© 2014. Published by The Company of Biologists Ltd | Disease Models & Mechanisms (2014) 7, 1165-1174 doi:10.1242/dmm.015321

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ABSTRACT Mutations affecting mitochondrial complex I, a multi-subunit assemblythat couples electron transfer to proton pumping, are the mostfrequent cause of heritable mitochondrial diseases. However, themechanisms by which complex I dysfunction results in diseaseremain unclear. Here, we describe a Drosophila model of complex Ideficiency caused by a homoplasmic mutation in the mitochondrial-DNA-encoded NADH dehydrogenase subunit 2 (ND2) gene. Weshow that ND2 mutants exhibit phenotypes that resemble symptomsof mitochondrial disease, including shortened lifespan, progressiveneurodegeneration, diminished neural mitochondrial membranepotential and lower levels of neural ATP. Our biochemical studies ofND2 mutants reveal that complex I is unable to efficiently coupleelectron transfer to proton pumping. Thus, our study providesevidence that the ND2 subunit participates directly in the protonpumping mechanism of complex I. Together, our findings support themodel that diminished respiratory chain activity, and consequentenergy deficiency, are responsible for the pathogenesis of complex-I-associated neurodegeneration.

KEY WORDS: Mitochondria, Drosophila, Mitochondrial disease,Respiratory chain, Leigh syndrome, Neurodegeneration

INTRODUCTIONMitochondria perform numerous vital cellular functions, includingproducing the majority of ATP in cells, through oxidativephosphorylation via mitochondrial respiratory chain complexes.NADH ubiquinone oxidoreductase, or complex I, is the largestcomplex of the mitochondrial respiratory chain. It consists of ~45subunits, including seven that are encoded by the mitochondrialgenome: NADH dehydrogenase subunit 1 (ND1)-ND6 and ND4L(Efremov et al., 2010). All mitochondrial DNA (mtDNA)-encodedsubunits of complex I span the mitochondrial inner membrane,

RESEARCH ARTICLE

1Department of Genome Sciences, University of Washington, Seattle, WA 98195,USA. 2Molecular and Cellular Biology Program, University of Washington, Seattle,WA 98195, USA. 3Center for Developmental Therapeutics, Seattle Children’sResearch Institute, Seattle, WA 98101, USA. 4Department of Pathology, Universityof Washington, Seattle, WA 98195, USA.*Present address: The Department of Biochemistry and Faculty of MedicineSiriraj Hospital at Mahidol University in Bangkok, Bangkok 10700, Thailand.‡These authors contributed equally to this work

§Authors for correspondence (pgm4@uw.edu; pallanck@uw.edu)

This is an Open Access article distributed under the terms of the Creative CommonsAttribution License (http://creativecommons.org/licenses/by/3.0), which permits unrestricteduse, distribution and reproduction in any medium provided that the original work is properlyattributed.

Received 27 December 2013; Accepted 23 July 2014

where a subset of them might function in proton pumping giventheir homology to characterized bacterial complex I subunits(Amarneh and Vik, 2003; Dieteren et al., 2008; Efremov et al.,2010; Roberts and Hirst, 2012).

Mutations in genes encoding complex I subunits are a commoncause of early-onset mitochondrial diseases such as Leigh syndrome,and mitochondrial myopathy, encephalomyopathy, lactic acidosisand stroke-like symptoms (MELAS) (Triepels et al., 2001). Thesediseases are highly debilitating, and are typically characterized byprogressive neurodegeneration, seizures and shortened lifespan(Schon and Manfredi, 2003). However, the molecular mechanismsunderlying the pathogenesis of diseases associated with complex Ideficiency remain unclear (Torraco et al., 2009).

Recently, Xu and colleagues developed a novel technology forcreating targeted mtDNA mutations in Drosophila melanogaster,and used this technology to generate strains that bear homoplasmicmtDNA mutations (Xu et al., 2008). Two of the strains that weregenerated using this approach harbor mutations in the ND2 gene (Xuet al., 2008). Because mutations in human ND2 have beendemonstrated to cause Leigh syndrome (Ugalde et al., 2007),Leber’s hereditary optic neuropathy (Brown et al., 1992) andexercise intolerance (Schwartz and Vissing, 2002), we hypothesizedthat Drosophila ND2 mutants might serve as an animal model ofcomplex-I-associated disease. Here, we show that ND2 mutantsexhibit a variety of phenotypes that parallel symptoms of complexI deficiency in humans, including stress-induced seizures,progressive neurodegeneration and shortened lifespan. Moreover,our biochemical studies of ND2 mutants reveal that their complex Iinefficiently couples electron transfer to proton pumping, resultingin decreased mitochondrial membrane potential and diminishedenergy production. Our findings provide the first evidence that theND2 subunit participates in the proton pumping activity of complexI, and suggest that symptoms associated with mitochondrial complexI deficiency derive from an energy deficit.

RESULTSDrosophila ND2 mutantsIn previous work, Xu and colleagues expressed a mitochondriallytargeted restriction enzyme in the germline of Drosophila femalesto select for surviving offspring that bear an mtDNA mutation in thecorresponding restriction site. Two of the strains that were generatedusing this approach harbor mutations in the ND2 gene. One of thesetwo mutants, mt: ND2ins1 (ND2ins1), bears a three-nucleotideinsertion in the ND2 coding sequence, and the other, mt: ND2del1

(ND2del1), bears a nine-nucleotide deletion in the ND2 codingsequence (Xu et al., 2008). The ND2ins1 mutation adds a serine atposition 189, whereas the ND2del1 mutation removes three aminoacids at positions 186-188 of the ND2 protein (supplementary

A Drosophila model of mitochondrial disease caused by acomplex I mutation that uncouples proton pumping from electron transferJonathon L. Burman1,‡, Leslie S. Itsara1,2,‡, Ernst-Bernhard Kayser3, Wichit Suthammarak3,*, Adrienne M. Wang4, Matt Kaeberlein4, Margaret M. Sedensky3, Philip G. Morgan3,§ and Leo J. Pallanck1,§

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material Fig. S1). These mutations reside in evolutionarilyconserved sequences (supplementary material Fig. S1), thus raisingthe possibility that flies bearing these mutations might serve asmodels of mitochondrial disease. Because the phenotypes of thesefly strains were not characterized in previous work, we proceededto test this hypothesis.

