Enhancing Fatty Acid Synthesis and Diverting Fatty Acids from
Membrane Lipids to Triacylglycerol in Arabidopsis Leaves
Role for Phospholipid:Diacylglycerol Acyltransferase:
Jilian Fan, Chengshi Yan, Xuebin Zhang, and Changcheng Xu1
Biosciences Department, Brookhaven National Laboratory, Upton, New York 11973
There is growing interest in engineering green biomass to expand the production of plant oils as feed and biofuels. Here, we
show that PHOSPHOLIPID:DIACYLGLYCEROL ACYLTRANSFERASE1 (PDAT1) is a critical enzyme involved in triacylglycerol
(TAG) synthesis in leaves. Overexpression of PDAT1 increases leaf TAG accumulation, leading to oil droplet overexpansion
through fusion. Ectopic expression of oleosin promotes the clustering of small oil droplets. Coexpression of PDAT1 with
oleosin boosts leaf TAG content by up to 6.4% of the dry weight without affecting membrane lipid composition and plant
growth. PDAT1 overexpression stimulates fatty acid synthesis (FAS) and increases fatty acid flux toward the prokaryotic
glycerolipid pathway. In the tgd1-1 mutant, which is defective in eukaryotic thylakoid lipid synthesis, the combined
overexpression of PDAT1 with oleosin increases leaf TAG content to 8.6% of the dry weight and total leaf lipid by fourfold. In
the act1 mutant, which is defective in the prokaryotic glycerolipid pathway, PDAT1 overexpression enhances TAG content at
the expense of thylakoid membrane lipids, leading to defects in chloroplast division and thylakoid biogenesis. Collectively,
these results reveal a dual role for PDAT1 in enhancing fatty acid and TAG synthesis in leaves and suggest that increasing FAS
is the key to engineering high levels of TAG accumulation in green biomass.
In plants, fatty acids (FAs), the building blocks for membrane
lipids and storage triacylglycerol (TAG), are almost exclusively
synthesized in the plastid (Ohlrogge and Browse, 1995). They
are incorporated into glycerolipids through two parallel path-
ways, with the prokaryotic pathway confined to plastids and the
eukaryotic pathway involving the endoplasmic reticulum (ER)
and plastids (Roughan and Slack, 1982; Ohlrogge and Browse,
1995). Both pathways begin with the stepwise acylation of
glycerol-3-phosphate (G3P), leading to the generation of phos-
phatidic acid (PA). Dephosphorylation of PA by PA phospho-
hydrolase gives rise to diacylglycerol (DAG), which in the plastid
serves almost exclusively as a precursor for assembly of pho-
tosynthetic membrane lipids, through the prokaryotic pathway.
DAG synthesized de novo in the ER through G3P acylation is
mostly used to synthesize extraplastidic membrane phospholi-
pids, such as phosphatidylcholine (PC) and phosphatidyletha-
nolamine (PE) in growing leaves (Bates et al., 2007) and in
developing seeds (Bates et al., 2009).
PC plays a central role in glycerolipid metabolism. It is the
precursor for the synthesis of glycolipids, the predominant lipid
species found in photosynthetic membranes. This synthesis
occurs through the eukaryotic pathway (Roughan and Slack,
1982; Ohlrogge and Browse, 1995; Somerville and Browse,
1996), although the extent to which the eukaryotic pathway
contributes to thylakoid lipid assembly varies depending on
plant species and tissue within the plant. For example, in pea
(Pisum sativum) plants (Mongrand et al., 1998) and in developing
seeds of Arabidopsis thaliana (Xu et al., 2005), the
pathway of thylakoid lipid synthesis predominates, whereas in
leaves of Arabidopsis and spinach (Spinacia oleracea; Browse
et al., 1986), about half of the photosynthetic membrane lipids
are derived from the eukaryotic pathway and half from the
prokaryotic pathway. Because the substrate specificity of acyl-
transferases in the plastid and ER differs, lipids synthesized via
the prokaryotic pathway are characterized by the almost ex-
clusive presence of a 16-carbon (C16) FA at the sn
the glycerol backbone, whereas those made via the eukaryotic
pathway have a C18 at the same position (Frentzen et al., 1983;
Heinz and Roughan, 1983; Frentzen, 1998). The Arabidopsis
act1 mutantsare defective in the first step of G-3-P acylation in
the plastid (Kunst et al., 1988; Xu et al., 2006). As a conse-
quence, these mutants lack prokaryotic thylakoid glycolipids
and instead synthesize their thylakoid lipids almost exclusively
via the eukaryotic pathway. On the other hand, the tgd1-1 mu-
tant isdeficient in eukaryotic thylakoid lipid synthesis, and most
of the photosynthetic membrane lipids in this mutant are as-
sembled via the prokaryotic pathway (Xu et al., 2003, 2005).
PC is the dominant entry point for acyl groups exported from
the plastid into glycerolipids through acyl editing (Bates et al.,
2007, 2009; Tjellström et al., 2012). It is the major site of the
eukaryotic pathway of FA desaturation (Sperling and Heinz,
1993; Shanklin and Cahoon, 1998), an important precursor for
the synthesis of eukaryotic thylakoid lipids (Ohlrogge and
-2 position of
1Address correspondence to firstname.lastname@example.org.
Theauthor 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: Changcheng Xu (cxu@bnl.
CSome figures in this article are displayed in color online but in black and
white in the print edition.
WOnline version contains Web-only data.
The Plant Cell, Vol. 25: 1–14, September 2013, www.plantcell.org ã 2013 American Society of Plant Biologists. All rights reserved.
Browse, 1995), and a potential carrier of newly synthesized FAs
from plastid envelope membranes to the ER (Tjellström et al.,
2012). In developing seeds, PC is the predominant source of DAG
for TAG synthesis via the acyl-CoA–dependent and –independent
pathway catalyzed by diacylglycerol:acyl-CoA acyltransferase
(DGAT) and phospholipid:diacylglycerol acyltransferase (PDAT),
respectively (Chapman and Ohlrogge, 2012). TAG is packaged in
a dynamic subcellular organelle termed oil droplets (ODs) or oil
bodies, which are composed of a central matrix of TAG sur-
rounded by a monolayer of phospholipids with a subset of spe-
cific embedded proteins (Huang, 1996). The most abundant
proteins coating seed ODs are oleosins, and genetic studies in
Arabidopsis mutants have suggested a key role for oleosins in
preventing ODs from coalescing, thereby maintaining ODs as
small discrete entities to facilitate TAG mobilization and confer
freezing tolerance during postgerminative seedling growth (Siloto
et al., 2006; Shimada et al., 2008).
In the model plant Arabidopsis, DGAT1 and PDAT1 play over-
lapping roles in TAG synthesis in developing seeds and pollen
(Zhang et al., 2009). Both DGAT1 and PDAT1 genes are ex-
pressed in leaves, roots, and stems, in addition to developing
seeds and flowers (Lu et al., 2003; Ståhl et al., 2004). DGAT1 has
been implicated in TAG biosynthesis in senescent leaves (Kaup
et al., 2002; Slocombe et al., 2009), but the functional role of
PDAT in leaf tissue remains largely unknown, although knockout
of PDAT1 has recently been shown to have a small negative effect
on TAG accumulation in roots of sdp1 mutants defective in
SUGAR-DEPENDENT1 TAG lipase (Kelly et al., 2013). Here, we
provide evidence that PDAT1 plays a critical role in TAG synthesis
in Arabidopsis leaves. Overexpression of PDAT1 enhances both
FA and TAG synthesis in leaves. The possible functional role of
PDAT1 in membrane lipid turnover is also discussed.
The Relative Contribution of PDAT1 and DGAT1 to TAG
Synthesis in Leaves
The Arabidopsis dgat1-1 and pdat1-2 mutants harbor an ethyl
methanesulfonate–induced lesion in the DGAT1 locus (Zou
et al., 1999) and a T-DNA insertion in the PDAT1 gene (Zhang
et al., 2009), respectively. As the first step toward understanding
the role of PDAT1 and DGAT1 in TAG synthesis in vegetative
tissues, we quantified TAG levels in developing and senescing
leaves of wild-type, dgat1-1, and pdat1-2 mutant plants. On
a dry weight (DW) basis, the TAG levels measured as fatty acid
methyl esters (FAMEs) were 0.04 and 0.15% in developing and
senescing leaves of wild-type plants, respectively (Figure 1).
