Skip Navigation


Human Molecular Genetics Advance Access originally published online on August 4, 2008
Human Molecular Genetics 2008 17(21):3291-3302; doi:10.1093/hmg/ddn225
This Article
Right arrow Abstract Freely available
Right arrow FREE Full Text (PDF) Freely available
Right arrow Supplementary Data
Right arrow All Versions of this Article:
17/21/3291    most recent
ddn225v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Add to My Personal Archive
Right arrow Download to citation manager
Right arrow Search for citing articles in:
ISI Web of Science (5)
Right arrowRequest Permissions
Google Scholar
Right arrow Articles by Spinazzi, M.
Right arrow Articles by Vergani, L.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Spinazzi, M.
Right arrow Articles by Vergani, L.
Social Bookmarking
Add to CiteULike   Add to Connotea   Add to Del.icio.us  
What's this?

© The Author 2008. Published by Oxford University Press. All rights reserved. For Permissions, please email: journals.permissions@oxfordjournals.org

A novel deletion in the GTPase domain of OPA1 causes defects in mitochondrial morphology and distribution, but not in function

Marco Spinazzi1,*, Silvia Cazzola1, Mario Bortolozzi2, Alessandra Baracca3, Emanuele Loro1, Alberto Casarin4, Giancarlo Solaini3, Gianluca Sgarbi3, Gabriella Casalena3, Giovanna Cenacchi5, Adriana Malena1, Christian Frezza6, Franco Carrara7, Corrado Angelini1, Luca Scorrano6, Leonardo Salviati4 and Lodovica Vergani1

1 Neurosciences Department, University of Padova, Italy 2 Venetian Institute of Molecular Medicine, Padova, Italy 3 Department of Biochemistry, University of Bologna, Italy 4 Department of Pediatrics, University of Padova, Padova, Italy 5 Department of Radiological and Histocytopathological Sciences, University of Bologna, Bologna, Italy 6 Dulbecco-Telethon Institute, Venetian Institute of Molecular Medicine, Padova, Italy 7 Istituto Neurologico C. Besta, Milano, Italy

* To whom correspondence should be addressed at: Clinica Neurologica II, Via Facciolati 71, 35100, Padova, Italy. Tel: +39 0498215315; Fax: +39 0498215310; Email: marco.spinazzi{at}unipd.it

Received May 15, 2008; Accepted July 31, 2008


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIAL AND METHODS
 SUPPLEMENTARY MATERIAL
 FUNDING
 REFERENCES
 
Autosomal dominant optic atrophy (ADOA), the commonest cause of inherited optic atrophy, is caused by mutations in the ubiquitously expressed gene optic atrophy 1 (OPA1), involved in fusion and biogenesis of the inner membrane of mitochondria. Bioenergetic failure, mitochondrial network abnormalities and increased apoptosis have all been proposed as possible causal factors. However, their relative contribution to pathogenesis as well as the prominent susceptibility of the retinal ganglion cell (RGC) in this disease remains uncertain. Here we identify a novel deletion of OPA1 gene in the GTPase domain in three patients affected by ADOA. Muscle biopsy of the patients showed neurogenic atrophy and abnormal morphology and distribution of mitochondria. Confocal microscopy revealed increased mitochondrial fragmentation in fibroblasts as well as in myotubes, where mitochondria were also unevenly distributed, with clustered organelles alternating with areas where mitochondria were sparse. These abnormalities were not associated with altered bioenergetics or increased susceptibility to pro-apoptotic stimuli. Therefore, changes in mitochondrial shape and distribution can be independent of other reported effects of OPA1 mutations, and therefore may be the primary cause of the disease. The arrangement of mitochondria in RGCs, which degenerate in ADOA, may be exquisitely sensitive to disturbance, and this may lead to bioenergetic crisis and/or induction of apoptosis. Our results highlight the importance of mitochondrial dynamics in the disease per se, and point to the loss of the fine positioning of mitochondria in the axons of RGCs as a possible explanation for their predominant degeneration in ADOA.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIAL AND METHODS
 SUPPLEMENTARY MATERIAL
 FUNDING
 REFERENCES
 
Autosomal dominant optic atrophy (ADOA, MIM165500) is the commonest cause of inherited optic atrophy. Its prevalence is estimated to be between 1/10000 and 1/50000 persons (1,2). Clinically, it is characterized by slowly progressive bilateral visual loss with frequent onset within the first two decades, dyschromatopsia, bilateral central scotoma and optic nerve atrophy. ADOA is caused by mutations in the optic atrophy 1 gene (OPA1) (3,4), mapped to chromosome 3q28–q29 and has variable clinical expression and penetrance (5).

OPA1 comprises 30 exons and is ubiquitously expressed, the highest level in the retina, but also abundantly in brain and muscle (4,6). Alternative splicing of exons 4, 4b, 5 and 5b results in at least eight isoforms, whose tissue distribution and relative abundance are scarcely known (7). Common to all isoforms of Opa1 are a GTPase, dynamin-like domain; a central domain; a C-terminal coiled coil domain presumably involved in protein–protein interaction; and an N-terminal mitochondrial targeting sequence that directs the protein to the inner mitochondrial membrane, facing the intermembrane space (8). Proteolytic cleavage of Opa1 occurs through distinct molecular pathways, both constitutively in a subset of Opa1 isoforms, and also by induction of apoptosis or loss of the mitochondrial membrane potential leading to Opa1 inactivation and mitochondrial fragmentation (9).

Various pathogenic mutations are dispersed throughout the gene coding sequence, but most occur in the catalytic GTPase domain (10). Almost 50% lead to premature truncation of the protein, and probably haploinsufficiency.

‘ADOA plus’ are rare complicated phenotypes with encephalomyopathic presentations, including sensorineural deafness (11), ataxia, ophthalmoplegia (12–14), peripheral neuropathy, associated with missense mutations in the GTPase domain through a possible semi-dominant mechanism. To date, only in these ADOA plus families, multiple mtDNA deletions were observed, which might explain defective energy metabolism and contribute to the atypical multisystemic phenotype.

OPA1 has been linked to two important mitochondrial activities: fusion (15), that in equilibrium with fission regulates mitochondrial morphology (16), and caspase-dependent apoptosis through mitochondrial cristae remodeling (17,18). The equilibrium between mitochondrial fusion and fission appears very dynamic and links mitochondrial morphology to mitochondrial function (16) and vice versa. Recent reports have also associated common OPA1 mutations, predicted to generate a truncated OPA1 protein and typical of uncomplicated ADOA phenotypes, to impaired oxidative phosphorylation with mitochondrial fragmentation in fibroblasts, and increased susceptibility to apoptosis (19,20). Moreover, the association of multiple mtDNA deletions with ‘ADOA plus’ phenotypes and missense mutations has suggested a putative role of OPA1 in mtDNA integrity maintenance (14). However, the relative contribution of these mechanisms to uncomplicated ADOA pathogenesis as well as their mutual relationships has not been unraveled. Thus, several important questions remain unanswered: is the mitochondrial fragmentation observed in OPA1-mutant cells a cause or a consequence of altered bioenergetics and apoptosis? Is pathology of ADOA really restricted to the optic nerve? What is the cause of the apparently exclusive susceptibility of the retinal ganglion cell (RGC) in ADOA, given that OPA1 is ubiquitously expressed?

