short title: leaf oil production rearranges membrane lipid flux … · 5 104 may be possible...

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Short title: Leaf oil production rearranges membrane lipid flux 1

Corresponding author: Philip D. Bates 2

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Title: Reorganization of acyl flux through the lipid metabolic network in oil-accumulating 4

tobacco leaves 5

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Xue-Rong Zhou1, Sajina Bhandari2, Brandon S. Johnson2, Hari Kiran Kotapati2, Doug K. 8

Allen3, Thomas Vanhercke1, Philip D. Bates2 9

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1. CSRIO Agriculture & Food, Canberra, ACT, Australia 11

2. Washington State University, Pullman, WA, USA 12

3. United States Department of Agriculture-Agricultural Research Service, Donald 13

Danforth Plant Science Center, St. Louis, MO, USA 14

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One-sentence Summary: 16

Engineering leaves to accumulate oils induced unexpected changes to fatty acid flux 17

through the leaf lipid metabolic network. 18

19

Footnotes: 20

Author contributions: 21

X.Z., T.V., and P.D.B. conceived the original research plans; X.Z., S.B., B.S.J., H.K., 22

and P.D.B performed the experiments and analyzed the data; D.K.A, analyzed the data 23

and contributed to the scope; P.D.B. wrote the article with contributions of all the 24

authors; P.D.B. agrees to serve as the author responsible for contact and ensures 25

communication. 26

27

Funding information: 28

This work is supported by Agriculture and Food Research Initiative Grant No. 2017-29

67013-26156, No. 2017-67013-29481, and the Hatch umbrella project #1015621 from 30

Plant Physiology Preview. Published on December 2, 2019, as DOI:10.1104/pp.19.00667

Copyright 2019 by the American Society of Plant Biologists

www.plantphysiol.orgon February 12, 2020 - Published by Downloaded from Copyright © 2019 American Society of Plant Biologists. All rights reserved.

http://www.plantphysiol.org

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the USDA National Institute of Food and Agriculture, and the National Science 31

Foundation Grant No. 1930559. 32

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Author of Contact: [email protected] 34

Abstract: 35

The triacylglycerols (TAGs; i.e., oils) that accumulate in plants represent the most 36

energy dense form of biological carbon storage, and are used for food, fuels, and 37

chemicals. The increasing human population and decreasing amount of arable land 38

have amplified the need to produce plant oil more efficiently. Engineering plants to 39

accumulate oils in vegetative tissues is a novel strategy, because most plants only 40

accumulate large amounts of lipids in the seeds. Recently, tobacco (Nicotiana tabacum) 41

leaves were engineered to accumulate oil at 15% of dry weight due to a push 42

(increased fatty acid synthesis) and pull (increased final step of TAG biosynthesis) 43

engineering strategy. However, to accumulate both TAG and essential membrane lipids, 44

fatty acid flux through non-engineered reactions of the endogenous metabolic network 45

must also adapt, which is not evident from total oil analysis. To increase our 46

understanding of endogenous leaf lipid metabolism and its ability to adapt to metabolic 47

engineering, we utilized a series of in vitro and in vivo experiments to characterize the 48

path of acyl flux in wild-type and transgenic oil-accumulating tobacco leaves. Acyl flux 49

around the phosphatidylcholine acyl editing cycle was the largest acyl flux reaction in 50

wild-type and engineered tobacco leaves. In oil-accumulating leaves, acyl flux into the 51

eukaryotic pathway of glycerolipid assembly was enhanced at the expense of the 52

prokaryotic pathway. However, a direct Kennedy pathway of TAG biosynthesis was not 53

detected as acyl flux through phosphatidylcholine preceded the incorporation into TAG. 54

These results provide insight into the plasticity and control of acyl lipid metabolism in 55

leaves. 56

57

Introduction 58

59

A finite supply of petroleum and a growing demand for energy to support 60

increasingly industrialized nations are global factors that emphasize the vital need to 61



3

develop renewable and sustainable sources of energy dense liquid fuels. The demand 62

is further exacerbated by growing populations and concerns linked to fossil fuel use and 63

associated waste streams. Seed-derived vegetative oil, mainly consisting of 64

triacylglycerol (TAG), provides a sustainable alternative. TAG-based plant oils are one 65

of the most energy-dense compounds found in nature. Plant oils are predominantly 66

used in the food industry (~80%), with the remainder supplying oleochemical production 67

(Carlsson et al., 2011). Due to their high energy density, they are increasingly viewed as 68

an attractive feed stock for production of biofuels (Lu et al., 2011). Breeding programs 69

and crop research in the last half century have substantially raised yields of oilseed 70

production, taking advantage of improved land, nutrient management, and more 71

efficient farming practices. Nevertheless, further gains in yield will require innovative, if 72

not disruptive, scientific approaches. The amount of arable land is finite and decreasing 73

with urban sprawl. As the world population continues to grow, agriculture production 74

must do more with less to meet food and energy demands. Non-seed derived plant oils 75

which can accumulate more lipids per acre of land are an attractive strategy, including 76

the production of oils in vegetative tissues of high biomass crops (Vanhercke et al., 77

2019). 78

Attempts to engineer oil in non-seed tissues have demonstrated increased TAG 79

levels by targeting different aspects of lipid biosynthesis, storage, and protection. These 80

include leaf, stem, tuber, root, or various vegetative tissues, in multiple plant species 81

including: Arabidopsis (Arabidopsis thaliana), tobacco (Nicotiana tabacum), potato 82

(Solanum tuberosum), sorghum (Sorghum bicolor), and sugarcane (Saccharum 83

officinarum) (reviewed in: Rahman et al., 2016; Xu and Shanklin, 2016; Vanhercke et 84

al., 2019). Our previous work generated a high oil (HO) tobacco line that accumulated 85

more than 15% dry weight TAG in leaf tissue by overexpressing: the Arabidopsis 86

transcription factor WRINKLED1 (AtWRI1) that upregulates glycolysis and fatty acid 87

synthesis (Focks and Benning, 1998; Cernac and Benning, 2004; Ma et al., 2013); the 88

Arabidopsis TAG biosynthetic enzyme ACYL-CoA: DIACYLGLYCEROL 89

ACYLTRANSFERASE 1 (AtDGAT1) (Katavic et al., 1995; Zou et al., 1999); and the 90

Sesamum indicum OLEOSIN gene in a combined push and pull strategy (Vanhercke et 91

al., 2014). The genetic changes in the HO line produced a large accumulation of fatty 92



4

acids in leaf TAG. However, the relationship between TAG synthesis and the underlying 93

leaf lipid metabolic network (Fig. 1), including effects on the accumulation of essential 94

leaf photosynthetic membranes, is unknown. The path (or flux) of fatty acids from 95

synthesis in the chloroplast to assembly into TAG in the endoplasmic reticulum (ER) is 96

critical to effectively control the amount and fatty acid composition of TAG without 97

detrimentally affecting membrane production. In particular for plant oil-based biofuels, 98

TAG containing high levels of monounsaturated fatty acids (e.g. oleate, 18:1 (# carbons: 99

# double bonds)) are desirable for the optimal mix of energy density, cold flow 100

properties, and oxidative stability of the fuels (Durrett et al., 2008). The HO leaves 101

accumulate TAG containing ~30% oleate and ~33% polyunsaturated fatty acids (PUFA) 102

(Vanhercke et al., 2014), indicating that substantial improvement of TAG composition 103



5

may be possible through further engineering. Changes to the PUFA level in plant TAG 104

are dependent on acyl flux through membrane lipid bound fatty acid desaturases 105

(Bates, 2016), however the impact of enhanced leaf oil production on acyl flux through 106

this biosynthetic network is less clear (Fig. 1). 107

Plant leaves have two parallel metabolic pathways of glycerol-3-phosphate (G3P) 108

acylation to produce membrane glycerolipids which have been characterized 109

biochemically and genetically over the past 50 years, and reviewed extensively, for 110

example: Roughan and Slack, 1982; Ohlrogge and Browse, 1995; Li-Beisson et al., 111

2013; Hurlock et al., 2014; Allen et al., 2015; LaBrant et al., 2018; Holzl and Dormann, 112

2019. In brief, plants synthesize fatty acids while esterified to acyl carrier proteins (ACP) 113

in the plastid. The plastid localized “prokaryotic” pathway of glycerolipid synthesis 114

utilizes acyl-ACPs to esterify 18:1 and 16:0 fatty acids to the sn-1 and sn-2 positions of 115

G3P respectively, producing first lysophosphatidic acid (LPA), then phosphatidic acid 116

(PA). Phosphatidylglycerol (PG) is produced from prokaryotic PA in the plastid, where 117

only “16:3” plants (including tobacco), also dephosphorylate PA to diacylglycerol (DAG) 118

producing a prokaryotic glycerolipid backbone containing a sn-2 16-carbon fatty acid for 119

synthesis of some of the plastid localized galactolipids, monogalactosyldiacyglycerol 120

(MGDG) and digalactosyldiacylglycerol (DGDG) (Mongrand et al., 1998). Plastid 121

localized desaturases produce the 16:3 by desaturation of the 16:0 incorporated into the 122

sn-2 position of MGDG and DGDG (Li-Beisson et al., 2013). The glycerolipid backbone 123

for the remaining galactolipids (or all galactolipids in 18:3 plants) is produced by the 124

“eukaryotic” pathway in the endoplasmic reticulum (ER) utilizing ER localized lipid 125

assembly enzymes. In the eukaryotic pathway, free fatty acids are exported from the 126

plastid and activated to acyl-CoAs prior to utilization by extra-plastidic acyltransferases. 127

The production of PA parallels that of the prokaryotic pathway except that 18-carbon 128

fatty acids are found at both sn-1 and sn-2. Any 16:0 present is localized to the sn-1 129

position (Frentzen et al., 1983), and is not further desaturated. Subsequent 130

dephosphorylation of PA produces the eukaryotic DAG backbone for synthesis of the 131

major ER membrane lipids phosphatidylcholine (PC) and phosphatidylethanolamine 132

(PE) (Li-Beisson et al., 2013). The production of eukaryotic galactolipids involves the 133

plastid localized assembly of MGDG from a eukaryotic DAG moiety derived from PC, 134



6

although the exact lipid that is transported from the ER to the plastid is unclear, but 135

could be PC, or the PC-derived intermediates PA or DAG (Hurlock et al., 2014; 136

Maréchal and Bastien, 2014; LaBrant et al., 2018; Karki et al., 2019). 137

Direct production of leaf TAG containing oleate in the HO tobacco line could 138

occur through utilization of newly synthesized oleoyl-CoA by the Kennedy pathway (Fig. 139

1A); however, the presence of PUFA in HO TAG indicates that other mechanisms of 140

acyl flux must be involved. Reactions which exchange acyl groups on and off PC are 141

integral to the eukaryotic pathway. PC is the site of ER localized fatty acid desaturation 142

of oleate (18:1Δ9) to make the PUFAs linoleate (18:2Δ9,12) and α-linolenate (18:3Δ9,12,15) 143

(Li-Beisson et al., 2013). PUFAs can enter the acyl-CoA pool to be used by the 144

eukaryotic pathway acyltransferases through a PC deacylation and lyso-PC acylation 145

cycle coined “acyl editing” (Fig. 1B) (Bates et al., 2007). Through acyl editing oleate is 146

incorporated into PC for desaturation, and the corresponding PUFA can reenter the 147

acyl-CoA pool to be used for the synthesis of glycerolipids by Kennedy pathway 148

reactions (Bates, 2016). Quantitative analysis of acyl flux through the eukaryotic 149

pathway with in vivo metabolic labeling has indicated that most nascent fatty acids first 150

are incorporated into PC through acyl editing prior to acylation of G3P, and that fatty 151

acid flux around the acyl editing cycle is the largest lipid metabolic flux in many plant 152

tissues including: developing pea (Pisum sativum) leaves (Bates et al., 2007); soybean 153

