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Brain Advance Access published online on February 21, 2007
Brain, doi:10.1093/brain/awm005

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A splice site mutation in the murine Opa1 gene features pathology of autosomal dominant optic atrophy

Marcel V. Alavi^1,*, Stefanie Bette^1,*, Simone Schimpf¹, Frank Schuettauf², Ulrich Schraermeyer³, Hans F. Wehrl⁴, Lukas Ruttiger⁵, Susanne C. Beck⁶, Felix Tonagel⁶, Bernd J. Pichler⁴, Marlies Knipper⁵, Thomas Peters⁷, Juergen Laufs⁷ and Bernd Wissinger¹

¹Molecular Genetics Laboratory, ²Department of Pathophysiology of Vision and Neuro-Ophthalmology, ³Section for Experimental Vitreoretinal Surgery, University Eye Hospital, ⁴Retinal Diagnostics Research Group, University Eye Hospital, Tuebingen, ⁵Molecular Neurobiology, Tuebingen Hearing Research Center (THRC), Department of Otorhinolaryngology, University of Tuebingen, ⁶Laboratory for Preclinical Imaging and Imaging Technology, Department of Radiology and ⁷Ingenium Pharmaceuticals AG, Martinsried, Germany

Correspondence to: E-mail: wissinger{at}uni-tuebingen.de

	Summary

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Autosomal dominant optic atrophy (adOA) is a juvenile onset, progressive ocular disorder characterized by bilateral loss of vision, central visual field defects, colour vision disturbances, and optic disc pallor. adOA is most frequently associated with mutations in OPA1 encoding a dynamin-related large GTPase that localizes to mitochondria. Histopathological studies in adOA patients have shown a degeneration of retinal ganglion cells (RGCs) and a loss of axons in the optic nerve. However little is known about the molecular mechanism and pathophysiology of adOA due to the lack of appropriate in vivo models. Here we report a first mouse model carrying a splice site mutation (c.1065 + 5G A) in the Opa1 gene. The mutation induces a skipping of exon 10 during transcript processing and leads to an in-frame deletion of 27 amino acid residues in the GTPase domain. Western blot analysis showed no evidence of a shortened mutant protein but a 50% reduced OPA1 protein level supporting haploinsufficiency as a major disease mechanism in adOA. Homozygous mutant mice die in utero during embryogenesis with first notable developmental delay at E8.5 as detected by magnetic resonance imaging (MRI). Heterozygous mutants are viable and of normal habitus but exhibit an age-dependent loss of RGCs that eventually progresses to a severe degeneration of the ganglion cell and nerve fibre layer. In addition optic nerves of mutant mice showed a reduced number of axons, and a swelling and abnormal shape of the remaining axons. Mitochondria in these axons showed disorganized cristae structures. All these defects recapitulate crucial features of adOA in humans and therefore document the validity and importance of this model for future research.

Key Words: adOA; OPA1; splice site mutation; mitochondria; mouse model

Abbreviations: adOA, autosomal dominant optic atrophy; ENU, N-ethyl-N-nitrosourea; MEFs, mouse embryonic fibroblasts; MRI, magnetic resonance imaging; OPA1, optic atrophy gene 1; RGC, retinal ganglion cell

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Received August 25, 2006. Revised December 3, 2006. Accepted January 5, 2007.

	Introduction

Top Summary Introduction Material and methods Results Discussion Supplementary material References

 
Autosomal dominant optic atrophy (adOA) is the most prevalent hereditary optic neuropathy with a reported frequency of up to 1 : 12 000 (Kivlin et al., 1983; Kjer et al., 1996). It is clinically characterized by a moderate to severe decrease in visual acuity, tritanopia, central visual field defects and optic nerve pallor. Disease onset is typically within the first two decades of life with a progressive course of visual function loss (Jaeger, 1974; Lorenz, 1994). However there is a considerable intra- and interfamilial variability in progression and severity of visual defects, ranging from functionally asymptomatic carriers to legally blind patients (Caldwell et al., 1971; Kline and Glaser, 1979; Roggeveen et al., 1985). In some families adOA is also associated with additional symptoms like hearing loss and adjunctive myopathic features including ptosis and progressive external ophthalmoplegia (Hoyt, 1980; Amati-Bonneau et al., 2003; Shimizu et al., 2003; Payne et al., 2004; Li et al., 2005). Histopathological post-mortem examinations of donor eyes have associated these visual impairments with a variable degeneration of retinal ganglion cells (RGCs) and optic nerve atrophy (Johnston et al., 1979; Kjer et al., 1983).

A major gene locus for adOA was identified on the chromosome region 3q28-q29 by linkage analyses (Eiberg et al., 1994) and subsequently mutations in a gene named optic atrophy gene 1 (OPA1) could be identified (Alexander et al., 2000; Delettre et al., 2000). Screening studies showed that 40–70% of unselected adOA patients carry mutations in the Opa1 gene, which probably is an underestimate given that larger heterozygous deletions and promoter mutations may not have been detected in these studies (Pesch et al., 2001; Toomes et al., 2001; Marchbank et al., 2002; Baris et al., 2003).

To date 117 different pathogenic mutations in OPA1 have been described (Ferre et al., 2005). Most mutations are predicted to give rise to truncated OPA1 polypeptides either by nonsense mutations, small deletions and/or insertions and splicing mutations. Therefore, haploinsufficiency is thought to be a major disease mechanism in OPA1-linked adOA (Marchbank et al., 2002).The Opa1 gene is composed of 30 coding exons distributed across more than 90 kb of genomic DNA on chromosome 3q28-q29. Alternative splicing of exons 4, 4b and 5b leads to eight transcript isoforms with open reading frames for polypeptides of 960–1015 amino acids.