Behavioral analyses of ND2 mutantsMitochondrial encephalopathies are typically characterized byneurological symptoms such as seizures (Pieczenik and Neustadt,2007). Similarly, Drosophila strains with mutations affectingmitochondrial function often show an analogous seizure-likeparalytic phenotype caused by mechanical stress, termed ‘bangsensitivity’ (Celotto et al., 2011; Fergestad et al., 2006; Ganetzkyand Wu, 1982). To determine whether ND2 mutants exhibited bangsensitivity, we subjected w1118 (control), ND2del1 and ND2ins1 mutantsto mechanical stress, and measured the time it took for them torecover from paralysis. Whereas control and ND2ins1 mutants wereunaffected by mechanical stress, ND2del1 mutants displayed bangsensitivity that progressively worsened with age (Fig. 1A;supplementary material Movie 1). Our experiments also revealedthat ND2del1 mutant females displayed substantial bang-sensitiveparalysis by 10 days of age, whereas ND2del1 mutant males did notexhibit significant bang sensitivity until 26 days of age (Fig. 1A).Because ND2ins1 mutants lacked a bang-sensitive phenotype, only

ND2del1 flies (hereafter referred to as ND2 mutants) were used in oursubsequent experiments.

The bang-sensitive phenotype of ND2 mutants could be aconsequence of reduced ND2 activity, or the acquisition of a novelactivity. To distinguish between these possibilities, we testedwhether restoring NADH dehydrogenase activity would rescue thebang-sensitive phenotype of ND2 mutants. Previous studies haveshown that ectopic expression of the yeast NADH dehydrogenase(Ndi1), which is composed of a single subunit, can rescue complexI deficiency in both mammalian cells and Drosophila (Cho et al.,2012; Seo et al., 1998). Thus, we tested whether expression of Ndi1could rescue the bang sensitivity of ND2 mutants. Ndi1 expressionpartially suppressed the bang-sensitive phenotype of ND2 mutants,demonstrating that the ND2del1 mutation results in a loss of ND2activity (supplementary material Fig. S2A).

Mitochondrial diseases are often associated with dramaticallyshortened lifespan (Distelmaier et al., 2009; Ugalde et al., 2007). Todetermine if this was true for ND2 mutants, we compared thelifespan of ND2 mutants and controls. We found that the medianlifespan of ND2 mutants was reduced compared with wild-typecontrols, and that survival of female ND2 mutants was shorter thanthat of male ND2 mutants (Fig. 1B). Because female ND2 mutantsexhibited stronger phenotypes than males, only females were usedin our subsequent experiments.

Another common symptom of mitochondrial disease is heatintolerance (Pieczenik and Neustadt, 2007). To test whether ND2mutants were sensitive to heat-induced paralysis, we exposed ND2mutants and controls to a temperature of 39°C, a conditionpreviously shown to result in premature heat-induced paralysis inmitochondrial mutants (Ganetzky and Wu, 1982). We found thatND2 mutants were fully paralyzed following exposure to 39°C for6 minutes, and required a prolonged recovery period after beingreturned to room temperature. In contrast, age-matched controlsfailed to undergo heat-induced paralysis under these conditions(Fig. 1C).

Mitochondrial diseases preferentially affect tissues with highenergetic demands, such as muscle and brain (Chinnery, 2000). Totest for impaired function in muscle and/or neural tissue, wemeasured the flight performance of young ND2 mutants using anestablished flight assay (Benzer, 1973; Greene et al., 2003). YoungND2 mutants displayed significantly reduced flying abilitycompared with age-matched controls (Fig. 1D). In total, theseexperiments support the hypothesis that the ND2del1 mutation affectsenergy-demanding muscle and/or brain tissue(s), mirroring deficitsfrequently seen in human mitochondrial disease.

Mitochondrial impairment has been shown to sensitizeDrosophila to hypercarbia-induced (Whelan et al., 2010) andhypoxia-induced (Feala et al., 2007) paralysis, so we tested whetherND2 mutants were also sensitive to these conditions. To induceparalysis, we exposed young and middle-aged ND2 mutants andage-matched controls to carbon dioxide or nitrogen, respectively. Wethen measured the time required for flies to recover from paralysisupon return to ambient conditions. Both young and middle-agedND2 mutants took longer to recover from exposure to hypercarbicand hypoxic conditions than controls (Fig. 1E,F). Conversely,hyperoxic conditions have been shown to shorten wild-typeDrosophila lifespan, and mitochondrial impairment enhances thiseffect (Whelan et al., 2010). To determine whether ND2 mutantswere sensitive to hyperoxic conditions, we housed young ND2mutants and controls in 100% oxygen and monitored their lifespan.As expected, ND2 mutants and controls both exhibited adramatically shortened lifespan under these conditions, but the ND2

RESEARCH ARTICLE Disease Models & Mechanisms (2014) doi:10.1242/dmm.015321

TRANSLATIONAL IMPACTClinical issueElectron transport chain (ETC) complex I dysfunction is the mostcommon cause of primary mitochondrial diseases. For example,mutations in the mitochondrial gene that encodes the complex I subunitNADH dehydrogenase subunit 2 (ND2) have been shown to be a causeof Leigh syndrome. Symptoms of Leigh syndrome include decreasedlifespan, seizures, heat intolerance and neurodegeneration. Notably,individuals with Leigh syndrome can exhibit decreased complex I activityand abundance. However, the molecular mechanisms leading from ETCdysfunction to disease pathogenesis remain unclear.

ResultsIn this study, the authors characterize a Drosophila melanogaster modelof complex I deficiency caused by a mutation in the mitochondrial-DNA-encoded gene ND2. The authors present biochemical and behavioraldata showing that ND2 mutants exhibit phenotypes that resemble Leigh syndrome. They report that these phenotypes include decreased lifespan, seizures, heat intolerance, movement deficits,neurodegeneration, and decreased complex I activity and abundance. Inaddition, the authors uncovered a defect in the ability of ND2 mutants topump protons through complex I. This defect was demonstrated by theobservation of a decrease in the efficiency of complex I, but not complexII, to power ADP conversion to ATP in ND2 mutants. The authorshypothesized that the pathogenesis of ND2 mutants resulted fromdecreased efficiency of proton pumping by complex I, leading to energydeficits, and age-related neurodegeneration. Supporting this hypothesis,the authors demonstrated decreased ATP levels and decreasedmitochondrial membrane potential in the brains of old ND2 mutants.

