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This is to certify that the
dissertation entitled

RE-EXAMINING THE INITIAL STEPS OF MEMBRANE
AND STORAGE LIPID ASSEMBLY IN PEA LEAVES

AND SOYBEAN EMBRYOS:
THE DOMINANT FLUX OF NEWLY SYNTHESIZED FATTY ACID
INCORPORATION INTO EXTRA-PLASTIDIC GLYCEROLIPIDS IS
THROUGH PHOSPHATIDYLCHOLINE ACYL EDITING

presented by
Philip David Bates

has been accepted towards fulfillment
of the requirements for the

Ph.D. degree in Biochemistry and Molecular
Biolgoy

 

 

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ABSTRACT

RE-EXAMINING THE INITIAL STEPS OF MEMBRANE AND STORAGE LIPID
ASSEMBLY IN PEA LEAVES AND SOYBEAN EMBRYOS:
THE DOMINANT FLUX OF NEWLY SYNTHESIZED FATTY ACID
INCORPORATION INTO EXTRA-PLASTIDIC GLYCEROLIPIDS IS THROUGH
PHOSPHATIDYLCHOLINE ACYL EDITING

By
Philip David Bates

In expanding pea leaves and developing soybean embryos over 95% of
fatty acids (FA) synthesized in the plastid are exported for assembly of eukaryotic
membrane and storage glycerolipids. It is often assumed that the major products
of plastid FA synthesis (18:1 and 16:0) are first acylated to glycerol-3-phosphate
(G3P) producing 1610/1821 and 18:1/18:1 initial molecular species of
phosphatidic acid (PA) which are then converted to phosphatidylcholine (PC), the
major eukaryotic phospholipid and site of acyl desaturation. However, by rapid
labeling of pea leaf and soybean embryo lipids with [“C]acetate and [14C]glycero|
we demonstrate that acyl editing is an integral component of eukaryotic
glycerolipid synthesis. First, no precursor-product relationship between
diacylglycerol (DAG) and PC [“C]acyl chains was observed at very early time
points. Second, analysis of PC molecular species and the positional localization
of nascent FA within PC at these early time points showed that the majority of
newly synthesized acyl groups (> 90% of the [“C]18:1 and [“C]saturated acyl

groups in pea, and >60% in soybeans) were incorporated into PC alongside a

 

previously synthesized unlabeled acyl group (18:2, 18:3 or 16:0). The nascent
FA occupied 62% of the sn-2 position in pea leaves and 86% in soybean
embryos. Third, [”Clglycerol labeling produced PC molecular species highly
enriched with 18:2, 18:3 and saturated acyl chains and not 18:1, the major
product of plastid fatty acid synthesis. And fourth, the kinetics of [“C]glycerol and
the positional analysis of newly synthesized FA incorporation into triacylglycerol
(TAG) of soybean embryos indicated that TAG is synthesized from a DAG
molecule generated by removal of the PC phosphocholine head group and not
de novo synthesized DAG. In conclusion, we propose that most newly
synthesized and plastid exported acyl groups are not immediately utilized for PA
synthesis, but instead are incorporated directly into PC through an acyl editing
mechanism that operates at both sn-1 and sn-2 positions. The acyl groups
removed by acyl—editing are largely used for the net synthesis of PC and other
glycerolipids through (G3P) acylation. Very little if any TAG is produced by three
consecutive acylations as in the traditional Kennedy pathway. These results
provide a more quantitative basis to demonstrate in vivo that FA flux in/out of PC
by acyl editing, and the inter-converstion of DAG with PC are dominate fluxes in

the assembly of eukaryotic membrane and storage glycerolipids in plants.

This dissertation is dedicated to my wife Cassandra. Her loving support,
strong encouragement, sacrifice of moving to Michigan, and ability to make me

fat and happy has made the perils of graduate school relatively easy to bear.

 

ACKNOWLEDGEMENTS

I would like to express my appreciation to my advisor John Ohlrogge. John
gave me the opportunity to work in his lab and entrusted me with interesting and
challenging research projects. His door was always open when I needed
assistance and he was always encouraging, even when the research was not
going well. I am especially grateful for his guidance that shifted my research
towards the subject of this dissertation. l have learned not only how to do good
science, but from his examples also how to be a good leader. Mike Pollard
provided me with enormous amounts of overall guidance and help with step by
step methods, scientific theory, and data analysis. Without his analytical mind
some of the major conclusions and finer details of many experiments would have
been passed over. Mike was my main advisor for many aspects of my research.
Plus, Mike kept me on my toes by providing a jovial remark to anything I said that
was less than very intelligent. I am grateful for all the members of the Ohlrogge
lab for their friendship and assistance, especially Doug Allen and Tim Durrett
who provided very valuable discussions and research assistance. Finally, I would
like to thank my guidance committee for their guidance during my graduate

research and their support to head into new directions when it was needed.

TABLE OF CONTENTS

 

 

LIST OF TABLES viii
LIST OF FIGURES ix
1 INTRODUCTION 1

 

 

1.1 Background overview 3
1.2 Fatty acid synthesis and desaturation 4
1.2.1 Acetyl-CoA carboxylase: Major FAS regulatory point ........................ 6
1.3 Esterification of fatty acids to glycerol-3—phosphate: de novo glycerolipid
synthesis. 9
1.3.1 The two pathways for de novo glycerolipid synthesis in plants ........ 10
1.3.2 Transfer of eukaryotic synthesized DAG to the plastid for glycolipid
14

 

 

 

 

 

 

synthesis
1.4 Long chain acyl-CoA pools for building glycerolipids ............................. 16
1.5 Synthesis of eukaryotic membrane lipid classes 18
1.6 TAG synthesis 19
1.6.1 Acyl transfer between lipids is required for a majority of TAG
synthesis 21
1.7 Suggestive evidence for acyl editing in plants 23
1.8 The in vivo labeling approach 26

2 INCORPORATION OF NEWLY-SYNTHESIZED FATTY ACIDS INTO
CYTOSOLIC GLYCEROLIPIDS IN PEA LEAVES OCCURS VIA ACYL

 

 

 

 

 

 

EDITING 30
2,1 Ahefmrt 31
2.2 Introduction 32
2.3 Experimental procedures 36
2.4 Results 41
2.5 Discussion 60
2.6 Footnotes 68

 

3 THE DOMINANT INITIAL FLUX OF FATTY ACIDS INTO SOYBEAN OIL IS

 

 

 

 

THROUGH PHOSPHATIDYLCHOLINE ACYL EDITING ............................. 70
3.1 Abstract 71
3.2 Introduction 72
3.3 Results 76
3.4 Discussion 101

vi

 

3.5 Conclusion 119

 

 

 

 

 

 

 

 

3.6 Materials and methods 120
3.7 Footnotes 126
4 CONCLUSION 127
4.1 Acyl editing conclusions 127
4.2 Additional significance of acyl editing and PC derived TAG synthesis.133
4.3 Future research on acyl editing and TAG synthesis pathways ............. 135
5 APPENDIX A: CHAPTER 2 SUPPLEMENT 143
6 APPENDIX B: CHAPTER 3 SUPPLEMENT 159
6.1 Supplement Figures for Chapter 3 Results 159
6.2 Supplement: Kinetic modeling of pool filling 170
6.3 Discussion of soybean TAG synthesis flux model Figure 3-10 ............ 178

7 APPENDIX C: PC ACYL EDITING IN ISOLATED PEA CHLOROPLASTS183

8 REFERENCES 187

vii

 

LIST OF TABLES

Table 3-1 Endogenous lipid pool sizes and fatty acid composition ..................... 78
Table 6-1 TAG Molecular Species Composition of Soybean Embryos ............. 167
Table 6-2 DAG Molecular Species Composition of Soybean Embryos ............. 168
Table 6-3 PC Molecular Species Composition of Soybean Embryos ................ 169
Table 6-4 TAG synthesis model: nascent FA flux value calculations ................ 179

viii

 

LIST OF FIGURES

Figure 1-1 Traditional textbook two pathway model of glycerolipid synthesis ..... 11
Figure 1-2 Possible enzymatic reactions leading to TAG synthesis in oilseeds ..20

Figure 2—1 Time course for [“C]acetate incorporation into the acyl groups of
glycerolipid classes by excised pea leaves. 43

Figure 2-2 [14C]Acyl composition of [14c1acetate labeled lipids ........................... 45

Figure 2-3 Time course for incorporation of label from [“C]glycerol into the
backbone and acyl groups of glycerolipids by excised pea leaves ...................... 47

Figure 2-4. Molecular species analyses of endogenous PC and PA from pea
leaves. 49

Figure 2-5 Analysis of endogenous PC molecular species and those labeled after
short time incubations of excised pea leaves with [“C]acetate or [“C]glycero| or
of seedlings with [“C]COz. 52

 

Figure 2-6 Stereochemical distribution of [“C]acyl chains in PC. ....................... 55

Figure 2-7 Comparison of unlabeled fatty acid compositions deduced from
[1 C]acetate and [14C]glycerol labeled PC molecular species to endogenous pea
leaf PC and lyso-PC compositions. 57

Figure 2-8 Distribution of lipid classes, labeled in their acyl chains, after
incubation of pea leaves with [1401002 or [“C]acetate. 59

 

Figure 2-9 Four possible models describing pathways and fluxes for the
incorporation of nascent FA, exported from the chloroplast immediately after de
novo FAS, into PC 62

 

Figure 3-1 Endogenous molecular species compositions of DAG, PC and TAG.
79

 

Figure 3-2 Time course for [‘4C]acetate incorporation into the acyl groups of the
major labeled soybean embryo lipids. 82

 

Figure 3-3 Fatty acid composition of [“Clacetate labeled lipids .......................... 85

Figure 3—4 Positional analysis of [”C]acetate labeled acyl groups in different
lipids. - 87

Figure 3-5 Molecular species compositions of [“C]acetate labeled PC and DAG.
89

Figure 3-6 Six min [“C]acetate labeled TAG molecular species ......................... 92

Figure 3-7 Time course for the incorporation of [14C]glycerol into the backbone

 

and acyl groups of the major labeled soybean embryo lipids. ............................. 96
Figure 3—8 Molecular species composition of [‘4C]glycerol backbone labeled PC,

DAG and TAG. 99
Figure 3-9 Representations of mathematical models for TAG synthesis ........... 107
Figure 3—10 Soybean TAG synthesis flux model. 111

 

Figure 5—1 Supplemental Figures S1A-S1M: Lipidomics of pea leaves and
acetate labeled pea leaves. 144

Figure 5-2 Estimates of single and dual acyl-labeled PC molecular species for
Model 3 (Figure 2-9)

Figure 6-1 [“C]Fatty acid composition of total [“Cjacetate labeled soybean
lipids. - 159

 

Figure 6-2 Alpha oxidation of [‘4C]acetate labeled saturated FA from TAG and
PC. 160

 

Figure 6-3 TLC image examples and flow chart for analysis of radiolabeled

 

 

 

 

glycerolipids. 161
Figure 6-4 [“Clacetate labeled PC molecular species time course. ................. 162
Figure 6-5 [“C]acetate labeled DAG molecular species time course. .............. 163
Figure 6-6 [“C]acetate labeled TAG molecular species time course ................ 164
Figure 6-7 [14C]glycerol labeled molecular species time course of PC and ONT-65
Figure 6-8 Fatty acid composition of [“C]glycerol labeled TAG, DAG and PC
molecular species. 166
Figure 6-9 Kinetic modeling of pool filling simulations 174
Figure 7-1 [”Cjacetate labeled PC and PG molecular species from isolated
chloroplasts. 184

 

 

1 INTRODUCTION

Fatty acid (FA) based lipids are essential to life and perform a wide variety
of functions. Living cells depend upon a hydrophobic bilayer of amphipathic
membrane lipids to separate the cell contents from the aqueous environment.
Eukaryotic cells accomplish complex metabolism by compartmentalization of
reactions within membrane bound organelles. Membrane lipids can be utilized as
substrates for metabolic signaling compounds [1-5]. Multi-cellular plants contain
fatty acid derived physical barriers against water, solutes, pathogens and solar
UV radiation in the form cutin, suberin, and wax [6-11]. Triacylglycerols (TAG)
contain three FA bound to a glycerol backbone and are the major form of
reduced carbon used for energy storage in plants and animals. In addition to the
biological importance of FA containing lipids within an individual organism, many
animals rely on plant TAG as a major source of nutrition.

The human population utilizes plant TAG for food, fuel and industrial
applications. In 2006, world harvest of plant oil was over 127 million tons [12].
Vegetable oil produced from soybeans represents the largest source of oil for
nutritional purposes at approximately 35 million tons produced in 2006 [13]. On
average the FA associated with common sources of vegetable oil are composed
of 16 and 18 carbon saturated FA (palmitic (16:0) and stearic acid (18:0),
respectively) and 18 carbon FA containing 1-3 double bonds (oleic (18:1), linoleic

(18:2) and linolenic acid (18:3), respectively). These common FA are useful for

 

cooking oils and high energy food sources but do not contain the most healthy,
oxidatively stable or most useful functional groups required in many highly
nutritional foods, fuels or industrial applications, respectively. Nutritionally
important unusual FA are the very long chain polyunsaturated fatty acids (PUFA)
such as docosahexaenoic acid (DHA) produced in the microalgae
Crypthecodinium cohnii [14] and usually consumed through fish. An example of
industrially important unusual FA is the hydroxyl containing ricinoleic acid
produced by the castor plant Ricinus communis [15, 16] which can be used in
lubricants and as a feedstock to polymers such as nylon.

Most plants cultivated for high yields of vegetable oil (e.g. soybeans and
rapeseed) were domesticated based on their value as food/feed and therefore do
not contain many of the industrially important unusual FA. Plant species that do
accumulate high levels of unusual FA (e.g. castor or tung) typically have
agronomical traits that make harvesting large yields difficult and expensive.
There is great interest in engineering unusual FA into some of the seasonal high
yielding crop species. Additionally, the rising human population and dwindling
petroleum reserves increases demand for renewable sources of chemical feed
stocks and fuel. Plant oils can be used as direct replacements for diesel fuel and
the wide variety of unusual FA could be the base for many industrial applications
[12, 13]. Even though there have been many attempts to engineer the FA
composition or the quantity of TAG in oilseeds, most cases have been met with
only limited success [17-27]. Most of the enzymes responsible for production of

the common, and many of the unusual, FA have been identified. However, not all

the factors that regulate FA synthesis or TAG accumulation are known and
questions remain about the sequence of reactions in the incorporation of FA into
membrane and storage lipids. A greater knowledge of the pathway of membrane
lipid and TAG synthesis will allow us to design better strategies for engineering
oilseeds in the future. This thesis will address the sequence of reactions that are

involved in incorporation of FA into extra-plastidic membrane and storage lipids.

1.1 Background overview

Assembly of most membrane or storage glycerolipids involves four basic
steps: 1) FA synthesis; 2) esterification of FA to a glycerol based backbone; 3)
addition of the sn-3 headgroup or FA; 4) desaturation of acyl chains and/or
transfer of modified acyl groups to other lipids. Although this process always
begins with fatty acid synthesis, the order of the remaining three steps may vary.
In plants these activities may be separated within a particular membrane system
or duplicated in different compartments, and thus may involve multiple pools of
substrates and trafficking between compartments. Additionally, some activities (in
particular desaturation) may be more active with a specific glycerolipid substrate
and thus require deacylation of the lipid after desaturation and further utilization
of the modified FA to make other lipids. An overview of the current state of
knowledge on plant membrane and storage glycerolipid synthesis is presented
as a background to the research in this dissertation and to set the stage for the
evidence that is presented in later chapters for changes in the current view of FA

flux through central plant lipid metabolism.

 

1.2 Fatty acid synthesis and desaturation

In plants the major site of de novo fatty acid synthesis (FAS) for
production of membrane and storage lipids is the plastid. This is significantly
different from animals and fungi where FAS is primarily cytosolic. The non-
cytosolic nature of plant FAS was first determined by subcellular-Iocalization of
acyl carrier protein (ACP), an essential component of FAS, to the chloroplasts of
spinach [28]. Mitochondria can also synthesize fatty acids, however it appears
that mitochondrial FAS is mostly for producing octanoate (8 C) for lipoic acid
synthesis [29]. Additionally, stable isotope labeling of Brassica napus embryos
demonstrated onlya plastidic labeling signature in total lipids [30], indicating that
any mitochondrial contribution to bulk lipids would be very minor compared to the
plastidic FAS. It is possible that mitochondrial FAS may play a more dominant
role in lipid synthesis in specific cell types such as epidermal cells involved in
wax and cutin synthesis where modified acyl chains are predominant [31, 32].
Since this work deals with membrane and storage lipids from leaves, total
seedings and developing embryos, FAS will refer to the major plastidic activity.

The first committed step in FAS is the production of malonyl-CoA by
acetyI-CoA carboxylase (ACCase) [33] in the plastid. The three carbon carboxylic
acid of malonyl-CoA is transferred to ACP and all further reactions take place
with the growing acyl chain on ACP. De novo synthesis of a fully reduced 16 or

18 carbon fatty acid takes place by the sequential condensation of the malonyl-

 

ACP with a growing fatty acyl—ACP. Release of CO; drives each condensation
reaction, elongating the acyl chain two carbons at a time. The four steps to make
the initial four carbon fully reduced acyl-ACP are [34]: First, condensation of the
initial malonyl-ACP wtih acetyl-CoA to produce the four carbon 3-ketobutyryl—
ACP by 3-ketoacyI-ACP synthase lll (KAS lll) releasing C02. Second, NADPH-
dependent reduction by 3-ketoacyI-ACP reductase produces 3-hydroxybutyryl-
ACP. Third, removal of water by 3-hydroxyacyl-ACP dehydratase produces a
trans-Az-butenoyl-ACP. Finally, reduction of the double bond by 2,3-trans-enoyl-
ACP reductase utilizing NADPH or NADH and producing the saturated four
carbon chain, butyryl-ACP. Additional condensations with malonyl-ACP are
catalyzed by different enzymes than the original condensation with acetyl-CoA.
During elongation from butyryl-ACP (4C) to palmitoyl-ACP (16C) condensation is
provided by 3—ketoacyl-ACP synthase l (KAS I). PalmltoyI-ACP can be
condensed once more with malonyl-ACP by 3-ketoacyl—ACP synthase ll (KAS II)
and reduced to produce the 180 saturated stearoyl-ACP. In this manner
saturated acyl chains are elongated two carbons at a time and after each
condensation the reduction, dehydration and further reduction of the keto-,
hydroxyl-, and enoyl- ACPs are catalyzed by the enzymes listed above. 16:0-
and 18:0-ACP are the major products of FAS however, the major newly
synthesized acyl chain that is incorporated into glycerolipids is the mono-
unsaturated oleic acid (18:1). Oleoyl-ACP is produced by the soluble stroma
localized stearoyl-ACP desaturase by inserting a double bond at the A 9 position

[35]. Thus de novo FAS and initial desaturation within the plastid of pea leaves,

 

for example, produces 18:1 (~75%), 16:0 (~20%), and 18:0 (~5%) acyl chains for
further lipid metabolism.

Plant membrane and storage glycerolipids mostly contain acyl chains
other than the initial products of plastid FAS. Leaf membrane lipid acyl groups
typically contain PUFA of either 16- or 18- carbons in length; most commonly
18:2, 18:3 and 16:3. Production of PUFA is accomplished after removal of the de
novo synthesized acyl groups from ACP and incorporation into membrane lipids
within the ER or plastid envelop membranes [36, 37]. Utilization of membrane
lipid acyl groups for desaturation in plants differs from that of animals, yeast and
fungi where the desaturases act on acyl Coenzyme A (CoA) thioesters as
substrates [38, 39]. Additionally, longer chain fatty acids (>180) can accumulate
in some minor membrane lipids such as phosphatidylserine (PS) or storage TAG.
Extra-plastidial elongation of acyl groups beyond 18 carbons may utilize
glycerolipids as a primer to donate the acyl substrate [40]. The donated acyl
chain is elongated two carbons at a time with the steps analogous to plastid FAS
except that the intermediates of elongation are bound to CoA instead of ACP

[41].

1.2.1 AcetyI-CoA carboxylase: Major FAS regulatory point

Production of malonyl-CoA by acetyl-CoA carboxylase (ACCase) is the
first committed step in de novo FAS [33] and regulation of ACCase activity
ultimately determines the flux through all of FAS [42]. All plants except the grass

family Gramineae contain two forms of ACCase, a homomeric and heteromeric

form [43]. The homomeric form produces the cytosolic malonyl—CoA pool, while
the heteromeric form is plastid localized and produces the malonyl-CoA pool for
de novo FAS. A second homomeric form that is plastid localized has been found
in Brassica napus, however its proportional contribution to FAS is unknown [44].
Since in most plants de novo FAS utilizes the products of the plastid localized
heteromeric ACCase, the discussion will be limited to this isoform and will be
referred to as just ACCase.

There are four subunits that make up ACCase which catalyzes two half
reactions. In the first half reaction the biotin carboxylase (BC) catalyzes the ATP
and H005 dependent carboxylation of a biotin prosthetic group attached to the
biotin carboxyl carrier protein (BCCP). The activated carboxy-biotin is transferred
to the active site of the carboxyltransferase (CT) made up of q-CT and B—CT
subunits. CT completes the second half reaction by carboxylation of acetyI-CoA
producing malonyl-CoA [45]. The B—CT subunit is encoded by the plastid genome
[46] and requires RNA editing for functionality [47], while the other three subunits
are all encoded by the nuclear genome and each contains a cleavable plastid
targeting peptide [48-50]. All four subunits are coordinately expressed and
assembled together in the plastid [51, 52]. It is believed that expression of the [3—
CT subunit by the plastid coordinates the expression of the other three subunits
[53]. The ACCase complex is mostly stroma localized but may be partially
associated to the inner envelope membrane in rapidly expanding leaves [54].

ACCase activity can be regulated developmentally or enzymatically. All

four subunits of ACCase are developmentally regulated with the highest

expression in the organs with the greatest demand for FA synthesis, for example
expanding leaves and developing oil seeds [52, 55]. Increased demand for long
chain FA induces expression of all subunits, as demonstrated in Brassica napus
seeds expressing a lauroyl—acyl carrier protein thioesterase which terminated
FAS at 12 carbons starving the cell of long chain FA [56]. The enzymatic activity
of ACCase can be activated by the substrate acetyl-CoA [57], and downstream
products of FAS such as long chain acyl-CoAs can cause feedback inhibition
[58]. Light is a strong regulator of FAS with over five times more FAS in the light
than the dark [59]. Some of lights effect on FAS may be due to increases in

' ACCase enzymatic activity without a change in gene expression [46, 57]. Light
causes the pH and Mg2+ concentration of the stroma to change toward the
enzymatic optima of ACCase [60]. A disulfide bond between the o-CT and B-CT
is reduced in the light which increases activity of ACCase. It is believed that the
disulfide reduction is mediated through a thioredoxin linked pathway [61-63]. The
B—CT subunit of ACCase is also reversibly phosphorylated in the light which has
been shown to increase enzymatic activity [64].

Since malonyl-CoA production commits carbon to FAS, this step has been
the subject of multiple attempts to increase FAS and thus TAG in oilseeds. Over
expression of individual nuclear encoded ACCase subunits has not increased
ACCase activity or FAS [65, 66]. Increased expression of the plastid operon that
includes B—CT increased leaf longevity and doubled seed yield in tobacco [53].
However, it is unknown if these effects are due to an increase in ACCase alone

or if the other gene products in the operon contributed to this phenotype.

 

Targeting the Arabidopsis homomeric ACCase to the plastid of Brassica napus
seeds increased in vitro ACCase activity, and caused a 5% increase in seed
TAG [67]. The strong control that ACCase has on the flux of carbon through FAS
indicates the importance of further understanding its regulation in relation to lipid

metabolism.

1.3 Esterification of fatty acids to glycerol-3—phosphate: de novo
glycerolipid synthesis.

Glycerolipids (membrane and storage) contain a glycerol moiety as a
central backbone in which fatty acyl groups and/or polar headgroups are
attached. Polar lipids that make up membranes contain two acyl groups esterified
to the sn-1 and sn-2 positions of glycerol reserving the sn—3 position for a polar
headgroup. TAG contains a third acyl group esterified to the sn-3 position. De
novo glycerolipid synthesis starts with two sequential acylations of glycerol-3-
phosphate (G3P). The first acylation by G3P acyltransferase (GPAT) produces 1-
acyl-2-lyso-glycerol-3—phosphate, commonly known as lyso-phosphatidic acid
(LPA). The second acylation by lyso-phosphatidic acid acyltransferase (LPAAT)
produces 1,2-diacylglycerol-3-phosphate, referred to as phosphatidic acid (PA).
These two acylation steps were first elucidated from microsomes of guinea pig
liver, with the energy of the reaction provided by activated acyl-CoA [68]. Later it
was shown by Eugene Kennedy in rat liver extracts that the diacylglycerol (DAG)
backbone produced from de novo glycerolipid synthesis and dephosphorylation

of PA could be used to make the common membrane lipids phosphatidylcholine

 

(PC) and phosphatidylethanolamine (PE) through enzymatic reaction with
nucleotide activated headgroups, CDP-choline and CDP-ethanolamine,
respectively [69]. Kennedy also demonstrated in chicken liver microsomes that
DAG could be acylated with acyl-CoA to produce TAG [70]. The fundamental
work by Eugene Kennedy in understanding aspects of glycerolipid synthesis has
led to the acylations of G3P en route to TAG synthesis in plants to be commonly

referred to as the Kennedy pathway.

1.3.1 The two pathways for de novo glycerolipid synthesis in plants

Plants contain two pathways for de novo glycerolipid synthesis (Figure 1-
1) involving the GPAT and LPAAT activities producing the common intermediate
PA. 1) The the ER localized “eukaryotic pathway" and, 2) the plastid stroma and
envelope localized “prokaryotic pathway". The prokaryotic pathway was named
because the plastid LPAAT specifically esterifies a 16 carbon FA to the sn-2
position of the glycerol backbone similar to that of cyanobacteria. In contrast, the
eukaryotic pathway acyltransferases localize 16 carbon FA, if present at all, to
the sn-1 position similar to animals [71].

The prokaryotic pathway utilizes the activated acyl-ACPs from FAS and
stearoyl-ACP desaturase, to make the initial PA molecular species 18:1/16:0
(molecular species refer the combination of FA esterified within a particular lipid).
Prokaryotic PA is used to synthesize phosphatidylglycerol (PG) in all plants and a

portion of the plastidic glycolipids in some plants [34]. Plastid localized

Eukagotic pathway I" PI,PS p5 Dammit,“

G3P Lyso-PA ? PA 5]) 1
2 q . .
a s'

Endoplasmic
reticulum

  
 
 

 
    

18:3 plants ~95% of FA
16:3 plants ~62% of FA ,

 
   
  
  
 

  
 
 
   

PA or DAG transferred
Into plastld for eukaryotic
glycollpld synthesis

  
 
 

Note: traditional textbook model
does not include acyl editing

  
 

Plastid

   

1 esp Lyso-PA PA
FAS 162/(2%:321- E —) {jg—D?
Acetyl-CoA m _’ 13:1 13:1 160

   

\ I 18:3 plants ~5% of FA
16:3 plants ~38% of FA

Figure 1-1 Traditional textbook two pathway model of glycerolipid
synthesis

 

 

The two pathway model of leaf glycerolipid synthesis as described in the text.
The model shows general localizationof prokaryotic pathway to the membranes
of the plastid and eukaryotic pathway to the endoplasmic reticulum membranes.
Estimations of the amount of fatty acids that flow through each pathway are
based on the fact that 18:3 plants only make PG by the prokaryotic pathway [71],
and the 16:3 plant Arabidopsis leaf measurements [72]. Abbreviations: glycerol-
3-phosphate, G3P; lyso-phosphatidic acid, Iyso-PA; phosphatidic acid, PA,
diacylglycerol, DAG; phosphatidylcholine, PC; phosphatidylinositol, PI;
phosphatidylserine, PS, phosphatidylethanolamine, PE; phosphatidylglycerol,
PG; monogalactosyldiacylglycerol, MGDG; digalactosyldiacylglycerol, DGDG;
suphoquinovosyldiacylglycerol, SQDG; fatty acid, FA; coenzyme A, CoA; acyl
carrier protein, ACP; fatty acid synthesis, FAS.

desaturases act on the initial 18:1/16:0 molecular species of PG and glycolipids
to produce the highly unsaturated molecular species (eg. 18:3/16:3) that are

characteristic of chloroplasts [37, 73].

Glycerolipid synthesis by the eukaryotic pathway is generally assumed to
utilize newly synthesized acyl groups exported from the plastid. Acyl chains are
hydrolyzed [74] from ACP in the stroma by a thioesterase specific for oleoyl or
saturated acyl—ACP [75, 76] and reactivated to acyl-CoAs by long chain acyl-CoA
synthetase (LACS) in the outer chloroplast envelope [77, 78]. Experiments with
isolated pea chloroplasts suggest a channeled mechanism to LACS rather than
simple diffusion across the plastid membranes [79]. It is then assumed that 18:1
and saturated acyl-CoAs are available within the cyctosol for the ER localized
eukaryotic pathway acyltransferases. Utilization of nascent exported 18:1 and
16:0 by the eukaryotic pathway predicts the initial PA molecular species to be
18:1/18:1 and 16:0/18:1. PA is rapidly converted to PC which is the major
extraplastidial phospholipid in multi—cellular plants [80] and main site of acyl
desaturation for production of all other molecular species [36, 37]. The
diacylglycerol backbone of PE, phosphatidylserine (PS), phosphatidylinositol (PI)
and storage TAG are also produced by the eukaryotic pathway. Additionally, the
eukaryotic pathway can produce the “DAG moiety" for plastid glycolipid
synthesis. New insights into the trafficking events by which the DAG moiety is
transported into the plastid have been achieved recently [80, 81], and will be

briefly discussed below.

12

 

The relative amounts of the eukaryotic or prokaryotic pathway that are
used to incorporate the acyl chain products of FAS into glycerolipids differs in
different types of plants and tissues within the same plant. The amount of
prokaryotic pathway in all plants has been related to the relative amount of PA
phosphatase activity in the plastid to produce DAG for glycolipid synthesis [73].
“18:3” plants, so called because the leaf galactolipids contain primarily 18:3 FA at
the sn-2 position, contain very little or no PA phosphatase activity. In 18:3 plants
only PG is produced from prokaryotic PA. Therefore, greater than 95% of the
FAs produced in the plastid are exported to the ER for acylation to G3P by the
eukaryotic pathway and all of the DAG moiety for plastidic glycolipids in 18:3
plants must be imported into the plastid from the eukaryotic pathway. “16:3"
plants, so called because the leaf galactolipids contain 16:3 FA at the sn-2
position can dephosphorylate PA in the plastid to provide a prokaryotic pathway
DAG moiety for glycolipid synthesis. The relative proportion of FAS products
utilized by the prokaryotic or eukaryotic pathways varies among different 16:3
plants [82]. In the case of the 16:3 plant Arabidopsis, the flux of acyl chains into
leaf glycerolipids is approximately 62% eukaryotic and 38% prokaryotic [72], with
about half of the galactolipid DAG moieties contributed by the eukaryotic
pathway. The eukaryotic pathway is the major pathway of glycerolipid synthesis
in non-leaf and oilseed tissue [83], and it is the dominant pathway among
angiosperms, of which 88% are 18:3 plants [82].

