# A comparison of natural and induced diversity in plant oils

Thierry Chardot

<figure><img src="image_1.png" /></figure>

## To cite this version:

Thierry Chardot. A comparison of natural and induced diversity in plant oils. OCL Oilseeds and fats crops and lipids, 2022, 29, pp.34. 10.1051/ocl/2022027. hal-04072697

HAL Id: hal-04072697

https://hal.inrae.fr/hal-04072697

Submitted on 18 Apr 2023

HAL is a multi-disciplinary open access archive for the deposit and dissemination of scientific research documents, whether they are published or not. The documents may come from teaching and research institutions in France or abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est destinée au dépôt et à la diffusion de documents scientifiques de niveau recherche, publiés ou non, émanant des établissements d'enseignement et de recherche français ou étrangers, des laboratoires publics ou privés.

<figure><img src="image_2.png" /></figure>

Distributed under a Creative Commons Attribution 4.0 International License
# A comparison of natural and induced diversity in plant oils

Thierry Chardot*

Université Paris-Saclay, INRAE, AgroParisTech, Institut Jean-Pierre Bourgin (IJPB), 78000 Versailles, France

Received 20 April 2022 – Accepted 5 September 2022

**Abstract** – Currently, there is a growing demand to replace the compounds in a given product that are of a petroleum origin with renewable resources. One of these compounds, called fatty acid (FA), is the main component of vegetable oils. FA composition is not only responsible for the physicochemical properties of plant oils, but it also determines their uses. For example, since time immemorial, products containing lipids have been used for lighting and heating purposes. They are also excellent lubricants and possess drying properties important molecules for painting, and wood preservation. In terms of nutrition, they have a high-energy content, are part of our daily health requirements, and are used for animal feed. We present here some lipids of interest, the plants that produce them naturally with high yield, the enzymes responsible for their synthesis when known, and their possible uses, as well as resources and ways that could allow the lipids of interest to be produced in quantity in different hosts.

**Keywords:** diversity / oleaginous plants / seeds / seed oil / biosynthesis / fatty acids

**Résumé** – Une comparaison de la diversité naturelle et induite dans les huiles végétales. Actuellement, il existe une demande croissante pour remplacer les composés d'origine pétrolière d'un produit donné par des ressources renouvelables. Depuis des temps immémoriaux, les produits contenant des lipides sont utilisés pour l'éclairage et le chauffage. Ils sont également d'excellents lubrifiants, possèdent des propriétés siccatives importantes pour la peinture, la préservation du bois. En termes de nutrition, ils ont un contenu énergétique élevé, font partie de nos besoins quotidiens en matière de santé ainsi que pour l'alimentation humaine et animale. Nous présentons ici quelques lipides d'intérêt, les plantes qui les produisent naturellement avec un rendement élevé, les enzymes responsables de leur synthèse lorsqu'elles sont connues, leurs utilisations possibles, ainsi que les ressources et les moyens qui pourraient permettre de produire en quantité les lipides d'intérêt dans différents hôtes.

**Mots clés** : diversité / plantes oléagineuses / graines / huile de graines / biosynthèse / acides gras

## 1 Introduction

World production of vegetable oils (2019–2020 crop year) is estimated at more than 200 million T<fnref n="1" />, 74% of which is used mostly for human nutrition and 5% for animals, with the remaining 20% used for energy purposes or chemistry. The plant kingdom, rich with more than 350 000 angiosperm species, represents an immense reservoir of unexploited natural product diversity, but only a very small number of plant species (palm, soybean, rapeseed, sunflower, coconut, etc.) are cultivated, and their oil is extracted mainly from fruits or seeds. Only a few

dozen among the hundreds of different FAs produced and stored in plant oils are exploited for different uses.

During the last 50 years, databases specifically dedicated to plant lipids have been developed. The lipid content of thousands of vegetable oils can be scrutinized using websites providing easy access to multiple databases. One online scientific database is Cyberlipid<fnref n="2" /> whose purpose is to collect, study and diffuse information on all aspects of lipidology. Seed oil fatty acids, or SOFA<fnref n="3" />, the oldest database and hosted by the Max Rubner-Institute (Karlsruhe, Germany), is a collection of the FA composition of wild plant seeds from the appropriate

<footnote>* Contribution to the Topical Issue “Minor oils from atypical plant sources / Huiles mineures de sources végétales atypiques”.</footnote>

<footnote>*Correspondence: thierry.chardot@inrae.fr</footnote>

<footnote n="1">1 http://www.statista.com/</footnote>

<footnote n="2">2 http://cyberlipid.gerli.com/</footnote>

<footnote n="3">3 https://plantfadb.org/</footnote>
pharmaceutical, botanical and chemical literature (Aitzetmüller et al., 2003). This database was the first to permit exploration of about 580 different FA structures, the composition from more than 7000 plant species, and lists about 130 000 individual percentage data for FA occurring in plant seeds. PlantFAdb<fnref n="4" /> (previously PhyloFAdb) (Ohlrogge et al., 2018) provides updates and enhancements to SOFA, and allows users to search and display FA composition data for over 9000 plants. It also includes more than 17 000 tables from more than 3000 publications and hundreds of unpublished analyses. All these databases represent valuable and unique tools to explore FA diversity in plants, chemotaxonomic relationships between FA structures, and plant species by displaying these relationships on dynamic phylogenetic trees (Aitzetmüller et al., 2003; Ohlrogge et al., 2018). The PlantFAdb catalogs just over 2% of the estimated species in the plant kingdom, with representative seed compositions for most phylogenetic clades available; thus a number of FAs remains to be discovered in seeds (Cahoon and Li-Beisson, 2020). Included in this database are also relatively ancient sets of data. Practical resources for exploring the diversity of plant lipids are also provided by valuable books (Gunstone et al., 1986; Ucciani, 1994).

In the following sections, we review a selection of FAs and focus our attention on the plants that contain the most FAs, as well as the enzymatic systems involved in their synthesis, when known. At the end, we give examples of oils from plants, some of which are already cultivated, and which have unique properties due to their content in specific FAs.

We also present examples of plants whose oil production has been modified by modern genome techniques such as gene expression modification and genome editing (CRISPR Cas9). As a complement to modern techniques we end by suggesting some possible ways to better exploit biodiversity, and try to identify some underlying challenges.

## 2 Biosynthesis and accumulation of oils with FAs of interest: from enzymes to plants that produce them naturally and in large quantity

Sugars are synthesized by plants through photosynthesis and converted into a variety of storage compounds, including oils (Baud, 2018). The main compounds of oil are FAs which are esterified to a glycerol skeleton. In plants, oil synthesis requires the collaboration of different metabolic pathways and subcellular compartments. The *de novo* synthesis of FAs requires malonyl-coenzyme A (CoA) which is the carbon donor, an acyl bound to an acyl carrier protein (ACP), and various cofactors. FA synthesis occurs in the plastids and is carried out by an enzyme complex, with four monofunctional proteins, termed fatty acyl synthase (type II FAS), which catalyzes cycles of successive reactions. It begins with a Claisen condensation between the acyl-ACP and a malonyl-ACP (3C, product from pyruvate oxidative decarboxylation) catalyzed by three β-ketoacyl-ACP synthases isoforms:

KASIII up to 4C-ACP, KASI up to 16C-ACP, and KASII catalyzes the last elongation up to 18C-ACP. The addition of two C requires the participation of three additional enzymes.

3-ketoacyl-ACP reductase (KAR) catalyzes a ketone hydrogenation, 3-hydroxyacyl-ACP dehydratase (HAD) catalyzes water elimination and enoyl-ACP reductase (ENR) catalyzes the hydrogenation of the alkene. Each synthesis cycle uses one malonate (3C) and results in the elongation of acyl-ACP by 2C (Baud and Lepiniec, 2010; Li-Beisson et al., 2010). Simultaneously in the chloroplast, elongation of FAs occurs until stearic acid (18:0) (Fig. 1.1). Stearoyl-ACP desaturase (SAD) desaturates stearic acid (18:0) bound to ACP into oleic acid (18:1Δ⁹) bound to ACP. SAD, a Δ9 acyl-ACP desaturase (Fig. 1.2), is a plastid-localized non-membrane-bound soluble desaturase which contains a di-iron center (Lindqvist et al., 1996). Similarly, Δ9 acyl-ACP desaturase transforms palmitic acid (16:0) bound to ACP into palmitoleic acid (C16:1Δ⁹) bound to ACP (Fig. 1.3). Acyl-ACP thioesterases from TE14 family (Swarbrick et al., 2020), located at the inner plastid envelope membrane, hydrolyze the newly formed acyl-ACP into free FA and ACP (Fig. 1.4). Free FAs, the chain length for which is dictated by enzyme specificity of acyl-ACP thioesterases, are transported through the membrane and are activated into the form of CoA esters by long-chain acyl-CoA synthetases (LACS) (Fig. 1.5) and transferred to the endoplasmic reticulum (ER) by acyl-CoA binding proteins (Fig. 1.6), feeding the eukaryotic pathway responsible for the formation of phospholipids, triglycerides, and other neutral lipids.