The Opa1 gene is expressed ubiquitously (Alexander et al., 2000; Bette et al., 2005) with the highest levels in retina, brain, testis, heart and muscle (Alexander et al., 2000). OPA1 is a dynamin-related GTPase, which is targeted into mitochondria by an N-terminal import sequence and is anchored mainly to the inner membrane facing the intermembrane space (Delettre et al., 2000; Olichon et al., 2002; Satoh et al., 2003). The OPA1 protein structure includes an N-terminal coiled-coil domain presumably for membrane association, a GTPase domain, a middle domain and C-terminal coiled-coil domain proposed to be a GTPase effector domain (GED). It has been shown that OPA1 together with mitofusin 1 plays a major role in mitochondrial fusion and that the fusion/fission equilibrium is critical to maintain the intracellular mitochondrial network (Cipolat et al., 2004). Down-regulation of OPA1 expression by RNA interference in HeLa cells causes the fragmentation of the mitochondrial network, loss of the mitochondrial membrane potential and a disorganization of the cristae. Furthermore, release of cytochrome c from mitochondria and caspase-dependent activation of the apoptotic cascade were observed in this experimental model (Olichon et al., 2003; Lee et al., 2004). Knockout mutants of the OPA1-homologue mgm1/msp1 in yeasts also show fragmentation of mitochondria and reduction of mitochondrial DNA content (Hales and Fuller, 1997; Sesaki et al., 2003; Wong et al., 2003). The presence of OPA1 homologues in yeast and all higher metazoans thus points out the essential and fundamental biological role of OPA1.

To date, only little is known about the pathological mechanism that gives rise to adOA, especially why a ubiquitously expressed gene leads to a phenotype that is in most instances restricted to visual defects.

Here we present for the first time a mouse model carrying a pathogenic mutation in the Opa1 gene. This splice site mutation leads to skipping of exon 10 and results in a polypeptide with an in-frame deletion in the GTPase domain. Heterozygous mutants show a reduction in OPA1 protein levels to about 50% compared with wild-type littermates due to rapid degradation of the mutant polypeptide. Our results also indicate an essential role of OPA1 during early embryogenesis since homozygous mice are not viable and show impaired development as early as E8.5. Retinal histological micrographs reveal a drastic loss of RGCs and gliosis of the optic nerve in a 17-month-old mutant resembling human adOA histopathology. Furthermore, electron micrographs of sections through the optic nerve of Opa1^enu/+ mice revealed a reduced number of axons, and swelling and distortions of the remaining axons. Although the extent of RGC loss is variable among heterozygous animals, we can quantitatively show a progressive RGC degeneration in long-term experiments over a time course of 13 months by retrograde labelling of RGC with hydroxystilbamidine. Taken together, our results show that the Opa1^enu/+ mouse is an animal model that presents typical features of adOA in humans.

	Material and methods

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Animals, genotyping and cDNA analyses
The generation of the sperm archive of ENU-mutagenized mice, screening of this archive by means of temperature gradient capillary electrophoresis and the establishment of new mouse lines by in vitro fertilization have been previously described (Augustin et al., 2005). Mice were kept in a 12 h light (10 lux)/12 h dark cycle with food and water available ad libitum in full-barrier facilities free of specific pathogens. Mouse breeding and all experimental procedures were done according to the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research and approved by legal authorities. Opa1^enu/+-mice were initial purebred C3HeB/FeJ animals. To eliminate the Pdeb^rd1 allele present in this strain mutants, we applied a genotype assisted breeding scheme with an outcross on C57/Bl6. Subsequent Opa1^enu/+ mice devoid of the Pdeb^rd1 allele were intercrossed to obtain Opa1^enu/+ and wild-type littermate controls. Genomic DNA was extracted from mouse ear punches and isolated with the DNeasy Tissue Kit (Qiagen, Hilden, Germany). Genotyping of the c.1065 + 5 G A in exon 10 of the Opa1 gene was done by PCR amplification with primers OPA1 exon 10 forward: 5'-CATGAAGGTGCAGGGATTG-3' and OPA1 exon 10 reverse: 5'-GGAAAAGAACAAAGTCCTAAGACAA-3' and subsequent heteroduplex analysis using a dHPLC system (Wave 3500HT, Transgenomic Inc., Omaha, NE, USA). Total RNA was isolated from liver with the RNeasy Kit (Qiagen), reverse transcribed into single-stranded complementary DNA (cDNA) using SuperScript III reverse transcriptase (Invitrogen, Carlsbad, CA, USA) and the OPA1 cDNA amplified as six overlapping fragments (oligonucleotide sequences available upon request). For DNA sequencing PCR and RT-PCR products were purified using ExoSAP-IT (USB, Cleveland, USA), cycle-sequenced using Big Dye Terminator chemistry 3.1 (Applied Biosystems, Weiterstadt, Germany) and separated on an ABI 3100 capillary sequencer (Applied Biosystems). For qualitative and quantitative analysis of mutant OPA1 transcript processing, cDNA fragments covering either exon 6 to exon 9 or exon 6 to exon 10 (using oligonucleotide primers OPA1 exon 6 forward: 5'-CAGACAAAGAAAAGATTGACCAACTT-3', OPA1 exon 9 reverse: 5'-TCACTGGAGAGCGTGTCATCAT-3' and OPA1 exon 10 reverse: 5'-TTCCCGAGAGCTATCTTTAAACAAG-3') were amplified from cDNA generated from total liver RNA of a F1 animal from a Opa1^enu/+ x Balb/CJ cross. Various numbers of PCR cycles were tested to obtain products from the late logarithmic stage of the amplification reaction. RT-PCR products were digested with DdeI (New England Biolabs, Frankfurt am Main, Germany) and fragments were separated on 3%-agarose gel.