In order to address the above questions, we investigated possible signs of neuromuscular involvement in a family with a non-syndromic form of ADOA harboring a novel OPA1 deletion. We then addressed its effects on mitochondrial function, morphology and viability in ex vivo biopsies.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIAL AND METHODS
 SUPPLEMENTARY MATERIAL
 FUNDING
 REFERENCES
 
A novel mutation in the Opa1 GTPase domain associated with extraocular signs
All patients were females of one family (Fig. 1A). The proband, Patient III-4 (47-year-old) developed chronic progressive visual loss from age 7, impaired color discrimination and later nocturnal distal paresthesias of the four limbs (Table 1). Neurological examination showed bilateral optic disc pallor (Fig. 1B) and pes cavus. Increased creatine kinase levels (208 U/L, nv 0–160) and neutropenia were found. Electromyography (EMG) showed a decrease in compound motor and sensory action potentials, slightly reduced conduction velocities in the lower extremities (sural nerve sensory conduction velocities 38 m/s) and initial denervation signs at the tibialis anterioris bilaterally indicating a moderate–severe predominantly axonal sensory-motor peripheral neuropathy at the lower extremities. Her father, affected by progressive visual loss, had gait difficulties in his old age. The 22-year-old daughter presented progressive visual loss from childhood, and type-I diabetes.


Figure 1
View larger version (28K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Figure 1. Clinical findings. (A) Pedigree of the studied family with autosomal dominant optic atrophy. Solid symbols, affected; arrow, index case. (B) Fundus photograph showing optic disc pallor, more evident on the temporal side. (C) Coronal Fast Spin Echo T2 weighted brain MRI image showing bilateral optic nerve atrophy (arrows).

 


View this table:
[in this window]
[in a new window]

  		Table 1. Clinical, electrophysiological and neuroradiological features of patients with ADOA

 
Patient III-6 (46-year-old) had chronic progressive visual loss from age 13. Neurological examination revealed bilateral optic disk pallor, moderate muscle deficit at the right tibialis anterioris and winging scapulae. EMG disclosed mild decrease of motor and sensory conduction velocities in the lower extremities and increased distal latencies, indicating a predominantly demyelinating peripheral neuropathy at the lower extremities.

Patient III-8 (44-year-old) presented chronic progressive visual loss from age 13. Neurological examination showed global optic disk pallor and winging scapulae. EMG was normal.

Brain MRI showed bilateral postlaminar optic nerve atrophy in all three patients (Fig. 1C). Audiometry was normal in all, as were lactate and pyruvate blood levels.

The clinical presentation of the patients, affected by prominent bilateral chronic progressive optic atrophy with childhood-onset, in association with a genetic inheritance consistent with an autosomal dominant pattern, prompted us to perform molecular analysis of the OPA1 gene.

Complete sequencing of OPA1 demonstrated a new 38-base-pair deletion (c1410_1443+4del38) spanning exon and intron 14 in the catalytic GTPase domain, with a predicted splice-site defect.

Expression of OPA1 c1410_1443+4del38
The mutation abolishes the physiological splice site at the 3'-end of exon 14. Amplification of cDNA extracted from patient fibroblasts revealed the presence of five different transcripts (Fig. 2A), one produced by the wild-type allele, the others resulting from the activation of cryptic splice sites in exon 14 or intron 14. The most abundant transcript results from the activation of a cryptic site 54 nucleotides downstream from the intron–exon boundary. It encodes for a truncated protein through the presence of an in-frame UAA termination signal in the retained intronic sequence, 11 codons after the deletion breakpoint, with a predicted protein product of 481 amino acids (Fig. 2A), undetectable by commercial antibodies targeted to epitopes between amino acid residue 500 and the C terminal. The three shorter transcripts, which are much less abundant, are produced by cryptic sites in intron 14 (five nucleotides after the intron–exon boundary), or in exon 14, 39 or 90 nucleotides upstream of the intron–exon boundary. These three transcripts maintain the correct reading frame, and are predicted to produce proteins lacking 11, 13 or 30 amino acids within the GTPase domain (Fig. 2A). It is not clear if these polypeptides are stable; however, it should be noted that the longest transcript accounts for the vast majority of aberrant transcripts seen in patient fibroblasts (>75% on densitometric analysis). Consistent with the molecular data is the abundance of full-length Opa1 protein in patient fibroblasts obtained by immunoblotting (~47% of controls) (Fig. 2B).


Figure 2
View larger version (28K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Figure 2. Opa1 in patient fibroblasts. (A) Analysis of cDNA in ADOA fibroblasts shows four altered transcripts, resulting in three shorter mRNA (217, 211 and 160) and one longer (266) than normal allele (250), owing to cryptic splice encompassing exon 14 and/or intron 14. Sequence analysis indicated the produced proteins, schematically represented at the side. (B) Expression levels of Opa1 protein in fibroblasts by western blotting, showing decreased levels of Opa1 in ADOA patients (P1–2) compared to controls (C1–5). Graph data of Opa1 bands, normalized to actin band density, represent mean ± SD from two independent experiments. Statistical significance: *P < 0.05.

 
Skeletal muscle abnormalities in OPA1 c1410_1443+4del38 patients
In order to investigate neuromuscular effects caused by OPA1 gene mutation, skeletal muscle biopsies were performed. Immunoblotting indicated a significant decrease of Opa1 (~31%) compared to controls (Fig. 3I). Morphological muscle analysis revealed mild abnormalities associated with denervation, e.g. atrophic type-I and -II fibers, angulated fibers, small-type grouping at ATPase staining and also some ring-shaped, split fibers (Fig. 3A). Muscle fibers had abnormal oxidative stain reactions with irregular central areas of loss of staining and subsarcolemmal and intermyofibrillar aggregates (Fig. 3B–D). Ragged red fibers or COX-negative fibers typical of mitochondrial DNA diseases were absent.


Figure 3
View larger version (112K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Figure 3. Muscle histopathological, Southern blot and protein analysis. Light microscopy analysis (A) H&E staining showing scattered atrophic fibers, fiber splitting (scale bar: 40 µm). (B) SDH reaction showing subsarcolemmal rims and focal areas of loss of staining. (C and D) NADH-TR and Gomori's trichrome staining showing intermyofibrillar aggregates of mitochondria. Ultrastructural analysis. (E) Low magnification shows clusters of mitochondria with lipid droplets (scale bar: 1 µm). (F and G) Polymorphous mitochondria with increased size variability and altered pattern of cristae organization with concentric or tubulo-vescicular arrangements (arrows). (H) Southern blot analysis of mtDNA from muscle of ADOA patients (P1–3), two controls (C), patient with a single large-scale mtDNA deletion (SD) and patient with multiple mtDNA deletions (MD). (I) Expression levels of Opa1 protein in skeletal muscle biopsies by western blotting, showing decreased levels of Opa1 in ADOA patients (P1–3) compared to controls (C1–3). Graph data of Opa1 bands, normalized to actin band density, represent mean ± SD from three independent experiments. Statistical significance: *P < 0.05.

 
However, ultrastructural analysis showed subsarcolemmal and intermyofibrillar aggregates of mitochondria with increased size variability, electron dense matrix (Fig. 3E–F). There were focal cristae abnormalities represented by tubulo-vescicular cristae and concentric abnormalities (Fig. 3G). Crystalline or paracrystalline inclusions were absent. Additionally, in Patient I, we observed myofibrillar disarray and lysis, and some atrophic and degenerated fibers with pycnotic nuclei and lipid droplets. OXPHOS activities were normal and both mitochondrial mass, expressed as citrate synthase activity, and mitochondrial DNA content, expressed as ratio cytB to Amyloid Precursor protein genes, were similar in ADOA compared to controls (58.29 ± 6.12 nmol/min/mg versus 63.88 ± 8.01; 1.65 ± 0.66 A.U. versus 1.76 ± 0.51, respectively). Southern blot analysis of mtDNA to screen for the presence of large-scale rearrangements was negative (Fig. 3H). In conclusion, these results showed that a decrease of Opa1 resulted in muscular atrophy and altered mitochondrial morphology, but not overt impairment of the respiratory chain or mtDNA instability.

