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This is to certify that the
dissertation entitled

Understanding Gene Expression and Metabolic Controls on

Fatty Acid Biosynthesis in Plants.

presented by
Gustavo Bonaventure

has been accepted towards fulfillment
of the requirements for the

 

 

Ph.D. /degc% in Genetics

 

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UNDERSTANDING GENE EXPRESSION AND METABOLIC CONTROLS ON
FATTY ACID BIOSYNTHESIS IN PLANTS

By

Gustavo Bonaventure

A DISSERTATION

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

DOCTOR OF PHILOSOPHY
Genetics Graduate Program

2004

ABSTRACT

UNDERSTANDING GENE EXPRESSION AND METABOLIC CONTROLS ON
FATTY ACID BIOSYNTHESIS IN PLANTS.

By
Gustavo Bonaventure
Plant fatty acids represent an important renewable source of highly reduced carbon chains
that can be used for purposes ranging from food production to industrial feedstocks. So
far, many attempts to engineer fatty acid production in plants have been based on current
knowledge of the fatty acid metabolic pathway and the structural genes involved in it.
However, plant tissues have proven to be not easily amenable to fatty acid engineering by
overexpression of single FAS enzymes. These experiments lead to the conclusion that
identification of regulatory factors limiting fatty acid production in plant tissues is crucial
for manipulation of this metabolic pathway. At present, these factors have not been
identified. Therefore, one major goal of this study has been to understand the signals and
mechanisms that control fatty acid biosynthesis and the expression of genes for this

primary metabolic pathway in plants.

The results presented in this study demonstrate that acyl carrier protein (ACP) genes in
Arabidopsis are under multiple levels of controls and suggest that some signals known to
activate fatty acid gene expression in non-plant organisms have been conserved in plants
(e. g., transcriptional activation by growth/cell cycle) whereas others differ (e.g.,
transcriptional activation by feedback mechanisms). Light has an important role on
expression of ACP genes, increasing ACP4 mRNA levels and promoting the association

of ACP transcripts with polyribosomes. Thus, this study demonstrates that different ACP

genes are regulated by specific signals depending on the tissue and its fatty acid

requirements.

In addition, this thesis has explored the production of saturated fatty acids and their
partitioning between diverse cellular processes by focusing on the characterization of an
Arabidopsis acyl-ACP thioesterase mutant (fatb-ko). Analysis of this mutant has revealed
the central role of FATB in the production of saturated fatty acids in all tissues.
Moreover, redistribution of these molecules between different biosynthetic pathways in
fatb-ko suggests that cells prioritize the synthesis of critical components for growth. In
addition, slower growth and production of non-viable seeds in the mutant demonstrate an

essential role of saturated fatty acids in plant growth and seed development.

Isotope labeling experiments and western blots were used to investigate fatty acid
metabolism and protein expression in fatb-ko leaves. The results indicate that fatb-ko
leaves increase the rate of fatty acid synthesis by 40 % compared to wild type. The
mutant also increases by 70% the initial rate of fatty acid breakdown. Western blot
analysis reveal that BCCP, ACPs and 18:0-ACP desaturase levels are increased by 1.5- to
2-fold in mutant leaves compared to wild type. Thus, plant cells appear to have
mechanisms that sense and respond to subnormal levels of saturated fatty acids in the
cytoplasm. These mechanisms probably coordinate the requirements for lipid synthesis in
the cytosol and fatty acid production inside the plastids. Moreover, the same mechanisms

may also activate expression of nuclear genes for fatty acid synthesis.

ACKNOWLEDGMENTS

Many people have contributed to this research and my graduate education. My thanks to
John Ohlrogge, my advisor, for giving me the opportunity to do my graduate studies in
his lab and for his support over the years. This work would not have been possible
without his guidance, the excellent working environment that his lab is, and the freedom I
found to follow my ideas. My committee members, David Amosti, Christoph Benning,
and Steve Triezenberg provided invaluable assistance, ideas and feedback over the years.
Amazingly, although they have extremely busy schedules they have always been willing
to meet with me, as well as with dozens of other grad students, to share their time and
knowledge. Members of the Ohlrogge lab taught me many things in my time here.
Special thanks to Mike Pollard for teaching me how to work with lipids on the bench and
sharing his time and knowledge with me. Special thanks to David Schultz, Xiaoming B30
and Abe Koo and also to Fred, Quino, Katrin, Joerg, Guiyun, Ajay, Sari, Vandana,
Naima, Troy, Linda, Anne, Jay, Phill, Greg, Yonghua, Sergei, etc. Jeannine Lee from the
Genetics Graduate program has also been of invaluable assistance over the years. My
thanks to Helmut Bertrand for being an excellent Director of the Genetics Program and
for helping me and my wife to establish at Michigan State University. Alicia Pastor-
Lecha and Ewa Danielewicz from the MSU Microscopy Facility provided me with
invaluable help with the Electron and Laser Confocal microscopes. Finally, my thanks to
my wife, Veronica, for all her love and for giving me total support and cheering me up
during the tough periods of research when nothing seems to work , and also for teaching

me to be more patient and tolerant in both science and life. To my daughter Marina for

iv

her unconditional love and teaching me that parenthood is totally compatible with science
and sometimes even inspiring. Without my parents and sisters none of my achievements
would have been possible. To all my friends at MSU that made all this possible: Rodrigo,
Maite, Hector, Lizzy, Gustavo, Claribel, Chris, Xiao, Emilio, Francisco, Setsuko, and

many many others.

TABLE OF CONTENTS

LIST OF FIGURES .......................................................................................................... IX
LIST OF TABLES ............................................................................................................ XI
CHAPTER 1 ....................................................................................................................... 1
INTRODUCTION .............................................................................................................. 1
Objectives ................................................................................................................... 6
Understanding regulation of F AS genes in Arabidopsis ............................................. 6
Transcriptional regulation of genes for fatty acid synthesis .................................... 7
Transcriptional regulation of F AS genes in microorganisms .................................. 7
Transcriptional regulation of FAS genes in animals ................................................ 9
Transcriptional regulation of FAS genes in plants ................................................ 10
Summary of Chapter 2 .............................................................................................. 12
Understanding regulation of fatty acid metabolism and FAS gene expression in a
fatb mutant of Arabidopsis ........................................................................................ 13
Fatty acid synthesis in plants ................................................................................. 15
The role of fatty acids in the synthesis of different cellular components .............. 22
Membrane glycerolipids ........................................................................................ 22
Other fatty acid-derived products .......................................................................... 24
Regulation of fatty acid biosynthesis in plants ...................................................... 25
Regulation by light ................................................................................................. 25
Regulation by feedback inhibition ......................................................................... 26
Regulation by demand ........................................................................................... 27
LITERATURE CITED ................................................................................................. 28
CHAPTER 2 ..................................................................................................................... 33
DIFFERENTIAL REGULATION OF MESSENGER RNA LEVELS OF ACP
ISOFORMS IN ARABIDOPSIS ...................................................................................... 33
ABSTRACT .................................................................................................................. 33
INTRODUCTION ........................................................................................................ 34
RESULTS ..................................................................................................................... 37
ACP4 mRNA Levels are Increased by Light ............................................................ 37
Polyribosomal Association of ACP mRNAs is Increased by Light ......................... 40
Polyribosomal association of ACP mRNAs in developing seeds of wild type
B.napus and transgenics over—expressing MCTE ..................................................... 42
Conserved Motifs occur in the 5’ Leader Region of ACP mRNAs .......................... 43

The 5’ Leader Sequences of ACP] and ACP2 Increase Reporter Gene Expression 46

ACP mRNA Levels are Affected by a Sucrose-derived Signal and/or Growth
Control in Cell Suspension Cultures ......................................................................... 49

vi

ACP2 mRNA Levels are Affected by a Sucrose-derived Signal and/or Growth

Control in planta ....................................................................................................... 52
Transcriptional regulation of Arabidopsis ACP2 gene by growth/cell cycle signals 54
Deletion analysis of Arabidopsis ACP2 promoter in tobacco BY-2 cells ................ 59
Conserved elements in Arabidopsis ACP promoters ................................................ 62
Changes in mRNA expression by inhibition of fatty acid synthesis in Arabidopsis
cells ........................................................................................................................... 65
FAS genes do not respond to inhibition of fatty acid biosynthesis in Arabidopsis
cells ........................................................................................................................... 67
DISCUSSION ............................................................................................................... 69
MATERIALS AND METHODS .................................................................................. 78
LITERATURE CITED ................................................................................................. 87
CHAPTER 3 ..................................................................................................................... 94
DISRUPTION OF THE FA TB GENE IN ARABIDOPSIS DEMONSTRATES AN
ESSENTIAL ROLE OF SATURATED FATTY ACIDS IN PLANT GROWTH .......... 94
ABSTRACT .................................................................................................................. 94
INTRODUCTION ........................................................................................................ 95
RESULTS ................................................................................................................... 100
Mutant Isolation and Complementation Analysis ................................................... 100
FATB mRNA Expression Analysis ........................................................................ 102
FATB is Essential for Normal Seedling Growth .................................................... 103
Leaf and chloroplast morphology of wild type Arabidopsis and fab-k0 ................ 106
F ATB is Essential for Normal Seed Morphology and Germination ....................... 109
Fatty Acid Composition of fatb-ko Tissues. ........................................................... 110
Fatty Acid Composition of Individual Leaf Glycerolipids. .................................... 112
Acyl-ACP Thioesterase Activity ............................................................................ 112
Total Palmitate Content in Arabidopsis Leaf Tissue .............................................. 114
Leaf and Stem Surface Wax Analysis .................................................................... 115
Sphingoid Base Analysis ........................................................................................ 116
Leaf and stem cutin analysis ................................................................................... 117
fatb-ko act] double mutant ..................................................................................... 119
DISCUSSION ............................................................................................................. 121
Seedling Growth and Seed Development ............................................................... 123
Reduced Export of Palmitate in fatb-ko plants and Other Sources of Palmitate and
Stearate in the Cell .................................................................................................. 125
Partition of Palmitate and Stearate to Non—glycerolipid Products. ......................... 127
Conclusions ............................................................................................................. 131
MATERIALS AND METHODS ................................................................................ 133
LITERATURE CITED ............................................................................................... 142
CHAPTER 4 ................................................................................................................... 146

vii

DISRUPTION OF THE FA TB GENE INCREASES THE RATE OF FATTY ACID
BIOSYNTHESIS,FATTY ACID AND LIPID TURNOVER, AND F AS PROTEIN

EXPRESSION IN LEAVES. .......................................................................................... 146
ABSTRACT ................................................................................................................ 146
INTRODUCTION ...................................................................................................... 147
RESULTS ............... . .................................................................................................... l 5 1

Rate of fatty acid biosynthesis in leaves of wild type Arabidopsis and fatb-ko. 151
Rate of fatty acid turnover in wild type and fatb-ko leaves .................................... 153
Analysis of polar lipid classes in wild type and fatb-ko leaves .............................. 154
Analysis of radiolabeled C16 and C18 fatty acids in polar lipids of wild type and fatb-
k0 leaves .................................................................................................................. 157
Mass distribution of C132C13 and ClozClg molecular species of polar lipids in wild
type and fatb-ko leaves ............................................................................................ 160
Immunoblot analysis of wild type and fatb-ko leaves. ........................................... 164
DISCUSSION ............................................................................................................. 166
Increased rates of fatty acid biosynthesis and turnover in fatb-ko .......................... 168
Turnover of Polar Lipids in wild type and fatb-ko ................................................. 173
Conclusions and Future Perspectives ...................................................................... 174
MATERIALS AND METHODS ................................................................................ 175
LITERATURE CITED ............................................................................................... 181
CHAPTER 5 ................................................................................................................... 185
FINAL CONCLUSIONS, PERSPECTIVES AND FUTURE DIRECTIONS ............... 185
Conclusions and future directions on FAS gene expression in Arabidopsis .......... 185
Major conclusions of this thesis research related to F AS gene expression are: .. 185
Future directions .................................................................................................. 188
Conclusions and future directions on fatty acid metabolism and its regulation in fatb-
k0. ............................................................................................................................ 192
Major conclusions of this thesis research related to the role of saturated fatty acid
are: ........................................................................................................................... 192
Future directions .................................................................................................. 194
LITERATURE CITED ............................................................................................... 197

viii

LIST OF FIGURES

FIGURE 1. FATTY ACIDS EXPORTED FROM PLASTIDS HAVE DIVERSE ROLES IN PLANT
CELLS. .......................................................................................................................... 3

FIGURE 2. SIMPLIFIED REPRESENTATION OF THE FATTY ACID BIOSYNTHESIS PATHWAY IN
ARABIDOPSIS (ACCORDING TO OHLROGGE AND BROWSE, 1995). .............................. 16

FIGURE 3. SCHEMATIC REPRESENTATION OF THE ENZYMATIC REACTION CATA LYZED BY
ACYL-ACP THIOESTERASES IN PLANTS. ..................................................................... 18

FIGURE 4. SIMPLIFIED SCHEME OF THE GLYCEROLIPID BIOSYNTHETIC PATHWAY IN
ARABIDOPSIS (ADAPTED FROM BROWSE AND SOMERVILLE, 1991). ........................... 20

FIGURE 5. LIGHT-MEDIATED INDUCTION OF ACP4 mRNA LEVELS IN ARABIDOPSIS LEAF
TISSUE. ....................................................................................................................... 39

FIGURE 6. LIGHT AFFECTS THE POLYRIBOSOME ASSOCIATION OF ACP MRNAS .............. 41

FIGURE 7. POLYSOME DISTRIBUTION OF ACP MRNAS IN WILD TYPE AND TRANSGENIC
(MCTE) B. NAPUS DEVELOPING SEEDS ....................................................................... 43

FIGURE 8. CONSTRUCTS USED FOR ARABIDOPSIS TRANSFORMATION. ............................. 46

FIGURE 9. REPORTER GENE EXPRESSION IN DIFFERENT TISSUES OF ARABIDOPSIS
TRANSGENIC PLANTS. ................................................................................................. 48

FIGURE 10. DIFFERENTIAL REGULATION OF ACP MRNA LEVELS IN STARVED
ARABIDOPSIS CELLS ................................................................................................... 51

FIGURE 11. DIFFERENTIAL REGULATION OF ACP MRNA LEVELS IN ARABIDOPSIS CELLS
GROWN IN THE PRESENCE OF SUCROSE. ...................................................................... 53

FIGURE 12. IN PLANTA REGULATION OF ACP2 MRNA BY CARBON .......................... 55

FIGURE 13. REGULATION OF ARABIDOPSIS ACP2 PROMOTER BY CARBON IN ARABIDOPSIS
CELL SUSPENSION CULTURE ................................................................. 56

FIGURE 14. REGULATION OF ACP2 PROMOTER BY DIFFERENT CARBON SOURCES IN
ARABIDOPSIS CELL SUSPENSION CULTURES ................................................. 58

FIGURE 15. ARABIDOPSIS ACP2 PROMOTER DELETION ANALYSIS IN TOBACCO BY-2
CELLS ............................................................................................ 61

FIGURE 16. CONSERVED ELEMENTS IN ARABIDOPSIS ACP PROMOTERS. ......................... 63

ix

FIGURE 17. Inhibition of fatty acid synthesis by cerulenin in Arabidopsis cells .......... 68

FIGURE 18. SCHEMATIC REPRESENTATION OF THE ENZYMATIC REACTION CATALYZED BY
ACYL-ACP THIOESTEASES (FAT). ............................................................................. 96

FIGURE 19. STRUCTURE OF THE ARABIDOPSIS FA TB GENE CARRYING THE T-DNA
INSERTION. ............................................................................................................... 102

FIGURE 20. GROWTH AND MORPHOLOGY OF ARABIDOPSIS WILD-TYPE AND FA TB—KO
PLANTS AND SEEDS ........................................................................... 105

FIGURE 21. GROWTH CURVES OF ARABIDOPSIS WILD-TYPE AND FA TB-KO PLANTS. ....... 107

FIGURE 22. MICROSCOPIC ANALYSIS OF LEAF SECTION AND CHLOROPLAST STRUCTURE OF
WILD TYPE AND FA TB-KO SEEDLINGS ....................................................... 108

FIGURE 23. LEAF AND STEM CUTIN COMPOSITION OF WILD TYPE ARABIDOPSIS AND FA TB-
KO(MOL%)... ....120

FIGURE 24. GROWTH AND MORPHOLOGY OF ARABIDOPSIS WILD-TYPE, FA TB-KO, ACT]
AND FA TB-KO ACT] PLANTS. ..................................................................................... 122

FIGURE 25. SIMPLIFIED SCHEME OF PREDICTED FLUXES OF C16 AND C18 FATTY ACIDS IN
MEMBRANE LEAF GLYCEROLIPIDS OF ARABIDOPSIS WILD TYPE, FA TB-KO AND FA TB-KO
A('TI MUTANTS ......................................................................................................... 128

FIGURE 26. FATTY ACID TURNOVER IN WILD TYPE ( o ) AND FA TB-KO( o ) LEAVES. ...... 154

FIGURE 27. REDISTRIBUTION OF RADIOACTIVITY AMONG GLYCEROLIPID CLASSES OF WILD
TYPE ( o ) AND FA TB-K0( o ) LEAVES ....................................................... 156

FIGURE 28. REDISTRIBUTION OF RADIOACTIVITY IN C”, FATTY ACIDS OF MEMBRANE
GLYCEROLIPID CLASSES FROM WILD TYPE ( o ) AND FA TB-K0( o ) LEAVES. . . . . . . . 1 59

FIGURE 29. REDISTRIBUTION OF C18: ng AND szcjg MOLECULAR SPECIES OF POLAR
LIPIDS IN WILD TYPE AND FA TB-KO ........................................................................... 162

FIGURE 30. SCHEME OF MASS LOSS FROM LABELED FATTY ACIDS IN LEAVES OF WILD TYPE
AND FATB-K0..... .................................................................................................... 165

FIGURE 31. IMMUNOBLOT ANALYSIS OF BCCP, ACP AND 18:0-ACP DESATURASE IN LEAF
TISSUE OF WILD TYPE AND FA TB—KO ARABIDOPSIS. .................................................. 167

FIGURE 32. COMPARATIVE SCHEME OF FATTY ACID METABOLISM IN WILD TYPE AND FA TB-
K0 ............................................................................................................................ 170

LIST OF TABLES
TABLE 1. FUNCTIONS OF LIPID MOLECULES IN HIGHER PLANTS ......................................... 2

TABLE 2. PROXIMAL UPSTREAM SEQUENCES OF ACP GENES IN DIFFERENT PLANT SPECIES
................................................................................................... 45

TABLE 3 . CHANGES IN mRNA ABUNDANCE OF GENES INVOLVED IN LIPID METABOLISM
AFTER INHIBITION OF FATTY ACID SYNTHESIS IN ARABIDOPSIS CELLS. ...................... 70

TABLE 4. EXAMPLE OF TRANSCRIPTS THAT INCREASE MORE THAN 2 FOLD AFTER 6 HOURS
OF CERULENIN TREATMENT. ....................................................................................... 71

TABLE 5. EXAMPLE OF TRANSCRIPTS THAT DECREASE MORE THAN 2-FOLD AFTER 6 HOURS
OF CERULENIN TREATMENT. ....................................................................................... 72

TABLE 6 RELATIVE AMOUNTS OF FATB MRNA IN LEAF TISSUE OF WILD-TYPE
ARABIDOPSIS AND FA TB-KO MUTANT ...................................................................... 103

TABLE 7. ARABIDOPSIS WILD-TYPE AND FA TB-KO BOLTING TIME AND GERMINATION
RATES. ..................................................................................................................... 1 10

TABLE 8. FATTY ACID COMPOSITION OF WILD-TYPE AND FA TB-KO ARABIDOPSIS TISSUES
(MOL%). .................................................................................................................. 1 1 1

TABLE 9. FATTY ACID COMPOSITION OF LEAF GLYCEROLIPIDS OF ARABIDOPSIS WILD-
TYPE (WS) AND FA TB-KO MUTANT (MOL%). ............................................................ 113

TABLE 10. MAJOR COMPONENTS OF EPICUTICULAR LEAF WAXES FROFROM WILD-TYPE
ARABIDOPSIS AND FA TB-KO MUTANT (MOL%). ....................................................... 116

TABLE 11. SPHINGOID BASE CONTENT OF LEAF TISSUE FROM WILD-TYPE ARABIDOPSIS
AND FA TB-KO PLANTS (MOL%) ................................................................................. 118

TABLE 12. LEAF FATTY ACID COMPOSITION OF WILD TYPE ARABIDOPSIS (COLUMBIA),
ACT] AND FA TB-KO A ("T1 DOUBLE MUTANT (MOL%) ................................................. 123

TABLE 13. RATES OF FATTY ACID SYNTHESIS (FAS) IN LEAVES OF ARABIDOPSIS WILD
TYPE AND FA TB-KO MEASURED BY DIFFERENT RADIOLABELED SUBSTRATES ............ 152

xi

CHAPTER 1

INTRODUCTION

”It takes a membrane to make sense out of disorder in biology. When the earth came
alive, it began constructing its own membrane, for the general purpose of editing the sun.
You have to be able to catch energy and hold it, storing precisely the needed amount and
releasing it in measured shares. A cell does this, and so do the organelles inside. Each
assemblage is poised in the flow of solar energy, tapping off energy from metabolic
surrogates of the sun. To stay alive, you have to be able to hold out against equilibrium,
maintain imbalance, bank against entropy, and you can only transact this business with
membranes in our kind ofworld Lewis Thomas.

 

 

Plant lipids are extremely diverse in structure and function (Table 1) and constitute the
products of several distinct biosynthetic pathways. These molecules serve several
essential functions in all organisms but their most critical role for life is the formation of
cellular membranes. These membrane structures form the major barriers that delineate
cells and their compartments and allow essential processes to occur (e.g., anabolic and
catabolic reactions, transport by vesiculation and permeation, photon capture) (Rausch

and Bucher, 2002; Bowsher and Tobin, 2001; Nebenfuhr and Staehelin, 2001).

Among the many types of plant lipids, glycerolipids are usually the most abundant in
plant cells (Ohlrogge and Browse, 1995). These lipids are the main constituent of cellular
membranes and are derived from the glycerol and fatty acid biosynthetic pathways
(Somerville et al., 2000). The fatty acid synthesis (FAS) pathway is a primary metabolic
pathway, because it is found in every cell of the plant and is essential for growth. In
addition to supplying fatty acids for the synthesis of membrane lipids, this pathway

provides substrates for several other essential cellular processes in plants. For example,

synthesis of sphingolipids, epicuticular waxes and cutin in epidermal cells and

triacylglycerols in seeds. Moreover fatty acids are substrates for acylation reactions (e. g.,

proteins, sterols) and also precursors of signaling molecules (Figure 1).

Table 1. Functions of lipid molecules in higher plants

 

Function

Lipid types

 

Membrane components

Glycerolipids, Sphingolipids, Sterols

 

Storage compounds

Triacylglycerols, Waxes

 

Compounds active in electron transfer

Chlorophylls, Ubiquinone, Plastoquinone

 

Photoprotection and free radical protection

Carotenoids, Tocopherols

 

Surface protection

Cutin, Suberin, Waxes, Triterpenes

 

Protein modification

14:0, 16:0, Farnesyl and GeranylGeranyl

 

 

Signaling

 

pyrophosphate, Phosphatidylinositol,
Ceramides, Dolichol

Jasmonate, Diacylglycerol, ABA, GA,
Brassinosteroids

 

As a result of the dependence of multiple cellular processes on the production of fatty

acids, the synthesis of these molecules is exquisitely controlled to balance supply and

demand for acyl chains. For plant cells, this means matching the level of fatty acid

synthesis to membrane biogenesis and to multiple cellular pathways depending on the

 

 

Figure 1. Fatty acids exported from plastids have diverse roles in plant cells.

In mesophyll cells fatty acids are used primarily for membrane glycerolipid synthesis. In
epidermal cells the bulk of fatty acids goes into waxes and cutin. In seeds,
triacylglycerols accumulate most of the exported fatty acids. In most cells saturated fatty
acids are precursors of ceramides and substrates of acylation reactions.

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tissue, growth rate, stage of development and time of the day. For example, leaves
increase by more than 10-fold the production of fatty acids during the day and some
seeds accumulate 60 % of their weight in oil (Ohlrogge and Jaworski, 1997). Thus, a
critical biochemical aspect of fatty acid biosynthesis in plants is to understand how cells
adjust the production of fatty acids according to different demands and how they partition
these molecules into several cellular pathways. Completion of the Arabidopsis genome
sequence has accelerated the identification of structural enzymes involved in the
synthesis of fatty acids and glycerolipids, and most of them have been either
experimentally described or have strong candidates (Beisson et al., 2003; Mekhedov et
al., 2000). By contrast, advances in the identification of regulatory factors and
mechanisms of control for this metabolic pathway have been slower. Thus, a number of

major question in the synthesis of fatty acids in plants remain unanswered:

> How are fatty acid and lipid biosynthetic genes regulated?

> Do global trans-acting factors exist that simultaneously control the expression of
most genes?

> What signals control the promoter activity of these genes?

> How is the production of fatty acids controlled in plants?

‘9 What signals are sensed to adjust fatty acid synthesis rates to different cellular
demands of fatty acids?

> How is the production of fatty acids in the plastids coordinated with their utilization

outside this organelle?

\\

r Do signals communicate between the cytosol and the plastid fatty acid synthesis

pathway?

Objectives

Based on these questions, the main objectives of the present work are to understand the
signals that control the rate of fatty acid synthesis in plants, the expression of genes for
this primary pathway and the production and partition of exported fatty acids from

plastids.

Understanding regulation of FAS genes in Arabidopsis

TO begin to answer some of these important aspects of plant gene regulation and
metabolism, Chapter 2 presents data on the signals and mechanisms that regulate the
expression of Arabidopsis ACP isoforms. Why do we study ACP gene expression? First,
these proteins are in the core of the fatty acids synthesis machinery and play a central role
during acyl chain synthesis. Due to this pivotal role, it is likely that signals and
mechanisms that affect ACP expression will also affect the expression of other fatty acid
genes. Second, ACP mRNA and protein levels are among the most abundant of the fatty
acid pathway, which in general are of low abundance. Third, ACP expression studies
such as tissue specific expression and promoter analysis have been previously reported

and provide partial but important information about regulatory signals and mechanisms

(Baerson et al., 1994; Hlousek-Radojcic et al., 1992; Battey and Ohlrogge, 1990;

Hannapel and Ohlrogge, 1988).

Transcriptional regulation of genes for fatty acid synthesis

Clues to the signals and mechanisms that activate plant FAS gene expression can be
obtained from known signals and mechanisms in non-plant organisms, if some of these
signals and mechanisms are conserved between distantly related living organisms.
Considering the universal role of lipids in cell structure and regulatory processes it may
be possible that fatty acid production and its genetic regulation Show, in some aspects,
close resemblance in divergent organisms. For the purpose of reviewing how FAS genes
are controlled in non-plant organisms, a short overview of transcriptional regulation of
FAS genes in these organisms is given below. In addition, a brief summary of previous

studies that analyzed different aspects of FAS gene expression in plants is presented.

Transcriptional regulation of FAS genes in microorganisms

The genes for fatty acid metabolism in E. coli are scattered about the genome with only
two clusters, the minimal accBC and the fab cluster (Cronan and Rock, 1996). All genes
are under regulation of F adR, a protein with dual functions that represses genes involved
in fatty acid degradation and activates genes involved in fatty acid synthesis. In the
absence of long chain acyl-COA, FadR directly binds to specific DNA sequences to

simultaneously activate transcription of biosynthetic genes and repress catabolic genes.

Thus, during exponential growth or in the absence of exogenous fatty acids, the pool of
acyl-COA remains very small and FadR activates biosynthetic genes and Shuts off
catabolic gene expression. When growth slows down or exogenous fatty acids are
abundant, the acyl-COA levels build up and FadR de-activates expression of FAS genes
and de-represses catabolic genes. Thus, in these organisms, FAS genes are almost
exclusively controlled by growth or exogenous fatty acids and respond to the demand for

new membranes (Cronan and Rock, 1996).

In B. subtilis the genes involved in fatty acid metabolism are located mainly in gene
clusters (e. g., the fabHAF operon) and are coordinately regulated by FapR. This regulator
is a transcription factor that represses the expression of lipid metabolic genes by binding
to a consensus sequence contained in their promoter regions (Schujman et al., 2003).
Thus, expression of fatty acid gene clusters is de-repressed when cells are in the
exponential phase of growth. Interestingly enough, addition of inhibitors of fatty acid
biosynthesis (e.g., cerulenin) can also de-repress these genes, suggesting that feedback
mechanisms of gene expression exist in this organism. During inhibition of FAS by
cerulenin, the intracellular concentration of malonyl-COA is increased (Heath and Rock,
1995) and it is proposed that intracellular levels of malonyl-COA regulates FapR activity.
Thus, if the rate of fatty acid synthesis falls below the normal levels (as a result of slow
growth or inhibition), a transient increase in the intracellular concentration of malonyl-
COA relieves FapR mediated repression of lipid biosynthetic genes (Schujman et al.,

2003).

In yeast, the synthesis of fatty acids is catalyzed by two multifunctional enzymes, acetyl
COA carboxylase (ACC) and fatty acid synthase (FAS). ACC is a tetramer of identical
subunits encoded by the FAS3 gene (Schuller et al., 1992). FAS consists of two
multifunctional proteins. a and [3, which are encoded by two unlinked genes, FAS] and
FAS2, respectively (Chirala, 1992). FAS], FASZ and FAS3 genes are coordinately
regulated by growth and exogenous fatty acids. Moreover, when FASZ is over-expressed,
expression of both FASl and FAS3 is also increased, suggesting that the FAS genes are
coordinately regulated in yeast. The conserved GCCAAA element (UASfas) is present in
all three FAS promoters and Specifically enhances the transcription of FAS genes. In
addition, FAS] and FASZ share an UAS.no element that is common to genes involved in
phospholipid biosynthesis. UAS,-no is a positive regulator of gene expression and is
required for efficient expression of these genes. UASfas and UASmo act synergistically for
Optimal expression of the FAS genes (Chirala, 1992; Schuller et al., 1992). In summary,
microorganisms possess global trans-acting factors that bind to consensus sequences
conserved in most of their FAS gene promoters and coordinately control the expression

of most of these genes.

Transcriptional regulation of FAS genes in animals

Animals obtain fatty acids almost exclusively from their diet, and they have developed

mechanisms that sense lipidic molecules such as cholesterol, fatty acids, fat-soluble

vitamins and other lipids present in their food. These molecules or their derivatives bind

to receptors in the nucleus or cytoplasm that directly or indirectly activate gene

expression (Chawla et al., 2001 ).

For example, one important group of receptors is the PPAR family (peroxisome
proliferator-activated receptor), that is activated by polyunsaturated fatty acids and
eicosanoids. PPARS belong to the nuclear receptor superfamily of transcription factors
that contain a large COOH-terminal region with a ligand binding domain. Upon ligand
binding, these transcription factors form heterodimers with the retinoid X receptor (RXR)
and bind to specific DNA sequences found in their target promoters known as hormone
response elements (HRE) (Chawla et al., 2001). Among their functions, PPARS can up-
regulate proteins that increase metabolism and transport of fatty acids into the

peroxisome and promote fat storage in the liver (Chawla et al., 2001).

A second group of regulatory factors of fatty acid metabolism in animals is the sterol
regulatory element binding protein (SREBP) family. These transcription factors are
located on ER membranes in an inactive form, and are released by proteolysis in order to
enter the nucleus. Proteolysis of SREBPS is inhibited by sterols and polyunsaturated fatty
acids in mammals, providing a feedback regulatory mechanism for lipid synthesis
(Dobrosotskaya et al., 2002). SREBPS interact with an escort protein, SCAP (SREBP
cleavage-activating protein) that serves as sensors of sterols, fatty acids or phospholipids

in animal cells (Seegmiller et al., 2002).

Transcriptional regulation of FAS genes in plants

10

Similarly to other higher eukaryotes, multiple cis elements and trans factors are expected
to be involved in regulation of plant fatty acid gene expression. However, in contrast to
animals, which obtain fatty acids mainly from their diets, plants depend on their own
production of fatty acids to subsist. In addition, the plant fatty acid machinery resembles
that from bacteria and is plastid localized, although almost entirely nuclear encoded.
Thus, regulatory mechanisms of fatty acid gene expression that communicate between
plastids and nucleus are probably present in plant cells (Jarvis, 2001). The essential
differences in the structure of fatty acid synthesis between animals and plants are most
likely reflected in the mechanisms and Signals by which fatty acid and lipid genes are
regulated in these organisms. For example, no obvious homologues of animal PPARS and

SREBPS are evident in the Arabidopsis genome.

One group of plant fatty acid genes that has received special attention is the ACP gene
group. Baerson and Lamppa (1993) fused an approximate 900 bp fragment upstream of
the Arabidopsis ACP2 gene to B-glucuronidase (GUS) and used the construct to generate
transgenic tobacco plants. GUS expression was detected during seed development, in
young leaves, in leaf epidermis, and flowers. In a second study, the same authors
analyzed a series of six deletions of the same promoter in tobacco transgenic plants
(Baerson et al., 1994). The results revealed distinct regions of the promoter involved in
vegetative and reproductive development. A —320 to —236 bp region was important for
expression of the reporter gene in leaves, whereas it did not alter expression in seeds and

flowers. Seed expression was reduced when a —235 to ~55 bp region was deleted. This

11

region was also essential for expression in flowers. Thus, transcriptional regulation of the
Arabidopsis ACP2 gene in different tissues involves different regions of the promoter and

presumably tissue specific factors.

Similarly, a deletion analysis of the Arabidopsis enoyl-ACP reductase promoter showed
that three domains of the promoter were important for differential tissue expression. First,
seed expression was unchanged by deletion to —47 bp of the transcription start Site,
suggesting that seed Specific cis elements are located close to the transcription starting
site. Second, removal of an intron in the 5’UTR resulted in increased expression in roots,
suggesting the presence of negative regulatory elements in this region. A third region was

important for high expression in young leaf tissue.

Other studies reported the light-dependent expression of FAD7 (plastidial desaturase)
(Nishiuchi et al., 1995), cold-dependent expression of FAD8 (also a plastidial desaturase)
(Vijayan and Browse, 2002), and low-phosphate induction of SQDI and SQD2 (Yu et al.,
2002; Essigmann et al., 1998). Thus, a subset of genes involved in lipid metabolisms
respond to specific Signals depending on their distinct functions. In addition to specific
environmental signals, the fatty acid and lipid genes that have been analyzed so far Show
a common denominator, high levels of expression in rapidly expanding tissue where cell
division is active (e.g., apical meristems), and also in developing seeds and flowers,

where lipid accumulation occurs.

