|Tissue-specific whole transcriptome sequencing in castor, directed at understanding triacylglycerol lipid biosynthetic pathways.|
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|PMID: 22319559 Owner: NLM Status: MEDLINE|
|BACKGROUND: Storage triacylglycerols in castor bean seeds are enriched in the hydroxylated fatty acid ricinoleate. Extensive tissue-specific RNA-Seq transcriptome and lipid analysis will help identify components important for its biosynthesis.
METHODOLOGY/FINDINGS: Storage triacylglycerols (TAGs) in the endosperm of developing castor (Ricinus communis) seeds are highly enriched in ricinoleic acid (18:1-OH). We have analysed neutral lipid fractions from other castor tissues using TLC, GLC and mass spectrometry. Cotyledons, like the endosperm, contain high levels of 18:1-OH in TAG. Pollen and male developing flowers accumulate TAG but do not contain 18:1-OH and leaves do not contain TAG or 18:1-OH. Analysis of acyl-CoAs in developing endosperm shows that ricinoleoyl-CoA is not the dominant acyl-CoA, indicating that either metabolic channelling or enzyme substrate selectivity are important in the synthesis of tri-ricinolein in this tissue. RNA-Seq transcriptomic analysis, using Illumina sequencing by synthesis technology, has been performed on mRNA isolated from two stages of developing seeds, germinating seeds, leaf and pollen-producing male flowers in order to identify differences in lipid-metabolic pathways and enzyme isoforms which could be important in the biosynthesis of TAG enriched in 18:1-OH. This study gives comprehensive coverage of gene expression in a variety of different castor tissues. The potential role of differentially expressed genes is discussed against a background of proteins identified in the endoplasmic reticulum, which is the site of TAG biosynthesis, and transgenic studies aimed at increasing the ricinoleic acid content of TAG.
CONCLUSIONS/SIGNIFICANCE: Several of the genes identified in this tissue-specific whole transcriptome study have been used in transgenic plant research aimed at increasing the level of ricinoleic acid in TAG. New candidate genes have been identified which might further improve the level of ricinoleic acid in transgenic crops.
|Adrian P Brown; Johan T M Kroon; David Swarbreck; Melanie Febrer; Tony R Larson; Ian A Graham; Mario Caccamo; Antoni R Slabas|
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|Type: Journal Article; Research Support, Non-U.S. Gov't Date: 2012-02-03|
|Title: PloS one Volume: 7 ISSN: 1932-6203 ISO Abbreviation: PLoS ONE Publication Date: 2012|
|Created Date: 2012-02-09 Completed Date: 2012-09-24 Revised Date: 2014-10-14|
Medline Journal Info:
|Nlm Unique ID: 101285081 Medline TA: PLoS One Country: United States|
|Languages: eng Pagination: e30100 Citation Subset: IM|
|APA/MLA Format Download EndNote Download BibTex|
Biosynthetic Pathways / genetics*
Castor Bean / enzymology, genetics*
Plants, Genetically Modified
Transcriptome / genetics*
Triglycerides / biosynthesis*
|BB/F004354/1//Biotechnology and Biological Sciences Research Council; BB/F004354/1//Department of Health|
|0/Ricinoleic Acids; 0/Triglycerides; I2D0F69854/ricinoleic acid|
Journal ID (nlm-ta): PLoS One
Journal ID (publisher-id): plos
Journal ID (pmc): plosone
Publisher: Public Library of Science, San Francisco, USA
Brown et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Received Day: 7 Month: 10 Year: 2011
Accepted Day: 9 Month: 12 Year: 2011
collection publication date: Year: 2012
Electronic publication date: Day: 3 Month: 2 Year: 2012
Volume: 7 Issue: 2
E-location ID: e30100
PubMed Id: 22319559
Publisher Id: PONE-D-11-19991
|Tissue-Specific Whole Transcriptome Sequencing in Castor, Directed at Understanding Triacylglycerol Lipid Biosynthetic Pathways Alternate Title:Tissue-Specific Transcriptome Sequencing in Castor|
|Adrian P. Brown1|
|Johan T. M. Kroon1|
|Tony R. Larson3|
|Ian A. Graham3|
|Antoni R. Slabas1*|
1School of Biological and Biomedical Sciences, Durham University, Durham, United Kingdom
2The Genome Analysis Centre, Norwich Research Park, Colney, Norwich, United Kingdom
3Department of Biology, Centre for Novel Agricultural Products, University of York, York, United Kingdom
|Max Planck Institute for Chemical Ecology, Germany
|Correspondence: * E-mail: email@example.com
Contributed by footnote: Conceived and designed the experiments: ARS. Performed the experiments: AB JK MF TL. Analyzed the data: DS MC AB JK. Contributed reagents/materials/analysis tools: DS MC TL IG. Wrote the paper: ARS AB JK.
Plant seeds contain a diverse range of fatty acids which have uses in the food, health and industrial sectors. These fatty acids accumulate in seed triacylglycerols (TAGs) in which each of the three carbons of the glycerol backbone can be linked to a different fatty acid. From an industrial perspective however it is desirable to have TAGs enriched with one specific fatty acid so they can be used directly following hydrolysis, circumventing the requirement to fractionate individual fatty acid species. Ricinoleic acid (12-hydroxyoctadec-cis-9-enoic acid; 18:1-OH) is the major fatty acid in Ricinus communis (castor bean) seeds where it constitutes up to 90% of the fatty acids found in TAGs. One of the principle uses of 18:1-OH is as a raw material for the production of Nylon-11 (N-11) which is resistant to organic solvents at high temperature. This property makes N-11 a highly desirable polymer for the manufacture of tubing used to carry hydraulic fluids in automotive engines. Over 50,000 tons of 18:1-OH, sourced from castor, is used annually for the production of N-11 but castor has a number of undesirable attributes as an industrial crop. It contains the highly allergenic 2S albumin and the toxin ricin, is restricted climatically to certain growth areas and is a non-determinant plant which is not conducive to mechanical harvesting. An added problem is the inconsistent price of castor oil in the commercial market which can greatly affect the economics of N-11 production. To overcome these problems, research has been directed at understanding the metabolic pathways of tri-ricinolein (TAG with 18:1-OH at all three positions on the glycerol backbone) biosynthesis from oleic acid in developing castor seeds with a view to engineering its synthesis in alternative oilseed crops.
Conversion of oleic acid to 18:1-OH in plant seeds occurs in the endoplasmic reticulum (ER) whilst the fatty acid is attached to the sn-2 position of phosphatidylcholine (PC) . This reaction is catalysed by oleate 12-hydroxylase (phosphatidylcholine 12-monooxygenase). The gene encoding this enzyme was discovered using an expressed sequence tag (EST) sequencing approach on developing castor beans . Identification was based on the knowledge that the reaction was not catalysed by a cytochrome P450 but probably by a protein related to acyl-lipid desaturases  and that 18:1-OH synthesis occurs exclusively in seeds. Subsequent transformation of tobacco and mass spectrometry of fatty acids demonstrated proof of function of the identified sequence. The amount of 18:1-OH produced in tobacco seeds was low, only 0.1%, but subsequent experiments with the castor and related Lesquerella fendleri hydroxylases produced seeds with 12.8 and 15.6% hydroxylated fatty acids respectively in Arabidopsis thaliana and Brassica napus.
Two sets of reactions can reduce 18:1-OH levels in both castor and transgenic plant TAGs: (a) competition for the pool of oleic acid by oleate desaturase and (b) further metabolism of 18:1-OH by elongation reactions and additional desaturases to give for example 18:2-OH and 20:1-OH fatty acids. The effect of reducing these competing reactions was studied by expressing oleate 12-hydroxylase in Arabidopsis with fad (oleate desaturase) and fae (fatty acid elongation) mutations . Double mutant lines accumulated 19.2% hydroxylated fatty acids in mature seeds which is substantially lower than the level in castor oil and points to a requirement for additional modification of TAG-synthetic reactions to increase 18:1-OH levels in transgenic plants.
The classical pathway for TAG biosynthesis in seeds is the acyl-CoA dependent Kennedy pathway which involves three membrane-bound acyltransferases and a phosphatidic acid phosphatase ,  (Figure 1). These enzymes are encoded by gene families in plants and each isoenzyme may have a different specificity for both acyl donor and acceptor. Individual acyltransferases can therefore influence incorporation of specific fatty acids into TAG and transfer of them between plant species has been shown to alter seed oil compositions , . Two diacylglycerol acyltransferase (DGAT) isoenzymes are present in castor: RcDGAT1 and RcDGAT2. In vitro experiments demonstrated that RcDGAT2 can utilise di-ricinolein and 18:1-OH-CoA substrates and its expression profile in developing castor bean is consistent with a specific involvement in tri-ricinolein biosynthesis . This was subsequently demonstrated by studies in which introduction of RcDGAT2 into Arabidopsis lines expressing oleate 12-hydroxylase resulted in elevation of 18:1-OH seed content from 17 to nearly 30% .
Identification of enzymes important in the formation of 18:1-OH-CoA substrates for acyl-CoA dependent reactions is complicated by the occurrence of enzyme isoforms and alternative biosynthetic routes. Acyl editing, the exchange of fatty acids between polar membrane lipids such as PC and the acyl-CoA pool without net glycerolipid synthesis , , , can occur by two pathways (Figure 1). Modified fatty acids may be removed from PC by phospholipase A2 (PL-A2) and the released fatty acids then activated to acyl-CoAs by long-chain acyl-CoA synthetase (LACS). Alternatively, the reverse reaction of acyl-CoA:lysophosphatidylcholine acyltransferase (LPCAT) can generate acyl-CoAs from PC .
