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Keywords:

  • Arabidopsis thaliana ;
  • triacylglycerol;
  • phospholipid:diacylglycerol acyltransferase;
  • premature cell death;
  • free fatty acid

Summary

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Phospholipid:diacylglycerol acyltransferase (PDAT) and diacylglycerol:acyl CoA acyltransferase play overlapping roles in triacylglycerol (TAG) assembly in Arabidopsis, and are essential for seed and pollen development, but the functional importance of PDAT in vegetative tissues remains largely unknown. Taking advantage of the Arabidopsis tgd1–1 mutant that accumulates oil in vegetative tissues, we demonstrate here that PDAT1 is crucial for TAG biosynthesis in growing tissues. We show that disruption of PDAT1 in the tgd1–1 mutant background causes serious growth retardation, gametophytic defects and premature cell death in developing leaves. Lipid analysis data indicated that knockout of PDAT1 results in increases in the levels of free fatty acids (FFAs) and diacylglycerol. In vivo 14C-acetate labeling experiments showed that, compared with wild-type, tgd1–1 exhibits a 3.8-fold higher rate of fatty acid synthesis (FAS), which is unaffected by disruption or over-expression of PDAT1, indicating a lack of feedback regulation of FAS in tgd1–1. We also show that detached leaves of both pdat1–2 and tgd1–1 pdat1–2 display increased sensitivity to FFA but not to diacylglycerol. Taken together, our results reveal a critical role for PDAT1 in mediating TAG synthesis and thereby protecting against FFA-induced cell death in fast-growing tissues of plants.


Introduction

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Plants store oils in the form of triacylglycerols (TAGs) in oil droplets in seeds and fruits, and the potential of these as food, feed, biodiesel and industrial chemicals has been explored (Carlsson et al., 2011; Lu et al., 2011). In addition to accumulation in these dedicated storage organs, TAGs also accumulate in non-seed tissues such as flower petals of Dianthus caryophyllus (Hudak and Thompson, 1997), mature pollen grains of flower plants (Murphy and Vance, 1999), stems and leaves of some plant species (Durrett et al., 2008), and in tubers of Cyperus esculentus (Turesson et al., 2010), albeit in lower abundance. Plant non-seed biomass represents the most abundant feedstock source for renewable bioenergy on earth (Schubert, 2006), and therefore a detailed understanding of TAG metabolism in vegetative tissues has important implications for the development of new energy crops for biofuel production (Durrett et al., 2008; Ohlrogge et al., 2009; Chapman et al., 2013; Troncoso-Ponce et al., 2013). In addition, studies in yeast and mammalian systems suggested that TAG accumulation fulfills specific physiological functions such as protection against cellular lipotoxicity induced by overload of free fatty acids (FFAs) and other lipid intermediates (Ducharme and Bickel, 2008; Yen et al., 2008; Kohlwein, 2010). The pathway and enzymes involved in TAG synthesis and the physiological importance of TAG homeostasis in non-seed tissues of plants remain largely unexplored.

TAG biosynthesis in plants occurs in the endoplasmic reticulum (ER) via the Kennedy pathway, involving three sequential acylations of glycerol-3–phosphate with acyl chains exclusively originating from the plastid (Ohlrogge and Browse, 1995; Chapman and Ohlrogge, 2012; Bates et al., 2013). The first two acylation reactions are shared between TAG and membrane lipid biosynthesis, and result in the generation of phosphatidic acid (PA). Dephosphorylation of PA by PA phosphatase generates diacylglycerol (DAG), which is used for the final acylation reaction to produce TAG, catalyzed by diacylglycerol:acyl CoA acyltransferase (DGAT) and phospholipid:diacylglycerol acyltransferase (PDAT) (Chapman and Ohlrogge, 2012; Bates et al., 2013). At least three distinct classes of DGATs, namely DGAT1 (Routaboul et al., 1999; Zou et al., 1999), DGAT2 (Shockey et al., 2006) and a soluble DGAT (Saha et al., 2006; Hernández et al., 2012) have been reported in plants, and biochemical and genetic evidence has established DGAT1 as a major player mediating TAG biosynthesis in developing seeds (Routaboul et al., 1999; Zou et al., 1999) and senescent leaves (Kaup et al., 2002; Slocombe et al., 2009) of Arabidopsis. Two close homologs of the yeast PDAT gene were identified in the Arabidopsis genome (Dahlqvist et al., 2000), but only PDAT1 activity has been confirmed by over-expression of PDAT1 in Arabidopsis (Ståhl et al., 2004), and PDAT1 has been shown to play overlapping roles with DGAT1 in TAG assembly in pollen grains and developing seeds (Zhang et al., 2009). Both DGAT1 and PDAT1 genes are expressed in leaves, roots and stems, in addition to developing seeds and flowers (Zou et al., 1999; Ståhl et al., 2004), but the functional role of PDAT in vegetative tissues remains to be elucidated. In addition to serving as an immediate precursor for TAG synthesis, DAG is also a substrate for synthesis of the major phospholipids phosphatidylcholine (PC) and phosphatidylethanolamine (PE) in the ER, catalyzed by aminoalcoholphosphotransferases (Dewey et al., 1994; Ohlrogge and Browse, 1995; Goode and Dewey, 1999).

PC plays a central role in the biosynthesis of TAG and membrane lipids (Bates et al., 2013). It is the major source of DAG for TAG synthesis in developing oilseeds (Bates et al., 2009; Bates and Browse, 2012). Importantly, this phospholipid undergoes constant acyl editing involving a deacylation and reacylation cycle, whereby the nascent fatty acids (FAs) exported from the plastid are desaturated prior to being used for de novo glycerolipid biosynthesis through stepwise glycerol-3–phosphate acylation (Bates et al., 2007, 2009; Tjellström et al., 2012).

