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Genetic analysis of tocopherol functions in Arabidopsis at low

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presented by

Wan Song

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5/08 KzlProlechresIClRC/Dateoue.indd

GENETIC ANALYSIS OF TOCOPHEROL FUNCTIONS IN ARABIDOPSIS
AT LOW TEMPERATURES

By

Wan Song

A DISSERTATION

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

DOCTOR OF PHILOSOPHY
Genetics

2009

ABSTRACT

GENETIC ANALYSIS OF TOCOPHEROL FUNCTIONS IN ARABIDOPSIS
AT LOW TEMPERATURES

By

Wan Song

Tocopherols (vitamin E) are lipid-soluble antioxidants that are synthesized only by
photosynthetic organisms. Molecular dissection of the tocopherol biosynthetic pathway in
Arabidopsis and Synechocystis and the availability of various mutants containing
different amounts and compositions of tocopherols have greatly facilitated studies
directed at elucidating tocopherol functions in photosynthetic organisms. Blockage in
phloem source-to-sink transportation is a common phenotype shared by multiple
tocopherol-deficient plant species but the molecular mechanism behind the phenomenon
has remained enigmatic. The phenotype in Arabidopsis thaliana tocopherol deficient vte2
mutant is inducible under low temperature (LT) treatment and thus provides an ideal
system to study the relevant tocopherol functions. A series of events occurs in vteZ
mutant, including abnormal transfer cell wall development, vascular callose deposition,
impaired photoassimilate export capacity, sugar and starch accumulation, which
eventually led to growth inhibition during LT adaptation. A forward genetic screen for
mutations that suppress the vte2 LT-induced phenotypes was undertaken in order to
understand the genetic basis of the vte2 phenotype and determine links between
tocopherol deficiency and the vte2 LT-induced phenotypes. Seven independent sve
(suppressor of 3th low temperature-induced phenotype) lines were identified from EMS
mutagenized vte2 population and three lines were selected for further detailed analysis

and molecular cloning based on biochemical characterization of the primary sve lines.

sveI completely suppressed all vteZ LT phenotypes and was found to be a novel allele of
fad2, the endoplasmic reticulum-localized oleate desaturase. sve2 showed partial
suppression and was found to be a new allele of trigalactosyldiacylglycero11 (tng), a
component of the ER-to-plastid lipid ATP-binding cassette (ABC) transporter.
Introduction of tgd2, tgd3, and tgd4 mutations into the vte2 background similarly
suppressed the vte2 LT phenotypes, indicating a key role for lipid transport in this
process. sve7 partially suppressed all vte2 LT phenotypes without impacting fatty acid
and lipid metabolism at permissive temperature. Analyses of the acyl composition of ER-
and plastid-derived lipids before and after LT treatment demonstrated the elevation of
18:2 in phosphotidylcholine is an early and key component in vte2 LT-induced responses
as all suppressors attenuated this change. Identification and characterization of sve loci
highlights the involvement of ER lipid metabolism in tocopherol function in plants. A
global transcript profiling study was carried out to further investigate the transcriptional
effect of tocopherol deficiency and understand the vte2 LT-induced phenotypes. By
comparing vte2 and wild type under different time period of LT treatment, it was shown
that tocopherol deficiency had no effect on gene expression at permissive conditions but
affected a limited number of specific genes after 48h of LT treatment. Based on gene
expression profiles, tocopherol deficiency appeared to result in some degree of oxidative
stress response and influenced the expression of genes involved in cell wall modification
and solute transport in LT-treated vte2. Statistical analyses also highlighted several
potentially important target genes for fiJture studies. These results together provide new

insights into tocopherol functions in LT adaptation in plants.

ACKNOWLEDGEMENTS

First of all I would like to express my deep and sincere gratitude to my advisor Dr.
Dean DellaPenna, professor of Department of Biochemistry and Molecular Biology,
Michigan State University. Dean gave me the opportunity to work in his lab for the past
five years and provided me with many helpful suggestions, important advice and constant
encouragement during the course of this work. I have not only learned how to solve
various problems and get smarter avoiding making the same mistakes the next time but
also from his management of time and handling of multitasks how to be an good and
efficient scientist. I am also grateful for his great support during my breastfeeding and
allowing me to occupy a separate office during my manuscript and thesis writing.

I would also like to thank my past and present guidance committee. Dr. Andreas
Weber, Dr. Rob Last, Dr. John Ohlrogge and Dr. Markus Pauly for their guidance,
encouragement and help through my graduate research.

Special thanks are due to Hiroshi Maeda, for his direct guidance during my rotation
period in the lab and his continuous support and friendship through the end of my thesis
research. His enthusiasm towards scientific research has greatly influenced me.

I would like to thank the former and present members of the DellaPenna Lab,
especially Laura Ullrich, for her patience, kindness and help in genetics; Scott Sattler for
providing the EMS mutagenized vte2 seed and helpful suggestions; Joonyul Kim for his
help in the protein alignment analysis; Payam Mehrshahi and Sabrina Jorge for their great
support and friendship; Laurent Mene-Saffrane for his generosity in sharing of new ideas

and all the other former and present members of the DellaPenna lab, Naoko Kobayashi,

iv

Eun-Ha Kim, Holly Little, Maria Magallanes, Jiying Li for their friendship and
assistance. Sincere thanks are extended to all former and present members of Dr.
Christoph Benning’s lab, especially Changcheng Xu and Eric Moellering for their great
technical assistance in lipid analysis and helpful discussions.

My special appreciation goes to my mother, Sizhen Yao, my sisters and my brother,
though living far away in another country, they always support and encourage me to
focus on scientific research.

I would like to express my heartiest thanks to my friends, Carlene Meijer and Herb
Meijer, for their kindness, affection and encouragement; Feng Shi, Ailing Zhou,
Xiangrong Guo, Xiaohua Wang, and for sharing the parental tips and their friendships
have made me feel that I am not too far away from my parents and country.

Finally, I would like to express special thanks to my husband Zhiyuan Jia. He took
the responsibility of taking care of our son and kept me away from most of family
responsibilities so that I could concentrate on completing this dissertation in the last few
months and supported mentally during the course of this work. Without his help and

encouragement, this study would not have been completed.

TABLE OF CONTENTS

 

 

 

 

 

 

LIST OF TABLES - - VIII
LIST OF FIGURES x
CHAPTER 1 LITERATURE REVIEW 1
TOCOPHEROLS 1
PLANT ADAPTATION TO LOW TEMPERATURE STRESS 12
AIM OF THIS STUDY 22
TABLES AND FIGURES 24

 

CHAPTER 2 GENETIC SCREEN AND PRELIMINARY CHARACTERIZATION
OF SUPPRESSOR OF VTEZ LOW TEMPERATURE-INDUCED PHENOTYPES
30

 

 

 

 

 

 

ABSTRACT 30
INTRODUCTION 32
RESULTS 37
DISCUSSION 43
MATERIALS AND METHODS 47
TABLES AND FIGURES 50

 

CHAPTER 3 CLONING OF ARABIDOPSIS S VE (S UPPRESSORS 0F VTEZ LOW
T EMPERA TURE-INDUCED PHENOTYPE) MUTANTS HIGHLIGHT THE

 

 

 

 

 

INVOLVEMENT OF TOCOPHEROLS IN LIPID METABOLISM 65
ABSTRACT 65
RESULTS 69
DISCUSSION 78
MATERIALS AND METHODS 84
TABLES AND FIGURES 88

 

CHAPTER 4 MICROARRAY ANALYSIS OF THE LOW TEMPERATURE

 

 

 

 

 

 

 

RESPONSE OF THE ARABIDOPSIS VTE2 MUTANT 118
ABSTRACT 1 l8
INTRODUCITON 120
RESULTS 123
DISCUSSION 134
MATERIALS AND METHODS 137
TABLES AND FIGURES - 142

CHAPTER 5 SUMMARY AND FUTURE PERSPECTIVES 161

 

vi

APPENDIX 171
TOCOPHEROLS PLAY A CRUCIAL ROLE IN LOW TEMPERATURE
ADAPTATION AND PHLOEM LOADING IN ARABIDOPSIS - 171

 

 

LITERATURE CITED 255

 

vii

LIST OF TABLES

Table 1.1 Phenotypes of lipid metabolism mutants at low temperature conditions. 24
Table 2.1 A summary of the primary sve lines identified from suppressor screens. 50

Table 2.2 Leaf fatty acid composition of the primary sve lines, Col, vte2 and vteI at
permissive conditions 52

 

Table 2.3 Leaf fatty acid composition of four primary sve lines and Col, vte2 and vte1
after 14 d of LT treatment 53

Table 3.1 Leaf fatty acid composition of sve1 vte2, sve2vte2 and sve7vte2 compared
with Col, vte2 and vte1. 88

 

Table 3.2 Leaf fatty acid composition and photoassimilate export capacity of
sve1vte2 backcross populations. 89

Table 3.3 Acyl compositions of PC, PE, DGDG, MGDG in indicated genotypes before
LT treatment 90

 

Table 3.4 Acyl compositions of PC, PE, DGDG, MGDG in indicated genotypes after 14d
LT treatment 94

 

Table 4.1 The 49 genes significantly upregulated in vte2 relative to Col at 48h of LT
treatment 142

 

Table 4.2 The 28 genes significantly downregulated in vteZ relative to Col at 48h of
LT treatment 144

 

Table 4.3 Genes containing W box sequence (C/T)TGAC(T/ G) in the 1000bp
upstream regions. 145 -

 

Table 4.4 Common 12 genes between the 77 significantly different genes in 48h-LT-
treated vte2 plant and 744 significantly different genes in 3-d-old vte2 seedling.
146

 

Table A.1 Tocopherol and DMPBQ Content in Leaves of Wild Types and Tocopherol
Biosynthetic Mutants Grown Under Permissive Conditions 209

 

viii

Table A2 Content of Photosynthetic Pigments and Tocopherols of Co] and the vte2
and vte1 Mutants After HL1800 Treatment at 22°C. 210

 

Table A.3 Yields and Abortion Rates of Seed Produced During Low Temperature
(75°C) Treatment. 2 1 1

 

Table A4 Content of Photosynthetic Pigments and Tocopherols of C01 and the vte2
and vte1 Mutants Grown at Permissive condition 212

 

Table A5 Content of Individual Photosynthetic Pigments of Col and the vte2 Mutant
During Low Temperature (75°C) Treatment. 213

 

ix

LIST OF FIGURES

Figure 1.1 Structure of tocopherols. ................................................................................. 25

Figure 1.2 Tocopherol biosynthetic pathway and locations of mutations in Arabidopsis
thaliana. .................................................................................................................... 27

Figure 1.3 An abbreviated diagram of two pathways of glycerolipid biosynthesis in

A rabidopsis leaves. ................................................................................................... 28
Figure 2.1 Suppressor screening scheme. ......................................................................... 54
Figure 2.2 An example flat from the first round of suppressor screening. ....................... 55
Figure 2.3 An example image of the second round of suppressor screening. .................. 56

Figure 2.4 Whole plant phenotype of primary lines of svelvteZ-sve 7vte2 compared with
C01, vte2 and vte1 before LT treatment. ................................................................... 57

Figure 2.5 Whole plant phenotype of the primary lines svelvteZ-sve 7vte2 compared with
C01, vte2 and vte1 after LT treatment. ...................................................................... 58

Figure 2.6 Leaf total soluble sugar content in primary svelvte2-sve 7vte2 lines compared
with C01, vte2 and vte1. ............................................................................................. 59

Figure 2.7 Total l4C02 fixation after 7 d of LT treatment of primary svelvteZ-sve 7vte2
lines compared with C01, vte2 and vte1. ................................................................... 60

Figure 2.8 Percent exudation of primary sverteZ-sve 7vte2 lines compared with C01, vte2
and vte1. .................................................................................................................... 61

Figure 2.9 Callose deposition in primary sve] vte2-sve 7vte2 lines compared with Co], vte2
and vte1 after 3 d of LT treatment. ........................................................................... 62

Figure 2.10 Callose deposition in primary sverteZ-sve 7vte2 lines compared with C01,
vte2, and vte1 after 7 d of LT treatment. ................................................................... 63

Figure 2.11 Molar ratios of linoleic acid (18:2) and linolenic acid (18:3) in primary
suppressor lines of sve3vte2, sve4vte2, sve6vte2, sve 7vte2 compared with Col, vte2
and vte1 after 14 d of LT treatment. ......................................................................... 64

Figure 3.1 Whole plant phenotypes of LT treated Col, vte2, svelvteZ, sve2vte2, sve 7vte2,
and the F1 progeny of sverteZ, sve2vte2, sve 7vte2 x vte2. ..................................... 98

Figure 3.2 A demonstration of leaf phenotypes of F2 population from svelvte2 x vte2
backcross. .................................................................................................................. 99

Figure 3.3 LT phenotypes of svelvte2, sve2vte2 and sve 7 vte2 compared with C01, vte2

and We]. .................................................................................................................. 100
Figure 3.4 Complementation test of svelvte2 and fad2-1vte2. ....................................... 102
Figure 3.5 sve2 carries a single recessive mutation in T GD] . ........................................ 105
Figure 3.6 Complementation test of sve2vte2 and tngvteZ. .......................................... 107
Figure 3.7 tgd2, tgd3 and tgd4 also suppress the vte2 LT phenotypes. .......................... 109
Figure 3.8 tgd mutations impact differently on vteZ LT-induced callose deposition. 111

Figure 3.9 Correlation of individual lipid species with photoassimilate export capacity.
................................................................................................................................. 112

Figure 3.10 Callose deposition in leaf vascular tissues of the indicated genotypes after 7 d
of LT treatment. ...................................................................................................... 114

Figure 3.11 Multiple sequence alignment of relevant regions of FAD2 (A) and TGDl (B)

from representative photosynthetic organisms. ...................................................... 115
Figure 3.12 Callose deposition in the indicated genotypes after 7 (1 LT treatment. ....... 116
Figure 3.13 Scattered plot of photoassimilate export capacity versus leaf total sugar

content ..................................................................................................................... 117
Figure 4.1 RNA degradation plot for the 18 arrays used in this study. .......................... 147

Figure 4.2 QC plot of 3’: 5’ ratios for control genes, percentage of present gene calls and
background levels. .................................................................................................. 148

Figure 4.3 Box plot of all perfect match (pm) intensities (Log2) bf unnormalized 18 array
data set. ................................................................................................................... 150

Figure 4.4 Histogram of kernel density estimates of all perfect match (pm) probe
intensities (Log2) of unnormalized l8 array data set .............................................. 151

Figure 4.5 Box plot of all perfect match probe intensities (Log2) of quantile normalized
array data ................................................................................................................. 152

Figure 4.6 Histogram of kernel density estimates of all perfect match probe intensities
(Log2) of quantile normalized 18 array data set. .................................................... 153

Figure 4.7 Cluster dendrogram of a correlation matrix for all-against-all chip
comparisons. ........................................................................................................... l 54

Figure 4.8 A gene tree of the 77 significantly different genes in 48h-LT-treated vte2
relative to C01 .......................................................................................................... 155

xi

Figure 4.9 Normalized expression levels of Msr genes in C01 (gray) and vte2 (purple) at
different time points of LT treatment ...................................................................... 156

Figure 4.10 A heat map of expression of the significantly upregulated 49 genes in 48h
LT-treated vte2 under different conditions. ............................................................ 157

Figure 4.1 1 Whole plant phenotype of Col, vte2, gsl4, gsl4vte2, gsII I, gsl] 1vte2. ....... 159

Figure 4.12 Vascular callose deposition in C01, vte2, gsl4, gsl4vte2, gslI 1 and gsll 1vte2

after 3 d of LT treatment. ........................................................................................ 160
Figure 5.1 A genetic model of vte2 responses under LT treatment. ............................... 170
Figure A.l Tocopherol Biosynthetic Pathway and vte Mutations in Arabidopsis thaliana.

................................................................................................................................. 215
Figure A.2 Phenotypic and Photosynthetic Responses of C01 and the vte2 and vte1

Mutants to HL stress. .............................................................................................. 216
Figure A.3 Visible Phenotype of vte Mutants During Extended Low Temperature

Treatment. ............................................................................................................... 221
Figure A.4 Tocopherol, Lipid Peroxide, Anthocyanin, and Photosynthetic Pigment

Content of C01 and the vte2 Mutant During Four Weeks of Low Temperature

Treatment. ............................................................................................................... 223

Figure A.5 Photosynthetic Status of C01 and the vte2 Mutant During Four Weeks of Low
Temperature Treatment. .......................................................................................... 226

Figure A.6 Changes in Starch and Soluble Sugar levels in C01 and the vte2 Mutant
During Four Weeks of Low Temperature Treatment. ............................................ 229

Figure A.7 Diurnal Changes in Starch and Soluble Sugar levels in C01 and the vte2
Mutant During the First Four Days of Low Temperature Treatment. .................... 231

Figure A.8 Biochemical Phenotypes in Mature and Young Leaves of C01, and the vte2
and vte1 Mutants After Four Weeks of Low Temperature Treatment .................... 233

Figure A.9 Translocation and Export of '4C Labeled Photoassimilates in Low
Temperature-Treated C01 and the vte2 and vte1 Mutants. ...................................... 235

Figure A.10 Aniline Blue Positive Fluorescence in Leaves of C01 and the vte2 Mutant
During Low Temperature Treatment. ..................................................................... 240

Figure A.11 Aniline Blue Positive Fluorescence in Leaves of the vte2 Mutant at Various
Light Intensities under Permissive and Low Temperature Conditions ................... 242

xii

Figure A. 12 Cellular Structure and Immunodetection of Callose in C01 and vte2-I Before
and After 14 Days of Low Temperature Treatment ................................................ 244

Figure A.13 Cellular Structure and Immunodetection of Callose in vteZ-I After 3 Days of
Low Temperature Treatment. ................................................................................. 246

Figure A. 14 Phenotypes of the vte2 Mutant and Col During HL, Drought and Salinity
Stress. ...................................................................................................................... 247

Figure A. 1 5 Phenotypic and Photosynthetic Responses of C01 and the vte2 and vte1
Mutants to HL stress. .............................................................................................. 250

Figure A.16 Aniline Blue Positive Fluorescence in Leaves of the vte2 and vte1 Mutant
During Low Temperature Treatment. ..................................................................... 252

Images in this dissertation are presented in color

xiii

CHAPTER 1 LITERATURE REVIEW

TOCOPHEROLS

Structure, physiochemical and physical properties

Vitamin E is a collective term for a group of eight amphipathic compounds derived
from tocopherols and tocotrienols (0-, 13-, y-, and b-tocopherol and a-, 13-, y-, and a-
tocotrienol). All tocopherol and tocotrienols (“tocochromanols”) possess a chromanol
ring head and 16-carbon hydrophobic side chain tail (Figure 1.1). The fully saturated 16-
carbon side chain of tocopherols is derived from phytol while 16-carbon side chain of
tocotrienols is derived from geranylgeranyl and contains three double bonds at C-3’, C-7’
and C-1 1’. The a-, 8-, y-, 6- isomer of tocochromanols designate the number and position
of methyl substituents attached to the chromanol head group (Schneider, 2005).

The physiochemical roles of tocochromanols as non-enzymatic antioxidants are well
established. Tocochromanols are can quench singlet oxygen I02 and scavenge a variety of
reactive oxygen species (ROS) and free radicals, including the superoxide anion radical

(02'), hydrogen peroxide (H202), and the hydroxyl radical (HO') (Brigelius-Flohe and

Traber, 1999; Wang and Quinn, 2000). The chromanol hydroxyl group is critical for the
antioxidant activity of tocochromanols as it allows donation of a hydrogen atom fi'om this
group (KamalEldin and Appelqvist, 1996). Donation of the chromanol ring hydrogen
generates the tocochromanol radical, which is relatively long lived and stable and can be
converted to tocochromanol by direct interaction with redox-active reagents like
ascorbate or other reductants (Liebler, 1993). In contrast, other aromatic antioxidants

must donate two hydrogen atoms to attain a stable state (Liebler and Burr, 2000). As they

are localized within membranes, tocochromanol primarily act as potent lipid—soluble
antioxidants to prevent free radical damage of polyunsaturated fatty acids (PUFAs) acyl
chains. The oxidation reactions by the free radicals can start a very destructive chain
reaction of lipid peroxidation. By reacting rapidly with fatty acid peroxyl radicals, the
primary products of lipid peroxidation, and converting the lipid peroxyl radicals to lipid
peroxides, tocochromanols terminate the propagation of lipid peroxidation chain
reactions (Burton and Ingold, 1981; Schneider, 2005). The lipid peroxides produced are
enzymatically or non-enzymatically converted to relatively inert hydroxy fatty acids or
other products.

The physical roles of tocochromanols in membranes are less well appreciated.
Various biophysical techniques have shown that the ring hydroxyl of tocochromanols
interact with the polar head groups of lipids while the prenyl chain anchors the molecule
firmly within the phospholipid bilayer orienting the chromanol ring moiety towards the
lipid-water interface of the phospholipid bilayer (Erin et al., 1983; Gorbunov et al., 1991;
Salgado et al., 1993; Atkinson et al., 2008). However, the orientation and depth of
chromonal head group of tocochromanols in the lipid bilayer is yet to be determined
(Fukuzawa, 2008). Studies have shown that tocopherols spontaneously and dynamically
associate with unsaturated lipids (Stillwell et al., 1992). In addition, when incorporated
into membranes, a-tocopherol assumes a negative curvature that is even higher than
cholesterol (Chen and Rand, 1997; Bradford et al., 2003; Atkinson et al., 2008), and as
such may have significant effects on membrane curvature stress and processes such as
vesicle fusion (Churchward et al., 2005). The matching curvatures between (It-tocopherol

(negative) and lysolipids (positive) also contribute to the well-known property of

tocopherols in countering the detergent-like, membrane-destabilizing effects of lysolipids
(Erin et al., 1986). These physical properties underlie the phenomena that tocopherols
preferentially partition into PUFA-enriched membrane domains (SanchezMigallon et al.,
1996) and potentially into those membrane domains sensitive to aspects of lipid packing
and curvature stress (Atkinson et al., 2008). Therefore, although tocopherols are usually
relatively minor membrane component, they may be able to exert a significant structural

role via localized effects.

Tocopherols in animals

Vitamin E is an essential nutrient for humans and animals that must be obtained
through the diet. Vitamin E is taken up as the free alcohol by the intestine, secreted into
chylomicrons and then taken up by the liver. Although all tocochromanols are taken up
and transported to the liver with similar kinetics (Brigelius-Flohe and Traber, 1999), in
the liver, (Jr-tocopherol is specifically bound and transferred by (Jr-tocopherol transfer
protein (orTTP) (Hosomi et al., 1997) and secreted into very-low-density lipoprotein
(VLDL). Metabolism of VLDL results in the delivery of a-tocopherol into low-density
lipoprotein (LDL) and high-density lipoprotein (HDL) (Traber et al., 2004). Chemically,
all tocochromanols have similar antioxidant activity but differ in biological activity as
vitamin E by orders of magnitude due largely to the preferential retaining of (it-tocopherol
by the body. (Jr-tocopherol is thus the biologically most active form of vitamin E,
accounting for over 90% of the vitamin E activity found in tissues (Cohn, 1997).

Vitamin E was first discovered as early as in 1922 as a micronutrient that was
indispensible for reproduction in female rats (Evans and Bishop, 1922). Since then,

symptoms of vitamin E deficiency have been produced experimentally in different animal

species, e.g. muscular dystrophy and encephalomalaecia in chickens (Shih et al., 1977)
and neurologic lesions in rats and monkeys (Towfighi, 1981). In addition, lack of a
functional orTTP gene causes delayed onset of ataxia and retinal degeneration in mice
(Yokota et al., 2001) and an autosomal recessive neurodegenerative disease called ataxia
with isolated vitamin E deficiency (AVED) with symptoms such as hyporeflexia, ataxia,
limitation of upward gaze and profound muscle weakness in humans (Gotoda et al.,
1995). The precise molecular mechanisms of these vitamin E deficiency symptoms,
especially the role of (l-tOCOphCI‘OI in the interplay between nerves and muscles, remain
elusive. While the mechanism for this is still not elucidated, in almost every major
chronic disease, free radical oxidative damage has been implicated (Pratico et al., 1998;
Yokota et al., 2001; Ahuja et al., 2004) and numerous studies suggest that activity of
vitamin E as scavenger of reactive metabolites of oxygen and as an inhibitor of lipid
peroxidation in membrane may play a mechanistic role in its impact on chronic diseases
(Brigelius-Flohe and Traber, 1999).

Increasing evidence is suggesting that in addition to its antioxidant role, vitamin E
also exerts biological effects through non-antioxidant mechanisms. Several studies have
shown that individual tocopherols have physiological activities beyond their action as
antioxidants. a-, but not B— tocopherol, was found to specifically inhibit the enzymatic
activity of protein kinase C (PKC), an important player in cellular signaling in vascular
smooth muscle cells leading to growth arrest (Boscoboinik et al., 1991), probably by
upregulating cytosolic protein phosphatase 2A (PP2A) which leads to increased
dephosphorylation and inactivation of PKC (Ricciarelli et al., 1998). (it-tocopherol was

also shown to specifically modulate expression of various genes, including the SR-A and

CD36 scavenger receptors (specific receptors for oxidized low density lipoprotein) in
smooth muscle cells (Ricciarelli et al., 2000), collagenase in human skin fibroblasts
(Ricciarelli et al., 1999) as well as a-tropomyosin (an actin-binding protein important for
muscle contraction) (Aratri et al., 1999). y-tocopherol also possesses unique features as it
specifically inhibits cyclooxygenase activity and thus has anti-inflammatory properties
(Jiang et al., 2001). All these effects are unrelated to the antioxidant activities of the
molecules, and possibly reflect specific interactions of the individual tocopherols with

enzymes, structural proteins, lipids, or transcription factors (Zingg and Azzi, 2004).

Tocopherol biosynthesis and distribution in plants

Vitamin E compounds are exclusively produced by photosynthetic organisms,
including all plants, algae and most cyanobacteria (Dasilva and Jensen, 1971; Grusak and
DellaPenna, 1999; Horvath et al., 2006). In most dicotyledonous species, tocopherols are
the principle vitamin E components in leaves and seeds while tocotrienols are found in

high amounts in seeds of certain cereal species (DellaPenna, 2005a).

The biosynthetic pathway of tocopherols in Arabidopsis has been elucidated from
radiotracer studies in isolated chloroplasts and cyanobacteria in the early 1970’s
(Whistanc and Threlfal, 1970). During the last 10 years, the enzymes of the core pathway
have been isolated by the combined approaches of genomics, genetics and biochemical
analyses (Shintani and DellaPenna, 1998; DellaPenna, 1999; Collakova and DellaPenna,
2001; Porfirova et al., 2002; Shintani et al., 2002; Cheng et al., 2003; Sattler et al., 2003b;
DellaPenna, 2005b, a). As shown in Figure 1.2, tocopherols are biosynthesized from two
converging pathways, with the head group (homogentisic acid, HGA) produced from

tyrosine catabolism via the shikimate pathway (Herrmann and Weaver, 1999), and the tail

5

group (phytyl diphosphate, PDP) synthesized via the non-mevalonate 2-C-methyl-D-
erythritol 4-phosphate/deoxy-xylulose phosphate (MEP/DOXP) pathway in plastids
(Schwender et al., 1996). In addition, an enzyme encoding a phytol kinase activity
(encoded by VTE5) was recently identified to provide an alternative source of PDP from
chlorophyll hydrolysis for tocopherol synthesis (Valentin et al., 2006). The first
committed step is catalyzed by homogentisate prenyl transferase (HPT, encoded by
VTE2), condensing PDP with HGA to produce 2—methyl-6-phytyl-1,4-benzoquinol
(MPBQ), the first intermediate for the synthesis of all forms of tocopherols. MPBQ can
be methylated by MPBQ methyl transferase (MTl, encoded by VTE3) to produce 2,3-
dimethyl-6-phytyl-l,4-benzoquinol (DMPBQ). Tocopherol cyclase (TC, encoded by
VTEI) can act on either MPBQ or DMPBQ to produce b-tocopherol or y-tocopherol,
respectively. The final production of [ES-tocopherol or or-tocopherol is catalyzed by a
second methyl transferase, y-tocopherol methyltransferase (yTMT, encoded by VTE4)
from 6-tocopherol and y-tocopherol, respectively. In Arabidopsis, or-tocopherol
predominates in photosynthetic tissues while y-tocopherol is the major tocopherol in seed
(Grusak and DellaPenna, 1999). Identification of the genes and rate-limiting activities for
vitamin E biosynthesis have greatly facilitated engineering efforts to dramatically elevate
vitamin E levels in crop species (Collakova and DellaPenna, 2003a; DellaPenna and Last,

2006).

There are large variations in the content and compositions of vitamin E in different
plants and tissues (Grusak and DellaPenna, 1999). Generally 10 to 50 ug vitamin E /g
FW exists in unstressed leaves, with the predominant form being tit-tocopherol. Seeds are

highly variable and contain many forms of tocotrienols or tocopherols (or, [3, y, a), with a

much wider range of content (300—2000 pg vitamin E /g oil(DellaPenna and Last, 2006).
Many studies indicate that tocopherols and tocotrienols are localized in plastids
(Lichtenthaler et al., 1981; Fryer, 1992; Kruk and Strzalka, 1995; Munne-Bosch and
Alegre, 2002). Though plastid envelopes are the primary sites of tocopherol synthesis,
roughly equal amount of a-tocopherol is found in envelopes (Lichtenthaler et al., 1981;
Arango and Heise, 1998) and thylakoid membranes, which include plastoglobuli
(Lichtenthaler et al., 1981; Wise and Naylor, 1987; Havaux, 1998). In addition, high
levels of tocopherols (as much as 38% of the total seed tocopherol content) have been
reported in oil bodies of soybean, oat and sunflower, which are extraplastidic organelles
derived from the ER (Yamauchi and Matsushita, 1976; Fisk et al., 2006; White et al.,
2006). Extra-plastidic localization of low amounts of (rt-tocopherol has also been reported
in vacuoles isolated from barley leaves, mitochondria in spinach leaves and microsomal
membranes in iron-supplemented soybean roots (Dilley and Crane, 1963; Rautenkranz et
al., 1994; Caro and Puntarulo, 1996). These studies have provided evidence that while
tocopherols are certainly synthesized in plastids their localization is not necessarily

restricted to this organelle.

T 0copherol functions in plants

The fully characterized tocopherol biosynthetic pathway and availability of various
mutants defective in specific steps of the core pathway (vte1 through vte4, Figure 1.2)
(Porfirova et al., 2002; Cheng et al., 2003; Sattler et al., 2003b; Sattler et al., 2004) have
greatly facilitated functional studies of tocopherols in plants and begun to uncover the
multi-facet functions (Dormann, 2007; Maeda and DellaPenna, 2007). The Arabidopsis

vte1 mutant, which lacks all forms of tocopherols but accumulate equivalent amounts of

redox active biosynthetic intermediate DMPBQ, and vte2 mutant, which lacks all forms
of tocopherols as well as DMPBQ, have been particularly useful in this regard. Both vte2
and vte1 have been shown to have significantly reduced seed longevity compared to wild
type and this is directly related to the antioxidant function of tocopherols. In addition,
vte2 mutants have severe seedling developmental defects, including impaired cotyledon
expansion and reduced root growth, and contain massive amounts of non-enzymatic lipid
peroxidation products, including lipid peroxides, lipid hydroperoxides, malondialdehyde
(MDA) and phytoprostanes, in comparison to wild type (Sattler etal., 2004; Sattler et al.,
2006). These combined results clearly indicate that a principle function of tocopherols in
plants is to protect PUFAs from nonenzymatic oxidation during seed storage,
germination, and early seedling development.

In mature photosynthetic tissues, tocopherols have long been assumed to provide
photoprotection (Fryer, 1992; Munne-Bosch and Alegre, 2002) since the chloroplast is
one of the major locations where ROS are produced (Knox and Dodge, 1985; Robinson,
1988). Consistent with this theory is the observation that tocopherol levels tend to
increase in response to various abiotic stresses including high-intensity light (Collakova
and DellaPenna, 2003b; Muller-Moule et al., 2003), cold (Maeda et al., 2006), drought
(Munne-Bosch and Alegre, 2000), heat (Bergmuller et al., 2003), osmotic (Abbasi et al.,
2007) and heavy metals (Collin et al., 2008). However, after germination vte2 and vte1

mutant plants are indistinguishable from wild type, suggesting tocopherols are not vital in

Arabidopsis plants under optimal growth conditions (22/18°C, 120 IE rn'2 s-l) (Maeda et

al., 2006). Similarly, vte2 plants were similarly indistinguishable from wild type plants

under drought and salt stress conditions (Maeda et al., 2006). Even when placed under

severe photooxidative stress conditions, the extent of bleaching and photoinhibition in
vte2 and vte1 was similar to wild type (Porfirova et al., 2002; Havaux et al., 2005;
Kanwischer et al., 2005; Maeda et al., 2006). Significant differences were only observed
when high intensity light was combined with low temperature (Havaux et al., 2005),
indicating a more limited role for tocopherols in photoprotection than had been assumed.
In tobacco, VTE2-RNAi lines containing <5% of WT tocopherol levels exhibited
increased lipid peroxidation, enhanced chlorosis and reduced fresh weight under salt or
sorbitol stress while those with >5% of WT tocopherol levels were indistinguishable
from WT (Abbasi et al., 2007). These combined results suggest that the tocopherol
threshold at which oxidative damage is observed is quite low and that the protective roles
of tocopherols in various oxidative stresses may be both stress- and species-specific.

In contrast, when vte2 plants are subjected to non-freezing low temperature (LT)

treatment (3~12°C with 15~200 uE m.2 3.1), they show severely inhibited grth

compared with wild type (Maeda et al., 2006). The visible phenotypes of vte2 plants
during the long-term low temperature treatment include inhibited growth, purple color in
mature leaves, shorter siliques, more aborted seeds and reduced seed yield (Maeda et al.,
2006). vte1 shows an intermediate phenotype between vte2 and wild type, suggesting
DMPBQ can partially replace the functions of tocopherols at LT conditions. Detailed LT
time course experiments defined a series of biochemical and physiological events occur
before visible phenotypes between vte2 and wild type are observed. After as short as 6 h
of LT treatment, photoassimilate export capacity from source (mature leaves) to sink
tissues (young leaves and roots) was impaired in vte2. This coincided with initiation of

callose deposition in leaf vasculature tissues in vte2. After 60 h of treatment, the amount

of soluble sugars in mature leaves of vte2 became significantly higher than in wild type.
After 14 d of LT treatment, the accumulation of starch and anthocyanins in vte2 also
became significantly higher than in wild type. At two weeks of LT treatment, vte2 started

to show a small but significant decrease in quantum yield of photosystem II electron

transport ((Dpsu), indicating a suppression of photochemistry. After 28 d, besides the

obvious inhibited growth, vte2 plants also contained significantly less chlorophylls and

carotenoids and (bpsu was decreased firrther. Interestingly these phenotypes occurred in

LT-treated vte2 without photoinhibition, as the maximum efficiency of photosystem II
(F v/F m) was identical in both vte2 and wild type during 4 weeks at LT. Furthermore, the
levels of lipid peroxides as measure by FOX assay were identical in vte2 and wild type
during the LT treatment (Maeda et al., 2006). Oxidized lipid profiling further confirmed
that there is no significant production of oxidized lipid species in LT-treated vte2 relative
to wild type (Maeda et al., 2008), indicating that vte2 plants are not experiencing severe
oxidative stress during low temperature treatment. These results together suggest an
important role of tocopherols in LT adaptation in plants that is independent of their
antioxidant roles.

Studies in plants other than Arabidopsis have also provided data pertinent to the
functional roles of tocopherols. The maize sde (sucrose export defective 1) mutant, now
known to be a mutant in the maize VTEI (tocopherol cyclase) ortholog, was reported to
have a defect in photoassimilate translocation and accumulate carbohydrate and
anthocyanins in mature leaves under normal grth conditions (Russin et al., 1996; Stitt,
1996; Provencher et al., 2001; Hofius et al., 2004). The potato VTEl-RNAi lines also

display constitutive carbohydrate and anthocyanin accumulation in mature leaves,

10

although the phenotype was only present in lines with >99% reduction in total
tocopherols (Hofius et al., 2004). In both sde and the potato VTEI-RNAi lines, callose
was constitutively deposited in the vasculature of mature leaves (Botha et al., 2000;
Hofius et al., 2004). However, the tobacco VTEI-RNAi line did not show similar
phenotypes either at normal or moderate LT (15°C) conditions (Abbasi et al., 2009),
which may be related with the ultra chilling-sensitivity of the species. Taken together, the
observed phenotypes of Arabidopsis vte2, and to a lesser degree Arabidopsis vte1, though
visible only under low temperature conditions, resemble the visible and biochemical
phenotypes of the maize and potato plants defective in VTEI at normal. temperatures,
indicating such functions of tocopherols may be conserved in plants.

Because tocopherols are synthesized in chloroplasts and photoassimilate
translocation is mediated through transporters at the plasma membrane, an intercellular
signaling function of tocopherols in plants was proposed (Munne-Bosch and Alegre,
2002; Sattler et al., 2003b; Munne-Bosch and Falk, 2004; Munne-Bosch, 2005b), but
evidence supporting such functions and the mechanism underlying the phenomenon is far
from being established. A recent study discovered that although the leaf fatty acid
compositions of vte2 and the wild type are identical at permissive conditions, 14 (1 LT-
treated vte2 had significantly higher and lower linoleic acid (18:2) and linolenic acid
(18:3), respectively, relative to wild type, predominantly in ER-derived lipids (Maeda et
al., 2008). In addition, introduction of fad2 and fad6, mutations in the ER and plastidic
localized oleate desaturases, respectively, suppressed the LT phenotypes of vte2 to
different degrees (Maeda et al., 2008). These new findings suggested a link between

tocopherol deficiency and extraplastidic LT responses of vte2. Further studies on the

11

effects of tocopherol deficiency on lipid metabolism could lead to a better understanding

of the roles of tocopherols in phloem loading and LT adaptation in plants.

PLANT ADAPTATION TO LOW TEMPERATURE STRESS

Exposure to low temperatures frequently occurs in nature and is one of the most
important factors affecting plant performance and distribution. Cold-hardy species,
including Arabidopsis, can increase their freezing tolerance by a period of pre-exposure
to low but non-freezing temperatures, a process known as cold adaptation or cold
acclimation (Thomashow, 1999; Xin and Browse, 2000). Low temperature causes
significant changes in the cell biophysics, and the acclimation process facilitates changes
to plant cell structure, metabolism, and biochemistry that help a plant to efficiently
operate under the new, colder conditions. During cold acclimation, multiple regulatory
and biochemical mechanisms are triggered to optimize grth at LT conditions, and it
has been difficult to determine which processes are directly affected or affected most
severely by LT, and differential responses between species generate complex indirect
effects. A number of genes and cell processes that respond to cold acclimation have been
described, yet there are many aspects of the cold acclimation process that have yet to be

explained.

LT perception and signal transduction

Plants first need to perceive the LT and transduce the signal to alter the expression of
appropriate genes to combat the diverse stresses that LT imposes on living cells.
Molecular and mutational analyses of temperature signaling in Synecocystis PCC6803

demonstrate the existence of at least two temperature sensors. One of the two component

12

regulators, Hik33, is activated by reduced membrane fluidity, allowing the
autophosphorylation of Hik33 and the subsequent transfer of a phosphate group to Hikl9
and finally to the response regulator Revl (Suzuki et al., 2000; Suzuki et al., 2001). In
higher plants, however, the cellular elements that sense LT and initiate the first stages of
LT signaling are not yet identified and details of the early LT-signaling pathway are
missing.