Different enzymatic systems further modify FAs through catalyzing sequential reactions in the ER, using FA belonging to different metabolic pools, such as acyl-CoA, phosphatidyl-choline (PC), or GL as substrates. FAD2 (Δ12 desaturase) transforms oleic acid bound to PC into linoleic acid (18:2Δ⁹c,12c) bound to PC (Fig. 1.7) (Zeng et al., 2017). FAD3 catalyzes the desaturation of linoleic acid bound to PC (18:2Δ⁹c,12c) acid into 18:3Δ⁹c,12c,15c bound to PC (Fig. 1.8) (Arondel et al., 1992). FADX, a fatty acid conjugase desaturase and a divergent form of Δ12 oleate desaturase FAD2, are responsible for the synthesis of the conjugated eleostearic acid (18:3Δ⁹c,11t,13t') (Fig. 1.9) (Dyer et al., 2002). Structurally, unrelated enzymes that have evolved separately are cytochrome P450 and delta 12 epoxygenase (Δ12 desaturase-like). These enzymes are capable of catalyzing the synthesis of 12-epoxy vernolic acid (Δ12,13-O-18:1Δ⁹c) by using PC-bound 18:3Δ⁹c,12c,15c as substrate (Fig. 1.10) (Cahoon et al., 2002).

Homologs of FAD2 are also capable of hydroxylating the Δ12 position of PC bound oleic acid (van de Loo et al., 1995), producing ricinoleic acid (Δ12-OH-18:1Δ⁹c) (Fig. 1.11). An activated form of ricinoleic acid, ricinoleoyl-CoA enters a discontinuous elongation process in which the 3-hydroxy C20 intermediate (not shown) is elongated into nebraskanic (7,18-(OH)2-24:1Δ¹⁵) by a specialized FAE (Li et al., 2018) (Fig. 1.12).

The synthesis of very long chain fatty acids (VLCFAs, longer than 18C), is catalyzed in the ER by the fatty acid elongase (FAE) complex which uses malonyl-CoA and long-chain acyl-CoA as substrates. The pool of long chain acyl-CoA is probably resulting from exchange of FAs between the acyl-

<footnote n="4">⁴ https://plantfadb.org/</footnote>
<figure><img src="image_3.png" /><figcaption>Fig. 1.</figcaption></figure>
<figure><img src="image_4.png" /><figcaption>Fig. 1. (Continued)</figcaption></figure>
<figure><img src="image_5.png" /><figcaption>**Fig. 1.** Overview of biosynthesis, accumulation, and deposition of selected fatty acids in plants.</figcaption></figure>

**Panel A:** In plants, synthesis of neutral lipids requires the collaboration of plastids and the endoplasmic reticulum while their accumulation occurs in ER derived organelles called lipid droplets, found in all organs. Blue arrows indicate flux toward neutral lipid accumulation in organelles and selected organs. **Panel B:** The synthesis and elongation of FAs, bound to ACP occur in the chloroplast till 18:0 (1, fatty acyl synthase) and its desaturation into 18:1<sup>9c</sup> by 9 stearoyl-ACP desaturase (SAD) (2). In a similar way, 16:0 desaturation by 9 palmitoyl-ACP desaturase (PAD) (3) produces palmitoleic acid (16:1<sup>9</sup>), which can be further elongated to produce cis vaccenic acid 18:1<sup>11c</sup>, (1). The synthesis of Acyl-ACP can be interrupted Acyl-ACP thioesterases (4) releasing free FA with chain lengths under 18C. **Panel C:** Free FAs are transported through the membrane, activated into the form of CoA esters by long-chain acyl-CoA synthetases (LACS) (5) and transferred to the endoplasmic reticulum (ER) by acyl-CoA binding proteins (6), where they are further processed. Oleic acid elongation and the other FA modifications take place mostly in the ER and involve different enzymes (elongases, hydroxylases, desaturases) which use acyl bound to ACP, glycerolipids (GL), PC and acyl-CoA as substrates. Oleic acid bound to PC or GL is transformed into linoleic acid (7, 12 desaturase). FAD3 catalyzes the desaturation of linoleic (18:2<sup>9c,12c</sup>) acid into linolenic (18:3<sup>9c,12c,15c</sup>) (8). FADX, a fatty acid conjugase desaturase is responsible for the synthesis of eleostearic acid (18:3<sup>9c,11t,13t</sup>) (9). Vernolic acid (12,13-O-18:1<sup>9c</sup>) is obtained upon the action of cytochrome P450 on 5 PC-bound linolenic acid or of delta 12 epoxygenase (12 desaturase like) (10). Hydroxylation of the 12 position of PC bound oleic acid (fatty acid hydroxylase) by homologs of fatty acid desaturase 2 (FAD2) (11) produces ricinoleic acid (12-OH-18:1<sup>9c</sup>). Ricinoleoyl-CoA, deriving from Ricinoleoyl-PC, is transformed into nebraskanic (7,18-(OH)2-24:1<sup>-15</sup>) acid upon the action of hydroxylases (12). Elongation of oleoyl-CoA by acyl-CoA elongase (13) produces eicosenoic (20:1<sup>11c</sup>) and erucic acid (22:1<sup>13c</sup>). **Panel D:** Esterification of FA to glycerol for accumulation into lipid droplets. Dihydroxyacetone phosphate (DHAP) originates from glycolysis and is converted to glycerol-3-phosphate (G3P) by glycerol-3-phosphate dehydrogenase (G3PDH) (14). G3P is acylated at *sn*1 positions by a glycerol phosphate acyltransferase (GPAT) (15) which produces lysophosphatidic acid (LPA). Acylation at *sn*2 position by a 1-acyl G3P acyltransferase (16) produces phosphatidic acid (PA). The dephosphorylation of PA by phosphatidic acid phosphatases (PAP) (17) produces 1,2-diacyl-*sn*-glycerol (DAG). Another acyl group is esterified at the *sn*-3 position of DAG either by acyl-CoA:1,2-diacyl-*sn*-glycerol acyltransferase (DGAT, acetyl-CoA dependent pathway) (18), or by the phospholipid:1,2-diacyl-*sn*-glycerol acyltransferase (PDAT) (19) which transfers acyl from PC to DAG. For the sake of clarity, we have focused our attention on the FAs which are exemplified in this review. For more detailed and comprehensive descriptions of FA metabolism in model plants and crops, please refer to the following very recent articles (Baud, 2018; Cahoon and Li-Beisson, 2020; Miklaszewska et al., 2021).
CoA and PC pools due to the action of lysophosphatidylcholine acyltransferase (LPCAT) (Stahl et al., 2008). FAE catalyze reactions similar to those occurring during *de novo* FA synthesis in the plastids (Haslam and Kunst, 2013). FAE which encodes the β-ketoacyl-CoA synthase is responsible for elongation of oleic acid (18:1) activated under the form of CoA ester to eicosenoic acid (20:1) (Fig. 1.13) and further to erucic acid (22:1) (Fig. 1.13) (Barret et al., 1998; Li et al., 2012).

The enzymatic esterification of FA to various molecules containing OH groups (glycerol, sterols, alcohols) leads to four different compounds: phospholipids, sterol esters, triacylglycerols (TAG), and waxes. Phospholipids are mostly used to build up membranes, while sterol esters and TAG are stored in specialized organelles called lipid droplets (Huang, 2018), and waxes which cover the epidermal cells.

TAG synthesis results from successive acylations of glycerol-3-phosphate (G3P). The conversion of dihydroxyacetone phosphate (DHAP), originating from glycolysis to glycerol-3-phosphate (G3P) is catalyzed by glycerol-3-phosphate dehydrogenase (G3PDH) (Fig. 1.14). G3P is sequentially acylated at *sn*1 and *sn*2 positions by a glycerol phosphate acyltransferase (GPAT) (Shockey et al., 2016) (Fig. 1.15) which produces lysophosphatidic acid (LPA), and by lysophosphatidic acyltransferase (LPAAT) (Maisonneuve et al., 2010) (Fig. 1.16) which produces phosphatidic acid (PA). The dephosphorylation of PA by phosphatidic acid phosphatases (PAP) (Fig. 1.17) produces 1,2-diacyl-*sn*-glycerol (DAG). Another acyl group is esterified at the *sn*-3 position of DAG either by acyl-CoA:1,2-diacyl-*sn*-glycerol acyltransferase (DGAT, acetyl-CoA dependent pathway) (Fig. 1.18), or by the phospholipid:1,2-diacyl-*sn*-glycerol acyltransferase (PDAT) (Fig. 1.19). which transfers acyl from PC to DAG to catalyze the production of TAG in the acetyl-CoA-independent pathway (Zhang et al., 2009).

WRINKLED1 (WRI1) transcription factor is encoded by a gene belonging to the APETALA transcription factor family. While it has been found that mutant *wri1* can cause an 80% reduction in seed oil content (Focks and Benning, 1998), recently, WRI1 has been shown to be a key regulator of genes involved in late glycolysis and FA biosynthesis in *Arabidopsis thaliana* seeds, and also in crops (Baud et al., 2007; Maeo et al., 2009; Li et al., 2015).

Among the more than 450 FA known in the plant kingdom, five FAs are termed usual, palmitic acid (16:0), stearic acid (18:0), oleic acid (18:1Δ⁹c), linoleic acid (18:2Δ⁹,¹²c), and α-linolenic acid (18:3Δ⁹,¹²,¹⁵c) (Li-Beisson et al., 2013). These are found in biological membranes, so mostly in phospholipids. The vast majority of FAs are stored in triacylglycerol (Cahoon and Li-Beisson, 2020), in plant polymers such as cutin and in waxes, and represent most of the diversity of these molecules, and are termed unusual. The asymmetric distribution of usual and unusual FAs between the different compartments without a doubt reflects the strong selection pressure exerted by evolution on biological membranes. The structure of FAs includes the number of carbon atoms in the acyl chain, and the presence of various modifications (unsaturation, hydroxyl-ation, epoxidation, and cyclopropane groups). This structure governs the physical-chemical properties of FAs, and gives them very different reactivities, which makes them unique, renewable raw materials for chemistry (Metzger, 2009).