Qualitative and quantitative analysis of the OPA1 protein
All investigated animals were 3 months old. Comparative analyses were done with animals from the same litter. Mice were sacrificed by decapitation, immediately dissected and the analysed organs or tissues (e.g. brain, heart, liver, kidney, lung, skeletal muscle and retina) removed. Tissues were homogenized with a Dounce homogenizer in a buffer comprising 10 mM Tris–HCl pH 7.5, 50 mM sucrose, 200 mM mannitol, 1 mM EDTA, 1 mM DTT and protease inhibitor cocktail mix (Calbiochem, San Diego, CA, USA), followed by centrifugation for 10 min at 1000 x g and 4°C to pellet cell debris. Crude mitochondrial fractions were obtained by centrifugation for 15 min at 15 000 x g and 4°C. All work has been done on ice. The supernatant was referred to as cytosolic fraction. Concentrations of total protein were determined using a Bradford assay (Bio-Rad Protein Assay, Bio-Rad, Munich, Germany) following the manufacturer's protocol. Ten micrograms of total protein from each fraction were loaded on a 8% SDS–PAGE gel and blotted on a nitrocellulose membrane (Bio-Rad). Blots were cut horizontally into sections and subsequently processed for immunodetection with antibodies against OPA1 (BD-Transduction, LA, CA, USA), actin (Chemicon, Temecula, CA, USA), Hsp60 (StressGen, Victoria, Canada) or ubiquitin (Santa Cruz, Heidelberg, Germany) using the ECL chemiluminescence system (GE Healthcare, Freiburg, Germany). For quantitative analysis 5, 10, 20 and 40 µg total protein of mitochondrial fractions or total cell fractions were separated on 8% SDS–PAGE or 10% SDS–PAGE and processed accordingly. Protein ratios were calculated based on densitometrical quantification of scanned films using ImageJ (http://rsb.info.nih.gov/ij/). OPA1 protein levels were compared to either Hsp60 or actin levels.

Cell culture
Mouse embryonic fibroblasts (MEFs) were prepared from single embryos at E15.5 as described elsewhere (Nagy, 2003) and cultured under standard conditions in Dulbecco's Modified Eagle Medium supplemented with 10% foetal calf serum and 50 µg/ml penicillin/streptomycin. For protease inhibition assays MEFs were grown for 7.5 h with 2 µM Epoxomicin (Sigma, Munich, Germany) or 200 µM 1,10-phenanthroline (Sigma).

Quantification of mitochondrial DNA content
Copy numbers of mitochondrial DNA were determined in 38 (20 Opa1^enu/+ and 18 Opa1^+/+) isogenic mice (purebred C3HeB/FeJ) by real time PCR following the comparative Ct method. For this, total genomic DNA was isolated from ear punches taken at P21 solely by boiling in 10 mM NaOH, 0.1 mM EDTA for 30 min. We noted that ear punch DNA isolated with methods that use proteinase K digestion causes inhibition of real time PCR as it is also described for tail tips (Burkhart et al., 2002). Real time PCRs for a segment of the mtDNA encoded cytochrome b gene (mt-Cytb) and a segment of the ß-actin gene (ActB) were done in parallel with an ABI 7500 instrument and SYBRgreen (Applied Biosystems) as reporter dye following the manufacturer's protocol. Oligonucleotide primers for mt-Cytb were the same as described (Kim et al., 2005) but adapted to the murine mtDNA sequence: cytB-F: 5'-GGTCTTTTCTTAGCCATACACTACA-3'; cytB-R: 5'-ATATCGGATTAGTCACCCGTAAT-3'; primers used for ActB were actin-F: 5'-GATCGATGCCGGTGCTAAGA-3' and actin-R 5'-CACCATCACACCCTGTGGAAG-3'. All measurements were done in triplicate and mitochondrial DNA content per cell was calculated based on two copies of ActB per cell.

Embryo preparation and magnetic resonance imaging
Embryos at stages E12 and E17 and placentas were dissected free of decidua and uterine muscle, and separated from the yolk sac. Parts of the embryonic brain were isolated and washed in PBS and DNA isolated using DNeasy-Kit (Qiagen). Blastocysts (E3.5) were isolated as described elsewhere (Nagy, 2003) and used directly in a whole genome amplification reaction (GenomiPhi, GE Healthcare) for subsequent genotyping. Embryonic development was monitored in vivo by MRI using a 7 T ClinScan in vivo animal MRI System (Bruker BioSpin, Ettlingen, Germany), which is equipped with a 300 mm bore magnet and a 300 mT/m gradient system with an inner diameter of 200 mm. The system utilizes the Syngo MR software (Siemens, Erlangen, Germany), which was used for data acquisition and processing. Mice were anaesthetized with isoflurane (1.5%/0.8 l/min oxygen) and positioned inside a 35 mm quadrature whole body mouse coil installed at the magnet isocenter used for transmission and reception. A series (18 slices with thickness of 1.0 mm) of T₂ weighted sequences in sagittal, transversal and coronal orientation [repetition time (TR): 2770 ms; echo time (TE): 42 ms; 4 averages; turbo factor: 7] were applied with a field of view (FOV) of 35 mm x 35 mm and a matrix size of 256 by 256 (interpolated to 512 by 512). An in-plane resolution of 0.14 mm was obtained with an acquisition time of 13 min and 43 s for each direction.

Histology/transmission electron micrographs
Paraffin sections (5 µm) of 4, 13, 17 and 23-month-old mouse eyes fixed in 4% paraformaldehyde were stained with haematoxylin and eosin following standard protocols. For electron microscopy enucleated eyes of 8-month-old mice were fixed in ice-cold 2.5% glutaraldehyde in 0.1 M cacodylate overnight and processed as described previously (Schraermeyer et al., 1999). Number of axons was determined by counting 30 squares 7 µm x 7 µm of representative axial sections of the optic nerve.

RGC labelling
For RGC labelling, Opa1^enu/+ and control mice of 2, 9 and 13 months of age were deeply anaesthetized with chloral hydrate (6 ml/kg body weight of a 7% solution), administered intraperitoneally. To determine RGC densities, cells were labelled retrogradely with the fluorescent tracer hydroxystilbamidine (Molecular Probes, Eugene, OR, USA) by stereotaxic injections into both superior colliculi as described previously (Schuettauf et al., 2002). Two days after labelling animals were sacrificed with CO₂, eyes were enucleated, retinas dissected, flat-mounted on cellulose nitrate filters (pore size 60 m; Sartorius, Long Island, NY, USA) and fixed in 2% PFA for 30 min. Visualization of RGC was performed on the same day by fluorescence microscopy. Images obtained using a digital imaging system (ImagePro 3.0, Media Cybernetics Inc., Silver Spring, MD) connected to a microscope were coded and analysed in a masked fashion. Counts taken in twelve distinct areas of 62 500 µm² each per retina were added and expressed as the cell density (cells per square millimetre). The data are presented as means with respective standard errors, unless stated otherwise. Statistical analysis has been performed with the use of Statistica for Windows software (StatSoft, Tulsa, OK, USA), version 6.0. To assess the significance of differences in the densities of RGC in retinas collected from Opa1^enu/+ and control mice, data were analysed using unpaired Student's t-test.