Summary of Chapter 2

l2

In Chapter 2, we tested signals known to activate fatty acid and lipid gene expression in
non-plant organisms for their ability to activate fatty acid genes in plant cells (e.g.,
growth-dependent expression, inhibition of fatty acid synthesis, sugar-sensing
mechanisms). Results suggested that some of these signals but not all differentially
increase expression of some ACP isoforms. Thus, it appears that some regulatory Signals
have been conserved throughout evolution between plants, animals and lower organisms.
For example, similarly to bacteria and yeast, ACP2 expression appears to respond to
growth/cell division Signals, whereas inhibition of fatty acid synthesis has no effect on
gene expression linked to fatty acid metabolism in plants. Analysis of the Arabidopsis
ACP2 promoter demonstrated that activation by growth signals acts at the transcriptional
level and identified promoter regions important for regulation and expression of ACP2.
Light, which stimulates growth and chloroplast development has an important role on
ACP gene expression at the levels of ACP4 mRNA abundance and association of ACP
transcripts with polysomes. We also demonstrated that the 5’UTR of ACP transcripts
increases gene expression and determines tissue specific expression. In summary,
Chapter 2 discloses multiple levels of control on ACP gene expression and provides the
basis for further exploration of gene regulation for fatty acid metabolism and also

identification of regulatory factors for these genes.

Understanding regulation of fatty acid metabolism and FAS gene expression in a
fatb mutant of Arabidopsis

13

Chapter 3 takes a different approach and centers on the characterization of an
Arabidopsis FA TB knock-out mutant (fatb—k0). FATB belongs to the acyl-ACP
thioesterase family of proteins and alteration of its function by either mutation or over-
expression has proved to be instructive in disclosing mechanisms of regulation of fatty
acid metabolism (Eccleston and Ohlrogge, 1998; Ohlrogge et al., 1995). These
observations together with our interest in understanding production of saturated fatty
acids in plants and their role in development, prompted us to characterize the fatb-k0
mutant. Thus, Chapter 3 describes the central role of FATB in producing saturated fatty
acids and the essential role of these molecules in plant growth and development. In
addition, analysis of saturated fatty acid derivatives provides evidence for the
involvement of F ATB in supplying fatty acids to distinct biosynthetic pathways and how
cells control the economy of saturated fatty acids. These results provide clues to
understand the effects of reduced saturated fatty acids on plant growth and set the basis

for the experiments describe in Chapter 4.

Chapter 4 illustrates the importance of altering acyl-ACP thioesterase expression in
disclosing regulatory signals and mechanisms for regulation of fatty acid metabolism.
fatb-k0 leaves respond to low levels of saturated fatty acids by increasing the rate of fatty
acid synthesis by 40 % compared to wild type. Moreover, western blot analysis revealed
that BCCP, ACPS and 18:0-ACP desaturase levels are increased by 1.5- to 2-fold in
mutant leaves compared to wild type. Thus, higher rates of fatty acid synthesis are
achieved at least in part by increasing fatty acid gene expression. In summary, plant cells

appear to have mechanisms that sense and respond to subnormal levels of saturated fatty

l4

acids in the cytoplasm. These mechanisms probably coordinate the requirements for lipid
synthesis in the cytosol and fatty acid production inside the plastids. Moreover, the same

mechanisms may also activate expression of nuclear fatty acid genes.

With the purpose of introducing the role of acyl-ACP thioesterases in fatty acid synthesis
in plants together with the role of fatty acids in the synthesis of different cellular
components and the current knowledge in regulation of fatty acid synthesis, a brief

overview ofthese topics is given below.

Fatty acid synthesis in plants

Plants are fundamentally different from other eukaryotes in the molecular organization of
the enzymes of fatty acids biosynthesis (FAS). The individual enzymes of this primary
pathway are dissociable soluble components located in the stroma of the plastids whereas
in other eukaryotes fatty acid synthesis is catalyzed by multifunctional polypeptide
complexes located in the cytosol (Somerville et al., 2000; Schuller et al., 1992; Chirala,

1992).

The central carbon donor for fatty acid biosynthesis is the malonyl-COA produced by
acetyl-COA carboxylase (ACCase). In dicot plants, plastidic ACCase is a mutisubunit
enzyme composed of four dissociated polypeptides (biotin carboxyl carrier protein
[BCCP], biotin carboxylase [BC], and a- and B-carboxyl transferases (CT). BCCP, BC
and d-CT subunits are nuclear encoded whereas B-CT is plastome encoded (Sasaki et al.,

1993). Before entering the fatty acid biosynthetic pathway, malonyl-COA is transferred

15

from coenzyme-A (COA) to an acyl carrier protein (ACP) by malonyl-CoAzACP
transacylase. From this point on, all the reactions of the pathway involve ACP (Figure 2).

After being transferred to ACP, the malonyl group enters a series of reactions that result

  
    

9° co

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acetyl-COA ' Carboxylase

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Trans-Z-butenoyl-ACP
n,o
Dehydration

Figure 2. Simplified representation of the fatty acid biosynthesis pathway in
Arabidopsis (according to Ohlrogge and Browse, 1995).

1, Acetyl-COA carboxylase; 2, Malonyl-CoAzACP transacylase; 3, 3-Ketoacyl-ACP
synthase (KAS); 4, 3-Ketoacyl-ACP reductase; 5, 3-Hydroxyacyl-ACP dehydratase; 6,
2,3-trans-enoyl-ACP reductase.

l6

in the formation of a carbon-carbon bond and in the release of C02. These reactions are
catalyzed by plastidial fatty acid synthase, which in plants is also a multisubunit enzyme
composed of 3-ketoacyl-ACP synthases I, II and III, 3-ketoacyl-ACP reductase, 3-
ketoacyl-ACP dehydrase, and enoyl-ACP reductase (Figure 2) (Somerville et al., 2000).
All of fatty acid synthase subunits are nuclear encoded. The FAS pathway produces
saturated fatty acids, however more than 75 % of plant fatty acids are unsaturated. The
first double bond is introduced by a soluble 18:0-ACP desaturase in the plastid.
Additional double bonds in fatty acids are incorporated by membrane-bound desaturases
that act once the fatty acid has been incorporated into glycerol (Browse and Somerville,

1991).

The elongation of fatty acids is terminated when the acyl group is removed from ACP.
This reaction can be catalyzed by acyl-ACP thioesterases that hydrolyze the acyl-ACPS to
release free fatty acids and ACPS or alternatively, the acyl group can be transferred to
g1ycerol-3-phosphate or monoacylglycerol-3-phosphate by the action of acyl-ACP
acyltransferases (Figure 3) (Somerville et al., 2000). More than half of the fatty acids
produced in the plastids flow through acyl-ACP thioesterases and are exported from this
organelle in the form of acyl-COAS. These molecules are later used for glycerolipid
synthesis at the level of the endoplasmic reticulum (ER). Based on its Similarity to the
animal pathway for glycerolipid synthesis it is named Eukaryotic Pathway (Figure 4).
The fraction of fatty acids that remains in the plastid is used for glycerolipid synthesis
within the plastid and based on its similarity to the bacterial pathway it is named the

Prokaryotic Pathway (Figure 4). Some of the lipids synthesized by the Eukaryotic

l7

Figure 3. Schematic representation of the enzymatic reaction catalyzed by acyl-
ACP thioesterases in plants.

Acyl-ACP thioesterases or FAT (fatty acid thioesterases) enzymes catalyze the
hydrolysis of the thioester bond between acyl carrier proteins (ACP) and fatty acids to
release free ACPS and free fatty acids inside the plastids. Fatty acid are then exported
from the plastid as acyl-COA molecules.

18

 

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19

Figure 3

Figure 4. Simplified scheme of the glycerolipid biosynthetic pathway in
Arabidopsis (adapted from Browse and Somerville, 1991).

Fatty acid synthesis (FAS) takes place inside the plastid and its acyl-ACP products
(16:0-ACP, 18:0-ACP and 18:1-ACP) are substrates of either acyl-ACP thioesterases
(FAT) or acyl-ACP acyltranferases (ACT). By the action of the latter, acyl groups enter
the Prokaryotic Pathway of lipid synthesis inside the plastids to produce MGDG,
DGDG, PG and SL. By the action of acyl-ACP thioesterases, acyl groups leave the
plastid as acyl-COA molecules and become substrates of the Eukaryotic pathway of lipid
synthesis in the endoplasmic reticulum (ER). Fatty acids can return to the plastid in the
DAG moiety of PC and are used for the synthesis of MGDG, DGDG and SL within the
plastid. Abbreviations: PC, phosphatidylcholine; PE, phosphatidylethanolamine;
MGDG, monogalactosyldiacylglycerol; DGDG, digalactosyldiacylglycerol; PG,
phosphatidylglycerol; PI, phosphatidylinositol; PS, phosphatidylserine; SL, sulfolipid;
DAG, diacylglycerol; PA, phosphatidic acid; LPA, lysophosphatidic acid.

20

 

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Figure 4

21

Pathway traffic back to the plastid and intermix with the plastid lipid pool (Figure 4)

(Browse and Somerville, 1991).

The role of fatty acids in the synthesis of different cellular components

Membrane glycerolipids

Membrane glycerolipids have fatty acids attached to the sn-1 and sn-2 positions of the
glycerol backbone and a polar head group in the sn-3 position. The combination of non-
polar acyl chains with polar head groups leads to the amphipathic physical properties of
these molecules, which are essential for the formation of membrane bilayers (Nagle et al.,
2000). Fatty acids are carboxylic acids of highly reduced hydrocarbon chains that in
plants can range from 8 to 32 carbons in length. However, the major fraction of
membrane glycerolipids contain chains of 16 and 18 carbons. Shorter and longer fatty
acids are usually found in storage lipids or epicuticular waxes (Post-Beittenmiller, 1996).
Fatty acids in lipid membranes contain from one to three cis double bonds and five of
these molecules (18:1, 18:2, 18:3, 16:0 and 16:3) make up over 90 % of membrane
glycerolipid acyl chains. The number and position of double bonds in fatty acids have
profound effects on their biophysical properties. For example, incorporation of a cis
double bond between carbons 9 and 10 of an 18 carbon saturated fatty acids decreases its
melting temperature from 70° C to 14° C. The addition of a second cis double bond

between carbons 12 and 13 reduces its melting point even further, from 14° C to —5° C.

22

The structure of the head group that is incorporated in the sn-3 position of glycerolipids
defines seven different classes of these molecules (e.g., phosphatidylcholine (PC),
phosphatidylethanolamine (PE), phosphatidylinositol (PI)). In addition, each class is
composed of distinct molecular species defined by the fatty acids attached to them. For
example, PC carries either a 16 carbon saturated fatty acid in the sn-1 position and an 18
carbon unsaturated in the sn-2 or unsaturated fatty acids in both the sn—1 and sn-2
positions. The fatty acid composition of individual glycerolipids is critical to confer the
required biophysical properties to cell membranes (Mc Elhaney, 1989). Those properties
are essential for fluidity, permeability, vesiculation, transport and protein activity (e.g.,
photosynthesis). Moreover, crystallization of intrinsic membrane proteins revealed that
specific lipids co-crystallize with them and are essential parts of their structures and

functions (Katona et al., 2003; Murtazina et al., 2002).

One central issue with respect to the role of glycerolipids in membranes is framed by the
observation that each membrane of the cell has a characteristic and distinct complement
of glycerolipids and that, within a Single membrane, each class of glycerolipid has a
distinct fatty acid composition (Somerville et al., 2000). Furthermore, the composition of
each membrane and lipid class is largely conserved throughout the plant kingdom. This
conservation implies that differences in lipid structure are important for membrane
function. However, the details of this structure/ function relation have remained elusive, in
part because plants can adapt to Significant changes in lipid composition and the effects
of those changes are not always obvious. For example, highly unsaturated 18:3 and 16:3

fatty acids constitute approximately 70 % of thylakoid membrane fatty acids.

23

Surprisingly, an Arabidopsis triple mutant fad3fad 7fad8 which completely lacks 18:3 and
16:3 exhibits normal rates of vegetative growth and photosynthesis at 22° C. These
results demonstrate that these polyunsaturated fatty acids are not essential for
photosynthesis in plants grown at standard conditions. The loss of 18:3 and 16:3 have
noticeable effects only at low (below 10°C) and high (above 30°C) temperatures (Wallis

and Browse, 2002).

Other fatty acid-derived products

In addition to their role in membrane glycerolipid biosynthesis, fatty acids are used for
several other biosynthetic pathways in plant cells. At the level of the ER, synthesis of
Sphingoid bases and ceramides consume two molecules of fatty acids. Ceramides are later
glycosylated at the level of the ER and Golgi apparatus to form the essential lipids called
sphingolipids. These lipids are found in the outer leaflet of the plasma membrane and
have critical functions in vesiculation, Golgi trafficking and Signaling (Sperling and
Heinz, 2003). Similarly to glycerolipids, a large number of Sphingolipid species exist that
differ in the number of double bonds and hydroxyl groups in the Sphingoid base, type of
acyl group in the ceramide moiety and type and number of sugars in the head group
(Sperling and Heinz, 2003). Most of the knowledge about Sphingolipid function in plants
has been inferred from information in animals and yeast and little direct experimental

data exist on their role in plants.

24

Fatty acids are also used in epidermal cells for the synthesis of epicuticular waxes and
cutin. Waxes are derived only from saturated fatty acids that are elongated at the level of
the ER to chains up to 32 carbons and later modified to alkanes, aldehydes, alcohols and
ketones (Post-Beittenmiller, 1996). Waxes are deposited outside cells and create an
hydrophobic barrier that protects plants from water loss and environmental stresses.
Cutins are derived from saturated and unsaturated fatty acids that become
polyhydroxylated and crosslinked to form a mesh that confers mechanical resistance to
the outer surface of plants (Kolattukudy, 2001). In seeds, fatty acids are precursors of
triacylglycerol (TAG) biosynthesis, the major sink of carbon in oilseeds. In addition, fatty
acids are the substrates for all sort of acylation reactions (e.g., proteins, sterols, etc) and

also precursors of signaling molecules (Farmer et al., 2003).

Regulation of fatty acid biosynthesis in plants

Regulation by light

The carboxylation of acetyl-COA to produce malonyl-COA by acetyl-COA carboxylase
(ACCase) is the first committed step of fatty acid synthesis and this enzyme is considered
most responsible for the primary regulation of this pathway (Ohlrogge and Jaworski,
1997). In animals and other non-plant systems, the activity of ACCase has been
determined to represent a primary regulatory step (Wakil et al., 1983; Goodridge, 1972).
Fatty acid synthesis is lO-fold or higher in illuminated leaves than non-illuminated ones
(Bao et al., 2000; Browse et al., 1981). Some of the light-dependence of this pathway can

be attributed to changes in the stroma] pH, Mg2+, ATP/ADP ratio and reductants (Hunter

25

and Ohlrogge, 1998; Sasaki et al., 1997; Eastwell and Stumpf, 1983). The in vitro
activity of ACCase is sensitive to all these conditions and in addition the formation of
malonyl-COA is light dependent. Thus, evidence suggests that light affects ACCase
activity and subsequently the rate of fatty acid production (Post-Beitenmiller et al.,
1991). However, the mechanism by which ACCase is activated by light is still under
debate. The B-CT subunit of ACCase is phosphorylated at a serine residue in the light and
evidence points to the B-CT of chloroplast ACCase as a candidate for regulation by

protein phosphorylation/dephosphorylation (Savage and Ohlrogge, 1999).

Regulation by feedback inhibition

Most biochemical pathways are controlled in part by feedback mechanisms in which
inhibition of enzyme activity occurs when the product of a pathway builds up to levels in
excess of need. In most cases this inhibition occurs at a regulatory enzyme which is often

the first committed step of the pathway.

In animals and yeast, it has been considered that fatty acid synthesis is partially
controlled by feedback on ACCase by long chain acyl-COAS (Wakil et al., 1983).
However, this model has been questioned because acyl-COA binding proteins are in
concentration high enough to sequester most acyl-COA molecules and therefore would be
present only at extremely low levels (Knudsen et al., 1994; Rasmussen et al., 1993).
Shintani and Ohlrogge (1995) demonstrated that fatty acid synthesis is inhibited by

feedback mechanisms in plant cells. However, potential feedback inhibitors such as acyl-

26

COA, free fatty acids and glycerolipids failed to strongly inhibit plastidial ACCase at
physiological concentrations in vitro. Because fatty acid synthesis takes place inside the
plastid but the major utilization of its products is at the ER membranes, it is likely that
feedback regulation must allow communication across the plastid envelope. At this time
it is not known what molecules are involved in feedback regulation of plastidial fatty acid

synthesis.

Regulation by demand

Evidence that utilization of fatty acids can increase rates of fatty acid biosynthesis comes
from different experiments. First, when a plant 12:0-ACP thioesterase (MCTE) is over-
expressed in E. coli cells, fatty acid synthesis is increased and cultures accumulate at least
lO-fold more total fatty acids than control cultures. In these experiments, the removal of
the products of fatty acid synthesis (acyl-ACPS) to free fatty acids by MCTE, appeared to
release feedback inhibition on ACCase, resulting in higher malonyl-COA and fatty acid
production (Ohlrogge et al., 1995). Second, over-expression of cloned membrane
proteins results in an increased production of fatty acids for membranes to accommodate
the excess proteins and maintain lipid/protein ratios constant (Nieboer et al., 1996).
Therefore, the rate of fatty acid synthesis can apparently be regulated by the demand for
fatty acids needed for membrane synthesis. Finally, analysis of developing Brassica
napus seeds expressing high levels of the California Bay 12:0-ACP thioesterase (MCTE)
shows that expression of fatty acid synthesis and B-Oxidation enzymes were induced.

These seeds appear to produce more 12:0 that can be metabolized to TAG and the excess

27

12:0 is degraded. To compensate for the loss of carbon by B-oxidation, cells increase
fatty acid production. Moreover, protein and activity levels of ACP, ACCase, 18:0-ACP
desaturase, and KAS HI increase 2 to 3-fold, indicating that concerted induction of these

enzymes has occurred (Eccleston and Ohlrogge, 1998).

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32

CHAPTER 2

DIFFERENTIAL REGULATION OF MESSENGER RNA LEVELS OF ACP
ISOFORMS IN ARABIDOPSIS
ABSTRACT
All higher plants express several different acyl carrier protein (ACP) isoforms in a tissue
specific manner. We provide evidence that expression of mRNA for the most abundant
ACP isoforrn in Arabidopsis leaves (ACP4) is increased several fold by light whereas
mRNA levels for ACP isoforms 2 and 3 are independent of light. The presence of
GATA-like motifs in the upstream region of the Acll.4 gene (encoding ACP4) and the
similarity in light-mediated induction to ferredoxin-A mRNA suggests a direct role of
light in Acll.4 gene activation. Polyribosomal analysis indicated that light also affects the
association of ACP transcripts with polysomes, similarly to mRNAs encoding ferredoxin-
A. ACP2, ACP3 and ACP4 mRNA levels were also examined in Arabidopsis cell-
suspension culture and found to be differentially controlled by metabolic and/or growth
derived signals. Moreover, Similar mechanisms operate in planta for ACP2 gene
expression. Fusion of the ACP2 promoter to luciferase demonstrated that regulation of
ACP2 mRNA levels was transcriptional and responded to growth/cell-cycle signals.
Arabidopsis ACP2 promoter deletion analysis in tobacco cells revealed conserved
mechanisms of ACP gene expression between plant Species and identified promoter
regions important for transcriptional regulation. Comparison of Arabidopsis ACP
promoters disclosed the presence of 2 conserved elements, an 11 bp element found in the
five Arabidopsis ACP genes and a 6 bp element found only in ACP2 and 3. Comparison

of proximal ACP 5’ upstream sequences between diverse plant species showed that two

33

motifs have been conserved during evolution, a CTCCGCC box and C-T rich sequences.
In Arabidopsis these regions are transcribed and are present in the 5’ untranslated regions
(UTRS) of ACP mRNAs. We tested the role of ACPl and ACP2 mRNA 5’-leader regions
in transgenic Arabidopsis plants and demonstrated that they enhanced several-fold the
expression of a reporter gene in a tissue Specific manner. By using large scale mRNA
profiling techniques we demonstrated that fatty acid and lipid synthesis genes did not

respond to inhibition of fatty acid synthesis in Arabidopsis cells.

INTRODUCTION

Despite our current understanding of the biochemistry of fatty acid and lipid synthesis in
plants, the signals and factors that direct the expression of genes in these pathways
remain largely unknown. Unlike in other organisms, the enzymes responsible for de novo
fatty acid synthesis in plants are not localized in the cytosol but rather in the plastid.
Although a portion of the newly synthesized acyl chains is utilized for lipid synthesis
within the plastid, a majority is exported into the cytosol for glycerolipid assembly at the
endoplasmic reticulum (ER) (Somerville et al., 2000). In addition, some extraplastidial
glycerolipids return to the plastid and extensive lipid interchange exists between these
two organelles (Ohlrogge and Jaworski, 1997). In order to adjust to changes in the
demand of lipids during tissue development and environmental influences, plant cells
must regulate and coordinate the expression of the many genes involved in fatty acid and
lipid synthesis pathways. An attractive possibility is the existence of a global

transcriptional control for these genes, perhaps comparable to the yeast system (Schuller

34

et al., 1992). For example, it has been shown that the expression of the different subunits
of acetyl-COA carboxylase (ACCase) is orchestrated during tissue development (Ke et
al., 2000). This enzyme catalyzes the first committed step in fatty acid synthesis and
available evidence points to its key role as a major regulatory enzyme in fatty acid
production (Ohlrogge and Jaworski, 1997). In addition, global post-transcriptional
mechanisms have also been suggested for regulation of fatty acid synthesis genes in

plants (Eccleston et al., 1998).

Analyses of gene expression under conditions where the demand for fatty acids is
elevated may unravel part of the signals and mechanisms that participate in the regulation
of genes involved in plant lipid biosynthesis. A Situation where the cells are fully
engaged in fatty acid production is during leaf expansion in illuminated plants (Browse et
al., 1981). In addition to cell division and growth, light induces the differentiation of the
proplastid to the specialized, membrane-rich chloroplast (Kasemir, 1979). This transition
involves not only a radical change in the structure and function of the plastid but also its
increase in Size and number per cell. Therefore, de novo synthesis of glycerolipids is
critical to provide chloroplasts with new lamellae to accommodate the photosynthetic
apparatus. During the process of leaf expansion light activates transcription and
translation of a large number of genes that participate in chloroplast differentiation
(Chory et al., 1994). Moreover, the interdependent relation between chloroplast
development and the activation of nuclear genes has been confirmed by mutant analyses
in Arabidopsis (Susek et al., 1993). As a consequence of the plastidic location of the fatty

acid synthesis machinery and its commitment to supplying lipids for new membrane

35

synthesis, it seems possible that at least some of the genes participating in this pathway
belong to the group of genes activated by light during leaf expansion. However, previous
studies argue against this hypothesis and suggest a minor role of light on the regulation of
the genes for fatty acid synthesis (Scherer et al., 1987; Battey et al., 1990; Baerson et al.,
1993). Presently, the only gene known to participate in glycerolipid production and to be
transcriptionally induced by light in Arabidopsis iS the chloroplast (0-3 fatty acid

desaturase (FAD7) (Nishiuchi et al., 1995).

A second condition where the fatty acid synthesis machinery is very active is during the
exponential growth of plant cells in culture. When plant cells are grown in liquid culture
in the presence of a carbon source, they Show rapid incorporation of labeled precursors
into phospholipids and neutral lipids (Weber et al., 1992). The presence of an energy
source such as carbohydrates in the media increases the cell growth rate and hence the
need for new membranes. Sugars not only function as substrates for growth but also
affect sugar-sensing systems that initiate changes in gene expression. In plant
metabolism, carbohydrate depletion generally enhances the expression of genes involved
in photosynthesis, reserve breakdown and export, whereas abundant carbon resources
favor genes for storage and utilization (Koch, 1996). For example, Graham et al (1994)
Showed that in cucumber, the mRNA levels for two enzymes of the glyoxylate cycle
(malate-synthase and isocitrate-lyase) are coordinately induced not only during post-
genninative seedling development but also by a metabolic signal derived from sugars.
The glyoxylate cycle plays an important role in the degradation of fatty acids during

seedling germination. In animal pancreatic cells a glucose-mediated signal activates the

36

transcription of the acetyl-COA carboxylase gene (Brunt et al., 1993). Thus, due to the
metabolic regulation via sugars of several genes involved in primary biochemical
pathways in plants, it is likely that at least some of the genes involved in fatty acid

synthesis are also under the control of Similar mechanisms.

Finally, in bacteria inhibition of fatty acid biosynthesis by drugs triggers the
transcriptional activation of genes involved in fatty acid and lipid metabolism (Schujman
et al., 2003). Thus, bacteria have the ability to sense a decrease in the activity of the FAS

pathway and to respond by adjusting the expression of F AS genes.

The regulation of acyl cam'er proteins (ACPS) has been the focus of several studies based
on their central role in lipid biosynthesis in plants (Hannapel et al., 1988; Hlousek-
Radojcic et al., 1992, Baerson et al, 1993). ACPS are small acidic proteins that carry the
nascent acyl chains during the synthesis of 16 and 18 carbon acyl groups. In this report
we demonstrate that the transcripts for several isoforms of Arabidopsis ACPS are subject
to differential regulation by light and also by metabolic and/or growth control. In
addition, we provide evidence for the role of the ACPl and ACP2 5’ untranslated
sequences (UTRS) in the enhancement and tissue specific expression of a reporter gene in

Arabidopsis plants.

RESULTS

ACP4 mRNA Levels are Increased by Light

37

The participation of light in the activation of fatty acid synthesis in young leaves is well
established (e. g. Browse et al., 1981; Bao et al., 2000). Light regulation can be attributed
in part to activation of ACCase by mechanisms that involve reduced thioredoxin,
phosphorylation, substrate activation, cofactors and pH changes (Sasaki et al., 1997;
Hunter and Ohlrogge, 1998; Savage et al., 1999). Although light-mediated mechanisms
of gene activation such as transcription are prominent during chloroplast biogenesis in
leaves, no available evidence connects these mechanisms to the activation of genes
responsible for de novo fatty acid synthesis. Therefore, we first examined the influence of
light on the mRNA steady state levels of distinct isoforms of Arabidopsis acyl carrier

proteins.

Based on protein analyses, ACPl, ACP2 and ACP3 have been identified in all tissues
examined (Hlousek-Radojcic et al., 1992) whereas a fourth isoforrn (ACP4) is expressed
predominantly in leaves (Shintani, 1996). As shown in Figure 5, the mRNA levels of
ACP4 increased approximately 4-fold in leaf tissue when two week-old Arabidopsis
seedlings were reilluminated for 4 hours after a 24 hours dark period. An even higher
increase (S-fold) was observed at eight hours of illumination with a decline to 4-fold
again after 12 hours (Figure 5). Interestingly, the ACP4 messenger RNA presented
similar kinetics of light-mediated induction to the Arabidopsis ferredoxin-A (FEDA)
mRNA, a 4-fold increase after 4 hours in white light with a subsequent decline after 12
hours (Figure 5). Previous studies demonstrated that the gene for FEDA is

transcriptionally activated by light, and part of this induction is initiated by phytochromes

38

A) B)

Time (hours)

 

0124812

Cl)
0
E 6 -
ACP2+3 , I.” w v '2 5 l
3 .
. ~8 ACP4
~ Il‘tx‘!’ .w. A: fins.” 4 ..
ACP4 *5 In . . . <1
, g 3 a FEDA
FEDA ... an .QW. «>2 2 .
‘3 1 ACP2+3
<1) i

0124812

time (hours)

Figure 5. Light-mediated induction of ACP4 mRNA levels in Arabidopsis leaf
tissue.

(A) Northern blot analyses of two week-old Arabidopsis plants dark treated for 24 hours
and reilluminated with fluorescent white light for the times indicated. Each lane of the
blot contains 5 ug of total RNA from leaf tissue. The eIF 4A probe was used as a loading
control. (B) Densitometry scanning of the blots in A. The signal intensities at each time
were normalized with the corresponding eIF4A Signal and expressed relative to time 0
(set arbitrarily to one).

(Caspar et al., 1993).

In contrast to ACP4 transcript levels, the signal presented by a probe corresponding to the
ACP2 coding sequence did not Show intensity variations in the conditions tested (Figure
5). This result agrees with a previous study that Showed that the Arabidopsis Acl].2 gene

promoter (ACP2) does not confer light responsiveness to a reporter gene in transgenic

39

tobacco seedlings (Baerson et al., 1993). Because the ACP2 and ACP3 isoforms are 82%
Similar at the nucleotide level, we assumed that the signal conferred by the ACP2
fragment represented the abundance of both, ACP2 and ACP3 transcripts (ACP2+3 in
Figure 5). This assumption is also based on the observation that both proteins are
expressed in similar amounts in Arabidopsis leaf tissue (Hlousek-Radojcic et al., 1992)
and that the corresponding promoters confer similar levels of expression to a reporter

gene in Arabidopsis leaves (Baerson et al., 1998).

Polyribosomal Association of ACP mRNAs is Increased by Light

Previous studies suggested that the expression of ACPS could be regulated at the level of
translation. First, transgenic Brassica napus plants over-expressing a 12:0-ACP
thioesterase Show an increase of approximately 2-fold in the ACP protein level with no
Significant changes in the corresponding messenger abundance (Eccleston et al., 1998).
Second, Hannapel et al (1988) reported that during seed development in soybean, the
relative abundance of ACP and lectin mRNAs is at most 28-fold different whereas the
corresponding protein levels differ at least ZOO-fold, suggesting a differential translational

efficiency between these two mRNAs.

Based on these observations, we compared the polyribosomal distribution of ACP
mRNAs with other transcripts highly expressed in leaf tissue of Arabidopsis plants. In
addition, the fact that Ii ght has a major effect on the polyribosomal association of several

messengers RNA encoding plastidic proteins (Berry et al., 1990; Dickey et al., 1998)

40

prompted us to investigate the influence of light on the association of ACP mRNAs with
ribosomes. Therefore, we analyzed Arabidopsis leaf tissue from plants either dark treated

for 24 hours or reilluminated for 12 hours with

 

 

 

 

Light Dark
0/o suc .fi-x-J‘f’m /’//’—’l
ACP2+3 a I 9 u I I -
FEDA ' jggfifi§$§“ .5 %_I_ i
“W e. H j t?!!""””

Figure 6. Light affects the polyribosome association of ACP mRNAs.

Leaf tissue extracts from plants grown in the dark for 24 h (Dark) or reilluminated for 12
h after a 12 h dark period (Light) were Spun in 15 to 60 % sucrose gradients. Sixty
aliquots were collected at identical volume intervals and absorbance at 260 nm measured.
Total RNA was extracted from 15 fractions of the gradients, loaded onto an agarose gel
and analyzed by Northern blot. The blots were hybridized with probes corresponding to
the sequences indicated. Fractions 7 to 15 correspond to the polyribosomal fractions
(black line).

white fluorescent light. The results demonstrated that the messengers for ACP2+3 and
ACP4 were associated with polyribosomes (fractions 10 to 15 of the gradient) after 12
hours of illumination (Figure 6). In contrast, after 24 hours in the dark, the transcripts for
the ACP isoforms appeared in the upper fractions (3 to 6) of the gradient, corresponding
to low molecular weight polyribosomes or ribosome-free mRNAs (Figure 6). The
polyribosomal distribution of the ferredoxin A (FEDA) mRNA was Similar to that of

ACP mRNAs in both conditions (Figure 6). The eukaryotic initiation factor 4A (eIF4A)

41

transcript also showed association with polyribosomes in the light, but in contrast to ACP
and FEDA mRNAs, a significant proportion of the eIF4A transcript was still bound to
polyribosomes after 24 hours in the dark (Figure 6). In summary, these observations
indicate that the transcripts encoding for ACP isoforms are associated with
polyribosomes in Arabidopsis leaf tissue in a light-dependent manner similar to FEDA

but differ Si gnificantly with respect to eIF4A mRNAs in the dark.

Polyribosomal association of ACP mRNAs in developing seeds of wild type B.napus
and transgenics over-expressing MCTE

To evaluate if translation of ACP mRNAs was altered in developing seeds of transgenic
Brassica napus over-expressing a medium-chain thioesterase (MCTE) compared to wild
type, the polyribosomal distribution of ACP mRNAs was analyzed in seeds of both plant
classes. Flowers from wild type plants and transgenic B. napus (event 198, Voelker et al.,
1992) were tagged and developing seeds harvested at 21, 28 and 35 days after flowering.
These time points correspond to mid-stage development of B.napus seeds, the period of

active Oil biosynthesis and accumulation.

The distribution profile of ACP transcripts on polysomes was Similar between B. napus
wild type and transgenic seeds during the three developmental stages analyzed (Figure 7).
These results suggested that translation of ACP mRNAs in seeds was not affected by
over-expression of MCTE and mechanisms affecting protein stability were more likely

explanations for higher FAS protein levels in transgenic seeds (Eccleston and Ohlrogge,

42

1998). In addition, association of ACP transcripts with high molecular weight polysomes

indicated that ACP mRNAs were efficiently translated in seeds.

 

 

 

 

 

 

MCTE Wild type
Polysomes Polysomes
o/o suc A///l

 

 

28DAF ,_ abun-
21w — Full.“ noon-a.

Figure 7. Polysome distribution of ACP mRNAs in wild type and transgenic
(MCTE) B. napus developing seeds.

Developing seeds from wild type B. napus and transgenic overexpressing MCTE were
harvested at 21, 28 and 31 days after flowering (DAF). Whole seeds were lised and
extracts loaded onto 15-60 % sucrose gradients. Fractions containing polysomes were
collected from the gradients after centrifugation and total RNA extracted and resolved by
electrophoresis. After transfer, the blots were probed with a seed Specific ACP cDNA
(Bn-ACP 28f10).

Conserved Motifs occur in the 5’ Leader Region of ACP mRNAs

The examination of Arabidopsis genomic and cDNA ACP sequences has revealed two
unusual motifs in the 5’ leader region of ACP mRNAs (Table 2 and Ohlrogge et al.,
1991). One feature is the presence of short sequences rich in pyrimidines. The high

cytosine and thymidine content is uncommon, and leader sequences of plant mRNAs tend

43

to be rich in adenines and thymidines (Joshi, 1987). A second conserved motif is an
element composed of seven nucleotides CTCCGCC (Table 2). In addition to Arabidopsis
sequences, these features are also conserved in the 5’ leader region of ACP messengers
from several diverse species (Table 2). The importance of 5’untranslated regions (UTRS)
as essential regulators of gene expression in plants has been described (Dickey et al.,

1998; Bolle et al., 1996).