An acyl-CoA independent TAG-synthetic pathway is active in a number of plant species, including castor . In this, phospholipid:diacylglycerol acyltransferase (PDAT) catalyses transacylation of the sn-2 fatty acid from PC onto the sn-3 position of diacylglycerol (DAG), with lysophosphatidylcholine (LPC) as a co-product (Figure 1). A variety of PDAT isoforms exist in both Arabidopis and castor and the effects of transferring castor PDATs into Arabidopsis co-expressing oleate 12-hydroxylase has been investigated. Transformation with RcPDAT1 produced up to 27% hydroxylated fatty acids in Arabidopsis seeds , . An alternative acyl-CoA independent process for the incorporation of hydroxylated fatty acids into TAG from PC is removal of the phosphocholine head-group to form diacylglycerol (which can be further acylated by PDAT). Three different mechanisms are possible for this step involving either phospholipase C, the reverse action of CDP-choline:diacylglycerol cholinephosphotransferase (CPT)  or the recently identified phosphatidylcholine:diacylglycerol cholinephosphotransferase (PDCT/ROD1) . Determining which isoform(s) of these enzymes are expressed in castor seed may be important for engineering of 18:1-OH synthesis. Metabolic labelling experiments have recently shown that in Arabidopsis thaliana the main flux of TAG synthesis (between 60 and 93%) is through PC and the majority of newly synthesised fatty acids in soybean embryos are incorporated into glycerolipids via PC acyl editing rather than the Kennedy pathway , . In castor the relative importance of acyl-CoA dependent and independent pathways and the proportions of de novo synthesised DAG versus PC-derived DAG contributing to TAG synthesis are unknown.
Although the activity of RcDGAT2 and RcPDAT1 go some way to explaining how 18:1-OH accumulates to such a large extent in castor seeds, there is still much to understand. This includes answers to questions such as: does the acyl-CoA pool size in developing seeds reflect the final oil composition (which might suggest little selectivity in TAG-synthetic enzymes); are there specific acyl-CoA pools used for TAG synthesis; is there metabolic channelling of intermediates in TAG-biosynthetic protein complexes and are there seed-specific enzyme isoforms for the synthesis of tri-ricinolein? Comparison of gene expression between developing castor seeds and other tissues, which can make TAG, but do not contain 18:1-OH, may help to pinpoint enzymes involved in a 18:1-OH specific TAG pathway if it exists.
We have previously generated ESTs from a normalised, subtracted castor seed library and obtained 4901 non-redundant sequences (EMBL/Genbank GE632284–GE637184). Normalisation via subtraction allows greater sequencing depth by compensating for highly expressed sequences but comparative information, which could be highly informative in selecting important genes in tissue specific metabolism, is lost. In this whole transcriptome sequencing study we sought to obtain quantitative gene expression data from a variety of castor tissues, selected on the basis of differing lipid metabolism. Developing seeds synthesise storage TAGs while germinating seeds use them as an energy source via β-oxidation and form new cellular membranes. TAG metabolism is limited in leaves but they require extensive photosynthetic membrane lipids. Male developing flowers were selected as they do contain TAGs but do not synthesise 18:1-OH. The castor genome has recently been sequenced , providing a reference genome for mapping reads from a next generation sequencing platform. Illumina's sequencing by synthesis technology was chosen for this study as it produces deep coverage of expressed sequences and reliable quantitative data . We anticipate comparison of gene expression will aid in selection of candidate genes for transformation studies to test their potential for increasing 18:1-OH levels in oilseeds.
Oil enriched in 18:1-OH is synthesised in the endosperm of developing castor seed. The fatty acid content of seed oils may directly reflect the composition of acyl-CoA pools available to TAG biosynthetic enzymes, or factors such as enzyme substrate selectivity and metabolon-formation could channel certain fatty acids into TAG. The acyl-CoA species present in developing castor bean endosperm were analysed using a fluorimetric method developed for plant tissues . The results show that 18:1-OH-CoA is not a dominant species in this tissue (Figure 2). It forms ∼4% of the detected acyl-CoA peak area at seed developmental stage III , the earliest obvious stage of endosperm formation, and the maximum value in later stages is 8% (Table 1). 18:1-OH-CoA is made from 18:1-CoA by acyl-editing on PC and the proportions of these species are broadly equivalent during stages of oil deposition (Table 1). During this period the most abundant acyl-CoA species in castor endosperm are 16:0 and 18:0, which is somewhat surprising, given that tri-ricinolein is the major TAG in castor beans. This implies that TAG synthesis must involve selective incorporation of 18:1-OH, possibly by acyl-CoA independent pathways, or the presence of a discrete acyl-CoA pool which is incorporated into TAG.
Castor oil consists of 80–90% 18:1-OH and FAME analysis of mature castor beans reveals an equivalent high proportion of this fatty acid (Table 2). Di- and tri-ricinoleoyl TAGs are synthesised in the endosperm and 18:1-OH can be detected in this tissue at an early stage of seed development (stage III, ). In an attempt to find an alternative castor tissue that makes TAG, but does not incorporate hydroxylated fatty acids into it, a number of lipid extracts were analysed (Table 2). Cotyledons from developing seeds contain substantial amounts of 18:1-OH and their lipid metabolism is thought to be similar to that of endosperm. Neither developing male flowers nor pollen however contain 18:1-OH and when neutral lipids from these were analysed by TLC a lipid with similar mobility to standard tri-olein was identified (Marked X in Figure 3). This was subsequently purified from pollen by Flash Chromatography (Figure S1) and FAME analysis revealed it contained predominantly 16:0, 18:2 and 18:3 fatty acids (Table 2, Lipid X).
To further investigate the nature of this neutral lipid species and to establish it was indeed TAG, electrospray mass spectrometry analysis in the presence of lithium was carried out . Addition of lithium enhanced formation of ion species at 830–900 amu in purified lipid-X which were not detected in other purified neutral lipids from pollen (Figure 4A, B). Collision associated dissociation of ions in this region led to formation of diagnostic fragments consistent with neutral loss of free fatty acids and fatty acid lithium salts from the parent ions, as previously described for MS analysis of TAGs . An example of such fragmentation is shown, together with the fatty acid composition of proposed TAG species which are consistent with the mass ions observed in this region of the spectrum (Figure 4C, D). We conclude that lipid X is a mixture of TAG species containing saturated and polyunsaturated fatty acids which are different from those found in castor oil. We have thus identified pollen-producing male flowers as a castor tissue which is capable of making TAG without 18:1-OH in it. Comparison of gene expression between this and developing endosperm might lead to the identification of genes and proteins involved in specific pathway(s) of 18:1-OH incorporation into TAG.
Selection of castor tissues was based on differences in their lipid metabolism, increasing the likelihood of informative results about enzymes important for seed oil synthesis. Two stages of developing seed endosperm, stages II/III and stage V/VI  were chosen as representative of early and mid-late stages of oil deposition, the level of lipid-biosynthetic transcripts might be expected to rise in the latter. Leaves synthesise fatty acids for photosynthetic and other membranes but oil does not accumulate in them to an appreciable extent. Some TAG is found in Arabidopsis leaves, but it accounts for less than 1% of lipid  and it remains to be determined whether TAG mobilisation plays an important role in this tissue. Germinating seeds were selected as they are degrading rather than synthesising storage lipids and transcriptome information from them might also be of use to those studying pathways of lipid degradation and in particular peroxisome metabolism and biogenesis. Finally, male flowers synthesise non-hydroxylated TAGs which are found in pollen.
Tissue samples from at least five different plants were pooled before RNA extraction to minimize the number of sequencing runs. RNA-Seq analysis was performed with the Illumina GAII sequencing by synthesis platform to obtain deep coverage of transcribed genes and statistically robust comparative data. The availability of a castor genome sequence provided a resource for data processing although it should be noted this was from a non-commercial castor line whereas here we use the commercial variety 99N89I. A summary of read numbers and mapping information from this study is in Table S1. Full data-sets are deposited and available at the European Nucleotide Archive (http://www.ebi.ac.uk/ena), accession number ERA047687.
RNA-Seq reads from the five castor tissues were mapped to release 0.1 of the castor assembly with expression detected for 18713 of the annotated 31221 gene models in the castor genome (see Materials and Methods). Normalised RNA-Seq fragment counts were used to measure the relative abundance of transcripts with expression reported in Fragments Per Kilobase of exon per Million fragments mapped (FPKM). Calculated FPKM expression values and results from a test statistic to determine significant changes in expression between pairs of samples are shown in Table S2. Our primary interest was in potential oil-biosynthetic enzymes in castor seeds and the results discussed below predominantly relate to lipid metabolism. Inspection of the castor genome identified 143 genes coding for enzymes of fatty acid synthesis and acyl (glycerolipid and wax) metabolism and expression data for these are in Table S3. This also includes 25 selected storage proteins, which provide a useful reference for transcriptional activity in developing seeds. A heatmap of FPKM normalised log2-transformed transcription values for genes in this table was generated (Figure S2), to aid analysis of lipid metabolic genes with similar expression patterns.