The two pathway scheme for de novo glycerolipid synthesis was first

elucidated by radiotracer studies of plant tissue and isolated organelles [71, 84]

and then confirmed genetically through analysis of pathway mutants [34, 85].
Critical experiments in the elucidation of the two pathways involved pulse—chase
labeling of FAS with [14C]acetate or 1"’COz, and [3H]glycerol labeling of the
backbone moiety of glycerolipids in leaf tissue of 18:3 and 16:3 plants [86, 87].
Over a period of hours it was evident in the 18:3 plant maize, that newly
synthesized FA first accumulated in PC; then during the chase, the labeled FA
and glycerol together were lost from PC and accumulated in MGDG. Thus the
DAG moiety of eukaryotic synthesized PC was incorporated into a plastid lipid.
Rapid synthesis of labeled PC and transfer to MGDG was also evident in
spinach, a 16:3 plant, but to a lesser degree due to a portion of the MGDG
synthesized by the prokaryotic pathway. Furthermore, isolated chloroplasts from
spinach could produce galactolipids from in situ synthesized DAG, however PC
was only produced from chloroplast-synthesized acyl-CoAs when microsomes
were present [84], indicating the physical separation of the two G3P acylation
pathways. The act1 mutant in the 16:3 plant Arabidopsis thaliana provided further
evidence of the two independent pathways by blocking prokaryotic glycolipid
synthesis by a severely reduced plastidial GPAT activity. This forced newly
synthesized FA to enter the eukaryotic pathway for acylation to GSP. In the act1
mutant the eukaryotic pathway provided all of the DAG for glycolipid synthesis,

essentially turning a 16:3 plant into a 18:3 plant [88].

1.3.2 Transfer of eukaryotic synthesized DAG to the plastid for glycolipid
synthesis

14

 

Glycolipids are synthesized from DAG in the plastid envelope. However, in
16:3 and 18:3 plants more than half and all, respectively, of the DAG moiety is
produced by the extraplastidial eukaryotic pathway (Figure 1-1). This separation
in glycerolipid synthesis activities has led to multiple hypotheses on the
mechanism of lipid transfer between the ER and plastid. Pulse-chase labeling in
maize demonstrated that the glycerol backbone and acyl chains of PC were a
precursor to plastid MGDG synthesis. Careful analysis of radiolabeled glycerol
and acyl chains in MGDG molecular species suggested that 18:2/18:2-PC was
the donor molecule for the DAG moiety incorporated into MGDG [86]. DAG was
hypothesized to be the transfer molecule when a cytosolic phospholipase D and
a hypothesized PA phosphatase from cytosolic extracts were added to isolated
pea chloroplasts inducing DAG accumulation in the chloroplast [89]. PA (possibly
derived from PC) was suggested as the transport molecule when a component of
an inner envelope ABC transporter, that could bind PA in vitro, was shown to
disrupt plastid import of eukaryotic DAG when mutated [90]. Furthermore, such
mutants accumulate PA with a eukaryotic fatty acid composition.

An alternative hypothesis to the transfer of a complete DAG moiety
originating from PC is the transfer of lyso-PC. In vitro experiments suggest that
lyso-PC can spontaneously partition through an aqueous environment from
isolated microsomes to isolated chloroplasts [91]. Additionally pulse chase
labeling of leek seedlings suggested that only the sn-1 acyl-chain from PC was
transferred to MGDG and DGDG [92, 93]. However the extremely long time

points of these experiments (24 hr increments) make it difficult to interpret the

15

labeling data in relation to the demonstrated rapid transfer of PC to MDGD in
other species [72, 86-88]. The transfer lipid may not have to traverse the
hydrophilic cytosol at all, but may travel through an ER plastid connection site.
Putative connection sites termed plastid associated membranes (PLAMs)
containing ER enzymatic activities have been found associated with isolated
chloroplasts [94]. Strong physical connections between the ER and chloroplasts
have been demonstrated in vitro utilizing optical tweezers [95]. Connection sites
may allow direct transfer of lipid molecules between the ER and plastid
membrane systems as has been demonstrated for ER-mitochondria connection
sites in rat liver and yeast [96, 97]. The connection site hypothesis, as opposed
to the lyso-PC hypothesis, could also give an explanation of how glycolipids are
exported from the plastid under times of phosphate stress [80, 98], and may also
be involved in the export of newly synthesized FA to the eukaryotic pathway

acyltransferases.

1.4 Long chain acyl-CoA pools for building glycerolipids

Eukaryotic pathway acyl transferases utilize Coenzyme A (CoA) activated
acyl groups to drive the esterification of FA to a glycerol backbone. The
composition of the FA esterified to 00A (e.g. 16:0, 18:3, etc) and acyltransferase
selectivity can affect the molecular species of lipids produced. Acyl-CoAs do not
freely diffuse across membranes and transport may be an active process as in

mammalian mitochondria [99]. Production of acyl-CoAs from FFA by LACS in

16

different subcellular compartments may generate multiple pools of acyl groups
with different compositions within the cell for different acyltransferase reactions.
LACS activity has been shown to take place in the ER [100], chloroplasts [77,
78], mitochondria [101], peroxisomes [102], and lipid bodies [103]. In Arabidopsis
LACS is encoded by a family of nine genes [104] of which two are involved in
activation of acyl groups for beta—oxidation in peroxisomes [105], at least two
located at the plastid envelope are candidates for involvement in export of acyl
groups from the plastid to the eukaryotic pathway [106], and one has been
shown to be very important for cuticle development [107]. The widespread
localization of LACS throughout the cell demonstrates that FFA are generated
throughout the cell, almost certainly by lipase activities, and the reutilization of FA
by acyltransferases requires the activation to acyl-CoAs. The multiple sites of
acyl-CoA synthesis may also be a mechanism for controlling the acyl
composition of multiple subpools of acyl-CoA for production of specific lipid
molecular species, such as highly saturated or polyunsaturated TAG species.
There is evidence that acyl-CoAs involved in lipid metabolism are not
freely soluble or present in membrane bound pools but are bound to acyl-CoA
binding proteins (ACBP) that deliver the activated acyl groups to
acyltransferases. ACBP have been found in all four eukaryotic kingdoms and
some bacteria [108]. ACBP are expressed in all plant organs [109]. High levels of
ACBP accumulate in oilseeds during oil deposition [110], suggesting their role in
TAG synthesis. Arabidopsis contains two ACBPs that are predicted as cytosolic

[111], two ACBPs have been localized to the plasma membrane [112, 113], and

one is targeted to the outside of the cell [114]. Much like the LACS proteins, the
wide distribution of ACBP suggests multiple roles in lipid metabolism. RNAi
knockdown of a membrane associated ACBP in Arabidopsis increased the
18:2/18:3 ratio of phospholipids [109]. Since acyl chains are desaturated while
bound to PC the knockdown of ACBP probably did not affect the desaturation of
18:2. Reduced ACBP levels may affect the delivery of recycled acyl-CoA to
acyltransferases after removal of desaturated FA from PC and reactivation to

acyl-CoA.

1.5 Synthesis of eukaryotic membrane lipid classes

Leaf glycerolipids are primarily made up of polar membrane lipids.
Membrane lipid synthesis starts with PA from de novo glycerolipid synthesis
(Figure 1-1) and follows one of two routes for acquiring the polar headgroup. (1)
Nucleotide—activated head group dependent sn-3 esterification of DAG produced
from PA by a phosphatase. Cytidine 5’-diphoshphate (CDP) activated choline or
ethanolamine is used for PC and PE synthesis and uridine 5'-diphosphate (UDP)
activated sugars are utilized in monogalactosyldiacylgylcerol (MGDG) and
sulfolipid (SL) synthesis [34]. A second galactose can be added to MGDG by
reaction of UDP-galactose with the 6-hydoxyl of the MGDG headgroup to
produce digalactosyldiacylgycerol (DGDG) [115]. (2) PA is reacted with cytidine
5’-triphosphate to make CDP-DAG. The activated CDP-DAG is then reacted with
serine or myo-inositol to produce PS or Pl, respectively. PG is synthesized by the

reaction of CDP-DAG with G3P, producing first phosphatidylglycerol-phosphate,

18

and then PG by a phosphatase reaction [34]. Subcellular localization of the
enzymes responsible for headgroup addition defines where each lipid is
produced. PC, PE, PS, and PI are produced from DAG and PA in the ER [34]
while MGDG, DGDG, SL and most of PG are produced within the plastid

envelope membranes [73, 81].

1.6 TAG synthesis

The traditional view of TAG synthesis (aka Kennedy Pathway) starts with
two consecutive acylations of G3P by the eukaryotic pathway producing PA
(Figure 1-2). PA dephosphorylation to form DAG and addition of a third acyl
group to the sn-3 position of the glycerol backbone by a diacylglycerol
acyltransferase (DGAT), produces TAG and completes the Kennedy Pathway
[116]. Since TAG is not a substrate for FA desaturation, the acyl-CoA pool that
feeds TAG synthesis acyltransferases must contain all the acyl groups found in
TAG. Alternatively, the DAG moiety for TAG synthesis can be produced from PC
by the reverse reaction of CDP-choline:diacylglycerol cholinephosphotransferase
(CPT) [16, 117, 118]. As with the Kennedy pathway final TAG synthesis can be
through an acyl-CoA dependent DGAT. Additionally, an acyl-CoA independent

mechanism can directly transfer an acyl group from the sn-2 position of PC to

19

pc~m

C

GaP.—-—> LPA PA-+ DAG TAG
Kennedy pathway 7
DAG MAG

Figure 1-2 Possible enzymatic reactions leading to TAG synthesis in
oflseeds

Reactions that may be involved in TAG synthesis as mentioned in the text.
Substrates are as text, enzymes are enclosed in boxes. Acyl-CoA pools and how
they are generated are not shown. Enzymes utilizing acyl-CoA as an additional
substrate are rectangles all other enzymes are in ovals. The traditional Kennedy
pathway is highlighted in gray. TAG, DAG and PC are bold because of their
central importance in TAG synthesis. Substrates: G3P, glycerol-3-phosphate;
LPA, Iyso-phosphatidic acid; PA, phosphatidic acid; DAG, diacylglycerol; PC,
phophatidylcholine; LPC, lyso-phosphatidylcholine; MAG, monoacylglycerol;
TAG, triacylglycerol. Enzymes: GPAT, G3P acyltransferase; LPAT, LPA
acyltransferase; LPCAT, LPC acyltransferase; PDAT, phospholipid:diacylglycerol
acyltransferase; DGAT, diacylglycerol acyltransferase; MAGAT,
monoacylglycerol acyltransferase; DDTA, DAG/DAG trans-acylase; PLD,
phospholipase D: PLC, phospholipase C; CPT, diacylglycerol:CDP-choline
cholinephosphotransferase.

20

DAG producing TAG and Iyso-PC by a phospholipid:diacylglycerol transferase
(PDAT) [119-121]. Although their role in oilseeds has been much less
investigated a phospholipase mediated pathway to produce DAG for TAG
synthesis can be proposed. DAG could be produced from PC by phospholipase
C or by phospholipase D plus PA phosphatase, similar to what has been
proposed for transfer of a eukaryotic DAG moiety from PC into the plastid for
galactolipid synthesis [89, 90]. A DAG/DAG transacylase may also be involved in
TAG synthesis [122] and could transfer FA from the sn-2 position of one DAG to
the sn-3 of another, producing TAG and a sn-1-acyl—monoacylglycerol (MAG).
Re-acylation of MAG by a monoacylglycerol-acyltransferase (MAGAT) would
generate a new DAG for TAG synthesis. Despite the multiple pathways that
could be involved in TAG synthesis as indicated by their in vitro enzymatic
activities, only the DGAT reaction has been conclusively shown to be involved in
TAG synthesis in vivo [123, 124]. Further knowledge of the actual enzymes and
pathways that lead to TAG accumulation will allow better strategies for

engineering of TAG quantity and/or FA composition.

1.6.1 Acyl transfer between lipids is required for a majority of TAG
synthesis
Synthesis of polyunsaturated TAG molecular species requires significant
transfer of acyl chains between lipid species. Phosphatidylcholine is the major
site of FA desaturation for membrane lipids and TAG [116], and thus the modified

acyl groups need to be removed for incorporation into TAG (at least for

21

incorporation into the sn-3 position). Additionally, production of many very long
chain (>18C) polyunsaturated FA requires desaturation on PC prior to elongation
and incorporation into TAG [125]. Desaturated FA may be removed by lipases,
reactivated to acyl-CoA and thus be utilized by the Kennedy pathway acyl
transferases to acylate the sn-1, -2, or -3 position of TAG glycerol backbone.
Alternatively, the reverse CPT mechanism of DAG formation is one possible
method for incorporation of desaturated acyl chains into the sn-1lsn-2 positions
TAG [16, 117, 118]. The relative amount of Kennedy pathway or reverse CPT
used to produce TAG in oilseeds is unknown. It has been suggested from
labeling of oilseed species that accumulate PUFA in TAG, that reverse CPT may
be a mechanism for channeling desaturated FA from PC into TAG [126, 127].
Further utilization of a PDAT that specifically transfers PUFA from sn-2 PC into
the sn-3 position of TAG [120] may complete a mechanism to specifically
produce tri-PU FA molecular species of TAG.

Many plants accumulate unusual FA in seed oil and these unusual FA are
mostly excluded from membrane lipids even though most acyl chain modification
takes place on the membrane lipid bound acyl chain [128]. An example is
ricinolelic acid (12-hydroxyoctadec-Q-enoate) which is produced in castor beans
(Ricinus communis) on PC [129, 130] and accumulates up to 90% of FA in seed
oil but is mostly excluded from bulk PC. The exact mechanisms that exclude
these unusual FA from membrane lipids and segregate them into TAGs are
unknown but identification of unusual FA specific lipases [131] suggest that they

may be selectively removed from PC and channeled into TAG through the

22

 

Kennedy pathway [17, 116]. In transgenic plants that produce unusual FA in a
non native plant species the unusual FA tend accumulate in PC in addition to
TAG suggesting that the mechanisms for selective removal of the unusual FA
from membrane lipids and incorporation into TAG are not available in the
transgenic plant [128]. The final step in TAG production may be provided by a
DGAT that is specific for the unusual FA, as demonstrated by the DGAT2
enzymes of castor and tung that appear to preferentially insert ricinoleic and
eleostearic acid, respectively into the sn-3 position of DAG [132, 133].

It is well documented that in oilseeds it is very important for the plant to be
able to transfer unusual modified FA from the site of synthesis in PC to the TAG
molecule where the modified FA accumulates. The transfer mechanisms and
enzymes could be the same or very similar to those for common desaturated FA.
Much of the oilseed data suggests that these deacylation-reacylation enzymatic
activities are specific for channeling unusual modified FA into TAG. However, a
large number of genes in the Arabidopsis genome are annotated as
acyltransferases and phospholipases with unknown metabolic function [134],
therefore it is possible that similar deacylation-reacylation reactions with common

FA may have a more central role in lipid metabolism.

1.7 Suggestive evidence for acyl editing in plants

Maintenance and fine tuning of the fatty acid composition of phospholipid

membranes appears to be essential in all organisms. Acyl editing also termed

23

 

"remodeling" or “retailoring” is one mechanism to achieve this. Acyl editing
involves the deacylation and reacylation of glycerolipids but does not by itself
result in net lipid synthesis. Acyl editing was first recognized as a significant
pathway of lipid metabolism in animal tissues [135]. In mammalian brain tissue in _
which phospholipid metabolism consumes 20% of cellular ATP [136], a quarter of
that is used for the deacylation-reacylation of phospholipids [137]. Evidence for
acyl editing has also been found in prokaryotes and yeast [138, 139].

In plants, acyl editing has not been previously studied as an integral part
of central lipid metabolism but there is ample suggestive evidence that it is
involved. The acyl editing reaction for phospholipids such as PC can take place
through two general mechanisms. The first mechanism involves deacylation by
exchange of acyl groups from PC directly onto CoA producing acyl-CoA and
lyso-PC (CoAzPC acyl exchange). Deacylation by CoAzPC acyl exchange has
been demonstrated in developing safflower cotyledon microsomes, and was
attributed to the reverse reaction of Iysoephosphatidylcholine acyltransferase
(LPCAT) [140]. LPCAT activity has been extensively reported in isolated plant
microsomes [141-144] and chloroplasts [91, 94, 145]. When reported, the
specificity of LPCAT activity is mostly at the sn-2 position of PC. The
identification of LPCAT activities in plant organelles presents the possibility for
CoArPC acyl exchange as an acyl-editing mechanism but does not indicate how
prevalent the reverse LPCAT activity is in vivo. The second possible mechanism
for acyl editing involves deacylation by hydrolytic cleavage of PC by a

phospholipase generating a FFA and a Lyso-PC. To complete the acyl editing

24

 

cycle the FA must be activated to acyl-CoA by LACS and PC is regenerated by
LPCAT activity. It is possible that both acyl chains could be removed by lipases
generating a glycerophosphocholine that could be reacylated to PC. However
glycerophosphocholine dependent acyl transferases have not been reported in
plants. Many plant phospholipases have been identified that could be involved in
an acyl editing cycle, of those the majority are PLAz, which remove acyl chains
from the sn-2 position [146]. Many of these PLAzS are specific for unusual FA
produced in oilseeds [131]. Removal of acyl chains from the sn-1 position has
been less prevalent in the plant literature but there are a few examples [147,
148]. Just as for the CoAzPC acyl exchange mechanism, evidence of lipase
enzymatic activity that could be involved in acyl editing is not evidence of an
actual acyl editing function in vivo.

More substantial evidence that acyl editing is involved in central lipid
metabolism has come from multiple isotopic labeling studies. Isolated
chloroplasts from pea plants could incorporate newly synthesized [14C]acetate
labeled FA into PC through a channeled pool of acyl-CoA [79]. Since chloroplasts
do not contain the eukaryotic pathway for de novo PC synthesis this result
suggests an acyl editing mechanism that can incorporate newly synthesized FA
into previously synthesized PC in the outer chloroplast membrane or an attached
PLAM region. Additionally, oxygen-18 labeling of the carbonyl oxygen of newly
synthesized FA demonstrated that the entire pool of PC acyl chains in spinach
leaves is under a substantial flux of hydrolytic deacylation and reacylation [74].

This hydrolytic turnover of PC acyl chains occurs at a rate similar to the overall

25

 

rate of FAS, and therefore may play a major role in phospholipid metabolism.
Additional insightful evidence on acyl editing came from careful comparison of
prokaryotic MGDG and eukaryotic PC molecular species after labeling leaf disks
of the 16:3 plant Brassica napus with [“Clcarbon dioxide. MGDG contained the '
predicted prokaryotic pathway initial molecular species of dual labeled 18:1/16:0.
The eukaryotic pathway predicts the major initial PC molecular species to be dual
labeled 18:1/18:1. However carbon dioxide pulse-chase labeling produced PC
with a large amount of scrambling of labeled FA with unlabeled FA. The authors
concluded that the acyl chains of the PC pool are under constant exchange
directly after de novo PC synthesis and during their prolonged 48 hr chase period
[149]. These three lines of evidence suggest that the plant PC pool is under a
deacylation-reacylation flux, so that the acyl groups esterified to G3P by the
eukaryotic pathway and incorporated into PC are not a permanent fixture of PC.
However, how acyl editing is tied in with central membrane and storage lipid

synthesis is unknown.

1.8 The in vivo labeling approach

Modern biological science benefits from various approaches to study the
metabolic pathways of life, each with its own strengths and weaknesses. The
pathways of plant lipid metabolism were originally elucidated by two approaches:
1) Discovery of enzymatic activities through in vitro assays of plant extracts, 2)

step by step pathway development through in vivo labeling. Through these

26

 

approaches the two pathway hypothesis for de novo glycerolipid synthesis was
discovered over 25 years ago and it is still valid today [71]. Since that time the
advent of genetic and molecular biology approaches has unleashed very
powerful tools for discovery of individual genes/enzymes involved in metabolic
pathways through mutational analysis. Additionally, the ability to clone and
overexpress genes has greatly increased the ability to study the effects of
changing one enzyme on metabolism in vivo and to purify enzymes for in vitro
assays. However, the redundancy of genes and metabolic pathways, the lethality
of certain gene knockouts, and the ability for plant metabolism to adapt to some
changes, can mean that key points of metabolism may not be revealed by
genetic or molecular biology techniques alone. Additionally, in vitro assays
identify what reactions are present but usually cannot determine their relative
contributions to metabolic fluxes in vivo. Recently, metabolic profiling techniques
have allowed us to study differences in metabolism due to different growth,
environmental or genetic changes. However, analysis of bulk pools of
metabolites may not reveal subcellular compartmentation, metabolic cycles or
rapid conversions of one metabolite to another.

In vivo labeling approaches are still the only way to study the kinetics of
metabolites moving through pathways in vivo and the relative contributions of
parallel or convergent pathways. Recently the in vivo labeling approaches have
been reinvigorated with the use of stable isotope labeling in conjunction with
mass spectrometry, nuclear magnetic resonance (NMR) and sophisticated

software for analysis. Stable isotope labeling over prolonged periods and

27

 

analysis of the composition of the label in metabolic end products (eg.
membrane and storage lipids, starch and cell walls, and proteins) allows a
steady-state determination of the relative fluxes through different known
metabolic pathways. Steady-state labeling that does not fit known metabolic
pathways can suggest areas of uncertainty in the pathway that need to be
revised [150]. The sensitivity of mass spectrometry and NMR often limit stable
isotope labeling to prolonged periods and analysis of steady-state metabolism. In
vivo labeling with radioisotopes is usually the most sensitive method for detecting
small amounts of labeled compounds as they move through metabolic pathways
[151]. Rapid dynamic labeling with radioisotopes, as opposed to long term
steady-state labeling, involves introducing a labeled compound to the plant or
tissue and rapidly stopping metabolism to determine how the labeled compound
has been metabolized. Multiple labelings of different time periods allows us to
generate kinetic curves for accumulation of label overtime in different metabolic
products. Metabolic pathway precursor-product relationships can then be
determined by the relative time it takes for each metabolite to accumulate label
linearly, which corresponds with the complete labeling of all precursor pools.

The research in this dissertation utilizes in vivo dynamic radiolabeling to
investigate if acyl editing is involved in central lipid metabolism. The textbook
eukaryotic pathway hypothesizes that newly synthesized FA are exported from
the plastid, activated to acyl-CoA, and ester'rfed to G3P producing initial PA
molecular species of 18:1/18:1 and 16:0/18:1. The PA is then converted to PC for

further desaturation of the nascent FA. Additionally, the initial PA may be used by

28

 

the Kennedy pathway to produce TAG in oilseed tissue. However, evidence that
nascent FA are channeled directly into PC in isolated chloroplast [79] through a
possible acyl editing mechanism suggest that there may be alternative
mechanisms at work for incorporation of newly synthesized FA into membrane
lipids. Additionally, some unusual FA produced in the plastid that are not further
modified on PC are transiently incorporated into PC prior to accumulation in TAG
[152, 153], suggesting there may also be an alternative to direct plastid export of
acyl groups into the Kennedy pathway for TAG synthesis. The goals of this
dissertation have been to test the glycerolipid synthesis mechanisms in vivo. The
next two chapters utilize in vivo kinetic labeling techniques to investigate the
initial events by which newly synthesized FA and the glycerol backbone are
incorporated into eukaryotic membrane and storage lipids. The results in pea
leaves and in developing soybean embryos demonstrate that the initial flux of FA
into glycerolipids occurs through PC acyl editing and that this pathway should be

considered a major part of eukaryotic glycerolipid synthesis.

29

 

2

INCORPORATION OF NEWLY-SYNTHESIZED FATTY ACIDS
INTO CYTOSOLIC GLYCEROLIPIDS IN PEA LEAVES
OCCURS VIA ACYL EDITING

Published in:
Journal of Biological Chemistry
Vol. 282, NO. 43, pp. 31206-31216, October 26, 2007

Philip D. Bates‘, John B. Ohlrogge’, Mike Pollard’

From Department of Biochemistry and Molecular Biology1 and Department of
Plant Biologyz, Michigan State University, East Lansing, MI 48824.

30

 

2.1 Abstract

In expanding pea leaves over 95% of fatty acids (FA) synthesized in the
plastid are exported for assembly of eukaryotic glycerolipids. It is often assumed
that the major products of plastid FA synthesis (18:1 and 16:0) are first
incorporated into 16:0/18:1 and 18:1/18:1 molecular species of phosphatidic acid
(PA) which are then converted to phosphatidylcholine (PC), the major eukaryotic
phospholipid and site of acyl desaturation. However, by labeling lipids of pea
leaves with [”C]acetate, [“C]glycerol and [”C]carbon dioxide we demonstrate
that acyl editing is an integral component of eukaryotic glycerolipid synthesis.
First, no precursor-product relationship between PA and PC [“C]acyl chains was
observed at very early time points. Second, analysis of PC molecular species at
these early time points showed that >90% of newly synthesized ["C]18:1 and
[“C]16:0 acyl groups were incorporated into PC alongside a previously
synthesized unlabeled acyl group (18:2, 18:3 or 16:0). And third, [“C]glycerol
labeling produced PC molecular species highly enriched with 18:2, 18:3 and 16:0
FA, and not 18:1, the major product of plastid fatty acid synthesis. In conclusion,
we propose that most newly synthesized acyl groups are not immediately utilized
for PA synthesis, but instead are incorporated directly into PC through an acyl
editing mechanism that operates at both sn-1 and sn-2 positions. Additionally,
the acyl groups removed by acyl-editing are largely used for the net synthesis of

PC through glycerol-3-phosphate acylation.

31

2.2 Introduction

In plants acyl carrier protein (ACP)-dependent de novo fatty acid synthesis
(FAS) is restricted to organelles [28]. Essentially all acyl chains for membrane
and storage lipid synthesis are produced in the plastid [30, 34]. The initial
products of FAS, 16- and 18-carbon fatty acids (FAs) esterified to ACP, are
incorporated into glycerolipids by one of two routes: (i) acyl-ACP is used directly
by acyltransferases of the “prokaryotic” pathway within the plastid, or (ii) the acyl-
ACP thioester bond is hydrolyzed during acyl export from the plastid prior to FA
reactivation and incorporation into glycerolipids by acyltransferases of the
“eukaryotic” pathway outside the plastid. This two pathway mechanism for
glycerolipid synthesis was first elucidated by radio-tracer studies [71], and was
confirmed genetically by analysis of Arabidopsis mutants [34, 154]. The
proportions of nascent FA (i.e. immediate products of FAS) incorporated into the
eukaryotic and prokaryotic pathways vary widely among plants and different
tissues within the same plant. The eukaryotic pathway predominates in non-
photosynthetic tissues of all higher plants. In 18:3 plants, so called because they
accumulate predominantly 18:3 and not 16:3 in their leaf galactolipids, 95% of
the FA produced in the plastid is exported for assembly into glycerolipids by the
eukaryotic pathway. Only phosphatidyl-glycerol (PG) is synthesized directly in the
plastid from acyl-ACPs. This is in contrast to leaves of 16:3 plants which can also
synthesize glycolipids by the prokaryotic pathway. Consequently, only about 60%

of FA produced in the chloroplast by 16:3 plants is exported to the eukaryotic

32

pathway [72]. 18:3 plants, which include pea, make up about 88% of angiosperrn

species [82].

It is generally assumed that the major exported products of chloroplast
FAS, namely 18:1 and 16:0 FA, are transferred to the outer envelope of the
plastid where they are activated to acyl-CoAs by a long chain acyl-CoA
synthetase [77, 78]. The eukaryotic pathway for de novo PC synthesis then
utilizes this pool of newly synthesized acyl-CoAs for sequential sn-1 and sn-2
acylations of glycerol-3-phosphate to produce 18:1/18:1 and 16:0/18:1 molecular
species of phosphatidic acid (PA). PA is rapidly converted to phosphatidylcholine
(PC) by the action of PA phosphatase and CDP-choline:1,2-diacyl-sn-glycerol
cholinephospho-transferase [34]. Desaturation of 18:1 to 18:2 and then 18:3 on
PC produces the abundant polyunsaturated molecular species of PC [36, 37].
However, several lines of evidence suggest that this model may be inadequate
and needs to take account of acyl editing. We define acyl editing, often also
termed “remodeling” or “re-tailoring," as any process that exchanges acyl groups
between polar lipids but which does not by itself result in the net synthesis of the
polar lipids. Acyl editing has long been considered an important facet of
phospholipid metabolism [135]. Acyl editing relevant to this work can occur
through two mechanisms. In plants acyl editing via a CoAzPC acyl exchange
mechanism was demonstrated in microsomes isolated from developing seeds
and was attributed to a reverse reaction of lyso-phosphatidylcholine acyl-
transferase (LPCAT) [140]. LPCAT activity has also been described in isolated

chloroplasts [91, 94, 145] as well as microsomes [141-144]. Thus LPCAT allows

33

for a mechanism for acyl editing, although the in vitm results do not indicate how
prevalent the reaction might be in vivo. In this context, isolated pea chloroplasts
incubated with [“Clacetate immediately label PC with newly synthesized FA
through a channeled pool of acyl-CoA [79]. As chloroplasts cannot synthesize PC
de novo, this indicated a functional mechanism for PC synthesis with nascent FA
through an acyl editing mechanism. A second mechanism for acyl editing
involves hydrolysis of the phospholipid, such as PC to lyso-PC or even to
glycerolphosphorylcholine (GPC), activation of the released free fatty acid and its
re—utilization for phospholipid synthesis from lyso-PC or GPC. Based on 1‘30
labeling there is some indication that acyl-chains esterified to bulk cellular PC are
under a constant flux of acyl editing that proceeds wholly or in part through a

hydrolytic deacylation- reacylation cycle [74].

