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11m WM“

 

 

TOWARD A BIOCHEMICAL AND MOLECULAR UNDERSTANDING
OF THE BIOSYNTHESIS OF FA'ITY ACID AND
TRIACYLGLYCEROL IN PLANTS
By

Xiaoming Bao

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

Doctor of Philosophy

Department of Botany and Plant Pathology

1 999

ABSTRACT
TOWARD A BIOCHEMICAL AND MOLECULAR UNDERSTANDING OF THE
BIOSYNTHESIS OF FATTY ACID AND TRIACYLGLYCEROL IN PLANTS
By
Xiaoming Bao

In plants, fatty acid synthesis is a primary metabolic pathway and
triacylglycerol (TAG) is one of the means for storing energy and carbon in seeds.
In this thesis, I addressed four questions dealing with plant lipid metabolisms which
span the pathway from the origin of acetyI-CoA to the ﬁnal accumulation of TAG in
oil seeds. First, acetyl-CoA is the ultimate precursor for fatty acid synthesis in all
kinds of tissues, but its origin has been a debated topic. Free acetate has been
suggested as a possible source of acetyl-CoA in leaf tissues. Pulse-chase and
continuous labeling with “CO2 were conducted in leaves ofArabidopsis, barley, and
pea, and the label rapidly appeared in fatty acids with a lag phase of less than 2
min. These data suggest that either the bulk pool of acetate is not involved in fatty
acid synthesis or the concentration of acetate must be less than 0.05 mM in light
grown leaves. Second, a genomic sequence of biotin carboxylase (BC), one of four
subunits of multisubunit acetyl-CoA carboxylase, was isolated from Arabidopsis
and its promoter was partially characterized. BC expression was found to be higher
in silique and ﬂower than in root and leaf. The region from -140 to +147 contained
necessary promoter elements which supported basal gene expression. A positive
regulatory region was found to be located between -2100 and -140, whereas a

negative element was possibly located in the ﬁrst intron. In addition, several

conserved regulatory elements were identiﬁed in the BC promoter. Third, to
examine whether newly synthesized oleic acid is directly elongated to erucic acid
in developing seeds of B. rapa, embryos were labeled with [“Clacetate, and the
ratio of radioactivity of carbon atoms C(5)-C(22) (from plastid) to C(1)-C(4) (from
cytosol) of erucic acid was monitored with time. This labeling ratio decreases with
time and therefore suggests the existence of an intermediate pool of oleate which
contributes at least part of the oleoyl precursor for the production of erucic acid.
The malonyl-CoA for elongation of oleate to erucate was produced by homodimeric
ACCase. Both light and haloxyfop increased the accumulation of [“C]oleate and
the parallel accumulation of [“Clphosphatidylcholine. Taken together, these results
show an additional level of complexity in the biosynthesis of erucic acid. Fourth,
to examine limiting factors on seed TAG production, developing C. lanceolata, U.
carpinifolia and U. parvifolia embryos were incubated with factors whose availability
might limit oil accumulation. The addition of glycerol or sucrose did not signiﬁcantly
inﬂuence the rate of TAG synthesis. However, the rate of 1“C-TAG synthesis, upon
addition of 2.1 mM 1“C-decanoic acid (10:0), was approximately four times higher
than the in vivo rate of TAG accumulation in Cuphea, and two times higher than the
respective in vivo rates in Ulmus. 14C-10:0 was incorporated equally well in all
three acyl positions of TAG. The results suggest that both Cuphea and Ulmus
embryos have sufﬁcient acyltransferase activities and glycerol-3-phosphate levels
to support rates of TAG synthesis in excess of those found in vivo. Consequently,
the amount of TAG synthesized in these oilseeds may be in part determined by the

amount of fatty acid produced in plastids.

Copyright by
Xiaoming Bao

1999

ACKNOWLEDGMENTS

As a foreign student with totally different culture background, especially the
language barrier, being able to reach this point should be credited to the people
around me. Firstly, I thank my major professor John Ohlrogge for taking the risk
to accept me into his lab. John assigned me an exciting and challenging research
platform, which gave me initial confidence, generated concrete results, and
allowed me to develop my own ideas in. His understanding and constant
encouragement enabled me to overcome some of the down times in the past ﬁve
years. I also want to thank my committee members, Ray Hammerschmidt, Lee
McIntosh, and Michael Thomashow for providing suggestions and guidance that
were invaluable to the completion of work presented in this thesis. My appreciation
also goes to the people who directly contributed to the work presented here; they
are Manfred Focke (chapter 1), Mike Pollard (chapter 1 and 3), and Basil Shorrosh
(chapter 2).

These past ﬁve years have been the most challenging and enjoyable years
of my life and for that I thank David Shintani, Linda Savage, Nicki Engeseth, Keith
Roesler, Sarah Hunter, Sergei Mekhedov, David Schultz, David Pan, Mi-Chung
Suh Jim Todd, Thomas Girke, and Jay Thelen. And to Yuanping, who sacriﬁced
her own career, has given me the support necessary to accomplish this, thank you
and I’m sorry for neglecting you for these years. Lastly I thank the American people
in the surrounding communities who provide intangible support and tolerance which

I am forever in debt to.

TABLE OF CONTENT

LIST OF TABLES .............................................................................................. Viii
LIST OF FIGURES .............................................................................................. ix
LIST OF ABBREVIATIONS ................................................................................. xiii
FATTY ACID NOMENCLATURE ........................................................................ XiV
CHAPTER 1 : '"trOdUCtion ........................................................................... 1
What is the Origin Of Plastidial AcetYI-COA? .............................................. 5

What Is the Nature of Regulation of ACCase
at transcriptioal level? .................................................................. 12

Biosynthesis of Erucic Acid: How do fatty acids
move from the pIastid to triacylglycerol ........................................ 17

What Are the Limiting Factors for the Accumulation

ofTAG in Oilseeds? ....................................................................... 23

Literature Cited ........................................................................................ 31
CHAPTER 2: Understanding in Vivo Carbon Precursor Supply

for Fatty ACld Synthesis in Leaf Tissue ................................ 39

Abstract .................................................................................................. 39

Introduction ............................................................................................. 41

Materia' and Methods ............................................................................. 44

Results ................................................................................................... 50

Discussion .............................................................................................. 63

Literature Cited ....................................................................................... 75
CHAPTER 3: Isolation and Characterization of an Arabidopsis

Biotin Carboxylase Gene and its Promoter ----------------------- 79

Abstract .................................................................................................. 79

I ntTOd UCtion ............................................................................................. 81

vi

Material and Methods ............................................................................. 82

Results ................................................................................................... 87
Discussion ............................................................................................ 101
Literature Cited ..................................................................................... 108
CHAPTER 4: The Biosynthesis of Erucic Acid in Developing
Embryo Of Brassica rapa L. ................................................. 1 13
Abstract ................................................................................................. 1 13
[ntroduction ........................................................................................... 1 15
Material and Methods ........................................................................... 117
Results .................................................................................................. 1 19
Discussion ............................................................................................. 132
Literature Cited ..................................................................................... 140
CHAPTER 5: Supply of Fatty Acid Is One Limiting Factor in the
Accumulation of Triacylgycerol in Developing
Embryos .............................................................................. 144
Abstract ................................................................................................. 144
Introduction ........................................................................................... 145
Materia| and Methods ........................................................................... 146
Results .................................................................................................. 149
Discussion ............................................................................................. 156
Literature Cited ..................................................................................... 161
CHAPTER 6: Conclusions and Future Research Perspective --------------- 164
COHC'USiOﬂS .......................................................................................... 164
Future Research perspective ................................................................... 165
Literature Cited ..................................................................................... 172

vii

LIST OF TABLES

Table 4.1 The ratio of radiolabel in oxidative cleavage fragments
of [“C]labeled fatty acids from incubation of B. rapa

embryos with [1_14C]acetate .................................................... 125

Table 5.1 Fatty acid composition of TAG from U. carpinifo/ia and

U. parvifolia embryos at 10 days after ﬁrst collection -------------------- 153

Table 5.2 Rate of 14C-TAG synthesis upon addition 2.1 mM

14C-decanoic acid .......................................................................... 153

viii

Figure 1.1

Figure 1.2

Figure 1.3

Figure 1.4

Figure 1.5

Figure 1.6

Figure 1.7

LIST OF FIGURES

The general pathway from acetyl-CoA to the

accumulation of TAG ................................................................. 6

Simpliﬁed scheme of proposed pathways that might
generate acetyl-CoA for fatty acid synthesis in leaf

tissue ....................................................................................... 8

Simulated models for kinetics of radioactivity
accumulated in fatty acids with different

concentrations of intermediate pools 11

The general pathway for the synthesis

of erucic aCid .......................................................................... 20

Flow chart of how distribution of radioactivity
within acyl chain is determined and the ratio of

cleaved fragments was obtained ............................................... 22

A schematic representation of the method and
logic used in Chapter 4 to understand the
movement of oleic acid from the plastid

and its elongation to erucic acid ............................................... 24

Proposed pathway for the synthesis of

tricylglycerols in oilseeds ............................................. . ............ 26

ix

Figure 2.1

Figure 2.2

Figure 2.3

Figure 2.4

Figure 2.5

Figure 2.6

Figure 3.1

Figure 3.2

Figure 3.3

The molecular ion distribution of 18:0 and
16:0 from Arabidopsis labeled by 13C02 at

one hour time point ................................................................. 52

Determining the rate of fatty acid synthesis in

Arabidopsis by continuos labeling with 13CO2 ------------------------------ 54

Some important parameters concerning plant
growth during the sampling period in the pulse-
chase experiment to determine the rate of fatty

acid synthesis in the dark ........................................................ 56

The pulse-chase labeling with 13002 to determine
the rate of fatty acid synthesis in the dark .................................... 58

Pulse-chase labeling of Arabidopsis with “‘002
to determine the lag phase ....................................................... 60

Continuos labeling of Arabidopsis with 1“C02 to

determine whether there is a slow ﬁlling pool ------------------------------ 62

Structure and sequence of the biotin carboxylase

gene from Arabidopsis ............................................................. 89

Northern analysis of BC mRNA levels in F (ﬂower),
L (leaf), R (root), and S (silique) from Arabidopsis ------------------------ 95

Structure of BC promoter-GUS deletion constructs

used for Arabidopsis and tobacco transformation ------------------------- 96

Figure 3.4 Localization of GUS expression in transgenic

Arabidopsis transformed by construct BF380 -------------------------------- 97

Figure 3.5 GUS activity in roots, leaves, ﬂowers and siliques

of independently transformed Arabidopsis plants ------------------------ 99

Figure 3.6 Comparison of GUS activity in transgenic tobacco
suspension cells transformed with different BC
promoter and 35S GUS constructs ............................................ 100

Figure 3.7 Changes of GUS activity during growth of tobacco

suspension cells transformed by different constructs ------------------ 102

Figure 4.1 Accumulation of fatty acids during embryo

development of B. rapa ......................................................... 121

Figure 4.2 Effects of light on the synthesis of 22:1 ,1821,
TAG, and PC ......................................................................... 123

Figure 4.3 Labeling of fatty acids by [“C]acetate in
developing B. rapa embryos ................................................... 129

Figure 4.4 Effect of increasing concentrations of haloxyfop

on fatty acid and lipid synthesis .............................................. 131

Figure 5.1 TAG accumulation and percentage of decanoic acid

in TAG in developing embryos of C. Ianceolata,

xi

U. ca’pir’ifoliaI and U. parvifolia ............................................... 150

Figure 5.2 Rate of 1“C-TAG synthesis under different conditions -------------- 155

xii

LIST OF ABBREVIATIONS

ACCase .............................................. aceyl-CoA camoxwase
ACP .................................................... acy. cam, mote,”

ADP .................................................... adenosine diphosphate

ATP .................................................... adenosinetriphosphate

BC ...................................................... biotin camoxymse

BCCP ................................................. biotin carboxy. came, mote,"

C16 and C18 ....................................... fatty acids with 16 o, 18 carbons

ch. ...................................................... Cmomphy"

CT ...................................................... camoxynmsfemse

DAF .................................................... days aﬂe, ﬂoweﬂng

DAFC .................................................. days aﬂe, the ms, conecﬁon

DAG .................................................... t,,acy,g,yce,o.

DAGAT ............................................... diacy.g.yce,o. acynmsfeme

FA ....................................................... fatty acid

FAME .................................................. my acid methy. we,

so ...................................................... gas chmmatogmphy

Gems ................................................. gas chmmatogmphymss spectmmetry
KAS ..................................................... 3_ketoacy._acy. came, pm,“ synthase
no ..................................................... thin ,aye, chmmamgmphy

pc ....................................................... phosphaﬁdymnne

pDC ..................................................... pymvate dehydmgenase comm
TAG ..................................................... t,,acy.g.yce,o.

GPAT .................................................. We“, 3_phosphate acymnsfeme
LPAAT ................................................. .ysophosphatmic add acyumnsfaase

xiii

FATTY ACID NOMENCLATURE

10:0 .................................................................... decanoic acid
12:0 .................................................................... lauric acid

14-0 .................................................................... myriStIC acid
16:0 .................................................................... palmitic acid
17:0 .................................................................... heptadecanoic acid
13:0 .................................................................... oleic acid

18.1 A9 ................................................................. decanoic acid
18:1 A6 ................................................................. petroselinic acid
18:1 A11 ................................................................ cis-vaccenic acid
18:2 A9.12 .............................................................. linoleic acid
1821“” 12.15 ........................................................... linolenic acid
20:1A11 ................................................................ eicosenoic aCId
22.1w ................................................................ erucic acid

xiv

CHAPTER 1
Introduction

The overall theme of this PhD dissertation has been to obtain a fuller
understanding at both the biochemical and molecular biological levels of the
biosynthesis of fatty acid and triacylglycerol in plants. The importance of fatty acid
and triacylglycerol synthesis is reﬂected in several aspects of plant biology. Firstly,
fatty acids are the essential component of all membrane glycerolipids and form the
central hydrophobic barrier at the core of the membrane. Fatty acids are attached
to both the sn-1 and sn-2 positions of glycerol backbone and a polar headgroup to
the sn-3 position. The resultant amphipathic properties of glycerolipids allow the
formation of membrane bilayers which delineate the cell and sub-cellular
compartments. These compartments not only are the sites of many essential
biochemical processes, like photosynthesis in chloroplasts and respiration in
mitochondria, but also separate certain reactions that make overall regulation and
coordination of metabolisms possible. Diverse glycerolipids species exists in plant
cells through the combination of different headgroups and fatty acids. The
membrane of each individual sub-cellular structure has it own unique lipids classes
that possess distinct fatty acid composition (Browse et al., 1993), like galactolipids
predominantly found in plastids and cardiolipin mainly in mitochondria. The
differences in lipid structure of each sub-cellular membrane system are important
for them to execute speciﬁc functions, even though the exact structure to function

relationships are not fully understood.

Secondly, fatty acids are precursors for the synthesis of cuticular wax that
are an important defense system against pathogens and prevent water loss for
terrestrial plants. A plant hormone, jasmonic acid, is also derived from fatty acid.

Thirdly, fatty acids are rarely found in the form of free fatty acids, instead,
and their carboxyl group is esteriﬁed, for the most part, to glycerol. In addition, fatty
acids cannot be transported between cells, so each cell has to synthesize its own.
Therefore, the fatty acid synthesis and utilization systems must be fully coordinated
and tightly regulated. The regulation of fatty acid synthesis occurs at several
levels. The de novo fatty acid synthesis in plastids determines the overall output
of total fatty acids to meet the demand at a given growth stage. After de novo
synthesis, further modiﬁcations are required to synthesize certain lipid classes.
Proportional distribution of fatty acids among lipid classes and organelles is
necessary for keeping the delicate balance in each living cell and the whole plant.
Currently, relatively little is known about the mechanism of regulation for fatty acid
synthesis in plants.

Finally fatty acid composition of glycerolipids in membranes changes in
responding to different environments. Plants grown at low temperature, have
higher percent of unsaturated fatty acids relative to that of grown at high
temperature. Wax synthesis increases signiﬁcantly when plants are transferred
from humid condition to drier environment. Fatty acid synthesis as a primary
metabolic pathway, its regulation and the origin of substrate for fatty acid synthesis

have been under extensive study in past twenty years, but many questions remain

to be answered.

Triacylglycerols (TAG), mainly accumulate in seeds, and unlike glycerolipids
in membranes, do not perform any structural role but rather serve primarily as
storage form of carbon and energy for the growth of next generation. TAGs are a
major agricultural commodities with an annual value of more than 50 billion USD
(Liihs and Friedt, 1994). More than 80% of vegetable oils are used in the food
industry. There are three practical goals which relate to this ﬁeld of research. First
is to increase the oil content and yield of existing oil crops to meet the demand of
population growth. Traditional and molecular markers guided breeding methods
have been used to achieve higher oil content and yield. However, because of
limited gerrnplasm of cultivated crops and extensive inbreeding, it is difﬁcult to
further increase yields through traditional plant breeding. Since most pathways and
enzymes related to carbon partition are now known, it looks more promising to
manipulate carbon distribution among different pathways in order to channel more
carbon into fatty acid synthesis, thereby increasing an oil yield. Second is to alter
the relative fatty acid compositions in seed oils that are beneﬁcial for human health
and replace some animal fat products. With the knowledge of pathways for TAG
synthesis and genetic engineering, also the health concerns animal fats in human
diets, substantial efforts have been given in altering the composition of vegetable
oil to improve nutritional value of oils consumed. Due to the low melting point of
vegetable oils, the production of margarine and shortening from vegetable oils

requires catalytic hydrogenation to reduce unsaturated fatty acids, and therefore

increase a melting point. This process not only increases the production cost, but
also converts certain percentage of natural cis conﬁgurations to trans isomer which
are suggested to be bad for health. By reducing the activity of stearoy-ACP
desaturase, the percent of saturated fatty acids increased about 20-fold in Brassica
(Knutzon et al., 1992), whose oil might be directly used for the production of
margarine and shortening without the hydrogenation process. On the other hand,
unsaturated fatty acids are beneﬁcial in reducing cholesterol levels. Increasing
unsaturated fatty acids in vegetable oil was achieved through enhancing the
elongation of palmitoyl-ACP to stearoyl-ACP in transgenic B. napus (Bleibaum et
al., 1993). A third goal is to produce special fatty acids in oil crops for industrial
usages and as a renewable raw material to replace fossil fuel. A diverse range of
fatty acid structures has been found in seed oils and a number of these are used
industrially. For example, eruciciacid is used as a slip agent in plastic ﬁlm and
might have a large potential as a precursor to nylon 13,13, a high-temperature
thermoplastic. Ricinolenic acid from castor beans is used in coatings and
lubricants. Lauric acid (12:0) from palm oil is a major ingredient in soaps and
detergents. Because of the limited geographical distribution of palm trees, the
incentive to produce Iauric acid in temperate oil crops resulted in the high Iauric
Brassica obtained by introducing lauroyl-ACP thioesterase from California bay into
Brassica (Voelker etal., 1992). Even though considerable progress has been made
to understand the synthesis of unusual fatty acids and TAG, we still have a lot of

difﬁculties to achieve the desired fatty acid levels that are high enough for industrial

usages.

In this thesis, four major questions were addressed, which span the lipid
biosynthetic pathway, from the origin of acetyl-CoA to the ﬁnal accumulation of
TAG. Each question will be discussed separately in the following paragraphs of
this chapter, which are organized to follow the biochemical pathway of carbon from
CO2 to TAG. An overview of these questions is presented schematically in Figure
1-1 to show their relationship to overall lipid metabolisms. Each question also

represents a chapter of this thesis and a publication or submitted manuscript.

I. What is the Origin of Plastidial Acetyl-CoA?

As illustrated in Figure 1.1, de novo fatty acid biosynthesis in higher plant
cells is exclusively located in the plastid (Ohlrogge et al., 1979), except a very small
contribution from the mitochondrion (Wada et al., 1997). The immediate carbon
precursor for fatty acid biosynthesis is acetyl-CoA, which is used as substrate by
KASlll for the acyl-chain primer reaction, and, after carboxylation to malonyl-CoA
and transacylation to malonyI-ACP, for the acyl-chain elongation steps. The ﬁrst
question we address in this thesis is: how is acetyl-CoA generated in plastids.
Because different tissues may not have the same mechanism to produce their
acetyl-CoA, we will limit our scope for this question to green leaf tissue.

The biosynthetic origin of acetyl-CoA for plastid fatty acid synthesis has been
addressed by many reports, but remains unsolved for the most part due to lack of

deﬁnitive evidence to prove or disprove the proposed possibilities (Givan, 1983;

 
   
 

 

Q 2: What is the nature of
regulation of ACCase at

= Q 1: What is the origin
of plastidial acetyl-CoA?

 

 

 

 

 

lllllll

 
 

Free Acetate
pool

   
 
 
 
  
 
 

 

Q 4: What are the limiting
factors for the accumulation
of TAG in oilseeds?

 

 

Cytosol

 
    

18:1:COA

Q 3:Biosynthesis of erucic
acid: How do fatty acids
move from the plastid to
TAG?

Intermediate
18:1 pool

20:1-CoA " l8:I-CoA

 

 

   
 

 

   

TAG

   

Oilbody

  

22: l -CoA

Figure 1-1. The general pathway from acetyl-CoA to the accumulation of TAG. The

major questions addressed in this thesis are indicated.

Liedvogel, 1986). Because acetyl-CoA does not readily cross the plastid
membrane (Brooks and Stumpf, 1965), it must be synthesized inside the plastid in
order to provide the carbon precursors for fatty avid synthesis. Several possible
routes generating acetyl-CoA for fatty acid synthesis have been suggested as
shown in Figure 1.2. Firstly, acetyl-CoA may be produced from pyruvate and 00A
by pyruvate dehydrogenase (PDC) inside the chloroplasts. PDC activity has been
clearly demonstrated in pea, spinach, and maize chloroplasts, even though at
relatively lower level than corresponding leaf mitochondria (Camp at al., 1985;
Liedvogel, 1985; Treede etal., 1986a; Trude and Heise, 1985). Mitochondrial PDC
is regulated by both end product inhibition and covalent modification through a
phosphorylation Idephosphorylation cycle, but the chloroplast PDC resembles the
E. coli PDC in that there is no evidence of any covalent modiﬁcation (Camp at al.,
1985). Pyruvate is most likely derived from 3-PGA which can be produced from
glycolysis or the initial reactions of theCalvin cycle. Although most of the enzymes
needed for glycolysis exists in the chloroplast stroma, phosphoglyceromutase,
required to convert 3-PGA to 2-PGA, may not have sufﬁcient activity in chloroplasts
to produce enough pyruvate in order to sustain the rate of fatty acid synthesis.
Therefore, the participation of cytosolic phosphoglyceromutase may be required
(Givan, 1983). This would explain the low incorporation of “‘002 into fatty acids by
isolated chloroplasts. An alternative source of pyruvate might be as a direct
product of ribulose-biphosphate carboxylase in chloroplasts. However, this source

can only count for a quarter of fatty acids synthesized (Andrews et al., 1991).

Mitochondria

Pyruvate

 

 

 

Figure 1-2. Simpliﬁed scheme of proposed pathways that might generate acetyl-

CoA for fatty acid synthesis in leaf tissue.

Secondly, acetyl-CoA may be synthesized from free acetate which is
imported into chloroplasts. Several lines of evidence support this view. First,
exogenous acetate is rapidly incorporated into fatty acids of isolated chloroplasts
and the rates of incorporation are equivalent to the in vivo rate of fatty acid
synthesis. In addition, acetate is three fold or more effective than pyruvate (or
“002 ) as an in vitro substrate for fatty acid synthesis in isolated chloroplasts
(Roughan et al., 1979; Springer et al., 1989). Second, high levels of acetyl-CoA
synthetase activity have been reported in isolated chloroplasts (Roughan and
Ohlrogge, 1994), and chloroplasts have been reported as the dominant site of
acetyl-CoA synthetase activity (Kuhn et al., 1981). Third, acetate concentrations
in leaf tissue are reported, ranging from 0.05to1.4 mM (Kuhn et al, 1981; Liedvogel
1985; Treede et al., 1986b; Roughan, 1995). Liedvogel and Stumpf (1982) were
ﬁrst to propose a mechanism to explain how free acetate was generated and used
for fatty acid synthesis. They suggested that pyruvate is converted to acetyl-CoA
in the mitochondrion. The acetyl-CoA is hydrolyzed to acetate by a mitochondrial
acetyl-CoA hydrolase and the free acetate then diffuses through the cytosol to the
chloroplast where it is converted to acetyl-CoA by the acetyl-CoA synthetase and
becomes available for fatty acid synthesis. This view is supported by the facts that
mitochondria have an active pyruvate dehydrogenase and acetyl-CoA hydrolase.
But this mechanism of acetyI-CoA production, as of yet, has not been supported by
in vivo experiments. Roughan (1995) concluded that it seems probable that

endogenous acetate would be the direct source of acetyl-CoA for fatty acid

synthesis in leaves. An important part of the argument for acetate as the dominant
substrate for fatty acid synthesis in vivo is that the bulk tissue concentrations are
above the concentration for acetate saturation for fatty acid synthesis. In addition
to the mechanisms discussed above, acetylcamitine was suggested as a possible
way to deliver acetyI-CoA into plastids (Woods et al., 1992).

It is clear from the above that a consensus is not yet available concerning
the origin of acetyl-CoA for fatty acid synthesis. Although several pathways can be
supported based on in vitro data, in vivo evidence is lacking in almost all cases. In
chapter 2, the question of whether free acetate is a substrate for fatty acid synthesis
in leaf tissue is examined in vivo. We approach this question ﬁrst by accurately
measuring the rate of fatty acid synthesis in leaf tissue. Knowing the synthesis rate
and the pool size of free acetate, the turnover time of free acetate pool can be
calculated, if it serves as the primary source for acetyl-CoA. As shown
schematically in Figure 1.3, when plants are given 1“002, the lag phase of
radioactivity accumulation in fatty acids will reﬂect the pool size of intermediate
substrates through which carbon must ﬂow. Thus, by examining the kinetics of 1“C
incorporation into fatty acids and the calculated turnover time of free acetate pool
as shown in Figure 1.3, we can test the possibility that the bulk acetate pool is the
main source for the synthesis of acetyl—CoA.

Accurate rate of fatty acid synthesis is crucial to calculate the turnover time
of the free acetate pool. In chapter 2 we also demonstrate a new method that can

accurately measure the absolute rate of fatty acid synthesis both in the light and in

10

Intermediate ‘

l l“C02 —> —> —-> I —> ——> —-> l“C-Fatty Acids
poo

   

2500 ,
l
‘ ‘ 0.05am .
l 2 2000. 1
; g 1500i 1
7 "3 0.50 mM
l “I
, .2 ,
j R 1000; ,
, 1'; 1.00 mM ‘
l 0
; .15
l E 500 l- .
l
l l
i 0 - I" ' v ' 7 1' v r TT—“T‘ l l ' "'—l

0 5101520253035404550556065

Labeling time (min) ;

Figure 1.3. Simulated models for kinetics of radioactivity accumulated in fatty acids
with different concentrations of intermediate pools. The rate of fatty acid synthesis
is 38nmols/min/gfw. The lag phase before linear kinetics is observed will depend
on the size of the pools through which carbon must ﬂow before incorporation into

fatty acids.

dark. And ﬁnally, short and long term carbon dioxide labeling is used to measure

the actual lag phase.

II. What Is the Nature of Regulation of ACCase at transcriptioal level?

Moving one step further down the pathway from acetyl-CoA as shown in
Figure 1.1, the ﬁrst committed reaction of de novo fatty acid biosynthesis is
catalyzed by acetyI-CoA carboxylase (ACCase). Malonyl-CoA is synthesized from
acetyl-CoA and bicarbonate via the two step reaction shown in the equations below.

Mg++

1. ATP + HCO3' + biotin-BCCP “=5 ADP + Pi +COz-biotin-E
biotin carboxylase

2. COz-biotin-BCCP + acetyl-CoA *—-‘ biotin-BCCP +malonyl-CoA
carboxyltransferase

Two types of ACCase are found in nature. Multisubunit or heteromeric ACCase is
composed of four dissociable subunits: biotin carboxylase (BC), biotin carboxyl
carrier protein (BCCP), or- and B-biotin transcarboxylase (a- CT and B—CT)
(Guchhait et al., 1974). Multifunctional or homomeric ACCase is a large single
polypeptide containing the same three function domains as the multisubunit
ACCase, but in one polypeptide.

