molecular identification and physiological ... of master of science ... discovered that the pool...
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Molecular Identification and Physiological Characterization of Alternative Oxidase
Gene Family Members in Nicotiana tabacum
by
Jia (Steven) Wang
A thesis submitted in conformity with the requirements for the Degree of Master of Science
Graduate Department of Cell & Systems Biology University of Toronto
© Copyright by Jia (Steven) Wang 2009
Molecular Identification and Physiological Characterization of Alternative Oxidase Gene Family Members in
Nicotiana tabacum Jia (Steven) Wang
Degree of Master of Science
Department of Cell & Systems Biology University of Toronto
2009
Abstract
Two projects were undertaken to study the non-energy conserving alternative pathway
present in the plant mitochondrial ETC. In the first project, a tobacco AOX2 gene was
cloned and characterized. AOX2 showed tissue specificity in expression and could not be
induced by common stresses. In the second project I carried out a physiological
characterization of transgenic tobacco plants with increased or decreased expression of
AOX1 subjected to cold stress. Under non-stress condition, a strong inverse relationship
between levels of AOX1 and levels of oxidative damage was observed, while after cold
treatment AOX1 transgenic lines and WT showed more complicated and differential
responses in aspects of oxidative damage and the capacity of antioxidant system. I also
discovered that the pool sizes of monosaccharides after temperature shift were proportional
to AOX1 levels. These results indicated that AOX1 might have crucial but complex
impacts on ROS balance and carbon metabolism during cold stress.
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Acknowledgements I would like to firstly thank my supervisor Prof. Greg Vanlerberghe for his excellent
supervision over my master project, seasoned guidance on my research work and
invaluable suggestion on my thesis writing. I also greatly appreciate the constant help from
my committee meeting members: Prof. Dan Riggs and Prof. Herbert Kronzucker.
I thank Dr. Sasan Amirsadeghi for generating AOX transgenic lines and providing
technical assistance in my research work. I also would like to thank Dr. Allison McDonald
for her preliminary work in cloning tobacco AOX2 gene.
I am grateful to Marina Cvetkovska and Melissa Cheung for their help during the course of
my project. I am also grateful to Nirusan Rajakulendran for his supporting work of my
project. I appreciate the kindly help from members of CSB department.
I acknowledge the financial support from the Natural Sciences and Engineering Research
Council of Canada.
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Table of Contents Abstract…………………………………………………………… ii
Acknowledgments…………………………………………...…....iii
List of Tables……………………………………………………..vii
List of Figures…………………………………………………....viii
List of Abbreviations………………………………………………x
Chapter 1: Introduction……………………………….................1 1.1 Mitochondrion and electron transport chain………………………..........1
1.2 Alternative oxidase (AOX)……………………………………………....5 1.2.1 Structure and classification of AOX…………………………………………....5
1.2.2 Function of AOX……………………………………………………………….7
1.3 Effect of low temperature on plants…………………………………….12
1.4 Reactive oxygen species (ROS)………………………………………...14 1.4.1 ROS identification and production…………………………………………… 14
1.4.2 Dual roles of ROS…………………………………………………………….. 17
1.4.3 Balance of ROS………………………………………………………………. 21 1.5 Background to project…………………………………………………..27
1.5.1 Cloning and characterization of tobacco AOX2 gene………………………... 27 1.5.2 Role of AOX in ROS balance and carbon metabolism under
cold stress…………………………………………………………………….. 32
Chapter 2: Materials and Methods…………………………….36 2.1 Cloning and characterization of tobacco AOX2 gene…………………. 36 2.1.1 Plant materials and growth conditions………………………………………...36
2.1.2 RNA extraction……………………………………………………………….. 36
2.1.3 Primer designing for 5’-RACE………………………………………………. 37
2.1.4 5’-RACE of AOX2 gene……………………………………………………….39
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2.1.5 Sequence analysis…………………………………………………………….. 39
2.1.6 Phylogenetic analysis…………………………………………………………. 40
2.1.7 RT-PCR assay………………………………………………………………… 40
2.1.8 Northern blot analysis………………………………………………………... 40
2.2 Role of AOX in ROS balance and carbon metabolism under
cold stress……………………………………………………………… 43 2.2.1 Generation of transgenic plants………………………………………………. 43
2.2.2 Plant materials and growth conditions……………………………………….. 44
2.2.3 Mitochondrial isolation……………………………………………………… 44
2.2.4 Western blot analysis of mitochondrial proteins……………………………....45
2.2.5 RNA extraction from polysaccharide-rich tissues…………………………….46
2.2.6 Northern blot analysis………………………………………………………… 47
2.2.7 ROS detection………………………………………………………………… 49
2.2.8 TBARS assay………………………………………………………………… 49
2.2.9 Enzyme assay………………………………………………………………… 50
2.2.10 Sugar assay………………………………………………………………….. 51
2.2.11 Statistical analysis…………………………………………………………... 54
Chapter 3: Results……………………………………………... 56 3.1 Cloning and characterization of tobacco AOX2 gene…………………. 56
3.1.1 Cloning of 5’-region of tobacco AOX2 gene…………………………………..56
3.1.2 AOX2 sequence was characterized by bioinformatic methods………………...58
3.1.3 Expression of AOX2 displayed tissue specificity……………………………...65
3.2 Role of AOX in ROS balance and carbon metabolism under
cold stress……………………………………………………………… 69 3.2.1 Transgenic tobacco plants with altered expression levels of AOX……………69
3.2.2 Change of oxidative damage after cold shift showed differential
patterns among different lines………………………………………………... 71
3.2.3 RI29 and two AOX-overexpressed mutants displayed higher
transcript levels of major antioxidant genes…………………………………...73
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3.2.4 The activity levels of ROS-scavenging enzymes partially
conformed to their transcript levels…………………………………………...83
3.2.5 Contents of soluble sugars were proportional to the AOX levels
after cold treatment……………………………………………………………85
Chapter 4: Discussion………………………………………….. 89 4.1 Cloning and characterization of tobacco AOX2 gene…………………..89
4.2 Role of AOX in ROS balance and carbon metabolism under
Cold stress……………………………………………………………...95 4.2.1 ROS gene network in response to different light intensities…………………..96
4.2.2 Role of AOX in ROS balance under cold stress………………………………98
4.2.3 “Threshold dose effect” of ROS signal in activating ROS-scavenging
system………………………………………………………………………...105
4.2.4 AOX is involved in inter-compartment signaling network…………………..110
4.2.5 AOX facilitates the accumulation of soluble sugars during
cold stress……………………………………………………………………..112
4.2.6 AOX plays crucial roles in both stress response and metabolic
homeostasis…………………………………………………………………..116
Reference……………………………………………………….119
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List of Tables 2.1 Primers used for 5’-RACE PCR and characterization of RACE products……….....38
2.2 Primer information for tobacco AOX1 and AOX2 genes……………………………42
2.3 Primer information for ROS-scavenging genes and AOX-related genes……………48
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List of Figures 1.1 Cartoon for respiratory metabolism in plants………………………………………..3
1.2 Generation of ROS by a series of reduction of oxygen…………………………… 15
1.3 Diagrammatic representation of a typical plant cell describing ROS-
related network…………………………………………………………………….. 24
1.4 AOX proteins detected by Western blot analysis………………………………… 28
1.5 Phylogenetic tree of AOX genes from various species……………………………. 31
2.1 Principle for the enzymatic cycling assay………………………………………… 55
3.1 DNA gels showing the products of 5’-RACE and the product
characterization…………………………………………………………………… 57
3.2 DNA sequence alignment between tobacco AOX1 and AOX2……………………..60
3.3 Protein sequence alignment between tobacco AOX1 and AOX2………………….62
3.4 Phylogenetic tree demonstrating the sequence homology between tobacco
AOX2 and other AOXs in different species………………………………………... 64
3.5 RT-PCR analysis on different tobacco tissues…………………………………….. 66
3.6 Northern blot analysis of tobacco AOX2 expression in different tissues………….. 67
3.7 Northern blot analysis of AOX2 expression in anther and ovary tissues
of WT and AOX1-silenced mutants……………………..………………………... 68
3.8 Northern and western blot analysis of AOX expression in WT and AOX
transgenic lines……………………………………………………………………..70
3.9 Lipid peroxidation in WT and transgenic plants before and after the
cold stress………………………………………………………………………….. 72
3.10 Northern blot analysis of H2O2-scavenging genes…………………………………76
3.11 Northern blot analysis of O2.--scavenging genes………………………………….. 78
3.12 Northern blot analysis of AOX-related genes……………………………………… 79
3.13 A representative ethidium bromide-stained RNA gel……………………………... 80
3.14 Relative transcript levels of ROS-scavenging genes and AOX-related
genes shown by the bar graphs with error bars………………………………….. 82
3.15 APX and SOD activities in WT and transgenic lines before and after
cold stress………………………………………………………………………….. 84
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3.16 Contents of soluble sugars and insoluble sugar in WT and AOX
transgenic lines……………………………………………………………………..86
3.17 Contents of soluble sugars and insoluble sugar after a long-term
cold stress………………………………………………………………………….. 88
4.1 “Threshold dose effect” model of ROS signal in activating defense
system during the abiotic stresses………………………………………………... 108
4.2 A working model describing the possible interrelationship between
alternative pathway and monosaccharide pool under the cold stress……………. 115
4.3 Model summarizing the relationship between AOX and ROS
balance/carbon metabolism during cold stress……………………………………118
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List of Abbreviations 6PG 6-phosphogluconate
ABA Abscisic acid
ADP Adenosine diphosphate
AOX Alternative oxidase
APX Ascorbate peroxidase
AsA Ascorbic acid
ATP Adenosine triphosphate
BLOSUM Blocks of amino acid substitution matrix
CaMV Cauliflower mosaic virus
cAPX Cytosolic APX
CAT Catalase
cDNA Complimentary DNA
CoA Coenzyme-A
COX6b Cytochrome c oxidase subunit 6b
Cu/ZnSOD Copper-zinc superoxide dismutase
Cys Cysteine
Cyt Cytochrome
DAB Diaminobenzidine
DEPC Diethyl pyrocarbonate
DMSO Dimethylsulfoxide
DNA Deoxyribonucleic acid
DTT Dithiothreitol
e- Electron
EST Expressed sequence tag
EtBr Ethidium bromide
ETC Electron transport chain
F6P Frucose-6-phosphate
FeSOD Iron superoxide dismutase
G6P Glucose-6-phosphate
G6PDH Glucose-6-phosphate dehydrogenase
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GHCL Guanidinium hydrochloride
GPx Glutathione peroxidase
GSP Gene specific primer
H+ Proton
H2O2 Hydrogen peroxide
HK Hexokinase
HR Hypersensitive response
IMM Inner mitochondrial membrane
INV Invertase
MAPK Mitogen-activated protein kinase
MDA Malondialdehyde
MFA Monofluoroacetic acid
mitoETC Mitochondrial electron transport chain
MnSOD Manganese superoxide dismutase
mROS Mitochondrial reactive oxygen species
MRR Mitochondrial retrograde regulation
MSO Murashige and Skoog
mTP Mitochondrial targeting peptide
NAD+ Nicotinamide adenine dinucleotide, oxidized form
NADH Nicotinamide adenine dinucleotide, reduced form
NADP+ Nicotinamide adenine dinucleotide phosphate, oxidized form
NADPH Nicotinamide adenine dinucleotide phosphate, reduced form
NBT Nitroblue tetrazolium
ND NAD(P)H dehydrogenase
NDex Alternate NAD(P)H dehydrogenase (external)
NDin Alternate NAD(P)H dehydrogenase (internal)
NO. Nitric oxide 1O2 Singlet oxygen
O2.- Superoxide
pAPX Plastidial APX
PCR Polymerase chain reaction
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PGI Phosphoglucose isomerase
PMF Proton motive force
PTOX Plastoquinol terminal oxidase
RACE Rapid amplification of cDNA ends
RNA Ribonucleic acid
RNAi RNA interference
ROS Reactive oxygen species
RT-PCR Reverse transcription PCR
RuBisCO Ribulose-1,5-bisphosphate carboxylase/oxygenase
RWC Relative water content
SDS-PAGE Sodium dodecyl sulfate polyacylamide gel electrophoresis
SHAM Salicylhydroxamic acid
SOD Superoxide dismutase
TBA Thiobarbituric acid
TBARS Thiobarbituric acid-reactive-substances
TCA Tricarboxylic acid
TF Transcription factor
UCP Uncoupling protein
UPM Universal primer mix
UTR Untranslated regions
WT Wild type
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Chapter 1
Introduction 1.1 Mitochondrion and electron transport chain
The mitochondrion is the main site responsible for energy production in both animals and
plants. Besides its important function in energy metabolism, the mitochondrion is also
believed to be widely involved in other physiological processes, such as production of
biosynthetic precursors, cellular redox balance, heat generation, regulation of second
messengers and programmed cell death (PCD) (Dmitry et al., 1997; Plaxton et al., 2006).
Increasing evidence from recent research revealed that respiratory metabolism plays
central roles in most of these processes (Plaxton et al., 2006). Respiratory metabolism is
basically comprised of three main pathways, namely glycolysis, the tricarboxylic acid
(TCA) cycle and the mitochondrial electron transport chain (mitoETC) (Fernie et al.,
2004) (Figure 1.1).
Light energy is harvested by chloroplasts and stored in the form of reduced carbohydrates
through photosynthesis (Plaxton et al., 2006). In glycolysis, carbohydrates are oxidized to
pyruvates via sequential reactions in the cytosol. The pyruvates are then transported into
mitochondria and converted into acetyl CoA by pyruvate dehydrogenase (Fernie et al.,
2004). Noticeably, carbohydrates are not the exclusive respiratory substrates. Malate,
formate, fatty acids and amino acids are all alternative substrates for respiration (Plaxton
et al., 2006). During some biotic or abiotic stresses when carbohydrate supply is limited
due to the decrease of capacity of photosynthesis, protein and lipid are broken down to
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amino acids and free fatty acids, which are further converted into acetyl CoA via
deamination/oxidation and β-oxidation, respectively (Dieuaide-Noubhani et al., 1997;
Brouquisse et al., 1998). The acetyl CoA generated from the above different sources is
thereafter oxidized to CO2 by a series of enzyme-catalyzed reactions (TCA cycle) in the
mitochondrial matrix (Figure 1.1), concomitant with the production of the reducing
equivalents (NADH/FADH2) (Fernie et al., 2004).
NADH/FADH2 generated from TCA cycle are then passed to mitoETC. The mitoETC,
located in the inner membrane of mitochondria, couples the respiratory electron transport
to the generation of proton motive force (PMF) across the inner membrane, which further
powers ATP synthesis (Fernie et al., 2004). Plant mitoETC is comprised of 5 basic
complexes (I to V), the same as is found in mammals, plus another four alternative
NADPH/NADH dehydrogenases and an additional terminal oxidase (AOX). In the basic
respiratory chain, NADH and FADH2 are oxidized by NADH dehydrogenase (complex I)
and succinate dehydrogenase (complex II) respectively, and the electrons are transferred
to ubiquinone pool, which are further passed through cytochrome c reductase (complex
III), cytochrome c and finally cytochrome c oxidase (complex IV) to reduce O2 to H2O.
During this process, protons are pumped from mitochondrial matrix to the intermembrane
space by complex I, III and IV to form PMF. The protons later flow back into the matrix
through the channel in ATP synthase (complex V) down the electro-chemical gradient,
by which the free energy generated is used to produce ATP (Fernie et al., 2004;
McDonald and Vanlerberghe, 2006b).
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Figure 1.1 Cartoon for respiratory metabolism in plants, which is comprised of glycolysis,TCA cycle and mitoETC. Non-energy conserving components in ETC (NDex, NDin, AOXand UCP) were shown by green color. Phosphorylating pathways were denoted by orangearrows; non-phosphorylating pathways were presented by green arrows. Complex I andcomplex III are the two major sites of ROS generation in mitochondria. IMS, inter-membrane space; IMM, inner mitochondrial membrane; TCA, tricarboxylic acid; CI,complex I (NADH dehydrogenase); CII, Complex II (succinate dehydrogenase), which isboth a TCA cycle enzyme and a mitoETC component; CIII, Complex III (cytochrome creductase); CIV, Complex IV (cytochrome c oxidase); CV, Complex V (ATP synthase);AOX, alternative oxidase; UQ, ubiquinone; NDex, Alternate NAD(P)H dehydrogenase(external); NDin, Alternate NAD(P)H dehydrogenase (internal); UCP, uncoupling protein.
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In addition to the energy-conserving ETC described above, plant mitoETC is also
integrated with several non-energy conserving pathways (Plaxton et al., 2006): (1)
Alternate NAD(P)H dehydrogenase (ND). NDs are located on both outer and inner
surfaces of inner membrane of mitochondria. Unlike the complex I (NADH
dehydrogenase), alternate NDs oxidize NAD(P)H without pumping protons into
intermembrane space and therefore no energy is conserved. Although the function of
NDs is far from being well understood, they may probably have an effect on the energy
conserving efficiency and redox balance in mitochondria (Rasmusson et al., 2004) (2)
Uncoupling protein (UCP). UCP in plants is homologous to thermogenin in animals. It
works as a proton transporter to dissipate the proton gradient from intermembrane space
to mitochondrial matrix, which bypasses ATP synthase and therefore uncouples electron
transport from ATP production (Plaxton et al., 2006). (3) Alternative oxidase (AOX).
AOX is a crucial component of the unique alternative pathway in plant mitochondria,
which is branching from ubiquinone (UQ) pool in ETC and transfers electrons to oxygen
without conservation of energy (Vanlerberghe et al., 1997a). In contrast to cytochrome
pathway, alternative pathway is resistant to cyanide and antimycin A (inhibitors of the
cytochrome pathway) but sensitive to substituted hydroxamic acids like
salicylhydroxamic acid (SHAM) (Schonbaum et al., 1971). Although these non-energy
conserving pathways seem to waste the energy, they do endow plants with metabolic
flexibility and help plants dampen the production of the toxic reactive oxygen species
(ROS) during adverse conditions by dissipating membrane potential (McDonald and
Vanlerberghe, 2006b). In the next part, AOX, which is our main object in this project,
will be discussed in more detail.
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1.2 Alternative oxidase (AOX)
1.2.1 Structure and classification of AOX
AOX, located on the matrix side of inner mitochondrial membrane (IMM), is proposed to
be a di-iron carboxylate protein containing four-helix bundle (Siedow et al., 1995). A
structural model of AOX proposed by Andersson et al. (1999) indicated that AOX is an
interfacial membrane protein with its N- and C-terminal hydrophilic regions exposed to
the mitochondrial matrix. It exists as either a covalently linked or non-covalently linked
dimer (Umbach et al., 1993) and the most N-terminal cysteine residue was proved by
site-directed mutagenesis to be responsible for the formation of the dimer through
disulfide-bond (Vanlerberghe et al., 1998). Further discussion regarding its crucial role in
the regulatory mechanism of AOX activity will be carried out in section 1.2.2 and 1.5.1.