Prominent mitochondrial morphology and distribution defects in cells derived from OPA1 c1410_1443+4del38 patients
To address the relative contribution of mitochondrial dysfunction versus morphological derangement in the observed muscular defects, we used primary cultures of fibroblasts and myotubes grown in glucose. Confocal microscopy of mitochondrially targeted red fluorescent protein (mtRFP) showed that a significantly increased proportion of OPA1-mutant fibroblasts presented fragmented mitochondria, appearing as spheres or very short rods (Fig. 4B–C; P < 0.005) compared to interconnected tubules as seen in controls (Fig. 4A and C). Aberrant mitochondrial shape was also retrieved in differentiated myotubes. Mitochondria of control myotubes displayed a tubular morphology with homogeneous reticular distribution along the whole cell (Fig. 4D). In ADOA myotubes, areas of tubular mitochondria alternated with areas of fragmented mitochondria even within the same cell, sometimes with a ‘beads on a string’ pattern (Fig. 4E) or abnormal large spherical morphology. Moreover, spatial distribution of mitochondria was remarkably irregular in ADOA myotubes, with areas of abnormal mitochondrial clustering while other areas were almost completely devoid of mitochondria (Fig. 4F–H). The visual appearance of altered mitochondrial distribution was corroborated by quantitative analysis calculated as parameter R, based on the ratio between average and maximal fluorescence on a given cell (examples are shown in Fig. 4H–I). This parameter was significantly decreased in OPA1 mutant myotubes compared to controls (R = 0.19 ± 0.01, n = 18 versus 0.41 ± 0.03, n = 17 in controls; P < 0.0001) (Fig. 4J) reflecting relatively lower average fluorescence intensity in areas of mitochondrial dearth and higher fluorescence values in areas of mitochondrial aggregation. Thus, our data indicate that mutated OPA1 results in abnormal mitochondrial shape and distribution under basal conditions.


Figure 4
View larger version (76K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Figure 4. Altered mitochondrial morphology and distribution in ADOA cells. Normal and ADOA fibroblasts labeled with DsRed plasmid and observed under confocal microscopy in glucose medium. (A) Representative epifluorescence image of typical tubular mitochondria of a control fibroblast (scale bar: 10 µm). (B) Representative epifluorescence image of fragmented, spherical mitochondria from ADOA fibroblast. (C) Bar graph showing quantitative analysis of mitochondrial shape changes in fibroblasts of patients compared to controls (n > 75 cells were counted for each group). Myotubes of controls and ADOA patients, after 10 days of differentiation in glucose, were loaded with MitoTracker Red and observed under confocal microscopy. (D) Representative epifluorescence image of a control myotube showing regular mitochondrial reticulum (scale bar: 20 µm). (E–G) Representative images of altered mitochondrial network in ADOA myotubes. (E) Bead-like mitochondria. (F–G) Mitochondrial network collapsed into clustered aggregates. (H) To compute the parameter R, regions of interests (ROIs), (white contours) are defined in controls (up) and OPA1-mutant (down) myotubes, basing on the myotube contours identified in the corresponding bright-field images. (I) Each pixel fluorescence value, F, of the representatives ROI in Fig. 4H generates a value between 0 and 1 for the distribution parameter R=F/Fmax, where Fmax is the highest F value of the ROI. The whole ROI has a Poisson distribution of R-values plotted as histogram. Abscissas are R intervals and ordinates the relative R frequency in the ROI. The ROI analysis shows that the OPA1-mutant distribution (black bars) is shifted towards lower values of R compared to WT (light gray bars). (J) Averaging R-values of 17 control myotubes led to a mean of 0.41 ± 0.03 (mean ± SE). This value is significantly higher compared to the OPA1-mutant R-value of 0.19 ± 0.01 (n = 18). Statistical significance: **P < 0.005; ***P < 0.0001.

 
Bioenergetics, ROS production and response to apoptotic stimuli are not impaired in cells derived from OPA1 c1410_1443+4del38 patients
Defects in mitochondrial morphology are often ancillary to primary impairment of mitochondrial function. To verify whether the observed organelle shape changes were a primary consequence of loss of pro-fusion activity of OPA1, we performed a functional analysis of the fibroblasts and myotubes isolated from the patients.

Preserved functions of both respiratory chain and ATP synthase, and integrity of the inner mitochondrial membrane were confirmed by evaluating the {Delta}{Psi}m of the fibroblasts under both state 3 and 4 respirations, and by measuring their ATP synthesis rate.

{Delta}{Psi}m was measured in resting fibroblasts cultured under different metabolic conditions: in glucose/pyruvate medium (data not shown) and in galactose/pyruvate medium (Fig. 5A–C) where galactose substituted for glucose as a substrate to push the aerobic mitochondrial ATP production. Though the ADOA cells show a slightly increased {Delta}{Psi}m compared to controls, the difference is below statistical significance both when ATP is actively synthesized (state 3) and when the ATP synthase proton channel is blocked by oligomycin (condition mimicking state 4) (Fig. 5B–C). We also evaluated {Delta}{Psi}m under state 3 and 4 conditions by TMRM probe using fluorescence microscopy, but no significant difference was observed in any tested culture condition between patients (Fig. 5B2 and C2) and controls (Fig. 5B1 and C1). The complex I-driven ATP synthesis rate was measured in resting fibroblasts cultured under different metabolic conditions, including the two above (Fig. 5D–E) and in galactose/pyruvate medium containing gramicidin to induce high energy demand (Fig. 5F). Gramicidin is an ionophore that stimulates ATP hydrolysis by the plasma membrane Na+/K+-ATPase. Our analysis indicates that the more energy is produced by OXPHOS, the more ATP synthesis rate falls in the ADOA compared to controls, but in none of the tested conditions was statistical significance reached, and the maximum difference between the two sets of samples was below 20% (Fig. 5E–F). To assess the energy availability of ADOA fibroblasts maintained in galactose/pyruvate medium, we determined the cellular ATP content (~30 nmol/mg protein), which was similar to controls (data not shown).


Figure 5
View larger version (49K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Figure 5. Normal mitochondrial membrane potential ({Delta}{Psi}m) and ATP synthesis rate in fibroblasts. (A) Typical fluorometric trace obtained in control-permeabilized fibroblasts cultured in galactose/pyruvate medium and incubated in ADP-containing buffer. {Delta}{Psi}m expressed by RH-123 fluorescence was evaluated under state 3 (+ Glu/Mal), state 4 (+ oligomycin) and uncoupled (+2 µM fluorogenic FCCP) respiratory conditions. (B) and (C) show the RH-123 steady-state fluorescence quenching (F.Q.) mean values measured in fibroblasts of both controls and ADOA patients under state 3 and 4 respiratory conditions, respectively. Images of fluorescence microscopy refer to ADOA and control fibroblasts maintained in galactose/pyruvate medium. Cells were loaded in respiratory buffer containing 0.2 µM fluorogenic TMRM, 20 µg/ml digitonin, 10 mM glutamate/10 mM malate, 0.4 mM ADP (state 3): (B1) control; (B2) ADOA patient. {Delta}{Psi}m measurements under state 4 respiratory condition were carried out adding 0.4 µM fluorogenic oligomycin to the samples: (C1) control; (C2) ADOA patient. (D), (E) and (F) show the complex I-driven ATP synthesis rate measured in fibroblasts of both controls and ADOA patients cultured in 25 mM glucose-110 mg/l pyruvate medium, 5 mM galactose-110 mg/l pyruvate medium and 5 mM galactose-110 mg/l pyruvate+40 ng/ml gramicidin medium, respectively. The mitochondrial ATP synthesis reaction was induced adding to digitonin-permeabilized fibroblasts (3 x 106 cells/ml) 10 mM glutamate/10 mM malate and 0.5 mM ADP. The data are averages ± SD of two independent triplicate experiments.

