3.... iv.

.5

1..
:1".

at!
. .

$.33

 

:_.:I . +5.73%. .[fvfimr .

 

was
. 2

 

,OV LIBRARY
5#398/ up? Michiga" State

University

 

 

 

This is to certify that the

dissertation entitled
IDENTIFICATION AND FUNCTIONAL
ANALYSES OF HOMOGENTISATE
PHYTYLTRANSFERASE; A KEY ENZYME
INVOLVED IN TOCOPHEROL SYNTHESIS

IN PHOTOSYNTHETIC ORGAN I SMS
presented by

Eva Collakova

has been accepted towards fulfillment
of the requirements for

Ph.D. degree in BlamLBinng

yaw/a '

Major professor

 

7/4/03

MS U is an Affirmative Action/Eq ual Opportunity Institution 0-12771

PLACE IN RETURN BOX to remove this checkout from your record.
To AVOID FINES return on or before date due.
MAY BE RECALLED with earlier due date if requested.

 

DATE DUE DATE DUE DATE DUE

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

6/01 c:/CIRC/DateDue.p65-p. 15

IDENTIFICATION AND FUNCTIONAL ANALYSES OF HOMOGENTISATE
PHYTYLTRANSFERASE; A KEY ENZYME INVOLVED IN TOCOPHEROL
SYNTHESIS IN PHOTOSYNTHETIC ORGANISMS
By

Eva Collakova

A DISSERTATION
Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of
DOCTOR OF PHILOSOPHY
Department of Plant Biology

2003

ABSTRACT
IDENTIFICATION AND FUNCTIONAL ANALYSES OF HOMOGENTISATE
PHYTYLTRANSFERASE; A KEY ENZYME INVOLVED IN TOCOPHEROL
SYNTHESIS IN PHOTOSYNTHETIC ORGANISMS
By

Eva Collakova

Tocopherols are lipid soluble antioxidants collectively known as Vitamin E, an
essential component of the human diet. Tocopherols consist of a polar chromanol ring
and a hydrophobic prenyl chain derived from homogentisate (HGA) and phytyl
diphosphate (PDP), respectively. Condensation of these two compounds represents the
committed step in tocopherol biosynthesis catalyzed by homogentisate phytyltransferase
(HPT). HPTS in Synechocystis Sp. PCC 6803 (San-IPT) and Arabidopsis thaliana
(AtHPT) were identified based on their sequence similarity to chlorophyll synthases,
which also utilize PDP. Both SynHPT and AtHPT showed prenyltransferase activity in
vitro with HGA and PDP as preferred substrates. Neither enzyme was active with
solanesyl diphosphate, the prenyl substrate used in plastoquinone-9 biosynthesis.
Plastoquinone-9 levels were unaffected in Synechocystis 5p. PCC 6803 HPT disruption
mutant, which showed a complete absence of tocopherols. These results suggest
existence of separate polyprenyltransferase activities involved tocopherol and
plastoquinone synthesis in Synechocystis Sp. PCC 6803.

Tocopherol levels fluctuate during plant growth, possibly due to an altered
expression of genes involved in tocopherol synthesis and accumulation. Because HPT

catalyzes the committed step of tocopherol biosynthesis, it is likely that HPT is a

regulated enzyme, activity of which may limit tocopherol synthesis in non-stressed and
stressed plants. This hypothesis was addressed by generating transgenic Arabidopsis
plants constitutively overexpressing AtHPT (358::HPT1). Increases in HPT specific
activity correlated with elevated total tocopherol levels observed in transgenic lines under
normal and high-light stress conditions. Non-stressed 3SS::HPT1 lines accumulated a
4.4-fold and 40 % increase in total tocopherol levels in leaves and seeds, respectively,
relative to wild type plants. Total tocopherol levels in 358::HPT1 increased 2- to 3.8-fold
relative to wild type during high light stress. These results indicate that HPT activity
limits tocopherol biosynthesis in non-stressed and stressed wild type Arabidopsis plants.
Stress resulted in an up-regulation of HPT and several other enzymes involved in
tocopherol accumulation and concomitantly in a dramatic increase in total tocopherol
levels in both wild type and 3SS::HPT1, suggesting that HPT is not the only enzyme
involved in regulation of tocopherol biosynthesis. The presence of high levels of other
tocopherols in stressed plants also suggested a possible limitation of (Ir-tocopherol
synthesis by other tocopherol-related enzymes. Identification of the steps limiting
tocopherol synthesis is crucial for understanding the regulation of the pathway, which can

be applied in plant metabolic engineering of the tocopherol biosynthetic pathway.

TABLE OF CONTENTS

LIST OF TABLES ............................................................................................................... vi
LIST OF FIGURES ........................................................................................................... vii
KEY TO ABBREVIATIONS .............................................................................................. ix
INTRODUCTION ............................................................................................................... l
Tocopherol And Tocotrienol Structures .................................................................... l
Tocopherol Antioxidant Properties in vitro ............................................................... 3
Quenching .................................................................................................... 4
Scavenging .................................................................................................... 5
Tocopherols And Membrane Stabilization ................................................................ 8
Vitamin E Function in Animals ............................................................................... 10
Antioxidant Function .................................................................................. l 1
Roles in Intracellular Signaling ................................................................... 12
Vitamin E And Degenerative Disease Prevention ........................................ l4
oz-Tocopherol Transport And Vitamin E Deficiency in Humans ................. l7
Occurrence And Subcellular Localization of Vitamin E .......................................... 18
Plant Tocopherol, Tocotrienol, And Plastoquinone-9 Biosynthesis ......................... 20
HGA Biosynthesis - Shikimate Pathway And Tyrosine Degradation ......... 20
Prenyl-DP Biosynthesis - Non-mevalonate Pathway .................................. 24
Tocopherol, Tocotrienol, And Plastoquinone Biosynthetic Pathways .......... 27
Tocopherol Function in Plants ................................................................................ 32
Antioxidant Function .................................................................................. 32
Roles in Signaling ....................................................................................... 38
Roles in Photosynthesis .............................................................................. 39
Nutritional Genomics .............................................................................................. 40
Natural Sources of Dietary Vitamin E ......................................................... 40
Regulation And Metabolic Engineering of Tocopherol Biosynthesis in
Plants .......................................................................................................... 42
CHAPTER 1

ISOLATION AND FUNCTIONAL ANALYSIS OF HOMOGENTISATE
PHYTYLTRANSFERASE FROM S YNECHOC YST IS SP. PCC 6803 AND

ARABIDOPSIS ................................................................................................................. 47
Abstract ................................................................................................................... 48
Introduction ............................................................................................................ 48
Results .................................................................................................................... 51

Identification And Disruption of A Polyprenyltransferase Involved in
Tocopherol Biosynthesis in Synechocystis sp. PCC 6803 .......................... 51
Biochemical Characterization of the SLR1736 Gene Product ..................... 55
Identification And Characterization of An HPT Homolog from
Arabidopsis ................................................................................................. 57
Physiological Consequences of Tocopherol Deficiency in Synechocystis sp.
PCC 6803 ................................................................................................... 60
Discussion .............................................................................................................. 61
Materials And Methods .......................................................................................... 67

CHAPTER 2
HOMOGENTISATE PHYTYLTRANSFERASE ACTIVITY IS LIMITING FOR

TOCOPHEROL BIOSYNTHESIS IN ARABIDOPSIS ................................................... 72
Abstract ................................................................................................................... 73
Introduction ............................................................................................................ 73
Results .................................................................................................................... 78

Biochemical And Molecular Characterization of Wild Type And 3SS::HPT1
Plants .......................................................................................................... 78
Efieicts of HPT] Overexpression on Tocopherol Levels In Arabidopsis
Se ............................................................................................................ 81
Tocopherol Analysis of Arabidopsis Leaves And Seeds Overexpressing
Both HPT] And rT MT Transgenes .......................................................... 83
Effects of Increased Tocopherol Levels on Chlorophyll, Carotenid, And
Plastoquinone-9 (PQ-9) Content in 358::HPT1 ......................................... 86
Discussion .............................................................................................................. 87
Outlook for Metabolic Engineering of Vitamin E Levels in Crops .............. 91
Materials And Methods .......................................................................................... 92

CHAPTER 3

HOMOGENTISATE PHYTYLTRANSFERASE ACTIVITY LIMITS TOCOPHEROL

SYNTHESIS IN STRESSED ARABIDOPSIS T HALIANA LEAVES ................................ 96
Abstract ................................................................................................................... 97
Introduction ............................................................................................................ 98
Results .................................................................................................................. 101

The Biochemical Response of Wild Type And 3SS::HPT1 Plants to High
Light Stress ............................................................................................... 101
HPT Expression And Enzyme Activity in Unstressed And Stressed Wild
Type And 35S::HPT1 ............................................................................... 103
Changes in mRNA Levels of Other Tocopherol Pathway-Related Genes in
Non-Stressed And Stressed Wild Type And 358::HPT1 Leaves .............. 106
Analysis of Individual Tocopherols And Phytquuinols in Non-Stressed
And Stressed Wild Type And Transgenic Plants ...................................... 108
Discussion ............................................................................................................ l l 1
Materials And Methods ........................................................................................ 117

SUMMARY AND FUTURE DIRECTIONS .................................................................. 121
Regulation And Metabolic Engineering of Tocopherol Biosynthesis in
Arabidopsis ........................................................................................................... l 25

APPENDIX ...................................................................................................................... 129

EXPERIMENTAL PROCEDURES ................................................................................ 129
Chemicals And Standards ..................................................................................... 129
Plasmids And Mutants .......................................................................................... 131
Oxygen Evolution Rates ....................................................................................... 136
Prenyllipid Analyses ............................................................................................. 137
Real-Time PCR ..................................................................................................... 138
Prenyltransferase Assays ...................................................................................... 139

Prenyltransferase Assays Using E. coli Protein Extracts .......................... 139
Prenyltransferase Assays Using Arabidopsis Chloroplasts And Seed Protein
Extracts ..................................................................................................... 142
LITERATURE CITED ..................................................................................................... 145

LIST OF TABLES

TABLE I. Prenyllipid contents of photoautotrophically grown wild type Synechocystis sp.

PCC 6803 and the SLR1736zszr mutant .......................................................................... 55
TABLE 11. Growth and Oz evolution rates of wild type Synechocystis sp. PCC 6803 and
SLR1736::Kmr ................................................................................................................... 60
TABLE III. HPT Specific activity in chloroplasts isolated from 2-, 4-, and 6-week—old wild
type and 35S::HPT1-ll and -54 leaves ............................................................................... 80
TABLE IV. HPT specific activity in seed of wild type and 358::HPTI overexpressers (lines
11 and 54) ........................................................................................................................... 83
TABLE V. Total chlorophyll, carotenoid, and PQ—9 levels in wild type and 3SS::HPTI
Arabidopsis leaves and seeds .............................................................................................. 86
TABLE VI. Vitamin E activity in wild type, single, and double overexpressers ................... 91
TABLE VII. Probes, primers and their final optimal concentrations used for real-time

PCR .................................................................................................................................. l 19
TABLE VIII. PCR primer pairs used in this study ........................................................... 131
TABLE IX. Sequencing primers used in this study .......................................................... 134

LIST OF FIGURES

FIGURE 1. Natural tocopherols and tocotrienols and their relative Vitamin E activities ........ 2

FIGURE 2. Example of lipid peroxidation and chain-breaking antioxidant activity

of tocopherols ....................................................................................................................... 7
FIGURE 3. Shikimate biosynthetic pathway ...................................................................... 21
FIGURE 4. The plastidic l-deoxy-D-xylulose-5-phosphate (DXP) biosynthetic

pathway ............................................................................................................................... 25
FIGURE 5. Tocopherol, tocotrienol, and plastoquinone-9 biosynthesis in cyanobacteria

and plants ............................................................................................................................ 28
FIGURE 6. Generalized overview of prenquuinone biosynthetic pathways in
photosynthetic organisms ................................................................................................... 50
FIGURE 7. Alignment of Synechocystis sp. PCC 6803 and Arabidopsis
polyprenyltransferases ........................................................................................................ 53
FIGURE 8. Construction of the Synechocystis Sp. PCC 6803 SLR1736zzKrnr mutant ....... 54
FIGURE 9. HPLC traces of tocopherol standards and lipids extracted from wild type
Synechocystis Sp. PCC 6803 and SLR1736::Kmr ............................................................... 54
FIGURE 10. Homogentisate polyprenyltransferase assays ................................................ 56
FIGURE 1 1. Normal phase HPLC separation of radiolabeled prenquuinones produced
from HGA and prenyl-DP substrates by HPTS .................................................................. 58
FIGURE 12. Phylogenetic analysis of various prenyllipid polyprenyltransferases ............. 65
FIGURE 13. The tocopherol biosynthetic pathway in plants .............................................. 75
FIGURE 14. Tocopherol composition and levels in leaves of 4-week-old wild type and
homozygous 35S::HPTI Arabidopsis plants ...................................................................... 79
FIGURE 15. Tocopherol composition and levels in seed of wild type and homozygous
35S22HPT1 Arabidopsis plants ........................................................................................... 82
FIGURE 16. Tocopherol composition and levels in leaves of 5-week-old wild type,
358::y—TMT, 358::HPT1, and double 358::y-TMT/358xHPT1 overexpressers ................. 84
FIGURE 17. Tocopherol composition and levels in seed of wild type, 35S::y-T MT ,
358::HPT1, and 358::y-TMT/BSSzzHPTI overexpressers .................................................. 85
FIGURE 18. Tocopherol biosynthesis in plants ................................................................. 99

FIGURE 19. Total anthocyanin, chlorophyll, carotenoid, and tocopherol levels in stressed
wild type and 35S::HPT1 leaves ....................................................................................... 102

FIGURE 20. HPT expression in control and stressed wild type and 3SS::HPT1 plants...104

FIGURE 21. Relative HPT specific activity in control and high light stressed wild type and
35S::HPT1 Arabidopsis chloroplasts ................................................................................ 105

FIGURE 22. Expression of other tocopherol—related genes in control and stressed wild type
and 358::HPT1 leaves ....................................................................................................... 107

FIGURE 23. Tocopherol levels and composition in stressed wild type and 358::HPT1
plants ................................................................................................................................ 1 10

FIGURE 24. Tocopherol levels and composition in stressed wild type, 35S::HPT1/
358::y-T M T double overexpressers, and the corresponding transgenic

parents .............................................................................................................................. 1 1 1
FIGURE 25. Phylogenetic analysis of related prenyltransferases involved in tocopherol,

tocotrienol, plastoquinone, chlorophyll, and ubiquinone biosynthesis ............................... 122
FIGURE 26. Constructs used in this study ...................................................................... 133

viii

KEY TO ABBREVIATIONS

12-OPDA - 12-oxo-phytodienoic acid

l3-HPOT - l3-hydroperoxy-9,l1,15-octadecatrienoic acid
358::1bTM T - transgenic lines overexpressing y-TMT
358::HPTI - transgenic lines overexpressing HPT

a—TE - (Ir-tocopherol equivalents

a—TTP - a-tocopherol transfer protein

y-TMT - y-tocopherol methyltransferase

ADP - adenosine diphosphate

ATBC - Alpha-Tocopherol Beta-Carotene

AtHPT - Arabidopsis thaliana homogentisate phytyltransferase
ATP - adenosine triphosphate

BLAST - basic local alignment search tool

bp - base pair

CaMV - cauliflower mosaic virus

cDNA - complementary deoxyribonucleic acid

CDP - cytidine diphosphate

CPME - 4-(cytidine-5’-diphospho)-2-C-methyl-D-erythn'tol
CTP - cytidine triphosphate

cyt b-559 HP - high potential form of cytochrome b-559
DAHP - 3-deoxy-D-arabino-heptulosonate-7-phosphate
DHQ - 3-dehydroquinate

DHS — dehydroshikimate

DMAPP - dimethylallyl diphosphate

DMGBQ - 2,3 -dimethyl-6-geranylgeranyl- 1 ,4-benzoquinol
DMPBQ - 2,3-dimethyl-6-phytyl- 1 ,4-benzoquinol
DMPQ-9 - 2-demethylplastoquinol-9

DXP - l-deoxy-D-xylulose-S-phosphate

DXPS - l-deoxy-D-xylulose-S-phosphate synthase

E4P - erythrose-4-phosphate

E. coli - Escherichia coli

EFla - elongation factor la

EPSP - 5-enolpyruvylshikimate-3-phosphate

FW - fresh weight

G3P - glyceraldehyde-3-phosphate

GGDP - geranylgeranyl diphosphate

GGDR - geranylgeranyl diphosphate reductase

GGPSl - geranylgeranyl diphosphate synthase 1

Glc - glucose

HEPES - [4-(2-hydroxyethyl)- l ~piperazine-ethanesu1fonic acid]
HGA - homogentisate or homogentisic acid

HGAD - homogentisate dioxygenase

HPF S - Health Professionals F ollow-up Study

HPLC - high performance liquid chromatography

HPP - p-hydroxyphenylpyruvate

HPPD - p-hydroxyphenylpyruvate dioxygenase

HPT - homogentisate phytyltransferase

IPP - isopentenyl diphosphate

IU - international units

JA - jasmonate

Kmr - kanamycin resistance

ix

LDL - low density lipoproteins

MECP - 2-C-methyl-D-erythitol-2,4-cyclodiphosphate
MEP - 2-C-methyl-D-erythritol—4-phosphate

MESP - 2-C-methyl-D-erythrose-4-phosphate
MGBQ - 2-methyl-6-geranylgeranyl- l ,4-benzoquinol
MPBQ - 2-methyl—6-phytyl- 1 ,4—benzoquinol

MPBQ MT - 2-methyl-6-phytyl- l ,4-benzoquinol methyltransferase
mRNA - messenger ribonucleic acid

NADPH - reduced nicotinamid adenosyl diphosphate
nd - not detected

NHS - Nurses’ Health Study

N MR - nuclear magnetic resonance

ORF - open reading frame

PCPME - 2-phospho-4-(cytidine-5’-diphospho)-2-C-methyl-D-erythritol
PCR - polymerase chain reaction

PDS - phytoene desaturation

PDP - phytyl diphosphate

PEP — phosphoenolpyruvate

PLA2 - phospholipase A2

prenyl-DP - prenyl diphosphate

PKC - protein kinase C

PPZA - protein phosphatase 2A

PQ-9 - plastoquinone-9

PS — photosystem

PUF A - polyunsaturated fatty acid

ROS - reactive oxygen species

rRNA - ribosomal ribonucleic acid

SAM - S-adenosyl-L-methionine

SD - standard deviation

SDP - solanesyl diphosphate

SDS - sodium dodecylsulfate

SELECT - Selenium and Vitamin E Cancer Prevention Trial
sn-l - stereospecifically numbered-1

SynHPT - Synechoqutis sp. PCC 6803 homogentisate phytyltransferase
TAP - (I-IOCOPhCI'OI associated protein

TAT - tyrosine transaminase

TC - tocopherol cyclase

TLC - thin-layer chromatography

Toc -— tocopherol

Tyr — tyrosine

UV — ultraviolet

VLDL - very low density lipoproteins

INTRODUCTION

TOCOPHEROL AND TOCOTRIENOL STRUCTURES

Vitamin E comprises a group of lipid-soluble antioxidants known as tocopherols
and tocotrienols, which are synthesized by photosynthetic organisms. Other organisms,
including humans, cannot make tocopherols and therefore acquire vitamin E fiom their diets
(Brigelius-Flohe and Traber, 1999; Bramley et al., 2000). Tocopherols can quench and
scavenge various reactive oxygen species including singlet oxygen and superoxide radicals
and also can deactivate alkyl peroxy radicals formed in the lipid bilayers. These combined
activities protect polyunsaturated fatty acids from lipid peroxidation (Fukuzawa et al., 1982;
Fukuzawa and Gebicki, 1983; Munne-Bosch and Alegre, 2002a).

Tocopherols are amphipathic compounds consisting of a polar chromanol ring that
is responsible for their antioxidant properties and a 15-carbon prenyl chain, which
contributes to their lipophilic character. Based on the number and relative positions of the
individual methyl groups attached to the chromanol ring, four different tocopherol vitamin
isomers (vitamers), a, B, y, and 5, are synthesized in photosynthetic organisms (Figure 1).
Besides the methyl substituent on carbon 8 of the chromanol ring, which is present in all
natural tocopherols, two additional positions, carbons 5 and 7, can be methylated. If neither
of the two positions is methylated then the resulting vitamer will be 8-tocopherol. A single
methylation on carbon 5 yields B-tocopherol, while methylation on carbon 7 results in the
formation of y—tocopherol. Methylation of both carbons yields a-tocopherol.

Tocotrienols have a chromanol ring that is identical to that in the corresponding
tocopherols. Tocopherols and tocotrienols differ only in the degree of saturation in their
prenyl chains (Figure l). Tocopherols contain a saturated tail derived from phytyl
diphosphate (PDP). In contrast, tocotrienols have an unsaturated isoprenoid tail based on

geranylgeranyl diphosphate (GGDP). In both tocopherols and tocotrienols, the prenyl

 

Tocotrienols

 

Relative vitamin E

 

R1 R2 activity

(%)
III-tocopherol (tocotrienol) CH3 CH3 100 (30)
B-tocopherol (tocotrienol) CH3 H 50 (5)
y-tocopherol (tocotrienol) H CH3 10 (ND)
5-tocopherol (tocotrienol) H H 3 (ND)

 

Figure 1. Natural tocopherols and tocotrienols and their relative vitamin E activities. a-
Tocopherol has the highest vitamin E activity in animals as determined by the rat fetal
resorption assay. Only a—tocotrienol showed significant vitamin E activity of all
tocotrienols. Vitamin E activities of B-, y-, and 8-tocotrienols was very low or below
detection (ND - not detected) (Food and Nutrition Board, 2000).

chain is attached to carbon 2 of the chromanol head-group. The resulting configuration of
all natural tocopherols is R, R, R (2-R, 4’-R, 8’-R). Similarly, prenyl chains of all naturally
synthesized tocotrienols are also present in a single, 2-R and 3’, 7’, and 11’ trans
configuration (Kamal-Eldin and Appelqvist, 1996). However, chemical synthesis of
tocopherols yields up to eight possible stereoisomers of each tocopherol (RRR, RSR, RRS,
RSS, SRR, SSR, SRS, SSS, also known as all rac-a-tocopherol), which vary significantly
in their vitamin B activity (Eitenrrriller, 1997; Food and Nutrition Board, 2000). Because
only 2R isomers are retained in plasma, these stereoisomers exhibit significant vitamin E
activity, while ZS isomers do not. The biological activity of all rac-a—tocopherol is
approximately 50% of the biological activity of RRR-a—tocopherol, the most potent form of
vitamin E. RRR-B—, -y-, and -8-tocopherols also show lower vitamin B activity than RRR-a-
tocopherol due to their reduced retention in plasma. Biological activity of individual
tocopherols is currently expressed in terms of (rt-tocopherol equivalents (or-TE). According
to this definition, 1 mg of RRR-a—, -B—, -y-, and -8-tocopherols have biological activities of l;
0.5; 0.1; and 0.03 mg a-TE, respectively (Bramley et al., 2000; Food and Nutrition Board,
2000). These values were determined using the rat fetal resorption-gestation assay (Food

and Nutrition Board, 2000).

TOCOPHEROL ANTIOXIDANT PROPERTIES IN VITRO

Vitamin E functions are associated predominantly with the antioxidant and lipophilic
properties of the molecule. The fiee hydroxyl group on the chromanol ring and prenyl tail
at carbon 2 with R stereochemistry are required for maximal tocopherol antioxidant function
and membrane localization, respectively (Nishikimi et al., 1980; Fukuzawa et al., 1981;
Kamal-Eldin and Appelqvist, 1996). Several mechanisms for tocopherol action in terms of

antioxidant properties have been proposed. These mechanisms include quenching and

scavenging of various radicals such as reactive oxygen species (ROS) and lipid peroxy

radicals (Fryer, 1992; Liebler, 1993; Liebler, 1998; Munne-Bosch and Alegre, 2002a).

Quenching

Tocopherols are effective quenchers of singlet molecular oxygen ('02), which can
cause damage to membranes (Fahrenholtz et al., 1974; Yamauchi and Matsushita, 1979b;
Neely et al., 1988). Quenching of '02 by tocopherols occurs through resonance energy
transfer and involves a physical deactivation of 'O2 (Fahrenholtz et al., 1974; Foote et al.,
1974; Stevens etal., 1974; Neely et al., 1988). An electron from the tocopherol molecule is
donated to the 'O2 molecule to form an excited charge transfer complex that dissociates into
tocopherol and triplet (ground state) molecular oxygen (302) (Y amauchi and Matsushita,
1977). The quenching rate was proposed to be dependent on the polarity of the solvent
used for these in vitro studies because hydrophobic solvents inhibited formation of these
charge transfer complexes (Fahrenholtz et al., 1974; Neely et al., 1988). The type of
tocopherol also affected the rate of radical quenching. (rt-Tocopherol was the most effective
'02 quencher, followed by B—, y-, and 8-tocopherols, in a hydrocarbon solvent that mimicked
a natural membrane environment (Neely et al., 1988). Similarly, (rt-tocopherol was the most
effective quencher of the perferryl ions in lecithin liposomes (Fukuzawa et al., 1982). A
single molecule of or-, B—, 'y-, 8—tocopherol, and tocol was able to protect 220, 120, 100, 30,
and 20 molecules of polyunsaturated fatty acids (PUFAS), respectively, from lipid
peroxidation (Fukuzawa et al., 1982). Based on kinetic studies utilizing autosensitized
rubrene photooxidation, it was estimated that a Single molecule of a—tocopherol quenches ~

120 molecules of '02 prior to its degradation (Fahrenholtz etal., 1974).

Scavenging

Scavenging of reactive oxygen species (ROS) and alkyl peroxy radicals is based on

the chemical reaction of a tocopherol molecule with the target radical molecule.

Tocopherols can scavenge ROS including 102, superoxide anion (02-), and hydroxyl (OH')

radical and perhydroxyl (OHZ') anion (F oote et al., 1974; Yamauchi and Matsushita, 1979b;

Nishikimi etal., 1980; Fukuzawa and Gebicki, 1983; Fukuzawa et al., 1997). Electron spin
resonance studies have demonstrated that scavenging of radicals by tocopherols involves the
donation of a hydrogen atom from tocopherols to the radical and formation of a
tocopheroxyl radical (Scarpa et al., 1984; Kagan and Packer, 1994; Singh et al., 1998).
Several other intermediates and final tocopherol degradation products have also been
identified. Scavenging of ROS by tocopherols usually results in the formation of
tocopheryl hydroperoxydienones and hydroxytocopherones of varying stability that
disintegrate into tocopheryl quinone and its epoxides in hydrocarbon solvents and model
membranes (Y amauchi and Matsushita, 1979b; Nishikimi et al., 1980; Fukuzawa and
Gebicki, 1983; Neely et al., 1988; Singh et al., 1998). Besides tocopheryl quinone

formation, nitration of yutocopherol to yield 5-NOZ-y-tocopherol was also observed during

scavenging of N02- radicals (Singh et al., 1998).

Tocopherols are potent protectors of PUFAs from lipid peroxidation in spite of a
relatively low tocopherol to PUFA ratio in membranes. This ability of tocopherols to
protect lipids from peroxidation may be due in part to recycling of tocopheroxyl radicals to
tocopherols via the ascorbate/glutathione/NADPH recycling system (Nishikimi et al., 1980;
Scarpa et al., 1984; Liebler, 1993; Kagan and Packer, 1994). Scavenging of lipid peroxy
radicals is the primary mechanism by which tocopherols protect polyunsaturated fatty acids
(PUFAS) from lipid peroxidation (Liebler, 1993). Figure 2 shows an example of

peroxidation of the PUFA linolenic acid, the scavenging of the corresponding lipid peroxy

radical by (rt-tocopherol, and recycling the a—tocopheroxyl radical back to (rt-tocopherol.
Lipid peroxidation is initiated by removing a hydrogen atom from PUFA by an electron-
deficient free radical, causing the formation of an alkyl radical, which is rearranged to a
conjugated diene. Interaction of the conjugated diene with molecular oxygen results in the
formation of a lipid peroxy radical. In the absence of chain-breaking antioxidants such as
tocopherols, these lipid peroxy radicals can remove hydrogen atoms from other PUFAS,
resulting in the formation of new radicals and propagation of lipid peroxidation.
Tocopherols are able to neutralize the highly reactive lipid peroxy radicals by donating a
hydrogen atom from their chromanol hydroxyl group thus leading to the termination of
lipid peroxidation by the formation of relatively stable alkyl hydroperoxide and
tocopheroxyl radical (Fukuzawa et al., 1981; Fukuzawa et al., 1982; Scarpa et al., 1984). In
plants, alkyl hydroperoxides can be metabolized to a wide variety of lipid alcohols and
epoxides, jasmonate, n-hexenal, or traumatic acids (Schaller, 2001; Blee, 2002; Howe and
Schilmiller, 2002).

Tocopheroxyl radicals are stabilized by delocalization of unpaired electrons on the
aromatic structure of the chromanol rings (Kamal-Eldin and Appelqvist, 1996; O'Malley,
2002). There are several ways to eliminate these radicals in vitro. Tocopheroxyl radicals do
not interact with PUF As, but they can be converted back to tocopherols via
ascorbate/glutathione/NADPH recycling system or react with another radical (Scarpa et al.,
1984; Kagan and Packer, 1994; Munne-Bosch and Alegre, 2002a). Reaction of a
tocopheroxyl radical with a lipid peroxy radical yields tocopherol/lipid adducts.
Tocopheroxyl radicals may react together to generate tocopherol dimers and trirners, while
interaction of a tocopheroxyl radical with ROS yields a variety of oxidation products
including epoxy tocopherones and epoxy tocopheryl quinones (Kamal-Eldin and
Appelqvist, 1996; Liebler et al., 1996; Liebler, 1998). The extent of tocopherol recycling or
whether all these oxidation products are formed in vivo is not known. In chloroplasts, the

final products of tocopherol oxidation were identified as tocopheryl quinol and the

- — - coo" Linolenic acid

(7 — — — Alkyl radical
l 3
We” Conjugated diene
3

HO
_ _ COOH
WW3 a-Tocopherol

Lipid peroxy radical

PUFA '0
\ Alkyl o 3"

 

Propagation of lipid peroxidation
o
,2.

 

 

radical
-T
H00 a ocopheroxyl.radicai
3
,' 13-HPOT \‘
.‘ 1 RH
' n-Hexenal,
Y Traumatic HO
12-OPDA a-Tocopheryl quinol

I ' 31

OH
JA a-Tocopheryl quinone

Figure 2. Example of lipid peroxidation and chain-breaking antioxidant activity of
tocopherols. Abbreviations: electron-deficient radical (R'); polyunsaturated fatty acid
(PUFA); l3-hydroperoxy-9,11,15-octadecatrienoic acid (l3-HPOT); 12-oxo-phytodienoic
acid (12-OPDA), jasmonate (JA)

corresponding quinone (Figure 2), which may be involved in cyclic photosynthetic electron
transport around photosystem II (Yamauchi and Matsushita, 1979a; Michalski and Kaniuga,
1981; Kruk and Strzalka, 1995; Fryer, 1992; Munne-Bosch, 2002a].

TOCOPHEROLS AND MEMBRANE STABILIZATION

A single cis double bond in a fatty acid introduces a “kink” in the hydrocarbon
chain of the fatty acid, interrupting some Van der Waals interactions with other fatty acids in
the membrane. Therefore, high levels of unsaturated fatty acids in the lipid bilayer result in
increased fluidity of such membranes, which is desirable at low temperatures in terms of
increasing the tolerance of living organisms to chilling (Wada et al., 1990; Gombos et al.,
1992; Wada et al., 1993; Ishizaki-Nishizawa et al., 1996; Los and Murata, 1998). Small
amphipathic molecules such as tocopherols and cholesterol fit between the gaps formed by
unsaturated fatty acids in the membranes and increase membrane rigidity by interacting with
these fatty acids (Stillwell et al., 1996). In 1973, Diplock and Lucy proposed a model
describing possible membrane-stabilizing interactions of (rt-tocopherol with arachidonic acid
as an example of a polyunsaturated fatty acid (PUFA) based on the complementary shapes
of these two compounds by using molecular models. The methyl groups at C-4’ and C-8’
of the vitamin fit in the pockets created by A5 and A” cis double bonds of the arachidonic
acid such that both polar parts of the tocopherol and phospholipid face the same direction
where they interact with the water-soluble cellular components. Using fluorescent probes, it
has been demonstrated that the chromanol ring of a—tocopherol does not protrude into the
1ipid-waterinterphase(Bisby and Ahmed, 1989). Diplock and Lucy (1973) also proposed
that at least two cis double bonds interrupted by a methylene group are required for pocket
formation and that the straight chains of the fully saturated fatty acids do not provide the
pockets to fit the phytyl methyl groups of tocopherols. Ideally, four double bonds would

provide two pockets for the interaction of both C-4’ and C-8’ methyl groups of the

tocopherol, which would lead to the stabilization of the lipid bilayers and contribute to the
rigidity of membranes (Diplock and Lucy, 1973).

To support the hypothesis of membrane stabilization by or-tocopherol, it was
necessary to demonstrate physical interaction of (rt-tocopherol with membrane lipids. Erin
et a1. (1984) Showed that a-tocopherol can form stable complexes with free saturated and
polyunsaturated fatty acids in ethanol or when incorporated into phosphatidylcholine
liposomes or sarcoplasmic reticulum membranes in vitro. In addition, this interaction was
independent of the length of these fatty acids, but it did increase with the degree of fatty acid
unsaturation (Erin et al., 1984), consistent with the hypothesis proposed by Diplock and
Lucy (1973). The following in vitro studies have demonstrated that a-tOCOpherol exerted
certain effects on various physical properties of the model membranes, including membrane
stability and rigidity as well as the transition of gel to liquid-crystalline structure of
membranes (Wassall et al., 1986; Stillwell et al., 1996; Wang and Quinn, 2000).

Stillwell et a1. (1996) compared the effects of or-tocopherol and cholesterol on the
properties of membranes composed of phosphatidylcholine containing saturated stearic acid
in the sn-1 position and various unsaturated fatty acids in the sn-2 position. Both
cholesterol and (rt-tocopherol broadened and lowered the main transition of gel to liquid-
crystalline structure of the membrane (Stillwell et al., 1996). Similar effects were observed
when a-tocopherol was added to the model multilamellar membrane consisting of egg yolk
lysophosphatidylcholine with [2H3l]—palmitic acid at the sn-2 position (Wassall et al., 1986).
Cholesterol and a—tocopherol had different effects on the membranes containing different
types of unsaturated fatty acids (Stillwell et al., 1996). a—Tocopherol had a larger and more
complex effect than cholesterol on the membranes containing a high content of PUFAs with
a double bond before the A5 position, suggesting that its mode of interaction with fatty acids
in lipid bilayers is different from that of cholesterol. (rt-Tocopherol decreased the
permeability to protons in all types of membranes tested, while cholesterol was effective

only with less unsaturated fatty acids. a—Tocopherol was shown to exert the highest effect

on membranes composed of highly unsaturated fatty acids, which coincides with its role in
protecting these PUFAS from lipid peroxidation (Stillwell et al., 1996).

It appears that (rt-tocopherol associates preferentially with PUFAS and this
interaction causes an increase in membrane rigidity. In all these in vitro studies,
phospholipid model membranes simulating animal membranes and non-physiologically
high concentrations of (rt-tocopherol were used (Erin et al., 1984; Wassall et al., 1986;
Stillwell et al., 1996; Wang and Quinn, 2000). No direct in viva evidence or studies using
membranes resembling the high galactolipid content of chloroplasts are available.
Photosynthetic organisms regulate their membrane fluidity through controlling unsaturation
of fatty acids (Wada et al., 1990; Gombos et al., 1992; Wada et al., 1993; Ishizaki—
Nishizawa et al., 1996; Los and Murata, 1998) and the contribution of a-tocopherol to the
regulation of membrane fluidity in plants is unclear. Cold-treated plants tend to have
reduced (rt-tocopherol levels relative to the plants grown at optimal temperatures (Fryer et al.,
1998; Leipner et al., 1999), which is expected because lower a-tocopherol levels translate
into higher membrane fluidity. However, this observation could also be a result of a-
tocopherol degradation due to oxidative damage during cold stress. Mutants and transgenic
plants with the altered levels and composition of a-tocopherol and PUFAS should be

helpful in addressing this question.

VITAMIN E FUNCTION IN ANIMALS

Tocopherol function in animals is mostly connected to their antioxidant properties,
especially their ability to protect PUFAS from lipid peroxidation. In animals, vitamin E
contributes to the normal muscular, neural, and immune function and was originally found
as a factor essential during pregnancy. In recent years, additional roles for (rt-tocopherol in

cellular signaling have been proposed in animals that may be unrelated to their antioxidant

10

properties because these functions are not performed by other tocopherols exhibiting

Similar antioxidant activities.

Antioxidant Function

The ability of tocopherols to quench and scavenge various free radicals in model
membranes predeterrnines their antioxidant activity in living organisms. In fact, there is an
inverse correlation between tocopherol levels and the degree of oxidative damage in
membranes. Male mouse livers, which were highly susceptible to oxidative damage induced
by 2-butoxyethanol, showed lower vitamin E levels than the less susceptible female mice
and rat livers (Siesky et al., 2002). In addition, tocopherol supplementation had protective
effects on lipids in different animal cells and tissues, in which oxidative damage was
induced by treatments with various prooxidants. For example, hydrogen peroxide-induced
oxidative stress resulted in a significant decrease in PUFA levels relative to control and
HzOz/vitamin E-treated guinea pig brains (Celik and Ozkaya, 2002). Vitamin E also
inhibited lipid peroxidation and cell death exerted by paraquat in isolated hepatocytes
(Watanabe et al., 1986). Skin treatment with curnene hydroperoxide caused an increased
radical adduct formation in keratocytes of vitamin E-deficient relative to vitamin E-sufficient
mice (Schvedova etal., 2002). Mitochondria in rat livers perfused with butylhydroperoxide
showed increased lipid peroxidation, which was inhibited by the addition of a—tocopherol
(Ham and Liebler, 1997). The degree of (rt-tocopherol degradation was similar in both
vitamin E treated and non-treated rat livers, indicating that the lipid peroxidation inhibition
occurred due to supplementary (rt-tocopherol (Ham and Liebler, 1997). Collectively, these
studies suggest that tocopherols reduce oxidative damage by protecting PUFAS from lipid

peroxidation in animals.

11

Roles in Intracellular Signaling

In animals, vitamin E has been proposed to affect signaling by modulating activities
of some components of the signal transduction pathways involved in regulation of
processes, such as cell proliferation, apoptosis, and inflammation (Azzi et al., 2000;
Meydani, 2001; Ricciarelli et al., 2001). Inflammatory processes are associated with a
phospholipase A2 (PLA2)-dependent release of arachidonic acid, followed by an induction
of lipoxygenases and cyclooxygenases, enzymes producing intermediates in oxylipin
synthesis (Balsinde et al., 1999). Vitamin E inhibits PLA2 activity in a non-competitive
manner by affecting physical properties of the membrane rather than directly interacting
with the enzyme (Grau and Ortiz, 1998). In contrast, vitamin E up-regulated PLA2 activity
in heart myoblasts and megakaryocytes (Tran et al., 1996; Chan et al., 1998). In mouse
macrophages, cyclooxygenase activity and prostaglandin formation can be induced by lipid
peroxidation and inhibited by (rt-tocopherol through its antioxidant function (Beharka et al.,
2002). Induction of platelet aggregation and cell proliferation involve protein phosphatase
2A (PP2A) inhibition and subsequent protein kinase C (PKC) activation. (rt-Tocopherol has
been shown to inhibit PKC activity by altering the PKC phosphorylation state by activating
PP2A in platelets and smooth muscle cells at the post-translational level (Ricciarelli et al.,
1998; Azzi et al., 2001; Freedman and Keaney, 2001). In mammary epithelial cells, all
tocotrienols, 6-tocopherol, but not or-, [3-, and y—tocopherols, were effective inhibitors of
PKCa activity and nritogenesis, suggesting an antioxidant-independent mechanism for these
vitamin E forms (Sylvester et al., 2001).

Apoptosis occurs through the induction of caspase cascade and also involves
modulation of PKC activity (Nicholson and Thomberry, 1997; Bang et al., 2001; Neuzil at
al., 2002). Differential effects of different vitamin E forms on apoptosis in distinct
experimental systems have been observed, which could be a result of the existance of

different PKC isofonns (Y u et al., 1999; Bang et al., 2001; Neuzil et al., 2002; Uemura et

12

al., 2002). Both a- and B-tocopherols efficiently scavenged reactive oxygen species,
reducing the degree of oxidized LDL-induced apoptosis in vascular endothelial cells, but
only a-tocopherol was the most effective form of vitamin E inhibiting caspase-3 activity
(Uemura et al., 2002). In human breast cancer cells, all tocotrienols and 6-tocopherol
efficiently induced apoptosis, while a-, B—, and y—tocopherols were ineffective even at high
concentrations (Yu et al., 1999). These differences in biological activities of the individual
forms of vitamin E point out a mechanism of tocopherol action beyond its antioxidant
properties (Yu et al., 1999; Azzi et al., 2001; Meydani, 2001; Ricciarelli et al., 2001;
Sylvester et al., 2001; Uemura et al., 2002).

A non-antioxidant a—tocopherol firnction may also involve an effect on gene
expression in animals. Both a- and 5-tocopherols induced expression of the gene encoding
a-tocopherol transfer protein (Fechner et al., 1998), which is responsible for specific
transport of a-tocopherol to lipoproteins in the liver (Arita et al., 1997). In rat vascular
smooth muscle cells, (rt-tocopherol, but not B—tocopherol, induced expression of a-
tropomyosin, which may play a role in early stages of atherosclerosis progression, by an
antioxidant-independent mechanism (Aratri et al., 1999). Neither study showed whether or-
tocopherol effects on the expression of these genes were direct or indirect. There is ample
evidence that the lipid soluble vitamins A and D directly regulate gene expression and
Similar roles have been proposed for the structurally similar a—tocopherol (Carlberg, 1999).
In recent years, a-tocopherol associated protein (TAP) has been shown to specifically bind
a-tocopherol, but not other tocopherols (Stocker et al., 1999; Yamauchi et al., 2001). Upon
binding a-tocopherol, TAP translocated from cytoplasm to the nucleus and activated
reporter gene expression, suggesting that TAP-a-tocopherol complex may function as a

transcriptional activator (Y arnauchi etal., 2001).

13

Vitamin E And Degenerative Disease Prevention

A positive correlation between oxidative damage and progression of certain
degenerative disease has been suggested. The ability of vitamin E to prevent oxidative
damage is thought to play a key role in decreasing the risk of cardiovascular disease and
certain types of cancer. Coronary heart disease is associated with the formation of
atherosclerotic depositions due to oxidized cholesterol present in low density lipoproteins
(LDL). Vitamin E may prevent atherogenesis by reducing oxidized cholesterol levels in
LDL (Buring and Hennekens, 1997; Brigelius-Flohe et al., 2002). Oncogenesis is initiated
by DNA damage caused by free radicals. Because tocopherols are able to quench and
scavenge fiee radicals, they may reduce the risk of cancer by preventing incidence of DNA
mutagenesis (Bramley et al., 2000; Brigelius-Flohe et al., 2002; Schwenke, 2002). Non-
antioxidant vitamin E functions may also provide a molecular basis for its mechanism in
chemoprevention of these diseases (Buring and Hennekens, 1997; Azzi et al., 2000; Azzi et
al., 2001; Meydani, 2001; Ricciarelli et al., 2001). This mechanism involves modulating
signal transduction pathways associated with the progression of atherosclerosis and cancer,
such as platelet aggregation, cell division, and apoptosis (Freedman and Keaney, 2001; Tang
and Meydani, 2001; Uemura et al., 2002).

Several studies have demonstrated positive effects of increased vitamin E intakes on
reducing atherosclerosis and cancer in animals. (rt-Tocopherol supplementation to
experimental animals helped to reduce plasma peroxide levels and subsequently
atherosclerotic plaque formation, thus decreasing the risk of heart disease (Smith and
Kummerow, 1989; Wojcicki et al., 1991; Schwenke et al., 2002). Increased a—tocopherol
intakes were correlated with a decreased DNA damage and the inhibition of tumor grth in
rodents (Fleshner et al., 1999; Factor et al., 2000; Slarnenova et al., 2002). The beneficial
effects of vitamin B were also associated with the inhibition of cell proliferation and

induction of tumor cell apoptosis in viva (Barnett et al., 2002; Malafa et al., 2002). While

14

all these studies suggested that high levels of vitamin E reduced the risk of these
degenerative diseases, conclusive evidence for beneficial vitamin E effects in humans is still
lacking.