Table 1 details the chemical formula, together with some properties and uses of selected usual and unusual FAs found in large quantities in the oil of cultivated and non-cultivated plants.

## 2.1 Lauric acid (12:0)

Palm kernel oil contains high amounts of octanoic (8:0), capric acid (10:0), and lauric acid which are extracted from the endosperm of the palm oil (up to 38.7%) (Dussert et al., 2013). Lauric acid in its sulfated form, is an ingredient found in many household products (toothpaste, shampoos, shaving foams, and bubble baths) (Vandeputte, 2012). Its global market was estimated at ~5 million T in 2015⁵, and strongly depends on the cultivation of palm trees. The accumulation of FAs with defined alkyl chain length is under the control of acyl carrier protein (ACP) thioesterases (Jones et al., 1995). These enzymes release free FAs, thus terminating alkyl chain elongation, and permitting esterification of the glycerol skeleton. Large amounts of octanoic, capric and lauric acid (up to 94.4, 93.8, and 85.9) are found in the oils of different cuphea shrub species. Cuphea is a *Lythraceae* common in tropical countries (Graham and Kleiman, 1992). A specialized diacylglycerol acyltransferase contributes to the extreme medium-chain FA content of cuphea oil (Iskandarov et al., 2017). However, despite the report of 93.8% capric content in seeds from *Cuphea avigera var pulcherrima* (Graham and Kleiman, 1992), the cultivation of this plant remains impaired by poor yield, seed shattering, and indeterminate growth (Zanetti et al., 1993).

## 2.2 Myristic acid (14:0)

*Pycnanthus Kombo* (African nutmeg) is a Magnoliaceae which can reach 15–20 m in size. Its drupes are rich in oil (up to 58%), with a high myristic acid content (14:0) (61.6%) (Banerji et al., 1984), making it an important source of oil and wax for fabricating candles, soap, fuel, and lubricants (PROTA)<fnref n="6" /> (van der Vossen and Mkamilo, 2007).

## 2.3 Palmitic acid (16:0)

Native to the coast and coastal plains of southeastern North America, the fruit of the shrub *Myrica carolinensis* accumulates up to 29% oil rich in palmitic (77.5%) and myristic (21.5%) acids (Harlow et al., 1965). Different derivatives of palmitate are used as emollients or plasticizers, and can be used to fabricate candles.

Thus, plants belonging to different botanical families capable of producing oils highly enriched in non-saturated FAs with up to (16:0) are found in nature. They represent alternatives to palm trees, and do not seem to be cultivated in large areas, or even domesticated. Key enzymes involved in the accumulation of specific FAs in these plants have, to our knowledge, not always been identified.

<footnote n="5">⁵ https://www.transparencymarketresearch.com/sodium-lauryl-sulfate-market.html</footnote>

<footnote n="6">⁶ https://www.prota4u.org/database/</footnote>
<table><caption>**Table 1.** Fatty acids of interest cited in this manuscript, some uses, and plants showing the highest content in their oil.</caption><thead><tr><th>Fatty acid common name</th><th>Nomenclature</th><th>Plant with highest known content (%)</th><th>Family</th><th>Uses</th><th>Reference</th></tr></thead><tbody><tr><td>Octanoic</td><td>8:0</td><td>Cuphea pulcherrima (94.4%)</td><td>Lythraceae</td><td>Perfume, antifungal</td><td>(Graham and Kleiman, 1992)</td></tr><tr><td>Capric</td><td>10:0</td><td>Cuphea koehne (93.8%)</td><td>Lythraceae</td><td>Flavors, perfumes, dyes</td><td>(Graham and Kleiman, 1992)</td></tr><tr><td>Lauric</td><td>12:0</td><td>Actinodaphne hookeri (96%)</td><td>Lauraceae</td><td>Detergent</td><td>(Banerji et al., 1984)</td></tr><tr><td>Myristic</td><td>14:0</td><td>Pycnathus kombo (61.6%)</td><td>Myristicaceae</td><td>Candles, soap, fuel, lubricants</td><td>(Banerji et al., 1984)</td></tr><tr><td>Palmitic acid</td><td>16:0</td><td>Myrica carolinensis (77.5%)</td><td>Myricaceae</td><td>Plasticizer</td><td>(Harlow et al., 1965)</td></tr><tr><td>Palmitoleic</td><td>16:1<sup>Δ9</sup></td><td>Doxhanta unguis (&gt; 60%)</td><td>Bignoniaceae</td><td>Production of octene</td><td>(Nguyen et al., 2010)</td></tr><tr><td>Stearic</td><td>18:0</td><td>Momordica charantia (74.5%)</td><td>Cucurbitaceae</td><td>Candles, soap, detergents</td><td>(Dave et al., 1985)</td></tr><tr><td>Vaccenic</td><td>18:1<sup>Δ11c</sup></td><td>Entandraphragma (62%)</td><td>Meliaceae</td><td>Production undecanoic acid (11:0, Nylon production)</td><td>(Kleiman and Payne-Wahl, 1984)</td></tr><tr><td>Vernolic</td><td>Δ12,13-O-18:1<sup>Δ9c</sup></td><td>Vernonia galamensis (79-81%)</td><td>Asteraceae</td><td>Adhesives, varnish, paintings, solvent</td><td>(Ucciani, 1994)</td></tr><tr><td>Ricinoleic</td><td>Δ12-OH-18:1<sup>Δ9c</sup></td><td>Ricinus communis (90%)</td><td>Euphorbiaceae</td><td>Polymers</td><td>(Gunstone et al., 1986)</td></tr><tr><td>Eleostearic</td><td>18:3<sup>Δ9c,11t,13t</sup></td><td>Vernicia fordii (75.2%)</td><td>Euphorbiaceae</td><td>Varnishes</td><td>(Zhang et al., 2014)</td></tr><tr><td>Linolenic</td><td>18:3cis-Δ<sup>9,12,15</sup></td><td>Fragaria vesca (43%)<br/>Linum usitatissimum (35-66%)<br/>Euphorbia niciciana (74.3%)<br/>Vitis vinifera (63-75.7%)</td><td>Rosaceae<br/>Linaceae<br/>Euphorbiaceae<br/>Vitaceae</td><td>Food, paintings, plastics</td><td>(Johansson et al., 1997; Ucciani, 1994)</td></tr><tr><td>Erucic</td><td>22:1<sup>Δ13c</sup></td><td>Crambe abyssinica (55-60%)</td><td>Brassicaceae</td><td>Emollients, tensio actifs</td><td>(Ortiz et al., 2020)</td></tr><tr><td>Nebraskanic</td><td>7,18-(OH)2-24:1<sup>Δ15</sup></td><td>Orychophragmus</td><td>Brassicaceae</td><td>Lubricant</td><td>(Romsdahl et al., 2019)</td></tr><tr><td>Wuhanic acid</td><td>7,18-(OH)2-24:2<sup>Δ15c,21c</sup></td><td>violaceus (38%)</td><td>Brassicaceae</td><td></td><td></td></tr></tbody></table>

## 2.4 Stearic acid (18:0)

Momordica charantia, a Cucurbitaceae, also called bitter gourd, accumulates up to 30% oil with a high stearic acid (18:0) content (74.5%) (Dave et al., 1985). The performance of this plant cultivated in Asia, Africa and in the Caribbean is remarkable when compared to sunflowers possessing two mutations. One of these is Es1 (the sunflower stearoyl-ACP desaturase [SAD named SAD17]), which cannot accumulate more than 50% stearate (Salas et al., 2021), and its oil must be refined before use.

## 2.5 Oleic acid (18:1<sup>Δ9c</sup>)

Oleic acid (18:1<sup>Δ9c</sup>) results from the desaturation of stearic acid (18:0) by Δ9 stearoyl-ACP desaturase, a chloroplastic enzyme (Fig. 1). Several examples of oleic acid uses for synthesis of chemicals are given in (Metzger, 2009), and high oleic sunflower has been commercialized since 1984 (Purdy, 1986).

FAs may be further elongated and modified by specific enzymes, mostly in the ER and contributes to the oil's useful properties for various uses. In the following sections, we detail selected examples of oils enriched in FA obtained after modifications of oleic acid (18:1<sup>Δ9c</sup>).

## 2.6 Erucic acid (22:1<sup>Δ13c</sup>)

Erucic acid is a long chain unsaturated FA accumulated in high erucic acid rape (HEAR), and its synthesis is under the control of the BnFAE1 gene (Brassica napus fatty acid elongase). This gene encodes the β-ketoacyl-CoA synthase responsible for the elongation of oleic acid (18:1) to eicosenoic acid (20:1) and further to (22:1) (Barret et al., 1998; Li et al., 2012). Erucic amide, a compound commonly used as a slip additive in the plastic industry, results from the condensation of erucic acid with ammonia. Cross-pollination of HEAR with food quality rapeseed (i.e., canola that does not produces erucic acid) could result in the production of oil with altered
quality. Thus, these two plants must be distantly cultivated. *Crambe abyssinica*, a Brassicaceae, also naturally produces an oil with a high erucic acid content (Samarappuli *et al.*, 2020) and could be an alternative crop.

