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. 2019 Jun 20;180(4):1962–1974. doi: 10.1104/pp.19.00600

The Plastid Lipase PLIP1 Is Critical for Seed Viability in diacylglycerol acyltransferase1 Mutant Seed1,[OPEN]

Karanbir Aulakh 1, Timothy P Durrett 1,2,3
PMCID: PMC6670099  PMID: 31221730

Global transcript profiling of developing Arabidopsis dgat1 seed reveals a critical role for the plastid lipase PLIP1 in providing polyunsaturated substrates for triacylglycerol synthesis.

Abstract

In developing Arabidopsis (Arabidopsis thaliana) seeds, the synthesis of triacylglycerol (TAG) is mediated primarily by the acyl-CoA-dependent enzyme diacylglycerol acyltransferase1 (DGAT1). In the absence of DGAT1 activity, phospholipid:diacylglycerol acyltransferase (PDAT1) plays an important role in TAG synthesis, consistent with the higher-than-expected oil content and altered fatty acid composition of dgat1 seed. Transcript profiling of developing wild type (Columbia-0) and dgat1-1 mutant seed identified 602 differentially expressed genes. Expression of genes important for the formation of phosphatidylcholine, including LYSOPHOSPHATIDYLCHOLINE ACYLTRANSFERASE2, and REDUCED OLEATE DESATURATION1 were strongly upregulated, consistent with increased substrate supply for PDAT1. In addition, several genes lacking a defined role in TAG biosynthesis were also upregulated, including the α/β-hydrolase family gene PLIP1, which encodes a plastid-localized lipase. In most tissues, PLIP1 was expressed at equivalent levels in wild-type and dgat1 plants, except for developing seed, where transcript levels were higher in the dgat1 mutant. Seeds from plip1 mutant plants possessed a 20% reduction in oil content and were smaller than seed from wild-type plants. Crosses between dgat1 and plip1 failed to generate double-homozygous mutant plants. Reciprocal crossing with wild-type plants demonstrated that both male and female gametophytes could transmit the dgat1 plip1 double-mutant genotype. Double-homozygous dgat1 plip1 seed formed but was green and failed to germinate. The synthetic lethal phenotype of dgat1 with plip1 indicates an important role for PLIP1 in the absence of DGAT1 activity, likely by supplying polyunsaturated fatty acid substrates for PDAT1.

Triacylglycerols (TAGs), neutral lipids with three fatty acids esterified to a glycerol backbone, are the major constituent of the seed oil produced by most plants. The highly reduced nature of the fatty acids means that TAGs are one of the most energy-dense products synthesized by plants, making them an ideal energy storage reserve during germination and early stages of seedling development. This high-energy density of TAG also means that plant oils have long represented a valuable source of calories for human and animal nutrition. More recently, societal concerns have spurred an interest in obtaining renewable and carbon-neutral fuels and chemicals from plant oils (Durrett et al., 2008; Dyer et al., 2008), expanding the utility and value of TAG. The synthesis of TAG in oil seeds therefore is important for many aspects of human nutrition and economic activity.

The synthesis and modification of fatty acids and their subsequent incorporation into TAG occurs in both the plastid and the endoplasmic reticulum (ER). In plastids, acetyl-CoA carboxylase (ACC) converts acetyl-CoA to malonyl-CoA, which is then used to synthesize fatty acids through multiple rounds of condensation, reduction, and dehydration reactions. Each round of synthesis adds two carbons to the growing fatty acid chain, which is connected to an acyl carrier protein (ACP). The introduction of the first double bond can also occur in the plastid, while the fatty acid is still conjugated to ACP, the FATTY ACID BIOSYNTHESIS2 (FAB2) ∆9-desaturase converts stearoyl (18:0)-ACP (fatty acids are abbreviated as X:Y, where X indicates the number of carbon atoms and Y represents the number of double bonds) to oleoyl (18:1)-ACP. Acyl-ACP thioesterases then terminate fatty acid synthesis by removing ACP from the nascent fatty acid, which is then exported from the plastid and conjugated to CoA. Most of these newly synthesized acyl-CoA molecules are used by ER-localized lysophosphatidylcholine acyltransferases (LPCATs) to acylate lysophosphatidylcholine (LPC) to form phosphatidylcholine (PC), a membrane lipid (Bates et al., 2009; Wang et al., 2012). Oleate (18:1) can be further desaturated while conjugated to PC through the activity of FATTY ACID DESATURASE2 (FAD2), a ∆12-desaturase, and FAD3, a ∆15-desaturase, to form the polyunsaturated fatty acids (PUFA) linoleate (18:2) and linolenate (18:3), respectively.

De novo incorporation of fatty acids into glycerolipids also occurs in the ER and involves the sequential acylation of a glycerol-3-phosphate backbone. Glycerol-3-phosphate acyltransferase, encoded by GPAT9, adds the first acyl group (Shockey et al., 2016; Singer et al., 2016). The product of this reaction, lysophosphatidic acid is further acylated by lysophosphatidic acyltransferases to form phosphatidic acid. Phosphatidic acid phosphohydrolases (PAH) then remove the phosphate group to form diacylglycerol (DAG), a key intermediate for both the synthesis of phospholipids as well as TAGs. In addition to the de novo DAG formed through this pathway, labeling experiments have suggested the presence of a second, PC-derived DAG pool that could be formed by multiple routes (Bates et al., 2009; Bates and Browse, 2011). For example, phosphatidylcholine:diacylglycerol cholinephosphotransferase (PDCT) transfers a phosphocholine head group between PC and DAG and plays a key role in the flux of PUFA into TAGs in Arabidopsis (Arabidopsis thaliana) seeds (Lu et al., 2009; Bates et al., 2012). In addition, PC can be converted to DAG through phospholipase C or the reverse activity of CDP-choline:diacylglycerol cholinephosphotransferase (Slack et al., 1983).

