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Effects of the expression of alternative oxidase on
oxidising pathway kinetics in Schizosaccharomyces pombe
Alexander Frans Johan van Aken
Submitted for the degree of Doctor of Philosophy
University of Sussex
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.
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
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-
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.
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
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
22.214.171.124 The Expression system 40
126.96.36.199 Yeast transformation 41
188.8.131.52 S. pombe growth 41
184.108.40.206 Isolation of mitochondria from S. pombe cultures 43
220.127.116.11 Spheroplast preparation 43
18.104.22.168 Isolation of mitochondria 43
22.214.171.124 S. pombe media 44
2.1.2 Saccharomyces cerevisiae 46
2.1.3 Potato tuber 46
126.96.36.199 Isolation of mitochondria from potato tubers 47
188.8.131.52 Potato tuber media 47
2.1.4 Arum maculatum 48
184.108.40.206 Isolation of mitochondria from Arum maculatum spadices 48
220.127.116.11 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
2.4.1 The oxygen electrode 53
2.4.2 The Q-electrode 54
2.4.3 The TPP+
18.104.22.168 The TPP+
-electrode setup 59
22.214.171.124 Detection of [TPP+
126.96.36.199 Construction of the TPP+
-electrode membrane 60
188.8.131.52 Conditioning of the TPP+
184.108.40.206 Calibrating the TPP+
-electrode correction 63
-electrode sensitivity 63
220.127.116.11 Durability of TPP+
-electrode response time 64
2.4.4 Respiratory measurements 65
18.104.22.168 Preparation of respiratory effectors 65
22.214.171.124 Nomenclature 66
126.96.36.199 Basic bioenergetic parameters 67
188.8.131.52 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
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
3.2.5 Stimulation of SDH by oligomycin 91
3.2.6 Adenine nucleotides and oligomycin inhibit succinate dependent
respiration in potato mitochondria
3.3 DISCUSSION 99
Chapter 4 Respiratory characteristics of
Schizosaccharomyces pombe mitochondria
4.1 INTRODUCTION 112
4.2 RESULTS 113
184.108.40.206 Respiratory rates with different substrates 113
4.2.2 Schizosaccharomyces pombe - cytochrome pathway kinetics 122
220.127.116.11 The relationships of Qr/Qt vs. vO2 and ∆ vs. vO2 under ADP
limited conditions with NADH as a substrate
18.104.22.168 The relationships of Qr/Qt vs. vO2 and ∆ vs. vO2 under state 3
conditions with NADH as a substrate
22.214.171.124 The relationship between Qr/Qt vs. vO2 under uncoupled
conditions with NADH as a substrate
126.96.36.199 A comparison of NADH and succinate dependent Qr/Qt vs. vO2
and ∆ vs. vO2 relationships under ADP limited conditions
188.8.131.52 A comparison of NADH and succinate dependent Qr/Qt vs. vO2
relationships under state 3 conditions
184.108.40.206 A comparison of NADH and succinate dependent Qr/Qt vs. vO2
relationships under uncoupled conditions
4.2.3 Schizosaccharomyces pombe - reducing pathway kinetics 134
220.127.116.11 SDH reducing pathway kinetics in sp.011 wt mitochondria under
state 2 and uncoupled conditions
18.104.22.168 External NADH dehydrogenase reducing pathway kinetics in sp.011
wt mitochondria under state 2 and uncoupled conditions
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
5.2.2 Oxidising pathway kinetics with NADH as substrate 157
22.214.171.124 Comparing sp.011 pREP and wt cytochrome pathway kinetics with
NADH as substrate
126.96.36.199 Comparing sp.011 AOX and sp.011 AOX + T oxidising pathway
kinetics with NADH as a substrate
5.2.3 Oxidising pathway kinetics with succinate as substrate 170
188.8.131.52 Comparing sp.011 AOX and sp.011 AOX + T oxidising pathway
kinetics with succinate as substrate
5.2.4 Oxidising pathway kinetics in sp.011 AOX mitochondria 176
184.108.40.206 Comparing sp.011 AOX oxidising pathway kinetics with either
NADH or succinate as a substrate
5.2.5 Cytochrome pathway kinetics in sp.011 AOX+T mitochondria 179
220.127.116.11 Comparing sp.011 AOX + T cytochrome pathway kinetics with
either NADH or succinate as a substrate
5.2.6 Alternative pathway kinetics in sp.011 AOX mitochondria 181
18.104.22.168 Comparing sp.011 AOX alternative pathway kinetics with either
NADH or succinate as substrate
5.2.7 Oxidising pathway kinetics in Arum maculatum mitochondria 183
22.214.171.124 Investigating substrate dependent differences in oxidising pathway
kinetics in Arum maculatum mitochondria
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
5.3.3 Comparing sp.011 AOX and sp.011 AOX+T oxidising pathway
kinetics with NADH as a substrate
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
5.3.6 Why are the oxidising pathway kinetics obtained in this study different
from Affourtit’s study?
