STUDIES ON THE ROLES AND REGULATION OF TWO GLYOXYLATE
REDUCTASES IN PLANTS
A Thesis
Presented to
The Faculty of Graduate Studies
of
The University of Guelph
by
WENDY LYNNE ALLAN
In partial fulfilment of requirements
for the degree of
Doctor of Philosophy
December, 2008
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While these forms may be included Bien que ces formulaires in the document page count, aient inclus dans la pagination, their removal does not represent il n'y aura aucun contenu manquant. any loss of content from the thesis. Canada ABSTRACT
STUDIES ON THE ROLES AND REGULATION OF TWO GLYOXYLATE REDUCTASES IN PLANTS
Wendy Lynne Allan Advisor: University of Guelph, 2008 Dr. Barry J. Shelp
Succinic semialdehyde (SSA) is a mitochondrial intermediate in the metabolism of
GABA, whereas glyoxylate is a peroxisomal metabolite generated from photorespiratory glycolate. Recent evidence indicates that distinct cytosolic and plastidial glyoxylate reductase isoforms from Arabidopsis thaliana (L.) Heynh (AfGLYRl and AfGLYR2, respectively) catalyse the in vitro NADPH-dependent conversion of SSA to gamma- hydroxybutyrate (GHB) and glyoxylate to glycolate. In this thesis, recombinant
Arabidopsis GLYR1 was demonstrated to simultaneously catalyze both reactions in vitro.
Time-course experiments revealed that GHB accumulated in leaves of Arabidopsis subjected to salinity, drought, submergence, cold or heat stress, and tobacco (Nicotiana tabacum L.) subjected to submergence. This was generally accompanied by higher
GABA and Ala levels, higher NADPH/NADP+ ratios, and lower Glu levels. Furthermore, expression analysis of Arabidopsis revealed that the relative abundance of GLYR1 transcript was enhanced under salinity, drought, submergence, cold and heat, as was the
GLYR2 transcript under cold and heat. Other time-course experiments revealed that Arabidopsis glyrl and glyr2 knockout mutants, unlike wild-type (wt) plants, did not have elevated levels of GLYR1 or GLYR2 transcript, GHB and NADPH in rosette leaves of plants subjected to submergence or low CO2 conditions (75 versus 380 [xmol mol"1 at
21% O2). Glycolate, glyoxylate and succinate also accumulated in glyrl and glyrl under low CO2, whereas Gly did not. Manipulation of redox balance via the use of Arabidopsis
NAD kinase (NADK) mutants (nadkl and nadk3 knockout mutants, NADK1 overexpressor, and NADK2 underexpressor) altered the plant response to submergence.
Thus, it can be concluded that: 1) GHB accumulation is a general response to abiotic stress and is regulated by both biochemical and transcriptional processes; and 2) GLYR activity is involved in the detoxification of both SSA and glyoxylate, and the recycling of phosphorylated pyridine nucleotides, thereby linking GABA metabolism and photorespiration to redox homeostasis. ACKNOWLEDGEMENTS
Many people have contributed to the development of this thesis. Firstly, I would like to give thanks to Dr. Barry Shelp for his unwavering support, guidance and expertise during my tenure as a PhD student in his laboratory. He largely contributed to my enthusiasm and determination to succeed and has been a continuing source of inspiration to me. I would also like to thank my committee members Dr. David Wolyn,
Dr. Chris Hall, Dr. Alan Bown and Dr. Barry Micallef for their advice, expertise and on-going support with a special thanks to Dr. Wolyn for his thoughtful comments on experimental design and statistical analysis. As well, I extend my gratitude to Dr.
Katerina Jordon and to Dr. Bernie Grodinski for their generosity in allowing me the use of their lab equipment from time to time.
I am also appreciative of the advice and help of greenhouse personnel and to the technical expertise and advice of many technicians and postdocs, including Alice
Barker, Rodger Tschanz, Owen Van Cauwenberghe, Amina Mahkmoudova, and
Gordon Hoover.
I had many years of enjoyment and laughs with my labmates and cohorts and I am very much appreciative of their technical help, scientific debates, and how they regularly boosted my morale. I want to especially thank Shawn Clark and Jeff Simpson for their support and the extra help they provided me when I really needed it.
Lastly, I would like to extend my love and gratitude to my husband and son,
George who have endured my long working hours, and my stress-filled moments over the years. You have both been patient and it is now finally finished.
i TABLE OF CONTENTS
ACKNOWLEDGEMENTS i
LIST OF TABLES vii
LIST OF FIGURES viii
LIST OF ABBREVIATIONS xi
CHAPTER 1 - INTRODUCTION 1
CHAPTER 2 - LITERATURE REVIEW 7
2.1. Metabolism and Toxicity of Aldehydes 7
2.1.1. Plant aldehydes 9
2.1.1.1. Detoxification of plant aldehydes 10
2.1.1.2. Aldehyde Dehydrogenases 11
2.1.1.3. Aldehyde Reductases 15
2.2. GABA Metabolism 17
2.2.1. Regulation 20
2.2.1.1 Glu decarboxylase (GAD) 20
2.2.1.2. y-aminobutyrate transaminase (GABA-T) 23
2.2.1.3. Succinic semialdehyde dehydrogenase (SSADH) 25
2.2.1.4. Glyoxylate/succinate semialdehyde reductase (GLYR) 26
2.2.2. Potential Roles 28
2.3. Pyridine Nucleotide Metabolism and Distribution 36
u 2.3.1. NAD+/NADH 37
2.3.2. NADP+/NADPH 41
2.4. Glyoxylate and Photorespiration 45
2.5 Stress and Photorespiration 50
CHAPTER 3 - y-HYDROXYBUTYRATE ACCUMULATION IN ARABIDOPSIS
AND TOBACCO PLANTS IS A GENERAL RESPONSE TO ABIOTIC STRESS:
PUTATIVE REGULATION BY REDOX BALANCE AND GLYOXYLATE
ISOFORMS 53
3.1. Contribution 53
3.2. Acknowledgements 53
3.3. Abstract 54
3.4. Introduction 55
3.5. Materials and Methods 56
3.5.1. Growth of plant material 56
3.5.2 Arabidopsis exposure to drought, salinity, submergence, cold or heat 57
3.5.3. Tobacco submergence and recovery 58
3.5.4. Extraction and analysis of GHB, amino acids, and phosphorylated pyridine nucleotides 59
3.5.5 Expression and analysis of GLYR1 and GLYR2 in Arabidopsis 59
3.6. Results 60
in 3.6.1. Stress responsiveness of metabolites and phosphorylated pyridine nucleotides in Arabidopsis and tobacco 61
3.6.2 Recovery of GHB and related metabolites, and phosphorylated pyridine nucleotides in tobacco after removal from submergence 64
3.6.3. Stress responsiveness of GLYR transcripts in Arabidopsis 66
3.7. Discussion 66
3.7.1. Impact of abiotic stress on GAB A, Ala and redox levels 66
3.7.2 GHB accumulation is a general response to abiotic stress 72
3.7.3 Recovery of metabolite and redox levels after exposure to abiotic stress 74
3.7.4 Role of GLYR isoforms in detoxification of SSA and glyoxylate 75
3.7.5 Summary 77
CHAPTER 4 - REDOX BALANCE AND DETOXIFICATION OF SUCCINIC
SEMIALDEHYDE AND GLYOXYLATE IN ARABIDOPSIS PLANTS SUBJECTED
TO ENVIRONMENTAL CONDITIONS THAT PROMOTE y-AMINOBUTYRATE
METABOLISM OR PHOTORESPIRATION 79
4.1. Contributions 79
4.2. Acknowledgements 80
4.3. Abstract 80
4.4. Introduction 81
4.5. Materials and Methods 83
iv 4.5.1. In vitro assay of simultaneous substrate utilization by recombinant
Arabidopsis GLYR1 83
4.5.2. Generation and description of transgenic Arabidopsis lines 84
4.5.3. Growth of Arabidopsis 87
4.5.4 Analysis of GLYR and NADK expression in Arabidopsis 87
4.5.5. Measurement of enzyme activity in wild-type Arabidopsis and mutant lines 88
4.5.6. Exposure of wild-type Arabidopsis and mutant lines to submergence 90
4.5.7. Exposure of wild-type Arabidopsis and glyr mutants to photorespiratory conditions 91
4.5.8 Extraction and analysis of GHB, amino acids, organic acids and pyridine nucleotides 92
4.6. Results 93
4.6.1 Simultaneous utilization of succinic semialdehyde and glyoxylate by recombinant GLYR 1 93
4.6.2 GLYR activity in cell-free extracts from leaves of wild-type Arabidopsis and glyr mutants 95
4.6.3 Response of wild-type Arabidopsis and glyr mutants to submergence 97
4.6.4 Response of wild-type Arabidopsis and NADK mutants to submergence.... 103
4.6.5. Response of wild-type Arabidopsis and glyr mutants to varying CO2/O2 ratios
110
4.7. Discussion 120
v 4.7.1. Redox homeostasis and detoxification of succinic semialdehyde in
Arabidopsis plants exposed to submergence 120
4.7.2. Redox homeostasis and detoxification of succinic semialdehyde and glyoxylate in Arabidopsis plants exposed to photorespiratory conditions 122
4.7.3. Summary 125
CHAPTER 5 - SUMMARY AND FUTURE PROSPECTS 127
5.1 Hypotheses Revisited 127
5.2 Future Prospects 135
CHAPTER 6 - LITERATURE CITED 138
vi OF TABLES
Page Biotic and abiotic stresses stimulating GABA and GHB
accumulation 32
4.1. Total glyoxylate-dependent GLYR activity in cell-free extracts of 96 rosette leaves from wt Arabidopsis and glyr mutants
4.2 NADK activities in cell-free extracts of tissue culture-grown wt
Arabidopsis and NADK mutants t nA
vii LIST OF FIGURES
Figures Page
2.1. Alternative pathways for GABA metabolism via succinic semialdehyde
(A). Glyoxylate reductase reaction (B) 19
3.1. Response of metabolites in mature rosette leaves of Arabidopsis plants (A)
and mature leaves of tobacco plants (B) subjected to salinity, drought,
submergence, cold or heat 63
3.2. Response of phosphorylated pyridine nucleotides in mature rosette leaves
of Arabidopsis plants (A) and mature leaves of tobacco plants (B)
subjected to cold, heat or submergence. 65
3.3. Recovery of GHB and related metabolites (A), as well as phosphorylated
pyridine nucleotides (B), in mature leaves of tobacco plants after being
subjected to submergence 68
3.4. Response of GLYR transcripts in mature rosette leaves of Arabidopsis
plants subjected to salinity, drought, submergence, cold or
heat 69
4.1. In vitro production of glycolate and GHB by recombinant GLYR1
incubated with subsaturating glyoxylate and SSA and saturating
NADPH 94
4.2. Response of GLYR transcripts in mature rosette leaves of wild-type
Arabidopsis and glyrl (A) and glyr2 (B) mutants subjected to
Vlll submergence 98
Response of pyridine nucleotides in mature rosette leaves of wild-type
Arabidopsis and glyrl (A) and glyr2 (B) mutants subjected to
submergence 100
Response of metabolites in mature rosette leaves of wild-type Arabidopsis
and glyrl (A) and glyr2 (B) mutants subjected to
submergence 102
Relative abundance of NADK1, NADK2 or NADK3 transcripts in wild-
type Arabidopsis and various NADK mutants 104
Response of pyridine nucleotides in mature rosette leaves of wild-type
Arabidopsis and various NADK mutants subjected to
submergence 107
Response of metabolites in mature rosette leaves of wild-type Arabidopsis
and various NADK mutants subjected to submergence... 108
Response of GLYR1 and GLYR2 transcripts in mature rosette leaves of
wild-type Arabidopsis and glyrl (A) and glyrl (B) mutants subjected to
various CO2 levels 112
Response of pyridine nucleotides in mature rosette leaves of wild-type
Arabidopsis and glyrl (A) and glyrl (B) mutants subjected to various CO2
levels 114
IX 4.10. Response of amino acids and GHB in mature rosette leaves of wild-type
Arabidopsis and glyrl (A) and glyrl (B) mutants subjected to various CO2
levels 116
4.11. Response of organic acids in mature rosette leaves of wild-type
Arabidopsis and glyrl (A) and glyr2 (B) mutants subjected to various CO2
levels 118
5.1 Subcellular model for the interaction between GABA and photorespiratory
gloxylate metabolism 131
x List of Abbreviations
AAP3 amino acid permease 3
AAAP3 amino acid/auxin transporter
ABA abscisic acid
ADH alcohol dehydrogenase
ALDH aldehyde dehydrogenase
ADP adenosine diphosphate
AKR aldo-keto reductase
AS Anti-sense
ATF amino acid transporter
ATP adenosine triphosphate
Ca2+ calcium
CaM calmodulin
Ci internal leaf carbon dioxide concentration
CMS cytoplasmic male sterility
CoA coenyzme A d day(s)
DM dry mass
Fd-GOGAT ferredoxin-dependent glutamine:2-oxoglutarate aminotransferase
FM fresh mass
GABA y-aminobutyrate
GABA-T y-aminobutyrate transaminase
GAD glutamate decarboxylase
xi GAT1 gamma-aminobutyrate transporter 1
GDC glycine decarboxylase
GFP green fusion protein
GGAT glutamate: glyoxylate aminotransferase
GHB y-hydroxybutyrate
GLYR glyoxylate reductase
GS glutamine synthetase h hours
HHE 4-hydroxy-(2E)-hexenal
HNE 4-hydroxy-(2E)-nonenal
KO knock-out
MDA malondialdehyde
NAD(H) oxidized and reduced forms of nicotinamide adenine dinucleotide
NADK nicotinamide adenine dinucleotide kinase
NADP(H) oxidized and reduced forms of nicotinamide adenine dinucleotide phosphate
NaMN nicotinic acid mononucleotide
OE overexpressor
OPPP oxidative pentose phosphate pathway
PARP poly(ADP-ribose) polymerase
PCR polymerase chain reaction
PMSF phenylmethylsulphonyl fluoride
xii ProTl proline transporter 1
PSI photosystem I
PSII photosystem II
PUFA poly-unsaturated fatty acids
QPRT quinolinic acid phosphoribosyl transferase
ROS reactive oxygen species
RT-PCR reverse transcription-polymerase chain reaction
RUBISCO ribulose-bisphosphate carboxylase/oxygenase
RuBP ribulose-bisphosphate s seconds
SE standard error
Ser serine
SSA succinic semialdehyde
SSADH succinic semialdehyde dehydrogenase
SGAT serine:glyoxylate aminotransferase
SSR succinic semialdehyde reductase
TAA total amino acids
TCA tricarboxylic acid
THF tetrahydrofolate
UV ultraviolet
Wt wild-type
xm CHAPTER 1 - Introduction
y-Aminobutyrate (GABA), a non-protein amino acid, and y-hydroxybutyrate (GHB), a short-chain fatty acid that closely resembles GABA, are found in virtually all prokaryotic and eukaryotic organisms. They are endogenous constituents of the mammalian nervous system, wherein GABA plays a role in neural transmission and development, and functions through interactions with specialized receptors (GABAA,
GABAB, GABAC) and transporters, and GHB serves as a neurotransmitter or neuromodulator postulated to act via a GABAB receptor or an independent GHB- specific receptor (see review by Fait et al, 2008). When administered, GABA does not cross the blood-brain barrier, whereas GHB does so with ease, penetrating the brain and producing diverse neuropharmacological and neurophysiological effects. For further details on the roles of GABA and GHB in animals, refer to reviews by Mamalek,
(1989) and Fait et al, (2008).
A large number of studies have reported the accumulation of GABA in plant tissues and transport fluids in response to many biotic and abiotic stresses (e.g., Bown and
Shelp 1989; Shelp et al, 1999; Kinnersley and Turano 2000; Bouche et al, 2004;
Bown et al, 2006). These include temperature shock, oxygen deficiency, cytosolic acidification, water stress and UV stress, as well as mechanical stimulation and damage, which are commonly associated with the activities of invertebrate pests during foraging and feeding. In some cases, the response is rapid, often within seconds, suggesting that the biochemical control, rather than transcriptional control, is involved, although there
1 is some evidence for the induction of Glu decarboxylase (GAD) and GAB A transaminase (GAB A-T) in the longer term.
Evidence for the occurrence and accumulation of GHB in plants recently became available. For example, oxygen deficiency increases GHB concentrations from about 10 to 155 nmol g"1 fresh mass (FM) in soybean sprouts, 273 to 739 nmol g"1 dry mass
(DM) in green tea leaves (Allan et al, 2003a), and 42 to 70 nmol g"1 FM in Arabidopsis leaves (Breitkreuz et al, 2003). Oxygen deficiency should increase the cellular
NADH:NAD+ ratio and decrease the adenylate energy charge, thereby inhibiting
SSADH activity and diverting carbon from succinate (Shelp et al, 1995, 1999; Busch et al, 2000). Subsequent work revealed that: 1) ssadh mutant Arabidopsis plants, grown under high UV light, have five times the normal level of GHB and high levels of reactive oxygen species (ROS) (Fait et al, 2005); and, 2) the pattern of GHB in cold- acclimated Arabidopsis plants is consistent with the rise and fall of GABA (Kaplan et al, 2007). Together, these data suggest that the accumulation of GHB in plants, as well as GABA, is a general response to abiotic stress.
In plants, GABA is derived primarily via the H+-consuming oc-decarboxylation of
Glu in an irreversible reaction catalyzed by cytosolic-localized GAD that has an acidic pH optimum (Shelp et al, 1999). While increasing cytosolic H+ concentration can result in GABA accumulation, there is abundant evidence for a mechanism involving Ca2+- dependent binding of calmodulin (CaM) to GAD proteins at neutral pH, thereby relieving the enzyme from autoinhibition and stimulating enzymatic activity (reviewed by Shelp et al, 1999). Thus, CaM links GABA accumulation and increasing cytosolic
Ca2+ that typically accompanies stress. Research has identified multiple GAD genes
2 from Petunia, tomato, tobacco, Arabidopsis and rice, and differential organ localization of two isoforms in both Arabidopsis and tobacco (Shelp et ah, 1999; Yevtushenko et al, 2003; Akama and Takaiwa, 2007; Miyashita and Good, 2008), implying that they may have specific functions. For example, GAD1 is predominantly expressed in roots, whereas GAD2 expression is evident in all organs; expression of the other three GAD genes is weak (Miyashita and Good, 2008). Phenotypic analysis of gadl knockout (KO) mutants revealed that GABA levels in roots are dramatically lower than in wild-type
(wt) roots, and that heat-induced GABA accumulation is prevented in gadl mutants
(Bouche and Fromm, 2004). Moreover, antisense suppression of GAD results in the accumulation of Glu in transgenic tomato fruit (Kisaka et al, 2006). Transcriptional induction of one or more GAD forms is often observed in response to low oxygen, water deficit, salinity or Agrobacterium infection (Klok et al, 2002; Deeken et al,
2006; Cramer et al, 2007; Pasentsis et al, 2007; Miyashita and Good, 2008).
GABA is then transaminated to succinic semialdehyde (SSA) via a mitochondrial- localized GABA transaminase (GABA-T) that is probably reversible (Van
Cauwenberghe and Shelp, 1999; Van Cauwenberghe et al, 2002). Both pyruvate- and
2-oxoglutarate-dependent activities are found in crude tobacco plant extracts; however, to date only the gene for pyruvate-dependent activity (GABA-T1) in Arabidopsis has been identified (Van Cauwenberghe et al, 2002). Highly homologous proteins in pepper and rice have been identified (Ansari et al, 2005; Wu et al, 2006), although protein function has not been examined. The expression of GABA-T1 is detected in all
Arabidopsis organs and the vegetative phenotype appears normal, but a gaba-tl mutant lacks a GABA gradient from the stigma to the embryo sac and pollen tube growth is
3 misdirected, thereby causing a reduced-seed phenotype, and GABA-T activity is decreased to negligible levels in both shoots and roots and GABA accumulates in roots
(Palanivelu et al, 2003; Miyashita and Good, 2008). There is typically significant transcriptional change in GABA-T1 under low oxygen, water deficit and salinity (Klok et al, 2002; Cramer et al, 2007), but not always (Miyashita and Good, 2007).
SSA dehydrogenase (SSADH) catalyzes the irreversible, NAD-dependent oxidation of SSA to succinate in the mitochondrion. The enzyme is competitively inhibited by NADH and AMP, non-competitively inhibited by ATP, and inhibited by
ADP via both competitive and non-competitive means (Busch et al,. 1999). SSADH occurs as a single-copy gene in Arabidopsis and ssadh mutants contain elevated levels of ROS, are hypersensitive to heat and light stress, and have a stunted and necrotic phenotype (Bouche et al, 2003).
Other research suggests an additional mechanism for the metabolism of SSA, which involves its reduction to GHB. A number of strategies, including complementation of a SSADH-deficient yeast mutant with an Arabidopsis cDNA library, recombinant expression in Escherichia coli, and transient expression in tobacco
BY-2 cells, were used to identify two highly homologous proteins (designated hereinafter as Arabidopsis glyoxylate reductases 1 and 2 or AfGLYRl and A/GLYR2) that catalyze the conversion of both SSA to GHB and glyoxylate to glycolate via an essentially irreversible, NADPH-based ordered Bi Bi mechanism, although they are located in different cellular compartments (cytosol, plastid) (Breitkreuz et al, 2003;
Hoover et al, 2007a, b; Simpson et al, 2008). NADP+ is an effective competitive
4 inhibitor with respect to NADPH, suggesting that the ratio of NADPH/NADP+ regulates the activity of both isoforms in planta.
Curiously, SSA is a mitochondrially-generated intermediate of GABA metabolism, whereas glyoxylate is a peroxisomally-generated intermediate of glycolate metabolism or photorespiration, a pathway believed to be particularly important under conditions of high light and temperature (Wingler et al, 2000). Glyoxylate is typically transaminated to Gly, but it can also undergo reduction to glycolate (Givan and
Kleczkowski, 1992). Both SSA and glyoxylate are aldehydes, which can accumulate and become toxic to the plant because of their extreme reactivity (Kotchoni et al,
2006). Aldehydes cause cellular and developmental problems that are related to their ability to react with DNA, oxidize membrane lipids, modify proteins, or influence the transcription of stress-related genes (Weber et al, 2004; Kotchoni et al., 2006).
Overexpression of oxidative or reductive enzymes that eradicate aldehydes can improve plant tolerance to various stresses (Oberschall et al., 2000; Sunkar et al., 2003).
Oxidation and reduction of the aldehyde grouping is accompanied, respectively, by the reduction of NAD(P)+ to NAD(P)H and the oxidation of NAD(P)H to NAD(P)+, indicating that the relative importance of these processes is probably dependent upon the cellular NAD(P)H:NAD(P)+ ratio (e.g., Busch and Fromm, 1999; Hoover et al,
2007a, b), and suggesting that aldehyde metabolism plays a role in redox homeostasis during stress (Bouche et al, 2003; Breitkreuz et al, 2003; Fait et al, 2005). The turnover of pyridine nucleotides involves other enzyme activities as well, including several distinct isoforms of NAD kinase (NADK) (Harding et al, 1997; Hunt et al,
2004; Turner et al, 2004).
5 In this thesis, two hypotheses were tested. The first hypothesis is GHB accumulation in Arabidopsis and tobacco plants is a general response to abiotic stress, and regulated by redox balance and glyoxylate reductase isoforms. In order to address this hypothesis, the responses of GHB and related amino acids (GABA, Ala, Glu), as well as NADP+ and NADPH, were monitored in mature leaves from Arabidopsis or tobacco plants subjected to various abiotic stresses (i.e., Arabidopsis during exposure to salinity, drought, submergence, cold or heat; tobacco during exposure to, and recovery from submergence) known to cause GABA accumulation. Other amino acids that are associated with primary nitrogen metabolism (Gin, Asp, Asn), photorespiratory nitrogen metabolism (Gly, Ser) and stress (Pro) were measured in order to gauge the broader impact of the stress. Furthermore, GLYR1 or GLYR1 and GLYR2 expression was monitored in Arabidopsis plants subjected to the various stresses.
The second hypothesis is glyoxylate reductase isoforms function in redox homeostasis and detoxification of both SSA and glyoxylate in planta. In this case, the simultaneous utilization of glyoxylate and SSA by recombinant Arabidopsis GLYR1 was investigated, and Arabidopsis GLYR1 and GLYR2 KO mutants (glyrl and glyrl, respectively) were utilized to investigate the role of these proteins in the detoxification of glyoxylate and/or SSA and in redox homeostasis under environmental conditions that promote GABA metabolism or photorespiration. Also, various NADK KO (nadkl, nadk3) and expression (NADK1 overexpressor (OE), NADK2 underexpressor (AS) mutants were utilized to investigate biochemical crosstalk between NADK and GLYR activities during submergence stress.
6 CHAPTER 2 - Literature Review
The following literature review provides an overview of aldehyde metabolism and toxicity, the pathway and regulation of GAB A and GHB metabolism in plants, pyridine nucleotide metabolism and distribution, as well as glyoxylate metabolism and photorespiration.
2.1. Metabolism and Toxicity of Aldehydes
Aldehydes belong to a large class of organic compounds containing the CHO functional group and are ubiquitous in nature. The electronegative character of the carbonyl carbon renders them reactive electrophiles that can deleteriously react with cellular nucleophiles, including proteins and nucleic acids (Lindahl, 1992; O'Brien et al, 2005).