The use of a gene stacking strategy is necessary to achieve a substantial increase in the amount of erucic acid in Crambe seed oil. First, it involves the expression of *Limnanthes douglasii* lysophosphatidic acid acyltransferase (LPAAT) (Cao *et al.*, 1990) to compensate for the inability of the endogenous LPAAT to use erucoyl-CoA as an acyl donor. A second gene is expressed, *B. napus* BnFAE1 (fatty acid elongase), which encodes the β-ketoacyl-CoA synthase, and is responsible for elongation of FAs from oleic acid (18:1Δ⁹c) to eicosenoic acid (20:1Δ¹¹c) and further to erucic (22:1Δ¹³c). In the end, FAD2 encoding a fatty acid desaturase catalyzing the desaturation of oleic (18:1Δ⁹c) to linoleic acid (18:2Δ⁹c,¹²c), is down-regulated (Li *et al.*, 2012). Taken together, these modifications lead to a decrease of oleic, linoleic and linolenic (18:3Δ⁹,¹²,¹⁵c) acid and permits a substantial increase in erucic acid (22:1Δ¹³c), from 55-60% in the wild type seeds (Ortiz *et al.*, 2020) to 73% in the best *C. abyssinica* transgenic line obtained by the authors (Li *et al.*, 2012).

## 2.7 Ricinoleic acid (Δ12–OH–18:1Δ⁹c)

Castor seeds (*Ricinus communis*) contain 39–59% of an oil rich (83–90%) in ricinoleic acid (Δ12–OH–18:1Δ⁹c) (Gunstone *et al.*, 1986). Castor fatty acid 12-hydroxylase (RcFAH12) (Fig. 1), a homolog of (FAD2) is the enzyme that hydroxylates the Δ12 position of PC bound oleic acid (van de Loo *et al.*, 1995), producing ricinoleic acid. Castor oil has for a long time been used as lamp oil, an ingredient for cosmetics, as a purgative, and more recently as a lubricant (Dumeignil, 2012). Castor oil also serves as the basis for the synthesis of many compounds, such as polyamide 11 (Rilsan), an agro-based bioplastic that is used in the hoses of the automotive industry (Winnacker and Rieger, 2016), and was classified as a strategic material critical to the US national defense by the Agricultural Materials Act P.L. 98-284 passed by Congress in 1984. Despite the industrial interest in castor oil, the poisons contained in the plant, as well as in many *Euphorbiaceae*, still represent a roadblock to its cultivation. The plant is not really cultivated in Europe; however approximately 1 443 588 T of castor seeds were produced in 2019 in India, the world’s largest producer (82% of world production).

## 2.8 Vernolic acid (Δ12,13–O–18:1Δ⁹c)

The seeds of various Asteraceae, such as *Vernonia galamensis* a plant native to East Africa, *Stokesia laevis*, native to the USA, or *Crepis biennis*, a plant commonly found in France, produce oils enriched (79–81%) in an epoxy FA called vernolic acid (Δ12,13–O–18:1Δ⁹c) (Ucciani, 1994). The seeds of *Euphorbia lagascae*, a Euphorbiaceae, also accumulate in their oil up to 63% vernolic acid (Cahoon *et al.*, 2002), due to the action of a cytochrome P450 on PC-bound linoleic acid (Cahoon *et al.*, 2002). *S. laevis* epoxidase exhibits high homology with *V. galamensis* epoxygenase, and transforms linoleic acid into vernolic acid (Hatanaka *et al.*, 2004). The genetic transformation of soybean by the *S. laevis* epoxidase gene together with *V. galamensis* diacylglycerol acyltransferase permits accumulation of up to 17.6% vernolic acid in the seeds (Li *et al.*, 2010).

The high content in the epoxy groups (68% vernolic acid) of vernonia oil permits its use in the manufacturing of industrial adhesives, varnishes, paints, and coatings (Baye *et al.*, 2005).

## 2.9 Linolenic acid (18:3Δ⁹,¹²,¹⁵c)

Flax seeds contain 35–50% of an oil rich in linolenic acid (18:3Δ⁹,¹²,¹⁵c) (35–66%) that is responsible for its drying properties. Two genes (LuFAD3A, LuFAD3B) that encode a flax seed microsomal desaturase capable of desaturating linoleic acid into linolenic have been identified and characterized (Vrinten *et al.*, 2005) (Fig. 1). Historically, oxidized flax seed oil was used for the synthesis of well-known linoleum flooring. In 2014, 2.6 million tons of flax seeds were produced worldwide, with 16 000 T produced in France alone. This country imports these seeds, mostly for food purposes (Labalette *et al.*, 2011). Seeds belonging to other botanical families may also contain significant amounts of linolenic acid. The performance of *Euphorbia niciciana*, not cultivated yet, seems to exceed that of flax, with similar to higher seed oil (39–44%) or linolenic acid (74–76%) content (SOFA) (Ucciani, 1994). As already stated; Euphorbiaceae notoriously accumulate toxic compounds in seeds and vegetative tissues, hindering their exploitation. Strawberry (Fragaria, Rosaceae) seeds represent around 1% of the fruit weight (Grzelak-Blaszczyk *et al.*, 2017) and their oil contains up to 43% linolenic acid (Johansson *et al.*, 1997). Strawberry production was 12 106 585 T in 2019⁷, and, so more than 121 065 T seeds were potentially available and could provide 52 058 T of linolenic acid. Grape seeds are rich in linolenic acid (63–75.7%) (Ucciani, 1994). California generates about 435 449 T/year of grape pomace, and a recent techno-economic assessment estimates the annual production of 1627 T of grape seed oil (Jin *et al.*, 2021).

## 2.10 Eleostearic acid (18:3Δ⁹c,¹¹t,¹³f)

*Vernicia fordii* tree, a Euphorbiaceae tree accumulates in its seeds an oil called tung oil. Native to Asia, it was introduced to South America, Thailand, and the southern United States, where due to climatic hazards, cultivated areas have fallen sharply (Zhang *et al.*, 2014). The consecutive action of two desaturases (FAD2 and FADX) converts PC-bound oleic acid into linolenic and eleostearic acid (18:3Δ⁹c,¹¹t,¹³f), a conjugated trans trienoic acid (Dyer *et al.*, 2002) (Fig. 1). Tung oil is easily oxidized due to its very high content in eleostearic acid, up to 75% (Zhang *et al.*, 2014), and is commonly used in formulations of inks, dyes, coatings, and resins because of its drying properties (Sonntag, 1979). 588 190 T of tung nuts were produced worldwide, mostly by China and Brazil (50% of 2019 world production, FAO). Production of oeleostearatic acid is not limited to Euphorbiaceae. As an example, *Momordica charantia*, a Cucurbitaceae is also capable of accumulating up to 56% eleostearic acid (Ucciani, 1994) in its seeds. As the plant is cultivated in the tropical Asian region, precise statistics about its cultivation are not available (PROTA). More examples of plants accumulating eleostearic acid have been reported (Hennessy *et al.*, 2016), such as *Punica granatum* (pomegranate), and *Fevillea trilobata* (*cucurbitale*).

<footnote n="7">7 http://www.fao.org/faostat/en/</footnote>
**3 Lipid accumulation in organs other than seeds**

Seeds and fruits are not the only organs that accumulate lipid. The use of tubercules from *Cyperus Esculentus* as food has been discovered in ancient Egypt (Vega-Morales et al., 2019). It has also been suggested that there were even earlier uses by hominids who lived in East Africa between 2.4–1.4 million years ago, who survived mainly on a diet based on grasses and Cyperaceae sedges (Macho, 2014). *C. Esculentus* produces an oil (17–29%) (Wang et al., 2020) rich in oleic acid (58.8%) (Ucciani, 1994) and is cultivated in several countries (Vega-Morales et al., 2019). *Tetraena mongolica* Maxim is a Zygophyllaceae found in inner Mongolia (China). Due to the high TAG content of the vegetative parts, and as the stems contain about 46 mg TAG/g dry matter (Wang et al., 2007), this plant was used as firewood.

**4 Plants producing oils with low viscosities**

As already stated, the oils used for food, health, chemistry, and energy are extracted from a very small number of plants when compared to existing diversity. The vast majority of oils have been characterized with analysis of FAs derivatives (generally methyl esters) using gas chromatography. The number of FAs contained in these oils is very small in comparison to the usual and unusual FAs, and the positional distribution of FAs on the glycerol skeleton remains largely unknown, except for oil from crops.