Compared with the wild type, the pdat1-2 mutant displayed
a 57% decrease in TAG content in developing leaves and 39%
in senescing leaves. By contrast, the dgat1-1 mutant showed
a less pronounced decrease in TAG level in developing leaves
(31%) but a more severe drop in TAG content in senescing
leaves (63%). These results suggest that PDAT1 has a more
important role in TAG synthesis in growing leaves than DGAT1,
whereas the opposite is true in senescing leaves. Consistent
with this notion, analyzing the database of lipid-related gene
expression during leaf senescence (Troncoso-Ponce et al.,
2013) revealed that PDAT1 is expressed at much higher levels
than DGAT1 during early leaf growth and development (see
Supplemental Figure 1 online). In addition, the PDAT1 transcript
level decreases with age in all three microarray studies compiled
in the database, whereas DGAT1 expression tends to increase
as the leaf senesces. The involvement of DGAT1 in TAG syn-
thesis during leaf senescence is consistent with previous reports
(Kaup et al., 2002; Slocombe et al., 2009).
Overexpression of PDAT1 Enhances TAG Accumulation,
Leading to OD Overexpansion in Leaves
A previous study showed that overexpressing PDAT1 driven by
the constitutive 35S cauliflower mosaic virus promoter did not
affect TAG levels or membrane lipid content and FA composition
in the young seedlings of three independent transgenic Arabi-
dopsis lines (Ståhl et al., 2004). We extended this study by ex-
amining TAG content in the leaves of a large number of adult
transgenic plants grown in soil. On average, there was a sev-
enfold increase in TAG content in 22 randomly chosen primary
Supplemental Figure 2A online). By comparison, overexpressing
DGAT1 driven by the same promoter resulted in only a marginal,
if any, increase in the average leaf TAG levels in 21 randomly
selected primary transformants tested. These results are in ac-
cordance with the absence of significant increases in total leaf
with thewildtype (see
Figure 1. Roles of DGAT1 and PDAT1 in TAG Synthesis during Leaf
Lipids were extracted from developing leaves (DL; the thirteenth leaf of 5-
week-old plants) and senescing leaves (SL; the fifth leaf of 7-week-old
plants) of the wild type (WT), dgat1-1, and pdat1-2. Values are means
and SD of three to five biological replicates.
[See online article for color version of this figure.]
2The Plant Cell
FA content in Arabidopsis transgenic plants constitutively
overexpressing DGAT1 as reported recently by others (Wi-
nichayakul et al., 2013). Together, these results indicate that
PDAT1 exerts a higher degree of regulation than DGAT1 over
TAG synthesis in Arabidopsis leaves.
We subsequently obtained three homozygous transgenic
lines overexpressing PDAT1 for detailed characterization. On
a DW basis, the TAG level in mature rosette leaves of 7-week-
old PDAT1 overexpressors grown in soil was 2.6%, a 28-fold
increase compared with the wild type (Figure 2A). TAG com-
positional analysis showed that the predominant acyl chains in
TAG derived from the leaves of PDAT1 overexpressors were
polyunsaturated FAs with 18:2 and 18:3 accounting for over
75% of the total acyl chains. By comparison, the FAs recovered
from wild-type plants by analysis of the TAG band from thin layer
chromatography (TLC) plates were largely saturated acyl chains
(Figure 2B). In contrast with TAGs isolated from Arabidopsis
seedlings ectopically overproducing a Chlamydomonas rein-
hardtii DGAT type-two enzyme (Sanjaya et al., 2013) or tran-
scription factors involved in storage product accumulation, such
as LEAF COTYLEDON2 (LEC2) (Santos Mendoza et al., 2005),
TAGs derived from leaves of PDAT1 overexpressors contained
very limited amounts of very-long-chain fatty acids (Figure 2B).
Consistent with the marked increase in TAG content, mi-
croscopy examination following Nile Red staining showed that,
whereas only a few or no OD-like structures were seen in the
leaves of wild-type plants under our growth conditions (Figures
3A and 3C), overexpression of PDAT1 led to OD accumulation in
leaves (Figure 3B). Those ODs were often irregular in shape and
frequently appeared to coalesce. Ultrastructural examination
using transmission electron microscopy (TEM) confirmed that
overexpression of PDAT1 led to the formation of large, irregu-
larly shaped ODs in leaves (Figure 3D).
Ectopic Expression of Oleosin Promotes the Clustering of
Small ODs and Boosts Oil Accumulation in PDAT1
Oleosins are known to play a key role in preventing ODs from
coalescing in oilseeds (Siloto et al., 2006; Shimada et al., 2008).
In addition, in yeast and mammalian model systems, the ectopic
expression of OD-associated proteins often leads to increased
TAG storage (Brasaemle et al., 2000; Froissard et al., 2009). To
test the functional role of oleosins in TAG accumulation in
vegetative tissues, OELOSIN1 (OLE1), the most abundant seed
OD-specific protein of Arabidopsis (Huang, 1996; Shimada et al.,
2008), was C-terminally tagged with green fluorescent protein
(GFP), and this fusion gene was expressed in Arabidopsis wild-
type plants under the control of the 35S cauliflower mosaic virus
promoter. Three independent transgenic lines exhibiting high
levels of green fluorescence signal in young seedlings were
obtained. Analyzing leaf lipid extracts from 3-week-old trans-
genic plants revealed an up to sevenfold increase in TAG con-
tent compared with the wild type (see Supplemental Figure 2B
online). TAGs isolated from OLE1 overexpressors were mainly
composed of 16:0, 18:2, and 18:3 FAs, whereas very-long-chain
fatty acids, which typically accumulate in Arabidopsis seeds,
were present at very low levels (see Supplemental Figure 2C
online), suggesting that TAG accumulation in 3-week-old
Figure 2. TAG Accumulation in Leaves of Transgenic Plants Over-
(A) Leaf TAG content in 7-week-old wild type (WT) and three independent
(B) FA composition of TAGs in the wild type and two independent
transgenic lines. Values are means and SD of three biological replicates.
Figure 3. OD Accumulation in Leaves of Transgenic Plants Over-
(A) and (B) Images of ODs in leaves of 7-week-old wild type (A) and
PDAT1-overexpressing line 16 (B) stained with Nile red. Bars = 5 µm.
(C) and (D) TEM images of leaf cells from 7-week-old wild type (C) and
PDAT1-overexpressing line 16 (D). Bars = 1 µm.
Role of PDAT1 Acyltransferase in Leaves3
seedlings is not a result of reduced hydrolysis of seed oil due to
the ectopic overexpression of OLE1.
TEM analysis showed that ectopic expression of OLE1 in-
duced the formation of clusters of small spherical ODs (Figure
4A), unlike the large aberrant ODs observed in transgenic plants
overexpressing PDAT1 (Figure 3). Using confocal microscopy,
we found that the OLE1-GFP fusion protein was exclusively
associated with OD clusters and appeared as a ring-like pattern
surrounding the perimeters of ODs (Figure 4B).
We next transformed the homozygous OLE1 overexpressor
line 1 with the PDAT1 construct. Three independent PDAT1/
OLE1 double transgenic lines were propagated to the T3 gen-
eration and used for detailed analysis. Using Fat Red 7B,
a nonfluorescent dye that specifically stains neutral lipids
(Brundrett et al., 1991), we found that the double transgenic
seedlings were stained red in both roots and leaves (Figure 5A),
whereas the wild-type seedlings did not stain. On soil, the
PDAT1/OLE1 double transgenic lines displayed no obvious
growth or developmental abnormalities (see Supplemental
Figure 3A online). Quantification of TAG levels in the rosette
leaves of 7-week-old soil-grown double transgenic plants
showed an up to 74-fold increase compared with the wild type.
On a DW basis, the TAG content in wild-type leaves was 0.05%,
while in the three PDAT1/OLE1 double transgenic lines, it was
between 5.7 and 6.4% (Figure 5B). There was also an up to
twofold increase in the total leaf FA content in 7-week-old
transgenic plants coexpressing PDAT1 and OLE1 compared
with the wild type (see Supplemental Figure 3B online). In ad-
dition, TAG accumulated up to 1.5% per DW in the stems of the
double transgenic plants, a 30-fold increase compared with the
Figure 4. Ectopic Expression of OLE1 Promotes the Clustering of Small ODs.
(A) TEM imaging of ODs in leaves of the OLE1-GFP–overexpressing line
1. Bar = 1 µm.
(B) Confocal microscopy of OD clusters in leaves of the OLE1-GFP–
overexpressing line 1. Bar = 2 µm.
Figure 5. TAG Accumulation in Transgenic Lines Coexpressing PDAT1
(A) Fat Red 7B staining of 3-week-old seedlings of the wild type (WT),
PDAT1, and OLE1-GFP double transgenic line 1 (PDAT1/OLE1 #1).