As the most external surface of the plant cell, the cell wall is the first element to
receive a stress signal and begin the signal transmission to the cell interior (Baluska et al.,
2003). The cell wall is intimately associated with both the plasma membrane and
cytoskeleton and communication across the cell wall-plasma membrane-cytoskeleton
continuum facilitates the transmission of cell wall and plasma membrane modifications to
microtubules (Nguema-Ona et al., 2007). Indeed, two of the earliest LT responses are
rigidification of the plasma membrane and remodeling of the cytoskeleton. Chilling
caused depolymerization and disassembly of microtubules (Wang and Nick, 2001). This
initial, partial disassembly of microtubules was shown to trigger efficient cold
acclimation (Orvar et al., 2000; Abdrakhamanova et al., 2003). Microtubule activity can
then mediate Ca2+ channel opening since disassembly of microtubules results in increased
activity of voltage-dependent Ca2+ channels (Thion et al., 1996). Cell wall proteins,
including kinases, extensins (Yoshiba et al., 2001) and arabinogalactan proteins with
glycosyl phosphatidylinositol (GPI) anchors (Humphrey et al., 2007) have also been

proposed to be potential temperature sensors.

Transient influx of Ca2+ into the cytosol is another early event occurring in plants in

response to various abiotic stresses (Monroy and Dhindsa, 1995; Xiong et al., 2002). It

13

was proposed that the opening of Ca2+ channels and Ca2+ influx occur immediately
following membrane and cytoskeleton changes, which in turn triggers protein kinases and
cold-specific signal transduction cascades, leading to the activation of cold-induced genes
and the acquisition of freezing tolerance (Murata and Los, 1997; Sangwan et al., 2001;
Wasteneys and Galway, 2003). The cytosolic Ca2+ burst is shown to be required for
regulating several cold-inducible genes (Monroy et al., 1993; Knight et al., 1996;
Polisensky and Braam, 1996; Tahtiharju et al., 1997). Recently, calmodulin binding
transcription activators (CAMRA) were shown to bind to a regulatory element in the
CBF2 gene promoter (Doherty et al., 2009), providing evidence for a link between
calcium signaling and cold induction of CBF pathway, a key signaling pathway in LT

adaptation.

Signaling pathways in low temperature adaptation

Extensive changes occur in the transcriptome during cold acclimation. Systematic
analysis of expression profiles of large numbers of genes using whole genome arrays
demonstrated that 45% of Arabidopsis transcripts could change in response to low
temperature (Zarka et al., 2003; Vogel et al., 2005). In a recent thorough transcript
profiling study using Affymetrix GeneChips that contain 24,000 genes, as many as 939
genes were determined to be cold- regulated after 24h of cold treatment at 0°C (Lee et al.,
2005)

Multiple regulatory pathways operate in LT adaptation. The best understood system
is the C-repeat Binding Factor (CBF)—dependent signaling pathway (Thomashow, 2001).
In Arabidopsis low temperature is believed to activate the Inducer of CBF Expression-l

(lCEl), a bHLH Goasic helix-loop-helix) transcription factor, which stimulates the

14

transcription of the CBF genes (Chinnusamy et al., 2003). The CBFl, 2, and 3
transcription activators are expressed rapidly in response to LT (within 15 min), binding
to the C-repeat (CRT)/dehydration response element (DRE) present in the promoters of
cold-regulated (COR) and other cold-responsive genes, inducing CBF-targeted genes
(CBF-regulon), which contributes to an increase in freezing tolerance (Thomashow et al.,
1997). The CBF transcription factors are the major regulators of cold acclimation (Cook
et al., 2004) and it was shown that about 10-15% of all the cold-regulated genes belong to
the CBF regulon (Hannah, et a1. 2005). Other transcription factors, including AtMY BIS
(Agarwal et al., 2006) and ZAT12 (Vogel et al., 2005), can negatively regulate the
expression of CBF genes. In addition, studies are identifying other regulatory pathways
that also activate during LT stress. For instance, the eskimoI mutant possesses
constitutive freezing tolerance under both acclimated and non-acclimated conditions
without constitutively expressing COR genes. The ESKIMO I (ESKI) gene product
controls transcription of a set of stress responsive genes that overlap with salt, osmotic
stress and abscisic acid (ABA) induced genes and are largely independent of the genes

regulated by cold acclimation and CBF (Xin and Browse, 1998).

Eflect of LT on plant carbohydrate metabolism

Traditional targeted metabolite studies have established correlations between LT
adaptation and the presence and abundance of certain metabolites. More recently,
metabolite profiling has emerged as an important tool in this regard and the rapid non-
targeted identification and quantification of a wide array of metabolites made it possible
to gain a thorough insight of the metabolic state of the plant and understand the effects of

low temperature stress on plant metabolism (Cook et al., 2004; Wang et al., 2006; Kaplan

15

et al., 2007). The non-targeted metabolite profiling approach revealed that extensive
changes in the metabolome occur in plants at low temperatures and expanded the
inventory of metabolites and pathways linked with cold acclimation (Cook et al., 2004;
Gray and Heath, 2005; Hannah et al., 2006; Kaplan et al., 2007). About 60% of
metabolites were shown to be affected by low temperature with central carbohydrate
metabolism being a particularly prominent component of the metabolome reprogramming

at low temperature (Cook et al., 2004).

It is well known that low temperatures inhibit phloem transport (Giaquinta and
Geiger, 1973; Chamberlain and Spanner, 1978; Krapp and Stitt, 1995; Strand et al.,
1997). Optimal rates of photosynthesis require an appropriate balance between the rates
of carbon fixation, sucrose synthesis and sucrose export (Stitt, 1991). Photosynthesis is
strongly inhibited in Arabidopsis leaves shifted to LT and this inhibition is associated
with the reduced expression of nuclear-encoded photosynthetic genes (Strand et al.,
1997). In the time period of hours to days, LT stress inhibits sucrose synthesis, inducing
accumulation of phosphorylated intermediates and Pi-limitation of photosynthesis. In the
longer term of weeks to months, carbon metabolism in photosynthetic organs is
reprogrammed and redirected into a variety of sugar phosphates and free sugars (Leegood
and Furbank, 1986; Sharkey et al., 1986; Stitt et al., 1988). Studies with Arabidopsis
showed that a sequence of events reverses the inhibition of sucrose synthesis and
photosynthesis as the plants acclimate to low temperatures. Full acclimation occurs in
leaves that developed at LT with the recovery of photosynthetic activity, which has been
shown to be associated with a strong increase in the activities of enzymes of the Calvin

cycle and of the sucrose biosynthetic pathway. These acclimative changes also result in a

16

pronounced shift in partitioning of newly fixed carbon into sucrose (Strand et al., 1999;
Hurry et al., 2000; Hurry et al., 2002). The sugars accumulated in cold-acclimated plants
were proposed to be important in osmoregulation or cryoprotection. Indeed, accumulation
of about 15 metabolites involving in central carbohydrate metabolism, including xylose,
glucose, fructose, galactose, sucrose and raffinose, were identified to be significantly

correlated with acclimated freezing tolerance (Hannah et al., 2006).

Plant glycerolipid biosynthesis and ER-plastid lipid trafficking

Two parallel pathways, prokaryotic and eukaryotic, Operate in plant cells for
glycerolipid biosynthesis and PUFA production (Figure 1.3). All acyl chains of
membrane and storage lipid synthesis are produced in the plastid (Ohlrogge and Browse,
1995). Glycerolipid synthesis starts with two sequential acylations of glycerol-3-
phosphate (G3P) catalyzed by G3P acyltransferase (GPAT) and lyso-phosphatidic acid
(LPA) acyltransferase (LPAAT) producing phosphatidic acid (PA). PA is used to
synthesize phosphatidylglycerol (PG) in all plants and a portion of the plastidic
glycolipids in some plants via the prokaryotic pathway (Ohlrogge and Browse, 1995).
Plastid localized acyl-lipid desaturases, including FAD4, FADS, FAD6, FAD7 and
FAD8, act on the initial 18:1/ 16:0 molecular species of PG or glycerolipids to produce
the highly unsaturated species that are characteristic of chloroplasts (Harwood, 1996).
Alternatively, the 18:1 is hydrolyzed and the resultant acyl-ACP thioesters are exported
from the plastid to enter the ER-localized eukaryotic pathway (Roughan and Slack, 1982;
Browse et al., 1986; Somerville and Browse, 1991). The determination of the membrane
phospholipid composition is completed in the ER compartment by the action of

microsomal acyl-lipid desaturases FAD2 and FAD3, and acyltransferases.

17

Phosphotidylcholine (PC) is the major extraplastidic phospholipid in multi-cellular plants
and main site of acyl desaturation for production of all other molecular species. The
diacylglycerol backbones of PE, PS, PI and storage TAG are also produced by the
eukaryotic pathway. The proportion of nascent FA incorporated into the eukaryotic and
prokaryotic pathways vary among plants and different tissues. In leaves of Arabidopsis, a
typical “16:3” plant, the flux of acyl chains into leaf glycerolipids is approximately 62%
eukaryotic and 38% prokaryotic, with about half of the galactolipid (plastidic) DAG

moieties contributed by the eukaryotic pathway (Browse et al., 1986).

The separation of glycerolipid synthesis activities makes it necessary for lipid
transfer between ER and plastid (Figure 1.3). Reversible exchange of lipids exists
between the eukaryotic and prokaryotic pathways (Browse and Somerville, 1991; Miquel
and Browse, 1992). The eukaryotic pathway produces the “DAG moiety” for plastid
glycolipid synthesis (Slack et al., 1977). Either PA (Ohlrogge and Browse, 1995; Awai et
al., 2006), diacylglycerol (DAG), or lyso-PC (Mongrand et al., 2000; Andersson et al.,
2004) has been hypothesized as the transfer molecule, however, the exact nature of the
lipid molecule transferred from the ER to the outer envelope of the plastid remains
unclear. Studies on the four Arabidopsis trigalactosyldiacylglycerol (tgd1-4) mutants in
recent years shed light on the process of ER-plastid lipid trafficking (Xu et al., 2005;
Awai et al., 2006; Lu et al., 2007a; Xu et al., 2008). A distinct processive galactolipid:
galactolipid galactosyltransferase was presumed to be induced, producing the diagnostic
oligogalactoglycerolipids accumulating in these mutants. TGDl, TGD2 and TGD3 have
been proposed to form an ABC lipid transporter complex in the inner chloroplast

envelope membrane responsible for phosphatidic acid (PA) transfer through the inner

18

envelope membrane (Xu et al., 2005; Awai et al., 2006; Lu et al., 2007a) whereas TGD4
likely associates with the ER membrane and transfers lipids between the ER and outer
plastid envelop membranes (Xu et al., 2008).

The recent demonstration of membrane contact sites (MCRs) between the ER and
plastid in leaf mesophyll cells, so called PLAMs for plastid associated membranes
(Andersson et al., 2007), provides a physical mechanism for the transfer of lipids between
the plastid and the ER (Figure 1.3). PLAMs are likely the plastidic structural and
functional equivalent of mitochondria associated membranes (MAMs, (Ruby et al., 1969;
Pichler et al., 2001; Choi et al., 2006; Goetz and Nabi, 2006) and plasma membrane
associated membranes (PAMs, (Pichler et al., 2001), ER membrane associations with
mitochondria and the plasma membrane, respectively, that are involved in non-vesicular
transport of hydrophobic molecules such as lipids and sterols (Hajnoczky et al., 2002;
Levine, 2004). Transfer lipid molecules may not have to traverse the hydrophilic cytosol
at all, but may travel through a PLAM which may allow direct transfer of lipid molecules

between the ER and plastid membrane systems.

In addition to transfer of lipids between subcellular components, numerous studies in
mammalian, yeast and bacterial systems showed that the acyl chains of phospholipids are
subjected to acyl editing or remodeling, which involves the deacylation and reacylation
of glycerolipids without net lipid synthesis (Macdonald and Sprecher, 1991; Schmid et
al., 1995; Boumann et al., 2003; Kol et al., 2004; de Kroon, 2007; Tanaka et al., 2008).
Recently, lipid remodeling has also been shown in leaves of pea and Brassica napus and
germinating embryos of soybean. Newly synthesized saturated or monounsaturated fatty

acids fi'om plastids are exchanged with PUFAs in PC through an acyl editing process

19

(Figure 1.3), producing initial molecular species of PC containing mixtures of previously
synthesized PUFA with newly synthesized FA (Williams et al., 2000; Bates et al., 2007;
Bates et al., 2009). Acyl editing may play an important role in fine-tuning the fatty acid
composition and optimizing membrane properties in plants under ever-changing growth
conditions and may act as a mechanism to allow enrichment of PUFAs for TAG synthesis

in oilseeds (Bates et al., 2009).

Effect of LT on plant lipid metabolism and lipid metabolism mutants

Normal functioning of integral membrane proteins such as transporters and receptor
proteins depends on maintaining the fluidity of the membrane, which is strongly
influenced at a given temperature by its lipid composition (reviewed in (Hazel, 1995).
One of the best-documented responses of plants to chilling stress is that activities of fatty
acid desaturase enzymes increase (Williams et al., 1992; Palta et al., 1993; Weiss et al.,
1993), resulting in an increase in polyunsaturated acyl chains of membrane lipids during
LT acclimation (Nishida and Murata, 1996; Welti et al., 2002). This kind of modification
promotes membrane fluidity at lower temperatures by decreasing the temperature at
which the membrane lipids experience a gradual phase change from fluid to semi-
crystalline and reduce, which in some cases may eliminate the propensity of cellular
membranes to undergo freezing-induced non-bilayer phase formation (Uemura et al.,

1995).

The importance of membrane lipid composition in temperature tolerance has also
been defined by transgenic studies and mutational analyses (Orvar et al., 2000; Sung et
al., 2003). Overexpressing Arabidopsis FAD7 under the control of 35S promoter caused a

decrease in dienoic fatty acids and an increase in trienoic fatty acids in tobacco, resulting

20

in reduced low-temperature-induced chlorosis in young seedlings (Kodama et al., 1994;
Kodama et al., 1995). Similarly, in transgenic tobacco plants expressing a A-9
cyanobacterial desaturase which introduces a cis—double bond at carbon 9 of both 16- and
18- carbon saturated fatty acids of various membrane lipids, the saturated fatty acid level
was greatly decreased, resulting in a significant increase in chilling resistance
(IshizakiNishizawa et al., 1996). Enhanced cold tolerance was observed in transgenic
tobacco expressing a chloroplast FAD7 gene under the control of a cold-inducible
promoter which resulted in a higher survival rate during a long-term exposure to cold
stress (0-3.5°C) (Khodakovskaya et al., 2006). Conversely, tobacco plants with the FAD7
gene silenced had a lower trienoic fatty acid content than the wild type and were better

able to acclimate to higher temperatures (Murakami et al., 2000).

Mutants affecting fatty acid and lipid metabolism in Arabidopsis (Figure 1.3) have
also provided important insight into the involvement of membrane lipid in LT adaptation.
Five lipid mutants were found to be indistinguishable from wild type at normal growth
conditions and have differing impacts at low temperatures (Table 1.1). The most dramatic
LT mutant phenotypes were observed in fab] and fad2. fab] mutants show reduced
photosynthetic capacity, chlorophyll and chloroplast glycerolipids after 7-10 (1, and
eventually dies after 5 to 7 weeks at 2°C (Wu and Browse, 1995; Wu et al., 1997), while
fad2 leaves gradually deteriorate, displaying patches of necrosis and extensive
accumulation of anthocyanins at 6°C and eventually leading to death of the whole plants
(Miquel et al., 1993). The other three mutants, fad5, fad6, and the triple mutant
fad3fad7fad8, have more moderate phenotypes under LT conditions. fad5 and fad6

develop chlorotic leaves when grown at 5°C with a 20-30% growth reduction and 70%

21

reduction in thylakoid membrane content (Kunst et al., 1989; Hugly and Somerville,

1992). When fad3fad7fad8 is subjected to short term LT, there is only a slight reduction

in (ngu, but under prolonged LT it shows decreased chlorophyll content and also a

reduction in thylakoid membrane content (McConn and Browse, 1996; Routaboul and
Browse, 2002). Surprisingly, a suppressor of fab] which rescued the collapse of
photosynthetic function and allowed survival of fab] plants at 2°C was found to be a new
allele of fad5 (Barkan et al., 2006). fablfad5 suppressors remained viable after 16 weeks
in the cold, were able to resume growth following a return to 22°C and subsequently
produced viable seed. The further increase in lipid saturation in the suppressor fablfad5
challenged the assumptions on the role of thylakoid unsaturation in photosynthetic
function and suggested the involvement of physical properties of different lipid species

and importance of lipid composition for the LT adaptation in plants.

AIM OF THIS STUDY

This thesis research involves a detailed characterization of the LT-induced
phenotypes in vte2 from multiple perspectives. During my rotation and first year of my
thesis research, I collaborated with a senior fellow graduate student (Hiroshi Maeda) to
assess aspects of the vte2 LT phenotypes in detail. I independently assessed the diurnal
changes of leaf soluble sugars and starch levels during the short term time courses of LT
treatment and also participated in the assessment of changes in other biochemical and
physiological parameters, including anthocyanins, tocopherols, carotenoids, lipid

peroxides, and chlorophyll fluorescence [maximum photosynthesis efficiency (Fv/Fm)

and quantum yield of PSII (<1>pgu)]. These time course measurements allowed a

22

biochemical order for the responses of vte2 to LT to be defined, with vascular callose
deposition and defective photoassimilate export capacity appearing first (6h of LT
treatment), followed by the elevated soluble sugars (60h), accumulation of starch and
anthocyanins (14d) and the distinctive growth inhibition (28d) in vte2. This work resulted
in a manuscript detailing these LT-induced phenotypes in vte2 published in Plant Cell in
2006 on which I was the second author (Maeda et al., 2006). This collaborative work laid
the foundation and logic for a suppressor screen to identify and understand the links
between tocopherol deficiency and the vteZ LT phenotypes. This screen became the
major focus of the work described in this thesis.

In fall 2005, I launched an unbiased genetic screen for second-site mutations that
alleviate the vte2 LT-induced phenotypes. Chapter 2 details the screening, identification
and preliminary biochemical, genetic and physiological analyses of a large number of
primary suppressors. Seven sve (suppressors of vte2 LT -induced phenotypes) loci were
defined from which three were selected for molecular cloning efforts detailed in Chapter
3. Chapter 3 provides the detailed characterization of these three sve loci (sveI, sve2 and
sve7) and demonstration of the molecular identities of sve] and sveZ, which together have
provided strong genetic evidence supporting roles for tocopherols in ER lipid
metabolism.

Chapter 4 describes an effort initiated in 2006 to further investigate the influence of
tocopherol deficiency on global gene expression and determine whether the
transcriptional responses could further our understanding of the differing responses of
vte2 and Col during LT treatment. Statistical analysis and insights gained from the

transcriptional profiling studies are detailed in Chapter 4.

23

TABLES AND FIGURES

Table 1.1 Phenotypes of lipid metabolism mutants at low temperature conditions.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

. Phenotype
Mutant Deficrent Effect on lipids at normal ET Phenotype at LT Citation
enzyme(s) . . ( C) condrtrons
conditions ,
fab] 3-ketoacyl- ~40% increase in Similar to 2 Reduced Wu and
acyl carrier 16:0 WT photosynthetic Browse,
protein (both at capacity, 1995; Wu
synthase II 22°C and chlorophyll, and et al.,
12°C) chloroplast 1997
glycerolipids;
Eventually die
fad2 ER o)-6 Substantial Similar to 6 Necrotic leaves Miquel et
desaturase decrease in 18:2 WT with al., 1993
and increase in anthocyanin
18:1 in all the accumulation;
major ER lipids Eventually die
fad5 Chloroplast Absence of 163- Similar to 5 Chlorotic Kunst et
16:0 6-7 6-7,10,13 WT (slight leaves; reduced al., 1989
desaturase chlorophyll growth and
fad6 Chloroplast Substantial ’educmn) thylako‘d Hugly and
. membrane .
(n-6 decrease In 18:3 content Somervrll
desaturase and 16:3 and e, 1992
increase in 18:1
and 16:1
fad3/ ER (fad3) Completely Similar to 4 Short term: Routaboul
fad7/ and deficient in 18:3 WT (slight reduced ‘I’Psu; and
fa d8 chloroplast and 16:3 chlorophyll Lon term' Brwose,
(fad7/8) (”—3 reduction g ' 2002
. decreased
desaturases and being
infertile) chlorophyll and
thylakord
membrane
content

 

24

 

 

0- CH3 CH3

Figure 1.1 Structure of tocopherols.

or-, 13-, y- and 6- tocopherols differ by the number and position of methyl groups on the
chromanol ring. a-tocopherol has a fully methylated chromanol ring while B- and y-
tocopherol contains two methyl groups and b-tocopherol contains only one methyl group.
Tocotrienols have the same basic structure except for the presence of double bonds on the

isoprenoid side-chain at C-3 ’, C-7’ and C-1 1 ’ positions.

25

Figure 1.2 Tocopherol biosynthetic pathway and locations of mutations in
Arabidopsis thaliana.

MPBQ, 2-methyl-6-phytyl—l,4-bcnzoquinol; DMPBQ, 2,3-dimcthyl-6-phytyl-l,4-
benzoquinol; PK, Phytol kinase; HPT, Homogentisate phytyl transferase; TC, tocopherol
cyclase; MT, MPBQ methyltransferase; y-TMT, y-tocopherol methyltransferase; vte1,
vte2, vte3, vte4 and vte5, mutants of TC, HPT, MTl, Y-TMT and PK, respectively. Bold
arrows indicate the major biosynthetic flux leading to accumulation of (Jr-tocopherol in

leaves.

26

 

Shikimate pathway Plastidic MEP/DOXP pathway

OH v v
Chlorophyll a
CH3 CH3 l
PPOWNH
CHZCOOH 3 Phytol

OH vte5 i- pK

Homogentisate Phytyl-diphosphate {—— Phytyl phosphate
vteZ HPT
CH3 CH3
HO / H
3
”93 H3C OH
CH3 CH3
MPBQ I "”1 DMPBQ
vte11- TC vte 1 004% TC

5- tocopherol y- tocophesml

CHavteHzlf Y'TMT vte4 whim-l y-TMT
HO CH3
W HO
H CH3
30 o 3
CH3 H
CH3 MGM
CH 3

B- tocopherol a- tocopherol

Figure 1.2

27

Figure 1.3 An abbreviated diagram of two pathways of glycerolipid biosynthesis in
Arabidopsis leaves.

Adapted from Browse and Somerville (1991) and Benning (2009). For clarity, only the
athways, genes and processes relevant to the work in this thesis are shown. Abbreviations
for the lipid structures: fatty acids - XzY, a fatty acid group containing X carbon atoms
and Y cis double bonds; t16:1, trans-hexadecenoic acid; G3P, glycerol-3-phosphate;
LPA, lysophosphatidic acid; PA, phosphatidic acid; DAG, diacylglycerol; PG,
phosphatidylglycerol; MGDG, monogalactosyldiacylglyccrol; DGDG,
digalactosyldiacylglycerol; SL, sulfoquivosyldiacylglycerol; PC, phosphatidylcholine;
PE, phosphatidylethanolamine; PI, phosphatidylinositol. Acyl editing is shown in which
PC is constantly turned over to LPC and newly exported fatty acids are exchanged
between the PC and the acyl-CoA pools. TGD4 is associated with the endoplasmic
reticulum (ER) and TGDl, TGD2, TGD3 are components of a transporter complex in the
inner envelope membrane. The exact nature and transporting mechanism of the lipid
precursor returned to the plastid is not known. Both acyl editing and ER-plastid lipid

trafficking could occur in the plastid envelop ER-contact sites (PLAMs).

28

 

 

 

 
       
  

Figure 1.3

    
 
  

   

PG, PI PE

 

Endoplasmic reticulum K
fad2

18 2(16 0) )
\raaisf l
\ 18 3(16 0)/ 1

 
   
 
     

 
   
      
   

   

LPA_, PA_., DAG DAG
- 7 " J l
13-1ACP L’ 18:2(16:0)/

G3P 18:1/16:0 18:1/16:0 918:1/16:0 18:2/18:29 18:0
P MGDG DGDG(SL) MGDG 0606

183$“? rad4$ fad5f

fab1 18:1/116:1 18:1/16:1

 

—16:0 ACP PG MGDG

fad6+ fad6+ fad6
Chloroplast
18:2/116:1 18:2/16:2 18:2/16:0 18:2/16:218:2/16:0
PG MGDG DGDG(SL) MGDG DGDG
fad7/8t fad7/8f fad7/8t RIM/8* tam/St

18:3/116:1 18:3/16:3 18:3/16:0 18:3/16:318:3/16:0
PG MGDG DGDG(SL) MGDG DGDG

  

29

 

CHAPTER 2 GENETIC SCREEN AND PRELIMINARY
CHARACTERIZATION OF SUPPRESSOR OF VT E2 LOW
TEMPERATURE-INDUCED PHENOTYPES

ABSTRACT

Tocopherols (vitamin E) are lipid-soluble antioxidants that are synthesized only by
photosynthetic organisms (plants, algae and some cyanobacteria). Molecular dissection of
the tocopherol biosynthetic pathway in Arabidopsis and Synechocystis has allowed the
isolation of mutants containing different amounts and compositions of tocopherols and
made possible studies directed at elucidating their functions in photosynthetic organisms.
Studies with the tocopherol-deficient Arabidopsis thaliana vte2 mutant demonstrated an
important role for tocopherols in transfer cell wall development and achievement of full
photoassimilate export capacity during low temperature (LT) adaptation. In order to
understand the genetic basis of the vte2 phenotype and determine the missing links
between tocopherol deficiency and the vte2 LT-induced phenotypes, a genetic screen for
mutations that suppress the vte2 LT-induced phenotypes was undertaken. This chapter
describes the primary genetic screen, identification and genetic determination of the
number of suppressor loci. Seven independent sve (suppressor of ytaZ low temperature-
induced phenotype) lines were identified from approximately 12,000 EMS mutagenized
M2 vte2 plants screened under low temperature conditions. Due to long time period
required for backcrossing and evaluation of allelism tests (4 weeks at permissive
conditions and 4 weeks at LT to determine suppression), primary mutants representing
individual from different pools were subjected to physiological and biochemical analysis

to gain insight into their impact on the vte2 LT phenotype and to provide data for

30

selecting a limited number of lines for detailed study. Biochemical characterization of the
primary sve lines demonstrated they have differing impacts on sugar accumulation,
photoassimilate export capacity and vascular-specific callose deposition in the vte2
background at LT. Significantly, five of the primary sve lines altered fatty acid
composition before or in response to LT treatment reinforcing an important role for lipid
metabolism in the initiation and development of the vte2 LT phenotypes. Preliminary
characterization of the primary sve lines in this chapter allowed three lines (sverteZ,
sve2vte2 and sve7vte2) to be selected and prioritized for further backcrossing, detailed

analysis, and cloning described in Chapter 3.

31

INTRODUCTION

Tocopherols (vitamin E) are amphipathic molecules consisting of a chromanol ring
head group and a phytyl-derived side chain (Figure 1.1) (Schneider, 2005). The best
characterized function of tocopherols in animal systems is their role as lipid-soluble
antioxidants that prevent free radical damage of membrane polyunsaturated fatty acids
(PUFAs) (Burton and Ingold, 1981; Liebler and Burr, 1992; Bramley et al., 2000;
Schneider, 2005). Studies have also shown that tocopherols can affect membrane
properties due to their physical structure and interactions with specific lipid acyls (Kagan,
1989; Wang and Quinn, 2000; Atkinson et al., 2008). In addition, in animals specific
tocopherols also appear to have functions that are independent of their antioxidant
activities such as the stimulation of protein phosphatase A2 and thereby inhibition of
protein kinase C activities by a-tocopherol but not y-tocopherol (Boscoboinik et al.,
1991; Tasinato et al., 1995; Ricciarelli et al., 1998) and inhibition of cyclooxygenase-2
activity and hence prostaglandin E2 synthesis by y-tocopherol but not (it-tocopherol (Jiang
et al., 2000).

Tocopherols are essential dietary component for humans and animals and are
produced exclusively by photosynthetic organisms: higher plants, algae and some
photosynthetic bacteria (Grusak and DellaPenna, 1999). The molecular genetic dissection
of the complete tocopherol biosynthetic pathway in Arabidopsis thaliana and
Synechocystis sp. PCC6803 in recent years (Shintani and DellaPenna, 1998; DellaPenna,
1999; Collakova and DellaPenna, 2001; Porfirova et al., 2002; Shintani et al., 2002;
Cheng et al., 2003; Sattler et al., 2003b; DellaPenna, 2005b, a; DellaPenna and Pogson,
2006) allowed manipulation of tocopherol content and composition in plant tissues and

32

also enabled studies of the biological functions of tocopherols in photosynthetic
organisms (Maeda and DellaPenna, 2007).

Tocopherols are localized in plastids where their primary function has long been
assumed to be protecting polyunsaturated fatty acids (PUFAs) from oxidative damage
(Knox and Dodge, 1985; Robinson, 1988; Fryer, 1992; Munne-Bosch and Alegre, 2002),
an assumption based largely on correlations of increased tocopherol levels in response to
various abiotic stresses (DeLong and Steffen, 1997; Munne-Bosch and Alegre, 2000;
Collakova and DellaPenna, 2003b; Abbasi et al., 2007; Collin et al., 2008). vitamin E
deficient (vte) mutants have been pivotal in defining tocopherol functions in plants. The
Arabidopsis vte2 mutant, which is deficient in homogentisate phytyl transferase and lacks
all forms of tocopherols and pathway intermediates, had significantly reduced seed
longevity and produced excessive amounts of lipid peroxidation byproducts during
germination (Sattler et al., 2004; Sattler et al., 2006), consistent with tocopherols playing
an important role in protecting PUFAs in seed from oxidation. The vte1 mutant, which is
deficient in tocopherol cyclase activity, also lacks all tocopherols but unlike vte2
accumulates the redox active pathway intermediate dimethylphytylbenzoquinone
(DMPBQ). vte1 has only modestly reduced seed viability and is otherwise
indistinguishable from wild type (WT) (Sattler et al., 2004), suggesting the DMPBQ
accumulated by vte1 suppresses these tocopherol-deficient seed phenotypes.

In contrast to the clear situation in seeds of tocopherol deficient plants, studies of
their mature photosynthetic tissues have been equivocal. For example, when placed under
severe photooxidative stress conditions, the extent of bleaching and photoinhibition in

vte2 and vte1 was similar to WT (Porfirova et al., 2002; Havaux et al., 2005; Kanwischer

33

et al., 2005; Maeda et al., 2006). Significant differences were only observed when high
intensity light was combined with low temperature, indicating a more limited role for
tocopherols in photoprotection than had been assumed. Similarly, vte2 and WT were
indistinguishable under drought and salt stress conditions (Maeda et al., 2006). In
tobacco, VTE2-RNAi lines containing <5% of WT tocopherol levels exhibited increased
lipid peroxidation, enhanced chlorosis and reduced fresh weight under salt or sorbitol
stress while those with >5% WT tocopherol levels were indistinguishable from WT
(Abbasi et al., 2007). These combined results suggest that the tocopherol threshold at
which oxidative damage is observed is quite low and that the protective roles of
tocopherols in various oxidative stresses may be both stress- and species-specific.

In contrast to the variable results observed for stress responses of tocopherol
deficient plant species, a phenotype observed in mature plants of most tocopherol
deficient species is a disturbance of carbohydrate metabolism in source leaves. The maize
sde (sucrose export defectivel) mutant was initially identified due to a constitutive
defect in photoassimilate translocation and carbohydrate and anthocyanins accumulation
in mature leaves under normal growth conditions (Russin et al., 1996; Provencher et al.,
2001). The SH)! gene was subsequently shown to encode tocopherol cyclase (VTE 1;
(Sattler et al., 2003b). Similarly, VTElzRNAi lines of potato display constitutive
carbohydrate and anthocyanin accumulation in mature leaves, although the phenotype
was only present in lines with >99% reduction in total tocopherols (Hofius et al., 2004).
In both sde and the potato VTE1:RNAi lines, callose was constitutively deposited in the
vasculature of mature leaves (Botha et al., 2000; Hofius et al., 2004). In Arabidopsis,

tocopherol-deficient mutants are indistinguishable from WT at permissive temperatures

34

but develop defective photoassimilate export capacity, sugar accumulation and vascular
callose deposition upon exposure to non-freezing low temperature (LT) treatment (Maeda
et al., 2006). vte1 shows an intermediate phenotype between vte2 and wild type,
suggesting DMPBQ partially replaces the functions of tocopherols at LT conditions.
These results demonstrated that tocopherols are required for the development of full
photoassimilate transport capacity at LT in Arabidopsis. Importantly, the inducible nature
of these phenotypes in Arabidopsis provides a unique platform to study this aspect of
tocopherol function (Maeda et al., 2006).

It was initially hypothesized that LT-treated vte2 mutants might be under severe
photo-oxidative stress considering the lack of antioxidant protection in the absence of
tocopherols. However, lipid hydroperoxide contents measured using the FOX assay
(Jiang et al., 1992; Wolff, 1994) were below background levels in both vte2 and WT
during the long term LT treatment (Maeda et al., 2006). Zeaxanthin and antheraxanthin,
two xanthophyll cycle carotenoids known to be induced during photooxidative stress
(Demmig-Adams and Adams, 2002; Bouvier et al., 2005), were not detected in either WT
or vte2. Furthermore, under normal grth conditions and also during LT treatment,
maximum photosystem II (PSII) efficiency (Fv/Fm) in both WT and vte2 was maintained
at approximately 0.85, the typical Fv/Fm value for healthy Arabidopsis plants (Krause
and Weis, 1984). In addition, a recent mass spectrometry (MS)-based lipid profiling did
not detect any significant production of lipid peroxidation products (Maeda et al., 2008).
These results together indicate that vte2 plants are not experiencing severe oxidative

stress during low temperature treatment (Maeda et al., 2006).

35

The vte2 LT-induced phenotypes, especially the significant carbohydrate
accumulation in mature leaves, have raised many intriguing questions: How does the lack
of tocopherols cause carbohydrate accumulation under LT conditions? Do any other
changes occur prior to elevation of soluble sugar accumulation in vte2 after 60 hours of
low temperature treatment? What role, if not as an antioxidant, does tocopherols play in
plants experiencing low temperature stress conditions?

To search for the missing links between the lack of tocopherols in vte2 and altered
carbohydrate metabolism at LT and begin to understand the phenotypes of vte2, I have
undertaken a genetic screen for second site mutations that can suppress some or all of the
vte2 phenotypes. As a forward genetic tool, a suppression screen is essentially unbiased,
in contrast to reverse genetic studies, which may be based on incorrect or biased
assumptions and hypotheses. Suppressor screening has been suecessfirlly applied to many
systems and has given unexpected insights into complicated genetic pathways, regulatory
networks, and gene interactions (Li et al., 1999; Carpinelli et al., 2004; Eriksson et al.,
2004; Collins and Cohen, 2005). Although more time-consuming than reverse genetic
screens, this screen approach is especially suitable for this study because we lacked
sufficient knowledge to propose obvious targets for reverse genetic approaches. The
overall aim of the suppressor screen is to identify genes whose mutation allow restoration
of a wild type or near wild type phenotype in the presence of the vte2 mutation.
Characterization and identification of the suppressor mutants at different developmental
and treatment stages will help to dissect the complex process whereby tocopherols

regulate carbohydrate metabolism at low temperatures.

36

Ethyl methane sulfonate (EMS), an alkylating agent that primarily produces
GC->AT transitions, was chosen to induce mutations since it is able to generate a greater
diversity and density of mutations than insertional mutagenesis (Jander et al., 2002). As
EMS typically introduces hundreds of mutations in each plant line, it is possible to
identify mutations in any given gene by screening fewer plants than in insertional
mutagenesis (Feldman, 1994). Compared to activation tagging (Weigel et al., 2000),
which mainly produces gain-of-function mutations, EMS induces point mutations that
primarily results in a total or partial loss of function. It is therefore possible to dissect the
genetic pathway involved in vte2 phenotype by analyzing the non-redundant mutants that

disrupt such a pathway.

RESULTS

Suppressor screen identified 7 independent suppressor loci

Previous studies have shown that exposing 4-week—old vte2 plants grown at
permissive conditions to 7°C results in rapid induction of callose deposition in phloem
transfer cells, decreased photoassimilate export and increased levels of soluble sugars,
starch and anthocyanins in source tissues culminating in plants that are somewhat smaller
than wild type (Maeda et al., 2006). When 2-week-old plants grown at permissive
conditions are treated for 4 weeks at 7°C, these same biochemical changes occur but the
size difference between wild type and vte2 is greatly exaggerated (e.g. see Figure 2.5).
Therefore the primary mutagenesis screen was performed for second site mutations that
attenuate this severe reduction in plant size of 2-week-old vte2 after 4 weeks of LT

treatment. All subsequent biochemical characterization was performed using LT

37

treatment of 4-week-old plants grown at permissive conditions to obtain sufficient
experimental materials and allow direct comparison to previous results (Maeda et al.,
2006; Maeda et al., 2008). Suppressor screens were performed on M2 plants, in which
homozygous recessive mutations can be detected. Approximately 12,000 M2 plants from
10 separate M. pools were screened (the suppressor screening scheme is depicted in
Figure 2.1) and the M2 plants were allowed to grow for two weeks at permissive
conditions before being transferred to low temperature conditions. After four weeks of
low temperature treatment, when the visible phenotypes between vte2 and Col were
readily distinguishable, a total of 97 individuals that were larger in plant size and had less
purple color in leaves than vte2-1, were selected as putative suppressor lines (an example
screening flat is shown in Figure 2.2). Two sets of plants were screened and the putative
suppressors lines labeled as Ol-XXX from set 1 and 02-XXX from set 2. Because the
vte2-1 mutation is a G to A transition converting a Trp (TGG) to a premature stop codon
(TGA) and the primary target of EMS is guanosines, it is highly unlikely that the stop
codon would be converted to that encoding an amino acid by EMS mutagenesis. Hence,
all the suppressors should still be deficient in HPT activity and thus lack tocopherols.
When tocopherol analysis was performed on each putative suppressor by HPLC (Sattler
et al., 2003b), three lines were identified that had wild type levels of tocopherols and
lacked the vte2-l polymorphism. These three lines most likely represent wild type seed
contamination and were discarded.

To confirm that the suppression phenotype is heritable, 10-15 M3 progeny from the
putative 94 suppressor lines were screened a second time (Figure 2.1) under the same LT

treatment (an example of screening pets is shown in Figure 2.3). Ultimately, 29 lines

38

originating from seven M. pools were identified that clearly inherited the suppressor
phenotype and were carried on for further analysis. The 29 suppressor lines obtained
from different pools, their various nomenclatures and their general biochemical and
developmental traits are listed in Table 2.1. Pool 4 and 9 had the most suppressor isolates,
with 10 and 7 isolated, respectively.