The structures of plant ACCases have been under intensive studies in the
past 15 years. Early attempts to purify ACCase from plant tissues yielded multiple

subunits on PAGE under denaturing conditions but these were thought to have

12

resulted from endogenous proteinase degradation (Hawvood, 1987). However, with
proteinase inhibitors ACCase was puriﬁed from both dicots and monocots (Egin-
Buhler et al., 1980; Finlayson and Dennis, 1983; Slabas and Hellyer, 1985; Hellyer
et al., 1986; Charles and Cherry, 1986) and in all cases the ACCases puriﬁed had
molecular mass in the range 220-240 kDa. Deduced amino acid sequence of
ACCase from Cyclotella cryptica (Roessler and Ohlrogge, 1993), alfafa ( Shorrosh
et al., 1994), and Arabidopsis (Roesler et al., 1994) also have a molecular mass
of 220-250 kDa. So until 1993, most researchers had concluded that plant ACCase
was a large (>200 kDa) mutifuntional protein similar to that of animal and yeast.
However, in 1993, Sasaki and co-worker demonstrated that the chloroplast genome
of peas encodes a subunit of an ACCase with structure related to the (3 subunit of
the carboxyltransferase found in the multisubunit ACCase of Escherichia coli
(Sasaki et al., 1993). In the following several years, a BC subunit (Shorrosh et al.,
1994), BCCP (Choi et al., 1995), and or-CT (Shorrosh et al.,1996) were cloned.
Among the four subunits, BC, BCCP, and oi-CT are nuclear encoded, whereas B—CT
is plastome encoded. The four subunits are assembled into a complex of 650-700
kDa which is easily dissociable. It has now been clariﬁed that dicot and most
monocots have both forms of ACCase, the multifunctional form localized in cytosol
and the mutisubunit form with at least four subunits localized in plastids (Alban et
al., 1994; Konishi et al., 1996; and Roesler etal.,1996). It is the mutisubunit plastid
form of ACCase that provides malonyl-CoA for fatty acid synthesis. The structures

of ACCase in Gramineae are different in that these species lack the multisubunit

13

form of ACCase and instead have two type of mutifunctional ACCase. An
herbicide-sensitive form is located in plastid and a resistant form in the cytosol.

It has been long known that fatty acid synthesis is tightly regulated. ACCase
is the ﬁrst committed step for de novo fatty acid synthesis, therefore, it is logical to
speculate that ACCase is an important site of regulation. In animals, ACCase is
regulated by covalent modiﬁcation (phosphoralation/dephosphoralation) and also
allosterically regulated by its reaction product and end products of fatty acid
synthesis. There are also complex transcriptional and posttranscriptional controls
of ACCase expression in animals which involve transcriptional induction and
differential splicing of mRNA in response to diet and developmental state (Lopez-
Casillas et al., 1989; Luo et al., 1989). The tight overall control of acyl lipid
synthesis by ACCase has also been shown in yeast and E. coli (Hasslacher et al.,
1993; Jackowski et al., 1991 ).

ln plant leaf tissues, the rate of fatty acid synthesis is at least six-fold higher
than in the dark. There has been great interest to ﬁnd out which enzyme(s) play the
regulatory role in the pathway of fatty acid synthesis. As mentioned earlier, ACCase
is located at the ideal site of regulation. There was early indirect evidence implying
that ACCase is an important regulatory enzyme. When isolated chloroplasts were
incubated with radio-labeled acetate or pyruvate, acetyI-CoA was formed in the
dark, but not malonyl-CoA and fatty acids. Only in the light were malonyl-CoA and
fatty acids labeled (Nakamura and Ymada, 1979). ACCase from chloroplast and

wheat germ were inhibited by ADP whose concentration was higher in the dark than

14

in the light (Eastwell and Stumpf, 1983). The direct evidence to prove that ACCase
is indeed the regulatory enzyme came from the work of Post-Beittenmiller et al.
(1990) through measuring the pools of intermediates during manipulation of the
rates of fatty acid synthesis by light in spinach leaves. When comparing light vs
dark incubations, it was found that there were large increases in the acetyl-ACP
and commensurate decrease of malonyl-ACP from light to dark. From these and
other observations it was concluded that light/ dark control of leaf fatty acid
synthesis resides in ACCase. In a following study (Post-Beittenmiller et al.,1992),
short chain acyl-CoA pools were measured in isolated pea and spinach
chloroplasts. Malonyl-CoA and malonyl-ACP could be detected only during light
incubations and increased as fatty acid synthesis increased. In contrast, both
acetyl-CoA and acetyl-ACP were detectable in the absence of fatty acid synthesis
and acetyl-ACP decreased with the increased rates of fatty acid synthesis. These
data provided direct in situ evidence that ACCase plays a regulatory role in fatty
acid synthesis. A quantitative measure of the importance of ACCase in the control
of ﬂux through the pathway of fatty acid synthesis was obtained by using the
metabolic control theory (Page et al., 1994). They took advantage of the fact that
plastidial ACCases of barley and maize leaves are sensitive to the herbicide
ﬂuazifop and sethoxydim. The sum of all the ﬂux control coefﬁents in a metabolic
pathway is one, and the closer an individual ﬂux control coefﬁcient is to one, the
more control that enzyme exerts in regulating the metabolic pathway. Apparent ﬂux

control coefﬁcient of about 0.58 and 0.52 were determined for acetyI-CoA

15

carboxylase in barley and maize leaves, respectively. These results clearly showed
that acetyI-CoA carboxylase is a major ﬂux controlling enzyme for light-stimulated
lipid synthesis in leaf tissues. ACCase is also the site of feedback regulation by
reducing fatty acid synthesis of tobacco suspension cell in response of to
exogenous lipids (Shintani and Ohlrogge, 1995). It is clear that plastid ACCase
activity is regulated at the biochemical level, but the mechanisms of how ACCase
is regulated has not been resolved yet. Savage and Ohlrogge (1999) observed that
the phosphorylation of B-CT of pea ACCase resulted in the decrease of ACCase
activity. So plant ACCase may be also regulated by covalent modiﬁcation
(phosphoralation/ dephosphoralation).

Control of ACCase gene expresiicm. In tobacco and castor, expression of
the BC gene is much higher in developing seeds than in leaf or root (Elborough et
al.,1996; Roesler et al.,1996; Shorrosh et al., 1995). In developing castor and
Brassica napus seeds, ACCase activity as well as expression levels of BC and
BCCP correlate with oil deposition (Roesler et al.,1996). Using in situ hybridization
analysis, BC and BCCP were found to be expressed in a coordinate fashion and
were correlated with oil accumulation during seed development in Arabidopsis (Choi
et al.,1996). The plastid ACCase subunits, BC, BCCP, and or-CT are encoded by
nuclear genes, whereas B-CT is encoded by a plastid gene. It is not known how
overall ACCase expression is controlled or how levels of the four subunits are
coordinately regulated. Nor is there research to investigate what cis acting

elements are responsible for the regulation of ACCase. To begin to address these

16

questions in chapter 3, we describe the isolation and characterization of a biotin
carboxylase genomic clone from Arabidopsis. Furthermore, we report initial studies

to determine the cis acting elements of the promoter of the BC subunit of ACCase.

lll. Biosynthesis of Erucic Acid: How do fatty acids move from the plastid to
triacylglycerol

The fatty acids synthesized in plastids have two fates; one is to remain inside
plastids for glycerolipid synthesis; the other is to be exported outside plastids. The
exported fatty acids are usually 16:0, 18:0, and 18:1, and they can be used in
several ways, such as the synthesis of membrane lipid, wax, and triacylglycerol.
Sometimes the fatty acids go through extensive modiﬁcation during these
processes. In oilseeds, most of the exported fatty acids are used for the synthesis
of triacylglycerol as a means of storing carbon and energy. A diverse range of fatty
acids can be found in seed oil. Some of the fatty acids are called unusual fatty
acids due to their limited distribution among plant species and their chemical
structures differing signiﬁcantly from those of the common C16 and C,8 acyl moieties
such as palmitic (16:0) and oleic acids (18:1A9). Structural divergence in these acyl
moieties include the variations in carbon chain lengths, position and numbers of
double bond, additional function group such as hydroxyl or epoxide ring, and the
combination of above. Here, we will limit our discussion to the biosynthesis of
erucic acid (221”), one of the unusual very long chain fatty acids.

Erucic acids is an important industrial material used as lubricants or as slip

17

agents in plastic ﬁlm manufacture. It may also have a larger potential as a
precursor of nylon 13, 13, a high temperature thermoplastic (Ohlrogge, 1994).
Erucic acid (cis-13-docosenoic acid) and its homolog, cis-11-eicosenoic acid, are
commonly found in the seed oils of the Crucifereae. The seed oil of Brassica spp.,
Crambe abyssinica, Sinapsis alba, and Lunan'a annua usually contain 40-60%
erucic acid while seed oil of nasturtium (Tropaeolum majus) contains as much as
80% erucic acid (Gustone et al., 1994; Luhs and Friedt, 1994). Commercially,
erucic acid has usually been derived from rapeseed oil. However, even for high
erucic rapeseed oil, the current level of 50% erucate in its triacylglycerol is not
sufﬁcient to compete well with alternative source of lubricant oils from
petrochemicals because of the high cost of puriﬁcation (Ohlrogge, 1994). Genetic
manipulation to increase the erucic acid content in rapeseed oil has attracted great
interests due to its economic potential. To achieve this goal, we have to better
understand the biosynthetic pathway of erucic acid.

The reactions leading to the synthesis of erucic acid are, for the most part,
well understood. In the developing embryos of several erucate accumulating
species, it was demonstrated that erucic acid is synthesized by the elongation of
oleic acid rather than by de novo synthesis (Appleby, 1974; Downey et al., 1964;
Ohlrogge et al., 1978; Pollard et al., 1980a and 1980b). This conclusion was
deduced from the observation that exogenous [“Clacetate was preferentially
incorporated into the carboxyl-terminal carbons of the long-chain fatty acids rather

than the methyl terminal 18 carbons. These results inferred that the addition of the

18

carboxyl-terminal four carbons were catalyzed by different enzymes and at different
compartment, comparing to the synthesis of methyl terminal 18 carbons of erucic
acid. Many reports have conﬁrmed that the location of oleoyl elongation system
was extra-plastidial, being associated with oil bodies or microsomal membranes
(Agrawal and Stumpf,1985a and 1985b; Fehling et al., 1991; lmai et al.,1995; von
Wettstein Knowles, 1993; Créach et al., 1993). In Arabidopsis, a mutation (FAE1)
result in reduced level of seed very long chain fatty acids and deﬁciencies in
elongation activities (James and Dooner, 1990). The gene has been cloned and
the predicted amino acid sequence shares homology with those of other
condensing enzymes, suggesting that it encode a fatty acid elongase (James at al.,
1995). Thus, as summarized in Figure 1.4, biosynthesis of erucic acid can be
described as oleate, synthesized in the plastid, is then exported to the cytosol,
where, presumably in the endo-membrane system, it is elongated via malonyl-CoA-
requiring elongases to C20 and longer-chain monounsaturated fatty acids.
However, the nature of the immediate18:1 donor needs to be clariﬁed.

It is generally believed that the end-product of newly synthesized oleic acid
exported from plastids is oleoyl-CoA. This oleoyl moiety can be elongated directly
to erucic acid in the cytosol through successive additions of two carbons derived
from malonyl-CoA. However, in an oil body fraction from developing rapeseed,
Hlousek-Radojcic et al.(1995) observed in vitm that radioactivity from oleoyl-CoA
was incorporated into eicosenoate and erucate at least 2.5-fold more slowly than

from malonyl-CoA. Furthermore, radioactivity from oleoyl-CoA was rapidly diluted

19

 

    
  

Plastid

Cytosol

 

O
Oleic acid (18:1) H II II
C C - C
/ \ / \ ACytosolic ACCase / \
Elongase ‘0 CH, S-CoA ‘ CH3 S-CoA
Malonyl-CoA Acctyl-CoA

9*: Si R A “k S'K at at 5': 0"
C/\

CH3 CH2 CH2 CH2 CH=CH CH2 CH2 CH2 CH2

 

O
Eicosenoic acid (20:1) 0 0 0
ill! l
/ \ / 4Cytosolic ACCase / \
Elongase -0 CH, S-CoA ‘ CH, S-CoA
C CH CH CH CH C H CH CH CH 0H
/’/\’/\2 /\’_ /\’/’§\’/\’/\’/\2
CH3 CH, CH, CH, CH—CH CH, CH, CH, CH, CH, Cg
O

Erucic acid (22:1)

Figure 1.4. The general pathway for the synthesis of erucic acid. Oleic acid is ﬁrst
produced in the plastid and later elongated outside the plastid to produce

eicosenoic and erucic acids.

20

upon the formation of eicosenoyl-CoA and the elongation could proceed without the
addition of exogenous oleoyl-CoA. Based on these in vitro observations, they
concluded that oleoyl-CoA is not the immediate substrate for elongation. Instead
they proposed that the intermediate oleoyl donor for the elongase may be either a
lipid or unesteriﬁed fatty acid. In addition, when intact Brassica (high erucic)
embryos were incubated with [“C]acetate, phosphatidylcholine (PC) was always
heavily labeled at early time points, with oleate constituting over 90% of [“C]labeled
fatty acid esteriﬁed to PC. We considered that this oleoyl-PC might contribute to the
synthesis of erucic acid, either via a mechanism of direct acyl transfer as proposed
by Hlousek-Radojcic et al (1995), or via the acyl-exchange between acyl-CoA and
PC, as ﬁrst reported by Stymne and Stobart (1984). The involvement of PC for the
synthesis of erucic acid can be envisioned for at least two physiological reasons.
First, the newly synthesized oleic acid is not directly accessible to the compartment
where the elongase is located, and PC is used as delivering carrier of oleic acid.
Second, because free fatty acids are toxic, PC may act as a temporary storage for
oleic acid before the elongation to erucic acids.

In this chapter, we have two goals in mind; one is to evaluate the existence
of intermediate oleoyl donor pool for the synthesis of erucic acid proposed by
Hlousek-Radojcic et al. (1995); the other is to examine the possibility that PC
serves as oleoyl donor. Developing Brassica rapa embryos were incubated with
1“C-acetate, labeled tissue would be collected at different time points, fatty acids

were separated from each time point, and the ratio of two fragments cleaved at the

21

(extracted from embryos laleled with [1-‘4C]acetate)
Total Lipids

l Fatty acid methyl-esters l

 

 

i Separated by argentation TLC
Argentation plate

ﬂ.
an. Individual fatty acid methyl esters

are located and quantiﬁed by
rim 2m

 

Instant Imager

  
    
 

 

m Fatty acid methyl esters are cleave at the
position of double bond by permanganate—
periodate oxidation.

Fragments are separated €59 C39 C? cn,.(cn,),.c.on
with TLC and quantiﬁed II :
by Instant Imager

 

 

 

 

 

 

 

 

 

 

 

   

C11
........................... Cg
HOf-cnrrcngrcnrcnrcnrcnrﬁao ~CH,
o i
v ...............
ﬂ HO-ﬁ-CHACHJrCHﬂrﬁ-O-Cﬂ, a
C13 0 ,
. , . Ho-c-CHACHM-ecn,
C9 0 ||
C] 1 O 0
RATIOC20:1:,—.,_ 11
C9
Calculate the ratio of radioactivity C9
in the two fragments of acyl chain. RATIOCHH:
C9

Figure 1.5. Flow chart of how distribution of radioactivity within acyl chain is

determined and the ratio of cleaved fragments was obtained.

22

double bond position would be calculated (Figure 1.5). The analytical method was
designed to elucidate these questions as explained in Figure 1.6. Since acetate
can diffuse into tissue within 5 seconds, if newly synthesized 18:1 is directly
elongated to 22:1 , radioactivity should be evenly distributed along the carbon chain.
Regardless of whether the carbons are added in plastids or cytosol, the distribution
of radioactivity along the acyl chain will remain constant with time course. However,
if newly synthesized 18:1 ﬁrst enters an intermediate pool and then is elongated to
22:1, an uneven distribution of radioactivity along the carbon chain will result from
different sites of the carbons added will reﬂect in the ratio decrease with course for
20: and 22:1.

In addition, in Chapter 4 we have examined the inﬂuence of light and of an
inhibitor of the homodimeric ACCase on erucic acid biosynthesis. Taken together,
our results show an additional level of complexity in the biosynthesis of erucic acid,
in that the supply of oleoyl groups for chain elongation is a combination of the
release of oleate from a large intermediate lipid pool, probably phosphatidylcholine

and the direct provision of newly synthesized oleate from the plastid.

IV. What Are the Limiting Factors for the Accumulation of TAG in Oilseeds?

After considering the origin of acetyl-CoA, the regulatory role of ACCase,
and the pathway of erucic synthesis, we move to the end of pathway shown in
Figure 1.1, and ask what determines how much oil accumulates in oilseeds. To

meet the increasing demands for vegetable oil, considerable research is oriented

23

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Plastld
(:33- (CI-12) 7-CH=CH— (m2) 7-C-OH Intennedlate 18.] p001
II >:::::::::=:::::::"
0 CytOSOI ooooooooo :ooooooo::
——13°1 Time °::::::::.
.e:s--... ooooooooo
= 1 HO-C-CH3-(CH2).,-C-OH = 1
,-t .. '9 II H 00.69%...
: 000000000
0 0 000000000
-‘ CH,-(CH,),-C-0H
.-L‘.p'. ..- 6-380108006
:1 || _
tab'aia-D'..- O GOOOOOOO.
0333133“ .33323333'
l V i
3‘ essoeeeaeee Tim 00000000000
3 M, m, , ,4, 3 ooooooooou
: tween...“ 20:1 ooooooooooo g»
.5 =1.2 — =17 3
=- "' e
g ., : .~ .- t .. -.~. HO-C-CHS-(CHQS-CHfC-OH 000000000 8-
g _ II II QIOOQOOODOO a
w - O 0 celea‘vboaeaw .3
= ‘La'u.-§ov'." c ‘wiivi’i‘uii a
,2, =12 =12 u)
3 '- C"3*¢"2)TC'°" ;::::3:::. .5.”
§ LOIUCO... I 0000.0... 0
'5 i 0 V I ‘3
'°".:‘?:"f“f - 'zzzzzzzzzm. ‘5
t.;;;::i:;3.3.‘ .22_-1. Tune 0000000000". :5
= 1.4 "2.3
.- f s- 27 .. .- c- »= ' HO-C—CH3-(CH,)8-(CH,)3-C-0H 0:;::;;;;.
..,-... H || 000000000
0 0 "straw-‘3'”.
663%;vaawsavv
_ CHJ—(CH,)7-C-OH = 1 4
— 1.4 .
.z»..r-l~----..r-.r Ii accessrceo‘
.. ~- ..... o
uUlt'Ibv44ﬁH
\/

 

 

 

 

 

 

 

Figure 1.6. A schematic representation of the method and logic used in Chapter
4 to understand the movement of oleic acid from the plastid and its elongation to
erucic acid. The ratio of radioactivity in the two fragments of fatty acid change

with time for indirect elongation pathway.

24

in two directions; one is to increase the oil content and yield of existing oil crops;
the other is to produce novel fatty acid in oil crop through genetic engineering in
anticipation to replace petroleum.

In plants, the biosynthesis of storage triacylglycerol (TAG) occurs at high
levels primarily in the seeds, but there is a wide range in the levels of TAG which
accumulate in different plant species. For example, seeds of species of Zea,
Hordeum or Pisum usually contain less than 5-10 % TAG by dry weight whereas
many other species such as castor accumulate over 50% TAG in seeds. On the
other hand, leaves, roots, and other vegetative tissues usually contain less 10%
lipids by dry weight, most of which is polar lipids. In contrast, neutral lipids
constitute over 95% of its lipids in oilseeds. The regulatory or metabolic factors
which inﬂuence this very wide range of oil accumulation in seeds are currently
unknown (Ohlrogge and Jaworski, 1997).

The pathway of TAG synthesis, orthe so-called Kennedy pathway (Kennedy,
1961), was found to operate in the synthesis of TAG in plants as early as 1962
(Barron and Stumpf, 1962) as shown in Figure 1.7. The ﬁrst reaction is catalyzed
by glycerolphosophate acyltransferase (E.C.2. 3.1 .1 5), the second by 1-acylglycerol-
3-phosphate acyltransferase (E.C.2.3.1.51), the third by phosphatidate
phosphatase (E.C.3.1.3.4), the fourth by diacylglycerol acyltransferase
(E.C.2.3.1.20) (Stymne and Stobart, 1987). Fatty acids are ﬁrst synthesized in
plastids, whereas the acyl-CoAs are formed on the outer membrane of the plastid

envelope by the action of acyl-CoA synthetase (Roughan and Slack, 1977; Joyard

25

Cell wall

 

Cytosol

 

lag—b ——> ——> —-> Glycero-S-phosphate

 

   
        
   
 

 

 

Endoplasmic reticulum Reaction 1

Glycerolphosphate
acyltransferase

 

Polar lipids ‘
Reaction 2 1-acylglycerol-3-phosphate
acyltransferase

Phosphatidate
phosphatase

     
  

Reaction 3
Reaction4 7 '
Diacylglycerol
acyltransferase

TAG

Figure 1.7. Proposed pathway for the synthesis of tricylglycerols in oilseeds. Some

possible limiting factors are marked by dotted box.

26

and Douce, 1977). The glycerol phosphate is mainly derived through
dihydroxyacetone oxidoreductase (Finlayson and Dennis, 1980), with perhaps a
minor contribution through glycerol kinase (Gurr et al., 1974). The site of sequential
addition of three acyl groups to glycerol backbone is located in ER or oilbody. The
diacylglycerol, can be used either for polar lipid synthesis or for the synthesis of
TAG. Therefore, diacylglycerol acyltransferase (DAGAT) is the only unique enzyme
for TAG synthesis. Since diacylglycerol acyltransferase locates at the branch point
that channels diacylglycerol to TAG synthesis, one potential explanation for TAG
synthesis primarily in seeds is that DAGAT is only expressed in this tissue.
However, there are several observations which argue against this view. For
example, DAGAT activity was found in spinach leaves (Martin et al.,1983) and was
primarily associated with chloroplast envelopes (Martin et al.,1984). Roughan et
al.(1987) reported that signiﬁcant amounts of TAG were synthesized when palmitic
acid was applied to the upper surface of expanding spinach leaves. The level of
neutral lipids (mainly TAG) increased at least 3 fold during protoplast isolation from
Arabidopsis leaves (Browse et al., 1988). Finally, ozone-fumigated spinach leaves
produced high proportions of TAG (Sakaki et al.,1990). These data together
suggest that DAGAT not only occurs in leaves, but also that leaves have the ability
to synthesize TAG.

Enormous efforts have been put into the isolation of DAGAT, but have not
been successful until recently. In 1998, a mouse cDNA sharing similarity with acyl-

CoA: cholestrol acyltransferase, when expressed in insect cells, resulted in high

27

levels of DAGAT activity and it is believed that this cDNA encodes a DAGAT
(Cases et al., 1998). Several Arabidopsis ESTs were found sharing signiﬁcant
similarity with the mouse DAGAT. Two putative DAGAT genes were isolated from
Mortierella ramanniana through protein sequencing, which are not related to mouse
DAGAT (Lardizabal et al., 1999). Stymne and co-workers (1999) reported that a
transacylase can transfer acylgroups from PC or DAG into TAG. So far, all the new
ﬁndings of plant DAGATs are at prelimary stage, and it is hard to draw any
conclusions at present time regarding the quantitative contributions of the enzymes
encoded by the newly discovered genes.

Considering that over 30 reactions are required to convert acetyl-CoA to
TAG, there could be many steps or genes which control the yield of end product
TAG. In order to begin to dissect possible limiting factor(s) in the pathway of TAG
biosynthesis, it is useful to conceptually divide the pathway into two parts: 1) the
production of acyl chains which occurs in plastids (source), and 2) the utilization of
acyl chains for glycerolipid synthesis in the ER and oilbody (sink). The
accumulation of TAG by oilseeds may be a response to the high rate of fatty acid
synthesis. There is so far no research speciﬁcally looking into this possibility, but
some observations are in agreement with the concept. In Chlamydomonas,
addition of exogenous PC Iiposomes to cultures caused a 10-fold increase in TAG
accumulation (Grenier et al., 1991). Another example comes from comparisons of
oleaginous (oil-accumulating) versus nonoleaginous yeast (Botham and Ratledge,

1979). They concluded that differences between these two types of species are

28

controlled by the production rather than utilization of fatty acids. On the other hand,
some evidence implies that utilization of fatty acids can increase the rate of fatty
acid synthesis. When a plant 12:0-ACP thioesterase was overexpressed in E. coli
cells, 10-fold more fatty acid accumulated than controls (Ohlrogge et al., 1995). In
another experiment, massive overexpression of membrane proteins resulted in no
change of membrane’s phospholipid to protein ratio, but rather more fatty acid was
produced to accommodate the excess proteins (von Meyenburg etal., 1984; Weiner
et al., 1984). So far, we don't know whether TAG accumulation is driven more
strongly by the supply or the demand for acyl chains or if both mechanisms operate
in oilseeds.

In chapter 5, we asked whether the supply of fatty acid can inﬂuence the
amount of TAG produced in oilseeds. As shown in Figure 1.7, the possible factors
that might limit TAG accumulation are marked red. If the supply of fatty acid is a
limiting factor for TAG biosynthesis, then providing exogenous fatty acid to the
developing embryos should increase the rate of TAG production. Unfortunately,
long-chain fatty acids (LCFA) have very low solubility in aqueous solution, and so
addition of LCFA at concentrations sufﬁcient to increase rates of lipid synthesis is
very difﬁcult. However, embryos of Cuphea Ianceolata, Ulmus carpinifolia, and
Ulmus parvifolia contain high levels of decanoic acid in their TAG (80%, 63%, and
71%, respectively). Decanoic acid is easily dissolved in water at mM
concentrations, and therefore, these species provided a convenient model system

to test the inﬂuence of fatty acid supply on TAG accumulation. The supply of

29

sucrose and glycerol are also evaluated in a similar fashion like fatty acids. Such
in vitro model experiments may provide a useful guide toward the selection of

targets for future metabolic engineering in transgenic plants.

30

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38

CHAPTER 2
Understanding in Vivo Carbon Precursor Supply for Fatty Acid
Synthesis in Leaf Tissue
Abstract

The principal supply of carbon precursors for fatty acid synthesis in leaf
tissue has been a debated topic with some experiments suggesting a direct supply
from the C3 products of photosynthetic carbon ﬁxation and others suggesting the
utilization of free acetate, for which concentrations in leaves in the range of 0.05 to
1 mM have been reported. To address this issue we ﬁrst reassessed the in vivo
rate of fatty acid synthesis using a new method, that of [13C]carbon dioxide labeling
of intact Arabidopsis thaliana plants with subsequent analysis of fatty acids by GC-
MS. This method gave an average value of 2.3 umoles carbon atoms h’1 mg
chlorophyll". The method was extended by the use of pulse labeling and isotopic
dilution analysis to measure the rate of fatty acid synthesis in the dark. There was
negligible fatty acid synthesis (<5% of the rate in the light) in the dark. These
results extend estimates from other workers. With the in vivo rate of fatty acid
synthesis in the light deﬁned, if the bulk tissue acetate concentration available for
fatty acid synthesis is 1 mM, this acetate pool can sustain fatty acid synthesis for
about 60 minutes. When leaves of Arabidopsis, barley, and pea were provided a
5 min pulse of [“C]carbon dioxide the label rapidly appeared in fatty acids with a
lag phase of 2-3 min. Continuous labeling with [“C]carbon dioxide, for up to one

hour, showed a similar result. Furthermore, 1“C-label in free acetate was less than

39

5% of that in fatty acids. In conclusion, these data suggest that either the bulk pool
of acetate is not involved in fatty acid synthesis or the concentration of acetate must

be less than 0.05 mM under strong illumination.

40

Introduction

Apart from a very small contribution from the mitochondrion (Wada et al.,
1997) de novo fatty acid biosynthesis is exclusively located in the chloroplast of leaf
mesophyll cells (Ohlrogge et al., 1979). The immediate carbon precursor for fatty
acid biosynthesis is acetyl-CoA, which is used as substrate by KAS-lll for the acyl-
chain primer reaction, and, after carboxylation to malonyl-CoA and transacylation
to malonyI-ACP, for the acyl-chain elongation steps catalyzed by KAS-l and KAS-ll.
The source of acetyl-CoA in chloroplasts and its relationship to the immediate
products of photosynthetic carbon dioxide ﬁxation continues to be a matter for
debate.

