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Plant Physiol. (1998) 118: 183-190
The Biosynthesis of Erucic Acid in Developing Embryos of
Brassica rapa1
Xiaoming Bao,
Mike Pollard, and
John Ohlrogge*
Department of Botany and Plant Pathology, Michigan State
University, East Lansing, Michigan 48824
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ABSTRACT |
The prevailing hypothesis on the
biosynthesis of erucic acid in developing seeds is that oleic acid,
produced in the plastid, is activated to oleoyl-coenzyme A (CoA) for
malonyl-CoA-dependent elongation to erucic acid in the cytosol. Several
in vivo-labeling experiments designed to probe and extend this
hypothesis are reported here. To examine whether newly synthesized
oleic acid is directly elongated to erucic acid in developing seeds of
Brassica rapa L., embryos were labeled with
[14C]acetate, and the ratio of radioactivity of carbon
atoms C-5 to C-22 (de novo fatty acid synthesis portion) to carbon
atoms C-1 to C-4 (elongated portion) of erucic acid was monitored with time. If newly synthesized 18:1 (oleate) immediately becomes a substrate for elongation to erucic acid, this ratio would be expected to remain constant with incubation time. However, if erucic acid is
produced from a pool of preexisting oleic acid, the ratio of C in the 4 elongation carbons to 14C in the
methyl-terminal 18 carbons would be expected to decrease with time.
This labeling ratio decreased with time and, therefore, suggests the
existence of an intermediate pool of 18:1, which contributes at least
part of the oleoyl precursor for the production of erucic acid. The
addition of
2-[{3-chloro-5-(trifluromethyl)-2-pyridinyl}oxyphenoxy] propanoic
acid, which inhibits the homodimeric acetyl-CoA carboxylase, severely
inhibited the synthesis of [14C]erucic acid, indicating
that essentially all malonyl-CoA for elongation of 18:1 to erucate was
produced by homodimeric acetyl-CoA carboxylase. Both light and
2-[{3-chloro-5-(trifluromethyl)-2-pyridinyl}oxyphenoxy]-propanoic acid increased the accumulation of [14C]18:1 and the
parallel accumulation of [14C]phosphatidylcholine. Taken
together, these results show an additional level of complexity in the
biosynthesis of erucic acid.
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INTRODUCTION |
Erucic acid (cis-13-docosenoic acid) and its homolog,
cis-11-eicosenoic acid, are commonly found in the seed oils
of the Cruciferae. The oils and the corresponding fatty acids are
produced by high-erucic acid cultivars of Brassica and
Crambe species and are oleochemical commodities. The
reactions leading to the synthesis of erucic acid are, for the most
part, well understood. In the developing embryos of Brassica
napus (Downey et al., 1964 ), Crambe abyssinica (Appleby
et al., 1974), Simmondsia chinensis (Ohlrogge et al., 1978), Tropaeolum majus (Pollard and Stumpf, 1980a),
and Limnanthes alba (Pollard and Stumpf, 1980b), it
was demonstrated that erucic acid is synthesized by the elongation of
oleic acid rather than by de novo synthesis. This conclusion was
deduced from the distribution of label in the long-chain fatty acids
after incubating seed tissue with exogenous
[14C]acetate. The label was preferentially
incorporated into the carboxyl-terminal carbons of the long-chain fatty
acids rather than the methyl-terminal 18 carbons. Subsequently, the
work of Ohlrogge et al. (1979) demonstrated that de novo fatty acid
synthesis was almost exclusively located in the chloroplast in spinach
leaves. Thus, the hypothesis on the biosynthesis of erucic acid was
extended to a description in which 18:1 (oleate), synthesized in the
plastid, was exported to the cytosol, where, presumably in the
endomembrane system, it was elongated via malonyl-CoA-requiring
elongases to C-20 and longer-chain monounsaturated fatty acids. Several
reports (e.g. von Wettstein-Knowles, 1993; Imai et al., 1995) have
confirmed that the location of the oleoyl elongation system was
extraplastidial, being associated with oil bodies or microsomal
membranes. Créach et al. (1993) and Fehling and Mukherjee
(1991) showed that oleoyl-CoA is readily elongated in vitro and that
the intermediates of the elongation reaction are acyl-CoA thioesters.
It is generally accepted that the end product of newly synthesized
oleic acid exported from plastids is oleoyl-CoA. This oleoyl moiety can
be elongated directly to erucic acid in the ER or in oil
body-associated membranes through successive additions of two carbons
derived from malonyl-CoA. However, in an oil body fraction from
developing rapeseed, Hlousek-Radojcic et al. (1995) observed in vitro
that radioactivity from oleoyl-CoA was incorporated into 20:1
(eicosenoate) and 22:1 (erucate) at least 2.5-fold more slowly than
from malonyl-CoA. Furthermore, radioactivity from oleoyl-CoA was
rapidly diluted upon the formation of eicosenoyl-CoA and the elongation
could proceed without the addition of exogenous oleoyl-CoA. Based on
these in vitro observations, they concluded that oleoyl-CoA is not the
immediate substrate for elongation. Instead, they
proposed that the intermediate oleoyl donor for the elongase may be
either a lipid or an unesterified acid. Furthermore when intact
Brassica (which are high in erucic acid) embryos were incubated with [14C]acetate, PC was always
heavily labeled at early time points, with 18:1 constituting more than
90% of the 14C-labeled fatty acid esterified to
PC (X. Bao and J. Ohlrogge, unpublished data). We considered that this
oleoyl-PC might contribute to the synthesis of erucic acid, either via
a mechanism of direct acyl transfer, as proposed by Hlousek-Radojcic et
al. (1995), or via the acyl exchange between acyl-CoA and PC, as first
reported by Stymne and Stobart (1984). In this study we examined
whether oleoyl-CoA produced by plastids is directly elongated to erucic acid, or if the oleoyl-CoA enters another intermediate pool before it
is elongated. In addition, we have examined the influence of light and
of an inhibitor of the homodimeric ACCase on erucic acid biosynthesis.