The central role of PC in glycerolipid biosynthesis is also indicated by the fact that PC-derived lipids serve as precursors for the synthesis of galactolipids via the eukaryotic pathway involving the ER and the plastid (Roughan and Slack, 1982; Ohlrogge and Browse, 1995), although the extent to which the eukaryotic pathway contributes to thylakoid lipid assembly varies depending on plant species and the various tissues within the plant. For example, in pea plants (Pisum sativum) (Mongrand et al., 1998) and green seeds of Arabidopsis (Xu et al., 2005), the eukaryotic pathway is responsible for the assembly of as much as 90% of thylakoid lipids, whereas in the leaves of Arabidopsis and spinach (Browse et al., 1986), approximately half of the photosynthetic membrane lipids are derived from this pathway. The remainder are assembled by a parallel pathway commonly referred to as the prokaryotic pathway, in which FAs are used directly for stepwise glycerol-3–phosphate acylation to generate PA in the plastid. The resultant PA and its dephospharylated product DAG serve almost exclusively as precursors for the synthesis of thylakoid membrane lipids involving the enzyme machinery present in envelope membranes (Dörmann et al., 1999; Awai et al., 2001; Benning and Ohta, 2005).

The Arabidopsis tgd1–1 mutant, so named because it accumulates unusual oligogalactolipids, particularly trigalactosyldiacylglycerol (TGDG), in leaves, is defective in the transport of ER-derived lipid precursors into plastids due to a point mutation in a gene encoding a permease-like component of an ABC transporter complex (Xu et al., 2003). This results in a drastic decrease in the amounts of thylakoid lipids produced by the eukaryotic pathway, with a compensatory increase in galactolipids produced by the prokaryotic pathway. Intriguingly, the tgd1–1 mutation also leads to accumulation of TAG in oil droplet-like structures in the cytosol in leaves (Xu et al., 2005), and, despite the drastic alterations in FA fluxes in tgd1 mutants, their growth and development are only slightly compromised (Xu et al., 2005). Thus, the tgd1 mutants offer a valuable tool to dissect TAG metabolism and homeostasis in non-seed tissues of plants, particularly at the molecular genetic level. Here we demonstrate that PDAT1 plays a critical role in mediating TAG synthesis in fast-growing tissues. Inactivation of PDAT1 in the tgd1–1 mutant background causes gametophytic defects and premature cell death in developing leaves, probably due to accumulation of cytotoxic FFAs. The possible functional role of PDAT1 in maintaining lipid homeostasis in plant non-seed tissues is discussed.

Results

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

PDAT1 plays a critical role in TAG synthesis in growing tissues

The tgd1–1 mutant accumulates TAG in leaves due to a defect in ER-to-plastid lipid trafficking (Xu et al., 2005). Because the activities of lipid synthesis and trafficking vary greatly between different plant tissues (Xu et al., 2005) or in the same tissues at different developmental stages (Andersson et al., 2001; Hellgren and Sandelius, 2001), we analyzed tissue-specific and development-related variations in TAG content in the tgd1–1 mutant. On a dry weight basis, the amounts of TAG were approximately three- and tenfold higher in developing leaves than in mature and senescing leaves, respectively (Figure 1a). Among the tissues examined, the highest TAG level was measured in flowers, with a level approximately tenfold higher than in developing leaves.

image

Figure 1. PDAT1 plays a critical role in TAG synthesis in growing tissues.

(a) TAG content of tgd1–1, tgd1–1 dgat1–1 and tgd1 pdat1–2 mutants.

(b) FA composition of TAG from developing leaves of tgd1–1, tgd1–1 dgat1–1 and tgd1 pdat1–2 mutants.

Lipids were extracted from senescing leaves (SL), mature leaves (ML), developing leaves (DL) and flowers (F) of 5–6-week-old plants. The inset in (a) shows the TAG content in SL and ML with a different y scale. Values are means and standard deviation of three to five replicates.

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To dissect the biochemical pathway leading to TAG accumulation in non-seed tissues, we constructed double mutants of tgd1–1 with either dgat1–1 or pdat1–2, which harbor an ethylmethanesulfonate-induced lesion in the DGAT1 locus (Zou et al., 1999) and a T–DNA insertion in the PDAT1 gene (Zhang et al., 2009), respectively. Compared with tgd1–1, the tgd1–1 pdat1–2 double mutant showed a 65% decrease in TAG in developing leaves and a 59% decrease in flowers, but only 27% in mature leaves and 21% in senescing leaves (Figure 1a). In comparison, the tgd1–1 dgat1–1 double mutant showed a less pronounced decrease in the TAG level in developing leaves (34%) and flowers (25%), but a more severe decrease in the TAG content in mature (51%) and senescing (59%) leaves.

In addition to changes in TAG content, disruption of PDAT1 in tgd1–1 also resulted in a marked increase in the proportions of saturated FAs at the expense of polyunsaturated acyl chains in TAG from developing leaves (Figure 1b) and flowers (Figure S1a) compared with tgd1–1, whereas there were limited changes in the FA profiles of TAG in tgd1–1 dgat1–1. A similar increase in the levels of saturated FAs, with a concomitant decrease in polyunsaturated acyl chains, was also observed in TAG from the flowers of the pdat1–2 single mutant compared with the wild-type (Figure S1b).

Over-expression of PDAT1 enhances TAG accumulation in the leaves of the tgd1–1 mutant

We next generated transgenic plants over-expressing PDAT1 or DGAT1 driven by the constitutive CaMV 35S promoter. Analysis of TAG content in developing leaves of 5-week-old soil-grown plants showed up to fivefold increases in three independent transgenic lines over-expressing PDAT1 compared with tgd1–1, but no significant increase in TAG content was observed in transgenic lines over-expressing DGAT1 (Figure 2a). Ultrastructural analysis using transmission electron microscopy (TEM) showed that, in contrast to minute oil droplet-like structures in tgd1–1 (Figure 2c) and tgd1–1 dgat1–1 (Figure S2b), over-expression of PDAT1 in tgd1–1 led to accumulation of large cytosolic oil droplets (Figure 2d). No oil droplets were observed in wild-type and tgd1–1 pdat1–2 (Figure S2a,c). Similar to TAG from developing leaves of tgd1–1 (Figure 1b), TAG from PDAT1 over-expressing lines was highly enriched in polyunsaturated FAs, particularly 18:2 and 18:3 (Figure 2b). Because in vitro biochemical assays have shown that PDAT1 has a strong preference for polyunsaturated acyl chains over FAs with fewer than two double bonds (Ståhl et al., 2004), the observations that knockout of PDAT1 decreased (Figure 1b), while over-expression of this gene increased (Figure 2b), the level of FA unsaturation of TAG strengthen the idea that PDAT1 plays a key role in TAG synthesis in growing leaves.

image

Figure 2. Over-expression of PDAT1 enhances TAG accumulation in tgd1–1.

