4 Mitochondrial ATP Synthase: Architecture, Function and Pathology 75

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2012 plaatse. Aula Major Paranimfen: 024-8444501 An Jonckheere ITNODIGING Merei Huigsloot mijn proefschrift Stefanie Henriet Comeniuslaan 2 Zwaluwstraat 155 6525 HP Nijmegen 6525 HP 6541 NV Nijmegen U om 10.30 uur precies application with special application with assay development and assay development [email protected] voor het bijwonen van de bijwonen van de voor het openbare verdediging van verdediging van openbare op maandag 17 september op maandag 17 ITOCHONDRIAL ITOCHONDRIAL [email protected] [email protected] Aansluitend bent u van harte uitgenodigd op de receptie ter Radboud Universiteit Nijmegen emphasis on human complex V emphasis on human M : An I. Jonckheere EDICINE M

ITOCHONDRIAL M assay development and application application and development assay with special emphasis on human complex V human complex emphasis on with special

Mitochondrial Medicine: assay development and application with special emphasis on human complex V An I. Jonckheere MITOCHONDRIAL MEDICINE:

assay development and application with special emphasis on human complex V

An Jonckheere

Mitochondrial Medicine: assay development and application with special emphasis on human complex V Thesis Radboud University Nijmegen with a summary in Dutch

The research presented in this thesis was performed at the Department of Pediatrics, Radboud University Nijmegen Medical Center, The Netherlands.

ISBN 978-94-6191-343-2

Cover: artistic representation of a mitochondrion embedding the five OXPHOS complexes, and detailed structure of complex V or ATP synthase.

Cover design and lay-out: An Jonckheere and Paul Van Herck

Printed by Ipskamp Drukkers Nijmegen

Mitochondrial Medicine:

assay development and application with special emphasis on human complex V

Proefschrift

ter verkrijging van de graad van doctor aan de Radboud Universiteit Nijmegen op gezag van de rector magnificus prof. mr. S.C.J.J. Kortmann, volgens besluit van het college van decanen in het openbaar te verdedigen op 17 september 2012 om 10.30 uur precies

door

An Isabel Jonckheere

geboren op 15 maart 1976 te Gent (België) Promotor Prof. dr. J.A.M. Smeitink

Copromotor Dr. R.J.T. Rodenburg

Manuscriptcommissie Prof. dr. M.A.A.P. Willemsen (voorzitter) Prof. dr. G. Vriend Prof. dr. F.A. Wijburg (AMC Amsterdam)

Zo volmaakt, zoveel geluk en toch nog maar zo klein. (uit: Als je slaapt – Glennis Grace)

Aan mijn ouders

Voor Paul

Contents

List of abbreviations 9

Part I General introduction

Chapter 1 Mitochondria and disease 13

Part II Characterization of human cell lines with mitochondrial dysfunction

Chapter 2 A high-throughput assay to measure oxygen consumption in digitonin-permeabilized cells of patients with mitochondrial disorders 29

Chapter 3 Toolbox for mitochondrial biochemical characterization of myogenic cell lines from Leigh (-like) patients 51

Part III Complex V in health and disease

Chapter 4 Mitochondrial ATP synthase: architecture, function and pathology 75

Chapter 5 A novel mitochondrial ATP8 (MT-ATP8) gene mutation in a patient with apical hypertrophic cardiomyopathy and neuropathy 111

Chapter 6 Restoration of complex V deficiency caused by a novel deletion in the human TMEM70 gene normalizes mitochondrial morphology 131

Chapter 7 Functional assays preceding exome sequencing: a rational strategy 161

Part IV General discussion

Chapter 8 General discussion, future perspectives and conclusions 187

Summary 199

Samenvatting (Dutch summary) 205

Dankwoord/ Acknowledgements 211

Curriculum Vitae 215

List of publications 219

Appendix 223

List of abbreviations

ANT adenine nucleotide transporter ADP adenosine diphosphate ATP adenosine triphosphate BN blue native Bp base pair CCCP carbonyl cyanide 3-chlorophenyl hydrazone cDNA coding DNA C.I - C.V complex I – complex V CN clear native CO2 carbon dioxide CoQ coenzyme Q or ubiquinone Cox8 mitochondrial targeting sequence of cytochrome c oxidase subunit 8A COX cytochrome c oxidase or complex IV CS citrate synthase Cyt c cytochrome c ∆ψm mitochondrial membrane potential DNA deoxyribonucleic acid FADH2 flavine adenine dinucleotide (reduced form) gDNA genomic DNA GFP green fluorescent protein HEK human embryonic kidney HHH-syndrome hyperammonemia-hyperornithinemia-homocitrullinuria - syndrome IGA in-gel activity kDa kilo Dalton MEGS mitochondrial energy-generating system MILS maternally inherited Leigh syndrome MRI magnetic resonance imaging Mt mitochondrial mtDNA mitochondrial DNA NAD(H) nicotinamide adenine dinucleotide (reduced form) NARP neuropathy, ataxia and retinitis pigmentosa nDNA nuclear DNA OSCP oligomycin sensitivity-conferring protein OXPHOS oxidative phosphorylation PAGE polyacrylamide gel electrophoresis PBS phosphate buffered saline PCR polymerase chain reaction PDHc pyruvate dehydrogenase complex Pi inorganic phosphate POLG polymerase gamma RFU relative fluorescence units RNA ribonucleic acid ROCR relative oxygen consumption rate RT-PCR reverse transcriptase PCR SDS sodium dodecyl sulphate TCA cycle tricarboxylic acid cycle TMEM70 transmembrane protein 70 tRNA transfer RNA

PART I

GENERAL INTRODUCTION

Mitochondria and disease

MITOCHONDRIAL BIOLOGY: THE OXPHOS SYSTEM

Mitochondria are subcellular organelles, present in virtually every human cell and essential for survival (1). The organelles have a double-membrane and are equipped to produce ATP and provide this as the energy source for intracellular metabolic pathways (1-2) (Figure 1).

Mitochondria are also involved in cell signaling, especially signaling for apoptotic cell death, and host other metabolic pathways including the fatty acid β-oxidation and lipid and cholesterol synthesis (1).

Figure 1 The mitochondrion. Mitochondria are enclosed by a double membrane. The inner membrane is composed of two subdomains: the inner boundary membrane and the cristae membrane. Cristae are invaginations of the inner membrane that are connected to the inner boundary membrane by narrow tubular structures, so-called crista junctions (3).

The mitochondrial oxidative phosphorylation (OXPHOS) system is the final biochemical pathway in the oxidative production of ATP (4). Enzymes or enzyme complexes directly involved in the generation of chemical energy by OXPHOS also comprise the pyruvate dehydrogenase complex (PDHc) and the tricarboxylic acid cycle (Figure 2). The OXPHOS system consists of five multiprotein complexes (I (EC 1.6.5.3), II (EC 1.3.5.1), III (EC

1.10.2.2), IV (EC 1.9.3.1), and V (EC 3.6.3.14)) and two electron carriers – coenzyme Q and cytochrome c -, embedded in the inner mitochondrial membrane (2, 4). The main function of the system is the coordinated transport of electrons and protons, which leads to the production of ATP (4). The first two events of respiration, i.e. electron transfer and proton

Chapter 1 pumping, are carried out by the mitochondrial respiratory chain, composed of the first four complexes (C.I-IV) (2, 4).

Figure 2 The energy metabolism. One of the main functions of mitochondria is to produce energy in the form of ATP by fatty acid oxidation, the oxidation of acetyl-CoA in the tricarboxylic acid (TCA) cycle, and oxidative phosphorylation (OXPHOS). The TCA cycle produces four NADH which are oxidized by complex I and one FADH2 which is oxidized by complex II. The redox reactions in complexes I to IV (the respiratory chain) generate a proton gradient across the inner mitochondrial membrane. The energy created by this proton electrochemical gradient is used by complex V to synthesize ATP from ADP. The OXPHOS complexes are, except for complex II, under dual genetic control, the number of subunits encoded by the nuclear (n) DNA and the mitochondrial (mt) DNA is indicated. Adapted from Zschocke/ Hoffman. Vademecum Metabolicum.

Basically, the respiratory chain accepts electrons from NADH (complex I) and FADH2

(complex II) and subsequently transfers them through a series of oxidation-reduction reactions (via coenzyme Q, complex III and cytochrome c) to molecular oxygen to produce water (complex IV). Simultaneously this reaction is coupled to the translocation of protons across the inner membrane (complex I, III, IV) (5-6). Synthesis of ATP from ADP is the second fundamental reaction of the OXPHOS system, a process catalyzed by complex V

Mitochondria and disease

(ATP synthase) (2). Electron transfer across the respiratory chain and ATP synthesis are coupled (2). The respiratory chain works as a proton pump which generates a proton gradient and a membrane potential of about -180 mV across the inner membrane (2).

Complex V uses the energy created by the proton electrochemical gradient to phosphorylate

ADP to ATP in the mitochondrial matrix (7-8). During this process the proton gradient is decreased and this activates respiration, i.e. electron transfer in order to maintain the membrane potential (5).

The formation of the OXPHOS system is under the control of two separate genetic systems, the nuclear genome and the mitochondrial genome (mitochondrial DNA (mtDNA)). Four of the five OXPHOS complexes (I, III, IV and V) contain both nuclear-encoded and mtDNA- encoded proteins. Human mtDNA is a circular double stranded molecule of 16.569 base pair

(bp) and codes for 22 transfer and two ribosomal RNAs, and 13 messenger RNAs (1).

Transmission of mtDNA occurs through the maternal lineage. There are hundreds to thousands of mtDNA molecules per cell, and mostly their sequence will be identical

(homoplasmy) (1). The coexistence of wild type and mutant mtDNA is called heteroplasmy.

During mitosis mtDNA molecules are randomly distributed to daughter cells (mitotic segregation). This means that the degree of heteroplasmy can change drastically between different generations of families, but also between tissues in one affected person, and even in one tissue through time. The OXPHOS system malfunctions if a critical number of mutated mtDNAs is present. This is called the threshold effect (4). MtDNA remains entirely dependent upon the nucleus for provision of enzymes for replication, repair, transcription and translation

(1). The protein subunits of the OXPHOS complexes are assembled by a set of assembly factors, encoded by the nuclear DNA (1). Accordingly, many different mutations in mtDNA- and nuclear DNA-encoding subunits, components or regulators of the OXPHOS system can produce a broad spectrum of OXPHOS diseases (9-10). Mitochondrial diseases are indeed among the most common genetic disorders with a prevalence of at least 1 in 5000 (11).

Chapter 1

MITOCHONDRIAL BIOCHEMISTRY

Because of the heterogeneous clinical presentation and organ involvement, the diagnosis of a mitochondrial disease is, in the group of inborn errors of metabolism, not straightforward.

Nevertheless, significant progress has been made over the past few years in the diagnostic possibilities for OXPHOS defects (12-14). Gold standard in the diagnostic process is a fresh muscle biopsy from the M. quadriceps in which the mitochondrial energy-generating system

(MEGS) capacity is studied. This can be done by measuring oxidation rates of [1-14C]- pyruvate + malate in the presence and absence of adenosine diphosphate (ADP), [1-

14C]pyruvate + carnitine, and the ATP production rate from oxidation of pyruvate + malate in fully coupled muscle mitochondria as described in (13), or by examining mitochondrial respiration with oxygen consumption measurements. In addition, complex I to V activities are analyzed using spectrophotometry (12), a radiochemical method is used to measure PDHc activity (15). Molecular genetic analysis is performed afterwards, oriented on the biochemical results. Currently, mtDNA can be analyzed quickly and completely with the Affymetrix Human

Mitochondrial Resequencing Array 2.0. Also the analysis of many nuclear encoded genes

(subunits of the respiratory chain complexes, assembly factors, transcription and translation genes) can be performed in the diagnostic laboratory. Despite this, the underlying genetic defect can hitherto be identified in only 50% of patients with a complex deficiency. This will probably be countered using whole genome and whole exome sequencing. Nowadays, these techniques are continuously optimized and become cheaper and faster, therefore it is expected that they will be implemented in a diagnostic setting on short notice. However, a considerable diagnostic challenge concerns the group of patients with a defect in the MEGS capacity outside the respiratory chain, complex V and PDHc. In our experience, this group forms a large subset of mitochondrial patients (1/3 of the patients in Nijmegen, diagnosed in the years 2005-2009 (16)). A mitochondriopathy characterized by isolated MEGS capacity disturbances can be primary or secondary. Therefore, it is first necessary to exclude secondary MEGS dysfunction in muscle tissue (e.g. as a consequence of disuse or poor

Mitochondria and disease

feeding (17)). Subsequently, it must be aimed to confirm the primary nature of the mitochondriopathy. Generally, the approach is two-fold: biochemical and functional studies in a second tissue, and genetic analysis. mtDNA mutations are now often found and it has been shown that the MEGS capacity has a greater sensitivity than respiratory chain enzymatic activities for the detection of rare or new mtDNA mutations, or mtDNA mutations with low mutation loads (18). Also the polymerase gamma (POLG) gene (mitochondrial deletions/depletion) is commonly screened in this setting. Measurements in a second tissue are commonly performed in fibroblasts out of combined skin-muscle biopsies. The respiratory chain, complex V and PDHc enzyme activities can routinely be determined. Measuring ATP synthesis in fibroblasts has been described (19). Different approaches for measuring the oxygen consumption rate are available. Polarographic studies using the Clark-type oxygen electrode are well established and can be performed in different cell types and tissues (20).

The relatively low sample throughput however is a limitation. A more advanced technique which measures the changes in oxygen and proton concentrations in the media surrounding cells is very informative but requires specialized equipment (21).

Despite this substantial diagnostic capacity approximately 50% of the patients with a primary

MEGS dysfunction still remain unsolved. This has been recognized earlier (22). It complicates diagnosis, prognosis, supportive treatment and counseling in the clinical setting.

In PART II of this thesis, we try to address this problem with appropriate additional functional studies in different human cell lines. First, we adapt a protocol using quenched-fluorescence oxygen sensing (23) to measure mitochondrial respiration in digitonin-permeabilized cells.

This high-throughput assay can identify patients with mitochondrial dysfunction in different cell types. Second, we adapt this oxygen consumption assay and other diagnostic biochemical assays to characterize muscle cells. Combined with general (whole genome or whole exome sequencing) and more targeted (e.g. homozygosity mapping) genetic studies we hope to facilitate the search for genetic defects localized on hitherto unknown loci involved in mitochondrial metabolism. Moreover, the application of an extended biochemical and genetic toolbox in human cell lines will help to improve the (patho)physiological

Chapter 1 knowledge of mitochondria and mitochondrial diseases. The study of muscle cells is important in this setting since this is a clinically important cell type. Understanding the nature of the broad and devastating group of mitochondrial diseases is indispensable for the ultimate goal, which is the development of (a) curative therap(y)ies.

AIMS

The first aim in this thesis is to expand current tools for biochemical diagnostics in patients suspected for a mitochondrial disease (PART II).

Unraveling the underlying genetic mutation and studying the functional impact of the defect in patients with a complex V deficiency is the second goal (PART III).

OUTLINE OF THE THESIS

PART I: General introduction

Chapter 1 introduces the OXPHOS system and describes current biochemical and molecular tools in mitochondrial diagnostics, which creates the framework for the study of additional functional assays in human cell lines (PART II of this thesis). Finally, the aims and outline of the thesis are given.

Mitochondria and disease

PART II: Characterization of human cell lines with mitochondrial dysfunction

Muscle biopsy analysis is regarded as the gold standard in diagnostic workups of patients with suspected mitochondrial disorders (12). Cultured fibroblasts are mostly used to confirm the defect in a second tissue. The measurement of individual OXPHOS complexes however does not always provide sufficient information about the functional state of the complete mitochondrial energy-generating system. Therefore, the aim of the study in chapter 2 is to adapt a protocol allowing quenched-fluorescence oxygen sensing to be used in the diagnostic evaluation of suspected mitochondrial disorders in patient-derived cells.

Fibroblasts are vital cell models for additional functional studies. Limitations however are their relatively low mitochondrial activity, the slow growth rate and a poor resemblance with muscle tissue. Muscle-derived cell lines can overcome these limitations. Therefore, the aim of the study in chapter 3 is two-fold: first, to establish OXPHOS-deficient myogenic cell lines and second, to test if several biochemical assays which assess the MEGS as a whole such as pyruvate oxidation, ATP production and oxygen consumption, can be adapted for both myoblasts and myotubes.

PART III: Complex V in health and disease

Mitochondrial ATP production is the main energy source for intracellular metabolic pathways

(1). The human mitochondrial ATP synthase, or complex V is the 5th multi subunit OXPHOS complex. It synthesizes ATP from ADP in the mitochondrial matrix using the energy provided by the proton electrochemical gradient (2, 7-8). Complex V comprises 16 subunits. Five of them (α, β, γ, δ, and ε) form the F1 sector, situated in the mitochondrial matrix, the other subunits (a, c-ring, b, d, F6, OSCP, e, f, g, and A6L) belong to the Fo sector, embedded in the

Chapter 1 inner mitochondrial membrane (24-25). Only two subunits, a and A6L, are encoded by the mtDNA ATP6 and ATP8 genes, respectively (26). The other subunits are nuclear encoded.

In chapter 4, we aim to review complex V with respect to architecture, function, assembly, pathology, and current therapeutic options. The structure and, intriguingly, the oligomerization of complex V determine its function, the former by the so-called “rotary catalysis” (24) and the latter by influencing mitochondrial and cristae morphology (27), as will be discussed.

Chapter 5 describes the first patient with a pathogenic mutation in the mitochondrial ATP8 gene. The clinical phenotype, the pathogenicity of the mutation and the molecular pathophysiology are discussed.

Complex V of mammalian mitochondria is arranged in long rows of dimeric supercomplexes

(27). Ribbons of complex V dimers are responsible for cristae formation, and essential for optimal performance of the mitochondrial enzyme (27). In addition, it has been demonstrated that transmembrane protein 70 or TMEM70 is required to maintain normal expression levels of complex V (28). Although mitochondrial structural abnormalities have been shown to occur in complex V deficiencies before, a direct causal relationship has never been shown.

Therefore, the main goal in chapter 6 is to study mitochondrial morphology in fibroblasts of a patient with a deletion in the TMEM70 gene before and after complementation of the genetic defect. Comparative genomics is used to study the evolution of the protein and to predict the subcellular localization of TMEM70. The latter is checked by immunogold labeling.

In chapter 7, the first two patients with complex V deficiency due to a mutation in the ATP5A1 gene, coding for complex V subunit α, are described. The whole exome sequencing technique is used in the search for the genetic mutation. The powerful combination of biochemical and genetic approaches to unravel a mitochondrial defect is demonstrated.

Mitochondria and disease

PART IV: General discussion

Chapter 8 provides the general discussion, future perspectives and conclusions regarding the different topics in this thesis.

REFERENCES

1. Schapira AH. Mitochondrial disease. Lancet. 2006 Jul 1;368(9529):70-82. 2. Zeviani M, Di Donato S. Mitochondrial disorders. Brain. 2004 Oct;127(Pt 10):2153-72. 3. Frey TG, Renken CW, Perkins GA. Insight into mitochondrial structure and function from electron tomography. Biochim Biophys Acta. 2002 Sep 10;1555(1-3):196-203. 4. Smeitink J, van den Heuvel L, DiMauro S. The genetics and pathology of oxidative phosphorylation. Nat Rev Genet. 2001 May;2(5):342-52. 5. Saraste M. Oxidative phosphorylation at the fin de siecle. Science. 1999 Mar 5;283(5407):1488-93. 6. Di Donato S. Disorders related to mitochondrial membranes: pathology of the respiratory chain and neurodegeneration. J Inherit Metab Dis. 2000 May;23(3):247-63. 7. Nijtmans LG, Klement P, Houstek J, van den Bogert C. Assembly of mitochondrial ATP synthase in cultured human cells: implications for mitochondrial diseases. Biochim Biophys Acta. 1995 Dec 12;1272(3):190-8. 8. Capaldi RA, Aggeler R, Turina P, Wilkens S. Coupling between catalytic sites and the proton channel in F1F0-type ATPases. Trends Biochem Sci. 1994 Jul;19(7):284-9. 9. DiMauro S, Schon EA. Mitochondrial respiratory-chain diseases. N Engl J Med. 2003 Jun 26;348(26):2656-68. 10. Zeviani M, Carelli V. Mitochondrial disorders. Curr Opin Neurol. 2003 Oct;16(5):585- 94. 11. Schaefer AM, Taylor RW, Turnbull DM, Chinnery PF. The epidemiology of mitochondrial disorders--past, present and future. Biochim Biophys Acta. 2004 Dec 6;1659(2- 3):115-20. 12. Janssen AJ, Smeitink JA, van den Heuvel LP. Some practical aspects of providing a diagnostic service for respiratory chain defects. Ann Clin Biochem. 2003 Jan;40(Pt 1):3-8.

Chapter 1

13. Janssen AJ, Trijbels FJ, Sengers RC, Wintjes LT, Ruitenbeek W, Smeitink JA, et al. Measurement of the energy-generating capacity of human muscle mitochondria: diagnostic procedure and application to human pathology. Clin Chem. 2006 May;52(5):860-71. 14. Janssen AJ, Trijbels FJ, Sengers RC, Smeitink JA, van den Heuvel LP, Wintjes LT, et al. Spectrophotometric assay for complex I of the respiratory chain in tissue samples and cultured fibroblasts. Clin Chem. 2007 Apr;53(4):729-34. 15. Sperl W, Trijbels JM, Ruitenbeek W, van Laack HL, Janssen AJ, Kerkhof CM, et al. Measurement of totally activated pyruvate dehydrogenase complex activity in human muscle: evaluation of a useful assay. Enzyme Protein. 1993;47(1):37-46. 16. Rodenburg RJ. Biochemical diagnosis of mitochondrial disorders. J Inherit Metab Dis. 2011 Apr;34(2):283-92. 17. Morava E, Rodenburg R, van Essen HZ, De Vries M, Smeitink J. Dietary intervention and oxidative phosphorylation capacity. J Inherit Metab Dis. 2006 Aug;29(4):589. 18. Janssen AJ, Schuelke M, Smeitink JA, Trijbels FJ, Sengers RC, Lucke B, et al. Muscle 3243A-->G mutation load and capacity of the mitochondrial energy-generating system. Ann Neurol. 2008 Apr;63(4):473-81. 19. Rizza T, Vazquez-Memije ME, Meschini MC, Bianchi M, Tozzi G, Nesti C, et al. Assaying ATP synthesis in cultured cells: a valuable tool for the diagnosis of patients with mitochondrial disorders. Biochem Biophys Res Commun. 2009 May 22;383(1):58-62. 20. Rustin P, Chretien D, Bourgeron T, Gerard B, Rotig A, Saudubray JM, et al. Biochemical and molecular investigations in respiratory chain deficiencies. Clin Chim Acta. 1994 Jul;228(1):35-51. 21. Ferrick DA, Neilson A, Beeson C. Advances in measuring cellular bioenergetics using extracellular flux. Drug Discov Today. 2008 Mar;13(5-6):268-74. 22. Trijbels FJ, Ruitenbeek W, Huizing M, Wendel U, Smeitink JA, Sengers RC. Defects in the mitochondrial energy metabolism outside the respiratory chain and the pyruvate dehydrogenase complex. Mol Cell Biochem. 1997 Sep;174(1-2):243-7. 23. Will Y, Hynes J, Ogurtsov VI, Papkovsky DB. Analysis of mitochondrial function using phosphorescent oxygen-sensitive probes. Nat Protoc. 2006;1(6):2563-72. 24. Devenish RJ, Prescott M, Rodgers AJ. The structure and function of mitochondrial F1F0-ATP synthases. Int Rev Cell Mol Biol. 2008;267:1-58. 25. Walker JE, Dickson VK. The peripheral stalk of the mitochondrial ATP synthase. Biochim Biophys Acta. 2006 May-Jun;1757(5-6):286-96. 26. Anderson S, Bankier AT, Barrell BG, de Bruijn MH, Coulson AR, Drouin J, et al. Sequence and organization of the human mitochondrial genome. Nature. 1981 Apr 9;290(5806):457-65.

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27. Strauss M, Hofhaus G, Schroder RR, Kuhlbrandt W. Dimer ribbons of ATP synthase shape the inner mitochondrial membrane. EMBO J. 2008 Apr 9;27(7):1154-60. 28. Cizkova A, Stranecky V, Mayr JA, Tesarova M, Havlickova V, Paul J, et al. TMEM70 mutations cause isolated ATP synthase deficiency and neonatal mitochondrial encephalocardiomyopathy. Nat Genet. 2008 Nov;40(11):1288-90.

PART II

CHARACTERIZATION OF HUMAN CELL LINES WITH

MITOCHONDRIAL DYSFUNCTION

A high-throughput assay to measure oxygen consumption

in digitonin-permeabilized cells of patients with

mitochondrial disorders

An I. Jonckheere, Merei Huigsloot, Antoon J.M. Janssen, Antonia J.H. Kappen, Jan A.M.

Smeitink, Richard J.T. Rodenburg

Clinical Chemistry 2010; 56(3):424-31

Chapter 2

ABSTRACT

Background: Muscle biopsy analysis is regarded as the gold standard in diagnostic workups of patients with suspected mitochondrial disorders. Analysis of cultured fibroblasts can provide important additional diagnostic information. The measurement of individual OXPHOS complexes does not always provide sufficient information about the functional state of the complete mitochondrial energy generating system. For this purpose, we optimized a high- throughput fluorescence-based methodology for oxygen consumption analysis in patient- derived cells.

Methods: Mitochondrial respiration in digitonin-permeabilized cells was analyzed in the presence of a substrate mix containing pyruvate and malate, using a phosphorescent probe,

96-wells plates and a fluorescence plate reader.

Results: In control fibroblasts, a clear stimulation of the pyruvate + malate driven respiration by ADP was observed. Known inhibitors of the OXPHOS system and the Krebs cycle significantly reduced respiration. In patient fibroblasts with different OXPHOS deficiencies,

ADP stimulated respiratory activity was decreased in comparison to control cells. In several patients with a reduced ATP production rate in muscle tissue but with normal OXPHOS enzyme activities, the fibroblasts displayed a reduced respiratory activity. Finally, a clear difference between control and complex I deficient transmitochondrial cybrid cells was observed.

Conclusion: These results confirm the validity of the assay as a high-throughput screening method for mitochondrial function in digitonin-permeabilized cells. The assay allows primary and secondary mitochondrial abnormalities in muscle to be differentiated, of great importance with respect to counseling, and also will facilitate the search for new genetic defects that lead to mitochondrial disease.

Oxygen consumption assay

INTRODUCTION

Mitochondria play a vital role in cellular energy metabolism. The enzymes of the mitochondrial energy-generating system (MEGS) are localized in the mitochondrial matrix space (pyruvate dehydrogenase complex (PDHc) and tricarboxylic acid (TCA) or Krebs cycle) and in the inner mitochondrial membrane (oxidative phosphorylation (OXPHOS) complex) (1). Enzyme analysis of individual OXPHOS complexes in skeletal muscle biopsy remains the mainstay of the diagnostic process for patients suspected of mitochondrial cytopathy (2). In addition, measurement of the MEGS capacity is a powerful tool for the assessment of mitochondrial function. The MEGS capacity can be examined in detail by

14 14 14 measurement of CO2 production rates from oxidation of [1- C]pyruvate and carboxyl- C- labeled TCA cycle intermediates in combination with measurement of ATP production in intact mitochondria from a muscle biopsy (1), or by examining mitochondrial respiration via oxygen consumption measurements using polarography. These methods have been demonstrated to be very valuable in the examination of fresh muscle tissue. For fibroblasts, several methods have been described to measure ATP production or oxygen consumption

(3, 4). Analysis of fibroblasts is useful for several reasons (5), including confirmation of the defects observed in muscle, to exclude secondary mitochondrial dysfunction e.g. due to poor feeding or disuse (6), and serving as a model system for follow up and functional testing.

One of the most informative ways of assessing mitochondrial function is by analysis of mitochondrial oxygen consumption by polarography using the Clark-type oxygen electrode

(7). With this method, oxygen consumption can be measured in isolated mitochondria, and in intact and detergent-permeabilized cells (8). Although this technique has proven very useful, a major limitation can be the relatively low sample throughput. An assay set-up with a higher throughput has recently become available, but requires specialized equipment (9).

Quenched-fluorescence oxygen sensing is an approach which may overcome these limitations. This measurement methodology allows the assessment of mitochondrial function using conventional instrumentation, thereby combining the high degree of information

Chapter 2 provided by oxygen consumption analysis with the simplicity, throughput and scaling up capabilities of standard microtitre plate assays (10). This approach (10) has previously been used to investigate mitochondrial toxicity of drugs in isolated mitochondria (7) and in cultured adherent cells (11).

The aim of our study was to adapt a protocol allowing quenched-fluorescence oxygen sensing to be used in the diagnostic evaluation of suspected mitochondrial disorders. We demonstrate that this assay provides a fast and non-laborious high-content screening method for mitochondrial function in different digitonin-permeabilized cell types.

Oxygen consumption assay

MATERIALS AND METHODS

This study was carried out in the Netherlands in accordance with the applicable rules concerning research ethics committee review (Commissie Mensgebonden Onderzoek Regio

Arnhem-Nijmegen) and informed consent.

Materials

Pyruvate and oligomycin were purchased from Sigma, malate from Fluka, and ADP from

Roche. All other chemicals were of the highest purity commercially available.

Phosphorescent oxygen-sensitive probe, MitoXpress™ (A65N-1), was from Luxcel

Biosciences. Black body clear bottom 96-well plates (Black Isoplate TC) and black backing tape were obtained from Perkin Elmer.

Cell cultures

After informed consent of the patients and their caretakers, skin biopsy specimens were obtained for diagnostic purposes and fibroblasts were cultured in E199 medium (Gibco,

Invitrogen Corporation). Transmitochondrial cybrids were cultured in high-glucose (4.5 g/L)

Dulbecco's Modified Eagle's Medium (DMEM). Both media were supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin (respectively 100 U/ml and 100

µg/ml). Before harvesting, fibroblasts were incubated for 48 hours in galactose medium, consisting of DMEM without glucose, without pyruvate, with L-glutamine 2 mmol/L (Invitrogen

Corporation) supplemented with 10% dialyzed FBS, 1mmol/L uridine, 1% penicillin/streptomycin and 5.5 mmol/L D-galactose. Cells were harvested and washed twice in ice cold 1% FBS in Phosphate Buffered Saline (PBS), and the cell pellet was subsequently

w 4 resuspended at 30% /v ( = 4-6 x 10 cells/ µl) in ice cold 1% FBS in PBS.

Chapter 2

Isolation of skeletal muscle mitochondria

Human skeletal muscle was obtained from healthy young adults after informed consent.

Muscle mitochondria were isolated as described (12). Briefly, the muscle specimens were placed in ice-cold 0.25 M sucrose, 2 mmol/L EDTA, 10 mmol/L Tris-HCl, 5x104 U/L heparin

(SETH)-medium. After chopping, homogenization was performed by hand with a Teflon-glass

Potter-Elvehjem type homogenizer. The resulting homogenate was centrifuged for 10 minutes at 600 x g. Mitochondria were isolated from the resulting supernatant by centrifugation for 10 minutes at 14000 x g. The mitochondrial pellet was finally taken up in

SETH-medium.

Fluorescence-based assay of mitochondrial respiration

MitoXpress™ (A65N-1) oxygen probe supplied as dry reagent in a vial, was dissolved in PBS to a concentration of 1 µM. Probe fluorescence is quenched by molecular oxygen via a non- chemical (collisional) mechanism, resulting in an increase in probe signal with decreasing concentration of dissolved oxygen due to mitochondrial respiration (10). Measurements were performed immediately after harvesting in digitonin-permeabilized non-fractionated cells with the addition of substrates. Also isolated fresh human skeletal muscle mitochondria were tested. The incubations were pyruvate + malate with or without ADP. The incubation volume in each well was 50 µl, containing 5 µl cell suspension (= 2-3 x 105 cells), 100 nM fluorescent probe, and a substrate mix consisting of 30 mM potassium phosphate , 75 mmol/L potassium

1 5 chloride, 8 mmol/L Tris, 1.6 mmol/L EDTA, 5 mmol/L MgCl2, 0.2 mmol/L p ,p -di(adenosine-

5’) pentaphosphate (myo-adenylate kinase inhibitor), 32.6 µmol/L digitonin for the permeabilization of the cell membranes, 1 mmol/L pyruvate, 1 mmol/L malate, and where indicated 2.0 mmol/L ADP, pH 7.4 (1). To regenerate ADP by creatine kinase in the cell preparations, 20 mmol/L creatine was added to all ADP-containing incubations. Compound solutions were added to the wells to give the indicated final concentrations. Finally, 100 µl of

Oxygen consumption assay mineral oil was added to each well to seal the samples from ambient oxygen which would otherwise destroy oxygen gradients and compromise assay performance (10), and the plate was placed in a time-resolved fluorescence plate reader (Viktor 2, Perkin Elmer) equilibrated to a temperature of 37 °C and monitored every 5 minutes over a period of 55 minutes.

Instrument settings were 340/642 nm excitation/emission filters, a delay time of 30 µs and a measurement window of 100 µs, and active temperature control of the microplate compartment at 37 °C. After completion of fluorescence measurements, time profiles of fluorescence intensity in each well were analyzed using Viktor (Perkin Elmer) and Excel

(Microsoft) software. All measurements were performed in duplicate and the results averaged. The relative oxygen consumption rate (ROCR), reflecting respiratory activity, was calculated from the maximal linearly increasing fluorescence intensity slope (Figure 1) to compare respiration rates of different samples. The results were normalized for citrate synthase (CS) activity (13).

Statistical methods

The different fibroblast groups were tested for the goodness of fit to the normal distribution using the Shapiro-Wilk test. The Kruskal-Wallis test was applied to study differences in respiration rates between the investigated groups. Differences were considered statistically significant if the p-value was lower than 0.05. When overall significance was found, the

Mann-Whitney test with Bonferroni correction (p-value < 0.0125) was used as a post-hoc test to compare each patient group to the control.

Chapter 2

RESULTS

In order to evaluate the maximal mitochondrial oxygen consumption capacity of the samples, measurements were done in non-fractionated digitonin-permeabilized cells under optimal substrate concentrations and in the presence of an excess of ADP. Most experiments were performed in fibroblasts to be able to exclude secondary mitochondriopathies. Therefore, different media were tested (data not shown). Galactose-based medium, which drives the cell towards the OXPHOS system for ATP production, thereby increasing the differences between control and patient fibroblasts (14, 15), was used for subsequent experiments in fibroblasts. Table 1 (see at the end of this article) gives an overview of the cell lines used for oxygen consumption measurements in fibroblasts.