The most direct line of evidence of acyl editing in plants comes from a
careful analysis of the molecular species of monogalactosyl-diacylglycerol and
PC after labeling leaf disks of the 16:3 plant Brassica napus with carbon dioxide
[149]. [“ClCarbon dioxide labeling produced initial acyl-labeled species as
expected for prokaryotic monogalactosyldiacylglycerol, namely dual labeled
16:0/18:1. By contrast for PC a high degree of scrambling between labeled and
unlabeled acyl chains was noted. The authors concluded that there was
continuous exchange of acyl groups between all molecular species of PC
immediately after labeling and during the prolonged pulse-chase period of 48
hours. In this paper we augment and extend these important observations and

conclusions in several ways:

34

(1) We perform rapid kinetic studies to more carefully address PC labeling and
that of its precursors, PA and 1,2-diacyl-sn-glycerol (DAG). We address
whether the initial incorporation of nascent fatty acids occurs via acyl editing,
or if there is a rapid incorporation by de novo PC synthesis via glycerol-3-P
(G3P) and DAG, which is followed by rapid acyl editing of PC.

(2) We perform both total molecular species and stereochemical analyses on
acyl-labeled PC using in vivo experiments with expanding pea leaves and
seedlings. Pea is an 18:3 plant, so this complements the analysis done
previously with the 16:3 plant B. napus [149].

(3) We track molecular species of PC labeled in the glycerol backbone. When
combined with the analysis of acyl labeling, this allows us to propose that sn-
1 acyl editing is as important a component as sn-2 acyl editing.

(4) As a control, we perform rapid labeling experiments in planta using carbon
dioxide. The results parallel those obtained with excised tissue assays,
indicating that there are no wound responses that compromise the metabolic
conclusions obtained from excised tissue experiments.

From these studies weanalyze possible models by which newly synthesized
FA are incorporated into eukaryotic lipids. The data presented in this paper and
from other studies do not allow us to unambiguously define one particular model,
but do narrow down the possible routes in which nascent FA are incorporated
into eukaryotic glycerolipids and suggest future directions to re-examine this

important yet poorly understood area of plant lipid metabolism.

35

2.3 Experimental procedures

Plant Materials—All experiments were performed with Garden pea (Pisum
sativum L. cv Little marvel) grown in soil/perlite/vermiculite (1 :1 :1) mixture at 22-
25 °C under a day/night 8/16-h photoperiod of white light at 185—210 pmol m”2 3“.

Leaves, or in some cases, whole seedlings were harvested 8 days after sowing.

RadiochemicaIs—[1-‘4C]Acetic acid, sodium salt (specific activity 50
mCi/mmol), [“C(U)]glycerol (specific activity 150 mCi/mmol), and [“Clsodium
bicarbonate (specific activity 50 mCi/mmol) were from American Radiolabeled

Chemicals, Inc. (St. Louis, MO).

Leaf [1 4C]Acetate or f4CJGlyceml Labeling —'Leaf labeling experiments
used ~ 0.3 g fresh weight of pea leaf strips per assay, incubated in the light (180-
220 pmol m'2 s") at 22-24 °C with reciprocal shaking in 5 ml media containing 20
mM MES pH 5.5, 0.1x MS salts, and 0.01% Tween-20. Cut leaves were placed in
media without radioisotope and pre-incubated in ambient light for 5 min. Labeling
was started by the addition of radioactive substrate and strong illumination. The
reaction was quenched by transfer of leaves into isopropanol at 80 °C for 10 min.
For labeling of acyl groups 250 uCi of [1-“C]acetic acid (1.0 mM) was used per
replicate. For [“Clglycerol labeling of PC molecular species each incubation
contained 47 pM [”C(U)]glycerol (39 110i). For [“Clglycerol time course the
labeling media contained 25 pM [“C(U)]glycerol (18 pCi) and the media was
reused for consecutive 1, 3, 6 and 9 min time points. In vivo labeling with excised

plant tissue can produce considerable variance between samples due to

36

differences in development and in uptake of substrate. To minimize such
variance each data point for total incorporation into lipids was normalized against
the trend line for all time points to allow improved kinetic plots (Figures 2-1 and 2-

3).

Seedling [14C]Carbon Dioxide Labeling— Assays were conducted with 8-
day potted pea seedlings in a closed 9.5 L glass desiccator under 250 umol m'2
s'1white light. [“C]COz was released by injection of 1 ml H2804 through the
desiccator lid into a vial containing aqueous [“C]NaHC03. The head space of the
vial was quickly flushed with 30 ml air and the desiccator sealed for the desired
time. Labeling was stopped by removal of the shoots at the base of the first
leaves and quenching in 80 °C isopropanol. The assay used ~ 700 uCi substrate
and ~ 40 seedlings. The seedlings from each labeling were split into two replicate
samples (~ 20 seedlings each) for analysis. Although each labeling was
nominally for 5 min, about 5 min was required to remove all the seedlings and

quench, so the assay is reported as of 5-10 minutes duration.

General Methods—Lipids were extracted from hot isopropanol quenched
tissue with hexane/isopropanol [155] after homogenization with a mortar and
pestle. Chlorophyll was determined spectrophotometrically at 652 nm in 20%
aqueous acetone [156]. Radioactivity in the total lipid samples, eluted lipids or
organic and aqueous phases recovered from transmethylation was quantified by
liquid scintillation counting (Beckman Instrument lnc., Fullerton, CA), while
radioactivity on TLC plates was visualized and quantified by electronic

radiography (Packard Instrument Co., Meriden, CT). AgN03-TLC plates were

37

prepared by impregnating Partisil® K6 silica gel 60 A TLC plates (Whatman,
Maidstone, UK) with 10% AgN03 in acetonitrile (wlv), drying in air and activating
at 110 °C for 5 min. Fatty acid methyl esters (FAMEs) were quantified by gas
chromatography (GC) using a flame ionization detection and a DB-23 capillary

column (30 m length x 0.25 mm inner diameter, 0.25 p film thickness; J&W).

Lipid Class Analysis—For kinetic analyses polar lipids were separated on
a K6 silica TLC plates impregnated with 0.15 M ammonium sulfate and heat
: activated (110 °C for 3 hr) prior to lipid loading. Plates were developed in
acetone/toluene/water (91 :30:8, v/vlv) [157]. DAG was analyzed by first
acetylating an aliquot of the total lipids in acetic anhydride/pyridine, (3:2, v/v) and
then separating the 1,2-diacyl-3—acetyl-glycerols by silica TLC developed in
hexane/diethyl ether/acetic acid (50:50:1, vlv/v). PA was isolated by a first
preparative silica TLC separation using a K6 plate developed in
chloroform/methanollwater (6522524, vlv/v), and further purified on a second K6
plate developed in chlorofonnlacetone/methanol/acetic acid/water (10:4:2:2:1,
v/vlvlv/v). For preparative TLC all solvents contained 0.01% butylated hydroxyl-
toluene antioxidant. Lipids were eluted from TLC silica with
chloroform/methanol/water (5:5:1, v/v/v). Chlorofonn and 0.88% aq. KCl were
added to give chloroform/methanollwater ratios of 2:1 :1, (v/v/v), resulting in a
phase separation. The aqueous phase was back extracted with chloroform and
lipids were recovered from the combined chloroform phases. For [“C]glycerol
and [“C]carbon dioxide labeled lipids the proportion of label in the acyl groups

versus the backbone/head group was determined by transmethylation [158] and

38

scintillation counting of the separated organic and aqueous phases. Aqueous
phase radioactivity from PC was determined to be in the glycerol backbone by
silica TLC developed in methanol/waterl28°/o NH4OH/3M NaCl (50:26.6:17.9:3.4,
vlvlv/v), as sample radioactivity co-migrated with glycerol and glycerol-3—

phosphate standards, but not choline.

Radiolabeled Acyl Group Composition— FAM Es were prepared from
purified individual lipids or total lipids by base-catalyzed transmethylation [158].
Recovered FAMEs were separated based on the number of double bonds by
AgNOa-TLC, the plates being developed to % height with hexane/diethyl ether

(1/1, vlv), then fully with hexane/diethyl ether (9:1, vlv).

PC Molecular Species Detennination—PC was separated from other lipids
by silica TLC (K6 plates developed with chloroform/methanol/acetic acid
(75:25:8, v/v/v)). DAG was produced from the purified PC by phospholipase C (B.
cereus, Sigma) digestion [159]. The DAG was acetylated as described above
and the 1,2-diacyl-3—acetyl-glycerols separated into molecular species based on
the number of double bonds by argentation-TLC [159], using a triple
development (‘/2 then ”/4 development in chloroform/methanol (96:4, vlv), then
fully in chloroform/methanol (99:1, vlv)). The proportion of radioactivity in each
band was determined by electronic autoradiography, then each band was eluted,
the recovered lipids transmethylated, and the [“C]FAME analyzed by AgNOa-
TLC, as described above. To determine endogenous acyl groups from isolated
PC molecular species, triheptadecanoin was added as an internal standard to

each fraction during elution and prior to transmethylation and GC analysis of

39

FAME. When necessary 1,2-diacyl-3-acetyl-glycerol fractions recovered from
AgNOa TLC plates were further purified by reverse phase TLC on Partisil® KC18
silica gel 60 A plates (Whatman, Maidstone, UK) developed with

methanol/acetonelwater (75:25:2, v/v/v).

PC Acyl Group Stereochemistry—PC was isolated as described above
and digested with phospholipase A2 (Crotalus atrox, Sigma) [159]. Briefly, PC
was dissolved in 1 ml diethyl ether and 0.5 unit PLAz in 0.1 ml of 50 mM Tris-HCI
pH 8.7, 5 mM CaClz. The reaction was mixed vigorously for 5 min then the ether
was evaporated under N2. To extract lipids 3.8 ml chloroforrnlmethanol (2/1, v/v)
and 1 ml 0.15 M acetic acid were added, the mixture vortexed, the chloroform
phase collected and the aqueous phase back extracted with 2.5 ml chloroform.
Reaction products were separated on silica TLC plates developed with
chloroform/methanol/acetic acid/water (50:30:8/4, vlvlvlv). Radioactivity in the
free fatty acid and lyso-PC fractions was quantified by electronic
autoradiography, then each product eluted and transmethylated by heating at 50
°C in 5% sulfuric acid in methanol for 30 min. Labeled and unlabeled FAME

compositions were determined as described above.

Lipid Molecular Species by Mass Spectrometry—Pea leaf samples were
analyzed by ESl-MS/MS by the Kansas Lipidomics Research Center. Extraction
of lipids was conducted by their standard Arabidopsis leaf protocol (Web site:
ELI/WWW.k-state.edu/lipid/lipidomics/leaf-extraction.html). The data set from this

analysis is available as a supplement in Appendix A, Figure 5-1: S1A-S1M.

40

Net Rate of Fatty Acid Deposition—The net rate of fatty acid synthesis by
pea leaves was determined by harvesting 10 leaves from pea seedlings at the
start and end of the light cycle for three days. After immediate weighing to
determine fresh weight, FAME and chlorophyll contents were measured as
described above. Triheptadecanoin was added during the lipid extraction as an
internal standard, providing methyl heptadecanoate after transmethylation for

FAME analysis by GC.

2.4 Results

Kinetics of Glycemlipid Labeling from [“C]Acetate— Relationships
between precursor and product pools in metabolic pathways are revealed by
kinetic labeling experiments. The linear accumulation of a product, once reached,
coincides with steady state labeling of all precursor pools. In this context
[“C]acetate is ideal for the study of acyl lipid metabolism as it is rapidly taken up
by excised leaf tissue and utilized for FAS, while the lipids are labeled in their
acyl groups, with minimal head group labeling [79]. The kinetics of [“Clacetate
labeling of glycerolipids from rapidly expanding excised pea leaves over a period
of 9 minutes is shown in Figure 2-1. PC was the major radiolabeled glycerolipid
with over 65% of the label at all time points. Furthermore, steady state labeling
was established very rapidly, with no detectable lag (Figure 2-1A) and thus
suggested that PC was a very early product of nascent FA incorporation into

eukaryotic glycerolipids. A second independent 9 min time course gave similar

41

results (data not shown). Phosphatidylethanolamine (PE) was labeled in a
similar manner (Figure 2-1C), with no obvious lag phase, but PC labeling was 15-
fold greater than PE labeling despite endogenous PC levels only being twice that

of PE (Appendix A, Figure 5-1:S1A).

Both the PA and the DAG pools were still filling after the onset of PC
steady state labeling. The label in PA reached a maximum by 3 min whereas
DAG labeling increased throughout the time course (Figure 2-1 B). Thus,
although PA and DAG are intermediates in the de novo synthesis of PC via
acylation of glycerol-3-P, these [‘4C]acyl labeled intermediates do not show a
precursor-product relationship relative to PC labeling, and thus represent
different glycerolipid pools. Of course, we cannot rule out that very small labeled
PA and DAG pools might contribute to PC labeling in a precursor-product
relationship, since experimentally, quenching in these kinetic assays limits the
temporal resolution of the method. A statistical analysis suggested about 0.5 min
variance (95% CI) in rate extrapolations to zero time. In conclusion, we were not

able to detect a [”Clacyl labeled glycerolipid precursor pool to the labeling of PC.

42

 

0/

pmol [14C1C2

w
via
a:
?e

 

 

 

2 3 4.5 6 7 s 910
Minutes

 

 

0/ mg chl

400q

 

pmol [14C1C2
i— [g DJ

 

 

 

 

 

0
pmol [14C]C2:0/ mg chl

 

 

 

Figure 2-1 Time course for
[“Clacetate incorporation
into the acyl groups of
glycerolipid classes by
excised pea leaves.

A-C: Results expressed as
mean pmoles [“Clacetate
incorporation per mg
chlorophyll (pmol [“0102:0/
mg chl) for each lipid class :I:
S.E. (triplicate determinations).
Glycerolipid classes are PC,
squares; DAG, triangles; PA,
inverted triangles; PG,
diamonds; PE, circles.

Acyl Compositions from [14C]Acetate Labeling of Lipids—Figure 2-2 shows
the composition of labeled fatty acids from the [“C]acetate time course, for total
lipids, PC, PA, DAG, PG and PE. Oleate was the major product in total lipids
(Figure 2-2A), with saturates, mainly palmitate, decreasing from 32% at 1 min to
about 24% for the remainder of the assay period. The reason for this decline is '
unknown. There was no detectable desaturation of 18:1 at the earliest time point
but desaturation to 18:2 was detectable from 3 min onwards. PC had less
saturates (11% falling to 8%) whereas PE had more saturates (54% falling to
38%). By the end of the time course desaturation produced 7% linoleate in PC
(Figure 2-28). In contrast to PC and PE, PA (Figure 2-20) contained high levels
of saturates (>60%) throughout the time course and had an acyl composition with
the closest match to PG. PA is also an intermediate of the prokaryotic
glycerolipid synthesis pathway, and so the labeled PA pool is likely the precursor
for prokaryotic PG labeling. In this context, close inspection of Figure 2-1B and 2-
1C shows a kinetic precursor-product relationship between PA (label reaching a
maximum at 3 min) and PG (lag phase ending at approximately 3 min),
confirming this conclusion. DAG contained an intermediate level of saturates
compared to PC and PE. The origin of the labeled DAG is not certain. It is
probably largely eukaryotic in origin, and may arise in part from the reverse
action of CDP-cholinezDAG cholinephospho-transferase or phospholipase C on
labeled PC, or from acyl groups edited from PC re-entering the eukaryotic de

novo glycerolipid synthesis pathway. Any of these explanations is supported by

44

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

A: Total Lipids B PC
a 100 g 103
o- n- F '- -— _
g 80- g 80-
0 C)
'3, 601 3:, 6o-
:: : »
i-Iu 40- _ 4 a
U U 0
V V
——-i 20" “—1 20“
G 6
1 3 . 6 9 1 3 . 6 9
Minutes Minutes

H:
>

 

 

H
”a
c:

A
Q
L

% [14Clacyl chains 9
a a

 

 

 

 

 

 

 

 

     

C
l

 

l
l 3 6 3 6

Minutes Minutes

 

9

l1!
'1
C)

 

 

S
H
sates:
eccca
gill

 

 

 

 

% [MCjacyI chain

 

 

 

 

3 . 6 9
Minutes

 

Figure 2-2 [“ClAcyl composition of [“Clacetate labeled lipids

Total lipids and isolated lipid classes from the time course assays run with
excised pea leaves, as shown in Fig. 1, were analyzed for their labeled acyl
group distribution. Results expressed as mean :t SE. (triplicate determinations).
Each bar represents the % of total lipid radioactivity present in saturates
(16:0>18:0) (black bars), monoenes (18:1) (white bars), and dienes (18:2)
(hatched bars). PG also contained 4% trienes (18:3) at 9 min. A, Total lipid
FAMEs; B, PC; C, PA; D, DAG; E, PG; F, PE.

45

the appearance of labeled linoleate in DAG by 9 min (Figure 2-20). Whatever its
origin, the difference in labeled acyl group composition of PA and DAG compared
to PC supports the conclusion from the kinetic data that they do not represent

precursors of initial PC acyl labeling.

The Kinetics of Glycerolipid Labeling from [’4CIGcherol—[14C]Glycerol is
rapidly taken up by pea leaves and incorporated into the glycerol backbone of
glycerolipids. In addition, acyl groups also become labeled because glycerol-3-
phosphate equilibrates with glycolytic precursors, leading to plastid acetyl-CoA
production [86]. To separately analyze the label from the backbone/head-group
and the acyl-chains, isolated lipid classes were transmethylated. Analysis of the
aqueous fraction from PC by TLC indicated that the radioactivity was contained
in the glycerol backbone and not the choline headgroup, as noted before [86]. In
marked contrast to acetate labeling, [“C]glycerol incorporation into lipids
exhibited a lag (Figure 2-3A). At the earliest time points, radioactivity in DAG was
approximately equal to PC. We assume that the labeled PA includes a large
contribution from the plastid component. However, chloroplast lipid assembly in
18:3—plants does not require PA conversion to DAG, so we also assume the

labeled DAG is largely associated with extra-plastidial membranes.

Two lines of argument support the notion that the biosynthesis of PC from
labeled acetate (Figure 2-1) and from labeled glycerol (Figure 2-3) report

different metabolic processes. First, for de novo PC synthesis the relative

46

A: [14C]glycerol backbone labeling

 

32129

b—I
00 c
c G
I l

   
    

N h a
c c a
I I I

 

 

 

G

   

015553655910
Minutes

CPM/ mg chloroph

B: Proportion of [14Clglycerol
incorporated into acyl groups

 

_‘ UPC .DAG

Mb)
2'?“

i—Ai—NN
Uh:
4I

¢
I

 

U!
I

 

% acyl group [14C]

3 . 6 9
Minutes

Figure 2-3 Time course for incorporation of label from [“Clglycerol into
the backbone and acyl groups of glycerolipids by excised pea leaves.

 

 

 

 

   

_-_

l

6
I
r

 

A: The amount of radioactivity in the glycerol backbone of each lipid. Results
expressed as mean :I: S.E. (triplicate determinations): PC, squares; DAG,
triangles; PA, inverted triangles. B: The % of total lipid [“Clglycerol labeling that
is in the acyl chains. The three samples of PC and DAG (Figure 3A replicates)
were each combined for the glycerol to acyl group determination. PC, white bars;
DAG, black bars.

47

movement of label from PA to DAG to PC should be same for both glycerol and
acyl group labeling strategies. However, at the earliest time points, it is clear that
acetate acyl chain labeling produces PC >> DAG (Figure 2-1A) whereas glycerol
backbone labeling produces DAG = PC (Figure 2-3A), suggesting separate
metabolic processes. Second, when we analyze the different kinetics of acyl
chain and glycerol labeling of PC and DAG from [“C]glycerol labeling (Figure 2-
3B), the amount of label in the acyl chains of DAG remained fairly constant (4-
5%) while the acyl label in PC fell from 32% at 1 min to 12% at 9 min. The
difference is explained by postulating two different pathways for PC synthesis.
About 10% of the label from exogenous glycerol is rapidly utilized for de novo
FAS and labeled acyl groups move rapidly to PC, and then to DAG, as described
for Figure 2—1. The remainder of the labeled glycerol moves through PA to DAG,
which has a half life for pool filling of about 3-4 minutes, and then to PC. Under
these conditions the fraction of PC which is acyl labeled is initially high but
declines steadily, while this simple model predicts that the acyl labeled fraction of
DAG remains low and fairly constant. This is what is observed. Together the
glycerol backbone and acyl chain labeling from [“C]glycerol and the [“C]acetate
acyl group labeling provide evidence for a separate (acyl-editing) pathway for
newly synthesized FA to rapidly enter PC without de novo PC synthesis by the

eukaryotic pathway.

48

A: PC molecular species by AgNO; TLC
35

 

 

 

 

88 so ST SM MM MD MI“ DD 01‘ TT
Molecular Species

B: PC molecular species by ESI-MS/MS

 

24

 

 

vavvesocsceoooooeoeoeocccc
MMMMMMMMMMMMMMMMVVVV

Molecular Species

C: PA molecular species by ESI-MS/MS
4o

GVMN—IQWVMN—I‘OWV’MNWVMN

 

 

 

 

49

Figure 24. Molecular
species analyses of
endogenous PC and PA from
pealeaves.

Data is presented as mole
percentage of each molecular
species within that lipid class
(100%). A: PC molecular
species as determined by
AgN03 TLC/GC (three
determinations). Molecular
species are reported as the
combination of two acyl
groups: S, saturates
(16:0>18:0); M, monoenes
(18:1); D, dienes (18:2); and T,
trienes (18:3). B-C: molecular
species as determined by ESI-
MS/MS (five separate
measurements). Molecular
species reported as (# total
carbons: # total double bonds).
B: PC. CzPA. All error bars are
standard error.

Glycerolipid Molecular Species in Expanding Pea Leaves—Molecular
species of PC from expanding pea leaves were analyzed by GC of FAMEs
following AgNOa-TLC separation of the corresponding 1,2-diacyl-3-acetyl-
glycerols. The data are shown in Figure 2-4A, and are very similar to a published
analysis of PC molecular species from pea leaf microsomes as determined by
reverse-phase HPLC [143]. In addition pea leaf PC molecular species from
expanding leaves were determined by ESl-MS/MS (Figure 2-4B). The major fatty
acids of pea leaf PC are 18:2 (44%), 18:3 (31%), 16:0 (18%), 18:0 (4.2%) and
18:1 (2.6%). The major molecular species that we observe are the expected
combinations of these fatty acids, given that saturates are confined to the sn-1
position. Oleic (18:1) and palmitic (16:0) are the major products of chloroplast
FAS (Figure 2-2A). We note that if these nascent FA were incorporated via de
novo PC synthesis without any contribution from acyl editing then 18:1/18:1-PC
would be the most abundant initial species formed. However this species
represents less than 1% of the total PC (MM, Figure 2-4A). Likewise, the PA
molecular species contain only trace amounts of 16:0/18:1 (034:1) and 18:1/18:1
(C36:2) based on ESl-MS/MS (Figure 2-4C). The majority of the C36:2 PA (and
PC) species will be 18:0/18:2. Total PA contributes 0.1-0.4 mole % to total pea
leaf membrane lipids. The PA profile is quite similar to that for PC, but it does
contain significantly more 16:0/18:3 (34:3). PA may include biosynthetic pools,
pools derived from lipid signaling via phospholipase D [4] and DAG kinase, and

the putative PA pool for delivery of eukaryotic molecular species to the plastid for

50

galactolipids synthesis [90]. PA also contains a small amount of 34:6, which is

presumed to be 18:3/16:3, and may therefore indicate a plastid contribution.

Analysis of Molecular Species of PC from (“CJAcetate Labeling—To
examine the initial molecular species of PC produced from incorporation of
nascent FA, pea leaves were labeled with [“C]acetate for 5 min. This time was a
compromise between allowing enough labeling of PC for the analysis, yet
keeping the amount of labeled 18:2 to a minimum (Figure 2-23). Each labeled
molecular species, analyzed as the 1,2-diacyl-3-acetyl-glycerol derivative, was
also analyzed for labeled fatty acid composition. Figures 2-5A and 2-58 show the
distribution of PC molecular species by mass and [“C]acyl group, respectively. If
acyl chains enter eukaryotic phospholipid synthesis via glycerol-3-phosphate
acylation without acyl editing the predicted initial molecular species of PC would
be 16:0/18:1 and 18:1/18:1 (SN and M/M, respectively). Each will be dual
labeled with newly synthesized FA, and if the saturates are 10% of the total label
(Figure 2-28) then the SIM to MlM labeling ratio will be approximately 1:4. This
does not occur. Also, the SIM molecular species was only labeled with oleate;
the corresponding palmitate was unlabeled. The MM molecular species which
would be predicted to contain 80% of the initial label actually represented only
6.5% of all the labeled molecular species. It is unclear if this species contains
one or both acyl groups labeled. The data in Figure 2-53 indicate that all other
labeled PC molecular species contain one labeled acyl group (18:1 or 16:0/18:0)

together with an unlabeled acyl group (18:2, 18:3, or 16:0/18:0). Thus > 90% of

51

A: Leaf PC mass composition

 

35

 

SS SD

 

 

ST SM MM MD MT DD DT TT
Molecular Species

B: 5 min ['4Clacetate labeled acyl chains

 

in 35
.E

a 30+
'5 25¢
g. 20-
2.154
10-
5-

% [“c

 

SS SD

;

H

 

 

 

 

 

 

 

 

t I I I l F—I—
ST SM MM MD MT DD DT TT
Molecular Species

C: 5 min [”Clglycerol labeled backbone

 

J

“-3
g 30-

 

SS SD

 

 

ST SM MM MD MT DD DT TT
Molecular Spec1es

D: 5-10 min l4C02 labeled acyl chains

1E

 

Mao

 

.Jlfln

SS SD

,1

 

 

 

 

 

 

T I

ST SM MM MD MT DD DT TT
Molecular Species

52

Figure 2-5 Analysis of
endogenous PC molecular
species and those labeled
after short time incubations
of excised pea leaves with
[“Clacetate or [“Clglycerol
or of seedlings with
[“c1coz.

A-D: Molecular species are
represented as pairs of FA,
with bar heights represent % of
species among the total. Bar
shading represent % of FA per
species (mass or 14C): S,
saturates (16:0>18:0), black
bars; M, monoenes (18:1),
white bars; D, dienes (18:2),
hatched bars; and T, trienes
(18:3), gray bars. A:
Endogenous PC molecular
species mass composition from
Figure 2-4A. B: 5 min
[“C]acetate labeled PC
molecular species. Results
expressed as mean 1 SE.
(triplicate determinations). C: 5
min [“Clglycerol labeled PC
molecular species, label was in
the glycerol moiety only (label
in acyl chains has been
subtracted). Results expressed
as mean :t: SE. (duplicate
determinations). D: 5-10 min
[“C]COz acyl chain labeled PC
molecular species. Results
expressed as mean :I: S.E.
(quadruplicate determinations).

the molecular species of [‘4C]PC contain one newly synthesized (labeled) FA in
the same molecule as a previously synthesized (unlabeled) FA. This finding

strongly suggests acyl editing as a mechanism of PC synthesis with nascent FA.

Analysis of Molecular Species of PC Synthesized de novo from
[14C]Glycerol—Because acyl editing only exchanges acyl groups it does not
result in net synthesis of phospholipid. However, in order for a leaf to grow net
(i.e. de novo) PC synthesis is required. To determine the PC molecular species
and hence ascertain which acyl chains are involved in de novo eukaryotic
glycerolipid synthesis, pea leaves were incubated with [“C]glycerol. The PC
glycerol backbone will be labeled regardless of whether the acyl chains are
nascent FA or from acyl-editing. Figure 2-5C shows the distribution of label
among PC molecular species after 5 min [“Clglycerol labeling. The data are
corrected to give only the glycerol backbone labeling by subtracting the small
amount of acyl labeling (Figure 2-3B) that occurs from [“Clglycerol. The PC
molecular species distribution obtained with glycerol backbone labeling (Figure 2-
5C) is quite different from that for acetate acyl labeling (Figure 2-58), but closely
resembles the endogenous mass distribution (Figure 2-5A), with the exception of
a significant reduction in 18:2-containing molecular species and an increase in
18:3-containing species. The major initial PC molecular species predicted from
incorporation of only nascent FAs, 18:1/18:1, was less than 0.2% of labeled
species. The total 18:1 content of the glycerol-labeled PC is 5%, some of which
may derive from nascent FA. We know that at this time point the ratio of

unsaturates to saturates in PC from nascent FA is about 10:1 (Figure 2-23) and

53

that there are negligible polyunsaturates. Thus we infer that at the most there is a
5.5% contribution of newly syntheSized FA to de novo PC biosynthesis, or,
conversely, that the bulk (2 95%) of the acyl groups in PC molecular species
labeled via glycerol backbone come from acyl editing. Figure 2-50, which
describes PC molecular species analysis from [“C]carbon dioxide labeling, will

be described in a later section.

Stereochemistry of PC acylation—Molecular species of PC can be defined
by acyl composition and/or by stereochemistry. To determine the stereospecific
incorporation of newly synthesized FA, PC isolated from 5 min incubations of pea
leaves with [“C]acetate was digested with phospholipase A2 and the reaction
products analyzed (Figure 2-6). 62% of the label was incorporated at the sn-2
position and 38% at the sn-1 position. Saturates were predominately found in the
sn—1 position, indicating that the PLA2 digestion was sn-2-specific. Although de
novo PC synthesis requires equimolar acylations at the sn-1 and sn-2 position
we cannot assume that this will be the case for the 16:0 and 18:1 labeled FA if
they are mixing with an endogenous acyl-CoA pool containing 18:2 and 18:3,
since we do not know the acyl transferase specificity at each position. Previous
workers have reported either a similar sn-1 to sn-2 distribution of acyl label, for

PC from 30 minute acetate labeling of spinach leaves [84], or, using a 2 hour

54

 

an 70
fl
'5' 60‘ "'T
'5 50-
5:40-
3. 30-
:0 20-
10-

0-

 

 

 

 

 

 

38 Elltl.__ail|-

sn-l snLZ

Stereochemical position

Figure 2-6 Stereochemical distribution of [“Clacyl chains in PC.

Pea leaves were labeled for 5 min with [“C]acetate. FAMEs generated from the
fatty acid and lyso-PC products of phospholipase A2 digestion of isolated PC
were separated based on the number of double bonds: saturates (16:0>18:0),
black bars; monoenes (18:1), white bars; dienes (18:2), hatched bars. Results
expressed as mean t SE. (triplicate determinations).

 

55

acetate labeling of leek leaves, a 2:1 to 3:2 enrichment in the sn-2 position [92,
93]. The unequal stereochemical labeling complements the conclusion that

nascent FAs are incorporated into PC along side previously synthesized FA.