AOX has been identified in many diverse species, including plants, protists, fungi and
more recently some animals (McDonald and Vanlerberghe, 2006a). In plants, AOX is
commonly encoded by a multi-gene family, the members of which can be generally
classified into two groups based on the sequence alignment and phylogenetic analysis
(Considine et al., 2002): AOX1 and AOX2. For example, in soybean there is one AOX1
gene and two AOX2 genes (AOX2a and AOX2b) (Patrick et al., 1997), while in
Arabidopsis four AOX1 genes (AOX1a, AOX1b, AOX1c, AOX1d) and one AOX2 gene
have been identified (Clifton et al., 2006).
Noticeably, while AOX1 gene family members are present in both monocot and eudicot
plants, AOX2 is only discovered in eudicot plants and absent in all monocot plants
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examined to date (Borecky et al., 2006; Costa et al., 2009). The AOX2 and AOX3 genes
identified in maize (Karpova et al., 2002) are actually AOX1a and AOX1b respectively
according to phylogenetic analysis. This distinct divergence of AOX2 across plant species
suggests that AOX2 may probably descend from AOX1 after the divergence of monocot
and eudicot plants and play a unique function in eudicot plant species (Considine et al.,
2002).
Despite their similar biochemical function in ETC, the expression patterns of AOX1 and
AOX2 are quite different. AOX1 gene is widely known for its induction by biotic or
abiotic stress stimuli such as pathogen attack, cold stress and chemical treatment
(Juszczuk et al., 2003). In contrast, AOX2 is usually expressed in certain tissues or
developmental stages and not affected by most stresses (Saisho et al., 2001; Considine et
al., 2002). For instance, Arabidopsis AOX2 showed a high expression level during seed
germination (Nakabayashi et al., 2005). In soybean, decrease of AOX2a expression was
paralleled by the increase of AOX2b expression during the development of cotyledons,
which suggested the complementary relationship between these two AOX2 genes
(McCabe et al., 1998). However, most stress conditions failed to induce the expression of
the AOX2 gene. The only two exceptions to date are AOX2 in Arabidopsis, which could
be induced by the treatments perturbing the chloroplast function, such as paraquat,
cysteine and norflurazon (Clifton et al., 2005) and AOX2b in cowpea, which could
respond to the osmotic stress (Costa et al., 2007). The existence of two AOX gene
subfamilies with different expression patterns raises an interesting question: what are the
specific functions of AOX1 and AOX2? One hypothesis is that AOX2 is needed for a
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generic, housekeeping function in respiration, while AOX1 is required for defense
response during stress conditions (Considine et al., 2002). However, this hypothesis is
still far from being confirmed.
1.2.2 Function of AOX
As a terminal oxidase, AOX transfers electrons from the ubiquinone pool directly to
oxygen, bypassing the last two proton-pumping sites (complexes III and IV). The energy
generated during the electron transfer through AOX is not conserved as PMF but
dissipated as heat (Vanlerberghe et al., 1997a). Therefore, AOX was proposed to play a
role in thermogenic respiration. In fact, the first confirmed function of AOX did relate to
thermogenesis in aroid (Meeuse, 1975) and some other species such as cycads (Skubatz
et al., 1993), where heat produced by respiration leads to the volatilization of the
aromatic compounds to attract pollinators. However, AOX was gradually found in more
and more non-thermogenic species (Kearns et al., 1992; Vanlerberghe et al., 1992b),
implying its functions in other physiological processes.
One of AOX functions proposed in the 1980s was “energy overflow” for Cyt pathway,
which was deduced from the observation that alternative pathway was not activated until
the degree of reduced ubiquinone reached to certain level and the Cyt pathway was
saturated with electrons (Lambers, 1982). Therefore, the alternative pathway was
proposed to consume excess carbohydrates when the supply of carbohydrates exceeds
what is required by respiration (Lambers, 1982). However, the biochemical regulation of
AOX discovered later suggested more complex functions of AOX rather than the simple
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“overflow” effect. As mentioned above, AOX in plants exists in either an oxidized form
or a reduced form, depending on the redox state in mitochondria. It has been proposed
that the AOX noncovalently linked dimer (reduced form) is more active compared with
the covalently linked dimer (oxidized form) (Umbach et al., 1993). Besides this redox
modification, studies showed that the reduced form of AOX could be further activated by
α-keto acid (e.g. pyruvate) through the interaction with a cysteine residue (Day et al.,
1994; Vanlerberghe et al., 1998). These biochemical controls enhance the affinity of
AOX for reduced ubiquinone and endow AOX with the ability to compete with Cyt
oxidase for electrons (Umbach et al., 1994). Therefore, it was believed that the non-
energy conserving nature of AOX together with these biochemical regulations of its
activity endowed plant respiration with more metabolic flexibility (Vanlerberghe et al.,
1998).
One proposed function of AOX during the disturbance of respiration process by the
adverse growth condition or stress is to modulate respiration to maintain the metabolic
and energetic homeostasis (Parsons et al., 1999; Jarmuszkiewicz et al., 2001; Moore et
al., 2002; Sieger et al., 2005; Fiorani et al., 2005). Under phosphate limitation, the Cyt
pathway, which is coupled with ATP generation, is inhibited because of the lack of
phosphate (Parsons et al., 1999; Plaxton, 2004), therefore impairing electron transport
and hindering the oxidation of NADH to NAD+. NAD+ is required for the continuation of
TCA cycle, which supplies carbon skeletons for biosynthesis (Millenaar et al., 2003). The
presence of the alternative pathway avoids the interruption of NADH oxidation and
electron transfer, therefore maintaining the function of TCA cycle and the supply of
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carbon intermediates for biosynthesis (Millenaar et al., 2003). In addition, AOX also
provides a flexible way for plants to regulate energetic poise. For instance, at the high
phosphate potential (high ATP/ADP ratio) or high NADPH status, alternative pathway is
activated to bring down the energy level (Jarmuszkiewicz et al., 2001). During nutrient
deficiency or low temperature conditions, plant growth, which is a highly energy-
consuming process, is optimized by the alternative pathway to maintain the homeostasis
of growth rate (Hanson et al., 2002; Sieger et al., 2005; Fiorani et al., 2005). Another
thing we should notice is that the alternative pathway is not simply an energy-wasting
pathway. Although the electron flow from reduced ubiquinone to AOX is not coupled
with ATP generation, the electron transfer in the upstream complex I produces PMF,
which means that when the Cyt pathway is inhibited, AOX supports ATP production,
albeit with low efficiency, by maintaining the electron flow to oxygen and allowing
energy production through respiration to some extent to sustain plant growth
(Vanlerberghe et al., 1997b).
Another putative function of AOX developed from its ability to maintain electron flow
during stress conditions is to prevent the production of reactive oxygen species (ROS) in
the mitoETC (Millenaar et al., 2003). ROS is generated from chloroplasts, mitochondria
and peroxisomes as normal products of metabolism and remains at basal level during the
normal condition (Suzuki et al., 2006). However, the level of cellular ROS could be
increased dramatically under various stress conditions such as temperature stress,
pathogen attack or nutrient deficiency (Dat et al., 2000), thus damaging the biomolecules
in cells and causing severe metabolic disorders (see below for details). In mitochondria,
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over-reduction of mitoETC components due to the increased metabolic activity or
decreased Cyt pathway capacity will increase the possibility of electron leakage from the
mitoETC and thus cause the formation of ROS (Navrot et al., 2007). Considering the
roles of AOX in maintaining electron transport and dissipating membrane potential, it
was hypothesized that AOX might help prevent the over-reduction of the mitoETC and
therefore dampen the ROS production (Purvis et al., 1993; Maxwell et al., 1999; Møller,
2001). In fact, AOX induction has been detected under various biotic and abiotic stresses,
most of which are concomitant with ROS accumulation (Juszczuk et al., 2003). This
correlation suggests that the alternative pathway may play certain roles in defense
responses against the oxidative damage. The function of AOX in dampening ROS
generation was further supported by the studies conducted on the mutants with altered
expression levels of AOX in tobacco and Arabidopsis (Maxwell et al., 1999; Yip et al.,
2001; Umbach et al., 2005; Giraud et al., 2008; Sugie et al., 2006). The transgenic
tobacco suspension cells with underexpressed AOX or overexpressed AOX showed a
higher or lower level of ROS compared with WT cells, respectively (Maxwell et al.,
1999). Besides the suspension cell system, similar results were also observed in whole
plants. In Arabidopsis plants treated with KCN, which is the inhibitor of Cyt pathway in
the mitoETC, an increased level of oxidative damage was observed in AOX anti-sense
line compared with WT (Umbach et al., 2005). These results strongly indicated the
crucial role of AOX in controlling ROS generation during adverse conditions.
It is also believed that AOX function is not only restricted in mitochondria but affects
other processes in other compartments based on the microarray studies (Umbach et al.,
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2005; Giraud et al., 2008). One well-studied example is the impact of AOX on
photosynthetic metabolism (Raghavendra et al., 2003; Yoshida et al., 2006 and 2007).
Photosynthesis in chloroplasts is widely involved in carbon metabolism, production of
reducing equivalents and energy balance. Therefore it was assumed that AOX might
influence the function of chloroplast (Raghavendra et al., 2003). Studies showed that the
inhibition of the alternative pathway in the leaves of drought-treated wheat caused the
over-reduction of photosystem II (PSII) (Bartoli et al., 2005) and in broad bean inhibition
of AOX lowered both of the photosynthetic rate and operating efficiency of photosystem
II (ΦII) (Yoshida et al., 2006). The possible reason for these phenomena is that the
alternative pathway in mitochondria can consume excess reducing power produced by
photosynthesis and therefore prevent ROS generation and photoinhibition in chloroplast
(Yoshida et al., 2006), which maintains the function of chloroplast especially under stress
condition.
In summary, the existence of the alternative pathway balances carbon metabolism and
electron transport, contributes to the modulation of both energy status and redox status in
plants especially during stress conditions and has crucial influence on other diverse
metabolic processes. Nonetheless, our understanding regarding its complex functions is
still far from complete.
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1.3 Effect of low temperature on plants
Unlike animals and other species, plants don’t have the capacity of locomotion to avoid
adverse environmental impacts. Thus, they face various environmental stresses during
growth, including biotic (virus, pathogen etc.) and abiotic ones (salt, heat, cold, light etc.).
Therefore, it is not surprising that plants have to develop more complicated mechanisms
to adapt to these stresses compared with other species.
Low temperature stress is one of the most typical and important abiotic stresses plants
encounter. In the world, two thirds of the agricultural industry is suffering from the great
loss in crop yield due to the low temperature (Beck et al., 2004). Therefore, study on
plant response to cold stress attracted more and more researchers and great efforts have
been made to improve the cold-resistant ability of plants. The concept “low temperature”
could be divided into two types: (1) chilling temperature (<20 ° C) (2) freezing
temperature (<0 ° C) (Chinnusamy et al., 2007). Under the cold stress, the cellular
homeostasis is disrupted, which leads to changes in various physiological processes,
including: (1) Modification of membrane lipid composition. Low temperature decreases
the fluidity of biomembranes. To maintain the fluidity and function of membranes, the
membrane lipid and fatty acid constituents during cold stress was rearranged through
either unsaturation of membrane lipid by fatty acid desaturases or the chain breakage and
shortening caused by ROS attack (Murata et al., 1997; Møller, 2006). (2) Reduction of
relative water content (RWC) in leaf. During cold stress loss of water happens in all
different species but more severely in cold-sensitive plants. The reduced water absorption
under low temperature condition is due to the decreased permeability and increased
- 12 -
viscosity of membranes (Lyons, 1973). In cold-sensitive plants, particularly, the transport
of water was also reduced during the exposure to the low temperature (Lyons, 1973). (3)
Accumulation of compatible solutes, such as sugars, amino acids, polyols and their
derivatives. These low-molecular-weight organic metabolites provide plants with
cryoprotection and osmoregulatory capacity (Kaplan et al., 2004). Particularly, the
soluble sugars, as the main substrates of metabolic and energy processes, are also
involved in reorganization of photosynthesis/respiration and formation of the resistance
of plant cell structure in response to the cold stress (Deryabin et al., 2005; Bogdanovic et
al., 2008). (4) Oxidative damage due to the formation of reactive oxygen species (ROS)
(e.g. 1O2, H2O2, O2.- and HO.) (Graham et al., 1982) and the concomitant induction of
ROS-detoxifying system (Kuk et al., 2003), which will be discussed in more detail later.
In the past decade, more and more evidence has indicated the importance of ROS
damaging effects on plants under cold stress (Prasad et al., 1994; Wise, 1995; Suzuki,
2006). In fact, one basic difference between chilling sensitive and chilling resistant plants
is the capacity of ROS-scavenging system (Walker et al., 1993; Hodges et al., 1997). In
chilling sensitive plants, ROS scavenging-system cannot handle the accumulation of ROS,
therefore leading to the chilling injury.
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1.4 Reactive oxygen species (ROS)
1.4.1 ROS identification and production
The term “reactive oxygen species” (ROS) is a collective one that includes not only the
oxygen-centered radicals like superoxide (O2.-) and hydroxyl radical (HO.), but also non-
radicals like hydrogen peroxide (H2O2) and singlet oxygen (1O2) (Mittler, 2002). These
highly active intermediates are produced during the process of O2 reduction in aerobic
organisms (Figure 1.2). ROS is well known as a toxic by-product of various cellular
metabolisms in aerobic organisms, which include photosynthesis in chloroplasts,
respiration in mitochondria and photorespiration in peroxisomes (Dat et al., 2000). Some
enzymes like NAD(P)H oxidases and peroxidases can also catalyze the production of
ROS (Gechev et al., 2006).
The chloroplast was believed to be the major source of ROS in plants (Asada, 2006),
especially when CO2 fixation is limited under certain stress conditions, such as cold,
drought, salt stress and combination of these stresses with high light stress (Mittler et al.,
2004). During these CO2-limiting conditions, alternative electron acceptors have to be
used to maintain the redox state of chloroplastic electron transport chain. The use of O2 as
the alternative acceptor will cause the generation of O2.- mainly from Fe-S centers of
photosystem I and reduced ferredoxin (Gechev et al., 2006). The O2.- generated will
further initiate the sequential reaction to produce other ROS such as H2O2 and OH. (Dat
et al., 2000). Nonetheless, the ability of O2 to accept electrons in this case helps reduce
the risk of over-reduction of the chloroplastic electron transport chain, during which the
activated singlet oxygen will be formed (Dat et al., 2000).
- 14 -
(H) O2.- e-
H+ H2O2 OH⋅e-
H2O e-
H+e-
H+ O2
Figure 1.2 Generation of ROS (superoxide [O2.-], hydrogen peroxide [H2O2] and hydroxyl
radical[OH.]) by sequential reduction of oxygen.
- 15 -
In the recent research, evidence has been accumulated that the electron transport chain of
mitochondrion is another main source of ROS in plant cell under stress conditions (Dat et
al., 2000). The major sites in the mitoETC responsible for ROS generation are complex I,
ubiquinone pool and complex III (Moller, 2001). Production of ROS in mitochondria is
usually due to the overreduction of electron transport chain (electron input exceeds ETC
capacity) when ETC is constrained or disrupted by stress (Moller, 2001). In comparison
with chloroplast, the production of ROS from mitochondrion is much lower, especially
during photoperiod when photosynthesis is active (Foyer and Noctor, 2003). However, in
the non-photosynthetic tissues or in the dark, mitochondria will make a major
contribution to ROS production (Puntarulo et al., 1988). In addition, it has been reported
that in maize the increase of H2O2 was independent of the light intensity during chilling
(Kingston-Smith et al., 1999) and in wheat mitochondrial proteins were more oxidized
than those from chloroplasts during abiotic stress (Bartoli et al., 2004). These
observations suggested that the abundant chloroplast-specific antioxidant system might
be able to minimize the effect of ROS produced from chloroplasts during stress
conditions (Maxwell et al., 1999). Therefore, ROS production from mitochondria should
not be underestimated.
Besides the above two ROS-generating sites, plants also have other sources which
contribute to ROS production. Oxygenation of ribulose-1,5-bisphosphate catalyzed by
Ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCO) causes the generation of
glycolates, which are transported from chloroplast to peroxisome. The later oxidation of
glycolates in the peroxisome yields H2O2 (Gechev et al., 2006). On the other hand, the
- 16 -
finding of plasmalemma-bound NAD(P)H oxidases and cell-wall peroxidases in plants,
which are responsible for active production of O2.- and H2O2 in the apoplast, revealed
another ROS-producing system (Sagi et al., 2006), in which these two enzymes allow
plants to regulate ROS-balance and ROS-related signaling network in response to certain
stresses more flexibly (Bailey-Serres et al., 2006).
1.4.2 Dual roles of ROS
As mentioned above, it is luxurious for plants to live in the optimal conditions at all times.
Exposure to adverse conditions is often associated with the accumulation of ROS, which
leads to oxidative damage. Increased level of ROS has been detected in plants under
stresses such as pathogen attack, high/low temperature exposure, high light treatment,
drought stress and heavy metal treatment (Dat et al., 2000). High level of ROS generated
by these stresses shows “phytotoxicity”. Its highly reactive property causes the oxidative
damage to a wide range of biomolecules (e.g. DNA, lipid and protein), which impairs the
integrity of cellular structure and the normal function of cells and may even lead to cell
death (Mittler, 2002). (1) ROS (particularly OH.) attack will cause various modification
of DNA, including oxidation of purines and pyrimidines, generation of alkali labile sites
and release of free bases (Mancini et al., 2006). These modifications will eventually lead
to DNA mutation, blocking of DNA replication or strand breaks. (2) ROS peroxidation
effect on polyunsaturated fatty acids in lipid results in lipid chain breakage and
shortening (Møller, 2006). Lipid peroxides can be generated directly by the combination
between polyunsaturated fatty acids (or their side chains) and 1O2 (Halliwell et al., 1993).
Alternatively, lipid peroxidation can also be initiated when HO. attacks membrane lipid
- 17 -
and abstracts hydrogen. H2O2 and O2.- are not able to initiate lipid peroxidation directly,
but they can promote lipid peroxidation by producing hydroxyl radical through transition-
metal irons reaction (Halliwell et al., 1993). (3) The damaging effect of ROS on protein
could be either direct or indirect (Møller, 2006). The direct effect is mainly due to the
chemical modification of certain amino acids (e.g. cysteine, proline and arginine) when
exposed to ROS. This change in amino acid property may affect the protein function (if
the protein is enzyme, it may lead to the inactivation of enzyme). The indirect effect is
related to the end product of lipid peroxidation, which can bind to the co-factor of
enzymes and inactivate the enzymes. Interestingly, Winger et al. (2005) has shown that
4-hydroxy-2-nonenal (HNE), one of the products of lipid peroxidation could inhibit the
activity of AOX, which suggests that during oxidative stress, AOX function may be
impaired by ROS indirectly.
