Research ArticleArticle
Open Access

Coexpressing Escherichia coli Cyclopropane Synthase with Sterculia foetida Lysophosphatidic Acid Acyltransferase Enhances Cyclopropane Fatty Acid Accumulation

Xiao-Hong Yu, Richa Rawat Prakash, Marie Sweet, John Shanklin

Published January 2014. DOI: https://doi.org/10.1104/pp.113.230953

Abstract

Cyclopropane fatty acids (CPAs) are desirable as renewable chemical feedstocks for the production of paints, plastics, and lubricants. Toward our goal of creating a CPA-accumulating crop, we expressed nine higher plant cyclopropane synthase (CPS) enzymes in the seeds of fad2fae1 Arabidopsis (Arabidopsis thaliana) and observed accumulation of less than 1% CPA. Surprisingly, expression of the Escherichia coli CPS gene resulted in the accumulation of up to 9.1% CPA in the seed. Coexpression of a Sterculia foetida lysophosphatidic acid acyltransferase (SfLPAT) increases CPA accumulation up to 35% in individual T1 seeds. However, seeds with more than 9% CPA exhibit wrinkled seed morphology and reduced size and oil accumulation. Seeds with more than 11% CPA exhibit strongly decreased seed germination and establishment, and no seeds with CPA more than 15% germinated. That previous reports suggest that plant CPS prefers the stereospecific numbering (sn)-1 position whereas E. coli CPS acts on sn-2 of phospholipids prompted us to investigate the preferred positions of CPS on phosphatidylcholine (PC) and triacylglycerol. Unexpectedly, in planta, E. coli CPS acts primarily on the sn-1 position of PC; coexpression of SfLPAT results in the incorporation of CPA at the sn-2 position of lysophosphatidic acid. This enables a cycle that enriches CPA at both sn-1 and sn-2 positions of PC and results in increased accumulation of CPA. These data provide proof of principle that CPA can accumulate to high levels in transgenic seeds and sets the stage for the identification of factors that will facilitate the movement of CPA from PC into triacylglycerol to produce viable seeds with additional CPA accumulation.

Modified fatty acids (mFAs; sometimes referred to as unusual fatty acids) obtained from plants play important roles in industrial applications as lubricants, protective coatings, plastics, inks, cosmetics, etc. The hundreds of potential industrial uses of mFAs have led to considerable interest in exploring their production in transgenic crop plants. mFAs are produced by a limited number of species, and the transfer of genes encoding mFA-producing enzymes from source plants to heterologous hosts has generally resulted in only modest accumulation, usually less than 20% of the desired mFA in transgenic seed (Napier, 2007) compared with levels found in the natural source. For example, ricinoleic acid accounts for more than 90% of the fatty acid of castor bean (Ricinus communis) seeds, and tung (Aleuites fordii) seeds accumulate more than 80% α-eleostearic acid (Thelen and Ohlrogge, 2002; Drexler et al., 2003). In order to elevate the content of mFAs in the engineered plants to that found in the native plant, it is necessary to (1) optimize the synthesis of mFA (Mekhedov et al., 2001), (2) minimize its degradation (Eccleston and Ohlrogge, 1998), and (3) optimize its incorporation into triacylglycerol (TAG; Bafor et al., 1990; Bates and Browse, 2011; van Erp et al., 2011).

Cyclic fatty acids (CFAs) are desirable for numerous industrial applications. The strained bond angles of the carbocyclic ring contribute to their unique chemistry and physical properties, and hydrogenation of CFAs results in ring opening to produce methyl-branched fatty acids. Branched chain fatty acids are ideally suited for the oleochemical industry as feedstocks for the production of lubricants, plastics, paints, dyes, and coatings (Carlsson et al., 2011). Cyclopropane fatty acids (CPAs) have been found in certain gymnosperms, Malvales, Litchi spp., and other Sapindales species. They accumulate to as much as 40% in seeds of Litchi chinensis (Vickery, 1980; Gaydou et al., 1993). Sterculia foetida accumulates the desaturated CFA (i.e. cyclopropene fatty acid) to more than 60% of its seed oil (Bohannon and Kleiman, 1978; Pasha and Ahmad, 1992). The first step in its synthesis is the formation of the CPA by the cyclopropane synthase (CPS) enzyme, which transfers a methyl group to C9 of the oleoyl-phospholipid followed by cyclization to form the cyclopropane ring (Grogan and Cronan, 1997; Bao et al., 2002, 2003). None of the known natural sources of CPA are suitable for its commercial production. Therefore, it would be desirable to create an oilseed crop plant that accumulates high levels of CPA by heterologously expressing CPS in seeds. However, to date, heterologous expression of plant cyclopropane synthase genes has led to only approximately 1.0% CPA in the transgenic seeds (Yu et al., 2011).

Two pathways for the biosynthesis of TAG exist in plants (Bates and Browse, 2012; Fig. 1). The de novo biosynthesis from glycerol-3-phosphate and acyl-CoA occurs via the Kennedy pathway and includes three acyltransferases: glycerol-2-phosphate acyltransferase, acyl-CoA:lysophosphatidic acid acyltransferase (LPAT), and acyl-CoA:diacylglycerol acyltransferase (DGAT; Kennedy, 1961). Alternatively, acyl-CoAs can be redirected from phosphatidylcholine (PC) via the action of a phospholipase C, choline phosphotransferase, phosphatidylcholine:diacylglycerol cholinephosphotransferase (PDCT; Hu et al., 2012; Lu et al., 2009), or phospholipid:diacylglycerol acyltransferase (PDAT; Dahlqvist et al., 2000). An acyl group can be released from PC to generate lysophosphatidylcholine (LPC) by the back reaction of acyl-CoA:LPC acyltransferase (Stymne and Stobart, 1984; Wang et al., 2012) or a phospholipase A/acyl-CoA synthase (Chen et al., 2011).

Figure 1.
Figure 1.

Schematic representation of the plant TAG biosynthesis network. Acyl editing can provide PC-modified fatty acids for de novo diacylglycerol/TAG synthesis. ACS, acyl-CoA synthase; CPT, CDP-choline:diacylglycerol choline phosphotransferase; G3P, glycerol-3-phosphate; GPAT, acyl-CoA:glycerol-3-phosphate acyltransferase; LPC acyltransferase, acyl-CoA:LPC acyltransferase; mFAS, modified fatty acid synthase (in this work, mFAS is CPS); PAP, phosphatidic acid phosphatase; PLA, phospholipase A; PLC, phospholipase C.

LPAT is a pivotal enzyme controlling the metabolic flow of lysophosphatidic acid (LPA) into different phosphatidic acids (PAs) in diverse tissues. Membrane-associated LPAT activities, identified in bacteria, yeast, plant, and animal cells, catalyze the transfer of acyl groups from acyl-CoA to LPA to synthesize PA. In plants and other organisms, LPAT activities have been identified in the endoplasmic reticulum (Kim et al., 2005), plasma membrane (Bursten et al., 1991), and mitochondria (Zborowski and Wojtczak, 1969). In higher plants, endoplasmic reticulum-localized LPAT plays an essential role transferring fatty acid from CoA esters to the sn-2 position of LPA in the synthesis of PA, a key intermediate in the biosynthesis of membrane phospholipids and storage lipids in developing seeds (Maisonneuve et al., 2010). LPAT from developing seeds of flax (Linum usitatissimum), rape (Brassica napus), and castor bean preferentially incorporate oleoyl-CoA, weakly incorporate cyclopropane acyl-CoA, and were unable to incorporate methyl-branched acyl-CoA when presented with an equimolar mix of these potential substrates (Nlandu Mputu et al., 2009). Thus, LPAT activity from agronomic plants constitutes a potential bottleneck for the incorporation of branched chain acyl-CoA into PA. In this work, we investigate the utility of an LPAT from a cyclopropanoid-syntheizing plant, S. foetida, with respect to its ability to enhance CPA accumulation. In our efforts to enhance CPA accumulation in transgenic plants, we screened CPS genes from diverse sources and identified Escherichia coli CPS (EcCPS) as an effective enzyme for the production of CPA in plants. However, EcCPS is reported to prefer the sn-2 position of E. coli phospholipid (Hildebrand and Law, 1964), and the data presented here show that its expression primarily leads to the accumulation of CPA at the stereospecific numbering (sn)-1 position. Moreover, coexpression of S. foetida lysophosphatidic acid acyltransferase (SfLPAT) results in the incorporation of CPA at the sn-2 position of LPA. Thus, coexpression of EcCPS and SfLPAT enables a cycle that enriches the accumulation of CPA at both sn-1 and sn-2 positions of PC and increases the accumulation of CPA. This work underscores the utility of coexpressing an acyltransferase from mFA-accumulating species with mFA-synthesizing enzymes to help mitigate bottlenecks in mFA TAG synthesis.