(B) TAG content in leaves of 7-week-old plants of the wild type and three
transgenic lines coexpressing PDAT1 and OLE1-GFP (PDAT1/OLE1).
Values are means and SD of three biological replicates.
(C) Images of ODs in leaves of 7-week-old PDAT1-overexpressing line 3
(PDAT1 #3) stained with Nile red. Bar = 5 µm.
(D) Confocal imaging of ODs in leaves of 7-week-old soil-grown trans-
genic line 1 coexpressing PDAT1 and OLE1-GFP (PDAT1/OLE1 #1).
Bar = 5 µm.
4 The Plant Cell
wild type (see Supplemental Figure 3C online). Microscopy
analysis showed that, unlike large and often irregularly shaped
ODs in plants carrying the PDAT1 transgene only (Figure 5C),
the double transgenic plants contained clusters of spherical
ODs in leaves (Figure 5D), similar to those seen in plants ec-
topically expressing OLE1 alone (Figure 4), thus confirming the
role of OLE1 in promoting the clustering of ODs.
The FA Composition of Membrane Lipids Is Altered in
PDAT1 Transgenic Plants
To investigate the potential impact of PDAT1 overexpression on
membrane lipids, we first surveyed the FA profiles of total polar
lipids in leaves of three independent transgenic lines over-
expressing PDAT1. The relative proportions of 16:3, 18:1, and
18:2 increased, with a concomitant decrease in the proportion of
18:3 (see Supplemental Figure 4A online). We next separated
individual polar lipids by TLC. Analysis by gas chromatography
of the FAMEs revealed no discernible differences in levels of the
individual classes of major polar lipids between PDAT1 over-
expressor 16 and the wild type (see Supplemental Figure 4B
online). However, there was a substantial decrease in 18:3 with
a corresponding increase in 18:1 and 18:2 in both extraplastidic
(PE and PC) and thylakoid (digalactosyldiacylglycerol [DGDG]
Supplemental Figures 4C to 4F online). The marked decrease in
18:3 in PC and PE may reflect the substrate preference of
PDAT1 as demonstrated by in vitro studies by Ståhl et al. (2004).
This is supported by the particular enrichment of 18:3 in TAG
accumulated in leaves of PDAT1 transgenic plants (Figure 2B).
On the other hand, the decrease in the proportion of 18:3 in
galactolipids, particularly in DGDG, may reflect the fact that PC
is the precursor of galactolipids synthesized by the eukaryotic
pathway. In addition to changes in C18 FAs, the levels of C16
acyl chains tended to decrease in PC but increase in DGDG and
MGDG when compared with the wild type (see Supplemental
Figures 4C to 4F online).
Overexpression of PDAT1 Alters the Positional Distribution
of FAs in Galactolipids
The increase in C16 FA content in galactolipids may suggest
that the prokaryotic pathway of thylakoid lipid synthesis is en-
hanced in PDAT1 overexpressors. To test this possibility, the
positional distribution of FAs in galactolipids was analyzed fol-
lowing Rhizopus lipase digestion. As shown in Figure 6, the
relative proportion of C16 FAs at the sn-2 position of MGDG
increased from 80% in the wild type to 94% in PDAT1 trans-
genic line 16. In the case of DGDG, the increase was from 34 to
57%. The finding that sn-2 C16 FA content increased to
a greater degree in DGDG than in MGDG reflects the fact that
the prokaryotic pathway makes a much smaller contribution to
the synthesis of DGDG than to MGDG in the leaves of wild-type
plants (Browse et al., 1986); consequently, DGDG is more af-
fected than MGDG when prokaryotic thylakoid lipid synthesis is
Rate of FA Synthesis Is Enhanced in PDAT1
The accumulation of large amounts of TAG without a concomi-
tant decrease in membrane lipid content in PDAT1 transgenic
plants suggests an increase in fatty acid synthesis (FAS). To test
this possibility, we conducted14C-acetate feeding experiments
with leaf strips from expanding leaves for a period of 1 h, which
was within the linear range of label incorporation (see
Supplemental Figure 5 online). Because acetate incorporation
into FAs can be influenced by levels of endogenous metabolites
(Cronan et al., 1975; Nunn et al., 1977), the labeling experiments
were performed in the presence of 1 mM unlabeled acetate in an
attempt to eliminate potential variations in endogenous sub-
strate pools between the transgenics and the wild type. We
detected 72 and 44% increases in radiolabeled lipids in the
PDAT1 overexpressor 9 and 16, respectively, compared with the
wild type (Figure 7A), suggesting an increase in the rate of FAS
due to PDAT1 overexpression. Analysis of label distribution
showed that overexpression of PDAT1 significantly increased
the label incorporation into galactolipids, phosphatidylglycerol
(PG), and TAG, whereas the proportions of label in PC, PE, and
sulfoquinovosyldiacylglycerol (SQDG)/phosphatidylinositol (PI)
decreased (Figure 7B). These results are consistent with an in-
crease in acyl fluxes toward galactolipids and PDAT1-mediated
TAG synthesis in response to PDAT1 overproduction.
Rate of FAS Strongly Correlates with Leaf TAG Content in
The leaf TAG levels in double transgenic plants coexpressing
PDAT1 and OLE1 (Figure 5B) were much higher than in
Figure 6. FA Composition Exclusively at the sn-2 Position of the Glyc-
erol Backbone of Galactolipids Isolated from Leaves of 7-Week-Old
Wild-Type and PDAT1-Overexpressing Line 16 (PDAT1 #16).
Values are means and SD of three biological replicates. WT, the wild type.
Role of PDAT1 Acyltransferase in Leaves5
transgenic plants carrying PDAT1 alone (Figure 2A). To begin to
dissect the biochemical basis for the increased TAG accumu-
lation in double transgenic lines, growing leaves of PDAT1 single
and PDAT1/OLE1 double transgenic plants were labeled with
14C-acetate. Compared with the transgenic plants over-
expressing PDAT1 alone (Figure 7A), the initial rate of acetate
incorporation into total lipids was much higher for the PDAT1/
OLE1 double transgenic lines (Figure 8A). In addition, in five
additional independent PDAT1/OLE1 double transgenic lines
tested, there was a strong correlation between the rate of FAS
(Figure 8B) and TAG content (Figure 8C). These results suggest
that increased TAG accumulation in leaves of PDAT1/OLE1
double transgenic lines may be due to enhanced FAS. On the
other hand, no apparent difference in acetate incorporation into
total lipids was found between the wild type and transgenic
plants overexpressing the OLE1 single transgene (Figure 8A),
suggesting that the increased TAG accumulation in transgenic
plants carrying OLE1 may be attributed to decreased TAG
turnover, likely because of the shielding of ODs by OLE1 from
the access of cytosolic lipases, in a manner similar to that ob-
served for mammalian OD-associated protein perilipins (Bra-
saemle et al., 2000).
Disruption of the Eukaryotic Thylakoid Lipid Pathway
Enhances TAG Accumulation in Double Transgenic Plants
Coexpressing PDAT1 and OLE1
In Arabidopsis, the prokaryotic and eukaryotic glycerolipid
pathway each consumes about half of acyl chains synthesized
de novo in the plastid (Browse et al., 1986). Disruption of either
glycerolipid pathway may therefore be expected to increase acyl
chains for TAG synthesis. To test this possibility, the tgd1-1
mutant, which is defective in the eukaryotic pathway of thylakoid
lipid synthesis (Xu et al., 2003), was crossed with the PDAT1/
OLE1 double transgenic line 1. Analysis of leaf lipid extracts
from mature leaves of 7-week-old soil-grown plants showed
that, on a DW basis, TAG content in the PDAT1/OLE1 double
transgenic line in the tgd1-1 background was increased to 8.6%
and total lipid content to 16.2%, a 97-fold and fourfold increase,
respectively, compared with tgd1-1 (Figure 9A). As expected,
coexpression of PDAT1 with OLE1 in tgd1-1 induced the
Figure 7. Overexpression of PDAT1 Enhances the Rate of FAS in
(A) Initial rates of FAS in growing leaves of 7-week-old wild-type (WT),
PDAT1-overexpressing line 3 (PDAT1 #3), and PDAT1-overexpressing
line 16 (PDAT1 #16) measured by14C-acetate labeling.
(B) The distribution of label into TAG and polar lipids after labeling of
detached leaves with14C-acetate for 60 min.
Values in (A) and (B) are means
indicate statistically significant differences from the wild type based on
Student’s t test (P < 0.05).
SD of three biological replicates. Asterisks
Figure 8. TAG Accumulation Positively Correlates with the Rate of FAS.
