A cohort of 34 paediatric patients with identified isolated complex I deficiency in skeletal muscle was selected for this study (selected patients are presented in Table 1). The cohort was clinically heterogeneous, presenting with a variety of phenotypes, including Leigh syndrome, isolated hypertrophic cardiomyopathy and congenital lactic acidosis. In each case, sequencing of the entire mitochondrial genome had previously excluded the possibility of mtDNA point mutations as the cause of the biochemical defect. Written informed consent was obtained from all families in accordance with the Declaration of Helsinki and the study was approved by the local Ethics Committee. Table 1

Mitochondrial respiratory chain complex I activities in patient skeletal muscle homogenates

The activities of the respiratory chain complexes I–IV and the matrix marker citrate synthase were determined in frozen skeletal muscle homogenates from all 34 patients as previously described (Kirby et al., 2007).

Total genomic DNA was extracted from either muscle or blood samples of 34 paediatric patients using the Qiagen DNA mini kit (Qiagen, Crawley, UK) or Nucleon BACC3 Blood and Cell Culture DNA kit (Tepnel, Manchester, UK), respectively.

Prior to polymerase chain reaction (PCR) amplification of nuclear targets, whole genomes were amplified by multiple displacement amplification using Φ29 polymerase (REPLI-g® Mini kit; Qiagen), as per the manufacturer’s instructions.

The entire mitochondrial genome was amplified using 36 sets of overlapping M13-tailed primers (Supplementary Tables 1 and 2). The coding regions of nine nuclear encoded complex I genes (NDUFS1-4, NDUFS7, NDUFS8, NDUFV1, NDUFV2 and NDUFAB1) were amplified using intronic M13-tailed primers, designed to flank the coding sequences of the genes (Hinttala et al., 2005) (Supplementary Table 3). All PCRs were carried out in 25 μl volumes containing 0.65 U AmpliTaq Gold polymerase (Applied Biosystems, Warrington, UK), 1× AmpliTaq Gold buffer, 0.8 μM of each primer, 100 μM deoxynucleoside triphosphates and 30-100 ng DNA (mtDNA PCR) or 1 μl of amplified DNA (nuclear gene PCR). Where appropriate, reactions were supplemented with 2% dimethyl sulphoxide (Supplementary Table 3).

All PCR products were purified (ExoSapIT, GE Healthcare, Buckinghamshire, UK) and sequenced with BigDye Terminator cycle sequencing chemistries (Applied Biosystems) on an Applied Biosystems ABI3100 Genetic Analyser. Sequence data were analysed using SeqScape software (v2.1.1, Applied Biosystems) and compared with the GenBank reference sequences: NC_012920 (revised Cambridge reference sequence for human mtDNA), NM_005006.5 (NDUFS1), NM_004550.3 (NDUFS2), NM_004551.1 (NDUFS3), NM_002495.1 (NDUFS4), NM_024407.3 (NDUFS7), NM_002496.1 (NDUFS8), NM_007103.2 (NDUFV1), NM_021074.2 (NDUFV2) and NM_004994.1 (NDUFAB1).

Patients 3, 19, 22 and 35 and their parents were genotyped for the microsatellite markers D1S2635, D1S2707, D1S2771, D1S2705, D1S2675, D1S2844 and D1S1677 and the single nucleotide polymorphisms (SNPs) rs686015, rs10594, rs11582932, rs1041068, rs12402879, rs352685, rs512645, rs836, rs11576830, rs3924264, rs2501875, rs3935401, rs382627, rs2007773, rs6683580, rs12407444, rs2341481, rs4657136, rs1932933 and rs2341744. SNPs with a minor allele frequency of 0.15–0.48 and located at ∼100 kb intervals up- and downstream of NDUFS2 were chosen. Primer sequences for the microsatellite markers and map positions were obtained from the UniSTS database of the National Center for Biotechnology Information (Supplementary Table 4); primer sequences for the SNPs were designed using Primer3 software (v0.4.0.) (Rozen and Skaletsky, 2000) (Supplementary Table 5). Forward primers for the microsatellite markers were 5′-6-FAM-labelled; forward and reverse primers for the SNP analysis were M13-tailed. Reactions were carried out in 25 μl volumes containing 1 U GoTaq® polymerase (Promega, Southampton, UK), 1× GoTaq® buffer, 0.2 μM of each primer, 100 μM deoxynucleoside triphosphates and 20–100 ng DNA. PCR products (1 μl) were electrophoresed on an ABI3130xl Genetic Analyser (Applied Biosystems) with the GeneScan 500 ROX size standard (Applied Biosystems). Data were analysed using GeneMapper v4.0 software (Applied Biosystems).