Unlabeled FA Composition of Endogenous PC, LPC, and Various Labeled
PCs—From Figure 2-53 we have estimated the composition of the unlabeled
FAs that are associated with the nascent, [“C]acetate-labeled FAs. The
calculated values are shown in Figure 2-7, along with the acyl composition of
endogenous PC (calculated from Figure 2-4A) and of endogenous lyso-PC
(obtained by ESl-MS/MS, Appendix A Figure 5-1 :81 K). To a first approximation
all three profiles are similar. The unlabeled, paired acyl groups from [“C]acetate-
labeled PC have a slightly reduced 18:2 and increased saturates compared to
endogenous PC. The unlabeled fatty acids must have had an inverted
stereochemical distribution to that of the labeled FA measured in Figure 2-6; that
is, 62% sn-1 and 38% sn-2. The fact that the FA profile of lyso-PC was similar to
that of total PC allows us to propose that the endogenous lyso-PC pool is
produced by approximately equal sn-1 and sn-2 deacylations, whether catalyzed
by hydrolysis or by PC acyl exchange with CoA. Thus the appropriate lyso—PC
pool is available for both sn-1 or sn-2 acylations with newly synthesized FA. The
unequal stereochemical acylation with nascent FA found in PC (Figure 26)
therefore may arise from sn-2 acyl migration in lyso-PC to the more

thermodynamically stable sn-1 position, or possibly a higher reacylation of the

56

 

UI
CD

  

A
c
I

N
c
I

% acyl composition
E $

 

 

;\§\\\§\\\\\\\w

   

c
I

Acyl composition

Figure 2-7 Comparison of unlabeled fatty acid compositions deduced from
[“Clacetate and [“Clglycerol labeled PC molecular species to
endogenous pea leaf PC and lyso-PC compositions.

Black bars: [120]FA in the same molecule as [‘4C]FA labeled molecular species,
as calculated from figure SB. White bars: endogenous PC FA mass composition,
as calculated from Figure 4A. Gray bars: endogenous pea leaf lyso-PC FA mass
composition, as calculated from ESl-MS/MS (supplemental Figure S1 K, 5
determinations). Hatched bars: FA composition calculated from [”C]glycerol
labeled PC molecular species, as calculated from Fig. 5D. All error bars are
standard error. S, saturates (16:0>18:0); M, monoenes (18:1); D, dienes (18:2);
T, trienes (18:3).

57

sn-1 position with the unlabeled FA that have been previously removed by acyl
editing. It is also possible that the bulk lyso—PC pool has little to do with the
incorporation of nascent FA. Figure 2-7 also shows the FA composition of

molecular species of PC labeled by [“C]glycerol (calculated from Figure 2-5C).

[1 4C]Carbon Dioxide Labeling—The use of excised tissue facilitates rapid
kinetic analysis. However, incubation of excised leaf tissue might cause a wound
response that could alter metabolism. In particular, lipase activities may be
rapidly and transiently induced that could alter phospholipid metabolism. To
control for this possibility whole pea seedlings were labeled for 5-10 minutes with
[“CJCOZ. Label is rapidly incorporated into both the acyl-chains and glycerol
backbone of lipids. Figure 2-8 compares the distribution of acyl group labeling
among lipid classes for [”C]COz labeling of whole seedlings and [“C]acetate
labeling of cut leaves at approximately equivalent assay times. Both labeling
experiments indicated that PC is the major recipient of newly synthesized FA and
that the relative acyl group labeling among other lipid classes is similar.
Additionally, molecular species of PC were analyzed for acyl-chain labeling from
whole seedlings with [14C1002 (Figure 2-50). This distribution should be
compared with that from [“Clacetate labeling of cut leaves (Figure 2-53). The
slightly longer time of assay (approximately 10 minutes including seedling

removal and quenching) for [“Clcarbon dioxide compared to [“Clacetate

58

 

co
6

I l4C02 I'J [14Cjacetate

as
‘P

 

N
©
I

 

 

 

 

% acyl chain labeling

PC PG DAG PE PA
Lipid Class

Figure 2-8 Distribution of lipid classes, labeled in their acyl chains, after
incubation of pea leaves with [“CJCOZ or [“Clacetate.

Black bars: ~5-10 min [1401002 labeling of intact pea seedlings, with results
expressed as mean :I: S.E. (quadruplicate determinations). White bars: 6 min
[“Clacetate labeling of excised pea leaves (Data from Fig. 2-1, results expressed
as mean :I: S.E. (triplicate determinations».

59

labeling allowed slightly greater desaturation of 18:1 to 18:2. However, it is clear
that the pattern of nascent FA acyl labeling from [”C]COZ matches that of
[“C]acetate. We conclude that the relative proportions of individual lipids and
initial PC molecular species labeled in incubations of excised leaves was not due
to a change in lipid metabolism caused by a wound response but is indicative of

the in planta lipid metabolism.

2.5 Discussion

Largely through analyses of the kinetics, molecular species and
stereochemistry of the immediately labeled products of eukaryotic glycerolipid
synthesis we demonstrate that, in rapidly expanding pea leaves, (1) there was no
detectable precursor-product relationship for PA and DAG in the incorporation of
nascent FA into PC, (2) nascent FA were incorporated into PC more rapidly than
the de novo synthesis of PC from glycerol, suggesting two systems for the
incorporation of acyl groups into PC, (3) greater than 90% of nascent FA were
incorporated into PC molecules in combination with an endogenous FA, (4)
greater than 95% of de novo synthesized PC was esterified by recycled acyl
groups, (5) acyl editing can take place at both the sn-1 and sn-2 positions, and
(6) that pea leaves contain an endogenous lyso-PC pool that could be involved in
acyl editing. Experiments with carbon dioxide labeling of whole seedlings
demonstrated that the metabolism seen in excised leaves is not due to a wound

response.

60

These results lead us to conclude that the major pathway for the
incorporation of newly synthesized FA into eukaryotic glycerolipids is largely
through acyl editing of PC and that de novo synthesis of eukaryotic glycerolipids
primarily utilizes acyl chains recycled from acyl editing. We will discuss these
results in terms of the possible mechanisms and flux models (Figure 2-9) that

can be proposed to drive acyl editing.

18:1/18:1 and 16:0/18:1 Are Not Initial Molecular Species of Eukaryotic
Glycerolipid Synthesis—The classical scheme for eukaryotic phospholipid
synthesis in leaves [34, 37], as summarized in Figure 2-9, Model 1, predicts the
biosynthesis of dual-labeled 16:0/18:1 and 18:1/18:1 PA, DAG and then PC
species sequentially. However, our kinetic analysis failed to detect precursor-
product relationships between PA, DAG and PC pools for acyl group labeling
. (Figures 2-1 and 2-2). Furthermore, we have not been able to observe the
expected, dual-labeled 16:0/18:1 and 18:1/18:1 PC species. Based on estimates
of FAS fluxes and endogenous PA and PC molecular species pool sizes (see
supplement Note 1, Appendix A), labeled 16:0/18:1 and 18:1/18:1 molecular
species of PC should have been easily detected by our analysis if Model 1 was
correct. Additionally, glycerol labeling of de novo synthesized PC revealed mostly
esterification with endogenous saturates and polyunsaturates. and not newly
synthesized saturates and oleate. Taken together these three lines of evidence
strongly suggest that 16:0/18:1 and 18:1/18:1 molecular species are not the first
products of nascent FA incorporation into eukaryotic glycerolipids and that Model

1 is incorrect.

61

 

G3P -> Lyso-PA -> PA -> DAG

V II
PC

FFA/acyl-CoA 3
Lyso-PC
sn-1Isn-2

 

 

 

 

 

G3P —> Lyso-PA —> PA —> DAG

Mildtl 3 Y
Lyso-PC

FFA/acyl-CoA

 

 

sn-l/sn-Z

 

  
 
 

 

 

 

G3P —’ Lyso—PA ‘9 PA "" DAG

Modtl 7 Y{

FFA/acyl-CoA

{LCyso-P
sn-l/sn-Z

 

 

G3P 9 Lyso-PA 9 PA 9 DAG

,
FFA/acyl-CoA $C

Lyso-PC
sn-1Isn-2

 

 

Figure 2-9 Four possible models describing pathways and fluxes for the
incorporation of nascent FA, exported from the chloroplast immediately

after de novo FAS, into PC.

For simplicity, the models are set up not to depend on enzyme specificity, and
show only PC. The mechanisms for removal of fatty acids from PC and provision
of acyl-CoA are not defined, while the width of the arrows gives an indication of
relative flux. The grey box represents the chloroplast; G3P, glycerol-3-phosphate;
FFA, free fatty acid. Model 1 shows a description lacking any acyl editing. Models
2 and 3 are considered the best representations for the acyl editing that
accompanies in the incorporation of nascent FA into PC, but a definitive choice of
the most correct model is not yet possible.

62

Most Newly Synthesized FAs Enter PC by sn-1 and sn-2 Acyl Editing,
while de novo PC Synthesis Primarily Uses Recycled Acyl Groups—We have
shown that the majority of newly synthesized (labeled) FAs in pea leaves are
immediately incorporated into PC molecular species where > 90% are paired with
endogenous (unlabeled) FAs. A similar observation of >60% incorporation of
nascent FA paired with endogenous FA is also reported by Williams et al. [149]
for Brassica napus leaves 1hr after a pulse of carbon dioxide labeling. Because
we have used rapidly expanding leaves from an 18:3 plant, whereas Williams et
al. [149] used mature leaves from a 16:3 plant, it appears that the observation
may be a general one. The result implies some form of acyl editing. Either the
nascent FA are used directly in an acyl editing process (Figure 2-9, Model 2), or
acyl groups are released from endogenous lipids and mix with the nascent FA
prior to completing the acyl editing cycle and/or acylation of glycerol-3-P for de
novo PC synthesis (Figure 2-9, Models 3 and 4). Models 2-4 will be discussed in
more detail later. Whatever the acyl editing mechanism, net (i.e. de novo)
phospholipid synthesis from glycerol-3-P is required for the leaf to grow. Glycerol
labeling showed that de novo PC synthesis utilized mainly endogenous palmitate
(+ stearate), linoleate and linolenate and not newly synthesized FA. Thus the FA
used for de novo PC synthesis must be largely (2 95%) recycled from acyl
editing, and, almost certainly, largely from PC. The fact that saturated FAs are
major acyl groups released by acyl editing suggests that sn-1 hydrolysis or
CoAzPC sn-1 acyl exchange contributes in a major way to this process. This is

confirmed by the results from the acetate labeling experiments shown in Figure

63

258 and Figure 2-6, namely that > 90% of the molecular species contain only
one labeled acyl group and 40% of the labeled acyl groups are at the sn-1
position. For Models 2 and 3, this implies that a sn-1 acyl editing process for
incorporation of nascent FA into PC represents 30-40% of the net flux. This
conclusion from our in vivo labeling represents a substantial biochemical activity
largely overlooked or underrepresented by previous in vitro analyses of -PC acyl

metabolism [94, 140] and in vivo labeling studies [149].

Proposed Models for Acyl Editing—Questions then arise as to the
pathways, fluxes and mechanisms for the metabolism of PC synthesis utilizing
nascent FA from the chloroplast. The puzzle is complex because of the large
number of unknowns, and our current data do not completely resolve the models
proposed. However, it is instructive to review possible models to highlight the
unknowns requiring resolution. Three models are considered which might
accommodate a flux of x moles of nascent FA from the chloroplast, producing
0.5x moles of net PC synthesis (Figure 2-9, Models 2-4). For simplicity, the
models were initially set up not to depend on enzyme specificity, and show only
PC. The mechanisms for removal of fatty acids from PC and provision of acyl-
CoA are not defined. They may be via phospholipase action with activation of the
released free fatty acid by an acyl-CoA synthetase, or via direct CoAzPC acyl

exchange.

Model 2 describes a situation whereby nascent FA are channeled to lyso-
PC. Their incorporation allows just one labeled FA per re-synthesized PC

molecule, as observed experimentally. The endogenous FA released from PC is

64

concomitantly used in the de novo synthesis of PC. The endogenous FA may
enter a general acyl-CoA pool, and the model does not rule out other sites of
phospholipid acyl editing also supplying this pool. Model 2 implies acyl transfer
reactions for nascent FA incorporation and de novo PC synthesis may be
spatially distinct metabolic processes. The model cannot accommodate the
catabolism of PC to GPC, because the acylation of GPC by the nascent, labeled
FA will lead to two labeled acyl groups per PC molecule. As the endogenous,
unlabeled acyl distribution in PC molecular species from acetate labeling closely
matches the endogenous PC acyl distribution (Figure 2-7), the lyso-PC and the
FA pools produced by PC editing must have the similar acyl profiles. This is
confirmed by the endogenous LPC analysis. The fact that both lyso-PC and
unlabeled FA in FAIFA*-PC have compositions identical to the bulk PC does not
absolutely require sn-1 and sn-2 deacylations to be on an equimolar basis.
However, the phospholipase/acyl exchange activities (and any lyso-PC
isomerization) will control the sn—1 and sn-2 distributions observed in acetate-
labeled acyl groups. Thus Model 2 can accommodate the excess sn-2 acylation
noted in acetate labeling. Model 2 requires an explanation for the reduction in
18:2 FA content for glycerol-labeled PC, by, for example, proposing a preference
for the transfer of 18:2-containing acylglycerol moieties to the chloroplast for

galactolipid synthesis [86].

Model 3 describes a situation whereby the nascent FAs are directed to a
general acyl-CoA pool, along with FAs from PC acyl editing. Thus the pairing of

labeled and unlabeled acyl groups in PC is caused by a high flux of acyl editing,

65

which greatly dilutes out nascent FA in the common acyl-CoA pool. In this model
it is possible to propose that PC editing stops at lyso-PC or instead goes through
two deacylation steps to produce GPC. However, because GPC-dependent
acyltransferases have not been reported in plants, we will focus on the lyso-PC
example, as shown in Figure 2-9 Model 3. An analysis of the molecular species
of PC labeled from acetate showed that 2 90% had single acyl group labeling.
Without any substrate specificity constraints, this requires that > 90% of the flux
of acyl groups in the acyl-CoA pool originate from unlabeled fatty acids
(Appendix A, Figure 5-2). Consistent with this observation, analysis of the
molecular species of PC with glycerol backbone labeling required that 2 95% of
the flux for de novo PC synthesis come from acyl editing. With this model the sn-
1 and sn-2 distributions for nascent FA do not have to fit a 1:1 ratio. Also, it is
easy to explain the deficit of 18:2 acyl groups in the de novo PC labeling with
glycerol. We can simply assert that there is some acyl-CoA selectivity in the de
novo synthesis at either sn-1 or sn-2 acylation. A moderate change in the de
novo synthesis pathway acyl composition will have little effect on the
endogenous acyl group composition measured by acetate labeling. Of course,

the simplicity of this explanation does not prove it.

In Models 2 and 3 the acyl editing cycle (phospholipid deacylation and
reacylation) is distinct from de novo phospholipid synthesis. It is, however,
possible to have de novo phsopholipid synthesis be part of the acyl editing cycle
(Figure 2-9, Model 4). This requires PC to be completely degraded to glycerol or

glycerol-3-P, and choline or choline-P. In this model there is only one pathway for

66

the synthesis of PC, and if the PC turnover rate is high relative to nascent FA
production, much of the labeling data is consistent with this model. However, a
comparison of labeling kinetics for DAG and PC using either acetate or glycerol
substrates strongly suggested that we were dealing with two distinct metabolic
processes for PC synthesis. This is inconsistent with Model 4, and on this basis,

we rule it out as a dominant pathway of leaf cytosolic glycerolipid synthesis.

In this Discussion we have arrived at the conclusion that one of a pair of
simple models (Figure 2-9, Models 2 and 3), offers the best description of
cytosolic glycerolipid synthesis in pea leaves. The models present quantitatively
very different magnitudes for the fluxes involved in acyl editing, and give different
views on how such acyl editing integrates with the incorporation into PC of
nascent FA exported from the chloroplast. Currently, we cannot categorically rule
out either Model 2 or Model 3. Furthermore, it is always possible to build more
complexity into models, including substrate specificity and the possibility of some
hybrid combination. Future oxygen-18 and acyl-CoA pool labeling studies may
yield pertinent information on the mechanism of acyl editing: lipolytic or through
CoAzPC acyl exchange. Enzymology with isolated chloroplasts and microsomes
may yield useful information on transacylation and de novo acyl transferase
mechanisms and selectivity. Such studies must take into account the very
substantial sn-1 component identified for acyl editing. Furthermore, endogenous
PC is 2-fold higher than PE, but acyl labeling of PC from acetate (and also from
glycerol) is 15-fold greater. This is despite the total leaf lyso-PE pool being

approximately equivalent to lyso-PC (Appendix A Figure 5-1 :81 B). Thus, there is

67

7.5 or 15 fold preference for incorporation into PC via acyl editing that will have to
be taken into account in defining any mechanism. A complete understanding may
require the study of metabolism in KO mutants. However, with so many genes
annotated as lipases, acyl hydrolases and transacylases [134], most with
unknown functions, this may not be a facile approach either to this tricky
question. Defining the mechanisms of the incorporation of nascent fatty acids into
glycerolipids will help our understanding of lipid turnover and be relevant to
research areas with more practical applications such as the integration of seed
triacylglycerol synthesis with phospholipid turnover and the effects of biotic and

abiotic stresses on phospholipid metabolism.

2.6 Footnotes

This work was supported by the US Department of Energy grant DE-FGOZ-
87ER13729. ESl-MS/MS analysis was provided by the Kansas Lipidomics

Research Center.

Abbreviations: ACP, acyl carrier protein; DAG, diacylglycerol; FA, fatty acid;
FAME, fatty acid methyl ester; FAS, fatty acid synthesis; GC, gas
chromatography; GPC, glycerophosphorylcholine; LPCAT, lyso-
phosphatidylcholine acyltransferase; PA, phosphatidic acid; PC,
phosphatidylcholine; PE, phosphatidylethanolamine; PG, phosphatidylglycerol.
Fatty acids are described by the convention “carbon number:number of double

bonds.” For example, oleic acid is represented as 18:1. The molecular species of

68

phospholipids are described by FAIFA. When known, the sn-1 position is
represented first, but often the stereochemistry is uncertain. For example, both 1-
oleoyl-2-linoleoyl-sn-phosphatidylcholine and 1-linoleoyl-2-oleoyl-sn-

phosphatidylcholine are described as 18:1/18:2-PC.

69

3 THE DOMINANT INITIAL FLUX OF FATTY ACIDS INTO
SOYBEAN OIL IS THROUGH PHOSPHATIDYLCHOLINE ACYL
EDITING

Philip D. Bates‘, Timothy P. Durrett’, John B. Ohlroggez, Mike Pollard2

From Department of Biochemistry and Molecular Biology1 and Department of
Plant Biologyz, Michigan State University, East Lansing, MI 48824.

The results of this chapter will be submitted for publication in plant physiology.

The ESl-MS/MS analysis of endogenous soybean lipid compositions was
provided by Tim Durrett. The data is presented in Appendix B Tables 6-2, 6-3, 6-
4. The ESl-MS/MS data was reformatted for discussion of soybean lipid
metabolism and comparison to labeled lipid data in Table 3-1 and Figures 3-1, 3-
6, and 3-8.

70

3.1 Abstract

The reactions leading to triacylglycerols (TAG) synthesis in oilseeds have
been well characterized. However, quantitative analyses of acyl group and
glycerol backbone fluxes that comprise extraplastidic phospholipid and TAG
synthesis, including acyl editing and phosphatidylcholine-diacylglycerol (PC-
DAG) inter-conversion, are lacking. To investigate these fluxes we rapidly labeled
soybean (Glycine max) embryos with [“Clacetate and [“Clglycerol. Cultured
intact embryos that mimic in planta growth were used to minimize any changes in
metabolism due to wounding. The initial kinetics of newly synthesized acyl chain
and glycerol backbone incorporation into PC, DAG and TAG were analyzed
along with their initial labeled molecular species and regiochemical compositions.
Over 60% of the newly synthesized fatty acids enter glycerolipids through PC
acyl editing, largely at the sn-2 position. This flux, mostly of oleate. was about 4-
fold greater than the flux of nascent [“C]fatty acids into sn-1 and sn-2 positions
of DAG through de novo glycerolipid synthesis. The total flux for PC acyl editing
was estimated to be 2 to 7 times the flux of nascent fatty acids entering the
eukaryotic pathway. Thus recycled acyl groups (16:0, 18:1, 18:2 and 18:3)
provide most of the acyl chains for de novo glycerolipid synthesis. Our results
also suggest that the DAG utilized for TAG synthesis is mostly derived from PC,
whereas de novo synthesized DAG is mostly used for PC synthesis. These
results provide a quantitative demonstration that acyl flux inlout of PC by acyl

editing is the dominate flux in soybean embryos during TAG synthesis.

71

3.2 Introduction

In plants essentially all acyl chains for membrane and storage lipid
synthesis are produced in the plastid by acyl carrier protein (ACP) dependent de
novo fatty acid synthesis (FAS) [28, 34, 160]. However, in oilseed tissue these
acyl groups are utilized almost completely (> 95%) by the extra-plastid pathways
of de novo glycerolipid synthesis, collectively termed the “eukaryotic pathway”
[34]. The plastid acyl-ACPs are hydrolyzed and the free fatty acid and products
(usually 18:1>>16:0>18:0) are transported across the plastid envelope [74]. Once
activated to acyl-CoAs [77, 78], these acyl groups are available within the
endoplasmic reticulum (ER) for incorporation by the acyltransferases of the
eukaryotic pathway. The acyl-CoAs may be used for the sequential sn-1 and sn-
2 acylations of G3P to produce phosphatidic acid (PA) [34]. PA is then converted
to sn-1,2-diacylglycerol (DAG) by the action of PA phosphatase. DAG represents
an important branch point between neutral and membrane lipid biosynthesis.
DAG may be acylated to produce TAG, or converted to phosphatidylcholine (PC)
by CDP-choline:1,2-diacyl-sn-glycerol cholinephosphotransferase (CPT), with an
analogous reaction to form phosphatidylethanolamine (PE) [34]. However, in
both leaves [161] and in oilseeds the flux of nascent fatty acids into PC
dominates over the synthesis of PE, PS and other phospholipids of the
eukaryotic pathway. Desaturation of 18:1 to 18:2 and then 18:3 on PC produces

the abundant polyunsaturated molecular species of PC [36, 37].

72

Recently we investigated the initial steps in eukaryotic membrane lipid
synthesis in expanding pea (Pisum sativum) leaves through in vivo labeling of the
acyl groups and backbones of glycerolipids [162]. The results were inconsistent
with a pathway in which newly synthesized fatty acids are directly esterified to
G3P during de novo glycerolipid synthesis. Instead almost all newly synthesized
fatty acids were esterified to a sn-1 or sn-2 lyso-PC through an “acyl editing”
process resulting in molecular species of PC with one newly synthesized and one
previously synthesized FA. In contrast, de novo glycerolipid synthesis in pea
leaves primarily utilized recycled acyl groups released from PC during acyl
editing. Acyl editing, often also termed “remodeling” or “re-tailoring,” is defined as
any process that exchanges acyl groups between polar lipids but which does not
by itself result in the net synthesis of the polar lipids. Acyl editing may proceed
by at least two mechanisms. The first mechanism involves CoAzPC acyl
exchange producing lyso-PC and acyl-CoA, which was demonstrated in
microsomes isolated from developing seeds and was attributed to a reverse
reaction of Iyso-phosphatidylcholine acyltransferase (LPCAT) [140]. There are
several reports of high LPCAT activity in seeds [130, 140, 142, 144, 163]. A
second mechanism for acyl editing involves hydrolysis of PC to lyso-PC,
activation of the released free fatty acid and its re-utilization for phospholipid

synthesis from lyso-PC by LPCAT.

Plants exhibit both acyltransferase and transacylase mechanisms for the
acylation of DAG to TAG. Three classes of diacylglycerolzacyl-CoA sn-3

acyltransferase have been identified, namely the ER membrane bound DGAT1

73

[164] and DGAT2 [165, 166] classes, and third a soluble cytosolic DGAT class
[167]. In addition, the transacylase PDAT (phospholipid:diacylglycerol
acyltransferase) allows the direct transfer of acyl groups from the sn-2 position of
PC to DAG, producing TAG and lyso-PC products [119]. A DAGzDAG
transacylase activity has also been described [122]. In addition to this diversity of
TAG synthesis mechanisms and biosynthetic genes, oil-accumulating tissues
may utilize different strategies to enrich for polyunsaturated fatty acids (PUFA) in
TAG. PC is the major site of fatty acid desaturation and related reactions, while
evidence for a direct role for DAG or TAG as a desaturase substrate is lacking.
The conversion of PC to DAG (for TAG synthesis) may enrich for PUFAs in TAG.
This conversion may be accomplished by a phospholipase or the reverse action
of CTP-choline:diayclglycerol cholinephosphotransferase [1 17, 1 18]. Secondly,
PUFA may be enriched in TAG by mechanisms of PC acyl editing. One such
general mechanism has already been described above, as PC acyl editing may
enrich the PUFA acyl-CoA pool by either a transacylase or lipase/LPCAT
mechanism [162]. A second possible mechanism arises from the action of PDAT
in TAG synthesis. The LPC produced by PDAT would be re-acylated by the

action of LPCAT.

From the above outline it is clear that there are a number of alternative
metabolic routes for TAG synthesis. These include the recycling of intermediates
of membrane lipid biosynthesis. The alternatives may vary with tissue, species
and development. Different metabolic labeling experiments have been used to

probe the sequence of reactions for TAG synthesis in vivo from a variety of

74

oilseed species. Utilization of a PC-derived DAG moiety has been proposed as
the major pathway for TAG synthesis in excised linseed and soybean cotyledons
[127]. Glycerol backbone labeling of developing safflower cotyledons suggested
that the DAG pool feeding TAG synthesis was in equilibrium with PC, however a
similar experiment with avocado mesocarp suggested that PC was not involved
in TAG synthesis [168]. This led the authors to propose that PC backbone
turnover is used primarily in high PUFA containing TAG synthesis.
Phosphocholine head group labeling in linseed cotyledons was also consistent
with the concept that DAG is in equilibrium with PC, allowing the production of a
highly unsaturated DAG pool prior to incorporation into TAG [117]. The authors
suggested that the reversible CPT reaction is most likely responsible for this PC-
DAG inter-conversion. Labeling of safflower cotyledon slices with 16:0 and 18:1
free fatty acids allowed an alternative mechanism for PUFA enrichment in TAG to
be proposed. An sn-2 specific LPCAT could direct 18:1 towards desaturation on
PC and release PUFA to the acyl-CoA pool for the Kennedy pathway [169].
Although there is now significant in vivo evidence for PC acyl editing in the
eukaryotic pathway in plant leaves [162], the in vivo evidence for such a cycle in

oilseeds is still minimal.

We took several steps to extend and improve on previous studies of
oilseed metabolism to determine the pathway of FA incorporation into membrane
and storage lipids in developing soybeans. To avoid potential artifacts associated
with wounding due to excision of tissues we utilized cultured soybean (Glycine

max) embryos which closely mimic in planta growth and oil accumulation [170].

75

To provide an analysis of the initial events in eukaryotic glycerolipid synthesis we
performed kinetic labeling studies with time points shorter than previous studies.
Labeling was conducted with both [“Clacetate and [14C]glycerol and the labeled
compositions, regiochemistry, and molecular species of labeled PC, DAG and
TAG, along with endogenous molecular species compositions were determined.
Together these approaches have allowed us to develop a more quantitative flux
model of glycerolipid synthesis in vivo that demonstrates that acyl fluxes in and
out of PC by acyl editing is a dominate flux in soybean during TAG synthesis. A
better understanding of the pathways of TAG biosynthesis in developing
soybeans may aid future efforts to engineer soybean oil with improved or novel

properties.

3.3 Results

Embryo culture and endogenous lipid compositions

Accurate interpretation of metabolic labeling studies requires that during
labeling the tissue functions as it would in vivo. In this context the use of cultured
embryos (as opposed to embryo or seed slices in simple buffered media)
minimizes lipase activation and other perturbations of lipid metabolism due to
wounding. Thus we have utilized a soybean embryo culture system that allows
dissected zygotic embryos to develop as in planta for over two weeks [170].
Radioactive precursors are incubated at low concentrations (acetate and glycerol

at 0.5-1 mM) relative to the culture media, which contains sucrose (140 mM),

76

glucose (70 mM) and amino acids (48 mM) at levels that mimic the apoplastic
concentration surrounding the embryo in planta. This minimizes changes in
metabolism due to alternative carbon source utilization during the labeling.
Relationships between precursor and product pools in metabolic pathways
are revealed by kinetic labeling experiments. The linear accumulation of a
product, once reached, coincides with steady state labeling of all precursor pools.
In this context the duration of labeling needs to be short enough so that precursor
pools have not filled before time point sampling begins. Soybean embryo DAG
and PC had very similar FA compositions (Table 3-1) and molecular species
profiles (Figure 3-1A). FA are desaturated on PC, but it is DAG that is the
precursor for TAG. The combined PClDAG pool is expected to provide precursor
DAG for TAG synthesis. TAG FA composition and molecular species distribution
are shown in Table 3-1 and Figure 3-1 B, respectively. Developing soybean
embryos of ~15-20 mg dry weight were pre-cultured for ~3 days before addition
of radioactive substrates, to allow metabolism to equilibrate past any transient
response due to the initiation of embryo culture. Cultured embryos accumulate
~6 mg total biomass per day [170] therefore each embryo doubles in biomass
during the pre-culture. If the ratio of DAG:PC:TAG (Table 3-1) stays the same
during pre-culture the initial pool sizes during labeling would be 208254229260
nmoles lipid embryo" respectively (~ 29230 nmoles total FA embryo").
Developing soybeans produce ~2.7 nmoles FA min'1 embryo‘1 [170]. As about

93% of newly synthesized FA are used to produce TAG, of which 1/3 are utilized

77

Table 3-1 Endogenous lipid pool sizes and fatty acid composition

nmol lipid DAG :l: PC :I: TAG :r

embryo'1

20.8 0.63 54.2 2.8 926 17.9
°/o mol FA
16:0 14.42 0.43 17.14 0.42 14.93 0.25
18:0 8.21 0.38 9.40 0.31 4.55 0.04
18:1 30.60 0.34 28.56 0.24 26.35 0.2
18:2 36.89 0.33 35.04 0.49 41.18 0.2
18:3 9.88 0.27 9.87 0.34 12.33 0.12
20:0 0.66 0.03

DAG, PC and TAG amounts as measured by ESl-MS/MS of 13-14 mg dry weight
developing embryos, before pre-culture. The fatty acid composition as calculated
from the molecular species presented in Figure 3-1. Fatty acid values are
represented as percent mol among each lipid class.

78

A: DAG and PC molecular species

 

 

 

 

 

 

25
IDAG

20 - oPc

g 15-

B

E 10 -
5 i L
0 ..