Implications and future directionsComposed of 45 subunits, complex I is the most intricate complex of theETC, but the functions of many of its subunits remain unclear. Theauthors’ findings show that ND2 has a crucial role in the proton pumpingmechanism of complex I, and point towards deficits in energy productionas etiological factors underlying the pathogenesis of complex I deficiency.The Drosophila model described in this study exhibits a number ofphenotypes amenable to genetic screening methodologies, and shouldtherefore provide a powerful genetic system in which to further dissectthe mechanisms underlying the pathogenesis of mitochondrial diseases.

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mutants had a significantly shorter lifespan relative to controls(supplementary material Fig. S2B).

Tissue integrity of ND2 mutantsA hallmark of mitochondrial disease is the progressive degenerationof muscle and/or neural tissue(s) (Finsterer, 2008). We thereforeanalyzed muscle and brain integrity by hematoxylin and eosinstaining of tissue from middle-aged and old ND2 mutants andcontrols. These analyses revealed no abnormalities in the flightmuscles of middle-aged or old ND2 mutants or in the brains ofmiddle-aged ND2 mutants (Fig. 2A,B). However, neurodegenerativevacuoles observed in the brains of old ND2 mutants were both

significantly larger and more numerous than those seen in age-matched controls (Fig. 2B-D), indicating that the mutants have aprogressive neurodegenerative phenotype.

To test whether ND2 mutants have subtle defects in muscleintegrity that were not evident from our histological analyses, we usedconfocal microscopy to compare cytoskeletal architecture,mitochondrial morphology and apoptotic cell death in the indirectflight muscles of ND2 mutants and controls by staining for actin,cytochrome c and cleaved caspase-3, respectively. We also includedparkin-null animals in these studies as a positive control because theyhave previously been shown to exhibit apoptotic degeneration ofindirect flight muscles, including disruption of actin organization, and

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Fig. 1. ND2 mutants exhibit behavioral abnormalities and shortened lifespan. (A) ND2 mutants exhibit mechanical-stress-induced paralysis. Both maleand female ND2del1 and ND2ins1 mutants, and male and female controls, were assayed for the length of time that flies remained paralyzed following mechanicalstress. Error bars represent standard error of the mean (s.e.m.); n=3 independent groups of 6-12 individual animals. Measurements for ND2ins1 mutants weretaken at 2, 10, 26 and 46 days of age. (B) Lifespan analysis depicting a decrease in the median lifespan of ND2 mutants. The median lifespan was 37 days forcontrol females, 47 days for control males, 23 days for ND2 mutant females and 27 days for ND2 mutant males. Error bars represent s.e.m. (P<0.0001); n=10independent groups of 12-20 animals for control and ND2 mutants. (C) ND2 mutants exhibit heat-induced paralysis. 14-day-old ND2 mutants were incubatedat 39°C, and the time for each individual fly to paralyze recorded in seconds. After 6 minutes, flies were placed in room-temperature vials, and the time torecover from paralysis recorded in seconds. Controls did not exhibit heat-sensitive paralysis over the time course of the assay (n/a). Histograms depict themedian time to paralyze and recover from paralysis (seconds); n=3 independent groups of 7-9 animals; P=4.37×10−17 and P=8.7×10−11 for paralysis andrecovery, respectively; ***P<0.001; error bars represent s.e.m. (D) ND2 mutants have reduced flight ability. 7-day-old ND2 mutants and controls were tested fortheir flying ability by measuring the distance that flies alighted when dispensed into a cylinder (P=0.001). Histograms represent the average flight index of theindicated genotypes, and error bars represent the s.e.m.; n=6 independent groups of 11-20 animals; **P<0.01. (E) ND2 mutants are hypersensitive to CO2

exposure. Histograms depict the time to recover (seconds) for control and ND2 mutant flies after hypercarbia-induced paralysis in young (3-day-old) andmiddle-aged (16-day-old) animals (n=2 groups of 12 animals; P=7.0×10−5 for 3-day-old animals; n=2 groups of 11 animals; P=0.0013 for 16-day-old animals;***P≤0.001; **P<0.01); error bars represent standard deviations. (F) ND2 mutants are hypersensitive to N2-induced hypoxia. Histograms depict the time torecover (seconds) for control and ND2 mutant flies after hypoxia-induced paralysis in young (3-day-old) and middle-aged (16-day-old) animals (n=3 groups of10-15 animals for each genotype); ***P=0.0004; *P=0.02; error bars represent the s.e.m.

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altered mitochondrial morphology (Greene et al., 2003; Klein et al.,2014). We did not detect significant differences in actin organization,mitochondrial size or cleaved caspase-3 staining between ND2mutants and aged-matched controls (supplementary material Fig. S3),thus providing further evidence that the ND2del1 mutation does notaffect the musculature. However, in agreement with previous work,we observed disruption of the actin cytoskeleton, enlargedmitochondria and cleaved-caspase-3 staining in parkin-null flightmuscle preparations (supplementary material Fig. S3).

Functional analyses of mitochondria isolated from ND2mutantsTo explore the biochemical mechanisms underlying the ND2 mutantphenotypes, we isolated mitochondria from middle-aged and oldND2 mutants and controls, and used complex-I-specific substratesto measure mitochondrial respiration. We performed theseexperiments using both limiting and saturating amounts of ADP tomeasure state 3 (ADP-stimulated) and maximal state 3 complex-I-dependent respiratory rates, respectively (supplementary materialFig. S4). As previously reported, state 3 and maximal state 3 ratesdecreased with age in control flies (Ferguson et al., 2005), and ND2mutants displayed similar kinetics of decreased activity with age

(Fig. 3A). Mitochondria isolated from either middle-aged or oldND2 mutants showed unimpaired state 3 respiration, but maximalstate 3 values were significantly decreased in ND2 mutants relativeto age-matched controls (Fig. 3A and Table 1). The ND2del1

mutation thus causes a complex-I-dependent respiratory defectspecifically under maximally demanding conditions. To ensure thatthere was no functional variability between mitochondrialpreparations caused by our isolation method, we measured complex-II- and complex-IV-dependent respiration in ND2 mutants andcontrols. We found no significant difference in complex-II-dependent respiration, complex-II-dependent ADP/O ratios orcomplex-IV-dependent respiration between ND2 mutant and controlmitochondria (Table 1). These results confirm the integrity of ourmitochondrial preparations, and the respiratory comparisons madebetween ND2 mutants and controls (Fig. 3A).