Initial rates of FAS in growing leaves of 5-week-old wild-type (WT),
OLE1-GFP–overexpressing line 1 (OLE1 #1), and PDAT1 and OLE1-GFP
double transgenic line 1 (PDAT1/OLE1 #1) and 3 (PDAT1/OLE1 #3)
measured by14C-acetate labeling is shown in (A). Initial rates of FAS in
growing leaves of 5-week-old OLE1 #1 and five double transgenic lines
coexpressing PDAT1 and OLE1-GFP (PDAT1/OLE1) measured by14C-
acetate labeling is presented in (B). TAG content in leaves of 7-week-old
OLE1 #1 and five double transgenic lines coexpressing PDAT1 and
OLE1-GFP (PDAT1/OLE1) is shown in (C). Values are means and SD of
three biological replicates.
6The Plant Cell
formation of OD clusters in leaves (Figure 9B). In addition to
leaves, TAG also accumulated in stems of tgd1-1/PDAT1/OLE1
transgenic plants to 3.0% 6 0.18% per DW (n = 3, 6SD), a 60-
fold increase relative to tgd1-1 (0.05% 6 0.01% per DW, n = 3,
6SD). Remarkably, despite a large increase in TAG and total lipid
levels, the growth and development of the double transgenic
plants were only slightly affected (see Supplemental Figure 6A
online). Furthermore, no apparent difference in total polar lipid
content was found between tgd1-1/PDAT1/OLE1 and tgd1-1
(see Supplemental Figure 6B online).
Disruption of PDAT1 Partially Alleviates the Phenotype of
the act1 Mutant
We next constructed a double mutant between pdat1-2 and
act1 defective in the prokaryotic thylakoid lipid pathway. When
grown in soil, the act1 pdat1-2 double mutant seedlings were
visibly darker in color compared with the act1 mutant (Figure
10A). Consistent with the dark-green phenotype, the total
chlorophyll content was increased by 9.8%, from 0.90 6 0.02 in
act1 to 0.99 6 0.04 mg/g fresh weight (FW) (n = 3, 6SD) in the
double mutant. Lipid analysis showed that disruption of PDAT1
in act1 resulted in small but significant increases in the levels of
thylakoid lipids MGDG and PG (Figure 10B). In addition, the
combined amount of SQDG and PI was slightly increased,
whereas the level of PC, PE, and DGDG stayed largely unaltered
in the act1 pdat1-2 double mutant relative to act1. The FA
composition of all polar lipid classes examined did not show
noticeable deviations from those in act1, with the exception of
PC, which exhibited a significant increase in relative proportions
of 18:2 and 18:3 with a corresponding decrease in 18:1 (Figure
10C). The mutation in act1 has been shown to result in de-
creases in the relative levels of chloroplast lipids PG, SQDG, and
MGDG and increases in the relative amounts of 18:1 in ex-
trachloroplast lipids, particularly in PC (Kunst et al., 1988), and
a slight reduction in the amount of chlorophyll on a FW basis
(Kunst et al., 1989). Thus, our lipid data, together with the
Figure 9. Disruption of the Eukaryotic Thylakoid Lipid Pathway Enhan-
ces TAG Accumulation in Transgenic Plants Coexpressing PDAT1 and
(A) TAG and total lipid levels in leaves of 7-week-old tgd1-1 and trans-
genic plants coexpressing PDAT1 and OLE1-GFP in the tgd1-1 back-
ground (tgd1-1/PDAT1/OLE1). Values are means and
biological replicates. Asterisks indicate statistically significant differ-
ences from tgd1-1 based on Student’s t test (P < 0.05).
(B) TEM imaging of ODs in leaves of 7-week-old tgd1-1/PDAT1/OLE1
plants. Bar = 2 µm.
SD of three
Figure 10. Disruption of PDAT1 Partially Complements the Phenotype of
the act1 Mutant.
Images of 4-week-old act1 and act1 pdat1-2 plants are shown in (A).
Polar lipid content in leaves of act1 and act1 pdat1-2 plants is shown in
(B). FA composition of PC isolated from leaves of 4-week-old wild-type
(WT) act1 and act1 pdat1-2 plants grown on agar plates is shown in (C).
Values are means and SD of three biological replicates. Asterisks indicate
a statistically significant difference from act1 based on Student’s t test
(P < 0.05).
[See online article for color version of this figure.]
Role of PDAT1 Acyltransferase in Leaves7
increased chlorophyll content in act1 pdat1-2 plants, suggest
that disruption of PDAT1 partially alleviates the phenotypes of
the act1 mutant, presumably as a consequence of the increased
diversion of FAS from TAG to eukaryotic thylakoid lipids in the
Overexpression of PDAT1 in act1 Enhances TAG
Accumulation at the Expense of Thylakoid Lipids
Next, the PDAT1 construct was introduced into act1 by Agro-
bacterium tumefaciens–mediated transformation. When grown
in soil, transgenic plants overexpressing PDAT1 in act1 (act1/
PDAT1) were frequently found to be pale in color and stunted in
growth (Figure 11A). Consistent with the pale phenotype, the
amount of the major thylakoid lipids MGDG, DGDG, and PG was
reduced by 25, 46, and 31%, respectively, in act1/PDAT1 line 7
compared with act1, whereas the levels of PC, PE, and SQDG/PI
remained largely unchanged (Figure 11B). On a FW basis, the
sum of MGDG, DGDG, and PG decreased from 2826 in act1 to
2065 µg/g FW in act1/PDAT1 line 7, resulting in a net loss of 761
µg/g FW, while the amount of TAG increased from 57.3 6 9.8 to
760.9 6 164.6 µg/g FW (n = 3, 6SD), a net gain of 704 µg/g FW.
These results suggest that TAG accumulation in act1 PDAT1
overexpressing lines is largely accounted for by the increased
diversion of acyl groups from thylakoid membrane lipid syn-
thesis to TAG. Consistent with this, the total leaf FA content was
comparable in act1/PDAT1 line 7 (5.61% 6 0.61% per DW, n =
3, 6SD) and act1 (5.91% 6 0.84% per DW, n = 4, 6SD).
To further investigate the nature of the pale-green phenotype
in act1 overexpressing PDAT1, microscopy examination of leaf
cross sections was performed. Compared with act1, the trans-
genic act1 plants displayed increased leaf thickness and in-
creased cell size, but decreased cell number (see Supplemental
Figure 7 online). The chloroplasts of the transgenic line ap-
peared to be larger. The number of chloroplasts per cell cross
section was reduced from 6.63 6 0.05 in act1 to 3.33 6 0.05 (n =
4) in the act1/PDAT1 line 7. At the ultrastructural level, the
amount of thylakoid membranes and thylakoid membrane
stacking per chloroplast in mature leaves of act1 plants over-
expressing PDAT1 were greatly reduced when compared with
chloroplasts from act1 leaves at the same developmental stage
(Figures 11C and 11D).
To gain more information on the metabolic changes leading to
TAG accumulation due to PDAT1 overproduction in act1, we
performed14C-acetate pulse-chase labeling experiments using
detached leaves. During the 1-h pulse, the rates of14C-acetate
incorporation were approximately threefold higher in the act1/
PDAT1 line 7 (232.0 6 18.9 DPM/mg FW/min, n
compared with act1 (636.6 6 127.7 DPM/mg FW/min, n = 3,
6SD), suggesting an increase in FAS due to PDAT1 over-
expression in the act1 background. Because the total leaf lipid
content remained largely unchanged in act1/PDAT1 line 7 rela-
tive to act1, the rate of FA degradation must be increased
concurrently such that a constant lipid level is maintained. In-
deed, the average label decay rate was increased by threefold in
act1/PDAT1 line 7 (22% per day) compared with act1 (7% per
day) (see Supplemental Figure 8A online). Analysis of label dis-
tribution showed that PC was the most radioactive lipid imme-
diately following the pulse, followed by PG and PE. Galactolipids
MGDG and DGDG, despite accounting for over 70% of the total
membrane lipid mass, contained <10% of total initial label in
act1 (Figure 12). Overexpression of PDAT1 led to a 19% drop in
initial label in PC, no change in PE, but a 25% decline in SQDG/
PI, 42% in DGDG, 35% in PG, and 44% in MGDG, whereas TAG
label increased from 3% in act1 to 25% in the act1/PDAT1 line
7, thus representing a nearly fourfold increase in the initial label
partitioning into TAG (Figure 12). During the chase, the PC label
declined, whereas labeled MGDG and DGDG increased in both
act1 and act1 transgenic plants, reflecting a well-documented
precursor-product relationship between PC and galactolipids,
but the extent of increases in label in MGDG and DGDG was
smaller in act1 transgenic plants (see Supplemental Figure 8C
online) than in act1 (see Supplemental Figure 8B online). In both
act1 and the act1 PDAT1 overexpressor, the label in TAG slightly
increased during the initial 24 h of chase. This was followed by
substantial decreases in labeled TAG during the remainder of
the chase, reflecting turnover of TAG in both act1 and act1
= 3, 6SD)
Figure 11. Overexpression of PDAT1 in act1 Promotes TAG Accumu-
lation at the Expense of Thylakoid Lipids.