 

SS SM SD ST MM MD MT DD DT TT

B: TAG molecular species

 

 

 

14
12-
1o-
3 3.
6
E 6' 'l
4‘ ll ll
2- l
0' l
(0:5l: F 2 CIF-CDI-I: 2 o P-cni-tZID F t
w «>02 0 0 Cl 0
m $ m m a a a 8 w w E g E 2 2 2 8 o 0

Figure 3-1 Endogenous molecular species compositions of DAG, PC and
TAG.

A-B, Endogenous glycerolipid molecular species from developing soybean
embryos of 13-14 mg dry weight per embryo as determined by ESl-MSIMS.
Molecular species are represented as two or three FA, with bar heights represent
% of species among the total (random stereochemistry). For comparison to
molecular species separated by argentation TLC the molecular species are
grouped based on the individual FA number of double bonds: Total saturates
(16:0, 18:0, 20:0, etc), S; monoenes (18:1), M; dienes (18:2), D; trienes (18:3), T.
Actual molecular species measurements of the non grouped lipid molecular
species for TAG, DAG, and PC are in Appendix B Tables 6-1, 6-2, 6-3,
respectively. A, DAG (black bars) and PC (white bars) were analyzed as their sn-
1,2-diacyl-3-acetyl-glycerol derivatives. B, TAG.

79

for sn-3 acylation, then ~1.7 nmoles FA min“ embryo" flux through the DAG
moiety precursor pool into TAG. Therefore turnover time of the combined
DAG/PC pool (1500 nmoles FA) would be 882 min (14.7 hrs) if all the DAG and
PC takes part in acyl metabolism. Thus our assays of 2-30 min will be effective
measures of initial rates and products but will not directly measure the long term
precursor-product relationships of FA flux through the DAG moiety into TAG.
However, these longer term relationships may be inferred through full analysis of
the molecular species of products observed and consideration of the end point

compositions.

[“ClAcetate labeled fatty acid products

[“Clacetate is an ideal substrate for the study of acyl lipid metabolism
because it rapidly enters plant tissue and is highly specific for incorporation into
newly synthesized FA . Transmethylation of total lipid extracts resulted in >96%
of the radioactivity recovered in the hexane soluble fraction. TLC analysis of total
lipids at 10 min of labeling indicated <9% of the radioactivity migrated with free
sterols or waxes. Thus, >87% of [“Clacetate incorporated into soybean lipids is
in the acyl moiety of glycerolipids (examples of TLC separation of labeled lipid
classes are in Appendix B Figure 6-3). The rate of [“Clacetate incorporation into
soybean lipid acyl fraction was 3.2 pmol [“C]acetate embryo“ day‘1 (data taken
from experiment shown in Figure 3-2), and is approximately 1/10 of total embryo
fatty acid synthesis rate expressed on a C2 unit incorporation basis (35 pmol
acetyl-CoA embryo'1 day'1 [170]. Therefore, assuming an equal distribution and

utilization of acetate within the embryo tissue, statistically most of the fatty acids

80

synthesized during the labeling period contained a single [“C]C2 unit.
[“C]Oleate (18:1) and [‘4clsaturated FA (16:0, 18:0) (Appendix B Figure 6-1) are
the immediate products of fatty acid synthesis. At 30 min of labeling less than 3%
of the labeled FA were desaturated to [“Cllinoleate (18:2); therefore short time
point labeling represents the initial acyl transferase reactions of newly
synthesized FA, prior to further metabolism. Total labeled saturates increased
from 23% of total FA at 2 min to 42% by 30 min of labeling. Most of the increase
was due to an increase in stearic acid (18:0) labeling. Chemical or-oxidation was
employed to determine the position of the label along the [“C]18:0 acyl chain
(Appendix B Figure 6-2). Label was evenly distributed along the chain. This result
suggested that the 18:0 was a direct product of fatty acid synthesis and not
labeled by a high specific activity cytosolic elongation as reported for other very
long chain FA [171, 172]. The reason for 18:0>16:0 production when
endogenous composition is 16:0>18:0 is unknown. The 18:0>16:0 labeling at
early time points in soybean embryos is not expected to affect the total saturates
acyl group fluxes as 16:0 and 18:0 behave in a similar fashion in acyl transfer

reactions, and have the similar positional distributions in the major lipid classes.

Kinetics of [“Clacetate labeling indicate independent rates of lipid class
acylation
To determine the initial rates and precursor-product relationships for the

incorporation of newly synthesized FA into soybean glycerolipids we followed

81

35

 

 

 

 

5 30 : PC

5‘ 25 .

E

g 20 -

.- TAG

tc . .
§ 15

I“ -

9' 1o .
a 5 - DAG
g 0 3 r T I I l I

C

O 5 10 15 20 25 30 35

Minutes

Figure 3-2 Time course for [“Clacetate incorporation into the acyl groups
of the major labeled soybean embryo lipids.

Results are based on the total radioactivity in each lipid class isolated by TLC
and measured by liquid scintillation counting. Results expressed as nmol of
["C]acetate incorporated per embryo for each lipid. Glycerolipid classes are PC,
diamonds; TAG, triangles; DAG, squares. Time points in minutes are, 2, 4, 6, 10,
20, 30 and 120. The graph shows the first 35 min to focus on initial kinetics.

82

[“Clacetate labeling into the major labeled soybean embryo lipids (Figure 3-2).
At all time points PC, DAG and TAG represent ~85% of labeled glycerolipids. PC
labeling was linear and gave the highest initial rate (1.15 nmol [“Clacetate
embryo“ min“). DAG labeling was also linear over the 30 minute period and
gave a lower initial rate (0.36 nmol [“Clacetate embryo“ min“). The initial rate of
TAG labeling was the lowest of the three major species (0.26 nmol [“Clacetate
embryo“ min“), however TAG labeling accelerated and accumulated more total
radioactivity than DAG over time. Other membrane lipids contained very little of
the newly synthesized FA. Phosphatidylethanolamine (PE) contained less than
8% of the radioactivity observed in PC even though PE mass abundance is
approximately half that of PC. Phosphatidic acid (PA) is an intermediate of de
novo glycerolipid synthesis, but PA labeling was barely detectable (less than 2%
of that for PC). The high rate of fatty acid incorporation into PC is similar to acyl
editing described in pea leaves where PC is the first incorporation product of

chloroplast exported nascent FA [162].

The rate of incorporation of nascent FA into PC is independent of the
incorporation of nascent FA into DAG or TAG indicating independent acylation
reactions. As it is reasonable to assume that the specific radioactivity of the fatty
acids for each process is identical, the relative initial acylation rates are about
10:3:2 respectively. The accelerating rate for TAG synthesis over the 30 minute
period may represent the combination of three possible effects: (i) increasing
contribution from very long-chain saturates, (ii) an accelerated initial rate,

possibly due to higher levels of stearate, and (iii) the movement of labeled

83

diacylglycerol moieties from DAG and PC pools to TAG becomes measurable,
although longer term assays will be required to detect concomitant reductions in
DAG and PC labeling. In the next five sections we describe the acetate labeled
PC, DAG and TAG products in more detail and summarize the product structures

and kinetic results.

[“ClAcetate labeled FA compositions differ among glycerolipid classes
Fatty acid methyl esters (FAME) of PC, DAG and TAG were separated by
argentation TLC to determine [“ClFA composition based on the number of
double bonds (Figure 3-3). At 2 min of labeling [14011821 made up >95% of the
labeled FA in PC and total labeled saturates slowly increased to approximately
25% at 30 min. By contrast total labeled saturates were relatively constant in
DAG and TAG over the first 30 min of labeling. The average amount of saturate
radioactivity between 4 and 30 min of labeling is 29% in DAG and 75% in TAG.
The lower amount of labeled saturates at 2 min most likely represented acyl
group pool filling. By 120 min radioactivity is detected in polyunsaturated fatty
acids (PUFA) in all lipids, and the relative proportions of saturates have dropped
as the newly synthesized FA are slowly equilibrated through metabolism to the
endogenous FA compositions. The very different initial [“ClFA compositions
among PC, DAG and TAG reflects different metabolic processes for incorporation
of nascent FA into glycerolipids, which is consistent with their independent acyl

labeling kinetics.

84

.3.’

% [1 4C] FAME

% [14C]FAME

F?

% [14C]FAME

A
res-0500017
000000

80

40
20

TAG

80
6O
40
20

 

 

 

 

 

6

10

Minutes

20

30

120

 

6

10

Minutes

20

30

120

 

6

10

Minutes

20

30

120

Figure 3-3 Fatty acid
composition of [“Clacetate
labeled lipids.

A-C. Lipid classes from the time
course assays were
transmethylated and the FAME
separated based on the number
of double bonds by AgNOa-TLC,
and quantified by electronic
autoradiography. Each bar
shows the distribution of
radioactivity among different acyl
groups at each time point. Bar
shading: total saturates, S, black;
monoenes (18:1), M, white;
dienes (18:2), D, gray. A, PC; B,
DAG; C, TAG.

Differential positional acylation of [“Clacetate labeled FA among lipid
classes

The regiochemistry of labeled FA distributions in PC, DAG and TAG was
determined by enzymatic digestion (Figure 3-4). At 2 min 86% of the newly
synthesized FA in PC was esterified at the sn-2 position (Figure 3-4A). Over 10
min of labeling, the label at sn-1 increased from 14% to 26%, which was mostly
due to an increase in saturates. The unequal distribution of nascent FA
incorporation into PC is reminiscent of our previous results in pea leaves at 5
minutes, in which acyl editing produced PC labeled at sn-1 and sn-2 positions of
38 % and 62%, respectively [162]. However, acyl editing in soybean embryos
appears to be more sn-2 specific than in pea leaves. The increase in PC sn-1
saturate labeling may reflect a slow conversion of de novo synthesized DAG into
PC. An alternative explanation is that the high [“C]sn-2 composition of early
labeled PC may reflect a large amount of sn-1 re-acylation with unlabeled
recycled saturated FA. As nascent [“Clstearate replaces unlabeled palmitate in
the acyl editing FA pool an increasing amount of labeled saturates are
incorporated into the sn-1 position of PC over time. In this scenario sn-1 acyl
editing may be just as active as sn-2 acyl editing.

The position of [14C]FA acylation in DAG differs significantly from PC, in
that at 2 min of labeling there is more sn-1 labeled FA than sn-2 (Figure3-43).
The relative sn-1 to sn-2 positional labeling equilibrated in DAG by 20 min. Two
possible causes for this equilibration are: (i) equilibration of newly synthesized FA

into a reacycled acyl group pool feeding de novo glycerolipid synthesis; or

86

E?

% [14C]FAME

% [14C]FAME

% 14C

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

PC ID UM IS
100
80- H " —
I
60-
40-
20- n a ll
O‘F'Di T
‘T ‘1‘ ‘7 ‘1‘ '7 ‘1‘ ‘T ‘1'
C C C C C C C C
In in In in m (D (D m
2 min 4 min 6 min 10 min
DAG
I
100 ID UM S
801
60-
40‘ ' "
20~
0.
‘TS'T‘P‘TS‘TS‘TW‘TS‘
CCCCCCCCCCCC
mmmmmmmmwmmm
Min: 2 4 6 10 20 30
' TAG IFFA DDAG IMAG
100
80-
60-
40-
205
0.
2 4 6 10 20 30
Minutes

87

Figure 3-4 Positional analysis
of [“Clacetate labeled acyl
groups in different lipids.

Major labeled lipids from the
[“Clacetate labeling time course
were subject to lipase positional
analysis. Bar heights represent
percent of lipid label in that
position. A-B. Bar shading
represents the labeled FA as a
percent of the whole lipid: total
saturates, S, black; monoenes,
M, (18:1), white; dienes, D,
(18:2), gray. A, PC PLAz
digestion. FAMEs generated
from lipase products lyso-PC
(sn-1) and FFA (sn-2) were
further analyzed by AgNOa-TLC.
B, Acetylated DAG pancreatic
1,3 TAG lipase digestion.
FAMEs generated from lipase
products, FFA (Sh-1) and 1-O-2-
acyl-3-acetyl glycerol (sn-2)
were further analyzed by
AgNOa-TLC. C. TAG pancreatic
1,3 TAG lipase digestion. Bar
shading: FFA (sn-1 or sn-3),
black; DAG (sn-1,2 DAG or sn-
2,3 DAG), white; MAG (sn-2),
gray.

(ii) highly sn-2 labeled PC conversion to DAG to balance out the higher sn-1 de
novo synthesized DAG. Since PC labeling at 20 min is over 4 times that of DAG
(Figure 3-2), the proportion of labeled PC moving back to DAG must be minor
othenlvise the DAG stereochemistry would reflect PC stereochemistry.

The results of the positional analysis of nascent [14C]FA incorporation into
TAG differ from both that of PC and DAG. Lipase digestion of TAG indicates that
almost the entire label at early time points was sn-1 or sn-3 (lipase product: free
fatty acid, Figure 3-4C). Labeled FA can be incorporated into only the sn-3
position of TAG by acylation of an unlabeled DAG; however any sn-1 labeling
must come from de novo synthesis of DAG. Since the DAG regiochemistry is
approximately equal at sn-1 and sn-2 positions, the amount of label at sn-1 of
TAG is predicted to be at most equal to the amount of label at sn-2. However, the
end product of the lipase digestion, MAG, which contains the sn-2 label, is less
than 2% of the total end products (Figure 3-4). If any of the highly sn-2 labeled
PC has been converted to DAG and then TAG, this would further reduce the
amount of label at sn-1. Therefore we conclude that initial incorporation of

nascent FA into TAG is almost completely localized to the sn-3 position.

88

A: 2 min labeled PC

40
35 q - ID

30. EIM
25- ‘ 'S
20-
15-
1o-
50 f]
0'1 1 .»

F—

2

U)
<08 5%

 

% 14C

 

 

 

 

 

 

 

 

 

 

MD

.—
(D

B: 6 min labeled DAG

40
35 1 ID
30 . BM
25 ~ ' IS
20 a
15 -
10 d

3; l .ll.

 

% 14C

 

 

 

 

 

SS
SD
ST
SM
MM
MD
MT
DD
DT
TT

Figure 3-5 Molecular species compositions of [“Clacetate labeled PC and
DAG.

A-B, molecular species are represented as pairs of FA, with bar heights
represent % of species among the total labeled lipid. Bar shading represent % of
[1 C]FA among species. In species with only one color it means that only the
colored FA is labeled the other FA within the molecular species is unlabeled. Bar
shading: S, total saturates, black bars; M, monoenes (18:1), white bars; D,
dienes (18:2), gray bars. Molecular species of labeled lipids were determined for
the earliest time points that had enough radioactivity for the analysis. Examples
of TLC systems used in molecular species analysis are in Appendix B Figure 6-3.
A, 2 min labeled PC. B, 6 min labeled DAG. Additional time point labeled
molecular species compositions for PC and DAG are in Appendix B Figures 6-4
and 6-5, respectively.

89

[“Clacetate labeled PC and DAG molecular species contain one newly
synthesized and one previously synthesized FA

The unequal [“C]FA positional labeling of DAG and PC suggest at least
some nascent, labeled FA are incorporated into DAG and PC next to an
unlabeled FA. To further investigate this facet of product structure the molecular
species of [“C]acetate labeled PC and DAG were analyzed (Figure 3-5).
Examples of the TLC systems used in molecular species analysis are given in
Appendix B Figure 6-3, while the shorthand nomenclature used to describe DAG,
PC and TAG molecular species is contained in the legend to Figure 3-1. At 2 min
the labeled molecular species SM made up 37% of total radioactivity in PC, with
almost all the label as [“C]M (Figure 3-5A). Thus 35% of the total PC must be
[‘Zcisr‘tcw, while only up to 2% of total PC could be the dual labeled species
[“C]S[“C]M. The majority of other species (40%) consisted of unlabeled PUFA
alongside a nascent labeled S or M. Assigning the labeled MM molecular species
of PC (22%) precisely as single or dual labeled is more problematic. Given that
labeled oleate was highly incorporated at the sn-2 position in total PC the
possibility that MM was largely the dual labeled species is highly unlikely.
Furthermore, at least 76% of nascent FA were incorporated into PC alongside
unlabeled S, D or T in proportions relative to their endogenous composition and
the position of acylation. Endogenous PC contained 30% M (Table 1), which was

almost as much as S and T together. Therefore it is highly likely that nascent

90

[”018 and [“C]M were mostly incorporated next to endogenous unlabeled M in
the labeled SM and MM molecular species, respectively.

[14C]Fatty acid positional analysis of DAG indicated that up to 90% of
molecular species could be produced solely from matching sn-1 and sn-2
acylations with nascent [“018 and [“ClM (Figure 3-4B). However, molecular
species of [“C]acetate labeled DAG at 6 min (3- 58) indicated that 56% of initial
labeled molecular species contained a nascent labeled FA next to an unlabeled
PUFA. In the labeled DAG molecular species SM, the unequal S and M labeling
indicated that at least an additional 15% of total DAG is [‘ZC]S[“C]M. Therefore
molecular species analysis of DAG indicates that at least 71% of DAG contains a
newly synthesized FA incorporated in the same molecule as previously
synthesized FA. As with the PC, this analysis cannot reveal how much of the
remaining SM and MM are dual labeled but it is probably much less than 29%.
We conclude that initial [“C]acetate labeled PC and DAG are comprised
predominantly of molecular species that contain one labeled newly synthesized

FA and one unlabeled previously synthesized FA.

[“ClAcetate labeled TAG molecular species indicates bulk DAG was used
for sn-3 acylation with nascent [“C]FA

Six minutes is the earliest time point where enough radioactivity has
accumulated in TAG to conduct a full molecular species analysis. At this time the

TAG molecular species distribution labeled with nascent FA does not match the

91

A: To

 

18
16 -
14 -
12 r
1

“lo 14C

 

ONAODQO
I l

 

 

 

tal 6 min [14C]acetate TAG
ITotal14C
UESI-MS/MS
mEDPEDFOPEEDPOPtDFt
(D a) o o o
magmgaggwwggggzzgoo

B: [14o]s and mom labeled TAG

 

 

 

 

C: [12C]DAG from [14C]acetate TAG

25

 

I 120 DAG
D DAG ESI
I PC ESI

     

 

 

 

SS SM SD ST MM MD MT DD DT TT

92

Figure 3-6 Six min
[“clacetate labeled TAG
molecular species.

A, Total 6 min
[14C]acetate labeled
molecular species (Black
bars), compared to
endogenous molecular
species (White bars).B, 6
min [“Clacetate labeled
TAG molecular species
with [“C]FA labeling. Bar
shading represents the
labeled FA within each
molecular species.
Saturates, 8, black;
monoenes (18:1), M,
white; dienes (18:2), D,
gray; trienes, T. C, the
calculated unlabeled
DAG molecular species
associated with labeled
TAG in panel B,
assuming one sn-3
labeled FA per molecular
species, black bars.
Endogenous molecular
species from Figure 3-1,
DAG (white bars), PC
(gray bars).

endogenous molecular species distribution (Figure 3-6A) but, as expected, is
enriched in species containing 8 and M and is deficient in species containing two
or three polyunsaturated fatty acids. The positional analysis described above
indicated that nascent FA incorporation into TAG was primarily in the sn-3
position. Therefore the [14C]acetate-labeled TAG molecular species in Figure 3-
6A each contain one labeled S or M and two unlabeled FA. The relative
proportions of [“013 and [“ClM labeled TAG are shown in Figure 353. The
calculated unlabeled ['ZClDAG molecular species composition associated with
the sn-3 [14C]FA labeled TAG was very similar to the endogenous bulk DAG and
PC molecular species compositions (Figure 3-6C). Therefore it appears that DAG
generated from the bulk PC pool is utilized for TAG synthesis with newly
synthesized FA. Furthermore, inspection of Figure 3-68 reveals that there is little
DAG selectivity when considering either [“cls or [“clm individually for
esterification at the sn-3 position of DAG. However, this does not rule out
selectivity of DAG molecular species for esterification of 18:2 or 18:3 to the sn-3

position.

Conclusions from [“Clacetate labeling

The [“Clacetate labeling kinetics, and the different labeled FA
composition, positional acylation and molecular species of DAG, PC and TAG
reveal that newly synthesized DAG is not a direct immediate precursor for
nascent FA incorporation into PC or TAG. Three different acyltransferase

pathways (de novo sn-1 and sn-2 acylations to produce DAG, PC acyl editing,

93

and sn-3 acyl transfer to produce TAG) act independently of each other for the
incorporation of nascent FA into glycerolipids. This clarifies ambiguities in
previous seed labeling studies about the overlap of these pathways, at least for
developing soybean embryos. The rapid labeling of PC largely at the sn-2
position suggests an acyl editing mechanism in which FA is largely removed from
the sn—2 position of PC to generate sn-1-acyl-2-Iyso-PC, which is then re-
esterified with nascent FA. The GPAT and LPAT of de novo glycerolipid
synthesis utilize nascent acyl groups mixed with unlabeled acyl groups to
produce the equal regiochemical labeling and mixed molecular species labeling
of [“C]aoetate labeled DAG. Presumably some of the unlabeled acyl groups
released from PC during acyl editing are recycled for de novo glycerolipid
synthesis. The labeled acyl composition, positional, and molecular species
analyses of TAG at early time points demonstrate clearly that nascent FA are
esterified to the sn-3 position of DAG by an acyltransferase system that is
selective towards saturates but not to DAG molecular species. Furthermore, the
DAG is likely derived from the bulk PC pool. Direct incorporation of nascent FA
by PC acyl editing, de novo DAG synthesis and TAG synthesis from preformed
DAG together account for about 85% of all newly synthesized FA incorporated
into cytosolic glycerolipids, while the ratio of initial rates of incorporation
(approximately 10:32, respectively), demonstrates that the major flux is a rapid
acyl editing mechanism with PC, similar to our previous results in pea leaves

[162].

94

[“Clglycerol labeling of lipid backbones and acyl chains indicate a
common backbone pathway and separate acylation pathways among lipid
classes

Lipids produced by de novo glycerolipid synthesis can be tracked with
[“Clglycerol, which is rapidly taken up by developing soybeans and incorporated
into the backbone of glycerolipids through G3P acylation. The polar head-group
of PC is typically not labeled from glycerol in short duration incubations (lasting a
few hours or less) [86, 162]. However, in addition to backbone labeling, acyl
groups are also rapidly labeled because G3P can provide precursors for plastidic
acetyl-CoA synthesis. The glycerol backbone and acyl moiety labeling can be
measured separately after lipid transmethylation. For cultured soybean embryos
the labeled nascent FA composition produced by [“C]glycerol was very similar to
that from [“Clacetate (data not shown) indicating that de novo fatty acid
synthesis products were not dependent on the substrate.

The kinetics of [“Clglycerol backbone labeling of PC, DAG and TAG
(Figure 3-7A) was distinctly different from acyl labeling of glycerolipids by
[14C]acetate (Figure 3-2). At 2 min of [“C]glycerol labeling the backbone
incorporation into DAG was already linear and was approximately four times the
initial rate for PC and more than eight times that of TAG. However, PC
[“Clbackbone labeling rapidly accelerated such that by 30 min PC contained
more label than DAG and by 120 min it was approaching the ~2:1 molar ratio of

endogenous PC:DAG. Thus the DAG and PC labeling showed the expected

95

A: [“Clglycerol backbone labeling

 

 

 

 

.- 700
g, 600 -

E 500 ~

g 400 « DAG

3 300 -

E 200 - PC 1
3 100 i TAG

E. 03 r T r r

E o 5 1o 15 20 25 30 35

Minutes

B: Relative acyllbackbone labeling

45
40 . IPC

35 - D DAG

30 ' ITAG
25 '
20 -
15 s
10 '
5 .
0 .

 

 

 

% [14C]acyl of total lipid

2 6 1O 20 30 120

Minutes

Figure 3-7 Time course for the incorporation of [“Clglycerol into the
backbone and acyl groups of the major labeled soybean embryo lipids.

[“Clglycerol labeled lipids were isolated by TLC and transmethylated to separate
the glycerol moiety from the acyl groups, each fraction quantified by liquid
scintillation counting. A, time course of only glycerol backbone moiety labeling.
Results expressed as pmol [“Clglycerol incorporated per embryo over the time
course (2, 6, 10, 20, 30 120 min). The graph shows the first 35 min to focus on
initial kinetics. B, % of total lipid [“clglycerol labeling that is in the acyl chains
over the time course. Bar shading: PC, black bars; DAG, white bars; TAG, gray
bars.

96

precursor-product relationship kinetics. Incorporation of [“C]glycerol into the
backbone of TAG lagged significantly behind that of DAG and PC. TAG labeling
was still accelerating after 120 min but had not reached the level of labeling
observed for PC or DAG even though endogenous levels of TAG accumulate to
~20 times that of PC. The extreme lag of TAG (as compared to PC) behind DAG
labeling suggests that the bulk of TAG synthesis (> 98%) is not related to rapidly
labeled de novo synthesis of DAG. TAG labeling is more consistent with a
precursor-product relationship with PC. The backbone labeling kinetics in
developing soybeans are consistent with a major TAG synthesis pathway of G3P
to DAG, to PC, to DAG, and finally to TAG and complement the molecular
species analysis of [“Clacetate labeled TAG, that had an unlabeled DAG moiety
very similar to bulk DAG and PC. The Discussion elaborates on this hypothesis
and includes evidence from mathematical modeling of the labeling data through

different possible pathways.

Approximately 5-10% of [“C]glycerol is rapidly incorporated into acyl
chains. Steady-state [“C]glycerol labeling of acyl and backbone precursor pools
coincides with a constant acyl to backbone labeling ratio. The dual acyl/backbone
labeling from [“C]glycerol confirms the conclusions made from separate
[14C]acetate acyl and [“C]glycerol backbone labeling experiments. Figure 3-7B
shows the percent of acyl labeling from [“C]glycerol acyl/backbone labeled lipids.
At 2 min PC contains 22% of the label in the acyl chains, a value which drops to
a steady-state level of ~5% by 20 min. This is consistent with rapid incorporation

of nascent FA into PC through acyl editing while the backbone label lags through

97

the de novo glycerolipid synthesis via the DAG pool. The constant ~5% acyl
labeling of DAG over the time course represents nascent FA and backbone
entering glycerolipids together by de novo G3P acylation. Initially TAG contains
the largest proportion of radiolabel in acyl chains and it takes over 120 min to
equilibrate to the level of PC and DAG. This result supports the hypothesis that
nascent FA can be esterified at the sn-3 position to a DAG pool that is not
immediately produced from de novo synthesized DAG and that TAG steady-state

labeling is consistent with backbone lagging through the large PClDAG bulk pool.

Analysis of acyl chains utilized by de novo glycerolipid synthesis through
molecular species separation of [“Clglycerol backbone labeled lipid
classes

Separation of the backbone-labeled lipid molecular species at early time
points allows for analysis of the acyl groups used for acylation of G3P. The initial
molecular species of backbone-labeled DAG and PC are very similar (Figure 3-8
A-B, respectively), indicating de novo synthesized DAG is converted to PC
without molecular species selectivity. The slightly higher SD and lower MM levels
in labeled DAG compared to labeled PC might indicate a differential utilization of
DAG molecular species by a low flux pathway, such as for PE or Pl synthesis.
The newly synthesized backbone labeled molecular species composition (Figure
3-8A, B) of DAG and PC contain relatively less polyunsaturated molecular

species than the endogenous PC and DAG molecular species,

98

A: [14C]glycerol labeled DAG
35

 

.6 min 140

_ 3° ‘ DESI-MS/MS

gn-

fw-

O

o 15 -

I".

°\o 10 "'
5 .
0 q I

88 SM so ST MM MD MT DD DT TT

 

 

 

 

B: [14C]glycerol labeled PC
35

30*

 

I6 min 14C

D ESI-MS/MS

 

 

 

 

SS SM SD ST MM MD MT DD DT TI'

C: [14C]glycerol labeled TAG
16

 

 

 

 

 

 

14. I30min14C
-12 . DESI-MS/MS
o
310-
58'
06'
3...
=22:
ol
WZOl-ZDl-Dl-tZOl-Ol-tOI-I:
a) a) o o o
w$8w3558mw§§§§228oo

99

Figure 3-8 Molecular
species composition of
[“Clglycerol backbone
labeled PC, DAG and
TAG.

A-C, molecular species
are represented as a
combination of two or
three FA. Total
saturates, S; monoenes
(18:1), M; dienes (18:2),
D; trienes (18:3), T.
[14C]glycerol labeled
molecular species are
backbone labeling only
(acyl chain labeling has
been subtracted), black
bars. Molecular species
were determined at the
earliest time point with
enough radioactivity for
analysis (Figure 3-7A).
The endogenous mass
molecular species
compositions are from
ESI-MS/MS
measurements, white
bars. A, 6 min labeled
DAG, 10 & 30 min time
points are in Appendix B
Figure 6-7B. B, 6 min
labeled PC, 10 & 30 min
time points in Appendix
Figure 6-7A. C, 30 min
labeled TAG was the
minimum labeling time
with enough radioactivity
for analysis.

respectively. Because acyl group desaturation is believed to occur on PC, but not
DAG, de novo DAG has to be converted to PC for further desaturation. And as
mentioned earlier the endogenous bulk PC and bulk DAG molecular species
compositions are very similar (Figure 3-1A). From this deduction and the
observation that the bulk DAG pool molecular species profile (Figure 3-1 A) is
similar to that used for TAG synthesis (Figure 3-6B) and not the de novo
synthesized DAG composition (Figure 3-8A), we conclude that the immediately
de novo synthesized DAG can contribute only a relatively small fraction to the
total endogenous DAG pool. Instead, the bulk of the endogenous DAG must be
generated by another route. That route involves the removal of the
phosphocholine head-group from PC to produce DAG, by one of several possible
mechanisms such as a reversal of the cholinephosphotransferase reaction or by
phospholipase C action.

[“ClGlycerol labeling of the TAG backbone was insufficient for molecular
species analysis until 30 min of labeling. However, at 30 min the [14C]glycerol
backbone labeled TAG molecular species (Figure 3-80) showed a
correspondence to that of initial [“Clglycerol labeled PC and DAG in that the
labeled molecular species composition was relatively less polyunsaturated as
compared to the endogenous composition of the corresponding lipid. The
calculated FA composition esterified to backbone labeled PC, DAG and TAG was
also very similar (Appendix B Figure 6-8). However, the FA composition of TAG

labeled with [14C]glycerol in the backbone was enriched in S and M, and depleted

100

in D and T, when compared to the FA composition of endogenous TAG. Further

interpretation of these results is left to the Discussion section.