Oxidative phosphorylation couplingThe respiratory defect of ND2 mutants offered the opportunity toinvestigate the functional role of the ND2 subunit in complex Iactivity. To test whether ND2 plays a role in the proton pumpingactivity of complex I, we measured the linkage of electron transportto ATP production by determining the ADP/O ratio. Because

RESEARCH ARTICLE Disease Models & Mechanisms (2014) doi:10.1242/dmm.015321

Fig. 2. ND2 mutants exhibit progressiveneurodegeneration, but lack muscle pathology.(A) Thoracic muscle integrity was analyzed by coronalsectioning of the thorax of middle-aged (26-day-old) or old(42-day-old) ND2 mutants and controls. Representativeimages of hematoxylin- and eosin-stained thoracic musclesections are depicted. (B) Brain integrity was analyzed bycoronal sectioning of the heads of middle-aged or old ND2mutants and controls. Representative images ofhematoxylin- and eosin-stained coronal brain sections aredepicted. Arrows demark neurodegenerative vacuoles.Scale bars: 200 μm. (C) Quantification of brain vacuole sizedemonstrates an increase in vacuole size in ND2 mutants(P=4×10e−5). Vacuoles were rarely detected in the brains ofmiddle-aged ND2 mutants and were, therefore, notsubjected to quantification. Error bars represent s.e.m.;n=26 animals for ND2 mutants; n=21 animals for controls;***P<0.001. (D) Quantification of the number of brainvacuoles demonstrates an increase in vacuole number inND2 mutants (P=0.008). Vacuoles were rarely detected inthe brains of middle-aged control or ND2 mutants andwere, therefore, not subjected to quantification. Error barsrepresent s.e.m.; n=26 animals for ND2 mutants; n=21animals for controls; **P<0.01.

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electron transport is mechanistically coupled to ADPphosphorylation via the proton circuit across the inner membrane,any defect that reduces the efficiency of complex I in pumpingprotons should lead to a decrease in ADP/O. We found a significantdecrease in complex-I-dependent ADP/O ratios in both middle-agedand old ND2 mutants (Fig. 3B and Table 1). This decrease is likelynot explained by a change in the efficiency of ATP production bycomplex V, nor a proton leak in the mitochondrial membrane,because ND2 mutant ADP/O ratios were normal when respirationwas dependent on complex II (Table 1). Moreover, themitochondrial respiratory rate following depletion of ADP (state 4),an indicator of proton leak across the mitochondrial innermembrane, was unaffected by the ND2del1 mutation (Fig. 3A andTable 1). Thus, our results demonstrate that electron flow throughcomplex I is inefficiently coupled to proton pumping in ND2mutants, indicating that the ND2 subunit functions in the protonpumping mechanism of complex I.

Complex I integrityAlthough the decrease in ADP/O in ND2 mutants indicates thatND2 is required for efficient coupling of electron transfer to protonpumping, it does not explain the decrease of maximal complex-I-dependent state-3 respiration in ND2 mutants. One possibleexplanation for this decrease is a reduction in complex I abundance.

To test this model, we examined mitochondria isolated from ND2mutants and controls using blue native gels. This analysis revealedthat the abundance of fully assembled complex I was decreased inND2 mutants relative to controls, without an accompanying decreasein complex V abundance (Fig. 3C). Western blot analysis alsorevealed a decrease in the abundance of the complex I subunitNDUFS3 in ND2 mutants, but little to no change in the abundanceof the β-subunit of complex V relative to age-matched controls (Fig.3D,E and supplementary material Fig. S5). These findings suggestthat ND2 is also required for the structural integrity of complex I.

Mitochondrial membrane potential and ATP productionThe decreased maximal complex-I-dependent state-3 activity andADP/O ratio of ND2 mutants suggested that the mutants might havediminished mitochondrial membrane potential and consequentlyreduced ATP synthesis. To test these possibilities, we measuredmitochondrial membrane potential in dissociated neurons from oldND2 mutants and controls, and we compared ATP levels in thethoraces and heads of old ND2 mutants and controls. We detected adecrease in the fraction of cells with polarized mitochondria in thebrains of old ND2 mutants relative to controls, and reduced ATPabundance in ND2 mutants that was specific to heads (Fig. 4A-C).These findings suggest that the ND2del1 mutation affects the protonpumping activity of complex I, which reduces the proton gradient

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Fig. 3. ND2 mutants exhibit decreased mitochondrial function. (A) Mitochondrial complex-I-dependent respiration rates were measured in mitochondriaisolated from middle-aged (14-day-old) or old (30- to 35-day-old) ND2 mutant or controls, revealing age-related declines in both state 3 and maximal state 3values within genotypes (P=0.08 and P=0.03 for comparison of middle-aged and old control or ND2 mutant state 3 values, respectively; P=0.001 and P=0.003for comparison of middle-aged and old control or ND2 mutant maximal state 3 values, respectively). State 4 respiration was similar under all conditions tested.No differences were found for state 3 values between control and ND2 mutants at either middle or old ages. However, maximal state 3 values were decreasedin both middle-aged and old ND2 mutants when compared with age-matched controls (P=0.04 for middle-aged and P=0.003 for old animals). Error barsrepresent standard deviations; n=3 independent mitochondrial preparations; **P<0.01; *P<0.05; #P<0.1. (B) Oxidative phosphorylation efficiency (ADP/O) isdecreased in ND2 mutants relative to controls at both middle and old ages (P=0.02 and P=0.01, respectively). Error bars represent standard deviations; n=3independent mitochondrial preparations; *P<0.05. (C) Blue native gel/complex I in-gel activity analysis of mitochondria isolated from middle-aged (14-day-old)ND2 mutants and controls reveals similar levels of complex V in both genotypes, but decreased fully assembled complex I abundance in ND2 mutants.(D) Western blot analysis of protein extracts from 14-day-old ND2 mutants and controls reveals a specific decrease in the abundance of the complex I subunitNDUFS3, with no decrease in the complex Vβ subunit. (E) Quantification of western blot data for NDUFS3 (n=3 independent tissue extracts; P=0.003) andcomplex Vβ (n=3 independent tissue extracts; P=0.46) levels normalized to actin, and expressed as the % of control values. Error bars represent s.e.m.;**P<0.01.