The NDUFS1 c.1222C>T mutation was confirmed by PCR/restriction fragment length polymorphism (RFLP) analysis. The assay was performed as previously described (Taylor et al., 2003) with modifications. A 218-base pair fragment encompassing the mutation site was amplified using the M13-tailed forward primer 5′-TGTAAAACGACGGCCAGTCATGGATTCTCTGTATGTCTTAAT-3′ and reverse primer 5′-ACCAACCTCTTTCGAATTCTAGC-3′. PCR amplifications were performed with an annealing temperature of 59°C and PCR products were digested with 10 U NlaIII (New England Biolabs, Hitchin, UK). Wild-type amplicons contain a single NlaIII recognition site and on digestion two fragments of 25 and 193 bp are generated. The c.1222C>T mutation creates an additional NlaIII site, and amplicons harbouring this mutation are digested into three fragments of 25, 45 and 148 bp.

Cultured skin fibroblasts from Patients 3, 5, 19, 22 and 27 and from two normal, paediatric controls were cultured as previously described (Taylor et al., 2003).

Total RNA was extracted from fibroblasts using Trizol (Invitrogen, Paisley, UK) according to the manufacturer’s instructions. Reverse transcription of the messenger RNA was performed using the Superscript® II reverse transcriptase kit (Invitrogen).

Reactive oxygen species levels in cultured skin fibroblasts from Patients 3, 5, 19, 22 and 27 were measured as described previously (Saretzki et al., 2008). Briefly, cells (2 × 10⁵) were stained at 37°C in serum-free Dulbecco’s modified Eagle’s medium (Gibco, Paisley, UK) with 5 μM MitoSOX (Molecular Probes, Eugene, OR, USA) for 15 min to measure mitochondrial superoxide levels or with 10 μM of nonyl acridine orange (Molecular Probes) for 10 min to ascertain mitochondrial mass. Cells were analysed in a flow cytometer (Partec, Münster, Germany) using blue excitation and the green (FL1) and red (FL3) emission channels. System alignment and gain settings were adjusted using fluorescent beads. Cells were gated in forward scatter/side scatter, and the median of the gated fluorescence peak was used. Data were corrected by subtraction of autofluorescence levels in unstained cells. Neonatal human skin fibroblasts (Control 1), MRC-5 lung fibroblasts (Control 3, both from the American Type Tissue Collection, Rockville, Maryland, USA) and paediatric skin fibroblasts from a healthy child (Control 2) were used as controls. To determine statistical significance, two-sample unpaired Student t-tests between each patient and control sample were performed. A Bonferroni corrected P-value ≤ 0.05 was considered statistically significant.

Mitochondria-enriched fractions were prepared from 80% confluent 225 cm² flasks of patient and control fibroblasts and processed as described elsewhere (Nijtmans et al., 2002). Sample protein concentrations were determined by Bradford assay. Protein samples (75 μg/lane) were separated on 4–13% blue-native gradient gels, cast and run as previously described (Nijtmans et al., 2002). To assess complex I assembly, electrophoresed native gels were incubated for 1 h with dissociating reagent (1% sodium dodecyl suphate, 1% β-mercaptoethanol) prior to protein transfer onto polyvinylidene fluoride membranes (Immobilon-P, Millipore Corporation, Watford, UK). Immunodetection was performed using primary monoclonal antibodies raised against the complex I NDUFA9 and complex II 70 kDa subunits (MitoSciences, Cambridge BioScience, Cambridge, UK). ECL Plus™ (GE Healthcare Life Sciences, Buckinghamshire, UK) chemiluminescence was detected on a Storm 480 PhosphorImager (Molecular Dynamics, Sunnyvale, California, USA). To determine in-gel complex I activity, assays of NADH dehydrogenase activity were performed in-gel with NADH and nitrotetrazolium blue (Sigma, Poole, UK) as substrates, as previously described (Calvaruso et al., 2008).