In Arabidopsis seeds, the final acylation of DAG to form TAG occurs primarily through the activity of the acyl-CoA:diacylglycerol acyltransferase DGAT1 (Lung and Weselake, 2006). Based on mutant phenotypes, DGAT1 plays a key role in the synthesis of TAG in Arabidopsis seeds. Arabidopsis dgat1 mutants possess a 25% to 45% reduction in seed oil content, as well as an altered fatty acid composition, with higher levels of 18:3 and lower levels of eicosenoic acid (22:1; Katavic et al., 1995; Routaboul et al., 1999; Zou et al., 1999). In addition to DGAT activity, phospholipid:diacylglycerol acyltransferases (PDATs), encoded by PDAT1 in Arabidopsis, can also synthesize TAG by transferring the acyl group at the sn-2 position of phospholipids to the sn-3 position of DAG (Dahlqvist et al., 2000; Ståhl et al., 2004). In contrast to dgat1, Arabidopsis pdat1 mutants do not possess any changes in oil content or fatty acid composition, suggesting an insignificant role in TAG synthesis in wild-type seeds relative to DGAT1 (Mhaske et al., 2005). However, DGAT1 and PDAT1 possess overlapping functions in TAG synthesis in seeds and pollen. A double dgat1 pdat1 mutant genotype in pollen prevents the formation of oil bodies and results in abnormal and sterile pollen. Importantly, silencing of PDAT1 using RNA interference in the dgat1 background results in a 70% to 80% decrease in oil content, demonstrating that PDAT1 plays a key role in TAG biosynthesis in the absence of DGAT1 function (Zhang et al., 2009). The same study demonstrated that other candidate acyltransferases present in Arabidopsis seed did not possess DGAT activity or lacked any seed oil phenotype when mutated, either individually or in combination with DGAT1.

The altered fatty acid composition and the importance of PDAT1 for TAG synthesis in dgat1 seed argue that developing Arabidopsis seed respond to the lack of DGAT1 activity. Given the importance of TAG production, we undertook a transcript profiling approach to better understand how mutant seeds respond to the inability to synthesize TAG through DGAT1. The use of RNA sequencing (RNA-Seq) offered advantages over previous microarray studies (Xu et al., 2012), including higher signal-to-noise ratios, a large dynamic range of expression levels and the ability to identify novel transcripts (Wang et al., 2009). Through this approach, we identified differentially expressed genes whose altered expression is consistent with the seed phenotypes observed in the dgat1 mutant. Importantly, we were not limited by the choice of transcripts detected by the standard Arabidopsis gene chip, enabling us to identify genes not previously associated with TAG biosynthesis in seeds. Mutations in one of these genes, PLIP1, that encodes a plastidial lipase, resulted in minor changes in oil content, fatty acid composition and seed size. However, when combined with mutations in dgat1, these mutations resulted in seed that failed to germinate. This synthetic lethal phenotype demonstrates the importance of PLIP1 in TAG accumulation, particularly when DGAT1 function is impaired.

RESULTS

Altered Lipid Composition and Gene Expression in Developing dgat1-1 Seeds

To identify differentially expressed genes in dgat1-1 important for regulating TAG accumulation, we isolated developing seeds during middle to late embryogenesis when storage lipid synthesis begins (Mansfield and Briarty, 1992). Similar to previous reports (Katavic et al., 1995), we observed a 35% reduction in the neutral lipid content of developing dgat1-1 seeds relative to those from Col-0 wild-type plants (Supplemental Fig. S1A), as well as increased polar lipids (Supplemental Fig. S1B). Developing mutant seed contained more polyunsaturated TAG molecular species and less molecular species with 56 or more acyl carbons (Supplemental Fig. S1C), consistent with the increased levels of 18:3 and lower levels of 20:1 characteristic of dgat1 seed (Katavic et al., 1995; Routaboul et al., 1999). In addition, PC levels were higher in dgat1-1 seed, with more polyunsaturated molecular species at 16 d after flowering (DAF; Supplemental Fig. S1, D and E).

Having confirmed the phenotype of the developing mutant seeds, we extracted RNA from the seeds at 8, 12, and 16 DAF, which corresponded to globular, torpedo, and mature embryo stages, respectively (Supplemental Fig. S2). Global changes in transcript levels across seed development were then quantified using RNA-Seq. After quality assessment and normalization, more than 95% of the reads mapped to genic regions of the Arabidopsis genome (Supplemental Table S1). We used the empirical analysis of differentially expressed genes test to identify differentially expressed (DE) genes with greater than 2-fold change (P < 0.05) that were expressed at levels higher than 0.1 reads per kilobase per million mapped reads. With these filtering parameters, the total number of statistically significant DE genes increased during seed development, with 7-fold more DE genes at 16 DAF compared to 8 DAF (Fig. 1A). We further classified the genes according to whether they were associated with lipid metabolism using existing detailed annotation (Fig. 1B; Li-Beisson et al., 2013).

Figure 1.

Figure 1.

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Differentially expressed (DE) genes in dgat1-1 mutant seed. A, Venn diagram showing the number of DE genes in dgat1-1 relative to wild type during different stages of seed development. Circle sizes are proportional to number of genes. Numbers in red and green refer to lipid-related and non-lipid-related genes, respectively. DAF, Days after flowering. B, Manhattan plot with distribution of DE genes across different chromosomes. The y axis shows log2 fold change of expression in dgat1-1 relative to Col-0, and the x axis indicates the chromosomal position of genes. The horizontal dotted lines represent the threshold for log2 fold change for selecting DE genes. Green and red colors represent lipid-related and non-lipid-related genes, whereas filled and open circles represent DE and non-DE genes, respectively.

Desaturases Are Strongly Up-regulated in dgat1

Analysis of genes associated with lipid biosynthetic pathways (Li-Beisson et al., 2013) revealed a striking up-regulation of a number of desaturases. For example, the transcript levels of the plastidial stearoyl-ACP desaturase genes FAB2 and DES2 were both 2-fold higher in dgat1-1; expression changes in their relatively uncharacterized homolog DES6 were even more extreme, with more than 6-fold up-regulation in dgat1-1 evident at 16 DAF (Fig. 2A; Supplemental Table S2). Interestingly, however, the expression of FAD4, FAD5, FAD6, and FAD7, all of which encode desaturases that introduce double bonds in various plastidial glycerolipids, were unaffected in dgat1-1 (Supplemental Table S2). Like FAB2 and DES2, the transcript levels of FAD2 and FAD3 encoding the ER-localized desaturases responsible for the synthesis of 18:2 and 18:3 were 50% higher in dgat1-1 at 16 DAF, making them the highest expressed lipid biosynthetic genes in the mutant seed. We confirmed the up-regulation of FAD2 in dgat1-1 using reverse transcription quantitative PCR (RT-qPCR), obtaining results highly consistent with those seen using RNA-Seq (Supplemental Fig. S3A).

Figure 2.

Figure 2.