To determine how many independent loci are represented by the 29 suppressor lines,
allelism tests were carried out by crossing representative lines from each M. pool, and in
selected cases between lines within an M. pool, and comparing the F. progeny with their
respective parental suppressor lines, and C01 and vte2. Evidence of allelism is provided
by the inability to complementation and the occurrence of suppression phenotypes similar
to both parents in the F. progeny. Efforts were made to perform the crosses that would
provide most information about allelism between suppressors from the different pools.
Suppressors from the same pool were invariably found to be allelic and likely descended
from the same M. plant while suppressor lines from separate pools were found to be non-
allelic. Therefore the representative members from each of the 7 pools were named “sve!
to sve 7”, respectively, for the “suppressor of ytaZ low temperature-induced phenotype 1—
2” (Table 2.1). It is important to stress that these are primary suppressors (denoted
“primary sve line”) and had not yet been backerossed. The remainder of this chapter
describes physiological and biochemical analysis with these primary sve lines that was
performed to gain information into their nature while in parallel the lines were being
backcrossed at least twice during a 9~12 month period to provide “clean” genetic lines
for detailed analysis in Chapter 3. Each representative primary sve line, vte2, vte1 and

Col had similar plant size and morphology before LT treatment (Figure 2.4). However,

39

after prolonged LT treatment, all primary sve lines exhibited enhanced growth and
reduced purple coloration of leaves relative to vte2, with the exception of sve6vte2, which

although was larger than Col, had leaf coloration similar to vte2 (Figure 2.5).

The primary sve loci show a range of suppression in sugar accumulation and

photoassimilate export capacity

Previously vte2, and to a lesser extent vte1, were found to accumulate sugars and
starch in mature leaves in response to LT treatment (Maeda et al., 2006). Since the
suppressors were selected primarily on the basis of plant size and lack of visible purple
coloration in the leaves, they may not necessarily suppress all other aspects of the vte2
LT phenotypes such as soluble sugar and starch accumulation. To test if the primary sve
loci affected the LT-inducible sugar accumulation phenotype of vte2, the leaf soluble
sugar content of each line was measured after two weeks of LT treatment (Figure 2.6).
The suppressors were found to have various degrees of suppression in soluble sugar
accumulation with the sverte2 and sve2vte2 primary suppressor lines completely
suppressing the vte2 sugar accumulation phenotype to the level of Col while the other
lines, with the exception of sve6vte2, showed at least a partial suppression. Primary
suppressor sve6vte2 contained sugar levels nearly equivalent to vte2 but had a plant size
similar to C01, suggesting carbohydrate accumulation and the vte2 grth phenotype can
be genetically uncoupled.

To investigate any impact of the primary sve lines on the impairment of

photoassimilate export in LT-treated vte2, photoassimilate export capacities were also
measured in all genotypes after 7 days of LT treatment. 14CO2 incorporation under LT
conditions was similar in all suppressor lines, vte2, vte1 and Col indicating that neither

40

the primary sve nor vte mutations significantly impacts the rate of carbon fixation in

leaves (Figure 2.7). Consistent with a prior report (Maeda et al., 2006), after 7 days of LT
treatment vte2 exhibited a dramatic reduction in l4C-labeled photoassimilate export to a

level 15% that of Col (Figure 2.8). This phenotype was partially suppressed in vte1 to a
level 35% that of C01. The seven primary sve lines exhibited varying degrees of
suppression of the vte2 LT-induced photoassimilate export phenotype (Figure 2.8).
sve1vte2 and sve2vte2 fully suppressed, sve6vte2 did not suppress, and suppression in
sve3vte2, sve4vte2, sve5vte2 and sve7vte2 was to a level similar or slightly higher than

that in vte1.

The primary sve lines display varying degrees of suppression in callose deposition

To investigate whether any of the primary sve lines have altered callose deposition
compared to vte2, two and four week-old sve plants were subjected to LT treatment and
aniline-blue positive fluorescence in leaf vascular tissues was examined and compared to
C01, vte2 and vte1. The most obvious differences in callose deposition were apparent
after 3 days of LT treatment (Figure 2.9). Primary sverteZ, sve2vt22 and sve5vte2 lines
completely suppressed callose deposition while primary sve3vte2, sve4vte2 and sve7vte2
lines showed partial suppression to levels less than or similar to vte1. Callose deposition
was not suppressed in sve6vte2 and was indistinguishable from or even stronger than
vte2. After 7 days of LT treatment, svelvte2 and sve2vte2 still showed almost complete
suppression of callose deposition while the other sve lines showed strong aniline-blue

fluorescence indistinguishable from vte2 (Figure 2.10).

41

Five of the primary sve lines show changes in fatty acid composition before or after LT

treatment

Recently it was shown (Maeda et al., 2008) that the levels of linolenic acid (18:3)
and linoleic acid (18:2) in vte2 leaves were significantly lower and higher than Col,
respectively, after two weeks of LT treatment. Furthermore, fad2 and fad6, the respective
ER- or chloroplast- localized w6-desaturase mutants (Miquel and Browse, 1992; Falcone
et al., 1994), were crossed to vte2, and the double mutants unexpectedly were found to
suppress both the morphological and biochemical phenotypes of the vte2 mutant at low
temperatures completely (fad2vte2) or partially (fad6vte2) (Maeda et al., 2008). Because
of these findings, we assessed the fatty acid (FA) composition of the primary sve lines.
To test whether any of the primary sve lines impacted fatty acid composition before LT
treatment, they and Col, vte2 and vte1 were grown at permissive conditions for four
weeks and their leaf fatty acid composition was assessed by analyzing the corresponding
fatty acid methyl esters using gas chromatograph (GC). Under permissive conditions
primary sve3vte2, sve4vte2, sve6vte2 and sve7vte2 lines had leaf fatty acid compositions
indistinguishable from Col, vte2 and vte1 (Table 2.2). Three other suppressors (primary
svelvteZ, sve2vte2, and sve5vte2 lines) had fatty acid compositions significantly different
from Col (or vte2) under permissive conditions. In comparison to both C01 and vte2, all
three lines had lower 18:3 and higher oleic acid (18:1) levels with the 18:1 level in
svelvteZ being particularly high and nearly 4-fold that in C01. The level of 18:2 was also
severely reduced in svelvte2 and moderately elevated in sve2vte2 and sve5vte2. Based on
the FA compositions in plants grown under permissive conditions, the suppressors were

categorized into two groups. Group I comprises the primary lines sverteZ, sve2vte2,

42

sve5vte2 from pools 1, 2 and 4, respectively, and all have altered FA compositions.
Group 11 includes suppressor lines from the rest of the four pools and they possess normal
leaf FA compositions compared with Col (and vte2) at permissive conditions (Table 2.1).

To further investigate if the LT-induced PUFA changes observed in vte2 were
affected in vte1 and the primary sve lines that showed fatty acid compositions similar to
C01 under permissive conditions (e.g. sve3vte2, sve4vte2, sve6vte2 and sve7vte2), four-
week-old plants were transferred from permissive to LT conditions for an additional two
weeks and fatty acid compositions analyzed by GC (Table 2.3). The LT-dependent PUFA
changes in these lines fell into two groups: primary suppressors sve4vte2, sve6vte2 and
vte1 had vte2-like PUFA changes and 18:3 and 18:2 levels not significantly different
from vte2, while the PUF A changes in primary lines sve3vte2 and sve 7vte2 were more
similar to C01 and had 18:3 and 18:2 levels significantly different from vte2 (Figure
2.11). These results suggest that the primary lines sve3vte2 and sve 7vte2 have suppressed

the LT-induced PUF A changes occurring in vte2.

DISCUSSION

LT treatment induced a series of responses in the tocopherol-deficient vte2 mutant,
and to a lesser extent the vte1 mutant, that were absent fiom wild type, including callose
deposition and altered transfer cell wall development in vascular parenchyma cells and
impairment in photoassimilate export followed by soluble sugar accumulation and growth
inhibition (Maeda et al., 2006). In this study, we employed a forward genetic approach to
identify a suite of suppressors of the LT-induced vte2 phenotypes in an attempt to
genetically define and dissect the pathway involved in the response of tocopherol-

deficient A rabidopsis mutants to LT treatment.

43

At least seven independent primary sve suppressor loci were identified based on
allelism tests and comparative biochemical characterization. This chapter reported
characterization of the primary suppressor lines. These data were used to select three sve
lines to target for future analysis and attempted cloning (Chapter 3). The growth
inhibition of LT-treated vte2 was at least partially suppressed in all the primary sve lines
(Figure 2.5) while other vte2-dependent responses were differentially impacted. The
primary sverteZ and sve2vte2 lines completely suppressed all LT-inducible vte2
phenotypes, including elevated sugar accumulation (Figure 2.6), reduced photoassimilate
export (Figure 2.8), and callose deposition (Figure 2.9). Primary sve3vte2, sve4vte2,
sve5vte2 and sve7vte2 lines showed a partial suppression of sugar accumulation and
photoassimilate export, similar to the levels observed in vte1. These four lines also had
partial or near total suppression of callose deposition after 3 days of LT treatment (Figure
2.9), but by 7 days of LT callose deposition in these lines was indistinguishable from vte2
(Figure 2.10). The primary sve6vte2 was unique in suppressing growth inhibition (Figure
2.5) but not impacting any other vte2-dependent responses (Figure 2.8, Figure 2.9)
indicating that grth inhibition can be genetically uncoupled from and is downstream of
the other LT-induced vte2 phenotypes.

Consistent with the observations of Maeda et al (2008), vte2 leaves show a fatty acid
composition identical to C01 under permissive conditions (Table 2.2) but have higher
18:2 and correspondingly lower 18:3 levels relative to C01 in response to LT treatment
(Table 2.3). Interestingly, three primary suppressor lines, svelvte2, sve2vte2 and sve5vte2
had fatty acid compositions that differed from Col or vte2 prior to LT treatment (Table

2.2). This result suggests that the constitutive alteration of fatty acids or membrane lipids

44

in these lines prevents the initiation and development of vte2 LT-induced phenotypes.
The fatty acid composition of the other four primary suppressor lines, sve3vte2, sve4vte2,
sve6vte2 and sve7vte2 were indistinguishable from Col under permissive conditions but
the responses of these suppressors to LT treatment fell in two distinct groups (Figure
2.12, Table 2.3). LT-treated sve3vte2 and sve7vte2 had fatty acid compositions most
similar to LT-treated Col, indicating these two lines suppress the vte2-dependent, LT-
induced PUFA changes, while LT-treated sve4vte2 and sve6vte2 showed PUFA
composition changes similar to LT-treated vte2 and thus genetically uncoupled vte2-
dependent, LT-induced PUFA changes from the other vte2 phenotypes. The fact that five
out of seven primary sve lines affect fatty acid composition either prior to (svelvteZ,
sve2vte2 and sve5vte2) or after LT treatment (sve3vte2 and sve7vte2) implies that LT-
induced PUFA changes is an important event involved in initiation and development of
LT phenotypes in vte2 and membrane lipid metabolism may play an early and central role
in the functions of tocopherols during LT adaptation.

Based on the physiological and biochemical analyses on the primary sve lines, three
lines (sverte2, sve2vte2 and sve7vte2), were selected for further detailed genetic,
physiological, biochemical analyses and molecular cloning (Chapter 3). The primary
lines of sve1vte2 and sve2vte2 almost completely suppressed all of the vte2 LT
phenotypes, including growth inhibition, sugar accumulation, photoassimilate export and
callose deposition (Figure 2.5, Figure 2.6, Figure 2.8 and Figure 2.9). In addition, both of
these lines have altered leaf fatty acid composition at permissive temperature (Table 2.1),
which may prove useful as a convenient selectable marker during genetic backcrossing

and mapping. More importantly, the biochemical processes which are disrupted in

45

svelvteZ and sve2vte2 may identify important links between tocopherol deficiency, the
abnormal fatty acid alteration and photoassimilate export defect in LT—treated vte2.
sve 7vte2, the third suppressor line, had a strong, though incomplete, suppression of all the
vte2 LT phenotypes (Figure 2.5, Figure 2.6, Figure 2.8 and Figure 2.9) and unlike
sverteZ and sve2vte2, did not display abnormal fatty acid composition at permissive
temperature (Table 2.1). sve7vte2 may have a distinctive suppression mechanism from
that of svelvteZ and sve2vte2 and thus it would be interesting to define the gene involved.
These three strongest suppressors lines selected for further studies should provide

valuable insights into tocopherol functions in plants.

46

MATERIALS AND METHODS

EMS mutagenesis

Mutagenesis was performed by exposing 52.4 mg of vte2-1 mutant seed to 0.3%
ethylmethane sulfonate (EMS) for 15 hours at room temperature. The seed were then
rinsed 10 times with distilled water over a 2h period, stratified for 5 days and separated
into 10 pools. Each M. pool was grown independently and the individual M2 seed pools

collected and used for suppressor screening.

Suppressor screening and low temperature treatment

For the first round suppressor screening, approximately 1200 seed from each of 10
M2 pool were surface sterilized, stratified for 5 days at 4°C, and then sown evenly on soil
along with a few seed of Columbia-0, vte1-1 and vte2-1 in each tray for comparison.
Plants were grown at permissive conditions (22°C day/ 18°C night) under 12h of 100
umol quanta m'zs'l illumination for two weeks. The temperature was then adjusted to 7°C
(: <3°C) and illumination lowered to 75 umol quanta m'zs'I for 4 weeks at which time
the visible phenotypes of vte2-I and Col-0 were readily distinguishable. 94 individual
seedlings that were judged larger than vte2-I and do not contain tocopherols were
selected as putative suppressors.

For the second round of suppressor screening, the putative suppressor lines were
allowed to self and grow to maturity at permissive conditions to produce M3 seed. About
15-20 progeny per line were re-screened as described above. 29 lines inheriting clearly

larger plant phenotypes at LT were selected and carried through additional analyses.

47

Allelism tests were carried out by performing multiple crosses between selected
suppressor lines from the same and different pools. 10-15 F. progeny plants were
subjected to low temperature treatment as described above and their phenotypes

compared with their parental lines, Col-0 and vte2-1.

Tocopherol analysis

To confirm the vte2 background in each putative suppressor, a leaf disk (~ 0.5cm in
diameter) was harvested from each plant, and lipids were extracted as described
(Collakova and DellaPenna, 2001). 0.01% (w/v) butylated hydroxytoluene (BHT) was
added for protecting lipids from oxidative damage and 0.5ug/ml tocol as an internal
standard. The lipid phase is dried and resuspended in the solvent (Methanol with 0.01%
w/v BHT) and used for reverse phase HPLC analysis to identify tocopherols as described

previously (Collakova and DellaPenna, 2001).

Analysis of soluble sugars

Quantification of leaf soluble sugar levels was based on the enzyme assay developed
by Jones et al (Jones et al., 1977) with minor modifications. A leaf disc (~ 0.50m in
diameter) from each plant was harvested at the end of the light cycle. The leaf punches
were quickly weighed and the soluble sugars were extracted twice in 80°C 80% ethanol
as previously described (Maeda et al., 2006). The extracts were combined, dried and
redissolved in 200 uL of distilled water. Dilutions were made on samples with higher
sugar contents and glucose, fructose and sucrose levels determined enzymatically as
previously described (Jones et al., 1977; Maeda et al., 2006). The total sugar content is

the sum of glucose, fructose and sucrose and expressed as % relative to vte2 level.

48

MC photoassimilate labeling

l4CO2 labeling of photoassimilate and measurement of phloem exudation were

performed as described (Maeda et al., 2006) with the exceptions that lOmM EDTA was
used for exudation buffer and 0.05 mCi of NaHMCOg. was used per labeling experiment.

Phloem exudates were collected after 5 hours of exudation.

Fluorescence microscopy

Leaves from each plant were harvested at the end of the light cycle and prepared for
aniline blue fluorescence microscopy as described (Maeda et al., 2006). At least 3 plants
for each genotype (n 2 3) were analyzed. The camera (SPOT FlexColor “C” mount
F X1520) settings for 3 (1 LT treated samples are: Exposure: 700 msec, Gain: 2, Gamma:
1.00 while those for 7 d LT treated samples are: Exposure: 900 msec, Gain: 4, Gamma:

1.40.

Analysis of fatty acid composition

Leaf punches from un-shaded mature leaf of each genotype were harvested and
lipids extracted as previously described (Dorrnann et al., 1995) and used to prepare fatty
acid methyl esters. The methyl esters were analyzed by gas-liquid chromatography using
myristic acid as an internal standard as previously described (James and Dooner, 1990;

Collakova and DellaPenna, 2001).

49

TABLES AND FIGURES

Table 2.1 A summary of the primary sve lines identified from suppressor screens.

Categorization of suppressors are based on fatty acid composition (Group), primary
suppressor locus name (Locus), the corresponding M. pool from which the suppressors
were identified (Pool), number of suppressor lines selected from each pool (Lines), the
names of the suppressor lines (suppressors), the leaf fatty acid composition under
permissive conditions (FA composition, 22°C) (refer to Table 2.2 for details), the type of
changes (“Col” or “vte2”) in linoleic acid (18:2) and linolenic acid (18:3) after 14 d of LT
treatment (18:2, 18:3 change, 7°C) (refer to Table 2.3 and Figure 2.11 for details), the
type of carbohydrate accumulation and photoassimilate export capacity (Sugars and
export) (“Col”, “vte2” or “intermediate”) in mature leaves after 14 d of LT treatment
(refer to Figure 2.6 and Figure 2.8 for details), the type of callose deposition (“Col”,
“vte2” or “intermediate”) after 3 d of LT treatment (refer to Figure 2.10 for details), and
the plant size after 28 d of LT treatment (Size). Representative suppressor lines at each

primary locus are marked with bold font

50

 

Table 2.1

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

FA 18:2,18:3
Locu Sugars and
Group Pool Lines: Suppressors composition change Callose Size
3 export
(22°C) (7°C)
01-034, 01-035,
Dramatically
01-036, 02-004,l
higher 18:1,
sve] 4 10 02-011, 02-017, NA C01 C01 C01
lower 18:2 and
02-018, 02-024,
18:3
1 02-025, 02-027
(altered Higher 18:1
01-016, 01-037,
FA) sve2 l 4 and 18:2, NA C01 C01 C01
02-007, 112-008
lower 18:3
Higher 18:1
111-050, 01-003,
sve5 10 4 and 18:2, NA Intermediate lntennediate Col
01-021, 02-001
lower 18:3
01-050, 01-003,
sve3 10 4 Normal Col lntennediate Intermediate Col
01-021, 02-001
01-047, 02-003,
11
01-048, 01-012,
(Normal sve7 9 7 Normal Col lntennediate lntennediate Col
01-013, 02-014.
FA)
02-015
sve4 3 2 01-032, 01-033 Normal vte2 lntennediate Intermediate Col
sve6 7 1 01-051 Normal vte2 vte2 vte2 Col

 

51

 

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53

{ ~ 12,000 M2 plants
(10 EMS-vte2 M1 pools)

l 22°C, 125uE

 

1*"t round screen < 2 week old plants

7.5°C, 75uE,
4 weeks

HPLC toc'
\ 94 putative suppressors

U

{ M3 progeny
22°C, 125m:

 

2 week old plants: 15-20 per line

7.5°c, 75uE,
2"cl round screen 4 weeks

 

29 confirmed suppressor lines

 

i 7 independent sve loci

Figure 2.1 Suppressor screening scheme.
The first round of suppressor screening identified 94 putative suppressor lines of which
29 were confirmed in a second screen. Allelism tests grouped these into seven

independent primary sve (suppressor 0fvte2 LT-induced phenotype) lines.

54

vte1

  

.g ..vun-m .

Figure 2.2 An example flat from the first roundof suppressor creening.

About 200 M2 plants from separate M. pools (in this image: pool 4) together with control
genotypes (Col, vte2 and vte1, marked with black circles) were planted in a flat and
allowed to grow for two weeks under permissive conditions and then transferred to low
temperature conditions. Pictures were taken after four weeks of low temperature
treatment. Individuals that were larger in plant size and had less purple color in leaves
than vte2 were selected as candidate suppressor lines (e.g. the four plants marked with red

circles). Size bar = 5 cm.

55

Col 01-013 01-016 vte2

 

Figure 2.3 An example image of the second round of suppressor screening.

The 94 putative suppressors were reassessed under LT conditions. About 15-20 M3 plants
from each putative suppressor line (eg. 01-013 and 01-16) were grown together with
control genotypes (Col, vte2) for two weeks under permissive conditions and then
transferred to LT conditions. Pictures are of plants after four weeks of low temperature
treatment. 29 lines whose N13 progeny inherited the suppressor phenotypes were

continued and used for further analysis. Size bar ; 2 cm.

56

sveZVtez

 

sve3wez sve 7Vtez
.3 \ b ’

Figure 2.4 Whole plant phenotype of primary lines of sverteZ-sve7vte2 compared

with Col, vte2 and vte1 before LT treatment.
Pictures were taken after growth under permissive growth conditions for two weeks. Size

bar = 2 cm.

57

 

Figure 2.5 Whole plant phenotype of the primary lines svelvte2—sve7vte2 compared

with Col, vte2 and vte1 after LT treatment.
All genotypes were grown for 2 weeks at permissive conditions and then transferred to

LT conditions. Pictures were taken after four weeks of LT treatment. Size bar = 2 cm.

58

160‘
140‘
120‘
100
80 ‘
60 i
40 i
20

 

l

 

 

Total sugar content (% of vteZ)

Col
vte2
vte1

sve1vte2
sve2vtez
sve3vteZ
sve4vte2
sve5vte2
sve6vte2
sve7vteZ

Figure 2.6 Leaf total soluble sugar content in primary sve1vte2-sve7vte2 lines
compared with Col, vte2 and vte1.

All the plants were grown for 2 weeks at permissive conditions and then transferred to
LT conditions. Sugar samples were taken from the 9-11'h leaves of plants at the end of the
light cycle after 2 weeks of LT treatment. Total sugar content is expressed relative to the
vte2 level. The dotted and dashed lines indicate the average sugar levels in vte2 and Col,
respectively. Data are means i SD (n = 5). Values significantly different from vte2

areindicated by * (P < 0.05) or ** (P < 0.01) (Student’s t-test).

59

140‘
120‘
100‘
80 ‘
60 '
4O ‘
20 ‘

1“C02 fixation rate (°/o of vte2)

 

3%E%%%%%‘3§‘3§
Usssssssss
3%‘33'33'6
>>>>>>>
WWWWMU’W

Figure 2.7 Total NC02 fixation after 7 d of LT treatment of primary sve1vte2-
sve7vte2 lines compared with Col, vte2 and vte1.

All lines were grown at permissive conditions for four weeks and transferred to low
temperature conditions. l4C02 labeling was done at the middle of the light cycle after 7 d
of LT treatment. Total fixed 14C02 is expressed as relative to vte2. Data are means i SD

(n 2 5).

60

 

 

Exudation (% total 14002 fixed)

Col
vte2
vte1

sve1vte2
sve2vte2
sve3vte2
sve4vteZ
sve5vteZ

sve6vte2
sve7vte2

Figure 2.8 Percent exudation of primary sve1vte2-sve7vte2 lines compared with Col,

vte2 and vte1.

All lines were grown at permissive conditions for 4 weeks and transferred to LT
conditions. MCOZ labeling of an individual leaf was done at the middle of the light cycle
following 7 d of LT treatment. Photoassimilate export capacity is expressed as the
percentage exudation (after 5 hours) of total fixed 14C02. The dotted and dashed lines
indicate the levels in vte2 and Col, respectively. Data are means i SD (n = 5). Values
significantly different from vteZ are indicated by * (P < 0.05) or ** (P < 0.01) (Student’s
t-test).

61

 

Figure 2.9 Callose deposition in primary sve1vte2-sve7vte2 lines compared with Col,
vte2 and vte1 after 3 d of LT treatment.

All lines were grown on ‘/2 MS plates for two weeks under permissive conditions and
whole seedlings were fixed at the middle of the light cycle after 3 additional days of LT
treatment. Aniline-blue positive fluorescence was examined in the lower portions of

leaves (11 23, size bar = 1 mm).

62

sve2vteZ

 

sve7vt92

 

sve3Vte2

Figure 2.10 Callose deposition in primary sve1vte2-sve 7vte2 lines compared with
Col, vte2, and vte1 after 7 d of LT treatment.

All lines were grown on soil under permissive conditions for four weeks and leaves were
fixed at the middle of the light cycle after an additional 7 days of low temperature
treatment. Aniline-blue positive fluorescence was examined in the lower portions of

leaves (n 2 3, size bar 2 1 mm).

I Col E] vte2 a vte1 m sve3vte2
40 - El sve4vte2 E sve6vte2 E3 sve7vte2

Molar ratio (%)

 

 

Figure 2.11 Molar ratios of linoleic acid (18:2) and linolenic acid (18:3) in primary
suppressor lines of sve3vte2, sve4vte2, sve6vte2, sve 7vte2 compared with Col, vte2 and
vte1 after 14 d of LT treatment.

Plants were grown under permissive conditions for 4 weeks and transferred to LT
conditions. Leaf samples were taken at the middle of the light cycle for fatty acid
analysis. sve3vte2 (vertical bars), sve4vte2 (dotted bars), sve6vte2 (horizontal bars),
sve7vte2 (hatched bars), Col (black), vte2 (white) and vte1 (gray). Data are means i SD

(n = 5).

64

CHAPTER 3 CLONING OF ARABIDOPSIS S VE
(S UPPRESSORS 0F VT E2 LOW T EMPERA T URE-INDUCED
PHENOTYPE) MUTAN TS HIGHLIGHT THE
INVOLVEMENT OF TOCOPHEROLS IN LIPID
METABOLISM

ABSTRACT

In the prior chapter, a genetic screen was carried out for sve mutations (suppressors
of the 2th low temperature-induced phenotype) and seven primary sve loci were
identified. In this chapter the three strongest sve loci (sveI, sve2 and sve7), which had
differing impacts on LT-induced sugar accumulation, photoassimilate export reduction
and vascular-specific callose deposition in vte2 were selected for further detailed analysis
and molecular cloning. sve] completely suppressed all vte2 LT phenotypes and was
found to be a novel allele of fad2, the endoplasmic reticulum-localized oleate desaturase.
sve2 showed partial suppression and was found to be a new allele of
trigalactosyldiacylglycerolI (tgdl), a component of the ER-to-plastid lipid ATP-binding
cassette (ABC) transporter. Introduction of tgd2, tgd3, and tgd4 mutations into the vte2
background similarly suppressed the vte2 LT phenotypes, indicating a key role for lipid
transport in the suppression of the phenotype. sve7 partially suppressed all vte2 LT
phenotypes without impacting fatty acid and lipid metabolism at permissive temperature.
Analyses of the acyl composition of ER- and plastid-derived lipids before and after LT
treatment demonstrated the elevation of 18:2 in phosphotidylcholine is an early and key
component in vte2 LT-induced responses as all suppressors attenuated this change and it
was the only lipid species significantly correlated with the suppressor photoassimilate

export capacity. Identification and characterization of sve loci highlights the involvement

65

of tocopherols in ER lipid metabolism in plants.

66

INTRODUCTION

Tocopherols are lipid-soluble antioxidants that are produced exclusively by
photosynthetic organisms (Grusak and DellaPenna, 1999). In plants, tocopherols are
synthesized in the plastid inner envelope (801] et al., 1980a; 8011 et al., 1980b) and the
majority of tocopherols are localized in plastids (Wise and Naylor, 1987; Havaux, 1998)
where their primary function has long been assumed to be protecting polyunsaturated
fatty acids (PUFAs) from oxidative damage (Knox and Dodge, 1985; Robinson, 1988;
Fryer, 1992; Munne-Bosch and Alegre, 2002). This was clearly shown to be a primary
role of tocopherols in dormant seed and germinating seedlings as in their absence the
seed PUFA pool suffers extreme non-enzymatic lipid peroxidation (Sattler et al., 2004;
Sattler et al., 2006). However in photosynthetic tissues, the role of tocopherols in
photoprotection was shown to be quite limited (Havaux et al., 2005; Maeda et al., 2006;
Abbasi et al., 2009) and studies in other plants are suggesting the roles of tocopherols in
various stresses may be both species and stress-specific (Maeda et al., 2006; Abbasi et al.,
2007; Maeda et al., 2008).

Similar to the maize sxd] and potato VTEI-RNAi lines (Russin et al., 1996;
Provencher et al., 2001; Hofius et al., 2004), the Arabidopsis tocopherol-deficient vte2
mutants develop vascular callose deposition, defective photoassimilate export capacity,
and sugar accumulation but only upon exposure to non-freezing low temperature (LT)
treatment (Maeda et al., 2006), suggesting that a role for tocopherols in development of
full photoassimilate transport capacity is evolutionarily conserved. A recent study
showed that LT-treated vte2 leaves have the levels of linolenic acid (18:3) and linoleic

acid (18:2) significantly lower and higher than wild type, respectively (Maeda et al.,

67

2008). However, exhaustive analysis by chemical and MS-based lipid profiling failed to
detect any significant differences in lipid peroxidation products in LT-treated vte2
relative to wild type. In addition, mutations in ER (13-6 fatty acid desaturasefadZ and to a
lesser degree plastidial 00-6 fatty acid desaturase fad6, suppressed the vte2 LT-induced
phenotypes, whereas mutations decreasing or eliminating 18:3, the fatty acid most
susceptible to oxidation (fad3, fad7fad8, fad3fad7fad8) did not (Maeda et al., 2008).
While these experiments have excluded a role for lipid oxidation in the vte2 LT
phenotype, the results have also clearly indicated the involvement of PUFA metabolism.
However, the cause and effect relationships between tocopherol deficiency, altered lipid
metabolism and the LT-induced vte2 phenotypes and the molecular mechanism
underlying this phenomenon remain unclear.

The previous chapter described a forward genetic approach to identify elements
linking tocopherol deficiency and carbohydrate metabolism in the LT-inducible
Arabidopsis vte2 system. Seven sve loci were identified but as it is impractical to move
forward and carry out map-based cloning with all the primary sve lines, the three
strongest lines (sve1vte2, sve2vte2 and sve7vte2) were prioritized based on their unique
features. The primary lines of sve1vte2 and sve2vte2 almost completely suppressed all of
the vte2 LT phenotypes (Figure 2.5, Figure 2.6, Figure 2.8 and Figure 2.9) and also have
altered leaf fatty acid composition at permissive temperature (Table 2.2). sve7vte2
showed a partial but strong suppression of all the vte2 LT phenotypes (Figure 2.5, Figure
2.6, Figure 2.8 and Figure 2.9) and has a normal fatty acid composition before LT
treatment (Table 2.2). In addition, following LT treatment, sve7vte2 showed changes in

fatty acids 18:2 and 18:3 similar to Co] (Table 2.3, Figure 2.11) suggesting that sve7vte2

68

may suppress the LT-induced vte2-specific PUFA changes. The biochemical processes
which are disrupted in sve1vte2, sve2vte2 and sve7vte2 are distinct and it is anticipated
that the detailed biochemical analysis and molecular identification of these suppressor
loci will reveal important links between tocopherol deficiency, fatty acid alterations and
photoassimilate export defect in LT-treated vte2 and thereby provide new insights into

tocopherol functions and their apparent involvement in lipid metabolism in plants.

RESULTS

sve], sve2 and sve7 are single recessive loci

Representative lines of the three strongest suppressors, sve1vte2, sve2vte2, and
sve7vte2 (Figure 2.5) were selected for fiirther genetic analysis and backcrossing to
remove other mutations in their backgrounds. F1 progeny of sve1vte2, sve2vte2 and
sve7vte2 backcrossed to vte2 showed LT phenotypes indistinguishable from LT-treated
vte2 (Figure 3.1), consistent with the three sve loci being recessive. This was confirmed
in F2 progeny of an sve1vte2 x vte2 cross which segregated 89:32 for vte2 and sve1vte2
LT plant phenotypes, respectively (Chi square test, p=0.67) (Figure 3.2). Similarly, F2
progeny of an sve 7vte2 x vte2 segregated 50:18 for vte2 and sve7vte2 LT phenotypes,
respectively (Chi square test, p=0.92) (data not shown). The two primary lines were
backcrossed at least twice and sve1vte2 and sve7vte2 were selected based on their visibly
distinguishable LT plant phenotype relative to vte2 and further confirmed by their
significantly lower sugar content (see later section). Segregation of sve2 in backcrossed
F2 population turned out to be more complex because of at least two other unlinked

mutations that resulted in yellowish leaf color and smaller plants and hence affected

69

sugar content and plant size making it difficult to select sve2 based on the LT sugar
phenotype (data not shown). Selection of sve2 was achieved using its other biochemical
phenotypes detailed in a later section. After segregating away the other mutations, unlike
the primary sve2 line which fully suppressed (Chapter 2 Figure 2.5, Figure 2.6, Figure 2.8
and Figure 2.9), the backcrossed sve2 now partially suppressed the vte2 LT phenotypes

(see later sections).

sve loci show different degrees of vte2 LT -induced phenotype suppression

When four-week-old sve1vte2 and sve7vte2 plants grown at permissive conditions
were subjected to an additional 4 weeks of LT treatment, they remained green and were
visibly similar to C01 while sve2vte2 plants showed a phenotype intermediate between
C01 and vte2 (Figure 3.3 A).

To assess whether the sve mutations affect the carbohydrate accumulation and export
phenotypes of LT-treated vte2, their soluble sugar contents and photoassimilate export
capacities were measured and compared with vte2, C01 and vte1. After two weeks of LT
treatment, the soluble sugar level in vte2 was nearly six times higher than C01 (232 and
40 umol/g FW, respectively). sve1vte2 had a sugar content identical to C01, while the
levels in sve2vte2 and sve7vte2 were intermediate and similar to vte1 (Figure 3.3 B).
Photoassimilate export capacity was measured aficr 7 d of LT treatment, when the
differences between vte2 and Col were sufficiently large (5-fold) for an intermediate
phenotype to be observed (Figure 3.3 C). The photoassimilate export capacity of sve1vte2
was indistinguishable from Co], while sve2vte2 and sve7vte2 were significantly different

from vte2 and Col but similar to vte1. These results indicate that sve], and to lesser extent

70

sve2 and sve 7, suppressed both the carbohydrate accumulation and export phenotypes of
LT-treated vte2.

To investigate whether sve lines altered the vasculature-specific callose deposition
phenotype of LT-treated vte2, aniline-blue positive fluorescence was examined in
response to LT treatment. Prior to LT treatment, all genotypes lacked callose deposition
in the vasculature (data not shown). Callose deposition remained absent in C01 at all LT
treatment time points (Figure 3.3 D, Figure 3.10). Afier 3 d of LT treatment, the
vasculature of vte2 and to a lesser extent vte1 contained significant levels of callose
(Figure 3.3 D), which intensified after 7 (1 LT treatment (Figure 3.10). sve1vte2 was
indistinguishable from Col after 3 (1 LT treatment (Figure 3.3 D) and had very few spots
of callose deposition after 7 d of LT treatment (Figure 3). After 3 d of LT treatment
callose deposition was delayed and decreased in sve2vte2 and sve7vte2 to a degree lesser
than vte1 (Figure 3.3 D) but by 7 d of LT treatment was similar to both vte2 and vte1

(Figure 3.10).

sve] and sve2 but not sve7 have constitutively altered fatty acid composition

A prior study showed that 14 d LT-treated vte2 had significantly higher and lower
linoleic acid (18:2) and linolenic acid (18:3), respectively, relative to C01 and esterified
primarily to phosphotidylcholine (PC) and phosphotidylethanolamine (PE) (Maeda et al.,
2008). To test the hypothesis that PUFA metabolism might be affected by sve mutations,
we assessed the leaf fatty acid composition of sve plants in comparison to C01, vte2, and
vte1 (Table 3.1). Under permissive conditions the fatty acid composition of Col, vte2 and
vte1 was nearly indistinguishable. sve7vte2 had slightly lower 18:2 and slightly higher

1623 but was otherwise similar to C01. In contrast, the fatty acid compositions of sve1vte2

71

and sve2vte2 were significantly different from Col. Both had elevated oleic acid (18:1)
levels with that of sve1vte2 nearly S-fold that of Col. 18:2 was severely reduced and 16:0
moderately reduced in sve1vte2 while both were moderately elevated in sve2vte2. The
level of 18:3 was also reduced significantly in sve2vte2. These results indicate that sve]
and sve2 but not sve7 significantly altered the fatty acid composition of vte2 at permissive

conditions.

sve] is a novel mutant allele of the ER localized oleic acid desaturase FAD2

sve] was the strongest suppressor isolated from the genetic screen (Figure 3.3) and
had significantly altered fatty acid composition (Table 3.1). The fatty acid composition of
F1 progeny from an sve1vte2 x vte2 cross at permissive temperature was identical to C01
and vte2 and photoassimilate export capacity after 7 (1 LT treatment was reduced to vte2
levels (Table 3.2). 32 F2 progeny of this cross that suppressed the LT—induced vte2
phenotype (Figure 3.2) also possessed a marked reduction in 18:2 and increase in 18:1
under permissive conditions, while 89 F2 progeny that did not suppress the vte2 LT
phenotypes (Figure 3.2) had fatty acid compositions identical to C01 and vte2 (Table 3.2).
These results confirmed that sve] is a single recessive locus and indicate its altered fatty
acid phenotype cosegregates with suppression of the vte2 LT phenotype.

The fatty acid composition in sve] suggested a defect in 18:1 to 18:2 desaturation
and was remarkably similar to the fatty acid profile of fatty acid desaturase 2 (fad2)
mutants (Lemieux et al., 1990; Miquel and Browse, 1992). To test if sve] carries a
mutation in FAD2, the coding and 1 kb upstream region of FAD2 in sve1vte2 was

sequenced and compared to C01. The FAD2 gene of sve1vte2 contains a single point

72

mutation (G to A) that converts A1a335 to Thr335, a highly conserved residue in FADZ
orthologs from evolutionary divergent photosynthetic organisms (Figure 3.11 A).

When sve1vte2 was crossed to an independently generated fad2-1vte2 double mutant
(Maeda et al., 2008) to test for genetic complementation, the leaf fatty acid composition
of F. progeny was nearly identical to both parents (Figure 3.4 A), indicating that sve]
carries a mutation at the same locus as fad2-1. Furthermore, the F . progeny of this cross
completely suppressed vte2 LT phenotypes including photoassimilate export deficiency
(Figure 3.4 B), callose deposition (Figure 3.4 C), and whole plant phenotype (Figure 3.4
D). These results indicate that the Ala335—>Thr335 mutation in FAD2 of sve1vte2 is

responsible for suppression of the vte2 LT phenotypes.

sve2 is a novel mutant allele of the ABC lipid transporter component T GD]

sve2vte2 partially suppressed all vte2 LT phenotypes to a degree similar to that of
vte1 (Figure 3.3) and also exhibited altered fatty acid composition under permissive
conditions (Table 3.1). The fatty acid composition of F1 progeny from a sve2vte2 x vte2
cross was identical to C01 and vte2 (Figure 3.5 A), consistent with the sve2 mutation
being recessive. The fatty acid profile of sve2vte2 did not resemble previously reported
fad mutants (Browse et al., 1993; Iba et al., 1993; Wallis and Browse, 2002). When the
polar lipid composition of sve2vte2, C01 and vte2 was analyzed by thin layer
chromatography, an additional spot was identified in sve2vte2 that is characteristic of
trigalactosyldiacylglycerol (TGDG) (Figure 3.5 B, (Xu et al., 2003b). Four
trigalactosyldiacylglycerol (tgd) loci, tgd] to tgd4, have been isolated in Arabidopsis and

found to accumulate TGDG due to defective lipid trafficking from the ER to plastid (Xu

73

et al., 2003b; Awai et al., 2006; Lu et al., 2007b; Xu et al., 2008). All tgd mutants exhibit
changes in fatty acid compositions similar to sve2vte2 (data not shown).