RESULTS

Expression of CPS in Yeast

Previously, we expressed four plant CPS genes, three from cotton (Gossypium hirsutum) and one from S. foetida, individually in yeast (Saccharomyces cerevisiae). Results from this work showed that the expression of GhCPS1, a CPS from cotton, led to the highest levels of CPA production in both yeast (5.3%) and plants (approximately 1.0%; Yu et al., 2011). To identify a CPS gene that leads to the accumulation of higher levels of CPA, CPS from E. coli, Agrobacterium tumefaciens, and five from Arabidopsis (Arabidopsis thaliana) were cloned and expressed in yeast. As shown in Figure 2, the fatty acid composition of yeast expressing EcCPS showed substantial CPA accumulation. In samples from EcCPS-expressing lines (Fig. 2B), two peaks corresponding to 17:0 CPA and 19:0 CPA (chemical structure shown in Supplemental Fig. S1) were identified based on their mass ions. The expression of EcCPS led to the accumulation of 27% 17:0 CPA and 17% 19:0 CPA, yielding a total of 44% CPA accumulation, which is about 8-fold that observed upon overexpression of the cotton CPS gene GhCPS1 (Fig. 2C). The expression of A. tumefaciens CPS and five putative CPS genes from Arabidopsis did not yield detectable levels of CPA products. These results demonstrate the efficacy of EcCPS relative to other CPS genes for converting both 16:1 and 18:1 fatty acid substrates to the corresponding 17-carbon and 19-carbon CPA products in yeast.

Figure 2.
Figure 2.

Analysis of yeast expressing E. coli and various plant CPS enzymes. GC-MS analysis of FAMEs is shown for yeast expressing empty vector pYES2 (A) or E. coli CPS, which produced both 17:0 CPA and 19:0 CPA (B), and CPA production in CPS expressed in yeast (C). DHSA, Dihydrosterculic acid. FAMEs were analyzed by GC-MS, and both 17:0 and 19:0 CPA accumulations were calculated as a percentage of the total fatty acids (FA). The values represent means and sd of at least three replicates.

Expression of CPS in Arabidopsis fad2fae1

The CPS open reading frames were transferred into plant expression vectors under the control of the seed-specific phaseolin promoter and transformed into Arabidopsis fad2fae1 plants (Meesapyodsuk and Qiu, 2008). This background was chosen because its seed contains more than 80% 18:1, the CPS fatty acid substrate. T1 seeds expressing EcCPS yielded the highest content of dihydrosterculic acid (19-carbon CPA; average 5.0%), and no 17-carbon CPA products were detected. Expression of GhCPS1 and S. foetida CPS led to the accumulation of at most 1% CPA (Yu et al., 2011), whereas the expression of five Arabidopsis and two cotton orthologs (GhCPS2 and GhCPS3) resulted in no detectable accumulation of CPA. T1 fad2fae1 seeds expressing EcCPS germinated with similar frequency to those of nontransformed seeds, and T2 lines with a single locus of insertion were identified and screened for CPA production. These T2 seed pools (containing a mixture of heterozygous and homozygous transgenic seeds) accumulated up to 5.8% CFA (Fig. 3).

Figure 3.
Figure 3.

CPA production in fad2fae1 T2 seeds. FAMEs were extracted from T2 EcCPS fad2fae1 seeds and analyzed by GC-MS. The values represent means and sd of at least three replicates. FA, Fatty acid.

Isolation of an LPAT from S. foetida Seed

RNA from S. foetida leaf and developing seed was extracted and subjected to 454 sequencing. The AtLPAT2 gene sequence encoding the ubiquitous endoplasmic reticulum-located LPAT (Kim et al., 2005) was used to BLAST the S. foetida EST sequences derived from S. foetida, which contains 21,362 seed, and 26,083 leaf EST assemblies. An S. foetida homolog was identified that showed preferential expression in seed, with 56 occurrences in the cotyledon and embryo of developing seeds versus 27 occurrences in leaf tissue. Oligonucleotides were designed to amplify the full-length complementary DNA (cDNA), which was cloned and designated SfLPAT (GenBank accession no. KC894726). SfLPAT has an 1,164-bp open reading frame that encodes a 387-amino acid protein with a predicted mass of 43,723 D and a theoretical pI of 9.63. The predicted amino acid sequence of SfLPAT shows strong homology to Arabidopsis LPAT2 (79.2%), rapeseed (Brassica napus) LPAT2 (79.1%), Arabidopsis LPAT3 (61.9%), and weaker homology to yeast (30.5%) and E. coli (23.5%) LPATs. A phylogenetic tree of the amino acid sequences of the various LPAT proteins was constructed by neighbor-joining distance analysis (Supplemental Fig. S2). S. foetida LPAT groups with AtLPAT2 and three LPATs from Brassica and forms a clade with maize (Zea mays) LPAT and Arabidopsis LPAT3.

Coexpression of EcCPS and SfLPAT in fad2fae1 Arabidopsis Seeds

In order to test if coexpression of the SfLPAT gene along with the EcCPS can enhance CPA accumulation, a single construct containing phaseolin (Phas):EcCPS and Phas:SfLPAT was transformed into the Arabidopsis fad2fae1 background. T1 seeds were analyzed individually for fatty acid composition. Independent T1 seeds accumulated up to 35% CPA. The 2.9% line appears to be an outlier, with all other lines exceeding 12.7% CPA (Fig. 4). The data clearly demonstrate that coexpression of SfLPAT with EcCPS significantly improves CPA accumulation relative to the expression of EcCPS alone.

Figure 4.
Figure 4.

CPA accumulation in individual T1 EcCPS-SfLPAT transgenic fad2fae1 seeds. FAMEs were analyzed by GC-MS, and CPAs are expressed as a percentage of the total fatty acid (FA).

CPA Accumulation in the Progeny of EcCPS-SfLPAT-Coexpressing Transgenic fad2fae1 Seeds

Only approximately 30% of the EcCPS-SfLPAT T1 seeds germinated and were able to develop into mature plants, compared with almost 100% for seeds expressing EcCPS alone. The approximately 70% nonviable seeds showed no radicle penetration of the seed coat, suggesting a failure of germination rather than of establishment. Transgenic lines containing single loci of insertion were identified and allowed to self-fertilize to obtain homozygous individuals. fad2fae1 T2 seeds containing EcCPS and SfLPAT that accumulated low levels of CPA, along with fad2fae1 seeds containing only the EcCPS, exhibited close to 100% germination rates, as did the untransformed fad2fae1 seeds. In contrast, T2 fad2fae1 seeds coexpressing EcCPS-SfLPAT that contained more than 11.5% CPA exhibited reduced germination rates even with the supplement of 1% (w/v) Suc in the medium. For example, no germination was observed for EcCPS-SfLPAT line 2, which accumulated 11.5% CPA in T2 seeds, and only one of 800 seeds of EcCPS-SfLPAT geminated and underwent establishment from line 17 (14.4% CPA) and line 40 (13.3% CPA), and the only T3 progeny identified were found to be heterozygous. Together, these data suggest that elevated accumulation of CPA rather than the presence of SfLPAT was responsible for the observed lack of germination.