3.4 Discussion

The goal of this study is to provide a quantitative analysis of acyl fluxes in
the developing embryo of an oilseed, from the point where free fatty acids are
exported from the plastid to the formation of TAG. It is a timely exercise, given
the advances in identification of acyltransferase genes in recent years. The
literature describing in vivo labeling of developing seeds suggests various
metabolic pathways. However, collectively, it suffers from a number of problems.
(1) The tissue is excised and immediately incubated, often in the absence of an
osmoticum or nutrients. Thus its physiological state relative to the in planta
situation is uncertain. (2) It has been particularly difficult to assess flux directions
and pools for the DAG-PC conversions. (3) Interpretation based on initial rates of
reaction is under-utilized, and there is often a lack of comprehensive product
analysis. (4) Quantitative conclusions have rarely been possible. In order to
undertake initial rate studies we have used C2/C3 precursors. These are rapidly
taken up and utilized, allowing the system to quickly reach a (quasi) steady state
labeling condition and hence the opportunity to obtain initial rate data. By
contrast labeled hexose substrates face very large dilutions and take too long to

reach a steady state. At the acetate concentrations employed a significant

101

absolute rate of incorporation (about 10%) relative to the endogenous rate of C2

utilization for fatty acid synthesis was measured.

Newly synthesized FA are incorporated into cytosolic glycerolipids through
three independent acyltransferase pathways

Through analysis of the kinetics of glycerolipid acyl labeling from acetate,
and analysis of the FA composition, position of acylation, and molecular species
of these products, we demonstrate that three different acyltransferase systems
are responsible for incorporation of newly synthesized FA into cytosolic
glycerolipids. 1) The major flux of nascent FA is through acyl editing of PC.
The highest rate of FA incorporation was into PC. There was no detectable
kinetic lag to the onset of steady-state labeling of PC, indicating that PC is the
immediate product of an acyltransferase utilizing nascent FA. Additionally the
labeled FA composition and stereochemistry of PC indicated it was not related to
the initial labeled DAG, a possible precursor. Molecular species and FA positional
analyses of PC demonstrated that at least 80% of newly synthesized FA are
esterified in the same molecule as pre-existing FA. Finally, [14C]glycerol acyl and
backbone labeling showed that initial labeled acyl groups and backbone are
incorporated into PC independently. Together these results reveal that about
57% of newly synthesized FA are directly esterified to the sn-1 or sn-2 position of
PC through acyl-editing mechanisms. Because GPCAT activity has not been
described in developing oilseeds, whereas microsomal sn-2 LPCAT and acyl

exchange activities are high, we ascribe the acyl editing to acylation of lyso-PC

102

[130, 140, 142, 144, 163]. 2) Newly synthesized FA mixed with recycled FA to
acylate G3P for de novo glycerolipid synthesis. Approximately 17% of newly
synthesized FAs in glycerolipids were incorporated into DAG, approximately
equally distributed between the sn-1 and sn-2 positions. At least 71% of DAG
molecular species contained newly synthesized and pre-existing FA in the same
molecule. [14C]glycerol acyl/backbone labeling demonstrated that nascent acyl
groups and backbone are incorporated into DAG simultaneously. These results
demonstrate that the GPAT and LPAT of the eukaryotic de novo glycerolipid
synthesis pathway utilize a mixed pool of acyl-CoA containing newly synthesized
and recycled FA. The recycled FA most likely originate from PC acyl editing. 3)
Newly synthesized FA were utilized for sn-3 acylation of pre-existing DAG
to generate TAG. Approximately 11% of newly synthesized FA was directly
incorporated into the sn-3 position of TAG, with negligible labeling in the sn-1 and
sn-2 positions. Nascent FA incorporation into TAG was kinetically independent of
de novo DAG synthesis. TAG molecular species analysis indicated that the DAG
used had the same composition as the endogenous bulk DAG derived from PC.
Therefore the direct incorporation of nascent FA into TAG utilized a pre-existing
DAG pool, not de novo synthesized DAG. In a later section below we use the
relative rates of these three independent acyl transferase pathways to construct a
more quantitative model of TAG synthesis in soybeans. This model takes
account of PC acyl editing, which has not be previously included in models of

oilseed TAG metabolism.

103

At least two systems of DAG sn-3 acylation producing TAG can be
idenfified

The first flux, as mentioned above, involves 11% of the total FA labeling
that is directly incorporated into the sn-3 position of TAG. Other nascent FA are
incorporated into glycerol lipids through acyl editing of PC, and de novo DAG
synthesis which constitute approximately 57% and 17% of the total fatty acid
labeling, respectively. The remaining 15% of acyl label unaccounted in the above
three glycerolipid classes is present largely in minor phospholipid species. As
TAG constitutes approximately 93% of the endogenous acyl lipid mass,
eventually about 31% of the nascent fatty acids must end up in the sn—3 position
of TAG. Thus about 20% of the nascent fatty acids incorporated into glycerolipids
other than TAG must eventually move through into the sn-3 position of TAG to
add to the 11% that are immediately incorporated. This represents a second,
distinctive flux of FA into the sn-3 position of TAG.

When compared to endogenous TAG, the molecular species analysis of
TAG from [14C]acetate labeling (Figure 3-60) also strongly suggests two
pathways to TAG, of approximately equal contribution. We estimated that TAG
labeled with nascent FA at the sn-3 position provides approximately 35% of total
TAG synthesis and has a total FA composition of 39% S, 29% M, 25% D and 7%
T, with the unlabeled DAG coming from the endogenous DAG pool (Figure 3-63)
and sn-3 made up of 75% S and 25% M (Figure 3-3C). Since the endogenous
TAG composition is 20% S, 27% M, 41% D and 12% T, it is likely that a separate

contribution to TAG synthesis is required to provide the remaining 65% of TAG.

104

This separate TAG synthesis must produce TAG of the composition 10% S, 26%
M, 49% D and 15% T to balance out the endogenous composition. It is clear that
this second TAG synthesis component is very different, containing reduced
saturates and much higher PUFA than the initial [14C]acetate labeled TAG. The
DAG moiety presumably comes from the bulk unsaturated DAG/PC pool, with a
selectivity for sn-3 acylation for PUFA.

The molecular species analysis of [‘4C]glycerol labeled TAG (Figure 3-8C)
is also consistent with two pathways to TAG. We can group the molecular
species of TAG into three groups; (1) high specific activity species SSS, SSM,
SSD, SST, SMM, MMM (about 16% of TAG endogenous mass); (2) low specific
activity species MTT, DDD, DDT, DTT (about 20% of endogenous TAG mass)
(and possibly STT, MDD and MDT), and (3) the other species, of intermediate
specific activity (about 64% of endogenous TAG mass). Clearly, rapidly labeled
(i.e. high specific activity) molecular species are dominated by high levels of S
and M, consistent with the nascent fatty acid labeling, while slowly labeled
species have high levels of D and T. Time is required for further desaturation of
the PC species derived from de novo synthesized DAG, to eventually produce
DAG species with both positions occupied by PU FA. These can then be utilized
by the TAG synthesis system that acylates the sn-3 position with PUFA, to
produce TAG molecular species with all three positions occupied by PUFA. A
PUFA selective DGAT or PDAT may be involved in this system.

A DAG/DAG transacylase [122] has been proposed for the synthesis of

TAG. Unsaturated FA would be transferred from the sn-2 position of one DAG

105

molecule to the sn-3 position of another, producing TAG and a sn-1-acyl-MAG.
Re-acylation of MAG by MAGAT would bring about a significant enrichment of
label in the sn-2 position of DAG. This is not observed. Therefore this pathway is

not a major route of TAG synthesis in developing soybean embryos.

Interconversion of DAG and PC and the supply of DAG for TAG synthesis

It is clear from previous studies on oilseed metabolism that DAG and PC
can be inter-converted. This is one way in which PUFA can be enriched for TAG
biosynthesis. This inter-conversion is usually postulated to result from a highly
reversible CPT [117, 118, 173]. It has been suggested that the PC and DAG
pools are in equilibrium. The kinetics of glycerol backbone labeling by soybean
embryos shown in Figure 3-7 are similar to those observed for detached linseed
cotyledons [127], and provides the strongest evidence that the majority of TAG
synthesis is from bulk DAG, via retro-conversion from PC, and not de novo
synthesized DAG. If TAG and PC synthesis competed for the de novo DAG pool,
with no opportunity for PC conversion back to DAG (Figure 3-9, Model A), TAG
should label up 20x more rapidly than PC, to generate the desired ratio of end
products. However, the converse is observed: PC labels up about 10x more
rapidly than TAG (Figure 3-7A). The observation that the DAG pool used for sn-3
acylation with [14C]acetate labeled nascent acyl groups closely resembles the
endogenous DAG and PC molecular species profiles (Figure 3-68), and not that
of the de novo DAG pool (Figure 3-88), is also consistent with DAG first moving

to PC, then back to DAG.

106

Model A

G3P —>__) DAG __> TAG

l

PC
M_od__el B

G3P—> _._)DAG __) TAG

ll

Model B, variant 1

G3P 9 DAG __) TAG

ll

PC(active) $ PC(bulk)

M_o_delc
—> <— <— DAG
33" 9D“; —> PC 9 (oil synth) PTAG

Model C, variant 1
G3P —_>_;.DAG _> PC _> DAG

(oil synth) TTAG
Model C, variant 2
cap ‘3‘ DAG -—> PC —> DAG
‘9 (active) (oil synth) —>TAG
PC (bulk)
Mggg! C, variant 3
G3P —> DAG —> PC —> DAG
-> (active) (active) -> TAG
PC (bulk) DAG (bulk)

107

Figure 3-9 Representations of
mathematical models for TAG
synthesis.

Mathematical models tested
against the kinetic labeling
data by running simulations.
Descriptions of the simulations
are in Appendix B. As
mentioned in the text the
mathematical model that most
fits the acetate and glycerol
labeling is Model C, variant 3.

The required DAG-PC inter-conversion might be supplied by different
mechanisms, each of which may display explicit kinetic consequences. The
simplest would be where both PC and DAG are the bulk biosynthetic pools
(Figure 3-9, Model B). However, the DAG pool could be split into de novo and oil
synthesis (oil synth) pools. Such a general mechanism (Figure 3-9, Model C)
might be simplified if the reversible reactions were replaced by unidirectional
reactions (Figure 3-9, Model 0, variant 1). Pool filling simulations for Models B
and C, and several variants, were run to see how closely they would conform to
the experimental kinetics of glycerol backbone labeling shown in Figure 3-7A.
The variants included splitting PC and DAG pools into two pools, which would
consist of a small biosynthetically active pool in kinetic equilibrium with a large
bulk pool, for example (Figure 3-9 Model B, variant 1). Details of these
simulations are provided in the Appendix B. Conditions were set to allow a
simulation of glycerol backbone labeling from DAG that produced continuous
DAG labeling but where PC labeling crossed over DAG labeling at about 30
minutes as in the experimental results (Figure 3-7A). The backbone labeling
models were then tested against the acyl chain labeling results (Figure 3-2) to
see if, when PC was labeled through acyl editing, how quickly would the acyl
label move from PC to DAG. If the equilibration of [14C]acetate acyl label at the
sn-1 and sn-2 positions of DAG is from PC conversion to DAG, in 30 minutes of
assay no more than about 10% of the labeled acyl groups that accumulated in
DAG can originate from PC (which is mostly sn-2 labeled through acyl editing,

Figure 3-4A-B). However, the total label in PC is 3 times that of DAG, so only

108

about 3% of the total labeled acyl groups in PC originating from acyl editing can
move through to DAG in 30 minutes. Simulations of acyl labeling with Model B
failed to meet this criterion. In a simulation of Model B, variant 1 with the acyl
editing input through PC(bulk), the movement of labeled acyl groups from acyl
editing into DAG was still far too fast. By contrast, models of type C were
generally much more successful at describing the DAG-PC precursor-product
glycerol labeling kinetics and yet having slow movement of acyl editing labeling of
PC move through into DAG.

A observation in setting up the model C concerns the possibility of splitting
the PC pool into a small, active pool and a large, bulk pool (Figure 3-9, Model C,
variant 2). If this is done, then acyl editing must be largely a function of the bulk
pool, not the active pool. Although several variants of Model C gave good
simulations, one problem was that they predicted movement of glycerol labeling
into TAG that was much slower than observed. This could be remedied by
splitting the DAG(oil synth) pool into a small active and a large bulk component
(Figure 3-9 Model C, variant 3). The de novo DAG pool must be small because it
is compositionally distinct from the bulk DAG pool. In addition, in leaves, the
endogenous DAG is very small, constituting less than 1% of the PC (Mhaske V.
and Pollard M., unpublished data). This DAG presumably encompasses the de
novo synthesized DAG pool we propose in Model C. In summary, the kinetics of
[“C]glycerol labeling suggest that the major flux of glycerol-3-P through de novo
synthesis into DAG is for PC synthesis, with almost none of the de novo

synthesized DAG being channeled directly to TAG. The conversion of de novo

109

synthesized DAG to PC is fairly rapid, but residence in the large PC pool is much
longer to allow for further desaturation, before conversion back to DAG for TAG
synthesis. The de novo synthesized DAG and the DAG pool destined for oil

biosynthesis are kinetically distinct and not in rapid equilibrium.

A flux model for glycerolipid synthesis in developing soybean embryos

Our analysis of glycerolipid acyl group and glycerol backbone labeling
allows us to generate a model of the flux of acyl groups during oil synthesis
(Figure 3-10). The individual elements of this model have already been discussed
in the three preceding sections. A detailed step-by-step logic of model
construction, along with its implicit assumptions and calculations, is presented in
Appendix B. The flux model structure is based on the kinetic model C, variant 3
(Figure 3-9) which fit the [“Clacetate and [“C]glycerol labeling data. The flux
model tracks 100 moles of fatty acids synthesized in the plastid through lipid
metabolism over a small time increment. The fluxes are determined based off of
initial rates of nascent FA incorporation into extra-plastidic glycerolipids (Figure 3-
2), their composition and molecular species (Figures 3-3, 3-4, 3-5, 3-6), along
with determinations of the acyl groups used by de novo glycerolipid synthesis
(Figure 3-8), and utilizes endogenous lipid compositions (Table 3-1, Figure 3-1)
and mass balance to predict fluxes. Assuming steady state metabolism the end
point reached in the developing soybean embryo accumulating oil will be 93

moles of acyl groups in TAG, 1.4 moles in DAG, 3.6 moles in PC and 2 moles in

110

  

DAG DA

G3P 2 i, \i’ 63%4
i 67 .__..._..PG 63.4 . ,. int: 62 . .. j

 

PA a (de ”OI/O) ‘. . . (Oil synth) 7" (Oil synth) .
59 pc (bulk) _ . PDAT 0-20 '

    
 

(57)

  
       
 

   

 

 

l

__ . E l

' FFA] Acyl editing '

Acyl-.CoA __ cycle I

89 I, \ “425427, 0 20 I11
,. II “x. \ LPC - I
(s <—=-' 100 mol FA: ‘- r \ PUFA selective 0W I
.\ '16:0,18:0,18:1 ‘ ~ ~ ___________ ' I

\ Plastid -..// I Channeled saturates :
"""""" ' or selective DGAT l

 

l ___________ ..
Figure 3-10 Soybean TAG synthesis flux model.

The model assumptions and flux calculations are in Appendix B. The model
follows 100 moles of newly synthesized FA in the plastid through the major fluxes
of lipid metabolism such that final accumulation of FA are TAG 93 moles, PC 3.6
moles, DAG 1.4 moles, other lipids 2 moles. Other membrane lipids in soybean
embryos are mostly made up of PE and PI. The arrows demonstrate the flux
reactions with the major metabolite pools highlighted in boxes. All of the arrows
represent steady-state fluxes, the black arrows are the route that nascent FA
take for incorporation into glycerolipids as measured by [14C]acetate labeling.
Dashed arrows represent uncertainty for that reaction. The numbers next to flux
arrows represent steady-state fluxes or flux regions as calculated in Appendix B
and utilizing mass balance for unknowns. The numbers in parentheses within the
substrate pools represent the major initial accumulation of nascent FA in
glycerolipids PC, DAG and TAG accumulate ~85% of initial FA. The * represents
small fluxes of acyl groups to other lipids which is s 2% of the total steady-state
acyl groups. ~15% of nascent FA initially accumulate in lipids other than PC,
DAG and TAG.

111

other lipids (based on the values in Table 3-1). The model shows the fluxes, or
expected flux ranges, for both the newly synthesized fatty acids and for the total
flux. It should be noted that in terms of an activity (expressed in moles fatty acid
accumulated unit time'1 unit fresh weight") the relative rate of lipid synthesis in
soybean is slow (about 5 fold less) compared to other oilseeds such as safflower
and rapeseed. Thus the model may or may not be a quantitative or qualitative
representation of other oilseeds which accumulate palmitate, stearate, oleate,
linoleate and linolenate.

The model has a single PC pool. Although it is possible to consider
organelles and specific domains of the endomembrane system giving rise to
distinct sub-pools of PC, to date we have no kinetic or metabolic evidence to
support such a conclusion. By contrast, our analysis strongly suggests three
DAG pools. Pools for the immediate product of de novo synthesis of DAG and for
TAG synthesis each contain only a small mole fraction of the total DAG, while the
bulk DAG pool may be associated with the oil body fraction. We cannot rule out
the possibility that a very small fraction of the de novo DAG pool is utilized
directly for TAG synthesis by the traditional Kennedy pathway, as indicated by
the dotted arrow. Likewise, the reversibility of PC-DAG inter-conversions is not
clear: back reactions are indicated as dotted lines. An important aspect of PC
metabolism is the acyl editing cycle. This is the largest acyl flux in the cytosol.
Our analysis cannot place an exact value on this total flux, but an analysis of
saturated fatty acid mass balance suggests a lower limit of over twice the

nascent fatty acid incorporation into PC (about 125 moles time increment“), while

112

an upper limit may be over seven fold (about 427 moles time increment"). In
considering an acyl editing cycle previously, from the data obtained from pea
leaves, we considered the possibility that there was a “channeled” mole of mole
exchange of nascent fatty acids with acyl groups released by acyl editing.
However, if the acyl editing cycle flux is significantly larger than the incorporation
of nascent acyl groups into PC via a PC:Iyso-PC acyl editing cycle, then the
simplest way to depict the process is via a bulk acyl-CoA pool where acyl groups
released by acyl editing and from de novo fatty acid synthesis mix, as shown in
Figure 3-10.

A surprise from the labeling was that we could make a kinetic distinction
for two TAG synthesis systems. TAG synthesis will require 31 nmoles of
acylation at the sn-3 position. About 11 moles are provided by the immediate
incorporation of nascent FA, with a high preference for saturates. Analysis of the
molecular species of [14C]FA labeled TAG suggests that this direct acylation
utilizes the bulk DAG pool, not the de novo synthesized DAG pool. The high sn-3
[”Clsaturated FA labeling of bulk DAG suggest that either a specific pool of acyl-
CoAs high in nascent saturates is delivered to a DGAT enzyme, or that the
DGAT has a strong selectively for saturates from the bulk acyl group pool (Figure
3-10). At least two mechanisms, both of which are drawn into Figure 10, may
provide the remaining 20 mol FA required for TAG synthesis, including
essentially all the tri-PUFA TAG molecular species. A PUFA selective DGAT2
may utilize the bulk acyl pool or a PDAT reaction may transfer sn-2 PUFA from

PC to DAG, generating TAG. The LPC produced by a PDAT reaction will add an

113

incremental flux to the PC acyl editing cycle (Figure 3-10). It is noteworthy that
the stereochemical analysis of TAG from soybean oil shows that the sn-1 and sn-
3 positions have quite similar acyl compositions. Such observations have lead to
speculations that the enzymes responsible for TAG synthesis are quite non-
specific. However, at least in soybean, there is the coincidental summation of
two routes with distinct specificities to give the overall composition. PC has long
been considered the site of FA desaturation. In developing soybeans acyl editing,
DAG-PC-DAG interconversion and a specific TAG synthesis system all contribute

to the biosynthesis of TAG containing PUFA at higher levels than found in PC.

Linking the Model to Questions of Biochemistry and Cell Biology

Acyl Editing: The LPCAT reactions which are hypothesized to produce the acyl
editing measured in pea leaves [162] and soybean embryos was first suggested
as a mechanism to allow enrichment of PUFA for TAG synthesis in oilseeds.
Subsequently, an in vitro mechanism, acyl exchange, a reversal of the LPCAT
reaction, was proposed as a mechanism for acyl editing. It was demonstrated
qualitatively by feeding free fatty acids to seeds [169]. In this study acyl editing is
demonstrated as a major flux for endogenously synthesized fatty acids. Our
previous identification of acyl editing in pea leaves and now in developing
soybeans suggests that PC acyl editing may be a ubiquitous part of plant lipid
metabolism. TAG synthesis in the cell must accommodate acyl editing, and

indeed TAG composition may be significantly modified by its action. However,

114

there are likely to be more fundamental reasons for acyl editing mechanisms.
First, it may be important in membrane dynamics and homeostasis. Control of the
low level of lyso-PC may be critically important due to its detergent effects on
membranes. Second, during periods of high fatty acid synthesis it ensures that
levels of saturated-oleoyl and dioleoyl molecular species of membrane
phospholipids (which may affect membrane fluidity, especially in the cold [174])
are kept low. Third, a high cycling rate allows for rapid metabolic changes in
times of stress. Thus there is an advantage of adaptability for membrane
biogenesis. Fourth, cycles in biology add robustness to metabolic networks. Fifth,
PC is appears to be the acyl flux bank of the plant cell, rather than other
phospholipids such as PE or Pl, which have more important and specific
functions. The idea that a PC:Iyso-PC cycle is the principle acyl acceptor system
in the cytosol of the plant cell is reinforced by the metabolic analysis we
performed on the fatB mutant, where additional acyl flux was directed into PC
and PC acyl cycling increased [161]. And finally, acyl editing may allow the
removal of oxidized acyl groups in membrane lipids.

ln seed, as in leaf [162], PC editing is an order of magnitude greater than
PE editing. Also, seed acyl editing of PC at the sn-2 position is particularly
dominant over that at the sn-1 position. This coincides with in vitro
measurements of acyl exchange in developing seeds, where over 90% of the
measured exchange is at the sn-2 position [143]. Assays of lyso-PL
acyltransferase activity in microsomes developing soybean cotyledons with

exogenous acceptor show a higher Vmax for LPEAT than LPCAT, but with

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endogenous lyso-Iipid acceptors the LPCAT activity was significantly higher
[175]. Thus further experiments are required to reveal whether lyso—PC
generation is by way of a phospholipase A and/or a transacylase (eg. acyl

exchange by a reversal of LPCAT) mechanism.

DAG-PC Inter-Conversion and Localization of DAG pools: In leaf tissue DAG
is a very minor glycerolipid. From this observation alone it may be inferred that
CPT operates in the direction DAG —+ PC, with very little reverse reaction. As
DAG is a component of lipid signaling Cascade [3], cellular control of DAG is
likely to be tight. Thus our finding that there are at least two, and probably three,
kinetically distinct DAG pools is hardly surprising. CPT would certainly be
associated with the de novo DAG synthesis pool. Whether the reverse CPT
reaction would also produce the DAG pool required for TAG synthesis is
questionable. If so, it may be another gene product or post-translationally
modified enzyme. In order for CPT to provide a flux running in the reverse
direction from that associated with de novo glycerolipid synthesis would require a
CPT with a locally different equilibrium position and /or a DAG removal
mechanism such a phase partitioning to another membrane domain. It is also
possible that a phospholipase C like activity provides the function of generating a
bulk DAG pool for TAG synthesis from bulk PC. Thus there is no requirement for
a rapidly reversible DAG-PC inter-conversion.

The identification of distinct kinetic DAG pools raises the question of

localization. In a comparative study of CPT and DGAT in microsomes from

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several different seeds [16] found little DAG selectivity for either enzyme. They
concluded that the exclusion of unusual fatty acids from membrane lipids was not
achieved on the basis of CPT or DGAT specificities and postulated distinctly
separate pools of DAG as an explanation. The concept of different
endomembrane domains is well known. However, the location of the de novo
DAG pool is unclear. This DAG pool may feed into a small, discrete PC pool, or
into the bulk PC pool. If the former is the case, then TAG synthesis may occur
from the bulk pool (which may itself have a more complex structure than we can
observe here). However, if the latter is the case, then TAG synthesis will occur
from a sub-domain. Or both DAG pools could each represent separate ER sub-
domains, connected by the bulk PC pool. However, the bulk DAG pool is most
likely associated with the oil bodies. It is a reasonable hypothesis that DAG is

also phase partitioned along with TAG into the oil bodies during their biogenesis.

TAG Synthesis: Despite the identification of three classes of genes that
encode for enzymes of TAG synthesis in plants, and the observation that DGAT1
and DGAT2 proteins from tung seeds, when tagged, localize to different ER
domains in tobacco and onion [133], the identification of kinetically distinct TAG
synthesis routes with a preference for saturated or polyunsaturated FA was
unexpected. It raises questions about acyltransferase genes, enzyme specificity
and acyl-CoA channeling.

Considering the synthesis of TAG highly enriched with saturates at the sn-

3 position, this is presumably a DGAT activity, and not PDAT, as it utilizes

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nascent fatty acids immediately. Furthermore, PDAT would transfer unsaturates
from the sn-2 position of PC. However, DGAT1 does not seem strong
candidates for this activity. The dgat1 mutant of Arabidopsis has a slight increase
in total saturates in TAG, with the fraction in the sn-3 position decreasing only
slightly [176]. DGAT2 has been implicated in the production of TAG containing
high amounts of novel unsaturated fatty acids, and on this basis also does not
seem to be a good candidate. Perhaps another DGAT is responsible? This
DGAT may have a very strong selectivity for saturates if it utilizes the mixed acyl-
CoA pool. A caveat concerns acyl group channeling via ACBP. The numerous
acyl-CoA binding proteins (ACBP) in plants [108-111, 113] may be involved in
shuttling acyl groups to different fates. Selective transfer of newly synthesized
saturated acyl groups to DGAT via a specific ACBP may have produced the
observed kinetics.

Considering the synthesis of TAG highly enriched with PUFA at the sn-3
position, including tri-PUFA TAG species, DGAT and PDAT mechanisms may be
invoked. Both are shown in Figure 3-10. TAG synthesis via various transacylases
of the PDAT family would transfer a sn-2 PUFA from PC to sn-3 DAG producing
TAG and lyso-PC [119, 120, 177]. Thus PDAT would leave a footprint
indistinguishable from sn-2 acyl editing. However, it cannot be the major
producer of lyso—PC for acyl editing because it would not allow recycling of FA to
feed de novo glycerolipid synthesis. For a putative DGAT to be involved, it would
either have to have a strong preference for PUFA acyl-CoAs, or such molecules

would have to be preferentially delivered to the site of the enzyme(s). It is

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noteworthy that highly sn-2 labeled PC from acyl editing is not immediately
available for conversion to DAG and then to TAG, otherwise TAG would contain
more sn-Z label at early time points. A simple explanation for this is that the bulk

PC pool that fills from acyl editing, creating a kinetic delay in its utilization.

3.5 Conclusion

Through in vivo labeling experiments we have shown that an acyl editing
cycle and generation of multiple sub pools of DAG are important for the synthesis
of TAG in developing soybeans. Similar research in the leaves of peas suggests
that acyl editing may be a ubiquitous part of plant acyl lipid metabolism. The
individual enzymes/genes that are involved in these processes are unknown and
thus represent an important area for further research on plant oil synthesis. As
there are many unknown genes in Arabidopsis annotated as acyl transferases
[134], further analysis of the acyl editing mechanism to determine if lipases or
transacylases are involved may be very useful for final identification of the
enzymes involved. Ultimate identification of acyl editing enzymes may require
analysis of mutants. The model of newly synthesized acyl group fluxes may be
useful in the interpretation of changes in lipid quantities or FA compositions
identified during mutant screens or in targeted knockouts. It is also possible that
not all the gene products/routes in the metabolism scheme have been identified.
In this context, the use of quantitative metabolic analysis taking into account acyl
editing with mutant lines may provide further clues to new genes with overlapping

roles in the processes. The fact that soybean oil is a major worldwide source of

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vegetable oil, with genetically engineered oil compositions in commercial
production and under development, adds a biotechnology dimension to this
study. Our identification of the major flux reactions of acyl groups from synthesis
in the plastid to accumulation in TAG may allow a more directed approach toward
identifying enzymes that might be useful in the engineering of TAG. The
enzymatic reactions involved in acyl editing may also be important for transferring
unusual FA from their site of synthesis on sn-2 PC to the three backbone
locations of TAG. The synthesis of TAG and phospholipids must be intricately
coordinated as both products require the synthesis of DAG. Production of
different lipids from DAG may be controlled by utilizing multiple DAG pools in
different locations. Identification of the sites and enzymes in each location may
allow more efficient engineering of novel lipid metabolizing enzymes to their site
of action in oilseed crop plants. Thus, a better understanding of the pathways of
TAG biosynthesis including acyl edting and DAG production in developing
soybeans may aid future efforts to engineer soybean oil with improved or novel

properties.

3.6 Materials and methods

Plant material-Developing soybean (Glycine max) embryos were
harvested from plants grown in the greenhouse at ~24° C, supplemented with
lights to maintain a 15 hr day. Embryos were harvested and cultured in media
containing the carbohydrates, amino acids, inorganic salts and light conditions

required for embryo growth to mimic in planta development as demonstrated

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previously [178]. Harvested embryos (~21 mg dry weight/embryo for [“Clacetate
labeling and ~15 mg dry weight/embryo for [14C]glycerol labeling) were
equilibrated in culture conditions for 2.5 days before starting labeling
experiments.

RadiochemicaIs—[1-“C]Acetic acid, sodium salt (specific activity 50
mCi/mmol), and [“C(U)]glycerol (specific activity 150 mCi/mmol) were from

American Radiolabeled Chemicals, Inc. (St. Louis, MO).

[ 1“C]acetate and [14C]glycerol Iabeling- Pre-cultured embryos were moved
to a single beaker containing fresh culture media plus radioactive substrate
[“C]acetate (1 mM) or [14C]glycerol (0.5 mM) to start the labeling reaction. The
media volume was just enough to cover all the embryos and was gently shaken
in a water bath at 27° C under 30-40 pmol rn'2 s" of white light. At each time
point the labeling reaction was quenched by transfering three embryos (four
during [14C]glycerol labeling) to 6 ml of 85° C isopropanol for 10 min.