Electroretinography and scanning laser ophthalmoscopy
ERGs and SLOs were performed according to previously described procedures (Seeliger et al., 2001, 2005). See supplementary material in Brain online for details.

Hearing measurements
Anaesthesia of animals, measurements of auditory brainstem responses (ABR) and the cubic distortion product of the otoacoustic emissions (DPOAE) and exposure to traumatic acoustic stimuli were performed as described before (Carnicero et al., 2004; Ruttiger et al., 2004) and outlined in detail as supplementary material.

	Results

Top Summary Introduction Material and methods Results Discussion Supplementary material References

 
OPA1 screening for mutations and cDNA analysis
Upon gene-based screening of a pre-made archive of N-ethyl-N-nitrosourea (ENU)-mutagenized mice we identified a mutant carrying a heterozygous mutation in intron 10 (c.1065 + 5 G A) of the Opa1 gene (Fig. 1A), which was subsequently revitalized from the corresponding sperm archive by in vitro fertilization. In the following we refer to this heterozygous mutant line as Opa1^enu/+-mice.

View larger version (32K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 1 The c.1065 + 5G A mutation in the Opa1enu/+ mutant induces a fully penetrant splicing defect. (A) Genomic sequence of the Opa1 gene showing the point mutation in the splice donor sequence of exon 10 (c.1065 + 5GA). (B) Overview about the localization of the mutation and its consequence upon transcript processing. (C) Direct sequencing of RT-PCR products from total liver RNA of the Opa1enu/+ mutant reveals a sequence overlay due to a skipping of exon 10 in mutant transcripts. Note that normal and mutant transcripts are at about the same level. (D) Overview about the strategy to analyse the penetrance of the splicing defect using F1 animals of an outcross to Balb/C. Transcripts from the mutant allele (on C3H background, lower panel) can be differentiated from Balb/C derived wild-type transcripts (upper panel) due to an SNP in exon 7 that abolishes a DdeI restriction site. The localization of primers used for RT-PCR of exon 6–9 and exon 6–10 cDNA fragments is given by horizontal arrows. Note that the RT-PCR applying the distal reverse primer in exon 10 specifically amplifies correctly spliced transcripts. (E) RT-PCR and RT-PCR/RFLP products of two OPA1 cDNA fragments (exon 6–9 and exon 6–10) amplified from liver RNA of a heterozygous F1 animal from the outcross to Balb/C. RT-PCR products from the mutant allele (C3H) can be differentiated from the wild-type allele (Balb/c) based on digestion with DdeI. While exon 6–9 products can be amplified for both alleles at about equal level (lane 3), exon 6–10 products are solely originating from the Balb/C allele (lane 4). The absence of exon 6–10 products for the C3H mutant allele indicate complete lack of correctly spliced products from mutant transcripts.

 
A guanosine nucleotide at position +5 in the splice donor sequence is well conserved and occurs in 80% of all vertebrate sequences (Padgett et al., 1986). To investigate whether this G to A substitution influences transcript processing, we analysed OPA1 cDNA synthesized from liver RNA of a heterozygous mutant animal. As shown in Fig. 1C we found correctly spliced OPA1 transcripts and transcripts that lack exon 10 in a roughly equal proportion. In order to assess whether skipping of exon 10 is obligate for transcripts derived from the mutant allele we crossed Opa1^enu/+ mice with Balb/C-mice that carry an A-to-C polymorphism in exon 7 (rs4167469). This SNP enabled us to discriminate between the mutant and the Balb/C wild-type allele in F1 animals. cDNA synthesized from liver RNA of these mice was investigated by semi quantitative PCR and PCR/RFLP analysis to discriminate both alleles (Fig. 1D). RT-PCR with primers that allow amplification of both the wild-type and the mutant cDNA (exon 6–9) results in roughly twice as much product than in an amplification restricted to correctly spliced mRNA (exon 6–10) (Fig. 1E, lanes 1 and 2). The complete digestion of the exon 6–10 RT-PCR product with DdeI clearly indicates that the mutant allele does not give rise to detectable amounts of correctly spliced transcripts (Fig. 1E, lane 4). Furthermore, our results show that OPA1 mRNAs from both alleles are about equally abundant (Fig. 1E, lane 3). Skipping of exon 10 in the mutant transcripts is predicted to result in an in-frame deletion of 27 amino acid residues (p.329-355del). These lacking 27 residues are an integral part of the GTPase domain of OPA1. Functionally homologous mutations, i.e. mutations that induce skipping of exon 10 and result in a homologous in-frame deletion have been identified in several adOA families (see Discussion).

