Phd thesis AFJ van Aken

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Phd thesis AFJ van Aken

  1. 1. Effects of the expression of alternative oxidase on oxidising pathway kinetics in Schizosaccharomyces pombe mitochondria Alexander Frans Johan van Aken Submitted for the degree of Doctor of Philosophy University of Sussex July 2006
  2. 2. ii I hereby declare that this thesis has not been and will not be submitted in whole or in part to this or any other University for a degree. Signed……………………………………………
  3. 3. iii Acknowledgements I would like to express my gratitude to professor Moore for giving me the opportunity to do a D.Phil. project in his laboratory which has been a bumpy ride at times but was overall a fruitful experience. Although I started working in a different field, having returned to neuroscience I still manage to find the time to further explore my bioenergetic research and being a tutor. I would like to thank the (past) members of the Moore laboratory (Charles, Jane, Paul, Alice, Sarah, Rob, Nick) for their help and support over the years. I would like to particularly thank Dr Mary Albury for her supervision with regards to the yeast expression system. I would also like to thank Dr David Whitehouse for helping me improve my English scientific writing and for having many useful scientific discussions. I would also like to thank professor Derek Lamport for many not so useful scientific discussions. Many thanks also to professor Kros for his scientific support. I would especially like to thank Judita a fellow PhD sufferer over the years for all the support and having spent several years together in suspended animation (hvala lijepa moja učiteljica). Many thanks to my parents and sister for the financial support and for putting up with not seeing me very much over the past four years. Also many thanks to Remon and Angelique for helping out financially at times. Many thanks to all the lovely people I met and am still in contact with after having lived in Kings Road for all the good times and for many visits to Poland, Croatia and Slovakia (Maja, Zuzana, Wojciech, Przemek, Ewa, Basha, Vasek, Marek, Zuzana, Sandra, Tamara, Hidemi, and Gang) hvala, d’akujem and dziękuję.
  4. 4. iv Summary The alternative oxidase (AOX) is a non-protonmotive terminal oxidase found in the respiratory chains of higher plants, various fungi and some protists. Its activity results in dissipation of free energy and affects efficiency of energy transduction in mitochondria. A plant AOX has been heterologously expressed previously in Schizosaccharomyces pombe mitochondria in this laboratory. The work presented in this thesis describes the effects of the expression of AOX on the respiratory kinetics of isolated yeast mitochondria. Succinate dehydrogenase (SDH) is the only respiratory complex which is both a component of the electron transfer chain and the citric acid cycle and possible has a strong regulatory role. It is still relatively unknown how this enzyme is regulated exactly. SDH in potato mitochondria can be activated by ADP, ATP and oligomycin. It has been hypothesized that these substances activate SDH indirectly via an effect on the membrane potential. This hypothesis was tested in a series of experiments using a multi- electrode setup. A characterisation of S. pombe mitochondrial respiratory kinetics is given and determinations of the membrane potential are presented for the first time. Oxidising pathway kinetics of S.pombe mitochondria are notably different from what is seen in mitochondria from other tissues. Results indicate that cytochrome bc1 complex activity is probably the underlying mechanism responsible for these kinetics. The expression of AOX in S. pombe mitochondria showed a substrate dependent difference in oxidising pathway kinetics. It was determined that neither cytochrome pathway or alternative pathway activity could account for these differences.
  5. 5. v Contents Acknowledgements iii Summary iv Contents v Abbreviations xiii Chapter 1 General Introduction 1.1 General background 1 1.1.1 Mitochondria 1 1.1.2 Energy transducing systems 2 1.2 The electron transfer chain 7 1.2.1 General background 7 1.2.2 Complex I 9 1.2.3 Alternative NAD(P)H dehydrogenases 10 1.2.4 Complex II 11 1.2.5 Ubiquinone / Ubiquinol 15 1.2.6 Complex III 17 1.2.7 Cytochrome c 21 1.2.8 Complex IV 22 1.2.9 Complex V 22 1.2.10 Uncoupling protein 23 1.2.11 Alternative oxidase 24 1.3 Schizosaccharomyces pombe 30 1.3.1 General background 30 1.3.2 The respiratory chain of S. pombe mitochondria 32
  6. 6. vi 1.3.3 SDH activation in S. pombe mitochondria 34 1.4 Summary energy transducing systems 35 1.5 A modular representation 36 1.6 Summary 38 1.7 AIMS 38 Chapter 2 Materials and Methods 2.1 Isolation and purification of mitochondria 40 2.1.1 Schizosaccharomyces pombe 40 2.1.1.1 The Expression system 40 2.1.1.2 Yeast transformation 41 2.1.1.3 S. pombe growth 41 2.1.1.4 Isolation of mitochondria from S. pombe cultures 43 2.1.1.5 Spheroplast preparation 43 2.1.1.6 Isolation of mitochondria 43 2.1.1.7 S. pombe media 44 2.1.2 Saccharomyces cerevisiae 46 2.1.3 Potato tuber 46 2.1.3.1 Isolation of mitochondria from potato tubers 47 2.1.3.2 Potato tuber media 47 2.1.4 Arum maculatum 48 2.1.4.1 Isolation of mitochondria from Arum maculatum spadices 48 2.1.4.2 Arum maculatum media 49 2.1.5 Specifics of plant mitochondrial isolation 50 2.2 Polyacrylamide gel electrophoresis & Western analysis 50 2.2.1 SDS-PAGE 50 2.2.2 Blotting to nitrocellulose 50 2.2.3 Immuno-detection of proteins 51 2.3 Protein estimations 51 2.4 Electrochemical techniques 52
  7. 7. vii 2.4.1 The oxygen electrode 53 2.4.2 The Q-electrode 54 2.4.3 The TPP+ -electrode 57 2.4.3.1 The TPP+ -electrode setup 59 2.4.3.2 Detection of [TPP+ o] 60 2.4.3.3 Construction of the TPP+ -electrode membrane 60 2.4.3.4 Conditioning of the TPP+ -electrode 61 2.4.3.5 Calibrating the TPP+ -electrode 62 2.4.3.6 TPP+ -electrode correction 63 2.4.3.7 TPP+ -electrode sensitivity 63 2.4.3.8 Durability of TPP+ -electrodes 64 2.4.3.9 TPP+ -electrode response time 64 2.4.4 Respiratory measurements 65 2.4.4.1 Preparation of respiratory effectors 65 2.4.4.2 Nomenclature 66 2.4.4.3 Basic bioenergetic parameters 67 2.4.4.4 Q-pool kinetics 67 2.4.5 Modelling of Q-pool data 68 2.6 Other methods 70 2.6.1 Spectroscopy 70 2.7 Bioinformatic resources 70 Chapter 3 New insights into the regulation of plant succinate dehydrogenase - revisited 3.1 INTRODUCTION 71 3.1.1 General background and aims 71 3.1.2 The membrane potential in mitochondria 73 3.1.3 Regulation of SDH 75 3.2 RESULTS 77
  8. 8. viii 3.2.1 General characterization 77 3.2.2 Stimulation of SDH by adenine nucleotides 81 3.2.3 Are the effects of ATP on , Qr/Qt and vO2 simultaneous? 86 3.2.4 Stimulation of SDH by adenine nucleotides in the presence of uncoupler 89 3.2.5 Stimulation of SDH by oligomycin 91 3.2.6 Adenine nucleotides and oligomycin inhibit succinate dependent respiration in potato mitochondria 93 3.3 DISCUSSION 99 Chapter 4 Respiratory characteristics of Schizosaccharomyces pombe mitochondria 4.1 INTRODUCTION 112 4.2 RESULTS 113 4.2.1.1 Respiratory rates with different substrates 113 4.2.2 Schizosaccharomyces pombe - cytochrome pathway kinetics 122 4.2.2.1 The relationships of Qr/Qt vs. vO2 and ∆ vs. vO2 under ADP limited conditions with NADH as a substrate 122 4.2.2.2 The relationships of Qr/Qt vs. vO2 and ∆ vs. vO2 under state 3 conditions with NADH as a substrate 124 4.2.2.3 The relationship between Qr/Qt vs. vO2 under uncoupled conditions with NADH as a substrate 127 4.2.2.4 A comparison of NADH and succinate dependent Qr/Qt vs. vO2 and ∆ vs. vO2 relationships under ADP limited conditions 130 4.2.2.5 A comparison of NADH and succinate dependent Qr/Qt vs. vO2 relationships under state 3 conditions 131 4.2.2.6 A comparison of NADH and succinate dependent Qr/Qt vs. vO2 relationships under uncoupled conditions 132 4.2.3 Schizosaccharomyces pombe - reducing pathway kinetics 134 4.2.3.1 SDH reducing pathway kinetics in sp.011 wt mitochondria under state 2 and uncoupled conditions 134
  9. 9. ix 4.2.3.2 External NADH dehydrogenase reducing pathway kinetics in sp.011 wt mitochondria under state 2 and uncoupled conditions 136 4.2.4 Are the biphasic patterns due to cytochrome bc1 complex kinetics? 137 4.2.5 Are biphasic respiratory kinetics a characteristic of yeast mitochondria? 141 4.3 Discussion 142 4.3.1 Respiratory characteristics of S. pombe mitochondria 142 4.3.2 Are the biphasic patterns due to an experimental artefact? 146 4.3.3 Are the biphasic patterns due to cytochrome bc1 complex kinetics? 147 4.3.4 Are biphasic respiratory kinetics a characteristic of yeast mitochondria? 148 Chapter 5 Functional expression of the alternative oxidase in Schizosaccharomyces pombe mitochondria 5.1 INTRODUCTION 150 5.2 RESULTS 153 5.2.1 General characterisation of sp.011 AOX, AOX + T, pREP and wt respiratory kinetics 153 5.2.2 Oxidising pathway kinetics with NADH as substrate 157 5.2.2.1 Comparing sp.011 pREP and wt cytochrome pathway kinetics with NADH as substrate 158 5.2.2.2 Comparing sp.011 AOX and sp.011 AOX + T oxidising pathway kinetics with NADH as a substrate 163 5.2.3 Oxidising pathway kinetics with succinate as substrate 170 5.2.3.1 Comparing sp.011 AOX and sp.011 AOX + T oxidising pathway kinetics with succinate as substrate 170 5.2.4 Oxidising pathway kinetics in sp.011 AOX mitochondria 176 5.2.4.1 Comparing sp.011 AOX oxidising pathway kinetics with either NADH or succinate as a substrate 176 5.2.5 Cytochrome pathway kinetics in sp.011 AOX+T mitochondria 179 5.2.5.1 Comparing sp.011 AOX + T cytochrome pathway kinetics with either NADH or succinate as a substrate 179 5.2.6 Alternative pathway kinetics in sp.011 AOX mitochondria 181
  10. 10. x 5.2.6.1 Comparing sp.011 AOX alternative pathway kinetics with either NADH or succinate as substrate 181 5.2.7 Oxidising pathway kinetics in Arum maculatum mitochondria 183 5.2.7.1 Investigating substrate dependent differences in oxidising pathway kinetics in Arum maculatum mitochondria 183 5.3 DISCUSSION 191 5.3.1 Differences between the various S. pombe mitochondria used 191 5.3.2 Does transformation of S. pombe mitochondria lead to changes in respiratory kinetics? 192 5.3.3 Comparing sp.011 AOX and sp.011 AOX+T oxidising pathway kinetics with NADH as a substrate 192 5.3.4 Does AOX activity affect  in S. pombe mitochondria? 193 5.3.5 Comparing sp.011 AOX and sp.011 AOX+T oxidising pathway kinetics with succinate as a substrate 195 5.3.6 Why are the oxidising pathway kinetics obtained in this study different from Affourtit’s study? 196 5.3.7 Are there any substrate dependent differences in sp.011 AOX oxidising pathway kinetics? 196 5.3.8 Are the substrate dependent differences a characteristic of the cytochrome pathway? 197 5.3.9 Are the substrate dependent differences a characteristic of the alternative pathway? 197 5.3.10 Are the substrate dependent differences a characteristic of the expression system used? 198 5.3.11 Can the alternative pathway compete with the cytochrome pathway in S. pombe mitochondria expressing AOX? 198 5.3.12 Does expression of the alternative oxidase lead to a change in Q-pool behaviour? 198 5.4 CONCLUSION 199
  11. 11. xi Chapter 6 Modelling of oxidising pathway kinetics in Schizosaccharomyces pombe mitochondria expressing the alternative oxidase 6.1 INTRODUCTION 200 6.1 RESULTS 201 6.1.1 Modelling of sp.011 AOX oxidising pathway kinetics 201 6.1.2 Are the oxidising pathway activities additive? 204 6.1.3 sp.011 AOX mixed titration studies 207 6.1.4 Applying Q-pool kinetics to fit sp.011 AOX mixed substrate oxidising pathway data 212 6.3 DISCUSSION 215 6.4 CONCLUSION 222 Chapter 7 General Discussion 7 General Discussion 223 7.1 Characterisation of the wild type S. pombe mitochondria 224 7.1.1 How does cytochrome bc1 complex activity lead to biphasic cytochrome pathway kinetics in S. pombe mitochondria? 225 7.1.2 Future work suggestions pertaining to the biphasic patterns in S. Pombe cytochrome pathway kinetics 230 7.2 Functional expression of AOX in S. pombe mitochondria yields substrate dependent differences in oxidising pathway kinetics 231 7.2.1 Does AOX activity affect  in S. pombe mitochondria ? 232 7.2.2 Substrate dependent differences in oxidising pathway kinetics of S. pombe mitochondria expressing AOX 232 7.2.3 What causes the substrate dependent differences in oxidising pathway kinetics in S. pombe mitochondria expressing AOX? 233 7.2.4 Are the substrate dependent differences in oxidising pathway kinetics 234