General methods- Lipids were extracted from hot isopropanol quenched
tissue with hexane/isopropanol [155] after homogenization with a mortar and
pestle. Radioactivity in the total lipid samples, eluted lipids or organic and
aqueous phases recovered from transmethylation was quantified by liquid
scintillation counting (Beckman Instrument lnc., Fullerton, CA), while radioactivity
on TLC plates was visualized and quantified by electronic radiography (Packard
Instrument Co., Meriden, CT). AgNO3-TLC plates were prepared by impregnating
Partisil® K6 silica gel 60 A TLC plates (Whatman, Maidstone, UK) with 10%

AgN03 in acetonitrile (wlv), drying in air and activating at 110 °C for 5 min.

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Unlabeled fatty acid methyl esters (FAMEs) were quantified by gas
chromatography (GC) using a flame ionization detection and a 08-23 capillary
column (30 m length x 0.25 mm inner diameter, 0.25 p film thickness; J&W). For
preparative TLC all solvents contained 0.01 % butylated hydroxyl-toluene
antioxidant.

Polar lipid class analysis- polar lipid separation by TLC, recovery of polar
lipids from TLC plates, PC molecular species separation, and PC FA positional
localization were done as reported previously [162].

Neutral lipid class analysis-DAG and TAG were isolated by first acetylating
an aliquot of the total lipids in acetic anhydridelpyridine, (3:2, v/v) and then
separating the 1,2-diacyl-3-acetyl-glycerols (acetylated DAG) from TAG by silica
TLC developed in hexane/diethyl ether/acetic acid (70:30:1, v/v/v). Individual
lipids were eluted from TLC silica by chloroforrnlmethanol (4:1, v/v). Methanol
and 0.88% aq. KCl were added to give chloroform/methanol/water ratios of 2:1:1,
(v/v/v), resulting in a phase separation. The aqueous phase was back extracted
with chloroform and lipids were recovered from the combined chloroform phases.

Radiolabeled acyl group analysis-FAM E were prepared from total lipid
extracts by heating to 80° C in 5% H2804 in methanol for 60 min. The organic
soluble FAME were separated from the aqueous soluble compounds by the
addition of water and hexane. FAME were prepared from purified individual lipids
by base-catalyzed transmethylation [158]. For [“Clglycerol labeled lipids the
proportion of label in the acyl groups versus the backbone/head group was

determined by transmethylation [158] and scintillation counting of the separated

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organic and aqueous phases. Aqueous phase radioactivity from PC was
determined to be in the glycerol backbone and not the choline moiety previously
[162]. [14C]FAME compositions were determined by separation based on the
number of double bonds by AgNOa-TLC, the plates being developed to % height
with hexane/diethyl ether (1 :1, vlv), then fully with hexane/diethyl ether (9:1, vlv).
Saturated FAME were further separated by removal from AgNO3-TLC with
CHClalMeOH (2:1, v/v) and separated on KC18 reversed phase silica gel 60 A
TLC plates (Whatman, Maidstone, UK) by development in

acetonitrile/methanol/water (65/35/0.5, vlv/v).

Molecular species separation of radiolabeled DAG and TA G- Acetylated
DAG isolated from TLC was separated into molecular species based on the
number of double bonds by AgNOa-TLC [159], using a triple development (‘/2
then ’/4 development in chloroform/methanol (96:4, vlv), then fully in
chloroform/methanol (99:1, vlv)). When necessary the saturate/diene and
monoene/monoene molecular species were eluted from the silica [159] together
and further separated on 15% AgNOa-TLC plates at -20° C with the same triple
development as above. TAG molecular species were separated by AgNOa-TLC
in two batches both at -20° C. The more saturated species (0-4 double bonds)
were separated by triple development (first 60% then 75% development in
chloroform/methanol (96:4, v/v) then full in chloroform/methanol (98:2, vlv)). The
more unsaturated species (5-9 double bonds) were separated by triple
development (60%, 75%, then full development in chloroform/methanol 94:6,

v/v). When necessary the saturate/monoene/monoene and

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saturate/saturate/diene molecular species were eluted together and separated by
AgN03 TLC at -20° C by triple development in chloroform/methanol (99:1, vlv).
The proportion of radioactivity in each molecular species band for DAG and TAG
was determined by electronic autoradiography, then each band was eluted, the
recovered lipids transmethylated, and the [‘4C]FAME analyzed by AgNOa-TLC,
as described above.

Regiochemistry of radiolabeled DAG and TAG-Collected acetylated DAG
and TAG were digested with porcine pancreatic lipase (sigma) [159]. The
digestion products were separated on silica TLC developed with hexane/diethyl
ether/acetic acid (70:30:2, v/v/v). The amount of radioactivity in MAG, DAG, free
fatty acids, and TAG was quantified by electrionic autoradiography, then each
band was eluted, the recovered lipids transmethylated, and the [14C]FAME
analyzed by AgNOa-TLC, as described above.

Endogenous molecular species compositions- Total lipids from four
soybean embryos (approx 13-14 mg dry weight), cultured under identical
conditions to those used for labeling, were extracted according to Hara and
Radin (1978). TAG concentrations were measured directly using electrospray
ionization mass-spectrometry (ESI-MS). DAG and PC were converted to their
sn-1,2-diacyl-3-acetyl-glycerol (ac-TAG) derivatives by digestion with pancreatic
lipase C (for PC) followed by acetylation prior to ESI-MS analysis.

ESI-MS in positive ion mode was performed by direct infusion with a Shimadzu
(Columbia, MD) SlL-5000 autosampler into a Waters (Milford, MA) Quattro micro

mass spectrometer. 10pl of sample in toluene was introduced to the electrospray

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source by flow injection analysis conveyed by a 65:32:3
chloroform:methanol:100mM ammonium acetate solution at a flow rate of 0.1
ml/min. The capillary, extractor, and cone voltages were 3.2 W, 2.0 V, and 40 V
respectively. The source and desolvation temperatures were 110 and 350 °C,
respectively. The desolvation gas flow rate was 400 l/hr. Mass spectra were
collected for 2 min; the m/z range scanned in the MS measurements was from
500 to 1000 (1 sec/scan) and in the MS2 measurements from 20 to the mass of
the parent ion. Collision-induced dissociation used argon as the collision gas (2
x 10'3 mbar) with the collision energy set at 22 eV. Mass spectra data was
acquired with MassLynx 4.0 software; [T AG-NH4]+ ion peaks were smoothed and
integrated using QuanLynx software.

In order to correct for the effect of the number of acyl chain carbons and
double bonds on the signal strength [179], TAG standards with varying acyl chain
length and number of double bonds were analyzed at different concentrations.
After correcting for natural isotope abundance effects, the ion peak intensities for
each TAG standard were normalized to the internal standard (10uM
triheptadecanoin). The normalized peak intensity was plotted against TAG
concentration. The slope of this standard curve was determined for each TAG
standard. Multiple linear regression was then used to create a correction
function relating the slope of the standard curve to the number of acyl chain
carbons and double bonds.

To determine the concentration of soybean TAG or ac-TAG molecular

species, the ion peak intensities were deisotoped and corrected for natural

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isotope abundance and then normalized to the 10uM triheptadecanoin internal
standard. The correction function for acyl chain carbons and double bonds was
then applied to determine the absolute concentration of each TAG species.
When necessary, ESI-MS2 was used to determine the relative abundance of
isomers with the same m/z value. The moles embryo" of each molecular species
measured for TAG, DAG, PC are in Appendix B Tables 6-1, 6-2, 6-3,

respectively.

3.7 Footnotes

Abbreviations: ACP, acyl carrier protein; CPT, CDP-choline:1,2-diacyl-sn-glycerol
cholinephosphotransferase ; DAG, sn-1,2-diacylglycerol; DGAT,
diacylglycerolzacyl-CoA sn-3 acyltransferase; ER, endoplasmic reticulum; FA,
fatty acid; G3P, glycerol-3-phosphate; LPCAT, lyso-phosphatidylcholine:acyl-
CoA acyltransferase; PC, phosphatidylcholine; PDAT,
(phospholipid:diacylglycerol acyltransferase); PE, phosphatidylethanolamine;

PUFA, polyunsaturated fatty acid; TAG, triacylglycerol.

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4 CONCLUSION

The two pathway hypothesis of eukaryotic and prokaryotic glycerolipid
assembly (Figure 1-1) was first proposed over 25 years ago based on radiotracer
studies of plant tissue and isolated organelles [71, 84]. The age of genetics and
molecular biology further confirmed this seminal work [85]. Since that time
identification of PC acyl exchange reactions [140], hydrolytic turnover of PC acyl
groups [74], channeling of nascent FA from the plastid into PC [79], and the
mixing of nascent FA with previously synthesized FA during PC synthesis [149]
have suggested that the steps of the eukaryotic pathway may be further refined.
A re-examination of plant glycerolipid synthesis requires that new investigations
circumvent the limitations of previous studies. Therefore, very rapid (as short as
1 min) in vivo kinetic labeling studies were conducted with [14C]acetate and
[14C]glycerol. Controls with [1401002 and media containing additional carbon
sources mimicking in planta conditions were utilized to limit changes in
metabolism due to wounding or utilization of exogenously applied metabolic
tracer compounds, respectively. Additionally, analysis included not only
precursor-product relationships between labeled lipid classes, but also analysis
of the regiochemistry and molecular species of lipids from the initial incorporation

of labeled nascent FA and glycerol backbones into eukaryotic glycerolipids.

4.1 Acyl editing conclusions

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Expanding pea leaves and developing soybean embryos were utilized for
in vivo of labeling of the initial events of eukaryotic membrane and storage lipid
synthesis. To ensure the in vivo relevance of the metabolic labeling data from
exogenously applied [14C]acetate or [14C]glycerol to excised pea leaves, intact
planted pea seedlings growing in pots were also labeled with 1“00;. The results
with the 1“C02 precursor confirmed the conclusions based on exogenously
applied [14C]acetate and [“Clglycerol metabolic tracers. Likewise, developing
soybean embryos were pre-cultured for three days in media that allows excised
embryos to mimic in planta growth. Trace amounts of [14C]acetate or [“Clglycerol
were added to the culture media for labeling experiments which allowed
exogenous tracers to be applied without wounding the embryos or changing the
carbon sources used for heterotrophic growth and represent methodological
improvements over most previous studies that utilized seed slices and a single
radiolabeled carbon source. These controls allow us to be more confident that
the labeling results reflect the reactions of central lipid metabolism that occur in

viva.

Kinetic analysis of rapid [14C]acetate and [“nglycerol incorporation into
lipids allowed precursor-product relationships to be established for newly
synthesized FA and glycerol backbone incorporation into eukaryotic glycerolipids,
respectively. Stereochemical analysis of [“Cjacetate labeled FA in different lipids
indicated that different mechanisms are employed for incorporation of nascent
FA into PC, DAG and TAG. Molecular species separation of labeled lipids

allowed determination of: (1) pre-existing FA that occur in the same molecule as

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the [14C]acetate labeled newly synthesized FA; (2) the FA esterifed to
[14C]glycerol during de novo glycerolipid synthesis. Finally, endogenous
compositions of different lipid molecular species were determined utilizing ESI-
MS/MS techniques which demonstrated differences between the steady state
molecular species and those of de novo synthesized lipids. Together these
analyses of de novo synthesized glycerolipids and incorporation of newly
synthesized FA into glycerolipids were used to develop a revised model of

eukaryotic membrane and storage lipid synthesis in plant leaves and seeds.

The pathway of plant eukaryotic membrane lipid synthesis shown in most
textbooks and reviews [34, 85] predicts that newly synthesized acyl groups
exported from the plastid are first esterified to G3P producing initial PA molecular
species of 18:1/18:1 and 16:0/18:1. PA is then converted to PC the major site of
acyl desaturation, and that desaturated PC may be in equilibrium with DAG to
produce other desaturated membrane or storage lipids (Figures 1-1 and 2-9
model 1). However, rapid labeling of the initial steps of eukaryotic lipid synthesis
in pea leaves indicated that this simple model is incorrect and the major flux of
newly synthesized FA into eukaryotic glycerolipids is not by G3P acylation but
through acyl editing of PC, producing initial molecular species of PC containing
one newly synthesized FA and one previously synthesized FA. Acyl editing
involves a cycle of deacylation and reacylation of PC that by it self does not lead
to net glycerolipid synthesis and produces an acyl group and lyso-PC as
intermediates. Two models of acyl editing were possible in pea leaves (Figure 2-

9, model 2 and 3). (1) Plastid FA export is tied to acyl editing in a mole for mole

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fashion, with newly synthesized FA directly esterified to the lyso-PC intermediate
and the released acyl groups utilized for de novo glycerolipid synthesis (Figure 2-
9 model 2). (2) The acyl editing cycle is at a much higher rate than nascent FA
synthesis such that a mixed acyl group pool is mostly FA recycled from acyl
editing, and nascent FA in the mixed pool are rapidly incorporated into PC
through the high flux of acyl editing (Figure 2-9 model 3). Labeling of initial
glycerolipid synthesis in developing soybean embryos also indicated acyl editing
as the major pathway for incorporation of newly synthesized FA into extra-
plastidial glycerolipids. However, only the high acyl editing flux pathway is
consistent with the soybean labeling results (Figure 3—10). Therefore,
incorporation of nascent FA into glycerolipids through a high flux of acyl editing is

probably the more accurate model in pea leaves and other plants.

Research in this thesis demonstrates that incorporation of newly
synthesized FA into extra-plastidial glycerolipids occurs through acyl editing in
leaves and developing embryos from peas and soybeans, respectively. Pea and
soybean are both “18:3” plants [82] and are both in the legume family Fabaceae
of the order Fabales. Similar PC molecular species labeling results have also
been observed in the leaves of the “16:3” plant Brassica napus [149] (order
Capparales, family Brassicaceae) suggesting that acyl editing may be a
ubiquitous part of plant leaf lipid metabolism. Additionally,
lysophosphatidylcholine acyltransferase enzymatic activity has been found in a

wide variety of plant leaf [91, 94, 143, 145] and oilseed tissues [130, 140, 142,

130

144, 163] further suggesting that acyl editing may be a central part of plant lipid

metabolism.

In soybean embryos the labeling experiments and endogenous PC, DAG
and TAG molecular species compositions suggested that TAG was produced
primarily from a PC derived DAG moiety generated by the removal of the
phosphocholine headgroup (Figure 3-10). A traditional Kennedy pathway
involving three consecutive backbone acylations was either very minor or
nonexistent. The utilization of the PC DAG moiety for TAG synthesis has been
suggested by in vitro assays of safflower [118] and sunflower [173] that
demonstrated the reversibility of CDP-choline:diacylglycerol
cholinephosphotransferase. In vivo pulse-chase backbone labeling demonstrated
the utilization of PC backbone for TAG synthesis in linseed [117, 127] and
safflower [168]. In each of these studies it is assumed that the DAG pool is in
equilibrium with the DAG moiety of PC and that this equilibrated DAG pool is
utilized for TAG synthesis. However, in the soybean embryo labeling reported in
chapter three the positional localization of nascent [14C]FA incorporation into PC
(>86% sn-2), DAG (equal sn-1 and sn-2) and TAG (>98% sn-3) suggested that
newly synthesized DAG and acyl edited PC are not in rapid equilibrium and

neither is rapidly used for TAG synthesis.

Safflower (Carthamus tincton’us L.) oil contains over 75% PUFA and
avocado (Persea amen’cana) oil less than 15%. The major initial product of
[“Clglycerol labeling in safflower cotyledons was PC yet it was TAG in avocado

mesocarp [168]. Thus, it has been suggested that utilization of the PC-DAG

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moiety for TAG synthesis may only take place in oilseeds that utilize PC for FA
modification and accumulate large amount of desaturated or unusual FA in TAG.
However, the unusual FA petroselinic acid (18:1cisA6) produced in coriander and
carrot endosperm [152] and the unusual FA 16:1-A6 produced in Thunbergia
alata [153] are both synthesized in the plastid, yet still flux through PC prior to
accumulation into TAG. If the PC-DAG moiety containing the unusual FA is used
for TAG synthesis it would imply that FA modification on PC is not. required for
conversion of PC to TAG. Additionally, in Brassica napus which contains 30%
PUFA in seed TAG, four hour labeling of developing embryos with acetate and
glycerol led the authors to suggest the Kennedy pathway is the major route to
TAG synthesis [180, 181]. lt is clear that multiple pathways may be involved in
TAG synthesis in different plant species and that utilization of the PC-DAG
moiety for TAG is a mechanism that may or may not be used by various oilseed
plants. However, the variability in the techniques of labeling studies in different
plants (variability of oil tissue and developmental stage, wounding effects of
excised tissue, different labeling time periods, FA or backbone labeling only,
labeling with or without stereochemical and/or molecular species analyses)
makes determinations of the sequence of reactions involved in the TAG
synthesis pathway difficult to deduce from most previous studies of oil
biosynthesis. The soybean inital glycerolipid synthesis studies presented here
demonstrate that very short time point labeling kinetics and analyses of both the
acyl and backbone components of TAG synthesis through regiochemistry and

molecular species analyses are essential to generate an accurate model of TAG

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synthesis. The soybean TAG synthesis model indicates that the PC-DAG moiety
utilized by TAG synthesis is not the same as the DAG produced by de novo
glycerolipid synthesis and therefore a direct Kennedy pathway may not be
required in oilseeds (Figure 3-10). Whether a direct Kennedy pathway is a more
major TAG biosynthesis route in other oilseeds will require additional in depth
investigations of TAG synthesis in different plant species.

4.2 Additional significance of acyl editing and PC derived TAG
synthesis

The process of acyl editing described in this thesis has possible
significance for understanding additional areas of basic plant lipid biology. The
mechanism of fatty acid export from the plastid for utilization by the eukaryotic
pathway acyltransferases has yet to be elucidated. Experiments with isolated pea
chloroplasts indicate that newly synthesized FA are channeled into PC, possibly
on the outer chloroplast envelope [79]. Isolated chloroplasts cannot synthesize
PC de novo; therefore acyl editing is very likely involved in incorporation of newly
synthesized FA into PC at the plastid envelope. Lateral diffusion of PC through
the PLAM [94, 95] to the cellular endomembrane system would allow a
mechanism of plastid FA export that would not require acyl-CoA to pass through

the cytosol.

Acyl editing may also be involved in ensuring that newly synthesized
glycerolipids do not disturb the integrity of cellular membranes. PUFA containing
lipids are required for proper membrane physiology and function, especially at

low temperatures [174, 182]. Desaturation of 18:1 to 18:2 and 18:3 lagged

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significantly behind the rate of 18:1 production in pea and soybean (Figures 2-2
and 3-3, respectively). However, acyl editing produced initial molecular species
of PC with mixtures of previously synthesized PUFA with newly synthesized
saturated or monounsaturated FA (Figures 2-5 and 3-5). The mixed FA
molecular species may benefit the plant by not allowing accumulation of relatively
saturated lipid molecular species produced from only newly synthesized FA (e.g.
16:0/18:1, 18:1/18:1). Without acyl editing, plant tissue that is rapidly producing
lipids (e.g. expanding leaves or developing oilseeds) may accumulate relatively
saturated molecular species in biosynthetic regions of the ER membrane that

could affect the membrane physical properties and function.

The supply of FA has been shown to be one of the limiting factors of TAG
accumulation in oilseeds [183]. High levels of externally supplied FA produces a
feedback control on fatty acid synthesis through modulation of plastidic ACCase
activity [58]. In soybean embryos most newly synthesized FA are initially
incorporated into glycerolipids through acyl editing (Figure 3-10). Acyl editing
enzymes may be able to regulate or sense free FA levels which could be
involved in the signaling to modify ACCase activity. In plant species that
accumulate unusual FA in TAG the amount of unusual FA in membrane lipids is
typically very minor. However, in transgenic plants producing unusual FA the
membrane lipids can sometimes accumulate more unusual FA than TAG [128].
An acyl editing scheme that selectively removes modified FA from PC for TAG
synthesis and incorporates newly synthesized FA into PC to be further modified

may be involved in plant species that naturally produce unusual FA. Likewise, in

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transgenic plants that express the biosynthetic enzymes for unusual FA but do
not express unusual FA selective acyl editing enzymes, the unusual FA may not
be efficiently removed from the site of synthesis on PC. A similar mechanism
selective for PUFA may be involved in plant species that accumulate large

amounts of PUFA in TAG.

The kinetic modeling of [14C]acetate and [14C]glycerol labeling of soybeans
(Figures 3-9 and 3-10) suggested at least three DAG sub-pools that are
kinetically distinct. The individual DAG sub-pools may suggest separate sub-
cellular localizations that are related to separate functions. Soybean TAG
synthesis primarily utilized the bulk DAG produced from bulk PC. The PC
headgroup removal to generate the bulk DAG may be localized to an ER region
associated with a developing oil body for TAG synthesis. A possible function of
utilizing bulk PC in a separate location for TAG synthesis may be to ensure the
use of desaturated PC (and thus DAG) and not de novo synthesized DAG/PC or
acyl edited PC which would contain at least one acyl group that was not
desaturated or otherwise modified. Utilization of PC that diffuses laterally through
the membrane from the site of desaturation to the proximity of the oil body may
be a mechanism to separate the steady-state desaturated bulk PClDAG for
polyunsaturated TAG synthesis without the need for a Kennedy pathway that

utilizes a high PUFA acyl-CoA pool or enzymes that are selective for PUFA.

4.3 Future research on acyl editing and TAG synthesis pathways

135

The next logical step in the investigation of lipid synthesis through acyl
editing is to identify the enzymes/genes responsible for the deacylation and
reacylation of both the sn-1 and sn-2 position of PC. The acyl editing model
presented (Figure 3-10) utilizes a sn-1/sn-2 lyso-PC intermediate. Therefore an
LPCAT enzyme is probably involved in the lyso-PC reacylation and possibly the
PC deacylation by the reverse action of LPCAT [140]. Arabidopsis thaliana
contains 11 gene candidates for lyso-phospholipid acyltransferases [134], and
recently two Arabidopsis genes have been expressed in yeast and the activities
demonstrated to be broad specificity lyso-phospholipid acyltransferases with the
highest activity for lyso—PC [184]. Alternatively phospholipases may be involved
in the deacylation of PC of which Arabidopsis has at least one PLA1 and six PLA2
gene candidates [134]. Assuming the knockout of acyl editing enzymes are not
lethal, the redundancy in Arabidopsis suggests that a fonrvard genetic screen is
unlikely to yield which gene products are involved in acyl editing. A reverse
genetic approach may be more suitable; however redundancy may also be a
problem and may require testing many combinations of gene knockouts.
Additionally, the labeling experiments and analysis methods presented in this
thesis to investigate acyl editing are laborious and time consuming and thus are
not suitable for screening of many different plants. A possible preliminary labeling
experiment to analyze reverse genetic candidates would be to rapidly pulse label
fatty acid synthesis with [“Clacetate and measure the ratio of [“ClDAG to

[“0ch. If acyl editing is reduced in the mutants the initial labeling ratio should

136

increase. Further analysis will be required to determine if a change in DAG and

PC labeling is actually due to a change in acyl editing.

A possible non metabolic labeling based analysis method to analyze
reverse genetic acyl editing mutant candidates would be to use mass
spectrometry to quantify lipid molecular species. If acyl editing is knocked out or
significantly reduced by a mutation then more nascent FA might enter de novo
glycerolipid synthesis and thus initially produce larger amounts of 18:1/18:1 &
16:0/18:1molecular species in PA, DAG and PC. Additionally, levels of lipid
molecular species that have nascent FA/previously synthesized FA composition
(e.g. 18:1/18:3) produced from acyl editing might be reduced. However,
desaturation of the initially produced molecular species would presumably not be
affected and any change in molecular species abundances may be small and
transient. Therefore to have the highest probability of success, tissue that is

rapidly producing lipids such as young expanding leaves should be utilized.

Since there are many possible gene candidates for acyl editing enzymes
further analysis of the mechanism of acyl editing to determine whether lipases or
transacylases are involved could narrow down the number of possibilities. In this
regard incorporation of oxygen-18 into the carbonyl oxygen of newly synthesized
FA through [180]acetate or H2180 labeling and measuring 180 content with mass
spectrometry would allow for measurements of FA hydrolysis. Each time a FA is
hydrolyzed by a lipase and re-esterified there is 50% exchange of the carbonyl
oxygen with the cellular H20 [74]. If acyl editing proceeds by way of a lipase then

there would be different amounts of 18O in the acyl-CoA pool and PC. However, if

137

acyl groups are removed from PC by a transacylation reaction such as reverse
LPCAT then 180 content of the acyl-CoA pool and PC should be the same.
Similar experiments utilizing [‘3O]phosphate may be able to determine if PC
headgroup removal to produced DAG for TAG synthesis is by way of a
phospholipase D and PA phosphatase (as suggested for eukaryotic galactolipid
DAG production [89, 90]) or reverse action of cholinephosphotransferase [16,

117,118].

Genetic approaches to identify the enzymes of acyl editing would be most
successful in the model species Arabidopsis thaliana, which has a fully
sequenced genome and many genetic/molecular biology tools available.
However, acyl editing was demonstrated in pea and soybean and not
Arabidopsis. Thus identification of the acyl editing enzymes from pea and
soybean by enzyme purification may be a logical approach. In this regard LPCAT
activity has been measured in microsomes of many different leaf [91, 94, 143,
145] and oilseed tissue [130, 140, 142, 144, 163], which suggest that at least
partial purification of enzymatic activity is possible. Preliminary experiments
suggest that an acyl editing activity is associated with isolated pea leaf
chloroplasts (Appendix C). Interestingly in five minutes of [14C]acetate labeling of
pea chloroplasts only nascent [14C]18:1 was incorporated into PC and was
located ~93% at the sn—2 position of PC. Similarly most in vitro LPCAT activities
measured in microsomes were also mostly sn—2 specific. Therefore the
substantial sn-1 acyl editing activity identified by in vivo labeling in pea leaves

(Figure 2-6) may be a soluble activity that is lost by isolation of chloroplasts or

138

microsomes alone. If an enzyme is able to be purified the amino acid sequence
may be identified by Edman degradation or mass spectrometry based
sequencing. Once the protein/gene sequence is known genetic and molecular
biology techniques could be utilized to confirm its function by analysis of acyl

editing in knockout/knockdown mutants and over expressers.

The models of pea leaf and soybean acyl editing presented describe an
acyl editing cycle involving nascent saturated and monounsaturated FA
incorporation into the sn-1 and sn-2 position of PC. Some evidence in chapter
three suggests that within the acyl editing cycle and glycerolipid synthesis there
could be different mechanisms involved in saturate and 18:1 incorporation into
glycerolipids which should be studied further. Soybean saturate labeling
increased with time in proportion to 18:1 in PC, while saturate labeling was
constant and in very different proportions to 18:1 in DAG and TAG (Figure 3-7B).
Additionally, the primarily sn-2 and 18:1 specific acyl editing in pea chloroplasts
(Appendix C) suggest that separate enzymatic activities and possibly locations
are involved with incorporation of nascent 18:1 into PC as compared to nascent
saturates in pea leaves. The high sn-3 saturate labeling of nascent FA
incorporation into TAG of developing soybeans may suggest that there is a
specific channeling of saturated FA to a DGAT. Release of newly synthesized
saturated FA and 18:1 from ACP at the termination of fatty acid synthesis is
carried out by two separate thioesterases, FATB and FATA respectively. The
Arabidopsis fatb mutant which reduces the amount of saturated FA exported

from the plastid [161, 185, 186] and increases total 18:1 plastid export and

139

breakdown may be useful for analysis of acyl editing mechanisms. If there is a
saturate specific acyl editing enzyme that is required for nascent saturated FA
incorporation into glycerolipids then a mutant should have a change in lipid
composition similar to fatb, and the double mutant lipid composition should be
similar to the single mutants. If the excess 18:1 exported from the plastid in the

. fatb mutant is incorporated into glycerolipids through acyl editing then the rate of
nascent FA incorporation into PC should increase. If acyl editing is required for
turnover and breakdown of the excess 18:1 in the fatb mutant, then B-oxidation in
an acyl editing and fatb double mutant might be reduced as compared to the fatb
mutant alone. Additionally, a fatb and 18:1 acyl editing mutant may accumulate

excess 18:1 containing PC molecular species as compared to wild type.

Soybean embryo fatty acid synthesis produced more newly synthesized
[“Clacetate labeled 18:0 than 16:0. However, the endogenous FA mass
composition contains more 16:0 than 18:0. The newly synthesized [“C]18:0 that
is initially more abundant than newly synthesized [“C]16:0 in soybean
glycerolipids may be a transient pool of 18:0 that is used for synthesis of other
products such as sphingolipids, cutin, or wax. Alternatively, 18:0 may be
desaturated while bound to PC or within the recycled acyl-CoA pool. The
plastidic stearoyl-ACP desaturase is considered the only known 18:0 desaturase
in plants [34]. Identification of an extra-plastidic stearate desaturase would be
novel to plant lipid metabolism. Further [14C]acetate soybean labeling with much

longer time points and/or pulse-chase [“Cjacetate labeling experiments may be

140

able to reveal the down steam products of the initially high 18:0 production for

additional investigation.

The identification of multiple sub-pools and DAG that are not in equilibrium
suggests that the enzymes producing each sub-pool are not in the same cellular
location. Even though we could not distinguish multiple pools of PC kinetically,
acyl edting may be at a separate location than de novo PC synthesis. Cell
biology localization techniques such as green fluorescent protein (GFP) fusions
of the enzymes for de novo glycerolipid synthesis, acyl editing, and TAG
synthesis could be employed to confirm the separate localization hypothesis. An
interesting facet of localization studies would be whether different isoforms of
enzymes localize to the same sub-cellular location. For instance if plastid
associated acyl editing is only sn-2 specific then a sn-2 LPCAT may be localized
to the PLAM region, but an sn-1 LPCAT that utilizes saturates may be soluble or
localized to an acyl-editing domain of the ER. A hypothesis as to why many
transgenic plants expressing FA modification enzymes do not accumulate the
same quantities of unusual FA in TAG as do the native species is that unknown
factors channel the unusual FA from site of synthesis to incorporation into TAG.
The GFP fusions used for localization studies could also be used for co-
lmmunoprecipitation studies to identify proteins interacting with the PC acyl
editing enzymes or the TAG synthesis DGATs and PDATs. Identification of the
enzymes required for incorporation of modified FA into TAG and the ability to

express them in the correct location to be efficiently utilized will greatly increase

141

our ability to generate transgenic crops that accumulate unusual FA of interest in

TAG.

This dissertation has demonstrated that FA incorporation into membrane
lipids and TAG involves large fluxes of acyl editing in and out of PC in pea leaves
and developing soybeans. The relation of acyl editing to previous studies suggest
that acyl editing may be involved in many different plant species and further
research may imply that it is a central part of plant glycerolipid metabolism. The
acyl editing models presented update the simple de novo glycerolipid synthesis
model found in most lipid biosynthesis reviews and textbooks. Further research
of the mechanisms of acyl editing and the enzymes/genes involved will deepen
the understanding of plant lipid synthesis and possibly increase our ability to
modify the incorporation of FA into plant lipids to suit our needs for food, fuel and

industrial applications.