Despite its destructive effects, ROS was also believed to act as important signaling
molecules (Dat et al., 2000; Gechev et al., 2006), which could be involved in regulation
of various biological processes such as development, plant stress response and
programmed cell death (Dat et al., 2000). For example, in root development, ROS was
shown to regulate the tip growth by triggering calcium-related signaling pathway through
the activation of the specific calcium channel (Foreman et al., 2003). During stress
conditions, the induced defense response system and enhanced protection were observed
in H2O2-treated plants or transgenic plants which displayed increased level of H2O2,
indicating the signaling role of H2O2 (Prasad et al., 1994; Wu et al., 1997; Gechev et al.,
2002). In addition to the direct evidence for the signaling role of ROS, the phenomenon
- 18 -
that tolerance to one oxidative stress endows the plants with a stronger ability to endure
another oxidative stress, which was known as “cross tolerance”, strongly suggests that
ROS may play the universal signaling role in response to various oxidative stresses
(Burke et al., 1985; Irigoyen et al., 1996; Dat et al., 2000).
One intriguing question concerning the signaling roles of ROS is how such simple
molecules precisely signal the various biological processes. Accumulated evidence in
recent researches indicated that the specificity of the ROS signal is achieved by different
mechanisms (Gechev et al., 2006). (1) Chemical property of ROS. Compared with other
ROS, H2O2 has longer half-life and crosses membranes more easily with the help of
peroxoporins (Bienert et al., 2006; Vranová et al., 2002). Therefore at the beginning most
of attentions were given to H2O2 to study its signaling function (Levine et al., 1994;
Prasad et al., 1994). However, observation from other studies found that H2O2 was not
the exclusive signal molecule in ROS family. In parsley O2.- rather than H2O2 can trigger
defense gene activation and phytoalexin accumulation (Jabs et al., 1997). More recently,
ROS-related microarray experiments were carried out by Gadjev et al. (2006), in which
they found that gene expressions were specifically regulated by different ROS (H2O2, O2.-
or 1O2). (2) Intensity of ROS signal. One good example for the dose effect of ROS is that
low level of ROS can cause the acclimation response to stress while high level of ROS
can initiate cell death or hypersensitive response (HR)-like symptom (Gechev et al., 2002
and 2006). However, how the dose of ROS affects signal transduction remains unknown.
(3) Sites of ROS generation. As mentioned above, ROS could be generated in different
intracellular compartments. It was believed that specific signaling components in a given
- 19 -
site might be involved in ROS signal transduction, which is known as “local detection
mechanism” (Rhoads et al., 2006). For instance, the accumulation of ROS in
mitochondria leads to the release of Cyt c to the cytosol, which will trigger programmed
cell death (Lam et al., 2001; Robson et al., 2002), while the ROS signal generated from
chloroplasts plays a central role in triggering the oxidative burst during ozone treatment
by activating membrane-associated NADPH oxidase (Joo et al., 2005). (4) Interaction
with other signaling molecules, such as NO., calcium and hormones. The combined
signal of ROS and NO. is involved in regulating hypersensitive disease resistance
response (Delledonne et al., 2001), while the interactions between ROS and calcium or
hormones like auxin and abscisic acid (ABA) are able to signal diverse plant
developmental processes, such as root hair growth, seed germination and root
gravitropism (Foreman et al., 2003; Kwak et al., 2003; Joo et al., 2001). All these
mechanisms contribute to the complexity of ROS signaling network and ensure the
specificity of ROS signal for regulating diverse biological processes. On the other hand,
although our understanding about ROS sensing is still far from complete, it was
suggested that ROS signal could be perceived by three modes: ROS receptors
(unidentified), activation of kinases (e.g. mitogen-activated protein kinase [MAPK]) and
redox-sensitive transcription factors (e.g. heat shock factor [HSF]), which may probably
be responsible for the communication between ROS and nucleus (Mittler et al., 2004;
Gechev et al., 2006) (Figure 1.3).
- 20 -
1.4.3 Balance of ROS
Considering the highly toxic property and important signaling role of ROS, it is no
wonder that there should exist an elaborate metabolic network which keeps ROS level
under strict control.
As described above, when the mitoETC is over-reduced under certain stress conditions
AOX is capable of avoiding the production of O2.- by maintaining electron flow to
oxygen, which we call “ROS avoidance mechanism”. Similarly, the other two non-energy
conserving bypasses in mitochondria: uncoupling protein and alternative NAD(P)H
dehydrogenase were also believed to dampen ROS generation by dissipating membrane
potential (McDonald and Vanlerberghe, 2006b). In addition, plastoquinol terminal
oxidase (PTOX), functionally analogous to AOX but located in plastid (Carol et al.,
2001), was believed to have the same function of dampening ROS generation as AOX
and work together with AOX in a coordinated manner to restrict the production of
cellular ROS level (Amirsadeghi et al., 2006). In addition to these molecular adaptations,
“ROS avoidance” could also be realized by physiological adaptation (e.g. C4 and CAM
metabolism) or anatomical adaptation (e.g. development of refracting epidermis and leaf
curling) (Mittler, 2001 and 2002; Mullineaux et al., 2002).
Besides the ROS avoidance mechanisms which reduce the generation of ROS, plants also
develop other strategies to scavenge the ROS which has been produced. One well-studied
system: antioxidant system, which contains both enzymatic components and non-
- 21 -
enzymatic components (Ajay et al., 2002), plays a crucial role in scavenging ROS in
plant cells.
Enzymatic components include superoxide dismutase (SOD), catalase (CAT) and various
peroxidase like ascorbate peroxidase (APX) and glutathione peroxidase (GPX). SODs,
which are the only known plant enzymes capable of removing O2.-, catalyze the
dismutation of O2.- to O2 and H2O2 (Gechev et al., 2006). Three SOD isozymes (MnSOD,
FeSOD and Cu/ZnSOD), which are classified by the metal co-factors involved in the
enzymes, have been widely found in different species (Arora et al., 2002). They were
typically localized in specific compartments: MnSOD (in mitochondrion and
peroxisome), FeSOD (in chloroplast) and Cu/ZnSOD (in cytosol). However, recent
studies also revealed their existence in other compartments, indicating their ubiquitous
involvement in different metabolic systems (Grene, 2002). The H2O2 produced from the
dismutation of O2.- is further detoxified to water and oxygen by APX, GPX and CAT,
which, again, are distributed in different compartments (Figure 1.3). Given the different
affinities of APX and CAT for H2O2, people classify them into two different groups:
APX is used for fine modulation of H2O2 for signaling purpose, while CAT is used for
removing excess H2O2 under severe stress (Mittler, 2002). The balance between SODs
and H2O2-scavenging enzymes is important for determining the homeostasis of O2.- and
H2O2, which was believed to help prevent the formation of highly toxic hydroxyl radicals
(Asada et al., 1987). Under stress conditions such as cold/heat, high light, salt or drought
that disrupt cellular redox homeostasis and enhance the production of ROS, most of these
- 22 -
ROS-scavenging enzymes will be induced to remove the excess ROS and help establish a
new ROS balance. (Mittler, 2002 and 2004; Rizhsky et al., 2002)
Besides these enzymatic components, non-enzymatic components like ascorbate, ß-
carotene, glutathione are also important in scavenging ROS (Noctor et al., 1998). On the
one hand, they act as ROS scavengers by donating electrons to oxygen radicals in both
non-enzymatic reactions and enzyme-catalyzed reactions (e.g. ascorbate-glutathione
cycle) (Grene, 2002). On the other hand, ascorbate and glutathione also act as potential
signals to regulate gene expression to respond to the stress conditions (Foyer et al., 2005).
The redox ratios of these antioxidants were believed to play crucial roles in the
modulation of ROS-scavenging system (Karpinski et al., 1997).
Research on transgenic plants with altered expression levels of antioxidant genes or
altered levels of ascorbic acid/glutathione further reveal the roles of antioxidant system
and the complex interrelationship between its different components. The transgenic
tobacco plants with overexpressed Cu/ZnSOD or GPX displayed stronger tolerance to
abiotic stresses (Ashima et al., 1993; Kazuya et al., 2004), which indicated the crucial
roles of antioxidant genes in scavenging ROS and preventing oxidative damage. However,
the study on loss-of-function antioxidant mutants bring us a much more complicated but
interesting story. In some cases, depression of antioxidant genes (e.g. APX, CAT and
Cu/ZnSOD) or antioxidant metabolites (e.g. ascorbic acid) increased the sensitivity of
transgenic plants to the adverse growth conditions (Orvar et al., 1997; Willekens et al.,
1997; Rizhsky et al., 2003; Conklin et al., 1996), while in other cases the plants with
- 23 -
Figure 1.3 Diagrammatic representation of a typical plant cell describing the cellular ROS-scavenging system and ROS-related signal for gene regulation. Chloroplast,mitochondrion and peroxisome are the major sites of ROS production. The major O2
.--scavenging enzymes (MnSOD, Cu/ZnSOD and FeSOD) and H2O2-scavenging enzymes(APX, GPX and CAT) are widely distributed in these compartments and also in cytosol,keeping ROS level under strict control. H2O2 was believed to be an important signalingmolecule in regulating gene expression through Mitogen-activated protein kinases(MAPK) kinase or ROS-responsive transcription factors (TF). cAPX, cytosolic APX; pAPX,plastidial APX.
- 24 -
underexpressed antioxidant genes paradoxically showed a stronger ability of tolerance to
stress conditions (Rizhsky et al., 2002; Miller et al., 2007). This phenomenon could be
explained by the up-regulation of other ROS-scavenging enzymes in these mutants,
which compensates for the function of the missing gene. The redundancy in ROS-
scavenging system guarantees the effective control of cellular ROS level. More
interestingly, recent studies in Arabidopsis reported that repression of APX isozyme
located in the cytosol or chloroplast triggered different response signals during stress
conditions (Miller et al., 2007), indicating that ROS-scavenging genes are not only
involved in removing excess ROS but also contribute to the regulation of plant signaling
network during stress conditions.
In addition to the aforementioned ROS avoidance and scavenging mechanism, more and
more evidence has implied the crucial role of soluble sugars in ROS balance under stress
conditions (Couee et al., 2006, Zhao et al., 2000). In vitro experiments indicated soluble
sugars might bind to ROS directly to reduce its damaging effects (Aver’yanov et al.,
1989). Furthermore, soluble sugars can feed the oxidative pentose-phosphate pathway,
which produces NADPH to help scavenge ROS (NADPH is a major cofactor in ROS-
scavenging pathways such as ascorbate-glutathione cycle) (Couee et al., 2006). Although
the experiments studying the exogenous sugar treatment showed the induction of
antioxidant genes by soluble sugars, it is still disputed because these experiments cannot
distinguish between induction of stress defense mechanisms by sugars and induction of
mechanisms against sugar-induced stress (Couee et al., 2006). On the other hand, soluble
sugars play a converse role in ROS balance in other processes: as energy providers,
- 25 -
soluble sugars are involved in ROS-producing metabolic pathway like mitochondrial
respiration and in photosynthetic system increased soluble sugars can negatively regulate
some Calvin cycle genes (Pego et al., 2000; Rolland et al., 2002), which may cause
excessive electron transfer, resulting in the production of ROS (Couee et al., 2006). The
dual roles of soluble sugars in ROS balance make it complicated to analyze their
functions in plant defense response.
In summary, plants have several mechanisms with different levels to respond to the
change of cellular ROS. To reduce the production of ROS, plants develop AOX in
mitochondria and PTOX in plastids, which are capable to dispose of excess electrons
during mitochondrial or chloroplastic electron transport respectively and therefore reduce
ROS generation. To remove the ROS which has been produced, plants have evolved a
very elaborate antioxidant system which contains antioxidant enzymes and antioxidant
metabolites. Besides, some other factors such as soluble sugars may also contribute to the
regulation of cellular ROS balance. In summary, these mechanisms together with ROS-
producing system (see section 1.4.1) compose a complicated reactive oxygen network
and work together to maintain ROS homeostasis.
- 26 -
1.5 Introduction to project
1.5.1 Cloning and characterization of tobacco AOX2 gene
In tobacco, which is the main research species in our lab, two AOX1 genes (AOX1a
[S71335], AOX1b [X79768]) and a partial sequence (~200bp) of a putative AOX2 gene
have been revealed by PCR techniques (Vanlerberghe et al., 1994; Whelan et al., 1995;
Norman et al., 2004). But to date, only the tobacco AOX1 gene has been widely
characterized in gene regulation and physiological function (Vanlerberghe et al., 1992b
and 1998; Robson et al., 2002; Sieger et al., 2005; Amirsadeghi et al., 2006).
Although little is known about other AOX genes in tobacco, the results from our several
previous experiments did suggest their/its existence and intriguing features. (1) In the
western blot analysis with monoclonal antibody recognizing AOX (Elthon et al., 1989),
AOX protein was almost undetectable in the leaf tissues of AOX1-silenced transgenic
plant RI9. However, we did detect a very strong expression of AOX gene in RI9 root
(Figure 1.4 A), indicating that another AOX gene might be expressed specifically in root
tissues which could not be silenced by AOX1 RNA interference. (2) In another
experiment studying the role of the critical cysteine residue in the formation of disulfide
bond between two AOX proteins (see above), all protein samples were pre-treated with
oxidant diamide and run on either a reducing gel (+DTT) or a non-reducing gel (-DTT)
(Dithiothreitol [DTT] is a sulfhydryl reductant which can break the disulfide bond
[Umbach et al., 1993]) (Figure 1.4 B). In the non-reducing gel, AOXs with the critical
cysteine were dimerized by diamide treatment and located at 70kD. In contrast, AOXs
- 27 -
A
B
Figure 1.4 (A) Western blot showing AOX protein levels in leaf and root tissues of WT andAOX1-silenced mutant RI9. In the leaves only tiny amount of AOX protein could bedetected in RI9 compared with WT. However, a very strong signal of AOX protein wasdetected in RI9 root. (B) Another experiment studying the role of cysteine residue in AOXin the formation of disulfide bond between two AOX proteins. In AOX1, Cys-126 wasproved to be responsible for the formation of disulfide bond between two AOX proteins.B9 is an overexpressor of native AOX1 gene as a negative control; C12 and F6 are twooverexpressors of mutated AOX1 in which Cys-126 was replaced by alanine. They servedas positive controls. MFA is an inhibitor of TCA cycle (Vanlerberghe et al., 1996) that caninduce ROS production and AOX expression. The sizes of AOX monomer and dimer are35kD and 70kD, respectively.
- 28 -
without the critical cysteine could not form the disulfide bond and therefore was
supposed to migrate faster than the covalently-linked dimer and located at 35kD.
Surprisingly, we found that part of the AOX proteins from monofluoroacetic acid
(MFA)-treated tobacco cell could not form covalently-linked dimer through S-S bond in
the non-reducing gel, which is different from AOX1 protein (Figure 1.4 B). This result
implied that there might be another AOX in tobacco which doesn’t contain the critical
regulatory cysteine residue. (3) Based on the phylogenetic analysis of most known AOX
genes, we found that two Cys-absent AOX genes identified from tomato and potato were
both located in the AOX2 clade of the phylogenetic tree (Figure 1.5). Considering these
two species are close relatives of tobacco (Kawagoe et al., 1991), we postulated that
tobacco AOX2 might also be a Cys-absent AOX.
Taken together, these aforementioned results suggested that an unknown AOX (probably
AOX2) expressed in tobacco root tissue and MFA-treated suspension cells might be a
Cys-absent AOX. If this is the case, one intriguing question we’d like to ask is that what
is the regulatory mechanism and physiological functions for this special AOX? We
believe that the naturally-existing Cys-absent AOX gene will provide an important hint
for the regulatory mechanism of AOX protein activity in nature and further reveal the
physiological function of AOX gene family.
To better understand the interrelationship between different AOX subfamilies in tobacco
and to learn more about the role of the putative Cys-absent AOX in plant metabolism, we
decided to clone the tobacco AOX2 gene. In the previous work of our lab, a 200 bp
- 29 -
Figure 1.5 A phylogenetic tree of AOX genes from various species constructed by Dr.Allison McDonald (unpublished). The most N-terminal cysteine residue existing in most ofAOX proteins was supposed to play important roles in redox modulation and pyruvateactivation of AOX activity. However, for some AOX genes identified so far, this criticalregulatory cysteine was substituted by serine (boxed in black color), which might changethe biochemical property of AOX. The so-called ‘serine-substituted” AOX were distributedin both AOX1 and AOX2 gene subfamilies. Particularly, one tomato AOX and one potatoAOX with the cysteine substituted by serine were located in AOX2 clade. The number ateach branching site stands for the frequency of reproduction in 100 bootstrap replicates.
- 30 -
AOX2
AOX1
- 31 -
fragment of putative AOX2 cDNA sequence was amplified with gene specific primers
(Norman et al., 2004), which was used to design the primers for 3’-RACE PCR. An 828
bp fragment was obtained, including 475 bp 3’-coding region and 273 bp 3’-untranslated
region (3’-UTR). In my first project, the full-length coding sequence and 5’-UTR of
tobacco AOX2 were cloned with RACE PCR and several critical structures and motifs
were identified in deduced AOX2 protein sequence with bioinformatic tools. Its relation
to other AOX genes in various plant species was characterized by phylogenetic analysis.
Furthermore, RT-PCR and northern blot were performed to investigate the expression
pattern of AOX2 in different tobacco tissues.
1.5.2 Role of AOX in ROS balance and carbon metabolism during cold stress
Exposure to cold stress will lead to the ROS production in plants, which contributes
largely to chilling damage (Suzuki et al., 2006). The elevated level of ROS in
mitochondria is mainly due to the reduced capacity of Cyt pathway by cold stress and the
resulting over-reduction of the mitoETC (McNulty et al., 1987; Collier et al., 1990;
Covey-Crump et al., 2007). In contrast, the capacity of alternative pathway is increased
during cold treatment, which was proposed to help maintain the electron flow to O2 and
prevent ROS accumulation (Vanlerberghe et al., 1992a; Yusuke et al., 1997; Gonzalez-
Meler et al., 1999; Fagoni et al., 2003). The researches on the transgenic plants with
altered AOX levels generally supported this idea: overexpression of wheat AOX in
Arabidopsis reduced the level of ROS in plants under cold treatment (Sugie et al., 2006)
and in AOX anti-sense transgenic Arabidopsis the level of lipid peroxidation (an index of
oxidative damage) was a bit higher than in wild type under long-term cold stress (Fiorani
- 32 -
et al., 2005). These results indicated the role of AOX in dampening ROS production
during cold stress. More interestingly, another study on comparing the different response
to cold/freezing stress between freezing-sensitive and freezing-tolerant wheat found that
the tolerant cultivar showed a higher level of AOX after the cold shift than the sensitive
one (Nobuyuki et al., 2007), which suggests the important function of AOX in the
acquirement of plant cold/freezing tolerance.
