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Plant Cell. Dec 2009; 21(12): 3885–3901.
PMCID: PMC2814504

DGAT1 and PDAT1 Acyltransferases Have Overlapping Functions in Arabidopsis Triacylglycerol Biosynthesis and Are Essential for Normal Pollen and Seed Development[W][OA]


Triacylglycerol (TAG) biosynthesis is a principal metabolic pathway in most organisms, and TAG is the major form of carbon storage in many plant seeds. Acyl-CoA:diacylglycerol acyltransferase 1 (DGAT1) is the only acyltransferase enzyme that has been confirmed to contribute to TAG biosynthesis in Arabidopsis thaliana seeds. However, dgat1 null mutants display only a 20 to 40% decrease in seed oil content. To determine whether other enzymes contribute to TAG synthesis, candidate genes were expressed in TAG-deficient yeast, candidate mutants were crossed with the dgat1-1 mutant, and target genes were suppressed by RNA interference (RNAi). An in vivo role for phospholipid:diacylglycerol acyltransferase 1 (PDAT1; At5g13640) in TAG synthesis was revealed in this study. After failing to obtain double homozygous plants from crossing dgat1-1 and pdat1-2, further investigation showed that the dgat1-1 pdat1-2 double mutation resulted in sterile pollen that lacked visible oil bodies. RNAi silencing of PDAT1 in a dgat1-1 background or DGAT1 in pdat1-1 background resulted in 70 to 80% decreases in oil content per seed and in disruptions of embryo development. These results establish in vivo involvement of PDAT1 in TAG biosynthesis, rule out major contributions by other candidate enzymes, and indicate that PDAT1 and DGAT1 have overlapping functions that are essential for normal pollen and seed development of Arabidopsis.


Triacylglycerols (TAGs) are major storage lipids that accumulate in developing seeds, flower petals, pollen grains, and fruits of a number of plant species (Stymne and Stobart, 1987; Murphy, 2005). TAGs from plants are important sources of human nutrition, provide precursors for chemical industries, and can serve as renewable biofuels (Weselake, 2005; Durrett et al., 2008; Dyer et al., 2008). In oilseeds, TAG bioassembly is traditionally thought to be catalyzed by membrane-bound enzymes that operate in the endoplasmic reticulum (Stymne and Stobart, 1987; Somerville et al., 2001), involving sequential acylation of the glycerol backbone via three sn-specific acyltransferases: glycerol-3-phosphate acyltransferase (EC, lyso-phosphatidic acid acyltransferase (EC, and, after removal of the phosphate group from the sn-3 position of the glycerol backbone by phosphatidate phosphatase (EC, the final acylation of sn-1,2-diacylglycerol (DAG) by diacylglycerol acyltransferase (DGAT; EC In many oilseeds, acyl chains produced in the plastid also rapidly enter phosphatidylcholine (PC) (e.g., Griffiths et al., 1988) via an acyl-editing pathway (Bates et al., 2009), allowing desaturation, hydroxylation, epoxidation, or other modifications.

The first DGAT gene was cloned from mouse and was a member of the DGAT1 family, which has high sequence similarity with sterol:acyl-CoA acyltransferase (Cases et al., 1998). A second family of DGAT genes (DGAT2) was first identified in the oleaginous fungus Morteriella ramanniana, but these have no sequence similarity with DGAT1 (Lardizabal et al., 2001). A DGAT2 has been cloned from tung tree (Vernicia fordii) and castor (Ricinus communis) (Kroon et al., 2006; Shockey et al., 2006) and appears to have a nonredundant function in TAG biosynthesis. Furthermore, the tung DGAT2 is localized to a different subdomain of the ER than the tung DGAT1 (Shockey et al., 2006).

In addition to DGAT1 and DGAT2, several other enzymes are reported to synthesize TAG. A bifunctional acyltransferase, wax ester synthase/DGAT that can use both fatty alcohols and DAGs as acyl acceptors to synthesize wax esters and TAGs, respectively, was identified from the bacterium Acinetobacter (Kalscheuer and Steinbuchel, 2003; Stoveken et al., 2005). A large number of wax ester synthase/DGAT homologs occur in plants, and some involved in cuticular wax synthesis have been characterized (King et al., 2007; Li et al., 2007). Other proposed additions to the traditional scheme of TAG biosynthesis in seeds include demonstrations that in developing castor and safflower (Carthamus tinctorius) seeds, TAG can also be generated from two molecules of DAG via a DAG:DAG transacylase (with monoacylglycerol as a coproduct) and that the reverse reaction participates in remodeling of TAGs (Lehner and Kuksis, 1996; Stobart et al., 1997), although genes encoding such enzymes have not been identified in plants. An enzyme encoding a soluble protein with DGAT activity has also been cloned from peanut (Arachis hypogaea; Saha et al., 2006).

In some species, it is apparent that TAG can also be formed by an acyl-CoA–independent enzyme, phospholipid:diacylglycerol acyltransferase (PDAT; EC, in which the transfer of an acyl group from the sn-2 position of PC to the sn-3 position of DAG yields TAG and sn-1 lyso-PC (Banas et al., 2000; Dahlqvist et al., 2000). In yeast, PDAT1 is a major contributor to TAG accumulation during the exponential growth phase (Oelkers et al., 2002). The two closest homologs to the yeast PDAT gene have been identified in Arabidopsis thaliana (Ståhl et al., 2004). It is not yet clear to what extent these enzymes may play a role in conventional TAG assembly in oilseeds. Overexpression of PDAT1 increased PDAT activity in Arabidopsis leaf and root microsomes, but no changes were found in lipid and fatty acid contents in these plants (Ståhl et al., 2004). Furthermore, Mhaske et al. (2005) isolated and characterized a knockout mutant of Arabidopsis (designated as pdat1-1), which has a T-DNA insertion in the PDAT1 locus At5g13640. In contrast with the situation in yeast, the fatty acid content and composition in seeds did not show significant changes in the mutant, suggesting that PDAT1 activity as encoded by At5g13640 may not be a major determining factor for TAG synthesis or is compensated by other reactions in Arabidopsis seeds.

We previously characterized an ethyl methanesulfonate–induced mutant of Arabidopsis, AS11 (also named tag1-1), which displayed a decrease in stored TAG and an altered fatty acid composition (Katavic et al., 1995). Since the first identification of a plant DGAT1 gene from Arabidopsis by three independent groups (Hobbs et al., 1999; Routaboul et al., 1999; Zou et al., 1999), genes encoding homologous microsomal DGAT1s have been cloned from several other plants (tobacco [Nicotiana tabacum], Bouvier-Nave et al., 2000; canola [Brassica napus], Nykiforuk et al., 2002; Castor-He et al., 2004; burning bush [Euonymus alatus], Milcamps et al., 2005; soybean [Glycine max], Wang et al., 2006; tung tree, Shockey et al., 2006; and nasturtium [Tropaeolum majus], Xu et al., 2008).

Metabolite analysis (Perry et al., 1999) and other studies (Zheng et al., 2008) suggested DGAT may be one of the rate-limiting steps in plant seed lipid accumulation. Thus, there was implied utility in manipulating the expression of DGAT genes for improving oil content. In this regard, overexpression of the Arabidopsis DGAT1 in wild-type plants led to an increase in seed oil content and seed weight (Jako et al., 2001). Subsequently, DGAT expression has been genetically manipulated to produce crops with increased oil content (Lardizabal et al., 2008; Weselake et al., 2008; Zheng et al., 2008; Taylor et al., 2009).

Because the Arabidopsis DGAT1 mutants AS11 (designated dgat1-1; Katavic et al., 1995) and ABX45 (designated dgat1-2 here and also referred to as tag1-2; Routaboul et al., 1999) show only a 20 to 40% decrease in oil content, it is apparent that other enzymes must contribute to TAG synthesis in the developing seeds (Lu and Hills, 2002). Therefore, to further investigate enzymes that may contribute to oil biosynthesis in Arabidopsis, we tested a number of candidate genes for their ability to complement a TAG-deficient yeast mutant. We also performed genetic studies by crossing dgat1-1 with pdat1-2 and other candidate mutants. Failure to obtain double homozygous dgat1-1 pdat1-2 mutant plants suggested a possible embryo- or gametophyte-lethal phenotype in the absence of both DGAT1 and PDAT1. Our studies described herein reveal that PDAT1 and DGAT1 have overlapping functions for TAG synthesis in both seed and pollen of Arabidopsis and that the absence of their function leads not only to a reduction in TAG, but also to critical defects in normal pollen and embryo development.


Identifying Genes for TAG Synthesis by Overexpressing Candidate Acyltransferase Genes in the Yeast H1246 Strain (TAG Quadruple Mutant)

The accumulation of TAG at 60 to 80% of wild-type levels in dgat1-1 and dgat1-2 mutants (Katavic et al., 1995; Routaboul et al., 1999) suggested other enzymes can contribute substantially to TAG biosynthesis in Arabidopsis. In order to identify additional candidate gene(s) involved in TAG synthesis, a genetic complementation strategy in yeast was attempted. Candidate acyltransferase genes from Arabidopsis were expressed in the yeast strain H1246, which lacks DGA1 (acyl-CoA:diacylglycerol acyltransferase 1), LRO1 (phospholipid:diacylglycerol acyltransferase 1), ARE1, and ARE2 (acyl-CoA:cholesterol acyl transferase related enzymes 1 and 2) and is devoid of TAG and sterol esters. DGAT1 (At2g19450) was used as a positive control and was able to restore TAG synthesis to the yeast mutant. DGAT2-like (At3g51520), PDAT2-like (At3g44830), and PDAT-like (At5g28910) genes failed to complement TAG synthesis in H1246. In addition, a number of acyltransferase candidates of unknown function (Beisson et al., 2003), which show high expression during seed development (At3g05510, At4g19860, At5g12420, At1g12640, At1g63050, Ag5g60620, At3g51970, At2g27090, At1g27480, and At5g55380), were expressed in H1246 but failed to restore TAG synthesis. Considering that lipases can also catalyze a reverse acyltransferase reaction, several putative TAG lipases that are expressed during seed development (At5g18630, At5g18640, and At1g10740) were also tested in H1246. Again, none of these could complement the TAG synthesis mutation in H1246. The above data are summarized in Supplemental Table 1 online.