(Glycine max) and camelina (Camelina sativa) embryos (Bates et al., 2009; Yang et al., 154

2017); and Arabidopsis seeds, leaves, and cell cultures (Bates et al., 2012; Tjellström et 155

al., 2012; Wang et al., 2012; Karki et al., 2019). Mechanisms of acyl transfer from 156

membrane lipids into TAG also include membrane lipid turnover resulting in DAG 157

containing PUFAs that are used for TAG biosynthesis (Fig. 1C) (Bates, 2016). In 158

various oilseed tissues PC-derived DAG is the major source for TAG synthesis (Bates 159

et al., 2009; Bates and Browse, 2011; Yang et al., 2017). In leaves, DAG derived from 160

chloroplast galactolipids can also be used to produce TAG by homeostatic mechanisms 161

(Xu and Shanklin, 2016), and during stress (Sakaki et al., 1990; Moellering et al., 2010; 162

Narayanan et al., 2016; Arisz et al., 2018). Thus, the composition of TAG produced in 163

leaves is a consequence of the relative rates of acyl flux through various membrane 164

lipid pools, and the relative rate of fatty acid desaturation within these lipid pools. 165



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The expression of the genes encoding AtWRI1, AtDGAT1, and sesame 166

OLEOSIN have led to an increased level of fatty acid synthesis and accumulation of 167

TAG in leaves of the HO tobacco line (Vanhercke et al., 2014). Transcriptomic analysis 168

of the HO line indicated upregulation of glycolysis and fatty acid synthesis (Vanhercke 169

et al., 2017), consistent with the function of WRI1 in plant tissues (Focks and Benning, 170

1998; Ma et al., 2013). However, there was little to no change in expression of the 171

acyltransferases involved in TAG and membrane lipid assembly (Vanhercke et al., 172

2017). Previous studies have indicated that transcript abundance may not correlate with 173

protein levels (Hajduch et al., 2010; Vogel and Marcotte, 2012), and transcript 174

abundance alone is a poor indicator of metabolic flux (Fernie and Stitt, 2012; 175

Schwender et al., 2014; Allen et al., 2015). Thus, the path of acyl flux through the lipid 176

metabolic network into TAG is unclear (Fig. 1A-C), including if turnover of the abundant 177

chloroplast lipids in leaves may be feeding TAG biosynthesis. Future leaf oil engineering 178

efforts may need to specifically target these facets of the lipid metabolic network to 179

optimize leaf TAG accumulation and composition. Therefore, to better understand the 180

pathways of acyl flux in wild-type tobacco leaves, and how these pathways are altered 181

when accumulating high levels of leaf TAG in the transgenic HO line, we performed a 182

series of in vitro enzymatic assays, and in vivo continuous pulse and pulse-chase 183

metabolic labeling studies that provide new insights into tobacco leaf lipid metabolism 184

and its engineering. 185

186

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Results 188

189

Engineering leaf TAG accumulation also effects the accumulation of leaf 190

membrane lipids 191

192

The HO line has a large increase in leaf TAG accumulation (Vanhercke et al., 193

2014), which was confirmed in the current effort. Our results indicate that the boost in 194

TAG was accompanied by changes to leaf membrane lipid abundance (Fig. 2). PC and 195

DAG increased whereas other membrane lipids including the galactolipids that are the 196

bulk of the chloroplast photosynthetic membranes, decreased, compared to the wild-197

type (WT) (Fig. 2A). The difference in lipid abundance was also reflected through 198

changes in fatty acid compositions (Fig. 3). Total leaf fatty acid composition (Fig. 3A) of 199

the HO line reflected alterations in the fatty acid composition of TAG (Fig. 3B) which 200

accumulated as the major lipid product (Fig. 2A). PC had a notable decrease in the 201

unsaturation index as 18:3 decreased and 18:1 significantly increased (Fig. 3C). MGDG 202

16:3 content decreased by half and 18:2 significantly increased (Fig. 3D). The change in 203

MGDG fatty acid composition was predominantly due to a reduction in 16-carbon fatty 204

acids at the sn-2 position, indicating an ~40% reduction in the proportion of prokaryotic 205

pathway produced MGDG (Supplemental Fig. S1A-C). 206

Considering that acyl and glycerol flux through PC is key to producing other 207

membrane lipids, we calculated the ratio of membrane lipids to PC (Fig. 2B). The 208

lipid/PC ratios dropped in the HO line and indicated a change in the redistribution of 209

fatty acid from PC to other lipids, though it is less clear if this is a consequence of 210

reduced biosynthesis, or enhanced turnover, or both. To better understand the changes 211

in the lipid metabolic network that accommodate TAG accumulation, the flux of acyl 212

groups through the lipid metabolic network was analyzed by both in vitro assays and in 213

vivo tracing of leaf lipid metabolism in the WT and HO line. 214

215

A direct linear Kennedy pathway of TAG biosynthesis is not active in WT or HO 216

leaf microsomes 217

218



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Microsomal studies have been invaluable in characterizing enzymatic functions 219



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that produce TAG through the linear Kennedy pathway in plants (Barron and Stumpf, 220

1962; Stymne and Stobart, 1984; Bafor et al., 1991). To determine if a direct Kennedy 221

pathway of TAG biosynthesis (Fig. 1A) is present, we assayed WT and HO tobacco leaf 222

microsomes for TAG production with [14C]G3P and 18:1-CoA (Fig. 4). No significant 223

TAG accumulation was detected within a 60 min assay, though the total label in lipids 224

produced by HO microsomes was approximately 5-fold higher than that in the WT 225

suggesting an overall upregulation in de novo glycerolipid assembly. In the HO line PA 226

was the major labeled product, suggesting that PA conversion to DAG may be limiting in 227

the isolated microsomes. The in vitro results indicate that efficient channeling of 228

substrates into TAG through a Kennedy pathway (Fig. 1A) may not occur in the HO line; 229

however, since some proteins can be lost during microsomal preparation, additional in 230

vivo pulse and pulse-chase metabolic labeling experiments were performed to further 231

study the acyl flux through lipids. 232

233



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TAG accumulation alters the relative flux of nascent fatty acids into the 234

eukaryotic and prokaryotic pathways of glycerolipid assembly 235

236

To understand how the push and pull engineering approach to produce leaf TAG 237

(Vanhercke et al., 2014) affects the initial flux of newly synthesized fatty acids into the 238

endogenous leaf lipid metabolic network (Fig. 1), we performed a continuous 239

[14C]acetate metabolic labeling of 3-120 minutes on leaf disks from 66-day-old WT and 240

HO plants. [14C]acetate is incorporated into the acetyl-CoA pool utilized for fatty acid 241



12

synthesis (Fig. 1), and short time point labeling is instructive for characterizing the initial 242

steps of nascent acyl flux into the lipid metabolic network (Allen et al., 2015). Total 243

incorporation of [14C]acetate into leaf lipids was linear for both the WT and the HO line 244

over the 120 min time course, and there was no statistical difference in total label 245

between genotypes at any time point (Supplemental Fig. S2). However, linear 246

regression indicated slopes of 50.5 ± 1.9 DPM µg chlorophyll-1 min-1 in the WT, and 43.1 247

± 1.3 DPM µg chlorophyll-1 min-1 in the HO line. The slopes were significantly different 248

with a p-value = 0.0035. The reason for the reduced slope of [14C]acetate incorporation 249

into lipids of the HO line is not immediately clear, however it could be due to dilution of 250

the exogenous [14C]acetate by the much larger flux of endogenous carbon into acetyl-251



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CoA and fatty acid production in the HO leaf cells compared to that in the WT. 252

Therefore, we normalized the total accumulation of HO lipids to the WT average total 253

lipid accumulation at each time point (Fig. 5) so that the relative rates of synthesis of 254

individual lipid classes between the genotypes could be compared. The normalization 255

slightly increased the total DPM µg chlorophyll-1 in each lipid class, but the pattern of 256

lipid synthesis essential for determining precursor-product relationships was unchanged 257

regardless of whether data was normalized (Fig. 5) or not (Supplemental Fig. S3). 258

In WT leaves most newly synthesized fatty acids accumulate in PC and MGDG 259

across the time course with only minor amounts in TAG (Fig. 5A). At early time points 260

PC is the major labeled lipid (Fig. 5C), and both PC and MGDG accumulate labeled 261

fatty acids at similar initial rates (Table 1) but by 120 min MGDG accumulates more 262

label (Fig. 5A, 5C). These results are consistent with: 1) PC as a first product of nascent 263

fatty acid incorporation into ER lipids (Bates et al., 2007; Tjellström et al., 2012); 2) de 264

novo synthesis of MGDG through the prokaryotic pathway; and 3) the precursor-product 265

relationship of PC and MGDG over time as acyl groups move through the eukaryotic 266

pathway of galactolipid synthesis (Li-Beisson et al., 2013). All other membrane lipids 267

initially accumulated little radiolabel, but slowly increased over time. This is consistent 268

with the redistribution of nascent fatty acids from PC to other lipids through acyl editing 269

(Bates et al., 2007; Bates, 2016), and the conversion of MGDG to DGDG within the 270

plastid (Kelly and Dormann, 2004; Hurlock et al., 2014; LaBrant et al., 2018). 271

The incorporation of nascent fatty acids into lipids of HO leaves was dramatically 272

different (Fig. 5B, 5D). Similar to the WT, in the HO line PC was the most labeled lipid at 273

the earliest time points (Fig. 5D) and had a similar rate of label accumulation (Table 1). 274

However, the next most labeled lipid was TAG (rather than MGDG of the WT, Fig. 5C) 275

and the initial rate of nascent fatty acid incorporation into TAG was 119-fold higher than 276

in the WT (Table 1). At the 3 min time point there was over twice the amount of nascent 277

fatty acids in PC (39.5 ± 3.9 DPM/µg chl) than TAG (17.0 ± 4.4 DPM/µg chl). However, 278

the accumulation of labeled fatty acids in TAG continued to accelerate surpassing PC 279

by 15 min, and by 120 min TAG accumulated 1.7-fold more labeled fatty acids than PC, 280

representing ~48% of total labeled lipids (Fig. 5B, 5D). This result is consistent with the 281

very large mass accumulation of TAG in HO leaves (Fig. 2). Despite the larger mass 282



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accumulation of TAG over time, the more rapid labeling of PC at initial time points 283

suggests a PC-TAG precursor-product relationship for fatty acid flux. 284

PE which is produced in the ER through the eukaryotic pathway similar to PC 285

and TAG, did not have a significant difference in the rate of synthesis with nascent fatty 286

acids in the WT and the HO line (Table 1). However, there was a significant decrease in 287

the rates of nascent fatty acid incorporation into chloroplast lipids MGDG (-4.9 fold), 288

DGDG (-2.5 fold), and PG (-1.9 fold) (Fig. 5, Table 1) in the HO line as compared to the 289

WT. For each of these lipids the initial rates of labeling represents synthesis through the 290

prokaryotic pathway, whereas eukaryotic pathway synthesis occurs over much longer 291

time scales as labeled fatty acids move through PC and ER-derived lipids and 292

eventually return to the chloroplast (Browse et al., 1986). Therefore, the results suggest 293

a shift in fatty acid allocation to the eukaryotic pathway over the prokaryotic pathway for 294

the production of TAG in the transgenic line. 295

In Figure 6 we estimated the relative flux of nascent fatty acids into the 296

eukaryotic and prokaryotic glycerolipid assembly pathways by comparing the 297

accumulation of label in ER localized (PC, PE, TAG) and plastid localized (MGDG, 298

