THE FUNCTIONAL SIGNIFICANCE of MITOCHONDRIAL SUPERCOMPLEXES in C. ELEGANS by WICHIT SUTHAMMARAK Submitted in Partial Fulfillment

Home , COX4I1, CYC1, Electron transport chain, NDUFA12, NDUFA5, NDUFA6

THE FUNCTIONAL SIGNIFICANCE OF

MITOCHONDRIAL SUPERCOMPLEXES in C. ELEGANS

WICHIT SUTHAMMARAK

Submitted in partial fulfillment of the requirements

For the degree of Doctor of Philosophy

Dissertation Advisor: Drs. Margaret M. Sedensky & Philip G. Morgan

Department of Genetics

CASE WESTERN RESERVE UNIVERSITY

January, 2011

CASE WESTERN RESERVE UNIVERSITY

SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

______

candidate for the ______degree *.

(signed)______(chair of the committee)

______

(date) ______

*We also certify that written approval has been obtained for any proprietary material contained therein.

Dedicated to my family, my teachers

and all of my beloved ones

for their love and support

ii ACKNOWLEDGEMENTS

My advanced academic journey began 5 years ago on the opposite side of the world. I traveled to the United States from Thailand in search of a better understanding of science so that one day I can return to my homeland and apply the knowledge and experience I have gained to improve the lives of those affected by sickness and disease yet unanswered by science. Ultimately, I hoped to make the academic transition into the scholarly community by proving myself through scientific research and understanding so that I can make a meaningful contribution to both the scientific and medical communities. The following dissertation would not have been possible without the help, support, and guidance of a lot of people both near and far. I wish to thank all who have aided me in one way or another on this long yet rewarding journey.

My sincerest thanks and appreciation goes to my advisors Philip Morgan and Margaret Sedensky. By introduction of Dr. Chanin Limwongse and Dr.

Shawn McCandless, I was given the opportunity to work with two intelligent and gracious individuals. They treated me with great respect and provided me with a nurturing and open work environment where I could speak my mind and share my ideas. They have also instilled in me the qualities of being a good scientist and thinker. Without their insightful scientific advice, guidance and support the completion of this project would not be possible. I would also like to thank them for having provided me with many opportunities to present my work at prominent conferences and speaking events. I am extremely fortunate to have been

iii blessed with the opportunity to have known and work with these two great

individuals.

I would like to thank my committee members Dr. Charles Hoppel, Dr.

Peter Harte, Dr. Shawn McCandless and Dr. Hua Lou for their guidance over the

years. They have offered me invaluable advice that challenged my way of

thinking. Their support and guidance has shaped me to become a better scientist

and for that I am truly grateful.

My appreciation also extends to my lab colleagues, both past and present.

Thanks to Dr. Marni Falk for her scientific discussion and project planning. Dr.

Bernhard Kayser for giving me helpful technical advice, sharing his tremendous

knowledge about mitochondria, and for showing me that mitochondria really

respire. Dr. Louise Steel for helping me correct the English in many of my

writings. Julie Rosenjack for her sense of humor and teaching me everyday

English. Thanks to Judith Preston for teaching me many lab techniques,

especially how to isolate intact mitochondria from worms, and for the delicious

cookies. Toni Portman, Katrina Elsaesser, Beatrice Predoi, Qiao-yun Jiang,

Melinda Hubbard, Yu-Ying Yang, Ching-Chun Yang and Vinod Singaram for all

your help in the lab.

The collaborative efforts by these individuals are also much appreciated.

Hiral Patel and David Kehres from Dr. Hoppel’s laboratory for their excellent

technical assistance. Dr. Michael Kinter and Belinda Willard from Cleveland for their input and advice, and Dr. Andrew Bauman and colleagues from Dr. Eugene

Kolker’s laboratory in Seattle for the mass spectrometric analysis of my BNGs.

iv On a personal level, special thanks goes to my family for their relentless love and support. Since youth, my parents have stressed the importance of getting a good education to my siblings and I. Although my dad is no long with us, I know he would be proud to see that I have lived up to his words. For my mom, I express special thanks and admiration for a woman who devoted most of her life to the care of her children and family. Her unwavering love and support is beyond anything I could ever express or describe in words. Above all, I am truly grateful for the greatest gift my parents have ever given me—life. I love you both wholeheartedly mom and dad.

I would like to thank Dr. Bundhit Tantiwongosi, my best friend, for offering his help and guidance when I first moved to the United States. My transition from

Thailand to the United States would not have been as smooth if it weren’t for all his help. Thank you for all your help, guidance, and support through all these years.

There are a few more people that I would like to thank for their contributions to my life. First is Ms. Ratree Rengsirikul, my favorite high school biology teacher. Ms. Ratree instilled in me the fundamental traits of a good scientist—curiosity and observance. She made me realize that if we want to see nature the way it really is, we have to look at it through science. Second is Ms.

Srijit Tungpong, my high school mathematics teacher. She treated me like her own son and helped me overcome personal difficulties and, until this day, still offers her love and support to me. Third is Dr. Patcharee Lertrit, my supervisor and the person who introduced me to the world of mitochondrial research. Last

v but not least, Ms. Rungnapa Sriwilai (P Aied) an excellent technician in Dr.

Lertrit’s laboratory who taught me good laboratory techniques.

Finally, I would like to thank all of my friends both in Thailand and the

U.S. They have supported me through both good and bad times and because of them, my life is more vibrant and meaningful. I am thankful their presence in my life—past, present, and future.

vi TABLE OF CONTENTS

Dedication………………………………………………………………………………..ii

Acknowledgements……………………………………………………………………..iii

Table of Contents…………………………………………………………………....…vii

List of Figures and Tables……………………………………………………………xiv

Abstract……………………………………………………………………………...…xvii

Chapter 1: Introduction 1

1.1. Mitochondrial respiratory chain complexes (MRC)………………...2

1.1.1. Structure and function...…………………………………………2

• Complex I (NADH-ubiquinone oxidoreductase)

• Complex II (succinate-ubiquinone oxidoreductase)

• Complex III (Ubiquinone-cytochrome c oxidoreductase)

• Complex IV (cytochrome c oxidase)

• Complex V (ATP synthase, F1F0-ATPase)

1.1.2. Oxidative phosphorylation………………………….………….10

1.2. Structural organization of respiratory chain complexes...... 11

1.2.1. Fluid-state model……………………………………………….11

• Lateral diffusion-random collision

• Ubiquinone pool behavior

1.2.2. Solid-state model……………………………………………….13

• Structural identification of respiratory supercomplex

• Metabolic flux control analysis

vii • Respiring mitochondrial supercomplexes

1.3 Implications of the mitochondrial supercomplex……………….…17

1.3.1. Functional advantage of supercomplexes…………………..18

• Possible mechanisms of enhanced respiratory chain

function by supercomplexes

• Supercomplex organization may prevent excessive

mitochondrial reactive oxygen species (ROS) production

1.3.2. Assembly and stability of complex I………………………….20

1.3.3. Decline of mitochondrial supercomplexes in impaired

respiratory chain function in mice and humans…………….21

• Supercomplex instability due to cardiolipin deficiency

• BCS1L mutations disrupt complex III and supercomplex

formation, causing Bjornstad and GRACILE syndromes

• Cytochrome b mutations and combined complex I/III

deficiency

• Complex IV defect and combined complex I/IV

deficiency

• Supercomplex dysfunction in heart failure

1.4. C. elegans as a model to study the function of mitochondrial

supercomplexes…………………………………………………..…....26

1.4.1. isp-1(qm150)…….……………………………………………...27

1.4.2. isp-1(qm150);ctb-1(qm189)..………………………………….28

1.4.3. Complex IV deficient C. elegans…..…………….……………29

viii 1.5. Conclusion……………………………………………………………….30

1.6. Specific Aims……………………………………………………..……..31

1.7. Figures…………………………………………...……………………….33

1.8. References……………………………………………………………….46

Chapter 2: The effects of COX IV- and COX Va-RNAi knockdowns on______

whole animal phenotypes, supercomplex function and respiratory______chain___ function in C. elegans ______57

2.1. Introduction……………………………………………………………...58

2.2. Results………………………………………………….…………………61

2.2.1. The effects of COX IV- and COX Va-RNAi knockdowns

on whole animal phenotypes……..…………..….……..…….61

2.2.2. Efficacy of RNAi knockdown…………………………..………61

2.2.3. The effects of COX IV- and COX Va-RNAi knockdowns

on respiratory chain function.……………....….....……...... …61

• Oxidative phosphorylation capacity

• Electron transport chain function

2.2.4. The effects of COX IV- and COX Va-RNAi knockdowns

on supercomplex formation……….…………….…………...63

• Supercomplex organization in C. elegans mitochondria

• COX IV and COX Va knockdowns altered

supercomplex profile

• Complex I formation was unchanged in COX IV and

ix COX Va worms

2.3. Discussion………………………………………………………………..66

2.4. Conclusion…………………………………………………………….…75

2.5. Figures and tables………………...…………...………………….……76

2.6. References……………………………………………………………….91

Chapter 3: Mutations in mitochondrial complex III uniquely affect______

complex I in C. elegans ______95

3.1. Introduction……………………………………………………………...96

3.2. Results…………………………………………………….………………99

3.2.1. Phenotypes of complex III mutants- delayed embryonic

development and egg laying.……………………………...... 99

3.2.2. Effect of complex III mutations on integrated

mitochondrial respiration…...... 100

3.2.3. Respiratory enzyme complex activity in complex III

mutants...... 100

3.2.4. Amounts of fully assembled respiratory complexes

in complex III mutants...... ….101

3.2.5. The effects of isp-1 on supercomplexes...... 102

3.2.6. Supercomplex profiles in complex III mutants...... 102

3.2.7. Complex I function from electroeluted supercomplexes....103

3.2.8. The stability of supercomplexes...... 104

x 3.2.9. The effect of sodium cholate on mitochondrial

supercomplexes……………………...………………………104

3.3. Discussion……………………………………………………………...106

3.4. Conclusion…………………………………………………..……...….114

3.5. Figures and tables...…………………………...……………………...115

3.6. References……………………………………………………………...131

Chapter 4: Discussion and future direction 135

4.1. Research summary……………………………………………………136

4.2. Discussion and future direction…………………………………….137

4.2.1. Supercomplex assembly requires proper quality and

quantity of each complex component….………………..…137

4.2.2. Supercomplex I:III versus I:III:IV…….……………………...141

• Supercomplex I:III:IV as a major contributor of

mitochondrial energetics

• Supercomplex I:III:IV, not complexes I, IIII and

IV, matters

4.2.3. Stoichiometric association of complexes I, III and IV in

supercomplexes…………….………………….……...... 145

4.2.4. Roles of supercomplex in mitochondrial OXPHOS

disorders and aging………………………………….…...... 146

• Combined respiratory complex deficiency

xi • Wide varieties of affected tissues in OXPHOS

disorders

• Mitochondrial supercomplexes and aging

4.3. Conclusion...... 150

4.4. Figures...... 151

4.5. References……………………………………………………………...155

Chapter 5: Material and methods 158

5.1. Worm cultures………………………………………..………………..159

5.1.1. Animals and normal growing condition...... 159

5.1.2. RNAi knockdown...... 159

5.2. Phenotypic study...... 160

5.2.1. Lifespan study...... 160

5.2.2. Embryonic developmental study...... 160

5.2.3. Egg laying assay...... 160

5.2.4. Anesthetic sensitivity...... 161

5.3. RNA extraction and quantitative reverse transcription PCR.....161

5.4. Mitochondrial functional assays…………………………..…….....162

5.4.1. Worm collection...... 162

5.4.2. Mitochondrial isolation...... 163

5.4.3. Oxidative phosphorylation assay...... 164

5.4.4. Electron transport chain assay...... 165

5.5. Mitochondrial supercomplex studies…………………………...... 166

xii 5.5.1. One-dimensional blue native gel electrophoresis

(1D BNG), in-gel activity assay (IGA) and

electroelution of supercomplexes...... 166

5.5.2. Two-dimensional blue native/high resolution clear

native gel electrophoresis (2D BN/hrCNE)...... 167

5.5.3 SDS-Western blot...... 167

5.5.4. Native-Western blot...... 168

5.5.5. Quantification of proteins/ protein complexes...... 168

5.5.6. Mass spectrometry...... 169

5.6. Statistical analysis………………………………………………….....170

5.7. References……………………………………………………………...171

Addendum: Copyright permissions for reuse of illustrations______in a dissertation______173

xiii List of Figures

Figure 1. The integrated function of the mitochondrial respiratory chain...…33

Figure 2. The functional modules of complex I………………………………...34

Figure 3. Structure of the membranous arm of E. coli complex I and

proposed model of proton translocation by complex I……………..35

Figure 4. Trimer association of E. coli complex II and its redox centers…....36

Figure 5. Q cycle of complex III………………………………………………….37

Figure 6. The catalytic core and redox centers of complex IV……………….39

Figure 7. ATP synthesis of complex V………………………………………….40

Figure 8. The structure of supercomplexes I1III2IV1 and the putative

binding sites of ubiquinone and cytochrome c……………...... 41

Figure 9. Simplified schematic explanation of metabolic flux control

analysis…………….………………………..…………………………43

Figure 10. A current model of complex I assembly pathway………………….44

Figure 11. Respiratory Supercomplexes in C. elegans…….………………….45

Figure 12. Deterioration of mitochondrial respiration in the RNAi-

knockdown worms………………………………………………..…...76

Figure 13. The effects of COX knockdown on electron transport chain

assays…………………...………………………..…….………………78

Figure 14. The organization of respiratory complexes in C. elegans…………80

Figure 15. Blue native electrophoresis and the in-gel activity assay (IGA)

of mitochondria from complex IV-deficient worms…...…………….81

xiv Figure 16. Complex IV disruption alters the ratio between supercomplex

I:III and I:III:IV but not the total amount of complex I…….………...83

Figure 17. Steady state level of the NUO-2 subunit of complex I……………..85

Figure 18. Number of eggs per worm per day in complex III mutants

and wild type………………………………………………………….115

Figure 19. Integrated mitochondrial function of complex III mutants

and wild types………………………………………………...... 116

Figure 20. Respiratory chain enzymatic activities of complex III mutants

and wild type………………………………………………………….117

Figure 21. Amounts of respiratory enzyme complexes……………...... 118

Figure 22. The stead-state level of respiratory enzyme subunits…………....120

Figure 23. Digitonin-based BNGs in complex III mutants…………………….121

Figure 24. Complex I activity in supercomplex I:III and I:III:IV……………….123

Figure 25. The ability of 2D-BN/hrCNE to dissociate wild-type and

complex III mutant supercomplexes………………………………..124

Figure 26. The effect of sodium cholate on mitochondrial

supercomplexes……………………………………………………...126

Figure 27. The yeast two-hybrid assay for detecting protein-protein

interactions...... 151

Figure 28. Comparison of the 3D map of the bovine heart supercomplex

I1III2IV1 with those of the individual complexes...... 153

xv List of Tables

Table 1. Phenotypical study of COX IV and COX Va knockdown

worms…………………………………………………………………...86

Table 2. Respiratory chain proteins identified by mass spectrometry in

each protein band in digitonin-based BNGs……………………...... 87

Table 3. Respiratory chain proteins identified by mass spectrometry in

each protein band in Triton X-100-based BNGs……..………….....89

Table 4. Development of complex III mutants and N2 at 20° C…………...128

Table 5. Relative expression of NUO-2, cytochrome c and COX I in

complex III mutants when ANT is used as a loading control…….129

Table 6. Proteomic analysis of ISP-A and ISP-B……………………………130

xvi The Functional Significance of

Mitochondrial Supercomplexes in C. elegans

Abstract

WICHIT SUTHAMMARAK

Recent evidence has indicated that respiratory enzyme complexes in the mitochondrial respiratory chain (MRC) are interconnected via supramolecular assemblies commonly called respiratory supercomplexes. Interdependence between complex components in a supercomplex has been regarded to play a role in respiratory chain dysfunction particularly in combined complex deficiencies of the electron transport chain (ETC). Furthermore, decline in supercomplexes has been found to be associated with some pathologies in humans such as Barth syndrome, heart failure, etc. However, the mechanism by which supercomplexes underlie the disease pathology has not been fully elaborated.

I take advantage of the nematode C. elegans to uncover the ill-defined aspects of supercomplexes. My study focuses on the complex III mutants, isp-1, isp-1;ctb-1 and ctb-1, and complex IV deficient RNAi-knockdowns. My work further investigates the effects of the mutation, which include:

xvii 1) Rate of embryonic development and egg laying.

2) Anesthetic sensitivity.

3) Mitochondrial respiration.

4) Enzymatic rate of respiratory complexes and supercomplexes.

5) The formation of respiratory complexes and supercomplexes.

6) Integrity of supercomplexes.

My study employs comprehensive mitochondrial functional assays, molecular biology techniques and genetic manipulations in C. elegans to address

the questions. I demonstrate that complex III is necessary for supercomplex

assembly/stability. Complex I assembly is, therefore, completed upon

supercomplex formation. The efficiency of ETC function is dictated by the

supercomplex profile and its integrity. Interestingly, complexes III and IV

influence complex I function through supercomplex structure in an allosteric

manner. These results provide new insights into the functional significance of

supercomplexes as well as the mechanism of combined complex deficiency.

xviii

Chapter 1

Introduction and background

1 1.1 Mitochondrial respiratory chain (MRC)

Every living organism harvests energy from food and converts it into ATP, the main energy currency of the cells. Eukaryotes posses specialized double membrane organelles, mitochondria which, under normal circumstances, faithfully produce ATP to supply cells throughout their life. Prokaryotes are more primitive, yet produce ATP by molecular machinery residing in an energy- transducing membrane which is a part of the plasma membrane. Regardless of location, the respiratory chain is composed of 5 multi-subunit complexes, complex I-V, and 2 mobile electron carriers, ubiquinone and cytochrome c.

Electrons from NADH and succinate (from other catabolic reactions) enter the respiratory chain at complex I and complex II, respectively. They are then transferred through a series of redox centers in complex III and IV until reaching the last electron acceptor, molecular oxygen (O2). Complexes I, III and IV couple electron transfer with the translocation of protons from the mitochondrial matrix to the intermembrane space, generating a proton gradient across the membrane.

Complex V utilizes the transmembrane proton gradient to produce ATP. This entire coupled process is called oxidative phosphorylation (Mitchell & Moyle

1967) (Fig. 1).

The structure and function of each respiratory complex as well as oxidative phosphorylation is discussed in more detail in 1.1.1. and 1.1.2.

1.1.1. Structure and function

• Complex I (NADH-ubiquinone oxidoreductase)

2 Complex I is the largest, yet the least understood, component of the respiratory chain. It couples the transfer of electrons from NADH to ubiquinone with the translocation of protons across the inner mitochondrial membrane. In

most prokaryotes, complex I weighs around 550 kDa and consists of 14 subunits

that are conserved in all organisms that possess a complex I. (Friedrich 2001).

These core subunits represent the minimal structural subunits necessary for

complex I function (Brandt 2006). The 14 complex I subunits of E. coli are named

NuoA to NuoN, where Nuo refers to NADH-ubiquinone oxidoreductase. In

contrast, the eukaryotic complex I has a molecular mass around 1,000 kDa and

can contain over 45 subunits. The functions of many of these additional proteins

remain largely uncharacterized (Brandt 2006; Lazarou et al. 2009). In bovine

complex I, most subunits are named according to their molecular weights (kDa),

whereas the human counterparts adopt another nomenclature. The human subunits of complex I that are encoded by mitochondrial DNA are termed ND, which refers to NADH dehydrogenase, followed by the subunit number (ND1-

ND6 and ND4L). In humans nuclear-encoded subunits of complex I are termed

NDU, which refers to NADH dehydrogenase ubiquinone, followed by a brief designation that refers to its predicted function, such as FA; subcomplex α, FB;

subcomplex β, FC; unidentified subcomplex, FS; iron-sulfer associated peptides

and FV; flavoprotein associated peptide (Lazarou et al. 2009).

Crystallographic and electron microscopic studies reveal that complex I is

L-shaped, with a membranous arm and a matrix arm (Yagi & Matsuno-Yagi

2003; Morgan & Sazanov 2008; Clason et al. 2010). Complex I can be divided

3 into 3 functional modules (Fig. 2). The input module (Brandt 2006), or N module,

oxidizes NADH and then passes electrons via flavin mononucleotide (FMN)

along a series of iron-sulfur redox centers. The output module (Brandt 2006), or

Q module, receives electrons from the N module through the iron-sulfur redox centers and transfers them to ubiquinone. Notably, both N and Q modules make up the matrix arm of complex I. The P module comprises the membranous arm and is believed to be largely involved in proton pumping. Transfer of 2 electrons from NADH is coupled to the translocation of 4 protons from the mitochondrial matrix to the intermembrane space. This H+/e- stoichiometry is also conserved

across species (Brandt 2006; Sazanov 2007).

The most recent study from Efremov et al. (Efremov et al. 2010) reveals

the unprecedented details of the architecture of respiratory complex I from E. coli

and T. thermophilus. Crystallographic data show that the 3 largest hydrophobic

subunits of complex I, subunits NuoL/M/N, in the membranous arm have the

antiporter-like structure (ion channel-like structure), and are therefore likely to

participate in proton pumping. Intriguingly, the NuoL subunit was found to have a

unique amphipathic α-helix spanning almost the entire length of the membranous

arm (Fig. 3). Efremov et al. suggested that the coupling mechanism between

electron transport and proton pumping should then be conformational-driven

(indirect) (Friedrich 2001; Yagi & Matsuno-Yagi 2003; Brandt 2006) rather than

redox-driven (direct) (Albracht & Hedderich 2000; Hellwig et al. 2000). As

previously described by others (Gondal & Anderson 1985; Belogrudov & Hatefi

1994; Mamedova et al. 2004), the complex I reduction that takes place in the

4 matrix arm changes the conformation of the joint region between the matrix arm

and the membranous arm. Efremov et al. then proposed that this change is

further conveyed to the membrane spanning helix of the NuoL subunit. This

movement could then trigger a conformational change of the antiporter-like

subunits NuoL/M/N and allow proton translocation across the inner mitochondrial

membrane (Efremov et al. 2010) (Fig. 3).

The regulation of the coupling mechanism (the coupling of NADH

oxidation to proton translocation) of complex I seems to be reciprocal, since

proton pumping can affect the reduction of complex I. Studies in E. coli have

shown that mutations in the Nuo subunits (Kao et al. 2004; Kao et al. 2005a; Kao

et al. 2005b), including the NuoM subunit (one of the proton pumping subunits)

(Torres-Bacete et al. 2007), decreased the rate of complex I reduction (as

determined by NADH-ubiquinone reductase activity) without impairing the

integrity of the matrix arm.

• Complex II (Succinate-ubiquinone oxidoreductase)

Complex II oxidizes succinate to fumarate and reduces ubiquinone.

Hence, it adopts another name, succinate dehydrogenase (SDH). The enzyme is composed of 4 subunits: a flavoprotein (SdhA), an iron-sulfur protein (SdhB) and two small membrane-anchor subunits (SdhC and SdhD). All are encoded by the nuclear genes in eukaryotes, while in prokaryotes all 4 genes are in the sdhCDAB operon (Bachmann 1990; Blattner et al. 1997).

A structural study of complex II in E.coli (Yankovskaya et al. 2003) demonstrated that complex II is associated as a trimer (~360 kDa) (Fig. 4).

5 Detailed experiments also suggested that this trimer association is physiologic; electron transfer independently occurs within each monomer. The hydrophilic domain (SdhA and SdhB) is responsible for the oxidation of succinate. The hydrophobic domain (SdhC and SdhD) provides the ubiquinone-binding site.

Interestingly, heme b (cytochrome b565), which is present in the hydrophobic domain, is not essential for ubiquinone reduction (Maklashina et al. 2001).

Complex II does not contribute to the transmembrane proton gradient; complex II has the sole function of transfer of electrons to ubiquinone.

• Complex III (Ubiquinol-cytochrome c oxidoreductase)

Complex III couples the transfer of electrons from ubiquinol (QH2) to

cytochrome c with proton translocation across the inner mitochondrial

membrane, with a 2e-/4H+ stoichiometry. It is composed of 11 subunits in eukaryotes, with fewer in prokaryotes (Schagger et al. 1986; Schagger et al.

1995). Cytochrome b is the only complex III subunit that is encoded by mitochondrial DNA. The catalytic core of complex III is composed of 3 conserved subunits: cytochrome b, the Rieske protein (iron sulfur protein or ISP) and cytochrome c1 (Robertson et al. 1993). Physiologically, complex III assembles

into a dimer; each monomer has a molecular mass of ~ 240 kDa (Iwata et al.

1998).

The electron transfer mechanism of complex III is also called the Q-cycle

(Trumpower 1990; Crofts 2004) (Fig. 5). The unique characteristic of the Q-cycle

is a bifurcation of the electron pathway upon the oxidation of QH2, which takes place at the QP site (the positive site or the intermembrane side). Upon QH2

6 oxidation, one electron is transferred to ISP ([2Fe-2S] cluster)  cytochrome c1

 cytochrome c, and is defined as a “high-potential chain”. The resulting

- semiquinone (SQ (or UQ⋅ ): an unstable ubiquinone intermediate that is partially

reduced with 1 electron) at the QP site then transfers the second electron to

cytochrome b by heme bL  heme bH, which is defined as a “low-potential chain”. Heme bH reduces ubiquinone to the SQ intermediate at the quinone reduction (QN) site (the negative site or the matrix side). SQ formed at the QN site

is stabilized until the second electron from another QH2 oxidation at the QP site

arrives. Therefore, under normal physiologic state, two turnovers of QH2

oxidation at the QP site are required to reduce a QN site-quinone to QH2.

Cytochrome c, however, carries one electron at a time.