Rates of fatty acid synthesis by intact leaf tissues have been assessed by
various methods. Browse et al. (1981) note that [3H] water incorporation into fatty
acids by spinach leaf discs was equivalent to 2.6 umol C h" mg chlorophyll".
However, that was an estimate made without knowledge of either a kinetic isotope
effect for tritium and assuming only 68% substitution of hydrogen by tritium. In the
same paper the authors measured the net fatty acid accumulation in maize leaves
during a 12 hour day at 80 nmol cm‘z. Using a conversion factor of 30 cm2 mg
chlorophyll", (Focke, personnal communication) this translates to 3.6 umol C h" mg
chlorophyll". Recently, using [‘30] acetate incubations with spinach leaf discs
Pollard and Ohlrogge (1999) showed that the total capacity to synthesis fatty acids,
that is, from a combination of both endogenous and exogenous carbon sources,

ranged from 2 to 8 umol C h" mg chlorophyll" depending on the age of the leaf,

41

with higher rates for younger leaves. Clearly, rates of synthesis of fatty acids will
vary depending on the age of the leaf, the physiological state and growth conditions
of the plant, and species, so exact comparisons between studies is difﬁcult, but it
is clear that endogenous rates of fatty acid synthesis in leaves of the order of 2-8
umol C h" mg chlorophyll" can be expected.

Exogenous acetate is rapidly incorporated into fatty acids of leaves and
isolated chloroplasts (Roughan et al., 1979; Springer et al., 1989; Pollard and
Ohlrogge, 1999). In addition high levels of acetyl-CoA synthetase activity have
been reported in isolated chloroplasts (Roughan and Ohlrogge, 1994), while
chloroplasts have been reported as the dominant site of acetyl-CoA synthetase
activity (Kuhn et al., 1981). The reported acetate concentration in leaf tissue varies
widely, ranging from 0.05 to1.4 mM (Kuhn et al., 1981; Liedvogel, 1985; Treede et
al., 1986; Roughan, 1995). Murphy and Stumpf (1981) and Liedvogel and Stumpf
(1982) were ﬁrst to propose a mechanism to explain how free acetate was
generated and used for fatty acid synthesis. They suggested that pyruvate is
converted to acetate via acetyl-CoA in the mitochondrion; that acetate then diffuses
through the cytosol to chloroplasts where it is converted to acetyl-CoA by the
acetyl-CoA synthetase and becomes available for fatty acid synthesis. Roughan
(1995) revisited this issue and compared different methods used to measure the
acetate concentration in leaf tissues. In his report, the acetate concentration
ranged from 0.5 to 0.8 mM in spinach, 0.5 to 0.8 mM in pea, and 0.9 to 1.4 mM in

Amaranthus, with higher values in younger leaves when compared to older ones.

42

The acetate concentration of leaves in the dark ranged from 0.35 to 0.70 mM, while
that of isolated chloroplasts was 0.4 mM. In conclusion, he suggested that it seems
probable that endogenous acetate would be the direct source of acetyl-CoA for fatty
acid synthesis in leaves. An important part of the argument for acetate as the
dominant substrate for fatty acid synthesis in vivo is that the bulk tissue
concentrations are above the concentration for acetate saturation of rate of fatty
acid synthesis.

Signiﬁcant labeling of fatty acids from other substrates has been reported for
both isolated chloroplasts and for leaf tissue. In leaf tissue Murphy and Leech
(1981) noted carbon dioxide was incorporated into lipids after a short (~5 min) lag
phase at an average rate of 2.1 nmol C h" mg chlorophyll", while in isolated
chloroplasts the rate was 0.6 umoles C h" mg chlorophyll". In a careful comparison
of highly-puriﬁed substrates Roughan et al. (1979) observed maximum rates of 2.7
umol C h" mg chlorophyll" for acetate, 0.8 umol C h" mg chlorophyll" for pyruvate
and 0.35 umol C h" mg chlorophyll" for bicarbonate. Clearly, isolated chloroplasts
can convert the carbon they ﬁx into fatty acids, but not at rates equivalent to the
endogenous rate of fatty acid accumulation.

In this report, we evaluate the relationship between the rate of fatty acid
synthesis in leaves and the total leaf pool of acetate. We demonstrate a new
method that can accurately measure the absolute rate of fatty acid synthesis both
in the light and in dark. We use this rate of synthesis in the light to estimate the

turnover time of the free acetate pool if it is the major source for carbon precursors

43

of fatty acid synthesis. The size of the free acetate pool will determine any lag
phase for added label. And ﬁnally, short term carbon dioxide labeling is used to
measure the actual lag phase. These results are inconsistent with the model of a
high concentration of free acetate (0.5-1.4 mM) supplying most of the carbon for

fatty acid synthesis in leaves.

Material and Methods

Plant Material

Arabidopsis thaliana (ecotype Columbia), Pea (Pisum sativum, cv Little
Marvel ), barley (Hordeum vulgare,cv Morax or Alexis ), and spinach (Spinacia
oleacera, hybrid spinach No. 7 R Early) were grown under the same environmental
conditions as used for the labeling experiments (13.5 h light and 10.5 h dark cycle,
at temperatures of 22-25°C). Cotyledons of the pea plants were sometimes removed
after 4 or 7 days after sowing to prevent the use of storage starch for fatty acid
biosynthesis. Pea plants were used at day 9 after sowing, barley plants at day 5 or
7. Arabidopsis plants were 2 to 3 weeks old and at the 4 to 6 leaf stage.
Labeling with Carbon Dioxide: General Conditions

All labeling experiments were performed in a transparent glove bag (40 liter
gas space, I2R®, Instruments for Research and Industry, Cheltenham, PA, USA) with
one gas inlet and one gas outlet and a larger sleeve closed by a clamp to put the
plant material inside the bag. Gloves were used to manipulate plants and to

liberate 1“CO, inside the bag. A CO2 atmosphere inside the bag was achieved by

44

injecting the appropriate mix of 1“C-NaHCO, (ICN) and unlabeled Na,CO, solutions
into a non-volatile acid (lactic or sulfuric acid). Air circulation was maintained by
using a small battery-driven fan inside the bag. The gas inlet and outlet of the bag
were connected via plastic tubes and stopcocks to an air pressure system and a
gas trap at the outlet in the case of the radioactive labeling experiments or with an
13CO,/air gas tank (compressed air, scrubbed to remove carbon dioxide and
replaced with 13CO2 at 450-480 ppm, BOC gases, Riverton, NJ, USA) at the inlet-
side in the case of the 13C-Iabeling experiments. During incorporation of labeled
CO2 the plants were illuminated (150 umols second " m2 ") with ﬂourescent tubes
(6 high-output light tubes, 8 feet long, Hubbell Lighting Inc.) at room temperature.
Labeling with ['3C]Carbon Dioxide

Continuous labelingto determine the Late of fgt_ty_a_cid synthesis in them
Two hundred and thirty 18-day-old Arabidopsis thaliana seedlings were transferred
to the glove bag under illumination. The ﬂow rate of 13CO2/air mix was 2L min". 20
seedlings each were taken at 30 or 60 min time intervals after the start of labeling,
beginning at 30 min and ending after 8 h. The tissue was frozen in liquid N,,
ground to a powder and stored at -70°C until aliquots were removed for lipid
analysis.

Pulse labeling to determine the rate of fatty acid jvnthesisji the da_r_l_<_,_
Arabidopsis thaliana plants (3-week-old) were labeled with 400uCi of 1“CO, for 20
min in the light; then labeled with 13CO2 for 12 hours under the light. The labeled

plants were removed from the bag and grown under a regular atmosphere for two

45

and half days with a cycle of 13.5 hr light and 10.5 hr dark, after which a sample of
20 plants was taken at several time points both during light and dark period. The
length of this sampling period was 5 days. The tissue was frozen in liquid N,,
ground to a powder and stored at -70°C until aliquots were removed for lipid
analysis.

Labeling with [“CjCarbon Dioxide

Pulse-chase. After a labeling phase of 5 min in which plants were exposed
to 700 ppm CO, generated with the inclusion of 2 mCi of 1“CO, (pulse period), the
radioactive atmosphere was rapidly vented and the plants were placed in a normal
(non-radioactive) atmosphere (chase period) both in the light and dark. Directly
after the 5 minute pulse period a sample of the plant material (20 plants for
Arabidopsis) was taken and frozen in liquid nitrogen. Further samples were taken
for up to 5 h and frozen.

Continuouilabeling. One hundred Arabidopsis plants were incubated in the
bag with 5.4 mmol CO, and 2 mCi 1“CO, to give a nominal carbon dioxide
concentration of 3050 ppm. At 5 or 10 minute intervals up to 60 minutes, 12 plants
were transferred into tubes with sulfuric acid, which were previously placed inside
the bag, to quench the reaction, thereby allowing a continuous labeling without
changing abruptly the labeling conditions.

Lipid Analysis
Before the extraction of fatty acids, 0.5 mg of pentadecanoic acid (15:0) was

added to each sample as an internal standard. Lipids were extracted with

46

hexane:isopropanol according to Radin (1981 ). The organic phase was evaporated
under a stream of nitrogen and the lipids were dissolved in methanol. Chlorophylls
and carotenoids were determined according to Lichtenthaler(1987). In experiments
with [“C]carbon dioxide, radioactivity in lipid, fatty acid and/or isoprenoid fractions
was determined by liquid scintillation counting. Lipids were saponiﬁed by heating
at 90°C for one hour in 2.2 M of KOH in 50% aqueous methanol. On cooling
isoprenoids were extracted three times with hexane. After acidiﬁcation with 10N
HCl the fatty acids were extracted three times with hexane. The pooled fatty acid
fractions were dried with nitrogen. Fatty acid methyl esters were prepared by
heating fatty acids at 90°C for 45 min in 0.3 ml of toluene and 1 ml of 10% (v/v)
boron trichloride/methanol.

To simplify GC-MS analysis, unsaturated FAME were converted to saturated
FAME by catalytic hydrogenation. After hydrogenation, the sum of 16:0 and 18:0
constitute over 99% percent of total fatty acids in Arabidopsis leaf tissue, allowing
rates of fatty acid synthesis to be expressed simply as the sum of C16 and C18 fatty
acids. FAME were hydrogenated using hydrogen at slightly greater than
atmospheric pressure with a platinum (IV) oxide catalyst in methanol, to convert all
16 and 18 carbon unsaturated fatty acids to 16:0 and 18:0, respectively. Total C16
and C18 fatty acids in tissues was determined by GC analysis of the hydrogenated
FAME, against the 15:0 internal standard.

Stable Isotope Analysis of [13C]-Fatty Acid Methyl Esters.

The percentage of newly synthesized fatty acids in total fatty acids was

47

determined by GC-MS analysis. A Hewlett Packard 5890 gas chromatography
conﬁgured with an autosampler and HP MSD 5972 mass analyzer (quadruple,
operating in electron impact mode) was used in this study. Separations of FAME
were carried out on a 30mXO.25mm i.d., 0.25pm thickness DB-23 capillary column
(J&W), with helium carrier gas and using a temperature ramp from 100°C to 240°C
at 5°C sec". Injector and GC-MS interface temperatures were 250°C, with G0
injection in the "splitless" mode. Mass spectra were obtained in "scan" mode
(scanning across 50 to 500 atomic mass units, 3 scans per data point, to give 1.03
spectra per second). Molecular ion clusters of 16:0 and18:0 were monitored for
each time point. The percent of newly synthesized fatty acids of 16 carbon was
calculated by the sum of intensities for (M+3)+ (mlz=273) to (M+16)+ (mlz=286)
divided by the sum of intensities over the entire molecular ion cluster, namely M+
(mlz=270) to (M+16)+ (mlz=286). For 18 carbon fatty acids, a similar calculation
was used, that is, the sum of the intensities for (M+3)* (mlz=301) to (M+18)”
(mlz=316) divided by the sum of intensities over the entire molecular ion cluster,
namely M“ (mlz=298) to (M+18)+ (mlz=316). The value of (M+3)“ (mlz=273) and
(mlz=301) was corrected by subtracting the natural abundance in the calculation
of percentage of newly synthesized fatty acids. The amount of newly synthesized
fatty acid was obtained by multiplying the percentage newly synthesized fatty acid
by the total amount fatty acid as measured by GC.

[“C] Acetate Analysis

To determine whether acetate was labeled during the 1“CO, pulse-chase

48

experiments Arabidopsis plants from the time points of 10, 20, 40, and 120 min
(chase period, after 5 min pulse with 1“CO,) were extracted and analyzed according
to the following protocol. 100 mg of frozen plant tissue was homogenized in 1.6 ml
isopropanol and then in 2.4 ml of hexane. The organic extracts were combined
and shaken with 150 mg of anhydrous sodium sulphate. After centrifugation the
supernatant was partitioned against 0.4 ml of 0.5 M glycine/HCI (pH2.0) saturated
with sodium sulphate and then against 0.2 ml of 0.1M potassium
hydrogencarbonate. The top phase containing the lipids were used to determine
the pigment content and the labeled fatty acids. The aqueous phase was used to
measure the radioactivity in acetate. As a control radioactive acetate was added
to frozen, un-labeled Arabidopsis seedling tissues which then undenlvent the same
extraction and derivatization procedure as the 1“CO, labeled materials. Acetate in
the aqueous extracts was analyzed by one of two methods. In the ﬁrst method,
aqueous extracts were derivatized by heating 0.1 ml of the extract with 0.9 ml of
acetonitrile containing 10 mM 2-bromoacetophenone, 1 mM 18-crownether-6 and
1% triethylamine, for one hour at 90°C in a closed tube. The phenacyl ester
reaction products were extracted with hexane/isopropanol (2:3, v/v) and separated
by TLC (Whatman K6F silica plates developed with hexanezdiethylether (7:3, v/v)).
The radioactive spots on the TLC plates were localized with an Instant lmager
(Canberra Instruments) using the lanes with the exogenously added acetate
samples as markers. The corresponding areas of acetate derivative were scrapped

and radioactivity assayed by liquid scintillation counting. In the second method,

49

the aqueous extracts were directly applied to silica TLC plates (Whatman K6) which
were then developed with 95% ethanol 225% ammonia: water 156:25z19, v/v/v). A
radioactive band with a similar but not identical Rf value to the authentic acetate
band was observed. In order to determine if this radioactive band was indeed
acetate, the band was scrapped out, derivatized and analyzed as its phenacyl

ester.

Results

Measurement of the rate of fatty acid synthesis in Arabidopsis in the light
using [“C]carbon dioxide

In order to determine whether the free acetate pool is involved in fatty acid
synthesis or not it is crucial to have accurate measurements of the rate of fatty acid
synthesis and the pool size of free acetate. The rate of fatty acid synthesis should
be the absolute rate, and not the net rate of fatty acid accumulation (rate of
synthesis less rate of degradation). Each of the methods described in the
Introduction for the measurement of rates of fatty acid synthesis in leaf tissues has
some drawback. Measuring rates with radioisotopes has the problem that although
the rate of synthesis from the exogenous carbon source can be readily quantiﬁed,
the contribution from endogenous substrates is much harderto measure accurately.
To gauge the endogenous contribution small mass increments occurring during the
labeling period (which are net accumulations) must be measured. Here,

[1°C]carbon dioxide labeling was employed in attempt to measure the rate of fatty

50

acid synthesis in Arabidopsis.

By monitoring the molecular ion clusters of 16:0 and18:0 FAME, which were
obtained after hydrogenation of total FAME, the newly synthesized fatty acids are
readily distinguishable from pre-existing fatty acids. As an example, the molecular
ion cluster of 18:0, at one hour labeling time point, is shown in Figure 2.1. An ion
corresponding to methyl [1°C,8] stearate (m/z = 316) is clearly visible, showing that
all the carbon atoms in the chain are derived from the added [13C]carbon dioxide.
More important, the distribution of species is such that methyl [130,4] stearate is the
dominant species in the distribution, indicating that dilution with endogenous carbon
is small (less than 30%). The source of this endogenous contribution may be from
leaks of 12CO, into the system, and from [‘2C1carbon dioxide released by the
system, especially through respiration of the plants themselves and soil
microorganisms. The point is that it can be easily seen and measured. Ignoring it
would underestimate the rate of fatty acid synthesis by-about 30%. In unlabeled
methyl stearate, natural abundance isotopic peaks can be readily observed for (M
+ 1), (M + 2) and (M + 3) ions, at relative intensities of 21.3%, 2.6% and 0.25%
respectively (experimental values, not calculated), when compared to the molecular
ion peak (M, m/z = 298, at 100% intensity). Thus the ions from 303 to 316 (m/z) are
clearly derived from only newly synthesized fatty acid containing 4 to 18 13C-
carbons. Ions from m/z = 298 to 302 have only a very small component from the
labeling period, which is less than 2% of the total newly fatty acid synthesis, and so

can be ignored. Thus the percent of newly synthesized fatty acids was obtained

51

 

 

 

 

 

 

 

 

                   

l
, 140°J| ‘1 1400. ‘
. l
i 1200i ‘ 1200. i
l
l 1000' l l 1000. l
E l i l 5 l
{ 600 i‘ 2 600- ,
l 400i ‘ 400 i l
”I l! I ””l l
i 0.1 , ,LL it OL- NvlﬁerLJngnguélLl a}
‘ §Ion§mlz§)for§mo§lecu§ritsér 017,18: ‘ 12n(;Iz):orr:ole:ular“clusterNof16:0 l
L, W , ,, ,, ,JL , 4 7 .,

 

Figure 2.1. The molecular ion distribution of 18:0 and 16:0 from Arabidopsis labeled
by 13CO2 at one hour time point. The molecular ion clusters of 18:0 (A) and16:0 (B)
FAME, which were obtained by the hydrogenation of total FAME from Arabidopsis
leaves, at one hour labeling time point with 13CO,. The 18:0 represents all 18

carbon fatty acids, and so as for 16:0, due to the hydrogenation.

52

through the sum of ion abundances derived from newly synthesized fatty acids (m/z
= 303 to 316) divided by the total ion abundance for the molecular ion cluster. The
percent of newly synthesized 16 and 18 carbon fatty acids were plotted with time
and are shown in Figure 2.2 (A) and (C). After 8 hour labeling with [1°C]carbon
dioxide, 16% of 16 carbon fatty acids and 14% of 18 carbon fatty acids are newly
synthesized. The data are also expressed as mg newly synthesized fatty acids mg
chlorophyll" (Figure 2.2 (B) and (C)). In both experiments shown in Figure 2.2 the
accumulation of both 16 and 18 carbon fatty acids showed linear increases
throughout the 8 hour labeling period. The synthesis rates of 16 and 18 carbon
fatty acids were 12 and 24 ug fatty acids h" mg chlorophyll", respectively.
Combining 16 and 18 carbon fatty acids, the rate of fatty acid synthesis is converted
to 2.3 pmol C h" mg chlorophyll" for the whole aerial part of the plant (stem plus
leaves)
Measurement of the rate of fatty acid synthesis in Arabidopsis in the dark with
[‘3C]carbon dioxide

It is well known that the rate of fatty acid synthesis of leaf tissues is lower in
the dark than in the light. Estimates made using labeled precursors fed to leaf discs
suggest 12-20% of the rate in the light (Browse et al., 1981). The same authors
show that fatty acid synthesis from acetate by isolated chloroplasts is essentially
zero. Since we can accurately measure the rate in the light as demonstrated in the
previous section, we should be able to determine the rate in the dark, if the dark

rate relative to light can be evaluated. This can be done with an isotope dilution

53

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‘9/9/98 _. - _ | 9/9/98

 

~_.—_ __ . _ ___—.I i-.—. _ ‘ .,.,—..__ _ —____ ‘,—.__ , _i

Figure 2.2. Determining the rate of fatty acid synthesis in Arabidopsis by continuos
labeling with ”CO, (A) (B) and (C) (D) represent two independent experiments. (A)
and (C) show the percentage of newly synthesized fatty acids in total fatty acids
with labeling time. The calculation of newly synthesized fatty acids was described
in Methods. (B) and (D) gave the amount of newly synthesized fatty acids with time
course. At given time point, the value was obtained by multiplying percent of newly
fatty acids with the absolute amount of total fatty acids measured by GC. (o)and (A)

represent 16 and 18 carbon fatty acids, respectively.

54

strategy, again employing [laclcarbon dioxide. Arabidopsis plants were labeled with
[“Cjcarbon dioxide for 20 min and then with [1°C]carbon dioxide for 12 hours under
the light. The labeled plants were removed from the labeling chamber and grown
under a normal atmosphere for two and half days prior to sampling both during light
and dark periods. The ﬁrst sample was taken just before the light was turned on,
and this time point was designated as zero time. The sampling period lasted for 5
days. The purpose for growing the labeled plants under a normal atmosphere for
60 hours was to deplete the labeled carbon intermediates, particularly sucrose and
starch, in order to minimize further incorporation of labeled carbons into fatty acids
during the sampling period. Thus it would be expected that any turnover of label
from any source into fatty acids would be highly diluted, contributing on average
only one or two 13C or 1"C atoms per newly synthesized fatty acid. As shown in
Figure 2.3, 1“C label in fatty acids measured on a per plant basis decreased slightly
during the sampling period, by about 22% (140 to110 units on Figure 2.3). The
turnover of fatty acids could be greater, however (since we are measuring a net
ﬁgure for total labeled fatty acids), if there is signiﬁcant recycling of label.
Considering C16 and C18 fatty acids, there is about a 100% increase in mass per
plant over the 120 h sampling period. Also in Figure 2.3, fatty acids, fresh weight
and chlorophyll all showed linear increase throughout the period, indicating the
labeled plants grew normally.

The ion distribution of species in the ion molecular cluster of 18:0 and 16:0

was analyzed for each sampling point. For the purpose of this analysis m/z = 309

55

l
l
l
l
l
l

.333. madam)

 

l
l 160 , —-A-fa 1&3019'931)
, , + 0” a+b (W)
l 1 ,1 + WWW)
’10 + /+ +,-+ _.+_. W
i\ / \ .+ +

 

16 and 18 carbon FAs, chlorophyll,
fresh weight, and CPM

 

Figure 2.3. Some important parameters concerning plant growth during the
sampling period in the pulse-chase experiment to determine the rate of fatty acid

synthesis in the dark.

56

was chosen to represent the 13‘C labeled C18 fatty acids, mlz = 280 to represent the
13C labeled C16 fatty acids. The base peak for each molecular ion cluster, namely
mlz = 298 for C18 fatty acids and mlz = 270 for C16 fatty acids, represents
unlabeled fatty acids. If there was any fatty acid synthesis during the sampling
period, it would be reﬂected as a percentage increase of mlz=298 for 18:0 and
mlz=270 for 16:0, or in a percentage decrease of mlz=309 for 18:0 and mlz=280 for
16:0 at any given time point. Conversely, if there was any degradation there would
be no change in the percentages. These data are plotted over the sampling period
in Figure 2.4. As demonstrated in Figure 2-4 (A) and (B), mlz=298 and mlz=270
(representing unlabeled C18 and C16) increased during the light periods, but not
during the dark periods. On the other hand, Figure 2.4 (C) and (D) showed that
mlz=309 and mlz=280 (representing labeled C18 and C16) decreased during the
light periods, but not during the dark periods. This result clearly demonstrated that
fatty acid synthesis was halted during the dark period. Averaging the data for four
dark periods gave a rate of zero.
Short term labeling with [“Clcarbon dioxide

As measured in the experiments described above, the rate of fatty acid
synthesis was 2.3 nmol C h" mg chlorophyll" for the whole aerial part of the plant.
If the free acetate concentration in aerial plant tissues is 1 mM (about 1.2 nmol mg
chlorophyll") and is completely available, this acetate pool would be able to sustain
63 min of fatty acid synthesis. This would cause a lag phase prior to incorporation

of exogenously labeled substrate. Since pool ﬁling is a ﬁrst order kinetic process,

57

 

 

 

 

 

 

 

 

 

 

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l o 20 4o 60 so 100 120 o 20 4o 60 80 100% 120i
I Sampling time(hr) Sampllng time (hr) I
l ____ 3 IL

 

 

Figure 2.4. The pulse-chase labeling with 13CO2 to determine the rate of fatty acid
synthesis in the dark. The same set of samples were used both in this ﬁgure and
ﬁgure 3. FAMEs were hydrogenated, so 18:0 and 16:0 represented 18 and 16
carbon fatty acids, respectively. The sampling time was the time after the ﬁrst
sample taken which designated as the zero time point. (A) (C) and (B) (D) are for
18:0 and 16:0, respectively. (-o-) and (-O-) represent light and dark period,

respectively.

58

with a rate constant k = 0.95 hr", calculated from the above values, the half life for
pool ﬁling is 44 minutes (t1 ,, = 0.693/k) (Segel, 1968). Thus the extrapolation of the
linear rate of fatty acid synthesis to the x-axis (time axis) would result in a lag
measured on the x-axis of 44 min. These calculations suggest that very signiﬁcant
lag phases would occur with [“Clcarbon dioxide labeling if a large free acetate or
other intermediate pool did indeed supply fatty acid synthesis in vivo.

Arabidopsis seedlings were pulse labeled for 5 min with [“Cjcarbon dioxide,
then chased with unlabeled carbon dioxide for up to 270 min. The time course for
label accumulating in fatty acids is shown in Figure 2.5 (A). Extrapolation of the
labeling rate at the end of the 5 minute pulse period to the beginning of the pulse
period gives a lag period of about 2-3 minutes. This can be explained, at least in
part, by label moving through C(1) to C(2) to C(3) in triose—phosphate intermediates
in the Calvin cycle. Such intermediates will be the source of pyruvate and then
acetyl-CoA. After 40 minutes of chase period movement of label from precursor
pools into fatty acids is almost complete. In addition to Arabidopsis a similar kinetic
behavior of fatty acid labeling was observed in the dicotyledonous plants spinach
and pea, and the monocotyledonous plant barley (data not shown). We therefore
believe that this labeling pattern is a general property of fatty acid biosynthesis in
higher plants.

In another experiment, Arabidopsis seedlings were labeled with [“Clcarbon
dioxide for 5 min, then chased with unlabeled carbon dioxide for 270 min in either

the light or the dark. These pulse-chase time courses are shown in Figure 2.5 (B).

59

 

 

 

 

 

l
l
l
, 100 . l
f 2: I
q a
E E 30: ‘
l a I
§ § I
3 3
.3 . I s I
_ 40 I
E : I 5 I
3 , I 3 ,0 I
r! . r! .
V 2‘" I I l
l I l I '
l 0 _ .3 . . . .. ,, ..., l 0 + _ _L_._‘ -_.. , #_ , !
I o 100 200 300 II o 100 200 300 I
I Pulse-chasing time (min) I i Pulse-chasing time (min) I
. l‘
I _. _ __ _. ___ .I _ __. ___ __ J

 

Figure 2.5. Pulse-chase labeling of Arabidopsis with 1"CO, to determine the lag
phase. (A) Arabidopsis seedlings were pulse labeling with for 5 min, then chase for
up to 280 min in the light. Radioactivity in fatty acids from two independent
experiments was plot with pulse-chase time. (B) Arabidopsis seedlings were pulse
labeling with for 5 min in the light, then chase for up to 260 min either in the light
(-o-) or in the dark (-0-).

6O

In the light, radioactivity accumulated in fatty acids continued increasing up to 40
min; but in dark, incorporation of radiolabel stopped immediately after plants are
transferred to the dark, demonstrating a complete cessation of fatty acid synthesis.

A third type of experiment used continuous [“C]carbon dioxide labeling for
one hour. This experiment was designed to look for a slow response pool in
addition to the rapid labeling kinetics seen in the previous two experiments by short
time pulses. The results are shown in Figure 2.6. The fatty acids shows a linear
labeling without upward hyperbolic kinetics at later time points, indicating that there
is not a second more slowly responding pool of precursors for fatty acid
biosynthesis in mesophyll cells. Consistent with [“C]carbon dioxide pulse-chase
experiment, extrapolation of the labeling rate to the x-axis gives a lag period of
about 3 minutes. Radioactivity accumulated into total isoprenoids (largely sterols
and plastidic isoprenoids) also showed a similar linear labeling pattern, with a
similar lag phase (Figure 2.6).
[“C] Acetate Analysis

In order to assay [“C]acetate recovered from the aqueous extracts of
incubations with [“C]carbon dioxide the aqueous extracts were derivatizaed to
convert volatile carboxylic acids into non-volatile phenacyl esters which could be
analyzed by TLC. Samples from the above pulse-chase experiment (shown in
Figure 2.5) at 10, 20, 40, and 120 minutes were analyzed using the Instant lmager
and by liquid scintillation counting. No radioactivity could be detected in the region

corresponding to the phenacyl acetate standard. Taking into account the efficiency

61

16000 _

 

14000 J .0
j 9
12000 .
10000 ‘
‘5'. l
E 8000. , '
a; ' 0
°- l
s 6000 1
a.
O
4000 l j __x
2000 l 3;. A.‘
1 ,". -'-‘_.-'
9‘ -"A
0 &""““.-f' " ""‘"T—' ‘TTT "*"“‘l r’ ”T‘TT‘I
0 10 20 30 ‘40 50 60

Labeling Time (min)

Figure 2.6. Continuos labeling of Arabidopsis with “C02 to determine whether
there is a slow ﬁlling pool. One hundred Arabidopsis plants were incubated in the
bag with 5.4 mmol CO2 and 2 mCi 1“C02. After 5, 10, 15, 20, 30, 40, 50, and 60
minutes 12 plants were transferred into tubes with sulfuric acid, then radioactivity

in fatty acids (0) and isopprenoids (A) in each sample was plot with labeling time.