DGDG, PG) lipids. The metabolic labeling of WT leaves showed that nascent fatty acids 299

accumulated into ER lipids at a slightly greater rate than plastid lipids (Fig. 6A). Linear 300

regression of the initial phase of glycerolipid assembly (first 10 min) indicated that flux of 301

newly synthesized fatty acids into glycerolipids was 20.3 ± 4.6 and 17.1 ± 1.5 DPM µg 302

chlorophyll-1 min-1 for ER and plastid lipids, respectively. The initial ER/plastid ratio at 3 303

min of labeling was ~2.4 but dropped to 1.3 by 10 min. This change is likely reflected by 304

the lipids quantified at these time points. In the eukaryotic pathway nascent fatty acids 305

exported from the plastid are initially directly incorporated into PC, but in the prokaryotic 306

pathway nascent fatty acids are first incorporated into LPA, PA, and DAG prior to 307

MGDG synthesis (Allen et al., 2015). Considering that lipid classes LPA, PA, and DAG 308

occur in both pathways (Fig. 1) they were not included in the analysis. Therefore, the 309

lag in acyl flux through intermediates of the prokaryotic pathway at short time points 310

may explain the ratio favoring the eukaryotic pathway at short time points. The changing 311

ratio of labeled fatty acids in ER/plastid lipids stabilized by 10 min, then slowly 312

decreased over the time course. However, in the HO line the relative initial rate of newly 313



15

synthesized fatty acid accumulation in ER and plastid lipids was 34.9 ± 4.6 and 3.6 ± 314



16

0.6 DPM µg chlorophyll-1 min-1, respectively. Thus, the eukaryotic pathway accounted 315

for a 9-fold higher flux of fatty acids into glycerolipids than the prokaryotic pathway of 316

the HO line. Similar to the WT, in the HO line the ER/plastid ratio for labeled fatty acid 317

accumulation stabilized by 10 min and then decreased over the time course (Fig. 6B). 318

The decrease in the ER/plastid ratio over time in both genotypes likely represents the 319

PC-galactolipid precursor-product relationship of the eukaryotic pathway. 320

In Figure 5 and 6, the accumulation of newly synthesized fatty acids in MGDG 321

can be due to both the prokaryotic and eukaryotic pathways. To determine if the 322

reduction in accumulation of labeled MGDG is due to reduced acyl flux through the 323

prokaryotic, eukaryotic, or both pathways we collected MGDG from the 30 and 120 min 324

time points and analyzed the radioactivity in individual molecular species (Supplemental 325

Fig. S4 and S5). The prokaryotic pathway initially produces the 18:1/16:0 molecular 326

species of MGDG which is further desaturated to predominantly 18:3/16:3 (Ohlrogge 327

and Browse, 1995). Eukaryotic MGDG is indicated to be synthesized from a 328

polyunsaturated-containing-DAG ultimately derived from PC, and is further desaturated 329

to predominantly 18:3/18:3 in the plastid (Slack et al., 1977; Ohlrogge and Browse, 330

1995). Therefore, 18/16-carbon-containing molecular species are representative of the 331

prokaryotic pathway, and 18/18-carbon molecular species are representative of the 332

eukaryotic pathway. The accumulation of MGDG through each pathway is summarized 333

in Table 2. The analysis of MGDG molecular species gave four insights into the acyl flux 334

through the prokaryotic and eukaryotic pathways: (1) the HO line had a reduced 335

proportion of prokaryotic MGDG molecular species (and thus increased eukaryotic 336

proportion) as compared to the WT; (2) however with the very large decrease in total 337 14C-MGDG accumulation (Fig. 5), the total acyl flux into MGDG through the eukaryotic 338

pathway was reduced by over 30% and acyl flux through the prokaryotic pathway was 339

reduced by over 70% (Table 2); (3) both lines had an increase in eukaryotic molecular 340

species from 30 to 120 min of labeling (Table 2) consistent with the role of the 341

eukaryotic pathway for PC turnover to produce plastid galactolipids, and consistent with 342

the decrease in the ER/plastid accumulation ratio of Fig. 6; (4) the profile of prokaryotic 343

and eukaryotic MGDG molecular species in the HO line suggested a reduced rate of 344

plastid desaturation, as compared to the WT (Supplemental Fig. S5). Therefore, an 345



17

increase in the total flux of newly synthesized acyl groups into ER lipids (mostly TAG, 346

Fig. 5, 6), and a reduction in acyl flux into plastid lipids through both the prokaryotic and 347

eukaryotic pathways (Table 2) contributed to the dramatic redistribution of acyl flux 348

through the lipid metabolic network in the HO tobacco line. 349

350

Regiochemical analysis indicates limited changes in pathway structure for initial 351

steps of ER glycerolipid assembly 352

353

To better understand which branches of the lipid metabolic network (Fig. 1) are 354

involved in the altered flux of nascent fatty acids into membrane lipids and TAG of the 355

HO line as compared to the WT, we performed regiochemical analysis (Fig. 7) of 356

labeled DAG, PC and TAG across the [14C]acetate labeling time course from Fig. 5. In 357

both the WT and the HO line newly synthesized fatty acids were initially incorporated 358

more on the sn-1 position relative to the sn-2 position of the total labeled DAG pool, but 359

this was quickly equilibrated to approximately equal distribution by 10 min (Fig. 7A, B), 360

and there was no statistical difference in stereochemical labeling in DAG between the 361

lines. The higher initial labeling of DAG sn-1 position over sn-2 has previously been 362

reported in the predominantly eukaryotic de novo DAG pools of developing soybean 363

embryos and Arabidopsis seeds (Bates et al., 2009; Bates et al., 2012). Rapid 364

equilibrium of labeling across stereochemical positions in tobacco plants may also 365

represent a substantial contribution of prokaryotic DAG which is produced from only 366

nascent fatty acids (Ohlrogge and Browse, 1995), and thus the labeled fatty acids will 367

be evenly distributed across both positions as demonstrated for prokaryotic lipids in 368

rapeseed (Brassica napus) leaves (Williams et al., 2000). In contrast to DAG, newly 369

synthesized fatty acids accumulated predominantly in the sn-2 position of PC across the 370

time course in both plants (~63-70% WT and 67-76% HO). The slightly more nascent 371

fatty acids at the sn-2 position of PC in the HO line was only significant at the 3- and 372

120-min time points (Fig. 7C, D). The PC stereochemical labeling is consistent with 373

previous leaf, seed, and cell culture analyses where a single nascent fatty acid is initially 374

incorporated next to a previously synthesized fatty acid within PC (with preference for 375

sn-2 over sn-1) through acyl editing as nascent fatty acids leave the plastid (Bates et al., 376



18

2007; Bates et al., 2009; Bates et al., 2012; Tjellström et al., 2012; Allen, 2016; Bates, 377



19

2016; Karki et al., 2019). 378

Partial TAG lipase digestions of labeled TAG from both plants revealed that most 379

labeled acyl groups were released by the lipase in the free fatty acid fraction (sn-1 or 380

sn-3) and little remained in the monoacylglycerol fraction (sn-2) (Figure 7E, F). 381

Considering the similar labeling of the sn-1 and sn-2 position of DAG, the low sn-2 382

labeling of TAG suggests that most TAG labeling within this short time course 383

represents incorporation of a newly synthesized radiolabeled fatty acid onto the sn-3 384

position of an unlabeled DAG molecule, and does not reflect the rapidly produced 385

eukaryotic de novo DAG that might be expected from a direct Kennedy pathway of TAG 386

synthesis (Fig. 1A). Together the DAG, PC and TAG regiochemical analysis suggests 387

that even though there are big differences between the WT and the HO line for the 388

quantity of acyl flux into eukaryotic pathway lipids, the initial steps of eukaryotic 389

glycerolipid assembly (or the initial structure of the eukaryotic lipid network) between 390

these genotypes do not vary considerably. 391

392

Nascent acyl groups initially incorporated into PC are redistributed differently 393

between the WT and the HO line. 394

395

The short time point [14C]acetate labeling in Fig. 5 demonstrated that a majority 396

of newly synthesized fatty acids are initially incorporated into PC of both genotypes. The 397

redistribution of fatty acids from PC to other lipids over time was assessed through an 398

additional pulse-chase experiment (Fig. 8, Fig. 9). Leaf disks of 73-day-old plants were 399

pulsed with [14C]acetate for 0.5 hours, rinsed and incubated without the radiolabel for up 400

to 22 hrs. In both plants the total 14C-labeled lipids increased throughout the pulse and 401

chase periods (Fig. 8). During the pulse, labeled lipids accumulated at the most rapid 402

rates (4800 ± 500 and 9600 ± 900, DPM µg chlorophyll-1 hr-1 in WT and HO, 403

respectively). During the first 2 hrs of the chase period the rate of labeled lipid 404

accumulation was reduced 6-7 fold to 840 ± 210 and 1300 ± 340 DPM µg chl-1 hr-1 in 405

WT and HO leaves respectively, which likely represents continued uptake of 406

[14C]acetate during the washes. Finally from 2-22 hrs radiolabel accumulated at even 407

slower but constant rates of 330 ± 90 and 250 ± 50 DPM µg chl-1 hr-1, which may 408



20

represent continued utilization of a pool of [14C]acetate that was taken up into the leaf 409



21

tissue during the pulse but utilized at a slower rate, as compared to the bulk of the 410

[14C]acetate substrate. Therefore, the experiment should be considered a rapid 14C 411

pulse that is followed by labeling with a significantly lower concentration of [14C]acetate 412

(15-38 fold lower based on initial and final rates). This distinction is relevant when 413

comparing the total accumulation of radiolabel in individual lipid classes (Fig. 9A-B) to 414

the relative radiolabel accumulation between lipid classes in each genotype (Fig. 9C-D). 415

Similar to the short time point pulse experiment (Fig. 5, Fig. 6), the HO line accumulated 416

labeled fatty acids predominantly in ER lipids across the time course, whereas labeling 417

of ER and plastid lipids was similar across the time course in the WT (Fig. 8). 418



22

In the WT all individual lipid classes accumulated 14C acyl groups during the 419

pulse-chase but at different rates across the time course (Fig. 9A). At the end of the 420

pulse, PC contained the most 14C with over 2.6-fold more 14C than any other lipid, but 421

the rate of labeled fatty acid accumulation in PC continued to slow down across the 422

chase time course. During the chase, the accumulation of 14C fatty acids increased in 423

MGDG relative to PC such that by the end of the time course they contained similar 424

amounts of total labeled fatty acids. The results are consistent with the PC-MGDG 425

precursor-product relationship of the eukaryotic pathway in leaves, and the 426

redistribution of nascent acyl groups from PC to other ER localized lipids through acyl 427

editing (Fig. 1). In HO leaves the [14C]acetate pulse-chase results are distinct. Initially 428

PC contained the most label after the pulse but was surpassed by TAG before the 2 hr 429

time point (Figure 9B, D). By the end of the 22 hr chase period TAG accumulated ≥ 4.9-430

fold more 14C fatty acids than any other lipid. Even though the total lipid labeling is 431

increasing over the chase period (Fig. 8B), the amount of labeled fatty acids in PC of 432

the HO line decreases over the whole chase period (Figure 9B). The difference in 433

accumulation of 14C fatty acids in PC between the WT and the HO line suggests that PC 434

turnover and redistribution of acyl groups occurs at a higher rate in the HO line. All other 435

membrane lipids in HO leaves increased slightly during the chase (Fig. 9B), but much 436

less than in WT leaves. In the WT, PC and MGDG had a clear precursor-product 437

relationship of acyl flux that is not directly evident in the HO line of Fig. 9B. To 438

determine if the small increases in HO MGDG 14C acyl accumulation are due to (1) the 439

continued synthesis of 14C fatty acids and their incorporation into the metabolic network 440

during the chase (Figure 8B); or (2) are due to a reduced redistribution of acyl label from 441