To analyze the participation of the 5’ leader sequences of the ACP mRNAs in the
regulation of ACP expression, we generated independent Arabidopsis transgenic lines in
which the 5’ UTR sequences of the Arabidopsis ACPl (lines atL-acpl) and ACP2 (lines
atL—acp2) mRNAs were fused to the luciferase reporter gene (LUC) under the control of
the CaMV35S promoter (Figure 8). To preserve the endogenous translation initiation site
a short portion of the corresponding ACP coding sequence was also included (see Figure
8 for details). The 5’ leader regions of ACPl and ACP2 mRNAs present sequence
variations but both have conserved CU rich regions and include the heptanucleotide motif
CTCCGCC (Table 2). However, ACPI and ACP2 transcripts contain distinct translation
initiation sites (Figure 8). The AUG context in the Acl].1 gene (ACPl) matches with one
of the most common translation initiation sequences found in plants, AAACAAU_GGC
whereas the translation initiation site in the Ac11.2 gene (ACP2), CUUCUAUQGC is
found in a smaller number of plant genes (Joshi et al., 1997). This suggests that the
efficiency of AUG recognition by the translation machinery might also influence ACP
expression. A third line of transgenic plants carrying the LUC gene fused to a deleted

version of the ACPl leader region (line atL-dell) was also generated. In this case, most

44

Table 2. Proximal upstream sequences of ACP genes in different plant species.

 

A. thaliana ACPl: pt_c_tt_tgtacaCTCCGCCctctctccccatctctttcgacagatctcttctctctctc gtgtttcacgaaaca atg
Athaliana ACP2 : mmCTCCGCCmatgtgaflgatgtggacgattcam atg

Athaliana ACP3: actgtttctcatctcttcgt_chTCCGCCMQaamachgagctcitacgattcattcgttct atg

Athaliana ACP4 : ccgaagataggcctgaatctccgagaacaaacCTCCGCCacaaaaacagaagacttgttgcttgcgtatcataat
cgacgccgtatctctacttgctcgaacaaacccaaaaagatacatatcaagagattaaaccttatccaactaagagaagccatttttattttttttgggtctctgag

ttgtgtattgagcttcatctccttcaa atg

 

Athaliana ACPS: acaaaatagtaattcacggtc_cttgaacaATCCGCCaMgatcagatcgata atg

B. campestris ACP-SQ: atcacgfltfigtacaCTCCGCCatctctctctctctcgagcagatctctctcgggaatatcgaca atg

B. campestris ACP-sfl: caagctaccatgggacatcachtacaCTCCGCCatctctctccathtctctcgtgaataacgaaa atg
B. napus ACP: atcacggmgtacaCTCCGCCatctctctctccttcgagcacagatctctctcgtgaatatcgaca atg

B. napus ACP-28f10: acgctctgtacaCTCCGCCatctctctccattctctctcgtgagtaacgaca atg

B. rapa ACP: gacatcacgflgtacaCTCCGCCatctctctctctctcgagcagatctctctcgggaatatcgaca atg

C. glauca ACP: gcggccgctngTCCGTthatttcmgmccctctctcaaaflagatctctctctctcgctttgtatct atg
Csativum ACP: tatcgtcacactctttgtgctCTCCGCCgtgtmgatcaataaacttttctcagatctaaactctatctatcaa atg

 

C. lanceolata AC Pl-3: cCTCCGTCgtflaMagctaccaa atg

C. lanceolata ACP l -l: cacggc_tCTCTCGCthatttgctcgctccctccctccctcccccatca atg

Spinach ACPII: atctctctcctctttctctCTCCGCCacattcatt_ctga%actttctctcccctctctctccgcttcttca atg
Spinach ACPI : aacttaataflactcaggataagcttctcactctctctctctctctctcttactacc atg

Barley ACP2: ccgccgcCTCCACCgccgcCTCCACCgcccaccgcgccggcctctccccctgtcccgtctccccc atg
Barley ACPl: aggaccagccggcctcttcccaccgcccccaaattctaccgagcagCTCAGCngccaaccc atg

 

A sequence of arbitrary length upstream the of translation initiation site was selected. The sequence
(CTCCGCC) and its derivatives are indicated in uppercase. The OT rich sequences are underlined. The
ATG start codon is separated by one space at the right end of the sequences. A.= Arabidopsis, B.=

Brassica, C. = Coriandrum

45

A) \ I ‘I

(L) NPTH +1 \\\\\\\\1 GUS (R)
m} Bl

B) ///’ “4L-

pCEtACPI IacctcIzaactctttgtacactccgccctctctccccatctctttcgacagatctcttctctctctcgtgtttcacgaaacaatggcgactcaattcagcgccatg

 

pCaAC P22 acctcgagactgtttctctatctctttgtcttctccgcctcctccgatctcactccgatctctctacgattcattcttctatggcttccattgccatg

pCadell I acctcgagctctttgtacaaacaatggcgactcaattcagcgccatg

Figure 8. Constructs used for Arabidopsis transformation.

(A) Scheme of the T-DNA constructs carrying the LUC and GUS genes. Black box: T-
DNA borders (L) left and (R) right ends, Open box: nopaline synthase 3’ polyadenylation
Signal, NPTII: neomycin phosphotransferase coding sequence, black triangle: CaMV35S
promoter, LUC: firefly luciferase coding sequence, GUS: B-glucuronidase coding
sequence, B: BamHI, P: PstI, Bl: BglII, +1 denotes the transcription initiation Site. (B)
Sequences of the 5’ UTRS derived from Arabidopsis ACPl (pCaACPl), ACP2
(pCaACP2) and the deleted version of ACP] (pCadell) used in the three independent
constructs. The ACP translation initiation codon is underlined. The displayed sequences
start at the predicted transcription initiation site and therefore the leaders generated by
these constructs have 8 additional bases from the vector sequence (underlined).

of the 5’ UTR was removed, conserving only the first nucleotides towards the 5’ end and
the ACPI translation initiation Site (see Figure 8 for sequence details). As an internal
control for position effect and copy number, all the T-DNA constructs also carried a
cassette expressing the B-glucuronidase (GUS) gene under the CaMV35S promoter

(Figure 8).

The 5’ Leader Sequences of ACP] and ACP2 Increase Reporter Gene Expression

46

We analyzed the LUC and GUS specific activities in expanding leaves of two week-old
transgenic plants. The atL-acpl and atL-acp2 transgenic lines Showed 10- and 20-fold
higher LUC/GUS ratios respectively compared to the atL-dell lines after 24 hours in the
dark (Figure 9A). From these results we conclude that the presence of the ACP 5’ UTRS

is essential for high expression of ACP] and ACP2 in Arabidopsis leaves.

Based on the polyribosomal distribution of the ACP transcripts (Figure 6) we also asked
whether the ACP 5’ UTRS could have a light regulatory role similar to other UTRS from
plastid proteins (Dickey et al., 1998; Bolle et al., 1994). After 6 hours of re-illumination,
the activity ratio of the reporter genes in the atL-acpl and atL-acp2 lines differed
approximately 20- and 30-fold with respect to atL-dell lines (Figure 9A). The additional
increase upon reillumination in the LUC/GUS ratios suggests that light-enhanced

expression of the ACP-LUC constructs is at least partly mediated by the ACP 5’UTR.

Arabidopsis developing seeds are green and posses photosynthetic capacity. However,
they are considered heterotrophic as imported sucrose is their major carbon source. As
shown in Figure 9B, the LUC/GUS ratios of specific activities in mid-stage developing
seeds differed from the ratios observed in leaves. The activity ratio in atL-acpl transgenic
lines was approximately 2.5-fold higher than the ratio in atL-acp2 plants and l3-fold
higher than the atL-dell transgenic lines (Figure 9B). Thus, in contrast to leaf tissue, the

5’ UTR of ACP] mRNA appeared to confer a preferential expression of the LUC gene in

47

A) Leaves B) Seeds C) Roots

 

 

LUC/GUS Ratio

 

 

 

 

I atL-dell Cl atL-acpl I atL-acp2

Figure 9. Reporter gene expression in different tissues of Arabidopsis transgenic
plants.

atL-dell. atL-acpl and atL-acp2 indicate the transgenic lines transformed with pCadell,
pCaAC P1 and pCaAC P2 respectively (see Figure 8). (A) Transgenic plants were dark
treated for 24 h (Dark) or reilluminated for 6h (Light) with white fluorescent light. LUC
and GUS specific activities were measured in leaves of 10 independent transgenic lines
for atL-dell, atL-acpl and atL-acp2. LUC/GUS ratios are the average of the 10
individual ratios from each transgenic line. The atL-acpl and atL-acp2 LUC/GUS ratios
are expressed with respect to atL-dell LUC/GUS ratio (set arbitrary to one). The bars
denote the standard deviation of the average. (B) LUC and GUS Specific activities were
measured in roots Of 10 independent lines for atL-dell, atL-acpl and atL-acp2.
LUC/GUS ratios were calculated and represented as in A. (C) Reporter gene activity was
measured in pools of developing seeds from 10 independent lines for atL-dell, atL-acpl
and atL-acp2. The LUC/ GUS ratios were calculated and represented as in A.

developing seeds compared to the 5’UTR of ACP2 mRNA. It is noteworthy that the
level of reporter gene expression in atL-acpl lines relative to control lines was similar in

leaves and seeds (between 10- and 20-fold) (Figure 9A and B).

48

The expression of the reporter genes was also evaluated in root tissue from 2 week-old
transgenic plants. In contrast to leaf and seed tissue, the influence of the AC P1 and ACP2
5’UTRS was less pronounced and for atL-acpl plants the ratio of reporter gene activity
was Similar to the ratio in atL-dell plants (Figure 9C). This result suggests that the 5’
leader of the ACPI mRNA does not affect the expression of the LUC mRNA in roots.
For atL-acp2 plants, the LUC/GUS ratio was 4-fold higher than the ratio in atL-dell
plants (Figure 9C). Similar to leaf tissue, the influence on LUC expression of the ACP2
leader was higher than the effect produced by the ACPI leader on the same reporter gene.
However, the relative ratios of reporter gene activity conferred by the ACP2 leader was

between 7 and lO-fold lower in roots compared to leaves (Figure 9A and C).

ACP mRNA Levels are Affected by a Sucrose-derived Signal and/or Growth
Control in Cell Suspension Cultures

In bacteria and yeast the expression of several genes involved in fatty acid and lipid
synthesis is tightly coupled to growth (Jiang et al., 1994; Carman et al., 1999). Cells
growing in the presence of a carbon source such as sucrose Show high rates of
transcription of these genes (Carman et al., 1999). In addition to their role as energy
sources, sugars have been demonstrated to control the expression of plant genes involved
in diverse processes such as starch metabolism (Nakamura et a1, 1991), storage protein
accumulation (Hattori et a1, 1990) and lipid degradation (Graham et al, 1994). Based on
these observations, we asked whether the expression of some of the fatty acid synthesis
genes in plants might also be under metabolic and/or growth control by sugars. For this

purpose we analyzed the mRNA levels of ACP2, ACP3, ACP4 and the endoplasmic

49

reticulum (ER) associated delta-12-desaturase (F AD2) in Arabidopsis mesophyll-derived
cell suspension cultures. In order to distinguish between the messengers corresponding to
ACP2 and ACP3 in this experiment, we synthesized a messenger specific probe based on
the 3’ UTR sequences of these genes. The transcript levels of ACP2 and ACP3 isoforms
declined approximately 2-fold after 48 hours of starvation (Figure 10). In contrast, the
ACP4 and FAD2 transcript levels showed no significant variation during this 48 hours
period (Figure 10). Thus, the absence of sucrose in the media and/or the reduced growth
rate preferentially affected the expression of the ACP2 and ACP3 mRNAs. In the same
experiment the transcript levels of the ribulose-1,5- biphosphate-carboxylase 1A small
subunit (RBCSlA) increased approximately 2-fold after 12 hours of starvation and
slightly declined afterwards (Figure 10). This result was in agreement with the fact that
the gene for the small subunit of Rubisco is activated by light and repressed by sugars
(Terzaghi et al., 1995). To evaluate the effect of addition of sucrose to starved cells in the
absence of light, the cells were grown in the dark and in the presence of 5 8 mM sucrose.
The transcript levels of ACP2 and ACP3 increased approximately 2.5-fold after 24 hours
(Figure 11). In contrast, if the cells were kept in an osmotic control media in the dark the
level of the same messenger RNA remained steady (Figure 11). Interestingly, we
observed that the incubation of starved cells in the dark and 58 mM sucrose had a
negative effect on the ACP4 mRNA levels, decreasing its abundance more than 2.5-fold.
Conversely, no variations in the levels of the same mRNA were observed With. the
osmotic control media in the dark (Figure 11). Thus, both the absence of light and the

presence of 58 mM sucrose were responsible for the down-regulation of the ACP4

50

A) B)

Time (hours)

 

06122448

3
i C‘.
. ‘2
ACP3 1" ‘2’ ‘ ‘ 3 2
,. . I j; , RBCSIA
ACP4 “pull. 5 1
. _ ,. E
, _~. . 01, A ACP4
FAD2 35* BF 3" ‘1‘ I“ is l \
j ~ .. g FAD2
RBCSIA Q.... “0 ACP2and3

eIF4A] a q as M n
4 Time (hours)

Figure 10. Differential regulation of ACP mRNA levels in starved Arabidopsis cells.

(A) Cells were starved for 48 hours in the presence of light and total RNA was extracted
at the times indicated. Each lane of the Northern blots contains 2.5 ug of total RNA. The
blots were hybridized with probes corresponding to the sequences indicated. The eIF4A
probe was used as a loading control. (B) The Signals in A were quantified by scanning
densitometry and the values are represented as in Figure 5.

transcript levels. The same mRNA profile was observed for RBCSIA and Similar
mechanisms of regulation might be operating for both genes (Figure 11). In the case of
FAD2 mRNAs, no differences in the relative levels of the corresponding messenger were
found in the conditions tested (Figure 10 and 11). Thus, in contrast to ACP, neither

starvation nor light altered FAD2 mRNA abundance in liquid cell culture.

To investigate whether sugars could have a direct role in the regulation of ACP transcript

levels, we examined if ACP expression could be altered by uncoupling growth from the

51

presence of sugars in the media. For this purpose, cells were starved in an osmotic control
media (58 mM mannitol) for 48 hours and subsequently transferred into media containing
either 58 mM 3-O-methyl-glucopyranose (3-OMG) or 2 mM 2-deoxy-glucose (2-d-Glc).
The effect of these two glucose analogs on gene expression has been previously studied
in Arabidopsis cell-suspension cultures (Fujiki et al., 2000). The first glucose analog is
taken up by cells but not phosphorylated whereas the second is phosphorylated but not
further metabolized (Dixon et al., 1979). The presence of 3-OMG and 2-d-Glc in the
media neither showed a positive effect on ACP2+3 transcript levels nor did it decrease
the ACP4 mRNA level after 24 hours (data not shown). These results suggested that cell
growth and/or a metabolic Signal generated by sucrose or downstream of glucose were

necessary to control ACP mRNA levels in Arabidopsis cell liquid culture.

ACP2 mRNA Levels are Affected by a Sucrose-derived Signal and/or Growth
Control in planta

To evaluate if mechanisms that regulate ACP2 transcript levels in Arabidopsis cell
cultures also operate in intact plants, Arabidopsis seedlings were grown in liquid media in
the presence or absence of carbon. The results in Figure 12 indicated that ACP2 mRNA
levels responded to carbon signals in intact plants similarly to cell suspension cultures.
Therefore, the underlying mechanisms were most likely physiological mechanisms of
regulation of ACP2 expression in plants rather than mechanisms acting only in cell

suspension cultures.

52

A)

Dark/Sucrose (hours) Dark/mannitol (hours)
ACP2 " “CU - '-
ACP3 a. ,
ACP4 a ? 21' ‘ M M ii a M
FAD2 r m A .
RBCSIA . M u - a . “
fl ” I" ~ '6 ~
eIF4A 9' 1‘"
B) 8
g 3 AC P2 and 3
5 l
B
a i
E
g 1 FAD2
E ACP4
g 0. RBCSIA

0 3 6 12 24

Time (hours)

Figure 11. Differential regulation of ACP mRNA levels in Arabidopsis cells grown
in the presence of sucrose.

(A) Cells were starved for 48 hours in the light and subsequently transferred to either
sucrose containing media (dark/sucrose) or an osmotic control media (dark/mannitol) and
incubated in the dark. Total RNA was extracted at the times indicated. Each lane of the
Northern blots contains 2.5 ug of total RNA. The blots were hybridized with probes
corresponding to the sequences indicated and the eIF4A probe was used as a loading
control. (B) The blots corresponding to the dark/sucrose treatment were quantified by
scanning densitometry and the values are represented as in Figure 5.

53

 

Transcriptional regulation of Arabidopsis ACP2 gene by growth/cell cycle signals

The increase in ACP2 mRNA abundance after carbon induction (Figures 10, l 1, 12) may
be brought about by transcriptional or post-transcriptional mechanisms (e.g., mRNA

stability).

Therefore, to investigate which mechanisms are involved in ACP2 gene expression, a 1.5
Kbp DNA region upstream of this gene was fused to luciferase (LUC) reporter and
transformed into Arabidopsis. In order to eliminate the effect on gene expression of the
ACP2 5’ leader (Figures 8 and 9), this region was not included in the construct (see
materials and methods). As a control, a 2 Kbp DNA region upstream of the eIF 4A1 gene
was fused to LUC. The steady-state levels of eIF4Al mRNA remained constant
regardless of the presence or absence of carbon in the media (Figures 10 and 11). In
addition, all constructs carried a second reporter gene (GUS) under the constitutive
CaMV35S promoter to normalize LUC activity for T-DNA copy number, position effect
and in this case also for differential transcriptional activity in cells with variant energetic

status (presence/absence of carbon sources).

Since Arabidopsis cells grown in liquid culture are recalcitrant to Agrobacteria-mediated
transformation, transgenic cell suspension cultures were induced from transgenic
Arabidopsis plants. Stable transformation was preferred for these experiments because
problems associated with transient transformation (e.g., loss of vector and reporter gene

activity) were avoided. This is particularly important when performing experiments that

54

- Sucrose + Sucrose

 

 

Hours: 0 12 24 48 12 24

At-ACPZ

 

rRNA

 

Figure 12. In planta regulation of ACP2 mRNA by carbon.

Arabidopsis seedlings were grown in liquid media for 2 weeks in the presence of
58 mM sucrose and light. Subsequently, seedling were transferred to media
containing 58 mM mannitol and incubated for 2 days in the light (-Sucrose).
Samples were taken at 0, 12, 24 and 48 hours and total RNA extracted. After 2 days
of starvation seedlings were transferred to media containing 58 mM sucrose for 1
day (+Sucrose). Samples were taken at 12 and 24 h after induction and total RNA
isolated. RNA was resolved by electrophoresis and blotted onto nylon membranes.
Filters were hybridized with an Arabidopsis ACP2 cDNA probe (At-ACP2).
Ribosomal RNA (rRN A) in the gel was visualized by ethidium bromide staining.

are carried out during several days. In addition, insertion Of the transgene in the genome
provides a better platform for regulation of ACP2 promoter activity. Interaction of
foreign DNA with nucleosomes in the chromosome context Should facilitate the
organization of the promoter region (e.g., nucleosome position, binding of factors) to

better reflect the actual promoter structure and regulation.

55

 

SUC

ACP2:LUC STV-48h
SUC-24 h

SUC
eIF4A] :LUC STV-48h

SUC-24 h

 

Fold Induction of LUC activity

Figure 13. Regulation of Arabidopsis ACP2 promoter by carbon in
Arabidopsis cell suspension culture.

Arabidopsis cell suspension cultures were generated from leaf tissue of transgenic
plants carrying 1.5 Kb of ACP2 promoter (ACP21LUC) and 2 kB of eIF4A]
promoter (eIF4AlzLUC). Vectors also carried the GUS reporter gene under a
constitutive promoter (CaMV35S:GUS) to control for T-DNA position and copy
number. Three independent cell liquid cultures for each construct were generated
from three independent transgenic plants. Cells were maintained in media
supplemented with 58 mM sucrose (SUC). At the beginning of the experiment,
cells were starved in media containing 58 mM mannitol for 2 days (STV-48h).
After this period cells were transfered to sucrose containing media for 1 day (SUC-
24h). Cell samples were taken at the beginning and end of each period and LUC
and GUS specific activities assayed in triplicate. LUC activity was normalized with
GUS activity (LUC/GUS) and expressed as fold induction of LUC activity.
LUC/GUS ratios were set to l for STV-48 to Simplify the interpretation of the
results.

*’*--‘

The results in Figure 13 indicated that the LUC/GUS ratio of specific activities was
reduced by 2-fold after 2 days of starvation and increased by more than 3-fold when
sucrose was added back. Changes in LUC expression in transgenic Arabidopsis cells
were Similar to those in ACP2 mRNA levels under the same growth conditions (Figures
10 and 11). Moreover, no Significant differences in LUC/GUS ratio of specific activities
were observed when LUC was under regulation of eIF4Al promoter. These results
agreed with previous experiments in which eIF4A] transcript levels did not change with
carbon availability (Figures 10 and 11). Thus, a 1.5 Kb DNA fragment upstream of the
ACP2 gene promoter was sufficient to confer carbon regulation of gene expression.
These results indicated that changes in ACP2 mRNA abundance were the result of
transcriptional activation/de-repression by carbon rather than increased transcript

stability.

Other carbon sources were tested for their ability to induce LUC expression in
Arabidopsis cells. In this regard, glucose and fructose either alone or together were
capable to activate LUC expression in cells carrying ACP22LUC construct (Figure 14).
These results demonstrated that metabolizable carbon sources other than sucrose were
also capable to regulate ACP2 gene promoter. To test if carbon sources different from
sugars could also increase LUC expression, Arabidopsis cells were incubated with 3 mM
sodium acetate after starvation. Acetate is rapidly taken up and used as substrate for fatty
acid biosynthesis, however it is not a good source of carbon to sustain Arabidopsis cell

growth (data not Shown). Nevertheless, acetate regulates gene expression of genes

57

involved in various cellular processes (Sheen,l990). The results in Figure 14 indicated

that acetate failed to induce LUC expression after starvation and suggested that growth

Fold Induction of LUC activity

[0
_.
'N
o.)
.Is

 

mannitol
sucrose
fructose
glucose
deoxy 2-glucose
3-O-M-glucose
acetate

sucrose + cerulenin

llllllll

Figure 14. Regulation of ACP2 promoter by different carbon sources in
Arabidopsis cell suspension cultures.

Transgenic Arabidopsis cells expressing ACP22LUC and CaMV3SS:GUS constructs
were starved for 2 days in mannitol containing media and then induced with different
carbon sources for 1 day. After this period, LUC and GUS Specific activities were
assayed in triplicate. LUC and GUS specific activities assayed in triplicate. LUC
activity was normalized with GUS activity (LUC/GUS) and expressed as fold induction
Of LUC activity. LUC/GUS ratios were set to 1 for mannitol treatment to Simplify the
interpretation Of the results. Sucrose, glucose and fructose were at 58 mM, 2-deoxy-
glucose (2-deoxy glc) at 5 mM, 3—O-methyl glucose (3-O—methyl glc) at 58 mM, acetate
at 3 mM, cerulenin at 0.01 mM . Bars denote Standard deviations of the average.

and not the sole presence of carbon sources was necessary to regulate ACP2 gene

promoter.

To further investigate the relationships between ACP2 gene transcription, carbon supply
and growth, two different strategies were followed. First, cells were incubated with sugar
analogs (3-O-methyl-glucose and 2-deoxy-glucose) that are not metabolized by cells yet
sensed as carbon and therefore provide a sugar-sensing signal (Fujiki et al., 2000).
Second, cells were incubated in the presence of sucrose plus 10 uM cerulenin. The
rationale for the latter experiment was to provide carbon for growth but at the same time
to halt growth/cell division by limiting supply of fatty acids. The results in Figure 14
indicated that both treatments, sugar analogs and sucrose plus cerulenin, failed to induce
ACP2:LUC expression after starvation. These data suggested that growth and/or cell
division is a requisite for regulation of transcription by the ACP2 gene promoter.
Possibly, transcription of ACP2 gene may be regulated by cell-cycle signals. In
agreement with this hypothesis, microarray profile analysis of synchronized Arabidopsis
cell cultures demonstrated that ACP2 mRNA expression is cell-cycle coordinated

(Menges et al., 2002).

Deletion analysis of Arabidopsis ACP2 promoter in tobacco BY-2 cells

To identify discrete DNA elements in the Arabidopsis ACP2 promoter responsible for

transcriptional regulation by grth and/or cell cycle, a promoter deletion analysis was

59

conducted. A promoter deletion series of the Arabidopsis ACP2 promoter was fused to
the luciferase reporter gene and the constructs used to transform tobacco BY-2 cells.
Tobacco cells, instead of Arabidopsis, were chosen for this experiment because the
former cells have the advantage of being transforrnable by Agrobacteria and
consequently more constructs could be conveniently analyzed. Nonetheless, this system
may have disadvantages. First, mechanisms existent in Arabidopsis to activate ACP2
gene expression could be different from those in tobacco cells. Second, factors in tobacco

cells may not recognize elements in the Arabidopsis ACP2 promoter.

The results in Figure 15 demonstrated that LUC expression driven by the 1.5 Kb ACP2
promoter responded to carbon Signals in tobacco BY-2 cells, similar to Arabidopsis cells.
Furthermore, the increase and decrease in LUC expression by carbon availability were in
the same range as observed in Arabidopsis cells (Figure 13 and 14). Thus, these data
indicated that mechanisms for ACP2 gene expression were conserved between tobacco
BY-2 cells and Arabidopsis cells and also that tobacco factors can recognize and regulate

Arabidopsis ACP2 promoter by growth/cell cycle signals.

The deletion series of the ACP2 promoter disclosed the presence of both positive and
negative domains for gene expression in the Arabidopsis ACP2 promoter (Figure 15).
Removal of the —1,500/—750 fragment reduced LUC expression by 6-fold, indicating the
presence of positive elements in this promoter area. Although overall LUC expression
was affected, the response to carbon was unaltered, suggesting that growth/cell cycle

responsive

60

LUC activity

- o

00

 

-1573 +1

Suc

LUC Stv-48h -
Sue-24h

 

 

 

 

 

 

Suc

LUC Stv-48h
Sue-24h

 

 

 

 

 

 

Suc

LUC Stv-48h
Suc-24h

 

 

 

 

 

 

Suc

LUC Stv-48h
Sue-24h

 

 

 

Figure 15. Arabidopsis ACP2 promoter deletion analysis in tobacco BY-2 cells.

Tobacco BY-2 cells were transformed via Agrobacteria with a binary vector
containing a deletion series of the Arabidopsis ACP2 promoter fused to luciferase
(LUC). Vector also carried GUS fused to CaMV35S. Three individual calli were
used to generate three independent cell suspension cultures for each construct. Cells
were maintained in media with 58 mM sucrose (Suc). At the beginning of the
experiment cells were starved in media containing 58 mM mannitol for 2 days (Stv-
48h). After this period cells were transferred to sucrose containing media for 1 day
(Sue-24h). Cell samples were taken at the beginning and end of each period and
LUC and GUS specific activities assayed in triplicate. Luc activity was normalized
by GUS activity and expressed as LUC activity (units of light emitted).

elements were still present in the 750 bp promoter fragment. Interestingly, deletion Of an
additional 250 bp (-750 to —500) reconstituted LUC expression to levels Similar to those

Obtained with the 1.5 Kb promoter fragment (Figure 15). This result indicated that

61

negative elements of expression were localized in the —750/—500 region. LUC expression
still responded to carbon signals and therefore growth/cell cycle elements were present in
the —500 promoter fragment. Further deletion of the promoter (-500 to —250) decreased
by 10-fold the overall expression of LUC, indicating the presence of positive elements in
the —500/-250 fragment. Induction of LUC activity by carbon was in the same range as
with the other constructs (~ 2-3 fold) indicating the presence of growth/cell cycle

responsive elements within the —250 promoter fragment (Figure 15).

Conserved elements in Arabidopsis ACP promoters

In addition to the conserved elements found in the 5’ UTRS (Table 2), computer
alignments of Arabidopsis ACP genes (plastidial isoforms) disclosed the presence of two
additional conserved elements (Figure 16). First, an 11 bp element was present in the
promoter regions of the five ACP genes (consensus sequence: CCTGCATCTCC). This
element was neither present in ACP genes of other plant species nor in other Arabidopsis
FAS and lipid biosynthetic genes (Beisson et al., 2003). Therefore, it may be that the 11
bp element is Specific for Arabidopsis ACP promoters. Nevertheless, it has to be
considered that information on ACP promoter sequences from different plant Species is
still limited. Second, a 6 bp element (GCCAAA) was found at —200 bp in ACP2 and
ACP3 promoter regions but not in the other Arabidopsis ACP promoters (Figure 16).

Interestingly, the same element is present in FASl and FASZ genes of yeast that are
regulated by growth (Chirala, 1992; Schuller et al., 1992) as well as in several

Arabidopsis genes which are cell-cycle regulated (e. g., At-CDC6, A t-MCM3) (Stevens et

Figure 16. Conserved elements in Arabidopsis ACP promoters.

Two conserved elements in the Arabidopsis ACP genes were identified by computer
alignments. An 11 bp element (highlighted in gray) was found in the promoter regions of
the five Arabidopsis ACP genes. The consensus sequence for this element is indicated at
the bottom of the figure together with its location (relative to the start codon [capital
letters]) in the different ACP genes. Interestingly, the 11 bp element in the ACPS gene is
located in the coding sequence. A 6 bp element GCCAAA (underlined) was found only
in ACP2 and ACP3 promoters. This element overlaps with the 11 bp element and this
observation may further suggest that these two elements are important for ACP gene
expression.

63

Iputer

ms of

red at
apital
me is
only
i this
gene

v

 

ACP1

ttgtcCCTGGTTCTCCgactgagagaagcagccatgatcttagtaaaccttgaggagaaag
atatatagaaacttaaccaaaaaacttcttcttgctcttccctcttatggtgactagtatt
gtgtttcacgaaacaATG

ACP2
ttccaaCCTGCATCTfiQQAAAgcacccaactccacctgacttgtctgtgtctgttgcgttca
ctttcattggaatctcagattttttattt‘ttttgtttgttttggttgttgcgatttgtgactataaaacctctcc
cacttggttcttcactctcactgtttctcatctcttcgtcttctccgcctccttcaatctcactccgatctc
tctacgattcattcgttctfi

AC P3

tttgttatttctctaCCTGCATCT_(3—__CCAAA tcacccacctcacctgacttgtctgtctgaattct
ctcattggaatctcagaaaagttttttttttttttttgtttaggttaaaatactataaaattaaaaataagtct
cccacttggattcttcactgtgtctcacatctctttgtcttctccgcctcctccgatctcactccgatctc
tctacgattcattcttc tATG

AC P4

gtgccgaagataggCCTGAATCTCCgagaacaaacctccgccacaaaaacagaagactt
gttgcttgcgtatcataatacgacgccgtatctctacttgctcgaacaaacccaaaaagatacatat
caagagattaaaccttatccaactaagagaagccatttttattttttttgggtctctgagttgtgtattga
gcttcatctccttcaafl

ACPS
ggatggtacttagaatagattttccaaaatgatggcagaatagaacgtggctctataaatacataaa

tcccagcagtgtttgccatcagctacaaaatagtaattcacqctccttaaacaatccgccatctctct
cgatcagatcgataATG gcgacaagtttctgcCCTCCATCTCC

Consensus sequence (location) CCTGCATCTCC

ACP1 (-615) CCTGGTTCTCC

1 1 bp element ACP2 (-21 l) CCTGCATCTCC
ACP3 (-202) CCTGCATCTCC

ACP4 (-188) CCTGAATCTCC

ACPS (+17) CCTCCATCTCC

Figure 16

64

al., 2002; de Jager et al., 2001).

Although ACP2 and ACP3 appear to derive from a recent gene duplication (these
isoforms are > 80 % identical. in their amino acid sequences and the respective genes lay
one next to the other in the Arabidopsis genome), the divergent amino acid sequences of
the other ACP isofomis (60-70 % identity) indicate that they diverged long ago.
Therefore, the 11 bp element present in the promoter region of the five ACP isoforms
represents a DNA motif that has been conserved for a long period throughout evolution.
Thus, this element is a good candidate for a regulatory element of ACP expression.
Because the presence of the 6 bp element in the ACP2 and ACP3 promoters is the result
of a more “recent” gene duplication event, its conservation in these two promoters may
only reflect its short existence in evolutionary terms. However, the presence of the same
element in other Arabidopsis cell-cycle regulated genes suggests that the 6 bp element is

also a good candidate for ACP gene regulation.

Changes in mRNA expression by inhibition of fatty acid synthesis in Arabidopsis
cells

To investigate Whether feedback mechanisms for FAS gene regulation Similar to those in
bacteria (Schujman et al., 2003) also exist in plant cells, Affymetrix GeneChip analysis
was used to follow changes in gene expression after inhibition of fatty acid synthesis in
Arabidopsis cells. Gene expression analysis of cell suspension cultures presents several
advantages over analysis of whole tissues. First, cell cultures are homogeneous systems

as opposed to tissues which are composed of different cell types. Second, growth

65

conditions can be rapidly changed and cells more readily take up components from the
media. Finally, cells Share the same environment and therefore they should perceive
Similar signals. Cerulenin, a potent inhibitor of fatty acid synthesis in plants was utilized
in the experiment. This drug crosses cell membranes rapidly and specifically inhibits

condensing enzyme KAS I (B-ketoacyl-ACP synthase) (Schneider and Cassagne,l995).

In order to evaluate the concentration of cerulenin required to inhibit fatty acid synthesis
significantly but with minimal cytotoxic effects, cells were incubated with 0, 10, 30 and
100 uM cerulenin for 0. 2 and 6 hours. Labeled acetate was added to the cultures and
after the incubation period, cells were quenched with hot isopropanol and lipids
extracted. Analysis of labeled fatty acids demonstrated that after 2 hours the inhibition of
F AS was 65%, 82% and 94% for 10, 30 and 100 uM cerulenin, respectively, compared to
the control (Figure 17). After 6 hours, inhibition of FAS in the presence of cerulenin
remained Similar to values measured at 2 hours (70%, 84% and 96% for 10, 30 and 100
uM cerulenin respectively) (Figure 17). These data indicated that the effect of cerulenin
on F AS persisted after 6 hours of treatment. In addition, the constancy in the percentage
of FAS inhibition throughout the experiment suggested no major cytotoxic effects by
cerulenin on Arabidopsis cells during this period. Based on these results, a concentration
of 10 uM of cerulenin was chosen for gene expression experiments. This low
concentration of inhibitor is enough to substantially reduce fatty acid synthesis (70%) and
yet provide small amounts of fatty acids that would minimize cytotoxic effects and stress

responses.