Complex III has the great propensity to produce reaction oxygen species

(ROS) due to the bifurcated electron transfer at the QP site (Cape et al. 2005;

Forquer et al. 2006). For instance, the partial inhibition of the QN site (heme bH) by antimycin A or the presence of rhodoquinol (a modified ubiquinone, where the methoxyl group in position 3 is replaced by an amino group) can result in an unfavorable state of the low potential chain. The SQ intermediates at both QP

and QN sites accumulate, and can reduce O2 to superoxide (Muller et al. 2002;

Sun & Trumpower 2003; Cape et al. 2005; Forquer et al. 2006).

• Complex IV (Cytochrome c oxidase)

Complex IV is the terminal enzyme in the respiratory chain, present in

eukaryotes and aerobic bacteria. It couples the transfer of electrons from

cytochrome c to molecular oxygen (O2) with proton translocation across the inner

7 mitochondrial membrane (Malmstrom 1990). In most eukaryotes complex IV is

composed of 10 nuclear and 3 mitochondrially encoded subunits (Capaldi 1990).

The latter three subunits form the catalytic core of the enzyme. The monomer of

complex IV has a molecular mass of ~ 204 kDa (Tsukihara et al. 1996) but usually exists as a dimer. Subunit VIa is responsible for the dimerization of complex IV (Tsukihara et al. 1996), which takes place under physiological conditions.

Complex IV transfers 1 electron at a time. The electron pathway of

cytochrome c oxidase can be simplified as: cytochrome c  CuA  heme a 

[heme a3-CuB]  O2 (Fig. 6). CuA, the electron entry site in cytochrome c oxidase

(Kobayashi et al. 1989; Hill 1991; Zhen et al. 2002), resides in subunit II. The

other 2 redox centers, heme a and [heme a3-CuB], are found in subunit I, the

largest and the most conserved subunit (Tsukihara et al. 1996). While the proton-

pumping pathway of complex IV is still inconclusive, it is generally accepted that

the stoichiometry of proton translocation is 1e-/1H+, i.e. 4 protons are

translocated per every O2 consumed (Wikstrom 1977; Wikstrom & Krab 1979;

Wikstrom 1988).

Cytochrome c oxidase controls cellular energy metabolism via allosteric

regulation of the enzyme (Kadenbach et al. 1997; Napiwotzki & Kadenbach

1998). The extramitochondrial ATP/ADP-ratio is the allosteric regulator of subunit

IV. During low cellular energy demand, when extramitochondrial ATP/ADP ratio

is high, the binding of ATP to the cytosolic domain of subunit IV leads to an

8 increased Km for cytochrome c, resulting in decreased cytochrome c oxidase activity (Napiwotzki & Kadenbach 1998).

• Complex V (ATP synthase)

Complex V does not contribute to the transmembrane proton gradient but utilizes it to mediate the phosphorylation of ADP to ATP. Complex V can be visualized under electron microscopy as distinct spherical knobs projecting from the matrix side of the inner mitochondrial membrane into the mitochondrial matrix

(Minauro-Sanmiguel et al. 2005; Dudkina et al. 2006). Complex V has 2 parts, an integral membrane F0, which controls proton translocation down the gradient,

and a water-soluble F1 which is a catalytic site for ATP synthesis (Boyer 1997;

Yoshida et al. 2001). The F0 domain is made of subunit a and a ring of 9-12 c

subunit (c-ring), forming a base structure of ATP synthase. Subunit γ provides a

central stalk of the F1 domain, attaching to the c-ring of the F0 domain. Subunits

OSCP and b (subunits δ and b2 in prokaryotes) also provide the peripheral

attachment between the F1 and F0 domains (Fig. 7).

The translocation of protons from the intermembrane space via channels

located at the interface between the c-ring and subunit a within the F0 domain

drives the synchronous rotation of the F0 and F1 domains (Cross 2004; Adachi et

al. 2007). The rotation of F0F1 occurs in steps of 120°; each facilitates phosphorylation of ADP to ATP at the catalytic head α3β3-hexamer of the F1

domain (Adachi et al. 2000; Yasuda et al. 2001; Cross 2004) (Fig. 7).

Translocation of 9-12 protons from the intermembrane space causes 1 complete

rotation of F1F0-ATP synthase, resulting in the production of 3 ATP molecules.

9 The number of c monomers per the c-ring determines the number of protons translocated per rotation (Junge & Nelson 2005; Dimroth et al. 2006).

1.1.2. Oxidative phosphorylation

The central concept of oxidative phosphorylation (Mitchell & Moyle 1967) is that complex I to complex IV of the respiratory chain catalyze the downhill

(passive) transfer of electrons from NADH or succinate to O2 (oxidative event)

and use the released energy to generate a transmembrane proton gradient.

Complex V then utilizes a gradient of protons to synthesize ATP (phosphorylation

event). The efficiency of oxidative phosphorylation is determined by the amount

of ATP synthesized per oxygen consumed during oxidative phosphorylation, a

P/O ratio. For complex I-dependent respiration, the oxidation of 1 molecule of

NADH yields 2 electrons, which are transferred through the respiratory chain to

- + reduce 1/2 molecule of O2. As the result, 10 protons are pumped (4H at

complex I, 4H+ at complex III and 2H+ at complex IV) into the intermembrane

space of a eukaryotic mitochondrion. Complex V requires a 120° rotation of F1F0

(Yasuda et al. 2001; Adachi et al. 2007) for each molecule of ATP that is synthesized. This requires the translocation of ~3-4 H+, depending on the number

of c subunits in the c-ring (Junge & Nelson 2005; Dimroth et al. 2006). Therefore, the maximal value of P/O ratio of complex I-dependent respiration is 2.5-3. For complex II-dependent respiration, succinate/O reaction results in 6H+/2e-

translocation, which brings ATP/O ratio to 1.5-2. However, these maximal P/O

values can only be sustained under the saturation of inorganic phosphate (Pi) and ADP in the in vivo situation (Devin & Rigoulet 2007). The actual P/O ratio

10 can vary according to the degree of proton leak, which happens in the presence

of membrane uncouplers (Ouhabi et al. 1991; Fitton et al. 1994), and the

functional state of mitochondria.

1.2 Structural organization of respiratory complexes

The organization of the respiratory complexes in the inner mitochondrial

membrane has been a controversial subject since the respiratory complexes

were successfully purified decades ago (Hatefi et al. 1962a; Hatefi et al. 1962c;

Hatefi et al. 1962b). Currently, there are 2 extreme models describing the organization of respiratory complex, the fluid-state model and the solid-state model.

1.2.1. Fluid-state model

Hackenbrock et al. described the arrangement of the respiratory complexes in a fluid-state model (Hackenbrock et al. 1986). They envisioned that the respiratory complexes are randomly dispersed in the phospholipids bilayer membrane where they can move freely by lateral diffusion. Electron transfer between complexes is facilitated by the movement of ubiquinone, connecting complex I/II and complex III, and cytochrome c, connecting complex III and IV.

This view had been widely accepted and was in fact a textbook view of the organization of the respiratory complexes for decades. The fluid-state model was based upon several key studies as described in the following:

• Lateral diffusion and random collision of respiratory chain components

11 Space is not a limiting factor for lateral diffusion since 60-70% of the inner mitochondrial membrane total area is unoccupied by proteins (Hackenbrock et al.

1976; Sowers & Hackenbrock 1981). This unoccupied space allows the lateral diffusion of the respiratory chain components to occur freely. A random distribution of the respiratory chain components was also supported by freeze- fracture electron microscopy and antibody studies (Segrest et al. 1974; Hochli &

Hackenbrock 1977; Hochli et al. 1985). Fluorescence recovery after photobleaching (FRAP) experiments convincingly demonstrated that cytochrome c (Hochman et al. 1982; Vanderkooi et al. 1985), ubiquinone (Lenaz et al. 1985;

Lenaz & Fato 1986), complex III (Gupte et al. 1984; Hochli et al. 1985), complex

IV (Hochli et al. 1985; Hochman et al. 1985) as well as membrane phospholipids

(Hochman et al. 1985; Vanderkooi et al. 1985) are mobile entities in the membrane. In addition, while complex III and IV diffuse in 2 dimensions, cytochrome c, which is not integral to the membrane, was demonstrated to diffuse predominantly in 3 dimensions in the membrane at physiological ionic strength (Gupte et al. 1984). The diffusion of cytochrome c then ensures electron transfer between complex III and IV. More importantly, the rates of diffusion of the respiratory components in the membrane was shown to not only directly affect the rate of electron transfer but also to be rate limiting (Hackenbrock et al.

1986).

• Ubiquinone pool behavior

The pool behavior of ubiquinone is another strong set of data that supports the fluid-state model of the ETC. It was derived using the assumption

12 that the steady-state respiration of mitochondria follows a two-enzyme system, the reduction of ubiquinone and the oxidation of ubiquinol. In order for quinone to kinetically behave as a homogenous pool in the membrane, the diffusion of both ubiquinone and ubiquinol must be faster than the chemical reactions of ubiquinone reduction and oxidation. Kroger and Klingenberg were the first to verify the pool behavior of ubiquinone from submitochondrial particles of bovine heart (Kroger & Klingenberg 1973). Further evidence of a ubiquinone pool was later provided in a variety of systems (see review (Lenaz & Genova 2009a)). In addition, it was shown that NADH-cytochrome c activities (complex I-III) and succinate-cytochrome c activities (complex II-III) were decreased after the mitochondrial membrane was subjected to phospholipid fusion since ubiquinone in the pool was diluted. Incorporation of excessive amounts of ubiquinone could restore these activities (Schneider et al. 1982).

1.2.2. Solid-state model

Despite the success of the studies supporting the fluid-state model, the use of blue native polyacrylamide gel electrophoresis (BN-PAGE or BNG) has resurrected the study of the organization of respiratory complexes. The solid- state model was originally proposed by Chance and Williams (Chance & Williams

1955), years before the fluid-state model. It rapidly lost credibility due to several kinetics and biophysics studies favoring the fluid state model. The solid-state model depicts the respiratory complexes I, III and IV as a very large macromolecular structure called a respiratory supercomplex. The term

“respirasome” is also used interchangeably with “supercomplex” merely to

13 implicate the functional relevance of these structures. It has been proposed that

the respirasome may increase the efficiency of the respiratory chain, bringing the

individual complexes and mobile electron carriers in close proximity (Chance &

Williams 1955). The following describes the important recent evidence supporting

a solid-state model of the ETC.

• Structural identification of respiratory supercomplex

BNGs have become the main experimental strategy for structural analysis

of respiratory supercomplexes. This technique deploys non-ionic detergents to

solubilize the membrane bound protein to preserve the native conformation of the

membrane protein. Coomassie blue dye is used to provide the cationic charges

to the solubilized native protein for running in the non-denaturing polyacrylamide

gel. Among the commonly used non-ionic detergents, digitonin is the mildest

detergent that keeps the supercomplex structure intact (Wittig et al. 2006).

Respiratory supercomplexes are found in both prokaryotes and eukaryotes. They exist in different stoichiometries, including I1III2, I1III2IV1-4, III2IV1-2 (the subscripts

indicate the copy number of a complex). Interestingly, complex II has rarely been

found to be a component of a supercomplex.

Besides detergent-based BNGs, supercomplexes can also be identified by

sucrose gradient fractionation (Acin-Perez et al. 2008), which substantiates that

supercomplexes are not an electrophoretic artifact of individual respiratory

complex as has been argued.

Recently, Schafer et al (Schafer et al. 2007) used electron microscopy to

reveal the positions and interactions of the complex components in bovine

14 supercomplex I1:III2:IV1 (Fig. 8). It was shown that a dimer of complex III is

attached to the midsection of the membranous arm of complex I, whereas a

monomer of complex IV binds at the distal end of the membranous part of

complex I. The ubiquinone binding sites of complex I (near the junction of the

matrix and membranous arms (Kashani-Poor et al. 2001; Zickermann et al.

2003)) and complex III (between the Rieske and cytochrome b subunits (Covian

& Trumpower 2005; Huang et al. 2005)) were postulated to face each other.

Similarly, cytochrome c1 and cytochrome c oxidase subunit II, the binding sites of

cytochrome c in complex III and IV, respectively, were shown to be in close proximity. This arrangement may enhance the efficiency of electron transfer from complex I via complex III to complex IV.

• Metabolic flux control analysis

A metabolic flux control analysis (Kacser & Burns 1979; Moreno-Sanchez et al. 1999) is a kinetic approach that differentiates the types of organization of

the enzymes that comprise a metabolic pathway of interest. The flux-control

coefficient (Ci) reflects the relative control of each enzyme over the global flux.

This can be calculated by the fractional change in the global flux through the pathway induced by a fractional change in the enzyme activity under particular circumstance, such as the in a presence of a specific inhibitor. If the metabolic pathway is comprised of freely diffusible enzymes as postulated by the fluid-state model, the relative kinetic control of the pathway could be different for each

enzyme within the metabolic pathway, i.e. Ci of each step in the pathway would be significantly different from each other. The sum of the Ci values for all

15 enzymes would approach 1. On the other hand, if the metabolic pathway

behaves as a single enzymatic unit, like the solid-state model, the inhibition of

any one of the enzyme components would equally affect the change in global

flux. The Ci values of any step in the pathway would be approximately 1. The sum of all Ci values would then be greater than 1. The schematic explanation of the metabolic flux control analysis is simplified in figure 9. Bianchi et al (Bianchi

et al. 2004) used metabolic flux analysis to test whether the bovine respiratory

chain behaves as a single enzyme unit. In their analysis, both complexes I and III

were highly rate controlling over aerobic NADH oxidation (CI = 1.06 and CIII

=0.99), favoring the existence of a supercomplex I:III. Complex IV, however, had a CIV of 0.26, most likely due to the pronounced abundance of a free complex IV dimer. For aerobic succinate oxidase, complex II is fully rate limiting for succinate oxidation (CII = 0.88, CIII = 0.34 and CIV = 0.20), suggesting the substrate

channeling of complex II towards complexes III and IV is absent.

• Respiring mitochondrial supercomplexes

Recently, Acin-Perez et al (Acin-Perez et al. 2008) has demonstrated that

mitochondrial supercomplexes are indeed the functional units of the respiratory

chain, capable of respiring when provided with specific electron donors.

Ubiquinone and cytochrome c were also identified within the supercomplex.

Respiration from supercomplex I:III:IV isolated from BNGs was shown as

electron donors of complexes I (NADH) and IV (N,N,N',N'-Tetramethyl-p-

Phenylenediamine; TMPD) were provided to the samples. NADH- and TMPD-

stimulated respirations were also inhibited by specific inhibitors of complexes I

16 and IV (rotenone and KCN). However, the simultaneous inclusion of each

individual component of the supercomplex (complexes I, III and IV mixture) isolated from BNGs did not show respiration in the presence of NADH. Not only are the respiratory complexes and electron donors required for integrated mitochondrial respiration, but also the proper arrangement of the respiratory complexes into a supercomplex is necessary for respiration.

1.3 Implications of the mitochondrial supercomplex

Organization of the mitochondrial respiratory chain into functional entities known as supercomplexes has gained acceptance as a model over the fluid- state model during the past decade, as discussed in 1.2.2. New evidence also suggests that besides normal expression of each individual respiratory complex, sufficient quantities of the assembled supercomplex is another crucial factor that determines respiratory chain function (D'Aurelio et al. 2006). Furthermore, there are a number of human patients with respiratory chain defects that are caused by a single mutation but exhibit combined complex deficiencies (Bruno et al. 2003;

Schagger et al. 2004; Blakely et al. 2005). Such defects stem from the interdependence between components within the supercomplex. When examining respiratory chain defects, consideration of the functional unit of the mitochondrion, the ETC supercomplexes, may be the optimal approach to understanding the patient’s pathology.

A discussion of the implications of supercomplexes as well as their role in respiratory chain defects follows.

17 1.3.1. Functional advantage of supercomplexes

• Possible mechanisms of enhanced respiratory chain function by

supercomplexes

Organization of the respiratory chain into supercomplexes is thought to improve the efficiency of the respiratory chain function by substrate channeling

(Bianchi et al. 2004). Supercomplexes bring together the components of the respiratory chain, assembling them into a single respiring unit (Acin-Perez et al.

2008). This unit provides a sufficiently short intercomplex distance for electron transfer between complexes, similar to electron tunneling (Moser et al. 2005) that facilitates electron transfer between the redox centers within a complex, i.e. intracomplex electron transfer. Electron transfer between components of the respiratory chain is therefore independent from the diffusion-coupled collision, which could be the rate-limiting factor of electron transfer in the respiratory chain

(Hackenbrock et al. 1986). Supercomplexes also sequester the mobile electron carriers (ubiquinone and cytochrome c) within them as previously shown (Acin-

Perez et al. 2008). The rate of electron transfer could also then be improved due to the instant availability of the mobile electron carriers.

Supercomplexes also improve respiratory chain function, by an allosteric mechanism. A study by Schafer et al. (Schafer et al. 2006) demonstrated that complex I in supercomplex I1III2IV1 is enzymatically more active than complex I in

supercomplex I1III2, implying that complex IV has a positive allosteric exertion on complex I function through supercomplex structure. Moreover, complex IV also exhibited a similar effect on complex III activity, i.e. complex III in supercomplex

18 I1III2IV1 is enzymatically more active than complex III in supercomplex I1III2.

However, understanding the mechanism by which individual members of

supercomplexes exert an allosteric effect on electron transfer within another

member of the supercomplex requires further investigation, which is included in

my study.

• Supercomplex organization may prevent excessive mitochondrial reactive

oxygen species (ROS) production

Panov et al. suggested that besides providing an enzymatic advantage,

supercomplexes are also an adaptive mechanism designed to prevent excessive

production of ROS (Panov et al. 2007). The FMN (flavin mononucleotide) and the

iron-sulfur cluster N2 are the potential sites for ROS production in complex I

(Lenaz et al. 2006; Lenaz & Genova 2007). Supercomplex structure may

facilitate important conformational changes in complex I (Radermacher et al.

2006) that can prevent exposing FMN and N2 to oxygen, thus alleviating

excessive ROS production from complex I. A study by Krause et al. (Krause et al.

2006) demonstrated that low ROS production was associated with longevity of

ex1, the long-lived strain of P. anserina. BNGs of ex1 mitochondria showed that

most of complex I was organized into a I2:III2 supercomplex rather than the usual

I1:III2 supercomplex. The complex I dimer may undergo a tighter interaction than a complex I monomer with the complex III dimer. FMN in the complex I dimer then became concealed and kept away from exposure to oxygen, resulting in the low ROS production.

1.3.2. Assembly and stability of complex I

19 Another crucial aspect of the supercomplex is its role in complex I stabilization/assembly. The maturation of complex I is linked to its assembly into supercomplexes (Fernandez-Vizarra et al. 2009; Lazarou et al. 2009) (Fig. 10). In bovine heart, only 14-16% of total complex I exist as a free form when assayed on digitonin-based BNGs, while the rest is found in supercomplexes. This suggests that in vivo all of complex I is bound to complex III, thus forming a I:III or I:III:IV supercomplex (Schagger & Pfeiffer 2001). In mammalian cell lines the presence of fully assembled complex III is required to maintain complex I. For instance, a missense mutation E373K in the cytochrome b subunit of complex III in a mouse cell line and a Δ4-CYT b mutation in a human cell line caused decreased levels of complex III. However, they also caused significant losses of complex I, leading to decreased enzymatic activities of both complexes I and III

(Acin-Perez et al. 2004). Complex I stability has also been shown to depend on the levels of complex IV. For example, a COX10 knockout mouse fibroblast cell line, which was unable to assemble complex IV, showed decreased amounts and activities of complex I (Diaz et al. 2006). Similarly, inhibition of COX IV expression caused impairment in complex I assembly in mouse cell lines (Li et al.

2007). In all cases, the depletion of complex I was caused by a decrease in the formation of complex I-containing supercomplexes due to a lack of complexes III and IV. D’Aurelio et al. (D'Aurelio et al. 2006) demonstrated a decrease in the levels of supercomplex I1:III2:IV1-4, in the hybrid human clones harboring high levels of MTCYB (complex III) and MTCO1 (complex IV) mutations. Either mutation decreased levels of complex I.

20 Complexes III and IV clearly affect the assembly/stability of complex I.

However this does not seem to be reciprocal, since neither qualitative nor

quantitative alteration of complex I has been found to affect levels of complex III

and IV. According to the current model of the complex I assembly process

(Fernandez-Vizarra et al. 2009; Lazarou et al. 2009), fully assembled complexes

III and IV are incorporated into the last pre-complex I intermediate, forming a

partially assembled supercomplex I:III:IV. The completion of supercomplex I:III:IV

formation is made by the addition of the N module of complex I (Fernandez-

Vizarra et al. 2009; Lazarou et al. 2009). The assemblies of complexes III and IV,

which each exist as dimers when not incorporated into a supercomplex, do not

require complex I (Fig. 10).

1.3.3. Decline of mitochondrial supercomplexes in impaired respiratory chain

function in mice and humans

Acin-Perez et al. showed that supercomplexes are the functional operating units of the respiratory chain (Acin-Perez et al. 2008). Following are several examples of human mitochondrial diseases that are caused by defects of

supercomplexes.

• Supercomplex instability due to cardiolipin deficiency

Cardiolipin, the most abundant phospholipid composition in the inner

mitochondrial membrane in most eukaryotes (Daum & Vance 1997;

Mileykovskaya et al. 2005; Joshi et al. 2009), is necessary for supercomplex

stability (Zhang et al. 2002; Pfeiffer et al. 2003; Zhang et al. 2005). In humans,

cardiolipin deficiency results in Barth syndrome, an X-linked recessive disease

21 that presents with cardioskeletal myopathy with neutropenia (Bione et al. 1996;

Barth et al. 2004; McKenzie et al. 2006; Schlame & Ren 2006). Mitochondria

from patients with Barth syndrome contain unstable supercomplexes. Digitonin

extraction of supercomplexes from the patients’ lymphoblasts results in

dissociation of complex IV from supercomplex I1:III2:IVn more readily than in normal cells (McKenzie et al. 2006). The association between complexes I and III in supercomplex I1:III2 was also compromised, as shown by maltoside extraction

(McKenzie et al. 2006). The instability of supercomplexes observed in Barth syndrome patients is speculated to be due to loss of mature cardiolipin species, which binds tightly to complex I (Sharpley et al. 2006) and the putative complex

III/IV interface that is comprised of cytochromes b and c1 (Zhang et al. 2002;

Pfeiffer et al. 2003). Interestingly, the activities of complexes I and IV were found to be reduced in skeletal muscle but not in fibroblasts of these patients, suggesting that respiratory complexes show variable tissue sensitivity to cardiolipin abnormalities. Supercomplex instability is thought to lead to decreased mitochondrial function, which underlies the disease phenotypes.

• BCS1L mutations disrupt complex III and supercomplex formation,

causing Bjornstad and GRACILE syndromes.

BCS1L is a chaperone protein in a member of the AAA family (ATPase

Associated with various cellular Activities) (Nobrega et al. 1992; Cruciat et al.

1999). BCS1L is an inner mitochondrial membrane protein that facilitates insertion of the Rieske iron-sulfur protein into a precursor of complex III (Cruciat et al. 1999). Mutations in BSC1L lead to a wide variety of clinical phenotypes that

22 range from the relatively mild Bjornstad syndrome (sensorineural hearing loss

and brittle hairs (Hinson et al. 2007) to the severe GRACILE syndrome (Growth

Retardation, Aminoaciduria, Cholestasis, Iron overload, Lactic acidosis and Early

death) (Fellman et al. 1998). Hinson et al. (Hinson et al. 2007) investigated

supercomplex formation in mitochondria from lymphocytes of a patient with

Bjornstad syndrome with an R318H mutation in BCS1L. The consequence of the

mutation was accumulation of a complex III precursor that lacked the Rieske

subunit, (apoIII2), as well as accumulation of an atypical intermediate of

supercomplex (apoCIII)2(CIV)n. However, a fully formed I1:III2:IVn supercomplex

was also observed in the patient’s lymphocytes. The overall function of the

electron transport chain, as determined by the rate of reduction of resazurin dye,

was decreased despite an increased mitochondrial content (Hinson et al. 2007).

GRACILE syndrome is the severe form of BCS1L mutation, characterized

by massive iron overload in the liver that develops during fetal life. Several

mutations in BCS1L have been found to be associated with GRACILE syndrome

(de Lonlay et al. 2001; De Meirleir et al. 2003; Moran et al. 2010). Recently

Moran et al. (Moran et al. 2010) demonstrated that some patients with GRACILE

syndrome not only had a defect in complex III biogenesis, but they also displayed

reduced levels of complex I and/or IV. Moran et al. suggested that the reduction

in levels of complex I may be caused by loss of complex III, since complex III has

been shown to stabilize complex I via assembly of supercomplexes (Acin-Perez

et al. 2004). Decreased levels of complex IV may suggest a secondary role of

BCS1L in complex IV maintenance.

23 • Cytochrome b mutations and combined complex I/III deficiency

Mutations in the mitochondrially encoded human cytochrome b gene

(MTCYB) are associated with a number of clinical presentations including

encephalopathy (De Coo et al. 1999), encephalomyopathy (Keightley et al. 2000;

Blakely et al. 2005), cardiomyopathy (Valnot et al. 1999; Andreu et al. 2000), and multiorgan dysfunction (Wibrand et al. 2001). The majority of patients with these mutations, however, present with exercise intolerance and myoglobinuria

(Dumoulin et al. 1996; Andreu et al. 1998; Lamantea et al. 2002; Mancuso et al.

2003). Interestingly, complex III defects caused by MTCYB mutations can be accompanied by a dramatic loss of complex I, resulting in combined complex I/III deficiency (Lamantea et al. 2002; Bruno et al. 2003; Blakely et al. 2005). A study by Schagger et al. (Schagger et al. 2004) indicated that a loss of complex III prevented supercomplex formation and led to the secondary loss of complex I, which explained the combined complex I/III deficiency in some of the MTCYB mutations.