5.3.7 Are there any substrate dependent differences in sp.011 AOX oxidising
5.3.8 Are the substrate dependent differences a characteristic of the
5.3.9 Are the substrate dependent differences a characteristic of the
5.3.10 Are the substrate dependent differences a characteristic of the
expression system used?
5.3.11 Can the alternative pathway compete with the cytochrome pathway in
S. pombe mitochondria expressing AOX?
5.3.12 Does expression of the alternative oxidase lead to a change in Q-pool
5.4 CONCLUSION 199
Chapter 6 Modelling of oxidising pathway kinetics in
Schizosaccharomyces pombe mitochondria expressing the alternative
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
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?
7.1.2 Future work suggestions pertaining to the biphasic patterns in S. Pombe
cytochrome pathway kinetics
7.2 Functional expression of AOX in S. pombe mitochondria
yields substrate dependent differences in oxidising pathway
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
7.2.3 What causes the substrate dependent differences in oxidising pathway
kinetics in S. pombe mitochondria expressing AOX?
7.2.4 Are the substrate dependent differences in oxidising pathway kinetics 234
in S. pombe mitochondria expressing AOX due to dehydrogenase
7.2.5 Future work suggestions pertaining to the substrate dependent oxidising
pathway kinetics in S. pombe mitochondria expressing AOX
7.3 Conclusion 237
Appendix 1 238
Appendix 2A 241
Appendix 2B 242
proton electrochemical gradient
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
CCCP carbonyl cyanide m-chlorophenylhydrazone
standard redox potential
ETC electron transfer chain
G6PD Glucose-6-Phosphate dehydrogenase
F Faraday constant (9.65104
FAD flavin adenine dinucleotide
FMN flavin mono-nucleotide
ISP iron-sulfur protein
IMM inner mitochondrial membrane
IMS intermembrane space
KCN potassium cyanide
NADH nicotinamide adenine dinucleotide, reduced form
OG octyl gallate
OMM outer mitochondrial membrane
Pi inorganic phosphate
pmf protonmotive force
PmitoKATP plant mitochondrial K+
PMS phenazine methosulfate
Qr/Qt Q-redox poise
R gas constant (8.31 J mol-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)
UCP uncoupling protein
vO2 oxygen consumption rate
z valence number
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  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 . 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.
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+
exchanger active, neurons in the central nervous
system have to expend as much as 70% . 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 . Mitochondria do not just generate ATP at a constant rate, ATP synthesis
is tightly regulated and mitochondrial activity is highly flexible depending on energetic
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  and excitotoxicity , 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.
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
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
 so there is a need to
partition the energy released during breakdown of glucose. ATP, releasing 57 kJ mol-1
hydrolysis (under cellular conditions) is used predominantly. Some ATP is generated
through substrate level phosphorylation (about 5% ), 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.