PC, we compared the labeled MGDG molecular species distribution at the 0 and 22 hr 442

time points (Supplemental Figure S6, summarized in Table 3). After the 30 min pulse, 443

the proportion of eukaryotic and prokaryotic MGDG molecular species in both the WT 444

and the HO line was similar to that of the 30 min continuous labeling time point (Table 445

2). During the 22 hr chase period in the WT, the eukaryotic MGDG molecular species 446

increased 3-fold as a proportion (Table 3), consistent with the PC-MGDG precursor-447

product relationship of the eukaryotic pathway. However, during the chase in the HO 448

line, the proportion of eukaryotic MGDG molecular species only increased 1.5-fold 449



23

(Table 3). Therefore, the reduced accumulation of MGDG in the HO line during the 450

chase (Fig. 9B) is also consistent with a reduced redistribution of acyl groups from PC 451

to MGDG through the eukaryotic pathway MGDG. 452

Pulse-chase experiments are commonly represented as the percent labeling in 453

the different products over time (Figure 9C, D), yet the interpretation is dependent on 454

the relative accumulation of the total lipids (Fig. 8A, B) and each individual lipid (Fig. 9A, 455

B) over the chase period. In both genotypes PC had the largest decrease in proportional 456

labeling, consistent with the conclusions from above that acyl groups are redistributed 457

from PC to other lipids over the time course. However, considering that nascent 14C acyl 458

groups continue to enter the system over the chase (Fig. 8) and are predominantly 459

incorporated into PC first (Fig. 5, Fig. 9), the actual turnover of PC is greater than the 460

apparent turnover of half of the labeled PC in the WT, and over 76% of the labeled PC 461

in HO leaves. In addition, the proportional labeling of MGDG in both genotypes also 462

decreased from 2-22 hrs of the chase. This represents both the MGDG-DGDG 463

precursor-product relationship of lipid synthesis (Li-Beisson et al., 2013), as well as the 464

continual incorporation of labeled acyl groups into predominantly PC of the WT, and 465

both PC and TAG of the HO line. Hence the apparent 32% decrease in MGDG 466

accumulation in the HO line does not indicate that MGDG is turning over to feed the 467

large increase in TAG accumulation, but it is the result of the labeled acyl group 468

accumulation predominantly in TAG as more fatty acids are synthesized over the time 469

course (Fig. 8). Therefore, the combined HO pulse-chase results indicate that TAG 470

synthesis draws acyl groups predominantly from PC turnover (Fig. 9B, 9D), which may 471

compete with eukaryotic pathway MGDG synthesis for acyl groups (Table 3), but there 472

does not appear to be evidence of galactolipid turnover providing substrates for TAG 473

biosynthesis. 474

475

476



24

Discussion 477

478

Biotechnology may help to meet societal needs by engineering metabolism to 479

enhance the production of biological resources for food or industry. Plant lipids can be 480

one part of this solution through increased oil yields per area of land for biofuel 481

production. The current state of vegetative oil engineering involves the expression of 482

only a few genes including: transcription factors to increase fatty acid synthesis, DGAT 483

to convert DAG to TAG, and oleosin to prevent TAG breakdown in a push-pull-protect 484

strategy (Vanhercke et al., 2014; Xu and Shanklin, 2016; Vanhercke et al., 2017). 485

However, TAG biosynthesis requires many additional enzymatic steps that directly 486

overlap with essential membrane lipid production (Fig. 1, (Bates and Browse, 2012)), 487

and quantitative analysis of the oil end product does little to explain the metabolic path 488

fatty acids take to accumulate in TAG. It is also unclear how an introduced DGAT fits 489

into the leaf lipid metabolic network designed to accumulate ER and chloroplast 490

membrane lipids, or which substrate pools are used in TAG biosynthesis (Fig. 1). For 491

biofuel production, newly synthesized 18:1 could be directly incorporated into TAG with 492

a minimal number of enzymatic steps using the Kennedy pathway (Fig. 1A), however 493

this would not account for the presence of 18:2 and 18:3 measured in TAG. To 494

understand the path of acyl flux through the lipid metabolic network in WT tobacco 495

leaves, and how the engineered changes in HO affect acyl flux, we analyzed the 496

mechanisms of acyl flux in WT and HO leaves. 497

498

A Kennedy pathway of TAG assembly is not present in HO leaves 499

500

TAG composed of oleate is a desirable quality for biofuel production (Durrett et 501

al., 2008). The least number of steps to incorporate oleate into TAG is directly through 502

the Kennedy pathway reactions: glycerol-3-phosphate acyltransferase (GPAT) and 503

lysophosphatidic acid acyltransferase (LPAT) to produce PA, dephosphorylation by 504

phosphatidic acid phosphatase (PAP) to produce DAG, and acylation of DAG by to 505

produce TAG. (Fig. 1A (Bates, 2016)). The large increase in 16:0 and 18:1 in HO TAG 506

suggests that a Kennedy pathway utilizing newly synthesized fatty acids could produce 507



25

at least some of the TAG in the HO line (Fig. 2, 3). The only Kennedy pathway 508

acyltransferase that was directly engineered into tobacco was AtDGAT1 (Vanhercke et 509

al., 2014). Therefore, TAG fatty acid composition is also dependent on the acyl 510

selectivity and substrate pools of the endogenous tobacco GPAT and LPAT. In vitro 511

assays did not produce TAG with microsomes from either the WT or the HO line (Fig. 512

4). This result may suggest that the four reactions of the Kennedy pathway in the HO 513

line are not associated together in the isolated microsomes for efficient shuttling of 514

substrates within the in vitro reactions. To further understand the path of acyl flux in WT 515

and HO leaves we utilized an in vivo labeling approach. 516

Multiple lines of evidence from the in vivo labeling results suggest a traditional 517

Kennedy pathway is not the major pathway of TAG synthesis in HO leaves. First, even 518

though fatty acids accumulate in HO TAG to levels that are 12-times that of PC (Fig. 519

2A), nascent fatty acids are incorporated into PC faster than into TAG (Fig. 5D). 520

Second, during the pulse-chase, fatty acids are redistributed predominantly from PC 521

into TAG (Fig. 9). Third, regiochemical analysis of de novo synthesized DAG indicated 522

an equal partitioning of labeled acyl chains at both sn-1 and sn-2 whereas TAG 523

contained nascent fatty acids only at sn-3 (Fig. 7). The regiochemical data indicates that 524

de novo DAG produced by Kennedy pathway GPAT/LPAT reactions (Fig. 1A) is not 525

directly used for TAG biosynthesis. In combination with the in vitro assay, the results 526

suggest that overexpressed AtDGAT1 does not produce a Kennedy pathway that 527

channels newly synthesized fatty acids directly into TAG. 528

The results in this study are most consistent with Fig. 1 option C which indicates 529

that a second pool of DAG (other than Kennedy pathway de novo DAG) is used for TAG 530

synthesis. It is not immediately clear how the second DAG pool is produced, it could be 531

derived from de novo DAG, or PC, or a combination of the two. The pulse-chase results 532

indicate that galactolipids, including MGDG, are not used for TAG production (Fig. 9). 533

Thus, the reported mechanisms that turn over chloroplast lipids to produce DAG for leaf 534

TAG under stress conditions (Vanhercke et al., 2019), are unlikely to be actively 535

contributing to TAG accumulation in HO tobacco leaves. Metabolic labeling with 536

[14C]glycerol in developing oil seed tissues has suggested that a PC-derived DAG pool 537

is utilized for TAG synthesis (Bates et al., 2009; Bates and Browse, 2011; Yang et al., 538



26

2017). The current [14C]acetate acyl labeling cannot directly confirm a PC-derived DAG 539

pool, but the acyl labeling results are consistent with the previous studies. It is also 540

possible that immediately synthesized de novo DAG may feed into a larger and more 541

slow turnover DAG pool such as in oil bodies where AtDGAT1 may co-localize with 542

oleosin proteins. DAG can phase partition into oil bodies (Slack et al., 1980; Kuerschner 543

et al., 2008). Thus, if the rapidly labeled de novo DAG mixes with a larger unlabeled 544

pool in the oil body it would slow the apparent flux of the sn-1/2 labeled de novo DAG 545

into TAG relative to the sn-3 TAG labeling of the total mixed DAG pool. 546

547

Both WT and HO leaf acyl fluxes are dominated by phosphatidylcholine acyl 548

editing 549

550

In both WT and HO leaves, most newly synthesized fatty acids are immediately 551

incorporated into PC (Fig. 5). The difference in stereochemical incorporation of newly 552

synthesized fatty acid in DAG and PC (Fig. 7) indicates there is no DAG-PC precursor-553

product relationship at the earliest labeling time points. PC labeling as a percent of ER 554

lipid labeling (Fig. 6) at 3 min indicates that PC is 94.9 ± 1.5% of the total labeled ER 555

lipids in the WT, and 70 ± 4% in the HO line. The simplest interpretation of this result is 556

a shift in acyl flux away from PC acyl editing in the HO line for direct incorporation of 557

nascent fatty acids into the sn-3 position of TAG (Fig. 7). However, the production of 558

TAG at heightened levels requires three acyl chains, of which a substantial percentage 559

are PUFAs. Acyl editing is a constant exchange of acyl groups in PC with the acyl-CoA 560

pool to accommodate desaturation. Thus, if the rate of acyl editing was increased in the 561

HO line, a proportion of the labeled fatty acids initially incorporated into PC at time zero 562

would be redistributed back to the acyl-CoA pool for use by AtDGAT1 to produce TAG 563

within 3 minutes. This concept is supported with linear regression data used to 564

determine labeling rates in Table 1. Extrapolating back to time zero the x-intercepts of 565

PC are 0.91 for WT, and 0.93 for HO. For TAG the x-intercepts are 2.5 for WT, and 2.4 566

for HO. The similar labeling lag times between the WT and the HO line suggest a 567

common path of nascent fatty acid incorporation into ER lipids, though at a higher rate 568

(1.7-fold) for the HO line (Fig. 6). Thus, the rate of acyl editing in the HO line was 569



27

enhanced by the same amount (i.e. 1.7-fold) to accommodate the increased rate of fatty 570

acid export from the plastid, and PC is the first product of nascent fatty acid 571

incorporation into glycerolipids of the eukaryotic pathway. 572

The stereochemical distribution of labeled fatty acids in PC indicates that the 573

initial incorporation of nascent fatty acids into PC can occur at both positions but with an 574

approximately 2-fold preference for sn-2 (Fig. 7). The slightly higher PC sn-2 labeling in 575

the HO line suggests that the increase in PC acyl editing favors sn-2 over sn-1 576

positions. Therefore, acyl flux around the PC acyl editing cycle (Figure 1, option B) is 577

the dominate acyl flux reaction in both WT and HO tobacco, similar to what has been 578

demonstrated in leaves of pea, Arabidopsis, and rapeseed (Williams et al., 2000; Bates 579

et al., 2007; Karki et al., 2019), and developing seeds of soybean, camelina, and 580

Arabidopsis (Bates et al., 2009; Bates and Browse, 2011; Yang et al., 2017). Both PC 581

acyl chains are the major extra-plastidic sites for fatty acid desaturation (Sperling and 582