• Complex IV defect and combined complex I/IV deficiency

Complex IV deficiency is one of the most common respiratory chain defects in human. Similar to complex III deficiency, defects in complex IV can be caused from mutations in its assembly factors, i.e. SCO1, SCO2 and SURF1, and mutations in the structural subunits. Supercomplex formation can be affected by loss of complex IV. For instance, a patient with decreased complex IV caused by a SURF1 mutation had a decreased ratio of supercomplex I:III:IV to supercomplex I:III (Schagger et al. 2004). Another study from Diaz et al. (Diaz et

24 al. 2006) demonstrated reductions in both supercomplex I:III and supercomplex

I:III:IV in fibroblasts from COX10 knockout mice. They proposed that the complete absence of complex IV in the COX10 knockout mice led to a global decrease of NADH oxidoreductase supercomplexes as determined by BNGs.

Interestingly, a combined I/IV defect is cited as the most common combined deficiency in patients (Korenke et al. 1990).

• Supercomplex dysfunction in heart failure

Heart failure is the pathological condition whereby the heart cannot supply sufficient blood flow for the body’s needs. A decrease in the amount of ATP in

the failing heart (Shen et al. 1999) is associated with reduction in mitochondrial respiration (Sharov et al. 1998; Sharov et al. 2000), indicating compromised respiratory chain function. Recently, Rosca et al. (Rosca et al. 2008) demonstrated that reduction in mitochondrial respiration in a mouse model of coronary microembilization-induced heart failure was caused by an alteration of the supercomplex profile. Mitochondria from the failing heart lost capacity for oxidative phosphorylation without impairment in either the phosphorylation system (ANT, ATP synthase, phosphate transporter) or levels of the respiratory chain components. However, a dramatic decrease in the amount of supercomplex I1:III2:IVn despite normal levels of individual respiratory complexes was found in the mitochondria from animals in heart failure. They concluded that

the decrease in oxidative phosphorylation of heart mitochondria was explained

by the decrease in the amount of I1:III2:IVn supercomplexes. This study clearly shows that abnormal respiratory chain function can occur despite normal levels

25 of components of the respiratory chain and its associated machinery. Normal expression of supercomplexes seems to be a crucial factor that determines mitochondrial respiration function.

1.4. C. elegans as a model to study the function of mitochondrial

supercomplexes

In 1963, Sydney Brenner introduced C. elegans, a free-living, non- parasitic soil nematode, as a model organism for pursuing research in behavior, developmental biology and neurology. Since its introduction, C. elegans has become a widely used animal model because of its physical and genetic simplicities, low maintenance, and short life cycle. C. elegans has been successfully used to study the mechanisms of several human diseases such as

Parkinson’s disease, Friedreich’s ataxia, muscular dystrophy, etc. (for a review see (Ventura et al. 2006)). Two most recognized research works involving C. elegans included the study of genetic regulation of organ development and programmed cell death—the Nobel Prize-winning research in Physiology and

Medicine in 2002—by Sydney Brenner, John E. Sulston and H. Robert Horvitz, and the discovery of RNA interference – gene silencing by double-stranded

RNA—the Nobel Prize-winning research in Physiology and Medicine in 2006—by

Andrew Z. Fire and Craig C. Mello.

There are many reasons that make C. elegans a useful model to study mitochondrial disorders. Similar to most eukaryotic mitochondria, C. elegans mitochondria house a number of metabolic and biosynthetic pathways such as

26 the Krebs cycle, urea cycle, β-oxidation of fatty acids, etc. Moreover, respiratory

chain subunits of C. elegans share extensive homology with their human

counterparts, i.e. 40-90% protein length identity (Falk et al. 2008). Several classic

mitochondrial mutants have already been isolated and characterized, which

include gas-1 (Kayser et al. 1999) and nuo-1 (Tsang et al. 2001); complex I

mutants, clk-1 (Lakowski & Hekimi 1996); ubiquinone deficiency mutant, mev-1

(Ishii et al. 1998); complex II mutant, and isp-1 and isp-1;ctb-1 (Feng et al. 2001);

complex III mutants. Mitochondrial supercomplexes also exist in C. elegans.

When digitonin was used to solubilize mitochondrial membrane, the functionally

intact supercomplexes I1:III2:IVn and I1:III2 were found to be the predominant form of ETC components. Notably, like mammalian mitochondria, there is no detectable free complex I in worm mitochondria, and free complexes III and IV appear to exist as dimers. (Suthammarak et al. 2009) (Fig. 11).

Mutations in individual members of supercomplexes may afford new insights into their function. A brief introduction of some of C. elegans

mitochondrial mutants that are of particular interest follows.

1.4.1. isp-1(qm150)

The isp-1 mutant allele qm150 was first identified by Feng et al. (Feng et al. 2001) as a long-lived mutant. All physiological rates of isp-1(qm150), including embryonic development, defecation cycle, and egg laying, were found to be 1.5-5 times slower than the wild-type. isp-1 also showed a decrease in whole animal oxygen consumption which was correlated with a high resistance to oxidative stress (Feng et al. 2001; Yang & Hekimi 2010).

27 The isp-1 mutation is a serine substitution of a highly conserved proline

residue that is located in the head domain of the Rieske iron-sulfur protein, ISP

(Iwata et al. 1998). ISP is one of the catalytic subunits of complex III and is

directly involved in the electron transfer of complex III or the Q-cycle as

discussed in 1.1.1. This proline residue has been shown to be crucial for the

secondary structure of the [2Fe-2S] cluster, for it creates an inward folding of the

backbone of the Rieske head domain immediately preceding strand β5 (Iwata et

al. 1996). Gatti et al. (Gatti et al. 1989) showed that a P146L substitution (a position that is equivalent to the worm’s isp-1 allele) in S. cerevisiae altered the midpoint potential of the [2Fe-2S] cluster. Although Feng et al. did not assay the enzymatic activities of isp-1, the mutant was very likely to suffer from, at least, a low complex III activity according to the significance of the mutation. Interestingly,

Feng et al. has also identified qm189 as a suppressor of qm150 phenotype, as the following discussion illustrates.

1.4.2. isp-1(qm150);ctb-1(qm189)

The allele qm189 of ctb-1 mutation, ctb-1(qm189), has been identified as a spontaneous suppressor of the isp-1(qm150) phenotype (Feng et al. 2001).

The qm189 arose spontaneously during the process of culturing qm150. isp-

1;ctb-1 exhibited improved physiological rates compared to isp-1. The ctb-1 mutation is an alanine to valine substitution at the N-terminal of helix αD of cytochrome b, another catalytic subunit of complex III. Although the mutation does not occur at a conserved residue, its location can potentially affect function of complex III. The N-terminal of helix αD of cytochrome b is part of the docking

28 site (helix αC, helix αcd1 and loop ef) of the Rieske head domain during electron transfer of complex III (Iwata et al. 1998). Interestingly, there are a few cases where mutations in mitochondrial genes are the suppressors of other mutations.

However, such suppressors have been reported only in mt-tRNA genes. For example, respiratory chain deficiency caused by the m.3243A>G mutation in the mt-tRNALeu(UUR) (MTTL1) gene was found to be suppressed by the m.12300A>G mutation in the mt-tRNALeu(CUN) of the (MTTL2) gene (El Meziane et al. 1998).

There is no other case where a mutation in a structural subunit of respiratory complex is a suppressor of another structural subunit mutation. Moreover, there is no report of a mutation that is equivalent to qm189 in other organisms.

It is interesting to note that the addition of the ctb-1 mutation in the already defective complex III, caused by isp-1, can render better whole animal phenotypes than isp-1 alone. One possibility is that ctb-1 may partially restore complex III function in isp-1 background. Unfortunately, Feng et al. (Feng et al.

2001) did not further investigate the mechanism of the suppression.

1.4.3. Complex IV deficient C. elegans

The repertoire of the classical mitochondrial mutants in C. elegans does

not include a complex IV deficiency mutant. However, it can easily be created

using RNA-mediated interference (RNAi), first introduced by Kamath el al.

(Kamath et al. 2001). This technique is a strategy to effectively reduce the

expression of a gene of interest in C. elegans. Lee et al. (Lee et al. 2003)

exploited the RNAi technique to screen thousands of genes across the genome

in order to identify genes that extended the lifespan of C. elegans when

29 inactivated. They found that COX IV knockdown extended the worm’s lifespan.

However, effects of the COX IV knockdown on the respiratory chain function were not further investigated. Complex IV deficiency in other animal models, including human, can perturb supercomplex formation and lower the amount and activities of respiratory complex I, as discussed in 1.3.3.

1.5 Conclusion

The intact oxidative phosphorylation system of the respiratory chain is the major determinant for normal mitochondrial function. Most defects in the mitochondrial respiratory chain result in energy insufficiency, which can affect any organ system in the body. At the moment, the mitochondrial supercomplex is accepted as a functional unit of the respiratory chain. Interdependence among components of supercomplexes has been showed to play a pivotal role in the respiratory chain function and the observed phenotypes. However, knowledge about the function of supercomplexes is still limited. Availability of the mitochondrial mutants in C. elegans allows us to acquire a better understanding of the structure and function of mitochondrial supercomplexes.

1.6 Specific Aims

My research work aims to answer the question regarding the functional significance of mitochondrial supercomplexes. To address this question, there are two hypotheses to test, which include:

30 1. Inactivation of mitochondrial complex IV genes in C. elegans causes alteration in respiratory complexes and supercomplexes formation, which results in some phenotypic changes.

2. The complex III mutant isp-1 exerts its effects by altering complex III function and by affecting supercomplex formation. The suppressor of isp-1, ctb-

1, restores the function of complex III and the formation of the supercomplexes.

The specific aims to test these hypotheses follow.

Aim 1 Characterize the effect of cytochrome c oxidase (complex IV) deficiency in C. elegans.

1.1. Perform RNAi knockdown of COX IV and COX Va, the two nuclear genes encoding the subunits of complex IV.

1.2. Study mitochondrial function in both RNAi-knockdown worms by the oxidative phosphorylation (OXPHOS) assay and electron transport chain

(ETC) assay.

1.3. Study the organization of mitochondrial supercomplexes in both knockdown worms by BNG.

1.4. Study lifespan, fecundity and anesthetic sensitivity of both knockdown worms.

Aim 2 Study the interaction between respiratory complexes I and III in the three complex III mutants, isp-1, isp-1;ctb-1 and ctb-1.

2.1. Study the phenotypes of isp-1, isp-1;ctb-1 and ctb-1.

2.2. Study mitochondrial function (OXPHOS and ETC assay) and organization of mitochondrial supercomplexes in isp-1, isp-1;ctb-1 and ctb-1.

31 2.3. Characterize the interaction between isp-1 and ctb-1.

These studies are intended to demonstrate the functional significance of mitochondrial supercomplexes by using a whole animal. The substantial amount of information that would be gained from my study will help us to understand more about the energy-producing machinery in our cells. It will potentially be useful for mitochondrial medicine and, in the future, may help many patients who suffer from mitochondrial disorders.

32 1.7 Figures

FIGURE 1

Figure 1. The integrated function of the mitochondrial respiratory chain.

Located in the inner mitochondrial membrane, the respiratory chain is composed of 5 multi-subunit complexes (complexes I through V) and 2 mobile electron carriers (Q; ubiquinone and Cyt c; cytochrome c). Electrons are transferred from

NADH or succinate to O2, the terminal electron acceptor. Complex I, III and IV generate a proton gradient across the membrane (grey arrows from matrix to intermembrane space). Complex V utilizes this gradient to phosphorylate ADP to

ATP. Figure adapted from Seelert et al. (Seelert et al. 2009).

33 FIGURE 2

Figure 2. The functional modules of complex I. A. All three functional modules of complex I, which are conserved across kingdoms, are illustrated: the electron input module (N module), electron output module (Q module) and the proton translocation module (P module). B. Electrons from NADH are transferred through complex I via the flavin mononucleotide (FMN) and a series of iron-sulfur redox centers (blue circles) to ubiquinone (Q). Complex I couples electron transfer with proton translocation across the membrane (dotted purple arrow).

Figure adapted from Lazarou et al. (Lazarou et al. 2009).

34 FIGURE 3

Figure 3. Structure of the membranous arm of E. coli complex I and proposed model of proton translocation by complex I. A. The crystallographic analysis of α-helical bundles of complex I in a side view (upper panel) and from the periplasm into the membrane (lower panel) represents the antiporter-like structure of subunits NuoL/M/N. NuoL also has a unique amphipathic α-helix (purple HL helix) that spans almost the entire length of the membranous arm of the complex. B. Electron transfer at the matrix arm of complex I is coupled to conformational changes of the joint region between the matrix arm and the membranous arm (arrows and green rod structures). These changes are transmitted to the membrane-spanning domain of NuoL, which then tilts three antiporter-like NuoL/M/N subunits, resulting in proton translocation.

Figure adapted from Efremov et al. (Efremov et al. 2010).

35 FIGURE 4

Figure 4. Trimer association of E. coli complex II and its redox centers.

Complex II forms a trimer under physiological conditions. viewed parallel to the inner mitochondrial membrane as shown in A, and from the cytoplasm along the membrane as shown in B. SdhA, SdhB, SdhC and SdhD subunits are shown in purple, orange, green and blue, respectively. C. Complex II monomer viewed parallel to the membrane. All redox centers are illustrated on the right. Heme b is not aligned with the rest of the redox centers of complex II. It is postulated that heme b is not directly involved in electron transfer of complex II. Figure adapted from Yankovskaya et al. (Yankovskaya et al. 2003).

36 FIGURE 5

Figure 5. Q cycle of complex III. A. The first half of the Q cycle. Ubiquinol

(UQH2) binds to the Qp site of complex III. Upon UQH2 oxidation (1), one electron is transferred to the iron-sulfur protein (2Fe/2S) of complex III, and then

- to cytochrome c (high-potential chain). The resulting semiquinone (UQ⋅ ) at the

37 QP site transfers a second electron to cytochrome b (bL to bH) (low potential

chain) (2) and (3). Heme bH reduces another ubiquinone (UQ) at the QN site to

- the UQ⋅ intermediate (4). B. The second half of Q cycle. Another UQH2 is

oxidized at the QP site and steps (1) (2) and (3) are repeated. At step (4), heme

- bH reduces the UQ⋅ intermediate from the first half of Q cycle at the QN site to

UQH2. Figure adapted from David G. Nicholls and Stuart J Ferguson.

Bioenergetics 3. Barcelona: Academic Press, 2002.

38 FIGURE 6

Figure 6. The catalytic core and redox centers of complex IV. The two key catalytic subunits of complex IV, COXI and COXII, are illustrated. Cytochrome c, docking at its binding site at COXII, is also depicted. An electron is transferred from cytochrome c  CuA  heme a (light grey arrows). The binuclear [heme a3-

CuB] center reduces O2 to water (not indicated). Cytochrome c oxidase couples electron transfer with proton translocation from the matrix (N-side) to the intermembrane space (P-side). Figure adapted from Lenaz et al. (Lenaz &

Genova 2009b).

39 FIGURE 7

Figure 7. ATP synthesis of complex V. A. Cross-section of the catalytic head

α3β3-hexamer of the F1 domain shows ATP synthesis. The 120° rotation of the γ- subunit leads to conformational changes in all three catalytic sites (O, L and T) of the F1 domain. As a result, one molecule of ATP is released from one catalytic site, new ADP and inorganic phosphate (Pi) are tightly bound to another catalytic

site, and another molecule of ATP is formed at the third catalytic site. B. Side

view of complex V. The a-subunit contains two proton channels. For a proton to cross the membrane, it must move through one channel, then bind to one of the c-subunits, and finally be carried to the other partial channel by rotation of the c- ring. The c-ring is anchored to the central stalk, which is composed of the γ-

subunit. The a-subunit is anchored, through b2, to the α3β3-hexamer of the F1

domain. Figure adapted from Cross et al. (Cross 2004).

40 FIGURE 8

Figure 8. The structure of supercomplex I1:III2:IV1 and the putative binding sites of ubiquinone and cytochrome c. A and F. Side views along the membrane plane. B. Top view from the matrix side. C-G. Side views along the membrane plan at different rotational angles as indicated. H. Bottom view from

41 the intermembrane space. All figures show the three individual complexes as they would assemble to form the supercomplex. Complex I is L-shaped, in yellow. Complex III dimer is in red and located at the mid-section of the membranous arm of complex I. Ubiquinone binding sites of complex I and complex III are facing each other as indicated in the black and grey rectangles.

Complex IV is illustrated in green and located at the end of the membranous arm of complex I. Cytochrome c binding sites of complex III and complex IV are in close proximity and facing towards each other (light blue rectangles). Figure adapted from Schafer et al. (Schafer et al. 2007).

42 FIGURE 9

Figure 9. Simplified schematic explanation of metabolic flux control

analysis. A. Liquid state model. In this model, the respiratory chain complexes

are organized according to the liquid-state model, where each individual complex

randomly disperses on the membrane as a separate entity. The relative control of

each respiratory complex may be different and the sum of all the flux coefficients

(Ci) is equal to 1. B. Supercomplex model. Any step in the pathway is regarded as a component of a single enzyme and must be completely rate-controlling.

Therefore each component elicits the maximal flux control coefficient (Ci = 1) and

the sum of all coefficients is higher than 1. Figure adapted from Lenaz et al.

(Lenaz & Genova 2009b).

43 FIGURE 10

Figure 10. A current model of complex I assembly pathway. (1) A nascent

form of the Q module is anchored to the membrane by ND1 and other membrane

embedded subunits forming a 400 kDa subcomplex. (2) Two assembly factors,

CIA30 and Ecsit, complete formation of a 460 kDa subcomplex of the P module.

(3) The 400 kDa and 460 kDa intermediates combine to form a 830 kDa

intermediate. Complexes III and IV are likely to incorporate into this intermediate

to form a 830 kDa/complex III:IV supercomplex intermediate. (4) The latter

stages of complex I assembly involve the addition of the N module and the

completion of the P module via the addition of a ND4:ND5 subcomplex. B17.2L and C6ORF66 are other factors involved in complex I assembly as indicated.

Figure from Lazarou et al. (Lazarou et al. 2009).

44 FIGURE 11

Figure 11. Respiratory supercomplexes in C. elegans. Mitochondrial proteins from wild-type animals were solubilized by digitonin and electrophoresed in 3.5–

11% gradient acrylamide gels. The gels were subsequently stained with

Coomassie blue or used to perform in-gel activity assays for complex I (CI-IGA) or complex IV (CIV-IGA) as shown. The numbers in the left panel indicate the approximate molecular masses (kDa) of the corresponding protein bands.

Respiratory complexes and supercomplexes are indicated on the right. Figure adapted from Suthammarak et al. (Suthammarak et al. 2009).

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Chapter 2

The effects of COX IV- and COX Va-RNAi

knockdowns on whole animal phenotypes, respiratory chain function and supercomplex

formation in Caenorhabditis elegans

(The work in this chapter has been published in the Journal of Biological

Chemistry, Vol 284, NO. 10, pp. 6425-6435, March 6, 2009)

2.1 Introduction

The mitochondrial respiratory chain consists of five multi-subunit

complexes termed complexes I through V. The physical organization of these

complexes is controversial. Two extreme models of their structure have been

proposed. A “liquid-state model” of the mitochondrial respiratory chain depicts the

respiratory complexes embedded in the inner mitochondrial membrane as

separate entities, functionally connected to each other by the mobile electron

carriers, coenzyme Q and cytochrome c. This model postulates that random collision among all respiratory components can account for measured electron transport rates in the inner mitochondrial membrane. Lateral diffusion of each

component is regarded as sufficient to generate contact between respiratory chain components (Hackenbrock et al. 1986; Chazotte & Hackenbrock 1988).

However, data also exist to support a “solid-state” model of respiratory complexes. This model proposes that mitochondrial respiratory complexes are organized into very large supercomplexes (Chance & Williams 1955; Schagger &

Pfeiffer 2000; Schagger & Pfeiffer 2001; Eubel et al. 2003; Stroh et al. 2004;

Krause et al. 2005). Stoichiometric architectures of supercomplexes have been identified in multiple organisms, from prokaryotes to humans, and include I1III2,

I1III2IV1-4 and III2IV4 (Cruciat et al. 2000; Schagger & Pfeiffer 2000; Schagger

2001; Schagger & Pfeiffer 2001; Bianchi et al. 2004; Schagger et al. 2004;

D'Aurelio et al. 2006; Schafer et al. 2006; Vonck & Schafer 2008). Supercomplex

architecture suggests a kinetic advantage which increases the respiratory rate by

providing substrate channeling between components of the supercomplex as well

as stabilizing the complexes (Schagger & Pfeiffer 2000; Genova et al. 2003;

Bianchi et al. 2004; Schagger et al. 2004).

There is also compelling evidence to show interdependence among the

individual components of supercomplexes. In mammalian cell lines complex III

assembly is required to maintain an intact complex I (Acin-Perez et al. 2004).

Structural defects in complex III affected the amount of complex I, whereas chemical inhibition did not. Patients with defects in cytochrome b not only lose complex III, but also show decreased amounts of complex I, while maintaining normal enzymatic activity of the complex (Budde et al. 2000; Bruno et al. 2003;

Schagger et al. 2004; Blakely et al. 2005). Conversely, the disruption of complex

I function caused by nonsense mutations in NDUFS4, a subunit of this large multimeric complex, leads to the partial loss of complex III activity in skin fibroblast cultures obtained from Leigh-like patients (Budde et al. 2000; Scacco et al. 2003). However defects in the complex I subunit ND5 did not cause a loss of complex III in the I-III supercomplex (Cardol et al. 2008).

In most eukaryotes cytochrome c oxidase (COX), or complex IV, contains

10 nuclear and 3 mitochondrial encoded subunits. In cell lines, complex IV stabilizes the assembly of complex I. COX10 knockout mouse cell lines, which were unable to assemble complex IV, showed decreased amounts of complex I, as detected by western blot analysis following blue native gel electrophoresis. In human cell lines, high COX I mutation levels can lead to destabilization of complex I (D'Aurelio et al. 2006). In addition, inhibition of COX IV expression caused impairment in complex I assembly in mouse cell lines (Li et al. 2007).

Although evidence supports interdependence among respiratory chain

complexes, the mechanism regulating this phenomenon remains unclear. I

hypothesized that supercomplexes exist in the nematode C. elegans. I also

hypothesized that decreasing amounts of complex IV subunits would inhibit the

assembly of supercomplexes that include complex IV. I expected that, ultimately,

reduced levels of complex IV subunits would in turn reduce amounts of

associated components of any supercomplex that includes complex IV. These

defects could lead to whole animal phenotypes characteristic of mitochondrial

dysfunction. Here we use RNA interference (RNAi) to knock down two different

predicted subunits of COX.

COX IV and Va are two nuclear encoded subunits of complex IV. Each

controls mitochondrial energy metabolism (Kadenbach et al. 1997; Napiwotzki et

al. 1997). COX IV does so through an allosteric mechanism when the

extramitochondrial ATP/ADP ratio is high. Binding of ATP to the cytosolic domain

of COX IV leads to an increase of the Km of cytochrome c (Napiwotzki &

Kadenbach 1998), which slows mitochondrial respiration. This mechanism can be abolished by the binding of 3,5-diiodothyronine to COX Va (Arnold et al.

1998).

No animals with nuclear encoded genetic defects in subunits of complex

IV have been reported. Genes predicted to encode the worm homologues of

COX IV and COX Va were each subjected to RNAi knockdown. After knockdown, whole animal phenotypes were recorded, as well as function and structure of the respiratory chian. Lifespan and fecundity of the animals were reduced, consistent

with mitochondrial dysfunction. Complex IV activity was reduced, commensurate

with decreased amounts of complex IV. Surprisingly, I found that the rate of electron transport within complex I was also decreased by knockdown of either complex IV transcript. Despite the decreased enzymatic activity of complex I in the complex IV knockdown animals, total amount of complex I remained normal.

In diagnosis of patients, our results caution that primary complex IV defects could be misinterpreted to be primary complex I /complex IV defects.

2.2 Results

2.2.1. The effects of COX IV- and COX Va-RNAi knockdowns on whole animal

phonotype

Nematodes treated with RNAi for either subunit COX IV (W09C5.8) or

COX Va (Y37D8A.14) required an additional day to reach adulthood compared to

wild type (N2). Lifespan was significantly shortened by 3-5 days, and fecundity

was less than half of normal (Table 1). No change in anesthetic sensitivity was

seen in either mutant.

2.2.2. Efficiency of RNAi knockdown

The average percentages of COX IV and COX Va knockdown determined by qPCR from 4 independent cultures were 53.1±7% and 27.4±4%, respectively

(Fig. 12, panel B).

2.2.3. The effects of COX IV- and COX Va-RNAi knockdowns on respiratory

chain function

• Oxidative phosphorylation (OXPHOS) capacity

OXPHOS capacity of intact mitochondria isolated from the knockdown

worms and wild-type animal was determined by rates of oxygen consumption. As

expected, complex IV respiration was decreased by both RNAi knockdowns (Fig.

12, panel A). In addition, the state 3 rates of both complex I- and complex II-

dependent respiration were significantly lower in COX IV knockdown worms than in N2. COX Va knockdown significantly decreased complex I-dependent rates but no significant change was seen in the complex II-dependent rates. The defects in respiratory chain function in both RNAi worms were consistent with complex IV deficiencies. The amount of knockdown of the target gene correlated with TMPD/ascorbate rates; respiratory rates were lowest in those preparations with the greatest RNAi effect (Fig. 12, panel B).