Generation of Double Mutant Plants by Crossing Candidate Mutants with dgat1-1 Mutant

As a parallel strategy to identify gene(s) encoding enzymes that might have complementary or redundant functions with DGAT1 for TAG synthesis in Arabidopsis, double mutant plants were created by crossing candidate mutants with dgat1-1, dgat2-like (SALK_067809), pdat1-2 (SALK_065334), pdat2-like (SALK_010854), and dgat1-1 (all are Columbia-0 [Col-0] ecotype). Single mutants dgat2-like and pdat2-like did not show any decrease in oil content compared with the wild type. Oil contents of the double homozygous lines of dgat1-1 dgat2-like and dgat1-1 pdat2-like were determined. Neither double mutant line showed significant further decreases in oil content beyond that already observed with the dgat1-1 single mutant (see Supplemental Figure 1 online). Combined with the results of the H1246 yeast expression studies, we concluded that the DGAT2 and PDAT2 genes do not play a substantial role in TAG synthesis in the dgat1-1 background.

Both Fertile and Sterile Pollen Were Found in Crossing of dgat1-1 and pdat1-2 Plants, and Genetic Analysis Indicates the Genotype of Sterile Pollen as dgat1-1 pdat1-2 Double Mutant

After several rounds of screening, it is noteworthy that we could get dgat1-1/dgat1-1 PDAT1/pdat1-2 or DGAT1/dgat1-1 pdat1-2/pdat1-2 single locus homozygous and one locus heterozygous plants, but no dgat1-1 pdat1-2 double homozygous mutant plants.

DGAT1 and PDAT1 are on two different chromosomes; therefore, these nonlinked genes will be randomly segregated and form male and female gametophytes. If female gametophytes or embryos were affected, abortion of seeds in the siliques would be observed. The developing embryos from the dgat1-1/dgat1-1 PDAT1/pdat1-2 or DGAT1/dgat1-1 pdat1-2/pdat1-2 plants were examined, and no obvious abortion in siliques was observed during embryo development. We deduced, therefore, that the reason for the failure to obtain double homozygous mutants might be related to pollen. TAG is an abundant storage material not only in seeds but also in pollen (Murphy, 2006), and DGAT1 has high expression not only in developing embryos but also in pollen (Lu et al., 2003). Furthermore, when the PDAT1 promoter was fused with β-glucuronidase (GUS), strong and specific expression was observed in anthers and pollen (Figure 1A). Based on this information, we hypothesized that the combined defects in both DGAT1 and PDAT1 expression might lead to lethality in the pollen.

Figure 1.
PDAT1 Expression Pattern in Arabidopsis Wild-Type (Col) Flower and Defective Pollen from dgat1-1/dgat1-1 PDAT1/pdat1-2 Plants.

Accordingly, anthers were detached from flowering wild-type, dgat1-1 mutant, pdat1-2 mutant, and dgat1-1/dgat1-1 PDAT1/pdat1-2 plants, and their mature pollen was directly observed by microscopy. We did not observe obvious morphological changes in pollen from dgat1-1 or pdat1-2 single mutants compared with the wild type. This was also confirmed in a second dgat1-2 mutant (ABX45; Routaboul et al., 1999) and a pdat1-1 mutant (Mhaske et al., 2005), both in the Arabidopsis Wassilewskija (Ws) background. However, we found that two distinct kinds of pollen occurred in dgat1-1/dgat1-1 PDAT1/pdat1-2 plants. Healthy pollen were observed that did not show any detectable changes compared with the wild type, while other pollen grains were smaller, deformed, and shrunken (Figure 1B). Histological analysis of pollen by Alexander's stain solution revealed that healthy pollen were the expected pink, while the deformed pollen could not effectively absorb stain solution and looked transparent or shrunken (Figure 1C). This suggested that deformed pollen were sterile and the pollen therefore could not transmit its genotype to the next generation.

Theoretically, there are two possible genotypes for pollen from dgat1-1/dgat1-1 PDAT1/pdat1-2 plants: one is dgat1-1 PDAT1 and the other is dgat1-1 pdat1-2. We hypothesized that the sterile pollen might be dgat1-1 pdat1-2. Viability and the genotypes of pollen and the female gametophyte from dgat1-1/dgat1-1 PDAT1/pdat1-2 were further investigated by self-pollination and reciprocal crossing with the wild type. Actual and theoretical ratios of genotypes are shown in Figure 2. When dgat1-1/dgat1-1 PDAT1/pdat1-2 plants were the pollen donor and the wild type was maternal plant, instead of the expected 1:1 ratio of offspring, nearly all seedlings were identified as DGAT1/dgat1-1 PDAT1/PDAT1 genotype and only 3.9% DGAT1/dgat1-1 PDAT1/pdat1-2 seedlings were found. In reciprocal crossing, with the wild type as pollen donor and dgat1-1/dgat1-1 PDAT1/pdat1-2 as the maternal plant, both DGAT1/dgat1-1 PDAT1/ PDAT1 and DGAT1/dgat1-1 PDAT1/pdat1-2 could be detected with the expected ratio 1:1 (Figure 2A). In the offspring population of dgat1-1/dgat1-1 PDAT1/pdat1-2 self-pollination, as mentioned above, no double homozygous plants were obtained, and the ratio of dgat1-1/dgat1-1 PDAT1/PDAT1 and dgat1-1/dgat1-1 PDAT1/pdat1-2 was 1:1 (P < 0.001), rather than the expected 1:2 (Figure 2B). These results showed that the dgat1-1 PDAT1 genotype, but not the dgat1-1 pdat1-2, could be transmitted by pollen, while both dgat1-1 pdat1-2 and dgat1-1 PDAT1 genotypes could be transmitted by female gametes. These results suggested that the sterile pollen had a double mutant dgat1-1 pdat1-2 genotype and that the inability to obtain a double homozygous phenotype was primarily due to pollen abnormalities.

Figure 2.
Test of Genetic Transmission of Arabidopsis dgat1-1 and pdat1-2 Gametophytes by Reciprocal Crossing and Self-Pollination.

If the dgat1-1 pdat1-2 double mutant led to sterile pollen, we speculated that this genotype of pollen should be observed in all plants with both dgat1 and pdat1 regardless of whether in the homozygous or heterozygous condition. Pollen from the DGAT1/dgat1-1 PDAT1/pdat1-2 F1 generation of the above cross were observed, and the same sterile pollen was found by both morphological observation and staining with Alexander's solution (see Supplemental Figure 2A online). In order to exclude the possibility that sterile pollen was dependent on a specific mutant line or ecotype, we conducted reciprocal crosses between dgat1-2 and pdat1-1. Again, the presence of sterile pollen from these F1 plants further suggested that this phenotype was caused by a dgat1 and pdat1 double mutation in pollen (see Supplemental Figure 2B online). Taken together, these data indicate that DGAT1 and PDAT1 have overlapping functions that are essential for pollen viability.

Mature Deformed Pollen Lacked Oil Bodies, but All Pollen Accumulated Exine Lipids and Appeared Normal at the Microspore Stage

Lipid is normally deposited both inside of pollen and externally as exine lipid on the pollen surface (Piffanelli et al., 1998; Murphy, 2006). Considering the high TAG content in mature pollen (Stanley and Linskens, 1974; Murphy, 2006), we postulated that, if DGAT1 and PDAT1 both contribute to oil synthesis in pollen, sterile pollen with a dgat1-1 pdat1-2 genotype should be deficient in storage oil. In order to test this hypothesis, anthers from wild-type, dgat1-1 mutant, pdat1-2 mutant, and dgat1-1/dgat1-1 PDAT1/pdat1-2 plants were examined by transmission electron microscopy (TEM) before flowering. dgat1-1 and pdat1-2 single mutants had no major changes in pollen oil bodies or other organelles compared with the wild type. By contrast, in dgat1-1/dgat1-1 PDAT1/pdat1-2 plants, major differences in pollen oil bodies were observed between normal and sterile pollen (Figures 3A to 3C). The healthy pollen, identified as dgat1-1 PDAT1 genotype by reciprocal crossing, showed normal oil bodies inside pollen and normal exine lipids, while the sterile pollen, identified as dgat1-1 pdat1-2 genotype by reciprocal crossing, showed normal exine lipids on the surface but no obvious oil bodies inside the deformed pollen. Furthermore, a diffuse intine and no obvious organelles were observed in the sterile pollen, while organelles, such as mitochondria, could be detected in healthy pollen (Figures 3D and 3E). These observations indicated that both DGAT1 and PDAT1 contribute to TAG synthesis inside pollen and that their expression was critical to pollen development.

Figure 3.
Oil Bodies Are Observed inside Arabidopsis Wild-Type (Col) Pollen and Normal Pollen, but Not in Deformed Pollen, from dgat1-1/dgat1-1 PDAT1/pdat1-2 Plants.

In order to determine that the observed pollen sterility and altered oil body phenotypes were truly dependent on the dgat1-1 pdat1-2 genotype of the male gametophyte, pollen were observed in the pollen sac of both young and mature flowers of dgat1-1/dgat1-1 PDAT1/pdat1-2 and the wild type. In contrast with the wild type, both healthy and deformed pollen were observed in the pollen sac of dgat1-1/dgat1-1 PDAT1/pdat1-2 at the mature pollen stage (Figures 4A and 4B). However, at the early-vacuolate microspore stage, all young microspores appeared to be identical and no oil bodies had yet appeared (Figures 4C and 4D). Organelles, such as mitochondria and Golgi bodies, could be detected in all microspores examined at this stage (Figure 4E). Collectively, these results indicated that disruption in pollen development occurred after the tetrad stage and was not dependent on the genotype of the maternal plant, but rather, depended on the genotype of the microspore/pollen.

Figure 4.
Abortion of Pollen in Arabidopsis dgat1-1/dgat1-1 PDAT1/pdat1-2 Plants Occurred between Microspore Stage and Pollen Maturity.