62

of extraction and derivatization, if the acetate concentration in leaves is 1 mM, the
detection limits for the protocol suggest that the amount of labeled acetate must be
less than 2% of the total acetate pool. The radioactive spot in the non—derivatized
samples running at a similar Rf value to acetate standard was not acetate, because
after derivatization with bromoacetophenone no corresponding phenacyl acetate

band could be detected by TLC.

Discussion

Previous considerations of the path of carbon into fatty acids in leaves have
almost entirely relied on in vitro measurements of enzyme activities and their
subcellular localization or on experiments with isolated chloroplasts. Although
useful to deﬁne possible reactions which occur in vivo, such studies have several
limitations related to inactivations or other disturbances which often occur upon cell
disruption. Therefore, in this study we attempted to re-examine the flux of carbon
into fatty acids by methods which could more closely reﬂect in vivo metabolism.
Measurements of In Vivo Rates of Fatty Acid Synthesis

The ﬁrst step in assessing the ﬂux of intermediates through a pool for any
pathway is to know the in vivo rate of synthesis of the product of interest, not just
the net rate of accumulation of the product. For fatty acids, the use of exogenous
tracers presents uncertainties because it is impossible to assess endogenous
contributions without measuring speciﬁc activities. Such measurements over short

periods give a high degree of statistical variability, since the increase in mass is

63

small, while over extended periods they introduce the uncertainty of net
accumulation. A way around the dilemma in the investigation of carbon ﬂuxes is to
perform in vivo [3H]water labeling to estimate the rate of fatty acid synthesis (Browse
et al.,1981). The water should exchange rapidly with the leaf tissue water and is
expected to label both endogenous and exogenous carbon substrates equally.
Injury to the tissue during preparation of the leaf discs may introduce some
uncertainty into applying the resultant rate to intact plants. Also, the use of
[3H]water requires corrections for isotope effects and a knowledge of the maximum
value for substitution of hydrogen by tritium. These corrections were not made by
Browse et al., (1981). Nevertheless, these workers obtained as estimated in vivo
rate of fatty acid synthesis of 2.6 pmol C h'1 mg chlorophyll“.

Using [13C]carbon dioxide labeling to measure the rate of fatty acid syntheses
by GC-MS has some obvious advantages. First, plants are labeled intact under
natural growth conditions, rather than as detached parts ﬂoating in incubation
medium. Second, the newly synthesized fatty acids are easily distinguishable from
pre-existing fatty acids (Figure 2.1). Thirdly, the method removes the problem of
measuring the small mass increments for fatty acid accumulation over short assay
periods. In the light, the rate of fatty acid synthesis of Arabidopsis obtained with
[13C]carbon dioxide labeling was 2.3 pmol C h'1 mg chlorophyll". Exact
comparisons with the 2.6 pmol C h'1 mg chlorophyll" obtained by [3H]water labeling
(Browse etal.,1981) are not possible because of the species difference and the fact

that our work represents a whole plant (aerial parts) average rather than selected

64

leaves. It is of interest that of the total rate of fatty acid synthesis about 30% was
not accountable to the added [‘3C]carbon dioxide label. What is important is that we
can see the mixing of the exogenous and endogenous label since the distributions
are essentially smooth curves. The conclusion that exogenously and endogenously
derived carbon substrates mix randomly for fatty acid synthesis was also reached
in a GC-MS analysis of fatty acids from the labeling of spinach leaf discs with [1-
13C] acetate (Pollard and Ohlrogge, 1999). The work presented here is based on
the validity of this assumption.

It is clear that the majority (at least 70%) of carbon in newly synthesized fatty
acids is derived from recently ﬁxed [‘3C]carbon dioxide. At this time we cannot
deﬁne whether the 30% 12C contribution represented dilution of the [‘3C]carbon
dioxide with unlabeled carbon dioxide (generated by respiration either internally
within the tissues or from the system, particularly the soil) or represented non-
photosynthetic parts of the tissues, or represented an additional carbon source for
fatty acid synthesis from previous ﬁxed carbon pools such as starch or sucrose.
Delwiche and Sharkey (1993) noted that isoprene labeling from carbon dioxide in
isolated oak leaves saturated at about 80% contribution from the carbon dioxide,
which was very similar to the saturation level of the phosphoglycerate pool in beet
leaves described by Canvin (1979). Both results suggested an approximately 20%
contribution of carbon from a source other than the immediately ﬁxed carbon in the
chloroplast.

The rate of fatty acid synthesis in the dark in leaf tissue is known to be much

65

reduced, but the question remains whether, in vivo, it is still signiﬁcant (10-20% of
light rate) or much closer to zero (<5%). Estimates made using labeled precursors
fed to leaf discs suggest 12-20% of the rate in the dark (Browse et al., 1981). The
same authors show that fatty acid synthesis from acetate by isolated chloroplasts
is essentially zero. Since we can accurately measure the rate in the light, we should
be able to determined the rate in the dark, if the dark rate relative to light can be
evaluated. This can be done with an isotope dilution strategy, again employing
[‘3C]carbon dioxide (Figures 2.3 and 2.4). In this experiment we demonstrate that
there is a slow fatty acid degradation, of the order of 4-5% per day. The label
released is not signiﬁcantly recycled back into fatty acids to complicate the analysis.
As shown in Figure 2.4, during the dark period, ions representative of the 13C
labeled C18 fatty acids (mlz = 309) and of the 13C labeled C16 fatty acids
(mlz=280), and ions representative of the unlabeled C18 fatty acids (mlz = 298) and
of unlabeled C16 fatty acids (mlz=280) remain unchanged in relative proportion.
Fatty acid synthesis will result in the latter set increasing at the expense of the
former, Whereas there was a very slight statistical average in the other direction.
These result clearly demonstrated that, for Arabidopsis seedlings, there was
negligible fatty acid synthesis in the dark. In support of this observation, when
Arabidopsis seedlings were transferred to dark after 5 min labeling with [“C]carbon
dioxide in the light, radioactivity in fatty acids remained constant throughout the
dark period (Figure 2.5(B)).

Rapid Movement of Label from Carbon Dioxide to Fatty Acids and lsoprenoids

66

The labeling from [‘3Cjcarbon dioxide suggests that at least 70% of the
carbon for fatty acid synthesis arises from newly fixed carbon, while the [“Clcarbon
dioxide pulse chase and time courses (Figures 2.5 and 2.6) show that the lag phase
for the movement of photosynthetically ﬁxed carbon into fatty acids occurs very
rapidly, within 2-3 minutes. This lag time is about half that reported by Murphy and
Leech (1981). Thus, if acetate is the dominant source of acetyl-CoA for fatty acid
synthesis, newly ﬁxed carbon (with all three triose—P carbon atoms labeled) will
have to move through to acetate very quickly, by a mechanism that is still
debatable. A particularly important consideration for this putative ﬂux is the rate at
which label from [“Cjcarbon dioxide moves into isoprenoids that are synthesized
in the chloroplast, most notably phytol (Figure 2.6). This discovery that isopentenyl
pyrophosphate for isoprenoid biosynthesis in the chloroplast is not derived from
acetyl-CoA via mevalonate, as it is in the cytosol, but via the glyceraldehyde-S-
phosphatezpyruvate pathway with 1-deoxy-D-xylulose-5-phosphate as an
intermediate, is important (Lichtenthaler et al., 1997). Because the synthesis of
phytol, via the chloroplastic isopentenyl pyrophosphate pathway used C3 precursor
that are "upstream" of acetyl-CoA any lag phase for phytol biosynthesis will include
part of the same lag phase for acetyl-CoA production, that is mixing time for
[“Cjcarbon dioxide in the bag, diffusion to chloroplasts and ﬂushing through of the
label in the Calvin cycle. In fact, as shown in Figure 2.6, the lag phases for the
movement of label from [“C]carbon dioxide into fatty acids and into isoprenoids are

experimentally indistinguishable. Thus the three minute lag phase for label moving

67

into fatty acids is an upper limit in considering an acetate pool, and may be
signiﬁcantly less than that value. Delwiche and Sharkey (1993) make estimates of
the lifetime of carbon in the photosynthetic carbon reduction cycle of the order of
two minutes. Pyruvate in plastids is not only used for the generation of acetyl-CoA
for fatty acid biosynthesis but also for isoprenoid biosynthesis and for the synthesis
of branched-chain amino acids (Hoppe et al., 1993; Schulze-Siebert et al., 1984).
Pyruvate concentrations of 40 to 120 (M in the leaves of dicot plants (Treede et al.,
1986) are low enough to allow the observed kinetic pattern of fatty acid labeling.
Issues of Acetate Supply of Fatty Acid Synthesis in Vivo

We know that the endogenous rate of fatty acid synthesis in Arabidopsis
plants is 2.3 pmol C h" mg chlorophyll" for the whole aerial part of the plant (stem
plus leaves), that most of the carbon for fatty acid synthesis comes from carbon
dioxide that has been recently ﬁxed, and that the movement of label from carbon
dioxide to fatty acids reaches a steady state level within about 2 minutes. If all this
carbon passed through an acetate pool, then the lifetime of acetate in this pool
would be about 3 minutes, and amount to 0.114 pmol C mg chlorophyll". In the
"Results" section we used a conversion factor of 1 mM acetate bulk tissue
concentration equivalent to 1.2 pmol acetate mg chlorophyll". Thus the putative
acetate pool supplying fatty acid synthesis will be 0.0475 mM, or more likely
<0.0475 mM. Consistent with this conclusion, the results of [“Cl-acetate analysis
demonstrated that if there was a 1 mM bulk acetate pool less than 2% of it was

labeled, which is equivalent to <0.02 mM. This result should be compared with

68

the range of bulk acetate concentrations measured in leaf tissues, from 0.05 to 1.4
mM (Kuhn et al., 1981; Liedvogel 1985; Treede et al., 1986; Roughan, 1995).
Roughan found higher values in younger leaves when compared to older ones, and
that acetate concentration of leaves in the dark ranged from 0.35 to 0.70 mM, while
that of isolated chloroplasts was 0.4 mM.

Several points must be kept in mind in interpreting the results. First, in
isolated chloroplasts the rate of fatty acid synthesis from exogenous acetate
saturates at between 0.1 and 0.2 mM (Roughan et al., 1979). Second, a bulk
concentration of 1 mM does not imply that each compartment in the cell will be at
1 mM. For example, the mesophyll cell has about 80% of its volume as vacuole
(Winter etal., 1994), with a pH of about 5.5. Thus at equilibrium most of the cellular
free acetate is likely to be partitioned into the cytoplasm, and even more so into the
chloroplast stroma. Thus, even at a bulk tissue concentration of 0.05 mM acetate,
it is probable that the concentrations of acetate in proximity to the chloroplast are
enough to sustain high rates of fatty acid synthesis. And third, high concentrations
of acetate can drive high rates of fatty acid synthesis in spinach leaf discs (Kosfor
exogenous acetate at pH 5.6 is about 2 mM, with Vmax of about 2 pmol C h" mg
chlorophyll"; Pollard and Ohlrogge, 1999). This conﬁrms the concept that acetate
will readily compete with endogenous substrate to supply fatty acid synthesis in
vivo, and is suggestive of the idea that acetate in leaf tissues is freely diffuseable,
not tightly bound, and therefore available.

Thus we reach a conclusion which seems almost counter-intuitive. If the in

69

vivo bulk concentration of acetate in leaf tissues is at the upper end of the range
reported in the literature (0.5-1.4 mM), we must conclude from our kinetic analyses
that this acetate is not a major substrate for fatty acid synthesis. The only possible
mechanism for this is that all this acetate is so tightly bound or sequestered that
it is unavailable. Because of the rapid movement of exogenously added acetate
into tissue fatty acids, regardless of how it is applied, these alternatives seems
unlikely. Possibly the measurements of the in vivo concentration of acetate in leaf
tissues are incorrect (Kuhn et al., 1981; Liedvogel, 1985; Treede et al., 1986;
Roughan, 1995). On the one hand, acetate is volatile and might be lost during the
preparation and clean-up of the extracts. On the other hand, during extract
preparation artiﬁcially high acetate concentrations might be generated due to
chemical or enzymatic hydrolysis of endogenous acetyl or to breakdown of
molecules yielding acetate (e.g. decarboxylation of malonate). All the methods
quoted above (Kuhn et al., 1981; Liedvogel, 1985; Treede et al., 1986; Roughan
1995) use heating steps at some stage in the extraction. By contrast, if the in vivo
bulk concentration of acetate in tissues is even lower than the range reported in the
literature, (e.g. <0.05 mM), it is still possible that acetate is a major supplier of
substrate for fatty acid biosynthesis. However, if the level of free acetate in tissues
is in actuality low, then the issue of its contribution to fatty acid synthesis becomes
a quantitative one. At a certain point the lower steady-state free acetate
concentration in leaf tissues will no longer be able to support the total carbon ﬂux

into fatty acids. The question then arises as to whether the rapid movement of

70

newly ﬁxed carbon into fatty acids is solely a ﬂux of intermediates within the plastid,
a ﬂux involving the plastid and the cytosol, or includes movement of carbon through
other organelles, especially the mitochondrion.

Origin of Acetyl-CoA in Vivo for Fatty Acid Synthesis.

The question regarding the origin of acetyl-CoA used for fatty acid
biosynthesis has been discussed for several decades. Starting with the observation
by Smimov (1960) that acetate is a very good precursor for incorporation into fatty
acids in isolated chloroplasts, acetate has been used for many labeling studies both
in vivo and with isolated plastids. Researchers then looked for mechanisms that
would explain how free acetate might be generated in the plant cell. In one model
acetyl-CoA is generated via oxidative decarboxylation of pyruvate in the
mitochondria. Acetyl-CoA, which is known not to be transported across
membranes, is hydrolyzed by an acetyl-CoA hydrolase in the mitochondrial matrix
to yield free acetate, which can diffuse freely through membranes and so enter the
chloroplast ( Murphy and Stumpf 1981 ; Liedvogel and Stumpf 1982). An alternative
pathway for acetate generation has also been proposed, namely through the action
of ATchitrate lyase (Nelson and Rinne, 1977; F ritsch and Beevers, 1979). In
chloroplasts, the existence of an acetyl-CoA synthetase using free acetate, CoA,
and ATP to synthesize acetyl-CoA, AMP, and pyrophosphate has been
demonstrated (Kuhn et al., 1981) and its maximum activity is over 5 fold higher than
the ﬂux of fatty acid synthesis (Roughan and Ohlrogge, 1994). It should be noted

that cellular anabolism produces free acetate by at least three mechanisms of which

71

we are aware. These are in the biosynthesis of cysteine via O-acetylserine, the
biosynthesis of ornithine and hence the polyamines via N-acetylglutamate, and the
acetylation and deacetylation of histones, each yielding acetate in a subsequent
deacetylation step. Thus the efﬁcient use of free acetate for acetyl-CoA
biosynthesis (and hence fatty acid biosynthesis) in the chloroplast may simply
reﬂect an efﬁcient scavenging mechanism for free acetate used by the plant cell.

An alternative pathway for the formation of acetyl-CoA in plastids has been
proposed via the plastidial pyruvate dehydrogenase complex (pPDC) (Reid et al.,
1977; Elias and Givan, 1979; V\ﬁlliams and Randall, 1979). There are two origins
for the pyruvate needed for the pPDC reaction. First the existence of a complete
glycolytic sequence inside the chloroplasts has been proposed (Schulze-Siebert
and Schulz, 1989; Hoppe et al., 1993). This is supported by the observation that
isolated chloroplasts can synthesize fatty acid from [“Clbicarbonate (Murphy and
Leech, 1977; Murphy and Leech, 1978), albeit with a lower rate than that required
for the in vivo rate of fatty acid synthesis (Roughan, 1993; Murphy and Leech, 1 981 ).
This pathway requires the conversion of 3—phosphoglyceric acid to acetyl-CoA via
2-phosphoglyceric acid, phosphoenolpyruvate and then pyruvate. It is sometimes
called the C,-C2 pathway. A modiﬁcation of this C3-C2 pathway occurs if not all the
glycolytic steps occur in the plastid but instead some reactions are located in the
cytosol and that PEP (Fischer et al., 1997) or pyruvate are reimported into the
chloroplast.

A third, but less generally accepted hypothesis is that acetate units may be

72

moved by an acetylcamitine-shuttle into different compartments. In such a scheme
pyruvate in the mitochondrial is converted to acetyl-CoA, which is exported from the
mitochondria as acetyl-carnitine, and imported by the chloroplast (Masterson et al.,
1990a and 1990b).

Although the majority of acetyl-CoA in plastids is used for fatty acid
biosynthesis, the supply of plastidial acetyl-CoA must be ﬂexible enough to support
plastidial polyhydroxybutyric acid biosynthesis in Arabidopsis thaliana transgenics
from acetyl-CoA, since it accumulates at up to 14% of dry weight of the leaf,
whereas no signiﬁcant changes of membrane lipids was observed (Nawrath et al.,
1994). In conclusion, the results presented in this paper provide in vivo data which
distinguish in part between the alternative pathways suggested for acetyl-CoA
production in leaves. Our data clearly indicate that fatty acid synthesis does not
depend on a high tissue concentration (0.5-1.4 mM) of acetate for a carbon source.

Furthermore, the involvement of other large pools of metabolic intermediates
between carbon dioxide and acetyl-CoA is ruled out. The kinetic analysis of the
rapid movement of label from carbon dioxide into both fatty acids and plastidial
isoprenoids is certainly suggestive of direct utilization of newly ﬁxed carbon via the
C3-C2 pathway into fatty acids. However, although unlikely, the type of experiments
we have performed do not provide enough data to eliminate with certainty the
possibility that a small pool of acetate, metabolically distinct from bulk acetate, is
an intermediate in leaf fatty acid synthesis. We do not know the time constant for

transport processes and for ﬁlling intermediate pools through other organelles,

73

especially the mitochondria, which is the only other site of PDC in the plant cell.
This will require rapid kinetic studies for which the use of leaf tissue may or may not
be adequate to give the required time resolution, and generally can only give bulk

cellular pool information.

74

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Kuhn DN, Knauf M, Stumpf PK (1981) Subcellular localization of acetyl-CoA
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Masterson C, Wood C, and Thomas DR (1990) L-acetylcamitine, a substrate for
chloroplast fatty acid synthesis. Plant Cell Environ. 13: 749-

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Murphy DJ, Leech RM (1978) The pathway of (“0) bicarbonate incorporation into
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Murphy D, Stumpf PK (1981) The origin of chloroplastic acetyl-CoA. Arch.
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Nawrath C, Poirier Y, and Somerville C (1994) Targeting of the
polyhydroxybutyrate biosynthetic pathway to the plastids of Arabidopsis
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Nelson DR, Rinne RW (1977) Role of citrate in lipid-synthesis in developing
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Ohlrogge JB, Kuhn DN, Stumpf PK (1979) Subcellular localization of acyl carrier
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Pollard MR, Ohlrogge JB (1999) Testing models of fatty acid transfer and lipid
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Radin NS (1981) Extraction of tissue lipids with a solvent of low toxicity. Methods
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Roughan PG (1995) Acetate concentrations in leaves are sufﬁcient to drive in vivo
fatty acid synthesis at maximum rates. Plant Sci. 107: 49-55.

Roughan PG, Hollad R, Slack CR, and Mudd JB (1979) Acetate is the preferred
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Roughan PG, Ohlrogge JB (1994) On the assay of acetyl-CoA synthetase activity
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Acetylcarnitine and fatty acid synthesis by isolated chloroplasts. Plant
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Schulze-Siebert D, Heineke D, Scharf H, Schultz G (1984) Pyruvate-derived
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78

CHAPTER 3‘
Isolation and Characterization of an Arabidopsis Biotin Carboxylase
Gene and its Promoter
Abstract
In the plastids of most plants, acetyl-CoA carboxylase (ACCase; EC 6.4.1.2) is a
multisubunit complex consisting of biotin carboxylase (BC), biotin-carboxyl carrier
protein (BCCP), and carboxyltransferase ( or-CT, B-CT ) subunits. To better
understand the regulation of this enzyme, we have isolated and sequenced a BC
genomic clone from Arabidopsis and partially characterized its promoter. Fifteen
introns were identiﬁed. The deduced amino acid sequence of the mature BC
protein is highly conserved between Arabidopsis and tobacco (92.6% identity). BC
expression was evaluated using Northern blots and BC/GUS fusion constructs in
transgenic Arabidopsis. GUS activity in the BCIGUS transgenics as well as
transcript level of the native gene were both found to be higher in silique and ﬂower
than in root and leaf. Analysis of tobacco suspension cells transformed with
truncated BC promoter/GUS gene fusions indicated the region from -140 to +147
contained necessary promoter elements which supported basal gene expression.
A positive regulatory region was found to be located between -2100 and -140,
whereas a negative element was possibly located in the ﬁrst intron. In addition,

several conserved regulatory elements were identiﬁed in the BC promoter.

 

lMaterial in this chapter was published previously [Bao X, Shorrosh BS, and Ohlrogge J (1997)
Plant Molecular Biology 35: 539-550].

79

Surprisingly, although BC is a low abundance protein, the expression of BC/GUS

fusion constructs was similar to 35S/GUS constructs.

80

Introduction

Acetyl-CoA carboxylase catalyzes the ATP-dependent carboxylation of
acetyl-CoA to produce malonyl-CoA. This reaction is the ﬁrst committed step of de
novo fatty acid biosynthesis and is a major point of ﬂux control for this pathway
(Harwood,1988; Ohlrogge and Jaworski, 1997; Wakil et al.,1983). Most plants
contain two structurally distinct forms of ACCase (Sasaki et al., 1995). One form,
termed “homodimeric” (HO), "eukaryotic" or "multifunctional” is a >200kD
polypeptide in which biotin carboxylase (BC), biotin carboxyl carrier protein (BCCP),
and carboxyltransferase (CT) occur as functional domains (Gornicki et al.,1994;
Roesler et al.,1994; Roessler and Ohlrogge, 1993; Shorrosh et al., 1994). The
other form of ACCase, termed “heteromeric” (HE), “multisubunit” or "prokaryotic" is
an enzymecomposed of easily dissociable subunits. Most evidence suggests the
HO-ACCase occurs in the cytosol of all plant families whereas HE-ACCase is
localized in the plastids. However, within the Gramineae family the HO-ACCase
exists in the plastid as well as the cytosol (Baldet et al.,1993; Gornicki and
Haselkom,1993; Konishi and Sasaki,1994; Roesler et al.,1994; Shorrosh et
al.,1995).

Plastid ACCase activity is regulated at the biochemical level by light and by
feedback regulation (Post-Beittenmiller et al.,1991; Post-Beittenmiller et al.,1992;
Shintani and Ohlrogge, 1995). There is also evidence which suggests plastid
ACCase is regulated at the level of gene expression. For example, in tobacco and

castor, expression of the BC gene is much higher in developing seeds than in leaf

81

or root (Elborough et al., 1996; Roesler et al.,1996; Shorrosh et al.,1995). In
developing castor and Brassica napus seeds, ACCase activity as well as
expression levels of BC and BCCP correlate with oil deposition (Roesler et
al.,1996). Using in situ hybridization analysis, BC and BCCP were found to be
expressed in a coordinate fashion and were correlated with oil accumulation during
seed development in Arabidopsis (Choi etal.,1996). The plastid ACCase subunits,
BC, BCCP, and or-CT are encoded by nuclear genes, whereas B-CT is encoded by
a plastid gene. It is not known how overall ACCase expression is controlled or how
levels of the four subunits are coordinately regulated. However, recent results
indicate that antisense or over-expression of the BC subunit in tobacco did not alter
expression of the BCCP subunit (Shintani et al., 1995). Nor is there research to
investigate what cis acting elements are responsible for the regulation of ACCase.
To begin to address these questions we describe the isolation and characterization
of a biotin carboxylase genomic clone from Arabidopsis. Furthermore, we report
initial studies to determine the cis acting elements of the promoter of the BC subunit

of ACCase.

Materials and Methods

Isolation of genomic clones and DNA sequencing
A biotin carboxylase cDNA clone from tobacco (Shorrosh et al., 1995) was
used as probe to screen an Arabidopsis thaliana 'Columbia' genomic library in A

GEM 11(Promega) vector. Approximately 106 plaques were screened using

82

Colony/Plaque Screen (New England Nuclear) membranes according to the
manufacturer's instructions. Hybridization with the ”P probe was carried out under
conditions as described (Shorrosh et al., 1994). Positive clones were puriﬁed and
then subcloned into pBluescript SK(+l-) phagemid (Stratagene) using Sacl, Hindlll,
and EcoRI sites. Both strands of plasmid clones were sequenced using universal
and synthetic primers and AmpliTaq DNA polymerase (Boehringer Mannheim).
Plant RNA isolation and Northern blotting

The RNAs from leaf, root, ﬂower, and silique, were prepared as described
(Sambrook et al., 1989). Total RNA (20 pg) was then fractionated on a
formaldehyde gel and blotted to Zeta-Probe nylon membranes (Biorad, Richmond
CA) as described (Sambrook et al., 1989). The membranes were prehybridized
overnight at 42°C in 5X SSC, 10X Denhardt's solution, 0.1% SDS. 0.1 M potassium
phosphate pH 6.8, 100 pg/ml salmon sperm DNA. The membranes were then
hybridized overnight with 1x10°CPMlml random primerazP-dCTP labeled probei n
5X SSC, 10X Denhardt's solution, 0.1 M potassium phosphate pH 6.8, 100 ullml
salmon sperm DNA, 10% dextran sulfate, 30% forrnamide. Blots were washed 3
times at 65°C in 0.1X SSC, 0.1% SDS.
Primer extension and RT-PCR

Primers were labeled by 32P in 10 pl labeling solution (1x Kinase buffer, 20
pmols of primer, 30 uCi v-32P-dATP, 10 units T4 polynucleotide kinase). The
reaction was incubated 30 min at 37°C and terminated by heating to 70°C for 10

min. Labeled primer (3 pmols) was added into annealing solution (10 mM Tris-Cl

83

pH 8.3, 50 mM KCl, 10 pg of total Arabidopsis RNA from developing seeds) in a
ﬁnal volume of 15 pl. The rest of the procedure was conducted as described
(Ausubel,1990). The cDNA was collected by centrifuging at top speed for 15 min
in a microcentrifuge and the pellet was washed twice with 70% ethanol and dried
in speed vacuum. The dried pellet was dissolved in 6 pl sequencing loading buffer
and denatured at 90°C for 5 min, then cooled quickly on ice. A 3 pl sample was
analyzed on 8% polyacrylamide sequencing gel containing 7 M urea.

For RT—PCR, ﬁrst-strand cDNA synthesis and cDNA ampliﬁcation were
performed using total RNA from developing seeds and CapFinderTMPCR cDNA
library construction kit (Clontech, Catalog# k1051-1). The primers used for
amplifying the fragment of interest were one near the transcription start (5') and a
set of 3'-primers located in the second and third exon (Figure 3.1). The PCR
products were cloned into pBluescript SK(+/-) for sequencing.

Promoter-GUS plasmid construction

As shown schematically in Figure 3.3, four promoter/GUS fusion constructs
were prepared to examine the expression pattern of the BC promoter and its cis
acting elements. Because there were no convenient restriction sites between -140
and +345, the Hindlll/EcoRl fragment (-140 to 748) was subcloned into the
polylinker of pBluescriptSK (+l-). Two PCR primers (JO380 and J0379) with
addition of Xhol and Xbal sites at the end (Fig. 1) in combination with T7 primer on
the vector, were used to amplify two promoter fragments (-140 to +345 and -140 to

+147) by PCR. The two PCR fragments, which were digested by Hindlll and Xbal,

84

were designated 380 and 379, and then directionally cloned into Hindlll/Xbal site
of pBl101.1 preceding GUS. Subsequently, the 380 and 379 fragments were
ligated back to the -2100 to -140 fragment at its Hindlll site and the two fragments,
-2100 to 345 and -2100 to 147, were then inserted into the Xbal site of pBl101.1
which generated the constructs designated as BF380 and BF379. The binary
vectors were transferred (An, 1987) from E. coli into Agrobacten'um tumefaciens
strain LBA4404 (for tobacco suspension cells) or strain C58C1 (for Arabidopsis).
Tobacco suspension cell and Arabidopsis transformation

Tobacco suspension cells (Nicotiana tabacum L. cv. bright yellow 2) were
maintained in liquid medium containing Murashige and Skoog basal salts (Gibco,
Grand Island, NY), 3% sucrose, 2.5 mM MES/KOH pH 5.7, 1 mglml thiamine, 1
mglml myo-inositol, and 1 pM 2,4-D. Cultures were subcultured weekly with a 5%
(vlv) inoculum from a 7-day-old culture and shaken at 28°C in 50 ml ﬂasks.
Agnobacten'um transformation was carried out as described (Rempel and Nelson
1991). The cells were pelleted by centrifuging at 10009 for 5 min at room
temperature in a swinging bucket rotor and were washed two additional times
before spreading on selection plates (liquid medium with addition of 0.7% phytagar,
100mglL kanamycin, and 500mg/L carbenicillin). After three weeks, independent
transforrnants were transferred to liquid medium containing 100mglL kanamycin
and maintained as described above.