Taken together, our results show an additional level of complexity in
the biosynthesis of erucic acid: The supply of oleoyl groups for chain
elongation is a combination of the release of 18:1 from a large,
intermediate lipid pool, probably PC, and the direct provision of newly
synthesized 18:1 from the plastid.
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MATERIALS AND METHODS |
Plant Material and Biochemicals
Developing embryos of Brassica rapa L., which are known
to have a high erucic acid content, were obtained from plants grown in
a growth chamber with 16 h of illumination at 25°C.
Four-week-old siliques were taken from plants and, after removal of
seed coats, the resulting embryos were used immediately for labeling
experiments.
[1-14C]Acetate (1.74 GBq/mmol) and
[U-14C]oleic acid (33.3 GBq/mmol) were
purchased from NEN-DuPont. The herbicide haloxyfop was a gift from
DowElanco (Indianapolis, IN).
[1-14C]Acetate Incubations of B. rapa
Embryos
In the [1-14C]acetate labeling
experiments, three 4-week-old embryos were incubated at 25°C with
gentle shaking either in light (300 µmol s 1
m2) or in the dark in 200 µL of 0.1 mM Mes-NaOH (pH 5.0) containing 5 mM sodium
[1-14C]acetate (1.74 GBq). Assays were
terminated by removing the incubation buffer, washing the embryos twice
with water, and initiating the lipid extraction. For experiments with
the herbicide haloxyfop the embryos were pretreated in 200 µL of 0.1 mM Mes-NaOH (pH 5.0) with the addition of different
concentrations of herbicide for 30 min before the addition of 2 mM [1-14C]acetate substrate.
Lipid Analysis
Lipids were extracted from the embryos according to the method of
Bligh and Dyer (1959). Radioactivity in lipids at each time point was
quantified by liquid-scintillation counting. Lipid classes were
separated by TLC (20- × 20-cm K6 silica, 60A plates, Whatman) to
heights of 4 and 12 cm in chloroform:methanol:acetic acid (75:25:8, v/v/v), allowing the plates to air-dry between developments. The TLC
plates were subsequently developed to 20 cm in hexane:diethyl ether:acetic acid (60:40:1, v/v/v). Radioactivity of the separated lipid classes on the plates was assayed with an Instant Imager (Packard
Instrument Co., Meriden, CT). Labeled TAG and PC bands were eluted from
the silica gel by elution with chloroform:methanol (1:2, v/v). For
transmethylation of total lipids or lipid classes, the lipids were
heated at 90°C for 45 min in 0.3 mL of toluene and 1 mL of 10% boron
trichloride/methanol (Sigma). The recovered C-fatty acid methyl esters were separated by
argentation TLC (Morris et al., 1967). Argentation plates (15% silver
nitrate) were developed sequentially at  20°C to heights of 10, 15, and 20 cm in toluene. Separated 14C-fatty acid
methyl esters were located and quantified with the Instant Imager. The
18:1, 20:1, and 22:1 bands were scraped into test tubes and recovered
by elution with 6 mL of hexane:ethyl ether (2:1, v/v). To characterize
the distribution of label in 18:1, 20:1, and 22:1, these fatty acid
methyl esters were cleaved at the position of the double bond by
permanganate-periodate oxidation (Christie, 1982). The resulting
nonanoic acid (C9A) and 1, -nonane-, undecane- or tridecane-dioic
monomethyl ester fragments (C9AE, C11AE, and C13AE, respectively) were
separated by silica TLC in hexane:ethyl ether:acetic acid (90:10:1,
v/v/v) and quantified using the Instant Imager. The relative amount of
[1-14C]acetate incorporated into the de novo
portion (C-5 to C-22) of 22:1 is calculated simply as C9A + 1.25 C9A or
2.25 C9A.
To measure the in vivo fatty acid accumulation rate, 20 embryos were
taken from plants at different stages from 20 to 43 DAF. At the
initiation of lipid extraction, 500 µg of
1,2-dipentadecanoyl-sn-glycero-3-phosphocholine (Sigma) was added to each sample as an internal standard. Lipid extraction was as described above. Fatty acid methyl esters from total
lipids at different times after flowering were separated and quantified
by GC analysis.
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RESULTS |
Accumulation of Fatty Acids during B. rapa Seed
Development
As shown in Figure 1, developing
B. rapa seeds accumulate fatty acids that are
primarily derived from oleic acid. The modifications of oleic acid fall
into two mutually exclusive types: further desaturation and further
chain elongation. Elongation is the more prevalent modification,
because erucic acid is the most abundant fatty acid, reaching a level
of 56 mol % in seeds at 43 DAF, whereas 18:2 (linoleate) plus 18:3
(linolenate) total <20% at the same stage. 16:0 (palmitate) and 18:0
(stearate), with a content of <3% in mature seeds, are the only
significant fatty acids that do not derive from oleic acid. The studies
described below were conducted on seeds at 28 DAF, when total fatty
acid and erucic acid were accumulating at maximum rates. At this stage, the rate of total 18:1 production, including its elongated and desaturated derivatives, was 16 nmol h1
embryo1.
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| Figure 1.
Accumulation of fatty acids during B. rapa development. Lipids were extracted from 20 pooled embryos
at the times indicated and fatty acid methyl esters were separated and
quantified by GC. The total 18:1 derivatives were obtained by adding
18:1, 18:2, 18:3, 20:1, and 22:1 together. The accumulation rate of
18:1 derivatives (16 nmol h1 embryo1) was
calculated for the embryos at 28 DAF when labeling experiments were
conducted.