Abnormal Germination and Deformed Embryos Were Observed in PDAT1 RNA Interference in a dgat1 Background

Because there was a few percent leakage of viable dgat1 pdat1 double mutant pollen in reciprocal crossing, it may have been theoretically possible to recover a double homozygous line in a larger population if those embryos could develop normally. However, we failed to obtain double homozygous mutant lines in hundreds of its offspring, which suggested that the double mutant might also be lethal to embryo development. Therefore, in order to further investigate the role of PDAT1 in TAG synthesis and embryo development, PDAT1 expression was reduced by RNA interference (RNAi) in a dgat1 background. The 35S promoter is not active in Arabidopsis pollen (Wilkinson et al., 1997) but is active in seeds, which makes it possible for further investigation of embryo development. Considering the variable effects of RNAi suppression, we expected a range of phenotypes of different severity. A PDAT1-specific fragment, driven by the 35S promoter, was tested for RNAi gene silencing. Adding sucrose in medium during germination can be used to rescue seedling development from seeds with low oil contents (Cernac et al., 2006) or blocked degradation of oil (Germain et al., 2001; Eastmond, 2006). Unexpectedly, we obtained almost no transgenic plants by Kanr selection, with or without sucrose in the medium. Based on seed germination, the transformation efficiency was <0.01% compared with 1%, typical for other constructs (Bent, 2006). As a result, we obtained only a few transgenic seedlings. Plants from these seedlings produced T2 seeds and when screened for Kanr, we were surprised to find many nongerminated seeds and abnormal seedlings during germination in several independent transgenic lines. While most of the abnormal seedlings did not survive, T3 seeds harvested from surviving T2 plants showed similarly consistent abnormal seedling phenotypes. The germination of these seeds could also not be enhanced by adding 0.1% Tween 80 to the medium. In order to exclude the possibility that the abnormal seedlings were due to the damaging effects of seed surface sterilization, nontreated seeds of two independent homozygous T3 lines, 2-8 and 6-9, which showed severe phenotypes, were incubated on wet filter paper in a Petri dish. Compared with wild-type seeds, which yielded 98% ± 0.4% normal seedlings, 68.1% ± 6.4% and 49.6% ± 7.7% of seeds of lines 2-8 and 6-9, respectively, could not germinate. Of the dgat1-1 PDAT1RNAi seeds that germinated, approximately one-third had abnormal cotyledons or were without cotyledons (Figures 5A and 5B). In order to determine whether this phenotype was due to abnormal germination or was a consequence of abnormal embryo development, some seeds from different independent homozygous T3 lines were imbibed on wet filter paper at 4°C for 20 h. Seed coats were removed and embryos were observed under a dissection microscope. Approximately 10 to 20% of the embryos showed different degrees of visible malformation (Figure 5C), which suggested that simultaneous disruption of expression of both DGAT1 and PDAT1 genes resulted in abnormal embryo development.

Figure 5.
Suppression of PDAT1 Expression in Arabidopsis dgat1-1 Results in Deformed Seedlings and Impaired Embryo Development.

Small Abnormal Embryos Developed in dgat1-1 PDAT1 RNAi Seeds and Only Small Amounts of Oil Accumulated

Considering the very low transformation efficiency and abnormal embryos, we considered that although transformed seeds might be produced, they may be defective in germination. In the flower dip method, female gametophyte cells are transformed (Bent, 2006). Because the 35S promoter was used for RNAi gene silencing, it was possible that the above abnormality of embryo development in T2 and T3 seeds was influenced by the maternal phenotype. We therefore repeated the RNAi strategy using a vector with a DsRed visible marker, which allows identification of T1 transgenic seeds harvested from wild-type maternal plants. In addition, nonviable seeds can be identified using DsRed and the level of fluorescence provides an approximation of transgene expression (Stuitje et al., 2003).

The DsRed fluorescence marker, driven by the cassava vein mosaic virus promoter, was inserted in the above PDAT1 RNAi binary vector construct, and this construct was introduced into both the wild type and dgat1-1 mutant. When we screened transgenic seeds for DsRed fluorescence, we found that most transgenic seeds in the dgat1 background were small, but with unambiguous fluorescence. However, in many cases, the fluorescence extended over only part of a seed. By contrast, nearly all transgenic seeds in the wild-type background showed stronger fluorescence that was distributed uniformly over the whole seed (Figure 6A). Although there were variations in degree of fluorescence in dgat1-1 PDAT1 RNAi seeds, few seeds showed fluorescence distributed over the whole seed as observed in transformants of the wild-type control.

Figure 6.
Suppression of PDAT1 Expression in Arabidopsis dgat1-1 Results in Very Low Seed Oil and Altered Seed Development.

Consistent with the unaltered oil content and fatty acid profile of the pdat1-1 seeds (Mhaske et al., 2005), there were no changes in PDAT1RNAi seeds under a wild-type background (Figures 6B and 6C). Although there were some variations from seed to seed of dgat1 PDAT1 RNAi, on average, absolute oil content per seed was decreased by 84% compared with the dgat1-1 control (Figure 6B). The oil content expressed as a percentage of dry weight of dgat1-1 PDAT1 RNAi seeds decreased by 63% compared with the dgat1-1 control (Figure 6C). In the dgat1 mutant, the characteristic changes in fatty acid profile, including an increase in the proportion of 18:3 and a decrease in 20:1, were obvious (Figure 6D; Katavic et al., 1995). By contrast, dgat1-1 PDAT1 RNAi seed oils exhibited a decrease in 18:3 compared with dgat1-1 background, but also retained the low 18:1c9 and 20:1 proportions, consistent with the dgat1 mutant background (Figure 6D). Because of the major reduction in oil in the dgat1-1 PDAT1 RNAi seeds (compared with dgat1-1) our results suggest a scenario in which PDAT1 is the gene responsible for most of the TAG synthesis in the dgat1-1 mutant background.

Abnormal Embryos and Obvious Decreases in Oil Content Are Also Observed in Reciprocal Studies of DGAT1 RNAi in a pdat1-1 Background

To further confirm and investigate the relationship between PDAT1 and DGAT1 during seed development, we conducted a reciprocal experiment in which DGAT1 expression was suppressed by RNAi in a pdat1-1 mutant background. Unlike the pdat1-1 mutant, dgat1-1 has clear phenotypes both in terms of oil content and distinctive fatty acid profile. This provided a method to check whether DGAT1 suppression was effective in providing phenotypes similar to the dgat1 mutant. Lipid analysis indicated that when DGAT1 RNAi was expressed in a wild-type background, similar to dgat1 mutants, oil content decreased by 24% and the proportion of 18:3 increased, while 20:1 substantially decreased (Figures 7B and 7D). This result confirmed that the DGAT1 RNAi construct worked effectively and could be used for evaluating DGAT1 function under a pdat1-1 mutant background.

Figure 7.
Suppression of DGAT1 Expression in Arabidopsis pdat1-1 Results in Very Low Seed Oil and Altered Seed Development.

The phenotype of T1 DGAT1 RNAi seeds in a pdat1-1 background was very similar to that of PDAT1 RNAi seeds in a dgat1 background (above), except for the degree of fluorescence and oil content. An analysis of the DsRed fluorescence of seeds showed that most transgenic seeds developed abnormally, although not as severely as PDAT1 RNAi seeds in the dgat1 background. As observed above, fluorescence of the abnormal seeds was also concentrated in a small region rather than evenly dispersed (Figure 7A). Both absolute and relative oil content measurements showed a major decrease compared with its pdat1-1 background (Figures 7B and 7C). The fatty acid changes observed in pdat1-1 DGAT1 RNAi seeds (Figure 7D) were qualitatively similar to those observed in dgat1-1 PDAT1 RNAi seeds. These results further confirmed that both DGAT1 and PDAT1, but not other genes, contribute to most of the TAG synthesis in Arabidopsis seeds and are important for normal embryo development.

Major Changes in Oil Bodies of Mature Seeds of dgat1-1 PDAT1 RNAi and pdat1-1 DGAT1 RNAi

As described above, seed oil content was strongly decreased in both dgat1-1 PDAT1 RNAi and pdat1-1 DGAT1 RNAi conditions, and in addition, many embryos appeared very small, suggesting impairment in embryo development. Mature seeds of the wild type, dgat1-1, pdat1-1, dgat1-1 PDAT1 RNAi, and pdat1-1 DGAT1 RNAi were further examined by TEM.

As expected, no phenotypic change in oil bodies or morphology was observed in pdat1-1 single mutant seeds (Figures 8A and 8F). However, we were surprised to find that oil bodies in cotyledons of the dgat1-1 single mutant had obvious changes in size and shape. Most oil bodies in dgat1-1 were smaller, rounder, and appeared dark gray (Figures 8A and 8C). The size of oil bodies in hypocotyls of the dgat1-1 mutant were also smaller and rounder than the wild type but less strikingly so than in cotyledons (Figures 8B and 8D). This result was confirmed by observations of a second dgat1-2 mutant (Figure 8E). The altered oil bodies were not simply the result of an oil decrease in the dgat1 mutants because although the oil content of pdat1-1 DGAT1 RNAi seeds was much lower than that of dgat1 seeds, its oil bodies showed a normal shape, similar to the wild type (Figure 8G). These data suggest that even low expression of DGAT1 in RNAi seeds may be enough to retain normal oil body morphology.

Figure 8.
Distorted Oil Bodies in Arabidopsis Seeds of dgat1 Mutants and Fewer Oil Bodies in pdat1-1 DGAT1 RNAi and dgat1-1 PDAT1 RNAi Seeds.