Six individual T3 plants from EcCPS lines 17, 38, and 43 and EcCPS-SfLPAT lines 8, 37, and 40 were grown along with parental fad2fae1 plants under identical conditions. There were no discernible morphological or developmental differences between transformed and nontransformed plants. There were no significant differences in flowering time, seed development, or seed numbers. As shown in Figure 5 and Supplemental Table S1, EcCPS T4 homozygous transgenic seeds yielded up to 9.1% CPA, and the progeny of EcCPS-SfLPAT-expressing seeds produced CPA ranging from 10.8% to 13.3%. SfLPAT expression was detected in lines coexpressing EcCPS and SfLPAT, and the EcCPS expression levels were slightly lower than those observed in two lines expressing only EcCPS (Fig. 6), providing further support that increased CPA accumulation results from SfLPAT expression.

Figure 5.
Figure 5.

CPA production in T3 seeds upon expression of the indicated EcCPS or EcCPS-SfLPAT in fad2fae1 plants. FAMEs were analyzed by GC-MS, and CPAs are expressed as a percentage of the total fatty acid (FA). The values represent means and sd of at least three replicates.

Figure 6.
Figure 6.

EcCPS and SfLPAT expression levels in transgenic plants. Quantitative reverse transcription-PCR analysis is shown for EcCPS (A) and SfLPAT (B) expression levels in seeds of fad2fae1 and three transgenic lines harboring EcCPS and EcCPS-SfLPAT as indicated. The expression levels are reported relative to the expression of the UBIQUITIN10 (Ubq10) transcript. The values represent means and sd of at least three replicates.

CPA Effect on Seed Weight and Oil Content

Fatty acids from mature seeds were trans-esterified and quantified via gas chromatography with the use of internal standards. Total seed fatty acid content of untransformed fad2fae1 was 6.45 ± 0.61 µg. The three homozygous lines expressing EcCPS showed no significant differences from the parental line, whereas lines coexpressing SfLPAT with EcCPS resulted in a significant (Student’s t test, P < 0.05) 18% decrease in total fatty acid (Fig. 7A). In addition to fatty acid content, seed weights were also determined. As shown in Figure 7B, parental fad2fae1 seeds weighed 20.0 ± 1.24 µg and T4 seeds of EcCPS-expressing lines showed no significant difference, whereas equivalent lines coexpressing EcCPS and SfLPAT showed a significant (Student’s t test, P < 0.05) decrease in seed weight of up to 11%, with the largest decreases occurring in lines that accumulate the most CPA. Together, these data show that CPA accumulation above 9% causes a decrease in seed weight that correlates with reduced accumulation of seed fatty acid (Fig. 7C).

Figure 7.
Figure 7.

Effects of the expression of EcCPS and EcCPS-SfLPAT on seed fatty acid (FA) content and weight. A, Total fatty acid content in transgenic Arabidopsis. Fatty acid content of seeds was quantified by gas chromatography of FAMEs. The values represent means and sd of at least three replicates. B, Seed weight based on the measurement of batches of 100 transgenic Arabidopsis seeds. The values represent means and sd of at least three replicates. C, Total fatty acid content as a proportion of seed weight. The graph is derived from the values used to generate A and B. Significance levels using Student’s t test are indicated with asterisks: **P < 0.01, *P < 0.05.

CPA Distribution in Transgenic Plants

In Arabidopsis, most mFAs in TAG originate from the PC pool (Bates et al., 2009). Consistent with this, the substrate for S. foetida CPS is 18:1 at the sn-1 position of PC (Bao et al., 2003). We found that CPA accumulates at 25% to 26% in the polar lipid fraction of EcCPS-expressing Arabidopsis lines that accumulate 5% to 9% CPAs in the seed oil at maturity (Fig. 5; Tables I). In order to investigate whether the expression of SfLPAT influences the amount of CPA in the polar lipids, we analyzed the CPA content of the polar lipid and TAG fractions of EcCPS- and EcCPS-SfLPAT-expressing seeds. CPA accumulation increased in both polar lipids and TAG when EcCPS was expressed along with SfLPAT (Tables I and II).

View this table:
Table I. Fatty acid composition of the polar lipids from transgenic Arabidopsis seeds expressing EcCPS or EcCPS-SfLPAT

The values represent means and sd of at least three replicates.

View this table:
Table II. Fatty acid composition of TAG from transgenic Arabidopsis seeds expressing EcCPS or EcCPS-SfLPAT

The values represent means and sd of at least three replicates.

sn-Positional Analysis of CPA in PC and TAG

Differences in sn-positional selectivity of plant and bacterial CPSs could potentially explain the increased abundance of CPA in transgenic seeds expressing the E. coli CPS relative to the plant enzymes. It is reported that SfCPS prefers the sn-1 position, whereas the EcCPS acts on the sn-2 position in E. coli (Hildebrand and Law, 1964; Bao et al., 2003). However, our positional analysis of fad2fae1 Arabidopsis seeds indicates that 18:1 is present at 85% in PC and that both the sn-1 and sn-2 positions of PC have more than 79% 18:1 (Table III). We selected EcCPS43 and EcCPS-SfLPAT37, which have similar total CFA compositions (approximately 9% and 11%, respectively), for detailed analysis to determine the positions occupied by CPA in the PC pool. To accomplish this, isolated PC was incubated with phopholipase A2. The identification of CPA at both the sn-1 and sn-2 positions implies that EcCPS acts on 18:1 at both sn-1 and sn-2 positions of PC in Arabidopsis (Table III); however, significantly higher levels of CPA are present at the sn-1 position, implying a possible preference of more than 5-fold for sn-1 over sn-2 for EcCPS. This results in the accumulation of 85% of the PC CPA in the sn-1 position in EcCPS43 and 70% of the PC CPA in EcCPS-SfLPAT37.

View this table:
Table III. Fatty acid composition of sn-1 and sn-2 positions of PC from transgenic Arabidopsis seeds of EcCPS43 and EcCPS-SfLPAT37

The values represent means and sd of at least three replicates.

In contrast to the different sn-positional distributions of CPA in PC for lines expressing EcCPS and coexpression lines expressing EcCPS and SfLPAT, TAG positional analysis shows that the sn-2 position lacks saturated fatty acids and contains over 90% 18:1. The CPA distribution is similar between the two lines, at approximately 4%, with approximately four times the level of CPA at the sn-1,3 positions relative to the sn-2 position (Table IV).

View this table:
Table IV. Fatty acid composition of sn-1, sn-3, and sn-2 positions of TAG from transgenic Arabidopsis seeds of EcCPS43 and EcCPS-SfLPAT37

The values represent means and sd of at least three replicates.

DISCUSSION

Several findings arise from this work: (1) the expression of EcCPS in transgenic seed leads to the accumulation of higher levels of CPA than the expression of various CPS enzymes from plant sources; (2) the coexpression of EcCPS with SfLPAT results in elevated levels of CPA in the sn-2 position of PC and significantly enhances CPA accumulation to as much as 35% of the total fatty acid in primary transformants; and (3) the accumulation of CPA correlates with reduced seed fatty acid content and germination rate.