Outdoor sources of exogenous aldehydes include the photochemical oxidation of hydrocarbons present in the atmosphere. Car exhaust, foliar terpenes and isoproprene emissions, forest fire and agricultural burns, incinerators, diesel engines and coal-based power plants all emit hydrocarbons or aldehydes into the atmosphere. The United States
Environmental Protection Agency and the Canadian Environmental Protection Act regulate the atmospheric aldehyde load. They report that the most significant aldehydes within the atmosphere include formaldehyde, acetaldehyde, acrolein, oc-oxoaldehydes
(glyoxyl, methyglyoxyl, glyoxylate) and acetone (O'Brien et al.,, 2005). Indoor air is also polluted with these compounds from emissions from furniture, carpets, particle boards, fabrics, paints, cooking fumes, fireplaces, and cigaratte smoke. Formaldehyde
7 levels have been reported to be as high as 1.38 mg l"1 and 6.8 mg l"1 in rain and fog water, respectively (WHO, 1989; Muir, 1991; O'Brien etal, 2005).
Despite the prevalence of aldehydes within the environment, the main exposure of organisms to aldehydes is from endogenous intermediary metabolism. Aldehydes are, for example, generated from the catabolism of Thr (acetoaldehyde), putrescine (y- amino-butyraldehyde), choline (betaine aldehyde), Pro (glutamic-Y-semialdehyde), vitamin A (retinal), and the GABA shunt (SSA). However, the major metabolic aldehydes are products of lipid peroxidation (Lindahl, 2002; O'Brien et al, 2005; Jin and Penning, 2007). Oxidative stress results in the excessive formation of ROS such as superoxide anion radical, hydroxyl radical, and hydrogen peroxide. One target of ROS is polyunsaturated fatty acids (e.g., linoleic acid, arachidonic acid) that form lipid hydroperoxides (Oberschall et al, 2000; Kotchoni et al, 2006; Jin and Penning, 2007), which decompose to form reactive lipid aldehydes (e.g., 4-hydroxy-2-nonenal, malondialdehyde, 4-oxo-2-nonenal). These lipid aldehydes are bifunctional electrophiles and undergo Schiff base formation with the e-NH2 group of reactive Lys and 1,4-Michael addition which results in protein cross-links. Thiohemiacetal/thioester intermediates between glyceraldehyde-3-phosphate and the sulfhydryl group of the active site Cys are formed during the turnover of glyceraldehyde 3-phosphate dehydrogenase, demonstrating that aldehydes also react with the sulfhydryl group of cysteines (Pederson and Jacobsen, 1980). Recently, Hao and Maret (2006) demonstrated that acetaldehyde and acrolein react with Cys ligands of zinc proteins, causing the zinc to be released, thereby affecting phosphorylation signaling and gene expression in mammals. Aldehydes also react with bases in DNA to form a series of
8 etheno- and heptano-etheno-DNA adducts, which are highly mutagenic (Esterbauer et al, 1991).
In humans, it has been shown that the production of alkenals (i.e., acrolein, crotonaldehyde, 2-nonenal, 2,4- decadienal, cinnamaldehyde), hydroxyalkenals (i.e. 4- hydroxynonenal, 4-oxonon-2-enal), dicarbonyls (i.e., glyoxal, methylglyoxal) and alkanals (formaldehyde, acetoaldehyde, nonanal, benzaldehyde, vanillin) occur upon lipid peroxidation decomposition (O'Brien et al.,, 2005). These compounds lead to the accumulation of DNA damage and DNA adducts, carbonylated proteins, and glycated proteins. Cellular damage from lipid aldehydes has been linked to many disease processes such as metal storage diseases (iron (hemochromatosis) or copper (Wilson's disease), liver cancer, muscular dystrophy, rheumatoid arthritis, Parkinson's disease,
Alzheimers and atherosclerosis. Dicarbonyls or oxoaldehydes such as glyoxal, methylglyoxal and deoxyglucosone are of particular concern because of their propensity to cross-link long-lived proteins. Cross-linking of proteins found in vivo is associated with long-term complications of diabetes and atherosclerosis (Lindahl, 2002;
Jin and Penning, 2007).
2.1.1. Plant aldehydes
In plants, the rapid and excessive accumulation of ROS caused by many environmental stressors, including high light, ultraviolet radiation, drought, flooding, salt stress, heat, cold, and heavy metals results in oxidative stress (Bouche et al. 2003; Sunkar et al.
2003; Hao and Maret, 2006; Kotchoni et al, 2006; Moller et al, 2007). The
9 amplification of ROS damage is further exacerbated by the accumulation of toxic aldehydes arising from reactions of ROS with lipids and proteins. The polyunsaturated fatty acids (PUFAs) linoleic acid (18:2) and linolenic acid 18:3) are the major fatty acids in the plant membrane galactolipids (thylakoids) and phospholipids (all other membranes). These fatty acids are susceptible to peroxidation by singlet oxygen and hydroxyl radical giving rise to a complex mixture of lipid hydroperoxides (Griffiths et al, 2000; Moller et al., 2007). In addition, lipoxygenase generates lipid peroxides when plants are attacked by phytopathogens and herbivores or during senescence (Berger et al, 2001; Mano et al, 2002). These lipid peroxides are degraded enzymatically or non- enzymatically to form toxic C3-C9 aldehydes such as a, (3-unsaturated aldehydes (2- alkenals) and malondialdehyde (MDA). MDA damages DNA by conjugation with guanine, creating an extra ring. The 2-alkenals, 4-hydroxy-(2E)-nonenal (HNE) and 4- hydroxy-(2E)-hexenal (HHE) are highly reactive. HNE inhibits decarboxylating enzyme complexes (e.g., pyruvate dehydrogenase, 2-oxoglutarate dehydrogenase,
NAD-malic enzyme) by forming an adduct with their lipoamide moiety (Miller and
Leaver, 2000; Mano et al, 2002; Moller et al, 2007). HNE also reacts with Cys and
Lys residues in the active sites of alternative oxidase (Winger et al, 2005), glucoses- phosphate dehydrogenase, glyceraldehyde-3-phosphate dehydrogenase, and cytochrome c oxidase (Millar and Leaver, 2000), effectively inhibiting their activity. HNE is further converted to 2, 3-epoxy-4-hydroxynonanal, which reacts with adenine and guanine
(Chung etal, 2006).
2.1.1.1. Detoxification of plant aldehydes
10 Because of their toxicity, cellular strategies have evolved to detoxify ROS and reactive aldehydes. ROS accumulation is largely counteracted by cellular antioxidant systems.
Enzymatic scavengers include superoxide dismutase, ascorbate peroxidase, glutathione peroxidase, glutathione-S-transferase, and catalase. Non-enzymatic scavengers involve low-molecular mass molecules like ascorbate, tocopherol, carotenoids and glutathione
(Bray et al, 2000; Sunkar et al, 2003; Moller et al, 2007). Reactive aldehydes are generally detoxified enzymatically. Four distinct pathways exist: the conjugation of aldehydes with glutathione via glutathione-S-transferase; the reduction to alcohols by aldo-keto reductases (AKR); the oxidation to carboxylates with alcohol or aldehyde dehydrogenases (ADH or ALDH, respectively); and, the hydrogenation of the a,P- unsaturated bond of 2-alkenals (Mano et al, 2002). Recently, an NADPH:quinone oxidoreductase Pl-£- crystallin was found to catalyze the hydrogenation of the a, (3- unsaturated bond of HNE in Arabidopsis. It was induced by oxidative stress treatment, suggesting that the enzyme plays a defensive role against lipid peroxidative damage
(Mano et al, 2002). This is the only known enzyme in plants which inactivates reactive aldehydes by hydrogenation.
2.1.1.2. Aldehyde dehydrogenases
Aldehyde dehydrogenases are NAD(P)+-dependent enzymes that catalyze the irreversible oxidation of aldehydes to their corresponding carboxylic acids (Kirch et al, 2004, 2005; O'Brien et al, 2005; Kotchoni et al, 2006). ALDH genes have been
11 identified in most organisms, including prokaryotes and eukaryotes. Across all taxa,
1000 genes have been identified. In eukaryotes, at least 172 genes have been identified,
which encode enzymes that belong to more than 21 protein families based on sequence
similarity (Kirch et al,, 2004; Fong et al, 2006). Protein sequences sharing > 40%
identity are placed into the same family, while those sharing > 60% are placed into the
same subfamily. Members of the same superfamily also share similar three-dimensional conformations, with three distinct domains in each monomer: a catalytic domain;
NAD(P)+ binding domain; and, an oligomerization domain (Kirch et al, 2004; Fong et al, 2006). In humans, ALDHs have been well-characterized; 16 genes and 3 pseudogenes have been identified and their chromosome locations determined in the human genome and their encoded enzymes have been studied (O'Brien et al, 2005). In plants, less is known about gene functions, but it is known that ALDH enzymes are represented in 11 families: ALDH2, ALDH3, ALDH5, ALDH6, ALDH7, ALDH10,
ALDH11, ALDH12, ALDH18, ALDH19, and ALDH21. Three of these families are unique to plants (ALDH11, ALDH 19, ALDH21) (Kirch et al, 2004).
Encoded ALDH enzymes range from being very substrate-specific to highly
variable in their substrate specificity (Sophos and Vasiliou, 2003). For example, in humans, aldehyde oxidase is expressed in the cytosol in the liver, kidney and intestine
and uses oxygen to oxidize all aromatic aldehydes, including benzaldehydes, indole-
aldehydes, retinal, and butyraldehyde with similar efficiency, whereas ALDH2 is
expressed in the mitochondria in the liver and has the strongest affinity for acetaldehyde
(O'Brien, 2005). In plants, the first ALDH2 gene (rf2) discovered encodes a nuclear
restorer of cytoplasmic male sterility (Kirch et al, 2005). Orthologues in maize, rice
12 and Arabidopsis have been characterized. The mitochondrial ALDH2 in rice detoxifies acetaldehyde during re-aeration after submergence and in Arabidopsis the two isoforms
ALDH2B7 and ALDH2B4 have been shown to oxidize acetoaldehyde and glycolaldehyde in vitro (Kirch et al, 2004). In contrast, three genes from family 3 are known in Arabidopsis and one particular isoform, ALDH3I1, is induced by various oxidative stresses and oxidizes several aldehydes from lipid peroxidation degradation
(Kirch et al, 2004).
In the last ten years, characterization of ALDH genes in plants has made progress. In the Arabidopsis genome, there are 14 aldehyde dehydrogenase genes which encode proteins that belong to nine ALDH protein families (Kirch et al, 2004, 2005).
ALDH3I1 and ALDH7B4 are known to be targeted to the chloroplast and cytoplasm, respectively, and both mRNA and protein accumulate in plants in response to salt stress, drought, ABA, hydrogen peroxide, and the heavy metals copper and cadmium.
The accumulation of protein is accompanied under stress by a decline in MDA levels, despite an increase in cellular hydrogen peroxide. In overexpressor transgenic lines, plants exhibit high tolerance to salt stress and dehydration, whereas in knockout (KO) lines, plants are unable to tolerate salt stress relative to the wild-type (Sunkar et al,
2003; Kotchoni et al, 2006). The functional homologue of ALDH3I1 from the dessication-tolerant plant Craterostigmaplantagineum,}loscht responds in a similar fashion to dehydration and salt stress (Kirch et al, 2001). Antiquitin is classified as family 7 in the ALDH superfamily and is subdivided into three subfamilies: ALDH7A,
ALDH7B, and ALDH 7C. The garden pea ALDH7B1 was the first antiquitin discovered in plants because of its increase in gene expression when the plant is
13 dehydrated. Orthologues in rapeseed, Arabidopsis, and Sorghum bicolor (L.) Moench have since been identified, and they are all dehydration-inducible. It has been suggested that antiquitin plays a role in osmoprotection by oxidizing aldehyde precursors to generate carboxylate-containing osmoprotectants. ALDH9 has a similar role, since it oxidizes betaine aldehyde to form glycine betaine. When ALDH9 is overexpressed, the plants are significantly improved in dehydration tolerance (Weretilnyk and Hanson,
1990). It has also been suggested that antiquitin contributes to the removal of HNE and
MDA because Arabidopsis and tobacco plants overexpressing soybean antiquitin exhibit higher tolerance against oxidative stress. A similar role is ascribed to ALDH3 in humans because of its higher activity towards HNE and MDA (Vasiliou et al, 2004;
Fong et al, 2006; Rodrigues et al, 2006).
Another aldehyde dehydrogenase which is prevalent in humans and plants is
SSADH. This enzyme is part of the GABA shunt and catalyzes the NAD+-dependent oxidation of SSA, a catabolite of GABA, to succinate (Shelp et al, 1999; Bouche et al,
2003; Breitkreuz et al, 2003; Fait et al, 2008). SSADH has been classified as a member of the ALDH family 5 (Kirch et al, 2004); it is located in the mitochondrion and in plants, there is only one isoform. It has been suggested that there is a second isoform based on the Arabidopsis genome, but cDNA and EST databases do not confirm this (Kirch et al, 2004). In humans, SSADH deficiency leads to severe neurodevelopmental and other physiologic abnormalities, concomitant with an elevation of GHB concentration in brain tissues (Hong et al, 1997; Nguyen et al,
2003). In plants, null mutants of SSADH in Arabidopsis are dwarfed and develop necrotic lesions upon exposure to UV-B radiation or heat, along with a rise in cellular
14 hydrogen peroxide (Bouche et al, 2003). The hypersensitivity to oxidative stress is also accompanied by elevated GHB levels. It has been suggested that the rise in GHB may be partially responsible for the effects seen with SSADH deficiency in humans and plants (Bouche et al, 2003; Nguyen et al, 2003).
2.1.1.3. Aldehyde reductases
The aldo-keto reductase superfamily (AKR) comprises a functionally diverse gene family in which the encoded enzymes catalyze the NADPH-dependent reduction of aldehydes and ketones to primary and secondary alcohols, respectively. The enzymes are cytosolic, monomeric (oc/P)8-barrel oxidoreductases of about 320 amino acid residues in length, which reduce a wide variety of metabolites including steroids, sugars, aldehydes and ketones (Hyndman et al, 2003). Enyzmes from this superfamily have been found in bacteria, fungi, nematodes, vertebrates and plants (Seery et al 1998;
Jez and Penning, 2001). There are currently 140 members in the superfamily across 15 families. There are 13 known human enzymes that play central roles in the detoxification of carbonyl-containing drugs (cancer therapeutic reagents such as doxorubicin and cyclophosphamide). Eight of the human AKRs belong to family 1 and these include aldehyde reductase (AKR1A1), aldose reductases (AKR1B1 and
AKR IB 10), hydroxysteroid dehydrogenases (HSDs) and steroid 5P-reductase
(AKRD1). Other human AKRs include the aflatoxin aldehyde reductases (AKR7A2,
AKR7A3) and SSAR (AKR7A1). The tissue distribution of AKRs in humans is widespread. AKR1 members are constitutive and ubiquitious with highest expression
15 found in the kidney, liver and brain. AKR1B10, on the other hand, is only expressed in the small intestine and colon. AKR7 members are also widely distributed and are equally abundant in all tissues.
AKRs are generally regulated by stress signals including osmotic shock and oxidative stress (Jin and Penning, 2007). For example, AKR1B1 (aldose reductase) is induced by osmotic shock. This response is expected because the enzyme catalyzes the conversion of glucose to sorbitol, which aids in water retention. The enzyme also detoxifies HNE and acrolein. In fact, AKR enzymes tend to catalyze reactions on a broad and overlapping array of substrates making it difficult to assign specific substrates and physiological functions to each enzyme.
Known plant AKRs are members of the AKR4 family and include aldose reductase, polyketide reductase (chalcone reductase), alkaloid reductase, and various oxidoreductases of poorly defined function (Hiroyuki et ah, 2007). Recently, a novel
AKR from the root of Aloe arborescens was found to reduce benzaldehyde and DL- glyceraldehyde using NADPH as co-factor (Hiroyuki et ah, 2007). In alfalfa cells, an
NADPH-dependent aldose/aldehyde reductase is expressed in response to osmotic shock, heavy metal toxicity, hydrogen peroxide, and abscisic acid (ABA), and alfalfa leaves exhibit a significant increase in mRNA and protein levels under drought conditions (Oberschall et ah, 2000). The purified enzyme reduces aldehyde substrates with NADPH as cofactor and the transgene overexpressed in tobacco confers stress resistance to paraquat, hydrogen peroxide, and water deficiency, as well as a decrease in lipid peroxidation-derived reactive aldehydes. In cultured bromegrass cells, freezing induces elevated levels of aldose reductase (Lee and Chen, 1993), and in monocots,
16 aldose reductases reduce glucose to sorbitol. Thus, AKRs in plants seem to function as detoxification pathways against reactive carbonyl-containing metabolites and may be involved in osmoregulation.
The occurrence of distinct NADPH-dependent glyoxylate reductase (GLYR) isoforms and activity in the cytosol and chloroplasts of spinach and pea has been documented (see refs in Givan and Kleczkowski, 1992). Recent research indicates that
Arabidopsis cytosolic GLYR1 and plastidial (GLYR2) are involved in the reduction of both glyoxylate and SSA to glyoxylate and GHB, respectively (Hoover et al., 2007a;
Simpson et al, 2008). Thus, plant GLYRs act as aldo-keto reductases; but, there is no significant homology with mammalian or bacterial NADPH-SSA reductases (SSARs), nor with members of the AKR superfamily (see Simpson et al., 2008 and references therein). There is however, 22-31% sequence identity to known |3-hydroxyacid dehydrogenases (HIBADH superfamily) such as tartronate semialdehyde reductase from Escherichia coli, and 6 phosphogluconate dehydrogenase from several species including E.coli, Cunninghamella, sheep, maize and spinach (see references in Simpson et al., 2008). GLYR1 and GLYR2 contain conserved residues implicated in substrate binding in 6-phosphogluconate dehydrogenase, and the ordered Bi Bi kinetic mechanism for recombinant GLYR1 is consistent with that for members of both the
AKR and HIBADH members (Hoover et ah, 2007b), These observations suggest that plant GLYR enzymes are members of the HIBADH superfamily (Hoover et al., 2007a;
Simpson et al., 2008).
2.2. GABA Metabolism
17 Glu can be metabolized via a short metabolic pathway (Fig. 2.1) that bypasses the
NAD+-dependent 2-oxoglutarate dehydrogenase and the ADP+-dependent succinyl-
CoA steps of the tricarboxylic acid cycle (TCA), and allows the Glu carbon backbone to enter the cycle as succinate (Shelp et ah, 1999). This pathway is found in virtually all organisms from bacteria to humans, underscoring its fundamental nature in metabolism.
GABA is a four-carbon non-protein amino acid with the amino group on the y-carbon, rather than the a-carbon. In plants, under a variety of abiotic and biotic stresses, GABA increases several fold (see Section 2.2.2). This is due to the decarboxylation of Glu via a reaction catalyzed via the cytosolic Ca2+/CaM-dependent GAD. The decarboxylation of Glu consumes a proton; thus, one possible role for the accumulation of GABA may be to ameliorate cytosolic acidification (Crawford et ah, 1994). The catabolism of
GABA involves pyruvate and 2-oxoglutatarate-dependent GABA-T activities (Shelp et ah, 1995; Van Cauwenberghe and Shelp, 1999; Van Cauwenberghe et ah, 2002). The pyruvate-dependent activity produces Ala which is known to rise under a variety of abiotic stresses (Breitkreuz et ah,, 2003; Allan and Shelp, 2006; Miyashita and Good,
2008). The 2-oxoglutarate-dependent activity produces Glu which likely feeds back into the GABA shunt (Shelp et ah, 1999; Fait et ah, 2008). Both GABA-T activities also produce SSA which is quickly oxidized via mitochondrial NAD+-dependent SSADH to succinate which feeds into the TCA cycle. (GAD, GABA-T and SSADH make up the pathway that is commonly known as the GABA shunt.) Under conditions such as
18 a. OH
OH
NH, Glutamate
GAD NADPH NADP+ b. OH OH N* CO, OH JL OH ^o GLYR Glyoxylate Glycolate
NH, y-Aminobutyrate
a-Ketoglutarate Pyruvate
Glutamate Alanine NADPH NADP*
SSAR/GLYR y-Hydrpxybutyrate Succinic semialdehyde
' NAD* SSADH S*. NADH OH o O
OH Succinate
Fig. 2.1. Alternative pathways for GAB A metabolism via succinic semialdehyde reductase (a) and glyoxylate reductase reaction (b). Enzymes are in italics. GAD,
glutamate decarboxylase; GABA-T, GABA transaminase; GLYR, glyoxylate reductase;
SSADH, succinic semialdehyde dehydrogenase; SSAR, succinic semialdehyde reductase. (Adapted from Hoover et al., 2007a).
19 anoxia wherein the TCA cycle is inhibited, SSA can be reduced to GHB via NADPH- dependent SSAR (Allan et al, 2003; Breitkreuz et al, 2003). Thus, the ratios of
NADH/NAD+ and NADPH/NADP+ probably play a pivotal role in the catabolism of
GABA and hence GABA levels in the cell. It has been suggested that the accumulation of GABA is a step in the Ca2+ signal transduction pathway, as well as a pH-stat mechanism (Cholewa et al, 1996; Shelp et al, 1999; Kinnersley and Turano, 2000).
Indeed, stresses that increase cytosolic Ca2+ or H+ levels, such as cold shock, mechanical touch, drought, salinity, or oxidative stress also elevate GABA.
2.2.1. Regulation
2.2.1.1. Glu decarboxylase (GAD)
GAD catalyzes the irreversible, proton-consuming, pyridoxal 5'-phosphate-dependent a-decarboxylation of L-Glu. This is shown in vivo by the production of 14CC>2 and unlabelled GABA from the decarboxylation of [l-14C]Glu (Chung et al, 1992; Tuin
and Shelp, 1996). This enzyme has maximum activity at pH 5.8 and little activity at pH
7.0, although it is activated by Ca2+-saturated CaM at physiological pH (Shelp et al,
1999). For example, Ling et al, (1994) demonstrated that GAD is a CaM-activated enzyme in fava bean roots using a I-CaM overlay assay and activity assays with the partially purified root GAD. Detailed molecular analysis of the CaM-binding domain in petunia GAD, and characterization of the partially-purified soybean GAD or purified recombinant petunia GAD showed that there is rapid stimulation of GAD activity in
20 response the addition of Ca and CaM (Arazi et al, 1995; Snedden et al, 1995, 1996).
Furthermore, a monoclonal antibody specific for the CaM C-terminal region of GAD activates the enzyme in the absence of Ca2+/CaM and CaM antagonists prevent the activation by Ca2+/CaM (Snedden et al, 1995,1996; Cholewa et al, 1996). Treatment of rice roots for 1 h with Ca2+ channel blockers and CaM antagonists prevents the accumulation of GAB A after 3 h of anoxia (Aurisano et al, 1995). Furthermore, treatment of isolated Asparagus densiflorus Kunth mesophyll cells with Ca2+ channel blockers and CaM antagonists prevented the accumulation of GABA accumulation after cold shock; in the absence of these agents, a fluorescent indicator revealed that within 2 s after cold shock, cytosolic Ca2+ increases, followed by GABA accumulation within 60 s (Cholewa et al., 1996). Without cold shock treatment, a Ca2+ ionophore elevates cytosolic Ca2+ and increases GABA pool sizes. Transgenic tobacco plants expressing a mutant petunia GAD without the CaM-binding domain exhibits stunted stems, along with elevated GABA and low Glu levels (Baum et al, 1993). Thus, GAD activity is linked to the Ca2+ signal transduction pathway.
GAD activity is also regulated transcriptionally and translationally (Shelp et al,
1999). Genes from petunia, tobacco, tomato, rice and Arabidopsis have been identified.
There are different expression patterns of mRNA and protein in different organs of petunia (Chen et al, 1994). Analysis of mRNA sequences from a tomato fruit cDNA library revealed a protein that is expressed during fruit color development and has 80% amino acid identity to the petunia GAD (Gallego et al, 1995). Bioinformatics or . genomic analysis indicated the presence of four to five GAD isoforms in the
Arabidopsis (Shelp et al, 1999), tobacco (Yevtushenko et al, 2003) and rice genomes.
21 AtGADl is root specific and AtGAD2 is present in all organs (Turano and Fang, 1998;
Zik et al., 1998). AtGADl is the major contributor for root GAB A pools under heat
stress (Miyashita and Good, 2008). If plants are fed NH4 as the nitrogen source,
AtGADl has higher transcript levels, encoded protein and specific activity in leaves.