To test if sve2 is allelic to any tgd locus, sve2vte2 was crossed to each tgd mutant
and the resulting F. progeny analyzed for polar lipid composition. Only the progeny of
the sve2vte2 x tgd] cross showed a TGDG band (Figure 3.5 B and data of others not
shown), indicating the mutation giving rise to TGDG production in sve2 is allelic to tgd].
Sequencing of the T GD] coding region in sve2vte2 identified a single G to A mutation
converting Ala3l9 to Thr319, a conserved residue in TGDl orthologs from evolutionary
divergent photosynthetic organisms (Figure 3.11 B). A CAPS marker was developed for
the sve2 (tgd!) allele and used to genotype F2 progeny from an sve2vte2 x vte2 that were
then scored for sugar content in response to LT treatment. The sugar content of 14
sve2/sve2 progeny was significantly lower than that of vte2 whereas that of the 22
S VE2/sve2 and 10 S VEZ/S VE2 progeny were not (Figure 3.5 C), indicating that the tgd]
point mutation in sve2 is recessive and cosegregates with the sve2vte2 suppression
phenotype.

The previously characterized tgd] -1 mutation (Xu et al., 2003a) was introduced into
the vte2 background and crossed to sve2vte2 to test for complementation. The leaf sugar
content (Figure 3.6 A), export capacity (Figure 3.6 B), callose deposition in mature
leaves after 3 (1 LT treatment (Figure 3.6 C) and purple coloration in mature plants after
prolonged LT treatment (Figure 3.6 D) of the resultant F. progeny were similar to both
parents and significantly different from that of vte2, indicating that tgd] mutation in sve2

is responsible for the suppression of the vte2 LT phenotypes. Notably, the effect of

74

sve2vte2 on the above phenotypes is less dramatic than that of tgd] -1 vteZ indicating that

sve2 is a weaker allele of tgd] .

Three other tgd loci also suppress the vte2 LT - induced phenotypes

TGDl, 2 and 3 have been proposed to form an ABC lipid transporter complex that is
responsible for phosphatidic acid (PA) transfer through the plastid inner envelope
membrane (Xu et al., 2005; Awai et al., 2006; Lu et al., 2007a). TGD4 associates with
the ER membrane and is hypothesized to transfer lipids between the ER and outer plastid
envelop membranes (Xu et al., 2008). To test if the other tgd loci (tgd2, tgd3 and tgd4)
also suppress the vte2 LT phenotypes, we introduced tgdZ-I (Awai et al., 2006), tgd3-1
(Lu et al., 2007b) and tgd4-I (Xu et al., 2008) into the vte2 background and tested the
single and corresponding double homozygous mutants for their LT phenotypes. Afier
transfer of plants to LT conditions, all tgd single mutants were slightly paler than Col but
had soluble sugar content and photoassimilate export capacity nearly identical to C01
(Figure 3.7), indicating the tgd mutations alone did not impact LT adaptation. All tgd
vte2 double mutants were larger and had less purple coloration than vte2, suggesting
partial suppression of the vte2 LT phenotype (Figure 3.7 A), which was also apparent at
the biochemical level. The soluble sugar levels of LT-treated tgd] vte2, tgd2vte2, tgd3vte2
and tgd4vte2 were significantly lower than vte2, at 38%, 42%, 48% and 76% of the vte2
level, respectively (Figure 3.7 B) and their photoassimilate export capacity significantly
higher than vte2 at 64%, 66%, 56% and 44% of the Col level, respectively (Figure 3.7 C).
These results indicate that all four tgd loci partially suppress the vte2 LT phenotypes in
the order tgd1=tgd2 >tgd3 >tgd4.

Callose deposition in single and double tgd vte2 mutants was also assessed in

75

comparison to C01 and vte2 before and during LT treatment. Under permissive
conditions, no significant deposition of callose was observed in any genotype (data not
shown). After 3 d (Figure 3.8) and 7 (1 (Figure 3.12) of LT treatment the four tgd single
mutants and Col lacked vascular callose deposition while in vte2 vascular callose
deposition was present throughout the leaf at 3 (1 (Figure 3.8) and intensified by 7 (1
(Figure 3.12). After 3 (1 LT treatment, tgdlvte2 and tgd2vte2 showed sporadic callose
deposition in the primary veins of petioles and a significant reduction in deposition in the
leaf blade relative to vte2 (Figure 3.8). Callose deposition in tgd3vte2 and tgd4vte2 was
also reduced but to a lesser degree and some callose deposition was also observed in the
leaf blade (Figure 3.8). By 7 d LT treatment callose deposition in all the tgd vte2 double
mutants was indistinguishable from vte2 (Figure 3.12). These time-course observations
indicated that tgd] vte2, tgd2vte2 and to a lesser extent tgd3vte2 and tgd4vte2 delayed the

development of vte2 LT-induced callose deposition.

sve lines alleviate the vte2 LT -induced increase in phospholipid 18:2

Changes in the PUF A composition of ER-derived lipids have been shown to occur in
LT-treated vte2 leaves (Maeda et al., 2008). To investigate if the suite of sve lines
impacts these changes, we separated the major groups of ER- and plastid- derived lipids
[PC and PE, DGDG (digalactosyldiacylglycerol) and MGDG
(monogalactosyldiacylglycerol), respectively] by thin layer chromatography (TLC) and
assessed their acyl composition before and after 14 d of LT treatment. Under permissive
conditions, vte2, vte1, sve7vte2 and Col had identical fatty acid compositions in all lipid
classes analyzed (Table 3.3), confirming that vte2, vte1 and sve7vte2 do not impact fatty

acid content at permissive temperature. The four tgd mutants had higher 18:1 and lower

76

18:3 in both ER- and plastid- derived lipids, and fad2 had 6-8 fold higher 18:1 and lower
18:2 in PC and PE, consistent with prior reports (Miquel and Browse, 1992; Xu et al.,
2003a; Awai et al., 2006; Lu et al., 2007b; Xu et al., 2008). Introduction of vte2 into tgd
backgrounds had no impact on these profiles (Table 3.3). The fatty acid composition of
fad2vte2 was also similar to fad2-I, with minor exceptions (slightly higher PE-18:3 and
DGDG-18:1 and slightly lower DGDG-16:3). These results reinforce that tocopherol
deficiency per se has little or no impact on either ER- or plastidial fatty acid and lipid
metabolism under permissive conditions.

After 14 d of LT treatment, all genotypes showed elevated 16:3 and 18:3 levels
(Table 3.4), a phenomenon commonly observed in LT-treated plants (Williams et al.,
1988; Johnson and Williams, 1989; Uemura and Steponkus, 1997), indicating that vte2
did not affect this general LT-induced response of fatty acid desaturation. Col, vte2, vte1
and sve7vte2 showed identical changes in all fatty acids esterified to galactolipids. The
compositions of fatty acids esterified to galactolipids in the four tgd vte2 double mutants
and fad2vte2 were similar to the corresponding tgd and fad2 single mutants, respectively
(Table 3.4). These results indicate that tocopherol deficiency had little impact on plastid
lipid metabolism under LT conditions.

In contrast, significant fatty acid composition changes occurred in 18 carbon PUFAs
of ER-derived phospholipids under LT conditions (Table 3.4). Consistent with a prior
report (Maeda et al., 2008), vte2 showed significantly lower 18:3 and higher 18:2 levels
in PC and PE relative to C01. These changes were intermediate in vte1 indicating that the
DMPBQ accumulated in this. genotype partially suppressed these vte2-dependent LT

PUFA changes. sve7vte2 fully suppressed these changes and had PC and PE 18:2 and

77

18:3 levels indistinguishable from those of C01. The four tgd vte2 double mutants also
attenuated these vte2-dependent LT PUFA alterations and had PC and PE 18:2 and 18:3
levels similar to their corresponding single mutants. The extreme impact of fad2 on 18
carbon PUFA content makes data interpretation challenging, but PC and PE 18:2 levels
were identical in fad2 and fad2vteZ. Thus, a common theme that emerges from the suite
of suppressors is that all alleviate the LT-induced increase in membrane phospholipid
18:2 that would otherwise occur in LT-treated vte2. When the levels of unsaturated 18
carbon fatty acids esterified to PC, PE, MGDG and DGDG after 14 (1 LT treatment were
plotted against the LT photoassimilate export capacity of the suppressors (Figure 3.9 A),
PC-18:2 was the only membrane lipid species that showed significant correlation (Figure
3.9 B), suggesting that PC-18:2 plays a central role in the induction and development of

the vte2 LT phenotype.

DISCUSSION

Tocopherol synthesis has been conserved during plant evolution but the biochemical
and physiological functions of these compounds in plants remains the subject of debate
(Grasses et al., 2001; Munne-Bosch and Alegre, 2002; Hofius et al., 2004; Havaux et al.,
2005; Munne-Bosch, 2005a; Maeda et al., 2006; Abbasi et al., 2007; Dormann, 2007;
Maeda and DellaPenna, 2007). Tocopherol deficient lines in maize and potato exhibit
constitutive vascular callose deposition, carbohydrate accumulation in source leaves and
impairment in photoassimilate translocation to sink tissues (Russin et al., 1996; Botha et
al., 2000; Provencher et al., 2001). In tocopherol-deficient Arabidopsis mutants, such

phenotypes are conditionally induced by LT treatment and identification of sve

78

suppressors in this study allowed us to genetically dissect this complex pathway and
provide further insight into this novel aspect of tocopherol function.

All suppressor mutations, sve1(fad2), sve2(tgd1), sve7, tdg2, tgd3, tgd4 and vte1
fully or partially alleviated the growth inhibition of LT-treated vte2 to degrees that
correlated with their suppression of sugar accumulation, photoassimilate export
impairment and callose deposition (Figures 3.3, 3.7 and 3.8). Given that none of the
suppressors uncoupled growth inhibition from reduced photoassimilate export and
soluble sugar accumulation, these phenotypes are tightly linked genetically. The facts that
impaired photoassimilate export occurs immediately before soluble sugar accumulation
in LT-treated vte2 (Maeda et al., 2006) and that these two traits are negatively correlated
across all suppressors (R2=-0.86, Figure 3.13) strongly suggest that defective
photoassimilate export is the direct cause of elevated sugar accumulation. The vascular—
specific callose deposition and photoassimilate export block were almost completely
eliminated by the strongest suppressor (sve1vte2) (Figure 3.3), while weaker suppressors
partially attenuated callose deposition during the first 3 d but not 7 d of LT treatment
(Figures 3.3, 3.7 and 3.8) and partially restored photoassimilate export. Because transfer
cell wall proliferation is rapidly initiated and largely completed by 3 d of LT treatment
(Maeda et al., 2006), the partial suppression of callose deposition by weaker sve loci [i.e.
sve2 (tng), sve 7, tgd2-4] during this critical developmental time window is apparently
sufficient to allow an intermediate level of photoassimilate export.

Prior and current studies consistently showed that under LT conditions vte2 develops
higher 18:2 and lower 18:3 levels in ER-derived lipids (Maeda et al., 2008). The

identification of sve] and sve2 as novel alleles of fad2 (encoding ER oleate desaturase;

79

Figure 3.4, Table 3.2) and tgd] (a component of ABC transporter system responsible for
ER to plastid lipid trafficking; Figures 3.5 and 3.6), respectively, provides genetic
evidence for involvement of ER PUFA/lipid metabolism in the vte2 LT phenotype.
Furthermore, because mutations in the other transporter components (TGDZ, T GD3,
T GD4) also suppressed the vte2 LT phenotype (Figures 3.7 and 3.8), we can conclude
that it is the disruption of the ER-plastid lipid trafficking process, rather than a specific
transporter component, that is responsible for the suppression of vte2 LT phenotypes.

Previous radioactive tracer studies and lipidomic analyses demonstrated that the
observed PUFA changes in LT-treated vte2 were consistent with reduced conversion of
18:2 to 18:3 in the ER catalyzed by FAD3 (Maeda et al., 2008). However, because
multiple changes were observed in LT-treated vte2 relative to C01 (e.g. elevated PC-l8:2
and PE-l822, decreased PC-1823, PE-l8:3 and DGDG-18:3) it was unclear which lipid
species were critical for development of the vte2 LT phenotype. The vte2 suppressor lines
isolated or generated in this study provide important insight into this question. The
common biochemical phenotype shared by all vte2 suppressing lines (i.e. sve], sve2,
sve 7, tdg2, tgd3, tgd4 and vte1) was found to be attenuation in the elevation of 18:2 in PC
and PE that would otherwise occur in vte2 at LT (Table 3.4). When the levels of 18
carbon fatty acids in individual lipid classes were plotted against the LT photoassimilate
export capacity of various genotypes, only PC-l8:2 levels were found to be significantly
correlated (R2=0.73, Figure 3.9). These results indicate that the increase in 18:2 esterified
to PC plays a key role in the onset of the vte2 LT phenotype.

Why would PC-18:2 be highly correlated with suppression of the vte2 LT

phenotype? In plants, ER-produced PC is a key intermediate in lipid metabolism: ER-

80

localized desaturases predominantly act upon PC (Arondel et al., 1992; Okuley et al.,
1994); PC is the precursor to PA, diacylglycerol (DAG) and lyso-PC (Ohlrogge and
Browse, 1995; Mongrand et al., 2000; Andersson et al., 2004); PC-18:2 species are the
major donors of the DAG moiety for plastidic galactolipid synthesis (Slack et al., 1977;
Browse ct al., 1986; Somerville and Browse, 1991) and acyl chains of PC are in a
constant state of remodeling or acyl editing (Williams et al., 2000; Bates et al., 2007;
Bates et al., 2009). The rapid development of transfer cell walls in LT-treated
Arabidopsis (i.e. during 3 (1 LT treatment, (Maeda et al., 2008) requires a highly-
regulated supply of membrane lipids to the plasma membrane through ER-derived
vesicles (Offler et al., 2002; Maeda et al., 2006; Maeda et al., 2008). In the absence of
tocopherols, the fatty acid acyl composition of ER-derived lipids is suboptimal or
unbalanced as manifested by increased PC- and PE-l8:2 pools (Table 3.4), resulting in
deformed vesicle formation, function and transfer cell wall development (Maeda 2006,
2008). The sve] mutation decreases the synthesis of 18:2 in the ER and may restore the
unbalanced PUFA composition of ER-derived lipids used for transfer cell wall
development. Likewise, sve2 and all tgd mutations indirectly reduce ER 18:2 synthesis as
indicated by their increased levels of PC- and PE-l8:l (Table 3.3, (Xu et al., 2003b;
Awai et al., 2006; Lu et al., 2007b; Xu et al., 2008), leading to partial restoration of ER
PUF A imbalance and suppression of vte2 LT phenotypes.

It is difficult to envision how the absence of tocopherols influence ER lipid
metabolism when tocopherols are synthesized and presumably localized in plastids (8011
et al., 1980a; Lichtenthaler et al., 1981; S011, 1987; Vidi et al.,,2006). However, the

presence of high levels of tocopherols in ER-derived extraplastidic oil bodies (Y amauchi

81

and Matsushita, 1976; Fisk et al., 2006; White et al., 2006) is evidence that tocopherols
are not necessarily restricted to plastids. The recent demonstration of ER/plastid
membrane connection sites (p_lastid associated membranes or PLAMs) (Kjellberg et al.,
2000; Andersson et al., 2007) and of an ERzplastid lipid transport system (Hartel et al.,
2000; Benning, 2009) provide physical mechanisms whereby tocopherols could directly
access components of the ER membrane and/or lipid biosynthetic machinery. Once one
accepts that tocopherols have access to the ER compartment, several mechanisms for
tocopherols impacting ER lipid metabolism become feasible.

In vitro studies have shown that tocopherols can stabilize membranes (Stillwell et al.,
1996; Atkinson et al., 2008), possess a negative membrane curvature greater than
cholesterol (Chen et al., 1997; Bradford et al., 2003) and, because they dynamically
associate with PUFAs (Urano et al., 1988), may partition into more fluid membrane
domains sensitive to curvature stress (Brown and London, 1998; Simons and Toomre,
2000). These properties would likely impact processes such as vesicle function and
membrane fusion (Churchward et al., 2005), both of which are highly active in
Arabidopsis transfer cells at LT (Aubert et al., 1996; Maeda et al., 2008; McCurdy et al.,
2008). In animal systems tocopherol levels have also been shown to impact lipid and
fatty acid metabolism (Okayasu et al., 1977; Buttriss and Diplock, 1988; Mahoney and
Azzi, 1988; Bell et al., 2000) by directly affecting enzyme activities (Douglas et al.,
1986; Chandra etal., 2002) or indirectly by affecting the association of enzymes with the
membrane (Cachia et al., 1998; Brigelius-Flohe, 2009). Whether analogous biophysical
or biochemical membrane processes are similarly affected in tocopherol deficient mutants

in plants will require additional studies.

82

The sve lines characterized in this study provide genetic evidence for the sequence of
physiological and biochemical events occurring in tocopherol deficient plants under LT
and have defined elevated PC-l8:2 lipid species as critical components of the vte2 LT
phenotype. The identification of sve] and sve2 as mutations in genes encoding an ER
fatty acid desaturase (FAD2) and a component of ER-plastid lipid transporter (TGDl)
highlights an important role for tocopherols in regulating ER PUFA metabolism. Future
studies focusing on the possible transport of tocopherols from plastid to extraplastidic
membranes, role(s) of ER lipid PUFAs (especially PC-l822) on transfer cell wall
development and the mechanistic basis of tocopherol function in ER PUFA metabolism
will eventually lead to a better understanding of these unanticipated tocopherol fimctions

in plants.

83

MATERIALS AND METHODS

Plant growth and low temperature treatment

Seed were surface sterilized, stratified for 5 days at 4°C, and then planted in a
vermiculite and soil mixture fertilized with 1x Hoagland solution, and grown in a

chamber under permissive conditions (22°C day/18°C night) under 12h of 100 mmol
quanta mos] illumination for 4 weeks. For low temperature treatment, 4-weck-old plants

grown at permissive conditions were transferred at the beginning of the light cycle to low

temperature conditions (12h day/12h night under 7: <3°C with light intensity of 75 mmol

-2 -l . . . .
quanta m s ) for Indicated time periods.

14C photoassimilate labeling and analysis of sugars and callose

14CO2 labeling of photoassimilate and measurement of phloem exudation after 5h

were as described (Maeda et al., 2006) except that exudation buffer contained lOmM

EDTA and 0.05mCi of NaHl4CO3 was used per labeling experiment. Photoassimilate

. . . l4 .
export capac1ty is the percent exudation of the total fixed CO2 and 1S expressed as % of

Col. Leaf glucose, fructose and sucrose analyses were performed as previously described
(Maeda et al., 2006) and leaf total sugar content is the sum of glucose, fi'uctose and
sucrose contents and expressed as % of vte2. Callose staining and visualization were

performed as described (Maeda et al., 2006).

84

Lipid and fatty acid composition analysis

Lipids were extracted and polar lipids analyzed on activated ammonium sulfate-
impregnated silica gel TLC plates (Si250 with pre-adsorbent layer; Mallinckrodt Baker)
as previously described (Dormann et al., 1995; Xu et al., 2005). Galactolipids were
visualized with a-naphthol spray. For fatty acid composition analysis, total lipids or
individual lipids isolated from TLC plates were converted to fatty acid methyl esters and

analyzed by gas-liquid chromatography using myristic acid as internal standard (James

and Dooner, 1990; Collakova and DellaPenna, 2001).

Generation of double mutants

tgd] -1, tgd2-1, tgd3, and tgd4-1 mutants were kindly provided by Dr. Christoph
Benning and introduced into the vte2 background. F2 plants homozygous for vte2 and
one tgd locus were identified and confirmed using published TLC, HPLC and genotyping
methods (Collakova and DellaPenna, 2001; Xu et al., 2003b; Awai et al., 2006; Lu et al.,

2007)

Genotyping analysis of sve2

For genotyping of sve2, a CAPS marker was developed based on the presence of the
point mutation in T GD]. A 300 bp T GD] PCR fragment was amplified using primers 5’-
CTGTTGGTATGGCTTCAA-3’ and 5’-TTCAAAGAATCTCCAGCACCT-3’. The

mutant produces bands of 220bp and 80bp when digested with Taal.

Sequencing and sequence alignment analysis of FAD2 and T GD]

The genomic regions of FAD2 in sve1vte2 and fad2-l, and T 601 in sve2vteZ were
amplified in triplicate, pooled and used as sequencing templates. Mutations were

85

identified by comparison to the published FAD2 and T GD] sequences. FAD2 and TGDI
protein sequences for alignments were obtained from cDNA or deduced from genomic
sequence for: Arabidopsis thaliana (P46313, PF02405), Oryza sativa (TC267364,
NP_001053506), Picea abies (CAC18722), Physcomitrella patens (Contig11227,
XP_001763740), Ostrecoccus tauri (Otl7g02110), Galdieria sulphuraria
(contig_818_Oetl3_2005_g2.t1_Ga), Chlorella vulgaris (BAB78716), Chlamydomonas
reinhardtii (TC40690, XP_001696522), Prochlorococcus marinus str. MIT 9313
(N C_005071), Synechococcus sp. PCC7002 (AAB61352) Synechococcus elongatus PCC
6301 (YP_l71846), Populus trichocarpa (XP_002303003), Picea sitchensis
(ABK25388). Sequences were aligned using the CLUSTAL X (1.81) multiple sequence

alignment program.

Statistical analysis

For comparing three or more groups, one-way ANOVA as implemented in the
CoStat program (Cohort Software, 1990. Berkeley, CA, USA) was used with genotype as
the fixed effects. When significance was observed (p<0.05), multiple pairwise

comparisons were performed using the Tukey-Kramer test (Hsu, 1996), which allows for

unequal sample sizes. Significance of the coefficient of determination (R2) in Figure 7

was assessed using a t-test with (n-2) degrees of freedom (n=number of genotypes).

Accession Numbers

Sequences can be found at TAIR or GenBank: VTEI (At4g32770), VTE2
(At2g18950), FADZ (At3g12120), TGDI (Atlgl9800), TGDZ (At3g20320), TGD3

(Atl g65410), T GD4 (At3g06960). Information for Arabidopsis mutants is at TAIR. Seed

86

homozygous for vte2 rapidly lose viability (see (Sattler et al., 2004)) and thus are

maintained in the authors’ lab and available upon request.

ACKNOWLEDGMENTS

We thank Dr. Scott Sattler for providing EMS mutagenized vte2 seed, Dr. Christoph
Benning for providing tgd mutant seed and members of the DellaPenna lab for their

critical advice, discussions and manuscript review. This work was supported by NSF

grant MCB-023529 to D.D.P.

87

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Figure 3.1 Whole plant phenotypes of LT treated Col, vte2, sve1vte2, sve2vte2,
sve 7vte2, and the F1 progeny of sve1vte2, sve2vte2, sve 7vte2 x vte2.

All genotypes were grown under permissive conditions for 4 weeks and then transferred
to LT conditions. Representative plants of the indicated genotypes are shown after 14 d
(A) and 28 d (B. C) of LT treatment. Bar = 2 cm.

A. Col, vieZ. sve/142, sve/1W2 x vte2 Fl

B. C 0], vte2, .svc'2vte2, sve2vte2 x vte2 F 1

C. Col. vte2, sve7vte2, sve7vte2 x vteZ F I

Col vteZ primary sve1

Mature leaves
of randomly
selected
individuals
from F2
population of
sve1vte2 X vte2
cross

 

 

Figure 3.2 A demonstration of leaf phenotypes of F2 population from sve1vte2 x vte2
backcross.

All genotypes were grown under permissive conditions for 4 weeks and then transferred
to LT conditions for 28 d. 16 mature leaves randomly cut from 32 sve1 -1ike and 89 vte2-
like plants in the 121 F2 population, together with mature leaves of Col. vte2 and the

primary sve1 vte}, are displayed.

99

sve1vt92 svetheZ sve7vt92

e , a ,, a

 

Figure 3.3 LT phenotypes of sve1vte2, sve2vte2 and sve 7vte2 compared with Col, vte2
and vte1.

All genotypes were grown under permissive conditions for 4 weeks before transfer to LT.
A Representative whole plant phenotypes after 4 weeks of LT treatment. Bar = 20m.

B Total soluble sugar content of mature leaves after 2 weeks of LT treatment. Samples
were taken from the 9—11th leaves at the end of the light cycle. Data are means iSD
(n=5). Non-significant groups are indicated alphabetically (Tukey-Kramer test, P<0.05).
C Photoassimilate export capacity from mature leaves after 7 d of LT treatment.
Photoassimilate export capacity was assessed at the middle of the light cycle. Data are
means iSD (n=5). Non-significant groups are indicated alphabetically (Tukey-Kramer
test, P<0.05).

D Callose deposition after 3 d of LT treatment. Leaves were fixed at the middle of the
light cycle and aniline-blue positive fluorescence was examined in the lower portions of

leaves (n23, Bar = 1mm).

100

 

 

522326
(103 JO %)
o Auoedeo uodxa ateliwisseotoqd

120 a

 

m (zapi to °/°)1uatuoo iefins mo; 0

Figure 3.3 (cont’d)

101

0 Col I vte2 Isve1vte2 Ifad2vte2 sve1vte2 x fad2vte2 F1

mol (%)

 

16:0 16:1 16:2 16:3 18:0 18:1 18:2 18:3
Fattyacids

Figure 3.4 Complementation test of sve1vte2 and fad2-1vte2.

Plants were grown 4 weeks under permissive conditions before transfer to LT conditions
for the indicated period of time prior to analysis.

A Comparison of leaf fatty acid composition. Leaf samples were taken for fatty acid

analysis at the middle of the light cycle before LT treatment. Data are means iSD (n=5).
Significant differences relative to C01 are marked with * (p<0.05) or ** (p<0.01)

(Student’s t test).

B Comparison of photoassimilate export capacity. Photoassimilate export capacity was
assessed using mature leaves at the middle of the light cycle after 7 d at LT. Data are
means iSD (n=5). Significant differences relative to C01 are marked with * (p<0.05) or
** (p<0.01) (Student’s t test).

C Comparison of callose deposition. Leaves were fixed at the middle of the light cycle
after 3 d at LT and aniline-blue positive fluorescence was examined around the mid-vein
(n23, Bar = 1 mm).

D Representative whole plant phenotypes after 4 weeks of LT treatment. Bar = 2 cm.

102

 

 

 

 

:8 .o a...
5833 tonxo o.m__E_mmmo.ocm

 

Figure 3.4 (cont’d)

103

. ...... . 9
_.u_ NQENBE
x Nmtzgm

_.n_ NmENBS
x Not. . gm

madam.

magnum.

 

I

Figure 3.4 (cont’d)

104

>

50 E’Col 'vte2 'sve2vte2 I=sve2vt‘e2x vtez F1

mo|%

    

16:0 16:1 16:2 16:3 18:0 18:1 18:3
Fattyacids

Figure 3.5 sve2 carries a single recessive mutation in TGDI.

Plants were grown 4 weeks under permissive conditions before transfer to LT conditions
for the indicated period of time prior to analysis.

A Comparisons of leaf fatty acid composition. Leaf samples were taken from the
indicated genotypes at the middle of the light cycle before LT treatment. Data are means
iSD (n=5). Significant differences relative to C01 are marked with * (p<0.05) or **
(p<0.01) (Student’s t test).

B Comparison of leaf polar lipid composition. Thin-layer chromatography of polar lipids
showing the accumulation of TGDG in sve2vte2, tgd] and the F 1 progeny of an sve2vte2
x tgd] cross. Lipids were visualized by a-naphthol staining. DGDG:

J' I
U

‘ J Li“, 151, -... -2, TGDG: trigalactosyldiacylglycerol.
C Segregation of leaf total sugar content in F2 progeny of an sve2vte2 x vte2 cross.
Samples were taken from the 9-11th leaves at the end of the light cycle. Data for C01 and

vte2 are means iSD (n=5). Significant differences relative to vte2 are marked with *

(p<0.05) or ** (p<0.01) (Student’s t test).

105

*

0000 0
20 0
1186m2

* \00

C Got. .0 *0 E350 .503 .30...

TGDG

at

* Origin

'i

sve2vt92 segregation population

Figure 3.5 (cont’d)

106

4.;
ON
OD

na0m
o0000

Photoassimilate export capacity In
(% of Col)

   

Figure 3.6 Complementation test of sve2vte2 and tgdlvteZ.

Plants were grown 4 weeks under permissive conditions before transfer to LT conditions
for the indicated periods of time prior to analysis.

A Comparison of leaf total soluble sugar content after 2 weeks of LT treatment. Samples
were taken from the 9-1 1th leaves at the end of the light cycle. Data are means iSD
(n=5). Non-significant groups are indicated alphabetically (Tukey-Kramer test, P<0.05).
B Comparison of photoassimilate export capacity. Photoassimilate export capacity was
assessed using mature leaves at the middle of the light cycle after 7 d of LT treatment.
Data are means iSD (n=5).

C Comparison of callose deposition. Leaves were fixed at the middle of the light cycle
afier 3 d at LT conditions and aniline-blue positive fluorescence was examined around
the mid-vein of the leaves (n23, Bar = 1 mm).

D Representative whole plant phenotypes after 4 weeks of LT treatment. Bar = 2 cm.

107

.0
E was 2%.
x NmENQG

.... was :3.
x NoENmSm

3
we: $9

main...

9

Not—No:

E

V

 

.

 

D

.
0

Figure 3.6. (cont’d)

108

tgd1vte2 tgd2vte2 tgd3vte2. tgd4 vteZ

 

Figure 3.7 tgd2, tgd3 and tgd4 also suppress the vte2 LT phenotypes.

Plants were grown 4 weeks under permissive conditions before transfer to LT conditions
for the indicated periods of time prior to analysis.

A Representative whole plant phenotypes after 4 weeks of LT treatment. Bar = 2 cm.

B Total soluble sugar content of mature leaves after 2 weeks of LT treatment. Samples
were taken from the 9-1 1th leaves at the end of the light cycle. Data are means iSD
(n=5). Non-significant groups are indicated alphabetically (Tukey-Kramer test, P<0.05).
C Photoassimilate export capacity. Photoassimilate export capacity was assessed using
mature leaves at the middle of the light cycle afier 7 d of LT treatment. Data are means
iSD (n=5). Non-significant groups are indicated alphabetically (Tukey-Kramer test,

P<0.05).

109

 
 

 

 

o no .0 _o .0 _ — do 4 11 ._l 10‘ a
1 1
fiat. .0 «.0 E250 .503. .22. 000 .o 95.

3333 toaxo o.a__E_mmmo.o..m

110

Figure 3.7 (cont’d)

 
 
 

 

tgm tgds
tgd1vte2 tgd2vte2 tgd3vt92

Figure 3.8 tgd mutations impact differently on vte2 LT-induced callose deposition.

tg d4 vteZ

   

All genotypes were grown at permissive conditions for 4 weeks and transferred to LT
conditions for 3 additional days. Leaves were fixed at the middle of the light cycle.
Aniline-blue positive fluorescence was examined in the lower portions of leaves (n23,

Bar = 1 mm).

111

>

 

’5‘ 45 1
% 32:0.73
5
4o -
'3
3
v- 35 -
_ 30 ..
2
2
a“. 25 i l l l l l

 

 

O 20 40 60 80 100 120

PC-18

Photoassimilate export capacity (% of Col)

Figure 3.9 Correlation of individual lipid species with photoassimilate export
capacity.

Data include sve 7vte2, tgd1vte2, tgd2vte2, tgd3vte2, tgd4vteZ, Col, vte2 and We].

A A scattered plot of PC-18:2 levels (average of n=3 or 4) after 14 (1 LT treatment and
photoassimilate export capacity (% of Col, average of n=5).

B Coefficient of determination (R2) of unsaturated 18C fatty acid levels (A 18:1, .182,
'18z3) esterified to PC, PE, MGDG and DGDG (average of n=3 or 4) after 14 (1 LT
treatment and the photoassimilate export capacity (% of Col, average of n=5). R2 is based

on linear regression. Significant R2 is marked with * (P<0.05).

112

 

0.81 I a ‘ 18:1
'18:2
0-6‘ ‘18:3
‘3: 0.44
I
0.2- A ‘
0 a 9 I #

 

Figure 3.9 (cont’d)

PC PE MGDG DGDG
Lipid species

113

sve 7 vteZ

 

Figure 3.10 Callose deposition in leaf vascular tissues of the indicated genotypes
after 7 d of LT treatment.

All genotypes were grown under permissive conditions for 28 d before being transferred
to LT conditions. Leaves were fixed at the middle of the light cycle after 7 d of LT
conditions and aniline-blue positive fluorescence was examined at the bottom part of the

leaves (n23, Bar=lmm). Representative images are shown.

114

  

A to T A to T
i i
Arabidopsis 100 IWVIAHECG iAIKP 333
Oryza 30 Amp 320
Picea 165 LFVLGHDCG I L EATEAVKP 395
Physcomitrella 94 SEATEAIKP 332
Ostreococcus 147 WVEAHECG - ‘Q ‘ATDALKE 397
Galdieria 55 IFVLGHDCGH L EAL-EALEP 293

Chlorella 95 IWVIAHECGH ..OEATEALKP 335

  

Synechococcus 82 LFVVGHDCGH L DATEAIKP 310

 
 
  

sve2 tgd1-1
A to T S to Y
i i

Arabidopsis 307 GAKGVGESTTSAVVMSL IFIAD LS 335
Oryza 301 GAKGVGESTTSAVVVSLVaIFVAD ‘LS 329
Populus 218 GAKGVGESTTSAWISLVgIFMADP‘LS 246
Picea 418 GAKGVGESTTSAVVISL IFIAD LS 446
Chlamydomonas 202 GAKGVGESTTSAWISL IFIAD LS 230
Physcomitrella 219 IFIAD LS 247

Synechococcus 221'“- - LS 249

Figure 3.11 Multiple sequence alignment of relevant regions of FAD2 (A) and TGDI
(B) from representative photosynthetic organisms.

Black boxes highlight identical, gray boxes highlight conserved amino acids. Black
arrows indicate amino acid substitutions converting A104 to T104 in fad2-1 and A335 to

T335 in sve1 (A); A319 to T319 in sve2 and S323 to Y323 in tng-I (B), respectively.

115

tgd1 tgdz tgd3 “
vtez tgd1 vteZ tgd2vte2 tgd3 vteZ tgd4 vte2

   

Figure 3.12 Callose deposition in the indicated genotypes after 7 d LT treatment.

All genotypes were grown under permissive conditions for 28 d before being transferred
to LT conditions. Leaves were fixed at the middle of the light cycle after 7 d of LT
conditions and aniline-blue positive fluorescence was examined at the bottom part of the

leaves (n23, Bar=1mm). Representative images are shown.

116

 

 

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2 no

0

£10m R2=0.86

E

0

3. 80*

8 L11

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0.

5

0 40'

5 L

.E 20.

U)

U)

(B

.9 o . . . , . .
g o 20 40 so so 100 120
0.

Total sugar content (% of vte2)

Figure 3.13 Scattered plot of photoassimilate export capacity versus leaf total sugar
content.

Data include vte1, sve1vt62, sve2vte2, sve 7 vteZ, tgd] vte2, tgd2vt82, tgd3vte2, tgd4vte2
(gray), Col (white) and vte2 (black). Photoassimilate export capacity and leaf total sugar
content are expressed relative to the 7 (1 LT treated C01 and 14 d LT treated vte2,

respectively. Unidirectional error bars (iSD, n = 4~5) are shown for clarity. Coefficient

. . 2 . . . .
of determmation (R ) 18 derived from linear regressron.

117

CHAPTER 4 MICROARRAY ANALYSIS OF THE LOW
TEMPERATURE RESPONSE OF THE ARABIDOPSIS VT E2
MUTANT

ABSTRACT

The contrasting phenotype displayed in germinating seed of vte2 relative to wild type
clearly demonstrated a primary role of tocopherols in limiting the damage of seed PUFA
pools from nonenzymatic lipid oxidation. Transcript profiling studies further confirmed
the importance of nonenzymatic lipid oxidation in triggering the oxidative and defense
responses in germinating seed of vte2. While mature plants of vte2 are biochemically and
physiologically indistinguishable from wild type at optimum growth conditions, they
develop a series of biochemical and physiological phenotypes, detailed in Chapter 2 and
3, which eventually led to growth inhibition. The identification and characterization of
suppressors of the vte2 LT-induced phenotypes (sve loci) in Chapter 2 and 3 have
indicated ER lipid metabolism is an early, upstream event and highlights an important
role for tocopherols and lipid metabolism in the initiation and development of the vte2
LT-induced phenotypes. To further understand the response of vte2 to LT at the
transcriptional level, a global transcript profiling study comparing vte2 and wild type
plants under different time period (0h, 48h, 120h) of LT treatment was carried out.
Tocopherol deficiency has no impact on global gene expression at permissive conditions
but affected a small number (77) of genes after 48h of LT treatment. Comparisons with
gene profiles of plants subjected to various stress conditions revealed that some degree of
oxidative stress response was likely occurring in 48h-LT-treated vte2. Consistent with its
biochemical and physiological phenotypes, LT-treated vte2 also showed enhancement in

expression of genes involved in cell wall modification and repression of genes related

118

with solute transport. Statistical analyses also highlighted several other potentially

important target genes for future studies.

119

INTRODUCITON

The Arabidopsis vte2 mutant, which is deficient in homogentisate phytyl transferase
(HPT) and lacks all forms of tocopherols and pathway intermediates, has significantly
reduced seed longevity compared with the wild type (Sattler et al., 2004). During
germination, vteZ seedlings exhibited severe seedling growth defects and contained
excessive amounts of the major classes of free radical-mediated lipid peroxidation
products: hydroxy fatty acids, malondialdehyde (MDA) and phytoprostanes (Sattler et al.,
2006). Sattler et al (2006) performed global transcript profiling on 3—d-old wild type, vte1
and vte2 seedlings and defined 160 genes induced by at least 2.5 fold in 3-d—old vte2
seedlings relative to wild type. Overall this gene list overlapped substantially with gene
expression profiles of oxidative and biotic stress (Sattler et al., 2006). These results
suggested that the non-enzymatic lipid peroxidation occurring in vte2 seedlings
contributed to triggering the transcriptional response of seedlings to pathogens.

Surprisingly, once past the germination and early seedling development stage, vte2
mutant plants are biochemically and physiologically indistinguishable from wild type at
optimum growth conditions (Maeda et al., 2006), suggesting that tocopherols are
dispensable in mature plants. However, when the mutant plants were subjected to low
temperature treatment, they displayed series of biochemical and phenotypic phenotypes,
as described in detail in Appendix (Maeda et al., 2006). The identification and
characterization of suppressors of the vteZ LT-induced phenotypes (sve loci) in Chapter 2
and 3 have further suggested that alteration in ER lipid metabolism is an early event and
highlighted an important role for tocopherols and lipid metabolism in the initiation and

development of the vte2 LT-induced phenotypes. However, it is still poorly understood

120

why tocopherol deficiency would alter ER lipid metabolism at LT conditions and what
might be the molecular links between tocopherol deficiency, lipid metabolism and LT
adaptation.