 
Latent mitochondrial dysfunction can be revealed by the real-time analysis of the changes in mitochondrial membrane potential following administration of ATPase inhibitor oligomycin. In cells where membrane potential is maintained by the reversal function of the ATPase, oligomycin causes a drop in TMRM fluorescence as opposed to the expected increase that follows inhibition of ATP synthesis in healthy mitochondria (21). Notably, in both control and patients' fibroblasts and myotubes, oligomycin resulted in an increase in mitochondrial TMRM fluorescence, indicating that membrane potential is maintained by proton pumping of fully functional respiratory chain complexes (Fig. 6A). Similarly, analysis of superoxide and H2O2 generation, detected by the fluorescence of the mitochondrially targeted MitoSOX (Supplementary Material, Fig. S1), and of the indicator Amplex Red (Fig. 6B), respectively, showed no difference between the two samples, indicating that the production of both ROS species, which often follows mitochondrial respiratory defects, was not increased in ADOA cells.


Figure 6
View larger version (28K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Figure 6. Preserved mitochondrial function, ROS production and response to apoptotic stimuli. (A) Normal TMRM uptake response to oligomycin in fibroblasts of controls (left) and ADOA patients (right). Data represent mean ± SE of five independent experiments. Where indicated (arrows), 2 µg/ml oligomycin and 2 µM fluorogenic carbonyl cyanide 4-(trifluoromethoxy)phenylhydrazone (FCCP) were added. (B) Normal H2O2 production in undifferentiated and differentiated cells from two ADOA patients and four controls. The values are means ± SD of two separate triplicate experiments. (C) Quantitative analysis of cell viability at baseline (without any pro-apoptotic stimuli) and after incubation with different apoptotic stimuli (staurosporine, etoposide, H2O2) showing a non-significant decrease in cell viability in ADOA fibroblasts. Data represent means ± SD of three independent experiments.

 
Finally, we verified if the abnormal mitochondrial network might be the result of a pre-apoptotic state, or lead to increased sensitivity to apoptosis, by testing fibroblast response to three different intrinsic apoptotic stimuli: etoposide, staurosporine and H2O2. A small, but not statistically significant increase in death was observed in ADOA fibroblasts both before and after administration of the apoptotic stimuli (Fig. 6C) indicating that at least the endpoint of apoptosis was unaffected by this ADOA mutation.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIAL AND METHODS
 SUPPLEMENTARY MATERIAL
 FUNDING
 REFERENCES
 
We report an ADOA family with a novel OPA1 gene deletion affecting the GTPase domain. Interestingly, we found subclinical neuromuscular involvement in addition to optic neuropathy, the cardinal feature of ADOA. These results were not unexpected considering the ubiquitous expression of OPA1, more prominent in the RGC but still considerable in skeletal muscle, brain and other tissues (4). Of note, skeletal muscle did not carry mtDNA deletions, suggesting no mtDNA instability. Overall, these data indicate that ADOA might have a predominant but not exclusive tropism for the optic nerves, and that neuromuscular abnormalities might be underestimated in classical ADOA and should be evaluated also in patients with different mutations.

The contribution of impaired OXPHOS and perturbed mitochondrial network to uncomplicated ADOA was uncertain. Our data show that mitochondrial network disruption is sufficient to cause ADOA without any appreciable respiratory deficiency.

Extensive bioenergetic evaluation did not detect any defect in ADOA fibroblasts even under stressed metabolic conditions, or in differentiated myotubes. These results differ from data recently reported by Zanna et al. (19) showing decreased complex I-driven ATP synthesis in ADOA-fibroblasts grown in glucose-free medium. However, this is in contrast with the reported normal cellular ATP content and cell growth that should indicate the absence of significant energetic defects. Moreover, being the mitochondrial mass of mutant cells grown in glucose-free medium ~20% higher than controls (19), the reduction of the ATP synthesis rate evaluated as ATP synthesis/CS ratio, results in an overestimation of the possible cellular energetic defect.

We propose that energy defect could variably arise to different extents from different mutations of the OPA1 gene, being more severe with missense mutations that determine ‘ADOA plus’ multisystemic phenotypes (11,13,14) with mtDNA multiple deletions, ragged red and cox-negative fibers, typical of classical mtDNA mitochondriopathies. However, bioenergetic failure alone would not explain why the high energy dependent pigmented epithelium of the retina and other types of neurons are not affected in the uncomplicated presentation of ADOA despite significant expression levels of Opa1 (6). In fact, pigmentary retinopathy, not observed in ADOA, occurs frequently in mitochondrial diseases with compromised energy metabolism (22) and severe complex-I deficiency associated with Leigh syndrome encephalo-cardiomyopathy is not necessarily associated with optic atrophy, even when caused by mutations in housekeeping genes in the nuclear DNA (23,24).

Moreover, while a putative pathogenetic analogy with Leber's hereditary optic neuropathy (LHON), associated with mtDNA mutations affecting complex I-subunits, would seem attractive, significant clinical and biochemical differences between the two conditions make this comparison unsatisfactory. In fact, while the end-stages of the two conditions are similar, ADOA gives a bilateral, slowly progressive visual loss in childhood, in contrast with LHON which is characterized by a later subacute and monolateral onset.

Moreover, in contrast with our data, LHON shows normal or slightly reduced redox activities of complex I (25), but induces severe complex I-driven ATP synthesis, impaired growth in galactose medium and ROS overproduction (26,27). In summary, inadequate energy production does not provide a comprehensive and simplistic explanation for ADOA pathogenesis nor for its predominant RGC involvement.

In contrast, we found a collapse of the mitochondrial network in ADOA fibroblasts and myotubes carrying an OPA1 deletion even under basal conditions in glucose medium, indicating that abnormal mitochondrial shape and distribution could be relevant per se in ADOA pathogenesis and in the relative tissue selectivity of the RGCs, as previously hypothesized (28,29). In contrast, in a previous study (19), mitochondrial fragmentation was observed in OPA1-haploinsufficient fibroblasts only under stressed metabolic conditions forcing oxidative phosphorylation, being possibly interpreted as a consequence of their defective complex I activity.

Moreover, we have provided, for the first time, data of mitochondrial architecture in differentiated OPA1-mutant myotubes: there were areas of mitochondrial clustering interposed with areas presenting very low mitochondrial density. The effect of OPA1 mutation on mitochondrial distribution seems to have some cell-type specificity, since it was evident in differentiated myotubes, rather than in fibroblasts. This may depend on relative Opa1 abundance and/or isoform expression, and different complexity of the mitochondrial network in different cell types.

Because of its unique ultrastructural and functional peculiarities, mitochondrial distribution is strictly regulated in the optic nerve to provide calibrated energy supply depending on different regional energy requirements. Mitochondrial concentration is, therefore, higher in the prelaminar unmyelinated optic nerve fibers, internodal regions and synapses where ATP consumption is higher and density of voltage-gated sodium channels is greater (30). Conversely, it is lower in the post-laminar myelinated region where energy requirement is reduced. This heterogeneous distribution is actively maintained to support normal function (28). As mitochondrial dynamics is crucial in providing the necessary energy at the right time and place (31), it is conceivable that the altered dynamics resulting from OPA1 mutation could trigger more pathological consequences in the RGCs than in other cell types.

Similar alterations in the architecture of the mitochondrial network were observed in OPA1 silenced rat RGCs (32), as well as COS-7 cells expressing OPA1 gene mutants (33), and patient's monocytes (3). However, the interpretation had been limited by lack of parallel mitochondrial functional analysis to rule out perturbed bioenergetics or apoptosis as the causes of these structural changes.

Moreover, abnormal mitochondrial distribution had also been observed in cell models of various neurodegenerative conditions, such as MFN2 mutant neurons (34), and myotubes (35), as well as in peripheral nerves of Charcot-Marie Tooth 2A (CMT2A) patients (36), and in optic nerves of patients affected by LHON (37).