Two types of clinical studies have been performed to assess whether increased
vitamin E intakes help to reduce the risk of heart disease and cancer in humans:
observational (prospective) studies and randomized trials. In the observational studies, a
large group of healthy individuals was divided into sub—groups based on the amount of
consumed vitamin E they reported along with other health-related information in
questionnaires. In randomized trials, 3 large group of randomly selected participants was
given daily a pharmacological dose of vitamin E or a placebo. In both types of trials,
various end-points such as death or manifestation of disease was observed within several
years and statistical analyses were performed to correlate high tocopherol consumption or
plasma levels with the decreased risk of the disease.

The Health Professionals Follow-up Study (HPFS), the Iowa Women’s Health
Study, and the Nurses’ Health Study (NHS) are examples of prospective cohort trials
(Rimm et al., 1993; Stampfer et al., 1993; Kushi et al., 1996). In the first two trials, a
statistically significant decrease in the incidence of myocardial infarction or mortality was
observed in participants with the highest vitamin E intakes (Rimm et al., 1993; Kushi et al.,
1996). In contrast, the NHS study did not Show this type of correlation (Stampfer et al.,
1993). lnconclusive results were also obtained from randomized trials. The Cambridge
Heart Antioxidant Study and the Secondary Prevention with Antioxidants of Cardiovascular
Disease in Endstage Renal Disease study suggested that pharmacological doses of RRR-a—
tocopherol (up to 800 IU daily) are beneficial for patients with heart disease in terms of
reducing the incidence of myocardial infarction, ischerrric stroke, and other heart disease-
related symptoms (Stephens et al., 1996; Boaz et al., 2000). In other studies, high a-
tocopherol intakes had no effect on cardiovascular disease progression (The ATBC Cancer

Prevention Study Group, 1994; The HOPE Study Investigators, 2000).

15

Several observational and randomized trials attempting to demonstrate a correlation
between high Vitamin E intakes and a decreased risk of various cancer types have also been
conducted. In many trials, high vitamin E consumption was associated with the decreased
risk of some cancers, while others failed to show this correlation. Some observational
studies showed the inverse relationship between increased vitamin E consumption and the
risk of non-melanoma skin, esophageal, and epithelial ovarian cancers (Bidoli et al., 2001;
Bollschweiler et al., 2002; Davies et al., 2002). In the Cancer Prevention Study H,
significantly lower mortality rates were associated only with regular vitamin E
supplementation periods exceeding ten years in patients suffering from bladder, but not
stomach cancer (Jacobs et al., 2002a; Jacobs et al., 2002b). Controversial results were also
obtained in the NHS and HPFS studies, in which high vitamin E intakes reduced the risk of
colon cancer in men, but not in women (Wu et al., 2002). In the randomized ATBC study,
besides fewer new cases of prostate cancer in the smokers receiving (rt-tocopherol relative to
the placebo group, there was no overall positive effect of vitamin E supplementation on the
gastric and lung cancer incidence or the mortality rate (The ATBC Cancer Prevention Study
Group, 1994; Malila et al., 2002).

Brigelius-Flohe et a1. (2002) pointed out that some factors such as dose and form of
vitamin E, initial vitamin E levels and genetic pre-disposition for the disease of selected
individuals, age and selection of individuals and stage of the disease have not been the same
or considered in these trials, which may explain some of the controversial results. Negative
results could also be due to low amounts of vitamin E or relatively short follow-up periods
used in some studies. For example, consumption 50 mg of all me a-tocopheryl acetate
daily for up to 8 years was probably too low to prevent the incidence of most cancers in the
ATBC study (The ATBC Cancer Prevention Study Group, 1994). Results from the
ongoing randomized trials, such as SELECT (the SELenium and vitamin E Cancer

prevention Trial), should provide more conclusive evidence than some previous trials about

16

the potential beneficial effects of vitamin E in degenerative disease chemoprevention (Klein

et al., 2001).

(rt-Tocopherol Transport And Vitamin E Deficiency in Humans

In humans, lipids and tocopherols are absorbed, assembled into chylomicrons in the
intestine and transported to the liver through the lymphatic system (Traber and Arai, 1999).
Discrimination between (rt-tocopherol and other forms of vitamin E occurs only after several
hours of tocopherol ingestion in the liver, where a-tocopherol transfer protein (a—TTP)
specifically binds (rt-tocopherol, which is subsequently packed into very low density
lipoproteins (VLDL) and distributed to the target organs through the bloodstream (Traber et
al., 1990; Traber and Arai, 1999). In addition to a-TTP, other proteins capable of binding
(rt-tocopherol have recently been identified in rats and humans (Stocker et al., 1999; Voegele
et al., 2002). Afamin is a protein found in human plasma able to specifically and efficiently
bind both a- and y-tocopherols at physiological concentrations. This albumin-type protein
has been proposed to play similar roles in the transport of the two tocopherols in the
bloodstream as chylomicrons and VLDL (Voegele et al., 2002). In contrast, (rt-tocopherol-
associated protein (TAP) belongs to a family of lipophillic ligand-binding proteins and is
most likely involved in regulation of gene expression (Stocker et al., 1999; Yamauchi et al.,
2001)

Genetic or metabolic disorders in any of these steps of tocopherol absorption and
transport may result in vitamin E deficiency, which is associated with severe neurological
disorders including cerebral ataxia, myopatlries, and sensory problems in humans
(Brigelius-Flohe and Traber, 1999). oc-TTP specifically facilitates a-tocopherol transport in
parenchymal hepatocytes, resulting in high plasma levels of (rt-tocopherol, but not other
forms of vitamin E (Arita et al., 1997). Patients with a non-functional a-TTP show low a-

tocopherol levels in plasma due to the impaired a-tocopherol transport in liver (Traber and

17

Arai, 1999). Besides defective genes encoding a-TTP and possibly other (rt-tocopherol
transport proteins, additional disorders such as lipid malabsorption or hyperlipidemia may
result in vitamin E deficiency (Burton and Traber, 1990; Traber and Arai, 1999). Because
a-tocopherol absorption efficiency is dependent on lipid intake, vitamin E deficiency may be
a secondary effect in patients suffering from lipid absorption disorders. Hyperlipidernic
patients may have normal plasma (rt-tocopherol levels, but they are actually deprived of
Vitamin B because their abnormally high lipid levels contribute to the reduced or-

tocopherol/lipid ratio (Burton and Traber, 1990; Traber and Arai, 1999).

OCCURRENCE AND SUBCELLULAR LOCALIZATION OF VITAMIN E

Vitamin E can be found nearly in all living organisms. However, only
photosynthetic organisms such as cyanobacteria, algae, and plants are able to synthesize
tocopherols and tocotrienols. Exceptions include some members of the cyanobacterial
Species such as Anacystis, Synechacaccus, and Prachlaracaccus, which completely lack and
are unable to synthesize tocopherols because their genomes do not contain the necessary
tocopherol biosynthetic genes (Dasilva and Jensen, 1971; Thomas et al., 1998; Sattler and
DellaPenna, 2003). The lack of tocopherols coincides with the traces or absence of PUFAS
observed in some of these cyanobacteria (Stanier and Cohen-Bazire, 1977; Merritt et al.,
1991). These observations are consistent with the proposed roles of tocopherols as
protectors of PUFAS from lipid peroxidation (Liebler, 1993). Most cyanobacteria
synthesize PU FAs and (rt-tocopherol and may accumulate some of the prenylchromanol and
quinol precursors of or-tocopherol (Dasilva and Jensen, 1971; Henry et al., 1987; Merritt et
al., 1991; Wada et al., 1993; Shintani and DellaPenna, 1998). In plants, tocopherol and
tocotrienol levels and composition vary widely among different tissues and between Species.

Tocopherols accumulate in photosynthetic tissues, flowers, fi'uits, and seeds of all plants,

18

while tocotrienols have been found in gymnosperrns and in the seeds of monocots (Piironen
etal., 1986; Franzen et al., 1991; Bramley et al., 2000; Ching and Mohamed, 2001).

In leaves and pine needles, (rt-tocopherol is the predominant form present, but small
amounts of y—tocopherol may accumulate as well (Piironen et al., 1986; Franzen and Haas,
1991; Molina-Torres and Martinez, 1991; Shintani and DellaPenna, 1998). In
photosynthetic tissues, tocopherols are localized mostly in membranes and plastoglobuli of
chloroplasts (Griffiths et al., 1967; Lichtenthaler et al., 1981; 8011 et al., 1984; S011 et al.,
1985). In spinach chloroplasts, (rt-tocopherol comprised approximately 64%, 20%, and
34% of the total prenquuinones in envelopes, thylakoids, and plastoglobuli, respectively
(Lichtenthaler et al., 1981). When expressed on a protein basis, the outer envelope of
spinach chloroplasts contained 50% more (rt-tocopherol than the inner envelope and or-
tocopherol was the major prenquuinone in both envelopes (Soll et al., 1984; 8011 et al.,
1985)

Plant seeds contain the highest tocopherol levels of all tissues, with y-tocopherol
being the major contributor to the total tocopherol in seeds of many plants including
soybean, mung bean, canola, maize, and Arabidopsis (Franzen and Haas, 1991; Shintani and
DellaPenna, 1998; Yoshida et al., 1998; Goffrnan and Mollers, 2000; Goffrnan and Bohme,
2001). In contrast, sunflower, safflower, and wheat seeds accumulate predominantly or-
tocopherol (Bramley et al., 2000; Delgado-Zamarreno et al., 2001). Other tocopherols,
namely [3- and 8-tocopherols, are present at small quantities in most seeds (Delgado-
Zamarreno et al., 2001; Goffrnan and Bohme, 2001; Goffman and Mollers, 2000). High
levels of tocotrienols can be found in seeds of monocots such as rice, maize, and palm
(Goffman and Bohme, 2001; Eitenmiller, 1997; Franzen and Haas, 1991), primary needles
and young seedlings of gymnosperms (Franzen and Haas, 1991; Franzen et al., 1991) and
some fruits and berries (Piironen et al., 1986). Within the seed, tocopherols and
tocotrienols accumulate in different parts. In oat, tocopherols and tocotrienols accumulated

preferentially in the germ and endosperm, respectively (Peterson, 1995). Tocopherol

l9

analysis of dissected seeds of three different soybean cultivars Showed that the axis had the
highest tocopherol content, followed by the cotyledons and seed coat (Yoshida et al., 1998).
Because cotyledons accounted for ~ 93% of total seed weight, they contributed to the
greatest extent to the final tocopherol levels of soybean seeds. In general, y-tocopherol was
the predominant tocopherol in all parts of the soybean seed, followed by 5- and a-
tocopherols (Yoshida et al., 1998).

PLANT TOCOPHEROL, TOCOTRIENOL, AND PLASTOQUINONE-9
BIOSYNTHESIS

Tocopherols consist of a chromanol ring and a lS-carbon prenyl chain derived from
homogentisate (HGA) and phytyl diphosphate (PDP), respectively. In tocotrienol
synthesis, geranylgeranyl diphosphate (GGDP) is used instead of PDP as a prenyl tail,
while plastoquinone-9 (PQ-9) is synthesized from HGA and solanesyl diphosphate (SDP).
HGA and these prenyl diphosphates (prenyl-DP) are synthesized by two independent
pathways (Figures 3 and 4). In plants, HGA originates from Shikimate pathway in the
cytosol (Figure 3) and PDP is synthesized from isopentenyl diphosphate (IPP) derived
from l-deoxy-D-xylulose-S-phosphate (DXP) in plastids (Figure 4).

HGA Biosynthesis - Shikimate Pathway And Tyrosine Degradation

Shikimate biosynthesis is found only in microorganisms and plants. In plants, the
Shikimate pathway provides intermediates for the synthesis of aromatic amino acids
(tyrosine, phenylalanine, and tryptophan), lignin, and phenylpropanoids and phenolic
compounds involved in protection of plants from pathogens (Hernnan, 1995; Herrrnan and
Weaver, 1999). Up to 20% of the photosynthetically fixed carbon can be directed to the
plant Shikimate pathway (Hernnann, 1995). In plants, all enzymes of the pathway are

predicted to be chloroplast localized because they contain chloroplast targeting sequences

20

(Herrmann and Weaver, 1999). Shikimate synthesis starts with the condensation of two
activated compounds, phosphoenolpyruvate (PEP) and erythrose—4-P (E4P) catalyzed by 3-
Deoxy-D-arabino-heptulosonate-7-P (DAHP) synthase to yield DAHP (Figure 3).
Antisense suppression of transketolase, which catalyzes E4P formation, resulted in a
reduction of aromatic amino acid and tocopherol levels in transgenic tobacco, confirming
that Shikimate pathway provides aromatic precursors for tocopherol biosynthesis (Henkes et
al., 2001). Genes encoding several isoforms of DAHP synthase have been cloned from a

variety of plants including potato, tobacco, Arabidopsis, and tomato (Dyer et al., 1989;

_ Pi HO coo" Pi HO coo
coo on o
,k + P0 CHO _oi‘—p P0 _o:p
PO on o on
PEP E4P °" DAHP °” DHQ 0"
PEP COO H 20

A} ”704%. Mme—z:

EPSP Shikimate-3P Shikimate
Pi
coo' cuzcocoo’

CH 2cocoo
:ocoo WMEZ :HPF; £0 Xi’: ,CHzcoo"
GA
OH COZ
Chortsmate Prephenate r0“

Plastoquinone 1 Degradation
Tocopherols

. . products
Tyrosme \ Tocotrienols

Figure 3. Shikimate biosynthetic pathway. The Shikimate pathway leads to the formation
of homogentisate (HGA) used in the tocopherol, tocotrienol, and PQ-9 biosynthesis in
photosynthetic organisms. The dashed arrow represents multiple biosynthetic steps.
Enzymes are represented by numbered circles: l. 3-deoxy-D-arabino-heptulosonate-7-P
(DAHP) synthase; 2. 3-dehydroquinate (DHQ) synthase; 3. DHQ dehydratase; 4.
Shikimate dehydrogenase; 5. Shikimate kinase; 6. 5-enolpyruvylshikimate-3-P (EPSP)
synthase; 7. chorismate synthase; 8. chorismate mutase; 9. prephenate dehydrogenase; 10.
p-hydroxyphenylpyruvate (HPP) dioxygenase; ll. tyrosine transaminase (TAT); 12. HGA
dioxygenase

     

21

Keith et al., 1991; Wang et al., 1991; Gorlach et al., 1993a). 3-Dehydroquinate (DHQ)
synthase is an oxidoreductase involved in the formation of DHQ from DAHP (Bartlett and
Satake, 1988). Tomato Dl-IQ synthase is similar to the bacterial homologues and the DHQ
synthase cDNA was shown to firnctionally complement an E. cali mutant lacking DHQ
synthase activity (Bischoff et al., 1996).

DHQ is converted in two steps to Shikimate by a single enzyme,
dehydroquinase/Shikimate dehydrogenase (Mousdale et al., 1987; Bonner and Jensen,
1994). In tobacco, the N terminus of this enzyme contains the dehydroquinase activity,
while the C terminus contains the shikirnate dehydrogenase activity (Bonner and Jensen,
1994). Several wound-inducible Shikimate dehydrogenase isozymes have been identified in
pepper leaves (Diaz and Merino, 1998). The next enzyme of the pathway is Shikimate
kinase, which uses ATP to phosphorylate the hydroxyl at carbon 3 on the Shikimate
molecule. Shikimate kinase is encoded by a single gene in tomato and is processed to a
mature form in chloroplasts (Schmid et al., 1992). The product of this enzyme, shikimate-
3-P, is a substrate for 5-enolpyruvylshikimate—3-P (EPSP) synthase, cDNAs for which have
been isolated from several plant species (Shah et al., 1986; Gasser et al., 1988; Gasser and
Klee, 1990). EPSP synthase is the target of the herbicide glyphosate and many plant
species have been subjected to metabolic engineering to improve their resistance to this
herbicide (Steinrucken and Amrhein, 1980; Shah etal., 1986).

After the condensation of shikimate-3-P and PEP, EPSP is dephosphorylated to
chorismate by the last enzyme of the Shikimate pathway, chorismate synthase (Hawkes et
al., 1990). Chorismate synthase genes have been isolated from Synechacystis sp. PCC
6803 and tomato (Gorlach et al., 1993b; Schmidt et al., 1993). Chorismate synthase is
targeted and processed in chloroplasts and only mature forms of this enzyme are firnctional
(Henstrand et al., 1995; Braun et al., 1996). Chorismate is the last product of the Shikimate
pathway and it is used as a substrate in the biosynthesis of many primary and secondary

metabolites. Chorismate mutase plays a central role in the biosynthesis of tyrosine and

22

phenylalanine as it catalyzes the first committed step in the synthesis of these aromatic
amino acids, conversion of chorismate to prephenate. In Arabidopsis, two out of three
isoforms of chorismate mutase contain chloroplast targeting sequences (Mobley et al.,
1999). Expression studies suggested that the three isoforms play different roles in
regulating chorismate mutase activity during development and pathogen attack in
Arabidopsis (Mobley et al., 1999).

Prephenate dehydrogenase oxidizes prephenate to form p-hydroxyphenylpyruvate
(HPP), a direct biosynthetic precursor of tyrosine and HGA. Transarnination of HPP
yields tyrosine, which is used in the biosynthesis of proteins or aromatic compounds
(Lopukhina et al., 2001). Excess tyrosine may be converted back to HPP because the
transamination is a reversible reaction. Radiotracer studies showed that HPP and tyrosine
were incorporated into tocopherols and plastoquinones in chloroplasts isolated from a
variety of plants, suggesting that HPP used for tocopherol and plastoquinone biosynthesis
can be formed not only directly from prephenate, but also from tyrosine (Whistance and
Threlfall, 1970; Fiedler et al., 1982; Schulz et al., 1993).

In plant tocopherol, tocotrienol, and plastoquinone synthesis, HPP is converted to
HGA by HPP dioxygenase (HPPD), a cytosolic enzyme that requires Fez“: ascorbate, and
molecular oxygen for its optimal activity (Fiedler et al., 1982; Schulz et al., 1993; Norris et
al., 1995; Garcia et al., 1997; Norris et al., 1998; Garcia et al., 1999; Tsegaye et al., 2002).
In Arabidopsis, HPPD is encoded by a single nuclear gene, PDS], disruption of which is
lethal due to a lack of plastoquinone, tocopherols, chlorophylls, and carotenoids (Norris et
al., 1995; Nonis et al., 1998). The lack of carotenoids and chlorophylls is a pleiotrophic
rather than a direct effect of the deficiency in HPPD activity. Plastoquinone is required as
an electron acceptor in the phytoene desaturation reactions in carotenoid biosynthesis
(Mayer et al., 1990). Because Arabidopsis pdsl mutants are unable to synthesize
carotenoids and tocopherols, the major lipid-soluble antioxidants in chloroplasts, they are

photobleached (Norris et al., 1995). In contrast to the Arabidopsis pdsl mutants,

23

Synechacystis sp. PCC 6803 HPPD disruption mutants also lack tocopherols, but have
normal levels of PQ-9 and other prenyllipids (Dahnhardt et al., 2002). These results
suggest that HPPD is required only for tocopherol biosynthesis and PQ-9 biosynthesis
most likely occurs through an HPPD-independent pathway in Synechacystis sp. PCC 6803
(Dahnhardt et al., 2002).

Prenyl-DP Biosynthesis - Non-mevalonate Pathway

Until 1993, it was widely accepted that all isoprenoids are derived from the
acetate/mevalonate pathway (Goldstein and Brown, 1990). In recent years, a novel pathway
for isoprenoid synthesis has been discovered in plants and other organisms, including
bacteria (Rohmer et al., 1993; Eisenreich et al., 1996; Schwender et al., 1996; Lichtenthaler
et al., 1997b; Disch et al., 1998; Proteau, 1998). In photosynthetic organisms, this
mevalonate-independent, or 1-deoxy-D-xylulose-S-phosphate (DXP), pathway (Figure 4) is
the major source of isopentenyl diphosphate (IPP) used for the synthesis of all plastidic
isoprenoids (carotenoids, isoprene, and side chains of tocopherols, chlorophylls,
plastoquinones, etc.), while the mevalonate pathway is used for the synthesis of cytosolic
isoprenoids (sterols, sesquiterpenes, and triterpenes) (Eisenreich et al., 1998; Lichtenthaler,
1998; Logan et al., 2000). However, a minimal IPP exchange between cytosol and
chloroplasts might occur (Arigoni et al., 1997; Estevez et al., 2000; Nagata et al., 2002).

The initial step in IPP synthesis is the condensation of g1yceraldehyde-3-phosphate
(G3P) and pyruvate for the synthesis of the five-carbon compound DXP (Rohmer et al.,
1996; Arigoni et al., 1997). This reaction is catalyzed by a transketolase-like enzyme, DXP
synthase, which utilizes thiamin diphosphate as a cofactor. Genes encoding DXP synthase
have been cloned and functionally characterized from a variety of plants including

Arabidopsis, bell pepper, tomato, and peppermint (Mandel et al., 1996; Bouvier et al., 1998;

24

Lange et al., 1998; Estevez et al., 2000; Lois et al., 2000). In Arabidopsis, DXP synthase is

encoded by a single nuclear gene, CLA] and the corresponding claI mutants

Y°°° °"‘H’\°—o"—»°fi"‘r\° 0PM

OH OH
Pyruvate MEP

G3P DXP
$0:
0,, .. o CMP

MECP |><(‘OP 4:9— ??oio—coPd—o— Ffi’cKO- CDP

mm, mm mm
9 ’. PCPME CPME
HM OPP 9
OH 9 HMBPP Chlorophylls
vaopw'p AA,"
DMAPP Phytol

Carotenoids {--O ”—0—;P

Tocotrienols GGDP

 
 

5 IPP
W Chlorophylls
PPO Phylloquinone
\ H Tocopherols
9 ‘ i
SDP ‘A Plastoquinone-9 r

Figure 4. The plastidic l-deoxy-D-xylulose-5-phosphate (DXP) biosynthetic pathway.
The DXP pathway leads to the formation of the prenyl diphosphates phytyl diphosphate
(PDP), geranylgeranyl diphosphate (GGDP), and solanesyl diphosphate (SDP) used for
tocopherol, tocotrienol, and PQ-9 synthesis. Dashed arrows represent multiple steps.
Enzymes involved are depicted as white numbers in black circles: 1. l-deoxy-D-xylulose-S-
P (DXP) synthase; 2. DXP reductoisomerase; 3. 2-C-methyl-D-erythritol (MEP)
cytidyltransferase; 4. 4-(cytidine-5’-diphospho)-2-C-methyl-D-erythritol (CPME) kinase;
5. 2-C-methyl- -D- erythitol- 2 ,4-cyclodiphosphate (MECP) synthase; 6. MECP reductase 7.
(E)-4- -hydroxy-3-methyl-but-2 -enyl diphosphate (HMBPP) reductase; 8. IPP/DMAPP
isomerase; 9. solanesyl diphosphate (SDP) synthase;10.geranylgeranyl diphosphate
(GGDP) synthase 1; ll. GGDP reductase; 12. prenol: ATP kinase; 13. chlorophyllase.
Additional abbreviations: 2-phospho-4-(cytidine-5’-diphospho)-2-C—methyl-D-erythritol
(PCPME)

25

show an albino phenotype as a result of impaired chlorophyll and carotenoid biosynthesis
(Mandel et al., 1996; Estevez et al., 2000). The CLAI gene is expressed in virtually all
tissues, but the highest CLAI expression levels are observed in young developing tissues
(Estevez et al., 2000).

DXP reductoisomerase catalyzes the molecular rearrangement of DXP and a
reduction of the resulting unstable intermediate 2-C-methyl-D-erythrose-4-phosphate
(MESP) to yield 2-C-methyl—D-erythritol-4-phosphate (MEP) (Takahashi et al., 1998).
Similar to the phenotype of Arabidopsis claI mutants, peppermint transgenic plants with
low DXP reductoisomerase expression due to cosuppression of the endogenous DXPR
gene were small and chlorotic, indicating that DXP reductoisomerase is also essential for
normal plant growth and development (Mahmoud and Croteau, 2001 ). The next enzyme in
IPP biosynthesis, MEP cytidyltransferase, is involved in the formation of 4-(cytidine-5’-
diphospho)-2-C-methy1-D—erythritol (CPME) (Rohdich et al., 1999; Rohdich et al., 2000a).
CPME is subsequently phosphorylated by CPME kinase to form 2-phospho-4-(cytidine—
5’-diphospho)-2-C-methyl-D-erythritol (PCPME) (Luttgen et al., 2000; Rohdich et al.,
2000b). 2-C-methyl-D-erythitol-2,4-cyclodiphosphate (MECP) synthase catalyzes the
conversion of PCPME to MECP (Herz et al., 2000).

In bacteria, MECP is converted to IPP and dimethylallyl diphosphate (DMAPP) in
two steps by the reductases chE and Lth through the intermediate (E)-4-hydroxy-3-
methyl-but-Z-enyl diphosphate (HMBPP) (Altincicek et al., 2002). In plants, these
enzymes remain to be identified. NMR studies have demonstrated that IPP synthesis
precedes DMAPP synthesis in plants (Arigoni et al., 1999). Similar to the mevalonate
pathway, IPP isomerase is thought to catalyze a reversible conversion of IPP to DMAPP in
plants, but branching at HMBPP may also occur in bacteria (Altincicek et al., 2002). The
genome of the cyanobacterium Synechacystis sp. PCC 6803 lacks any apparent
homologues of IPP isomerase and it is not clear whether this enzyme is absent from this

organism or structurally distinct from the known IPP isomerases and therefore

26

unrecognizable in BLAST searches (Ershow et al., 2000). The steps subsequent to IPP
synthesis involve a family of related prenyl diphosphate (prenyl-DP) synthases (Chen et al.,
1994). In geranylgeranyl diphosphate (GGDP) synthesis, chloroplastic GGDP synthase 1
combines one molecule of DMAPP with three molecules of IPP in a head-to-tail manner
(Okada et al., 2000). GGDP can be used directly as a prenyl—DP precursor in tocotrienol
biosynthesis in seeds of monocots and gymnosperms or it can serve as a substrate in SDP
and PQ-9 synthesis (Lichtenthaler et al., 1997a). Alternatively, GGDP can be reduced to
PDP by a multifunctional GGDP reductase (GGDR) identified in cyanobacteria and plants
(Soll et al., 1983; Addlesee et al., 1996; Keller et al., 1998; Addlesee and Hunter, 1999). In
chloroplasts, PDP can also be formed from phosphorylation of phytol with ATP, which is

catalyzed by an unknown soluble kinase present in the stroma (8011 et al., 1984).

Tocopherol, Tocotrienol, And Plastoquinone Biosynthetic Pathways

Approximately thirty years ago, radiotracer studies using isolated chloroplasts and
seedlings of various plants delineated possible tocopherol, tocotrienol, and plastoquinone
biosynthetic intermediates and enzymatic activities (Threlfall and Whistance, 1971; Schulze-
Siebert et al., 1987). The tocopherol, tocotrienol, and plastoquinone biosynthetic pathways
are shown in Figure 5. The first committed step in tocopherol, tocotrienol, and
plastoquinone synthesis involves the condensation of the aromatic precursor HGA and
PDP, GGDP, or SDP catalyzed by homogentisate prenyltransferase/decarboxylases
(Threlfall and Whistance, 1971; Hutson and Threlfall, 1980; 8011 et al., 1980; Schulze-
Siebert et al., 1987; Collakova and DellaPenna, 2001). The products of these prenylation
reactions are C02, inorganic pyrophosphate, and the first prenquuinol intermediates in the
respective tocopherol and plastoquinone pathways leading to 2-methyl-6-phytyl-l,4-
benzoquinol (MPBQ) and 2-demethylplastoquinol-9 (DMPQ-9), respectively (Hutson and

27

"WM"

 

SDP
OH
: 'CHZCOO- :H Hl" ‘T |;:AM ::: E’ T "91'1
0" co +PP
HGA 2 i DIIIIPCIH2 .9 pm.2 .9
FPO/\)\ R PDP or GGDP 0
if Wu
co2 +PP, 0 PQ-9 9
V SAM
HO HO
1
MPBQ DMPBQ

 

? ? H
Ho Ho ”Ma for PDP

; :O|\R : ; :OI\R H
Ian-Ms forGGDP

5-tocopherol y-tocopherol

4! SA" s”?

B-tocopherol a-tocopherol

 

 

 

Figure 5. Tocopherol, tocotrienol, and plastoquinone-9 biosynthesis in cyanobacteria and
plants. Enzymes are shown as numbers in black circles. l. homogentisate (HGA)
prenyltransferase(s); 2. HGA solanesyltransferase; 3. 2-methyl-6-prenyl-1,4-benzoquinol
(MPBQ) methyltransferase(s) (MT); 4. tocopherol cyclase (TC); 5. y-tocopherol
methyltransferase (y—TMT). Other abbreviations: solanesyl diphosphate (SDP); phytyl
diphosphate (PDP), geranylgeranyl diphosphate (GGDP); demethyl plastoquinol-9
(DMPQHz); plastoquinol-9 (PQHZ); plastoquinone-9 (PQ-9); 2,3-dimethyl-6-prenyl-1,4—
benzoquinol (DMPBQ); S-adenosyl-L-methionine (SAM)

Threlfall, 1980; 8011 et al., 1980; Marshall et al., 1985; Collakova and DellaPenna, 2001). In
tocotrienol synthesis, a similar reaction would yield 2-methyl-6-geranylgeranyl-1,4-
benzoquinol (MGBQ) (Threlfall and Whistance, 1971).

Biochemical studies using isolated chloroplasts and radiolabeled precursors of
tocopherol and plastoquinone biosynthesis could not distinguish between separate
prenyltransferase activities (Schulze-Siebert et al., 1987). Phytylation of HGA occurred
primarily in the envelopes, whereas prenylation of HGA with SDP took place both in the
envelopes and thylakoids, suggesting that separate prenyltransferase activities might be
responsible for tocopherol and plastoquinone synthesis (8011 et al., 1980). However, these
differences could also be due to different availability of the endogenous substrates in the
two membranes. Arabidopsis homogentisate phytyl/prenyltransferase was postulated to be
encoded by the PDSZ gene because disruption of PDS2 led to the same albino phenotype as
the pdsI mutation (Norris et al., 1995). The Arabidopsis pdsZ mutants did not accumulate
detectable levels of tocopherols, PQ-9, chlorophylls, or carotenoids. Norris et a1. (1995)
proposed that the PDS2 gene product may represent a homogentisate phytyl/
prenyltransferase involved in tocopherol and PQ-9 biosynthesis. Chapter 1 reports cloning
and characterization of the gene encoding homogentisate phytyltransferase in Synechacystis
sp. PCC 6803 and Arabidopsis thaliana. This work was also published in Plant
Physiology (Collakova and DellaPenna, 2001). Genetic and biochemical data presented in
Chapter I suggested that homogentisate phytyltransferase (HPT) is involved only in
tocopherol biosynthesis and that a separate homogentisate solanesyltransferase (HST) must
exist in these two photosynthetic organisms. Similarly, tocopherol and tocotrienol synthesis
may involve separate prenyltransferase activities utilizing PDP and GGDP, respectively, to
prenylate HGA in monocots, which accumulate tocotrienols. Many plant species contain
several different genes encoding putative prenyltransferases that may be involved in
tocopherol or tocotrienol synthesis (DellaPenna et al., 2000). However, I have demonstrated

that the Svnechacystis HPT can utilize both PDP and GGDP, suggesting that a single

29

enzyme can contain both prenyltransferase activities (Collakova and DellaPenna, 2001).
This observation was surprising because Synechacystis sp. PCC 6803 does not accumulate
any detectable tocotrienol levels (Collakova and DellaPenna, 2001 ). More research is
required to address this question in plants.

Some of the subsequent steps in tocopherol, tocotrienol, and plastoquinone
synthesis may also involve the same types of enzymes and intermediates. The three
prenquuinols formed by the HGA prenyltransferases can be methylated to 2,3-dimethyl-6-
phytyl-1,4-benzoquinol (DMPBQ), 2,3-dimethyl-6-geranylgeranyl-l,4-benzoquinol
(DMGBQ), and plastoquinol-9, respectively (8011 and Schult, 1979; 8011 et al., 1980;
Schultz et al., 1985; Shintani et al., 2002). These reactions may be catalyzed by a single
methyltransferase (MT), which utilizes S-adenosyl-L-methionine (SAM) as a methyl donor.
In Synechocystis sp. PCC 6803, the MT encoded by the ORF SLL0418 can efficiently
methylate both MPBQ and DMPQ-9 in vitro, indicating that the methylation of these two
prenquuinols may be performed by the single enzyme MPBQ/DMPQ-9 MT (Shintani et
al., 2002). SLL0418 disruption mutants showed altered tocopherol composition and B-
tocopherol in addition to (rt-tocopherol, the predominant tocopherol form in Synechacystis
sp. PCC 6803, was detected. Consistent with the hypothesis that SLL0418 is also involved
in PQ-9 synthesis in this cyanobacterium, only partial 8110418 disruption mutants were
isolated presumably because PQ-9 is essential for survival of photosynthetic organisms
(Shintani et al., 2002).

Spinach chloroplasts can catalyze all three methylation reactions when radiolabeled
SAM and the corresponding 2-methylprenquuinols are used as substrates (S011 and Schult,
1979; 8011 et al., 1980; Schultz et al., 1985). Based on studies with purified chloroplast
envelopes and thylakoids, the prenquuinol methylation reaction in tocopherol synthesis has
been associated with the envelope membranes, while that in PQ-9 synthesis occured in both
envelopes and thylakoids, suggesting that separate MT activities may operate in plant

tocopherol and PQ-9 synthesis (8011 et al., 1980; Schultz et al., 1985). Geranylgeranylated

30

precursors were preferred substrates relative to the phytylated ones for the
methyltransferase(s) present in spinach chloroplasts (8011 and Schultz, 1979). Because no
tocotrienols have been detected in spinach, it is possible that GGDP is not used in viva to
prenylate HGA or that any newly formed tocotrienol precursors are efficiently reduced to
tocopherol precursors (Collakova and DellaPenna, 2001). These questions are further
discussed in Chapter 1.

The next steps in tocopherol and tocotrienol synthesis are catalyzed by tocopherol
cyclase (TC), a relatively non-specific enzyme that can utilize both single and double
methylated tocopherol intermediates and both phytylated and geranylgeranylated
intermediates as substrates (Stocker et al., 1996; Porfirova et al., 2002). Cyclization of
MPBQ and DMPBQ yields 5- and y-tocopherol, respectively, whereas cyclization of
MGBQ and DMGBQ results in the formation of 8- and y—tocotrienols, respectively. TC
activity has been shown in Anabeana variabilis and bell pepper chromoplasts (Stocker et
al., 1993; Stocker etal., 1994) and TC has been cloned in Synechacystr‘s sp. PCC 6083,
Arabidopsis, and maize (Porfirova et al., 2002; Sattler and DellaPenna, 2003). TC is the
best studied enzyme of the tocopherol and tocotrienol biosynthetic pathway in terms of
reaction mechanism and substrate specificity. The prenquuinol enters the active site of the
enzyme with its tail (Stocker et al., 1996). The chromanol ring is formed by a nucleophillic
attack of carbon 1 phenolic hydroxyl group on the protonated double bond of the phytyl tail
(Stocker et al., 1994). Tocopherol cyclase requires prenquuinol substrates, a prenyl tail in
the 6th position on the benzoquinol ring, the double bond in the (E) configuration, and
prenyl tails longer than 10 carbons (Stocker et al., 1993; Stocker et al., 1994; Stocker et al.,
1996). The last step of tocopherol synthesis involves methylation of y— and 5-tocopherols
to a— and B—tocopherols, respectively. In Arabidopsis and Synechacystis, these reactions are
catalyzed by y—tocopherol methyltransferase (y-TMT), a membrane-bound enzyme that

utilizes SAM as a methyl donor and requires an intact chromanol ring with carbon 5

31

available for methylation (Shintani and DellaPenna, 1998). Unfortunately, tocotrienols have

not been used as substrates to test y-TMT substrate specificity.

TOCOPHEROL FUNCTION IN PLANTS

Antioxidant Function

Chloroplasts are the major source of 'O2 and Oz- in plant cells (Mittler, 2002). In

chloroplasts, 'O2 originates by the interaction of ground-state molecular oxygen (3O2 -
triplet oxygen) and excited triplet chlorophyll during photosynthesis. During this reaction,
the energy is exchanged between these two molecules and ground-state singlet chlorophyll
and excited '02 are formed (Melis, 1999; Mittler, 2002). Excess light energy may result in
generation of excited triplet chlorophyll when the photosynthetic electron transport is

overreduced. On the other hand, ferredoxin and other components of the photosynthetic

electron transport are generators of Oz- radicals (Mittler, 2002). In chloroplasts,

tocopherols are most likely involved in Oz- scavenging rather than directly in 102

quenching, which is efficiently performed by carotenoids and superoxide dismutase
(Y amauchi and Matsushita, 1979a; Michalski and Kaniuga, 1981). Carotenoids are prone
to degradation by the products of lipid peroxidation and one mechanism, by which
tocopherols can indirectly reduce 102 levels, is to prevent carotenoid degradation by
inhibiting lipid peroxidation. In isolated spinach chloroplasts, visible light-induced lipid
peroxidation was accompanied by (rt-tocopherol disappearance and a-tocopheryl quinone
formation, suggesting the active oxidation of (rt-tocopherol (Y amauchi and Matsushita,
1979a). Illumination of detached cold-treated tomato leaves led to a several-fold increase of
ot-tocopherol and (rt-tocopheryl quinone levels relative to leaves stored at room temperature

(Michalski and Kaniuga, 1981). Because cold-treated tomato leaves contain inactive

32

superoxide dismutase, another important scavenger of 02' in plant chloroplasts, 02'

oxidized (rt-tocopherol to form (rt-tocopheryl quinone to a greater extent in cold-treated
leaves than in the fresh leaves containing active superoxide dismutase. Addition of
exogenous a—tocopherol reduced malondialdehyde formation in isolated chloroplasts,
suggesting that a-tocopherol is involved in protection of membrane lipids from peroxidation
in plant chloroplasts (Michalski and Kaniuga, 1981).

Interaction of tocopherols with other antioxidants may play an important role in
protecting plants from photooxidative damage. F inckh and Kunert (1985) subjected nine
different plant species to an herbicide-induced lipid peroxidation. A decrease in oxidative
damage correlated well with high tocopherol and ascorbate levels. Mustard and sicklepod
plants, which contained tocopherols and ascorbate in the ratio of 1 to 10 - 15 Showed the
lowest degree of cell damage. Although action of other protective mechanisms could not be
excluded, the authors suggested that tocopherols were recycled by ascorbate because the
plants that showed high tocopherol and ascorbate levels, as well as a certain ratio of these
two antioxidants, were most resistant to oxidative damage (Finckh and Kunert, 1985). The
Arabidopsis vtc-I mutant, which has levels of ascorbate reduced to ~ 60% of the
corresponding wild type levels, was more sensitive to lipid peroxidation and accumulated
less (rt-tocopherol and B—carotene than wild type during drought stress (Munne-Bosch and
Alegre, 2002b). Ascorbate, which is present in the cytosol, is unlikely to scavenge ROS
within membranes and the reduced a-tocopherol and B-carotene levels in the vtc-I mutant
can be caused by impaired recycling (Munne-Bosch and Alegre, 2002b). Because
ascorbate is required as a cofactor for the tocopherol biosynthetic enzyme HPPD (Fiedler et
al., 1982), low tocopherol levels in the vtc—I mutant may also be a result of impaired
tocopherol biosynthesis. Because tocopherols protect B-carotene by scavenging lipid
peroxy radicals (Y amauchi and Matsushita, 1979a), reduced B—carotene levels in this mutant

may be due to a secondary effect of insufficient tocopherol levels. Direct evidence that

33

ascorbate plays a key role in recycling of lipid-soluble antioxidants such as a-tocopherol
and B-carotene is still lacking.

Plants have developed numerous antioxidant systems to protect them from oxidative
damage. Oxidative stress is associated with up-regulation of multiple antioxidants
(carotenoids, ascorbate, glutathione, or anthocyanins) and ROS detoxifying enzymes
(glutathione reductase, ascorbate peroxidase, catalase, or superoxide dismutase) (Leipner et
al., 1999; Ye and Gressel, 2000). Tocopherol levels also increase in plants exposed to
various abiotic stress conditions such as high light, low temperature, drought, and high
salinity (Munne-Bosch and Alegre, 2002a). Cultivars showing increased tolerance to stress
accumulate increased (rt-tocopherol levels relative to the stress-sensitive cultivars (Gossett et
al., 1994; Streb et al., 1998; Leipner et al., 1999). Mountain plants growing at high altitudes
are exposed to low temperatures and high irradiance. Several alpine plant species growing
at high altitudes showed increased tocopherol levels relative to their counterparts growing at
low altitudes or lowland plants (Wildi and Lutz, 1996; Streb et al., 1998). High light and/or
low temperatures led to an increase in a—tocopherol levels in other plants including
Arabidopsis and maize (Fryer et al., 1998; Havaux and Niyogi, 1999; Leipner et al., 1999;
Havaux et al., 2000; Havaux and Kloppstech, 2001). Drought and/or high light stress also
resulted in a several-fold increase in a—tocopherol levels in rosemary and wheat leaves
(Bartoli et al., 1999; Munne-Bosch and Alegre, 2000). It appears that various types of
oxidative stress lead to an increase in or-tocopherol levels in a variety of plants, which, along
with other antioxidant systems, may protect those plants from oxidative damage and death.
However, this correlation has not been shown in all studies.

In some instances, (rt-tocopherol levels decreased or did not change with oxidative
stress (Gossett et al., 1994; Havaux et al., 2000; Kranner et al., 2002). These observations
can be explained in part by the progressive degradation of tocopherols, which may occur
when oxidative stress becomes detrimental to the plant. a—Tocopherol levels gradually

increased in Arabidopsis leaves exposed to high light intensities from 250 to 1,000 IE, but

34

rapidly decreased when the plants were exposed to 1,500 pE (Havaux et al., 2000). An
eight-month desiccation of the resurrection plant Myrathamnus flabellifalia resulted in a
nearly complete depletion of the (rt-tocopherol levels as well as other antioxidants, which
were recovered upon rehydration (Kranner et al., 2002). It is also possible that the stress
conditions used in some studies were not high enough to induce tocopherol synthesis in
some plant species or that these plants use other components of the antioxidant system to
cope with particular types of stress. Cotton plants treated with 150 mM NaCl did not
accumulate increased tocopherol levels, but showed increased glutathione reductase,
peroxidase, and superoxide dismutase activities (Gossett et al., 1994).

Abiotic stress is not the only factor affecting tocopherols in plants. Tocopherol
levels and composition change during different stages of plant growth and development
(Franzen and Haas, 1991; Franzen et al., 1991; Molina-Torres and Martinez, 1991,
Tramontano et al., 1992). These changes may contribute to optimal plant growth and
development by controlling the levels of reactive oxygen species and lipid peroxidation.
During seed germination, light induces the synthesis of various components of
photosynthetic apparatus including chlorophylls, carotenoids, and plastoquinone in
developing chloroplasts, which are becoming photosynthetically active and generate high
ROS levels (Lichtenthaler, 1969). Light also induced tocopherol accumulation in
developing barley seedlings, thus most likely protecting these chloroplasts from oxidative
damage (Lichtenthaler, 1969). Tocopherol levels and composition vary among seeds and
seedlings of different plant species. In general, the tocopherol, which predominates in dry
seeds, is also initially found in germinating seeds and primary leaves or needles of
angiosperrns or gymnosperms (Franzen and Haas, 1991; Franzen et al., 1991). The levels
of a-tocopherol, the major tocopherol in gymnosperm seeds, gradually increased in
developing Norway spruce seedlings (Franzen and Haas, 1991; Franzen et al., 1991). y-
Tocopherol was the major tocopherol present in the seeds and primary leaves of l-, 2-, and

3-day old monocot and dicot seedlings (Franzen and Haas, 1991). In these seedlings, the

35

gradual decrease in y-tocopherol levels was accompanied by an increase in the levels of a-
tocopherol (Franzen and Haas, 1991), the major tocopherol present in mature monocot and
dicot leaves (Piironen et al., 1986; Molina-Torres and Martinez, 1991; Tramontano et al.,
1992; Shintani and DellaPenna, 1998).