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Expression profiles of lipid-related genes in developing seeds. Expression levels of selected lipid-related genes as determined by RNA-Seq in developing wild-type (WT) and dgat1-1 seed at 8, 12, and 16 d after flowering (DAF). Genes involved in acyl desaturation (A), fatty acid synthesis (B), TAG synthesis (C), phospholipid synthesis (D), and TAG hydrolysis and β-oxidation (E) are shown. For some enzyme activities, the expression levels of multiple encoding gene isoforms is summed. Data represents the mean of three biological replicates ± sd. *P < 0.05; **P < 0.01 (Student’s t test).

We observed expression changes in other lipid biosynthetic pathways, but with the few exceptions described below, few genes underwent changes on the level of the desaturases. For example, as exemplified by the expression of genes involved in ACCase or acyl-ACP thioesterase activity, genes encoding components of fatty acid synthesis were expressed about 20% higher at 12 DAF and 40% higher at 16 DAF in dgat1 (Fig. 2B). The expression profiles of the regulatory proteins PII and WRINKLED1 followed similar patterns, consistent with their role in controlling fatty acid synthesis (Cernac and Benning, 2004; Baud et al., 2007, 2010). Given the lower oil content of dgat1-1, we also examined expression of different enzymes involved in TAG hydrolysis and β-oxidation. Here, the expression of most genes was unchanged or slightly lower in dgat1-1 (Fig. 2E).

Increased Expression of Genes Important for PC Metabolism in dgat1 Seed

Of particular interest, we observed no large differences in expression of genes involved in the de novo synthesis of DAG in developing dgat1-1 seeds. GPAT9 and genes encoding lysophosphatidic acyltransferase activity were slightly upregulated at different time points; combined expression of PAH genes was lower in dgat1-1 seed at 16 DAF (Fig. 2C). Consistent with previous work (Zou et al., 1999), we were able to detect expression of the mutant dgat1-1 transcript, which was slightly higher at 12 DAF but otherwise unchanged. Particularly surprisingly, the low transcript levels of PDAT1 in wild type seeds were unchanged in dgat1-1. Analysis of PDAT1 expression using RT-qPCR confirmed that the gene was expressed at the same low levels in wild-type and dgat1-1 seed (Supplemental Fig. S3B).

In contrast to the relatively low changes in expression of de novo DAG synthesis, ROD1 encoding PDCT activity that interconverts DAG and PC was almost 2-fold higher in dgat1-1 at 16 DAF (Fig. 2D). Similarly, LPCAT2 was also upregulated in dgat1-1, though its closely related homolog LPCAT1 was significantly down regulated at 16 DAF. Consistent with an increased role of flux through PC, NMT1 encoding the methyltransferase responsible for the de novo synthesis of PC was also strongly upregulated in dgat1-1.

Selection and Confirmation of Genes Upregulated in dgat1 Seed

When selecting candidate genes for further characterization, we decided to focus on genes upregulated more than 2-fold for at least one time point. In addition, the role of the gene product needed to be relatively unknown. With these criteria, we identified four genes of particular interest. Two encoded lipases for which little was known at the beginning of the study. PLIP1 (At3g61680) is a member of the large α,β-hydrolase gene family, of which some members play a role in lipid metabolism (Ghosh et al., 2009; James et al., 2010; Vijayakumar et al., 2016). Expression of PLIP1 is similar in dgat1-1 and wild-type seeds at 8 and 12 DAF but is more than 2-fold increased in the mutant at 16 DAF (Fig. 3A). This pattern of expression was confirmed using RT-qPCR (Fig. 3B). The second lipase is encoded by NPC6 (At3g48610), a member of the nonspecific phospholipase C gene family. This lipase gene is expressed at lower levels than PLIP1. During wild-type seed development, NPC6 transcript levels decrease from 8 to 12 DAF and then remain stable to 16 DAF. In contrast, after an initial decrease in dgat1-1 seeds, NPC6 expression increases at 16 DAF (Fig. 3A). As with the other genes of interest, RT-qPCR was used to confirm the expression patterns of NPC6 seen with RNA-Seq (Fig. 3B).

Figure 3.

Figure 3.

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Expression levels of differentially expressed genes in developing seeds. Expression levels of PLIP1, MFT, and NPC6 in developing wild-type (WT) and dgat1-1 seed were quantified using RNA-Seq (A) and RT-qPCR (B). For RT-qPCR, gene expression was normalized to the reference genes ASAR1, UBC21, and PP2AA3. Data presented is the mean ± sd of three biological replicates. *P < 0.05 (Student’s t test).

We also were intrigued by the expression of MOTHER OF FT (MFT; At1g18100), which is upregulated more than 2-fold at both 12 and 16 DAF in dgat1-1 (Fig. 3A). MFT has previously been shown to be important for germination (Xi et al., 2010), but a role in TAG accumulation has not yet been demonstrated. As with all the other genes analyzed in this manner, RT-qPCR confirmed the higher expression of MFT in dgat1-1 (Fig. 3B). Finally, given its strong up-regulation in dgat1-1 (Fig. 2D), we were also curious to see if NMT1 was important for TAG biosynthesis.

Mutant plip1 Seeds Are Smaller and Possess Reduced Oil Content

Homozygous T-DNA mutant lines containing insertions in these DE genes were obtained. For all the candidate genes, we identified at least two different mutant alleles (Supplemental Fig. S4). Given our subsequent interest in PLIP1, we sequenced the flanking regions of the T-DNA insertions in the two alleles of this gene to confirm the location of the mutations (Supplemental Fig. S4A). Consistent with their T-DNA insertions in the 5′ and 3′ untranslated regions of PLIP1, previous characterization suggests that both plip1 mutant alleles are leaky with transcript detected in both mutant lines (Wang et al., 2017). Based on expression levels, plip1-2 appears to be the stronger allele and therefore was prioritized in many of our subsequent experiments. All homozygous mutants were then grown alongside wild type and dgat1-1 until maturity, and seeds were collected. The fatty acid content of seeds from both plip1-1 and plip1-2 plants was significantly lower than wild-type seed with levels similar to that of dgat1-1 plants (Fig. 4A). The fatty acid composition of both plip1 mutant lines was also distinct from wild type and dgat1-1, with slightly lower levels of 18:1 and slightly higher levels of 18:3 compared to wild type (Fig. 4B). When harvesting and analyzing the seeds produced by the different mutant lines, we noticed that a substantial proportion of the seeds from both plip1-1 and plip1-2 plants were smaller than typical wild-type seeds (Fig. 5A). When quantified, wild-type seed had an average size of 93,100 µm2. Consistent with our initial observations, the mean seed size of plip1-1 seeds was 84,500 µm2 and that of plip1-2 seeds was 83,534 µm2 (Fig. 5B).