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available for ATP generation, leading to energy deficiency in thebrains of ND2 mutants.

Reactive oxygen species damageMitochondrial complex I deficiency is believed to trigger theformation of reactive oxygen species (ROS), and increased ROSproduction is a widely believed cause of the symptomsaccompanying complex-I-associated diseases (Lin et al., 2012). Totest whether the phenotypes of ND2 mutants could derive fromexcessive ROS production, we measured the abundance of acommon byproduct of oxidative stress: 4-hydroxynonenal (4-HNE)-containing protein adducts. Levels of 4-HNE protein adducts areelevated under conditions of increased oxidative stress (Poli andSchaur, 2000), and their abundance can be easily measured using anantiserum that detects 4-HNE adducts, as has been previously shownin Drosophila (Sun et al., 2014; Tsai et al., 1998). The use of thisantiserum on a western blot of protein extracts from ND2 mutantsand age-matched controls revealed no significant alteration in theabundance of 4-HNE protein adducts between these genotypes inyoung, middle-aged or old animals (supplementary material Fig.S6). These data argue that the phenotypes of ND2 mutants are not aconsequence of increased ROS-mediated damage.

DISCUSSIONA number of hypotheses have been proposed to explain thepathological consequences of complex I deficiency. Among the mostenduring of these models are that complex I deficiency results in theelevated production of toxic ROS and/or an energy deficit (Torracoet al., 2009; Lin et al., 2012). To address these and other possiblemodels of pathogenesis, we studied a Drosophila strain with a

mutation affecting the mitochondrial-encoded respiratory chaincomplex I subunit ND2. We show that ND2 mutants exhibit manyof the behavioral and histological characteristics of mitochondrialdisease, suggesting that ND2 mutants represent a valid model of thisspectrum of disorders. Moreover, our work shows that ND2 mutantshave diminished ATP abundance, mitochondrial membranepotential, complex I abundance, and complex I activity. However,we detected no evidence of increased ROS in ND2 mutants. Thesefindings support a model in which the pathogenesis accompanyingcomplex I deficiency is caused by diminished energy production.

Several of our findings contrast with previously published workusing other models of complex I deficiency. For example, studies ofNdufs4 knockout mice, and transgenic mice harboring a pathogenicpoint mutation in ND6, revealed decreased maximal complex Iactivity, and increased levels of ROS-mediated damage (Quintanaet al., 2010; Lin et al., 2012). These studies suggest that ROS is acontributor to the pathogenesis of complex I deficiency. However,ROS levels are thought to increase as a result of decreased basalcomplex I activity (Turrens, 2003). Therefore, it is possible that ND2mutants do not show changes in ROS-mediated damage because oftheir normal basal complex I activity, whereas the Ndufs4 and ND6mouse models might exhibit increased ROS-mediated damage as aresult of decreased basal complex I activity. Further analyses of thebasal activity of complex I in these mouse models of complex Ideficiency will be required to test this hypothesis. Anotherdiscrepancy of our work concerns the mild phenotypes of ND2mutants relative to other fly and mouse models of complex Ideficiency. We believe this discrepancy is best explained bydifferences in the severity of the mutations analyzed in eachrespective study. The ND2del1 allele is an in-frame deletion predicted

RESEARCH ARTICLE Disease Models & Mechanisms (2014) doi:10.1242/dmm.015321

Table 1. Mitochondrial oxygen consumption measurementsGenotype Measurement Respiration rate (nmol O2 /min/mg) or ratio Standard deviation P-value

Middle-aged flies (14 days old)Control CI:State 3 86.2 ±27.2 0.50 ND2del1 CI:State 3 99 ±14.6Control CI:State 4 27.2 ±5.3 0.32 ND2del1 CI:State 4 32.5 ±6.4Control CI:Max.State3 180.2 ±13.6 0.04 ND2del1 CI:Max.State3 150.5 ±13.4Control CI:ADP/O 2.7 ±0.2 0.02 ND2del1 CI:ADP/O 1.9 ±0.1Control CII:State 3 42 ±1.95 0.19 ND2del1 CII:State 3 49.9 ±6.2Control CII:Max.State3 43.2 ±4 0.16 ND2del1 CII:Max.State3 52.2 ±7.2Control CII:ADP/O 1.7 ±0.1 0.42 ND2del1 CII:ADP/O 1.6 ±0.1Control CIV 426.2 ±100 0.39 ND2del1 CIV 546.2 ±198.25

Old flies (35 days old)Control CI:State 3 48.4 ±5.3 0.45 ND2del1 CI:State 3 54.5 ±12.5Control CI:State 4 19.6 ±9.85 0.39 ND2del1 CI:State 4 25.7 ±5.65Control CI:Max.State 3 119.5 ±25.2 0.003 ND2del1 CI:Max.State3 82.6 ±8.8Control CI:ADP/O 2.9 ±0.1 0.01 ND2del1 CI:ADP/O 1.8 ±0.3Control CIV 313 ±75.4 0.32 ND2del1 CIV 406 ±62.2

Respiratory rates are expressed in units of O2 consumed nmol/min/mg (n=3 independent mitochondrial preparations). ADP/O ratios are unit-free (n=3independent mitochondrial preparations). The corresponding standard deviations and P-values for each measurement are indicated. CI, complex I; CII,complex II; CIV, complex IV.

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to make a full-length protein lacking three amino acids. Thismutation could therefore represent a hypomorphic allele of ND2. Bycontrast, many animal models of complex I deficiency, withphenotypes more severe than those of ND2 mutants, harbor severeloss-of-function or null alleles (Cho et al., 2012; Kruse et al., 2008;Zhang et al., 2013). In further support of this model, the magnitudesof the decreases in complex I activity and abundance in ND2del1

mutants are similar to those seen in Caenorhabditis elegans withpartial inactivation of complex I subunits (Falk et al., 2009). Therelatively mild phenotypes of ND2 mutants potentially make themwell-suited to genetic modifier screens aimed at the identification ofcurrently unknown genetic factors that influence the phenotypesassociated with complex I deficiency.