66

FAS genes do not respond to inhibition of fatty acid biosynthesis in Arabidopsis cells

Affymetrix gene chips (8K) representing approximately one third of the Arabidopsis
genome were used to evaluate changes in gene expression afier inhibition of fatty acid
synthesis. Two flasks with 100 mL of cells growing at exponential phase were incubated
in the presence of 10 BM cerulenin for 6 hours. Samples (20 mL) were taken at 0, 2 and 6
hours after drug treatment from each of the flasks (two‘biological replications per time
point). Total RNA was isolated from each sample and used to synthesize cDNA with
poly(T) primers fused to the T7 promoter. Subsequently, T7:cDNAs were used as
templates for cRNA synthesis by in vitro transcription in the presence of biotinylated-

ATP. Finally, biotinylated-cRNA was used for GeneChip hybridization reactions.

The results in Table 3 indicated that mRNA levels from some genes involved in fatty acid
and lipid metabolism changed slightly after 2 and 6 hours of inhibition of fatty acid
synthesis. For example, the mRNA levels corresponding to the biotin containing subunit
(BCCP) of acetyl-COA carboxylase increased ~ 2-fold after 6 h of cerulenin treatment.
A few other mRNAs showed slight increases (e.g., 1.4-fold for ACP2 and cytosolic
acetyl-COA carboxylase). The standard deviations of these values were within 5 % of the
average value and therefore the fold increases were reliable. In contrast, the mRNA levels
of other lipid related genes were reduced by cerulenin treatment. Transcripts for acyl-

ACP

67

140000

120000

100000

80000

DPM

60000

40000 ~

20000

0 ,

 

fItfiM
10 UM ,
112130 uM
C1100 uM

 

 

 

0 hours 2 hours 6 hours

Figure 17. Inhibition of fatty acid synthesis by cerulenin in Arabidopsis

cells.

Arabidopsis cells were incubated with 0, 10, 30 and 100 uM cerulenin for 0, 2
and 6 hours. Labeled acetate (0.025 mCi) was added to the cultures. After the
incubation periods cells were quenched with hot isopropanol and lipids
extracted. Radiolabeled fatty acids were transmethylated and separated by thin
layer chromatography. Quantitation of bands corresponding to fatty acids was
performed by scanning in an Instant-Imager and scintillation counting. Results
are expressed in desintegrations per minute (DPM).

thioesterases declined approximately 1.6-fold after 6 h of drug treatment (Table 3). Thus,

although small differences could be seen in the mRNA levels of some fatty acid synthesis

genes, mRNA levels for most FAS and lipid synthesis genes were not significantly

altered by inhibition of fatty acid biosynthesis.

68

Significant changes in mRNA levels were observed for genes related to cellular processes
such as stress response, metabolism, drug resistance and several regulatory proteins such
as transcription factors and kinases. Approximately 90 genes presented increases in
transcript abundance of more than 2-fold after 6 h of cerulenin treatment (Table 4).
Changes in the transcript levels of genes related to stress response, metabolism and drug
resistance most likely reflected the stress conditions imposed by inhibition of fatty acid
synthesis. Approximately 110 genes presented lower mRNAs levels than control cells

after 6 h of cerulenin treatment (Table 5).

DISCUSSION

One objective addressed by the current work is the nature of the signals that give rise to
changes in the expression of genes involved in fatty acid synthesis. Although a limited
number of previous studies suggest a minor role of light on the expression of these genes
(Battey et al., 1990; Baerson et al., 1993, Scherer et al., 1987), the mutually dependent
relation between cthTOplast biogenesis and de novo production of glycerolipids envisage
a closer connection between light and at least some of the genes for this pathway. In this
report we demonstrated that light affects the expression of mRNAs encoding acyl carrier

protein isoforms from Arabidopsis.

First, we demonstrated that the levels of the messenger for ACP4 increased 4- to 5-fold
after light treatment of dark grown plants. The similarity in the kinetics of mRNA
induction between ACP4 and FEDA in Arabidopsis suggests Similar mechanisms of

activation for both genes. Nevertheless, it remains to be examined whether the increase of

69

Table 3 . Changes in mRNA abundance of genes involved in lipid metabolism after

inhibition of fatty acid synthesis in Arabidopsis cells.

 

 

 

descriptions Fold chan e
at 2 h at 6 h

AT5G16390 acetyl-COA carboxylase (BCCP subunit) 1.7 i 2.1
AT4G38570 putative phosphatidylinositol synthase 1.7 r 2.1
AT4G22340 CDP-diacylglycerol synthetase-like protein 1.5 2.0
AT1G3617O acetyl-COA carboxylase subunit 1.2 1.5
AT1G54580 acyl carrier protein isoform 2 1.0 1.4
AT1G54580 acyl carrier protein isoform 2 1.2 1.4
AT3G51840 Short-chain acyl COA oxidase 1.2 1.2
AT3G12120 delta-12 desaturase (Fad2) -2.1 1.2
AT4G30950 omega-6 fatty acid desaturase (fad6) 1.0 1.2
DL4775C thioesterase like protein 1.0 1.1
glycerol-3-phosphate acyltransferase (ACTI) 1.0 1.1

AT1G01480 acetyl-COA carboxylase (ACC2) 1.0 1.1
ATZGZS710 biotin holocarboxylase synthetase -1.1 1.1
ATSGOSSSO omega-3 fatty acid desaturase (fad8) 1.0 1.1
AT2G44620 acyl carrier protein precursor (mitochondrial) -1.0 1.1
AT2G32260 putative phospholipid cytidylyltransferase 1.0 1.1
AT4G23850 acyl-COA synthetase-like protein 1.3 1.1
AT1G67730 b—keto acyl reductase (glossy8) 1.3 1.1
AT4G13840 fatty acid elongase-like protein (cer2-like) 2.0 1.1
DL4770C thioesterase like protein 1.6 1.1
AT5G35360 biotin carboxylase subunit (CAC2) -l .3 1.1
AT4020870 fatty acid hydroxylase-like protein 1.0 1.1
ATZG31360 delta 9 desaturase 1.0 1.1
AT2G33150 3-ketoacyl-COA thiolase -1.3 1. 1
AT3G11170 omega-3 fatty acid desaturase 1.0 1.1
AT4G34510 putative ketoacyl-COA synthase -l.3 1.1
AT1G76490 3-hydroxy-3-methylglutaryl COA reductase -8.3 1.1
AT3G1 1 170 fatty acid desaturase -1.9 - l .l
AT4G34250 fatty acid elongase-like protein -1.0 -1.1
DL4435W triacylglycerol lipase like protein - l .2 - l .1
AT1G68530 very-long-chain fatty acid condensing enzyme CUTl -l .0 -1.1
AT4G25050 acyl carrier-like protein (ACP-4) 1.0 -1.2

 

 

 

 

*Fold change: fold increase in signal intensity compared to time o h

70

 

Table 4. Example of transcripts that increase more than 2 fold after 6 hours of

cerulenin treatment.

 

 

 

 

 

 

 

descriptions Fold chan e
at 2 h at 6 h

T020048 jasmonate inducible protein isolog 9.0 11.2
AT5G13930 chalcone synthase (CHS) 1.9 10.8
AT2G34660 multidrug resistance-associated protein 2 7.5 9.3
AT4GI9030 nodulin 26 like protein 14.2 8.7
AT4G12360 putative lipid transfer protein 6.5 5.4
AT1G17740 Phosphoglycerate dehydrogenase 2.6 5.3
AT1G14900 high mobility group protein a (HMGa) 1.9 5.1

AtMYB42 R2R3-MYB transcription factor. 9.0 5.0
AT2G33810 squamosa promoter binding protein-like 3. 4.5 4.7
AT5G18170 glutamate dehydrogenase 1 (GDHl) 2.4 4.6
ATl 610460 germin-like protein (GLP7) 4.7 4.6
AT4G21880 membrane-associated salt-inducible protein 4.0 4.6
AT5G08640 flavonol synthase 1.6 4.2
AT2G16590 putative protein 24.7 4.1
AT3G22490 LEA D34 protein homologue typel. 2.6 4.0
AT1G78820 receptor like ser/threo kinase ARK3 3.7 4.0
AT2G47000 multi drug resistance proteins 4.4 3.8
AT2G01830 putative histidine kinase (CREl) 3.0 3.6
F13P17.32 cytochrome P450 homolog 2.4 3.5
F16N3.5 prolamin box binding factor (PBF) 3.7 3.4
AT1G09530 phytochrome-associated protein 3 (PAP3). 2.0 3.3
AT1G19050 ARR7 mRNA for response regulator 7 3.8 3.2
AT2G1 1620 putative 3.3 3.2
AT2G18650 RING zinc finger protein 2.8 3.1

 

*Fold change: fold increase in signal intensity compared to time o h

71

 

Table 5. Example of transcripts that decrease more than 2-fold after 6 hours of
cerulenin treatment.

 

 

 

 

 

 

 

72

descriptions Fold change
at 2 h at 6 h
ATSGS756O xyloglucan endotransglycosylase (TCH4) -3.45 -8.85
AT4G27280 calcineurin B-like protein 3 -4.05 -8.45
AT3G47380 putative -2.35 -6.95
AT2G19800 putative -0.55 -6.40
AT1G32170 xyloglucan endotransglycosylase-related protein -3.55 -6.35
AT4GZ7450 glutamine-dependent asparagine synthetase -1.20 -5.75
late embryogenesis abundant protein homolog -2.90 -5.65
AT2G37170 aquaporin -4. 50 -5 .40
AT4G23550 putative protein -1 .60 -5.40
AT3G47340 glutamine-dependent asparagine synthetase -3.55 -5.40
AT2G33830 putative auxin-repressed protein -3.25 -5.30
AT1G131 10 cytochrome P450 -0.15 -5.20
AT2G44380 putative CHP-rich zinc finger protein -0.40 -5.05
AT2G28630 putative fatty acid elongase -1.60 -5.00
class 1 non-symbiotic hemoglobin -l.15 -5.00
AT5G59520 zinc transporter (ZIP2) -4.50 -4.95
AT4G02380 putative -2.80 -4.95
AT5G24090 acidic endochitinase gene -3.55 -4.95
class IV chitinase -3.70 -4.90
AT3G57700 protein kinase-like protein -3.85 -4.85
DL4265W membrane transporter like protein -5. 15 -4.80
A_TM017A05.3 putative -0.75 -4.80
AT2G44840 ethylene response element binding protein -4.50 -4.75
AT4G08950 putative phi-l-like phosphate-induced protein -2.20 -4.65
AT4G22590 trehalose-6-phosphate phosphatase-like protein -2.50 -4.65

*Fold change: fold decreased in signal intensity compared to time o h

 

the ACP4 transcript levels is direct due to light (e.g. via phytochromes) or indirect via
cell growth. The analyses of 1 Kb of genomic sequence upstream of the Acll.4 gene
disclosed the presence of several GATA-like motifs. The GATA (or I) boxes are
regulatory elements that are functionally important in many light-regulated promoters.
The core element is defined as GATAA and related GATA motifs with variable flanking
sequences are found in several promoters (Terzaghi et al., 1995). For instance, the le
upstream region of the ferrredoxin-A gene contains seven copies of related GATAA
elements. Although the actual transcription initiation site for the ACP4 gene has not been
mapped yet, we localized the GATA-like motifs respective to the AUG initiation codon
of ACP4. Thus, the upstream region of this gene presents two AAGATAA elements at —
715 and -812, one AGATAA at —505 and three GATAA at —70, -526 and —908 base pairs
respective to the translation initiation codon. The presence of these elements in the
upstream region of the gene for ACP4 suggests the direct role of light on the transcription
of this gene. In contrast, only one and two copies (expected by chance) of these elements
are present in the upstream regions (1 Kb) of AclI.2 and AclI.3 genes respectively. This
observation is consistent with results that demonstrate that light has no impact on mRNA

levels of ACP2 and most likely ACP3 (Figure 5 and Baerson et al., 1993).

A second aspect of the influence of light on ACP gene expression was observed at the
level of ribosome association. A general effect of light on polysome formation has been
previously described in several studies (Giles et al., 1977; Mosinger et al., 1983). These
works Show that the proportion of ribosomes present as polyribosomes increases

substantially in response to light. Moreover, continuous far—red (FR) light mediates a

73

strong increase in the relative level of polysomes, demonstrating the participation of
phytochromes in the response. Despite this general effect on translation, there is evidence
that particular mRNAs are less affected by the decrease in the translation rate in the
absence of light. For example, Berry et al (1990) showed that in Amaranth cotyledons the
expression of the small and large subunits of Rubisco are not found associated with
polyribosomes in cotyledons of dark grown plants but rapidly recruited onto them upon
illumination. In contrast, mRNAs encoding non-light regulated proteins are associated
with polysomes regardless of the light-dark treatment. Our results (Figure 6)
demonstrated that the transcripts for the ACP isoforms tested and FEDA largely
dissociated from polysomes during the dark period in contrast to the non-plastidic eIF 4A
mRNA, which remained associated with high molecular weight polyribosomes in the
dark. Thus, these observations reinforce the notion that cytoplasmic mRNAs are
differentially associated with polyribosomes during the light-dark period (Berry et al.,

1990; Petracek et al., 1997).

In chloroplasts of Chlamydomones and plants, both translation and mRNA stability are
enhanced by nuclear encoded factors that bind to the 5’ UTR of several chloroplast-
encoded RNAS (Danon et al., 1991; McCormac et al., 2000). Likewise, several studies
confirmed the major role of the 5’UTR in translation and mRNA stability of cytoplasmic
mRNAs (Dickey et al., 1998; Lukaszewicz et al., 1998). A previous report demonstrated
that these regions can also be important for transcription activation (Bolle et al., 1994 and
1996). Analyses of the 5’ leader sequences of several genes encoding for thylakoid

proteins in spinach, disclosed the presence of CT-rich regions, designated CT-leader

74

boxes (CT-LB and CT-B) (Bolle et al., 1994). The deletion of these elements from the
PsaF (subunit III of photosystem I) and PetH (plastocyanin) upstream gene sequences
severely reduces the transcription of a reporter gene in transgenic tobacco (Bolle et al.,
1994). It is noteworthy that the CT-LB box is also present in the 5’UTR of the spinach
ACP II (Bolle et al., 1996). The presence of several CT-rich elements and the CTCCGCC
box in the ACP 5’UTRS of several diverse plant species (Table 2) suggests that these
motifs might be important for ACP gene expression. The results presented in Figure 9
provide convincing evidence that these regions have a positive effect on reporter gene
expression in transgenic Arabidopsis plants. Moreover, this effect was enhanced by light,
suggesting that the ACP 5’UTRs may contain light-responsive elements that increase
gene expression post-transcriptionally and/or transcriptionally in the context of the
CaMV3SS promoter (Figure 9). We also demonstrated that the 5’ UTRS of Arabidopsis
ACP1 and ACP2 conferred differential reporter gene expression in a tissue specific
manner (Figure 9). At this point it is not possible to distinguish between a transcriptional,
translational or mRNA stability mechanism to explain these differences in reporter gene

activity.

Gallie et al (1992) showed that 5’ leader length alters the expression of a reporter gene in
carrot protoplasts. In particular, translation is influenced by leader length, and a 74 base
leader construct is expressed 6-fold more highly than a 29 base leader construct.
Conversely, Bolle et al (1994) found that in transgenic tobacco, the expression of a
reporter gene under CaMV35S promoter is not altered when the 5’ leader of PsaF (188

bases) is added to the 5’ UTR (24 bases) of the native reporter gene mRNA. Based on

75

these results, it is possible that part of the differences observed in the LUC/GUS ratios in
transgenic Arabidopsis tissues could be attributed to differences in the length of the ACP
and control leaders (Figure 8 and 9). However, the tissue specific differences in reporter
gene activity conferred by the three constructs analyzed, that ranged from almost no
effect in roots to 10 to 30-fold increase in leaves and seeds, supports a sequence-

dependent enhancement by the ACP 5’ UTRS.

The results presented in Figures 10 and 11 for Arabidopsis mesophyll-derived cell culture
indicate that the messengers for ACP2, ACP3 and ACP4 are under growth and/or
metabolic control. The induction of the ACP2 and ACP3 transcript levels in the presence
of 58 mM sucrose together with the inability of sugar analogs to reproduce this effect
demonstrates that cell growth and/or a signal generated by sucrose or downstream of
glucose is required for up-regulation of these messenger RNAS. In contrast to the
response of these messenger RNAS, the ACP4 mRNA levels remained steady in starved
cells and in the presence of 3-O-methyl-glucopyranose whereas they were down-
regulated in the presence of 58 mM sucrose in the dark (Figure 10 and 11). The
observation that mRNA levels for the ER localized F AD2 desaturase Showed no variation
under the conditions analyzed indicates that the response of genes encoding plastid
components of fatty acid synthesis may differ from those controlling later steps such as

cytosolic desaturation.

Regulatory mechanisms of ACP2 gene expression also responded to carbon signals in

planta, suggesting that these mechanisms were physiologically relevant and not the

76

consequence of cell culture induction. Analysis of the ACP2 promoter demonstrated that
carbon-mediated regulation of ACP2 gene expression was transcriptional and that
promoter elements responsible for this regulation were located within —250 bp from the
transcription initiation site. Comparison of Arabidopsis ACP promoters identified a
GCCAAA element located at —200 bp of ACP2 and 3 genes. This element was also found
in yeast FAS genes (Chirala, 1992; Schuller et al., 1992) and Arabidopis cell-cycle
regulated genes (Stevens et al., 2002; de Jager et al., 2001). Therefore, it is a good

candidate element for growth/cell cycle regulation of A CPZ gene expression.

Analysis of gene expression after inhibition of fatty acid synthesis suggested that, in
contrast to bacteria, fatty acid and lipid synthesis genes in plants did not respond to
feedback mechanisms of gene expression after blocking this primary pathway. Thus,
mechanisms other than regulation of FAS mRNA abundance may be involved in
counteracting the inhibitory effect of cerulenin in plant cells (e.g., post-translational

modifications of gene expression, metabolic regulation of FAS enzymes).

In summary, results of this study demonstrate that expression of genes involved in plant
fatty acid synthesis is under multiple levels of control. Light clearly has a major impact
on ACP4 mRNA levels, but this impact does not extend to the other ACP isoforms
examined here. These different responses to light on ACP are probably transcriptional,
based on the presence of GATA boxes in the ACP4 but not other ACP upstream regions.
Light control is also exerted post-trancriptionally because all ACP isoforms are more

highly associated with polysomes in the light than the dark. Experiments with suspension

77

cultures indicate that differences in expression patterns of ACP isoforms are also
observed in their response to sugar supplements. A common element in both the leaf and
tissue culture experiments is that ACP4, the most leaf-specific isoform, behaves in a
manner Similar to mRNA for genes involved in photosynthesis (FEDA or RBCSIA). In
contrast, expression of the other ACP isoforms may be most responsive to demands for
fatty acid synthesis brought about by enhanced growth. This level of complexity
demonstrated by multiple ACP genes under different controls may reflect the need of
plant cells to tightly regulate both the amount and the cellular destination of fatty acids
produced in plastids (Ohlrogge and Browse, 1995) and to match the supply of fatty acids

to different tissue and environmental demands.

MATERIALS AND METHODS

Nomenclature

There are five genes for plastidial ACP in the Arabidopsis thaliana genome (Mekhedov
et al., 2000). In this study we report results for Ac11.1, AclI.2, AclI.3 and AclI.4 genes,
corresponding to the gene products ACP1, 2, 3 and 4 respectively. Acl].1 (GenBank
accession number X13708) was previously described in Post-Beittenmiller et al., 1989
and Ac112.2 and AclI.3 (GenBank accession numbers X57698 and X57699 respectively)
in Lamppa et al., 1991. We propose that the ACP major leaf isoform (Shintani, 1996)
gene be named AclI.4 (isoform ACP4, At4g25050 or F13M23.190) and that Ac11.5

(ACPS) referred to GenBank accession number: A_TM021B04.6 or AF 007271.

78

Plant Growth and Lighting conditions

Wild type Arabidopsis thaliana plants (ecotype Columbia) were grown on soil at 20°C in
a 12:12 hours lightzdark photoperiod (80-100 umol m'2 s"). For the analyses of light-
mediated induction of mRNA levels, 2 week-old plants were left in the dark for 24 hours
and reilluminated with white fluorescent light (80-100 umol m'2 s”) for the times
indicated in Figure 5. Leaf tissue was harvested and immediately frozen in liquid nitrogen
prior to RNA extraction. For polyribosomal association analyses, 2 week-old plants were
dark incubated for 12 hours and then incubated in the dark for an additional 12 hours or
illuminated for 12 hours. Leaf tissue was harvested at the end of each period and

processed as indicated above.

Constructs and Plant Transformation

Sequences corresponding to the 5’ untranslated regions of ACP1 (GenBank accession
number X13708) and ACP2 (X57698) mRNAs were obtained by PCR amplification of
Arabidopsis (ecotype Columbia) genomic DNA using the primers:
5 ’CATGCTCGAGCTCTTTGTACACTCCGCCCT3’ and 5 ’-
CATGCCATGGCGCTGAATTGAGTCGCCAT-3’ for ACP1 (gene AclI.1); 5’-
CATGCTCGAGACTGTTTCTCTATCTCTTTG-3’ and 5’-
CGCGCCATGGCAATGGAAGCCATAGAAGAAT-3’ for ACP2 (gene AclI.2). The

PCR amplification products were digested with 20101 and Ncol and cloned in frame to the

79

firefly luciferase gene (LUC) in pUC-LUC-BT2 (WeiBhaar et al., 1991) to give pTACPl
and pTAC2 respectively. A truncated version of the ACP1 5’ leader sequence was
generated by in vitro annealing of two synthetic oligonucleotides: 5’-
TCGAGCTCTTTGTACAAACAATGGCGACTCAATTCAGCGC-3’ and 5’-
CATGGCGCTGAATTGAGTCGCCATTGTTTGTACAAAGAGC-3’. The double-
stranded fragment was cloned in frame to the LUC coding region in pUC-LUC-BT2 to
give pTdell. All constructs were confirmed by sequencing. The vector pTACPl was
digested with BamHI and the vectors pTACP2 and pTdell with BglII. The resulting
fragments were cloned independently in the BamHI site of the binary vector pCAMBIA-
2201 (Hajdukiewicz et al., 1994) to generate pCaACPl, pCaACP2 and pCadell
respectively (Figure 8). The orientation of the inserts was confirmed by restriction
mapping using Pstl. Agrobacterium tumefaciens strain C58Cl (pM90) (Koncz et al.,
1986) was transformed with the pCaACPI, pCaACP2 and pCadell binary vectors by the
freeze/thaw transformation protocol (An, 1987). The leaf-vacuum infiltration method was
used for Arabidopsis transformation (Bechtold et al., 1993). The transgenic lines
transformed with pCaACPl , pCaACP2 and pCadell were named atL-acpl, atL-acp2 and

atL-dell respectively.

A 1.5 Kb DNA fragment upstream of the Arabidopsis ACP2 gene was amplified by PCR
using the following primers: fwd, 5’ GACTGGATCCTGCATTAGCTGGTAGTTCAG
3’ and rvs, 5’ GACTCTCGAGTGAGAAACAGTGAGAGTGAA 3’. The resulting PCR
fragment was digested with BamHI and Xhol and ligated into the pUC-LUC-BT2 vector

upstream of the luciferase coding sequence (Td-2-ACP2-LUC). A 2 Kbp upstream region

80

corresponding to the Arabidopsis eIF 4A1 gene was amplified by PCR using the following
primers: fwd, 5’ GACTGGATCCGAAAACGAACCTCAGTTCAG 3’ rvs, 5’
GACTCTCGAGGAGAGACTGGTGGAATATCC 3’. The resulting PCR fragment was
digested with BamHI and Xhol and ligated into the pUC-LUC-BT2 vector upstream of
the luciferase coding sequence (Td-2-eIF4A1-LUC). The constructs were digested with
Bglll and independently subcloned into the BamHI site of pCAMBIA-2201 binary vector.

The orientation of the insert was analyzed by digestion with Pst-I and sequencing.

For construction of the ACP2 promoter deletion series, the vector Td-2-ACP2-LUC
carrying 1.5 Kbp upstream region of the ACP2 gene was used as template for PCR. A
combination of forward primers with the same reverse primer (same sequence as rvs
primer described above) was used to generate the deletions. The forward primers were as
follows: D750, 5’ GACTGGATCCGTAGTA AACCACTATTGAAA 3’; D500 5’
GACTGGATCCCACATAGTAAATTGATGCGT 3’; D250 5’
GACTGGATCCGACGTAGTAGAATTTACAAA 3 ’; D l 00 5’
GACTGGATCCTGACTTGTCTGTGTCTGTTG 3’. The PCR products were digested
with BamHI and Xhol and ligated into Td-2 vector. The different constructs were
digested with BglII and ligated into the BamHI Site of pCAMBIA 2201 as indicated

above. All constructs were confirmed by restriction mapping and sequencing.

Selection of Transgenic Plants and Tissue Collection

81

Seeds (T1) from transformed Arabidopsis plants were surface-sterilized and sown on
seed germination media in the presence of 50 ug/mL kanamycin. After one week,
resistant seedlings (T1 plants) were transplanted individually to soil and grown at 20°C in
a 18:6 hours lightzdark period until they set seeds. For reporter gene analyses in leaf
tissue, seeds from T2 plants were selected in germination media under 50 ug/mL
kanamycin and transferred to soil. Plants from 10 independent transgenic lines for each
construct (see Figure 9) were grown in a 12:12 hours lightzdark cycle for 2 weeks. The
last day the plants were kept in the dark for 24 hours and the leaf tissue from half of the
plants per line was collected and immediately frozen in liquid nitrogen for subsequent
enzyme analyses. Leaf tissue from the remaining plants was harvested as above after 6 h
of reillumination with white fluorescent light (80-100 umol m"2 s"). For reporter gene
analyses in developing seeds, T2 seedlings were selected as above and transferred to soil.
Ten independent lines per construct (see Figure 9) were grown in a 16:8 hours lightzdark
cycle. Flowers from the primary stem were tagged in the morning every day during one
week. Flower anthesis was considered as day zero in our experiment, and mid-stage
siliques (8 to 10 days after anthesis) were dissected and the seeds immediately frozen in

liquid nitrogen for subsequent enzyme analyses.

Cell Suspension and Root Liquid Cultures

Arabidopsis (ecotype Columbia) suspension cultures (Axelos et al., 1992) were a gift

from N. Raikhel laboratory (Michigan State University, DOE Plant Research

Laboratory). Suspension cultures were maintained in a 12:12 hour lightzdark photoperiod

82

at 22°C on a rotary Shaker (120 rpm) in CSM-sue media (0.32% (w/v) Gamborgs with
minimal organics (Sigma G5893), 58 mM sucrose, 0.05% (w/v) MES, 1.1 pg mL'l 2,4D,
pH 5.7). For the analyses of mRNA levels in different growth conditions, 100 mL cell
cultures were grown for 3 days in CSM-sue in the presence of white light (100 umol m’2
3"). After the third day, the cells were pelleted at 700 rpm and washed twice with CSM-
ma (0.32% (w/v) Gamborgs with minimal organics (Sigma G5893), 58 mM mannitol,
0.05% (w/v) MES, 1.1 ug mL" 2,4D; pH 5.7). Cells were finally resuspended in 150 mL
of CSM-ma and incubated in the presence of light for 2 days (starved cells). An aliquot of
10 mL was taken at the times indicated in Figure 10 and the cell pellet stored at -80°C
until RNA extraction. The remaining 100 mL of the culture was pelleted as before,
resuspended in either 50 mL of CSM-sue or 50 mL of CSM-ma and incubated in the dark
for 24 hours. An aliquot of 10 mL was taken at the times indicated in Figure 11 and the
cell pellet stored at —80°C until RNA extraction. In the experiment with glucose analogs,
cells were starved with CSM-man for 2 days and subsequently transferred to CSM
supplemented with either 58 mM of 3-O-methyl-D-glucopyranose (Sigma) or 0.2 mM of
2-deoxy-D-glucose (Sigma). Cells were incubated in the dark and an aliquot of 10 mL
was taken at 0, 3, 6, 12 and 24 hours and the cell pellet stored at -80°C until RNA

extraction.
Induction of Arabidopsis cell suspension culture from leaf tissue was performed as

described by Axelos et al (1992). Tobacco bright yellow-2 cells (BY-2) were transformed

via A grobacterium tumefaciens strain LBA4404 as described by BaO et al (2002).

83

For root liquid culture, approximately 50 seeds from ten independent transgenic lines (T2
plants) per construct (see legend Figure 9) were surface sterilized and incubated in root
liquid media (0.43% (w/v) MS salts, 0.05% (w/v), MES, 0.1% (v/v) 1000X B5 vitamin
stock, 50 ug mL‘I kanamycin, pH 5.7) at 4°C for two days. The seeds were then
incubated at 22°C in continuous fluorescent white light on a rotary shaker (120 rpm) for 2
weeks. Root tissue from approximately 20 seedlings per transgenic line was collected
under the dissecting microscope and frozen in liquid nitrogen for subsequent enzyme

analyses.

RNA Extraction and Northern Analyses

Total RNA was isolated from plant tissue and cell pellets by standard phenol/chloroform
extraction and lithium chloride precipitation (Sambrook et al., 1989). The RNA was
fractionated in formaldehyde-agarose gels and blotted onto Hybond-N filters
(Amersham-Pharmacia Biotech, NJ). In all cases hybridization reactions were done at
42°C in 50% formamide with a-[32P]-dCTP radiolabeled fragments corresponding to
ACP2 (X57698), ACP3 (X57699), ACP4 (RXW18), FEDA (M35868), eIF4A (X65052),
FAD2 (L26296) and RBCSIA (X13611). After autoradiography, the relative mRNA
levels from the northern blots were quantified by scanning densitometry (Molecular
Dynamics). Messenger Specific probes for ACP2 and ACP3 transcripts were generated by
PCR amplification of Arabidopsis (ecotype Columbia) genomic DNA corresponding to
the 3’UTR of these transcripts. The primers used were: 5’-

TGAAAAGGCCAAGTAGAAT-3’ and 5’-GTCAGATACAAGCCTTGTA-3’ for

84

ACP2; 5’-GGAAAAGGCCAAGTAGAAA-3’ and 5’-CAAGCCTTGTAATAATTATC-

3’ for ACP3.

Polyribosome Analyses

The protocol for polyribosome isolation was adapted from Petracek et al (1997) with the
following modifications: approximately 1 g of Arabidopsis leaf tissue or B.napus
developing seeds were homogenized in 4 mL of U buffer (200 mM Tris-HCl , pH 8.5, 50
mM potassium chloride, 25 mM magnesium chloride, 2 mM EGTA, 100 ug/mL heparin,
2% polyoxyethylene, and 1% deoxycholic acid), centrifuged at 13,000 g for 15 min at
4°C and the complete supernatant (4 mL) was loaded onto a 30 mL linear sucrose
gradient (15 to 60%). After gradient centrifugation, fifteen 2 mL fractions were collected
by dripping directly into 2 mL of phenol-chloroform, 50 uL of 10% SDS, 40 uL Of 0.5M
EDTA, and 10 uL of 100 mM aurin-tricarboxylic acid (Sigma). The final RNA pellets
were washed with 70% ethanol, dried and resuspended in 10 LIL water, 20 uL forrnamide,
6.5 uL formaldehyde, and 3.5 uL MOPS buffer (0.2 M MOPS, pH 7, 50 mM sodium
acetate, 5 mM EDTA). Samples were heated for 15 min at 68°C and loaded onto a
formaldehyde-agarose gel. Northern analyses were performed as described above. For
gradient UV profile analyses, two gradients were spun in parallel and one used to
measure UV absorbance. Fifty 100 uL samples were taken at identical volume intervals

and the absorbance measured at 260 nm.

Tissue Extraction and Enzyme Assays in Transgenic Plants and Cells.

85

Leaf tissue or cell pellets (~ 0.25 g) were ground with a mortar and pestle and
resuspended in either 300 LIL of GUS buffer (200 mM Trizma-HCL, pH 7.0, 10 mM 13-
mercaptoethanol, 10 mM EDTA, 0.01% (w/v) SDS, and 0.1% (v/V) Triton X-100) or 300
uL of LUC buffer (100 mM potassium phosphate, pH 7.8, 1 mM EDTA, 7 mM 13-
mercaptoethanol, 10 % (v/v) glycerol) and further homogenized on ice. Root tissue (~ 0.2
g) was homogenized and resuspended in IOOuL of either GUS buffer or LUC buffer.
Developing seeds (~ 0.1 g) were homogenized in either 200 LIL of GUS buffer or LUC
buffer. In all cases, cell debris was removed by centrifugation for 15 min at 13,000g at
4°C. The supernatant was transferred to a clean tube and kept on ice. For GUS activity,
50 uL of the sample supernatant were diluted in 400 LIL of GUS buffer and mixed with
50 uL of 10 mM 4-methylumbelliferyl-b-D-glucuronide (Sigma). The reaction was
incubated at 37°C and a 100 uL aliquot was diluted in 0.2 M NazCO3 at time 0 and every
15 min for 60 minutes. UV fluorescence was measured at 365-mn excitation and 455-nm
emission with a Hitachi F -2000 fluorometer. GUS Specific activity is expressed as nmol
methylumbelliferone produced min'1 mg"1 protein. For LUC analyses, a 25 uL aliquot Of
the sample was mixed with 100 uL of LUC reagent (20 mM Tricine, pH 7.8, 5 mM

MgC12, 0.1 mM EDTA, 3.3 mM DTT, 270 uM COA, 500 uM luciferin (Promega), 500
uM ATP). The linear range of the reaction was determined by mixing increasing extract
volumes with 100 LIL Of LUC reagent. Luminescence was measured (3 s delay, 10 5
integration time) in a Tumer TD-20e luminometer. LUC Specific activity is expressed as
light units/mg protein. In all cases the GUS and LUC specific activities and LUC/GUS

ratios were measured in triplicate and independently for each transgenic line. Protein

86

concentration was determined using a protein assay kit (Bio-Rad) with BSA (Sigma) as

the standard.

Determination of FAS inhibition by cerulenin in Arabidopsis cell cultures.