Qualitative and quantitative analysis of the OPA1 protein
Western blot analyses were performed to characterize and quantify OPA1 protein in Opa1^enu/+ mice and control mice. First cellular localization of OPA1 was tested in cell fractionations obtained by differential centrifugation using a monoclonal antibody raised against the c-terminal portion of OPA1. Two well described bands (OPA1^L and OPA1^S) of OPA1 migrating between 85 and 100 kDa are clearly present only in mitochondrial fractions in all analysed tissues (brain, heart, liver, kidney, lung, skeletal muscle, retina) of Opa1^enu/+ and control mice. Figure 2A shows the analysis of whole brain protein extracts as an example. No qualitative differences could be detected in Opa1^enu/+ mice in respect to size even after separation on high resolution SDS–PAGE (data not shown) suggesting that either no protein is translated from mutant transcripts or that the mutant protein is unstable and rapidly degraded. We therefore quantified OPA1 protein levels in these tissues. Serial dilutions of mitochondrial protein or total protein were separated by SDS–PAGE, blotted on nitrocellulose membranes and sections of the same blot were assayed with antibodies either against OPA1 and Hsp60 (as mitochondrial marker protein) or antibodies against OPA1 and actin (as a cytosolic marker). Analyses of mitochondrial fractions of brain tissue of Opa1^enu/+ and control mice tested with OPA1 and Hsp60 antibodies are shown exemplarily in Fig. 2B and total retinal protein of the same animals analysed with OPA1 and actin antibodies in Fig. 2D. For the latter experiment a higher concentrated SDS–PAGE system was used to better separate OPA1 and actin, which precluded the clear separation of the two OPA1 isoforms. Yet both OPA1 isoforms are present in the retina as shown in Fig. 2C. Densitometric quantification of these western blots show a 50% reduction of OPA1 in Opa1^enu/+-mice with actin as calibrator (Fig. 2F). Normalized to the mitochondrial marker Hsp60 we found an even higher reduction to 42% of the wild-type mouse level (Fig. 2F), which might be due to an up-regulation of Hsp60 or a change in the number of mitochondria per cell in Opa1^enu/+ mice. Quantitative densitometric comparisons between the small versus the large OPA1 isoform showed no difference between Opa1^enu/+ and control mice, indicating that processing of OPA1 is not impaired by this mutation (Fig. 2G). To further investigate the absence of the mutant OPA1 protein, we established mouse embryonic fibroblasts (MEFs) of Opa1^enu/+ and control mice and cultured them in the presence of either epoxomicin, a specific inhibitor of the proteasome (Meng et al., 1999) or 1,10-phenanthroline, a chelator similar in its function to EDTA (Fig. 2F). OPA1 protein levels in MEFs treated with epoxomicin remain unchanged both in Opa1^enu/+ and in wild-type cells (Fig. 2H) although a massive accumulation of ubiquitinated proteins could be observed (Fig. 2F). On the other hand we noted a small increase of the total level of OPA1 protein only in Opa1^enu/+ MEFs upon treatment with phenanthroline. However the protein pattern did not differ from treated wild-type MEFs and smaller sized mutant OPA1 protein if present was still below the level of detection (Fig. 2F). Recently it has been shown that inhibition of the mitochondrial AAA-protease paraplegin with phenanthroline affects OPA1 processing (Ishihara et al., 2006). We also found a phenanthroline induced accumulation of the large OPA1 isoform (OPA1^L) in both Opa1^enu/+ and wild-type MEFs (Fig. 2G, arrows).

View larger version (59K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 2 Western blot analysis and quantitative analysis of OPA1 protein levels in the Opa1enu/+ mutant. (A) Both forms of OPA1 (OPA1L and OPA1S) can clearly be assigned to mitochondrial fractions in wild-type and Opa1enu/+ mice as shown exemplarily for whole brain protein fractions. An anti-Hsp60 antibody is used as mitochondrial marker. (B, E) Increasing amounts of whole brain mitochondrial protein from wild-type and the Opa1enu/+ mutant were used to generate a titration series for the quantification of OPA1. Using Hsp60 as calibrator we found that in Opa1enu/+ mutants the level of OPA1 protein is reduced to a relative ratio of 0.42 compared to wild-type littermates. (C) Assaying total cell lysate of different parts of the eye, we could show that OPA1 is highly enriched in the retina and presenting the typical pattern of two bands (OPA1L, OPA1S) that were not resolved in the gel shown in caption D. (D, E) Increasing amounts of total retinal protein from wild-type and the Opa1enu/+ mutant were used to generate a titration series. Using actin as cytosolic protein calibrator the level of OPA1 protein in mutants was reduced to a relative ratio of 0.5 compared to wild-type littermates. (F) MEFs were grown in the presence of either epoxomicin or phenanthroline to inhibit the proteasome or metal ion dependent proteases, respectively. Although there is a massive accumulation of ubiquitinated proteins, OPA1 levels remained unchanged in wild-type cells and Opa1enu/+ cells. (G) Densitometric scans of anti-OPA1 immunoblots in caption F show that the ratio of the two OPA1 isoforms (OPA1L and OPA1S) of the Opa1enu/+ mutant does not differ from that of wild-type littermates. The proportion of OPA1L is similarly increased in both wild-type and mutant MEFs upon treatment with phenanthroline, which inhibits the proteolytic cleavage of OPA1.

 
Quantification of mitochondrial copy number
Recently Kim and coworkers reported a reduction in mtDNA copy number in adOA patients with defined OPA1 mutations (Kim et al., 2005). Therefore and with respect to a reduced OPA1 protein level in the mutant, we investigated the mitochondrial DNA content in Opa1^enu/+ and control mice. DNA from ear punches from 20 Opa1^enu/+ mice and 18 wild-type littermates was used for relative quantification of cytochrome b (mt-Cytb) versus ß-actin (ActB) gene copy number by real time PCR. We obtained a mean value of 238 (SD 24.3255) copies of mt-Cytb per cell in Opa1^enu/+ mice compared to 236 (SD 33.0934) copies in wild-type littermates, excluding any significant differences between these two groups (t-test: t = 0.2502; P = 0.8055; Fig. 3).

View larger version (9K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 3 Quantification of mitochondrial DNA copy numbers in Opa1enu/+ by real time PCR. mtDNA content was determined by real time PCR amplification of a cytochrome b gene fragment and mtDNA copy number calculated relative to ct values obtained for the ß-actin gene. The box–whisker plot summarizing data of 20 Opa1enu/+ and 18 littermate controls of isogenic C3H background indicate that there is no significant difference in mtDNA copy number between the two groups.

 
Embryonic lethality in Opa1^enu mice
Heterozygous Opa1^enu/+ mice demonstrate regular viability and fertility with a normal habitus and a comparable behaviour to wild-type animals. They also do not differ in size or weight from their wild-type littermates. However of more than 25 progeny obtained from crosses of heterozygous mutants we never obtained homozygous mutant mice, suggesting lethality of homozygous Opa1^enu/enu in utero (Table 1). Examination of different embryonic states (E12–E17) revealed no homozygous embryos, but a large number of ‘moles’ representing dead placentas at a proportion equal to what would be expected for homozygous mutants. Unfortunately no embryonic tissue could be identified in these structures anymore, which prevented genotyping of these moles. Nevertheless, although we were unable to obtain genotypes for these moles, we assume that they represent the remnants of homozygous mutant embryos (Fig. 4C). However, we were able to obtain homozygous Opa1^enu/enu blastocysts at embryonic stage E3.5, which did not show any apparent morphological differences compared to heterozygous or wild-type blastocytes (Fig. 4A and B). These data indicate that homozygous Opa1^enu/enu embryos die between E3.5 and E12. To narrow the time point of embryonic death we followed embryonic development in vivo from day 6 to 15 post-fertilization in two pregnant animals using MRI. First conspicuities in embryonic development were observed at E8.5 for a total of six embryos, which were apparently retarded in their development (Fig. 4D, white arrows). This stall in development becomes more obvious at E9.5 (Fig. 4E). Still there was an accumulation of liquid over time in these retarded embryonic structures until E11.5 that finally is resorbed at E12.5 (Fig. 4F and G). These structures presumably represent the amnion which subsequently disintegrates upon ultimate death of the embryos. Taken together, these data show that first defects of embryogenesis could be assigned to E8.5 and that the homozygous mutant embryos die before E12.5 marked by the resorption of the amnion.