It would be more correct to use the term ‘reducing equivalents’ as will be explained further in section
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:
R: gas constant (8.31 J mol-1
T: absolute temperature (K)
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 (9.65104
Protons in the matrix and the intermembrane space normally will be affected by both a
concentration gradient and an electrical gradient which gives:
This Gibbs energy difference is generally referred to as the proton electrochemical gradient:
And with the definition for pH (pH = - log [H+
]) the equation can be further simplified:
To facilitate comparison with redox potential differences in the electron transfer chain
(ETC) Mitchell  defined the term protonmotive force (p) which is:
FmVp H /)(
p is expressed in units of V and substituting values for R and T at 25 C the equation
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.
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 ). 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
. 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
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 . 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 .
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 .
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
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+
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 . 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  and rhein . Recently, complex I defects caused by pathogenic
mutations in mtDNA and nDNA have been linked to various neurodegenerative diseases
such as Parkinson’s disease . Defective complex I functioning leads to a decreased H+
and a concomitant decrease in ATP production whereas ROS formation is stimulated.
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 ) 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 126.96.36.199.
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 188.8.131.52). SQORs couple
the oxidation of succinate to fumarate to the reduction of quinone to quinol :
succinate fumarate + 2H+
quinone + 2H+
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 .
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 Three dimensional structure of the E. coli SDH taken from Figure 1C in . 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
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 . 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.
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 . Intracellular
oxidation of succinate is inhibited by fluoride .
Regulation of SDH:
SDH in isolated mitochondria is in a partially deactivated state due to bound OAA .
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 . SDH can be activated by ATP
and ADP . Chapter 3 deals with SDH activation and regulation of SDH is discussed
more in-depth in its introduction, see section 3.1.3.
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+
reduction to form ubiquinol
(QH2). Because of its long hydrocarbon side chain both Q and QH2 are highly hydrophobic
. Figure 1.7 shows the chemical structure of Q. The side chain can vary in length
depending on species, n = 10 in mammalian mitochondria , whereas plant mitochondria
contain a mixture of ubiquinone molecules with n = 9 and 10 . In yeast mitochondria n
= 6 . 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 . 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
and n = 9 in amoebal mitochondria.
This figure does not hold for all yeasts, e.g. Candida utilis mitochondria contain both UQ7 and UQ9 .
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.  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.  showed that S. cerevisiae mitochondria do show
homogenous pool behaviour (cf. Figure 3B in ). Several studies showed that AOX
(from either Candida albicans or Lycopersicon esculentum (tomato)) expressed in S.
cerevisiae mitochondria  and  respectively, could utilise QH2 as a substrate, which
implies Q-pool behaviour in S. cerevisiae.
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
. 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 
and in potato  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  (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
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 . 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 .
In order to understand the pathway of electron flow through the bc1 complex an
understanding of the Q-cycle  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 . The Q-pool in the IMM
exists in large molar excess over the bc1 complexes with a ratio of 21:1 .
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
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 .
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.
Figure 1.12 The Q-cycle in mitochondria, adapted from Figure 5.14 in . P: positive N: negative
See text for explanation.
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-
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 .
1.2.7 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
) . Cytochrome c is nuclear encoded and
has a molecular weight of about 13 kDa . 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 .
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) 
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+
. 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 . 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 .
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
unfavourable reaction of ATP synthesis  and complex V can be considered a p
consumer . 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 .
The ATP synthase is regulated by the natural inhibitor protein IF1 which binds to a -
subunit from the F1 complex . 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
 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 . 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  and yeast
mitochondria . Inhibitory proteins can cross-react with mitochondria from other sources
1.2.10 Uncoupling protein:
The uncoupling proteins (UCP) are a subfamily of the mitochondrial carriers (MC)  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 .
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 . 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].
1.2.11 Alternative oxidase (AOX):
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 . 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 .
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 ) and alkyl gallates (such as octyl gallate and dodecyl gallate
). 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 ) and protists (e.g. Trypanosoma brucei and
Chlamydomonas reinhardtii ).