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Light Alters Relative Proportions of Oleic and Erucic Acid
Synthesized
To monitor how light influences the accumulation of 18:1 and 22:1,
and TAG versus PC, incubations with
[14C]acetate were carried out either in light
or in the dark. As shown in Figure 2,
incubation of embryos in the light increased radioactivity in 18:1
approximately 2-fold, whereas the radioactivity in 22:1 was only
fractionally higher in light incubations. The comparatively small
effect of light on radioactivity of 22:1, which is highly labeled at
the carboxyl end, suggests that the homodimeric acetyl-CoA carboxylase
and fatty acid elongation are not strongly influenced by light, whereas
the 50% reduction of radioactivity in 18:1 in dark versus light
implies that a major site of light regulation is located in the
plastids, which are responsible for the de novo fatty acid synthesis.
An alternative interpretation of these experiments is that reduction in
label of 18:1 in the dark reflects changes in the endogenous pools of acetate. This explanation was ruled out because very similar light dependence was also observed when [14C]Suc or
[3H]water was used as the precursor for fatty
acid synthesis (not shown). These observations suggest that an
endogenous pool of 18:1 might contribute to the synthesis of
cis-11-20:1 and 22:1. If the newly synthesized 18:1 was the
predominant source of oleoyl moieties for chain elongation and the
synthesis of 18:1 was reduced by one-half in dark, we would expect that
the labeling of 22:1 would also be reduced similarly. In fact, very
little reduction of 22:1 labeling occurs in the dark.
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| Figure 2.
Effects of light on the synthesis of 22:1 (A),
18:1 (B), TAG (C), and PC (D).  , Dark;  , light. Radioactivity is
expressed per embryo.
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Analysis of the Distribution of Label from
[1-14C]Acetate in Long-Chain Fatty Acids with Time
Erucic acid is synthesized from 18:1 by the addition of two
2-carbon units from two molecules of malonyl-CoA. When exogenous [14C]acetate is used as a substrate for the
biosynthesis of 22:1, the 2-carbon units from the chain elongation of
18:1 have a higher specific activity than the 2-carbon units of the
methyl-terminal 18 carbons, which are derived from de novo fatty acid
synthesis of 18:1. The same applies to cis-11-20:1, except
that only one elongation cycle occurs from 18:1. This differential
labeling of C-20 and longer fatty acids from exogenous acetate was
previously documented for four different oilseed species:
Brassica napus (Downey et al., 1964), Simmondsia
chinensis (Ohlrogge et al., 1978), Tropaeolum majus
(Pollard and Stumpf, 1980a), and Limnanthes alba
(Pollard and Stumpf, 1980b), and has been interpreted as reflecting different pools of acetate supplying the de novo fatty acid
synthesis and the chain-elongation reaction. What has not been
examined, however, is the time dependency of this differential labeling. As we will demonstrate below, this time dependency provides information on the pool of 18:1 supplying chain elongation.
Oxidative cleavage at the double bond of monounsaturated fatty acid
methyl esters allows determination of the relative specific activity of
14C in the acid and the acid-ester fragments of
the acyl chain because both of the fragments can be quantitatively
recovered. Table I presents the ratio of
radioactivity in each fragment for isolated 18:1,
cis-11-20:1, and 22:1 when 4-week-old developing B. rapa embryos were incubated with
[1-14C]acetate. The ratios were measured after
various incubation times during a 1-hour period. It is expected that
18:1 will be uniformly labeled, and if the substrate is
[1-14C]acetate, the theoretical distribution
between the C-9 acid-ester and the C-9 acid fragments will be 1.25 (5/4). The measured ratios decreased in the range of 1.24 to 1.30, with
an average value of 1.259 ± 0.016. As an additional control, we
used permanganate-periodate cleavage of commercial
[U-14C]18:1. The theoretical ratios of 1.0 (C9AE/C9A) in 18:1 was obtained exactly (1.00 ± 0.002). For
further confirmation that this method is suitable for quantitative
analysis, the radioactivity in fatty acids was measured before the
oxidation, and was measured again after extraction of cleavage products
from the reaction solution. Loss of radioactivity was negligible
throughout the procedure. All of these controls indicate the high
degree of accuracy and reproducibility of the technique, which is
essential, because the variations in the ratios with time are the key
experimental data.
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Table I.
Ratio of radiolabel in oxidative cleavage fragments
of 14C-labeled fatty acids from incubation of B. rapa
embryos with [1-14C]acetate
Ratios of radioactivity in the 13-C, 11-C, and 9-C carbon fragments
were determined after permanganate-periodate oxidation at the double
bond and isolation of the cleavage products.
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Table I presents the direct measurements of the ratio of
14C label in acid and ester fragments. The
results indicate that the C ratio in the
oxidative cleavage fragments of 18:1 remained constant with time and
was close to the expected value. In contrast, the ratios of
14C in the C13AE/C9A fragments of 22:1 and the
C11AE/C9A fragments of 20:1 both decreased with time in both light- and
dark-incubated embryos. The experiment was repeated 10 times, and
although the absolute values of the acid-to-acid-ester ratios at each
time point showed some degree of variation between experiments, the trend was always the same. Also noted in Table I and consistent with
other experiments described below was the observation that the
acid-ester-to-acid ratios for 22:1 and 20:1 were always higher in dark-
than in light-incubated embryos at any given time point. Thus, in the
dark the relative specific activity of C-2 units used for elongation
when compared with C-2 units derived from de novo fatty acid synthesis
was increased by a factor of 1.5 to 1.6.
