. Biol. Chem., 281,
12344–12351.
16. Acquaviva, F., De Biase, I., Nezi, L., Ruggiero, G., Tatangelo, F.,
Pisano, C., Monticelli, A., Garbi, C., Acquaviva, A.M. and Cocozza,
S. (2005) Extra-mitochondrial localisation of frataxin and its association
with IscU1 during enterocyte-like differentiation of the human colon
adenocarcinoma cell line Caco-2. J. Cell. Sci., 118, 3917–3924.
17. Condo, I., Ventura, N., Malisan, F., Tomassini, B. and Testi, R. (2006) A
pool of extramitochondrial frataxin that promotes cell survival. J. Biol.
Chem., 281, 16750–16756.
18. Lu, C. and Cortopassi, G. (2007) Frataxin knockdown causes loss of
cytoplasmic iron –sulfur cluster functions, redox alterations and induction
of heme transcripts. Arch. Biochem. Biophys., 457, 111 –122.
19. Fosset, C., Chauveau, M.J., Guillon, B., Canal, F., Drapier, J.C. and
Bouton, C. (2006) RNA silencing of mitochondrial m-Nfs1 reduces Fe–S
enzyme activity both in mitochondria and cytosol of mammalian cells.
J. Biol. Chem., 281, 25398–25406.
20. Biederbick, A., Stehling, O., Rosser, R., Niggemeyer, B., Nakai, Y.,
Elsasser, H.P. and Lill, R. (2006) Role of human mitochondrial Nfs1 in
cytosolic iron –sulfur protein biogenesis and iron regulation. Mol. Cell.
Biol., 26, 5675–5687.
21. Pondarre, C., Antiochos, B.B., Campagna, D.R., Clarke, S.L., Greer, E.L.,
Deck, K.M., McDonald, A., Han, A.P., Medlock, A., Kutok, J.L. et al.
(2006) The mitochondrial ATP-binding cassette transporter Abcb7 is
essential in mice and participates in cytosolic iron–sulfur cluster
biogenesis. Hum. Mol. Genet., 15, 953–964.
22. Seznec, H., Simon, D., Bouton, C., Reutenauer, L., Hertzog, A., Golik, P.,
Procaccio, V., Patel, M., Drapier, J.C., Koenig, M. et al. (2005) Friedreich
ataxia: the oxidative stress paradox. Hum. Mol. Genet., 14, 463 –474.
23. 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.
24. Harrison, R. (2002) Structure and function of xanthine oxidoreductase:
where are we now? Free Radic. Biol. Med., 33, 774 –797.
25. Zhou, G., Broyles, S.S., Dixon, J.E. and Zalkin, H. (1992) Avian
glutamine phosphoribosylpyrophosphate amidotransferase propeptide
processing and activity are dependent upon essential cysteine residues.
J. Biol. Chem., 267, 7936–7942.
26. Ide, H. and Kotera, M. (2004) Human DNA glycosylases involved in the
repair of oxidatively damaged DNA. Biol. Pharm. Bull., 27, 480 –485.
27. Takao, M., Kanno, S., Kobayashi, K., Zhang, Q.M., Yonei, S.,
van der Horst, G.T. and Yasui, A. (2002) A back-up glycosylase in Nth1
knock-out mice is a functional Nei (endonuclease VIII) homologue.
J. Biol. Chem., 277, 42205–42213.
28. Schoenfeld, R.A., Napoli, E., Wong, A., Zhan, S., Reutenauer, L.,
Morin, D., Buckpitt, A.R., Taroni, F., Lonnerdal, B., Ristow, M. et al.
(2005) Frataxin deficiency alters heme pathway transcripts and decreases
mitochondrial heme metabolites in mammalian cells. Hum. Mol. Genet.,
14, 3787–3799.
29. 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.
30. Bulteau, A.L., Ikeda-Saito, M. and Szweda, L.I. (2003) Redox-dependent
modulation of aconitase activity in intact mitochondria. Biochemistry, 42,
14846–14855.
31. Layer, G., Heinz, D.W., Jahn, D. and Schubert, W.D. (2004) Structure and
function of radical SAM enzymes. Curr. Opin. Chem. Biol., 8, 468–476.
32. Hanzelmann, P., Hernandez, H.L., Menzel, C., Garcia-Serres, R.,
Huynh, B.H., Johnson, M.K., Mendel, R.R. and Schindelin, H. (2004)
Characterization of MOCS1A, an oxygen-sensitive iron-sulfur protein
involved in human molybdenum cofactor biosynthesis. J. Biol. Chem.,
279, 34721–34732.
33. Tong, W.H. and Rouault, T. (2000) Distinct iron-sulfur cluster assembly
complexes exist in the cytosol and mitochondria of human cells.
EMBO J., 19, 5692–5700.
34. Tong, W.H. and Rouault, T.A. (2006) Functions of mitochondrial ISCU
and cytosolic ISCU in mammalian iron –sulfur cluster biogenesis and iron
homeostasis. Cell. Metab., 3, 199–210.
35. Tan, G., Napoli, E., Taroni, F. and Cortopassi, G. (2003) Decreased
expression of genes involved in sulfur amino acid metabolism in
frataxin-deficient cells. Hum. Mol. Genet., 12, 1699– 1711.
36. Rudolf, J., Makrantoni, V., Ingledew, W.J., Stark, M.J. and White, M.F.
(2006) The DNA repair helicases XPD and FancJ have essential iron–
sulfur domains. Mol. Cell. Biol., 23, 801–808.
37. Paulson, H.L. and Miller, V.M. (2005) Breaks in coordination: DNA
repair in inherited ataxia. Neuron, 46, 845–848.
38. Postic, C., Shiota, M., Niswender, K.D., Jetton, T.L., Chen, Y., Moates,
J.M., Shelton, K.D., Lindner, J., Cherrington, A.D. and Magnuson, M.A.
(1999) Dual roles for glucokinase in glucose homeostasis as determined
by liver and pancreatic beta cell-specific gene knock-outs using Cre
recombinase. J. Biol. Chem., 274, 305–315.
39. Takao, M., Kanno, S., Shiromoto, T., Hasegawa, R., Ide, H., Ikeda, S.,
Sarker, A.H., Seki, S., Xing, J.Z., Le, X.C. et al. (2002) Novel nuclear and
mitochondrial glycosylases revealed by disruption of the mouse Nth1
gene encoding an endonuclease III homolog for repair of thymine glycols.
EMBO J., 21, 3486–3493.
40. Beckman, J.S., Parks, D.A., Pearson, J.D., Marshall, P.A. and
Freeman, B.A. (1989) A sensitive fluorometric assay for measuring
xanthine dehydrogenase and oxidase in tissues. Free Radic. Biol. Med., 6,
607–615.
41. Gray, E.G. and Whittaker, V.P. (1962) The isolation of nerve endings
from brain: an electron-microscopic study of cell fragments derived by
homogenization and centrifugation. J. Anat., 96, 79 –88.