(A) Images of 6-week-old wild type (WT), act1, and three PDAT1-over-
expressing lines in the act1 background (act1/PDAT1).
(B) Polar lipid levels in leaves of act1 and the act1/PDAT1 #7. Values are
means and SD of three to four biological replicates. Asterisks indicate
statistically significant differences from act1 based on Student’s t test
(P < 0.05).
(C) and (D) TEM analysis of chloroplasts in leaves of act1 (C) and the
act1/PDAT1 #7 (D). Bars = 1 µm.
[See online article for color version of this figure.]
8 The Plant Cell
PDAT Is Functionally Conserved among Yeast, Microalgae,
In plants as well as in microalgae, mammals, and fungi, the
terminal and committed step in the pathway of TAG bio-
synthesis is catalyzed by multiple DAG acyltransferases en-
coded by distinct gene families. Although the basic enzymes
involved in the DAG esterification reaction have been extensively
characterized at the biochemical and molecular level in several
model systems, fundamental questions remain regarding the
roles of the individual DAG acyltransferases and the functional
significance of highly redundant activities. At least three classes
of DAG acyltransferases consisting of acyl-CoA–dependent
DGAT1 and DGAT2 and acyl-CoA–independent PDAT, contrib-
ute to TAG synthesis in yeasts (Kohlwein, 2010), microalgae (Liu
and Benning, 2012 ), and plants (Chapman and Ohlrogge, 2012).
Studies in both yeast (Oelkers et al., 2000, 2002) and the green
alga C. reinhardtii (Yoon et al., 2012) have indicated that PDAT
plays a major role in TAG synthesis during phases of active cell
growth and division, while DGATs appear to be more important
in cells entering the stationary phase of growth (Oelkers et al.,
2002; Sandager et al., 2002). In Arabidopsis, most evidence
suggests that DGAT1 is a major contributor to TAG synthesis in
developing seeds (Katavic et al., 1995; Routaboul et al., 1999;
Jako et al., 2001) and in leaves during senescence (Kaup et al.,
2002; Slocombe et al., 2009). On the other hand, although
PDAT1 is known to have an overlapping role in TAG synthesis in
seeds and pollen (Zhang et al., 2009), neither disruption (Ståhl
et al., 2004; Mhaske et al., 2005) nor overexpression of PDAT1
(Ståhl et al., 2004) appears to affect lipid content and FA com-
position in seeds and in young seedlings. Through extensive
analyses of PDAT1 knockout mutants and overexpressing lines,
particularly in the act1 mutant background, this work provides
insight into the biological functions of PDAT1. We show that
PDAT1 plays an important role in TAG synthesis in leaves;
overexpression of PDAT1, but not DGAT1, increased leaf TAG
content, whereas disruption of PDAT1 substantially decreased
TAG levels with the most pronounced decrease occurring in
rapidly growing leaves and the least in senescing leaves. Col-
lectively, these results point to evolutionary conservation of
PDAT functions associated with rapid cell growth and mem-
brane proliferation in yeast, microalgae, and plants.
According to current knowledge about glycerolipid metabo-
lism in plants, most of the acyl chains exported outside the
plastid are first incorporated into PC, the substrate of PDAT,
through acyl editing in growing leaf cells (Bates et al., 2007,
2009; Tjellström et al., 2012). Because lipid metabolism in
growing leaf cells is directed primarily toward membrane lipid
synthesis, the acyl chains released as a result of acyl editing of
PC are primarily channeled into G3P acylation reactions to
generate DAG for de novo PC synthesis (Bates et al., 2007) and
therefore may be limiting for DGAT1-mediated TAG synthesis.
As leaf tissues age, the rate of FAS declines (Bao et al., 2000;
Hellgren and Sandelius, 2001) and fewer nascent FAs are fluxed
through PC and more are used directly in the chloroplast to
sustain thylakoid lipid turnover (Hellgren and Sandelius, 2001).
This may explain why the PDAT1-mediated acyl-CoA–in-
dependent route is more involved in TAG synthesis in growing
but not in senescing leaves of Arabidopsis than the DGAT1-
catalyzed acyl-CoA–dependent reaction.
PDAT Plays a Role in Membrane Lipid Turnover
In vitro biochemical assays showed that Arabidopsis PDAT1
displays the highest activity toward oxygenated acyl groups,
such as hydroxyl acyl chains (Ståhl et al., 2004), much as does
the enzyme from castor (Ricinus communis) oil seeds that ac-
cumulates high levels of hydroxyl FAs (Dahlqvist et al., 2000). It
has thus been hypothesized that Arabidopsis PDAT1, like PDAT
in castor, may play a role in cell membrane repair by removing
oxygenated FAs from membrane phospholipids (Ståhl et al.,
2004). However, direct evidence in support of this hypothesis is
Recent studies in other model organisms have indicated that
membrane lipid and TAG metabolism are tightly linked pro-
cesses (Rajakumari et al., 2010; Horvath et al., 2011; Yoon et al.,
2012). In yeast acyl-CoA–deficient mutants, disruption of PDAT
blocked phospholipid deacylation, resulting in an increase in
levels of PC and PE (Mora et al., 2012). Likewise, knockdown of
PDAT in C. reinhardtii has been shown to alter the molecular
species composition of major thylakoid membrane lipids (Yoon
et al., 2012). In this study, we showed that disruption of PDAT1
in the act1 mutant increased the proportions of polyunsaturated
FAs in PC with a corresponding decrease in 18:1, whereas the
reverse trend was found for PDAT1-overexpressing plants. Be-
cause PC is the site of FA desaturation, the opposite effects of
PDAT1 knockout and overproduction on levels of FA unsatura-
tion are in line with PDAT1’s function in the removal of acyl
groups from PC. In addition to changes in the FA composition of
PC, knockout of PDAT1 increased, whereas overexpression of
this gene decreased levels of thylakoid lipid content in the act1
mutant. These results provide direct in vivo evidence that Ara-
bidopsis PDAT1, like its homologs in yeast and C. reinhardtii, is
also involved in the turnover of membrane lipids in leaves.
In the pdat1-2 single mutant, no appreciable change in
membrane lipid content and FA composition was observed,
despite the marked reduction in TAG content compared with the
wild type. The lack of detectable phenotype in pdat1-2 may
Figure 12. The Distribution of Radioactivity in Polar Lipids and TAG after
Labeling of Detached Leaves with14C-Acetate for 60 min in act1 and
act1 PDAT1-Overexpressing Line #7 (act1/PDAT1 #7).
Values are means and SD of three biological replicates.
Role of PDAT1 Acyltransferase in Leaves9
reflect the operation of lipid homeostatic mechanisms that fully
compensate for the changes in membrane compositions re-
sulting from the loss of PDAT1 function in the wild type. In this
context, in PDAT1-overexpressing lines in the wild-type back-
ground, the increased diversion of acyl groups from membrane
lipid synthesis to TAG was entirely compensated for by en-
hanced FAS and increased thylakoid lipid assembly via the
prokaryotic pathway, thus resulting in little net change in the
membrane lipid content and composition. Although the factors
responsible for such metabolic adjustments are unknown, it is
tempting to speculate that biochemical feedback regulation of
FAS plays a key role. In this scenario, PDAT1-mediated TAG
accumulation consumes, in the cytosol, acyl moieties destined
for eukaryotic thylakoid lipid synthesis. This could (1) provide
a driving force for FA export outside the chloroplast through
vectorial acylation and (2) increase the demand for FAs needed
for the prokaryotic thylakoid lipid synthesis. Both of these
metabolic responses could lead to a depletion of 18:1-acyl
carrier protein in the plastid, thereby releasing its feedback in-
hibition on the plastidic acetyl-CoA carboxylase (Andre et al.,
2012). This could in turn lead to an increase in the rate of FAS
and an increase in flow of FAs toward the prokaryotic pathway
to compensate for the loss of lipid precursors normally shuttled
from the ER to the chloroplast for eukaryotic thylakoid lipid
In contrast with PDAT1 overexpressors in the wild-type
background, overexpression of PDAT1 in act1 resulted in sub-
stantial reductions in levels of thylakoid lipids, leading to defects
in chloroplast biogenesis, despite marked increases in the rates
of FAS. These results indicate that the prokaryotic pathway
plays a critical role in maintaining membrane lipid homeostasis
when the eukaryotic pathway is severely compromised. In this
regard, we showed previously that the genetic defects in the
eukaryotic pathway of thylakoid lipid synthesis in tgd1 mutants
can be almost fully compensated for by the augmented pro-
karyotic pathway without major growth and developmental
consequences (Xu et al., 2005).