The other three other putative isoforms possess 75-82% amino acid identity with GAD1
over the entire amino acid sequence (Shelp et al., 1999). The C-terminal residues
making up the CaM-binding domain have more variability. Isoforms 2 to 5 have 35-
43% identity with the C-terminal domain of GAD1, and only the CaM-binding domain
of GAD2 does not contain Ser, Thr or Tyr residues, which are potential phosphorylation
sites. To date, the C-terminal domain of all GADs from dicotyledonous plants are
autoinhibitory, contain a basic, amphiphilic a-helix, are hydrophobic, contain a highly
conserved Trp and Lys cluster and are Ca2+/CaM dependent (Turano and Fang, 1998;
Zik et al, 1998; Liu et al., 2004; Oh et al., 2005). This is in contrast to those from
bacteria and animals, which do not contain a CaM-binding domain. Recently, however,
a rice isoform (OsGAD2), which is not activated by Ca /CaM, was identified (Akama
et al., 2001; 2007). Nevertheless, the C-terminal extension of this isoform is
autoinhibitory, and when the C-terminal extension is removed, the truncated protein has
7-40-fold higher GAD activity in vitro with or without Ca2+/CaM (Akama et al, 2007).
Rice calli cells transformed with the truncated GAD have up to 100-fold higher levels
of GAB A than the wt, whereas calli transformed with the full-length GAD have only
six-fold higher GABA levels. Rice and tobacco plants transformed with the truncated
GAD are sterile with pale green curly leaves and GABA levels are elevated several-fold
in various organs.
22 GAD activity may also be stimulated by substrate availability. Elevated Glu levels in Asparagus mesophyll cells are accompanied by increased GABA levels (Chung et al, 1992; Cholewa et ah, 1997), and long-term conditions that increase Glu pool sizes, such as limited Gin synthesis, reduced protein synthesis or enhanced protein degradation, are correlated with increased GABA levels (Satya Narayan and Nair,
1990; Shelp et al, 1999). Scott-Taggart et al. (1999) demonstrated that when photorespiratory NH3 recycling is restricted by aminoacetonitrile, a Gly decarboxylase
(GDC) inhibitor, 14C-radioactivity in Gin decreases and 14C-radioactivity in Glu, succinate and TCA cycle organic acids increases. Precursor-product relations also showed that succinate is derived primarily from the GABA shunt (Scott-Taggart etal,
1999). Thus, GAD is stimulated by increasing Glu availability, and more carbon enters the TCA cycle via GABA than via 2-oxoglutarate.
2.2.1.2. y-Aminobutyrate transaminase (GABA-T)
The catabolism of GABA is initiated with the activity of GABA-T, which catalyzes the pyruvate-dependent or 2-oxoglutarate-dependent transamination of GABA to SSA and
Ala or Glu, respectively. GABA-T activity was first demonstrated in nodulated roots of pea in the 1960; however, that data was controversial because activity could not be demonstrated in non-nodulated pea roots or in several species (see Satya Narayan and
Nair, 1990). In 1972, Streeter and Thompson convincingly demonstrated GABA-T activity in radish leaves using a radiometric assay. Wallace et al. (1984) later showed activity in soybean cotyledons and seeds, tomato, alfalfa, cowpea, barley and yellow
23 foxtail using a similar assay and more thorough characterization. The reaction is freely reversible, is pyridoxal-phosphate-dependent and has a pH optimum of 8.6-9.0
(Wallace et al, 1984; Satya Narayan and Nair, 1990; Shelp et al, 1995). In bacteria, yeast, fungi, and mammals, GABA-T seems to be specific for 2-oxoglutarate as the amino group acceptor and is located in mitochondria (Van Cauwenberghe and Shelp,
1999; Van Cauwenberghe et al, 2002). However, in plants two distinct GABA-T activities have been detected: 2-oxoglutarate-dependent and pyruvate-dependent, although pyruvate appears to be the preferred amino acceptor. GABA-T isolated from peanut-seedlings, radish leaves, soybean leaves, wheat embryos, potato tuber and soybean seeds have 1-19 times more activity with pyruvate than 2-oxoglutarate
(Streeter and Thompson, 1972; Wallace et al, 1984; Galleschi et al, 1985; Satya Naran and Nair, 1986; Shelp et al, 1995). Both activities are found in tobacco leaf crude extracts and can be separated using ion chromatography, whereas pyruvate -dependent activity only is found in lysed mitochondrial preparations (Van Cauwenberghe and
Shelp, 1999). Recently, Clark (2008) and DiLeo (2004) discovered the existence of three GABA-T genes in tomato, which are located in the mitochondrion, cytosol or plastid. The mitochondrial isoform exhibited the highest activity which was comparable to GABA-T activity from Arabidopsis. Interestedly, all three tomato isoforms and the mitochondrial GABA-T from Arabidopsis utilized pyruvate and glyoxylate as amino acceptors. The use of glyoxylate as an amino acceptor by GABA-T suggests that
GABA metabolism is linked to photorespiration. Despite the existence of multiple isoforms of GABA-T, the discovery of gene(s) for 2-oxoglutarate-dependent activity remains elusive.
24 2.2.1.3. Succinic semialdehyde dehydrogenase (SSADH)
SSADH is a NAD+-dependent enzyme localized in the mitochondrial matrix (Satya
Narayan and Nair, 1989; Busch and Fromm, 1999). It efficiently catalyzes the irreversible oxidation of SSA to succinate at a pH optimum of 9.0-9.5 (Streeter and
Thompson, 1972b; Satya Narayan and Nair, 1989; Shelp et al, 1995) and the succinate enters the TCA cycle. The plant enzyme has been purified to homogeneity from barley seeds and potato tubers and enzyme activity has been detected in crude extracts from
Arabidopsis, radish cotyledons, wheat embryos, barley, soybean and potato tubers
(Satya Narayan and Nair, 1989; Breitkreuz and Shelp, 1995; Shelp et al, 1995). A
SSADH gene has been cloned and the recombinant protein purified and characterized fxora Arabidopsis (Busch and Fromm, 1999). The gene encodes a 53 kDa subunit that associates into a homotetramic structure of 197 kDa within the mitochondrion. The protein is about 59% identical at the amino acid level to E. coli, rat and human SSADH.
The N-terminus consists of a 33-amino acid mitochondrial targeting peptide that is cleaved inside the mitochondrion to produce mature protein. Activity is inhibited by
NADH, ATP and AMP and thus SSADH acts as a sensor of respiratory stress (Busch and Fromm, 1999; Busch et al, 2000). In mammals, deficiency in this enzyme causes a significant rise in GHB, which has been linked to severe neurological abnormalities
(Chambliss et al, 1998; Gibson et al, 1998). In plants, KO mutants in Arabidopsis exhibit chlorotic lesions and an accumulation of hydrogen peroxide and GHB, thereby
25 linking the GABA shunt to redox homeostasis in the cell (Bouche et ah, 2003; Fait et al, 2005). It likely also plays a role in preventing a rise in the SSA pool.
2.2.1.4. Glyoxylate/succinate semialdehyde reductase (GLYR)
In plants, NADPH-dependent glyoxylate/SSAR (designated officially as GLYR) catalyzes the essentially irreversible reduction of glyoxylate and SSA to glycolate and
GHB, respectively (Breitkreuz et al, 2003; Hoover et al, 2007a; Simpson et al, 2008).
Two isoforms are known in Arabidopsis, a cytosolic (AtGLYRl) and a plastidial
(AtGLYR2) form. In vitro kinetic measurements of recombinant GLYR1 revealed that it
1 1 has a 250-fold higher preference for glyoxylate over SSA (kcJKm = 2870 s" mM" for glyoxylate; 11.6 s"1 mM"1 for SSA), and 100-fold higher preference for SSA than other substrates tested (formaldehyde, acetaldehyde, butyraldehyde, 2-carboxybenzaldehyde, glyoxal, methylglyoxal, phenylglyoxal, and phenylglyoxylate) (Hoover et al, 2007a).
Further characterization via initial velocity, dead-end inhibiton and product inhibition studies demonstrated that the kinetic mechanism is ordered Bi Bi, involving the complexation of NADPH to the enzyme before glyoxylate or SSA and the release of
NADP+ before the release of glycolate or GHB, respectively (Hoover et al, 2007b).
NADP+ is also a competitive inhibitor with respect to NADPH {K{ =1-3 |iM), suggesting that the enzyme functions in vivo as a regulator of NADPH/NADP4 ratios.
GLYR2 has a 57% identity at the amino acid level to the cytosolic form. The sequence of the full-length cDNA encodes an N-terminal plastid targeting sequence of 43 amino acids, and when both isoforms are fused separately to the green fluorescent protein and
26 transiently expressed in tobacco suspension cells, the GLYR2 is targeted to the plastid, whereas GLYR1 is targeted to the cytosol (Simpson et al, 2008). The truncated, recombinant protein of GLYR2 also prefers glyoxylate over SSA (350-fold higher)
1 1 1 based on performance constants (kcat/Km= 660 s" mM" for glyoxylate and 1.89 s" mM for SSA) and uses NADPH as its preferred cofactor. Thus, it too may function in vivo as a regulator of reducing equivalents. Both enzymes have kinetic characteristics that are consistent with the members of the AKR and HIBADH superfamilies (Simpson et al., 2008). Thus, they play a role in aldehyde detoxification as well.
Glyoxylate (oxaldehydic acid), an intermediate in photorespiratory metabolism, is produced in the peroxisomes by the oxidation of glycolate via glycolate oxidase.
Glyoxylate is quickly aminated to Gly. If glyoxylate accumulates, ribulose- bisphosphate carboxylase/oxygenase (RUBISCO) can be inhibited (Campbell and
Ogren, 1990). Glyoxylate is also a reactive aldehyde that can modify Lys and Arg side chains, and form glyoxylated DNA adducts (Schmitt et al, 2005; Maekawa et al,
2006). Thus, glyoxylate reductase activity in cells may function in scavenging excess glyoxylate.
In Arabidopsis cell-free extracts, total GLYR activity using SSA as substrate is found at all developmental stages and in all tissues; however, activity is generally higher in vegetative and reproductive organs than in roots (Hoover et al, 2007a).
SSADH-deficient yeast complemented with an Arabidopsis GLYR1 grows on 20 mM
GABA as the sole nitrogen source and contain elevated levels of GHB (Breitkreuz et al, 2003). These studies suggest that SSA-dependent AfGLYR activity potentially occurs in planta, despite the fact that glyoxylate is the preferred substrate in vitro.
27 2.2.2. Potential Roles
The accumulation of GAB A in response to various stresses in planta is well documented (Bown and Shelp, 1989; Satya Narayan and Nair, 1990; Shelp et al, 1999;
Kinnersley and Turano, 2000; Bouche and Fromm, 2004; Fait et al, 2008), yet the role of the GABA shunt remains speculative. In mammals, the prominent role of GAB A is as a signaling molecule since it is a major inhibitory neurotransmitter in the central nervous system (CNS), and it influences proliferation, migration, and differentiaton of cells during development of the CNS (Kinnersley and Turano, 2000; Owens and
Kriegstein, 2002; Bouche et al, 2003; Bown et al, 2006). In plants, the role of GABA as a signaling molecule is not as clearly defined. In Arabidopsis, a large pool of GABA is generated by the root specific GAD1 and GABA has been found in the xylem stream and phloem exudate of many species (Bown and Shelp, 1989; Chung et al, 1992;
Beuve et al, 2004; Fait et al, 2008). Potential GABA transporters in the plasma membrane of several tissues have been identified, including amino acid permease 3
(AAP3), Pro transporter 1 (ProTl), Pro transporter 2 (ProT2) and GABA transporter 1
(GAT1) (Rentsch et al, 1996; Breitkreuz et al, 1999; Schwack et al, 1999; Meyer et al, 2006). AAP3 and the ProTs, which belong to the amino acid/auxin transporter
(AAAP) or amino acid transporter (ATF) superfamily however, might not transport
GABA in planta as their affinity for amino acids or for the compatible solutes Pro and glycine betaine are considerably higher than for GABA. The recently discovered GAT1 transporter has high specific affinity for GABA when expressed in yeast and Xenopus
28 oocytes and is targeted to the Arabidopsis plasma membrane as shown by fusion to a green fluorescent protein (Meyer et al, 2006). Transcript levels are highest in
Arabidopsis flowers and are only induced in other tissues during senescence and wounding. Thus, AtGKIl probably transports GABA from the apoplast under certain conditions or during pollination. Indeed, Palanivelu et al, (2003) showed that a GABA gradient is needed for pollen tube growth and guidance through female tissue to the micropyle, implying a role for GABA in cellular migration. However, AtGATl expression is very low in pollen and only slightly elevated in carpels, confounding its role in this process. Recently, a role for long distance signaling in Brassica napus L. has been indicated (Beuve et al., 2004). Exogenous GABA supplied to the roots induces mRNA expression of a nitrate transporter, BnNrt2. This observation was interpreted in conjunction with a correlation between GABA in phloem exudate and nitrate uptake, suggesting a putative role for GABA as a long-distance inter-organ signal. Furthermore, plant development can be affected by elevated GABA. High
GABA concentration inhibits stem elongation in Stellaria, soybean hypocotyl tissue, and tobacco (see refs in Shelp et al, 1999). Agonists and antagonists of animal GABA receptors exert contrasting effects on the growth of duckweed (Lemna minor L.
(Kinnersley and Lin, 2000). Together, these studies provide circumstantial evidence for the inter- and intra-cellular transport and signaling of GABA. Nevertheless, the existence of specific GABA receptors is elusive. It is noteworthy that Arabidopsis Glu receptors share sequence and structural homology with the mammalian ionotropic Glu receptors, and possess a domain known to bind modulatory ligand molecules which share structural homology with domains of mammalian GABAB receptors (Bouche and
29 Fromm 2004). These receptors bind Glu or Gly and mediate calcium ion influx into the plant cell. This process may be modulated by GABA; however, direct evidence is lacking.
Shelp et al, (2006) recently reviewed evidence indicating that GABA mediates communication between plants and other organisms. For example, crustose red algae secrete GABA-mimetic molecules onto its surface and larvae of Haliotis rufescens
Swainson (abalone of eastern Pacific) which contain GABA receptors, settle and metamorphose due to chemosensory recognition of the GABA molecules when they graze the algal surface (Morse et al, 1979). Crawling of insect larvae on plant surfaces is known to increase leaf GABA concentration by 4-12 fold due to wounding (see refs in Bown et al, 2006). It was hypothesized that that ingestion of elevated GABA levels disrupts the normal physiological and developmental processes of the larvae because
GABA activates chloride channels at neuromuscular junctions. Plants overexpressing full-length or truncated GAD lacking the autoinhibitory CaM domain also reduce feeding by tobacco budworm larvae and impair the development of the northern root- knot nematode (MacGregor etal, 2003; McLean et al, 2003). Infection by the fungus,
Cladosporiumfulvum, induces GAD and GABA levels rise in the apoplast of tomato
(Solomon and Oliver, 2002, 2001). As a consequence, GABA-T and SSADH are induced in the fungus. When Agrobacterium tumefaciens is incubated with wounded tomato stems, elevated GABA levels are subsequently transported into the bacterium via the GABA transporter Bra (Chevrot et al, 2006), thereby inducing the expression of the attLKM operon in the bacterium and stimulating the production of lactonase AttM, which in turn inactivates N-(3-oxooctanoyl) homoserine lactone. The lactone controls
30 the conjugation of the Ti plasmid and the severity of the tumoral symptoms. Also, transgenic tobacco plants with higher GABA levels are less susceptible to
Agrobacterium infection.
Several other roles have been postulated for the GABA shunt. GABA rapidly accumulates upon exposure to various stresses (Table 2.1), suggesting that de novo
GAD synthesis is not involved. Indeed, the stimulation of GAD activity by Ca2+/CaM in vitro at neutral pH and in vivo places GAD in a signal transduction pathway (Aurisano et al, 1995; Cholewa et al, 1997). Ca2+ influx into the cell is stimulated by touch
(Knight et al, 1991), cold shock (Cholewa et al, 1997), heat shock, salinity, drought, and osmotic stress (Kinnersley and Turano, 2000 and references therein) and GABA does accumulate. However, at acidic pH, GAD is stimulated independently of
Ca2+/CaM, and in vivo, cytosolic acidification does stimulate GAD activity and GABA production. (Crawford et al, 1994; Shelp et al, 1995). Stresses that disrupt cellular membranes such as wounding, freezing or heat stress, could cause vacuolar contents to be released into the cytosol or inhibit ATPase enzymes which pump protons from the cytosol into the vacuole or apoplast causing a concomitant decrease of cellular pH and stimulation of GAD activity. Hypoxia is known to decrease cytosolic pH by 0.3-0.6 pH units and GABA does increase under oxygen deficiency in soybean, tobacco and
Arabidopsis (Shelp et al, 1995; Allan et al, 2003; Breitkreuz et al, 2003). It has been suggested that GABA accumulation helps to regulate cytosolic pH because GAD activity consumes a proton. Two independent studies demonstrated the direct effect
31 Table 2.1. Biotic and abiotic stresses stimulating GABA and GHB accumulation
Metabolite Treatment Plant Source Reference
GABA Mechanical Soybean leaves Wallace et ah, 1984 stimulation Hypocotyl tissue Bown and Zhang, 2000 Mechanical Soybean and Ramputh and damage tobacco leaves Bown, 1996; Bown et ah, 2002; Hall et ai, 2004 Alfalfa and Girousse et ah, 1996; tomato phloem Valle etah, 1998 exudate Fungal infection Tomato cell Solomon and Oliver, apoplast 2001 Agrobacterium Arabidopsis Deeken et ah, 2006 infection tumors Rhizobium Legume nodule Vance and Heichel, infection 1991 Cold stress Soybean and Wallaces ah, 1984; Arabidopsis leaves Kaplan et ah, 2007 Asparagus mesophyll cells Cholewa^aZ., 1997 Barley and wheat seedlings Mazzucotelli et ah, 2006. Heat stress Cowpea cell Mayer et ah, 1990 cultures Oxygen Rice roots Reggiani et ah, 1988; deficiency Aurisano et ah, 1995 Tea leaves, Tsushida and Murai, 1987 Soybean sprouts, Allan etah, 2003; Tobacco and Breitkreuz etah, Arabidopsis 2003 leaves Medicago Ricoult et ah, 2005 seedlings Rice cotyledons Kato-Noguchi and Ohashi, 2006 Broccoli florets Hansen et ah, 2004
32 .Table 2.1 con't.
Water stress Tomato roots and Bolarin et al, 1995 leaves Soybean nodules Serraj et al, 1998 and xylem sap Wheat seedlings Bartyzel et al, 2003/4 Phytohormones Datura root Ford et al, 1996 cultures Carbon dioxide Cherimoya fruit Merodio et al, 1998 enrichment broccoli florets Hansen et al, 2001
UV stress Arabidopsis plants Fait et al, 2005
GHB Oxygen Tea leaves, Allan et al., 2003; deficiency soybean sprouts, Bretikeuz et al, 2003 tobacco and Arabidopsis leaves Cold stress Arabidopsis Kaplan et al, 2007 leaves UV stress Arabidopsis plants Fait et al, 2005
33 cytosolic pH on GAD activity. Crawford et al, (1994) used a fluorescent probe to monitor the cytosolic pH changes when A. densiflorus cells are exposed to permeant weak acids. Within 2 s, the pH declines by 0.6 units and within 15 s, GABA accumulates by 200%, with the accumulation of GABA accounting for 50% of the imposed acid load within 45 s of exposure to the weak acids. Carroll et al. (1994) monitored cytosolic pH changes in Daucus carota L. cells upon initiation of ammonium assimilation using 31P-and 15N-NMR spectroscopy. The decline in pH by 0.2 units is followed by an increase in GAD activity and GABA accumulation and a decrease in
GAD activity when pH recovers. Thus, GABA production does ameliorate cytosolic acidification.
GABA could act as a compatible osmolyte (Shelp et al., 1999). During water, salt or freezing stress, tissues become dehydrated and cells can compensate by increasing the number of solutes within the cytoplasm to regulate their osmotic potential.
Compatible osmolytes are a group of small, organic compounds that are highly soluble and do not interfere with cellular processes. They are neutrally charged at physiological pH, either non-ionic or zwitterionic, are excluded from the hydration shell of macromolecules, and generally accumulate to high cellular concentrations upon dehydration. GABA shares these characteristics, along with Pro and Gly betaine, which are well-studied compatible osmolytes (Breitkreuz et al., 1999; Schwacke et al., 1999).
AfProT2 and LeProTl have the capacity to transport Pro, Gly betaine and GABA, and
AfProT2 is induced by water deficit. At high concentrations, GABA stabilizes and protects isolated thylakoids from freezing damage in the presence of salt (Heber et al.,
34 1971). In vitro, GAB A has higher hydroxyl-radical-scavenging activity than Pro or Gly betaine (Turano et al, 1997).
Another significant role for GABA is as a transient, nitrogen storage molecule (see refs in Shelp et al, 1999). Factors that induce a decline in nitrogen in the cell over time,
such as inhibition of Gin synthesis, reduction of protein synthesis, or an increase in protein degradation elevate Glu levels which are then rapidly decarboxylated to GABA.
Glu is a central molecule in amino acid metabolism in planta. The a-amino group of
Glu is directly involved in both the assimilation and dissimilation of ammonia and is transferred to all other amino acids. In addition, both the carbon skeleton and the oc- amino group form the basis of not only GABA, but also Arg and Pro (Forde and Lea,
2007). Glu concentrations tend to remain remarkably constant during the diurnal cycle under different environmental conditions (Allan and Shelp, 2006; Forde and Lea,
2007). However, the stimulation of GAD activity markedly decreases Glu levels,
suggesting GABA acts as a nitrogen storage molecule in times of stress. Indeed,
Masclaux-Daubresse et al (2002) reported a 6-h shift between GABA and Glu over a
2-d night cycle in tobacco. Since higher GABA levels always precede Glu levels, they hypothesized that GABA buffers Glu. Allan and Shelp (2006) also confirmed a close relationship between the fluctuations of Glu and GABA over a 2-d night cycle in
different aged leaves and nitrogen supply in Arabidopsis, although elevated GABA pools do not precede Glu pools.
The GABA shunt could act as a bypass of the TCA cycle for carbon. The bypass of the succinyl-CoA ligase and the 2-oxoglutarate dehydrogenase step provides metabolic flexibility to the cell by providing Glu rather than pyruvate carbon for use in the TCA
35 cycle. The oxidation of Glu is theroretically less energetically productive than pyruvate, however, because one mole of NADH and ATP are produced at the 2-oxoglurate dehydrogenase and succinyl-CoA steps, respectively, whereas only one mole of NADH is produced at the SSADH step in the GABA shunt. Nevertheless, stresses that restrict the oxidation of NADH diminish TCA cycle intermediates and/or inhibit TCA cycle enzymes such as isocitrate dehydrogenase, permit greater flux through the GABA shunt. For example, oxygen deficiency inhibits the mitochondrial electron transport chain, thereby stimulating glycolysis and fermentation to recycle NADH. Hypoxia and submergence does decrease Glu and increase GABA and Ala (Shelp et al, 1995; Allan et al, 2003; Breitkreuz et al, 2003; Miyashita and Good, 2008). Others showed that manipulation of TCA enzymes such as NAD+- dependent isocitrate dehydrogenase or aconitase does alter GABA shunt activity (Lemaitre et al, 2007; Nunes-Nesi et al,
2007). If succinyl-CoA ligase gene expression is downregulated, the GABA shunt can complement this deficiency (Fait et al, 2008). The relative carbon flux to TCA succinate through Glu or pyruvate is unknown.
2.3. Pyridine Nucleotide Metabolism and Distribution
The "division of labour" hypothesis concerning the metabolic roles of NAD and NADP was first formulated by Kaplan et al in 1956. Essentially, it states that NAD is reduced to NADH via a variety of catabolic dehydrogenase reactions and that this NADH is reoxidized by the mitochondrial respiratory chain to yield ATP. NADP on the other hand is reduced by a relatively few number of dehydrogenase reactions and the
36 resultant NADPH is used by the cell as reducing power for biosynthetic reactions. The knowledge of pyridine nucleotides has greatly expanded since this basic hypothesis was formulated and their role in the cell is evidently much more complex. Indeed, NAD and its derivative NADP are ubiquitous in living cells. They mediate hundreds of redox reactions, and thus influence virtually every metabolic pathway in the cell.
Furthermore, they are involved in ROS-producing and ROS-consuming systems in plants, and thus are key players in signaling events (Moller, 2001; Noctor et ah, 2006).
ROS production is influenced by NADPH/NADP"1" ratios in the chloroplast and mitochondrion and NAD(P)H oxidases are key players in the generation of ROS at the plasma membrane. ROS scavenging and quenching partly depends on glutathione and ascorbate pools, which are fundamentally maintained by NAD(P)H. Pyridine nucleotides play other defensive and signaling roles such as production of nitric oxide and metabolism of reactive aldehydes. Therefore, in addition to the basic roles outlined by Kaplan, NAD+ and NADP+ are crucial in redox metabolism linked to stress and development.
In plants, the biosynthetic pathway of NAD+ and NADP+ is not fully understood
(Noctor et ah, 2006): However, it is known that NAD+ pools are maintained by de novo and salvage pathways and NADP+ are synthesized by NADKs.