It is also instructive to consider these labeling data in terms of the
proportion of total plastid-produced [14C]18:1
units that appear in erucic acid. As calculated from the fatty acid
compositions shown in Figure 1, more than 55% of 18:1 fatty acids
synthesized by 28 DAF in developing seeds are elongated to erucic acid.
Determination of the 14C in the oxidative
cleavage products of 22:1 allows calculation of the
14C content of the de novo-synthesized 18 carbons. Comparison of this value with the total 18:1 radioactivity
accumulated in the incubation (18:1 plus 18-carbon portion of 20:1 and
22:1) gave the values plotted in Figure
3A. After 5 min of incubation in the
light, only 21% of the total 18:1 produced in the incubation appeared
in 22:1, and this value increased to 35% by 60 min. Thus, there was a
substantial lag in the appearance of 14C in the
18:1 portion of 22:1. In dark incubations, a higher proportion of
[14C]18:1 initially appeared in erucic acid, but
the increase with time was similar. The labeling of the 20:1 de novo
and elongation carbons showed parallel patterns in both light and dark
(data not shown).
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| Figure 3.
Labeling of fatty acids by
[14C]acetate in developing B. rapa
embryos. A, The percentage of [14C]acetate incorporated
into total 18:1 (18:1 plus the derived 18:1 portion of 20:1 and 22:1)
that appears in 22:1 is plotted versus time. The incubation was
conducted either in the light () or in the dark (). B, Time
course of 14C accumulation into total 18:1 derivatives
(18:1 plus the derived 18:1 portion of 20:1 and 22:1) in the light. All
of the fatty acids were derived from total lipid extracts, and the
numbers represent radioactivity calculated per embryo.
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Two general explanations can be proposed for the change in labeling
within the very-long-chain fatty acids over time. The first is that
there are different-sized acetate-accessible pools supplying
malonyl-CoA for de novo fatty acid synthesis and for chain elongation.
The pool supplying chain elongation would be relatively small, rapidly
reaching a steady-state contribution from exogenous acetate. The pool
supplying de novo fatty acid biosynthesis would be large and
equilibrate more slowly. A variant of this hypothesis is that the pools
for elongation and de novo fatty acid synthesis that use exogenous
acetate directly are both small, but that acetate can also be used to
sustain de novo fatty acid synthesis via an indirect metabolic pathway
that slowly reaches steady-state labeling. In all cases, 18:1
synthesized at early time points would be of a lower specific
radioactivity compared with later time points, and thus the
acid-ester-to-acid ratio would decrease with time. A corollary of this
hypothesis is that the synthesis of total
[14C]18:1 would show a lag at early time points
with respect to elongation, and with a time scale similar to that of
the change in differential labeling within the very-long-chain fatty
acids. Figure 3B shows the time-dependent accumulation of label in
total 18:1 derivatives (18:1 plus 18:1 portion of 20:1 and 22:1; 18:2
and 18:3 contributed less than 2% of the total label in these
short-term incubations and were not included). The accumulation of
[14C]18:1 was essentially linear, with no lag
phase. Furthermore, the ratio of label in total 18:1 to the label in
the elongation portion of 22:1 plus cis-11-20:1 remained
approximately constant (data not shown), indicating that the first
general mechanism, acetate-accessible pool sizes, was not responsible
for the changes in 14C ratios of de novo to
elongation carbons (Table I).
The second general explanation for the changing ratios of label in the
de novo and elongation carbons considers that another 18:1 source in
addition to the newly synthesized [14C]18:1 was used as a
substrate to synthesize the very-long-chain fatty acids. As shown in
Figure 3A, the radioactivity accumulated in the de novo (18:1) portion
of 22:1 lagged significantly compared with that of total 18:1. This
result supports the concept that there is an endogenous source of 18:1
for the synthesis of 22:1 in addition to the newly synthesized
[14C]18:1. If only newly synthesized 18:1 was
used directly for elongation, the 14C ratios of
C13AE/C9A and C11AE/C9A (Table I) and the percentage of total labeled
18:1 in 22:1 would remain constant with time. The contribution of newly
synthesized 18:1 is gauged by the extrapolation of the curves of Figure
3A to 0 time. At 0 time the amount of the total labeled 18:1 produced,
which contributes to erucic acid biosynthesis, was only 20% in the
light (32% in the dark), whereas the theoretical maximum was 55%.
Thus, 36% (light) and 58% (dark) of the 18:1 flux through the
chain-elongation system to 22:1 was used directly from newly
synthesized (14C-labeled) 18:1 in the plastid, since at 0 time any large intermediate pools of 18:1 had yet to fill.
Inhibition of the Homodimeric ACCase Blocks Erucic Acid
Production
The synthesis of erucic acid from 18:1 has been demonstrated in
vitro to require malonyl-CoA, and it has been assumed that this
malonyl-CoA is produced by the cytosolic homodimeric ACCase. This
assumption has never been tested. To evaluate this hypothesis in vivo
we have used the herbicide haloxyfop, which specifically inhibits the
homodimeric acetyl-CoA carboxylase (Burton et al., 1987, 1991), and
examined how it influences the synthesis of erucic acid and other lipid
species. In the experiment shown in Figure 4 embryos were pretreated with different
concentrations of haloxyfop for 30 min and then incubated with 2 mM [14C]acetate in the light for
1 h. Incorporation of radioactivity into 22:1 decreased with
increased concentrations of haloxyfop. In contrast, radioactivity in
18:1 increased with haloxyfop concentrations lower than 100 µM. At a concentration of 50 µM haloxyfop,
synthesis of 22:1 was inhibited by 70%, but 14C
accumulation in 18:1 increased almost 2-fold. These results demonstrate
that the elongation of 18:1 to 22:1 is dependent on homodimeric ACCase
to supply malonyl-CoA, and that haloxyfop can inhibit the elongation
without inhibition of de novo synthesis of 18:1. Also plotted in Figure
4 are the accumulations of 14C in PC and TAG. In
response to the addition of haloxyfop, radioactivities in 22:1 and TAG
were inhibited in parallel, whereas the accumulation of radioactivities
in PC and 18:1 showed parallel patterns of increase.