In animals, multiple studies have investigated transcriptome level responses to
vitamin E in different organs using either experimental animals fed with tocopherol -
sufficient and -deficient diets (Barella et al., 2004; Rota et al., 2004; Rota et al., 2005;
Nell et al., 2007; Oommen et al., 2007) or a-tocopherol transfer protein knock out mice
(aTTP(-/-)), which have extremely low cellular tocopherol contents (Gohil et al., 2003;
Vasu et al., 2007; Vasu et al., 2009). While a few studies showed that vitamin E
deficiency elicits pronounced changes in the expression of known antioxidant—responsive
genes, such as catalase, superoxide dismutase and glutathione S-transferase (Gohil et al.,
2003; Jervis and Robaire, 2004; Hyland et al., 2006), other studies failed to detect
significant changes in the expression of these classical antioxidant genes and instead
showed alteration of expression in other distinctive groups of genes, such as those
involved in testosterol synthesis and cancer development in testes (Rota et al., 2004),
hormone metabolism and apoptosis in the hippocampus (Rota et al., 2005), lipid
metabolism, inflammation and immune system in the heart (Vasu et al., 2007), transport
processes, synaptic vesicular trafficking in particular, in livers (Nell et al., 2007),
cytoskeleton modulation in lungs (Oommen et al., 2007) and muscle contractility and
protein degradation pathways in muscles (Vasu et al., 2009). The impact of dietary or
mutation induced a-tocopherol deficiency may explain the observed whole animal
dysfunctions in muscle, neural and reproductive organs that seem to be independent of

tocopherol’s antioxidant fimctions.

121

In plants, no global gene expression profile for the effect of tocopherols in
photosynthetic tissues has hitherto been undertaken. Comparing wild type and vte2 plants
at permissive and low temperatures should provide an ideal system to investigate the
transcriptome level responses of plants to tocopherol deficiency in plants. DNA
microarray analysis with Affymetrix’s ATHl GeneChip® technology was used to
investigate the effects of tocopherol deficiency on global gene expression at both
permissive and LT conditions. Because our goal was to identify early changes in gene
expression that might be directly related to absence of tocopherols at LT, these time
points were selected for the experimental design: 0, 48 and 120 hours of LT treatment.
Based on the previous time course physiological and biochemical analyses detailed in
Appendix (Maeda et al., 2006), at 0h (before LT treatment), vte2 and Col plants are
physiologically and biochemically indistinguishable. After 48h of LT treatment, vascular
callose deposition has been induced and photoassimilate export capacity in mature leaves
of vte2 is significantly lower than Col but soluble sugar accumulation between the two do
not differ. 120h of LT treatment represents a relatively late response of vteZ where
soluble sugars are significantly higher in the mutant and callose deposition is extensive
and wide spread. Afler 48 h of LT treatment, it should be possible to identify early
responses to tocopherol deficiency that are distinct from those due to the elevated sugar
levels in vte2 after 120h of LT treatment. The zero (permissive) 0h time point is a critical
control to compare gene expression differences between vte2 and C01 in the absence of
LT treatment. The experiment comprised 18 chips and is a factorial design, with three

independent biological replicates for Co] and vte2 at each of the three time points.

122

RESULTS

Data evaluation and preprocessing

Preprocessing, including data evaluation, cleaning and the analytical methods that
are applied to obtain reliable estimates of the relative abundance of each gene in a
sample, is an essential requirement before any downstream statistical analysis (Russell et
al., 2009). The quality of RNA used for GeneChip hybridization can be crucial in
determining the quality of the obtained data. Because perfect match (PM) probe
intensities should be systematically elevated at the 3' end of a probe set when compared
to the 5' end when RNA degradation is sufficiently advance, the 3’/5’ ratios of expression
have been used as a global indicator of RNA degradation (Bolstad et al., 2005). When the
3'/5' expression ratios of PM probes were used to evaluate the quality of the 18 arrays
(Figure 4.1), all the arrays showed similar slopes, indicating the RNA used for the
experiments is of good and comparable quality.

The 3’: 5’ ratios of the control genes on the chips were also assessed as another
quality check. The 3’: 5’ values of GAPDH and B-aetin (Figure 4.2) of all the chips were
within the recommended ranges (Affymetrix’s recommendation is ~1 for GAPDH and <3
for B-actin). In addition, the percentage of present genes on all the chips were similar,
ranging from 49.6 ~ 61.7% (Figure 4.2). The average background of most of the chips
ranged from 86.3 ~ 109.8, except for chips CO3, v01 and v02 (which represents Col-Oh-
replicate 3, vte2-0h—replicate l, and vte2—Oh-replicate 2, respectively) (Figure 4.2), which
therefore necessitated up-scaling as indicated by the positive scale factors in Figure 4.2.

Different arrays have different distributions and it is important to assess the effect of

background noise and to diagnose outliers. As seen in the boxplot (Figure 4.3) and the

123

 

histogram (Figure 4.4) of the chip data, most chips had a similar distribution range, with
the exception of chips v01 and v02 which were significantly lower than the others. The
low intensity of these chips is probably due to attenuated biochemical reactions during
labeling or a reduction in detection efficiency. Therefore the data needed to be
preprocessed with background correction and normalization to ensure chip-chip
comparisons.

The Robust Multichip Average (RMA) technique for background adjustment,
normalization and summarization functions (Irizarry et al., 2003b) as implemented in the
affy package was chosen for preprocessing the array data because it performs better than
the scaling method and is computationally fast (Russell et al., 2009). It consists of three
preprocessing steps: convolution and background correction, quantile normalization, and
a summarization based on a multi-array model fit robustly using the median polish
algorithm. Because of various problems associated with Mismatch (MM) probes, only
PM intensities are used in the RMA convolution method (Bolstad et al., 2003; Irizarry et
al., 2003a; Irizarry et al., 2003b). Quantile normalization imposes the same empirical
distribution of intensities to each array and summarization combines the multiple probe
intensities for each probe set to produce an expression value. The distributions of the chip
data afier these three steps were similar as shown in the boxplot (Figure 4.5) and
histogram (Figure 4.6), suggesting that expression values from different chips are now
directly comparable.

To further investigate the relationship of the chips, a cluster dendrogram was
generated (Figure 4.7). The chips grouped into three clusters, which are consistent with

the three time points of low temperature treatment, with the chips for 48h and 120h LT-

124

treated samples being more closely related than to the Oh data, suggesting that the effect
of LT treatment was greater than the differences between the genotypes. The three
replicates of genotype in each treatment generally clustered together, suggesting the good

biological repetition of the experiments.

T ocopherol deficiency had little impact on global gene expression at permissive

conditions

Previous biochemical and physiological analyses have shown that except for an
absence of tocopherols, vte2 mutant plants grown at permissive conditions contain
identical levels of chlorophylls, carotenoids, anthocyanins and lipid peroxides, have

normal photosynthesis as indicated by maximum photosynthetic efficiency (F v/F m) and

quantum yield of PSII ((1313311), and like wild type, contain identical soluble sugar and

starch and have no callose deposition in vasculature tissues (Maeda et al., 2006).
Recently it was further shown that under permissive conditions vte2 plants have fatty acid
compositions identical to wild type indicating tocopherol deficiency does not impact the
fatty acid and lipid metabolism prior to LT treatment (Maeda et al., 2008).

To investigate if there are any transcriptional changes in vte2 plants relative to C01
before LT treatment, limma analysis was performed to detect differently expressed genes
between vte2 and Col at the Oh time point. Only two genes [Nth (Atcg01080, NADH
dehydrogenase ND6) and Ath (AtchOISO, a subunit of ATPase complex CFO], both of
which are encoded by and located in chloroplast, showed significant differences (adjusted
P value = 0.05) between vte2 and Col before LT treatment. For comparison, a different
method was used for data analysis with GeneSpring sofiware. To reduce false positives,

the data of 6 chips at Oh time point were filtered using the detection call (Present/Absent)

125

generated by the Affymetrix microarray suite version 5 software (MASS). 13572 probe
sets called “Present” in either Col or vte2 were further filtered with “2 fold change”
between vte2 and C01 and 3 genes were identified. Afier t test, the same 2 genes as
detected by limma analysis (Nth and Ath) were identified to be significantly different
between the 2 genotypes (False discovery rate < 0.05). The near identical results obtained
from the two different analysis methods indicated that lack of tocopherols had little, if

any, impact on the global gene expression in vte2 at permissive conditions.

77 genes are significantly different in 48h-L T-treated vte2 and Col

At 48 hours of LT treatment, 77 probe sets were found to be significantly different
(adjusted p value<0.05) between vte2 and Col using the limma analysis. The genes
represented by the probe sets can be grouped into two categories: 49 genes that are
significantly induced (Table 4.1), and 28 genes that are significantly repressed in vte2
relative to C01 (Table 4.2). A gene tree was generated to further investigate the
expression patterns of the genes across different time points and based on this, the
significant genes in vte2 at 48h of LT treatment were further classified into 4 groups
(Figure 4.8). The majority of the genes (43 genes, group I) showed temporary and
moderate induction in C01 while their induction in vte2 was higher than Col at 48b of LT
treatment and generally remained higher than Col at 120h of LT treatment. Group IV
contains 16 genes that were temporarily repressed in 48h-LT-treated C01 and were
continually repressed in LT-treated vte2. Group II (11 genes) are repressed in LT-treated
Col but induced in LT-treated vte2 whereas group III (18 genes) generally had increased
expression in LT-treated C01 and decreased expression in LT-treated vte2. Because

groups 11 and 111 genes have opposite expression patterns in vte2 and Col, they may be

126

used as potential “marker genes” differentiating the transcriptional responses between
LT-treated vte2 and Col.
Significantly induced genes in 48h-LT-treated vte2 and possible roles of WRK Y

Among the list of 49 significantly upregulated genes in vte2 after 48h of LT
treatment, 6 are annotated as glycosyl transferases (At3g14750, At3g59100, At4g19460),
UDP-glucosyl transferase (At3g21780) or glucanases (Atlg26450, At1g65610) (Table
4.1). These genes are likely involved in different aspects of cell wall modification, which
is consistent with the major modifications of cell wall structure in phloem parenchyma
cells of LT-treated vte2. Notably, LT treatment induced significant, albeit low, expression
of two putative callose synthase genes, GSL4 (At3g14570) and GSLII (AGGS9IOO) in
vte2. These two members of the 12 GSL family in Arabidopsis may contribute to the
substantial callose deposition in transfer cells of vte2 and are investigated further in a
later section.

Some of the upregulated genes in LT-treated vte2 are involved in stress and
senescence responses, including a senescence-associated family protein (At4g23410) and
three glutathione S-transferases (GSTs), which have well-established roles in xenobiotic
and ROS detoxification (Foyer et al., 1997). Furthermore, several upregulated genes are
involved in various signaling pathways, including two auxin-responsive genes
(Atlg59500, At4g27260, both are auxin-responsive GH3 family proteins) and two genes
potentially involved in calcium signaling (At3g22910, a calcium-transporting ATPase,
and At1g68620, a putative calmodulin-related protein).

Notably, 5 transcription factors are among the induced genes in vteZ, including three

WRKY, one NAM and a zinc finger (C2H2 type) family protein (Table 4.1). WRKY

127

factors belong to the zinc-finger-type class of proteins and these plant-specific
transcription factors contain a DNA-binding region of approximately 60 amino acids in
length (the WRKY domain), which comprises the absolutely conserved WRKY motif
adjacent to a novel zinc-finger motif (Ulker and Somssich, 2004). The WKRY family has
74 members in Arabidopsis and the physiological fimctions of most are not known,
however, some have been implicated in plant stress responses (Eulgem et al., 2000),
especially biotic stress responses (Eulgem and Somssich, 2007). The three WRKY
factors that were induced in vte2 relative to C01 after 48h LT treatment are WRKY75
(At5gl3080), WRKY8 (At5g46350) and WRKY38 (At5g22570). Based on microarray
analysis, WRKY 75 has been shown to be involved in pathogen infection (Dong et al.,
2003) and senescence (Guo et al., 2004) and is strongly induced during phosphate
deprivation in Arabidopsis (Devaiah et al., 2007). WRKY38 was shown to be involved in
regulating cold and drought stress response in barley (Mare et al., 2004), is induced by
both pathogen infection and SA treatment and thought to function as a negative regulator
of plant basal defense in Arabidopsis (Dong et al., 2003; Kim et al., 2008). The function
of WRKY8 is currently unknown.

It is possible that some of the induced genes in 48h-LT-treated vte2 might be
regulated by WRKY factors. WRKY family members have been shown to bind to a DNA
sequence designated the W box, (C/T)TGAC(T/G) (Eulgem et al., 2000). When the
promoter regions of the 46 induced genes (excluding three WRKY from the total of 49
induced genes) in 48h-LT-treated vte2 were analyzed for this sequence motif, 22 were
found to have at least one hit for a W box element in their promoter regions (Table 4.3).

Notably, the promoters of all six cell wall-related genes have one or more W-box

128

sequences and could potentially be regulated by the WRKY transcription factors. Some
of the genes involved in stress (two GSTs) and auxin homeostasis (At1g59500) also have

the binding sites of W box.

Significantly repressed genes in 48h-L T -treated vte2

One of the significantly repressed gene in vte2 encodes VTE2/HPT (Table 4.2), the
basis of the mutant and key limiting enzyme activity for tocopherol biosynthesis
(Collakova and DellaPenna, 2003b). One of the downregulated genes of interest is MsrB5
(At4g04830), which encodes methionine sulfoxide reductase B5. It is known that a
variety of oxidants generated in metabolic stress conditions readily oxidize methionine
(Met) residues in proteins to Met sulfoxide (MetSO) (R or S configuration) (Brot and
Weissbach, 1991), resulting in modification of protein activity and conformation. MetSO
can be reduced back to Met by Met sulfoxide reductase (MSR, EC 1.8.4.6). MSR activity
is present in most organisms and has been included in the minimal set of proteins
sufficient for cell life (Moskovitz et al., 1995; Mushegian and Koonin, 1996). MsrA
enzymes utilize thioredoxin to reduce the S stereoisomer of MetSO back to Met
(Moskovitz et al., 1995) while MsrB can catalyze the reduction of the R isomer of MetSO
to Met (Lowther et al., 2002; Weissbach et al., 2002). Interestingly, plants have the
largest number of Msr isoforms, with five MsrA (Sadanandom et al., 2000) and nine
MsrB genes (Dos Santos et al., 2005) identified in Arabidopsis. Certain Msr genes
displays organ-specific expression patterns and may protect plants from oxidative
damage under different conditions (Sadanandom et al., 2000; Bechtold et al., 2004;

Romero et al., 2004).

129

Altogether, 12 genes, five MsrA and seven MsrB, are represented on the Affymetrix
chip. The expression levels of MsrA3 (cytosolic), MsrA4 (plastidic) and MsrB7
(cytosolic) increased continuously, while that of MsrB] @lastidic) continuously decrease
during LT treatment (Figure 4.9). The expression of MsrAI, MsrAZ, MsrB] and MsrBZ
were temporarily repressed at 48h but tended to recover to pre-treatment levels at 120h of
LT (Figure 4.9). Among all these Msr genes, only MsrB5, which likely localizes in the
cytosol (Rouhier et al., 2006), showed significantly different expression between vte2 and
Col at 48h of LT treatment. Before LT treatment, vte2 and Col had similar levels of
MsrB5 expression, however, after 48h of LT treatment, vte2 showed a more than 3 fold
decrease relative to C01, and remained repressed in vte2 by 120h of LT. The transcript of
MsrBS was reported to be most abundantly expressed in roots (Rouhier et al., 2006), it
would be interesting to locate the transcription signals of MsrB5 in LT-treated leaf tissue.

The other group of genes that was specifically repressed in 48h-LT-treated vte2
include four genes in the nodulin drug/metabolite transporter (DMT) superfamily, a
family of structurally homologous membrane proteins with diverse transport fimctions.
Three of the four nodulins belong to the MtN21 family, which have 10 putative (it-helical
transmembrane spanners (TMSs) and are proposed to be localized in plasma membranes
(Rouanet and Nasser, 2001). Several members of the DMT family in bacteria have been
implicated in transport of various solutes (Ferguson and Krzycki, 1997; Tucker et al.,
2003). The functions of nodulins in plants are largely uncharacterized, except for a few
involved in bacterial nodulation in plants (Vandewiel et al., 1990; Hohnjec et al., 2009).
Considering the enormous deposition of vascular callose and defective export capacity of

both sugars and amino acids in LT-treated vte2 (Maeda et al., 2006) and potato VTEI-

130

RNAi lines (Hofius et al., 2004), repression of the expression of these nodulins might
reflect the declined transport of certain solutes from source to sink tissues in toc0pherol-

deficient plants.

Diflerent stress responses of the 49 induced genes in vte2

To investigate how the 49 induced genes in LT-treated vte2 respond to other types of
stress conditions, their expression patterns in response to various selected abiotic and
biotic stress conditions were examined (Figure 4.10). Roughly half of the induced genes
in vte2 were also induced in response to treatment with the necrotrophic fungus Botrytis
cinerea and the pathogenic leaf bacterium Pseudomonas syringae but not with phloem-
feeding aphid Myzus persicae. Interestingly many of the same genes induced by pathogen
treatments were also induced by ozone treatment and less so by hydrogen peroxide
treatment. These genes include most of the transcription factors (WRKY75, WRKY 8,
NAM and zinc finger family proteins) and some of the stress and signal-related genes
(GST, auxin-responsive GH3 family protein). In contrast, very few induced genes in LT-
treated vte2 are responsive to abiotic stress conditions including heat, drought or cold
treatments or hormone treatments such as zeatin, IAA, ethylene or ABA. The lack of
induction of these genes by cold treatment suggests their response in vte2 is due to
tocopherol deficiency rather than cold stress per se.

Previously, the transcriptome of germinating vte2 seedlings was shown to be
substantially influenced by the elevation in nonenzymatic lipid peroxidation in the mutant
(Sattler et al., 2006). The transcriptional responses of LT-treated vte2 were further
compared with those of 3-d-old vte2 seedlings. When the same procedure of

preprocessing and statistical analysis was applied for the microarray data of germinating

131

C01 and vte2 seedlings, 744 genes were identified as significantly different (adjusted
p<0.05) in 3-d-old vteZ seedlings relative to C01 (gene list not shown). When the list of
744 genes was compared with the list of 77 significant genes in 48h-LT-treated vte2
plants, only 12 genes were in common (Table 4.4). The overlapping genes are 10
upregulated genes and two downregulated genes in both lists, including all three induced
GSTs, WRKY8 and NAM transcription factors, and a chloroplast TAG lipase
(At2g30550). Notably, these genes were also induced by Botrytis cinerea, Pseudomonas
syringae and ozone treatments (Figure 4.10), suggesting that there was at least some
degree of oxidative stress response at the transcriptional level in vte2 at 48h of LT

treatment.

Preliminary work on putative callose synthase genes

Previously, callose deposition was observed in phloem vascular parenchyma cells of
LT-treated vte2 as early as 6h of LT treatment, which coincided with the impairment of
photoassimilate export in vte2 (Maeda et al., 2006). There are 12 putative glucan synthase
like (GSL) genes encoding callose synthases in Arabidopsis (Richmond and Somerville,
2000; Hong et al., 2001). Five GSL family members (GSLl, GSL2, GSLS, GSL8,
GSLlO) have been shown to be required in microgametogenesis (Dong et al., 2005; Enns
et al., 2005; Nishikawa et al., 2005; Toller et al., 2008; Huang et al., 2009). GSL5
(PA/024) was also shown to be responsible for deposition of callose after wounding or
pathogen attack (Jacobs et al., 2003; Nishimura et al., 2003). When the gsl5 mutation was
introduced into the vte2 background, the double mutant showed a significant but
incomplete reduction of callose deposition relative to vte2 in response to LT treatment

but this did not impact photoassimilate export defect, soluble sugar accumulation and leaf

132

purple coloration and plant growth inhibition (Maeda, 2006). These data indicate that
GSL5 is responsible for a large portion of callose deposited in LT-treated vte2, but that
GSL5-dependent callose deposition is not the root cause for the subsequent
photoassimilate export defect and may be secondary effect brought about by other factors

or callose deposition by other GSL family members.

Two putative callose synthase genes, GSL4 (At3gl4570) and GSL]! (At3G59100)
were induced in 48h-LT-treated vte2 (Table 4.1). To investigate if either of these callose
synthase like genes is responsible for the GSL5-independent callose deposition in LT-
treated vte2, homozygous mutants of gsl4 and gslI] fi'om available T-DNA inactivation
lines were selected and introduced into the vte2 background. The single gsl and vte2
mutants and gsl4vte2 and gslI Ivte2 double mutants were subjected to LT treatment to
assess their LT phenotypes. Before being transferred to LT conditions, all the single and
double mutants appeared visually similar to C01 and vte2 (Figure 4.11, panel A). After
prolonged LT treatment (28d), the single mutants appeared similar to C01 while both of
the double mutants were smaller and purple, similar to vte2 (Figure 4.11, panel B),
suggesting that neither gsl4 nor gslI] mutation alone has any impact on the vte2 LT
phenotype. This is also reflected at the level of vascular callose deposition (Figure 4.12),
indicating that gsl4 or gslI] mutation is not responsible for the callose deposited in LT-
treated vte2. It is possible that GSL4 and GSL] 1 are functionally redundant in this regard
or alternatively other GSL genes, which are not regulated at the transcriptional level, are
responsible for the non-GSL5-dependent callose deposition in LT-treated vte2.
Generating gs14gslIIvte2, gsl4gs15vte2 and gslI Igsl5vte2 triple mutants and a

gsl4gsl5gsll Ivte2 quadruple mutant would be required to address this question.

133

 

of

58'

Elm

11M

DISCUSSION

This study indicates that tocopherol deficiency does not impact the transcriptome of
Arabidopsis at permissive conditions. Compared with the transcriptional responses of
vte2 seedlings after 3d of germination where accumulation of massive amounts of non-
enzymatic lipid peroxidation products had a strong impact on the expression of a
relatively large amount of genes, relatively very few genes in vte2 showed different
expression relative to C01 after 48h of LT treatment.

The biochemical phenotype of 48h-LT-treated vte2 plants includes phloem transfer
cell-specific callose deposition that has started to spread from the petiole to the upper part
of the mature leaves and impact on the capacity of source to sink photoassimilate
transportation (Maeda et al., 2006). Consistent with these phenotypes, the expression of
several genes which are likely involved in the cell wall modification in phloem transfer
cells were significantly upregulated in vte2 (Table 4.1). Further analysis suggested that
these cell-wall modifying genes contain W box elements in the promoter regions (Table
4.3) and may be regulated by WRKY transcription factors. In addition, the expression
levels of several nodulin genes which may be involved in solute transport, were repressed

(Table 4.2), probably as a response to the blockage of phloem transport process.

Previous reports (Maeda et al., 2006; Maeda et al., 2008) and the characterization of
suppressors in Chapter 2 and Chapter 3 have highlighted the involvement of tocopherols
in ER lipid metabolism under low temperature conditions. Based on these results, it
might be expected that some genes related to lipid metabolism might be differentially
expressed in vte2. Surprisingly, only 2 of the 77 genes differentially expressed in vte2 are

involved in lipid metabolism. Both of them are lipase class 3 family proteins and

134

proposed to have triacylglycerol lipase activities, with one of them (At2g30550) localized
in the chloroplast and the other (At1g30370) in mitochondria. Previous studies have
shown that the majority of fatty acid desaturase genes in Arabidopsis do not respond to
decreases in temperature via changes in transcription (Iba et al., 1993; Okuley et al.,
1994; Heppard et al., 1996), or to alterations in the membrane fatty acid composition due
to mutations in the desaturase pathway (Falcone et al., 1994). Only the relatively unusual
case of the desaturation step catalyzed by FAD8 (Gibson et al., 1994), and other highly
specific examples for particular plant species (Berberieh et al., 1998), have been shown to
be regulated at the level of transcription by temperature. It seems likely that the effect of
tocopherol deficiency on ER lipid metabolism in LT-treated vte2 plants does not occur
primarily through transcriptional regulation but rather at the post-transcriptional and

enzymatic activity levels.

Although no obvious elevation of lipid peroxidation was detected in LT-treated vte2
(Maeda et al., 2006; Maeda et al., 2008), comparisons of the expression of the
significantly different genes in vte2 across various abiotic and biotic treatments (Figure
4.10, Table 4.4) suggests that at least some degree of oxidant stress was occurring in vte2
afier 48h of LT treatment. Because all biochemical assays of lipid oxidation required
whole leaf tissue for practical reasons, it is possible that oxidation was occurring only in
small fraction of cells (e.g. transfer cells) and that this signal is diluted in whole leaf
assays. It is possible that these oxidative stress responses reflected on the level of protein
oxidation, as MsrB5 was significantly repressed in LT-treated vte2 (Figure 4.9) and such

misregulation could induce covalent modification of the firnctions of related enzymes.

135

As with biochemical analyses, the experimental materials for this microarray
analysis were of necessity taken from whole leaves (see Methods) and it is possible that
the transcriptional “signatures” present in a small portion of specialized cell types are
diluted and thus difficult to identify in bulk leaf samples. Consistent with this idea, most
of the genes differentially expressed in LT-treated vte2 have low expression levels and
attempts to verify their expression by traditional RNA gel blot analysis failed (data not
shown). It is possible that transcriptional responses may only be present in certain cell
types of vte2, especially in its transfer cells where endomembrane biogenesis is largely
enhanced and the deposition of callose and abnormal cell wall ingrowths occur. In situ
hybridization or laser-mierodissection of specific cells especially the vascular
parenchyma cells will help to further confirm the expression levels and possible
localization specificity of some of the 77 genes identified in this study. Future
experimental investigations with real-time PCR and analysis of post-transcriptional
processes, such as protein oxidation, will be necessary to elucidate how these genes may

be regulated by tocopherols and contribute to the LT-induced phenotype of vte2.

136

MATERIALS AND METHODS

Experimental design

A factorial design comprising of two genotypes (Col, vte2) at three different time
points (Oh, 48h, 120h of LT treatment) was chosen. Three independent biological
replicates were included for each genotype at each time point. Experiments for the first
two replicates for C01 and vte2 at Oh time point were performed with the first set of plants
in May-June of 2005 and the rest of the experiments were finished with a second set of
plants during July-Sep of 2006. Fresh seed were used for both sets, plants were grown in

the same chamber with identical conditions.

Plants and growth conditions

Arabidopsis thaliana seeds from wild type (eeotype Columbia-O) and vte2-1 were
planted on vermiculite and potting soil mixture and stratified in the dark for 5 days at 4°C
before being transferred to permissive conditions with 12 hours of light (lOOuE, 22°C)
and 12 hours of dark period (18°C). Plants from both genotypes were randomly assigned
to different locations in the chambers. Plants were fertilized with leoagland solution
every week. For Oh time point, plants of vte2 and Col were grown to 4 weeks at
permissive temperatures and the 9th to 11th leaves from 3 plants of the same genotype
were harvested. For 48h and 120h time points, 4 week old plants were transferred to LT
conditions (75uE, 7.5:3°C, 12 hour light) and the 9th to 11th leaves from 3 plants of the
same genotype were harvested when plants were treated for 48 and 120 hours,
respectively. All leaf materials were harvested directly into tubes filled with liquid

nitrogen when plants were at 1 hour of light cycle.

137

RNA extraction, labeling and hybridization

Total RNA was extracted using the RNAqueous RNA extraction kit and the Plant
RNA Isolation Aid (Ambion) according to the manufacturer’s protocol. Labeling and
hybridization of RNA were conducted using standard Affymetrix protocols by the
Michigan State University DNA Microarray Facility. ATHl Arabidopsis GeneChips
(Affymetrix, Santa Clara, CA) were used for measuring changes in gene expression
levels. Total RNA was converted into cDNA, which was in turn used to synthesize
biotinylated cRNA. The cRNA was fragmented into smaller pieces and then was
hybridized to the GeneChips. After hybridization, the chips were automatically washed
and stained with streptavidin phycoerythrin using a fluidics station. The chips were
scanned by the GeneArray scanner by measuring light emitted at 570nm when excited

with 488-nm wavelength light.

Data evaluation and preprocessing

The raw chip data were analyzed with R software (version 2.9, http://www.r-
project.org/). The RNA degradation plot for all 18 chips was generated using the
AffyRNAdeg function in the simpleAffy package. Boxplot and Histogram tools included
in the affy and simpleaffy packages were used to investigate the data distribution of the
18 chips. RMA function as implemented in the affy package was used for background
adjustment, normalization and summarization. The cluster dendrogram (Figure 4.8) was
generated applying hclust function using average linkage clustering of Euclidean distance

based on the normalized expression values from 18 chips.

138

Statistical analysis for significant genes

As we are interested in genes differentially expressed between genotypes at different
time points of cold treatment and also the differential response of genotypes to different
durations of cold treatments. Considering this complexity, the limma package
implemented in Bioconductor of R software was used for statistical analysis, because it is
designed to analyze complex experiments involving comparisons between many RNA
targets simultaneously. Another consideration is that use of empirical Bayes method in
limma can borrow the information across genes making the analysis stable even for
experiments with small number of arrays, as in this case. For statistical analysis, the
signal intensity data were analyzed with use of a linear statistical model and an empirical
Bayes method for assessing differential expression in the limma package (Smyth, 2005).
Genes with <5% of adjusted P values were considered significant. Cluster analysis of the
77 significantly different genes in vte2 at 48h of LT treatment was performed by
Pearson’s method.

Data of Oh chips were also analyzed with Agilent’s GeneSpring analysis platform
(version 7.2). The data files generated by the Affymetrix microarray suite version 5
software (MASS) were imported and subjected to recommended normalization process
(the expression values <0.01 were set to 0.01; the data are normalized to 50% percentile
per chip and median per gene). The data were then filtered using the detection call
(Present/Absent) and probe sets called “Present” in either genotype were further filtered
with “2 fold change” between the genotypes. The probe sets after the above advanced

filtering were finally analyzed with t test (False discovery rate < 0.05).

139

The 49 genes which are significantly induced in 48h LT-treated vte2 relative to wild-
type plants were examined using the Meta-Analyzer feature of GENEVES'IIGATOR
(Zimmermann et al., 2004) to assess their responses to various conditions or treatments.
The gene expression responses were calculated as ratios between treatment and negative
control samples and the resulting values thus reflect up- or down-regulation of genes.
Twelve different conditions were chosen: Biotic stress includes treatments with Bottytis

cinerea (6 treatment chips + 6 control chips), Pseudomonas syringae (9+9) and Myzus
persicae (3+3). Chemical stress includes hydrogen peroxide (H202) (3+2) and ozone

treatments (3+3). Hormone stress includes treatments with ABA (2+2), ethylene (3+3),
1AA (3+3), zeatin (3+3). Abiotic stress includes treatments with cold (6+6), drought
(3+1) and heat (2+2). Only high-quality 22K chips were used for the analysis. For Figure
4.12, the data for Od- and 3d- old seedlings of C01 and vte2 (three biological replicates at
each treatment, see (Sattler et al., 2006) were subjected to the same procedure of
preprocessing and statistical analysis as described above for vte2 plants under LT

treatments.

Promoter analysis

The conservative four amino acid sequence of W box (TGAC) was used for
promoter regulatory sequence analysis using the promoter analysis tool implemented in
GeneSpring. 1000bp upstream of 49 induced genes in 48h-LT—treated vte2 was searched.
The promoters containing TGAC were further checked for the presence of the consensus

sequence of W box (C/T)TGAC(T/G).

140

Generating double homozygous mutants and LT treatment

The following T-DNA insertion lines were obtained from the Arabidopsis Biological
Resource Center at Ohio State University: SALK_OOOSO7 (exon) for GSL4 (At3g14570)
and SALK_Ol9534 (exon) for GSL]! (At3G59100). Homozygous mutant lines were
screened with PCR and crossed into vte2. Double homozygous mutants were obtained by
PCR screening for the presence of the insertions and vte2 tocopherol deficiency with
HPLC. The vte2 allele was confirmed by CAPs marker developed for vte2-l point
mutation (Maeda et al., 2006). Plants of wild type (Col), vte2, single mutants of gsl4,
gslI] and double mutants of vte2gsl4 and vte2gslII were grown for 28 d at permissive
conditions and then transferred to LT conditions for various time for evaluation of LT-

induced phenotypes.
ACKNOLEDGEMENTS

We would thank Maria Magallanes—Lundback for performing RNA extraction and
labeling and the Research Technology Support Facility (RTSF) at Michigan State

University for performing microarray analysis.

141

TABLES AND FIGURES

Table 4.1 The 49 genes significantly upregulated in vte2 relative to C01 at 48h of LT
treatment

M-value (M) is the value of the contrast and represents a log; fold change between 48h-
LT-treated vte2 and Col. adj-P.value is the p-value adjusted for multiple testing with
Benjamini and Hochberg’s method to control the false discovery rate. B-statistic (B) is
the log-odds that the gene is differentially expressed. Description of gene functions was

obtained from the Gene Ontology section of The Arabidopsis Information Resources.

 

AGI Adj.P.
number Val
At1g26450 2.62 0.00 1 1.64 beta-l,3—glucanase-related
At4g23410 1.53 0.00 10.08 Senescence-associated family protein
Atlg68290 2.39 0.00 7.97 bifunctional nuclease, putative
At5g22860 1.38 0.00 7.04 serine carboxypeptidase S28 family protein
_A_t3g14570 1.85 0.00 6.88 glycosyl transferase family 48 protein
Atl $59500 1.58 0.00 6.79 auxin-responsive GH3 family protein
At3g17690 1.80 0.00 6.74 cyclic nucleotide-binding transporter 2 / CNBT2
Atl g74590 1.50 0.00 6.31 Glutathione S-transferase, putative
At4g20320 1.08 0.00 5.80 CTP synthase, putative / UTP-ammonia ligase, putative
A t3g2291 0 1.82 0.00 5.65 fjétipgporting ATPase, plasma membrane-type, putative
At5gl3080 1.88 0.00 5.51 WRKY family transcription factor (WRKY75L
At5g47920 1.06 0.00 5.13 expressed protein
At1g68620 2.08 0.00 4.95 expressed protein
Atlg65500 2.13 0.00 4.84 expressed protein
Atlg76640 2.52 0.00 4.54 calmodulin-related protein, putative
At2g26020 2.01 0.00 4.40 plant defensin-fusion protein, putative (PDF1.2b)
Atl g74055 1.00 0.00 4.18 expressemotein
Atl g30370 1.60 0.01 3.80 lipase class 3 family protein
At5g13880 1.45 0.01 3.60 expressed protein
At3g49l30 0.94 0.01 3.16 hypothetical protein
At5 g46590 1.31 0.01 3.09 no apical meristem (NAM) family protein
At3g21780 0.91 0.01 3.09 UDP-glucosyl transferase family protein
At1g17180 1.21 0.02 2.82 lutathione S-transferase, putative
Atl g65610 1.33 0.02 2.70 endo-l,4-betajlucanase, putative / cellulase, putative
‘ At3g53600 0.70 0.02 2.39 zinc finger (C2H2 type) family protein
At5g13170 1.01 0.02 2.37 nodulin MtN3 family protein

B Annotated gene function

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

142

Table 4.1 (cont’d)

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

AGI M Ad‘I'P' B Annotated gene function

number Val
At5g46350 1.02 0.02 2.34 WRKY family transcrjnion factor (WRKY8)
At5g09470 0.93 0.02 2.26 mitochondrial substrate carrier family protein
At4g35730 1.24 0.02 2.23 guessed protein
At2g29090 1.07 0.03 2.15 cytochrome P450 family protein
At2g30550 0.70 0.03 2.08 lipase class 3 family protein
At4g3 8420 1.58 0.03 2.02 multi-copper oxidase type 1 family protein
At4g27260 0.76 0.03 1.98 auxin-responsive GH3 family protein
At3g59100 1.04 0.03 1.81 l cosmransferase family 48 protein
At4g19460 0.80 0.03 1.80 glycosyl transferase family 1 protein
At3g09270 2.07 0.03 1.73 glutathione S-transferase, putative
At5g22570 1.18 0.03 1.72 WRKY family transcription factor (WRKY38)
Atl 332350 1.45 0.04 1.64 alternative oxidase, putative
At5gl7330 0.74 0.04 1.54 glutamate decarboxylase 1 (GAD l)
At4g28550 1.05 0.04 1.43 RabGAP/TBC domain-containing protein
At5g65600 1.27 0.04 1.38 legume lectin family protein / protein kinase family protein
At4g36430 0.79 0.04 1.37 peroxidase, putative
At5g04080 0.63 0.04 1.32 expressed protein
At5g64905 1.78 0.04 1.25 expressed protein
At5g66920 1.09 0.04 1.24 multi-copper oxidase type 1 family protein
At5g63970 0.97 0.05 1.20 copine-related
At2g23270 0.95 0.05 1.20 expressed protein
At1g19250 0.86 0.05 1.10 flavin-containing monooxygenase family protein
At5g67080 1.69 0.05 1.08 protein kinase family protein

 

143

 

Table 4.2 The 28 genes significantly downregulated in vte2 relative to C01 at 48b of

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

LT treatment
AGI M Adj. B Annotated Gene function
number P.Val

At2g18950 -2.36 0.00 7.93 HPT: tocopherol phytyltransferase

At5g14740 -0.77 0.00 6.32 carbonate dehydratase 2 (CA2) (CA18)

App 1930 -1.47 0.00 6.10 Universal stress protein (USP) family protein

At2g36830 -1.01 0.00 5.94 major intrinsic family protein / MIP familmotein

At1g04680 -0.74 0.00 5.41 pectate lyase family protein

Atlg76800 -l.43 0.00 5.36 nodulin, putative

At4L08300 -2.98 0.00 4.58 nodulin MtN21 family protein

At2g22330 -l.67 0.00 4.44 cytochrome P450, putative

At3g10080 -0.72 0.01 3.86 ermin-like protein, putative

At5g44720 -1.05 0.02 2.89 molybdenum cofactor sulfurase familyprotein

At5g23020 -3.81 0.02 2.88 2-isopropylmalate synthase 2 (IMSZ)

At3g47470 -0.84 0.02 2.77 chlorophyll A-B binding protein 4, chloroplast / LHCI type
III CAB-4 (CAB4)

Atlg51400 -0.88 0.02 2.61 photosystem II 5 kD protein

At4g08290 -1.22 0.02 2.56 nodulin MtN21 family protein

At1g21440 -1.17 0.03 2.18 mutase family protein

At1g01620 -0.83 0.03 2.02 plasma membrane intrinsic protein 1C (PIPlC) / aquaporin
PIP1.3 (PIPl .3L/transmembrane protein B (TMPB)

At3g08940 -0.82 0.03 1.95 chlorophyll A-B binding protein (LHCB4.2)

At5g24490 -1.22 0.03 1.91 30S ribosomal protein, putative

At4g04830 - l .57 0.04 1.58 methionine sulfoxide reductase domain-containing protein

A%460 -1.93 0.04 1.54 nodulin MtN21 family protein

At4g04040 -0.69 0.04 1.44 pyrophosphate--fructose-6-phosphate l-phosphotransferase
beta subunit, putative

At1g31 180 -0.85 0.04 1.41 3-isopropy1ma1ate dehydrogenase, chloroplast, putative

Atli78370 -l .07 0.04 1.40 glutathione S-transferase, putative

At5g02260 -1.23 0.04 1.38 expansin, putative (EXP9)

At5g67070 -0.51 0.04 1.28 rapid alkalinization factor (RALF) family protein

At1g13280 -0.94 0.04 1.26 allene oxide cyclase family protein

At3g09580 -0.73 0.05 1.08 amine oxidase family protein

flg03600 -0.48 0.05 1.08 photosystem 11 family protein

 

 

 

M-value (M) is the value of the contrast and represents a log; fold change between 48h-
LT-treated vte2 and Col. adj-P.value is the p-value adjusted for multiple testing with
Benjamini and Hochberg’s method to control the false discovery rate. B-statistic (B) is
the log-odds that the gene is differentially expressed. Description of gene functions was

obtained from the Gene Ontology section of The Arabidopsis Information Resources.