Pyruvate-malate driven oxygen consumption rate

In control fibroblasts, pyruvate-malate driven respiration yielded substantially higher respiratory activity upon ADP stimulation (Figure 1a), indicating proper functional coupling of the mitochondria. This was also clearly seen in isolated mitochondria obtained from control fresh muscle biopsies. Muscle showed both a higher respiratory activity rate and a higher coupling efficiency than fibroblasts (Figure 1b).

Figure 1 ADP stimulation of pyruvate-malate driven respiration in control fibroblasts (a) and in

Oxygen consumption assay isolated mitochondria obtained from a control fresh muscle biopsy (b). The relative oxygen consumption rate (ROCR) of the linearly increasing fluorescence intensity slope, reflecting respiratory activity, is presented by the straight line. (Normalized time profile of fluorescence intensity; RFU = relative fluorescence units; CS = citrate synthase; ● = +ADP; ○ = -ADP)

During the measurement, the initial reduction in raw signal reflects plate temperature equilibration (11). This is followed by a linear phase of oxygen consumption that can be used to analyze mitochondrial respiration. The linear phase of oxygen consumption ends due to oxygen exhaustion (10), which can be concluded from results of pyruvate oxidation rate measurements following the methods outlined in (1). In cultured control fibroblasts, the pyruvate oxidation rate is between 0.1 and 0.15 nmol/min for 5 µl cells. At this rate, 40 minutes of pyruvate oxidation produces 16-24 nmol of NADH and reduces 8-12 nmol O2, whereas the reaction volume of 50 µl at 37 ºC and 1 bar (100 kilopascals) pressure contains only 10.7 nmol O2. In control muscle mitochondria the pyruvate oxidation rate is 0.55-0.65 nmol/min for 5 µl mitochondrial suspension, producing 28-32 nmol NADH and reducing 14-16 nmol O2 in 12 minutes. The addition of a layer of mineral oil to each sample limited back diffusion of oxygen (10). The presence of low rate back diffusion through the mineral oil seal and polystyrene plate (oxygen diffuses through polystyrene at considerable rates) determined the lower limit of the assay sensitivity, i.e. mitochondrial oxygen consumption below this rate cannot be analyzed (10). This is in line with previous observations (7, 11).

The signal stabilizes when equilibrium is reached between the rates of oxygen consumption and back diffusion, so successful analysis in the microplate assay requires an oxygen consumption rate sufficient to produce a measurable signal change (10). Due to the back diffusion the rates of change of dissolved oxygen measured using this assay are lower than the actual mitochondrial oxygen consumption rate. Therefore, they were expressed as relative oxygen consumption rate (ROCR) in relative fluorescence units per unit citrate synthase (RFU/ U CS). The correlation between initial rates of increase of probe signal and the corresponding mitochondrial protein concentration has been demonstrated to be close to linear (7), which made it possible to compare respiration rates of different samples by analyzing the ROCR of the linearly increasing respiratory activity slope (Figure 1). Control

Chapter 2 fibroblasts typically exhibit a ROCR above 3 RFU/ U CS (mean ± standard deviation = 5.0 ±

2.0). It was seen that the oxygen consumption rate can decrease when reaching a higher cell culture passage number (Figure 2 and Table 1).

Figure 2 Normalized respiration rates showing the influence of the passage number on the measurements. The oxygen consumption rate decreases when reaching a higher cell culture passage number. (ROCR: relative oxygen consumption rate; CS = citrate synthase; 8015, 8128 and 5119 = control fibroblasts)

Effect of inhibitors

To study the sensitivity of the assay, several classical inhibitors of the mitochondrial energy generating system were added to control fibroblasts: rotenone (for complex I), antimycin A

(complex III), azide (complex IV), oligomycin (complex V), atractyloside (adenine nucleotide transporter (ANT)), α-hydroxycyanocinnamate (pyruvate carrier), and arsenite (PDHc and α- ketoglutarate dehydrogenase). Figure 3a shows that all inhibitors significantly reduced oxygen consumption. Similar results were obtained with isolated mitochondria from a control fresh muscle biopsy (Figure 3b). This indicates that oxygen consumption measured in this assay is dependent on a fully functional MEGS system. Moreover, it should be feasible to pick up deficiencies in any of its components.

Oxygen consumption assay

Figure 3 Inhibitors of the mitochondrial energy generating system reduce oxygen consumption in control fibroblasts (a) and in isolated mitochondria obtained from a control fresh muscle biopsy (b). (Normalized time profile of fluorescence intensity; RFU = relative fluorescence units; CS = citrate sythase; ● = +ADP + 1 µM rotenone; ○ = +ADP + 1 µM antimycin a; ▼= +ADP + 2 mM azide; ∆ = 0.2 mM oligomycin; ■ = +ADP + 20 µM atractyloside; □ = +ADP + 0.025 mM α- hydroxycyanocinnamate; ♦ = +ADP + 2 mM arsenite; ◊ = + ADP; ▲ = -ADP)

Diagnostics

Analysis of oxygen consumption in fibroblasts from patients with mitochondriopathies

For diagnostic purposes, it is important that deficiencies in the MEGS system can be diagnosed or confirmed in fibroblasts. Therefore, patient fibroblasts with various known

OXPHOS defects were measured in this assay (Figure 4).

Figure 4 Comparison of (mean, where applicable) relative oxygen consumption rates in fibroblasts (see Supplemental Data Table 1) after ADP stimulation using the Mann-Whitney test with Bonferroni correction (p < 0.0125). In the presence of ADP, complex I and IV deficient fibroblasts showed a significantly lower respiratory activity rate compared to control cells. There

Chapter 2 was no significant difference when comparing the controls with the complex V deficient fibroblasts. Fibroblasts of patients with a decreased MEGS capacity in fresh muscle show a range of activities, varying from normal to clearly reduced. (ROCR: relative oxygen consumption rate; CS = citrate synthase; each dot represents one cell line; control = control fibroblasts; C.I = complex I deficient fibroblasts; C.IV = complex IV deficient fibroblasts; C.V = complex V deficient fibroblasts; other = non-OXPHOS deficient fibroblasts, *: p = 0.001; ^: p = 0.008; NS = not significant)

Patient cell lines with a high passage number and a low respiratory activity were not included in this study to prevent possible bias by aging (see Figure 2). There was an overall difference in respiration rates between the groups (Kruskal-Wallis test, p = 0.002). All complex I and complex IV deficient fibroblasts showed an ADP-stimulated respiratory activity rate below the lowest control value. There was a significant difference between the control group and the complex I (p = 0.001) and complex IV (p = 0.008) deficient fibroblasts, and no significant difference when comparing the controls with the complex V deficient fibroblasts (p

= 0.427) (Figure 4). A characteristic feature of the complex V deficient cells was the clear stimulation of oxygen consumption by addition of an uncoupler (2 µM carbonyl cyanide 3- chlorophenyl hydrazone: CCCP) (Figure 5a). This phenomenon was not seen in control cells

(Figure 5b) or in cells with other OXPHOS enzyme deficiencies (data not shown). These data indicate that the fluorescence-based oxygen sensing probe assay is a sensitive tool to measure mitochondrial function in fibroblasts.

Figure 5 Complex V deficient fibroblasts showed clearly higher rates after the addition of an uncoupler (2 µM carbonyl cyanide 3-chlorophenyl hydrazone or CCCP) (a), a phenomenon not seen in control cells (b). (Normalized time profile of fluorescence intensity; RFU = relative fluorescence units; CS = citrate synthase; ● = +ADP ○ = -ADP; ▼= +ADP + CCCP; ∆ = -ADP + CCCP)

Oxygen consumption assay

For patients with non-OXPHOS related decreases in MEGS capacity in fresh muscle biopsies, the oxygen consumption rate would be an important tool to confirm the deficiency since traditional OXPHOS enzyme assays are not discriminative. Several fibroblasts cell lines from this patient category were included. The level of oxygen consumption upon ADP stimulation ranged from control respiratory activity rates to, predominantly, low to very low rates (Figure 4). As a group they were not statistically significant (p = 0.089) but nevertheless several of the non-OXPHOS related MEGS deficiencies could be confirmed in cultured fibroblasts, suggesting that these are most likely caused by (a) primary defect(s).

Cybrid cells

Since transmitochondrial cybrid cells are often used to verify the pathogenicity of mitochondrial DNA (mtDNA) mutations, both control and complex I deficient (ND1 mutation) cybrid cells were included. The clear difference in respiratory activity between control and complex I deficient cybrids (Figure 6; ROCR after ADP stimulation 10.1 and 1.1 RFU/ U CS, respectively) shows that the assay can also be used to rapidly screen cybrid clones for mitochondrial dysfunction due to mtDNA mutations.

Figure 6 Analysis of oxygen consumption in cybrid cells showing a clear difference between control and complex I deficient transmitochondrial cybrid cells. (Normalized time profile of fluorescence intensity; RFU = relative fluorescence units; CS = citrate synthase; cy 73 = control cybrids, cy ND1 = complex I deficient cybrids due to the mutation m.3460G>A in the MT-ND1 gene (known LHON mutation); ● = cy 73 +ADP ○ = cy 73 -ADP; ▼= cy ND1 +ADP; ∆ = cy ND1 -ADP)

Chapter 2

DISCUSSION

Despite the availability of many tools to diagnose mitochondrial disease (16, 17), current methods are still insufficient for the evaluation of a large subgroup of patients suspected to have a mitochondriopathy. Here, we show that quenched-fluorescence oxygen sensing provides a fast and efficient additional diagnostic tool that can be used to examine the mitochondrial energy generating system in cultured patient-derived cells.

The assay showed a clear difference between control and patient cell lines: control cells showed a substantial increase in oxygen consumption rate upon ADP stimulation, in contrast to the patient cell lines. This was illustrated in particular by the cybrid cells. Complex I and IV deficient fibroblasts showed a significantly decreased oxygen consumption rate. Complex V deficient fibroblasts could be recognized because of a substantial increase after uncoupling.

Addition of an uncoupler leads to a decrease in the mitochondrial membrane potential which subsequently gives rise to a maximal stimulation of the respiratory rate (18). Uncoupling makes the oxygen consumption rate independent of complex V and this explains the initial increase in complex V deficient cells. It has also been shown that CCCP stimulates apoptosis via loss in mitochondrial transmembrane potential and reactive oxygen species

(ROS) production (19). Cell death, combined with back diffusion of oxygen, could explain the subsequent decrease of the respiratory activity slope that was sometimes observed.

Inhibitors of the entire pathway from pyruvate import to ATP export, and several steps in between (the MEGS), reduced oxygen consumption in fibroblasts and in isolated muscle mitochondria, confirming that the integrity of the entire OXPHOS machinery can be tested successfully both in permeabilized cells and in isolated mitochondria. A peculiar finding was a slight loss of inhibition of oligomycin in the fibroblasts after 35 minutes. This was not seen in isolated muscle mitochondria. Further research is needed to understand this finding.

Some cautionary notes must be taken into account when measuring the oxygen consumption rate in fibroblasts. It was seen that the rate can decrease when reaching a higher cell culture passage number (Figure 2 and Table 1). To get reliable results it is very important to

Oxygen consumption assay measure cells at an early passage stage and to measure samples in at least two independent experiments. Usually, the passage number increases by 1-2 between measurements. In case of poor reproducibility of multiple measurements with increasing passage number, a possible negative effect of the passage stage should be considered.

Results should then be distrusted and cells from an earlier passage should be tested, if available.

The assay can also be used to support the search for new defects in mitochondrial energy metabolism. In a group of patients with a decreased ATP production in fresh muscle, a subgroup could be identified with a clearly reduced oxygen consumption rate, despite normal

OXPHOS enzymes and PDHc activities. Such data indicate that another still unidentified defect in the mitochondrial energy metabolism may be responsible for the reduced oxygen consumption rates. This will encourage further studies to uncover new genetic defects causing mitochondrial disease. Another subgroup of patients showed normal or near-normal oxygen consumption rates in fibroblasts that may be a reflection of a tissue specific mitochondriopathy, which is not uncommon, e.g. in case of mtDNA mutations with uneven tissue distribution or in case of polymerase gamma (POLG)1 gene mutations. A normal oxygen consumption rate in fibroblasts can also be expected when the MEGS dysfunction in muscle tissue is secondary (poor feeding, disuse) (6). Differentiating between primary and secondary mitochondrial abnormalities is of great importance with respect to counseling.

Thus, this assay provides a valuable tool in the diagnostics of mitochondrial disorders.

Advantages of the assay compared to substrate oxidation measurements include its simplicity, the requirement of less cell material, and the elimination of the need of an isotope laboratory. When comparing oxygen-sensitive fluorescence-based methods to classical polarography it has been shown that, due to the low rate of back diffusion, the rates of change of dissolved oxygen measured using the microplate approach are lower than the consumption rates measured using a sealed polarographic system (7).

In conclusion, quenched-fluorescence oxygen sensing provides a simple, high-throughput, high-content screening method for mitochondrial function in different cell types. It requires a

Chapter 2 relatively low sample volume and allows for simultaneous testing of multiple assay conditions. In addition, this approach is also suitable for the analysis of cell lines with a relatively low mitochondrial activity (e.g. fibroblasts). Patients with a primary mitochondriopathy that does not affect the complexes of the respiratory chain can be identified. In combination with functional studies and other diagnostic assays (for instance homozygosity mapping) the availability of this approach will facilitate the search for hitherto unknown genetic defects leading to mitochondrial disease.

Oxygen consumption assay

Acknowledgments

AJ and JS are supported by a grant from the Prinses Beatrix Fonds and MH by grant

IGE05003 from Senternovem.

REFERENCES

1. Janssen AJ, Trijbels FJ, Sengers RC, Wintjes LT, Ruitenbeek W, Smeitink JA, et al. Measurement of the energy-generating capacity of human muscle mitochondria: diagnostic procedure and application to human pathology. Clin Chem 2006;52:860-71. 2. Janssen AJ, Smeitink JA, van den Heuvel LP. Some practical aspects of providing a diagnostic service for respiratory chain defects. Ann Clin Biochem 2003;40:3-8. 3. Wanders RJ, Ruiter JP, Wijburg FA. Studies on mitochondrial oxidative phosphorylation in permeabilized human skin fibroblasts: application to mitochondrial encephalomyopathies. Biochim Biophys Acta 1993;1181:219-22. 4. Rizza T, Vazquez-Memije ME, Meschini MC, Bianchi M, Tozzi G, Nesti C, et al. Assaying ATP synthesis in cultured cells: a valuable tool for the diagnosis of patients with mitochondrial disorders. Biochem Biophys Res Commun 2009;383:58-62. 5. van den Heuvel LP, Smeitink JA, Rodenburg RJ. Biochemical examination of fibroblasts in the diagnosis and research of oxidative phosphorylation (OXPHOS) defects. Mitochondrion 2004;4:395-401. 6. Morava E, Rodenburg R, van Essen HZ, De Vries M, Smeitink J. Dietary intervention and oxidative phosphorylation capacity. J Inherit Metab Dis 2006;29:589. 7. Hynes J, Marroquin LD, Ogurtsov VI, Christiansen KN, Stevens GJ, Papkovsky DB, Will Y. Investigation of drug-induced mitochondrial toxicity using fluorescence-based oxygen- sensitive probes. Toxicol Sci 2006;92:186-200. 8. Rustin P, Chretien D, Bourgeron T, Gerard B, Rotig A, Saudubray JM, Munnich A. Biochemical and molecular investigations in respiratory chain deficiencies. Clin Chim Acta 1994;228:35-51. 9. Ferrick DA, Neilson A, Beeson C. Advances in measuring cellular bioenergetics using extracellular flux. Drug Discov Today 2008;13:268-74. 10. Will Y, Hynes J, Ogurtsov VI, Papkovsky DB. Analysis of mitochondrial function using phosphorescent oxygen-sensitive probes. Nat Protoc 2006;1:2563-72.

Chapter 2

11. Hynes J, Hill R, Papkovsky DB. The use of a fluorescence-based oxygen uptake assay in the analysis of cytotoxicity. Toxicol In Vitro 2006;20:785-92. 12. Bookelman H, Trijbels JM, Sengers RC, Janssen AJ, Veerkamp JH, Stadhouders AM. Pyruvate oxidation in rat and human skeletal muscle mitochondria. Biochem Med 1978;20:395-403. 13. Srere PA. Citrate synthase. In: Methods Enzymology; 1969. p. 3-11. 14. van der Westhuizen FH, van den Heuvel LP, Smeets R, Veltman JA, Pfundt R, van Kessel AG, et al. Human mitochondrial complex I deficiency: investigating transcriptional responses by microarray. Neuropediatrics 2003;34:14-22. 15. Rossignol R, Gilkerson R, Aggeler R, Yamagata K, Remington SJ, Capaldi RA. Energy substrate modulates mitochondrial structure and oxidative capacity in cancer cells. Cancer Res 2004;64:985-93. 16. Wolf NI, Smeitink JA. Mitochondrial disorders: a proposal for consensus diagnostic criteria in infants and children. Neurology 2002;59:1402-5. 17. Taylor RW, Schaefer AM, Barron MJ, McFarland R, Turnbull DM. The diagnosis of mitochondrial muscle disease. Neuromuscul Disord 2004;14:237-45. 18. Mracek T, Pecina P, Vojtiskova A, Kalous M, Sebesta O, Houstek J. Two components in pathogenic mechanism of mitochondrial ATPase deficiency: energy deprivation and ROS production. Exp Gerontol 2006;41:683-7. 19. Chaudhari AA, Seol JW, Kim SJ, Lee YJ, Kang HS, Kim IS, et al. Reactive oxygen species regulate Bax translocation and mitochondrial transmembrane potential, a possible mechanism for enhanced TRAIL-induced apoptosis by CCCP. Oncol Rep 2007;18:71-6.

Oxygen consumption assay

Table 1: Overview of the cell lines used for oxygen consumption measurements in fibroblasts

Cell line Passage Activity ROCR Mean Gene-Mutation

number (normal 3.0- ROCR

11.0 RFU/

U CS)a

Controls

9022 11 normal 5.64 -

9022 12 normal 3.00 4.09 -

9022 13 normal 3.62 -

8015 12 normal 4.37 -

8015 16 normal 5.82 6.94 -

8015 18 normal 10.63 -

7972 11 normal 7.37 - 6.22 7972 16 normal 5.06 -

8128 10 normal 11.00 - 9.50 8128 20 normal 8.00 -

6869 x+20 normal 3.15 -

8088 15 normal 8.79 -

8088 16 normal 9.29 7.72 -

8088 16 normal 3.99 -

5119 12 normal 6.59 -

9481 20 normal 4.15 -

4752 12 normal 3.87 -

6855 13 normal 3.42 -

9966 4 normal 3.79

9968 7 normal 4.35

Chapter 2

10004 x+4 normal 3.06

10047 6 normal 3.55

Complex I Complex I deficiency (normal

110-260

mU/UCOX)

8328 x+12 26 0.53 NDUFS1-

(del211Q+V228A)

8328 x+13 26 0.43 NDUFS1- 1.04 (del211Q+V228A)

8328 x+14 26 2.17 NDUFS1-

(del211Q+V228A)

5737 16 62 0.00 NDUFS4-

VPEEKH167/

VEKSIstopb

5175 16 65 1.12 NDUFS7-V122Mb

7719 12 59 0.00 ND3-S45P

9270 x+10 56 0.52 NDUFS7-V122M

6603 x+14 16 0.57 NDUFS8-R94Cb

Complex IV Complex IV deficiency (normal

680-1190

mU/UCS)

9353 10 256 0.76

300 13 427 0.42

Oxygen consumption assay

4987 11 155 2.20 SURF1-

(312insATdelTCTGC

CAGCC)c

Complex V Complex V deficiency (normal

209-935

mU/UCOX)

8836 12 14 7.06 TMEM70-317-2A>Gd

8836 14 14 1.41 4.40 TMEM70-317-2A>Gd

8836 16 14 4.72 TMEM70-317-2A>Gd

8835 x+8 113 0.18

Non- ATP+CrPe

OXPHOS (normal defects 42.1-81.2

nmol/h.mU

CS)

7928 14 15 6.86

9322 8 6 12.01

5133 9 8 5.07

8948 11 21 7.89

9354 7 8 2.25

8240 11 12 4.03

5415 x+9 0 3.43

6191 12 30 0.47

9121 x+9 18 1.83

Chapter 2

9492 x+5 n.d.f 0.59

9493 x+5 12 0.95

9158 x+6 10 1.42 1.77 9158 x+8 10 2.12

9164 9 11 1.03

ROCR: relative oxygen consumption rate; RFU: relative fluorescence units; SD: standard deviation; COX: cytochrome oxidase; CS: citrate synthase; CrP: phosphocreatine; x: unknown number of previous passages, typically 2-4 a mean ± SD = 5.0 ± 2.0 (calculated for each cell line, thus taking the mean ROCR where applicable) b Distelmaier F et al. Brain 2009;132(4):833-42 c Coenen MJ et al. J Child Neurol 2006;21(6):508-11 d Cizkova A et al. Nat Genet.2008;40(11):1288-90 e ATP+CrP: production from pyruvate (Janssen AJ et al. Clin Chem 2006;52 (5):860-71) f n.d.: not determined; but severely decreased substrate oxidation rate of [1-14C]pyruvate: 0,61 (normal 3,61 – 7,48 nmol CO2/h.mUCS)

Toolbox for mitochondrial biochemical characterization of

myogenic cell lines from Leigh (-like) patients

Merei Huigsloot1, An I. Jonckheere1, Antoon J.M. Janssen, Antonia J.H. Kappen, Emmy

Zeetsen, Frans C. van den Brandt, Jan A.M. Smeitink, Richard J.T. Rodenburg

1 Joint first authors

In preparation

Chapter 3

ABSTRACT

Two OXPHOS-deficient myogenic cell lines were established, one containing a mutation in the nuclear encoded structural complex I subunit NDUFS7, the other showing a combined complex I and IV deficiency of unknown genetic origin. Biochemical assays assessing the mitochondrial energy generating system (MEGS), such as pyruvate oxidation, ATP production, and oxygen consumption, could be adapted successfully for both myoblasts and myotubes. Combined with spectrophotometric enzyme activity measurements and immunoblotting, a complete biochemical toolbox is provided for the identification and characterization of mitochondrial disorders in muscle cells. This resulted in the first description of an isolated complex I deficiency due to a nuclear genetic mutation in muscle cells and paves the way for multiple applications, e.g. the study of OXPHOS defects that are only expressed in muscle tissue, in-depth research on mitochondrial physiology using biological assays in living cells, and targeted pharmacological intervention studies.

Toolbox for characterization of myogenic cell lines

INTRODUCTION

Most mitochondrial (mt) disorders affect skeletal muscle which has led to the preference of using fresh muscle biopsies for diagnostic purposes. Fibroblasts are mostly used to investigate the defect in a second tissue (1). They are also important cell models for additional functional studies. Limitations of fibroblasts for functional studies are their relatively low mitochondrial activity, the slow growth rate and a poor resemblance with muscle tissue.

For example, a defect in the Polymerase gamma (POLG) gene does not lead to pathologic alterations in fibroblasts (2). Muscle-derived cell lines can overcome some of these limitations. Cultured myoblasts obtained from satellite cells isolated from fresh muscle biopsies grow fast and can be differentiated into myotubes, which display a higher mitochondrial activity than myoblasts (3). Myogenic cell cultures, providing a back-up of muscle tissue, could also preclude a second muscle biopsy should additional testing be required.

In mitochondrial medicine, myogenic cell lines have been used in the study of combined

OXPHOS deficiency due to mtDNA mutations to elucidate the relationship between mutation load and pathology as well as the mechanism of mtDNA segregation in myogenesis (4-8). To study the effect of mtDNA mutations on myogenesis and muscle-specific pathophysiology myogenic cybrids (9) and rhabdomyosarcoma cybrids (10) have been used. However, due to the possibility of genetic drift of heteroplasmic cell lines as well as the switch to cancer- derived nuclear genetic control in cybrids, these are not ideal model cell lines to study the pathophysiology of a genetic defect in a patient. So far only few myogenic cell lines have been established from OXPHOS deficient patients resulting from nuclear DNA mutations.

These include cytochrome c oxidase – deficient SCO2 mutant cells (11) and two cell lines with decreased mitochondrial translation under nuclear genetic control (12). The latter defect was only expressed in myotubes and did not result in decreased complex I and IV enzyme activities measured in vitro in myoblasts and myotubes (12). Since many mitochondrial patients have an isolated complex I deficiency, the establishment of a complex I deficient

Chapter 3 myogenic cell line could provide vital clues to the development of mitochondriopathy in these patients as well as provide a clinically important cell type to test therapeutic agents.

In the present study, we established two OXPHOS deficient myogenic cell lines. One contains a mutation in the nuclear encoded structural complex I subunit NDUFS7. A second myoblast cell line showed combined deficiency of complex I and complex IV, which was not due to mtDNA mutations or several other genes known to affect mtDNA integrity. These cell lines were used to demonstrate a proof-of-principle that several biochemical assays which assess the mitochondrial energy generating system (MEGS), such as pyruvate oxidation,

ATP production, and oxygen consumption, could be adapted for both myoblasts and myotubes.

Toolbox for characterization of myogenic cell lines

PATIENTS AND METHODS

Patients

Patient 1 presented at the age of 14 months with bilateral ptosis. Psychomotor development had been unremarkable. Clinical investigation revealed nystagmus, intention tremor and head titubation. Cerebral MRI showed symmetrical lesions in the medulla oblongata. Lactic acid was elevated in plasma and liquor. A muscle and skin biopsy was performed because of the suspicion of a mitochondrial disorder in this patient with a Leigh-like phenotype.

Myoblasts and myotubes were established out of the muscle biopsy.

The clinical course initially showed a remission marked by an absence of symptoms.

However, an acute disease exacerbation occurred at the age of 2 years and 9 months, and the patient deceased due to hypoventilation and central apneas, leading to respiratory failure.

Patient 2 was born prematurely at 32 weeks and suffered from multiple apnea and bradycardia episodes. He has a developmental retardation and epilepsy. Cerebral MRI performed at the age of 6 years showed white matter lesions in both hemispheres and periaqueductal, suggesting Leigh syndrome. He developed nocturnal apnea and perceptive hearing loss, pointing to central neurological disturbances. A muscle and skin biopsy was performed, and muscle cells were isolated.

At the age of 9 years, the clinical picture is stable. There is a slow developmental progression.

Skeletal muscle cell culture

Primary myoblast cultures were established from quadriceps muscle biopsies of the two patients and seven controls. Satellite cells were released from the sarcolemma by enzymatic digestion as described by Yasin et al. (13) and Shoubridge et al. (14). Cells were separated

Chapter 3 from debris by using a 100 μm nylon mesh cell strainer (Becton Dickinson, Breda, The

Netherlands) and pelleted at 290 g for 5 minutes at 4°C. Myoblasts were cultured in growth medium consisting of Skeletal Muscle Growth Medium (SKGM® BulletKit®, Lonza, Belgium) supplemented with 15% fetal calf serum and penicillin/ streptomycin (respectively 100 U/ml and 100 μg/ml). Fibroblast contamination of bulk cultures was minimized by preplating the cells on collagen (type I from calf skin at 2 μg/cm2 in phosphate-buffered saline (PBS);

Sigma, Zwijndrecht, The Netherlands) for 45 minutes and subsequent transfer of non- adherent cells to a second collagen coated flask (adapted from (15)). Growth medium was changed after 24 hours and every 2-3 days thereafter. Purity of cells was analyzed by immunocytochemical staining with an antibody against desmin (monoclonal mouse anti- human desmin, Dako, Glostrup, Denmark) and cultures routinely contained > 90% desmin positive cells. Myotubes were generated by culturing the myoblasts for 4-6 days in differentiation medium, containing Dulbecco's Modified Eagle's Medium (DMEM) with 1 g/L glucose, 4% Ultroser G® (Pall, France), 4 mM L-glutamine (Invitrogen, Breda, The

Netherlands), and penicillin/streptomycin (respectively 100U/ml and 100 µg/ml). Cells were used at passage 3-7.

In the diagnostic setting in Nijmegen, satellite cells are now isolated from all sufficiently large fresh muscle biopsies. Cells are stored frozen directly following enzymatic digestion.

Myoblast cultures are started when indicated by biochemical and genetic analysis of muscle tissue and/ or fibroblasts. To reduce fibroblast contamination, cells are preplated at the first passage as described above.

Immunofluorescence

Myoblasts or myotubes were fixed in 4% w/v paraformaldehyde in PBS at pH 7.4, permeabilized with 0.1% w/v Triton X100 in PBS and stained with myogenin (F5D, Becton

Dickinson), myosin heavy chain (MHC, clone MF-20 developed by Fishman DA was obtained from the Developmental Studies Hybridoma Bank developed under the auspices of the

Toolbox for characterization of myogenic cell lines

NICHD and maintained by The University of Iowa, Department of Biology, IA (16)) or desmin

(clone D33, Dako) primary antibodies. Secondary antibodies used were FITC-conjugated goat anti-mouse IgG1, Texas Red- conjugated goat anti-mouse IgG2b (both Mitosciences,

Eugene, OR), Alexa488-conjugated goat anti-mouse (Invitrogen, Breda, The Netherlands).

Nuclei were stained with the fluorescent dye Hoechst 33258 (Sigma, Zwijndrecht, The

Netherlands).

Biochemical assays

Measurement of the MEGS capacity was performed on myoblasts or myotubes collected as described for fibroblasts (17). Substrate oxidation and ATP production were subsequently determined using the same buffers as in (18) except that 5 μl of cell suspension was used in a total reaction volume of 50 μl, which contained 0.04% w/v digitonin (Sigma, Zwijndrecht,

The Netherlands) to permeabilize the cells, similar to (17). Mitochondrial respiration in digitonin-permeabilized cells was analyzed in the presence of a substrate mix containing pyruvate and malate, using the phosphorescent probe MitoXpress (Luxcel, Cork, Ireland),

96-wells plates and a fluorescence plate reader as described (17). The activities of the mitochondrial respiratory chain enzymes and complex V were measured spectrophotometrically in mitochondria isolated from frozen pellets of myoblasts, myotubes, muscle tissue and fibroblasts of the patients according to described protocols (19-21) and were expressed in milliunits per unit cytochrome c oxidase (COX) (22) or the mitochondrial matrix enzyme citrate synthase (CS) (23).

Blue Native and SDS-PAGE analysis

One-dimensional 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-

PAGE) and 5%-15% blue native (BN)-PAGE was performed as described previously (24-25).

Lanes were loaded with 30 (SDS analysis) or 40 (BN analysis) µg of solubilized

Chapter 3 mitochondrial protein (26). After electrophoresis, blue native gels were processed further for immunoblotting as described by (25). For Western blotting, antibodies used were raised against NDUFA9 (complex I; Invitrogen, Breda, The Netherlands), Core 2 (complex III;

Invitrogen), COX 1 (complex IV; Mitosciences, Eugene, OR, USA), subunit alpha (complex

V; Mitosciences), CoII-70kDa (complex II, loading control, Mitosciences), myogenin and

MHC (see 2.3). Secondary antibodies used were peroxidase-conjugated anti-mouse or anti- rabbit IgGs (Invitrogen). Creatine kinase (CK) isoenzymes were analyzed in 2 μg cell lysate using the CK isoenzyme Titan gel-kit (Helena Biosciences, Tyne and Wear, UK).

Toolbox for characterization of myogenic cell lines

RESULTS

High quality myoblast and myotube cultures established from patient muscle biopsies

For correct interpretation of biochemical measurements in the muscle cell cultures, it is important that little to no contaminating fibroblasts are present. The purity of the myoblast cultures was determined by staining the cells for the muscle-specific protein desmin. Very high purity was seen in all cultures established from fresh muscle biopsies (> 90% desmin positive cells) (Figure 1, left). Myoblasts could be differentiated into myotubes as shown by fusion of the myoblasts into multinucleated tubes and by the induction of myogenin (green) and MHC (red) (Figure 1).

Figure 1 High quality myoblast and myotube cultures were established from patient muscle biopsies. Myoblast cultures were stained for desmin (left) and all cultures were > 90% desmin positive. Both patient and control myoblasts could be differentiated into myotubes as shown by the induction of myogenin (green) and myosin heavy chain (MHC, red). Arrowheads indicate striation of MHC, nuclei were stained blue with Hoechst.

Biochemical assays and molecular genetic analysis in muscle tissue and fibroblasts

Analysis of the MEGS capacity revealed that the pyruvate oxidation rate in muscle tissue

14 from both patients was severely decreased (1.60 and 1.63 nmol CO2/h.mU citrate synthase

(CS), respectively, normal 3.45-7.99). The ATP production rate from oxidation of pyruvate

Chapter 3 was also severely decreased (19.8 and 18.4 nmol (ATP+CrP)/ h.mU CS, respectively, normal 36-81.7). For patient 1, spectrophotometric analysis showed an isolated complex I deficiency in muscle tissue (53 mU/ U CS, normal 100-401) and in fibroblasts (118 mU/ U

CS, normal 163-599). Genetic analysis revealed compound heterozygous mutations in the

NDUFS7 gene (c.364G>A and c.449T>C), a structural subunit of complex I. Patient 2 showed a complex I deficiency in muscle tissue (36 mU/ U CS, normal 100-401) and a combined complex I (70 mU/ U CS, normal 163-599) and IV (178 mU/U CS, normal 288-954) deficiency in fibroblasts. Complex IV activity in muscle tissue was in the lower normal range

(886 mU/ U CS, normal 810-3120). Hitherto, genetic analysis (mtDNA, POLG, PUS1,

MPRS16, GFM1, TSFM) showed no mutations. The activities of the remaining OXPHOS enzymes and citrate synthase were normal in both patients.

Biochemical assays in myoblasts and myotubes

To ensure that cultured myoblasts and myotubes can be used as a valid model system for complex I (patient 1) or complex I and IV (patient 2) deficiency, they were fully characterized.

To mimic the biochemical screening used for fresh muscle tissue, we adapted the existing

MEGS capacity assays (18). Digitonin was added to permeabilize the cells and the total reaction volume was reduced. With these modifications functional measurements could be performed successfully in muscle cells. Results showed that the pyruvate oxidation rate and

ATP production rate from oxidation of pyruvate were decreased in the myoblasts of patient 2

(Figure 2a+2b). The relative oxygen consumption rate was decreased in myoblasts as well as myotubes of both patients (Figure 2c).