(a) TAG levels in developing leaves of 5-week-old transgenic lines over-expressing (OE) DGAT1 or PDAT1 in the tgd1–1 background.

(b) FA composition of TAG from developing leaves of three independent transgenic lines over-expressing PDAT1.

(c, d) TEM micrographs of leaf cells from the tgd1–1 mutant (c) and tgd1–1 PDAT1 over-expressing line 1 (PDAT1-OE1) (d). The white arrow in (c) indicates an oil droplet. Scale bar = 1 μm.

Values in (a, b) are means and standard deviation of three biological replicates. Asterisks indicate a statistically significant difference compared with tgd1–1 based on a two-tailed Student's t-test (< 0.05).

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Disruption of PDAT1 in the tgd1–1 mutant compromises plant growth and development and causes premature cell death in growing leaves

Although the incidence and severity of symptoms varied among individuals, disruption of PDAT1 in the tgd1–1 background resulted in pleiotropic phenotypes, including severe growth retardation compared with tgd1–1 (Figure 3a), premature primary shoot meristem arrest and aberrant leaf morphology (Figures 3b), while tgd1–1 dgat1–1 plants were visually indistinguishable from tgd1–1 (Figure 3a). Furthermore, the leaves of tgd1–1 pdat1–2 plants grown under aseptic conditions often displayed spontaneous necrotic lesions (Figure 3b). After transfer to soil, approximately 10% of 3-week-old tgd1–1 pdat1–2 seedlings (35/378) died within 2 weeks. Those that survived often displayed the necrotic phenotype that appeared during leaf expansion (Figure 3c). The necrotic lesions in the emerging leaves did not appear to increase substantially in size during leaf maturation (Figure 3c,d).

image

Figure 3. Growth defects and spontaneous cell death in tgd1–1 pdat1–2.

(a) Size comparison of 3-week-old tgd1–1, tgd1–1 dgat1–1 and tgd1–1 pdat1–2 plants grown on half-strength agar-solidified MS medium.

(b) Three-week-old tgd1–1 pdat1–2 plants grown on MS medium showing primary shoot meristem arrest and spontaneous necrotic lesion formation on leaves (arrows).

(c) Five-week-old tgd1–1 pdat1–2 plants showing the appearance of necrotic lesions in developing leaves (arrows).

(d) Photograph of the same tgd1–1 pdat1–2 plant as shown in (c) at 6 weeks old.

Note the similar size of necrotic lesions (arrows) on the same leaf at the different stages of growth in (c) and (d). Scale bars = 1 mm.

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The tgd1–1 pdat1–2 double mutant is defective in gametophytic development

The adult tgd1–1 pdat1–2 plants were severely reduced in stature, showing a bushy, compact appearance (Figure S3a). They did not produce siliques, but did produce numerous flower buds that in most cases turned yellow and eventually aborted before opening (Figure S3b,c). Visual examination of the flowers showed that, whereas there were no obvious morphological differences in anthers between tgd1–1 and tgd1–1 dgat1–1 (Figure 4a,b), the tgd1–1 pdat1–2 anthers were smaller, shrunken, dry and brownish in color and lacked any obvious pollen grains (Figure 4c). In addition, cross-pollination of the tgd1–1 pdat1–2 stigmas with wild-type pollen failed to produce siliques, indicating that female reproductive development is also compromised in tgd1–1 pdat1–2.

image

Figure 4. Defects in anther development and dehiscence in tgd1–1 pdat1–2.

(a–c) Flower phenotypes of tgd1–1 (a), tgd1 dgat1–1 (b) and tgd1–1 pdat1–2 (c).

(d, e) Anthers at stage 12.

(f, g) Anthers at stage 13. DPG, degenerated pollen grain; Den, degenerated endothecium; E, epidermis; En, endothecium; Sm, septum; St, stomium.

Scale bars = 20 μm.

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The lack of obvious pollen grains on anthers may be a result of pollen abortion or defects in anther dehiscence or both. To distinguish between these possibilities, histochemical analysis was performed to examine the detailed morphology of tgd1–1 and tgd1–1 pdat1–2 anthers at stages 12 and 13 as defined by Sanders et al. (1999). As shown in Figure 4(d), anthers at stage 12 in tgd1–1 were bi-locular and contained tricellular round pollen grains, which were eventually released following anther dehiscence (Figure 4f), whereas pollen grains in the double mutant anthers were shrunken and aggregated (Figure 4e), with cell debris remaining in anther locules at late stage 13 (Figure 4g). Both the septum and stomium failed to degenerate, resulting in indehiscent anthers in the double mutant. Taken together, these results suggest that disruption of PDAT1 in the tgd1–1 mutant background causes defects in both male and female gametophytic development and anther dehiscence.

FFA and DAG accumulate in tgd1–1 pdat1–2

To obtain insight into the biochemical basis that underlies the cell death phenotypes in tgd1–1 pdat1–2, we first measured the levels of FFA and DAG in developing leaves and flowers of the single and double mutants. Compared with tgd1–1, the tgd1–1 pdat1–2 double mutant had a 1.8-fold increase in FFA content in developing leaves and a 2.3-fold increase in FFA in flowers (Figure 5a). There were also 2.3-fold and 1.4-fold increases in the DAG level in developing leaves and flowers, respectively, in tgd1–1 pdat1–2 relative to tgd1–1 (Figure 5b). In contrast to tgd1–1 pdat1–2, the increases in amounts of FFA and DAG in developing leaves and flowers of tgd1–1 dgat1–1 were much less pronounced, despite the substantial decrease in TAG content in tgd1–1 dgat1–1 (Figure 1a).