144

Table 4.3 Genes containing W box sequence (C/T)TGAC(T/G) in the 1000bp

upstream regions.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Probe AGI 1D Gene function Distance Sequence
(b9)
248896_at At5 g463 50 beta-l ,3-glucanase-related 929 TTGACT
766 TTGACT
758 TTGACT
717 CTGACT
252989_at At4g38420 Multi-copper oxidase type I family 879 CTGACT
rotein
254575_at At4g19460 glycosyl transferase family 1 protein 802 TTGACT
211 TTGACG
248777_at At5g47920 expressed protein 681 TTGACT
544 TTGACG
262229_at At1g68620 exmssed protein 664 CTGACT
264685_at At1g65610 Endo-1,4-beta-glucanase, putative / 531 TTGACT
cellulase
251950_at At3g53600 zinc finger (C2H2 type) family protein 521 TTGACT
509 TTGACT
247026_ at At5g67080 protein kinase family protein 476 CTGACT
251499 at At3g59100 glycosyl transferase family 48 protein 458 TTGACG
252289_at At3g49130 hypothetical protein 433 TTGACT
160 TTGACT
256833_at At3g22910 Ca” ATPase, plasma membrane-type 401 TTGACT
(ACA13)
253768_at At4g28550 beta-1,3-glucanase-related 399 TTGACT
256306 at Atlj30370 Lipase class 3 family protein 375 TTGACT
250211 at At5g13880 expressed protein 316 TTGACT
261004’at At1g26450 beta-1,3-glucanase-related 280 TTGACT
245982_at At5g13170 nodulin MtN3 family protein 195 TTGACT
248855__at At5g46590 no apical meristem (NAM) family 169 TTGACT
protein
262517_at Atlg17180 Glutathione S-transferase, putative 153 CTGACT
260225_at At1g74590 Glutathione S-transferase, putative 134 TTGACT
247145 at At5g65600 legume lectin family protein 127 TTGACT
262099_s At1g59500 Auxin-responsive GH3 family protein 125 TTGACT
__at 108 CTGACG
258377_at At3gl 7690 cyclic nucleotide-binding transporter 2 / 124 TTGACG
CNBT2 (CNGC19)

 

145

Table 4.4 Common 12 genes between the 77 significantly different genes in 48h-LT-
treated vte2 plant and 744 significantly different genes in 3-d-old vte2 seedling.

Data of seedling of LT-treated plants were subjected to the same process of limma
analysis for significant genes (see Methods). The overlapping 12 genes between the 744
differentially expressed genes in 3—d-old vte2 seedlings and 77 differentially expressed
genes in 48h-LT-treated vte2 plants were listed. M-value (M) is the value of the contrast
and represents a log fold change, and adj. P, the p-value adjusted for multiple testing
with Benjamini and Hochberg’s method to control the false discovery rate, were shown
for 48h-LT-treated vte2 plant and 3-d-old vte2 seedling, respectively. Descriptions of

gene function and cellular component were obtained from the Gene Ontology section of

The Arabidopsis Information Resources.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

48h-LT“ 3-d-old vte2
AGI treated vtez . . GO molecular GO cellular
seedllng Gene title .
number plant function component
M adj.p M adj.p
At1g68290 2.39 0.00 2.01 0.00 bifunctional . nucleic acrd binding /// endomembrane
nuclease, putative endonuclease actlvrty system
At1g65500 2.13 0.00 4.33 0.00 expressed protein end‘m‘embme
system
glutathione S- glutathione transferase
At1g74590 1.50 0.00 2.19 0.00 transferase activity cytoplasm
no apical transcription factor
At5g46590 1.31 0.01 1.93 0.00 meristem (NAM) activity /// DNA ---
family protein binding
Atlgl7180 1.21 0.02 1.93 0.03 glu‘a‘h'm’e S“ glulalhmnetmnfiemse cytoplasm
transferase actrvrty
WRKY family transcription factor
At5g46350 1.02 0.02 1.69 0.00 transcription activity /// DNA Nucleus
factor (WRKY8) bindinL
At3g09270 2.07 0.03 1.29 0.02 glutamwm 5‘ gluialhwne "a“Sfe’ase Cytoplasm
transferase aCtIVlty
At2g30550 0.07 0.03 1.02 0.05 "We Class? mtclf'g'yc‘imHPase chloroplast
family protein actlv1ty
eroxidase peroxidase activity M Endomembrane
At4g36430 0.79 0.04 3.20 0.00 p . ’ calcium ion binding ///
putative . . . system
oxrdoreductase actrvrty
A15j64905 1.78 0.04 1.10 0.01 expressed motein --- ---
At1g76800 -1.43 0.00 -0.74 0.02 nodulin, putative --- «-
At3g09580 -073 0.05 -090 0.02 “ml“? °"'d‘i‘:°’e oxidoreductase activity Chloroplast
famllpproteln

 

146

 

30—

20“

Mean intensity: sifted and scaled

 

 

 

l l l I I l
0 2 4 6 8 10

5' ( ----- > 3'
Probe number

Figure 4.1 RNA degradation plot for the 18 arrays used in this study.
For each array, the probes are ordered from the 5’ end of the targeted transcript. Three

replicates for each treatment were shown in the same color for clarity.

147

Figure 4.2 QC plot of 3’: 5’ ratios for control genes, percentage of present gene calls
and background levels.

Dotted horizontal lines separate the plot into rows, one for each chip. Dotted vertical lines
provide a scale from —3 to 3. Each row shows the array index (C0.1-3 stands for Col 0h
replicates 1-3, C48.1-3 stands for Col 48h replicates 1-3, and C120.1-3 stands for Col
120h replicates 1-3 while v01-3 stands for vte2 Oh replicates 1-3, v48.1-3 stands for vte2
48h replicates 1-3, and v120.1-3 stands for vte2 120h replicates 1-3, respectively), %
present, average background, scale factors (plotted as a red line from the center line of
the image. A line to the left corresponds to a down-scaling, to the right, to an up—scaling),

3’:5’ ratios of GAPDH (blue circles) and B-actin (blue triangles) for an individual chip.

148

V1203
V1202
V1201
01203
C1202
01201
V483
V482
V401
C483
C482
C401
403
v0.2
401
C03
C02
C01

Figure 4.2

81.71%
98.19
57.8%
105.25
55.87%
98.49
81.48%
102.88
80%
109.82
58.94%
108.8?
59.38%
91.92
80.14%
102.52
55.18%
99.8
58.8%
109.34
57.93%
105.84
58.01%
108.78

149

 

w __-__.._.___.._______.._____-----------—----—_______-_--....-....._--

 

 

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I NON;
I ...ON;
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Array Index

 

 

8
Emcee: N.33

Figure 4.3 Box plot of all perfect match (pm) intensities (Log;) of unnormalized 18

array data set.

C0.1-3 stands for Col Oh replicates 1-3, C48.1-3 stands for Col 48h replicates 1—3, and

C120.1-3 stands for Col 120h replicates 1-3 while v01-3 stands for vte2 Oh replicates l-

3, v48.1-3 stands for vte2 48h replicates 1-3, and v120.1-3 stands for vte2 120h

replicates 1-3, respectively.

150

 

 

 

 

 

 

 

 

 

1.4 _
1.2 _ 1': --- 00.1
,'. '“ C02
1‘. 00.3
"ti “‘ v01
10 _, ll‘l ---
l g_. ‘7“ v0.2
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: ' "‘11, C401
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0.4 .1 .':
0.2 —I ‘1‘]:
I!"
0.0 ——t —l.“l __a-4_
l l l l l
6 8 10 12 14
L092 intensity

Figure 4.4 Histogram of kernel density estimates of all perfect match (pm) probe

intensities (Log2) of unnormalized 18 array data set.

C01—3 stands for Col 0h replicates 1-3, C48.1-3 stands for Col 48h replicates 1-3, and
C120.1-3 stands for Col 120h replicates 1-3 while v01-3 stands for vte2 0h replicates 1-3,

v48.1-3 stands for vte2 48h replicates 1-3, and v120.1-3 stands for vte2 120h replicates 1-

3, respectively.

151

 

 

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INng
I VON;
ImdNFO
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Imdvo
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3323:: $3

Array Index

Figure 4.5 Box plot of all perfect match probe intensities (Log) of quantile

normalized array data.

C0.l-3 stands for Col Oh replicates 1-3, C48.1-3 stands for Col 48h replicates 1-3, and

C120.1-3 stands for Col 120h replicates 1-3 while v01-3 stands for vte2 0h replicates 1-3,

v48.1-3 stands for vte2 48h replicates 1-3, and v120.1-3 stands for vte2 120h replicates 1-

3, respectively.

152

 

06

 

 

 

 

 

 

 

 

ii‘ a-
2 0' 7
CD
‘0
(\1
c5-
0. .4
O
I l l
6 8 1O 12 14
logintensity

Figure 4.6 Histogram of kernel density estimates of all perfect match probe

intensities (Log2) of quantile normalized 18 array data set.

153

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

140
120‘
100‘
E
09 _.
'65 80
.: F?—
60 l E N
9
N
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048.3
v48
v48 1
v48.2
C120 2
C1201
C1203
V1201 ___._

__l
v120.3 __|
co 3

V1202

Figure 4.7 Cluster dendrogram of a correlation matrix for all-against-all chip
comparisons.

C01-3 stands for Col 0h replicates 1-3, C48.1-3 stands for C01 48h replicates 1-3, and
C120.1-3 stands for Col 120h replicates 1-3 while v01-3 stands for vte2 0h replicates 1-3,
v48.1-3 stands for vte2 48h replicates 1-3, and v120.1-3 stands for vte2 120h replicates 1-

3, respectively.

154

Expression

Li

 

0001020304050607080910 1.215 2.02.510 4.0 5.0

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gem» ‘ are

Col_ 0 Col_ 48 Col_ 120 vte2_ 0 vte2_ 48 vte2_ 120

Figure 4.8 A gene tree of the 77 significantly different genes in 48h-LT-treated vte2
relative to Col.

The color bar represents different levels of expression (logz). The groups (labeled as I, II,
III and IV) were based on general expression patterns across C01 and vte2 at three time

points of LT treatment.

155

200
L50
LOO
050
000

ZOO
L50
LOO
050
000

250
200
L50
LOO
050
000

300

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Oh 48h 120h
A4

Oh 48h 120h
82

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L50

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050

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250
200
L50
LOO
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250
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L50
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050
000

500
000
300
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0h 48h

Oh 48h

/

120h
B1

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120h
35

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Oh 48h 120h
39
0h 48h 120h

Figure 4.9 Normalized expression levels of Msr genes in Col (gray) and vte2 (purple)

at different time points of LT treatment.

Al-AS, Bl-B3, BS-B7 and B9 represents the individual MsrA or MsrB genes on the
Affymetrix chip, respectively. Based on targeting prediction software, MsrAI-A3 are
cytosolic and MsrA4 is plastidic. MsrB] and MsrBZ are plastidic, MsrB3 is addressed to a

secretory pathway and the remainder of the MsrB genes are likely located in the cytosol.

The expression of MsrB5 in vte2 (red) is significantly different from Col (blue) (adjusted

p value <0.05) at 48h of LT treatment and is highlighted. Standard errors are indicated

(n=3).

156

Figure 4.10 A heat map of expression of the significantly upregulated 49 genes in
48h LT-treated vte2 under different conditions.

Genes (AGI number) significantly induced in 48h LT-treated vte2 relative to C01 plants
were examined using the Meta-Analyzer feature of GENEVESTIGATOR to assess their
responses to various conditions or treatments relative to each corresponding negative
control. The conditions and treatments were sorted by increasing ratios from the left to

the right. The color bar represents different levels of expression (logz)

157

 

Figure 4.10

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158

0'3 9‘ l O'l 9'0 0'0 9'0‘ 0' L' 9' L' 0'3' 9'3-

9'3

A Before LT treatment
Col vte2 gsl4vte2 95/11 95/11 vteZ

p

B After 14d LT treatment
Col vte2 gsl4 gsl4vt92 gsl11 gsl11vt92

4 O 7‘0 Q -¢; 9

 

Figure 4.1] Whole plant phenotype of Col, vte2, gsl4, gsl4vte2, gslI], gslIIvte2.
All genotypes were grown under permissive conditions for 4 weeks and then transferred
to LT conditions. Representative plants of the indicated genotypes are shown before LT

treatment (A) and after 28d of LT treatment (B). Bar=20m.

159

gs,“
vte2 gsl4vteZ gsl11vt92

Figure 4.12 Vascular callose deposition in Col, vte2, gsl4, gsl4vte2, gslI] and

 

gslIIvteZ after 3 d of LT treatment.
All genotypes were grown under permissive conditions for 4 weeks and then transferred
to LT conditions. Samples for callose staining were fixed in the middle of the light cycle

after 3 d of LT treatment. Representative images are shown (n=3). Bar : 1mm.

160

CHAPTER 5 SUMMARY AND FUTURE PERSPECTIVES

Tocopherol levels in plant tend to increase in response to various abiotic stresses
including high intensity light, salt, cold, drought, heat, osmotic and heavy metals
(Munne-Bosch and Alegre, 2000; Collakova and DellaPenna, 2003b; Muller-Moule et al.,
2003; Maeda et al., 2006; Abbasi et al., 2007; Collin et al., 2008). Some phenotypes of
tocopherol deficient mutants are consistent with an important role for tocopherols in
controlling lipid peroxidation, however in many cases the lack of expected phenotypes
consistent with enhanced oxidative stress in tocopherol deficient mutants indicates long
held assumptions about the role and function(s) of these compounds in plants are
erroneous (Porfirova et al., 2002; Havaux et al., 2005; Kanwischer et al., 2005; Abbasi et
al., 2007). For instance, the absence of enhanced lipid peroxidation in Arabidopsis vteZ
plants indicates the photoprotective role of tocopherols is limited and studies indicate
many functions of tocopherols may be both stress- and species-specific. However, one
common phenotype shared by maize (sxdl), potato (VTEl-RNAi) and Arabidopsis (vte2
ad vte1) tocopherol deficient lines is disruption in photoassimilate transportation from the
source leaves to sink tissues (Russin et al., 1996; Provencher et al., 2001; Hofius et al.,
2004; Maeda et al., 2006). This common phenotype between dicotyledous and
monocotyledous species suggests that tocopherols are required for the development of
full photoassimilate transport capacity, and this function is evolutionarily conserved.
However, the mechanism of how tocopherol deficiency induces the photoassimilate
export phenotype was elusive. The inducible nature of the phenotypes in Arabidopsis has

provided a unique and ideal platform to study this aspect of tocopherol functions.

161

In this thesis work, a suppressor screen of EMS mutagenized vte2 was carried out to
identify second site mutations which impacted the vte2 LT-induced phenotypes (Chapter
2, Chapter 3). Transcript profiling studies were also performed on mature plants of vte2
and Col prior to and during LT treatment to assess the whole genome responses (Chapter
4). The unbiased approaches of both studies were aimed at identifying the molecular links
between tocopherol deficiency and the vte2 LT-induced phenotypes.

Seven suppressors of vte2 LT -induced phenotype (sve loci) were identified.
Preliminary characterization of these primary sve loci has provided information for
categorizing them and to prioritize and select the three most significant loci for the
subsequent studies and as targets for molecular cloning.

sve1 was the strongest suppressor, had a unique fatty acid profile at permissive
conditions and was found to be a novel allele of fad2, the endoplasmic reticulum-
localized oleate desaturase. Coincidentally an independent study aiming to investigate the
interactions of various PUFA species and tocopherols (Maeda et al., 2008) also identified
fad2 as a suppressor of the vte2 LT-induced phenotype. This common finding from both
physiological and genetic screen studies reinforces the importance of the ER PC-18:1
desaturation step as an important component in the development of vte2 LT-phenotypes.
The second suppressor, sve2, showed partial suppression and also had a fatty acid
composition that differed from wild type before the LT treatment. sve2 was found to
contain a unique lipid species, trigalactosyldiacylglycerol (TGDG) and was determined to
be a new allele of trigalactosyldiacylglycerol1 (tgdl), a component of the ER-to-plastid
lipid ATP-binding cassette (ABC) transporter. Introduction of the other available tgd

mutations, tgdZ, tgd3, and tgd4, into the vte2 background similarly suppressed the vte2

162

LT phenotypes, indicating a key role for lipid transport in this process. The finding that
all tgd mutations were sve suppressors indicated that the disruption of the ER-plastid lipid
trafficking process is responsible for the suppression of vte2 LT phenotypes. The
identification of sve1 and sve2 provides clear and unbiased genetic evidence for
involvement of ER PUFA/lipid metabolism in the vte2 LT phenotype. The third sve
locus, sve7, partially suppressed all vte2 LT phenotypes, but unlike sve1 and tgd
mutations, had no detectable impact on fatty acid and lipid metabolism at permissive
temperature. sve7 suggests that processes other than constitutive alteration of fatty acid
and lipid metabolism can also impact on the vte2 LT phenotypes. sve7 has not been
cloned.

Maeda et al (2008) found that the lower 18:3 and higher 18:2 levels consistently
observed in LT-treated vte2 was due to a reduced conversion of 18:2 to 18:3 in ER
derived lipids and lipidomics further showed that a large number of lipid species were
identified to be altered in vte2 relative to wild type. However, it was not clear from this
work which lipid species alteration was critical for the development of vte2 LT
phenotypes or whether all were equally important. To gain further insight, a detailed
study on the changes of the acyl composition of ER- and plastid-derived lipids in
suppressors before and after LT treatment was carried out. The common biochemical
phenotype shared by all vte2 suppressing lines (i.e. sve1, sve2, sve7, tdg2, tgd3, tgd4 and
vte1) was an attenuation in the elevation of 18:2 in PC and PE that would otherwise occur
in vte2 at LT. Interestingly, when the levels of 18 carbon fatty acids in individual lipid
classes were plotted against the LT photoassimilate export capacity of various genotypes,

only PC-18:2 levels were found to be significantly correlated. These results indicate that

163

the increase in 18:2 esterified to PC plays a key role in the onset of the vte2 LT
phenotype.

ER-produced PC is a key intermediate in lipid metabolism because it is the
predominant substrate for ER-localized desaturases (Arondel et al., 1992; Okuley et al.,
1994), the precursor of the transport lipid molecules from ER to plastid (Ohlrogge and
Browse, 1995; Mongrand et al., 2000; Andersson et al., 2004), subjected to constant acyl
editing (Williams et al., 2000; Bates et al., 2007; Bates et al., 2009), and especially PC-
1812 species are the major donors of the DAG moiety for plastidic galactolipid synthesis
(Slack et al., 1977; Browse et al., 1986; Somerville and Browse, 1991). Therefore it is
likely that an elevated ER PC-18:2 level in LT-treated vte2 could influence many ER-
localized biochemical processes. Such suboptimal or unbalanced fatty acid acyl
compositions may affect the formation and function of ER-derived vesicles and thus
influence the coordinated and rapid development of transfer cell wall upon LT treatment.
In this regard, mutations directly (sve1) or indirectly (sve2 and all tgd) decreasing PC-
l8:2 synthesis may completely or partially restore the unbalanced PUFA composition of
ER-derived lipids used for transfer cell wall development.

Global transcript profiling study comparing vte2 and wild type plants under different
time periods (Oh, 48h, 120h) of LT treatment showed that tocopherol deficiency has no
impact on global gene expression at permissive conditions but affects a limited number of
specific genes (77 genes) after 48h of LT treatment. Gene expression profiles were
consistent with a degree of oxidative stress response by vte2 that was not reflected at the
biochemical level (Maeda et al., 2006; Maeda et al., 2008). LT-treated vteZ was also

shown to have an enhancement in expression of genes involved in cell wall modification

164

and repression of genes related with solute transport. These responses seem to be
consistent with the biochemical and physiological phenotypes observed in vte2 at the
corresponding time period of LT treatment.

The results from these biochemical, genetic and transcriptome studies are
summarized in a genetic model of vte2 LT-responses (Figure 5.1). This study has made
the involvement of tocopherols in ER lipid metabolism clear but the exact mechanism of
how tocopherols influence the ER lipid metabolism still remains unknown and there are
many questions to be answered to further understand the involved tocopherol functions.
Is PC—18:2 elevation the root cause of the vte2 LT phenotype?

All the characterized lines in this study (sve1, sve2, tgd2, tgd3, tgd4, sve7 and vte1)
suppressed the elevation in PC-18:2 in LT-treated vte2. However it remains to be
determined whether the PC-18:2 elevation is the root cause of the abnormal transfer cell
wall development, vascular callose deposition and the other downstream vte2 LT
phenotypes. Previously the fad3 single mutant was shown to have reduced export
capacity under LT (Maeda et al., 2008), which may be due to the increased 18:2—PC level
in fad3. If this is the case, it would mean that the PC-18:2 elevation causes the defective
photoassimilate export capacity independent of tocopherol status and is a general rather
than tocopherol-specific phenomenon. A 358 promoter-driven overexpressor of the ER
lineolate desaturase FAD3 was shown to have reduced 18:2 and increased 18:3 (Arondel
et al., 1992) and its analysis for the photoassimilate export phenotype could be
informative. Similarly introduction of 358::FAD3 into vte2 and the assessment of the LT

phenotype of the single and double mutants could also be informative.

165

Assessing the acyl composition of ER-derived lipid species in tocopherol-deficient
mutants in other plant species, such as maize sxdl mutant will also provide an
independent assessment of whether the alteration in ER lipid metabolism is a general
consequence from tocopherol deficiency, although in this case, the constitutive nature of
sde mutation may make data interpretation difficult.

Are tocopherols located in ER or PLAMs to affect the ER lipid metabolism and vesicle
function? .

One possible mechanism that tocopherols could affect the ER lipid metabolism is
that they are localized to the PLAM regions or even in the ER membranes so that they
have access to the ER compartment. Although there are several lines of evidence that
tocopherols are present in ER-derived oil bodies in seed (Yamauchi and Matsushita,
1976; Fisk et al., 2006; White et al., 2006), it still remains to be determined if tocopherols
are also distributed to extraplastidic sites in photosynthetic tissues. It might be possible to
isolate PLAMs, ER and chloroplast and assess the tocopherol content in the different
subcellular compartments. However such fractioning is often difficult because
tocopherols are easily oxidized during fractioning and the process is also complicated
with the likely contamination especially between ER and chloroplast.

General lipid peroxidation is clearly not involved in the vte2 LT phenotypes (Maeda
et al., 2008). Based on the suppressor studies, I have proposed that tocopherols influence
the lipid or membrane proteins involved in lipid metabolism via their other biophysical or
biochemical properties, which would likely impact processes such as vesicle function and
membrane fusion in Arabidopsis transfer cells at LT (Chapter 3). It is difficult to directly

test these possibilities. However, it is possible to use the combination of stable isotope

166

pulse labeling and ESI-MS/MS detection to evaluate the flux of phospholipid species and
any alteration of acyl editing process in LT—treated vte2. The possible role of tocopherols
in vesicle formation and function might be tested in vte2 at other process or
developmental stages which also require massive formation of vesicles, such as during
the fast elongation of root.

What is the nature of the other sve loci?

The cloning of sve1 and sve2 has provided important insight into the mechanisms of
tocopherol function. The third locus, sve7, has been backcrossed and characterized in
detail but not yet cloned. sve7 had an incomplete but strong suppression on all the vte2
LT phenotypes. It would be possible to physically clone the locus by map-based cloning
based on its visible and sugar phenotypes. However, the sugar accumulation phenotype
has large biological variations and may be technically difficult to assess in a single plant
among the segregation populations. Unlike sve1 or sve2, sve7 did not have any alteration
in fatty acid composition before LT treatment. sve7 is especially interesting as it may be
involved in regulating ER lipid metabolism under LT conditions and this feature may
help to narrow down the candidate genes for fine mapping process. A fourth interesting
sve locus is sve5, which had partial suppression of all vte2 LT phenotypes and an
abnormal fatty acid composition before LT treatment that is distinct from that of sve1 and
sve2. Preliminary backcrossing work on sve5 suggested that the fatty acid profile is
tightly linked to the suppression mutation (data not shown) and thus it may also be
possible to map and clone this locus based on its fatty acid profile. Map-based cloning of

sve7 and sve5 will be pursued by others in the lab afier my departure.

167

How to validate the results from microarray analysis and identifi/ tocopherol target
genes?

Studies of global transcript profiling comparing LT-treated vte2 and wild type have
highlighted several potentially important target genes for future studies. Several
experiments are ongoing to test the roles of some of these genes.

Although lacking direct experimental evidence for the genes identified, several genes
might associate with abnormal transfer cell wall development and callose deposition in
vte2. Isolation and introduction of single mutations for the induced callose synthase genes
(gsl4, gslI 1) into vte2 background did not yield a clear impact on the vte2 LT phenotype,
but this may be due to genetic redundancy and generation and analysis of the triple
mutant (gsl4gs111v162) and double mutants with other callose synthase genes will further
provide information in this regard.

In addition, analysis of differentially expression genes between vte2 and Col found
that six cell wall related genes were induced in 48h-LT-treated vte2 and subsequent
promoter analysis further identified the presence of W-box motif in their promoter
regions. It would be interesting to investigate whether the WRKY transcription factors
are regulators involved in cell wall modification of transfer cells. WRKY 75 showed the
strongest induction among the three induced WRKY factors. The possible regulating
roles of WRKY75 involved in vte2 LT-induced phenotype is being tested by evaluating
the phenotypes of homozygous wrky75 and the double mutant of wrky75vte2 at both
permissive and LT conditions.

Transcript profiling suggested that some degree of oxidative stress response might

occur in LT-treated vte2, with MSRBS being significantly repressed in 48h-LT-treated

168

vte2 relative to C01. Transcriptional repression of this antioxidant repair enzyme may lead
to oxidation and malfunction of certain proteins and it would be interesting to assess
protein oxidation in LT-treated vte2. The msrb5 and msrb5vte2 double mutant have been
generated and their roles in seedling germination and plant LT adaptation will be tested.
Enhanced endomembrane biogenesis and the deposition of callose and abnormal cell
wall ingrowths were only observed in the phloem parenchyma transfer cells of LT-treated
vte2 and is a remarkably cell type-specific response. In this regard it would be most
useful to isolate transfer cells by laser-microscopy dissection to evaluate the expression
or apply the in situ hybridization to localize the expression of the more interesting genes
already identified. Such studies may be able to help identify the transcriptional elements
linking tocopheorl deficiency and the manifested LT phenotypes in vte2 and lead to

further understanding on tocopherol fimctions.

169

vteZ

lack of tocopherols
22°C

No transcriptional changes
No biochemical &physiological phenotype

sve1/fad2——|4Normal Fatty acid composition I.— sve2/tgd1, tgd2, tgd3, tgd4

(PUFA synthesi5)>/ ! I7C (lipid trafficking)

14 sve7
(DMPvga)—Ii Phospholipid 18 2 increase i— (lipid metabolism regulator?)

(Some degree of oxidative stress response)

I
I

Callose deposition
(upregulation of cell-wall related genes)

I
l

Photoassimilate export defect
(repression of solute transport-related genes)

1

Sugar accumulation

1

Growth reduction

Figure 5.1 A genetic model of vte2 responses under LT treatment.

The biochemical and physiological responses are in black lettering, transcriptional
changes are in green lettering and the suppressors and the (potential) involved process are
in red lettering. 22°C and 7°C represents permissive and low temperature conditions,
respectively. The vte2 LT responses are connected with double arrows to indicate

multiple steps from one response to the other.

170

APPENDIX

TOCOPHEROLS PLAY A CRUCIAL ROLE IN LOW
TEMPERATURE ADAPTATION AND PHLOEM LOADING IN
ARABIDOPSIS

The work presented in this section has been published:
Hiroshi Maeda, Wan Song, Tammy L. Sage and Dean DellaPenna

(2006) Plant Cell 18, 2710-2732.

Author’s contributions:

Hiroshi Maeda led all the physiological and biochemical assessment except for the
diurnal carbohydrate and TEM analyses, which were performed independently by Wan
Song and Tammy L. Sage, respectively. Wan Song and Tammy L. Sage also contributed
to writing the method part for carbohydrate analysis, and the method and result portion of
TEM microscopy analysis. Wan Song also participated the assessment of various
biochemical and physiological parameters, including long-term carbohydrate,

anthocyanins, tocopherols, carotenoids, lipid peroxides, and maximum photosynthesis

efficiency (Fv/Fm) and quantum yield of PSII ((ppsu). Dean DellaPenna supervised the

entire project and involved in all aspects of manuscript writing.

l7l

ABSTRACT

To test whether tocopherols (vitamin E) are essential in protection against oxidative
stress in plants, a series of Arabidopsis yitamin E (vte) biosynthetic mutants that
accumulate different types and levels of tocopherols and pathway intermediates were
analyzed under abiotic stress. Surprisingly subtle differences were observed between the
tocopherol-deficient vte2 mutant and wild type during high light, salinity and drought
stresses. However, vte2, and to a lesser extent vte1, exhibited dramatic phenotypes under
low temperature, i.e., elevated anthocyanin levels and reduced growth and seed
production. That these changes were independent of light level and occurred in the
absence of photoinhibition or lipid peroxidation suggests the mechanisms involved are
independent of tocopherol functions in photoprotection. Compared to wild-type, vte1 and
vte2 had reduced rates of photoassimilate export as early as 6 h into low temperature
treatment, elevated soluble sugar levels by 60 h, and increased starch and reduced
photosynthetic electron transport rate by 14 days. The rapid reduction in photoassimilate
export in vte2 coincides with callose deposition exclusively in phloem parenchyma
transfer cell walls adjacent to the companion cell/sieve element complex. Together these
results indicate that tocopherols have a more limited role in photoprotection than
previously assumed but play crucial roles in low temperature adaptation and phloem

loading.

172

INTRODUCTION

Tocopherols are the best-studied class of lipid soluble antioxidants and are produced
only by photosynthetic organisms including all plants and algae, and some cyanobacteria.
Structurally, all four tocopherols (a, B, y and 5-tocopherols) consist of a chromanol head
group attached to a phytyl tail and differ only in the number and positions of methyl
groups on the chromanol ring (Figure A.l). Tocopherols are amphiphatic molecules and
in vitro studies using artificial membranes have shown that tocopherols form complexes
with specific lipid constituents and physically stabilize membranes (W assall et al., 1986;
Stillwell et al., 1996; Wang and Quinn, 2000; Bradford et al., 2003). Tocopherols can
efficiently quench singlet oxygen, scavenge various radicals, particularly lipid peroxy
radicals, and thereby terminate lipid peroxidation chain reactions (Liebler and Burr,
1992; Bramley et al., 2000; Schneider, 2005). In animals, vitamin E deficiency results in
muscular weakness and neurological dysfunction, which often coincide with elevated
lipid peroxidation (Machlin et al., 1977; Yokota et al., 2001). Recent studies have shown
that tocopherols also have functions in animals unrelated to their antioxidant activity,
such as modulation of cell signaling and transcriptional regulation (Ricciarelli et al.,
1998; Jiang etal., 2000; Rimbach et al., 2002; Kempna et al., 2004).

In contrast to the extensive studies of tocopherol functions in animals, we are only
beginning to understand tocopherol functions in the photosynthetic organisms in which
they are produced. In plants, tocopherols are synthesized and localized in plastid
membranes that are also highly enriched in polyunsaturated fatty acids (PUFA) (Bucke,
1968; 8011 et al., 1980c; Lichtenthaler et al., 1981; 8011 et al., 1985; S011, 1987; Vidi et

al., 2006) and increased tocopherol content has been correlated in the response of

173

photosynthetic tissues to a variety of abiotic stresses, including high intensity light (HL),
salinity, drought and low temperatures (Keles and Oncel, 2002; Bergmuller et al., 2003;
Collakova and DellaPenna, 2003b). Such data, together with the evolutionary
conservation of tocopherol synthesis among photosynthetic organisms, has led to the
assumption that a primary function of tocopherols is to protect photosynthetic membranes
from oxidative stresses by acting as lipid-soluble antioxidants (Foyer et al., 2002; Munne-
Bosch and Alegre, 2002). While plausible, such hypotheses are based primarily on
correlations and circumstantial evidence and have yet to be rigorously tested in planta.
The isolation of Arabidopsis mutants disrupting steps of the tocopherol biosynthetic
pathway provides powerfirl tools to directly investigate tocopherol functions in plants
(Figure A.l) (Shintani and DellaPenna, 1998; Collakova and DellaPenna, 2001; Porfirova
et al., 2002; Cheng et al., 2003; Sattler et al., 2003a).

The vte2 (vitamin E 2) mutant is defective in homogentisate phytyl transferase
(HPT) and lacks all tocopherols and pathway intermediates (Figure A.1, Table A.l). vte2
mutants are severely impaired in seed longevity and early seedling development due to
the massive and uncontrolled peroxidation of storage lipids (Sattler et al., 2004),
consistent with loss of the lipid-soluble antioxidant functions of tocopherols (Ham and
Liebler, 1995, 1997). Interestingly, the vte2 mutants that do survive early seedling
development become virtually indistinguishable from wild type under standard growth
conditions (Sattler et al., 2004), suggesting that unlike seed longevity and germination,
tocopherols are dispensable in mature plants in the absence of stress. Consistent with this,
constitutive over-expression of VT E2 in Arabidopsis increased total leaf tocopherols 4.5-

fold but had no discernible effect relative to wild type on plant grth or chlorophyll and

174

carotenoid content in the absence of stress or under combined nutrient and HL stress
(Collakova and DellaPenna, 2003b).

The vte1 mutant is defective in tocopherol cyclase activity and deficient in all
tocopherols but unlike vte2, accumulates the redox active biosynthetic intermediate 2,3-

dimethyl-6-phytyl-l,4-benzoquinol (DMPBQ) (Figure A.l, Table A.l, (Sattler et al.,
2003a). When grown at 100 to 120 umol photon m-2 s-1 vte1 plants are virtually identical

to wild type at all developmental stages (Porfirova et al., 2002; Sattler et al., 2003a;
Sattler et al., 2004). The lipid peroxidation phenotype observed in germinating vte2
seedlings was not observed in vte1 indicating the DMPBQ can fully compensate for

tocopherols as a lipid-soluble antioxidant in seedlings (Sattler et al., 2004). Under HL

stress (5 days at 850 umol photon m.2 s"; (Porfirova et al., 2002) or a combination of

low temperature and HL stress (5 days at 6 to 8°C and 1100 umol photon m.2 5-1(Havaux

et al., 2005), vte1 was nearly identical to wild type for all parameters measured, including
lipid peroxidation, with the exception of a slight decrease in maximum photosynthetic

efficiency (Fv/Fm). Only under extreme conditions (24 h at 3°C and continuous 1500-
1600 umol photon rn'2 s‘l) did vte] show a more rapid induction of lipid peroxidation

than wild type, although this difference was transient and after 48 h of treatment lipid
peroxidation was similarly elevated in vte1 and wild type (Havaux et al., 2005). These
studies with vte1 and vte2 mutants suggest that a primary function of tocopherols is to
control non-enzymatic lipid oxidation, especially during seed storage and early
germination, and also probably in photosynthetic tissues but only under the most extreme

of combined HL and low temperature stress.

175

Interestingly, a maize tocopherol cyclase mutant (sxdl, sucrose export defective I)
was identified several years prior to the identification of Arabidopsis vte1, not due to its
impact on tocopherol synthesis, but because of accumulation of carbohydrates and
anthocyanins in sde source leaves, which coincided with aberrant plasmodesmata
between the bundle sheath and vascular parenchyma cells (Russin et al., 1996). Cloning
of the SXDI locus did not provide insight into the biochemical activity of the nuclear—
encoded chloroplast-localized protein (Provencher et al., 2001) and it is only in retrospect
that SXDI has been demonstrated to have tocopherol cyclase activity (Porfirova et al.,
2002; Sattler et al., 2003a). The maize sde carbohydrate accumulation phenotype was
intriguing as it suggested an unexpected link between the tocopherol pathway and
primary carbohydrate metabolism, though the mechanism involved was unclear. A
similar carbohydrate phenotype did not occur in the orthologous Arabidopsis vte1 mutant
(Sattler et al., 2003a) but was observed in VTEI RNAi knock-down lines in potato
(Hofius et al., 2004).

In the current study, we firrther define and clarify the physiological role(s) of
tocopherols in photosynthetic plant tissues by subjecting and analyzing the response of a
suite of Arabidopsis tocopherol mutants to a variety of abiotic stresses. We report that in
contrast to long-held assumptions about tocopherol functions in plants, tocopherol-
deficient mutants are remarkably similar to wild type in their response to most abiotic
stresses with the notable exception being an increased sensitivity to non-freezing low
temperatures. Detailed physiological, biochemical and ultrastructural data demonstrate
that the earliest impact of tocopherol deficiency during low temperature treatment is an

inhibition of photoassimilate transport associated with dramatic structural changes in

176

phloem parenchyma transfer cells, a bottleneck for photoassimilate transport. The
resulting accumulation of carbohydrates in source leaves impacts the physiology and
response of the entire plant to low temperatures.

RESULTS

T ocopherol Biosynthetic (vte) Mutants Used in This Study

vte1-1, vte1-2 and vte2-1 are previously isolated and characterized ethyl
methanesulfonate mutants in the Columbia (Col) ecotype that are deficient in the
tocopherol cyclase and HPT enzymes, respectively (Sattler et al., 2003a; Sattler et al.,
2004); Figure A.l). vte2-2 and vte4-3 are T-DNA insertion mutants in the Wassilewskija
(Ws) ecotype in genes encoding HPT and y-tocopherol methyltransferase (y-TMT),
respectively. Leaves of all mutants, vte2-1, vte2-2, vte1-1, vte1-2 and vte4-3, lack a-
tocopherol, the major tocopherol in wild type Arabidopsis leaves (Figure A. 1, Table A. 1 ,
(Sattler et al., 2003a). vte2-1 and vte2—2 lack all tocopherols and pathway intermediates.
vte1-1 and vte1-2 lack all tocopherols but accumulate the biosynthetic pathway
intermediate DMPBQ at a level comparable to d-tocopherol in Col. The vte4-3 mutant
accumulates y-tocopherol at an equivalent or slightly higher level than (It—tocopherol in

Ws. Three to five-week-old plants of all mutant genotypes grown under permissive
conditions (12 h 120 pmol photon m-2 3.1 light at 22°C / 12 h darkness at 18°C) were
virtually identical to their respective wild type backgrounds, consistent with previous
reports that mutations disrupting tocopherol synthesis have little impact on the normal

growth of mature plants (Porfirova et al., 2002; Bergmuller et al., 2003; Sattler et al.,

2003a; Sattler et al., 2004).