Cells were incubated in the presence of 0, 10, 30 and 100 uM of cerulenin and 25 uCi of
[l-MC] sodium acetate for 0, 2 and 6 hours. Total lipids were extracted by hexane-
isopropanol method and fatty acids transmethylated with 5% (v/v) sulfuric acid in
methanol for 1 hour at 80° C. FAMES were separated by thin layer chromatography
(TLC) with 90/10 (v/v) hexane/diethyl-ether. Radiolabeled bands corresponding to fatty
acids were quantitated by scanning in an Instant Irnager (Packard, Meriden, CT) and

scintillation counting (Beckman Instruments, Fullerton, CA).

Affymetrix GeneChip analysis.

All the experimental protocols were performed according to the Affymetrix GeneChip®
Expression Analysis Manual (http://www.affymetrix.com). Data analysis was performed

with GeneChip® Data Mining Tool Software (Affymetrix, Inc; Santa Clara, CA).

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93

CHAP TER 3

DISRUPTION OF THE FA TB GENE IN ARABIDOPSIS DEMONSTRATES AN
ESSENTIAL ROLE OF SATURATED FATTY ACIDS IN PLANT GROWTH

ABSTRACT

Acyl-ACP thioesterases determine the amount and type of fatty acids that are exported
from the plastids. In order to better understand the role of the FATB class of acyl-ACP
thioesterases we identified an Arabidopsis mutant with a T-DNA insertion in the FA TB
gene. Palmitate (16:0) content of glycerolipids of the mutant was reduced 42% in leaves,
56% in flowers, 48% in roots and 56% in seeds. In addition, stereate (18:0) was reduced
by 50% in leaves and 30% in seeds. The grth rate was reduced in the mutant resulting
in 50 % less fresh weight at 4 weeks compared to wild-type plants. Furthermore, mutant
plants produced seeds with low viability and altered morphology. Analysis of individual
glycerolipids revealed that the fatty acid composition of prokaryotic plastid lipids was
largely unaltered whereas the impact on eukaryotic lipids varied but was particularly
severe for phosphatidylcholine (PC) with more than a 4-fold reduction of 16:0 and 10-
fold of 18:0 levels. The total wax load of fatb-k0 plants was reduced 20 % in leaves and
50% in stems, implicating FATB in the supply of saturated fatty acids for wax
biosynthesis. Analysis of C13 Sphingoid bases derived from 16:0 indicated that, despite a
50 % reduction in exported 16:0, the mutant cells maintained wild-type levels of
Sphingoid bases presumably at the expense of other cell components. Cutin composition
of the mutant was reduced by more than 85 % in CH, derivatives, however its total cutin

load was unaltered compared to wild type. The growth retardation caused by the fatb

94

mutation is enhanced in a fatb-k0 act] double mutant where saturated fatty acid content is
further reduced. Together, the results demonstrate the in vivo role of FATB as a major
determinant of saturated fatty acid synthesis and the essential role of saturates for the
biosynthesis and/or regulation of cellular components critical for plant growth and seed

development.

INTRODUCTION

In plants, de novo fatty acid synthesis in plastids can be terminated either by the action of
plastidial acyltransferases that transfer the acyl group of acyl-ACP to produce
glycerolipids within the plastid (prokaryotic pathway) or by acyl-ACP thioesterases
(FAT) that release free fatty acids and ACP. After export from the plastid, free fatty
acids are re-esterified to coenzyme-A (COA) to form the cytosolic acyl-COA pool, which
is primarily used for glycerolipid biosynthesis at the endoplasmic reticulum (ER)
(eukaryotic pathway) (Figure 4) (Browse and Somerville, 1991). In Arabidopsis leaves,
18:1 and 16:0 are the major products of plastid fatty acid synthesis and approximately 60
% of these products are exported to the cytosol as free fatty acids. In other tissues or plant
Species, flux through the acyl-ACP thioesterase to the eukaryotic pathway is more
predominant with 90 % or greater contribution. Therefore, thioesterases play an essential
role in the partitioning of de novo synthesized fatty acids between the prokaryotic and
eukaryotic pathways. Moreover, thioesterase substrate specificity determines the chain
length and saturation level of fatty acids exported from the plastid (Pollard et al., 1991).

Based on

95

Figure 18 Schematic representation of the enzymatic reaction catalyzed by acyl-
ACP thioesteases (FAT).

Acyl-ACP thioesterases or FAT enzymes (fatty acid thioesterase) hydrolyze the thioester
bond between acyl canier proteins (ACP) and fatty acids to release free ACP and free
fatty acids inside the plastid. Free fatty acids are then exported as acyl-COA molecules.
In most plants the are two classes of FAT enzymes, A and B. The FATA class has
higher in vitro specificity for unsaturated fatty acids (18:1) whereas FATB has more
activity towards saturated fatty acids (14:0, 16:0 and 18:0). The X axis of the histograms
represent the fatty acid attached to ACP to form the acyl-ACP substrate.

96

 

 

 

 

 

 
 

 

 

 

 

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Figure 18

amino acid sequence comparisons and substrate Specificity, two different classes of acyl-
ACP thioesterases have been described in plants (Voelker et al., 1997). The FATA class
has highest in vitro activity for 18:1-ACP and much lower activity for saturated acyl-
ACP substrates. Members of the second class of thioesterases, FATB, prefer saturated
acyl groups but also have activity for unsaturated acyl-ACPS (Figure 18) (Doermann et

al., 1995; Voelker et al., 1997; Salas and Ohlrogge, 2002).

In the Arabidopsis genome, there are two genes for FATA and a single gene for FA TB
(Beisson F., http://plantbiologymsu.edu/gene_survey/front_page.htm). All other higher
plants that have been examined appear to express both classes of thioesterase (Mekhedov
et al., 2000). One salient question is why plants require two classes of acyl-ACP
thioesterase and what individual role does each provide. The major exported fatty acids in
Arabidopsis is 18:1 and based on in vitro activity, it can be predicted that FATA
determines the in vivo levels of 18:1 that move out from the plastid (Salas and Ohlrogge,
2002). In the case of FATB, a previous antisense and over-expression study in
Arabidopsis demonstrated that this enzyme is involved, at least in part, in the in vivo
production of saturates in flowers and seeds (Doermann et al., 2000). Similarly,
downregulation of FA TB expression in soybean also demonstrates partial reduction of
seed palmitic acid (Buhr et al., 2002; Wilson et al., 2001) However, the origin of palmitic
acid which remains following gene Silencing procedures and the extent to which each
class of thioesterase contributes in vivo to the production of exportable fatty acids by

different tissues remains unresolved.

98

One possible role for two thioesterases is to provide control over the
saturated/unsaturated balance of membrane fatty acids. The composition of almost all
plant, animal and microbial membranes contain a mixture of saturated and unsaturated
fatty acids. Such a mixture is believed essential to provide a balance of physical
properties (e.g. fluidity), as well as a method to adapt to changes in environment (e.g.
temperature) and to prevent phase transitions or lateral phase separations which are
promoted by lipids with uniform fatty acid composition. However, as demonstrated by
extensive feeding studies with microbial cells that depend on exogenous fatty acids for
membrane synthesis (Walenga and Lands, 1975; Mc Elhaney, 1989) most organisms can
accommodate a surprising range of fatty acid structures in their membranes without
impairments in growth. Similarly, a wide range of mutations in plant fatty acid
desaturases demonstrate that fatty acid composition of plant membranes can be
considerably altered with no apparent phenotype under normal growth conditions. For
example, Arabidopsis mutants with elimination of 16:1 trans A3, 16:3, 18:3, or large
reductions in 18:2 grow normally at 25°C (Wallis and Browse, 2002). These studies
suggest that within a certain range, the composition of fatty acids in membranes is not
critical as long as a mixture of acyl chains is provided. However, beyond this range, e. g.
under temperature extremes or when major changes in fatty acid composition occur,
growth can be severely impacted. For example, the fab2 mutant of Arabidopsis is
severely reduced in growth due to an increase in18:0 leaf content to 10-15 % (Lightner et
al., 1994), and in the fad2 fad6 double mutant, the complete elimination of
polyunsaturated fatty acids leads to loss of photosynthetic ability (Wallis and Browse,

2002). In addition to a role as bulk components of membranes, where exact structures

99

 

 

seem less critical, some unsaturated fatty acids have more Specialized roles as precursors
for signal molecules (e. g. linolenic for jasmonate) and saturated fatty acids are precursors
for Sphingolipids, surface waxes and cutin, and are involved in protein acylation. Such
roles are clearly vital because mutants completely lacking trienoic acids, for example, are
male sterile and impaired in pathogen defense (Wallis and Browse 2002). To date such

critical roles for saturated fatty acids have not been described.

In this study we describe the isolation and analysis of a mutant disrupted in the FATB
thioesterase gene. Surprisingly, despite only an overall 40-50 % reduction of saturated
fatty acid content, the mutant grew slowly and produced deformed seeds with low
viability. These results have revealed an essential role of FATB in the supply of saturated
fatty acids for the biosynthesis and/or regulation of components vital for plant growth and

seed development.

RESULTS

Mutant Isolation and Complementation Analysis

In an effort to better understand the in vivo functions of the Arabidopsis FA TB acyl-ACP
thioesterase, a PCR-based strategy was used to screen a population of T-DNA tagged
Arabidopsis plants (Sussman et al., 2000) for disruption of the FA TB gene. PCR analysis
of pooled leaf genomic DNA identified a template from plants containing a T-DNA
insertion in the second intron of the FA TB gene (fatb-k0) (Figure 19). Genetic segregation

analysis of heterozygote FA TB T-DNA insertion lines selected one line segregating with

100

the expected ratio (3:1) for a single T-DNA insertion (280:105 BASTA
resistantzsusceptible) (x2 = 0.53, P > 0.4). However, since approximately half of the
homozygous fatb-k0 plants were lost during germination (see below) an expected ratio of
2.5:1 (resistantzsusceptible) better fit the observed ratio (x2 = 0.18, P > 0.6). One hundred
and ten individuals of these 280 resistant plants were grown to full maturity and of these,
25 had a slow-growth phenotype (see below). Again, the observed phenotypic
segregation ratio agreed with the expected 2:0.5 ratio (x2 = 0.2, P > 0.6) of segregation
for a single T-DNA insertion when considering the lower germination rate of the mutant.
A subset of the 110 BASTA resistant plants were randomly selected and subjected to
PCR and GC analysis to determine the genotype and the fatty acid composition,
respectively. All plants with wild-type visual phenotype and fatty acid composition were
heterozygous for the FA TB T-DNA insertion, whereas plants with mutant visual
phenotype and fatty acid composition were homozygous for the same insertion (data not
shown). The confirmation that the visual phenotype and the genetic (T-DNA) and the
biochemical (fatty acid composition) markers all co-segregated suggested that the T-
DNA insertion in the FA TB gene was responsible for both the observed growth and fatty
acid phenotypes. In order to rule out the possibility that a second-site mutation closely
linked to the FA T B T-DNA insertion was responsible for the phenotype, the wild-type
FATB cDNA was inserted under the control of the constitutive CaMV35S promoter and
expressed in homozygous fatb-k0 plants. Transgenic lines resistant to both hygromycin-B
(transgene T-DNA) and BASTA (knockout T-DNA) were indistinguishable from wild-

type and showed normal growth and biochemical characteristics (see below). Therefore,

101

J L-202

(__ T-DNA (5.5 Kb)
LB ‘

 

 

 

 

 

 

 

 

 

5’.. cttgcaccctga attatatgctc.. .3’

Figure 19. Structure of the Arabidopsis FA TB gene carrying the T-DNA insertion.

A, The FA TB gene is composed of 5 exons and the T-DNA is located in the second
intron. The arrows above the gene scheme represent the primers used for the initial
screening of thefatb-knockout plants (F w: forward, Rv: reverse, JL-202: left border of T-
DNA). The arrowheads underneath exons 2 and 3 represent the primers used for mRNA
quantification by real-time PCR. B, Sequence of a portion of intron 2 encompassing the
Site of integration of the T-DNA (arrow)

we conclude that disruption of the FA TB gene is responsible for the phenotypes Observed

in the mutant plants.

FATB mRN A Expression Analysis

The T-DNA was located in the second intron of the FA TB gene and therefore it was
possible that the cells could correctly Splice out at least a fraction of the precursor FATB
RNA to yield mature mRNA. Indeed, by using RT-PCR with a set Of primers that

spanned the second intron of the FA TB gene (see Figure 19), small amounts of correctly

102

spliced mRNA were detected (data not shown). To determine the extent of gene
disruption, the F ATB transcript levels were quantified by real-time PCR in wild-type and
mutant leaf tissue and found to be more than ISO-fold lower in mutant than in Wild-type
(Table 6). Therefore, although PCR could detect correctly spliced FATB mRNA, this
transcript represented less than 0.7 % of wild-type levels. Furthermore, western blot
analysis developed with anti-Arabidopsis FATB antibodies did not detect FATB protein
in the insertion mutant (data not shown). These results indicated that the T-DNA insertion

generated an essentially complete knock-out mutant.

 

Table 6 Relative amounts of FATB mRNA in leaf tissue of wild-type Arabidopsis and
fatb-k0 mutant

 

 

FATB mRNA EIF4A1* mRNA ratio FATB/eIF4A]
C, value 2Ct Ct value 2Cf
WT (Ws) 23.5 i 0.4 1.2 107 21.4 a 0.36 2.8 106 0.24
fatb-k0 (het) 24.3 3: 0.1 1.0 107 21.2 i 0.15 2.3 106 0.23
fatb-k0 (horn) 30.4 a. 0.3 1.4 10" 21.2 a 0.18 2.4 106 0.0016

 

 

 

 

*The eukaryotic protein synthesis initiation factor A1 (eIF4Al) mRNA was used as an internal control
since the levels of this transcript did not differ in leaf tissue of wild-type and fatb-k0 plants. Het and Hom:

heterozygous and homozygous for T-DNA insertion respectively.

FATB is Essential for Normal Seedling Growth

The first visual characteristic offatb-knockout plants was their Size compared to wild-
type (Figure 20A and B). The rosettes offatb-ko plants were approximately half the
diameter of wild-type rosettes during the first weeks of growth at 22°C. In addition, the
bolting time was delayed in the mutant. More than 90% of the wild-type plants bolted
after 4 weeks under our growing conditions, whereas development was delayed in fatb-k0

such that only after the Sixth week did more than 90% offatb-ko plants bolt (Table 7).

103

The morphology of the different organs from mutant plants was unchanged compared to
wild-type. However the stems of the mutant elongated more slowly than wild-type stems.
AS shown in Figure 21A and B, a plot of the fresh weight of the aerial parts of wild-type
and fatb-k0 mutant plants indicated that during the first 4 weeks after germination the
plants grew at a constant rate. However, the rate was slower for the mutant. Results in
Figure 21B (log scale) indicated that wild-type plants increased their fresh weight 10.6-
fold (i 0.4) per week, whereas for the fatb-k0 plants the increase was only 8.8-fold (i
0.6). This 17 % reduction in growth rate led to a reduction of more than 50 % in the fresh
weight of the mutant after 4 weeks (Figure 20A). Growth of both wild-type and mutant
plants slowed after the fourth week, but more so for the former, such that by the sixth
week the fatb-knockout plants differed in size and fresh weight by about 25% (Figure
2 1 A). During the growing phase the percent ratio of dry to fresh weight remained at ~9 %
while during the drying period it increased to ~14 % for both wild-type and mutant
plants. Taken together, the similar morphology but different growth rates suggested that
differences in wild-type and mutant plants were the consequence of a reduced grth rate
and not altered development of the mutant. Wild-type and fatb-k0 plants were grown in
the presence of l % sucrose on either plates or liquid culture to determine if the normal
growth rate could be recovered. Sucrose availability did not eliminate the slower growth
rate of the mutant (data not shown) suggesting that photosynthetic capacity or carbon

limitations of the fatb-knockout plants were not causes of the reduced growth rate.

104

 

Figure 20. Growth and morphology of Arabidopsis wild-type and fatb-k0 plants and
seeds. A, Four weeks old wild-type (left) and fatb-k0 (right) plants. B, Two weeks Old
wild-type (left) and fatb-k0 (right) plants. C, wild-type Arabidopsis seed. D, E and F,
wild-type-like, intermediate deformed and very deformed seeds from fatb-k0 plants
respectively.

105

Plant grth rate is modified by temperature and part of this effect may be associated
with variations in the physical properties of cell membranes. To test the possibility that
the growth retardation of the fatb-k0 mutant was a function of temperature, wild-type and
fatb-k0 plants were grown for 2 weeks at 22°C and then transferred to three different
temperatures (16°C, 22°C and 36°C). The fatb-k0 plants Showed the same percent
reduction (~50 %) of fresh weight per seedling compared to wild-type at the three
different temperatures. Therefore, the slower relative growth of the mutant plants was not
altered within the range of temperatures used. In addition we tested wether adding
exogenous saturated fatty acids by either spraying plants or supplementing seedlings
grown in liquid culture could overcome the slower growth of the fatb-k0 mutant. These
procedures were not sufficient to chemically complement the fatb-k0 phenotype. In fact,
addition of higher amounts of exogenous fatty acids showed deleterious effects on plant

growth (data not Shown).

Leaf and chloroplast morphology of wild type Arabidopsis and fab-k0

The leaves offatb-ko average 50 % of the area and fresh weight of wild type leaves
(Figure 20 and data not shown). These differences may be brought about by reduced cell
Size or less number of cells per leaf. In addition, leaves are composed of different cell
types (e.g., mesophyll, epidermal) and some of those might be differentially affected by
the genetic lesion. We analyzed optical sections of leaf tissue from wild type and mutant
plants by laser confocal microscopy (LCM) and leaf epidermal cells by scanning electron

microscopy (SEM) (Figure 22A to D). The structures of leaves and epidermis together

106

 
 
 
  

if A
:1 3-5; Wild type
'6 l
s 31
m
E 2.5..
a 2 l
"' 1 51 ~
on ‘1 . fatb-k0

0.5'

O " ' r +—#+-—-~*—+~*l 4' t t ~ws~r—+—~t
1 2 3 4 5 6 7 8
weeks after germination
10 Wild type

B

0.1

0.01

 

grams/seedling (log scale)

0.001

 

weeks after germination

Figure 21. Growth curves of Arabidopsis wild-type and fatb-k0 plants.

A, The fresh weight of aerial parts of Arabidopsis plants was measured on a weekly basis
for a period of 8 weeks. Each time point is the average value of at least 7 individual
plants. B, Logarithm of the fresh weight versus time.

with Sizes of epidermal and mesophyll (palisade and spongy) cells were similar between
both plant classes (Figure 22A to D). Thus, these observations indicated that the smaller
size offatb-ko leaves compared to wild type was the result of fewer cells per leaf rather
than smaller cell Sizes. These results contrasted with those Obtained with the Arabidopsis

fab2 mutant, in which decrease in leaf size was the consequence of reduced

107

 

Figure 22. Microscopic analysis of leaf section and chloroplast structure of wild
type and fatb-k0 seedlings. A—B, Scanning electron micrographs of wild type (A) and
fatb-k0 (B) leaf epidermal cells. C-D, leaf sections of wild type (C) and fatb-k0 (D). E-F,
Transmission electron microscopy (TEM) of wild type (E) and fatb-k0 (F) leaf
chloroplasts. G-H, TEM of thylakoid membranes from wild type (G) and fatb-k0 (H) leaf
chloroplasts.

108

expansion of leaf cells (Lightner et al., 1994).

Growth of fatb-k0 in the presence of sucrose did not revert its growth phenotype,
suggesting that photosynthetic capabilities of the mutant did not differ substantially from
wild type plants. Analysis of leaf chloroplast ultrastructure by transmission electron
microscopy (TEM) Showed no obvious differences between organelles of wild type and
fatb-k0 (Figure 22E and F). The size and number of chloroplasts per cell of the mutant
were Similar to wild type. Moreover, structure and number of thylakoid membranes were
also Similar between the two plant classes (Figure 22G and H). In summary, these
observations agreed with similar photosynthetic capabilities between wild type and

mutant plants.

FATB is Essential for Normal Seed Morphology and Germination

Germination of seeds produced by fatb—k0 plants was reduced by approximately 50 %
either on soil or 1 % sucrose (Table 7). Close examination of mature seeds produced by
the fatb-k0 mutant revealed a continuous range of deformity in seed morphology, with
wild-type-like seeds in one extreme to very deformed seeds (approximate fi'equency of 20
%) in the other (Figure 20C-F and Table 7). The germination rate of very deformed seeds
was only 16 %. These observations suggested that some stage of seed or embryo
development may be substantially affected in the mutant. By analyzing developing
siliques it was not evident that deformed seeds were located in Specific segments of this

organ. Upon surface sterilization, some mutant seeds also lost the seed coat, suggesting

109

 

 

Table 7. Arabidopsis wild-type and fatb-k0
bolting time and germination rates.
Bolting time* (%)

 

 

Time WT (WS) fatb-k0
4th week 92.3 27.6
2 5‘h week 7.7 72.4

 

Germination rate (%)

 

WT (WS) fatb-k0
Soil 96.7 45 .6
1% sucrose 94.4 45.8

 

Deformed seeds in total seeds (%)
WT (WS) fatb-k0
0.8 21.7

 

 

Germination on soil of deformed seeds (%)
fatb-k0
16

 

 

* 18:6 h lightzdark; 22 C

 

that the structure of this tissue could be altered. Scanning electron microscopy (SEM)
analysis of the seed coat from mutant plants with wild-type-like or intermediate
morphology did not Show any obvious structural differences compared to wild-type seeds
(Figure 20C-F). Many deformed seeds from the mutant displayed a shriveled seed coat

(Figure 20F).

Fatty Acid Composition of fatb-k0 Tissues.

110

_-.I_IE|

1.3;..1‘

AS indicated in Table 8, palmitate (16:0) in homozygous fatb-k0 plants was reduced 42 %
in leaves, 56% in flowers, 48% in roots and 56% in seeds compared to wild-type.
Stearate (18:0) decreased almost 50 % in leaves and 30 % in seeds, with negligible
changes in flowers and roots. The fatb-k0 plants also showed an increase of 150-200 %
in oleate (18:1) and 40-60 % in linoleate (18:2) in leaves, flowers and roots. Linolenate
(18:3) declined by 15-20 % in leaves, flowers and roots. Seed unsaturated fatty acids
were less affected. Together, these results demonstrate the in vivo role of FATB as a
major determinant of 16:0 in all the tissues analyzed and also indicate that FATB
contributes to the level of 18:0 in leaves and seeds. fatb-k0 plants transformed with the
wild-type FATB cDNA under the CaMV35S constitutive promoter had a fatty acid
composition very similar to wild type, confirming that the FATB cDNA complemented

the biochemical phenotype of the mutant (Table 8).

 

Table 8. Fatty acid composition of wild-type and fatb-k0 Arabidopsis tissues (mol%).
16:0 16: 1(9) 1621(3) 16:3 18:0 18:1(9) 18:2 18:3 Other

 

 

 

 

 

Leaftissue
WT (WS) 17.5 0.6 3.7 11.8 1.5 2.6 14.4 47.7
fatb-k0 10.1 1.3 3.4 12.4 0.8 8.2 23.2 40.1
3SS-FATB** 15.5 0.6 4.0 11.9 1.3 4.6 16.8 45.0
Flowers
WT (WS) 27.4 1.7 1.2 2.4 3.2 30.6 31.3
fatb-k0 12.0 4.6 2.6 2.6 8.5 41.9 24.7
Roots
WT (WS) 25.3 0.9 6.6 3.6 28.8 21.3 13.2
fatb-k0 13.4 1.1 6.0 9.2 42.5 17.4 10.1
Seeds
WT (WS) 8.3 0.3 3.5 13.0 27.5 18.6 28.6
fatb-k0 3.6 0.4 2.4 11.9 33.1 16.2 32.1

 

* Average of at least 3 samples. Standard deviation values are not shown and represent less than 5% of

the average values. ** fatb-k0 plants transformed with CaMV35S-FATB cDNA.

 

Ill

Fatty Acid Composition of Individual Leaf Glycerolipids.

The fatty acid composition of individual glycerolipids from homozygous fatb-k0 and
wild-type leaves are presented in Table 9. Palmitate reductions occurred mainly in extra-
plastid lipids. While phosphatidylethanolamine (PE), phosphatidylserine (PS) and
phosphatidylinisitol (PI) had an approximately 50 % reduction in 16:0 compared to wild
type, in phosphatidylcholine (PC) the reduction was almost 80 %. The palmitate levels in
plastid lipids were less affected with a significant reduction (40 %) only in
sulfoquinovosyldiacylglycerol (SQDG). All of the extra-plastid glycerolipids except PI
had reduced 18:0. Again PC was the most affected with a 10-fold reduction in 18:0. The
characteristic changes in unsaturated fatty acids seen in Table 8 for total leaf lipids,
namely increases in 18:1 and 18:2, and a decrease in 18:3, were most pronounced for the
phospholipids and SQDG. The data in Table 9 also indicated that despite the changes in
fatty acid composition there were no major differences in the relative proportions of leaf
glycerolipids between wild—type and fatb-k0 plants. And finally, the total amount of fatty
acid methyl esters produced by acid-catalyzed transmethylation of Arabidopsis leaves
was the same for the wild-type (11.5 umoleS/gfw) and fatb-k0 (11.6 umoles/gfw),

indicating that the fatb-k0 did not affect net fatty acid accumulation per fresh weight.

AcyI-ACP Thioesterase Activity

To determine if any compensatory changes occurred in acyl-ACP thioesterase activity,
mutant and wild-type plants were assayed for hydrolysis of 18:1-ACP and 16:0-ACP.

The FA TA gene product has an acyl specificity 18:1 >> 18:0 >> 16:0 while the FA TB

112

 

Table 9. Fatty acid composition of leaf glycerolipids of Arabidopsis wild-type (WS)
and fatb-k0 mutant (mol%).
% total 16:0 1621(3) 16:3 18:0 18:1(9) 18:1(11) 18:2 18:3

 

PC
WT (WS) 11.2 i 0.3 21.1 2.4 6.1 - 34.7 35.5
fatb-k0 13.6 e 0.7 4.5 0.2 17.8 1.3 46.1 30.1
PE
WT (WS) 10.0 i 0.2 29.0 2.1 2.8 - 35.1 30.8
fatb-Ito 7.9 :r 0.6 11.6 1.1 11.3 1.2 50.8 24.9
PS
WT (WS) 0.9 i 0.1 30.8 6.8 2.9 27.1 32.3
fatb-k0 0.8 i 0.1 17.2 2.9 10.2 44.3 25.2
PI
WT (WS) 5.5 e 0.1 40.0 2.0 2.2 25.3 30.3
fatb-k0 4.4 i 0.1 21.2 1.8 9.2 36.9 29.3
PG
WT (WS) 12.0 i 0.3 29.2 22.0 1.1 4.5 - 10.8 32.1
fatb-k0 12.6 i 0.3 24.3 20.7 - 8.5 0.9 16.9 27.6
SQDG
WT (WS) 4.3 i 0.1 34.6 1.5 3.6 0.7 21.5 37.8
fatb-k0 4.2 i 0.6 20.4 0.5 10.5 1.2 36.4 30.8
DGDG
WT (WS) 20.2 e 0.3 12.8 4.2 1.0 1.1 - 6.7 73.9
fatb-k0 19.4 e 0.1 12.0 5.4 0.5 1.2 0.5 7.9 70.5
MGDG
WT (WS) 35.5 e 0.1 1.6 38.1 0.8 3.0 53.4
fatb-k0 36.7 i 0.2 1.3 40.3 1.6 3.2 50.7

 

 

 

gene product has a specificity 16:0 > 18:1 > 18:0 (Doermann et al., 1995; Salas and
Ohlrogge, 2002). The activity from the FA TA gene product dominates acyl-ACP

thioesterase activity measurements made With crude extracts. OleoyI-ACP hydrolytic

113

activity in leaf extracts of wild-type and mutant plants was similar (~l25 pmol*min'
1"‘mg") and hydrolytic activity on 16:0-ACP was close to background levels and
therefore difficult to quantify. These results indicate that measurable acyl-ACP hydrolytic
activity does not change in mutant leaf extracts compared to wild-type and therefore

endogenous levels of FATA activity are not up-regulated in the mutant.

Total Palmitate Content in Arabidopsis Leaf Tissue

Most acid or base-catalyzed transmethylation methods for fatty acid analysis efficiently
convert O-acyl groups such as found in glycerolipids to fatty acid methyl esters.
However, N-acyl groups such as found in Sphingolipids react only very slowly using such
methods. In order to evaluate the total palmitic and stearic content (0- and N-linked) of
Arabidopsis cells, a strong alkaline hydrolysis on total leaf tissue, on lipids extracted with
multiple chloroform methanol extractions, and on the solvent-extracted residue was
performed. The total 16:0 content in leaves of wild-type plants was 1.87 i 0.02
umoles/gfw, whereas leaves offatb-ko plants contained 1.14 i 0.03 umoles/gfw. Thus, a
39 % reduction in total 16:0 was observed in the mutant, similar to the 42 % reduction of
16:0 in glycerolipids (Table 8). The amount of total stearic acid was also reduced by 50

% in leaf tissue, from 0.16 i 0.01 umoles/gfw in wild-type plants to 0.075 i 0.005

umole'S/gfw in the mutant. The analysis of extracted lipids and solvent-extracted residue
indicated that almost all the 16:0 and 18:0 in the cells was co-extractable from leaf tissue,
since the lipid fraction contained similar amounts of saturated fatty acids to the total
tissue (data not shown). In contrast, the solvent-extracted residue which may contain

acylated proteins and other insoluble lipids, contained 3 % of the total 16:0 and no

114

detectable 18:0. Similar reductions of 16:0 and 18:0 were observed in all the fractions
analyzed, indicating that absence of FATB reduced saturated fatty acid levels in both

organic soluble and insoluble components.

Leaf and Stem Surface Wax Analysis

The very long chain fatty acids (VLCFA) required for wax synthesis are produced by
elongation of 16 and 18 carbon saturated fatty acids (Post-Beittenmiller, 1996). To
determine if the 40-50 % reduction in saturated fatty acids influences total wax load or
composition in the fatb-k0 mutant, leaf and stem epicuticular waxes were analyzed. The
results in Table 10 indicated that at 5 weeks total wax load per fresh weight in leaves was
reduced by 20 % in the fatb-k0 mutant. However, no novel components or substantial
changes in the distribution of wax components were observed at this stage. A 20 %
reduction in leaf wax load was consistently observed at different stages of plant
development (data not Shown). Analysis of primary stems indicated a 50 % reduction in
wax load per fresh weight in the fatb-k0 mutant compared to wild type (1.1 i 0.1
umol/gfw versus 2.2 i 0.3 umOl/gfw respectively), again without changes in the
distribution of wax components. These data indicate that supply of saturated fatty acids
by FATB is one factor limiting wax biosynthesis but that reduction of this supply does
not result in the replacement of 16:0 by 18:1 or other precursors for surface wax
structures. The role of FATB in supplying wax precursors was more evident in stems
than in leaves as the former tissue accumulates higher amounts of epicuticular waxes.
Similar tissue specific wax reductions are observed in most of the Arabidopsis wax

biosynthetic mutants (eceriferum) in which reductions in stem waxes are larger than in

115

 

 

Table 10. Major components of epicuticular leaf waxes
from wild—type ArabidJorsiS and fatb-k0 mutant (mol%).

 

 

 

(carbon chain length) WT (WS) fatb-k0
Alkanes
27 1.45.0.1 l.2:l:0.05
29 21.93:].2 22.1:E0.l
30 1.23:0] 1.0:t0.03
31 39.4 i 1.1 40.5 :1: 0.5
32 0.9 i 0.1 0.9 i 0.03
33 11.7 i 0.4 12.7 :t 0.03
Free fatty acids
24 3.5 i 0.2 3.3 i 0.06
26 5.7 i 0.2 5.3 i 0.09
28 2.2 31:04 1.8101
30 0.7 d: 0.1 0.5 i 0.1
Primary alcohols
26 3.1 i 0.4 2.3 i 0.3
28 3.6i 0.5 3.63:0.3
30 2.2 i 0.1 2.2 i 0.02
32 2.3 3: 0.1 2.2 :1: 0.02
Total (pg/gm) 104.1 A 6.6 83.9 :1: 4.5
Total (umol/gfw) 0.24 :t 0.01 0.19 i 0.01

 

leaf waxes (Rashotte et al., 2001).

Sphingoid Base Analysis

Since sphingoid base and Sphingolipid synthesis are initiated by serine
palmitoyltransferase (Lynch, 1993) an analysis of sphingoid bases was conducted.
Leaves of wild-type (0.54 i 0.09 umoles/gfw) and fatb-k0 (0.50 i 0.08 umoles/gfw)

plants did not differ significantly in the total amount of sphingoid bases. However,

116

differences could be observed in the relative abundance of the individual sphingoid bases
(Table 11). Most significantly, trihydroxy-18:0 (118:0) increased by almost 4-fold in the
total Sphingoid bases of the fatb-k0 mutant. The total sphingoid base composition was
similar to that reported by Sperling et al. (1998): the most abundant sphingoid base in
Arabidopsis leaf tissue was t18:1(8E), followed by t18:1(8Z). Monoglucosylceramide
(MGC) is considered one of the most abundant Sphingolipids and therefore the sphingoid
base composition in leaf MGC was analyzed. The base composition in MGC of wild-type
and mutant plants was indistinguishable, and was Similar to that reported by Imai et al
(2000) (Table 11). The difference in composition between total and MGC sphingoid base
compositions indicates that MGC is not the predominant Sphingolipid in Arabidopsis
leaves. As suggested by Imai et al. (2000), complex Sphingolipids such as
phosphoinositolceramides could be more abundant than MGC in Arabidopsis leaf tissue.
Sphingoid bases in extracted lipids and solvent-extracted residue were also analyzed. The
latter fraction can contain sphingoid bases from highly glycosylated
phophoinositolceramides and GPI moieties from GPI-anchored proteins. Again, no major
changes in sphingoid base composition between Wild-type and fatb-k0 plants were

observed except in trihydroxy 18:0 (Table l 1).

Leaf and stem cutin analysis

The monomers of cutin are synthesized from the COA esters of palmitic acid (16:0) and
oleic acid (18:1), respectively. To determine if the reduction in saturated fatty acids in
fatb-k0 influences cutin composition, leaf and stem cutin were analyzed. The results

shown in Figure 23 indicate that 16 carbon derivatives of cutin were reduced by more

117

 

Table 11. Sphingoid base content of leaf tissue from wild-type Arabidopsis and fatb-k0
plants (mol%)

 

118:1(8E) t18:1(8Z) 118:0 d18:1(8E) d18:1(8Z) d18:2(4,8)

 

Total Tissue

WT (WS) 60 32 2 4 t 2

fatb-k0 57 32 8 2 l 1
Monoglucosylceramide

WT (WS) 24 59 t 11 t 5

fatb-ko 25 59 t 1 0 2 5

Solvent Extracted Lipids
WT (WS) 62 33 2 2 1 t
fatb-k0 57 36 5 2 t t
Solven t-Extracted Residue
WT (WS) 85 7 3 t 3 l

fatb-k0 76 7 13 t 3 1

 

Dihydroxy bases: 8-sphingenine (d18:1[8E or Z]), 4,8-sphingadienine (d18:2[4E,EZ or
4E,8Z]); trihydroxy bases: 4-hydroxysphinganine (t18:0) and 4-hydroxy-8-sphingenine
(t18:1[8E or 2]). t, traces.