Toolbox for characterization of myogenic cell lines

Figure 2 MEGS capacity of cultured muscle cells is affected in both patients. Pyruvate oxidation rate (a), ATP production rate (b) and relative oxygen consumption rate (ROCR) (c) were determined in myoblasts and myotubes of both patients. The ROCR of patient 2 myoblasts was below the detection limit (indicated as ‘n.d.’, not detectable) (Controls: n=14 for (a), n=15 for (b), n=15 for (c) blasts, n=13 for (c) tubes, bars represent the 25th and 75th percentile of control values; Pyruvate oxidation: pyruvate oxidation rate (expressed as nmol 14CO2/h.mU CS); ATP production: ATP production rate from oxidation of pyruvate (expressed as nmol (ATP+CrP)/h.mU CS); ROCR: relative oxygen consumption rate (expressed as relative fluorescence units (RFU)/ U CS))

Respiratory chain enzyme activity analysis in myoblasts and myotubes of patient 1 revealed an isolated complex I deficiency (Figure 3a). A combined complex I and IV deficiency was present in the myoblasts and myotubes of patient 2 (Figure 3a+3b).

The activities of the remaining OXPHOS enzymes and citrate synthase were normal in both patients (data not shown).

Chapter 3

Figure 3 Cultured muscle cells show isolated complex I deficiency for patient 1 and combined complex I and IV deficiency for patient 2. The activity of (a) complex I and (b) complex IV was analyzed spectrophotometrically in myoblasts and myotubes and expressed relative to citrate synthase activity. (Controls: n=7, bars represent the 25th and 75th percentile of control values; CS: citrate synthase; COX: cytochrome c oxidase = complex IV)

Together, the enzymatic activity measurements confirm the isolated complex I deficiency found in muscle tissue and fibroblasts of patient 1. For patient 2, the enzymatic activity measurements in myoblasts and myotubes match the results found in fibroblasts. The complex IV activity in muscle tissue of patient 2 was situated in the lower normal range. In conclusion, these data describe the first isolated complex I deficiency due to a nuclear DNA mutation demonstrated in cultured muscle cells. They also show that different biochemical assays can be applied successfully to fully characterize patient muscle cell lines.

Immunoblotting in myoblasts and myotubes (Figure 4)

In order to obtain more information about the structure and amount of the different complexes in relation to their activity, BN-PAGE was performed in myoblasts and myotubes of the two patients.

Toolbox for characterization of myogenic cell lines

Figure 4 Holocomplex I (and IV for patient 2) levels are reduced in patient myoblasts and myotubes. BN-PAGE analysis (rows 1-5) of complex I to V of the OXPHOS system in patient and control myoblasts and myotubes. Expression of the differentiation markers myogenin, MHC and CK-MM was analyzed by SDS-PAGE (rows 6- 7) or in-gel activity (row 8), respectively. (Con: control; P1: patient 1; P2: patient 2; CI: complex I; CII: complex II; CIII: complex III; CIV: complex IV; CV: complex V; MHC: Myosin Heavy Chain; CK-MM: muscle fraction of creatine kinase).

Immunoblotting after BN-PAGE showed a decreased amount of complex I in myoblasts and myotubes of patient 1. Holocomplex I and IV were nearly absent in the myoblasts of patient

2, while the amount of these complexes was strongly decreased in the myotubes. The complex III panel showed three bands, the lowest represents complex III, the two slower migrating bands represent supercomplexes containing complex III. The middle band is absent in myoblasts of patient 2. Most likely, this band represents a supercomplex containing complex III and complex IV (27). These results demonstrate that both patients have a decreased amount of complex I and that patient 2 also has a decreased amount of complex

IV in muscle cells, which is in line with the enzymatic activity data measured spectrophotometrically. For patient 1, the complex IV band is slightly decreased compared to the control. However, the middle band in the complex III panel, representing a supercomplex of complex III and complex IV, is normal, which is compatible with the spectrophotometric enzyme activity data, showing an isolated complex I deficiency. The differentiation markers

Chapter 3 myogenin, MHC and the CK muscle fraction (CK-MM) were more pronounced in myotubes than in myoblasts. The control cell line showed some differentiation in myoblasts, which was also seen during culture (not shown).

Toolbox for characterization of myogenic cell lines

DISCUSSION

A fresh muscle biopsy is the gold standard for detailed biochemical analysis of the MEGS capacity and enzymatic activity measurements in patients suspected for a mitochondrial disorder (19). Additional analysis of fibroblasts is useful to confirm the defect observed in muscle tissue, to exclude a possible secondary nature of the alterations found in muscle e.g. due to poor feeding or disuse (28), and to perform functional assays in a diagnostic follow-up or research setting (1, 17). Cultured myoblasts and myotubes, however, could be a advantageous alternative since they are directly derived from the gold standard and could thus be a source for in-depth research in muscle-derived cells. For this aim, we optimized a myoblast culture protocol for diagnostic purposes. The preplating technique (15) was adapted to improve the purity of the culture. Different media for the culture of myoblasts and myotubes were compared and the ones generating the best cultures were selected. These protocol modifications resulted in mostly round myoblasts and fairly differentiated myotubes with, overall, a high degree of purity. Limitations of these cultures are the degree of differentiation which in vitro cannot reach to the stage of a mature muscle, a variable growth- and differentiation rate among different cell lines (data not shown) and the viability of the cultures, which is restricted to 8-9 passages. Differences in the degree of fusion and a restricted proliferative potential of myoblasts have been observed previously (14). It was also stated that cells have to be used at an early passage if fusion to myotubes is wanted (14).

Immortalization of the muscle cells, as performed in (29) can solve the latter issue. Since the biochemical and genetic diagnosis is not yet known when isolating satellite cells from the fresh muscle biopsy, it was decided in Nijmegen to further adapt the protocol and to freeze satellite cells from fresh muscle biopsies directly following their enzymatic digestion. Starting the muscle cell culture after thawing of these frozen fractions did not lead to a loss of quality of the cultured cells (data not shown).

We selected 2 patients with OXPHOS deficiencies demonstrated in muscle tissue and fibroblasts to study the muscle-derived cells. For the first time, the entire biochemical toolbox,

Chapter 3 including pyruvate oxidation and ATP production rates, oxygen consumption, spectrophotometric analysis of single enzymes and BN-PAGE, was applied to myogenic cells. This resulted, to our knowledge, for the first time in the confirmation of an isolated complex I deficiency due to a nuclear genetic mutation in human myoblasts and myotubes

(patient 1). Patient 2 presented with a combined complex I and IV deficiency in fibroblasts. In muscle, complex I was also decreased but the complex IV activity was situated in the lower normal range. The complex I and IV deficiencies were also observed in myoblasts and myotubes thus confirming the presence of a combined OXPHOS deficiency. Since the genetic defect is not known, one can only speculate why the complex IV activity in muscle tissue differs to a certain extent from that in fibroblasts and cultured muscle cells. Possibly, a developmentally regulated factor important for mitochondrial translation in a late stage of muscle cell differentiation could play a role (12).

The reduction in MEGS function, as demonstrated by several functional assays (pyruvate oxidation and ATP production rates, oxygen consumption), correlates with the severity of the deficiency found by single enzyme measurements and BN-PAGE. This is illustrated by the two patients, patient 1 having a milder defect than patient 2. Furthermore, it was observed that the relative oxygen consumption rate is the most sensitive assay, which picked up both deficiencies in cultured myoblasts as well as in differentiated myotubes. Decreased pyruvate oxidation and ATP production rates were only detected in myoblasts of patient 2. This suggests that these two assays are less sensitive in myogenic cells, which may be caused by their stem cell properties or by alterations induced by cell culture conditions.

Another possible application of the functional assays lies in the characterization of the large subset of patients with a decreased ATP production without a defect in the OXPHOS enzymes in muscle tissue. When these assays show a decrease in energy production in cultured muscle cells or fibroblasts of this patient group, it most probably reflects a primary mitochondrial disease and not secondary mitochondrial dysfunction. The latter can be caused by e.g. poor feeding or inactivity (17, 28, 30) and is characterized by a decreased

Toolbox for characterization of myogenic cell lines

MEGS capacity solely in fresh muscle tissue which can normalize in muscle tissue taken after clinical improvement.

Altogether, this study indicates that the myoblast and myotube cell model together with the biochemical assays provide a complete toolbox for the identification and characterization of mitochondrial disorders in a clinically important cell type. The fact that the muscle cells are primary, non-immortalized or otherwise genetically modified makes them more genuine than, for example, myoD transformed fibroblasts (31). This paves the way for multiple applications.

These cells could be suited to study tissue specific OXPHOS defects that are only expressed in muscle tissue and not in fibroblasts. The physiology of mitochondrial disease can be studied more thoroughly by applying known biological assays like Reactive Oxygen Species

(ROS) measurements, membrane potential and calcium uptake measurements in living muscle cells (32-36). A better knowledge of the pathophysiology of mitochondrial disease can help to develop targeted pharmacological intervention studies, which could be tested in vitro using the cultured muscle cells. Moreover, the myoblasts containing a mutation in

NDUFS7 could serve as a model to test whether cells with mutated structural subunits are susceptible for therapy.

Chapter 3

Funding

This work was supported by a grant from the Prinses Beatrix Fonds [grant number OP-05-04] to AIJ and JS and grant IGE05003 from Senternovem to MH. The authors confirm independence from the sponsors; the content of the article has not been influenced by the sponsors.

Conflicts of interest

None.

The study has been carried out in the Netherlands in accordance with the applicable rules concerning the review of research ethics committees (Commissie Mensgebonden Onderzoek

Regio Arnhem-Nijmegen) and informed consent.

REFERENCES

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Toolbox for characterization of myogenic cell lines

5. Pye D, Watt DJ, Walker C, Lightowlers RN, Turnbull DM. Identification of the RAG-1 as a suitable mouse model for mitochondrial DNA disease. Neuromuscul Disord. 2004 May;14(5):329-36. 6. Boulet L, Karpati G, Shoubridge EA. Distribution and threshold expression of the tRNA(Lys) mutation in skeletal muscle of patients with myoclonic epilepsy and ragged-red fibers (MERRF). Am J Hum Genet. 1992 Dec;51(6):1187-200. 7. Suomalainen A, Majander A, Pihko H, Peltonen L, Syvanen AC. Quantification of tRNA3243(Leu) point mutation of mitochondrial DNA in MELAS patients and its effects on mitochondrial transcription. Hum Mol Genet. 1993 May;2(5):525-34. 8. Sasarman F, Antonicka H, Shoubridge EA. The A3243G tRNALeu(UUR) MELAS mutation causes amino acid misincorporation and a combined respiratory chain assembly defect partially suppressed by overexpression of EFTu and EFG2. Hum Mol Genet. 2008 Dec 1;17(23):3697-707. 9. Sobreira C, King MP, Davidson MM, Park H, Koga Y, Miranda AF. Long-term analysis of differentiation in human myoblasts repopulated with mitochondria harboring mtDNA mutations. Biochem Biophys Res Commun. 1999 Dec 9;266(1):179-86. 10. Vergani L, Malena A, Sabatelli P, Loro E, Cavallini L, Magalhaes P, et al. Cultured muscle cells display defects of mitochondrial myopathy ameliorated by anti-oxidants. Brain. 2007 Oct;130(Pt 10):2715-24. 11. Jaksch M, Paret C, Stucka R, Horn N, Muller-Hocker J, Horvath R, et al. Cytochrome c oxidase deficiency due to mutations in SCO2, encoding a mitochondrial copper-binding protein, is rescued by copper in human myoblasts. Hum Mol Genet. 2001 Dec 15;10(26):3025-35. 12. Sasarman F, Karpati G, Shoubridge EA. Nuclear genetic control of mitochondrial translation in skeletal muscle revealed in patients with mitochondrial myopathy. Hum Mol Genet. 2002 Jul 1;11(14):1669-81. 13. Yasin R, Van Beers G, Nurse KC, Al-Ani S, Landon DN, Thompson EJ. A quantitative technique for growing human adult skeletal muscle in culture starting from mononucleated cells. J Neurol Sci. 1977 Jul;32(3):347-60. 14. Shoubridge EA, Johns T, Boulet L. Use of myoblast cultures to study mitochondrial myopathies. Methods Enzymol. 1996;264:465-75. 15. Yaffe D. Retention of differentiation potentialities during prolonged cultivation of myogenic cells. Proc Natl Acad Sci U S A. 1968 Oct;61(2):477-83. 16. Bader D, Masaki T, Fischman DA. Immunochemical analysis of myosin heavy chain during avian myogenesis in vivo and in vitro. J Cell Biol. 1982 Dec;95(3):763-70.

Chapter 3

17. Jonckheere AI, Huigsloot M, Janssen AJ, Kappen AJ, Smeitink JA, Rodenburg RJ. High-throughput assay to measure oxygen consumption in digitonin-permeabilized cells of patients with mitochondrial disorders. Clin Chem. 2010 Mar;56(3):424-31. 18. Janssen AJ, Trijbels FJ, Sengers RC, Wintjes LT, Ruitenbeek W, Smeitink JA, et al. Measurement of the energy-generating capacity of human muscle mitochondria: diagnostic procedure and application to human pathology. Clin Chem. 2006 May;52(5):860-71. 19. Janssen AJ, Smeitink JA, van den Heuvel LP. Some practical aspects of providing a diagnostic service for respiratory chain defects. Ann Clin Biochem. 2003 Jan;40(Pt 1):3-8. 20. Janssen AJ, Trijbels FJ, Sengers RC, Smeitink JA, van den Heuvel LP, Wintjes LT, et al. Spectrophotometric assay for complex I of the respiratory chain in tissue samples and cultured fibroblasts. Clin Chem. 2007 Apr;53(4):729-34. 21. Jonckheere AI, Hogeveen M, Nijtmans LG, van den Brand MA, Janssen AJ, Diepstra JH, et al. A novel mitochondrial ATP8 gene mutation in a patient with apical hypertrophic cardiomyopathy and neuropathy. J Med Genet. 2008 Mar;45(3):129-33. 22. Cooperstein SJ, Lazarow A. A microspectrophotometric method for the determination of cytochrome oxidase. J Biol Chem. 1951 Apr;189(2):665-70. 23. Srere PA. Citrate synthase 1969. 24. Schagger H, von Jagow G. Blue native electrophoresis for isolation of membrane protein complexes in enzymatically active form. Anal Biochem. 1991 Dec;199(2):223-31. 25. Nijtmans LG, Henderson NS, Holt IJ. Blue Native electrophoresis to study mitochondrial and other protein complexes. Methods. 2002 Apr;26(4):327-34. 26. Smith PK, Krohn RI, Hermanson GT, Mallia AK, Gartner FH, Provenzano MD, et al. Measurement of protein using bicinchoninic acid. Anal Biochem. 1985 Oct;150(1):76-85. 27. Huigsloot M, Nijtmans LG, Szklarczyk R, Baars MJ, van den Brand MA, Hendriksfranssen MG, et al. A mutation in C2orf64 causes impaired cytochrome c oxidase assembly and mitochondrial cardiomyopathy. Am J Hum Genet. 2011 Apr 8;88(4):488-93. 28. Morava E, Rodenburg R, van Essen HZ, De Vries M, Smeitink J. Dietary intervention and oxidative phosphorylation capacity. J Inherit Metab Dis. 2006 Aug;29(4):589. 29. Lochmuller H, Johns T, Shoubridge EA. Expression of the E6 and E7 genes of human papillomavirus (HPV16) extends the life span of human myoblasts. Exp Cell Res. 1999 Apr 10;248(1):186-93. 30. Jonckheere AI, Huigsloot M, Janssen AJ, Kappen AJ, Smeitink JA, Rodenburg RJ. High-throughput assay to measure oxygen consumption in digitonin-permeabilized cells of patients with mitochondrial disorders. Clinical chemistry. Mar;56(3):424-31. 31. Choi J, Costa ML, Mermelstein CS, Chagas C, Holtzer S, Holtzer H. MyoD converts primary dermal fibroblasts, chondroblasts, smooth muscle, and retinal pigmented epithelial

Toolbox for characterization of myogenic cell lines cells into striated mononucleated myoblasts and multinucleated myotubes. Proc Natl Acad Sci U S A. 1990 Oct;87(20):7988-92. 32. Distelmaier F, Koopman WJ, Testa ER, de Jong AS, Swarts HG, Mayatepek E, et al. Life cell quantification of mitochondrial membrane potential at the single organelle level. Cytometry A. 2008 Feb;73(2):129-38. 33. Koopman WJ, Distelmaier F, Esseling JJ, Smeitink JA, Willems PH. Computer- assisted live cell analysis of mitochondrial membrane potential, morphology and calcium handling. Methods. 2008 Dec;46(4):304-11. 34. Distelmaier F, Visch HJ, Smeitink JA, Mayatepek E, Koopman WJ, Willems PH. The antioxidant Trolox restores mitochondrial membrane potential and Ca2+ -stimulated ATP production in human complex I deficiency. J Mol Med. 2009 May;87(5):515-22. 35. Koopman WJ, Verkaart S, van Emst-de Vries SE, Grefte S, Smeitink JA, Willems PH. Simultaneous quantification of oxidative stress and cell spreading using 5-(and-6)- chloromethyl-2',7'-dichlorofluorescein. Cytometry A. 2006 Dec 1;69(12):1184-92. 36. Verkaart S, Koopman WJ, van Emst-de Vries SE, Nijtmans LG, van den Heuvel LW, Smeitink JA, et al. Superoxide production is inversely related to complex I activity in inherited complex I deficiency. Biochim Biophys Acta. 2007 Mar;1772(3):373-81.

PART III

COMPLEX V IN HEALTH AND DISEASE

Mitochondrial ATP synthase: architecture,

function and pathology

An I. Jonckheere, Jan A.M. Smeitink, Richard J.T. Rodenburg

Adapted from: J Inherit Metab Dis 2012; 35(2):211-25

Chapter 4

ABSTRACT

Human mitochondrial (mt) ATP synthase, or complex V consists of two functional domains:

F1, situated in the mitochondrial matrix, and Fo, located in the inner mitochondrial membrane.

Complex V uses the energy created by the proton electrochemical gradient to phosphorylate

ADP to ATP. This review covers the architecture, function and assembly of complex V. The role of complex V di-and oligomerization and its relation with mitochondrial morphology is discussed. Finally, pathology related to complex V deficiency and current therapeutic strategies are highlighted.

Mitochondrial ATP synthase

Mitochondrial ATP production is the main energy source for intracellular metabolic pathways

(1). The human mitochondrial (mt) ATP synthase, or complex V (EC 3.6.3.14) is the 5th multi subunit oxidative phosphorylation (OXPHOS) complex. It synthesizes ATP from ADP in the mitochondrial matrix using the energy provided by the proton electrochemical gradient (2-4).

To date, numerous mutations in the mtDNA encoded subunits a and A6L have been found, next to mutations in a structural subunit (subunit epsilon, (5)), an assembly factor (ATP12,

(6)) and an ancillary factor (TMEM70, (7)) which are nuclear-encoded. Most of these mutations give rise to severe mitochondrial disease phenotypes, ranging from NARP

(Neuropathy, Ataxia, and Retinitis Pigmentosa) or MILS (Maternally Inherited Leigh

Syndrome) for subunit a mutations to neonatal mitochondrial encephalo(cardio)myopathy and dysmorphic features in patients with an ATP12 or TMEM70 gene mutation (6-7). In this article, we have aimed to review the 5th OXPHOS complex with respect to architecture, function, assembly, pathology, current therapeutic options and future perspectives. The structure and, intriguingly, the oligomerization of complex V determine its function, the former by the so-called “rotary catalysis” (8) and the latter by influencing mitochondrial and cristae morphology (9), as will be discussed below.

ATP synthase: architecture (Figure 1)

ATP synthase consists of two well defined protein entities: the F1 sector, a soluble portion situated in the mitochondrial matrix, and the Fo sector, bound to the inner mitochondrial membrane. F1 is composed of three copies of each of subunits α and β, and one each of subunits γ, δ and ε. F1 subunits γ, δ and ε constitute the central stalk of complex V. Fo consists of a subunit c-ring (probably comprising eight copies, as shown in bovine mitochondria (10)) and one copy each of subunits a, b, d, F6 and the oligomycin sensitivity- conferring protein (OSCP). Subunits b, d, F6 and OSCP form the peripheral stalk which lies to one side of the complex. A number of additional subunits (e, f, g, and A6L), all spanning

Chapter 4

the membrane, are associated with Fo (8, 11). Two of the Fo subunits, subunit a and subunit

A6L are encoded by the mtDNA ATP6 and ATP8 genes, respectively (12). The detailed structure of most of the bovine mitochondrial complex V subunits, which will not be covered in this review, has been resolved by X-ray crystallography by John Walker and his group

(see, among others, (11, 13-19)).

Figure 1 Human mitochondrial ATP synthase, or complex V, consists of two functional domains, F1 and Fo. F1 comprises 5 different subunits (three α, three β, and one γ, δ and ε) and is situated in the mitochondrial matrix. Fo contains subunits c, a, b, d, F6, OSCP and the accessory subunits e, f, g and A6L. F1 subunits γ, δ and ε constitute the central stalk of complex V. Subunits b, d, F6 and OSCP form the peripheral stalk. Protons pass from the intermembrane space to the matrix through Fo, which transfers the energy created by the proton electrochemical gradient to F1, where ADP is phosphorylated to ATP. One β subunit is taken out to visualize the central stalk.

For comparison, the subunit composition of human, yeast and E. coli ATP synthase is presented in Table 1 (10, 20).

Table 1: Subunit composition of human, yeast and E. coli ATP synthase (10, 20).

Bacteria Mitochondria

Stoichiometry E. coli S. cerevisiae H. sapiens

F1 3 α α α

3 β β β

1 γ γ γ

1 ε δ δ

1 - ε ε

Mitochondrial ATP synthase

1 δ OSCP OSCP

Fo 1 a 6 a

1 - 8 A6L

10-15 c10-12 910 c8

1-2 b2 4 b

1 - d d

1 - h F6

1 - f f

t.b.d. - e e

t.b.d. - g g

1 - i -

1 - k -

Regulators 1 - Inh1p IF1

t.b.d. - Stf1p -

t.b.d. - Stf2p - t.b.d.: to be determined

How structure relates to function: the rotary F1Fo ATP synthase

The function of ATP synthase is to synthesize ATP from ADP and inorganic phosphate (Pi) in the F1 sector. This is possible due to energy derived from a gradient of protons which cross the inner mitochondrial membrane from the intermembrane space into the matrix through the

Fo portion of the enzyme. The proton gradient establishes a proton-motive force, which has two components: a pH differential and an electrical membrane potential (∆ψm) (21). The released energy causes rotation of two rotary motors: the ring of c subunits in Fo (22)

(relative to subunit a), along with subunits γ, δ and ε in F1 (23), to which it is attached.

Protons pass Fo via subunit a to the c-ring (24). Rotation of subunit γ within the F1 α3β3

Chapter 4 hexamer provides energy for ATP synthesis. This is called “rotary catalysis” (8) and can be explained by the “binding-change” mechanism, first proposed by Boyer (25). This mechanism describes ATP synthesis and ATP hydrolysis at the catalytic sites, located in each of the three β subunits, at the interface with an adjacent α subunit. In the case of ATP synthesis, each site switches cooperatively through conformations in which ADP and Pi bind, ATP is formed, and then released. ATP hydrolysis uses the same pathway, but in reverse (26).

These transitions are caused by rotation of the γ subunit. The α3β3 hexamer must remain fixed relative to subunit a during catalysis, this occurs through the peripheral stalk. Complex

V can therefore mechanically be divided into “rotor” (c-ring, γ, δ, ε) and “stator” (α3β3, a, b, d,

F6, OSCP) components (8).

Oligomycin is an inhibitor of proton translocation in ATP synthase, putatively binding the Fo subunits a and c (27).

F1Fo ATP synthases, or F-type ATPases, belong to a bigger rotor protein family, consisting of

F-, V-, and A-type ATPases (28-29). The different ATPases are categorized on the basis of their function and taxonomic origin. They are related in both structure and mechanism (30).

F-type ATPases synthesize ATP, but are also capable of the reversed reaction and can hydrolyze ATP. Vacuolar or V-type ATPases use the energy derived from ATP hydrolysis to pump protons through membranes (28). V-ATPases are localized to intracellular membranes where they acidify intracellular compartments (e.g. lysosomes). They also lie in the plasma membrane, to transport protons out of highly specialized cells, like kidney cells or osteoclasts

(30). Eukaryotes contain both F- and V-ATPases. In contrast, archaea typically contain only one complex which is evolutionarily closer to V-ATPases, called A-ATPase/synthase (28). A-

ATPases are structurally simpler than F- or V-ATPases, but functionally more versatile, performing both ATP synthesis and ATP hydrolysis (28). F-type ATP synthases contain only one peripheral stalk, while A- and V-ATPases have two and three peripheral stalks, respectively, each made of an E-G heterodimer (28).

Mitochondrial ATP synthase

Proteins related to mammalian ATP synthase

Coupling factors: the Inhibitor Protein IF1 and Factor B

In normally respiring mitochondria, ∆ψm is high (estimated at between 150 and 180 mV negative to the cytosol), favoring ATP synthesis by F1Fo ATP synthase (21). However, when mitochondrial respiration is compromised and ∆ψm falls below a threshold, F1Fo ATP synthase can reverse, hydrolyzing ATP to pump protons through the membrane. Since depletion of ATP precipitates cell death, wasteful hydrolysis of ATP is not desirable, and must be prevented during inhibition of respiration (8). This regulation is carried out by the inhibitor protein IF1, which inhibits mitochondrial F1-ATPase activity (31) in a pH-dependent manner (32). The active form predominates at pH values < 6.5 (33). IF1 conserves ATP at the expense of ∆ψm, which has been shown to be protective to cells during ischemia (21, 34). IF1 acts as a homodimer, binding to two F1-ATPase sectors via subunits β and γ. IF1 has no prokaryotic counterpart but is highly conserved through evolution in eukaryotic species, which indicates its functional importance (21). Remarkably, an increased IF1 expression - as found in certain tumors and cancer cells – will inhibit both the synthetic and hydrolytic activities of complex V (35).

Factor B, which has no prokaryotic homologue nor a counterpart in Saccharomyces cerevisiae, appears to bind to Fo on the matrix side (36). It seems to have a regulatory function, like IF1. In addition to the classic proton-translocating pathway in Fo, situated at the interface of subunit a and the c-ring, it has been proposed that the membrane sector Fo of animal mitochondria could harbor a second, latent proton-translocating pathway, to the assembly of which supernumerary subunits e, f, g, and A6L, as well as the ADP/ATP carrier could contribute their transmembrane segments (37). It has further been proposed that factor

B occludes the latent proton-translocating pathway (37). Doing so, factor B blocks a proton leak, keeping ∆ψm high and thus favoring ATP synthase activity (37).

Chapter 4

Assembly factors: ATP11 and ATP12

ATP11 and ATP12 bind to unassembled β and α subunits, respectively, by shielding their hydrophobic surfaces (38-39). In the absence of these assembly factors, the α-F1 and β-F1 aggregate within large inclusion bodies in the mitochondrial matrix (40-41).

TMEM70

Transmembrane protein 70 (TMEM70) has been shown to be localized in mitochondria (42).

The N-terminal part of the cytosolic 29 kDa precursor is cleaved to a 21 kDa mature mitochondrial protein (43). Analysis of submitochondrial fractions has shown that TMEM70 is associated with the inner mitochondrial membrane (43). It has been demonstrated that

TMEM70 is required to maintain normal expression levels and activity of complex V (7). It however does not interact directly with holocomplex V (43). A transient binding to complex V assembly intermediates has been proposed (44). Since TMEM70 is a low abundant mitochondrial protein, like the ATP11 and ATP12 assembly factors, a regulatory role of

TMEM70 in complex V biogenesis has been suggested (43). Further studies are necessary to clarify the exact mechanism by which TMEM70 defects lead to complex V deficiency.

Proteins associated with mammalian ATP synthase

Two ATP synthase-associated membrane proteins were identified a few years ago (45). Both proteins had been identified earlier in a different context. The first one is a 6.8-kDa mitochondrial proteolipid (MLQ protein), the second one has been denoted as diabetes- associated protein in insulin-sensitive tissue (DAPIT) (AGP protein). It has been shown recently that DAPIT has a role in maintaining ATP synthase amount in mitochondria and

Mitochondrial ATP synthase therefore possibly can influence cellular energy metabolism (46). The functional role of the

6.8-kDa proteolipid as ATP synthase-associated protein still has to be elucidated (45).

Complex V assembly

Current knowledge about the assembly of ATP synthase is mainly based on research performed on assembly-deficient yeast mutants (20). Nevertheless complex V assembly remains puzzling and partially hypothetical because the biogenesis of ATP synthase is not easily studied biochemically (47). This is due to rapid turnover of subunits of Fo and the stator in these yeast mutants (47). This problem has been addressed lately by labeling and tracking the mitochondrial gene products of Fo in isolated mitochondria (48).

It is known that the assembly of F1, the stator and the c-ring occurs separately, but the assembly sequence of subunits into the different modules and the sequence between the modules is not entirely clear (49-50). Using Clear Native Polyacrylamide Gel Electrophoresis

(CN-PAGE), performed with milder detergents than Blue Native PAGE (BN-PAGE), it has been shown in human mitochondria lacking mtDNA (and therefore subunits a and A6L) that

ATP synthase can assemble into a complex with a mass of 550 kDa (49). This is a little bit smaller than holocomplex V (597 kDa), suggesting that the only subunits lacking in the 550 kDa complex are subunit a (24.8 kDa) and A6L (8 kDa), and possibly the small associated proteins AGP and MLQ (49). This implies that the so-called F1-z complex (or V* (3)) contains the c-ring and that it is a breakdown product of the 550 kDa complex under the harsher conditions of BN-PAGE instead of an assembly intermediate (49). Based on these findings and on previous yeast work (reviewed in (20, 47)) a proposal regarding the assembly of mammalian ATP synthase has been made: assembly of the c-ring followed by binding of F1, the stator arm, and finally of subunits a and A6L (49) (Figure 2).

Chapter 4

Figure 2 Complex V assembly and dimerization. The current working model is based on assembly of the c-ring followed by binding of F1, the stator arm, and finally of subunits a and A6L. Two ATP synthase monomers dimerize via the Fo sector, where subunits a, e, g, b and A6L stabilize the monomer-monomer interface.

A recent yeast study indicated that ATP synthase is formed from three different modules: the c-ring, F1 and the Atp6p/Atp8p complex (48). It is proposed that ATP synthase assembly in yeast involves two separate pathways (F1/Atp9p and Atp6p/Atp8p/2 stator subunits/Atp10p chaperone) that converge at the end stage (48). The final addition of the mitochondrial encoded subunits to mammalian complex V is in line with the yeast studies describing that the expression of yeast subunits 6 and 8 is translationally regulated by the F1 sector (48, 51).

Therefore a balanced output can be achieved between the nuclear encoded and mtDNA encoded subunits. The role of complex V subunits in the assembly process has been studied. The peripheral stalk is important for the stability of the c-ring/F1 complex (47). It has been described that subunit A6L provides a physical link between the proton channel and the other subunits of the peripheral stalk (52). F1 subunit ε is indispensable for the assembly of holocomplex V (53). More specifically, subunit ε plays a role in the biosynthesis and

Mitochondrial ATP synthase

assembly of the F1 sector and seems to be involved in the incorporation of subunit c to the rotor structure of ATP synthase (5). Subunit c isoforms (P1, P2, and P3) are not functionally interchangeable since their targeting peptides have different roles, varying from mitochondrial import to the regulation of the assembly and function of other OXPHOS complexes (54). On the contrary, an example of over engineering is the F1 subunit γ, which only needs its N-terminal helix together with subunit δ in an upward position to catalyze ATP synthesis (55-56).

Complex V di- and oligomerization

An important role of subunits a and A6L is the stabilization of holocomplex V (49). It has been shown that ATP synthase is organized in dimers and higher oligomers (24, 57-62). The interaction between two ATP synthase monomers mainly takes place via the Fo sector

(Figure 2). It has been suggested that subunit a forms the most important basis for dimerization since it has a high number of predicted transmembrane helices (24). Next to subunit a, subunits of the stator stalk and accessory subunits (e, g, b, and A6L) stabilize the monomer-monomer interface (24, 49, 63). Another protein that has a role in the stabilization of di- and oligomers is IF1. Mature IF1 is only bound to di- and oligomeric ATP synthase in mammals (49). IF1 links two ATP synthases via the F1 sector (8).

Monocomplex V is fully capable of ATP synthesis. Nevertheless forming di- and oligomers has been proven to be beneficial to the cell. First, di- and oligomers provide stabilization of complex V, which is continuously subject to dynamic rotor/ stator interactions and can therefore be dissociated more easily (64). Secondly, oligomerization of complex V facilitates

ATP synthesis (9). This happens because dimer ribbons of ATP synthase shape the inner mitochondrial membrane (9). Because of the angular association of two monomers, dimerization leads to bending of the inner mitochondrial membrane, creating protrusions of the membrane in the matrix, called mitochondrial cristae. Clustering of ATP synthase dimers

Chapter 4 at the apex of the cristae creates a strong local positive curvature which generates a proton trap. This facilitates ATP synthesis (9). IF1 contributes to this mechanism and is therefore beneficial during ischemia, since ATP synthesis can be preserved when mitochondrial respiration is compromised (21, 34).

Complex V and mitochondrial morphology

The association of ATP synthase dimers as generating the tubular cristae has been hypothesized by Allen (65). The link between dimerization of mitochondrial ATP synthase, through subunits e, g, and –as described just recently, subunits i, k, and the biogenesis of cristae has been provided by studying yeast cells (62, 66-67). Due to cristae formation, the inner mitochondrial membrane can be divided into the inner boundary membrane and the cristae membrane. The cristae are connected to the inner boundary membrane by narrow structures, called crista junctions. In yeast, it has been suggested that the formation of cristae and crista junctions in mitochondria depends on antagonism between Fcj1 (formation of crista junction 1) and subunits e/g (68). IF1 contributes to cristae formation (34). Other components such as prohibitins or OPA1 or others yet to be identified also could contribute to crista junction and cristae tip formation (68-69).

It has been shown in yeast that in the absence of complex V dimerization (by depleting subunits e and g) normal cristae formation is hampered, resulting in onion-like structures

(66). Inhibition of F1 synthesis in yeast leads to an absence of cristae (41, 66). Another yeast study describes that the lack of Atp6p rather than an ATP production deficit modifies the overall structure of the mitochondria (70). Taken together, the complex V structure and not its enzymatic activity seems to modify cristae morphology, which is in agreement with the necessity of ATP synthase dimerization for mitochondrial cristae formation (9, 34, 66).

Mitochondrial ATP synthase

Complex V deficiency

Complex V deficiency is one of the rarer OXPHOS deficiencies. Based on the results of mitochondrial biochemical diagnostics in Nijmegen in the years 2005-2009, complex I, IV and combined enzyme deficiencies are encountered most frequently (8%, 5% and 7%, respectively of 1,406 fresh muscle samples examined), followed by complex III (3%), complex II (2%) and complex V (1%) (71).