Plant mitochondria play important roles in sensing and responding to stresses (Butow et
al., 2004). It was believed that mitochondria which were functionally altered could send
certain signals to the nucleus to modulate the expression of genes, which is known as
mitochondrial retrograde regulation (MRR) (Rhoads and Vanlerberghe, 2004). Although
the mechanism of MRR is relatively poorly understood compared with chloroplastic
retrograde regulation, several possible signaling pathways have been proposed (Rhoads et
al., 2007), in which mROS was believed to play crucial roles. It may act as a signal or
part of the signaling pathway by changing the redox status or producing certain
secondary signals (local detection mechanism) during stresses (Rhoads et al., 2006;
Amirsadeghi et al., 2007). This hypothesis was supported by the fact that in tobacco
suspension cells treated with antimycin A which caused the ROS generation from
mitochondria, adding of antioxidants or inhibition of ROS-transition pores could
dramatically reduce the gene induction in the nucleus (Maxwell et al., 1999 and 2002). In
addition, mROS may also contribute to the signaling network between mitochondria and
other compartments such as chloroplasts (Rhoads et al., 2006).
- 33 -
Considering both the damaging effect and signaling role of mROS and the ROS-avoiding
function of AOX in mitochondria, we supposed that AOX might play crucial roles not
only in protecting plants from oxidative damage but also in modulating the ROS
signaling network under oxidative stress, in particular, cold stress. Interestingly, in
salicylic acid-treated tobacco suspension cells (Amirsadeghi et al., 2006) and in
Arabidopsis treated with combined drought and light stress (Estelle et al., 2008), higher
capacity of antioxidant system was observed in AOX anti-sense mutant than in WT,
indicating the complementary relationship between AOX and antioxidant system and also
suggesting the existence of inter-compartment communication in ROS-controlling system.
However, our understanding regarding the interrelationship between AOX and
antioxidant system in ROS balance during cold treatment is still quite ambiguous.
Besides the interaction between AOX and the ROS gene network during cold stress,
another objective of this project is to understand the role of AOX in carbon metabolism
under cold stress. In Sieger et al. (2005) the carbon use efficiency of wild-type tobacco
suspension cells was found to decrease dramatically under nutrient limitation, while in
AOX-silenced mutant it did not change. In addition, a nice negative correlation between
respiration rate and amount of carbohydrates in cells was observed. These results
suggested that AOX could regulate the balance between respiration rate and the supply of
carbohydrate. As discussed above, during stress conditions AOX was also believed to
play essential roles in maintaining the function of photosynthesis (Yoshida et al., 2006),
which is the main process responsible for the production of carbohydrates. Therefore,
considering the influence of AOX on both downstream respiratory system and upstream
- 34 -
photosynthetic process, we believe that there may be a certain interrelationship between
the alternative pathway and carbon metabolism during cold stress, which, however, is
largely unknown. The important roles of soluble sugars in ROS balance and cold
tolerance (see above) add more interests to the research concerning the influence of AOX
on carbon metabolism in cold-treated plants.
In this part of project, I hope to reveal the relationship between AOX and ROS balance,
AOX and carbon metabolism under cold stress. The cold stress we applied on tobacco
plants was a low temperature above freezing point (chilling), which mimicked the early
response of plants to cold in fall. We believe that this experimental system might help us
understand the mechanism behind the plant chilling acclimation, which involves various
physiological and biochemical adjustments (Hughes et al., 1996; Francois, 2007). Our lab
has generated AOX1-silenced transgenic tobacco lines (RI9 and RI29) and AOX1-
overexpressed transgenic tobacco lines (B7 and B8), which were compared with wild
type tobacco in order to analyze their different responses to cold stress. Lipid
peroxidation levels (an indicator for chilling damage) were measured to evaluate the
oxidative damage caused by cold stress. On the other hand, the expressions and activities
of major antioxidant enzymes were analyzed to evaluate plant responses to the change of
redox status in vivo. In addition, the contents of soluble sugars (glucose, fructose and
sucrose) and insoluble sugar (starch) were compared between WT and transgenic plants
with the hope to reveal the effect of altered AOX expressions on carbon metabolism.
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Chapter 2
Materials and Methods 2.1 Cloning and characterization of tobacco AOX2 gene
2.1.1 Plant materials and growth conditions
Wild-type tobacco (Nicotiana tabacum) and AOX1-silenced transgenic lines (termed RI9
and RI29) were used for AOX2-related research. The various plant materials used for
RNA extraction were prepared as follows: To get the imbibed seeds, dry seeds were
surface-sterilized with 70% ethanol and 10× diluted bleach sequentially, rinsed with
sterile water for 5 times (1min for each) and then kept in sterile water at room
temperature for 24 h and 72 h, respectively. Young seedlings were obtained by
germinating seeds on Murashige and Skoog (MSO) medium for 10 days. For collecting
leaf and root tissues, seeds were germinated in vermiculite. Two weeks later seedlings
were transferred into the hydroponic tank filled with the 10 × diluted full-strength
Hoagland’s solution and cultivated at room temperature with continuous light. After
another two weeks, leaf tissues and root tissues were harvested separately. To obtain the
tissues at the reproductive stage, young seedlings were transferred into the growing
medium (Pro-mix : vermiculite [4 : 1]) and kept in the growth chambers (Model PGR-15,
Conviron, Winnipeg, Canada) with a 16 h photoperiod, a temperature of 28 °C/22 °C
(light/dark) and a relative humidity of 60%. Buds, anthers, ovaries, petals and sepals were
collected separately.
2.1.2 RNA extraction
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RNA was extracted from leaves, roots, buds, anthers, ovaries, petals, sepals, imbibed
seeds and young seedlings by “TRIZOL method” (Invitrogen). For extracting leaf RNA,
high salt solution (0.8 M sodium citrate and 1.2 M NaCl) was added into the sample at
the step of RNA precipitation, which prevents some contaminating compounds like
polysaccharides and proteoglycans from precipitating together with RNA (TRIZOL
Reagent instruction, Invitrogen). The RNA sample was quantified with
spectrophotometer at 260 nm.
C [μg/μl] = V × A260nm / (ε× d ×v)
V = total volume [μl]
A260nm= absorbance at 260nm
ε (extinction coefficient of RNA at 260nm) = 25 [μl×μg-1 cm-1]
d= light path [cm]
v = sample volume [μl]
2.1.3 Primer designing for 5’-RACE
By aligning the partial sequence of AOX2 with tobacco AOX1 sequences obtained from
National Center for Biotechnology Information (NCBI) database with Clustalx 1.8, low-
conserved regions were identified and used for designing primers (see figure 3.2).
5’RACErev1, 5’RACErev2, 5’RACErev3 and P1rev were designed for 5’-RACE PCR.
The Universal Primer Mix (UPM) which binds to 5’-end of 5’-RACE-ready cDNA was
provided in the RACE cDNA Amplification Kit. P1fwd and 3’fwd primers were used
together with other reverse primers to characterize RACE products (Table 2.1).
- 37 -
Table 2.1 The primers used for 5’-RACE PCR and characterization of RACE products.UPM: universal primer mix, which contains a long universal primer (UPM [L]) and a shortuniversal primer (UPM [S]). The annealing position and orientation of AOX2-specificprimers on AOX2 template were shown in figure 3.2.
- 38 -
2.1.4 5’-RACE of AOX2 gene
The main cloning procedure followed the instruction in SMART™ RACE cDNA
Amplification Kit produced by Clontech. Total RNA (2 μg) extracted from anther was
used for first-strand cDNA synthesis. Four 5’-RACE reverse primers were designed
according to the partial AOX2 sequence. They were expected to work together with UPM
to amplify AOX2 fragments with distinguishable size differences, which could facilitate
the analysis of RACE product. To increase the specificity of amplification in 5’-RACE,
touchdown-PCR was performed by starting with high annealing temperature (>70 °C).
The PCR program is as follows: 5 cycles: 94 °C 30 sec, 72 °C 3 min; 5 cycles: 94 °C 30
sec, 70 °C 30 sec, 72 °C 3 min; 25 cycles: 94 °C 30 sec, 65 °C 30 sec, 72 °C 2 min. PCR
products were fractionated in 1% agarose gel. The promising bands were gel-purified
with QIAquick Gel Extraction Kit (Qiagen) and verified by PCR with fwd primers and
5’RACErev primers, which were then cloned into pGEM-T easy vector (Promega,
Madison, WI) for sequencing.
2.1.5 Sequence analysis
The AOX2 sequence I cloned was translated with on-line translation Tool in Expert
Protein Analysis System (ExPASy) and analyzed with various bioinformatics tools. DNA
and protein sequence alignments were conducted with Clustalx 1.8. The mitochondrial
targeting sequence was predicted with MITOPROT. The membrane binding domain was
predicted with TMHMM Server 2.0. The putative motifs and the critical cysteine residue
in tobacco AOX2 protein sequence were analyzed by comparing with tobacco AOX1.
- 39 -
2.1.6 Phylogenetic analysis
Full-length protein sequences of AOX from tobacco and other species were retrieved
with Blastx search on the website of NCBI and aligned by Clustalx 1.8. The phylogenetic
tree was calculated by neighbor-joining method with 1000-replicate bootstrap, which
revealed the phylogenetic relationship between tobacco AOX2 and AOX genes in other
species.
2.1.7 RT-PCR assay
Total RNA (1 μg) extracted from leaves, roots, buds, anthers, ovaries, petals, sepals
imbibed seeds and young seedlings were applied in reverse transcription-PCR (RT-PCR)
analysis using Access RT-PCR System (Promega) to reveal the expression pattern of
AOX2 gene in various tissues. To eliminate DNA contamination, RNA samples (1 μg)
were incubated with 1U DNAse for 15 min at room temperature followed by adding 1 μl
of 25 mM EDTA. The samples were then heated at 65 °C for 10min to inactivate the
DNAse. The primer pair used in RT-PCR was Probe fwd and rev (Figure 3.2). The RT-
PCR program is as follows: 1 cycle: 45 °C 45 min; 1 cycle: 94 °C 2 min; 20 cycles: 94 °C
30 sec, 60 °C 1 min, 68 °C 2 min; 1 cycle: 68 °C 7 min.
2.1.8 Northern blot analysis
Northern blot analysis was performed as described by Sieger et al. (2005). Total RNA
(20 μg) from different tissues were fractionated in 1% denaturing agarose gel and
transferred to Hybond N+ nylon membranes (Amersham Pharmacia). Membrane-bound
RNA was firstly pre-hybridized with denatured salmon sperm DNA (100 μg/ml) for 4~5
- 40 -
hours and then hybridized with AOX1 or AOX2 specific probes which were 32P-labeled
by random priming using a Random Prime Labeling System (Amersham) for overnight at
65 °C or 60 °C respectively in hybridization buffer (0.25 M Na2HPO4, pH 7.2, 7% SDS).
On the next day, AOX1 and AOX2 blots were washed with Wash buffer (Wash buffer I:
20 mM Na2HPO4, pH 7.2, 5% SDS; Wash buffer II: 20 mM Na2HPO4, pH 7.2, 1% SDS)
under 65 °C and 60 °C respectively until background was low and then the blots were
exposed to films for autoradiography (Sieger et al., 2005). The gene specific primers used
for probe synthesis were listed in table 2.2.
- 41 -
Table 2.2 The primers used for synthesizing cDNA probes for northern blot analysis. Thelengths of cDNA products and gene accession number were also included in the table.Primer designing for AOX2 gene was based on its known partial sequence. The annealingposition and orientation of AOX2-specific primers on AOX2 template were shown in figure3.2.
- 42 -
2.2 Role of AOX in ROS balance and carbon metabolism under cold
stress
2.2.1 Generation of transgenic plants
AOX-silenced transgenic lines (Amirsadeghi et al., 2006) and AOX-overexpressed
transgenic lines (Vanlerberghe et al., 1998) were generated in our previous research work.
For generating AOX-silenced transgenic lines, a T-DNA construct containing two
tobacco AOX1 cDNA copies (around 1.4kb) in an inverted repeat orientation separated by
pyruvate orthophosphate dikinase intron (encoding an intron-spliced hairpin RNA) was
integrated into pKANNIBAL vector. The resulting construct together with the upstream
cauliflower mosaic virus (CaMV) 35S promoter and the downstream octopine synthase
transcription termination sequence was cut and subcloned into binary plant
transformation vector pART27, which was then introduced into Agrobacterium
tumefaciens LBA4404 and used for transformation of tobacco leaf discs. The progenies
from the primary transformed plants that showed a 3:1 mendelian segregation ratio for
resistance to kanamycin were selected, which indicated the single-locus insertion of T-
DNA. Thereafter, the homozygous progenies from these selected second generations of
the primary transformed plants were screened by injecting leaves with antimycin A. The
leaf tissues with AOX effectively silenced were supposed to die under antimycin A
treatment because of the complete inhibition of respiration.
For generating AOX-overexpressed transgenic plants, an AOX1 cDNA driven by CaMV
35S promoter was built into the binary expression vector pGA748, which was then used
- 43 -
to transform Agrobacterium tumefaciens LBA4404 (Vanlerberghe et al., 1998). The rest
steps were basically the same as described above.
2.2.2 Plant materials and growth conditions
Five tobacco lines (Nicotiana tabacum): wild-type (WT), two AOX-silenced transgenic
lines (termed RI9 and RI29) and two AOX-overexpressed transgenic lines (termed B7
and B8) were applied for all of the experiments in this project.
Seeds were germinated in vermiculite and three weeks later were transferred into 4-inch
plastic pots containing a general purpose growing medium (Pro-mix: vermiculite = [4: 1]),
which were kept in growth chambers (Model PGR-15, Conviron, Winnipeg, Canada)
with a 16 h photoperiod (irradiance is 110~130 μmol/m2 /s), a temperature of 28 °C/22 °
C (light/dark) and a relative humidity of 60%. Plants were irrigated with 10x diluted full-
strength Hoagland’s solution everyday. After three weeks fully-developed leaves were
sampled 12 h after light restoration. For short-term cold stress, plants were transferred to
another growth chamber with the same conditions as the previous chamber except for the
temperature (12 °C/5 °C, light/dark) and sampled at 0 h (ctrl), 24 h, 48 h and 72 h after
cold shift, respectively. For long-term cold stress, seeds were initially germinated under
low temperature condition in growth chamber (12 °C/5 °C, light/dark) and leaf tissues
were sampled in around 90 days.
2.2.3 Mitochondrial isolation
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The procedure of mitochondrial isolation was described in Vanlerberghe et al. (1995)
with slight modification. Around 40 g of leaf tissues with the main vein removed were
collected and homogenized with homogenization buffer (0.3 M sucrose, 25 mM N-tris
(hydroxymethyl) methyl-2-aminoethane-sulfonic acid (TES), 2 mM EDTA, 10 mM
KH2PO4, 1% PVP-40, 0.5% BSA, 20 mM ascorbic acid, 4 mM cysteine, pH 7.5) in pre-
cooled mortar and pestle. The mixture was filtered through 2 layers of miracloth
(Calbiochem) and centrifuged at 3,000 rpm for 5 min at 4 °C (All subsequent procedures
were carried out at 4 °C). The supernatant was then transferred to the fresh tubes and
centrifuged at 12,000 rpm for 20min. The pellet was washed with 1X wash (+BSA) (0.3
M sucrose, 10 mM TES, 0.1% BSA, pH 7.2) and centrifuge at 3,000 rpm for 5 min. The
supernatant was collected and centrifuged at 12,000 rpm for 20 min. Thereafter, the pellet
was suspended with 2 ml of 1X wash (+BSA), loaded onto the PVP-percoll gradients
(Day et al., 1985) and then centrifuged at 18,000 rpm for 40 min. The mitochondrial
fraction in the gradient was collected and washed with 1X wash (+BSA). Finally,
mitochondrial extract was suspended in 1X wash (-BSA) (0.3 M sucrose, 10 mM TES,
pH 7.2) containing 5% dimethylsulfoxide (DMSO) and stored in -80 °C. Mitochondrial
proteins were quantified by modified Lowry assay (Larson et al., 1986).
2.2.4 Western blot analysis of mitochondrial proteins
Reducing SDS-PAGE was carried out with an SE 600 electrophoresis unit (Hoefer
Pharmacia Biotech, San Francisco, CA). A 5% (w/v) polyacrylamide stacking gel and 10
to 17.5% polyacrylamide gradient resolving gel were used. Mitochondrial proteins (100
μg) were mixed with 3X sample buffer (125 mM Tris-HCl, 6% SDS, 6% β-
- 45 -
mercaptoethanol and 30% glycerol) and incubated in the boiling water for 2 min,
followed by adding 0.08% bromophenol blue. The loaded gel was allowed to run at
constant current (50mA) for 4 h. A TE 50X electrotransfer unit (Hoefer Pharmacia
Biotech, San Francisco, CA) was then applied to transfer the resolved proteins to
nitrocellulose membrane (at 0.5 to 1 amp for 1 to 2 h). After gel transfer, the blot was
dried briefly and washed twice with PBS-tween (10 mM NaH2PO4, 150 mM NaCl and
0.3% Tween 20, pH7.2) for 15 min. For immunoanalysis, the blot was incubated in
SuperBlock blocking buffer (Pierce) for 2 h before hybridizing with the monoclonal
antibody recognizing either AOX (Elthon et al., 1989) or Cyt oxidase II (a gift from Dr.
Tzagaloff, Columbia University, New York) at a dilution rate of 1:2,000 for 1 h. After
that, the blot was washed 6 times (5 min for each) with PBS-Tween and incubated with
second antibody from Goat anti-mouse IgG (H+L; Pierce Laboratories) at a dilution rate
of 1:25,000 for 1 h. After another 6 times wash with PBS-Tween, the blot was incubated
with Supersignal West Pico Chemiluminescent detection reagent (Pierce) for 15 min and
exposed to films for autoradiography.
2.2.5 RNA extraction from polysaccharide-rich tissues
To remove the excessive polysaccharides accumulated in leaf tissues during cold stress,
which usually co-precipitate with RNA, the method described in Vanessa et al. (2008)
was used for RNA extraction with some revision. The frozen leaf tissues were ground
using mortar and pestle and homogenized with guanidinium hydrochloride (GHCL)
extraction buffer (6.5 M guanidinium hydrochloride, 100 mM Tris-HCl pH 8.0, 500 mM
sodium acetate pH 5.5, 0.1 M β-mercaptoethanol). The mixtures were divided into
- 46 -
several tubes which were then placed in liquid nitrogen until finishing all samples. Then
1 M potassium acetate was added into the tube to get the final concentration of 0.2 M
potassium acetate. The tubes were vortexed for 15 sec and incubated at room temperature
for 10 min to precipitate polysaccharides. After that, the mixtures were centrifuged at
12,000 g at 4 °C for 10 min. The supernatant containing RNA was pipetted into a new
tube and 0.5 ml TRIZOL was added into each tube. After incubated for 5 min at room
temperature, each tube received 0.2 ml chloroform and was centrifuged at 12,000 g at 4 °
C for 10 min. The upper aqueous phase was transferred to a new tube and then was mixed
with 0.5 ml of isopropyl alcohol. RNA was allowed to precipitate at -20 °C for 1 h and
then was collected by centrifugation at 12,000 g at 4 °C for 20 min~30 min. The RNA
pellets were washed with 1 ml 70% ethanol, dried with vacuum dryer for 5 min and
suspended in 40 μl of DEPC-treated water. The RNA sample was then quantified with
spectrophotometer at 260 nm.