Heinz, 1993; Sperling et al., 1993), therefore 18:1 flux through PC acyl editing at both 583

sn-1 and sn-2 likely contributes to a PUFA containing acyl-CoA pool that leads to the 584

incorporation of PUFA in TAG of HO leaves. The decrease in the PC desaturation index 585

(Fig. 3C) is also consistent with an increased rate of acyl flux through PC, because 586

membrane lipid desaturation is dependent on both the rate of desaturation and the rate 587

of acyl flux through the membrane lipid. Increases in the fatty acid synthesis rate have 588

been demonstrated to increase 18:1 and decrease PUFA content of membrane lipids 589

(Maatta et al., 2012; Meï et al., 2015; Botella et al., 2016). Considering that the 590

engineering of a very large pull of acyl chains into TAG in the HO line only increases PC 591

acyl editing instead of drawing acyl chains away from it, PC acyl editing may be 592

considered a key part of fatty acid export from the plastid into the eukaryotic pathway. 593

Interestingly, both the [14C]acetate continuous pulse and the pulse-chase 594

experiments produced similar initial labeling in lipids for the WT and the HO line (Fig. 6, 595

Fig. 8), but the pulse-chase experiment showed a more dramatic labeling in the 596

immediate chase period in the HO line relative to the WT (Fig. 8). Such a description of 597

initial labeling is consistent with hypothesized transport of acyl chains out of the 598

chloroplast and directly into PC that subverts the large bulk acyl-CoA pool as has been 599

previously documented through bulk pool kinetic measurements with time course 600



28

labeling experiments (Tjellström et al., 2012; Allen, 2016) and isotopically labeled 601

mutant analysis (Bates et al., 2009; Karki et al., 2019), and is likely part of the acyl 602

editing mechanism where rapid labeling in PC from [14C]acetate was initially observed 603

(Bates et al., 2007; Bates et al., 2009). During the pulse-chase experiment, it may be 604

that the bulk acyl-CoA pool in the HO line is larger and becomes more labeled over the 605

duration of the pulse by enhanced flux through the acyl editing cycle, and therefore can 606

make a greater contribution to total lipid labeling during the initial phase of chase. 607

608

Reduced prokaryotic pathway and altered redistribution of acyl chains from PC to 609

other lipids in the HO line 610

611

Engineering the accumulation of TAG in the HO line reduced the steady-state 612

accumulation of chloroplast localized galactolipids by approximately 24% (Fig. 2). Total 613

MGDG content in the HO line was reduced ~19%, and the proportion of prokaryotic 614

pathway produced MGDG was reduced ~40% (Supplemental Fig. S1). DGDG is 615

produced mostly by eukaryotic pathway derived substrates, and total DGDG levels were 616

reduced by ~32% in the HO line as compared to the WT. Therefore, the mass 617

accumulation of galactolipids indicates that TAG accumulation in the HO line negatively 618

affects galactolipid production through both the prokaryotic and eukaryotic pathways. 619

The reductions in galactolipid levels in the HO line could be due to reduced 620

synthesis, increased turnover, or both which cannot be determined from the 621

quantification of steady-state lipid levels; but are reflected in time course-based acyl flux 622

experiments. At short time points [14C]acetate labeling of nascent fatty acid flux into 623

MGDG represents predominantly prokaryotic MGDG, which is reduced almost 5-fold in 624

the HO line (Fig. 5, Tables 1-2). Therefore, the reduction in prokaryotic MGDG 625

accumulation is primarily due to reduced synthesis. It’s also possible that homeostatic 626

turnover of galactolipids was reduced to allow higher accumulation of MGDG than 627

would be expected from the low rates of synthesis. To track the PC-MGDG precursor-628

product relationship of eukaryotic MGDG synthesis, we used pulse-chase analyses with 629

longer time points. The [14C]acetate pulse-chase labeling indicated that the 630

redistribution of acyl groups from PC in the HO line was predominantly into TAG with 631



29

reduced flux into eukaryotic MGDG synthesis as well as other lipids when compared to 632

the WT (Figure 9, Table 3). As there was no reduction in total 14C-MGDG accumulation 633

during the chase period in the HO line, the reduced eukaryotic MGDG accumulation 634

was due to reduced redistribution of acyl groups from PC to MGDG through the 635

eukaryotic pathway. Thus there is no evidence to suggest enhanced galactolipid 636

turnover in the HO line. 637

There are likely multiple alterations in enzymatic activity that led to the 638

redistribution of acyl flux through the lipid metabolic network in the HO line. From the 639

acyl flux analysis, we can propose several related hypotheses for future studies. First, 640

the massive increase in fatty acid accumulation in TAG of the HO line combined with 641

the reduced prokaryotic pathway are likely both related to an increase in acyl-ACP 642

thioesterase activity, which removes the substrate for the prokaryotic pathway and 643

initiates fatty acid export from the chloroplast (Bates et al., 2013; Li-Beisson et al., 644

2013). The gene expression of both thioesterases FATA and FATB were up regulated in 645

the HO line (Vanhercke et al., 2017). The reduced prokaryotic pathway flux (Fig 5, 9, 646

Tables 2-3), combined with the reduced rates of MGDG desaturation in the HO line 647

(Supplemental Figs. S5-S6) also suggest a possible general down regulation of 648

prokaryotic pathway enzymatic activity. 649

Second, within the eukaryotic pathway 1/3 of the fatty acids in TAG are 650

incorporated into TAG directly from the acyl-CoA pool by the acyltransferase activity of 651

the overexpressed AtDGAT1. In the WT the exchange of fatty acids from PC into the 652

acyl-CoA pool would be mostly used for de novo glycerolipid synthesis that would 653

produce the molecular species of PC used for eukaryotic galactolipid synthesis (Karki et 654

al., 2019). Therefore, the increased flux around the PC acyl editing cycle combined with 655

enhanced DGAT activity in the HO line would draw acyl flux away from PC and into sn-3 656

TAG, and reduce the amount of fatty acids available for de novo PC and galactolipid 657

synthesis. 658

Third, the reduction in eukaryotic galactolipid synthesis of the HO line may also 659

be due to reduced turnover of PC to produce the PC-derived substrate for galactolipid 660

synthesis, or the commandeering of that PC-derived substrate for TAG biosynthesis. 661

The identity of the eukaryotic pathway substrate that is transferred from the ER to the 662



30

plastid is not clear, and leading candidates include PC, and PC-derived PA and/or DAG 663

(Hurlock et al., 2014; LaBrant et al., 2018; Karki et al., 2019). If PC-derived DAG is the 664

substrate that is transferred from the ER to the chloroplast for galactolipid synthesis, 665

then the overexpressed AtDGAT1 may compete for the PC-derived DAG substrate in 666

the ER and reduce its transfer to the chloroplast for eukaryotic galactolipid synthesis. 667

However, if PA is the PC-derived species that is transferred to the chloroplast, then it 668

would not be a substrate for AtDGAT1 activity unless PA phosphatase activity was also 669

upregulated to convert PA to DAG. Our previous transcriptomics in the HO line 670

indicated increased expression of two phospholipase D isoforms which could produce 671

PA from PC, however an increase in PA phosphatase expression was not detected 672

(Vanhercke et al., 2017). In Arabidopsis the TRIGALACTOSYLDIACYLGLYCEROL 1 673

mutant (tgd1) or overexpression of (PDAT1) increases WT leaf TAG content from 674

<0.1% of dry weight to ~0.5% and ~1% of dry weight, respectively. Mutation of PA 675

phosphatase activity in the PHOSPHATIDIC ACID HYDROLASE 1 and 2 double mutant 676

(pah1 pah2) reduces this TAG accumulation in both the tgd1 and AtPDAT1 677

overexpression backgrounds suggesting that PA phosphatase activity may be involved 678

in leaf TAG production (Fan et al., 2014). However, the pah1 pah2 mutant has 679

increased synthesis and double the accumulation of leaf PC and PE content (Eastmond 680

et al., 2010). In WT Arabidopsis leaves PC and PE accumulate 5- to 10-fold more fatty 681

acids than TAG (Fan et al., 2014), therefore the effect of the pah1 pah2 double mutation 682

on leaf TAG accumulation in the tgd1 and AtPDAT1 overexpression lines may be due to 683

a shift in fatty acid allocation from TAG to ER membrane lipids, rather than a reduction 684

in TAG biosynthetic capacity. Therefore, the previous results in Arabidopsis and our 685

transcriptomics have not fully elucidated the role of PA phosphatases in leaf TAG 686

production. In addition, our analysis of acyl fluxes alone could not confirm if the DAG 687

pool for TAG synthesis was derived from PC or not. Therefore, further [14C]glycerol 688

labeling experiments to confirm if HO leaf TAG is derived from PC, combined with 689

analysis of changes in PC lipase and PAP enzymatic activities would be beneficial to 690

determining both the altered pathway fluxes in the HO line, as well as identifying the 691

PC-derived substrate that is used for eukaryotic galactolipid synthesis. 692

693



31

The tobacco leaf acyl flux analysis suggests strategies to reduce PUFA 694

accumulation in leaf oil 695

696

To reduce the PUFA content of oilseed crops, research has focused on reducing 697

seed specific FATTY ACID DESATURASE 2 and 3 (FAD2 and FAD3) activity through 698

mutations of isoforms mostly expressed in seeds but not vegetative tissue (Pham et al., 699

2010), or through seed specific RNA interference (Wood et al., 2018; Islam et al., 2019). 700

The purpose of the seed specific reduction in desaturase activity is to increase the 701

oleate content of the seed oil, but not effect leaf membrane lipid compositions in 702

vegetative tissue. ER membrane based FAD2 activity is required for proper leaf 703

membrane function, especially at low temperatures (Miquel et al., 1993). Due to the 704

importance of leaf desaturases for vegetative growth, a similar reduction of desaturase 705

activity would likely be counterproductive in a vegetative oil crop. The analysis of acyl 706

fluxes in WT and HO tobacco leaves presented here indicate that fatty acid flux through 707

the PC acyl editing cycle is the dominate reaction in the WT, and is enhanced at least 708

1.7-fold in the HO line. Because PC is the site for ER localized fatty acid desaturation 709

this movement of acyl groups through PC contributes to accumulation of PUFA in HO 710

leaf TAG. Therefore, an alternative strategy may be to alter acyl flux away from PC. In 711

Arabidopsis The LYSOPHOSPHATIDYLCHOLINE ACYLTRANSFERASEs (LPCAT1, 712

LPCAT2) are responsible for the direct incorporation of nascent fatty acids into PC 713

through acyl editing in both leaves (Karki et al., 2019) and seeds (Bates et al., 2012). 714

The lpcat1 lpcat2 double mutant alters acyl flux such that nascent fatty acids are first 715

esterified into glycerol lipids through the GPAT and LPAT reactions of the Kennedy 716

pathway, rather than PC acyl editing (Bates et al., 2012; Karki et al., 2019). In seeds, 717

this leads to an increase in the seed oil monounsaturated/polyunsaturated fatty acid 718

ratio from 0.72 to 0.84. When reduced exchange of DAG in and out of PC of the 719

PHOSPHATIDYLCHOLINE: DIACYLGLYCEROL CHOLINEPHOSPHOTRANSFERASE 720

mutant rod1 is combined with the lpcat1 lpcat2 double mutant, the ratio is further 721

increased to 3.95 in the lpcat1 lpcat2 rod1 triple mutant (Bates et al., 2012). From our 722

current analysis it was unclear if leaf TAG was produced from PC-derived DAG, but if 723

PC-derived DAG also contributes to leaf TAG a similar approach reducing acyl editing 724