Aging and senescence are programmed stress-inducing processes involving
chlorophyll, nucleic acid, protein, and lipid degradation and nutrient mobilization from
senescing parts of plants to developing leaves (Buchanan-Wollaston, 1997). These
processes are also associated with an increase in lipid peroxidation, a decrease in
photosynthetic activity and chlorophyll levels, and changes in antioxidant status (Munne-
Bosch et al., 2001; Munne-Bosch and Alegre, 2002c). (rt-Tocopherol accumulation
increased with both leaf and plant age in soybean, pinto bean, and Xanthium strumarium
leaves (Tramontano et al., 1992; Molina-Torres and Martinez, 1991). In addition, an
increase in y—tocopherol levels with age was also observed in soybean plants (Tramontano et
al., 1992). Rise et a1. (1989) reported that (Ir-tocopherol levels increased in the senescing
leaves in a variety of plant species, which was also correlated with elevated chlorophyll
degradation. It has been postulated that the phytol tail released by chlorophyllase during
chlorophyll degradation is phosphorylated in chloroplasts and the resulting phytyl
diphosphate used for (rt-tocopherol synthesis in senescing leaves (801] et al., 1980; Rise et
al., 1989). Other studies Showed that (rt-tocopherol levels decreased during aging and
senescence in several Mediterranean plant species (Munne-Bosch et al., 2001; Munne-
Bosch and Alegre, 2002c). These Mediterranean field-grown plants were also subjected to
a combination of high light and drought stress and the reduced a-tocopherol levels observed
were likely due to increased tocopherol degradation.

Because plants regulate tocopherol levels and composition as a response to oxidative
stress, specific tocopherols may play important roles in controlling oxidative damage during
plant grth and development. Several cyanobacterial and plant mutants or transgenic

plants showing altered tocopherol levels have been isolated and characterized (Henry et al.,

36

1986; Nonis et al., 1995; Tanaka et al., 1999). Because some mutants were also impaired in
the synthesis of other vitally important compounds, photobleached phenotypes observed in
these organisms could not be ascribed to the lack of tocopherols. Geranylgeranyl
diphosphate reductase (GGDR) is a multifunctional enzyme involved in the conversion of
GGDP to PDP, a substrate used for tocopherol, phylloquinone, and chlorophyll
biosynthesis (Soll et al., 1983; Addlesee et al., 1996; Keller et al., 1998; Addlesee and
Hunter, 1999). Besides tocopherols, a Scenedesmus mutant lacking GGDR activity or
tobacco GGDR antisense plants were unable to synthesize chlorophylls and phylloquinone
(Henry et al., 1986; Tanaka et al., 1999). Similarly, the Arabidopsis PDS] gene encodes p-
hydroxyphenylpyruvate dioxygenase (HPPD), an enzyme involved in tocopherol and
plastoquinone biosynthesis (Norris et al., 1995). The homozygous pds] mutants in
Arabidopsis was seedling-lethal and required a carbon source for its growth and completely
lacked tocopherols, plastoquinone-9, chlorophylls, and carotenoids (Norris et al., 1995).

In Synechacystis sp. PCC 6083, disruption of HPPD or HPT completely abolished
tocopherol synthesis, while the levels of other prenyllipids were unaffected (Collakova and
DellaPenna, 2001; Dahnhardt et al., 2002; Savidge et al., 2002). The absence of tocopherols
had no effects on the growth or photosynthesis under normal or high-light conditions in
these mutants (Collakova and DellaPenna, 2001; Dahnhardt et al., 2002; Savidge et al.,
2002). Some cyanobacterial species naturally lack tocopherols as indicated by the absence
of several tocopherol biosynthetic genes in their genomes, which suggests that tocopherols
are not required for normal growth in these organisms (Dasilva and Jensen, 1971; Thomas
et al., 1998; Sattler et al., 2003). However, tocopherols may protect photosynthetic
organisms from specific types of stress. Experiments using the Synechacystis and
Arabidopsis HPT mutants are underway in Dr. Dean DellaPenna’s laboratory to address

this question.

37

Roles in Signaling

In animals, eicosanoid Signaling is associated with the phospholipase AZ-dependent
release of arachidonic acid, a precursor in the synthesis of various oxylipins involved in
inflammatory processes (Balsinde et al., 1999). Tocopherols can affect phospholipase A2
activity and regulate oxylipin production in animals (Tran et al., 1996; Chan et al., 1998;
Grau and Ortiz, 1998). In plants, linolenic acid is cleaved off by an analogous
phospholipase and used for the synthesis of phyto-oxylipins including jasmonate (JA) in
octadecanoid signal transduction pathway (Schaller, 2001; Blee, 2002; Howe and
Schilmiller, 2002). Other enzymes downstream of phospholipases, which include
cytochromeS P450, lipoxygenases, and other oxygenases, are also functionally similar in
animals and plants, indicating a resemblance between animal eicosanoid and plant
octadecanoid signaling (Blee, 2002). Although there is no evidence that tocopherols may
directly modulate enzyme activities involved in plant octadecanoid signaling, the possibility
exists that they may affect plant signal transduction pathways by mechanisms involving
their antioxidant properties.

ROS are by-products of aerobic metabolism considered to be harmful to
chloroplasts. ROS levels or production are controlled by a variety of means, including
antioxidant levels. However, animals and plants also produce ROS as signaling molecules
to regulate various cellular processes including stress responses, plant-pathogen interaction,
and programmed cell death (Mittler, 2002). Tocopherols may affect some of these
processes by controlling ROS and lipid hydroperoxide levels in plants (Munne-Bosch and
Alegre, 2002a). Lipid peroxidation propagation is associated with the formation of a lipid
peroxy radical, which can interact with another PUF A to generate a lipid hydroperoxide and
another lipid peroxy radical (Figure 2). For example, if the target molecule is linolenic acid,
the resulting lipid hydroperoxide formed during lipid peroxidation will be 13-hydroperoxy-

9,1 1,15-octadecatrienoic acid (13-HPOD), which is also a product of l3-lipoxygenase and a

38

substrate for several enzymes including allene oxide synthase involved in IA biosynthesis
(Schaller, 2001; Howe and Schilmiller, 2002; Blee, 2002). Uncontrolled propagation of
lipid peroxidation could lead to a massive JA and other oxylipin synthesis during oxidative
stress. As chain-breaking antioxidants, tocopherols can scavenge lipid peroxy radicals to
form lipid hydroperoxides such as l3-HPOD. Therefore, by scavenging l3-HPOD
radicals and by controlling the levels of lipid peroxidation, tocopherols may interfere with
the octadecanoid pathway signaling by inhibiting synthesis of jasmonate and other

oxylipins.

Roles in Photosynthesis

The oxidation products of (rt-tocopherol, (rt-tocopheryl quinol and quinone (Figure
2) have been suggested to be involved in photosynthetic electron transport (Lichtenthaler,
1993; Kruk and Strzalka, 1995). a-Tocopheryl quinol/quinone can be oxidized/reduced to
the corresponding quinone/quinol and this redox state change may allow the molecules to
perform similar roles as ubiquinones and plastoquinones in electron transport. Several in
vitro studies have suggested that or-tocopheryl quinone may participate as an electron carrier
in cyclic electron transport of photosystem 11 (PS 11) in chloroplasts (Michalski and
Kaniuga, 1981; Kruk et al., 2000). Stable picrylhydrazyl radicals can Specifically oxidize
and extract a-tocopherol present in chloroplasts. (rt-Tocopherol oxidation with the
picrylhydrazyl radicals resulted in an inhibition of PS 11 activity, which was partially
reversed by the addition of a—tocopherol to isolated chloroplasts (Michalski and Kaniuga,
1981). It was proposed that at least two (rt-tocopherol pools exist independently in
chloroplasts and that only ~ 70% of (rt-tocopherol may function in PS 11 electron transport
(Michalski and Kaniuga, 1981).

Experiments with the Scenedesmus abliquus GGDP reductase mutant suggested

that (rt-tocopherol and its quinone may be involved in the cyclic electron transport chain

39

around PS 11 when the PS 11 is over-reduced (Kruk et al., 2000). Besides tocopherols, this
mutant also lacks the high-potential form of cytochrome b-559 (cyt b-559 HP), which
participates in the cyclic electron transport in PS 11 (Henry et al., 1986; Kruk et al., 2000).
Both PQ-A and (rt-tocopherol reduced b-559 HP, while (rt-tocopheryl quinone oxidized the
reduced form of this cytochrome in wild type Scenedesmus abliquus, but not in the mutant,
which suggests a specific interaction between these prenyllipids and the cyt b-559 HP. or-
Tocopheryl quinone also efficiently quenched chlorophyll fluorescence in thylakoid
membranes isolated from both wild type and mutant, suggesting that this firnction was
independent of the cyt b-559 HP and that (rt-tocopheryl quinone interacts with another
component of the cyclic electron transport besides cyt b-559 HP (Kruk et al., 2000).
Although all these studies suggest that (rt-tocopherol and a-tocopheryl quinone may be
involved in cyclic electron transport, no studies have been performed to confirm whether this

function exists in viva.

NUTRITIONAL GENOMICS

Natural Sources of Dietary Vitamin E

As mentioned before, humans cannot synthesize tocopherols and must acquire
vitamin E from their diets, mainly from plants and their products (Brigelius-Flohe and
Traber, 1999; Bramley et al., 2000). Plant seed oils are the major source of dietary vitamin
E in the average American diet (Eitenmiller, 1997). Soybean oil, the most commonly
consumed plant oil in the world, contains up to 115 mg of total tocopherols per 100 g oil
(Eitenmiller, 1997; Bramley et al., 2000). However, the predominant tocopherol in soybean
oil is y-tocopherol, which has only 10% the biological activity of a—tocopherol. Therefore,
the maximal vitamin E activity of soybean oil is relatively low, only 20 mg a-TE per 100 g

oil (Eitenmiller, 1997; Bramley et al., 2000). Seeds of other plants such as canola, com, and

40

Arabidopsis also accumulate predominantly y-tocopherol and their vitamin B activity is also
very low (Eitenmiller, 1997; Bramley et al., 2000; Goffman and Mollers, 2000; Goffman
and Bohme, 2001). Oils with the highest percentage and content of a-tocopherol have the
highest vitamin B activity. These oils include safflower, sunflower, and wheat germ oil,
vitamin E activities of which varies between 41 and 174 mg a—TE per 100 g of oil,
respectively (Eitenmiller, 1997). In most vegetables, berries, and fruits, (rt-tocopherol is the
predominant form of vitamin E, but its levels are very low, between 1 and 6 mg (rt-tocopherol
per 100 g fresh weight (Piironen et al., 1986; Ching and Mohamed, 2001).

Currently, the recommended dietary intakes for a-tocopherol are 15 mg (35 mol)
of RRR-a-tocopherol daily (Food and Nutrition Board, 2000). The maximal vitamin E
activities of spinach leaves, soybean oil, sunflower oil, and wheat-germ oil are 3.7, 20, 63,
and 174 mg a-TE per 100 g fresh weight or oil, respectively (Ching and Mohamed, 2001;
Eitennriller, 1997; Bramley et al., 2000). Based on these values, it can be estimated that one
should daily consume one of the following to meet the dietary requirements for or-
tocopherol: 405 g fresh spinach leaves, 75 g soybean oil, 24 g sunflower oil, or 8.6 g wheat-
germ oil. Pharmacological doses of up to 800 IU (537 mg of a-TE) of RRR—a—tocopherol
or its acetate were administered daily to the patients suffering from atherosclerosis in the
heart disease prevention trials CHAOS and SPACE (Stephens et al., 1996; Boaz et al.,
2000), while the upper tolerable limit for a—tocopherol is 1000 mg per day (Food and
Nutrition Board, 2000). Even the best natural (rt-tocopherol sources are clearly insufficient
to provide these extremely high doses of vitamin E. The limitation of a-tocopherol in the
average American diet may be overcome through metabolic engineering of tocopherol

biosynthesis in plants.

41

Regulation And Metabolic Engineering of Tocopherol Biosynthesis in Plants

Enzymes of tocopherol biosynthetic pathway can be divided into two groups. The
first group includes enzymes that may be involved in regulating flux through the pathway
and subsequently total tocopherol levels. This group contains some enzymes committed to
the tocopherol biosynthetic pathway as well as the enzymes of the Shikimate and non-
mevalonate pathways. The second group includes only three enzymes determining
tocopherol composition, namely TC, MPBQ MT and y—TMT (Grusak and DellaPenna,
1999). It is desirable both to increase total tocopherol levels and to alter tocopherol
composition in plants towards (rt-tocopherol, the most potent form of vitamin E. In order to
genetically manipulate plants to produce elevated a—tocopherol levels, it is important to
understand the regulation of tocopherol biosynthetic pathway, identify enzymatic steps
limiting tocopherol synthesis, and clone and study the relevant genes. The regulation of
tocopherol biosynthesis appears to be a very complex process and may involve several
enzymes of the pathway.

In plants, the Shikimate pathway (Figure 2) provides chorismate, an aromatic
precursor in the biosynthesis of aromatic amino acids, tocopherols, plastoquinones,
phylloquinones, salicylic acid and other phenolics involved in plant pathogen defense and
cell wall lignification (Herrrnan, 1995; Herrman and Weaver, 1999; Diaz et al., 2001). This
pathway is probably regulated at the transcriptional level in plants and can be globally up-
regulated during biotic and abiotic stresses (Gorlach et al., 1995; Jones et al., 1995;
Bischoff et al., 1996; Batz et al., 1998; Diaz and Merino, 1998; Diaz et al., 2001).
Unfortunately, the effects of the Shikimate pathway up-regulation on tocopherol levels was
not investigated in these reports. However, at least two enzymes downstream of chorismate,
TAT and HPPD, have been shown to regulate HGA production and tocopherol levels in
plants (Sandorf and Hollander-Czytko, 2002; Lopukhina et al., 2001; Klebler-Janke and
Krupinska, 1997; Tsegaye etal., 2002). In plant chloroplasts, TAT catalyzes transamination

42

of tyrosine to HPP and its activity can be induced by a variety of factors including
coronatine, wounding, various octadecanoids, high light, UV light, and the herbicide
oxyfluorfen in wild type Arabidopsis (Lopukhina et al., 2001; Sandorf and Hollander-
Czytko, 2002). Elevated a- and y—tocopherol levels were associated with the increase in
TAT protein levels induced by all these elicitors (Sandorf and Hollander-Czytko, 2002). In
the Arabidopsis ddeI mutant impaired in JA synthesis due to a block in the octadecanoid
enzyme 13-OPDA reductase 3, TAT levels were lower and tocopherol levels were higher
than in wild type. The high total tocopherol content in the non-stressed ddel mutant did not
correlate with the low TAT levels, which suggests that TAT is not the only factor affecting
regulation of tocopherol synthesis (Sandorf and Hollander-Czytko, 2002).

HPPD activity may also be involved in regulation of tocopherol synthesis because
HPPD mRN A levels were elevated during stress senescence in barley leaves (Klebler-Janke
and Krupinska, 1997). HPPD catalyzes the conversion of HPP to HGA and its gene has
been cloned and characterized from several plant species (Garcia et al., 1997; Norris et al.,
1998; Klebler-Janke and Krupinska, 1997; Garcia et al., 1999). The HGA pool may limit
tocopherol synthesis because the addition of HGA to safflower cultures moderately
stimulated tocopherol synthesis in spite of a rapid HGA oxidation in the media (Furuya et
al., 1987). Thus, HPPD activity may also limit tocopherol synthesis in other plants
(Tsegaye et al., 2002). Constitutive HPPD overexpressing in Arabidopsis resulted in a 10-
fold increase in HPPD specific activity, but only a 10 to 30 % increase in total tocopherol
levels in both leaves and seeds. The minor effect of HPPD overexpression on tocopherol
levels might be due to enhanced degradation of HGA in transgenics. It is also possible that
HPPD does not present a significant limitation for tocopherol biosynthesis (Tsegaye et al.,
2002)

In plastids, the non-mevalonate pathway provides PDP, which is used as a substrate
in tocopherol biosynthesis (Rohmer et al., 1993; Schwender et al., 1996; Lichtenthaler et al.,
1997b; Proteau, 1998). Feeding phytol alone to safflower cell cultures induced tocopherol

43

synthesis to even a higher extent than HGA, indicating that phytol pool is also limiting for
tocopherol biosynthesis (Furuya et al., 1987). Genes encoding some of the enzymes of the
non-mevalonate pathway providing the IPP pool for the plastidic isoprenoid synthesis have
been cloned and some of these genes overexpressed in plants (Lange et al., 1998; Estevez et
al., 2000; Estevez et al., 2001; Mahmoud and Croteau, 2001). Only the first enzyme of the
DXP pathway, DXP synthase, has been studied in terms of tocopherol biosynthesis
(Estevez et al., 2001). Constitutive overexpression of DXP synthase in Arabidopsis leaves
resulted in elevated levels of several plastidic isoprenoids. Tocopherol levels increased up to
2-fold in leaves of transgenics relative to wild type plants, indicating that DXP synthase
limits tocopherol synthesis in Arabidopsis chloroplasts (Estevez et al., 2001). Therefore,
tocopherol synthesis may be regulated in part by the availability of HGA and phytol or
PDP. In chloroplasts, these precursors are also used in the syntheses of other prenyllipids
such as plastoquinone, phylloquinone, and chlorophyll (Threlfall and Whistance, 1971;
Schulze-Siebert et al., 1987; Oster et al., 1997). Because of the involvement of HGA and
PDP in multiple biosynthetic pathways, the actual regulation most likely occurs at the level
of the next biosynthetic steps, the prenylation reactions that are catalyzed by structurally and
functionally related prenyltransferases.

HPT catalyzes the first committed step of tocopherol biosynthesis, the condensation
of HGA and PDP (Hutson and Threlfall, 1980; S011 et al., 1980; Marshall et al., 1985;
Collakova and DellaPenna, 2001; Savidge et al., 2002). A recent report of seed-specific
overexpression of HPT in Arabidopsis seeds demonstrated a 75% increase in total seed
tocopherol levels, suggesting that HPT activity is a limiting factor for tocopherol
biosynthesis in Arabidopsis seeds (Savidge et al., 2002). Because the seed-specific napin
promoter was used, the effects of HPT overexpression on tocopherol levels in leaves could
not be determined (Savidge et al., 2002). In Chapter 2 (Collakova and DellaPenna, 2003), I
described and discussed the effects of constitutive HPT overexpression on the levels of

tocopherols and other prenyllipids in Arabidopsis leaves and seeds. HPT overexpressers

(358::HPT1) showed up to a 100-fold increase in HPT mRNA levels and a lO-fold increase
in HPT activity relative to wild type. These increases in HPT expression resulted in a 4.4-
fold and 40% increase in total tocopherol levels in leaves and seeds, respectively. The levels
of other prenyllipids was not affected by increasing flux to the tocopherol pathway
(Collakova and DellaPenna, 2003).

In Chapter 3, I describe the use of high light stress to increase flux through the
tocopherol biosynthetic pathway. During high light stress, total tocopherol levels rapidly
increased in both wild type and 358::HPT] leaves. Stressed 3SS::HPTI leaves accumulated
elevated total tocopherol levels relative to the corresponding wild type leaves. The increase
in total tocopherol levels was most likely due to an up-regulation of HPT mRNA levels and
activity and increased expression of two other tocopherol-related genes, TAT and HPPD.
Enhanced tyrosine and chlorophyll degradation have been proposed to provide increased
pool of HGA and PDP that could be used to synthesize tocopherols in stressed plants
(Chapter 3).

The last three enzymes of the pathway are involved in the controlling tocopherol
composition in plants. Tocopherol cyclase catalyzes the formation of the first tocopherols,
y— and 6- tocopherols (Stocker et al., 1993; Stocker et al., 1994; Stocker et al., 1996;
Porfirova et al., 2002; Sattler et al., 2003). Based on the kinetic studies of HGA
incorporation in yellow bell pepper chromoplasts, TC does not seem to limit tocopherol
biosynthesis in chromoplasts (Arango and Heise, 1998). MPBQ MT methylated MPBQ to
DMPBQ and the corresponding gene has been cloned from Synechacystis ps. PCC 6803
(Shintani et al., 2002). The Arabidopsis MPBQ MT is currently being cloned and
characterized in Dr. DellaPenna’s laboratory. The Arabidopsis y—TMT gene has been
cloned and overexpressed in leaves and seeds (Shintani and DellaPenna, 1998). Wild type
Arabidopsis leaves accumulated predominantly (rt-tocopherol, while the major tocopherol
present in seeds was y—tocopherol. In Arabidopsis leaves, y—TMT overexpression had no

effect on total tocopherol levels or composition. In contrast, overexpression of y—TMT in

45

Arabidopsis seeds resulted in the conversion of more than 95% of the y-tocopherol to a-
tocopherol. Similar to leaves, total tocopherol content remained unchanged in the transgenic
seed. These results indicated that y—TMT was a key enzyme controlling seed tocopherol
composition, but had no effect on flux through the pathway (Shintani and DellaPenna,
1998). If the entire pool of y-tocopherol can be methylated to a—tocopherol in HPT
overexpressers (Savidge et al., 2001), then a combined overexpression of HPT and y—TMT

will increase vitamin E activity in Arabidopsis seeds up to lS—fold.

46

CHAPTER 1

Collakova E, DellaPenna, D (2001) Plant Physiol 127: 1113-1124. The following work is

copyrighted by the American Society of Plant Physiologists and is presented here with the

Society’s permission.

47

ISOLATION AND FUNCTIONAL ANALYSIS OF HOMOGENTISATE
PHYTYLTRANSFERASE FROM S YNECHOC YST IS SP. PCC 6803 AND
ARABIDOPSIS THALIANA

Abstract

Tocopherols, collectively known as Vitamin E, are lipid soluble antioxidants
synthesized exclusively by photosynthetic organisms and are required components of
mammalian diets. The committed step in tocopherol biosynthesis involves condensation of
homogentisic acid and phytyl diphosphate (PDP) catalyzed by a membrane—bound
homogentisate phytyltransferase (HPT). HPTS were identified fiom Synechacystis sp. PCC
6803 and Arabidopsis thaliana based on their sequence similarity to chlorophyll synthases,
which utilize PDP in a similar prenylation reaction. HPTS from both organisms used
homogentisic acid and PDP as their preferred substrates in vitro but only Synechacystis
HPT was active with geranylgeranyl diphosphate as a substrate. Neither enzyme could
utilize solanesyl diphosphate, the prenyl substrate for plastoquinone-9 synthesis. In
addition, disruption of Synechacystis HPT function causes an absence of tocopherols
without affecting plastoquinone-9 levels, indicating that separate polyprenyltransferases
exist for tocopherol and plastoquinone synthesis in Synechacystis. Surprisingly, the
absence of tocopherols in this mutant had no discernible effect on cell grth and

photosynthesis.
Introduction
Tocopherols are a group of amphipathic compounds synthesized only by
photosynthetic organisms. The best characterized and probably most important function of

tocopherols is to act as recyclable chain-reaction terminators of polyunsaturated fatty acid

48

free radicals generated by lipid oxidation. Tocopherols have a well—docurnented role in
mammals both as an essential nutrient (vitamin E) and general antioxidant (Fryer, 1993;
Liebler, 1998; Brigelius-Flohe and Traber, 1999). A similar, though less well documented
antioxidant role is also proposed for tocopherols in photosynthetic organisms (Fryer, 1992;
Niyogi, 1999).

From a biosynthetic perspective, tocopherols are members of a large, multifunctional
family of lipid soluble compounds called prenquuinones that also include tocotrienols,
plastoquinones, and phylloquinones (vitamin K1). Structural features shared by all
prenquuinones include hydrophobic prenyl tails of various lengths attached to aromatic
head-groups that can reversibly change their redox states. Tocopherols contain a chromanol
head-group and lipophillic tail derived from the 20-carbon alcohol phytol, while
plastoquinones contain a quinone head-group and isoprenoid tails of 40, 45, or 50 carbons.
Such structural features are essential for the diverse biochemical and physiological roles
fulfilled by various prenquuinones.

The committed step in the synthesis of all prenquuinones is the condensation of
various aromatic precursors and prenyl diphosphate (prenyl-DP) substrates in reactions
catalyzed by a small family of related polyprenyltransferases (Lopez et al., 1996). Most
aromatic and prenyl-DP substrates are utilized by more than one polyprenyltransferase
(Figure 6). For example, the aromatic compound homogentisic acid (HGA) is used for
condensation with PDP, geranylgeranyl diphosphate (GGDP), or solanesyl diphosphate
(SDP) in tocopherol, tocotrienol, and plastoquinone synthesis, respectively, while PDP is
used as the isoprenoid-derived tail in the synthesis of tocopherols, phylloquinones, and
chlorophylls (Threlfall and Whistance, 1971; Schulze-Siebert et al., 1987; Oster et al.,
1997). Thus, polyprenyltransferases act at biosynthetic branch-points and are potential key
regulatory enzymes for the synthesis of many essential compounds in photosynthetic

organisms.

49

In plant chloroplasts, the synthesis of tocopherols and plastoquinones is closely
related. Biochemical studies have shown that condensation of HGA with PDP or SDP
yields 2-methyl-6-phytyl-l,4-benzoquinol (MPBQ) and 2-demethylplastoquinol-9, the first
prenquuinol intermediates in tocopherol and plastoquinone-9 (PQ-9) synthesis,

respectively (Hutson and Threlfall, 1980; S011 et al., 1980; Marshall et al., 1985). Although

HO U
cuzcocog
PPM p—hydroxyphenyl- PP M H
pyruvate

GGDP pd” 4? Isopentenyl-DP

e .. we
P" H chzco; PPMH

 

3 OH
Chlorophyllide PhytyI-DP Homogentisate SolmeeyI-DP
(GGDP) ¥
1 ,4-dihydroxy 0" pgsz -|

2-naphtoate w H
OH
OH 3
I 2-methyl-6- % H
phytyl benzoquinol OH 9

+ 2-demathyl-
Chlorophylls I plastoquinoI-s

i i I

Phylloqmnone W H o I
HO 3 m H

(rt-tocopherol o 9
(Tocotrienol) Plastoquinone-9

 

Figure 6. Generalized overview of prenquuinone biosynthetic pathways in photosynthetic
organisms. Prenylation steps and substrates in tocopherol and plastoquinone synthesis are
shown in detail, while those for other prenyllipids are incomplete for clarity. Aromatic and
prenyl-DP substrates are shared among the various polyprenyltransferases (see text for
details). Enzymes are depicted as numbers in black circles: 1. Homogentisate
phytyltransferase; 2. Homogentisate solanesyltransferase; 3. Chlorophyll synthase; 4. 1,4-
dihydroxy-Z-naphthoate phytyltransferase; 5. GGDP reductase; 6. p-
hydroxyphenylpyruvate dioxygenase; 7. SDP synthase. Compounds in parentheses
indicate where GGDP may be used in place of PDP by HPT resulting in a tocotrienol
product.

50

these studies could not distinguish whether one or more polyprenyltransferases catalyzed
these reactions, it was suggested that separate enzymes might be involved (Schulze—Siebert
et al., 1987). In contrast, recent genetic data from Arabidopsis thaliana suggested
involvement of a single polyprenyltransferase activity in tocopherol and PQ-9 synthesis.
Two loci were identified, PDS] and PDS2 (PDS = Phytoene DeSaturation), which when
mutated, decreased the levels of both tocopherols and plastoquinones below detection
(Norris et al., 1995), consistent with the disruption of enzymes shared in their synthesis.
The PDS] locus has been cloned and encodes p—hydroxyphenylpyruvate dioxygenase
(HPPD) (Norris et al., 1998), which catalyzes formation of HGA. The pdsZ mutation was
proposed to disrupt another shared pathway enzyme, most likely a polyprenyltransferase,
which could utilize either PDP or SDP as substrates for condensation with HGA (Norris et
al., 1995). Unlike PBS], the PDSZ locus has not yet been cloned.

As an altemative to purifying the membrane-bound PDS2 gene product or walking
to the PDS2 locus, we attempted to clone an orthologous gene from the cyanobacterium
Synechacystis sp. PCC 6803, which also synthesizes or-tocopherol. In this paper, we report
the cloning and functional analysis of gene products from Synechacystis and Arabidopsis
encoding polyprenyltransferases specific to tocopherol biosynthesis. We also present
biochemical and physiological characterization of the corresponding Synechacystis

polyprenyltransferase knockout mutant, which completely lacks tocopherols.

Results

Identification And Disruption of A Polyprenyltransferase Involved in Tocopherol
Biosynthesis in Synechacystis sp. PCC 6803

Due to the metabolic synteny observed for the prenyllipid biosynthetic pathways in
photosynthetic organisms, we decided to utilize a genomics-based approach to identify the

gene encoding the homogentisate polyprenyltransferase involved in tocopherol synthesis,

51

first from cyanobacteria, and subsequently from plants. We hypothesized that this
polyprenyltransferase would show some similarity to previously characterized
polyprenyltransferases from cyanobacteria and plants that utilize similar prenyl-DPS as
substrates.

Chlorophyll synthase is a polyprenyltransferase that attaches PDP or GGDP to the
tetrapyrrole core of chlorophyllide during chlorophyll biosynthesis (Lopez et al., 1996;
Oster et al., 1997). The Synechacystis chlorophyll synthase ORF (ChlG, GenBank
accession number BAA10281) was used to query CyanoBase, which contains the complete
Synechacystis sp. PCC 6803 genome sequence (Kaneko et al., 1996). Several ORFs
showing varying degrees of similarity were identified and SLR1736 was selected as a
putative HPT based on its protein identity to ChlG (21%) and the presence of prenyl
diphosphate and divalent cation-binding motifs characteristic of polyprenyltransferases
(Lopez et al., 1996; Figure 7A). SLR1736 is also a highly hydrophobic protein (Figure
7B), as would be expected for a membrane-bound HPT (Soll et al., 1980; 8011 et al., 1984).

To test the hypothesis that SLR1736 is involved in tocopherol biosynthesis, a
disruption mutant (SLR1736::Km') was generated by homologous recombination of the
kanamycin cassette-disrupted SLR1736 gene into the wild-type SLR1736 locus (Figure 8).
If the SLR1736::Kmr mutation disrupted HPT activity, one would expect a complete
absence of tocopherols and their prenquuinol intermediates. HPLC analysis shows that
wild type Synechacystis lipid extracts contain predominantly or-tocopherol (Figure 9 and
Table I). In contrast, a-tocopherol and its prenylchromanol and quinol precursors are
absent from SLR1736::Kmr lipid extracts (Figure 9, Table I and data not shown), consistent
with the hypothesis that SLR1736 encodes a polyprenyltransferase involved in tocopherol
synthesis.

As shown in Figure 6, the various polyprenyltransferases in photosynthetic
organisms utilize many of the same aromatic and prenyl-DP substrates. Homogentisic acid

is the aromatic precursor in both tocopherol and plastoquinone synthesis and PDP is a

52

substrate for tocopherol, phylloquinone, and chlorophyll polyprenyltransferases (Threlfall
and Whistance, 1971; Schulze-Siebert et al., 1987; Oster et al., 1997). Given this
biosynthetic relationship, disrupting SLR1736 activity could directly or indirectly affect the
synthesis of other prenylated compounds in pathways that also utilize these substrates. To

determine the effect of the SLR1736 gene disruption on the synthesis of other prenylated

A

AtHPT msLLssssLvsxxccrcwonLansLsunancnssxvvxxrxrnNNLvarncocssLLLernx

ChIG - - - - MSDTQNTG- onox
SynHPT ---------------- Mir -------- I di'rrhiiiirsar 33- - liffifrfilvfifivrfirr mains mam
AtHPT snrnvnxrxcbnu ns nsxoxsr ansLmnvnrsnna - - vrervzs its Ms rrcgvsxvs n- - LI-ILL
cme KA- - - - - - - noLLcMKGAAPc- . - -- - as sr Mr nLomepiirwr Ini- r‘wc vccmrssflvr wsvrrnr

_ . . 7 _ ..... , ; ‘ ‘ _
SynHPT hDLvrchndctlcii 1 Ivcuror. a vet. RINKP NI. PLA 150F131AQBRWVGLCGUKfiLAIAIWLdL
AtHPT PTGI Lithvvaxnmvm Tremor. 1» var IKVNKPYLPL cnsVnrcI AILUASFSIMUFMGlfl vns
ChlG IKALTC MLH’SGPIIHI‘ TQT‘I..I'.€DF .1. an], Araer‘vnzr P 2A1 :vrflvvror Ll LL'Qllmcr cvrrvemnv

    

 

:0 .
SynHPT thLTVGIBLII61-?7'77'1'TX'Y'S'T77'FP anIrnrs LLAA LICK ET'V’IGI‘SWNLGLP PPRIILGYPWI
“HPT .. ‘FWALFV‘FNLGI° ° - - - 5W5! NLPLLIV‘KRIALVMNCI LAVRAIIIMQI Arv nrornvrcnmr LP
ChIG br- AQIIDPthvancc/IrvmnTailgrsugoncvtcnunaasv- - - - - - -1A PWAEHALFGI‘N

SynHPT 3 If wring: ivflfl‘ifli' .fg’rc‘iiv tsunami Ii‘i‘orjflr or fixonfi,firtfiiinrcc$uflr fiLWA
AtHPT n. Linn Mr Straw AL. KIN Paracnxrrcrnsrsflrr‘ our: Icvrfwom" VAI r. ATS
01116 P- rammingm Y’LAGLG‘éVVNrF KS vggprtmm. Kilrr'iMI-‘fir crAAiIII Emma pvr GIHGYLI vv

SynHPT AMPIII‘NTAPLINPsniticuLALnWNS an IIQ1. as warm PYO'PI‘VKBIIEL‘IYLEYPLADV."rs N'l‘l r
AtHPT H I wsxvr s vamp I EA'I'TIIIAIAKS upswflltric‘vwn Intramural . .
ChlG Hool- - - YATI VL L.l P-QI H on» YrgnNPLignnvxzoAs- Aoru‘rvr cmxrflflcnxcr . - .

 

4
B AtHPT

    

 

 

 

8
§
’3‘
§
§
§
§
Hydro phi Iicity

»2
-o
1.2

 

 

 

Figure 7. Alignment of Synechacystis and Arabidopsis polyprenyltransferases. A,
Homogentisate phytyltransferases from Synechacystis sp. PCC 6803 (SynHPT, GenBank
accession number 874813) and Arabidopsis (AtHPT, AF 324344) share 41% protein
identity, while SynHPT and ChlG (Synechacystis sp. PCC 6803 chlorophyll synthase,
BAA10281) share 22% protein identity. Residues conserved in at least two of three
sequences are shaded in gray, while residues identical in all three proteins are labeled by
black dots. The conserved prenyl-DP and divalent cation binding domains are indicated by
dashed and solid boxes, respectively. The predicted AtHPT chloroplast targeting domain
cleavage site is indicated by a black arrow. B, Kyte/Doolittle hydrophillicity profiles of
AtHPT and SynHPT. The two profiles are nearly identical. Negative values indicate
hydrophobicity.

53

A Kmr - 886 bp

 

 

4- 1.85 kb

”,3 4.. 0.97 kb

Figure 8. Construction of the Synechacystis PCC 6803 SLR1736::Kmr mutant. A,
Simplified scheme of the wild type SLR1736 ORF in the Synechacystis genome. Insertion
of a kanamycin (Km') cassette into Mfel site of the SLR1736 ORF and the SLR1736F and
R PCR primers (F and R, small arrows) are indicated. B, Autoradiograph of the PCR
products amplified from wild type (lane 1) and the SLR1736::Kmr mutant (lane 2) genomic
DNA. No wild type copies ofSLR1736 were detected in the SLR1736zszr mutant.

 

 

 

 

 

 

O

0

c

0

g

a II
10‘ F , .,

E (I Wlld type \___
5‘ _- A, W ‘

-w: “' SLR1736::KI'nr
$2 1'6 23 22
Minubs

Figure 9. HPLC traces of tocopherol standards and lipids extracted from wild type
Synechacystis sp. PCC 6803 and SLR1736::Km'. Equivalent weights of fresh cells were
extracted for the analysis Shown. Tocopherol analysis was performed on a normal phase
column using 8% di-isopropyl ether in hexane as a solvent. A, Separation of a-, B—, y-, and
8-tocopherol (Toc = tocopherol) and tocol standards. B, Wild type cells accumulate
predominantly (rt-tocopherol (gray trace). No tocopherols were detected in the
SLR1736::Kmr mutant (black trace). Tocol was used as an internal standard.

54

Table I. Prenyllipid contents afphatoautatraphically grown wild type Synechacystis sp.
PCC 6803 and the SLR1736::Km' mutant. Other than the absence of tocopherols in the
SLR1736::Kmr mutant, no significant differences were observed between wild type and
mutant cells for the presented parameters. Each value is the mean i SD of at least five
separate measurements per experiment. Each experiment was repeated at least three times.
8 mmol mol'l chlorophyll a; b mmol mg‘I cells

 

Prenyllipid wild type SLR1736::Kmr
a—tocopherol “ 18.7 i- 1.4 not detected
Plastoquinone-9 a 31.6 i 4.3 28.4 i 1.7
Phylloquinone ’ 13.8 i 1.1 12.5 i 1.2
Chlorophyll a b 4.3 i 0.3 4.6 i 0.4

 

 

compounds, we analyzed plastoquinone, phylloquinone, and chlorophyll levels in the
SLR1736::Kmr mutant relative to wild type. No significant differences were observed in the

levels of these compounds (Table 1).

Biochemical Characterization of the SLR1736 Gene Product

The SLR173622Kmr phenotype strongly suggests that SLR1736 encodes a
polyprenyltransferase specific to tocopherol synthesis. To determine the activity and
substrate specificity of the SLR1736 gene product, HGA polyprenyltransferase assays were
performed using SLR1736 protein expressed in E. cali. These assays are based on TLC
separation and subsequent autoradiography or HPLC separation of prenylated quinones
formed from radioactive HGA and various unlabeled prenyl-DPS in the presence of a
putative polyprenyltransferase.

When various prenyl-DPS at the same molar concentrations were tested as potential
substrates for the SLR1736 protein, PDP was used most efficiently, though GGDP could
also be utilized (Figures 10, 11B and 11D). The amount of geranylgeranylated
benzoquinone product formed was approximately 18% that of the phytylated product. No

products were observed when SDP, the prenyl-DP substrate for PQ-9 synthesis, was used

55

(Figure 10). In the case of PDP, the main reaction product co-migrated with 2 ’-trans-
MPBQ in both TLC and HPLC analyses (Figures 10 and 113).

 

 

  
   

Control SynHPT AtHPT
Q
Q Q QOQ
#6"? (59° °e (1°08 triage;
{AI-Kr
+ riff-a? fi- 1.11.!» ..i....‘... ‘II'.'.: +1,- "n- use W'

123456789101112

Figure 10. Homogentisate polyprenyltransferase assays. Individual reactions contained the
indicated prenyl-DP and protein extracts from E. cali expressing empty vector or the
indicated phytyltransferases. Radiolabeled prenquuinol reaction products were extracted,
oxidized to corresponding quinones, separated by TLC, and subjected to autoradiography.
SynHPT can utilize both PDP and GGDP as prenyl-DP substrates (lanes 6 and 7), while
AtHPT can only use PDP (lane 10). Neither enzyme could catalyze condensation of HGA
and SDP (lanes 8 and 12). No prenquuinone products were detected in control reactions
(lanes 1 - 5, and 9). The arrow indicates the origin.

We also observed a couple of minor products on TLC and HPLC (Figure 10 and
l 1). On TLC, a band (Figure 10, lane 6, Rf~ 0.3) likely corresponds to the quinol form of
MPBQ, since intensity of this band increases when the samples are not oxidized with AgO
prior to TLC (data not shown). A small radioactive peak eluting before the major 2 ’-trans-
MPBQ peak was also observed in HPLC analysis (Figure 118). This peak probably
corresponds to 2 ’-cis-MPBQ formed by isomerization of the trans-isomer as previously
reported (Hutson and Threlfall, 1980; Henry et al., 1987). It is unlikely that this peak
represents the corresponding quinol as quinols are eluted much later than quinones in the

HPLC system used. Due to their low abundance, firrther analyses of these minor peaks
could not be performed.

56

When GGDP was used as a substrate the SLR1736 enzyme also produced a major
and minor product. Unfortunately, 2-methyl-6-geranylgeranyl-l,4-benzoquinone (MGBQ),
the expected product of HGA and GGDP condensation, was not available. However,
indirect evidence suggests that the major GGDP reaction product is MGBQ. First,
consistent with previous reports (S011 and Schultz, 1979; Hutson and Threlfall, 1980), this
product migrates slightly slower than its phytylated counterpart in the TLC system used
(Compare lanes 6 and 7 of Figure 10). Moreover, the chromatographic properties of the
major product in normal-phase HPLC are consistent with those previously published for
MGBQ (Hutson and Threlfall, 1980). Specifically, it elutes 9 minutes after the internal
control 2 ’-trans-MPBQ (Figure 11C), which is in good agreement with the previously
reported relative elution difference between MPBQ and MGBQ (Hutson and Threlfall,
1980). As with PDP, an unknown minor GGDP product migrating prior to the major peak
is also present in HPLC analysis. Unfortunately, neither GGPD products are produced in
sufficient quantity to allow further analytical characterization. Based on the combined
results of these polyprenyltransferase assays and the tocopherol-specific phenotype of the
SLR1736::Kmr mutant, the protein encoded by the SLR1736 gene was named SynHPT, for

.Sjlrechacystis Homogentisate Bhytleransferase.

Identification And Characterization of An HPT Homologue from Arabidopsis
thaliana

To identify an HPT homologue from plants, we used the SynHPT protein sequence
as a database query for BLAST searches (Altschul et al., 1990). A single predicted
Arabidopsis gene on chromosome 2 (BAC clone F 19F24) containing regions of significant
similarity to SynHPT was identified. Subsequently, the corresponding cDNA was isolated
from an Arabidopsis seed cDNA library and fully sequenced. The predicted protein
encoded by this cDNA (GenBank accession number AF324344) shares 41% identity with

SynHPT. In addition, both proteins have remarkably similar hydrophobicity profiles and

57

 

100‘

19447

 

 

 

 

 

Absorbance (252 nm)
8

§§1
or
or

o 3 10 15 2'0 2'5

 

 

 

 

 

 

 

-A“‘-A‘-

 

 

 

I fl 1 fl 1 17

0 5 10 15 20 25 3O 35

Radioactivity (10‘3 DPM)
C d N (0 b 01 O}

 

2‘ t

wt“

9
I
r
i
i
i
i
I

 

 

 

 

I I I r T

0 5 10 15 20 25 30 35
minutes

Figure 11. Normal phase HPLC separation of radiolabeled prenquuinones produced from
HGA and prenyl-DP substrates by HPTS. Homogentisate polyprenyltransferase reactions
were performed in a total volume of 0.5 mL for SynHPT with PDP (panel B) and 5 mL for
SynHPT with GGDP, and AtHPT with PDP or GGDP (panels C, D, or E, respectively) as
described in Materials and Methods. Elution of the internal standard 2 ’-trans-MPBQ was
monitored at 252 nnr, while that of the prenquuinones formed during the assay was
monitored by scintillation counting of collected fractions. The UV traces are not Shown for
B, C and D for clarity, but when aligned with the radioelution profiles Shown, the major
radiolabeled phytylated products co-chromatographed with authentic 2 ’-trans-MPBQ
standard (indicated by arrows). A, Elution of 2 ’-trans-MPBQ; B and C, SynHPT and
AtHPT catalyzed formation of 2 '-trans-MPBQ from HGA and PDP; D, SynHPT catalyzed
formation of MGBQ from HGA and GGDP (circles). AtHPT did not produce a product
with HGA and GGDP as substrates (triangles).

58

contain prenyl-diphosphate and divalent cation binding motifs conserved in both location
and sequence (Figure 7). The Arabidopsis protein also contains an additional 95 amino
acid N-terminal extension that is not present in SynHPT. The first 36 amino acids of this
domain exhibit features of a chloroplast targeting sequence (Emanuelsson et al., 1999),
consistent with the reported chloroplast envelope localization of HPT activity in plants (8011
et al., 1980; 8011 et al., 1984). The Arabidopsis protein was tentatively named AtHPT for
Arabidopsis thaliana Homogentisate _Phytleransferase.

In order to determine the activity and substrate specificity of the putative AtHPT and
compare it to SynHPT, AtHPT was expressed in E. cali and HGA polyprenyltransferase
assays were performed. Like SynHPT, AtHPT catalyzed condensation of HGA and PDP
to form 2 ’—trans-MPBQ as a major product and was not active with the substrates HGA
and SDP. Unlike SynHPT, no products were observed when HGA and GGDP were used
as substrates (Figures 10 and 11D). To test whether this difference between the two
enzymes was due to the presence of chloroplast targeting sequences in AtHPT, we also
tested two truncated versions of the protein. One truncation removed the predicted 36
amino acid chloroplast transit peptide, while the second removed 95 N-terminal amino acids
not present in SynHPT. Neither truncation altered the specific activity or substrate
specificity of AtHPT (results not shown).