• Electron transport chain (ETC) function

KCN-sensitive-cytochrome c oxidase activities in COX IV and COX Va knockdown worms were significantly lower than in the wild type, as expected

(Fig. 13, panel A). Interestingly, the average rotenone-sensitive NADH- decylubiquinone oxidoreductase activities (electron flow through complex I) in the

COX IV and COX Va worms were 59% and 48% lower than in the wild type, respectively (Fig. 13, panel B). NADH-ferricyanide reductase (NFR), a measure of proximal complex I function, was not decreased (Fig. 13, panel C). However, rotenone-sensitive NADH-cytochrome c oxidoreductase (CI-III), a measure of

electron flow through complex I to III was decreased in COX Va knockdowns

(Fig. 13, panel D). However, complex II-III rates in the knockdowns were

statistically unchanged compared to control (Fig. 13, panel E). Antimycin A-

sensitive decylubiquinol-cytochrome c oxidoreductase activities (complex III) in

the knockdown worms were also not statistically different than wild type (Fig. 13,

panel F). Since II-III and III activity were unchanged, we concluded that complex

II is unaffected by knockdown of complex IV. Complex I and complex IV activity

were specifically decreased in the mutants; we therefore asked whether the

amounts of these complexes were also changed.

2.2.4. The effects of COX IV- and COX Va-RNAi knockdowns on supercomplex

formation

• Supercomplex organization in C. elegans mitochondria

Blue native gels (BNGs) were used to determine the amount of respiratory complexes in the worms with and without RNAi treatment. To optimize the BNG conditions, we first tested dodecyl-maltoside (MS), Triton X-100 (TX), and digitonin (DT), as detergents in our isolation. I also varied the protein to detergent mass ratio in multiple attempts to optimally isolate supercomplexes as discreet entities and found that digitonin at a detergent/protein mass ratio of 6/1 best preserved most mitochondrial supercomplexes (Fig. 14). Mass spectrometry was

used to identify the polypeptides in each band appearing in digitonin (6/1) BNGs.

Of the respiratory chian proteins, 26 complex I subunits, 5 complex III subunits, 6

complex IV subunits and 12 complex V subunits were identified (Table 2).

Mitochondrial matrix proteins were found in the same bands as mitochondrial

supercomplexes, although the ratio of non-respiratory chain/respiratory chain

peptides was less than ¼, and most commonly seen in isolated complex III. The

presence of specific subunits led to the identification of the complexes contained

in each band.

In the digitonin gel, multiple bands of greater than 1,236 kDa contained

subunits of supercomplex I:III:IV (Fig. 14, panel A, DT) (Table 2). Two other

bands at 1,048 and 1,000 kDa were also identified as supercomplex I:III:IV. The

950 kDa band contained only supercomplex I:III. Three bands at 880, 800, and

720 kDa were identified as complex V. Molecular weight analysis predicts that

the 880 kDa band represents dimeric complex V while the 720 kDa band is the

monomer. Each band contained peptides from all complex V subunits. Relative

amounts of these three bands to each other varied slightly in digitonin based

gels, but the sum of their staining was relatively constant (data not shown). The

band at 600 kDa corresponds to dimeric complex III. Complex IV is located at

420 kDa, predicted by molecular weight to be dimeric complex IV. A band at 500

kDa, which is positive for CIV-in gel activity (IGA), was not analyzed by mass

spectrometry. However, its molecular weight matches the summation of complex

III and complex IV (~300 kDa and ~200 kDa, respectively). Although complex III is usually found only in dimeric form, I cannot rule out that a monomer may form with complex IV. Thus, the band at 500 kDa may represent a supercomplex III:IV but with the monomeric form of each. Mass spectrometry was also use to analyze bands from the TX-treated lane (Table 3). It was shown that that the band at about 800 kDa contained only complex I subunits, with no proteins identified from other complexes. Thus, Triton X-100 can be used to isolate complex I, although it is not known whether it contains a complete form of the

complex. The two bands at approximately 860-900 kDa each contained complex

I and III subunits. No bands were seen above 900 kDa in both TX and MS lanes.

I was not able to detect complex II in any of my BNGs.

I used in-gel activity assays (IGA) of BNGs of N2 mitochondria to verify

the presence and amount of the respiratory complexes (Fig. 14, panels B and C).

IGAs corroborated the proteomic results (Tables 2 and 3). No complex I or complex IV activity was seen outside of bands that were predicted by mass spectrometry to contain subunits of these complexes. Complex I remained sufficiently intact in all three detergents to maintain enzymatic function, while complex IV activity was only seen in the digitonin treated lane.

• COX IV and COX Va knockdowns altered supercomplex profile

RNAi treatment for COX IV and COX Va disrupted supercomplex formation, as seen in BNGs of digitonin treated mitochondria (Fig. 15). Not surprisingly, COX IV and COX Va knockdown decreased amounts of dimeric complex IV (Fig. 15, panel A). Supercomplex I:III:IV also decreased in both knockdowns, compared to the wild type, whereas supercomplex I:III increased.

RNAi treatment did not affect amounts of isolated complex III or complex V.

CIV-IGA verified loss of complex IV in the knockdown animals (Fig. 15, panel B). Compared to wild-type activity, I observed a decrease in COX activity at 420 kDa, the dimeric complex IV, in both RNAi worm strains. The activity of the complex IV-component in supercomplex I:III:IV was also decreased in the

RNAi treated worms. CI-IGA was decreased in supercomplex I:III:IV after RNAi treatment in the knockdown worms, consistent with the decrease in

supercomplex I:III:IV formation (Fig. 15, panel C). However, CI-IGA of supercomplex I:III increased. The ratio of CI-IGA activity in I:III:IV to the total CI-

IGA, averaged over 4 independent gels, was significantly decreased in the knockout animals compared to N2. (Fig. 16, panel A)

• Complex I formation was unchanged in COX IV and COX Va worms

Since ETC activity of complex I was decreased with COX subunit knockdown, we attempted to quantify complex I more precisely. In the BNGs solubilized with digitonin (as in Fig. 15), total CI-IGA, normalized for protein loading, was the same in N2 and the RNAi knockdown strains (Fig. 16, panel B).

At a detergent to protein mass ratio of 5/1, Triton X-100-based BNG revealed one individual complex I band and two supercomplex I:III bands (Fig. 16, panel

C) as determined by mass spectrometry and corroborated by CI-IGA (Fig. 16, panel D). Both individual complex I and supercomplex I:III were not significantly decreased from N2 in COX IV or COX Va knockdown animals. The total complex I in each of the complex IV knockdown worms was the same as in the wild-type animals. Western blot analysis of mitochondrial protein probed with an antibody to a complex I subunit (orthologue of NUO-2) showed no differences in levels of expression between the knockdown strains and N2 (Fig. 17).

2.3. Discussion

Nematode COX IV (encoded by the gene W09C5.8) has 24% identity to human COX IV in amino acid sequence, while nematode COX Va (encoded by

Y37D8A.14) has 30.5% identity to human COX Va. RNAi treatment of COX IV

and COX Va inactivated the expression of their respective genes as shown by

qPCR data. Knockdown of either gene impaired respiration and electron

transport in a manner consistent with COX deficiency. Therefore, knockdown of

these two highly conserved components of the MRC indicate that they are

orthologues of their mammalian counterparts in complex IV of the MRC.

This study demonstrated that COX IV- and COX Va- depleted worms grew

slowly, had shortened lifespans, and decreased fecundity, consistent with

mitochondrial dysfunction. Lifespans of COX IV and COX Va knockdown worms

were 14±0.9 days and 12±1.5 days, respectively (compared to 17 + 1.1 days for

N2). This differed from the report of Lee et al. (Lee et al. 2003), which found that

COX IV knockdown worms lived longer than the wild type. However, they used

FUDR (fluorodeoxyuridine) to limit offspring and allow an RNAi screen of 5,690 genes. I also studied lifespan in the presence of FUDR and found results similar to those reported by Lee et al. I did not use FUDR here in order to assay conditions under normal life stresses. Lifespan may be directly affected by sterility per se. My data indicate that disruption of complex IV makes the animals unable to meet the increased stress of oocyte production, leading to shortened lifespan.

BNGs of mitochondria revealed that mitochondrial supercomplexes exist in the nematode, much as described in other organisms such as supercomplexes

I:III:IV and I:III (Cruciat et al. 2000; Schagger & Pfeiffer 2000; Schagger 2001;

Eubel et al. 2003; Bianchi et al. 2004; Schagger et al. 2004; Stroh et al. 2004;

Krause et al. 2005; D'Aurelio et al. 2006; Schafer et al. 2006; Vonck & Schafer

2008). Interestingly, the I:III:IV supercomplex is the most abundant supercomplex

in C. elegans’s mitochondria as well as in human and bovine. I observed

decreased amounts of complex IV and decreased CIV-IGA for both complex IV

mutants. As COX IV and Va encode core subunits of complex IV, the decrease in

functional complex IV, seen in oxidative phosphorylation capacity, ETC and IGA

assays, is likely caused by an insufficient amount of this subunit for full complex

IV assembly. Our results are similar to the study from Li et al. (Li et al. 2006)

which found that the inhibition of COX IV expression led to a loss of complex IV

assembly in mouse cell lines. Interestingly, I found that CIV-IGA activity was

either in a dimeric complex IV (420 kDa) or a supercomplex of complexes I, III, and IV; I did not detect CIV-IGA in a monomeric form. The dimeric form of COX is generally accepted as its functionally active form (Yoshikawa 1997). My

observation is in contrast to the report of Grad et al. (Grad & Lemire 2006),

which found both the monomeric and the dimeric forms of cytochrome c oxidase

of C. elegans in their CIV-IGA. However, Grad et al. did not report the detergent

used to solubilize mitochondria. Digitonin may keep more respiratory chain

complexes and supercomplexes intact (Fig. 14).

To identify the proteins in the bands appearing on BNG as completely as

possible, I took advantage of the high sensitivity and specificity of mass

spectrometry. Supercomplex I:III:IV appeared as a stack of multiple bands at the

upper most part of the gel (1,236 kDa and higher). The previous study by

D’Aurelio, et al (D'Aurelio et al. 2006) showed that supercomplex I:III:IV in human cybrid cell lines can be assembled into 4 different stoichiometries. Each

contained 1 copy of complex I and 2 copies of complex III, i.e. I:III2, as well as

different copy numbers of complex IV, varying from 1-4 copies. The uppermost

bands in our BNGs are likely to be I:III2:IV4, I:III2IV3, I:III2:IV2, and I:III2:IV.

However, the molecular weight of each band could not be accurately defined by its mobility in the gel. In addition, it would not have been exactly the cumulative molecular weight of all complex components because not every subunit of complex I, III and IV has been identified by mass spectrometry in my gels (Table

2). For example, supercomplex I:III:IV was also identified at both 1,048 kDa and

1,000 kDa, but each band was composed of fewer ETC subunits than the higher molecular band at greater than 1,236 kDa. This most probably represents relative binding strength of different subunits to the complexes, resulting in the dissociation of those subunits during the isolation.

The isolation of supercomplexes and their stoichiometries in worms are dependent on the amount and type of detergent used for mitochondrial solubilization. Like other reports, I obtained very different BNGs using digitonin,

Triton X-100, and dodecly-maltoside as detergents (Fig. 14) (Schagger & Pfeiffer

2000; Eubel et al. 2003). Digitonin showed the maximal preservation of supercomplexes, as well as of IGA of complex IV, both as part of a supercomplex and as a homodimer.

I did not observe a band in BNGs that contained only complex I subunits in any gel that used digitonin as a detergent. I interpret these results to indicate that under physiologic conditions virtually all complex I is bound to complex III in

C elegans. Similar observations have been made in plants, where 50-90% of

complex I forms parts of supercomplex I:III2 (Eubel et al. 2003; Eubel et al. 2004;

Dudkina et al. 2005). In bovine heart mitochondria only 15% of complex I was found in free form (Schagger & Pfeiffer 2001). All human complex I in skeletal muscle was shown to be exclusively assembled into supercomplexes (Schagger et al. 2004). It may then be a general feature among mitochondria that almost all complex I is associated with supercomplexes. In contrast, I did not find complex

II, the smallest of the ETC complexes, in any supercomplex on BNGs, nor did I identify it as an isolated entity.

RNAi inactivation of COX IV and COX Va demonstrated a relationship between supercomplex I:III and I:III:IV, as well as a linked regulation of complex I and IV. The amount of supercomplex I:III increased when the amount of complex

IV and supercomplex I:III:IV decreased (Fig. 15, panels A and C). Similarly to my findings, Schagger et al. demonstrated that a patient with decreased complex IV caused by a SURF1 mutation had a decreased ratio of supercomplex I:III:IV to supercomplex I:III (Schagger et al. 2004). They did not report ETCs for this patient. In the report by Schagger, amounts of complex I did not decrease, even though the residual complex IV levels were only 10% of normal. However,

SURF1 is not a structural subunit of complex IV, but rather a protein involved in its assembly. In studies of patients with missense mutations in COX 10, also a

COX assembly factor (Antonicka et al. 2003), complex I levels were normal. In a separate study, fibroblast lines from two patients with no measurable complex IV

(McKenzie et al. 2007), contained normal amounts of complex I. In contrast,

Diaz et al. (Diaz et al. 2006), found that a knockout of COX10 in a mouse

fibroblast line drastically reduced both supercomplex I:III, and supercomplex

I:III:IV. They proposed that the complete absence of complex IV in a COX10

knockout led to a global decrease of NADH oxidoreductase supercomplexes as

determined by BNGs. Li et al. (Li et al. 2007) established mouse cell lines (Li et

al. 2006) that decreased COX IV levels more than 75% and COX activity by more

than a half as shown by CIV-IGA (Li et al. 2006). These cells exhibited more than a 50% decrease in the levels of two nuclear-encoded complex I subunits, GRIM-

19 and NDUFA9 (Li et al. 2007); supercomplex formation was not investigated.

They concluded that unstable complex assembly, as opposed to decreased

protein synthesis, led to a reduction in complex I subunits. Although D’Aurelio et

al. (D'Aurelio et al. 2006) have argued that a threshold decrease in the level of

complex IV is necessary to decrease complex I level, results from patients would

indicate that even very low levels of complex IV do not necessarily decrease

complex I levels.

In a recent review (Lazarou et al. 2008), it has been concluded that the

role of complex IV in complex I assembly is unclear. It is interesting that in all the

data from intact animals, even animals as far apart in the animal kingdom as

nematodes and humans, defects in complex IV do not decrease amounts of

complex I. However, clearly, in the nematode, low levels of complex IV shift the

ratio of complex I from supercomplex I:III:IV to I:III. Even with normal amounts of

complex I, this shift in supercomplex formation results in a striking loss of

enzymatic activity of complex I itself (see below).

Complex I enzymatic function, as assayed by standard ETC assays, is

decreased in our complex IV knockdowns. ETC assays are not done on

complexes isolated from BNGs, but rather from mitochondria treated with cholate

which allows entry of substrates specific for the complexes studied (e.g. NADH for complex I). It is possible that defects in complex IV destabilize complex I such that cholate treatment negatively affects complex I function. In either case, complex IV has an effect on complex I function that is measured as per standard protocols for patient diagnosis (Sottocasa et al. 1967; Hoppel & Cooper 1969;

Hatefi 1978; Krahenbuhl et al. 1991).

Two assays revealed defects in complex I function: flow of electrons through complex I to the artificial electron acceptor decylubiquinone, and electron transport through complex I, coenzyme Q, and complex III, to cytochrome c (for

COX Va alone). This decreased enzymatic activity was seen despite the fact that the overall amount of complex I, as assayed by Coomassie blue staining, by CI-

IGA activity, or by measurement of NUO-2 levels, was not significantly different in these animals compared to N2. Electron movement within the matrix arm of complex I (NFR), was not affected by knockdown of either complex IV subunit.

Although each RNAi experimental culture varied slightly in amount of measured knockdown, knockdown was always verified either by qPCR or measurements of complex IV dependent respiration. Therefore it is not possible that lack of change in levels of complex I was due to a lack of knockdown in the particular culture. In addition, at least 4 gels from different cultures were used to assess supercomplex organization in knockdown animals.

The total amount of CI-IGA in supercomplexes is not decreased in the

mutants, similar to our finding that NFR of complex I is not decreased in the

knockdown animals. It is not known exactly what step of electron transport is

measured by IGA of BNGs. My data indicate that CI-IGA involves a proximal part

of complex I rather than the entire complex. However, transport of electrons

through the entire complex I, as measured using NADH as an electron donor and

decylubiquinone as the electron receptor (Fig. 13, panel B) is significantly

decreased in both knockdowns. Further complex I-III activity is significantly

decreased in one knockdown and approaches significance in the second

knockdown (p = 0.07). This indicates then that the I:III:IV supercomplex, which is

reduced in both knockdown animals, is crucial for normal rates of electron

transport within complex I.

My results are consistent with the model that supercomplexes represent

an advantage in electron transfer, in this case even affecting movement through

another member of the associated supercomplex. Schafer et al. (Schafer et al.

2006) showed that NADH:ubiquinone reductase was more active in

supercomplex I:III2:IV than in supercomplex I:III2. This is the same step of electron transfer that is measured in our complex I assay. They concluded that complex IV enhanced the function of complex I in supercomplex I:III:IV. Their enzyme assays were done with samples directly electroluted from BNGs. My results corroborate and extend these findings. I have produced a defective complex IV through knockdown of two nuclear genes that encode different subunits of the complex. Both knockdowns lead to a decreased rate of electron

transport through complex I, while leaving enzymatic activity within the matrix

arm unchanged. This is in keeping with the model in which the membrane portion

of complex I, in animals, is in extensive contact with complex IV, while the matrix

arm is not (Vonck & Schafer 2008). I attribute the decreased rates to loss of the

I:III:IV supercomplexes, with relatively decreased activity of the I:III supercomplex in passing electrons within complex I. It appears that complex IV then plays a role in maintaining maximal rates of electron flow within complex I itself through its contact within the membrane bound arm.

Multiple deficiencies of the ETC have been reported in patients in conditions like peripheral arterial disease (I+III (Brass et al. 2001)) or Parkinson’s disease (I+IV+V (Cardellach et al. 1993), I+IV (Benecke et al. 1993)), in multiple

infantile defects (I+IV, I+V (von Kleist-Retzow et al. 2003)), in children with

myopathies (I+IV (Korenke et al. 1990)), and in infants with intractable lactic

acidosis (I+IV (van Straaten et al. 2005)). A combined I/IV defect is cited as the

most common combined deficiency in patients (Korenke et al. 1990). My study

demonstrates that a single primary defect can have wide ranging effects on

function of multiple different protein complexes of the ETC. This finding adds to

the complexity of interpreting ETC assays when used in isolation to diagnose

mitochondrial defects. Patients presenting with puzzling multiple defects of

electron transport involving complexes I and IV may, in fact, harbor a single

genetic defect affecting complex IV.

2.4. Conclusion

Here I show that partial depletion of complex IV as a result of RNAi

knockdown of COX IV and COX Va can lead to combined complex I/IV deficiency

in C. elegans, despite normal level of complex I. Evidence from bovine heart mitochondria indicate that complex I in supercomplex I:III:IV is significantly more active than complex I in supercomplex I:III. Therefore, I suggest that decrease in the relative amount of supercomplex I:III:IV to supercomplex I:III in the RNAi- knockdown worms is the mechanism that underlies the decreased complex I function observed in the complex IV-deficient worms in my study.

2.5. Figures and tables

FIGURE 12

A 400 N2 COX IV COX Va 300 ! 200 !! State 3 rate 100 ! !! ! (nanoAtom (O)/min/mg protein) 0 TMPD/ascorbate Malate Succinate B 400 COX IV COX Va 300

200

respiratory rate 100 TMPD/ascorbate driven (nanoAtomO/min/mg prot)

0 20 40 60 80 100 % knockdown

Figure 12. Deterioration of mitochondrial respiration in the RNAi- knockdown worms. A. The integrated mitochondrial functions of RNAi worms were determined by the OXPHOS assay, using malate, succinate and

TMPD/ascorbate as the electron donors to drive complex I-, complex II- and

complex IV-dependent respirations, respectively. Complex IV-dependent

respiratory rates in both knockdown worms were significantly lower than N2.

Similarly, the state 3 rates of complex I-and complex II-dependent respiration

were significantly lower in the knockdown worms. Error bars represent standard

error of mean (SEM) from four independent isolations. * and ** indicate statistical

significance as p < 0.05 and < 0.005, respectively. B. The RNAi effect on the

target genes (COX IV and COX Va) was measured by quantitative RT-PCR. The

correlation between the target gene expression level and complex IV-dependent

respiratory rate was shown. Respiratory rates were measured as the rate of

disappearance of oxygen in nanoAtomO/min/mg of mitochondrial protein. The

extents of RNAi knockdown were determined by relative quantification of the

level of mRNA expression of the target genes to the housekeeping gene and

then normalized to that of control worms (N2). Error bars represent standard

error of mean (SEM) from four independent isolations. * and ** indicate statistical

significance as p < 0.05 and < 0.005, respectively. Figure adapted from

Suthammarak et al. (Suthammarak et al. 2009)

FIGURE 13

A B C 1200 240 2400 N2 COX IV COX Va 900 180 1800

600 120 ! ! 1200 !!!! 300 60 600 nmole substrate/min/mg prot nmole substrate/min/mg prot nmole substrate/min/mg prot

0 0 0 CIV CI NFR D E F 1200 120 1600

900 ! 90 1200

600 60 800

300 30 400 nmole substrate/min/mg prot nmole substrate/min/mg prot nmole substrate/min/mg prot

0 0 0 CI-III CII-III CIII

Figure 13. The effects of COX knockdown on electron transport chain assays. A. Enzymatic activities of KCN-sensitive cytochrome c oxidase (CIV).

B. Enzymatic activity of rotenone-sensitive NADH-decylubiquinone oxidoreductase (CI). C. Enzymatic activity of NADH-ferricyanide reductase

(NFR). D. Enzymatic activity of rotenone-sensitive NADH-cytochrome-c oxidoreductase (CI-III). E. Enzymatic activity of antimycin A-sensitive succinate- cytochrome c oxidoreductase (CII-III). F. Enzymatic activity of antimycin A-

sensitive decylubiquinol-cytochrome c oxidoreductase (CIII) were spectrophotometrically measured in N2 as wild type and in both complex IV knockdown worms. Error bars represent SEM from three to five independent worm cultures. * and ** indicate statistical significance as p < 0.05 and < 0.01, respectively. Figure adapted from Suthammarak et al. (Suthammarak et al. 2009)

FIGURE 14

A B C MW (kDa) DT TX MS DT TX MS DT TX MS 1236 1048 1000 950 880

800

720

600

500

420

Figure 14. The organization of respiratory complexes in C. elegans.

Mitochondrial proteins from N2 were solubilized by digitonin (DT), Triton X-100

(TX) or dodecyl-maltoside (MS) and were electrophoresed in 3.5-11% gradient acrylamide gel. The gels were subsequently further stained with Coomassie blue in A or used to perform in-gel activity (IGA) assays for complex I (CI-IGA) or complex IV (CIV-IGA) as shown B and C, respectively. The numbers in the left panel indicate the approximate molecular weights (kDa) of the corresponding protein bands. Figure adapted from Suthammarak et al. (Suthammarak et al.

2009)

FIGURE 15

A B C

MW (kDa) N2 COX IVCOX VaN2 COX IVCOX Va N2 COX IVCOX ComplexesVa 1236

1048 I:III:IV 1000 950 I:III 880

800 V

720

600 III

420 IV MW (kDa) Complexes 1236 1048 I:III:IV 1000 I:III 950 880

800 V

720

600 III

420 IV

Figure 15. Blue native electrophoresis and the in-gel activity assay (IGA) of mitochondria from complex IV-deficient worms. Two representative BNGs

(upper and lower) to show the degree of reproducibility in the amount of supercomplexes between isolations. Each gel was loaded in triplicate and cut into thirds for the assays shown in A, B, and C. In each set of three lanes, N2 is

the left lane, COX IV is the middle lane and COX Va is the right lane. A.

Coomassie blue stained BNGs. Twelve distinct blue protein bands were

consistently observed by Coomassie blue staining. Mass spectrometry identified

three bands out of twelve, between molecular weights 450 – 500 KDa, as

containing non-OXPHOS proteins. The other nine bands contained primarily

subunits of MRC complexes and were analyzed as follows: supercomplex I:III:IV,

1,236, 1,048 and 1000 KDa; supercomplex I:III, 950 KDa; complex V, 880, 800

and 720 KDa; complex III, 600 KDa; complex IV, 420 KDa. B. Complex IV-In

Gel Activity (CIV-IGA). The dark brown bands indicate the activity of

cytochrome c oxidase which reduces diaminobenzidine. Note that activity is

found in those bands shown to contain complex IV by mass spectrometry and

that activity is decreased in the knockdown strains. C. Complex I-In Gel

Activity (CI-IGA). The activities of complex I are marked by the deep blue color

of formazan crystals resulted from the reduction of nitroblue tetrazolium by

complex I. The positive bands appearing in CI-IGA corresponded to the

identification of complex I by mass spectrometry. Note that the amount of

complex I staining is relatively unchanged in the knockdown strains. Figure adapted from Suthammarak et al. (Suthammarak et al. 2009)

FIGURE 16

A B 1 6 N2 COX IV COX Va 0.75 ! ! 4.5

0.5 I 3 I:III:IV

total V

I:III + I:III:IV

0.25 1.5

0 0

C D MW (kDa) N2 COX IV COX Va N2 COX IV COX Va Complexes 900 860 I:III

780 I

Figure 16. Complex IV disruption alters the ratio between supercomplex I:III and I:III:IV but not the total amount of complex I. A. The amounts of complex

I as present in supercomplex I:III and I:III:IV were measured from 4 independent digitonin-based CI-IGAs. Quantitative analysis shows significant changes in the amount of CI-IGA in supercomplex I:III to the total amount of CI-IGA in both complex IV-depleted worms compared to N2. B. The total amount of CI-IGA present in both supercomplex I:III and I:III:IV, after normalization to a total complex V is not perturbed by complex IV disruption. C. Triton X-100-based

BNGs also confirm that the total amount of complex I in complex IV-depleted worms is unchanged as shown by Coomassie blue stain. D. Triton X-100-based

BNGs corroborates that the total amount of complex I in complex IV-depleted

worms is unchanged as shown by CI-IGA. Figure adapted from Suthammarak et al. (Suthammarak et al. 2009)

FIGURE 17

A 2 N2 COX IV COX Va 1.5 B N2 COX IV COX Va 1 NUO-2 NUO-2/ANT ANT 0.5

Figure 17. A. Steady state level of the NUO-2 subunit of complex I. A. NUO-

2/ANT ratios in both complex IV knockdown worms were unchanged compared

to that of N2. Error bars represent SEM from four independent experiments. B.