Biotechnological Implications for PDAT
FAs are the predominant component of TAG. Therefore, in-
creasing FAS is a prerequisite for attaining high oil yield in ra-
tional genetic engineering studies aimed at enhancing oil
accumulation in vegetative tissues of plants. To date, much
attention has been focused on the combined overexpression of
transcription factors involved in seed storage product accumu-
lation and seed maturation, such as WRIKLED1
LEC2 (Slocombe et al., 2009; Andrianov et al., 2010; Kelly et al.,
2013; Vanhercke et al., 2013). Aside from the growth and de-
velopmental defects associated with the ectopic overexpression
of such seed-specific master regulators (Stone et al., 2001;
Cernac and Benning, 2004; Baud et al., 2007), recent genetic
evidence suggests that the WRI class of transcription factors are
not involved in the regulation of FAS in vegetative tissues of
Arabidopsis (To et al., 2013 ). In addition, it has been reported
that WRI1-mediated TAG accumulation is dependent on the
addition of soluble sugar in growth media (Cernac and Benning,
2004), and ectopic overexpression of WRI1 alone only results in
marginal increases in TAG accumulation in leaves of soil-grown
plants (Sanjaya et al., 2011). Foliar tissues constitute a major
portion of the harvestable biomass of dedicated bioenergy
crops such as switchgrass. So far, however
of leaf TAG have been achieved (Troncoso-Ponce et al., 2013),
with the highest level among genetically modified Arabidopsis
plants being 5% per DW found in the sdp1 TAG lipase mutant
coexpressing WRI1 and DGAT1 (Kelly et al., 2013). Our study
shows that PDAT1 has a dual role in enhancing FAS and di-
recting FAs from membrane lipids to TAG in Arabidopsis leaves.
We show that the combined expression of PDAT1 and OLE1
increases leaf TAG to 6.4% per DW in the wild type and 8.6%
per DW in tgd1 without major negative growth consequences.
Given the growing recognition of the potential benefits of max-
imizing TAG content in vegetative tissues of crops (Durrett et al.,
2008; Ohlrogge et al., 2009; Ohlrogge, 2011
2013; Troncoso-Ponce et al., 2013), this clarification of the role
of PDAT1 in plants may enable new strategies for future genetic
engineering efforts aimed at enhancing oil accumulation in
biomass crops used for biofuel production.
, only modest levels
; Chapman et al.,
Plant Materials and Growth Conditions
The Arabidopsis thaliana plants used in this study were of the Columbia
ecotype. The tgd1 mutant was previously described (Xu et al., 2003), as
were dgat1-1 and pdat1-2 (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) (Murashige and
Skoog, 1962) medium supplemented with 1% (w/v) Suc in an incubator
with a photon flux density of 80 to 120 µmol m–2s–1, a light period of 16 h
(22°C), and a dark period of 8 h (18°C). For growth in soil, plants were first
grown on MS medium for 10 d and then transferred to soil and grown
under a photosynthetic photon flux density of 150 to 200 mmol m22s21at
22/18°C (day/night) with a 16-h-light/8-h-dark period.
Generation of Plant Expression Vectors and Plant Transformation
The full-length coding regions of PDAT1 and DGAT1 were amplified by
RT-PCR using the primers 59-GCGTGGTACCATGCCCCTTATTCATCG-
GA-39 and 59-ACGTCTGCAGTCACAGCTTCAGGTCAATACGCTC-39 for
PDAT1 and 59-ACCTGGAGCTCATGGCGATTTTGGATTCTGC-39 and 59-
CCGAGGTACCTCATGACATCGATCCTTTTCGGT-39 for DGAT1. The
resulting PDAT1 and DGAT1 PCR products were digested with KpnI-PstI
or SacI-KpnI, respectively, and inserted into the respective sites of a bi-
the OLE1-GFP fusion construct, the entire genomic DNA encoding the
Arabidopsis OLE1 was amplified using the primers 59-ATGGCGGATA-
CAGCTAGAG-39 and 59-AGTAGTGTGCTGGCCACC-39 and ligated into
pCR8 topo-cloning entry vector (Invitrogen). The gene was then fused in
frame with GFP at the C terminus through the LR reaction to the
nation vector pGKPGWG (Zhong et al., 2008). After confirming the in-
tegrity of the constructs by sequencing, plant stable transformation was
performed according to Clough and Bent (1998). Transgenic plants were
selected in the presence of the respective antibiotics for the vectors on
MS medium lacking Suc.
10The Plant Cell
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 vol-
ume) and 1 M KCl-0.2 M H3PO4according to Dörmann et al. (1995).
Neutral and total polar lipids were separated on silica plates (Si250 with
a preadsorbant layer; Mallinckrodt Baker) by 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 spraying 5% H2SO4fol-
lowed by charring. For quantitative analysis, lipids were visualized by brief
exposure to iodine vapor and identified by cochromatography with lipid
standards. Individual lipids were scraped from the plate and used to
prepare FAMEs. Separation and identification of the FAMEs was per-
formed onanHP5975gaschromatography–massspectrometer (Hewlett-
Packard) fitted with 60 m 3 250-µm SP-2340 capillary column (Supelco)
with helium as a carrier gas. The methyl esters were quantified using
heptadecanoic acid as the internal standard as described by Fan et al.
(2011). The TAG content was calculated as described previously (Li et al.,
2006). The FA composition at the sn-2 position of the glycerol backbone
was determined by Rhizopus arrhizus lipase digestion as described by
Härtel et al. (2000). Pigments were quantified according to Lichtenthaler
In Vivo Acetate Labeling
Invivo labeling experiments with14C-acetate weredone according to Koo
et al. (2005). Briefly, rapidly growing leaves of 7-week-old plants were cut
in strips and then incubated in the light (60 µmol m22s21at 22°C with
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 the addition of 0.1 mCi
mmol; American Radiolabeled Chemicals). At the end of the incubation,
leaf strips were washed three times with water and blotted onto filter
paper. For the chase period, leaf tissue was incubated in the same
medium lacking14C-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 or different lipid classes was
determined by liquid scintillation counting.
of14C-acetate (106 mCi/
Staining with Fat Red 7B
Seedlings were stained with a 0.1% (w/v) solution of Fat Red 7B (Sigma-
Aldrich) overnight essentially as described by Brundrett et al. (1991).
Following brief rinsing with distilled water, samples were visualized with
a Wild Heerbrugg dissecting microscope.
For OD imaging, leaf tissues were stained with a neutral lipid-specific
fluorescent dye, Nile red (Sigma-Aldrich), at a final concentration of 10 µg/
mL and observed under a Zeiss epifluorescence microscope (Carl Zeiss
Axiovert 200M) with a GFP filter. For TEM, leaf tissues were fixed with
2.5% (v/v) glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.4, for 2
h and then postfixed with 1% osmium tetroxide in the same buffer for 2 h
at room temperature. After dehydration in a graded series of ethanol, the
tissues were embedded in EPON812 resin (Electron Microscopy Scien-
ces) and sectioned and stained with 2% uranyl acetate and lead citrate
before viewing under a JEOL JEM-1400 LaB6 120KeV transmission
electron microscope. For light microscopy observation, the sections were
stained with 1% toluidine blue and examined using a Zeiss epifluor-
escence microscope as described above.
Sequence data from this article can be found in the Arabidopsis Genome
Initiative or GenBank/EMBL databases under the following accession
numbers:DGAT1, At2g19450; OLE1,
The following materials are available in the online version of this article.
Supplemental Figure 1. Changes in Levels of DGAT1 and PDAT1
Transcript during Leaf Development.
Supplemental Figure 2. TAG Levels in Transgenic Plants Over-
expressing DGAT1, PDAT1, or OEL1.
Supplemental Figure 3. Growth and Lipid Phenotypes of the Trans-
genic Lines Coexpressing PDAT1 and OLE1.
Supplemental Figure 4. FA Composition of Individual Membrane
Lipids from Leaves of PDAT1 Overexpressors.
Supplemental Figure 5. Time Course of14C-Acetate Incorporation
into Total Lipids of Leaf Strips from 7-Week-Old Wild-Type Plants
Grown in Soil.