The specific activity of AtHPT expressed in E. cali was approximately 3% that of
SynHPT expressed from the same vector. Several explanations are plausible for this
difference, including decreased protein stability, poor protein expression in E. cali due to
codon bias, or an improper lipid environment relative to that of chloroplasts. Neither
AtHPT nor SynHPT could be visualized on coomassie-stained gels following induction,
indicating both are expressed at low levels in E. cali. Addition of lipids extracted from
Arabidopsis leaves or seeds to reactions had no discernible effect on AtHPT activity (data
not Shown). Finally, addition of Tween 80 and CHAPS to final concentrations of 0.2, 0.5,

1, or 2% moderately stimulated both AtHPT and SynHPT activities without appreciably

59

changing their Specific activities relative to each other (data not shown). It appears that a
combination of lower expression and/or lower stability of AtHPT relative to SynHPT may

be the cause of limited AtHPT activity in E. cali.

Physiological Consequences of Tocopherol Deficiency in Synechacystis sp. PCC
6803

Given the importance of tocopherols in free radical scavenging, a photosynthetic
organism lacking tocopherols might be expected to be compromised in growth or exhibit
increased sensitivity to high light stress. To address this question, we compared growth of
the tocopherol deficient SLR1736::Kmr mutant and wild type Synechacystis under low light
or high light conditions (~ 30 and 110 pE, respectively). Surprisingly, the doubling times

of both strains under photoautotrophic or heterotrophic conditions in both low and

Table [1. Growth and 02 evolution rates of wild type Synechacystis sp. PCC 6803 and
SLR1736::Km’. Doubling times and photosynthetic activity of wild type and mutant cells
are similar. Each value is the mean i SD of three independent measurements in a

representative experiment. Each experiment was repeated at least three times. a The cells
were grown at high light (105 - 110 uE); b The cells were grown photoautotrophically, 02
evolution rates are in mmol 02 mg'1 chlorophyll h'I and measured at 5 mE.

 

 

.- U“ ‘l»'{‘W-l “0"h ' .1. 4“ ‘. ‘9‘, P - nn‘n‘

 

Parameter wild'ty‘pe SL—Rl 736::Kmmr

Growth, doubling time (h) a “m“
Photoautotrophic 22 .i: 3 19 i 3
Photoheterotrophic 12 i- l 13 i 1

Oxygen evolution rates b
Whole chain 156 i 6 163 i 8

 

high light were comparable (Table II and data not shown). Whole-chain oxygen evolution
measured at 0.75, 2 and 5 mE was also found to be similar in both strains, indicating that the

initial rates of photosynthesis in SLR1736::Kmr and wild type are comparable (Table II).

60

Discussion

Due to the conserved evolution of photosynthetic organisms, many biosynthetic
pathways in cyanobacteria and plants are often quite similar (Whistance and Threlfall, 1970;
Marechal et al., 1997). The rapid growth of EST and genome databases in a wide variety of
organisms allows plant biochemists to utilize such inter-kingdom conservation in their
research more effectively. In particular, the availability of the fully sequenced Synechacystis
sp. PCC 6803 genome in a searchable on line database, CyanoBase (Kaneko et al., 1996),
coupled with straightforward gene disruption methods for analysis of gene firnction in this
organism (Williams, 1988) makes Synechacystis an attractive system to complement studies
of tocopherol synthesis in plants. Two tocopherol biosynthetic enzymes, GGDP reductase
(Addlesee et al., 1996; Keller et al., 1998) and y-tocopherol methyltransferase (Shintani and
DellaPenna, 1998) have already been cloned and characterized from Synechacystis and used
successfully as database probes to identify orthologs from Arabidopsis. The y-tocopherol
methyltransferases from both organisms were shown to have nearly identical activities
(Shintani and DellaPenna, 1998). We have employed a similar genomics-based approach to
identify and characterize a third enzyme of the tocopherol pathway from Synechacystis and
Arabidopsis, HGA phytyltransferase, and assess whether this step in tocopherol synthesis is
also conserved between cyanobacteria and plants.

In photosynthetic organisms, condensation of HGA with either a 20 or 45 carbon
prenyl-DP is the branch point in tocopherol and plastoquinone synthesis, respectively.
Early biochemical studies established that the tocopherol and plastoquinone pathways are
remarkably similar in oxygenic cyanobacteria, algae, and plants (Whistance and Threlfall,
1970). Although these studies could not distinguish separate polyprenyltransferase
activities for tocopherol and plastoquinone synthesis, it was suggested that separate
prenylation enzymes might be involved (Schulze—Siebert et al., 1987). More recently,

genetic analysis of the pathways in Arabidopsis identified two loci whose mutant

61

phenotypes are consistent with the disruption of enzymes shared in the synthesis of
tocopherols and plastoquinones (Norris et al., 1995). Indeed, this was found the case for
the PDS] locus, which encodes HPPD, the enzyme that produces the aromatic head-group
HGA in both the plastoquinone and tocopherol pathways (Norris et al., 1998). The PDS2
locus was suggested to encode a similarly shared polyprenyltransferase that could utilize
either PDP or SDP for tocopherol and plastoquinone synthesis, respectively (Norris et al.,
1995). The cloning of plant and cyanobacterial HPTS now allows us to directly address the
nature of polyprenyltransferases involved in tocopherol and plastoquinone synthesis in
oxygenic photosynthetic organisms.

The Synechacystis SLR1736::Kmr mutant lacks tocopherols but accumulates wild
type levels of PQ-9, consistent with the existence of separate HGA polyprenyltransferases
in tocopherol and plastoquinone synthesis. This conclusion is also supported by enzymatic
studies showing that SynHPT can utilize the 20 carbon tocopherol substrates, PDP or
GGDP, but is inactive with the 45 carbon PQ-9 substrate, SDP. Thus, it appears that
Synechacystis contains a single, polyprenyltransferase specific to tocopherol synthesis,
SynHPT, and a separate, yet to be characterized, polyprenyltransferase specific to
plastoquinone synthesis.

In Arabidopsis, the prenyltransferase reaction involving HGA and PDP substrates
appears to be nearly identical to that in Synechacystis. The AtHPT and SynHPT enzymes
share 61% protein similarity and both enzymes use PDP as their preferred prenyl-DP
substrate in vitro to form 2 ’-trans-MPBQ as a major product. This is in agreement with
previous biochemical studies of HPT activity in isolated spinach, lettuce, and pea
chloroplasts where MPBQ was the only product detected (Hutson and Threlfall, 1980; S011
et al., 1980; Marshall et al., 1985). Like SynHPT, AtHPT did not generate detectable
prenquuinone products with SDP as a substrate, suggesting Arabidopsis likely contains
separate polyprenyltransferases for tocopherol and plastoquinone synthesis. An

Arabidopsis HPT knockout mutant is needed to rigorously address this question.

62

In considering the nature of homogentisate polyprenyltransferase reactions in plants,
it is important to note that our original goal of cloning the Arabidopsis PDS2 locus has not
been achieved. Tocopherol and plastoquinone levels are reduced below detection in pds2,
leading to the hypothesis that PDS2 encodes a polyprenyltransferase shared in tocopherol
and plastoquinone synthesis (Norris et al., 1995). However, given that AtHPT is a
phytyltransferase encoded by a single-copy gene on chromosome 2 while PDS2 maps to
chromosome 3, this proposal now seems unlikely. If PDS2 is not a polyprenyltransferase
shared in tocopherol and plastoquinone synthesis, what does it encode? One explanation is
that PDS2 encodes an enzyme specific to plastoquinone synthesis (i.e., HGA
solanesyltransferase or SDP synthase) and that the absence of tocopherols in pds2 is a
pleiotropic effect of this mutation. In this scenario, the absence of plastoquinone, the main
lipid soluble electron carrier in plastids, results in such high levels of oxidative stress in
pds2 that any tocopherols produced are rapidly oxidized and degraded, and hence
undetectable. Alternatively, plastoquinone may be a cofactor required for the synthesis of
tocopherols and its absence arrests tocopherol synthesis. Regardless of mechanism, it
appears likely that the tocopherol deficiency in pds2 is an indirect, rather than a direct effect
of the pds2 mutation.

Though SynHPT and AtHPT are similar in their substrate specificities, there is one
notable exception: SynHPT can use both PDP and GGDP as substrates, while AtHPT only
uses PDP. The utilization of both PDP and GGDP as substrates by a
polyprenyltransferase is not unprecedented. An analogous reaction occurs in chlorophyll
biosynthesis where chlorophyll synthase can attach either PDP or GGDP to the tetrapyrrole
moiety, and in cyanobacteria, PDP is the preferred substrate (Oster et al., 1997). We
observed a similarly strong preference of SynHPT for PDP over GGDP. The use of
GGDP by SynHPT in viva would yield tocotrienol intermediates and endproducts that only
differ from their tocopherol counterparts in having an unsaturated rather than saturated

hydrophobic tail. This would necessitate subsequent enzymes in the pathway being active

63

towards geranylgeranylated substrates. At least one other tocopherol biosynthetic enzyme
from cyanobacteria has been shown to utilize both phytylated and geranylgeranylated
intermediates in vitro, tocopherol cyclase from Anabaena variabilis (Stocker et al., 1996).
However, Synechacystis sp. PCC 6803 does not accumulate tocotrienols, suggesting that
any geranylgeranylated intermediate produced by SynHPT is either efficiently reduced
(most likely by GGDP reductase), or that GGDP is not a substrate in viva. Additional work
is required to delineate the in viva substrate(s) and product(s) of SynHPT.

Within the limits of our assay sensitivity (~ 3 % of PDP product levels), AtHPT did
not utilize GGDP as a substrate. Other researchers also failed to demonstrate condensation
of HGA and GGDP using isolated Spinach, lettuce, and pea chloroplasts (Hutson and
Threlfall, 1980; 8011 et al., 1980). These data are consistent with the general observation that
dicots do not produce tocotrienols (Piironen et al., 1986; Franzen and Haas, 1991).
However, many monocots and gymnosperms do produce both tocopherols and tocotrienols
(Piironen et al., 1986; Franzen et al., 1991; Franzen and Haas, 1991) and we speculate that
HPTS from such organisms would utilize GGDP and PDP as substrates, analogous to
SynHPT. Phylogenetic analysis of polyprenyltransferases fiom various photosynthetic
organisms shows that HPTS from cyanobacteria, monocots and dicots form separate groups
(Figure 12), which probably represents taxonomic differences, although it may in part
reflect differences in the substrate specificities of these enzymes. As with Synechacystis,
we would also anticipate that pathway enzymes after HPT in monocots would be active
toward both geranylgeranylated and phytylated intermediates. The substrate specificity of
tocopherol biosynthetic enzymes from monocots have not been characterized, however, in
spinach (a dicot), where enzymology of the pathway has been best-studied, later
methyltransferases of the pathway are active toward various geranylgeranyl intermediates
(8011 and Schultz, 1979). It appears that at some point in evolution, dicots, like monocots,
could likely produce tocotrienols but have lost this ability as their HPTS have evolved

substrate specificity for PDP over GGDP.

64

0.167

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

0.241 0 167 Synechacystis } Chlorophyll
' Arabidopsis synthases
0.059
0.057 Wheat
E: cho Monocot
— HPT
0.175 0044
L— Maize
0.096
0117 _L 0 096 Soybean } Dicot
' Arabidopsis HPT
0.223
0069 Synechacystis
l 0.05 I o 093 0125 Nostoc Cyanobacterial
' 0425 HPT
Anabaena

Figure 12. Phylogenetic analysis of various prenyllipid polyprenyltransferases. Sequence
alignment and phylogenetic analysis were performed using MacVector software (Genetic
Computer Group, Madison, WI). Numbers indicate distances between protein sequences
estimated by the uncorrected p-distance method. Chlorophyll synthases form a separate
clade from homogentisate phytyltransferases. Within HPTS, cyanobacterial, monocot, and
dicot HPTS also form distinct sub-groups.

Numerous studies suggest that tocopherols are important antioxidants involved in
photoprotection of plants. Tocopherol levels correlate well with the degree of oxidative
stress in numerous plant species grown under various stress conditions, including high
light, drought, and low temperatures (Havaux et al., 2000; Wildi and Lutz, 1996; Streb et al.,
1998; Bartoli et al., 1999; Munne-Bosch and Alegre, 2000). Given this suggestive role of
tocopherols in antioxidant and photoprotective function, a mutation that eliminates
tocopherol synthesis in a photosynthetic organism would be anticipated to increase
sensitivity to oxidative stress, reducing growth or viability of this organism under stressful
conditions. Various mutants and transgenic plants with decreased tocopherol levels have
been reported to exhibit photobleaching phenotypes and compromised growth (Henry et al.,
1986; Norris et al., 1995; Tanaka et al., 1999). However, in all these cases, other

prenyllipids such as phylloquinone, chlorophylls, carotenoids, or plastoquinone were

affected in addition to tocopherols (Henry et al., 1986; Norris et al., 1995; Tanaka et al.,

65

1999). Therefore, it was impossible to specifically attribute these phenotypes to tocopherol
deficiency. The tocopherol-specific phenotype of SLR1736::Kmr provides a unique tool to
begin to specifically address the question of tocopherol function in photosynthetic
organisms.

Surprisingly, SLR1736::Kmr growth rates under photoautotrophic and
photoheterotrophic conditions in low and high light were indistinguishable from wild type.
Mutant and wild type whole-chain oxygen evolution rates were also similar. The
observation that the absence of tocopherols did not appreciably affect growth,
photosynthetic electron transport, and tolerance to high light stress seemingly contradicts
the concept that tocopherols are essential lipid soluble antioxidants. However, there are
several possible explanations for these apparently incongruous results. First, (rt-tocopherol
is not the only antioxidant present in photosynthetic membranes. Photosynthetic organisms
have evolved multiple mechanisms for protection from oxidative stresses (carotenoids,
ascorbate, superoxide dismutases, etc.) which if up-regulated, could partially or fully
compensate for the absence of tocopherols in SLR1736::KmIr under certain conditions.
Alternatively, tocopherols may protect from a specific type of lipid peroxidation or at a
particular Site. Finally, the lirrrited oxidative treatments used in this report may not be
sufficient to produce detectable differences between wild type Synechacystis and
SLR1736::Kmr at the level of culture growth rates. A detailed analysis of membrane lipids,
lipid peroxidation products, and other component of oxidative stress compensation and
adaptation are needed to discern any effects of tocopherol deficiency in SLR1736zsz'.

Finally, one potential application of the described work would be to allow
engineering of elevated tocopherol levels in food crops for nutritional purposes. Given the
central position of HPT in tocopherol synthesis, it seems likely that the enzyme may be an
important step for controling flux into the pathway. A crucial observation is that eliminating
HPT activity in Synechacystis does not affect the levels of other biosynthetically related

compounds (plastoquinone, phylloquinone, and chlorophylls). This suggests that altering

66

HPT enzyme levels in plants may also be tocopherol-specific and have little effect on the
synthesis of other prenquuinone compounds in the plastid. Experiments are underway to
positively modify AtHPT expression in Arabidopsis to test whether the activity is a target

for engineering tocopherol levels in plants.

Materials and Methods

Chemicals and Bacterial Strains. Prenyl-DPS were more than 99% pure. PDP was
kindly provided by Dr. Stephanie Sen (Purdue University, Indianapolis, IN), while GGDP
and SDP were purchased from American Radiolabeled Chemicals (St. Louis, MO). (U-
l4C)-HPP (0.6 - 1.5 uM) was prepared from (U-MC)-tyrosine (specific activity 464 mCi
mmol", Amersham, Arlington Heights, IL) as described (Schulz et al., 1993). Tocol was a
gift from H. Banno (Eisai Company, Japan). A mixture of various cis- and trans-methyl-
phytyl-1,4-benzoquinone isomers was synthesized by Dr. Daniel Liebler (University of
Arizona, Tucson, AZ). 2 ’-trans-2-methyl-6-phytyl-1,4-benzoquinone was purified fi‘om the
mixture by a combination of TLC and HPLC (Henry et al., 1987). Plastoquinone-9 was
extracted from Iris hallandica bulbs and purified by both TLC (Pennock, 1985) and HPLC
(Johnson et al., 2000). Wild type Synechacystis sp. PCC 6803 was grown on BG-ll plates
or liquid media (Williams, 1988) either photoheterotrophically (with 15 mM glucose) or
photoautotrophically (without glucose) at 20 - 30 IE and 30 0C unless otherwise stated.
Synechacystis cells were sub-cultured at least three times in liquid media prior to growth
experiments. E. cali strains DHSa (Stratagene, La Jolla, CA) and BL-2l (DE3) (Novagen,

Milwaukee, WI) were used for conventional sub-cloning and protein expression,

respectively.

Plasmids and Mutants. Primers 5’-TATTCATATGGCAACTATCCAAGCI'I'I'I'IG-3’
(SLR1736F) and 5’-GGATCCTAATTGAAGAAGATACTAAATAGTI‘C-3’ (SLR1736R)

67

containing engineered Ndel and BamHI sites (underlined) and Vent DNA polymerase (New
England Biolabs, Beverly, MA) were used to amplify the SLR1736 ORF (GenBank
accession number BBA17774) from Synechacystis genomic DNA. The amplified fi'agment
was sub-cloned into the EcaRV site of pBluescript 11 KS (+) to generate pKSl736.
pKS l 736 was digested with Mfel and ligated with the EcoRI digested kanamycin resistance
cassette from pUC4K (Taylor and Rose, 1988). Two constructs with opposite orientation
of the kanamycin resistance cassette relative to the SLR1736 ORF were used to transform
wild type Synechacystis and generate disruption mutants by homologous recombination
(Williams, 1988). Transfonnants were sub-cultured on kanamycin containing media for
several plating cycles and the absence of wild type SLR1736 gene copies was confirmed by
PCR using SLR1736F and R primers followed by Southern blot analysis (Figure 8A and

B). Because the two orientation disruption mutants were phenotypically indistinguishable

(data not shown), that with the Kmr cassette in the same orientation as the SLR1736 ORF,
referred to hereafter as the SLR1736::Kmr mutant (Figure 8A), was used for further
analyses. The Ndel-BamHI fragment from pKSl736 encoding the entire SLR1736 ORF
was ligated into Ndel-BamHI digested pET30b (Novagen) to create pSynHPT, which was
transformed into BL-21 (DE3) cells for protein expression.

The SLR1736 protein sequence was used to search the A. thaliana database and
identify a single genomic clone, F19F24, containing a homologous sequence. Primers 5’-
TTGTTITCAGGCTGTTGTTGCAGCTCTC-3 ’ and 5 ’ -
CGT'ITCTGACCCAGAGTTACAGAGAATG-3’ were used to amplify a 977-bp fragment
from F 19F24 for use as a probe to screen an Arabidopsis seed cDNA library (a gift of Dr.
John Ohlrogge, Michigan State University, East Lansing, MI). The longest of 15 positive
clones was sequenced and shown to encode a protein similar to SLR1736 that was
designated AtHPT. For protein expression purposes the full-length clone encoding AtI-IPT
was amplified using primers (5’-CCATGGAGTCTCTGCTCTC-3’ and 5’-

68

GGATCCCAAGCAGAGACTTCI I IACC-3’) and sub-cloned into Ncal-BamHI digested

pET3d vector (Novagen) to generate pAtHPT.

Prenyllipid Analysis. Fifteen to twenty mg of 14-day old plate-grown, Synechacystis sp.
PCC 6803 cells were harvested, their lipids extracted (Bligh and Dyer, 1959) and dissolved
in 100 uL of hexane or ethyl acetate. Ten uL of each sample was withdrawn for chlorophyll
determination (Lichtenthaler, 1987), while 50 uL was subjected to HPLC (Hewlett Packard
1100, Wilmington, DE) on a LiChrosorb 5 Si60A 4.6 X 250 mm normal phase column
(Phenomenex, Torrance, CA) at 42 °C as described (Syvaoja et al., 1986). Tocopherols
were detected by fluorescence using 290 nm excitation and 325 nm emission. For
plastoquinone and phylloquinone analysis, separation was achieved on a reverse phase
column (Spherisorb 5 ODSZ 4.6 X 250 mrrr, Waters, Marlborough, MA) as described
(Johnson et al., 2000). Plastoquinone-9 and phylloquinone were detected at 250 nm and

275 nm, respectively.

Homogentisate polyprenyltransferase assay. Each 0.2 mL reaction contained freshly
prepared (U-'4C)-HPP (~ 0.2 uM, specific activity 464 mCi mmol“), 50 mM HEPES (pH
7.6), 4 mM MgC12, 50 mM potassium ascorbate, 100 uM KF, 0.2 % (w/v) CHAPS, and 0.1
mg of total protein extracted from E. cali expressing HPPD (Norris et al., 1995). Individual
reactions contained 100 pM PDP, GGDP, or SDP and the insoluble protein fraction from
E. cali expressing pSynHPT (0.03 mg protein), pAtHPT (1 mg protein), or the empty pET
vector (0.03 or 1 mg protein). Reactions were incubated for 1 hour at room temperature,
extracted with two volumes of methanolzchloroforrn (1:1), and any newly formed
prenquuinols were oxidized with AgO (Pennock, 1985). The organic phase was
transferred to a fresh tube, evaporated to dryness, dissolved in ethyl acetate, and subjected to

TLC on Silica gel (J .T.Baker, Phillisburg, NJ) developed with 20% ethyl ether in petroleum

69

ether (Pennock, 1985). Labeled prenquuinones were detected by autoradiography for 14
days.

For HPLC analyses, the polyprenyltransferase assays were performed as described
above except that larger volumes were used for the individual reactions to ensure formation
of sufficient products: 0.5 mL for SynHPT with PDP and 5 mL for SynHPT with GGDP,
and AtHPT with PDP or GGDP. At the end of incubation, reactions were spiked with 2 '-
trans-MPBQ and extracted prenyllipids resolved by TLC as above. The areas
corresponding to prenquuinones (Rf 0.36 - 0.67) were scraped from the TLC plates, eluted
with ethyl ether, dried under nitrogen, and dissolved in hexane. Samples were then
subjected to HPLC on a normal phase column (LiChrosorb 5 Si60A, 4.6 X 250 m) using
0.1% dioxane in hexane as a mobile phase to separate various methyl-phytyl benzoquinone
isomers (Henry et al., 1987). For geranylgeranylated quinone products, a mobile phase
consisting of 0.15% dioxane in iso-octane was used (Hutson and Threlfall, 1980).
Prenquuinones were detected at 252 nm. Eluents were collected at 30- to 60—second

intervals and the associated radioactivity determined by liquid scintillation counting.

Growth curves. Wild type Synechacystis and the SLR1736::Kmr mutant were inoculated
to a final OD730 of 0.05 in 50 mL of liquid BG-ll medium and grown at 30 °C with
vigorous shaking in four possible combinations: with or without 15 mM glucose and at 20-
30 (low light) or 105-1 10 (high light) uE. The OD730 was measured every 6-12 hours and

used to calculate cell density (Williams, 1988).

Oxygen evolution. Liquid cultures of photoautotrophically grown wild type and mutant
Synechacystis cells were washed twice and resuspended in fresh BG-ll medium at a
concentration of 3 mg chlorophyll per mL. The cells were exposed to three different high

light intensities for 5 minutes (0.75, 2 and 5 mE). Oxygen measurements were performed

with a Clark-type electrode at 250 C using a Hansatech CB 1 -D3 recording unit with Minirec

70

recording software (Hansatech Instruments, King’s Lynn, England). The oxygen evolution

rate was calculated from the slope within the'linear region of the curves.

Phylogenetic analysis. Sequence alignment (ClustalW alignment using the BLOSUM
series matrix) and subsequent phylogenetic analysis were performed using MacVector
software (Genetic Computer Group, Madison, WI). The N-terrninal 96-amino acid
extension of AtHPT and the corresponding N-termini of the other polyprenyltransferases
were not included in the phylogenetic analysis. The following protein sequences were used:
Synechacystis ChlG (BAA10281), Arabidopsis ChlG ($60222), Synechacystis HPT
($74813), Nostoc HPT (480 - 1445 bp of contig 566), Anabaena HPT (6672 - 7625 bp of
contig 0295), rice HPT (AX046728), maize HPT (AX046716), Arabidopsis HPT
(AF324344), soybean HPT (AX046734), and wheat HPT (BE471221 - overlapped
BE471221 and BG604641 corresponded to the C-tenninal part of AtHPT starting at Asp-
160). For phylogenetic analysis, distances between the amino acid sequences were
estimated by using the uncorrected p-distance method with gaps distributed proportionally.
The best tree was constructed by the Unweighted Pair-Group Method with Arithmetic Mean
with random tie breaking. Bootstrapping (10,000 reps.) confirmed the confidence of the

best tree structure.

71

CHAPTER 2

Collakova E, DellaPenna, D (2003) Plant Physiol 131: 632-642. The following work is
copyrighted by the American Society of Plant Physiologists and is presented here with the

Society’s permission.

72

HOMOGENTISATE PHYTYLTRANSFERASE ACTIVITY IS LIMITING FOR
TOCOPHEROL BIOSYNTHESIS IN ARABIDOPSIS

Abstract

Tocopherols are essential components of the human diet and synthesized
exclusively by photosynthetic organisms. These lipophilic antioxidants consist of a
chromanol ring and a lS-carbon tail derived from homogentisate (HGA) and phytyl
diphosphate (PDP), respectively. Condensation of HGA and PDP, the committed step in
tocopherol biosynthesis, is catalyzed by homogentisate phytyltransferase (HPT). To
investigate whether HPT activity is limiting for tocopherol synthesis in plants, the gene
encoding Arabidopsis thaliana HPT, HPT], was constitutively overexpressed in
Arabidopsis. In leaves, HPT] overexpression resulted in a ten-fold increase in HPT
specific activity, and a 4.4-fold increase in total tocopherol content relative to wild type. In
seeds, HPT] overexpression resulted in a four-fold increase in HPT specific activity and a
total seed tocopherol content that was 40% higher than wild type, primarily due to an
increase in y—tocopherol content. This enlarged pool of y-tocopherol was almost entirely
converted to (rt-tocopherol by crossing HPT] overexpressing plants with lines constitutively
overexpressing y—tocopherol methyltransferase (y-TMT). Seed of the resulting double
overexpressing lines had a 12-fold increase in vitamin E activity relative to wild type. These
results indicate that HPT activity is limiting in various Arabidopsis tissues and that total
tocopherol levels and vitamin E activity can be elevated in leaves and seeds by combined

overexpression of the HPT] and y-T M T genes.

Introduction

Tocopherols, collectively known as Vitamin E, are a class of lipid soluble
antioxidants synthesized exclusively by photosynthetic organisms. Tocopherols are

essential components of the human diet as they perform numerous critical functions

73

including quenching and scavenging various reactive oxygen species and free radicals and
protecting polyunsaturated fatty acids from lipid peroxidation (F ukuzawa and Gebicky,
1983; Neely et al., 1988; Fryer, 1993; Bramley et al., 2000). Due to these and other
activities, dietary tocopherols are thought to play an important role in improving immune
firnction and in limiting the incidence and progression of several degenerative human
diseases including certain types of cancer, cataracts, neurological disorders, and
cardiovascular disease (Brigelius-Flohe and Traber, 1999; Bramley et al., 2000; Pryor,
2000).

In plants, indirect evidence suggests that tocopherols perform antioxidant and radical
quenching functions similar to those in animals (Fryer, 1992), and that tocopherols may
have additional roles related to photosynthesis (Munne-Bosch and Alegre, 2002a). Plants
alter their tocopherol levels during development (Molina-Torres and Martinez, 1991;
Tramontano et al., 1992) and in response to a variety of stresses, including high light, low
temperature, drought, and salt stress (Gossett et al., 1994; Streb et al., 1998; Leipner et al.,
1999; Havaux et al., 2000; Munne—Bosch and Alegre, 2000). In addition, during leaf
senescence, a process accompanied by chlorophyll degradation and oxidative damage in
photosynthetic membranes, there is an increased accumulation of tocopherols (Rise et al.,
1989). These combined studies suggest that the synthesis of tocopherols is highly
regulated during plant grth and development.

Tocopherols are arnphipatic molecules, consisting of a polar chromanol head group
and a lipophilic isoprenoid tail derived from HGA and PDP, respectively. The tocopherol
biosynthetic pathway is depicted in Figure 13. In plants, the aromatic precursor of
tocopherols, HGA, is synthesized from p-hydroxyphenyl pyruvate (HPP) by a cytosolic
HPP dioxygenase (HPPD) (Nonis et al., 1995; Garcia et al., 1997; Garcia et al., 1999).
The biosynthetic source of HPP in tissues is unclear as HPP can originate from the

Shikimate pathway via prephenate or by transamination of tyrosine (Threlfall and

74

Shikimate

Tyrosine
V
CHzcocoo' glyceraldehyde-3P
,0 + pyruvate
Ho
p-hydroxyphenyl
pyruvate
°2 1-deoxy-D-xylylose-5P
co2
OH V
""9 . , Degradation 3 isopentenyl-PP
Q Products + dimethylallyl-PP
(2112000-
OH
homogentisate
GGDP
CO2 + PP
2-methyl-6-phytyI-BQ3 2 ,3-dimethyI-6-phytyl-BQ

? t
°W: °mw:

5-tocopherol y-tocopherol

? SAM SAM
HO HO
O H O H
3

B—tocopherol 3 a-tocopherol

Figure 13. The tocopherol biosynthetic pathway in plants. Dashed arrows represent
multiple steps. Enzymes are indicated by circled numbers: (1) homogentisate
phytyltransferase, (2) p-hydroxyphenylpyruvate dioxygenase, (3) homogentisate
dioxygenase, (4) geranylgeranyl diphosphate reductase, (5) geranylgeranyl diphosphate
synthase, (6) l~deoxy-D-xylulose-5-phosphate synthase, (7) 2-methyl-6-phytyl-l,4-
benzoquinol methyltransferase, (8) tocopherol cyclase, (9) y-tocopherol methyltransferase

75

Whistance, 1971; Fiedler et al., 1982; Garcia et al., 1999; Lopukhina et al., 2001). Phytyl
diphosphate is formed by the sequential action of the plastidic geranylgeranyl diphosphate
synthase 1 (GGPSI) and geranylgeranyl diphosphate reductase (GGDR) (Kuntz et al.,
1992; Addlesee et al., 1996; Keller et al., 1998; Addlesee and Hunter, 1999; Okada et al.,
2000). As with other plastid-synthesized isoprenoids, PDP is derived from the l-deoxy-D-
xylulose-S-phosphate (DXP) pathway (Eisenreich et al., 1998; Lichtenthaler, 1998).

In photosynthetic organisms, condensation of HGA and PDP, the committed step in
tocopherol biosynthesis, is catalyzed by homogentisate phytyltransferase (HPT) (8011 et al.,
1980; 8011 et al., 1984; 8011, 1987; Collakova and DellaPenna, 2001; Savidge et al., 2002).
The product of this reaction, 2-methyl-6-phytyl-l,4-benzoquinol (MPBQ), is the first
prenquuinol intermediate in the pathway and can be methylated to 2,3-dimethyl-6-phytyl-
1,4-benzoquinol (DMPBQ) by MPBQ methyltransferase (MPBQ MT) (8011 and Schultz,
1979; Hutson and Threlfall, 1980; S011 and Schultz, 1980; Marshall et al., 1985; 8011, 1987;
Shintani et al., 2002). Both MPBQ and DMPBQ are substrates for tocopherol cyclase
(TC) to yield the first tocopherols of the pathway, y-tocopherol and 8-tocopherol,
respectively (Stocker et al., 1996; Arango and Heise, 1998; Porfirova et al., 2002). Both 7-
and 8-tocopherol can be methylated by y-tocopherol methyltransferase (y-TMT) to yield a-
and B-tocopherol, respectively (D' Harlingue A and Camara, 1985; Shintani and DellaPenna,
1998)

Due to the importance of vitamin E in human and plant physiology, the tocopherol
biosynthetic pathway has become a focus for plant metabolic engineering. In order to
successfully manipulate the tocopherol content and/or composition of various plant tissues,
enzymes with high flux coeflicients must be identified in the pathway. This requires the
cloning of individual tocopherol biosynthetic enzymes and a detailed understanding of the
molecular and biochemical regulation of individual steps of the pathway. In recent years,
several genes encoding enzymes directly or indirectly involved in tocopherol biosynthesis

have been cloned and overexpressed in plants to test whether they are limiting for

76

tocopherol synthesis in various tissues (Garcia et al., 1997; Norris et al., 1998; Shintani and
DellaPenna, 1998; Garcia et al., 1999; Collakova and DellaPenna, 2001; Estevez et al., 2001;
Schledz et al., 2001; Savidge et al., 2002; Tsegaye et al., 2002).

Precursor feeding studies with safflower cell cultures suggested that levels of HGA,
PDP, or both might be limiting for flux through the tocopherol biosynthetic pathway
(Furuya et al., 1987). HPPD produces HGA from HPP and has been cloned and
characterized from a variety of plants (Garcia et al., 1997; Norris et al., 1998; Garcia et al.,
1999). Overexpression of HPPD in Arabidopsis leaves and seeds resulted in a lO-fold
increase in HPPD specific activity, but only a 10 to 30% increase in total tocopherol levels
(Tsegaye et al., 2002). This result may be due to enhanced degradation of HGA in
transgenics or to a low flux coefficient of HPPD in the tocopherol pathway (Tsegaye et al.,
2002). Plastidic phytol levels may also be limiting for tocopherol synthesis as feeding
phytol to safflower cell cultures stimulated tocopherol levels to an even higher extent than
HGA (Furuya et al., 1987). Other studies suggest that DXP pathway-derived IPP, which is
used for the synthesis of phytol and PDP, may limit isoprenoid synthesis in Arabidopsis
chloroplasts (Estevez et al., 2001). Constitutive overexpression of the first enzyme of the
DXP pathway, DXP synthase (DXPS) in Arabidopsis leaves resulted in elevated levels of
several plastidic isoprenoids including tocopherols, which were increased up to 2-fold
relative to wild type (Estevez et al., 2001).

Arabidopsis leaves accumulate predominantly or-tocopherol, while the major
tocopherol present in Arabidopsis seeds is y-tocopherol (Shintani and DellaPenna, 1998).
This difference was shown to be the result of low seed y—TMT activity as overexpression of
y—TMT in Arabidopsis seeds led to the conversion of more than 95% of the y-tocopherol to
(rt-tocopherol. In these experiments, total tocopherol levels remained unchanged in the
transgenic seed. These results indicated that y-TMT was a key enzyme controlling seed
tocopherol composition, but had no effect on flux through the pathway (Shintani and

DellaPenna, 1998).

77

Homogentisate phytyltransferase is one of the most recent tocopherol biosynthetic
enzyme to be cloned and characterized (Collakova and DellaPenna, 2001; Schledz et al.,
2001; Savidge et al., 2002). Homogentisate phytyltransferase catalyzes the committed step
of tocopherol biosynthesis (Figure 13), making it a likely candidate for an enzyme with a
high flux coefficient. To test whether HPT activity is limiting for tocopherol synthesis in
different tissues, HPT was overexpressed in Arabidopsis leaves and seeds and the resulting
transgenic lines were characterized at both the molecular and biochemical levels. In
addition, simultaneous overexpression of HPT and y—TMT in Arabidopsis leaves and seeds
was performed to determine whether the transgenic phenotypes conferred by these two

genes were additive.

Results

Biochemical And Molecular Characterization of Wild Type And 358::HPTI Plants

Using a genomicS-based approach, we have recently cloned HPT], the gene
encoding HPT in Arabidopsis, which catalyzes the condensation of HGA and PDP in
tocopherol synthesis (Collakova and DellaPenna, 2001). Because both HGA and PDP are
substrates for other prenyltransferases (Threlfall and Whistance, 1971; Schulze-Siebert et
al., 1987; Oster et al., 1997) and HPT activity is low in plant chloroplasts, we hypothesized
that HPT could be a highly regulated enzyme with a high flux coefficient (Collakova and
DellaPenna, 2001). To assess whether HPT activity is limiting for tocopherol synthesis in
different tissues, the enzyme was expressed under the control of the Cauliflower Mosaic
Virus 358 rRNA (CaMV 358) promoter in Arabidopsis. Sixty-six independent primary
transforrnant lines (358::HPT1) were generated and analyzed (data not shown). 358::HPT1
lines showing antibiotic resistance segregation ratios consistent with a single insertion locus

and exhibiting leaf tocopherol levels higher than wild type were taken to homozygosity.

78

To address the question of whether HPT activity was limiting for tocopherol
synthesis in photosynthetic tissue, tocopherol composition and levels were analyzed in
leaves of wild type and homozygous 358::HPT1 lines. Plants overexpressing HPT]
contained 3 to 4.4-fold higher total tocopherol levels than wild type that were highly
significant (p < 0.01, Figure 14). Both wild type and 358::HPT1 leaves contained
predominantly (rt-tocopherol, though y-tocopherol levels were also elevated in 3SS::HPT1.
Wild type leaves accumulated a-tocopherol at 19.1 i 1.5 pmol mg", while a-tocopherol
levels in leaves of 358::HPT1 ranged between 55 and 80 pmol mg". Leaf y-tocopherol
content was 0.6 i- 0.1 pmol mg‘1 in wild type and l - 9 pmol mg"l in HPT] overexpressers.
The levels of B- and 8-tocopherols in both wild type and 358::HPT1 were below detection.
MPBQ and DMPBQ, prenquuinol intermediates in tocopherol biosynthesis, were not
detectable in wild type and 358::HPT1 leaves (data not shown). These results suggest that

120

 

887 Clalpha
' 1 I amma 3"
81.0 9

9°“ lair

.7 87.0

60.0
I 61.16359 59.1

Ii

20.1

wt11548162135373940

358::HPT1

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Tocopherols (pmol mg'1 FW)
8

 

 

 

 

 

 

 

Figure 14. Tocopherol composition and levels in leaves of 4-week old wild type and
homozygous 358::HPT1 Arabidopsis plants. Thirty mg of leaf tissue was extracted and
individual tocopherols were separated and quantified by reverse phase HPLC. Each line is
represented as an average i SD of tocopherol levels in 3 plants analyzed in duplicate. Total
tocopherol levels in pmol mg" leaf are indicated above the error bar of each line. Total
tocopherol levels of 3SS::HPT1 lines were significantly higher than wild type levels (p <
0.01). B- and 5—tocopherols were not detected.

79

MPBQ MT and TC are not limiting activities for tocopherol synthesis in leaf tissue. Two
independent transgenic lines exhibiting relatively high a- and y-tocopherol levels,
358::HPT1-1 l and -54 (Figure 14), were selected for subsequent analyses.

To confirm that the increased tocopherol levels in transgenic lines were due to
elevated HPT activity resulting from overexpression of HPT], HPT mRNA levels and
specific activity were measured in wild type and transgenic lines. Real time PCR showed
that HPT mRNA levels were extremely low in wild type and 20- to lOO-fold higher in
35S::HPT1 lines (data not shown). Using isolated chloroplasts, prenyltransferase assays
demonstrated that HPT specific activity in 2, 4, and 6 week old wild type plants was also
very low (0.11 - 0.34 pmol h" mg" protein) and that transgenic lines had four- to ten-fold
higher HPT specific activity than the corresponding wild type plants (Table III). In general,
the highest HPT specific activity in both wild type and 358::HPT1 was observed

Table III. HPT specific activity in chloroplasts isolated fiam 2, 4, and 6 week old wild type
and 35S::HPT1-11 and -54 leaves. HPT specific activity was determined using
radiolabeled HGA and unlabeled PDP and expressed as an average i SD of 2 - 3
independent experiments performed in triplicate. Values in parentheses represent fold
increases in the specific activity of transgenics compared to wild type plants of the
corresponding age. HPT specific activity in leaves of both 35S::HPT1 overexpressers was
significantly higher than wild type (p < 0.05). Though there was a general trend of HPT
specific activity decreasing in older relative to younger plants in each line, this was only
statistically Significant between 2 and 4 week old wild type plants (p < 0.01)*.

 

Plant age Specific activity in pmol h'l mg" protein (fold change)

 

"[1566161 " wild type 358::HPT1-11 35S::HPT1-54
2 0.34 i 0.06* 1.69 i 0.39 (5.1) 1.36 i 0.11 (4.1)
4 0.11 i 003* 1.14 i 0.48 (10.3) 0.84 i 0.27 (7.6)
6 0.15 i 0.10 0.71 i 0.41 (4.7) 0.75 i 0.36 (5.0)

 

 

Wlm\fiw “\‘I ‘- \

in chloroplasts isolated from young, 2 week old plants. The specific activity of both wild
type and HPT] overexpressing lines decreased approximately by 2-fold by 6 weeks of age.

However, the elevated specific activity in transgenics relative to wild type was maintained

80

throughout plant development (Table 111). These results are consistent with previous
findings that HPT activity is highest in young plants and diminishes with age (Hutson and
Threlfall, 1980).

In addition to HPT, the enzymes shown in Figure 13, HPPD, HGAD, GGPSl,
GGDR, and y-TMT are also directly or indirectly involved in tocopherol synthesis. Given
the elevated levels of tocopherols in 35S::HPTI lines, it is possible that the expression of
other enzymes involved in tocopherol synthesis might also be indirectly affected in the
transgenics. To test this hypothesis, the mRNA levels of all available genes of the pathway
in wild type and 3SS::HPTI leaves were measured. With the exception of the 20- to 100-
fold increase in HPT mRNA levels in 3SS::HPTI, no significant differences in the
expression of these other tocopherol-related genes were observed between wild type and

358::HPT1 Arabidopsis plants (data not shown).

Effects of HPT] Overexpression on Tocopherol Levels in Arabidopsis Seed

Molecular and biochemical analyses of the tocopherol biosynthetic pathway in
leaves of wild type and 358::HPT1 transgenic plants demonstrated that HPT expression and
activity are limiting for tocopherol synthesis in Arabidopsis leaf tissue. To test whether
HPT is also limiting in seeds, total and individual tocopherol levels were determined in
seeds of wild type and homozygous 355::HPT1 plants. Total tocopherol levels in wild type
Arabidopsis seeds were 838 i 18 pmol mg" seed. In 8 of the 9 homozygous 35S::HPT1
lines shown in Figure 3, 358::HPT1-16 being the exception, highly significant increases (p
< 0.001) in total seed tocopherol content were observed. The best HPT] overexpressing
lines (11 and 54) contained up to 37% higher seed tocopherol levels than wild type,
corresponding to an additional 310 pmol tocopherols mg" seed (Figure 15). The majority
of this increase was due to elevation of y—tocopherol levels, the predominant tocopherol in

wild type Arabidopsis seeds, though in some transgenic lines, 5-tocopherol levels also

81

 

1 800

A l gamma
1500 _ I delta
[:1 alpha
1148 1156
1200 _ 1144 109 04310801108 1140

1048

1029
900 - 835 ‘ 885
600 -
300 -
o _l
I delta
13() a
l
l
60 e
40 I
20 -
0

Dalpha
wt 11 54 8 16 21 28 35 37 39 40 53
358::HPT1

Figure 15. Tocopherol composition and levels in seed of wild type and homozygous
35S::HPT1 Arabidopsis plants. Seed was extracted and analyzed for tocopherols by
normal phase HPLC. Tocopherol levels are expressed as an average i SD of 3 analyses
per each line in a representative experiment. Total tocopherol levels in pmol mg" seed are
indicated above the error bar of each line. Arabidopsis seeds accumulate predominantly y-
tocopherol, which increased up to 40% in 35S::HPTI compared to wild type. The levels of
a-tocopherol did not change significantly, whereas 8-tocopherol increased up to 2-fold in
some transgenic lines. B—Tocopherol was not detected. With the exception of 35S::HPT1-
16, statistical Significance for total tocopherol levels of all transgenic lines relative to wild
type wasp < 0.001.

 

 

Tocopherols (pmol mg'1 seed)

 

 

 

 

82

doubled. a—Tocopherol levels did not change significantly (Figure 15).

We have previously demonstrated that 358::HPT 1-11 and -54 have significantly
higher HPT specific activity relative to wild type (Table III). HPT specific activity in seeds
was also determined to assess whether the observed differences in seed tocopherol levels
between wild type and 358::HPT1 lines were correlated with increased HPT activity. Both
transgenic lines (-11 and -54) exhibited nearly a 4-fold increase in HPT specific activity
relative to wild type seeds (Table IV). When compared to leaf HPT specific activities, the
HPT specific activity in seeds was similar to that of chloroplasts from 6-week old plants
(Tables III and IV). Collectively, these results suggest that, as in photosynthetic tissues,
HPT activity is limiting for tocopherol synthesis in Arabidopsis seed.