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The Response of T ocopherol—Deficient Mutants to High Intensity Light Stress

High intensity light (HL) stress results in excessive excitation of chlorophyll and
consequently generates reactive oxygen species (ROS), which in turn attack various
biochemical targets in the cell including PUFA-enriched photosynthetic membranes.
Tocopherols are most abundant in these photosynthetic membranes (Bucke, 1968;
Lichtenthaler et al., 1981; S011 et al., 1985) and leaf tocopherol levels increase up to 18-
fold during HL stress in Arabidopsis (Collakova and DellaPenna, 2003b; Havaux et al.,
2005). Therefore, it has been presumed that the elimination of tocopherols from
photosynthetic membranes would have dramatic impacts on plant survival during HL
stress. To test this hypothesis, Col, vte2-I, vte1-1, and vte1-2 were grown for four weeks

under permissive conditions and then subjected to two levels of HL stress, 1000 and 1800

umol photon m-2 s'1 16 h light/ 8 h darkness at 22°C (hereafter referred to as HLlOOO

and HL1800, respectively). HLIOOO did not result in differential visible or biochemical
phenotypes between vte2-1 and Col (Figure A.l4). When Col, vte2-1, vte1-1 and vte1-2
were subjected to HL1800, which approaches the intensity of full sunlight, this led to
bleaching of some mature leaves in all genotypes. vte2-1 had a slight tendency toward
more bleached leaves than Col but this was not reproducible or significant, while vte1-l
and vte1-2 reproducibly had as many or more bleached mature leaves than Col or vte2-I
(Figure A.2A and Figure A.lS). vte2-2 and vte4-3 subjected to HL1800 responded
similarly to Ws, the corresponding wild type (data not shown).

To assess changes in photosynthetic pigment and tocopherol levels in response to HL
stress, the 7th to 9th oldest leaves were harvested before and after 4 days of HL1800 for

HPLC analysis. Before HL1800 all levels were similar between genotypes except for

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slightly lower B-carotene content in vte2-I and vte1-1 relative to C01 and the absence of
tocopherols in all vte genotypes (Table A.4). After 4 days of HL1800, the vte mutants
generally had lower total and individual chlorophyll levels than Col but these differences
were not significant in all cases after 4 days of HL1800, even with n = 19 (Figure A.ZB,
Table A2, Figure A.15). Total carotenoids were consistently and significantly lower than
C01 in vte1-1 and vte1-2, but not always in vte2-1 after 4 days of HL1800. Neoxanthin
and violaxanthin were significantly lower in all vte mutants, while lutein was
significantly lower only in vte1-1 and vte1-2. Interestingly, zeaxanthin was 70 % higher
than C01 in vte2-1 but unchanged relative to C01 in both vte1 alleles (Table A2).

In vivo chlorophyll a fluorescence was also analyzed to assess PSII function during
HL stress. Typically, when plants are under oxidative stress, PSII is inactivated due to
enhanced turnover of the D1 protein, a process termed photoinhibition, and maximum
photosynthetic efficiency (F v/F m) decreases (Maxwell and Johnson, 2000). Four-week-
old Col, vte2-1, vte1-1 and vte1-2 plants grown under permissive conditions had identical
Fv/F m values of between 0.8 and 0.85, typical values for healthy leaves (Maxwell and
Johnson, 2000)data not shown). After 24 h of HL1800 (8 h HL1800, 8h darkness, and 8h
HLISOO), a few vte2-1 leaves showed a dramatic reduction in Fv/Fm (< 0.5), but the
majority had values similar to C01 and the average Fv/Fm of vte2-I was not significantly
different from Col, even with n = 30 (Figure A.2D, Figure A.15). In contrast, vte1—I and
vte1-2 both had more leaves with Fv/Frn < 0.5 and average Fv/Fm values that were
significantly lower than Col (Figure A.2D, Figure A.15).

These combined results indicate that the elimination of tocopherols in vte2 has

surprisingly little impact on the response of the photosynthetic apparatus to HL stress in

179

comparison to C01, with the exception of altered xanthophyll cycle carotenoids. Equally
surprising is the fact that though the vte2 and vte1 genotypes are identical with regard to
their tocopherol deficiencies, vte1 alleles are slightly more susceptible to HL1800 than
vte2. As the primary biochemical difference between these two genotypes is that vte1
mutants accumulate the redox active intermediate DMPBQ while vte2 mutants do not, the
presence of DMPBQ in vte1 may have negative impacts on HL stress tolerance in

Arabidopsis.

T ocopherol—Deficient Mutants Exhibit a Low Temperature Sensitive Phenotype.

In searching for a condition that more obviously impacts wild type and the
tocopherol—deficient mutants in a differential fashion, vte2-1 and Col plants were
subjected to abiotic stress treatments other than HL, including salinity (100, 150 and 200
mM NaCl), drought, and various low temperature treatments. Like HL stress, the salinity
and drought stress conditions used also did not result in obvious phenotypic differences
between vte2-1 and Col (Figure A.l4) and further analyses will be required to determine
any consequences of tocopherol-deficiency during these stresses. However, when plants
were transferred from permissive conditions to non-freezing low temperature conditions,
both vte2-1 and vte2-2 grew more slowly than their respective wild types, C01 and Ws,
and their mature leaves changed color to purple (Figure A.3). These phenotypic

differences were consistently observed in conditions ranging from 3 to 12°C and light

. . . -2 -1 .
1ntensrt1es from 15 to 200 umol photon m 3 (data not shown). Differences were most

obvious and consistent under 75°C, 12 h 75 umol photon m-2 s-1 light/12h darkness

(Figure A.3) and this low temperature regime (hereafter termed 7.5°C-treated) was used

180

for all subsequent experiments.

Following transfer to 75°C, vte2-1 and Col did not differ in time to bolting (53 i4
and 51 i3 days, respectively, after transfer to 75°C) or number of leaves produced at the
start of bolting (32 i3 and 31 i2 leaves, respectively), indicating that the process of
vernalization was not affected by the lack of tocopherols. However, after prolonged
growth at 75°C vte2-1 siliques were shorter, produced significantly fewer seeds per
silique and per plant compared to C01, and 35 % of the seeds in vte2-I siliques were
aborted compared to less than 1 % in Co] siliques (Figure A.3E, Table A.3). These results
indicate tocopherols play a crucial role in low temperature adaptation in Arabidopsis.

Subjecting vte1-I to 75°C treatment resulted in a phenotype intermediate between
vte2-I and Co] in terms of overall growth, mature leaf color, silique size, number of
seeds/silique, percentage of aborted seed, and seed yield per plant (Figure A.3, Table
A.3). These phenotypes in 7.5°C-treated vte4-3 were virtually indistinguishable from
wild type (W s) (Figure A.3A, Figure A.3B and Figure A.3C). These results indicate that
during low temperature adaptation in Arabidopsis the quinol biosynthetic intermediate
DMPBQ partially compensates for the lack of tocopherols in vte1-1 while the y-
tocopherol accumulated in vte4-3 leaves can functionally replace a-tocopherol in this

regard.

Photooxidative Damage Is Not Associated with the vte2 Low Temperature Phenotype

To further examine changes during the time course of 7.5°C treatment, vte2-1 and

Col were subject to detailed comparative biochemical analyses. Plants grown for four

. . . . . . th th
weeks at permrssrve conditions were transferred to 75°C conditions and the 7 to 9

oldest fully expanded rosette leaves were harvested at various time points for analyses.

181

The tocopherol content in C01 started to increase after 3 days of 7.5°C treatment reaching
levels five-fold higher than initial levels by 28 days, while vte2-1 lacked tocopherols at
all time points (Figure A.4A).

Consistent with the purple color of mature leaves of 7.5°C-treated vte2 mutants
(Figure A.3B), vte2-I accumulated significantly higher level of anthocyanins than Col
after 14 days of 7.5°C (Figure A.4D). In Col anthocyanins were detected only at 7 days.
Because anthocyanin accumulation is often associated with plant responses to stress
(Leyva et al., 1995; Chalker-Scott, 1999) and tocopherols are well-characterized lipid-
soluble antioxidants in animals (Ham and Liebler, 1995, 1997), it seemed plausible that
elevated lipid peroxidation might be occurring in vte2-1 during low temperature
treatment. However, the lipid peroxide levels of vte2-1 and Col analyzed by the ferrous
oxidation xylenol orange (FOX) assay were found to be similar and near background
levels at all time points (Figure A.4B), indicating that the observed phenotypic
differences between 7.5°C-treated vte2-I and Col are not associated with a detectable
increase in lipid peroxidation.

Given the reported localization of tocopherols and tocopherol biosynthetic enzymes
to plastids (Bucke, 1968; S011 et al., 1980c; Lichtenthaler et al., 1981; 8011 et al., 1985;
S011, 1987), it seems reasonable to hypothesize that tocopherol deficiency might affect
the components and function of the photosynthetic apparatus during 7.5°C treatment.
Under permissive growth conditions, the levels of individual and total photosynthetic
pigments (chlorophylls and carotenoids) were nearly identical in vte2-1 and Col (Figure
A 4C and Figure A 4E, Table A.5 at 0 day). The chlorophyll and carotenoid content of

both vte2-I and Col changed in parallel during the first two weeks of 7.5°C treatment and

182

became significantly different only at 28 days (Figure A.4C and Figure A.4E, Table A.5).
It is especially noteworthy that zeaxanthin, a xanthophyll cycle carotenoid that
accumulates under HL stress (Muller et al., 2001), was not detectable at any time point in
7.5°C-treated C01 and vte2-I (Table A.5), suggesting that the plants were not
experiencing photooxidative stress under the low temperature conditions used.

To assess the response of the photosynthetic apparatus to 7.5°C, changes in
photosynthetic parameters were analyzed. Fv/Fm was unchanged in both C01 and vte2-1

at any time point (Figure A.5A), indicating that photoinhibition is not occurring in either
genotype during permissive or 7.5°C conditions. The quantum yield of PSH ((Dpsu) was

also identical between C01 and vte2-1 under permissive growth conditions (Figure A.5B
at 0 day), suggesting that tocopherol deficiency also does not affect the efficiency of

electron transport via PSII in the absence of stress (Genty et al., 1989). During the first 7
days of 7.5°C treatment, 0)ng responded identically in C01 and vte2-1: (bpgn-decreased
sharply during the first day followed by a gradual recovery by 7 days. However, at 14

days the vte2-1 (DPSII was significantly lower than C01 and declined further by 28 days,

while the (DPSII of Col remained stable from day 14 and onward (Figure A.5B).

T ocopherol-Deficient Mutants Accumulate Carbohydrates During Low Temperature

Treatment

The reduced (bpsn in vte2-1 after 14 days could result from feedback inhibition of

photosynthesis due to the accumulation of downstream carbon metabolites (Goldschmidt

and Huber, 1992; Koch, 1996; Paul and Foyer, 2001; Paul and Peliny, 2003). To assess

183

this possibility, starch, glucose, fructose and sucrose contents were analyzed during the
time course of 7.5°C treatment. Starch represents the main plastidic carbohydrate storage
pool, sucrose and fructose are cytosolic pools, while glucose is present in both subcellular
compartments. C01 and vte2-1 had identical carbohydrate contents at the end of the light
period under permissive growth conditions (Figure A.6 at 0 day). During the first 7 days
of 7.5°C treatment starch content increased similarly in both C01 and vte2-I to
approximately 120 pmol glucose equivalents/ g FW. After 7 days vte2-1 starch content
steadily increased to 680 umol glucose equivalents/g FW while Col starch levels
decreased to near initial levels (Figure A.6A). Likewise, the glucose, fructose and sucrose
content of C01 and vte2-I increased similarly during the first 3 days of low temperature
treatment (Figure A.6B, Figure A.6C and Figure A.6D), likely as a component of the
well-documented cold acclimation response(s) in Arabidopsis (Wanner and Junttila,
1999; Taji et al., 2002). After 3 days, Col soluble sugar levels decreased, while vte2-I
continued to rise reaching 35, 43 and 255 times the initial levels of glucose, fructose and
sucrose, respectively, after 28 days of low temperature treatment. The timing of the

increase and accumulation of carbohydrates in vte2-1 is consistent with this being the

root cause of the reduction in (bpgu observed after 14 days at 7.5°C (Figure A.SB).

To further investigate any differences in carbohydrate accumulation between vte2-1
and Col during the initial 5 days of low temperature treatment, diurnal changes in
carbohydrate content were analyzed one hour before the end of the light and dark cycles.
During the 25 h prior to low temperature treatment (Figure A.7; -25 h, —13 h and -1 h,
with O h being the transfer of plants to low temperature at the start of the light cycle),

starch, glucose, fructose and sucrose content were identical in vte2-1 and Col. These data

184

indicate the lack of tocopherols does not have a significant impact on carbohydrate
metabolism under permissive growth conditions. Following transfer to low temperature
the soluble sugar content increased similarly in vte2-I and Col for the first two diurnal
cycles with significant differences first being observed between genotypes at the end of
the third low temperature light period (59 h in Figure A.7B, Figure A.7C and Figure
A.7D). In contrast, starch levels did not become significantly different between genotypes
until 14 days of low temperature treatment (Figure A.6A and Figure A.7A). The
differential elevation of soluble sugars prior to starch accumulation in vte2-1 indicates
that the increase in cytosolic soluble sugars precedes starch accumulation in the
chloroplast and that soluble sugars are not being efficiently metabolized or mobilized in

7.5°C-treated vte2-l .

The vte2 and vte1 Cold Sensitive Phenotypes Are Attenuated in Young Leaves
th th th th
Mature (7 to 9 oldest) and young (13 to 16 oldest) leaves of vte2-1 and vte1-1

mutants showed obvious visible differences in their responses to low temperature; young
leaves of vte2-1 and vte1-I did not change their color to purple even after two months of
7.5°C treatment (Figure A.3C).—Consistent with this visual observations, mature leaves of
vte1-I had an anthocyanin content 10 % that of vte2-I but still higher than Col, while
young vte2-I and vte1-1 leaves accumulated much less anthocyanins compared to their
respective mature leaves after 28 days of 7.5°C treatment (Figure A.8A). Fv/Fm was

above 0.8 in all cases, indicating that photoinhibition was not occurring in either young or

mature leaves of any genotype (data not shown). The (bpsn of mature vte2-1 leaves was

reduced to 70 % of mature Col leaves, consistent with Figure A.5, while the (1313311 of

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mature vte1-1 leaves was only slightly decreased relative to mature Col leaves. However,
the (bpsu of mature and young Col leaves and young vte2-I and vte1-I leaves were not

significantly different (Figure A.88). Levels of all carbohydrates in mature vte2-1 leaves
were greatly elevated in comparison to C01, consistent with Figure A.6. Mature vte1-I
leaves contained intermediate levels of starch, glucose, fructose and sucrose (51, 53, 68
and 58 % of vte2-1 levels, respectively) (Figure A.8C to Figure A.8F). Young vte2-1 and
vte1-1 leaves contained substantially reduced starch, glucose, and sucrose levels
compared to their respective mature leaves. These results indicate that the initiation and
development of young vte2-1 and vte1-1 leaves under 7.5°C conditions attenuates the
biochemical phenotypes observed in mature leaves of both genotypes and that the
DMPBQ accumulated in vte1-1 further suppresses these biochemical phenotypes in both

mature and young leaves.

Tocopherol-Deficient Mutants Have Reduced Source Leaf Photoassimilate Export

Capacity at Low Temperatures

The reduced seed yield (Table A.3) and attenuated carbohydrate accumulation in
young leaves relative to mature leaves (Figure A8) in 7.5°C-treated vte2-1 suggested

impaired translocation of photoassimilates from mature source tissues to young sink

tissues. To test this possibility 14C02 labeling experiments were conducted. C01 and

vte2-1 were grown on plates under permissive conditions for three weeks and then

transferred to 7.5°C for an additional 7 days. Whole plants were labeled with 14C02 at

7.5°C, transferred to high humidity in darkness at 7.5°C for 2 h to allow for

photoassimilate transport and subsequently exposed to a phosphor screen to visualize the

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movement of 14C labeled photoassimilate. Immediately after labeling >99% of the 14C02
incorporated was present in leaf tissue (data not shown). C01 and vte2-1 incorporated

. . 14 . . . .
Similar amounts of C02 into photosynthate suggesting therr carbon fixation rates do not
differ, consistent with the similar (Dpsu within the first 7 days at 7.5°C (data not shown

and Figure A.SB). Following the 2 h dark period Col had translocated 13.2 % of the 14C

labeled photoassimilate fixed in leaves to roots, whereas only 2.7 % was translocated in
vte2-1 (Figure A 9A). These results demonstrate that vte2-1 translocates significantly less
photoassimilate from source to sink than Col after 7 days of 7.5°C-treatment.

Impaired photoassimilate translocation in 7.5°C-treated vte2-1 could be due to
reduced sink strength or impaired photoassimilate export from source leaves (Gottwald
2000, Stitt 1996, Herbers and Sonnewald 1998). To address these possibilities, phloem
exudation experiments were conducted (King and Zeevaart 1974). C01 and vte2-1 were

grown for four weeks at permissive conditions and transferred to 7.5°C for an additional

0, 1, 3, or 7 days. Mature (7th to 9th oldest) leaves were excised from plants and labeled

with 14C02. The petioles of labeled leaves were then transferred to an EDTA solution to
induce phloem exudation and radioactivity in the EDTA solution was determined at
various time points (King and Zeevart, 1974). Again, total 14CO; fixed in mature leaves

were similar in all genotypes at each time point (Figure A.9C). Prior to 7.5°C treatment

(0 day), Col, vte1-l and vte2-l leaves exuded similar amounts of labeled

photoassimilates, accounting for approximately 34 % of the total l4C02 fixed in each

genotype. During 7.5°C treatment, the percent exudation by Col slightly decreased after 3

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and 7 days (to 27 and 31 % of the total l4C02 fixed, respectively), whereas that of vte2-I

was greatly reduced (to 11% and 4% at 3 and 7 days, respectively). Even more
intriguingly, exudation in vte2-1 was significantly lower than Col during the first day of

7.5°C treatment, which corresponds to only 6 h at 7.5°C treatment. The vte1-I mutant

exuded l7 and 15 % of the total 14COZ fixed after 3 and 7 days at 7.5°C, respectively,

levels intermediate between C01 and vte2-1 (Figure A.9C).

In apoplastic loaders like Arabidopsis sucrose is almost the exclusive translocated
photoassimilate (Vanbel, 1993). To assess the chemical nature of the labeled compounds
exuded from C01 and vte2-I, phloem exudates were collected and separated by anion-
exchange chromatography together with sugar standards. As shown in Figure A.9B,
approximately 85 % of the label in C01 and vte2-1 exudates comigrated with the sucrose
standard and 10 % with glucose/fructose standards. The high proportion of sucrose
indicates that the label collected is almost entirely from phloem exudate rather than

sugars from the cytosol of damaged cells.

Overall, the results obtained from 14C02 labeling experiments indicate that
tocopherol deficiency in both vte1-1 and vte2-1 results in dramatically reduced capacity
of photoassimilate export from source leaves in response to 7.5°C treatment. The rapidity
of the reduction in photoassimilate export in 7.5°C-treated vte2-I strongly suggests that
impairment of photoassimilate export is the root cause of the sugar accumulation
phenotype observed in mature leaves of 7.5°C-treated tocopherol-deficient mutants.
Structural Changes in Low Temperature- Treated T ocopherol-Deficient Mutants

Previously, callose was reported to accumulate at the bundle sheath/vascular

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parenchyma interface of the maize sxdl mutant and in vascular tissue of potato VT E1 -
RNAi lines, both of which are defective in tocopherol cyclase (Botha et al., 2000; Hofius
etal., 2004). To determine whether callose deposition also occurs in C01, vte2-1 and vte1-
], leaves were harvested at 0, l, 3 and 13 days of 7.5°C treatment and aniline blue-
positive fluorescence assessed. Under permissive conditions, aniline blue-positive
fluorescence was absent or sporadic and no significant differences were observed in any
genotypes. Aniline blue-positive fluorescence was also not altered in C01 during the
entire 7.5°C treatment period (e.g., 13 days at 7.5°C, Figure A.10C). In contrast, aniline
blue-positive fluorescence strongly increased in the vascular tissue of 7.5°C-treated vte2-
] (Figure A.10), and to a slightly lesser extent vte1-I (Figure A.l6). In both vte2-1 and
vte1-1, fluorescence initially appeared in a limited number of vascular cells in the petiole
as early as 6 h after transfer to 7.5°C conditions (Figures A.lOD and Figures A.10F,
Figures A.16A and Figures A.16B). The number of aniline blue fluorescing cells in the
vasculature and their fluorescent intensity subsequently increased in an acropetal fashion
in both vte2-1 and vte1-I during the course of 7.5°C treatment (Figure A.10, Figure
A.l6). Intriguingly, the induction, intensity and acropetal spread of vasculature-specific

aniline blue positive fluorescence in vte2-1 at 7.5°C was unaffected by light levels
ranging from 1 to 800 umol photon m-2 s.1 (Figure A.1 1A to Figure A.l 1F). Aniline blue
positive fluorescence was not observed in the vasculature of Col at any light level at
7.5°C (data not shown) and was also absent from the vasculature of both C01 and vte2-I
subjected to HL1800 at 22°C for up to 4 days (Figure A.1 1G and Figure A.l 1H and data

not shown).

To confirm whether or not aniline blue-positive fluorescence was cell specific and

189

could be attributed to callose deposition, serial sections of 0 and 14 day 7.5°C-treated C01
and vte2-I vascular tissue were examined at the level of the transmission electron
microscope (TEM). The spatial organization of cells and types of cells comprising the
phloem and xylem of both C01 and vte2-I were identical to what has been previously
described for Arabidopsis (Haritatos et al., 2000, data not shown). Notably, at day 0
phloem vascular parenchyma cells of both C01 and vte2-1 contained transfer cell wall
ingrowths adjacent to sieve elements and companion cells (e.g. Figure A.12A). 7.5°C-
treatment of Col for 14 days did not result in obvious ultrastructural changes in any
vascular cell type except for a noticeable increase in phloem parenchyma transfer cell
differentiation and transfer cell wall deposition exclusively adjacent to sieve elements
and companion cells in all vascular tissue (Figure A.IZB) although to a lesser degree in
the midvein.

During the same time course of 7.5°C-treatment in vte2-1, changes in cell fine
structure occurred exclusively within the phloem parenchyma transfer cells of all
vascular traces. Phloem parenchyma transfer cells in 14 day-treated vte2-I exhibited
irregularly thickened cell wall depositions with ultrastructural features characteristic of
callose (Nishimura et al., 2003); Figure A.12C to Figure A.12F). Large callosic-like
masses that dissected the cell lumen corresponded in shape to aniline blue-positive
fluorescent regions (compare Figure A.12C and Figure A.12E to Figures A.10F, Figures
A.lOI and Figures A.lOL). The callosic-like wall material also formed a sheath around
the cells (Figure A.12D), was deposited over transfer cell wall ingrowths (Figure A.12F)
and between the end walls of adjoining transfer cells including plasmodesmata (data not

shown). Immunolocalization using monoclonal antibodies against callose confirmed the

190

presence of callose at each location (Figure A.12G to Figure A.121) and at
plasmodesmata between the phloem parenchyma transfer cells and bundle sheath (Figure
A.12K). No immunolabelling was present in controls using secondary antibody only
(Figure A.12D to Figure A.12F) and immunolabelling was rare to absent in all cell types
of untreated Col and vte2-I and in 14-day 7.5°C-treated Col, including phloem
parenchyma transfer cells (e.g. Figure A.12L).

Serial sections of vascular tissue from C01 and vte2-1 treated at 7.5°C for 3 and 7
days were subsequently examined at the level of the TEM to determine the spatial and
temporal development of callose deposition within phloem parenchyma cells. At 3 days,
phloem parenchyma transfer cell wall deposition in C01 was confined to the sieve
element or companion cell boundary (data not shown) but in vte2-1 wall deposition was
present around the entire transfer cell periphery (Figure A.13A). Cell wall deposition in
3 day-treated vte2-1 resulted in abnormally thickened and irregular shaped ingrowths
with callose-like depositions adjacent to sieve elements and companion cells (Figure
A.13A) that grew increasingly prominent by day 7 (data not shown). In 3-day 7.5°C-
treated vte2-1 positive immunolocalization with monoclonal antibodies to callose was
present exclusively at the phloem parenchyma transfer cell wall-sieve element boundary
(Figure A.13B) and included phloem parenchyma transfer cell-sieve element
plasmodesmata] connections (Figure A.13C). In contrast to 14 days, plasmodesmata
between bundle sheath and phloem vascular parenchyma cells in 3 day 7.5°C-treated
vte2-1 were continuous and immunonegative for callose (compare Figure A. 12K and
Figure A.13D).

DISCUSSION

l9l

The chemistry of tocopherols as lipid soluble antioxidants and terminators of PUFA
free radical chain reactions has been well established from analyses in artificial
membranes and animal-derived membrane systems (Liebler and Burr, 1992; Ham and
Liebler, 1995). It has long been assumed that similar chemistry occurs in the tocopherol-
containing PUFA-enriched membranes of photosynthetic organisms, and this assumption
has recently been supported by studies of tocopherol deficient photosynthetic organisms
(Sattler et al., 2004; Havaux et al., 2005; Maeda et al., 2005). During the first several
days of germination, a period of high oxidative metabolism, Arabidopsis vte2 seedlings
contain levels of oxidized lipids >100-fold higher than wild type or vte1, which were
indistinguishable (Sattler et al., 2004 and Sattler et a1 2006). Thus, a chemical role for
tocopherols (or DMPBQ in vte1) in limiting lipid peroxidation appears to be conserved
between photosynthetic organisms and animals. By 18 days of growth the lipid peroxide
levels of vte2 seedlings had decreased to near that of wild type and vte1 (Sattler et al.,
2004) and_became indistinguishable from wild type and vte1 after four weeks of grth
(Figure A.4B), consistent with tocopherols being dispensable in mature photosynthetic
tissues in the absence of stress (Porfirova et al., 2002; Bergmuller et al., 2003; Sattler et
al., 2003a; Sattler et al., 2004).

While a chemical role for tocopherols in controlling lipid peroxidation at specific
points of the plant life cycle now seems clear, the physiological roles of tocopherols
during plant stresses do not. In the current study, we have utilized a suite of Arabidopsis
vte mutants that accumulate different types and levels of tocopherols and pathway
intermediates (Table A.1) to directly assess tocopherol specific functions in

photosynthetic tissues in planta in response to abiotic stress treatments.

192

A Limited Role for T ocopherols in Protecting Arabidopsis Plants from High Intensity

Light Stress.

Tocopherols have long been assumed to play crucial roles in HL protection
presumably by acting as singlet oxygen quenchers and lipid peroxy radical scavengers
(Fryer, 1992; Munne-Bosch and Alegre, 2002; Trebst et al., 2002). In the current study
the biochemical and photosynthetic responses of the tocopherol-deficient vte2 mutant
exposed to HLIOOO and HL1800 at 22°C for up to eleven days was surprisingly similar to
wild type (Figure A.2, Figures A.l4 and Figures A.15). Likewise, the mutation
corresponding to Arabidopsis vte2 in the cyanobacterium Synechocystis sp. PCC6803
(slr1736) had a similarly limited impact on growth and photosynthesis under permissive
growth conditions and during HL stress (Collakova and DellaPenna, 2001; Maeda et al.,
2005). In both organisms it is only when HL stress has been combined with other
stresses, in combination with lipid peroxidation inducing chemicals in the Synechocystis
slrI 736 mutant (Maeda et al., 2005) or in combination with low temperatures (2-3°C) in
Arabidopsis vte2 and vte1 mutants that differential impacts on photosynthetic parameters
or lipid oxidation are observed (Havaux et al., 2005). These combined data indicate that
tocopherols are not essential for adaptation and tolerance of photosynthetic tissues
subjected to BL stress alone. Such a conclusion runs counter to long-held presumptions
that a primary function of tocopherols is to protect photosynthetic tissues against HL
stress (Fryer, 1992; Munne-Bosch and Alegre, 2002).

One possible explanation for this surprisingly limited role of tocopherols during HL
stress is that other mechanisms compensate for their absence. The zeaxanthin level of

HL1800 vte2 was nearly twice that of Col (Table A2). vte2 (and vte1) also had a higher

193

xanthophyll de-epoxidation state (A+Z/A+Z+V) after HL1800 treatment (Table A2), as
did vte1 during HL stress combined with low or high temperatures (Havaux et al., 2005).
Growth of a tocopherol-deficient Synechocystis mutant was also much more susceptible
than wild type to treatment with a biosynthetic inhibitor of carotenoid synthesis during
HL stress (Maeda et al., 2005). Similarly, a double mutant of vte1 and nqu (non-
photochemical quenching I), which cannot accumulate zeaxanthin in response to HL and
hence cannot induce non-photochemical quenching (Niyogi et al., 1998), was reported
more susceptible than either single mutant to the combination of HL and low temperature
stress (Havaux et al., 2005). Conversely, the young nqu leaves were also tolerant to
short and long term HL stress (up to HL1800) and accumulated higher level of
tocopherols than wild type (Havaux et al., 2000; Golan et al., 2006). These data suggest
that tocopherols and carotenoids, particularly zeaxanthin, have overlapping functions in
protecting photosynthetic organisms against HL stress.

In prior studies it was demonstrated that, although vte1 and vte2 are both tocopherol
deficient, the two genotypes behaved quite differently during early seedling development:
vte2 exhibited a >100 fold increase in non-enzymatic lipid peroxidation during
germination, whereas lipid peroxidation in vte1 was identical to wild type (Sattler et al.,
2004; 2006). In the current study vte] was again found to behave differently from vte2.
In response to HL1800 stress vte1 had a slightly, but reproducibly, higher degree of
photoinhibition and higher level of photobleaching than either vte2 or wild type (Figure
A.2, Table A2 and Figure A.lS), suggesting the vte1 mutation negatively impacts HL
stress tolerance beyond its tocopherol deficiency. Why would vte1 respond so differently

from vte2 during germination and HL1800 given that both mutant genotypes are

194

tocopherol deficient? The most likely explanation resides in the singularly unique
biochemical feature of vte1: it accumulates the redox active quinol biosynthetic
intermediate DMPBQ in place of tocopherols. DMPBQ is absent from vte2 and Col
(Table A.l and Sattler et al., 2003) and its presence in vte1 can clearly have significant,
unintended experimental consequences that are independent of the tocopherol deficiency
in vte1. Thus, when attempting to define tocopherol functions based on vte mutant
phenotypes one must be careful to delineate genuine tocopherol functions, which would
occur in both vte1 and vte2, from potentially confounding artifacts due to the presence of
DMPBQ, which would only occur in vte1. Such DMPBQ-dependent artifacts can have
negative (HL1800), positive (seedling germination) or partially positive (low temperature
adaptation) consequences depending on the treatment condition and phenotype assessed.

These concerns are not relevant for vte2.

Arabidopsis T 0copherol-Deficient Mutants Exhibit a Cold Sensitive Phenotype

Independent of Photooxidative Damage

In contrast to the equivocal results of HL, salinity and drought stress treatments with
tocopherol-deficient photosynthetic organisms (Figure A.2, Figure A. 14 and Figure A.15,
(Porfirova et al., 2002; Havaux et al., 2005; Maeda et al., 2005)), both tocopherol-
deficient vte1 and vte2 genotypes were found susceptible to non-freezing low temperature
treatments in comparison to their respective wild types (Figure A.3). These results clearly
indicate that tocopherols play a critical role in the responses of mature Arabidopsis plants
to non-freezing low temperatures. Given the well-defmed role of tocopherols as lipid
soluble antioxidants, we initially hypothesized that tocopherol-deficiency at low

temperature would result in increased photooxidative damage relative to wild type and

195

that this might lead to the low temperature sensitive phenotype observed in vte2.
However, the hallmarks of photooxidative stress and photoinhibition: decreased Fv/Fm
and chlorophyll levels and increased zeaxanthin accumulation and lipid peroxidation
(Havaux and Niyogi, 1999; Maxwell and Johnson, 2000; Broin and Rey, 2003) were not
observed during the first two weeks of low temperature treatment (Figure A.4B, Figure

A.4C, Figure A.5A, and Table A.5), the timeframe during which the vte2 carbohydrate

accumulation phenotype firlly develops (Figure A.6 and Figure A.7). Likewise, (DPSIIs

though altered by low temperature, was identical between wild type and vte2 during the
first week of low temperature treatment (Figure A.5B). These data indicate that the
tocopherol-deficiency has no discemable impact on photosynthesis under the low
temperature conditions used and that the low temperature sensitive phenotype of vte2 is
not associated with increased photooxidative damage or photoinhibition due to the

absence of tocopherols.

Tocopherols Are Required for Photoassimilate Export from Source Leaves During Low

Temperature Adaptation

A well-documented response of plants to low temperatures is the accumulation of
soluble sugars and other osmoprotectants, which are critical components for the process
of cold acclimation leading to freezing tolerance (Wanner and Junttila, 1999; Gilrnour et
al., 2000). The subsequent recovery of photosynthesis and sucrose metabolism is an
important component of low temperature adaptation in that it provides carbon to sustain
growth under low temperatures (Strand et al., 1997; Strand et al., 1999). During the first
two days of low temperature treatment, soluble sugar levels increased similarly in both

vte2 and wild type (Figure A.7), suggesting tocopherols have little impact on the initial

196

accumulation of soluble sugars in response to low temperature. ”However, the
accumulation of sucrose and other soluble sugars was much higher in vte2 than wild type
after 60 h of low temperature treatment (Figure A.7B - Figure A.7D), although the rates
of photosynthesis and carbon fixation were indistinguishable between the two genotypes
until 14 days at low temperature (Figure A.4B and Figure A.9C). vte2 also reduced
soluble sugar levels more slowly at night than wild type after 3 days of low temperature
treatment (Figure A.7B- Figure A.7D). These results suggest that tocopherol deficiency
affects carbohydrate utilization/mobilization rather than the supply of fixed carbon from

photosynthesis during low temperature adaptation.

l4C02-labeling experiments demonstrated that in comparison to wild type, low

temperature-treated vte2 translocated significantly less l4C-labeled photoassimilates from

leaves (source tissue) to roots (sink tissue, Figure A.9A). The long distance transport of
photoassimilates occurs through phloem and the transport rate is determined either by the
rate of export from source leaves to phloem (loading) or by removal into sink tissues
(unloading) (V anbel, 1993; Stitt, 1996; Herbers and Sonnewald, 1998). Phloem exudation

experiments with excised leaves showed that vte2 source leaves exported significantly
less l4C labeled photoassimilates than wild type as early as 6 h following transfer to low

temperature (Figure A.9C). The rapidity of this reduction in photoassimilate export in
comparison to the elevated sugar accumulation in vte2 starting at 60 h (compare Figure
A.9 and Figure A.7) indicates that impaired photoassimilate export is an early, upstream
event in the vte2 low temperature phenotype and the likely root cause of the elevated
sugar accumulation in low temperature-treated vte2. Taken together, these analyses

demonstrate that tocopherols are required for proper regulation of photoassimilate export

197

from source leaves and thereby play a critical role in low temperature adaptation in
Arabidopsis.

Previous studies of the maize sde mutant and potato VI‘EI-RNAi lines (both
affecting tocopherol cyclase activity) had suggested a linkage between carbohydrate
metabolism and tocopherol biosynthesis, as in both cases carbohydrates accumulated to
high levels in mature leaves at normal growth temperatures (19 to 30°C, (Russin et al.,
1996; Provencher et al., 2001; Hofius et al., 2004). The absence of this phenotype in
Arabidopsis vte1 and vte2 at 22°C raised questions of the universality of any interaction
between tocopherol synthesis and carbohydrate metabolism (Sattler et al., 2003a), Figure
A.6 and Figure A.7 at 0 day). We now know that tocopherol-deficient Arabidopsis
mutants do indeed exhibit a phenotype that is analogous to sxdl but which is inducible
only at low temperatures. Thus, the linkage between tocopherol biosynthesis and
carbohydrate metabolism is conserved among all tocopherol-deficient mutants identified
in higher plants to date (maize sxd], potato VTEI-RNAi and low temperature-treated
Arabidopsis vte1 and vte2).

Although the maize sde mutant and potato V T E1 -RNAi line suggested tocopherol
chromanol ring cyclization was somehow related to regulation of carbohydrate
metabolism, it was unclear whether the phenotype was due to the lack of tocopherols or
accumulation of the redox active quinol intermediate DMPBQ (Sattler et al., 2003a;
Hofius et al., 2004). Analysis of the full suite of Arabidopsis vte mutants now allows a
conclusive answer to this question. Given that vte2 lacks DMPBQ (Table A.1) and
exhibits a more severe carbohydrate accumulation phenotype than vte1 (Figure A.8), we

can conclude that it is the absence of tocopherols rather than accumulation of DMPBQ

198

that causes the carbohydrate accumulation phenotype. The reduced severity of the
carbohydrate accumulation phenotype in vte1 suggests that DMPBQ partially suppresses

the low temperature-inducible vte2 carbohydrate accumulation phenotype.

T ocopherol Deficiency Results in a Cell Specific Response by Phloem Parenchyma

Transfer Cells at Low Temperature

The carbohydrate accumulation phenotype of maize sde was reported to be
associated with altered structural features within vascular tissue. Plasmodesmata at the
sde bundle sheath/vascular parenchyma boundary were reported occluded by wall
materials (Russin et al., 1996; Provencher et al., 2001) and subsequently suggested to
correspond to aniline blue-positive fluorescence (Botha et al., 2000). This structural
aberration in sde plasmodesmata was posited to be the basis of the sde carbohydrate
accumulation phenotype because it would lead to a block in the symplastic movement of
photoassimilate. Callose was also observed in vascular tissue of potato VTEI-RNAi
plants by light microscopy with monoclonal antibodies against [3-1,3 glucan (Hofius et
al., 2004). In the absence of high-resolution microscopy, Hofius et a1. (2004) also
suggested this vascular-associated callose somehow interrupts photoassimilate transport.
However, in both the sde and potato VTEI-RNAi studies it was impossible to determine
whether callose deposition was a cause or effect of carbohydrate accumulation. A critical
observation from the present study is that the low temperature-inducible photoassimilate
export defect in Arabidopsis tocopherol-deficient mutants is temporally associated with
callose deposition in a specific vascular tissue cell type (compare Figure A.9C and Figure
A.lO). These results are significant as they provide a direct link between defective

photoassimilate export and callose deposition (or events tightly associated with callose

199

deposition) in tocopherol-deficient mutants and exclude the possibility that callose
deposition is a secondary effect caused by carbohydrate accumulation.

The low temperature-inducible callose deposition in Arabidopsis vte2 selectively
occurred in phloem parenchyma transfer cells (Figure A.12 and Figure A.13).
Importantly, initial callose deposition was site specific within these cells and resulted in a
callose boundary between the phloem parenchyma transfer cell and sieve
element/companion cell complex where transfer cell wall ingrowths occur (Figure A.13).
We saw no evidence of callose deposition or occlusion of plasmodesmata at the bundle
sheath-vascular parenchyma boundary during induction of the export defective phenotype
in vte2 (e.g., 3 days of low temperature treatment, Figure A.13D). However, by 14 days
of low temperature treatment, when vte2 contains high levels of starch and anthocyanins
(Figure A.4D and Figure A.6A) and more closely resembles the phenotype of maize sxd],
the entire parenchyma transfer cell became encased in a callose sheath associated with
abnormally shaped transfer cell wall ingrowths and it is at this point that callose
deposition is also observed in vte2 plasmodesmata at the bundle sheath-vascular
parenchyma boundary (Figure A.12K). When one compares the development, polarity
and morphology of transfer cell walls in 7.5°C-treated vte2 with C01 (e.g. compare Figure
A.12G and Figure A.12L), it becomes clear that tocopherols play an important role in
transfer cell wall synthesis at low temperatures.