The Source of CPS Affects CPA Accumulation in Transgenic Plants

To achieve the goal of accumulating CPA in higher plants, our first approach was to heterologously express CPS genes from other plants, namely cotton, S. foetida, and Arabidopsis, in the fad2fae1 Arabidopsis background tailored to accumulate the CPS substrate oleate. Of the plant CPS genes tested, expression of only S. foetida and a cotton CPS resulted in the accumulation of detectable CPA (up to approximately 1%; Yu et al., 2011). Expression of the remaining seven plant CPSs yielded no detectable CPA. Our screen was then broadened to include CPS genes from the microbial sources E. coli and A. tumefaciens. Expression of the E. coli CPS resulted in the accumulation of approximately 5.0% CPA in primary transformants. These hemizygous T1 seeds were apparently limiting for the CPS enzyme, because CPS accumulation showed a strong dose dependency (i.e. T3 homozygous seeds accumulated approximately twice as much [9.1%] CPA as hemizygous T1 seeds). It is apparently paradoxical that a microbial CPS gene would outperform higher plant CPSs when heterologously expressed in a higher plant. However, a similar occurrence has been reported for the engineering of the hydroxy fatty acid (HFA) ricinoleic acid in plants. In this case, the expression of an oleate hydroxylase from the fungus Claviceps purpurea yielded more hydroxy product than the expression of a plant homolog (Meesapyodsuk and Qiu, 2008). Reasons for the disparity between the efficacy of plant and bacterial CPSs are currently unknown; however, the accumulation of up to 35% CPA upon the coexpression of EcCPS with SfLPAT suggests that the substrate supply (i.e. 18:1 fatty acid and S-adonosyl-Met), is not limiting. Among the possible explanations for the stronger performance of EcCPS relative to the plant CPSs are that the EcCPS may have a higher Vmax or a lower Km for its substrates than the plant CPS enzymes, or the bacterial enzyme may be more stable than the plant enzymes. Understanding why EcCPS outperforms plant CPSs when expressed in plants is the subject of ongoing investigation.

Enhanced Cyclopropane Accumulation upon Coexpression of SfLPAT and EcCPS

A preferentially seed-expressed S. foetida LPAT with homology to the Arabidopsis LPAT2 was identified. Because sterculic acid preferentially accumulates in seeds relative to vegetative tissues, we hypothesized that SfLPAT may have evolved to accommodate the transfer of cyclic fatty acid. A large increase of up to 35% CPA accumulated in the T1 seeds of Arabidopsis fad2fae1 when SfLPAT was coexpressed with EcCPS. We found that E. coli CPS acts on 18:1 primarily at the sn-1 position of PC; coexpression of SfLPAT results in the introduction of CPA at the sn-2 position of LPA. This enables a cycle that enriches for the accumulation of CPA at both sn-1 and sn-2 positions of PC and increases the accumulation of CPA. CPA accumulation is also associated with a decrease in 18:1 fatty acid and a small increase in 16:0 fatty acid in both TAG and total seed fatty acid fractions. That SfLPAT facilitates the accumulation of CPA is interesting because S. foetida accumulates cyclopropene rather than CPAs. The mechanism and substrate of CPA desaturation to produce cyclopropene fatty acids in S. foetida remain elusive; therefore, it is unresolved whether SfLPAT has preference for CPA, cyclopropene fatty acids, or both, relative to normal FA.

Decreased Fatty Acid Content, Seed Weight, and Poor Germination Are Associated with the Coexpression of SfLPAT and EcCPS

When SfLPAT was coexpressed with EcCPS in Arabidopsis, seeds with elevated CPA content exhibited a mildly wrinkled phenotype, similar to that associated with reduced oil content (Focks and Benning, 1998). Quantitative analysis confirmed that wrinkled seeds were lighter than nontransformed control seeds and contained reduced fatty acid content. In transformants expressing only EcCPS that showed high levels of CPA accumulation, oil content was decreased similar to that of EcCPS- and SfLPAT-coexpressing lines, confirming that the developmental deficits arose from elevated CPS rather than the expression of SfLPAT per se. However, at this time, we are unable to preclude the possibility that the expression of SfLPAT results in reduced seed germination. Similar decreases in seed weight and fatty acid content of 10% to 20% have been reported for transgenics accumulating hydroxy or conjugated fatty acid (Cahoon et al., 2006; van Erp et al., 2011). Reduction of seed weight and fatty acid content of ricinoleic acid-accumulating transgenics has been attributed to feedback inhibition of fatty acid synthesis (FAS) upon the accumulation of hydroxy fatty acid lipid intermediates (Bates and Browse, 2011; van Erp et al., 2011). Acyltransferase enzymes facilitate the conversion of membrane lipids to TAG, the sink for fatty acid. Coexpression of castor seed DGAT2, PDCT, or PDAT along with RcFAH12 increased ricinoleic acid accumulation in TAG and decreased ricinoleate levels in membrane lipids along with partial reversion of the inhibition of FAS (Burgal et al., 2008; van Erp et al., 2011; Hu et al., 2012). We hypothesize that elevated levels of CPA in membrane lipids could result in a wrinkled phenotype by a similar feedback inhibition of FAS to that observed upon the accumulation of ricinoleate in membrane lipids (i.e. that the fatty acid released from the turnover of CPA-containing lipids reduce the total rate of fatty acid synthesis; Bates and Browse, 2011; van Erp et al., 2011; Andre et al., 2012). RcFAH-expressing seeds exhibit poor germination, but upon coexpression of castor DGAT2 or PDAT1, higher levels of ricinoleoyl TAG accumulate; however, germination improves because elevated ricinoleoyl-TAG levels were associated with reduced hydroxy fatty acid content in membrane lipids. In contrast, coexpression of EcCPS with SfLPAT resulted in increased CPA in both TAG and membrane lipids, and germination rates decreased as CPA levels increased. In addition to feedback inhibition of FAS, the accumulation of mFA in membrane lipids may result in impaired physiological function of membranes during seed set or desiccation. However, T1 seeds coexpressing EcCPS and SfLPAT accumulate up to 35% CPA with near-normal development and mildly depressed oil accumulation, suggesting that germination is more sensitive to CPA accumulation than seed maturation. Upon germination, mobilization of fatty acid from TAG, or fatty acid transport and catabolism, could be compromised by the presence of mFA in TAG. That the transgenics germinate poorly on Suc-containing medium, under conditions in which β-oxidation mutants germinate well, is consistent with the view that the buildup of mFA degradation intermediates could underlie the poor germination (Eastmond, 2006).

EcCPS Expression Results in the Accumulation of CPA at the sn-1 Position of PC in Plants

SfCPS prefers the sn-1 position of PC in plants (over 90% of the CPA is found in the sn-1 position), whereas EcCPS acts mainly on the monoenoic fatty acid in the sn-2 position of E. coli phospholipid (Hildebrand and Law, 1964; Bao et al., 2003). The reported preference of EcCPS for the sn-2 position of phospholipid may not reflect its actual preference and likely results from the low occupancy of 18:1 substrate at the sn-1 position, which contains predominantly (79%) saturated fatty acid. In fad2fae1 Arabidopsis seeds, 18:1 is present at approximately 84% of the total fatty acid, occurring at similar levels in both the sn-1 and sn-2 positions of PC. It is interesting that EcCPS results in the accumulation of CPA at the sn-1 position of PC in Arabidopsis when presented with equal levels of 18:1 substrate in both the sn-1 and sn-2 positions. S. foetida accumulates sterculic acid equally at all sn-positions, whereas malvic acid is present at only 3% at the sn-2 position (Howarth and Vlahov, 1996). That coexpression of SfLPAT with EcCPS increases the level of CPA in the sn-2 position of PC indicates that CPA from the CoA pool must become esterified to LPA to form PA with CPA at the sn-2 position (Fig. 1). However, that coexpression of SfLPAT does not affect the CPA distribution in TAG implies a preference for normal fatty acid relative to CPA in the sn-2 position by enzymes downstream of PC, such as DGAT, PDCT, and PDAT in the TAG assembly network (Fig. 1). We also note that 16:0 is elevated at the sn-1 and sn-2 positions of PC in the EcCPS-SfLAT37 line (Table III).