In addition to exploring the pathogenic mechanisms underlyingcomplex I deficiency, our work provides insight into the role of ND2in complex I function. Although the ND2 subunit has beenhypothesized to pump protons based on its membrane localizationand sequence homology to a family of cation antiporters (Efremovet al., 2010), only a study of the bacterial ND2 homolog, NuoN, hasprovided support for this model (Amarneh and Vik, 2003). Ourfinding that the ND2del1 mutation results in uncoupling of protonpumping from electron transfer [i.e. decreased oxidativephosphorylation efficiency (ADP/O) with normal basal complex-I-mediated electron flux] strongly supports a direct role for ND2 inthe proton pumping mechanism of eukaryotic complex I.

Although our work advances the understanding of theconsequences of ND2 deficiency, and the functional role of the ND2subunit in eukaryotes, it also raises several questions. One importantquestion raised by our work concerns the mechanisms underlyingthe progressive nature of the ND2 mutant phenotypes. One possibleexplanation of the progressive phenotypes of ND2 mutants is thatenergy deficiency does not occur until a critical threshold ofcomplex I activity is crossed. As previously reported (Ferguson etal., 2005), and confirmed in our current study, complex I activitynaturally declines with age in Drosophila. This decline in complexI activity might not reach the point where it results in an energydeficiency over the natural lifespan of Drosophila. However, theND2 mutation exacerbates this natural decline in complex I activity,and thus could account for the progressive nature of the ND2 mutantphenotypes.

Another important question raised by our study is why a systemicdeficit in complex I function results in selective neurodegeneration.One possible explanation could be that neurons are more sensitive toloss of mitochondrial function because they rely primarily onoxidative phosphorylation for ATP, whereas other tissues, such asmuscle, are capable of using both oxidative phosphorylation andglycolysis (Chan et al., 2009). Although neural selective vulnerabilityis not uncommon in mitochondrial-associated diseases, includingLeigh syndrome, Leber’s hereditary optic neuropathy and Alpers’syndrome (Bianchi et al., 2011; Koopman et al., 2013), further studieswill be required to understand the mechanisms underlying theprogressive tissue-specific nature of mitochondrial disease.

In summary, our work suggests an important role for ND2 in theproton pumping mechanism of complex I, and provides a geneticallytractable animal model of complex I deficiency. This model shouldprove a valuable tool in which to further explore the pathologicalmechanisms underlying human mitochondrial disease.

MATERIALS AND METHODSDrosophila strains and maintenanceAll Drosophila strains were maintained on standard cornmeal/molassesmedium at 25°C with a 12-hour light-dark cycle. The ND2del1 and ND2ins1

stock was obtained from the laboratory of Dr Patrick O’Farrell (Universityof California, San Francisco, CA) (Xu et al., 2008). The isogenic w1118 stockwas obtained from the Bloomington Drosophila Stock Center at IndianaUniversity. To control for differences in nuclear genetic background, weoutcrossed ND2 mutants to the w1118 strain. F1 offspring derived fromcrossing ND2 mutant females to w1118 males were used as the experimentalgroup, given that they inherit mtDNA from the ND2 mutant strain; F1

offspring derived from crossing ND2 mutant males to w1118 females wereused as the control group, given that they inherit mtDNA from the w1118

strain. The homoplasmic status of the ND2del1 mutation from the outcrossedND2 mutant strain was reconfirmed by PCR and restriction digest analysis(data not shown). Parkin-null animals harbored the previously describedpark25 null allele (Greene et al., 2003), and were composed of the genotypeCyO/IF; park25/park25. Previously described transgenic flies capable ofexpressing yeast Ndi1 were obtained from Dr David Walker (Cho et al.,2012).

Mechanical-stress-induced paralysisFlies were assayed for bang sensitivity using a modification of a previouslypublished protocol (Ganetzky and Wu, 1982). Briefly, flies were vortexedat maximum speed for 10 seconds in inverted glass vials containing cottonstoppers, and the time required for each individual animal to right itself wasrecorded (supplementary material Movie 1).

Heat-induced paralysis14-day-old flies were assayed for heat-induced paralysis by placing groups offive to ten animals into pre-warmed vials maintained at 39°C. The time for the

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Fig. 4. ND2 mutants exhibit decreased mitochondrial membranepotential and ATP production. (A) Neurons from 30- to 35-day-old ND2mutants and controls labeled with the mitochondrial membrane potential dyeTMRE were analyzed by flow cytometry. Histograms depict the percentmaximal number of cells measured at each TMRE fluorescence intensity.(B) Quantification of the average percent of isolated neurons exhibitingmitochondrial membrane-potential-dependent fluorescence above 103

arbitrary units (% polarized mitochondria) is shown for control and ND2mutants (P=0.004). Error bars represent standard deviation; n=3independent brain preparations; **P<0.01. (C) ATP levels were determined inheads of 35-day-old ND2 mutants and controls, and displayed relative tocontrol values (P=0.05). Error bars represent s.e.m.; n=3 groups of 5individuals; *P≤0.05 for a one-tailed t-test.

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flies to become paralyzed was recorded. After exposure to 39°C for 6 minutesthe animals were then placed in new room-temperature vials (~20°C), and therecovery time of ND2 mutants from paralysis recorded.

LifespanLifespan assays were performed using vials containing 12-20 flies. Flieswere transferred to new vials every 2 days throughout the assay period. Atleast 60 flies were monitored for each experiment. For sex-specific lifespanassays, ND2 mutant males and females or gender- and age-matched controlflies were separated within 1 day of eclosion, and the assay conducted asdescribed above.

FlightFlight assays were performed as previously described (Benzer, 1973; Greeneet al., 2003). Briefly, an acetate sheet was divided into five parts, coated withvacuum grease and inserted into a 1 liter graduated cylinder. 7-day-old flieswere gently tapped into a funnel at the top of the cylinder, and became stuckto the vacuum grease where they alighted. The acetate sheet was removed,and the number of flies in each section was counted. Flies that alighted in ahigher section of the acetate sheet received a correspondingly higher valuein scoring. The number of flies alighting in the top section was summed andmultiplied by four, the number of flies alighting in the second highest sectionwas summed and multiplied by three, and so on. Finally, the weighted sumfor the entire group of flies was determined, and normalized to four timesthe total number of flies used in the assay (the maximum possible score),and this normalized value was reported as the flight index. Flies that alightedat the top of the cylinder received a flight index score of 1, whereas flies thatfell to the bottom of the cylinder received a flight index score of 0. At leastsix groups of 11-20 flies per genotype were tested.