Results from previous structural studies on the minor vein structure of Arabidopsis
have suggested that phloem parenchyma transfer cells are the site of apoplastic unloading
0f photoassimilates arriving symplastically from bundle sheath cells (Haritatos et al.,

2000). The coincidence in reduction of photoassimilate export with callose deposition in

200

these spatially distinct subcellular sites in the vte2 mutant during low temperature
treatment (Figure A.9, Figure A. 10 and Figure A.l3) provides direct support for the role
of transfer cells in photoassimilate export from source leaves via delivery to the phloem
apoplast. This callose deposition (or events associated with the callose deposition) in
phloem parenchyma transfer cells of 7.5°C-treated vte2 would form a barrier to symplast-
to-apoplast but not symplast-to-symplast transport. The limited export that still occurs in
7.5°C-treated vte2 source leaves (Figure A.9) may be due to apoplastic unloading from
bundle sheath cells and subsequent loading to the sieve element/companion cell complex
(Haritatos et al., 2000). The special characteristics of phloem parenchyma transfer cells
that lead them, in comparison to other cell types in the leaf, to be so specifically and
differentially impacted by tocopherol-deficiency during low temperature treatment

remains to be determined.

T ocopherol Functions in Plant Stress Physiology

In the current study the tocopherol-deficient vte2 mutant was found to be remarkably
similar to wild type in its response to most abiotic stresses with the notable exception of
non-freezing low temperature treatments. Tocopherol-deficiency specifically results in
abnormal phloem parenchyma transfer cell wall development at low temperature. This
leads to rapid impairment of photoassimilate export that profoundly impacts cellular
metabolism and whole plant physiology during both short and long term low temperature
treatments. That this occurs in both vte2 and vte1 strongly argues that tocopherols play a
crucial, previously unrecognized role in low temperature adaptation, specifically in
phloem loading. Several studies have suggested that vascular tissues, including vascular

parenchyma, are metabolically distinct, sensitive to changing environmental conditions

201

and hence critical sites for stress responses (Orozco-Cardenas et al., 2001; Hibberd and
Quick, 2002; Fryer et al., 2003; Koiwai et al., 2004; Narvaez-Vasquez and Ryan, 2004).
Our data are consistent with this thesis and suggest tocopherols have important
fimction(s) in regulating the response of these specific cell types to environmental stress,
such as low temperatures.

Our findings that photooxidative damage and photoinhibition are not associated with
the vte2 low temperature phenotype and that HL1800 (which approaches full sunlight) at
22°C has little impact on vte2 compared to wild type suggest a more limited role for
tocopherols in protecting plants from photooxidative stress than has been assumed. This
seems in direct contradiction with a recent report using Arabidopsis vte mutants that
concluded tocopherols protect Arabidopsis against photoinhibition and photooxidative
stress (Havaux et al., 2005). However, the conclusions of this prior work were based

entirely on the differential responses of wild type and vte mutants exposed to low

temperatures (2-8°C) in combination with HL (1000 to 1600 umol photon m-2 s-1) for

durations of up to seven days. We now know that such low temperature treatments would
rapidly block photoassimilate export in tocopherol-deficient genotypes, but not in wild
type, independent of any light regime imposed (Figure A.9 to Figure A.l 1) and likely
confound any interpretations with respect to previously proposed HL-specific tocopherol
functions. Thus, the photoprotective functions of tocopherols in plants remain an open

question and a critical reassessment is needed to clarify this issue.

202

METHODS

Growth Conditions and HL and Low Temperature Treatment

Seeds were stratified for four to seven days (4°C), planted in a vermiculite and soil

mixture fertilized with 1 x Hoagland solution, and grown in a chamber under permissive

conditions: 12h, 120 umol photon m-2 5.1 light at 22°C / 12h darkness at 18°C with 70 %

relative humidity. Plants were watered every other day and with 0.5 x Hoagland solution

once a week. For HL treatments, four-week-old plants were transferred in the middle of

the light cycle to 1800 umol photon m-2 3'1 16h light / 8h darkness at 22°C. For low

temperature treatments, three to four-week-old plants were transferred at the beginning of

light cycle to 12h 75 umol photon m-2 5'1 light / 12h darkness at 7.5°C (d:< 3°C).

Tocopherol, Anthocyanin, Chlorophyll and Carotenoid Analyses

Leaf samples ( 12-15 mg) were harvested directly into liquid, nitrogen at the end of
the light cycle and lipids extracted in the presence of 0.01 % (w/v) butylated
hydroxytoluene (BHT) using tocol as an internal standard as described (Collakova and
DellaPenna, 2001). After phase separation, the aqueous phase was transferred to a new
tube, acidified by adding an equal volume of 1N HCl and anthocyanin content measured
spectrophotometrically at 520 nm as described (Merzlyak and Chivkunova, 2000). The
lipid phase was used for reverse-phase HPLC analyses to identify and quantify each
tocopherol, chlorophyll and carotenoid species as described previously (Collakova and

DellaPenna, 2001; Tian and DellaPenna, 2001).

203

Lipid Peroxide Analysis

Lipid peroxide content was measured using the ferrous oxidation-xylenol orange
(FOX) assay as previously described (Sattler et al., 2004) with the following
modifications. Leaf samples (25-30 mg) harvested at the end of light cycle were
immediately extracted with 200 uL of methanol containing 0.01 % (w/v) BHT, 200 uL of
dichloromethane, and 50 uL of 150 mM acetic acid using three 3-mm glass beads and a
commercial paint shaker. After shaking for 4 min, 100 uL of water and 100 uL of
dichloromethane were added for phase separation. Half of the organic phase was
incubated with an equal volume of 50 mM triphenyl phosphine in methanol for 30 min to
reduce lipid peroxides and half was incubated with an equal volume of methanol for 30
min. The triphenyl phosphine-treated and untreated samples (100 uL) were incubated
with 900 pL of FOX reagent [90% (v/v) methanol, 4 mM BHT, 25 mM sulfuric acid, 250

uM ferrous ammonium sulfate, 100 uM xylenol orange] at room temperature for exactly

20 min and A560 was measured. Lipid peroxide content was calculated based on a
standard curve of hydrogen peroxide as previously described (DeLong et al., 2002).

Chlorophyll Fluorescence Measurements

In vivo chlorophyll a fluorescence was measured in the middle of the light cycle
using a pulse amplitude modulation (PAM) fluorometer FMSZ (PP Systems, Haverhill,
MA). Attached leaves were dark adapted for at least 15 minutes prior to measurements

and fluorescence parameters were determined according to (Maxwell and Johnson, 2000).

Quantum yield of PSII ((Dpsu) was calculated as (F’m-Ft)/F’m, where F’m and Ft are

maximum fluorescence and steady state fluorescence in the light, respectively.

204

Carbohydrate Analyses

Soluble sugar (glucose, fructose and sucrose) and starch levels of leaves were
quantified as described (Jones et al., 1977; Lin et al., 1988) with minor modifications.
Unshaded leaf tissue (<50mg) was harvested, immediately frozen in liquid nitrogen and
extracted twice with 700 uL of 80% ethanol at 80°C. The ethanol extract was evaporated
and redissolved in 200 uL of distilled water (Jones et al., 1977). For starch analysis, the
extracted leaf residue was ground in 200 pL 02N KOH and boiled at 95°C for 45 min.

After cooling, the sample was neutralized to pH 5 with 50 uL of 1N acetic acid,

centrifuged and 50 pL of supernatant mixed with 492.5 uL of 0.2 M sodium acetate (pH

4.8), 150 uL of H20, 4 uL of a-amylase (4 units) and 3.5 uL of amyloglucosidase (2

units) and incubated at 37°C overnight. Glucose, fructose and sucrose levels in the
soluble sugar extract and the glucose level of the digested starch extract were determined
enzymatically (Jones et al., 1977).

14C02 Labeling Experiments

For phloem exudation experiments, approximately 20 mature leaves (7th to 9th

oldest) were detached in the middle of the day, 0.5 cm of petiole was re-cut under water,

the petiole of each leaf was submerged in water and placed in a tightly sealed 10 L glass

chamber. 14C02 was generated in the chamber by adding 3 mL of 0.25 N H2SO4 to 0.1
mCi (7 umol) NaHl4CO3 and unlabeled 93 umol NaHCO3 to give a carbon dioxide

concentration of 522 ppm. After labeling for 30 min at 120 pmol photon m.2 5-1, the

petiole of each leaf was submerged in 0.45 mL of 20 mM disodium-

205

ethylenediaminetetraacetic acid (EDTA) (pH 7.0) and kept in the dark with high humidity
to induce phloem exudation (King and Zeevart 1974). All of the aforementioned
procedures were performed at 7.5°C for low temperature-treated leaf samples. Exudation
of radiolabel into the EDTA solution was then periodically measured over the course of
10 h by liquid scintillation counting (Tri-Carb 28OOTR; PerkinElmer, Wellesley, MA,
USA). After 10 h of exudation, radiolabel remaining in the leaves was determined by
liquid scintillation counting. Total radiolabel fixed per leaf was calculated by adding total
radiolabel exuded and remaining in each leaf.

For analyzing translocation of radiolabeled photoassimilates in whole plants, plants
were grown on 1/2 MS plates under permissive conditions for 3 weeks and transferred to

low temperature conditions for seven days with lids partially ajar to supply atmospheric
C02. Whole plants were labeled at 7.5°C for 40 min as described above, placed in

darkness at 7.5°C and high humidity for 2 h to allow for translocation. Roots were
excised from leaves and both exposed to a phosphor screen to visualize the location of
radioactivity (Storm; GE Healthcare, UK).

Carbohydrate analysis of phloem exudate was performed by high-pH anion exchange
chromatography (HPAEC). Excised leaves were treated as described for phloem
exudation experiments except at the end of the initial 2 h exudation period the petiole of
each leaf was transferred to 0.45 mL water for 4 h to collect exudates for analysis. The
water exudates were dried under vacuum, dissolved in 50 uL water and 20 uL of the
sample was mixed with 5 uL of standards (25 nmol glucose, 50 nmol fructose, 125 nmol
sucrose and 100 nmol raffinose) and injected onto the HPAEC. The mixtures were

separated on a CarboPac PA-lO column (DIONEX, Sunnyvale, CA, USA) using a 30 min

206

linear gradient of 20 to 140 mM NaOH with a flow rate of 1 mL min]. One mL fractions

were collected and radioactivity was determined by liquid scintillation counting, while

sugar standards were detected by pulsed amperometric detection.

Fluorescence and Transmission Electron Microscopy

Leaves were prepared for aniline blue fluorescence microscopy (n = 2 leaves/plant,
4-6 plants/sample time; (Martin, 1959) and transmission electron microscopy (TEM; n =
1 leaf/plant, 2-3 plants/sample time; (Sage and Williams, 1995) at the same sampling
times as above for export studies. The presence or absence of callose was determined
using immunolocalization at the level of the TEM as described by Lam et al. (2001) with
monoclonal antibodies to B-1, 3 glucan (Biosupplies, Australia). Primary and secondary
(anti-mouse IgG gold conjugate 18 nm, Jackson Immunoresearch, West Grove, PA,
USA) antibody dilutions were 1: 100 and 1:20 respectively. Incubation time in the 1°
and 2° antibodies were 2 and l h, respectively. Controls were run by omitting 1°
antibody. Images were captured on the Leica MZ 16F fluorescence microscope (Wetzlar,
Germany) and the Phillips 201 TEM equipped with an Advantage HR Camera System

(Advanced Microscopy Techniques Corp. Danvers, MA, USA).

Statistical Analysis

One-factor ANOVA was used for the data in Table A2 and Figure A.2 using
genotype as a factor. Two-factor ANOVA was used for the data in Figure A.9C using
days of cold treatment and genotype as factors. When significance was observed (P <
0.05), pair-wise comparison of least square means was evaluated. SAS software was used
for these analyses (SAS Institute, Cary, NC, USA). Student t test was used for the rest of

the data to compare statistical significance of mutants relative to C01 (P < 0.05) using

207

Microsoft Excel.

Accession Numbers

Sequence data for the Arabidopsis thaliana tocopherol biosynthetic enzymes
described in this article can be found in the GenBank nucleotide sequence database under
the following accession numbers: VTEI (At4g32770), NM119430; VTE2 (At2g18950),

NM179653; VTE4 (Atlg64970), NM105171.

ACKNOWLEDGMENTS

We thank Maria Magallanes-Lundback for isolation of the vte4—3 allele, Kathy Sault
for technical assistance with microscopy, Ashok Ragavendran at MSU CANR Biometry
Group for assistance with statistical analyses and Drs. Andreas Weber, Dr. Lars V011 and
members of the DellaPenna lab for their critical advice, discussions and manuscript
review. This work was supported by NSF grant MCB-023529 to D.D.P. and a Connaught

Award and NSERC of Canada Discovery Grant to T.L.S.

208

TABLES AND FIGURES
Table A.l Tocopherol and DMPBQ Content in Leaves of Wild Types and

Tocopherol Biosynthetic Mutants Grown Under Permissive Conditions.

 

 

 

6° B y 6 total DMPBQ
Col 15.6 13.0 0" 0.6 20.1 o 16.2 13.0 o
vte2-1° o o o o o
vte1-1° 0 o o o 19.6 11.3
vte1-2° o o o o 17.3 :18
Ws 12.3 1.2.2 o o o 12.3 :22 o
vte2-2° o o o o
vte4-3d o 17.7 :l:1.2 o 17.7 11.2 o

 

Plants were grown for four weeks under permissive conditions and mature leaves were
harvested for analysis. Data are means :1:SD (n = 3 or 4) and are expressed as pmol/mg

FW.

a or, B, y and 6 indicate on, [3, y and 6-tocopherol, respectively.
b 0 indicates the compound was below detection (typically _<_ 0.1 pmol/mg F W).
0

Col background.

d Ws background.

209

Table A.2 Content of Photosynthetic Pigments and Toc0pherols of C01 and the vte2

and vte1 Mutants After HL1800 Treatment at 22°C.

 

 

 

After HL1800

Col vteZ-t vte1-1 vte1-2

tocggfi'mls 1.58 10.25" 0 10" 0 1:0" 0 10"
chlo::‘:;ylls 17.28 11.01 16.60 11.37 16.51 1:1.84 16.21 12.05
Chla 12.25 10.87 11.69 11.04 11.61 11.38 11.52 11.58
Chlb 5.02 1:026 4.90 10.35 4.90 10.51 4.69 10.50
ChlaIChlb 2.44 10.16 2.38 10.09 2.37 10.13 2.45 10.14
“rlgtzgids 5.37 1035‘" 4.98 10.36" 4.85 10.54" 4.75 1:0.70"
B-car 0.66 10.08 0.64 10.07 0.63 1:009 0.64 10.11
Lutein 2.45 10.14" 2.39 10.15” 2.26 10.22"C 2.20 10.27c
N 0.57 10.03" 0.51 10.04" 0.51 :I:0.06b 0.50 10.07"
V 1.01 1:012a 0.57 10.100 0.78 1017" 0.77 10.22"
A 0.39 10043" 0.39 1004“ 0.36 10.04"c 0.36 10040
Z 0.28 10.05" 0.48 10.06" 0.30 :t0.06b 0.29 10.03"
A+Z+V 1.69 1013a 1.44 10.12" 1.44 1:0.18b 1.42 1:0.26b
A+ZIA+Z+V 0.40 10050 0.61 10.05" 0.46 1:007" 0.47 10.07"

 

Plants were grown for four weeks under permissive growth conditions (120 nmol photon

rrf2 s-]) and pigment contents were analyzed after 4 days of HL (1800 nmol photon m-2

s-l) treatment. Data are means iSD (ug/cmz, n = 19). When significance is observed

between genotypes (ANOVA, P < 0.05), pair-wise comparison of least square means is
evaluated and non-significant groups are indicated by a, b or c with a being the highest
group. N, neoxanthin; B-car, B-earotene; V, violaxanthin; A, antheraxanthin; Z,

zeaxanthin; Chlb, chlorophyll b; Chla, chlorophyll a.

210

Table A.3 Yields and Abortion Rates of Seed Produced During Low Temperature

 

 

(7.5°C) Treatment.
Col vte2-1 vte1-1
seeds I silique 31.1 :I:1.6 19.5 13.4“ 27.3 12.5”
aborted seeds I sillique 0.1 10.4 6.8 12.4" 2.01-10*
percentage abortion (%) 0.4 34.6“ 73*
yield (mg seeds I plant) 373 1:77 87 1:27“ 2238

 

Data are means iSD (n = 3 for yields, 11 = 7 for aborted seed). Student’s t tests relative to

C01 (*P < 0.05, **P < 0.01).

a An average of duplicate plants (225.7 and 219.4 mg seeds/plant)

211

Table A.4 Content of Photosynthetic Pigments and Tocopherols of C01 and the vteZ

and vte1 Mutants Grown at Permissive condition

 

 

 

Before HL1800

Col vte2-1 vte1-1 vte1-2

,ocgggg'm, 0.11 10.01" 0 10" 0 10" 0 1:0"
0111;31:1in1 21.90 11.14 20.51 11.70 20.15 11.12 21.26 11.19
Chla 15.26 10.77 14.25 11.18 14.02 10.79 14.78 10.84
Chlb 6.64 10.37 6.25 10.53 6.13 10.33 6.49 10.35
Chla/Chlb 2.30 10.03 2.28 10.02 2.29 10.01 2.28 10.02
“(121:3 d8 3.56 10.23 3.32 10.28 3.28 10.18 3.44 10.22

B-Car 0.61 10.05" 0.57 10.03" 0.55 10.03" 0.59 10.05""

Lutein 1.85 10.13 1.72 10.16 1.70 10.10 1.79 10.12
N 0.59 10.03 0.55 10.05 0.54 10.03 0.57 10.03
v 0.51 10.04 0.48 10.05 0.49 10.03 0.50 10.03

A 0 10 o 10 0 10 0 10

z 0 10 0 10 0 10 0 10
A+Z+V 0.51 10.04 0.48 10.05 0.49 10.03 0.50 10.03

A+Z/A+Z+V 0 1:0 0 1:0 0 1:0 0 1:0

 

Plants were grown for four weeks under permissive growth conditions (22°C, 120 nmol

photon m-2 s-l) and mature leaves were analyzed. Data are means iSD (pg/cmz, n = 7).
When significance is observed between genotypes (ANOVA, P < 0.05), pair-wise
comparison of least square means is evaluated and non-significant groups are indicated
by a or b with a being the highest group. N, neoxanthin; B-car, B-carotene; V,

violaxanthin; A, antheraxanthin; Z, zeaxanthin; Chlb, chlorophyll b; Chla, chlorophyll a.

212

Table A.5 Content of Individual Photosynthetic Pigments of C01 and the vte2

Mutant During Low Temperature (7.5°C) Treatment.

 

 

 

 

 

Days Lutein B-car V
J; Col Vte2-1 Col vtez-t Col vt92-1 Col vte2-1
0 154 116 165 110 66 110 79 16 43 13 46 12 37 13 40 13
1 148 17 167 121 65 14 73 114 39 12 45 16 27 13 31 15
2 154 17 170 18 64 17 72 18 42 12 43 13 34 12 41 13
5 167 18 168 19 63 13 66 14 40 11 42 13 46 13 50 13
7 160 111 160 111 58 14 58 14 42 13 42 13 46 18 46 18
14 159 113 146 126 61 13 54 113 45 12 40 19 46 14 40 16
28 184 114 105 19** 86 113 48 17** 55 13 27 15" 46 13 30 11"
Days A Z total Car Chlb
cb'id Col vte2-1 Col 17162-1 Col 17162-1 Col vte2-1
T 1 10 1 10 0 10 0 10 301 132 330 121 363 123 388 118
1 4 11 4 12 0 10 0 10 283 114 319 143 320 111 359 141
2 3 1:0 2 11 0 10 0 10 297 118 329 119 316 16 317 130
5 2 12 2 11 0 10 0 10 319 114 327 118 258 17 262 116
7 4 11 4 11 o 10 0 10 309 125 309 125 246 117 246 117
14 2 12 5 12 0 10 0 10 313 123 286 154 251 19 209 139
28 211 3 11 010 010 374130 213119" 304127 138121"

 

213

Table A.5 (continued)

 

 

Days Chle Total Chl ChlalChlb
'" Col vte2-1 Col vte2-1 Col 17192-1
cold
0 842 175 931 148 1205 198 1319 165 2.32 10.06 2.40 10.04
1 759 134 854 1107 1079 145 1212 1148 2.37 10.03 2.38 10.03
2 753 138 790 168 1069 144 1107 198 2.39 10.07 2.49 10.03
5 593 118 613 130 851 124 875 146 2.30 10.02 2.34 10.05
7 559 21:32 559 1:32 805 149 805 149 2.28 10.03 2.28 10.03

14 581 118 499 195 832 1:26 708 i134 2.32 10.03 2.39 10.02
28 729 189 357 149” 1033 3:114 495 169” 2.39 10.12 2.80 1018“

Values are expressed as pmol/mg F W. Data are means :tSD (n = 4 or 5). Student’s t tests
of vte2-1 relative to C01 (* P < 0.05, ** P < 0.01). N, neoxanthin; B-car, B-carotene; V,
violaxanthin; A, antheraxanthin; Z, zeaxanthin; total Car, total carotenoids; Chlb,

chlorophyll b; Chla, chlorophyll a; total Chl, total chlorophylls.

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215

Figure A.2 Phenotypic and Photosynthetic Responses of C01 and the vte2 and vte1
Mutants to HL stress.

Plants were grown under permissive conditions for four weeks and then transferred to HL
stress in the middle of the day. When significance is observed between genotypes
(ANOVA, P < 0.05), pair-wise comparison of least square means is evaluated and non-
significant groups are indicated by a, b or c with a being the highest group.

(A) Six representative plants after 3 days of HL1800. Bars = 2 cm.

(B) and (C) Individual values of total chlorophyll (B) and carotenoid

(C) contents from 19 leaves after 4 days of HL1 800.

(D) Individual values of Fv/Fm from 30 leaves after 24 h of HL1800.

216

 

 

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219

Figure A.3 Visible Phenotype of vte Mutants During Extended Low Temperature
Treatment.

Plants were grown under permissive conditions for three weeks and then subjected to
7.5°C treatment for the indicated time periods. (A) to (D) Representative plants of three-
week-old wild type (C01 and W5), vte1-1 and vte2-1 (Col background), vte2-2 and vte4-3
(Ws background) after 0 day (A), one month (B), two months (C) and four months (D) of
7.5°C treatment. Bars = 2 cm. (E) Representative siliques from Col, vte1-1 and vte2-1
plants after five months of low temperature treatment. Bar = 0.2 cm. Arrows denote

aborted seeds in vte1—1 and vte2-1 siliques.

220

 

Figure A.3

221

 

Figure A.3 (eont’d)

222

Figure A.4 Tocopherol, Lipid Peroxide, Anthocyanin, and Photosynthetic Pigment
Content of Col and the vte2 Mutant During Four Weeks of Low Temperature
Treatment.

Col (closed circles) and vte2-I (open squares) were grown under permissive conditions
for four weeks and then transferred to 7.5°C conditions at the beginning of the light cycle
for the indicated time. Data are means iSD (n = 3 or 4). Student’s t tests of vte2-I
relative to C01 at each time point (*P < 0.05, **P < 0.01).

(A) Total tocopherols; 4

(B) Lipid peroxides;

(C) Total chlorophylls;

(D) Anthocyanins;

(E) Total carotenoid

223

 

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Days of 7.5 °C treatment

226

25

 

30

Figure A.5 Photosynthetic Status of Col and the vte2 Mutant During Four Weeks of
Low Temperature Treatment.

Col (closed circles) and vte2-1 (open squares) were grown under permissive conditions
for four weeks and then transferred to 7.5°C conditions at the beginning of the light cycle
for the indicated time. Analysis was conducted in the middle of the light cycle. Data are
means iSD (n = 4). Student’s t tests of vte2-l relative to Col at each time point (*P <
0.05).

(A) Maximum photosynthetic efficiency (Fv/F m)

(B) Quantum yield of PSII ((ngu)

227

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0.7

0.65

 

(I) PSII
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0 5 10 15 20 25 30
Days of 7.5 °C treatment

Figure A. 5 (cont’d)

228

I

 

Starch (nmol glngFW)

 

 

Figure A.6 Changes in Starch and Soluble Sugar levels in Col and the vte2 Mutant
During Four Weeks of Low Temperature Treatment.

Col (closed circles) and vteZ-l (open squares) were grown under permissive conditions
for four weeks and then transferred to 7.5°C conditions at the beginning of the light cycle
for the indicated time. Samples were harvested at the end of light cycles. 0 days of cold
treatment indicates the end of the light cycle of the day prior to initiating 7.5°C treatment.
Starch is expressed as umol glucose equivalents / g FW. Data are means iSD (n = 3 or
4). Student’s ttests of vte2-1 relative to C01 at each time point (*P < 0.05, "P < 0.01).
(A) Starch

(B) Glucose

(C) Fructose

(D) Sucrose

229

Glucose (pmollgFW)

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230

 

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Starch (nmol glulgFW)

 

0L ' v I T T v V V I I I

-25 -13 -1 11 23 35 47 59 71 83 95 10

Figure A.7 Diurnal Changes in Starch and Soluble Sugar levels in Col and the vte2
Mutant During the First Four Days of Low Temperature Treatment.

Col (closed circles) and vte2-1 (open squares) were grown under permissive conditions
for four weeks and then transferred to 7.5°C conditions at the beginning of the light cycle
for the indicated time. Samples were harvested at the end of dark and light cycles. Gray
shadows indicate 12 h dark cycles. 0 h of cold treatment indicates the beginning of the
first light cycle of low temperature treatment. Starch is expressed as pmol glucose
equivalents / g FW. Data are means iSD (n = 5). Student’s t tests of vte2-1 relative to C01
at each time point (*P < 0.05, **P < 0.01).

(A) Starch

(B) Glucose

(C) Fructose

(D) Sucrose

231

Glucose (p mol/gFW)

 

 

 

‘ U ' I

-25 -13 -1 11 23 35 47 59 71 83 95 10

C 50 - u

Fructose (umoIIgFW)

 

 

-25 ~13 -1 11 23 35 47 59 71 83 95 107

 

 

Sucrose (nmol/gFW)

 

0' I I U U U U I U I I '

-25 -13 -1 11 23 35 47 59 71 83 95 10

Hours of 7.5°C treatment

Figure A.7 (cont’d)

232

 

 

 

 

 

 

2 q ** 0.7 '
*

I: l mature 0.6 ,
g 1.5 - Dyoung 0.5 ‘ **
T:
E 1 §O.4 -
.E . '6‘ o_3 .
C
a o 2 -
g 0.5 - -
5 0.1
g l

O ' 0 d —

Col vteZ-1 vte1-1 Col vte2-1 vte1-1

Figure A.8 Biochemical Phenotypes in Mature and Young Leaves of Col, and the
vte2 and vte1 Mutants After Four Weeks of Low Temperature Treatment.

Col, vte2-I and vte1-1 mutants were grown under permissive conditions for four weeks
and then transferred to 7.5°C conditions at the beginning of the light cycle for an
additional four weeks. Mature leaves (7th to 9th oldest, black bars) and young leaves
(13th to 16th oldest, white bars) were harvested at the end of the light cycle for analyses
in (A), (C), (D), (E) and (F). Photosynthetic parameter in (B) was measured in the middle
of the light cycle. Data are means iSD (n = 4 or 5). Student’s t tests of mutant leaves
relative to corresponding Col young or mature leaves are indicated (*P < 0.05; **P <
0.01)

(A) Anthocyanin content
(B) Quantum yield of PSII ((Dpsn)

(C) Starch content expressed as mmol glucose equivalents / g FW

(D) to (F) Glucose, fructose and sucrose content, respectively.

233

O
U

a

8
It-
It

 

 

 

 

A ** 140-
E 6001 E 120'
9 a
2 :0 100- **
3 ** E 80'
g “’0' 3 *1:
e g 60.
.r: ** o
2 200. g 40-
g * ‘9 20‘

OJ OJ

Col vte2-1 vte1-1 Col vteZ-1 vte1-1
140- 140-

Eno- E 120« H
€100- €100-
: 80. M g 80- u u
g 60' ** ** § 60:
§ 40- g 40-
h
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0 0‘

Col vt92-1 vte1-1 Col vte2-1 vte1-1

Figure A.8 (cont’d)

234

Figure A.9 Translocation and Export of 14C Labeled Photoassimilates in Low
Temperature-Treated Col and the vte2 and We] Mutants.
(A) 14C labeled photoassimilate translocation of C01 and vte2-1 treated for 7 days at

7.5°C. Percent label detected in leaves (top) and roots (bottom) are indicated as means
iSD (n = 3). Student’s ttests relative to C01 (*P < 0.05, **P < 0.01).

(B) HPLC analysis of phloem exudates collected from mature leaves of C01 and vte2-I
treated for 10 days at 7.5°C. The HPLC trace of sugar standards is shown as dotted grey
lines. The percentage of label detected in the glucose/fructose or sucrose fractions are

indicated as means iSD (n = 3). Glu, glucose;~Fru, fructose; Sue, sucrose; Raf, raffinose.

(C) Phloem exudation of 14C labeled photoassimilates from C01 and vteZ-I and vteI-I

mature leaves during 7 days of 7.5°C treatment. Total 14C fixed per mg fresh weight of

each sample at the indicated time following transfer to 7.5°C is shown below each graph.
Data are means :tSD (n = 6 to 8). Two-factor ANOVA using end points (values at 10 h of
exudation) indicates interactions are significant (P < 0.05, days of 7.5°C treatment and
genotype as factors). The pair-wise comparisons of least square means between
genotypes at l, 3 and 7 days of 7.5°C treatment are indicated as a, b or c, while 0 day is

not significant. N.A., data not available

235

A
Col vte2-1

 

 

86.8 1: 2.4 % 97.3 1 0.8 %*
« '11 .2 g :‘
l
l .2“ .1
l
3 _; .4.
13.2 1 2.4 % — 2.7 1 0.8 %*
Figure A.9

236

B Glu+Fru Suc
Col 11.0 3: 2.8 85.8 1 4.0

 

(95 de)
V1924 8.9 :l: 5.1 83.2 1 9.8
10000 -
50
3 .-
9ooo ‘ 6 1
8°00 ‘ E E + COI E-so
C 7000 - E { -D' "92'1
O f! :; T40
g 0. 0
ch
E
Q
.0

 

 

 

 

 

o 5 1o 15 20 25 30
elutlon (mln)

Figure A.9 (cont’d)

237

 

 

 

 

401 40 1
C 30- 301
.2
fl
:5
'U
:1
x
‘1’ 20. 201
.\°
10 t 10 1
G I I I I ' 0L I v I I I
0 2 4 6 8 10 0 2 4 6 8 10
Hours of exudation Hours of exudation
14C fixed (dpmlmgFW)
Col 13868 11177 7839 11090
vte2-1 15195 11197 7716 11080
vte1-1 14491 11370 N.A.

Figure A. 9 (cont’d)

238

30-
201

101

 

40¢

20¢

 

-0— Col
-Ci— vte2-1
—A— vte1-1

 

T I I V U

0 2 4 6 8 10
Hours of exudation

5915 :t 180
5381 t 209“
6314 t 305‘

Figure A. 9 (cont’d)

 

239

2 4 6 8
Hours of exudation

5192 1 490
4898 1 847
5167 1 336

10

Figure A.10 Aniline Blue Positive Fluorescence in Leaves of Col and the vte2 Mutant
During Low Temperature Treatment.

Col (C) and vte2-1 (all panels except C) were grown under permissive conditions for four
weeks and then transferred to 7.5°C at the beginning of the light cycle. Leaves were
harvested in the middle of the day before 7.5°C treatment (0 day; A and B) and after 1
day (6 h; D to F), 3 days (G to I) and 13 days (C and J to L) of 7.5°C treatment and
aniline blue positive fluorescence were observed at leaf petioles (A, D, G and J), the
lower half of leaves (B, C, E, H and K) and vein junctions (F, I and L). Arrows in (D and
F) denote highly fluorescent spots that initially appear in side veins of vteZ—I petioles

after 6 h of 7.5°C treatment. Bars = 50 pm for F, l and L and 500 pm for all other panels.

240

W
a
d
3
1
d
W

 

Figure A.10

241

 

Figure A.“ Aniline Blue Positive Fluorescence in Leaves of the vte2 Mutant at
Various Light Intensities under Permissive and Low Temperature Conditions.

vte2-I were grown under permissive conditions for four weeks and then transferred at the
beginning of the light cycle to 7.5°C 12h light/ 12 h darkness at the indicated light levels
(A to F) and in the middle of the day to HL1800 at 22°C (G and H). Leaves were
harvested in the middle of the day. Aniline blue positive fluorescence was observed at
leaf petioles (A, C, E and G) and the lower half of leaves (B, D, F and H). Bars = 500

pm.

(A) and (B) 7.5°C at 1 nmol photon m-2 5‘1 for 3 days (54h).
(C) and (D) 7.5°C at 75 nmol photon m'2 s‘1 for 2 days (30 h).
(E) and (F) 7.5°C at 800 pmol photon m'2 s'1 for 2 days (30h)

(G) and (H) 22°C at 1800 nmol photon m_2 s.1 for 4 days (72h).

242

 

Figure A.“ (cont‘d)

243

Figure A.12 Cellular Structure and Immunodetection of Callose in Co] and vte2-1
Before and After 14 Days of Low Temperature Treatment.

(G) to (L) are immunolabeled with anti-[34,3 glucan antibody. (D) to (F) are controls
with only 2° antibody. Single arrows denote phloem parenchyma transfer cell wall
ingrowths. Double arrows denote abnormal thickening of phloem parenchyma transfer
cell wall ingrowths. Single asterisks (*) mark massive wall ingrowths of phloem
parenchyma transfer cells. Double asterisks (**) mark wall ingrowths immunolabeled
with anti-[34,3 glucan. Paradermal (C and G) and transverse (E and I) sections show
entire phloem parenchyma transfer cell occluded with callose. Paradermal (C and G) and
transverse (D and H) sections show the peripheral callose sheath of phloem parenchyma
transfer cells. Note callose at boundary between phloem parenchyma transfer cell and
sieve element (F and J). Plasmodesmata between bundle sheath (upper cell) and phloem
parenchyma transfer cell immunolabeled with anti-[54,3 glucan (K). b, bundle sheath; 0,
companion cell; 5, sieve element; v, vascular parenchyma. Bars = 1 pm (all panels except
C) and 5 pm (C)

(A) vteZ-I before 7.5°C treatment.

(B) to (L) vte2-I (C to K) and Col (B and L) after 14 days of 7.5°C treatment.

244

.....yzp,._....fi

.. . . to! ~
... Ir: 1.1. Is. ...3}.ng
.. no . 1
Q.

C.

 

Figure A.12

245

 

‘ ‘ K‘ a. ~> ‘51,..A. 4: f“ .

.. 2._ v J.‘ I, ‘-
4' :1». “V k.“ "1 .. -
Figure A. 13 Cellular Structure and Immunodetection of Callose in vte2—1 After 3 Days of

Low Temperature Treatment.

(B) to (D) are immunolabeled with anti-[34,3 glucan antibody. Single arrows denote
phloem parenchyma transfer cell wall ingrowths adjacent to bundle sheath. Double
arrows denote abnormal thickening of phloem parenchyma transfer cell wall ingrowths
adjacent to companion cell. Note that transfer cell wall ingrowths are present around the
entire phloem parenchyma transfer cell (A). Transverse section of wall ingrowths
immunolabeled with anti-[34,3 glucan at phloem parenchyma transfer cell and sieve
element boundary (B) and plasmodesmata between phloem parenchyma transfer cell
(upper cell) and sieve element (C). Plasmodesmata between bundle sheath (upper cell)
and phloem parenchyma cell are continuous, lack callose-like wall depositions and are
immunonegative for anti-[34,3 glucan (D). b, bundle sheath; c, companion cell; 3, sieve

element; v, vascular parenchyma transfer cell. Bars = 0.5 pm

246

Figure A.14 Phenotypes of the vte2 Mutant and Col During HL, Drought and
Salinity Stress.

The vte2 mutant and Co] were grown at permissive conditions for four to five weeks prior
to the indicated stress treatments.

(A) Four-week-old plants were grown under permissive conditions and then transferred
to HLlOOO stress (16 h 1000 mmol photon In2 S-1 light/8h darkness at 22°C) in the

middle of the day. The graph shows the F v/F m of C01 and vte2-1 grown under permissive
conditions or after 24 h of HLIOOO. The image shows representative plants of C01 and
vte2-1 after 11 days of HLIOOO.

(B) Five-week-old plants grown under permissive conditions were subjected to drought
stress. The image shows representative plants of Co] and vteZ-I that had water withheld
for 10 days.

(C) Four-week-old plants grown under permissive conditions were subjected to salinity
stress. The image shows representative C01 and vte2-1 plants that were watered with 200

mM NaCl every other day for three weeks.

247

Dcontrol (120,113
I 100011E for 24h

vteZ- 1

~ we»!

1000,11E for 11 days

vteZ-1 vteZ-1

 

Figure A. 14

248

 

Figure A.14 (cont’d)

249

 

Figure A.lS Phenotypic and Photosynthetic Responses of Col and the vte2 and vte1
Mutants to HL stress.

Plants were grown under pemiissive conditions for four weeks and then transferred to HL
stress in the middle of the day. When significance is observed between genotypes
(ANOVA, P < 0.05), pair—wise comparison of least square means is evaluated and non—
significant groups are indicated by a, b or c with a being the highest group.

(A) Six representative plants after 3 days of HL1800. Bars = 2 cm.

(B) and (C) Individual values of total chlorophyll (B) and carotenoid (C) contents from
19 leaves afier 4 days of HL1800. ’

(D) Individual values of F v/Fm from 19 leaves after 24 h of HL1 800.

250

@808 O O

880 088 O O

6088 O O

038

 

. ‘ ‘ ‘ q ‘ ‘ C C

6 4 2 0 8 6 4 2 0

.. ....Eowueyafieozofiop

bc

C

b
Col vte2-a1 vte1-1 vte1-2

8

C

 

REG 8 0
880800 0 o

Rho I88 0

0.6950

...o h u. m. u m

r829: 33:89.3 .33

b b

vte2-1 vte1-1 vte1-2

8

Col

Figure A.15 (cont’d)

251

 

Figure A.15 (cont’d)

ocmocmmoo
CW
00 ammo

COWQDCDCIDG)

O

a a b b
Col vte2-1 vte1-1 vte1-2

252

Figure A.16 Aniline Blue Positive Fluorescence in Leaves of the vte2 and vte1
Mutant During Low Temperature Treatment.

vte2-1 (A, C and E) and vteI-I (B, D and F) were grown under permissive conditions for
four weeks and then transferred to 7.5°C at the beginning of the light cycle. Leaves were
harvested in the middle of the day after 1 day (6 h) (A and B), 2 days (C and D) and 3
days (E and F) of 7.5°C treatment and aniline blue positive fluorescence were observed at
leaf petioles (A and B), the lower half of leaves (C to F). Arrows in (A and B) denote

fluorescence spots initially appeared at side veins of petioles. Bars = 500 pm.

253

vte1-1

vteZ-1

 

E 8 son F sou N sou m

Figure A.l6

254

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