2.2.6 Northern blot analysis
The procedure for gel electrophoresis, membrane transfer and probe hybridization
followed the protocol described in section 2.1.8. To generate the probes for hybridization,
partial sequence for each designated gene was amplified with RT-PCR from tobacco total
RNA and cloned into pGEM-T easy vector. The cDNA fragments were then excised from
these plasmids and used for radioactive labeling. The primer sequences used for probe
synthesis were shown in table 2.3.
- 47 -
Table 2.3 The primers used for synthesizing cDNA probes for northern blot analysis. Thesubcellular locations of genes, lengths of cDNA products and gene accession numbers werealso included in the table. The sequence of PTOX was obtained from our previous work(Amirsadeghi et al., 2006). The sequence of COX6b was obtained from tobacco ExpressedSequence Tag (EST) database (http://compbio.dfci.harvard.edu/tgi/cgi-bin/tgi/gimain.pl?gudb=Tobacco).
- 48 -
2.2.7 ROS detection
H2O2 and O2.- levels in leaves were detected with Diaminobenzidine (DAB) and Nitroblue
tetrazolium (NBT) method respectively, as described in Dutilleul et al. (2003) with some
modification. Leaf discs were punched out with a cork borer (1.5 cm in diameter) from
two fully developed leaves of each plant and vacuum infiltrated 5~6 times with syringe
immediately with either 1 mg/ml DAB in ddH2O (pH 3.8) or 0.25 mg/ml NBT in 10 mM
potassium phosphate buffer (pH7.8). Then the leaf discs were incubated at room
temperature in the dark for 16 h (for DAB) or 1 h (for NBT), during which H2O2 reacted
with DAB to form a deep brown product while O2.- reacted with NBT to generate a dark
blue insoluble compound. Thereafter, the leaf discs were cleared with boiling 95%
ethanol to completely remove all the chlorophyll and were then stored in 30% glycerol
for color intensity analysis with densitometer.
2.2.8 TBARS assay
Lipid peroxidation is usually used to evaluate chilling injury of plants (Lukatkin, 2002).
The level of lipid peroxidation in tobacco leaves, denoted by malondialdehyde (MDA)
content was determined by thiobarbituric acid-reactive-substances (TBARS) assay (Fryer
et al., 1998) with slight modifications. MDA, formed in the lipid peroxidizing system,
can react with thiobarbituric acid (TBA) at low pH with heat and yield a pink chromagen
with an absorbance maximum at 535 nm, which could be detected by spectra assay. Six
leaf discs (1.5 cm in diameter) were weighed and homogenized with 4 ml of 5 mM
potassium phosphate buffer (pH 7.0) in a pre-cooled mortar and pestle and centrifuged at
1,000g for 10 min at 4 °C. The supernatant (0.9 ml) was mixed with 0.6 ml TBA reaction
- 49 -
solution containing 0.45% (w/v) SDS, 250 μl of 20% (w/v) acetic acid (pH 3.5) and 250
μl of 0.8% (w/v) TBA. The control group was a mixture of 0.9 ml potassium phosphate
buffer and 0.6 ml TBA reaction solution. The mixture were incubated at 98 °C for 1 h
and centrifuged at 12,000 g for 5 min after being cooled to room temperature. The
supernatants were applied for the spectra assay. The subtracted absorbance (A535-A600)
was used for the calculation of MDA contents (extinction coefficient: 1.56×105 M-1 cm-1).
2.2.9 Enzyme assay
APX activity
The procedure for the assay was described in Panchuk et al. (2002). One-hundred
milligrams of frozen leaf tissues stored at -80 °C were ground in liquid nitrogen, mixed
with 0.5 ml of extraction buffer (50 mM Na-phosphate (pH 7.0), 0.25 mM EDTA, 2%
(w/v) polyvinylpyrrolidone-25, 10% (w/v) glycerol, and 1 mM ascorbic acid [AsA]) in
the tube for 30 sec and centrifuged at 13,200 g for 10 min at 4°C. The supernatant (25 μl)
was immediately applied for the measurement of APX activity by mixing with 975 μl
reaction buffer (25 mM Na-phosphate [pH 7.0], 0.1 mM EDTA, 1 mM H2O2, 0.25 mM
AsA). The oxidation rate of AsA was detected by monitoring the decrease of absorbance
at 265 nm (extinction coefficient: 14 mM -1 cm -1) every 30 sec for 4 min in total. Protein
concentration was determined by Lowry assay (Larson et al., 1986). APX activity could
be lost quite easily at room temperature. Hence all the extraction steps were carried on at
4 °C and the assay was performed within 30 min (Panchuk et al., 2002).
SOD activity
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Total SOD activity in leaf was measured according to the method described in Martinez
et al. (2001) with some modification. Three-hundred milligrams of frozen leaf tissues
stored at -80 °C were ground in liquid nitrogen and homogenized in 100 mM potassium
phosphate buffer (pH 7.8) containing 0.1 mM EDTA, 1% (w/v) PVP, and 0.5% (v/v)
Triton X-100 (Janknegt et al., 2007). After incubation on ice for 10 min, the mixtures
were centrifuged at 13,200 rpm for 10 min at 4 °C. The supernatant was aliquoted into
fresh tubes and flash-frozen with liquid nitrogen. One aliquot was used for quantification
of protein content with Lowry assay (Larson et al., 1986). To measure the activity of
SOD, ten milligrams of protein sample (100 μl) was mixed with 2.9 ml reaction buffer
(50 mM potassium phosphate (pH7.8), 0.1 mM EDTA (pH 7.8), 13 mM methionine,
0.075 mM NBT and 0.002 mM riboflavin) in transparent glass tube (75×10 mm, Fisher).
The control group was a mixture of 100 μl extraction buffer and 2.9 ml reaction buffer.
Reaction was initiated by placing the tube between two light banks consisting of two 15
W fluorescent lamps. The absorbance of the solution at 560 nm was measured every two
minutes. The data of A560 was plotted against the reaction time (min) and the slope was
used to calculate % inhibition, as shown in the formula below. 50% inhibition was
defined as one enzyme unit.
% inhibition = (slope of buffer control - slope of sample)×100 / slope of buffer control
2.2.10 Sugar assay
Sugar extraction
The methods of Stitt et al. (1989) and Jones (1981) were used as main templates for the
procedure of sugar assay with some revision. Frozen leaf tissues were ground into
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powder in liquid nitrogen, freeze-dried at -50 °C for 5 h (with vacuum of 7 microns Hg).
Leaf powder was weighed (around 6mg) and extracted three times with 80% (v/v)
ethanol for 20 min at 80 °C. For each round of extraction, the mixture was centrifuged at
14,000 rpm for 5 min and the supernatant was then transferred into a new tube.
Eventually the supernatants from three rounds of extraction were pooled and 5mg
activated charcoal was added to get rid of the particle and pigment. The sample was
vortexed for 1 min and centrifuged at 4 °C for 10 min at 16,000 g. The supernatant was
transferred to a fresh tube. Thereafter, the activated charcoal was suspended twice in 0.2
ml 80% (v/v) ethanol and centrifuged for 10 min at 16,000 g to collect the residual sugars.
The supernatants were pooled together with the previous sugar extract and dried in the
rotary vacuum system at 40 °C for 2 h to remove all the ethanol. After that, 1 ml ddH2O
was added into the tube to dissolve the soluble sugars. Before the assay, the soluble sugar
extract was membrane-filtered with 25 mm syringe filter (pore: 0.2 μm) (Pall) in order to
produce a clear sample.
For starch analysis, the insoluble pellet during the ethanol extraction was washed twice
with ddH2O and solubilized by heating at 95 °C in 0.1 M NaOH for 60 min. After
acidification to pH 4.9 with 1 M acetic acid, the suspension was digested overnight at 55
°C with 1 ml enzyme solution (0.2 M sodium acetate, pH 5.0, 2 U/ml amyloglucosidase
[Sigma, A7420], 10 U/ml α-amylase [Sigma, A3403]). After centrifuged at 16,000 g for
5 min, the supernatant was diluted by 20 times and membrane-filtered. The glucose in the
diluted solution was used to assess the starch content of the sample.
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Enzyme preparation
Glucose-6-phosphate dehydrogenase (G6PDH) (Sigma, G8404) and phosphoglucose
isomerase (PGI) (Sigma, P5381) were both supplied as suspension in ammonium sulfate.
To remove the ammonium sulfate, the suspensions were centrifuged for 5 min at 16,000 g
and supernatants were carefully removed. The precipitated enzymes G6PDH and PGI
were dissolved in G6PDH buffer (100 mM Tris-HCl, pH 8.1, 5 mM MgCl2) and PGI
buffer (100 mM Tris-HCl, pH 8.1), respectively. The enzymes were then aliquoted into
fresh tubes, flash-frozen with liquid nitrogen and stored at -80 °C. Hexokinase (HK)
(Sigma, H6380) and invertase (INV) (Sigma, I9274) were dissolved in enzyme buffer
(100 mM Tris-HCl, pH 8.1, 5 mM MgCl2, 50% glycerol) and stored at -20 °C.
Determination of sugar contents with enzymatic cycling assay
Soluble sugar extract (40 μl) was added to 760 μl of assay medium (100 mM imidazole,
pH 6.9, 1.5 mM MgCl2, 0.5 mM NADP+, 1.1 mM ATP) to give a reaction volume of 800
μl. Samples were mixed and then placed in the spectrophotometer. The absorbance at
340nm representing the conversion from NADP+ to NADPH was monitored (Figure 2.1).
One unit of G6PDH was added to assay glucose-6-phosphate (G6P) content, followed by
an addition of 0.5 U of HK to assay the glucose content. Once the A340 had leveled off,
2 U of PGI were added for determination of fructose and fructose-6-phosphate (F6P).
After that 20 U of INV were added for the determination of sucrose. For another sample,
1 U of G6PDH was added followed immediately by an addition of 2 U of PGI to assay
F6P content.
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For starch assay, 20 μl of 20 times-diluted digested samples were added to 780 μl of
assay medium to obtain 800 μl final volume. 1 U of G6PDH was added followed
immediately by an addition of 0.5 U of HK to assay the glucose content, which acts as
the equivalent of starch.
After obtaining the absorbance change at 340 nm for each kind of sugar, the
concentrations of them were determined with the formula below:
C [mol/L] = V × ∆A / (ε× d ×v)
V = final reaction volume [ml]
∆A= absorbance change
ε (extinction coefficient of NADPH at 340 nm) = 6300 [M-1 cm-1]
d= light path [cm]
v = sample volume [ml]
2.2.11 Statistical analysis
The statistical calculations in this project were carried out with two-way ANOVA
integrated in Graphpad Prism 5.0 to determine the significant difference between
different tobacco lines (WT versus AOX1 transgenic lines) or different temperature
treatments (control versus low temperature).
- 54 -
Figure 2.1 Principle for the enzymatic cycling assay. The change in A340nmmonitored by spectrophotometer represents the reduction of NADP+ to NADPH duringthe conversion of G6P To 6PG. The amounts of different sugars could be determinedby adding G6PDH, HK, PGI and INV sequentially into the assay system. G6PDH,glucose-6-phosphate dehydrogenase; PGI, phosphoglucose isomerase; HK,hexokinase; INV, invertase; G6P, glucose-6-phosphate; F6P, fructose-6-phosphate;6PG, 6-phosphogluconate.
- 55 -
Chapter 3
Results 3.1 Cloning and characterization of tobacco AOX2 gene
3.1.1 Cloning of 5’-region of tobacco AOX2
RNA extracted from WT anther with the “TRIZOL method” served as the template for
5’-RACE PCR. Four AOX2 gene-specific 5’-RACE rev primers were designed and used
for 5’-RACE cloning (Figure 3.1 A).
DNA gel analysis on the 5’-RACE PCR products showed that multiple bands appeared in
all different groups (figure 3.1A). After making a further analysis on the size of each
band, I found that the size difference between most intensive band in each group (boxed
in figure 3.1A), which was also the most promising one, conformed well to my prediction
based on their different primer binding positions on AOX2 template (Figure 3.2). To
further confirm if these bands were AOX2 fragments, they were gel-purified and served
as templates in PCR testing (Figure 3.1B). The result showed that bands with expected
sizes (160 bp, 199 bp and 480 bp) were amplified from these three selected fragments
respectively with the nested primers and no band was amplified if one of the primers was
beyond the templates (negative control), which indicated that the three fragments I
selected were probably all AOX2 gene fragments. These putative AOX2 fragments were
then cloned into pGEM-T easy vector and sent for sequencing. The sequencing results
were verified and combined with known 3’-fragment of AOX2 to generate full-length
sequence (Figure 3.2).
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A B
Figure 3.1 DNA gels showing the products of 5’-RACE (A) and the productcharacterization (B). Three RACE products with the highest intensity in eachamplification group were identified and tested with AOX2-specific primers. Lane 2, 4and 6 in (B) were the PCR products amplified with AOX2 nested primers (thepredicted size of each product was shown above each band). Lane 3 and 5 werenegative controls whose reverse primer binding positions were beyond thecorresponding template regions. The primer pair used for each PCR reaction wasshown at the top.
- 57 -
3.1.2 AOX2 sequence was characterized by bioinformatic methods
DNA sequence analysis
Eventually the complete coding sequence and flanking untranslated regions (UTR) of
tobacco AOX2 gene were cloned, which contain 311 nt 5’-UTR, 1098 nt coding sequence
and 273 nt 3’-UTR. The full length DNA sequence alignment of AOX1 and AOX2 genes
in tobacco indicates that their 3’-coding regions share high similarity, while the 5’-UTR,
5’-coding regions and 3’-UTR are quite distinct from each other (Figure 3.2).
- 58 -
Figure 3.2 DNA sequence alignment of tobacco AOX1 and AOX2 with Clustalx 1.8. Theblack arrow pair (P1 fwd and P1 rev) indicates the primers used for initial amplification ofthe 200bp fragment. The red arrow pair indicates the primers used for RT-PCR and probesynthesis. Three blue-colored primers (5’RACE rev1, 2 and 3) and P1 rev were used asgene specific primers (GSP) in 5’-RACE. 3’fwd primer (green color) and P1 fwd wereused together with gene-pecific reverse primers for PCR testing of RACE products. All theprimers were designed with Oligo 6.0. The start codons and stop codons were boxed inblack and red color, respectively. The codons encoding the critical regulatory cysteineresidue were boxed in blue color. The asterisk “*” indicates the identical nucleotide.
- 59 -
- 60 -
Protein sequence analysis
AOX2 protein sequence predicted from its nucleotide sequence was aligned with tobacco
AOX1 (Figure 3.3). The deduced AOX2 protein contains 365 residues, which is 12
residues longer than AOX1. Their protein sequences show low similarity at the N-
terminus but high similarity at the C-terminus. With the help of the bioinformatic tools,
some crucial motifs and structures were identified in their sequences. Mitochondrial
targeting peptide (mTP), which is rich in positively charged residues but lack of acidic
residues (Hartl et al., 1989; Claros et al., 1996), was detected at the N-terminus of both
AOX1 and AOX2 proteins. The critical regulatory cysteine that is involved in the
covalent linkage of the two AOX monomers via an S-S bond and stimulation by pyruvate
was identified in both of AOX1 and AOX2 sequences. Consistent with the structural
model proposed by Andersson et al. (1999), a four-helix bundle at the C-terminus was
identified in both of AOX1 and AOX2 protein sequences. The hydrophobic region
between helix 2 and helix 3 was proposed to be inserted into the membrane. Two
conserved Glu-X-X-His motifs are located in helix 2 and helix 4, respectively, which
were assumed to be involved in the formation of the binuclear iron center (Siedow et al.,
1995).
Another interesting thing I noticed through sequence analysis is that the second amino
acid in the antibody-recognizing region (RADEAHHRDVNH, Finnegan et al., 1999) of
AOX2 is threonine rather than alanine (Figure 3.3), which is usually quite conserved in
other AOX proteins (Finnegan et al., 1999). This variation may affect the AOX antibody
recognition of tobacco AOX2 protein.
- 61 -
Figure 3.3 The protein sequences of tobacco AOX1 and AOX2 were deduced fromcDNA sequences and aligned by Clustalx 1.8. Mitochondrial targeting peptides (mTP)boxed in green color are located at the N-terminus of both AOX1 and AOX2. Thehighly conserved regulatory cysteine residue was boxed in black color with “∇”. Fourpredicted helices are designated as “H1”, “H2”, “H3” and “H4”. Membrane bindingregion is designated as “M”. The conserved diiron-binding motifs (E-X-X-H) areindicated with “♦”. The AOX antibody-binding site is denoted by a black bar with “ ”indicating the divergent site between AOX1 and AOX2. The asterisk “*” indicates theidentical residue; “:” indicates conserved substitutions; “.” indicates semi-conservedsubstitutions.
- 62 -
Phylogenetic analysis of AOX2 gene
A bootstrap phylogenetic tree based on full-length protein sequences was constructed to
reveal the relationship between tobacco AOX2 and other AOX genes. All the sequences
were retrieved from the NCBI database. The phylogenetic analysis clearly showed that
the AOX genes from different species were classified into either AOX1 or AOX2 clade.
The gene I cloned was grouped into AOX2 gene clade (Figure 3.4).
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A.thaliana aox1b NM 113134
A.thaliana aox1a NM 113135
N.attenuata aox1 AY422688
N.tabacum aox1a S71335
N.tabacum aox1b X79768
S.tuberosum aox AB176953
C.roseus aox AB009395
G.hirsutum aox DQ250028
L.esculentum aox1b AY034149
S.officinarum aox AY644465
Z.mays aox2 AY059647
O.sativa aox1a AB004864
T.aestivum aox AB078882
Z.mays aox3 AY059648
P.bipinnatifidum aox AB190213
D.vulgaris aox AB189673
G.max aox2b U87907
A.thaliana aox2 NM 125817
M.indica aox X79329
C.sativus aox2 AY258276
G.max aox2a U87906
V.unguiculata aox2a AJ319899
D.carota aox2 EU286575
V.vinifera aox2 EU523224
N.tabacum aox2
AOX2
AOX1
987
999727
1000
479
495
976
445
1000973
997
642
999
599
994
836
521
449
462
1000
640
195
0.05
Figure 3.4 Phylogenetic tree demonstrating the sequence homology between tobaccoAOX2 and other AOXs in different species. Protein sequences rather than DNAsequences were used for the alignment, which performed a better differentiationbetween the AOX1 clade and AOX2 clade. Sequences were aligned with Clustalx 1.8using the BLOcks of Amino Acid SUbstitution Matrix (BLOSUM) and the phylogenictree was calculated by neighbor-joining method with 1000-replicate bootstrap. Thenumber at each branching site stands for the frequency of reproduction in 1000bootstrap replicates. All gene sequences were retrieved from NCBI database and onlycomplete sequences were used for tree construction.