Engineering Metabolism to Further Increase CPA Production

In this work, we have shown that coexpression of SfLPAT along with EcCPS results in the accumulation of up to 35% CPA in T1 seeds. However, high levels of CPA in membrane lipids result in unwanted decreases in seed weight and TAG accumulation, resulting from a bottleneck in the conversion of CPA to TAG. Heterologous coexpression of two castor acyltransferases, DGAT2 and PDAT1, with the castor 12-hydroxylase resulted in increased HFA accumulation in Arabidopsis (Burgal et al., 2008; Kim et al., 2011; van Erp et al., 2011). Both by analogy with hydroxy fatty acid accumulation and the bias against CPA in the TAG synthetic network observed in this study, it is likely that stacking TAG-synthesizing enzymes such as DGAT, PDCT, or PDAT with preference for PC CPAs into the EcCPS-SfLPAT transgenic lines will result in enhanced CPA accumulation in TAG (Burgal et al., 2008; Kim et al., 2011; van Erp et al., 2011; Hu et al., 2012; Li et al., 2012). It is interesting that when acyltransferases are expressed in the absence of an mFA-synthesizing enzyme, more rapid movement of fatty acid into TAG generally stimulates higher levels of fatty acid accumulation. For example, increased TAG levels have been observed upon the expression of rapeseed microsomal LPAT or yeast LPAT genes (SLC1 and SLC1-1) in Arabidopsis and rape (Maisonneuve et al., 2010). These reports are consistent with the removal of the feedback inhibition of FAS by reducing the levels of intermediates in the pathway of TAG.

CONCLUSION

Expression of the E. coli CPS gene in fad2fae1 Arabidopsis results in the accumulation of as much as 9.3% CPA in Arabidopsis seeds. Coexpression of SfLPAT can increase CPA accumulation to as high as 35% and elevate levels of CPA in the sn-2 position of PC. However, the seeds with greater than 7% CPA accumulation appear wrinkled and are reduced in size and seed weight. Seeds with more than 11% CPA exhibit reduced germination rates. E. coli CPS acts on 18:1 primarily at the sn-1 position of PC; coexpression of SfLPAT results in the incorporation of CPA at the sn-2 position of LPA. This enables a cycle that enriches the accumulation of CPA at both sn-1 and sn-2 positions of PC and increases the accumulation of CPA. The findings presented here suggest that further optimization of CPA accumulation can potentially be made by stacking additional genes encoding enzymes that will enhance the movement of CPA from membrane lipids into TAG.

MATERIALS AND METHODS

Vector Construction

For expression in yeast, full-length cDNAs of CPSs from Escherichia coli, Agrobacterium tumefaciens, Sterculia foetida, cotton (Gossypium hirsutum), and Arabidopsis (Arabidopsis thaliana) At3g23460, At3g23470, and At3g23480 were amplified and cloned into yeast expression vector pYES2 by restriction of SacI and EcoRI. At3g23510 and At3g23530 were amplified from Arabidopsis cDNA and cloned into pYES2 by restriction of SacI and EcoRI. Primers for Arabidopsis CPS were designed according to their sequences in The Arabidopsis Information Resource (http://www.arabidopsis.org/). For expression in plants, E. coli CPS was amplified from E. coli strain K-12 (Substrain MG1655) using primers ECPS-5′PacI and ECPS-3′XmaI and cloned into the pDsRed plant expression vector (Pidkowich et al., 2007) to form pPhasECPS. Another expression cassette of E. coli CPS was constructed using overlap-extension PCR (Horton et al., 1990). Overlapping fragments of the phaseolin promoter (Pidkowich et al., 2007), E. coli CPS, and the phaseolin terminator were amplified in separate PCRs using appropriate primer pairs (Supplemental Table S2). The PCR products were gel purified and assembled in a PCR primed with terminal primers Phas5′EcoRI and Phas3′EcoRI and then cloned into the pPhasECPS vector with the EcoRI restriction site (New England Biolabs). Further restrictions screen p2PhasECPS, in which the two sets of promoters are in the same direction. S. foetida LPAT was amplified from a native plant. S. foetida LPAT was further cloned into p2PhasECPS through restriction of PacI and XmaI. Primer sequences are listed in Supplemental Table S2.

Plant Growth Conditions and Transgenic Analyses

Developing seeds and leaves of S. foetida were collected from the Montgomery Botanical Center in Florida. The seed coats were removed, and the cotyledons and embryos were frozen with liquid nitrogen and stored at −80°C for RNA extraction and lipid analysis.

Arabidopsis plants were grown in walk-in-growth chambers at 22°C with a 16-h photoperiod. Binary vectors were introduced into A. tumefaciens strain GV3101 by electroporation and were used to transform Arabidopsis via the floral dip method (Clough and Bent, 1998). Seeds of transformed plants were screened under fluorescence emitted upon illumination with green light from an X5 light-emitting diode flashlight (Inova) in conjunction with a Quantaray 25A red camera filter (Pidkowich et al., 2007).

S. foetida EST Analysis

RNA from S. foetida leaves and seeds at different development stages was extracted according to Schultz et al. (1994). RNA quality and concentration were determined by gel electrophoresis and Nanodrop spectroscopy. EST analysis was performed after reverse transcription and size fractionation of the cDNA. mRNA was purified using the Illustra mRNA purification kit (GE Healthcare), and cDNA synthesis was carried out with the use of the Creator SMART cDNA Library Construction Kit (Clontech). The first-strand cDNA was synthesized from 2.0 μg of mRNA using SuperScript II reverse transcriptase (Invitrogen) and Clontech first-strand buffer along with a modified CDS III/3′ cDNA synthesis primer, 5′-TAGAGGCCGAGGCGGCCGACATGTTTTGTTTTTTTTTCTTTTTTTTTTVN-3′. Fourteen thermal cycles were used for cDNA amplification. The cDNA fractions containing sequences of more than 500 bp were used for subsequent 454 sequencing and assembly by the Research Technology Support Facility at Michigan State University.

Phylogenetic Analysis

Phylogenetic analysis of the LPAT family was conducted by using full-length protein sequences from S. foetida, Saccharomyces cerevisiae, E. coli, coconut (Cocos nucifera), rape (Brassica napus), maize (Zea mays), rice (Oryza sativa), and Arabidopsis. Full-length amino acid sequences were first aligned by ClustalW version 2.0.12 (Thompson et al., 1994) with default parameters and imported into the Molecular Evolutionary Genetics Analysis (MEGA) package version 5.0 (Tamura et al., 2007). Phylogenetic and molecular evolutionary analyses were conducted using the neighbor-joining method (Saitou and Nei, 1987) implemented in MEGA, with the pairwise deletion option for handling alignment gaps and the Poisson correction model for computing distance. The final tree graphic was generated using TreeView (Page, 1996).