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| Figure 4.
Effect of increasing concentrations of haloxyfop
on fatty acid and lipid synthesis. B. rapa embryos were
pretreated with the indicated concentrations of herbicide for 30 min,
then incubated for 1 h in the light with the addition of
[14C]acetate. Radioactivity of lipid and fatty acid are
quantified on the basis of dpm per embryo.
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DISCUSSION |
The radioactivity incorporated into 18:1 of embryos incubated in
the light was approximately 2-fold higher than that in the dark. These
results extend the observations of Browse and Slack (1985), who, in a
study using isolated plastids from linseed seeds (green embryos) and
safflower seeds (white embryos), concluded that linseed plastids are
photosynthetically active and provide a source of ATP and NAD(P)H for
fatty acid synthesis. Similarly, Eastmond et al. (1996) and Asokanthan
et al. (1997) recently concluded that although photosynthesis is
unlikely to provide substantial net carbon for B. napus seed
anabolism, light-driven electron transport may provide ATP and reducing
equivalents for storage-product synthesis. Taken together with the data
in Figure 2, these studies suggest that cofactor supply and/or
plastidial acetyl-CoA carboxylase regulation by light rather than
carbon precursors may be a major limiting factor in the rate of fatty
acid accumulation in Brassica embryos. This hypothesis is
consistent with the observation that acetyl-ACP and malonyl-ACP pools
are relatively high at all stages of active fatty acid synthesis in
developing spinach seeds and castor endosperm (Post-Beittenmiller et
al., 1991, 1992a).
The strong inhibition of acetate incorporation into 22:1 by haloxyfop,
a known noncompetitive inhibitor of the cytosolic homodimeric acetyl-CoA carboxylase, confirms the hypothesis that the acetate that
is used for chain elongation is in the cytosolic compartment and is
distinct from the acetate used for de novo fatty acid synthesis. In all
dicots examined the cytosolic acetyl-CoA carboxylase is the homodimeric
form, whereas the plastid acetyl-CoA carboxylase is the heteromeric
form (Konishi and Sasaki, 1994). However, a homodimeric form of
ACCase has also recently been reported in B. napus plastids
(Roesler et al., 1997; Schulte et al., 1997). Although the function of
this ACCase is unclear at this time, it appears to be less abundant
than the heteromeric form. Furthermore, if this plastid homomeric
ACCase is as sensitive to haloxyfop as the cytosolic form, our
observation that a 50 µM concentration of this herbicide
caused no inhibition of de novo fatty acid synthesis while strongly
inhibiting elongation suggests that the homomeric ACCase in the plastid
has a quantitatively minor role in de novo fatty acid synthesis.
As presented in Table I and Figure 3, we observed differential labeling
of the elongation and de novo carbons within the very-long-chain fatty
acids with time. We considered two explanations for these changes. The
explanation that different-sized acetate-accessible pools supplied
malonyl-CoA for de novo fatty acid synthesis and for chain elongation
was ruled out based on kinetic and pool-size considerations. This is
not surprising if acetate is being used directly for acetyl-CoA
synthesis in both plastids and the cytosol. Data are not available for
oilseeds, but in leaf tissue the chloroplast pool of acetyl-CoA, the
direct substrate for the synthesis of malonyl-CoA, ranges from 10 to 20 µM, and it can be equilibrated with exogenously supplied
acetate within several seconds (Post-Beittenmiller et al., 1992b;
Roughan, 1997). The chloroplast pool of acetyl-CoA dominates the total
leaf acetyl-CoA pool, and, indeed total leaf CoA, so the cytosolic pool
of acetyl-CoA is also expected to be small. Extrapolating to seed
tissue, even if the cytosolic pool of acetyl-CoA is depleted at a rate
of about one-tenth of that of the plastid pool (as calculated from the
mass composition of the oil) it is expected that the direct utilization
of exogenous acetate by either pool will reach steady state very
quickly.
Because endogenous free acetate pools have not been measured in
developing seeds, it is not possible to estimate the endogenous contribution of free acetate to fatty acid synthesis in seeds. However,
utilization of exogenous acetate clearly reaches a steady-state rate
very quickly (Fig. 3B), within 1 min. Acyl-CoA and acyl-ACP pools might
also contribute to a lag in labeling. In Brassica embryos,
and in developing seeds in general, there is limited information on
acyl-ACP levels and a dearth of information on acyl-CoA levels. We can
make some extrapolations from the situation in leaves and from the
limited seed data. In chloroplasts isolated from leaf tissues (Soll and
Roughan, 1982; Roughan and Nishida, 1990), it was estimated that
acyl-ACP half-lives were approximately 10 s. In seeds ACP levels
of approximately 1 µg/g fresh weight were noted (Hannapel and
Ohlrogge, 1988), whereas in spinach seeds and leaves up to 60% of the
ACP is in the free form. Similarly, long-chain acyl-CoA pools in
Cuphea lutea (Singh et al., 1986) and developing B. napus (V. Eccleston and J. Ohlrogge, unpublished observations)
indicate levels of <20 µM. The B. rapa seeds
in the present study accumulated about 16 nmol of fatty acid
h1 seed1. Using these numbers, the
acyl-CoA-pool and acyl-ACP-pool turnover time is calculated to be less
than 1 min. Clearly, accumulation of 18:1 in the acyl-thioester pools
cannot explain any lag in the labeling kinetics of 22:1. This indicates
that the second general explanation, that another 18:1 source in
addition to the newly synthesized [14C]18:1 is used
as substrate to synthesize the very-long-chain fatty acids, is the
correct one.