High expression of lipid-related genes in a particular tissue suggested a possible specific role in lipid metabolism and warranted further investigation. Analysis of interesting candidate enzymes included identification of the closest Arabidopsis homologue, in order to gain insight into possible gene function. Although some lipid metabolic genes have been manually annotated , designation of most castor gene models is based on automated homology- and domain-searches and more reliable functional information may be present in database entries for Arabidopsis orthologues.
Gene expression between pairs of tissues was assessed using the Cuffdiff algorithm  in order to try and identify genes important for tri-ricinolein synthesis and preferably those co-expressed with oleate 12-hydroxylase in castor seed.
Early (free nuclear, stage II/III) and later (cellular, stage V/VI) endosperm samples of developing seeds reveal which lipid metabolic genes are transcribed in a tissue synthesising tri-ricinolein, their changes during seed development and how expression relates to storage protein transcript levels. We have divided selected gene models into functional groups including fatty acid synthase, TAG synthesis and seed storage proteins for analysis (Table S3) and the main findings are discussed below.
From Table S3, under the heading of fatty acid synthesis, it can be seen that transcription of all the enzymes required for fatty acid biosynthesis from acetyl-CoA is elevated in the endosperm. Read counts for constituents of both acetyl-CoA carboxylase (ACCase) and fatty acid synthase (FAS) are substantially higher than in other tissues. Genes encoding subunits of heteromeric ACCase, (α-CT, β-CT, BC and BCCP) are much more highly expressed than those for the homomeric, multi-functional ACCase. All FAS components have at least one isoform with transcript levels comparable to heteromeric ACCase and enzyme isoforms more highly expressed in castor endosperm are easily identifiable. Thioesterase A terminates fatty acid synthesis at a chain length of 18 carbons and its 5-fold higher FPKM value compared to thioesterase B, which releases C16 fatty acids, reflects demand for C18 fatty acids for seed-oil synthesis. Stearoyl-ACP desaturase completes oleic acid synthesis in the plastid and elevated transcription of several genes encoding it occurs in the endosperm. Transcription levels for all of these enzymes in stage II/II are apparently sufficient to meet demands for fatty acid synthesis in developing castor seeds and read counts generally decrease in the later endosperm stage.
The situation is different for the oleate 12 hydroxylase (Table S3 - Fatty acid modification) where transcription nearly doubles in the later stage. Differential regulation of oleate 12-hydroxylase and the related oleate 12-desaturase genes occurs during seed development; expression of the latter decreases in stage V/VI and there is 6-fold difference in calculated FPKM values at this stage, compared to a 2 fold difference at the earlier stage. Our data for these two genes is in agreement with previous expression profiling in castor seeds , . If this transcriptional control is reflected at the protein and enzyme activity level, there would be decreased competition between hydroxylation and desaturation reactions in later stage endosperm. The putative cytochrome P450 (29791.m000529) is highlighted here due to high ratios of FPKM values in endosperm:leaf and endosperm:male flower. The Arabidopsis orthologue of this is a fatty acid (omega1) hydroxylase and while it is likely this enzyme is involved in cutin or suberin synthesis in castor, its substrates and products are unknown at present.
Fatty acids synthesised in the plastid are exported and activated to acyl-CoA thioesters by acyl-CoA synthetase (ACS) enzymes, alternatively known as fatty acid CoA ligases. Transcription of seven castor ACS genes was detected and one, 29908.m006186, is clearly the most highly expressed in endosperm, with an FPKM value of 232 in both developmental stages (Table S3 - Acyl-CoA activation). This encodes a homologue of long chain acyl-CoA synthetase 9 (LACS9) from Arabidopsis, which has recently been shown to functionally overlap with the ER-localised LACS1 in seed-oil synthesis . Acyl-CoAs are thought to be bound by acyl-CoA-binding proteins as they move between organelles, such as the plastid and endoplasmic reticulum, and expression of the castor acyl-CoA binding protein encoded by gene 29827.m002594 is highest in the endosperm.
The first steps of the acyl-CoA dependent Kennedy pathway are catalysed by glycerol-3-phosphate acyltransferase (GPAT) and lysophosphatidic acid acyltransferase (LPAT). No clear GPAT candidate with a specific role in seed TAG synthesis emerges from the RNA-Seq data as read counts for predicted GPATs are relatively low in the two developing endosperm stages (Table S3 - Kennedy pathway). GPAT9 (30122.m000357) is the most likely to be important in castor oil synthesis as it has the highest FPKM value amongst GPATs in endosperm, although equivalent expression in other tissues is evident. Read counts for 27810.m000646, the most highly expressed LPAT in the endosperm, suggest an involvement in storage oil metabolism but again, it is also transcribed to an appreciable level in other tissues. Somewhat surprisingly gene 29666.m001430, the closest orthologue of LPAT genes encoding seed-specific microsomal LPATs found in species that accumulate unusual fatty acids in their seed oil , , has low transcript levels in castor endosperm.
Phosphatidic acid phosphatase (PAP) converts PA to diacylglycerol (DAG) in the Kennedy pathway. Multiple PAP types and isoforms exist in plants but their exact role in TAG biosynthesis is unclear. No PAP candidate for TAG synthesis was identified from in this study and the castor homologues of Arabidopsis PAH1 and 2, which are expressed strongly in seeds , are transcribed in all tissues. The final step in TAG assembly by the Kennnedy pathway is the acylation of DAG by diacylglycerol acyltransferase (DGAT). In castor, single-copy gene models encode two classes of ER-associated DGAT as well as an orthologue of a soluble DGAT in peanut , , . Expression of membrane-bound RcDGAT2 - 29682.m000581, is substantially higher than the two other DGATs in developing castor seeds (Table S3).
Removal of 18:1-OH from PC and formation of 18:1-OH-CoA may be critical steps in efficient tri-ricinolein synthesis in castor seeds. Lysophosphatidylcholine acyltransferase (LPCAT) catalyses these reactions but expression of candidate genes for this enzyme, 30170.m014002 or 30174.m008937, is broadly equivalent in all tissues (Table S3 - Acyl editing/alternative TAG synthesis). A better candidate for this reaction from these results is the membrane-bound O-acyltransferase (MBOAT) protein encoded by 30190.m011126, which has significantly raised FPKM values in endosperm. This gene belongs to a sub-family of MBOATs containing lysophospholipid acyltransferases with a preference for lysophosphatidylcholine , .
Formation of 18:1-OH-CoA may be catalysed by a lipase followed by ACS. Our data indicate that the LACS9 orthologue 29908.m006186 is probably important in acyl-CoA synthesis in castor seeds, but it is harder to identify a lipase that could produce 18:1-OH. Two triacylglycerol lipases, 29935.m000048 and 30183.m001305, have specific strong expression in endosperm with transcript levels increasing at the later developmental stage (Table S3 - Phospholipases/TAG lipases). Gene 29935.m000048 has previously been characterised as an acidic lipase of castor beans which is active on TAG  and although these two genes may not produce enzymes which remove sn-2 acyl groups from PC, their expression patterns suggests a role in seed development or lipid accumulation. An additional lipase candidate, 28885.m000108, was also strongly expressed in early endosperm. The protein encoded by this gene has homology to an Arabidopsis GDSL lipase (GLIP), but it remains to be determined if it has a phospholipase activity that may release modified fatty acids for TAG synthesis.
Analysis of the seven castor genes related to phospholipid:diacylglycerol acyltransferase (PDAT) shows PDAT1A and PDAT2 (29912.m005286 and 29991.m000626 respectively) have the highest FPKM values and are expressed most strongly and specifically in the endosperm (Table S3 - Acyl editing/alternative TAG synthesis). It has recently been demonstrated that castor PDAT1A increases hydroxylated fatty acid amounts in transgenic seed oil , . Two other enzymes in pathways of acyl-CoA independent TAG synthesis, CDP-choline:diacylglycerol choline-phosphotransferase (CPT) and phosphatidylcholine:diacylglycerol choline-phosphotransferase (PDCT) do not have such clear seed-specific expression as PDAT1A and 2. FPKM values for PDCT (29841.m002865) in the endosperm are only approximately double those in leaf or male flowers and CPT (30138.m003845) has equivalent, relatively low-level, expression in four of the samples (Table S3). This does not immediately suggest a major role for these two enzymes in tri-ricinolein synthesis and their importance for seed-oil synthesis in castor, as has been shown for PDCT in Arabidopsis , remains to be determined.
Selected genes encoding lipid metabolic and seed storage proteins were clustered based on tissue expression patterns (Figure S2). The major clade which includes oleate-12 hydroxylase contains 58 proteins, listed in Table S4. Most of the proteins discussed above have similar expression patterns to oleate-12 hydroxylase and are contained within it, the exceptions being GPAT9 (30122.m000357), LPCAT (30170.m014002 and 30174.m008937) and CPT (30138.m003845). Genes encoding these three proteins are more ubiquitously expressed and likely to be controlled by different regulatory systems.
Storage-protein genes exhibit strong induction of expression in the endosperm and their FPKM values are among the highest in this study (Table S3 - Seed storage proteins/ricin). Oil bodies in plant seeds are surrounded by oleosin proteins and transcription of two oleosins is high in the endosperm. In contrast to most of the lipid metabolic genes, expression of storage proteins and oleosins increases in the later stage of endosperm development. Our results show that transcriptional activation of lipid synthesis occurs very early in castor seed, before development of cellular endosperm, and does not coincide with high levels of ricin or storage protein expression.