Transgene Expression Analysis

Arabidopsis RNA from Arabidopsis fad2fae1 seeds and three lines each from EcCPS and EcCPS-SfLPAT transgenic plants were extracted according to Wu et al. (2002). RNA quality and concentration were determined by gel electrophoresis and Nanodrop spectroscopy. Reverse transcription-PCR and quantitative reverse transcription-PCR analysis of EcCPS and SfCPS gene expression were carried out as described (Yu et al., 2011). Primers AtUbiq10-F (5′-TCTTCGCCGGAAAGCAACTTGA-3′) and AtUbiq10-R (5′-TGGCCTTCACGTTGTCAATGGT-3′) were used to amplify UBIQUITIN10 as an internal standard. Gene-specific primers used were qEcCPS-F (5′-GTACCGTATCGCCAACGAATTAC-3′) and qEcCPS-R (5′-CAATACGCAGCGTGTCTTTGA-3′) for EcCPS and qSfLPAT-F (5′-ACTTCTTGGGCATGCTTTGT-3′) and qSfLPAT-R (5′-CTACTGCTGTTTGTCTCGTCTTG-3′) for SfLPAT.

Fatty Acid Analyses

Yeast culture, expression, and fatty acid analyses were carried out as described (Broadwater et al., 2002). Lipids were extracted in methanol:chloroform (2:1) from 0.1 g fresh weight of cotton tissue, and heptadecanoic acid was added as an internal standard. The isolated lipid was methylated in 1% sodium methoxide at 50°C for 1 h and extracted with hexane. Fatty acid methyl esters (FAMEs) from single seeds were prepared by incubating the seed with 30 µL of 0.2 m trimethylsulfonium hydroxide in methanol (Butte et al., 1982). Lipid profiles and acyl group identification were analyzed on a Hewlett-Packard 6890 gas chromatograph equipped with a 5973 mass selective detector for gas chromatography-mass spectrometry (GC-MS) and an Agilent J&W DB 23 capillary column (30 m × 0.25 mm × 0.25 μm). The injector was held at 225°C, and the oven temperature was varied from 100°C to 160°C at 25°C min−1, then to 240°C at 10°C min−1. The percentage values were converted to mole percent and presented as means of at least three replicates.

CPA Distribution in the TAG

Total lipids were extracted from 20 seeds of each T4 line by homogenizing in 500 μL of methanol:chloroform:formic acid (20:10:1, v/v). The organic solvent was extracted with 250 μL of 1 m KCl and 0.2 m H3PO4 twice. The organic phase was dried under N2 and suspended in hexane. Lipids were separated by thin-layer chromatography (TLC) with hexane:diethylether:acetic acid (80:20:1, v/v). The internal standard heptadecanoic acid was added to each fraction, and FAMEs were prepared with 1 mL of methanol:HCl at 90°C for 1 h and extracted with hexane. FAMEs were quantified by GC-MS as described previously (Yu et al., 2011).

Stereospecific Analysis of the Fatty Acid Composition of PC and TAG from Transgenic Arabidopsis

Polar lipid and TAG were separated through a 3-mL Supelco Supel Clean LC-Si SPE column (Sigma), and polar lipid was analyzed with the use of activated ammonium sulfate-impregnated silica gel TLC plates developed with acetone:toluene:water (91:30:7, v/v/v) and stained with 0.05% primuline to isolate PC. PC was digested with phospholipase A2 from Naja mossambica mossambica (Sigma) in borate buffer (0.5 m boric acid, 0.4 mm CaCl2, pH 8.2) for 20 min at room temperature. Reaction products were separated as described previously (Bates et al., 2007). The stereospecific analysis of the fatty acid composition of TAG was performed as described by Cahoon et al. (2006), except that the TLC mobile phase was replaced with chloroform:methanol:acetic acid (70:30:1, v/v/v).

Sequence data from this article can be found in the GenBank/EMBL data libraries under accession number KC894726.

Supplemental Data

The following materials are available in the online version of this article.

Acknowledgments

We thank Dr. M. Patrick Griffith (director of the Montgomery Botanical Center) for assistance with S. foetida tissue collection; Dr. Sean McCorkle (Brookhaven National Laboratory) and Dr. Kevin Carr (Michigan State University) for bioinformatics support; and Dr. Changcheng Xu (Brookhaven National Laboratory), Dr. Phil Bates (University of Southern Mississippi), and Xiangjun Li (University of Nebraska) for technical advice on sn-positional analysis.

Footnotes

  • The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: John Shanklin (shanklin{at}bnl.gov).

  • www.plantphysiol.org/cgi/doi/10.1104/pp.113.230953

  • ↵1 This work was supported by the U.S. Department of Energy (Office of Basic Energy Sciences funding to J.S. and 2010 and 2011 Science Undergraduate Laboratory Internship program funding to M.S.) and by the National Science Foundation (grant no. DBI 0701919 to X.-H.Y. and R.R.P.).

  • ↵2 Present address: Department of Natural Sciences, Suffolk County Community College, Grant Campus, Brentwood, NY 11717.

  • ↵[OPEN] Articles can be viewed online without a subscription.

  • ↵[W] The online version of this article contains Web-only data.

Glossary

mFA
modified fatty acid
TAG
triacylglycerol
CFA
cyclic fatty acid
CPA
cyclopropane fatty acid
LPA
lysophosphatidic acid
LPAT
lysophosphatidic acid acyltransferase
DGAT
diacylglycerol acyltransferase
PC
phosphatidylcholine
PDCT
phosphatidylcholine:diacylglycerol cholinephosphotransferase
PDAT
phospholipid:diacylglycerol acyltransferase
PA
phosphatidic acid
CPS
cyclopropane synthase
cDNA
complementary DNA
FAS
fatty acid synthesis
FAMEs
fatty acid methyl esters
GC-MS
gas chromatography-mass spectrometry
TLC
thin-layer chromatography
  • Received October 21, 2013.
  • Accepted November 5, 2013.
  • Published November 7, 2013.