Cell lysates were prepared from 80% confluent 75 cm² flasks of patient and control fibroblasts in 200 μl lysis buffer (42.5 mM Tris–HCl pH 7.5, 127.5 mM NaCl, 1.7 mM MgCl₂, 1% Nonidet P-40 and Roche ethylenediaminetetraacetic acid protease inhibitor cocktail). Samples were vortexed briefly, spun at 560 g for 2 min at 4°C and the supernatants retained. Protein concentrations were determined by Bradford assay. Equal amounts of protein (10–15 μg) were loaded and resolved on 10% sodium dodecyl sulphate polyacrylamide gel electrophoresis (PAGE) gels. Following wet transfer onto polyvinylidene fluoride membranes western blotting was performed using monoclonal antibodies against complex I NDUFS9, NDUFS3 and NDUFB8 and complex II 70 kDa subunits (MitoSciences). A monoclonal antibody raised against β-actin, clone AC-15, was also used (Sigma). ECL Plus (GE Healthcare Life Sciences) chemiluminescence was detected on a Storm 480 PhosphorImager (Molecular Dynamics).

Nine nuclear-encoded complex I subunits (NDUFS1, NDUFS2, NDUFS3, NDUFS4, NDUFS7, NDUFS8, NDUFV1, NDUFV2 and NDUFAB1) were sequenced in a cohort of 34 paediatric patients with demonstrated isolated complex I deficiency in skeletal muscle. Twenty-four sequence variants were identified, almost half of which are synonymous substitutions (Table 2). Of the remaining 13 non-synonymous substitutions, 4 are described SNPs (NDUFS2 p.P20T and p.P352A, NDUFS7 p.P23L and NDUFV2 p.V29A), whilst the homozygous NDUFS8 sequence change in Patient 34 (p.P79L; Fig. 1) has been previously described by Loeffen et al. (1998a) as one of two heterozygous mutations implicated in the pathogenesis of a patient with Leigh-like syndrome. The final 8 sequence changes are, to our knowledge, reported here for the first time. One homozygous NDUFS1 sequence variant, c.1222C>T (predicting p.R408C), was identified in Patient 5 (Fig. 2). Two heterozygous NDUFS2 sequence variants were discovered in Patient 27, c.413G>A (p.R138Q) and c.998G>A (p.R333Q) (Fig. 3A), and a single heterozygous NDUFS2 substitution, c.422A>G (p.Y141C), was detected in Patient 16. Patients 3, 19 and 22 all had an identical NDUFS2 transition, c.875T>C (p.M292T), in addition to secondary unique heterozygous sequence variants in the same gene, namely c.353G>A (p.R118Q), c.1328T>A (p.M443K) and c.442G>A (p.E148K), respectively (Fig. 3A). Interestingly, sequencing of a separate cohort of patients with isolated complex I deficiency (n = 34) in a diagnostic centre in Melbourne (Australia) also identified the NDUFS2 c.875T>C transition in a white Caucasian patient (Patient 35), in conjunction with a second unique heterozygous substitution, c.866+4A>G, located within the 5′-splice site of intron 6. DNA from this patient was included in the haplotype studies described below. Figure 2

NDUFS1 homozygous mutation, c.1222C>T (p.R408C). (A) Family 5 pedigree, (B) sequence electropherograms of NDUFS1 gene with c.1222 position highlighted in patient and family members, (C) confirmation of homozygous c.1222C>T mutation by PCR–RFLP. U = uncut; C1 = Control 1; C2 = Control 2. RFLP products were separated through a 12% non-denaturing polyacrylamide gel. The wild-type PCR product contains a single NlaIII site, which cuts the 218-bp amplicon into two fragments of 193 and 25 bp. In amplicons harbouring the c.1222C>T mutation, a second NlaIII site is created that cuts the 193-bp fragment into two smaller fragments of 148 and 45 bp. Fragment sizes (bp) are shown on the left; (D) amino acid alignment of NDUFS1 orthologues. Alignments were generated with ClustalW using GenBank sequences from Mus musculus (NP_663493.1), Rattus norvegicus (NP_001005550.1), Homo sapiens (NP_004997.4), Pan troglodytes (XP_516047.2), Canis lupus familiaris (XP_859697.1), Bos taurus (NP_777245.1), Danio rerio (NP_001007766.1), Drosophila melanogaster (NP_727255.1), Arabidopsis thaliana (NP_568550.1), Oryza sativa (NP_001051072.1) and Caenorhabditis elegans (NP_503733.1). Position p.408 (Homo sapiens) is indicated. Conserved sites are indicated (asterisk).