The number of oil bodies also appeared to be substantially lower in both dgat1-1 PDAT1 RNAi and pdat1-1 DGAT1 RNAi seeds. Furthermore, there appeared to be only about one layer of oil bodies distributed near the plasma membrane, with very few oil bodies located near the middle of cells. As with dgat1-1, oil bodies in dgat1-1 PDAT1 RNAi seeds were smaller and darker (Figures 8G and 8H). Other organelles, including storage protein bodies, appeared normal. Additionally, we did not observe accumulation of starch in dgat1-1 PDAT1 RNAi and pdat1-1 DGAT1 RNAi seeds as has been observed in another low oil mutant (Lin et al., 1999). Higher-magnification TEM micrographs of oil bodies from cotyledons further showed above differences in ultrastructure (see Supplemental Figure 3 online).


The biosynthesis of TAG is a common metabolic pathway that occurs in essentially all plants, animals, fungi, and some bacteria. The major function of the pathway is considered to be the storage of acyl chains as a reserve of carbon and energy. Despite its central and conserved features, progress in understanding several aspects of TAG biosynthesis has been slow. Much effort has focused on DGAT since it is an enzyme unique to TAG synthesis. The involvement of other acyltransferases in the assembly of TAG has remained uncertain. For example, the glycerol-3-phosphate acyltransferase that initiates extraplastidial glycerolipid synthesis has not been clearly identified in plants. Furthermore, although a large number of mutants that influence fatty acid composition have been identified by extensive forward genetic screening, so far, DGAT1 is the only acyltransferase identified in Arabidopsis for which mutation results in low oil content. As noted above, dgat1 mutants are reduced only 20 to 40% in seed oil content. The simplest interpretation to account for the remaining TAG in dgat1 is that other enzymes contribute to acylation of the sn-3 position and compensate for the loss of DGAT1 activity. Distinguishing between alternative pathways for TAG synthesis is important not only for a further understanding of fundamental aspects of plant oil synthesis, but also to improve strategies for manipulating oil synthesis.

DGAT1 and PDAT1 Have Complementary Functions Essential for Normal Pollen Development

In this study, the failure to obtain double homozygous plants led to an unexpected discovery that the dgat1-1 pdat1-2 double mutant resulted in gametophytic mutations in pollen development. The Cruciferae family deposits lipids and proteins during pollen maturation (Baker and Baker, 1979). There are several types of lipidic structures in pollen grains, which play important roles in pollen development, dispersal, and pollination (Preuss et al., 1993; Wolters-Arts et al., 1998). Extracellular lipids include exine, a complex mixed polymer of acyl lipid with phenylpropanoid precursors, deposited on the pollen surface in some species. Intracellular lipids are found as components of internal membrane systems and are also deposited in lipid bodies (Piffanelli et al., 1998; Murphy, 2006). TAGs are known to be the main component in intracellular oil bodies of pollen (Stanley and Linskens, 1974; Murphy, 2005). These lipid bodies gradually appear shortly after pollen mitosis I and increase in number in the vegetative cell of pollen (Park and Twell, 2001) together with organelles, such as microbodies and mitochondria (Kuang and Musgrave, 1996). Consistent with those results, we observed abundant oil bodies in mature pollen but not in young microspores (Figures 3D and 3E).

It has been proposed that precursors of extracellular pollen lipids are determined by the sporophytic tapetum, while internal pollen lipids are determined by expression of the haploid genome of pollen (Mascarenhas, 1989; Ottaviano and Mulcahy, 1989; Piffanelli et al., 1997). This is supported by reports on tapetum development where a number of mutants are known to influence extracellular pollen lipids and impact fertility (Zheng, et al., 2003; Ma, 2005; Blackmore et al., 2007). However, to our knowledge, no mutants have been characterized that are deficient in oil bodies in Arabidopsis pollen, and genetic evidence is not yet available that internal storage lipids are synthesized by haploid-encoded enzyme(s). Here, we demonstrate that heterozygous dgat1-1/dgat1-1 PDAT1/pdat1-2 plants produced both fertile and sterile pollen. Furthermore, both types had normal extracellular lipids, but the shrunken and sterile pollen were devoid of oil bodies (Figures 3D and 3E). Our data from heterozygous crossing strongly support the hypothesis that lipid bodies in mature pollen are primarily determined by the gametophytic genome.

Although DGAT1 was reported to have high expression in pollen (Lu et al., 2003), other candidate acyltransferases are also expressed in pollen, and no direct evidence indicated DGAT1 involvement in TAG synthesis in pollen. Here, we showed that PDAT1 also had high expression in pollen (Figure 1A), and no obvious oil bodies were found in pollen with the dgat1-1 pdat1-2 double mutant genotype. Thus, we conclude that DGAT1 and PDAT1 have overlapping functions essential for TAG synthesis during pollen development and also for pollen viability.

No obvious difference was found among newly released microspores of dgat1-1/dgat1-1 PDAT1/pdat1-2 plants, whereas shrunken and deformed sterile pollen grains without obvious organelles were found in mature sterile pollen (Figures 3D, 4D, and 4E). The disappearance of organelles in addition to a lack of oil bodies suggested that disruption of TAG synthesis may lead to other pleiotropic effects (discussed below). Although mutation in both genes appeared to completely block TAG deposition and resulted in sterile pollen, the results of reciprocal crosses showed that viability of female gametophytes with a dgat1-1 pdat1-2 genotype was not affected (Figure 2), which indicated either that a deficiency in TAG synthesis did not impact female gametophyte development or that other genes conferred this function.

DGAT1 and PDAT1 Have Complementary Functions Essential for Normal Embryo Development

Because we could not obtain double homozygous dgat1-1 pdat1-2 mutant plants, an RNAi strategy was used to enable us to test their role in TAG synthesis in seeds. PDAT1 RNAi under a dgat1-1 background and its reciprocal- DGAT1 RNAi under a pdat1-1 background both resulted in strongly reduced TAG content in seeds (Figures 6 to 88),), indicating that both enzymes contribute to embryo TAG synthesis. Taken together, a key conclusion of these results is that the other candidate acyltransferase or transacylase genes, as mentioned in the Introduction, can be excluded as playing a major role in TAG synthesis in Arabidopsis developing seeds. This raises questions regarding the function of, for example, DGAT2-like and PDAT2-like (Ståhl et al., 2004) genes, both of which are expressed in Arabidopsis seeds. These and other genes might participate at some level in TAG synthesis, but either the level, location, or timing of expression might not be adequate to replace the functions provided by DGAT1 and PDAT1. Although some TAG was present in the RNAi mutant lines, it is worth noting that because RNAi strategies rarely provide a complete knockout of mRNA, it is possible that the double mutant of DGAT1 and PDAT1 may completely block TAG biosynthesis in Arabidopsis seeds.

The major reduction of oil content in RNAi mutant seeds, which indicates DGAT1 and PDAT1 have complementary functions in seed TAG synthesis, also led to the discovery that TAG synthesis (or possibly other functions of DGAT1/PDAT1) are essential for normal embryo development. Previously, dgat1-2 seeds were found to develop more slowly than the wild type (Routaboul et al., 1999), and there was a 30% lag time (4 weeks versus 3 weeks) reported in seed development in the AS11 mutant (Katavic et al., 1995). Disruption of both DGAT1 and PDAT1 in this study led to much more substantial defects in embryo maturation. Using DsRed as a visible marker to identify T1 transgenic seeds (from wild-type maternal plants) allowed us to exclude maternal effects as an explanation for these embryo phenotypes.

Suppression of PDAT1 and DGAT1 under dgat1-1 and pdat1-1 backgrounds, respectively, resulted in disruptions of embryo development. Seeds with DsRed fluorescence were shrunken and wrinkled, and fluorescence extended over only a small part of seeds in most cases (Figures 7A and and8A).8A). Only ~1% of these seeds could develop into seedlings (by comparing number of seedlings surviving during seed screening), and this was not increased by adding 1.5% sucrose or 0.1% Tween 80 to the germination medium. This contrasts with other mutants of Arabidopsis, such as wrinkled1, which has an 80% reduction in oil content but germinates efficiently on sucrose medium (Cernac et al., 2006). In addition, several mutants blocked in oil utilization (e.g., 3-ketoacyl-CoA thiolase 2 and sugar-dependent 1) can germinate on media containing sugars (Germain et al., 2001; Eastmond, 2006). Thus, low levels of TAG alone, as in wrinkled1, or the inability to utilize TAG during germination lead to less detrimental phenotypes than we observed.

In this study, we also found that DGAT1 was important for oil body size and shape, which was confirmed in both ethyl methanesulfonate and T-DNA insertion dgat1 mutants (Figure 8). It is interesting to note that oil bodies were normal in seeds of DGAT1 RNAi, which suggests that low level expression of DGAT1 was apparently sufficient to maintain normal oil body size. Other organelles appeared normal in dgat1-1 and also in the RNAi mutant lines where oil content was severely decreased. No starch grains were observed in either dgat1-1 PDAT1 RNAi or pdat1-2 DGAT1 RNAi oil- deficient seeds, which fails to support the suggestion that starch formation is a default storage deposition pathway in Arabidopsis (Lin et al., 1999).

The fact that the dgat1-1 mutant showed a 20 to 30% decrease in oil content (Katavic et al., 1995) while no changes of oil in pdat1-1 were observed (Mhaske et al., 2005) might suggest that DGAT1 can completely compensate for the lack of PDAT1 function, whereas PDAT1 only partially complements the function of DGAT1 in developing seeds. Such compensations are known to occur through a wide variety of transcriptional and posttranscriptional mechanisms, such as increased translation, mRNA or protein stability, and enzyme activation. In addition, because the products of PDAT1 are TAG and lysophosphatidylcholine (LPC), a lysophospholipid acyltransferase (e.g., LPCAT) is necessary to cooperate with PDAT1 to regenerate PC from LPC. Thus, either PDAT1 or LPCAT could limit the final TAG accumulation in dgat1 seeds. If so, it may be necessary to overexpress lysophospholipid acyltransferase(s) together with PDAT1 in developing embryos to better understand this issue.