The use of classic TLC or more elaborate nuclear magnetic resonance techniques allows for the detection of unknown FA or new esterified FA that could provide an explanation for the low viscosity of oil. Members of the genus Lesquerella, are Brassicaceae known to be rich in hydroxy FAs. The analysis of TAG structure revealed the presence of molecules containing two or more acyl groups joined via ester linkages between an –OH moiety on the hydrocarbon chain of one acyl group and the –COOH moiety of another acyl group, termed estolides (Hayes et al., 1995). *Orychophragmus violaceus* (Chinese violet Cress) is a Brassicaceae originating from China used for ornamental purposes. The seed oil was believed to be rich in linoleic acid, until the discovery of two major components, C24 FAs containing hydroxyl groups at carbon atoms 7 and 18, and the deciphering of a new pathway for the biosynthesis of hydroxyl FAs (Li et al., 2018). A premature or “discontinuous” elongation of a 3–OH intermediate by a divergent 3-ketoacyl-CoA synthase during a chain extension cycle explains the origin of the presence of dihydroxy FAs in *O. violaceus* (nebraskanic and wuhanic acids) (Li et al., 2018) (Fig. 1). *O.s violaceus* oil contains up to 38% nebraskanic and wuhanic acids, and its coefficient of friction for the sliding steel surfaces at 100°C is three times lower in comparison with castor oil (Li et al., 2018). The biosynthetic basis for estolides in *O. violaceus* seeds remains unknown (Cahoon and Li-Beisson, 2020).

The seed of the plant *Euonymus alatus*, a flowering shrub in the Celastraceae family (also called winged spindle or burning bush), contains around 44% oil (Earle, 1966) composed of more than 90% 3-acetyl-1,2-diacylglycerol

(also termed acetyl TAGs). *E. alatus* accumulates oil specifically in the albumen and the embryo of seeds. This plant grows mostly in gardens for ornamental purposes, and the question of its industrial cultivation has not yet been considered. Developments in nucleic acid sequencing techniques allowed for the identification of an acyltransferase that is both expressed in the genesis of *E. alatus* albumen (Durrett et al., 2010) and involved in accumulation of acetyl TAGs. Acetyl TAGs possess a low viscosity and can be used either directly as fuel or as lubricants of renewable origin.

**5 Transformation of plants**

Many plants that synthesize interesting oils possess characteristics that are unfavorable to their cultivation (low yield, presence of toxins, some are invasive species). Crop engineering offers solutions to overcome these issues.

**5.1 Achieve competitive yields**

Crops modified by transgenic techniques can accumulate oils that approach the level and quality of native plants. The efficient accumulation of oils with a given composition relies on the expression of enzymes synthesizing specific FAs, as well as on the availability of acyl transferases and acyl acceptor molecules. The biosynthesis and accumulation of lipids requires the collaboration of several metabolic pathways and branches, as well as different compartments and even possibly TAG remodeling. Because these requirements are complex, efficient lipid engineering is very difficult (Bates, 2016; Bhandari and Bates, 2021). Only a small number of studies have shown that modifying the expression of a single gene is enough to significantly improve the oil content and nature of seeds. To achieve this goal, additional specific genes usually need to be expressed in order to both synthesize FAs and to accumulate them efficiently in target organs.

Leaves represent targets for increasing oil accumulation using the concepts of FA synthesis (“Push”), TAG assembly (“Pull”), and lipid turnover (“Protect”) (Vanhercke et al., 2014). The next section presents successful examples of this concept.

The synthesis of TAGs requires a glycerol backbone, acyl acceptor on which the FAs will be esterified. Increasing the availability of reaction precursors will increase the final yield of TAG synthesis. Glycerol 3-phosphate dehydrogenase allows the transformation of dihydroxyacetone phosphate into glycerol 3-phosphate (G3P) (Push), the first substrate for TAG synthesis. Expression of foreign acyl carrier protein (ACP) thioesterases modifies the availability of specific FAs for esterification on the glycerol backbone and these enzymes are also targeted (Push). WRINKLED1 (WRI1), transcription factors are involved in transcriptional regulation for adapting the rate of acyl chain production to cell requirements (Push) (Vanhercke et al., 2014).

DGATs are classified into three distinct classes, sharing no sequence homology, and probably result from convergent evolution. DGAT1 and 2 types are integral membrane proteins of the endoplasmic reticulum (Stone et al., 2006; McFie et al., 2010). In *A. thaliana*, AtDGAT1 makes a major contribution to seed oil content (Routaboul et al., 1999). In crops producing
non-edible oils, members of the DGAT2 family incorporate
unusual FAs containing hydroxy- (Ricinus communis) and
epoxy groups (V. galamensis) into seed TAGs (Kroon et al.,
2006; Li et al., 2010). DGAT3 are soluble proteins (Saha et al.,
2006; Chi et al., 2014) and the *A. thaliana* isoform possesses a
[2Fe–2S] cluster (Ayme et al., 2018). Due to the slow velocity
of the catalyzed reaction, DGAT has been regarded as a
limiting step in TAG accumulation at the time of high lipid
synthesis during oil accumulation and seed maturation in
*B. napus* (Perry and Harwood, 1993). Efficient incorporation
of unusual FAs into TAG not only requires enzymes specific
for their production but also the expression of DGATs, which
selectively use them (Bates et al., 2014). Recent results
obtained upon expression of one DGAT3 isoform from
*Camelina sativa* demonstrated the high DGAT activity of the
enzyme upon expression in leaves and its preference for
unsaturated FAs (Gao et al., 2021). This makes DGAT3 a
promising candidate for increasing both oil yield (Pull) and
quality in plants. DGATs from different families thus represent
a reservoir of enzymes allowing the incorporation of specific
FAs into TAGs.

Two different research teams have overexpressed yeast
glycerol 3-phosphate dehydrogenase (gdp1) in different plants,
alone or in combination with DGAT1 from *A. thaliana* (Pull).
In oilseed rape, overexpression of gdp1 increased the amount
of G3P available for acylation by successive acyltransferases,
resulting in a 40% increase in the lipid content of the mature
seed (Vigeolas et al., 2007). Oleosins are the major proteins
found at the surface of LDs, which are oil storage organelles.
The role of oleosins in lipid accumulation (Protect) was
demonstrated in seeds (Siloto et al., 2006), and more recently
in leaves (Zhai et al., 2021). The co-expression of three genes
involved in different aspects of TAG production; WRI1 (Push),
DGAT1 (Pull), and OLEOSIN (Protect) permitted the
accumulation of more than 15% TAG (dry weight basis) in
*Nicotiana tabacum* leaves (Vanhercke et al., 2014). More
details about TAG metabolism and accumulation in plant
vegetative tissues are extensively detailed by Xu and Shanklin
(2016).

Seeds from *C. sativa* co-expressing the *A. thaliana* DGAT1 and yeast *gdp1* accumulate up to 13% higher oil content and up to 52% higher seed mass compared to wild-type plants (Chhikara *et al.*, 2018). Similarly, co-expression of specific acyltransferases and acyl-ACP thioesterases (Wiberg *et al.*, 1997; Iskandarov *et al.*, 2017) in Brassicaceae improved the accumulation of medium-chain saturated FAs in the seeds.

Plant oils containing ω-7 FAs have potential as sustainable feedstocks. However, plants with oil containing a very high content (> 60%) of palmitoleic acid (*Doxhanta unguis* or *Kermadecia sinuate*) are not cultivated and even considered as invasive (*D. unguis*). Using a strategy, consisting of strongly suppressing KASII (elongase activity) and increasing the desaturase activity in the host plant, two teams were able to obtain plants that accumulate high quantities of ω-7 FAs in seeds. Amounts are around 2–3% in *A. thaliana* and *C. Sativa* wild-type plants. Transgenic *A. thaliana* accumulate up to 71% ω-7 FAs, (levels equivalent to those found in Doxhanta seeds (71.9%) (Nguyen *et al.*, 2010) and transgenic *C. sativa* 44.3% (Rodriguez-Rodriguez *et al.*, 2021).

*Euonymus alatus* diacylglycerol acetyltransferase (*EaDAcT*) was cloned and expressed in different plants

(Arabidopsis, Camelina, Soya). As a consequence, it was
found that wild type *A. thaliana* plants expressing *EaDAcT*
accumulated 45% acetyl-TAGs in their oil (Durrett *et al*.,
2010). Expression of *EaDAcT* in plants affected the expression
of DGAT1, the enzyme limiting TAG accumulation, permitting
the accumulation of up to 70% acetyl TAG in camelina and
soybean. In transgenic camelina, grown in the field, with the
silenced DGAT1 gene and expressing *EaDAcT*, it was found
that the oil contained up to 85% acetyl TAG (Liu *et al*., 2015a).

5.2 Accounting for undesired effects

Seeds store lipidic compounds in their reserves, which act
as energetic molecules and/or building blocks for the
establishment of autotrophic plantlets. The modification of
the nature and quantities of accumulated lipids can affect the
fate of seeds sometimes with unexpected or even deleterious
effects of which we give some examples.

Transgenic rice plants engineered to overproduce linoleic
acid have been developed by different research groups. Plants
overexpressing delta-12 fatty acid desaturase have unexpec-
tedly shown enhanced cold tolerance during the reproductive
stage and an increase of grain yield (46%) under cold
conditions (Shi et al., 2012). Similarly, overexpression in rice
of an ω-3 fatty acid desaturase from *Glycine max* (GmFAD3A)
has been shown to enhance the total polyunsaturated FAs
(PUFAs) content in seeds, as well as seed germination rate at
low temperature (Wang et al., 2019).

Although inactivation of rapeseed stearoyl-acyl-desaturase
has resulted in increased production of stearic acid in Brassica
seeds (up to 40%) compared to conventional varieties, the
transgenic seeds have shown poor germination rates (Knutzon
et al., 1992). A similar situation was observed in *A. thaliana*
seeds where the high amounts of stearate (up to 25%) were
negatively linked to germination performance (Kazaz et al.,
2020). In genetically transformed *Brassica juncea*, a negative
correlation between seed stearate content (up to 31%) and seed
germination performance was also observed (Bhattacharya
et al., 2015).