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3.1.3 Expression of AOX2 displayed tissue specificity
Total RNA was extracted from different tobacco tissues and AOX2 expression was
analyzed with RT-PCR and northern blot.
RT-PCR performed with AOX2 gene-specific primers showed that AOX2 could be
amplified from buds, ovaries, anthers, petals, sepals, young seedlings, leaves and roots,
but not in imbibed seeds (Figure 3.5). Northern blot analysis with AOX2 specific probe
indicated that compared with tobacco AOX1 gene, the transcript levels of AOX2 gene
were much lower in all selected tissues and the hybridization signal was stronger in
anther than in other tissues (Figure 3.6).
To better understand the expression pattern of AOX2 in reproductive tissues and to see if
knockdown of AOX1 has any influence on AOX2 expression, I collected anther and
ovary tissues of WT and AOX1-silenced transgenic lines (RI9 and RI29) with different
stages based on the length of buds (Koltunow et al., 1990) and analyzed AOX2
expression with northern blot. The results showed that AOX2 transcript levels in anther
were generally a bit higher than in ovary. However, no obvious pattern was detected
between different lines or different stages (Figure 3.7).
- 65 -
A
200bp 100bp
B
200bp 100bp
Figure 3.5 (A) Expression analysis of AOX2 gene in different tobacco tissues with RT-PCR. AOX2-specific primers (probe fwd and probe rev in Figure 3.2) were used for thisexperiment. The expected size of RT-PCR product is 122bp. (B) An example showingthat DNA contamination was eliminated by adding DNase to RNA samples. Negativecontrols were performed without adding reverse transcriptase into the reaction systemto make sure that there was no DNA contamination.
- 66 -
A
AOX1 AOX2
B
Figure 3.6 (A) Northern blot analysis of tobacco AOX2 expression in seedling, bud,ovary, anther, petal, sepal, leaf and root. AOX1 blot was used as a control group. (B)Relative transcript levels of AOX2 gene determined by densitometer analysis ofnorthern blot.
- 67 -
WT RI9 RI29 WT RI9 RI29 WT RI9 RI29 Ovary
Anther
Length of bud <22mm 22mm~43mm >43mm
Figure 3.7 Northern blot analysis of AOX2 expression in ovary and anther tissues ofWT and AOX1-silenced transgenic lines RI9 and RI29 with different stages (based onthe length of bud).
- 68 -
3.2 Role of AOX1 in ROS balance and carbon metabolism under cold
stress
3.2.1 Transgenic tobacco plants with altered expression levels of AOX
The levels of AOX transcript and protein in WT, two AOX-silenced transgenic plants
(RI9 and RI29) and two AOX-overexpressed transgenic plants (B7 and B8) before and
after the cold stress were analyzed by northern and western blot, respectively. Under the
normal condition, both AOX transcript and protein were almost undetectable in WT,
while after exposure to cold stress, AOX in WT was induced dramatically at 24 h time
point and then declined slightly at the later time point (72 h), but was still well above the
control level. Compared with WT, AOX protein levels in RI9 and RI29 were barely
detectable before and after the cold stress, except that in RI9 the expression of AOX was
moderately induced by cold stress and a small amount of AOX protein could be detected
in 48 h cold-treated sample. For northern blot, the smear underneath AOX bands in RI9
and RI29 was probably an indication of RNA degradation due to RNA interference. In
B7 and B8, AOX transcript and protein were much more abundant than in WT at both
control and low temperature conditions. The levels of Cyt oxidase subunit II protein
(COXII), which served as an internal control, were quite constant throughout all the lines.
1 The word “AOX” used in describing the experiments or results of second project refers to tobacco AOX1 unless otherwise mentioned.
- 69 -
A
B
Figure 3.8 Northern (A) and western (B) blot analysis of AOX in WT, AOX-silencedmutants (RI9 and RI29) and AOX-overexpressed mutants (B7 and B8). Total RNAand purified mitochondrial proteins from tobacco leaves were subjected to northernand western blot analysis, respectively. The bottom row of numbers refers to the timeafter plants were transferred to the cold environment. The AOX and COXII proteinwere detected by probing with antibodies against AOX and COXII protein,respectively. COXII was used as an internal control indicating the equal loading ofproteins.
- 70 -
3.2.2 Change of oxidative damage after cold shift showed differential patterns
among different lines
To evaluate the oxidative damage caused by ROS generation during cold stress, the levels
of lipid peroxidation represented by malondialdehyde (MDA) contents were determined
by TBARS assay. Under the normal condition (0 h), contents of MDA in two AOX-
silenced mutants were higher than in WT (significant difference was detected between
WT and RI29), while in B8 were lower than in WT. In another AOX overexpressor B7,
MDA level was similar to WT (Figure 3.9 A). Once exposed to the cold stress, lipid
peroxidation levels in WT and RI9 were increased significantly at the early time point (24
h) and then decreased steadily in the following two days (Figure 3.9 B). In comparison
with WT, lipid peroxidation levels in RI9 always stayed at a relatively higher level
(Figure 3.9 A). Contrary to RI9, lipid peroxidation levels in the other AOX-silenced
mutant RI29 was surprisingly decreased after cold treatment (24 h) and in the next two
time points (48 h and 72 h) lipid peroxidation levels kept decreasing and became even a
bit lower than WT (Figure 3.9 A and B). On the other hand, in B7 and B8 lipid
peroxidation levels slightly increased after cold shift and then returned to the normal
levels in the following two days (Figure 3.9 B), which, compared with WT, were
consistently lower throughout the 72 h cold treatment (Figure 3.9 A). Figure 3.9 C clearly
showed that the line displaying the highest level of lipid peroxidation under control
condition (RI29) was the only line to exhibit a decrease in lipid peroxidation after cold
shift.
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A B
C
Figure 3.9 The levels of lipid peroxidation in WT and transgenic plants before andafter the cold stress. MDA, one major product of lipid peroxidizing system was used topresent the lipid peroxidation level. Data were presented in two different ways (A) and(B) to facilitate the comparison among different lines or different time points,respectively. Graph (C) showed the change of MDA levels after the cold shift (24 h) indifferent lines. In graph (A) and (C), open column, black column and hatched columndenoted AOX-silenced mutant, WT and AOX-overexpressed mutant, respectively. Ingraph (B), control groups were denoted by black column while cold-treated groupswere denoted by open column. The values shown in graph are the mean±SE fromthree independent experiments. The statistical analysis was performed by two-wayANOVA. In graph (A) and (B), the comparison was carried out between black columnand other columns within each group and the significant difference was denoted by“*”, “**” or “***” (representing P<0.05, P<0.01 or P<0.001, respectively). In graph (C),bars with different letter are significantly different.
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3.2.3 RI29 and two AOX-overexpressed mutants displayed higher transcript levels
of major antioxidant genes
The transcript levels of key ROS-scavenging genes including three H2O2-scavenging
genes (ascorbate peroxidase [APX], glutathione peroxidase [GPX] and catalase [CAT])
(Figure 3.10), three O2.--scavenging genes (manganese superoxide dismutase [MnSOD],
copper-zinc superoxide dismutase [Cu/ZnSOD] and iron superoxide dismutase [FeSOD])
(Figure 3.11) and some other AOX-related genes (Cyt c oxidase subunit 6b [COX6b] and
plastoquinol terminal oxidase [PTOX]) (Figure 3.12) were analyzed by northern blot. The
RNA samples subjected to northern analysis were extracted from the leaves of all
different lines treated either with or without cold stress. Two or three independent
experiments were performed and the relative transcript levels of each gene from each
experiment determined by densitometer analysis were combined and shown in both line
graphs without error bars (Figure 3.10 - Figure 3.12) and bar graphs with error bars
(Figure 3.14).
Before the cold stress, the transcript levels of most genes were generally similar among
all different lines except for CAT and FeSOD, the transcript levels of which in two AOX
overexpressors were slightly higher than WT. After cold treatment, the transcriptions of
the key ROS-scavenging genes (APX, GPX, Cu/ZnSOD and maybe FeSOD), COX6b and
AOX analog PTOX were up-regulated in both WT and transgenic plants. However, the
induction patterns among these genes were different. For genes APX, COX6b and PTOX
their transcript levels kept increasing during the whole 72 h time-course, while for GPX
and Cu/ZnSOD the transcriptions were dramatically up-regulated at 24 h and then down-
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regulated in the following two days. Contrary to these genes mentioned above, the
transcriptions of MnSOD and CAT were down-regulated significantly by the cold stress.
On the other hand, when comparing the transcription patterns between different lines, I
found that the transcript levels of most ROS-scavenging genes (APX, GPX, Cu/ZnSOD
and FeSOD) and AOX analog PTOX in RI29 and two AOX overexpressors were
generally increased faster and were higher than in WT after cold treatment. However,
different from RI29, another AOX-silenced mutant RI9 showed the similar transcript
levels for most of genes to WT.
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Figure 3.10 Northern blot analysis of H2O2-scavengning genes (A) APX, (B) GPX and(C) CAT in WT and AOX transgenic plants before and after the cold stress. Arepresentative blot was shown for each gene. Relative transcript levels for all thegenes were determined by densitometer analysis of northern blots. WT, RI9, RI29, B7and B8 were presented with black ( ), orange ( ), red ( ), blue ( ) and greencolor ( ), respectively. The values shown in the line graphs are the means from twoto three independent experiments.
- 75 -
A. APX
B. GPX
C. CAT
- 76 -
Figure 3.11 Northern blot analysis of O2.--scavenging genes (A) Cu/ZnSOD, (B)
MnSOD and (C) FeSOD in WT and AOX transgenic plants before and after the coldstress. A representative blot was shown for each gene. Relative transcript levels for allthe genes were determined by densitometer analysis of northern blots. WT, RI9, RI29,B7 and B8 were presented with black ( ), orange ( ), red ( ), blue ( ) andgreen color ( ), respectively. The values shown in the line graphs are the meansfrom two to three independent experiments.
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A. Cu/ZnSOD
B. MnSOD
C. FeSOD
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A. PTOX
B. COX6b
Figure 3.12 Northern blot analysis of AOX-related genes (A) PTOX and (B) COX6b inWT and AOX transgenic plants before and after the cold stress. A representative blotwas shown for each gene. Relative transcript levels for all the genes were determinedby densitometer analysis of northern blots. WT, RI9, RI29, B7 and B8 wererespectively presented with black ( ), orange ( ), red ( ), blue ( ) and greencolor ( ). The values shown in the line graphs are the means from two to threeindependent experiments.
- 79 -
Figure 3.13 A representative ethidium bromide-stained RNA gel indicating the equalloading of RNA.
- 80 -
Figure 3.14 Relative transcript levels of ROS-scavenging genes and AOX-relatedgenes in WT and AOX transgenic plants before and after the cold stress shown by thebar graphs with error bars. (A) APX, (B) GPX, (C) CAT, (D) Cu/ZnSOD, (E) MnSOD,(F) FeSOD, (G) PTOX and (H) COX6b. Open column, black column and hatchedcolumn denoted AOX-silenced mutant, WT and AOX-overexpressed mutant,respectively. The values shown in graphs are the mean±SE from two to threeindependent experiments. The statistical comparison was performed by two-wayANOVA between WT and the transgenic lines within each group and the significantdifference was denoted by “*” or “**” (representing P<0.05 or P<0.01, respectively).
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B. GPX A. APX
C. CAT D. Cu/ZnSOD
Rel
ativ
e tra
nscr
ipt l
evel
E. MnSOD F. FeSOD
H. COX6b G. PTOX
- 82 -
3.2.4 The activity levels of ROS-scavenging enzymes partially conformed to their
transcript levels
To further confirm the results I obtained by northern blot analysis, the activities of the
key H2O2-scavenging enzyme APX and O2.--scavenging enzyme SOD were examined.
Similar to APX transcript data, a steady increase of APX activities after the cold shift was
observed in all five lines (Figure 3.15 A [2]), which, however, was not so dramatic as the
increase at the transcript level. APX activities were generally higher in AOX-silenced
mutants and AOX-overexpressed mutants than in WT before and after cold stress,
especially for the samples with 72 h cold treatment (Figure 3.15 A [1]). In addition, APX
activities in RI29 and B7 were increased faster than other lines after cold stress (Figure
3.15 A [2]), which coincided with what I observed in northern blot analysis.
SOD activities in all five lines were also increased after cold stress (Figure 3.15 B [2]).
Compared with WT, SOD activities in two AOX-overexpressed transgenic lines were
consistently lower (not statistically significant) both before and after cold stress (Figure
3.15 B [1]), but the speed at which the activities were increased by cold shift in
overexpressors was faster than that in WT (Figure 3.15 B [2]). On the other hand, SOD
activities in two AOX-silenced transgenic lines, which were lower than WT under the
normal condition, were enhanced faster than WT after cold stress (Figure 3.15 B [2]) and
became even a little bit higher than WT at the 72 h time point (Figure 3.15 B [1]).
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A. APX (2) A. APX (1)
B. SOD (1) B. SOD (2)
Figure 3.15 APX (A) and SOD (B) activities in WT and transgenic lines before andafter cold stress. For each enzyme activity, data were presented in two different ways(1) and (2) to facilitate the comparison among different lines or different time points,respectively. APX activity was evaluated by the capacity of protein extract from eachsample to oxidize ascorbic acid in the present of H2O2. SOD activity was presented bythe capacity of the inhibition of NBT reduction. One unit of SOD activity was definedas the amount of enzyme required to inhibit NBT reduction by 50%. Proteinconcentrations were determined by Lowry assay. In graph A (1) and B (1), opencolumn, black column and hatched column denoted AOX-silenced mutant, WT andAOX-overexpressed mutant, respectively. In graph A (2) and B (2), control groupswere denoted by black column while cold-treated groups were denoted by opencolumn. The values for APX and SOD shown in the graphs are the mean±SE fromfour and three independent experiments, respectively. The statistically significantdifference between black column and other columns within each group was denotedby “*”, “**” or “***” (representing P<0.05, P<0.01 or P<0.001, respectively), which wasperformed by two-way ANOVA.
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3.2.5 Contents of soluble sugars were proportional to the AOX levels after cold
oluble and insoluble sugars in WT and transgenic plants before and after cold
treatment
The major s
stress were quantified by enzymatic cycling assays based on the dry weights of leaf
tissues. Under the normal condition both the glucose and fructose contents in all five
lines were extremely low and could not be distinguished from each other. Once exposed
to cold stress (24 h), the contents of these two monosaccharides were increased
dramatically and kept going up in the following two days. Noticeably, after 72 h cold
treatment, both glucose and fructose contents in two AOX-silenced mutants RI9 and
RI29 were significantly lower than WT, while in two AOX-overexpressed mutants B7
and B8 were significantly higher than WT (Figure 3.16 A and B). The pool size of
disaccharide (sucrose) was not affected by cold stress too much. But interestingly, like
the monosaccharides, a similar pattern for sucrose, which was not so apparent as what I
observed for monosaccharides though, was detected in the later time points of cold stress:
AOX overexpressors B8 contained more sucrose than WT at 48 h and in AOX-silenced
transgenic line RI29 the level of sucrose was significantly lower than in WT at 72 h
(Figure 3.16 C). Measurement of the insoluble sugar starch, which was presented by
glucose equivalent, showed that the amount of starch was also increased by cold stress
like monosaccharides. However, no significant difference was detected among different
lines (Figure 3.16 D). The contents of G6P and F6P were not detectable in the assay.
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Figure 3.16 The contents of glucose (A), fructose (B), sucrose (C) and insolublesugar starch (D) (denoted by glucose equivalent) in all five lines before and after cold
A. Glucose B. Fructose
C. Sucrose D. Starch
stress. The results are the mean±SE from three independent experiments. Thestatistically significant difference between WT and transgenic line was denoted by “*”,“**” or “***” (representing P<0.05, P<0.01 or P<0.001, respectively), which wasperformed by two-way ANOVA.
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To confirm the correlation between AOX level and amount of soluble sugars observed in
the short-term cold stress, a long-term cold stress experiment was carried out. All the
plants were grown in the cold (12/5 °C) for around 90 days before the sampling.
Interestingly, the results were generally consistent with the short-term cold stress
experiments: the contents of glucose and fructose in AOX-overexpressed transgenic lines
and AOX-silenced transgenic lines were respectively higher and lower than in WT.
However, no obvious difference in sucrose or starch contents was found between WT and
transgenic plants (Figure 3.17).
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A. Soluble sugars B. Starch
Figure 3.17 Contents of soluble sugars (glucose, fructose and sucrose) (A) andinsoluble sugar starch (B) (denoted by glucose equivalent) in WT and transgenicplants grown under low temperature (12/5 °C) for around 90 days. The results are themean±SE from two independent experiments.
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Chapter 4
Discussion My master’s project focused on the molecular and functional aspects of the tobacco AOX
gene family. In the first part, research addressing the distribution and interrelationship of
tobacco AOX gene subfamilies was carried out by cloning tobacco AOX2 gene and
comparing it with AOX1 in their DNA/protein sequence features and expression patterns.
In the second part, the function of AOX in stress defense response and carbon
metabolism was investigated by comparing WT with the transgenic lines with altered
expression levels of AOX1 during the low temperature treatment. In this chapter, these
two sub-projects will be discussed separately.
4.1 Cloning and characterization of tobacco AOX2 gene
Different from the well-characterized AOX1 gene, which attracted much attention
because of its stress-induced expression and wide distribution in all different plant
species, the AOX2 gene generally shows tissue and developmental specificity in
expression pattern and is exclusively confined to eudicot plants so far (Costa et al.,
2009). To better understand the interrelationship between these two AOX gene
subfamilies, in the first project I focused my attention on tobacco AOX2 gene. Full-length
coding sequence and flanking untranslated regions of tobacco AOX2 gene were
successfully cloned from WT anther with RACE method and its sequence was analyzed
by bioinformatic tools. Furthermore, the expression pattern of AOX2 was investigated by
RT-PCR and northern blot analysis.
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RACE PCR technology is designed to amplify 5’- and 3’-end of target gene when only
partial sequence of the gene is available. One of the important properties of PowerScript
Reverse Transcriptase I used in the RACE PCR is that the smart sequence is only added
to the complete first-strand cDNA (SMART™ RACE cDNA Amplification Kit User
Manual. 2006), which allows us to amplify the cDNA with maximum amount of 5’-
sequence. However, whether the cloned sequence is full-length cDNA sequence depends
on the quality of RNA template, which was carefully handled and turned out to be
qualified for cloning in this project. Besides the quality of RNA sample, another factor
determining a successful cloning is the specificity of cloning primers. To avoid the
interference caused by other known tobacco AOX genes, primer designing was performed
with the help of sequence alignment between tobacco AOX1 gene and the known region
of AOX2 gene. All primers for AOX2 were designed at relatively low-conserved regions
to avoid non-specific amplification of AOX1 gene in tobacco. However, multiple bands
still appeared in the final products of RACE, which might be due to non-specific
amplification of other members of AOX multigene family or alternative splicing (Kong et
al., 2003; SMART™ RACE cDNA Amplification Kit User Manual. 2006). To further
reduce the false positive results, the stringency of experiment was improved by using
touch-down PCR (Don et al., 1991), raising annealing temperature (Roux, 1995) and
adjusting primer concentration (Robertson et al., 1998). For the multiple bands still
existing, PCR testing with the nested AOX2 gene-specific primers was used to further
verify the RACE products.