Western blot analysis of mitochondrial protein probed with an antibody to

the NUO-2 bovine orthologue. 25 ug of mitochondrial protein from the wild-type worm, COX IV knockdown and COX Va knockdown worms were resolved by western blot in the left, the middle and the right lane, respectively. The amounts of probable NUO-2 subunit and adenosine nucleotide translocase (ANT), as a loading control, were determined by monoclonal antibody against each protein.

Figure adapted from Suthammarak et al. (Suthammarak et al. 2009)

TABLE 1

Phenotype N2 COX IV COX Va Mean Lifespan (days) 17 + 1.1 14 + .9 12 + 1.5 Fecundity (eggs/worm) 254 + 22 124 + 14 107 + 17 Days to Adulthood (days) 3 + 0.1 4 + 0.3 4 + 0.4 EC50 (halothane) 3.2 + 0.2 2.9 + 0.2 2.8 + 0.2

Table 1: Phenotypical study of COX IV and COX Va knockdown worms. The phenotypes of worms after two generations of exposure to RNAi. Table adapted

from Suthammarak et al. (Suthammarak et al. 2009)

TABLE 2

C. elegans Protein band’s molecular weight (kDa) Human’s homolog peptide (HGNC’s symbol) (CGC’s name) 1,236 1,048 1,000 950 880 800 720 600 420

NUO-1 NDUFV1 + + + + NUO-2 NDUFS3 + + + + NUO-4 NDUFA10 + + + + NUO-5 NDUFS1 + + + + F53F4.10 NDUFV2 + + + + C33A12.1 NDUFA5 + + + + Y54F10AM.5 NDUFA8 + + + + Y53G8AL.2 NDUFA9 + + + + Y94H6A.8 NDUFA12 + + + + C25H3.9a NDUFB5 + + + + ZK809.3 NDUFB6 + + + + Y51H1A.3b NDUFB8 + + + + C16A3.5 NDUFB9 + + + + F59C6.5 NDUFB10 + + + + GAS-1 NDUFS2 + + + + Y54E10BL.5 NDUFS5 + + + + W01A8.4 NDUFB4 + + + T20H4.5 NDUFS8 + + C18E9.4 NDUFB3 + + F42G8.10a NDUFB11 + + NUO-3 NDUFA6 + + + W10D5.2 NDUFS7 + + + F45H10.3 NDUFA7 + + TAG-99 NDUFS2 + F44G4.2 NDUFB2 + D2030.4 NDUFB7 + UCR-2.1 UQCRC2 + + + + + UCR-2.2 UQCRC2 + + + + + UCR-2.3 UQCRC2 + + + + + T02H6.11 UQCRB + + + + + F45H10.2 UQCRQ + + + + + W09C5.8 COX4I1 + + + + CCO-2 COX5A + + + + CCO-1 COX5B + + + TAG-174 COX6A + + Y71H2AM.5 COX6B1 + + F26E4.6 COX7C + + H28O16.1d ATP5A1 + + + + + + ATP-2 ATP5B + + + + + + + Y69A2AR.18a ATP5C1 + + + + + ATP-5 ATP5H + + + + ASB-2 ATP5F1 + + + ASB-1 ATP5F1 + + + ASG-2 ATP5L + + + ASG-1 ATP5L + + + R04F11.2 ATP5I + + + F58F12.1 ATP5D + + + + F32D1.2 ATP5E + + + + R05D3.6 ATP5E +

Table 2. Respiratory chain proteins identified by mass spectrometry in each protein band in digitonin-based BNGs. Only MRC proteins are listed.

Protein names are obtained from C. elegans Genetics Center (CGC) and HUGO

Gene Nomenclature Committee (HGNC), respectively. Absence of a subunit does not preclude its presence in the protein band as mass spectrometry may miss individual proteins. These data are used to indicate which complexes are in each band and not to identify all subunits present. Turquoise shading indicates complex I subunits, yellow complex III, pink complex IV, and grey complex V.

Table adapted from Suthammarak et al. (Suthammarak et al. 2009)

TABLE 3

C. elegans Human’s Protein band’s molecular weight (kDa) peptide homolog (CGC’s name) (HGNC’s symbol) 900 860 780 700 550 530 450

NUO-1 NDUFV1 + + + NUO-2 NDUFS3 + + + NUO-3 NDUFA6 + NUO-4 NDUFA10 + + + NUO-5 NDUFS1 + + + C33A12.1 NDUFA5 + + + Y54F10AM.5 NDUFA8 + + + Y53G8AL.2 NDUFA9 + + + Y94H6A.8 NDUFA12 + + + W01A8.4 NDUFB4 + ZK809.3 NDUFB6 + + + Y51H1A.3b NDUFS8 + C16A3.5 NDUFB9 + + + F59C6.5 NDUFB10 + + + Y71H2AM.4 NDUFC2 + + GAS-1 NDUFS2 + + + Y54E10BL.5 NDUFS5 + + W10D5.2 NDUFS7 + + + T20H4.5 NDUFB8 + + + F53F4.10 NDUFV2 + + UCR-1 UQCRC1 + + UCR-2.1 UQCRC2 + + UCR-2.2 UQCRC2 + + T02H6.11 UQCRB + F45H10.2 UQCRQ + CYC-1 CYC1 + + ISP-1 UQCRFS1 H28O16.1d ATP5A1 + + + + ATP-2 ATP5B + + + + ASB-2 ATP5F1 + ATP-3 ATP5O + ATP-4 ATP5J + ATP-5 ATP5H + Y69A2AR.18a ATP5C1 + + + ASB-1 ATP5F1 + ASG-1 ATP5L + R53.4 ATP5J2 + F58F12.1 ATP5D + + +

Table 3. Respiratory chain proteins identified by mass spectrometry in each protein band in Triton X-100-based BNGs. Only MRC proteins are listed.

Protein names are obtained from C. elegans Genetics Center (CGC) and HUGO

Gene Nomenclature Committee (HGNC), respectively. Absence of a subunit

does not preclude its presence in the protein band as mass spectrometry may miss individual proteins. These data are used to indicate which complexes are in each band and not to identify all subunits present. Turquoise shading indicates complex I subunits, yellow complex III, and grey complex V. Table adapted from

Suthammarak et al. (Suthammarak et al. 2009)

2.6. References

Acin-Perez R, Bayona-Bafaluy MP, Fernandez-Silva P, Moreno-Loshuertos R, Perez-Martos A, Bruno C, Moraes CT, Enriquez JA (2004). Respiratory complex III is required to maintain complex I in mammalian mitochondria. Mol Cell. 13, 805-815. Antonicka H, Leary SC, Guercin GH, Agar JN, Horvath R, Kennaway NG, Harding CO, Jaksch M, Shoubridge EA (2003). Mutations in COX10 result in a defect in mitochondrial heme A biosynthesis and account for multiple, early-onset clinical phenotypes associated with isolated COX deficiency. Hum Mol Genet. 12, 2693-2702. Arnold S, Goglia F, Kadenbach B (1998). 3,5-Diiodothyronine binds to subunit Va of cytochrome-c oxidase and abolishes the allosteric inhibition of respiration by ATP. Eur J Biochem. 252, 325-330. Benecke R, Strumper P, Weiss H (1993). Electron transfer complexes I and IV of platelets are abnormal in Parkinson's disease but normal in Parkinson- plus syndromes. Brain. 116 ( Pt 6), 1451-1463. Bianchi C, Genova ML, Parenti Castelli G, Lenaz G (2004). The mitochondrial respiratory chain is partially organized in a supercomplex assembly: kinetic evidence using flux control analysis. J Biol Chem. 279, 36562- 36569. Blakely EL, Mitchell AL, Fisher N, Meunier B, Nijtmans LG, Schaefer AM, Jackson MJ, Turnbull DM, Taylor RW (2005). A mitochondrial cytochrome b mutation causing severe respiratory chain enzyme deficiency in humans and yeast. FEBS J. 272, 3583-3592. Brass EP, Hiatt WR, Gardner AW, Hoppel CL (2001). Decreased NADH dehydrogenase and ubiquinol-cytochrome c oxidoreductase in peripheral arterial disease. Am J Physiol Heart Circ Physiol. 280, H603-609. Bruno C, Santorelli FM, Assereto S, Tonoli E, Tessa A, Traverso M, Scapolan S, Bado M, Tedeschi S, Minetti C (2003). Progressive exercise intolerance associated with a new muscle-restricted nonsense mutation (G142X) in the mitochondrial cytochrome b gene. Muscle Nerve. 28, 508-511. Budde SM, van den Heuvel LP, Janssen AJ, Smeets RJ, Buskens CA, DeMeirleir L, Van Coster R, Baethmann M, Voit T, Trijbels JM, Smeitink JA (2000). Combined enzymatic complex I and III deficiency associated with mutations in the nuclear encoded NDUFS4 gene. Biochem Biophys Res Commun. 275, 63-68. Cardellach F, Marti MJ, Fernandez-Sola J, Marin C, Hoek JB, Tolosa E, Urbano- Marquez A (1993). Mitochondrial respiratory chain activity in skeletal muscle from patients with Parkinson's disease. Neurology. 43, 2258-2262. Cardol P, Boutaffala L, Memmi S, Devreese B, Matagne RF, Remacle C (2008). In Chlamydomonas, the loss of ND5 subunit prevents the assembly of whole mitochondrial complex I and leads to the formation of a low abundant 700 kDa subcomplex. Biochim Biophys Acta. 1777, 388-396. Chance B, Williams GR (1955). A method for the localization of sites for oxidative phosphorylation. Nature. 176, 250-254.

Chazotte B, Hackenbrock CR (1988). The multicollisional, obstructed, long-range diffusional nature of mitochondrial electron transport. J Biol Chem. 263, 14359-14367. Cruciat CM, Brunner S, Baumann F, Neupert W, Stuart RA (2000). The cytochrome bc1 and cytochrome c oxidase complexes associate to form a single supracomplex in yeast mitochondria. J Biol Chem. 275, 18093- 18098. D'Aurelio M, Gajewski CD, Lenaz G, Manfredi G (2006). Respiratory chain supercomplexes set the threshold for respiration defects in human mtDNA mutant cybrids. Hum Mol Genet. 15, 2157-2169. Diaz F, Fukui H, Garcia S, Moraes CT (2006). Cytochrome c oxidase is required for the assembly/stability of respiratory complex I in mouse fibroblasts. Mol Cell Biol. 26, 4872-4881. Dudkina NV, Eubel H, Keegstra W, Boekema EJ, Braun HP (2005). Structure of a mitochondrial supercomplex formed by respiratory-chain complexes I and III. Proc Natl Acad Sci U S A. 102, 3225-3229. Eubel H, Heinemeyer J, Braun HP (2004). Identification and characterization of respirasomes in potato mitochondria. Plant Physiol. 134, 1450-1459. Eubel H, Jansch L, Braun HP (2003). New insights into the respiratory chain of plant mitochondria. Supercomplexes and a unique composition of complex II. Plant Physiol. 133, 274-286. Genova ML, Bianchi C, Lenaz G (2003). Structural organization of the mitochondrial respiratory chain. Ital J Biochem. 52, 58-61. Grad LI, Lemire BD (2006). Riboflavin enhances the assembly of mitochondrial cytochrome c oxidase in C. elegans NADH-ubiquinone oxidoreductase mutants. Biochim Biophys Acta. 1757, 115-122. Hackenbrock CR, Chazotte B, Gupte SS (1986). The random collision model and a critical assessment of diffusion and collision in mitochondrial electron transport. J Bioenerg Biomembr. 18, 331-368. Hatefi Y (1978). Preparation and properties of NADH: ubiquinone oxidoreductase (complex I), EC 1.6.5.3. Methods Enzymol. 53, 11-14. Hoppel C, Cooper C (1969). An improved procedure for preparation of inner membrane vesicles from rat liver mitochondria by treatment with digitonin. Arch Biochem Biophys. 135, 173-183. Kadenbach B, Frank V, Rieger T, Napiwotzki J (1997). Regulation of respiration and energy transduction in cytochrome c oxidase isozymes by allosteric effectors. Mol Cell Biochem. 174, 131-135. Korenke GC, Bentlage HA, Ruitenbeek W, Sengers RC, Sperl W, Trijbels JM, Gabreels FJ, Wijburg FA, Wiedermann V, Hanefeld F, et al. (1990). Isolated and combined deficiencies of NADH dehydrogenase (complex I) in muscle tissue of children with mitochondrial myopathies. Eur J Pediatr. 150, 104-108. Krahenbuhl S, Chang M, Brass EP, Hoppel CL (1991). Decreased activities of ubiquinol:ferricytochrome c oxidoreductase (complex III) and ferrocytochrome c:oxygen oxidoreductase (complex IV) in liver

mitochondria from rats with hydroxycobalamin[c-lactam]-induced methylmalonic aciduria. J Biol Chem. 266, 20998-21003. Krause F, Reifschneider NH, Goto S, Dencher NA (2005). Active oligomeric ATP synthases in mammalian mitochondria. Biochem Biophys Res Commun. 329, 583-590. Lazarou M, Thorburn DR, Ryan MT, McKenzie M (2008). Assembly of mitochondrial complex I and defects in disease. Biochim Biophys Acta. Lee SS, Lee RY, Fraser AG, Kamath RS, Ahringer J, Ruvkun G (2003). A systematic RNAi screen identifies a critical role for mitochondria in C. elegans longevity. Nat Genet. 33, 40-48. Li Y, D'Aurelio M, Deng JH, Park JS, Manfredi G, Hu P, Lu J, Bai Y (2007). An assembled complex IV maintains the stability and activity of complex I in mammalian mitochondria. J Biol Chem. 282, 17557-17562. Li Y, Park JS, Deng JH, Bai Y (2006). Cytochrome c oxidase subunit IV is essential for assembly and respiratory function of the enzyme complex. J Bioenerg Biomembr. 38, 283-291. McKenzie M, Lazarou M, Thorburn DR, Ryan MT (2007). Analysis of mitochondrial subunit assembly into respiratory chain complexes using Blue Native polyacrylamide gel electrophoresis. Anal Biochem. 364, 128- 137. Napiwotzki J, Kadenbach B (1998). Extramitochondrial ATP/ADP-ratios regulate cytochrome c oxidase activity via binding to the cytosolic domain of subunit IV. Biol Chem. 379, 335-339. Napiwotzki J, Shinzawa-Itoh K, Yoshikawa S, Kadenbach B (1997). ATP and ADP bind to cytochrome c oxidase and regulate its activity. Biol Chem. 378, 1013-1021. Scacco S, Petruzzella V, Budde S, Vergari R, Tamborra R, Panelli D, van den Heuvel LP, Smeitink JA, Papa S (2003). Pathological mutations of the human NDUFS4 gene of the 18-kDa (AQDQ) subunit of complex I affect the expression of the protein and the assembly and function of the complex. J Biol Chem. 278, 44161-44167. Schafer E, Seelert H, Reifschneider NH, Krause F, Dencher NA, Vonck J (2006). Architecture of active mammalian respiratory chain supercomplexes. J Biol Chem. 281, 15370-15375. Schagger H (2001). Respiratory chain supercomplexes. IUBMB Life. 52, 119- 128. Schagger H, de Coo R, Bauer MF, Hofmann S, Godinot C, Brandt U (2004). Significance of respirasomes for the assembly/stability of human respiratory chain complex I. J Biol Chem. 279, 36349-36353. Schagger H, Pfeiffer K (2000). Supercomplexes in the respiratory chains of yeast and mammalian mitochondria. EMBO J. 19, 1777-1783. Schagger H, Pfeiffer K (2001). The ratio of oxidative phosphorylation complexes I-V in bovine heart mitochondria and the composition of respiratory chain supercomplexes. J Biol Chem. 276, 37861-37867. Sottocasa GL, Kuylenstierna B, Ernster L, Bergstrand A (1967). An electron- transport system associated with the outer membrane of liver

mitochondria. A biochemical and morphological study. J Cell Biol. 32, 415- 438. Stroh A, Anderka O, Pfeiffer K, Yagi T, Finel M, Ludwig B, Schagger H (2004). Assembly of respiratory complexes I, III, and IV into NADH oxidase supercomplex stabilizes complex I in Paracoccus denitrificans. J Biol Chem. 279, 5000-5007. Suthammarak W, Yang YY, Morgan PG, Sedensky MM (2009). Complex I function is defective in complex IV-deficient Caenorhabditis elegans. J Biol Chem. 284, 6425-6435. van Straaten HL, van Tintelen JP, Trijbels JM, van den Heuvel LP, Troost D, Rozemuller JM, Duran M, de Vries LS, Schuelke M , Barth PG (2005). Neonatal lactic acidosis, complex I/IV deficiency, and fetal cerebral disruption. Neuropediatrics. 36, 193-199. von Kleist-Retzow JC, Cormier-Daire V, Viot G, Goldenberg A, Mardach B, Amiel J, Saada P, Dumez Y, Brunelle F, Saudubray JM, Chretien D, Rotig A, Rustin P, Munnich A , De Lonlay P (2003). Antenatal manifestations of mitochondrial respiratory chain deficiency. J Pediatr. 143, 208-212. Vonck J, Schafer E (2008). Supramolecular organization of protein complexes in the mitochondrial inner membrane. Biochim Biophys Acta. Yoshikawa S (1997). Beef heart cytochrome c oxidase. Curr Opin Struct Biol. 7, 574-579.

Chapter 3

Mutations in mitochondrial complex III

uniquely affect complex I in Caenorhabditis

elegans

(The work in this chapter has been accepted for publication in the Journal

of Biological Chemistry on October 22, 2010)

3.1. Introduction

A solid-state model (Chance & Williams 1955) of the mitochondrial

respiratory chain (MRC) within the mitochondrial membrane was proposed a

half-century ago. In this model the respiratory complexes are assembled into

multi-complex structures, supercomplexes. Supercomplexes are capable of

substrate channeling, and thus facilitate transfer of electrons from one complex

to the next (Bianchi et al. 2004). This is in contrast to the random collision model

(Hackenbrock et al. 1986), which proposes that the complexes of the MRC are embedded in the inner mitochondrial membrane as separate entities. Individual complexes are functionally connected to each other by the small, mobile electron carriers, CoQ and cytochrome c. The random collision model became more generally accepted as kinetic studies demonstrated homogenous pool behavior of CoQ (Kroger & Klingenberg 1973), rates of electron transfer that did not require substrate channeling, and the successful isolation of individual respiratory complexes that were enzymatically active (Hatefi et al. 1962; Hackenbrock et al.

1986). However, supercomplex structures containing complexes I, III, and IV,

have been investigated by blue native polyacrylamide gel electrophoresis (BN-

PAGE), sucrose gradient centrifugation and single particle analysis (Dudkina et

al. 2005; Schafer et al. 2007). A recent study by Acin-Perez et al. (Acin-Perez et al. 2008) convincingly showed that supercomplexes are functional units capable of consuming oxygen when provided appropriate electron donors. They concluded that supercomplexes are in fact the functional respiratory unit of the mitochondrion in vivo. Schagger and Pfeiffer demonstrated that supercomplex

I:III:IV appears on blue native gels (BNGs) with increasing stoichiometries of

complex IV. They termed these entities S0-S4 where the numeral denotes the

number of complex IVs within the supercomplex (Schagger & Pfeiffer 2000).

Supercomplexes provide a logical explanation for combined deficiencies

of the electron transport chain (ETC) observed in patients. It is estimated that

about 30% of all ETC disorders involve multiple complexes (Smits et al. 2010).

Combined I:IV deficiencies (Korenke et al. 1990; Benecke et al. 1993; Brass et

al. 2001; van Straaten et al. 2005), as well as multiple examples of combined I:III

deficiencies have been reported (Budde et al. 2000; Lamantea et al. 2002;

Schagger et al. 2004). In most eukaryotes, complex III is composed of 11 subunits (Xia et al. 1997; Iwata et al. 1998), of which only cytochrome b is encoded by the mitochondrial genome. The catalytic core of complex III is comprised of cytochrome b, ISP (iron sulfur protein; Rieske protein) and cytochrome c1. Most of the mutations in cytochrome b cause isolated complex III deficiencies. However, there is a subset of cytochrome b mutations that cause combined I:III deficiencies, both in patients (Budde et al. 2000; Lamantea et al.

2002; Schagger et al. 2004; Blakely et al. 2005) and in mammalian cell lines

(Acin-Perez et al. 2004). In addition, loss of cytochrome c has been reported to lead to loss of both complexes I and IV in cell lines (Vempati et al. 2009). From these findings, it has been suggested that fully assembled complex III, functional or not, is required for the assembly and stability of complex I. Notably, no structural mutations in ISP or cytochrome c1 have been reported in animals.

However complex III deficiencies in human patients are caused by mutations in

BCS1L (de Lonlay et al. 2001; De Meirleir et al. 2003; Fernandez-Vizarra et al.

2007), a chaperon of ISP in the nascent complex III (Nobrega et al. 1992; Cruciat et al. 1999). There are no reports that defects in complex III can affect the function of fully assembled complex I without decreasing the amount of complex

In C. elegans, two different complex III mutants confer intriguing phenotypes. isp-1 changes a highly conserved amino acid (P225S) residue in the head domain of ISP. Hekimi’s group reported that isp-1 exhibits low oxygen consumption, decreased sensitivity to ROS, extended lifespan, and delayed embryonic development (Feng et al. 2001). A mutation (ctb-1) in a complex III subunit cytochrome b, A170V, was discovered as a suppressor of a delayed development phenotype of isp-1. isp-1;ctb-1 shows an improvement in the rate of development compared to isp-1 (Feng et al. 2001). However, the molecular cause of this improved phenotype is unknown.

I show here that a structural defect of complex III, caused by a mutation in

ISP, can reduce the amount of fully assembled complex I without a reduction in the amount of fully assembled complex III. Furthermore, I demonstrate that ctb-1 can decrease complex I function without decreasing either the amount of fully assembled complex I, or the distribution of complex I in supercomplexes. In addition, I show that the improved whole animal phenotypes of isp-1;ctb-1 compared to isp-1 alone, do not stem from improved complex III function. Rather salutary allosteric effects on complex I result from the introduction of a second mutated subunit into complex III. In addition, this double mutant heightens the

weakened association of supercomplex I:II:IV such that the composition of

supercomplexes is altered on BNGs. The detailed mechanisms of complex I and

III interdependence seen in a model system indicate that supercomplexes play a

major role in modulating intrinsic enzymatic activities of complexes I and III. The complexity of the interdependence between components of the ETC has important implications for the diagnosis of patients with defects in multiple complexes.

3.2 Results

3.2.1. Phenotypes of complex III mutants- delayed embryonic development and

egg laying defect

All complex III mutants that hatched did so within 24 hours of being laid.

The subsequent rate of development, however, was severely affected. Wild type

(N2) and ctb-1 reached adulthood at about 3.5 days. isp-1 and isp-1;ctb-1 required 7.5 and 5 days to develop from eggs to adults, respectively. Specifically,

L3 to L4 development lasted approximately 12 hours for N2 and ctb-1, and 24 hours for isp-1 and isp-1;ctb-1. The development from L4 to adult in N2, isp-

1;ctb-1 and ctb-1 was complete within 12 hours, whereas isp-1 required another

24 hours for L4s to develop into adults (Table 4).

Complex III mutations also affected the number of eggs laid, with isp-1 < isp-1;ctb-1

3.2.2. Effect of complex III mutations on integrated mitochondrial respiration

The state 3 rates of both complex I- and complex II-dependent respiration of all complex III mutants were significantly decreased compared to that of wild type. Complex IV-dependent respiration rates of isp-1 and isp-1;ctb-1 were greater than wild type, (Fig. 19). Overall, mitochondrial respiration in complex III mutants was consistent with a complex III deficiency.

3.2.3. Respiratory enzyme complex activity in complex III mutants

A large decrease in CIII (antimycin A-sensitive decylubiquinol-cytochrome

c reductase) activity was observed in isp-1, (Fig. 20, panel A) the mutant that

presented the slowest rate of development. Surprisingly, this same decrease in

CIII activity was also observed in isp-1;ctb-1, although this mutant had a much

improved phenotype compared to isp-1. CIII activity in ctb-1 was greater than

isp-1 or isp-1;ctb-1, but was still significantly less than wild type. Compared to

N2, all complex III mutants exhibited significantly decreased rotenone-sensitive

CI (NADH-decylubiquinone reductase) activities (Fig. 20, panel B). However, CI

activities of both isp-1;ctb-1 and ctb-1 were significantly improved from that of

isp-1. NFR (NADH-ferricyanide reductase) activities were decreased only in isp-

1, whereas isp-1;ctb-1 and ctb-1 had normal NFR activities (Fig. 20, panel C). CI-

III activities (Fig. 20, panel D) were significantly lower in all complex III mutants

than in wild type, consistent with a complex III defect. CII-III activity of isp-1;ctb-1

was significantly lower than that of wild type (Fig. 20, panel E). CII activities in

complex III mutants were normal (Fig. 20, panel F). COX (CIV) activities,

although higher in absolute value, were not significantly different than those for

100

N2 in any complex III mutants (Fig. 20, panel G). In summary, the only

improvement we measured in mitochondrial function in isp-1;ctb-1 compared to

isp-1 was improved CI function.