Supplemental Figure 6. Growth Phenotype and Total Leaf Polar Lipid
Content of Transgenic Plants Coexpressing PDAT1 and GFP-OLE1 in
the tgd1-1 Background.
Supplemental Figure 7. Overexpression of PDAT1 in act1 Leads to
Defects in Chloroplast Division and Thylakoid Biogenesis.
Supplemental Figure 8. Pulse-Chase Acetate Labeling of FAs
Associated with Individual Lipids of the act1 Mutant and act1
We thank John Ohlrogge for providing pdat1-2 mutant seeds. We also
thank John Shanklin, John Ohlrogge, and Jitao Zou for critical reading of
the article. 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 DE-
AC0298CH10886 (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.
C.X. and J.F. designed the experiments. C.X., J.F., C.Y., and X.Z.
performed the research. C.X. and J.F. analyzed the data and wrote the
Received August 12, 2013; revised August 12, 2013; accepted Septem-
ber 12, 2013; published ▪▪▪.
Andre, C., Haslam, R.P., and Shanklin, J. (2012). Feedback
regulation of plastidic acetyl-CoA carboxylase by 18:1-acyl carrier
Role of PDAT1 Acyltransferase in Leaves 11
protein in Brassica napus. Proc. Natl. Acad. Sci. USA 109: 10107–
Andrianov, V., Borisjuk, N., Pogrebnyak, N., Brinker, A., Dixon, J.,
Spitsin, S., Flynn, J., Matyszczuk, P., Andryszak, K., Laurelli, M.,
Golovkin, M., and Koprowski, H. (2010). Tobacco as a production
platform for biofuel: Overexpression of Arabidopsis DGAT and LEC2
genes increases accumulation and shifts the composition of lipids
in green biomass. Plant Biotechnol. J. 8: 277–287.
Bao, X., Focke, M., Pollard, M., and Ohlrogge, J. (2000).
Understanding in vivo carbon precursor supply for fatty acid
synthesis in leaf tissue. Plant J. 22: 39–50.
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–
Bates, P.D., Ohlrogge, J.B., and Pollard, M. (2007). Incorporation of
newly synthesized fatty acids into cytosolic glycerolipids in pea
leaves occurs via acyl editing. J. Biol. Chem. 282: 31206–31216.
Baud, S., Mendoza, M.S., To, A., Harscoët, E., Lepiniec, L., and
Dubreucq, B. (2007). WRINKLED1 specifies the regulatory action of
LEAFY COTYLEDON2 towards fatty acid metabolism during seed
maturation in Arabidopsis. Plant J. 50: 825–838.
Brasaemle, D.L., Rubin, B., Harten, I.A., Gruia-Gray, J., Kimmel, A.
R., and Londos, C. (2000). Perilipin A increases triacylglycerol
storage by decreasing the rate of triacylglycerol hydrolysis. J. Biol.
Chem. 275: 38486–38493.
Browse, J., Warwick, N., Somerville, C.R., and Slack, C.R. (1986).
Fluxes through the prokaryotic and eukaryotic pathways of lipid
synthesis in the ‘16:3’ plant Arabidopsis thaliana. Biochem. J. 235:
Brundrett, M.C., Kendrick, B., and Peterson, C.A. (1991). Efficient
lipid staining in plant material with sudan red 7B or fluorol
[correction of fluoral] yellow 088 in polyethylene glycol-glycerol.
Biotech. Histochem. 66: 111–116.
Cernac, A., and Benning, C. (2004). WRINKLED1 encodes an AP2/
EREB domain protein involved in the control of storage compound
biosynthesis in Arabidopsis. Plant J. 40: 575–585.
Chapman, K.D., Dyer, J.M., and Mullen, R.T. (2013). Commentary:
Why don’t plant leaves get fat? Plant Sci. 207: 128–134.
Chapman, K.D., and Ohlrogge, J.B. (2012). Compartmentation of
triacylglycerol accumulation in plants. J. Biol. Chem. 287: 2288–
Clough, S.J., and Bent, A.F. (1998). Floral dip: A simplified method for
Agrobacterium-mediated transformation of Arabidopsis thaliana.
Plant J. 16: 735–743.
Cronan, J.E., Jr., Weisberg, L.J., and Allen, R.G. (1975). Regulation
of membrane lipid synthesis in Escherichia coli. Accumulation of
free fatty acids of abnormal length during inhibition of phospholipid
synthesis. J. Biol. Chem. 250: 5835–5840.
Dahlqvist, A., Ståhl, U., Lenman, M., Banas, A., Lee, M., Sandager,
L., Ronne, H., and Stymne, S. (2000). Phospholipid:diacylglycerol
independent formation of triacylglycerol in yeast and plants. Proc.
Natl. Acad. Sci. USA 97: 6487–6492.
Dörmann, P., Hoffmann-Benning, S., Balbo, I., and Benning, C.
(1995). Isolation and characterization of an Arabidopsis mutant
deficient in the thylakoid lipid digalactosyl diacylglycerol. Plant Cell
triacylglycerols as feedstocks for the production of biofuels. Plant J.
Fan, J., Andre, C., and Xu, C. (2011). A chloroplast pathway for the de
novo biosynthesis of triacylglycerol in Chlamydomonas reinhardtii.
FEBS Lett. 585: 1985–1991.
Frentzen, M. (1998). Acyltransferases from basic science to modified
seed oils. Fett-Lipid 100: 161–166.
Frentzen, M., Heinz, E., McKeon, T.A., and Stumpf, P.K. (1983).
acyltransferase and monoacylglycerol-3-phosphate acyltransferase
from pea and spinach chloroplasts. Eur. J. Biochem. 129: 629–636.
Froissard, M., D’andréa, S., Boulard, C., and Chardot, T. (2009).
Heterologous expression of AtClo1, a plant oil body protein,
induces lipid accumulation in yeast. FEMS Yeast Res. 9: 428–438.
Hajdukiewicz, P., Svab, Z., and Maliga, P. (1994). The small,
versatile pPZP family of Agrobacterium binary vectors for plant
transformation. Plant Mol. Biol. 25: 989–994.
Härtel, H., Dörmann, P., and Benning, C. (2000). DGD1-independent
deprivation in Arabidopsis. Proc. Natl. Acad. Sci. USA 97: 10649–
Heinz, E., and Roughan, P.G. (1983). Similarities and differences in
lipid metabolism of chloroplasts isolated from 18:3 and 16:3 plants.
Plant Physiol. 72: 273–279.
Hellgren, L.I., and Sandelius, A.S. (2001). Age-dependent variation in
membrane lipid synthesis in leaves of garden pea (Pisum sativum
L.). J. Exp. Bot. 52: 2275–2282.
Horvath, S.E., Wagner, A., Steyrer, E., and Daum, G. (2011).
triacylglycerol metabolism in the yeast Saccharomyces cerevisiae.
Biochim. Biophys. Acta 1811: 1030–1037.
Huang, A.H.C. (1996). Oleosins and oil bodies in seeds and other
organs. Plant Physiol. 110: 1055–1061.
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.
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 seedfattyacid
affecting diacylglycerol acyltransferase activity. Plant Physiol. 108:
Kaup, M.T., Froese, C.D., and Thompson, J.E. (2002). A role for
Physiol. 129: 1616–1626.
Kelly, A.A., van Erp, H., Quettier, A.L., Shaw, E., Menard, G., Kurup,
S., and Eastmond, P.J. (2013). The sugar-dependent1 lipase limits
triacylglycerol accumulation in vegetative tissues of Arabidopsis.
Plant Physiol. 162: 1282–1289.
Kohlwein, S.D. (2010). Triacylglycerol homeostasis: Insights from
yeast. J. Biol. Chem. 285: 15663–15667.
Koo, A.J.K., Fulda, M., Browse, J., and Ohlrogge, J.B. (2005).
Identification of a plastid acyl-acyl carrier protein synthetase in
Arabidopsis and its role in the activation and elongation of
exogenous fatty acids. Plant J. 44: 620–632.
Kunst, L., Browse, J., and Somerville, C. (1988). Altered regulation
of lipid biosynthesis in a mutant of Arabidopsis deficient in
chloroplast glycerol-3-phosphate acyltransferase activity. Proc.
Natl. Acad. Sci. USA 85: 4143–4147.
Kunst, L., Browse, J., and Somerville, C. (1989). Altered chloroplast
structure and function in a mutant of Arabidopsis deficient in plastid
glycerol-3-phosphate acyltransferase activity. Plant Physiol. 90:
12 The Plant Cell
Li, Y., 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.