Interestingly, MFN2, such as OPA1, is a dynamin-related GTPase involved in mitochondrial fusion. MFN2 mutations give rise to CMT2A, characterized by severe axonal peripheral neuropathy, sometimes with subacute optic neuropathy (38). An interesting speculation might be that the partial clinical overlap is caused by partial functional homologies between the two fusion proteins.

Finally, rearrangements of the mitochondrial distribution have also been related to increased ROS formation or apoptosis (39). However, in our experiments, mitochondrial network disruption was not associated with increased ROS production or significant vulnerability to pro-apoptotic stimuli, in contrast with a previous study (20).

We, therefore, suggest that abnormal mitochondrial network is the primary defect, at least in this pedigree of ADOA patients, whereas different mutation types could be associated with different bioenergetic profiles and susceptibility to apoptosis. An imbalance between the topographically different energy requirements of the RGCs and the supply of energy provided by mislocated mitochondria could lead to subcellular energy depletion and eventually progressive axonal impairment and cell death. In addition, the pathological fragmentation of the mitochondrial network could also lead to impaired calcium metabolism, abnormal electro-chemical connection between mitochondria and mtDNA complementation defects in susceptible cells (e.g. postmitotic) increasing interorganellar functional heterogeneity (40).

Biochemical effects might vary according to different OPA1 mutations, but also to the different tissues examined, owing to their specific histological organization, cell metabolism and expression of different OPA1 isoforms. Therefore, we suggest that extensive multiparametric mitochondrial analysis in different ADOA pedigrees should be not restricted to fibroblasts but extended to different cell types, and detailed neuropathological examinations of patients will be crucial in finding a common denominator of the pleiotropic effects of OPA1-mutants as the main pathogenetic mechanism.

Finally, these findings raise questions about the mechanism behind the mitochondrial spatial abnormalities in ADOA. It is now clear that the relocation of mitochondria requires competent fusion and fission machineries, as observed in leukocytes (41) and in neurons (42). One might speculate that mitochondrial clusters could result in OPA1-mutated cells from partially impaired fusion due to unopposed fission or from inter-mitochondrial tethering, as found in cells expressing truncated mitofusin-1 (43). Dissection of the biological machinery underlying altered mitochondrial dynamics and its functional consequences in differentiated cells is likely to have a relevant impact in understanding a number of neurodegenerative diseases.


   MATERIAL AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIAL AND METHODS
SUPPLEMENTARY MATERIAL
 FUNDING
 REFERENCES
 
Genetic analysis
After informed consent, genomic DNA was obtained using standard extraction protocols. OPA1 gene analysis was performed by PCR DNA amplification, DHPLC and direct DNA sequencing as described (44).

Analysis of mRNA splicing in fibroblasts
Total RNA was extracted from primary cultured skin fibroblasts of patients using Trizol (Invitrogen) according to the manufacturer's protocol. cDNA was synthesized using superscript III reverse transcriptase (Invitrogen) and random hexameres. A fragment encompassing the exons 13, 14 and 15 was amplified using primers 5'-TGCAGAATCCTAATGCCATC-3' and 5'-GAGCTGTTCCCTTTTCCTGT-3' and Taq DNA polymerase (Roche) and 5% DMSO. PCR conditions were 94°C 3 min, 35 cycles of 94°C 1 min, 55°C 1 min and 72°C 1 min, and a final extension step of 72°C for 7 min. PCR products were separated on a 4% agarose gel, excised and sequenced using amplification primers and the Big Dye Ready reaction kit and the 310 Genetic Analyzer (Applied Biosystems).

Muscle tissue analysis
Samples of skeletal muscle biopsies, stored at –80°C, were used for morphological, biochemical and immunoblotting analysis. Electron microscopy analysis was performed on fixed muscle specimens as described (45). The activities of NADH dehydrogenase (C-I), succinate dehydrogenase (C-II), NADH-cytochrome c oxidoreductase (C-I/III), SDH- cytochrome c oxireductase (C-II/III), cytochrome c oxidase (C-IV) and citrate synthase were measured in 600 g supernatant of tissue homogenates as described elsewhere (46). Homogenates of muscle biopsies were analyzed by western blotting using mouse anti-Opa1 antibody raised against the amino acid residues 708–830 (BD Transduction Laboratories) and visualized by ECL (Amersham). Protein quantity was determined by densitometry and normalized by the actin band.

MtDNA was quantified by RT–PCR as described (47). Total DNA samples were obtained from muscle tissue of three patients and four controls. DNA extraction was performed using GenEluteTM Mammalian Genomic DNA Miniprep Kit (Sigma, USA).

The mtDNA copy number was estimated by amplifying a portion of the cytochrome b (cyt b) gene of mtDNA and comparing it to the amplification profile of a nuclear single copy gene, Amyloid Precursor protein (APP). Primers for cyt b were: forward 5'-GCC TGC CTG ATC CTC CAA AT-3', reverse 5'-AAG GTA GCG GAT GAT TCA GCC-3'. Primers for APP were: forward 5'-TTT TTG TGT GCT CTC CCA GGT CT-3', reverse 5'-TGG TCA CTG GTT GGT TGG C-3'. ABI PRISM 7000 Sequence Detection System and Platinum SYBR green qPCR SuperMix-UDG (Invitrogen, USA) were used.

Southern blot analysis was carried out on PvuII digested total DNA using a non-radioactive hybridization system based on direct labeling of probe with horseradish peroxidase and detection system based on chemiluminescence (ECL Direct Nucleic Acid Labeling and Detection System RPN3000 GE Healthcare Amersham). Ten micrograms of genomic DNA extracted from muscle tissue with standard methods were southern blotted as described (48). Overnight hybridization was carried out following the protocol of the ECL Kit, using, as probes, two different PCR fragments of mtDNA. All procedures were performed following the ECL Kit instructions. Chemiluminescent reaction was carried out on the filter, which was then immediately used for autoradiography on Hyperfilm ECL Amersham (RPN3103K).

Mitochondrial morphology and distribution in cultured cells
Primary fibroblasts cultures were obtained from two patients (patient III-4, III-6) and three adult controls as described (49). The experiments were performed on cells with similar passage numbers 3 to 15. Myoblast cultures were obtained from muscle biopsies of ADOA patients and controls, and differentiated as previously described (49). Mitochondrial morphology was assayed in: (i) fibroblasts transfected with plasmids for expression of mitochondrially targeted DsRed (mtRFP) using Lipofectamine (Invitrogen); (ii) myotubes grown on coverslips loaded with 125 nM MitoTracker Red (Molecular Probes) in growth medium for 30' at 37°C, and observed in Hanks Buffered Salt Solution (HBSS). Fluorescence images were captured on living cells using a confocal Leica TCF SP5 microscope. For morphological quantification, fibroblasts were classified into two categories based on appearance of their mitochondrial network: (i) fragmented-intermediate length mitochondria when they predominantly appeared as spheres (<2 µm) or very short rods (<5 µm); (ii) tubular mitochondria when they appeared as elongated cables (>5 µm).

Analysis of mitochondrial subcellular distribution
We defined a distribution parameter R, independent of variability of MitoTraker Red loading and fluorescence emission intensity in relation to myotube position along the z-axis within cell layers. For a given region of interest (ROI) of the myotube, the distribution parameter was defined as R{equiv}F/Fmax, where F is mean fluorescence of the ROI (background subtracted image) and Fmax its highest fluorescence value. Images with saturated Fmax values were excluded from the analysis. For each myotube, the ROI was drawn with comparable area, based on the contours identified in bright-field (Fig. 4H). R (between 0 and 1) is expected to have higher values in cells with homogeneous mitochondrial distribution, as F is closer to Fmax compared to cells displaying mitochondrial clustering (Fig. 4I). Using the ratio F to Fmax helps to eliminate errors due to variability in fluorescence intensity.