2.3.1. NADVNADH
NAD+ is a coenzyme that switches between oxidized and reduced forms without any net consumption. NADH is mainly involved in catabolic reactions, and is mainly
37 oxidized by the mitochondrial respiratory chain. Under aerobic conditions in yeast and
bacteria, there is a large NAD+ pool which serves to drive oxidative catabolic reactions
that support respiration (Grose et al, 2006). NADH pools are large under anaerobic
conditions, which may help to support growth when the rate-limiting step in providing
energy is reoxidation and recycling of NADH by using electron acceptors that are less
avid than oxygen. NAD+ is also utilized irreversibly as a substrate by the silent
information regulator family enzymes, which couple the degradation of NAD+ to
nicotinamide and the deacetylation of a substrate such as acetylated histones (Katoh et
al, 2006; Noctor et al, 2006). ADP-ribose units from NAD+ can be transferred to
target proteins as a post-translational modification catalyzed by poly(ADP-ribose)
polymerases (PARPs) and mono-(ADP-ribosyl) transferases. PARP activity is stress-
induced and hyperactivation of PARP-1 by ROS or nitric oxide has been shown to
deplete NAD+ and ATP pools in mammalian and plant cells and decreasing PARP
inhibition can diminish death in both types of cells (Heller et al., 1995; Amor et al.,
1998; De Block et al, 2005).
NAD+ is synthesized de novo from amino acid precursors in two distinct pathways:
the aspartate pathway and the kynurenine pathway (Noctor et al, 2006). In bacteria and
Arabidopsis, aspartate is the starting amino acid (Katoh et al, 2006). Asp is oxidized
via Asp oxidase to a-iminosuccinic acid, which is subsequently condensed with
glyceraldehyde-3-P and cyclized to produce quinolinic acid by quinolinate synthase.
The next step is catalyzed by quinolinic acid phosphoribosyl transferase (QPRT) which
forms nicotinic acid mononucleotide (NaMN) from quinolinic acid and phosphoribosyl
pyrophosphate. NaMN is then converted to NAD+ in two steps. In the kynurenine
38 pathway, which is found in mammals and fungi, NAD+ synthesis begins with Tip,
which is degraded via kynurenine to quinolinic acid in five enzymatic steps. The pathway from quinolinic acid to NAD+ is conserved among prokaryotes and eukaryotes.
In most organisms, including plants, the degradative metabolites of NAD+ and NADP+,
nicotinamide and nicotinic acid, are reutilized for the synthesis of pyridine nucleotides by salvage pathways (Ashihara et al, 2005).
NAD(H) concentrations in leaf mitochondria, chloroplasts, and cytosol are approximately 2.0, 0.4, and 0.6 mM, respectively (Takahama et al, 1981; Igamberdiev and Gardestrom, 2003) and total leaf pool sizes are typically 20-100 nmol g"1 FM
(Dutilleul et al, 2005; Shen et al, 2006). The synthesis of NAD(H) is compartmentalized, but characterization of the full pathway is incomplete. Quinolinate production is thought to be in the chloroplast because functional GFP-fused aspartate oxidase, quinolinic acid synthase, and QPRT have been localized to plastids in
Arabidopsis (Katoh et al, 2006). However, QPRT NaMN formation may also be in the cytosol and mitochondrion based on tobacco QPRT cDNA sequences that include a
shorter transcript and a longer transcript with a putative mitochondrial targeting
sequence (Sinclair et al, 2000). In Arabidopsis, a single gene is predicted to encode each of the enzymes necessary for the conversion of NaMN to NAD+ and neither
sequence encodes an apparent transist peptide sequence (Noctor et al, 2006). In mammals, three isoforms of NaMNAT have been found in the nucleus, in an undefined location and in the cytosol and mitochondrion (Magni et al, 2004). Therefore, it is probable the last two steps of NAD+ synthesis are not plastidial. This implies a requirement for mitochondrial and plastid inner membrane transporters able to transport
39 quinolinate and NAD+, but very little genetic and molecular evidence is available on
such transporters (Noctor et al, 2006).
NAD+ pools are plastic in plants and undergo significant changes in response to the
environment. For example, NAD+ contents are increased when barley leaves are
infected with powdery mildew, but this is not due to de novo synthesis (Noctor et al,
2006). NAD+ pools can be modified by degradation and recycling through salvage pathways and in turn, NAD+ recycling can be influenced by the removal of recycling
metabolites for pyridine alkaloid synthesis, degradation of nicotinamide or the
formation of nicotinamide and nicotinate conjugates during stress. Interestingly,
increases in NAD+ concentration are associated with stress resistance. Tobacco lines,
deficient in mitochondrial complex I, respire through alternative respiratory
dehydrogenases and are resistant to abiotic and biotic stress. This resistance is
accompanied by a two-fold increase in leaf NAD+ and NADH concentrations without
an appreciable change in NADPH/NADP+ ratio (Dutilleul et al, 2005). In the
cytoplasmic male sterile mutant (CMS), which displays stress resistance, NAD+
concentration is also elevated, but the biochemical and molecular processes for this
increase are not fully understood (Dutilleul et al, 2003).
It is unclear how quickly the oxidized and reduced states of NAD+ and NADP+
pools respond to the environment. Increasing light causes progressive over-reduction at
the reducing side of photosystem I (PSI), causing NADP+ levels to fall, and so favoring
the diversion of electrons to oxygen to produce ROS. Yet, the stromal redox state seems
to be constant in spite of irradiance. Direct measurements of fractionated pea leaf
protoplasts showed that cytosolic NADP+ redox states are independent of light, dark or
40 + + + C02 availability, yet cytosolic NAD and mitochondrial NAD and NADP are more reduced under photorespiratory conditions (Igamberdiev and Gardestrom, 2003).
Several other studies indicate that changes in cytosolic and mitochondrial NAD+ reduction are important in nitrate reduction and nitrogen assimilation (Igamberdiev and
Gardestrom, 2003; Rachmilevitch et al, 2004; Dutilleul et al, 2005).
2.3.2. NADP+/NADPH
The pyridine nucleotide NADP+ is synthesized by ATP-dependent 2'-phosphorylation of NAD(H). The enzyme responsible, NADK, is essential therefore, because NADPH supports reductive biosynthetic reactions and defends cells against oxidative stress
(Grose et al, 2006; Noctor et al, 2006). It is only recently that genes encoding NADKs have been cloned from human, yeast, and bacteria (Kawai et al, 2001; Lerner et al,
2001; Outten and Culotta, 2003). Eukaryotes possess multiple NAD(H)K genes, whereas bacteria apparently have only one gene. The enzymes are divided into NADKs
(EC 2.7.123) or NAD(H)Ks (EC 2.7.1.86) depending on their substrate preference for the oxidized or reduced forms of NAD (Strand et al, 2003; Berrin et al, 2005).
In general, plant NADK activities are present in all cell compartments, revealing the existence of several isoforms. In maize, the CaM-dependent enzyme is found in the cytoplasm, in the outer membrane of mitochondria and in the chloroplast envelope, whereas the CaM-independent form is located in the stroma of the chloroplast
(McGuiness and Bulter, 1985; Sauer and Robinson, 1985). In Avena sativa L. seeds, soluble and mitochondrial NADK activities are present in dependent and independent
41 forms (Gallais et ah, 2001; Pou de Crescenzo et ah, 2001). CaM-dependent activity has also been detected in pea seedlings, Solarium lycopersicum L. var. pimpinellifolium
(Juslen) Mill., and green bean (Delumeau et ah, 2000; Ruiz et ah, 2002 ). In
Arabidopsis, there are three isoforms; two are cytosolic (A^NADKl, AfNAD(H)K3) and one is targeted to the chloroplast (AtNADK.2) (Turner et ah, 2004, 2005; Chai et ah,
2005, 2006). A*NAD(H)K3 is homologous to POS5, a mitochondrial isoform from yeast recently described as an NADK which prefers NAD(H) as substrate, but it has negligible amino acid identity to other NADKs (Strand et ah, 2003; Turner et ah,,
2005). A?NAD(H)K3 has a pH optimum of 7.9, is partially inhibited by Ca2+/CaM , has a subunit molecular mass of 35 kDa and exists as a dimer. It can phosphorylate NADH or NAD+ in an ATP-dependent manner, but the specificity constant (Vmax/^m) for
NADH is several-fold higher than for NAD+ (Berrin et ah, 2005; Turner et ah, 2005).
This is in contrast to other known NADK isoforms in plants which exclusively prefer
NAD+ as substrate and are generally Ca2+/CaM-dependent (Sauer and Robinson, 1985;
Delumeau et ah, 2000; Turner et ah, 2004). Recently, Berrin et ah, (2005) demonstrated that A?NADK1 is induced by oxidative stress such as ionizing radiation and paraquat treatment and by compatible and incompatible plant-pathogen interactions. Another group showed that NAD(H)K3 is also induced by a variety of abiotic and biotic stresses and exogenous abscisic acid (Chai et ah, 2006); their data imply that this isoform has a large impact on the cytosolic NADPH level under stress.
Manipulation of AfNADK2 levels affects chloroplastic NADPH levels, and null mutants are stunted, with a pale yellow color, and hypersensitive to abiotic stress (Chai
42 et al, 2005). It was suggested that this isoform plays a role in chlorophyll synthesis and chloroplast protection against oxidative damage.
In the stroma, the concentration of NADP+ available for reduction by ferredoxin-
NADP+ reductase is considered important in determining electron flow to alternative acceptors such as oxygen (Scheibe et al., 2005). Cytosolic NADPH provides the reductant for production of ROS through NADPH oxidases. ROS interact with nitric oxide in defence signaling and growth responses (Noctor, 2006). Processing of ROS is also very heavily dependent on NADPH. Detoxification of ROS and organic peroxides occurs through the ascorbate and glutathione pools, which are regenerated through
NADPH dependent reactions such as monohydroascorbate reductase, glutathione reductase and thiol-disulfide thioredoxin (Benin et al., 2005; Noctor, 2006). In addition, reactive aldehydes generated from lipid peroxides are detoxified by NADP+- dependent oxidation or NADPH-dependent reduction catalysed by aldehyde dehydrogenases, aldo-keto reductases or alkenal reductases (Sunkar et al., 2003; Mano et al., 2005). NADPH-dependent hydroxylation reactions catalyzed by cytP450s are important in xenobiotic detoxification.
In the light, generation of NADPH in the chloroplast depends on the reduction of
NADP+ via ferredoxin-NADP+ reductase and by the oxidative pentose phosphate pathway (OPPP) dehydrogenases, glucose-6-P dehydrogenase and 6-phosphogluconate dehydrogenase, in the dark. Glucose-6-P and 6-phosphogluconate dehydrogenases also play roles in other compartments, such as the peroxisomes (Corpas et al., 1998). In the cytosol and mitochondria, NAD(P)-dependent isocitrate dehydrogenases (ICDH) play a major role in NADPH pool sizes (Igamberdiev and Gardestrom, 2003; Marino et al.,
43 2007). NADP-ICDH activity in mitochondria is higher in photosynthetic than heterotrophic tissues, implying an important role during photosynthesis. It has been suggested that NADP-ICDH in mitochondria provides NADPH for redox regulation of the alternative oxidase via thioredoxin (Vanlerberghe et al, 1995). Igamberdiev and
Gardestrom (2003) found that the NADPH/NADP"1" ratio is high during photorespiratory conditions in mitochondria due to oxidation of Gly. Indeed, GDC activity provides more NADH than the TCA cycle (Atkin et al, 2000). They suggested that this higher ratio stimulates transhydrogenation activity between NADH and NADP+. Two distinct mitochondrial transhydrogenation activities have been detected (Bykova et al, 1999,
2001). As a result, mitochondrial NADPH levels rise, NAD-ICDH in the TCA Cycle is inhibited and the mitochondrial NADP-ICDH operates in the reverse direction.
Consequently, citrate is primed to be exported from the mitochondrion, whereas 2- oxoglutarate is imported and enters the reverse NADP-ICDH reaction, as 2- oxoglutarate dehydrogenase activity is probably low during photorespiration. Thus,
NADP-ICDH reverse activity may recycle NADPH and 2-oxoglutarate in the mitochondrion during photorespiration.
+ + NADH/NAD and NADPH/NADP ratios at limiting C02 concentration are higher than at saturating CO2 concentrations in pea leaves and barley protoplasts
(Igamberdiev et al, 2001). GDC is inhibited by NADH (Kt 15 (iM), as is SSADH.
Under photorespiratory conditions, therefore, there is a necessity to remove NADH from mitochondria. Possible pathways for this include the export of reducing equivalents via the malate/oxaloacetate shuttle, oxidation of NADH in the electron transport chain, or transhydrogenation with NADP+.
44 In the dark, mitochondrial NADP+ and NAD+ pools are more oxidized (Igamberdiev and Gardestrom, 2003). Under non-photorespiratory conditions, the NADH/NAD+ratio is not significantly higher in nonphotorespiratory conditions as in the dark. This represents the oxidation of NADH mainly through the TCA cycle and not through
GDC.
2.4. Glyoxylate and Photorespiration
Photorespiration results from the oxygenation of ribulose-l,5-bisphosphate (RuBP) catalyzed by RUBISCO. This reaction produces glycolate-2-phosphate which is metabolized through the photorespiratory pathway to form glycerate-3-phosphate, which enters the Calvin cycle. The rate of carboxylation versus oxygenation is dependent on the ambient CO2 and O2 concentrations, their Km values, and the velocity of the overall reaction (Farquhar et al, 1980). Sharkey (1988) estimated that under ambient conditions, the rate of photorespiratory CO2 release is about 25% of the rate of net CO2 assimilation. When temperature rises, the specificity of RUBISCO for CO2 decreases, and the solubility of CO2 relative to O2 decreases, resulting in enhanced photorespiratory rates under higher temperatures. This effect is further augmented by lowered intercellular CO2 concentration, which can occur upon stomatal closure during drying conditions such as high light or drought (Noctor et al., 2002). Due to higher photorespiratory CO2 release, photorespiration becomes energetically inefficient, imposing a strong carbon loss on plants, and possibly leading to the depletion of carbohydrates and accelerated senescence. However, the suppression of
45 photorespiration over the long term results in decreased rates of photosynthetic CO2 assimilation, poor plant growth, and chloroplast aberrations (Tolbert et al, 1995; Migge et al, 1999). Thus, photorespiration is a useful process. It has been suggested that photorespiration is needed for energy dissipation to prevent photoinhibition under high light conditions and as a source of Ser and Gly (Madore and Grodzinski, 1984; Wingler et al, 2000).
The photorespiratory pathway involves several enzymes compartmentalized across chloroplasts, peroxisomes and mitochondria. Glycolate-2-phosphate is first hydrolyzed to glycolate by a chloroplastic phosphoglycolate phosphatase, which is in turn transported to peroxisomes where it is oxidized to glyoxylate and hydrogen peroxide via glycolate oxidase. H2O2 is removed by catalase and glyoxylate is transaminated to
Gly by Ala-, Ser- or Glu-dependent aminotransferases (AGAT, SGAT or GGAT)
(Nakamura and Tolbert, 1983; Liepman and Olsen, 2001). One-half of the Gly is converted to N5, N10-methylene tetrahydrofolate via NAD+-dependent GDC in the mitochondrion. In this reaction, CO2 and NH3 are released. The other half of the Gly reacts with N5-N10-methylene-THF to form Ser in a reaction catalyzed by Ser hydroxymethyltransferase. Ser is transported back to the peroxisomes where it is transaminated by Ser:glyoxylate aminotranferase to hydroxypyruvate, which is reduced to glycerate by NADH-dependent hydroxypyruvate reductase. Glycerate is then phosphorylated by glycerate kinase in the chloroplasts and the resulting glycerate-3- phosphate enters the Calvin cycle. Because of the transamination of glyoxylate to Gly and the large release of NH3 in the GDC reaction, photorespiratory metabolism is intimately linked to leaf nitrogen metabolism. It has been estimated that
46 photorespiratory ammonia release is an order of magnitude greater than the primary assimilation of reduced nitrogen (Keys et al, 1978). Thus, the reassimilation of photorespired NH3 by the glutamine synthetase/ferredoxin-dependent glutamate synthase (GS/GOGAT) system is essential for the nitrogen status in plants (Wingler et a/., 2000).
Genes involved in the photorespiratory pathway were discovered by growing mutagenized plants under high CO2 conditions to suppress photorespiration and permit growth similar to wt plants, and then under air to permit expression of severe stress symptoms such as chlorosis ( Somerville, 1986; Leegood et al, 1995). Upon return to high CO2, the mutant plants recover. A plethora of enzymes and transporters involved in photorespiratory metabolism in a wide range of species has been exposed in this way.
However, the peroxisomal glycolate oxidase and GGAT have not been isolated by this screening process (Wingler et al, 2000; Ruemann and Weber, 2006). It may be that other enzymes compensate under low CO2. Subsequent studies revealed that plants with reduced activities in photorespiratory pathway enzymes can reduce photosynthetic rates by 1) depletion of Calvin cycle intermediates, 2) reduction of photorespiratory nitrogen reassimilation, ultimately leading to a depletion in photosynthetic proteins, and 3) accumulation of photorespiratory metabolites, such as glyoxylate, which inhibit photosynthetic and Calvin cycle enzymes (Wingler et al, 2000).
Glyoxylate is the only photorespiratory metabolite to exert a feedback effect on photosynthesis in heterozygous barley mutated in GS2, Fd-GOGAT, GDC or SGAT
(Wingler et al, 2000). In vitro, glyoxylate can inhibit the activation of RUBISCO
(Campbell and Ogren, 1990) and in vivo, there is a negative relationship between
47 glyoxylate content in the leaves of heterozygous GS2 mutants and the activation state of
RUBISCO (Hausler et al, 1996). Other evidence suggests that glycerate and glycolate-
2-phosphate inhibit fructose-l,6-bisphosphatase/sedoheptulose-l,7-bisphosphatase and triose-phosphate isomerase, respectively (Anderson, 1971).
There are several possible pathways for the metabolism of glyoxylate (also see
Section 2.1.1.3). Glyoxylate can be reduced back to glycolate by an NADH or NADPH-
dependent hydroxypyruvate reductase in the peroxisome, or by an NADPH-dependent
glyoxylate reductase in the chloroplast or cytosol (Tolbert et al, 1970; Kleczkowski et
al, 1986, 1990; Hoover et al, 2007a; Simpson et al, 2008). Glyoxylate can also be
oxidized to oxalate ((Halliwell and Butt, 1974). It has long been proposed that glycolate oxidase is able to oxidize glyoxylate to oxalate based on kinetic studies, but this view is contentious (Franceschi et al, 1987, 2005). Oxalate can accumulate in leaves of rice independent of glycolate oxidase activity since antisense suppression of glycolate
oxidase has no effect on oxalate accumulation (Xu et al, 2006). Glyoxylate can also be
oxidatively decarboxylated to formate non-enzymatically (Igamberdiev et al, 1999). It has been suggested that the decarboxylation of glyoxylate to formate can form a GDC-
independent bypass to the normal photorespiratory pathway, as shown in Euglena
gracilis (Yokota et al, 1985a, b; Wingler et al, 2000). This is because formate can
react with tetrahydrofolate in the CI- tetrahydrofolate synthase pathway, thereby providing one-carbon units for the synthesis of Ser, and other products such as purines,
thymidylate, methionine and formylmethionyl-tRNA. Since hydrogen peroxide is rapidly degraded by catalase in the peroxisomes, the rate of CO2 formation by
glyoxylate decarboxylation is likely to be low compared with the rate of CO2 formation
48 by GDC (Yokota et al, 1985a). Nevertheless, because this pathway bypasses transaminase reactions in the peroxisomes, it may play a role under low nitrogen supply
(Niessen et al, 2007).
Recently, NAD+-dependent glycolate dehydrogenase activity from Arabidopsis mitochondria was characterized (Niessen et al, 2007). This enzyme oxidizes glycolate to glyoxylate in the mitochondria, and contributes to photorespiration. T-DNA insertions within the glycolate dehydrogenase gene reduce the magnitude of the post- illumination CO2 burst, as well as the Gly/Ser ratio, but are not conditionally lethal under low CO2. This pathway is similar to the oxidation of glycolate in chlorophytes. A
Chlamydomonas strain harbouring an insertion within the mitochondrial glycolate dehydrogenase is conditionally lethal at ambient CO2 levels and is capable of growing under elevated CO2, a results reminiscent of higher plant photorespiratory mutants
(Niessen et al, 2007). The presence of a mitochondrial enzyme to oxidize glycolate in plants confers more flexibility and adaptability in the photorespiratory response. Other examples of plasticity include the ability of chloroplast extracts to oxidize glycolate and glyoxylate to CO2 without the involvement of peroxisomes or mitochondria or to reduce glyoxylate to glycolate (Zelitch and Gotto, 1962; Zelitch, 1972; Mulligan et al.,
1983; Goyal and Tolbert, 1996). A glycolate-quinone oxidoreductase system, associated with the chloroplastic photosynthetic electron transport chain catalyzes the oxidation of glycolate to glyoxylate (Goyal and Tolbert, 1996).
Photorespiratory metabolism has higher energy requirements than in the absence of photorespiration (5.375 mol ATP and 3.5 mol NADPH per CO2 fixed in photorespiration versus 3 mol ATP and 2 mol NADPH per CO2 fixed without
49 photorespiration) (Wingler et ah, 2000). Thus, it has been suggested that photorespiration is important for maintaining electron flow to prevent photoinhibition under stress. Indeed, the reduction of glyoxylate via NADPH-dependent glyoxylate reductase would provide a sink for electrons by the recycling of reducing equivalents.
Kebeish et al., (2007) introduced the Escherichia coli glycolate catabolic pathway into
Arabidopsis to reduce the loss of fixed carbon and nitrogen during photorespiratory metabolism by diversion of glycolate from the normal photorespiratory pathway. They transformed chloroplasts with glycolate dehydrogenase only, or with glycolate dehydrogenase, glyoxylate carboligase, and tartronic semialdehyde reductase, and found a substantial increase in rosette diameter under ambient and photorespiratory conditions, a decrease in the CO2 compensation point, and lower Gly/Ser ratios under high light and temperature relative to the wild-type. This suggests that glycolate is oxidized in the chloroplast and glyoxylate is successfully metabolized by glyoxylate carboligase and tartronic semialdehyde reductase, releasing CO2 near RUB IS CO.
Interestingly, the plants transformed only with glycolate dehydrogenase also showed evidence of chloroplastic glyoxylate metabolism.
2.5. Stress and Photorespiration
Photoinhibition occurs when the photochemical processes of photosynthesis do not have enough ADP or NADP+ to enable continued flow of energy through the electron transport system (Nilsen and Orcutt, 1996). Electron flow becomes blocked as molecules within the electron transport chain become reduced. Then the electrons of the
PSII centre become highly excited as light energy continues to be absorbed;
50 photoinhibion occurs as the excess light energy damages the Dl protein in the PSII reaction centre. Photoinhibition is usually temporary because of the turnover rate of the repair mechanisms of DI protein. However, if other molecules within the photochemistry system are permanently damaged by excess energy, chlorophyll is bleached, a process known as photo-oxidation.
The risk of photo-oxidation occurs under conditions in which stomata close and the internal leaf C02 concentration (Q) falls (Osmond, 1981; Osmond et al, 1997). In this situation, the regeneration of RuBP cannot keep up with the excess energy absorption, resulting in over-reduction of the electron transport chain. It has been suggested that photorespiration is important to maintain electron flow to prevent photoinhibition and/or photo-oxidation under stress.
Under mild to moderate drought or salt stress, Ci falls as stomata close to prevent water loss (Wingler et al, 2000 and references therein). Under salt stress, photorespiration has been observed to rise as indicated by higher CO2 compensation points (Fedina et al, 1993), higher light to dark ratio of CO2 production (Rajmane and
Karagde, 1986), higher glycolate oxidase activity, and the formation of photorespiratory metabolites such as Gly, Ser and glycolate (Downton, 1977). Under moderate drought stress in heterozygous barley mutants inhibited in various photorespiratory enzymes,
CO2 assimilation declines quickly than in wt, demonstrating the control over carbon fixation by photorespiratory metabolism under stress (Wingler et al, 2000). As well, in
Casuarina equisetifolia, L. Rubisco and photorespiratory enzymes activities are sustained under drought stress as Ci decreases, suggesting that photorespiration rates would increase relative to photosynthetic rates and in absolute terms (Sanchez-
51 Rodriguez et al, 1999). These studies suggest photorespiration can maintain electron flow under under stress.
52 CHAPTER 3 - y-Hydroxybutyrate Accumulation in
Arabidopsis and Tobacco Plants is a General Response to
Abiotic Stress: Putative Regulation by Redox Balance and
Glyoxylate Isoforms
3.1. Contributions
A slightly modified version of this chapter was published in the Journal of
Experimental Botany 59: 2555-2564 (2008). This manuscript derives from my primary
research project and I had an integral role in planning and deciding the direction of the
research. I performed all the experiments and data analysis, and prepared the entire
manuscript in conjunction with Barry J. Shelp. Co-authors Jeffrey Simpson and Shawn
Clark provided technical assistance and contributed to the planning of the research, as
well as interpretation of the data..
Wendy L. Allan, Jeffrey P. Simpson, Shawn M. Clark, and Barry J. Shelp.
Department of Plant Agriculture, University of Guelph, Guelph, Ontario, Canada NIG 2W1
3.2. Acknowledgments
This work was supported by funds to BJS from the Natural Sciences and Engineering
Research Council of Canada. SMS was the recipient of an Ontario Graduate
Scholarship.