Three distinct scenarios can be envisaged for the supply of 18:1 for
chain elongation to cis-11-20:1 and 22:1 in developing oilseeds. In the first model, oleoyl moieties are exported from the
plastid, activated to oleoyl-CoA, and can immediately become substrates
for elongation. In this scenario the newly synthesized oleoyl groups
can be considered channeled directly to the cytosolic elongation
system. In the second model, the newly exported oleoyl-CoA is rapidly
equilibrated with the bulk cytosolic pool of oleoyl-CoA. The
acyl-exchange mechanism first reported by Stymne and Stobart (1984) can
be envisaged to dilute the [14C]oleoyl-CoA pool
with oleoyl groups from the sn-2 position of PC, whereas
[14C]18:1 will enter the sn-2
position of PC and be diluted. The bulk pool of oleoyl-CoA is then
available for chain elongation. The third hypothesis is based on the
observations of Hlousek-Radojcic et al. (1995) and requires that newly
synthesized [14C]18:1 acylate an acceptor
lipid, as yet unidentified, and then be transferred directly or
indirectly from this lipid to the elongase. An analysis of the kinetics
of differential labeling within the acyl chain of very-long-chain fatty
acids suggests that an endogenous 18:1 pool contributes to the
biosynthesis of 22:1, although the labeling itself cannot be used to
distinguish between models two and three. Our results cannot rule out
model one or other combinations of these models as a contributing
pathway until the details of the endogenous pool can be demonstrated
and quantified. However, extrapolating Figure 3A to 0 time indicates
that only 20% of total 18:1 synthesis in light (or 32% in the dark)
is directly elongated to 22:1. Comparison of this with the in vivo
conversion of >55% of 18:1 to 22:1 (Fig. 1) suggests that at least
one-half of the 18:1 enters an intermediate pool before elongation to
22:1.
What is the endogenous pool that can accept newly synthesized 18:1 from
the plastid and also provide 18:1 as a substrate for elongation? A
logical approach to obtaining information on such intermediate pools is
to perform pulse-chase experiments. However, with intact B. rapa embryos we found that it was not possible to remove the
[14C]acetate sequestered inside the embryos,
which continued to sustain fatty acid synthesis and hence confound
interpretation. In the absence of pulse-chase data, we decided to take
an indirect approach in which we inhibited the elongation from 18:1 to
22:1 with haloxyfop without reducing the de novo synthesis of 18:1; we
then monitored where the extra 18:1 accumulated. As shown in Figure 4,
there was an increasing amount of 18:1 accumulated in PC when 22:1
synthesis was inhibited, indirectly supporting the concept that the
18:1 esterified to PC could be a source of oleoyl moieties for the synthesis of 22:1. Consistent with this observation, Hlousek-Radojcic et al. (1995) found that when [14C]oleoyl-CoA
was incubated with B. napus oil bodies, more than 50% of
radioactivity was found in PC.
In a study of petroselinic acid biosynthesis in developing coriander
and carrot endosperm, Cahoon and Ohlrogge (1994) concluded that PC was
an intermediate in the movement of petroselinic acid from its site of
biosynthesis in the plastid into TAG. Similar observations have
recently been made for the accumulation of 16:1 6 in developing
seeds of Thunbergia (D. Shultz, E.B. Cahoon, and J. Ohlrogge, unpublished data). Because neither of these unusual fatty
acids is synthesized or further modified on PC, a rationale for their
movement through a PC pool before incorporation into TAG is not
immediately obvious. The present study on erucic acid biosynthesis adds
another example in which PC may be an "intermediate" in the flux of
fatty acids into TAG, even though no metabolism of the fatty acid may
be directly associated with the PC. The observation of a large flux of
fatty acids through PC without modification in three diverse oilseed
species may simply indicate that acyl exchange between the acyl-CoA
pool and PC is very rapid. The low accumulation of unusual fatty acids
in PC may further reflect specific mechanisms for their removal (e.g.
phospholipases). However, it is interesting to speculate that PC may
play some more general role in TAG assembly, perhaps as a carrier of
acyl chains toward a subcellular site of TAG assembly. This role might be analogous to the major flux of acyl chains through PC in leaves, followed by their movement from the ER to the chloroplast.
Because of its commercial value, several attempts have been made to
increase erucic acid content in transgenic plants. However, the factors
that limit erucic acid content in Cruciferae species are still largely
unknown. Expression of a sn-2 acyltransferase from
Limnanthes in transgenic B. napus led to the
accumulation of erucic acid in the sn-2 position, but no
increase in total 22:1 content of the oil (Lassner et al., 1995).
Similarly, overexpression of elongases has resulted in increased chain
length but not an increase in mol % of very-long-chain fatty acids
(Lassner et al., 1996). The recent report that a mutated yeast
SLC1 gene expressed in Arabidopsis or B. napus
gave increased levels of erucic acid (Zou et al., 1997) is at this time
difficult to interpret in light of the results with the
Limnanthes enzyme. Reduction of 18:1 desaturation by
mutation of the oleoyl desaturase might be expected to increase 18:1
availability for elongation. However, in a mutant of Arabidopsis that
is deficient in desaturation of 18:1 (Lemieux et al., 1990), 18:2 and
18:3 content of seeds decreased from 53% to 8.7%, 18:1 content
increased from 15.4% to 53.5%, but 20:1 and 22:1 content just
slightly increased from 20.2% to 26.7%. Likewise, elimination of the
elongation of 18:1 might be expected to increase 18:1 availability for
desaturation. However, neither low-erucic-acid lines of
Brassica (Daun, 1983) nor the fae1 mutant of
Arabidopsis (Kunst et al., 1992) exhibit corresponding increases in
18:1 desaturation. Although several interpretations are possible, one
hypothesis consistent with these results is that the pathways for 18:1
elongation and desaturation may draw on different pools of 18:1. In
summary, our results, together with those of Hlousek-Radojcic et al.