Purification of the ER from castor endosperm (stage IV/V) enabled proteomic identification of membrane-bound TAG-synthetic enzymes in both acyl-CoA dependent and independent pathways . Proteins encoded by several of the genes described above were detected in the ER, showing that gene products from loci with high endosperm FPKM values were present in this tissue. Based on the number of spectra detected in the MudPIT-type proteomic analysis, oleate-12 hydroxylase was substantially more abundant than oleate-12 desaturase in endosperm ER, reflecting the FPKM ratios observed for their genes. The putative cytochrome P450 (29791.m000529), a fatty acid ω-1 hydroxylase homologue, was also present in the ER. The most abundant ACS protein in enriched endosperm ER was the LACS9 homologue encoded by 29908.m006186. Other genes which are highly-expressed in the endosperm and encode proteins detected in enriched ER from this tissue are: GPAT9 - 30122.m000357; LPAT2 - 27810.m000646; DGAT2 - 29682.m000581; MBOAT - 30190.m011126; PDAT1A - 29912.m005286; PDCT - 29841.m002865; GLIP-1 lipase - 28885.m000108 and triacylglycerol lipase - 30183.m001305. These results support the hypothesis that analysis of gene expression in endosperm and other castor tissues may help to pinpoint enzymes involved in an 18:1-OH specific TAG pathway.
Germinating seeds degrade lipids derived from storage TAG and convert them into sugars which support early development . TAGs are first hydrolysed by lipases, releasing free fatty acids which enter peroxisomes. Here they undergo β-oxidation to form acetyl-CoA which is converted to oxaloacetate by the glyoxylate pathway. Following export from the peroxisome oxaloacetate is used in gluconeogenic reactions to synthesise carbohydrates. It is not therefore unexpected that expression of fatty acid synthetic (FAS and ACCase) genes is greatly decreased in germinating compared to developing castor seeds (Table S3 - Fatty acid synthesis). Amongst other lipid metabolic genes in castor, β-oxidation pathway components including 3-hydroxyacyl-CoA dehydrogenase, 3-ketoacyl-CoA thiolase B, acyl-CoA oxidase and acyl-CoA dehydrogenase are highly expressed in germinating seed, together with proteins involved in fatty acid desaturation in the ER: FAD2 and 3, cytochrome b5 and NADH-cytochrome b5 reductase (Table S5).
It is evident that differential regulation of ACS genes occurs in developing and germination seeds (Table S3 - Acyl-CoA activation). Transcription of ACS2, 29844.m003365, is strongly induced in germinating seed and it is clearly the dominant isoform expressed in this tissue, followed by 29732.m000322. In contrast, the FPKM value for the most highly expressed LACS in developing seed, 29908.m006186, is dramatically reduced during germination. This pattern is analogous to that seen in Arabidopsis, where two specific LACS isoforms are associated with peroxisomal fatty acid activation during germination (LACS6 and 7) and different enzymes (LACS1 and LACS9) are induced during oil synthesis in developing seeds.
Significant differences in expression of lipases between these tissues are also apparent. Transcripts for the three lipases induced in seed development (29935.m000048, 30183.m001305 and 28885.m000108) are essentially absent in the germinating seed and expression of others, such as 29884.m000182, 29884.m000183, 30167.m000881 and 28470.m000422 is increased (Table S3 - Phospholipases/TAG lipases). The latter is a homologue of the SDP1 lipase required for TAG breakdown and normal germination in Arabidopsis seeds  but the substrates of these castor lipases are not proven.
One of the most abundant transcripts in germinating seeds is that coding for isocitrate lyase (Table S6) which generates glyoxylate and succinate from isocitrate as part of the glyoxylate cycle. Other members of the glyoxylate cycle such as malate synthase, malate dehydrogenase and aconitase are also highly expressed in this tissue. Full lists of genes which have sequencing reads mapped to them in each tissue are in Table S7.
Transcripts encoding photosynthetic proteins for both light-harvesting and Calvin cycle reactions are abundant in leaf RNA (Table S6). No lipid metabolic pathway is obviously up-regulated in leaf (such as occurs with β-oxidation in germinating seed) and FPKM values for almost all of the fatty acid biosynthetic enzymes that are expressed in developing seed are substantially lower in leaf. A leaf-specific acyl carrier protein (30147.m014425) can easily be identified (Table S3 – Fatty acid synthesis) and increased expression of both palmitoyl-ACP thioesterase (29848.m004677) and plastidial desaturases (29666.m001456, 29696.m000105 and 28176.m000273) reflects an increased demand for photosynthetic membrane fatty acids. Unsurprisingly, read counts for oleate-12 hydroxylase are orders of magnitude lower in leaf than seed.
Three GPAT genes have higher expression in the leaf than the seed, one of which is the plastidial GPAT. The other two are homologues of Arabidopsis GPAT4 and 6 which were recently shown to be required for synthesis of cutin, a polyester found in the extracellular surface barrier of leaves and other plant aerial parts . Transcript levels of three TAG-synthetic acyltransferases are noticeably reduced in leaf compared to seed. Expression of two PDATs, 29912.m005286 and 29991.m000626, is low in leaf and the FPKM value for DGAT2 is reduced by 7-fold in this tissue. These three enzymes have expression patterns which suggest they are important in seed oil metabolism. Finally gene 29889.m003411, a soluble DGAT/wax synthase, is up-regulated in leaf although it is currently unclear whether its product is involved with TAG or wax synthesis.
Lipids play a number of roles in the formation and viability of pollen. TAG is present in pollen and it is thought mobilization and β-oxidation of its constituent fatty acids provides an energy source for pollen tube growth during fertilisation . Lipids such as sterol esters and TAG accumulate in the tapetal cell layer which surrounds developing pollen grains and components such as sterol esters are released and incorporated into the pollen surface when the tapetum breaks down , . Precursors required for sporopollenin and tryphine formation, derived mainly from saturated precursors such as long-chain fatty acids or long aliphatic chains, are also provided by the tapetum . The importance of lipid synthesis to pollen development is shown by the fact that knockout of either trienoic fatty acid or TAG synthesis results in non-viable pollen , 
Castor pollen and male flowers contain TAG which is enriched in 16:0, 18:2 and 18:3 fatty acids. Developing male flowers were selected as a negative type of control to aid identification of genes important for tri-ricinolein synthesis in the endosperm. Clearly there is major difference in expression of the oleate-12 hydroxylase gene, with no reads mapped in male flowers (Table S3), but differential regulation of other lipid metabolic genes is also apparent.
Expression of FAS and ACCase components is generally reduced in male flowers compared to the endosperm, although exceptions are the two FATB thioesterases which terminate fatty acid synthesis at 16 carbons (Table S3 - Fatty acid synthesis). One of the stearoyl-ACP desaturase genes (29929.m004514) is the most highly expressed fatty acid synthetic gene in male flowers, although it is not specific for this tissue and is strongly induced in endosperm as well. Elevated transcription of other desaturases is apparent. Expression of both ω-3 (Fad3) and ω-6 (Fad2) ER desaturases and a homologue of Arabidopsis FAD5, 29841.m002863, is also up-regulated in this tissue (Table S3 - Fatty acid modification), perhaps to be expected given the high proportion of polyunsaturated (18:2 and 18:3) fatty acids present (Table 2).
Two ACS isoforms, the LACS1 homologue 30076.m004616 and ACS1, 30190.m10831, have higher FPKM values in male flowers, with expression of the former specifically induced in this tissue (Table S3 – Acyl-CoA activation). ACS enzymes encoded by these genes may important for synthesising pollen lipids or surface components and a requirement for specific ACS isoforms in pollen coat formation has been demonstrated in Arabidopsis . It is notable that transcription of the LACS9 homologue 29908.m006186, which is most highly expressed in endosperm, is greatly reduced in male flowers.
A homologue of Arabidopsis GPAT6 (29969.m000267) has the highest expression of any TAG-synthetic gene with an FPKM value of 566 in male flowers. GPAT6 is involved in cutin biosynthesis and tapetum development in Arabidopsis , . Another acyltransferase with strongly elevated transcription only in male flowers, 29637.m000766 (designated as phosphatidylcholine acyltransferase in the castor database) is homologous to phospholipid sterol O-acyltransferases and is likely to be required for sterol ester synthesis in the tapetum. Several genes discussed in previous sections, with high FPKM values in the endosperm and possible roles in TAG synthesis, have reduced expression in male flowers (Table S3). This is particularly true for PDAT1 and 2, but DGAT2, MBOAT protein (30190.m01126) and PDCT have 4.6, 5.4 and 2.4-fold respectively higher transcript levels in the endosperm which suggests these enzymes may be important for tri-ricinolein synthesis in endosperm.
Tissue specific expression of four TAG lipases is evident in male flowers and endosperm (Table S3 Phospholipases/TAG lipases). Two of these, 29646.m001089 and 29646.m001091, are strongly activated in male flowers and may be associated with mobilization of pollen oil reserves during tube growth and fertilisation. Degradation of TAG during anther development in Brassicaceae has been reported  and lipases specifically expressed in male flowers could play a role in this process, releasing lipids required for pollen maturation. Transcription of TAG lipases 29935.m000048 and 30183.m001305, which were strongly transcribed in the endosperm, was not detected in male flowers. Although this is consistent with a role in tri-ricinolein synthesis, and they may be involved in re-modelling of TAGs after synthesis in developing castor bean , the function of these lipases remains unclear.