 

than 85 % in fatb-k0 leaves and stems compared to wild type. These results
demonstrated that FATB provides 16:0 for cutin biosynthesis in Arabidopsis. Although
fatb-k0 has reduced 16 carbon derivatives in cutin the total cutin load was unaffected in
the mutant (approximately 5 umol/g residue in leaves and stems of both plant classes).
Thus, the mutant compensates for the decrease in CH, derivatives by producing more C18

derivatives (Figure 23).

118

fatb-k0 act] double mutant

The lipid analysis demonstrated that despite the absence of FATB in the plastids, mutant
plants contain approximately 50 % of the saturated fatty acids found in wild-type plants.
What is the origin of these remaining saturated fatty acids? Palmitate and stereate, as
16:0-ACP and 18:0-ACP respectively, may be utilized directly by acyltransferases in the
plastid for prokaryotic lipid synthesis (Somerville et al., 2000). In order to evaluate how
much of the saturated fatty acids remaining in the fatb-k0 plants derive from acyl group
fluxes through plastidial acyltransferases, a cross between fatb-k0 plants and the
Arabidopsis act] (atsl) mutant was performed. The act] mutant has reduced plastidial
glycerol-3-phosphate:acyl-ACP acyltransferase activity, the first step in the plastid
pathway of glycerolipid biosynthesis. Although act] plants contain reduced amounts of
16:3 and higher amounts of 18:1, grth of this mutant is normal (Kunst et al., 1988).
However, the fatb-k0 act] double mutant was severely impaired in growth when
compared to wild-type,fatb—ko and act] plants (Figure 24). Leaf fatty acid analysis of the
double mutant indicated that this tissue contained approximately 3-4 mol% 16:0,
representing a 70 % reduction compared to wild type (Table 8 and 12). The levels of 18:1
in leaves were increased in the double mutant compared to fatb-k0 whereas 18:2 and 18:3
levels remained almost unchanged (Table 8 and 12). Interestingly, leaf levels of 18:0 in
the double mutant were Similar to those in fatb-k0 leaves, indicating no further decrease
in overall 18:0 by the second mutation. Analysis of individual lipid classes from leaf
tissue of the double mutant revealed that the C H, acyl composition of extraplastidial

glycerolipids and PG did not differ substantially from those in fatb-k0 leaves (data not

119

A Composition of leaf cutin

 

 

 

f Hf _.
80 i .WS 1
70 . i
60 Ufatb-ko l
50 -
.\°
T: 40
E
30 ..
20 .
‘° I. ll
0 I: LLtLj:
I I 7 I 7 I I 7
[069 761/0676: ’IKO’Q/OIIKO .90 0,0
760 4); [04) 6); 6’7 7 If ’Q/
o o
‘90 76106-13 ‘97 (5)0 ’39 4),
7 (20
B Composition of stem cutin
80 I“ —
. .WS
70 l
60 - Qfatb-ko
50 . ‘ 7 "i’
o\° l
a 40 l
E l
30
20
A . l -. [1112.6
7 7 I I I 7 7
70%? *5? Yo o’e’e’Q 9
’2, F0 761/ "a ’59, Yo 70 ’Q, ’0
6.0/302,3 ,2’962 <2
’6‘. 7 76>. ’29 {O {O
.0 .9 , ’59, A»,
‘9'; 63.0

Figure 23. Leaf and stem cutin composition of wild type Arabidopsis and fatb-k0
(mol%).

Cutin components were reduced with LiAlH4 and analyzed by GLC. A, cutin monomer
composition of four-week old leaves from wild type and fatb-k0 plants. B, cutin
monomer composition of six-week old stems from wild type and fatb-k0 plants. Bars
denote standard deviations.

120

shown). Moreover, the relative abundance of each glycerolipid was similar. These results
demonstrated that the second mutation (actI) affected primarily 16:0 accumulation in

plastidial glycerolipids with minor effects on extraplastidial lipids compared to fatb-k0.

In summary, double mutant plants blocked in both F ATB and ACTI were further reduced
in saturated fatty acid content to levels ~ 30 % of wild-type plants. These plants displayed
a smaller size and thus this more severe grth phenotype associated with a greater
reduction in saturated fatty acids further demonstrates the essential role of saturated fatty

acids in maintaining normal rates of plant growth.

DISCUSSION

Acyl-ACP thioesterases are responsible for the export from the plastid of fatty acids
produced by the de novo fatty acid synthesis system. In this study an Arabidopsis
insertion mutant of the FA TB gene was isolated,‘and we describe its effects on plant
growth and on the production and utilization of saturated fatty acids. In a previous study,
antisense of Arabidopsis FATB using the CaMV3SS promoter resulted in a substantial
reduction of 16:0 only in flowers and seeds and minimal difference in leaves and roots
and did not Show any visual phenotype (Doermann et al., 2000). The lack of anti-sense
impact on leaves suggested either FATB was not the major controller of 16:0 in leaves or
that the reduction of FATB mRNA was not sufficient to reduce 16:0 levels in certain
tissues. The characterization of an Arabidopsis fatb-knockout mutant in this study

clarifies that the second interpretation is correct and that FATB is a major control point

121

 

Figure 24. Growth and morphology of Arabidopsis wild-type,fatb-ko, act] and fatb-
ko act] plants.

Four weeks Old wild-type (WS) (bottom right), fatb-k0 (bottom left), act] (top right) and
fatb-k0 act] double mutant (top left) plants. The size of act] mutants is similar to the size
of wild type Columbia (Col) plants (not shown). The size of wild type plants derived
from the cross between fatb-k0 (WS) and act] (C01) is intermediate to the parental
ecotypes (not shown).

for saturated fatty acid fluxes in all tissues. The more extensive biochemical phenotype
and the reduced growth rate and seed viability observed in the mutant compared to

antisense plants suggest that the FATB enzyme or its mRNA may be in large excess and

122

 

Table 12. Leaf fatty acid composition of wild type Arabidopsis (Columbia), act] and
fatb-k0 act] double mutant (mol%)
16:0 16:1 16:1(3) 16:3 18:0 18:1(9) 18:2 18:3

Wild type 16.2 1.4 3.0 11.3 1.5 3.0 13.8 49.8
(Col)

act] 14.3 1.1 3.1 0.4 1.6 11.9 24.1 43.1

fatb-k0 act] 3.1 1.2 3.2 0.5 0.8 23.4 24.8 41.5

 

 

difficult to reduce sufficiently by antisense methods to produce a growth phenotype.

Seedling Growth and Seed Development

A large number of mutants with diverse changes in fatty acid composition have been
isolated in Arabidopsis (Wallis and Browse, 2002). Most of these are not readily
distinguishable from wild-type plants when grown at standard temperatures (15°C-25°C).
One of the few examples in which a mutation affecting fatty acid composition has
consequences for plant growth is the fab? mutant in Arabidopsis, in which elevated
levels of 18:0 in the membrane lipids results in dwarf plants (Lightner et al., 1994).
However, the phenotype can be partially ameliorated by growing the mutant at high
temperature, suggesting that membrane fluidity or a related physical property reduces

growth of the mutant at 22°C.

The Arabidopsis fatb-knockout is the first example of an Arabidopsis mutant with
reduced levels of saturated fatty acids where a reduction in vegetative growth at standard

growth conditions occurred. Low temperature did not alleviate nor did high temperature

123

exacerbate the Slow growth phenotype offatb-ko plants suggesting that effects other than
changes in bulk membrane physical properties were limiting the growth of the mutant.
Hence, in contrast with fab2 in which high 18:0 levels may disrupt the proper function of
membranes, reduced saturate levels in fatb-k0 plants may be altering the biosynthesis and
function of critical cell components. However, we can not rule out that lower amounts of
saturates may be associated with more subtle changes in the physical properties of
cellular membranes that could affect functions such as transport or vesicle formation.
The fatb-k0 mutant with a ca. 50% reduction in palmitate also contrasts with the fab]
mutant that is increased approximately 50% in palmitate but displays normal growth (Wu
et al., 1994). The fatb-k0 mutant is also distinguished from other fatty acid mutants in its
effect on seed development and germination (Figure 20C-F and Table 7). However, in
contrast to the Slow growth phenotype that occurred in all seedlings, the penetrance of the
seed phenotype was incomplete. At this stage it is not clear whether the seed defects are a
consequence of alterations during specific seed developmental phases, or perhaps an

indirect effect due to deficiencies in supply of nutrients from maternal tissues.

Palmitoyl-ACP pools in the plastid are subject to three major reactions; acyltransfer to
glycerol, elongation to 18:0-ACP or hydrolysis by FATB. Mutations that block or reduce
all three of these fates are now available. In the fab] mutant (Wu et al., 1994), reduction
in 16:0-ACP elongation results in increased 16:0 in both the prokaryotic and eukaryotic
lipids suggesting that flux into both these pathways can be increased by increased
availability of 16:0-ACP. Moreover, in both fatb-k0 and fab] leaves there is an increase

in 16:1(9) levels, suggesting that 16:0-ACP pools are increased within chloroplasts of

124

these two mutants. In contrast, in the act] mutant the loss of the acyltransferase pathway
results in increased 16:0-ACP elongation to 18:0 rather than increased flux to the
eukaryotic path via the FATB thioesterase (Kunst et al., 1989). Similarly, in the fatb-k0,
the reduction in flux via the thioesterase also primarily increased elongation to C18 rather
than increased flux of C K, into prokaryotic lipids. These contrasting responses suggest
that the elongation rate of 16:0-ACP is regulated primarily by availability of substrate but
that the contributions of the FATB and acyltransferase reactions to 16:0 flux likely have

additional levels of control.

Reduced Export of Palmitate in fatb-k0 plants and Other Sources of Palmitate and
Stearate in the Cell

Lipid analysis (Table 8) demonstrated that despite the homozygous fatb-k0 insertion,
mutant plants still produce approximately 50 % of the palmitate found in wild-type
plants. How and where is the remaining palmitate balance in a plant cell produced? A
Similar statement and question can be made for stearate. Palmitate is both an intermediate
and an end product of de novo fatty acid synthesis in the plastid. Palmitate, as 16:0-ACP,
may be utilized directly by acyltransferases in the plastid for prokaryotic lipid synthesis,
in particular by the lyso-phosphatidic acid sn-2 acyltransferase (Frentzen et al., 1983).
Alternatively, after hydrolysis by acyl-ACP thioesterases, free fatty acids including
palmitic acid may be exported from the plastid, a proposed mechanism now substantiated
by in vivo labeling (Pollard and Ohlrogge, 1999). And finally, plant mitochondria have
the capacity for de novo synthesis of fatty acids (Wada et al., 1997) and, although
considered a minor pathway this organelle could partially compensate for low 16:0 levels

in the fatb mutant. Could (I) the transfer of palmitate from prokaryotic lipids to

125

eukaryotic lipids, (2) the FATA acyl-ACP thioesterase, and/or (3) mitochondrial fatty

acid synthesis account for the remaining exported palmitate production?

The cross offatb-ko with act] plants demonstrates that the prokaryotic pathway provides
about 60 % of the saturated fatty acids in leaves offatb-ko. Approximately half of the
saturates that are still produced in the double mutant can be attributed to plastidial PG
(produced with prokaryotic character by an unknown path) and to FATA activity. The
Arabidopsis FATA-encoded thioesterase has a small but measurable in vitro activity
towards 16:0 and 18:0-ACP (about 2 % and 16 % of the activity towards 18:1-ACP,
respectively) and our results suggest that these enzymes have in vivo hydrolytic activity
towards 16:0- and 18:0-ACP, a conclusion also drawn by Nadev et al. (1992). The
mutants studied here demonstrate that FATA, mitochondrial FAS or other sources of 16:0
are minor contributors to leaf saturated fatty acid flux in comparison to FATB and the

plastid acyltransferases.

As summarized in Figure 25, the results of this study allow a better estimate of the
relative contributions of alternative pathways for saturated fatty acid supply in plants. In
wild-type plants, the total C16 fatty acids incorporated into membrane glycerolipids is
33.3 units, which includes 16:1(3)-trans in PG and 16:2 and 16:3 in MGDG. About 23
units are used for prokaryotic lipid synthesis, assuming that all PG and MGDG are
derived from this pathway, and that the proportion of 16:0-containing DGDG that is
prokaryotic is one third, based on its sn-2 distribution (Kunst et al., 1989). Assuming that

half the SQDG is of prokaryotic origin, of the remaining 10.3 units which are exported

126

from the plastid, about 2.5 units return to this organelle as SQDG and DGDG while the
remaining 7.8 units are used for phospholipid synthesis (Figure 25). In the fatb-k0 line
we do not know what proportion of the 16:0-containing DGDG and SQDG pools are now
of prokaryotic origin, so estimates are ranges. About 2.6-4.2 units of palmitate (25-45 %
of wild-type) are exported and 2.6 units (33 % of wild-type) are retained for
phospholipid synthesis while 0-1.6 units (0-65 % of wild type) are returned for plastid
lipid synthesis. Palmitate used for prokaryotic lipid synthesis is barely affected by the
mutation, increasing from 23 to 23.6-25.6 units. In the fatb-k0 act] double mutant, about
3.5 units of palmitate (35 % of wild type) are exported and 2.5 units (33 % of wild type)
are retained for phospholipid synthesis while 0-1 units (0-40 % wild type) are returned to

the plastid.

Partition of Palmitate and Stearate to Non-glycerolipid Products.

Palmitate constitutes the primary saturated fatty acid exported from plastids and
incorporated into membrane glycerolipids. In addition to glycerolipids, several other
cellular components derived from the exported palmitate and/or stearate have essential
structural and perhaps signaling roles for cell growth. First, sphingoid bases in plants are
18 carbon amino alcohols that are synthesized outside the plastid from palmitoyl-COA

and serine by the action of serine palmitoyltransferases (Lynch, 1993). In addition, 16:0

127

Figure 25. Simplified scheme of predicted fluxes of C16 and C13 fatty acids in
membrane leaf glycerolipids of Arabidopsis wild type, fatb-k0 and fatb-k0 act]
mutants.

The numbers in the figure represent average values of mol% units of C16 and C13 fatty
acids accumulated in membrane glycerolipids. A range of values is given when
alternative biosynthetic pathways are considered (see text for details). The average mol%
units were calculated based on the mol% abundance of a particular fatty acid in a Specific
lipid, the relative abundance of the lipid and the contribution of each pathway
(prokaryotic and eukaryotic) to the synthesis of that lipid (Browse et al., 1986) (see
Discussion for more details). Arrows represent the flux of acyl molecules through FATB
and FATA (acyl-ACP thioesterases B and A respectively), KAS-II (3-ketoacyl-ACP
synthase-II) and LPAAT (lyso-phosphatidate sn-2 acyltransferase) that are either used for
lipid biosynthesis in the plastid (prokaryotic pathway) or exported from the same
organelle. ACTl (glycerol-3-phosphate acyltransferase) transfers 18:1 to glycerol-3-P in
the prokaryotic pathway. Question marks indicate that: l, the actual flux of 16:0 through
FATA in wild type leaves is not known and an upper limit of 4 mol% units can be
estimated based on fatb-k0 data (however, flux through FATA in fatb-k0 could be a
compensatory mechanism); and 2, alternative pathways for biosynthesis of PG in the
fatb-k0 act] double mutant can be considered.

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129

Figure 25

or its hydroxylated derivatives can be N-linked to sphingoid bases to form ceramides and
sphingolipids. Second, in epidermal cells, saturates (16:0 and 18:0) are precursors of the
cutin/suberin monomers and wax components. (Post-Beittenmiller,1996). Third,
myristoylation (14:0) and palmitoylation (16:0) of proteins is critical for the localization

and regulation of protein activity (Yalovsky et al., 1999).

In wild-type leaves the total 16:0 content is about 1.9 umoles/gfw. Deducting the
contribution from prokaryotic lipid synthesis, about 1.2 umoles/gfw of 16:0 must be
exported from the plastid to fuel glycerolipid synthesis. Sphingolipid base synthesis
requires another 0.5 umoles/gfw of palmitate export. In addition, there are significant
levels of 16:0 and 2-hydroxy-1620 N-acyl groups in sphingolipids, so the total flux of
palmitate into Sphingolipids is actually higher. Thus Sphingolipid synthesis may
consume 30-40 % of the total cytosolic palmitate pool. However, despite a reduction of
about 50 % in extraplastidial 16:0 in the fatb mutant, the total amount per fresh weight of
sphingoid bases in leaf tissue was similar to wild-type. One interpretation of the
constancy of Sphingolipid production is that sphingoid base synthesis is tightly
maintained at the expense of acyl composition changes in other glycerolipids.
Furthermore, because Sphingolipids are essential lipids for cell growth (Wells and Lester,
1983) the slow grth of the fatb-k0 plants could result from a slower supply of the

critical 16:0 component to sphingolipids.

130

Although in leaf mesophyll cells the major fraction of fatty acids is used for biosynthesis
of membrane glycerolipids, in epidermal cells most newly produced fatty acids are
directed toward the biosynthesis of cutin and epicuticular waxes. In Arabidopsis leaves,
an epicuticular wax load of 0.2 pmoles/gfw represents only approximately 10 % of the
total leaf pool of palmitate plus stearate, but within the epidermal cells the proportions
will be much higher. Analysis of leaf and stem epicuticular waxes in the fatb-k0 mutant
showed a 20 % and 50 % reduction in total wax load respectively, indicating that the
FATB thioesterase is one source for production of wax precursors in epidermal cells.
Moreover, a reduction of about 85 % in 16 carbon derivatives from cutin demonstrated

that FATB supplies palmitic acid for cutin biosynthesis.

In leaf tissue of the fatb mutant, all extra-plastidial phospholipids showed a reduction of
approximately 50 % in the relative 16:0 levels (Table 9), except PC that showed a 78 %
reduction. The fact that PC was most affected may suggest that this phospholipid is a key
pool in the delivery or partition of palmitate in the cell, for example as an indirect donor
of saturated groups to sphingolipid biosynthesis. In addition, the lO-fold reduction in the
relative 18:0 levels also implicated that PC may play a role in 18:0 partitioning. PC plays
a major role in the flux of glycerol and fatty acids during membrane glycerolipid
biosynthesis and therefore an intriguing question is whether the larger changes in the
fatty acid composition of PC could affect its function and be responsible, at least in part,

for the slower growth rate of the fatb-knockout plants.

Conclusions

131

 

The fatb-k0 line shows a reduction in saturated fatty acids exported to the cytosol, a 17 %
reduction in the rate of growth, and altered seed morphology and germination. Although
this study clearly demonstrates the requirement for the FATB gene and saturated fatty
acids for normal rates of Arabidopsis growth and viable seed formation, the specific
function(s) supplied by saturated fatty acids to sustain normal growth are still uncertain.
Other than the reduction in saturated fatty acid content, and a decrease in wax load,
alterations in glycerolipids and sphingoid base compositions were minor. Future work
will focus on whether the growth rate of the mutant is linked to the biosynthesis of
specific cellular components, subtle variations in membrane properties or changes in
F AS/ lipid tumover-degradation rates or a combination of effects. The recent development
of new isotope labeling techniques to investigate lipid synthesis and tumover-degradation
will be critical to answer these questions (Pollard and Ohlrogge, 1999; Bao et al., 2000).
However, it is important to note that a lack of change of a critical component for growth
may point to its essential nature more than change. Thus, the slower production of an
essential lipid in the fatb mutant could slow growth to a balance point between synthesis
of that component and growth and therefore, no change in the level of the key component
per plant weight would be expected in the mutant. For example, although we observed
no overall reduction in sphingoid base accumulation, the essential nature of sphingolipid
synthesis for growth in other systems (Wells and Lester, 1983) suggests compositional
changes might not be expected. Similarly, if rates of protein acylation, cutin biosynthesis
or synthesis of other saturate derived components are essential to growth, a biochemical
phenotype in these components may not be observed. Thus, the isolation of suppressor

mutants for the fatb-k0 phenotype may also provide insights into the underlying

132

mechanisms which connect the supply of saturated fatty acids to the biosynthesis and/or

regulation of essential plant growth components.

MATERIALS AND METHODS

Plant Material and Growth Conditions

Wild-type Arabidopsis thaliana and fatb-k0 mutant plants (ecotype Wassilewska) were
grown at 80-100 umol m'2 s", 22°C under l8-h light/6-h dark photoperiod. Seeds were
always estratified for three days at 4°C. Selection of T-DNA tagged plants was carried
out by soaking the soil with 50 ug/mL of commercial BASTA (Finale) (AgrEvo,
Montvale, NJ). Surface sterilized seeds of Arabidopsis were germinated on 0.8 % (w/v)
agar solidified MS medium supplemented with l % (w/v) sucrose. For the experiments at
different temperatures the plants were grown for 2 weeks at 22°C (16 hours light) and

then transferred to 16°C and 36°C or kept at 22°C under identical lighting conditions.

Arabidopsis fatb-k0 act] double mutant was generated by using Arabidopsis fatb-k0
plants as a pollen donor and Arabidopsis act] (Kunst et al., 1988) as pollen recipient.
Seeds obtained from the crosses were sown on soil in the presence of 50 ug/mL BASTA
in order to select for heterozygous act]/+ fatb—k0/+ seedlings. F2 seedlings from F1
heterozygous plants were screened by GC and PCR in order to detect homozygous,

heterozygous and wild types for act] and fatb-k0 loci.

Mutant Isolation

133

 

A transfer DNA (T-DNA, 5.5 Kb) insertion into the FATB gene was identified by
screening pooled genomic DNA prepared from a T-DNA tagged Arabidopsis thaliana
(ecotype Wassilewska) collection (Sussman et al., 2000). The gene-specific primers used
for the screening of insertions into the FA TB gene were 5’-
CTCATATCCACATATATCTCTCTCTCACC-3’ forward and 5’-
CAAGCAAGCAAGGTGGTAGTAGCAGATAT-3’ reverse, and the T-DNA specific
primer matching the left end of the T-DNA (IL-202) was 5’-
CATTTTATAATAACGCTGCGGACATCTA C-3’. A FATB genomic fragment was
labeled using random priming and used to detect specific PCR products by Southern blot
hybridization (Sambrook et al., 1989). The T-DNA/FA TB junctions from both ends of the
insertion were PCR amplified, subcloned and sequenced from both ends to determine the

insertion point for the 5.5 kb T-DNA.

Complementation Analysis

The binary vector pBINAR-Hyg (Becker D, 1990) carrying the wild-type FATB cDNA
was a gift from P. Doermann. The vector was used to transform Arabidopsis fatb-
knockout plants by Agrobacterium vacuum infiltration (Bechtold et al., 1993). The
transformation of homozygote plants for the FATB T-DNA insertion did not render
transgenic seedlings and therefore heterozygote fatb-k0 plants were transformed.
Transgenic seedlings were first selected on agar plates in the presence of 25 ug/mL of
hygromycin-B. After 7 days, seedlings were transferred to soil pre-soaked with 50 ug/mL

of BASTA in order to select against wild-type plants for the fatb-knockout T-DNA

134

insertion. DNA was extracted from hygromycin-B (transgene T-DNA) and BASTA
(knock out T-DNA) resistant plants by using the Qiagen Plant DNA extraction kit
(Chatsworth, CA). Homozygote fatb-k0 plants were identified by PCR using the same

primers as for the PCR originally used to isolate the fatb-k0 mutant.

Real-Time PCR Quantification of mRNAs

Total RNA was prepared from leaf tissue of wild-type (Wassilewska) and fatb-k0 mutant
Arabidopsis by using the Qiagen Plant RNA extraction kit (Chatsworth, CA) according to
the instructions. A 5 ug aliquot was used as a template for cDNA synthesis employing the
SuperScript First Strand Synthesis system and oligo dT primers (Stratagene). Specific
primers for the second and third exons of the FATB gene were designed with Primer
Express sofiware (PE Applied Biosystems). The sequences of forward and reverse
primers are: 5 ’ -AATCATGTTAAGACTGCTGGATTGC-3 ’ and S ’-
ATACCATTCTTTCCAGACTGACTGA-3’ respectively (Figure 19). Primers were
verified by showing that the PCR reaction product produced a single band after agarose
gel electrophoresis. Real Time Quantitative PCR analysis was performed according to the
manufacturer’s instructions (PE Applied Biosystems). The reaction contained, in a final
volume of 30 uL, 250 ng of reverse transcribed total RNA, 1.5 uM of the forward and
reverse primers, and 2X SYBR Green PCR Master Mix. All reactions were performed in
triplicate. The relative amounts of all mRNAs were calculating using the Comparative
CT method as described in User Bulletin #2 (PE Applied Biosystems). Arabidopsis
eukaryotic protein synthesis initiation factor 4A1 (eIF4A1) mRNA was used as an

internal control for variations in amounts of mRNA. Levels of FATB mRNA were

135

normalized to eIF 4A1 mRNA levels and presented as a ratio between wild-type and fatb-
ko mutant plants. The forward and reverse primers used to amplify eIF4A] mRNA were
5 ’ -CCAGAAGGCACACAGTTTGATGCA-3 ’ and 5 ’ -

AGACTGAGCCTGTTGAATCACATC-3’ respectively.

Fatty Acid Analysis of Glycerolipids from Different Tissues of Arabidopsis

Approximately 0.1 g fresh weight (gfw) of tissue from 5-week old Arabidopsis plants
was heated at 90°C for 1 hour in 0.3 mL of toluene and 1 mL of 10% (v/v) boron
trichlodide/methanol (Sigma) with heptadecanoic acid (17:0) as an internal standard.
After acidification with aqueous acetic acid fatty acid methyl esters were extracted two
times with hexane and analyzed by gas chromatography with a flame ionization detector

(GC-FID) on a DB-23 capillary column.

Individual Glycerolipid Analysis

One gfw of leaf tissue from 5-week old Arabidopsis plants was ground in liquid nitrogen.
Lipids were extracted in hexane-isopropanol and glycerolipid classes were separated by
thin layer chromatography (TLC) on K6 silica plates (Whatman Inc, Clifton, PA)
impregnated with 0.15 M ammonium sulphate and activated for 3 hours at 110° C (Kahn
and Williams, 1977). The TLC plates were developed three times with 912308 (vzvzv)
acetoneztoluenezwater and lipids were detected after spraying with 0.2 % (w/v) 2’-7’-
dichlorofluorescein/methanol and viewing under UV light. Standards were used to

identify the different glycerolipid classes. Lipids were eluted from the silica with

136

chloroform-methanol and fatty acid methyl esters prepared and analyzed as described

above.

Leaf Epicuticular Waxes

Approximately 3 gfw of leaf tissue from 5-week old Arabidopsis plants were used for
epicuticular wax analysis. The tissue was dipped in chloroform for 30 seconds and then
the following internal standards (IS) were added; n-octacosane at 20 pg per gr of fresh
weight, docosanoic acid and l-tricosanol both at 10 pg per gr of fresh weight. All the
compounds were purchased from Sigma. After evaporation of the chloroform under
nitrogen the epicuticular waxes were silylated to convert free alcohols and carboxylic
acids to their trimethylsilyl(TMS)-ethers and -esters respectively. The epicuticular waxes
were heated at 110° C for 10 min in 100 uL of pyridine and 100 uL of N,O-
bis(trimethylsilyl)trifluoroacetamide (Sigma). After cooling, the solvent was evaporated
under nitrogen and the product was resuspended in 1:1 (volzvol) heptaneztoluene for GC
analysis. GC conditions were the following: a HP-5 capillary column (30 m x 0.32 mm x
0.25 pm film thickness) with helium carrier gas at 2 ml/min was used; injection was in
split mode; injector and FID-detector temperatures were set at 360°C; and the oven
temperature was programmed at 150°C for 3 minutes, followed by a 10°C/min ramp to
350°C, and then held for an additional 20 min at 350°C. GC/MS analysis was also

performed to identify components of the mixture.

Sphingoid Base Analysis

137

Approximately 1 gfw of leaf tissue from 5-week old Arabidopsis plants was heated at
110°C for 24 hours with 4 mL of dioxane (Sigma) plus 3.5 mL of 10 % (w/v) aqueous
Ba(OH)2 (Sigma) (Sperling et al., 1998). D-erythro-sphingosine (Matreya Inc, Pleasant
Gap, PA) at 25 ug/gfw and heptadecanoic acid (Sigma) at 400 ug/gfw were added as
internal standards. After saponification the sample was extracted with chloroform to
obtain the sphingoid bases in the organic phase and fatty acids in the alkaline aqueous

phase. Each phase was independently analyzed as described below.

The chloroform fraction containing the sphingoid bases was back-extracted with an equal
volume of 0.4 M aqueous HCl. The acid aqueous phase (containing the protonated
sphingoid bases) was then titrated with KOH to pH = 10. The sphingoid bases were re-
extracted into chloroform and after evaporation under nitrogen resuspended in 1 mL of
methanol. The sphingoid bases were oxidized to their corresponding aldehydes by stirring
the sample with 100 uL of 0.2 M sodium periodate (Sigma) at room temperature for 1
hour in the dark (Kojima et al., 1991). The aldehydes were recovered by hexane
extraction and used directly for GC analysis. GC conditions were the following: a HP-5
capillary column (30 m x 0.32 mm x 0.25 mm film thickness) with helium carrier gas at 2
ml/min was used; injection was in split mode; injector and FID-detector temperatures
were set at 250°C; and the oven temperature was programmed at 100°C for 3 minutes,
followed by a 10°C/min ramp to 260°C, and then held for an additional 10 min at 260°C.

GC/MS analysis was also performed to identify components of the mixture.

138

 

The basic aqueous phase was acidified with HCl to pH < 4 and extracted twice with
hexane to recover fatty acids. Fatty acids were transmethylated and analyzed by GC using

the same protocol as indicated above.

Monoglucosylceramide (MGC) Analysis

Approximately 10 g of Arabidopsis leaf tissue was quenched with 50 mL of hot
isopropanol and ground in a polytron. The extract was filtered and the residue extracted
with 25 mL of 2:1 (vzv) chloroformzmethanol and re-filtered. This residue was re-
extracted with 25 mL of 1:2 (vzv) chloroformzmethanol and again filtered. All the three
filtrates were combined and evaporated to dryness on a rotary evaporator. The lipid
fraction was finally dissolved in 5 mL of chloroform (“Lipid Fraction” in Table 10). The
solvent extracted residue dried under vacuum (“Solvent Extracted Residue” in Table 10).
The lipid fraction was subjected to a partial base transmethylation to convert most of the
O-acyl glycerolipids to fatty acid methyl esters under conditions which leave the N-acyl
groups intact. This was achieved by vortexing the lipids (100 mg) with 2M KOH in
methanol (0.6 mL) plus hexane (4 mL) for 2 min at room temperature. The reaction was
quenched by adding 4 mL of 1M aqueous acetic acid. The hexane phase was removed
and the acidified aqueous phase extracted with 2:] then 1:2 (vzv) chloroformzmethanol.
The hexane and chlorofonn2methanol fractions were combined and evaporated to dryness
under nitrogen. Lipids were analyzed by TLC as described for “Individual Glycerolipid
Analysis” above. A band that co-migrated with beta-D-Glucosyl Ceramide standard
(Matreya Inc, Pleasant Gap, PA) was eluted from the silica with 2:1 and 1:2 (vzv)

chloroformzmethanol. The solvent was dried under nitrogen and the MGC sample was

139

cleaved by alkaline hydrolysis in 1 mL dioxane (Sigma) and 1 mL 10% (w/v) aqueous
Ba(OH)2 (Sigma) for 24 hours at 110°C. Sphingoid bases were recovered and their

aldehyde-derivatives analyzed as indicated above for total sphingoid base analysis.

Cutin Analysis

Approximately 20 g of leaf or stem tissue was ground in liquid nitrogen and extracted
with 200 mL of isopropanol. The extract was filtered and the residue re-extracted with
200 mL 2/1 (v/v) chlorofomr/methanol. After filtering the residue was re-extracted with
200 mL 1/2 (v/v) chloroform/methanol and air-dried. An excess (2.5 times by weight) of
LiAlH4 plus 6 mL of tetrahydrofuran were added to 100 mg of residue. Hydrolysis was at
80°C for 48 hours with periodic vortexing. After hydrolysis, the excess of LiAlH4 was
decomposed by careful addition of ethyl-acetate to the reaction mixture. The mixture was
acidified by the addition of 5 mL of water plus 0.8 mL of concentrated HCL. Cutin
components were extracted two times with 6 mL of diethyl-ether. The ethereal solution
was dried under nitrogen and the sample dissolved in 0.1 mL pyridine plus 0.1 mL of
BSTFA (SIGMA). Silylation was at 110° C for 10 min. The excess of reagent was
evaporated under nitrogen and the sample dissolved in 1/1 (v/v) heptane/toluene for
GLC-MS analysis in an HP-5 capillary column with the oven temperature programmed at
90°C for 5 min, followed by 10°C/min ramp to 300°C, and then held for an additional 10

min at 300°C.

Acyl-ACP Thioesterase Activities

140

Assays for 16:0-ACP and 18:1-ACP hydrolysis were performed according to Eccleston

and Ohlrogge, 1996.

Scanning electron microscopy

Plants were harvested at 3 weeks of age and fixed in 3% glutaraldehyde in 0.1 M PIPES
(pH 7.2) overnight at 4°C. Samples were washed three times with 0.1 M PIPES (pH 7.2),
10 min per wash, before dehydration through graded ethanol series (50, 60, 70, 80, 90,
100%). Dehydrated samples were transferred to a drying apparatus and critical point
dried at C02. Dehydrated specimens were affixed to stubs with double-sided tape and
coated with gold in argon. SEM pictures were taken at the Center for Advanced
Microscopy at Michigan State University in a J EOL (Japan Electron Optics Laboratories)

J SM-6400V scanning electron microscope.

Transmission electron microscopy

For TEM small leaf samples were cut from the middle of the oldest true leaves and
immersed in 3%glutaraldehyde in 0.1 M PIPES buffer (pH 7.2). Samples were then fixed
overnight at 4°C as described above. After three washes in 0.1 M PIPES buffer, the
tissues were postfixed in 2% aqueous osmium tetroxide overnight at 4°C. Samples were
again washed 3 times in 0.1M PIPES (pH 7.2) and then dehydrated through graded
ethanol series as above. TEM samples were embedded in epoxy resin and thin sectioned

before staining.