Figure 4.

Figure 4.

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Fatty acid content and composition of mutant seed. Fatty acid content (A) and composition (B) of wild-type (WT) and mutant seed were quantified by whole-seed acid-catalyzed transmethylation of dry seeds followed by gas chromatography. Values are the mean ± sd of seeds harvested from three individual plants. *P < 0.05 (A, Student’s t test; B, two-way ANOVA with Holm-Sidak correction for multiple comparisons). The results shown are representative of two independent experiments.

Figure 5.

Figure 5.

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plip1 mutants produce smaller seed than wild-type (WT) plants. A, Representative seed from WT, plip1-1, and plip1-2 plants. Bars represent 1 mm, arrows indicate smaller or unusual plip1 seed. B, Size distribution for seeds harvested from WT, plip1-1, and plip1-2 plants. Data represents the mean of three biological replicates.

While the oil content from all three npc6 mutant lines was lower compared to wild-type plants, none of the reductions were statistically significant. Similarly, the fatty acid composition of the npc6 mutant seed was essentially the same as wild type (Fig. 4B). The seed oil content of mft-2 plants was ∼80% that of wild type; however, that of mft-3 was essentially the same as that of wild type. Both mft-2 and mft-3 possessed lower amounts of 18:1 and slightly higher levels of 18:2 compared to wild type (Fig. 4B). Finally, both nmt1 mutant lines produced seed with a 10% reduction in oil, but the fatty acid composition was unchanged (Supplemental Fig. S5).

PLIP1 Is Upregulated in dgat1-1 Seeds

As PLIP1 transcripts are not detected by the ATH1 GeneChip used by the Arabidopsis eFP Browser (Winter et al., 2007), we quantified the expression of PLIP1 in different tissues of wild-type plants to determine whether the gene played a role in tissues other than seeds. PLIP1 transcripts were detected in all tissues examined but were more abundant in flowers, siliques, and roots (Fig. 6). We also measured PLIP1 transcript levels in dgat1-1 tissues to investigate whether the gene was upregulated in different organs of the plant in this mutant background. Here, we observed a more than 2-fold increase in PLIP1 expression in siliques, consistent with our results from developing seeds (Fig. 3). However, in all the other tissues, PLIP1 expression did not differ between wild type and dgat1-1 (Fig. 6), suggesting that any role for PLIP1 in dgat1 is seed specific.

Figure 6.

Figure 6.

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PLIP1 is upregulated in siliques of dgat1-1. Expression profile of PLIP1 in different tissues of wild-type (WT) and dgat1-1 plants. The expression of PLIP1 was quantified using RT-qPCR and normalized to the expression of the reference gene ASAR1. Data represents the mean ± sd for three biological replicates. *P < 0.05 (Student’s t test).

The dgat1 plip1 Double Mutant Is Synthetically Lethal

In addition to quantifying the oil content of mutants in candidate genes, we also crossed some of the mutant lines with dgat1-1 to determine the role of the upregulated genes in a dgat1 background. Double mutants homozygous for dgat1-1 and mft-2, mft-3, or npc6-1 were successfully isolated and the seed fatty acid content quantified. There was no difference in the fatty acid content and composition between these double mutant lines and the dgat1-1 single mutant (Supplemental Fig. S6), indicating that DGAT1 is epistatic to NPC6 and MFT.

Interestingly, after genotyping 87 plants, we did not obtain any double homozygous dgat1-1 plip1-2 mutants in the F2 generation. Similarly, the genotyping of 63 F2 plants derived from crosses between dgat1-1 and plip1-1 also failed to identify any double-homozygous mutants (Supplemental Fig. S7). To confirm this observation, we collected seed from dgat1-1/dgat1-1 PLIP1/plip1-2 and DGAT1/dgat1-1 plip1-2/plip1-2 plants and genotyped the resulting progeny. Again, no double-homozygous mutants were obtained from either parental genotype (Fig. 7). For the plant with genotype dgat1-1/dgat1-1 PLIP1/plip1-2, we observed a segregation ratio close to 1:2 for dgat1-1/dgat1-1 PLIP1/PLIP1 and dgat1-1/dgat1-1 PLIP1/plip1-2, respectively. Similarly, for the plant with genotype DGAT1/dgat1-1 plip1-2/plip1-2, we observed a segregation ratio of approximately 1:2 for DGAT1/DGAT1 plip1-2/plip1-2 and DGAT1/dgat1-1 plip1-2/plip1-2, respectively. These 1:2 ratios observed for the detected genotypes and the absence of any double homozygous mutants provides additional evidence that the combination of dgat1 and plip1 genotypes is lethal.

Figure 7.

Figure 7.

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A dgat1 plip1 genotype is synthetically lethal. Genotyping results of progeny derived from self-pollinated dgat1-1/dgat1-1 PLIP1/plip1-2 (A) and DGAT1/dgat1-1 plip1-2/plip1-2 (B) plants.

Interestingly, seed from dgat1-1/dgat1-1 PLIP1/plip1-2 plants possessed a lower oil content than dgat1-1 seed, with higher levels of 18:3 and reduced 18:1 and 18:2 content (Supplemental Fig. S8). The oil content of seeds from plants homozygous for plip1-2 and heterozygous for dgat1-1 was lower than that of plip1-2 seed, consistent with the gene dosage effect previously observed for dgat1-1 heterozygotes (Katavic et al., 1995).

Both Male and Female dgat1-1 plip-2 Gametophytes Are Viable

We considered three scenarios that might cause lethality of the dgat1-1 plip1-2 double mutant: (1) that pollen or egg cells with a dgat1-1 plip1-2 genotype are nonviable, (2) that DGAT1 and PLIP1 are required for normal embryo development, or (3) that both genes are required for seed germination. To test the first hypothesis of gametophyte viability, a series of crosses were made with wild-type plants. When wild-type plants were the pollen donor and dgat1-1/dgat1-1 PLIP1/plip1-2 was the maternal plant, both the expected genotypes DGAT1/dgat1-1 PLIP1/plip1-2 and DGAT1/dgat1-1 PLIP1/PLIP1 were obtained in a 1:1 ratio (Fig. 8A). The DGAT1/dgat1-1 PLIP1/plip1-2 genotype could only result from a dgat1-1 plip1-2 egg cell, thus indicating that a double-mutant genotype does not affect the viability of the female gametophyte. Similar results were obtained when DGAT1/dgat1-1 plip1-2/plip1-2 plants were used as the maternal plant (Fig. 8B). When dgat1-1/dgat1-1 PLIP1/plip1-2 plants were the pollen donor and wild type was used as female receptacle, both the expected genotypes DGAT1/dgat1-1 PLIP1/plip1-2 and DGAT1/dgat1-1 PLIP1/PLIP1 were obtained in a 1:1 ratio (Fig. 8C). The DGAT1/dgat1-1 PLIP1/plip1-2 genotype could only result from dgat1-1 plip1-2 pollen, thus indicating that the double mutant does not affect pollen viability. Similar results were obtained when DGAT1/dgat1-1 plip1-2/plip1-2 was used as the pollen donor, providing additional evidence that dgat1-1 plip1-2 pollen is viable. (Fig. 8D). Together, these results suggest that the inability to obtain a double homozygous genotype is not the result of nonviable male or female gametophytes.