Arabidopsis thaliana ‘Columbia' plants were grown with 16 hours light/8

hours dark at a temperature of 22°C. After 4-6 weeks, when the primary

85

inﬂorescence were 10-15 cm tall and the secondary inﬂorescence were appearing
at the rosette, the vacuum inﬁltration method was used for transformation as
described (van Hoof and Green,1996). The transformed plants were grown until
harvest under the same conditions as before transformation. Collected seeds were
screened on kanamycin (50mg/L) plates and resistant seedlings were transferred
to soil for further GUS assays.

Fluorometric B-glucuronidase (GUS) assays and histochemical assays
Protein extraction and GUS assays were carried out as described (Jefferson, 1987)
with Arabidopsis tissues (root, leaf, ﬂower, and silique) of independent
transforrnants and transformed tobacco suspension cells. Assays were conducted
at four time points (0, 15, 30, 45 min) with each time point containing 20 pl protein
extract and 20 pl assay buffer. Reactions were terminated by addition of 1 ml of
stop buffer and after all reactions were terminated, another 0.5 ml of stop buffer was
added to each tube. Methyl umbelliferyl (MU) concentration was determined with
a spectroﬂuorimeter (excitation at 365 nm, emission at 455 nm) and GUS activity
was expressed in terms of pmol MU/minlmg protein.

GUS activity was localized in different organs of F2 transgenic Arabidopsis
transformed with the BF380 construct by histochemical staining with X-gluc as
described (Jefferson, 1987) with the following modiﬁcations. Fresh tissue was
incubated in reaction solution (1 mM X—gluc, 50 mM sodium phosphate buffer pH
7.0, 1 mM K“ ferricyanide/ferrocyanide mixture, 0.01% Triton X-100, 10 mM 8-

Mercaptoethanol, 20% methanol, 1 mM EDTA) at 37°C for times which varied from

86

1 to 12 hour. After staining the tissue and rinsing with 70% ethanol, observation

and photographs were taken under a dissecting microscope.

Results

Analysis of the biotin carboxylase genomic clone

The complete Arabidopsis BC genomic sequence was obtained from multiple
overlapping genomic subclones (Figure 3.1). To determine the position of introns
and the translation start site, we compared the Arabidopsis genomic sequence to
the tobacco BC cDNA deduced amino acid sequence (Shorrosh et al., 1995). The
predicted amino acid sequence of Arabidopsis BC deduced from the genomic DNA
sequence shared a very high (92%) sequence identity to the tobacco BC deduced
amino acid sequence. Further evidence substantiating the placement of introns is
that the predicted protein from the genomic sequence is 100% identical to a
deduced 120 amino acid sequence of an Arabidopsis EST (150M20T7). Based on
these analyses, we concluded that the isolated genomic clone is the Arabidopsis
BC gene and contains 15 introns. Placements of introns-2 through intron-14 were
made based on comparison to the tobacco BC protein, in addition to the presence
of the conserved intron boundary sequence GT/AG (Figure 3.1). Because of a lack
of sequence similarity between Arabidopsis BC and tobacco BC in the transit
peptide region, intron-1 was identiﬁed by comparison of the genomic sequence to
that of RT-PCR products prepared from developing seed RNA (see Materials and

Methods). The Arabidopsis genomic BC predicted protein and the tobacco BC

87

sequence also shared no signiﬁcant similarity over the last 15 amino acids.
Therefore, placement of the 3'-most intron was based on comparison with additional
sequence obtained from EST clone 150M20T7.

The translation start site (indicated in Figure 3.1) begins with the ﬁrst Met
codon after the predicted transcriptional start site. The ﬁrst 70 amino acids following
this MET have a high SERIT HR content, only one acidic amino acid, and a high
proportion of hydrophobic amino acids. These characteristics correspond to a
typical chloroplast transit peptide. We have tentatively identiﬁed the transit peptide
cleavage site between amino acids 71 and 72 based on the consensus cleavage
site described by Gavel et al. (1990). The transcriptional start site was identiﬁed
by primer (JO381, Figure 3.1) extension of RNA extracted from seedlings. A
putative TATA motif was found 49 bp upstream from the transcriptional start site
and a putative polyadenylation signal was identiﬁed at +4353. Several interesting
sequence motifs were also found in the BC promoter and the 5' untranslated region.

First, there are three short CT stretches (TCTCTGTI'TCTCTG'ITTCTCTGTTT,
TCTCTCTCTCTCTCTC, and TCTCTCTCTCTCTC) located at position +310, +1 15,
and -745, respectively (Figure 3.1). Second, two clusters similar to the SEF4
(soybean embryo factor) binding motif (Lessard, 1991) were identiﬁed in the BC
promoter. Each cluster contains three copies of binding sites and the two clusters
were separated by about 950 bp (Figure 3.1). Lastly, two large (approximately 480
and 280 bp long) segments of repetition were found in the BC promoter; one repeat

was between -1289 to -888 and -478 to -84, and the two regions share 75%

88

-2051
-2028
~1968
-1908
-1848
~1788
-1728
-1668
-1608
-1548
-l488

-1428

-1388
-1328
-1268

-1208
~1148
-1088
-1028
-968
-908
-848
-788
-728
-668
-608
-548
-488

-428

-368

-308

-248

-188

-128
-68

52

EcoRI
GAATTCAAATTATTTTTCTATGA
TTTTGTCCCTTAAATTATAACTGATCTACTTCATATAGACTATTATAACTAAAATCATTT
CTAGTCAATAATAGAAGATGCTACGAACAAAAGCACTTAATTTTGAAACCTTAAATAAAA
TACATAGTATAACGATTTTTATATACTTATCTATTAGATCACAAAAACTAAATTCAATGT
TAAAAATAAAGTACATAAATTTATTTGAGATATGGAAGAGGTAAGAAAATATTATTTGAT
AAATATATCACAACAATTATTTTGTTAAATATTACATAAATAAAATGACACACGAATTAT
TATCGAAAATAGTCAAATATCCAAAATAGATAATAGAGATAGATCTTTTCAACTTCATTT
CCTTGCATGATGGTCAAAATTGATATTTTGTTACATGTATCAAGACTTCCAGTTTACAAA
GGTGAATTTGAACTAAAATAAATATGAAAATCAAAGAAGAAATAAGTATAATCTAGTAAC
ACAATATAAATAAAGGGTTTTGTGCAAAAAAAACTGAAATTGTATATTGTTGCAGCTTTA
CCCTTCACGTTAAAAAAATGCAAATAATCCTATAAATTTTAAAAATAATGCGAAATATAG
SEF-4
CCATTTTTTTGCAAATTTACATATATTGTAAATATTTTCATGATCACATACATTTGTTTT
SEF-4
TGAATTAGACTCTTCCAATATAGCAATCTATTGAAGAAATAAGTATTATTTATCAAAAAA
TTGAATTTTTGACAATAGCAAAAAAGTTAATTTGCGTATTATTTTGAAATTCATAAGTAT
TTTGCATTTTTTTATGTGAAAGATAAAATTGCAAAAATAGACGAATTTGAAAGTTGTTTT
SEF—4
TGCACAAAACCCAAAATAAAATACAACAATATAGGATATATAGATCATAGCCTCTAAAAA
TAAATTTAAAATTAGATTCTTCCAATGATGTTTCAACCCATATAACAACAGAAAAGCAAA
ACTAAAATATTTAAAATTTGAGATAGGTTCGAAGATGATTATCTTTGGAACTTGGTTTTC
TTGATGTATATGATTAAATTTGGTGCATATGTAATTAAGTGTGCACTGGACCAATAAAAA
ACATTGAAACCTTATATAATATATCTTCTTATGACACATTATTATTTAAATAAACAAACA
AACAAAACTTTAAAAGCGTTTGCGTTCAGCGTTCAGCGTTCAGCGTTTGCGTTCGACGTT
TAGAGTTGGCACAAAATCCACCAAATTAAAACCATTCNGGCGGAGGATTCAATATATCTC
ACAGTTTCACTCTCAACATTTCAATATCTCTCTCTCTCTCTGGGAAGAACAAATACAGCG
TTTCTTCTGCGATTAGATCTATTTCGATACTCACGAGGATTGCCTCCTGAGGAGACTACA
TATACTAAACAAAATACAAGCTGAGGAGACTAAATATATTTAGTTAATACCAATAATATA
TGATATACAACTACATATTTACAAACAAAGAAAATATAGTAGATTTGTAATATATACATA
ATTGAGATTTGATGATTGATTAAACCAAGAATACCGTGCGTAGCACGGGTACTGACATAG
TATGTGTATTATTCTGAACTTCATAAGATTATTTTTCATTTTTGATGTGAAAGATAAAGT
SEF-4
TGCAAAAATGGTCGAATTTGAAGGTTTTTTGCACAAACCCAGAAATAAAATAAAACAATA
SEF—4
TAAGATATGATCATAGCCTCTGAAAATAAATTTAGAACTAGACTCTTCCATTGATGTTTC
AACCCATATAACAACATAAAAGCAAAGCAATAAGTTGAAAGATGATAATCAAAATTAAAA
AAATGAGTATCTTTGCAACTTGGTTTTCTTGATGTATATAATTAAATTTGTTACATATGT
SEF-4 HindIII
AATTAAGTGTAAACTGGACCCATAAGACCTTTTTCATCCTTGTAAGCTTAATGGGCTTTA
TCAATATAATGTTGGCCGACAAAAAAAAAAAAAAAAAGACCCATAAGAAACATTGAAACC
AGTCTGGCCCATGACCCATATAATTTATCTTTAAAAGAGTCTGCGTTCCACGTTTGCGTT
V Jo381 3 ' -GGTC
QQGAGTTTGCGTTCCGTGTTTGCGTTTAGAGTTGGCACAAAATCCACCAAATCAAACCAG
AGCCCGCCTCCTAAGTTATAG-S'
TCGGGCGGAGGATTCAATATCTCAGAGTTTCACTCTCAACATTTCAngggggggzgggg
J0379

89

112
172

232

292

352

412

472

532

99

592

652

712

772

832

892

952
1012

1072

3'-ATCTAGACAAAGCTATTAGTGCCTTagatctgagctcggc-5'
QTQAACAATTAGATCTGTTTCGATAATCACGAGGTTTCCCTCACTTCCTTTCCATTGCTA
AACCAGGATTTGCCTCCTGCATTTGGAAATGGACGCTTCTATGATTACCAATTCCAAATC
M’ D A S .M I T .N S K S
CATTACTTCTCCACCCgtaagcaccaaaacgctattaatttgaaattcgtcaatctctgt
I T S P P
J0380
3'-AGAGACTTAGACAATTTAGACAATATACTTC

11

16

agatctgagctcggc-S'

ttctctqtttctctqtttcatgtctctgaatctgttaaatctgttatatgaagTCTTTAG
S L A
CCTTAGGGAAGTCAGGAGGAGGAGGAGTTATCAGAAGTTCACTATGTAACTTAATGATGC
L G K S G G G G V I R S S L C N' L M’ M’ P
CAAGCAAAGTTAACTTCCCTAGACAAAGAACTCAAACTCTAAAGGTTAGCCAGAAGAAAC
S K V .N F P R Q R T Q T L K V S O K K L
TCAAGAGAGCTACTAGTGGTGGTCTTGGTGTTACATGCAGTGGTGGTGATAAGATTCTCG
K R A T S G G L G V T CV/S G G D K I L V
TAGCGAATCGAGGTGAAATTGCAGTTCGTGTTATCAGAACTGCTCATGAAATGGGGATTC
A N R G E I A V R V I R T A H E M G I P

CTTGTGTTGCTGTGTATTCTACTATAGATAAAGATGCTCTTCATGTTAAATTAGCTGATG
C V A V Y s T I D K D A L H V K L A D E

AAGCTGTTTGTATTGGTGAAGCTCCTAGTAACCAGTCgtaagtgtggaacaaaatgattg
A V C I G E A P S N Q s

S - - _ - - - - _ - - -
EcoRI
atcttgntgtgtagagaatgaactagtgttgaattcttgtccttttgtgtgtagGTATTT

Y L
GGTGATTCCGAATGTTCTGTCTGCGGCTATTAGCCGTGGATGTACAATGCTTCATCCTGG
V I P N V L S A A I S R G C T M L H P G
ATATGGTTTTCTTTCGGAGAACGCTCTCTTTGTTGAAATGTGTAGAGACCATGGGATCAA

Y G F L S E N A L P V E M C R D H G I N
- - - - A - - - V - - - - - - E - - - -
TTTTATTGGACCTAATgtaagtgtcttatatctcttcaaagactttatagactttgcaga
F I G P N
acttcattagtctcagttcagccctagctttcttttgttggagagaaggaagatttcagt
atctgatgtatcgtattcgtctttcctatctgtagCCTGATACCATCCGTGTTATGGGAG
P D S I R V M G D
ACAAAGCGACTGCAAGAGAGACAATGAAAAATGCAGGGGTTCCCACTGTACCAGGGAGTG
x A T A R E T M K N A G V P T V P G s D

90

19

39

59

79
76

Ara

96

119
116

131

128

133

130

153
150

173
170

178

175

187

184

207

1132

1192

1252

1312
1372
1432
1492

1552

1612
1672
1732
1792
1852
1912

1972

2032
2092
2152

2212

2272
2332
2392
2452

2512

2572

_ S _ _ - D - _ _ - - - - - - - - - _ _
ATGGGCTATTGCAthatttttttagattatcagtttagnatttcaagtttttgcatgtc
G L L Q
aagagangtcttgggatttgaatgttttattttgtcttattgcagAGTACAGAAGAAGCT
8 T E E A
- - - - G
GTCAGGGTCGCCAATGAGATTGGTTTCCCTGTAATGATCACGgtaatattaacttgattg
V R V A N E I G P P V N I T
- - L - E - — - Y - - — - K
gatttggagttaaattcgtatttcatactttcttttatgtaaccgataaactggattctc
aaaattggtctgcttgaaaagacacaaatttaagtatgtttattttatgctcagcttgtc
tccctgtacttattgatatgcttatagttacaaaaactgactgttctcacttttctggtt
tcaagGCAACAGCTGGTGGTGGTGGACGTGGAATGCGTCTTGCTAAGGAACCGGGAGAGT
A T A G G G G R G M R L A R E P G E F
- - .. - - .. _ .. .. - - .. - _ - .. D .. -
TTGTAAAACTGTTGCAthatgacaatcatgtaaatctgaaccaacaattcctctgatag
V K L L Q
aatttttaatgagtccagggatcaagttctccataagcataattgcagnattgncacaga
ttctcnatnnctnnctaactcagacaatgtaaaggtgtnatctcatgatgnttncccaca
nnncgnaaannaagccncctgnatagttgtgaaanctggaagtgtagtancntaaatcta
gtaatnctgaccattgatatcaantacnccaaccagattgtatgctttccgcttgccttt
atagtcacagttttcttaaatgaaaattccacaaaagattcgacttccctttattgaact
tggcttaatttttttatcgacagCAAGCTAAGAGCGAGGCTGCTGCTGCTTTTGGGAATG
Q A R s E A A A A P G N D
ATGGATGTTATCTGGAGAAGTTCGTTCAAAACCCTAGACATATTGAGTTCCAGgtatgca
G C Y L E K P V Q N P R H I R P Q
- V _ - _ _ Y - _ _ - - _ - .. .. -
ttttatcaggagaagttgncaaaacagttgaggttttancaaaagatgattcttgtgagt
tgatggtctagtgatacacattattgatcccaaagccatattttctctcngtacctctng
tatttcagGTGCTGGCAGATAAATTTGGAAATGTTGTTCACTTTGGTGAGCGTGACTGCA
V L A D K P G N V V H P G E R D C 8
- .. .. - .. Y - .. - - .. - .. _ .. _ - ..

GCATCCAthatcttcctcttaggttttttataatttcattaccatcaattgttgcatag
I Q
tgctccaagaataaatgtaactgaatatatcaaaacaggaataaatttacttcagcctaa
ttactgcacttgcatgtcatatcccagctgcatgatctgctaaactgattttttcctgta
ctaattgctaagataactcgaacttcgatactagttcacagaatttgagtatacctcttg
ctgctttctttcaattcttgtatgacatctaaaaccatctggatcatttaattatatctt
HindIII
cccctgacaacagAGGCGTAACCAAAAGCTTCTGGAAGAACCTTCTCCAGCATTGACCGC
R R N Q R L L E R P 8 P A L T A
- - - - - - - - - A- - - - - - P
TGAATTGCGAAAAGCCATGGGTGATGCAGCAGTCGCAGCAGCAGCATCCATTGGGTACAT

91

204

211
208

216
213

230
227

249
246

254
251

267
264

284
281

302
299

304
301

320
318

2632

2692

2752

2812

2872

2932

2992

3052
3112

3172

3232

3292

3352

3412

3472

3532
3592
3652

TGGTGTTGGTACTGTGGAGTTTCTTTTAGATGANAGAGGTTCCTTCTACTTCATGGAAAT
G V G T V E F L L D E R G S F Y F M E M

GAACACTAGGATCCAthtgaattttattatgactgctaagtatatttataagcatctga
N T R I Q
agtatactgaatgcgaaccatttcttgaatctaaactcagGTGGAGCATCCTGTGACAGA
V E H P V T E
GATGATTTACTCTGTTGATTTGATAGAGGAGCAGATTCGTGTTGCAATGGGAGAGAAACT
M I Y S V D L I B E Q I R V A M G E K L

_ _ S - _ _ - _ _ - _ _ - - _ _ _ _ - _

TCGTTACAAACAthttgaatagtgttgtatatgatgttttgagtggataccaaaaaagt
R Y R Q
tatttcaaaatttaattttgtttgtttacagGAAGATATTGTGCTCAGAGGGCACTCAAT
E D I V L R G H 8 I
TGAATGTCGTATCAATGCAGAAGATCCATTTAAAGGATTCAGACCTGGACCTGgtatatt
E C R I N A E D P P R G P R P G P G
_. _ _ .. _ .. _ - A .. - N _ .. - .. - ..
tcagtagacttttatatgttaacgtcttcgaattttagcagattggctttctttaaaata
acgtcactcttacaacaacagGCAGAATAACATCATACCTGCCATCTGGAGGTCCTTTCG
R I T S Y L P S G G P P V
- - - A - - - A - .. - - ..
TTAGAATGGACAGCCATGTCTATTCCGACTATGTTGTGCCTCCAAGCTATGATTCTCTTC
R M D S H V Y S D Y V V P P 8 Y D s L L
- - - N - - - P - - - - - - - D - - - -
TTGGGAAthaatatctgtcttttactttctctggatagccatttagtatctgtttggtt
G K
atttaggaacaaattttgtggcttcatggtaaaataatcactcgggtgttttggaatctt
aaacaaatattggaaaagtcttcctggtgttgttcatttgcttctatagcagcagcttac

tggaacttatagctgatgtgttgacacactaaatctcttgtagCTTATTGTGTGGGCTCC

L I V W A P
AACGAGGGAAAAAGCAATTGAACGGATGAAGCGGGCGCTTAATGACACTATCATTACAGQ

T R E K A I E R M R R A L N D T I I T G

- - .. G - - - - _ - - .. - - - .. .. - _ ..
tatgtcgagtactactttttgttaattccatattgtttatgcttcttcattgtctgtttt
gtttttcttttattctgtgtaagatgtgacatttcgttacattggacatctggtggaatg
ttaaatcaaatttcgttttcttggtttcatcagGGGTTCCAACGACTATCAATTACCACA
V P T T I N Y H K

92

340
338

360
358

365
363

372
370

392

390

396

394

406
404

423

422

437

435

457
455

458
457

465
463

485
483

494

3712

3772

3832

3892
3952
4012
4072

4132

4192
4252
4312
4372
4432
4492

- _ - _ _ E - _ _
AACTTATCCTTGATGTTGAthgtgtgaaataagtaaagctatagtttagcgaattttgt
L I L D V E
- - - - I _
aatgaaaatgttggtttttttgtaatccaattgttgtgttggtcacagGATTTCAAAAAT
D P K N
GGAAAAGTTGATACAGCTTTCATCGTCAAGCATGAAGAGGAGCTAGCTGAthaaattcg
G K V D T A P I V K H 8 E E L A E
- - F — P S - — P - — G G — - — p
aatgtctctgtggtcggctaaagacacaggccattggatgtattcattttcaatgcttct
aagttagtgggatatgaagcctttggtaaagctgaacatagagataggagaattgcatga
gaagaatgaaactatatatagaggaagacagattaaattgtgtttgtaatagtaaggttg
gttaaaccaaatggtgggttctaacatggttatattgtatatgcagCCTCAAGAAATTGT
P Q E I V
- H K M —
GGCAGTGAAAGATCTGACAAACGCAACGGTTTAGAATGATACATCAGCTCTGAAAGACCA

A V K D L T N A T V *

P A A T K E M V N A S A *
CCCAATTATTGCAATCTTCGTCTCCTTTTGTGTGTTCATTGGTACAAACTTCGGTAACGA
TAAGACTGTTTTACTAGAGTCTGTGGTTTTATCAATGTTCTTCTTGTATCATAAACAAGA
GGAAACACTTTTGTTTGTAGTCTCTATGTCAAGATTTTTCAATAATTTTCATATATAGTA
GAGATTATATGAACAAGGCCTATCACTATTAACAAAGTAACCAAAACCAAATAAAACAAA
AGACAAAAATAATTCACATTTAGGTGATCTCATCGCGGGCCGGACACTTCCTGAGACGCT
ATGTAAGACGCAGACTGCCTAGAGCCGCCACTATTAGAGCTC

SacI

492

500
498

504
502

521
519

526
524

536
536

Figure 3.1. Structure and sequence of the biotin carboxylase gene from

Arabidopsis. The transcription start site is marked with a V, and a putative TATA

box is shown in bold and underlined. A putative polyadenylation signal is shown in

bold. Underlined regions are discussed in the text. lntrons are shown in lower case

and GT/AG nucleotides corresponding to the intron boundaries are shown in bold.

The derived amino acid sequence is presented in single-letter code.

In the

alignment with tobacco BC, identical amino acids are presented by‘ -‘ otherwise the

single-letter code is used. Primers J0380 and JO379 were used to make

promoter/GUS constructs. Primer JO381 was used for primer extension.

93

identity. A different repeat occurs between -972 to -670 and -86 to 181, and these
regions share 67% identity.

Gene expression pattern and BC promoter expression analysis

An Arabidopsis Northern blot was probed with the 2.7 kb Hindlll genomic fragment
of BC and a single band was detected in all tissues tested (Figure 3.2). Expression
levels of BC were quantiﬁed, and variability attributable to unequal RNA loading
was estimated by probing with ElF4a (eukaryotic initiation factor). The relative
expression level of biotin carboxylase based on the ratio of BC/ElF4a signal was
found to be highest in siliques (0.54), ﬁve fold lower in leaves (0.10), and at
intermediate levels in ﬂowers (0.39) and roots (0.27).

The expression pattern of the BC promoter during early seedling
development and in mature plants was examined in Arabidopsis plants transformed
with the BF380 GUS construct (Figure 3.3). To locate BC promoter activity by GUS
expression, seedlings were stained with 5-bromo-4-chloro-3-indolyI-B-glucuronide
(X-gluc) at various development stages. GUS staining was found in roots of young
seedlings (48 hours after germinating), but not in cotyledons (Figure 3.4). After 3
days, GUS staining appeared in roots and radicles, but still not in cotyledons. At
4 days, GUS activity could be detected throughout the seedlings, but was still
highest in root tips. GUS staining was observed in all tissues of mature plants but
the most intense staining occurred in anthers. Wrthin leaves, staining was
strongest in the vascular tissues. Because staining rates varied in tissues of

mature plants, and may in part reﬂect rates of substrate access to the enzyme, it is

94

 

 

IEF4a U

 

 

 

BC

 

 

 

 

Figure 3.2. Northern analysis of BC mRNA levels in F (ﬂower), L (leaf), R (root), and
S (silique) from Arabidopsis. Total RNA (20 pg/sample) was separated on a
formaldehyde gel and blotted to a nylon membrane. This blot was hybridized with
a 32P-Iabeled probe of the Hindlll fragment from BC genomic clone. The same blot
was stripped and hybridized again with 32P-labeled ElF4a probe as a loading

control.

95

Transcription
start site

 

 

 

 

 

Er: RI Hindlll t ‘- u; in:
(J
I H) II)
SEN SEF4
TATAAT
-49
ECORI Hindlll Xbal Xbal
-2051
I BF
I BF

 

 

Figure 3.3. Structure of BC promoter-GUS deletion constructs used for Arabidopsis

and tobacco transformation. The numbers are relative to the transcription start site;

a putative TATA box at position -49 is also indicated. ATG represents the

translation start codon of the GUS gene.

96

 

Figure 3.4. Localization of GUS expression in transgenic Arabidopsis transformed
by construct BF380. (A) Two-day-old seedling stained for 3 hr. (B) Three— (left)
and four-day-old seedlings stained for 2 hr. (C) Mature leaf stained for 12 hr. (D)

Flower stained for 1 hr.

97

difﬁcult to make an accurate comparison of staining levels in different tissues.
However, as judged by the time required for staining, the order from the fastest to
the slowest was anther, silique and embryo, root, stern, and leaf (Figure 3.4).

To further characterize the relative expression level of the BC promoter in
different tissues, ten independent F2 transgenic plants of Arabidopsis (transformed
by BF380) were screened for GUS activity. Although total GUS activity varied from
plant to plant, each independent transfonnant had a similar expression pattern with
the highest activity in the silique, followed by ﬂower and root, and the lowest activity
was in leaf (Figure 3.5).
BC promoter deletion studies
To identify functionally signiﬁcant domains within the BC promoter, deletions were
made from the 3' and/or 5' ends of the promoter and these constructs were fused
to GUS in the plant transformation vector, pBl101.1. The deletions and structure
of each derivative are diagramed in ﬁgure 3-3. These constructs and a 358/GUS
control were used to transform tobacco suspension cells. Fifteen independently
transformed lines were recovered for each construct, and then grown in liquid
medium with subculturing every 7-8 days. Constructs BF380 (full length), 380 (5'
deletion), 379(5‘ and 3' deletion) and 358 all had similar levels of GUS activity.
However, construct BF379 which is deleted in the promoter 3' end but not 5'
showed approximately 5 fold higher GUS activity (Figure 3.6).

The expression of ACCase has previously been found to be tightly

associated with growth rate in E. coli (Li and Cronan, 1993). Therefore, we

98

m
aBadivityindffererttise l

 

n)
83%

 

i

 

 

 

5”

GBadivity (pr'dlm'rmgp'mi

 

 

 

 

Figure 3.5. GUS activity in roots, leaves, ﬂowers and siliques of independently
transformed Arabidopsis plants. The construct used fortransformation was BF380.
The insert (upright) presents the average GUS activity (iS.E.) in different tissues
from the ten independent plants.

99

Expresion level of different constructs

 

 

 

 

U}
21
8000 ﬂ 4500 3
l‘ 4000
l l
l 7000! 3500 ‘
l L 3000
l 2500
6000 .(
‘i g 1 2000 .8 a E:
1500 .V’ w 0)
c; 50°°l g S
L g: ‘ 1000 j :
E l
i i: 4000 l 500 1 [i1 l
‘ o l
l E l '7 7 7 i
l v l {
l .5 l
. {3 l
l g l i
l 2000l l
i ; l
1000 l .
l l l
l .
i o.‘ I ‘
1 358 BF380 BF379 380 379 E
i Constructsusedfortransfonnation ‘
L... J , a ,- ., - 7 ,7V.7,, ,. v.7 #l

Figure 3.6. Comparison of GUS activity in transgenic tobacco suspension cells
transformed with different BC promoter- and 35S GUS constructs. GUS activity of
15 independent lines from each construct was ﬂuorometrically assayed and the
speciﬁc activities calculated based on extract protein concentration. The insert
(upright) presents the average (iS.E.) GUS activity of 15 cell lines transformed by

different constructs.