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Figure 5. Accumulation of FFAs and DAG in tgd1 pdat1–2.

FFAs (a) and DAG (b) were isolated from developing leaves (DL) and flowers (F) of 5-week-old plants of the tgd1–1, tgd1–1 dgat1–1 and tgd1 pdat1–2 mutants grown on soil. Values are means and standard deviation of three replicates. Asterisks indicate a statistically significant difference compared with tgd1–1 based on a two-tailed Student's t-test (< 0.05).

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Similar to what has been observed in young rosette leaves (Li et al., 2011), 16:0 and 18:3 were the most abundant FFA in developing leaves of tgd1–1, followed by 18:2 and 18:0 (Figure S4a). Disruption of DGAT1 or PDAT1 did not lead to substantial changes in FFA composition in tgd1–1. The major FAs in DAG derived from developing leaves of tgd1–1 and double mutants were 18:2, 16:0, 18:3, 18:1 and 18:0 (descending order of abundance) (Figure S4b). Compared with tgd1–1, both tgd1–1 dgat1–1 and tgd1–1 pdat1–2 showed decreased levels of 18:0 with corresponding increases in 16:0.

Phospholipid levels are elevated in tgd1–1 pdat1–2

To assess to what extent the deficiency of TAG synthesis affects membrane lipids in non-seed tissues, major polar lipid classes in developing leaves of the single and double mutants were quantified. Although no significant differences in polar lipid content were found between tgd1–1 and tgd1–1 dgat1–1, the tgd1–1 pdat1–2 double mutant showed significant increases in levels of PC and PE compared with tgd1–1 (Figure 6). The FA composition of major membrane lipids was also altered. Notably, there were substantial increases in 18:1 across all lipid species examined (Figure S5), particularly PC and PE. In PC, the increase in 18:1 was accompanied by decreases in 18:0 and 18:3; In PE, there was a decrease in 18:2. There were substantial decreases in 16:3 in digalactosyldiacylglycerol and monogalactosyldiacylglycerol.

image

Figure 6. Elevated phospholipid levels in tgd1–1 pdat1–2.

Lipids were extracted from developing leaves of 5-week-old plants grown on soil. Values are means and standard deviation of three biological replicates. Asterisks indicate a statistically significant difference compared with tgd1–1 based on a two-tailed Student's t-test (< 0.05). MGDG, monogalactosyldiacylglycerol; PG, phosphatidylglycerol; DGDG, digalactosyldiacylglycerol; PI, phosphatidylinositol; SL, sulfoquinovosyldiacylglycerol; PE, phosphatidylethanolamine; PC, phosphatidylcholine.

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The rate of FA synthesis drastically increases in tgd1–1 and tgd1–1 pdat1–2 mutants and PDAT1 transgenic plants

As fatty acid synthesis (FAS) is known to be subjected to feedback inhibition by 18:1-acyl carrier protein in plants (Andre et al., 2012), the accumulation of FFA raises questions regarding the regulation of FAS in tgd1–1 and tgd1 pdat1–2. To assess whether the in vivo rate of FAS is altered due to genetic modifications in tgd1–1, initial rates of 14C-acetate incorporation into growing leaves of single and double mutants and transgenic lines over-expressing PDAT1 were analyzed. To minimize the potential difference in endogenous substrate pool size (Cronan et al., 1975; Nunn et al., 1977), the labeling experiment was performed in the presence of 1 mm cold acetate. This analysis showed a 3.8-fold increase in the rate of FAS in tgd1–1 relative to that in wild-type (Figure 7a). In addition, neither disruption nor over-expression of PDAT1 in tgd1–1 resulted in substantial changes in the rate of FAS (Figure 7a), despite large alterations in leaf TAG content (Figures 1a and 2a). The implications of these results are twofold: (i) a block in ER-to-plastid lipid trafficking and a consequent enhancement of the prokaryotic pathway of thylakoid lipid synthesis results in relief of feedback inhibition on FAS in tgd1–1, presumably because of the increased consumption of 18:1-acyl carrier protein in the plastid by the prokaryotic pathway, and (ii) the rate of FAS in tgd1–1 is not regulated in response to large changes in the demand for FAs outside the plastid.

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Figure 7. Rates of FA synthesis and degradation in the tgd1–1 mutant, and effects of PDAT1 alterations.

Developing leaves of 5-week-old plants were labeled with 14C-acetate for 1 h, and the label chased for a period of 3 days. Values are means and standard deviation of three replicates.

(a) Initial rates of 14C-acetate incorporation into wild-type (Col–2), tgd1–1, tgd1–1 dgat1–1, tgd1 pdat1–2 and PDAT1 over-expressing (OE) lines 1 and 8.

(b, c) FA turnover in Col–2 and tgd1–1 (b) and tgd1–1, tgd1–1 pdat1–2 and PDAT1-OE1 (c).

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The rate of FA degradation increases in the tgd1–1 mutant and remains unaltered in the tgd1–1 pdat1–2 mutant, but decreases in PDAT1 transgenic plants

A 3.8-fold increase in the rate of FAS in the tgd1–1 mutant is expected to result in a corresponding increase in total lipid content, assuming that the rate of FA degradation remains constant. However, quantification of the total amounts of FAs in developing leaves of 5-week-old plants by gas chromatography revealed no substantial difference between wild-type (5.26 ± 0.67 mg g−1 FW, = 3) and tgd1–1 (4.65 ± 0.24 mg g−1 FW, = 3), suggesting that the rate of FA breakdown in tgd1–1 increases in proportion to FAS. To verify this assumption, detached developing leaves were pulsed with 14C-acetate for 1 h, and changes in the amount of total label in FAs were then followed for 3 days. The results showed a 3.3-fold higher mean rate of labeled FA degradation in tgd1–1 (20% per day) compared with wild-type (6% per day) (Figure 7b).