Table IV. HPT specific activity in seed of wild type and 35S:.°HPTI overexpressers (lines
#11 and 54). Protein extracts from dry Arabidopsis seeds were used to determine HPT
specific activity as described. Values are an average i SD of 3 independent experiments

performed in triplicate. HPT specific activity in seed of both 358::HPT1 overexpressers
was significantly higher than wild type (p < 0.01).

 

 

 

n HPT specific activity
Line [pmol h‘I mg" protein]
’ wild typem “ 0.17 i 0.05 ”

358::HPTl-ll 0.66 i 0.10
358::HPTl-54 0.67 i 0.08

 

Tocopherol Analysis of Arabidopsis Leaves And Seeds Overexpressing Both HPTI
And rTM T Transgenes

Constitutive HPT] overexpression in Arabidopsis led to a maximal 4.4-fold and
40% increase in total leaf and seed tocopherol levels, respectively. The increase in leaves
was primarily due to an elevation in a—tocopherol levels and to a lesser extent y—tocopherol.
In Arabidopsis seed, y-tocopherol was the major contributor to the 40% increase (Figure

15). This is a result of a limitation in seed y—TMT activity, which catalyzes methylation of

83

y- and 8-t000pherols to a- and B—tocopherols, respectively, and this metabolic block can be
overcome by overexpressing y-TMT during seed development (Shintani and DellaPenna,
1998).

To test whether the traits conferred individually by y-TMT and HPT overexpression
are additive, individual 358::HPT1 and 358::y-T MT lines were crossed, double
homozygotes selected, and their leaf and seed tocopherol content and composition
determined. The levels of individual tocopherols in leaves of wild type, double
overexpressers, and the corresponding single transgene homozygous parent lines are shown
in Figure 16. The total leaf tocopherol levels of 3SS::HPTI—ll and —54 in Figure 16 are
somewhat lower than in Figure 14. Leaf tocopherol content is extremely sensitive to
environmental stimuli and plant developmental stage (Munne-Bosch and Alegre, 2002a),
and these differences are consistent with the normal range of inter-experiment variation

observed in tocopherol content (data not shown).

 

so
Dalpha
Igamma 566
6° " 52-9 53.0 54.3 a; 51.3
I J: — I
40 -

21.5 21.1

.E!gll_a_

11 54 49/11 49/54 18]"
y—TM T HP T1 r—TM 17HP T1

 

 

Tocopherols (pmol mg" FW)
N
o
l

 

 

 

 

 

 

 

 

 

 

 

 

Figure 16. Tocopherol composition and levels in leaves of S-week old wild type, 358:27-
TMT, 358::HPT1, and double 358::rTMT/3SS::HPT1 overexpressers. Leaf tissue (~ 70
mg) was extracted and individual tocopherols were separated and quantified by normal
phase HPLC. Each line is represented as an average 3: SD from 3 plants. Total tocopherol
levels of 358::HPT1 and 3SSzzy-T MT /358::HPT1 plants were significantly higher than wild
type levels (p < 0.005). y-tocopherol in double transgenics was methylated to a-tOCOphCI‘Ol.

84

There was no significant difference in tocopherol composition or levels between
wild type and 358::y-T MT leaves. However, the leaf Hocopherol present in 358::HPT1-11
and -54 was converted to a—tocopherol in 3SS::HPTI/3SS::y-TMT double homozygotes
(Figure 16). Similarly, in seed of double overexpressers, nearly the entire pool of y- and 6-
tocopherols was methylated to a- and B-tocopherols, respectively (Figure 17A and B).

These data indicate that the traits conferred by each single transgene are additive.

 

   

  

 

 

 

 

 

 

 

 

 

 

 

 

1800
Igamma A
1500 J Dalpha
Idelta 1164
1200 .. Ibeta 1076 a 1043 1058
397

’6 90° ' 301 ,___.
8
3’ 600 -
b
E 300 -
O
E
e. 0 d
.3 70
E 60‘ Idelta B
8. Ibeta
O
O
.—

 

 

 

3 .

wt 18

11 54 49/11 49/54 18I11
r—TMT HP r1 r—TMTIHP T1

Figure 17. Tocopherol composition and levels in seed of wild type, BSSzzy-T MT ,
358::HPT1, and 358::rTMT/3SSzzHPT1 overexpressers. A, a—, l}, y-, and 6- tocopherols,
B, B- and 8- tocopherols. Tocopherol analysis of mature dry seed was performed as
described in Materials and Methods. Each line is presented as an average of 3 independent
measurements of 3 replicates. Total tocopherol levels of 358::HPT1 and 358::7-
TMT/3SS::HPT1 lines were significantly higher than wild type (p < 0.05). The majority of
y- and 8- tocopherols was converted to a- and B- tocopherols in 358::y-TMT/3583HPT1
overexpressers.

85

Effects of Increased Tocopherol Levels on Chlorophyll, Carotenoid, And
Plastoquinone-9 Content in 358::HPT1

In plastids, the HGA and IPP (as GGDP or PDP) used in tocopherol synthesis are
also utilized in the synthesis of chlorophylls, carotenoids, and plastoquinone-9 (PQ-9)
(Threlfall and Whistance, 1971; Schulze-Siebert et al., 1987; Oster et al., 1997). Several
studies suggest that the plastidic GGDP pool originating from the DXP pathway may be
limiting for isoprenoid synthesis (Fray et al., 1995; Shewmaker et al., 1999; Estevez et al.,
2001). Increased flux through the tocopherol pathway may therefore have an effect on the
levels of other compounds formed from GGDP or HGA. To test this hypothesis, levels of
HGA-derived PQ-9 and two major GGDP-derived compounds, chlorophylls and
carotenoids, were measured in leaves and seeds of wild type and 358::HPT1 -11 and -54.
No statistically significant differences in total chlorophyll, carotenoids, or PQ-9 levels were

observed between wild type and transgenic plants (Table V). In leaf, the average chlorophyll

Table V. Total chlorophyll, carotenoids, and PQ-9 levels in wild type and 35S::HPT1
Arabidopsis leaves and seeds. In leaves, chlorophyll and carotenoid levels were determined
in 90% methanol as described previously (Lichtenthaler, 1987). Twenty mg of seed was
extracted and pigments were analyzed by reverse phase HPLC. Lutein comprised > 95% of
total seed carotenoids (data not shown) and its levels were taken as total carotenoid levels.
Chlorophylls and PQ-9 were not detected (nd) in dry Arabidopsis seed. Chlorophyll and
carotenoid levels in 3SS::HPTI leaves and seeds were comparable to wild type levels.
Pigment levels are an average 1- SD of 3 measurements.

 

 

' Leaf content Seed content
Line Chlorophyll] Carotenoid PQ-9 -1 Chlorophyll Carotenoid
(nmol mg ) (nmol mg ) (pmol mg ) and PQ-9 (pmol mg )
wild type 1.00 i 0.06 0.43 i 0.04 82 i 8 nd 30.3 i 0.2
HPT1-ll 1.01 i 0.10 0.45 i 0.04 80 i- 5 nd 28.3 i 0.6

HPT1-54 0.98 i 0.08 0.43 i 0.05 83 i 4 nd 35.9 i 0.2

 

 

 

n- .—-—

86

and carotenoid levels were ~ 1 nmol mg" and 0.44 nmol mg" tissue, respectively. In wild
type and 3SS::HPT1, PQ-9 levels were ~ 82 pmol mg" leaf tissue. In seed, chlorophylls
and PQ-9 were not detected, while carotenoid levels were ~ 31 pmol mg" seed (Table V).

Discussion

Homogentisate phytyltransferase is a likely candidate for an enzyme with a high
flux coefficient in tocopherol biosynthesis. HPT utilizes HGA and PDP to catalyze
formation of the first committed prenquuinol intermediate in the tocopherol biosynthetic
pathway, MPBQ. Because HGA and PDP are also used in the synthesis of plastoquinones,
phylloquinones, and chlorophylls (Threlfall and Whistance, 1971; Schulze-Siebert et al.,
1987; Oster et al., 1997) and HPT acts at the branch-point for these plastidic prenyllipid
pathways, it could be a key enzyme controlling flux through the tocopherol pathway. To
test this hypothesis, the Arabidopsis HPT1 cDNA was constitutively expressed under the
control of the CaMV 35$ promoter. Significant increases in HPT specific activity and total
tocopherol levels were observed in both leaf and seed of 358::HPT1 plants compared to
wild type.

In leaves, HPT1 overexpression resulted in a 4.4-fold increase in total tocopherols,
the highest percentage increase in engineering leaf tocopherol content yet reported in
Arabidopsis. Three other enzymes affecting tocopherol biosynthesis, DXPS, HPPD, and y-
TMT, have been overexpressed in Arabidopsis leaves with varying degrees of success in
altering leaf tocopherol levels. Estevez et a1. (2001) reported significantly elevated levels of
several plastidic isoprenoids, including a 2-fold increase in tocopherols, when DXPS was
constitutively expressed in Arabidopsis leaves. Only a 30% increase in total leaf tocopherol
content was observed in transgenic Arabidopsis lines overexpressing HPPD (Tsegaye et al.,
2002). Constitutive overexpression of y—TMT in Arabidopsis had no effect on tocopherol

composition or total tocopherol levels in leaves (Shintani and DellaPenna, 1998). With the

87

exception of y-TMT, it appears that HPT, DXPS, and HPPD activities all limit tocopherol
accumulation to some degree in wild type Arabidopsis leaves.

In seeds, HPT1 overexpression resulted in accumulation of 37% more total
tocopherols than wild type, which corresponds to an additional 310 pmol mg" seed versus
70 pmol mg" tissue for the 4.4 fold increase in leaf tocopherol levels. The increase in the
absolute tocopherol levels in leaf tissue appears less significant than in seed until one
considers the relative water content of the two tissues. Because Arabidopsis leaves contain
91% water, total tocopherol levels increased from 230 to 1010 pmol mg" dry weight in wild
type and 358::HPT1 leaves, respectively. In contrast, because mature Arabidopsis seed
contain 11% water, total tocopherol content increased from 940 to 1290 pmol mg" dry
weight in wild type and 3SS::HPT1 leaves, respectively. Thus, overexpressing HPT1
elevated the HPT specific activity of seeds and leaves to similar levels, leading to similar
final levels of total tocopherols in the two tissues but a proportionately higher impact on
tocopherol accumulation in leaves. These data suggests that HPT has a relatively high flux
coefficient in both leaves and seed, though indirect evidence suggests HPT activity is still
limiting for tocopherol synthesis in 358::HPT1 seeds. Savidge et al. (2002) reported that
HPT1 overexpression driven by the strong, seed-specific napin promoter resulted in a 75%
increase in seed tocopherol levels relative to wild type, which would correspond to
approximately 1650 pmol mg" dry weight. Although HPT activity was not determined in
these studies (Savidge et al., 2002), the higher tocopherol levels obtained are consistent with
reports that the CaMV 358 promoter is less effective in developing seed than the napin
promoter (Eccleston and Ohlrogge, 1998). Despite these differences, both studies
demonstrate that HPT activity is limiting for tocopherol synthesis in wild type Arabidopsis
seed and is a key enzyme regulating tocopherol accumulation in plant tissues.

By alleviating the HPT limitation in the tocopherol pathway, it is possible that other
enzyme activities downstream of HPT (TC, MPBQ MT, or y—TMT) or their substrates

might become limiting in 358::HPT1 plants. A limitation in TC activity would result in

88

accumulation of DMPBQ, while a limitation in MPBQ MT activity would lead to the
increased levels of B- and 8—tocopherols. If both enzymes were limiting, MPBQ in addition
to DMPBQ and B- and til-tocopherols would accumulate. None of these compounds were
detected in leaves of wild type or 3SS::HPT1 plants. In 358::HPT1 seed, 5-tocopherol
levels were doubled, but still accounted for less than 5% of the total tocopherol pool. These
data indicate that neither TC nor MPBQ MT are limiting in 358::HPT1 seeds and leaves.
Recent radiotracer studies using HGA and PDP also suggested that TC and MPBQ MT are
not limiting for a-tocopherol synthesis in chromoplasts of yellow pepper (Arango and
Heise, 1998).

In wild type Arabidopsis seeds, but not in leaves, y—TMT activity has previously
been shown to be limiting for (at-tocopherol synthesis (Shintani and DellaPenna, 1998). In
leaves of 3SS::HPT1, y-tocopherol levels were up to 15-fold higher than wild type, while a
37% increase in y-tocopherol levels was observed in 358::HPTI seeds. To test whether 7-
TMT or S-adenosyl-L-methionine (SAM) were limiting in 358::HPT 1, the previously
characterized 358::y-T MT lines (Shintani and DellaPenna, 1998) were crossed to
358::HPT1 lines and the resulting double homozygous transgenic progeny selected and
analyzed for tocopherol content and composition. Overexpression of y—TMT in the
358::HPTI background resulted in almost complete methylation of the excess y—tocopherol
to a-tocopherol in both leaves and seeds. The persistence of small quantities of y-
tocopherol in 3SSzzy-TMT and the double transgenics may reflect a distinct functional role
for y—tocopherol or indicate that a fraction of the y-tocopherol pool is not accessible for
further methylation. Collectively, these data indicate that the biochemical phenotypes
conferred individually by HPT and y—TMT overexpression are additive in both seeds and
leaves, and that SAM is not limiting for a—tocopherol synthesis in 358::HPT1 lines.

Altering flux through a metabolic pathway can have unanticipated effects on
compounds synthesized by biochemically related pathways, especially when some of the
substrates are shared among these pathways (Fray et al., 1995; Shewmaker et al., 1999;

89

Estevez et al., 2001). Overexpression of DXPS in Arabidopsis leaves increased the levels
of several isoprenoid-derived compounds including carotenoids, chlorophylls, and
tocopherols, suggesting that availability of chloroplastic GGDP, a common precursor in
their biosynthesis, may be limiting in wild type plants (Estevez et al., 2001). In canola seed,
overexpression of phytoene synthase led to a 50-fold increase in carotenoids and a 50%
decrease in tocopherol levels relative to wild type (Shewmaker et al., 1999). Constitutive
overexpression of the GGDP-utilizing carotenoid biosynthetic enzyme phytoene synthase
in tomato caused chlorosis and dwarfism, most likely by redirecting the limited GGDP pool
from the gibberellin and chlorophyll biosynthetic pathways to carotenoid synthesis (Fray et
al., 1995). In light of these reports, the effects of HPT1 overexpression on the levels of
other HGA- and GGDP-derived compounds in Arabidopsis tissues were also assessed.
Although we did not directly measure gibberellin levels in leaves of 358::HPT1
plants, gibberellin metabolism appeared to be unaffected as no obvious differences in
growth, flowering time, dormancy, or physical appearance were observed relative to wild
type (data not shown). PQ-9, chlorophyll, and carotenoid levels in leaf and seed were also
unaffected by HPT1 overexpression in Arabidopsis. One molecule of GGDP is used in the
synthesis of each chlorophyll molecule, while carotenoids contain two molecules of GGDP
or “GGDP equivalents”. The total chlorophyll and carotenoid content in leaves
corresponds to 1900 pmol mg" tissue of “GGDP equivalents”; the 70 pmol mg" increase
in incorporation of “GGDP equivalents” into leaf tocopherols in 358::HPT 1-11 is
negligible by comparison. In mature wild type Arabidopsis seeds, this composition is
reversed, with tocopherols predominating (up to 1000 pmol mg" seed) and carotenoids
being a minor component (~ 30 pmol mg" seed). The 310 pmol mg" seed increase in
“GGDP equivalents” incorporated into tocopherols in 358::HPT1-11 seed is highly
significant relative to the level of carotenoids, but apparently did not affect GG DP
availability for carotenoid synthesis. This increased demand for GGDP in seeds may be

compensated in part by increased flux through the DXP pathway. A similar phenomenon

90

was observed in Brassica seed engineered for elevated carotenoid synthesis (Shewmaker et

al., 1999). There seems to be a relatively high flexibility for flux through the seed DXP
pathway leading to GGDP, likely by feedback regulation.

Outlook for Metabolic Engineering of Vitamin E Levels in Crops

Tocopherols are essential components of the human diet. Due to the relatively low
levels of a—tocopherol, the most effective form of vitamin E, in most commonly consumed
vegetables and oils (Eitenmiller, 1997; Bramley et al., 2000; Ching and Mohamed, 2001), a-
tocopherol is limiting in the average American diet (Grusak and DellaPenna, 1999; Horwitt,
2001). It is recommended that 35 mo] (15 mg) of a—tocopherol be consumed daily (Food
and Nutrition Board, 2000). Although Arabidopsis is not of direct agricultural importance,
it has become a model for metabolic engineering of the tocopherol pathway and can be used
to extrapolate the impact of engineering on dietary vitamin E intakes. In Table VI we
estimate the vitamin B activity of wild type and various transgenic lines reported in this

study in terms of the a-tocopherol equivalents (ct-TE) per “serving size” of 100 g leaves

Table VI. Vitamin E activity in wild type, single, and double overexpressers. Vitamin B
activity was calculated for each line using the absolute levels of individual tocopherols from
Figures 16 and 17 and the relative biological activities of R,R,R-tocopherols. One mg of or-
, B-, y—, and 8-tocopherol corresponds to l, 0.5, 0.1, and 0.03 mg a-TE, respectively (Food
and Nutrition Board, 2000).

 

- s'“?rllwtd?1'lmfi’b"u’\l r—w -

 

 

 

Vitamin B activity
.. .. (mg a-TE per 100 g tissue)”
Line leaves seeds
wild type 0.75 3.9
35S::HPT1 1.81 5.2
358::y-TMT 0.87 35.6
358::HPT1/35822y-TMT 2.39 47.4

 

91

and seeds (for definition of a-TE see Table VI). In leaves of 358::HPT1/3SSzzy-TMT
double overexpressers, which accumulate predominantly a-tocopherol, vitamin E activity
was 3.2-fold higher than wild type. Because y-tocopherol is the major tocopherol in mature
wild type Arabidopsis seed, the vitamin E activity is relatively low (3.9 mg a—TE) despite the
high levels of total tocopherols. In seed of 358::HPT1/3SS::y-TMT lines, a-tocopherol is
the major tocopherol and the vitamin B activity increased 12-fold relative to wild type (Table
VI). Similar increases in seed vitamin B activity are anticipated for genetically engineered
crops (Grusak and DellaPenna, 1999).

Materials And Methods

Chemicals. Tocopherol standards and tocol were purchased from Matreya (Pleasant Gap,
PA). Phytyl diphosphate was a gift from Dr. Stephanie Sen (Purdue University, West
Lafayette, IN). [U- l4C]HPP was prepared from L-[U- l4C]tyrosine (sp. activity 464 mCi
mmol", Amersham, Arlington Heights, IL) according to Schulz et a1. (1993). Minor
modifications were introduced to obtain more concentrated [U- ”C]HPP. Briefly, L-[U-
l“C]tyrosine was dried under a stream of nitrogen and dissolved in the original volume of
phosphate buffer (0.5 M, pH 6.5). Catalase (Roche, Indianapolis, IN) and L-amino acid
oxidase type IV (Sigma, St. Louis, M0) were both added to the final concentration 0.4 mg
mL". After 2 hours of incubation at room temperature, [U- ”C]HPP was purified on an ion
exchange column (Dowex, Sigma) equilibrated with 0.1 N HCl and used immediately for

prenyltransferase assays.

Generation of HPT1 overexpressing lines. A cDNA encoding HPT was excised from
pSKHPTI (Collakova and DellaPenna, 2001) by digestion with EcoRI and KpnI and sub-
cloned into EcoRI and Kpnl digested pART7 (Gleave, 1992). The resulting construct was
mobilized by NotI digestion and ligation into the pART27-based vector pMLBART (Gleave,

92

1992), which contains the bar gene for selection of transformed plants. This construct was
introduced into Agrobacterium tumafaciens and used to transform wild type Arabidopsis
plants (ecotype Columbia) by the floral dip method (Clough and Bent, 1998) to obtain
3SS::HPTI T0 sense plants.

Sixty-six independent transforrnants were selected by glufosinate (120 mg L")
resistance and analyzed for tocopherol content. Seeds of lines exhibiting elevated leaf
tocopherol levels were harvested and subjected to segregation analysis. All T2 3SS::HPT1
transgenic plants segregating 3:1 for BASTA resistance were carried through the next
generation and plants homozygous for BASTA resistance were selected for further
experiments. Plants overexpressing y—T M T (Shintani and DellaPenna, 1998) under the
control of 358 CaMV promoter were crossed with 358::HPT] and double homozygotes
were selected based on their dual resistance to kanamycin and BASTA. Transgenic lines,

358::HPT1-1 1 and ~54 and 358217-TMT-l8 and -49 were used for these crosses.

Prenyllipid analyses of transgenic plants. For leaf prenyllipid analyses, plants were
grown in a 16-hour photoperiod (70 - 100 uE) at 22/19 °C day/night cycle for 4 to 5 weeks.
For tocopherol analysis, lipids from 30 to 35 mg of leaf tissue or 10 to 13 mg of seeds were
extracted in the presence of butylated hydroxytoluene (2 mg mL") to prevent tocopherol
degradation (Bligh and Dyer, 1959). Tocol was used as an internal standard. For leaf
analysis, tocopherols were separated on a reverse phase HPLC (Cm, ODSZ, 4.6 X 250 mm,
Column engineering, Ontario, CA; Shirnadzu VP HPLC system, Japan) using an isocratic
solvent system of 5% (v/v) isopropanol in methanol at 2 mL min". For leaf and seed
analyses, where separation of B- and y—tocopherols was required, a normal phase HPLC
system (LiChrosorb 5 Si60A 4.6 X 250 mm Silica column, Column engineering, Ontario,
CA and HP 1100 series HPLC system, Agilent Technologies, Palo Alto, CA) and a 10-
minute isocratic method using 17% di-isopropyl ether in hexane (v/v) at 42 °C at 2 mL min"

were utilized. Tocopherols were detected by fluorescence at 290 nm excitation and 325 nm

93

emission. Chlorophylls, carotenoids, and PQ-9 were analyzed as described (Collakova and
DellaPenna, 2001). For PQ-9 analysis, 50 mg of leaves or 15 mg of seeds was used for

extractions. Detection limit for PQ-9 was 0.04 pmol in seed.

Real time PCR. To obtain sufficient amounts of tissue, plants (wild type and 358::HPT1
#1 l and 54) were grown at a lO-hour photoperiod at 75 - 100 uE for 6 weeks. Tissue from
6 representative plants was harvested 2 - 3 hours after the start of the light cycle and
immediately frozen in liquid nitrogen. Total RNA was isolated and any contaminating
genomic DNA was removed by treatment with RQl-RNase free DNase (Promega,
Madison, WI). Ten pg of total RNA was reverse transcribed to generate cDNA in two 50-
uL reactions for each sample using a TaqMan kit according to manufacturer
recommendations (Applied Biosystems, Foster City, CA). An aliquot of cDNA
corresponding to 100 - 200 ng of total RNA was used in each TaqMan real-time PCR
assays (Applied Biosystems). Elongation factor EF la was used to normalize RNA
concentrations. Standard curves were constructed for each gene and used to calculate the
corresponding mRNA concentrations. Sequences of the primers and probes used in the

real-time PCR assays and their optimal final concentrations are available upon request.

Prenyltransferase assays. Homogentisate phytyl transferase assays were performed
using crude chloroplast preparations or seed protein extracts from wild type plants and
358::HPT1 #11 and 54 lines grown as described for real-time PCR assays. Chloroplasts
were prepared from 25 - 30 g of leaf tissue. Tissue was disrupted using a blender in a
buffer containing 0.6 M sorbitol, 0.5 M HEPES pH 8.4, 4 mM EDTA, 4 mM EGTA, 10
mM NazCO3, 0.2% Bovine Serum Albumin (w/v), 1 uM benzarnidine (BA), and 5 11M 4-
aminocaproic acid (4-ACA). Chloroplasts were filtered through 2 layers of miracloth and

centrifuged at 4,000 g for 5 minutes at 4 °C. Crude chloroplasts were washed once in a

94

buffer containing 50 mM HEPES pH 7.6, 4 mM MgCl,, 1 pM BA, and 5 11M 4-ACA and
once more in the same buffer lacking proteinase inhibitors.

Each prenyltransferase reaction (0.1 mL) contained 0 or 100 pM PDP,
approximately 5 pM [U- l4C]HPP (l uCi), 20 pg HPPD protein, and chloroplasts
corresponding to about 0.45 - 0.65 mg of protein (0.10 — 0.15 mg of chlorophyll) in HPT
reaction buffer (50 mM HEPES pH 7.6, 4 mM MgCl,, 1 mM KF, 0.2% (v/v) TWEEN 80,
and 50 mM potassium ascorbate). Reactions were incubated at room temperature for 2
hours with shaking and quinones were extracted with two volumes of acetone: light
petroleum ether (1:1 v/v). After centrifugation, the ether phase was loaded on a 4-mL Silica
column (pore size 60 um, Sigma) and quinones were eluted with 4 volumes of 20% (v/v)
diethyl ether in petroleum ether. After drying, 5 mL of scintillation liquid was added to each
sample and radioactivity was determined by scintillation counting. Background activities
obtained from control reactions containing l4C labeled HGA without exogenous PDP were
subtracted fi'om the activities when both substrates were added, which was always at least
3.5-fold higher than the background activities to obtain HPT specific activity. Chloroplasts
were extracted with 90% methanol (v/v) and centrifirged to determine chlorophyll
concentration (Lichtenthaller, 1987). Proteins present in the remaining pellet were
solubilized by NaOH in the presence of SDS and determined by the method of Lowry
(Stoscheck, 1990).

In seeds, protein extracts were prepared as follows: Dry mature seeds were ground
in 50 mM HEPES (pH 7.6) containing 4 mM MgCl2 and centrifuged. The resulting pellet
of water insoluble proteins was resuspended in HPT reaction buffer to a protein
concentration of 20 - 30 mg mL". HPT assays containing 2 - 3 mg seed protein were
performed in a volume of 0.2 mL as described for chloroplasts. Seed HPT specific

activities were about 3-fold higher than the corresponding background activities.

95

CHAPTER 3

Collakova E, DellaPenna, D (2003) submitted for publication. The following work will be

copyrighted by the American Society of Plant Physiologists and is presented here with the

Society’s permission.

96

HOMOGENTISATE PHYTYLTRANSFERASE ACTIVITY LIMITS
TOCOPHEROL SYNTHESIS IN STRESSED ARABIDOPSIS THALIANA
LEAVES

Abstract

Tocopherols are amphipathic antioxidants synthesized exclusively by photosynthetic
organisms. Tocopherol levels change significantly during plant growth and development
and in response to stress, likely as a result of altered expression of pathway-related genes.
Homogentisate phytyltransferase (HPT) is a key enzyme limiting tocopherol biosynthesis
in unstressed Arabidopsis leaves (Collakova and DellaPenna (2003) Plant Physiol. 131:
632-642). Wild type and transgenic Arabidopsis plants constitutively overexpressing HPT
(358::HPT1) were subjected to high light stress (up to 1 mE) for up to 15 days and
tocopherol levels, composition, and expression of several tocopherol pathway-related genes
were determined. High light stress resulted in an 18- and 8-fold increase in total tocopherol
content in wild type and 35S::HPTI leaves, respectively, with levels in 358::HPT1 being 2-
to 4-fold higher than wild type at all experimental time points. Increased total tocopherol
levels correlated with elevated HPT mRNA levels and HPT specific activity in 358::HPT1
and wild type leaves, suggesting that HPT activity limits total tocopherol synthesis during
high light stress. In addition, substrate availability, and expression of pathway enzymes
prior to HPT also contribute to increased tocopherol synthesis during stress. The
accumulation of high levels of B-, y-, and 5-tocopherols in stressed tissues suggested that the
methylation of phytquuinol and tocopherol intermediates limit a—tocopherol synthesis
during stress. Overexpression of y-tocopherol methyltransferase in the 358::HPT1
background resulted in nearly complete conversion of y— and 8-tocopherols to a- and B-
tocopherols, respectively, indicating that y-tocopherol methyltransferase activity limits or-

tocopherol synthesis during high light stress.

97

Introduction

Tocopherols are a group of lipid soluble antioxidants collectively known as vitamin
E that are essential components of animal diets. Dietary vitamin E is required for
maintaining proper muscular, immune, and neural function and may be involved in reducing
the risk of cancer, cardiovascular disease, and cataracts in humans (Pryor, 2000; Brigelius-
F lohe et al., 2002). In plants, tocopherols are believed to protect chloroplast membranes
from photo-oxidation and help to provide an optimal environment for the photosynthetic
machinery (Fryer, 1992; Munne-Bosch and Alegre, 2002a). Many of the proposed
tocopherol functions in animals and plants are related to their antioxidant properties, the
most prominent of which is protection of polyunsaturated fatty acids from lipid
peroxidation by quenching and scavenging various reactive oxygen species (ROS)
including singlet oxygen, superoxide radicals, and alkyl peroxy radicals (Fukuzawa and
Gebicky, 1983; Munne—Bosch and Alegre, 2002a).

Tocopherols are only synthesized by photosynthetic organisms and consist of a
polar chromanol ring and a lS-carbon lipophilic prenyl chain derived from homogentisic
acid (HGA) and phytyl diphosphate (PDP), respectively (Figure 18). In plants, HGA is
formed from p-hydroxyphenyl pyruvate (HPP) by the cytosolic enzyme HPP dioxygenase
(HPPD) (Garcia et al., 1997; Norris et al., 1998; Garcia et al., 1999). Based on radiotracer
studies, HPP can originate either from prephenate or tyrosine by the Shikimate pathway, but
the relative contribution of these two precursors to the total HPP pool is unknown (Threlfall
and Whistance, 1971; Fiedler et al., 1982; Lopukhina et al., 2001). In plastids, isopentenyl
diphosphate (IPP) derived from the 1-deoxyxylulose-S-phosphate (DXP) pathway
(Eisenreich et al., 1998; Lichtenthaler, 1998) is utilized by geranylgeranyl diphosphate
synthase 1 (GGPSl) for the synthesis of geranylgeranyl diphosphate (GGDP) (Okada et
al., 2000). Three of the four double bonds in the GGDP molecule are reduced to form PDP
through partially reduced intermediates by a multifunctional GGDP reductase (GGDR)

98

(Addlesee etal., 1996; Keller et al., 1998). Alternatively, PDP can be generated from phytol

and ATP by a kinase activity present in chloroplast stroma (8011 et al., 1980).

tyrosine prephenate
DXP pathway
_ v
Omzcmm 3 isopentenyl-PP
HO + dimethylallyl-PP
p-hydroxyphenyl
pyruvate degradation
0; products
4 PM
002 4
0" _°_> maleylaceto- GGDP
acetate
cflzcoo- chlorophyll

OH
homogentisate

"WM" +0: PW
PDP ATP

602+Pfi
HO 3AM HO
I H %_.> | H
OH 3 OH 3
2-methyl-6-phytyl-BQ 2,3-dlmethyI-6-phytyl-BQ

.. t .. ‘9

5-tocopherol y-tocopherol
? SAM SAM ‘?
Ho W Ho W
H H
o 3 o 3

B-tocopherol a-tocopherol

Figure 18. Tocopherol biosynthesis in plants. Dashed arrows represent multiple steps.
Enzymes are indicated as circled numbers: 1. homogentisate phytyltransferase, 2. tyrosine
transaminase, 3. prephenate dehydrogenase, 4. p-hydroxyphenylpyruvate dioxygenase, 5.
homogentisate dioxygenase, 6. geranylgeranyl diphosphate reductase, 7. geranylgeranyl
diphosphate synthase 1, 8. unspecified kinase, 9. chlorophyllase, 10. 2-methyl-6-phytyl-l,4-
benzoquinol methyltransferase, ll. tocopherol cyclase, 12. y-tocopherol methyltransferase

99

HPT is a membrane-bound chloroplast enzyme, which catalyzes the committed step
of tocopherol biosynthesis, the condensation of HGA and PDP, to form 2-methyl-6-phytyl-
1,4-benzoquinol (MPBQ) (8011, 1987; Collakova and DellaPenna, 2001; Savidge et al.,
2002). MPBQ can be methylated to 2,3-dimethyl-6-phytyl-1,4-benzoquinol (DMPBQ) by
MPBQ methyltransferase (MPBQ MT) (Marshall et al., 1985; 8011, 1987; Shintani et al.,
2002). MPBQ and DMPBQ can be cyclized by tocopherol cyclase (TC) to form 6- and y-
tocopherol, respectively (Stocker et al., 1996; Arango and Heise, 1998; Porfirova et al.,
2002). The last enzyme of the pathway, y-tocopherol methyltransferase (y—TMT), catalyzes
methylation of y- and 8—tocopherol to a- and B-tocopherol, respectively (D' Harlingue and
Camara, 1985; Shintani and DellaPenna, 1998).

In plants, tocopherol levels and composition vary in different tissues and fluctuate
during development and in response to abiotic stresses. Dry and germinating seeds of
many plants accumulate predominantly y—tocopherol, while a-tocopherol is the major
tocopherol in leaves, which may reflect distinct roles of individual tocopherols in these
tissues (Bramley et al., 2000; Franzen and Haas, 1991; Shintani and DellaPenna, 1998).
Significant increases in leaf a-tocopherol levels are observed during aging and senescing of
plants (Rise et al., 1989; Molina-Torres and Martinez, 1991; Tramontano et al., 1992),
possibly to protect cellular components from increased oxidative stress (Munne-Bosch and
Alegre, 2002c). Enhanced tocopherol accumulation also occurs in response to a variety of
abiotic stresses including high light, drought, salt, and cold and may provide an additional
line of protection from oxidative damage (Havaux et al., 2000; Munne-Bosch and Alegre,
2002a).

While there is a growing body of knowledge about the individual enzymes required
for tocopherol biosynthesis in plants, the mechanisms that regulate the overall pathway and
result in differential tocopherol content and composition during plant development or stress
remain poorly understood. Regulation of tocopherol biosynthesis in senescing and stressed

plants may occur at multiple steps of the pathway. HPPD activity limits tocopherol

100

synthesis in non-stressed Arabidopsis plants (Tsegaye et al., 2002) and HPPD mRNA
levels are up—regulated in senescing barley leaves (Klebler-Janke and Krupinska, 1997).
Similarly, various biotic and abiotic stresses elevate tyrosine aminotransferase (TAT)
mRNA and protein levels and enzyme activity in Arabidopsis (Lopukhina et al., 2001;
Sandorf and Hollander-Czytko, 2002). Whether other steps of the tocopherol pathway are
also involved in the regulation of tocopherol biosynthesis during stress remains to be
determined.

It has been recently demonstrated that HPT activity limits tocopherol synthesis in
non-stressed Arabidopsis leaves (Collakova and DellaPenna, 2003). The gene encoding
HPT, HPT1, has been cloned from Synechocystis sp. PCC 6803 and Arabidopsis thaliana
(Collakova and DellaPenna, 2001; Schledz et al., 2001). Overexpression of HPT in
Arabidopsis increased leaf and seed tocopherol content by 4.4-fold and 75%, respectively
(Collakova and DellaPenna, 2003; Savidge et al., 2002). The current study was undertaken
to further define the role of HPT in regulating tocopherol biosynthesis in stressed
photosynthetic tissues. By combining high light stress with molecular and biochemical
analyses, we have also identified additional enzymes and/or substrates that limit a-

tocopherol synthesis in stressed Arabidopsis leaves.

Results

The Biochemical Response of Wild Type and 358::HPT1 Plants To High Light
Stress

Stress is associated with increased total tocopherol levels in a variety of plants
(reviewed in Munne-Bosch and Alegre, 2002a). We have shown previously that HPT
activity is limiting for tocopherol synthesis in non-stressed Arabidopsis leaves (Collakova
and DellaPerma, 2003). To investigate whether HPT activity also limits tocopherol

synthesis in stressed Arabidopsis leaf tissue, 6-week-old wild type and two well-

101

characterized 358::HPT1 lines (lines 11 and 54, Collakova and DellaPenna, 2003) were
subjected to high light stress (0.8 - 1 mE) for up to 15 days and tocopherol content and
composition were analyzed during the treatment. The general response to stress was
monitored by assessing anthocyanin accumulation and the decrease in chlorophyll and

carotenoid levels (Figure 19A and B). Total anthocyanin levels rapidly increased fiom

 

   
  
 

 

 

 

 

  

 

 

 

 

A 40 r A Anthocyanins
‘2‘
E 30-
o
'6 20‘
0
18*
5“ 15
E
o
3
E
5
3.
o Carotenoids
18
C Total tocopherols
g“ 15‘
E 12~
u
a 9-
E 6‘
c
v 3-l
0

 

o 3 6 9 12 15
Days of treatment

Figure 19. Total anthocyanin, chlorophyll, carotenoid, and tocopherol levels in stressed wild
type and 3SS::HPT1 leaves. Plants of the indicated genotypes were grown in a 10/14 hour
light/dark cycle at 75 to 100 uE for 6 weeks and then transferred to approximately 900 uE
growth conditions. A, Anthocyanin accumulation in leaves of stressed wild type and
3SS::HPT1 Arabidopsis plants. Anthocyanin synthesis increased within the first 3 days of
hi gh-li ght stress and reached high steady-state levels afier 6 days of high light treatment. B,
Chlorophyll and carotenoid degradation in leaves of stressed wild type and 358::HPTI
plants. Total chlorophyll and carotenoid levels decreased gradually to approximately 50 %
of the initial levels in all stressed lines. C, Total tocopherol levels in leaves of stressed wild
type and 358::HPT I plants. High-light stress resulted in a significant elevation of total
tocopherol levels in both 358::HPTI transgenic and wild type plants.

102

below detection to approximately 35 nmol cm'2 leaf area by day six and were maintained at
this level throughout the stress treatment (Figure 19A). Total chlorophyll and carotenoid
levels decreased gradually during the 15-day stress treatment to approximately half of their
initial levels (Figure 193). There were no significant differences in chlorophyll, carotenoid,
or anthocyanin content between wild type and 3SS::HPT1 throughout the course of the
experiment (Figure 19A and B).

Total tocopherol levels increased in a near linear manner during exposure of both
wild type and 3SS::HPT1 plants to high light (R2 2 0.97, Figure 19C). Prior to stress
treatment, the total tocopherol levels of 358::HPTI leaves were 3-fold higher than wild type
(1.06 i 0.22 and 0.36 i 0.05 nmol cm'2 leaf area, respectively). In response to 15 days of
high light stress, total tocopherol levels increased 24—fold to 8.7 i 0.3 nmol cm‘2 leaf area in
wild type, while the total tocopherol levels of 35S::HPTI increased 15.5-fold to 16.4 i 0.8
nmol cm‘2 leaf area. After 15 days, the respective final total tocopherol levels in non-
stressed wild type and 358::HPT1 leaves were 0.49 i 0.04 and 2.00 i 0.50 nmol cm"2 leaf
area, respectively. At any time point during stress, total tocopherol levels in 358::HPTI
were 1.9- to 38-fold higher than wild type (p < 0.006, Figure 19C), suggesting that HPT

activity limits tocopherol synthesis in stressed wild type Arabidopsis leaves.

HPT Expression And Enzyme Activity in Unstressed And Stressed Wild Type And
358::HPT1

We have shown previously that non-stressed 358::HPT1 Arabidopsis plants
accumulated 20- to 100-fold higher HPT1 mRNA levels than wild type and showed 4- to
lO-fold increases in HPT specific activity, which resulted in up to 4.4-fold increased
tocopherol levels in leaves of transgenic lines compared to wild type (Collakova and
DellaPenna, 2003). In non-stressed wild type leaves, average HPT mRNA levels ranged
from 0.3 to 0.6 frnol mg" total RNA during the 12-day experimental time course (Figure

103

20A). Consistent with our previous study (Collakova and DellaPenna, 2003), HPT mRNA
levels in non-stressed 3SS::HPT1 leaves were at least 20—fold higher than wild type and
ranged from 7 to 15 fmol mg" total RNA during the course of the experiment (Figure 20B).

—O—wt
+11 } stressed

+54

w<>—-wt
.4}... 11 } non-stressed

+54

 

 

 

 

 

 

30a

 

HPT mRNA levels (fmol mg" total RNA)

 

 

O
o 3 6 9 12
Days of treatment

Figure 20. HPT expression in control and stressed wild type and 358::HPT1 plants.
Plants were grown and stressed as in Figure 2, total RNA extracted, and HPT mRNA levels
determined by real-time PCR. Data are normalized for EF-la mRNA levels and presented
as average 1 SD of three independent experiments. A, Wild type HPT mRNA levels. B,
Wild type and 358::HPT1 HPT mRNA levels. High light stress resulted in an up-
regulation of HPT mRNA levels in wild type, while no trend was observed in stressed
358::HPT1 and the corresponding control plants.

104

To assess any correlation between elevated total tocopherol levels and changes in
HPT expression or activity during stress, HPT mRNA levels and specific activity were
determined in non-stressed and stressed wild type and 358::HPTI plants. HPT mRNA
levels were induced up to 3.5-fold in wild type by 3 days of high light treatment and
remained elevated throughout the course of the experiment (Figure 20A). No clear trend
was observed for HPT mRNA levels in stressed 358::HPT1 lines (Figure 20B), though
there was significant biological variation in HPT mRNA levels in both wild type and
358::HPT1 plants during stress treatments (Figure 20A and B). Consistent with prior
studies, HPT specific activity in the absence of high light stress was 6—fold higher in
358::HPT1 lines compared to wild type (Figure 21; p < 0.0005). In response to 3 and 6
days of high light stress, HPT specific activity in wild type and 35S::HPT1 lines increased
approximately 3-fold and up to 4.4-fold relative to their respective unstressed controls. In

35S::HPT1, the relative HPT specific activity after 3 days of stress was 9-fold that of

 

 

 

 

 

 

Relative HPT specific activity

0 3 6
Days of treatment

Figure 21. Relative HPT specific activity in control and high light stressed wild type and
35S::HPT1 Arabidopsis chloroplasts. Six week-old plants were transferred to high light
(0.8 - l mE) for three and six days, chloroplasts isolated, and assayed for HPT activity.
Results from two independent experiments are presented as an average i SD of the activity
increase relative to wild type non-stressed chloroplasts (0.15 i 0.10 pmol h" mg" protein).
High light stress resulted in the strong induction of HPT activity relative to control plants in
both wild type and 3SS::HPTI. Relative HPT specific activity in 358::HPT1 was
significantly higher than in wild type (p < 0.0005).

105

comparably treated wild type. After 6 days of high light stress, HPT specific activity was
5.6- and 3.5-fold higher than the corresponding wild type for 358::HPTI lines I] and 54,

respectively (Figure 21).

Changes in mRNA Levels of Other Tocopherol Pathway-Related Genes in Non-
Stresscd And Stressed Wild Type And 3SS::HPTI Leaves

In addition to HPT, several other tocopherol biosynthetic enzymes (TAT, HPPD,
HGAD, GGPSI, GGDR, TC, and y-TMT) may play roles in regulating tocopherol
synthesis in Arabidopsis leaves. To assess the role of these enzymes in regulating
tocopherol accumulation during high light stress, their mRNA levels were measured in wild
type and 358::HPTI leaves using real-time PCR (Figure 22). TAT and HPPD catalyze
formation of the tocopherol biosynthetic precursors HPP and HGA, respectively, whereas
HGAD is involved in HGA degradation (Figure 18). GGPSI and GGDR catalyze
synthesis of PDP, a prenyl substrate used in tocopherol biosynthesis, while TC and y—TMT
are involved in regulating tocopherol composition (Figure 18). The steady-state mRNA
levels of TAT, HPPD, HGAD, GGPSI, and y—TMT in non-stressed tissues were similar in
the different genotypes and varied between 1 and 6 fmol mg" total RNA during the 12—day
experimental time course (Figure 22A, B, C, D, G). GGDR showed the highest steady-state
mRNA levels of all tested genes and fluctuated between 20 and 30 fmol mg" total RNA in
non-stressed Arabidopsis leaves (Figure 22E). TC mRNA levels were quite low (0.3 - 0.6
frnol mg" total RNA) and comparable to wild type HPT mRNA levels (Figures 22F and
21A). Although leaf mRNA levels of most tested genes varied 2- to 3-fold across all
genotypes and time-points the overall trends indicated relatively constant steady-state
mRNA levels for each gene and the differences observed probably represent the biological
variation of the system (Figure 22).