Biochemical diagnosis

Measurement of the mitochondrial energy-generating system (MEGS) capacity in fresh muscle tissue is a powerful tool to assess mitochondrial function and to detect deficiencies of complex V and other OXPHOS complexes. The MEGS capacity can be examined in detail by measurement of 14CO2 production rates from oxidation of [1-14C]pyruvate and carboxyl-14C- labeled TCA cycle intermediates in combination with measurement of ATP production in intact mitochondria from a muscle biopsy (72), or by examining mitochondrial respiration by measuring oxygen consumption in digitonin-permeabilized cells (73). A reduced pyruvate oxidation rate that is normalized by addition of an uncoupler, e.g. carbonyl cyanide 3- chlorophenyl hydrazone (CCCP), indicates a defect in complex V, the adenine nucleotide transporter, or the phosphate carrier (71). Enzyme analysis of complex V (mtATPase) in skeletal muscle biopsy and in cultured fibroblasts remains the mainstay of the diagnostic process (74). In Nijmegen, mtATPase activity is measured spectrophotometrically in isolated mitochondria from fresh muscle tissue and fibroblasts as described (71): a solution of 980 µl containing 250 mM sucrose, 50 mM KCl, 30 mM phosphate buffer pH 7.4, 25 mM phosphate buffer pH 8.0, 0.1 mM phosphoenolpyruvate, 5 mg/L Ap5A, 0.3% BSA, 0.2 mM EGTA, 3 mM

ATP, 7.5 mM MgCl2, 250 µM NADH, 2.5 U/ml lactate dehydrogenase, and 1.5 U/ml pyruvate kinase is incubated for 10 min at 37°C. A mitochondrial suspension is freeze-thawed three times and 20 µl is added to the reaction mixture, mixed and transferred to a cuvette, after

Chapter 4 which the absorption at 340 nm is measured for 8 min. Simultaneously, the absorption is measured in a second cuvette after addition of 1 µl of an 8 mg/ml oligomycin solution in ethanol to the mitochondrial suspension and the reaction mixture. The oligomycin-sensitive

3 activity of complex V is calculated using an ε340 for NADH of 6.22x10 L/mol.cm and is expressed in units (the amount of enzyme required to convert 1 µmol NADH per min) per unit cytochrome oxidase (COX) (75) activity. Analysis of complex V in frozen tissue is considered to be less reliable than in fresh muscle tissue ((76), and personal experience (data not shown)). In addition, BN-PAGE followed by western blotting (77-78) and in-gel activity measurements of ATP hydrolysis (78) can be performed to assess the amount and activity of complex V. BN-PAGE can also provide information about the assembly of complex V (7, 79).

Molecular genetics

Hitherto, complex V mutations have been described in the mtDNA encoded ATP6 (MT-

ATP6) and ATP8 (MT-ATP8) genes. ATP synthase deficiency due to the nuclear encoded

ATP12 and TMEM70 genes has also been described (6-7). Interestingly, only one mutation has hitherto been described in a nuclear encoded structural complex V subunit, ATP5E (5).

ATP synthase subunit a; MT-ATP6 (MIM ID +516060)

Mutations in MT-ATP6 were the first complex V genetic defects to be reported (80) and have been described most frequently to date. The classical clinical picture is found in m.8993T>G/C and m.9176T>G/C mutations and covers a spectrum which varies between isolated ataxia, NARP, bilateral striatal necrosis, to Leigh or Leigh-like syndromes

(www.mitomap.org). Also other clinical features have been associated with MT-ATP6 mutations (www.mitomap.org). Below, an overview of a part of the hitherto described mutations in this gene is given, i.e. the mutations that have been well studied in humans and in yeast and those that, in our opinion, add interesting information regarding the clinical

Mitochondrial ATP synthase expression of mitochondrial complex V diseases. They have been outlined in relation to their clinical picture. m.8993T>G. This mutation leads to NARP (typically < 90-95% heteroplasmy) or MILS (when

> 95% heteroplasmy). Biochemically a decreased ATPase activity (measured spectrophotometrically) and a severely decreased ATP production rate (until 70% when >

90% heteroplasmy) is observed (81-87). Subunit a has two functions: (i) it conducts protons from the inner mitochondrial membrane to the matrix, which (ii) leads to the rotation of the ring of c-subunits that contacts subunit a in the membrane. The resulting mechanical energy is used to induce conformational changes at the level of the catalytic sites in the F1 extra- membrane domain that favor ATP synthesis (81, 88). The m.8993T>G point mutation leads to p.Leu156Arg. Leu156 is highly conserved in eukaryotes (81, 89) and is situated in the region of subunit a that regulates proton translocation coupled to rotation of the c-ring (20,

81). The introduction of a positively charged amino acid (arginine) instead of the hydrophobic leucine causes a structural change that has been shown to affect rotation of the c-ring (81).

The proton flux through Fo is slower, but not blocked (81). Furthermore, it has been shown that ATP synthase still assembles and oligomerizes correctly. This is confirmed by an intact oligomycin sensitivity (90), a property that is lost when enzyme structure is severely altered

(91-92). The assembly of F1 with Fo has only been shown to be delayed because of a slower subunit a synthesis or an increased subunit a instability and degradation (93). Taken together, the severe impairment of ATP synthesis is due to functional inhibition in a correctly assembled ATP synthase (90). m.8993T>C. The clinical picture is similar to the m.8993T>G mutation, but is generally milder

(87, 94-96). The mutation results in p.Leu156Pro. Opposite to the positive charged arginine in the m.8993T>G mutation, proline is a non-charged amino acid. Biochemically, ATP synthesis is not severely affected (81). So this does not fully explain the NARP or MILS phenotypes seen in patients. It has been shown that the m.8993T>C mutation favors

Reactive Oxygen Species (ROS) production, which is believed to be the major pathogenic

Chapter 4 mechanism (81). ROS plays a major role in the pathogenesis of neurological diseases related to mitochondrial dysfunction (97-98). m.9176T>G/C. The clinical picture is characterized by familial bilateral striatal necrosis

(FBSN) and Leigh syndrome (99-100). A hereditary spastic paraplegia-like disorder has been described in a family carrying the homoplasmic m.9176T>C mutation (101). The mutations also change a highly conserved hydrophobic leucine residue into arginine (m.9176T>G) or proline (m.9176T>C) at position 217 of the protein. Position 217 of subunit a also lies in proximity to the c-ring (70). It has been shown in human cells with a high degree of heteroplasmy that the m.9176T>G mutation severely decreases the rate of ATP production

(102). In yeast carrying the m.9176T>G mutation, it has been described that that incorporation of subunit a into ATP synthase was almost completely prevented (70). This resulted in a disturbed assembly which profoundly altered mitochondrial morphology (70).

This finding is in contrast to the m.8993T>G mutation, where the assembly is not altered (as mentioned before). Taken together, the pathogenicity of m.9176T>G may not be limited to a bio-energetic deficiency, but may also be attributed to an altered mitochondrial and cristae morphology (70). Also the m.9176T>C mutation has been studied in yeast, only showing a mild decrease in ATP production (88). There is a poor correlation between genotype and phenotype for the m.9176T>C mutation (88). Additional determinants may be responsible and are a plausible explanation for this finding. One of these determinants has been proposed to be the higher degree of oligomycin vulnerability in the mutant than in wild type

(88).

In tightly coupled mitochondria the oxygen consumption rate and ATP synthesis depend on each other (103). Therefore it is not very surprising to see a decreased respiratory activity in many ATP synthase defective mutants (103). In several yeast models with NARP-MILS mutations (atp6-p.Leu183Pro (104), atp6-p.Leu183Arg (105), atp6-p.Leu247Arg (70), atp6- p.Leu247Pro (88), and ∆atp6 (103)), it has been shown that there is a linear correlation between ATP synthase activity and complex IV activity (88). So it appears that complex IV is

Mitochondrial ATP synthase an important target for the co-regulation of electron transfer and ATP synthesis in mitochondria (88). m.9035T>C, m.9185T>C, m.9191T>C, m.8851T>C. These mutations are illustrations of variants of the earlier described NARP-MILS clinical spectrum. m.9035T>C has been found in a family with maternally transmitted cognitive developmental delay, learning disability, and progressive ataxia (106). Mutant cybrids had less than half of the steady-state content of

ATP and a substantial higher basal level of Reactive Oxygen Species (ROS) (106). m.9185T>C (p.Leu220Pro) results in a phenotype varying from mild learning difficulties and foot deformities, ataxia, an acute neurological presentation with complete recovery, to Leigh syndrome (107-108). m.9191T>C (p.Leu222Pro) has been described in a patient with Leigh syndrome (108). The pathogenic mechanism of the m.9185T>C and m.9191T>C mutations is unknown, but they are situated in a highly conserved region of the protein. The observed phenotypical differences are proposed to result from additional genetic and/or environmental modifying factors (107). A patient suffering from bilateral striatal necrosis and harboring a m.8851T>C mutation has also been described (109). This phenotype is less severe than found in typical Leigh disease (109).

∆TA9205. This mutation has been reported in a patient with seizures and lactic acidemia

(110). ∆TA9205 is situated at the junction between the two genes MT-ATP6 and MTCO3

(which codes for COX3, a complex IV subunit). It removes the termination codon from

RNA14, the bi-cistronic RNA unit encoding MT-ATP8 and MT-ATP6, which has been shown to cause a decrease in the steady-state level of the mutated RNA14 (111). The level of

RNA15, the RNA transcribed from MTCO3, is not affected (111). This mutation illustrates how a mtDNA mutation can influence the turnover of a human mitochondrial mRNA.

Modifiers. Phenotypical variations between patients harboring the same mtDNA mutation have classically been attributed to mtDNA heteroplasmy. However, a discrepancy between the levels of mutant mtDNA and disease severity is sometimes observed (93). The mtDNA background has been shown to play an important role in modulating the biochemical defects and clinical outcome (93). MT-ATP6 mutations m.8741T>G and m.8795A>G have been

Chapter 4 shown to be protective factors for the m.8993T>G MILS mutation (112). A MT-ATP6 polymorphism, m.9055G>A, significantly reduces the risk of Parkinson disease, especially in women (113). The mitochondrial haplogroup J has been shown to be associated with longevity and protection against certain diseases (101). Vice versa, respiratory chain defects upstream of ATP synthase contribute to ATP synthesis impairment, worsening the biochemical and clinical phenotype e.g. of m.8993T>G/C and m.9176T>G mutations (93).

The germ-line mutation m.8932C>T has, together with somatic mtDNA mutations in the cytochrome oxidase subunit I (COI) gene, been associated with prostate cancer (114-115).

The mtDNA variants m.8836T>G, m.9016A>G, m.9101T>C, m.9139G>A have been linked with patients who had Leber Hereditary Optic Neuropathy (LHON) or LHON-like optic neuropathies (116-119). Hence, mtDNA background can explain the biochemical and clinical variations observed between patients with the same mutation and a comparable mutation load. This also illustrates the role of mtDNA variants in the complexity of mtDNA disease expression. In general, phenotypic variability is proposed to be the result of interactions between the casual genes, genetic background or modifier genes (mitochondrial or nuclear), and probably environmental factors (120).

ATP synthase subunit A6L; MT-ATP8 (MIM ID +516070)

m.8528T>C. This homoplasmic mutation has been found in four unrelated patients who presented as infants with isolated hypertrophic cardiomyopathy and congestive heart failure, evolving to multisystem disease (121). Electron microscopy of muscle tissue showed increased variation in size and shape of mitochondria with dense parallel cristae in one patient (121). For A6L, m.8528T>C (p. Trp55Arg) is a pathogenic missense mutation, replacing a hydrophobic tryptophan into a basic arginine in a conserved region of the subunit.

In subunit a, the nucleotide alteration results in the change of the initiation methionine to threonine, but the effect of this mutation on the function of this subunit is not clear (121).

Mitochondrial ATP synthase m.8411A>G. The de novo mutation with 97% heteroplasmy has been described in a patient with psychomotor delay, epilepsy, tetraplegia, congenital deafness, central blindness and swallowing difficulties (122). He died at the age of 10 years. Brain MRI showed involvement of the interpeduncular nucleus, the central tegmental tract, the white matter and the cerebellum. This leucodystrophy phenotype differs from the other mutations described in MT-

ATP8, where cardiac involvement predominates. Leucodystrophy has been described in two siblings with Leigh syndrome harboring the m.9176T>C mutation in MT-ATP6 (123). It should be noted that complex V enzyme analysis was not performed in this patient (122), and therefore the pathogenicity of the m.8411A>G mutation has not been firmly established yet.

ATP12 (MIM ID *608918)

A homozygous T>A mutation, changing an evolutionary conserved tryptophan to an arginine at position 94 (p.Trp94Arg) of the nuclear encoded complex V assembly gene ATP12 has been found in a patient who presented with a severe neonatal encephalopathy and dysmorphic features, evolving to basal ganglia atrophy within months and death at the age of

14 months (6, 124). The mutation resulted in a severely decreased complex V amount and activity. It was hypothesized that the change of a neutral polar amino acid (tryptophan) into a basic one (arginine) resulted in an ATP12 protein that was no longer able to mediate F1 assembly (6). Later, it has been shown in a yeast model that the p.Trp94Arg mutation decreases the solubility of the protein, implying that the primary impact of the mutation was a change in physical rather than functional parameters (124). Mitochondrial morphology has been studied in fibroblasts, but no alterations compared to control cells could be observed

(124).

TMEM70 (MIM ID *612418)

Chapter 4

The TMEM70 gene encodes a mitochondrial protein, transmembrane protein 70. A common splice site mutation and an isolated frameshift mutation have been described in the TMEM70 gene particularly in a homogeneous ethnic group (Romanies), with a clinical phenotype characterized by neonatal mitochondrial encephalocardiomyopathy, lactic acidosis and dysmorphic features (7). The description of four novel mutations in the TMEM70 gene has confirmed and expanded this classical clinical picture with early onset cataract, gastrointestinal dysfunction, congenital hypertonia and a fetal presentation of the syndrome

(125). One patient with a milder clinical phenotype carrying the common splice site mutation and a missense variant has also been described (126). This phenotype corroborates with two reported patients who are compound heterozygous for the common splice site mutation and a frame-shift mutation in exon 1 (44, 127). It was shown that TMEM70 is required to maintain normal expression levels of complex V (7). The exact mechanism behind this remains to be elucidated, although it has been suggested that TMEM70 is involved in complex V biogenesis (7).

ATP synthase subunit epsilon; ATP5E (MIM ID *606153)

Recently, a homozygous missense mutation c.35A>G (p.Tyr12Cys) has been described in exon 2 of the ATP5E gene (5). The patient was a 22 year old woman presenting with neonatal onset, lactic acidosis, 3-methylglutaconic aciduria, and mild mental retardation. She developed peripheral neuropathy (5). The mutation caused a decrease in the amount and activity of holocomplex V. Remarkably, subunit c was found to accumulate, in contrast to the other investigated complex V subunits, and in contrast to what was found in patients with a mutation in the TMEM70 (127) and ATP12 (6) genes (5). Using pulse-chase experiments, this study pointed to the crucial role of subunit ε in the biosynthesis and assembly of the F1 part of ATP synthase. Moreover, subunit ε seems to be involved in the incorporation of subunit c into the rotor (5).

Mitochondrial ATP synthase

Therapy

Current available treatment options for patients with mitochondrial diseases are mainly supportive. Therefore, a lot of effort is put into the search for both pharmacological and genetic approaches to cure this devastating group of disorders (for reviews, see (128-130)).

Focusing here on complex V deficiency, current research has mainly covered the therapy of mtDNA mutations.

Antioxidants

As mentioned above, complex V mutations can increase ROS production which is deleterious for the cell. The antioxidants N-acetylcysteine (NAC) and dihydrolipoic acid

(DHLPA) have therefore been tested in fibroblasts harboring 97% m.8993T>G mutant mtDNA. It has been shown that they significantly improved mitochondrial respiration and ATP synthesis in these cells (131).

Substrates: α-Ketoglutarate/Aspartate

In yeast studies, it has been suggested that forcing substrate-level phosphorylation to work overtime may counteract the energy crisis due to OXPHOS impairment (132).

For the application of this approach in OXPHOS-deficient human cells, exogenous substrates capable of stimulating the Krebs cycle flux while at the same time removing the excess of reduced nicotinamide adenine dinucleotide (NADH) have been chosen (133). It has been demonstrated that homoplasmic cybrids harboring the m.8893T>G mutation were protected from cell death and had an ATP content similar to controls after supplementation of the culture medium with α-ketoglutarate and aspartate (133).

Chapter 4

Affecting heteroplasmy of the mtDNA (gene-shifting)

This genetic approach aims to force a shift in heteroplasmy, reducing the ratio of mutant to wild-type genomes (also called gene-shifting) (128). Various methods can be applied to achieve this goal.

Allotopic expression. Here, a normal version of a mutant mtDNA-encoded protein is imported into the nucleus. For example, MT-ATP6 can be converted from the mitochondrial into the nuclear genetic code (128, 134). To be transported to the mitochondrion, it has to be provided with a mitochondrial targeting signal, of which the genetic sequence can be borrowed from another mtDNA-encoded protein. The biochemical defect in cybrid cell lines harboring the m.8993T>G mutation has been corrected successfully using this strategy

(134).

Xenotopic expression. The correction here implies the transfection of mammalian cells with either mitochondrial or nuclear genes from other organisms encoding the protein of interest

(128). This has also been applied to human cybrids harboring the m.8993T>G mutation by expressing the nDNA encoded ATPase6 protein of the alga Clamydomonas reinhardtii, which already possessed the mitochondrial targeting sequence since it is nuclear encoded

(135). Also this approach could correct the biochemical defect in these cells (135).

Restriction endonucleases. These are specific proteins that cut mutant mtDNA but not wild type mtDNA (128). For example, the m.8993T>G mutation creates a unique cleavage site for the restriction endonuclease SmaI. The gene for SmaI was fused to a mitochondrial targeting sequence and expressed in heteroplasmic mutant cybrid cells, which lost mutant mtDNA and recovered biochemically (136-137).

Oligomycin. It has been shown that culturing heteroplasmic m.8993T>G cybrid cells in medium containing oligomycin (complex V inhibitor) and galactose (which forces the cells to rely on oxidative metabolism for ATP production) allowed for the selection of wild type over mutant mtDNA (138).

Mitochondrial ATP synthase

Germline therapy

It has been proposed that nuclear transfer techniques may be an approach for the prevention of transmission of human mtDNA disease (139-140). Briefly, this means that the pronucleus of an oocyte, or zygote, of a woman carrying mutated mtDNA could be transferred to a donor enucleated oocyte, or zygote, carrying wild type mtDNA (128, 141). In that way, the offspring will carry all the nuclear – and physiognomonic - traits of the parents, but not the mutated mtDNA of the mother (128). Two successful approaches have recently been described.

Metaphase II spindle transfer between unfertilized metaphase II oocytes. It has been demonstrated in mature non-human primate oocytes (Macaca mulatta) that the mitochondrial genome can be efficiently replaced by spindle-chromosomal complex transfer from one egg to an enucleated, mitochondrial-replete egg (142). Subsequently, it was possible to have normal fertilization and embryo development. The offspring was healthy (142).

Pronuclear transfer between zygotes. This is essentially the same procedure, except that the nuclear material, both the male and female pronucleus, is removed after fertilization

(142). It has been shown that transfer of pronuclei between abnormally fertilized human zygotes resulted in minimal carry-over of donor zygote mtDNA and is compatible with onward development of the blastocyst stage in vitro (141).

There have been few randomized controlled trials for the treatment of mitochondrial disease

(143). To date, there is no clear evidence supporting the use of pharmacological agents, non-pharmacological treatments (vitamins and food supplements), and physical training in patients with mitochondrial disorders (143). Although very promising, all genetic techniques are still in an experimental phase and different technical, ethical and safety issues still have to be solved (20, 128, 141-142). Nevertheless, they do allow cautious optimism for the future.

Chapter 4

Prenatal and preimplantation diagnosis

Since current therapeutic options for mitochondrial diseases are insufficient, the possibility of prenatal diagnosis for fetuses at risk is a valuable alternative. If it concerns a known nuclear genetic defect, the mutation can directly be searched for in fetal tissue. If the complex V deficiency is caused by a mtDNA defect (in MT-ATP6 or MT-ATP8), prenatal diagnosis is complicated by several factors. First, the correlation between mutant mtDNA load and disease severity is poor in many mtDNA diseases. Second, the heteroplasmy level differs between tissues and in one tissue through time (144). In this context, the m.8993 mutation

(NARP mutation) is an exception. There is a correlation between mutant mtDNA load and disease severity (144). Moreover, it has been shown that the placental/ amniotic mutant loads do reflect the NARP mutant mtDNA load in the whole fetus (145-146). Finally, is suggested that the heteroplasmy level remains stable after 10 weeks of gestation (146). A mutation load <30% gave rise to healthy children at 2-7 years of age, while the correlation between an intermediate mutant load (> 30% and < 80%) and disease severity still needs to be assessed (146). Hitherto termination of pregnancy has been preferred in case of intermediate mutant loads (146). Remarkably, the intermediate mutant loads question the observation that the m.8993T>G mutation has a skewed segregation during oogenesis (147).

Post-zygotic drift might explain this discrepancy (146). Current options for women with MT-

ATP6 or MT-ATP8 mutations can be oocyte donation or preimplantation genetic diagnosis

(PGD), since the different heteroplasmy levels between tissues are not yet present in blastomeres (144, 148-149). The interpretation of PGD results nevertheless demands a known correlation between mutation load and clinical phenotype. In addition, caution is warranted since some pathogenic mutations could exhibit different segregation behavior

(149). In case the genetic examination of an index case has revealed no mutations in both mtDNA and nDNA, prenatal diagnosis could still be possible. In Nijmegen, complex V activity can be measured spectrophotometrically in native chorionic villi, cultured chorionic cells or

Mitochondrial ATP synthase cultured amniotic cells if there is a clear isolated complex V deficiency in fibroblasts and muscle tissue (or other tissue) of the index patient (71, 150).

REFERENCES

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Chapter 4

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132. Schwimmer C, Lefebvre-Legendre L, Rak M, Devin A, Slonimski PP, di Rago JP, et al. Increasing mitochondrial substrate-level phosphorylation can rescue respiratory growth of an ATP synthase-deficient yeast. J Biol Chem. 2005 Sep 2;280(35):30751-9. 133. Sgarbi G, Casalena GA, Baracca A, Lenaz G, DiMauro S, Solaini G. Human NARP mitochondrial mutation metabolism corrected with alpha-ketoglutarate/aspartate: a potential new therapy. Arch Neurol. 2009 Aug;66(8):951-7. 134. Manfredi G, Fu J, Ojaimi J, Sadlock JE, Kwong JQ, Guy J, et al. Rescue of a deficiency in ATP synthesis by transfer of MTATP6, a mitochondrial DNA-encoded gene, to the nucleus. Nat Genet. 2002 Apr;30(4):394-9. 135. Ojaimi J, Pan J, Santra S, Snell WJ, Schon EA. An algal nucleus-encoded subunit of mitochondrial ATP synthase rescues a defect in the analogous human mitochondrial- encoded subunit. Mol Biol Cell. 2002 Nov;13(11):3836-44. 136. Tanaka M, Borgeld HJ, Zhang J, Muramatsu S, Gong JS, Yoneda M, et al. Gene therapy for mitochondrial disease by delivering restriction endonuclease SmaI into mitochondria. J Biomed Sci. 2002;9(6 Pt 1):534-41. 137. Alexeyev MF, Venediktova N, Pastukh V, Shokolenko I, Bonilla G, Wilson GL. Selective elimination of mutant mitochondrial genomes as therapeutic strategy for the treatment of NARP and MILS syndromes. Gene Ther. 2008 Apr;15(7):516-23. 138. Manfredi G, Gupta N, Vazquez-Memije ME, Sadlock JE, Spinazzola A, De Vivo DC, et al. Oligomycin induces a decrease in the cellular content of a pathogenic mutation in the human mitochondrial ATPase 6 gene. J Biol Chem. 1999 Apr 2;274(14):9386-91. 139. Sato A, Kono T, Nakada K, Ishikawa K, Inoue S, Yonekawa H, et al. Gene therapy for progeny of mito-mice carrying pathogenic mtDNA by nuclear transplantation. Proc Natl Acad Sci U S A. 2005 Nov 15;102(46):16765-70. 140. Brown DT, Herbert M, Lamb VK, Chinnery PF, Taylor RW, Lightowlers RN, et al. Transmission of mitochondrial DNA disorders: possibilities for the future. Lancet. 2006 Jul 1;368(9529):87-9. 141. Craven L, Tuppen HA, Greggains GD, Harbottle SJ, Murphy JL, Cree LM, et al. Pronuclear transfer in human embryos to prevent transmission of mitochondrial DNA disease. Nature. 2010 May 6;465(7294):82-5. 142. Tachibana M, Sparman M, Sritanaudomchai H, Ma H, Clepper L, Woodward J, et al. Mitochondrial gene replacement in primate offspring and embryonic stem cells. Nature. 2009 Sep 17;461(7262):367-72. 143. Chinnery P, Majamaa K, Turnbull D, Thorburn D. Treatment for mitochondrial disorders. Cochrane Database Syst Rev. 2006(1):CD004426.

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144. Poulton J, Marchington DR. Segregation of mitochondrial DNA (mtDNA) in human oocytes and in animal models of mtDNA disease: clinical implications. Reproduction. 2002 Jun;123(6):751-5. 145. Dahl HH, Thorburn DR, White SL. Towards reliable prenatal diagnosis of mtDNA point mutations: studies of nt8993 mutations in oocytes, fetal tissues, children and adults. Hum Reprod. 2000 Jul;15 Suppl 2:246-55. 146. Steffann J, Gigarel N, Corcos J, Bonniere M, Encha-Razavi F, Sinico M, et al. Stability of the m.8993T->G mtDNA mutation load during human embryofetal development has implications for the feasibility of prenatal diagnosis in NARP syndrome. J Med Genet. 2007 Oct;44(10):664-9. 147. Blok RB, Gook DA, Thorburn DR, Dahl HH. Skewed segregation of the mtDNA nt 8993 (T-->G) mutation in human oocytes. Am J Hum Genet. 1997 Jun;60(6):1495-501. 148. Jenuth JP, Peterson AC, Shoubridge EA. Tissue-specific selection for different mtDNA genotypes in heteroplasmic mice. Nat Genet. 1997 May;16(1):93-5. 149. Dean NL, Battersby BJ, Ao A, Gosden RG, Tan SL, Shoubridge EA, et al. Prospect of preimplantation genetic diagnosis for heritable mitochondrial DNA diseases. Mol Hum Reprod. 2003 Oct;9(10):631-8. 150. Niers L, van den Heuvel L, Trijbels F, Sengers R, Smeitink J. Prerequisites and strategies for prenatal diagnosis of respiratory chain deficiency in chorionic villi. J Inherit Metab Dis. 2003;26(7):647-58.

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A novel mitochondrial ATP8 (MT-ATP8) gene mutation in a

patient with apical hypertrophic cardiomyopathy and

neuropathy

An I. Jonckheere, Marije Hogeveen, Leo G.J. Nijtmans, Mariël A.M. van den Brand, Antoon

J.M. Janssen, Heleen H.S. Diepstra, Frans C. van den Brandt, Lambert P. van den Heuvel,

Frans A. Hol, Tom G.J. Hofste, Livia Kapusta, Ulrich Dillmann, Mohammed G. Shamdeen,

Jan A.M. Smeitink, Richard J.T. Rodenburg

Journal of Medical Genetics 2008; 45(3):129-33

Chapter 5

ABSTRACT

Purpose: To elucidate the biochemical and molecular genetic defect in a 16-year-old patient presenting with apical hypertrophic cardiomyopathy and neuropathy suspected for a mitochondrial disorder.

Methods: Measurement of the mitochondrial energy-generating system (MEGS) capacity in muscle and enzyme analysis in muscle and fibroblasts were performed. Relevant parts of the mitochondrial DNA were analyzed by sequencing. Transmitochondrial cybrids were obtained by fusion of 143B206 TK- rho zero cells with patient-derived enucleated fibroblasts.

Immunoblotting techniques were applied to study the complex V assembly.

Results: A homoplasmic nonsense mutation m.8529G>A (p.Trp55X) was detected in the mitochondrial ATP8 gene in the patient’s fibroblasts and muscle tissue. A decreased complex V activity was measured in the patient’s fibroblasts and muscle tissue, and was confirmed in cybrid clones containing patient derived mitochondrial DNA. Immunoblotting after Blue Native Polyacrylamide Gel Electrophoresis (BN-PAGE) revealed a lack of holocomplex V and increased amounts of subcomplexes of mitochondrial ATP synthase. In- gel activity assay of ATP hydrolysis showed activity of free F1-ATPase in the patient’s muscle tissue and cybrid clones.

Conclusion: We describe the first pathogenic mutation in the mitochondrial ATP8 gene, resulting in an improper assembly and a decreased activity of the complex V holoenzyme.

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INTRODUCTION

Mitochondrial (mt) ATP synthase, or complex V (EC 3.6.3.14), uses the proton gradient provided by the activity of the respiratory chain enzymes complexes I, III and IV for ATP synthesis, thereby generating over 95% of cellular ATP (1). Complex V is a multisubunit complex consisting of two functional domains, F1 and Fo, connected by a stalk. The F1 domain contains 5 different subunits (3 alpha’s, 3 beta’s, and one gamma, delta, and epsilon), is situated in the mitochondrial matrix and acts as the catalytic domain. The Fo domain is embedded in the mitochondrial inner membrane and consists of 8 subunits

(a,b,c,d,e,f,g and A6L). The stalk contains the subunits OSCP, F6, b and d (2). Protons pass from the intermembrane space to the matrix through Fo, which transfers the energy created by the proton electrochemical gradient to F1, where ADP is phosphorylated to ATP (2-3). It has been proposed that a shaft (“rotor” consisting of the ring of 10 c subunits, plus the subunits gamma, delta and epsilon that form the central stalk) linked to Fo rotates relative to the catalytic subunits (3 alpha, 3 beta) of F1, thereby sequentially changing the conformation of these subunits, creating the “binding change mechanism of energy coupling” proposed by

Boyer (4-6). Two of the Fo subunits, subunit a (or subunit 6) and subunit A6L (or subunit 8) are encoded by the mtDNA ATP6 and ATP8 genes, respectively (7). Hitherto mtDNA complex V mutations have only been described in the mitochondrial ATP6 gene (MT-ATP6).

The point mutations m.8993T>G/C are the most frequently encountered MT-ATP6 mutations and, depending on the degree of heteroplasmy, mainly lead to the clinical picture of NARP

(Neuropathy, Ataxia, and Retinitis Pigmentosa) or, more severe, MILS (Maternally Inherited

Leigh Syndrome) (1, 8). ATP synthase deficiency due to nuclear genetic defects has also been described (1, 9). To our knowledge pathogenic mutations in the mitochondrial ATP8 gene have not been published before. Here we describe a 16-year-old patient presenting with apical hypertrophic cardiomyopathy and neuropathy. Biochemical analysis performed in muscle tissue and fibroblasts showed an isolated complex V deficiency. MtDNA mutation

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Chapter 5 analysis revealed a pathogenic nonsense mutation in MT-ATP8. We studied the molecular pathophysiology of this novel mtDNA mutation using transmitochondrial cybrid cells.

CASE HISTORY

The patient is the third child of healthy, non-consanguineous Caucasian parents. His older brother and sister are healthy. The family history is unremarkable. His motor development has been slower than the sibs since infancy. He walked independently at the age of 1 ½ - 2 years, and learned to ride a bicycle at the age of 7-8 years. During childhood, he fell easily, and his gross motor skills were clumsy. According to the parents his speech development was delayed but turned to normal after an adenotomy at the age of 4 years. He is wearing glasses since the age of 3 years for hypermetropia and astigmatism, and one eye was temporarily covered because of amblyopia. At the age of 9-10 years the balance problems became prominent. Since then he also mentions decreased muscle strength, walking problems, and tingling in the extremities. These symptoms appear to be progressive.

Exercise intolerance, shortness of breath (dyspnea) during exercise and angina were also common symptoms. The clinical examination at the age of 16 years revealed a slender young man. His speech was dysarthric. He had an ataxic walking pattern and revealed a clear Trendelenburg sign. Decreased tendon reflexes and a Babinski sign bilaterally were present. A moderate external ophthalmoplegia was noted. The metabolic tests performed at the age of 14 years revealed normal blood lactic acid concentrations, with increased lactic acid in the cerebrospinal fluid (4000 µmol/l, normal values 1350-1900 µmol/l). The organic acid pattern in urine was normal. The ophthalmologic evaluation confirmed the astigmatism and hypermetropia, and showed no signs of retinitis pigmentosa. Audiology was normal. The

MRI of the brain did not demonstrate any abnormalities besides a cisterna magna.

Neuropsychological evaluation revealed a moderate learning disorder. Cardiac evaluation showed an extensive left ventricular hypertrophy on the ECG, without signs of arrhythmia.

Echocardiography was abnormal: left ventricular hypertrophy was concentrated around the

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A novel mitochondrial ATP8 gene mutation apex of the left ventricle (Figure 1) with discrete asymmetric hypertrophy of the echogenic interventricular septum and without left ventricular outflow tract obstruction. A small atrial septal defect with left to right shunt was detected. A sensor and motor axonal polyneuropathy was seen on an electromyogram at the age of 16 years: the sensible potentials were not evocable in the sural nerve bilaterally, and there were slightly delayed motor nerve conduction velocities with decreased amplitudes in the peroneal and tibial nerves. Finally a muscle (M. quadriceps femoris) and skin biopsy was performed.

Figure 1 Echocardiography showing apical left ventricular hypertrophy. LA = Left Atrium; RA = Right Atrium; LV = Left Ventricle; RV = Right Ventricle. The arrows indicate the myocardial hypertrophy.

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METHODS

Cell cultures

Fibroblasts were cultured in M199 medium (Gibco®, Invitrogen Corporation) supplemented with 10% fetal calf serum and penicillin/streptomycin (respectively 100U/ml and 100 µg/ml).

Transmitochondrial cybrids were obtained by fusion of 143B206 TK- rho zero cells with control or patient-derived enucleated fibroblasts as described before (10). Seven independent colonies (clones) of the patient transmitochondrial cybrid cell line were randomly picked for further study. For the control cybrid cell line the colonies were pooled, resulting in a clone mixture.