(1995) and those cited above, suggest that the pathway for erucic acid biosynthesis may be more complex than originally thought. Furthermore, the flux of 18:1 through distinct intermediate lipid pools before elongation or desaturation may be one factor that limits the
availability of 18:1 for elongation.
| |
FOOTNOTES |
1
This work was supported by a grant from the
Department of Energy (no. DE-FG02-87ER12729).
*
Corresponding author; e-mail ohlrogge{at}pilot.msu.edu; fax
1-517-353-1926.
Received March 18, 1998;
accepted June 2, 1998.
| |
ABBREVIATIONS |
Abbreviation:
ACCase, acetyl-CoA carboxylase.
ACP, acyl-carrier
protein.
DAF, days after flowering.
haloxyfop, 2-[{3-chloro-5-(trifluromethyl)-2-pyridinyl}oxyphenoxy]-propanoic
acid.
PC, phosphatidylcholine.
TAG, triacylglycerol.
X:Y, a fatty acyl
group containing X carbon atoms and Y cis double
bonds.
| |
ACKNOWLEDGMENT |
We thank the Michigan Agricultural Experiment Station for its
support of this research.
| |
LITERATURE CITED |
Appleby RS,
Gurr MI,
Nichols BW
(1974)
Studies on seed-oil triglycerides: factor controlling biosynthesis of fatty acid and acyl lipids in subcellular organelles of maturing Crambe abyssinica seeds.
Eur J Biochem
48:
209-216
[Medline]
Asokanthan PS,
Johnson RW,
Griffith M,
Krol M
(1997)
The photosynthetic potential of canola embryos.
Physiol Plant
101:
353-360
[CrossRef]
Bligh EG,
Dyer WJ
(1959)
A rapid method of total lipid extraction and purification.
Can J Biochem Physiol
37:
911-917
Browse J,
Slack CR
(1985)
Fatty-acid synthesis in plastids from maturing safflower and linseed cotyledons.
Planta
166:
74-80
[CrossRef]
Burton JD,
Gronwald JW,
Keith RA,
Somers DA,
Gengenbach BG,
Wyse DL
(1991)
Kinetics of inhibition of plant acetyl-coenzyme A carboxylase by sethoxydim and haloxyfop.
Pestic Biochem Physiol
39:
100-109
Burton JD,
Gronwald JW,
Somers DA,
Connelly JA,
Gengenbach BG,
Wyse DL
(1987)
Inhibition of plant acetyl-coenzyme A carboxylase by the herbicides sethoxydim and haloxyfop.
Biochem Biophys Res Commun
148:
1039-1044
[CrossRef][Web of Science][Medline]
Cahoon EB,
Ohlrogge JB
(1994)
Apparent role of phosphatidylcholine in the metabolism of petroselinic acid in developing Umbelliferae endosperm.
Plant Physiol
104:
845-855
[Abstract]
Christie WW
(1982)
Lipid Analysis, Ed 2.
Pergammon Press, Oxford, UK
Créach A,
Lessire R,
Cassagne C
(1993)
Kinetics of C18:1-CoA elongation and transacylation in rapeseeds.
Plant Physiol Biochem
31:
923-930
Daun JK
(1983)
The introduction of low erucic acid rapeseed varieties into Canadian production.
In
JKG Kramer,
FD Sauer,
WJ Pigden,
eds, High and Low Erucic Acid Rapeseed Oils: Production, Usage, Chemistry, and Toxicological Evaluation.
Academic Press, Toronto, Canada, pp 161-180
Downey RK,
Craig BM
(1964)
Genetic control of fatty acid biosynthesis in rapeseed (Brassica napus L.).
J Am Oil Chem Soc
41:
475-478
Eastmond P,
Kolacna L,
Rawsthorne S
(1996)
Photosynthesis by developing embryos of oilseed rape (Brassica napus).
J Exp Bot
47:
1763-1769
Fehling E,
Mukherjee KD
(1991)
Acyl-CoA elongase from higher plant (Lunaria annua): metabolic intermediates of very-long-chain acyl-CoA products and substrate specificity.
Biochim Biophys Acta
1082:
239-246
[Medline]
Hannapel DJ,
Ohlrogge JB
(1988)
Regulation of acyl carrier protein messenger RNA levels during seed and leaf development.
Plant Physiol
86:
1174-1178
[Abstract/Free Full Text]
Hlousek-Radojcic A,
Imai H,
Jaworski JG
(1995)
Oleoyl-CoA is not an immediate substrate for fatty acid elongation in developing seeds of Brassica napus.
Plant J
8:
803-809
Imai H,
Hlousek-Radojcic A,
Matthis A,
Jaworski J
(1995)
Elongation system involved in the biosynthesis of very long chain fatty acids in Brassica napus seeds: characterization and solubilization.
In
JC Kader,
P Mazliak,
eds, Plant Lipid Metabolism.
Kluwer Academic Publishers, Dordrecht, The Netherlands, pp 118-120
Konishi T,
Sasaki Y
(1994)
Compartmentalization of two forms of acetyl-CoA carboxylase in plants and the origin of their tolerance towards herbicides.