Accession Numbers

141

The accession numbers for the Arabidopsis thaliana proteins described in Figure 25 and
text are: acyl-ACP thioesterase B (FATB) (At1g08510); acyl-ACP thioesterase A
(FATA) (At3g25110 and At4gl3050), glycerol-3-phosphate acyltransferase (ACTl)
(At1g32200 ), 3-ketoacyl-ACP-synthase-II (KAS-II) ( Atlg74960), lyso-phosphatidate

sn-2 acyltransferase (LPAAT) (At4g30580) and eIF4Al (At3gl3920).

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Kunst L., Browse J., Somerville C. (1989) A mutant of Arabidopsis deficient in
desaturation of palmitic acid in leaf lipids. Plant Physiol, 90: 943-947.

Kunst L., Browse J., Somerville C. (1988) Altered regulation of lipid biosynthesis in a
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Lightner J., Wu J.R., Browse J. (1994) A mutant of Arabidopsis with increased levels
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Rashotte AM, Jenks MA, Feldmann KA (2001) Cuticular waxes on eceriferum
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8011 J., Roughan G. (1982) Acyl Acyl carrier protein pool sizes during steady state fatty
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Sperling P., Zahringer U., Heinz E. (1998) A sphingolipid desaturase from higher
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Wada H., Shintani D., Ohlrogge J. (1997) Why do mitochondria synthesize fatty acids?
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CHAPTER 4

DISRUPTION OF THE FA TB GENE INCREASES THE RATE OF FATTY ACID
BIOSYNTHESIS,FATTY ACID AND LIPID TURNOVER, AND FAS PROTEIN
EXPRESSION IN LEAVES.

ABSTRACT

Disruption of the FATB gene in Arabidopsis (fatb-k0) results in ~ 60 % reduction in
saturated fatty acid export from plastids and 17 % reduction in the growth rate of the
mutant compared to wild type (Chapter 3 and Bonaventure et al., 2003). In this study we
report that although fatb-k0 seedlings grow more slowly than wild type the rate of fatty
acid synthesis (FAS) in leaves of the mutant increases by 40 %. Moreover, the protein
levels of BCCP, ACPS and 18:0-ACP desaturase are increased by 1.5-2 fold in fatb-k0
leaves compared to wild type, suggesting concerted up-regulation of FAS protein
expression. To maintain constant amounts of fatty acids in leaves, thereby
counterbalancing their higher rate of production, the mutant also increases by 70 % the
initial rate of fatty acid breakdown. However, although fatb-k0 leaves have higher rates
of fatty acid synthesis and turnover, the composition of membrane lipids is similar to
wild type. Thus, homeostatic mechanisms to preserve membrane compositions are
functional and compensate for substantial changes in rates of fatty acid and glycerolipid
metabolism in the mutant The results suggest that fatb-k0 cells attempt to increase
saturated fatty acid production, however a surplus of fatty acids above amounts needed
for leaf growth is created and leads to increased turnover. In the mutant, the surplus of

fatty acids exported from leaf plastids (largely C18 unsaturated fatty acids) and

146

1: AF. I
n
I

In'. .2 o

1

incorporated into polar lipids is mainly lost via degradation or interconversion of C182C18-
PC species. In addition, Cu, fatty acids are also rapidly removed from this phospholipid in
fatb-k0 leaves, suggesting that mechanisms are not present which can preferentially

preserve the saturated fatty acids.

INTRODUCTION

In plants, the major pathway for the de novo fatty acid synthesis occurs in the plastid
(Ohlrogge et al., 1979) and this organelle exports fatty acids to supply a diverse array of
extraplastidial biosynthetic pathways and cellular processes in the cell (Kolattukudy,
2003; Yalovsky et al., 1999; Post-Beitenmiller, 1996; Lynch, 1993). Production of fatty
acids to be exported from plastids depends on the activity of acyl-ACP thioesterases
(F ATs) that hydrolize the acyl group from acyl-acyl carrier protein (acyl-ACP) to release
free fatty acids and ACP (reviewed by Voelker et al., 1997). After export from the
plastids, free fatty acids are re-esterified to coenzyme-A (CoA) to form the cytosolic
acyl-COA pool (Pollard and Ohlrogge, 1999). In most plant cells, these molecules are
primarily used for the biosynthesis of membrane glycerolipids at the level of the
endoplasmic reticulum (ER) (Browse and Somerville, 1991). However, in other tissues
exported acyl-COA molecules have different destinations. For example, in oilseeds, the
major fraction of acyl groups is incorporated into triacylglycerols (TAG) that may
constitute more than 90 % of total lipids. In epidermal cells, a significant fraction of acyl-
CoA molecules is utilized for the synthesis of waxes and cutin (Kolattukudy, 2003; Post-

Beittenmiller, 1996). Furthermore, all cells synthesize sphingolipids, and we recently

147

estimated that as much as 30-40 % of exported 16:0-COA is needed for sphingoid base
synthesis in leaves (Bonaventure et al., 2003). Finally, exported 14:0-CoA and 16:0-COA

participates in other acylation reactions, such as protein acylation (Yalovsky, 1999).

Because acyl-ACP thioesterases terminate fatty acid synthesis and allow the export of
fatty acids from plastids, these enzymes are critical regulators of cellular metabolism,
controlling the partitioning of de nova-synthesized fatty acids between plastids and
cytosol and therefore their supply to diverse cellular processes. Two classes of FAT
enzymes has been described in most plants, namely FATA and FATB (Voelker et al.,
1997). The FATA class has highest in vitro activity for 18:1-ACP and lower for saturated
acyl-ACP substrates. In contrast, the FATB class prefers saturated acyl-ACP but also
shows activity for unsaturated acyl-ACPS (Salas and Ohlrogge, 2002; Voelker et al.,
1997; Doermann et al., 1995). The Arabidopsis genome encodes two FATA genes and a
single FATB gene (Beisson et al., 2003; Mekhedov et al., 2000). We previously
described the isolation and analysis of an Arabidopsis mutant disrupted in the FA TB gene
(fatb-k0) (Bonaventure at al., 2003). In this mutant, the overall amounts of saturated fatty
acids in different tissues are reduced by 40 to 50 % compared to wild type. This reduction
occurs only in the cytosolic pool of saturated fatty acids, affecting the fatty acid
composition of extraplastidial glycerolipids, synthesis of wax in leaves and stems, and the
fatty acid composition of TAGS in seeds. However, although sphingolipid synthesis is
initiated from 16:0-COA, no reductions were observed in total sphingoid base content,
suggesting that plants may prioritize the synthesis of these essential lipids (Wells and

Lester, 1.983). These observations also suggested the existence of a metabolic hierarchy

148

 

 

in the economy of saturated fatty acids in plant cells, which prioritizes synthesis of
essential components for growth (Bonaventure et a] 2003). The reduction in the pool of
cytosolic saturated fatty acids also slows down the growth of the mutant, resulting in
seedlings approximately half the size of wild type seedlings (Bonaventure et al, 2003).
Thus, reduction in the export of saturated fatty acids from plastids affects cellular

processes that are critical for plant growth.

Complete suppression or disruption of any of the core enzymes of FAS would be
expected to reduce fatty acid synthesis and affect plant performance. In support of this,
analysis of tobacco plants engineered to constitutively express an antisense transcript of
the tobacco biotin carboxylase (BC) or the Arabidopsis biotin carboxylase carrier protein
(BCCP), showed reductions of leaf fatty acids together with a stunted phenotype
(Shintani et al, 1997). Growth phenotypes together with reduction in leaf lipids were also
obtained with antisense constructs directed against stearoyl-ACP desaturase and ACP4,
and with induced mutations in the fab2 and enoyl-ACP reductase genes (Branen et al.,
2003; Mon at al., 2000; Lightner et al., 1994). It is apparent from these studies that even
slight reductions in leaf fatty acids synthesis can have pleiotropic effects on plant growth

and development. Some of these effects come from changes in chloroplast membrane

structure and therefore loss of photosynthetic capability. By contrast, a wide range of

mutations in plant fatty acid desaturases, acyltransferases and condensing enzymes
demonstrate that the fatty acid composition of plant membranes can be altered
considerably with no apparent phenotype under normal growth conditions (Wallis and

Browse, 2002; Kunst et al., 1989; Wu et al., 1994). These mutants reveal the plasticity of

149

 

lipid metabolism in plants and the capacity of these organisms to adjust to alternative

lipid compositions.

Based on these observations, one possible mechanism responsible for the slower growth
of fatb-k0 plants could be a reduced synthesis of fatty acids and therefore a decline in the
rate of membrane lipid biosynthesis (Branen et al., 2003; Shintani et al., 1997).
Alternatively, slower growth of the mutant may be the result of reduced synthesis of
other critical components such as sphingolipids, cutin and waxes or lower rates of
acylation reactions. In addition, lipid-derived signaling molecules that affect growth
could be affected in the mutant (Nandi et al., 2003). When fatb-k0 plants are grown at a
range of different temperatures, no suppression of the fatb-k0 grth phenotype is
observed, suggesting that changes in the bulk properties of cellular membranes are not
the main factor affecting plant performance (Bonaventure et al., 2003). However, more
subtle changes in transport and vesiculation cannot be ruled out. Likewise, when the
mutant is grown at different relative humidity levels and even in liquid culture, no
reversion of the growth phenotype occurs, suggesting that increased water loss through
reduced wax content is also not a major factor affecting growth. Similar results are
obtained when fatb-k0 is supplemented with exogenous sugar and in this case its
photosynthetic capability appears not to limit its growth (Bonaventure et al., 2003). In
addition, the chlorophyll content and the ultrastructure of fatb-k0 chloroplasts are similar

to wild type (Chapter 3, Figure 22)

150

g

 

To further elucidate the role of saturated fatty acids in plant growth and to understand
fatty acid partition and F AS regulation in fatb-k0, a series of isotope labeling experiments
were conducted. Unexpectedly, the rates of both fatty acid synthesis and turnover were
higher in fatb-k0 than wild type leaves. In addition, up-regulation of three proteins
involved in FAS was detected, suggesting that higher F AS rates were achieved at least in
part by higher FAS protein expression. Thus, fatb-k0 plants appear to induce a futile
cycle of fatty acid production and degradation, perhaps as an attempt to increase saturated

fatty acid synthesis.

RESULTS

Rate of fatty acid biosynthesis in leaves of wild type Arabidopsis and fatb-k0.

The rate of fatty acid biosynthesis in leaves correlates mainly with the expansion rate of
this organ, being higher in younger leaves and reflecting the demand for new membranes
to sustain cell division and chloroplast biogenesis (Bao et al., 2000; Browse et al., 1981).
It was previously reported that fab-k0 plants grow slowly resulting in a 50 % decline in
their fresh weight compared to wild type (Bonaventure et al., 2003). To evaluate if the
disruption of the FATB gene reduces the rate of fatty acid synthesis in leaves,
incorporation of labeled precursors was used to assess the in vivo rate of this pathway in
Arabidopsis wild type and fatb-k0. Because isotope incorporation rates can be influenced
by internal pools which might differ between mutant and wild type (Nunn et al., 1977;

Cronan et al., 1975), three different labeled substrates were tested.

151

aura-ruin”
h.

 

First, the rate of 3 H incorporation into fatty acids from 3HzO was measured in leaf tissue
of wild type and fatb-k0. The results in Table 13 indicate that mutant leaves incorporated
3 H into fatty acids at a rate 40 % higher than wild type tissue. The incorporation of 3 H
was linear for the first two hours of the assay and followed non-linear kinetics thereafter
(data not shown). Similar to other systems, a lag phase of 10-15 min for 3 H incorporation
into fatty acids was observed (data not shown and Browse et al., 1981). Second, leaf
tissue of wild type Arabidopsis and fatb-k0 were incubated with exogenous [MC] acetate
as the label substrate. Similar to the results obtained with radiolabeled water, mutant
leaves incorporated 14C from acetate into fatty acids at a rate 30 % higher than wild type
(Table 13). The kinetic of 14C acetate incorporation into fatty acids was linear for at least
1 hour (data not shown). Finally, intact wild type and mutant seedlings were labeled with
14C02. As shown in Table 13, fatb-k0 leaves incorporated 14C from C0; into fatty acids at
a higher rate than wild type (ca. 50 %). In all the experiments, the chlorophyll content per
gram fresh weight (gfw) in leaves of the two plant classes was within 5 % of one another

(on average: 1.05 i 0.01 mg/gfw for wild type and 1.07 i 0.03 mg/gfw for fatb-k0).

 

Table 13. Rates of fatty acid synthesis (FAS) in leaves of
Arabidopsis wild type and fatb-k0 measured by different
radiolabeled substrates

 

Substrate Wild type* fatb-k0* IncreaseI (%)
3H20 2801 i 139 3878 i 145 40
14C02 42.0 i 1.6 63.6 a: 2.9 50

"‘C acetate 39.0 i 1.2 50.9 i 1.4 30

 

Units are in nmol of 3 H or 14C / h / mg Chl incorporated into fatty
acids and values represent initial rates of fatty acid synthesis (see
Materials and Methods). lPercent increase of fatb-k0 rates versus
wild type rates.

 

152

 

 

gm

 

In conclusion, by using three different radiolabeled substrates it was demonstrated that

fatb-k0 leaves synthesized fatty acids at an average rate 40 % higher than wild type.

Rate of fatty acid turnover in wild type and fatb-k0 leaves

We previously reported that leaves from fatb-k0 plants contained the same amount of
fatty acids per gram of fresh weight than wild type (Bonaventure et al., 2003). Therefore,
if the FAS rate in fatb-k0 leaves is on average 40 % higher than wild type, the rate of
fatty acid tumover must concomitantly increase to maintain fatty acid amounts constant
in this tissue. Therefore, to test this conclusion fatty acid turnover was evaluated by
labeling intact seedlings with a 30 min pulse of 14CO; and then determining the level of

MC in fatty acids at different times up to 120 hours (Figure 26).

During the first 6 hours of the chase period, there was net accumulation of '4C into fatty
acids of both wild type and mutant (Figure 26). This net accumulation of label most
likely reflected the use of 14C labeled carbohydrates as substrates for fatty acid synthesis
(Bao et al., 2000). After 6 h of the initial l4CO; pulse, the amount of label in fatty acids
from wild type and mutant began to decay (Figure 26). In the 6-120 h period, the
radioactivity in fatty acids dropped by 70 % in fatb-k0 leaves compared to 47 % in wild
type leaves (Figure 26). Therefore, wild type and mutant leaves have an average turnover
rate of 9 % and 14 % per day respectively (55 % increase in fatb-k0). However, as the
label in fatty acids decayed following exponential kinetics in the mutant, the average
rates did not properly reflect the actual fatty acid turnover rates (Figure 26). Thus, if

initial rates were considered instead (6-24 h period), wild type and mutant plants lost

153

1

 

 

0.39 and 0.67 nmol 14C/mg Chl/hour, respectively (Figure 26). These results demonstrate
that the initial rate of fatty acid turnover in the mutant was approximately 70 % higher

that in wild type.

 

80

 

 

nmoles 14C in fatty acids/mg Chl

 

 

time (hours)

Figure 26. Fatty acid turnover in wild type ( O ) and fatb-k0 ( O ) leaves.

Intact plants were pulsed for 30 min with 2 mCi of 14C02 and fatty acids chased for a
period of 5 days in a non-radioactive atmosphere at 22° C. At the times indicated leaf
samples were harvested and the fatty acids purified and their radioactivity measured as
described in “Materials and Methods”.

Analysis of polar lipid classes in wild type and fatb-k0 leaves

To investigate whether higher rates of fatty acid turnover in fatb-k0 leaves affected the
tunover of particular membrane glycerolipids, we analyzed leaf glycerolipid classes after

pulsing wild type and fiztb-ko seedlings with 14CO; and chasing lipids for different times

154

 

 

 

up to 120 h. The results shown in Figure 27 indicate that the distribution of radiolabeled
plastidial and extraplastidial glycerolipids was similar between wild type and fatb-k0

leaves.

The label in phosphatidylcholine (PC) from wild type and fatb-k0 declined by
approximately 65 % in the 0-120 h period (Figure 27). This high rate of label
disappearance in PC is explained by the donation of diacylglycerol moieties (DAG) from
PC to galactolipids and sulfolipids in chloroplasts (Roughan, 1970).
Phosphatidylethanolamine (PE) was the only glycerolipid analyzed with lower label in
fatb-k0 than wild type leaves (Figure 27). In wild type tissue, PE accounted for
approximately 7 to 9 % of labeled lipids throughout the 5 days period whereas in the
mutant represented only 5 to 6 %. Moreover, these levels remained constant throughout
the chase period (Figure 27). This observation agreed with the lower steady state amounts
of PE observed in fatb-k0 leaves compared to wild type (Bonaventure et al., 2003).
Lower incorporation of radioactivity in PE of the mutant was also observed at early time
points during continuous labeling of leaves with [HQ-acetate (data not shown). These

data suggest reduced synthesis rather than increased degradation of this phospholipid in

fatb-k0 leaves.

In the case of plastidial lipids, the percentage of label in monogalactosyldiacylglycerol

(MGDG) and digalactosyldiacylglycerol (DGDG) increased by approximately 25 % and

155

 

 

 

 

Percentage of radioactivity

 

 

 

Percentage ofradioactivity

 

 

 

time (hours)

Figure 27. Redistribution of radioactivity among glycerolipid classes of wild type ( O

) and fatb-k0 ( O ) leaves. Seedlings were pulsed with 14C02 and leaf glycerolipids
separated by TLC and their activities quantitated as described in “Materials and
Methods”. Abbreviations: PC, phophatidylcholine; PE, phosphatidylethanolamine;
MGDG, monogalactosyldiacylglycerol; DGDG, digalactosyldiacylglycerol; PG,
phosphatidylglycerol.

150 % respectively in both wild type and fatb-k0 during the chase period (Figure 27).
These increments in labeled MGDG and DGDG at longer times represented the

contribution of the eukaryotic pathway to the plastidial pool of these lipids (Browse et al.,

156

1989). These data together with the decline of labeled PC are consistent with the parallel
operation of the prokaryotic and eukaryotic pathways of lipid biosynthesis in wild type
and fatb—k0 leaves. In contrast, the percentage of label in phosphatidyldiacylglycerol
(PG), which is mainly produced in the plastid, declined in both plant classes by

approximately 30 % (Figure 27).

The kinetics of l4C labeling in the different lipid classes of wild type Arabidopsis leaves
were in accordance with previous studies in which the same tissue was labeled with 14C-
acetate (Wu et al., 1994; Browse et al., 1989; Kunst et al., 1989). In conclusion, the
similarity in the distribution of label among lipid classes in wild type and fatb-k0 leaves
indicate that despite higher rates of fatty acid turnover cells maintained a constant

composition of polar lipids.

Analysis of radiolabeled C16 and C13 fatty acids in polar lipids of wild type and fatb-
ko leaves

The fatty acid composition of leaf membrane glycerolipids is different between wild type
andfatb-ko. In this regard, PE, PS and P1 in the mutant have reductions of approximately
50 % in their 16:0 content compared to wild type, whereas in PC the reduction is 80 %
(Bonaventure et al., 2003). To understand how the lack of FATB activity affects fatty
acid partitioning into leaf glycerolipids, we evaluated redistribution of labeled fatty acids
in these lipids. Wild type and mutant seedlings were pulsed with 14CO; and radiolabeled
fatty acids in polar lipids chased for different times up to 120 b (Figure 28). In order to

obtain an accurate estimate of 14C18 and 14C”, levels, fatty acids were first hydrogenated

157

 

~

I.‘
l;

 

 

 

to remove double bonds and then separated by reverse phase-thin layer chromatography
(TLC) according to their chain lengths. The results of the analysis revealed that in the
mutant, redistribution of labeled C13 and Cu, fatty acid was similar to wild type in

plastidial lipids but differed substantially in extraplastidial lipids (Figure 28).

In PC of wild type leaves, labeled C16 fatty acids increased by 25 % in the 0-120 h period
(from 20 % to 25 %) whereas in PC of the mutant they decreased by 60 % (from 15 % to
6 %) (Figure 28). These results indicated that in PC of fatb-k0 leaves, C16 fatty acids were
first incorporated at 30 % lower levels than wild type and later removed to reach 75 %
lower levels than wild type. Moreover, PC in the fatb-k0 lost almost 50 % of its labeled
C16 fatty acids in 6 hours, suggesting a rapid removal of this fatty acid from PC (Figure
28). Therefore, the ~80 % reduction in the amount of C 10 in PC of fatb-k0 compared to
wild type (Bonaventure et al., 2003), results from both reduced incorporation but mainly

from higher removal of this fatty acid (Figure 28).

Labeled C10 fatty acids in PE redistributed differently from those in PC. In both, wild

type and fatb-k0 leaves their levels decreased after the initial incorporation (Figure 28).

158

 

 

MGDG

 

k.-

 

Percentage ome fatty acids

 

 

 

 

 

 

 

s.

 

Percentage 0me fatty acids

 

 

 

0 6 12 24 48 72

 

time (hours)

 

 

40 i 1 i r i r in r i I i r i
0 6 12 24 48 72 96 120

 

time (hours)

Figure 28. Redistribution of radioactivity in C16 fatty acids of membrane

glycerolipid classes from wild type ( O ) and fatb-k0 ( O ) leaves. Seedlings were
pulsed with 14C02, lipid classes separated by TLC and their corresponding fatty acids
transmethylated, hydrogenated and separated by reverse phase TLC. Abbreviations: PC,

phosphatidylcholine; PE, phosphatidylethanolamine; MGDG,
monogalactosyldiacylglycerol; DGDG, di galactosyldiacyl glycerol; PG,
phosphatidyl glycerol.

159

 

 

 

 

Nevertheless, PE from mutant leaves incorporated lower levels of C u, fatty acids than
wild type (Figure 28). Thus, in contrast to PC, the steady state differences in the C16
content of PE between wild type and fatb-k0 was primarily the result of lower

incorporation instead of increased removal of this fatty acid (Bonaventure et al., 2003).

In contrast to extraplastidial glycerolipids, the incorporation and redistribution of label in
C18 and C16 fatty acids of plastidial glycerolipids was similar between wild type and fatb-
ko (Figure 28). These observations agreed with previous studies in which no significant
changes in the fatty acid composition of MGDG, DGDG and PG were found between
wild type and the mutant (Bonaventure et al., 2003). These results also suggest that the
ratio of degradation or interconversion to other components of C13 and Cu, fatty acids
from MGDG, DGDG and PG was not altered in fatb-k0 compared to wild type. In both,
MGDG and DGDG, the C16 fatty acid levels declined in accordance with their
dependence on the prokaryotic and eukaryotic pathways for their synthesis (Figure 28).
Moreover, the more pronounced decline of labeled C16 in DGDG reflected the higher
dependence of this lipid on the eukaryotic pathway (Browse et al., 1986). In the case of
phosphatidylglycerol (PG), the situation was different from the other major plastidial
lipids and reflected its primary synthesis by the prokaryotic pathway (Browse and
Somerville, 1991). The level of C16 fatty acids incorporated into this phospholipid at time
0 was 55 % and increased to more than 60 % at 120 h (Figure 28). These data suggest

that Cu, fatty acids in PG were more stable than C18 fatty acids.

Mass distribution of C133C13 and szClg molecular species of polar lipids in wild
type and fatb-k0 leaves

160

_ .T'm'mf

 

 

 

Arabidopsis membrane glycerolipids are composed of either C163C18 or C182C13 fatty
acids in a 1:1 molar ratio. Whereas lipids derived from the prokaryotic pathway are
predominantly C1(,:C13, those derived from the eukaryotic pathway are either C lg:C .8 or
ClozCrx (Browse and Somerville, 1991). In order to better understand the fluxes of C13
and CH, fatty acids into lipids and their degradation or interconversion to other lipids, the
decay in the mass of labeled fatty acids was calculated for the different molecular species

of polar lipids (Figure 29).

The results shown in Figure 29A indicate that in wild type leaves, PC lost 4.8 nmol
l“C/mg, Ch] of c.8138 and 3.3 nmol 14C/mg (3111 of (3,613.8 during the 6-120 h time
frame. In parallel, MGDG and DGDG together gained 3.5 nmol l4C/mg Chl of C13:C13
and lost 12.4 nmol l4C/mg Chl as C162C13 (Figure 29A). Thus, gain of Clnglg species in
MGDG and DGDG could be accounted for by transfer of these molecules from PC. The
remaining 1.3 nmol 14C/mg Chl of C132C18 lost from PC could either be degraded in the
cytosol or interconverted into other lipid-derived components (e.g., membrane lipids).
Thus, in wild type, most of C182C13 loss in PC (73 %) was via MGDG and DGDG
biosynthesis in the plastid (Figure 30). The fate of lost szClg species in PC could be
explained by trafficking of these molecules back to the plastid, degradation in the cytosol

or interconversion into other molecules (Figure 30).

161

”15"“

 

 

 

 

A. wild type

   
    

I)’. fatb-I10

 

 

 

PC PC
20
[j C16:C18
10
0 __
20 7 PE PE

10

 

 

 

 

20 i DGDG DGDG

10

 

nmol l4C lmg Chl

 

 

20

 

 

 

 

 

20-

10

 

 

 

 

 

0

.;
N v o
v- N 19

N60
NON
.—

Time (hours)

Figure 29. Redistribution of C13: C13 and C16:C13 molecular species of polar lipids in
wild type and fatb-k0. Absolute masses for radiolabeled C13: C13 and C162C13 species
were calculated using total label in fatty acids, polar lipid abundance and percentage of
radiolabeled C13 and C16 in the different polar lipids. Abbreviations: PC,

phosphatidylcholine; PE, phosphatidylethanolamine; MGDG,
monogalactosyldiacylglycerol; DGDG, digalactosyldiacylglycerol; PG,
phosphatidyl glycerol.

162

 

 

 

In fatb-k0 leaves, PC lost 10.4 nmol MC/mg cm of cure18 and 2.4 nmol MC/mg of
C1sz 18 in the 6-120 h period (Figure 29B). MGDG and DGDG together gained 3.1 nmol
l4C/mg Chl of C182C18 and lost 21.9 nmol l4C/mg Chl of CmiClg (Figure 29B). Thus,
from the total mass of labeled Clnglg lost from PC only 3.1 nmol 14C/mg Chl contributed
to galactolipid synthesis whereas 7.3 nmol l4C/mg Chl were not accounted for. Thus,
these results indicate that in contrast to wild type, the major fraction (70 %) of C132C13
lost from PC was not incorporated into galactolipids in the mutant. Importantly, this
fraction represented 62 % of the label lost from extraplastidial lipids in fatb-k0 whereas
only 18 % in wild type (Figure 30). The fate of this mass of labeled C182C13 together with
lost 06sz from PC could be degradation in the cytosol, interconversion to other cellular

components or trafficking to the plastid in case of the latter (Figure 30).

A second important observation of these experiments was that disappearance of C162C13
species from MGDG, DGDG and PG represented the main loss of mass from labeled
fatty acids in wild type and fatb-k0 leaves. In leaves of the former, total loss of labeled
fatty acids was 23.4 nmol l4C/mg Chl in the 6-120 h period (Figures 1 and 5).
Disappearance of C16:C13 species from MGDG, DGDG and PG accounted for 16.4 mnol
l4C/mg Chl, and corresponded to ~70 % of total mass loss (Figure 30). In the mutant, the
total loss of labeled fatty acids was 40.8 nmol l4C/mg Chl in the same period and in this
case 29 nmol l4C/mg Chl were lost through plastidial Cloiclg species (~70 % of total

mass loss) (Figure 26 and 5). These results indicate that a major fraction of labeled fatty

163

 

acids was eliminated by turnover of C162C18 Plastidial species and that the percentage of

total label loss by this route was similar between wild type and fatb-k0 (~70 %).

In summary, these results demonstrate that increased fatty acid synthesis in fatb-k0 was
compensated by increased absolute rates of lipid turnover in a way that preserved the
ratio of polar lipids. Furthermore, the results suggest that in the mutant the surplus of
fatty acids exported from plastids and incorporated into polar lipids is lost at the level of
PC via degradation or interconversion of this phospholipid into other components. In
addition, this higher degradation/interconversion rate of PC in fatb-k0 leaves may
explained why Cu, fatty acids are rapidly removed from this phospholipid and accumulate
at levels 80 % lower than wild type, whereas in the rest of the extraplastdial lipids they

accumulate at levels 50 % lower than wild type (Bonaventure et al., 2003).

Immunoblot analysis of wild type and fatb-k0 leaves.

To determine if the increase in the rate of fatty acid biosynthesis in fatb-k0 leaves was
correlated with an up-regulation of FAS protein expression, specific antibodies against
BCCP (biotin carboxylase carrier protein), acyl carrier proteins (ACP) and stearoyl-ACP
desaturase were used for immunoblot analysis of protein extracts from wild type and
fatb-k0 leaves. As shown in Figure 31 protein levels of the BCCP subunit of plastidic
acetyl-COA carboxylase (ACCase) were increased by 1.5-fold in fatb-k0 leaves compared
to wild type. Similarly to BCCP, the protein levels of ACPS and stearoyl-ACP desaturase

were induced by approximately 2—fold in the mutant (Figure 31). Arabidopsis leaves

164

I

 

express several isoforms of plastidic ACPS with ACP-4 as the most abundant in this

tissue followed by ACP—2 and 3 (Hlousek-Radojcic et al.. 1992). The immunoblot results

 

 
  

Total
+42 wt
+60fatb I

1

 
        
 

plastid

 

     
 
  
 
 

 

 

 

 

 

 

 

 

 

Croicrs
-l6.4 Wt
-29fatb
Degradation? :
lnterconversion?
1 1‘
Cisicrs Crsicrs
- 3.3 Wt - 1.3 wt
9.41am M
l——_.
C18:C18 C16:Cl8
- 2.4 wt
- 0.3fatb - 1.8 atb

 

 

 

 

Figure 30. Scheme of mass loss from labeled fatty acids in leaves of wild type and
fatb-k0. The arrows depict the flux of either fatty acids (16:0/18:1) exported from the
plastid or €111:ng and C 162C”; in the different lipids (gray: wild type (wt); black:fi1tb-ko
(fatb)). The values inside the boxes are nmol l4C/mg Chl of C13: Cu; and ClézClg species.
Initial positive values represent total label input at time 6 hours and negative values loss
of labeled C13: C 13 and CmICIg species after 120 hours. Loss of label from galactolipids
and PG most likely represent degradation. Loss of labeled C16:C13 from PC could not be
assigned to a specific pathway, it may occur by conversion to galactolipids, degradation
or interconversion to other cellular components. Similar fates can be assigned to lost
ClgICm-PC. C lg:C jg-PE and C miClg-PE. Abbreviations: GaL: galactolipids (MGDG and
DGDG); PC, phosphatidylcholine; PE, phosphatidylethanolamine; PG,
phosphatidyl glycerol, FAS: fatty acid synthesis pathway.

165

 

 

indicated that most of ACP isoforms were up—regulated in leaves of the mutant (Figure
31). In contrast, a single band in the blot probed with anti-stearol-ACP desaturase
antibody appeared up-regulated in fatb-k0 leaves, suggesting that one major leaf isoform

of this enzyme was induced (Figure 31).

Changes in the expression of some FAS proteins occur during leaf development, being
higher in young leaves and declining after this tissue completes its expansion (J.
Ohlrogge, unpublished results). Thus, to determine whether the increased levels of
BCCP, ACPS and 18:0-ACP desaturase in fatb-k0 were the result of differences in the
developmental stage of wild type and mutant leaves, immunoblot analysis was also
performed on leaf extracts at different stages of development (2, 3 and 4 weeks old
seedlings). The results showed a consistent increase (1 .3-2 fold) in the levels of the three
FAS proteins in fatb-k0 leaves compared to wild type at the different stages of

development (data not shown).

DISCUSSION

Acyl-ACP thioesterases (FAT) are responsible for the export of fatty acids from the
plastid produced by the de novo fatty acid synthesis. In this study, isotope labeling
experiments and western blots were performed to investigate changes in lipid metabolism
and protein expresSion brought about by disruption of the FATB gene in Arabidopsis

(Bonaventure et al., 2003). These experiments suggest that fatb-k0 alters its fatty acid and

166

sneer-‘91

 

glycerolipid metabolism together with FAS protein expression to compensate for the
deficiency in saturated fatty acids. However, higher FAS rate in fatb-k0 increases the

production and export from the plastids of 18:1 over 16:0 fatty acids (Bonaventure et al.,

 

fatb-k0 wild type
A 3.75 7.5 15 30 3.75 7.5 15 30 (ug)
BCCP -5 "— ‘- —-— -—--
ACPS M.,
18:0-ACP , .......
Desaturase
2.5

B
2.0
Fold increase 1.5
(fatb—k0 vs wt)
1.0
0.5
0.0

i BCCP ACPs 18:0-Aer
Desaturase

Figure 31. Immunoblot analysis of BCCP, ACP and 18:0-ACP desaturase in leaf
tissue of wild type and fatb-k0 Arabidopsis. A, total Arabidopsis protein was extracted
from leaves of wild type and mutant and increasing amounts loaded on the gel (3.75, 7.5,
15 and 30 ug). BCCPs were detected with anti-biotin antibodies, ACPs and 18:0-ACP
desaturases with specific antibodies against spinach ACP-I and avocado stearoyl-ACP
desaturase, respectively. Relative molecular weight for BCCP-l is 35 kD, for ACP 10 kD
and for 18:0-ACP desaturase 35 kD. When known, major leaf isoforms are noted; B,
amount of each polypeptide was quantitated with ImageQuant software and is expressed
as relative fold increase of mutant versus wild type. The fold values shown are the
average of the fold values for each protein concentration and the bars denote the standard
deviation of the average. Fold increase values correspond to BCCP-l, all ACPS detected
and major 18:0-ACP desaturase band. The intensity of coomasie—blue stained bands was
used to normalize western blot signals.

167

2003) and the resulting excess of C18 unsaturated fatty acids may surpass the capacity of
cells to use them for membrane biosynthesis. This surplus leads then to increased fatty
acid turnover (Figure 32). Importantly, although fatb-k0 leaves undergo substantial
changes in fatty acid and glycerolipid metabolism, the cells maintain a constant
composition of polar lipids. These observations demonstrate the importance of lipid
homeostasis in plant cell membranes and how cells adapt glycerolipid metabolism to
alternative fatty acid fluxes. Similar results were obtained with the Arabidopsis act]
mutant in which a major disruption of the prokaryotic pathway for lipid biosynthesis does

not affect significantly the lipid composition of the plant (Kunst et al., 1989).