Biochemical assays

Measurement of MEGS capacity was performed as described before (11). The complex V (or mtATPase) activity was measured spectrophotometrically in mitochondria isolated from frozen pellets of the patient and control cybrids as described (8): a solution of 980 µl containing 250 mM sucrose, 50 mM KCl, 50 mM K2HPO4, 50 mM KH2PO4, 1 mM phosphoenolpyruvic acid, 11 nM P1-P5-di(adenosine-5’) pentaphosphate, 0.2 mM EGTA, 5 mM ATP, 5 mM MgCl2, 0.2 mM oubaine, 2 µM carbonyl cyanide 3-chlorophenyl hydrazone

(CCCP), 250 µM NADH, 1.25 µM rotenone, 2.5 U/ml lactate dehydrogenase, and 1.5 U/ml pyruvate kinase was incubated for 10 min at 30°C. A mitochondrial suspension was freeze- thawed three times and 20 µl was added to the reaction mixture, mixed and transferred to a cuvette, after which the absorption at 340 nm was measured for 8 min. Subsequently, 1 µl of a 8 mg/ml oligomycin solution in ethanol was added and the absorption was measured for another 8 min. The oligomycin-sensitive activity of complex V was calculated using an ε340 for

NADH of 6.22x103 L/mol.cm and was expressed in units (the amount of enzyme required to convert 1 µmol NADH per min) per unit cytochrome oxidase (COX) (12) and the

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A novel mitochondrial ATP8 gene mutation mitochondrial matrix enzyme citrate synthase (CS) (13) activities, respectively. The activities of the mitochondrial respiratory chain enzymes were measured spectrophotometrically in muscle tissue and fibroblasts of our patient according to described protocols (14-15).

Molecular genetic analysis

Total DNA from frozen skeletal muscle tissue and from cultured fibroblasts was extracted using the Gentra genomic isolation kit according the protocol of the manufacturer (Biozym,

Landgraaf, The Netherlands). Major DNA deletions or other rearrangements in the mtDNA were screened for using standard long template PCR. The presence of the MELAS m.3243A>G, MERFF m.8344A>G and Leigh/NARP m.8993T>G/C point mutations was investigated using Pyrosequencing™ technology as described previously (8, 16). Sequence analysis of PCR amplified products of the mtDNA encoded tRNAGln, tRNAGly, tRNAIle, tRNALeu(cun), tRNALeu(uur), tRNALys, tRNASer(agy), mitochondrial ATP6 and ATP8 genes was performed on an ABI3730 automatic capillary sequencer using BigDye® terminator chemistry (Applied Biosystems). Primers used for amplification of tRNA genes are available upon request. MT-ATP6 and MT-ATP8 were PCR amplified as 3 overlapping fragments (a, b and c) using the following primers: For-a: 5’-cggtcaatgctctgaaatctgtg-3’; Rev-a: 5’- gagatatttggaggtggggatc-3’; For-b: 5’-ctgttcgcttcattcattgcc-3’; Rev-b: 5’-gtggcgcttccaattaggtg-

3’; For-c: 5’-cccacttcttaccacaaggc-3’; Rev-c: 5’-gtgctttctcgtgttacatcg-3’. PCR conditions were

92 ºC for 30 seconds, 55 ºC for 30 seconds and 72 ºC for 60 seconds for a total of 35 cycles.

Subsequently, mtDNA was analyzed in the patient and control cybrids looking for the nonsense mutation m.8529G>A following the protocol described above.

Complex V assembly and activity

BN-PAGE, blotting and complex V in-gel activity was performed as described (17-18) loading

60 µg of protein (19) of OXPHOS complexes isolated from a mitoplast fraction of the patient and control cybrids and muscle homogenates. For Western blotting, monoclonal antibodies

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Chapter 5 against mitochondrial ATP synthase subunit alpha and CoII-70 kDa (complex II) (Invitrogen,

Breda, The Netherlands) were used. For the complex V in-gel activity assay the gel was incubated overnight at room temperature with the following solution: 35 mM Tris, 270 mM glycine, 14 mM MgSO4, 0.2% Pb(NO3)2, and 8 mM ATP, pH 7.8 (18).

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A novel mitochondrial ATP8 gene mutation

RESULTS

Biochemical assays

Measurement of MEGS capacity in muscle tissue showed a decreased pyruvate oxidation rate, which improved after addition of CCCP, which could point to a complex V defect (11).

The ATP production rate from oxidation of pyruvate was decreased. Spectrophotometric analysis confirmed the decreased complex V (mtATPase) activity in muscle tissue and in fibroblasts. The activities of the respiratory chain enzymes and CS were normal. (Table 1)

Table 1: Measurement of MEGS capacity in muscle tissue and enzymatic analysis of the activities of complex V (mtATPase), the respiratory chain enzymes and CS in muscle tissue and fibroblasts.

Muscle tissue Fibroblasts

[1-14C]Pyruvate + malate 2.25 n.d.

14 (nmol CO2/h.mU CS; normal 3.45-7.99)

[1-14C]Pyruvate + malate-ADP + CCCP 4.39 n.d.

14 (nmol CO2/h.mU CS; normal 3.76-8.37)

ATP production rate 22 n.d.

(nmol (ATP+CrP)/h.mU CS; normal 36-81.7)

Complex V 54 mU/UCOX 122 mU/UCOX

(normal 169-482) (normal 209-935)

Complex I 73 mU/UCS 223 mU/UCOX

(normal 70-251) (normal 110-260)

Complex II 518 mU/UCS 812 mU/UCOX

(normal 335-749) (normal 536-1027)

Complex III 3438 mU/UCS 1998 mU/UCOX

(normal 2200-6610) (normal 1270-

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2620)

Complex IV 2134 mU/UCS 797 mU/UCS

(normal 810-3120) (normal 680-1190)

CS 231 mU /mg 398 mU/mg

(normal 37.4-162) (normal 144-257)

MEGS: mitochondrial energy-generating system; CCCP: carbonyl cyanide 3-chlorophenyl hydrazone; COX: cytochrome oxidase; CS: citrate synthase; n.d.: not determined.

Molecular genetic analysis

The diagnosis of a mitochondriopathy was further substantiated by mutation analysis of mtDNA from muscle tissue and fibroblasts, showing a homoplasmic (>90%) nonsense mutation m.8529G>A (p.Trp55X) in the mitochondrial ATP8 gene. The mutation is located in an overlap region of MT-ATP6 and 8 (nucleotides m.8527-8572). It results in a silent change in MT-ATP6 (Met1Met; ATG>ATA), whereas it introduces a premature stopcodon in the C- terminal domain of MT-ATP8, which is a conserved region (20) (Figure 2). This probably results in a truncated protein lacking the last 14 amino acids.

Figure 2 The C-terminal domain of mt ATP synthase subunit 8 is a conserved region in mammals. p.Trp55X indicates the mutation site at the protein level.

Cell biological consequences

DNA analysis in the cybrid cells confirmed the presence of the mitochondrial nonsense mutation m.8529G>A in each of the clones containing patient-derived mtDNA. Sequence

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A novel mitochondrial ATP8 gene mutation analysis showed that all clones were homoplasmic for the mutation. The control cybrid cell line showed no mutation (Figure 3).

Figure 3 Molecular genetic analysis of the cybrid clones containing patient-derived mtDNA reveals the mt nonsense mutation m.8529G>A (arrow). Top row: control cybrid cell line; subsequent row: patient cybrid clone. Sequence analysis shows that all the clones are homoplasmic for the mutation (data not shown).

Enzymatic analysis of complex V was performed in the patient cybrid clones and in the control cybrid clone mixture. A distinctly decreased activity was found in the patient cell line relative to the control (Table 2). CS and COX activities were normal in the patient and control cybrids (data not shown).

Table 2: Enzymatic analysis of the complex V (mtATPase) activity in cybrids.

Complex V/COX (mU/U COX) Complex V/CS (mU/U CS)

Control cybrid clone 2.0 0.3 mixture

Patient cybrid clone 1 0.45 0.09

Patient cybrid clone 2 0.08 0.02

Patient cybrid clone 3 0 0

Patient cybrid clone 4 0.56 0.09

Patient cybrid clone 5 0 0

Patient cybrid clone 6 0.87 0.07

Patient cybrid clone 7 0.23 0.07

COX: cytochrome oxidase; CS: citrate synthase

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A higher range of residual activities in individual cybrids is seen when normalized to the COX activity because the COX assay has a broader normal range than the CS assay.

Immunoblotting after BN-PAGE (Figure 4) with anti-ATP synthase subunit alpha antibody clearly revealed a lack of holocomplex V and increased amounts of subcomplexes of mitochondrial ATP synthase in the patient cybrid clones and muscle homogenate. This was not found in the control cybrid and control muscle which only showed the complex V holoenzyme and dimer. The subcomplexes most likely are the F1-ATPase and the subcomplex V*, as denoted before (2, 21).

Figure 4 Immunoblotting after BN-PAGE shows increased amounts of subcomplexes of mt ATP synthase in the patient cybrid clones and muscle. The control cybrid and muscle only show the complex V holoenzyme and dimer. Antibodies used are against ATP synthase subunit alpha and CoII-70 kDa (complex II, loading control). Lane 1: control cybrid cell line (coCy); lanes 2-8: patient cybrid clones (1->7); lane 9: patient muscle homogenate (ptMu); lane 10: control muscle homogenate (coMu).

The in-gel activity assay of ATP hydrolysis showed activity of free F1-ATPase in the patient cybrid clones and muscle. The holoenzyme showed no activity in neither the patient nor control samples (Figure 5).

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A novel mitochondrial ATP8 gene mutation

Figure 5 The in-gel activity (IGA) assay of ATP hydrolysis shows activity of free F1-ATPase in the patient cybrid clones and muscle. Lane 1: control cybrid cell line (coCy); lanes 2-8: patient cybrid clones (1->7); lane 9: patient muscle homogenate (ptMu); lane 10: control muscle homogenate (coMu).

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DISCUSSION

Clinical phenotype

To our knowledge we describe the first pathogenic mutation in the mitochondrial ATP8 gene.

It has been stated before that the clinical phenotype of complex V deficiency due to nuclear genetic defects (hypertrophic cardiomyopathy, hypotonia, facial dysmorphism and microcephaly) markedly differs from the severe central system changes observed in NARP or MILS (1). The clinical phenotype of our patient is characterized by signs and symptoms that have mainly been described both in mitochondrial (neuropathy, ataxia) and nuclear

(hypertrophic cardiomyopathy) complex V defects. Hypertrophic cardiomyopathy can manifest with negligible to extreme hypertrophy, minimal to extensive fibrosis and myocyte disarray, absent to severe left ventricular outflow tract obstruction, and distinct septal contours/morphologies with extremely varying clinical course (22). The apical variant- hypertrophic cardiomyopathy reported here represents a genotype-phenotype relationship that has not been described before. Taken together, this indicates that the clinical variability among the patient group carrying mitochondrial complex V mutations is greater than previously assumed.

Pathogenicity

Our results strongly suggest that the MT-ATP8 mutation is the cause of the complex V deficiency in our patient. Several lines of evidence point to this. (I) The mutation leads to a premature stopcodon situated in the C-teminal region of subunit 8. (II) The C-terminal domain of mt ATP synthase subunit 8 is a conserved region in yeast (20) and in mammals

(www.mitomap.org) (Figure 2) (III) Hitherto no polymorphisms have been described at the position of the mutation (23). (IV) The mutation is homoplasmic (>90%) in muscle tissue and fibroblasts and in both tissues the complex V activity is clearly decreased (Table 1). (V)

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A novel mitochondrial ATP8 gene mutation

Transmitochondrial cybrids retained the complex V deficiency, confirming the mitochondrial origin of the pathogenic mutation.

Molecular pathophysiology

Immunoblotting after BN-PAGE showed a lack of holocomplex V and increased amounts of subcomplexes and the in-gel activity assay revealed an isolated activity of F1-ATPase in the cybrids and in muscle. The results of the in-gel activity assay of ATP hydrolysis probably point to the free rotation capacity of the F1 subcomplex in the patient. It was reported that F1 can be detected as a single entity (2) and that it retains ATP hydrolysis activity (21). The higher in-gel ATP hydrolysis activity of F1-ATPase has been described before (21). It was suggested that subunits of the Fo portion of ATP synthase restrict in-gel ATP hydrolysis (21), which results in an apparently low activity of the holocomplex in this assay. Whereas the in- gel activity assay showed an isolated F1-ATPase activity, the spectrophotometric assay demonstrated a very low FoF1-ATPase activity. The enzymatic assay measures the oligomycin-sensitive activity of complex V (8). This is the result of the subtraction of the mtATPase holoenzyme activity and the activity after addition of oligomycin, an inhibitor of proton translocation in mtATPase (24). Although the ATPase activity in the absence of oligomycin was in the same order of magnitude in our patient as in controls, the activity in the patient was not sensitive to oligomycin inhibition, whereas in control cells, usually a 90% inhibition is seen. In fact, detachment of the F1 from the Fo subcomplex – as seen on the BN gel data of the patient cybrid clones and muscle homogenate- leads to a decrease in oligomycin-sensitive complex V activity. ATP hydrolysis by F1 takes place, but without the coupling of ATP hydrolysis to extrusion of protons through Fo, the sensitivity to oligomycin is no longer evident (24).

How can the increased amounts of subcomplexes of complex V be explained? It has been described that subunit 8, which is unique to mtATPase of fungi and mammals, consists of three functional domains: the N-terminus, a central transmembrane hydrophobic domain and

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Chapter 5 the C-terminus localized to the mitochondrial matrix side (25-26). In yeast, the positively charged C-terminal region has been shown to be required for assembly of yeast subunit 8

(Y8) into the Fo-sector (27). Since Y8 interacts with other subunits in the assembly of the Fo- sector – subunit 6 assembly into mtATPase requires the presence of assembled Y8 in the enzyme complex (28)- it could be postulated that mutations in the C-terminal region – as present in our patient: p.Trp55X, subunit 8 having 68 residues- are associated with a decreased complex V assembly, which appears to be similar to the effect of human MT-

ATP6 mutations (21). Our immunoblotting results confirm this hypothesis.

The impaired complex V assembly very likely has a repercussion on the function of the holoenzyme, as was demonstrated in our patient by enzymatic analysis. Impaired subunit 6 assembly into the Fo-sector possibly plays a role. It has been described that subunit 6 has two functions: first, it leads the protons through a hydrophilic half-channel into the matrix, and secondly, it turns the rotor for ATP synthesis (29). The MT-ATP6 m.8993T>G mutation interferes with the rotation of the ring of c subunits, and therefore it uncouples ADP phosphorylation in F1 from proton transfer through Fo (29). The assembly problem is most likely not the sole pathogenic factor. It is also likely that the subunit 8 protein is decreased due to the nature of the mutation in our patient, namely a premature stopcodon, most likely resulting in a truncated protein. In that case a direct negative effect on the function of the enzyme could be assumed. It has indeed been described in yeast that Y8 also plays an important role in determining the mtATPase activity (25). In particular, the hydrophobic nature of amino acids in the center of the transmembrane domain of Y8 is essential for coupling proton transport through Fo to ATP synthesis on F1 (25). Thus, Y8 may take part in conformational changes that occur between the Fo- and F1-sectors of the enzyme during catalysis (25). However, further investigation is warranted in order to elucidate this latter issue.

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Acknowledgments

AIJ and JS are supported by a grant from the Prinses Beatrix Fonds.

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11. Janssen AJ, Trijbels FJ, Sengers RC, Wintjes LT, Ruitenbeek W, Smeitink JA, et al. Measurement of the energy-generating capacity of human muscle mitochondria: diagnostic procedure and application to human pathology. Clin Chem. 2006 May;52(5):860-71. 12. Cooperstein SJ, Lazarow A. A microspectrophotometric method for the determination of cytochrome oxidase. J Biol Chem. 1951 Apr;189(2):665-70. 13. Srere PA. Citrate synthase 1969. 14. Smeitink J, Sengers R, Trijbels F, van den Heuvel L. Human NADH:ubiquinone oxidoreductase. J Bioenerg Biomembr. 2001 Jun;33(3):259-66. 15. Janssen AJ, Smeitink JA, van den Heuvel LP. Some practical aspects of providing a diagnostic service for respiratory chain defects. Ann Clin Biochem. 2003 Jan;40(Pt 1):3-8. 16. Lowik MM, Hol FA, Steenbergen EJ, Wetzels JF, van den Heuvel LP. Mitochondrial tRNALeu(UUR) mutation in a patient with steroid-resistant nephrotic syndrome and focal segmental glomerulosclerosis. Nephrol Dial Transplant. 2005 Feb;20(2):336-41. 17. Schagger H, von Jagow G. Blue native electrophoresis for isolation of membrane protein complexes in enzymatically active form. Anal Biochem. 1991 Dec;199(2):223-31. 18. Nijtmans LG, Henderson NS, Holt IJ. Blue Native electrophoresis to study mitochondrial and other protein complexes. Methods. 2002 Apr;26(4):327-34. 19. Smith PK, Krohn RI, Hermanson GT, Mallia AK, Gartner FH, Provenzano MD, et al. Measurement of protein using bicinchoninic acid. Anal Biochem. 1985 Oct;150(1):76-85. 20. Straffon AF, Nagley P, Devenish RJ. Rescue of yeast defective in mitochondrial ATP synthase subunit 8 by a heterologous gene from Aspergillus nidulans. Biochem Biophys Res Commun. 1994 Sep 30;203(3):1567-73. 21. Nijtmans LG, Henderson NS, Attardi G, Holt IJ. Impaired ATP synthase assembly associated with a mutation in the human ATP synthase subunit 6 gene. J Biol Chem. 2001 Mar 2;276(9):6755-62. 22. Bos JM, Ommen SR, Ackerman MJ. Genetics of hypertrophic cardiomyopathy: one, two, or more diseases? Curr Opin Cardiol. 2007 May;22(3):193-9. 23. Ingman M, Gyllensten U. mtDB: Human Mitochondrial Genome Database, a resource for population genetics and medical sciences. Nucleic Acids Res. 2006 Jan 1;34(Database issue):D749-51. 24. Devenish RJ, Prescott M, Boyle GM, Nagley P. The oligomycin axis of mitochondrial ATP synthase: OSCP and the proton channel. J Bioenerg Biomembr. 2000 Oct;32(5):507-15. 25. Devenish RJ, Prescott M, Roucou X, Nagley P. Insights into ATP synthase assembly and function through the molecular genetic manipulation of subunits of the yeast mitochondrial enzyme complex. Biochim Biophys Acta. 2000 May 31;1458(2-3):428-42.

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26. Belogrudov GI, Tomich JM, Hatefi Y. Membrane topography and near-neighbor relationships of the mitochondrial ATP synthase subunits e, f, and g. J Biol Chem. 1996 Aug 23;271(34):20340-5. 27. Papakonstantinou T, Galanis M, Nagley P, Devenish RJ. Each of three positively- charged amino acids in the C-terminal region of yeast mitochondrial ATP synthase subunit 8 is required for assembly. Biochim Biophys Acta. 1993 Aug 16;1144(1):22-32. 28. Hadikusumo RG, Meltzer S, Choo WM, Jean-Francois MJ, Linnane AW, Marzuki S. The definition of mitochondrial H+ ATPase assembly defects in mit- mutants of Saccharomyces cerevisiae with a monoclonal antibody to the enzyme complex as an assembly probe. Biochim Biophys Acta. 1988 Mar 30;933(1):212-22. 29. Sgarbi G, Baracca A, Lenaz G, Valentino LM, Carelli V, Solaini G. Inefficient coupling between proton transport and ATP synthesis may be the pathogenic mechanism for NARP and Leigh syndrome resulting from the T8993G mutation in mtDNA. Biochem J. 2006 May 1;395(3):493-500.

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Restoration of complex V deficiency caused by a

novel deletion in the human TMEM70 gene

normalizes mitochondrial morphology

An I. Jonckheere, Merei Huigsloot, Martin Lammens, Jitske Jansen, Lambert P. van den

Heuvel, Ute Spiekerkoetter, Jürgen-Christoph von Kleist-Retzow, Marleen Forkink, Werner

J.H. Koopman, Radek Szklarczyk, Martijn A. Huynen, Jack A. Fransen, Jan A.M. Smeitink,

Richard J.T. Rodenburg

Mitochondrion 2011; 11(6):954-63

Chapter 6

ABSTRACT

We report a fragmented mitochondrial network and swollen and irregularly shaped mitochondria with partial to complete loss of the cristae in fibroblasts of a patient with a novel

TMEM70 gene deletion, which could be completely restored by complementation of the

TMEM70 genetic defect. Comparative genomics analysis predicted the topology of TMEM70 in the inner mitochondrial membrane, which could be confirmed by immunogold labeling experiments, and showed that the TMEM70 gene is not restricted to higher multi-cellular eukaryotes. This study demonstrates that the role of complex V in mitochondrial cristae morphology applies to human mitochondrial disease pathology.

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INTRODUCTION

Complex V (or the mitochondrial (mt) ATP synthase, EC 3.6.3.14), uses the proton gradient to generate ATP during oxidative phosphorylation. Complex V is a multisubunit complex consisting of two functional domains, F1 and Fo, connected by two stalks. The catalytic F1 domain is composed of subunits α, β, γ, δ, ε and a loosely attached IF1 inhibitor protein. The membrane-embedded Fo domain consists of an additional ten subunits, a (or subunit 6), b, c, d, e, f, g, OSCP, A6L (or subunit 8), and F6 and functions as a proton channel (1-2).

Subunits 6 and 8 of the Fo domain are encoded by the mtDNA genes MT-ATP6 and MT-

ATP8, respectively. All other complex V subunits are encoded by the nuclear DNA.

Mitochondria are enclosed by a double membrane. The inner membrane is composed of two subdomains: the inner boundary membrane and the cristae membrane. Cristae are invaginations of the inner membrane that are connected to the inner boundary membrane by narrow tubular structures, so-called crista junctions (3). It has been shown that complex V of mammalian mitochondria is arranged in long rows of dimeric supercomplexes (4). Ribbons of complex V dimers are common to all eukaryotes (4). The role of complex V dimers in the formation of tubular cristae has been hypothesized by Allen (5). The link between the dimerization of mitochondrial complex V, through subunits e, g, and –as described just recently- subunits i, k, and the biogenesis of cristae has been provided by studying yeast cells (6-7). Complex V oligomers are found either on the crest of lamellae, or along the length of tubular cristae and introduce a positive curvature to the inner mitochondrial membrane (4).

In a recent yeast study, this effect has been shown to be attributed to the action of subunit e and subunit g (8). Moreover, it has been suggested that the formation of cristae and crista junctions in mitochondria depends on antagonism between Fcj1 (formation of crista junction protein 1) and subunits e and g (8). In HeLa cells, it has been shown that IF1 overexpression increases mitochondrial cristae formation and dimerization of complex V (9). Other components such as prohibitins or OPA1 or others yet to be identified also could contribute to crista junction and cristae tip formation (8, 10). Complex V mutations have been described

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in MT-ATP6 (see, amongst many others, (11)), in MT-ATP8 (12), in the nuclear encoded F1- specific assembly gene ATP12 (13), and in the nuclear encoded structural subunit ε (ATP5E)

(14). A common splice site mutation and an isolated frameshift mutation were described in the TMEM70 gene particularly in a homogeneous ethnic group (Romanies), and it has been shown that TMEM70 is required to maintain normal expression levels of complex V (15). The

TMEM70 gene, located on chromosome 8q21.11, consists of three exons and encodes a

260 amino acid (AA) protein, transmembrane protein 70. GFP-tagged TMEM70 has been shown to be localized in mitochondria (16). The exact molecular function of the TMEM70 protein is not known at present (2), although it has been suggested that TMEM70 is involved in complex V biogenesis (15). TMEM70 homologues have been found in genomes of multi- cellular eukaryotes and plants, while it has been described to be absent from yeast and other fungi (2).

CASE HISTORY

The boy, born at 40 weeks, was the third child of healthy consanguineous Iraqi parents with unremarkable family history. Two older siblings are healthy. Prenatal ultrasound revealed fetal ascites and oligohydramnion. He was small for gestational age (birth weight 2090 g, length 47 cm, head circumference 33 cm). Postnatal echocardiography was normal. He presented on day four of life with feeding difficulties and fever. Lactic acidosis was diagnosed

(base excess -14.5 mmol/l) with a blood lactate of 13.9 mmol/l (normal < 2). Also hyperammonemia (557 µg/dl, normal < 45) occurred. Plasma amino acid analysis revealed elevation of ornithine (651 µmol/l, normal 0-235) and alanine (971 µmol/l, normal < 450). In urine, excretion of homocitrulline was observed, together suggesting presence of hyperammonemia-hyperornithinemia-homocitrullinuria (HHH)- syndrome. After supplementation of sodium-benzoate and L-arginine and protein restriction, the clinical condition stabilized and hyperammonemia resolved. However, lactic acidosis and

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TMEM70 gene deletion hyperalaninemia persisted, suggesting mitochondrial impairment. During follow-up lactate was between 3.5 and 7.8 mmol/l.

In the first months of life, hypertrophic cardiomyopathy developed. During follow-up motor and especially speech development was impaired. Cerebral magnetic resonance spectroscopy revealed an increased lactate peak within the basal ganglia and the white matter region as well as an increased choline peak.

All symptoms were initially interpreted as either a primary mitochondrial disease or HHH- syndrome with secondary mitochondrial impairment. After HHH –syndrome was excluded by molecular analysis, skin and muscle biopsies were performed in order to investigate a primary mitochondrial enzyme defect. Currently, at the age of 6 years, the cardiomyopathy has remained stable under treatment with calcium antagonists. The child’s development is progressive but he retains a distinctive motor and mental retardation. He has an ataxic gait and ptosis.

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MATERIALS AND METHODS

Cell cultures

Fibroblasts were cultured in medium 199 (Gibco®, Invitrogen Corporation) supplemented with 10% fetal calf serum and penicillin/streptomycin (respectively 100 U/ml and 100 µg/ml).

The amphotropic packaging cell line PA317 (# CRL-9078, LGC, Middlesex, UK) and Flp-In T-

REx293 cells (Invitrogen) were grown in Dulbecco's Modified Eagle's Medium (DMEM) containing 4.5 g/L glucose, 10% fetal calf serum and penicillin/streptomycin (respectively 100

U/ml and 100 µg/ml). Inducible cell lines were selected on 5 μg/ml blasticidin (Invitrogen) and

200 μg/ml hygromycin (Calbiochem), and expression of the transgene was induced by addition of 1 μg/ml doxycycline (Sigma Aldrich) for 24 hours.

Biochemical assays

Measurement of the mitochondrial energy-generating system (MEGS) capacity was performed as described before (17). The complex V (or mtATPase) activity was measured spectrophotometrically in mitochondria isolated from frozen pellets of the patient and control fibroblasts as described (11). The activities of the mitochondrial respiratory chain enzymes and citrate synthase (CS) were measured spectrophotometrically in muscle tissue and fibroblasts of the patient according to described protocols (18-19).

Molecular genetic analysis

Total DNA from blood and cultured fibroblasts was extracted using the salting out procedure for human DNA extraction (20). The coding region in genomic DNA of the TMEM70 gene was polymerase chain reaction (PCR) amplified using three primer pairs (Table 1a). PCR conditions were 95°C for 5 minutes (denaturation), 92°C for 45 seconds, 60°C for 45

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TMEM70 gene deletion seconds and 72°C for 45 seconds for a total of 35 cycles. Major deletions in genomic DNA were screened for by long template PCR (Expand Long Template PCR system, Roche,

Germany) using new primers (Table 1a). PCR conditions were 95°C for three minutes

(denaturation), 93°C for 10 seconds, 60°C for 30 seconds, 68°C for seven minutes, for a total of 9 cycles followed by 93°C for 10 seconds, 60°C for 30 seconds, 68°C for seven minutes, plus 20 seconds each cycle for a total of 24 cycles. Sequence analysis of PCR amplified products was performed on an ABI3730 automatic capillary sequencer using BigDye® terminator chemistry (Applied Biosystems, Nieuwerkerk a/d Ijssel, The Netherlands). RNA from the patient cultured fibroblasts was extracted as described (20). Reverse Transcriptase

PCR (RT-PCR) was performed to produce cDNA followed by PCR amplification (Table 1b).

Sequence analysis was subsequently performed as described above.

Table 1a: Primers for PCR amplification of the genomic DNA

Exon(s) Forward primer Reverse primer Comment

1 5'-GGCATGCGCCACTTGTGCG-3' 5'-GCTGGGGCTCCCCAGGCC-3' Conventional

PCR

2 5'-AGGTTAGTTGACCATAATGATCCCTG- 5'-ACGTCTTGTAATTAAGGGATGCCA-3' Conventional

3' PCR

3a* 5'-GCAGATTTCCTGCCTGGAGAGG-3' 5'-TCTGGAATCTTCACATCATTCTGGTG-3' Conventional

PCR

3b* 5'-TGATTGGCCTTACATTTCTGCCA-3' 5'-AACAATCAGTCACTAACGGAATGCAA- Conventional

3' PCR

2 5'-GGATTGGCAAACCTGGTTAAAA-3' 5'-GGCTGAGGCAGGAAAATCCTT-3' Long-template

PCR

*Exon 3 of TMEM70 is PCR amplified in two overlapping fragments (a and b).

Table 1b: Primers for PCR amplification of the cDNA

Fragment Forward primer Reverse primer

1 5'-GGCAGTCGGGTGGGAAGCC-3' 5'-ACCCGGACGCCGGAGAAGG-3'

2 5'-CCTTCTCCGGCGTCCGGGT-3' 5'-CAGTCACTAACGGAATGCAAAAG-3'

3 5'-GGTGATCACCCCAGTGCTGC-3' 5'-CAGTCACTAACGGAATGCAAAAG-3'

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Complex V assembly and activity

Blue Native Polyacrylamide Gel Electrophoresis (BN-PAGE), blotting and in-gel activity of

ATP hydrolysis was performed as described (21). Gel lanes were loaded with 40 µg of protein (22) of OXPHOS complexes isolated from a mitoplast fraction of the patient and control fibroblasts. For Western blotting, monoclonal antibodies against mitochondrial complex V subunit alpha and CoII-70 kDa (complex II) (Invitrogen, Breda, The Netherlands) were used. For the in-gel activity assay of ATP hydrolysis the gel was incubated overnight at room temperature with the following solution: 35 mM Tris, 270 mM glycine, 14 mM MgSO4,

0.2% Pb(NO3)2, and 8 mM ATP, pH 7.8 (21).

Cloning of human TMEM70 gene, generation of inducible cell lines, and retroviral complementation

Human full length TMEM70 cDNA clone IOH40588 (Imagenes, Berlin, Germany) (without stop codon) flanked by Gateway AttB sites was amplified by PCR according to the manufacturer’s protocol and cloned into pDONR201 by the use of Gateway BP Clonase II

Enzyme Mix. The pDONR201-TMEM70-vector was recombined with the AcGFP1-destination vector by use of the LR Clonase II Enzyme Mix (Invitrogen). The obtained constructs were transfected into Flp-In T-REx293 cells (Invitrogen) by using SuperFect Transfection Reagent

(QIAGEN) according to the manufacturer’s manual. The inducible COX8 leader sequence

(first 210 base-pairs of sequence NM_00004074)-GFP cell line has been described previously (23). For complementation, the TMEM70 cDNA clone IOH40588 was cloned into the GATEWAY retroviral destination vector pDS_FBneo (# MBA-295, LGC, Middlesex, UK). pDS_FBneo retroviral vector with or without TMEM70 was used to transfect the amphotropic packaging cell line PA317 using Lipofectamine 2000 according to the manufacturer’s protocol (Invitrogen, Breda, The Netherlands) (24). At 24 hours after transfection, fresh medium was added and supernatants containing retroviral particles were collected at 40

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TMEM70 gene deletion hours after transfection and filtered through a 0.45 µm filter. Fibroblasts in 75 cm2 flasks were incubated for 24 hours with a 1:1 mixture of growth medium and virus-containing medium in the presence of 4 µg/ml polybrene (24). The medium was then changed to selection medium consisting of 199 medium supplemented with 15 % fetal calf serum, penicillin/streptomycin

(respectively 100 U/ml and 100 µg/ml) and 500 µg/ml geneticin (G418, PAA, Pasching,

Austria). Cells were selected for at least 14 days. G418-resistant cells were used for biochemical analyses within 5 passages after transduction.

Immunogold labeling

Cells were fixed in 1% formaldehyde and 0,01% glutaraldehyde in 0,1M PHEM buffer for 1 h at RT. Next, cells were pelleted in 10% gelatin and post-fixed in 1% formaldehyde for another

24 h. Ultra-thin cryosections were prepared on a Leica EMFCS, incubated with a homemade polyclonal antiserum raised against GFP (25) followed by incubation with protein A complex to 10 nm gold particles according to standard procedures (26). Sections were observed in a

JEOL 1010 electron microscope.

Quantitative mitochondrial morphology and membrane potential assessment

Mitochondrial morphology and membrane potential was quantified in single living cells using tetramethyl rhodamine methyl ester (TMRM; Invitrogen, Breda, The Netherlands) staining and videomicroscopy applying a protocol described previously (27), with some adaptations.

To prevent retrovirus contaminations, cells were cultured for three days and imaged in 35 mm glass bottom dishes with a 22 mm glass aperture (WillCo Wells BV, Amsterdam, The

Netherlands). Immediately after loading and washing, the dishes were sealed using paraffin.

The TMRM loading solution was prepared using 1 µl of a 20 µM “working” stock solution in

DMSO in 1 ml of medium 199. Image acquisition was performed using a Zeiss 40x/1.3 NA

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Chapter 6 fluor objective instead of a Zeiss 63x/1.25 NA Plan NeoFluar objective. Statistical significances were assessed by a Student’s two-population, independent t-test.

Electron microscopy

Trypsinized fibroblast suspensions in culture medium were centrifuged for 10 minutes at 120 g. The medium was carefully removed and 2% glutaraldehyde buffered with 0.1 M sodium cacodylate pH 7.4 was gently added. After at least 4 hours fixation the pellets were very carefully taken of and post-fixed in 1% osmium tetroxide in Palade’s buffer pH 7.4 with 1% potassium hexacyanoferrat(III)-trihydrate and after dehydration in ethanol and propylene oxide, embedded in Epon. Semi-thin, 1µm thick transverse sections were stained with 1% toluidine blue. Ultrathin sections were stained with uranyl acetate and lead citrate and examined in a JEOL 1200. Electron microscopy was solely performed in fibroblasts because muscle tissue of the patients was no longer available.

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RESULTS

Biochemical assays

Measurement of the MEGS capacity in muscle tissue showed both a severely decreased pyruvate oxidation rate and ATP production rate from oxidation of pyruvate (Table 2).

Spectrophotometric analysis showed an undetectable complex V (mtATPase) activity in muscle tissue (0 mU/U Cytochrome Oxidase (COX), normal 169-482) and a combined deficiency of complex V (113 mU/U COX, normal 209-935) and complex I (60 mU/U COX, normal 110-260) in fibroblasts. The activities of the respiratory chain enzymes complex II, III, and IV, and citrate synthase (CS) were normal (Table 2).

Table 2: Measurement of MEGS capacity in muscle tissue and enzymatic analysis of the activities of complex V (mtATPase), the respiratory chain enzymes and CS in muscle tissue and fibroblasts.