View larger version (125K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 4 Embryonic lethality of homozygous mutants. (A, B) Prior to nidation homozygous Opa1enu/enu blastocysts (E3.5) could be identified that morphologically do not differ from heterozygous blastocysts. Bar scale: 50 µm. (C) Overview that shows the dissected uterus of one female mouse that underwent MRI analyses. Embryos are spread according to their native position in the uterus. Embryos 3, 4, 5, 9, 10 and 11 were heterozygous for the OPA1 mutation, while embryo 1 is wild-type (black arrows). No homozygous mutant embryos were observed at this stage of foetal development (E15.5). However several embryonic remnants (so-called moles, white arrows) can be seen that most likely originate from stalled homozygotes. Note that mole 2 is buried in the adipose tissue. (D–G) In vivo monitoring of the development of the same embryos shown in caption C at different embryonic stages by MRI. Developmentally delayed embryos can be identified at E8.5 (D, white arrows), which becomes more prominent at E9.5 (E, white arrows). These amnions still accumulate liquid until E11.5 but are finally resorbed at E12.5 (F and G, white arrows).

 

View this table: [in this window] [in a new window]   Table 1 Numbers of progeny by genotype obtained from crosses of heterozygous Opa1enu/+ mice

 
In vivo imaging of the Opa1^enu/+ mutant retina and electroretinography
Fundus visualization with scanning laser ophthalmoscopy (SLO) at 2 and 9 months of age did not reveal any gross changes in the nerve fibre layer or signs of retinal degeneration in Opa1^enu/+ compared to wild-type retinas (Supplementary Material, S1). Since Lebers hereditary optic neuropathy –– another form of optic neuropathy caused by mutations in the mtDNA –– is often associated with alterations of the retinal blood vessels we applied fluorescence angiography to assess retinal and choroidal vasculature in Opa1^enu/+ mice. Yet this analysis revealed no changes in the inner retina or in the choroid of mutant mice (data not shown).

To investigate the effect of the deletion of Opa1^enu/+ on retinal function, ERGs were recorded from 2- and 9-month-old Opa1^enu/+ and age-matched wild-type littermates under scotopic and photopic conditions. Under both conditions, no significant differences were observed between the retinal electric signals obtained from Opa1^enu/+ and wild-type retinae at 2 and 9 months (Supplementary Material, S1). It should be noted that adOA patients do also show mostly normal responses in standard electroretinographic recordings.

Retinal histology and retrograde labelling of RGCs
Retinal sections of Opa1^enu/+ mice of different ages were investigated histologically. 2- and 4-months-old Opa1^enu/+ animals showed no differences compared to age-matched wild-type littermates while 13-month-old Opa1^enu/+ mice showed a slight reduction in the number of nuclei in the retinal ganglion cell layer compared with controls. Intriguingly one 17-month-old Opa1^enu/+ mouse showed a prominent loss of cells in the retinal ganglion cell layer to about 20–40% compared with wild-type littermates (Fig. 5C and D). In addition the retinal nerve fibre layer was drastically thinned in this mouse (Fig. 5A and B). On the other hand retinal cross sections of a 23-month-old mutant did not show this prominent reduction in the number of cells in the RGC layer. The murine ganglion cell layer does not only contain RGCs but up to 50% of displaced amacrine cells (Pesch et al., 2004). In order to assess and quantify specifically the loss of RGCs and to monitor its course and progression, RGCs were labelled retrogradely by injection of hydroxystilbamidine into both superior colliculi of Opa1^enu/+ and wild-type littermate mice at different ages (Fig. 6). Opa1^enu/+ mice showed a significant age-related decline in retinal ganglion cell number through 13 months. At two months, Opa1^enu/+ mice had 3022 ± 86 cells per mm² (n = 12; P = 0.0007), 2378 ± 71 cells per mm² at 9 months (n = 6; P = 0.0002), and 1467 ± 31 cells per mm² at 13 months, (n = 4; P < 0.0001). Control mice did not show any significant age-related changes with 3578 ± 113 cells per mm² (n = 10) at 2 months, 3204 ± 44 cells per mm² (n = 4) at 9 months and 3063 ± 219 cells per mm² (n = 2) at 13 months. These significant changes were consistent within the different areas of the mouse retina at 2 and 13 months. Yet at 9 months, significant changes were found solely in the mid-periphery and periphery of the respective retinae (Table 2).

View larger version (133K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 5 Massive loss of RGC layer nuclei and thinning of the nerve fibre layer in a 17-month-old Opa1enu/+ mutant. (A, B) Eosin–haematoxylin stained retinal sections of a Opa1enu/+ mutant and a wild-type littermate control. While the outer retinal layers (outer segments, outer and inner nuclear layer, outer plexiform layer) are normal, there is an extensive loss of RGC layer nuclei, a thinning of the nerve fibre layer and a gliosis of the optic nerve head in the Opa1enu/+ mutant. (C, D) Enlarged sections of A and B documenting the extensive loss of nuclei in the ganglion cell layer and an apparent thinning of the inner plexiform layer.

 

View larger version (104K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 6 Age-dependent loss of retinal ganglion cells in Opa1enu/+ mice. (A–F) Whole mount retina preparation of Opa1enu/+ and control mice following injection of hydroxystilbamidine into the superior colliculus that enable the specific labelling of retinal ganglion cells. Opa1enu/+ mice (A, C, E) show a reduced number of labelled cells in comparison with age-matched wild-type littermates (B, D, F) that progresses with age (A, B: 2 months; C, D: 9 months; E, F: 13 months). See also Table 2 for a statistical analysis.