Bioenergetic analysis in cultured cells
Mitochondrial membrane potential ({Delta}{Psi}m) response to oligomycin
Fibroblasts and myotubes were loaded with 20 nM tetramethyl rhodamine methyl ester (TMRM), a potentiometric fluorescent probe. Changes in TMRM fluorescence were monitored as described (21).

{Delta}{Psi}m measurement
Fluorescence micrographs of oligomycin-treated (state 4 respiratory condition) and untreated (state 3) adherent permeabilized fibroblasts were obtained as reported (50). Images were acquired using a fluorescence inverted microscope (Olympus IX50 equipped with a color CCD camera). Multiple high-power (20x) images were acquired with IAS2000 software (Delta Sistemi, Italy). To quantitate {Delta}{Psi}m under different metabolic conditions, steady-state fluorescence quenching measurements of permeabilized cells incubated with rhodamine 123 (RH-123) were carried out as described (50,51).

ATP synthesis
Mitochondrial ATP synthase activity was determined in permeabilized fibroblasts as described (52).

Citrate synthase
Citrate synthase activity was assayed according to Trounce's method (53). The cell sample protein concentration was quantified as described (52).

ROS determination
The selective detection of superoxide in the mitochondria of fibroblasts was obtained using MitoSOX Red (Molecular Probe) (54). Fibroblasts, grown on cover glass, were allowed to load MitoSOX for 20 min at 37°C at final concentration of 0.25 µM in HBSS. Before analysis with inverted microscope (Ex:510-Em:560), the fluorescent probe was washed out and cells were kept 20 min in incubator at 37°C. To test if the fluorescence was effectively produced by superoxide, cells were pre-incubated overnight with 0.5 mM N-acetyl cysteine and for 30 min with 0.1 mM Trolox. The fluorescent signal was abolished by 5 min pre-treatment with 20 µM FCCP. Rate of mitochondrial superoxide production, expressed in arbitrary units (AU)/hour/106 cells, was carried out by flow cytometry (FACS Calibur) using cells loaded in the same way with MitoSOX. The average intensity of MitoSOX fluorescence was recorded at time 0 and 10 min. The rate of H2O2 formation in living fibroblasts and myotubes was determined using the oxidation in the extracellular medium of 20 µM fluorogenic indicator Amplex Red (Invitrogen) in presence of 1 unit/ml horseradish peroxidase (Sigma) as described (55).

Analysis of cell death
A total of 3 x 105 fibroblasts of the indicated genotype were incubated for 24 h with 10 µM etoposide (Sigma), 300 µM H2O2, 750 nM staurosporine (Sigma) and stained with Annexin-V-Fluos (Boeringher) and propidium iodide, according to manufacturer's protocol. Apoptosis was measured by flow cytometry (FACS Calibur) as percentage of Annexin-V-positive events.

Statistical analysis
Data were expressed as mean ± SE or SD as indicated. Statistical analysis of group differences was performed using the Student t-test. Differences were considered significant at 95% confidence level (P < 0.05). Where appropriate, one-way ANOVA was applied with multiple comparisons followed by Bonferroni post hoc test.


   SUPPLEMENTARY MATERIAL
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIAL AND METHODS
 SUPPLEMENTARY MATERIAL
FUNDING
 REFERENCES
 
Supplementary Material is available at HMG Online.


   FUNDING
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIAL AND METHODS
 SUPPLEMENTARY MATERIAL
 FUNDING
REFERENCES
 
This study was supported by Association Française Contre le Myopathies (11032 to L.V.); Telethon Bank (GTB07001 to A.C.); Eurobiobank (QLRT 2001-027769 to A.C.); Telethon Italy (TELU02016 to L.S.).


   ACKNOWLEDGEMENTS
 
We would like to thank Prof. Bernd Wissinger, University of Tübingen, Germany for the OPA1 gene mutation analysis, Dr Piero Nicolao for the electrophysiological studies. Luca Scorrano is a Senior Telethon Scientist of the Dulbecco-Telethon Institute.

Conflict of Interest statement. None declared.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIAL AND METHODS
 SUPPLEMENTARY MATERIAL
 FUNDING
 REFERENCES
 

  1. Kjer B., Eiberg H., Kjer P., Rosenberg T. Dominant optic atrophy mapped to chromosome 3q region. II. Clinical and epidemiological aspects. Acta Ophthalmol. Scand. (1996) 74:3–7.[Web of Science][Medline]

  2. Bette S., Schlaszus H., Wissinger B., Meyermann R., Mittelbronn M. OPA1, associated with autosomal dominant optic atrophy, is widely expressed in the human brain. Acta Neuropathol. (2005) 109:393–399.[CrossRef][Medline]

  3. Delettre C., Lenaers G., Griffoin J.M., Gigarel N., Lorenzo C., Belenguer P., Pelloquin L., Grosgeorge J., Turc-Carel C., Perret E., et al. Nuclear gene OPA1, encoding a mitochondrial dynamin-related protein, is mutated in dominant optic atrophy. Nat. Genet. (2000) 26:207–210.[CrossRef][Web of Science][Medline]

  4. Alexander C., Votruba M., Pesch U.E., Thiselton D.L., Mayer S., Moore A., Rodriguez M., Kellner U., Leo-Kottler B., Auburger G., et al. OPA1, encoding a dynamin-related GTPase, is mutated in autosomal dominant optic atrophy linked to chromosome 3q28. Nat. Genet. (2000) 26:211–215.[CrossRef][Web of Science][Medline]

  5. Toomes C., Marchbank N.J., Mackey D.A., Craig J.E., Newbury-Ecob R.A., Bennett C.P., Vize C.J., Desai S.P., Black G.C., Patel N., et al. Spectrum, frequency and penetrance of OPA1 mutations in dominant optic atrophy. Hum. Mol. Genet. (2001) 10:1369–1378.[Abstract/Free Full Text]

  6. Wang A.G., Fann M.J., Yu H.Y., Yen M.Y. OPA1 expression in the human retina and optic nerve. Exp. Eye Res. (2006) 83:1171–1178.[CrossRef][Web of Science][Medline]

  7. Olichon A., Elachouri G., Baricault L., Delettre C., Belenguer P., Lenaers G. OPA1 alternate splicing uncouples an evolutionary conserved function in mitochondrial fusion from a vertebrate restricted function in apoptosis. Cell Death Differ. (2007) 14:682–692.[CrossRef][Web of Science][Medline]

  8. Olichon A., Emorine L.J., Descoins E., Pelloquin L., Brichese L., Gas N., Guillou E., Delettre C., Valette A., Hamel C.P., et al. The human dynamin-related protein OPA1 is anchored to the mitochondrial inner membrane facing the inter-membrane space. FEBS Lett. (2002) 523:171–176.[CrossRef][Web of Science][Medline]

  9. Griparic L., Kanazawa T., van der Bliek A.M. Regulation of the mitochondrial dynamin-like protein Opa1 by proteolytic cleavage. J. Cell Biol. (2007) 178:757–764.[Abstract/Free Full Text]

  10. Ferre M., Amati-Bonneau P., Tourmen Y., Malthiery Y., Reynier P. eOPA1: an online database for OPA1 mutations. Hum. Mutat. (2005) 25:423–428.[CrossRef][Web of Science][Medline]

  11. Amati-Bonneau P., Guichet A., Olichon A., Chevrollier A., Viala F., Miot S., Ayuso C., Odent S., Arrouet C., Verny C., et al. OPA1 R445H mutation in optic atrophy associated with sensorineural deafness. Ann. Neurol. (2005) 58:958–963.[CrossRef][Web of Science][Medline]