Transcription of genes encoding enzymes which elongate fatty acids and make wax, cutin and alkane precursors is increased in male flowers. 3-ketoacyl-CoA synthase (KCS) enzymes catalyse the initial condensation steps in microsomal fatty acid elongation. KCS transcripts are among the most abundant from lipid metabolic genes in male flowers (Table S5) and homologues of Arabidopsis KCS 10 and 21 (29726.m004062 and 29777.m000271) are only expressed in this tissue (Table S3 – FA elongation/wax synthesis). Other components are required to complete an elongation cycle and increased expression of microsomal 3-ketoacyl-CoA reductase and 3-hydroxyacyl-CoA dehydratase isoforms also occurs. Strong induction of 29709.m001216, a homologue of Arabidopsis CER1 required for wax synthesis and necessary for pollen development , and expression of two proposed alcohol O-fatty-acyltransferases (wax synthases), 27613.m000611 and 29629.m001370, reflect a demand for surface and cuticular lipids in developing male flowers. Homologues of several Arabidopsis lipid metabolic genes, with proven roles in pollen formation and male fertility, are significantly up-regulated in male flowers compared to the endosperm (Table S8).
Transcription factors (TFs) are DNA binding proteins that regulate gene expression which are encoded by distinct and conserved multigene families in plants. The number of genes and gene transcripts encoding members of a particular TF family may differ between plant species and specific tissues on account of specific function(s) and/or evolutionary expansion/contraction. The expression of TFs in plant seeds is of substantial interest as a route to identify potential master regulators that control genes with roles in pathways of reserve accumulation, especially lipids. Analysis of microarray datasets indicates that a number of plant lipid metabolism genes are transcriptionally co-regulated , , but specific factors controlling gene transcription for TAG metabolism in castor and other oilseeds remain to be uncovered.
A complex network, including several TFs, determines aspects of seed and embryo development . In Arabidopsis thaliana, LEAFY COTYLEDON1 (LEC1) and LEC1-LIKE (L1L) are key regulators of fatty acid biosynthesis. The transcription factor WRINKLED1 (WRI1), which belongs to the APETALA2-ethylene responsive element-binding protein (AP2-EREBP) family, controls fatty acid, but not TAG, synthesis via LEC2 and possibly LEC1  and it is likely that additional TFs are involved. Complete regulatory mechanisms controlling other lipid metabolic pathways, such cuticular wax and cutin biosynthesis, are also not known. The Arabidopsis AP2-EREBP TF WIN1/SHN1 activates cuticular wax biosynthesis  and genes involved in cutin biosynthesis such as GPAT4 and possibly LACS2 but much remains to be discovered.
We analysed expression of TF in the five castor tissues. Castor protein sequences were searched against the Pfam library of HMMs to identify Pfam domains. A custom perl script was used to identify and classify transcription factors according to rules utilised in the plant transcription factor database PlnTFDB (http://plntfdb.bio.uni-potsdam.de/v3.0/) . A total of 1310 TFs were identified in the castor genome, which can be divided into 60 families (Table S9, TF families sheet). The largest four TF families were AP2-EREBP, bHLH, MYB-related and NAC domain, with 115 (8.8%), 103 (7.9%), 97 (7.4%) and 95 (7.2%) genes respectively. BLAST P alignment against the TAIR9.0 database (expect score 1×10−5) detected Arabidopsis homologues for 1289 out of these 1310 castor TF. Only 919 Arabidopsis TF were represented in these alignments versus 1289 castor proteins, which may reflect their evolutionary relationship.
Transcripts were detected in at least one of the RNA samples for 1060 castor TF, 157 of which were specific for one tissue (Table S9, TF expression sheet). Expression of 41 TF was confined to the endosperm. In order to associate TF expression to 18:1-OH and tri-ricinolein metabolism, data for TFs in endosperm stages and male developing flowers were compared (Table S9, Endosperm – flower TF sheet). In addition to the 41 TF which were only transcribed in endosperm, 26 with endosperm FPKM values >5 were not expressed in flowers and a further 31 had this FPKM cut-off and (FPKM EII-III/FPKM Male flowers) values >10. Investigation of the targets for some of these TF may be important to determine factors linked to high levels of ricinoleic acid in castor oil.
In this study we have examined the fatty acid composition of a variety of castor tissues and shown that ricinoleic acid (18:1-OH) is present in both developing endosperm and cotyledons but absent from male developing flowers or pollen. Triacylglycerol (TAG) accumulates in both of these tissues, but its fatty acid composition is different to TAG synthesised in developing castor seeds. TAG in the seed-endosperm predominantly contains 18:1-OH, but this is absent from pollen and flower TAG which is instead mainly composed of 16:0, 18:2 and 18:3 fatty acids. Analysis of the acyl-CoAs in developing endosperm revealed that 18:1-OH-CoA is not the major acyl-CoA species. This is consistent with high levels of 18:1-OH accumulation in TAG being driven by one, or a combination of, the following: [a] restricted access to different in vivo pools of acyl-CoAs, [b] strict metabolic channelling and [c] substrate selectivity of enzymes involved in their biosynthesis.
TAG biosynthesis in castor endosperm occurs in the endoplasmic reticulum (ER). A previous proteomic study from our group identified proteins in purified endosperm ER and detected TAG-biosynthetic proteins in developing oil-seeds for the first time. We also reported quantitative RT-PCR data comparing expression in seed and leaf of selected genes potentially involved in tri-ricinolein biosynthesis. In order to extend that study and gain a better understanding of gene expression in castor, we performed RNA-Seq studies in a variety of tissues. This report focuses on genes associated with lipid metabolism and a key aim was comparison of transcription in endosperm and male developing flowers. We believe it unlikely that the different TAG compositions in these tissues is solely due to oleate-12 hydroxylase activity, since expression of this enzyme in a number of plant species does not result in synthesis of high 18:1-OH TAG. Comparison of expression between tissues allowed identification of candidate genes which may be important for tri-ricinolein synthesis in castor seed in addition to the oleate-12 hydroxylase – Table 3. Gene products from nine of these eleven genes were detected in purified endosperm ER, the site of TAG synthesis. Expression of two of these genes, DGAT2 and PDAT1A, in plants expressing oleate-12 hydroxylase elevated 18:1-OH incorporation into seed oils and co-expression of additional genes in Table 3 might increase this level further. We are currently performing pull-down and native blue PAGE experiments to identify interacting partners and complexes involved in TAG synthesis in endosperm ER, which should advance understanding of metabolic compartmentalisation in this tissue. A combination of these two approaches could lead to higher 18:1-OH levels in future transgenic plants.
A milestone in castor research was the recent sequencing of its genome . Here we report transcriptome analysis of five tissues from this species, using a paired end sequencing approach with read lengths of 120 bases. Previous castor expression studies ,  (http://www.ncbi.nlm.nih.gov/sra accession SRX007402 to SRX007408) were limited to developing seeds. High-quality RNA-Seq data allows identification and accurate quantification of transcription and the results presented here supplement information on expression of lipid metabolic genes in soybean Jatropha curcus, bitter melon  and Arabidopsis .
Our data provides sequence information that could be used to update the current castor genome assembly and identify new genes that were not originally annotated by largely automated processes. To indicate the potential for this approach, RNA-Seq reads were aligned to the reference genome and output from each of the five tissue samples separately assembled using the Cufflinks package. Resulting assemblies were then merged and compared to the JCVI reference annotation. In total the RNA-Seq reads assembled into 75090 transcripts corresponding to 29759 ‘genes’ which compares to the 31221 genes in version 0.1 of the JCVI assembly. Of the 29759 cufflinks genes, 2847 were located intergenic to the JCVI annotation and hence may represent novel genes. A total of 218147 splice junctions were identified from mapping the RNA-Seq reads, with 112337 supported by at least 10 reads. Comparison of these junctions to the JCVI annotation shows over 30000 are distinct, with ∼4000 mapping intergenic. While the Cufflinks assemblies will be incomplete and contain errors, such as incorrect merges and retained introns, it is clear that integrating the RNA-Seq data into the JCVI gene build would allow the annotation of several thousand alternative splice variants and potentially several hundred novel genes. Such integration is outside the scope of this work, but the RNA-Seq data reported here clearly provides a valuable resource for evaluation of current gene models (http://castorbean.jcvi.org) and investigation of alternative splicing or single nucleotide polymorphisms in castor.
Castor (Ricinus communis) variety 99N89I was grown in John Innes No.3 compost with a 16/8 h light/dark cycle at 23°C and 18°C. Plants were watered daily and inflorescences tagged on emergence of the first female flowers. Complete flowering stems were removed 15 to 20 or 25 to30 days after flowering and endosperm samples removed for immediate freezing in liquid nitrogen. Fruit were cut transversely and the endosperm appearance enabled developmental staging according to Greenwood and Bewley . Cotyledons for lipid analysis were dissected out of whole developing beans at stage V/VI. All material collected for RNA preparation was frozen in liquid nitrogen and stored at −80°C. For RNA extraction, equal amounts of endosperm at stages II/III (endosperm free-nuclear stage) or stages V/VI (onset of cellular endosperm development) were pooled from at least five different castor plants. Expanding true leaves, appearing after the first cotyledons and leaf-pair, were harvested when they were 10–15 cm in size. Germinating seed (cotyledon) tissue was obtained by soaking dry beans in running water overnight followed by germination in the dark for 3 days in moist vermiculite at 30°C and material from >30 separate seeds was pooled. Developing male developing flowers were harvested from at least 10 castor plants. The perianth of the male flower (comprised of sepals) was removed using a razorblade and reproductive organs, including pollen and anthers, collected.