Parental genomic DNA samples were sequenced to confirm recessive inheritance of the NDUFS1 and NDUFS2 alleles. Both the mother and father of Patient 5 were revealed to be carriers of the NDUFS1 c.1222C>T transition (Fig. 2B and C). Unfortunately, DNA was unavailable from her unaffected brother; however, her clinically affected sister was similarly homozygous for this sequence variant. For all five NDUFS2 patients, each parent was shown to be heterozygous for only one of the two substitutions identified in respective patients (data not shown). During our investigations, the mother of Patient 3 conceived a third pregnancy and the parents requested prenatal genetic testing. Analysis of a chorionic villus biopsy failed to identify either of the familial NDUFS2 mutations (c.353G>A or c.875T>C).

To determine whether the repeatedly observed NDUFS2 c.875T>C transition could be attributed to a common founder event, Patients 3, 19, 22 and 35 and their parents were initially genotyped for microsatellite markers located upstream (D1S2635, D1S2707, D1S2771 and D1S2705) and downstream (D1S2675, D1S2844 and D1S1677) of NDUFS2 (Supplementary Table 4). D1S2705 was the only common flanking marker in all four patients. Due to the shortfall of microsatellite markers within the genomic region surrounding the NDUFS2 gene, a more exhaustive investigation was subsequently carried out by SNP analysis, using SNPs spaced ∼100 kb apart upstream (rs686015, rs10594, rs11582932, rs1041068, rs12402879, rs352685, rs512645, rs836 and rs11576830) and downstream (rs2501875, rs3935401, rs382627, rs2007773, rs6683580, rs12407444, rs2341481, rs4657136, rs1932933 and rs2341744) of NDUFS2. The intragenic NDUFS2 SNPs, rs3924264 and rs1136207, were also assessed (Supplementary Table 5). Up to 12 SNPs, spanning a region ∼480-kb upstream and ∼520-kb downstream of NDUFS2, appeared to co-segregate with the c.875C>T sequence variant in all four patients, suggesting a mutation-specific haplotype [rs352685 (G), rs512645 (A), D1S2705 (156), rs836 (G), rs11576830 (C), rs3924264 (A), rs1136207 (T), rs2501875 (G), rs3935401 (T), rs382627 (C/T), rs2007773 (A), rs6683580 (A) and rs12407444 (A); Supplementary Fig. 1].

Mitochondrial DNA haplotypes were determined for each c.875T>C patient from mtDNA polymorphisms. All patients belonged to independent Caucasian mitochondrial haplogroups, namely V (Patient 3), K (Patient 19), J (Patient 22) and U (Patient 35).

Investigation of complex I assembly and stability in muscle was not possible due to insufficient patient samples. However, cultured skin fibroblasts were available from Patients 3, 5, 19, 22 and 27. As cell lines do not consistently exhibit the complex I deficient phenotype (Mazzella et al., 1997; Benit et al., 2001; Antonicka et al., 2003), the activity of the complex was first assessed in these fibroblasts. In-gel complex I activity assays were performed on protein samples separated on 1D blue-native PAGE. Decreased complex I activity was identified in cell lines from Patients 3, 5 and 27, and to a lesser extent in cells from Patient 22 (Fig. 4A). Fibroblasts of Patient 19 demonstrated complex I activity comparable to controls. Figure 4