Figure 8.

Figure 8.

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Test of genetic transmission of dgat1-1 and plip1-2 gametophytes by reciprocal crossing. Genotyping results of progeny derived from reciprocal crossing between dgat1-1/dgat1-1 PLIP1/plip1-2 (A and B) and wild-type (WT) plants and DGAT1/dgat1-1 plip1-2/plip1-2 (C and D) and wild-type plants.

The dgat1-1 plip1-2 Double Mutant Fails to Germinate

No obvious abortion of seeds in siliques was observed, and developing seeds in dgat1-1/dgat1-1 PLIP1/plip1-2 or DGAT1/dgat1-1 plip1-2/plip1-2 siliques all appeared normal (Supplemental Fig. S9), suggesting the inability to obtain a double-homozygous mutant is not caused by defects during embryogenesis. To validate the existence of double-mutant seed, we genotyped individual seeds derived from dgat1-1/dgat1-1 PLIP1/plip1-2 or DGAT1/dgat1-1 plip1-2/plip1-2 plants. We were able to identify seed homozygous for both mutations (Fig. 9A), confirming that such seeds are produced. These double-homozygous mutant seeds represented one-quarter of the total seeds from both parental genotypes (Table 1), consistent with the normal genetic transmission of both mutant alleles.

Figure 9.

Figure 9.

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Double mutant dgat1-1 plip1-2 seed is germination defective. A, Genotyping of individual seeds from a self-pollinated DGAT1/dgat1-1 plip1-2/plip1-2 plant. Arrows show allele-specific PCR products from primer combinations indicated to the right of each gel. Asterisks highlight double homozygous mutant dgat1-1/dgat1-1 plip1-2/plip1-2 genotypes. B, Seeds from self-pollinated dgat1-1/dgat1-1 PLIP1/plip1-2, dgat1-1/dgat1-1 PLIP1/ PLIP1 plants, DGAT1/dgat1-1 plip1-2/plip1-2, and DGAT1/ DGAT1 plip1-2/plip1-2 plants were germinated on plates. Arrows indicate seed that failed to germinate. C, Individual seeds from self-pollinated dgat1-1/dgat1-1 PLIP1/plip1-2 plants were photographed under a microscope and then genotyped. Six representative seeds from each genotype are shown. Bars indicate 1 mm.

Table 1. Genotypes of individual seed from double-mutant plants.

Parent Genotype Seed Genotype Expecteda Observeda χ2
dgat1-1/dgat1-1 PLIP1/plip1-2 dgat1-1/dgat1-1 PLIP1/PLIP1 9 10 0.167 (P = 0.920)
dgat1-1/dgat1-1 PLIP1/plip1-2 18 17
dgat1-1/dgat1-1 plip1-2/plip1-2 9 9
DGAT1/dgat1-1 plip1-2/plip1-2 DGAT1/DGAT1 plip1-2/plip1-2 10 6 2.15 (P = 0.341)
DGAT1/dgat1-1 plip1-2/plip1-2 20 23
dgat1-1/dgat1-1 plip1-2/plip1-2 10 11
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a

Based on the genotyping of 36 seeds from dgat1-1/dgat1-1 PLIP1/plip1-2 and 40 seeds from DGAT1/dgat1-1 plip1-2/plip1-2.

We therefore tested the ability of the double-homozygous mutant seeds to germinate. When seeds from dgat1-1/dgat1-1 PLIP1/plip1-2 or DGAT1/dgat1-1 plip1-2/plip1-2 genotypes were germinated on plates, approximately one-quarter of the seeds failed to germinate (Fig. 9B). No defects were noted in the germination of plip1-2 and dgat1-1 seed. Closer observation revealed that almost all the seeds that failed to germinate were green. Genotyping of green seed revealed that they were homozygous mutant for both DGAT1 and PLIP1 (Fig. 9C). Genotyping of the resulting seedlings from dgat1-1/dgat1-1 PLIP1/plip1-2 resulted in a 1:2 ratio of dgat1-1/dgat1-1 PLIP1/PLIP1 and dgat1-1/dgat1-1 PLIP1/plip1-2, respectively. Likewise, genotyping of the resulting seedlings from DGAT1/dgat1-1 plip1-2/plip1-2 resulted in 1:2 ratio of DGAT1/DGAT1 plip1-2/plip1-2 and DGAT1/dgat1-1 plip1-2/plip1-2, respectively. Together, based on the segregation ratios of all seed (Table 1), the green phenotype and the genotype of the nongerminating seed, and the genotypes of the germinating seedlings, we conclude that the quarter of seeds that failed to germinate possess the dgat1-1 plip1-2 genotype.

DISCUSSION

The synthesis of TAG in developing seeds is a complex mechanism that involves the movement of substrates between different pathways and organelles. Not only is TAG biosynthesis valuable for agricultural production, but it is also intrinsically connected with seed development, with seeds unable to convert DAG to TAG possessing aberrant morphology and low germination rates (Zhang et al., 2009). Given the value of TAG synthesis at these different levels, we obtained global gene expression profiles to gain insights into how developing dgat1-1 mutant seeds respond to reduced levels of these important storage lipids.