53 3.3. Abstract
Enzymes that reduce the aldehyde chemical grouping (i.e. H-C=0) to its corresponding alcohol are probably crucial in maintaining plant health during stress. SSA is a mitochondrially-generated intermediate in the metabolism of GAB A, which
accumulates in response to a variety of biotic and abiotic stresses. SSA can be reduced to GHB under oxygen deficiency and high light conditions. Recent evidence indicates that distinct cytosolic and plastidial glyoxylate reductase isoforms from Arabidopsis
(A/GLYR1 and AfGLYR2, respectively) catalyse the in vitro conversion of SSA to
GHB, as well as glyoxylate to glycolate, via NADPH-dependent reactions. In the present report, the responses of GHB and related amino acids, as well as NADP+ and
NADPH, were monitored in leaves from Arabidopsis or tobacco plants subjected to various abiotic stresses (i.e. Arabidopsis during exposure to salinity, drought,
submergence, cold, or heat; tobacco during exposure to, and recovery from,
submergence). Time-course experiments revealed that GHB accumulated in both
Arabidopsis and tobacco plants subjected to stress, and that this accumulation was generally accompanied by higher GABA and Ala levels, higher NADPH/NADP+ ratio,
and lower Glu levels. Furthermore, the analysis of gene expression in Arabidopsis revealed that the relative abundance of GLYR1 (salinity, drought, submergence, cold,
and heat) and GLYR2 (cold and heat) transcripts was enhanced by the stress tested.
Thus, GHB accumulation in plants is a general response to abiotic stress and appears to be regulated by both biochemical and transcriptional processes.
54 3.4. Introduction
Under some stress conditions, aldehydes can accumulate in plants and react with DNA, oxidize membrane lipids, modify proteins, or influence the transcription of stress- related genes, thereby causing cellular and developmental problems (Weber et al,
2004; Kotchoni et al, 2006). Consequently, metabolic pathways and enzymes that reduce the aldehyde chemical grouping (i.e., H-C=0) to its corresponding alcohol are probably essential for maintaining plant health.
SSA is a mitochondrially-generated intermediate in the metabolism of GABA, which accumulates in response to a variety of biotic and abiotic stresses (Bown and
Shelp, 1997; Shelp et al, 1999). SSA is typically oxidized to succinate (Tuin and
Shelp, 1994; Bouche et al, 2003), but evidence for the reduction of SSA to GHB is also available (Breitkreuz et al, 2003), at least under oxygen deficiency and high light
(Allan et al, 2003a, b; Breitkreuz et al, 2003; Fait et al, 2005). Glyoxylate is a peroxisomally-generated intermediate of glycolate metabolism or photorespiration, a pathway believed to be particularly important under conditions of high light and temperature (Wingler et al, 2000). Glyoxylate is typically transaminated to Gly, but it can also undergo reduction to glycolate (Givan and Kleczkowski, 1992). Recently, recombinant expression of a cytosolic enzyme from Arabidopsis thaliana (L.) Heynh
(glyoxylate reductase 1 or AfGLYRl; EC 1.1.1.79) revealed that it effectively catalyzes the in vitro NADPH-dependent reduction of both SSA and glyoxylate (Hoover et al,
2007a). Part way through the present study, the existence of a plastidial enzyme
55 (glyoxylate reductase 2 or Art3LYR2) that catalyzed the same reactions in vitro was also demonstrated (Simpson et al, 2008).
Herein, the responses of GHB and related amino acids (GABA, Ala, Glu), as well as
NADP+ and NADPH, were monitored in mature leaves from Arabidopsis or tobacco plants subjected to various abiotic stresses {i.e., Arabidopsis during exposure to salinity, drought, submergence, cold or heat; tobacco during exposure to, and recovery from submergence) known to cause GABA accumulation. Other amino acids that are associated with primary N metabolism (Gin, Asp, Asn), photorespiratory N metabolism
(Gly, Ser) and stress (Pro) were measured in order to gauge the broader impact of the stress. Furthermore, GLYR1 and GLYR2 expression was monitored in Arabidopsis plants subjected to the various stresses. The results indicate that GHB accumulation is a general response to abiotic stress, and suggest that both redox balance and GLYR isoforms are involved in the regulation of SSA detoxification in planta.
3.5. Materials and Methods
3.5.1. Growth of plant material
Arabidopsis thaliana (L.) Heynh ecotype Columbia seeds were sown on Fox sandy loam (pH 6.5) in 5-cm pots, stratified at 4 °C in the dark for 48 h and grown under an
11-h photoperiod (06.00-17.00 h) in environmentally-controlled growth chambers
(Controlled Environments Ltd., Winnipeg, Manitoba) set at 22/18 °C day/night temperatures, a photosynthetic photon flux density (PPFD) of 150 ixmol m"V1 at the
56 top of the pots (supplied by a combination of cool white fluorescent and incandescent lighting; Sylvania, Mississauga, Canada), and 65% relative humidity. Arabidopsis plants were sub-irrigated with tap water as needed and fertilized once weekly with 20-
20-20 fertilizer. Tobacco (Nicotiana tabacum L. cv. Samsun NN) plants were grown under a 16-h photoperiod (06.00- 22.00 h) in a greenhouse with supplemental lighting and temperature as described previously (Scott-Taggart et al, 1999). Seeds were sown on Fox sandy loam soil (pH 6.5) in 5-cm diameter pots and grown for 3 weeks.
Seedlings were watered with tap water as needed and fertilized once weekly with 20-
20-20- fertilizer (Plant Products Co. Ltd., Brampton, Ontario, Canada). Individual seedlings were transferred to 30-cm diameter pots containing the same soil type, and grown for six more weeks with once daily watering with tap water and bi-weekly fertilization with 20-20-20.
3.5.2 Arabidopsis exposure to drought, salinity, submergence, cold or heat
Five different stresses (drought, salinity, submergence, cold, heat) were individually applied in separate experiments. In all experiments, pots were individually arranged in trays that were randomly located within a single growth chamber; water and nutrients were supplied via sub-irrigation, and treatments commenced when plants were 18-27 d old. For the drought experiment, sub-irrigation was withheld from one-half of the plants for up to 9 d, and plants were harvested after 0-9 d at 11.00 h. For the salinity experiment, one-half of the plants were sub-irrigated with 250 mM NaCl solution, with the other half receiving tap water, and plants were harvested at 0-24 h after first light
57 (06.00 h). For the submergence experiment, plants were divided into two basins, one of which was filled with water at ambient temperature. The basins were covered tightly with tinfoil, and plants were harvested under dim light for up to 6 h, beginning at 11.00 h. For the cold and hot experiments, the plants were divided into two covered basins at
11.00 h and 13.00 h, respectively; one each was placed in a Kelvinator refrigerated cabinet set at 4 °C or an incubator (model 1535, VWR Scientific Products,
Mississauga, Canada) set at 40 °C, and on the adjacent benchtop, and plants were harvested at 0-6 h thereafter. In all experiments, the shoots were rapidly frozen by dipping into liquid nitrogen for 10 s and then clipped at soil level into liquid nitrogen.
The rosette leaves were separated from the stem and stored at -80 °C until further analysis. Each sample consisted of 3-4 replicates, containing 3-4 pooled plants. While the submergence experiment was repeated in its entirety, the results for one experiment are shown herein.
3.5.3. Tobacco submergence and recovery
Beginning at 18.00 h, experimental plants were submerged in water equilibrated to the temperature of the greenhouse, for up to 9 h and then allowed to recover from the stress for a further 9 h. Experimental and control plants were placed in 50-1 garbage pails with tight fitting lids for the duration of the experiment. Recovering plants were transferred in the unlighted greenhouse at 3, 6 and 9 h after the onset of submergence from water- filled pails to dry pails. The most fully expanded leaf was sampled from experimental and control plants (n=3 for each datum) over the 18-h time course by dipping that leaf
58 into liquid nitrogen for 20 s and then snipping it from the shoot directly into liquid nitrogen. All samples were subsequently stored at -80 °C until further analysis.
3.5.4. Extraction and analysis of GHB, amino acids, and phosphorylated pyridine nucleotides
GHB and soluble amino acids were extracted from frozen leaves (300 mg aliquot) and analyzed by high performance liquid chromatography essentially as described previously (Allan et al, 2003; Allan and Shelp, 2006). Total amino acids (TAA) were estimated from the amino acid profiles. Phosphorylated pyridine nucleotides {NADP+,
NADPH) were extracted from frozen leaf tissue (50 mg aliquot) and assayed as described in Gibon and Larher (1997) except for the following modifications. After the neutralization of the acidic and basic extracts, 0.1 ml 16 mM phenazine ethosulfate was added and the extracts were incubated on ice for 30 min before being added to the assay solution. After the enzyme-cycling assay was completed, the reactions were stopped by adding 0.5 ml 6 M NaCl, followed by cooling on ice for 10 min. These solutions were centrifuged for 5 min (10 000 g, 4 °C) and the supernatant was carefully siphoned off. The pellet was then washed in 1 ml distilled water and solubilized in 1 ml
96% cold ethanol, and the absorbance of the formazan from 3-(4,5-dimethylthiazolyl-
2)-2,5-diphenyltetrazolium bromide was determined at 570 nm.
3.5.5. Expression and analysis of GLYR1 and GLYR2 in Arabidopsis
59 Quantitative real-time reverse transcription-polymerase chain reaction (RT-PCR) was preformed using the Platinum SYBR Green qPCR SuperMix UDG (Invitrogen) with a
BioRad iCycler (Hercules, CA). The following pairs of gene specific primers were used: GLYR\ gene (5TCGTAGAAGGTCCGGTTTCA3',
5'AAACTCCTTAGTTAGGGTCG3'); GLYR2 gene (5'AAG GAT ACC GGA GCC
TTG TT3', 5TCCACTGTTCGGCGATATGC3') and the 18S ribosomal RNA gene
(5*TCTGGCTTGCTCTGATGATT 3',5TCGAAAGTTGATAGGGCAGA3'). For analysis of gene expression for GLYR1 and GLYR2 under stress, total RNA was isolated from the leaves of the Arabidopsis plants under each experimental condition for each time point and treated with Turbo DNase (Ambion, Austin, TX). For cDNA synthesis 300 ng of total RNA was incubated with 1 |il of random hexamer primers, brought up to 15 |^1 with RNase free water, incubated at 75 °C for 10 min and chilled on ice. Two microlitres of lOx reaction buffer, 40 units of RNase inhibitor (Ambion), and 10 nM dNTPs were added and then incubated at 25 °C for 10 min. One hundred units M-MLV reverse (Ambion) were added to bring the final reaction volume to 20 jil, then the reaction was incubated at 37 °C for 1 h and inactivated at 95 °C for 10 min.
Each quantitative PCR reaction used lx SYBR Green qPCR mix, 0.2 JJM forward and reverse primers and 1 fil of cDNA in a 20-|il volume. All tubes were subjected to 3 min at 95°C, followed by 40 cycles of 95 °C for 20 s, 60 °C for 20s and 72 °C for 20 s.
SYBR Green absorbance was detected at 72°C and all reactions were conducted in triplicate. GLYR1 or GLYR2 transcript abundance in each sample was normalized to the corresponding level of 18S ribosomal RNA.
60 3.6. Results
3.6.1. Stress responsiveness of metabolites and phosphorylated pyridine nucleotides in Arabidopsis and tobacco
Six separate experiments were conducted, five with Arabidopsis (salinity, drought, submergence, cold and heat) and another with tobacco (submergence). For those involving Arabidopsis, the pool sizes of metabolites and phosphorylated pyridine nucleotides in control plants varied somewhat because of differences in plant age at the beginning of the experiments, the duration of the experiments, and the time of harvest
(see Allan and Shelp, 2006). It is significant that where appropriate (i.e., submergence, cold, heat), control plants were placed in the dark to eliminate the complicating effects of the dark environment that was superimposed during treatment.
With one exception (i.e., Arabidopsis during submergence), leaf Pro accumulated in plants of both species subjected to the various treatments, indicating that these treatments initiated the stress response (Fig. 3.1). In all cases and regardless of the plant species, the imposition of stress increased the GHB concentration over the duration of the time course used. With only two obvious exceptions (i.e., Ala under salinity, Glu under cold), the patterns for GHB-related metabolites were consistent, with GABA and
61 Fig. 3.1. Response of metabolites in mature rosette leaves of Arabidopsis plants (A) and mature leaves of tobacco plants (B) subjected to salinity, drought, submergence, cold or heat. Closed and open circles represent control and experimental plants, respectively. Data represent the mean ± SE
(n =3, each sample consists of leaves from 3-4 pooled rosettes); where the bar is not shown, it is within the symbol. Note that the drought data only are expressed on a DM basis, rather than a FM basis. TAA represents total amino acids.
62 A, Arabidopsis B. Tobacco
Salinity Drought Submergence 4°C 40 V Submergence TAA , 140 80 80 - 30 100 '^y^\ 40 - *#f^"* 70 15 40 S*< 50 xy V^ 0 i 0 0 0 14 •Gin] 1.6 16 28 . / A 7 0.8 8 \X 14 \Z— 1 f ° •y 0 /A ^v^^ 0.0 0 0 16 24 10 Glu 6 o—»w£^»0 <•/< j>i ^*~6 s ?fyr^ 8 .V ' 12 3h
0 0 0 0 3 LA f f 0.12 4 6 2 /: /\ vi\/ 3 1 0.06 r 2 ^>- °^*^— •* ^ Y 3 0.05 • 4 4 \Ac^ V \V^-^^^^ o 0 0.00 0 V><2 0 6 1.6 14 14 • ; Vs. / 0.2 3 /M 0.8 7 7 i **~*~~~-—~* »^sW-* •\ 0 \f —*~* • I • I 0.0 0 L V-r-° ? 0 Pro] 2 3 3 4 A 0.6 2 • r\ 2 <^ ••/• \ • A 2 1 •L^> 1 4 ••+ »r» r* o^» •—-^ 0 10 20 0 4 8 o 3 6 0 3 6 0 3 6 Time(h) Time (d) Time (h) Time(h) Time(h) 63 Ala being positively related, and Glu being negatively related with GHB. The patterns for other amino acids, as well as TAA, were not consistent across the various stresses. In three of the six experiments, the concentrations of phosphorylated pyridine nucleotides were also determined (Arabidopsis under cold or heat conditions, tobacco during submergence) (Fig. 3.2). In these cases, the plants exhibited a temporary increase in NADPH concentration together with general decline in NADP+, resulting in enhanced NADPH/NADP+ ratios over most of the time course. 3.6.2. Recovery of GHB and related metabolites, and phosphorylated pyridine nucleotides in tobacco after removal from submergence In the submergence experiment with tobacco, plants were removed from the stress at various times to test their ability to recover within a subsequent 9-h period. When plants were removed 3 h after submergence, the GHB concentration decreased immediately in a linear fashion, although it was still higher than that in control plant 64 A. Arabidopsis B. Tobacco 4°C 40 °C Submergence NADPH NADPH 60 40 ^ * 5 •£ b ^—-* i—t 20 x!_ "tan ,_O^ | o 0 «i NADP* 20 NADP 0 2 4 6 0 2 4 6 0 3 6 9 Time (h) Time (h) Time (h) Fig. 3.2. Response of phosphorylated pyridine nucleotides in mature rosette leaves of Arabidopsis plants (A) and mature leaves of tobacco plants (B) subjected to cold, heat or submergence. Closed and open circles represent control and experimental plants, respectively. Data represent the mean ± SE (n =3, each sample consists of leaves from 3-4 pooled rosettes); where the bar is not shown, it is within the symbol. 65 Ala continued to increase as in submerged plants before declining to control values, whereas the concentration of Glu exhibited a temporary increase and then approached control values. Also, there was a temporary increase in NADPH and NADPH/NADP+, which returned to control values; curiously NADP+ exhibited a longer term increase followed by a decline (Fig. 3.3B). Increasing the time of submergence before transfer to air generally decreased the capacity of plants to recover from the stress; particularly evident were the unwavering GHB and NADPH concentrations and fluctuating NADPH/NADP+ ratios at the end of the time course. 3.6.3. Stress responsiveness of GLYR transcripts in Arabidopsis In three of the five experiments involving Arabidopsis (salinity, drought, submergence), GLYR1 transcript abundance was determined (Fig. 3.4A), whereas in the other two (cold and heat), the levels of both GLYR1 and GLYR2 transcripts were determined (Fig. 3.4B). In these cases, GR transcript abundance was consistently higher in treated plants than in control plants. 3.7. Discussion 3.7.1. Impact of abiotic stress on GABA, Ala and redox levels 66 Fig. 3.3. Recovery of GHB and related metabolites (A), as well as phosphorylated pyridine nucleotides (B), in mature leaves of tobacco plants after being subjected to submergence. The column headings indicate the duration of submergence (3,6 or 9 h) before transfer to air for an additional 9 h. Data for control and submerged plants for the first 9 h are taken from figures IB and 2B. Closed circles, open circles and closed triangles, respectively, represent control plants, submerged plants, and plants returned to air after a period of submergence. Data represent the mean ± SE (n=3, each sample consists of the most recently expanded leaf) ; where the bar is not shown, it is within the symbol. 67 A. Tobacco GABA • »i « Ala .. B. Tobacco 3 6 9 160 NADPH 0*e&-~*—° ^—•—•—• -o ' IS o + •73 40 NADP * u 20 J> ^^ NADPH/ NADP* .2 4 Sf^—0 °^*-»-«-*-* 0 4 8 12 0 4 8 12 16 0 4 Time(h) 68 A. Arabidopsis Drought 0.08 0.04 0.00 0 3 6 0 3 6 Time(h) Time (h) B. Arabidopsis 4°C i 0.04 \V- 0.02 : level s GLYR1 > 0.00 i ' •• . H3 GZ77J.2 4e-6 Rel ; 2e-6 /•>^: 0 2 4 6 Time(h) Fig. 3.4. Response of GLYR transcripts in mature rosette leaves of Arabidopsis plants subjected to salinity, drought, submergence, cold or heat. Closed and open circles represent control and experimental plants, respectively. Data represent the mean ± SE (n= 3, each sample consists of the leaves from 3-4 pooled rosettes); where the bar is not shown, it is within the symbol. 69 GABA is a 4-carbon non-protein amino acid found in virtually all prokaryotic and eukaryotic organisms (Shelp et al, 1999). In plants, GABA is derived from Glu via the enzyme Glu decarboxylase (GAD). GAD is a cytosolic enzyme with an acidic pH optimum, which is stimulated by the increasing cytosolic H+ and Ca + levels that often accompany stress. Ca2+ in turn forms a complex with calmodulin, which binds to GAD, thereby relieving the enzyme from autoinhibition and causing the accumulation of GABA (Baum et al, 1993; Ling et al, 1994; Arazi et al, 1995; Snedden et al, 1995, 1996). A large number of studies have reported the accumulation of GABA and the concomitant loss of Glu in plant tissues and transport fluids in response to many abiotic stresses, including temperature shock, oxygen deficiency, cytosolic acidification, water stress and UV stress (Girousse et al, 1996; Valle et al, 1998; Allan et al, 2003a, b; Bartyzel et al, 2003/4; Breitkreuz et al, 2003; Fait et al, 2005; Ricoult et al, 2005; Kato-Noguchi and Ohashi, 2006; Mazzucotelli etal, 2006; Kaplan et al., 2007; see also refs in Bown and Shelp, 1989, 1997; Shelp etal, 1999). Moreover, antisense suppression of GAD results in the accumulation of Glu in transgenic tomato fruit (Kisaka et al, 2006). In the present report, oxygen deficiency (submergence in water), water stress (salinity, drought) or temperature stress (4 °C, 40 °C) caused similar changes in the pool sizes of GABA and Glu in leaves of Arabidopsis and tobacco plants; the only possible exception was the relatively stable pool size of Glu at 4 °C (Fig. 3.1). It is noteworthy that all stresses tested, with the exception of Arabidopsis under submergence, resulted in the accumulation of Pro, findings that are consistent with the impact of abiotic stress in the literature (e.g., Kaplan et al. 2004, 2007). 70 SSA is produced from GABA via GABA transaminase (GABA-T), an enzyme that can apparently utilize either pyruvate or 2-oxoglutarate as an amino acceptor, resulting in the production of Ala and Glu, respectively (Shelp et al, 1995; Van Cauwenberghe and Shelp, 1999; Van Cauwenberghe et al, 2002). Evidence indicates that 1) Ala accumulation, rather than Glu accumulation, typically occurs in response to oxygen deficiency (see references in Bown and Shelp, 1989 and Miyashita et al, 2007), 2) the primary role of an inducible Ala aminotransferase is to degrade Ala when it is in excess, for example, during recovery from oxygen deficiency (Miyashita et al, 2007), and 3) hypoxia-induced Ala accumulation is partially inhibited in roots of gadl and gaba-tl KO mutants (Miyashita and Good, 2008), indicating that pyruvate-dependent GABA-T reaction contributes to Ala formation during oxygen deficiency. In the present paper, Ala accumulation accompanied the accumulation of GABA during exposure to four of the five stresses (the exception being salinity) (Fig. 3.1). Since GABA-T appears to be mitochondrial (Breitkreuz and Shelp, 1995; Van Cauwenberghe et al, 2002) GABA probably enters the mitochondrion, where it is converted to SSA, and then to succinate via an NAD-dependent SSA dehydrogenase (SSADH) (Breitkreuz and Shelp, 1995; Busch and Fromm, 1999). Biochemical characterization of plant SSADH revealed strong inhibition by NADH, as well as ADP and ATP (Busch and Fromm, 1999). Thus, under stress conditions such as oxygen deficiency, when the mitochondrion exhibits higher ratios of NADH:NAD+ and ADP:ATP, SSADH activity is probably restricted, contributing to the accumulation of SSA and feedback inhibition of GABA-T (Shelp et al, 1995; Busch and Fromm, 1999; Van Cauwenberghe and Shelp, 1999). Loss-of-function ssadh mutants are 71 phenotypically less fit than wt Arabidopsis, exhibiting dwarf stature and less leaf area and chlorophyll concentration (Bouche et al, 2003). When these mutant plants are exposed to abiotic stresses, specifically high UV light and high temperature, they develop necrotic lesions and produce elevated levels of hydrogen peroxide, which is a reactive oxygen species. Stress can cause the influx of Ca2+ and H+ into plant cells, thereby activating a number of metabolic pathways such as that involving GAB A (Shelp et al. 1999; Buchanan et al, 2000). In addition, Ca2+/calmodulin activates NADK, an enzyme responsible for adding phosphate to NAD+ or NADH to make NADP+ and NADPH, respectively (Harding et al, 1997; Hunt et al., 2004). In stressed plants, NADPH is required as a substrate for NADPH oxidase, a membrane protein that generates reactive oxygen species and results in activation of many stress tolerance genes and pathways in plants (Hunt et al, 2004). A balanced NAD(P)H:NAD(P)+ ratio is essential for efficient functioning of the electron transport chains on the mitochondrial and chloroplast membranes (Noctor, 2006). In the mitochondrion and chloroplast, NAD(P)+ acts as an electron acceptor for generating NAD(P)H, and when these reducing equivalents accumulate under oxidative stress damage could occur due to limited availability of the NAD(P)+ (Scheibe et al, 2005). Our studies confirmed that the NADPH/NADP"1" ratios increase in response to oxygen deficiency and temperature stress (Fig. 3.2), and provide support for the involvement of redox balance in the response to stress. 3.7.2. GHB accumulation is a general response to abiotic stress 72 Other research suggests an additional coping mechanism for the detoxification of SSA, which involves its reduction to GHB. In animals, GAB A functions as a signaling molecule during neurotransmission. Like plants, GABA in humans is catabolized in mitochondria to provide succinate to the Krebs cycle through an oxidative pathway (Maitre, 1997). Under oxygen deficiency, SSA is diverted to the production of GHB (Mamelak, 1989), a reaction catalyzed via the enzyme SSA reductase, which is localized in the cytosol and uses NADPH as a cofactor (Schaller et al, 1999). GHB, a short chain fatty acid similar in structure to GABA, is normally present at about 1% of the GABA level (Pearl et al, 2003). This reductive pathway is proposed to minimize energy demands of the brain under stress (Mamelak, 1989). Accumulation of GHB in the brain due to a non-functional SSADH (i.e., recessive disorder called GHB aciduria) is associated with progressive mental retardation in humans. This phenotype is thought to be attributed to GHB competition with GABA for GABA receptors in the brain, thereby disrupting normal neural transmission (Pearl et al, 2003). The first evidence for the occurrence of GHB in plants was presented in 2003 (Allan et al, 2003 a, b; Breitkreuz et al, 2003). Oxygen deficiency increases GHB concentrations from about 10 to 155 nmol g"1 FM in soybean sprouts, and from 273 to 739 nmol g"1 DM in green tea leaves (Allan et al, 2003 a). A cDNA (initially designated as y- hydroxybutyrate dehydrogenase, but later renamed glyoxylate reductase 1 (GLYR1; Hoover et al,2007a)), which encodes a 289-amino acid polypeptide, was identified via complementation of an SSADH-deficient yeast mutant with an Arabidopsis cDNA library (Breitkreuz et al, 2003). Overexpression of the cDNA enables growth of the mutant yeast on GABA and results in the accumulation of 73 GHB. Furthermore, the concentrations of GHB and GABA increase in Arabidopsis plants during submergence, a condition that should increase the cellular NADH:NAD+ ratio, thereby inhibiting SSADH activity. Other work revealed that 1) ssadh mutant Arabidopsis plants, grown under high UV light, have five times the normal level of GHB and high levels of ROS (Fait et al, 2005), and 2) the pattern of GHB in cold- acclimated Arabidopsis plants is consistent with the rise and fall of GABA (Kaplan et al, 2007). The present study provided the first evidence for GHB accumulation in response to water stress (salinity, drought) and heat stress, and confirmed that GHB accumulates under oxygen deficiency (submergence) and cold stress (Fig. 3.1). Together, these data indicate that GHB accumulation is a general response to abiotic stress. 