  12. 12. xii in S. pombe mitochondria expressing AOX due to dehydrogenase characteristics? 7.2.5 Future work suggestions pertaining to the substrate dependent oxidising pathway kinetics in S. pombe mitochondria expressing AOX 236 7.3 Conclusion 237 Appendix 1 238 Appendix 2A 241 Appendix 2B 242 References 244
  13. 13. xiii Abbreviations  H ~  proton electrochemical gradient  membrane potential G Gibbs free energy p protonmotive force AA antimycin A ADH alcohol dehydrogenase ADP adenosine 5'-diphosphate AK adenylate kinase AMP adenosine 5'-monophosphate ANC adenine nucleotide carrier AOX alternative oxidase ATP adenosine 5'-triphosphate BCA bicinchoninic acid BSA bovine serum albumin CAT carboxyatractyloside CCCP carbonyl cyanide m-chlorophenylhydrazone DCCD N,N’-dicyclohexylcarbodiimide DCIP 2,6-dichlorophenolindophenol DNP 2,4-dinitrophenol E° standard redox potential ETC electron transfer chain G6P Glucose-6-Phosphate G6PD Glucose-6-Phosphate dehydrogenase F Faraday constant (9.65104 C mol-1 ) FAD flavin adenine dinucleotide FMN flavin mono-nucleotide ISP iron-sulfur protein IMM inner mitochondrial membrane
  14. 14. xiv IMS intermembrane space KCN potassium cyanide NADH nicotinamide adenine dinucleotide, reduced form OAA oxaloacetate OG octyl gallate OMM outer mitochondrial membrane MK menaquinone Pi inorganic phosphate pmf protonmotive force PmitoKATP plant mitochondrial K+ ATP channel PMS phenazine methosulfate Q ubiquinone QH2 ubiquinol Qr/Qt Q-redox poise R gas constant (8.31 J mol-1 K-1 ) RCR respiratory control ratio ROS reactive oxygen species SDH succinate dehydrogenase SHAM salicyl hydroxamic acid SMP submitochondrial particle SQOR succinate:quinone oxidoreductase T absolute temperature (K) TPP+ tetraphenyl phosphonium UCP uncoupling protein vO2 oxygen consumption rate z valence number
  15. 15. 1 Chapter 1 General introduction 1.1 General background 1.1.1 Mitochondria—Mitochondria are double walled organelles found in eukaryotic cells (Figure 1.1). The innermost compartment, the matrix, is separated from the intermembrane space (IMS) by the inner mitochondrial membrane (IMM) which is relatively impermeable to ions and large solutes. The outer mitochondrial membrane (OMM) on the other hand is relatively permeable to most solutes with a molecular weight less than 10 kDa [1] and because of this the intermembrane space is assumed to be continuous with the cytosol. The IMM shows numerous invaginations (cristae). Whether or not the cristae are continuous with the intermembrane space is still under investigation [2]. In this study it is assumed that under in vitro conditions (isolated mitochondria in solution) there are only two compartments, the matrix and outside of the matrix. Figure 1.1 A schematic representation of a mitochondrion. (Source: courtesy of Dr. Michael W. Davidson, Florida State University). Mitochondria are typically depicted in this ‘sausage’ form but the shape can vary dramatically depending on tissue and/or developmental state.
  16. 16. 2 Mitochondria are known as the powerhouses of cells responsible for the generation of ATP, the cellular energy currency. ATP is used to drive many thermodynamically unfavourable processes in the cell and a continuous supply is needed in order for the cell to survive. For instance, to continuously maintain a resting membrane potential most cells expend as much as 30% of cellular ATP to keep the Na+ /K+ exchanger active, neurons in the central nervous system have to expend as much as 70% [3]. A healthy complement of mitochondria is therefore vital for cellular functioning. The energy released upon hydrolysis of the terminal anhydride bond of the ATP molecule is used to drive the uphill process to which ATP hydrolysis is coupled. Hydrolysis of one mole of ATP under normal cellular conditions, i.e. where the mass action ratio of the ATP synthesis reaction is kept away 10 orders of magnitude from equilibrium, can release 57 kJ of energy. The ratio of ATP to ADP concentration in the cytosol is typically maintained at a value of 1000 [1]. Mitochondria do not just generate ATP at a constant rate, ATP synthesis is tightly regulated and mitochondrial activity is highly flexible depending on energetic conditions. Given that cells rely heavily on the efficient generation of ATP by the mitochondria it is curious that respiratory chains of many organisms contain respiratory complexes which actively reduce this efficiency by dissipating energy. Two of these complexes are the uncoupling protein (UCP) (see section 1.2.10) and the alternative oxidase (AOX) (see section 1.2.11). In order to understand how these complexes can reduce the efficiency with which ATP is synthesized an understanding of energy transducing systems is needed. 1.1.2 Energy transducing systems—Returning to the example of neurons, the brain needs a continuous supply of oxygen and glucose; temporary shortages of either of them (e.g. ischaemia) can lead to disastrous results in which cells may go into apoptosis. Upon restoring the supply of oxygen and glucose things may get even worse as happens under the conditions of reperfusion injury [4] and excitotoxicity [3], during which mitochondrial processes set off a series of unfortunate events which lead to cell death. During oxygen deprivation mitochondria become ‘highly reduced’; when oxygen becomes available again this increased reduction level leads to the formation of reactive oxygen species (ROS) which leads to the breakdown of membranes jeopardizing cellular integrity.
  17. 17. 3 In order to understand how the synthesis of ATP, the consumption of glucose and oxygen and the formation of ROS are related a brief description of energy transducing systems will be given. All organisms need a continuous energy supply in order to prevent a state of thermodynamic equilibrium (death). Energy is readily available in the form of electromagnetic rays from the sun for those organisms which can trap this form of energy to subsequently transduce it into another form. Most other organisms derive energy from the breakdown of ‘energy rich’ compounds, such as glucose. The breakdown of glucose under standard conditions yields 2870 kJ mol-1 . Most energy utilising reactions in the cell require between 10 to 50 kJ mol-1 [5] so there is a need to partition the energy released during breakdown of glucose. ATP, releasing 57 kJ mol-1 upon hydrolysis (under cellular conditions) is used predominantly. Some ATP is generated through substrate level phosphorylation (about 5% [6]), in the presence of molecular oxygen however the bulk of cellular ATP is generated via energy transducing reactions. Basically all energy transducing systems operate along the same principles: two proton pumps, located in the same membrane (which is relatively impermeable to protons and other ions) their activities coupled to each other via a proton current. In Figure 1.2 the situation as it occurs in mitochondria is shown schematically. By convention the matrix is considered the N side (N for negative) and the intermembrane space the P side (P for positive). The so called primary pump utilises electrons1 to drive the transport of protons against their concentration gradient from the matrix to the intermembrane space. This creates a protonmotive force (pmf) which is subsequently utilised by the secondary pump to drive the synthesis of ATP via the influx of protons from the intermembrane space to the matrix. 1 It would be more correct to use the term ‘reducing equivalents’ as will be explained further in section 1.2.
  18. 18. 4 Figure 1.2 A schematic representation of an energy transducing membrane containing two proton pumps communicating with each other via a proton circuit. N: negative P: positive. The pmf, or p, is a driving force with units of V, which consists of two components: a concentration gradient of protons (pH) and an electrical potential difference (). Displacement of ions across a membrane generates an electrochemical potential which is expressed in kJ mol-1 (units of energy). The change in free energy (G) upon the transport of 1 mol of protons across a membrane (in the absence of a ) is given by the following equation: [1.1] R: gas constant (8.31 J mol-1 K-1 ) T: absolute temperature (K) i: inside o: outside          o i H H RTmolkJG ][ ][ log3.2)1(
  19. 19. 5 The free energy change associated with the separation of 1 mol of univalent ions across a membrane (in the absence of a concentration gradient) is given by:  zFmolkJG )1( [1.2] z: valence number (1 in this case) F: Faraday constant (9.65104 C mol-1 ) Protons in the matrix and the intermembrane space normally will be affected by both a concentration gradient and an electrical gradient which gives:            o i H H RTzFmolkJG ][ ][ log3.2)1( [1.3] This Gibbs energy difference is generally referred to as the proton electrochemical gradient:  H ~  And with the definition for pH (pH = - log [H+ ]) the equation can be further simplified: pHRTFmolkJH    3.2)( ~ 1  [1.4] To facilitate comparison with redox potential differences in the electron transfer chain (ETC) Mitchell [1] defined the term protonmotive force (p) which is: FmVp H /)( ~         [1.5]
  20. 20. 6 p is expressed in units of V and substituting values for R and T at 25 C the equation simplifies to: pHmVp  59)( [1.6] A good understanding of these basic equations is necessary to appreciate the method with which membrane potentials were determined in this study. This topic will be discussed in detail in chapter 2.
  21. 21. 7 1.2 The electron transfer chain 1.2.1 General background—Figure 1.3 shows the ETC as it is organised in the mitochondria of mammals. The various components within the chain are organised according to their redox potentials in order of increasing value. Substrates (e.g. NADH or succinate) can be oxidised at specific locations where they donate reducing equivalents (a reducing equivalent can be defined as 1 mole of hydrogen atoms, one proton and one electron per H atom [6]). The red arrows indicate the transfer of electrons through the chain, which eventually reduce oxygen to water at complex IV. At three sites (complexes I, III and IV) the transfer of electrons is coupled to the translocation of protons from the matrix to the intermembrane space (blue arrows), this generates the aforementioned pmf, the matrix being negative with respect to the IMS. Protons can re-enter the matrix via complex V, a process which is coupled to the synthesis of ATP from ADP and Pi. Another inward pointing arrow indicates the passive leak of protons back into the matrix, it is postulated that protons can traverse the IMM via the junctions between lipid and protein [1]. Apart from leak and ATP synthesis there are many transporters (symporters and antiporters) which utilise the proton electrochemical gradient to drive the translocation of metabolites across the IMM (not shown in the figure). Overall, oxygen is consumed and substrates are oxidised, as a result of this, energy is stored in a pmf, which can be utilised by the ATP synthase to drive the reaction of ATP synthesis, this process is known as oxidative phosphorylation. The components of the mammalian ETC are: complexes I to V, the Q pool and cytochrome c. With respect to the relative abundance of complexes within the ETC the following stoichiometry is currently accepted: complexes I : II : III : IV : cytochrome c : ubiquinone = 1 : 2 : 3 : 7 : 14 : 63 [7]. The plant ETC contains the same components but is more complicated than its mammalian counterpart due to the presence of some extra respiratory proteins, see Figure 1.4. The plant ETC contains several alternative NAD(P)H dehydrogenases, which are non-protonmotive, two of them located on the inner leaflet of the IMM and two on the outer leaflet. Another component is the alternative oxidase, which like complex IV catalyses the reduction of molecular oxygen to water [8].