3.2.4. Amounts of fully assembled respiratory complexes in complex III mutants

The amount of fully assembled complex I was determined from both Triton

X-100-based and digitonin-based BNGs as shown in Fig. 21, panels A, and Fig.

23, panel B, respectively (see Methods). The amounts of complex I in

supercomplexes were normalized to the total amount of complex V as a loading

control and presented as fold changes relative to wild type. Among the complex

III mutants, only isp-1 had a decreased amount of fully assembled complex I,

which was 50% that of N2 (Fig. 21, panel C).

Native-Western blots of digitonin-based BNGs (n=3) probed with anti-

Rieske mAb were used to measure fully assembled complex III (representative

blots are shown in Fig. 21, panels B and C). The amount of fully assembled

complex III was the same in the mutants as in N2 (Fig. 21, panel C). We

observed another signal from the position of dimeric complex IV. This signal was

not included in the quantification since its molecular weight indicates that it is a

partially formed III2 co-migrating with dimeric complex IV. Identical results were obtained using an antibody to the complex III Core 2 protein (data not shown). In addition, we quantified the amount of fully assembled complex IV by densitometry scanning of in gel activity of digitonin BNGs. Complex IV was not significantly increased in any of the mutants (Fig. 21 panel C and Fig. 23, panel

101

C). Western blots from SDS-Western corroborated our results from native-

Westerns (Fig. 22 and Table 5).

3.2.5. The effects of isp-1 on supercomplexes

isp-1 decreased amounts of both I:III and I:III:IV supercomplexes as seen

on digitionin-based BNGs (Fig. 23, panels A-C). III2 appeared more diffuse in

isp-1 than in N2 (Fig. 23, panels A and D), consistent with native-Western blot

analysis (Fig. 21, panel B).

Coomassie staining and Western blots of digitonin-based BNGs revealed

three unique bands in isp-1 which were not readily discernable in wild type

mitochondria (Fig. 23, panel D). These bands were labeled as ISP-A, ISP-B and

ISP-C. The components of ISP-A and ISP-B were identified by mass

spectrometry (Table 6) and consisted of subunits of complexes I, III and IV. In keeping with our proteomic data, ISP-A stained negative for CI-IGA (Fig. 23, panel B), suggesting that it lacks the N module of complex I (NADH-oxidase module, see (Lazarou et al. 2009) for a review). Similarly, ISP-B is an intermediate that consisted of most of the P module (proton pumping module, subcomplex Iβ, see (Lazarou et al. 2009)), and the Q module (coenzyme Q reduction module, see (Lazarou et al. 2009)) as determined by the presence of the NDUFS3 subunit (Fig. 3D, right panel). ISP-C was not characterized by mass spectrometry due to its close proximity to IV2, but was most likely the Q and P modules incorporated into a subcomplex, as determined by its size and the presence of NDUFS3 subunit (Fig. 23, panel D right).

3.2.6. Supercomplex profiles in complex III mutants

102

Since decreased CI ETC activities in isp-1;ctb-1 and ctb-1 are not due to decreased amounts of complex I, we asked if changes in their supercomplex profiles might underlie a complex I deficiency. In mammalian mitochondria, complex I is much less active in the I:III form than in supercomplex I:III:IV

(Schafer et al. 2006). Identifications were inferred from our previous proteomic data for wild type and from in gel activity staining for complexes I and IV

(Suthammarak et al. 2009). isp-1;ctb-1, displayed increased amounts of I:III supercomplex and decreased amounts of I:III:IV supercomplexes as determined by optical density scanning of digitonin-based BNGs (representative gels in Figs.

23, panels A and B, quantification in Fig. 23, panel E). In contrast, the supercomplex profile of ctb-1 was in every way identical to that of wild type (Fig.

23, panel E).

3.2.7. Complex I function from electroeluted supercomplexes

The decrease in CI activity in ctb-1 was not explained by a decrease in either total complex I or in the ratio of complex I:III:IV to total complex I. However, as in mammalian mitochondria (Schafer et al. 2006), after electroelution from

BNGs, complex I in I:III:IV supercomplexes was significantly more active than

complex I in the I:III supercomplex from N2. Complex I in the I:III supercomplex

of isp-1;ctb-1 and ctb-1 was as active as that of wild type. Surprisingly however,

complex I in I:III:IV supercomplexes from isp-1;ctb-1 and ctb-1 were significantly

less active than in N2 (Fig. 24).

103

3.2.8. The stability of supercomplexes

If components of the supercomplex are displaced because of altered subunits, the strength of association between components supercomplex might be altered. Dodecyl maltoside dissociated increased amount of complex IV from the I:III:IV supercomplexes (S1-S4 differ by the number of complex IVs in the supercomplex) of isp-1;ctb-1 and ctb-1 from the 1D-gel (digitonin-treated) into smaller supercomplexes in the second dimension (Fig. 25, panel A). The original

S4 from the 1D-gel was partially dissociated into 3 smaller supercomplexes whose molecular weights were comparable to the original S3, S2 and S1 that migrated from the 1D-gel. Similarly, S3, S2, and S1 were partially dissociated and gave the smaller supercomplexes. All dissociated supercomplexes stained positive for complex I by in gel staining (data not shown). Silver staining visualized a band in the second dimension (Fig. 25, panel B) at the size expected for dimeric complex IV, below the smaller, disassociated supercomplexes. In wild type mitochondria, dissociated supercomplexes and dimeric IV also appeared in the second dimension, but not to the extent of the mutants (Fig. 25, panels A and

B). Quantitative measurement of the amount of complex IV2 (normalized to the amount of complex V) indicated a significant loss of complex IV from the I:III:IV supercomplexes in isp-1;ctb-1 and ctb-1, compared to wild type (Fig. 25, panel

C).

3.2.9. The effect of sodium cholate on mitochondrial supercomplexes

The decrease in complex I activity that was observed in the complex III mutants was determined by electron transport chain assays, in which

104

mitochondria were solubilized by sodium cholate. However, the explanation for decreased complex I activity was based upon supercomplex structure that was observed in BNGs where mitochondria were solubilized by digitonin. Therefore, it is crucial to demonstrate that the cholate that is used in electron transport chain assay does not disrupt supercomplex structure or break apart supercomplexes.

Figure 26 illustrates BNGs that were performed on wild-type mitochondria solubilized by digitonin, cholate or both. Digitonin was titrated to a digitonin/protein mass ratio of 6/1. The amount of sodium cholate was the same as that in electron transport chain assays, which is 1% in final concentration. I found that cholate alone did not extract supercomplexes from the inner mitochondrial membranes. However, abundant material that stained dark blue with Coomassie remained at the bottom of the well of the stacking gel (lane 2 in

Fig. 26). This material retained complex I and IV activities as determined by IGAs

(lane 2 in Fig. 26, panels B and C). When mitochondria were treated with cholate for 10 minutes, followed by digitonin solubilization, a few bands containing complexes I and IV were extracted and were observed in BNGs (lane 3 in Fig.

26). Those bands were most likely supercomplexes according to the migration on the gel. There were still a considerable amount of the material remaining at the bottom of the well, similar to lane 2. In contrast, when mitochondria were treated with digitonin followed by cholate (lane 4 in Fig. 26), the band pattern was identical to that of digitonin-treated BNG in lane 1. This result shows that whereas cholate lacks the ability to extract supercomplexes from the inner mitochondrial membranes (lane 2), it does not appreciably disrupt

105

supercomplexes (lane 4). The material at the bottom of the well of the sample gel in lanes 2 and 3 suggests that cholate might simply break apart the inner mitochondrial membranes into smaller pieces. Those pieces contain complexes I and IV, most likely in a form of supercomplexes, but are too large to enter the gel.

3.3. Discussion

I have shown that, in the nematode, primary changes in complex III can significantly affect the function of complex I. This can occur without decreasing the amount of complex III (isp-1) or the amount or the distribution of complex I in supercomplexes (ctb-1).

Originally I hypothesized that in isp-1;ctb-1, ctb-1 would suppress the slow

development of isp-1 (Feng et al. 2001) by restoring complex III activity since in

general a genetic suppressor attenuates negative effect of a causative mutation

by improving function of a protein that they both involve. Surprisingly, compared

to isp-1, complex III activity in isp-1;ctb-1 was not improved by the suppressor

mutation, ctb-1. Rather complex I activity was improved, which was the only

improvement in mitochondrial function that we could measure. Our results

indicate that improving physical interaction between complexes I and III in

supercomplexes is the basis of ctb-1’s suppression of the severe developmental

phenotype of isp-1.

Development from L3 to L4, and L4 to adult is accompanied by a dramatic

increase in mitochondrial DNA content (Tsang & Lemire 2002; Bratic et al. 2009),

106

which has been linked to increased cellular energy expenditure (Tsang & Lemire

2002). isp-1 and isp-1;ctb-1 spent twice the time in L3-L4 as did ctb-1 or wild

type, implying that the mitochondrial deficiencies in isp-1 and isp-1;ctb-1 are

each more severe than in ctb-1. These results were corroborated by decreased

fecundity in isp-1 and isp-1;ctb-1. Interestingly, although ctb-1 developed at the

same rate and laid a similar number of eggs as wild type, (Table 4), its maximum

rate of egg laying was 1 day later than wild type. This was the only whole animal

phenotype observed in ctb-1, which we interpret it as a mild form of mitochondrial

deficiency.

The isp-1 allele we studied is a serine substitution of a highly conserved

proline, which is located in the head domain of ISP (Iwata et al. 1998). This

proline residue has been shown to be crucial for the secondary structure of the

[2Fe-2S] cluster, for it creates an inward folding of the backbone of the Rieske

head domain immediately preceding strand β5 (Iwata et al. 1996). Such an amino acid change could negatively affect redox properties of the ISP by altering the position of the [2Fe-2S] residues and significantly impede complex III activity.

Furthermore, a P146L substitution (a position that is equivalent to the worm’s isp-

1 allele in our study) in S. cerevisiae was found to alter the midpoint potential of the [2Fe-2S] cluster (Gatti et al. 1989). Therefore it was not unexpected to observe a large decrease in complex III activity of the isp-1 mutant. isp-1(qm150) is the first example of a defect in the ISP itself that affects complex III function in animals. All of the complex III deficiencies in animals, which are associated with the ISP, are reportedly caused by the mutations of BCS1L (de Lonlay et al. 2001;

107

De Meirleir et al. 2003; Fernandez-Vizarra et al. 2007), a chaperon protein involved in ISP synthesis.

The ctb-1 mutation is a conservative alanine to valine substitution at the N-

terminal of helix αD of cytochrome b that is adjacent to the docking site (helix αC,

helix αcd1 and loop ef) of the Rieske head domain during electron transfer (Iwata et al. 1998). The mutated alanine is not a highly conserved residue and may not significantly alter the conformation of cytochrome b. Thus, it is not surprising that the decrease in complex III activity in ctb-1 was not as dramatic as in isp-1. In

addition, the locations of the ctb-1 and isp-1 mutations are not close (Xia et al.

1997; Iwata et al. 1998), and not predicted to directly interact. Therefore it may

not be surprising that the two mutations do not combine to repair complex III

function. All complex III mutants had normal amounts of fully assembled complex

III, but decreased complex III activity, including the conservative change in ctb-1,

which significantly affected complex III function.

Only isp-1 decreased the amount of fully assembled complex I, which

undoubtedly led to its decreased CI activity compared to N2. CI activity in isp-

1;ctb-1 is decreased relative to N2 despite having normal amounts of CI.

However, I have shown that complex I is normally more active in the I:III:IV

supercomplex than in the I:III form, as in bovine heart (Schafer et al. 2006). Thus,

in the double mutant the decreased amount of supercomplex I:III:IV relative to

supercomplex I:III is likely responsible for the decreased complex I activity, as is

the case for COX mutations in C. elegans (Suthammarak et al. 2009). It most

likely represents a increase of the detrimental effect of ctb-1 on complex IV

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binding within the supercomplex (see below). Nevertheless, the increased

amount of total complex I in the double mutant is sufficient to improve complex I

function compared to isp-1 alone.

ctb-1 has decreased complex I activity despite a wild-type distribution of supercomplex I:III and I:III:IV, and a normal amount of complex I. Therefore, ctb-

1 negatively affected complex I in its most active form, the I:III:IV supercomplex.

The simplest explanation may be that this mutation changes the conformation of the supercomplex I:III:IV such that complex IV is less tightly bound to the supercomplex, reducing complex I activity toward enzymatic rates of its I:III form.

If so, the increased proportion of supercomplex I:III relative to I:III:IV in isp-1;ctb-

1 may be a reflection of this effect of ctb-1 in the double mutant, and the reason that the activity in the double mutant was approximately the same as in ctb-1.

I hypothesized that if complex III altered interactions between components of the I:III:IV supercomplex in this manner, I could detect an increased vulnerability of the supercomplex to disassociation. Both complex III mutants lost complex IV from supercomplexes (S1-S4) more readily than wild type. In addition, S4 was partially dissociated into smaller supercomplexes whose molecular weights were comparable to S3, S2 and S1 suggesting, as I originally inferred, that S4 was supercomplex I:III:IV4 containing 4 copies of complex IV.

Wild-type supercomplex resisted dissociation relative to either isp-1;ctb-1 and ctb-1, suggesting that these two mutations produce conformational changes that disrupt supercomplex integrity.

How can we link the finding that complex III mutations alter

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supercomplexes such that complex IV loosens its grip on supercomplex I:III:IV,

and decreases CI activity? According to a current 3D map of supercomplex

I:III:IV, complex III and IV saddle the membranous arm of complex I (Schafer et

al. 2006) where the proton-pumping machinery of complex I is thought to reside

(Brandt 1997; Efremov et al. 2010). NADH-Q reductase in the matrix arm provides the driving force for proton pumping via conformational coupling

(Friedrich 2001; Brandt 2006; Efremov et al. 2010). Mutations in the homologues of ND subunits in the membranous arm of complex I, which form a core of the proton-pumping machinery, result in decreased CI activity but normal NFR activity (Kao et al. 2004; Kao et al. 2005a; Kao et al. 2005b; Torres-Bacete et al.

2007). The relaxed interaction between complex IV and other components of the supercomplexes in the mutants may lead to an imperfect conformation and dysfunction of the proton-pumping module, which then impedes NADH-Q reductase in the matrix arm. A similar logic may explain how complex I in wild- type supercomplex I:III is not as active as complex I in wild-type supercomplex

I:III:IV.

As expected, complex III defects affected both complex I and complex II- dependent respiration; complex IV respiration was increased in all mutants, most dramatically in isp-1 and isp-1;ctb-1 even though the amount of complex IV appeared unchanged in these mutants. Cytochrome c oxidase controls mitochondrial energy metabolism (Kadenbach et al. 1997; Napiwotzki &

Kadenbach 1998) by acting as a sensor of intramitochondrial ATP/ADP (Arnold et al. 1998). A low ATP/ADP ratio, as is expected in the complex III mutants, may

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increase complex IV driven respiration. However, COX activity in ETC assays was not significantly increased in complex III mutants compared to wild type.

Since the mitochondrial membranes are solubilized in ETC assays, ATP/ADP differences between strains are eliminated. However, this argument loses

traction since the complex IV respiration was measured in the presence of an

uncoupler DNP (2,4-dinitrophenol). The uncoupler can dissipate the proton

gradient across the membrane and allows complex V to hydrolyze ATP to ADP.

Difference in ATP/ADP ratio between wild type and the complex III mutants may no longer exist. Nonetheless, our unpublished data for the complex II mutant

(mev-1) show that uncoupled TMPD/ascorbate rates following succinate stimulation are significantly higher than uncoupled TMPD/ascorbate rates after malate stimulation of respiration. Since mev-1 is a complex II mutant and has almost no measurable respiration with succinate alone, malate as a fuel should yield more ATP than succinate. Therefore, inhibition of uncoupled complex IV activity by ATP in malate stimulated respiration is more potent than in succinate stimulated respiration. How can ATP-inhibition of complex IV still happen in the presence of the uncoupler? A possible explanation is that DNP may not cause complex V to immediately hydrolyze all ATP. Maintenance of high ATP, which causes inhibition of complex IV is due to slow ATP hydrolysis. This theory is corroborated by a study from Ramzan et al. They demonstrated that in rat heart mitochondria, addition of exogenous ATP, which increased ATP/ADP ratio could inhibit complex IV even in the presence of the uncoupler (Ramzan et al. 2010). A recent study by Yang et al. (Yang & Hekimi 2010) demonstrated decreased

111

enzymatic activities of CI-III and CII-III in isp-1, although rates of CII-III were depressed more than CI-III. This is qualitatively different than our data. They did not measure complex III activity per se, nor complex I activity, and all rates were normalized to citrate synthase. It is difficult to compare techniques between our studies and those of Yang et al., so the differences between them are not fully interpretable.

I also identified three unique bands in the BNGs of isp-1 that may represent intermediates in complex I assembly or degradation. A current model of supercomplex assembly proposes that the subcomplex Iβ of complex I associates first with the partially assembled complex III and IV, leading to a module upon which the mature supercomplex forms (Lazarou et al. 2009). ISP-A represents a nearly complete sub-complex consisting of the P and Q modules already combined with ND4:ND5 intermediates and partially assembled complex

III and IV. This suggests that the mutation in the ISP halted supercomplex maturation at the final step prior to incorporation of the NADH-oxidase module (N module). In turn, two smaller intermediates accumulate, ISP-B (an 830 kDa intermediate P/Q complex, with partially assembled complex III and IV), and ISP-

C (likely a 400 kDa intermediate consisting of the Q module and an early form of

P, the membranous arm). Even though the bands we see correspond well to proposed assembly modules of complex I, I have not ruled out that they may represent steps in degradation of a supercomplex.

The physical association between complex I and III is thought to involve

ISP of complex III and the NDUFS2 and NDUFS4 subunits of the Q module of

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complex I. Missense mutations in those complex I subunit decreased levels of

complex III (Ugalde et al. 2004). Since the mutation in isp-1(qm150) alters an

amino acid in the highly mobile peripheral domain of the protein, it is difficult to

envision how this might alter a complex I-III interaction. However, such a change

could easily affect assembly or stability of the supercomplex resulting in

misaligned complexes. The appearance of abnormal subcomplexes in the isp-1

BNGs supports this model.

It is intriguing that ctb-1 improves complex I function in an isp-1 background without affecting complex III function. The effects of complex III subunits on complex I represent an allosteric mechanism wherein subunits of the catalytic core can regulate supercomplex assembly (isp-1), ratios of supercomplexes (isp-1;ctb-1) or complex I electron flow in the face of normal amounts and profiles of supercomplexes (ctb-1). This is the first report of a complex III mutation that can affect complex I in this manner.

This study provides a new insight into the role of supercomplexes in ETC defects. Complexes I and III have intricate structural and allosteric interactions revealing a complicated inter-relatedness between different complexes of the mitochondrial respirasome (Schagger & Pfeiffer 2000; Schagger et al. 2004).

Diagnosis of patients with mitochondrial disease, which is clearly problematic,

must take into account the myriad of consequences in the ETC that may result

from a single mutation. Clearly a defect in a single subunit of the ETC can have

wide ranging and unanticipated effects on mitochondrial function.

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3.4. Conclusion

Complexes III and IV can influence the function of complex I via their intricate association in the supercomplex. Complex III is necessary for complex I assembly since a mutation in complex III subunit such as isp-1 can prevent the

formation of complex I. The physical association of complex IV with the

membranous part of complex I in the supercomplex can also be weakened by

mutation in complex III subunit(s) as demonstrated in isp-1;ctb-1 and ctb-1.

Consequence from unstable supercomplexes includes a decreased complex I function. Here I provide evidence suggesting that complexes III and IV influence complex I function in an allosteric manner via supercomplex structure.

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3.5. Figures and tables

FIGURE 18

100 N2 isp-1 isp-1;ctb-1 75 ctb-1

50 Eggs/worm

0 2 4 6 8 10 12 14 Days

Figure 18. Number of eggs per worm per day in complex III mutants and wild type. Plates were examined every 24 hours. Day 0 is the day that parental

animals were hatched. Data are represented as mean ± SEM from three independent worm cultures (totaling 30 adult animals for each strain).

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FIGURE 19

120 120 600 N2 isp-1 isp-1;ctb-1 # * # ctb-1 90 # * 90 450 * * # * # 60 60 # 300 * * 30 30 150 nano atom (O)/min/mg prot nano atom (O)/min/mg prot nano atom (O)/min/mg prot

0 0 0 Malate Succinate TMPD/ascorbate

Figure 19. Integrated mitochondrial function of complex III mutants and wild type. Intact mitochondria were isolated from complex III mutants and wild type, then assayed for oxygen consumption rates. Malate, succinate and

TMPD/ascorbate were the substrates to stimulate complex I-, complex II- and complex IV-dependent respiration, respectively. Data are represented as mean ±

SEM from four independent experiments. * and # indicate statistical significances as p < 0.05 in comparison to wild type and isp-1, respectively.

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FIGURE 20

A B C N2 isp-1 400 800 1800 isp-1;ctb-1 # # ctb-1 600 300 1350 # # * # * 400 * 200 * 900

200 100 * 450 * *

nmole substrate/min/mg prot 0 nmole substrate/min/mg prot 0 nmole substrate/min/mg prot 0 CIII CI NFR

D E F G

1200 160 32 5000

900 120 24 3750 # 600 * 80 16 # 2500 * 300 * 40 8 1250 *

nmole substrate/min/mg prot 0 nmole substrate/min/mg prot 0 nmole substrate/min/mg prot 0 nmole substrate/min/mg prot 0 CI-III CII-III CII COX

Figure 20. Respiratory chain enzymatic activities of complex III mutants and wild type. A. Enzymatic activity of CIII. B. Enzymatic activity of CI. C.

Enzymatic activity of NFR. D. Enzymatic activity of CI–III. E. Enzymatic activity of

CII–III. F. Enzymatic activity of CII. G. Enzymatic activities of CIV. Data are represented as mean ± SEM from four to eight independent worm cultures. * and

# indicate statistical significance as p < 0.05 in comparison to wild type and isp-1, respectively.

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FIGURE 21

A B

N2 isp-1isp-1;ctb-1ctb-1 N2 isp-1isp-1;ctb-1ctb-1 N2 isp-1isp-1;ctb-1ctb-1 N2 isp-1isp-1;ctb-1ctb-1 I:III S4 S3 S2 I S1 S0 V2 V CI in-gel staining

V F1 domain of III2 ATPase F1 domain of Coomassie ATPase stain anti-Rieske anti-ATPase Va C 1.5 N2 isp-1 isp-1;ctb-1 1 ctb-1

* 0.5 * Fold changes relative to N2 0 CI CI CIII CIV Triton X-100 Digitonin native-Westerns Digitonin based-BNGs based-BNGs based-BNGs

Figure 21. Amounts of respiratory enzyme complexes. A. Triton X-100 based-BNGs demonstrate the fully assembled complex I in an isolated form (I) and in the supercomplex form (I:III). CI-IGA is performed in a duplicate gel. B.

Native Western blot probed with anti-Rieske mAb (left) or with anti-ATPase subunit Va mAb (right, loading control in duplicate gel). Abbreviations: S0, supercomplex I:III; S1, I:III:IV1; S2, I:III:IV2; S3, I:III:IV3; S4, I:III:IV4. C.

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Quantitative analysis of the amount of fully assembled complexes I, III and IV.

Complexes I and IV were measured as densitometry scans of their IGAs on

BNGs, complex III as densitometry scans of native Westerns. All values were

normalized to complex V (see Methods). Data are represented as mean ± SEM

from three to four independent experiments. * indicates statistical significance as p < 0.05 in comparison to wild type.

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FIGURE 22

N2 isp-1 isp-1;ctb-1 ctb-1

NUO-2

COXI

ATPase Va

cytochrome c

ANT

Figure 22. The stead-state level of respiratory enzyme subunits. SDS-

PAGE/Western blot analysis of NUO-2 subunit of complex I, COX I subunit of complex IV, ATPase subunit Va and cytochrome c; ANT was a loading control.

(Quantification of the bands is shown in Table 5).

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FIGURE 23

A B C

N2 isp-1 isp-1;ctb-1ctb-1 N2 isp-1 isp-1;ctb-1ctb-1 N2 isp-1 isp-1;ctb-1ctb-1 4 S4 S4 S 3 S3 S3 S 2 S2 S2 S S1 S1 S1 S0 S0 S0 V2 V2 V2

V V V

III2

2 IV2 IV

D E anti-NDUFS3

N2 isp-1 N2 isp-1 S4 S4 0.95 S3 S3 N2 S2 S2 S1 S1 isp-1 S0 S0 0.90 isp-1;ctb-1 V2 ISP-A ISP-A ctb-1 ISP-B ISP-B 0.85 *

V I:III:IV

I:III + I:III:IV + I:III 0.80 III2

0.75

IV2 ISP-C 0.70

Figure 23. Digitonin-based BNGs in complex III mutants. A. Coomassie stain.

B. CI-IGA and C. CIV-IGA. D. Left; the Coomassie stain of gel slices of N2 and isp-1 are compared side by side to reveal ISP-A and ISP-B, which appear only in isp-1. Right; native-Western blots performed using anti-NDUFS3 mAb in order to better visualize ISP-A, ISP-B and ISP-C. E. The amounts of complex I in

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supercomplexes I:III (S0) and I:III:IV (S1-S4) were measured from four independent digitonin-based CI-IGAs and presented as the ratio between them.