Altered Pathways Provide Substrate for PDAT

Our results demonstrate that in the absence of functional DGAT1, the synthesis of TAG is adjusted to provide increased substrate, particularly PC, so that PDAT1 can now synthesize the TAG found in dgat1 seed. The absence of any lipid phenotype in pdat1 mutant seed implies that the enzyme plays a very minor role in TAG accumulation when DGAT1 is present (Mhaske et al., 2005). In contrast, the lethal phenotype of dgat1 pdat1 double mutants argues that PDAT1 is responsible for TAG synthesis in dgat1 mutants (Zhang et al., 2009). Consistent with this, in dgat1 mutant seed, the expression of specific genes is upregulated to increase supply of substrate for PDAT1 to compensate for the lack of DGAT1. In particular, we observed increased expression of genes involved in the formation of PC, which acts as the acyl donor for the formation of TAG by PDAT1, from various sources. These include LPCAT2, whose gene product regenerates PC from LPC, one of the products of the PDAT1 reaction (Fig. 2). The importance of LPCAT2 in regenerating PC for PDAT1 has been previously demonstrated, with seed from dgat1 lpcat2 double mutants possessing an oil content 35% of that of seeds from wild-type plants (Xu et al., 2012). The related LPCAT1 acyltransferase appears to play a less-important role in this PC generation for PDAT1 compared to LPCAT2. Our results indicate that the encoding gene is downregulated in dgat1 at 16 DAF (Fig. 2). This result is generally consistent with the fact that the oil content of the dgat1 lpcat1 double mutant is the same as that of dgat1 but differs from the microarray results in the same study that showed an up-regulation of LPCAT1 in dgat1 seed (Xu et al., 2012).

Other genes upregulated in developing dgat1 seed also play a role in the synthesis of PC, consistent with the increased levels of this phospholipid in the mutant (Supplemental Fig. S1D). For example, all three Arabidopsis phosphoethanolamine N-methyltransferases that methylate phosphoethanolamine to form the PC precursor phosphocholine (Chen et al., 2018) are upregulated, with NMT1 expression increased almost 5-fold in 16 DAF dgat1-1 seed (Fig. 2; Supplemental Table S2). Likewise, transcript levels of ROD1, which encodes the PDCT that interconverts DAG and PC through transfer of the choline headgroup (Lu et al., 2009) are higher in dgat1-1 seed.

In addition to increasing synthesis of PC for PDAT, dgat1 seeds also increase the proportion of lipid molecular species containing PUFA (Supplemental Fig. S1E), the preferred substrates of PDAT1 (Ståhl et al., 2004). These increased levels of polyunsaturated PC are consistent with the strong up-regulation of different desaturases (Fig. 2; Supplemental Fig. S3A), including FAD2 and FAD3, which add double bonds to 18:1 and 18:2, respectively, conjugated to PC and other phospholipids found in the ER. The plastidial desaturase FAB2 that adds double bonds to fatty acids destined for the eukaryotic pathway is also upregulated (Fig. 2), thus providing more monounsaturated substrate for FAD2 (Fig. 10). Interestingly, DES6, a homolog of FAB2, was upregulated to an even greater extent (Fig. 2). The function of DES6 is relatively unknown, with one study suggesting a role for the enzyme in increasing levels of unsaturated fatty acids in Agrobacterium-derived crown galls under hypoxic and drought conditions (Klinkenberg et al., 2014). Our results also imply a role in TAG synthesis, though additional work is needed to demonstrate this. Importantly, desaturases such as FAD4 that catalyze the synthesis of PUFA in plastidial glycerolipids are not upregulated (Fig. 2). Instead, increased desaturase activity appears focused on lipids in, or destined for, the eukaryotic pathway. In this manner, the concomitant up-regulation of FAB2, FAD2, and FAD3 leads to overall higher levels of PUFA-PC, enabling increased synthesis of TAG by PDAT1 (Fig. 10).

Figure 10.

Figure 10.

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The role of PLIP1 in TAG biosynthesis during development of wild-type (WT) and dgat1 seed. The top depicts an overview of TAG biosynthesis in wild-type seed; the bottom depicts the modified pathway in dgat1 seed. The bold arrows indicate increases in flux in the different seed. In wild-type seed, TAG is synthesized primarily through the acyl-CoA-dependent acylation of DAG by DGAT1. Polyunsaturated fatty acids (PUFA) are synthesized by FAD2 and FAD3 in the ER and incorporated into DAG through the action of PDCT. The plastid-localized lipase PLIP1 plays a minor role in the incorporation of PUFAs into PC by releasing 18:3 from 18:3/16:1-PG. In dgat1-1 seed, PDAT1 is responsible for the synthesis of TAG in the absence of DGAT1. Increased expression of the desaturases FAB2, FAD2, and FAD3 result in more polyunsaturated substrate for PDAT1. Upregulation of LPCAT2 also allows the regeneration of PC for the synthesis of TAG. In addition, PLIP1 is upregulated to increase the flow of 18:3 to PUFA-PC pool, thereby increasing the flux of PUFAs for incorporation into TAG via PDAT1.

Taken together, the alteration of gene expression in developing dgat1 seed is consistent with an increased role for PDAT activity in the absence of DGAT function. Somewhat surprisingly, therefore, PDAT1 expression was not increased in developing dgat1-1 mutant seed relative to that in wild-type seeds, both when quantified using RNA-Seq or RT-qPCR (Fig. 2C; Supplemental Fig. S3B). Previous work has demonstrated that many other potential DAG acyltransferases, including DGAT2 and PDAT-like, do not contribute to TAG synthesis in Arabidopsis seed, even in the absence of DGAT1 function (Zhang et al., 2009). In addition, while the dgat1-1 allele produces an aberrant transcript, this has been proven to encode a nonfunctional enzyme, thus demonstrating that the mutant is a complete knockout (Xu et al., 2012). In the absence of additional acyltransferases or partial activity from mutant DGAT1 protein, the likely explanation is that increases in PDAT1 activity occur posttranscriptionally in dgat1 seed. Further work is therefore needed to confirm this hypothesis and determine the nature of the regulation of PDAT1.

PLIP1 Is Essential for TAG Synthesis in Developing dgat1 Seed

We also demonstrated that MFT and NPC6, genes that have not been previously shown to play a role in TAG biosynthesis were upregulated in dgat1 seed (Fig. 3). However, the fatty acid content of npc6 mutant seed was only slightly reduced compared to wild type, and the fatty acid composition was unchanged (Fig. 4). Minor changes in mft mutant seed (Fig. 4) imply a role for the transcription factor in TAG biosynthesis, but demonstration of any direct involvement by MFT requires additional work. Further, when combined with dgat1-1, mutations in NPC6 and MFT did not result in a seed fatty acid phenotype different from dgat1-1 (Supplemental Fig. S6), indicating a minimal role for these genes in TAG synthesis.