3.7.3. Recovery of metabolite and redox levels after exposure to abiotic stress Wallace et al. (1984) reported that changes in amino acid composition of leaflets of soybean plants exposed to 6 °C for 8 min are fully reversible within 1 h after being returned to their original growing conditions at 33 °C. Recently, Miyashita et al. (2007) showed that Ala levels in Arabidopsis plants return to normal within 24 h following a 24-h period in 5% oxygen; GABA was not assayed and Glu was unaffected by the stress. In the present study, the levels of GABA, Ala, GHB and phosphorylated pyridine nucleotides (but not Glu) in mature tobacco leaves approached normality within 9 h following a 3-h period of submergence; 9 h was clearly inadequate for recovery of metabolite and redox levels after 6 or 9 h of submergence (Fig. 3.3). 74 3.7.4. Role of GLYR isoforms in detoxification of SSA and glyoxylate Recently, Hoover et al (2007a) expressed the Arabidopsis GLYR1 cDNA in E. coli and purified the recombinant protein to homogeneity. Kinetic studies confirmed that the protein reduces SSA to GHB (Km = 0.87 mM) in an NADPH-dependent and essentially irreversible reaction. Unexpectedly, further tests of substrate preference revealed that the protein also reduces glyoxylate (Km = 4.5 fj,M), with a 250-fold greater preference than for SSA. The enzyme is dramatically inhibited by NADP+, but not GHB (Hoover et al, 2007b). A combination of web-based and recombinant expression tools revealed the existence of a highly homologous protein, designated as Arf3LRY2, which also catalyzes the irreversible, NADPH-dependent reduction of both SSA and glyoxylate to glycolate; GLYR2 has a 350-fold higher preference for glyoxylate than SSA, but the affinity for both substrates was an order of magnitude lower than that for GLYR1 (Simpson et al, 2008). Analysis of the transient expression of recombinant GLYR1 and GLYR2 in tobacco Bright Yellow 2 suspension cells revealed that they are localized to the cytosol and chloroplast, respectively (Simpson et al, 2008). These results are consistent with earlier biochemical evidence, which revealed the existence of distinct cytosolic (Km for glyoxylate of 70-100 uM) and chloroplastic (^m for glyoxylate of 85 u.M) NADPH-dependent GLYRs in spinach (Kleczkowski et al, 1986; also see refs in Givan and Kleczkowski, 1992). Also, the results lend support for the hypothesis that any glyoxylate escaping transamination to Gly in peroxisomes during photorespiration is scavenged (Givan and Kleczkowski 1992; Hoover et al, 2007a; Simpson et al, 2007), thereby preventing the deactivation of the primary 75 photosynthesis enzyme, ribulose bisphosphate carboxylase/oxygenase, by glyoxylate (Campbell and Ogren, 1990). Analysis of specific gene expression using DNA micro-array technology or quantitative real-time RT-PCR, revealed that the abundance of GAD1, GABA-T1 and Ala aminotransferase! transcripts is often elevated in plants subjected to low oxygen, water deficit or salinity (Klok et al., 2002; Cramer et al, 2007). One study, which used semi-quantitative RT-PCR, indicated that the global expression of several putative homologs of GAD and GABA-T are induced in barley during cold acclimation, but only in a frost-resistant cultivar is it maintained during subsequent freezing (Mazzucotelli et al, 2006). Another study utilized micro-array analysis to investigate gene expression in cold-acclimated Arabidopsis; increases in the transcript levels of two GAD genes precede the peak in GABA, thus demonstrating a characteristic transcript abundance- regulated response (Kaplan et al, 2007). GABA-T transcript abundance is unchanged during cold acclimation, whereas SSADH transcript abundance peaks after GABA and succinic semialdehyde reductase (presumably this identification is based on our earlier manuscript; Breitkreuz et al, 2003) is slightly downregulated. Unfortunately, in these two studies it is unclear whether the control plants can adequately account for the conditions present during plant exposure to the cold treatment (i.e., dark). Rice GABA- T expression, as indicated by northern blot analysis, is induced by UV and abscisic acid, as well as blast fungal infection, mechanical damage and salicylic acid, which are commonly associated with biotic stresses (Wu et al, 2006). Previous research from our laboratory suggested that the accumulation of GABA and GHB in Arabidopsis during 76 submergence can not be attributed to the abundance of GABA-T and GLYR1 transcripts, as determined by relative RT-PCR (Breitkreuz et al, 2003). In the present study, quantitative real-time RT-PCR was utilized to improve the specificity, sensitivity and reliability of the gene expression analysis (Gachon et al, 2004), and the transcript abundance of GLYR1 or GLYR2 in Arabidopsis was consistently enhanced in response to the various abiotic stresses tested (submergence, drought, salinity, cold, heat) (Fig. 3.4). Since these abiotic stresses are clearly associated with the accumulation of both GAB A and GHB, the results support the hypothesis that GLYR1 and GLYR2 are involved in the detoxification of SSA in planta, despite their higher affinity for glyoxylate than for SSA in vitro (Hoover et al, 2007a; Simpson et al, 2008). In Chapter 4, KO mutants and additional environmental conditions (i.e., high light and temperature, altered CO2/O2 ratio) that influence photorespiration were utilized to further resolve the roles of GLYR1 and GLYR2 in the detoxification of SSA and glyoxylate in plants. 3.7.5 Summary The aldehyde chemical grouping confers high reactivity on molecules (Bartels, 2003; Kotchoni et al, 2006). Thus, it is believed that: 1) aldehydes that accumulate under some stress conditions are highly toxic, reacting with DNA, oxidizing membrane lipids and modifying proteins (Kotchoni et al, 2006), or influencing the transcription of stress-related genes (Weber et al, 2004), thereby causing cellular and overall developmental problems in the plant; and 2) the induction and operation 77 of reductases under stress conditions should contribute to eradication of aldehydes and redox balance (Oberschall et ah, 2000; Sunkar et ah, 2003). Herein, the imposition of abiotic stress on plants was associated with the accumulation of GHB, the product of SSA reduction, as well as altered redox levels and the induction of GLYR1 and GLYR2 genes. Thus, GHB levels appear to be regulated by a combination of biochemical and transcriptional processes. 78 CHAPTER 4 - Redox Balance and Detoxification of Succinic Semialdehyde and Glyoxylate in Arabidopsis Plants Subjected to Environmental Conditions that Promote y- Aminobutyrate Metabolism or Photorespiration 4.1. Contributions This chapter, which is in preparation for submission to the Journal of Experimental Botany, derives from my primary research project and I had an integral role in planning and deciding the direction of the research. Kevin E. Breitkreuz and Doris Rentsch generated and supplied the homozygous glyrl mutant lines, and Jeffrey C. Waller and Wayne A. Snedden generated and measured NADK activities in nadkl, NADK2 AS and NADK1 OE mutants (Table 4.2), and supplied seeds for my experiments. Jeffrey Simpson and Amanda Rochon analyzed protein expression in glyrl and glyr2 mutant lines (Table 4.1), and Gordon J. Hoover produced and purified the recombinant GLYR1 for in vitro study of substrate specificity (Fig. 4.1). I performed the remaining experiments and data analysis, and prepared the entire manuscript in conjunction with Barry J. Shelp. Wendy L. Allan, Kevin E. Breitkreuz, Jeffrey C. Waller, Jeffrey P. Simpson, Gordon J. Hoover, Amanda Rochon, Doris Rentsch, Wayne A. Snedden, and Barry J. Shelp 79 Department of Plant Agriculture, University ofGuelph, Guelph, ON NIG 2W1, Canada (W.L.A., K.E.B., J.P.S., G.J.H., A.R., B.J.S.); Institute of Plant Sciences, University of Bern, 3013 Bern, Switzerland (K.E.B., D.R.); and Department of Biology, Queen's University, Kingston, ONK7L3N6, Canada (J.C.W., W.A.S.). 4.2. Acknowledgments This work was supported by funds from the Swiss National Foundation (D.R.) and the Natural Sciences and Engineering Research Council of Canada (W.A.S., B.J.S.). K.E.B. and J.C.W. were recipients of an NSERC Canada Postgraduate Scholarship, and J.P.S. was the recipient of an Ontario Graduate Scholarship. 4.3. Abstract SSA is a mitochondrially-generated intermediate in the metabolism of GABA, which accumulates in response to a variety of biotic and abiotic stresses. SSA can be reduced to GHB under oxygen deficiency, drought, salinity, temperature stress and high light conditions. Glyoxylate is a metabolite generated from photorespiratory glycolate. Recent evidence indicates that distinct cytosolic and plastidial glyoxylate reductase isoforms from Arabidopsis thaliana (L.) Heynh (AfGLYRl and AJGLYR2, respectively) catalyse the in vitro conversion of SSA to GHB, as well as glyoxylate to glycolate, via NADPH-dependent reactions. In the present study, recombinant Arabidopsis GLYR1 was demonstrated to catalyze the NADPH-dependent reduction of both glyoxylate and SSA simultaneously to glycolate and GHB, respectively. Time- 80 course experiments revealed that Arabidopsis GLYR KO mutants (glyrl and glyrl), unlike wt plants, have elevated levels of GLYR1 or GLYR2 transcripts, GHB and NADPH in rosette leaves during submergence in water or exposure to low CO2 conditions (75 versus 380 [imol mol"1 at 21% O2). The mutants accumulated glycolate, glyoxylate and succinate as well, but not GLY, under low CO2. Furthermore, manipulation of NAD kinase (NADK) levels and redox balance via the use of Arabidopsis mutants (nadkl and nadk3 KO mutants, NADK overexpressor, and NADK2 underexpressor) altered the plant response to submergence. It can be concluded that in planta GLYR activity is involved in the detoxification of both SSA and glyoxylate, as well as the recycling of phosphorylated pyridine nucleotides, thereby linking GABA metabolism and photorespiration to redox balance. 4.4. Introduction The aldehyde chemical grouping (i.e., H-C=0) is found in many molecules involved in cellular functions. These molecules, which are known as aldehydes, can accumulate and become toxic to the plant because of their extreme reactivity (Kotchoni et al, 2006). They cause cellular and developmental problems that are related to their ability to react with DNA, oxidize membrane lipids, modify proteins, or influence the transcription of stress-related genes (Weber et al, 2004; Kotchoni et al, 2006). Overexpression of oxidative or reductive enzymes that eradicate aldehydes can improve plant tolerance to various stresses (Oberschall et al, 2000; Sunkar et al, 2003). Oxidation and reduction of the aldehyde grouping is accompanied, respectively, 81 by the reduction of NAD(P)+ to NAD(P)H and the oxidation of NAD(P)H to NAD(P)+, indicating that the relative importance of these processes is probably dependent upon the cellular NAD(P)H:NAD(P)+ ratio (e.g., Busch and Fromm, 1999; Hoover et al, 2007a, b), and suggesting that aldehyde metabolism plays a role in redox balance during stress (Bouche et al, 2003; Breitkreuz et al, 2003; Fait et al, 2005). The turnover of pyridine nucleotides involves other enzyme activities as well, including several distinct isoforms of NADK (Harding et al, 1997; Hunt et al,, 2004; Turner et al, 2004). SSA is a mitochondrially-generated intermediate in the metabolism of GABA, which accumulates in response to a variety of biotic and abiotic stresses (Bown and Shelp, 1997; Shelp et al, 1999). SSA is typically oxidized to succinate (Tuin and Shelp, 1994; Bouche et al, 2003), but evidence for the reduction of SSA to GHB during exposure to abiotic stress or high light is also available (Allan et al, 2003a, b; Breitkreuz et al, 2003; Fait et al, 2005). Glyoxylate is a peroxisomally-generated intermediate of glycolate metabolism or photorespiration, a pathway believed to be particularly important under conditions of high light and temperature (Wingler etal, 2000). Glyoxylate is typically transaminated to Gly, but it can also undergo reduction to glycolate (Givan and Kleczkowski, 1992). Recently, we cloned and identified separate Arabidopsis cytosolic and chloroplastic NADPH-dependent GLYR isoforms (A/GLYR1 and A*GLYR2, respectively) (Breitkreuz et al, 2003; Simpson et al, 2008). Kinetic studies revealed that the recombinant proteins of both isoforms prefer glyoxylate (GLYR activity), although SSA can also serve as an effective substrate in vitro (SSAR activity) (Hoover et al, 82 2007a; Simpson et al., 2008). Furthermore, in planta studies of wt Arabidopsis revealed that the accumulation of GHB, like GAB A, is a universal response to abiotic stress, and suggested that GLYR1 and GLYR2 are involved in SSA detoxification (Allan etal., 2008). In this chapter, recombinant A rabidopsis GLYR1 was demonstrated to utilize both glyoxylate and SSA simultaneously, and the use of Arabidopsis GLYR1 and GLYR2 KO {glyrl and glyr2, respectively) established that in planta the proteins are involved in pyridine nucleotide balance and the detoxification of glyoxylate and/or SSA under environmental conditions that promote GABA metabolism or photorespiration. Also, the use of various NADK KO (nadkl, nadk3) and expression (NADK1 overexpressor (OE), NADK2 underexpressor (AS)) mutants revealed biochemical crosstalk between NADK and GLYR activities during submergence stress. 4.5. Materials and Methods 4.5.1. In vitro assay of simultaneous substrate utilization by recombinant Arabidopsis GLYR1 The open reading frame of the Arabidopsis thaliana (L.) Heynh GLYR1 cDNA (GenBank Ace. No. AY044183) was expressed in a recombinant bacterial system as described elsewhere (Hoover et al, 2007a). The recombinant protein was purified to homogeneity and its final concentration estimated using the extinction coefficient computed by the ProtParam tool (http://ca.expasv.org/tools/protparam.html). Enzymatic 83 activity was determined over a 2-min time course as the production of glycolate from glyoxylate or GHB from SSA, which was monitored using HPLC methods that are described below. The 0.8-ml reaction mixture contained 100 mM HEPES (pH 7.8), 20% sorbitol, 10 jiM glyoxylate, 1.0 mM SSA and 0.5 mM NADPH. The assays, which were conducted in capped 1.0-ml plastic tubes in a 30 °C waterbath, was initiated by the addition of 3 nM purified enzyme, and then stopped at 10-s intervals by the addition of cold ethanol to a final concentration of 95%. The samples were placed on ice for 1 h, dried to a residue overnight in a speedvac concentrator, resuspended in 0.5 ml distilled water, and stored at -80°C until analysis. The assays were conducted in triplicate using a single typical enzyme preparation. 4.5.2. Generation and description of transgenic Arabidopsis lines Dr. Kevin Breitkreuz (Universities of Guelph and Bern) identified two dependent AtGLYRl (at locus AT3G25530) KO lines of Arabidopsis ecotype Wassilewskija (hereinafter designated as Atglyrl-lmd Atglyrl-2). A mutant carrying a T-DNA insert in GLYR1 was initially identified by screening superpools of genomic DNA, each representing 1,000 independent transformants, by PCR using Ex-Taq polymerase (Takara Shuzo, Kyoto). Four pairs of primers were used, each including a AtGLYRl gene-specific primer {AtGLYRMX 5'-ATGGAAGTAGGGTTTCTGGGTTTGGG-3' and AtGLYRl-rl 5'-CTCAGCCAATCCAAATGAGTGGATG-3'), and a T-DNA- specific primer (5'-GATGCACTCGAAATCAGCCAATTTTAGAC-3' for the left border of the T-DNA and 5'-TCCTTCAATCGTTGCGGTTCTGTCAGTTC-3' for the right border of the T-DNA). One superpool was identified (AtGLYRl-rl/Left border), 84 which gave a positive signal after southern hybridization with an AtGLYRl cDNA probe. Genomic DNA from each of the 10 pools of 100 plants from which the positive superpool was constituted was screened by PCR. Then, subpools of 20 plants were screened, leading to the identification of a single subpool containing the AtGLYRl disruption mutant. Finally, PCR screening of DNA samples from individual plants led to identification of homozygous plants carrying the insertion in the AtGLYRl gene. The integration of the T-DNA was determined by sequencing PCR product amplified from the region adjacent to the T-DNA (AtGLYR 1-rl/Left border and AtGLYRMl 5'- CTCAGCCAATCCAAATGAGTGGATG-37Left border). This revealed that the T- DNA replaced nucleotides -109 to -67 upstream of the ATG. The plants were grown to homozygosity and then backcrossed twice to the wt to minimize spurious insertions Homozygous seed derived from the two different backcrossing events was a gift of Dr. Doris Rentsch (University of Bern). Homozygous seed of the T-DNA insertional mutant of AtGLYR2 at locus ATIGl7650.1 was available from the Arabidopsis Biological Resource Centre in Arabidopsis ecotype Columbia (SALK_047412.34.95.x KO). The T-DNA strand had inserted within the promoter region at 300 bp from the 5' end of the gene. Dr. Jeff Waller generated homozygous seeds for all the Arabidopsis NADK lines used (except AtNADK3) and seeds were kindly supplied by Dr. Wayne Snedden (Queen's University). The two AtNADK cDNAs for the overexpressed cytosolic Arabidopsis NADK (AtNADKl OE) and for the antisense plastidic Arabidopsis NADK (AtNADK2 AS) were amplified by PCR and cloned into either the BamHl/Sacl or Xbal/Saclsites of the binary vector pBI121 (Clonetech) using the primers 85 (GGATCCATGTCGTCGACCTACAAG-0E-F1 and CATCAGCAGA- TTGGAGCTCTAAGGTC-OE-R1) or (GAGCTCATGTTCCT- ATGCTTTTGCCCTTGC-AS-F2 and TCTAGATCAGAGAGCCTTTTGATCAAG- ACG-AS-R2), respectively. Oligonucleotide primers were designed to amplify the fragments containing the cDNAs with restriction endonuclease sites introduced at the 5'- and 3'-ends. The plasmids containing the chimeric CAM35S..NADKgenes were introduced into Arabidopsis by Agrobacterium tumefaciens (LBA4404)-mediated transformation by the floral dip method (Clough and Bent, 1998). The ArNADKl homozygous T-DNA insertional mutant lines (AfNADKl-11-19-1 and ArfSfADKl-11-4- 3, which were designated hereinafter as nadkl-1 and nadkl-2, respectively), were provided by Dr. Wayne Snedden. The T-DNA strand inserted 722 bp upstream of the ATG start site. Homozygous transgenic lines were selected using kanamycin (Chiasson, 2005). Homozygous seed for the T-DNA insertional mutants in cytosolic NAD(H)K3 (Atnadk3) at locus AtlG78590 (SALK_079342) was obtained from ABRC. This KO was generated in Arabidopsis ecotype Columbia genetic background. Atnadk3 mutant contains a T-DNA strand within an annotated intron. Homozygosity of transgenic lines obtained was confirmed by re-screening for kanamycin resistance. Seeds were surface-sterilized with 20% bleach, washed 3X in sterile distilled water and sown on 0.8% agar plates containing l/2x MS salts and 5 (ig ml"1 kanamycin. After 48 h in the dark at 4 °C, the plates were placed in a growth chamber (Model EY15, 9 1 Controlled Environments Ltd., Winnipeg, Manitoba) providing 100 (JJIIOI m" s" PPFD at the plate level, 22 °C/18 °C temperature, and a 16-h/8-h photoperiod for 2 weeks. 86 Kanamycin-resistant plants were transferred to soil and allowed to grow to maturity. This seed was further screened for kanamycin resistance. 4.5.3. Growth of Arabidopsis Typically, all Arabidopsis lines, but nadk3, were sown in Perlite soil mix (Schundler Co. Edison, New Jersey) in 40-cm3 pots and placed in the dark at 4 °C for 48 h. Then the pots were transferred to environmentally-controlled growth chambers (model EY15, Controlled Environments Ltd., Winnipeg, Manitoba) that supplied a 11-h photoperiod (beginning at 0600 h), 150 pimol m"2 s"1 PPFD at pot level, 22 °C/18 °C temperature, and 65% humidity for 21-28 d. Plants were sub-irrigated every second day and fertilized once weekly with 20-20-20 fertilizer. Seeds of nadk3 were surface-sterilized by immersion in 95% ethanol for 30 s, then soaked in 20% bleach for 15 min and thoroughly washed 3x in sterile distilled water. Individual seeds were sown onto sterile 0.8% agar plates (10-20 seeds/plate) containing 0.5X Murashige and Skoog salts and 3% sucrose, and the plates were sealed with parafilm. The plates were placed in the dark at 4 °C for 48 h and then placed in growth chambers set at the same conditions as the potted plants. To assess NADK activity, wt and mutants plants were sown onto sterile 0.8 % agar plates containing 0.5X Murashige and Skoog media and 1.0% sucrose, and grown with a 8-h photoperiod at 120 fxmol m" s" PPFD and 22 °C. 4.5.4. Analysis of GLYR and NADK expression in Arabidopsis 87 Total RNA was extracted from 3-week-old rosette leaves of soil-grown and surviving plants growing on kanamcyin plates, using Qiagen RNeasy kit (Qiagen, Inc. Mississauga, Ontario), and DNA was digested by DNase I (Invitrogen Canada Inc., Burlington, ON.). Three hundred micrograms of the DNA-free RNA was then converted to cDNA via reverse transcription according to Sambrook and Russell (2001). The RNA was incubated with 1 nmol ml"1 random primers at room temperature for 10 min, and then reverse transcribed using MLV-reverse transcriptase. Levels of mRNA transcripts for GLYR1 and GLYR2 were monitored via real-time PCR according to Chapter 3. For the NADK lines, generated cDNA was amplified via AmpiTaq Applied Biosystems SYBR Green Master Mix Kitpolymerase (Applied Biosystems, Foster City, CA, USA) following the manufacturer's directions. The primers were 5'-GGATGGAGCACTTAGCAAAGTCTCCGCGC-3*- F/3'CCCATAGCA-CTGTGCCATCCCCACCAAGAG-5'-R for AtNADKl; 5'- GGAGGGGAGGACAGG-TAACCCAAGAAGG-3'-F/3'-CCAAGCTTA- GTTTCAGAAACCTCCCGACGC-5'-R for AtNADK2; and, 5'-TCTTCAATA- TCTAGAAAGCAGG-3'-F/3*-CAGCCGATGGCTGT-GCGATCAAAATCGTC-5'-R for AtNADK3. Transcript abundance in each sample was normalized to the corresponding level of 18S ribosomal RNA (Chapter 3). 4.5.5. Measurement of enzyme activity in wild-type Arabidopsis and mutant lines GLYR activity was measured in cell-free crude extracts of wt and GLYR mutant plants using an assay described previously (Hoover et ai, 2007a). Briefly, 1 g of leaf tissue 88 from soil-grown 21-d-old wt, glyrl or glyr2 plants was ground in three volumes of 100 mM Hepes, (pH 7.8), 20% sorbitol, 0.5 mM phenylmethylsulphonyl fluoride, 9 |JM leupeptin, and 1.5 |JM pepstatin using a chilled mortar and pestle. The extract was centrifuged for 10 min (14,000 g, 4 °C), and the supernatant was collected and then desalted using a Sephadex G-25 PD10 column (Bio-Rad Laboratories (Canada) Ltd., Mississauga, Canada) equilibrated with 100 mM Hepes (pH 7.8) and 20% sorbitol. Enzymatic activity was measured continuously as the oxidation of NADPH using a Cary 300 double-beam absorption spectrophotometer (Varian Canada Inc., Mississauga, ON, Canada) set at 340 nm wavelength. The 0.8-ml reaction contained 0.2 ml extract, 100 mM Hepes (pH 7.8) buffer containing 20% sorbitol, and saturating NADPH (50 |iM); activity was determined from the rate before and after the addition of 50 |iM glyoxylate. All measurements were performed in triplicate and three biological replicates were used. Total protein concentration was measured by the Bio- Rad assay method (Bio-Rad Laboratories (Canada) Ltd.), according to the manufacturer's protocol using bovine serum albumin, fraction V (Pierce, Rockford, 111.) as a standard. NADK activity in the presence of Ca2+/CaM was measured in cell-free extracts of seedlings of 2-week-old wt and NADK mutants essentially as described by Turner et al. (2004). The seedlings were quickly harvested, snap-frozen in liquid nitrogen and 100 mg tissue was ground in extraction buffer with sterile sand in a pre-chilled mortar and pestle. The extraction buffer contained 50 mM Tris-Cl, pH 7.5, 200 mM KC1, 3 mM MgCl2,1 mM EGTA, 0.5 mM EDTA, 2% (w/v) polyvinylpolypyrrolidone, 1 mM dithiothreitol, 1 mM phenylmethylsulfonylfluoride, 1 mM benzamidine, 50 fiM 89 leupeptin, and 10/aMN-(N-(L-3-transcarboxyoxirane-2-carbonyl)-L-leucyl) agmatine (E-64). The extract was centrifuged for 5 min at 14,000 g at 4 °C. NADK activity was assayed in microliter plates. The standard assay mixture contained 50 mM Tris-Cl (pH + 7.9), 4 mM ATP, 4 mM NAD , 10 mM MgCl2, 1 mM CaCl2, and 300 nM recombinant petunia CaM81. The reaction was initiated with up to 18 n 1 of extract and allowed to proceed for 30 min at 25°C. NADP formed was immediately detected by adding 20 fi\ of the NADK assay mixture to a cycling assay (250 fil final volume) containing 50 mM Tris-HCl (pH 7.9), 5 mM glucose-6-P, 1 mM EGTA, 0.5 units of yeast glucose-6-P dehydrogenase, 1 mg ml-1 2,6-dichlorophenolindophenol (DCIP), and 1 mg ml-1 methyphenazinium methylsulfate. Reduction of DCIP was monitored at 600 nM using a SpectroMax Plus microplate reader (Molecular Devices, Sunnyvale, CA) and the amount of NADP quantified by comparison to a standard curve produced using analytical-grade NADP. 4.5.6. Exposure of wild-type Arabidopsis and mutant lines to submergence Individually-potted Arabidopsis plants from all lines (i.e., GLYR mutants and corresponding wt; NADK mutants and corresponding wt) were grown in a random arrangement within the growth chamber described above and then selected for size uniformity at the time of experimentation (3-4 weeks). At zero time, five replicate samples, comprised of three to four pooled plants, were collected for each line by rapidly dipping the shoot into liquid nitrogen for 10 s and then clipping the plant at soil level into liquid nitrogen. The remaining plants from all lines were quickly 90 arranged into five blocks, each comprised of two opaque 4-1 containers containing 3-4 plants; one of each pair was filled with tap water at room temperature, and the chamber lights were turned off. The containers were tightly covered with tin foil, and at 2,4 or 6 h, plants were withdrawn from each containers and quickly frozen in liquid nitrogen in situ. Only the rosette leaves were retained and stored at -80 °C until analysis. 