We next tested whether knockout or over-expression of PDAT1 affects the rate of FA turnover in tgd1–1. The results in Figure 7(c) show that, while no substantial difference was noted between tgd1–1 and tgd1–1 pdat1–2, over-expression of PDAT1 resulted in a marked decrease in the rate of labeled FA degradation in tgd1–1. The decreased rate of FA turnover may explain why over-expression of PDAT1 in tgd1–1 enhances TAG accumulation (Figure 2a) without a concomitant increase in the rate of FAS (Figure 7a). At present, we do not know why over-expression of PDAT1 in tgd1–1 led to decreased FA turnover. A likely explanation is that TAGs in large oil droplets in the leaves of PDAT1 over-expressors (Figure 2d) may be less susceptible to lipase attack due to a decrease in the surface area to volume ratio of oil droplets. In line with this possibility, previous studies have showed that increases in the size of oil droplets are associated with decreases in the rate of TAG hydrolysis during seed germination (Siloto et al., 2006; Shimada et al., 2008).

Disruption of PDAT1 increases the sensitivity of growing leaves to exogenous unsaturated FAs

In yeast and mammalian cells, deficiency of TAG synthesis often leads to increased sensitivity to exogenous FA supplementation (Garbarino et al., 2009; Petschnigg et al., 2009). To investigate whether this is also the case in plants, detached developing leaves of single and double mutants were incubated in solutions containing various FFAs for 24 h in the light at 22°C. As shown in Figure 8(a), while tgd1–1 and tgd1–1 dgat1–1 leaves remained largely normal in appearance after treatment with 2.5 mm 18:1, incubation with the same concentration of 18:1 led to severe chlorosis in developing leaves of tgd1–1 pdat1–2. Compared with 18:1, polyunsaturated FAs such as 18:2 are apparently more destructive, causing small chlorotic lesions, particularly at the leaf edge, even in tgd1–1 and tgd1–1 dgat1–1 and a severe bleaching phenotype in tgd1–1 pdat1–2 at a much lower concentration. Similar to double mutants, developing leaves of the pdat1–2 single mutant also showed increased susceptibility to 18:1 (Figure 8b). In contrast to polyunsaturated acyl chains, long-chain saturated FAs, possibly because of their extremely low solubility in water, did not have any obvious effect even after extended treatment for 3 days (Figure S6a).

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Figure 8. Disruption of PDAT1 results in increased sensitivity to exogenous FAs.

Detached developing leaves of 5-week-old tgd1–1, tgd1–1 dgat1–1 and tgd1 pdat1–2 (a) or Col–2, dgat1–1 and pdat1–2 (b) plants were floated in water containing 0.0005% Tween–20 and 0.25% ethanol and 2.5 mm 18:1 or 1.25 mm 18:2. Photographs were taken after 24 h of treatment. Controls were treated with water containing 0.0005% Tween–20 and 0.25% ethanol.

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In addition to FFAs, the DAG content also increased in tgd1 pdat1–2. To test whether the increased DAG levels contribute to the cell death phenotype, detached developing leaves were incubated with either 18:1/18:1 DAG or the more water-soluble short-chain 8:0/8:0 DAG. Compared with controls, treatment with 18:1/18:1 or 8:0/8:0 DAG did not induce any visible phenotype in tgd1–1, tgd1–1 dgat1–1 and tgd1 pdat1–2 even after extended incubation for 3 days (Figure S6a,b). Together, these results suggest that elevated levels of FFAs, rather than DAG, may be responsible for premature cell death in growing tissues of tgd1 pdat1–2.

We next tested whether over-expression of PDAT1 alters the performance of tgd1–1 following challenge with exogenous FAs. To this end, developing leaves of tgd1–1 and two independent transgenic lines over-expressing PDAT1 in the tgd1–1 background were incubated with 18:1. Treatment with 2.5 mm 18:1 for 36 h led to a severe bleaching phenotype, and no obvious difference in appearance was observed between tgd1–1 and PDAT1 over-expressors (Figure S6c). At present, we do not understand why PDAT1 over-expression does not increase the resistance of tgd1–1 to exogenous FA treatment, despite markedly enhanced TAG synthesis in PDAT1 over-expressors (Figure 2a). One interpretation is that the protective effects of PDAT1 over-expression on FA-induced cell death may be limited by regeneration of PC from lysophosphatidylcholine (LPC). In line with this possibility, it has recently been shown that reacylation of LPC catalyzed by acyl CoA acyltransferase 2 (LPCAT2) is critical for PDAT1-mediated TAG synthesis in the absence of DGAT1 function, and disruption of LPCAT2 severely compromises the growth and development of the dgat1–1 mutant (Xu et al., 2012).

Discussion

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

The function of TAG synthesis in protection against FA-induced cell death is conserved among eukaryotes

Previous studies in yeast and mammalian model systems have firmly established that, in addition to serving as a means for efficient carbon and energy storage, TAG synthesis plays a critical role in detoxifying FFAs and other lipid metabolic intermediates (Yen et al., 2008; Kohlwein, 2010). This conclusion is based primarily on two key observations. First, deficiency of TAG synthesis genes renders both yeast (Garbarino et al., 2009; Petschnigg et al., 2009) and mammalian cells (Listenberger et al., 2003) highly sensitive to exogenous FA supplementation in terms of cell growth and survival. Second, disruption of TAG synthesis induces programmed cell death (Buszczak et al., 2002; Zhang et al., 2003; Stone et al., 2004), which has been linked to increased levels of FFAs (Fakas et al., 2011) or DAG (Zhang et al., 2003) in yeast. Although direct evidence for a role of TAG synthesis in protection against toxic lipid intermediates is still lacking in plants, it has been shown that Arabidopsis double mutants defective in DGAT1 and PDAT1 lack viable pollen, and RNA interference-mediated silencing of DGAT1 or PDAT1 in the pdat1 or dgat1 mutant background, respectively, results in defects in pollen and embryo development (Zhang et al., 2009). In addition, deficiency of DGAT1 activity in Brassica napus (Lock et al., 2009) and in Arabidopsis wild-type plants (Katavic et al., 1995; Zou et al., 1999) and mutants defective in FA breakdown (Slocombe et al., 2009) has been shown to have negative effects on plant growth and development. The results of the present study show that knockout of PDAT1 increases the sensitivity of developing leaves to exogenous FFAs. In addition, disruption of PDAT1 in the tgd1–1 mutant background causes premature cell death in developing leaves and floral organs, with concomitant increases in the levels of FFAs and DAG. Taken together, these results demonstrate an evolutionally conserved role for TAG synthesis in protection against the toxicity induced by FFAs and possibly other lipid metabolic intermediates in yeast, plants and mammals.