Instead, we focused on the role of PLIP1 due to its expression pattern and phenotype of mutant alleles, particularly in combination with mutations in DGAT1. In wild-type plants, PLIP1 expression was not confined to seeds, as transcripts were detected in other tissues (Fig. 6). However, PLIP1 was not upregulated in these other tissues in dgat1 plants, consistent with the fact that DGAT1 is primarily expressed in developing seeds and suggesting that the role of PLIP1 in dgat1 mutants is seed specific. Seed from plip1 mutants possessed slightly reduced fatty acid levels compared to wild-type seed (Fig. 4A). The seed fatty acid composition was also slightly changed in the mutants, with higher levels of 18:3 and lower levels of 18:1 and 18:2. A large portion of mutant plip1 seeds were smaller than wild type (Fig. 5), providing additional support for the role of PLIP1 in seed embryo development. Importantly, we were unable to recover homozygous double dgat1 plip1 plants with both plip1 alleles, consistent with PLIP1 playing an important role in the seed of dgat1 (Fig. 7; Supplemental Fig. S7). At this point, we cannot exclude the possibility that this synthetic lethal phenotype is caused by another mutation present in the dgat1-1 mutant line. However, such a mutation would need to be closely linked to DGAT1, making such an event unlikely. While this synthetic lethal phenotype is reminiscent to that of the dgat1 pdat1 double mutant, the latter is caused by nonviable pollen cells (Zhang et al., 2009). In contrast, reciprocal crossing experiments demonstrated that both male and female gametophytes possessing a dgat1 plip1 genotype are viable (Fig. 8). Instead, dgat1 plip1 double mutant seed is formed but fails to germinate (Fig. 9), suggesting the importance of PLIP1 in seed development, particularly in a dgat1 background.

PDAT1 Is Dependent on PLIP1 for the Supply of PUFA Substrate

A recent report has demonstrated that PLIP1 encodes a plastid-localized lipase, PLIP1, that releases PUFA from phosphatidylglycerol (PG; Wang et al., 2017). These released fatty acids are incorporated into PC and eventually into TAG, providing an additional pathway for the incorporation of PUFA into TAG. Results from other studies suggest additional routes besides acyl editing for the incorporation of PUFA into TAG. In dgat1 lpcat2 mutants, where the acyl editing cycle to regenerate PC for PDAT1 is disrupted, TAG is still synthesized and contains high levels of 18:3 (Xu et al., 2012). Likewise, the seed from lpcat1 lpcat2 rod1 triple mutants still contains appreciable quantities of 18:3 (Bates et al., 2012).

Our results suggest that the PUFA substrates resulting from PLIP1 activity are required for use by PDAT1 in dgat1 seed. In wild-type seeds, PDAT1 makes only a minor contribution to the overall synthesis of TAG (Mhaske et al., 2005). However, when DGAT function is eliminated, PDAT1 activity becomes important for TAG synthesis. In dgat1 seed, therefore, increased PLIP1 expression results in more PUFA-PC for PDAT1, which is then incorporated into TAG (Fig. 10). Elimination of the source of substrate for PDAT1 in dgat1 seed results in nonviable seed (Fig. 7; Supplemental Fig. S7), similar to previous work knocking down PDAT1 expression in a dgat1 background (Zhang et al., 2009). Consistent with this idea, despite relatively unchanged levels of PG in dgat1 seed, we observe lower levels of PG 34:4, the proposed substrate of PLIP1 (Wang et al., 2017) and higher levels of 18:3 containing PC and TAG molecular species such as PC 36:6 and TAG 54:9 (Supplemental Fig. S10). Further, seed from dgat1 plants heterozygous for PLIP1 possess a lower oil content compared to dgat1 seed (Supplemental Fig. S8), providing additional evidence that PLIP1 plays a role in TAG synthesis that is independent of DGAT1. The Arabidopsis genome contains two putative paralogs of PLIP1, PLIP2 (At1g02660) and PLIP3 (At3g62590), which encode plastid-localized glycerolipid A1 lipases that release fatty acids from chloroplast membrane lipids to produce jasmonic acid (Wang et al., 2018). The expression of these two genes is downregulated in developing dgat1-1 seed (Supplemental Fig. S11), arguing against any redundancy with PLIP1 in providing substrate for PDAT. The existence of metabolically distinct PC pools has been noted (Bates et al., 2009), and it will be interesting to determine whether one of these specifically receives 18:3 via PLIP1 and is essential for PDAT activity. However, we also cannot rule out the possibility that PLIP1 activity establishes a PUFA-DAG substrate pool required by PDAT1. Multiple routes for the interconversion of DAG and PC exist; here, the up-regulation of PDCT (Fig. 2) might be relevant. Indeed, the lower oil content of plip1 mutant seed (Fig. 4) suggests that fatty acids provided by PLIP1 are incorporated in a DAG pool accessible by DGAT1 to form TAG.

CONCLUSION

Our transcript profiling results provide additional evidence that PDAT1 synthesizes TAG when DGAT1 activity is removed. In particular, the transcriptome of dgat1 seed is altered to provide more PC, the acyl donor used by PDAT1 to acylate DAG, and to increase the levels of PUFA molecular species, the preferred substrates of PDAT1. In addition, a critical role for the plastidial lipase PLIP1 in providing substrate for PDAT1 was established, adding further complexity to the flux of fatty acids from the plastid into TAG. Detailed analysis of this flux of fatty acids, as well as the purpose of different pools of DAG and PC in providing substrate for different enzymes, will assist in the mechanistic understanding of TAG synthesis in seeds.

MATERIAL AND METHODS

Plant Materials and Growth Conditions

T-DNA insertion mutant lines from the SALK (Alonso et al., 2003), SAIL (Sessions et al., 2002), and GABI-Kat (Rosso et al., 2003) collections were obtained from the Arabidopsis Biological Resource Center at the Ohio State University. Seeds were cold stratified for 2 d at 4°C before growing on soil mixture (4:2:1 mixture of Metromix Professional Growing Mix:vermiculite:perlite) in growth chambers at 23°C with 16 h light/8 h dark. For growth on plates, seeds were surface sterilized before plating on half-strength Murashige and Skoog media with 5% (w/v) phytoblend agar and 1% (w/v) Suc. When making crosses, immature flower buds were emasculated and manually cross-pollinated with pollen from the flower of the donor parent. To isolate seeds at different developmental stages, emerging flowers were tagged with cotton thread at specific time intervals. Whole siliques were flash frozen in liquid nitrogen and stored at −80°C. Seed harvesting from siliques was performed on dry ice followed by filtering with liquid nitrogen cooled sieves (Bates et al., 2013).