100

examined this relationship in the tobacco suspension cells. The growth rate of
suspension cells increased over the ﬁrst four days after subculturing and then
decreased (Figure 3.7). GUS speciﬁc activity of all constructs remained almost
unchanged in the ﬁrst three days after subculturing. After day three all cultures
showed a several fold increase in GUS speciﬁc activity as the growth rate
increased. Although the general patterns of increase are similar, the timing of the

increases and the peak speciﬁc activities varied among the constructs (Figure 3.7).

Discussion
Because of the importance of ACCase activity for regulation of fatty acid synthesis
we are interested in understanding the control of expression of ACCase genes. As
a starting point we have begun characterization of the BC promoter. In this paper
we describe the isolation, sequence and expression of the Arabidopsis BC gene
and its promoter. We were able to identify a number of cis-genetic elements,
including transcriptional start site, translation start codon, distribution of introns and
exons, and polyadenylation signal. Northern blot analysis demonstrated that BC
is expressed at higher levels in silique and ﬂower and lower levels in root and leaf.
The histochemical staining of Arabidopsis plants transformed with the BF380
construct demonstrated that anthers (pollen) stained most rapidly followed by
silique, root and leaf. Quantitative GUS assays of tissue extracts provided
comparable results (Figure 3-5). The higher level of GUS activity in silique and

embryo is understandable as a high rate of fatty acid synthesis is required to meet

101

asmmdmutdaysmawﬁdﬁ

 

3 g

'3

FL

.5535

 

aBadiu'tyWnirmg pain)

 

0

Figure 3.7. Changes of GUS activity during growth of tobacco suspension cells
transformed by different constructs (A). The growth rate of suspension cells is

shown in the right box in terms of mg weight increase/mg cells/day (B).

102

triacylglycerol synthesis in oil storing Arabidopsis seeds.

Histochemical analysis and quantitative GUS assays presented here are
consistent with previous analysis of expression of ACCase and other lipid
biosynthetic genes. For example, BC expression levels were found to be directly
correlated with ACCase activity and oil deposition in castor and B. napus seeds
(Roesler et al.,1996). The high level of GUS activity in ﬂower may be due to higher
amount of lipids required during pollen formation and the synthesis of some
polyketides (Evans et al., 1992). Similar patterns of expression in pollen grains
were observed when the A1 ACP gene from Arabidopsis and the stearoyl-ACP
desaturase gene from B. napus were expressed in tobacco (Baerson and Lamppa,
1993; Slocombe et al., 1994). In addition to the high expression level in oleogenic
tissues, high levels of activity were also observed in tissues undergoing rapid
growth (e.g. root tip, meristem). These results are consistent with the notion that
genes involved in FAS would be expressed at high levels in tissues requiring
elevated levels of lipid synthesis. In summary, the BC gene is regulated, at least
in part, at the level of gene transcription in a pattern similar to other FAS genes and
which is apparently correlated to the rate of tissue fatty acid synthesis.

To date only a few studies have focused on fatty acid biosynthesis gene
promoters (Baerson et al., 1994; de Beer et al.,1996; de Silva et al.,1992;
Slocombe et al., 1994). One consistent trend, found with Arabidopsis BC and
previous studies of promoters of FAS genes, is that such promoters show the

highest expression levels in rapidly growing tissues and developing seeds. Two

103

conserved elements (GCCCAT and ATGGGC), found in the B. napus ACPO5,
ACP09, and an Arabidopsis ACP gene were also identiﬁed in the Arabidopsis BC
promoter (located at -62 to -57 and -138 to -133, respectively) (Figure 3.1). These
elements have previously been observed in seed-expressed genes (Doyle et
al.,1986). The (CT)n stretch, which we found in the Arabidopsis BC promoter, also
occurs in the 5'-untranslated leader sequences of BC from tobacco ( Shorrosh et
al.,1995). three ACP genes from Arabidopsis (Lamppa and Jacks,1991; Post-
Beittenmiller et al., 1989), two ACP genes from B. napus (de Silva et al., 1990), four
ACP genes from other species (Ohlrogge et al.,1991), stearoyl-acyl carrier protein
desaturase gene (Slocombe et al., 1994), 1-acyl-sn-glycerol-3-phosphate
acyltransferase gene (Knutzon et al., 1995), and Arabidopsis HMG2 gene (Enjuto
et al., 1995). The d(CT)n tracts in eukaryotic genomes may be involved in
recombination (Weinreb et al., 1990), replication ( Caddle et al., 1990), transcription
( Wells et al.,1988), and chromatin structure (Lu et al., 1993). Proteins which
speciﬁcally bind (CT)n(GA)n promoter elements have also been reported (Gilmour
et al., 1989). So far, almost all the characterized promoters or 5' UTR of genes
involved in the pathway of fatty acid synthesis possess this (CT)n tract. The
signiﬁcance of these CT tracts is unknown, but its conservation in genes involved
in fatty acid synthesis suggests that it may have a universal regulatory function.
The importance of the SEF4 binding motif found in the BC promoter is also not
clear. SEF (soybean embryo factor) is a nuclear DNA binding protein whose

expression begins to appear in mid-maturation soybean embryo and increases

104

 

moderately during embryo development (Lessard et al., 1991). Based on the
change in SEF4 activity during seed development, in parallel with accumulation of
the B—subunit mRNA of B-conglycinin, it was proposed that the binding of SEF4 to
the promoter region could activate the expression of the [3 subunit of B—conglycinin.
These motifs may also play some role for the higher expression level of BC in the
seed.

To our knowledge there are no previous reports showing large segment
repeats in promoter regions such as those found in the BC promoter (Figure 3.1).
We do not yet know any regulatory function of these promoter repeats; however,
in comparing the GUS activity of construct 379 and BF379, the repetition region
located in region from -2048 to -140 may enhance the expression of the BC gene.

As a ﬁrst approach to identify cis regulatory elements involved in the control
of Arabidopsis BC gene expression, we analyzed the effect of BC promoter
deletions/GUS fusion constructs in transgenic tobacco suspension cells. We found
that deletions removing sequences from either the 5' and/or the 3' end of the full
length BC-GUS construct BF 380 resulted in altered expression. The ﬁrst exon and
intron were included in this construct because there are two repeat elements in the
ﬁrst intron. The ﬁrst element (CTCTGTTI') has three repeats and the second
(ATCTGTTAA) has two repeats. We suspect that the ﬁrst intron may play a
functional role in the regulation of BC expression and in this regard, the deletion of
the ﬁrst exon and ﬁrst intron (construct BF379) resulted in a several-fold increase

of GUS activity. There are at least two possible reasons for the increase of GUS

105

activity. The ﬁrst is that the intron can not be recognized and properly excised in
tobacco cells. This seems unlikely because the ﬁrst intron possesses conserved
intron boundary sequences (Figure 3.1) and in numerous previous examples introns
from one dicot species can be properly processed in other species. Furthermore,
intron junctions throughout the plant kingdom are highly conserved (Laliberte et
al.,1992; Paszkowski etal.,1992; Quigley etal.,1988; Vancanneyt etal.,1990). The
second possibility is that the intron does play a role in the regulation of BC
expression and functions to depress the expression of BC. The GUS activity of
construct 379 (5' deletion) compared with construct BF379 is decreased several-
fold. This implies that other regulatory element(s) which could enhance the GUS
expression may exist between the region corresponding to -2048 and-140. In
addition, this also suggests that the region between +140 and -147 contains the
basal promoter elements which can support the expression of BC gene. Construct
380 (-140 to +344) shows essentially the same GUS expression level as BF 380 and
379. Taken together, it appears as though positive element(s) are located between
-2048 and -147, and a negative element(s) is located between +147 and +344.
Considering the relatively low GUS expression of construct BF380 (full length
promoter), we conclude that the negative element is dominant over the positive
element. The presence of a negative element in the ﬁrst intron is similar to a recent
report that deletion of the ﬁrst intron from the enoyl-ACP reductase promoter of
Arabidopsis resulted in increased expression in roots (de Boer et al.,1996). All our

deletion experiments were performed in tobacco suspension cells, Thus we do not

106

yet know if these elements regulate BC expression in Arabidopsis tissues in the
same manner.

We have observed that GUS activity of BF380 is comparable to the GUS
activity observed in constructs under the control of the 358 promoter. This is
surprising because BC is a low abundance protein in plant cells with levels
estimated to be <0.05% of protein in leaf or root (Roesler et al.,1996). The fact that
GUS activity under the control of the BC promoter is comparable to that of 358
promoter suggests that translation of the BC mRNA may be inefﬁcient or that the

protein may have a high rate of turnover in vivo.

107

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112

CHAPTER 42

The Biosynthesis of Erucic Acid in Developing
Embryo of Brassica rapa L.
Abstract

The prevailing hypothesis on the biosynthesis of erucic acid in developing
seeds is that oleic acid, produced in the plastid, is activated to oleoyl-CoA for
malonyl-CoA-dependent elongation to erucic acid in the cytosol. Several in vivo
labeling experiments designed to probe and extend this hypothesis are reported
here. To examine whether newly synthesized oleic acid is directly elongated to
erucic acid in developing seeds of Brassica rapa L, embryos were labeled with
[”Cjacetate, and the ratio of radioactivity of carbon atoms C(5)-C(22) (de novo fatty
acid synthesis portion) to C(1)-C(4) (elongated portion) of erucic acid was
monitored with time. If newly synthesized oleate immediately becomes a substrate
for elongation to erucic acid, this ratio would be expected to remain constant with
time. However, if erucic acid is produced from a pool of pre-existing oleic acid the
ratio of 1"C in the four elongation carbons to 1“C in the methyl terminal 18 carbons
would be expected to decrease with incubation time. This labeling ratio indeed
decreases with time and therefore suggests the existence of an intermediate pool
of oleate which contributes at least part of the oleoyl precursor for the production

of erucic acid. Addition of haloxyfop, which inhibits the homodimeric acetyl-CoA

 

2 Material in this chapter was published previously [ Bao X, Pollard M, and Ohlrogge J (1998)
Plant Physiol. 118: 183-190].

113

carboxylase, severely inhibited the synthesis of [“Cjerucic indicating that
essentially all malonyl-CoA for elongation of oleate to erucate was produced by
homodimeric ACCase. Both light and haloxyfop increased the accumulation of
["C]oleate and the parallel accumulation of [“ijhosphatidylcholine. Taken
together, these results show an additional level of complexity in the biosynthesis of

erucic acid.

114

Introduction

Erucic acid (cis-13-docosenoic acid) and its homolog, cis-1 1-eicosenoic acid,
are commonly found in the seed oils of the Crucifereae. The oils and the
corresponding fatty acids are produced by high erucic acid cultivars of Brassica and
Crambe species and are oleochemical commodities. The reactions leading to the
synthesis of erucic acid are, for the most part, well understood. In the developing
embryos of Brassica napus (Downey et al., 1964), Crambé abyssinica (Appleby,
1974), Simmondsia chinensis (Ohlrogge et al., 1978), Tropaeolum majus (Pollard
et al., 1980a) and Limnanthes alba (Pollard et al., 1980b), it was demonstrated that
erucic acid is synthesized by the elongation of oleic acid rather than by de novo
synthesis. This conclusion was deduced from the distribution of label in the long-
chain fatty acids after incubating seed tissue with exogenous [“Clacetate. The
label was preferentially incorporated into the carboxyl-terminal carbons of the long-
chain fatty acids rather than the methyl terminal 18 carbons. Subsequently, the
work of Ohlrogge et al. (1979) demonstrated that de novo fatty acid synthesis was
almost exclusively located in the chloroplast in spinach leaves. Thus the
hypothesis on the biosynthesis of erucic acid was extended to a description in
which oleate, synthesized in the plastid, was exported to the cytosol, where,
presumably in the endo-membrane system, it was elongated via malonyl-CoA-
requiring elongases to C20 and longer-chain monounsaturated fatty acids. Several
reports (e.g. Imai et al. .1995; von Wettstein Knowles, 1993) have conﬁrmed that

the location of oleoyl elongation system was extra-plastidial, being associated with

115

oil bodies or microsomal membranes. Créach et al. (1993) and Fehling et al. (1991)
showed that oleoyl-CoA is readily elongated in vitro and the intermediates of the
elongation reaction are acyl-CoA thioesters.

It is generally accepted that the end-product of newly synthesized oleic acid
exported from plastids is oleoyl-CoA. This oleoyl moiety can be elongated directly
to erucic acid in the endoplasmic reticulum or oil body-associated membranes
through successive additions of two carbons derived from malonyl-CoA. However,
in an oil body fraction from developing rapeseed, Hlousek-Radojcic et al. (1995)
observed in vitro that radioactivity from oleoyl-CoA was incorporated into
eicosenoate and erucate at least 2.5-fold more slowly than from malonyl-CoA.
Furthermore, radioactivity from oleoyl-CoA was rapidly diluted upon the formation
of eicosenoyl-CoA and the elongation could proceed without the addition of
exogenous oleoyl-CoA. Based on these in vitro observations, they concluded that
oleoyl-CoA is not the immediate substrate for elongation. Instead they proposed
that the intermediate oleoyl donor for the elongase may be either a lipid or
unesteriﬁed acid. Furthermore, in unpublished experiments in our lab, when intact
Brassica (high erucic) embryos were incubated with [“Cjacetate,
phosphatidylcholine was always heavily labeled at early time points, with oleate
constituting over 90% of [“leabeled fatty acid esteriﬁed to phosphatidylcholine.
We considered that this oleoyl-phosphatidylcholine might contribute to the
synthesis of erucic acid, either via a mechanism of direct acyl transfer as proposed

by Hlousek-Radojcic et al. (1995), or via the acyl-exchange between acyl-CoA and

116

phosphatidylcholine, as ﬁrst reported by Stymne and Stobart (1984). In this study,
we examined whether oleoyl-CoA produced by plastids is directly elongated to
erucic acid, or if the oleoyl-CoA enters another intermediate pool before it is
elongated. In addition we have examined the inﬂuence of light and of an inhibitor
of the homodimeric ACCase on erucic acid biosynthesis. Taken together, our
results show an additional level of complexity in the biosynthesis of erucic acid, in
that the supply of oleoyl groups for chain elongation is a combination of the release
of oleate from a large intermediate lipid pool, probably phosphatidylcholine and the

direct provision of newly synthesized oleate from the plastid.

Materials And Methods
Plant material and biochemicals

Developing embryos of Brassica rapa L. (high erucic) were obtained from
plants grown in growth chamber with 16 hours illumination at 25°C. Four-week-old
siliques were taken from plants and after removal of seed coats, the resulting
embryos were used immediately for labeling experiments.

[1 -"'C]acetate (1 .74GBq/mmol) and [U-“C]oleic acid (33.3GBq/mmol) were
purchased from New England Nuclear-DuPont (Vlﬁlmington, DE). The herbicide,
haloxyfop [2-((3-chIoro-5-(triﬂuoromethyl)-2-pyridinyl)oxy)phenoxy) propanoic acid],
was a gift from DowEIanco (Indianapolis, IN 46268).

[1 -“C]Acetate incubations of rapeseed embryos

In the [1 -“'C]acetate labeling experiments, three four-week-old embryos were

117

incubated at 25°C with gentle shaking either in light (300 umols/s/mz) or in the dark
in zoom of 0.1 mM MES-NaOH (pH 5.0) containing 5 mM sodium [1-“C]acetate
(1 .74qu). Assays were terminated by removing the incubation buffer, washing the
embryos twice with water, and initiating the lipid extraction. For experiments with
the herbicide haloxyfop the embryos were pretreated in 200pl of 0.1 mM MES-
NaOH ( pH 5.0) with addition of different concentration of herbicide for 30 minutes
before the addition of 2 mM [1-“Cjacetate substrate.
Lipid Analysis

Lipids were extracted from the embryos according to the method of Bligh and
Dyer (1959). Radioactivity in lipids at each time point was quantiﬁed by liquid
scintillation counting. Lipid classes were separated by TLC (20 x 20 cm K6 silica,
60A plates, Whatman) to heights OM and 12 cm in chloroformzmethanolzacetic acid
(75:25:8, v/v), allowing the plates to air dry between developments. The TLC plates
were subsequently developed to 20 cm in hexanezdiethyl etherzacetic acid (60:40:1,
v/v). Radioactivity of the separated lipid classes on the plates was assayed with an
Instant lmager (Packard Instrument Company). Labeled triacylglycerol and
phosphatidylcholine bands were eluted from the silica gel by elution with
chloroformzmethanol (1 :2, v/v). For transmethylation of total lipids or lipid classes,
the lipids were heated at 90°C for 45 minutes in 0.3 ml of toluene and 1 ml of 10%
boron trichloridelmethanol (Sigma). The recovered [“0] fatty acid methyl esters
were separated by argentation TLC (Morris et al.,1967). Argentation plates (15%

silver nitrate) were developed sequentially at -20°C to heights of 10, 15, and 20 cm

118

in toluene. Separated ["0] fatty acid methyl esters were located and quantiﬁed by
Instant Imager. The oleate, eicosenoate and erucate bands were scraped into test
tubes, and recovered by elution with 6 ml of hexanezethyl ether (2:1, v/v). In order
to characterize the distribution of label in 18:1, 20:1, and 22:1, these fatty acid
methyl esters were cleaved at the position of the double bond by perrnanganate-
periodate oxidation (Christie, 1982). The resulting nonanoic acid (09A) and 1,0)-
nonane-, undecane- or tridecane-dioic monomethyl ester fragments (CQAE, C1 1AE
and C13AE respectively) were separated by silica TLC in hexanezethyl etherzacetic
acid (90:10:1) and quantiﬁed using the Instant Imager. The relative amount of [1-
1"Clacetate incorporated into the de novo portion (CS-022) of 22:1 is calculated
simply as 09A+1.25C9A or 2.25C9A.

In order to measure the in vivo fatty acid accumulation rate, twenty embryos
were taken from plants at different stages from 20 DAF (days after ﬂowering) to 43
DAF. At the initiation of lipid extraction, 500 pg 1,2-dipentadecanoyl-sn-glycero-3-
phosphocholine (Sigma) was added to each sample as internal standard. Lipid
extraction was as described above. Fatty acid methyl esters from total lipids at

different DAF were separated and quantiﬁed by GC analysis.

Results
Accumulation of fatty acids during B. rapa L. seed development.
As shown in Figure 4.1, developing B. rapa seeds accumulate fatty acids which are

primarily derived from oleic acid. The modiﬁcations of oleic fall into two mutually

119

exclusive types: further desaturation and further chain-elongation. Elongation is the
more prevalent modiﬁcation, as erucic acid is the most abundant fatty acid,
reaching a level of 56 mol % in seeds at 43 DAF, whereas 18:2 plus 18:3 total
<20% at the same stage. Palmitate and stearate with a content of <3 % in mature
seeds are the only signiﬁcant fatty acids which do not derive from oleic acid. The
studies described below were conducted on seeds at 28 DAF when total fatty acid
and erucic acid were accumulating at maximum rates. At this stage, the rate of total
18:1 production, including its elongated and desaturated derivatives, was 16 nmol
hr "embryo“.

Light alters relative proportions of oleic and erucic acid synthesized

To monitor how light inﬂuences the accumulation of oleate and erucate, and
triacylglycerol versus phosphatidylcholine, incubations with [“Cjacetate were
carried out either in light or dark. As shown in Figure 42, incubation of embryos
in the light increased radioactivity in oleate approximately two-fold, whereas the
radioactivity in erucate was only fractionally higher in light incubations. The
comparatively small impact of light on radioactivity of erucate, which is highly
labeled at the carboxyl end, suggests that the homodimeric acetyl-CoA carboxylase
and fatty acid elongation are not strongly inﬂuenced by light, whereas the 50%
reduction of radioactivity in oleate in dark versus light implies that a major site of
light regulation is located in the plastids which are responsible for the de novo fatty
acid synthesis. An alternative interpretation of these experiments is that reduction

in label of oleate in the dark reﬂects changes in the endogenous pools of acetate.

120

7000 - t
5 - Total FAs l
' Total 18:1

6000. ° derivatives '
l A 5000. I
0
Z‘
l n l ' i
E
l s
l '5 22:1 l
E
I 5
‘ a
E
O
N
l s
(I
. IL
I
18:1
l 18:2
‘ 18:3

 

 

20:1 1

 

 

Days after ﬂowering {

Figure 4.1. Accumulation of fatty acids during embryo development of B. rapa.
Lipids were extracted from 20 pooled embryos at times indicated and fatty acid
methyl-esters were separated and quantiﬁed by GC. The total 18:1 derivatives
were obtained by adding 18:1, 18:2, 18:3, 20:1, and 22:1 together. The
accumulation rate rate of 18:1 derivatives (16 nmol/hr/embryo) was calculated for

the embryos at 28 DAF when labeling experiments were conducted.

121

This explanation was ruled out because very similar light dependence was also
observed when ["stucrose or [3H]-water were used as the precursors for fatty acid
synthesis (not shown). These observations suggest that an endogenous pool of
oleate might contribute to the synthesis of cis-11-eicosenoate and erucate. If the
newly synthesized oleate was the predominant source of oleoyl moeities for chain
elongation and the synthesis of oleate was reduced by half in dark, we would
expect that the labeling of erucate would also be reduced similarly. In fact, very
little reduction of erucate labeling occurs in the dark.

Analysis of the distribution of label from [1-“C]acetate in long-chain fatty
acids with time.

Erucic acid is synthesized from oleate by addition of two two-carbon units
from two molecules of malonyI-CoA. When exogenous [“C]acetate is used as
substrate for the biosynthesis of erucate the two-carbon units from the chain
elongation of oleate have a higher speciﬁc activity than the two-carbon units of the
methyl terminal 18 carbons, which are derived from de novo fatty acid synthesis of
oleate. The same applies to cis-11-eicosenate, except that only one elongation
cycle occurs from oleate. This differential labeling of C20 and longer fatty acids
from exogenous acetate was previously documented for four different oilseed
species: Brassica napus (Downey et al., 1964), Simmondsia chinensis (Ohlrogge
et al., 1978) Tropaeolum majus (Pollard et al., 1980) and Limnanthes alba (Pollard
et al., 1980) and has been interpreted as reﬂecting different pools of acetate

supplying the de novo fatty acid synthesis and the chain elongation reaction. What

122

   

 

mdoactivity (CPM In 221
, mammrm in 13:1

10D
0 I . - ~ EE,.“ . 1"l"“ ' OI —4--4-a-~ ~—+~--+—-~--+——--l»-
ommso120150180 omaosomtsom
' I "mum (Hill) . ' m" ,al....lvlwfﬂzlna(ﬂn) . .
D

   

 

Moaclivitﬂa-‘M in m;
magawvrtyrm In PC

 

0306090120150180 obsoeoeomiwwo
lmmﬁmﬂmmln) vrmmmammny.

Figure 4.2. Effects of light on the synthesis of 22:1 (A) 18:1 (B), TAG (C), and PC

(D). 0, light; 0, dark. Radioactivity is expressed per embryo.

123

has not been examined, however, is the time dependency of this differential
labeling. As we will demonstrate below, this time dependency provides information
on the pool of oleate supplying chain elongation.

Oxidative cleavage at the double bond of monounsaturated fatty acid methyl
esters allows determination of the relative speciﬁc activity of 1“C in the acid and the
acid-ester fragments of the acyl chain since both of the fragments can be
quantitatively recovered. Table 4.1 presents the ratio of radioactivity in each
fragment for isolated oleate, cis-11-eicosenoate and erucate, when four-week-old
developing Brassica embryos were incubated with [1-“C]acetate. The ratios were
measured after various incubation times over a one hour period. It is expected that
oleate will be uniformly labeled, and if the substrate is [1 -“C]acetate, the theoretical
distribution between the C9 acid-ester and the 09 acid fragments will be 1.25 (5/4).
The measured ratios fell in the range of 1.24-1.30, with an average value of 1.259
$0.016. As an additional control, we used permanganate-periodate cleavage of
commercial [U-“C]oleate. The theoretical ratios of 1.0 (C9AE/C9A) in oleate was
obtained exactly (1.0010002). For further conﬁrmation that this method is suitable
for quantitative analysis, the radioactivity in fatty acids was measured before the
oxidation, and was measured again after extracting cleavage products from the
reaction solution. Loss of radioactivity was negligible throughout the procedure.
All these controls indicate the high degree of accuracy and reproducibility of the
technique which is essential, since the variations in the ratios with time are the key

experimental data.

124

Table 4-1. The ratio of radiolabel in oxidative cleavage fragments of [“leabeled

fatty acids from incubation of B. rapa embryos with [1-“Cjacetate.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

5 min 10 min 20 min 30 min 40 min 50 min 60 min
Light:C13/C9 of 22:1 9.81'I 9.68 8.65 8.29 7.92 7.42 6.42
011/09 of 20;1 5.58 4.77 4.52 4.39 4.38 4.14 3.63
09/09 of 13;1 1.28 1.25 1.27 1.26 1.24 1.25 1.25
Dark: C13/C9 of 22:1 14.42 13.23 10.87 9.73 9.51 9.38 8.93
C11/CQ of 20:1 8.01 7.21 5.48 5.44 5.34 5.28 5.02
cg/cg of 1&1 1.30 1.27 1.26 1.25 1.25 1.26 1.24

 

8. Ratios of radioactivity in the 13C, 1 1C, and QC carbon fragments was determined

after permanganate-periodate oxidation at the double bond and isolation of the

cleavage products.

125

 

Table 4.1 presents the direct measurements of the ratio of [“C]Iabel in acid
and ester fragments. The results indicate that the [“Clratio in the oxidative
cleavage fragments of oleate remained constant with time and was close to the
expected value. In contrast, the ratio of 1"C in the C13AE/C9A fragments of erucate
and the C11AE/C9A fragments of eicosenoate both decreased with time in both
light and dark incubated embryos. The experiment was repeated ten times, and
although the absolute values of the acid to acid-ester ratios at each time point
showed some degree of variation between experiments, the trend was always the
same. Also noted in Table l and consistent with other experiments described below
was the observation that the acid-ester to acid ratios for erucate and eicosenoate
were always higher in dark incubated embryos than in light at any given time point.
Thus, in the dark the relative speciﬁc activity of C2 units used for elongation when
compared to CZ units derived from de novo fatty acid synthesis was increased by
a factor of 1.5-1.6.

It is also instructive to consider these labeling data in terms of the proportion
of total plastid-produced [“C]oleate units which appear in erucic acid. As
calculated from the fatty acid compositions shown in ﬁgure 4.1, over 55 % of 18:1
fatty acids synthesized by 28 DAF developing seeds are elongated to erucic acid.
Determination of the 1"C in the oxidative cleavage products of 22:1 allows
calculation of the 1"C content of the de novo synthesized 18 carbons. Comparison
of this value to the total 18:1 radioactivity accumulated in the incubation (18:1 plus

18 carbon portion of 20:1 and 22:1) gave the values plotted in Figure 4.3A. After

126

5 min incubation in the light, only 21% of the total 18:1 produced in the incubation
appears in 22:1 and this value increases to 35% by 60 min. Thus, there is a
substantial lag in the appearance of 1"C in the 18:1 portion of 22:1. In dark
incubations, a higher proportion of [“C]18:1 initially appears in erucic, but the
increase with time is similar. The labeling of the 20:1 de novo and elongation
carbons showed parallel patterns in light and dark (data not shown).