In response to high light treatment, TAT, HPPD, and HGAD mRNA levels were
elevated several-fold in both wild type and 3SS::HPT1 lines (Figure 22A, B, and C),

106

 

+ wt
+11 } stressed

+54

 

 

—~<>- wt
.--..:1--—— 11 } non stressed
—--{_-—- 54

 

 

 

 

B HPPD

 

 

 

 

 

 

 

 

mRNA levels (fmol mg‘1 total RNA)

 

 

 

 

 

 

 

 

 

 

 

 

 

o 3 e 9 12 o 3 e 9 12
Days of treatment

Figure 22. Expression of other tocopherol-related genes in control and stressed wild type
and 358::HPTI leaves. Experiments were performed as described in Figure 3. A, tyrosine
transaminase (TAT); B, p—hydroxyphenyl pyruvate dioxygenase (HPPD); C, homogentisate
dioxygenase (HGAD); D, geranylgeranyl diphosphate synthase 1 (GGPSI); E,
geranylgeranyl diphosphate reductase (GGDR); F, tocopherol cyclase (TC); G, 7-
tocopherol methyltransferase (y-TMT). TAT, HPPD, and HGAD mRNA levels increased,
while GGDR mRNA levels decresed during stress. GGPSI, TC, and y—TMT mRNA levels
were not up-regulated in response to stress.

107

suggesting that these enzymes may also be involved in regulating tocopherol levels during
stress. TAT mRNA levels increased 3- to 5-fold relative to non-stressed leaves within the
first 6 days of high light stress before returning near control levels by the end of the time
course (Figure 22A). HPPD and HGAD mRNA levels were elevated throughout the time
course of stress treatment (Figure 228 and C) and in this regard showed expression
profiles similar to HPT mRNA levels in stressed wild type (Figure 20A). The maximal
increase for HGAD and HPPD in stressed relative to non-stressed plants was 2.7- and 5.4-
fold, respectively (Figure 22B and C). GGPSI, TC, and y—TMT steady-state mRNA levels
were not significantly altered in wild type or 358::HPT1 leaves in response to high light
stress (Figure 22E, F, and G). The GGDR expression profile was unique in showing a
downward trend during high light stress (Figure 22D). As with HPT, mRNA levels of the
tocopherol pathway-related genes showed a large biological variation between experiments,
especially in stressed plants. Consistent with our previous study (Collakova and
DellaPenna, 2003), there were no significant differences between genotypes for mRNA
levels of these genes in stressed or unstressed plants (Figure 22). These results suggest
that increased HPT expression and tocopherol levels in 358::HPT1 transgenic lines does

not have a significant impact on the expression of other tocopherol pathway-related genes.

Analysis of Individual Tocopherols And Phytquuinols in Non-Stressed And
Stressed Wild Type And Transgenic Plants

Like most plants, Arabidopsis can synthesize four different tocopherols, 01-, B—, y-,
and 8-tocopherols from the phytquuinols MPBQ and DMPBQ by the routes shown in
Figure 18. The current study (Figure 19C, 20, and 21) and previous report (Collakova and
DellaPenna, 2003) make it clear that HPT activity is a major limitation in total tocopherol
synthesis and accumulation in both non-stressed and stressed leaves. Analyses of
tocopherol and phytquuinol levels and compositions were performed to identify steps

subsequent to HPT that might limit a—tocopherol synthesis in non-stressed and stressed

108

leaves of wild type and 358::HPT1 Arabidopsis plants. Non-stressed wild type
Arabidopsis leaves accumulate more than 95% (rt-tocopherol and less than 5% y-tocopherol
(Collakova and DellaPenna, 2003). In non-stressed 358::HPT1 leaves, a- and y-
tocopherols constituted 90% and 10% of total tocopherol levels, respectively. B-Tocopherol,
and 8-tocopherol were below detection levels in non-stressed wild type and 358::HPT1
Arabidopsis leaves. The phytquuinols MPBQ and DMPBQ were also below detection in
non-stressed and stressed wild type and 358::HPT1 leaves (data not shown), suggesting
that TC activity is not limiting for tocopherol synthesis during normal growth or high light
stress.

Tocopherol composition changed significantly in both wild type and 358::HPT1
leaves during high light stress. Though (rt-tocopherol was still the major tocopherol in
stressed leaves, B-, y—, and 8-tocopherols accumulated to high levels as well (Figure 23).
This effect was most pronounced in 3SS::I-IPTI leaves after 15 days of stress treatment,
where (rt-tocopherol constituted only 57% of total tocopherols, as opposed to 84 % in wild
type (Figure 23A). This difference was due to the presence of high levels of other
tocopherols in 358::HPT1 leaves throughout the experimental time course. B- and 6-
tocopherols accumulate when MPBQ is directly cyclized to yield 6—tocopherol rather than
being methylated to DMPBQ (Figure 18). Collectively, B- and 6-tocopherols accounted for
0.48 nmol cm’2 leaf area (5.6% of total tocopherols) in stressed wild type, but accounted for
3 and 6 nmol cm'2 leaf area (20 and 35% of total tocopherols) in stressed 358::HPT1 lines
54 and 11, respectively (Figure 23C and D). Similarly, stressed 358::HPTI lines
accumulated yutocopherol, the immediate precursor of (rt-tocopherol, to a greater extent than
wild type (Figure 238 and D). After 15 days of high light stress, y—tocopherol constituted
10% of total tocopherols in wild type and 14 and 17% in 358::HPT1-11 and -54,
respectively (Figure 23B and D). These results collectively indicate that both the
methylation of MPBQ to DMPBQ and y-tocopherol to (rt-tocopherol limit (rt-tocopherol

synthesis in stressed 358::HPT1 leaves.

109

 

 

1 , A a-tocopherol C B-tocopherol

 

 

 

 

 

 

 

B y-tocopherol D 6-tocopherol

 

 

 

 

Tocopherol levels (nmol cm'2 leaf area)

 

 

0 3 6 9 12 15 0 3 6 9 12 15

Days of treatment

Figure 23. Tocopherol levels and composition in stressed wild type and 358::HPT1 plants.
A, (rt-tocopherol B, y-tocopherol C, B-tocopherol D, 6-tocopherol. Leaves of stressed
358::HPT1 plants accumulated slightly more a-tocopherol than those of wild type plants.
Large differences are evident in the levels of B—, y-, and 8-tocopherol between 3SS::HPT1
and wild type.

To distinguish between a limitation in y—TMT activity and SAM levels, 358::HPT1
lines were crossed to previously characterized lines overexpressing y-TMT (3SSzzy-T MT),
which should eliminate any limitation to y—TMT activity in vivo (Shintani and DellaPenna,
1998; Collakova and DellaPenna, 2003). We have shown previously that under optimal
growth conditions, excess y—tocopherol synthesized in 358::HPTI was methylated to a-
tocopherol in 358::HPTI/35822y-TMT double overexpressers (Collakova and DellaPenna,
2003). During high light stress, y—TMT overexpression resulted in reduced 7— and 8-
tocopherol levels in 35$er MT and 358::HPT1/3SS:2y-TMT lines relative to wild type and
3SS::HPT1, respectively, without affecting total tocopherol levels (Figure 24). In stress
treated 358::HPT1/3SS::y-T MT double overexpressers, the change in tocopherol

composition was significant as y— and 6—tocopherols constituted less than 5% of total

110

tocopherols as opposed to 34% in stressed 3SS::HPT1. This change was only marginal in
3SSzzy-T MT parental lines, where y— and 5—tocopherols accounted for less than 6% of total
tocopherols compared to 14% in wild type leaves during high light stress (Figure 24).
These results suggest that y-TMT activity and not SAM levels limits a- and B-tocopherol

synthesis in stressed wild type and 358::HPT1 plants.

 

 

 

g. 13 — Ialpha 15.3 "5'8
' Elbeta
E 12 1
2 15 ‘ Igamma '
o
Idelta
E 12 -
8.6

CD
2 9 4 71 67
3 ii
a. 5 "
§ r

3 - l
1— y I i

o _. :7; _h (V.- g .3

   

wt 19 49 11 4911119111
r-TMT HPT1 rmnnprr

 

 

Figure 24. Tocopherol levels and composition in stressed wild type, 358::HPT1/3SS::y-
T M T double overexpressers, and the corresponding transgenic parents. Plants were
stressed for 15 days and individual tocopherols were analyzed by normal phase HPLC.
After 15 days of high light stress, 358::HPT1 leaves accumulated 2-fold higher total
tocopherol levels relative to wild type. The conversion of y- and 6-tocopherols to a- and B-
tocopherols occurred to a much greater extent in 35S::HPTl/3SS::y-TMT than in
3SS::HPT1, suggesting that y—TMT activity limits a- and B-tocopherol synthesis in
3SS::HPT1 leaves during high light stress.

Discussion

Chloroplasts generate increased levels of excited chlorophylls and ROS including
singlet oxygen, superoxide anions, and hydroxyl radicals during stress (Fryer, 1992;
Mittler, 2002). Plants have developed numerous highly regulated mechanisms for

protection from excessive ROS including a variety of different antioxidants and ROS

lll

detoxifying enzymes that are generally up—regulated in their levels and/or activity during
stress (Foyer et al., 1994). Tocopherols are thought to represent a key antioxidant
protecting polyunsaturated fatty acids from photooxidation by quenching and scavenging
ROS and alkyl peroxy radicals generated during photosynthesis (Fryer, 1992). In plants,
tocopherol levels can change significantly depending on tissue type, developmental stage,
and environmental conditions (Rise et al., 1989; Havaux et al., 2000; Munne-Bosch and
Alegre, 2002a). We have used light stress as a treatment to increase tocopherol synthesis in
wild type and 358::HPT1 plants in order to address basic questions about regulation of the
pathway in response to stress.

We have shown previously that phytylation of HGA is a limiting step in tocopherol
biosynthesis in non-stressed Arabidopsis leaves (Collakova and DellaPenna, 2003). To
determine whether HPT activity is also limiting for tocopherol synthesis under stress
conditions, wild type and 3SS::HPT1 plants were subjected to high light stress and
analyzed. Stressed 358::HPT1 leaves accumulated 2- to 4-fold elevated total tocopherol
levels relative to wild type throughout the experimental time course. These increases
correlated with significantly elevated HPT mRN A levels and HPT specific activity in
3SS::HPT1 compared to wild type leaves during high light stress. Collectively, these results
indicate that, as in unstressed leaves, HPT activity limits tocopherol synthesis in stressed
wild type leaves. It appears that HPT limits plant tocopherol biosynthesis under both
normal and stressed conditions.

While a several-fold difference in total tocopherol levels between stressed wild type
and 3SS::HPTI plants was maintained during the experimental time course, the 15-day high
light treatment also resulted in an 18- and 8-fold increase in wild type and 3SS::HPT 1,
respectively, relative to unstressed controls. In non-stressed wild type and 358::HPT1
plants, tocopherol levels nearly doubled during the 15-day time course of the experiment.
This increase is likely due to aging of the 6-week-old plants during the experiment as

increasing plant age is associated with a modest increase in total tocopherol content (Rise et

112

al., 1989; Molina-Torres and Martinez, 1991; Tramontano et al., 1992). Despite this
increase, the absolute tocopherol levels in unstressed wild type and 3SS::HPT1 plants
remained very low relative to stressed plants. In wild type leaves, both HPT mRNA levels
and HPT specific activity were up-regulated in response to high light treatment throughout
the experimental time course. In stressed 358::HPT1 lines, HPT mRNA levels were not
significantly elevated, whereas HPT specific activity was, suggesting that the HPT enzyme
may also be regulated at the post-transcriptional level during high light stress. Collectively,
these results indicate that elevated HPT expression and activity contribute significantly to
the increased tocopherol accumulation in stressed wild type and 358::HPT1 leaves.

Although HPT specific activity is clearly a key factor regulating total tocopherol
synthesis during stress, it appears that other factors are also involved. This hypothesis is
supported by the observation that HPT specific activity did not correlate with the linear
increase in total tocopherol levels at all time points. First, HPT specific activity was similar
in wild type leaves stressed for 3 and 6 days, though total tocopherol levels nearly doubled
during this period. Similarly, in 35S::HPT1 lines, HPT specific activity was reduced
significantly in 6-day stressed plants relative to those stressed for 3 days, whereas
tocopherol levels doubled during this time. Finally, though HPT specific activity increased
nearly 3-fold in wild type leaves stressed for 6 days, it still did not reach the levels of non-
stressed 358::HPTI plants, yet total tocopherol levels in these stressed wild type plants were
2.3-fold higher than non-stressed transgenics. These results suggest that one or more
HPT-independent limitation(s) that cannot be overcome by HPT1 overexpression alone in
non-stressed Arabidopsis plants are alleviated by high light stress treatment. Increased
tocopherol levels in stressed leaves may simply be due to the increased availability of
substrates for tocopherol synthesis as well as up-regulation of other tocopherol pathway
enzymes upstream of HPT.

HPT catalyzes the committed step of tocopherol synthesis, condensation of two

potentially limiting substrates, HGA and PDP. The aromatic precursor HGA is derived

113

from prephenate or tyrosine, both products of the Shikimate pathway (Threlfall and
Whistance, 1971; Fiedler et al., 1982; Lopukhina et al., 2001). In plants, the Shikimate
pathway has been shown to be regulated at the transcriptional level and globally up-
regulated during biotic and abiotic stresses (Gorlach et al., 1995; Herman and Weaver,
1999; Diaz et al., 2001). Thus, the Shikimate pathway may provide an increased pool of
prephenate that is at least partially utilized for elevated tocopherol synthesis in stressed
plants. In addition, tyrosine is released during protein degradation and may also provide an
increased pool of HPP available for tocopherol synthesis during high light stress. Elevated
tyrosine catabolism in stressed plants is consistent with our findings that TAT, HPPD, and
HGAD mRNA levels were all up-regulated during stress in both wild type and 358::HPTI .
These results are consistent with prior studies that TAT expression was up—regulated in
Arabidopsis leaves during various biotic and abiotic stresses (Lopukhina et al., 2001;
Sandorf and Hollander-Czytko, 2002) and that HPPD mRNA levels were elevated during
senescence-induced stress in barley leaves (Klebler-Janke and Krupinska, 1997). Although
increases in HPPD mRNA levels are associated with elevated tocopherol levels during
stress, HPPD activity is probably a minor component regulating tocopherol biosynthetic
pathway because constitutive HPPD overexpression alone or in a HPT1 overexpressing
background had no effect on total tocopherol levels in high light stressed Arabidopsis leaves
(data not shown).

As with HGA, there are two known metabolic sources of the other HPT substrate,
PDP. First, excess PDP required for tocopherol biosynthesis may be synthesized de novo
from DXP pathway-derived IPP during stress. In plastids, flux through the DXP
isoprenoid biosynthetic pathway is regulated at the level of DXP synthase and DXP
reductoisomerase activities, which have been shown to limit isoprenoid biosynthesis in non-
stressed plants (Estevez et al., 2001; Mahmoud and Croteau, 2001). Enzymes downstream
of the DXP pathway, such as GGPSI and GGDR were not up—regulated at mRNA levels in

response to high light stress, suggesting that they are not major regulators of tocopherol

ll4

biosynthesis or, if they are, the activities of these two enzymes are translationally or post-
translationally regulated. The second potential source of PDP is phytol, which is derived
from chlorophyll degradation, a process involving several steps including the removal of the
phytol tail by chlorophyllase (Matile et al., 1999; Tsuchiya et al., 1999). Chlorophyllase
gene expression is induced by treatments with stress-associated compounds such as
coronatine and methyl jasmonate (Tsuchiya et al., 1999). A correlation between chlorophyll
degradation and tocopherol accumulation during leaf senescence has also been reported
(Rise et al., 1989). We observed a large decrease in chlorophyll levels in wild type and
358::HPT1 during high light stress, which coincided with the rise in tocopherol levels. If
one only considers degradation of existing chlorophyll, the maximal pool of phytol released
during the 15-day stress treatment would allow for the synthesis of ~ 6 nmol cm‘2 leaf area
of total tocopherols in wild type and 358::HPT1 leaves. While this possibility is intriguing,
it is still not clear what contribution chlorophyll degradation makes to tocopherol
biosynthesis during high light stress because we lack direct evidence for the incorporation
of chlorophyll-derived phytol into tocopherols.

The increase in total tocopherol levels was paralleled by a corresponding increase in
a-tocopherol levels in response to high light stress. However, large increases in the levels
of other tocopherols in stressed wild type and 3SS::HPT1 plants suggested that steps
downstream of HPT that regulate tocopherol composition, including TC, MPBQ MT, and
y—TMT, may limit (rt-tocopherol accumulation during stress. The absence of the
phytquuinol intermediates MPBQ and DMPBQ suggests that TC activity is sufficient in
both stressed and unstressed plants and the phytquuinol intermediates are rapidly converted
to the corresponding tocopherols, or rapidly degraded by unknown mechanisms. The fact
that TC mRNA levels are very low and not up-regulated significantly during stress suggests
TC is either very active and/or very stable in stressed Arabidopsis leaves. The presence of
[3- and 8—tocopherols in stressed wild type and transgenic Arabidopsis leaves suggests that

either MPBQ MT activity or SAM limits (rt-tocopherol synthesis in these plants. This

115

limitation appears to be more severe in 3SS::HPT1 leaves, which accumulate [3- and 8-
tocopherol at several-fold higher levels than wild type at all time points during stress. Based
on these results, it appears that the excess MPBQ produced by HPT in stressed transgenic
plants is not methylated to DMPBQ, but rather cyclized to form 5-tocopherol.

The final enzyme of the tocopherol pathway is y-TMT, which utilizes SAM to
generate a- and B-tocopherol from y- and 5-tocopherol, respectively (D'Harlingue and
Camara, 1985; Shintani and DellaPenna, 1998). Relatively high levels of y— and 5-
tocopherols were detected in stressed wild type and much higher levels in 358::HPT1
leaves, suggesting that yuTMT activity or SAM may be limiting for (rt-tocopherol synthesis
during high light stress. To differentiate these possibilities, tocopherol composition and
levels were analyzed in stressed transgenic plants overexpressing HPT1 and y-TM T in
combination (358::HPT1/3SSzzy-TMT). Conversion of nearly the entire pool of y- and 6-
tocopherols to a- and B-tocopherols, respectively, in stressed 358::HPT 1/3SSzzy-T M T
indicates that y-TMT activity, and not SAM, limits (rt-tocopherol biosynthesis during stress.
The persistence of small but similar amounts of y- and 8-tocopherols in stressed leaves of
wild type and all transgenic genotypes implies that these tocopherols are either not
accessible to y—TMT or accumulate at these levels for a distinct functional role.

The current study represents an initial step toward understanding the molecular and
metabolic regulation of tocopherol biosynthesis during high light stress in plants. We have
focused on determining the changes in steady-state tocopherol levels and pool sizes and
correlating this with changes in gene expression for various steps of the pathway. Because
we cannot yet address tocopherol turnover, our data do not represent total tocopherol
synthesis or fluxes through the pathway. HPT catalyzes HGA phytylation, the first and
committed step in the tocopherol pathway, and limits total tocopherol synthesis in both non-
stressed and stressed wild type Arabidopsis plants. Although stressed leaves showed
increased HPT specific activity relative to non-stressed leaves, HPT activity is not the only

component that contributes to the elevated total tocopherol levels observed during high light

116

stress. The differential production and availability of aromatic and prenyl diphosphate
precursors in non-stressed and stressed plants may also play an important role in regulating
tocopherol levels under various growth conditions. Up-regulation of the Shikimate pathway
and protein and tyrosine degradation may all contribute to the HPP pool used for elevated
tocopherol synthesis in stressed relative to non-stressed plants. Similarly, elevated
chlorophyll degradation and de novo isoprenoid synthesis may both contribute to the
increased PDP pool needed for tocopherol synthesis during high light stress. All enzymes
downstream of HPT (TC, MPBQ MT and y-TMT) regulate leaf tocopherol composition.
Our results indicate that methylation of both MPBQ and y-tocopherol, but not the
cyclization of tocopherol intermediates, also appears to limit (rt-tocopherol synthesis in

stressed wild type and 358::HPT1 Arabidopsis leaves.

Materials And Methods

Plant growth and high-light stress. The following lines were used in this study: wild
type Arabidopsis thaliana (ecotype Columbia) and the previously characterized
homozygous transgenic lines 35837-7” MT -18 and -49 (Shintani and DellaPenna, 1998) and
3SS::HPT1-11 and -54, and the corresponding homozygous double overexpressers
obtained by crossing these lines (Collakova and DellaPenna, 2003). To obtain a sufficient
amount of tissue for all analyses, seeds were planted 3 - 4 cm apart and grown at a 10-hour
photoperiod at 75 - 100 11E (22/19 °C day/night cycle) for 6 weeks. At 6 week age, plants
were transferred to high light (0.8 - 1 mE, lO—hour photoperiod) for stress experiments or
maintained at 75 - 100 11E. To exacerbate oxidative stress, a combination of high light, low
nutrient, and water stress was used such that plants grown at high light were watered every 2
days with distilled water. For simplicity, this combination of abiotic stresses is referred to
high light stress. At indicated time-points, tissue was harvested for HPLC, real-time PCR,

or HPT enzyme assays, within 3 to 5 hours after the start of the light cycle.

117

Prenyllipid and anthocyanin analyses. Each genotype was represented by three plants
and each plant was analyzed twice. Three representative leaf discs (total surface area: 1.507
cmz, 30 - 35 mg of fresh weight) per plant were harvested and subjected to lipid extraction
(Bligh and Dyer, 1959). Afier phase separation, an aliquot of the aqueous phase was
acidified with 0.1 N HCl and the amount of anthocyanins present was measured
spectrophotometrically at 520 nm as described previously (Merzlyak and Chivkunova,
2000). The organic phase was dried under vacuum and lipids were dissolved in hexane. An
aliquot of the resulting lipid extract was dried, dissolved in 90% (v/v) methanol, and used to
determine chlorophyll and carotenoid content by the spectrophotometric method of
Lichtenthaler (1987). Tocopherols were analyzed and quantified by normal phase HPLC as
described (Collakova and DellaPenna, 2003).

Real-time PCR. Tissue harvested from three representative plants was ground in liquid
nitrogen, total RNA isolated, and real-time PCR was performed as previously described
(Collakova and DellaPenna, 2003). The procedure was modified such that 3 pg of total
RNA for each sample was reverse transcribed in triplicate. Each reaction was diluted with 2
volumes of water and cDNA corresponding to 200 ng of total RNA was subjected to real-
tirne PCR using ABI PRISM Sequence Detection System 7000 (Applied Biosystems,
Foster City, CA). Standard curves were constructed for each gene and used to determine
the absolute mRNA levels. Relative mRNA levels of elongation factor EF-la were used to
normalize each sample. EF-la mRNA levels varied over a 2.5-fold range during the high
light treatment with no apparent trend (data not shown). Results from three independent
experiments are presented as average i SD for each line and time point. Taqman probes,

primers and their final optimized concentrations are presented in Table VII.

118

Table VII. Probes, primers and their final optimal concentrations used for real-time
PCR. Each Taqman probe was labeled with 6-fluorescein at the 5’- and the quencher
TAMRA at the 3’-end.

 

Primers/

 

, . Conc.

Enzyme Probes Sequence (5 -> 3 ) [nM]
Forward TCTCTAAAAGACI'ICTG‘I'I'IGCTATTCG 900
H PT Reverse AGTCGAGA I'I'I CGGGTTAATGC 900
Probe AAAGCCTCAGGCTGACCCGCAGT 1 75
Forward CAAGGAGTGTGAGGAATI‘AGGGATT 300
H P P D Reverse TCGTCGGCCTGTCACCTAGT 300
Probe ATGATCAAGGGACGTTGC’I‘I‘CAAATC’ITCA 1 50
Forward ACCAACCATCGAGGAGGAAA 900
HGAD Reverse TAAATACAGTCCTT ACAGCACCAACT G 900
Probe ATGACAAAATCAAGCAAGGCCACACCA l 75
Forward GTA'I'I'I'GAGCCTCATTGGCAGAT 900
TC Reverse TCACCGCCCCATTCTATCC 900
Probe ATGGCAGGAGGCC‘I'I'I'CCACAGG 200
Forward TCCGGTGAGAGTGGTTCGAG 900
GGP S 1 Reverse CTTGACCCGCCACTAACCC 900
Probe TTGGAGAATTGGCTAAAGCGATAGGAACAGA 200
Forward CAC I ' I AGGGAACACCCAACCA 900
GGDR Reverse AC 1 'l ACTATGAGGA' I ' I 'I AGCTGAGATGTATG 900
Probe AGAAATCCGGCGACACATCATCTCCA l 50
Forward AACCGAAAGCCAACG‘ I ' I ' I ' I 'G 900
TAT Reverse TCTTGTAGATGGAGCGGACTAGGT 600
Probe TTCCGAGTCCCGGCTTCCCATG 200
Forward GGCCAAGAGAGCCAATGATC 900
y—TMT Reverse CGCATCCGCAAC I'I'GGA 900
Probe CGGCTGCTCAATCACTCTCTCATAAGGCT 200
Forward CGAAC‘ I 'I CCATAGAGCAATATCGA 300
EF - 1 a Reverse GCATGGGTGTTGGACAAACTT 300
ACCACGGTCACGCTCGGCCT l 75

Probe

 

vfimfi“. '

Prenyltransferase assays. Homogentisate phytyl transferase assays were performed

using [U-'4C]HGA and PDP as substrates and crude chloroplast preparations from wild

119

type and transgenic plants as described previously (Collakova and DellaPenna, 2003). Each
high-light experiment involved analysis of control (0 days) and stressed (3 and 6 days) wild
type and 358::HPT1 plants. Chloroplast isolation and HPT activity assays of all plants
were performed the same day to ensure equal treatment of all samples in the experiment.
Thus, a set of 6-week-old plants grown under non-stress conditions were transferred to high
light and 3 days later another set of non-stressed plants was transferred to high light. After
additional 3 days, chloroplasts were isolated from a set of non-stressed, 3-day-stressed, and
6-day-stressed wild type and 3SS::HPTI plants. HPT assays with plants stressed for more
than 6 days could not be reliably performed because of inconsistent and insufficient
chloroplast yields obtained fi'om such plants. HPT specific activity was calculated by
subtracting background HPT activities obtained from control reactions containing
radiolabeled HGA without exogenously added PDP from the HPT activities when both
substrates were present. HPT specific activity is presented as relative HPT specific activity,
which is expressed as a fold difference relative to HPT specific activity of non-stressed wild

type chloroplasts.

120

SUMMARY AND FUTURE DIRECTIONS

Homogentisate phytyl transferase plays an important role in vitamin E biosynthesis,
because it catalyzes the committed step of the tocopherol biosynthetic pathway,
condensation of HGA and PDP. The isolation and biochemical characterization of the
genes encoding HPT in Synechocystis sp. PCC 6803 and Arabidopsis thaliana were
described in Chapter 1. Phenotypic characterization of the Synechocystis sp. PCC 6803
HPT disruption mutant lacking tocopherols (SLR1736::Km') is also presented. In Chapters
2 and 3, molecular and biochemical characterization of Arabidopsis transgenic plants
overexpressing HPT were presented to address the question whether HPT activity is
limiting for tocopherol synthesis in non-stressed and stressed Arabidopsis plants.

I identified the genes encoding SynHPT and AtHPT based on their similarity to
chlorophyll synthase from Synechocystis sp. PCC 6803, an analogous prenyltransferase in
chlorophyll biosynthesis. Both SynHPT and AtHPT utilized HGA and PDP, but not SDP,
to yield MPBQ indicating that HPT is specific to tocopherol, but not plastoquinone
biosynthesis. This hypothesis was further supported by the finding that the tocopherol-
deficient mutant SLR1736::Kmr showed wild-type PQ-9 levels. The gene encoding
homogentisate solanesyltransferase in cyanobacteria and plants remains to be identified.
There are several ORFs encoding prenyltransferases similar to SynHPT in Synechacystis
genome, which has been fully sequenced. In Synechacystis sp. PCC 6803, the ORFs
SLR1518 and SLL0057 encode 1,4-dihydroxy-2-naphtoate phytyltransferase involved in
phylloquinone synthesis and chlorophyll synthase, respectively (Addlesee et al., 1996;
Reategui et al., 1998; Addlesee and Hunter, 1999). The only remaining putative HST ORF
is SLR0096, which shows the highest homology to the E. coli hydroxybenzoate
octaprenyltransferase (Figure 25), which is known to catalyze the condensation of 4-
hydroxybenzoate and octaprenyl diphosphate in the biosynthesis of ubiquinone, an electron

carrier involved in mitochondrial respiration (Soballe and Poole, 1999).

121

Several lines of evidence suggest that SLR0096 encodes putative HST. Because
Synechacystis sp. PCC 6803 does not synthesize ubiquinones, both respiratory and
photosynthetic electron transport involve PQ-9 as an electron carrier in this cyanobacterium

(Friedrich and Weiss, 1997). In addition to SLR0096, Svnechocvstis genome contains

 

AtUBlA (NP567688) —-> Ubiquinone

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

AtPDsz (BABO3104) g f
GmHPT (AX046732) g g
— TaHPT (AX046740) §§
'1: OsHPT (AX046730) g g
ZmHPT (AX046726) E t.
Gm HPT (AX046736) ..
_‘1: OsHPT (AX046728) 2 c;
ZmHPT (AX046716) g g
—- "CAtHPT (AF324344) E g
GmHPT(AXO46734) 3 g
, AvHPT (ALR3448) '- t
1 SynHPT (slr1736)
RcChlG (P26170)
creme (CT1270) 2
— —_1: AsChlG (AJ277210) E
AtChIG (NP190750) §
TeChlG (TLL1539) g
Q

 

 

 

C AvChlG (ALL4480)
SynChlG (SLR0056)
EcUBlA (X66619) —> Ubiquinone

TeUBIA (TLRO402)
NpUBIA (NCOO3272)} Plastoquinone ?

0.1 _l
' ' SynUBlA (SLR0926)

 

 

 

 

 

 

 

Figure 25. Phylogenetic analysis of related prenyltransferases involved in tocopherol,
tocotrienol, plastoquinone, chlorophyll, and ubiquinone biosynthesis. Numbers in
parentheses represent GenBank accession numbers or ORF numbers from the online
database CyanoBase. Analysis was performed as described for Figure 12 in Chapter 1.
Arabidopsis UbiA was used as an outgroup. The transit peptides were not included in the
analysis. Plants: As — Avena sativa, At - Arabidopsis thaliana, Gm — Glycine max, Os -
OIjvza sativa, Ta — Triticum aestivum, Zm — Zea mays; Eubacteria: Anabaena sp. PCC
7120, Ct — Chlorobium tepidum TLS, Ec — Escherichia coli, Nos — Nostoc sp. PCC 7120,
Rc — Rhodobacter capsulatus, Syn — Synechocystis sp. PCC 6803, Te —

T hermosvnechococcus elongatus BP-l

122

other ORFs that show remarkable similarity to other E. coli ubiquinone biosynthetic
enzymes such as UbiX and UbiH (Dahnhardt et al., 2002). PQ-9 synthesis in
Synechacystis sp. PCC 6803 does not require HPPD, the enzyme involved in the formation
of the aromatic precursor HGA (Dahnhardt et al., 2002). HPPD ORF is absent from the
genomes of cyanobacteria that can synthetize PQ-9, but not tocopherols (Sattler et al.,
2003). Thus, PQ-9 synthesis is independent of HPPD and HGA and probably proceeds
through a ubiquinone-like pathway using chorismate as a precursor and the ubiquinone-like
solanesyltransferase SLR0096 in this cyanobacterium. Initially, feeding experiments using
various radiolabeled aromatic precursors in tocopherol and plastoquinone synthesis
including tyrosine, HPP, HGA, chorismate, prephenate, and hydroxybenzoate should be
performed with wild type Synechocystis sp. PCC 6803 and the deletion HPPD mutant
Aslr0090 (Dahnhardt et al., 2002) to delineate the cyanobacterial PQ-9 biosynthetic
pathway.

In contrast to Synechocystis sp. PCC 6803, HPPD plays a key role in plant
plastoquinone biosynthesis by providing HGA for HST (Norris et al., 1995; Norris et al.,
1998; Garcia et al., 1998). In Arabidopsis, HST is probably encoded by the PDS2 gene,
disruption of which leads to the lethal albino phenotype identical to that of the pdsl mutants
(Norris et al., 1995). It is unlikely that the PDS2 gene encodes SDP synthase or other
enzymes of isoprenoid synthesis upstream of SDP synthase, disruption of which would
also result in a lethal albino phenotype (Estevez et al., 2000; Mahmoud et al., 2001), because
the PDS2 gene product strongly resembles the prenyltransferases. Phylogenetic analysis of
prenyltransferases from several plants has identified a separate clade containing putative
HSTs, which includes PDSZ. This clade is more closely related to HPTs than to other
prenyltransferases such as 4-hydroxybenzoate polyprenyltransferases (ubiquinone
synthesis) or chlorophyll synthases (Figure 25). Confirmation that the PDSZ gene encodes
HST can be accomplished by cloning the PDS2 gene, showing the prenyltransferase activity

123

using radiolabeled HGA and SDP, and functionally complementing the Arabidopsis pds2
mutant.

SynHPT, but not AtHPT, was also able to catalyze the condensation of HGA and
GGDP in vitro to form MGBQ, a prenquuinol precursor in tocotrienol biosynthesis.
These results were surprising because Synechocystis sp. PCC 6803 does not accumulate
detectable levels of tocotrienols. Thus, the geranylgeranyl tails of tocotrienols or their
prenquuinol precursors may be efficiently reduced by GGDP reductase (ChlP) in
Synechocystis sp. PCC 6803 (Keller et al., 1998). Further research is required to determine
the significance of the GGDP utilization for tocopherol/tocotrienol biosynthesis in vivo and
the Synechocystis GGDR mutant, which is unable to produce PDP may be helpful in
addressing this question. The presence of tocotrienols in this mutant would suggest that
SynHPT can use GGDP as a substrate in vivo.

Besides AtHPT, we have identified numerous putative prenyltransferases in other
plants (DellaPenna et al., 2000). Based on their sequence similarity, these
prenyltransferases fall into two main functional clades (Figure 25). First, as mentioned
above, proteins similar to PDSZ are probably solanesyltransferases involved in PQ-9
synthesis. Second, sub-clades of closely related monocot and dicot prenyltransferases
similar to AtHPT are most likely involved in tocopherol biosynthesis. Some of the monocot
HSTs or HPTS may represent prenyltransferases that utilize HGA and GGDP as substrates
in tocotrienol biosynthesis. Additional partial clones that were identified in some monocots
and are not shown in the phylogenetic tree may be responsible for tocotrienol synthesis.
Biochemical analysis needs to be done to reveal substrate specificities of these putative
prenyltransferases. All these enzymes should be tested for the prenyltransferase activity in
vitro using PDP, GGDP, and SDP. To determine whether dicots retained their ability to
synthesize tocotrienols, some of the confirmed monocot geranylgeranyltransferases may be
overexpressed in a dicot such as Arabidopsis. If successful, this experiment will represent

the first step in the engineering of tocotrienol synthesis in plants. Tocotrienols have

124

cholesterol lowering properties in animals (Packer et al., 2001) and engineering dicot crops
for high tocotrienol content would provide novel sources of dietary tocotrienols.

In addition to wild type Arabidopsis, the geranylgeranyltransferase cDNAs should
also be transformed into the tocopherol-free background of the HPT T-DNA tagged
Arabidopsis mutant or GGDR antisense transgenic Arabidopsis plants, both isolated in Dr.
Dean DellaPenna’s laboratory (unpublished data). These plants may be particularly helpful
in determining whether the geranylgeranyl tail of tocotrienols or their intermediates is
converted to phytyl tail of tocopherols by GGDR and in testing for both geranylgeranyl and
phytyltransferase activities in vivo. Tocotrienols are present in seeds, but not in leaves in
monocots (Franzen and Haas, 1991), which suggests that the two types of tissues may be
metabolically distinct. These distinction could be a result of the differences in GGDP
availability or expression of the tocotrienol biosynthetic genes in the two tissues. Therefore,

both leaf and seed tocotrienol analyses should be performed in these transgenic plants.

Regulation And Metabolic Engineering of Tocopherol Biosynthesis in Arabidopsis

HPT catalyzes phytylation of HGA and appears to be a highly regulated enzyme in
the tocopherol biosynthetic pathway. In Chapters 2 and 3, I presented data showing that
HPT activity is limiting for tocopherol synthesis in non-stressed wild type Arabidopsis
leaves and seeds as well as in leaves subjected to high light stress. Overexpression of HPT
in Arabidopsis resulted in increased HPT mRNA levels and activity and subsequently in
elevated total tocopherol levels in non-stressed leaves and seeds compared to wild type.
Similarly, stressed 358::HPTI leaves exhibited increased HPT activity and accumulated up
to 3.8-fold increased total tocopherol levels relative to stressed wild type Arabidopsis leaves.
Analysis of prenquuinol and tocopherol composition in stressed wild type and 3SS::HPT1
leaves was useful in identifying two other enzymatic steps that limit (rt-tocopherol synthesis,

MPBQ and y-tocopherol methylation reactions. Collectively, these data contribute

125

significantly to the current knowledge about the regulation of tocopherol synthesis.
However, there are several questions that need to be addressed about the regulation of
tocopherol biosynthetic pathway under normal and stress conditions.

From the developmental point of view, HPT activity decreases with age in non-
stressed wild type and 35S::HPT1 leaves as young, 2-week old plants exhibited higher HPT
activity than 4— and 6-week old plants. However, a more detailed analysis during initial
stages of plant development (during and within the first 2 weeks after seed germination) and
in senescing plants (more than 8 weeks old) would complement the current knowledge
about tocopherol biosynthetic pathway and its regulation. Wild type Arabidopsis seeds
contain approximately 4—fold more tocopherols than leaves, when expressed on a dry weight
basis. Seed germination is accompanied by an increase in oxidative stress as photosynthetic
apparatus begins to function (Lichtenthaler, 1969). Similarly, leaf senescence is also
associated with increased oxidative stress in chloroplasts (Munne-Bosch and Alegre,
2002c). Increased oxidative stress translates into increased tocopherol degradation and a
need for tocopherol synthesis to sustain oxidative damage. As described in Chapter 2, dry
Arabidopsis seeds contain detectable HPT activity, suggesting that HPT may contribute to
the de novo tocopherol synthesis in young developing seedlings. It would be interesting to
study the role of HPT in tocopherol synthesis and to identify other possible bottlenecks of
the pathway in germinating seeds, developing seedlings, and senescing plants by using wild
type and 3SS::HPT1 Arabidopsis lines.

In Chapter 3, steady-state levels of tocopherols and their prenquuinol intermediates
in control and high light stressed wild type and 3SS::HPT1 leaves are used to shed some
light into regulation of the tocopherol pathway. There are several processes that may
contribute to the steady-state tocopherol levels in chloroplasts: tocopherol biosynthesis,
degradation, and recycling. Recycling of the tocopheroxyl radicals back to tocopherols has
been demonstrated in vitro, but there is only indirect evidence that it occurs in vivo (Munne-

Bosch and Alegre, 2002b). The lack of evidence for the in vivo tocopheroxyl radical

126

recycling is primarily due to inability to distinguish between tocopherol recycling and
scavenging of ROS by ascorbate (Kagan and Packer, 1994).

Tocopherol oxidation yields the corresponding tocopheryl quinone. In non-stressed
tissues, tat-tocopheryl quinone comprises from 4 to 10% a-tocopherol levels (Lichtenthaler,
1969; Lichtenthaler et al., 1981; Michalski and Kaniuga, 1981). The levels of (rt-tocopheryl
quinone increased up to 3-fold in stressed tomato leaves relative to non-stressed leaves
(Michalski and Kaniuga, 1981). Therefore, total tocopherol levels presented for the high
light time course for both wild type and 358::HPT1 are likely underestimated and do not
represent total tocopherol synthesis because tocopheryl quinone levels were not measured.
In addition, the steady-state tocopherol levels alone cannot be used to determine flux
through the pathway, hence a detailed flux analysis in non-stressed and stressed wild type
and 358::HPT1 leaves should be performed. This analysis would require an effective
separation, identification, and quantitation of major tocopherol biosynthetic precursors and
breakdown products. Arabidopsis tocopherol biosynthetic mutants in HPT, TC, MPBQ
MT, and y—TMT, which are currently available in Dr. Dean DellaPenna’s laboratory, should
be helpfull in identifying the individual intermediates in tocopherol biosynthesis and
degradation.

Real-time PCR experiments suggested that some tocopherol biosynthetic genes are
up-regulated during high light stress. All genes encoding the enzymes of the Shikimate
pathway were also transcriptionally up-regulated during biotic and abiotic stress (Jones et
al., 1995; Bischoff et al., 1996; Diaz and Merino, 1998; Gorlach et al., 1995; Batz et al.,
1998). To obtain a complete picture about the transcriptional regulation of the whole
pathway during stress, all genes, including these from the shikirnate and DXP pathways as
well as the genes encoding TAT, chlorophyllase, and tocopherol enzymes presented in
Figures 3, 4, and 5 should be used in a microarray experiment. Total RNA isolated from
leaves of non-stressed wild type Arabidopsis plants and those subjected to various stress

treatments including high light and JA can be reverse-transcribed, labeled, and utilized to

127

probe the microarray slide. A global induction of transcription of the tocopherol-related
genes in stressed plants would indicate an existence of one or more transcriptional
regulators in tocopherol biosynthesis.

Valuable data contributing to the understanding of the regulation of tocopherol
biosynthetic pathway in Arabidopsis plants under normal and stressed conditions are
presented in this dissertation. As outlined in this section, additional research is required to
further characterize mechanisms that regulate plant tocopherol levels. Besides studying
these mechanisms, future tocopherol research should also focus on determining role(s) of

tocopherols in plants and interaction of tocopherols with other antioxidants.

128

APPENDIX

EXPERIMENTAL PROCEDURES

In this section, procedures used in this study are described in a great detail such that
all experiments presented can be repeated using these protocols without the necessity of
consulting other publications. Unpublished observations or problems that one may
encounter when using some of the procedures or their limitations are also presented. All

relevant primers, their sequences, and use are summarized in Tables VIII and IX.

Chemicals And Standards

Large-scale quinone extraction. The purified PQ-9 standard was used to create standard
curves for PQ-9 quantification in Synechocystis sp. PCC 6803, while 2-DMPQ-9 was used
as an internal standard for quantitative quinone determinations. PQ-9 and 2-DMPQ-9
standards were extracted from Iris hollandica bulbs and purified by both TLC (Pennock,
1985) and HPLC (Johnson et al., 2000). PQ-9 can also be extracted from leaves of many
plants such as spinach, peas, or ficus, but due to the lack of contaminating chlorophylls, Iris
hollandica bulbs are an ideal quinone source. In addition, other quinones such as PQ-8, 2-
DMPQ-8, and ubiquinones can also be obtained from these bulbs. Bulbs were cleaned and
cut into small pieces. Tissue was frozen in liquid nitrogen and stored at - 80 °C or used
immediately for lipid extractions. After adding 100 mL of 50% methanol (v/v) to 50 g of
bulbs, tissue was homogenized in a blender. The homogenate was transferred to a clean
glass beaker containing 50 mL of chloroform. After 5 - 10 minute incubation at room
temperature, the homogenate was filtered through 6 layers of miracloth and the filtrate was
centrifuged at 8000 g for 10 minutes. The organic phase (bottom phase) was dried using a

rotary evaporator at 30 0C, lipids were dissolved in l - 2 mL of ethylacetate and quinols

129

were oxidized by shaking with 50 mg of silver oxide in glass tubes at room temperature for
2 h. Lipids were subjected to separation by TLC (Silica) in 20% ethyl ether (v/v) in
petroleum ether. PQ-8 and -9 migrated together in this TLC system (Rf 0.88; right below [3- .
carotene). 2-DMPQ-8 and -9 also migrated together, but at Rf 0.77. Silica corresponding
to the two areas was scraped and transferred to 5 mL syringe columns; quinones were then
eluted with 5 volumes of ethyl ether, dried under nitrogen, and subjected to reverse phase
HPLC. The following gradient was used to purify these prenquuinones (Johnson et al.,
2000): isocratic flow from 100% methanol for 10 minutes changed to a gradient of 3% (v/v)
methanol in isopropanol within next 20 minutes, which was held for additional 10 minutes,
all at flow 0.5 mL min". The column was equilibrated for 15 minutes with 100% methanol
at 2 mL min". Quinones were dried, dissolved in 0.5 mL of hexane, and stored at - 20 °C
for at least 3 years without detectable degradation. Purity of the quinones was confirmed by
HPLC and the PQ-9 concentration was determined spectrophotometrically at 255 nm in
hexane using 8 18,030 M" cm" (Pennock, 1985).