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) . 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 . A recent search in a metagenomic dataset from
marine microbes in the Sargasso Sea uncovered 69 different AOX genes  which
indicates that AOX may be widespread in aquatic environments.
Cyanide resistant respiration in higher plants has been reported since the early 1900’s 
but only with the advent of antibodies raised against the partially purified alternative
oxidase from Sauromatum guttatum  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 , 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 . Several other genes (aox2a,
aox2b) have been identified since . In plants, AOX is found as a homodimeric enzyme
whereas in fungi it is a monomer . Apart from a structural difference both types of
AOX are also very different with respect to activation mechanisms . In this study a
plant alternative oxidase from S. guttatum was heterologously expressed in S. pombe
mitochondria , 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 . In any case plant AOX affinity for oxygen is lower than that of
complex IV (0.14 M ). 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 . In this
section only the latest consensus model will be discussed. The current consensus model is
the Andersson Nordlund (AN) model . The alternative oxidase is believed to be an
interfacial di-iron carboxylate protein  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
. Their results were the first experimental evidence supporting the hypothesis that AOX
is indeed a member of the di-iron carboxylate proteins , 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 .
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 . In many
fruits alternative pathway activity is known to increase during the ripening process. In
mango fruit alternative pathway activity, amounts of protein and mRNA levels all increase
in parallel . 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 . 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 . In S. guttatum it was found that application of salicylic acid led
to an increase in AOX mRNA levels . AOX expression is also shown to be
developmentally regulated .
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 .
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 . In transgenic tobacco plants expressing AOX it was found that
certain TCA intermediates (citrate, isocitrate and malate) could reduce AOX . 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.  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
dependent on pyruvate metabolism which implies that pyruvate has a direct effect on AOX
. As opposed to activation of AOX via pyruvate formation an alternative mechanism
was hypothesised by Wagner et al.  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 . 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  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 . Conversely, if pyruvate is present, significant AOX activation can only be
achieved when the dimer is reduced .
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 . A pH effect on AOX
mediated respiration is seen in certain plant species , for instance with external NADH
as substrate S. guttatum mitochondria displayed a pH optimum for cyanide-resistant
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 . Interestingly, in recent work where the protist AOX from Ciona
intestinalis was expressed in human cells, pyruvate was found to activate the AOX protein
. Purine nucleotides, such as AMP, GMP and IMP are reported to have an activating
effect on fungal and protist AOX . Also an effect of pH on AOX activation in
Acanthamoeba castellanii mitochondria was found . 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 , the Q-pool redox
poise  and the amount of AOX protein present [85, 87].
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 . 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 . 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 ; 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 .
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
enzymes in one system . 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 . Furthermore it
has been shown in plant mitochondria that activity of UCP appears to be coordinated with
AOX activity . 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  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’ 
is increasingly the preferred model system to investigate a wide range of processes such as
the cell cycle , DNA repair , microtubule formation, meiotic differentiation,
cellular morphogenesis and stress response mechanisms  over the traditionally used
Saccharomyces cerevisiae (which recently has also been used to heterologously express a
plant alternative oxidase ).
S. pombe divides by fission and is one of the few free-living eukaryotic species
whose genome has been completely sequenced  at the time of writing. The S. pombe
genome is haploid and contains three chromosomes 13.8 Mb in size . The
mitochondrial genome is 20 kb in size . It has been reported that S. pombe resembles
mammalian cells more closely than does S. cerevisiae , for instance, S. pombe
recognizes various mammalian promoters, splices mammalian introns and shares the same
polyadenylation signals with mammalian cells, unlike S. cerevisiae . It is also reported
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 . 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 . 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
. 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 . 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  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 . 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
. Mitochondrial ADH couples the oxidation of ethanol to the reduction of endogenous
to NADH, which can subsequently be oxidised by an internal NADH
dehydrogenase, as happens in S. cerevisiae . The presence of a gene encoding for
ADH in S. pombe was confirmed as far back as 1983 , it was concluded that the
protein was a cytosolic one . The current hypothesis in this laboratory is that having
both a cytosolic and a mitochondrial ADH can function as a means to equilibrate
/NADH ratios on both sides of the IMM in a way similar to the situation in S.
cerevisiae . 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  the discovery of a S. pombe mitochondrial ADH may have
potential medical implications .