6 GM crops improved for oil traits

6.1 Cultivated

GM crops have been cultivated in the field for more than
twenty years. In 2018 the surface planted with these crops was
184 million hectares total (Brookes and Bargfoot, 2020),
mostly located in different countries outside of the European
Union. In 2016, most GM crops grown had traits that allowed
them to resist herbicides or insect (Napier et al., 2019);
however, there are other aspects of oil crops to consider.

Because of their composition, oils can have beneficial or
deleterious effects for consumers, whether they are humans or
animals. Coriander seed oil (CSO) is highly enriched in
petroselenic acid, and in a scientific opinion, the European
Food Safety Authority (EFSA) panel concluded that the novel
food ingredient, CSO, was safe under the proposed uses and
use levels (Agostoni et al., 2016) as a food supplement for
healthy adults. Since the presence of erucic acid in rapeseed
oils can be harmful to the heart muscle, varieties with little or
no erucic acid have been developed, and EFSA has issued a
scientific opinion on the daily intake level for erucic acid (Knutsen et al., 2016).

A search performed using the International Service for the Acquisition of Agri-biotech Applications (ISAAA) website<fnref n="8" /> indicates that at least three crop plants (B. napus, Carthamus tinctorius, Glycine max) with traits improved for oil amount or composition using GM techniques are commercially available. It should be noted that modified plants can accumulate FAs that they do not naturally synthesize. For example, B. napus expressing *Umbellularia californica* 12:0-ACP thioesterase are intended for palmitic acid production. When considering the accumulation of DHA, seven foreign genes (desaturases and elongases) are expressed in other events. To achieve the production of docosapentaenoic acid (C22:5n-3), a FA not naturally found in plants, the introduction of ten foreign genes was necessary. Modifications of *Carthamus tinctorius* L. are available as well, in which the synthesis of delta-12 desaturase enzyme is suppressed by RNA interference, or in which the production of FATB enzymes or acyl-acyl carrier protein thioesterases are suppressed by RNA interference.

## 6.2 Camelina, an emerging crop, capable of synthesizing and accumulating non-plant FA

Given that some modified plants have already been successfully grown in the field, *C. sativa* represents an excellent example of a plant for which seed oil can be improved using modern techniques (transformation, gene edition).

*C. sativa* is a hexaploid and possesses three delta-12-desaturase (FAD2) genes. Using similar approaches based on CRISPR-Cas9 gene editing, (Morineau et al., 2017; Jiang et al., 2017) performed selectively targeted mutation of the three FAD2 genes. Both teams obtained transgenic Camelina lines with lipid profiles, ranging from 10% to 62% oleic and 16% to over 50% oleic acid in their seeds. High oleic acid content was associated with a decrease of polyunsaturated FAs.

Camelina plants expressing *EaDAcT* were grown in the field and produced up to 70 mol% acetyl-dioleoyl-glycerol. The overall features (seed weight, oil content, seed yield, and harvest index) of camelina accumulating acetyl-TAG were not modified (Liu et al., 2015b). A camelina construct was also designed to accumulate in its seeds eicosapentaenoic acid (EPA) and docosahexaenoic acid (DHA), FAs not naturally present in plants. Camelina has now been evaluated in the field, in separate geographic and regulatory locations (the UK, US, and Canada). Its ability to grow in the field, accumulating EPA and DHA only in its seeds confirms the promise of this modified plant as a new source of omega-3 rich oils normally found only in marine organisms (Han et al., 2020).

## 7 From the rationale exploration to the uses of biodiversity

Compared to other organs, seeds store the widest repertoire of FAs in their oil, and the number of accumulated FA species certainly exceeds the 580 found in the SOFA database.

The plants listed in PlantFAdb (over 9000) represent a small fraction of the plant kingdom (over 350 000 different species), even if most phylogenetic clades have representative seed compositions. Thus, a large number of unusual FAs and their assemblies into glycerolipids remains to be discovered in seeds, and even in other organs. Further esterifications of acyl chains can lead to networks of FAs, increasing the number of molecules potentially produced and stored in oils in a combinatorial manner. Approaches using combinations of liquid chromatography (LC) and mass spectrometry (MS) are very efficient and allow for the identification of many molecules. In addition to the already cited examples of the detection of nebraskanic and wuhanic acids in (Li et al., 2018), an analysis of the leaf lipidome has also yielded 393 molecular species within 23 different lipid classes (Tarazona et al., 2015) that cannot be deduced from the FA composition alone. A very recent article (Gan et al., 2022) report on the exploration of Thunbergia genus, which revealed that numerous species accumulate petroselenic (18:1Δ⁶) instead of sapienic acid (16:1Δ⁶). Thus, detailed study of a genus may reveal the accumulation of unexpected FAs, resulting from probable evolutionary divergence in the genus.

The databases cited here represent useful tools to identify plant species storing unusual FAs. The 1000 plants (oneKP or 1KP) initiative<fnref n="9" /> is an international multi-disciplinary consortium that has generated large-scale gene sequencing data for over 1000 species of green plants (Leebens-Mack et al., 2019). The exploitation of this sequence database permits the exploration of genomes to identify candidates' genes involved in the synthesis of these FAs and their assembly in the form of neutral lipids. To efficiently use these databases, difficulties arise in linking and integrating the different levels of data (genomic, transcriptomic, lipidomic) that can be used to identify the genes or metabolic pathways involved in the biosynthesis of specific FAs for further metabolic engineering. Many enzymes that allow the accumulation of original FAs in seed oils remain to be discovered that would allow these compounds to be accumulated in plants whose metabolic pathways have been altered.

Biotechnology approaches, including gene editing, have led to convincing and fairly rapid results in terms of modification of the FA composition of oils. In all cases, these approaches should not be excluded from the exploration and the better use of existing plants, whether cultivated or found in existing commercial collections, botanical gardens, or even in the wild.

## Conflict of interest

The author declares that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

## Funding

The IJPB benefits from the support of Saclay Plant Sciences-SPS (ANR-17-EUR-0007).

<footnote n="8">⁸ https://www.isaaa.org/</footnote>

<footnote n="9">⁹ https://sites.google.com/a/ualberta.ca/onekp/</footnote>
*Acknowledgments*. The author warmly thanks Dr Sébastien Baud, Dr Anne-Marie Le-Coq, Dr Betty Cottyn and Dr Philippe Guerche (INRAE IJPB) as well as Dr Radka Koncitikova (INRAE BBF Marseille) for their helpful discussions during the redaction of the manuscript. With thanks to Melissa-Ann Thomas, the Academic Writing Center, Université Paris-Saclay for proofreading this article.

## References

Agostoni C, Berni Canani R, Fairweather-Tait S, et al. 2016. Scientific opinion on the safety of “coriander seed oil” as a novel food ingredient. *EFSA J*. https://doi.org/10.2903/j.efsa.2013.342.

Aitzetmüller K, Matthäus B, Friedrich H. 2003. A new database for seed oil fatty acids – The database SOFA. *Eur J Lipid Sci Technol* 105: 92–103.

Arondel V, Lemieux B, Hwang I, et al. 1992. Map-based cloning of a gene controlling omega-3 fatty acid desaturation in Arabidopsis. *Science* 258(5086): 1353–5. https://doi.org/10.1126/science.1455229.

Ayme L, Arragain S, Canonge M, et al. 2018. Arabidopsis thaliana DGAT3 is a [2Fe-2S] protein involved in TAG biosynthesis. *Sci Rep* 8(1): 17254. https://doi.org/10.1038/s41598-018-35545-7.

Banerji R, Chowdhury AR, Misra G, Nigam SK. 1984. Butter from plants. *Fette Seifen Anstrichmittel* 86: 279–284. https://doi.org/10.1002/lipi.19840860706.

Barret P, Delourme R, Renard M, et al. 1998. A rapeseed FAEE1 gene is linked to the E1 locus associated with variation in the content of erucic acid. *Theor Appl Genet* 96(2): 177–186. https://doi.org/10.1007/s001220050725.

Bates PD. 2016. Understanding the control of acyl flux through the lipid metabolic network of plant oil biosynthesis. *Biochim Biophys Acta* 1861(9PtB): 1214–1225. https://doi.org/10.1016/j.bbalip.2016.03.021.

Bates PD, Johnson SR, Cao X, et al. 2014. Fatty acid synthesis is inhibited by inefficient utilization of unusual fatty acids for glycerolipid assembly. *Proc Natl Acad Sci USA* 111(3): 1204–9. https://doi.org/10.1073/pnas.1318511111.

Baud S. 2018. Seeds as oil factories. *Plant Reprod* 31(3): 213–235. https://doi.org/10.1007/s00497-018-0325-6.

Baud S, Lepiniec L. 2010. Physiological and developmental regulation of seed oil production. *Prog Lipid Res* 49(3): 235–49. https://doi.org/10.1016/j.plipres.2010.01.001.

Baud S, Mendoza MS, To A, et al. 2007. WRINKLED1 specifies the regulatory action of LEAFY COTYLEDON2 towards fatty acid metabolism during seed maturation in Arabidopsis. *Plant J* 50(5): 825–38. https://doi.org/10.1111/j.1365-313X.2007.03092.x.