  22. 22. 8 Figure 1.3 Schematic representation of the mammalian ETC. I : complex I (NADH dehydrogenase), II : complex II (succinate dehydrogenase), III: complex III (ubiquinol:cytochrome c oxidoreductase), IV: complex IV (cytochrome c oxidase), V: complex V (ATP synthase), c: cytochrome c, Q: the Q-pool (ubiquinone + ubiquinol). Blue arrows: proton flow. Red arrows: electron flow. Also indicated is the non specific leak of protons across the IMM. The route taken by electrons transferred from QH2 to complex III (and subsequently to cytochrome c to complex IV) is referred to as the cytochrome pathway. Electrons transferred to AOX are said to use the alternative pathway. The main difference between these two pathways is that the alternative pathway is non-protonmotive [8]. Figure 1.4 Schematic representation of the plant ETC which contains several additional components compared to the mammalian system (see Figure 1.3). NDH (non-protonmotive NADH dehydrogenase), AOX (alternative oxidase).
  23. 23. 9 A physical description of the components of the ETC will be given in the remainder of this section. The alternative oxidase will be discussed in detail given its importance in this study. Also complexes II and III will be discussed in somewhat more detail because a thorough understanding of the functioning of these respiratory proteins is necessary in order to interpret the acquired experimental results. 1.2.2 Complex I (NADH:quinone oxidoreductase, NADH dehydrogenase): Complex I catalyses the transfer of two electrons to ubiquinone in a reaction coupled to proton translocation across the IMM. Currently the proton translocation stoichiometry is believed to be 4H+ /2e- [1]. Of all the complexes involved in oxidative phosphorylation, complex I is by far the largest. In mammalian mitochondria it consists of 43 subunits with a total molecular weight in the range of 750-1000 kDa. Not all of these subunits are required for electron transfer as it was found that bacteria contain a minimal functional unit of just 14 subunits [1]. Complex I is normally taken to be L-shaped with a hydrophilic and a hydrophobic part. The hydrophilic part contains a flavin mono-nucleotide (FMN) moiety which is reduced by NADH, electrons subsequently are transferred through 8 or 9 iron sulfur clusters (FeS) where a molecule of ubiquinone (Q) accepts the electrons. Complex I is both nuclear (nDNA) and mitochondrial (mtDNA) encoded and is potently inhibited by rotenone, piericidin A [9] and rhein [10]. Recently, complex I defects caused by pathogenic mutations in mtDNA and nDNA have been linked to various neurodegenerative diseases such as Parkinson’s disease [9]. Defective complex I functioning leads to a decreased H+ and a concomitant decrease in ATP production whereas ROS formation is stimulated.
  24. 24. 10 1.2.3 Alternative NAD(P)H dehydrogenases [11, 12] : In mammalian mitochondria the only ETC complex able to accept reducing equivalents from NADH is complex I. In the mitochondria of plants and fungi (including S. pombe) one or more alternative NAD(P)H dehydrogenases can be found. Like complex I these dehydrogenases catalyse the transfer of two electrons to ubiquinone, however this reaction is not coupled to proton translocation across the IMM, therefore no energy is conserved. Another difference is the use of a flavin adenine dinucleotide molecule (FAD) as a redox prosthetic group instead of FMN. The external NADH dehydrogenase (Ext. NDH) and the external NAD(P)H dehydrogenase (Ext. N(P)DH) are situated at the outer leaflet of the IMM facing the IMS. The internal NADH dehydrogenase (Int. NDH) and the internal NAD(P)H dehydrogenase (Int. N(P)DH) are situated at the inner leaflet of the IMM facing the matrix. Unlike complex I all the alternative NADH dehydrogenases are believed to be relatively small with only one to four subunits. Complex I inhibitors have no effect on the alternative NAD(P)H dehydrogenases and any inhibitors which do affect these complexes are rare and mostly unspecific. It is hypothesized that alternative NADH dehydrogenases can be employed as a dynamic response to changing metabolic needs. Given their small size they can be made readily available as opposed to complex I which requires 43 subunits to be expressed. Varied expression and activity of the alternative NAD(P)H dehydrogenases and the alternative oxidase provides flexibility in regulating the redox state of cytoplasmic and mitochondrial matrix NAD(P)H pools. Mitochondria of some organisms lack complex I completely (e.g. S. cerevisiae and S. pombe [13]) and they are dependent on alternative NADH dehydrogenases to oxidise matrix generated NADH. External NADH dehydrogenase: The Ext. NDH dependent oxygen uptake can be stimulated by the presence of divalent cations which electrostatically screen negative membrane charges. Also Ext. NDH has a high affinity for calcium binding which is believed to affect the interaction with ubiquinone. Early work done on external NADH oxidation gave ADP/O* values between 1.2 and 1.4 whilst NADH oxidation could be inhibited with antimycin A (AA, complex III inhibitor) and cyanide (complex IV inhibitor). These observations indicate that electrons enter the ETC just before complex III. Its molecular weight is estimated to be 32 kDa. * the amount of ADP molecules converted to ATP molecules per atom of oxygen , see section 2.4.4.3.
  25. 25. 11 External NAD(P)H dehydrogenase: The Ext. N(P)DH has similar ADP/O values as the Ext. NDH and it is also inhibited by AA and cyanide indicating a point of entry in the ETC just before complex III. The Ext. NDH and Ext. N(P)DH have different pH profiles. Also Ext. N(P)DH is more calcium dependent than Ext. NDH. A protein doublet with molecular weight 58 kDa, localized to the outer surface of the IMM, was found to oxidize both NADH and NADPH. Internal NADH dehydrogenase: Internal NADH oxidation, in the presence of rotenone, showed ADP/O values of 1.5 in plant mitochondria. It was also found that the Int. NDH had a ten times lower affinity for NADH than complex I. No calcium activation has been found so far. In S. cerevisiae mitochondria a single polypeptide with a weight of 53 kDa was identified as an internal NADH dehydrogenase. Internal NAD(P)H dehydrogenase: Unlike the Int. NDH the Int. N(P)DH is activated by calcium. Apart from NAD(P)H it possibly also oxidizes NADH. Its molecular weight is estimated to be 43 kDa. A complex identified as an NADPH dehydrogenase in one species may be found in another species where it can only oxidise NADH, this illustrates that the alternative NADH dehydrogenases still require a lot of research. 1.2.4 Complex II (succinate dehydrogenase) [14, 15]: Succinate dehydrogenase (SDH) is the only ETC complex which is also a component of the citric acid cycle and fulfils therefore a dual role, being active in both the process of energy transduction and the generation of carbon intermediates for biosynthetic metabolism. SDH is a member of the succinate:quinone oxidoreductases (SQOR, EC 1.3.5.1). SQORs couple the oxidation of succinate to fumarate to the reduction of quinone to quinol [16]: succinate  fumarate + 2H+ + 2e- quinone + 2H+ + 2e-  quinol
  26. 26. 12 This oxidoreduction reaction is not coupled to proton translocation therefore complex II does not contribute to the conservation of energy. In vitro, SQORs can catalyse both succinate oxidation and fumarate reduction, be it at different rates. By providing substrate in excess, directionality is achieved under experimental conditions. SQORs consist of four subunits referred to as A, B, C and D, see Figure 1.5. Figure 1.5 Succinate dehydrogenase. The hydrophilic subunits A and B are exposed to the matrix (negative side). The hydrophobic subunits C and D are situated within the IMM. The SDH shown here is a type C SQOR (class 3) which is normally found in eukaryotic mitochondria. Adapted from Figure 2C in [14]. The presence of a single heme group (indicated by the rectangle within subunits C and D) and the presence of two hydrophobic subunits are indicative of an eukaryotic SDH. Other classes show variations in the amount of heme groups and hydrophobic subunits. Another way of classifying SQORs is on the basis of quinone substrate. In this study the respiratory activity of yeast and plant mitochondria was investigated therefore no description of archeal and bacterial SQORs will be given here, for more information on these complexes see Refs 14 and 15. Although most bacteria express SDH of a form different from what is found in eukaryotic species, recently acquired X-ray structures show that SDH in E. coli would be classified equivalent to the mammalian complex [16, 17], see Figure 1.6.
  27. 27. 13 Figure 1.6 Three dimensional structure of the E. coli SDH taken from Figure 1C in [17]. Subunits A and B are coloured teal and purple respectively. Subunits C and D are shown in orange and yellow respectively. Prosthetic groups shown are covalently bound FAD (subunit A), [2Fe-2S], [4Fe-4S] and [3Fe-4S] iron-sulfur centers (subunit B). Subunits C and D display bound quinone (black) and heme b556 (magenta). The E. coli SDH is equivalent to the SQOR type normally found in eukaryotic mitochondria. Subunit A (also known as the flavoprotein Fp or CII-1) contains a covalently bound FAD prosthetic group and the dicarboxylate binding site; its molecular weight is 70 kDa. Subunit B (also known as the iron-sulfur protein or CII-2) contains three iron-sulfur clusters, [2Fe- 2S], [4Fe-4S] and [3Fe-4S] (also known as Centers 1-3) and weighs 27 kDa. Subunits C and D (also referred to as anchor proteins or CII-3 and CII-4 respectively) contain the quinone reduction and oxidation sites and one heme group (in eukaryotes), their molecular
  28. 28. 14 weights are 15 and 13.5 kDa respectively. Subunits A and B have a high sequence homology amongst species, whereas this is much lower for subunits C and D. All subunits are nuclear encoded making complex II unique in the sense that the other main ETC complexes (I, III, IV and V) are all partially encoded by mitochondrial DNA. Subunits A and B are hydrophilic and extend into the matrix. Subunits C and D are hydrophobic and span the IMM, i.e. parts of subunits C and D are accessible from the cytosolic side. Electron transfer though complex II is linear and experimental data have shown that electron transfer through SDH is not sensitive to uncouplers [17]. Succinate binds to the Fp unit where it subsequently donates two electrons and two protons to the FAD group reducing it to FADH2. From there electrons are transferred to the IP unit where they pass through the three iron-sulfur centres. The electrons end up reducing ubiquinone to ubiquinol where two protons are taken up from the matrix. At present the role of heme in the electron transfer from succinate to ubiquinone is unclear. Inhibitors: Malonate and oxaloacetate (OAA) are potent competitive inhibitors of SDH. Several inhibitors interfere with quinone binding such as 2-thenoyltrifluoroacetone (TTFA) and 3- methyl-carboxin and 2-n-heptyl-4-hydroxyquinoline-N-oxide (HQNO). These inhibitors do not interfere with the activity of the solubilised enzyme. The site of action is between Center 3 and the Q-pool. Apparently cyanide disrupts Center 3 in SDH [18]. Intracellular oxidation of succinate is inhibited by fluoride [19]. Regulation of SDH: SDH in isolated mitochondria is in a partially deactivated state due to bound OAA [15]. The slowness of the reaction and the high energy of activation (35.6 kcal/mol* ) of SDH was interpreted as a conformational change in the enzyme [20]. SDH can be activated by ATP and ADP [21]. Chapter 3 deals with SDH activation and regulation of SDH is discussed more in-depth in its introduction, see section 3.1.3. * 149 kJ/mol
  29. 29. 15 1.2.5 Ubiquinone / Ubiquinol (Q-pool): The electron transfer chain has two mobile pools of electron carriers: the cytochrome c pool (see section 1.2.7) and the Q-pool. The Q-pool consists of both oxidised and reduced forms of ubiquinone. Ubiquinone (Q) undergoes an overall 2H+ + 2e- reduction to form ubiquinol (QH2). Because of its long hydrocarbon side chain both Q and QH2 are highly hydrophobic [1]. Figure 1.7 shows the chemical structure of Q. The side chain can vary in length depending on species, n = 10 in mammalian mitochondria [1], whereas plant mitochondria contain a mixture of ubiquinone molecules with n = 9 and 10 [22]. In yeast mitochondria n = 6 [1]. Figure 1.8 shows the two step reduction of Q. After the first step a highly reactive intermediate (semiquinone) is formed, if allowed to react with molecular oxygen it would lead to ROS formation. Hence the necessity to firmly bind semiquinone during reduction of Q to QH2 as will be discussed in 1.2.6. It is generally assumed that the Q-pool functions as a homogenous pool of electron carriers which forms the linkage between dehydrogenases (complexes I and II and the alternative NADH dehydrogenases) and ubiquinol oxidases (complex III and AOX). In mammalian submitochondrial particles (SMP) the activity of both complex I and II were found to be linearly dependent on the Q redox poise (QH2 / (Q+QH2)). A linear relationship was also found between the dependency of complex III activity on Q redox poise. These linear dependencies from both dehydrogenases and ubiquinol oxidases on the level of Q reduction are commonly referred to as quinone pool behaviour [23, 24]. Figure 1.7 Structure of ubiquinone as adapted from Figure 5.6 in [1]. The length of the side chain (R) can vary, n = 10 in mammalian mitochondria, in plant mitochondria a mixture of n=9 and n=10 is found. n = 6 in yeast mitochondria2 and n = 9 in amoebal mitochondria. 2 This figure does not hold for all yeasts, e.g. Candida utilis mitochondria contain both UQ7 and UQ9 [25].