Muscle tissue Fibroblasts

[1-14C]Pyruvate + malate * 0.68 (3.45-7.99) n.d.

ATP production rate ** 3.3 (36-81.7) n.d.

Complex V, mU/U COX 0 (169-482) 113 (209-935)

Complex I, mU/U CS (m), mU/U COX (f) 75 (70-251) 60 (110-260)

Complex II, mU/U CS (m), mU/U COX (f) 394 (335-749) 635 (536-1027)

Complex III, mU/U CS (m), mU/U COX (f) 3800 (2200-6610) 2175 (1270-2620)

Complex IV, mU/U CS 2049 (810-3120) 762 (680-1190)

CS, mU/mg 298 (37.4-162) 114 (144-257)

MEGS: mitochondrial energy-generating system; COX: cytochrome oxidase; CS: citrate synthase; n.d.: not determined ; m : muscle tissue ; f : fibroblasts. Figures in brackets are normal ranges. *Expressed as nmol 14CO2/h.mU CS **Expressed as nmol (ATP+CrP)/h.mU CS

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Molecular genetic analysis

Homozygosity mapping revealed several homozygous regions, including a 9.2 cM region on chromosome 8q13.3-21.13, that includes the TMEM70 gene (data not shown). PCR of the coding region of the TMEM70 gene showed a normal fragment for exon 1 and exon 3 but no fragment for exon 2. Fragments of known sizes for all three exons were seen in control DNA and in DNA of the parents (Figure 1a). Long template PCR from exon 1 to 3 resulted in a smaller fragment in the patient’s DNA compared to control DNA. Long template PCR on DNA samples of the parents showed two fragments, one with the same size as the control, the other with the same size as the patient (Figure 1b). Taken together, these observations suggest the presence of a homozygous deletion in the patient’s TMEM70 gene, for which the parents are heterozygous carriers. This was confirmed by sequence analysis of the long template PCR product, which showed a homozygous 1353 base pair deletion containing exon 2 (g. 2436-3789) in the TMEM70 gene of the patient (Figure 1c). To evaluate the effect of the deletion on the TMEM70 transcript, the patient’s cDNA was analyzed, revealing a smaller transcript compared to wild type TMEM70 cDNA (Figure 1d). Sequence analysis of the patient’s cDNA confirmed that exon 2 was deleted from the transcript, resulting in a mRNA consisting of exon 1 spliced to exon 3 (Figure 1e). No mutations were found in exons

1 and 3. At the protein level, this is an in-frame deletion resulting in a theoretical protein of

226 AA, missing 34 AA (AA 71-105) encoded by exon 2.

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Figure 1 Molecular genetic analysis of the TMEM70 gene in fibroblasts. (a) PCR of the coding region in genomic DNA shows no fragment for exon 2 in the patient, in contrast to the control and both parents. (F: father; M: mother; P: index patient; -: negative control; +: wild type). (b) Long template PCR in genomic DNA shows a smaller fragment in the patient compared to wild type DNA. DNA of the parents shows two fragments, which points to heterozygosity. (F: father; M: mother; P: index patient; -: negative control; +: wild type). (c) Genomic sequence analysis of the patient’s DNA reveals a homozygous 1353 base pair deletion (g. 2436-3789) containing exon 2. (d) Reverse Transcriptase PCR shows a smaller product in the patient for the fragment containing all three exons (1F+3R). Fragments of normal size are present for exon 1 (1F+1R) and for exon 3 (3F+3R). (F: forward primer, R: reverse primer, see Table 1b; P: patient; -: negative control; +: wild type). (e) cDNA sequence analysis confirms a deletion of exon 2 in the patient.

Complementation of complex V deficiency by wild type TMEM70

Immunoblotting after BN-PAGE (Figure 2a) with anti-complex V subunit alpha antibody was performed on protein extracts from fibroblasts of the patient, two controls and a patient harboring the previously described homozygous c.317-2A>G mutation (15, 28). The blot clearly revealed a nearly absent holocomplex V in the patient samples, in contrast to the

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Chapter 6 control samples. The in-gel activity assay of ATP hydrolysis showed activity in both controls, but not in the patient samples (Figure 2b).

To confirm that deletion of TMEM70 exon 2 caused the strong reduction of complex V holocomplex amount and activity, both index patient and control fibroblasts were retrovirally transduced with human full length TMEM70. Complementation of patient fibroblasts with wild type TMEM70 induced a normalization of fully assembled complex V on BN-PAGE (Figure

2c). Moreover, in-gel activity of ATP hydrolysis at the position of complex V in the gel was detectable in extracts from the TMEM70-complemented patient cell line at a similar level as in the control (Figure 2d).

Figure 2 Complementation of the TMEM70 gene defect in fibroblasts. (a) Immunoblotting after BN-PAGE shows a nearly absent holocomplex V in the fibroblasts of the patients, in contrast to the control samples. (b) The in-gel activity assay of ATP hydrolysis demonstrates activity in both controls, but not in the fibroblasts of the patients. (8835: index patient; 8836: patient with homozygous c.317-2A>G mutation in the TMEM70 gene). (c) Immunoblotting after BN-PAGE shows a normalization of the concentration of the fully assembled complex V in the complemented fibroblasts of the index patient. (d) In-gel activity assay of ATP hydrolysis demonstrates activity in the control and in the complemented index patient cell line. (Lane 1: control; lane 2: control + pDS_FBneo; lane 3: control + pDS_FBneo + TMEM70 gene; lane 4: index patient; lane 5: index patient + pDS_FBneo; lane 6: index patient + pDS_FBneo + TMEM70 gene). Antibodies used are against complex V subunit alpha and CoII-70 kDa (complex II, loading control).

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TMEM70 comparative genomics

To shed more light on the evolution and function of TMEM70 we analyzed its sequence and relevant genomics data. Profile based homology detection using PSI-Blast (5 iterations, E <

0.001) shows that orthologs of TMEM70 are not only present in multicelullar species and in the closest single cell relative of the metazoa, the choanoflagellate Monosiga brevicollis (15).

The gene is also present in fungi (Schizosaccharomyces pombe), plants (Arabidopsis thaliana) and even stramenopiles (Phytophthora infestans) (Figure 3a). This indicates that

TMEM70 was already present in the ancestor of all eukaryotes. Consistent with its function in respiration, TMEM70 cannot be found in obligatory anaerobic species. However, it does not specifically co-occur with complex V, as it is also absent from many species that do have complex V, like Saccharomyces cerevisiae. Most genomics data on TMEM70 are consistent with a role in OXPHOS. Orthologs of TMEM70 have been identified in the proteomic analysis of Caenorhabditis elegans mitochondria (29) as well as in the fluorescent-tag study of

Schizosaccharomyces pombe - organelles (30). Furthermore, large-scale co-expression studies of mammalian cells (31-33) show that three nuclear encoded subunits of complex V genes (ATP5J, ATP5C1, ATP5A1) are among the top 10 genes that co-express with

TMEM70.

TMEM70 has been predicted to have two transmembrane regions located in the well- conserved part of the protein (15). Based on a multiple sequence alignment and a consensus-based tool for the prediction of transmembrane topology (MetaTM, (34)), the N- and the C-terminus are predicted to be located in the mitochondrial matrix, with the region between the two transmembrane regions located in the inter-membrane space (Figure 3b).

The deletion observed in the patient overlaps with the predicted transmembrane region

(Figure 3a).

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Figure 3 TMEM70 comparative genomics. (a) Alignment of the human TMEM70 protein with orthologs from eukaryotic crown groups (animals, fungi, plants, chromalveolates) suggests the early origin of the protein. Predicted mitochondrial targeting signal (MTS), transmembrane regions (TM), as well as the exon deleted in the index patient (DEL) are marked. (b) Likely topology of TMEM70 in the inner mitochondrial membrane. Transmembrane topology was predicted with MetaTM (34).

Immunogold labeling

Immunogold labeling of Flp-In T-Rex293-TMEM70-GFP cells after incubation with an antibody against GFP demonstrated that TMEM70 is localized in mitochondria predominantly in the cristae membrane (Figure 4a+b). As a control, the localization of the COX8 leader sequence attached to GFP, a known matrix-soluble protein (23), was found predominantly in the mitochondrial matrix (Figure 4c).

Figure 4 Immunogold labeling. (a+b) Immunogold labeling of Flp- In T-REx293-TMEM70-GFP cells after incubation with antibody against GFP shows that TMEM70 is localized in mitochondria predominantly in the cristae membrane and is absent from the mitochondrial matrix (asterix). (c) In contrast, immunogold staining of the COX8 leader sequence attached to GFP, a known matrix- soluble protein, shows that this construct is present in the mitochondrial matrix (asterix). (d) The negative control, Flp-In T- REx293 cells without GFP-tag, shows no label. Scale bar is 0.5 μm.

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Live cell mitochondrial morphology and membrane potential

The mitochondrial network is fragmented in TMEM70 patient fibroblasts (Figure 5a). This was demonstrated by quantification of mitochondrial morphology, revealing that TMEM70 patient fibroblasts contained mitochondria with a reduced aspect ratio ('AR', Figure 5b) and a reduced length and degree of branching ('F', Figure 5c). This was paralleled by an increased number of mitochondria per cell ('Nc', Figure 5d). The mitochondrial network parameters were reversed after complementation with wild type TMEM70 (Figure 5b,c,d). In addition, it was demonstrated that the mitochondrial membrane potential in patient cells is increased.

This was also reversed by complementation (Figure 5e), which is in agreement with previously published observations (15).

For the parameters TMRM intensity and mitochondrial form factor, the relatively strong effect of complementation of patient cells with wild type TMEM70 was probably due to the relatively high levels of expression caused by the retroviral promoter that drives expression of

TMEM70 in the complemented cells. In summary, these observations show that TMEM70 affects mitochondrial network parameters, indicating a role of TMEM70 in mitochondrial morphology.

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Figure 5 Live cell mitochondrial morphology and membrane potential (a) The mitochondrial network is fragmented in TMEM70 patient fibroblasts (8835: index patient; 8836: patient with homozygous c.317-2A>G mutation in the TMEM70 gene). (b+c+d) Quantitative mitochondrial morphology assessment. (b) Mitochondrial aspect ratio (AR), being the ratio between mitochondrial length and width, in control cells, patient cells (8835, 8836) and complemented patient cells (8835 + empty: index patient + pDS_FBneo, 8835 + TMEM70: index patient + pDS_FBneo + TMEM70 gene). (c) Mitochondrial form factor (F), being a combined measure of mitochondrial length and degree of branching, in control cells, patient cells (8835, 8836) and complemented patient cells (8835 + empty, 8835 + TMEM70). (d) The number of mitochondria per cell in control cells, patient cells (8835, 8836) and complemented patient cells (8835 + empty, 8835 + TMEM70). (e) Mitochondrial membrane potential, expressed by TMRM intensity, in control cells, patient cells (8835, 8836) and complemented patient cells (8835 + empty, 8835 + TMEM70). All data were expressed as % of the average control value (i.e. that measured the control in cell line) measured on the same day. Numerals indicate the number of cells (panel B) analyzed in four independent experiments. Statistics: ***(p<0.001), **(p<0.01) and *(p<0.05) relative to control.

Electron microscopy

Mitochondria in fibroblasts of the index case (Figure 6a) and in cases with complex V deficiency due to TMEM70 c.317-2A>G (Figure 6b) and MT-ATP8 8529G>A (Figure 6c)

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TMEM70 gene deletion mutations were consistently swollen and irregularly shaped with disruption of the central part to complete loss of the cristae. Fibroblasts with a MT-ATP6 8993T>G mutation predominantly showed normal cristae (Figure 6d). The mitochondrial morphology of the fibroblasts with the TMEM70 defect normalized after complementation with wild type

TMEM70 (Figure 6f).

Figure 6 Electron microscopy in fibroblasts of patients with complex V deficiency compared to control cells and complemented cells. Mitochondria in fibroblasts of the index case (a, scale bar: 0.5 µm) and in cases with complex V deficiency due to TMEM70 c.317-2A>G (b, scale bar:

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0.2 µm) and MT-ATP8 8529G>A (c, scale bar: 0.2 µm) mutations are swollen and irregularly shaped with disruption of the central part to complete loss of the cristae. Fibroblasts with a MT-ATP6 8993T>G mutation predominantly show normal cristae (d, scale bar: 0.5 µm). Control fibroblasts show normal shaped mitochondria (e, scale bar: 0.5 µm). Fibroblasts of the index patient complemented with wild type TMEM70 show normal mitochondria (f, scale bar: 0.5 μm). (m: mitochondrion; v: vacuole; n: nucleus).

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DISCUSSION

Clinical phenotype

The patient described in this report shares common clinical features with the Gypsy patients harboring the common splice site mutation c.317-2A>G in the TMEM70 gene described before (15). He presented during the first week of life with lactic acidosis and hyperammonemia. Later, he developed non-progressive hypertrophic cardiomyopathy and psychomotor delay. The description of four novel mutations in the TMEM70 gene has confirmed and expanded this classical clinical picture with early onset cataract, gastrointestinal dysfunction, congenital hypertonia and a fetal presentation of the syndrome

(35). A milder clinical course has been found in a patient who was compound heterozygous for the c.317-2A>G and a c.494G>A mutation (36). These clinical features differ from those in a patient with a mutation in the nuclear encoded complex V assembly gene ATP12, presenting with a severe encephalopathy and basal ganglia involvement (13). Patients with mtDNA mutations in MT-ATP6 usually show central nervous system involvement, presenting with neuropathy, ataxia, and retinitis pigmentosa (NARP), maternally inherited Leigh syndrome (MILS), or bilateral striatal necrosis (2). Apical hypertrophic cardiomyopathy and neuropathy has been found in a case harboring a MT-ATP8 mutation (12). Recently, the first case of complex V deficiency due to a mutation in the nuclear encoded structural subunit ε has been described (14). This 22 year old patient presented with neonatal onset, lactic acidosis, 3-methylglutaconic aciduria, mild mental retardation and developed peripheral neuropathy (14). Taken together, the clinical picture described here resembles that of previously described Gypsy patients with a defect in the TMEM70 gene.

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Pathogenicity

In addition to the fact that the clinical symptoms resemble those of the previously described

Romany patients with TMEM70 gene mutations (15), several additional lines of evidence point to the pathogenicity of the novel TMEM70 deletion described here. Both parents are heterozygous carriers of the deletion. The homozygous deletion in the patient leads to loss of exon 2 from the full length transcript which includes the first 5 amino acids of a conserved

DUF1301 domain and the first 3 amino acids of the putative transmembrane region 1

(www.ensembl.org). This results in a severely decreased amount and activity of holocomplex

V. Complementation of the TMEM70 gene defect normalized the amount of complex V, the in-gel activity of ATP hydrolysis and the mitochondrial morphology in the patient fibroblasts.

Immunogold labeling confirms TMEM70 localization to the inner mitochondrial membrane

The exact localization of TMEM70 within mitochondria by immunogold labeling experiments has not been described before. The results confirm both the prediction that was based on the comparative genomics data presented in this report, and the recently described observations in submitochondrial fractions (37). The apparent localization of TMEM70 in the mitochondrial cristae membrane corresponds to the location of complex V and supports the involvement of

TMEM70 in complex V biogenesis, as has been proposed before (38).

Mitochondrial morphology

The role of complex V in mitochondrial cristae formation has previously been studied in yeast

(6, 8, 39-42) and in animal tissue (4). In yeast, it has been shown that in the absence of dimerization of complex V, the organization of the tubular cristae is abolished and the inner membrane is produced in an uncontrolled fashion, resulting in onion-shaped structures (6).

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The importance of mitochondrial cristae morphology has been investigated in beef heart and rat liver (4). Ribbons of complex V dimers are responsible for cristae formation, and are essential for optimal performance of complex V (4). Although mitochondrial structural abnormalities have been shown to occur in complex V deficiencies before, a direct causal relationship has never been demonstrated. In this study, we show that the mitochondrial abnormalities that are present in fibroblasts with a TMEM70 genetic defect causing complex

V deficiency, can be completely restored by complementation with wild type TMEM70.

Electron microscopy observations showed similar mitochondrial structural abnormalities in fibroblasts of patients with complex V deficiency caused by different genetic defects.

Fibroblasts carrying a TMEM70 defect contain swollen and irregularly shaped mitochondria with disruption of the central part to complete loss of the cristae. This was also seen in fibroblasts of patients with complex V deficiency harboring the TMEM70 c.317-2A>G and

MT-ATP8 8529G>A mutations, respectively. By contrast, fibroblasts of a patient with a MT-

ATP6 8993T>G mutation predominantly showed normal cristae. Previous findings in yeast have shown mitochondria with an absence of cristae in mutants in which F1 synthesis had been virtually abolished (6, 41). It has also been described in yeast that the lack of Atp6p protein, rather than an ATP synthesis deficit, modifies the overall structure of mitochondria

(43). In addition, it has been shown that the MT-ATP6 8993T>G mutation does not impair

F1Fo assembly and oligomerization in human fibroblasts (44). The MT-ATP8 8529G>A mutation gives rise to an improper assembly of the complex V holoenzyme (12). It has also been assumed that subunits e/g and a/A6L have dimer/oligomer stabilizing effects that are additive (45). Mutations in the TMEM70 gene lead to a nearly absent holocomplex V. Taken together, it is the complex V structure rather than its enzymatic activity that seems to modify cristae morphology. This is in agreement with the necessity of complex V dimerization for mitochondrial cristae formation (4, 6, 9). It could be argued that the role of complex V in cristae morphology is at odds with the high mitochondrial cristae density in brown adipose tissue (BAT), which has relatively low amounts of complex V protein (46-48). However, the structure of complex V in BAT is normal (49-50), which is a prerequisite for dimerization. We

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Chapter 6 postulate that these complex V dimers, in spite of the relatively low levels, are sufficient to lead to a high cristae density in BAT, in particular in the presence of mitogenic stimuli, such as cold exposure or norepinephrine (47, 51). Quantitative mitochondrial morphology assessment in fibroblasts of the TMEM70-deletion patient and of the patient harboring the

TMEM70 c.317-2A>G mutation revealed a disruption of the mitochondrial network. Although mitochondrial enzyme deficiencies are not systematically associated with fragmentation of the mitochondrial network (52-53), these data show that TMEM70 gene mutations affecting complex V structure result in mitochondrial network fragmentation. Complex V structure, mitochondrial morphology and the mitochondrial network were restored by complementation of the fibroblasts of the index patient with wild type TMEM70, which proves the causal relationship between TMEM70, human complex V integrity, and mitochondrial morphology, as has been recently suggested (38). This demonstrates that the previously established role of complex V in mitochondrial cristae morphology is highly relevant to human complex V pathology. The role of complex V in mitochondrial cristae biogenesis differs from other complexes of the respiratory chain, where a mutation leads to secondary loss of cristae due to degeneration of the mitochondria (54-55).

Implications

Hitherto, in most genotyped cases a substitution (c.317-2A>G) has been found (2, 15, 36). In addition, four novel TMEM70 mutations were described, confirming the distinct clinical phenotype caused by mutations in this gene (35). This report describes the first pathogenic deletion in the TMEM70 gene, further extending the mutation spectrum of this gene and confirming the classical clinical phenotype and pan-ethnicity of the disorder. Comparative genomics data point to the functional importance of conserved transmembrane regions of

TMEM70 in the evolution of eukaryotes. The predicted localization in the inner mitochondrial membrane corresponds to the immunogold labeling data. The complementation study demonstrates a causal relationship between TMEM70, complex V structure and

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TMEM70 gene deletion mitochondrial morphology in human fibroblasts. This proves that the role of complex V in mitochondrial cristae morphology that has been previously studied in yeast and in animal tissues applies to human complex V pathology.

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Funding

This work was supported by a grant from the Prinses Beatrix Fonds [grant number OP-05-04] to AIJ and JS, grant IGE05003 from Senternovem to MH, a grant from the IGMD (Institute for

Genetic and Metabolic Disease) of the Radboud University Nijmegen Medical Centre

(RUNMC) to WJHK, an equipment grant of NWO (Netherlands Organization for Scientific

Research, number: 911-02-008) and a grant from the Netherlands Genomics Initiative

(Horizon Program) to RS. The authors confirm independence from the sponsors; the content of the article has not been influenced by the sponsors.

Acknowledgements

The authors thank Ms L. Eshuis for technical assistance with the electron microscopy of the fibroblasts and Ms. M. Wijers-Rouw for technical assistance with the immunogold labeling study.

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35. Spiegel R, Khayat M, Shalev SA, Horovitz Y, Mandel H, Hershkovitz E, et al. TMEM70 mutations are a common cause of nuclear encoded ATP synthase assembly defect: further delineation of a new syndrome. J Med Genet. 2011 Mar;48(3):177-82. 36. Shchelochkov OA, Li FY, Wang J, Zhan H, Towbin JA, Jefferies JL, et al. Milder clinical course of Type IV 3-methylglutaconic aciduria due to a novel mutation in TMEM70. Mol Genet Metab. 2010 Oct-Nov;101(2-3):282-5. 37. Hejzlarova K, Tesarova M, Vrbacka-Cizkova A, Vrbacky M, Hartmannova H, Kaplanova V, et al. Expression and processing of the TMEM70 protein. Biochim Biophys Acta. 2011 Jan;1807(1):144-9. 38. Cameron JM, Levandovskiy V, Mackay N, Ackerley C, Chitayat D, Raiman J, et al. Complex V TMEM70 deficiency results in mitochondrial nucleoid disorganization. Mitochondrion. 2011 Jan;11(1):191-9. 39. Arselin G, Vaillier J, Salin B, Schaeffer J, Giraud MF, Dautant A, et al. The modulation in subunits e and g amounts of yeast ATP synthase modifies mitochondrial cristae morphology. J Biol Chem. 2004 Sep 24;279(39):40392-9. 40. Gavin PD, Prescott M, Luff SE, Devenish RJ. Cross-linking ATP synthase complexes in vivo eliminates mitochondrial cristae. J Cell Sci. 2004 May 1;117(Pt 11):2333-43. 41. Lefebvre-Legendre L, Salin B, Schaeffer J, Brethes D, Dautant A, Ackerman SH, et al. Failure to assemble the alpha 3 beta 3 subcomplex of the ATP synthase leads to accumulation of the alpha and beta subunits within inclusion bodies and the loss of mitochondrial cristae in Saccharomyces cerevisiae. J Biol Chem. 2005 May 6;280(18):18386-92. 42. Thomas D, Bron P, Weimann T, Dautant A, Giraud MF, Paumard P, et al. Supramolecular organization of the yeast F1Fo-ATP synthase. Biol Cell. 2008 Oct;100(10):591-601. 43. Kucharczyk R, Salin B, di Rago JP. Introducing the human Leigh syndrome mutation T9176G into Saccharomyces cerevisiae mitochondrial DNA leads to severe defects in the incorporation of Atp6p into the ATP synthase and in the mitochondrial morphology. Hum Mol Genet. 2009 Aug 1;18(15):2889-98. 44. Cortes-Hernandez P, Vazquez-Memije ME, Garcia JJ. ATP6 homoplasmic mutations inhibit and destabilize the human F1F0-ATP synthase without preventing enzyme assembly and oligomerization. J Biol Chem. 2007 Jan 12;282(2):1051-8. 45. Wittig I, Meyer B, Heide H, Steger M, Bleier L, Wumaier Z, et al. Assembly and oligomerization of human ATP synthase lacking mitochondrial subunits a and A6L. Biochim Biophys Acta. 2010 Jun-Jul;1797(6-7):1004-11.

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46. Houstek J, Tvrdik P, Pavelka S, Baudysova M. Low content of mitochondrial ATPase in brown adipose tissue is the result of post-transcriptional regulation. FEBS Lett. 1991 Dec 9;294(3):191-4. 47. Rodriguez-Cuenca S, Pujol E, Justo R, Frontera M, Oliver J, Gianotti M, et al. Sex- dependent thermogenesis, differences in mitochondrial morphology and function, and adrenergic response in brown adipose tissue. J Biol Chem. 2002 Nov 8;277(45):42958-63. 48. Loncar D. Convertible adipose tissue in mice. Cell Tissue Res. 1991 Oct;266(1):149- 61. 49. Houstek J, Andersson U, Tvrdik P, Nedergaard J, Cannon B. The expression of subunit c correlates with and thus may limit the biosynthesis of the mitochondrial F0F1- ATPase in brown adipose tissue. J Biol Chem. 1995 Mar 31;270(13):7689-94. 50. Kramarova TV, Shabalina IG, Andersson U, Westerberg R, Carlberg I, Houstek J, et al. Mitochondrial ATP synthase levels in brown adipose tissue are governed by the c-Fo subunit P1 isoform. FASEB J. 2008 Jan;22(1):55-63. 51. Bronnikov G, Houstek J, Nedergaard J. Beta-adrenergic, cAMP-mediated stimulation of proliferation of brown fat cells in primary culture. Mediation via beta 1 but not via beta 3 adrenoceptors. J Biol Chem. 1992 Jan 25;267(3):2006-13. 52. Sauvanet C, Duvezin-Caubet S, di Rago JP, Rojo M. Energetic requirements and bioenergetic modulation of mitochondrial morphology and dynamics. Semin Cell Dev Biol. 2010 Aug;21(6):558-65. 53. Guillery O, Malka F, Frachon P, Milea D, Rojo M, Lombes A. Modulation of mitochondrial morphology by bioenergetics defects in primary human fibroblasts. Neuromuscul Disord. 2008 Apr;18(4):319-30. 54. Dinopoulos A, Smeitink J, ter Laak H. Unusual features of mitochondrial degeneration in skeletal muscle of patients with nuclear complex I mutation. Acta Neuropathol. 2005 Aug;110(2):199-202. 55. Stadhouders AM, Sengers RC. Morphological observations in skeletal muscle from patients with a mitochondrial myopathy. J Inherit Metab Dis. 1987;10 Suppl 1:62-80.

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Functional assays preceding exome sequencing: a rational

strategy

An I. Jonckheere, Herma Renkema, Maaike Bras, Lambert P. van den Heuvel, Alexander

Hoischen, Sander B. Nabuurs, Martijn A. Huynen, Maaike C. de Vries, Jan A.M. Smeitink,

Richard J.T. Rodenburg

In preparation

Chapter 7

ABSTRACT

Whole exome sequencing (WES) is a powerful tool to detect novel pathogenic mutations.

The interpretation of WES data however remains challenging. Here, we present two siblings with a severe neonatal encephalopathy caused by complex V deficiency. WES analysis revealed a heterozygous mutation the ATP5A1 gene, coding for complex V subunit α, in the two patients, but also in their father. Due to the pathogenic susceptibility of the mutation, cDNA was checked and revealed a homozygous mutation in the patients, while father remained heterozygous. Complementation with wild type ATP5A1 restored complex V, confirming the pathogenicity of the mutation. At protein level, there is a disturbed interaction of the α subunit with the β subunit of complex V, which interferes with the stability of the complex. In summary, we describe the first human complex V ATP5A1 mutation and demonstrate the value of functional assays preceding and following WES analysis.

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INTRODUCTION

To date, mitochondrial (mt) ATP synthase (complex V) deficiencies have been described in the mtDNA encoded ATP6 (MT-ATP6) (MIM +516060) and ATP8 (MT-ATP8) (MIM +516070) genes (www.mitomap.org). ATP synthase deficiency due to the nuclear encoded ATP12

(MIM 604273) and TMEM70 (MIM 612418) genes has also been reported (1-2). Interestingly, only one mutation has hitherto been found in a nuclear encoded structural complex V subunit

(3), ATP5E (MIM 606153). Most of these genes however are not routinely screened in a diagnostic setting since complex V deficiency is not a common OXPHOS deficiency and since there is a large set of candidate genes (4).

Recent advances in next-generation sequencing can counter the latter issue. Since most

Mendelian disorders are caused by exonic mutations or splice-site mutations, whole exome sequencing (WES) can be applied to detect genetic defects underlying monogenic inherited disorders (5). On one hand, WES can be initiated “blindly” based only on a clinical phenotype. The interpretation of millions of genomic variants however is very challenging.

Bioinformatics analysis tools and filter criteria (6-7) are used to identify causative mutations.

For instance, when a recessive mode of inheritance is suspected, only the genes carrying homozygous or compound heterozygous variants are selected (6-7). On the other hand, biochemical data can be used to guide WES analysis. The latter approach has been applied in this study, describing two siblings with severe neonatal encephalopathy and an isolated complex V deficiency. WES was subsequently performed to uncover the genetic defect. We show that WES data have to be interpreted with caution in certain cases and that mitochondrial biochemistry preceding WES analysis can be very helpful to do so.

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PATIENTS AND METHODS

Patients

We report two patients from a Dutch non-consanguineous family with the same clinical phenotype. Pregnancy and delivery were uneventful in both cases. They both died in the first weeks of life due to a severe encephalopathy characterized by intractable seizures (apnoeic spells). Clinical examination showed no dysmorphic features or organomegaly. Neurologic examination at day one revealed irritability, a high-pitched cry, a horizontal and vertical nystagmus, abnormal primitive reflexes, and tonus dysregulation. Brain MRI demonstrated a progressive and severe encephalopathy characterized by hyperdense thalami and subcortical densities in the eldest sibling. The EEG showed abnormal slow waves in the right hemisphere. Postmortem anatomopathological examination revealed extensive cerebral damage: the cerebellum was too small, the consistency of the occipital lobe was weakened.

There was a cystic degeneration of the white matter, especially occipital and temporoparietal.

Also the pons and the brain stem were damaged. The lungs were hypoplastic, there were small renal cysts, and skeletal muscle contained small lipid droplets. The cerebral, renal and skeletal muscle lesions indicated a mitochondrial disease. The MRI of the younger patient was characterized by progressive frontal and parieto-occipital damage including the posterior limb of the internal capsule (PLIC), the pyramidal tract and basal ganglia, mainly in the right hemisphere. Also the cerebellum and pons were damaged. Metabolic screening in blood, urine and liquor showed no abnormalities in both cases.

Cell cultures

Fibroblasts were cultured in medium 199 (Gibco®, Invitrogen Corporation) supplemented with 10% fetal calf serum and penicillin/streptomycin (respectively 100 U/ml and 100 µg/ml).

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293FT cells (Invitrogen, Breda, The Netherlands) were grown in Dulbecco's Modified Eagle's

Medium (DMEM) containing 4.5 g/L glucose, 10% fetal calf serum, 4 mM L-glutamine, 1 mM

MEM sodium pyruvate, 0.1 mM MEM non-essential amino acids, 1% penicillin/streptomycin, and 500 μg/ml geneticin (G418). During transfection, the medium does not contain penicillin/streptomycin and geneticin (according to Invitrogen recommendations).

Biochemical assays

Mitochondrial respiration in digitonin-permeabilized cells was analyzed in the presence of a substrate mix containing pyruvate and malate, using the phosphorescent probe MitoXpress

(Luxcel, Cork, Ireland), 96-wells plates and a fluorescence plate reader as described (8). The complex V (or mtATPase) activity was measured spectrophotometrically in mitochondria isolated from patient and control fibroblasts as described (9). The activities of the mitochondrial respiratory chain enzymes and citrate synthase (CS) were measured spectrophotometrically in fibroblasts of the patients according to described protocols (4, 10-

11).

Complex V assembly and activity

One-dimensional (1D) 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-

PAGE), 5%-15% blue native (BN)-PAGE and two-dimensional (2D) 10% SDS-PAGE after

BN-PAGE were performed as described previously (12-13). Lanes were loaded with 30 (SDS analysis) or 40 (BN analysis) µg of solubilized mitochondrial protein (14). After electrophoresis, gels were processed further for immunoblotting as described by (13). For

Western blotting, antibodies used were raised against V5 (Invitrogen), mitochondrial complex

V subunit alpha, beta, OSCP, d; complex I subunit NDUFA9, complex IV subunit IV, complex

III subunit core 2, and CoII-70 kDa (complex II) (MitoSciences). For the in-gel activity assay of ATP hydrolysis the gel was incubated overnight at room temperature with the following

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solution: 35 mM Tris, 270 mM glycine, 14 mM MgSO4, 0.2% Pb(NO3)2, and 8 mM ATP, pH

7.8 (13).

Molecular genetic analysis

Whole exome sequencing

We sequenced the exomes (Sure Select v2, 50Mb, Agilent, Santa Clara, CA, USA) of the two affected relatives, patient 2 (DDNA10-0926) and patient 1 (DDNA10-0976) on a SOLiD 4 system (Life Technologies, Carlsbad, CA, USA). We sequenced a total of 157,698,947 and

126,253,568 50 base pair (bp) reads, respectively. Sequence data were mapped to the hg19 reference genome with the SOLiD bioscope software version 1.3 (Table 1a). This resulted in

83% and 80% of the targets being covered at least 10 times, and a median coverage of 71- and 55-fold respectively (see Table 1a).

Variants were called by Bioscope v1.3 and annotated using a custom-made bioinformatics pipeline (15-16). Variants were prioritized based on predicted amino acid consequences and overlap with common variation (dbSNP, in-house variant database) (Table 1b). All private non-synonymous and splice site variants that were present in both affected individuals are shown in Table 1c (see at the end of this article).

Table 1a: Exome sequencing performance DDNA10-0926 DDNA10-0976 Total mapped bases in regions: 4.999Gb (80.3%) 3.987Gb (75.3%) Total mapped bases near regions: 0.488Gb (7.8%) 0.492Gb (9.3%) Total mapped bases outside regions: 0.740Gb (11.9%) 0.812Gb (15.3%) Total reads: 157,698,947 126,253,568 Total mapped reads: 129,880,994 109,891,958 Median target coverage 71.0 55.2 % Targets covered >=10x 83% 80% % Targets covered >=20x 76% 72%

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Table 1b: Variant prioritization DDNA10-0926 DDNA10-0976 # Variants called 29,063 28,434 Exonic and splice sites 14,700 14,141 Non-synonymous and splice 8,101 7,951 sites Novel dbSNP v134 625 614 Novel in-house database 127 108 Of those: Shared variants 22 (Table 1c)

ATP5A1 genomic and coding DNA analysis

Total DNA from blood and cultured fibroblasts was extracted using the salting out procedure for human DNA extraction (17). The coding region in genomic DNA of the ATP5A1 gene was polymerase chain reaction (PCR) amplified using ten primer pairs (Table 2a). PCR conditions were 95°C for 10 minutes (denaturation), 95°C for 30 seconds, 60°C for 30 seconds and 72°C for 1 min for a total of 35 cycles. The promotor region in of the ATP5A1 gene was amplified under the same PCR conditions using seven primer pairs (Table 2b).

Sequence analysis of PCR amplified products was performed on an ABI3730 automatic capillary sequencer using BigDye® terminator chemistry (Applied Biosystems). RNA from blood and cultured fibroblasts was extracted as described (17). Reverse Transcriptase PCR

(RT-PCR) was performed to produce cDNA followed by PCR amplification (Table 2c).