 

View this table: [in this window] [in a new window]   Table 2 Retinal ganglion cell survival in Opa1enu/+ and littermate controls of different ages

 
Ultrastructural analysis of the optic nerve
Axial sections of the optic nerve of 8-month-old Opa1^enu/+ and wild-type littermate mice were prepared and examined by transmission electron microscopy. In the mutant we observed a complete loss of large axons, a disorganized structure, axonal swelling and distorted shapes of axons. Irregular myelination of axons as well as formation of membranous whorls and loss of myelin sheets were also noted (Fig. 7A and B). Furthermore, the optic nerve of the Opa1^enu/+ mutant showed an increased content of collageneous material and a decreased number of neurofibrils (Fig. 7A and B). We also observed a significant loss of small axons in Opa1^enu/+ mice based on quantitative analysis (Fig. 7C). In higher magnifications, there were also morphologically abnormal mitochondria with disorganized cristae in Opa1^enu/+ mice (Fig. 7E), that were not found in controls (Fig. 7D).

View larger version (137K): [in this window] [in a new window] [Download PowerPoint slide]   Fig. 7 Transmission electron microscopy of the optic nerve of Opa1enu/+ and control mice. (A, B) EM micrographs of axial section through the optic nerve of an Opa1enu/+ mutant and a wild-type littermate control. The mutant shows a reduced number of small axons, and a swelling and abnormal shape of the remaining axons. Bar scale: 7 µm. (C) Counting of axons on EM micrographs revealed a significant reduced number of axons in Opa1enu/+ mice. Dashed lines indicate mean values. (D, E) High magnification EM micrographs document that axons of the Opa1enu/+ mutant do contain a large number of structurally abnormal mitochondria with disorganized cristae. Bar scale: 100 nm.

 
Hearing function in Opa1^enu/+ mice
It is now well documented that adOA patients with the R445H mutation in OPA1 do also develop a sensorineural hearing loss (Amati-Bonneau et al., 2003; Shimizu et al., 2003; Payne et al., 2004; Li et al., 2005). We therefore tested hearing function in Opa1^enu/+ mice (Supplementary Material, S2). Hearing thresholds were determined in Opa1^enu/+ mice and controls in 2- and 8-month-old animals by click-induced and frequency specific auditory brain stem responses (ABR). Hearing thresholds were normal for both 2-month-old mice (hearing threshold mean ± SD 14.9 ± 2.14 dB SPL, n = 10 ears and 17.1 ± 7.71 dB SPL, n = 10 ears for wild-type and Opa1^enu/+ mice, respectively) and 8-month-old mice (18.7 ± 3.47 dB SPL, n = 11 ears and 15.4 ± 2.77 dB SPL, n = 14 ears for wild-type and Opa1^enu/+ mice, respectively). Hearing thresholds for frequency specific stimuli were also similar for both genotypes and ages. The active cochlear mechanics was investigated by measuring the distortion products of the otoacoustic emissions (DPOAEs). Again, no impairment or alteration of regular hearing function could be observed in Opa1^enu/+ mice. There was also no significant difference in hearing thresholds and DPOAE measurements between mutants and controls after exposing animals to acoustic trauma. These data indicate that this specific mutation in the Opa1 gene has no effect on the auditory system and its regenerative potential in mice.

	Discussion

Top Summary Introduction Material and methods Results Discussion Supplementary material References

 
This study introduces and characterizes a first mouse mutant with a pathogenic mutation in the murine Opa1 gene. We were able to show that the mutation induces a splicing defect that predicts an in-frame deletion in the polypeptide. Yet in heterozygotes we did not observe a shortened mutant OPA1 but a reduction on the level of OPA1 protein to about 50%. The mtDNA level however is not altered in the mutants. Homozygous mutants die in utero with first signs of developmental delay at E8.5. Heterozygous Opa1^enu/+ mice are vital and fertile and do not differ from their wild-type littermates in size, weight or lifespan. Hearing and outer retina (photoreceptors and bipolar cells) function are not impaired. However, heterozygous mice do show an age-related progressive loss of RGCs and optic nerve axons. Though variable in expression, the pathology in mutant mice eventually advances to a stage of nearly complete loss of RGCs and gliosis of the optic nerve that is very similar to histopathological studies in patients with severe adOA.

The c.1065 + 5G A mutation in the splice donor sequence of intron 10 that is present in the mouse mutant causes the skipping of exon 10 during transcript processing and is predicted to result in an in-frame deletion of 27 amino acid residues in the OPA1 polypeptide (p.329-355del). By crossing the mutant with Balb/CJ we could also show that the splicing defect is non-permissive, i.e. the mutant allele does not give rise to correctly spliced transcripts up to the level of detection (Fig. 1). Interestingly functional identical splicing defects of exon 10 of the human Opa1 gene have been observed in several families with adOA. Homologous to the mouse mutant skipping of exon 10 was also seen in the OPA1 cDNA of a family with a c.1065 + 3A C mutation (Pesch et al., 2001) and a family segregating a c.1065 + 1G > A mutation (unpublished results). Another pathogenic OPA1 mutation in that splice donor sequence, c.1065 + 2T C, has been identified in a Finish adOA family, however its consequence at the transcript level has not been investigated to date (Puomila et al., 2005). Therefore the Opa1^enu/+ mouse mutant represents a true model for OPA1 associated adOA at the molecular level.

Deletion of exon 10 in OPA1 transcripts predicts an in-frame deletion of 27 amino acid residues in the OPA1 protein. The deleted region (amino acid residues 329–355) covers parts of the highly conserved GTPase domain of OPA1. However we were unable to detect shortened OPA1 protein in tissue extracts of Opa1^enu/+ mutant mice neither in cytosolic nor in mitochondrial fractions using high resolution SDS–PAGE. Instead we could show that in mutant mice the level of OPA1 protein is reduced to about 50% of that in wild-type littermates in all investigated tissues and also for mitochondrial protein fractions (Fig. 2). Since the steady-state amounts of mutant and wild-type transcripts in Opa1^enu/+ mutant mice do not differ (Fig. 1), we reason that the mutant OPA1 protein is unstable and rapidly degraded. However the nature of this degradation (and the turnover of mitochondrial proteins in general) is not solved yet. The inhibition of proteasomal protein degradation does not alter the OPA1 protein levels in Opa1^enu/+ as well as in wild-type MEFs. Whether the small increase in total OPA1 protein in phenanthroline treated Opa1^enu/+ MEFs reflects the accumulation of mutant protein needs to be further investigated (Fig. 2).