Two general explanations can be proposed for the change in labeling within
the very long chain fatty acids over time. The ﬁrst considers that there are different
sized acetate-accessible pools supplying malonyl-CoA for de novo fatty acid
synthesis and for chain elongation. The pool supplying chain elongation would be
relatively small, rapidly reaching a steady state contribution from exogenous
acetate. The pool supplying de novo fatty acid biosynthesis would be large and
equilibrate more slowly. A variant of this hypothesis is that the pools for elongation
and de novo fatty acid synthesis that utilize exogenous acetate directly are both
small, but that acetate can also be used to sustain de novo fatty acid synthesis via
an indirect metabolic pathway that slowly reaches steady state labeling. In all
cases, oleate synthesized at early time points would be of lower speciﬁc
radioactivity compared with later time points, and thus the acid-ester to acid ratio
would decrease with time. A corollary from this hypothesis is that the synthesis of
total [“Cloleate would show a lag at early time points with respect to elongation,
and with a time scale similar to the change in differential labeling within the very

long-chain fatty acids. Figure 438 shows the time-dependent accumulation of label

127

in total oleate derivatives (18:1 plus 18:1 portion of 20:1 and 22:1. 18:2 and 18:3
contributed less than 2% of the total label in these short term incubations and were
not included). The accumulation of ["C]oleate is essentially linear, with no lag
phase. Furthermore, the ratio of label in total oleate to the label in the elongation
portion of erucate plus cis-1 1-eicosenoate remains approximately constant (data not
shown), indicating that the ﬁrst general mechanism, acetate-accessible pool sizes,
is not responsible for the changes in [“C]ratios of de novo to elongation carbons
(Table 4.1)

The second general explanation for the changing ratios of label in the de
novo and elongation carbons considers that another oleate source besides the
newly synthesized [“Cjoleate was also used as substrate to synthesize the very
long-chain fatty acids. As shown in Figure 4.3A, the radioactivity accumulated in
the de novo (18:1) portion of erucate lagged signiﬁcantly to the radioactivity
accumulation of total oleate. This result supports the concept that there is an
endogenous source of oleate for the synthesis of erucate besides the newly
synthesized [“Cjoleate. If only newly synthesized oleate was directly used for
elongation, the [“C]ratios of C13AE/C9A and C11AE/C9A (Table 4.1), and the
percentage of total labeled oleate in erucate would remain constant with time. The
contribution of newly synthesized oleate is guaged by the extrapolation of the
curves of Figure 4.3A to zero time. At zero time the amount of the total labeled
oleate produced which contributes to erucic acid biosynthesis is only 20% in the

light (32% in the dark), whereas the theoretical maximum is 55%. Thus 36% (light)

128

 

45

l
E‘
".1 1 II-

N l

N

.5 4°! :8
o: I I;
F o

t l
2 I '1’:
3 35.! 1’:
" 3

._
66 I '.':
‘I— Q
9 I
1 30 . 5
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C 25 5
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° .2
n. 1:
20 1 r . r ‘ v t V ‘ ' r U r l "r l
0 10 20 30 40 50 60 O 10 20 30 40» 5O 60

Incubation time (min) Incubation time (min)

Figure 4.3. Labeling of fatty acids by [“Clacetate in developing B. rapa embryos.
[A]. The percentage of ["C]acetate incorporated into total 18:1 (18:1 plus derived
18:1 portion of 20:1 and 22:1) which appears in erucate is plotted vs. time. Light (0)
or dark (6) indicates that the incubation was conducted either under the light or in
dark. [B] Time course of 1"C accumulation into total 18:1 derivatives (18:1 plus 18:1
derived portion of 20:1 and 22:1) under the light. All the fatty acids were derived
from total lipid extracts, and the numbers in the ﬁgure represent radioactivity

calculated per embryo.

129

and 58% (dark) of the oleyl ﬂux through the chain elongation system to erucate is
directly utilized from newly synthesized (“C-labeled) oleate in the plastid, since at
zero time any large intermediate pools of oleate have yet to ﬁll.
Inhibition of the homodimeric ACCase blocks erucic acid production

The synthesis of erucic acid from oleate has been demonstrated in vitro to
require malonyl-CoA, and it has been assumed that this malonyI-CoA is produced
by the cytosolic homodimeric ACCase. This assumption has never been tested.
To evaluate this hypothesis in vivo we have used the herbicide haloxyfop, which
speciﬁcally inhibits the homodimeric acetyl-CoA carboxylase (Burton et al.,1987 and
1991) and examined how it inﬂuences the synthesis of erucic acid and other lipid
species. In the experiment shown in Figure 4.4 embryos were pre-treated with
different concentrations of haloxyfop for 30 min and then incubated with 2 mM
[“Clacetate under light for another hour. Incorporation of radioactivity into erucate
decreased with increased concentrations of haloxyfop. In contrast, radioactivity in
oleate increased with haloxyfop concentrations lower than 100 uM. At a
concentration of 50 pM haloxyfop, synthesis of erucate was inhibited by 70%, but
1"C accumulation in oleate increased almost two-fold. These results demonstrated
that the elongation of oleate to erucate is dependent on homodimeric ACCase to
supply malonyl-CoA, and that haloxyfop can inhibit the elongation without inhibition
of de novo synthesis of oleate. Also plotted in Figure 4.4 are the accumulations of
1"C in PC and TAG. In response to the addition of haloxyfop, radioactivities in

erucate and triacylglycerols were inhibited in parallel whereas the accumulation of

130

4500 . , , L ,. L. _z#_,_ 1400" I
-1260 2
1- 1090‘
4800"

.600

Radi°a°“""y 0“ “0 (DPM) in TAG and 22:1
Radioactivity of “C (PPMIJIn PCifend 15:1

 

 

l
I , . , . , , , , . . 1 0
o 20 4o 60 so 100 -120 140 ,160. 16.0.. 12.00. 220

 

Herbicide concentration (11M)

Figure 4.4. Effect of increasing concentrations of haloxyfop on fatty acid and lipid
synthesis. B. rapa embryos were pre-treated with indicated concentration of
herbicide for 30 min, then incubated for another hour under light with addition of
[“Clacetate. Radioactivity of lipid and fatty acid are quantiﬁed on a basis of DPM
per embryo.

131

radioactivities in PC and oleate showed parallel patterns of increase.

Discussion

The radioactivity incorporated into oleate of embryos incubated under light
was approximately two fold higher than that in the dark. These results extend the
observations of Browse and Slack (1985), who, in a study using isolated plastids
from linseed seeds (green embryos) and safﬂower seeds (white embryos)
concluded that linseed plastids are photosynthetically active and provide a source
of ATP and NAD(P)H for fatty acid synthesis. Similarly, Eastmond et al. (1996) and
Asokanthan etal. (1997) recently concluded that although photosynthesis is unlikely
to provide substantial net carbon for B. napus seed anabolism, light driven electron
transport may provide ATP and reducing equivalents for storage product synthesis.
Taken together with the data in Figure 4.2, these studies suggest that cofactor
supply and/or plastidyl acetyl-CoA carboxylase regulation by light rather than
carbon precursors may be a major limiting factor in the rate of fatty acid
accumulation in Brassica embryos. This hypothesis is consistent with the
observation that acetyl-ACP and malonyl-ACP pools are relatively high at all stages
of active fatty acid synthesis in developing spinach seeds or castor endosperm
(Post-Beittenmiller et al., 1991 and 1992a).

The strong inhibition of acetate incorporation into erucate by haloxyfop, a
known non-competitive inhibitor of the cytosolic homodimeric acetyl-CoA

carboxylase, conﬁrms the hypothesis that acetate that is utilized for chain

132

elongation is in the cytosolic compartment and distinct from acetate utilized for de
novo fatty acid synthesis. In all dicots so far examined the cytosolic acetyl-CoA
carboxylase is the homodimeric form, whereas the plastid acetyl-CoA carboxylase
is the heteromeric form (Konishi et al., 1994). However, a homodimeric form of
ACCase has also recently been reported in B. napus plastids (Schulte et al., 1997;
Roesler et al., 1997). Although the function of this ACCase is at this time unclear,
it appears to be less abundant than the heteromeric form. Furthermore, if this
plastid homomeric ACCase is as sensitive to haloxyfop as the cytosolic form, our
observation that 50 [M herbicide caused no inhibition of de novo FAS while strongly
inhibiting elongation suggests that the homomeric ACCase in the plastid has a
quantitatively minor role in de novo FAS.

As presented in Table 4.1 and Figure 4.3 we observed differential labeling
with time of the elongation and de novo carbons within the very long chain fatty
acids. We considered two explanations which could cause these changes. The
explanation whereby different sized acetate-accessible pools supplied malonyl-CoA
for de novo fatty acid synthesis and for chain elongation was ruled out from kinetic
and pool size considerations. This is not surprising if acetate is being utilized
directly for acetyl-CoA synthesis in both plastids and the cytosol. Data are not
available for oilseeds, but in leaf tissue the chloroplast pool of acetyl-CoA, the direct
substrate for the synthesis of malonyl-CoA, ranges from 10 to 20 pM, and it can be
equilibrated with exogenously supplied acetate within several seconds (Post-

Beittenmiller et al., 1992b; Roughan, 1997). The chloroplast pool of acetyl-CoA

133

dominates the total leaf acetyl-CoA pool, and, indeed total leaf CoA, so the cytosolic
pool of acetyl-CoA is expected to also be small. Extrapolating to seed tissue, even
if the cytosolic pool of acetyl-CoA is depleted at a rate of about one tenth of that of
the plastid pool, as calculated from the mass composition of the oil, it is expected
that the direct utilization of exogenous acetate by either pool will reach steady state
very quickly. As endogenous free acetate pools have not been measured in
developing seeds it is not possible to estimate the endogenous contribution of free
acetate to fatty acid synthesis in seeds. However, utilization of exogenous acetate
clearly reaches a steady state rate very quickly (Figure 4.38), within one minute.
AcyI-CoA and acyl-ACP pools might also contribute to a lag in labeling. In Brassica
embryos, and in developing seeds in general, there is limited information on acyl-
ACP levels and a dearth of information on acyl-CoA levels. We can make some
extrapolations from the situation in leaves and from the limited seed data. In
chloroplasts isolated from leaf tissues (Soll and Roughan, 1982; Roughan and
Nishida, 1990), it was estimated that acyl-ACP half lives were of the order of 10
seconds. In seeds, ACP levels of the order of 1 pg/gfw were noted (Hannapel and
Ohlrogge, 1988), while in spinach seeds and leaves up to 60% of the ACP is in the
free form. Similarly, acyl-CoA pools in Cuphea (Singh et al., 1986) and developing
B. napus (unpublished observations) indicate levels of <20 uM. The B. rapa seeds
in the present study accumulated about 16 nmoles fatty acid per hour per seed.
Using these numbers, the acyl-CoA and acyl-ACP pool turnover time is calculated

to be less than one minute. Clearly, accumulation of oleate in the acyl-thioester

134

pools cannot explain any lag in the labeling kinetics of erucate. This indicates that
the second general explanation, that another oleate source besides the newly
synthesized [“C]oleate is also used as substrate to synthesize the very long-chain
fatty acids, is the correct one.

Three distinct scenarios can be envisaged for the supply of oleate for chain
elongation to cis-11-eicosenoate and erucate in developing oilseeds. In the ﬁrst,
oleoyl moieties are exported from the plastid, activated to oleoyl-CoA, and can
immediately become substrates for elongation. In this scenario the newly-
synthesized oleoyl groups can be considered channeled directly to the cytosolic
elongation system. In the second model, the newly exported oleoyl-CoA is rapidly
equilibrated with the bulk cytosolic pool of oleoyl-CoA. The acyl exchange
mechanism ﬁrst reported by Stymne and Stobart (1984) can be envisaged to dilute
the [“Cjoleoyl-CoA pool with oleoyl groups from the sn-2 position of
phosphatidylcholine, while [“Cjoleate will enter the sn-2 position of
phosphatidylcholine and be diluted. The bulk pool of oleoyl-CoA is then available
for chain elongation. The third hypothesis is based on the observations of Hlousek-
Radojcic et al. (1995) and requires that newly synthesized [“Cjoleate acylate an
acceptor lipid, as yet unidentiﬁed, and then be transferred directly or indirectly from
this lipid to the elongase. An analysis of the kinetics of differential labeling within
the acyl chain of very long-chain fatty acids suggests that an endogenous oleate
pool contributes to the biosynthesis of erucate, though the labeling itself cannot be

used to distinguish between models two and three. Our results do not rule out

135

model one or other combinations of these models as a contributing pathway until
the details of the endogenous pool can be demonstrated and quantiﬁed. However,
extrapolating ﬁgure 4-3A to zero time indicates that only 20 % of total 18:1 synthesis
in light (or 32% in dark) is directly elongated to 22:1. Comparing this to the in vivo
conversion of >55% of 18:1 to 22:1 (Figure 4-1) suggests that at least half of the
oleate enters an intermediate pool before elongation to erucate.

The question follows as to what is the endogenous pool which can accept
newly synthesized oleate from the plastid and also provide oleate as substrate for
elongation. A logical approach to obtain information on such intermediate pools is
to perform pulse chase experiments. However, with intact B.rapa embryos we found
it was not possible to remove the [“Clacetate sequestered inside the embryos,
which continued to sustain fatty acid synthesis and hence confound interpretation.
In the absence of pulse-chase data, we decided to employ an indirect approach in
which we inhibited the elongation from 18:1 to 22:1 with haloxyfop without reducing
the de novo synthesis of oleate, and then monitored where the extra 18:1
accumulated. As shown in Figure 4.4, there is an increasing amount of oleate
accumulated in phosphatidylcholine when erucate synthesis is inhibited, indirectly
supporting the concept that the oleate esteriﬁed to phosphatidylcholine could be a
source of oleoyl moeities for the synthesis of erucate. Consistent with this
observation, Hlousek-Radojcic et al. (1995) found that when [”Cloleoyl-CoA was
incubated with B. napus oil bodies, over 50% of radioactivity was found in

phosphatidylcholine.

136

In a study of petroselinic acid biosynthesis in developing coriander and carrot
endosperm, Cahoon and Ohlrogge (1994) concluded that phosphatidylcholine was
an intermediate in the movement of petroselinic acid from its site of biosynthesis in
the plastid into triacylglycerol. Similar observations have recently been made for
the accumulation of 16:1“3 in developing seeds of Thunbergia (Shultz, Cahoon and
Ohlrogge, unpublished). Because neither of these unusual fatty acids are
synthesized or further modiﬁed on PC, a rationale fortheir movement through a PC
pool before incorporation into TAG is not immediately obvious. The present study
on erucic acid biosynthesis adds another example where PC may be an
“intermediate” in the ﬂux of fatty acids into TAG, even though no metabolism of the
fatty acid may be directly associated with the PC. The observation of a large ﬂux
of fatty acids through PC without modiﬁcation in three diverse oilseed species may
simply indicate that acyl exchange between the acyl-CoA pool and PC is very rapid.
The low accumulation of unusual fatty acids in PC may further reﬂect speciﬁc
mechanisms for their removal (e.g. phospholipases). However, it is interesting to
speculate that PC may play some more general role in TAG assembly, perhaps as
a carrier of acyl chains toward a subcellular site of TAG assembly. This role might
be analogous to the major ﬂux of acyl chains through PC in leaves followed by their
movement from the ER to the chloroplast.

Because of its commercial value, several attempts have been made to
increase erucic acid content in transgenic plants. However, the factors which limit

erucic acid content in Cruciferae species are still largely unknown. Expression of

137

an sn-2 acyltransferase from Limnanthes in transgenic B. napus led to accumulation
of erucic in the sn-2 position, but no increase in total erucate content of the oil
(Lassner et al., 1995). Similarly, over-expression of elongases has resulted in
increased chain-length of VLCFAs but not an increase in mole % VLCFA (Lassner
et al., 1996). The recent report that a mutated yeast SLC1 gene expressed in
Arabidopsis or B. napus gave increased erucic levels (Zou et al., 1997) is at this
time difﬁcult to interpret in light of the results with the Limnanthes enzyme.
Reduction of 18:1 desaturation by mutation of the oleoyl-desaturase might be
expected to increase 18:1 availability for elongation. However, in a mutant of
Arabidopsis which is deﬁcient in desaturation of 18:1 (Lemieux et al., 1990) 18:2
and 18:3 content of seeds decreased from 53% to 8.7%, 18:1 increased from 15.4%
to 53.5%, but 20:1 and 22:1 just slightly increased from 20.2% to 26.7%. Likewise,
elimination of the elongation of 18:1 might be expected to increase 18:1 availability
for desatu ration. However, neither low erucic lines of Brassica (Daun, 1983) nor the
fae1 mutant of Arabidopsis; (Kunst et al., 1992) exhibit corresponding increases in
18:1 desaturation. Although several interpretations are possible, one hypothesis
consistent with these results is that the pathways for 18:1 elongation and
desaturation may draw on different pools of 18:1. In summary, our results together
with those of Hlousek-Radojcic et al. (1995) and those cited above suggest that the
pathway for erucic acid biosynthesis may be more complex than originally
envisaged. Furthermore, the ﬂux of oleate through distinct intermediate lipid pools

prior to elongation or desaturation may be one factor which limits the availability of

138

 

oleate for elongation.

139

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Pollard MR, Stumpf PK (1980b) Biosynthesis of C20 and C22 fatty acid by
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of the Arabidopsis homomeric acetyl-CoA carboxylase to plastids of
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acetyl-CoA carboxylase from Brassica napus is encoded by a multi-gene

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Singh SS, Nee TY, Pollard MR (1986) Acetate and mevalonate labeling studies
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Soll J, Roughan PG (1982) Acyl-acyl carrier protein pool sizes during steady-state
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Stymne S, Stobart AK (1984) The biosynthesis of triacylglycerols in microsomal
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Cell 9: 909-923.

143

 

 

CHAPTER 53
Supply of Fatty Acid Is One Limiting Factor in the Accumulation of
Triacylgycerol in Developing Embryos
Abstract

The metabolic factors which determine oil yield in seeds are still not well
understood. To begin to examine limits on triacylglycerol (TAG) production,
developing Cuphea Ianceolata, Ulmus carpinifolia and Ulmus parvifolia embryos
were incubated with factors whose availability might limit oil accumulation. The
addition of glycerol or sucrose did not signiﬁcantly inﬂuence the rate of TAG
synthesis. However, the rate of 1"C-TAG synthesis, upon addition of 2.1 mM 1“C-
decanoic acid (10:0), was approximately 4 times higher than the in vivo rate of TAG
accumulation in Cuphea, and two times higher than the respective in vivo rate in
Ulmus. In Cuphea embryos, the highest rate of 1“C-TAG synthesis (14.3
nmols/hr/embryo) was achieved with addition of 3.6 mM of 10:0. 1“C-10:0 was
incorporated equally well in all three acyl positions of TAG. The results suggest
that both Cuphea and Ulmus embryos have sufﬁcient acyltransferase activities and
glycerol-3-phosphate levels to support rates of TAG synthesis in excess of those
found in vivo. Consequently, the amount of TAG synthesized in these oilseeds may

be in part determined by the amount of fatty acid produced in plastids.

 

3Material in this chapter was published previously [ Bao X and Ohlrogge J (1999) Plant Physiol.
120: 1057-1062].

144

Introduction

In plants, the biosynthesis of storage triacylglycerol (TAG) occurs at high
levels primarily in the seeds, but there is a wide range in the levels of TAG which
accumulate in different plant species. For example, seeds of species of Zea,
Hordeum or Pisum usually contain less than 5-10 % TAG by dry weight whereas
many other species such as castor accumulate over 50% TAG in seeds. The
regulatory or metabolic factors which inﬂuence this very wide range of oil
accumulation in seeds are currently unknown (Ohlrogge and Jaworski, 1997).
Considering that over 30 reactions are required to convert acetyl-CoA to TAG, there
could be many steps or genes which control the yield of end product TAG. In order
to begin to dissect possible limiting factor(s) in the pathway of TAG biosynthesis,
it is useful to conceptually divide the pathway into two parts: 1) the production of
acyl chains which occurs in plastids, and 2) the utilization of acyl chains for
glycerolipid synthesis in the ER and oilbody. In the second part, the only unique
enzyme for TAG synthesis is diacylglycerol acyltransferase (DAGAT), which is
responsible for the acylation of 1,2-diacylglycerol at the sn-3 position (Roughan et
al., 1982; and Stymne et al., 1987). Since diacylglycerol acyltransferase locates at
the branch point that channels diacylglycerol to TAG synthesis, some reports have
suggested that DAGAT may be one rate limiting enzyme for the accumulation of
TAG (Grifﬁths et al., 1988 and 1991; lchihara et al., 1988; Perry of al., 1993a and
1993b). In this study we asked whether the supply of fatty acid can inﬂuence the

amount of TAG produced in oilseeds. If the supply of fatty acid is a limiting factor

145

for TAG biosynthesis, then providing exogenous fatty acid to the developing
embryos should increase the rate of TAG production. Unfortunately, long-chain
fatty acids (LCFA) have very low solubility in aqueous solution, and so addition of
LCFA at concentrations sufﬁcient to increase rates of lipid synthesis is very difﬁcult.
However, embryos of Cuphea Ianceolata, Ulmus carpinifolia, and Ulmus parvifolia
contain high levels of decanoic acid in their TAG (80%, 63%, and 71%,
respectively). Decanoic acid is easily dissolved in water at mM concentrations, and
therefore, these species provided a convenient model system to test the inﬂuence
of fatty acid supply on TAG accumulation. Such in vitro model experiments may
provide a useful guide toward the selection of targets for future metabolic

engineering in transgenic plants.

Materials And Methods

Plant materials and Chemicals

Cuphea Ianceolata plants were grown in Beal garden on the Michigan State
University campus. Cuphea plants typically begin to ﬂower in mid-July and our
experiments were performed in the month of August. Flowers were hand-pollinated
and seeds were harvested at various stages of development. After removal of the
seed coat, the resulting embryos were used immediately for labeling experiments
or stored at -20°C for later lipid analysis. Embryos were also collected from two
species of elm trees (Ulmus carpinifolia and Ulmus parvifolia) growing on the

Michigan State University campus. Elm trees begin ﬂowering in early May and we

146

collected embryos when they were big enough to dissect, until embryo maturity.
The embryos were removed from seed coat and used for labeling experiments
immediately or stored at -20°C for later lipid analysis.

[1-“C]Octanoic acid (SSpCi/mmol), [1-“C]decanoic acid (550Ci/mmol), and
[1-“C]oleic acid (55pCi/mmol) were purchased from American Radiolabeled
Chemicals, lnc.(11624 Bowling Green Drive, St. Louis, MO 63146). Tritridecanoin
(C1320), L-dipentadecanoyl (C15:0) or-phosphatidylcholine, and lipase (from
Rhizopus arrhizus) were obtained from Sigma.

Lipid analysis

Lipids were extracted from 20 embryos at each developmental stage
according to the method of Bligh and Dyer (1959). Priorto extraction, tritridecanoin
(C1320) and L-dipentadecanoyl (C15:0) a-phosphatidylcholine were added to each
sample as internal standards for GC analysis. Triacylglycerol (TAG) was separated
from polar lipids by TLC (20XZ0 cm K6 silica, 60A plates, Whatman) in
hexanezdiethyl etherzacetic acid (70:30:1, vlv). TAG bands were eluted from the
silica gel with chloroformzmethanol (1:2, vlv). Fatty acid methyl esters from TAG
were prepared by heating lipids at 90°C for 45 min in 0.3 ml of toluene and 1 ml of
10% (vlv) boron trichloridelmethanol (Sigma). The resulting fatty acid methyl esters
were separated and quantiﬁed by GC analysis.

Feeding developing embryos with exogenous fatty acid
Ten pairs of Cuphea cotyledons (10 days after ﬂowing) were cut in half and

incubated at 28°C with gentle shaking in 200 pl of 0.1 M phosphate (pH7.2)

147

containing 2.1 mM [1-“C]decanoic acid with the presence or the absence of 0.125
mM of glycerol. The incubation buffer was changed once after one hour incubation.
Assays were terminated by removing the incubation buffer, washing the embryos
twice with water, and initiating lipid extraction. Triacylglycerol was separated by
TLC using a solvent system composed of hexanezdiethyl etherzacetic acid (70:30:1,
v/v). Labeled TAG was quantiﬁed with both Instant Imager (Packard Instrument
Company) and liquid scintillation counting. In some experiments, other factors
which may inﬂuence TAG synthesis, such as, exogenous fatty acid concentration
(2.1 to 80.1 mM), glycerol (with/without 0.125mM), sucrose (0 to 200 mM), and pH
(6 to 8), were tested. Only one factor was changed in each different treatment. The
position of exogenous fatty acid incorporated in TAG was determined using TAG
lipase from Rhizopus anhizus which cleaves fatty acid from sn-1 and sn-3 positions
of TAG, and then the radioactivity remaining in the sn-2-monoacylglycerol was
compared to free fatty acids released from the sn-1 and sn-3 of TAG by the action
of TAG lipase. Puriﬁed TAG was dissolved in 0.5 ml diethyl ether in 13 ml screw
cap glass tube. 1 ml of 0.1 M Tris-HCI (pH7.8) buffer containing 5 mM CaCl2 was
added to the tube, then 43,000 units lipase was added to the bottom of the tube,
bubbled in N2, and shaken for ten min at room temperature. Monoacylglycerol and
free fatty acid products were extracted and resolved by TLC in hexanezdiethyl
ether2acetic acid (35:70:1 .5, vlv). The radioactivity in the monoacylglycerol and free
fatty acid bands on the TLC plate was quantiﬁed using an Instant Imager.

In similar experiments, ten pairs of cotyledons from U. carpinifolia and U.

148

 

parvifolia embryos at the mid-stage of development were also used for feeding
experiments as described above, except incubation buffer was changed every half

houn

Results

Fatty acid deposition, composition, and in vivo rate of TAG accumulation
during embryo development

Mature C. Ianceolata seeds accumulate TAG in which decanoic acid is the
predominant fatty acid, reaching a level of 80 mol% (Bafor et al., 1990). Embryos
were large enough to isolate at 6 days after ﬂowering (DAF), and the seeds
reached maturity about 20 DAF. As shown in ﬁgure 5-1, TAG deposition in the
Cuphea embryos was linear from 8 to 12 DAF and during this period TAG
accumulated at the rate of 2.9 nmols/hr/embryo. This result was close to the 2.3
nmols/hr/embryo measured by Bafor et al. (1990). During this same period, the
relative amount of decanoic acid in TAG increased from 40% to 75% mol percent.
The fatty acid composition of TAG in Cuphea embryos at the mid-stage of TAG
accumulation (11 DAF) are similar to the results obtained by Bafor et al. (1990).

In U. carpinifolia and U. parvifolia, it is difﬁcult to tag ﬂowers on a daily basis,
because the elm trees are very tall. Therefore, embryos were collected only when
they reached a stage when lipid analysis was feasible. The ﬁrst collection was
designated as the zero time point, and latertlme points were recorded as days after

the ﬁrst collection (dafc). The rate of TAG accumulation of U. carpinifolia and U.

149

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150

 

 

content (9), and the right scale the percent of 10:0 in TAG (A)

 

 

 

 

 

 

__ #‘L ,_.ﬁ_l

Figure 5.1. TAG accumulation and percentage of decanoic acid in TAG in
developing embryos of C. Ianceolata, U. carpinifolia, and U. parvifolia. 20 embryos

at each stage were analyzed for TAG content. The left scale represents the TAG

parvifolia embryos was linear from 7 to 10 dafc, and the relative amount of decanoic
acid in TAG increased from 40% to 65% (Figure 5.1). The fatty acid composition
of TAG in both elm species at 10 dafc is given in Table 5.1. At this stage, medium-
chain fatty acids constitute 77% and 85% of the TAG fatty acids in U. carpinifolia
and U. parvifolia, respectively.
Rates of TAG synthesis by developing embryos with addition of exogenous
fatty acid

Bafor et al. (1990) observed that, when Cuphea developing embryos were
incubated with exogenous decanoic acid and glycerol, the rate of exogenous
decanoic acid incorporated into TAG was 33.9 nmol/hr/embryo. Assuming that
decanoic acid could be esteriﬁed to all the three position of glycerol, the rate of 1“C-
TAG synthesis was at least 1 1.3 nmols/hr/embryo which is four time higher than the
in vivo rate of TAG accumulation (2.9 nmols/hr/embryo). To further examine and
extend these results, we tested whether addition of exogenous fatty acid alone
could increase the rate of 1"C-TAG synthesis in Cuphea developing embryos, and
whether these results could be extended to other species. Ten pairs of cotyledons
of C. Ianceolata, U. carpinifolia, and U. parvifolia, harvested when TAG
accumulation was in the linear range, were incubated in buffer containing 2.1 mM
[1 -'4C]decanoic acid with or without glycerol. As shown in Table 5.2, in the absence
of glycerol, the average rates of 1"C-TAG synthesis were 12.5, 20.4, and 12.9
nmoIs/hr/embryo for C. Ianceolata, U. carpinifolia, and U. pan/ifolia, respectively.

Compared to their respective in vivo rates of TAG accumulation, which were 2.9 (C.

151

Ianceolata), 9.1 (U. carpinifolia), and 7.7 (U. parvifolia) nmols/hr/embryo, the rates
of 1“C-TAG synthesis were approximately 4 times higher for C. Ianceolata and 2
times for elm tree embryos upon the addition of exogenous decanoic. The addition
of 0.125 mM glycerol to the incubation solution did not strongly inﬂuence the rates
of 14C-TAG synthesis (Table 5.2). This suggests that there is an adequate
endogenous supply of glycerol for higher TAG synthesis in developing Cuphea and
Ulmus embryos. The level of 1“C-decanoic acid incorporated into TAG was used
to calculate the rate of 1“C-TAG synthesis in each species (Table 5.2). This
calculation did not include the contribution derived from the endogenous de novo
TAG synthesis from non-radioactive precursors, and so the values given in Table
II represent an underestimation of the total TAG synthesis from both exogenous and
endogenous fatty acids.

The TAG derived from Cuphea embryos incubated with [1 -“C] decanoic acid
for one hour was treated with TAG lipase (from Rhizopus arrhizus). The resulting
sn-2-monoacylglycerol and the non-esteriﬁed fatty acid (derived from sn-1 and sn-3
positions of TAG) were separated by TLC. Radioactivity was detected in both sn-2-
monoacylglycerol and free fatty acid fractions, indicating that labeled decanoic acid
was esteriﬁed to the sn-2 as well as to the sn-1 and sn-3 positions of the glycerol
backbone. The ratio of radioactivity from sn-2-monoacylglycerol to that of free fatty
acid was 1:2. This suggests that exogenous decanoic acid was equally distributed
among the three positions of TAG.