Sensitivity to CO2 exposureYoung (3-day-old) or middle-aged (16-day-old) control or ND2 mutantswere exposed to 100% CO2 (Praxair) on a gas-permeable mesh pad for 1minute, rendering them completely immobile. The gas was then shut off,leaving the flies exposed only to room air, and the time to recover fromparalysis was measured.

Hypoxia sensitivityThree separate sets of 10-15 age-matched young (3-day-old) and middle-aged (16-day-old) flies were placed into 70 ml chambers and 100% N2

(Praxair) certified at less than 10 ppm O2 was flowed through the chamberat 350 ml/minute for 1.5 minutes to induce paralysis. After 30 seconds, flieswere exposed to room air at the same rate, and the time from theintroduction of room air to recovery from paralysis was recorded.

Hyperoxia sensitivityFemale flies were collected within 48 hours of eclosion and placed into fourvials of 25 flies per genotype. Vials were then placed into a temperature-controlled hyperoxia chamber and maintained in a 100% O2 environment at25°C. Flies were transferred to new vials every 24 hours and scored forsurvival.

Confocal microscopyIndirect flight muscles were dissected from 40-day-old control or ND2mutant animals, or from 7-day-old parkin-null animals in phosphatebuffered saline (PBS). Dissected muscles were placed in 0.3% Triton X-100(Sigma), 4% paraformaldehyde (Ted Pella Inc.) in PBS, and rotated at roomtemperature for 40 minutes. Muscles were then washed twice in PBS andplaced in blocking buffer [0.3% Triton X-100, 5% normal goat serum(Fisher Scientific) in PBS] for 45 minutes. Muscles were then incubatedwith a mouse primary antibody raised against cytochrome c (BD Labs#556433) (1/1000), and a rabbit primary antibody raised against cleavedcaspase-3 (Cell Signaling #9661) (1/400) for 2 days at 4°C in 0.3% TritonX-100 and 2.5% normal goat serum in PBS. Tissue was then washed twicein 0.3% Triton X-100 in PBS, for 20 minutes each, at room temperature. Thetissue was then placed in secondary antibodies: mouse-Alexa-Fluor-647 andrabbit-Alexa-Fluor-488 (Molecular Probes), and incubated for 1 hour

rotating at room temperature. At 20 minutes prior to removal fromsecondary antibodies, phalloidin-568 (Life Technologies) was added at aconcentration of 1/400. Tissue was removed and washed two times for 10minutes each in 0.3% Triton X-100 in PBS. The muscles were then placedin PBS containing DAPI (Sigma-Aldrich) at 2 μg/ml for 5 minutes. Cellswere washed in PBS for 10 minutes, mounted between two glass slides withFluoromount (Sigma-Aldrich), and imaged using an Olympus FV-1000 witha 60× lens and a 4× digital zoom.

Image quantificationImages of muscles stained with cytochrome c were opened in ImageJ,converted to 8-bit, thresholded, watershed, and analyzed using the analyzeparticle tool. The average perimeter of distinct mitochondria was used as anindicator of mitochondrial morphology and mass. For cleaved-caspase-3staining, the number of cleaved-caspase-3-positive myocytes, as indicatedby DAPI-positive/cytochrome c/cleaved-caspase-3-positive staining, weremanually counted.

Tissue preparation and sectioningMiddle-aged (14-day-old) or old (42-day-old) flies were anesthetized,mounted in fixing collars and placed in Carnoy’s fixation solution (10% aceticacid; 30% chloroform; 60% absolute ethanol) for 3.5 hours. Fixed flies werethen placed in 95% ethanol twice for 30 minutes each, in 100% ethanol for 45minutes, then in methyl benzoate overnight. Flies were then incubated in a 1:1mixture of methyl benzoate:paraffin for 1 hour at 60°C, transferred to 100%liquid paraffin and incubated at 60°C for 30 minutes, then transferred to fresh100% liquid paraffin four times for 15-30 minutes each. The flies were nexttransferred to 100% liquid paraffin in a plastic mold, placed at 60°C for 15minutes, and the paraffin was allowed to harden at room temperature. 5-μm-thick coronal histological sections were cut on a Shandon Finesse 325Microtome, and processed for hematoxylin and eosin staining. Images werecollected on a Nikon Optiphot-2 using a 20× objective. Vacuole size wasquantified in ImageJ (Abramoff et al., 2004) by converting pixel values intomicrometers (0.439 μm/pixel). Only vacuoles present in at least twoconsecutive sections were quantified to avoid potential artifacts derived fromtissue sectioning.

Mitochondrial preparationsMitochondria were obtained from 0.25-0.5 g of 14-day-old or 30- to 35-day-old flies. Flies were chilled on ice, and then homogenized using a chilledPotter/Elvehjem homogenizer with 10 manual strokes in 20 ml ofhomogenization buffer (200 mM mannitol, 70 mM sucrose, 5 mM MOPS,2 mM EDTA; 0.4% defatted BSA; pH 7.4), avoiding shearing. Thehomogenate was then filtered through cotton gauze, and spun at 300 g for 4minutes at 4°C. The supernatant was then filtered through gauze and spunat 10,000 g for 10 minutes at 4°C. The pellet was resuspended in 10 ml ofresuspension buffer (200 mM mannitol, 70 mM sucrose, 5 mM MOPS, 2mM EDTA; pH 7.4), and spun at 10,000 g for 10 minutes at 4°C. Themitochondrial pellet was then resuspended in 10-50 µl of final buffer (200mM mannitol, 70 mM sucrose, 5 mM MOPS; pH 7.4), to a finalconcentration of ~50 mg protein/ml. 150-300 mg of mitochondria was usedfor oxidative phosphorylation assays, and 250 mg for blue native gelanalysis. All reagents were obtained from Sigma (St Louis, MO).