Another typical protein involved in bioenergetic processes is the adenylate kinase
(AK) which catalyses the reaction: ATP + AMP 2 ADP . In potato mitochondria
its activity was shown to be responsible for the relatively high respiratory rate under ADP
limited conditions . 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  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.
The S. pombe ETC, just like S. cerevisiae  does not contain complex I , therefore
the only means of generating a pmf is via the cytochrome pathway. Work done in this
laboratory  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 ) respectively.
The aforementioned findings on the S. pombe mitochondrial ADH indicate the presence of
an internal NADH dehydrogenase .
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 .
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)  AOX expression cannot be induced in S.
pombe by incubation of the cells with antimycin A . 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) .
It was found, in general, that yeasts which do not display cyanide resistant respiration also
do not express complex I in their ETC . 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.  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 ) 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 . Given the similarities between the make up of the
Within older literature  but also in recent textbooks  (page 213) S. pombe mitochondria are
reported to display cyanide resistant respiration, which is incorrect.
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 . 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 .
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 . Upon performing a BLAST search4
(Basic Local Alignment Search Tool )
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 .
In this study S. pombe is used to functionally express AOX . 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 . It is therefore relevant to know
whether or not S. pombe expresses an uncoupling protein. It was found by Stuart et al.
 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  and a recent BLAST search did not reveal any
1.3.3 SDH activation in S. pombe mitochondria:
Comparable to plant mitochondria, activation of SDH in yeast requires the addition of ATP
. It is known that the mechanism of ATP activation is not due to unbinding of OAA
The same study also showed that the S. cerevisiae genome does not contain any UCP homologs.
. 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 . 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  in a way similar
to plant succinate dependent respiration .
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 .
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
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.
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  or ADP exchanged for ATP by the adenine nucleotide
carrier (ANC) ).
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
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 , 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:
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) . The approach used in this study
assumes the pmf to be constant .
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  to hearing disorders .
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  could place the alternative oxidase back in the picture as a
clinical tool for investigating human diseases.
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.
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.
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
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-
. 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 184.108.40.206.
220.127.116.11 The Expression system—Functional expression of a plant alternative oxidase in S.
pombe was achieved by using a system developed by Albury et al. . A Sauromatum
guttatum cDNA clone (pAOSG81 ) which represents the nuclear gene aox1  was
cloned into the expression vector pREP  in which it is under the control of the nmt1
. 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
no message in thiamine
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 .
18.104.22.168 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 . 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
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.
22.214.171.124 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
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 ) 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:
Where is the growth rate, N0 is the starting concentration, Nx the final concentration and
t the period of growth. After rearranging this gives:
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 . 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.
126.96.36.199 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 188.8.131.52. 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 .
184.108.40.206 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
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.
220.127.116.11 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
Seikagaku corporation, code number: 120491
Sigma, code number: L1412
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.
18.104.22.168 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
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.
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.
Minimal medium :
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
Spheroplast wash (pH 6.8)
0.75 M sorbitol
0.4 M mannitol
10 mM MOPS
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 22.214.171.124) 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 126.96.36.199).
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 . Media used in this
protocol are described in section 188.8.131.52. All operations were performed on ice to minimize
184.108.40.206 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
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.
220.127.116.11 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
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 . Media used in this protocol are described in section 18.104.22.168. All
operations were performed on ice to minimize enzymatic activity.
22.214.171.124 Isolation of mitochondria from Arum maculatum spadices—Spadices from local
Sussex woods were isolated from the leaf tissue and chopped into small ~1 cm3
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
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.
126.96.36.199 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
The same as for potato, see section 188.8.131.52.
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 . 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
. 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.
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  (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  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).