Proc Natl Acad Sci USA
91:
3598-3601
[Abstract/Free Full Text]
Kunst L,
Taylor DC,
Underhill EW
(1992)
Fatty acid elongation in developing seeds of Arabidopsis thaliana.
Plant Physiol Biochem
30:
425-434
Lassner MW,
Lardizabal K,
Metz JG
(1996)
A jojoba beta-ketoacyl-CoA synthase cDNA complements the canola fatty acid elongation mutation in transgenic plants.
Plant Cell
8:
281-292
[Abstract]
Lassner MW,
Levering CK,
Davies HM,
Knutzon DS
(1995)
Lysophosphatidic acid acyltransferase from meadowform mediates the insertion of erucic acid at the sn-2 position of triacylglycerol in transgenic rapeseed oil.
Plant Physiol
109:
1389-1394
[Abstract]
Lemieux B,
Miquel M,
Somerville C,
Browse J
(1990)
Mutants of Arabidopsis with alterations in seed lipid fatty acid composition.
Theor Appl Genet
80:
234-240
[Web of Science]
Morris LJ,
Wharry DM,
Hammond EW
(1967)
Chromatographic behaviour of isomeric long-chain aliphatic compounds. II. Argentation thin-layer chromatography of isomeric octadecenoates.
J Chromatogr
31:
69-76
[CrossRef]
Ohlrogge JB,
Kuhn DN,
Stumpf PK
(1979)
Subcellular localization of acyl carrier protein in leaf protoplasts of Spinacia oleracea.
Proc Natl Acad Sci USA
76:
1194-1198
[Abstract/Free Full Text]
Ohlrogge JB,
Pollard MR,
Stumpf PK
(1978)
Studies on biosynthesis of waxes by developing jojoba seed tissue.
Lipids
11:
651-662
Pollard MR,
Stumpf PK
(1980a)
Long-chain (C20 and C22) fatty acid biosynthesis in developing seeds of Tropaeolum majus: an in vivo study.
Plant Physiol
66:
641-648
[Abstract/Free Full Text]
Pollard MR,
Stumpf PK
(1980b)
Biosynthesis of C20 and C22 fatty acid by developing seeds of Limnanthes alba.
Plant Physiol
66:
649-655
[Abstract/Free Full Text]
Post-Beittenmiller D,
Jaworski JG,
Ohlrogge JB
(1991)
In vivo pools of free and acylated acyl carrier proteins in spinach: evidence for sites of regulation of fatty acid biosynthesis.
J Biol Chem
266:
1858-1865
[Abstract/Free Full Text]
Post-Beittenmiller D, Jaworski JG, Ohlrogge JB (1992a) Regulation
of lipid synthesis in castor seeds: analysis of the in vivo acyl-acyl
carrier protein pools. In SL MacKenzie, DC Taylor, eds, Seed
Oils for the Future. American Oil Chemists' Society, Champaign, IL, pp
44-51
Post-Beittenmiller D,
Roughan PG,
Ohlrogge JB
(1992b)
Regulation of fatty acid biosynthesis: analysis of acyl-coenzyme A and acyl-acyl carrier protein substrate pools in spinach and pea chloroplasts.
Plant Physiol
100:
923-930
[Abstract/Free Full Text]
Roesler KR,
Shintani D,
Savage L,
Boddupalli S,
Ohlrogge JB
(1997)
Targeting of the Arabidopsis homomeric acetyl-CoA carboxylase to plastids of rapeseed.
Plant Physiol
113:
75-81
[Abstract]
Roughan PG
(1997)
Stromal concentrations of coenzyme A and its esters are insufficient to account for rates of chloroplast fatty acid synthesis: evidence for substrate channelling within the chloroplast fatty acid synthase.
Biochem J
327:
267-273
Roughan PG,
Nishida I
(1990)
Concentration of long-chain acyl-acyl carrier proteins during fatty acid synthesis by chloroplasts isolated from pea (Pisum sativum), safflower (Carthamus tinctoris), and amaranthus (Amaranthus lividus) leaves.
Arch Biochem Biophys
276:
1-9
[Medline]
Schulte W,
Topfer R,
Stracke R,
Schell J,
Martini N
(1997)
Multi-functional acetyl-CoA carboxylase from Brassica napus is encoded by a multi-gene family: indication for plastidic localization of at least one isoform.
Proc Natl Acad Sci USA
94:
3465-3470
[Abstract/Free Full Text]
Singh SS,
Nee TY,
Pollard MR
(1986)
Acetate and mevalonate labeling studies with developing Cuphea lutea seeds.
Lipids
21:
143-149
Soll J,
Roughan PG
(1982)
Acyl-acyl carrier protein pool sizes during steady-state fatty acid synthesis by isolated spinach chloroplasts.
FEBS Lett
146:
189-192
[CrossRef]
Stymne S,
Stobart AK
(1984)
The biosynthesis of triacylglycerols in microsomal preparations of developing cotyledons of sunflower (Helianthus annus L.).
Biochem J
220:
481-488
[Medline]
von Wettstein-Knowles PM (1993) Wax, cutin, and suberin.
In TS Moore Jr, ed, Lipid Metabolism in Plants. CRC Press,
Boca Raton, FL, pp 127-166
Zou JT,
Katavic V,
Giblin EM,
Barton DL,
Mackeenzie SL,
Keller WA,
Hu X,
Taylor DC
(1997)
Modification of seed oil content and acyl composition in the Brassicaceae by expression of a yeast sn-2 acyltransferase gene.
Plant Cell
9:
909-923
[Abstract/Free Full Text]
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Top
Abstract
Introduction