REFERENCES

  1. ↵
    1. Andre C,
    2. Haslam RP,
    3. Shanklin J
    (2012) Feedback regulation of plastidic acetyl-CoA carboxylase by 18:1-acyl carrier protein in Brassica napus. Proc Natl Acad Sci USA 109: 10107–10112
    OpenUrlAbstract/FREE Full Text
  2. ↵
    1. Bafor M,
    2. Jonsson L,
    3. Stobart AK,
    4. Stymne S
    (1990) Regulation of triacylglycerol biosynthesis in embryos and microsomal preparations from the developing seeds of Cuphea lanceolata. Biochem J 272: 31–38
    OpenUrlAbstract/FREE Full Text
  3. ↵
    1. Bao X,
    2. Katz S,
    3. Pollard M,
    4. Ohlrogge J
    (2002) Carbocyclic fatty acids in plants: biochemical and molecular genetic characterization of cyclopropane fatty acid synthesis of Sterculia foetida. Proc Natl Acad Sci USA 99: 7172–7177
    OpenUrlAbstract/FREE Full Text
  4. ↵
    1. Bao X,
    2. Thelen JJ,
    3. Bonaventure G,
    4. Ohlrogge JB
    (2003) Characterization of cyclopropane fatty-acid synthase from Sterculia foetida. J Biol Chem 278: 12846–12853
    OpenUrlAbstract/FREE Full Text
  5. ↵
    1. Bates PD,
    2. Browse J
    (2011) The pathway of triacylglycerol synthesis through phosphatidylcholine in Arabidopsis produces a bottleneck for the accumulation of unusual fatty acids in transgenic seeds. Plant J 68: 387–399
    OpenUrlCrossRefPubMed
  6. ↵
    1. Bates PD,
    2. Durrett TP,
    3. Ohlrogge JB,
    4. Pollard M
    (2009) Analysis of acyl fluxes through multiple pathways of triacylglycerol synthesis in developing soybean embryos. Plant Physiol 150: 55–72
    OpenUrlAbstract/FREE Full Text
  7. ↵
    1. Bates PD,
    2. Browse J
    (2012) The significance of different diacylgycerol synthesis pathways on plant oil composition and bioengineering. Front Plant Sci 3: 147
    OpenUrlPubMed
  8. ↵
    1. Bates PD,
    2. Ohlrogge JB,
    3. Pollard M
    (2007) Incorporation of newly synthesized fatty acids into cytosolic glycerolipids in pea leaves occurs via acyl editing. J Biol Chem 282: 31206–31216
    OpenUrlAbstract/FREE Full Text
  9. ↵
    1. Bohannon MB,
    2. Kleiman R
    (1978) Cyclopropene fatty acids of selected seed oils from Bombacaceae, Malvaceae, and Sterculiaceae. Lipids 13: 270–273
    OpenUrlCrossRefPubMed
  10. ↵
    1. Broadwater JA,
    2. Whittle E,
    3. Shanklin J
    (2002) Desaturation and hydroxylation: residues 148 and 324 of Arabidopsis FAD2, in addition to substrate chain length, exert a major influence in partitioning of catalytic specificity. J Biol Chem 277: 15613–15620
    OpenUrlAbstract/FREE Full Text
  11. ↵
    1. Burgal J,
    2. Shockey J,
    3. Lu C,
    4. Dyer J,
    5. Larson T,
    6. Graham I,
    7. Browse J
    (2008) Metabolic engineering of hydroxy fatty acid production in plants: RcDGAT2 drives dramatic increases in ricinoleate levels in seed oil. Plant Biotechnol J 6: 819–831
    OpenUrlCrossRefPubMed
  12. ↵
    1. Bursten SL,
    2. Harris WE,
    3. Bomsztyk K,
    4. Lovett D
    (1991) Interleukin-1 rapidly stimulates lysophosphatidate acyltransferase and phosphatidate phosphohydrolase activities in human mesangial cells. J Biol Chem 266: 20732–20743
    OpenUrlAbstract/FREE Full Text
  13. ↵
    1. Butte W,
    2. Eilers J,
    3. Hirsch K
    (1982) Trialkysulfonium-hydroxides and trialkylselonium-hydroxides for the pyrolytic alkylation of acidic compounds. Anal Lett 15: 841–850
    OpenUrlCrossRef
  14. ↵
    1. Cahoon EB,
    2. Dietrich CR,
    3. Meyer K,
    4. Damude HG,
    5. Dyer JM,
    6. Kinney AJ
    (2006) Conjugated fatty acids accumulate to high levels in phospholipids of metabolically engineered soybean and Arabidopsis seeds. Phytochemistry 67: 1166–1176
    OpenUrlCrossRefPubMed
  15. ↵
    1. Carlsson AS,
    2. Yilmaz JL,
    3. Green AG,
    4. Stymne S,
    5. Hofvander P
    (2011) Replacing fossil oil with fresh oil: with what and for what? Eur J Lipid Sci Technol 113: 812–831
    OpenUrlCrossRefPubMed
  16. ↵
    1. Chen GQ,
    2. Snyder CL,
    3. Greer MS,
    4. Weselake RJ
    (2011) Biology and biochemistry of plant phospholipases. Crit Rev Plant Sci 30: 239–258
    OpenUrlCrossRef
  17. ↵
    1. Clough SJ,
    2. Bent AF
    (1998) Floral dip: a simplified method for Agrobacterium-mediated transformation of Arabidopsis thaliana. Plant J 16: 735–743
    OpenUrlCrossRefPubMed
  18. ↵
    1. Dahlqvist A,
    2. Stahl U,
    3. Lenman M,
    4. Banas A,
    5. Lee M,
    6. Sandager L,
    7. Ronne H,
    8. Stymne S
    (2000) Phospholipid:diacylglycerol acyltransferase: an enzyme that catalyzes the acyl-CoA-independent formation of triacylglycerol in yeast and plants. Proc Natl Acad Sci USA 97: 6487–6492
    OpenUrlAbstract/FREE Full Text
  19. ↵
    1. Drexler H,
    2. Spiekermann P,
    3. Meyer A,
    4. Domergue F,
    5. Zank T,
    6. Sperling P,
    7. Abbadi A,
    8. Heinz E
    (2003) Metabolic engineering of fatty acids for breeding of new oilseed crops: strategies, problems and first results. J Plant Physiol 160: 779–802
    OpenUrlCrossRefPubMed
  20. ↵
    1. Eastmond PJ
    (2006) SUGAR-DEPENDENT1 encodes a patatin domain triacylglycerol lipase that initiates storage oil breakdown in germinating Arabidopsis seeds. Plant Cell 18: 665–675
    OpenUrlAbstract/FREE Full Text
  21. ↵
    1. Eccleston VS,
    2. Ohlrogge JB
    (1998) Expression of lauroyl-acyl carrier protein thioesterase in Brassica napus seeds induces pathways for both fatty acid oxidation and biosynthesis and implies a set point for triacylglycerol accumulation. Plant Cell 10: 613–622
    OpenUrlAbstract/FREE Full Text
  22. ↵
    1. Focks N,
    2. Benning C
    (1998) wrinkled1: a novel, low-seed-oil mutant of Arabidopsis with a deficiency in the seed-specific regulation of carbohydrate metabolism. Plant Physiol 118: 91–101
    OpenUrlAbstract/FREE Full Text
  23. ↵
    1. Gaydou EM,
    2. Ralaimanarivo A,
    3. Bianchini JP
    (1993) Cyclopropanoic fatty acids of Litchi (Litchi chinensis) seed oil: a reinvestigation. J Agric Food Chem 41: 886–890
    OpenUrlCrossRef
  24. ↵
    1. Grogan DW,
    2. Cronan JE Jr.
    (1997) Cyclopropane ring formation in membrane lipids of bacteria. Microbiol Mol Biol Rev 61: 429–441
    OpenUrlAbstract/FREE Full Text
  25. ↵
    1. Hildebrand JG,
    2. Law JH
    (1964) Fatty acid distribution in bacterial phospholipids: specificity of cyclopropane synthetase reaction. Biochemistry 3: 1304–1308
    OpenUrlCrossRefPubMed
  26. ↵
    1. Horton RM,
    2. Cai ZL,
    3. Ho SN,
    4. Pease LR
    (1990) Gene splicing by overlap extension: tailor-made genes using the polymerase chain reaction. Biotechniques 8: 528–535
    OpenUrlPubMed
  27. ↵
    1. Howarth OW,
    2. Vlahov G
    (1996) C-13 nuclear magnetic resonance study of cyclopropenoid triacylglycerols. Chem Phys Lipids 81: 81–85
    OpenUrlCrossRef
  28. ↵
    1. Hu ZH,
    2. Ren ZH,
    3. Lu CF