PDAT plays a critical role in lipid homeostasis

A working model for a critical role of PDAT1 in buffering cells against the cytotoxic effects of FFAs in non-seed tissues is shown in Figure 9. FAs are almost exclusively synthesized in the plastid (Ohlrogge et al., 1979). In Arabidopsis leaves, approximately 40% of these newly synthesized acyl chains are directly utilized in the plastid to support the prokaryotic pathway of thylakoid lipid synthesis (Browse et al., 1986). The remainder are exported outside the plastid, and first used to acylate LPC to produce PC, catalyzed by LPCATs (Bates et al., 2007, 2012; Wang et al., 2012). After 18:1 desaturation to 18:2 and 18:3 by ER-resident desaturases, the resulting polyunsaturated PC has multiple possible metabolic fates including (i) deacylation to LPC and FAs as part of the acyl-editing cycle (Bates et al., 2007, 2009; Tjellström et al., 2012), (ii) conversion to thylakoid lipids through the eukaryotic pathway, which brings approximately 40% of the exported FAs (24% of the total newly synthesized FAs) back into chloroplasts (Browse et al., 1986), (iii) conversion to polyunsaturated DAG for TAG synthesis (Bates and Browse, 2012), or (iv) transacylation catalyzed by PDAT1, resulting in TAG and LPC. In growing leaves of wild-type plants, <3% of nascent acyl chains are used for TAG synthesis (Figure S7). In the tgd1–1 mutant, a defect in the transport of PC-derived lipid precursors into chloroplasts results in a 3.8-fold increase in the rate of FAS and a threefold increase in the rate of FA breakdown, presumably through β–oxidation. In addition, there is a 2.4-fold increase in the acyl flux into TAG (Figure S7), largely via the acyl CoA-independent reaction catalyzed by PDAT1 (Figure 1a). Reacylation of LPC generated by PDAT1 using nascent FAs exported from the plastid regenerates PC and thus creates a deacylation and reacylation cycle that consumes toxic lipid intermediates such as fatty acyl chains and DAG, with net production of inert TAG.

image

Figure 9. Working model for the function of PDAT1 in maintaining lipid homeostasis in non-seed tissues.

In the wild-type plants, a major fraction of PC or its derivative is transported into the chloroplast for synthesis of glycolipids (GLs). This process is blocked (double lines) in the tgd1–1 mutant, which leads to (i) enhanced prokaryotic thylakoid lipid synthesis, and (ii) constitutive activation of fatty acid synthesis (FAS). FAs in excess of cellular needs are (i) degraded by β–oxidation or (ii) sequestered into inert TAG through the PDAT1-mediated acyl CoA-independent route, thereby protecting cells from detrimental effects of FFAs and possibly other lipid intermediates in the tgd1–1 mutant. Lighter arrows indicate pathways and metabolism that are enhanced in the tgd1–1 mutant. Dashed lines indicate potential sources of DAG and PC for TAG synthesis mediated by PDAT1. PL, phospholipids.

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In developing leaves of the tgd1–1 mutant, disruption of PDAT1 resulted in no substantial changes in the rates of FAS or FA degradation. However, there were significant increases in the PC and PE levels in developing leaves of the tgd1–1 pdat1–2 mutant. Increases in membrane lipid content were also observed in yeast mutants defective in TAG synthesis upon FA supplementation (Garbarino et al., 2009; Petschnigg et al., 2009; Connerth et al., 2010) or even without FA supplementation (Fakas et al., 2011), and such increases have been postulated as an alternative means for protection against toxic effects of FAs. Apparently, the increased phospholipid synthesis is insufficient to fully compensate for the buffering function conferred by PDAT1-mediated TAG synthesis, which may explain why FFA levels increase in tgd1–1 pdat1–2.

Initiation and propagation of the cell death phenotype in tgd1–1 pdat1–2 were observed in rapidly growing cell types but not in mature and senescing tissues (Figure 3). The underlying reason for the observed developmental variations in phenotypes may lie in the fact that both the rates of FAS (Kannangara et al., 1973) and the proportions of de novo synthesized acyl chains exported outside the chloroplast (Andersson et al., 2001) are much higher in expanding leaves than in mature leaves. In addition, transcripts encoding key enzymes of β–oxidation, the major pathway for FA degradation in plants (Gerhardt, 1992), are normally expressed at very low levels during leaf growth but are induced during leaf senescence (Troncoso-Ponce et al., 2013), which may explain why disruption of PDAT1 in growing leaves of tgd1–1 did not lead to a further increase in the rate of FA degradation (Figure 7c). Thus, a block in the return of acyl chains from the ER to chloroplasts results in much more FAs being available for TAG synthesis, and higher accumulation of TAG in developing leaves than in expanded and senescent leaves, and disruption of both acyl chain return and TAG synthesis causes higher accumulation of toxic lipid intermediates, thereby resulting in more deleterious effects in the growing leaves than in mature leaves. A situation similar to that in young growing leaves appears to exist in floral organs, which accumulate almost nine times more TAG compared with the growing leaves of the tgd1–1 mutant, and TAG accumulation in flowers is, to a major extent, mediated by PDAT1 (Figure 1a). Consequently, disruption of PDAT1-mediated TAG synthesis in the tgd1–1 mutant background results in marked increases in levels of FFA and DAG, leading to the observed defects in the development of male and female gametophytes in tgd1–1 pdat1–2 double mutant plants. In this regard, it is worth noting that loss of both PDAT1 and DGAT1 functions in dgat1 pdat1 double mutants affects the viability of male but not female gametophytes (Zhang et al., 2009). Thus, although the amount of TAG is markedly reduced in the flowers of the tgd1–1 pdat1–2 mutant, it seems unlikely that the gametophytic developmental defects in tgd1–1 pdat1–2 are mainly due to the deficiency in TAG synthesis, as both male and female gametophytes are affected in the double mutant.