  12. Payne M., Yang Z., Katz B.J., Warner J.E., Weight C.J., Zhao Y., Pearson E.D., Treft R.L., Hillman T., Kennedy R.J., et al. Dominant optic atrophy, sensorineural hearing loss, ptosis, and ophthalmoplegia: a syndrome caused by a missense mutation in OPA1. Am. J. Ophthalmol. (2004) 138:749–755.[CrossRef][Web of Science][Medline]

  13. Hudson G., Amati-Bonneau P., Blakely E.L., Stewart J.D., He L., Schaefer A.M., Griffiths P.G., Ahlqvist K., Suomalainen A., Reynier P., et al. Mutation of OPA1 causes dominant optic atrophy with external ophthalmoplegia, ataxia, deafness and multiple mitochondrial DNA deletions: a novel disorder of mtDNA maintenance. Brain (2008) 131:329–337.[Abstract/Free Full Text]

  14. Amati-Bonneau P., Valentino M.L., Reynier P., Gallardo M.E., Bornstein B., Boissiere A., Campos Y., Rivera H., de la Aleja J.G., Carroccia R., et al. OPA1 mutations induce mitochondrial DNA instability and optic atrophy ‘plus’ phenotypes. Brain (2008) 131:338–351.[Abstract/Free Full Text]

  15. Cipolat S., Martins de B.O., Dal Z.B., Scorrano L. OPA1 requires mitofusin 1 to promote mitochondrial fusion. Proc. Natl. Acad. Sci. USA (2004) 101:15927–15932.[Abstract/Free Full Text]

  16. Ishihara N., Fujita Y., Oka T., Mihara K. Regulation of mitochondrial morphology through proteolytic cleavage of OPA1. EMBO J. (2006) 25:2966–2977.[CrossRef][Web of Science][Medline]

  17. Frezza C., Cipolat S., Martins de B.O., Micaroni M., Beznoussenko G.V., Rudka T., Bartoli D., Polishuck R.S., Danial N.N., De S.B., Scorrano L. OPA1 controls apoptotic cristae remodeling independently from mitochondrial fusion. Cell (2006) 126:177–189.[CrossRef][Web of Science][Medline]

  18. Olichon A., Baricault L., Gas N., Guillou E., Valette A., Belenguer P., Lenaers G. Loss of OPA1 perturbates the mitochondrial inner membrane structure and integrity, leading to cytochrome c release and apoptosis. J. Biol. Chem. (2003) 278:7743–7746.[Abstract/Free Full Text]

  19. Zanna C., Ghelli A., Porcelli A.M., Karbowski M., Youle R.J., Schimpf S., Wissinger B., Pinti M., Cossarizza A., Vidoni S., et al. OPA1 mutations associated with dominant optic atrophy impair oxidative phosphorylation and mitochondrial fusion. Brain (2008) 131:352–367.[Abstract/Free Full Text]

  20. Olichon A., Landes T., Arnaune-Pelloquin L., Emorine L.J., Mils V., Guichet A., Delettre C., Hamel C., Amati-Bonneau P., Bonneau D., et al. Effects of OPA1 mutations on mitochondrial morphology and apoptosis: relevance to ADOA pathogenesis. J. Cell Physiol. (2007) 211:423–430.[CrossRef][Web of Science][Medline]

  21. Irwin W.A., Bergamin N., Sabatelli P., Reggiani C., Megighian A., Merlini L., Braghetta P., Columbaro M., Volpin D., Bressan G.M., et al. Mitochondrial dysfunction and apoptosis in myopathic mice with collagen VI deficiency. Nat. Genet. (2003) 35:367–371.[CrossRef][Web of Science][Medline]

  22. Biousse V., Newman N.J. Neuro-ophthalmology of mitochondrial diseases. Curr. Opin. Neurol. (2003) 16:35–43.[CrossRef][Web of Science][Medline]

  23. Martin M.A., Blazquez A., Gutierrez-Solana L.G., Fernandez-Moreira D., Briones P., Andreu A.L., Garesse R., Campos Y., Arenas J. Leigh syndrome associated with mitochondrial complex I deficiency due to a novel mutation in the NDUFS1 gene. Arch. Neurol. (2005) 62:659–661.[Abstract/Free Full Text]

  24. Benit P., Chretien D., Kadhom N., de Lonlay-Debeney P., Cormier-Daire V., Cabral A., Peudenier S., Rustin P., Munnich A., Rotig A. Large-scale deletion and point mutations of the nuclear NDUFV1 and NDUFS1 genes in mitochondrial complex I deficiency. Am. J. Hum. Genet. (2001) 68:1344–1352.[CrossRef][Web of Science][Medline]

  25. Lenaz G., Baracca A., Carelli V., D'Aurelio M., Sgarbi G., Solaini G. Bioenergetics of mitochondrial diseases associated with mtDNA mutations. Biochim. Biophys. Acta (2004) 1658:89–94.[Medline]

  26. Baracca A., Solaini G., Sgarbi G., Lenaz G., Baruzzi A., Schapira A.H., Martinuzzi A., Carelli V. Severe impairment of complex I-driven adenosine triphosphate synthesis in leber hereditary optic neuropathy cybrids. Arch. Neurol. (2005) 62:730–736.[Abstract/Free Full Text]

  27. Carelli V., Rugolo M., Sgarbi G., Ghelli A., Zanna C., Baracca A., Lenaz G., Napoli E., Martinuzzi A., Solaini G. Bioenergetics shapes cellular death pathways in Leber's hereditary optic neuropathy: a model of mitochondrial neurodegeneration. Biochim. Biophys. Acta (2004) 1658:172–179.[Medline]

  28. Yu Wai Man C.Y., Chinnery P.F., Griffiths P.G. Optic neuropathies—importance of spatial distribution of mitochondria as well as function. Med. Hypotheses (2005) 65:1038–1042.[CrossRef][Web of Science][Medline]

  29. Chen H., Chan D.C. Critical dependence of neurons on mitochondrial dynamics. Curr. Opin. Cell Biol. (2006) 18:453–459.[CrossRef][Web of Science][Medline]

  30. Barron M.J., Griffiths P., Turnbull D.M., Bates D., Nichols P. The distributions of mitochondria and sodium channels reflect the specific energy requirements and conduction properties of the human optic nerve head. Br. J. Ophthalmol. (2004) 88:286–290.[Abstract/Free Full Text]

  31. Huang P., Yu T., Yoon Y. Mitochondrial clustering induced by overexpression of the mitochondrial fusion protein Mfn2 causes mitochondrial dysfunction and cell death. Eur. J. Cell. Biol. (2007) 86:289–302.[CrossRef][Web of Science][Medline]

  32. Kamei S., Chen-Kuo-Chang M., Cazevieille C., Lenaers G., Olichon A., Belenguer P., Roussignol G., Renard N., Eybalin M., Michelin A., et al. Expression of the Opa1 mitochondrial protein in retinal ganglion cells: its downregulation causes aggregation of the mitochondrial network. Invest. Ophthalmol. Vis. Sci. (2005) 46:4288–4294.[Abstract/Free Full Text]

  33. Misaka T., Miyashita T., Kubo Y. Primary structure of a dynamin-related mouse mitochondrial GTPase and its distribution in brain, subcellular localization, and effect on mitochondrial morphology. J. Biol. Chem. (2002) 277:15834–15842.[Abstract/Free Full Text]

  34. Baloh R.H., Schmidt R.E., Pestronk A., Milbrandt J. Altered axonal mitochondrial transport in the pathogenesis of Charcot-Marie-Tooth disease from mitofusin 2 mutations. J. Neurosci. (2007) 27:422–430.[Abstract/Free Full Text]