  30. 30. 16 Quinone Semiquinone Quinol Figure 1.8 Two step reaction of quinone reduction to ubiquinol. The homogeneity of the Q-pool was deduced from the observation that antimycin A titrations (which inhibit complex III) do not result in a linear response. Figure 1.9 shows a plot with fictitious data representing two situations: one in which every Q molecule is in a fixed relationship with a bc1 complex () and one in which all Q molecules are free to engage with different bc1 complexes (). In the first situation inhibition titrations with AA would lead to a linear relationship. In the second situation Q molecules can still donate electrons to uninhibited bc1 complexes and overall electron flux, expressed as respiratory activity is seen to be ‘antimycin resistant’. The antimycin resistant respiratory kinetics are normally seen in mitochondria [24, 26]. In an article on pool behaviour of Q (and cytochrome c) in S. cerevisiae mitochondria Boumans et al. [27] reported non-homogenous pool behaviour and a linear relationship between respiratory activity and complex III inhibition was found; from this it was concluded that in yeast mitochondria the respiratory components of the ETC are arranged as an ordered macromolecular assembly which does not allow for diffusion based collisions between components. Another study done in the same year by Rigoulet et al. [28] showed that S. cerevisiae mitochondria do show homogenous pool behaviour (cf. Figure 3B in [28]). Several studies showed that AOX (from either Candida albicans or Lycopersicon esculentum (tomato)) expressed in S. cerevisiae mitochondria [29] and [30] respectively, could utilise QH2 as a substrate, which implies Q-pool behaviour in S. cerevisiae. e- e- + 2H+
  31. 31. 17 Figure 1.9 Theoretical antimycin A titration data points illustrating homogenous () and non-homogenous Q pool () behaviour in mitochondria, see text for details. 1.2.6 Complex III (ubiquinol:cytochrome c oxidoreductase, bc1 complex): The bc1 complex is a protonmotive homodimer catalysing the oxidation of ubiquinol to the reduction of cytochrome c. In bovine heart and S. pombe mitochondria each monomer consists of 11 subunits of which 8 do not have a catalytic role in the oxidation of ubiquinol [31]. Complex III in S. cerevisiae mitochondria contains 10 subunits. A presequence targeting the Rieske protein is cleaved from the protein; in bovine heart and S. pombe mitochondria this cleaved presequence is retained as a subunit whereas in S. cerevisiae [31] and in potato [32] it is degraded. The redox groups consist of a 2Fe/2S centre which is located on the iron-sulfur protein (ISP), two B-type heme groups (bL and bH) located on a single polypeptide and the heme of cytochrome c1 [1] (see Figure 1.10). In many bacteria a functionally similar but structurally simpler version of the bc1 complex is found in the plasma membrane. These complexes have the same electron transfer and proton translocation functionality as their mitochondrial counterparts. Paracoccus for instance
  32. 32. 18 only has a basic three subunit enzyme similar to the protein complex in Figure 1.10. This indicates that the supernumerary subunits are not required for electron transfer or proton translocation [31]. Crystal structures of the bc1 complex have become available in recent years, see Figure 1.11 for the S. cerevisiae bc1 complex structure. Figure 1.10 Schematic representation of the bc1 complex. Only the subunits containing redox groups are shown. The iron-sulfur protein (ISP) also referred to as the Rieske protein. The b-type hemes containing polypeptide and the cytochrome c1 subunit. The midpoint potentials at pH 7 for the redox centres in the yeast bc1 complex are: ISP +280 mV, cytochrome c1 +240 mV, bL –30 mV and bH +120 mV [33]. In order to understand the pathway of electron flow through the bc1 complex an understanding of the Q-cycle [32] is needed, see Figure 1.12. In a complete turnover of the Q-cycle two molecules of ubiquinol are oxidised, one molecule of ubiquinone is reduced, 2 protons are taken up from the matrix, 4 protons are released in the IMS and two cytochrome c1 groups are reduced [1]. The Q-pool in the IMM exists in large molar excess over the bc1 complexes with a ratio of 21:1 [7]. In stage 1 a molecule of ubiquinol diffuses to the binding site Qp (p for positive as it is situated near the positive site of the IMM) where it is oxidised in several stages: One electron is transferred to the ISP, two protons are released to the cytosol and a semiquinone molecule (see Figure 1.7) remains temporarily bound at Qp. The second electron is transferred to bL. The electron transferred to the ISP passes down the ETC to ISP cyt c1 bL bH IMS matrix
  33. 33. 19 cytochrome c1, cytochrome c and cytochrome c oxidase. The electron on bL passes onto bH. This electron is used to reduce a molecule of ubiquinone, at another binding site Qn, to semiquinone which remains bound there until a next molecule of ubiquinol comes along in the second part of the cycle. Figure 1.11 Structure of the S. cerevisiae bc1 complex taken from Figure 1A in [34]. The bc1 complex shown in its homodimeric form. Cytochrome c1 is shown in red, the Rieske protein in green, cytochrome b in blue, the hinge domain in cyan. Antibodies binding to the bc1 complex are shown in orange. Cytochrome c bound to one monomer is shown in yellow. All redox prosthetic groups are shown in black. IMS: intermembrane space. IM: inner membrane. MA: matrix.
  34. 34. 20 Figure 1.12 The Q-cycle in mitochondria, adapted from Figure 5.14 in [1]. P: positive N: negative See text for explanation.
  35. 35. 21 When a second molecule of ubiquinol binds to Qp some of the steps in stage 1 are repeated. One electron is transferred to ISP, again 2 protons are released into the cytosol. One electron is transferred to bL. The electron on ISP is passed down the ETC to cytochrome c1, cytochrome c and cytochrome c oxidase. The electron on bL is transferred to bH. The semiquinone molecule still bound at Qn is reduced by bH and to complete the full reduction of semiquinone to ubiquinol 2 protons are taken up from the matrix. This completes the Q- cycle [1]. Inhibitors: Figure 1.12 shows two inhibition sites in the bc1 complex indicated as red bars in stage 1. Myxothiazol blocks events at Qp and stigmatellin inhibits electron transfer to the Rieske protein. Antimycin A acts at Qn, preventing reduction of ubiquinone by bH [1]. 1.2.7 Cytochrome c: Cytochrome c, a mobile redox carrier, is a peripheral protein located on the P-face of the IMM which transfers electrons between complex III and IV and can be readily solubilised from intact mitochondria. Electrons can be donated artificially from molecules such as tetramethyl-p-phenylene diamine (TMPD) and electrons can leave the ETC via cytochrome c through reduction of ferricyanide (Fe(CN)6 3- ) [1]. Cytochrome c is nuclear encoded and has a molecular weight of about 13 kDa [35]. Apart from being an electron carrier, cytochrome c plays a role in the process of apoptosis. Upon induction of cell death cytochrome c leaves the confinement of the IMM and diffuses to the cytosol where it initiates the activation of caspases, a family of cysteine proteases [35].
  36. 36. 22 1.2.8 Complex IV (cytochrome c oxidase): Cytochrome c oxidase in mitochondria consists of up to 13 subunits in mammalian mitochondria (11 in yeast) [36] of which only two (subunits I and II) are involved in electron transfer and proton translocation. Complex IV is present in the IMM as a homodimer. The complex catalyses the complete reduction of oxygen to water and pumps protons across the IMM with a stoichiometry of 2H+ /2e- . Four electrons are transferred sequentially from the cytochrome c pool to complex IV. Subunit II contains a copper centre (CuA) which has two copper ions in a cluster with sulfur atoms. This complex accepts electrons from cytochrome c one at a time. Subunit I contains two heme groups (heme a and heme a3) and another copper centre (CuB). Electrons from CuA are transferred to heme a onto heme a3 and finally onto CuB where oxygen is reduced to water. Heme a3 is also the binding site for several complex IV inhibitors: cyanide, azide, nitric oxide and carbon monoxide [1]. A regulatory effect of adenine nucleotides on complex IV is well known, addition of ATP to yeast mitochondria leads to an increase in enzymatic capacity of cytochrome c oxidase but does not stimulate respiration rate [36]. 1.2.9 Complex V (ATP synthase, F1.Fo-ATPase): Complex V is a proton pump which couples the hydrolysis of one molecule of ATP to ADP and Pi to the translocation of three protons across the IMM. The name F1.Fo-ATPase indicates the two mayor components of this complex, the hydrophobic Fo complex (160kDa) which translocates protons and the F1 complex (370 kDa) which contains the catalytic and regulatory sites. The Fo complex is located in the IMM whereas the F1 part extends into the matrix. The electrochemical gradient of protons generated through ETC activity can be used to drive the synthesis of ATP and the ATP synthase is seen to operate in reverse. The influx of protons drives the thermodynamically
  37. 37. 23 unfavourable reaction of ATP synthesis [1] and complex V can be considered a p consumer [37]. The Fo and F1 complexes can be targeted directly by specific inhibitors. Oligomycin (hence the o in Fo) and venturicidin bind at the Fo complex. Aurovertin and efrapeptin bind at the F1 complex. Dicyclohexylcarbodiimide (DCCD) has inhibitory effects on both complexes [1]. The ATP synthase is regulated by the natural inhibitor protein IF1 which binds to a - subunit from the F1 complex [38]. The binding interferes with the cooperative ATP synthesis process of complex V. Binding of the protein inhibits hydrolytic activity and it is suggested that it can also inhibit ATP synthesis [39, 40]. High p induces release of IF1 [40] and the off-rate (rate of release) is high. High concentration of ATP induces binding of IF1 and the on-rate (rate of binding) is high under these conditions [39]. The equilibrium between the on-rate and off-rate determines the steady state inhibition of the ATP synthase by IF1. The IF1 protein was found in mammalian [39, 40], plant [41] and yeast mitochondria [42]. Inhibitory proteins can cross-react with mitochondria from other sources [38, 42]. 1.2.10 Uncoupling protein: The uncoupling proteins (UCP) are a subfamily of the mitochondrial carriers (MC) [43] of which 5 types have been identified so far: UCP1-5. The first UCP to be found was UCP1 in brown adipose tissue where it functions to drive non-shivering thermogenesis in hibernators, cold-adapted rodents and newborn mammals [1]. The activity of the protein is stimulated by fatty acids and inhibited by nucleotides. By catalysing proton transport into the matrix it dissipates the p by increasing the proton conductivity across the IMM [43]. The p dissipation would lead to an increase in body temperature, another beneficial purpose of stimulating UCPs would be to control ROS production [43, 44].