Induction of a futile cycle of fatty acid production and degradation is also observed when
the california bay 12:0-ACP thioesterase (MCTE) is over-expressed in developing
Brassica napus seeds (Eccleston and Ohlrogge, 1998). In this system, the re-direction of
fatty acids towards 12:0 reprogram fatty acid metabolism in a way that both the catabolic
and biosynthetic pathways are increased to remove the surplus of 12:0 and to maintain
normal levels of C16 and C13 fatty acids, respectively. Thus, although by means of
different mechanisms, the missexpression of acyl-ACP thioesterases either in mutant or
transgenic plants has major impacts in lipid metabolism and triggers cellular responses to
lessen those effects. These observations demonstrate the central role of acyl-ACP
thioesterases in balancing and providing fatty acids for several different cellular

processes.

Increased rates of fatty acid biosynthesis and turnover in fatb-k0

168

11W!

3

 

 

 

Different labeled substrates have been used to assess the in vivo rate of fatty acid
biosynthesis in plants, with [l-MC]-acetate, l4COz and 3H20 the most commonly used
(Bao et al., 2000; Pollard and Ohlrogge, 1999; Browse et al., 1981; Jungas, 1968). The
advantages of using labeled water over acetate or carbon dioxide to determine in vivo
rates of product biosynthesis is that water crosses all membranes and rapidly equilibrates
with all body water pools (Kelleher, 2001). Moreover, using labeled water eliminates the
problem of the dilution of carbon tracers in inaccessible pools and also of different
metabolite pool sizes in wild type versus mutant plants (Kelleher, 2001). For example,
initial differences in the intracellular pools of acetate between wild type and mutant
organisms could result in differential dilution of the tracers inside cells and therefore in
different specific radioactivities in labeled fatty acids (Nunn et al., 1977; Cronan et al.,
1975). A second important parameter when performing labeling experiments with plants
is the use of intact plants versus tissue sections (e. g., leaf discs). A drawback of using the
latter is that some artifacts may be introduced by induction of wounding responses that

alter fatty acid metabolism (Nishiuchi et al., 1997; Conconi et al., 1996).

Therefore, the observation that three different labeled substrates gave consistent increases
in FAS rate of fatb-k0 compared to wild type, demonstrates that this mutant increases the
rate of fatty acid production in leaves. Moreover, the use of cut and intact tissue shows
that this effect is independent of differential wounding responses between the mutant and
wild type. An important conclusion of these results is that fatty acid production appears
not to limit the amount of total membrane lipid biosynthesis and consequently grth of

fatb-k0 plants. Thus, the slower grth of the mutant is most likely the consequence of a

169

Figure 32. Comparative scheme of fatty acid metabolism in wild type and fatb-k0.
FAS activation signals are triggered by mechanisms that sense low levels of saturated
fatty acids in the cytosol. The same mechanisms may induce the up-regulation of FAS
protein expression by transcriptional or post-transcriptional mechanisms. In addition,
biochemical and metabolic activation of the FAS machinery are also considered.

170

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171

reduced supply of saturated fatty acids for cellular processes other than membrane lipid
biosynthesis. However, changes in membrane properties as a result of changes in their
fatty acids composition can not be ruled out at this point as a mechanism affectting

growth of fatb-k0.

Wild type and fatb-k0 leaves presented an average rate of fatty acid turnover of 9 % and
14 % per day respectively (Figure 26). Bao et al., (2000) previously reported a turnover
rate of 4-5 % per day in wild type Arabidopsis plants. The higher rate obtained in our
experiment may be the result of using different growth conditions and plant ages. In
particular, the lightzdark period in our experiment was 18:6 hours whereas it was
13.52105 hours in the study by Bao et al., (2000). During light periods the rate of fatty
acid turnover is accelerated and therefore longer light periods will increase decay of

labeled fatty acids (Bao ct al., 2000).

In addition, because fatb-k0 seedlings have slower growth rate than wild type
(Bonaventure et al., 2003) and the decay of labeled fatty acids was followed for a period
of 5 days, differences in the net accumulation of fatty acids per seedling were significant
between wild type and mutant plants. Wild type seedlings increased by 2-fold their fatty
acid content whereas fatb-k0 seedlings by 1.88-fold in this period of time (data not
shown). Thus, the differences between the fatty acid turnover rates of wild type and fatb-
ko (either 55 % when the average is considered or 70 % when the initial rate is

considered) are slightly underestimated.

172

Turnover of Polar Lipids in wild type and fatb-k0

Analysis of the absolute turnover rates of C132C13 and C16:C18 molecular species of polar
lipids reveal that after 5 days of chase period, approximately 70 % of the label lost in
both wild type and fatb-k0 leaves was as C102C18 species of galactolipids and PG (Figures
4 and 5). This observation agrees with the fact that this pool of lipids contain more than
70 % of the fatty acids in leaves. Moreover, the similar percentage of label lost via
galactolipids and PG in wild type and fatb-k0 firrther demonstrate the conservation of

lipid homeostasis despite higher absolute rates of lipid turnover in the mutant.

One intriguing question is why PC in fatb-k0 leaves has a bigger reduction in 16:0
content compared to the rest of extraplastidial glycerolipids (Bonaventure et al., 2003). In
this study we demonstrate that C16 fatty acids are rapidly removed from PC afier
incorporation in the mutant whereas they accumulate in wild type (Figure 28). One
possibility is that the increased turnover of PC in the mutant is not selective for the type
of fatty acid and therefore Clnglg as well as szClg species of this phospholipid are
either degraded or interconverted to other components. Alternatively, more Clszlg
species from PC may be used for plastid lipid biosynthesis in the mutant or C10 fatty acids
selectively removed from PC to supply other cellular processes (e.g., sphingolipid

synthesis or acylation reactions).

The kinetics of fatty acid labeling also indicate that the ratio of flux of total fatty acids

(C18 plus C16) through the prokaryotic and eukaryotic pathways is not substantially

173

altered in mutant leaves compared to wild type. Hence, the ratio of the distribution of
initial masses of l4C-fatty acids into the different polar lipids between these two pathways
was similar when wild type and fatb-k0 were compared (data not shown). Moreover, the
data agree with previous results obtained from lipid biochemical analysis (Figure 25 in

Chapter 3 and Figure 5 in Bonaventure et al., 2003).

Up-regulation of F AS protein expression

Disruption of the FATB gene in Arabidopsis has provided new insights into some of the
regulatory mechanisms that affect expression of F AS enzymes. What are the mechanisms
involved in up-regulation of BCCP, ACPS and 18:0-ACP desaturase protein levels in
fatb-k0 leaves? Are there other fatty acid proteins up-regulated in fatb-k0 leaves? The use
of large scale techniques for mRNA profiling will help to investigate whether up-
regulation of FAS protein expression occurs at the mRNA level and also to identify
additional FAS transcripts with altered expression in mutant versus wild type leaves.
Moreover, this analysis could give additional information on changes in the expression of
non-lipid genes that may help to understand changes in overall metabolism and perhaps

to identify regulatory genes (Ruuska et al., 2002; Girke et al., 2000).

Conclusions and Future Perspectives

The rate of FAS in fatb-k0 leaves is increased by 40 % compared to wild type and this

increment may be partially achieved by up-regulation of FAS protein expression. The

174

 

resulting increased supply of fatty acids above amounts needed for leaf growth leads to
increased fatty acid turnover. Importantly, despite major changes in fatty acid and
glycerolipid metabolisms in the mutant a constant composition of polar lipids is
maintained in leaves. The increase in fatty acid synthesis and up-regulation of BCCP,
ACPs and 18:0-ACP desaturase protein levels suggest that plant cells have mechanisms
capable of sensing subnormal levels of saturated fatty acids and signaling the activation
of the FAS machinery and protein expression in order to increase their production. The
understanding of the mechanisms responsible for activation of fatty acid synthesis and
lipid turnover as well as up-regulation of FAS protein expression in fatb-k0, may provide
important clues for the understanding of the regulation of fatty acid and lipid synthesis in

plants.

MATERIALS AND METHODS

Plant Material and Growth Conditions

In all the experiments wild type Arabidopsis thaliana and fatb-k0 mutant plants (ecotype

Wassilewska) (Bonaventure et al., 2003) were grown on a mixture of soilzvermiculite

(1:1) for three weeks under white fluorescent light (100 nmol m"2 s") in a 18h:6h
lightzdark photoperiod at 22°C in growth chambers. Sowed seeds were always stratified

for four days at 4°C.

Rate of Fatty Acid Biosynthesis by Arabidopsis Leaves

175

Rapidly expanding leaves from wild type and fatb-k0 plants (3 weeks old) were cut in
strips (0.5 cm wide) and transferred to pre-weighed glass flasks containing 4.75 mL of
incubation buffer (2.5 mM sodium MES (2-[N-Morpholino] ethanesulfonic acid) pH: 5.7,
0.0075 % (w/v) Tween-20 and 2.15 mg/mL of MS salts. Leaf strips from the same plant
were randomly distributed in different flasks to ensure sample homogeneity between
flasks. After sufficient samples had been prepared (approximately 0.2 g fresh weight per
flask), the flasks were re-weighed to obtain gram fresh weight (gfw) values. The assay
started by the addition of either 0.25 mL of3H20 (100 mCi/mL, 3.7 GBq/mL) (ARC Inc,
American Radiolabeled Chemicals Inc; St. Louis, MO) or 0.025 mCi of [l-MC] sodium
acetate (56 mCi/mmol) (ARC Inc, American Radiolabeled Chemicals Inc; St. Louis, M0)
to each flask and incubating them for different times at 22°C in a temperature-controlled
water bath with gentle agitation and continues illumination. All data points were
performed in duplicate and the values presented in Table 13 represent initial rates of fatty
acid synthesis (calculated using data from 0, 10, 20, 40, 60 min of continuous labeling).
At the end of the assay period the incubation medium was removed and the tissue quickly
washed twice with de-ionized water and quenched by heating in 10 mL of isopropanol for
10 min at 80°C. Lipids were extracted with hexane-isopropanol method (Hara and Radin,
1978). An aliquot of the lipid extract was suspended in acetone:water (4:1, v/v) to
determine chlorophyll content (Amon, 1949). The methodology used to determine the
rate of fatty acid biosynthesis using 14CO; was identical to the methodology used to
determine the rate of fatty acid turnover (see below) with the only difference that plants

were removed at different time points from the sealed bag using a double sealed air trap.

176

Rate of Fatty Acid Turnover by Arabiodpsis Leaves

Wild type and fatb-k0 plants were grown for 3 weeks as indicated above. One day prior
to the experiment, a total of 12 pots (6 pots with wild type and 6 withfatb-ko plants (15
plants per pot)) were randomly placed inside a transparent glove bag (40 litre gas space,
12R® Instruments for Research and Industry, Cheltenham, PA) with circulating air and
same lighting and temperature conditions as indicated above. A 30 min pulse of 14C02
was given to the plants by mixing 2 mCi of 14C—NaHCO3 (56 mCi/mmol) (ARC Inc; St.
Louis, MO) with concentrated sulfuric acid inside the sealed bag and air circulated by
using a small battery-driven fan. For the chase period, the radioactive atmosphere was
rapidly vented and the plants were placed in a normal (non-radioactive) atmosphere for
different times in the same growth conditions as described above. At each time point , 15
wild type and 15 fatb-k0 plants were randomly removed from different pots and separated
in two individual samples (7-8 plants/sample for wild type and mutant). Leaf tissue was
immediately weighed, frozen in liquid nitrogen and stored at -80°C for subsequent lipid
extraction with hexane—isopropanol method (Hara and Radin, 1978) and for chlorophyll

content determination (Amon, 1949).

Fatty Acid Analysis of Lipid Extracts from Leaf Tissue

Transmethylation of fatty acids from total lipid extracts was performed similarly for all

the experiments. An aliquot of lipid extracts was heated at 90°C for 1 hour in 1 mL of

177

10% (v/v) boron-trichloride/methanol (Sigma). After acidification with aqueous acetic
acid, fatty acid methyl esters (FAMES) were extracted two times with hexane and
radioactivity in the sample (either 3 H or 14C) analyzed by scintillation counting (Beckman
Instruments Inc, Fullerton, CA). A second aliquot of total lipid extract from each sample
was transmethylated and F AMES were separated by thin layer chromatography (TLC) on
K6 silica plates (Whatman Inc, Clifton, PA) using 90/10 (v/v) hexane/diethyl-ether.
Radioactive bands corresponding to FAMES were localized by scanning in an Instant
Imager system (Packard, Meriden, CT). The bands corresponding to FAMES were
recovered from the plates and radioactivity measured by scintillation counting (Beckman

Instruments Inc, Fullerton, CA).

Individual Glycerolipid Analysis.

Glycerolipid classes from total lipid extracts were separated by thin layer
chromatography (TLC) on K6 silica plates (Whatman Inc, Clifton, PA) impregnated with
0.15 M ammonium sulphate and activated for 3 hours at 1100 C (Kahn and Williams,
1977). The TLC plates were developed three times with 91/30/8 (v/v/v)
acetone:toluene:water and scanned for radioactivity using an Instant Imager (Packard,
Meriden, CT), both to quantitate radioactivity and to locate the appropiate bands for
recovery. Lipids were also detected after spraying with 0.2 % (w/v) 2’-7’-
dichlorofluorescein/methanol and viewing under UV light. Standards were used to
identify the different glycerolipid classes. Lipids were eluted from the silica with

chloroform/methanol/water (5/5/1) and fatty acid methyl esters prepared as described

178

above. FAMES from the different lipid classes were hydrogenated using hydrogen at
slightly greater than atmospheric pressure with a platinum (IV) oxide catalyst in
methanol. The reaction was performed for at least 2 hours giving complete reduction of
unsaturated to saturated FAMES. Analysis and isolation of 18 and 16 carbon FAMES
were by KC18 reverse phase TLC (Whatman, Clifton, PA), developed half and then fully
with acetonitrile/methanol/water (130/70/1, v/v/v). Plates were scanned for radioactivity
using an Instant Imager (Packard, Meriden, CT) to quantitate radioactivity and to locate
the appropiate bands for recovery. Scintillation counting was also used to measure
radioactivity after scrapping-off the bands corresponding to 16 and 18 carbon fatty acids

from the plates.

SDS-PAGE and Immunoblotting

Extraction of proteins from leaves was performed as follows. Up to 0.1 g of leaf tissue
was harvested and placed into a 1.5 mL plastic microcentrifuge tube. Plant material was
pulverized with a microcentrifuge pestle in the presence of liquid nitrogen. Powder was
immediately reconstituted in 0.2 mL of extraction buffer (2 % (v/v) 2-mercaptoethanol,
50 mM HEPES (pH: 7.8), 100 mM NaCl, 0.05 % (w/v) SDS) by vortexing. Insoluble
debris was collected by centrifugation for 15 min at 12,000g. Supernatant was removed
and placed into a fresh microcentrifuge tube. Protein concentration was determined by

dye-binding protein assay using bovine serum albumin as the standard (Bradford, 1976).

179

 

 

 

SDS-PAGE and protein transfer to nitrocellulose was performed using standard
conditions. Nitrocellulose membranes were blocked for at least 1 h with 10 mM Tris-
HCl, pH: 8.0, 0.15 M sodium chloride, 0.3 % (v/v) Tween 20 (TBS-T), and 2 % (w/v)
nonfat dry milk. Anti-biotin antibodies conjugated to alkaline phosphatase (Kirkegaard
and Perry Laboratories, Gaithersburg, MD) were directly added at a 1:5,000 dilution to
detect BCCP. Antisera raised against avocado stearoyl-ACP desaturase (Shanklin and
Somerville, 1991) and spinach acyl carrier protein-I (ACP-I) (Post-Beittenmiller et al.,
1991) were added at a 1:1,000 dilution. Probing proceeded for 16 h at 4° C followed by 1
h at 25° C after which membranes were briefly rinsed with TBS-T. Antibody bound
proteins were detected by incubating blots for l h at 250 C with alkaline-phosphatase-
conjugated anti-rabbit secondary antibodies (Kirkegaard and Perry Laboratories,
Gaithersburg, MD). The membranes were then washed 6 times for 5 min each with TBS-
T. Blots were then washed 5 min with developing solution (0.1 M Tris-HCl, pH: 9.5, 0.1
M sodium chloride and 5 mM magnesium chloride) before colorimetric detection in
developing solution containing 0.33 mg/mL p-nitro blue tetrazolium chloride and 0.17
mg/mL 5-bromo-4-chloro-3-indoyl phosphate p-toluidine salt. Immunoblot signals were
quantitated using ImageQuant software (Molecular Dynamics, Sunnyvale, CA), which
accounted for band area plus intensity. Blot signals were corrected for differences in
protein loading (as determined by Coomasie Brilliant Blue R 250 staining) and
normalized against values of the control plants. Increasing amounts of total protein were
resolved by SDS-PAGE for immunoblot analyses to ensure that both major and minor

bands were within the linear sensitivity range of the detection system.

180

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Chapter 5

FINAL CONCLUSIONS, PERSPECTIVES AND FUTURE DIRECTIONS

The work presented herein used several approaches to increase our understanding of the
signals and regulatory mechanisms that influence fatty acid biosynthesis (FAS) in plants.
These approaches range from gene expression analysis to detailed biochemical analysis
of lipid metabolism in a mutant and the results have provided new insights into regulation
of this primary metabolic pathway. In addition, work has focused on the partition of fatty
acids from their source inside plastids to an array of biosynthetic pathways outside this
organelle. One goal of plant biotechnology is to use these organisms as large scale
chemical factories for production of edible and industrial products and as an alternative to
petroleum-derived products (Thelen and Ohlrogge, 2002). The understanding of the
regulatory mechanisms of fatty acid synthesis and partitioning should contribute to the

development of better strategies to manipulate fatty acid production in plants.

Conclusions and future directions on F AS gene expression in Arabidopsis

Major conclusions of this thesis research related to FAS gene expression are:

1. Different environmental and metabolic signals regulate ACP expression in

Arabidopsis depending on the ACP isoforms and their tissue specific expression.

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2. Up-regulation of a subset of fatty acid and lipid genes and the steady expression of
others appear to be sufficient to match new requirements for fatty acids in plant
tissues.

3. Mechanisms that sense low levels of saturated fatty acids may signal the up-

regulation of FAS proteins in Arabidopsis leaves.

In Chapter 2, the genes encoding for acyl carrier proteins (ACPS) were selected for the
initial experiments and results presented in that chapter demonstrate that regulation of
these genes depends on the protein isoforms and their tissue specific expression. For
example, ACP4 is the most abundant isoform in Arabidopsis leaves and it is regulated by
light and carbon, similarly to photosynthetic genes. By contrast, the less abundant leaf
isoforms ACP2 and ACP3 are not light-regulated but instead respond to growth,

suggesting a more critical role for them in rapidly dividing tissue (e.g., meristems).

One obvious question that arises from the analysis of ACP expression is whether ACP4 is
the only fatty acid gene regulated by light in Arabidopsis. A recent report based on
GeneChip analysis of phytochrome mutants and different lighting conditions indicates
that at least 4 additional genes involved in fatty acid and lipid synthesis respond also to
light (Tepperrnan et al., 2001). These genes include ACP4, chloroplast linoleate
desaturase (FAD7), omega-3-desaturase (FAD3), chloroplast omega-6 desaturase
(FAD6) and stearoyl-ACP desaturase. Since the microarrays used in that study covered
approximately one third of the predicted genes in the Arabidopsis genome (Affymetrix

8K GeneChip), additional fatty acid and lipid genes may also be light regulated.

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Are ACP2 and ACP3 the only fatty acid genes that are growth/cell cycle regulated in
Arabidopsis? A recent report describes a GeneChip analysis of synchronized Arabidopsis
cell suspension cultures used to identify cell-cycle controlled genes (Menges et al.,
2002). The results indicate that only ACP2 and 3, two putative acyl-CoA synthetases and
chloroplast omega-3-desaturase (F AD7-8) are under cell-cycle regulation in Arabidopsis.
Again, only one third of the Arabidopsis genome was covered by the GeneChip used in
that study and therefore the list of fatty acid and lipid genes regulated by cell-cycle may

extend in the future.

When an Arabidopsis cDNA microarray representing 3,500 genes was used to examine
changes in gene expression during Arabidopsis seed development and in particular during
the phase of oil accumulation (8-12 days after flowering), it was found that only a subset
of genes involved in fatty acid and lipid metabolism showed significant changes in
expression (Ruuska et al., 2002). For example, ACCase subunits, KAS I and II, ACP1,

and F AD2 and 3 show peaks of expression between 8-12 days after flowering.

In summary, it appears from these observations that rather than up-regulating all or most
genes for fatty acid and lipid synthesis, plant cells up-regulate a limited number of them
according to the tissue and its specific needs for fatty acids and lipids. Thus, instead of
“global” regulatory mechanisms for all or most FAS genes, up-regulation of a subset of
them and the steady expression of others seem to be sufficient to match new requirements

for fatty acids in different tissue or under changing environmental conditions. Although

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we focused our research on ACP genes, a logical extension can be applied to other fatty
acid genes and we speculate that most likely all of them respond differentially to diverse
signals and can be up-regulated according to tissue requirements (Menges et al. 2002,
Ruuska et al., 2003; Tepperrnan et al., 2001). In Arabidopsis, most enzymes involved in
fatty acid and lipid synthesis are encoded by multiple genes, and one intriguing question
has been why Arabidopsis requires so many different FAS enzyme isoforms (Beisson et
al., 2002). Their differential tissue expression and response to signals under variable

conditions sheds some light on this question.

Analysis of gene expression in fatb-k0 leaves, indicates that the protein levels of BCCP,
ACPS and 18:0-ACP desaturase are increased between 1.5 to 2-fold in young leaves.
Thus, these results suggest that mechanisms that sense low levels of saturated fatty acids
can increase the expression of at least some F AS proteins. Therefore a new signal can be
added to the repertoire of signals that, by still unknown mechanisms, activate fatty acid
protein expression. In addition, it remains to be tested whether additional fatty acid gene
products also respond to this regulatory mechanism. Thus, is there a global up-regulation
of fatty acid proteins in fatb-k0 leaves or, are only a subset of critical genes up-regulated?
An answer to this question will provide interesting information to correlate induction of

specific FAS genes with increased rates of fatty acid production in leaves.

Future directions

Although most of the structural genes for fatty acid and lipid biosynthesis are known or

have strong putative candidates, information about regulatory factors for this metabolic

188

 

 

 

pathway is still poor (Beisson et al., 2003; Mekhedov et al., 2000). Thus, one
breakthrough step in plant fatty acid and lipid research will be to isolate regulatory
factors for the expression of FAS proteins. One important consequence of the
identification of these factors would be to manipulate lipid metabolism in genetically
engineered plants. The advantage of expressing regulatory factors rather than single
enzymes is that an entire metabolic pathway could be up-regulated and for example
increase the yield of desired products (Gantet and Memelink, 2002; Memelink et al.,
2001; Grotewold et al., 1998). Nevertheless, one difficulty presented by primary
metabolism is that it often is not very amenable to metabolic engineering, in part because
it operates in parallel with multiple pathways and therefore has complex interconnections
with them (Carman and Henry, 1999). For example, carbon and nitrogen metabolism are
closely interconnected and genes involved in nitrogen metabolism are regulated by
carbon (Oliveira and Coruzzi, 1999). In addition, alteration of primary metabolism can
cause unwanted secondary effects that have to be controlled before engineered plants can
be used for commercial purposes (e.g., deformed morphology and size of seeds with
reduced saturated fatty acids, Chapter 3-Figure 20). Thus, in order to successfully
manipulate fatty acid metabolism in plants, it is helpful to know and consider the

complex interconnections between metabolic pathways and their regulation.

The studies performed in Chapter 2 open the possibility to identify transcription factors
involved in regulation of fatty acid gene expression. One procedure to achieve this goal is
to develop a system in which signal—specific (e.g., growth) changes in gene expression

can be followed to perform functional promoter analysis (Sun et al., 2003; Chen et al.,

189

2002; Fujiki et al., 2000). Thus, in Chapter 2 (Figure 15) a system to identify. elements
involved in expression and regulation of the Arabidopsis ACP2 promoter by growth was
developed. Further characterization (deletions and site-specific mutations) of this
promoter in tobacco BY-2 cells will allow the identification of short DNA elements
important for growth-regulation of this gene. Once these elements have been identified,
the isolation of transcription factors can be approached by techniques such as yeast one-
hybrid system, oligo-affinity chromatography, and electromobility shift assays (EMSA)
(Carles et al., 2002; Uno et al., 2000; Jarrett, 1993). Keeping in mind our longer-term
interest in increasing oil production in seeds, one evident question is whether the isolation
of factors involved in regulation of ACP2 by growth will also be important for expression
of ACP2 or fatty acid genes in seeds. On the one hand, the same transcription factor
could be expressed in different tissues and have similar functions perhaps by association
with alternative partners. On the other hand, completely different regulatory factors may
be used in different tissues. In this case however, the ectopic expression in seeds of, for
example, a meristem specific factor, could still have effects on fatty acid metabolism in

seeds.

Similarly, a functional promoter analysis of ACP4 can allow the identification of
transcription factors involved in light-regulation of this gene. For this purpose an ACP4
promoter deletion series fused to luciferase can be created and analyzed in leaf tissue of
transgenic plants. In parallel, the same plants can be used for the analysis of mechanisms

involved in sugar-repression of the ACP4 gene.

190

um
. 1‘

 

I

 

What are the mechanisms involved in up-regulation of BCCP, ACPS and 18:0-ACP
desaturase protein levels in fatb-k0 leaves? Is this up-regulation at the level of
transcription or at downstream processes such as protein turnover? Are there other fatty
acid genes up-regulated in fatb-k0 leaves?

One experiment that may provide important information about increases in mRNA
abundance of BCCP, ACPs and 18:0-ACP desaturase together with additional fatty acid
genes is to compare transcript profiles between fatb-k0 and wild type leaves by
microarray analysis. Moreover, this analysis could give additional information on
changes in the expression of non-lipid genes that may help to understand changes in
overall metabolism and perhaps to identify regulatory genes. One limitation of
microarrays is that this technique only searches for differences in transcript abundance
and information on translational control or protein degradation are not considered. In
addition, transcript abundance for regulatory proteins is usually low and difficult to detect
by this method (Vainrub and Pettitt, 2003; Watson and Akil, 2002). Thus, additional
experiments will be required to obtain a more complete picture of changes in fatty acid
gene expression including translational control (e.g., polysome association) and protein

turnover.

What is the role of the C T C CGCC box and polypyrimide tracts in the 5 ’ leader of A CP
mRNAs?

The results in Chapter 2 (Figure 9) indicate that the 5’UTR of ACPS have an active role
on gene expression. One possible scenario is that specific factors bind to the leader and

facilitate the interaction between ribosomes and ACP transcripts. Alternatively, a tertiary

191

“T‘mr‘

I
-4!!!

 

 

structure in the RNA may form to facilitate ribosome association and presumably
translation initiation (Bolle et al., 1996; Staub and Maliga, 1994; Danon and Mayfield,
1991). Several approaches can be followed to answer these questions, for example the
conserved elements in the ACP leaders can be replaced with random sequences to test
how the association with polysomes and expression of a reporter gene are affected in
transgenic plants. Second, the binding capacity of factors present in cytosolic fractions to
ACP 5’UTRs can be investigated by electromobility-shift assays (EMSA) (Danon and
Mayfield, 1991). Transcript stability can be analyzed by techniques such as mRNA
turnover after inhibiting RNA Pol II activity with a-amanitin in seedlings grown in liquid

culture (Hua et al., 2001).

Conclusions and future directions on fatty acid metabolism and its regulation in
fatb-k0.

Major conclusions of this thesis research related to the role of saturated fatty acid are:
1. FATB is a major regulator of saturated fatty acid production in all tissues.

2. Saturated fatty acids have a critical role in plant growth and development.

3. Low levels of saturated fatty acids in the cytosol triggers signals to increase the rate

of fatty acid biosynthesis in leaf plastids.

Chapter 3 investigates the production and partition of saturated fatty acids from plastids
to other cellular pathways. Analysis of the fatb-k0 mutant revealed that FATB is a major
enzyme responsible for export of saturated fatty acids from plastids in all tissues.

Moreover, reduction of cytosolic saturated fatty acid pools brought about by the lack of

192

if“..-

 

 

FATB caused changes in the distribution of saturated fatty acids between different
pathways that reveal a hierarchy in the metabolism of these molecules. For example,
although there is more than a 50 % reduction in exported saturated fatty acids in the
mutant, sphingoid base amounts in leaves were similar to wild type. In contrast, changes
in wax amounts could be detected between these two plant classes. These observations
are interpreted to indicate that maintaining sphingolipid levels is more essential than
maintaining wax amounts for the plant. Some of the changes in saturated fatty acid-
derived pathways have clearly negative effects in plant performance and seed
development at standard growth conditions. Thus, saturated fatty acids have a critical role
in plant growth and development and the understanding of the cellular steps affected will
be important to manipulate fatty acid composition in plants without affecting their

performance (see below).

Analysis of fatty acid metabolism in fatb-k0 also gave new insights into the mechanisms
and signals that increase the rate fatty acid biosynthesis in plant cells (Chapter 4). Low
levels of saturated fatty acids in the cytosol not only triggers signals to up-regulate fatty
acid protein expression but also to increase the rate of fatty acid biosynthesis. These
results demonstrate that plants can uncouple rate of growth from rate of fatty acid
production and therefore they have the inherent capacity to increase fatty acid production
although growing at lower rates than normal. This is an important observation because it
suggests that cells can be programmed to produce more fatty acids independent of a

developmental program or rate of growth that sets the amount of fatty acids to be made.

193

Future directions

What signal(s) activate fatty acid synthesis in plants? How is the rate of fatty acid
synthesis increased?

These are crucial questions that if answered will give important insights in the regulation
of fatty acid biosynthesis in plants. This thesis research suggests that fatb-k0 provides a
new system to tackle these questions. The rate of fatty acid biosynthesis in fatb—k0 leaves
is increased by approximately 40 % compared to wild type and unraveling the underlying
activation mechanisms could provide a breakthrough in understanding regulation of fatty
acid biosynthesis. One critical issue to begin to answer these questions is to understand
where in the cell the signal(s) to activate FAS are being generated. In Chapter 4, several
mechanisms have been proposed to explain possible signal sources such as sensing of
extraplastidial acyl-COA pools, rate of sphingoid base synthesis, rate of protein acylation,
changes in membrane properties that affect transport or vesiculation. One possible
approach to identify the actual mechanisms is to isolate suppressors of fatb-k0 (see

below).

What is the target of these activation mechanisms in fatb-k0 leaves? Is AC Case activity
altered in fatb—k0 leaves?

Several lines of evidence suggest that acetyl-COA carboxylase (ACCase) is a major
regulatory enzyme in fatty acid production in plants and therefore it could be one target
of the activation mechanisms (Ohlrogge and Jaworski, 1997). The B-CT subunit of
ACCase is phosphorylated and this event is correlated with activation of its activity

(Savage and Ohlrogge, 1999). Thus, analysis of the phosphorylation status of B—CT in

194

fatb-Ito and wild type leaves could provide information on ACCase as a target of the

activation mechanism induced in the mutant.

In addition, evidence for a negative feedback regulatory mechanism acting on ACCase
comes from experiments where tobacco cells were incubated in the presence of
exogenous fatty acids (Shintani and Ohlrogge, 1995). Could ACCase in fatb-k0 leaves be
released from some sort of feedback control? It has been proposed that in bacteria
ACCase is inhibited by acyl-COA products, and this inhibition can be released by
overexpression of acyl-CoA thioesterases (Jiang and Cronan, 1994). However, plant
ACCase appears to be non responsive to long acyl-ACP products in vitro (Roesler et al.,
1996). Other potential feedback inhibitors of FAS enzymes could be acyl-CoA, free fatty
acids and glycerolipids. Because FAS occurs inside the plastids but the major utilization
of the products of fatty acid synthesis is at the ER membranes, it is possible that feedback
regulation requires communication across the plastid envelop. Could cytosolic saturated

fatty acids be feedback controllers of FAS synthesis?

What is limiting growth of the fatb-k0 mutant?

Different hypothesis can be proposed to explain the reduced growth of fatb—k0 mutant as
outlined in Chapter 3. However, pleiotropic effects brought about by the genetic lession
could be difficult to interpret and one approach to study this complex system is to
identify suppresors of the grth phenotype of fatb-k0. The mutant grows slower than the
wild type plants and identification of seedlings that performed better than the mutant

could be identified in mutageneized fatb-k0 populations. Mutagenesis of fatb-k0 can be

195

performed chemically by using EMS or by T-DNA activation tagging. One advantage of
using EMS is that it creates point mutations that could lead to amino acid substitutions
and change protein properties without killing its activity (J ander et al., 2003; Me Callum
et al., 2000). On the other hand, activation tagging finds suppressors by overexpression
of targeted genes and therefore a different group of genes from those targeted by EMS is
selected (Memelink, 2003; van der Fits et al., 2001). Suppresors of fatb-k0 may fall in
different categories, for example genes that increase saturated fatty acids levels, genes
involved in biochemical pathways different from FAS, genes involved in regulation of
FAS, genes that participate in membrane transport or vesiculation, and changes in protein
acylation or properties of acylated proteins. Thus, as mentioned above the suppressor
approach could help to unravel both: mechanisms that limit fatb-k0 growth as well as
regulatory mechanisms of fatty acid synthesis. For instance, if a particular cellular
process is affected and limits growth, a mutation that bypass this limitation could reside
in a regulatory protein of an interdependent process. To illustrate this idea, suppressor
mutants of yeast deficient in Golgi vesicle formation were mapped in genes
corresponding to transcription factors (HACl) and kinases (IREl) that are involved in the
unfolded protein response (UPR) signaling pathway (Higashio and Kohno, 2002). Thus,
activation of UPR indirectly stimulates ER to Golgi vesiculation and reconstitutes growth
of yeast affected in this process. In a different experiment, yeast suppressors of
sphingolipid biosynthesis where isolated and named SLC (sphingolipid compensation)
(Lester et al., 1993). These strains are viable without synthesizing sphingoid bases and
therefore lack ceramides. SLC mutants cells synthesize novel PI (phosphatidylinositol)

substituted with long (26 carbons) fatty acids in the sn-2 position of glycerol. Thus, the

196

SLC suppressors overcome the essential function of sphingolipids by producing novel PI

that structurally mimics sphingolipids.

In conclusion, the identification of regulatory factors and the unraveling of the regulatory
mechanisms responsible for FAS activation in fatb-k0 together with the origin and nature
of the inducing signal(s) may provide essential clues for the understanding of the

regulation of fatty acid and lipid synthesis in plants.

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