Error bars represent mean ± SEM from three independent experiments. * indicates statistical significance as p < 0.05 in comparison to wild type.

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FIGURE 24

0.15 I:III I:III:IV

0.1 *

CI/NFR * 0.05 *

0 N2 isp-1;ctb-1 ctb-1

Figure 24. Complex I activity in supercomplex I:III and I:III:IV. Electroeluted

I:III and I:III:IV supercomplexes were assayed for CI and NFR activities. Data are represented as mean ± SEM. from four independent experiments. * indicates statistical significance as p < 0.05.

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FIGURE 25

digitonin 2D-maltoside A N2 isp-1;ctb-1 ctb-1

S4 S4 S3 S3 S4 S2 S2 S3 S1 S2 S1 S0 S1 S0 S0 V2 2 V V2

V V V

B a b c d a b c d a b cd

6 N2 * isp-1;ctb-1 / V 4.5 ctb-1 2 *

dissociated IV 1.5

Figure 25. The ability of 2D-BN/hrCNE to dissociate wild-type and complex

III mutant supercomplexes. A. Coomassie stain of the 2D gel over the area of supercomplex dissociation. B. Silver stain of the duplicates of the gels in A, which are cut from the 420 kD region of the second dimension. This size is identical to that of IV2. C. The amount of complex IV2, measured by densitometry scanning of bands a-d in 5B, relative to the amount of Coomassie staining of monomeric complex V. Error bars represent mean ± SEM from three

124

independent experiments. * indicates statistical significance as p < 0.05 in comparison to wild type.

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FIGURE 26

A B C 1 2 3 4 1 2 3 4 1 2 3 4 2nd 1st 2nd 1st 2nd 1st Digitonin + + + + + + + + + 1st 2nd 1st 2nd 1st 2nd Sodium cholate + + + + + + + + + Coomassie + + + + + + + + + + + +

sample gel

I:III:IV I:III V2

III2

IV2

Coomassie stain CI in-gel staining CIV in-gel staining

Figure 26. The effect of sodium cholate on mitochondrial supercomplexes.

Wild-type mitochondria (250 ug protein each lane) were solubilized by digitonin alone (lane 1), sodium cholate alone (lane 2), cholate followed by digitonin (lane

3), and digitonin followed by cholate (lane 4). These samples were resolved in

3.5-11% non-denaturing polyacrylamide gel. Digitonin was titrated to a digitonin/protein mass ratio of 6/1. The amount of sodium cholate was the same as that in electron transport chain assays, which is 1% in the final concentration.

Gels in panels A, B and C were stained coommasie, complex I in-gel activity and complex IV in-gel activity, respectively. A stacking gel was included to reveal

126

material that did not enter the gel, which accumulated at the bottom of the wells

as seen in lanes 2 and 3. The bands were labeled according to their identification

from digitonin-based BNGs; I:III:I, supercomplexes I:II:IV; I:III, supercomplex I:III;

V2; dimeric complex V; V, monomeric complex V; III2; dimeric complex III, and

IV2; dimeric complex IV.

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TABLE 4

Strain L3 L4 Adult Total eggs/worm N2 D2.5 D3 D3.5 223.8±3.9 isp-1 D5.5 D6.5 D7.5 79.6±10.7 isp-1;ctb-1 D3.5 D4.5 D5 126.4±9.3 ctb-1 D2.5 D3 D3.5 209±9.2

Table 4. Development of complex III mutants and N2 at 20° C. Synchronized cultures of worms were studied for the rate of development to adulthood and subsequent fecundity. Three independent cultures totaling 30 animals for each strain were studied. Error is ± S.D.

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TABLE 5

N2 isp-1 isp-1;ctb-1 ctb-1 68.77±5.44% 100.78±1.94% 92.21±3.91% NUO-2 100% (0.01) (0.56) (0.08) 107.59±7% 116.38±23.84% 103.13±9.58% Cytochrome c 100% (0.2) (0.35) (0.63) 83.32±27.1% 96.44±28.12% 109.94±34.49% COX I 100% (0.4) (0.85) (0.73)

Table 5. Relative expression of NUO-2, cytochrome c and COX I in complex

III mutants when ANT is used as a loading control. N2 = 100% expression

(n= 3, p-values in the parentheses).

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TABLE 6

C. elegans Human isp-1’s unique bands ISP-A ISP-B Y56A3A.19 NDUFB1 + C18E9.4 NDUFB3 + W01A8.4 NDUFB4 + + ZK809.3 NDUFB6 + + D2030.4 NDUFB7 + + Y51H1A.3a NDUFB8 + + C16A3.5 NDUFB9 + + F59C6.5 NDUFB10 + + Y71H2AM.4 NDUFC2 + + Y54F10AM.5 NDUFA8 + Y54E10BL.5 NDUFS5 + MTCE.25 ND4 + MTCE.35 ND5 + F42G8.12 UQCRFS1 + + MTCE.21 MT-CYB + C54G4.8 CYC1 + + F56D2.1 UQCRC1 + + T10B10.2 UQCRC2 + + VW06B3R.1a UQCRC2 + T02H6.11 UQCRB + + F45H10.2 UQCRQ + + MTCE.31 MT-CO2 + + W09C5.8 COX4l1 + + F26E4.6 COX7C + + cco-2 COX5A + Y71H2AM.5 COX6B + +

Table 6. Proteomic analysis of ISP-A and ISP-B. Only MRC encoding genes are listed. C. elegans and human gene names encoding the identified proteins are obtained from the C. elegans Genetics Center and HUGO Gene

Nomenclature Committee respectively. Absence of a subunit does not preclude its presence in the protein band as mass spectrometry may miss individual proteins. Turquoise shading indicates complex I subunits, yellow complex III, and peach complex IV.

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134

Chapter 4

Discussion and Future Direction

135 4.1. Research summary

I used C. elegans as a model for the study of respiratory supercomplexes.

In the first part of my work, RNA interference was used to inactivate two genes encoding complex IV subunits to create complex IV-deficient animals. These animals exhibited abnormal phenotypes such as delayed development and decreased fecundity, and exhibited decreased complex IV-dependent respiration.

Interestingly, complex IV knockdown not only resulted in a decreased complex IV activity but also led to decreased complex I activity despite a normal amount of complex I. The study of the supercomplex profile in complex IV-deficient animals revealed that these animals had a decreased amount of I:III:IV supercomplexes and an increased amount of the I:III supercomplex. According to a previous study by Schaffer et al. (Schafer et al. 2006) performed in bovine heart mitochondria, complex I was more enzymatically active when in I:III:IV supercomplexes than when in I:III supercomplex. I concluded that the decreased complex I activity in complex IV-deficient C. elegans was caused by the decreased amount of I:III:IV supercomplex, a secondary outcome of complex IV knockdown. Therefore, complex IV has an allosteric effect on complex I function.

In the second part of my thesis, an extension of the first, I took advantage of three different complex III mutants in C. elegans to understand how defective complex III affects complex I function and supercomplex formation. isp-1, a missense mutation in the Rieske subunit of complex III, causes a dramatic decrease in complex III activity without decreasing the amount of complex III.

Interestingly, isp-1 exhibited a significant decrease in the amount of complex I by

136 disrupting supercomplex formation/stability, resulting in an incompletely

processed complex I. ctb-1, a missense mutation in the cytochrome b subunit of complex III, is a suppressor of the whole-animal phenotype of isp-1. The isp-

1;ctb-1 double mutant grows much more quickly than isp-1. This second mutation, however, did not improve complex III function. The only measurable improvement in respiratory complex activity in the double mutant, compared to isp-1, was improved complex I activity. However, overall complex I activity was still less than wild type, despite normal amounts of complex I. ctb-1 as a single mutation had both decreased complex III activity and decreased complex I activity, although not as severe as seen in isp-1. Like the double mutant, the amount of complex I was normal in ctb-1. Electron transport chain (ETC) activity was tested in supercomplexes I:III and I:III:IV isolated from native gels.

Surprisingly, complex I activity within the supercomplex I:III:IV is significantly reduced in ctb-1 and the double mutant compared to the wild type. Two- dimensional native gel electrophoretic profiles of the mutant supercomplexes suggest vulnerability for dissociation of complex IV from the I:III:IV supercomplex in the complex III mutant animals. I concluded that both complexes III and IV play a role in supercomplex formation, and the formation and function of complex I.

4.2. Discussion and future direction

4.2.1. Supercomplex assembly requires proper quality and quantity of each

complex component

137 Availability of complex components that make up the supercomplex is a key factor for successful formation of supercomplexes. Lack of complex IV caused decreased formation of supercomplexes I:III:IV together with an

increased amount of supercomplex I:III (chapter 2), suggesting that

supercomplex I:III is a building block for supercomplexes I:III:IV. A similar

observation in a study of Acin-Perez et al. (Acin-Perez et al. 2008) supports this model. Acin-Perez et al. performed pulse-chase experiments by labeling the mtDNA-encoded proteins and analyzed the time course of their incorporation into respiratory complexes and supercomplexes. The labeling of large supercomplexes appeared to have a different pattern depending on whether they contained complex IV. Supercomplexes I:III:V appeared as soon as after 0.5 hour of chase, whereas supercomplexes I:III:IV were not assembled until 24 hours after a chase phase began, despite the presence of fully assembled complex IV from the onset of the chase phase (0.5-1 hours).

Availability of normal complex subunits is another important factor that determines supercomplex formation. Complex III that contains a mutant subunit can interrupt supercomplex formation; the P225S mutation in the Rieske subunit of complex III halted supercomplex maturation. However, such a negative effect of mutant complex III on supercomplex formation appears to be limited to some mutations. The A170V mutation of cytochrome b subunit of complex III did not affect supercomplex formation/stability at all.

One challenging question regarding supercomplex structure is how components of the supercomplexes are held together. All of the structural

138 visualization of supercomplexes available up to date was obtained from electron microscopic studies, which do not provide enough resolution to localize individual subunits. To advance our understanding of intercomplex interactions in supercomplexes, an X-ray crystallographic study of supercomplexes may be a worthwhile investigation. Unfortunately, supercomplex purification and crystallization, which are the necessary procedures prior to X-ray crystallography, has never been achieved. However, an alternate way to solve the question may be to observe to effects of mutations of the complex components of supercomplexes.

We might anticipate that any significant amino acid substitution in the components of supercomplexes, such as a critical proline substitution, or an alteration of a highly conserved residue, would disrupt supercomplex structure and/or assembly, like the P225S mutation in isp-1. On the other hand, we might also expect that a mutation in a peripheral subunit of the complex would affect supercomplex structure/formation less than mutation in an interface between complexes. However, this is not necessarily true. For example, the E373K substitution in the cytochrome b subunit of complex III was shown to cause decreased levels of both complexes I and III in mouse (Acin-Perez et al. 2004).

This mutation is located at the carboxy end of cytochrome b, which is located in the mitochondrial matrix (Xia et al. 1997; Zhang et al. 1998). It is a very peripheral residue in the peptide and is also most likely not at the complex I- complex III interface. Nonetheless, it can affect the structural interdependence between complexes I and III. Therefore, to predict an outcome of a mutation on

139 supercomplex structure/formation by only considering the position and

conservation of a mutation might not be accurate.

Another approach to identify complex-complex interactions in

supercomplexes is to use the yeast two-hybrid (Y2H) screening assay (Fields

& Song 1989). This technique is based on the modular structure of transcription factors where close proximity of the DNA-binding domain to the activation domain induces transcription of a reporter gene, reflecting a putative protein- protein interaction (Fig. 26). Putative interactions from the Y2H assay have to be confirmed or eliminated by further biological experimentations such as a knockout experiment, which is discussed below. To use this assay to study, for example, complex I-complex III interactions in the supercomplexes, potential candidates for the assay should be the subunits of complexes I and III that are likely to be localized at the complex I-III interface. Data from X-ray crystallographic studies of individual complexes and from electron microscopic studies of the supercomplexes should provide this information. The Y2H assay

has been successfully used to map a large-scale interactome network in C.

elegans by Li et al. (Li et al. 2004)

RNAi may be another useful method to identify complex-complex interactions in the supercomplexes in C. elegans. RNAi knockdown just creates hypomorphic phenotype as opposed to the missense mutations which produce defective protein. However, suppression of a gene of interest by RNAi has to fulfill two criteria. First, a subunit of interest has to be largely depleted (knocked out), which may be achieved by constitutively expressing RNAi targeting the

140 gene. Second, a knockout can still assemble such complex. For example, a

knockout of the Rieske subunit should be able to assemble the Rieske-depleted

complex III. This approach may be useful to test whether the disruption in

supercomplex formation in isp-1 is a result of that particular mutation (P225S) or

because the Rieske subunit is necessary for supercomplex formation since it is a subunit interface. If the disruption in supercomplex formation in isp-1 is a result of

that particular mutation (PS225), e.g. a consequence of the mutation through

other complex III subunits, the Rieske-depleted complex III in the knockout

should be able to maintain supercomplex formation. Alternatively, if the Rieske

subunit is a subunit interface and is necessary for supercomplex formation, I

would expect to see that the Rieske-depleted complex III in the knockout should

not be able to maintain supercomplex formation. Partial suppression of an

OXPHOS subunit by RNAi knockdown may not be useful for this purpose

because residual expression of a subunit of interest may allow full assemble of

the complex regardless.

4.2.2. Supercomplex I:III versus I:III:IV

In many animals the amount of supercomplexes I:III:IVn is much higher

than that of supercomplex I:III such as (Schagger & Pfeiffer 2000; D'Aurelio et al.

2006; Schafer et al. 2006; Rosca et al. 2008; Suthammarak et al. 2009).

Supercomplexes I:III:IVn should provide more electron flux through the

respiratory chain than supercomplex I:III does and, thus, supercomplex I:III:IV is

likely to be a major contributor of mitochondrial membrane potential and ATP

pool. How is supercomplex I:III:IV such an important player in the respiratory

141 chain? Is it possible that supercomplex I:III and free IV2 together can substitute for the function of supercomplex I:III:IV? A short answer is that supercomplex

I:III:IV may not be functionally replaceable by supercomplex I:III and free IV2. A few reasons to support my statement follow.

• Supercomplex I:III:IV as a major contributor of mitochondrial energetics.

Supercomplex I:III:IV executes the NADH-Q reductase activity 3 times

faster than complex I in supercomplex I:III (Schafer et al. 2006; Suthammarak et

al. 2009) and presents in mitochondria 9 times more abundant than supercomplex I:III (Suthammarak et al. 2009). Taking these two factors into account, supercomplex I:III:IV may contribute mitochondrial membrane potential and ATP production up to 27 times of the extent that supercomplex I:III does. But how is supercomplex I:III:IV so active? Two possible mechanisms follow.

First, the close proximity of complex IV in the supercomplex may sustain complexes I and III in the reduced state almost all the time, which would ensure rapid oxidation of NADH and steady electron flow through complex I, thus, increasing complex I activity. Second, complex IV enhances complex I function by allosteric exertion through supercomplex structure. Attachment of complexes

III and IV to the membranous arm of complex I in the supercomplex results in the bending of the matrix arm toward to the membranous arm of complex I in supercomplex I:III:IV (Schafer et al. 2007) (Fig. 27). This structural change in the matrix arm of the supercomplex may augment the rate of electron flow from

NADH to ubiquinone, which takes place mainly in the matrix arm of complex I, and increases complex I activity, hence, increasing overall activity of

142 supercomplex I:III:IV. Unfortunately, however, there is no direct comparison between the shape of the matrix arms from supercomplex I:III and I:III:IV from the

same species. Structural studies of supercomplex I:III and supercomplex

I:III:IV from the same species may suggest whether the conformational change of the matrix arm of complex I really plays a role in the difference in complex activity in these two supercomplexes or not.

• Supercomplex I:III:IV, not complexes I, IIII and IV, matters.

Theoretically, complexes I, III and IV should be able to perform similar

electron transfer activity to that of supercomplex I:III:IV. However, according to

the study from Acin-Perez et al. (Acin-Perez et al. 2008), the simultaneous

inclusion of the individual complexes I, III and IV to mimic supercomplex I:III:IV

did not exhibit an integrated respiration when NADH was provided. They

concluded that not only are the respiratory complexes and electron donors

required for mitochondrial respiration, but also the proper arrangement of the

respiratory complexes into a supercomplex is necessary for respiration.

However, I found experimental flaws that might have prevented complexes I, III

and IV from respiring. O2 consumption determinations were done from gel slides

containing complexes I, III and IV that were cut into small pieces. Those complexes might have not been exposed to each other at all since they were still in the gels. Furthermore, coenzyme Q and cytochrome c were not supplemented in the assay. In order to improve the reliability of this experiment, I suggest that

O2 consumption measurement has to be done from electroeluted

143 complexes with supplementation of coenzyme Q and cytochrome c to allow

the interaction between them and ensure complete electron carriers.

What is the driving force of supercomplex I:III:IV formation? Supercomplex

I:III is a building block of supercomplex I:III:IV. Added to the fact that

supercomplex I:III:IV is much more active than supercomplex I:III, the formation of supercomplex I:III:IV is likely to be driven by high cellular energy requirement.

Under normal physiological conditions, supercomplex I:III may be functionally idling yet exists as a reserve capacity of the respiratory chain. When the cellular energy expenditure surges, supercomplex I:III shifts to supercomplex I:III:IV by addition of complex IV to keep up with the cellular energy requirement. To prove that energy requirement is the driving force of supercomplex formation, a comparison between the amounts of supercomplexes I:III and I:III:IV in different tissue types that have different energy requirements may be worth investigating such as supercomplexes in brain versus fat tissue or exercised muscle versus paralyzed muscle. In addition, during larval development of C. elegans where the energy requirement seems to be steeply increasing during L3 to L4 and L4 to adult, a comparison of the amounts of supercomplexes I:III and I:III:IV at each larval stage may be another worthwhile experiment to perform. Along with such study, other parameters that can reflect the energetic status of the cells, e.g. ATP levels, redox states of NADH and FAD+, should also

be monitored.

144 4.2.3. Stoichiometric association of complexes I, III and IV in supercomplexes

The stoichiometries within supercomplexes were first investigated in

bovine heart mitochondria by Schagger et al. (Schagger & Pfeiffer 2000). 1D BN-

PAGE was used to isolate supercomplexes from mitochondrial membrane then

2D BN-PAGE dissociated supercomplexes from 1D into the individual

complexes. A subsequent 3D SDS-PAGE was used to dissolve polypeptides in

each individual complex obtained from 2D to determine the molar ratios between

complexes I, III and IV in supercomplexes. They finally characterized I1III2IVx

complexes, each containing monomeric complex I, dimeric complex III and 0-3

copies of complex IV. After that, several studies have documented respiratory

supercomplexes I1III2IV3 and I1III2IV4 in other organisms (Eubel et al. 2003; van

Lis et al. 2003; D'Aurelio et al. 2006; Duarte & Videira 2009) in addition to those

that have been shown by Schagger et al. (Schagger & Pfeiffer 2000).

The presence of complex IV in supercomplexes obviously benefits the electron transport chain function since complex IV affects complex I function through supercomplex structure (Schafer et al. 2006; Suthammarak et al. 2009).

However, what could be (an) advantage(s) of different stoichiometries of complex

IV in supercomplexes? According a study from Schagger et al. (Schagger &

Pfeiffer 2001), the molar ratio of OXPHOS components in bovine heart

mitochondria (whole OXPHOS system, not supercomplex) is 1 : 1.3 : 3 : 6.7 for

complex I : II : III : IV. The high molarities of the terminal OXPHOS components

ensure the reduced state of the respiratory chain, which makes the respiratory

chain become more efficient and produce fewer ROS (Skulachev 1996; Lenaz &

145 Genova 2010). Therefore, if this rationale were applied to the stoichiometry of supercomplexes, I would expect that the bigger supercomplex I:III:IV is, the more efficient electron transfer is which produces less ROS. To prove that, the integrated function, i.e. O2 consumption in the presence of NADH, as well as rates of ROS production, has to be measured in each of I1III2IVx that is

eluted from BNGs.

4.2.4. Roles of supercomplex in mitochondrial OXPHOS disorders and aging

• Combined respiratory complex deficiency

A mutation in complex III or IV not only affects catalytic activity of the resident complex, but can also compromise the stabilizing effect of complex III or

IV to complex I, and lead to combined complex deficiency. For example, Blake et al. showed that the K319P mutation in human cytochrome b decreases complex

III activity and secondarily reduces the amount of complex I, resulting in combined complex I/III deficiency (Blakely et al. 2005). A combined complex I/IV deficiency in mice can be caused by COX10 knockout (COX10 is an assembly factor of complex IV) (Diaz et al. 2006). In this case, Diaz et al. demonstrated that the absence of complex IV affects complex I assembly and/or stability. The interdependence of supercomplex components in such manner clearly explains many cases of combined complex deficiency in both humans and mice (Andreu et al. 1999; Lamantea et al. 2002; Bruno et al. 2003; Acin-Perez et al. 2004; Diaz et al. 2006; Li et al. 2007).

Combined complex deficiency can also be caused by two other mechanisms. First is the redistribution of supercomplexes I:III and I:III:IV which

146 leads to combined complex I/IV deficiency in complex IV-deficient C. elegans

(Suthammarak et al. 2009). The second mechanism is the weakened physical

association of complex IV in supercomplex I:III:IV caused by complex III

mutations, leading to combined complex I/III deficiency without destabilization of complex I (Chapter 3).

However, there are several more cases of combined complex deficiency that are diagnosed without a known mechanism. I would like to suggest that

BNGs and in-gel activity staining of the complexes should be performed in addition to the established investigation panel to diagnose a cause of a combined complex deficiency in a patient. BNGs and in-gel activity staining are easy procedures, relatively inexpensive, yet provide accurate quantification of the amounts of fully assembled complexes. The amounts of fully assembled complexes will suggest whether combined complex deficiency is caused by the decreased amounts of respiratory complexes or by redistribution of supercomplexes. This information will also be useful to determine whether further investigations such as molecular testing for mutation will be worthwhile or not.

• Wide varieties of affected tissues in OXPHOS disorders

Abnormalities in the mitochondrial OXPHOS system affect organs that have high-energy demand such as skeletal muscle, nervous system and endocrine gland, etc. However, a mitochondrial OXPHOS disorder does not always affect every organ with high-energy requirement altogether. Distelmaier et al. (Distelmaier et al. 2009) have performed a comprehensive analysis of clinical, biochemical and cell physiological information of 15 children harboring known

147 mutations that cause complex I deficiency. While they found that some mutations

share certain clinical features such as mutations in NDUFS4 and NDUFS8 that tend to manifest a cardiomyopathy, obvious mutation-related phenotypes were

not obtained from their study. However, among the parameters that they

investigated, supercomplex formation was overlooked. In my point of view,

different tissue types may express different kinds/patterns of supercomplexes

due to difference in energy requirements. Complex I deficiencies may lead to

wide varieties of clinical presentations by affecting supercomplex formation in different manners. Each tissue type may rely on different types of supercomplexes, thus, be differentially susceptible to the same mutation. A study in supercomplex expression in different tissue types in normal physiological and disease condition may be another worthwhile investigation for a better understanding of genotype-phenotype correlations.

• Mitochondrial supercomplexes and aging

A decline in mitochondrial supercomplexes has been implicated as one of the principal underlying factors of aging. Gomez et al. (Gomez et al. 2009) have demonstrated that a decline in the amounts of supercomplexes I1III2IVn in rat heart mitochondria was associated with aging ,while the amounts of individual respiratory complexes remained unchanged with age. In addition, supercomplexes I1III2IVn displaying the highest molecular masses (high n) were

the most severely affected. They suggested that the decline in supercomplexes

I1III2IVn decreases oxidative capacity of the respiratory chain and increases ROS

production. These consequences are thought to underlie manifestation of aging

148 at some degree. However, their speculations would be more solid if mitochondrial

function and ROS production were assayed from mitochondria of the aging rat heart. The precise mechanism involved in the age-associated loss of supercomplexes has not been investigated in this study. Gomez et al. noted that cardiolipin is essential to anchor the respiratory supercomplexes to the mitochondrial membrane (Zhang et al. 2002; McKenzie et al. 2006). Since addition of exogenous cardiolipin has been demonstrated to reverse the age- linked decrease in the function of rat heart mitochondria especially complex IV function (Paradies et al. 1997), its loss may be responsible for a destabilization of supercomplexes in aging rat heart. However, it is still controversial whether cardiolipin plays a role here or not. Aged-associated decrements in cardiolipin were not observed in the aging rat heart from in a study from Moghaddas et al.

(Moghaddas et al. 2002). Finally, cytochrome c content should be considered since depletion of cytochrome c has been shown to destabilize supercomplexes in mouse fibroblasts as shown by Vempati et al. (Vempati et al. 2009). The decline in supercomplex in the aging rat heart might be due to a decrease in cytochrome c expression with age.

A potential treatment of OXPHOS disorders and rejuvenation therapy might be to prevent supercomplex destabilization and promote supercomplex formation.

149 4.3. Conclusion

Mitochondrial supercomplexes, containing complexes I, III and IV of the electron transport chain, are now regarded as an established entity.

Supercomplex I:III:IV has been theorized to improve respiratory chain function by allowing quinone channeling between complexes I and III. My work documents that the role of the supercomplexes extends beyond channeling. Physical association of complexes III and IV augments complex I function through supercomplex structure in an allosteric manner. Further investigation of the function of supercomplexes with different stoichiometries as well as the driving force of supercomplex formation will provide a better understanding of the mitochondrial respiratory chain and pathophysiology of mitochondrial diseases.