We recently observed that Iauric acid produced in transgenic B. napus can

152

Table 5.1. Fatty acid composition of TAG from U. carpinifolia and U. parvifolia embryos

at 10 days after ﬁrst collection.

 

Fatty acid distribution in TAG (mol%)1

 

TAG‘(nmol/embryo) 8:0 10:0 12:0 14:0 16:0 18:0 18:1 18:2 18:3
U. carpinifolia 2750 6.2 62. 4.0 3.8 6.7 0.8 7.4 6.6 1.8

 

 

 

 

 

 

 

 

 

 

 

 

 

 

U. parvifolia 3170 7.5 71 4.2 2.6 4.2 0.9 4.9 4.4 0.6

 

120 embryos were used for lipid extraction and isolation of TAG.

Table 5.2. Rate of 1“C-TAG synthesis upon addition 2.1 mM 14C-decanoic acid.

 

 

 

C. Ianceolata U. carpinifolia U. parvifolia
1“C-1020 (mM) 2.1 2.1 2.1 2.1 2.1 2.1
Glycerol (mM) 0 0.125 0 0.125 0 0.125

 

Rate of TAG

synthesis 12.5:t1.3 11.1119 20.4:22 21511.0 12.9120 13.8113

 

 

 

 

 

 

 

 

(nmol/hr/embryo)

 

Notes:

1. Ten pairs of cotyledons at mid-stage of development were incubated with 2.1 mM decanoic acid
in 0.2 ml of 0.1 M phosphate buffer (pH7.2) at 28 °C for 2 hours in the absence or presence of 0.125
mM glycerol. The incubation buffer was changed once at one hour for Cuphea, and every half hour
for Elm.

2. Rates of 1"C-TAG synthesis are calculated as nmols of 1"C-decanoic acid found in TAG (per hour)
divided by three.

3. Data represent the average of three independent experiments.

153

be subject to B-oxidation (Eccleston and Ohlrogge, 1998). If this had occurred in
the embryos supplemented with decanoic acid, a loss of lipid soluble 1“C would
occur and 1"C would be detected not only in decanoic, but also in other fatty acids
isolated after the incubations. Neither event was observed and furthermore
recoveries of added decanoic acid were at least 75-85%. Therefore B-oxidation
was not a major fate of the added decanoic acid.

Factors which inﬂuence the incorporation of exogenous fatty acid into TAG.

As shown in Table 5.2, addition of glycerol to the incubation buffer had no
signiﬁcant effect on the rate of 1“C-TAG synthesis. We also examined if other
factors such as exogenous fatty acid concentrations, sucrose (0 to 200 mM), and
pH may inﬂuence the rate of 1“C-TAG synthesis by Cuphea developing embryos.
Decanoic acid concentrations were varied over the range from 2.1 to 80 mM. As
shown in Figure 5.2A, the highest rate of 1“C-TAG synthesis (14.3 nmol/hr/embryo)
was obtained with addition of 3.6 mM of decanoic acid. Concentrations higher than
3.6 mM of decanoic acid apparently had a deleterious effect on embryos and
resulted in lower rates of TAG deposition.

Developing Cuphea embryos are non-photosynthetic tissue, so the ultimate
carbon source for TAG synthesis is derived from sucrose. To determine if sucrose
concentrations may limit TAG formation in these experiments, different
concentrations of sucrose (0 to 200 mM) were added to the basic incubation
solution. Despite minor variations among samples, the rates of 1“C-TAG synthesis

in all samples were close to 12 nmol/hr/embryo. This result suggests that carbon

154

a

l

l

l

l

l

l

l

l

l

I

44-41

A-..

a

 

Rte of TAG synthesis (nmollhrlembryo)
Me of TAGsyrtthesis (nmollhrlembryo)
0')

 

 

4
2..
0‘ _, 7 *1- ~74
6 65 7.2 75 8
Damoic acid ooncentrdion (mM) pl-lof incubdion buffer

4. i 4 f 4.r ELL 4’.LI#.I4 4 .4 4 j

 

Figure 5.2. Rate of 1“C-TAG synthesis under different conditions. (A) ten pairs of
cotyledon incubated in 0.2 ml of phosphate buffer (pH7.2) with concentration of 1"C-
decanoic acid as indicated. (B) ten pairs of cotyledon incubated in 2.1 mM 1‘0-
decanoic acid plus 0.2 ml of phosphate buffer, whose pH ranged from 6 to 8.

155

supply in the form of sucrose is not limiting TAG accumulation in these short term
experiments.

Developing Cuphea embryos were also incubated under a range of pH from
6.0 to 8.0. As shown in Figure 5.23, the optimal pH for TAG deposition was 7.2
whereas at pH6.5 or 7.5, the rate of 1"C-TAG synthesis decreased to half of that at
pH7.2.
Utilization of other exogenous fatty acids by Cuphea cotyledons for TAG
synthesis

Each ten pairs of cotyledons of developing Cuphea embryos were separately
supplied with 2.1 mM of [1-“C]octanoic acid, [1-“C]decanoic acid, or [1-"C] oleic
acid in 200 pl 0.1 M phosphate (pH7.2). 2.0 mM Triton X-100 was used to increase
solubility of the oleic acid. After one hour incubation, 5.8 nmols of octanoic acid,
37.5 nmols of decanoic acid, and a trace amount of oleic acid were incorporated
into TAG per embryo, respectively. One simple interpretation of this result is that
one or more of the acyltransferases of Cuphea display a strong selectivity in favor
of decanoic acid (Bafor etal., 1992; Vogel etal., 1996). However, Brassica napus
embryos were also incubated with oleic acid under the same conditions and the
incorporation of oleic acid into TAG was signiﬁcantly lower than the in vivo rate
(data not shown). So, for oleic acid, low aqueous solubility or transport into the

tissue may also prevent its rapid incorporation into TAG by embryos.

Discussion

156

TAG normally accumulates to a high level only in seeds but a metabolic
understanding of the tissue speciﬁcity of oil accumulation is not yet available. One
potential explanation is that DAGAT, which catalyzes the acylation of position 3 of
1,2-diacyl-sn-glycerol, is speciﬁcally expressed in seed. However, there are
several observations which argue against this view. For example, DAGAT activity
was found in spinach leaves (Martin et al.,1983) and was primarily associated with
chloroplast envelopes (Martin et al.,1984). Roughan et al. (1987) reported that
signiﬁcant amounts of TAG were synthesized when palmitic acid was applied to the
upper surface of expanding spinach leaves. The level of neutral lipids (mainly
TAG) increased at least 3 fold during protoplast isolation from Arabidopsis leaves
(Browse et al., 1988). Finally, ozone-fumigated spinach leaves produced high
proportions of TAG (Sakaki et al.,1990). These data together suggest that DAGAT
not only occurs in leaves, but also that leaves have the ability to synthesize TAG.

Although expressed in several tissues, higher expression of DAGAT during
seed development might represent one explanation for TAG accumulation in
oilseeds. lchihara et al. (1988) measured the speciﬁc activity of DAGAT from
safﬂower in vitro and found DAGAT activity was lower than expected. They
concluded that the DAGAT reaction may be rate-limiting. When developing
safﬂower and sunﬂower cotyledons were incubated with exogenous radiolabeled
fatty acid tracers, substantial amounts of labeled fatty acids were esteriﬁed to DAG
(Grifﬁths et al., 1988). Since DAG is the direct substrate of DAGAT, it was

suggested that DAGAT could be a rate-limiting step. Perry et al. (1993a, 1993b)

157

 

 

found that, when developing seeds of Brassica napus were incubated with [1-
1“Clacetate and [2-3nglycerol, very low accumulation of the Kennedy pathway
intermediates occurred apart from DAG. These results were also interpreted as
indicating that DAGAT is likely to exert signiﬁcant ﬂux control over TAG
accumulation. A similar conclusion was drawn by Grifﬁths et al. (1991) from studies
of TAG synthesis in cocoa. However, the accumulation of DAG might also be
explained as a shortage of acyl chain supply rather than ﬂux control at DAGAT.
Because both DAG and acyl-CoA are direct substrates of DAGAT, lack of one
substrate (acyl-CoA) can lead to the accumulation of the other (DAG) if the DAGAT
K,n for acyl-CoA is higher than the other acyltransferases. Thus, the accumulation
of DAG does not necessarily imply that DAGAT exerts ﬂux control for TAG
synthesis.

In order to begin to examine the limiting step(s) in TAG production for this
study we considered the pathway of TAG biosynthesis in two parts. The ﬁrst half
can be characterized as fatty acid production inside plastids; the second half can
be considered as the assembly of TAG in the ER or oilbodies (Cao et al., 1986;
Settlage et al., 1995). If the supply of fatty acid is a limiting factor for TAG
synthesis, addition of excess exogenous fatty acid should increase the rate of TAG
synthesis. As shown in Figure 5.1, TAG accumulated at the rate of 2.9, 9.07, and
7.65 nmols/hr/embryo in vivo for developing embryos of Cuphea Ianceolata, U.
carpinifolia, and U. pan/ifolia, respectively. With addition of exogenous decanoic

acid, their rate of 1"C-TAG synthesis was two to four fold higher than the in vivo

158

 

 

accumulation rate. This result clearly indicates that the supply of fatty acid can be
one limiting factor for TAG accumulation. In agreement with these observations in
seeds, in Chlamydomonas, addition of exogenous PC Iiposomes to cultures caused
10-fold increases in TAG accumulation (Grenier et al., 1991). Thus, it appears that
the capacity of these systems for TAG accumulation is greater than actually used
and that fatty acid supply, rather than the utilization enzymes may limit TAG
accumulation in Cuphea, elm trees, and Chlamydomonas. In addition, we found
that sucrose and glycerol had no signiﬁcant inﬂuence on the rate of 1"C-TAG
synthesis in Cuphea. This implies that both the endogenous carbon source and
glycerol backbone are in excess and not limiting 1“C-TAG synthesis during these
incubations. However, it is important to emphasize that such short-term incubations
may not reﬂect factors which control overall long-term accumulation of storage oils.
For example, over the time scale of seed development, many other factors such as
the ability of oilbodies to accommodate increased TAG might become limiting.

In this study, the exogenous decanoic acid was almost equally distributed
among the three positions of TAG from Cuphea. This result implied that not only
diacylglycerol acyltransferse, but also glycerol 3-phosphate acyltransferse and
lysophosphatidic acid acyltransferase activities could incorporate exogenous
decanoic acid at rates several fold above their in vivo activity with endogenous
substrates. Although our studies support the concept that increased fatty acid
supply can increase TAG accumulation, they do not rule out that other factors or

enzyme expression levels may have a similar effect. Flux through a metabolic

159

pathway can often be driven by either stronger source inputs and/or by stronger
sinks pulling on the pathway. The observations of Zou et al. (1997) that in B. napus
expression of a yeast acyltransferase can increase oil yields may represent an
example of sink driven increases in oil accumulation. Furthermore, we recently
found that transgenic B. napus seeds which express very high levels of medium
chain acyl-ACP thioesterase and produce high levels of Iauric acid induce the beta-
oxidation pathway to degrade some of the Iauric acid (Eccleston and Ohlrogge,
1998). Because oil yields are not reduced in these seeds, fatty acid synthesis
apparently increased to provide a constant oil yield.

In oilseeds, fatty acids esteriﬁed to TAG can be generally divided into two
groups. One (such as 18:3, 22:1, 18:1-HO) needs post-plastidial modiﬁcation, while
the other (10:0, 12:0, 18:1) does not. This work clearly shows that the supply of
fatty acids is one limiting factor for the rate of 1‘C-TAG synthesis in Cuphea and
Ulmus, which might be generalized representatives of the second category.
However, for the synthesis of TAG containing high level of post-plastidially modiﬁed
fatty acids, the involvement of phospholipids, desaturases, elongases, hydrolases,
etc. may be additional factors which limit TAG accumulation. In addition, the
observation that increased expression of acetyl-CoA carboxylase resulted in
increased oil content of high-erucic rapeseed (Roesler et al., 1997) suggests that
also in rapeseed, where fatty acids are modiﬁed by elongation, increased fatty acid

supply can increase TAG accumulation.

160

Literature Cited

Bafor M, Stymne S (1992) Substrate speciﬁcities of glycerol acylating, enzymes
from developing embryos of two Cuphea species. Phytochemistry 31 : 2973-
2976.

Bafor M, Jonson L, Stobart K, Stymne S (1990) Regulation of triacylglycerol
biosynthesis in embryos and microsomal preparation from the developing
seeds of Cuphea Ianceolata. Biochem. J. 272: 31-38.

Bligh EG, Dyer WJ (1959) A rapid method of total lipid extraction and puriﬁcation.
Can. J. Biochem. Physiol. 37: 911-917.

Browse J, Somerville CR, Slack CR (1988) Changes in lipid composition during
protoplast isolation. Plant Science. 56: 15-20.

Cao YZ, Huang AHC (1986) Diacylglycerol acyltransferase in maturing oil seeds
of maize and other species. Plant Physiology 82: 813-820.

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

Grenier G, Guyon D, Roche O, Tremolieres A (1991) Modiﬁcation of the
membrane fatty acid composition of Chlamydomonas reinhardtii cultured in
the presence of liposomes. Plant Physiol. Biochem. 29: 429440.

Grifﬁths G, Hamood JL (1991) The regulation of triacylglycerol biosynthesis in
cocoa (Theobroma cacoa) L. Planta 184: 279-284.

Grifﬁths G, Stymne S, Stobart AK (1988) The utilization of fatty-acid substrates
in triacylglycerol biosynthesis by tissue-slices of developing safﬂower
(Carthamus tincton'us L.) and sunﬂower (Helianthus annuus L.) cotyledons.
Planta 173: 309-316.

lchihara K, Takahashi T, Fujii S (1988) Diacylglycerol acyltransferse in maturing
safﬂower seeds: its inﬂuences on the fatty acid composition of triacylglycerol
and on the rate of triacylglycerol synthesis. Biochim Biophys Acta 958: 125-
129.

Martin BA, Wilson RF (1983) Subcellular location of triacylglycerol synthesis in

161

spinach leaves. Lipids 19: 117-121.

Martin BA, Wilson RF (1984) Properties of diacylglycerol acyltransferase from
spinach leaves. Lipids 18:1-6.

Ohlrogge JB, Jaworski JG (1997) Regulation of fatty acid synthesis. Annu. Rev.
Plant Physiol. Plant Mol. Biol. 48: 109-136.

Perry HJ, Harwood JW (1993a) Radiolabeling studies of acyl lipids in developing
seeds of Brassica napus: use of [1-“C] acetate precursor. Phytochemistry
33: 329-333.

Perry HJ, Harwood, JW (1993b) Use of [2-3H] glycerol precursor in radiolabeling
studies of acyl lipids in developing seeds of Brassica napus. Phytochemistry
34: 69-73.

Roesler, KR, Shintani, D, Savage, L, Boddupalli, S, Ohlrogge, JB (1997)
Targeting of the Arabidopsis homomeric acetyl-CoA carboxylase to Brassica
napus seed plastids. Plant Physiol 113: 75-81.

Roughan PG, Slack CR (1982) Cellular organization of glycerollipid metabolism.
Annu Rev Plant Physiol 33: 97-132.

Roughan PG, Thompson GA Jr, Cho SH (1987) Metabolism of exogenous long-
chain fatty acids by spinach leaves. Archiv Biochem Biophys. 259: 481-496.

Sakaki T, Saito K, Kawaguchi A, Kondo N, Yamada M (1990) Conversion of
monogalactosyldialglycerols to triacylglycerols in ozone-fumigated spinach
leaves. Plant Physiol. 94: 766-972.

Settlage SB, Wilson RF, Kwanyien P (1995) Localization of diacylglycerol
acyltransferase to oil body associated endoplasmic reticulum. Plant Physiol.
Biochem. 33: 399-407.

Stymne S, Stobart AK (1 987) Triacylglycerol biosynthesis. In PK Stumpf, EE Conn,
eds, The Biochemistry of Plants, Vol 9, Lipids: Structure and Function,
Academic Press, pp175-211.

Vogel G, Browse J (1996) Cholinephosphotransferase and diacylglycerol
acyltransferase. Substrate speciﬁcities at a key branch point in seed lipid
metabolism Plant Physiol. 110: 932-931.

Zou J T, Katavic V, Giblin E M, Barton D L, Mackenzie S L, Keller W A, Hu X,

162

Taylor DC (1997) Modiﬁcation of seed oil content and acyl composition in
the Brassicaceae by expression of a yeast sn-2 acyltransferase gene. Plant
Cell 9: 909-923.

 

 

163

CHAPTER 6

Conclusions and Future Research Perspective

Conclusions
Studies presented in this thesis have clariﬁed several questions along the pathway
from the origin of plastidial acetyl-CoA to the ﬁnal accumulation of triacylglycerol.
A method was developed to accurately measure the net rate of fatty acid synthesis,
in photosynthetic leaf tissues in short period time. Using the combination of 1:"CO2
labeling and GC/MS analysis, the rate of fatty acid synthesis for 3-week-old
Arabidopsis seedlings (whole aerial part of plant) was 2.3pmol C h’1 mg chlorophyll"
with 150 pmol s"m""1 Iumination. Employing isotope dilution strategy, the rate of
fatty acid synthesis in the dark was less than 5% of light rate. Both in pulse-chase
and continuous “002 labeling experiments, the maximum of lag phase was less
than two minutes, not the predicted 60 min lag iffree acetate is involved in fatty acid
synthesis and its concentration is 1 mM. Also in consistent with this observation,
radioactivity recovered in free acetate after one hour continuous labeling was below
detection limits or below a concentration of 0.02mM. Based on our kinetic analysis
and radioactivity in free acetate, we conclude that the bulk pool of acetate is not a
major substrate for fatty acid synthesis or the concentration of acetate is below 0.05
mM.

A biotin carboxylase genomic clone from Arabidopsis was isolated and its

promoter was partially characterized. The gene contains 16 exons and 15 introns;

164

the transit peptide is 71 amino acid long and there is one intron inside the transit
peptide. Northern blot and BC/GUS fusion constructs indicated that BC was
regulated at expression levels which correlated with the rate of fatty acid synthesis.
For example, the highest expression level was found in seeds. A positive and a
negative element were also identiﬁed in the promoter region.

Work described in chapter 4 let to the revision of the biosynthetic pathway
for erucic acid in the developing embryos of Brassica rapa. This revised pathway
indicates that newly synthesized 18:1 enters into an intermediate pool before it is
elongated to 20:1, then to 22:1. The results of manipulating the rate 18:1 synthesis
and blocking the elongation from 18:1 to 22:1 imply that PC might serve as the
intermediate pool of 18:1 for elongation reactions.

In chapter 5, the pathway of TAG synthesis was dissected into the supply
and the utilization of fatty acids. The addition of exogenous fatty acid resulted in
a 2 to 4 fold increase of the rate of TAG synthesis in Cuphea and elm developing
embryos. This result suggests that the amount of TAG synthesized in oilseeds may
be a response of a high rate of fatty acid synthesis. The acyltrasferases and
glycerol-3-phosphate levels seem to not limit the accumulation of TAG in the

embryos studied.

Future Research Perspective
1. The source of acetyl-CoA for fatty acid synthesis in plastids

Even though our results indicate that the bulk acetate pool is not involved in

165

fatty acid synthesis, the question of how the acetyl-CoA in plastids is generated
remains unclear. We still can not rule out the possibility that a very small pool of
acetate (less than 0.05mM) serves as intermediate for fatty acid synthesis. Due to
the short lag of carbon ﬂowing from CO2 to fatty acids, pool size analysis with
isotope labeling can not distinguish acetyl-CoA generated from different pathways.
In addition, because common precursors are shared by pathways that produce
acetyl-CoA, carbon position analysis can not provide informative data either. Wrth
these limitations, manipulating the key enzymes in each individual pathway is a
logical next approach to further clarify the relative contribution of plastidial acetyl-
CoA from several pathways. As shown in ﬁgure 1-2, pyruvate and acetate are the
two main candidates as immediate precursors to generate acetyl-CoA inside
chloroplast for fatty acid synthesis. The plastidial pyruvate dehydrogenase and
acetyl-CoA synthetase are responsible to convert pyruvate and acetate to acetyl-
CoA, respectively. It is ideal to inhibit each enzyme, and then measure the rate of
fatty acid synthesis to evaluate its relative contribution. Unfortunately, no inhibitors
that speciﬁcally inhibit acetyl-CoA synthetase or plastidial PDC have been reported.
Therefore, more emphasis should be placed on isolating and analyzing mutants of
acetyl-CoA synthetase and plastidial PDC. The drawback of this approach is that
mutations could be lethal if plastidial acetyl-CoA is generated solely by one
pathway, but it is still possible to obtain leaky mutants. If there are more than one
pathway contributing to acetyl-CoA in plastids, it should be easier to obtain mutants

for each enzyme, considering that different pathways might compensate for each

166

other. Alternatively, since the DNA sequences of acetyl-CoA synthetase and most
subunits of plastidial PDC are available, antisense of either or both enzymes should
provide more information about the origin of plastidial acetyl-CoA. Back at al.
(1999) reported that they obtained transgenic Arabidopsis containing a sense or
antisense construct of acetyl-CoA synthetase, pPDHor or pPDHB created from
Arabidopsis cDNA. Their research is still at the preliminary stage, information about
fatty acid synthesis and lipid contents of those transgenic plants is not available yet.
2. Analyzing promoters related to lipid synthesis

The BC promoter was only broadly analyzed in this study, and ﬁne deletions
are needed to further deﬁne regulatory factors. Promoter analysis primarily serves
two purposes; one is to identify cis elements responsible for certain expression
patterns and further to isolate the trans regulation factors; the other is to design a
promoter under whose control the target gene can be expressed as desired. Wrth
the onset of Arabidopsis genomic project and microarray technique, conceptual
changes are necessary to adapt new technologies, so it may not be a wise
approach to analyze single promoter. To establish the relationship between gene
expression level and enzymatic activity, several layers of regulation have to be put
into consideration. At a given condition, the transcript level, the stability of the
transcript, the efﬁciency of translation, the stability of the translated polypeptide,
and proper targeting are all contributing factors for the ﬁnal enzyme activity.
Therefore, high transcript level does not necessarily mean high enzymatic activity.

It is difﬁcult to obtain information about the stability of the transcript, the efﬁciency

167

of translation, the stability of the polypeptide for large number of genes, but it is now
relatively easy to get transcript levels and the ﬂux through corresponding enzymes.
The microarray chips containing all the known ESTs from Arabidopsis will soon be
available. As a result, the transcript levels of all the genes related to fatty acid and
TAG synthesis in roots, leaves, and seeds can be obtained very quickly; followed
with the comparison of gene expression in different tissues. We are interested in
seed speciﬁc promoters and cis and trans elements that are responsible for the up
regulation of certain genes in oilseeds. Future promoter analysis should be
conducted , at least in my opinion, in step wise fashion as outlined below. 1) The
transcript levels from seeds will be compared with that from roots or leaves. 2) The
genes will be classiﬁed into several categories according to changes of expression
patterns as following: speciﬁcally expressed, upregulated, unchanged, and down
regulated, and completely shut down in seeds. 3) Promoter regions of seed speciﬁc
genes will be retrieved from Arabidopsis genomic data bank, multiple comparison
of these promoters will be done to identify seed speciﬁc cis elements, and the
secondary structures and locations of these genes will be examined to determine
whether position effects also play a role for seed speciﬁc gene expression. 4) The
strong seed speciﬁc promoter will be made through combinations of different cis
elements from different seed speciﬁc expressed gene promoters. 5) For the
promoters that are completely shut down in seeds, the similar analysis will be
performed. In this group, we are interested in ﬁnding out whether these promoters

lack certain seed speciﬁc elements or have certain repression elements. 6) For the

168

up, unchanged, and down regulated genes in the seeds, we need calibrate
transcript leVel on per cell basis, and compare their corresponding enzyme activity
in terms of ﬂux. Only those gene promoters where transcript level changes are
correlated with their enzyme activity changes are deemed for further cis elements
analysis. Genes that their transcript levels are not correlated with their enzyme
activities, are candidates subject to other levels of regulation rather than on the
transcription level. In summary, with the complete genomic sequence of
Arabidopsis and microarray technique, above goals can be achieved in relatively
short period of time, for which we dare not dream two years ago.
3. Development of transgenic Brassica with high erucic acid content

As mentioned in chapter 4, the level of 50% erucic acid in high erucic
rapeseed oil is not sufﬁcient to compete well with alternative source of lubricant oils
from petrochemicals because of the high cost of puriﬁcation (Ohlrogge, 1994). In
order to replace petrochemicals, erucic acid content should be over 75% in the
rapeseed oil. In the TAG of high erucic Brassica, because most of sn-2 position is
occupied by 18 carbon fatty acids such as 18:1, 18:2, or 18:3, most research is
oriented toward replacing 18 carbon fatty acids with erucic acid at sn-2 position
hoping to achieve higher overall erucic acid content. Unusual fatty acids can be
esteriﬁed to sn-2 position of TAG. Decanoic acid composes 80% of the total fatty
of Cuphea and elm seeds, 85% petroselinic acid in coriander seeds, 80% A6
hexadecanoic acid in Thunbergia alata. This group of fatty acids are synthesized

in plastids and do not need any further modiﬁcation in the cytosol before esteriﬁed

169

to glycerol and they occupy about 30-50% of sn-2 position in TAGs. Castor oil
contains 85% of ricinoleic acid, 65% of vernolic acid in Euphorbia seed, 65%
linolenic in linseed oil. These fatty acids need further modiﬁcation before
incorporated into TAG, and the modiﬁcation reactions take place on the sn-2
position of PC. Then the unusual fatty acids would be presumably cleaved from the
sn-2 position and made available for TAG synthesis. This group of fatty acids also
constitute about 20-50% at sn-2 positions in their respective TAGs. The enzyme
responsible to remove fatty acid from sn-2 position of PC is still a debated issue.

Lysophosphatidylcholine: acyl-CoA acyltransferase (LPCAT) catalyzes the transfer
of a acyl group from acyl-CoA to the sn-2 lysophosphatidylcholine as a alternative
to form PC (Stymne and Stobart, 1984; lchihara et al., 1995). This reaction is fully
reversible and does not require ATP (Stymne ans Stobart, 1984). In safﬂower, the
activity of LPCAT was correlated with TAG synthesis during seed development
(lchihara et al., 1995). So it is suggested that LPCAT also catalyze the removal of
modiﬁed fatty acid from sn-2 position of PC. The pathway of erucic acid synthesis
is different from above two group of unusual fatty acids. As indicated in chapter 4,
the newly synthesized 18:1 ﬁrst enters the PC pool, then it is later removed from
PC, and ﬁnally elongated to erucic acid. We speculate that insufﬁcient LPCAT
activity might be the cause of high 18 carbon fatty acid content at sn-2 position of
TAGs in high erucic rapeseed. If there is not sufﬁcient LPCAT activity, the 18:1
esteriﬂed to the sn-2 position of PC most likely have two fates; one is being further

desaturated to 18:2 or 18:3; the other is along with PC being converted to DAG

170

directly and then to TAG. Both scenarios would result in higher 18 carbon fatty
acid content than othenrvise. Our hypothesis may explain why the failure to increase
erucic content by expression of an sn-2 acyltransferase from Limnanthes (Lassner
et al., 1995), over-expression of elongases (Lassner et al., 1996), and reduction of
18:1 desaturation (Lemieux et al., 1990). On the contrary, nasturtium (Tropaeolum
majus) seed oil contains as much as 80% erucic acid (Gustone et al., 1994; Luhs
and Friedt, 1994) . In order to increase erucic acid content in Brassica, we need to
ﬁrst carefully compare the elongation system of nasturtium with Brassica, especially
determining whether 18:1 also enters PC pool before being elongated. Second,
LPCAT activities from developing seeds of both species should be assayed. Third,
if LPCAT activity of nasturtium is signiﬁcantly higher than that of Brassica, we
should try to isolate the LPCAT gene from nasturtium (Tropaeolum majus) and
transfer this gene into high erucic acid Brassica lines. The bigest difﬁculty for
isolating lyso—PC acyltransferase from nasturtium is that there is not single gene
sequence from plant available yet. But considering the economic potential and
understanding the differences of the two elongation system, it is worth for further

invesﬁgaﬁon.

171

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Stymne S and Stobart AK (1984) Evidence for the reversibility of the acyl-
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Zou JT, Katavic V, Giblin EM, Barton DL, Mackeenzie SL, Keller WA, Hu X,
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173

   

               

 

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