In addition to reduced oil content, both the AS11 and ABX45 dgat1 mutants display characteristic fatty acid profiles with an increase in proportions of 18:3 and decreases in proportions of 18:1 and very-long-chain fatty acids (>C18), especially at the sn-3 position (Katavic et al., 1995). This phenotype is consistent with PDAT1's function in TAG synthesis as determined in this study. In the wild type, both DGAT1 and PDAT1 contribute to sn-3, drawing from the acyl-CoA and PC pools, respectively. However, in dgat1-1, nearly all acyl groups on sn-3 must be derived from phospholipids via PDAT, and there is a greater chance that those acyl groups are desaturated (e.g., 18:2 or 18:3) and a lesser chance that very-long-chain (>C18; e.g., 20:1) acyl groups will be available from the PC. However, given our current knowledge, it is not possible to conclude how much of the acyl flux is carried by PDAT in wild-type developing seeds. Over 70% of the fatty acids at the sn-3 position of Arabidopsis TAG are either saturated or >C18 in length (Katavic et al., 1995), and these fatty acids are generally excluded from the sn-2 position of PC, which is one of the substrates involved in the PDAT reaction. This would suggest that in wild-type Arabidopsis seeds, DGAT1 contributes more to the sn-3 position than does PDAT1. A recent analysis of fluxes into TAG of developing soybeans could distinguish two kinetically distinct sn-3 acylations of DAG that used either saturated or polyunsaturated fatty acids, and these likely reflect the activities of DGAT and PDAT reactions, respectively (Bates et al., 2009). Similar kinetic studies of dgat1, pdat1, and wild-type Arabidopsis developing seeds may allow a better estimate of relative flux through the alternative pathways.

Possible Other Functions of TAG Synthesis or DGAT1 and PDAT1 Related to Diverse Phenotypes Observed

The general concept that the major role of TAG synthesis is to provide a neutral storage material is supported by studies of the yeast dga1 lro1 are1 are2 quadruple mutant, which is completely devoid of TAGs. Normal growth of this mutant suggested that TAGs were not essential for yeast growth under laboratory culture conditions (Sandager et al., 2002). However, this traditional point of view is being modified by several reports from different species. In the fission yeast Schizosaccharomyces pombe, a double knockout of Plh1 (phospholipid:diacylglycerol acyltransferase) and Dga1 (acyl-CoA:diacylglycerol acyltransferase) led to almost complete absence of TAG, and these mutant cells lost viability after entering the stationary stage (Zhang et al., 2003). A mutant of DGAT (gene ID: W01A11.2) in Caenorhabditis elegans showed increased sensitivity to hypoxic injury (Mabon et al., 2009). Drosophila mutants in the midway gene (identified as a DGAT1) displayed severely reduced levels of neutral lipids in the germline and showed premature apoptosis and degeneration of nurse cells (Buszczak et al., 2002). Also, no mutant completely devoid of TAG has been found in oleaginous bacteria, such as Streptomyces coelicolor (Arabolaza et al., 2008), which may hint that deficiency of TAG synthesis genes may be lethal. As discussed by Daum et al. (2007), these and other reports (e.g., Lock et al., 2009) suggest that TAG synthesis in several species plays an important role not only as a storage reserve, but also in growth and development. Furthermore, other studies suggest that TAG synthesis may serve to prevent its substrates, DAG and acyl-CoA (or fatty acids), from reaching potentially damaging levels (Listenberger et al., 2003; Zhang, et al., 2003). Deficiency of sterol and TAG synthesis triggers fatty acid–mediated cell death in yeast (Garbarino et al., 2009; Siloto et al., 2009). DAG is generally believed to be important in several processes as a lipid precursor and as a lipid second messenger, and its level is strictly regulated (Carrasco and Merida, 2006; van Herpen, and Schrauwen-Hinderling, 2008). In dgat1-1 mutant seeds, DAG increased up to 12% of total lipid, compared with <1% in mature wild-type seeds (Katavic et al., 1995). Given its known role in signal transduction processes, we also cannot rule out that a possible change in the relative DAG concentration leads to cascading mechanisms that affect normal embryo morphological development.

The range of defects we observed in embryo and pollen structure and morphology and in seed germination reinforce the conclusion that TAG synthesis in plants functions in additional roles besides simply providing a carbon/energy reserve. Although the need for TAG homeostasis and/or avoidance of disruptive concentrations of intermediates are likely explanations for the pleiotropic effects observed in this study, other possibilities cannot be ruled out. For example, an N-terminal–deleted yeast PDAT1 could catalyze a number of transacylation reactions in addition to PDAT activity, including acylation of long-chain alcohols (Ghosal et al., 2007). Thus, it is possible that some other functions of DGAT1 and/or PDAT1 have not been identified that are essential in plant growth and development. The mutants and transgenic lines obtained in this study may allow a more focused examination of these alternatives and lead to a better understanding of roles played by TAG synthesis in plant biology.


Plant Materials and Growth Conditions

Arabidopsis thaliana ecotypes Col and Ws were used as control wild-type plants for the different ecotype mutants, respectively. SALK mutant lines, Col ecotype, were obtained from the Salk Institute via the ABRC (Ohio State University, Columbus, OH). Seeds in pots or plates were cold-treated for 3 d at 4°C in the dark before being transferred to a controlled growth chamber. Arabidopsis plants were grown in a soil mixture (3:1:1 mixture of peat moss–enriched soil:vermiculite:perlite) in a growth chamber with 16 h light (200 μmol m−2 s−1 radiation) and 8 h dark at 22°C. For growth on plates, seeds were surface-sterilized for 20 min in 20% (v/v) bleach and rinsed three times with sterile water. Seeds plated on agar medium containing half-strength Murashige and Skoog medium salts and 0.75% phytoblend with/without 1.5% (w/v) sucrose, adjusted to pH 5.7 using KOH before autoclaving. Kanamycin (50 mg/L) was added or omitted after media were autoclaved. In embryo rescue experiments, filter-sterilized Tween 80 was added to the medium at a final concentration of 0.1% (v/v). Nonsterilized seeds were also germinated on wet filter paper in a Petri dish and washed every day to reduce possible microbial growth. Germination was determined after 7 d. All seedlings developing on plates were cultured under the same conditions as were plants in pots (see above). Six-week-old plants, with inflorescences trimmed once, were used for transformation by the floral dip method (Bent, 2006). For all crosses between mutants or between a mutant and the wild type, immature flower buds were emasculated and manually cross-pollinated with pollen from a parental blooming flower.

Genotyping of Mutants and Alleles by PCR

All PCR primers used in this study are presented in Supplemental Table 2 online. Isolation of dgat1-1 (AS11, Col background), dgat1-2 (ABX45, Ws background), and pdat1-1 mutants (Ws background) were previously described (Katavic et al., 1995; Routaboul, et al., 1999; Mhaske et al., 2005, respectively). We use dgat1- to describe AS11 and ABX45 mutants because it is more specific and avoids overlap with the Arabidopsis TAG1 transposase. Forward primer AS11a and reverse primer AS11b were used to identify the dgat1-1 mutant allele (Zou et al., 1999). Forward primer PD1F, reverse primer PD1R1010, and T-DNA primer PDTDNA were used to identify the pdat1-1 mutant allele. Primers, used for identifying SALK T-DNA insertion mutants, were designed by the SIGnAL T-DNA Express Arabidopsis Gene Mapping Tool (http://signal.salk.edu/tdnaprimers.2.html, provided by the Salk Institute Genomic Analysis Laboratory; Alonso et al., 2003). The pdat1-2 mutant, designated in the SALK collection as line S065334, was isolated using left genomic primer S065334LP, right genomic primer S065334RP, and T-DNA left border primer LBb1. dgat2, designated in the SALK collection as line S067809, was isolated using left genomic primer S067809LP, right genomic primer S067809RP, and LBb1. pdat2 designated in the SALK collection as line S010854 was isolated using left genomic primer S010854LP, right genomic primer S010854RP, and LBb1. Double mutants, created by crossing, were identified by two relevant pairs of primers. All homozygous lines were confirmed in their offspring. Genotyping of seedlings from seeds of reciprocal crosses were conducted using two pairs of primers to identify dgat1-1 and pdat1-2 mutant alleles, respectively, and identification of heterozygous DGAT1/dgat1-1 was used as a marker for true hybrids.

Candidate Genes Expressed in H1246 Yeast Stain

Total RNA was extracted from Arabidopsis leaves or siliques using the RNeasy plant mini kit, and cDNA was synthesized using Superscript II reverse transcriptase according to the product instructions. Gene-specific primers (see Supplemental Table 2 online) were designed to amplify the coding regions of candidate acyltransferase genes: At3g51520, At3g44830, At5g28910, At3g05510, At4g19860, At5g12420, At3g03520, At1g12640, At1g63050, Ag5g60620, At3g51970, At2g27090, At2g44080, At1g27480, and At5g55380. To the blunt-end PCR products obtained by Pfu Turbo DNA polymerase, an adenine overhang was added at the 3′ end using Taq DNA polymerase, and the amplicons were cloned into the yeast shuttle vector pYES2.1. After confirming cloning integrity by plasmid sequencing, the vectors were transformed into yeast mutant stain H1246 (dga1, lro1, are1, and are2 TAG quadruple mutants; Sandager et al., 2002). The DGAT1 gene was used as a positive control in this complementation experiment, and the self-ligated pYES2.1 was used as negative control. The resultant yeast stains were cultured in yeast nitrogen base medium with Brent Supplement mix –URA. Yeast cells were induced for gene expression in 2% β-galactose medium and were harvested at the stationary phase. Complementation of TAG synthesis was tested in transformed yeast strains by lipid analysis (see below).

Pollen Observation and Staining

Anthers from flowering plants were detached and squeezed between glass slides. Pollen were spread on the slide and sealed with a cover slip. Slides were observed under a Leica DM2000 light microscope (with ×400 magnification), and digital pictures were captured with a Leica DFC290 camera. For testing pollen viability, pollen slides were prepared from anthers as above, and 200 μL of Alexander's staining solution was applied to the pollen spot and after staining for 5 min at room temperature, a cover slip was put in place. Slides were observed under a light microscope and digital pictures were taken as above. Anthers from plants homozygous for the ProPDAT1:GUS construct were used for histochemical GUS staining with the method of Jefferson et al. (1987).