  38. 38. 24 1.2.11 Alternative oxidase (AOX): General background: Plant mitochondria exhibit cyanide resistant respiration to various degrees, potato tuber mitochondria show only a little resistance to inhibition with cyanide whereas mitochondria isolated from aroid spadix tissues seem to be fully resistant [8]. This cyanide resistant respiration is associated with the presence of an extra terminal oxidase (apart from complex IV) which functions as an ubiquinol:oxygen oxidoreductase and is referred to as the alternative oxidase (AOX). It catalyses the complete reduction of oxygen to water [45]. Although generally referred to as cyanide resistant, AOX is insensitive to cytochrome pathway inhibitors in general (e.g. antimycin A, azide and carbon monoxide). AOX is sensitive to hydroxamic acid derivatives (such as salicylhydroxamic acid (SHAM) which interferes with Q-binding [46]) and alkyl gallates (such as octyl gallate and dodecyl gallate [47]). AOX accepts electrons from the Q-pool, bypassing the cytochrome pathway. Given the non-protonmotive nature of AOX no energy is conserved during this step which is reflected in a decreased ADP/O ratio. In a situation where reducing equivalents are donated to either complex II (or any of the alternative NAD(P)H dehydrogenases) and if the alternative pathway is the only available oxidising pathway, all energy freed by the oxidation reactions is dissipated as heat. Apart from higher plants, AOX is found in several other species which have branched respiratory pathways, such as various fungi (e.g. Neurospora crassa and Pichia anomala [48]) and protists (e.g. Trypanosoma brucei and Chlamydomonas reinhardtii [48]). Quite recently (2004) the occurrence of AOX encoding genes was found to extend into the animal kingdom as well. Sequences coding for AOX were found in the genomes of a mollusc (Crassostrea gigas), a nematode (Meloidogyne hapla) and in chordates (Ciona intestinalis and Ciona savignyi) [49]. The belief that AOX only occurs in eukaryotes has been challenged recently by reports on the occurrence of AOX in prokaryotes such as Novosphingobium aromaticivorans [50]. A recent search in a metagenomic dataset from
  39. 39. 25 marine microbes in the Sargasso Sea uncovered 69 different AOX genes [51] which indicates that AOX may be widespread in aquatic environments. Cyanide resistant respiration in higher plants has been reported since the early 1900’s [52] but only with the advent of antibodies raised against the partially purified alternative oxidase from Sauromatum guttatum [53] was it demonstrated positively that AOX was a genuine component of the ETC of cyanide-resistant mitochondria. The plant alternative oxidase is nuclear encoded and the first identified gene was named aox1 [54], importantly this gene encodes for a protein which includes a pre-sequence targeting it to mitochondria with an approximate weight of 39 kDa. Cleaving the target sequence yields a protein of approximately 32 kDa [55]. Several other genes (aox2a, aox2b) have been identified since [56]. In plants, AOX is found as a homodimeric enzyme whereas in fungi it is a monomer [57]. Apart from a structural difference both types of AOX are also very different with respect to activation mechanisms [58]. In this study a plant alternative oxidase from S. guttatum was heterologously expressed in S. pombe mitochondria [59], therefore the remainder of this section will focus on plant AOX and only significant differences between plant AOX and non-plant AOX will be discussed. Contrasting values for the plant AOX apparent KM for oxygen have been reported from ~1.7 M to 10-20 M [26]. In any case plant AOX affinity for oxygen is lower than that of complex IV (0.14 M [60]). The partitioning of reducing equivalents between the alternative and the cytochrome pathway will be discussed in-depth in chapter 5. Structure of AOX: Several models of the AOX structure have been proposed over the years [61]. In this section only the latest consensus model will be discussed. The current consensus model is the Andersson Nordlund (AN) model [62]. The alternative oxidase is believed to be an interfacial di-iron carboxylate protein [63] attached to the inner leaflet of the IMM facing the matrix space, see Figure 1.13. Getting structural information has been notoriously difficult and continuous efforts at spectroscopic detection were not successful until Berthold et al. in 2002 managed to get an EPR signal from a membrane fraction of E. coli expressing the Arabidopsis thaliana AOX
  40. 40. 26 [52]. Their results were the first experimental evidence supporting the hypothesis that AOX is indeed a member of the di-iron carboxylate proteins [64], a group of nonheme iron proteins that contain a coupled binuclear iron center. Figure 1.13 Structure of the plant AOX according to the AN model, adapted from Figure 1 in [63]. Regulation of AOX: Regulation of the alternative oxidase can be divided into two categories, regulation through expression and post-translational regulation. Regulation of the plant AOX is quite different from its counterpart in fungi. For this study a plant alternative oxidase (S. guttatum) was expressed in S. pombe, therefore in this section emphasis will be on plant AOX regulation. Regulation through expression: AOX expression can be increased in many ways. Stress conditions such as chilling, wounding, injury and osmotic stress are all known to increase expression [65]. In many fruits alternative pathway activity is known to increase during the ripening process. In
  41. 41. 27 mango fruit alternative pathway activity, amounts of protein and mRNA levels all increase in parallel [66]. In Hansenula anomala (now Pichia anomala) incubation with cytochrome pathway inhibitors, such as antimycin A or KCN, led to increased transcription of AOX, similar behaviour was seen in tobacco cells [66]. How inhibition of the cytochrome pathway is perceived by and transmitted to the nucleus to activate AOX expression is unclear. One suggested mechanism is via generation of ROS. In tobacco suspension cells it was found that upon addition of H2O2 within the span of two hours the level of AOX mRNA was increased [67]. In S. guttatum it was found that application of salicylic acid led to an increase in AOX mRNA levels [68]. AOX expression is also shown to be developmentally regulated [69]. Post-translational regulation: As mentioned previously, the plant AOX is a homodimer whereas the fungal AOX is a monomer. The plant AOX is subject to two interrelated post-translational mechanisms of regulation. In plants the alternative oxidase can be in an oxidised or a reduced form [61]. Plant AOX can be activated by reduction of a dimer-forming disulphide bridge. The reduced (active) form is a non-covalently linked dimer whereas the oxidised (inactive) form is covalently linked [70]. In transgenic tobacco plants expressing AOX it was found that certain TCA intermediates (citrate, isocitrate and malate) could reduce AOX [71]. It was hypothesized that the aforementioned intermediates may be involved in NADP reduction in plants and that NADPH mediates reduction of plant AOX in vivo. This could be a means of regulating AOX in response to changing matrix reduction levels. The second mechanism is direct activation of AOX by certain organic acids. A study by Millar et al. [72] showed that the plant AOX can be activated by a range of organic acids, most of them -keto acids: pyruvate, hydroxypyruvate, glyoxylate, -ketoglutarate, oxaloacetate, L-malate and succinate. It was determined that from these acids L-malate and succinate did not activate AOX directly. It was found that in the absence of malic enzyme (in SMP’s) succinate and malate could no longer activate AOX, which implies that it is in fact generation of pyruvate via malic enzyme which causes activation. It was concluded that this type of plant AOX activation is restricted to -keto acids. It was also determined that pyruvate activation is not
  42. 42. 28 dependent on pyruvate metabolism which implies that pyruvate has a direct effect on AOX [66]. As opposed to activation of AOX via pyruvate formation an alternative mechanism was hypothesised by Wagner et al. [73] where addition of succinate or malate led to changes in membrane fluidity which could facilitate the diffusion of QH2 from dehydrogenase to AOX. A substrate dependent difference in AOX activity is commonly seen where succinate dependent cyanide resistant respiratory rates are higher than NADH ones [8, 73-76]. This could be explained by a change in membrane fluidity, however it is more generally accepted that the higher cyanide resistant respiration rate with succinate is due to production of endogenous pyruvate [77]. The activating effect of pyruvate on AOX initially was assumed to change the affinity of AOX for QH2. In the absence of pyruvate, plant AOX is known to activate only at relatively high levels of Q-reduction (between 35- 50% reduced) [78, 79]. Addition of pyruvate was seen to reduce the threshold level of Q- pool reduction at which AOX becomes engaged [80] whilst pyruvate showed no effect on the redox status of the AOX protein disulfide bond. The two mechanisms are interrelated because pyruvate can only significantly activate AOX when the dimer is in the reduced form [80]. Conversely, if pyruvate is present, significant AOX activation can only be achieved when the dimer is reduced [81]. Interestingly enough many studies indicate that pyruvate can activate AOX activity with succinate as a substrate [79, 80] although it has been reported that succinate itself can activate AOX. This suggests that the amount of pyruvate generated indirectly from succinate is not sufficient to fully activate AOX and that further addition of pyruvate is required to fully activate AOX. It was concluded from experiments in which malic enzyme was inhibited that differences in the generation of intramitochondrial pyruvate can explain differences in AOX activity between tissues and substrates [77]. A pH effect on AOX mediated respiration is seen in certain plant species [82], for instance with external NADH as substrate S. guttatum mitochondria displayed a pH optimum for cyanide-resistant respiration [83]. Activation of fungal and protist AOX is quite different from their plant counterpart. AOX in fungi and protists are generally found as monomers and are not subject to organic acid stimulation [61]. Interestingly, in recent work where the protist AOX from Ciona
  43. 43. 29 intestinalis was expressed in human cells, pyruvate was found to activate the AOX protein [84]. Purine nucleotides, such as AMP, GMP and IMP are reported to have an activating effect on fungal and protist AOX [58]. Also an effect of pH on AOX activation in Acanthamoeba castellanii mitochondria was found [82]. Despite differences between plant and non-plant AOX at the level of regulation, monoclonal antibodies raised against Sauromatum guttatum AOX cross-react with fungal and protist AOX proteins Other factors regulating AOX are the amount of ubiquinone present [85], the Q-pool redox poise [86] and the amount of AOX protein present [85, 87]. AOX Function: The only function of AOX which has been commonly accepted is that of heat generation in order to volatilise odiferous compounds in order to attract insects during pollination in thermogenic plants [88]. Its function in non-thermogenic plants, let alone in non-plant species to date is still a matter of debate. Several, not mutually exclusive, hypotheses have been proposed. Continued turnover of the TCA cycle during any condition that inhibits or decreases the activity of the cytochrome pathway, such as under ADP limited conditions or during stress (wounding, chilling, drought etc.) will allow continuous production of biosynthetic precursors [89]. Another possible function of AOX is to scavenge harmful ROS produced under conditions (limited ADP, stress conditions) where components within the respiratory chain become highly reduced [65]; by keeping the ETC relatively oxidised AOX activity could prevent synthesis of ROS. Both hypotheses have in common that inhibition of the cytochrome pathway leads to activation of AOX. A more recent hypothesis suggests that AOX activity in plants serves as a means to keep plant growth relatively stable under variable environmental conditions [90]. AOX in relationship to UCP: Both AOX and UCP dissipate free energy as heat. It is interesting to note that some organisms express both enzymes in their mitochondria [91, 92]. Affourtit et al. raise the question as to what the physiological need could be for having two energy dissipating