32

and PC-derived DAG production may be valuable to alter acyl flux around PC to 725

increase the oleate content of leaf TAG while maintaining the PUFA content of 726

membranes. Therefore, the acyl flux analysis presented here has improved our 727

understanding of how leaf lipid metabolism responds to an increased push and pull of 728

fatty acids into TAG, as well as provided new hypotheses on how to further enhance 729

vegetative oil engineering. 730

731

In summary the analysis of acyl fluxes in WT and HO tobacco leaves indicate: (1) 732

The push and pull leaf oil production in the HO line reduces acyl flux into the prokaryotic 733

and enhances flux into the eukaryotic glycerolipid assembly pathways. (2) Fatty acids 734

entering the eukaryotic pathway are first incorporated into PC through acyl editing in 735

both the WT and the HO tobacco plants. (3) The high flux of nascent acyl groups 736

directly into PC acyl editing, and the initial labeled TAG regiochemical analysis both 737

indicate that a direct Kennedy pathway of TAG biosynthesis with nascent fatty acids is 738

not occurring in HO leaves. (4) In HO leaves acyl groups are redistributed from PC 739

mostly into TAG, rather than eukaryotic MGDG production as in the WT. (5) The 740

enhanced flux of fatty acids into TAG combined with the reduced flux of fatty acids into 741

both the prokaryotic and eukaryotic pathways of galactolipid synthesis reduced the 742

steady-state accumulation of MGDG and DGDG. (6) The pulse-chase did not indicate 743

TAG synthesis from galactolipid turnover. (7) Characterization of the high rates of PC 744

acyl editing in the HO line suggests that limiting PC acyl editing may be a future 745

engineering strategy to increase the monounsaturated fatty acid content of leaf derived 746

biofuels. 747

748

749



33

METHODS 750

751

Plant growth 752

753

For lipid mass analysis and metabolic labeling experiments, WT and HO tobacco 754

(Nicotiana tabacum) plants were grown in Percival E-41HO growth chambers set at 755

16/8 hours light/dark, 26/22°C, and fluorescent white light intensity at pot level across 756

the chamber was 300-400 µmol photons m-2 s-1. Pots were watered three times a week, 757

with one watering replaced by Peters 20/20/20 NPK fertilizer at 0.97 g/L once a week. 758

For microsomal assays, tobacco plants were grown in the glasshouse during summer 759

condition at 24°C/18°C for 16 hr/8 hr light/dark. 760

761

Chemicals and supplies 762

763

Unless specified all chemicals were purchased from Fisher Scientific 764

(www.fishersci.com), and solvents were at least HPLC grade. [14C]acetate sodium salt 765

50 mCi/mmol (American Radiolabeled Chemicals, Inc. St. Louis, MO). Glass TLC 766

plates: Analtech HL 250 µm, 20 x 20 cm. Liquid scintillation fluid: EcoScint Original 767

(National Diagnostics, Atlanta, GA). Lipase from Rhizomucor miehei and Phospholipase 768

A2 lipase from Apis mellifera (Sigma-Aldrich, St. Louis, MO). 769

770

Microsomal assays 771

772

Leaves were harvested from 66-day-old tobacco plants, and the microsomal 773

proteins were prepared as described (Zhou et al., 2013). Protein content of the 774

microsomal preparations was measured with BCA reagents (Pierce Chemical 775

Company) with BSA as a standard. The enzyme assay was essentially done as 776

described (Guan et al., 2014). Microsomal proteins (100 µg) were incubated at 30°C 777

with gentle shaking in 0.1 M Tris buffer pH 7.2 containing 4 mM MgCl2, 10 mg/ml BSA, 778

12.5 nmol of [14C]glycerol-3-phosphate (8000 dpm/nmol) and 25 nmol 18:1-CoA in a 779

final assay volume of 100 µL for 15 or 60 min. The assays were terminated by addition 780



34

of 250 µL of methanol/chloroform/acetic acid (50:50:1) (v/v/v), followed by extraction of 781

the lipids into a chloroform phase. The total lipids were separated on silica TLC plates 782

by developing with chloroform/methanol/acetic acid/water, 90:15:10:3 (v/v/v/v) to half-783

way of the plate to separate the polar lipids. After air drying for a few minutes, the plates 784

were redeveloped with hexane/diethyl either/acetic acid, 70:30:1 (v/v/v) to separate the 785

neutral lipids. Radioactive labels of 1000 dpm were spotted three times on each plate as 786

reference, before exposing to a phosphor image screen for overnight. The radioactivity 787

of each band was quantified with Fujifilm FLA-5000 Phosphor Imager. 788

789

Continuous pulse and pulse-chase [14C]acetate metabolic labeling 790

791

The continuous pulse metabolic labeling of WT and HO leaf disks was done for 792

3, 6, 10, 30, 120 minutes in triplicate, within 20 mM MES pH 5.5, 0.1X MS salts, 0.01% 793

Tween 20, and 1 mM [14C]acetate. Procedure: 10 mm diameter leaf disks were 794

collected from multiple plants (two WT and three HO) randomized across all horizontal 795

leaves. For each time point replicate, 12 disks were collected directly into 10 mL 796

incubation media (without [14C]acetate) in 100 mL beakers and placed in a 26°C water 797

bath under ~330 µmol photons m-2 s-1 white light with gentle shaking for 10 min to 798

equilibrate temperature. To start the labeling time course the media was removed and 799

replaced with 5 mL of incubation media with [14C]acetate. At each time point the media 800

was removed and the 12 leaf disks were placed into 85°C 2.5 ml isopropanol, 0.01% 801

(w/v) butylated hydroxytoluene for 10 min to quench metabolism. Each replicate time 802

course for each plant line used three 5 ml aliquots of 1 mM [14C]acetate labeling media. 803

The remaining [14C]acetate media after the 6 min time point was used for the 120 min 804

labeling, and the remaining media from the 10 min time point was used for the 30 min 805

labeling. The remaining [14C]acetate media from the 3 and 10/30 labeling time points 806

were mixed and used for the pulse-chase [14C]acetate labeling. 807

For each of the triplicate pulse-chase labeling time courses, 24 leaf disks were 808

collected as described above and pulsed with [14C]acetate labeling media independently 809

for 30 min. The 14C media was removed and the disks were washed three times (10 ml 810

each) in media without [14C]acetate and a final 10 mL media was added for chase 811



35

incubations. At each chase time point of 0, 2, 6, and 22 hours six leaf disks were 812

collected from each time course incubation and quenched as described above. 813

814

Lipid extraction and lipid class TLC separations 815

816

Lipids were extracted from isopropanol quenched tissue following a previous 817

method (Hara and Radin, 1978). After drying total extracts under N2, lipids of each 818

extract were dissolved in 0.5 ml toluene and aliquots were used for various analytical 819

procedures. Total 14C extracts were quantified by liquid scintillation counting on a 820

Beckman Coulter LS 6500 liquid scintillation counter. Neutral lipids were separated on 821

silica TLC plates in hexane/diethyl either/acetic acid, 70:30:1 (v/v/v). Polar lipids were 822

resolved with toluene/acetone/water (30/91/7, v/v/v) on silica TLC plates pre-treated 823

with 0.15 M ammonium sulfate and baked at 120°C for 3 hours prior to loading lipids. 824

Lipid classes were identified based on co-migration with standards. Relative 825

radioactivity of lipids separated by TLC was measured by phosphor imaging on a GE 826

Typhoon FLA7000, and ImageQuant analysis software. 827

828

Leaf lipid mass analysis 829

830

Leaf lipids extracted from 86-day-old plants were separated by TLC as described 831

above, and stained with 0.05% primulin in acetone/water 80:20 (v/v) and visualized 832

under UV light. Scraped bands were transmethylated along with a 17:0 TAG internal 833

standard of fatty acid methyl esters (FAMEs) in 2.5% (v/v) sulfuric acid in methanol at 834

80°C for 1 hr. FAMEs were collected into hexane by adding hexane and 0.88% (w/v) 835

NaCl to force a phase separation. FAMEs were separated and quantified by gas 836

chromatography with flame ionization detection on a Restek Stabilwax column: 30 m, 837

0.25 ID, 0.25 µm film thickness. 838

839

Regiochemical analysis of DAG, PC, and TAG 840

841



36

Total lipids extracted as described above from the WT and the HO line were co-842

loaded with 30 µg PC and 30 µg DAG. For the WT, 30 µg TAG was also co-loaded. 843

Polar lipid and neutral lipid TLC and primulin staining was performed as described 844

above. PC bands were scrapped off and eluted with chloroform/methanol/acetic acid 845

(5:5:1, v/v/v). Partial digestion of PC was performed with bee venom (Apis mellifera) 846

phospholipase PLA2 (Sigma) (Bates et al., 2007). The digested products were 847

separated by TLC in chloroform/methanol/acetic acid/water 50:30:8:4 (v/v/v/v). 848

Regiochemical analysis of neutral lipids were performed as described (Cahoon et al., 849

2006). DAG and TAG were digested with 0.2 ml of the Rhizomucor meihei lipase 850

(Sigma) for 30 and 60 min respectively. Digested products were separated by TLC in 851

hexane/diethyl ether/acetic acid (35:70:1, v/v/v). Lipid standards were stained with 852

iodine vapor and marked with 14C. Identification of unknowns was based on co-853

migration with standards. Radioactivity was quantified by using phosphor imaging as 854

described above. 855

856

Analysis of [14C]acetate labeled MGDG molecular species 857

858

MGDG was isolated by normal phase HPLC on an Agilent 1260 Infinity II 859

(quaternary pump, autosampler, column thermostat, DAD set to 210 nm, fraction 860

collector, running OpenLAB CDS Version C.01.09). The method is an adaption of 861

(Kotapati and Bates, 2018), with modifications as follows: injection volume 5-15 µL in 862

toluene; flow rate 1 ml/min; mobile phases (A: 2-Propanol, B: hexanes, C: Methanol, 863

D:25 mM triethylamine + 25 mM Formic acid (pH 4.1)). Linear gradients between steps 864

from: 0 min 19.3%A/80%B/0.5%C/0.20%D; 3 min 73.6%A/25%B/1%C/0.4%D; 6 min 865

87.5%A/10%B/1.5%C/1%D; 15 min 65%A/0%B/25%C/10%D; held for 3 min; 20 min 866

100%A; held for 3 min; 24 min is the starting composition, and equilibration between 867

samples is 10 min. MGDG was collected between 5.3 and 6.2 min. MGDG molecular 868

species were separated by HPLC on a Thermo Scientific Accucore C18 column (150 869

mm x 3mm; 2.6 µ particle size), according to (Yamauchi et al., 1982) except that the 870

flow rate was 0.35 ml/min for 35 minutes. Vial sampler was maintained at 20°C and the 871

column compartment at 35°C. Samples were injected in 8-15 µL methanol and 872



37

contained 5000-20000 CPM. To measure radioactivity the column eluent flowed into a 873

LabLogic β-Ram 6 flow liquid scintillation detector, flow cell volume set at 300 µL, 874

eluant:scintillation cocktail (LabLogic FloLogic-U) ratio was 1:3, with a residence time of 875

12.9 s. Laura 6.0.1.40 software was used to acquire and process the 14C data. To 876

confirm the identity of labeled molecular species, each fraction was collected, converted 877

to FAME as above, and separated by argentation TLC as in (Bates et al., 2009). 878