142

5 APPENDIX A: CHAPTER 2 SUPPLEMENT

NOTE 1

Discussion: 18: 1/18:1 and 16:0/18:1 Are Not Initial Molecular Species of

Eukaryotic Glycerolipids

Rapidly expanding pea leaves at day 8 and day 9 produce about 2.2 and
0.6 pmol FAlhr/g fresh wt. respectively in the light, and contain 20-25 pmol FAlg
fresh wt. Although endogenous 16:0/18:1 and 18:1/18:1 molecular species of PA
and PC can arise by non-biosynthetic mechanisms, if we treat them as
essentially all biosynthetic, produced by de novo synthesis from glycerol-3-P, and
estimating their PA and PC levels at 0.002 and 0.2 mole % respectively, the
residence time for any labeled FA passing through these molecular species pools
is calculated as 0.7-3 sec and 1.1-5 minutes respectively. Thus, a pool of
16:0/18:1 and 18:1/18:1 molecular species of PC should have been easily

detected by our analysis.

143

SUPPLEMENTAL FIGURES

Figure 5-1 Supplemental Figures S1A-S1M: Lipidomics of pea leaves and
acetate labeled pea leaves.

Pea leaf lipid samples were analyzed by ESI-MS/MS by the Kansas Lipidomics
Research Center. Extraction of lipids was conducted by their standard
Arabidopsis leaf protocol (Web site: h_ttp:/lwww.k-state.ed u/ligid/lipidomics/leaf-
extraction.html). Data is represented as mol percent of total lipids measured.
Error bars are standard error, n = 5. Black bars: 8 day old pea leaves out and
quenched immediately. White bars: 8 day old pea leaves out into strips and
labeled with 1 mM cold (‘20) acetate for 5 min and quenched immediately. Mock
labeling conditions were the same as for [“Clacetate labeling. A-B: The sum of
total molecular species for each lipid class measured. C-M: Individual molecular
species for each lipid class measured. A: Digalactosyldiacylglycerol (DGDG),
Monogalactosyldiacylglycerol (MGDG), PC, PE, PG, phosphatidylinositol (Pl). B:
phosphatidylserine (PS), PA, Lyso-PC, Lyso-PE, Lyso-PG. C: DGDG. D: MGDG.
E: PC. F: PE. G: PG. H: Pl. l: PS. J: PA. K: Lyso—PC. L: Lyso-PE. M: Lyso-PG.

144

SlA: Total molecular species by lipid class

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Lipid class

145

SIB: Total molecular species by lipid class

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Lipid Class

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Molecular Species
147

SID: MGDG

 

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148

SIE: PC

 

 

 

 

 

 

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Molecular Species

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mol %

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Molecular Species

155

 

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Molecular Species

156

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Molecular Species

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157

 

Figure 5-2 Estimates of single and dual acyl-labeled PC molecular species
for Model 3 (Figure 2-9).

Calculation of PC molecular species containing one labeled acyl group
(PC[FA*FA]) or two labeled acyl groups (PC[FA*FA*]), based on x moles of
labeled fatty acid (FA*) and y moles of unlabeled fatty acid (FA) distributed
randomly between sn-1 and sn-2 positions. The distribution is calculated as
PC[FA*FA]/(PC[FA*FA*] + PC[FA*FA]). The small amount of labeled saturates
largely at the sn-1 position (Figure 2-6) will not greatly skew these results.

 

100

 

 

 

 

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158

6 APPENDIX B: CHAPTER 3 SUPPLEMENT
6.1 Supplement Figures for Chapter 3 Results

 

 

 

 

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Figure 6-1 [“C]Fatty acid composition of total [“C]acetate labeled soybean
lipids.

Total lipid FAMEs from each [14C]acetate labeling time point were produced by
acid catalyzed transmethylation. FAME were separated based on the number of
double bonds by AgNOa-TLC and based on chain length/unsaturation by RP-
TLC. In AgNO3-TLC the saturated FAME co-migrated and in RP-TLC 16:0/18:1
co-migrated. FAME bands that co-migrated with standards were quantified by
electronic autoradiography. The individual FA compositions were calculated
based on their relative quantities in each TLC system. Color labels: Black, 16:0;
hatched, 18:0; white, the sum of 20:0, 22:0, 24:0, 26:0; light gray, 18:1; dark
gray, 18:2.

159

 

Alpha oxidation of [“Clacetate labeled saturated FA

 

 

 

 

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Alpha oxidation substrate

Figure 6-2 Alpha oxidation of [“Clacetate labeled saturated FA from TAG
and PC.

Individual saturated FA acids from TAG and PC were subject to chemical alpha
oxidation. Each reaction is a mixture of FA from 20, 30, 120 & 360 min acetate
labeling. The reaction produced a series of FA each shortened by 10 from the
carboxyl end. Results are expressed as relative specific activity to that of the full
length reaction substrate. The X-axis groups each individual substrate reaction
products with decreasing chain length from left to right. Individual FA products
bar shading is in the key.

Method: Individual saturated FAME from TAG and PC were collected by first
AgNOa then reversed phase TLC (as in methods) and saponified to FA. 100 ug
of non labeled carrier FA was added to each reaction with the respective FA
chain length. The reaction was started by adding 10 mg of finely ground KMnO4
and 0.5 ml acetone to each dried FA sample and heated to 60° C. After six hours
remaining KMnO4 was reduced in the cooled sample by adding 1 ml of 0.5M
FeSO4 in 0.5 N H2804. FA were extracted with hexane and FAME generated by
reaction in 5% sulfuric acid in MeOH at 50° C for 30 min. FAME extracted by
hexane were split into two aliquots, mass quantified by GC-FID, and radioactivity
quantified by reversed phase TLC and electronic autoradiography.

160

 

 

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m

Figure 6-3 TLC image examples and flow chart for analysis of radiolabeled
glycerolipids.

A-E. Examples of the major TLC systems used during analysis of radiolabeled
glycerolipid classes, molecular species, and FA composition. Molecular species
are represented as a combination of two or three FA, total saturates, S;
monoenes (18:1), M; dienes (18:2), D; trienes (18:3), T. A, Neutral lipid class
TLC, pictured 6 min [14C]acetate. B, Polar lipid class TLC, pictured 6 min
[“Clacetate. C, TAG molecular species argentation TLC, pictured 30 min
[“Clglycerol. D, Molecular species separation of PC and DAG as 1,2-diacyl-3-
acetyl-glycerols, pictured 6 min [14C]acetate PC. D, FAME argentation TLC,
pictured FAMEs from different 6 min [“Clacetate PC molecular species.

161

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Figure 6-4 [“Clacetate labeled PC molecular species time course.

A-D, molecular species are represented as pairs of FA, with bar heights
represent % of species among the total. Bar shading represent % of [“ClFA
among species. In species with only one color it means that only that FA is
labeled the other FA within the molecular species is unlabeled. Bar shading: S,
total saturates, black bars; M, monoenes (18:1), white bars; D, dienes (18:2),
gray bars. A-D Time of [14C]acetate labeling: A, 2 min; B, 6 min; c, 10 min; 0, 30

min.

162

A; 6 min Figure 6-5 [“Clacetate
labeled DAG molecular

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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Figure 6-6 [14C]acetate
labeled TAG molecular
species time course.

A-C, molecular species
are represented as three
of FA, with bar heights
represent % of species
among the total. A-B,
bar shading represent %
of [“C]FA among
species. In species with
only one color it means
that only that FA is
labeled the other FA in
the molecular species is
unlabeled. Bar shading:
S, total saturates, black
bars; M, monoenes
(18:1), white bars; D,
dienes (18:2), gray bars.
A 6 min labeling; B, 10
min labeling. C, Only the
total [“C] label per
molecular species is
reported.

A: PC

 

  
 

   

 

 

      

 

  
 

   

    

40
35 -
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SS SM SD ST MM MD MT DD DT ‘lT
Molecular species
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30 -
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SS SM SD ST MM MD MT DD DT TT

Molecular species

Figure 6-7 [“Clglycerol labeled molecular species time course of PC and
DAG.

A-B, molecular species are represented as a combination of two FA. Total
saturates, S; monoenes (18:1), M; dienes (18:2), D; trienes (18:3), T.
[“C]glycerol labeled molecular species are backbone label only (acyl chain
labeling hase been subtracted). Bar shading represents labeling time: 6 min,
black bars; 10 min, white bars; 30 min, gray bars. The endogenous mass
molecular species compositions are from ESI-MS/MS measurements, hatched

bars. A, PC. B, DAG.

165

45

 

 

 

40 1 ~
§
35 J §
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30
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E. 25 . g Q ‘ C16 min DAG
g 20 - % % § I30 min DAG
15 § % $ US min PC
§ g $ .30 min PC
1° ‘ Q S S V
§ § § §
5- n t % |t
0 _ \\ .lk § &

 

Figure 6-8 Fatty acid composition of [14C]glycerol labeled TAG, DAG and
PC molecular species.

The fatty acid composition of acyl groups esterified during de novo glycerolipid
synthesis with glycerol-3-phosphate acylation. Values are calculated from the
[14C]glycerol backbone labeled molecular species in figures 70 (TAG) and S7
(PC and DAG). Fatty acids are represented as number of double bonds:
saturates, S; monoenes, M; dienes, D; trienes, T. Bar shading: 30 min TAG,
black bars; 6 min DAG, white bars; 30 min DAG, light gray bars; 6 min PC,
hatched bars; 30 min PC, dark gray bars.

166

 

TAG Molecular Species Enomoles TAG/embryo
PPI 4.19 1 0.17
PPL 12.92 :I: 0.35
PPO 4.52 :t 0.35
PPS 0.66 :l: 0.27
PLL 8.03 :l: 0.28
PLI 26.86 :t 1.23
PLL 38.60 t 1.11
POI 15.78 :I: 0.45
POL 37.72 1: 2.13
PSI 2.16 :l: 0.12
POO 13.70 :I: 0.26
PM 9.26 :l: 0.18
PSO 6.02 i 0.78
Lll 14.10 i 0.77
LLI 34.12 :l: 1.82
OH 7.40 d: 0.39
OLT 39.06 :l: 1.75
LLL 28.94 :t 1.29
Sll 1.33 i: 0.05
OLL 43.15 :I: 1.75
OOl 12.88 :I: 0.52
SLI 7.69 :l: 0.31
OOL 28.44 :I: 2.06
OLL 10.49 i: 0.76
SOI 5.27 :l: 0.38
SOL 16.20 :1: 0.76
000 11.95 :l: 0.56
SSI 0.79 :l: 0.03
800 8.23 :l: 0.40
SSL 2.97 :l: 0.14
SSO 3.35 :l: 0.21
ALL 2.43 :l: 0.09
AOl 2.85 :l: 0.11
AOL 3.70 :l: 0.20

Table 6-1 TAG Molecular Species Composition of Soybean Embryos

Individual TAG molecular species of developing soybean embryos were
quantified using ESI-MS and ESI-M82. Values represent nanomoles of TAG per
embryo (1: standard deviation). Descriptions of the various molecular species do
not imply any specific stereochemical arrangement of the different acyl chains.
Abbreviations for fatty acids: P = palmitate (16:0), S = stearate (18:0), O = oleate
(18:1), L = linoleate (18:2), l = linolenate (18:3) and A = arachidonate (20:0)

167

DAG Molecular Sgecies picomoles DAG/embryo

PP 19.75 :I: 2.94
Pl 384.64 :I: 22.84
PL 1629.70 :1: 146.31
PO 878.23 :I: 80.62
PS 80.59 d: 11.76
M 125.62 :I: 10.41
Ll 751.03 :I: 39.99
OI 537.02 :I: 30.00
LL 1364.04 :l: 76.21
SI 130.86 :I: 8.31
OL 1884.44 :1: 119.75
SL 692.70 :I: 43.88
00 1239.94 :l: 78.56
$0 594.25 :I: 46.74
SS 105.53 :I: 11.42

Table 6-2 DAG Molecular Species Composition of Soybean Embryos

DAG molecular species of developing soybean embryos were converted to their
sn-1,2-diacyly-3-acetyl-glycerol derivatives and then quantified using ESI-MS and
ESI-M82. Values represent picomoles of DAG per embryo (i standard
deviation). Descriptions of the various molecular species do not imply any
specific stereochemical arrangement of the different acyl chains. Abbreviations
for fatty acids: P = palmitate (16:0), 8 = stearate (18:0), O = oleate (18:1), L =
linoleate (18:2), and l = linolenate (18:3).

168

PC Molecular Species gicomoles Pgembrvo

PP 952.25 :1: 118.71
Pl 832.32 :I: 97.16
PL 3827.04 :1: 456.43
PO 2141.75 1245.10
PS 524.49 :I: 76.10
M 523.51 :I: 98.42
LI 1836.70 :l: 218.43
OI 1244.15 :l: 128.81
LL 3483.64 :t 360.67
SI 358.86 :I: 37.87
0L 4360.23 :l: 460.21
SL 1912.57 :I: 186.51
00 2964.48 :l: 289.10
SO 1730.68 :l: 165.72
SS 259.02 :I: 43.78

Table 6-3 PC Molecular Species Composition of Soybean Embryos

DAG molecular species of developing soybean embryos were converted to their
sn-1,2-diacyly-3-acetyI-glycerol derivatives and then quantified using ESI-MS and
ESI-M82. Values represent picomoles of PC per embryo (:l: standard deviation).
Descriptions of the various molecular species do not imply any specific
stereochemical arrangement of the different acyl chains. Abbreviations for fatty
acids: P = palmitate (16:0), 8 = stearate (18:0), 0 = oleate (18:1), L = linoleate
(18:2), and l = linolenate (18:3).

169

6.2 Supplement: Kinetic modeling of pool filling

To choose between alternative models of TAG synthesis which take into
account various options for DAG-PC interconversions simulations of kinetic
labeling were undertaken (Figure 3-9, and6-9). The expectation was not that any
of the more complex models would exactly match the actual kinetics measured
(Figures 3-2, 3-7). Instead the modeling is used to formulate the best

descriptions of acyl metabolism.

In such modeling of this in vivo data there is a list of implicit assumptions:
(1) The cells are in a uniform physiological state.
(2) Although exogenous “non-physiological” substrates may penetrate differently,
may perturb metabolism slightly differently, they effectively report the bulk tissue
metabolism.
(3) In vivo variance, including that generated by using different experiments used
for acetate and glycerol labeling is small.
(4) We assume that there is no futile cycling of acyl groups, that TAG is a static
end product, and that different lipid molecular species have similar time
constants for pool filling.

(5) Minor acyl fluxes, sUch as to sphingolipids, plastid lipids, PE etc. are ignored.

The modeling is set up with 1000 moles FA in total PC and 400 moles in
total DAG (based off of Table 3-1), which are invariant in size over the hour

period the simulation is run. The inputs are either 100 moles/hr of FA into the

170

system into the DAG(input) pool, or 300 moles via acyl editing into a bulk or small
active PC pool. The output is 100 moles/hr of FA into TAG. A simple Excel
spreadsheet is used to run the simulation, using 0.01 hour increments, that is,
with 1 mole FA input per time increment when modeling for de novo DAG input.
The models simulated are based off those represented in Figure 3-9. In the case
of the simplest model tested, model B (Figure 6-9a), the rate constant for DAG to
TAG conversion k1 will be 0.25 hr", to allow a fixed rate of TAG synthesis of 100
moles/hr (400 moles DAG x 0.25 hr"). The rate constants for the DAG to PC (kg)
and PC to DAG conversions (k3) can be set at any value, but because the DAG
and PC pool sizes must be kept constant, the ratio of k2:k3 = 2.5. The values of
kg and k3 were then varied to allow a simulation of glycerol backbone labeling
from DAG that produced continuous DAG labeling but where PC labeling crossed
over DAG labeling at about 30 minutes as in the experimental results (Figure 3-
7A). This occurred when the forward and back reactions for the DAG to PC
interconversion were 20 times the rate of TAG synthesis (Figure 6-9a). These
parameters were then used to test what would be predicted for an acyl editing
input (Figure 6-9b). At 30 minutes about 21% of the PC would be converted to
DAG. This is inconsistent with experimental observations that show about 3% of
the PC have actually moved from PC to DAG. Thus model B fails to predict the
slow movement of labeled PC to DAG. If at 0.5 hr 21 % PC was converted to
DAG that would change the composition and stereochemistry of the DAG product

dramatically.

171

Additional pool filling simulations for Models B and C (including variants)
were run to see how closely they would conform to the experimental kinetics of
glycerol backbone labeling, of TAG labeling, and of acyl editing. Some of these
are shown in Figure 6—9. Variants on model B suffered the same problem as the
simplest form of model B (too much back conversion of acyl labeled PC to DAG
at early time points). By contrast, variants on model C was generally much more
successful at describing the DAG-PC precursor-product glycerol labeling kinetics
and yet having slow movement of acyl editing labeling of PC move through into
DAG. Model C variants have as a common theme at least two kinetically distinct
DAG pools, DAG(in) for the immediate products of de novo DAG synthesis, and
DAG(out), which supplies TAG synthesis. The DAG input pool is kept small (2%
of PC), in keeping with the observation of very small DAG pools observed in leaf,
because the de novo DAG pool must be compositionally distinct from the bulk
DAG pool, and because glycerol labeled DAG moves through to PC very
significantly within 30 minutes (PC pool filling to TAG takes 10 hours, so if 15 min
pool filling of DAG occurs, it will be 1/40 of PC). To fit for the most simplified
version of Model C (—> DAG -—> PC —> DAG —>TAG) gives are rather flattened
DAG filling curve (6- 90) compared to Figure 3-7, although this can be attenuated
by adding a direct low flux loop from DAG(in) to DAG(out). Thus Model C,

variant 1 seems unlikely.

Splitting the PC pool into a small, active pool and a large, bulk pool Model

C, variant 2 (Figure 6-9d) greatly alleviates the problem of slow DAG pool filling

172

at later times seen in Model C, variant 1. Keeping the active PC pool at 2% of
the total PC, and therefore at the same size as DAG(in), requires that the
interchange between the active and bulk PC pools be 3 times the rate of TAG
synthesis. If this is done, then acyl editing must be largely a function of the bulk
pool (Figure 6-9e), not the active pool (Figure 6-9f). In proposing two PC pools, it
is not clear whether the desaturases act on PC during its residence in the small,
active PC pool, or the bulk PC pool, or both. One problem with simulations of
Model C, variants 1 (Figure 6-9c) and 2 (Figure 6-9d), was that they predicted
movement of glycerol labeling into TAG that was much slower than observed
(Figure 3-7A). This could be remedied by splitting the DAG(out) pool into a small
active and a large bulk component (Figure 6-9g, Model C, variant 3). Of course,
with multiple TAG synthesis routes, this partitioning of DAG(out) may still be a
simplification. More generally, the splitting of the intermediate pools of PC and
DAG allows for a large degree of flexibility in modeling TAG synthesis, with many
combinations of pool sizes and exchange rates. The exercise of modeling initial
kinetics has lead to Model C, variant 3 as the preferred model, giving a
reasonable though not an exact simulation. However, many more complex
variants are possible and cannot be adequately distinguished. Clearly, additional
methods, such as longer term and pulse-chase labeling, will be required to refine

the model.

173

Figure 6-9 Kinetic modeling of pool filling simulations

Graphs of simulations of pool filling, either from de novo DAG acyl labeling
(it will be the same for glycerol backbone de novo labeling), or from acyl editing
inputs. The pool filling represents the labeling experiments, where nascent,
labeled acyl groups are available from fatty acid synthesis for glycerolipid
synthesis.

Figure 6-9a: Pool filling from de novo DAG synthesis using model B.

Figure 6-9b: Pool filling from acyl editing using model B.

Figure 6-9c: Pool filling from de novo DAG synthesis using model C, variant 1.
Figure 6-9d: Pool filling from de novo DAG synthesis using model C, variant 2.
Figure 6-9e: Pool filling from acyl editing via the bulk PC pool using model C,
vaflant2.

Figure 6-9f: Pool filling from acyl editing via the active PC pool using model C,
vanant2.

Figure 6—99: Pool filling from de novo DAG synthesis using model C, variant 3.

Figure 6-9a: Pool filling from de novo DAG synthesis
using model B.

 

70

 

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174

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176

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

   

 

 

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177

 

6.3 Discussion of soybean TAG synthesis flux model Figure 3-10

Model assumptions

The following assumptions used to generate a model of acyl group fluxes
in soybean embryos are based on the endogenous lipid compositions and the
[“C]acetate/[“C]glycerol labeling results and conclusions. 1) De novo synthesis 3
of DAG and acyl editing of PC are metabolically distinct processes. 2) Nascent
FA directly enter PC through sn-1/sn-2 acyl editing. 3) Nascent FA can mix with
recycled FA released from acyl editing for de novo glycerolipid synthesis. 4) The
major flux of de novo synthesized DAG is for PC synthesis. 5) TAG synthesis
utilizes the bulk DAG pool (not de novo DAG) generated from bulk PC. 6) The
bulk DAG generating reaction is in a different location or selective such that de
novo synthesized PC and acyl edited PC are not rapidly used for TAG synthesis.
7) In a soybean embryo rapidly accumulating oil 90% of FAS is ultimately used
for TAG synthesis and most of the PClDAG produced is turned over for TAG
synthesis. 8) Newly synthesized FA are incorporated into PC:DAGzTAG at
relative initial rates of approximately 4:1 :1, respectively. Thus the initial FAS
product incorporation into glycerolipids: 60% PC, 15% DAG, 15% TAG (total
90%).

178

 

Table 6-4 TAG synthesis model: nascent FA flux value calculations

Total S (% 14C) M (% 14C) Nascent Nascent

[“C]FA s Flux M Flux
PC 57 5 95 2.9 54.1
DAG 17 29 71 4.9 12.1
TAG 1 1 75 25 8.2 2.3
Other 15 29 71 4.4 10.6

[14C]acetate labeling values used to calculate the flux of newly synthesized
saturated (S, 16:0, 18:0) and monounsaturated (M, 18:1) FA through the lipid
metabolism model towards TAG. Total [“C]FA values are based on the fact that
85% of newly synthesized FA are incorporated into PC, DAG and TAG at a ratio
of 10:3:2. The remaining 15% most likely is incorporated into other membrane
lipids. Nascent S & M flux values are The model assumes that 93% of FA will
ultimately flux into TAG to generate the endogenous lipid composition.

Supplement: steflw step logic in flpx moc_Lel construction

Model development assumes 100 moles of FAS and utilizes mass balance
along with the initial nascent FA incorporation rates and compositions (Table 6-
4). Newly synthesized FA are incorporated into PC:DAGzTAG at relative initial
rates of 10:3:2, respectively. Thus of initial FAS product incorporation into
glycerolipids: 57% PC, 17% DAG, 11% TAG (total 85%). If 93 moles of FA are
ultimately incorporated into TAG (based on Table 1, and that PC represents ~1/2
of soybean membrane lipids) and the remaining 7 moles for membrane lipid
synthesis then 31 and 3.5 moles of G3P are required for acyl group esterification,
respectively. De novo synthesis of 34.5 mol of total glycerolipids would require 69
mol of FA to flux through PA and DAG into the sn-1/sn-2 positions of membrane

lipids and TAG. The remaining 31 moles from FAS will ultimately be esterifed to

179

the sn-3 position of TAG. If 32 moles of nascent [“C]FA are used by de novo
glycerolipid synthesis (Table 6-4, DAG + other lipids), then an additional 37
moles of recycled [‘ZC]FA are required to balance de novo synthesis.
Presumably this is provided by the acyl chains released by acyl editing to allow
57 mol of nascent [‘4C]FA incorporation into PC through Lyso-PC acylation.
However, 57 mol of PC acyl editing will not provide enough saturates to sustain)
de novo glycerolipid synthesis. Saturates make up ~29% of acyl groups entering
de novo glycerolipid synthesis, as estimated from [“C]glycerol labeling (Figure 6-
8). Thus total saturates would comprise 20 moles of the 69 moles acylated to
G3P. Newly synthesized FA would contribute 9.3 moles of saturates (Table 6-4),
requiring 10.7 moles of recycled saturates. However, 57 mol of PC acyl editing
with equal sn-1/sn-2 contribution could only provide up to 7.6 mol of saturated FA
(PC 26.5% S x 28.5 mol sn-1 FA release). Additionally, nascent FA incorporation
into PC was 14% sn-1 and 86% sn-2. Saturates are predominately at the sn-1
position, therefore if the labeling reflects the positional release of FA during acyl
editing much less saturates would be released and recycled into de novo
synthesis. Therefore the flux of acyl editing cannot be one acyl group released
from PC for each nascent acyl group incorporated, and must be 1.4 times that
required for incorporation of nascent FA into PC with the possibility of being
much greater. The excess flux of recycled acyl groups produced during acyl
editing may be re-incorporated into PC through the acyl editing cycle (Figure 3-
10), with the entire process reducing the saturated concentration of PC and

redistributing FA between the sn-1/sn-2 positions.

180

It is very unlikely that nascent acyl groups are channeled directly into
Lyso-PC that is under a high flux of reacylation with recycled acyl groups, but
more likely the nascent acyl groups mix with the recycled acyl groups of the rapid
acyl editing cycle (Figure 3-10). Initial PC and DAG contain different [14C]FA
incorporation rates and compositions indicating there is either different mixed
pools of new and recycled acyl groups feeding de novo synthesis and acyl editing
or that the acyl editing flux is selective and fast enough that over three times as
much nascent FA enter PC than DAG (Figure 3-2) through de novo synthesis
from the same mixed pool. To determine the relative flux of acyl groups around
the acyl editing cycle we consider how many more FA must be removed from the
mixed acyl pool by acyl editing compared de novo synthesis to produce the
different [”C]FA labeling amounts and stereochemistry of PC and DAG. To
simplify the compositional and stereochemical differences of acylation we
consider the relative flux of [“C]M into the sn-2 position of both lipids, assuming
initial labeling of PC to be 100% sn-2 M and DAG to be 50% sn-2 M. In this
scenario 54.1 moles nascent [“C]M enter sn-2 PC and 11.4 moles enter de novo
glycerolipid synthesis (de novo DAG + other membrane lipids, Table 6-4).
Therefore the flux of nascent [“CJM into sn-2 PC is a maximal 4.8 times faster
than esterification of sn-2 G3P, any greater flux would lower the amount of
nascent acyl groups entering de novo glycerolipid synthesis. However, if there
are any enzymatic selectivites of LPAT or acyl editing enzymes there is a

possibility that the flux could be lower.

181

TAG synthesis will require 31 nmoles of sn-3 acylation. About 11 moles of
this flux is provided by the immediate incorporation of nascent FA, with a high
preference for saturates (Table 64). Analysis of the molecular species of [“ClFA
labeled TAG suggests that this direct acylation utilizes the bulk DAG pool, not the
de novo synthesized DAG pool, and provides about 35% of the molecular
species in endogenous TAG. The other 65% of TAG synthesis would be
compensated by utilizing the recycled acyl-CoA pool from acyl edting and also

PDAT reactions.

182

7 APPENDIX C: PC ACYL EDITING IN ISOLATED PEA
CHLOROPLASTS

Introduction

Isolated pea chloroplasts have been shown to channel newly synthesized
FA into PC, even though chloroplasts cannot synthesized PC de novo [79]. To
determine if acyl editing is involved in this channeling of newly synthesized FA
into PC, isolated pea leaf chloroplasts were labeled with [“C]acetate. The
[“CJFA labeled PC was analysed for labeled FA composition, regiochemistry,
and molecular species.
Results and Discussion

After 5 min of [14C]acetate labeling of isolated pea chloroplasts over half
the radioactivity was in free fatty acids and acyl CoA as reported previously [79].
PC contained the most labeled FA out of all labeled polar lipids and was entirely
[“C]18:1. The stereochemical analysis indicated that ~93% of labeled FA were
localized to the sn-2 position. Separation of labeled molecular species (Figure 7-
1) demonstrated that nascent [14C]FA were incorporated next to unlabeled
saturates, mono-, di- and tri-unsaturates. Together these results are very similar
to the acyl editing measured by in vivo labeling of pea leaves, with the exception

of a highly sn-2 specific FA incorporation.

183

 

60 1
IPC

50
I PG

40

i

30

J

20

L

10

% 14C of total molecular species

 

 

 

 

 

0 .
SS SD ST SM MM MD MT DD DT TT

 

 

 

Figure 7-1 [“C]acetate labeled PC and PG molecular species from isolated
chloroplasts.

Isolated chloroplasts were labeled with [14C]acetate for five minutes. Molecular
species of PC and PG were separated by argentation TLC. Bar heights represent
percent of labeled species among each lipid. Molecular species are represented
as a pair of FA. S, saturates; M, monounsaturates; D, diunsaturates,
triunsaturates. Black bars, PC. Gray bars, PG.

184

The radiolabeled FA composition and molecular species of the prokaryotic
lipid PG were analyzed to ensure that the PC labeling results were from acyl
editing and not due to a transfer of the PC phosphocholine headgroup to a
prokaryotic pathway synthesized DAG. PG from five minute [“C]acetate labeled
pea leaf chloroplast contained approximately equal amounts of saturated and
monounsaturated radiolabeled FA. The molecular species labeled were
predominantly fully saturated and saturated/monounsatured (Figure 7-1).
Together the PG labeling is the expected result of a de novo synthesized
prokaryotic lipid. The difference in PG and PC labeling confirms that
incorporation of newly synthesized FA into PC of isolated chloroplasts is not from
prokaryotic pathway produced DAG.

Therefore, incorporation of newly synthesized FA into PC of isolated pea
chloroplasts is through acyl editing of the sn-2 position. In pea leaves PC sn-2
acyl-editing may be located within the outer chloroplast envelope or associated
with the chloroplast through the PLAM region. The lack of PC sn-1acyl editing in
isolated chloroplasts compared to intact leaves may be because this process
takes place within a different cellular membrane, involves soluble enzymes, or
the activity may be disrupted by chloroplast isolation.

Methods

Pea plants were grown, chloroplasts isolated, labeling conditions and lipid
extraction were as done previously [79]. Labeling was started by adding 5 pCi of
50 mCi/mmol [“C]acetate resuspened in labeling media to 0.4 mg chlorophyll of

chloroplasts with a final volume of 0.2 ml. Chloroplasts were incubated at room

185

temperature under ~180 mol of photons/m2/s of white light, with gentle shaking
for 5 min. The labeling reaction was stopped by adding 2 ml of
chloroform/methanol (1 :1, v/v) to start the lipid extraction. Lipid separation,
radioactive fatty acid composition, regiochemistry and molecular species were

determined as done previously [162].

186

10.

11.

12.

13.

14.

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