  44. 44. 30 enzymes in one system [37]. It has been demonstrated in Acanthamoeba castellanii mitochondria that combined activity of both AOX and UCP leads to stronger reduction in ROS formation than with either of the complexes being active alone [91]. Furthermore it has been shown in plant mitochondria that activity of UCP appears to be coordinated with AOX activity [92]. These observations suggest that when both energy dissipating mechanisms are present in the same system their coordinated activities are involved in reducing ROS concentration. 1.3 Schizosaccharomyces pombe 1.3.1 General background: In this study S. pombe mitochondria were used as a model system to heterologously express a plant alternative oxidase [59] in order to investigate its respiratory characteristics within the respiratory chain. The same system has been used previously in our laboratory to investigate structure-function relationships [26, 45, 46, 59, 63, 70, 93, 94]. Yeast systems are a useful tool to investigate protein characteristics, it is relatively easy to express a foreign gene and within a short time a large amount of the protein of interest can be harvested. The yeast Schizosaccharomyces pombe also referred to as ‘the other yeast’ [95] is increasingly the preferred model system to investigate a wide range of processes such as the cell cycle [96], DNA repair [97], microtubule formation, meiotic differentiation, cellular morphogenesis and stress response mechanisms [98] over the traditionally used Saccharomyces cerevisiae (which recently has also been used to heterologously express a plant alternative oxidase [30]). S. pombe divides by fission and is one of the few free-living eukaryotic species whose genome has been completely sequenced [99] at the time of writing. The S. pombe genome is haploid and contains three chromosomes 13.8 Mb in size [98]. The mitochondrial genome is 20 kb in size [99]. It has been reported that S. pombe resembles mammalian cells more closely than does S. cerevisiae [100], for instance, S. pombe recognizes various mammalian promoters, splices mammalian introns and shares the same polyadenylation signals with mammalian cells, unlike S. cerevisiae [101]. It is also reported
  45. 45. 31 that S. pombe genes have longer upstream regions on average than those of S. cerevisiae which may mean that they are more complex and possibly more like those of higher eukaryotes [98]. S. pombe has proportionally more genes conserved in metazoans than does S. cerevisiae which is another argument in favour of the claim that S. pombe as an organism is more closely related to higher eukaryotes than S. cerevisiae. On the other hand each yeast shares genes with metazoans which the other lacks [102]. Furthermore a significant number of chromosome associated proteins are absent in S. cerevisiae but shared between S. pombe and metazoans making S. pombe the preferred system to study chromosome dynamics [102]. In 2002 a total of 172 S. pombe proteins were found to have similarities to human disease proteins whereas 182 such proteins were identified in S. cerevisiae. Most of the genes coding for these proteins are shared between the two yeasts [99]. Therefore both yeasts appear to be similarly useful as model organisms for the study of human disease gene function although given their different biologies one organism could be preferred for certain genes over the other and vice versa. Although S. pombe has been extensively used to investigate the cell cycle and genome repair mechanisms, in comparison to S. cerevisiae, relatively little work has been done on S. pombe metabolism [103] and even less has been done on the respiratory characteristics of its mitochondria [104, 105]. Recently however S. pombe mitochondria have been used to investigate several bioenergetic processes. S. pombe is the preferred system to investigate F1-ATPase (complex V) catalysis. The F1 part of complex V has several nuclear encoded subunits (the  and  units). S. cerevisiae cannot produce mutants of these subunits, leading to the production of “petite” colonies, i.e. cells with impeded oxidative phosphorylation. Hence F1 mutants cannot be studied in S. cerevisiae [106]. It has been a long held belief that S. pombe does not express a mitochondrial alcohol dehydrogenase (ADH), recent work done in this laboratory however indicates otherwise [26]. Mitochondrial ADH couples the oxidation of ethanol to the reduction of endogenous NAD+ to NADH, which can subsequently be oxidised by an internal NADH dehydrogenase, as happens in S. cerevisiae [107]. The presence of a gene encoding for ADH in S. pombe was confirmed as far back as 1983 [108], it was concluded that the protein was a cytosolic one [26]. The current hypothesis in this laboratory is that having both a cytosolic and a mitochondrial ADH can function as a means to equilibrate
  46. 46. 32 NAD+ /NADH ratios on both sides of the IMM in a way similar to the situation in S. cerevisiae [26]. The S. pombe gene SPAC5H10.06c was identified as a likely candidate encoding the mitochondrial ADH isozyme. Because of a recent discovery of a homolog of this protein in human liver [109] the discovery of a S. pombe mitochondrial ADH may have potential medical implications [26]. Another typical protein involved in bioenergetic processes is the adenylate kinase (AK) which catalyses the reaction: ATP + AMP  2 ADP [110]. In potato mitochondria its activity was shown to be responsible for the relatively high respiratory rate under ADP limited conditions [110]. Work done in this laboratory showed that continuous regeneration of ADP (from either endogenous or added nucleotides) led to constant activity of the ATP synthase affecting both membrane potential and oxygen consumption rate [38, 110, 111]. A gene coding for AK in S. pombe has been identified [112] and results suggested that the enzyme was found both in the cytosol and in the mitochondria. 1.3.2 The respiratory chain of S. pombe mitochondria: The respiratory chain of S. pombe mitochondria is rather similar to the mammalian one, see Figure 1.14: Figure 1.14 Schematic representation of the S. pombe ETC. See legends of figures 1.3 and 1.4.
  47. 47. 33 The S. pombe ETC, just like S. cerevisiae [27] does not contain complex I [13], therefore the only means of generating a pmf is via the cytochrome pathway. Work done in this laboratory [104] showed that isolated S. pombe mitochondria could respire on either succinate or NADH (in a rotenone-insensitive manner) indicating the presence of complex II and an external NADH dehydrogenase (which is nuclear encoded [113]) respectively. The aforementioned findings on the S. pombe mitochondrial ADH indicate the presence of an internal NADH dehydrogenase [26]. To the best of our knowledge the membrane potential across the IMM in isolated S. pombe mitochondria had not been measured prior to this study, but results by Moore et al. showed the occurrence of protein import into the matrix. This process could be abolished by addition of valinomycin which confirmed the presence of a membrane potential [104]. Respiration in S. pombe mitochondria can be completely inhibited by cytochrome pathway inhibitors which indicates the absence of an alternative oxidase3 [26, 104]. Unlike the yeast Hansenula anomala (now Pichia anomala) [116] AOX expression cannot be induced in S. pombe by incubation of the cells with antimycin A [59]. It has been proposed that cyanide resistant respiration (due to the presence of AOX) is found only in non-fermentative and Crabtree-negative yeasts (capable of fermentation but not under aerobic conditions) [117]. It was found, in general, that yeasts which do not display cyanide resistant respiration also do not express complex I in their ETC [118]. Non-fermentative yeasts under conditions where the cytochrome pathway is inhibited can still generate an electrochemical gradient of protons via complex I using the alternative oxidase as a terminal oxidase. The pmf generated could be utilised by complex V to drive the synthesis of ATP. It was hypothesized by Veiga et al. [117] that non-fermentative yeasts express both complex I and AOX as an alternative to cytochrome pathway respiration, whereas yeasts such as S. cerevisiae and S. pombe (which do not express complex I [13]) use fermentation as an alternative to cytochrome pathway respiration. In section 1.2.5 it was mentioned that in S. cerevisiae mitochondria the Q-pool displayed non-pool behaviour [27]. Given the similarities between the make up of the 3 Within older literature [114] but also in recent textbooks [115] (page 213) S. pombe mitochondria are reported to display cyanide resistant respiration, which is incorrect.
  48. 48. 34 respiratory chains of both yeasts this had implications for S. pombe. In this laboratory experiments were done, employing the same techniques which were used in the S. cerevisiae study and it was found that in S. pombe mitochondria the Q-pool does show pool behaviour [26]. Also, it was found previously that S. pombe mitochondria display antimycin resistant respiratory kinetics (see section 1.2.5) during NADH dependent respiration, cf. Figure 2A in [119]. It was reported that in addition to the respiratory components described so far S. pombe contains genes for several other respiratory linked proteins, namely: Gut2 encoding a glycerol-3-phosphate dehydrogenase, hmt2 encoding a sulphide dehydrogenase and ura3 encoding a dihydroorotate dehydrogenase. All three proteins can donate electrons to the Q- pool [26]. Upon performing a BLAST search4 (Basic Local Alignment Search Tool [120]) through the S. pombe genome another gene coding for a respiratory linked protein was found. SPAC20G8.04c codes for an electron transfer flavoprotein-ubiquinone oxidoreductase (ETF) which is a water-soluble matrix based complex that contains a FAD moiety and can accept electrons from several dehydrogenases containing flavin [1]. In this study S. pombe is used to functionally express AOX [59]. The alternative oxidase is non-protonmotive and its activity dissipates free energy. It was described in section 1.2.11 that AOX and UCP have the capacity to act in synergism. The presence of an uncoupling protein in yeast has been demonstrated [121]. It is therefore relevant to know whether or not S. pombe expresses an uncoupling protein. It was found by Stuart et al. [122] that the S. pombe genome did not contain any UCP homologs5 . However, at the time of that study (1999) the S. pombe genome was only partially sequenced (55%). At present the whole S. pombe genome is known [99] and a recent BLAST search did not reveal any UCP homologs. 1.3.3 SDH activation in S. pombe mitochondria: Comparable to plant mitochondria, activation of SDH in yeast requires the addition of ATP [123]. It is known that the mechanism of ATP activation is not due to unbinding of OAA 4 http://www.genedb.org/genedb/pombe/index.jsp 5 The same study also showed that the S. cerevisiae genome does not contain any UCP homologs.
  49. 49. 35 [124]. And addition of ATP to S. pombe mitochondria only partially activates SDH. For full activation the addition of glutamate (which leads to removal of OAA) is required [93]. Also an inhibitory effect of the uncoupler CCCP (which leads to dissipation of the pmf) on succinate dependent respiration in S. pombe mitochondria was found [26] in a way similar to plant succinate dependent respiration [21]. 1.4 Summary energy transducing systems To recapitulate; energy transducing systems can be defined in terms of the chemiosmotic theory put forward by Peter Mitchell [125]. An energy transducing system: 1) has a set of membrane located components which reversibly couples the translocation of protons to oxido- reduction reactions which generates an electrochemical potential of protons. (the ETC being an example of such a set of components). 2) has a membrane located ATP hydrolysing proton pump which can work in reverse driven by the aforementioned electrochemical potential of protons which leads to the catalysis of the thermodynamically unfavourable reaction of ATP synthesis. (complex V). 3) can use the aforementioned electrochemical potential of protons to directly or indirectly drive the transport of substrates across the membrane. (e.g. succinate via the dicarboxylate carrier [43] or ADP exchanged for ATP by the adenine nucleotide carrier (ANC) [126]). 4) the systems of postulates 1,2 and 3 are located in a specialised coupling membrane which has a low permeability to protons and to other ions in general. (e.g. the IMM).