Baye T, Becker HC, Von Witzke-Ehbrecht S. 2005. Vernonia galamensis, a natural source of epoxy oil: Variation in fatty acid composition of seed and leaf lipids. *Ind Crops Prod* 21(2): 257–261. https://doi.org/10.1016/j.indcrop.2004.04.003.

Bhandari S, Bates PD. 2021. Triacylglycerol remodeling in *Physaria fendleri* indicates oil accumulation is dynamic and not a metabolic endpoint. *Plant Physiol* 187(2): 799–815. https://doi.org/10.1093/plphys/kiab294.

Bhattacharya S, Sinha S, Das N, Maiti MK. 2015. Increasing the stearate content in seed oil of *Brassica juncea* by heterologous expression of M1FatB affects lipid content and germination frequency of transgenic seeds. *Plant Physiol Biochem* 96: 345–55. https://doi.org/10.1016/j.plaphy.2015.08.015.

Brookes G, Bargfoot P. 2020. GM crop technology use 1996–2018: Farm income and production impacts. *GM Crops & Food* 11(4): 142–261. https://doi.org/10.1080/21645698.2020.1779574.

Cahoon EB, Li-Beisson Y. 2020. Plant unusual fatty acids: Learning from the less common. *Curr Opin Plant Biol* 55: 66–73. https://doi.org/10.1016/j.pbi.2020.03.007.

Cahoon EB, Ripp KG, Hall SE, McGonigle B. 2002. Transgenic production of epoxy fatty acids by expression of a cytochrome P450 enzyme from Euphorbia lagascae seed. *Plant Physiol* 128 (2): 615–24. https://doi.org/10.1104/pp.010768.

Cao YZ, Oo KC, Huang AH. 1990. Lysophosphatidate acyltransferase in the microsomes from maturing seeds of Meadowfoam (*Limnanthes alba*). *Plant Physiol* 94(3): 1199–206. https://doi.org/10.1104/pp.94.3.1199.

Chhikara S, Abdullah HM, Akbari P, Schnell D, Dhankher OP. 2018. Engineering *Camelina sativa* (L.) Crantz for enhanced oil and seed yields by combining diacylglycerol acyltransferase1 and glycerol-3-phosphate dehydrogenase expression. *Plant Biotechnol J* 16(5): 1034–1045. https://doi.org/10.1111/pbi.12847.

Chi X, Hu R, Zhang X, et al. 2014. Cloning and functional analysis of three diacylglycerol acyltransferase genes from peanut (*Arachis hypogaea* L.). *PLoS One* 9(9): e105834. https://doi.org/10.1371/journal.pone.0105834.

Dave RG, Patel RM, Patel RJ. 1985. Characteristics and composition of seeds and oil of wild variety of *Momardica chrantiu* L. from Gujarat, India. *Fette Seifen Anstrichmittel* 8: 326–327.

Dumeignil F. 2012. Propriétés et utilisation de l'huile de ricin. *OCL* 19 (1): 10–15. https://doi.org/10.1051/ocl.2012.0427.

Durrett TP, McClosky DD, Tumaney AW, et al. 2010. A distinct DGAT with sn-3 acetyltransferase activity that synthesizes unusual, reduced-viscosity oils in Euonymus and transgenic seeds. *Proc Natl Acad Sci USA* 107(20): 9464–9. https://doi.org/10.1073/pnas.1001707107.

Dussert S, Guerin C, Andersson M, et al. 2013. Comparative transcriptome analysis of three oil palm fruit and seed tissues that differ in oil content and fatty acid composition. *Plant Physiol* 162 (3): 1337–58. https://doi.org/10.1104/pp.113.220525.

Dyer JM, Chapital DC, Kuan JC, et al. 2002. Molecular analysis of a bifunctional fatty acid conjugase/desaturase from tung. Implications for the evolution of plant fatty acid diversity. *Plant Physiol* 130(4): 2027–38. https://doi.org/10.1104/pp.102.010835.

Earle FR. 1966. Optically active aceto-triglyceride of *Euonymous verrucosus*. *J Am Oil Chem Soc* 36: A102.

Focks N, Benning C. 1998. WRINKLED1: A novel, low-seed-oil mutant of Arabidopsis with a deficiency in the seed-specific regulation of carbohydrate metabolism. *Plant Physiol* 118(1): 91–101.

Gan L, Park K, Chai J, et al. 2022. Divergent evolution of extreme production of variant plant monounsaturated fatty acids. *Proc Natl Acad Sci USA* 119(30): e2201160119. https://doi.org/10.1073/pnas.2201160119.

Gao H, Gao Y, Zhang F, et al. 2021. Functional characterization of an novel acyl-CoA:diacylglycerol acyltransferase 3-3 (CsDGAT3-3) gene from *Camelina sativa*. *Plant Sci* 303: 110752. https://doi.org/10.1016/j.plantsci.2020.110752.

Graham SA, Kleiman R. 1992. Composition of seed oils in some Latin American Cuphea (Lythraceae). *Ind Crops Prod* 1: 31–34. https://doi.org/10.1016/0926-6690(92)90042-T.

Grzelak-Blaszczyk K, Karlinska E, Grzeda K, Roj E, Kolodziejczyk K. 2017. Defatted strawberry seeds as a source of phenolics, dietary fiber and minerals. *Lwt-Food Sci Technol* 84: 18–22. https://doi.org/10.1016/j.lwt.2017.05.014.
Gunstone FD, Harwood JL, Padley FB. 1986. The Lipid Handbook. CRC Press.

Han L, Usher S, Sandgrind S, et al. 2020. High level accumulation of EPA and DHA in field-grown transgenic Camelina-A multi-territory evaluation of TAG accumulation and heterogeneity. *Plant Biotechnol J* 18(11): 2280-2291. https://doi.org/10.1111/pbi.13385.

Harlow RD, Litchfield C, Fu H-C, Reiser R. 1965. The triglyceride composition of Myrica carolinensis fruit coat fat (bayberry tallow). *J Am Oil Chem Soc* 42(9): 747-750. https://doi.org/10.1007/BF02631853.

Haslam TM, Kunst L. 2013. Extending the story of very-long-chain fatty acid elongation. *Plant Sci* 210: 93-107. https://doi.org/10.1016/j.plantsci.2013.05.008.

Hatanaka T, Shimizu R, Hildebrand D. 2004. Expression of a Stokesia laevis epoxygenase gene. *Phytochemistry* 65(15): 2189-96. https://doi.org/10.1016/j.phytochem.2004.06.006.

Hayes DG, Kleiman R, Phillips BS. 1995. The triglyceride composition, structure, and presence of estolides in the oils of lesquerella and related species. *JAOCS* 72: 559-569.

Hennessy AA, Ross PR, Fitzgerald GF, Stanton C. 2016. Sources and bioactive properties of conjugated dietary fatty acids. *Lipids* 51 (4): 377-97. https://doi.org/10.1007/s11745-016-4135-z.

Huang AHC. 2018. Plant lipid droplets and their associated proteins: Potential for rapid advances. *Plant Physiol* 176(3): 1894-1918. https://doi.org/10.1104/pp.17.01677.

Iskandarov U, Silva JE, Kim HJ, et al. 2017. A specialized diacylglycerol acyltransferase contributes to the extreme medium-chain fatty acid content of cuphea seed oil. *Plant Physiol* 174 (1): 97-109. https://doi.org/10.1104/pp.16.01894.

Jiang WZ, Henry IM, Lynagh PG, et al. 2017. Significant enhancement of fatty acid composition in seeds of the allohexaploid, *Camelina sativa*, using CRISPR/Cas9 gene editing. *Plant Biotechnol J* 15: 648-657.

Jin Q, O'Keefe SF, Stewart AC, et al. 2021. Techno-economic analysis of a grape pomace biorefinery: Production of seed oil, polyphenols, and biochar. *Food Bioprod Process* 127: 139-151. https://doi.org/10.1016/j.fbp.2021.02.002.

Johansson A, Laakso P, Kallio P. 1997. Characterization of seed oils of wild, edible Finnish berries. *Z Lebensm Unters Forsch* 203: 300-307.

Jones A, Davies HM, Voelker TA. 1995. Palmitoyl-acyl carrier protein (ACP) thioesterase and the evolutionary origin of plant acyl-ACP thioesterases. *Plant Cell* 7(3): 359-71. https://doi.org/10.1105/tpc.7.3.359.

Kazaz S, Barthole G, Domergue F, et al. 2020. Differential activation of partially redundant delta9 stearoyl-acp desaturase genes is critical for omega-9 monounsaturated fatty acid biosynthesis during seed development in arabidopsis. *Plant Cell* 32(11): 3613-3637. https://doi.org/10.1105/tpc.20.00554.

Kleiman R, Payne-Wahl KL. 1984. Fatty acid composition of seed oils of the meliaceae, including one genus rich in cis-vaccenic acid. *JAOCS* 61: 1836-1838.

Knutsen HK, Alexander J, Barregard L, et al. 2016. Erucic acid in feed and food. *EFSA J*. https://doi.org/10.2903/j.efsa.2016.4593.

Knutzon DS, Thompson GA, Radke SE, et al. 1992. Modification of Brassica seed oil by antisense expression of a stearoyl-acyl carrier protein desaturase gene. *Proc Natl Acad Sci USA* 89(7): 2624-8. https://doi.org/10.1073/pnas.89.7.2624.