Sequence analysis was subsequently performed as described above.

Table 2a: Primers for PCR amplification of the genomic DNA Exon(s) Forward primer Reverse primer Comment 2 5'-CAGTACTTCCGGGTCAGGTG-3' 5'-ATTCTGCCTCTGCGCAGGTGAC-3' Conventional PCR 3 5'-GATCACACTAAGCTGGCATTTG-3' 5'-CAACTCACCACAGTTATACGATG-3' Conventional PCR 4 5'-AAGTGCTGGGATTACAGCTGTGAG-3' 5'-GAATAACTGAGAAGACTTAGATCC-3' Conventional PCR 5-6 5'-TTGGTGTATGGGGAGGAGTG-3' 5'-ACCATTCCAAGACTAAAATATAATCC-3' Conventional PCR 7 5'-TGCATAGACACAGTAACAGTCTG-3' 5'-CTACAATCAGCAGCAATGGGAC-3' Conventional PCR 8-9* 5'-TTTTATAGTAGCCTCCAATTTAAAACC- 5'-CGGTAAGCAACAGCCTATGGTAC-3' Conventional 3' PCR 9* 5'-GTACCATAGGCTGTTGCTTACCG-3' 5'-CAACCAATGCCTTAGAATTATCC-3' Conventional PCR 10 5'-TTGGGGTACTGATGCTTTGC-3' 5'-GTACTGTAACTCAGTGACACAGAGC-3' Conventional PCR 11 5'-GCTCTGTGTCACTGAGTTACAGTAC-3' 5'-TGTGATGCAGCTCTGTAGGC-3' Conventional PCR 12-13 5'-GACCTTGGTAACATGGCATTC-3' 5'-CTGGTATTTGATGTGAATCC-3' Conventional PCR *Exon 9 is amplified in two overlapping fragments

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Table 2b: Primers for PCR amplification of the promoter region Fragment Forward primer Reverse primer Comment 1 5'-CCAGGTAAGCATAGAAAGTG-3' 5'-CCCATCTGGTGCTAGAGGA-3' Conventional PCR 2 5'-CCAGGTAAGCATAGAAAGTG-3' 5'-ATATTCCGGCTTTGGTCCTG-3' Conventional PCR 3 5'-CTATCATCCCATCACCTAGG-3' 5'-CCCATCTGGTGCTAGAGGA-3' Conventional PCR 4 5'-CTATCATCCCATCACCTAGG-3' 5'-ATATTCCGGCTTTGGTCCTG-3' Conventional PCR 5 5'-CACGGTCTAACACTACCGAC-3' 5'-ATATTCCGGCTTTGGTCCTG-3' Conventional PCR 6 5'-CACGGTCTAACACTACCGAC-3' 5'-ATTCTGCCTCTGCGCAGGTGAC-3' Conventional PCR 7 5'-GACACAGGTGCCCTTTTGCTC-3' 5'-GGGTATTAATCCTTACAGCC-3' Conventional PCR

Table 2c: Primers for PCR amplification of the cDNA Fragment Forward primer Reverse primer 1 5'-CTCCTTACTCTGGCTGTTCC-3' 5'-CTGGCAAAGCAGTCAAGGAG-3'

Multiplex Ligation-dependent Probe Amplification (MLPA) analysis

To detect exon deletions or duplications in ATP5A1, MLPA analysis was performed as described (18).

Comparative genomic hybridization (CGH) array

Affymetrix CytoScan HD array was performed to search for genome-wide deletions or duplications with a median resolution of 20 kilo base (Human Genome Build hg19, UCSC genome browser February 2009).

Cloning of human ATP5A1 gene and lentiviral complementation

The full length ORF for ATP5A1 was amplified by PCR from I.M.A.G.E. cDNA clone

IRATp970G0784D (Source BioScience) using specific primers with Gateway AttB flanking sites without a stop codon. The product was cloned into pDONR201 using the Gateway BP

Clonase II Enzyme Mix (Invitrogen). The resulting entryclone was verified by sequence

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(Invitrogen) using the Gateway LR Clonase II Enzyme Mix (Invitrogen). The Gateway BP and

LR reactions were performed according to the manufacturer’s instructions. The resulting pLenti6.2-V5-ATP5A1 or the control construct pLenti6.2-V5-AcGFP were transfected into 10 cm dishes 293FT cells together with a packaging mix consisting of pLP1, pLP2, and pLP/VSVG using Lipofectamine 2000 (Invitrogen). At 24 hours after transfection the medium was refreshed and supernatants containing the viral particles were harvested at 72 hours after transfection. Supernatants were cleared by centrifugation (5 min. 300 g) and the viral particles were stored at -80°C prior to use. Infections were performed on fibroblasts in 75 cm2 flasks with 2 ml of virus containing supernatant in the presence of 6 µg/ml polybrene (Sigma).

At 24 hours after infection the medium was refreshed and at 48 hours after transfection the selection medium was added (M199, 20 % fetal calf serum, penicillin/streptomycin

(respectively 100 U/ml and 100 µg/ml), 2 µg/ml blasticidin (InvivoGen)). Cells were selected for 14 days, in which time the mock infected cells (without virus) died. Blasticidin resistant cells were used for biochemical analysis within six passages after transduction.

Three-dimensional (3D) Modeling

A 3D model of the human F1-ATPase, including the Arg329Cys mutation, was created using the automatic homology modeling script in the YASARA & WHAT IF Twinset (19-20). As a template for this model we used PDB structure 1BMF (21), containing the structure of the bovine F1-ATPase, of which the alpha-subunit is 98% identical to the human one.

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RESULTS

Biochemical assays

Oxygen consumption measurements in fibroblasts as described by (8) showed a decreased oxygen consumption rate in cells of both patients (Figure 1a,b) compared to control cells

(Figure 1c). A substantial increase was seen after addition of an uncoupler (2 µM carbonyl cyanide 3-chlorophenyl hydrazone: CCCP) in the fibroblasts of both patients (Figure 1a,b).

This is not observed in control cells (Figure 1c) and indicates a complex V deficiency (8).

Figure 1 Oxygen consumption rate in fibroblasts. A decreased oxygen consumption rate is seen in both patients (a, b) compared to the control (c). A clearly higher rate is observed in the cells of both patients (a, b) after addition of an uncoupler (2 μM carbonyl cyanide 3- chlorophenyl hydrazone (CCCP)). This phenomenon is not observed in control cells (c) and indicates a complex V deficiency. (Normalized time profile of fluorescence intensity; RFU = relative fluorescence units; CS = citrate synthase; fibroblasts of (a) patient 1 and (b) patient 2; ● = +ADP ○ = -ADP; ▼= +ADP + CCCP; ∆ = -ADP + CCCP).

Measurements of the mitochondrial respiratory chain enzyme activities were performed in cultured fibroblasts (no muscle tissue was available for both patients) and displayed an isolated complex V deficiency in both cases (Table 3).

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Table 3: Enzymatic analysis of the activities of complex V (mtATPase), the respiratory chain enzymes and CS in fibroblasts of patients 1 and 2.

Patient 1 Patient 2

Complex V, mU/U COX a 116 (209-935) c 76 (209-935)

Complex I, mU/U COX 140 (110-260) 126 (110-260)

Complex II, mU/U COX 661 (536-1027) 698 (536-1027)

Complex III, mU/U COX 2311 (1270-2620) 2431 (1270-2620)

Complex IV, mU/U CS b 929 (680-1190) 918 (680-1190)

CS, mU/mg 171 (144-257) 179 (144-257) a COX: cytochrome c oxidase; b CS: citrate synthase; c figures between brackets are reference ranges.

Complex V assembly and activity

We performed 1D and 2D BN-PAGE analysis on fibroblasts of the two patients and a healthy control. 1D BN-PAGE showed that the amount of complex V was strongly reduced in the patients compared to the control (Figure 2a) The in-gel activity assay (IGA) of ATP hydrolysis demonstrated a reduced complex V activity (Figure 2a, top panel). 2D BN-PAGE analysis showed holocomplex V, with no accumulation of subcomplexes (Figure 2b). 1D

SDS-PAGE showed that the levels of individual complex V subunits were decreased (Figure

2c), which is in accordance with (22) describing proteolytic removal of unassembled F1 subunits in HeLa cells depleted in the β subunit of complex V. In conclusion, these data indicate a disturbed complex V assembly in an early stage (at the F1 level).

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Figure 2 Complex V assembly and activity in fibroblasts. (a) BN-PAGE shows a reduced amount and activity (top panel, in-gel activity (IGA)) of complex V in both patients. (b) Two dimensional BN-PAGE analysis shows holocomplex V, with no accumulation of subcomplexes. (c) Immunoblot after SDS-PAGE shows decreased levels of complex V subunits. Note that blots stained with different antibodies can only be compared qualitatively since exposure times are not the same. Antibodies used are against complex V subunits α, β, OSCP and d, complex I subunit NDUFA9, and complex II-70 kDa (loading control). (8834: patient 2 and 8618: patient 1 fibroblasts; H: holocomplex).

Molecular genetic analysis

Genetic analysis did not reveal mutations in the mtDNA or nuclear genes known to cause complex V deficiency (ATP12, TMEM70, ATP5E). Therefore, whole exome sequencing was performed and revealed a heterozygous mutation at c.985C>T (p.Arg329Cys) (Ref Seq accession number NM_001001937) in exon 9 of ATP5A1 (chromosome 18q21, MIM

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*164360). Father was also heterozygous, whereas mother did not have this mutation (Figure

3, left panel). The mutation was not present in more than 120 healthy Caucasian controls.

Next, cDNA was checked and revealed a homozygous c.985C>T mutation in both patients, father was heterozygous and mother showed the wild-type coding sequence (Figure 3, right panel).

Figure 3 Molecular genetic analysis in fibroblasts. Left panel: genomic (g)DNA analysis shows a heterozygous mutation at c.985C>T (pArg329Cys) in exon 9 of ATP5A1 in both patients and in the father. Mother shows no mutations. Right panel: coding (c)DNA analysis shows a homozygous mutation in both patients. Father is heterozygous, and mother does not have this mutation.

This means that one allele of mother is not expressed and that, in fact, both patients are hemizygous for the mutation at cDNA level. The loss of this transcript was not due to promoter region mutations (checked from g.41931368 – g.41933253 (Chr18, NCBI Build

36)). MLPA analysis did not show deletions or duplications in ATP5A1 alleles (data not shown). CytoScan HD array showed no alterations (data not shown).

Complementation of complex V deficiency by wild type ATP5A1

To confirm the pathogenicity of the mutation, we transduced the index patients and a healthy control using the lentiviral system. Complementation of patient fibroblasts with wild-type

ATP5A1 completely normalized complex V amount (Figure 4) and activity (Table 4).

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Figure 4 Complementation with wild type ATP5A1. (a) BN-PAGE shows that complex V amount normalizes in patient fibroblasts after complementation with wild type ATP5A1. Incubation with anti-V5 antibody demonstrates that the wild type ATP5A1 construct is incorporated correctly. Other OXPHOS complexes are expressed normally. Of note, complementation of the control fibroblasts is less clear since these cells don’t need the complemented construct (competitive inhibition with wild type complex V). (b) Immunoblot after SDS-PAGE again demonstrates that the wild type ATP5A1 construct is incorporated correctly. Incubation with subunit α antibody shows two bands, the lower band is native subunit α, the upper band is subunit α tagged with V5. Antibodies used are against V5, complex V subunits α and β, complex IV subunit IV, complex III core 2, complex I NDUFA9, and complex II-70 kDa (loading control). (8834: patient 2 and 8618: patient 1 fibroblasts; WT: wild type; GFP: negative control construct pLent6.2-V5- AcGFP; 5A: wild type ATP5A1 construct pLent6.2-V5-ATP5A1).

Table 4: Complementation with wild type ATP5A1 restores complex V activity in patient fibroblasts.

Complex V activity (mU/U CS, reference

range 193-819)c

Patient 1 - GFPa 118

Patient 1 – ATP5A1b 337

Patient 2 - GFP 118

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Patient 2 – ATP5A1 381

Control - GFP 748

Control – ATP5A1 665 a Patient 1 - GFP: negative control construct pLent6.2-V5-AcGFP; b Patient 1 – ATP5A1: wild type ATP5A1 construct pLent6.2-V5-ATP5A1.

Modeling of p.Arg329Cys in ATP5A1

The pathogenicity of the mutation was also corroborated by protein modeling. The arginine

329 of the α subunit that is mutated in the patients is predicted to normally form a hydrogen bond with a carbonyl group of the glycine 323 of the β subunit (Figure 5). As the F1-ATPase has three α subunits and three β subunits, the Arg329Cys mutation would break three stabilizing interactions in the complex. This predicted loss of stability of the complex is in line with the Western blot results that show degradation of unassembled subunits. Furthermore, the arginine 329 is very well conserved, even among bacterial orthologs of ATP5A1 (data not shown), underlining its importance for the functioning of the complex.

Figure 5 Impact of the ATP5A1 c.985C>T mutation at the protein structure level. p.Arg329Cys disturbs the interaction of the α subunit with the β subunit. Normally arginine 329 is predicted to interact with a carbonyl group of glycine 323 in the backbone of the β subunit. The Arg329Cys mutation is thus predicted to interfere with the stability of complex V. (R: arginine; C: cysteine; G: glycine)

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DISCUSSION

Clinical phenotype

The severe neonatal encephalopathy of the patients described here resembles the clinical picture of the patient with a mutation in the complex V assembly gene ATP12 (1). The latter patient however also presented with dysmorphic features, an enlarged liver and hypoplastic kidneys, features that were not present here. It should be noted that postmortem anatomopathological examination of the eldest sibling revealed renal cysts and hypoplastic lungs. Remarkably, there is a predominance of the cerebral lesions in the right hemisphere in the patients described here. Additional patients with mutations in ATP5A1 are needed to confirm this clinical phenotype. Hypertrophic cardiomyopathy, which is often seen in complex

V deficiency (2, 9), was not documented. A 4 base pair duplication in exon 3 of ATP5A1 leading to a stopcodon in exon 4 and therefore an instable transcript, has been described in an animal model and this resulted in death in utero (23).

Pathogenicity

Several lines of evidence point to the pathogenicity of the ATP5A1 mutation. Clinically, there is a severe and fatal neonatal encephalopathy, while biochemically, an isolated complex V deficiency was found. Modeling of the mutation showed a disturbed interaction of the α and β subunit of complex V. This leads to instability of the complex which is in line with the Western blot results, showing a severely decreased holocomplex V and degradation of unassembled subunits. Complementation of the ATP5A1 defect normalized the amount and activity of complex V. Finally, the mutation is situated in a conserved region of the protein.

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Value of functional assays before and after whole exome sequencing

In the routine diagnostic setting for mitochondrial disorders, we first perform functional

(mitochondrial energy-generating system (MEGS) capacity, oxygen consumption and enzymatic activity) assays. Genetic analysis is subsequently performed, oriented on these results. This approach was successful here, since the heterozygous mutation in ATP5A1 which was found in both patients and in their father was only investigated further because of the known complex V deficiency in the patients. Therefore we want to emphasize the importance of functional assays preceding exome sequencing. Biochemical data can help to identify causative mutations which would have been filtered out when only WES data were considered. Indeed, the patients described here nicely show that also heterozygous mutations found in the index patient(s) and in one of the parents warrant further investigation if the mutation is suspected to be pathogenic. This differs from whole exome sequencing strategies which include homozygosity as a filter criterion when a recessive mode of inheritance is suspected (6-7). After WES, defining the true impact of genomic variants by functional follow-up (cDNA analysis, complementation studies) is mandatory. This is also emphasized in (24), where a branch-site mutation instead of a homozygous missense mutation in the NUBPL gene was shown to cause complex I deficiency.

Together, we currently recommend functional studies before and after WES analysis to identify and confirm the pathogenicity of a mutation. As genetic knowledge and technology is rapidly evolving, time will tell whether functional assays remain valuable in a whole exome or whole genome sequencing setting.

Implications

In summary, we report the first mutation in ATP5A1, which codes for complex V subunit α and gives rise to an isolated complex V deficiency with a distinct clinical phenotype, i.e. a severe neonatal encephalopathy leading to early death. Moreover, this study demonstrated

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Funding

This work was supported by a grant from the Prinses Beatrix Fonds [grant number OP-05-04] to AIJ and JS. The authors confirm independence from the sponsors; the content of the article has not been influenced by the sponsors.

Acknowledgements

The authors thank Marthe HendriksFranssen, Marion Antoine, Mariël van den Brand and

Karen Janssen for technical assistance.

Conflicts of interest

None.

The study has been carried out in the Netherlands in accordance with the applicable rules concerning the review of research ethics committees (Commissie Mensgebonden Onderzoek

Regio Arnhem-Nijmegen) and informed consent.

REFERENCES

1. De Meirleir L, Seneca S, Lissens W, De Clercq I, Eyskens F, Gerlo E, et al. Respiratory chain complex V deficiency due to a mutation in the assembly gene ATP12. J Med Genet. 2004 Feb;41(2):120-4. 2. Cizkova A, Stranecky V, Mayr JA, Tesarova M, Havlickova V, Paul J, et al. TMEM70 mutations cause isolated ATP synthase deficiency and neonatal mitochondrial encephalocardiomyopathy. Nat Genet. 2008 Nov;40(11):1288-90. 3. Mayr JA, Havlickova V, Zimmermann F, Magler I, Kaplanova V, Jesina P, et al. Mitochondrial ATP synthase deficiency due to a mutation in the ATP5E gene for the F1 epsilon subunit. Hum Mol Genet. 2010 Sep 1;19(17):3430-9. 4. Rodenburg RJ. Biochemical diagnosis of mitochondrial disorders. J Inherit Metab Dis. 2011 Apr;34(2):283-92. 5. Majewski J, Schwartzentruber J, Lalonde E, Montpetit A, Jabado N. What can exome sequencing do for you? J Med Genet. 2011 Sep;48(9):580-9.

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6. Gotz A, Tyynismaa H, Euro L, Ellonen P, Hyotylainen T, Ojala T, et al. Exome sequencing identifies mitochondrial alanyl-tRNA synthetase mutations in infantile mitochondrial cardiomyopathy. Am J Hum Genet. 2011 May 13;88(5):635-42. 7. Gilissen C, Hoischen A, Brunner HG, Veltman JA. Disease gene identification strategies for exome sequencing. Eur J Hum Genet. 2012 Jan 18. 8. Jonckheere AI, Huigsloot M, Janssen AJ, Kappen AJ, Smeitink JA, Rodenburg RJ. High-throughput assay to measure oxygen consumption in digitonin-permeabilized cells of patients with mitochondrial disorders. Clin Chem. 2010 Mar;56(3):424-31. 9. Jonckheere AI, Hogeveen M, Nijtmans LG, van den Brand MA, Janssen AJ, Diepstra JH, et al. A novel mitochondrial ATP8 gene mutation in a patient with apical hypertrophic cardiomyopathy and neuropathy. J Med Genet. 2008 Mar;45(3):129-33. 10. Janssen AJ, Smeitink JA, van den Heuvel LP. Some practical aspects of providing a diagnostic service for respiratory chain defects. Ann Clin Biochem. 2003 Jan;40(Pt 1):3-8. 11. Srere PA. Citrate synthase 1969. 12. Schagger H, von Jagow G. Blue native electrophoresis for isolation of membrane protein complexes in enzymatically active form. Anal Biochem. 1991 Dec;199(2):223-31. 13. Nijtmans LG, Henderson NS, Holt IJ. Blue Native electrophoresis to study mitochondrial and other protein complexes. Methods. 2002 Apr;26(4):327-34. 14. Smith PK, Krohn RI, Hermanson GT, Mallia AK, Gartner FH, Provenzano MD, et al. Measurement of protein using bicinchoninic acid. Anal Biochem. 1985 Oct;150(1):76-85. 15. Gilissen C, Arts HH, Hoischen A, Spruijt L, Mans DA, Arts P, et al. Exome sequencing identifies WDR35 variants involved in Sensenbrenner syndrome. Am J Hum Genet. 2010 Sep 10;87(3):418-23. 16. Hoischen A, van Bon BW, Rodriguez-Santiago B, Gilissen C, Vissers LE, de Vries P, et al. De novo nonsense mutations in ASXL1 cause Bohring-Opitz syndrome. Nat Genet. 2011 Aug;43(8):729-31. 17. Miller SA, Dykes DD, Polesky HF. A simple salting out procedure for extracting DNA from human nucleated cells. Nucleic Acids Res. 1988 Feb 11;16(3):1215. 18. Schouten JP, McElgunn CJ, Waaijer R, Zwijnenburg D, Diepvens F, Pals G. Relative quantification of 40 nucleic acid sequences by multiplex ligation-dependent probe amplification. Nucleic Acids Res. 2002 Jun 15;30(12):e57. 19. Vriend G. WHAT IF: a molecular modeling and drug design program. J Mol Graph. 1990 Mar;8(1):52-6, 29. 20. Krieger E, Koraimann G, Vriend G. Increasing the precision of comparative models with YASARA NOVA--a self-parameterizing force field. Proteins. 2002 May 15;47(3):393- 402.

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21. Abrahams JP, Leslie AG, Lutter R, Walker JE. Structure at 2.8 A resolution of F1- ATPase from bovine heart mitochondria. Nature. 1994 Aug 25;370(6491):621-8. 22. Rak M, McStay GP, Fujikawa M, Yoshida M, Manfredi G, Tzagoloff A. Turnover of ATP synthase subunits in F(1)-depleted HeLa and yeast cells. FEBS Lett. 2011 Aug 19;585(16):2582-6. 23. Baran AA, Silverman KA, Zeskand J, Koratkar R, Palmer A, McCullen K, et al. The modifier of Min 2 (Mom2) locus: embryonic lethality of a mutation in the Atp5a1 gene suggests a novel mechanism of polyp suppression. Genome Res. 2007 May;17(5):566-76. 24. Tucker EJ, Mimaki M, Compton AG, McKenzie M, Ryan MT, Thorburn DR. Next- generation sequencing in molecular diagnosis: NUBPL mutations highlight the challenges of variant detection and interpretation. Hum Mutat. 2012 Feb;33(2):411-8.

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Table 1c: Summary of all private non-synonymous and splice site variants identified in both affected individuals by WES Tota Vari Genomic % Referenc Mutation AminoA mRNA Granth Patien Chr Refere Vari l ant Gene phyl position variati Gene id e Amino Amino cid chang am t ID . nce ant read read name oP [hg19] on Acid Acid changes es Score s s chr NM_001009 P818S 2380C DDNA- 23,409,678 C T 75 34 45.3 KDM1A P S 5.83 74 100976 1 999 P794S >T DDNA- chr NM_001009 P794S 2452C 23,409,678 C T 110 44 40.0 KDM1A P S 5.83 74 100926 1 999 P818S >T chr 1009A DDNA- 113,215,017 A T 55 22 40.0 TTC12 NM_017868 I L I337L -0.73 5 100976 11 >T DDNA- chr 1009A 113,215,017 A T 117 47 40.2 TTC12 NM_017868 I L I337L -0.73 5 100926 11 >T DDNA- chr 581A> 9,501,029 A G 47 23 48.9 ZNF143 NM_003442 D G D194G 3.70 94 100976 11 G chr 581A> DDNA- 9,501,029 A G 60 28 46.7 ZNF143 NM_003442 D G D194G 3.70 94 100926 11 G DDNA- chr 710G> 73,714,986 G A 15 5 33.3 UCP3 NM_003356 P L P237L 5.87 98 100976 11 A chr 710G> DDNA- 73,714,986 G A 14 4 28.6 UCP3 NM_003356 P L P237L 5.87 98 100926 11 A DDNA- chr KIAA17 3514A 93,431,592 A G 155 74 47.7 NM_033395 I V I1172V 0.54 29 100976 11 31 >G chr KIAA17 3514A DDNA- 93,431,592 A G 198 86 43.4 NM_033395 I V I1172V 0.54 29 100926 11 31 >G DDNA- chr BDKRB 173T> 96,730,192 T C 16 8 50.0 NM_000710 V A V58A 2.26 64 100976 14 1 C DDNA- chr BDKRB 173T> 96,730,192 T C 28 14 50.0 NM_000710 V A V58A 2.26 64 100926 14 1 C chr 184G> DDNA- 43,308,025 G A 22 13 59.1 FMNL1 NM_005892 E K E62K 4.95 56 100976 17 A DDNA- chr 184G> 43,308,025 G A 52 26 50.0 FMNL1 NM_005892 E K E62K 4.95 56 100926 17 A chr 1451G DDNA- 40,052,881 G A 25 11 44.0 ACLY NM_001096 S F S484F 2.28 155 100976 17 >A DDNA- chr 1451G 40,052,881 G A 43 17 39.5 ACLY NM_001096 S F S484F 2.28 155 100926 17 >A chr ATP5A 985G> DDNA- 43,667,165 G A 28 12 42.9 NM_004046 R C R329C 5.70 180 100976 18 1 A DDNA- chr ATP5A 985G> 43,667,165 G A 25 12 48.0 NM_004046 R C R329C 5.70 180 100926 18 1 A DDNA- chr NM_001032 potential 2392> 44,681,807 - A 17 13 76.5 ZNF226 E E798EX 1.36 NA 100976 19 373 fs A chr NM_001032 potential 2392> DDNA- 44,681,807 - A 15 9 60.0 ZNF226 E E798EX 1.36 NA 100926 19 373 fs A DDNA- chr NM_001164 3716C 152,534,137 C A 396 155 39.1 NEB S I S1239I 5.93 142 100976 2 508 >A chr NM_001164 3716C DDNA- 152,534,137 C A 582 248 42.6 NEB S I S1239I 5.93 142 100926 2 508 >A DDNA- chr TACTT KIDINS splice 8,888,010 - 21 13 61.9 NM_020738 NA NA 100976 2 AC 220 site chr TACTT KIDINS splice DDNA- 8,888,010 - 52 21 40.4 NM_020738 NA NA 100926 2 AC 220 site DDNA- chr SEC14 1021G 30,887,620 G A 39 25 64.1 NM_174977 R C R341C 1.22 180 100976 22 L4 >A DDNA- chr SEC14 1021G 30,887,620 G A 39 24 61.5 NM_174977 R C R341C 1.22 180 100926 22 L4 >A chr 542A> DDNA- 88,189,002 A G 99 37 37.4 ZNF654 NM_018293 N S N181S 2.41 46 100976 3 G DDNA- chr 542A> 88,189,002 A G 91 46 50.5 ZNF654 NM_018293 N S N181S 2.41 46 100926 3 G chr MAP3K 421G> DDNA- 185,146,790 G A 38 16 42.1 NM_004721 V I V141I 6.10 29 100976 3 13 A DDNA- chr MAP3K 421G> 185,146,790 G A 77 29 37.7 NM_004721 V I V141I 6.10 29 100926 3 13 A

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chr NEURO DDNA- 113,436,601 T G 11 3 27.3 NM_024019 K Q K11Q 31T>G 0.93 53 100976 4 G2 DDNA- chr NEURO 113,436,601 T G 50 20 40.0 NM_024019 K Q K11Q 31T>G 0.93 53 100926 4 G2 chr 581A> DDNA- 155,530,867 A G 170 106 62.4 FGG NM_021870 F S F194S 2.16 155 100976 4 G DDNA- chr 581A> 155,530,867 A G 182 89 48.9 FGG NM_021870 F S F194S 2.16 155 100926 4 G chr PCDHG 1216A DDNA- 140,740,918 A T 82 28 34.1 NM_018923 T S T406S 2.45 58 100976 5 B2 >T DDNA- chr PCDHG 1216A 140,740,918 A T 88 28 31.8 NM_018923 T S T406S 2.45 58 100926 5 B2 >T DDNA- chr 4411G 33,648,392 G A 23 21 91.3 ITPR3 NM_002224 V I V1471I 3.34 29 100976 6 >A chr 4411G DDNA- 33,648,392 G A 42 32 76.2 ITPR3 NM_002224 V I V1471I 3.34 29 100926 6 >A DDNA- chr KIAA15 NM_001164 4807G 138,552,843 G A 60 23 38.3 R W R1603W 3.03 101 100976 7 49 665 >A chr KIAA15 NM_001164 4807G DDNA- 138,552,843 G A 37 12 32.4 R W R1603W 3.03 101 100926 7 49 665 >A DDNA- chr 1043T 101,601,143 T C 76 33 43.4 SNX31 NM_152628 Q R Q348R 3.43 43 100976 8 >C chr 1043T DDNA- 101,601,143 T C 94 39 41.5 SNX31 NM_152628 Q R Q348R 3.43 43 100926 8 >C DDNA- chr 1037G 101,980,430 G A 66 26 39.4 ALG2 NM_033087 S L S346L 4.24 145 100976 9 >A DDNA- chr 1037G 101,980,430 G A 57 27 47.4 ALG2 NM_033087 S L S346L 4.24 145 100926 9 >A

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PART IV

GENERAL DISCUSSION

General discussion, future perspectives and conclusions

General discussion and conclusions

PART II: CHARACTERIZATION OF HUMAN CELL LINES WITH MITOCHONDRIAL

DYSFUNCTION

Establishing a biochemical diagnosis in the mitochondrial medicine field is of utmost importance since it guides the search for the underlying genetic defect. Therefore, we extended current tools for biochemical diagnostics in patients suspected for mitochondrial disease in this thesis. Functional measurements in fresh muscle tissue, comprising substrate oxidation, ATP production and oxygen consumption measurements, evaluate the capacity of the whole mitochondrial energy-generating system (MEGS). It is important to be able to perform these functional measurements in other tissues than skeletal muscle. A decreased

MEGS capacity in other tissues than skeletal muscle, and this is certainly the case for confirming a defect in cultured skin fibroblasts, points to a primary mitochondriopathy, which is mostly inherited. A decreased MEGS capacity solely in muscle tissue can be secondary due to poor feeding or inactivity and can normalize in muscle tissue taken after clinical improvement, or can still indicate a primary mitochondriopathy, which is not expressed in the other tissue(s) examined. This means that functional assays in several tissues can be helpful to determine if it concerns an inherited disease. This is important for patient counseling.

Chapter 2 described a simple, high-throughput, high-content screening method for oxygen consumption analysis in different digitonin-permeabilized cell types. It was demonstrated that this assay provides a valuable tool in the diagnostics of mitochondrial disorders. It can differentiate between primary and secondary mitochondrial abnormalities. Patients with a primary mitochondriopathy that does not affect the complexes of the respiratory chain can be identified. At present, the method is validated in order to implement these measurements in the routine diagnostic setting. In a research context, the assay is used in the search for and characterization of new genetic defects, and can be applied to evaluate the effects of therapeutic interventions.

In chapter 3, two OXPHOS-deficient myogenic cell lines were established. It was shown that the complete biochemical toolbox, including substrate oxidation, ATP production rate and

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Chapter 8 oxygen consumption measurements, as well as spectrophotometric enzyme analysis and

BN-PAGE could be adapted successfully for the study of mitochondrial energy metabolism in myoblasts and myotubes. Mitochondrial disorders can thus be identified and characterized in muscle cells. This paves the way for multiple applications, e.g. the study of OXPHOS defects that are only expressed in muscle tissue, in-depth research for mitochondrial physiology using biological assays in living cells, and targeted pharmacological intervention studies. The immortalization of myoblasts can provide a back-up of muscle cells, which circumvents novel muscle biopsies.

Together, these new tools expand current diagnostic and counseling possibilities.

Mitochondrial biochemical assays are continuously being optimized, validated and extended.

In Nijmegen a micro method has been developed to measure the substrate oxidation and

ATP production rates in human muscle mitochondria requiring less than one-tenth of the original amount of muscle tissue. Spectrophotometric enzyme activity measurements are now automated and can be performed much faster, more accurate and also requiring less sample. This shortens completion time of diagnostic assays and therefore the time of uncertainty in patients and their families. Methods for the measurement of the pyruvate carrier, the glutamate carrier and the α-ketoglutarate carrier in muscle tissue using radioactive substrates have been developed. These assays will be adapted soon for fibroblasts. The development of methods to measure different carriers is important because it can identify patients with described defects (1-2), but also, when expanding the number of carrier assays, could lead to the discovery of novel defects, further unraveling the complexity of mitochondrial diseases. Finally, mitochondrial biochemical diagnostics does not only serve patients suspected for a mitochondriopathy, but also other patient groups e.g. Multiple

Sclerosis (MS) patients (3), Complex Regional Pain Syndrome type I (CPRS I) patients (4), and patients with several other disorders. Multiple mtDNA deletions are present in the gray matter of MS patients. It was proposed that these deletions are an important contributor to neurodegeneration in MS (3). Mitochondrial dysfunction was seen in muscle tissue of CPRS I patients; it remains to be established whether this is a primary or secondary phenomenon

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(4). So mitochondrial biochemical assays undoubtedly also serve research goals.

Biochemical measurements can guide the search for genetic defects. In that way, the genetic search can be performed step-wise and directed to specific genes, e.g. complex V genes in case of a complex V deficiency. Afterwards, biochemical diagnostic assays are again needed to evaluate the pathogenicity of genetic variants. Together with bioinformatics tools, biochemical assays are mandatory in the interpretation of whole exome sequencing data.

PART III: COMPLEX V IN HEALTH AND DISEASE

Unraveling the underlying genetic mutation and studying the functional impact of the defect in patients with a complex V deficiency was the second goal in this thesis.

Chapter 4 reviewed the architecture, function and assembly of complex V. The role of complex V di- and oligomerization and its relation with mitochondrial morphology was discussed. Finally, pathology related to complex V deficiency and current therapeutic strategies were highlighted. Thanks to the extensive research over the last decades, nowadays much is known regarding the structure and function of the world’s smallest rotary nanomotor. Most of the structure of the bovine mitochondrial enzyme has been resolved. The structures of the membrane domain of subunit b, subunit a, and the accessory subunits e, f, g, and A6L remain to be determined (5). The mechanism of the rotary F1Fo ATP synthase has been described by Boyer (6). Still, understanding the enzyme fully at a molecular level will require further efforts, both experimental and theoretical (for a review, see (7)). Next to structure and function of the monocomplex, also the role of di- and oligomerization of complex V, shaping the inner mitochondrial membrane, has been addressed in many studies both in yeast and in mammalian mitochondria (8-9). The role of IF1 in this process has been shown to be important (10-11).

In chapter 5, we studied the clinical features and molecular pathophysiology in a patient with the first described mitochondrial ATP8 gene mutation. The clinical picture was characterized

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Chapter 8 by apical hypertrophic cardiomyopathy and neuropathy. At the molecular level, we found an improper assembly and reduced activity of complex V.