Lichtenthaler, H.K. (1987). Chlorophylls and carotenoids: Pigments
of photosynthetic membranes. Methods Enzymol. 148: 350–382.
Liu, B., and Benning, C. (2013). Lipid metabolism in microalgae
distinguishes itself. Curr. Opin. Biotechnol. 24: 300–309.
Lu, C.L., de Noyer, S.B., Hobbs, D.H., Kang, J.L., Wen, Y.C.,
Krachtus, D., and Hills, M.J. (2003). Expression pattern of
triacylglycerol biosynthesis, in Arabidopsis thaliana. Plant Mol. Biol.
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.
Mongrand, S., Bessoule, J.J., Cabantous, F., and Cassagne, C.
(1998). The C-16:3/C-18:3 fatty acid balance in photosynthetic
tissues from 468 plant species. Phytochemistry 49: 1049–1064.
Mora, G., Scharnewski, M., and Fulda, M. (2012). Neutral lipid
metabolism influences phospholipid synthesis and deacylation in
Saccharomyces cerevisiae. PLoS ONE 7: e49269.
Murashige, T., and Skoog, F. (1962). A revised medium for rapid
growth and bio assays with tobacco tissue cultures. Physiol. Plant.
Nunn, W.D., Kelly, D.L., and Stumfall, M.Y. (1977). Regulation of
biosynthesis in Escherichia coli. J. Bacteriol. 132: 526–531.
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–
Oelkers, P., Tinkelenberg, A., Erdeniz, N., Cromley, D., Billheimer,
J.T., and Sturley, S.L. (2000). A lecithin cholesterol acyltransferase-
like gene mediates diacylglycerol esterification in yeast. J. Biol.
Chem. 275: 15609–15612.
Ohlrogge, J., Allen, D., Berguson, B., Dellapenna, D., Shachar-Hill,
Y., and Stymne, S. (2009). Energy. Driving on biomass. Science
Ohlrogge, J., and Browse, J. (1995). Lipid biosynthesis. Plant Cell 7:
Ohlrogge, J., and Chapman, K.D. (2011). The seeds of green energy
- Expanding the contribution of plant oils as biofuels. The
Biochemist 33: 34–38.
Triacylglycerol lipolysis is linked to sphingolipid and phospholipid
metabolism of the yeast Saccharomyces cerevisiae. Biochim.
Biophys. Acta 1801: 1314–1322.
Roughan, P.G., and Slack, C.R. (1982). Cellular organization of
glycerolipid metabolism. Annu. Rev. Plant Physiol. 33: 97–132.
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–
Sandager, L., Gustavsson, M.H., Ståhl, 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:
Sanjaya, D., Durrett, T.P., Weise, S.E., and Benning, C. (2011).
Increasing the energy density of vegetative tissues by diverting
carbon from starch to oil biosynthesis in transgenic Arabidopsis.
Plant Biotechnol. J. 9: 874–883.
an enzyme involved in
the cessationof phospholipid
R., andDaum, G.
Sanjaya, M., Miller, R., Durrett, T.P., Kosma, D.K., Lydic, T.A.,
Muthan, B., Koo, A.J., Bukhman, Y.V., Reid, G.E., Howe, G.A.,
Ohlrogge, J., and Benning, C. (2013). Altered lipid composition
and enhanced nutritional value of Arabidopsis leaves following
introduction of an algal diacylglycerol acyltransferase 2. Plant Cell
Santos Mendoza, M., Dubreucq, B., Miquel, M., Caboche, M., and
Lepiniec, L. (2005). LEAFY COTYLEDON 2 activation is sufficient to
trigger the accumulation of oil and seed specific mRNAs in
Arabidopsis leaves. FEBS Lett. 579: 4666–4670.
Shanklin, J., and Cahoon, E.B. (1998). Desaturation and related
modifications of fatty acids. Annu. Rev. Plant Physiol. Plant Mol.
Biol. 49: 611–641.
Shimada, T.L., Shimada, T., Takahashi, H., Fukao, Y., and Hara-
Nishimura, I. (2008). A novel role for oleosins in freezing tolerance
of oilseeds in Arabidopsis thaliana. Plant J. 55: 798–809.
Siloto, R.M.P., Findlay, K., Lopez-Villalobos, A., Yeung, E.C.,
Nykiforuk, C.L., and Moloney, M.M. (2006). The accumulation of
oleosins determines the size of seed oilbodies in Arabidopsis. Plant
Cell 18: 1961–1974.
Slocombe, S.P., Cornah, J., Pinfield-Wells, H., Soady, K., Zhang,
accumulation in leaves directed by modification of fatty acid
breakdown and lipid synthesis pathways. Plant Biotechnol. J. 7:
Somerville, C., and Browse, J. (1996). Dissecting desaturation:
Plants prove advantageous. Trends Cell Biol. 6: 148–153.
Sperling, P., and Heinz, E. (1993). Isomeric sn-1-octadecenyl and sn-
2-octadecenyl analogues of lysophosphatidylcholine as substrates
for acylation and desaturation by plant microsomal membranes.
Eur. J. Biochem. 213: 965–971.
Ståhl, U., Carlsson, A.S., Lenman, M., Dahlqvist, A., Huang, B.,
Banas, W., Banas, A., and Stymne, S. (2004). Cloning and
acyltransferase from Arabidopsis. Plant Physiol. 135: 1324–1335.
Stone, S.L., Kwong, L.W., Yee, K.M., Pelletier, J., Lepiniec, L.,
Fischer, R.L., Goldberg, R.B., and Harada, J.J. (2001). LEAFY
COTYLEDON2 encodes a B3 domain transcription factor that
induces embryo development. Proc. Natl. Acad. Sci. USA 98:
Tjellström, H., Yang, Z., Allen, D.K., and Ohlrogge, J.B. (2012).
Rapid kinetic labeling of Arabidopsis cell suspension cultures:
Implications for models of lipid export from plastids. Plant Physiol.
To, A., Joubès, J., Barthole, G., Lécureuil, A., Scagnelli, A.,
Jasinski, S., Lepiniec, L., and Baud, S. (2012). WRINKLED
transcription factors orchestrate tissue-specific regulation of fatty
acid biosynthesis in Arabidopsis. Plant Cell 24: 5007–5023.
Troncoso-Ponce, M.A., Cao, X., Yang, Z., and Ohlrogge, J.B.
(2013). Lipid turnover during senescence. Plant Sci. 205-206: 13–
Vanhercke, T., El Tahchy, A., Shrestha, P., Zhou, X.R., Singh, S.P.,
and Petrie, J.R. (2013). Synergistic effect of WRI1 and DGAT1
coexpression on triacylglycerol biosynthesis in plants. FEBS Lett.
Winichayakul, S., Scott, R.W., Roldan, M., Hatier, J.H., Livingston,
S., Cookson, R., Curran, A.C., and Roberts, N.J. (2013). In vivo
packaging of triacylglycerols enhances Arabidopsis leaf biomass
and energy density. Plant Physiol. 162: 626–639.
Xu, C., Fan, J., Froehlich, J.E., Awai, K., and Benning, C. (2005).
Mutation of theTGD1 chloroplast
phosphatidate metabolism in Arabidopsis. Plant Cell 17: 3094–
envelope protein affects
Role of PDAT1 Acyltransferase in Leaves13
Xu, C., Fan, J., Riekhof, W., Froehlich, J.E., and Benning, C. (2003).
A permease-like protein involved in ER to thylakoid lipid transfer in
Arabidopsis. EMBO J. 22: 2370–2379.
Xu, C., Yu, B., Cornish, A.J., Froehlich, J.E., and Benning, C. (2006).
Phosphatidylglycerol biosynthesis in chloroplasts of Arabidopsis
acyltransferase. Plant J. 47: 296–309.
Yoon, K., Han, D., Li, Y., Sommerfeld, M., and Hu, Q. (2012).
Phospholipid:diacylglycerol acyltransferase is a multifunctional
enzyme involved in membrane lipid turnover and degradation while
synthesizing triacylglycerol in the unicellular green microalga
Chlamydomonas reinhardtii. Plant Cell 24: 3708–3724.
Zhang, M., Fan, J., Taylor, D.C., and Ohlrogge, J.B. (2009). DGAT1
Arabidopsis triacylglycerol biosynthesis and are essential for normal
pollen and seed development. Plant Cell 21: 3885–3901.
Zhong, S.L., Lin, Z.F., Fray, R.G., and Grierson, D. (2008). Improved
plant transformation vectors for fluorescent protein tagging.
Transgenic Res. 17: 985–989.
14 The Plant Cell
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