Plasmid Constructs for Study of PDAT1 Expression Patterns and for Creating RNAi Transformants

In general, standard methods were used in DNA and RNA isolation and manipulation as prescribed by Sambrook and Russell (2001). All primers are summarized in Supplemental Table 2 online. All vectors were confirmed by sequencing at each step.

The PDAT1 promoter (834 bp upstream of the PDAT1 gene) and 466 bp of the 5′ untranslated region were amplified from genomic DNA of the wild type (Col) using primers PPD1F and PPD1R. The amplicons were cloned into the Gateway pENTR vector. The PDAT1 promoter with its 5′ untranslated region was recombined into the Gateway pMDC162 destination vector via an LR reaction in which the GUS reporter gene was fused downstream of the above sequence.

Total RNA was extracted from leaves or siliques of the wild type (Col) and cDNA molecules synthesized by reverse transcription were used as templates for the following cloning. A BLAST analysis of PDAT1 conducted against the Arabidopsis genomic and EST databases showed that the fragment from 1607 to 1855 bp is a highly specific region, which will maximally avoid off-targeting of gene silencing. A 249-bp PDAT1 mRNA-specific fragment was amplified using primers PD1F1607 and PD1R1855. The amplicon was cloned into the Gateway pENTR vector. PDAT1 fragment was recombined into RNAi destination vector pK7GWIWG2 (II) via an LR reaction and designated as PDAT1RNAi. This destination vector was used for plant transformation or further constructs. The DsRed gene with cassava vein mosaic virus promoter and agropine synthase terminator were amplified using primers DsRedF and DsRedR. The amplicon was cloned into the pGEM T easy vector. This fragment was released from the vector by HindIII digestion and ligated into the above PDAT1RNAi vector, which was also digested by HindIII. A 179-bp DGAT1 mRNA-specific fragment was amplified using primers DG1F763 and DG1R941. Similar steps were taken for construction of the DGAT1RNAi vector with DsRed marker, except the pH7GWIWG2 (II) destination vector was used. All plasmids were transferred to Agrobacterium tumefaciens strain C58CI and used for plant transformation by the floral dip method (Bent, 2006).

Lipid and Fatty Acid Analysis

To identify complementation of candidate genes in the yeast H1246 mutant (devoid of TAGs), cells were collected and washed twice with PBS (10 mM phosphate, 2.7 mM KCl, and 137 mM NaCl, pH7.4) and homogenized with an equal volume of glass beads using the BeadBeater at 4°C using three bursts of 1 min. Total lipids were extracted by the chloroform:methanol:KCl/HNO3 (1:2:0.8,v/v/v) method. Lipid fractions were resolved on silica G60 thin layer chromatography plates developed in hexane:diethyl ether: acetate acid (70:30:1). The TAG bands were identified according to standards.

Seed oil content measurements followed the method of Li et al. (2006) with minor modification. Thirty seeds per replicate were used for measuring seed weights after stabilizing seed water content in desiccators for 48 h. Ten randomly picked seeds were used for measuring oil contents and fatty acid profiles. One microgram of triheptadecanoin was used as a TAG internal standard. Two milliliters of 2.5% (v/v) concentration of sulfuric acid in methanol was added to each tube and kept at 90°C for 90 min. The fatty acid methyl ester extracts were analyzed by gas chromatography with a flame ionization detector on a DB23 column GC. Assuming most of the fatty acids are in the neutral lipid fraction in Arabidopsis seeds, total fatty acids measured as methyl esters to approximate the total oil for comparison purposes.

Fluorescence Microscopy

After transformation of PDAT1RNAi and DGAT1RNAi with DsRed marker, seeds were harvested from T0 plants. T1 transgenic seeds were screened under a dissection microscope with a red filter under green light. Digital pictures of transgenic seeds were captured under a fluorescent microscope with ×100 magnification, with a Zeiss #20 filter set (excitation 546 ± 12 nm, emission 575 to 640 nm).

Light Microscope and TEM

Mature seeds were imbibed on wet filter paper at 4°C for 16 h and seed coats were removed. Mature embryos were observed under a Leica MZ125 dissection microscope, and digital pictures were captured by a Leica DC480 camera. Young flower buds at different developmental stages were fixed in 0.1 M sodium cacodylate buffer, pH 7.4, with 2.5% paraformaldehyde and 2.5% glutaraldehyde for 24 h at 4°C. After washing three times with 0.1 M sodium cacodylate buffer to remove fixative, samples were postfixed in 1% osmium tetroxide containing in 0.1 M sodium cacodylate buffer for 3 h, washed three times, and dehydrated in a graded acetone series and infiltrated into a graded resin using an Ultrabed low viscosity embedding kit (Electron Microscopy Sciences) for 4 d. Samples were then embedded in gelatin molds and polymerized for 1 d. Sections of 1 μm thickness were obtained with a Power Tome XL ultramicrotome (Boeckeler Instruments), placed on glass slides, and stained with 1% toluidine blue for 2 min at 50°C. Sections were observed under a Leica DM2000 light microscope and digital pictures were captured with a Leica DFC290 camera. Sections of 70-nm thickness were cut with the same ultramicrotome, picked and placed onto 200 mesh cooper grids, and stained with 2% uranyl acetate in 50% ethanol for 10 min and then with Reynold's lead citrate solution for 15 min. The sections were observed and pictures were taken with the JEOL 100CX transmission electron microscope at 100 kV accelerating voltage. For seed sample preparations, dry seeds were imbibed on wet filter paper at 4°C for 16 h, seed coats were peeled, and cotyledons and hypocotyls were separated under a dissection microscope. Because seeds of both PDAT1 and DGAT1 RNAi in the dgat1-1 and pdat1-1 mutant background, respectively, were small and distorted, we could not remove seed coats and separate cotyledons and hypocotyls before fixing. Lipid bodies and other organelles in pollen were identified according to Kuang and Musgrave (1996). Similar steps as above were taken for preparing sections of TEM except with 7 d of infiltration. The same methods as above were used for thin section and TEM observation.

Accession Numbers

Sequence data from this article can be found in the Arabidopsis Genome Initiative or GenBank/EMBL databases under the following accession numbers: DGAT1, At2g19450; PDAT1, At5g13640; DGAT2-like, At3g51520; PDAT2-like, At3g44830; PDAT-like, At5g28910; At3g05510, At4g19860, At5g12420, At1g12640, At1g63050, Ag5g60620, At3g51970, At2g27090, At1g27480, At5g55380, At5g18630, At5g18640, and At1g10740.

Supplemental Data

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

  • Supplemental Figure 1. Neither dgat1-1 dgat2-Like nor dgat1-1 pdat2-Like Double Mutant Lines Showed Significant Oil Content Decreases beyond That Observed in the dgat1-1 Mutant.
  • Supplemental Figure 2. Defective Pollen Were Observed in both F1 Crossings of dgat1-1 with pdat1-2 and dgat1-2 with pdat1-1.
  • Supplemental Figure 3. High-Magnification TEM Micrographs of Oil Bodies in Arabidopsis Wild-Type, pdat1-1, dgat1-1, dgat1-2, pdat1-1 DGAT1 RNAi, and dgat1-1 PDAT1 RNAi Seeds.
  • Supplemental Table 1. Complementation of TAG Synthesis by Expressing Arabidopsis Genes in Yeast Quadruple Mutant H1246 Strain.
  • Supplemental Table 2. Primers Used in This Study.

Supplementary Material

[Supplemental Data]


We thank Mike Pollard (Michigan State University) for helpful advice, discussions, and suggestions during this study. We also thank Alicia Pastor (Michigan State University) for TEM sample dissection, Betty Lockerbie (Plant Biotechnology Institute, Canada) for oil content measurement of two double mutants, Sten Stymne (Swedish University of Agricultural Sciences, Sweden) for his gift of the H1246 yeast strain, and Timothy P. Durrett (Michigan State University) for providing the DsRed vector. This study is supported by grants from the U.S. Department of Energy (DE-FG02-87ER13729) and the Great Lakes Bioenergy Research Center (Cooperative Agreement DE-FC02-07ER64494) and from the National Science Foundation (DBI-0701919).


The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantcell.org) is: John B. Ohlrogge (ude.usm@eggorlhO).