An important question arising from this work relates to the metabolic origin of DAG for TAG synthesis. In plants, two major pathways for DAG synthesis are (i) de novo synthesis through glycerol-3–phosphate acylation, and (ii) conversion of PC into DAG (Bates and Browse, 2012). In developing seeds, PC is the major DAG donor for TAG synthesis, whereas DAG derived from glycerol-3–phosphate acylation reactions is used for the synthesis of PC (Bates et al., 2009). Similarly, two lines of evidence suggest that PC may be the major source of DAG for TAG synthesis in growing leaves. First, the FA composition of both TAG (Figure 1b) (Xu et al., 2005) and DAG (Figure S4b) in leaves of the tgd1–1 or tgd1–1 pdat1–2 mutants, respectively, resembles that of PC (Figure S5). Second, disruption of PDAT1 results in increases in the levels of PC and PE in growing leaves of tgd1–1 (Figure 6). If PC is the DAG donor for TAG synthesis in developing leaves, the net result of the deacylation and reacylation cycle driven by PDAT1 and LPCAT is production of TAG at the expense of PC and acyl CoA. This is in line with the proposed function of PDAT in the regulation of membrane lipid composition (Dahlqvist et al., 2000).

Potential acyltransferases responsible for LPC reacylation to PC in leaves include LPCAT1 and 2, which have recently been shown to play a major role in PC acyl editing in developing seeds (Bates et al., 2012; Wang et al., 2012). In addition, recent genetic evidence also suggests that LPCAT2 is a key enzyme catalyzing LPC reacylation to support PDAT1-mediated TAG synthesis in seeds of the dgat1–1 mutant (Xu et al., 2012). In rapidly expanding pea leaves, substantial LPCAT activity is associated with chloroplast envelope membranes (Bessoule et al., 1995; Tjellström et al., 2012). It has been proposed that the chloroplast-associated LPCAT is involved in channeled incorporation of nascent acyl chains into PC via acyl editing (Tjellström et al., 2012). Although the exact subcellular localization of PDAT1 remains to be determined, PDAT1 activity has been shown to reside in microsomal membranes of Arabidopsis roots and leaves (Ståhl et al., 2004). Given the amphiphilic nature of LPC, its rapid partitioning between the ER and chloroplast membranes (Bessoule et al., 1995), and the close physical association between the ER and the plastid (Andersson et al., 2007; Xu et al., 2008), it is likely that the PDAT1/LPCAT cycle may be directly involved in directing newly synthesized acyl chains into TAG without mixing with the bulk cytosolic acyl CoA pool.

In summary, our results show that PDAT plays a critical role in mediating TAG synthesis in young growing tissues of plants. Loss of function of PDAT disrupts lipid homeostasis, leading to gametophytic defects and premature cell death in growing cell types. Studies in yeast and mammalian systems suggested that disruption of TAG synthesis often triggers lipoapoptosis, a type of FA-induced programmed cell death that has a distinct set of physiological and morphological features, including nuclear condensation, DNA fragmentation and elevated oxidative stress (Buszczak et al., 2002; Zhang et al., 2003; Fakas et al., 2011). Future studies using pdat1–2 and tgd1–1 pdat1–2 mutants will address how TAG synthesis deficiency affects the cell structure and function, and assess the mechanistic basis underlying cytotoxic effects of FFAs in plant model systems.

Experimental procedures

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

Plant materials and growth conditions

The Arabidopsis thaliana plants used in this study were of the Columbia ecotype (Col–2). The tgd1–1 mutant was previously described by Xu et al. (2003), and the dgat1–1 mutant (originally named ASII, Katavic et al., 1995) and the pdat1–2 mutant were previously described by Zhang et al. (2009). For growth on plates, surface-sterilized seeds of Arabidopsis were germinated on 0.6% w/v agar-solidified half-strength Murashige and Skoog (MS) medium (Murashige and Skoog, 1962) supplemented with 1% w/v sucrose in an incubator with a photon flux density of 80–100 μmol m−2 sec−1 and a light period of 16 h (22°C) and a dark period of 8 h (18°C). For growth on soil, plants were first grown on MS medium for 10 days, and then transferred to soil and grown under a photosynthetic photon flux density of 150–200 μmol m−2 sec−1 at 22/18°C (day/night) with a 16 h light per 8 h dark period.

Generation of plant expression vectors and plant transformation

The full-length coding regions of PDAT1 (At5g13640) and DGAT1 (At2g19450) were amplified by RT–PCR using primers 5′-GCGTGGTACCATGCCCCTTATTCATCGGA-3′ and 5′-ACGTCTGCAGTCACAGCTTCAGGTCAATACGCTC-3′ for PDAT1, and 5′-ACCTGGAGCTCATGGCGATTTTGGATTCTGC-3′ and 5′-CCGAGGTACCTCATGACATCGATCCTTTTCGGT-3′ for DGAT1. The resulting PDAT1 and DGAT1 PCR products were restriction-digested using KpnI/PstI or SacI/KpnI, respectively, and inserted into the respective sites of a binary vector derived from pPZP212 (Xu et al., 2008). After confirming the integrity of the constructs by sequencing, stable plant transformation was performed as described by Clough and Bent (1998). Transgenic plants were selected in the presence of the appropriate antibiotics for the vector on MS medium.