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The obtained AOX2 sequence was examined with bioinformatic methods including
DNA/Protein sequence alignment, motif searching and secondary structure identification.
Despite their difference in DNA and protein sequences (particularly at 5’/N-terminal
region), they do share the similar motifs and structures, including mitochondrial targeting
peptides (mTP) and a four-helix bundle containing diiron-binding centers. Noticeably,
unlike most of other AOX proteins, the second amino acid in the antibody-recognition
region of AOX2 is threonine rather than alanine. This alanine was shown to play an
essential role in the recognition by AOX monoclonal antibody (Finnegan et al., 1999).
Therefore, we supposed that AOX2 protein would likely not be detected by the widely-
used AOX antibody. We need to keep this point in mind when analyzing AOX2 gene
expression at protein level.
According to our early analysis (see section 1.5.1), we hypothesized that the tobacco
AOX2 might not contain the critical regulatory cysteine residue at the N-terminus, which
is responsible for the formation of covalently-linked dimer and activation effect by
pyruvate (Vanlerberghe et al., 1998). However, based on the sequence alignment with
tobacco AOX1 I did identify this critical cysteine in AOX2, which indicates that AOX2
should have similar biochemical properties and regulatory mechanisms to AOX1.
Combining the hypothesis that AOX antibody we used may not be able to recognize
AOX2 (See above) with the results we obtained in our previous western blot analysis
(See section 1.5.1), we suppose that the AOX protein we detected with western blot
analysis was probably not AOX2 gene product but some other AOX, which doesn’t
contain the critical cysteine. Admittedly, we cannot rule out the possibility that the
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existence of reduced AOX in MFA-treated cell sample may be simply due to the
incomplete oxidation of protein sample by oxidant diamide. In order to go on pursuing
this so-called “Cys-absent” AOX in tobacco, I did the DNA sequence alignment between
the known Cys-absent AOX genes (in tomato and potato) and regular AOX genes in other
species and designed the degenerate primers at the region which is quite specific for Cys-
absent AOX family. Thereafter, these degenerate primers were used in RT-PCR with the
RNA extracted from MFA-treated cells as template. Although the size of RT-PCR
product was roughly consistent with the predicted size, the sequence analysis on the
cloning product indicated that it belonged to some other gene family (Data not shown).
Therefore, the existence of Cys-absent AOX in tobacco is still questionable. There are
some other possible methods worth a try in the future to clone this “MFA-inducible Cys-
absent AOX”. For example, the highly conserved region across the whole AOX family
could be used to design the degenerate primer for RACE PCR so that no AOX gene will
be missed in cloning. The multiple bands obtained in RACE PCR product may represent
different AOX genes and Cys-absent AOX gene should be included (if it exists). Another
possible way to test the existence of this special AOX gene is to collect the inoxidizable
AOX protein band from MFA-treated sample, digest it into peptides and sequence them
with mass spectrometry (Syka et al., 2004). The peptide sequence could be compared
with known tobacco AOX to verify its identity.
Phylogenetic analysis was also performed on tobacco AOX2 sequence. To more precisely
show its relationship with other AOX genes, only AOX genes with the full-length protein
sequences were selected from the database for constructing the phylogenetic tree. The
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results showed that the gene I cloned was located in the same clade as AOX2 genes in
other species on the phylogenetic tree of the AOX family, which further confirms its
identity as a member of AOX2 gene subfamily.
The expression pattern of tobacco AOX2 gene was investigated with RT-PCR and
northern blot. AOX2 could be amplified by RT-PCR from all different kinds of tissues
except imbibed seeds, indicating its universal expression pattern. However, northern blot
analysis showed that AOX2 could only be clearly detected in bud and anther. This
discrepancy is simply because of the relatively lower sensitivity of northern blot
compared with RT-PCR (Hernandez et al., 2000; Dean et al., 2002). AOX2 expression in
anther was dramatically higher than the other tissues, which implied that AOX2 might
have a function in anther development. Interestingly, Kitashiba et al. (1999) observed the
reduction of pollen viability in transgenic tobacco plants with AOX knock-downed by
transforming an anti-sense fragment of Arabidopsis AOX1a into tobacco. The sequence
alignment between Arabidopsis AOX1a and tobacco AOX1/AOX2 showed that this
Arabidopsis fragment displayed 77% and 70% identity with the corresponding tobacco
AOX1 and AOX2 fragment, respectively (Data not shown). Although we cannot
determine for now which AOX was knock-downed in the transgenic tobacco line they
generated, considering the high transcript level of AOX2 in anther and absence of
abnormal pollen development in our AOX1-silenced transgenic lines (RI9 and RI29), we
assumed that tobacco AOX2 might be the one that was knock-downed, which resulted in
the reduction of pollen viability.
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To test if knockdown of AOX1 is compensated for by up-regulation of AOX2 gene, I
compared the transcript levels of AOX2 in ovary and anther of WT and AOX1-silenced
mutants (RI9 and RI29) with different developmental stages. However, no obvious
difference was detected. Similarly, in Arabidopsis the suppression or overexpression of
AOX1a also has no effect on the expression of the other four AOX genes (including
AOX2) (Umbach et al., 2005). These results indicate that AOX2 may not be able to
respond to the change of AOX1 expression and therefore complement the lack of AOX1.
The expressions of AOX1 and AOX2 are probably controlled independently.
To test the stress response property of AOX2, I also treated the plants/suspension cells
with various chemicals and stresses (e.g. paraquat, salicylic acid, MFA and cold
treatment). AOX1 gene as a control was highly induced by all these treatments, but AOX2
expression could not be induced by any of these treatments (data not shown). Besides, the
lack of induction of AOX2 by MFA is further evidence that the putative MFA-induced
Cys-absent AOX (see section 1.5.1) is probably encoded by a different AOX gene. The
totally different expression pattern between AOX1 and AOX2 gene strongly implied that
their promoter regions might be quite different, which endows them with different
regulatory mechanisms at the transcriptional level. All these results suggest that AOX2
expression only shows the tissue specificity, which is similar to AOX2 genes in other
species (Saisho et al., 2001; Considine et al., 2002). The expression patterns of tobacco
AOX2 analyzed in this study together with our previous understanding concerning
tobacco AOX1 expression further support the hypothesis discussed in the introduction,
that is, AOX1 may be required for plant stress response while AOX2 may function in
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certain developmental events, in particular, anther development in the case of tobacco
AOX2.
Although the transcript level of tobacco AOX2 was always low and it could not be
induced by the several stresses I tested, we cannot reach a conclusion that AOX2 is
useless in plant growth and defense response. Low transcript level does not necessarily
mean low protein and low activity level considering the regulation at post-transcriptional
or post-translational level (Dutilleul et al., 2003). In addition, it is possible that AOX2
may be induced and function under specific conditions that I haven’t tried. Therefore,
more work is needed to further understand the physiological roles of AOX2 and its
interrelationship with other AOX genes.
4.2 Role of AOX in ROS balance and carbon metabolism under cold
stress
In the second part of my project, the roles of tobacco AOX in ROS balance and carbon
metabolism during low temperature stress were investigated by comparing wild-type and
transgenic lines with altered expression levels of AOX1.
No obvious difference in visual phenotype was detected between WT and the other four
transgenic lines throughout the 72 h cold stress or the long-term cold stress, which was
consistent to the previous studies on the response of AOX mutants in other species to
cold stress (Sugie et al., 2006; Watanabe et al., 2008). The only exception so far was
described by Fiorani et al. (2005), in which the AOX anti-sense and AOX-overexpressed
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Arabidopsis displayed smaller versus larger leaf areas and rosettes respectively compared
with WT at the early growth stage in cold stress. Most of AOX-related studies together
with my observation indicate that in most cases the effects caused by underexpression or
overexpression of AOX may be compensated by other mechanisms in plants and
therefore largely change in growth will not appear during cold stress.
4.2.1 ROS gene network in response to different light intensities
All the plants used for this project were grown under a light intensity of 110~130 μmol
m-2 s-1. Based on the northern and western blot analysis, I found that both AOX transcript
and protein were barely detectable in WT (during the normal temperature), which was
quite different from what was observed when plants grew under a relatively higher light
intensity (~400 μmol m-2 s-1) (Amirsadeghi et al., 2006). Apparently, this inconsistency
indicated that AOX expression in tobacco could be induced by the “high light”, which in
fact, is still much lower compared with the natural light (around 1000~2000 μmol m-2 s-1).
A further comparison between “high-light” plants and “low-light” plants brought more
interesting findings. Under high light condition used in the previous research
(Amirsadeghi et al., 2006), two AOX-silenced transgenic lines (RI9 and RI29) displayed
lower ROS level (H2O2 and O2.-) compared with WT. This result was contrary to the
previous hypothesis considering the ROS-dampening function of AOX but was supported
by northern blot analysis, which showed that the expressions of most ROS-scavenging
genes (APX, CAT, MnSOD, Cu/ZnSOD and FeSOD) were higher in AOX-
underexpressed lines than in WT (Amirsadeghi et al., 2006). The resulting higher
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capacity of ROS-scavenging system might complement the lack of AOX in RI9 and RI29
and lead to a lower level of ROS in plant tissues.
In contrast, under the low-light condition with normal temperature (28/22 °C) used in my
experiments, lipid peroxidation levels (an index of oxidative damage) in AOX-silenced
mutants (RI9 and RI29) and AOX-overexpressed mutant (B8) were respectively higher
and lower than in WT (Figure 3.9). Contrary to the observation in the “high light” plants,
almost all the antioxidant genes had no response to the alteration of AOX level and their
transcript levels were similar among different lines. Therefore, the higher lipid
peroxidation level in AOX-silenced mutants was probably because the lack of AOX
increased the ROS generation from mitoETC, which was consistent with the idea that one
important physiological function of AOX is to dampen the production of ROS (Millenaar
et al., 2003).
The contrary results concerning the relationship of AOX level and oxidative damage
between these two experiments strongly indicate that light intensity affects AOX-related
mitochondrial signaling in ROS controlling network. Not much information addressing
this issue is available so far (Giraud et al., 2008). But considering the fact that both AOX
and intensity of light can affect ROS homeostasis in plants (Vanlerberghe et al., 1997a;
Jiao et al., 2004) and the role of ROS as pervasive signaling molecules in plant stress
response (see introduction), I hypothesize that the different level of AOX and light
intensity may result in ROS signals with different strengths, which activate the ROS-
scavenging system (antioxidant genes) to different extents. A further discussion
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concerning the roles of AOX and light intensity in signaling plant antioxidant defense
system will be undertaken later in the section 4.2.3.
4.2.2 The role of AOX in ROS balance under cold stress
One of the most obvious adverse effects of low temperature stress on plants is the
oxidative damage caused by ROS accumulation. Although the major sites responsible for
ROS production during cold stress have not been clearly determined, it was assumed that
the disruption of ETC in chloroplasts and mitochondria may contribute largely to the
ROS accumulation given that the membrane-associated processes like photosynthesis and
respiration are more susceptible to temperature stress than other processes due to the
temperature-sensitive property of membrane (Suzuki et al., 2006). As mentioned above,
the alternative pathway in plant mitochondrion was supposed to reduce the generation of
ROS in mitochondrial ETC by preventing the over-reduction of ETC when Cyt pathway
is depressed under cold stress condition (See introduction). Therefore, it is believed that
AOX may play an important role in cold tolerance. Although some researches have
already addressed this issue (see introduction), the understanding concerning the role of
AOX under cold stress is still far from complete.
In this part of project, I exerted a short-term cold treatment (from 28/22 °C to 12/5 °C) on
both WT and transgenic tobacco lines with altered levels of AOX. Compared with the
long-term stress, the “shift” experiment was believed to be a powerful system to analyze
the function of genes and the nature of signaling pathway responding to the stress
condition.
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To evaluate the oxidative damage caused by cold stress, I firstly measured the in vivo
ROS level in all five lines before and after cold stress. The H2O2 and O2.- levels detected
by in-situ staining assay with DAB and NBT respectively didn’t show any stable pattern
in different lines under control or low temperature condition (data not shown). Given the
low resolution of this in-situ assay due to the highly reactive property and very short half-
life of ROS, I further applied another technique: measurement of lipid peroxidation to
indirectly judge the ROS level during cold treatment. The results indicated that lipid
peroxidation levels (presented by MDA contents) did show different pattern between
different lines. Under the normal condition, as mentioned above, MDA levels in two
AOX-silenced mutants and two AOX-overexpressed mutants (with the possible
exception of B7) were respectively higher and lower than in WT, which meant that the
levels of AOX could affect the total redox state in plant cell even under normal condition
(here we should remember that there is no clear cut distinction between the so-called
“normal condition” and “stress condition”). This result supported the idea that AOX
could reduce the ROS generation and therefore relieve the oxidative damage in plant
tissue. Considering the important role of AOX in dampening the generation of ROS
under stress condition, we hypothesized that after the cold treatment more oxidative
damage should appear in AOX-silenced mutants while less damage should appear in
AOX overexpressors. For two AOX-overexpressors (B7 and B8), the lipid peroxidation
levels after cold stress were both consistently lower compared with WT and the change of
lipid peroxidation in B7 and B8 showed the similar pattern. This result was consistent
with our previous hypothesis that overexpressed AOX should be capable of dampening
the ROS generation more efficiently than WT, which however, needs to be confirmed by
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other experiments. For both WT and RI9, a large increase of lipid peroxidation was
detected after 24h cold treatment followed by a steady decrease in the next two days,
which suggested that these two lines might experience a transient oxidative damage
during the early stage of cold stress and then respond to the damage probably by
activating the ROS-scavenging system. The consistently higher lipid peroxidation levels
in RI9 compared with WT is a good indication that reduced AOX expression can lead to
more ROS generation under stress condition. But interestingly, for another AOX-silenced
mutant RI29 I saw something totally different from all the other four lines (Figure 3.9 C):
the level of lipid peroxidation in RI29 strikingly decreased after cold treatment (24h),
followed by a further steady decline in the next two time points and became even lower
than WT, which meant that less oxidative damage appeared in RI29 during cold stress
and RI29 surprisingly showed a higher capacity to resist oxidative damage compared
with WT, which was contrary to our previous hypothesis.
To explain the different levels of oxidative damage (lipid peroxidation) observed in WT
and the transgenic plants, the transcript levels of key ROS-scavenging genes and AOX-
related genes and the activities of several ROS-scavenging enzymes were analyzed.
Before the cold stress, the transcript levels of most genes in all five lines were similar to
each other (expressions of CAT and FeSOD were a bit higher in two AOX
overexpressors), indicating that under normal condition (low light in this case) altered
levels of AOX had no obvious influence on the regulation of capacity of ROS-scavenging
system. Therefore the levels of oxidative damage (lipid peroxidation) before the cold
shift were inversely proportional with the amount of AOX in plants, which indicated the
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function of AOX in dampening ROS generation. After exposure to cold stress, both the
transcript level and protein level of AOX in WT were increased dramatically, suggesting
that AOX may play an important role in maintaining electron flux to oxygen when cold
stress inhibits other downstream ETC pathway. For genes MnSOD and CAT, their
transcript levels were down-regulated by cold stress, which were in accordance with a
recent report in which the enzyme activities of these two genes were decreased in cold-
treated tobacco plants (Zhang et al., 2009). In contrast, the transcripts of other ROS-
scavenging enzymes (APX, GPX and Cu/ZnSOD and maybe FeSOD) were all increased
after cold stress in both WT and transgenic plants but the increase was much greater in
RI29 and two AOX overexpressors than in WT and RI9. Similarly, although the
transcript levels of CAT were decreased after cold stress, its expressions in RI29, B7 and
B8 were also generally higher than the other two lines.
Corresponding to the northern blot data, the APX activities in RI29 and B7 were also
higher than in WT after the cold shift. The partial inconsistency between APX transcript
and activity data was probably due to the post-transcriptional/post-translational regulation
and the different responses of other APX isozymes to stress condition (Pasqualini et al.,
2007; Watanabe et al., 2008). For total SOD activity, it is difficult to align it with the
transcript pattern of certain SOD gene considering that the total SOD activity I measured
was a combined effect of different SOD isozymes from different compartments. After the
cold treatment, SOD activities in all five lines were steadily increased and generally no
significant difference was detected between different lines. However, we did reveal that
the increases of SOD activities in AOX-silenced mutants and AOX-overexpressed
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mutants were more rapid than in WT after cold treatment (Figure 3.15 B [2]), which
basically corresponded with the conclusion we obtained by the northern blot analysis of
antioxidant system.
As mentioned in the introduction, the antioxidants such as ascorbic acid and glutathione
also make crucial contributions to the detoxification of ROS. The research work
conducted by another student in our lab (Nirusan Rajakulendran) showed that after cold
stress total ascorbate pool and total glutathione pool in all five lines were increased. More
interestingly, the ratio of reduced glutathione to oxidized glutathione (GSH/GSSG) and
perhaps the ratio of reduced ascorbate to dehydroascorbate (ASC/DHA) in RI29 were
higher than WT and RI9 after cold treatment, suggesting the higher ROS-detoxifying
capacity of non-enzymatic antioxidant components in RI29, which corresponded with
what I had observed in lipid peroxidation analysis and northern blot analysis of ROS-
scavenging genes.
Despite certain discrepancies between some of the results discussed above, the analysis
on the plant antioxidant system generally indicated that RI29 and two AOX-
overexpressed transgenic plants displayed stronger activation of antioxidant system than
WT and RI9 after exposure to the cold stress.
The highly induced transcript levels of ROS-scavenging genes in two AOX
overexpressors (B7 and B8) indicated that their consistently low levels of oxidative
damage after cold stress were probably the result of the collaboration of both
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overexpressed AOX and antioxidant system with higher capacity. Interestingly, the
research work by Fiorani et al. (2005) also showed that the expression levels of
antioxidant genes (peroxiredoxin IIC and IIF) in AOX overexpressors were up-regulated
faster than in WT during cold treatment. However, these results were contrary to the
general concept that overexpressed AOX might block the up-regulation of antioxidant
genes by stress considering their complementary relationship in ROS balancing (Maxwell
et al., 1999; Pasqualini et al., 2007). Apparently, the strikingly strong induction of
antioxidant genes in AOX overexpressors suggested the existence of another stress
signaling system besides the ROS-related one. Although we can not determine what
components are involved in this new signaling system and how it works, we did raise two
hypothesis: (1) Considering that AOX is one of the earliest responsive factors to various
stresses (Arnholdt-Schmitt et al., 2006), we suppose that AOX itself may probably act as
a stress sensor and is involved in the signaling network related to the activation of stress
response system (e.g. ROS-scavenging system) under certain stress condition. The
overexpressed AOX during cold stress may strengthen this signal which triggers a
stronger response of antioxidant system. (2) NADPH is crucial for the function of
antioxidant system (e.g. glutathione/ascorbate cycle) (Couee et al., 2006). High AOX
level may compromise the NADPH pool by maintaining the electron transport from
NADPH to oxygen and therefore result in a more oxidized NADPH/NADP+ pool, which
may serve as a positive feedback signal to activate the capacity of antioxidant system.