Endosperm samples were pooled according to developmental stage from a number of different inflorescences. Mixed samples were ground in a pestle and mortar and 10–20 mg transferred into chilled eppendorfs. Acyl-CoA extraction and measurement were as reported previously using the modified mobile phase composition for extended HPLC separation , .
A ricinoleoyl-CoA standard to confirm peak identity was synthesised from an ammonium salt of ricinoleic acid (18:1-OH). This was made by adding 0.5 ml 2 M NH3 solution to 10 mg 18:1-OH (Sigma R7257), incubating at 60°C for 30 minutes and adding 0.5% Triton X-100 after cooling and removal of NH3 under nitrogen. The acyl-CoA was made in a 1 ml reaction containing 0.1 M Tris/HCl pH 7.5, 10 mM MgCl2, 10 mM ATP, 1 mg 18:1-OH, 0.25% Triton X-100, 10 mM CoASH and 1 U acyl-CoA synthetase (Sigma). After incubation for 3 hour at 30°C, 18:1-OH-CoA was purified using a 1 ml Sep-Pak C18 Plus column (Waters) which had been equilibrated with 6 ml methanol and 6 ml Tris/HCl pH 7.5. The reaction was loaded and the column washed with 1.5 ml Tris/HCl pH 7.5 and 1.5 ml 50% MeOH before elution with 5 ml HPLC-grade MeOH. Progression of synthesis and eluted products were checked by TLC on silica K6F plates (Whatman) developed with butanol/acetic acid/water (5∶2∶3). Methanol was removed by vacuum centrifugation, the 18:1-OH-CoA resuspended in 10 mM NaOAc pH5.9 and concentration determined by A260. Samples ranging from 1 to 100 pmoles were analysed as described above.
Lipid extraction from castor tissues used the Folch method . Pollen was incubated in chloroform/methanol 2∶1 at 50°C for 10 minutes before extraction at room temperature, while other tissues were homogenised in this solvent to start the procedure. Aliquots of the final extracts were dried and fatty acid composition determined by fatty acid methyl ester (FAME) analysis as described .
Total lipids were prepared from 4.5 ml volume pollen using the Folch method. Inhibition of lipases with isopropanol before extraction did not alter the lipid profiles observed. Lipid-X was purified with two Flash chromatography runs on silicic acid columns (100–200 mesh Sigma) developed with hexane/diethylether/acetic acid (70∶30∶1): see Figure S1. Eluting fractions were analysed by thin layer chromatography on silica gel 60 aluminium-backed plates (Merck) developed in the solvent system above and fractions containing lipid-X pooled. FAME analysis was as for the tissue lipid extracts. For analysis by mass spectrometry, purified lipids were re-suspended in methanol/chloroform (4∶1) and 2.5 mM lithium acetate added before static infusion into a hybrid quadropole-TOF mass spectrometer (QStar Pulsar i, Applied Biosystems). Selected ions were fragmented by collision induced dissociation following introduction of nitrogen and products analysed by time of flight.
The castor tissues for RNA-Seq analyses were ground to powder in liquid nitrogen and total RNA was prepared using a minor modification of the instructions of the ToTALLY RNA Kit (Ambion). To selectively deplete abundant rRNA molecules, mRNA was enriched using the MicroPoly(A)Purist Kit with two rounds of oligo dT purification (Ambion). The mRNA quality was checked on a Nano chip of the Bioanalyser 2100 (Agilent) and the quantity measured with the Qubit RNA kit and Qubit fluorometer (Invitrogen). Transcriptome libraries were constructed using the Illumina mRNA Seq sample preparation kit with some modifications. In brief, purified mRNA was fragmented by addition of 5× fragmentation buffer (Illumina, Hayward, CA) and was heated for 2 min at 94°C in a thermocycler. A shorter fragmentation time was used to yield library size of 350–400 bp. First strand cDNA was synthesised using random primers to eliminate the general bias towards 3′ end of the transcript. Second strand cDNA synthesis was done by adding GEX second strand buffer (Illumina, Hayward, CA), dNTPs, RNaseH and DNA polymerase I followed by incubation for 2.5 h at 16°C. Second strand cDNA was further subjected to end repair, A-tailing, and adapter ligation in accordance with the manufacturer supplied protocols. Purified cDNA templates were enriched by 15 cycles of PCR for 10 s at 98°C, 30 s at 65°C, and 30 s at 72°C using PE1.0 and PE2.0 primers and with Phusion DNA polymerase (Illumina, Hayward, CA). The samples were cleaned using QIAquick PCR purification columns and eluted in 30 µl EB buffer as per manufacturer's instructions (QIAGEN, CA). Purified cDNA libraries were quantified using Bioanalyser DNA 100 Chip (Agilent). All libraries were sequenced on the Illumina Genome Analyzer II as paired end 120 bp reads following the manufacturer's recommendations.
The Illumina processing pipeline version 1.3 was employed for processing of raw images to make base calls and generate sequence reads, the default quality filter (CHASTITY≥0.6) was applied. FASTQ files were generated for each of the five tissues. The number of paired end reads that passed the quality filter for each tissue is shown in Table S1 total reads. Sequence data has been submitted to the ENA sequence read archive (SRA) under accession ERA047687.
Reads for each sample were aligned to the TIGR castor bean WGS assembly release 0.1 (http://castorbean.jcvi.org/index.php) using Tophat v1.2.0 . Tophat aligns RNA-Seq data to the genome first using Bowtie  to identify coverage islands. In this initial step reads are split into smaller segments and mapped independently. The resulting coverage islands are used to generate a junction database that is supplemented with junctions directly inferred from paired end data. Additionally we supplied Tophat with all annotated junctions in the TIGR castor bean genome release 0.1 though the –G parameter. Tophat was run with default alignment parameters, allowing multiple mappings per read (40 default) and 2 mismatches in each segment alignment, the max intron length was changed to 20,000 bp.
Differential expression testing was carried out using the Cuffdiff program from the Cufflinks package version 0.9.3 . Transcript abundance was determined for the gene models annotated in the TIGR castor bean genome release 0.1 using the Tophat RNA-Seq BAM alignments for each of the five tissue samples. Cuffdiff reports an expression value for each transcript and gene in Fragments Per Kilobase of exon model per Million mapped fragments (FPKM). A test statistic is also calculated which after Benjamini-Hochberg correction for multiple-testing  was used to determine significant changes in expression between each pair of samples.
Purification of lipid for GC and MS analysis.
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Heatmap of selected lipid metabolism and storage protein genes differentially expressed in five castor tissue samples.
Click here for additional data file (pone.0030100.s002.pdf)
Summary of sequence read mapping.
Click here for additional data file (pone.0030100.s003.doc)
Expression values of castor gene models and significant differences between castor tissues.
Click here for additional data file (pone.0030100.s004.xls)
Read counts of candidate genes for fatty acid, glycerolipid and wax metabolism and for storage proteins.
Click here for additional data file (pone.0030100.s005.xls)
Genes in the heatmap made from lipid synthetic and storage proteins that are co-expressed with oleate hydroxylase in Ricinus tissues.
Click here for additional data file (pone.0030100.s006.xls)
Expression of lipid metabolic genes in castor germinating seeds, developing endosperm and male flowers.
Click here for additional data file (pone.0030100.s007.xls)
Castor genes with the highest FPKM values in the five castor tissues.
Click here for additional data file (pone.0030100.s008.xls)
Castor gene models ordered on FPKM values in individual tissues.
Click here for additional data file (pone.0030100.s009.xls)
Lipid metabolism genes with proven roles in male gametophyte and floral development.
Click here for additional data file (pone.0030100.s010.doc)
Expression of castor transcription factors.
Click here for additional data file (pone.0030100.s011.xls)
Competing Interests: This study was partly funded by a LINK grant with Linnaeus, Inc. and Arkema as industrial partners. This does not alter the authors' adherence to all the PLoS ONE policies on sharing data and materials, as detailed online in the guide for authors..