Oxidative phosphorylation assaysOxygen consumption of isolated mitochondria was monitored using a Clark-type electrode (Oxytherm with analysis software Oxyg32, HansatechInstruments, Pentney, Norfolk, UK) by an adaptation of previous methods(Kayser et al., 2001). Briefly, 150-300 μg of purified mitochondria werestirred in 500 μl of assay buffer (100 mM KCl, 50 mM MOPS, 1 mMEGTA, 5 mM potassium phosphate, 1 mg/ml defatted BSA; pH 7.4) at30°C. 10 mM malate and 20 mM pyruvate were added as complex-I-specificelectron donor substrates, 20 mM succinate was added as a complex-II-specific electron donor substrate or 25 mM TMPD and 250 mM ascorbate(pH 7.0) were added as complex-IV-specific electron donor substrates. Alow dose of ADP (90.7 nmol) was added to determine state 3 mitochondrialrespiration rates and, following depletion of ADP, ensuing state 4 respiration

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rates. A saturating dose of ADP (1 μmol) was used to assess maximalcomplex-I-dependent electron transport capacity under phosphorylatingconditions (maximal state 3). Saturating amounts of TMPD/ascorbate wereused to measure complex IV activity. ADP/O was defined as the number ofADP molecules phosphorylated per oxygen atom reduced to water, andADP/O was calculated by dividing the amount of ADP added (90.7 nmol)by the amount of oxygen consumed between the time of ADP addition andthe onset of state 4 (supplementary material Fig. S4).

ATP determinationHeads or thoraces were sectioned from five 35-day-old flies and homogenizedin 6 M guanidine HCl (Sigma); 10 mM TRIS (Sigma), pH 7.3, as previouslydescribed (Xu et al., 2008). ATP content was measured using an ATPdetermination kit (Molecular Probes, Eugene, OR). Protein abundance wasmeasured using a Bradford protein assay, and ATP abundance was normalizedto total protein abundance as previously described (Xu et al., 2008).

Mitochondrial membrane potentialMeasurement of neural mitochondrial membrane potential from 30- to 35-day-old flies was conducted as previously described (Burman et al., 2012),except that dissections were performed in supplemented DME/Ham’s F-12High Glucose media lacking phenol red (Sigma), and all incubation stepswere carried out at 25°C. In addition, 10 nM tetramethylrhodamine ethylester (TMRE) was utilized throughout the protocol. The % of cells harboringpolarized mitochondria was determined using FloJo software (Treestar Inc.,Ashland, OR), and defined as the % of cells exhibiting relative TMREfluorescence above 103 arbitrary fluorescence units, as previously described(Burman et al., 2012). All flow cytometry measurements were performed ona Becton-Dickinson LSR II flow cytometer.

Blue native gel electrophoresis and in-gel activity stainingAssays were carried out using mitochondrial preparations from 14-day-oldflies as previously described (Suthammarak et al., 2009; Wittig et al., 2006).All reagents were purchased from Sigma.

Western blots14-day-old whole flies were flash-frozen in liquid nitrogen andhomogenized with a pestle in 2× lysis buffer (50 mM TRIS, pH 8.0; 300mM NaCl; 2 mM EDTA; 1% SDS; 2% Triton X-100) with proteaseinhibitor cocktail (Sigma). Homogenates were centrifuged at 21,000 g for 5minutes, and the supernatant subjected to western blot processing andanalysis. Blots were labeled with monoclonal antibodies to actin (Millipore#MAB1501) diluted 1/25,000, the β-subunit of ATP synthase (Invitrogen#A21351) diluted 1/1500, the NDUFS3 subunit of complex I (Abcam#17D95) diluted 1/800, and an anti-HNE fluorophore antibody (Calbiochem393206) diluted 1/2500. This antibody detects a stable, crosslinking productbetween two lysyl residues and 4-HNE: 2:1 Nα-acetyllysine-HNEfluorophore. The antigen is taken as a proxy for accumulated 4-HNEdamage, or accumulated ROS damage in general (Sun et al., 2014; Tsai etal., 1998). All reagents were purchased from Sigma unless otherwise noted.Western blot band intensities were quantified using ImageJ gel analysistools, and the intensities were expressed as the % of control values followingnormalization to actin (Abramoff et al., 2004). Complete western blots areshown in supplementary material Fig. S5 as a demonstration of samples runin parallel.

StatisticsUnless otherwise stated, statistical significance tests were calculated usingan unpaired two-tailed Student’s t-test.

AcknowledgementsWe thank Dr Patrick H. O’Farrell (University of California, San Francisco) forproviding the Drosophila ND2 mutant stocks (Xu et al., 2008); Dr David Walker(University of California, Los Angeles) for providing a transgenic Drosophila strainexpressing the yeast Ndi1 protein; Dr Ying-Tzang Tien (Department of Pathology,University of Washington) for histological staining of Drosophila tissue sections; DrRenata de Lima Sales Goncalves (Buck Institiute of Aging, Novato, USA), LaurieAndrews and Selina Yu (University of Washington) for technical assistance; Dr

Ruth E. Thomas, and Tarannum and Afshan Dhillon, Stephanie N. Yu and RyanHau (University of Washington) for maintaining Drosophila stocks. We thank DrEvelyn Vincow for critical reading of the manuscript.

Competing interestsThe authors declare no competing financial interests.

Author contributionsJ.L.B., L.S.I., A.M.W., M.K., E.-B.K., P.G.M., M.M.S. and L.J.P. designed research;J.L.B., L.S.I., A.M.W., W.S. and E.-B.K. performed research; J.L.B., L.S.I., W.S.,E.-B.K., P.G.M., M.M.S. and L.J.P. analyzed data; J.L.B., L.S.I., E.-B.K., M.M.S.,P.G.M. and L.J.P. wrote the paper.

FundingThis work was supported by a Parkinson Society Canada Research Fellowship (toJ.L.B.), and a subsequent Canadian Institute of Health Research Fellowship (toJ.L.B.), National Institute of Neurological Disorders and Stroke FellowshipF31NS071857 (to L.S.I.), National Institute of Health grant 1RO1GM086394 andgrant 5R01GM104990 (to L.J.P.) and a grant from the Muscular DystrophyAssociation (to L.J.P.). L.S.I. and A.M.W. were supported on the GeneticApproaches to Aging Training Grant T32AG000057 (to Peter Rabinovitch). W.S.was supported by a NW Mitochondrial Guild Postdoctoral Fellowship. Thesestudies were also supported by the University of Washington School of Medicineand Department of Pathology (M.K.).

Supplementary materialSupplementary material available online athttp://dmm.biologists.org/lookup/suppl/doi:10.1242/dmm.015321/-/DC1

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