  35. Bach D., Pich S., Soriano F.X., Vega N., Baumgartner B., Oriola J., Daugaard J.R., Lloberas J., Camps M., Zierath J.R., et al. Mitofusin-2 determines mitochondrial network architecture and mitochondrial metabolism. A novel regulatory mechanism altered in obesity. J. Biol. Chem. (2003) 278:17190–17197.[Abstract/Free Full Text]

  36. Verhoeven K., Claeys K.G., Zuchner S., Schroder J.M., Weis J., Ceuterick C., Jordanova A., Nelis E., De V.E., Van H.M., et al. MFN2 mutation distribution and genotype/phenotype correlation in Charcot-Marie-Tooth type 2. Brain (2006) 129:2093–2102.[Abstract/Free Full Text]

  37. Carelli V., Ross-Cisneros F.N., Sadun A.A. Optic nerve degeneration and mitochondrial dysfunction: genetic and acquired optic neuropathies. Neurochem. Int. (2002) 40:573–584.[CrossRef][Web of Science][Medline]

  38. Zuchner S., De J.P., Jordanova A., Claeys K.G., Guergueltcheva V., Cherninkova S., Hamilton S.R., Van S.G., Krajewski K.M., Stajich J., et al. Axonal neuropathy with optic atrophy is caused by mutations in mitofusin 2. Ann. Neurol. (2006) 59:276–281.[CrossRef][Web of Science][Medline]

  39. Pletjushkina O.Y., Lyamzaev K.G., Popova E.N., Nepryakhina O.K., Ivanova O.Y., Domnina L.V., Chernyak B.V., Skulachev V.P. Effect of oxidative stress on dynamics of mitochondrial reticulum. Biochim. Biophys. Acta (2006) 1757:518–524.[Medline]

  40. Chan D.C. Mitochondria: dynamic organelles in disease, aging, and development. Cell (2006) 125:1241–1252.[CrossRef][Web of Science][Medline]

  41. Campello S., Lacalle R.A., Bettella M., Manes S., Scorrano L., Viola A. Orchestration of lymphocyte chemotaxis by mitochondrial dynamics. J. Exp. Med. (2006) 203:2879–2886.[Abstract/Free Full Text]

  42. Li Z., Okamoto K., Hayashi Y., Sheng M. The importance of dendritic mitochondria in the morphogenesis and plasticity of spines and synapses. Cell (2004) 119:873–887.[CrossRef][Web of Science][Medline]

  43. Koshiba T., Detmer S.A., Kaiser J.T., Chen H., McCaffery J.M., Chan D.C. Structural basis of mitochondrial tethering by mitofusin complexes. Science (2004) 305:858–862.[Abstract/Free Full Text]

  44. Pesch U.E., Leo-Kottler B., Mayer S., Jurklies B., Kellner U., Apfelstedt-Sylla E., Zrenner E., Alexander C., Wissinger B. OPA1 mutations in patients with autosomal dominant optic atrophy and evidence for semi-dominant inheritance. Hum. Mol. Genet. (2001) 10:1359–1368.[Abstract/Free Full Text]

  45. Cenacchi G., Fanin M., De Giorgi L.B., Angelini C. Ultrastructural changes in dysferlinopathy support defective membrane repair mechanism. J. Clin. Pathol. (2005) 58:190–195.[Abstract/Free Full Text]

  46. Russell A.P., Schrauwen P., Somm E., Gastaldi G., Hesselink M.K., Schaart G., Kornips E., Lo S.K., Bufano D., Giacobino J.P., et al. Decreased fatty acid beta-oxidation in riboflavin-responsive, multiple acylcoenzyme A dehydrogenase-deficient patients is associated with an increase in uncoupling protein-3. J. Clin. Endocrinol. Metab. (2003) 88:5921–5926.[Abstract/Free Full Text]

  47. Tyynismaa H., Sembongi H., Bokori-Brown M., Granycome C., Ashley N., Poulton J., Jalanko A., Spelbrink J.N., Holt I.J., Suomalainen A. Twinkle helicase is essential for mtDNA maintenance and regulates mtDNA copy number. Hum. Mol. Genet. (2004) 13:3219–3227.[Abstract/Free Full Text]

  48. Zeviani M., Moraes C.T., DiMauro S., Nakase H., Bonilla E., Schon E.A., Rowland L.P. Deletions of mitochondrial DNA in Kearns-Sayre syndrome. Neurology (1988) 38:1339–1346.[Abstract/Free Full Text]

  49. Martinuzzi A., Vergani L., Rosa M., Angelini C. L-carnitine uptake in differentiating human cultured muscle. Biochim. Biophys. Acta (1991) 1095:217–222.[Medline]

  50. Solaini G., Sgarbi G., Lenaz G., Baracca A. Evaluating mitochondrial membrane potential in cells. Biosci. Rep. (2007) 27:11–21.[CrossRef][Web of Science][Medline]

  51. Baracca A., Sgarbi G., Mattiazzi M., Casalena G., Pagnotta E., Valentino M.L., Moggio M., Lenaz G., Carelli V., Solaini G. Biochemical phenotypes associated with the mitochondrial ATP6 gene mutations at nt8993. Biochim. Biophys. Acta (2007) 1767:913–919.[Medline]

  52. Sgarbi G., Baracca A., Lenaz G., Valentino L.M., Carelli V., Solaini G. Inefficient coupling between proton transport and ATP synthesis may be the pathogenic mechanism for NARP and Leigh syndrome resulting from the T8993G mutation in mtDNA. Biochem. J. (2006) 395:493–500.[CrossRef][Web of Science][Medline]

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

  54. Robinson K.M., Janes M.S., Beckman J.S. The selective detection of mitochondrial superoxide by live cell imaging. Nat. Protoc. (2008) 3:941–947.[CrossRef][Medline]

  55. Vergani L., Malena A., Sabatelli P., Loro E., Cavallini L., Magalhaes P., Valente L., Bragantini F., Carrara F., Leger B., et al. Cultured muscle cells display defects of mitochondrial myopathy ameliorated by anti-oxidants. Brain (2007) 130:2715–2724.[Abstract/Free Full Text]


Add to CiteULike CiteULike   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us    What's this?


This article has been cited by other articles:


Home page
 IOVSHome page
P. Yu-Wai-Man, V. J. Davies, M. J. Piechota, L. M. Cree, M. Votruba, and P. F. Chinnery
Secondary mtDNA Defects Do Not Cause Optic Nerve Dysfunction in a Mouse Model of Dominant Optic Atrophy
Invest. Ophthalmol. Vis. Sci., October 1, 2009; 50(10): 4561 - 4566.
[Abstract] [Full Text] [PDF]

Home page
 Cardiovasc ResHome page
L. Chen, Q. Gong, J. P. Stice, and A. A. Knowlton
Mitochondrial OPA1, apoptosis, and heart failure
Cardiovasc Res, October 1, 2009; 84(1): 91 - 99.
[Abstract] [Full Text] [PDF]

Home page
 Physiol. Rev.Home page
M. Liesa, M. Palacin, and A. Zorzano
Mitochondrial Dynamics in Mammalian Health and Disease
Physiol Rev, July 1, 2009; 89(3): 799 - 845.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow FREE Full Text (PDF) Freely available
Right arrow Supplementary Data
Right arrow All Versions of this Article:
17/21/3291    most recent
ddn225v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Add to My Personal Archive
Right arrow Download to citation manager
Right arrow Search for citing articles in:
ISI Web of Science (5)
Right arrowRequest Permissions
Google Scholar
Right arrow Articles by Spinazzi, M.
Right arrow Articles by Vergani, L.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Spinazzi, M.
Right arrow Articles by Vergani, L.
Social Bookmarking
Add to CiteULike   Add to Connotea   Add to Del.icio.us  
What's this?