879

Data analysis 880

All calculations from raw data were done in Microsoft Excel. Graphing and 881

statistical analysis done with GraphPad Prism version 7.05. 882

883

Accession Numbers 884

AtWRI1, AT3G54320; AtDGAT1, AT2G19450; SiOLEOSIN; EU999158; AtFAD2, 885

AT3G12120; AtFAD3, AT2G29980; AtLPCAT1, AT1G12640; AtLPCAT2, AT1G63050; 886

AtPAH1, AT3G09560; AtPAH2, AT5G42870; AtPDAT1, AT5G13640; AtROD1, 887

AT3G15820; AtTGD1, AT1G19800. 888

889

Supplemental Data 890

Supplemental Figure S1. Stereochemical fatty acid composition of MGDG and DGDG 891

from WT and HO leaves. 892

Supplemental Figure S2. Total incorporation of [14C]acetate into WT and HO leaves. 893

Supplemental Figure S3. Initial incorporation of nascent [14C]acetate labeled fattyacids 894

into lipids, non-normalized. 895

Supplemental Figure S4. Example of 14C-MGDG molecular species analysis. 896

Supplemental Figure S5. Labeled MGDG molecular species from 30 and 120 minute 897

continuous [14C]acetate labeling. 898

Supplemental Figure S6. Labeled MGDG molecular species from 0 and 22 hours 899

pulse-chase [14C]acetate labeling. 900

901

ACKNOWLEDGEMENTS 902

We thank Bei Dong for technical support. 903



38

904

905

906

907

TABLES 908

909

PC PE PG MGDG DGDG DAG TAG

WT 18.59 ± 2.97

1.53 ± 0.17

0.31 ± 0.004

16.49 ± 1.79

0.28 ± 0.0001 2.08 ± 0.35 0.14 ± 0.01

HO 17.2 ± 1.7 1.04 ± 0.13

*0.16 ± 0.02

*3.36 ± 0.59

*0.11 ± 0.01 1.57 ± 0.37 *16.68 ±

2.83

p-value 0.7234 0.1444 0.0263 0.0199 0.0075 0.4212 0.0281

F.C. -1.9 -4.9 -2.5 119.1

910

Table 1. Initial rates of nascent fatty acid incorporation into individual lipid 911

classes. 912

Rates in DPM µchlorophyll-1 min-1 are the slope best fit ± SE from the linear regression 913

of the first 10 minutes of [14C]acetate labeling from Fig. 5. The p-values indicate if the 914

slopes are significantly different (p-value < 0.05), those that are significant are marked 915

with an asterisk. The fold change (F.C.) for lipids with significantly different rates are 916

indicated. 917

918

Samples: WT 30 min

HO 30 min

HO % of WT DPM

WT 120 min

HO 120 min

HO % of WT DPM

Ave. total DPM/µg chl 335.2 86.7 2066.4 646.6 Eukaryotic proportion

DPM/µg chl 9.6% 32.2

20.9% 18.1

56.3%

11.4% 234.5

25% 161.7

69%

Prokaryotic proportion DPM/µg chl

90.4% 303.0

79.1% 68.2

22.6%

88.6% 1831.9

75% 484.6

26.5%

919

Table 2. Acyl flux into eukaryotic and prokaryotic molecular species of MGDG 920

The average total MGDG DPM/µg chl at 30 and 120 min is from Fig. 5. The proportion 921

of eukaryotic and prokaryotic molecular species are from Supplemental Figure S5. The 922

DPM/µg chl of eukaryotic and prokaryotic MGDG molecular species are calculated from 923



39

the total label and the relative proportion of each. The “HO % of WT DPM” is the 924

amount of HO eukaryotic or prokaryotic MGDG as compared to the WT at each time 925

point. 926

927

928

929

930

Samples: WT 0 hr WT 22 hr 0-22 hr F.C. HO 0 hr HO 22 hr 0-22 hr

F.C. Eukaryotic proportion Prokaryotic proportion

12.8% 87.2%

38.6% 61.4%

3.0 0.7

20.4% 79.5%

31.5% 68.5%

1.5 0.86

931

Table 3. Change in MGDG eukaryotic and prokaryotic molecular species over the 932

[14C]acetate pulse-chase. The proportion of eukaryotic and prokaryotic molecular 933

species are from Supplemental Figure S6. F.C., fold-change. 934

935

Figure Legends 936

937

Figure 1. Model of leaf lipid pathways and hypotheses for acyl flux into TAG. 938

Plastid localized fatty acid synthesis (F.A.S.) and chloroplast localized prokaryotic 939

pathway are in the green box. All other reactions represent extra-plastidial metabolism. 940

Filled arrowheads represent flux of the glycerol backbone, open arrowheads represent 941

acyl transfer reactions. Dashed lines and boxes represent uncertainty in acyl flux in HO 942

tobacco lines, and large blue letters represent 3 hypotheses for altered acyl flux in HO 943

tobacco lines: A, the use of de novo DAG by AtDGAT1 for a Kennedy pathway TAG 944

synthesis; B, uncertain quantitative flux through acyl editing that affects incorporation of 945

PC-modified fatty acids in TAG; C, the use of a membrane lipid derived DAG by 946

AtDGAT1 for TAG synthesis. Abbreviations: DAG, diacylglycerol; DGDG, 947

digalactosyldiacylglycerol; G3P, glycerol-3-phosphate; LPA, lysophosphatidic acid; 948

MGDG; monogalactosyldiacylglycerol; PA, phosphatidic acid; PC, phosphatidylcholine; 949

PE, phosphatidylethanolamine; PG, phosphatidylglycerol; TAG, triacylglycerol. 950

951



40

Figure 2. Accumulation of lipids in WT and HO leaves. 952

The abundance of polar membrane lipids and neutral lipids in leaves of 86-day-old 953

plants. A, mass abundance of each lipid. B, ratio of mass abundance of lipid to PC. WT, 954

blue. HO, red. Data is average ± SEM for 3-4 replicates. Significant differences in the 955

HO line from the WT (Student’s t-test, p-value < 0.05) are marked with an asterisk. 956

Abbreviations: DAG, diacylglycerol; DGDG, digalactosyldiacylglycerol; MGDG; 957

monogalactosyldiacylglycerol; PC, phosphatidylcholine; PE, phosphatidylethanolamine; 958

PG, phosphatidylglycerol; PI, phosphatidylinositol; PS, phosphatidylserine; TAG, 959

triacylglycerol. 960

961

Figure 3. Fatty acid composition of WT and HO leaf lipids. 962

Weight percent fatty acid composition of lipid classes isolated from leaves of 86-day-old 963

plants. A, total leaf extract. B, TAG. C, PC. D, MGDG. Abbreviations as in text. WT, 964

blue. HO, red. Data is average ± SEM for 3-4 replicates. Significant differences in the 965

HO line from the WT (Student’s t-test, p-value < 0.05) are marked with an asterisk. 966

Abbreviations: PC, phosphatidylcholine; MGDG, monogalactosyldiacylglycerol; TAG, 967

fatty acids, # carbons: # double bonds, d#, delta double bond position. 968

969

Figure 4. In vitro assay of the Kennedy pathway in leaf microsomes. 970

Assay conditions: 100 ug of leaf microsomal protein, 12.5 nmole [14C]G3P + 25 nmole 971

of 18:1-CoA, incubated for 15 or 60 min. For each line and assay length, data is the 972

average and SEM of 3 replicates. Abbreviations: DAG, diacylglycerol; LPA, 973

lysophosphatidic acid; PA, phosphatidic acid; MAG, monoacylglycerol; TAG, 974

triacylglycerol. 975

976

Figure 5. Initial incorporation of nascent [14C]acetate labeled fatty acids into 977

lipids. 978

A-B, major labeled lipids from continuous [14C]acetate labeling for 3-120 min in leaf 979

disks of 66-day-old plants of the WT and the HO line. C-D, an expanded view of the 0-980

35 min portion of A & B. All data points are average and SEM of 3 biological replicates. 981

Abbreviations: DAG, diacylglycerol; DGDG, digalactosyldiacylglycerol; MGDG; 982



41

monogalactosyldiacylglycerol; PC, phosphatidylcholine; PE, phosphatidylethanolamine; 983

PG, phosphatidylglycerol; TAG, triacylglycerol; DPM, disintegrations per minute; chl, 984

chlorophyll. 985

986

987

Figure 6. Relative incorporation of nascent fatty acids into the eukaryotic and 988

prokaryotic pathways of WT and HO leaves. A, the WT. B, the HO line. The 989

endoplasmic reticulum (ER) localized lipids (PC, PE, TAG) and the plastid localized 990

lipids (PG, MDGD, DGDG) from Fig. 5 were added together, and the ratio of the two. 991

DAG was not included because it can be localized to multiple compartments. All data 992

points are average and SEM of 3 biological replicates. Abbreviations: DPM, 993

disintegrations per minute; chl, chlorophyll. 994

995

Figure 7. Regiochemical analysis of initial incorporation of nascent [14C]acetate 996

labeled fatty acids into DAG, PC, and TAG. A-B, TAG lipase digestion of DAG from 997

the WT and the HO line. C-D, phospholipase A2 digestion of PC from the WT and the 998

HO lines. E-F, TAG lipase digestion of TAG from the WT and the HO line. All data 999

points are average and SEM of 3 biological replicates. In A-D, statistical significance of 1000

the HO line from the WT (Student’s t-test, p-value < 0.05) is marked with an asterisk at 1001

each data point. Abbreviations: DAG, diacylglycerol; FFA, free fatty acid; MAG, 1002

monoacylglycerol; PC, phosphatidylcholine; TAG, triacylglycerol. 1003

1004

Figure 8. Pulse-Chase [14C]acetate accumulation in lipids of the WT and the HO 1005

line. In all panels the pulse starts at -0.5 hours, and the chase starts at 0 hours. The 1006

leaves of 73-day-old plants were used for both plants. A-B, the WT and the HO line 1007

demonstrating the total 14C incorporated into the lipid extract; the endoplasmic reticulum 1008

(ER) localized lipids (PC, PE, TAG) and the plastid localized lipids (PG, MDGD, DGDG) 1009

were added together, and the ratio of the two. DAG was not included because it can be 1010

localized to multiple compartments. All data points are average and SEM of 3 biological 1011

replicates. Abbreviations: DPM, disintegrations per minute; chl, chlorophyll. 1012

1013



42

1014

Figure 9. Pulse-Chase [14C]acetate labeling of acyl flux through the lipid metabolic 1015

network. In all panels the pulse starts at -0.5 hours, and the chase starts at 0 hours. 1016

The leaves of 73-day-old plants were used for both plants. A-B, accumulation of 1017

individual radiolabeled lipids as DPM/µg chlorophyll from the total labeled samples in 1018

Fig. 8. C-D, the labeled lipids in A-B, represented as a percentage of the sum. All data 1019

points are average and SEM of 3 biological replicates. Abbreviations: DAG, 1020

diacylglycerol; DGDG, digalactosyldiacylglycerol; MGDG; monogalactosyldiacylglycerol; 1021

PC, phosphatidylcholine; PE, phosphatidylethanolamine; PG, phosphatidylglycerol; 1022

TAG, triacylglycerol; DPM, disintegrations per minute; chl, chlorophyll. 1023

1024

1025

1026

1027



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Bates PD (2016) Understanding the control of acyl flux through the lipid metabolic network of plant oil biosynthesis. Biochimica etBiophysica Acta (BBA) - Molecular and Cell Biology of Lipids 1861: 1214-1225


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Copyright © 2019 American Society of Plant Biologists. All rights reserved.

https://www.ncbi.nlm.nih.gov/pubmed?term=Allen DK %282016%29 Assessing compartmentalized flux in lipid metabolism with isotopes%2E Biochimica et Biophysica Acta %28BBA%29 %2D Molecular and Cell Biology of Lipids&dopt=abstract

https://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Allen&hl=en&c2coff=1