4.5.7. Exposure of wild-type Arabidopsis and glyr mutants to photorespiratory conditions Wt and glyr lines, grown as described above, were exposed to varying ratios of C02:02 for 6 h. Three 4-1 gas-tight, transparent plastic containers (Starfrit, Atlantic Promotions, Inc. Longueuil, Quebec) were connected in series via tygon tubing and placed into each of three growth chambers set at 150 (imol m"2 s"1 PPFD, 22 ° C and 65% humidity. Individually-potted plants from each genotype were placed in the open container and allowed to pre-condition for 1 h before the start of gas flow. A prepared gas mixture (Praxair, Mississauga ON, Canada) consisting of 21% O2 and 75, 380 or 1000 |xmol mol"1 CO2, with the balance N2, was passed through the containers in one chamber at a flow-rate of 100 ml min"1. Each container corresponded to one time point in which all the plants were harvested at one time. The experiment was initiated at 1100 h, and plants were harvested every 2 h as described above. Each sample consisted of rosettes from 5-6 pooled plants and the experiment was repeated three times on successive days, 91 with the C02:02 regimes being randomized among the three chambers (N =3 replicates). 4.5.8. Extraction and analysis of GHB, amino acids, organic acids and pyridine nucleotides GHB, amino acids and NADPH/NADP+ pool sizes were extracted and analyzed as previously described (Allan and Shelp, 2006; Chapter 3). For the analysis of NADH/NAD"1", plant tissues were extracted and treated as for the NADPH/NADP"1" cycling assay, except that 100 units of alcohol dehydrogenase from baker's yeast (Sigma-Aldrich, Oakville, ON) and 5.0 mM ethanol were used in the assay instead of glucose-6-phosphate dehydrogenase and glucose-6-phosphate. Organic acids were prepared from frozen tissues as were amino acids and then resuspended in 1.0 ml distilled water. This fraction was applied to a prefilled 2-ml polyprep ion exchange column containing AG50W-X8 resin 200-400 mesh (BioRad Laboratories, Hercules, California, USA) that had been previously equilibrated with 10 ml 1.0 N NH4OH. The column was washed with 5 ml distilled water and the flow- through applied to a similar column containing AG1-X8 resin that had been equilibrated with 1.0 N formic acid. Organic acids were eluted from this column with 10 ml 6.0 N formic acid, dried down in a Speedvac at room temperature overnight, and the dried residue resuspended in 1.0 ml distilled water. The sample was filter-sterilized through a 0.45-|im syringe filter and frozen at -20 °C until analysis. Organic acids were separated on a Ultra-Aqueous C18 column (Restek Corp., Beliefonte, Pennsylvania, 92 USA) with an 1100 series Agilent HPLC system (Agilent Technologies, Inc. Mississauga, ON, Canada) described previously (Allan and Shelp, 2006). The isocratic separation was run with 50 mM potassium phosphate, pH 2.5, as the mobile phase, at 25 °C and a 1.0 ml min"1 flow rate. Organic acids were detected with a diode array detector at 210 nm. Prior to the analysis, standard curves for malate, succinate, 2- oxoglutarate, glycolate and glyoxylate were generated from 0-1000 nmol for each organic acid. Three independent standards were analyzed and averaged to generate each curve. Recovery for these acids was estimated by spiking 40 ml boiling ethanol with 500 nmol each of glycolate and glyoxylate. The aqueous extracts (three) were treated exactly the same as the analyzed plant samples, and the amount of glycolate and glyoxylate recovered in the final extract determined. An average of 480 + 0.2 nmol glycolate and 482 + 0.5 nmol glyoxylate was found, which corresponded to a recovery of 96 % glycolate and glyoxylate. 4.6. Results 4.6.1. Simultaneous utilization of succinic semialdehyde and glyoxylate by recombinant GLYR1 Previous research on the recombinant GLYR1 measured enzymatic activity as the turnover of NADPH in response to the addition of SSA or glyoxylate (Hoover et ah, 2007a). Herein, enzymatic activity was monitored as the simultaneous production of both GHB and glycolate from SSA and glyoxylate, respectively (Fig. 4.1). When both substrates were provided at concentrations near their respective Km values, it is clear 93 0 40 80 120 Time (s) Fig. 4.1. In vitro production of glycolate and GHB by recombinant GLYR1 incubated with subsaturating glyoxylate and SSA and saturating NADPH. Data are from a typical enzyme preparation and measurements were done in triplicate. Closed and open circles represent glycolate and GHB, respectively. 94 that they were utilized by the recombinant protein over the first 60 s of the time course. These results indicate the potential for competition between the two substrates in planta. 4.6.2. GLYR activity in cell-free extracts from leaves of wild-type Arabidopsis and glyr mutants When this work began, glyrl and glyr2 mutants were only available in Wassilewskija and Columbia ecotypes, respectively. Glyoxylate-dependent GLYR activity in cell-free extracts from 21-d-old Arabidopsis rosette leaves was measured as the oxidation of NADPH (Table 4.1). Total GLYR activity in wt plants of both ecotypes was readily detected, but activities in the glyrl-1 and glyrl-2 mutants were decreased by 70% and 64 %, respectively, and in the glyrl mutant, total activity was equivalent to the activity in the Columbia wt. These data indicate that the GLYR1 and GLYR2 activities add up to more than 100% of the activity in the wt lines, and that the isoforms were probably being expressed at higher levels when the complementary form was absent. This interpretation is supported by the rise in GLYR transcript abundance when plants are submerged (see section 4.3.3), and further suggest that the two isoforms compensate for each other in planta. 95 Table 4.1. Total glyoxylate-dependent GLYR activity in cell-free extracts of rosette leaves from wt Arabidopsis and glyr mutants. Data represent the mean + SE of three biological replicates. Ecotype/mutant iGLY R activity (nmol"1 min"1 mg"1 protein] ) (nmol1 min"1 g"1 FM) Wt Wassilewskija 16.1+0.9 74.4 ±4.2 Glyr 1-1 4.8 ± 0.5 22.7 ±3.1 Glyrl-2 5.4 + 0.3 24.3 ±1.4 Wt Columbia 18.2 ±3.9 82.9 ±12.3 Glyrl 18.2 + 2.0 77.8 ± 10.8 96 4.6.3. Response of wild-type Arabidopsis and glyr mutants to submergence A preliminary experiment (4 h after transfer from light to dark) confirmed previous findings (Chapter 3) that the concentrations of GHB and GAB A and the ratio of NADPH/NADP+ in rosette leaves of wt Wassilewskija grown in pots were markedly higher in submerged plants than in control plants stored in the dark at the same temperature (data not shown). In addition, it revealed that GHB concentrations in glyrl- 1 and glyr 1-2 mutants were unaffected by submergence, although the GAB A concentrations did accumulate; for glyrl-1 the NADPH/NADP"1" ratio was unaffected, whereas for glyr 1-2 the ratio was increased, albeit less than that in the wt. Consequently, herein the response of the glyr mutants to submergence was further characterized by determining GLYR transcript abundance and the pool sizes of all pyridine nucleotides (NAD+, NADH, NADP+, NADPH), as well as various metabolites including GHB and its closely related metabolites (GABA, Ala, Glu). A time-course experiment was conducted, involving the rapid transfer of pot-grown plants from light conditions (0 h) to dark (2-, 4- and 6-h controls) or a combination of dark and submergence in water, a design which should distinguish the effect of submergence from that due to the dark environment alone. The expression of GLYR1 and GLYR2 was readily detected in both wt Wassilewskija (Fig. 4.2A) and wt Columbia (Fig. 4.2B) ecotypes using real-time quantitative RT-PCR. Expression of GLYR1 could not be detected in the two glyrl 97 A. Ecotype Wassilewskija B. Ecotype Columbia Ws glyri-i gtyrl-2 Col glyrl 0.04 GLYR: GLYR1 0.02 0.02 0.01 - ^ « 0.00 0.00 GLYR2j 13 5.0e-10 • at 8e-8 4e-8 I 2.5e-10 \ 0.0 O O—1—O O- 0 3 60 3 60 3 6 i0 3 60 3 6 Time (h) Time (h) Fig. 4.2. Response of GLYR transcripts in mature rosette leaves of wild-type Arabidopsis and glyrl (A) and glyrl (B) mutants subjected to submergence. The closed and open circles represent dark control and dark submerged treatments, respectively; if closed symbols are not evident, they are within the open symbol. Data represent the mean ± SE (n= 3, each sample consists of the leaves from 5-6 pooled rosettes); where the bar is not shown, it is within the symbol. 98 mutants, whereas expression of GLYR2 was weak. Furthermore, the expression of GLYR1 but not GLYR2 was evident in glyr2. For all genotypes, submergence transiently increased the expression of GLYR1 and/or GLYR2 to levels above those found in the corresponding dark controls. These results demonstrated that the two Arabidopsis ecotypes under study contain both GLYR isoforms, established that the putative KOs of GLYR1 and GLYR2 indeed lack the appropriate gene function as expected and contain a functional gene for the alternative GLYR isoform, and indicated that transcriptional induction of both GLYR1 and GLYR2 occurs in response to submergence. In wt plants of both ecotypes, submergence led to reductions in the concentrations of NAD+ and NADP+ relative to the dark controls, whereas the NADH and NADPH concentrations increased, resulting in enhanced ratios of NADH/NAD+ and NADPH/NADP+ (Fig. 4.3). In the glyr mutants, the concentrations of NADP+ and NADPH and the ratios of NADPH/NADP"1" in the two mutant types were more variable and less consistent over time than those in the corresponding wt, and therefore the trends were somewhat ambiguous. In contrast to wt plants, the glyrl mutants no longer accumulated GHB in response to submergence, and the glyr2 mutant exhibited a much smaller increase (Fig. 4.4). GABA and Ala accumulation in the mutants also appeared to be more transient or lower in magnitude than in the corresponding wt. In contrast to these metabolites, the concentrations of most primary and photorespiratory amino acids in the wt and glyrl mutants were reduced by submergence; the only exception was Asn in wt Wassilewskija. Furthermore, the concentrations of Glu, Asn, Asp and Gly in the glyr2 mutant were not affected by submergence, but the dark-induced accumulations found in wt Columbia were not evident. 99 A. Ecotype Wassilewskija B. Ecotype Columbia Ws glyrl-1 glyrl-2 Col g/^2 43 500 600 250 300 '^^ ^^< NAD+V- NAD+^~° N^i 0 0 300 300 £—-o-—• « 150 o^r^A 150 OH *3=$4 NADH NADH ^o^ "° 0 0 NADH#v NADH/ I 2 2 o NADV ? NAD+/ 1 1 ^^=P^ HV4 o—*=^=-=8 0 0 80 200 8 ^ 40 -^^3 100 PL, Fig. 4.3. Response of pyridine nucleotides in mature rosette leaves of wild-type Arabidopsis and glyrl (A) and glyr2 (B) mutants subjected to submergence. The closed and open circles represent dark control and dark submerged treatments, respectively; if closed symbols are not evident, they are within the open symbol. Data represent the mean ± SE (n= 3, each sample consists of the leaves from 5-6 pooled rosettes); where the bar is not shown, it is within the symbol. 100 Fig. 4.4. Response of metabolites in mature rosette leaves of wild-type Arabidopsis and glyrl (A) and glyr2 (B) mutants subjected to submergence. The closed and open circles represent dark control and dark submerged treatments, respectively; if closed symbols are not evident, they are within the open symbol. Data represent the mean ± SE (n= 3, each sample consists of the leaves from 5-6 pooled rosettes); where the bar is not shown, it is within the symbol. 101 A. Ecotype Wassilewskija B. Ecotype Columbia Ws glyrJ-J glyrl-2 Col glyri Gin Gin C N^° •^ "^ • Asn O <£ d °~r-o-i--9: Asp /)tt -Csi A: <*^s* GABA OA|A^__5 •-4^ ^=* fc V •°=^=S^ V—°—° 0 3 6 0 3 6 0 3 6 0 3 6 0 3 6 Time (h) Time (h) 102 From these results it is clear that the overall reducing environment in wt plants was enhanced by submergence and mechanisms associated with pyridine nucleotide balance and SSA catabolism during submergence was altered in the glyr mutants. Furthermore, GLYR1 was probably more important than GLYR2 in GHB production, but caution is required when making direct comparisons between the ecotypes. 4.6.4 Response of wild-type Arabidopsis and NADK mutants to submergence NADK mutants were chosen as a strategy to investigate the impact of redox balance on the production of GHB and related pathways. Nadkl and nadk3 mutants were readily available from public resources, whereas an AS mutant of NADK2 and an OE mutant of NADK1 were available from collaborators at Queen's University; all were in the ecotype Columbia background. Two separate experiments were conducted, one dealing with two of the NADK isoforms in pot-grown plants, and another dealing with the third isoform in tissue culture-grown plants. Specific gene expression was checked, and the pool sizes of the pyridine nucleotides and various metabolites were determined under submergence as described above. Real-time quantitative RT-PCR was used to measure the abundance of NADK1, NADK2 or NADK3 transcripts in each mutant. The NADK1 and NADK3 transcripts, respectively, were indeed absent in the nadkl and nadk3 mutants (Fig. 4.5), whereas the NADK1 OE mutant had about three times the level of NADK1 transcript found in the wt. The NADK2 transcript in the NADK2 AS mutant was about one-half the level 103 Wt nadkl-1 nadkl-2 NADK10E Wt NADK2AS Wt nadk3 Fig. 4.5. Relative abundance of NADK1, NADK2 or NADK3 transcripts in wild-type Arabidopsis and various NADK mutants. Three separate experiments were conducted, each consisting of Wt Columbia and the mutants immediately plotted to its right in the graph. Data represent the mean ± SE (n= 3, each sample consists of the leaves from 5-6 pooled rosettes); where the bar is not shown, it is within the symbol. 104 found in the wt. As well, Ca /calmodulin dependent enzyme activity was reduced by 95% in the nadkl mutants and by 26% reduced in the NADK2 AS mutant, and stimulated by 51% in the NADK1 OE line (Table 4.2). These results confirmed that the mutants contain the desired levels of specific gene expression and corresponding enzyme activity. In nadkl and nadk3 mutants, the initial ratios of NADH/NAD+ and ratios of NADPH/NADP"1" in the dark control or under submergence were generally higher and lower, respectively, than those in wt Columbia, a result associated with dramatic declines in NADPH (Fig. 4.6). It is noteworthy that the ratio of NADH/NAD+ at 2 h and the ratio of NADPH/NADP"1" over most of the time course were actually higher under submergence than in the dark, despite the actual decline in NADPH concentration. Antisense expression of NADK2 did not lower the ratio of NADPH/NADP+ in the dark, despite strong declines in both NADP+ and NADPH; however, submergence did enhance the NADPH concentration. Overexpression of NADK1 increased the levels of all pyridine nucleotides in dark and under submergence compared to the wt, although NADPH/NADP"1" actually declined with time in the dark and increased with time under submergence. The concentrations of GHB and related amino acids in wt Columbia were elevated in response to submergence, regardless of whether the plants were pot- or tissue-culture-grown, and the concentrations of primary and photorespiratory amino acids declined (Fig. 4.7). In nadkl, nadk3 and NADK2 AS mutants, the submergence-induced increase in GHB was markedly reduced or delayed, whereas in the NADK1 OE the GHB concentration was dramatically enhanced. The 105 Table 4.2. Ca2+/CaM-dependent NADK activities in cell-free extracts of tissue culture-grown wt Arabidopsis and NADK mutants. Data represent the mean + SE of four biological replicates; each replicate consisted of several pooled seedlings from one Petri dish. Ecotype/mutant NADK activity (jxmol min"1 mg"1 protein) Wt Columbia 0.3824 + 0.0133 nadkl-1 0.0165 + 0.0031 nadkl-2 0.0190 ±0.0017 NADK1OE 0.5782 + 0.0099 NADK2 AS 0.2846 + 0.0388 106 A. Ecotype Columbia (pot-grown) B. (Tissue-culture-grown) Col nadkl-1 nadkl-2 NADK2AS: NADK1 OH Col nadk3 •jg ,-,400 NAD"1' 800 400 NAD+ 8 fe 200 400 200 n ,6» o 1200 600 y "3 600 NADH NADH 300 600 300 A^ PL, «<5r-—• 0 0 0 o 3 .NADH/ 3 NADH/ ,+ 2 a! 2 NAD' NAD+ J, 1 1 c^^: *^fc=* 0 0 1 100 t INADP"1" 400 100 NADP+ 8 E 200 "*N^3 50 ;^x 0 0 1 S-^ 1000 40 i e. B 40 500 NADPH NADPH 0-=*=*=- o 0 0 NADPH/ NADPH/ o 4 E. NADP+ 2 'NADPt^ 1 2 o^-*—* • 0 0 ^ 0 3 60 3 60 3 60 3 60 3 6 0 3 60 3 6 Time(h) Time(h) Fig. 4.6. Response of pyridine nucleotides in mature rosette leaves of wild-type Arabidopsis and various NADK mutants subjected to submergence. The plants were grown in pots (A) or in tissue-culture (B). The closed and open circles represent dark control and dark submerged treatments, respectively; if closed symbols are not evident, they are within the open symbol. Data represent the mean ± SE (n= 3, each sample consists of the leaves from 5-6 pooled rosettes); where the bar is not shown, it is within the symbol. 107 Fig. 4.7. Response of metabolites in mature rosette leaves of wild-type Arabidopsis and various NADK mutants subjected to submergence. The plants were grown in pots (A) or in tissue-culture (B). The closed and open circles represent dark control and dark submerged treatments, respectively; if closed symbols are not evident, they are within the open symbol. Data represent the mean ± SE (n= 3, each sample consists of the leaves from 5-6 pooled rosettes); where the bar is not shown, it is within the symbol. 108 A. Ecotype Columbia (pot-grown) B. Ecotype Columbia (tissue culture) Col nadkl-1 nadkl-2 NADK2 ASl NADK1OE Col nadki Gin 8 Gin 2 1 4 »^\^ 0 '^Cusxsi ^*^. 0 12 Glu 3.0 1.5 °^fc^^ Glu aS 0.0 Asn 0.06 0.4 « 1.5 0 0.0 0.2 Pro 0.2 0.1 0.1 0.0 ~*£s 0.0 0 3 60360 3 60 3 60 36 0 3 60 3 6 Time (h) Time (h) 109 increases in GABA concentrations were smaller and sometimes temporary in the nadkl and nadkS mutants and no increases were found in the NADK2 AS; the NADK1 OE exhibited a strong transient increase in GABA. The accumulation of Ala was delayed or suppressed in all NADK mutants. The concentrations of the primary and photorespiratory amino acids in pot-grown plants of nadkl and NADK2 AS mutants were reduced by submergence much in the same manner as in the corresponding wt plants; the only exception was Gly. The concentrations of these amino acids in tissue- culture-grown plants were relatively unaffected by submergence; the only real exceptions were Ser and Pro in wt and nadk3, respectively. Taken together, these data demonstrated that altered NADPH pools influence GABA metabolism, and that biochemical crosstalk exists between NADK activity and the GABA shunt. 4.6.5. Response of wild-type Arabidopsis and glyr mutants to varying CO2/O2 ratios In these experiments, the impact of varying CO2/O2 ratios on GLYR transcript abundance and the pool sizes of pyridine nucleotides and various metabolites in rosette leaves of Arabidopsis in the light was investigated with two different ecotypes (Wassilewskija and Columbia for assessing glyrl and glyr2 mutants, respectively), and involved the rapid transfer of plants from air to an atmosphere containing 75 (high photorespiration), 380 (moderate photorespiration) or 1000 (negligible photorespiration if any) urnol mol"1 CO2 and 21% O2. As reported above, the expression of GLYR1 was 110 readily detected in both wt Wassilewskija (Fig. 4.8A) and wt Columbia (Fig. 4.8B) ecotypes, whereas expression of GLYR2 was much weaker. Also, GLYR1 and GLYR2 expression could not be detected in the glyrl and glyr2 mutants, respectively, and the expression of GLYR1 in glyr2 was considerably lower than in the wt. It is clear that the expression in wt plants, but not in the glyr mutants, of both GLYR forms was elevated by the lowest CO2 condition; this impact was most evident with GLYRL These results confirmed that the two Arabidopsis ecotypes under study contain both GLYR isoforms, established that the putative KOs of GLYR1 and GLYR2 indeed lack the appropriate gene function as expected and contain a functional gene for the alternative GLYR isoform, and indicated that transcriptional induction of both GLYR1 and GLYR2 occurs in response to low CO2. Exposure of wt Wassilewskija and Columbia to the various gas mixtures demonstrated that the ratios of NADPH/NADP+, but not the NADH/NAD+, are inversely correlated to the CO2 concentration, a relationship that disappeared in the glyr mutants (Fig. 4.9). Furthermore, the concentrations of Gly (Fig. 4.10 and glycolate (Fig. 4.11) in wt plants of both ecotypes increased as a function of decreasing CO2 concentration; the difference was most pronounced between 75 and 380 (xmol mol" CO2. On the other hand, the concentrations of Ser (Fig. 4.10) and glyoxylate (Fig. 4.11) tended to be lower at 75 ^imol mol"1 than at 380 pimol mol"1 CO2. Since increasing Gly/Ser and glycolate/glyoxylate ratios are well-known markers of photorespiratory 111 A. Ecotype Wassilewskija Ws 0.38 .^ 0:36 GLJR1 0.34 > 3.3e4 :•§• 3 3.0e4 2.7e4 0 3 6 B. Ecotype Columbia slw2 0.04 * 0.03 "•^Sl •—o 0.02 le-4 5e-5 n 6 0 Time (h) Fig. 4.8. Response of GLYR1 or GLYR2 transcripts in mature rosette leaves of wild- type Arabidopsis and glyrl (A) and glyr2 (B) mutants subjected to various CO2 levels. The open circles, closed circles and closed inverted triangles represent 75, 380 and 1000 [j,mol mol"1 CO2, respectively. Data represent the mean ± SE (n= 3, each sample consists of the leaves from 5-6 pooled rosettes); where the bar is not shown, it is within the symbol. 112 Fig. 4.9. Response of pyridine nucleotides in mature rosette leaves of wild-type Arabidopsis and glyrl (A) and glyr2 (B) mutants subjected to various CO2 levels. The open circles, closed circles and closed inverted triangles represent 75, 380 and 1000 [Amol mol"1 CO2, respectively. Data represent the mean ± SE (n= 3, each sample consists of the leaves from 5-6 pooled rosettes); where the bar is not shown, it is within the symbol. 113 Ratio Pyridine nucleotides (nmol g"* FM) Ratio Pyridine nucleotides (nmol g"* FM) tt> Ratio Pyridine nucleotides (nmol g"* FM) ^atl° Pyridine nucleotides (nmol g"* FM) tJ4s- u> Oi P P *- to 'Ji $ I s a H + + s H » •% CD O n © S" 3 9 ST Fig. 4.10. Response of amino acids and GHB in mature rosette leaves of wild-type Arabidopsis and glyrl (A) and glyrl (B) mutants subjected to various CO2 levels. The open circles, closed circles and closed inverted triangles represent 75, 380 and 1000 [xmol mol"1 CO2, respectively. Data represent the mean ± SE (n= 3, each sample consists of the leaves from 5-6 pooled rosettes); where the bar is not shown, it is within the symbol. 115 A. Ecotype Wassilewskija B. Ecotype Columbia Ws glyrl-1 glyrJ-2 Col gtyr2 TAA 140 140 •^q^-* 70 ;*<^ 70 0 0 J l_ 14 14 7 7 • Gin Gin 0 0 20 20 10 10 *£ Glu Glu 0 0 0.2 0.10 •j^S S 0.1 0.05 r^S^^ Asn **f£=r~Z GO 0.0 0.00 4 6 *-=*-^3. !/2 Asp 0 0 o 0.6 1.0 0.3 0.5 ^^^ I GABA GABA 0.0 0.0 12 6 Ala 0 0 0.10 0.08 0.05 0.04 0.00 0.00 8 10 0 12 Ser 6 * 1^-* D U^ Ser 0 0 3 60 3 6036 0 3 60 3 6 Time (h) Time (h) 116 Fig. 4.11. Response of organic acids in mature rosette leaves of wild-type Arabidopsis and glyrl (A) and glyr2 (B) mutants subjected to various CO2 levels. The open circles, closed circles and closed inverted triangles represent 75, 380 and 1000 jimol mol"1 CO2, respectively. Data represent the mean ± SE (n= 3, each sample consists of the leaves from 5-6 pooled rosettes); where the bar is not shown, it is within the symbol. 