Construction of double mutants

To generate the double mutants, tgd 1–1 was used as the pollen donor in crosses with dgat1–1 and pdat1–2 mutants. F2 plants derived these crosses were screened for homozygous tgd1–1 mutants by analyzing lipid patterns for the diagnostic presence of TGDG. The tgd1–1 dgat1–1 and tgd1–1 pdat1–2 double mutants were identified from the homozygous tgd1–1 mutants by genotyping the dgat1–1 and pdat1–2 mutations, respectively, using primer sets as described by Zhang et al. (2009).

Treatments with exogenous FFAs and DAG

FFAs were purchased from Sigma-Aldrich (http://www.sigmaaldrich.com) and 18:1/18:1 DAG and 8:0/8:0 DAG were purchased from Avanti Polar Lipids (http://www.avantilipids.com). FFAs were dissolved in ethanol, and DAG was prepared by drying the chloroform solvent under nitrogen gas and then suspending in water by sonication. For treatments, detached developing leaves of 5-week-old plants were floated in water containing 0.0005% Tween–20 and 0.25% ethanol and various concentrations of FFAs or DAG in the light (50 μmol m−2 sec−1) at 22°C with gentle shaking. Controls were treated with water containing 0.0005% Tween–20 and 0.25% ethanol.

In vivo acetate labeling

In vivo labeling experiments with 14C-acetate were performed as described by Bonaventure et al. (2004). Briefly, rapidly growing leaves of 5-week-old plants were cut into strips and then incubated in the light (50 μmol m−2 sec−1) at 22°C with gentle shaking in 10 ml of medium containing 1 mm unlabeled acetate, 20 mm MES pH 5.5, one-tenth strength of MS salts and 0.01% Tween–20. The assay was started by addition of 0.1 mCi of 14C-acetate (106 mCi mmol−1; American Radiolabeled Chemicals, http://www.arc-inc.com). At the end of incubation, leaf strips were washed three times for 3 min at room temperature in water and blotted onto filter paper. For the chase period, leaf tissue was incubated in the same medium lacking 14C-acetate under the same conditions as used for the pulse. Total lipids were extracted and separated as described above, and radioactivity associated with total lipids was determined by liquid scintillation counting.

Lipid and FA analyses

Plant tissues were frozen in liquid nitrogen, and total lipids were extracted by homogenization in chloroform/methanol/formic acid (1:1:0.1 by volume) and 1 m KCl 0.2 m−1 H3PO4. FFA was extracted as described by Welti et al. (2002). Neutral lipids were separated on silica plates (Mallinckrodt Baker, http://www.avantormaterials.com) by thin layer chromatography (TLC) using a solvent system of hexane/diethyl ether/acetic acid (70:30:1 by volume). Polar lipids were separated using a solvent system consisting of acetone/toluene/water (91:30:7 by volume). Lipids were visualized by brief exposure to iodine vapor, and identified by co-chromatography with lipid standards. Individual lipids were scraped from the plate and used to prepare FA methyl esters. To eliminate saturated fatty acid contamination from the silica gel, the TLC plates used for FFA separation were pre-developed using methanol/chloroform (1:1 by volume). Preparation and quantification of the FA methyl esters were performed as described by Fan et al. (2011).

Transmission electron microscopy and light microscopy

For electron microscopy, leaf tissues were fixed with 2.5% v/v glutaraldehyde in 0.1 m sodium phosphate buffer (pH 7.2) for 2 h at room temperature, and then post-fixed with 1% osmium tetroxide in the same buffer. After this double fixation, samples were dehydrated in a graded series of ethanol and propylene oxide, embedded in EPON812 resin (Electron Microscopy Sciences, http://www.electronmicroscopysciences.com), and sectioned. The thin sections were stained with 2% uranyl acetate and lead citrate before viewing under a JEM–1400 LaB6 120KeV transmission electron microscope (JEOL Inc., http://www.jeolusa.com). For light microscopic observation, the sections were stained with 1% toluidine blue and examined using a Leica DM 5500B microscope (http://www.leica-microsystems.com).

Acknowledgements

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information

We thank Prof. John Ohlrogge (Department of Plant Biology, Michigan State University) for providing pdat1-2 mutant seeds. We also thank Dr. John Shanklin for critical reading of the manuscript and Prof. John Ohlrogge for advice on how to eliminate saturated fatty acid contamination from TLC plates. This work was supported by the Division of Chemical Sciences, Geosciences, and Biosciences, Office of Basic Energy Sciences of the U.S. Department of Energy through Grant DEAC0298CH10886 (BO-163) to C. X. Use of the transmission electron microscope and Confocal microscope at the Center of Functional Nanomaterials was supported by the Office of Basic Energy Sciences, U.S. Department of Energy, under Contract DEAC02-98CH10886.

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  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information
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Supporting Information

  1. Top of page
  2. Summary
  3. Introduction
  4. Results
  5. Discussion
  6. Experimental procedures
  7. Acknowledgements
  8. References
  9. Supporting Information
FilenameFormatSizeDescription
tpj12343-sup-0001-FigS1-S7.pdfapplication/PDF4069K

Figure S1. Alterations in FA profiles of TAG in flowers of pdat1–2 and tgd1–1 pdat1–2.

Figure S2. TEM micrographs of leaf cells from the wild-type, tgd1–1 dgat1–1 and tgd1–1 pdat1–2.

Figure S3. Defects in reproductive development in tgd1–1 pdat1–2.

Figure S4. FA composition of FFA and DAG from developing leaves of tgd1–1, tgd1–1 dgat1–1 and tgd1 pdat1–2 plants.

Figure S5. FA composition of major membrane lipids from developing leaves of tgd1–1, tgd1–1 dgat1–1 and tgd1 pdat1–2 plants.

Figure S6. Images of detached tgd1–1, tgd1–1 dgat1–1 and tgd1 pdat1–2 leaves treated with exogenous FAs or DAG.

Figure S7. Distribution of label into TAG and polar lipids after labeling of detached developing leaves with 14C-acetate for 1 h.

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