150 4.4. Figures

FIGURE 27

Figure 27. The yeast two-hybrid assay for detecting protein-protein interactions. The target protein fused to a DNA-binding domain that can bind to

the regulatory region of a reporter gene (bait). If the target protein binds to its

binding partner that express as a fusion protein to a transcription domain (prey),

their association will bring together two halves of a transcriptional activator,

resulting in expression of the reporter gene. The reporter gene is often one that permits growth on a selective medium. Bait and prey fusion proteins are generated by standard recombinant DNA techniques. Figure adapted from Bruce

151 Alberts, Alexander Johnson, Julian Lewis, Martin Raff, Keith Roberts, and Peter

Walter. Molecular Biology of the Cell. 4th ed. New York; Garland Science; 2002.

152 FIGURE 28

Figure 28. Comparison of the 3D map of the bovine heart supercomplex

I1III2IV1 with those of the individual complexes. The left column shows a surface representation in blue of the supercomplex I1III2IV1. The middle column

displays the semitransparent supercomplex and the fitted structures of the

complexes I (yellow), III (red) and IV (green). The right column illustrates

complex I, III and IV as they would assemble to form the supercomplex. The two

upper rows display side views along the membrane plane. The third row shows

the particles as seen from the matrix space and the lower row as seen from the

intermembrane space. The location of the membrane in a side view is displayed

153 in purple. IMS; intermembrane space. Figure adapted from Schafer et al.

(Schafer et al. 2007).

154 4.5. References

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155 Lenaz G, Genova ML (2010). Structure and organization of mitochondrial respiratory complexes: a new understanding of an old subject. Antioxid Redox Signal. 12, 961-1008. Li S, Armstrong CM, Bertin N, Ge H, Milstein S, Boxem M, Vidalain PO, Han JD, Chesneau A, Hao T, Goldberg DS, Li N, Martinez M, Rual JF, Lamesch P, Xu L, Tewari M, Wong SL, Zhang LV, Berriz GF, Jacotot L, Vaglio P, Reboul J, Hirozane-Kishikawa T, Li Q, Gabel HW, Elewa A, Baumgartner B, Rose DJ, Yu H, Bosak S, Sequerra R, Fraser A, Mango SE, Saxton WM, Strome S, Van Den Heuvel S, Piano F, Vandenhaute J, Sardet C, Gerstein M, Doucette-Stamm L, Gunsalus KC, Harper JW, Cusick ME, Roth FP, Hill DE, Vidal M (2004). A map of the interactome network of the metazoan C. elegans. Science. 303, 540-543. Li Y, D'Aurelio M, Deng JH, Park JS, Manfredi G, Hu P, Lu J, Bai Y (2007). An assembled complex IV maintains the stability and activity of complex I in mammalian mitochondria. J Biol Chem. 282, 17557-17562. McKenzie M, Lazarou M, Thorburn DR, Ryan MT (2006). Mitochondrial respiratory chain supercomplexes are destabilized in Barth Syndrome patients. J Mol Biol. 361, 462-469. Moghaddas S, Stoll MS, Minkler PE, Salomon RG, Hoppel CL, Lesnefsky EJ (2002). Preservation of cardiolipin content during aging in rat heart interfibrillar mitochondria. J Gerontol A Biol Sci Med Sci. 57, B22-28. Paradies G, Ruggiero FM, Petrosillo G, Quagliariello E (1997). Age-dependent decline in the cytochrome c oxidase activity in rat heart mitochondria: role of cardiolipin. FEBS Lett. 406, 136-138. Rosca MG, Vazquez EJ, Kerner J, Parland W, Chandler MP, Stanley W, Sabbah HN, Hoppel CL (2008). Cardiac mitochondria in heart failure: decrease in respirasomes and oxidative phosphorylation. Cardiovasc Res. 80, 30-39. Schafer E, Dencher NA, Vonck J, Parcej DN (2007). Three-dimensional structure of the respiratory chain supercomplex I1III2IV1 from bovine heart mitochondria. Biochemistry. 46, 12579-12585. Schafer E, Seelert H, Reifschneider NH, Krause F, Dencher NA, Vonck J (2006). Architecture of active mammalian respiratory chain supercomplexes. J Biol Chem. 281, 15370-15375. Schagger H, Pfeiffer K (2000). Supercomplexes in the respiratory chains of yeast and mammalian mitochondria. EMBO J. 19, 1777-1783. Schagger H, Pfeiffer K (2001). The ratio of oxidative phosphorylation complexes I-V in bovine heart mitochondria and the composition of respiratory chain supercomplexes. J Biol Chem. 276, 37861-37867. Skulachev VP (1996). Role of uncoupled and non-coupled oxidations in maintenance of safely low levels of oxygen and its one-electron reductants. Q Rev Biophys. 29, 169-202. Suthammarak W, Yang YY, Morgan PG, Sedensky MM (2009). Complex I function is defective in complex IV-deficient Caenorhabditis elegans. J Biol Chem. 284, 6425-6435. van Lis R, Atteia A, Mendoza-Hernandez G, Gonzalez-Halphen D (2003). Identification of novel mitochondrial protein components of

156 Chlamydomonas reinhardtii. A proteomic approach. Plant Physiol. 132, 318-330. Vempati UD, Han X, Moraes CT (2009). Lack of cytochrome c in mouse fibroblasts disrupts assembly/stability of respiratory complexes I and IV. J Biol Chem. 284, 4383-4391. Xia D, Yu CA, Kim H, Xia JZ, Kachurin AM, Zhang L, Yu L, Deisenhofer J (1997). Crystal structure of the cytochrome bc1 complex from bovine heart mitochondria. Science. 277, 60-66. Zhang M, Mileykovskaya E, Dowhan W (2002). Gluing the respiratory chain together. Cardiolipin is required for supercomplex formation in the inner mitochondrial membrane. J Biol Chem. 277, 43553-43556. Zhang Z, Huang L, Shulmeister VM, Chi YI, Kim KK, Hung LW, Crofts AR, Berry EA, Kim SH (1998). Electron transfer by domain movement in cytochrome bc1. Nature. 392, 677-684.

157

Chapter 5

Material and Methods

158 5.1. Worm cultures

5.1.1. Animal and normal growing condition

Wild type C. elegans (N2), isp-1(qm150), isp-1(qm150);ctb-1(qm189) were

obtained from the Caenorhabditis Genetics Center (Minneapolis, MN, USA).

Worms were grown on nematode growth media (NGM) agar with a lawn of E. coli

OP50 as food source and maintained at 20°C. To propagate cultures in a large quantity for subsequent mitochondrial preparation, adult worms were transferred from NGM plates to liquid culture where E. coli OP50 was food source.

5.1.2. RNAi knockdown

The control worms were fed HT115 on NGM plates (3 days) and

transferred to liquid culture, (3 additional days) when most animals were young

adults. The knockdown worms were grown on either W09C5.8 or Y37D8A.14,

the 2 clones of E. coli containing the plasmids which express RNAi

corresponding to the target genes, COX IV and COX Va, respectively. They

required 4 days and 6 days on the plates and in liquid culture, respectively. In

this manner two generations of worms were exposed to RNAi or control bacteria.

5 mM isopropyl β -D-1-thiogalactopyranoside (IPTG) was used to induce RNAi

synthesis from the bacteria. Temperature was controlled at 20°C throughout.

Gene knockdown was checked by qPCR. RNA isolation and quantitative RT-

PCR were done by standard techniques (see below). HT115 bacteria containing

plasmids used for RNAi knockdown were obtained from GeneService

(Cambridge, UK).

159 5.2. Phenotypic study

5.2.1. Lifespan study

Adult nematodes were allowed to lay eggs for 4-6 hours on plates containing lawns of the control or RNAi producing bacteria. The adults were removed and the eggs were allowed to hatch. The day of egg laying for this F1 generation was defined as day 0 of life. Animals were moved to new plates on day 3 of life and plated at a density of 20-22 animals per 35 mm plate. The animals were then moved every two days. Death, defined as failure to respond to a light touch, was scored each day. Each experiment consisted of 100-180 animals and was repeated in triplicate. The total number of animals for the three experiments was recorded.

5.2.2. Embryonic developmental study

Adult nematodes were allowed to lay eggs for 2-4 hours on lawns of E. coli OP50. The adults were removed and the eggs were allowed to hatch. The day of hatching for this F1 generation was defined as day 0 of life. The first day of adulthood was recorded by the appearance of F2 eggs on the plate.

5.2.3. Egg laying assay

Immediately after the nematodes become adults, and before any eggs were seen on the plates, 25-30 adult nematodes were transferred to new NGM plates and allowed to lay eggs. The worms were subsequently transferred to the new plates every 24 hours and eggs counted after the adults were removed.

160 5.2.4. Anesthetic sensitivity

Freshly washed young adult worms from liquid cultures of each strain were transferred to agar plates and exposed to varying concentrations of the volatile anesthetic, halothane to determine the EC50s (effective concentration at which 50% of the animals are immobilized). Dose response analysis is performed as previously described (Morgan et al. 1988).

5.3. RNA Extraction and Quantitative reverse transcription PCR

Total RNA was extracted using Trizol® reagent, following the protocol from

Invitrogen. All RNA samples were subject to DNase digestion by using Ambion

DNA-freeTM Kit, (Applied Biosystems) and stored at -80°C until used. The integrity and purity of RNA samples were checked by measuring an absorbance

ratio at 260/280 nm. The total amount of 200 µg of RNA from each sample was

used to synthesize cDNA using M-MuLV RNase H+ reverse transcriptase,

random hexamer and RT buffer from DyNAmoTM, SYBR® Green 2-step qRT-PCR

kit (Finnzymes). Subsequent qPCR was performed in 50 µl reactions following

the manufacturer protocol. Primer concentration and annealing temperature were

optimized to obtain the amplification efficiency for each primer pair in a range of

100±5%.

To calculate the relative expression of each target genes, i.e. W09C5.8,

Y37D8A.14 in the RNAi-treated worms, the CT (threshold cycle), which is the cycle number at which the fluorescent signal increases exponentially, was determined from the spectrograms. In both RNAi-treated worms and wild-type

161 worms, the CT of the tested gene was normalized to that of ACT-3 (reference gene) and was presented as ΔCT. The relative expression of the target gene in

knockdown worm compared to wild-type worms was determined by normalizing the ΔCT of knockdown worms to the ΔCT of wild type. This calculation method is a modification of Livek et al (Livak & Schmittgen 2001). Specific primers for each gene used in the qPCR step were designed to span the exons to minimize the chance of amplifying genomic DNA of the corresponding genes. In addition this minimizes primer-dimer formation, self-priming and secondary structure interference, maximizing efficiency of PCR amplification. Primer concentrations and annealing temperatures used in the qPCR step were optimized to ensure that amplification efficiencies of the test and ACT-3 gene were similar (±1% from each other). No fluorescent signal was detected in any negative control and the melting curves indicated specificity of the amplification. All procedures were performed by using DNase/RNase-free consumables.

5.4. Mitochondrial functional assays

5.4.1. Worm collection

Young adult worms in liquid culture were subject to cleaning from bacteria for mitochondrial isolation. The liquid culture was first spin down at 2250 rpm for

5 minutes. The supernatant was removed and the worm pellets were washed 3 times with S-basal. The pellet was resuspended in 32.5% sucrose (dissolved in S basal solution) followed by a centrifugation of 1500 rpm for 5 minutes to separate adult worms from bacteria, debris, young larvae and eggs that should be at the

162 bottom of the tube. A light brown disc of healthy adult worms should be floating

on the top. Adult worms were transferred to a clean tube and washed with S-

basal twice. Worm pellets were condensed in one 15 ml tube and weighted after

washing.

5.4.2. Mitochondrial isolation (the protocol was used in Dr. Hoppel’s laboratory for isolating mitochondria from rat liver and was modified by Dr. EB Kayser in our laboratory for worm mitochondrial isolation)

All preparations were done at 4°C. First the clean worms were suspended with 9 ml MSM-E (220 mM mannitol, 70 mM sucrose, 5 mM MOPS (3-(N- morpholino)propanesulfonic acid), 2 mM EDTA (ethylenediaminetetraacetic acid), pH 7.4) and the pellet was spun at 350g Sorvall SS-34 rotor for 10 minutes. The supernatant was removed and the above step was repeated. The clean worms were resuspended in MSM-E (~10 ml total), followed by 3 strokes in a Potter/Elvehjem (glass tube/teflon pestle) homogenizer at 400 rpm. Sigma

Proteinase VIII (Subtilisin) was added to the homogenate in a ratio of 5 mg

Subtilisin/ 1 g fresh worm weight, and gently stirred for 10 minutes on ice. After incubation with proteinase, the worm pellet was homogenated again with 10 slow strokes in a Potter/Elvehjem (glass tube/teflon pestle) homogenizer at 400 rpm.

The worm homogenate was diluted with equal volume of MSM-EB (MSM-E and

0.4% bovine serum albumin) and spun down debris at 350g for 10 minutes. After centrifugation, supernatant was transferred into a clean centrifuge tube and spun at 7000g for 10 minutes. The mitochondrial pellet was resuspended in MSM-E and washed at 7000g for 10 minutes. After centrifugation, the pellet was

163 resuspended in MSM and washed at 7000g for another 10 minutes. The final

mitochondrial pellet was resuspended in MSM. The protein concentration was

determined by the Lowry assay (Lowry et al. 1951).

5.4.3. Oxidative phosphorylation assay

The capacity of oxidative phosphorylation of the mitochondrial electron transport chain was determined by recording rate of disappearance of oxygen from an assay medium when specific electron donors were provided to intact mitochondria. At the beginning of each assay, the initial oxygen content in the assay medium was measured. Intact mitochondria were then added, a small amount of oxygen was consumed until all endogenous substrates were exhausted or all endogenous ADP was converted to ATP. At this point 25 pmole of ADP was added to ensure that all internal substrates were consumed. After a low constant oxygen consumption rate was obtained, 5 µmole of exogenous substrates and 50 pmole of ADP were added to stimulate mitochondria respiration. Mitochondria exhibited state 3 respiration, the maximum respiration in the presence of ADP and substrate. When ADP was exhausted, a lower but also constant respiration rate was measured and was defined as state 4 respiration. It represented the residual reduction of oxygen when no phosphorylation took place. The ratio (state 3 rate/state 4 rate) is termed the respiratory control ratio (RCR). RCR is a measure of the quality of mitochondrial preparation. A high RCR (> 3) reflects good coupling of respiration and phosphorylation, i.e. is indicative of intact mitochondria.

Complex I-, II-, and IV- dependent respirations were measured by using

164 complex specific substrate; malate, succinate and TMPD/ascorbate, respectively.

5.4.4. Electron transport chain assay (protocol was obtained from Dr. Hoppel’s

laboratory)

Enzyme activities of individual respiratory complexes were measured at

30°C, exclusively using cholate treated mitochondria (1 mg of mitochondrial

protein solubilized in 0.1 M potassium phosphate buffer containing 1% cholate

(w/v), pH 7.4 to a final volume of 1 ml). CI: NADH-decylubiquinone

oxidoreductase (rotenone-sensitive and -insensitive) (Estornell et al. 1993;

Rauchova et al. 1997), NFR: NADH-ferricyanide oxidoreductase (with

background correction) (Hatefi 1978), CI-III: NADH-cytochrome c oxidoreductase

(rotenone-sensitive and -insensitive) (Hoppel & Cooper 1969), CII-III: succinate-

cytochrome c oxidoreductase (antimycin-sensitive and -insensitive) (Sottocasa et al. 1967), CIII: Ubiquinol-cytochrome c oxidoreductase (antimycin-sensitive and – insensitive) (Krahenbuhl et al. 1994) and CIV: cytochrome c-oxygen oxidoreductase (KCN-sensitive and -insensitive) (Krahenbuhl et al. 1991) were determined by established spectrophotometric methods as mentioned. The activity of CIV was determined by first order kinetic and reported as AU/min/mg of protein. The activities of the other assays were determined by zero order kinetic and reported as nmole of substrate/min/mg protein. Agilent 8453 UV- visible Spectroscopy System and Agilent ChemStation for UV-visible

Spectroscopy (Agilent Technologies Inc, Santa Clara, CA, USA) were the hardware and the software used for ETC assay, respectively.

165 5.5. Mitochondrial supercomplex studies

5.5.1. One-dimensional blue native gel electrophoresis (1D BNG), in-gel activity

assay (IGA) and electroelution of supercomplexes.

150-250 µg of mitochondrial protein as determined by Lowry assay were subjected to solubilization by digitonin with a detergent/protein mass ratio of 6/1.

The solubilization was performed at room temperature for 10 minutes, followed by centrifugation at 15,000 rpm, 4°C for 20 minutes. After centrifugation, supernatants were collected and Coomassie blue G-250 was added to the supernatants to obtain a dye/detergent mass ratio of 8/1 before loading onto a

3.5-11% polyacrylamide gradient gel, 0.15 x 14 x 16 cm (Hoefer Inc, Holliston,

MA, USA). The gels were run for at 100 V until all samples entered the separating gel and then at 300 V for the remainder. When the dye reached one- third of the gel, the first cathode buffer was replaced by the second cathode buffer which contained a 10-fold dilution of Coomassie blue G-250. Complex I in-

gel activity (CI-IGA) was visualized by incubating the gel with 0.5 mM nitroblue

tetrazolium and 5 mM NADH in 50 mM Tris-HCl pH 7.4 at room temperature for

60 minutes. Complex IV in-gel activity (CIV-IGA) was visualized by incubating the

gel with 0.1 % (w/v) 3,3’ diaminobenzidine, 0.1% (w/v) cytochrome c and 24

units/ml catalase in 50 mM Tris-HCl pH 7.4 at 37 °C for 3-6 hours. Both protocols

are modifications of Grad et al (Grad & Lemire 2006). To visualize complex I as

an individual complex in BNGs, triton X with a detergent/protein mass ratio of 5/1

was used instead of digitonin.

166 Electroelution of supercomplexes from BNG was accomplished using

eletroelution tubes equipped with dialysis membranes with a molecular weight

cutoff of 3.5 kDa (Gerard Boitech, Oxford, OH) and an electrode buffer (Wittig et al. 2006). The bands from BNGs containing I:III and I:III:IV supercomplexes were excised from the gels and transferred to electroelution tubes that were then submerged in the electrophoresis apparatus containing electrode buffer.

Electroelution was performed at 300 V, 4 °C. Eluent was collected from the tubes after 3 hours and was kept on ice to use for immediate assay of enzymatic

activity.

5.5.2. Two-dimensional blue native/high resolution clear native gel

electrophoresis (2D BN/hrCNE)

Gel strips from 1D BNG were subjected to 2D BN/hrCNE following previously established protocol (Wittig et al. 2007) with my minor modifications.

In brief, a vertical gel strip excised from a 1D BNG was placed horizontally on the

3.5-11% native gradient polyacrylamide gel. The cathode buffer to run the second dimension contained 0.05% deoxycholate, and 0.02% dodecyl maltoside.

Electrophoresis was performed at 125 V, 4 °C for 15 hours. CI-IGA and CIV-IGA,

Coomassie blue staining and silver staining (Color Silver Stain Kit, Thermo

Fisher Scientific, Inc., Rockford, IL) were subsequently performed as previously described (Suthammarak et al. 2009).

5.5.3. SDS-Western blot

25 µg of mitochondrial protein were separated by 12% SDS-PAGE and transferred to PVDF membranes. Membranes were blocked with 5% nonfat milk

167 in phosphate-buffered saline/Tween 20 (0.05%) and then incubated with the

following monoclonal antibodies (Mitosciences, Eugene, OR): anti-complex I

subunit NDUFS3 (MS112,), anti-complex IV subunit I (MS 404), anti-cytochrome

c (MSA06), and anti-adenosine nucleotide transporter (ANT) (MSA02). I also

used anti-complex III subunits Rieske (MS305) and anti-Core 2 (both gifts from

Mitosciences). Secondary antibodies were from Santa Cruz Biotechnology.

Chemiluminescence substrate (SuperSignal West Pico, Thermo Fisher Scientific,

Inc., Rockford, IL) was used to develop the reactions.

5.5.4. Native-Western blot

Supercomplexes resolved by BNGs were transferred to polyvinylidene difluoride (PVDF) membranes following a previously published protocol (Wittig et al. 2006). Membranes were blocked with 5% nonfat milk in phosphate-buffered saline/Tween 20 (0.05%) and incubated with monoclonal antibodies as described above.

5.5.5. Quantification of Proteins/Protein Complexes

The amount of complex I in each genotype was quantified by densitometry scanning of complex I IGA staining of both digitonin-based and Triton X-100- based BNGs. These measurements were normalized to densitometry scans of

Coomassie staining of total complex V. Quantification of complex IV was done by densitometry scanning of complex IV IGA in digitonin-based BNGs, also normalized to Coomassie staining of total complex V. Complex III was quantified from native Westerns of digitonin-based BNGs probed with anti-Rieske mAb then normalized to duplicate native-Westerns probed for complex V. The amount of

168 fully assembled complex III was the sum of optical density scanning of complex

III in both supercomplexes and in dimeric (III2) forms. In all cases at least 3 gels

were scanned.

5.5.6. Mass spectrometry

Coomassie blue bands excised from BNGs were analyzed by LCMS

(liquid chromatography mass spectrometry), using an LTQ Velos MS coupled to an Eksigent 1D-plus nano-LC. In brief, 20 ul of each in-gel tryptic digest of the

BNG slices were injected onto a reverse phase column (Magic C18 200 A,

Michrom Bioresources, Auburn, CA) and washed for 10 minutes in stationary phase at flow rate of 3 uL/min (5% acetonitrile) prior to separation on a 10 cm reverse phase analytical column (HALO C18, Michrom Bioresources, Auburn

CA) at 300 nl/min over a 30 minute gradient of either 10-50% or 10-70% acetonitrile. MS spectra were collected in data-dependent mode, with each full scan followed by MS scans of the 10 most intense ions. MS Peak lists were generated using Xcalibur software (Thermo Fisher Scientific, Inc., Rockford, IL) and ReAdW to produce peak lists in MzXML format. Peak lists were searched against a C. elegans proteome database with SPIRE (Systematic Protein

Investigative Research Environment, http://protein.spire.org/) using the search

algorithm X!Tandem, fully tryptic enzyme specificity, and 2.5 Da mass accuracy.

A 1% global false discovery rate (FDR) was used as the basis for protein

identification. Protein identifications were made using the methods and tools

outlined elsewhere in (Hogan et al. 2005), where a reshuffled C. elegans

proteome database was used to estimate the global FDR.

169 5.6. Statistical analysis

Analysis of Variance (ANOVA) was used to analyze groups of data for significant differences. Unpaired Student-T test was employed to calculate statistical significance of specific pairs if a difference was noted with ANOVA.

Error bars in the figures represent standard error of the mean (SEM) for each experiment.

170 5.7. References

Estornell E, Fato R, Pallotti F , Lenaz G (1993). Assay conditions for the mitochondrial NADH:coenzyme Q oxidoreductase. FEBS Lett. 332, 127- 131. Grad LI , Lemire BD (2006). Riboflavin enhances the assembly of mitochondrial cytochrome c oxidase in C. elegans NADH-ubiquinone oxidoreductase mutants. Biochim Biophys Acta. 1757, 115-122. Hatefi Y (1978). Preparation and properties of NADH: ubiquinone oxidoreductase (complexI), EC 1.6.5.3. Methods Enzymol. 53, 11-14. Hogan JM, Higdon R, Kolker N , Kolker E (2005). Charge state estimation for tandem mass spectrometry proteomics. OMICS. 9, 233-250. Hoppel C , Cooper C (1969). An improved procedure for preparation of inner membrane vesicles from rat liver mitochondria by treatment with digitonin. Arch Biochem Biophys. 135, 173-183. Krahenbuhl S, Chang M, Brass EP , Hoppel CL (1991). Decreased activities of ubiquinol:ferricytochrome c oxidoreductase (complex III) and ferrocytochrome c:oxygen oxidoreductase (complex IV) in liver mitochondria from rats with hydroxycobalamin[c-lactam]-induced methylmalonic aciduria. J Biol Chem. 266, 20998-21003. Krahenbuhl S, Talos C, Wiesmann U , Hoppel CL (1994). Development and evaluation of a spectrophotometric assay for complex III in isolated mitochondria, tissues and fibroblasts from rats and humans. Clin Chim Acta. 230, 177-187. Livak KJ , Schmittgen TD (2001). Analysis of relative gene expression data using real-time quantitative PCR and the 2(-Delta Delta C(T)) Method. Methods. 25, 402-408. Lowry OH, Rosebrough NJ, Farr AL , Randall RJ (1951). Protein measurement with the Folin phenol reagent. J Biol Chem. 193, 265-275. Morgan PG, Sedensky MM, Meneely PM , Cascorbi HF (1988). The effect of two genes on anesthetic response in the nematode Caenorhabditis elegans. Anesthesiology. 69, 246-251. Rauchova H, Fato R, Drahota Z , Lenaz G (1997). Steady-state kinetics of reduction of coenzyme Q analogs by glycerol-3-phosphate dehydrogenase in brown adipose tissue mitochondria. Arch Biochem Biophys. 344, 235-241. Sottocasa GL, Kuylenstierna B, Ernster L , Bergstrand A (1967). An electron- transport system associated with the outer membrane of liver mitochondria. A biochemical and morphological study. J Cell Biol. 32, 415- 438. Suthammarak W, Yang YY, Morgan PG , Sedensky MM (2009). Complex I function is defective in complex IV-deficient Caenorhabditis elegans. J Biol Chem. 284, 6425-6435. Wittig I, Braun HP , Schagger H (2006). Blue native PAGE. Nat Protoc. 1, 418- 428.

171 Wittig I, Karas M , Schagger H (2007). High resolution clear native electrophoresis for in-gel functional assays and fluorescence studies of membrane protein complexes. Mol Cell Proteomics. 6, 1215-1225.

172

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If you find copyrighted material related to this license will not be used and wish to cancel, please contact us referencing this license number 2526061403012 and noting the reason for cancellation.

Questions? [email protected] or +1-877-622-5543 (toll free in the US) or +1-978-646-2777.

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