Complex I in-gel activity and assembly in patient fibroblasts. (A) Complex I in-gel activity. Mitochondria-enriched protein fractions (75 μg) extracted from patient (P) and control (C) fibroblasts were separated on 5–13% 1D blue-native PAGE gels. Gels were histochemically stained for 2 h with 2 mM Tris–HCl (pH 7.4), 0.1 mg/ml NADH and 2.5 mg/ml nitrotetrazolium blue to assess complex I activity. (B) Complex I assembly. Mitochondria-enriched protein fractions (75 μg) extracted from patient and control fibroblasts were separated on 5–13% 1D blue-native PAGE gels. Proteins were transferred to a polyvinylidine fluoride membrane and probed with antibodies raised against complex I NDUFS3 and complex II 70 kDa subunits. (C) Complex I subunit expression. Total cellular protein lysates (10 μg) of patient and control fibroblasts were separated on 10% sodium dodecyl sulphate–PAGE gels then transferred to a polyvinylidine fluoride membrane. Western blotting was performed with antibodies raised against complex I subunits NDUFS9, NDUFS3 and NDUFB8, complex II 70 kDa subunit and β-actin.

To ensure that all NDUFS2 alleles (both wild-type and mutant) were being transcribed, total RNA was extracted from fibroblasts of Patients 3, 19, 22 and 27. Following DNase1 treatment and reverse transcription of the RNA, regions of complementary DNA encompassing the putative pathogenic NDUFS2 mutations were sequenced. In all cases, both NDUFS2 alleles were found to be expressed (data not shown).

Complex I assembly in patient and control fibroblasts was assessed by western analysis of a 1D blue-native PAGE gel (Fig. 4B). An antibody against NDUFS3 was used to visualize complex I. To confirm loading, an antibody against the 70 kDa subunit of complex II was used. No complex I was detected in fibroblasts of Patient 3. Fully assembled complex I was present in fibroblasts of Patient 19 at levels comparable to controls. Intact complex I was also observed in fibroblasts of Patients 22 and 27, albeit at noticeably lower levels than controls. Whilst a marked decrease in fully assembled complex I was evident in fibroblasts of Patient 5, high levels of a lower molecular weight sub-complex were also observed.

Protein expression levels of complex I subunits NDUFA9, NDUFS3 and NDUFB8 were assessed by sodium dodecyl sulphate–PAGE in patient and control fibroblasts (Fig. 4C). To check loading, the expression levels of the 70 kDa subunit of complex II and β-actin were also analysed. No difference in NDUFS3 protein levels was observed in patient cells relative to controls. NDUFA9 and NDUFB8 protein levels were reduced in fibroblasts of Patients 3, 22 and 27 when compared with controls. No marked changes in NDUFA9 protein levels were evident in fibroblasts of Patient 5, whereas NDUFB8 levels were slightly increased. In fibroblasts of Patient 19, the expression levels of both NDUFA9 and NDUFB8 subunits were strongly increased relative to controls.

Three control fibroblasts [healthy neonatal (C1), paediatric (C2) or MRC-5 embryonic lung fibroblasts (C3)] demonstrated consistent values of MitoSOX and nonyl acridine orange staining, which were used as standards for cells without any mitochondrial defect. Cells from Patients 3, 5, 19 and 22 did not show any elevated levels of mitochondrial superoxides, while Patient 27 fibroblasts displayed an approximate 2.5-fold increase in MitoSOX staining (Bonferroni corrected P < 0.05; Fig. 5). Analysis of mitochondrial mass revealed no significant differences in patient fibroblasts when compared with controls, although nonyl acridine orange staining of fibroblasts of Patient 27 was slightly elevated. Figure 5

Reactive oxygen species levels and mitochondrial mass in patient fibroblasts. Mitochondrial superoxide levels were determined using MitoSOX (white bars) and mitochondrial mass was assessed with nonyl acridine orange (black bars) in fibroblasts of Patients 3, 5, 19, 22 and 27. Values are mean and standard error from at least three independent experiments. a.U. = arbitrary units; C1 = Control 1 (human neonatal skin fibroblasts); C2 = Control 2 (healthy paediatric skin fibroblasts); C3 = Control 3 (MRC-5 lung fibrobasts); NAO = nonyl acridine orange; *Bonferroni corrected P < 0.05.