BG—ll medium. BG-ll liquid medium can be prepared as follows: One liter of BG-ll
contained 10 mL 100X BG-ll and 1 mL of each 1000X solutions. 100X BG-ll without
iron, phosphate, carbonate hadl49.6 g NaNO3, 7.5 g MgSO4, 3.6 g CaC12.2HzO, 0.6 g citric
acid, 1.12 mL 0.25 M NaEDTA pH 8, 100 mL trace minerals per 1 L. Trace minerals were
present at 2.86 g H3BO3, 1.81 g MnC12.4HZO, 0.22 g ZnSO4.7HzO, 0.39 g
NazMoO4.2HzO, 0.079 g CuSO4.5HzO, 0.049 g Co(NO3)2.6H,O per 1 L. Other
components included 1000X stocks of 6 mg mL" fenic ammonium citrate, 20 mg mL"
NazCO3, 30.5 mg mL" KzHPO4. For agar plates, 10 mL 1M TES/NaOH buffer pH 8.2, 3
g sodium thiosulfate, and 10 g phytagar were added. For photoheterotrophically grown

cyanobacteria, glucose was added to a final concentration 15 mM.

130

Plasmids And Mutants

Generation of constructs, mutants, and transgenic plants. Primers SLR1736F and R
(Table VIII) were used to amplify the SLR1736 ORF (GenBank accession number

BBA17774) from Svnechocystis genomic DNA using Vent DNA polymerase (New

Table VIII. PCR primer pairs used in this study.

 

Sequence.(from 5’ to 3’) of

 

 

Name primer parrs Thrs primer parr was used to.

SLR1736F TATTCATATGGCAACTATCCA Amplify SLR1736 ORF in
AGCTI'ITTG Synechocystis and to show the

SLR1736R GGATCCTAATTGAAGAAGAT absence of wild type copies of
AC1" AAATAGTTC this ORF in the SLR1736::Kmr

mutant.

ATI736F TTGT'ITI'CAGGCTGTTGTTGC Amplify a part of AtHPT
AGCTCTC genomic sequence for use as a

ATI736B CGT'ITCTGACCCAGAGTTAC probe to screen an Arabidopsis
AGAGAATG cDNA library.

HPTF CCATGGAGTCTCTGCT CT C Amplify full-length HPT1

HPTB GGATCCCAAGCAGAGACTTC cDNA for protein expression in
'ITTACC E. coli.

HPTAF AGCCATGGGTTGTGATTC Create a truncated version of

HPTB GGATCCCAAGCAGAGACTT C AtHPT lacking the first 37 aa.
TITACC

HPT96 CCATGGACTCGTTAGATGCG Create a truncated version of

HPTB GGATCCCAAGCAGAGACTT C AtHPT lacking the first 96 aa.
TTTACC

 

“$555!”. -

England Biolabs, Beverly, MA). The following program was used: 95 °C for 5 minutes (1
cycle); 95 °C for 45 seconds, 45 °C for 45 seconds, and 68 0C for 45 seconds (5 cycles); 95
0C for 45 seconds, 52 0C for 45 seconds, and 72 °C for 45 minutes (30 cycles); 72 °C for
10 minutes (1 cycle). Note that no PCR products were obtained without the 5-cycle
“priming”. ATG in the SLR1736F represents the start codon of the SLR1736 ORF. The
Ndel and BamHI sites (underlined) were engineered in the two primers to facilitate firrther

sub-cloning procedures for protein expression. The amplified SLR1736 ORF was sub-

l3l

cloned into the EcoRV site of pBluescript 11 KS (+). The resulting construct pKS 1736 and
pUC4K (Taylor and Rose, 1988) were digested with Mfel and EcoRI. Ligation of
pKSl7 36 with the kanamycin resistance cassette released from pUC4K yielded two
constructs, one with the same (pKS-l736-KanF) and one with the opposite (pKS-1736-
KanR) orientation of the kanamycin resistance cassette relative to the SLR1736 ORF. Both
constructs were used to transform wild type Synechocystis sp. PCC 6803 and generate
disruption mutants by homologous recombination (Williams, 1988). Wild type cells (0.1
mL, 109 cells mL", OD730 of 0.25 corresponds to 108 cells mL") were mixed with the
knockout construct DNA (~ 1 pg) and incubated at 20 - 30 pE at 30 °C with shaking. After
a 4-hour incubation, the cells were pelleted and plated on sterile nitrocellulose filters
(without detergent) placed onto BG-ll plates with 15 mM glucose and without kanamycin.
After 2 days of incubation at 20 - 30 pE at 30 °C, the filters were transferred onto fresh BG-
11 plates containing 15 mM glucose and 15 pg mL" kanamycin. Selected transforrnants
were sub-cultured on kanamycin containing media for 6 plating cycles and the absence of
wild type SLR1736 gene c0pies in the mutants was confirmed by PCR using SLR1736F
and R primers followed by Southern blot analysis. Because there was no phenotypic
difference between the two orientation disruption mutants (data not shown), the disruption
mutant derived fiom pKS-1736-KanF (referred to hereafter as the SLR1736::Kmr mutant)
was used for firrther analyses.

A single genomic clone, F19F24, was identified in the Arabidopsis thaliana
database based on its protein similarity to the SLR1736 protein. Primers ATI736F and B
(Table VIII) were used to amplify a 977-bp fragment from F19F 24 with the following PCR
program: 95 0C for 5 minutes (1 cycle); 95 °C for 45 seconds, 54 °C for 45 seconds, and 72
0C for 1 minute (30 cycles); 72 0C for 10 minutes (1 cycle). The resulting PCR product
was utilized as a probe to screen a pBluescript II SK-based Arabidopsis seed cDNA library
(a gift of Dr. John Ohlrogge, Michigan State University, East Lansing, MI). The longest of

15 positive clones (#1) was sequenced (see Table IX for sequencing primers) and

132

designated pSKHPTI (Figure 26A), which is also labeled as pSK-ATI736. The HPT1

gene encodes a prenyltransferase similar to SLR1736, AtHPT.

A
1.7 kb

 

 

F W
Sacl EcoRl Xhol Kpnl

 

 

p358::HPT1
15.43 kb

      
 

Figure 26. Constructs usedin this study. A, pSKHPTI B, p358: HPT1 Amp— ampicillin;
kb— kilobases; LB— left border; ori — origin of replication; RB — right border; Sp —
spectinomycin; St — streptomycin

For protein expression in E. coli, the entire SLR1736 ORF was released from
pKSl736 by Ndel and BamHI digestion and sub-cloned into Ndel/BamHI digested
pET30b (Novagen) to create pSynHPT. Similarly, primers HPT1 and 2 (Table VIII) were
used to amplify the firll-length Arabidopsis HPT1 gene from pSKHPTI. To facilitate sub-
cloning into the pET vectors, the Ncol and BamHI sites were introduced into these primers
(Table VIII, underlined). The PCR product was sub-cloned into the EcoRV site of
pBluescript 11 KS (+) to generate pKS-HPT. After digestion of pKS-HPT and pET3d with
N001 and BamHI and ligation, pAtHPT was generated. The pSynHPT and pAtHPT
constructs were transformed into BL-21 (DE3) cells (Novagen, Milwaukee, WI) to express
both prenyltransferases. for the enzyme assays. For enzyme activity purposes, two

additional truncated versions of AtHPT were used. The first truncation removed the first 37

133

amino acids corresponding to a transit peptide predicted by PSORT (primers used: HPTAF
and HPTB) and the second truncation removed the first 96 amino acids corresponding to a

putative transit peptide missing from SynHPT, (primers used: HPT96 and HPTB).

Table 1X. Sequencing primers used in this study.

 

 

_.-—.~wa

 

 

 

Primers for SynHPT

1736T3S CCAGAGTAAGGCTAATAAGCAC
l 736T7S TGACCGCATCAATAAGCC
Primers for AtHPT

ATl 736T3C CGTTTATGAGCTT’ITI‘CTCTGTCG
ATI 736T7C GACATATT’ITTGCAGTCTGCC
0AT1736F TTCGAGTAAAGTTGTCGC
1F1736A AGAGACTCGTTAGATGCG
2FAT1 736 T'ITCACTGGCATCTTGGAGG
3FAT1 736 TGCAGCAATGTGTATCCTCG
4FAT l 736 CTCGCAACAACTTTGTGGGC
0ATI 736R TGCCAATGGAAGATAGGG
lRATl 736 TTCCAACAAAACCCACCAGC
2RAT1 736 AAAATGCTAAGCACTGTGCC
3RAT1 736 GTACCGAGCATGAAACTCAC
4R1736 GCCACTGCGTTTATGAGC
4R1736A AGTGGCGAAAATAAGAGG
SRATl 736 CAAGCAGAGACITCT'ITACCC

 

To generate transgenic plants in Arabidopsis overexpressing the HPT1 gene,
p3SS::HPT1 (Figure 26B) was created by sub-cloning the cDNA encoding HPT fiom
pSKHPTI into pART7 and then into pMLBART, a pART27-based vector (Gleave, 1992).
Both pSKHPTI and pART7 were subjected to digestion with SacI and Kpnl and
subsequent ligation to yield pARTHPTI. Digestion of pARTHPTI with NotI released a
fragment containing the CaMV 35S rRNA promoter followed by the HPT1 gene in the
sense orientation. This fragment was ligated into the Notl site of pMLBART containing the
bar gene for selection of transformed plants on glufosinate. The resulting construct

p3SS::HPTI was introduced into Agrobacterium tumafaciens. 358::HPT1 T0 sense plants

134

were obtained by transforming wild type Arabidopsis plants (ecotype Columbia) with
A grobacterium containg p3SS::HPTI using the floral dip method (Clough and Bent, 1998).

Briefly, the floral dip medium contained 5% sucrose and 0.05% Silwet L-77. Plants
were submerged into this medium for ~ 5 seconds and left covered overnight in dark to
keep high humidity. Tl seeds were planted in soil and seedlings were sprayed with the
glufosinate solution (120 mg of ammonium glufosinate and 0.25 mL of silwett per 1 liter of
water) three times after their germination to ensure the elimination of wild type seedlings:
before and after most of them developed the first pair of true leaves and one more time after
the sixth leaf development (it takes about 5 days to visually observe the effects of the
herbicide sensitivity). Surviving plants (66 independent transformants) were analyzed for
tocopherol content. Lines 3SS::HPT1-8, ll, 16, 21, 35, 37, 53, 54 showed increased leaf
tocopherol levels and their T2 plants segregated 3:1 for glufosinate resistance. These lines
were carried through to the homozygosity. Lines 358::HPT1-ll and -54 were selected for
further analyses and for the crosses with 35$er M T-18 and —49, the two best
characterized Arabidopsis lines overexpressing y-T M T under the control of 358 CaMV
promoter (Shintani and DellaPenna, 1998).

Seed from crosses #9 (3SS::HPT1-ll X 3SS::y-TMT-49), #14 (3SS::HPT1-54 X
358::y-TMT-18), #16 (35$:2HPT1-54 X 3SSzzy-TMT-49), and #20 (358::HPT1-11 X
3SS::y-TMT-l8) were surface sterilized, plated on MS plates containing 50 pg mL" Km,
transferred to soil, and sprayed with glufosinate after 5 days of adaptation to soil to select
for dual resistance. These plants were hemizygous for Km and glufosinate resistance.
Their seed was germinated on plates and 50 - 60 seedlings for each cross were selected for
dual resistance as described above. Seeds of each of these plants were harvested
individually and planted on soil. After glufosinate selection, seeds of the lines homozygous
for glufosinate resistance were plated on MS plates containing Km. Double homozygotes

were selected for crosses #9, #14, #16, and #20. In crosses #14 and #16, Km and

135

glufosinate resistances were linked, suggesting that 358::HPT1-54, 3SSzzy-TMT-18, and

358::y-T MT -49 are localized on the same chromosome.

Oxygen Evolution Rates

Oxygen evolution rates. Liquid cultures of photoautotrophically grown wild type and
mutant Synechacystis cells were washed twice with fresh BG-ll medium and resuspended
in BG-l 1 medium. For O, evolution experiment, the cells were diluted to a concentration of
3 mg chlorophyll per mL. The cells were incubated in the dark for l h. Cell suspension ( 1
mL) was placed into the cell and O, consumption was measured in the dark for one minute.
Subsequently, the cells were exposed to high light for 5 minutes and three different light
intensities were used: 0.75, 2 and 5 mE. O, measurements were performed with a Clark-
type electrode at 25 0C using a Hansatech CBl-D3 recording unit with Minirec recording
software (Hansatech Instruments, King’s Lynn, England). The O, evolution rate was
calculated from the slope within the linear region of the curves.

Gloves must be worn while handling the electrode. The electrode was prepared as
follows: A square of 1.5 cm cigarette paper was pre-wet in saturated KCl solution, placed
on the top of the electrode, and then covered by a same sized piece of S4 PTFE membrane
(Hansatech Instruments). The small rubber O-ring was used to secure the paper and
membrane to the electrode without any bubbles or membrane damage. The electrode was
attached to the cell and the large rubber O-ring was used to prevent leakage from the cell at
the junction with the electrode. Temperature was controlled by a thermostat, while a
magnetic stirrer was used to ensure homogeneity of the cell suspension. The electrode was
allowed to equilibrate for l - 2 h in air-saturated water or until a stable maximal baseline
(1400 - 1600 U) was observed. Sodium dithionate was added to the cell to remove O, and
to determine the minimal baseline (~ 40 U). If the electrode was prepared properly, the

response to sodium thiosulfate should be very fast and the reading of the lower baseline

136

should be obtained within less than 1 minute. Because sodium dithionate tends to bind to
the electrode, it was quickly removed by washing with air-saturated water. O, evolution
rates obtained from dark measurements were subtracted fiom the rates obtained during the
high light treatment and the resulting values represent the net 0, evolution rates. The

following equation was used to calculate 0, evolution rates [pmol O, mg" Chlorophyll h"]:

O, evolution rate = (S X 60 X 60 X dy)/(Ay X C),

where S is the solubility of O, in water at 25 °C and corresponds to 0.253, dy is the slope of
the linear response obtained from measurements collected every second, Ay can be obtained
by subtracting the minimal baseline from the stable maximal baseline, and C is the total

chlorophyll amount in mg.

Prenyllipid Analyses

Tocopherol analyses. Wild type Synechocystis and the SLR1736::Kmr mutant was grown
on BG-ll plates either photoheterotrophically (with 15 mM glucose) or
photoautotrophically (without glucose) at 20 - 30 pE at 30 °C for 2 weeks. The cells (15 -
20 mg) were harvested and their lipids were extracted in eppendorf tubes by adding 200 pL
of 2:1 methanolzchlorofonn (v/v). Tocol (50 - 100 ng) was added and the cells were
disrupted by grinding with a pestle. Chloroforrn (70 pL) and water (200 pL) were added
and the tubes were centrifuged at 13,000 rpm for 5 minutes after vortexing. The organic
phase was transferred to a fresh tube and dried under vacuum. The lipids were dissolved in
100 pL of hexane and 10 pL of each sample was withdrawn for chlorophyll determination
(Lichtenthaler, 1987), while 50 pL of the lipid extract was subjected to HPLC (Hewlett
Packard 1100, Wilmington, DE) on a LiChrosorb 5 Si60A 4.6 X 250 mm normal phase

column (Phenomenex, Torrance, CA) at 42 °C and 1 mL min" using a gradient of isopropyl

137

ether and hexane (v/v) according to Syvaoja et a1. (1986). Within the first 30 minutes,
isopropyl ether content was increased from 8% to 18% and kept at 18% for an additional 10
minutes. The column was then equilibrated with 8% of isopropyl ether in hexane (v/v) for
15 minutes. Tocopherols were detected by fluorescence using 290 nm excitation and 325
nm emission.

To analyze tocopherol content in non-stressed wild type and transgenic Arabidopsis
leaves and seeds, plants were grown in a l6-hour photoperiod (70 - 100 pE) at 22/19 °C
day/night cycle. Lipids from 30 - 35 mg of leaf tissue or 10 - 15 mg of seeds were
extracted in the presence of butylated hydroxytoluene (2 mg mL") to prevent tocopherol
degradation (Bligh and Dyer, 1959). Tocol was used as an internal standard. For leaf
analysis, tocopherols were separated on a reverse phase HPLC (Clg, ODSZ, 4.6 X 250 mm,
Column engineering, Ontario, CA; Shimadzu VP HPLC system, Japan) using an isocratic
solvent system of 5% (v/v) isopropanol in methanol at 2 mL min". For seed analysis, a 10-
minute isocratic method using 17 % di-isopropyl ether in hexane (v/v) at 42 °C at 2 mL min'
' was utilized on a normal phase HPLC system (LiChrosorb 5 Si60A 4.6 X 250 mm Silica
column, Column engineering, Ontario, CA and HP 1100 series HPLC system, Hewlett
Packard, Wilmington, DE). Tocopherols were detected by fluorescence at 290 nm

excitation and 325 nm emission.

Real-Time PCR

Real-time PCR. Plants (wild type and 358::HPT1 #11 and 54) were grown at a lO-hour
photoperiod at 75 - 100 pE for 6 weeks. Tissue from three representative plants was
harvested 2 - 3 hours after the start of the light cycle and immediately frozen in liquid
nitrogen. Total RNA was isolated and any contaminating genomic DNA was removed by
treatment with RQl-RNase free DNase (Promega, Madison, WI) and re-extracted with

phenol and chloroform to remove the DNase. Total amount of 9 pg of total RNA was

138

reverse transcribed to generate cDNA in three 30-pL reactions for each sample using a
TaqMan kit according to manufacturer recommendations (Applied Biosystems, Foster City,
CA) in a 96-well format. Briefly, each 30-pL reaction contained 3 pL 10X RT buffer, 6.6
pL 25 mM MgCl,, 6 pL dNTPs, 1.5 pL oligo dTm, 1.2 pL RNasin, 0.75 pL Multiscribe
Reverse Transcriptasem, and 3 pg total RNA (in 10.95 pL water). The reactions were pre-
incubated at 25 °C for 10 minutes and reverse transcription was carried out at 48 °C for 30
minutes. The enzyme was inactivated by incubation of samples at 95 °C for 10 minutes.
An aliquot of cDNA corresponding to 200 ng of total RNA was used in each TaqMan real-
time PCR assays (Applied Biosystems). Each 30-pL reaction contained 15 pL of 2X
mastennix purchased from Applied Biosystems, 3 pl of each primer and the corresponding
probe, and 6 pL cDNA. Standard curves were constructed for each gene and used to

calculate the corresponding mRNA concentrations.

Prenyltransferase Assays

Prenyltransferase Assays Using E. coli Protein Extracts

Preparation of the E. coli protein extracts (adapted from pET vector manual). E.
coli cells harboring pETl736, pETAtHPT, pACHPPD, and vectors alone (pET3d and
pET30b) were grown overnight at 30 °C with shaking. Each of these cultures was used to
inoculate 50 mL LB media containing appropriate antibiotics (50 mg L" kanamycin for
pET30b and pETAtHPT and 100 mg L" ampicillin for pET3d, pETl736, and pACHPPD).
The cells were grown for 3 - 4 h at 30 °C with shaking until the cell density reached OD600
0.6. With the exception of the cells harboring pACHPPD, 240 mg L" IPTG was added to
induce protein expression from the lacZ operone. Cells containing pETl 736 and pET3d
were shaken at 30 °C, while those transformed with pETAtHPT, pACHPPD, and pET30b

were transferred to room temperature. After 4h of shaking, the cells were washed once with

139

the ice-cold HPT reaction buffer (50 mM HEPES (pH 7.6) containing 4 mM MgC12) and
sonicated on ice by 3 or 4 short, lO-second pulses at 4 - 5 output control using a microtip
probe (W-220F sonicator, Ultrasonics, INC., Plainview, NY). After centrifugation, the
protein pellets (all expressed proteins were previously found to be associated with insoluble
inclusion bodies) were washed twice and resuspended in the HPT reaction buffer. These
crude E. coli protein extracts were stored at - 20 °C for up to 3 weeks and used for the

following prenyltransferase assays.

Preparation of U-[”C] HPP. U-[MC] HPP was prepared from U-[MC] tyrosine (sp.
activity 464 mCi mmol", Amersham, Arlington Heights, IL) according to Schulz et al.
(1993) with minor modifications. Briefly, 2 mL of 0.5 M potassium phosphate buffer (pH
6.5), bovine liver catalase (Boehringer Mannheim, Germany), and L-amino acid oxidase
type IV (Sigma, St. Louis, M0) were added to 1 mL of U-["’C] tyrosine (10 pCi). The
final concentration of both enzymes was 0.4 mg mL". After 2 hours of incubation at room
temperature, U-[‘4C] HPP was purified on an ion exchange column (Dowex, Sigma)
equilibrated with 0.1 N HCI as follows: the first 2 - 3 drops were discarded and the next 3
mL were collected. HPPD assays using eluted fiactions and HPPD demonstrated that the
first 3 mL of eluate contained most U-[”C] HPP (data not shown). The eluted solution pH

was adjusted to ~ 7 with KOH and used immediately for prenyltransferase assays.

Prenyltransferase assays. Each 0.2 mL reaction was performed in the HPT reaction
buffer (50 mM HEPES (pH 7.6) and 4 mM MgC12) containing 100 pM KF, 0.2 % (w/v)
CHAPS, 0.1 mg of total protein extracted fi'om E. coli expressing HPPD, and freshly
prepared 0.2 pM U-[MC] HPP and 50 mM potassium ascorbate. To test HPT substrate
specificity, individual reactions contained either 100 pM PDP, GGDP, or SDP and the
insoluble protein fraction from E. coli expressing pSynHPT (0.03 mg protein), pAtHPT (1

mg protein), or the corresponding empty vectors (0.03 or 1 mg protein from cells

140

transformed with pET3d and pET30b, respectively). Reactions were incubated for an hour
at room temperature, extracted with two volumes of methanolzchloroform (1:1), and any
newly formed prenquuinols were subsequently oxidized at room temperature with AgO for
2 h with shaking (Pennock, 1985). The organic phase was transferred to a fresh tube,
evaporated to dryness, dissolved in ethyl acetate, and subjected to TLC on silica gel
(J.T.Baker, Phillisburg, NJ). The TLC plate was developed with 20% (v/v) ethyl ether in
petroleum ether (Pennock, 1985), dried, and subjected to autoradiography. The film was
exposed for 14 days.

For HPLC analysis, the polyprenyltransferase assays contained all-of-the-above
components except that larger reaction volumes were needed to ensure formation of
detectable prenquuinone products. The following final reaction volumes were used: 0.5 mL
for SynHPT with PDP and 5 mL for SynHPT with GGDP, AtHPT with PDP, and AtHPT
with GGDP. After 2 hours of incubation at room temperature, the reactions were spiked
with non-labeled 2 ’-trans-MPBQ used as a canier and internal standard and extracted
prenyllipids were separated by TLC as described above. For each sample, silica
corresponding to the area of RF 0.36 - 0.67 was scraped from the TLC plates, eluted with 5
mL of ethyl ether, dried under nitrogen, and dissolved in 100 pL of hexane. Samples (50
pL) were then injected on a normal phase HPLC column (LiChrosorb 5 Si60A, 4.6 X 250
mm; Shimadzu) to separate various methyl—phytyl benzoquinone isomers in 0.1% (v/v)
dioxane in hexane (Henry et al., 1987). For geranylgeranylated quinone products, a mobile
phase consisting of 0.15% dioxane (v/v) in iso-octane was used (Hutson and Threlfall,
1980). Isocratic separation was performed at 30 °C with a constant flow of 2 mL min'l and
the prenquuinones were detected at 252 nm for both methods. Eluents were collected at
30- to 60-second intervals and the associated radioactivity was determined in 5 mL of

scintillation liquid by scintillation counting.

I41

Prenyltransferase Assays Using Arabidopsis Chloroplasts And Seed Protein
Extracts

Preparation of crude chloroplasts and seed protein extracts. Wild type and
3SSzzHPTI-ll and -54 Arabidopsis plants were used for all the prenyltransferase assays.
To determine HPT specific activity during plant development, chloroplasts were prepared
from 25 - 30 g of leaf tissue of 2, 4, and 6 week old plants. For the high-light experiment,
6-week-old wild type and 358::HPT1 plants were either kept at normal light conditions or
transferred to high light for 6 days. Three days later, some of the non-stressed plants were
transferred to high light for 3 days, so that analysis of control plants and the plants stressed
for 3 and 6 days was performed the same day to ensure equal treatment of all samples in the
experiment. All procedures were performed at 4 °C and all buffers were pre-chilled and
kept on ice. Tissue was disrupted in 200 mL of buffer containing 0.5 M HEPES pH 8.4,
0.6 M sorbitol, 4 mM EDTA, 4 mM EGTA, 10 mM Na,CO,, 0.2% BSA (w/v), 1 pM
benzamidine (BA), and 5 pM 4-aminocaproic acid (4-ACA) using a blender. Crude
chloroplasts were filtered through two layers of miracloth and centrifuged at 4,000 g for five
minutes. Chloroplasts were washed in the HPT reaction buffer containing 1 pM BA and 5
pM 4-ACA. After centrifugation, an additional wash with the HPT reaction buffer without
the proteinase inhibitors BA and 4-ACA was performed and the chloroplast membranes
were resuspended in the HPT reaction buffer to a final protein concentration of 5.5 - 6.5 mg
mL". Seed protein extracts were prepared as follows: Dry mature seeds (~ 100 mg) were
ground in 1 mL of the HPT reaction buffer using a mortar and pestle and centrifuged. The
resulting pellet of water insoluble proteins was washed once and resuspended in the HPT
reaction buffer to a protein concentration of 20 - 30 mg mL". Preliminary HPT assays

showed that over 95% of HPT activity was associated with the pellet (data not shown).

Prenyltransferase assays using Arabidopsis chloroplasts and seed protein extracts.

To obtain more concentrated U-[MC] HPP, U-[MC] tyrosine (1 mL, 50 pCi, specific activity

142

464 mCi mmol") was dried under a stream of nitrogen and dissolved in 0.8 mL of
phosphate buffer (0.5 M, pH 6.5). U-[MC] HPP was prepared and purified as described
above. For HPT assays in non-stressed and stressed chloroplasts, each prenyltransferase
reaction (0.1 mL) was performed in HPT reaction buffer containing approximately 5 pM U-
[MC] HPP (l pCi), 0 or 100 pM PDP, 1 mM KF, 0.2% (v/v) TWEEN 80, 50 mM
potassium ascorbate, 20 pg HPPD protein, and chloroplasts corresponding to 0.45 — 0.65
mg of protein (0.10 — 0.15 mg of chlorophyll for non-stressed plants). HPT assays using
seed protein extracts were performed as these using chloroplasts except that each reaction
(performed in a reaction volume 0.2 mL) contained 2 - 3 mg seed protein. Reactions were
incubated at room temperature for 2 hours with shaking and quinones were extracted with 2
volumes of acetonezlight petroleum ether (1 :1 v/v). After centrifugation, the quinones (in
ether phase) were purified on hand-made 4-mL Silica columns (pore size 60 pm, Sigma) in
lO-mL syringes and eluted directly into scintillation vials with 4 volumes of 20% (v/v)
diethyl ether in petroleum ether. After drying, 5 mL of scintillation liquid was added to each
vial and radioactivity was determined by scintillation counting. Background activities
obtained from control reactions containing l“C-labeled HGA with no exogenously added
PDP were subtracted from the activities when both substrates were added, which was always
at least 3.5- and 3-fold higher than the background activities in chloroplasts and seeds,
respectively. HPT specific activity was calculated as PDP-stimulated HPT activity per mg
of protein. Chloroplasts and seed protein extracts were extracted with 90 % methanol (v/v)
and centrifuged. Total chlorophyll concentrations were determined spectrophotometrically
on the 90 % methanol (v/v) chloroplast extracts (Lichtenthaller, 1987). Proteins present in
the remaining pellet were solubilized by NaOH in the presence of SDS and determined by
the method of Lowry (Stoscheck, 1990) in a 96-well format. Briefly, three 4-mm beads
(Sigma) and 0.2 mL 1N NaOH were added. The protein pellets were disrupted by using a
paint shaker for 5 minutes, incubated at 37 0C for 10 minutes, and this procedure was

repeated three times. The resulting NaOH protein extract was briefly centrifuged and 80 pL

143

of the supernatant was added to a flesh tube containing 80 pL 10% SDS. After adding 240
pL copper reagent and 400, pL water, the samples were incubated 10 minutes at room
temperature and 200 pL 0.2 N Folin reagent (Sigma, sold as 2N solution) was added. The
absorbance was measured at 750 nm against a blank consisting of all reagents lacking
proteins. The standard curve was constructed using BSA as a standard. The copper reagent
must be prepared fresh and contains 2 g Na,CO3, 0.04 g CuSO4.5H,O, and 0.02 g sodium

potasium tartrate per 30 mL water.

144

LITERATURE CITED

Addlesee HA, Gibson LCD, Jensen PE, Hunter CN (1996) Cloning, sequencing and
functional assignment of the chlorophyll biosynthesis gene, chlP, of Synechacystis sp. PCC
6803. FEBS Lett 389: 126-130

Addlesee HA, Hunter CN (1999) Physical mapping and fimctional assignment of the
geranylgeranyl-bacteriochlorophyll reductase gene, bchP, of Rhodobacter sphaeroides. J
Bacteriol 181: 7248-7255

Altincicek B, Duin EC, Reichenberg A, Hedderich R, Kollas A-K, Hintz M,
Wagner S, Weiser J, Beck E, Jomaa H (2002) Lth protein catalyzes the terminal step of
the 2-C—methyl-D-erythritol-4-phosphate pathway of isoprenoid biosynthesis. FEBS Lett
532: 437-440

Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ (1990) Basic local alignment
search tool. J Mol Biol 215: 403-410

Arango Y, Heise K-P (1998) Tocopherol synthesis from homogentisate in Capsicum
anuum L. (yellow pepper) chromoplast membranes: evidence for tocopherol cyclase.
Biochemical Journal 336: 531-533

Aratri E, Spycher SE, Breyer I, Azzi A (1999) Modulation of a-tropomyosin
expression by a—tocopherol in rat vascular smooth muscle cells. F EBS Lett 447: 91-94

Arigoni D, Eisenreich W, Latzel C, Sagner S, Radykewicz T, Zenk MH, Bacher A
(1999) Dimethylallyl pyrophosphate is not the committed precursor of isopentenyl

pyrophosphate during terpenoid biosynthesis from l-deoxyxylulose in higher plants. Proc
Natl Acad Sci USA 96: 1309-1314

Arigoni D, Sagner S, Latzel C, Eisenreich W, Bacher A, Zenk MH (1997) Terpenoid
biosynthesis from l-deoxy-D-xylulose in higher plants by intrarnolecular skeletal
rearrangement. Proc Natl Acad Sci USA 94: 10600-10605

Arita M, Nomura K, Arai H, Inoue K (1997) a-Tocopherol transfer protein stimulates
the secretion of a-tocopherol from a cultured liver cell line through a brefeldin A-insensitive
pathway. Proc Natl Acad Sci USA 94: 12437-12441

Azzi A, Breyer I, Feher M, Pastori M, Ricciarelli R, Spycher S, Staffieri M, Stocker
A, Zimmer S, Zingg J-M (2000) Specific cellular responses to (rt-tocopherol. J Nutr 130:
1649-1652

Azzi A, Breyer I, Feher M, Ricciarelli R, Stocker A, Zimmer S, Zingg J-M (2001)
Nonantioxidant functions of a-tocopherol in smooth muscle cells. J Nutr 131: 3788-381 S
Balsinde J, Balboa MA, Insel PA, Dennis ED (1999) Regulation and inhibition of
phospholipase A,. Annu Rev Pharmacol Toxicol 39: 175-189

Bang O-S, Park J-H, Kang S-S (2001) Activation of PKC but not of ERK is required for
vitamin E-succinate—induced apoptosis of HL-60 cells. Biochem Biophys Res Comm 288:
289-297

Barnett KT, Fokum FD, Malafa MP (2002) Vitamin E succinate inhibits colon cancer
liver metastases. J Surg Res 106: 292-298

145

Bartlett PA, Satake K (1988) Does dehydroquinate synthase synthesize dehydroquinate?
J Am Chem Soc 110: 1628-1630

Bartoli CG, Simontacchi M, Tambussi E, Beltrano J, Montaldi E, Puntarulo S
(1999) Drought and watering-dependent oxidative stress: effect on antioxidant content in
T riticum aestivum L. leaves. J Exp Bot 50: 375-383

Batz O, Logemann E, Reinold S, Halbrock K (1998) Extensive reprogramming of
primary and secondary metabolism by fungal elicitor of infection in parsley cells. Biol
Chem 379: 1127-1135

Beharka AA, Wu D, Seraf’mi M, Meydani SN (2002) Mechanism of vitamin E
inhibition of cycloxygenase activity in macrophages from old mice: Role of peroxynitrite.
Free Rad Biol Med 32: 503-511

Bidoli E, LaVecchia C, Talamini R, Negri E, Parpinel M, Conti E, Montella M,
Carbone A, Franceschi S (2001) Micronutrients and ovarian cancer: A case-control study
in Italy. Annals Oncol 12: 1589-1593

Bisby RH, Ahmed S (1989) Transverse distribution of (rt-tocopherol in bilayer membranes
studied by fluorescence quenching. Free Rad Biol Med 6: 231-239

Bischoff M, Rosier J, Raesecke HR, Gorlach J, Amrhein N, Schmid J (1996)
Cloning of a cDNA encoding a 3-dehydroquinate synthase from a higher plant, and
analysis of the organ-specific and elicitor-induced expression of the corresponding gene.
Plant Mol Biol 31: 69-76

8199 E (2002) Impact of phyto-oxylipins in plant defense. Trends Plant Sci 7: 315-321
Boaz M, Smetana S, Weinstein T, Matas Z, Gafter U, Iaina A, Knecht A,
Weissgarten Y, Brunner D, Fainaru M, Green MS (2000) Secondary prevention with
antioxidants of cardiovascular disease in endstage renal disease (SPACE): randomized
placebo-controlled trial. Lancet 356: 1213-1218

Bligh EG, Dyer WJ (1959) A rapid method of total lipid extraction and purification. Can J
Biochem Physiol 37: 91 1-917
Bollschweiler E, Wolfgarten E, Nowroth T, Rosendahl U, Monig SP, Holscher AH
(2002) Vitamin E intake and risk of subtypes of esophaegal cancer in Germany. J Cancer
Res Clin Oncol 128: 575-580

Bonner CA, Jensen RA (1994) Cloning of cDNA encoding the bifunctional
dehydroquinase Shikimate dehydrogenase of aromatic amino acid biosynthesis in Nicotiana
tabacum. Biochem J 302: ll-l4

Bouvier F, d'HarIingue A, Suire C, Backhaus RA, Camara B (1998) Dedicated roles
of plastid transketolases during the early onset of isoprenoid biogenesis in pepper fruits.
Plant Physiol 117: 1423-1431

Bramley PM, Elrnadfa I, Kafatos A, Kelly FJ, Manios Y, Roxborough HE, Schuch
W, Sheehy PJA, Wagner K-H (2000) Vitamin E. J Sci Food Agric 80: 913-938

Braun M, Henstrand JM, Gorlach J, Amrhein N, Schmid J (1996) Enzymatic

properties of chorismate synthase isoenzyrnes in tomato (Lycopersicon esculentum Mill.).
Planta 200: 64-70

146

Brigelius-Flohe R, Kelly FJ, Salonen JT, Neuzil J, Zingg J-M, Azzi A (2002) The
European perspective on vitamin E: current knowledge and future research. Am J Clin Nutr
76: 703-716

Brigelius-Flohe R, Traber MC (1999) Vitamin E: function and metabolism. FASEB J
13: 1145-1155

Buchanan-Wollaston V (1997) The molecular biology of leaf senescence. J Exp Bot 48:
181-199

Buring JE, Hennekens CH (1997) Antioxidant vitamins and cardiovascular disease. Nutr
Rev 55: 853-860

Burton GW, Traber MG (1990) Vitamin E: Antioxidant activity, biokinetics, and
bioavailability. Annu Rev Nutr 10: 357-382

Carlberg C (1999) Lipid soluble vitamins in gene regulation. BioFactors 10: 91-97

Celik S, Ozkaya A (2002) Effects of intraperitoneally administered lipoic acid, vitamin E,
and linalool on the level of total lipid and fatty acids in guinea pig brain with oxidative stress
induced by H202. J Biochem Mol Biol 35: 547-552

Chan AC, Wagner M, Kennedy C, Chen E, Lanuville O, Mezl VA, Tran K, Choy
PC (1998) Vitamin E up-regulates arachidonic acid release and phospholipase A, in
megakaryocytes. Mol Cell Biochem 189: 153-159

Chen A, Kroon PA, Poulter CD (1994) Isoprenyl diphosphate synthases: Protein
sequence comparisons, a phylogenetic tree, and predictions of secondary structure. Prot Sci
3: 600-607

Ching LS, Mohamed S (2001) Alpha-tocopherol content in 62 edible tropical plants. J
Agric Food Chem 49: 3101-3105

Clough SJ, Bent AF (1998) Floral dip: a simplified method for Agrobacterium-mediated
transformation of Arabidopsis thaliana. Plant J 16: 735-743

Collakova E, DellaPenna D (2001) Isolation and functional analysis of homogentisate
phytyltransferase from Synechacystis sp. PCC 6803 and Arabidopsis. Plant Physiol 127:
1113-1124

Collakova E, DellaPenna D (2003) Homogentisate phytyltransferase activity is limiting
for tocopherol biosynthesis in Arabidopsis thaliana. Plant Physiol 131: 632-642

D' Harlingue A, Camara B (1985) Plastid enzymes of terpenoid biosynthesis:
purification and characterization of a gamma-tocopherol methyltransferase. J Biol Chem
260: 15200—15203

Dahnhardt D, Falk J, Appel J, Kooij TAW, Schulz-Friedrich R, Krupinska K

(2002) The hydroxyphenylpyruvate dioxygenase from Synechocystis sp. PCC 6803 is not
required for plastoquinone biosynthesis. F EBS Lett 523: 177-181

147

Dasilva EJ, Jensen A (1971) Content of a—tOCOpherol in some blue-green algae. Biochim
Biophys Acta 239: 345-347

Davies TW, Treasure F P, Welch AA, Day NE (2002) Diet and basal cell skin cancer:
results from the EPIC-Norfolk cohort. Brit J Dermatol 146: 1017-1022

Delgado-Zamarreno MM, Bustamante—Rangel M, Sanchez-Perez A, Hernandez-
Mendez J (2001) Analysis of vitamin E isomers in seeds and nuts with and without
coupled hydrolysis by liquid chromatography and coulometric detection. J Chromatogr A
935: 77-86

DellaPenna D, Collakova E, Coughlan SJ, Helentjaris T (2000)
Phytyl/prenyltransferase nucleic acids, polypeptides and uses thereof. Patent Application
WO 0068393

Diaz J, Bernal A, Pomar F, Merino F (2001) Induction of Shikimate dehydrogenase and
peroxidase in pepper (Capsicum annuum L.) seedlings in response to copper stress and its
relation to lignification. Plant Sci 161: 179-188

Diaz J, Merino F (1998) Wound-induced Shikimate dehydrogenase and peroxidase
related to lignification in pepper (Capsicum annuum L.). J Plant Physiol 152: 51-57

Diplock AT, Lucy JA (1973) The biochemical modes of action of vitamin E and
selenium: A hypothesis. F EBS Lett 29: 205-210

Disch A, Schwender J, Muller C, Lichtenthaler HK, Rohmer M (1998) Distribution
of the mevalonate and glyceraldehyde phosphate/pyruvate pathways for isoprenoid
biosynthesis in unicellular algae and the cyanobacterium Synechacystis PCC 6714.
Biochem J 333: 381-388

Dyer WE, Henstrand JM, Handa AK, Herrmann KM (1989) Wounding induces the
first enzyme of the Shikimate pathway in Solanaceae. Proc Natl Acad Sci USA 86: 7370-
7373

Eccleston V, Ohlrogge J (1998) Expression of lauroyl-acyl carrier protein thioesterase in
Brassica napus seeds induces pathways for both fatty acid oxidation and biosynthesis and
implies a set point for triacylglycerol accumulation. Plant Cell 10: 613-621

Eisenreich W, Menhard B, Hylands P, Zenk MH, Bacher A (1996) Studies on the
biosynthesis of taxol: the taxane carbon skeleton is not of mevalonoid origin. Proc Natl
Acad Sci USA 93: 6431-6436

Eisenreich W, Schwarz M, Cartayrade A, Arigoni D, Zenk MH, Bacher A (1998)
The deoxyxylulose phosphate pathway of terpenoid biosynthesis in plants and
microorganisms. Chem Biol 5: R221-R233

Eitenmiller RR (1997) Vitamin E content of fats and oils - nutritional implications. Food
Technol 51: 78-81

Emanuelsson O, Nielsen H, von Heijne G (1999) ChloroP, a neural network-based

method for predicting chloroplast transit peptides and their cleavage sites. Prot Sci 8: 978-
984

148

Erin AN, Spirin MM, Tabidze LV, Kagan VE (1984) Formation of a-tocopherol
complexes with fatty acids. A hypothetical mechanism of stabilization of biomembranes by
vitamin E. Biochim Biophys Acta 774: 96-102

Ershow Y, Gantt R, Cunningham FX, Gantt E (2000) Isopentenyl diphosphate
isomerase deficiency in Synechocystis sp. strain PCC 6803. F EBS Lett 473: 337-340

Estevez JM, Cantero A, Reindl A, Reichler S, Leon P (2001) l-Deoxy-D-xylulose-S-
phosphate synthase, a limiting enzyme for plastidic isoprenoid biosynthesis in plants. J Biol
Chem 276: 22901-22909

Estevez JM, Cantero A, Romero C, Kawaide H, Jimenez LF, Kuzuyama T, Seto H,
Kamiya Y, Leon P (2000) Analysis of the expression of CLAI, a gene that encodes the l-
deoxyxylulose S-phosphate synthase of the 2-C-methyl-D-erythritol-4-phosphate pathway
in Arabidopsis. Plant Physiol 124: 95-103

Factor VM, Laskowska D, Jensen MR, Woitach JT, Popescu NC, Thorgeirsson SS
(2000) Vitamin E reduces chromosomal damage and inhibits hepatic tumor formation in a
transgenic mouse model. Proc Natl Acad Sci USA 97: 2196-2201

Fahrenholtz SR, Doleiden FH, Trozzolo AM, Lamola AA (1974) On the quenching
of singlet oxygen by a—tocopherol. Photochem Photobiol 20: 505-509

Fechner H, Schlame M, Guthmann F, Stevens PA, Rustow B (1998) 01- And 6-
tocopherol induce expression of hepatic (rt-tocopherol-transfer-protein mRNA. Biochem J
331: 577-581

Fiedler E, Sol] J, Schultz G (1982) The formation of homogentisate in the biosynthesis
of tocopherol and plastoquinone in spinach chloroplasts. Planta 155: 51 1-515

F inckh BF, Kunert KJ (1985) Vitamins C and E: An antioxidative system against
herbicide-induced lipid peroxidation in higher plants. J Agric Food Chem 33: 574-577

F leshner N, Fair WR, Huryk R, Heston WDW (1999) Vitamin E inhibits the high-fat
diet promoted groowth of established human prostate LNCaP tumors in nude mice. J Urol
161: 1651-1654

Food and Nutrition Board, Institute of Medicine (2000). Vitamin E. In Dietary
reference intakes for vitamin C, vitamin E, selenium, and carotenoids. National Academy
Press, Washington DC, pp186-283

Foote CS, Ching T-Y, Geller GG (1974) Chemistry of singlet oxygen. XVIII. Rates of
reaction and quenching of (rt-tocopherol and singlet oxygen. Photoch Photobiol 20: 511-
513

Foyer CH, Lelandais M, Kunert KJ (1994) Photooxidative stress in plants. Physiologia
Plantarum 92: 696-717

Franzen J, Bausch J, Glatzle D, Wagner E (1991) Distribution of vitamin E in spruce
seedling and mature tree organs, and within the genus. Phytochemistry 30: 147-151

Franzen J, Haas MM (1991) Vitamin E content during development of some seedlings.
Phytochemistry 30: 291 1-2913

149

Fray RG, Wallace A, Fraser PD, Valero D, Hedden P, Bramley PM, Grierson D
(1995) Constitutive expression of a fruit phytoene synthase gene in transgenic tomatoes
causes dwarfism by redirecting metabolites from the gibberellin pathway. Plant J 8: 693-
701