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  • Alonso, J.M., et al. (2003). Genome-wide insertional mutagenesis of Arabidopsis thaliana. Science 301 653–657. [PubMed]
  • Arabolaza, A., Rodriguez, E., Altabe, S., Alvarez, H., and Gramajo, H. (2008). Multiple pathways for triacylglycerol biosynthesis in Streptomyces coelicolor. Appl. Environ. Microbiol. 74 2573–2582. [PMC free article] [PubMed]
  • Baker, H.G., and Baker, I. (1979). Starch in angiosperm pollen grains and its evolutionary significance. Am. J. Bot. 66 591–600.
  • Banas, A., Dahlqvist, A., Stahl, U., Lenman, M., and Stymne, S. (2000). The involvement of phospholipid: diacylglycerol acyltransferases in triacylglycerol production. Biochem. Soc. Trans. 28 703–705. [PubMed]
  • Bates, P.D., Durrett, T.P., Ohlrogge, J.B., and Pollard, M. (2009). Analysis of acyl fluxes through multiple pathways of triacylglycerol synthesis in developing soybean embryos. Plant Physiol. 150 55–72. [PMC free article] [PubMed]
  • Beisson, F., et al. (2003). Arabidopsis thaliana 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 Physiol. 132 681–697. [PMC free article] [PubMed]
  • Bent, A.F. (2006). Arabidopsis thaliana floral dip transformation method. In Agrobacterium Protocols, 2nd ed, K. Wang, ed (Totowa, NJ: Humana Press), pp. 87–103.
  • Blackmore, S., Wortley, A.H., Skvarla, J.J., and Rowley, J.R. (2007). Pollen wall development in flowering plants. New Phytol. 174 483–498. [PubMed]
  • Bouvier-Nave, P., Benveniste, P., Oelkers, P., Sturley, S.L., and Schaller, H. (2000). Expression in yeast and tobacco of plant cDNAs encoding acyl CoA:diacylglycerol acyltransferase. Eur. J. Biochem. 267 85–96. [PubMed]
  • Buszczak, M., Lu, X.H., Segraves, W.A., Chang, T.Y., and Cooley, L. (2002). Mutations in the midway gene disrupt a Drosophila acyl coenzyme A:diacylglycerol acyltransferase. Genetics 160 1511–1518. [PMC free article] [PubMed]
  • Carrasco, S., and Merida, I. (2006). Diacylglycerol, when simplicity becomes complex. Trends Biochem. Sci. 32 27–36. [PubMed]
  • Cases, S., Smith, S.J., Zheng, Y.W., Myers, H.M., Lear, S.R., Sande, E., Novak, S., Collins, C., Welch, C.B., Lusis, A.J., Erickson, S.K., and Farese, R.V. (1998). Identification of a gene encoding an acyl CoA: diacylglycerol acyltransferase, a key enzyme in triacylglycerol synthesis. Proc. Natl. Acad. Sci. USA 95 13018–13023. [PMC free article] [PubMed]
  • Cernac, A., Andre, C., Hoffmann-Benning, S., and Benning, C. (2006). WRI1 is required for seed germination and seedling establishment. Plant Physiol. 141 745–757. [PMC free article] [PubMed]
  • Dahlqvist, A., Stahl, U., Lenman, M., Banas, A., Lee, M., Sandager, L., Ronne, H., and Stymne, S. (2000). Phospholipid:diacylglycerol acyltransferase: An enzyme that catalyzes the acyl-CoA-independent formation of triacylglycerol in yeast and plants. Proc. Natl. Acad. Sci. USA 97 6487–6492. [PMC free article] [PubMed]
  • Daum, G., Wagner, A., Czabany, T., and Athenstaedt, K. (2007). Dynamics of neutral lipid storage and mobilization in yeast. Biochimie 89 243–248. [PubMed]
  • Durrett, T.P., Benning, C., and Ohlrogge, J. (2008). Plant triacylglycerols as feedstocks for the production of biofuels. Plant J. 54 593–607. [PubMed]
  • Dyer, J.M., Stymne, S., Green, A.G., and Carlsson, A.S. (2008). High-value oil from plants. Plant J. 54 640–655. [PubMed]
  • Eastmond, P.J. (2006). SUGAR-DEPENDENT1 encodes a patatin domain triacylglycerol lipase that initiates storage oil breakdown in germinating Arabidopsis seeds. Plant Cell 18 665–675. [PMC free article] [PubMed]
  • Garbarino, J., Padamsee, M., Wilcox, L., Oelkers, P.M., D'Ambrosio, D., Ruggles, K.V., Ramsey, N., Jabado, O., Turkish, A., and Sturley, S.L. (2009). Sterol and diacylglycerol acyltransferase deficiency triggers fatty acid-mediated cell death. J. Biol. Chem. 284 30994–31005. [PMC free article] [PubMed]
  • Germain, V., Rylott, E.L., Larson, T.R., Sherson, S.M., Bechtold, N., Carde, J.P., Bryce, J.H., Graham, I.A., and Smith, S.M. (2001). Requirement for 3-ketoacyl-CoA thiolase-2 in peroxisome development, fatty acid beta-oxidation and breakdown of triacylglycerol in lipid bodies of Arabidopsis seedlings. Plant J. 28 1–12. [PubMed]
  • Ghosal, A., Banas, A., Stahl, U., Dahlqvist, A., Lindqvist, Y., and Stymne, S. (2007). Saccharomyces cerevisiae phospholipid:diacylglycerol acyl transferase (PDAT) devoid of its membrane anchor region is a soluble and active enzyme retaining its substrate specificities. Biochim. Biophs. Acta 1771 1457–1463. [PubMed]
  • Griffiths, G., Stobart, A.K., and Stymne, S. (1988). Delta-6-desaturase and delta-12-desaturase activities and phosphatidic-acid formation in microsomal preparations from the developing cotyledons of common borage (Borago officinalis). Biochem. J. 252 641–647. [PMC free article] [PubMed]
  • He, X., Turner, C., Chen, G.Q., Lin, J.T., and McKeon, T.A. (2004). Cloning and characterization of a cDNA encoding diacylglycerol acyltransferase from castor bean. Lipids 39 311–318. [PubMed]
  • Hobbs, D.H., Lu, C., and Hills, M.J. (1999). Cloning of a cDNA encoding diacylglycerol acyltransferase from Arabidopsis thaliana and its functional expression. FEBS Lett. 452 145–149. [PubMed]
  • Jako, C., Kumar, A., Wei, Y.D., Zou, J.T., Barton, D.L., Giblin, E.M., Covello, P.S., and Taylor, D.C. (2001). Seed-specific over-expression of an Arabidopsis cDNA encoding a diacylglycerol acyltransferase enhances seed oil content and seed weight. Plant Physiol. 126 861–874. [PMC free article] [PubMed]
  • Jefferson, R.A., Kavanagh, T.A., and Bevan, M.W. (1987). GUS fusions: Beta-glucuronidase as a sensitive and versatile gene fusion marker in higher plants. EMBO J. 6 3901–3907. [PMC free article] [PubMed]
  • Kalscheuer, R., and Steinbuchel, A. (2003). A novel bifunctional wax ester synthase/acyl-CoA:diacylglycerol acyltransferase mediates wax ester and triacylglycerol biosynthesis in Acinetobacter calcoaceticus ADP1. J. Biol. Chem. 278 8075–8082. [PubMed]
  • Katavic, V., Reed, D.W., Taylor, D.C., Giblin, E.M., Barton, D.L., Zou, J.T., Mackenzie, S.L., Covello, P.S., and Kunst, L. (1995). Alteration of seed fatty-acid composition by an ethyl methanesulfonate-induced mutation in Arabidopsis thaliana affecting diacylglycerol acyltransferase activity. Plant Physiol. 108 399–409. [PMC free article] [PubMed]
  • King, A., Nam, J.W., Han, J.X., Hilliard, J., and Jaworski, J.G. (2007). Cuticular wax biosynthesis in petunia petals: Cloning and characterization of an alcohol-acyltransferase that synthesizes wax-esters. Planta 226 381–394. [PubMed]
  • Kroon, J.T.M., Wei, W.X., Simon, W.J., and Slabas, A.R. (2006). Identification 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. Phytochemistry 67 2541–2549. [PubMed]
  • Kuang, A., and Musgrave, M.E. (1996). Dynamics of vegetative cytoplasm during generative cell formation and pollen maturation in Arabidopsis thaliana. Protoplasma 194 81–90. [PubMed]
  • Lardizabal, K., Effertz, R., Levering, C., Mai, J., Pedroso, M.C., Jury, T., Aasen, E., Gruys, K., and Bennett, K. (2008). Expression of Umbelopsis ramanniana DGAT2A in seed increases oil in soybean. Plant Physiol. 148 89–96. [PMC free article] [PubMed]
  • Lardizabal, K.D., Mai, J.T., Wagner, N.W., Wyrick, A., Voelker, T., and Hawkins, D.J. (2001). DGAT2 is a new diacylglycerol acyltransferase gene family - Purification, cloning, and expression in insect cells of two polypeptides from Mortierella ramanniana with diacylglycerol acyltransferase activity. J. Biol. Chem. 276 38862–38869. [PubMed]
  • Lehner, R., and Kuksis, A. (1996). Biosynthesis of triacylglycerols. Prog. Lipid Res. 35 169–201. [PubMed]
  • Li, Y.H., Beisson, F., Koo, A.J.K., Molina, I., Pollard, M., and Ohlrogge, J. (2007). Identification of acyltransferases required for cutin biosynthesis and production of cutin with suberin-like monomers. Proc. Natl. Acad. Sci. USA 104 18339–18344. [PMC free article] [PubMed]
  • Li, Y.H., Beisson, F., Pollard, M., and Ohlrogge, J. (2006). Oil content of Arabidopsis seeds: The influence of seed anatomy, light and plant-to-plant variation. Phytochemistry 67 904–915. [PubMed]
  • Lin, Y., Sun, L., Nguyen, L.V., Rachubinski, R.A., and Goodman, H.M. (1999). The pex16p homolog SSE1 and storage organelle formation in Arabidopsis seeds. Science 284 328–330. [PubMed]
  • Listenberger, L.L., Han, X.L., Lewis, S.E., Cases, S., Farese, R.V., Ory, D.S., and Schaffer, J.E. (2003). Triglyceride accumulation protects against fatty acid-induced lipotoxicity. Proc. Natl. Acad. Sci. USA 100 3077–3082. [PMC free article] [PubMed]
  • Lock, Y.Y., Snyder, C.L., Zhu, W.M., Siloto, R.M.P., Weselake, R.J., and Shah, S. (2009). Antisense suppression of type 1 diacylglycerol acyltransferase adversely affects plant development in Brassica napus. Physiol. Plant. 137 61–71. [PubMed]
  • Lu, C., and Hills, M.J. (2002). Arabidopsis mutants deficient in diacylglycerol acyltransferase display increased sensitivity to abscisic acid, sugars, and osmotic stress during germination and seedling development. Plant Physiol. 129 1352–1358. [PMC free article] [PubMed]