Seed Phenotypic Analysis

Freshly harvested seeds from different stages of development were collected from wild type and dgat1-1 plants and fixed for 1 h in ethanol:acetic acid (6:1, v/v) at room temperature. Samples were washed using 70% (v/v) ethanol, mounted in chloral hydrate:glycerol:water (8:1:2, v/v/v), and cleared for about 1 h at room temperature (Anderson, 1954; Berleth and Jurgens, 1993). Cleared samples were visualized using a Zeiss Axioplan 2 upright microscope equipped with a Lumenera Infinity 3S-1UR monochrome CCD camera. The size of mature seeds was quantified by photographing approximately 1000 seeds under a light microscope and then using ImageJ particle analysis software to process the images (Herridge et al., 2011).

Genotyping of T-DNA Lines

Genomic DNA was extracted from leaves and seed as previously described (Edwards et al., 1991). Primers used for the PCR-based genotyping of the SALK and SAIL mutant lines were designed using the SIGnAL T-DNA Express Arabidopsis Gene Mapping Tool (Alonso et al., 2003); primers for the GABI-Kat lines were designed using the GK primer tool (Huep et al., 2014). All genotyping primers used in this study are listed in Supplemental Table S3. The location of the T-DNA insertion for plip1-1 and plip1-2 was confirmed by isolating and sequencing DNA adjacent to the insertion. Chi squared (χ2) tests were used to quantify the goodness of fit between expected and observed allelic ratios.

Lipid Extraction and Quantification

Total lipids were extracted from developing wild-type and dgat1-1 seed using a hexane-isopropanol method (Li et al., 2006) with di-15:0 PC and tritridecanoin (Nu-Check Prep) as internal standards. Total lipids were loaded onto a small silica column and neutral lipids eluted with 5 mL 99:1 (v/v) chloroform:methanol. Polar lipids were then recovered with 5 mL methanol. These lipid extracts were then analyzed using electro-spray ionization mass spectrometry as described previously (Bansal and Durrett, 2016). Seed fatty acids were quantified by transmethylating intact seeds (Li et al., 2006) using tripentadecanoin (Nu-Chek Prep) as an internal standard. The resulting fatty acid methyl esters were quantified on a Shimadzu GC-2010 Plus gas chromatograph equipped with a 30 × 0.25 mm DB-23 column (Agilent) as described previously (Aznar-Moreno and Durrett, 2017).

RNA Isolation and Sequencing

Total RNA was extracted from the pooled seeds of 30 to 40 siliques using the RNeasy Plant Mini Kit (Qiagen) according to the manufacturer’s instructions, followed by additional purification with sodium citrate to improve RNA quality (Nybo, 2011). We obtained total RNA yields in the range of 1 to 3 µg from ∼50-mg seeds. Using an Agilent Bioanalyzer, we confirmed that the RNA Integrity Number for all samples was greater than 9. Total RNA (500 ng) was sequenced at the Genomics Core at the University of Kansas Medical Center. The mRNA fraction was enriched with oligo dT capture, sized, reverse transcribed into complementary DNA and ligated with the appropriate indexed adaptors using the TruSeq Stranded mRNA Sample Preparation Kit (Illumina). Following Agilent Bioanalyzer QC of the library preparation and library quantification using the Roche LightCycler 96 with the KAPA SYBR Fast Universal qPCR kit (KAPA Biosystems), the RNA-Seq libraries were adjusted to a 4-nM concentration and pooled for multiplexed sequencing. Libraries were denatured and based on qPCR results, diluted to the appropriate concentration, followed by clonal clustering onto the sequencing flow cell using the TruSeq Paired-End Cluster Kit v3-cBot-HS (Illumina). The clonal clustering procedure was automated using the Illumina cBOT Cluster Station. The clustered flow cell was sequenced on an Illumina HiSeq 2500 Sequencing System using the TruSeq SBS Kit v3-HS (Illumina) to obtain 100-bp pair-end reads, which were trimmed and quality assessed before assembly against the Arabidopsis (Arabidopsis thaliana) reference genome (Araport 11) using CLC Genomics Workbench v 7.5.1 (Qiagen). To identify DE genes, the empirical analysis of differentially expressed genes test was used, which implements the “exact test” for two-group comparisons. This test assumes that all samples have negative binomial distribution and accounts for biological variability of replicates.

RT-qPCR

Complementary DNA was synthesized from 1 µg total RNA using the QuantiTect Reverse Transcription kit (Qiagen) according to the manufacturer’s instructions. Relative mRNA levels were quantified with iTaq Universal SYBR Green Supermix (Bio-Rad) on a CFX-96 real-time PCR system (Bio-Rad) and normalized to the reference genes ASAR1, UBC21, and PP2AA3. The primers used for RT-qPCR are listed in Supplemental Table S4.

Supplemental Data

The following supplemental materials are available.

Large Data Sets

Supplemental Table S2 contains transcript levels for all expressed genes in developing wild type and dgat1-1 seed.

Accession Numbers

The sequences from this study are deposited in the Gene Expression Omnibus (GEO) under accession number GSE123303.

Acknowledgments

The authors would like to thank Kathrin Schrick and her research group for assistance with microscopy and for many helpful suggestions during the research and Dan Boyle for helping to obtain the images of developing seeds. The authors are grateful to Clark Bloomer at the Genomics Core at the University of Kansas Medical Center for advice concerning the RNA-Seq experiments. The plip1-1 mutant line was kindly provided by Christoph Benning. The authors would also like to thank Nathan Henderson for his diligent work looking after plants, harvesting seed, and analyzing fatty acid content. This is contribution 19-064-J from the Kansas Agricultural Experiment Station.

Footnotes

1

This work was partially supported by the USDA National Institute of Food and Agriculture Multistate Research Project (grant no. NC1203) and the United States Agency for International Development (under prime agreement ESP-A-00-05-00001-00) with the National Academy of Sciences (subgrant award PGA-2000003650 to T.P.D.). RNA-seq was supported by the Kansas IDea Network for Biomedical Research Excellence (grant no. GM103418), by the Kansas Intellectual and Developmental Disabilities Research Center (grant no. HD002528), and by Illumina. ESI-MS analyses were performed at the Kansas Lipidomics Research Center, where instrument acquisition and lipidomics method development was supported by the National Science Foundation (grant nos. EPS-0236913, MCB-1413036, DBI-0521587, DBI-1228622), the Kansas Technology Enterprise Corporation, K-IDeA Networks of Biomedical Research Excellence (INBRE) of the National Institutes of Health (grant no. P20GM103418), and Kansas State University.

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