117 A. Ecotype Wassilewskija B. Ecotype Columbia Col Glyoxylate 0.02 0.00 V^ 0.2 KJlycolate 0.1 0.0 2-OG 0.6 0.3 0.0 6 Succ 0 10 LMal J-—..- -1- 0 3 6 0 3 6 Time (h) 118 conditions (Zelitch, 1973; Novitskaya et al, 2002), it is clear that the wt plants of both ecotypes responded to the CO2 treatments as generally expected. However, it is particularly noteworthy that the Gly concentration (Fig. 4.10) in the glyr mutants did not increase at all in response to low CO2, even though its immediate precursor glyoxylate (Fig. 4.11) exhibited a transient increase; a more dramatic transient increase in glyoxylate was evident at the high CO2 concentration. Glycolate seemed to increase over the time course in the glyr2 mutant, but not the glyrl mutants. Exposure to the low CO2 also resulted in GHB accumulation in wt plants of both ecotypes, but not in the glyrl mutants (Fig. 4.10). Curiously, GHB accumulation was evident in the glyrl mutant but it was associated with the high CO2 concentration. There was no evidence of GAB A and Ala accumulation as an inverse function of CO2 level in wt Wassilewskija and glyrl mutants; indeed the accumulation, if any, seemed to be positively related to CO2 level, an effect that was enhanced in the mutant, at least in the initial part of the time course. By contrast, GABA accumulated in wt Columbia and the glyrl mutant in response to both low and high CO2 levels, whereas Ala accumulation was positively related to CO2 level in the wt, but not in the mutant. There was little impact of CO2 levels on the concentrations of succinate in wt Wassilewskija and wt Columbia, although there were dramatic increases in succinate in the glyr mutants and this was apparently inversely related to CO2 level, particularly in the glyrl mutant. The malate concentrations were slightly increased in wt plants in response to the high CO2, and this was markedly increased in glyrl but not glyrl (Fig. 4.11). Thus, the response of Gly, Ser, glyoxylate and glycolate in both wt ecotypes to varying CO2 conditions, confirmed photorespiratory or non-photorespiratory 119 conditions. The response to varying CO2 conditions of GHB, organic acids, amino acids and reducing equivalents in the glyr mutants differed from that in the wt, indicating that the detoxification of glyoxylate and the regulation of GLYR activity are influenced by the CO2 environment. 4.7. Discussion 4.7.1. Redox balance and detoxification of succinic semialdehyde in Arabidopsis plants exposed to submergence Previous work demonstrated that GHB and NADPH accumulate in response to submergence and abiotic stress in general, and that this response is catalyzed by GLYR1 (Allan et al, 2008). Submergence was utilized to generate oxygen-deficient conditions, thereby restricting mitochondrial electron transport chain activity and the associated recycling of NADH and ATP formation. Under these conditions, NADH would be oxidized by lactate or alcohol dehydrogenase, and the resultant NAD+ would be available for generation of ATP via glycolysis (Sachs et al, 1980). Furthermore, the increasing NADH levels could inhibit SSADH activity (Busch and Fromm, 1999), resulting in elevated levels of SSA. Since SSA is a toxic reactive aldehyde, it would be quickly reduced by NADPH-dependent GLYR activity to GHB. The present study verified that: the NADH/NAD+ ratio rises in the rosette leaves of wt Arabidopsis under submergence (Fig. 4.3), establishing the existence of cellular oxygen deprivation; GHB accumulates in leaves under submergence and this is accompanied by increasing 120 GABA and Ala levels and NADPH/NADP+ ratios (Figs. 4.3 and 4.4); and, the abundance of GLYR1, as well as GLYR 2, transcripts increases under oxygen deficiency (Fig. 4.2). Thus, it can be concluded that both biochemical and transcription mechanisms appear to be involved in the regulation of GLYR activity in planta under the experimental conditions. This conclusion is supported by the use of glyrl and glyr2 mutants (Fig. 4.2; Table 4.1), which did not accumulate GLYR1 and GLYR2 transcripts, respectively, under submergence, and the correlation of these results to the suppression or reduction of GHB accumulation (compare Figs. 4.2 and 4.4). Furthermore, the NADPH/NADP+ ratios and the NADPH levels in glyrl and glyr2 mutants were occasionally altered by submergence, but the trends were quite erratic compared to wt plants (Fig. 4.3), indicating that the loss of GLYR1 or GLYR2 activity influences the recycling of cellular NADPH, thereby linking the reduction of SSA and the GABA shunt to cellular redox balance. SSADH is also known to be involved in cellular redox balance (Bouche et ah, 2003). Arabidopsis ssadh mutants are dwarfed, chlorotic and develop necrotic lesions and reactive oxygen species (ROS) when exposed to high light and heat stress, and the regulation of ROS levels and ROS processing depends on the availability of ascorbate and glutathione, which are maintained by NADPH (Noctor et al., 2006). Interestingly, NADH/NAD+ ratios were not elevated in either glyrl or glyr2 under submergence (Fig. 4.3). Since the NAD+ levels increased under these conditions and the NADP+ pools differed from those in the wt, it seems likely that NAD(H) phosphorylation and turnover was affected. The phosphorylation of NAD+ is catalyzed by NADK activity (Kawai and Murata, 2008), which can be regulated by Ca2+/CaM 121 (Turner et al, 2004), and the transcription of several isoforms is stress-responsive (Berrin et al, 2005; Chai et al, 2005, 2006). Three NADK isoforms are present in Arabidopsis: NADK1 (Berrin et al, 2005; Turner et al, 2004)); NADK2 (Chai et al, 2005); and, NADK3 (Turner et al, 2005; Chai et al, 2006). NADK1 and NADK3 occur in the cytosol and catalyze the phosphorylation of NAD+ and NADH, respectively, whereas NADK2 catalyzes the phosphorylation of NAD+ in the chloroplast. NADK3 is also postulated to be in mitochondria based on its similarity to yeast mitochondrial NADK3 (Pollack et al, 2007). Manipulation of redox balance via the use of a variety of NADK mutants (e.g., Table 4.2) markedly influenced the production of GHB and related amino acids, particularly during submergence (Figs. 4.6 and 4.7), lending further support for the hypothesis that aldehyde detoxification is regulated by redox balance (Hoover et al, 2007a; Simpson et al, 2008) 4.7.2. Redox balance and detoxification of succinic semialdehyde and glyoxylate in Arabidopsis plants exposed to photorespiratory conditions Previously, Hoover et al (2007a) and Simpson et al (2008) demonstrated that recombinant GLYR1 and GLYR2 reduce both glyoxylate and SSA to glycolate and GHB, respectively, in an irreversible NADPH-dependent reaction, although glyoxylate is the preferred substrate. In the present study, it is evident that GLYR1 and GLYR2 could simultaneously reduce glyoxylate and SSA in vitro (Fig. 4.1). Furthermore, glyrl and glyr2 transcripts, as well as glycolate and GHB accumulated in wt plants exposed to low CO2 level, whereas glyoxylate accumulated together with glycolate in glyrl and 122 glyr2 mutants, transcript levels were effectively diminished, and the GHB pool increased in glyr2 (Figs. 4.8,4.10 and 4.11). Concomitantly, the NADPH/NADP+ ratio increased in the wt, but declined in glyrl and was unresponsive to CO2 in glyr2 (Fig. 4.9). These results provide in planta evidence for the detoxification of glyoxylate and the recycling of NADPH by both GLYR1 and GLYR2 under photorespiratory conditions, and for a more prominent role for GLYR1 than GLYR2 in the detoxification of SSA. Photorespiratory conditions place a higher energy demand on CO2 fixation (5.4 mol ATP and 3.5 mol NADPH per CO2 fixed) than non-photorespiratory conditions (3.0 mol ATP and 2.0 mol NADPH per C02 fixed) (Sharkey, 1988; Wingler et al, 2000). It has been suggested that photorespiration is required to maintain electron flow and protect against photoinhibition, especially under stress (Wingler et al, 2000). The reduction of glyoxylate and the concomitant recycling of NADPH may therefore act as an important electron sink under photorespiratory conditions. As well, the C flux through GABA to GHB would further contribute to this sink by the recycling of NADPH in the cytosol. The NADH/NAD+ ratios were reduced under low CO2 in all lines (Fig. 4.9). Thus, SSADH activity is probably not inhibited. Indeed, the significant increase in the succinate pool, in the glyr mutants exposed to low CO2 (Fig. 4.11) suggests that SSA is being detoxified via SSADH when GLYR activity is inhibited. These data lend further support for a role of GABA metabolism in aldehyde detoxification and redox balance. Previously, Fait et al. (2005) utilized ssadh mutants to demonstrate that flux through GABA is involved in SSA detoxification, with the production of ROS and GHB 123 increasing. Inhibition of GABA-T activity by y-vinyl-GABA reduces ROS and GHB accumulation under high light. The prevention of GAB A transamination therefore prevents toxic SSA accumulation and the resultant increase in ROS. Interestingly, the response to low CO2 of the Gly pools in the glyr mutants markedly differed from that in wt plants (Fig 4.10). The absence of Gly accumulation in glyr mutants under low CO2 , together with the continual rise in glycolate, suggests that glycolate provided the carbon backbone for Gly and that GLYR1 and GLYR2 activities were involved in regulating glycolate pools. However, it has been shown that excess glyoxylate can react with ammonium ions to form a carbinolamine, a Ser analog that inactivates Ser:glyoxylate or Glu:glyoxylate aminotransferase and prevents Gly biosynthesis (Igamberdiev and Kleczkowski, 1997). Alternatively, glycolate oxidase may be inhibited by excess glyoxylate, thereby restricting the C flux to Gly synthesis. In any case, the accumulation of glyoxylate in the glyr mutants is associated with the suppression of low CCVinduced Gly synthesis. The impact of GLYR activity on metabolite pools and reducing equivalents in Arabidopsis exposed to high CO2 is unclear. The decline in Gly: Ser ratio and the smaller glyoxylate and glycolate pools are suggestive of suppressed photorespiration in wt plants (Fig. 4.10 and 4.11). However, glyoxylate accumulated in the glyr mutants, albeit transiently in glyrl. Therefore, there may be another source of glyoxylate such as the oxidation of Gly via Gly oxidase in the chloroplast and cytosol as found in wheat or the cleavage of isocitrate via isocitrate lyase into glyoxylate and succinate (Igamberdiev and Kleckowski, 1997). The lower NADPH/NADP+ and NADH/NAD+ ratios in wt plants under high CO2 (Fig. 4.9) indicate a more oxidized cellular environment and 124 possibly inhibition of respiration (Backhausen and Scheibe, 1999). The increases in NADH and NADPH pools in glyrl, and NADH/NAD+ ratio in glyrl (Fig 4.9) suggest that GLYR activity contributes to redox balance under this condition. The dramatic accumulation in malate in glyrl (Fig. 4.11) may suggest that the reduction in GLYR1 activity stimulates NADPH-dependent malate dehydrogenase activity to facilitate the recycling of excess cytosolic NADPH. Furthermore, the accumulation of Ala (Fig. 4.10) is similar to results found under high NAD(P)(H) conditions induced by hypoxia, due to the upregulation of Ala aminotransferase or GABA-T (Limami et al, 2008; Miyashita et al, 2007, 2008). However, in glyrl, NADPH does not accumulate, GHB unexpectedly rises and the malate pool declines. Thus, there is a differential response of the two isoforms to high CO2. While an explanation for this result is not readily apparent, GLYR1 activity may compensate for the lack of GLYR2 activity. Double mutants of glyrl/glyrl are being generated to better understand the roles of the two GLYR isoforms. 4.7.3. Summary In this chapter, recombinant Arabidopsis GLYR1 was demonstrated to competitively utilize both glyoxylate and SSA in vitro, and the use of Arabidopsis glyrl and glyrl mutants indicated that the cytosolic GLYR1 and plastidial GLYR2 are involved in the detoxification of glyoxylate or SSA, as well as redox balance under environmental conditions that promote photorespiration or GABA metabolism. Together, these data indicate that the GLYR isoforms function in the detoxification of aldehydes that are 125 produced in two separate and distinct metabolic pathways. Also, the use of various NADK KO (nadkl, nadk3) and expression (NADK1 overexpressor (OE), NADK2 underexpressor (AS)) mutants revealed biochemical crosstalk between NADK and GLYR activities during submergence stress. 126 CHAPTER 5 - Summary and Future Prospects 5.1. Hypotheses Revisited It is widely acknowledged that in plants, under a plethora of abiotic and biotic stresses, GABA levels rise severalfold due to the oc-decarboxylation of Glu by the Ca /CaM- activated, cytosol-localized GAD (Shelp et al, 1999, and references therein). The first step of GABA catabolism occurs in the mitochondrion by transamination to SSA and Ala or Glu in a reaction catalyzed by pyruvate- or 2-oxoglutarate-dependent GABA-T, respectively (Shelp et al., 1999; Van Cauwenberghe and Shelp, 1999). SSA, a reactive aldehyde, is subsequently oxidized to succinate via mitochondrial NAD+-dependent SSADH (Busch and Fromm, 1999; Shelp et al, 1999). In 2003, Breitkreuz et al. showed, using a yeast complementation strategy, that yeast deficient in SSADH activity, could be complemented with a cDNA from Arabidopsis, allowing the yeast to grow using GABA as it sole nitrogen source. The cDNA was further characterized and shown to encode an NADPH-dependent dehydrogenase that produces GHB. Further work revealed that GHB accumulates in soybean sprouts, tea leaves and Arabidopsis leaves in response to oxygen deficiency, demonstrating for the first time that GHB is produced in planta (Allan et al, 2003; Breitkreuz et ah, 2003). A search of the Genbank database revealed a full-length homolog (GLYR2) with 57% amino acid identity to GLYR1 (Simpson et al, 2008). Recombinant expression of the two cDNAs and biochemical characterization of the resultant proteins revealed that both isoforms are capable of reducing glyoxylate and SSA to glycolate and GHB, respectively, in an 127 irreversible reaction that utilizes NADPH as the preferred cofactor. However, glyoxylate (Km of 4.5 (GLYR1) and 34 (GLYR2) jiM) is preferred over SSA (Km of 0.87 (GLYR1) and 8.96 (GLYR2) mM) (Hoover et al., 2007a, 2007b; Simpson et al, 2008); thus, the two isoforms were renamed glyoxylate reductases. Localization studies illustrated that one isoform (GLYR1) localized to the cytosol and the other (GLYR2) localized to the plastid (Simpson et. al., 2008). The high affinity of both recombinant enzymes for NADPH (Kms of 2.2-2.6 (GLYR1) and 1.2-1.4 |JM (GLYR2)) and the sensitivity of GLYR1 to competitive inhibition by NADP+ suggests that the NADPH/NADP"1" ratio regulates activity in vivo. In this thesis, to investigate the role of the glyoxylate reductases in planta, two hypotheses were formulated and tested: 1) GHB production is a general response to abiotic stress, and is regulated by redox balance and glyoxylate reductase isoforms; and, 2) glyoxylate reductase isoforms function in redox balance and detoxification of both SSA and glyoxylate. As expected, GABA consistently accumulated in Arabidopsis and tobacco leaves in response to oxygen deprivation, drought, temperature stress and salinity in Arabidopsis and tobacco (Chapter 3). Concomitantly, Glu pools declined, Ala and GHB accumulated, and NADPH/NADP+ ratios were enhanced, suggesting that the production of GHB is associated with redox balance. Furthermore, the time required for recovery of GABA and GHB levels in tobacco was positively correlated with the duration of oxygen deficency, lending further support for a link between GHB production and the turnover of reducing equivalents. GLYR isoforms were implicated 128 in GHB production since their transcript levels increased under the various stresses tested. Followup studies, which used glyrl and glyr2 knockout mutants, unequivocally demonstrated that GLYR activity is responsible for the reduction of SSA to GHB under oxygen deficiency and low CO2, and that redox balance is altered in the absence of GLYR activities (Chapter 4). InArabidopsis, the three isoforms of NADK, cytosolic NADK1 and NAD(H)K3 and chloroplast NADK2, are essential for the phosphorylation of NAD+ and have been linked to the plant stress response (Turner et al, 2004, 2005; Chai et al, 2005, 2006). Herein, different NADK mutants were used to assess the impact of altered redox balance on GABA metabolism and GLYR activity. Clearly, GABA and GHB pools were attenuated when NADK1 and NAD(H)K3 activities were suppressed and GHB pools were dramatically elevated when NADK1 was overexpressed, demonstrating crosstalk between NADK activity and GABA metabolism. Glycolate is produced in the chloroplast from the oxygenation of ribulose bisphosphate in a reaction catalyzed by RUBISCO, and is further metabolized to glycerate-3-P in the photorespiratory pathway. Glyoxylate is primarily an oxidized metabolite of glycolate within the peroxisomes. Accumulation of this metabolite is believed to be toxic and therefore it is rapidly metabolized. Herein, it was established that GLYR activity has the capacity to simultaneously reduce both glyoxylate and SSA, while recycling reducing equivalents. In the glyr mutants, unlike wt plants, glyoxylate + increased under low C02, but GHB and the NADPH/NADP ratio did not, demonstrating that GLYR utilizes both SSA and glyoxylate in planta and recycles reducing equivalents (Chapter 4). Thus, GLYR activity serves to detoxify aldehydes 129 that are typically generated in two distinct metabolic paths, GABA metabolism and photorespiration. A subcellular model for the detoxification of photorespiratory gloxylate and GABA-derived SSA is shown in Figure 5.1. When stress is imposed on the cell, cytosolic NADK3 activity is stimulated and the intracellular Ca2+ concentration increases. The Ca2+ complexes with calmodulin, which binds to cytosolic NADK1, plastidial NADK2 and cytosolic GAD, thereby stimulating their activity and elevating cellular NAD(P)H and GABA. GABA is transported into the mitochondrion where it is catabolized to SSA by GABA-T activity. The rise in NADH inhibits SSADH activity and SSA diffuses to the cytosol where it is reduced to GHB via GLYR1. Under photorespiratory conditions, glycolate is oxidized to glyoxylate in the peroxisomes, and it is likely that some glyoxylate diffuses into the cytosol and/or chloroplast to be detoxified by GLYR1 and/or GLYR2, respectively. The activity of GLYR1 and GLYR2 recycles NADPH and thus contributes to redox balance upon recovery from stress. Other enzymatic and non-enzymatic mechanisms exist to maintain a more electronegative redox potential to protect the cell from oxidative stress and to maintain balance between the generation and removal of ROS. The cellular redox potential is usually lower than -200 mV and is maintained by millimolar concentrations of glutathione (Desikan et ah, 2004). In turn, glutathione pools are sustained by the ascorbate-glutathione cycle. Ascorbate is produced from the reduction of monohydroascorbate via NADPH-dependent monodehydroascorbate reductase in the 130 Fig. 5.1. Subcellular model for the detoxification of SSA and glyoxylate produced during GABA metabolism and photorespiration, respectively. Abbreviations: GAD, glutamate decarboxylase; GABA-T, GABA transaminase; GDC, glycine decarboxylase; GGAT, glutamaterglyoxylate aminotransferase; GLYR, glyoxylate reductase; GO, glycolate oxidase; HPR, hydroxypyruvate reductase; NADK, NAD kinase; RUBISCO, ribulose-bisphosphate decarboxylase/oxygenase; SGAT, serinerglyoxylate aminotransferase; SHMT, serine hydroxymethyl transferase; SSADH, succinic semialdehyde dehydrogenase. 131 I Ca^/CaM NAD(H) • NADP(H) Glutamate HH+ ~~N Ca*7CaO M Hi' CO, Glycolate GfflJ GABA Glycolate l\ i-iiiv G /Glycerate 1 ° / vSSA Glyoxylate' NAI>' HPR \ ^GGAr V .P»\l»ll Hydroxypyruvate SGAT^ Glycine. "•2 Glycine GDC" kk* ISADISADIH . TCA Serine AH.V«T Succinate —«, Cycle Peroxisome Serine Miliicliinulriiiii 132 chloroplast or by the spontaneous dismutation of monohydroascorbate to dehydroascorbate which reacts with glutathione to produce ascorbate and oxidized glutathione (GSSG) via dihydroascorbate reductase. The GSSG is reduced by glutathione reductase which consumes NADPH. Ascorbate and glutathione are antioxidants which are involved in ROS removal. Thus, redox balance underpins the enzymatic control of glutathione levels and helps to control ROS levels. Non-enzymatic mechanisms of ROS removal include their neutralization by carotenoids and tocopherols. (Bray et al.,2000; Desikan et al, 2004). 5.2. Future Prospects Like GABA metabolism, photosynthesis is highly susceptible to stress. The flow of electrons through the chloroplast electron transport chain (ETC), from photosystem II to I, must be regulated in order to maintain redox balance. Temperature and water stresses, for examples, can limit the CO2 supply to the Calvin cycle, thereby preventing NADP+ recycling and leading to NADPH accumulation and ETC overreduction (Huner etal., 1998). Light stress can also cause an overreduction of the chloroplast ETC if NADPH is produced in excess of the cell's requirement (i.e., photoinhibition). Future research could investigate whether GLYR1 and GLYR2 contribute to the balance between energy input through photochemistry and energy utilization through metabolism (i.e. photostasis), and prevent the accumulation of glyoxylate and SSA. My previous work used glyrl and glyr2 KOs in Wassilewskija and Columbia backgrounds, respectively. Both glyrl and glyrl mutants are now available in the Columbia ecotype, 133 so that a glyrl/glyr2 double KO could be generated. The levels of GABA/GHB, glyoxylate/glycolate and related amino and organic acids, as well as the redox (including state of photosystems) and energy balance and photosynthetic efficiency could be monitored in mature leaves of wt and single and double KOs subjected to stresses such as temperature (hot and cold), drought, high light, and altered CO2/O2 ratios. If the KOs are more susceptible to stress than wt, it could be concluded that the GLYR isoforms are involved in photostasis and prevent the accumulation of toxic aldehydes. Art3LYR2 is localized to the chloroplast; however, its affinity for glyoxylate and SSA is ten-fold lower than that for GLYR1. Typically, the over-reduction of the chloroplast ETC by NADPH accumulation is probably alleviated by GLYR2, but transgenic plants in which GLYR1 overexpression is targeted to the cytosol or chloroplast could provide additional protection against photoinhibition and other stresses. If GLYR overexpression provides enhanced protection against photoinhibition and other stresses, it could become a strategy for engineering stress resistance in important crop plants. While the the fate of GHB in plants has not yet been studied, research from other organisms provides a useful starting point. For example, in anaerobic bacteria such as Clostridium kluyveri (Kenealy and Waselefsky, 1985; Wolff et ah, 1993; Sohling and Gottschalk, 1996) and Clostridium aminobutyricum (Hardman and Stadtman, 1963), GHB is catabolized to acetate and butyrate. In Alcaligenes eutrophus, GHB is a precursor for poly(3-hydroxybutyrate -co-4-hydroxybutyrate) (Valentin et al., 1995). By contrast, in mammals GHB is catabolized to succinate which feeds into the Krebs 134 cycle (Lyon et al, 2007). The enzyme(s) involved in GHB catabolism are not clear; however, preliminary evidence suggests that the main enzyme is a mitochondrial NAD+ -dependent alcohol dehydrogenase. Also, a mitochondrial 2-oxoglutarate-dependent hydroxyacid-oxoacid transhydrogenase that oxidizes GHB to SSA and D-2- hydroxyglutatarate has been detected in rat liver (see Lyon et al., 2007 and references therein). Future research could compare enzymes involved in bacterial or mammalian pathways to known plant genomic sequences using bioinformatics tools to investigate the existence of highly similar sequences or any annotated electronic data and if so, these sequences could be cloned, corresponding recombinant enzymes expressed and characterized in vitro, and enzyme activities measured in cell-free extracts from plants. Radiolabeled tracer studies could be utilized to determine the flux from GABA to succinate or GHB and to determine precursor-product relations for GHB catabolites. 135 CHAPTER SIX - Literature Cited Akama K, Akihuro T, Kitigawa M, Takaiwa, F. 2001. Rice (Oryza sativa) contains a novel isoform of glutamate decarboxylase that lacks an authentic calmodulin- binding domain at the C-terminus. Biochimica et Biophyica Acta 1522,143-150. Akama K, Takaiwa F. 2007. 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