    (2012) The phosphatidylcholine diacylglycerol cholinephosphotransferase is required for efficient hydroxy fatty acid accumulation in transgenic Arabidopsis. Plant Physiol 158: 1944–1954
    OpenUrlAbstract/FREE Full Text
  29. ↵
    1. Kennedy EP
    (1961) Biosynthesis of complex lipids. Fed Proc 20: 934–940
    OpenUrlPubMed
  30. ↵
    1. Kim HU,
    2. Lee KR,
    3. Go YS,
    4. Jung JH,
    5. Suh MC,
    6. Kim JB
    (2011) Endoplasmic reticulum-located PDAT1-2 from castor bean enhances hydroxy fatty acid accumulation in transgenic plants. Plant Cell Physiol 52: 983–993
    OpenUrlAbstract/FREE Full Text
  31. ↵
    1. Kim HU,
    2. Li Y,
    3. Huang AH
    (2005) Ubiquitous and endoplasmic reticulum-located lysophosphatidyl acyltransferase, LPAT2, is essential for female but not male gametophyte development in Arabidopsis. Plant Cell 17: 1073–1089
    OpenUrlAbstract/FREE Full Text
  32. ↵
    1. Li RZ,
    2. Yu KS,
    3. Wu YM,
    4. Tateno M,
    5. Hatanaka T,
    6. Hildebrand DF
    (2012) Vernonia DGATs can complement the disrupted oil and protein metabolism in epoxygenase-expressing soybean seeds. Metab Eng 14: 29–38
    OpenUrlCrossRefPubMed
  33. ↵
    1. Lu CF,
    2. Xin ZG,
    3. Ren ZH,
    4. Miquel M,
    5. Browse J
    (2009) An enzyme regulating triacylglycerol composition is encoded by the ROD1 gene of Arabidopsis. Proc Natl Acad Sci USA 106: 18837–18842
    OpenUrlAbstract/FREE Full Text
  34. ↵
    1. Maisonneuve S,
    2. Bessoule JJ,
    3. Lessire R,
    4. Delseny M,
    5. Roscoe TJ
    (2010) Expression of rapeseed microsomal lysophosphatidic acid acyltransferase isozymes enhances seed oil content in Arabidopsis. Plant Physiol 152: 670–684
    OpenUrlAbstract/FREE Full Text
  35. ↵
    1. Meesapyodsuk D,
    2. Qiu X
    (2008) An oleate hydroxylase from the fungus Claviceps purpurea: cloning, functional analysis, and expression in Arabidopsis. Plant Physiol 147: 1325–1333
    OpenUrlAbstract/FREE Full Text
  36. ↵
    1. Mekhedov S,
    2. Cahoon EB,
    3. Ohlrogge J
    (2001) An unusual seed-specific 3-ketoacyl-ACP synthase associated with the biosynthesis of petroselinic acid in coriander. Plant Mol Biol 47: 507–518
    OpenUrlCrossRefPubMed
  37. ↵
    1. Napier JA
    (2007) The production of unusual fatty acids in transgenic plants. Annu Rev Plant Biol 58: 295–319
    OpenUrlCrossRefPubMed
  38. ↵
    1. Nlandu Mputu M,
    2. Rhazi L,
    3. Vasseur G,
    4. Vu TD,
    5. Gontier E,
    6. Thomasset B
    (2009) Identification of a potential bottleneck in branched chain fatty acid incorporation into triacylglycerol for lipid biosynthesis in agronomic plants. Biochimie 91: 703–710
    OpenUrlCrossRefPubMed
  39. ↵
    1. Page RD
    (1996) TreeView: an application to display phylogenetic trees on personal computers. Comput Appl Biosci 12: 357–358
    OpenUrlFREE Full Text
  40. ↵
    1. Pasha MK,
    2. Ahmad F
    (1992) Analysis of triacylglycerols containing cyclopropene fatty acids in Sterculia foetida (Linn) seed lipids. J Agric Food Chem 40: 626–629
    OpenUrlCrossRef
  41. ↵
    1. Pidkowich MS,
    2. Nguyen HT,
    3. Heilmann I,
    4. Ischebeck T,
    5. Shanklin J
    (2007) Modulating seed beta-ketoacyl-acyl carrier protein synthase II level converts the composition of a temperate seed oil to that of a palm-like tropical oil. Proc Natl Acad Sci USA 104: 4742–4747
    OpenUrlAbstract/FREE Full Text
  42. ↵
    1. Saitou N,
    2. Nei M
    (1987) The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol 4: 406–425
    OpenUrlAbstract
  43. ↵
    1. Schultz DJ,
    2. Craig R,
    3. Cox-Foster DL,
    4. Mumma RO,
    5. Medford JI
    (1994) RNA isolation from recalcitrant plant tissue. Plant Mol Biol Rep 12: 310–316
    OpenUrlCrossRef
  44. ↵
    1. Stymne S,
    2. Stobart AK
    (1984) Evidence for the reversibility of the acyl-CoA:lysophosphatidylcholine acyltransferase in microsomal preparations from developing safflower (Carthamus tinctorius L.) cotyledons and rat liver. Biochem J 223: 305–314
    OpenUrlAbstract/FREE Full Text
  45. ↵
    1. Tamura K,
    2. Dudley J,
    3. Nei M,
    4. Kumar S
    (2007) MEGA4: Molecular Evolutionary Genetics Analysis (MEGA) software version 4.0. Mol Biol Evol 24: 1596–1599
    OpenUrlAbstract/FREE Full Text
  46. ↵
    1. Thelen JJ,
    2. Ohlrogge JB
    (2002) Metabolic engineering of fatty acid biosynthesis in plants. Metab Eng 4: 12–21
    OpenUrlCrossRefPubMed
  47. ↵
    1. Thompson JD,
    2. Higgins DG,
    3. Gibson TJ
    (1994) CLUSTAL W: improving the sensitivity of progressive multiple sequence alignment through sequence weighting, position-specific gap penalties and weight matrix choice. Nucleic Acids Res 22: 4673–4680
    OpenUrlAbstract/FREE Full Text
  48. ↵
    1. van Erp H,
    2. Bates PD,
    3. Burgal J,
    4. Shockey J,
    5. Browse J
    (2011) Castor phospholipid:diacylglycerol acyltransferase facilitates efficient metabolism of hydroxy fatty acids in transgenic Arabidopsis. Plant Physiol 155: 683–693
    OpenUrlAbstract/FREE Full Text
  49. ↵
    1. Vickery JR
    (1980) Fatty-acid composition of seed oils from 10 plant families with particular reference to cyclopropene and dihydrosterculic acids. J Am Oil Chem Soc 57: 87–91
    OpenUrlCrossRef
  50. ↵
    1. Wang LP,
    2. Shen WY,
    3. Kazachkov M,
    4. Chen GQ,
    5. Chen QL,
    6. Carlsson AS,
    7. Stymne S,
    8. Weselake RJ,
    9. Zou JT
    (2012) Metabolic interactions between the Lands cycle and the Kennedy pathway of glycerolipid synthesis in Arabidopsis developing seeds. Plant Cell 24: 4652–4669
    OpenUrlAbstract/FREE Full Text
  51. ↵
    1. Wu YR,
    2. Llewellyn DJ,
    3. Dennis ES
    (2002) A quick and easy method for isolating good-quality RNA from cotton (Gossypium hirsutum L.) tissues. Plant Mol Biol Rep 20: 213–218
    OpenUrlCrossRef
  52. ↵
    1. Yu XH,
    2. Rawat R,
    3. Shanklin J
    (2011) Characterization and analysis of the cotton cyclopropane fatty acid synthase family and their contribution to cyclopropane fatty acid synthesis. BMC Plant Biol 11: 97
    OpenUrlCrossRefPubMed
  53. ↵
    1. Zborowski J,
    2. Wojtczak L
    (1969) Phospholipid synthesis in rat liver mitochondria. Biochim Biophys Acta 187: 73–84
    OpenUrlPubMed
View Abstract
Back to top

Table of Contents

Print
Download PDF
Email Article
Citation Tools
Request Permissions
Coexpressing Escherichia coli Cyclopropane Synthase with Sterculia foetida Lysophosphatidic Acid Acyltransferase Enhances Cyclopropane Fatty Acid Accumulation
Xiao-Hong Yu, Richa Rawat Prakash, Marie Sweet, John Shanklin
Plant Physiology Jan 2014, 164 (1) 455-465; DOI: 10.1104/pp.113.230953
del.icio.us logo Digg logo Reddit logo Twitter logo CiteULike logo Facebook logo Google logo Mendeley logo

Jump to section

In this issue

View this article with LENS