Freedman JE, Keaney JF (2001) Vitamin E inhibition of platelet aggregation in
independent of antioxidant activity. J Nutr 131: 3748-3773

Fryer MJ (1992) The antioxidant effects of thylakoid Vitamin E (ct-tocopherol). Plant Cell
Environ 15: 381-392

Fryer MJ (1993) Evidence for the photoprotective effects of vitamin E. Photochem
Photobiol 58: 304-312

Fryer MJ, Andrews JR, Oxborough K, Blowers DA, Baker NR (1998) Relationship
between CO, assimilation, photosynthetic electron transport, and active 0, metabolism in
leaves of maize in the field during periods of low temperature. Plant Physiol2 116: 571-580

F ukuzawa K, Chida H, Tokumura A, Tsukatani H (1981) Antioxidative effect of oz-
tocopherol incorporation into lecithin liposomes on ascorbic acid- Fe -induced lipid
peroxidation. Arch Biochem Biophys 206: 173- 180

Fukuzawa K, Gebicki JM (1983) Oxidation of a-tocopherol in micelles and liposomes
by the hydroxyl, perhydroxyl, and superoxide free radicals. Arch Biochem Biophys 226:
242-251

F ukuzawa K, Matsuura K, Tokumura A, Suzuki A, Terao J (1997) Kinetics and
dynamics of singlet oxygen scavenging by (rt-tocopherol in phospholipid model
membranes. Free Rad Biol Med 22: 923-930

Fukuzawa K, Tokumura A, Ouchi S, Tsukatani H (1982) Antioxidant activities of
tocopherols on iron(2+)-ascorbate-induced lipid peroxidation in lectithin liposomes. Lipids
17: 511-513

F uruya T, Yoshikawa T, Kimura T, Kaneko H (1987) Production of tocopherols by
cell culture of safflower. Phytochemistry 26: 2741-2747

Garcia 1, Rodgers M, Lenne C, Rolland A, Sailland A, Matringe M (1997)
Subcellular localization and purification of a p—hydroxyphenylpyruvate dioxygenase from
cultured carrot cells and characterization of the corresponding cDNA. Biochem J 325: 761 -
769

Garcia 1, Rodgers M, Pepin R, Hssich T, Matringe M (1999) Characterization and
subcellular compartmentation of recombinant 4-hydroxyphenylpyruvate dioxygenase from
Arabidopsis in transgenic tobacco. Plant Physiol 119: 1507-1516

Gasser CS, Klee HJ (1990) A Brassica napus gene encoding 5-enolpyruvyl-shikimate 3-
phosphate synthase. Nucleic Acids Res 18: 2821-2821

Gasser CS, Winter JA, Hironaka CM, Shah DM (1988) Structure, expression, and

evolution of the 5-enolpyruvyl-shikimate 3-phosphate synthase genes of petunia and
tomato. J Biol Chem 263: 4280-4289

150

Gleave AP (1992) A versatile binary vector system with a T-DNA organisational structure
conductive to efficient integration of cloned DNA into the plant genome. Plant Mol Biol 20:
1203-1207

Goffman FD, Bohme T (2001) Relationship between fatty acid profile and vitamin E
content in maize hybrids (Zea mays L.). J Agric Food Chem 49: 4990-4994

Goffman FD, Mollers C (2000) Changes in tocopherol and plastochromanol-8 contents in
seeds and oil of oilseed rape (Brassica napus L.) during storage as influenced by
temperature and air oxygen. J Agric Food Chem 48: 1605-1609

Goldstein JL, Brown MS (1990) Regulation of the mevalonate pathway. Nature 343:
425-430

Gombos Z, Wada H, Murata N (1992) Unsaturation of fatty acids in membrane lipids
enhances tolerance of the cyanobacterium Synechocystis PCC6803 to low-temperature
photoinhibition. Proc Natl Acad Sci USA 89: 9959-9963

Gorlach J, Beck A, Henstrand JM, Handa AK, Herrmann KM, Schmid J, Amrheim
N (1993a) Differential expression of tomato (Lycopersicon esculentum L.) genes encoding
Shikimate pathway isoenzymes. I. 3-Deoxy-D-arabino-heptulosonate 7-phosphate synthase.
Plant Mol Biol 23: 697-706

Gorlach J, Raesecke H-R, Rentsch D, Regenass M, Roy P, Zala M, Keel C, Boiler T,
Amrheim N, Schmid J (1995) Temporally distinct accmnulafion of transcripts encoding
enzymes of the prechorismate pathway in elicitor-treated, cultured tomato cells. Proc Natl
Acad Sci USA 92: 3166-3170

Gorlach J, Schmid J, Amrheim N (1993b) Differential expression of tomato
(Lycopersicon esculentum L.) genes encoding Shikimate pathway isoenzymes. II.
Chorismate synthase. Plant Mol Biol 23: 707-716

Gossett DR, Millhollon EP, Lucas MC (1994) Antioxidant response to NaCl stress in
salt-tolerant and salt-sensitive cultivars of cotton. Crop Sci 34: 706-714

Grau A, Ortiz A (1998) Dissimilar protection of tocopherol isomers against membrane
hydrolysis by phospholipase A,. Chem Phys Lipids 91: 109-1 18

Griffiths WT, Threlfall DR, Goodwin TW (1967) Nature, intracellular distribution and
formation of terpenoid quinones in maize and barley shoots. Biochem J 103: 589-600

Grusak MA, DellaPenna D (1999) Improving the nutrient composition of plants to
enhance human nutrition and health. Annu Rev Plant Physiol Plant Mol Biol 50: 133-161

Ham AJ, Liebler DC (1997) Antioxidant reactions of vitamin E in the perfused rat liver“.
product distribution and effect of dietary vitamin E supplementation. Arch Biochem
Biophys 339: 157-164

Havaux M, Bonfils J-P, Lutz C, Niyogi KK (2000) Photodamage of the photosynthetic
apparatus and its dependence on the leaf development stage in the npql Arabidopsis mutant
deficient in the xanthophyll cycle enzyme violaxanthin de-epoxidase. Plant Physiol 124:
273-284

151

Havaux M, Kloppstech K (2001) The protective firnctions of carotenoids and flavonoid
pigments against excess visible radiation at chilling temperature investigated in Arabidopsis
npq and tt mutants. Planta 213: 953-966

Havaux M, Niyogi K (1999) The violaxanthin cycle protects plants from photooxidative
damage by more than one mechanism. Proc Natl Acad Sci USA 96: 8762-8767

Hawkes TR, Lewis T, Coggins JR, Mousedale DM, Lowe DJ, Thorneley RNF
(1990) Chorismate synthase, pre-steady-state kinetics of phosphate release from 5-
enolpyruvylshikimate 3-phosphate. Biochem J 265: 899-902

Henkes S, Sonnewald U, Badur R, Flachmann R, Stiff M (2001) A small decrease of
plastid transketolase activity in antisense tobacco transfonnants has dramatic effects on
photosynthesis and phenylpropanoid metabolism. Plant Cell 13: 535-551

Henry A, Powls R, Pennock JF (1986) Scenedesmus obliquus P828: 01 tocopherol-free
mutant which cannot form phytol. Biochem Soc Trans 14: 958-959

Henry A, Powls R, Pennock JF (1987) Intermediates of tocopherol synthesis in the
unicellular alga Scenedesmus obliquus. Biochem J 242: 367-373

Henstrand JM, Schmid J, Amthein N (1995) Only the mature form of the plastidic
chorismate synthase is enzymatically active. Plant Physiol 108: 1127-1132

Herrman KM (1995) The Shikimate pathway as an entry to aromatic secondary
metabolism. Plant Physiol 107: 7-12

Herrman KM, Weaver LM (1999) The shikirnate pathway. Annu Rev Plant Physiol Plant
Mol Biol 50: 473-503

Herz S, Wungsintaweekul J, Schuhr CA, Hecht S, Luttgen H, Sagner S, Fellermeier
M, Eisenreich W, Zenk MH, Bacher A, Rohdich F (2000) Biosynthesis of terpenoids:
ngB protein converts 4—diphosphocytidyl-2C-methyl-D-erythritol 2-phosphate to 2C-
methyl-D-erythritol 2,4-cyclodiphosphate. Proc Natl Acad Sci USA 97 : 2486-2490

Horwitt MK (2001) Critique of the requirement for vitamin E. Am J Clin Nutr 73: 1003-
1005

Howe GA, Schilmiller AL (2002) Oxylipin metabolism in response to stress. Curr Opin
Plant Biol 5: 230-236

Hutson KG, Threlfall DR (1980) Synthesis of plastoquinone-9 and phytylplastoquinone
from homogentisate in lettuce chloroplasts. Biochim Biophys Acta 632: 630-648

Ishizaki-Nishizawa O, Fujii T, Azuma M, Sekiguchi K, Murata N, Ohtani T,
Toguri T (1996) Low-temperature resistance of higher plants is significantly enhanced by
a nonspecific cyanobacterial desaturase. Nature Biotech 14: 1003-1006

Jacobs EJ, Connell CJ, McCullough ML, Chao A, Jonas CR, Rodriguez C, Calle
EE, Thun MJ (2002a) Vitamin C, vitamin E, and multivitamin supplement use and stomach
cancer mortality in the Cancer Prevention Study 11 cohort. Cancer Epidem Biomarker
Prevention 1]: 35-41

152

Jacobs EJ, Henion AK, Briggs PJ, Connell CJ, McCullough ML, Jonas CR,
Rodriguez C, Calle EE, Thun MJ (2002b) Vitamin C and vitamin E supplement use and

bladder cancer mortality in a large cohort of US men and women. Am J Epidemiol 156:
1002-1010

Johnson TW, Shen G, Zybailov B, Kolling D, Reategui R, Beauparlant S, Vassiliev
IR, Bryant DA, Jones AD, Golbeck J, Chitnis PR (2000) Recruitment of a foreign
quinone into the Al site of photosystem I. J Biol Chem 275: 8523-8530

Jones JD, Henstrand JM, Handa AK, Herrmann KM, Weller SC (1995) Impaired
wound induction of 3-deoxy-D-arabino-heptulosonate-7-phosphate (DAHP) synthase and
altered stem development in transgenic potato plants expressing a DAHP synthase antisense
construct. Plant Physiol 108: 1413-1421

Kagan VE, Packer L (1994) Light-induced generation of vitamin E radicals: Assessing
vitamin E regeneration. Methods Enzyrnol 234: 316-320

KamaI-Eldin A, Appelqvist L-A (1996) The chemistry and antioxidant properties of
tocopherols and tocotrienols. Lipids (Review) 31: 671-701

Kaneko T, Sato S, Kotani H, Tanaka A, Asamizu E, Nakamura Y, Miyajima N,
Hirosawa M, Sugiura M, Sasamoto S, Kimura T, Hosouchi T, Matsuno A, Muraki
A, N akazaki N, Naruo K, Okumura S, Shimpo S, Takeuchi C, Wada T, Watanabe
A, Yamada M, Yasuda M, Tabata S (1996) Sequence analysis of the genome of the
unicellular Cyanobacterium Synechocystis sp. strain PCC6803. 11. Sequence detennination
of the entire genome and assignment of potential protein-coding regions. DNA Res 3: 109-
136

Keith B, Dong X, Ausubel FM, Fink GR (1991) Differential induction of 3-deoxy-D-
arabino-heptulosonate 7-phosphate synthase genes in Arabidopsis thaliana by wounding
and pathogenic attack. Proc Natl Acad Sci USA 88: 8821-8825

Keller Y, Bouvier F, D'Harlingue A, Camara B (1998) Metabolic compartrnentation of
plastid prenyllipid biosynthesis. Evidence for the involvement of a multifunctional
geranylgeranyl reductase. Eur J Biochem 251: 413-417

Klebler-Janke T, Krupinska K (1997) Isolation of cDNA clones for genes showing
enhanced expression in barley leaves during dark-induced senescence as well as during
senescence under field conditions. Planta 203: 332-340

Klein EA, Thompson IM, Lippman SM, Goodman PJ, Albanes D, Taylor PR,
Coltman C (2001) SELECT: The next prostate cancer prevention trial. J Urol 166: 1311-
1315

Kranner I, Beckett RP, Wornik S, Zorn M, Pfeifhofer HW (2002) Revival of a
resurrection plant correlates with its antioxidant status. Plant J 31: 13-24

Kruk J, Schmid GH, Strzalka K (2000) Interaction of ct-tocopherol quinone, a-
tocopherol and other prenyllipids with photosystem II. Plant Physiol Biochem 38: 271-277

Kruk J, Strzalka K (1995) Occurrence and function of ot-tocopherol quinone in plants. J
Plant Physiol 145: 405-409

153

Kuntz M, Romer S, Suire C, Hugueney P, Schantz JHWR, Camara B (1992)
Identification of a cDNA for the plastid-located geranylgeranyl pyrophosphate synthase

from Capsicum annuum: correlative increase in enzyme activity and transcript level during
fruit ripening. Plant J 2: 25-34

Kushi LH, Folsom AR, Prineas RJ, Mink PJ, Wu Y, Bostick RM (1996) Dietary
antioxidant vitamins and death from coronary heart disease in postmenopausal women. N
Engl J Med 334: 1156-1162

Lange BM, Wildung MR, McCackiIl D, Croteau R (1998) A family of transketolases
that directs isoprenoid biosynthesis via a mevalonate-independent pathway. Proc Natl Acad
Sci USA 95: 2100-2104

Leipner J, Fracheboud Y, Stamp P (1999) Effect of growing season on the
photosynthetic apparatus and leaf antioxidative defenses in two maize genotypes of different
chilling tolerance. Environ Exp Bot 42: 129-139

Lichtenthaler HK (1969) Light-stimulated synthesis of plastid quinones and pigments in
etiolated barley seedlings. Biochim Biophys Acta 184: 164-172

Lichtenthaler HR (1987) Chlorophylls and carotenoids: Pigments of photosynthetic
biomembranes. Methods Enzyrnol 148: 350-382

Lichtenthaler HK (1993) The plant prenyllipids, including carotenoids, chlorophylls, and
prenquuinones. Lipid metabolism in plants: The plant prenyllipids. CRC press, pp 427-450

Lichtenthaler HK (1998) The plants' 1-deoxy-D-xylulose-5-phosphate pathway for
biosynthesis of isoprenoids. Petr/Lipid 100: 128-138

Lichtenthaler HK, Prenzel U, Douce R, Joyard J (1981) Localization of
prenquuinones in the envelope of spinach chloroplasts. Biochim Biophys Acta 641: 99-
105

Lichtenthaler HK, Rohmer M, Schwender J (1997a) Two independent biochemical
pathways for isopentenyl diphosphate and isoprenoid biosynthesis in higher plants.
Physiologia Plantarum 101: 643-652

Lichtenthaler HK, Schwender J, Disch A, Rohmer M '(1997b) Biosynthesis of
isoprenoids in higher plant chloroplasts proceeds via a mevalonate-independent pathway.
FEBS Lett 400: 271-274

Liebler DC (1993) The Role of Metabolism in the Antioxidant Function of Vitamin E.
Critical Rev Tox 23: 147-169

Liebler DC (1998) Antioxidant chemistry of a-tocopherol in biological systems. Roles of
redox cycles and metabolism. Subcell Biochem 30: 301-317

Liebler DC, Burr JA, Phillips L, Ham AJ (1996) Gass chromatography-mass
spectrometry analysis of vitamin E and its oxidation products. Analytic Biochem 236: 17-
34

Logan BA, Monson RK, Potosnak MJ (2000) Biochemistry and physiology of foliar
isoprene production. Trends Plant Sci 5: 477-481

154

Lois LM, Rodrigues-Concepcion M, Gallego F, Campos N, Boronat A (2000)
Carotenoid biosynthesis during tomato fiuit development: regulatory role of l-deoxy-D-
xylulose 5-phosphate synthase. Plant J 22: 503-513
Lopez JC, Ryan S, Blankenship RE (1996) Sequence of the bchG gene from
Chloroflexus aurantiacus: Relationship between chlorophyll synthase and other
polyprenyltransferases. J Bacteriol 178: 3369-3373

Lopukhina A, Dettenberg M, Weiler EW, Hollander-Czytko H (2001) Cloning and
characterization of a coronatine-regulated tyrosine aminotransferase from Arabidopsis.
Plant Physiol 126: 1678-1687

Los DA, Murata N (1998) Structure and expression of fatty acid desaturases. Biochim
Biophys Acta 1394: 3-15

Luttgen H, Rohdich F, Herz S, Wungsintaweekul J, Hecht S, Schuhr CA,
Fellermeier M, Sagner S, Zenk MH, Bacher A, Eisenreich W (2000) Biosynthesis of
terpenoids: YchB protein of Escherichia coli phosphorylates the 2-hydroxy group of 4-
diphosphocytidyl-ZC-methyl-D-erythritol. Proc Natl Acad Sci USA 97: 1062-1067

Mahmoud SS, Croteau RB (2001) Metabolic engineering of essential oil yield and
composition in mint by altering expression of deoxyxylulose phosphate reductoisomerase
and menthofirran synthase. Proc Natl Acad Sci USA 98: 8915-8920

Malafa MP, Fokum FD, Mowlavi A, Abusief M, King M (2002) Vitamin E inhibits
melanoma in mice. Surgery 131: 85-91

Malila N, Taylor PR, Virtanen MJ, Korhonen P, Huttunen JK, Albanes D,
Virtamo J (2002) Effects of alpha-tocopherol and beta-carotene supplementation on
gastric cancer incidence in male smokers (ATBC Study, Finland). Cancer Causes Contr 13:
617-623

Mandel MA, Feldmann KA, Herrera-Estrella L, Rocha-Sosa M, Leon P (1996)
CLAI , a novel gene requered for chloroplast development, is highly conserved in evolution.
Plant J 9: 649-658

Marechal E, Block MA, Dome AJ, Douce R, Joyard J (1997) Lipid synthesis and
metabolism in the plastid envelope. Physiologia Plantarum 100: 65-77

Marshall PS, Morris SR, Threlfall DR (1985) Biosynthesis of tocopherols: A re-
examination of the biosynthesis and metabolism of 2-demethyl-6-phytyl-l,4-benzoquinol.
Phytochemistry 24: 1705- l 71 l

Matile P, Hortensteiner S, Thomas H (1999) Chlorophyll degradation. Annu Rev Plant
Physiol Plant Mol Biol 50: 67-95

Mayer MP, Beyer P, Kleirrig H (1990) Quinone compounds are able to replace molecular
oxygen as terminal electron acceptor in phytoene desaturation in chromoplasts of Narcissus
pseudonarcissus L. Eur J Biochem 191: 359-363

Melis A (1999) Photosystem II-damage and repair cycle in chloroplasts: what modulates
the rate of photodamage in vivo. Trends Plant Sci 4: 130-135

155

Merritt MV, Rosenstein SP, Loh C, Hsui—sui Chou R, Allen MM (1991) A
comparison of the major lipid classes and fatty acid composition of marine unicellular
cyanobacteria with freshwater species. Arch Microbiol 155: 107-113

Merzlyak MN, Chivkunova OB (2000) Light-stress-induced pigment changes and
evidence for anthocyanin photoprotection in apples. J Photochem Photobiol B: Biol 55:
155-163

Meydani M (2001) Vitamin E and atherosclerosis: Beyond prevention of LDL oxidation. J
Nutr 131: 3668-3688

Michalski WP, Kaniuga Z (1981) Photosynthetic apparatus of chilling-sensitive plants.
IX. The involvement of a-tocopherol in the electron transport chain and the anti-oxidizing
system in chloroplasts of tomato leaves. Biochim Biophys Acta 635: 25-37

Mittler R (2002) Oxidative stress, antioxidants and stress tolerance. Trends Plant Sci 7:
405-410

Mobley EM, Kunkel BN, Keith B (1999) Identification, characterization and comparative
analysis of a novel chorismate mutase gene in Arabidopsis thaliana. Gene 240: 1 15-123

Molina-Torres J, Martinez ML (1991) Tocopherols and leaf age in Xanthium
strumarium L. New Phytol 118: 95-99

Mousdale DM, Campbell MS, Coggins JR (1987) Purification and characterization of
bifunctional dehydroquinase-shikimatezNADP oxidoreductase from pea seedlings.
Phytochemistry 26: 2665-2670

Munne-Bosch S, Alegre L (2000) Changes in carotenoids, tocopherols and diterpenes
during drought and recovery, and the biological significance of chlorophyll loss in
Rosmarinus oflicinalis plants. Planta 210: 925-931

Munne-Bosch S, Alegre L (2002a) The function of tocopherols and tocotrienols in plants.
Crit Rev Plant Sci 21: 31-57

Munne-Bosch S, Alegre L (2002b) Interplay between ascorbic acid and lipophilic
antioxidant defences in chloroplasts of water-stressed Arabidopsis plants. FEBS Let 524:
145-148

Munne—Bosch S, Alegre L (2002c) Plant aging increases oxidative stress in chloroplasts.
Planta 214: 608-615

Munne-Bosch S, Jubany-Mari T, Alegre L (2001) Drought-induced senescence is
characterized by a loss of antioxidant defences in chloroplasts. Plant Cell Environ 24: 1319-
1327

Nagata N, Suzuki M, Yoshida S, Muranaka T (2002) Mevalonic acid partially restores
chloroplast and etioplast development in Arabidopsis lacking the non-mevalonate pathway.
Planta 216: 345—350

Neely WC, Martin JM, Barker SA (1988) Products and relative reaction rates of the
oxidation of tocopherols with singlet molecular oxygen. Photochem Photobiol 48: 423-428

156

Neuzil J, Weber T, Schroder A, Lu M, Ostermann G, Gellcrt N, Mayne GC,
Olejnicka B, Negre-Salvayre A, Sticha M, Coffey RJ, Weber C (2002) Induction of
cancer cell apoptosis by a-tocopheryl succinate: molecular pathways and structural
requirements. F ASEB J 15: 403-415

Nicholson DW, Thornberry NA (1997) Caspases: Killer proteases. Trends Biochem Sci
22: 299-306

Nishikimi M, Yamada H, Yagi K (1980) Oxidation by superoxide of tocopherols
dispersed in aqueous media with deoxycholate. Biochim Biophys Acta 627: 101-108

N iyogi KK (1999) Photoprotection revisited: Genetic and molecular approaches. Annu Rev
Plant Physiol Plant Mol Biol 50: 333-359

Norris SR, Barrette TR, DellaPenna D (1995) Genetic disection of carotenoid synthesis
in Arabidopsis defines plastoquinoe as an essential component of phytoene desaturation.
Plant Cell 7: 2139-2149

Norris SR, Shen X, DellaPenna D (1998) Complementation of the Arabidopsis pdsI
mutation with the gene encoding p-hydroxyphenylpyruvate dioxygenase. Plant Physiol
117:1317-1323

O'Malley PJ (2002) Hybrid density firnctional studies on phenoxyl free radicals modeling
a-tocopheroxyl. J Phys Chem B 106: 12331-12335

Okada K, Saito T, Nakagawa T, Kawamukai M, Kamiya Y (2000) Five
geranygeranyl diphosphate synthases expressed in different organs are localized into three
subcellular compartments in Arabidopsis. Plant Physiol 122: 1045-1056

Oster U, Bauer CE, Rudiger W (1997) Characterization of chlorophyll a and
bacteriochlorophyll a synthases by heterologous expression in Escherichia coli. J Biol
Chem 272: 9671-9676

Pennock JF (1985) Biosynthesis of plastoquinone. Methods Enzymol 110: 313-319

Peterson DM (1995) Oat tocols: concentration and stability in oat products and distribution
within kernels. Cereal Chem 72: 21-24

Piironen V, Syvaoja E-L, Varo P, Salminen K, Koivistoinen P (1986) Tocopherols
and tocotrienols in finnish foods: Vegetables, fruits, and berries. J Agric Food Chem 34:
742-746

Porfirova S, Bergmuller E, Tropf S, Lemke R, Dormann P (2002) Isolation of an
Arabidopsis mutant lacking vitamin E and identification of a cyclase essential for all
tocopherol biosynthesis. Proc Natl Acad Sci USA 99: 12495-12500

Proteau PJ (1998) Biosynthesis of phytol in the cyanobacterium Synechocystis sp. UTEX
2470: Utilization of the non-mevalonate pathway. J Nat Prod 61: 841-843

Pryor WA (2000) Vitamin E and heart disease: Basic science to clinical intervention trials.
Free Rad Biol Med 28: 141-164

Ricciarelli R, Tasinato A, Clement S, Ozer NK, Boscoboinik D, Azzi A (1998)

Alpha-tocopherol specifically inactivates protein kinase C alpha by changing its
phosphorylation state. Biochem J 334: 243-249

157

Ricciarelli R, Zingg J-M, Azzi A (2001) Vitamin E: protective role of a Janus molecule.
FASEB J 15: 2314-2325

Rimm EB, Stampfer MJ, Ascherio A, Giovannucci E, Colditz GA, Willett WC
(1993) Vitamin E consumption and the risk of coronary heart disease in men. N Engl J
Med 328: 1450-1456

Rise M, Cojocaru M, Gottlieb HE, Goldschmidt EE (1989) Accumulation of a-
tocopherol in senescing organs as related to chlorophyll degradation. Plant Physiol 89:
1028-1030

Rohdich F, Wungsintaweekul J, Eisenreich W, Richter G, Schuhr CA, Hecht S,
Zenk MH, Bacher A (2000a) Biosynthesis of terpenoids:4-diphosphocytidyl-2-C-methyl-
D-erythritol synthate of Arabidopsis thaliana. Proc Natl Acad Sci USA 97: 6451-6456

Rohdich F, Wungsintaweekul J, Fellermeier M, Sagner S, Herz S, Kis K,
Eisenreich W, Bacher A, Zenk MH (1999) Cytidine 5'-triphosphate-dependent

biosynthesis of isoprenoids: ngP protein of Escherichia coli catalyzes the formation of 4-
diphosphocytidyl-Z-C-methylerythritol. Proc Natl Acad Sci USA 96: 11758-11763

Rohdich F, Wungsintaweekul J, Luttgen H, Fischer M, Eisenreich W, Schuhr CA,
Fellermeier M, Schramek N, Zenk MH, Bacher A (2000b) Biosynthesis of terpenoids:
4-diphosphocytidyl-Z-C-methyl-D-erythritol kinase from tomato. Proc Natl Acad Sci USA
97: 8251-8256

Rohmer M, Knani M, Simonin P, Sutter B, Sahm H (1993) Isoprenoid biosynthesis in
bacteria: a novel pathway for the early steps leading to isopentenyl diphosphate. Biochem J
295: 517-524

Rohmer M, seemann M, Horbach S, Bringer-Meyer S, Sahm H (1996)
Glyceraldehyde 3-phosphate and pyruvate as precursors of isoprenic units in an alternative
non-mevalonate pathway for terpenoid biosynthesis. J Am Chem Soc 118: 2564-2566

Sandorf I, Hollander-Czytko H (2002) Jasmonate is involved in the induction of tyrosine
aminotransferase and tocopherol biosynthesis in Arabidopsis thaliana. Planta 216: 173-
179

Sattler S, DellaPenna D (2003) submitted to Plant Physiology

Savidge B, Weiss JD, Wong Y-HH, Lassner MW, Mitsky TA, Shewmaker CK,
Post-Beittenmiller D, Valentin HE (2002) Isolation and characterization of
homogentisate phytyltransferase genes from Synechocystis sp. PCC 6803 and Arabidopsis.
Plant Physiol 129: 321-332

Scarpa M, Rigo A, Maiorino M, Ursini F, Gregolin C (1984) Formation of 01-
tocopherol radical and recycling of a—tocopherol by ascorbate during peroxidation of
phosphatidylcholine lipsomes. Biochim Biophys Acta 801: 215-219

Schaller F (2001) Enzymes of the biosynthesis of octadecanoid-derived signaling
molecules. J Exp Bot 52: 11-23

Schledz M, Seidler A, Beyer P, Neuhaus G (2001) A Novel phytyltransferase from
Synechocystis sp. PCC 6803 involved in tocopherol biosynthesis. FEBS Lett 499: 15-20

158

Schmid J, Schaller A, Leibinger U, Boll W, Amrhein N (1992) The in vitro
synthesized tomato Shikimate kinase precursor is enzymatically active and is imported and
processed to the mature enzyme by chloroplasts. Plant J 2: 375-383

Schmidt J, Bubunenko M, Subramanian AR (1993) A novel operon organization
involving the genes for chorismate synthase (aromatic biosynthesis pathway) and ribosomal
GTPase center proteins (L11, L1, L10, L12: rleAJL) in cyanobacterium Synechocystis
PCC 6803. J Biol Chem 268: 27447-27457

Schultz G, 8011 J, F iedler E, Schulze-Siebert D (1985) Synthesis of prenquuinones in
chloroplasts. Physiol Plant 64: 123-129

Schulz A, Ort O, Beyer P, Kleinig H (1993) SC-0051, a 2-benzoyl-cyclohexane-l,3-
dione bleaching herbicide, is a potent inhibitor of the enzyme p-hydroxyphenylpyruvate
dioxygenase. FEBS Lett 318: 162-166

Schulze-Siebert D, Homeyer U, 8011 J, Schultz G (1987) Synthesis of plastoquinone-9,
(rt-tocopherol and phylloquinone (vitamin K1) and its integration in chloroplast carbon
metabolism of higher plants. In PK Stumpf, JB Mudd, WD Nes, eds, The metabolism,
structure, and function of plant lipids. Plenum Press, New York, pp 29-36

Schvedova AA, Kisin ER, Murray AR, Kommineni C, Castranova V, Mason RP,
Kadiiska MB, Gunther MR (2002) Antioxidant balance and fiee radical generation in
vitamin E-deficient mice after dermal exposure to curnene hydroperoxide. Chem Res Tox
15: 1451-1459

Schwender J, Seemann M, Lichtenthaler HK, Rohmer M (1996) Biosynthesis of
isoprenoids (carotenoids, sterols, prenyl side-chains of chlorophylls and plastoquinone) via
a novel pyruvate/glyceraldehyde 3-phosphate non-mevalonate pathway in the green alga
Scenedesmus obliquus. Biochem J 316: 73-80

Schwenke DC (2002) Does lack of tocopherols and tocotrienols put women at increased
risk of breast cancer. J Nutr Biochem 13: 2-20

Schwenke DC, Rude] LL, Sorci-Thomas MG, Thomas MJ (2002) alpha-Tocopherol
protects against diet induced atherosclerosis in New Zealand white rabbits. J Lipid Res 43:
1927-1938

Shah DM, Horsch RB, Klee HJ, Kishore GM, Winter JA, Turner NE, Hironaka
CM, Sanders PR, Gasser CS, Aykent S, Siegel NR, Rogers SG, F raley RT (1986)
Engineering herbicide tolerance in transgenic plants. Science 233: 478-481

Shintani D, DellaPenna D (1998) Elevating the Vitamin E content of plants through
metabolic engineering. Science 282: 2098-2100

Shintani DK, Cheng Z, DellaPenna D (2002) The role of 2-methyl-6-
phytylbenzoquinone methyltransferase in determining tocopherol composition in
Synechocystis sp. PCC6803. FEBS Lett 511: l-5

Shewmaker CK, Sheehy JA, Daley M, Colburn S, Ke DY (1999) Seed-specific

overexpression of phytoene synthase: increase in carotenoids and other metabolic effects.
Plant J 20: 401-412

159

Siesky AM, Kamendulis LM, Klauning JE (2002) Hepatic effects of 2-butoxyethanol
in rodents. Tox Sci 70: 252-260

Singh RJ, Goss SPA, Joseph J, Kalyanaraman B (1998) Nitration of y—tocopherol and

oxidation of (rt-tocopherol by copper-zinc superoxide dismutase/H,O,/NO,': Role of
nitrogen dioxide free radical. Biochemistry 95: 12912-12917

Slamenova D, Chalupa I, Robichova S, Gabelova A, Farkasova T, Hrusovska L,
Bacova G, Sebova L, Eckl P, Bresgen N, Zeitheim P, Schneider P, Wsolova L,
Barancokova M, Kazimirova A, Navarova J, Bezek S (2002) Effects of dietary intake
of vitamin A or E on the level of DNA damage, chromosomal aberrations, and micronuclei
induced in freshly isolated rat hepatocytes by different carcinogens. Nutr Cancer 42: 117-
124

Smith TL, Kummerow FA (1989) Effect of dietary vitamin E on plasma lipids and
atherogenesis in restricted anovulatory chickens. Atherosclerosis 75: 105-109

8011 J (1987) (rt-tocopherol and plastoquinone synthesis in chloroplast membranes.
Methods Enzymol 148: 383-392

8011 J, Kemmerling M, Schultz G (1980) Tocopherol and plastoquinone synthesis in
spinach chloroplasts subfractions. Arch Biochem Biophys 204: 544-550

Soil J, Schultz G (1979) Comparison of geranylgeranyl and phytyl substituted
methquuinols in the tocopherol synthesis of spinach chloroplasts. Biochem Biophys Res
Comm 9]: 715-720

Sol] J, Schultz G (1980) 2-methyl-6-phytquuinol and 2,3-dimethyl-5-phytquuinol as
precursors of tocopherol synthesis in spinach chloroplasts. Phytochemistry 19: 215-218

8011 J, Schultz G, Joyard J, Douce R, Block MA (1984) Localisation and synthesis of
prenquuinones in isolated outer and inner envelope membranes from spinach chloroplasts.
In P-A Siegenthaler, W Eichenberger, eds, Structure, function and metabolism of plant
lipids. Elsevier Science Publishers B. V., Amsterdam, pp 263-266

Sol] J, Schultz G, Joyard J, Douce R, Block MA (1985) Localization and synthesis of
prenquuinones in isolated outer and inner envelope membranes of spinach chloroplasts.
Arch Biochem Biophys 238: 290-299

Sol] J, Schultz G, Rudiger W, Benz J (1983) Hydrogenation of geranylgeraniol:
Two pathways exist in spinach chloroplasts. Plant Physiol 71: 849-854

Stampfer MJ, Hennekens CH, Manson JE, Colditz GA, Rosner B, Willett WC
(1993) Vitamin E consumption and the risk of coronary disease in women. N Engl J Med
328: 1444-1449

Stanier RY, Cohen-Bazire G (1977) Phototrophic prokaryotes: The cyanobacteria. Ann
Rev Microbiol 31: 225-274

Steinrucken HC, Amrhein N (1980) The herbicide glyphosate is a potent inhibitor of 5-

enolpyruvylshikimic acid 3-phosphate synthase. Biochem Biophys Res Comm 94: 1207-
1212

160

Stephens NG, Parson A, Schofield PM, Kelly F, Cheeseman K, Mitchinson MJ,
Brown MJ (1996) Randomized controled trial of vitamin E in patients with coronary
disease: Cambridge Heart Antioxidant Study (CHAOS). Lancet 347: 781-786

Stevens B, Small RD, Perez SR (1974) The photoperoxidation of unsaturated organic
molecules. XIII. Molecular oxygen (‘Ag) quenching by (rt-tocopherol. Photochem Photobiol
20: 515-517

Stillwell W, Dallman T, Dumaual AC, Crump T, Jenski LJ (1996) Cholesterol
versus (rt-tocopherol: Effects on properties of bilayers made fi'om heteroacid
phosphatidylcholines. Biochemistry 35: 13353-13362

Stocker A, Fretz H, Frick H, Ruttimann A, Woggon W-D (I996) The substrate
specificity of tocopherol cyclase. Bioorg Med Chem 4: 1 129-1134

Stocker A, Netscher T, Ruttimann A, Muller RK, Schneider H, Todaro LJ, G. D,
Woggon W-D (I994) The reaction mechanism of chromanol-ring formation catalyzed by
tocopherol cyclase from Anabaena variabilis Kutzing (cyanobacteria). Helv Chim Acta 77:
1721-1737

Stocker A, Ruttimann A, Woggon W-D (1993) Identification of the tocopherol-cyclase
in the blue-green algae Anabena variabilis Kutzing (Cyanobacteria). Helv Chim Acta 76:
1729-1738

Stocker A, Zimmer S, Spycher SE, Azzi A (1999) Identification of a novel cytosolic
tocopherol-binding protein: Structure, specificity, and tissue distribution. IUBMB Life 48:
49-55

Stoscheck CM (1990) Quantitation of protein. In MP Deutscher, ed, Guide to protein
purification. Academic Press, San Diego, pp 57-60

Streb P, Shang W, Feierabend J, Bligny R (1998) Divergent strategies of
photoprotection in hi gh-mountain plants. Planta 207: 313-324

Sylvester PW, McIntyre BS, Gapor A, Briski KP (2001) Vitamin E inhibition of normal
mammary epithelial cell growth is associated wit a reduction in protein kinase Cor activation.
Cell Prolif 34: 347-357

Syvaoja EL, Piironen V, Varo P, Koivistoinen P, Salminen K (1986) Tocopherols
and tocotrienols in finish foods - oils and fats. J Am Oil Chem Soc 63: 328-329

Takahashi S, Kuzuyama T, Watanabe H, Seto H (1998) A l-deoxy-D-xylulose 5-
phosphate reductoisomerase catalyzing the formation of 2-C-methyl-D-erythritol-4-
phosphate is an altemative non-mevalonate pathway for terpenoid biosynthesis. Proc Natl
Acad Sci USA 95: 9879-9884

Tanaka R, Oster U, Kruse E, Rudiger W, Grimm B (1999) Reduced activity of
geranylgeranyl reductase leads to loss of chlorophyll and tocopherol and to partially
geranylgeranylated chlorophyll in transgenic tobacco plants expressing antisense RNA for
geranylgeranyl reductase. Plant Physiol 120: 695-704

Tang F-Y, Meydani M (2001) Green tea catechins and vitamin E inhibit angiogenesis of

human microvascular endothelial cells through suppression of lL-8 production. Nutr
Cancer 41: 119-125

161

 

Taylor LA, Rose RE (1988) Correction in the nucleotide sequence of the TN903
kanamycin resistance determinant in pUC4K. Nucl Acid Res 16: 358-358

The ATBC Cancer Prevention Study Group (1994) The effect of vitamin E and beta
carotene on the incidence of lung cancer and other cancers in male smokers. N Engl J Med
330: 1029-1035

The HOPE Study Investigators (2000) Vitamin E supplementation and cardiovascular
events in high-risk patients. N Engl J Med 342: 154-160

Thomas DJ, Avenson TJ, Thomas JB, Herbert SK (1998) A cyanobacterium lacking
superoxide dismutase is sensitized to oxidative stress induced with methyl viologen but is
not sensitized to oxidative stress induced with norflurazon. Plant Physiol 116: 1593-1602

Threlfall DR, Whistance GR (1971) Biosynthesis of isoprenoid quinones and
chromanols. In T Goodwin, ed Aspects of terpenoid chemistry and biochemistry. Academic
press, Liverpool, England, pp 357-404

Traber MG, Arai H (1999) Molecular mechanisms of vitamin E transport. Annu Rev Nutr
19: 343-355

Traber MG, Burton GW, Ingold KU, Kayden HJ (1990) RRR- and SRR-or-
tocopherols are secreted without discrimination in human chylomicrons, but RRR-a-
tocopherol is preferentially secreted in very low density lipoproteins. J Lipid Res 31: 675-
685

Tramontano WA, Ganci D, Pennino M, Dierenfeld ES (1992) Age dependent or-
tocopherol concentrations in leaves of soybean and pinto beans. PhytochemistryI 31: 3349-
3351

Tran K, Wong JT, Lee E, Chan AC, Choy PC (1996) Vitamin E potentiates
arachidonate release and phospholipase A2 activity in rat heart myoblastic cells. Biochem J

319: 385-391

Tsegaye Y, Shintani DK, DellaPenna D (2002) Over-expression of the enzyme p-
hydroxyphenylpyruvate dioxygenase in Arabidopsis and its relation to tocopherol
biosynthesis. Plant Physiol Biochem 40: 913-920

Tsuchiya T, Ohta H, Okawa K, Iwamatsu A, Shimada H, Masuda T, Takamiya K
(1999) Cloning of chlorophyllase, the key enzyme in chlorophyll degradation: Finding of a
lipase motif and the induction by methyl jasmonate. Proc Natl Acad Sci USA 96: 15362-
15367

Uemura M, Manabe H, Yoshida N, Fujita N, Ochiai J, Matsumoto N, Takagi T,
Naito Y, Yoshikawa T (2002) a-Tocopherol prevents apoptosis of vascular endothelial
cells via a mechanism exceeding that of mere antioxidation. Eur J Pharmacol 456: 29-37

Voegele AF, Jerkovic L, Wellenzohn B, Eller P, Kronenberg F, Liedl KR,

Dieplinger H (2002) Characterization of the vitamin E-binding properties of human
plasma afamin. Biochemistry 41: 14532-14538

162

 

Wada H, Gombos Z, Murata N (1990) Enhancement of chilling tolerance of a
cyanobacteriumby genetic manipulationof fatty acid desaturation. Nature 347 : 200-203

Wada H, Gombos Z, Sakamoto T, Higashi S, Los DA, Heinz E, Schmidt H,
Nishida I, Murata N (1993) Fatty acid desaturation in cyanobacteria. In N Murata, CR
Somerville, eds, Biochemistry and Molecular Biology of Membrane and Storage Lipids in
Plants. The American Society of Plant Physiologist, Rockville, pp 67-78

Wang X, Quinn PJ (2000) The location and function of vitamin E in membranes. Mol
Membrane Biol 17: 143-156

Wang Y, Herrmann KM, Weller SC, Goldsbrough PB (1991) Cloning and nucleotide
sequence of a complementary DNA encoding 3-deoxy-D-arabino-heptulosonate 7-
phosphate synthase from tobacco. Plant Physiol 97: 847-848

Wassall SR, Thewalt JL, Wong L, Gorrissen H, Cushley RJ (1986) Deuterium NMR
study of the interaction of (rt-tocopherol with a phospholipid model membrane.
Biochemistry 25: 319-326

Watanabe N, Shiki Y, Morisaki N, Saito Y, Yoshida S (1986) Cytotoxic effects of
paraquat and inhibition of them by vitamin E. Biochim Biophys Acta 883: 420-425

Whistance GR, Threlfall DR (1970) Biosynthesis of phytoquinones. Homogentisic acid:
A precursor of plastoquinones, tocopherols and a—tocopherolquinone in higher plants, green
algae and blue-green algae. Biochem J 117: 593-600

Wildi B, Lutz C (1996) Antioxidant composition of selected high alpine plant species
from different altitudes. Plant Cell Environ 19: 138-146

Williams JGK (1988) Construction of specific mutations in photosystem II
photosynthetic reaction center by genetic engineering methods in Synechocystis 6803.
Methods Enzymol 167: 766-778

Wojcicki J, Rozewicka L, Barcew-Wiszniewska B, Samochoviec L, Juzwiak S,
Kadlubowska D, Tustanowski S, Juzysyn Z (1991) Effect of selenium and vitamin E
on the development of experimental atherosclerosis in rabbits. Atherosclerosis 87: 9-16

Wu K, Willett WC, Chan JM, Fuchs CS, Colditz GA, Rimm EB, Giovannucci EL
(2002) A prospective study on supplemental vitamin E intake and risk of colon cancer in
women and men. Cancer Epidemiol Biomarker Prevention 11: 1298-1304

Yamauchi J, lwamoto T, lKida S, Masushige S, Yamada K, Esashi T (2001)
Tocopherol-associated protein is a ligand-dependent transcriptional activator. Biochem
Biophys Res Comm 285: 295-299

Yamauchi R, Matsushita S (1977) Quenching effect of tocopherols on the methyl
linoleate photooxidation and their oxidation products. Agric Biol Chem 41: 1425-1430

Yamauchi R, Matsushita S (1979a) Light-induced lipid peroxidation in isolated
chloroplasts and role of a-tocopherol. Agric Biol Chem 43: 2157-2161

Yamauchi R, Matsushita S (1979b) Products formed by photosensitized oxidation of
tocopherols. Agric Biol Chem 43: 2151-2156

163

 

Ye B, Gressel J (2000) Transient, oxidant-induced antioxidant transcript and enzyme levels
correlate with greater oxidant-resistance in paraquat-resistant Conyza bonariensis. Planta
211: 50-61

Yoshida H, Takagi S, Ienaga H, Tsuchiya C (1998) Regional distribution of
tocopherols and fatty acids within soybean seeds. J Am Oil Chem Soc 75: 767-774

Yu WP, Simmons-Menchaca M, Gapor A, Sanders BG, Kline K (1999) Induction of

apoptosis in human breast cancer cells by tocopherols and tocotrienols. Nutr Cancer 33:
26-32

164