  50. 50. 36 The chemiosmotic theory is generally accepted in the field of bioenergetics and it therefore is considered paradigmatic in this work. Although generally accepted, as recently as 2005, the theory is still questioned, see references [127-129]. 1.5 A modular representation: In order to study oxidative phosphorylation in this study we used a modular approach in which ETC components are lumped into Q-pool reducing pathways and Q-pool oxidising pathways [37], see Figure 1.15. The external NADH dehydrogenase and the combined activity of both SDH and the dicarboxylate carrier are the reducing pathways. The cytochrome pathway and the alternative pathway are the oxidising pathways. The Q-pool can be viewed as a reservoir which can accept electrons from the reducing pathways and can donate electrons to the oxidizing pathways. When all ubiquinone is reduced to ubiquinol the reservoir is ‘full’ and when all ubiquinol is completely oxidized it is ‘empty’. Under steady state conditions the rate with which electrons flow into the Q-pool equals the rate with which they leave. The overall electron flux through the respiratory chain can then be assessed by measuring the oxygen consumption rate. The steady state Q redox poise and oxygen consumption rate values are dependent on the interplay between the activities of the reducing and oxidising pathways. Activity of the cytochrome pathway leads to the formation of a pmf, due to the backpressure of this proton gradient the activity of this pathway can be limited. If now an uncoupler were added, the backpressure would be relieved and the activity of the cytochrome pathway increases, which is reflected in an increased steady state oxygen consumption rate with a concomitant oxidation of the Q pool. Some reducing pathways are not fully active upon addition of substrate, e.g. SDH becomes more active upon addition of ATP, which is reflected also in an increased steady state oxygen consumption rate, but in this situation the Q pool would become more reduced. Things are not as straightforward when a condition changes which affects reducing and oxidising pathway activities simultaneously, as will be discussed in chapter 3. But for now these examples illustrate clearly the general idea of the modular approach that is used in this study and which has been used successfully in previous studies [76, 93, 130, 131]. This approach has two main tenets:
  51. 51. 37 1) the Q-pool is homogenous, i.e. ubiquinone (ubiquinol) molecules can freely interact with different dehydrogenases and oxidases. 2) Reducing pathways only interact with oxidising pathways via the Q-pool as an intermediate, i.e. there is no direct interaction between these pathways. Figure 1.15 A modular representation of the ETC with the Q-pool as the central intermediate between pathways. Reducing pathways: external NADH dehydrogenase and the combined activities of SDH and the dicarboxylate carrier. Oxidising pathways: the cytochrome pathway (complexes III, IV and cytochrome c) and the alternative pathway consisting of the alternative oxidase only. A different modular approach called ‘top-down’ metabolic control analysis using p as the central intermediate has been successfully applied to both mammalian and plant mitochondria [132, 133]. In that approach energy transducing processes are classified as either p producers or p consumers. Three components are defined which communicate with one another via p: the ‘respiratory chain’ (dicarboxylate carrier + ETC), the ‘proton leak’ (proton leak, cation cycles etc.) and the ‘phosphorylating system’ (ATP synthase, the phosphate carrier and the adenine nucleotide carrier) [132]. The approach used in this study assumes the pmf to be constant [37].
  52. 52. 38 1.6 Summary Hopefully this introduction has managed to illustrate that mitochondria are a bit more than just little cellular batteries, they are indeed very important in regulating cellular physiology. Not only in plants or yeasts but as much in mammalian cells. Another trend of recent years, it appears, is the realisation that mitochondria are more and more involved in many human medical afflictions ranging from diabetes [134] to hearing disorders [135]. The alternative oxidase is quite often considered a typical plant protein and therefore according to current funding body standards maybe not so ‘fashionable’ however given the recent finding that AOX is also found in the animal kingdom and the fact that it has been expressed in human cells [84] could place the alternative oxidase back in the picture as a clinical tool for investigating human diseases. 1.7 Aims: To set up a three electrode system which allows for simultaneous determination of oxygen consumption rate (vO2), Q-redox poise (Qr/Qt) and membrane potential () in isolated mitochondria (chapter 3). This was achieved and successful recordings were made. SDH activation in potato mitochondria by ADP, ATP and oligomycin was hypothesized to occur indirectly using  as an intermediate. This was investigated (chapter 3) and it was determined that SDH activation did not occur indirectly via . S. pombe respiratory kinetics have previously been characterised in terms of oxygen consumption rate and Q-redox poise under various energetic conditions, but to the best of our knowledge the membrane potential had not yet been determined. A further characterization of the S. pombe respiratory kinetics was undertaken (chapter 4). This yielded some interesting oxidising pathway kinetics which have not been seen previously in mitochondria from other species.
  53. 53. 39 S. pombe has been used previously in this laboratory to heterologously express AOX. It was found then that AOX expression led to a change in respiratory kinetics under various energetic conditions. Kinetic curves were fitted to data obtained from malonate titrations done on mitochondria respiring on succinate, which has yielded a limited amount of data points. In this study a novel titration method (an NADH regenerating system) was used to obtain a larger dataset and a possible effect of AOX expression on  generation was studied (chapter 5). This approach demonstrated that S. pombe mitochondria expressing AOX display substrate dependent differences in oxidising pathway kinetics. Furthermore an effect of  on AOX activity could not be demonstrated.
  54. 54. 40 Chapter 2 Materials and Methods 2.1 Isolation and purification of mitochondria In this study mitochondria were isolated from (transformed) Schizosaccharomyces pombe cultures, Saccharomyces cerevisiae cultures, fresh potato tubers and Arum maculatum spadices. 2.1.1 Schizosaccharomyces pombe—The S. pombe strain used in this study is the so called sp.011, ade6-704, leu1-32, ura4-D18,h- [104]. From this strain three types of yeast cultures were grown, a wild-type (sp.011 wt) and two transformants. One type of transformant had the S. guttatum AOX expressed, depending on the presence of thiamine in the growth medium AOX was either expressed or repressed in the mitochondria (sp.011 AOX and sp.011 AOX+T respectively). Another type of transformant had an empty vector expressed (sp.011 pREP). Therefore a total of four different types of S. pombe mitochondria were investigated in this study. All media used for yeast transformation, yeast growth and yeast mitochondrial isolation are described in section 2.1.1.7. 2.1.1.1 The Expression system—Functional expression of a plant alternative oxidase in S. pombe was achieved by using a system developed by Albury et al. [59]. A Sauromatum guttatum cDNA clone (pAOSG81 [136]) which represents the nuclear gene aox1 [54] was cloned into the expression vector pREP [137] in which it is under the control of the nmt1 promoter6 . A transformed S. pombe culture will not express AOX when grown in the presence of thiamine. Apart from aox1 several other genes are present on the plasmid. The LEU2 gene (from S. cerevisiae) codes for a protein involved in leucine biosynthesis. Using a S. pombe strain in 6 no message in thiamine
  55. 55. 41 which the equivalent gene (leu1- ) is disrupted the plasmid becomes essential for growth in a medium lacking leucine. The ars1 gene (autonomously replicating sequence) is required for initiation of replication in yeast, whereas the ori gene is required for initiation of replication in bacteria. The AMP gene (resistance against amphicillin) provides a method to select for the plasmid in bacteria. When expressed in S. pombe (grown in the absence of thiamine and leucine) AOX is targeted to and incorporated into the IMM as a functional enzyme [59]. 2.1.1.2 Yeast transformation—S. pombe cells were transformed with pREP-AOX (coding for the S. guttatum AOX) or with pREP (just the vector) using a modified lithium acetate method [101]. A single sp.011 wt colony was inoculated in 5 ml YES medium and incubated overnight (150 rpm, 30 C) which normally grew to ~2.5x107 cells/ml. 800 l of this culture was used to inoculate 100 ml of minimal medium, which was grown overnight (150 rpm, 30 C). The cells were harvested by bench-top centrifugation (2 minutes at 3000 rpm at room temperature) and washed in distilled water. The cells were then resuspended in 0.1 M lithium acetate / Tris-EDTA (LiA/TE) in 10 0.1 ml aliquots at a density of ~1x109 cells/ml. The cells were then incubated at 30 C (waterbath) for one hour with occasional mixing. To each aliquot 1-2 g DNA (in ~10 l) and 290 l 50% polyethylene glycol (PEG) dissolved in LiA/TE was added. The cells were then again incubated for one hour in the waterbath at 30 C with occasional mixing. This was followed by a heat shock step of 15 minutes at 43 C (waterbath) for 15 minutes. Cells were then pulsed to a pellet and the supernatant removed. The pellet was gently resuspended in 100 l 0.1 M LiA/TE. The cells were then plated on minimal medium agar plates and grown for 3-5 days at 30 C. Colonies grown on these plates were subsequently used to inoculate larger cultures. 2.1.1.3 S. pombe growth—A starter culture was set up by picking a colony from a yeast plate and adding it to a 200 ml solution of minimal media. Supplements were added depending on yeast type. 0.4 mM adenine and 0.7 mM uracil were added to all cultures. In the case of sp.011 AOX+T 0.5 mM thiamine was added to repress expression of AOX. The wild type sp.011 WT requires addition of 1.1 mM leucine. A culture was incubated under
  56. 56. 42 aerobic conditions for three days at 30 C (150 rpm) for the cells to reach stationary phase. Cell concentration was assessed spectrophotometrically (light scattering at A595 [93]) using a 15-fold dilution of the cell culture in distilled water. This value was used to calculate the volume of culture required to inoculate each of four fresh 1 litre culture flasks (minimal medium with the appropriate supplements) to give cell concentrations ~ 40% of the stationary phase density at the anticipated time of harvesting. For this the following exponential growth equation was used: [2.1] Where  is the growth rate, N0 is the starting concentration, Nx the final concentration and t the period of growth. After rearranging this gives: [2.2] In this form N0 represents the starting cell density which gives the required Nx density after t time (hours) with cells growing at a rate of . In a previous study values of  of ~0.14 and 0.12 h-1 for non-transformed and transformed cells were determined [26]. In this study similar values were obtained. The inoculated 1 litre flasks were typically incubated for 19- 21 hours under aerobic conditions at 30 C and 150 rpm. t NNx    )log(log303.2 0  ) 303.2 (log 0 10 t Nx N    
  57. 57. 43 2.1.1.4 Isolation of mitochondria from S. pombe cultures—On the day before isolation four 1 litre flasks were sub-cultured by adding a certain volume of starter culture, the amount of which calculated as described in 2.1.1.3. The isolation of mitochondria from S. pombe cultures can be broadly divided into two stages. 1: Spinning down of the yeast cells from the four 1 litre cultures and treatment of the cells with lysing enzymes to remove the outer membranes and induce spheroplast formation. 2: Inducing spheroplast lysis by osmotic shock and subsequent harvesting of the mitochondria through differential centrifugation. The isolation protocol used here is based on the method of [104]. 2.1.1.5 Spheroplast preparation—Cells were harvested by centrifugation (10 minutes, 7000 rpm). Cells were washed by resuspension in distilled water at 4 C and again spun down (10 minutes, 7000 rpm). The wet weight was recorded (typically between 15-20 g). Cells were resuspended in 200 ml spheroplast buffer (SB) and incubated at 30 C, 150 rpm for 15 minutes in the presence of the cell wall-digesting enzyme preparation Zymolyase 20T7 (5 mg / g wet weight). Upon addition of a second preparation, ‘lysing enzyme’8 (15 mg / g wet weight) the suspension was incubated for a further 45 minutes at 30 C, 150 rpm. Spheroplast formation was subsequently assessed spectrophotometrically by diluting a suspension aliquot 100x in distilled water. The level of scattering (A800) due to cells and intact spheroplasts compared to cells not treated with digestive enzymes was used to indicate the proportion of osmotically sensitive spheroplasts in the sample. Also, a 5 l aliquot of cell suspension was osmotically shocked by addition of 5 l distilled water, the effect of which was observed using a light microscope. Due to removal of the cell wall this led to lysis. 2.1.1.6 Isolation of mitochondria—All procedures described from here on were performed on ice to minimize enzymatic activity. The spheroplast suspensions were diluted 2-fold in spheroplast wash (SW) and spun down for 10 minutes at 1600 rpm at 4 C. As a washing step the pellets were resuspended in SW and again spun down for 10 minutes at 1600 rpm at 4 C. Pellets were resuspended in ~2 ml mannitol wash (MW) and transferred to a glass 7 Seikagaku corporation, code number: 120491 8 Sigma, code number: L1412
  58. 58. 44 homogeniser. With two gentle strokes the resuspended pellets were homogenised, the total volume was subsequently increased to 600 ml with MW to lyse the spheroplasts (osmotic shock). Cell debris and unlysed cells were removed by centrifugating at 3500 rpm for 15 minutes at 4 C. The supernatant was subsequently centrifuged at 13000 rpm for 10 minutes at 4 C. Pellets highly enriched with mitochondria were pooled and centrifuged at 10000 rpm for 10 minutes at 4 C to yield a final mitochondrial pellet which was resuspended in a small volume of ~1-2 ml of MW and kept on ice throughout the remainder of the experimental day. 2.1.1.7 S. pombe media—The following media were used for the transformation of S. pombe cells, the growth of S. pombe cultures and the isolation of mitochondria from these cultures. YES medium (Yeast Extract with Supplements): amt component final conc 5 g/l yeast extract 0.5% w/v 30 g/l glucose 3.0% w/v Supplements: 225 mg/l adenine, histidine, leucine, uracil and lysine hydrochloride. Solid media (for plates) was made by adding 2% Difco Bacto Agar. Transformation media: 0.1 M LiA/TE : 0.1 M lithium acetate in Tris-HCl (10 mM, pH 7.6) and EDTA (1 mM). 50% PEG in 0.1 M LiA/TE made up fresh on the day of transformation, not sterilised.