  • Lu, C.L., de Noyer, S.B., Hobbs, D.H., Kang, J., Wen, Y., Krachtus, D., and Hills, M.J. (2003). Expression pattern of diacylglycerol acyltransferase-1, an enzyme involved in triacylglycerol biosynthesis, in Arabidopsis thaliana. Plant Mol. Biol. 52 31–41. [PubMed]
  • Ma, H. (2005). Molecular genetic analyses of microsporogenesis and microgametogenesis in flowering plants. Annu. Rev. Plant Biol. 56 393–434. [PubMed]
  • Mabon, M.E., Mao, X., Jiao, Y., Scott, B.A., and Crowder, C.M. (2009). Systematic identification of gene activities promoting hypoxic death. Genetics 181 483–496. [PMC free article] [PubMed]
  • Mascarenhas, J.P. (1989). The male gametophyte of flowering plants. Plant Cell 1 657–664. [PMC free article] [PubMed]
  • Mhaske, V., Beldjilali, K., Ohlrogge, J., and Pollard, M. (2005). Isolation and characterization of an Arabidopsis thaliana knockout line for phospholipid: Diacylglycerol transacylase gene (At5g13640). Plant Physiol. Biochem. 43 413–417. [PubMed]
  • Milcamps, A., Tumaney, A.W., Paddock, T., Pan, D.A., Ohlrogge, J., and Pollard, M. (2005). Isolation of a gene encoding a 1,2-diacylglycerol-sn-acetyl-CoA acetyltransferase from developing seeds of Euonymus alatus. J. Biol. Chem. 280 5370–5377. [PubMed]
  • Murphy, D.J. (2006). The extracellular pollen coat in members of the Brassicaceae: Composition, biosynthesis, and functions in pollination. Protoplasma 228 31–39. [PubMed]
  • Murphy, D.J., ed (2005). Plant Lipids: Biology, Utilization and Manipulation. (Oxford, UK: Blackwell Publishing).
  • Nykiforuk, C.L., Furukawa-Stoffer, T.L., Huff, P.W., Sarna, M., Laroche, A., Moloney, M.M., and Weselake, R.J. (2002). Characterization of cDNAs encoding diacylglycerol acyltransferase from cultures of Brassica napus and sucrose-mediated induction of enzyme biosynthesis. Biochim. Biophys. Acta 1580 95–109. [PubMed]
  • Oelkers, P., Cromley, D., Padamsee, M., Billheimer, J.T., and Sturley, S.L. (2002). The DGA1 gene determines a second triglyceride synthetic pathway in yeast. J. Biol. Chem. 277 8877–8881. [PubMed]
  • Ottaviano, E., and Mulcahy, D.L. (1989). Genetics of angiosperm pollen. Adv. Genet. 26 1–64.
  • Park, S.K., and Twell, D. (2001). Novel patterns of ectopic cell plate growth and lipid body distribution in the Arabidopsis gemini pollen1 mutant. Plant Physiol. 126 899–909. [PMC free article] [PubMed]
  • Perry, H.J., Bligny, R., Gout, E., and Harwood, J.L. (1999). Changes in Kennedy pathway intermediates associated with increased triacylglycerol synthesis in oil-seed rape. Phytochemistry 52 799–804.
  • Piffanelli, P., Ross, J.H.E., and Murphy, D.J. (1997). Intra- and extracellular lipid composition and associated gene expression patterns during pollen development in Brassica napus. Plant J. 11 549–562. [PubMed]
  • Piffanelli, P., Ross, J.H.E., and Murphy, D.J. (1998). Biogenesis and function of the lipidic structures of pollen grains. Sex. Plant Reprod. 11 65–80.
  • Preuss, D., Lemieux, B., Yen, G., and Davis, R.W. (1993). A conditional sterile mutation eliminates surface components from Arabidopsis pollen and disrupts cell signaling during fertilization. Genes Dev. 7 974–985. [PubMed]
  • Regan, S.M., and Moffatt, B.A. (1990). Cytochemical analysis of pollen development in wild-type Arabidopsis and a male-sterile mutant. Plant Cell 2 877–889. [PMC free article] [PubMed]
  • Routaboul, J.M., Benning, C., Bechtold, N., Caboche, M., and Lepiniec, L. (1999). The TAG1 locus of Arabidopsis encodes for a diacylglycerol acyltransferase. Plant Physiol. Biochem. 37 831–840. [PubMed]
  • Saha, S., Enugutti, B., Rajakumari, S., and Rajasekharan, R. (2006). Cytosolic triacylglycerol biosynthetic pathway in oilseeds. Molecular cloning and expression of peanut cytosolic diacylglycerol acyltransferase. Plant Physiol. 141 1533–1543. [PMC free article] [PubMed]
  • Sambrook, J., and Russell, D. (2001). Molecular Cloning: A Laboratory Manual, 3rd ed. (Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press).
  • Sandager, L., Gustavsson, M.H., Stahl, U., Dahlqvist, A., Wiberg, E., Banas, A., Lenman, M., Ronne, H., and Stymne, S. (2002). Storage lipid synthesis is non-essential in yeast. J. Biol. Chem. 277 6478–6482. [PubMed]
  • Shockey, J.M., Gidda, S.K., Chapital, D.C., Kuan, J.C., Dhanoa, P.K., Bland, J.M., Rothstein, S.J., Mullen, R.T., and Dyer, J.M. (2006). Tung tree DGAT1 and DGAT2 have nonredundant functions in triacylglycerol biosynthesis and are localized to different subdomains of the endoplasmic reticulum. Plant Cell 18 2294–2313. [PMC free article] [PubMed]
  • Siloto, R.M., Truksa, M., He, X., McKeon, T., and Weselake, R.J. (2009). Simple methods to detect triacylglycerol biosynthesis in a yeast-based recombinant system. Lipids 44 963–973. [PubMed]
  • Somerville, C., Browse, J., Jaworski, J.G., and Ohlrogge, J.B. (2001). Lipids. In Biochemistry and Molecular Biology of Plants, B.B. Buchanan, W. Gruissem, and R.L. Jones, R.L., eds (Rockville, MD: American Society of Plant Biologists), pp. 456–527.
  • Ståhl, U., Carlsson, A.S., Lenman, M., Dahlqvist, A., Huang, B., Banas, W., Banas, A., and Stymne, S. (2004). Cloning and functional characterization of a phospholipid:diacylglycerol acyltransferase from Arabidopsis. Plant Physiol. 135 1324–1335. [PMC free article] [PubMed]
  • Stanley, R.G., and Linskens, H.F. (1974). Pollen. (New York: Springer-Verlag).
  • Stobart, K., Mancha, M., Lenman, M., Dahlqvist, A., and Stymne, S. (1997). Triacylglycerols are synthesised and utilized by transacylation reactions in microsomal preparations of developing safflower (Carthamus tinctorius L) seeds. Planta 203 58–66.
  • Stoveken, T., Kalscheuer, R., Malkus, U., Reichelt, R., and Steinbuchel, A. (2005). The wax ester synthase/acyl coenzyme A:diacylglycerol acyltransferase from Acinetobacter sp strain ADP1: Characterization of a novel type of acyltransferase. J. Bacteriol. 187 1369–1376. [PMC free article] [PubMed]
  • Stuitje, A.R., Verbree, E.C., van der Linden, K.H., Mietkiewska, E.M., Nap, J.P., and Kneppers, T.J.A. (2003). Seed-expressed fluorescent proteins as versatile tools for easy (co)transformation and high-throughput functional genomics in Arabidopsis. Plant Biotechnol. J. 1 301–309. [PubMed]
  • Stymne, S., and Stobart, A.K. (1987). Triacylglycerol biosynthesis. In The Biochemistry of Plants, Vol. 9, Lipids: Structure and Function. P.K. Stumpf, ed (New York: Academic Press), pp 175–214.
  • Taylor, D.C., et al. (2009). Molecular modification of triacylglycerol accumulation by over-expression of DGAT1 to produce canola with increased seed oil content under field conditions. Botany 87 533–543.
  • van Herpen, N.A., and Schrauwen-Hinderling, V.B. (2008). Lipid accumulation in non-adipose tissue and lipotoxicity. Physiol. Behav. 94 231–241. [PubMed]
  • Wang, H.W., Zhang, J.S., Gai, J.Y., and Chen, S.Y. (2006). Cloning and comparative analysis of the gene encoding diacylglycerol acyltransferase from wild type and cultivated soybean. Theor. Appl. Genet. 112 1086–1097. [PubMed]
  • Weselake, R. (2005). Storage lipids. In Plant Lipids, D.J. Murphy, ed (Oxford, UK: Blackwell Publishing), pp. 162–206.
  • Weselake, R.J., et al. (2008). Metabolic control analysis is helpful for informed genetic manipulation of oilseed rape (Brassica napus) to increase seed oil content. J. Exp. Bot. 59 3543–3549. [PMC free article] [PubMed]
  • Wilkinson, E.J., Twell, D., and Lindsey, K. (1997). Activities of CaMV 35S and nos promoters in pollen: Implications for field release of transgenic plants. J. Exp. Bot. 48 265–275.
  • Wolters-Arts, M., Lush, W.M., and Mariani, C. (1998). Lipids are required for directional pollen-tube growth. Nature 392 818–821. [PubMed]
  • Xu, J.Y., Francis, T., Mietkiewska, E., Giblin, E.M., Barton, D.L., Zhang, Y., Zhang, M., and Taylor, D.C. (2008). Cloning and characterization of an acyl-CoA-dependent diacylglycerol acyltransferase 1 (DGAT1) gene from Tropaeolum majus, and a study of the functional motifs of the DGAT protein using site-directed mutagenesis to modify enzyme activity and oil content. Plant Biotechnol. J. 6 799–818. [PubMed]
  • Zhang, Q., Chieu, H.K., Low, C.P., Zhang, S.C., Heng, C.K., and Yang, H.Y. (2003). Schizosaccharomyces pombe cells deficient in triacylglycerols synthesis undergo apoptosis upon entry into the stationary phase. J. Biol. Chem. 278 47145–47155. [PubMed]
  • Zheng, P., et al. (2008). A phenylalanine in DGAT is a key determinant of oil content and composition in maize. Nat. Genet. 40 367–372. [PubMed]
  • Zheng, Z.F., Xia, Q., Dauk, M., Shen, W.Y., Selvaraj, G., and Zou, J.T. (2003). Arabidopsis AtGPAT1, a member of the membrane-bound glycerol-3-phosphate acyltransferase gene family, is essential for tapetum differentiation and male fertility. Plant Cell 15 1872–1887. [PMC free article] [PubMed]
  • Zou, J.T., Wei, Y.D., Jako, C., Kumar, A., Selvaraj, G., and Taylor, D.C. (1999). The Arabidopsis thaliana TAG1 mutant has a mutation in a diacylglycerol acyltransferase gene. Plant J. 19 645–653. [PubMed]

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