However, the observation of a reduced OPA1 protein level in Opa1^enu/+ mutant mice strengthens current views on the pathological basis of adOA that argue for haploinsufficiency as a main disease mechanism. In fact Opa1 alleles with nonsense mutations close to the initiation codon like W2X, R38X or Q61X are unlikely to give rise to functional products or polypeptides that exert a dominant effect. Moreover we have observed that Opa1 transcripts that carry nonsense mutations or frame-shift mutations undergo non-sense-mediated mRNA decay in patient lymphocytes (Pesch et al., 2001 and unpublished results). And finally Marchbank and co-workers described an Australian adOA family that segregate a complete deletion of the Opa1 gene (Marchbank et al., 2002). Taken together, we propose that also other OPA1 mutations observed in adOA patients like those functionally homologous to the murine mutation (see above) or other in-frame deletions may exert their dominant effect through haploinsufficiency.

Our analyses show that the reduced level of OPA1 protein in the Opa1^enu/+ mutant does not affect mtDNA copy number. These data were obtained with purebred, age-matched mutants and wild-type littermates that were kept under controlled conditions. Although the ear punches used to quantify mtDNA content are not a homogeneous tissue we observed only little variance between samples indicating reliability of the comparison between Opa1^enu/+ mutant mice and their wild-type littermates. A singular study in adOA subjects has shown a reduced mtDNA copy number in leukcocytes (Kim et al., 2005). It remains to be shown in replicate studies whether this discrepancy is tissue specific or represent a principal difference between humans and mouse. However, these data also demonstate that mtDNA content and OPA1 protein levels are largely independent.

Our genotyping data demonstrate that homozygous Opa1^enu/enu mutants die in utero. No homozygous mutant embryos were observed at later embryonal stages (E12 and E17) but moles at a number as expected for homozygous mutants. However we still found homozygous mutant blastocysts at 3.5 days post coitum that do not differ morphologically from heterozygous mutant or wild-type blastocysts (Fig. 4). In vivo imaging of embryonic development by MRI revealed that embryogenesis is impaired as early as embryonic stage E8.5 (Fig. 4). At this time point the neural folds which develop with E7.5 start closing (Nagy, 2003). The embryonic lethality of homozygous mutants emphasizes the essential biological function of OPA1 as was also noted for other proteins involved in the mitochondrial fusion/fission process like mitofusin 1 and mitofusin 2 (Chen et al., 2003).

The outer retinal layers (photoreceptor outer segments, outer nuclear layer, outer plexiform layer and inner nuclear layer) of heterozygous Opa1^enu/+ mice are morphologically normal. ERGs responses that mainly assess function of the outer retina were also not impaired in Opa1^enu/+ mice. This is not a surprising finding since ERG recordings are mostly normal in adOA patients. In fact normal ERG responses are used to differentiate adOA from other clinical pictures like cone dystrophies that exhibit similar visual symptoms.

However, Opa1^enu/+ mice showed an age-dependent progressive loss of RGCs (Fig. 7). At 13 months of age mutant mice retain on average less than 50% of the number of RGCs compared to controls. The extent of RGCs loss was rather uniform throughout the retina, however at younger ages there is a tendency for a preferential loss in the peripheral retina (Table 1). An even more prominent loss of RGCs and a thinning of the nerve fibre layer was observed in retinal sections of a 17-month-old Opa1^enu/+ mouse (Fig. 5). This picture very closely resembles the retinal histopathology of adOA patients (Johnston et al., 1979). However we noted that the late stage of RGC degeneration in Opa1^enu/+ mutants is variable and not essentially as prominent as seen in Fig. 6. Phenotypic variability is also a hallmark of adOA in humans and has been attributed to modifier effects. This may hold true for our Opa1^enu/+ mutant, which has been kept and cross-bred on a mixed C3H x C57/Bl6 background in this study. Differences in the expression level of OPA1 as recently noted between mouse strains may be a factor that contributes to the variability in phenotypic expression (Pollak et al., 2006). Electron micrographs of axial sections of the optic nerve revealed a complete loss of large axons and a significant reduction of the number of small axons in Opa1^enu/+ mice. In addition we observed irregular or reduced myelin sheets, swelling and unusual shapes of axons and formation of membranous whorls (Fig. 7). There were also a number of irregular mitochondrial structures, i.e. mitochondria with disorganized or reduced cristae (Fig. 7). Similar morphological changes of mitochondria were also reported in siRNA induced OPA1 depletion in HeLa cells (Olichon et al., 2003). Interestingly, recent work suggests that OPA1 and its proteolytic processing plays a crucial role in stabilizing cristae junctions that govern cytochrome c release from mitochondria upon apoptotic stimuli (Cipolat et al., 2006; Frezza et al., 2006). Reduced amounts of the OPA1 protein, as observed in the Opa1^enu/+ mutant may therefore render cells more susceptible to apoptosis.

Our study shows that the Opa1^enu/+ mutant features the main characteristic of adOA in humans, that is the degeneration of retinal ganglion cells and loss of axons in the optic nerve. We therefore reason that the Opa1^enu/+ mutant is a valuable model to study this neurodegenerative process and to develop and evaluate future therapeutic strategies for adOA.

	Supplementary material

Top Summary Introduction Material and methods Results Discussion Supplementary material References

 
Supplementary material is available at Brain Online.

	Footnotes

 
*These authors contributed equally to this work.

	Acknowledgements

 
We are deeply grateful for the excellent technical assistance by N. Rieger, S. Bernhard-Kurz and S. Schultheiss. We appreciate comments and helpful discussion by M. and U. Sausbier, P. Heiduschka, S. Gerold, M. Seeliger, T. Schmidt and N. Fuhrmann (all University of Tuebingen). This work was supported by the Fritz Thyssen Stiftung für Wissenschaftsfoerderung, Koeln, Germany.

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