  59. 59. 45 Minimal medium [138]: 0.11 M glucose 19 µM FeCl3.6H2O 93 mM NH4Cl 0.9 µM Na2MoO4.H2O 15 mM Na2HPO4 0.6 µM KI 15 mM KH-Phthalate 0.2 µM CuSO4.5H2O 5.2 mM MgCl2.6H2O 5 µM citric acid 0.1 mM CaCl2 1 µM Na pantothenate 14 mM KCl 80 µM nicotinic acid 0.3 mM Na2SO4 55 µM inositol 8.0 µM H3BO3 40 nM biotin 1.8 µM MnSO4.4H2O 1 mM NaOH 1.4 µM ZnSO4.7H2O Solid media (for plates) was made by adding 2% Difco Bacto Agar. Spheroplast buffer (pH 5.8) 1.35 M sorbitol 1 mM EGTA 10 mM Citrate/phosphate Citrate/phosphate: 100 mM citric acid and 100 mM Na2HPO4 pH 5.8 mixed in ratio ~50:200 Spheroplast wash (pH 6.8) 0.75 M sorbitol 0.4 M mannitol 10 mM MOPS
  60. 60. 46 Mannitol wash (pH 6.8) 0.65 M mannitol 2 mM EGTA 10 mM MOPS Yeast reaction medium (pH 6.8) 0.65 M mannitol 1 mM MgCl2 5 mM Na2HPO4 10 mM NaCl 20 mM MOPS 2.1.2 Saccharomyces cerevisiae—The S. cerevisiae strain used in this study was ordinary baker’s yeast ‘Carrs – breadmaker yeast’ purchased at a local supermarket. A small quantity of dried yeast was dissolved in distilled water and subsequently plated on YES medium (section 2.1.1.7) based agar plates to grow S. cerevisiae colonies. The same protocols used for growing S. pombe cultures and isolating S. pombe mitochondria were used with S. cerevisiae with some minor alterations. The starter culture was grown for one day only, as opposed to three days and in the degradation step only Zymolyase was used (5 mg / g wet weight), the lysing enzymes were omitted. For electrochemical experiments the yeast reaction medium was used (section 2.1.1.7). 2.1.3 Potato tuber—Fresh potato tubers were bought at a local supermarket. The protocol to isolate and purify mitochondria from this tissue is based on [139]. Media used in this protocol are described in section 2.1.3.2. All operations were performed on ice to minimize enzymatic activity.
  61. 61. 47 2.1.3.1 Isolation of mitochondria from potato tubers—Potatoes (~1.5 kg) were peeled thickly, cut into large chip-sized pieces and homogenised in grinding medium using a juice extractor (Moulinex type 140). The juice was collected directly in 1.3 liter of grinding medium (GM). The homogenate was then filtered through a moistened muslin (to remove large starch particles). The pH was adjusted to 7.4. Subsequently, three centrifugation steps were performed, all at 4 C. First the homogenate was centrifugated for 5 minutes at 1500 rpm (to completely get rid of starch). The supernatant was centrifugated for 10 minutes at 4000 rpm. The supernatant was then centrifugated for 15 minutes at 10000 rpm. The pellet was resuspended in 2-5 ml washing medium (WM). The suspension was transferred to two 50 ml centrifuge tubes and WM was added to fill the tubes. This was followed by another centrifugation step of 10000 rpm for 10 minutes at 4 C. The pellets were resuspended in a small volume (2-5 ml) of WM and pipetted on top of a self-forming PercollTM gradient in 25 ml of purification medium (PM). The tubes were centrifugated at 18000 rpm for 30 minutes at 4 C. Using a pastette the mitochondria were removed from the gradient and diluted in WM (at least 1:10). Purified mitochondria were pelleted by a centrifugation step of 10000 rpm for 10 minutes at 4 C. The mitochondrial pellet was then resuspended in a small amount of WM (1-2 ml) and kept on ice for the remainder of the experimental day. 2.1.3.2 Potato tuber media: Grinding medium (pH 7.4) 0.3 M mannitol 0.1% w/v BSA 40 mM MOPS 2 mM EDTA 0.6% w/v PVP 40 10 mM cysteine
  62. 62. 48 Washing medium (pH 7.4) Identical to grinding medium apart from the fact that cysteine is omitted. Purification medium (pH 7.4) 21% v/v PercollTM 0.3 M sucrose 5 mM MOPS 0.1% w/v BSA Potato reaction medium (pH 7.2) 0.3 M mannitol 1 mM MgCl2 5 mM K2HPO4 10 mM KCl 20 mM MOPS 2.1.4 Arum maculatum—The protocol to isolate and purify mitochondria from this tissue was based on [88]. Media used in this protocol are described in section 2.1.4.2. All operations were performed on ice to minimize enzymatic activity. 2.1.4.1 Isolation of mitochondria from Arum maculatum spadices—Spadices from local Sussex woods were isolated from the leaf tissue and chopped into small ~1 cm3 slices and added to ice-cold grinding medium (GM). The slices were homogenised in a WaringTM blender in 2x3 s bursts. The homogenate was filtered through a wetted muslin and centrifugated at 4000 rpm for 10 minutes at 4 C. The supernatant was then centrifugated at 10000 rpm for 10 minutes at 4 C. The pellet was resuspended in washing medium (WM) which was then centrifugated at 12000 rpm for 10 minutes at 4 C. The pellet was resuspended in a minimal amount of WM and loaded onto a 21% PercollTM self-forming
  63. 63. 49 gradient. The gradient was centrifugated at 14000 rpm for 30 minutes at 4 C. The mitochondrial band was removed using a pastette and transferred to WM. The mitochondrial suspension was then centrifugated at 12000 rpm for 10 minutes at 4 C. Mitochondria were gently resuspended in a small volume (1-2 ml) of WM and kept on ice during the remainder of the experimental day. Mitochondria were isolated either immediately after picking of the spadices or after storing the spadices overnight at 4 C. 2.1.4.2 Arum maculatum media: Grinding medium (pH 7.5) 0.3 M mannitol 0.2% w/v BSA 20 mM MOPS 2 mM EDTA 2 mM pyruvate 7 mM cysteine Washing medium (pH 7.5) Identical to grinding medium apart from the fact that cysteine is omitted. Purification medium (pH 7.5) 21% v/v PercollTM 0.3 M sucrose 5 mM MOPS 0.1% w/v BSA 2 mM pyruvate Reaction medium The same as for potato, see section 2.1.3.2.
  64. 64. 50 2.1.5 Specifics of plant mitochondrial isolation [38, 140]: Plant cells have a rigid cell wall which requires the use of shearing forces to disrupt, in order to liberate cytoplasmic organelles. This leads inevitably to the rupture of the cell vacuole thereby releasing harmful compounds, such as hydrolytic enzymes, phenolic compounds, tannins, alkaloids and terpenes, which can interact with the mitochondrial membranes. In order to minimise interaction of these compounds with the mitochondria several precautionary measures can be taken. Phenolic compounds and their oxidation products (quinones) being highly reactive can react strongly with mitochondrial membranes. Bovine serum albumin (BSA) is routinely used, not only to bind free fatty acids, but also because it can bind to quinones. Cysteine which is added to the grinding medium of potato and arum preparations is another protective agent preventing quinone interactions. Polyvinylpyrrolidone acts as a scavenger of phenols and tannins. pH is kept between 7.2-7.5 as alkaline pH will increase phenol autooxidation and acid pH will increase interaction between phenols and protein functional groups. 2.2 Polyacrylamide gel electrophoresis & Western analysis 2.2.1 SDS-PAGE—Proteins were separated using 1-D SDS PAGE. Mitochondrial samples (stored at –80 C) were defrosted on the day of electrophoresis. Mitochondrial protein (15 µg per lane) was separated on 0.75 mm thick 10% SDS-polyacrylamide gels according to the method of [141]. Electrophoresis was performed for ~ 1 hour at 150 V. Samples were run under non-reducing conditions through the omission of -mercaptoethanol in the gel. 2.2.2 Blotting to nitrocellulose—Separated proteins were transferred from SDS- polyacrylamide gels to nitrocellulose membranes using standard electrophoretic methods [142]. Transfer was carried out in ice cold transfer buffer (25 mM Tris-192 mM glycine, 10% v/v methanol, pH 8.8) for 1 hour at 100 V.
  65. 65. 51 2.2.3 Immuno-detection of proteins—After blotting, nitrocellulose membranes were washed in Tris-buffered saline (TBS; 140 mM NaCl, 20 mM Tris-HCl pH 7.6) and gently agitated overnight at 4 °C in blocking solution (BS; 2% w/v milk powder 3% w/v BSA and 0.1% v/v Tween 20 in TBS). This was followed by 6 x ~ 5 min washes in TBS, filters were incubated for 1 hour at room temperature in BS containing mouse monoclonal, anti-AOX (from Sauromatum guttatum) antibodies [53] (1:2000 dilution). Filters were washed in TBS, as before, and then incubated for 1 hour in BS containing a 1:1000 dilution of secondary antibody (linked to horseradish peroxidase). Antibodies were detected on light- sensitive film using an enhanced chemiluminescence kit (Amersham International plc). S. guttatum AOX antibodies were a gift from Dr. Tom Elthon (University of Nebraska). 2.3 Protein estimations Due to the time consuming nature of the isolation of mitochondria and subsequent experiments; protein estimations were normally done on a different day. Therefore isolated mitochondria were kept frozen at –80 C and defrosted on the day of protein estimation. Protein concentrations were determined using a bicinchoninic acid (BCA) assay [143] in the form of a kit (BCA, Pierce, Rockford, UK) with bovine serum albumin (BSA) as a standard. In this assay, a solution of protein is incubated with a solution containing cupric sulfate and BCA. The cupric ion (Cu2+ ) is reduced to the cuprous ion (Cu+ ) by proteins in an alkaline medium. This reaction is often referred to as the Biuret reaction. The cuprous ion then forms a purple-colored complex with BCA that strongly absorbs light at 562 nm. All samples were kept for 30 minutes at 37 C before determining absorbances in a spectrophotometer (CARY 400 Scan). A calibration curve (absorption vs. mg protein) was made using a series of BSA standards by diluting a stock solution of BSA of 2 mg/ml with distilled water, including one cuvette filled with distilled water as a blank. This was done in duplicate and the absorption values were measured in a spectrophotometer. The acquired values were plotted in Kaleidagraph (version 3.02) and fitted using a linear fit. The mitochondrial samples with unknown protein weight were diluted 20 and 50 times (both in duplicate) and the acquired absorbance values were inserted into the equation which was derived from fitting the calibration curve (also done in Kaleidagraph).

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