Admittedly, the results obtained by Fiorani et al. (2005) as well as my project are the
only evidence so far implying that overexpressed AOX may activate the antioxidant
system somehow, which needs further research to reveal the mechanism behind it.
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The greater capacity of ROS-scavenging system in RI29 compared with WT can well
explain why oxidative damage in RI29 after the cold stress kept decreasing even without
the help of AOX to dampen ROS generation. I hypothesize that faster response to the
cold stress and higher expression and activity levels of antioxidant genes in RI29 might
over-compensate for the lack of AOX and result in a higher capacity of cold defense
system than WT. Actually, this so-called “over-compensation” effect concerning AOX
has already been revealed in some other papers. In Amirsadeghi et al. (2006), the higher
transcription levels of antioxidant genes in three transgenic tobacco lines lacking AOX
were detected under high light condition (~400 μmol m-2s-1) compared with WT, which
resulted in a lower O2.- and H2O2 in these transgenic lines. In another paper also
addressing AOX function under cold stress in Arabidopsis (Watanabe et al., 2008), they
found the similar pattern that in AOX-knockout mutant several antioxidant defense genes
were induced and MDA content was lower than WT. In addition to AOX-underexpressed
mutants mentioned above, more recently this “overcompensation” effect was also
discovered in AOX-overexpressed transgenic tobacco treated with ozone (Pasqualini et
al., 2007), in which plant sensitivity to ozone treatment (another kind of oxidative stress)
was paradoxically increased due to its suppression of antioxidant system. All these results
indicate that AOX may be involved in the regulation of antioxidant system in response to
stress condition. Noticeably, besides AOX, the ‘overcompensation effect” was also
discovered in other ROS-controlling network. One recent research studying on
Arabidopsis showed that the plants lacking of both cytosolic and chloroplastic H2O2-
scavenging enzyme APXs strikingly displayed a stronger tolerance to heat stress than
WT, which was probably due to the activation of redundant ROS removal pathway
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(Miller et al., 2007). In spite of the fact that all these reports support the existence of
overcompensation effect, we have to concede that in most cases lack of AOX gene or
ROS-scavenging genes will lead to more ROS generation and oxidative damage (see
introduction). The possible reason for this conflict will be discussed later.
In contrast to RI29, the transcript levels of almost all the ROS-scavenging genes in
another AOX-silenced mutant RI9 were not distinguishable from WT, despite the fact
that APX activities were a bit higher in RI9 than in WT during cold stress. This result
could well explain why RI9 displayed a higher oxidative damage (lipid peroxidation)
than WT, which was due to its reduced level of AOX and poor induction of antioxidant
genes. But it also brings us a new question: why did these two AOX-silenced transgenic
lines (RI9 and RI29) show different defense response to the cold stress? The western blot
analysis on the expression of AOX remind us that compared to RI29, RI9 is a poor AOX
silencer. Is it possible that this “leaky expression” of AOX could change the signal
transduction from mitochondria to other compartments?
4.2.3 “Threshold dose effect” of ROS signal in activating ROS-scavenging system
As mentioned in the introduction, ROS is one of the most important signaling molecules
during stress condition, which we call “ROS signal”. The “ROS signal” could be the ROS
molecule itself (e.g. H2O2) or some other signaling molecules generated by ROS (Rhoads
et al., 2007). So far, the site responsible for the generation of the “ROS signal” has not
been determined yet because of the technical difficulty of measuring the in vivo ROS
production from different compartments (Dutilleul et al., 2003). Considering the
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important signaling roles of mROS and the role of AOX in maintaining mitochondrial
function (see introduction), I assumed that the “ROS signal” we discussed in this AOX-
related project might come from mitochondria. However, we cannot rule out the
possibility of other sources because AOX can also affect the ROS balance in other
compartments (e.g. chloroplast) (see section 1.2.2).
ROS was hypothesized to act as a signal to activate antioxidant system during plant stress
response (Dat et al., 2000). Therefore, we supposed that any factor which could affect
ROS level might have certain influence on regulation of capacity of antioxidant system.
In this project, several ROS-related factors were involved. (1) AOX, dampening ROS
generation from mitochondria by preventing over-reduction of mitoETC. (2) Cold stress,
causing the malfunction of different compartments and the consequential ROS generation
(See introduction). (3) Light stress, causing ROS generation from photo-inhibited
photosynthetic system (Murata et al., 2007). Accordingly, we assume that AOX and
environmental stresses can negatively and positively regulate the intensity of ROS
signaling pathway, respectively. In other words, both the lack of AOX and environmental
stress (cold or light) contribute to ROS generation and may strengthen the intensity of
ROS signal. When plants with different levels of AOX expose to different environmental
conditions (normal temperature vs. low temperature or low light vs. high light), ROS-
scavenging system is supposed to be activated to different degrees (Figure 4.1). During
the normal temperature condition, although more ROS signal is generated in RI9 and
RI29 compared with WT because of their underexpressed AOX, the ROS signal is not
strong enough to activate ROS-scavenging system, which we consider stays at “silent”
status (shown by the green area [level 1] in Figure 4.1 A). Once exposed to low
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temperature stress, ROS signals in all different lines are increased. For WT and RI9, the
ROS signal exceeds the first threshold between level 1 and level 2, activating a higher
capacity of defense system than control condition (shown by the yellow area [level 2] in
Figure 4.1 A). In this case, the level of ROS signal in RI9 is higher than WT because of
its underexpressed AOX but is still not strong enough to exceed the second threshold
between level 2 and level 3. However, for the other AOX silenced mutant RI29, an even
stronger ROS signal is accumulated due to its complete silence of AOX, which activates
the capacity of defense system to a higher level than WT and RI9 (shown by orange area
[level 3] in Figure 4.1 A). Interestingly, when I compared the plant responses to different
light intensities (see section 4.2.1), I found that this model could be applied to the light
stress as well (Figure 4.1 B), in which small modifications were made considering that
the involvement of light stress rather than cold stress might change the intensity of ROS
signal as well as the threshold level needed for activation of defense system.
From the analysis above, I assume that the level of ROS signal may have a “threshold
dose effect” on activating ROS-scavenging system, which means that stronger ROS
signal may lead to higher capacity of defense system when its intensity exceeds certain
threshold.
Noticeably, the model proposed above may also be used to interpret the conflicting
results obtained in some previous AOX-related papers. The research work by Fiorani et
al. (2005) showed that under the low temperature growth condition the oxidative damage
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A
B
Figure 4.1 “Threshold dose effect’ model of ROS signal in activating defense systemduring the abiotic stresses: low temperature stress (A) and light stress (B). The “Basallevel” means the intensity of ROS signal before stress. The different intensities ofROS signals in WT and AOX-silenced transgenic lines under various conditions weredenoted by the different heights of horizontal lines. The capacity of defense systemwas differentiated by level 1, 2 and 3, shown with green, yellow and orange colors,respectively.
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(presented by MDA level) in AOX antisense Arabidopsis lines were higher than in WT
and the expression level of several antioxidant genes could not be distinguishable
between these two lines. On the contrary, Watanabe et al. (2008) reported that under cold
stress the MDA level in AOX knock-out Arabidopsis mutant was actually lower than in
WT and capacity of antioxidant system in AOX knock-out mutant was higher than in
WT. Comparison of these two independently-generated AOX knock-out lines
interestingly showed that the remaining CN-resistant respiration capacities in the former
transgenic line (around 27%) was higher than the latter one (almost undetectable), which
was coincidentally similar to the two AOX silenced mutants I used in this project (RI9 vs.
RI29), indicating that under certain stress condition completely silenced AOX is able to
trigger the ‘overcompensation effect” of antioxidant system through ROS signaling
system while incomplete silence of AOX may have no effect or little effect on the
activation of antioxidant system because of its weak ROS signal. These similar results in
both tobacco and Arabidopsis suggest that the ‘dose effect” of ROS signal related to AOX
gene expression in activating defense system may be an universal mechanism for plants
to respond to the stress condition, but this needs to be further confirmed.
Another question we would like to ask is if this “dose effect” of ROS signal exists in
other ROS-controlling system. Interestingly, Rizhsky et al. (2002) found that plants
lacking of both APX and CAT displayed stronger capacity of tolerance to oxidative stress
compared with WT or the single mutants lacking APX or CAT. Correspondingly, the
defense system in the double mutant was activated while in the single mutant was not.
The authors suggested that this was probably because the signaling pathways activated by
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the lack of APX or CAT were integrated in the double mutant, which lead to a different
outcome. Nonetheless, based on the model we made in this project, another possible
explanation for the different stress tolerance between double mutant and single mutants
could be that more ROS signal was accumulated in the double mutant and in turn
activated a stronger capacity of antioxidant system to complement the lack of two
antioxidant genes, while the ROS signal activated by the lack of single antioxidant gene
was not high enough to exceed the threshold needed for the activation of antioxidant
system, which lead to a weaker tolerance to the oxidative stress.
From the discussion above, I assume that the communication among the different parts of
the ROS-related network and the balance between ROS production and ROS scavenging
greatly rely on the “ROS signal”. Either the disturbance of this ROS network (e.g.
knockdown of AOX or ROS-scavenging genes) or the stress condition (e.g. cold or high
light) can enhance the ROS signal, but whether or not the defense system will be
activated and to what extent the system will be activated depend on the intensity of the
ROS signal. In some cases (weak ROS signal), the response of defense system cannot
compensate for the disturbance of the ROS-related network and therefore lead to more
oxidative damage; while in some other cases (strong ROS signal), the enhanced defense
system can overcompensate for the disturbance and endow the plants with stronger
tolerance to the stress.
4.2.4 AOX is involved in inter-compartment signaling network
Interestingly, from the gene expression data I obtained above, it seems that during cold
stress altered AOX expression levels had more impact on the expression of the proteins
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outside of mitochondria (APx, GPx, Cu/ZnSOD in cytosol, FeSOD in chloroplast and
PTOX in plastid) than the proteins within the mitochondria (MnSOD and COX6b).
Admittedly, this phenomenon was only based on the analysis of a relatively small sample
of genes. However, a similar conclusion was also made according to the microarray
experiments in Umbach et al. (2005) and Giraud et al. (2008), suggesting that other
compartments might be more susceptible to the altered levels of AOX than mitochondria.
Among all the genes being affected by AOX levels during cold stress, PTOX attracted
special attention given its close relationship with AOX. As mentioned in introduction,
PTOX is a functional analog of AOX in plastids, which was believed to be capable of
removing excess electrons from photosynthetic electron transport during stress condition
(Peltier et al., 2002). The observation in this project that PTOX expression was induced
by cold stress suggested its function of protecting chloroplast metabolism. Furthermore,
in the transgenic plant with underexpressed AOX (RI29), the transcript level of PTOX
was increased more dramatically than WT after cold treatment, supporting the hypothesis
that these two genes work in a coordinated manner (Amirsadeghi et al., 2006; Moseley et
al., 2006).
Based on these observations together with the discussion above, we assume that AOX
may be able to influence the expressions of antioxidant genes in other compartments
through either ROS-based signaling pathway (in the case of RI9 and RI29) or non-ROS-
based signaling pathway (in the case of B7 and B8) to respond to the environmental
stress.
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4.2.5 AOX facilitates the accumulation of soluble sugars under cold stress
To understand the function of AOX in carbon metabolism during cold stress, the amounts
of major carbohydrates including monosaccharides (glucose and fructose), disaccharide
(sucrose) and starch were measured with enzymatic cycling assay and compared among
the different lines. AOX was considered to maintain the electron flux to oxygen and
therefore keep TCA cycle operating under cold stress when the function of Cyt pathway
was impaired (see introduction). Accordingly, we originally assumed that if the
production rates of carbohydrates by photosynthesis in WT and transgenic lines were
similar to each other under cold stress, the plants with higher level of AOX should have
smaller pool sizes of carbohydrates because more carbohydrates should be consumed by
the respiration with higher capacity due to the existence of more AOX. Actually, the
research by Sieger et al. (2005) working on the tobacco suspension cell system indicated
that under nutrient-limited condition AOX anti-sense line (AS8) contained a larger pool
size of carbohydrates than WT, which supported the idea that AOX could balance
between the carbon metabolism and electron transport and relieve the build-up of
carbohydrate pool when Cyt pathway is inhibited. However, my results were contrary to
what we expected: the pool sizes of monosaccharides (glucose and fructose) and probably
disaccharides (sucrose) in the plants with overexpressed AOX (B7 and B8) or suppressed
AOX (RI9 and RI29) were respectively larger or smaller than in WT after cold stress.
The plants with the long-term cold treatment also showed the similar pattern.
One possible explanation for this result may be that the prerequisite for our previous
hypothesis (the production rates of carbohydrates by photosynthesis in different
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genotypes were similar) is incorrect. Photosynthesis is the main source for carbohydrate
generation. Although the breakdown of starch can partially contribute to the
accumulation of soluble sugars during cold stress, it has been documented that the plants
with greater photosynthetic efficiency did display more remarkable accumulation of
soluble sugars (Keller et al. 1995). On the other hand, it has been reported that the
inhibition or knock-down of AOX could lower the photosynthetic rate and impair carbon
assimilation (Yoshida et al., 2006; Padmasree et al., 2001; Giraud et al., 2008). This is
probably because AOX is able to consume the excess reducing power generated by
photosynthesis which may otherwise lead to the photo-inhibition of photosynthetic
process (Yoshida et al., 2007). Taken together, I supposed that higher level of AOX
could protect photosynthesis more efficiently and therefore more carbohydrates could be
produced. Although some extra monosaccharides were consumed by respiration in the
plants with higher level of AOX, the reduction of monosaccharides by this process might
be complemented by a larger influx of carbohydrates from photosynthesis, which taken
together resulted in a larger size of monosaccharide pool (Figure 4.2). Based on the
results I obtained and the model I constructed in Figure 4.2, it could be further predicted
that even though in WT AOX was already greatly accumulated after cold stress,
protection of chloroplast function could be better improved by increasing AOX levels
even further (AOX-overexpressed lines).
Another possible interpretation for the correlation between AOX levels and contents of
soluble sugars is that the change of respiration due to the altered levels of AOX might be
buffered by other ATP-uncoupling pathways such as NAD(P)H dehydrogenases (NDs)
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and uncoupling protein (UCP) (see introduction). Watanabe et al. (2008) found that in
Arabidopsis the expression of NDB2 and UCP1 were induced by the lack of AOX and
correspondingly the total respiration rate in AOX-knockout mutants was actually higher
than WT during cold stress, which indicated that the lack of AOX could be
complemented by other ATP-uncoupling pathways and this compensation effect might
lead to a higher respiration rate. However, these two possible explanations still need to be
further confirmed by measuring photosynthetic rate and respiratory rate of WT and AOX
transgenic lines.
- 114 -
Figure 4.2 A working model describing the possible interrelationship betweenalternative pathway and monosaccharide pool under the cold stress. In brief, AOX caneither facilitate the accumulation of monosaccharide by protecting the function ofphotosynthesis or impair its accumulation by maintaining the electron flow and therespiration rate during cold stress. Based on the results we obtained, it seems that theformer effect probably exceeds the latter effect, which leads to a larger pool size ofmonosaccharides in the plants with higher level of AOX.
- 115 -
4.2.6 AOX plays crucial roles in both stress response and metabolic homeostasis
From what we observed in this project and what we discussed above, we conclude that
AOX in mitochondria does play crucial yet complicated roles in both ROS balance and
carbon metabolism under cold stress (Figure 4.3).
The roles of AOX in ROS balance could be divided to three parts according to the data I
obtained in the project: Firstly, as what we have already known, AOX is capable of
dampening the generation of ROS from mitoETC when ETC is over-reduced under
certain stress condition (e.g. low temperature) and probably helping consume excess
reducing power from other compartments which may otherwise cause the ROS
generation. Secondly, AOX is involved in ROS-related signaling pathway. It can
influence the capacity of plant defense response (ROS-scavenging system) by negatively
regulating the intensity of ROS signal, which is able to activate the defense system by a
“threshold dose effect”. Thirdly, the strikingly stronger induction of defense response in
AOX overexpressors compared with WT during cold stress suggests that the excess AOX
may activate another unknown signaling pathway (different from the above ROS-related
signaling pathway) to enhance the capacity of plant defense response.
Different from what we hypothesized before, the role of AOX in carbon metabolism
during the cold stress is more than just a carbon consumer to bring down the
carbohydrate levels (particularly glucose and fructose levels) through respiration by
maintaining electron flow to oxygen. Another hypothesized role of AOX in carbon
metabolism intertwines with its function in balancing the cellular ROS level, namely that
- 116 -
AOX may protect photosynthesis from over-reduction and the consequential
photoinhibition caused by the cold stress and therefore maintain the production of
carbohydrates. The result of combining these two opposite effects of AOX on
carbohydrate balance during cold stress turns out to be that AOX has a positive impact on
monosaccharides (perhaps disaccharide) accumulation in response to cold stress.
In summary, we conclude that AOX in mitochondrion can serve as both a metabolic
modulator and a signaling modulator at the whole-cell level, maintaining the metabolic
and signaling homeostasis in plants under cold stress.
- 117 -
Figure 4.3 Model summarizing the relationship between AOX and ROS balance / carbon metabolism during the exposure to cold stress. Low temperature results in theaccumulation of soluble sugars (1), inhibition of photosynthesis (2), ROSaccumulation (3) and induction of ROS-related genes (AOX [4] and ROS-scavenging enzymes [5]). Besides the antioxidant system which scavenges the excess ROSgenerated during cold stress (6), AOX can also control the cellular ROS level by dampening the production of ROS from MitoETC (7) or by indirectly reducing ROSgenerated from some other sources. In addition, AOX may negatively regulate the intensity of ROS signal, which is capable of activating stress defense system (8) by a “threshold dose effect”. Noticeably, we found that the excess AOX could activate the defense system through certain unknown signaling pathway (9) rather than the ROS-related one (8). On the other hand, AOX is also involved in carbohydrate balance. Bymaintaining electron transport during cold stress, AOX can enhance the consumption of carbohydrates (10). Meanwhile, another role of AOX is to protect the photosyntheticprocess from photoinhibition (11), which may facilitate the production of carbohydrates (12).
- 118 -
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