Funding: This work was funded by the Biotechnology and Biological Sciences Research Council (http://www.bbsrc.ac.uk/), grant no. BB/F004354/1, a LINK grant with Linnaeus, Inc. and Arkema as industrial partners. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
|1.||Bafor M,Smith MA,Jonsson L,Stobart K,Stymne S. Year: 1991Ricinoleic acid biosynthesis and triacylglycerol assembly in microsomal preparations from developing castor-bean (Ricinus communis) endosperm.Biochem J2805075141747126|
|2.||van de Loo FJ,Broun P,Turner S,Somerville C. Year: 1995An oleate 12-hydroxylase from Ricinus communis L. is a fatty acyl desaturase homolog.Proc Natl Acad Sci USA92674367477624314|
|3.||Smith MA,Jonsson L,Stymne S,Stobart K. Year: 1992Evidence for cytochrome b5 as an electron donor in ricinoleic acid biosynthesis in microsomal preparations from developing castor bean (Ricinus communis L.).Biochem J2871411441417766|
|4.||Broun P,Boddupalli S,Somerville C. Year: 1998A bifunctional oleate 12-hydroxylase: desaturase from Lesquerella fendleri.Plant J132012109680976|
|5.||Smith MA,Moon H,Chowrira G,Kunst L. Year: 2003Heterologous expression of a fatty acid hydroxylase gene in developing seeds of Arabidopsis thaliana.Planta21750751614520576|
|6.||Napier JA. Year: 2007The production of unusual fatty acids in transgenic plants.Annu Rev Plant Biol5829531917472567|
|7.||Brown AP,Slabas AR,Rafferty JB. Year: 2009Fatty acid biosynthesis in plants – metabolic pathways, structure and organization.Wada H,Murata NLipids in Photosynthesis: Essential and Regulatory Functions: Springer1134|
|8.||Lassner MW,Levering CK,Davies HM,Knutzon DS. Year: 1995Lysophosphatidic acid acyltransferase from meadowfoam mediates insertion of erucic acid at the sn-2 position of triacylglycerol in transgenic rapeseed oil.Plant Physiol109138913948539298|
|9.||Brough CL,Coventry JM,Christie WW,Kroon JTM,Brown AP,et al. Year: 1996Towards the genetic engineering of triacylglycerols of defined fatty acid composition: Major changes in erucic acid content at the sn-2 position affected by the introduction of a 1-acyl-sn-glycerol-3-phosphate acyltransferase from Limnanthes douglasii into oil seed rape.Mol Breeding2133142|
|10.||Kroon JT,Wei W,Simon WJ,Slabas AR. Year: 2006Identification and functional expression of a type 2 acyl-CoA:diacylglycerol acyltransferase (DGAT2) in developing castor bean seeds which has high homology to the major triglyceride biosynthetic enzyme of fungi and animals.Phytochemistry672541254917084870|
|11.||Burgal J,Shockey J,Lu CF,Dyer J,Larson T,et al. Year: 2008Metabolic engineering of hydroxy fatty acid production in plants: RcDGAT2 drives dramatic increases in ricinoleate levels in seed oil.Plant Biotechnol J681983118643899|
|12.||Bates PD,Ohlrogge JB,Pollard M. Year: 2007Incorporation of newly synthesized fatty acids into cytosolic glycerolipids in pea leaves occurs via acyl editing.J Biol Chem282312063121617728247|
|13.||Bates PD,Durrett TP,Ohlrogge JB,Pollard M. Year: 2009Analysis of acyl fluxes through multiple pathways of triacylglycerol synthesis in developing soybean embryos.Plant Physiol150557219329563|
|14.||Li-Beisson Y,Shorrosh B,Beisson F,Andersson MX,Arondel V,et al. Year: 2010Acyl-Lipid Metabolism.165 The Arabidopsis Book: The American Society of Plant Biologists.|
|15.||Stymne S,Stobart AK. Year: 1984Evidence for the reversibility of the acyl-CoA:lysophosphatidylcholine acyltransferase in microsomal preparations from developing safflower (Carthamus tinctorius L.) cotyledons and rat liver.Biochem J2233053146497849|
|16.||Dahlqvist A,Stahl U,Lenman M,Banas A,Lee M,et al. Year: 2000Phospholipid: diacylglycerol acyltransferase: An enzyme that catalyzes the acyl-CoA-independent formation of triacylglycerol in yeast and plants.Proc Natl Acad Sci USA976487649210829075|
|17.||van Erp H,Bates PD,Burgal J,Shockey J,Browse J. Year: 2011Castor phospholipid: diacylglycerol acyltransferase facilitates efficient metabolism of hydroxy fatty acids in transgenic Arabidopsis.Plant Physiol15568369321173026|
|18.||Kim HU,Lee KR,Go YS,Jung JH,Suh MC,et al. Year: 2011Endoplasmic reticulum-located PDAT1-2 from castor bean enhances hydroxy fatty acid accumulation in transgenic plants.Plant Cell Physiol5298399321659329|
|19.||Slack CR,Campbell LC,Browse JA,Roughan PG. Year: 1983Some evidence for the reversibility of the cholinephosphotransferase-catalysed reaction in developing linseed cotyledons in vivo.Biochim Biophys Acta7541020|
|20.||Lu C,Xin Z,Ren Z,Miquel M,Browse J. Year: 2009An enzyme regulating triacylglycerol composition is encoded by the ROD1 gene of Arabidopsis.Proc Natl Acad Sci USA106188371884219833868|
|21.||Bates PD,Browse J. Year: 2011The pathway of triacylglycerol synthesis through phosphatidylcholine in Arabidopsis produces a bottleneck for the accumulation of unusual fatty acids in transgenic seeds.Plant J doi:10.1111/j.1365-313X.2011.04693.x.|
|22.||Chan AP,Crabtree J,Zhao Q,Lorenzi H,Orvis J,et al. Year: 2010Draft genome sequence of the oilseed species Ricinus communis.Nat Biotechnol2895195620729833|
|23.||Metzker ML. Year: 2010Sequencing technologies - the next generation.Nat Rev Genet11314619997069|
|24.||Larson TR,Graham IA. Year: 2001Technical Advance: a novel technique for the sensitive quantification of acyl CoA esters from plant tissues.Plant J2511512511169187|
|25.||Greenwood JS,Bewley JD. Year: 1982Seed development in Ricinus communis (castor bean) I. Descriptive morphology.Can J Bot6017511760|
|26.||Hsu FF,Turk J. Year: 1999Structural characterization of triacylglycerols as lithiated adducts by electrospray ionization mass spectrometry using low-energy collisionally activated dissociation on a triple stage quadrupole instrument.J Am Soc Mass Spectr10587599|
|27.||Yang Z,Ohlrogge JB. Year: 2009Turnover of fatty acids during natural senescence of Arabidopsis, Brachypodium, and switchgrass and in Arabidopsis beta-oxidation mutants.Plant Physiol1501981198919561121|
|28.||Trapnell C,Williams BA,Pertea G,Mortazavi A,Kwan G,et al. Year: 2010Transcript assembly and quantification by RNA-Seq reveals unannotated transcripts and isoform switching during cell differentiation.Nat Biotechnol2851151520436464|
|29.||Lu CF,Wallis JG,Browse J. Year: 2007An analysis of expressed sequence tags of developing castor endosperm using a full-length cDNA library.BMC Plant Biol742 doi:10.1186/1471-2229-7-42. 17672910|
|30.||Chen GQ,Turner C,He X,Nguyen T,McKeon TA,et al. Year: 2007Expression profiles of genes involved in fatty acid and triacylglycerol synthesis in castor bean (Ricinus communis L.).Lipids4226327417393231|
|31.||Zhao L,Katavic V,Li F,Haughn GW,Kunst L. Year: 2010Insertional mutant analysis reveals that long-chain acyl-CoA synthetase 1 (LACS1), but not LACS8, functionally overlaps with LACS9 in Arabidopsis seed oil biosynthesis.Plant J641048105821143684|
|32.||Brown AP,Brough CL,Kroon JTM,Slabas AR. Year: 1995Identification of a cDNA that encodes a 1-Acyl-sn-glycerol-3-phosphate acyltransferase from Limnanthes douglasii.Plant Mol Biol292672787579178|
|33.||Testerink C,Munnik T. Year: 2011Molecular, cellular, and physiological responses to phosphatidic acid formation in plants.J Exp Bot622349236121430291|
|34.||Eastmond PJ,Quettier AL,Kroon JTM,Craddock C,Adams N,et al. Year: 2010PHOSPHATIDIC ACID PHOSPHOHYDROLASE1 and 2 regulate phospholipid synthesis at the endoplasmic reticulum in Arabidopsis.Plant cell222796281120699392|
|35.||He X,Turner C,Chen GQ,Lin JT,McKeon TA. Year: 2004Cloning and characterization of a cDNA encoding diacylglycerol acyltransferase from castor bean.Lipids3931131815357018|
|36.||Saha S,Enugutti B,Rajakumari S,Rajasekharan R. Year: 2006Cytosolic triacylglycerol biosynthetic pathway in oilseeds. Molecular cloning and expression of peanut cytosolic diacylglycerol acyltransferase.Plant Physiol1411533154316798944|
|37.||Shindou H,Shimizu T. Year: 2009Acyl-CoA:lysophospholipid acyltransferases.J Biol Chem2841518718904|
|38.||Stahl U,Stalberg K,Stymne S,Ronne H. Year: 2008A family of eukaryotic lysophospholipid acyltransferases with broad specificity.FEBS Lett58230530918154737|
|39.||Eastmond PJ. Year: 2004Cloning and characterization of the acid lipase from castor beans.J Biol Chem279455404554515322116|
|40.||Brown AP,Kroon JT,Topping JF,Robson JL,Simon WJ,et al. Year: 2011Components of complex lipid biosynthetic pathways in developing castor (Ricinus communis) seeds identified by MudPIT analysis of enriched endoplasmic reticulum.J Proteome Res103565357721657795|