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https://www.ncbi.nlm.nih.gov/pubmed?term=Allen DK %282016%29 Quantifying plant phenotypes with isotopic labeling %26 metabolic flux analysis%2E Current opinion in biotechnology&dopt=abstract


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https://www.ncbi.nlm.nih.gov/pubmed?term=Allen DK%2C Bates PD%2C Tjellstr�m H %282015%29 Tracking the metabolic pulse of plant lipid production with isotopic labeling and flux analyses%3A Past%2C present and future%2E Progress in Lipid Research&dopt=abstract


https://scholar.google.com/scholar?as_q=Tracking the metabolic pulse of plant lipid production with isotopic labeling and flux analyses: Past, present and future&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

https://scholar.google.com/scholar?as_q=Tracking the metabolic pulse of plant lipid production with isotopic labeling and flux analyses: Past, present and future&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Allen&as_ylo=2015&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Arisz SA%2C Heo J%2DY%2C Koevoets IT%2C Zhao T%2C van Egmond P%2C Meyer J%2C Zeng W%2C Niu X%2C Wang B%2C Mitchell%2DOlds T%2C Schranz ME%2C Testerink C %282018%29 Diacylglycerol acyltransferase 1 contributes to freezing tolerance%2E Plant Physiology&dopt=abstract

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https://www.ncbi.nlm.nih.gov/pubmed?term=Bafor M%2C Smith MA%2C Jonsson L%2C Stobart K%2C Stymne S %281991%29 Ricinoleic acid biosynthesis and triacylglycerol assembly in microsomal preparations from developing castor%2Dbean %28Ricinus%2Dcommunis%29 endosperm%2E Biochemical Journal&dopt=abstract

https://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Bafor&hl=en&c2coff=1

https://scholar.google.com/scholar?as_q=Ricinoleic acid biosynthesis and triacylglycerol assembly in microsomal preparations from developing castor-bean (Ricinus-communis) endosperm&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

https://scholar.google.com/scholar?as_q=Ricinoleic acid biosynthesis and triacylglycerol assembly in microsomal preparations from developing castor-bean (Ricinus-communis) endosperm&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Bafor&as_ylo=1991&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Barron EJ%2C Stumpf PK %281962%29 Fat metabolism in higher plants%2E XIX%2E The biosynthesis of triglycerides by avocado%2Dmesocarp enzymes%2E Biochim Biophys Acta&dopt=abstract

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https://scholar.google.com/scholar?as_q=Fat metabolism in higher plants&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

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https://www.ncbi.nlm.nih.gov/pubmed?term=Eastmond PJ%2C Quettier A%2DL%2C Kroon JTM%2C Craddock C%2C Adams N%2C Slabas AR %282010%29 PHOSPHATIDIC ACID PHOSPHOHYDROLASE1 and 2 Regulate Phospholipid Synthesis at the Endoplasmic Reticulum in Arabidopsis%2E The Plant Cell Online&dopt=abstract

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https://www.ncbi.nlm.nih.gov/pubmed?term=Hajduch M%2C Hearne LB%2C Miernyk JA%2C Casteel JE%2C Joshi T%2C Agrawal GK%2C Song Z%2C Zhou M%2C Xu D%2C Thelen JJ %282010%29 Systems analysis of seed filling in Arabidopsis%3A using general linear modeling to assess concordance of transcript and protein expression%2E Plant physiology&dopt=abstract

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https://scholar.google.com/scholar?as_q=Mutant alleles of FAD2-1A and FAD2-1Bcombine to produce soybeans with the high oleic acid seed oil trait.&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Pham&as_ylo=2010&as_allsubj=all&hl=en&c2coff=1

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https://www.ncbi.nlm.nih.gov/pubmed?term=Vanhercke T%2C Divi UK%2C El Tahchy A%2C Liu Q%2C Mitchell M%2C Taylor MC%2C Eastmond PJ%2C Bryant F%2C Mechanicos A%2C Blundell C%2C Zhi Y%2C Belide S%2C Shrestha P%2C Zhou XR%2C Ral JP%2C White RG%2C Green A%2C Singh SP%2C Petrie JR %282017%29 Step changes in leaf oil accumulation via iterative metabolic engineering%2E Metab Eng&dopt=abstract

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https://scholar.google.com/scholar?as_q=Step changes in leaf oil accumulation via iterative metabolic engineering&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

https://scholar.google.com/scholar?as_q=Step changes in leaf oil accumulation via iterative metabolic engineering&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Vanhercke&as_ylo=2017&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Vanhercke T%2C Dyer JM%2C Mullen RT%2C Kilaru A%2C Rahman MM%2C Petrie JR%2C Green AG%2C Yurchenko O%2C Singh SP %282019%29 Metabolic engineering for enhanced oil in biomass%2E Progress in Lipid Research&dopt=abstract


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https://scholar.google.com/scholar?as_q=Metabolic engineering for enhanced oil in biomass&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Vanhercke&as_ylo=2019&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Vanhercke T%2C El Tahchy A%2C Liu Q%2C Zhou X%2DR%2C Shrestha P%2C Divi UK%2C Ral J%2DP%2C Mansour MP%2C Nichols PD%2C James CN%2C Horn PJ%2C Chapman KD%2C Beaudoin F%2C Ruiz%2DL�pez N%2C Larkin PJ%2C de Feyter RC%2C Singh SP%2C Petrie JR %282014%29 Metabolic engineering of biomass for high energy density%3A oilseed%2Dlike triacylglycerol yields from plant leaves%2E Plant Biotechnology Journal&dopt=abstract


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https://scholar.google.com/scholar?as_q=Metabolic engineering of biomass for high energy density: oilseed-like triacylglycerol yields from plant leaves&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Vanhercke&as_ylo=2014&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Vogel C%2C Marcotte EM %282012%29 Insights into the regulation of protein abundance from proteomic and transcriptomic analyses%2E Nature reviews genetics&dopt=abstract

https://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Vogel&hl=en&c2coff=1

https://scholar.google.com/scholar?as_q=Insights into the regulation of protein abundance from proteomic and transcriptomic analyses.&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

https://scholar.google.com/scholar?as_q=Insights into the regulation of protein abundance from proteomic and transcriptomic analyses.&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Vogel&as_ylo=2012&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Wang L%2C Shen W%2C Kazachkov M%2C Chen G%2C Chen Q%2C Carlsson AS%2C Stymne S%2C Weselake RJ%2C Zou J %282012%29 Metabolic Interactions between the Lands Cycle and the Kennedy Pathway of Glycerolipid Synthesis in Arabidopsis Developing Seeds%2E The Plant Cell Online&dopt=abstract

https://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Wang&hl=en&c2coff=1

https://scholar.google.com/scholar?as_q=Metabolic Interactions between the Lands Cycle and the Kennedy Pathway of Glycerolipid Synthesis in Arabidopsis Developing Seeds&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

https://scholar.google.com/scholar?as_q=Metabolic Interactions between the Lands Cycle and the Kennedy Pathway of Glycerolipid Synthesis in Arabidopsis Developing Seeds&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Wang&as_ylo=2012&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Williams JP%2C Imperial V%2C Khan MU%2C Hodson JN %282000%29 The role of phosphatidylcholine in fatty acid exchange and desaturation in Brassica napus L%2E leaves%2E Biochemical Journal&dopt=abstract

https://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Williams&hl=en&c2coff=1

https://scholar.google.com/scholar?as_q=The role of phosphatidylcholine in fatty acid exchange and desaturation in Brassica napus L&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

https://scholar.google.com/scholar?as_q=The role of phosphatidylcholine in fatty acid exchange and desaturation in Brassica napus L&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Williams&as_ylo=2000&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Wood CC%2C Okada S%2C Taylor MC%2C Menon A%2C Mathew A%2C Cullerne D%2C Stephen SJ%2C Allen RS%2C Zhou XR%2C Liu Q %282018%29 Seed�?�specific RNAi in safflower generates a superhigh oleic oil with extended oxidative stability%2E Plant biotechnology journal&dopt=abstract

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https://scholar.google.com/scholar?as_q=Seedâ�?�specific RNAi in safflower generates a superhigh oleic oil with extended oxidative stability&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

https://scholar.google.com/scholar?as_q=Seedâ�?�specific RNAi in safflower generates a superhigh oleic oil with extended oxidative stability&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Wood&as_ylo=2018&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Xu C%2C Shanklin J %282016%29 Triacylglycerol Metabolism%2C Function%2C and Accumulation in Plant Vegetative Tissues%2E Annual Review of Plant Biology&dopt=abstract

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https://scholar.google.com/scholar?as_q=Triacylglycerol Metabolism, Function, and Accumulation in Plant Vegetative Tissues&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

https://scholar.google.com/scholar?as_q=Triacylglycerol Metabolism, Function, and Accumulation in Plant Vegetative Tissues&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Xu&as_ylo=2016&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Yamauchi R%2C Kojima M%2C Isogai M%2C Kato K%2C Ueno Y %281982%29 Separation and Purification of Molecular Species of Galactolipids by High Performance Liquid Chromatography%2E Agricultural and Biological Chemistry&dopt=abstract

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https://scholar.google.com/scholar?as_q=Separation and Purification of Molecular Species of Galactolipids by High Performance Liquid Chromatography&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

https://scholar.google.com/scholar?as_q=Separation and Purification of Molecular Species of Galactolipids by High Performance Liquid Chromatography&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Yamauchi&as_ylo=1982&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Yang W%2C Wang G%2C Li J%2C Bates PD%2C Wang X%2C Allen DK %282017%29 Phospholipase Dzeta Enhances Diacylglycerol Flux into Triacylglycerol%2E Plant Physiol&dopt=abstract

https://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Yang&hl=en&c2coff=1

https://scholar.google.com/scholar?as_q=Phospholipase Dzeta Enhances Diacylglycerol Flux into Triacylglycerol&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

https://scholar.google.com/scholar?as_q=Phospholipase Dzeta Enhances Diacylglycerol Flux into Triacylglycerol&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Yang&as_ylo=2017&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Zhou XR%2C Shrestha P%2C Yin F%2C Petrie JR%2C Singh SP %282013%29 AtDGAT2 is a functional acyl%2DCoA%3Adiacylglycerol acyltransferase and displays different acyl%2DCoA substrate preferences than AtDGAT1%2E FEBS Lett&dopt=abstract

https://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Zhou&hl=en&c2coff=1

https://scholar.google.com/scholar?as_q=AtDGAT2 is a functional acyl-CoA:diacylglycerol acyltransferase and displays different acyl-CoA substrate preferences than AtDGAT1&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

https://scholar.google.com/scholar?as_q=AtDGAT2 is a functional acyl-CoA:diacylglycerol acyltransferase and displays different acyl-CoA substrate preferences than AtDGAT1&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Zhou&as_ylo=2013&as_allsubj=all&hl=en&c2coff=1

https://www.ncbi.nlm.nih.gov/pubmed?term=Zou JT%2C Wei YD%2C Jako C%2C Kumar A%2C Selvaraj G%2C Taylor DC %281999%29 The Arabidopsis thaliana TAG1 mutant has a mutation in a diacylglycerol acyltransferase gene%2E Plant Journal&dopt=abstract

https://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Zou&hl=en&c2coff=1

https://scholar.google.com/scholar?as_q=The Arabidopsis thaliana TAG1 mutant has a mutation in a diacylglycerol acyltransferase gene&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

https://scholar.google.com/scholar?as_q=The Arabidopsis thaliana TAG1 mutant has a mutation in a diacylglycerol acyltransferase gene&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Zou&as_ylo=1999&as_allsubj=all&hl=en&c2coff=1


short title: leaf oil production rearranges membrane lipid flux … · 5 104 may be possible...

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