In chapter 6, we studied mitochondrial morphology in a patient with a deletion in the

TMEM70 gene, causing complex V deficiency. We saw a fragmented mitochondrial network and swollen and irregularly shaped mitochondria with partial to complete loss of the cristae in fibroblasts of this patient, which could be completely restored by complementation of the

TMEM70 genetic defect. Comparative genomics analysis predicted the topology of TMEM70 in the inner mitochondrial membrane, which could be confirmed by immunogold labeling experiments, and showed that the TMEM70 gene is not restricted to higher multi-cellular eukaryotes. This study demonstrated that the role of complex V in mitochondrial cristae morphology applies to human mitochondrial disease pathology.

In chapter 7, we described two siblings with intractable epilepsy and complex V deficiency due to a mutation in ATP5A1, coding for complex V subunit α. The mutation gave rise to a disturbed interaction of the α subunit with the β subunit of complex V, which interfered with the stability of the complex. This study also demonstrated that biochemical diagnostics preceding and following whole exome sequencing are indispensable for the correct interpretation of these data, since heterozygous mutations found in the index case and in one of the parents warrant cDNA analysis if the mutation is suspected to be pathogenic.

Lots of questions regarding complex V remain to be answered. The assembly of the different subunits into the holocomplex continues to be puzzling. Most of the research has been done in yeast. But the yeast assembly process probably differs from the one in mammalian mitochondria, since there are substantial differences between higher and lower eukaryotes such as the number of Fo subunit c-genes, ATP synthase-specific assembly factors, and factors regulating transcription of ATP synthase genes (12). To gain further insight into the assembly of complex V, techniques like blue native and clear native PAGE, combined with incorporation and knock-down experiments of different subunits as described in (13) could refine our current knowledge. Further, only two assembly factors, ATP11 and ATP12, are

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General discussion and conclusions

hitherto known in mammalian ATP synthase. They both have a role in F1 assembly. TMEM70 maintains normal expression levels of complex V, and has been suggested to have a role in complex V biogenesis (14). The exact mechanism however still remains to be elucidated.

Tandem-affinity purification (TAP) experiments can be performed to identify TMEM70 co- purifying proteins. This information can be used to set up additional experiments in order to gain further insight into the TMEM70 mechanism. Moreover, the existence of specific factors involved in mammalian Fo formation is probable (12). A possible approach could be to study the evolution of complex V subunits and complex V chaperones by comparative genomics.

For example, the yeast Fo assembly factor Atp23p has a human homolog for which, however, no involvement in ATP synthase assembly could be demonstrated (15). Also a homology of complex V chaperones with other human proteins could be of interest in the search of specific assembly factors. Another intriguing fact is that to date, mutations have only been found in two nuclear encoded structural complex V genes (16). It could be possible that mutations in some of the structural subunits are incompatible with life. On the other hand, given the lower frequency of complex V deficiency compared to the other OXPHOS deficiencies, routine screening of all nuclear structural genes is rarely implemented in a diagnostic setting. Whole genome or whole exome screening (WES) could possibly solve some of the hitherto unknown genetic defects causing complex V deficiency. A current drawback of WES is an incomplete coverage of the whole exome (17). Furthermore, WES misses structural variation and variants situated in highly conserved non-coding regions of the genome (17). Whole genome sequencing (WGS) can overcome these issues. However, since the amount of data generated from WGS is 100 times more than WES, data analysis is, next to the cost of this assay, still a major challenge. Already for WES, an overwhelming set of data is obtained. In this context, functional biochemical characterization of newly discovered genetic variants or of genes with unknown function is, next to bioinformatical analysis, mandatory to define their pathogenicity and pathophysiological consequences. This means that biochemical diagnostic assays are needed both in the search for genetic defects and in defining the impact of found genetic variants in an individual patient. This is well

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Chapter 8 illustrated in chapter 7, where the knowledge of an isolated complex V deficiency in the patients stimulated further study of the heterozygous mutation found in the ATP5A1 gene by whole exome sequencing, even if the same mutation was also found in the father. The pathogenicity of the mutation was confirmed by complementing the patient cells with wild type ATP5A1. Finally, the biggest challenge will be to find a tailored curative therapy for this patient group. Compound screening can be applied to find a possible pharmacological approach. For example, the compound 5-aminoimidazole-4-carboxamide ribonucleoside

(AICAR) activates mitochondrial biogenesis via activation of the AMPK/PGC1α axis (18).

Stimulation of mitochondriogenesis nevertheless implies some residual activity of the deficient complex. If this is not the case, mitochondrial targeted delivery of the protein of interest via a liposome-based carrier system could be an alternative (19). For mtDNA defects, gene-shifting and germline techniques are promising. Nevertheless, much more and thorough experimental research is needed before these approaches can be implemented in the patient setting. In conclusion, mitochondrial ATP synthase has been and still is a popular research topic. Thanks to sustained effort, many aspects of this intriguing protein have been elucidated. This knowledge will guide further physio(patho)logical studies, paving the way for future therapeutic interventions.

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General discussion and conclusions

BRIEF SUMMARY AND IMPLICATIONS OF THIS THESIS

New tools for the biochemical characterization of cell lines of patients with a

(suspected) mitochondriopathy are described (see PART II). Providing the most accurate biochemical diagnosis is important for the identification of new genetic defects. PART III of this thesis focused on complex V deficiency, and again used biochemical functional assays e.g. complementation studies to determine the pathogenicity of the genetic defect. Furthermore, (patho)physiology was studied using

BN-PAGE, microscopy for mitochondrial morphology and for immunogold labeling

(TMEM70 mutation), and biomolecular informatics (TMEM70 and ATP5A1 mutations).

In conclusion, mitochondrial biochemistry and genetic analysis are complementary in the search for and in the characterization of novel mutations. Well-characterized defects are important for the development and testing of targeted therapeutic agents.

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Chapter 8

REFERENCES

1. Brivet M, Garcia-Cazorla A, Lyonnet S, Dumez Y, Nassogne MC, Slama A, et al. Impaired mitochondrial pyruvate importation in a patient and a fetus at risk. Mol Genet Metab. 2003 Mar;78(3):186-92. 2. Wibom R, Lasorsa FM, Tohonen V, Barbaro M, Sterky FH, Kucinski T, et al. AGC1 deficiency associated with global cerebral hypomyelination. N Engl J Med. 2009 Jul 30;361(5):489-95. 3. Campbell GR, Ziabreva I, Reeve AK, Krishnan KJ, Reynolds R, Howell O, et al. Mitochondrial DNA deletions and neurodegeneration in multiple sclerosis. Ann Neurol. 2011 Mar;69(3):481-92. 4. Tan EC, Janssen AJ, Roestenberg P, van den Heuvel LP, Goris RJ, Rodenburg RJ. Mitochondrial dysfunction in muscle tissue of complex regional pain syndrome type I patients. Eur J Pain. 2011 Aug;15(7):708-15. 5. Rees DM, Leslie AG, Walker JE. The structure of the membrane extrinsic region of bovine ATP synthase. Proc Natl Acad Sci U S A. 2009 Dec 22;106(51):21597-601. 6. Boyer PD. The ATP synthase--a splendid molecular machine. Annu Rev Biochem. 1997;66:717-49. 7. Junge W, Sielaff H, Engelbrecht S. Torque generation and elastic power transmission in the rotary F(O)F(1)-ATPase. Nature. 2009 May 21;459(7245):364-70. 8. Paumard P, Vaillier J, Coulary B, Schaeffer J, Soubannier V, Mueller DM, et al. The ATP synthase is involved in generating mitochondrial cristae morphology. EMBO J. 2002 Feb 1;21(3):221-30. 9. Strauss M, Hofhaus G, Schroder RR, Kuhlbrandt W. Dimer ribbons of ATP synthase shape the inner mitochondrial membrane. EMBO J. 2008 Apr 9;27(7):1154-60. 10. Campanella M, Casswell E, Chong S, Farah Z, Wieckowski MR, Abramov AY, et al. Regulation of mitochondrial structure and function by the F1Fo-ATPase inhibitor protein, IF1. Cell Metab. 2008 Jul;8(1):13-25. 11. Campanella M, Parker N, Tan CH, Hall AM, Duchen MR. IF(1): setting the pace of the F(1)F(o)-ATP synthase. Trends Biochem Sci. 2009 Jul;34(7):343-50. 12. Houstek J, Pickova A, Vojtiskova A, Mracek T, Pecina P, Jesina P. Mitochondrial diseases and genetic defects of ATP synthase. Biochim Biophys Acta. 2006 Sep-Oct;1757(9- 10):1400-5. 13. Wagner K, Perschil I, Fichter CD, van der Laan M. Stepwise assembly of dimeric F(1)F(o)-ATP synthase in mitochondria involves the small F(o)-subunits k and i. Mol Biol Cell. 2010 May 1;21(9):1494-504.

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14. Cizkova A, Stranecky V, Mayr JA, Tesarova M, Havlickova V, Paul J, et al. TMEM70 mutations cause isolated ATP synthase deficiency and neonatal mitochondrial encephalocardiomyopathy. Nat Genet. 2008 Nov;40(11):1288-90. 15. Kucharczyk R, Zick M, Bietenhader M, Rak M, Couplan E, Blondel M, et al. Mitochondrial ATP synthase disorders: molecular mechanisms and the quest for curative therapeutic approaches. Biochim Biophys Acta. 2009 Jan;1793(1):186-99. 16. Mayr JA, Havlickova V, Zimmermann F, Magler I, Kaplanova V, Jesina P, et al. Mitochondrial ATP synthase deficiency due to a mutation in the ATP5E gene for the F1 epsilon subunit. Hum Mol Genet. 2010 Sep 1;19(17):3430-9. 17. Majewski J, Schwartzentruber J, Lalonde E, Montpetit A, Jabado N. What can exome sequencing do for you? J Med Genet. 2011 Sep;48(9):580-9. 18. Viscomi C, Bottani E, Civiletto G, Cerutti R, Moggio M, Fagiolari G, et al. In vivo correction of COX deficiency by activation of the AMPK/PGC-1alpha axis. Cell Metab. 2011 Jul 6;14(1):80-90. 19. Yamada Y, Akita H, Kogure K, Kamiya H, Harashima H. Mitochondrial drug delivery and mitochondrial disease therapy--an approach to liposome-based delivery targeted to mitochondria. Mitochondrion. 2007 Feb-Apr;7(1-2):63-71.

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Summary

Metabolism is the buildup and degradation of nutrients by chemical reactions at the cellular level. The degradation of nutrients generates metabolites, which can be used for energy production in the form of ATP. ATP is needed for growth, development and maintaining the living state of the cell. The main source of ATP production is the oxidative phosphorylation

(OXPHOS) system. The OXPHOS system is situated in the mitochondria, double membrane cell organelles harboring their own DNA. This system consists of five multiprotein complexes

(complex I to V). The first four complexes form the respiratory chain and are responsible for electron transfer and proton pumping. Complex V uses the energy created by the proton electrochemical gradient to synthesize ATP from ADP. Part of the OXPHOS proteins (called subunits) is encoded by the mitochondrial (mt) DNA, part by the nuclear (n) DNA, except for complex II which is entirely encoded by the nDNA. mtDNA depends on enzymes encoded by the nDNA for replication, repair, transcription and translation. Moreover, a set of assembly factors, encoded by the nDNA, is needed to assemble the complexes. As a consequence, many gene defects can lead to mitochondrial dysfunction. Mitochondrial diseases are therefore common and heterogeneous genetic disorders. Because of this complexity, the diagnosis of these diseases is not straightforward. The first step in this process is to define the defect biochemically as precise as possible. This can guide the search for the underlying genetic defect. Therefore, the first aim in this thesis was to expand current tools for biochemical diagnostics in patients suspected for a mitochondrial disease.

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In chapter 3, two OXPHOS-deficient myogenic cell lines were established. It was shown that the complete biochemical toolbox, including substrate oxidation, ATP production rate and oxygen consumption measurements, as well as spectrophotometric enzyme analysis and

Taken together, biochemical assays can help in understanding the (patho)physiology of mitochondria and mitochondrial diseases and can guide the identification of the underlying genetic defect e.g. mutation analysis of complex V genes in case of an isolated complex V deficiency.

The human mitochondrial ATP synthase, or complex V is the last multi subunit OXPHOS complex. Complex V comprises 16 subunits. Five of them (α, β, γ, δ, and ε) form the F1 sector, situated in the mitochondrial matrix, the other subunits (a, c-ring, b, d, F6, OSCP, e, f, g, and A6L) belong to the Fo sector, embedded in the inner mitochondrial membrane. Only two subunits, a and A6L, are encoded by the mtDNA ATP6 and ATP8 genes, respectively.

The other subunits are nuclear encoded.

Unraveling the underlying genetic mutation and studying the functional impact of the defect in patients with a complex V deficiency was the second goal in this thesis. To this end, genetic mutation analysis was performed step-wise. First, the mtDNA was checked in all patients.

Second, the complex V genes with reported mutations were examined. Third, whole exome sequencing was applied.

As an introduction, chapter 4 reviewed the architecture, function and assembly of complex V.

The role of complex V di- and oligomerization and its relation with mitochondrial morphology was discussed. Finally, pathology related to complex V deficiency and current therapeutic strategies were highlighted.

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Summary

Chapter 5 described the first patient with a mutation in the mitochondrial encoded ATP8 gene

(step 1). He presented with apical hypertrophic cardiomyopathy and neuropathy. At the molecular level, we found an improper assembly and reduced activity of complex V.

In chapter 6, we described a patient with complex V deficiency due to a deletion in the

TMEM70 gene (step 2). We saw a fragmented mitochondrial network and swollen and irregularly shaped mitochondria with partial to complete loss of the cristae in fibroblasts of this patient, which could be completely restored by complementation of the TMEM70 genetic defect. Comparative genomics analysis predicted the topology of TMEM70 in the inner mitochondrial membrane, which could be confirmed by immunogold labeling experiments, and showed that the TMEM70 gene is not restricted to higher multi-cellular eukaryotes. This study demonstrated that the role of complex V in mitochondrial cristae morphology applies to human mitochondrial disease pathology.

In chapter 7, we performed whole exome sequencing to discover the mutation in two siblings with intractable epilepsy and complex V deficiency (step 3). We found a heterozygous mutation in ATP5A1, coding for complex V subunit α. This mutation was also found in the father, but because of the clear clinical picture and biochemical results, we decided to continue this study and checked the coding (c) DNA, revealing that the patients expressed the mutation homozygous while the father remained heterozygous. The mutation gave rise to a disturbed interaction of the α subunit with the β subunit of complex V, which interfered with the stability of the complex. This study demonstrated that biochemical diagnostics are indispensable for the correct interpretation of whole exome sequencing data.

In conclusion (chapter 8), mitochondrial biochemistry is needed to define mitochondrial dysfunction. It guides the search for the genetic defect. Mitochondrial biochemistry is also needed to confirm the pathogenicity of (a) found genetic variant(s). As both biochemical and genetic assays are continuously evolving, this powerful combination of approaches to unravel mitochondrial defects will undoubtedly enhance the physiopathological knowledge of mitochondrial diseases. This knowledge is crucial for the development of targeted therapies.

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Samenvatting (Dutch summary)

Samenvatting

Stofwisseling is de aanmaak en afbraak van bouwstoffen in ons lichaam op celniveau. Bij afbraak ontstaan metabolieten, die gebruikt worden als brandstof om het lichaam te voorzien van energie onder de vorm van ATP. ATP is nodig voor groei, ontwikkeling en voor het dagelijkse metabolisme. De belangrijkste bron van ATP productie is het oxidatieve fosforyleringssysteem (OXFOS). Het OXFOS ligt in de mitochondriën, dit zijn celorganellen met een dubbele membraan die hun eigen DNA bevatten. Het OXFOS systeem bestaat uit vijf complexen (complex I tot en met V), elk bestaande uit meerdere eiwitten, subunits genaamd. Complex I tot en met IV vormen de respiratoire keten en zorgen voor de transfer van elektronen naar O2 en het pompen van protonen. Complex V gebruikt de energie die door deze proton electrochemische gradiënt gecreëerd wordt om ADP om te zetten tot ATP.

Een deel van de OXFOS subunits wordt gecodeerd door het mitochondriële (mt) DNA, een deel door het nucleaire (n) DNA, met uitzondering van complex II wat volledig nucleair gecodeerd is. Het mtDNA hangt voor replicatie, herstel, transcriptie en translatie af van enzymen gecodeerd door het nDNA. De assemblage van de verschillende subunits tot complexen is eveneens afhankelijk van nucleair gecodeerde eiwitten. Dit betekent dat mutaties in veel verschillende genen kunnen leiden tot mitochondrieel disfunctioneren.

Mitochondriële ziekten komen daarom vrij frequent voor en zijn heterogeen van karakter.

Omwille van deze complexiteit is de diagnosestelling in deze groep van ziekten niet eenvoudig. De eerste stap in dit proces is de zo precies mogelijke biochemische karakterisatie van het defect. Dit is van belang bij het zoeken naar het verantwoordelijke gendefect. Het eerste doel in dit proefschrift was dan ook het uitbreiden van de huidig beschikbare mitochondrieel biochemische diagnostiek.

In hoofdstuk 2 werd een snelle en eenvoudige methode beschreven om zuurstofconsumptie te meten in verschillende celtypes na lysis van de celmembraan met digitonine. Er werd aangetoond dat dit een waardevolle methode is om mitochondriële disfunctie aan te tonen.

Door de zuurstofconsumptie te meten in verschillende celtypes of weefsels kan er een

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onderscheid gemaakt worden tussen primaire en secundaire mitochondriële afwijkingen. Een primaire mitochondriële aandoening zonder complexdeficiëntie kan aangetoond worden. Dit is van belang voor (erfelijkheids)advies aan de patiënt.

In hoofdstuk 3 werden twee OXFOS-deficiënte spiercellijnen geïsoleerd uit spierweefsel.

Verschillende mitochondrieel biochemische methodes konden succesvol worden toegepast op myoblasten en myotubes. Dit betekent dat mitochondriële aandoeningen kunnen geïdentificeerd en gekarakteriseerd worden in spiercellen. Dit geeft de mogelijkheid tot tal van toepassingen, zoals het bestuderen van OXFOS defecten die enkel tot expressie komen in spierweefsel, de studie van mitochondriële fysiologie in de levende cel, en het testen van doelgerichte therapeutische interventies.

Samenvattend kunnen biochemische testen helpen om inzicht te verwerven in de

(patho)fysiologie van mitochondriën en mitochondriële ziekten. Bovendien zijn ze zeer zinvol om het onderliggende genetische defect te helpen identificeren, bijvoorbeeld bij een geïsoleerde complex V deficiëntie kan de aandacht in een eerste tijd gericht worden op de bekende complex V genen.

Het humane mitochondriële ATP synthase, of complex V, is het vijfde OXFOS complex. Het bestaat uit 16 subunits. Vijf subunits (α, β, γ, δ, and ε) vormen het F1 deel. De andere subunits (a, c-ring, b, d, F6, OSCP, e, f, g, and A6L) vormen het Fo deel. Fo ligt verankerd in de mitochondriële binnenmembraan en zorgt voor de transfer van protonen naar F1, gelegen in de matrix, waar ATP wordt gesynthetiseerd. Slechts twee van de complex V subunits, subunit a en subunit A6L, worden gecodeerd door het mtDNA, namelijk respectievelijk door de ATP6 en ATP8 genen. De andere subunits worden nucleair gecodeerd.

De identificatie en functionele impact van de onderliggende mutatie bij patiënten met een geïsoleerde complex V deficiëntie ontrafelen was het tweede doel in dit proefschrift.

Hiervoor werd de genetische mutatie analyse stapsgewijs uitgevoerd. Eerst werd het mtDNA

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Samenvatting (Dutch summary) nagekeken (stap 1). Vervolgens werden alle “bekende” complex V genen onderzocht (stap

2). Tenslotte werd whole exome sequencing toegepast (stap 3).

Als introductie gaf hoofdstuk 4 een overzicht van de architectuur, functie en assemblage van complex V. De rol van di- en oligomerizatie van complex V in de mitochondriële morfologie werd besproken. Tenslotte werden de aan complex V gerelateerde pathologie en de huidige therapeutische denkpistes belicht.

In hoofdstuk vijf vond de eerste beschrijving plaats van een patiënt met een complex V deficiëntie ten gevolge van een mutatie in het mitochondrieel gecodeerde ATP8 gen (stap 1).

Klinisch werd er een apicaal hypertrofe cardiomyopathie en een neuropathie vastgesteld. Op moleculair niveau vonden we een verstoorde assemblage en een verminderde activiteit van complex V.

Hoofdstuk 6 behandelde een patiënt met een complex V deficiëntie ten gevolge van een deletie in het TMEM70 gen (stap 2). Bij deze en ook andere complex V deficiënte patiënten die een structurele afwijking in het complex hebben, werd in huidcellen een verstoorde mitochondriële morfologie gezien, gekenmerkt door gezwollen mitochondriën met een verlies van cristae. Het mitochondriële netwerk was gefragmenteerd. Deze afwijkingen konden hersteld worden door complementatie (toevoegen van het normale gen) van het TMEM70 gendefect. Comparatieve genoom analyse voorspelde dat TMEM70 ook voorkomt bij lagere eukaryoten en dat het eiwit gelegen is in de mitochondriële binnenmembraan. Deze lokalisatie kon bevestigd worden met behulp van immunogold labeling en electronenmicroscopie. Deze studie toonde aan dat de rol van complex V in de mitochondriële morfologie ook geldt voor humane complex V pathologie.

In hoofdstuk 7 werd whole exome sequencing toegepast bij twee broers met onbehandelbare epilepsie en complex V deficiëntie (stap 3). Er werd een heterozygote mutatie gevonden in het ATP5A1 gen, wat codeert voor de structurele complex V subunit α. Diezelfde mutatie werd ook gevonden bij vader, maar omdat het klinische beeld gecombineerd met de complex

V deficiëntie zo overtuigend waren bij de patiënten, werd besloten de mutatie verder te onderzoeken. En inderdaad, op coderend (c) DNA niveau werd de mutatie homozygoot

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teruggevonden bij beide kinderen en heterozygoot bij vader. Functioneel resulteerde dit defect in een verstoorde interactie van de α met de β subunit, wat aanleiding geeft tot instabiliteit van het complex. Deze studie toonde mooi aan dat een biochemische diagnose onontbeerlijk is voor de juiste interpretatie van whole exome sequencing data.

In conclusie (hoofdstuk 8) kan gesteld worden dat mitochondriële biochemie nodig is om mitochondriële disfunctie te definiëren. Het helpt de zoektocht naar het verantwoordelijke gendefect. Mitochondriële biochemie is eveneens nodig om de pathogeniciteit van (een) gevonden genetische variant(en) te bevestigen. Gezien zowel de biochemische als genetische technieken voortdurend in ontwikkeling zijn, zal deze sterke combinatie ongetwijfeld leiden tot het verder ontrafelen van oorzaken van mitochondriële ziekten. Dit draagt bij tot de fysiopathologische kennis van deze aandoeningen, wat cruciaal is voor het ontwikkelen van gerichte behandelingen.

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Dankwoord / Acknowledgements

Dankwoord/Acknowledgements

Dit proefschrift kan ik pas afronden nadat ik collega’s, vrienden en familie die er – direct of indirect – voor gezorgd hebben dat dit project is kunnen slagen, heb bedankt.

Mijn promotor, Prof. Dr. Jan Smeitink. Beste Jan, zes jaar geleden schonk je me je vertrouwen en bood je me de unieke kans om een klinisch fellowship metabole ziekten te combineren met promotieonderzoek. Ik heb je leren kennen als een uitzonderlijk gedreven dokter en wetenschapper met één doel: het beter maken van je patiënten. Het is een voorrecht om me in zo’n rijke omgeving te ontwikkelen en verder te groeien in “het vak”. Bedankt dat je er steeds was en dat je op het juiste moment het juiste zei.

Mijn copromotor, Dr. Richard Rodenburg. Beste Richard, zonder jou was dit boekje er niet gekomen. Je gaf me ruimte en rust om prettig te kunnen werken. Voor mij was dit erg belangrijk. Bedankt voor je continue betrokkenheid en voor de vele inspirerende ideeën, gesprekken en correcties van mijn manuscripten.

Leden van de manuscriptcommissie, Prof. Dr. Michèl Willemsen, Prof. Dr. Gert Vriend, en Prof. Dr. Frits Wijburg. Dank voor de beoordeling van mijn manuscript binnen een voor u allen reeds zeer druk werkschema.

Een speciale dank aan mijn collega’s van het lab Kindergeneeskunde en Neurologie (LKN), thans Laboratorium voor Genetische, Endocriene en Metabole ziekten (LGEM). Frans, Virginie, Jacqueline en Özlem van de kweekgroep, bedankt voor jullie inzet en flexibiliteit om af en toe in te springen en voor mijn cellen te zorgen, en eveneens voor jullie persoonlijke belangstelling. Collega’s van de spiergroep, in het bijzonder Antoon en Antonia, ik vond het heel aangenaam om met jullie samen te werken en verschillende hoofdstukken van dit boekje tot een goed einde te brengen. (Ex-) collega’s van de DNA-groep, in het bijzonder Heleen, Roel en Edwin, jullie leerden me de PCR en sequencing technieken. Jitske, Maaike en Marthe, jullie DNA-bijdrage aan verschillende artikelen heb ik erg gewaardeerd. Prof. Dr. Bert van den Heuvel, je aanmoedigingen tijdens gesprekjes op de gang hebben me gestimuleerd. Dr. Leo Nijtmans, je interesse in complex V en je nuttige suggesties aangaande mijn onderzoek waren inspirerend. Mariël, Bas, Jessica en Thatjana van de eiwitgroep, jullie stonden steeds klaar om te helpen bij alles wat met blotten te maken heeft. Een extra woordje richt ik graag aan Mariël. Beste Mariël, dankjewel voor het aanleren van en je continue hulp en raad bij het uitvoeren van Blue Native en andere blots. Je bent een geweldig research analist. Rutger, je immer heldere en kritische blik aangaande het onderzoek in bredere zin verrasten me keer op keer. Ook alle “menselijke” gesprekken zijn me bijgebleven. (Ex-) collega’s van “het hok”, Cindy en Monique, ik heb genoten van jullie aangename aanwezigheid en de vele leuke gesprekken. Cindy, dank voor het wegwijs maken binnen CorelDraw. Monique, het was heel fijn om een maatje te hebben voor alle babybabbels tijdens en na onze zwangerschap! Aan alle andere collega’s hierboven niet vernoemd, de warme en goede sfeer op het lab zorgden ervoor dat ik er met heel veel plezier heb gewerkt. Aan alle promoverende collega’s: veel succes!

Collega’s die hebben bijgedragen aan dit proefschrift: Prof. Dr. Martijn Huynen, Dr. Radek Szklarczyk en Dr. Sander Nabuurs, Prof. Dr. Martin Lammens en Lilian Eshuis, Dr. Jack Fransen en Mietske Wijers-Rouw, Dr. Werner Koopman en Marleen Forkink, Dr. Alexander

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Hoischen, en Dr. Herma Renkema, jullie bijdrage aan verschillende artikelen in dit manuscript was heel waardevol.

(Ex-) collega’s uit de kliniek, het “metabole team”: Eva, Maaike, Lonneke, Saskia, Marion, Anita, Annette, Leo, Marleen, Renske, Annemiek. Dank jullie wel voor de gezelligheid, de vele leermomenten, jullie begrip en mee-leven. Maaike, Marleen en Renske: heel leuk en bijzonder om samen voor de tweede keer zwanger te zijn!

Fokje, je bracht me rust in mijn hoofd. Ik denk dat je weet hoeveel dit voor me betekende.

Mijn paranimfen, Merei en Stefanie. Merei, als ervaren postdoc heb je me alle kneepjes in het lab geduldig bijgebracht. Gaandeweg deelden we ook lief en leed. Ik heb je erg gemist na meer dan drie jaar samenwerken, maar ben heel blij dat we er een goede vriendschap aan hebben overgehouden. Dank voor alles. Stefanie, wij delen onze studieachtergrond. Met dezelfde idealen zijn we in Nijmegen terecht gekomen. Ik bewonder je werkkracht en doorzettingsvermogen. Bovendien ben je een fantastische vriendin. De gezellige avonden met thee en Pim’s brachten telkens weer nieuwe energie. Bedankt dat je steeds voor me klaarstond, zowel praktisch als met goede raad.

Het thuisfront. Onze goede vrienden. Vicky en Iris, Katrien en Martine, jullie zijn er voor ons. Bedankt voor alle goede gesprekken over werk en leven, en de balans daartussen. Ik hecht heel veel waarde aan jullie vriendschap. Mijn schoonbroer. Jozef, het is gezellig om af en toe bij te kletsen aan tafel. Ik wens je het allerbeste toe. Mijn broer en zussen. Pieter, Katherine en Marilyn, ik ben heel blij dat jullie er zijn en me mijn werk af en toe laten relativeren. Tante, bedankt voor de gezelligheid en back-up voor Alexander in Nijmegen! Mijn schoonouders. Jullie staan elke dag klaar voor ons gezin, zowel voor verwachte als voor onverwachte situaties. Niets is teveel gevraagd. Zonder jullie steun was dit hele project niet haalbaar geweest. Heel welgemeend en van harte dank hiervoor. Mijn ouders. Jullie hebben me alle kansen gegeven. Dank voor de vrijheid van keuzes die jullie me steeds hebben gelaten. Dank voor jullie nooit aflatende vertrouwen en steun in alles wat ik ondernam. Tot slot, Paul en Alexander. Paul, ik bewonder je inzicht, je zorgzaamheid, je doorzettingsvermogen en je geduld. Je bent mijn steun en toeverlaat. Bij jou kan ik thuiskomen. Bedankt voor je jarenlange flexibiliteit. Alexander, je bent ons mooie en vrolijke mannetje. Bij jullie heb ik geluk mogen vinden. Samen uitkijken naar de komst van een broertje of zusje is het allermooiste.

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Curriculum Vitae

An Isabel Jonckheere werd geboren op 15 maart 1976 te Gent, België. Ze volgde de middelbare school in de Sint-Bavohumanioria te Gent, afdeling Latijn-Wiskunde van 1988 tot

1994. Van 1994 tot 2001 studeerde ze geneeskunde aan de Rijksuniversiteit te Gent, waar ze afstudeerde met grootste onderscheiding. Van 2001 tot 2006 was ze geneesheer- specialist in opleiding in de pediatrie in het Universitair Ziekenhuis te Gent (stagemeester

Prof. dr. D. Matthys). Deel van de opleiding vormden een jaar in het Centre Hospitalier

Régional te Namen (België) onder supervisie van Dr. M. Verghote, en een jaar in het

Wilhelmina Kinderziekenhuis te Utrecht, onder Prof. dr. J. Kimpen. Haar interesse in de aangeboren stofwisselingsziekten ontstond tijdens de opleiding tot kinderarts dankzij de samenwerking met Prof. dr. D. Carton en Prof. dr. L. De Meirleir in het Universitair

Ziekenhuis te Gent. Deze interesse werd bevestigd tijdens de stage bij Dr. T. de Koning en

Dr. G. Visser in het Wilhelmina Kinderziekenhuis te Utrecht. Na afronden van de opleiding kindergeneeskunde kon ze tussen 2006 en 2012 een fellowship metabole ziekten combineren met promotieonderzoek in het Universitair Medisch Centrum Sint-Radboud te

Nijmegen (promotor en opleider Prof. dr. J. Smeitink, copromotor Dr. R. Rodenburg). Het promotieonderzoek resulteerde in de artikelen welke gebundeld zijn in dit proefschrift.

Ze is getrouwd met Paul Van Herck en samen zijn ze de trotse ouders van een zoon,

Alexander (° januari 2011). In oktober 2012 verwachten ze hun tweede kind.

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List of publications

Mitochondrial ATP synthase: structure, function and pathology. Jonckheere AI, Smeitink JA, Rodenburg RJ. J Inherit Metab Dis 2012; 35(2):211-25.

Restoration of complex V deficiency caused by a novel deletion in the human TMEM70 gene normalizes mitochondrial morphology. Jonckheere AI, Huigsloot M, Lammens M, Jansen J, van den Heuvel LP, Spiekerkoetter U, von Kleist-Retzow J-C, Forkink M, Koopman WJH, Szklarczyk R, Huynen MA, Fransen JA, Smeitink JAM, Rodenburg RJT. Mitochondrion 2011; 11(6):954-63.

PKU: niet altijd ‘PKU’. Croonen EA., Jonckheere AI, Morava E. Tijdschrift voor Kindergeneeskunde 2010; 78(5):197-201.

A high-throughput assay to measure oxygen consumption in digitonin-permeabilized cells of patients with mitochondrial disorders. Jonckheere AI, Huigsloot M, Janssen AJM, Kappen AJH, Smeitink JAM, Rodenburg RJT. Clin Chem 2010; 56(3):424-31.

Biochemical and genetic analysis of 3-methylglutaconic aciduria type IV: a diagnostic strategy. Wortmann SB, Rodenburg RJ, Jonckheere A, de Vries MC, Huizing M, Heldt K, van den Heuvel LP, Wendel U, Kluijtmans LA, Engelke UF, Wevers RA, Smeitink JA, Morava E. Brain 2009; 132(Pt 1):136-46.

A novel mitochondrial ATP 8 (MT-ATP8) gene mutation in a patient with apical hypertrophic cardiomyopathy and neuropathy. Jonckheere AI, Hogeveen M, Nijtmans LGJ, van den Brand MAM, Janssen AJM, Diepstra JHS, van den Brandt FCA, van den Heuvel LP, Hol FA, Hofste TGJ, Kapusta L, Dillmann U, Shamdeen MG, Smeitink JAM, Rodenburg RJT. J Med Genet 2008; 45(3):129-33.

Normal pregnancy outcome in L-2-hydroxyglutaric aciduria. Jonckheere A, Carton D, Jaeken J, Gerlo E. J Inherit Metab Dis 2006; 29(4):588.

Successful liver transplantation for argininosuccinate lyase deficiency. Robberecht E, Maesen S, Jonckheere A, Van Biervliet S, Carton D. J Inherit Metab Dis 2006; 29(1):184-5.

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Appendix

Thesis series of the Institute for Genetic and Metabolic Disease Radboud University Medical Centre

1. Guillard, M. (2012). Biochemical and clinical investigations in the diagnosis of congenital disorders of glycosylation. Radboud University Nijmegen, Nijmegen, The Netherlands.

2. Markus, K. (2012). Strain or no strain. The application of two-dimensional strain echocardiography in children. Radboud University Nijmegen, Nijmegen, The Netherlands.

3. Jonckheere, A. (2012). Mitochondrial Medicine: assay development and application with special emphasis on human complex V. Radboud University Nijmegen, Nijmegen, The Netherlands.

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ITOCHONDRIAL M assay development and application application and development assay with special emphasis on human complex V human complex emphasis on with special

Mitochondrial Medicine: assay development and application with special emphasis on human complex V An I. Jonckheere