|41.||Graham IA. Year: 2008Seed storage oil mobilization.Annu Rev Plant Biol5911514218444898|
|42.||Quettier A-L,Eastmond PJ. Year: 2009Storage oil hydrolysis during early seedling growth.Plant Physiol Biochem4748549019136267|
|43.||Yang W,Pollard M,Li-Beisson Y,Beisson F,Feig M,et al. Year: 2010A distinct type of glycerol-3-phosphate acyltransferase with sn-2 preference and phosphatase activity producing 2-monoacylglycerol.Proc Natl Acad Sci USA107120401204520551224|
|44.||Footitt S,Dietrich D,Fait A,Fernie AR,Holdsworth MJ,et al. Year: 2007The COMATOSE ATP-binding cassette transporter is required for full fertility in arabidopsis.Plant Physiol1441467148017468211|
|45.||Hsieh K,Huang AHC. Year: 2005Lipid-rich tapetosomes in Brassica tapetum are composed of oleosin-coated oil droplets and vesicles, both assembled in and then deatched from the endoplasmic reticulum.Plant J4388989916146527|
|46.||Hsieh K,Huang AHC. Year: 2007Tapetosomes in Brassica tapetum accumulate endoplasmic reticulum-derived flavonoids and alkanes for delivery to the pollen surface.Plant Cell1958259617307923|
|47.||Ariizumi T,Toriyama K. Year: 2011Genetic regulation of sporopollenin synthesis and pollen exine development.Annu Rev Plant Biol6243746021275644|
|48.||McConn M,Browse J. Year: 1996The critical requirement for linolenic acid is pollen development, not photosynthesis, in an Arabidopsis mutant.Plant Cell840341612239389|
|49.||Zhang M,Fan J,Taylor DC,Ohlrogge JB. Year: 2009DGAT1 and PDAT1 acyltransferases have overlapping functions in Arabidopsis triacylglycerol biosynthesis and are essential for normal pollen and seed development.Plant Cell213885390120040537|
|50.||Jessen D,Olbrich A,Knufer J,Kruger A,Hoppert M,et al. Year: 2011Combined activity of LACS1 and LACS4 is required for proper pollen coat formation in Arabidopsis.Plant J doi:10.1111/j.1365-313X.2011.04722.x.|
|51.||Li-Beisson Y,Pollard M,Sauveplane V,Pinot F,Ohlrogge J,et al. Year: 2009Nanoridges that characterize the surface morphology of flowers require the synthesis of cutin polyester.Proc Natl Acad Sci USA106220082201319959665|
|52.||Li XC,Zhu J,Yang J,Zhang GR,Xing WF,et al. Year: 2011Glycerol-3-phosphate acyltransferase 6 (GPAT6) is important for tapetum development in Arabidopsis and plays multiple roles in plant fertility.Molecular Plant doi:10.1093/mp/ssr057.|
|53.||Mancha M,Stymne S. Year: 1997Remodelling of triacylglycerols in microsomal preparations from developing castor bean (Ricinus communis L.) endosperm.Planta2035157|
|54.||Aarts MG,Keijzer CJ,Stiekema WJ,Pereira A. Year: 1995Molecular characterization of the CER1 gene of Arabidopsis involved in epicuticular wax biosynthesis and pollen fertility.Plant Cell7211521278718622|
|55.||Schmid M,Davison TS,Henz SR,Pape UJ,Demar M,et al. Year: 2005A gene expression map of Arabidopsis thaliana development.Nat Genet3750150615806101|
|56.||Peng FY,Weselake RJ. Year: 2011Gene coexpression clusters and putative regulatory elements underlying seed storage reserve accumulation in Arabidopsis.BMC Genomics1228621635767|
|57.||Santos-Mendoza M,Dubreucq B,Baud S,Parcy F,Caboche M,et al. Year: 2008Deciphering gene regulatory networks that control seed development and maturation in Arabidopsis.Plant J5460862018476867|
|58.||Baud S,Mendoza MS,To A,Harscoet E,Lepiniec L,et al. Year: 2007WRINKLED1 specifies the regulatory action of LEAFY COTYLEDON2 towards fatty acid metabolism during seed maturation in Arabidopsis.Plant J5082583817419836|
|59.||Broun P,Poindexter P,Osborne E,Jiang CZ,Riechmann JL. Year: 2004WIN1, a transcriptional activator of epidermal wax accumulation in Arabidopsis.Proc Natl Acad Sci USA1014706471115070782|
|60.||Kannangara R,Branigan C,Liu Y,Penfield T,Rao V,et al. Year: 2007The transcription factor WIN1/SHN1 regulates cutin biosynthesis in Arabidopsis thaliana.Plant Cell191278129417449808|
|61.||Perez-Rodriquez P,Riano-Pachon DM,Correa LGC,Rensing SA,Kersten B,et al. Year: 2009PlnTFDB: updated content and new features of the plant transcriptiuon factor database.Nucleic Acids Res38 doi:10.1093/nar/gkp805.|
|62.||Severin AJ,Woody JL,Bolon YT,Joseph B,Diers BW,et al. Year: 2010RNA-Seq Atlas of Glycine max: a guide to the soybean transcriptome.BMC Plant Biol10160 doi:10.1186/1471-2229-10-160. 20687943|
|63.||Costa GG,Cardoso KC,Del Bem LE,Lima AC,Cunha MA,et al. Year: 2010Transcriptome analysis of the oil-rich seed of the bioenergy crop Jatropha curcas L.BMC genomics11462 doi:10.1186/1471-2164-11-462. 20691070|
|64.||Yang P,Li X,Shipp MJ,Shockey JM,Cahoon EB. Year: 2010Mining the bitter melon (momordica charantia l.) seed transcriptome by 454 analysis of non-normalized and normalized cDNA populations for conjugated fatty acid metabolism-related genes.BMC plant biology10250 doi:10.1186/1471-2229-10-250. 21080948|
|65.||Beisson F,Koo AJK,Ruuska S,Schwender J,Pollard M,et al. Year: 2003Arabidopsis genes involved in acyl lipid metabolism. A 2003 census of the candidates, a study of the distribution of expressed sequence tags in organs, and a Web-based database.Plant Physiol13268169712805597|
|66.||Larson TR,Edgell T,Byrne J,Dehesh K,Graham IA. Year: 2002Acyl CoA profiles of transgenic plants that accumulate medium-chain fatty acids indicate inefficient storage lipid synthesis in developing oilseeds.Plant J3251952712445123|
|67.||Folch J,Lees M,Sloane Stanley GH. Year: 1957A simple method for the isolation and purification of total lipides from animal tissues.J Biol Chem22649750913428781|
|68.||Trapnell C,Pachter L,Salzberg SL. Year: 2009TopHat: discovering splice junctions with RNA-Seq.Bioinformatics251105111119289445|
|69.||Langmead B,Trapnell C,Pop M,Salzberg SL. Year: 2009Ultrafast and memory-efficient alignment of short DNA sequences to the human genome.Genome Biol10R25 doi:10.1186/gb-2009-10-3-r25. 19261174|
|70.||Benjamini Y,Hochberg Y. Year: 1995Controlling the false discovery rate - a practical and powerful approach to multiple testing.J Roy Stat Soc B Met57289300|
Table 1 Acyl-CoA levels in developing castor endosperm.
|Stagea||18:1-OH CoA||18:1 CoA||16:0 CoA||18:0 CoA||18:2 CoA|
|III||4.25 (0.34)||7.18 (0.89)||32.25 (3.04)||31.73 (4.36)||7.52 (0.62)|
|IV||5.17 (0.84)||6.59 (0.42)||23.74 (1.88)||31.34 (0.35||6.21 (0.64)|
|V||4.90 (0.22)||7.97 (0.52)||33.05 (3.21)||26.65 (4.44)||4.45 (0.31)|
|VI||8.01 (0.89)||6.61 (0.71)||28.89 (3.25)||28.08 (1.23)||7.89 (1.71)|
|VII||3.82 (0.42)||5.17 (0.38)||33.63 (0.40)||31.35 (4.09)||4.68 (0.50)|
|VIII||7.03 (0.37)||4.23 (0.51)||27.93 (1.49)||26.12 (3.45)||4.13 (0.74)|
The percentage of the total acyl-CoA peak area represented by specified acyl-CoAs are shown for castor endosperm stages during seed development. Analysis of fluorescent acyl-CoA derivatives was as in  and average values from three extractions and analyses for each stage are shown, together with the standard error in brackets (n = 3).
aEndosperm samples were staged according to .
Table 2 Fatty acid analysis of castor tissues and purified lipid X.
|Fatty acid||Early endo-sperm (4)||Mature bean (2)||Cotyledon (4)||Developing ♂ Flowers (2)||Pollen (2)||Lipid X (2)|
Average peak-area percentages of fatty acid methyl ester derivatives are listed. The number of separate sample extractions and analyses are listed in brackets, except for lipid X, where the result from two GC injections of the purified fraction is shown.
aItalicised entries are probable fatty acid designations, although the species did not exactly co-chromatograph with standards.
bIndicates the identity of the molecular species is not known but likely wax or hydroxylated fatty acid derivatives from pollen protective layers.
Table 3 Candidate genes that may be important in tri-ricinolein synthesis based on RNA-Seq data.
|Gene ID||Gene Identifier||Detected in ER|
|27810.m000646||Lysophosphatidic acid acyltransferase (LPAT2)||x|
|29682.m000581||Type 2 diacylglycerol acyltransferase (DGAT2)||x|
|29791.m000529||Cytochrome P450, Putative||x|
|29841.m002865||Phosphatidylcholine diacylglycerol cholinephosphotransferase (PDCT)||x|
|29908.m006186||Long-Chain-Fatty-Acid CoA Ligase, Putative (ACS9)||x|
|29912.m005286||Phospholipid:diacylglycerol acyltransferase 1 (PDAT1A)||x|
|29935.m000048||Triacylglycerol Lipase, Putative|
|29991.m000626||Phosphatidylcholine:Diacylglycerol Acyltransferase 2 (PDAT2)|
|30183.m001305||Triacylglycerol Lipase, Putative||x|
|30190.m011126||O-Acyltransferase (Membrane Bound) Domain Containing Protein||x|
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