Kroon JT, Wei W, Simon WJ, Slabas AR. 2006. Identification and functional expression of a type 2 acyl-CoA:diacylglycerol acyltransferase (DGAT2) in developing castor bean seeds which has high homology to the major triglyceride biosynthetic enzyme of fungi and animals. *Phytochemistry* 67(23): 2541-9. https://doi.org/10.1016/j.phytochem.2006.09.020.

Labalette F, Land N, Wagner D, Roux-Duparque M, Saillet E. 2011. La filière lin oléagineux française: panorama et perspectives. *OCL* 18(3): 113-122.

Leebens-Mack JH, Barker MS, Carpenter EJ, et al. 2019. One thousand plant transcriptomes and the phylogenomics of green plants. *Nature* 574(7780): 679. https://doi.org/10.1038/s41586-019-1693-2.

Li-Beisson Y, Shorrosh B, Beisson F, et al. 2010. Acyl-lipid metabolism. *Arabidopsis Book* 8: e0133. https://doi.org/10.1199/tab.0133.

Li-Beisson Y, Shorrosh B, Beisson F, et al. 2013. Acyl-lipid metabolism. *Arabidopsis Book* 11: e0161. https://doi.org/10.1199/tab.0161.

Li R, Yu K, Hatanaka T, Hildebrand DF. 2010. Vernonia DGATs increase accumulation of epoxy fatty acids in oil. *Plant Biotech J* 8: 184-195.

Li X, van Loo EN, Gruber J, et al. 2012. Development of ultra-high erucic acid oil in the industrial oil crop Crambe abyssinica. *Plant Biotechnol J* 10(7): 862-70. https://doi.org/10.1111/j.1467-7652.2012.00709.x.

Li Q, Shao J, Tang S, et al. 2015. WRINKLED1 accelerates flowering and regulates lipid homeostasis between oil accumulation and membrane lipidabolism in *Brassica napus*. *Front Plant Sci* 6: 1015. https://doi.org/10.3389/fpls.2015.01015.

Li X, Teitgen AM, Shirani A, et al. 2018. Discontinuous fatty acid elongation yields hydroxylated seed oil with improved function. *Nat Plants* 4(9): 711-720. https://doi.org/10.1038/s41477-018-0225-7.

Lindqvist Y, Huang W, Schneider G, Shanklin J. 1996. Crystal structure of delta9 stearoyl-acyl carrier protein desaturase from castor seed and its relationship to other di-iron proteins. *EMBO J* 15(16): 4081-92.

Liu J, Rice A, McGlew K, et al. 2015a. Metabolic engineering of oilseed crops to produce high levels of novel acetyl glyceride oils with reduced viscosity, freezing point and calorific value. *Plant Biotechnol J* 13(6): 858-65. https://doi.org/10.1111/pbi.12325.

Liu J, Tjellstroem H, McGlew K, et al. 2015b. Field production, purification and analysis of high-oleic acetyl-triacylglycerols from transgenic *Camelina sativa*. *Ind Crops Prod* 65: 259-268. https://doi.org/10.1016/j.indcrop.2014.11.019.

Macho GA. 2014. Baboon feeding ecology informs the dietary niche of Paranthropus boisei. *PLoS One* 9(1): e84942. https://doi.org/10.1371/journal.pone.0084942.

Maeo K, Tokuda T, Ayame A, et al. 2009. An AP2-type transcription factor, WRINKLED1, of *Arabidopsis thaliana* binds to the AW-box sequence conserved among proximal upstream regions of genes involved in fatty acid synthesis. *Plant J* 60(3): 476-87. https://doi.org/10.1111/j.1365-313X2009.03967.x.

Maisonneuve S, Bessoule JJ, Lessire R, Delseny M, Roscoe TJ. 2010. Expression of rapeseed microsomal lysophosphatidic acid acyltransferase isozymes enhances seed oil content in Arabidopsis. *Plant Physiol* 152(2): 670-84. https://doi.org/10.1104/pp.109.148247.

McFie PJ, Stone SL, Banman SL, Stone SJ. 2010. Topological orientation of acyl-CoA:diacylglycerol acyltransferase-1 (DGAT1) and identification of a putative active site histidine and the role of the n terminus in dimer/tetramer formation. *J Biol Chem* 285(48): 37377-87. https://doi.org/10.1074/jbc.M110.163691.

Metzger JO. 2009. Fats and oils as renewable feedstock for chemistry. *Eur J Lipid Sci Technol* 111: 865-876. https://doi.org/10.1002/ejlt.200900130.

Miklaszewska M, Zienkiewicz K, Inchana P, Zienkiewicz A. 2021. Lipid metabolism and accumulation in oilseed crops. *OCL* 28. https://doi.org/10.1051/ocl/2021039.
Morineau C, Bellec Y, Tellier F, et al. 2017. Selective gene dosage by CRISPR-Cas9 genome editing in hexaploid *Camelina sativa*. *Plant Biotechnol J* 15(6): 729–739. https://doi.org/10.1111/pbi.12671.

Napier JA, Haslam RP, Tsalavouta M, Sayanova O. 2019. The challenges of delivering genetically modified crops with nutritional enhancement traits. *Nat Plants* 5(6): 563–567. https://doi.org/10.1038/s41477-019-0430-z.

Nguyen HT, Mishra G, Whittle E, et al. 2010. Metabolic engineering of seeds can achieve levels of omega-7 fatty acids comparable with the highest levels found in natural plant sources. *Plant Physiol* 154(4): 1897–904. https://doi.org/10.1104/pp.110.165340.

Ohlrogge J, Thrower N, Mhaske V, et al. 2018. PlantFAdb: a resource for exploring hundreds of plant fatty acid structures synthesized by thousands of plants and their phylogenetic relationships. *Plant J* 96(6): 1299–1308. https://doi.org/10.1111/tpj.14102.

Ortiz R, Geleta M, Gustafsson C, et al. 2020. Oil crops for the future. *Curr Opin Plant Biol* 56: 181–189. https://doi.org/10.1016/j.pbi.2019.12.003.

Perry HJ, Harwood JL. 1993. Changes in the lipid-content of developing seeds of *Brassica napus*. *Phytochemistry* 32(6): 1411–1415. https://doi.org/10.1016/0031-9422(93)85148-k.

Purdy RH. 1986. High oleic sunflower-Physical and chemical characteristics. *J Am Oil Chem Soc* 63(8): 1062–1066. https://doi.org/10.1007/bf02673799.

Rodriguez-Rodriguez MF, Moreno-Perez AJ, Makni S, et al. 2021. Lipid profiling and oil properties of *Camelina sativa* seeds engineered to enhance the production of saturated and omega-7 fatty acids. *Ind Crops Prod* 170. https://doi.org/10.1016/j.indcrop.2021.113765.

Romsdahl T, Shirani A, Minto RE, et al. 2019. Nature-guided synthesis of advanced bio-lubricants. *Sci Rep* 9. https://doi.org/10.1038/s41598-019-48165-6.

Routaboul JM, Benning C, Bechtold N, Caboche M, Lepiniec L. 1999. The TAG1 locus of Arabidopsis encodes for a diacylglycerol acyltransferase. *Plant Physiol Biochem* 37(11): 831–840. doi:S0981-9428(99)00115-1.

Saha S, Enugutti B, Rajakumari S, Rajasekharan R. 2006. Cytosolic triacylglycerol biosynthetic pathway in oilseeds: molecular cloning and expression of peanut cytosolic diacylglycerol acyltransferase. *Plant Physiol* 141(4): 1533–1543. https://doi.org/10.1104/pp.106.082198.

Salas JJ, Bootello MA, Martinez-Force E, Venegas Caleron M, Garces R. 2021. High stearic sunflower oil: Latest advances and applications. *OCL* 28. https://doi.org/10.1051/ocl/2021022.

Samarappuli D, Zanetti F, Berzuini S, Berti MT. 2020. Crambe (Crambe abyssinica Hochst): A non-food oilseed crop with great potential: A review. *Agronomy-Basel* 10(9). https://doi.org/10.3390/agronomy10091380.

Shi JL, Cao YP, Fan XR, et al. 2012. A rice microsomal delta-12 fatty acid desaturase can enhance resistance to cold stress in yeast and *Oryza sativa*. *Mol Breed* 29(3): 743–757. https://doi.org/10.1007/s11032-011-9587-5.

Shockey J, Regmi A, Cotton K, et al. 2016. Identification of Arabidopsis GPAT9 (At5g60620) as an essential gene involved in triacylglycerol biosynthesis. *Plant Physiol* 170(1): 163–79. https://doi.org/10.1104/pp.15.01563.

Siloto RM, Findlay K, Lopez-Villalobos A, et al. 2006. The accumulation of oleosins determines the size of seed oilbodies in Arabidopsis. *Plant Cell* 18(8): 1961–74. https://doi.org/10.1105/tpc.106.041269.

Sonntag, ed. 1979. Composition and characteristics of individual fats and oils. In: Swern D, ed. *Bailey's industrial oil and fat products*. New York: John Wiley & Sons.

Stahl U, Stalberg K, Stymne S, Ronne H. 2008. A family of eukaryotic lysophospholipid acyltransferases with broad specificity. *FEBS Lett* 582(2): 305–9. https://doi.org/10.1016/j.febslet.2007.12.020.

Stone SJ, Levin MC, Farese RV Jr. 2006. Membrane topology and identification of key functional amino acid residues of murine acyl-CoA:diacylglycerol acyltransferase-2. *J Biol Chem* 281(52): 40273–82. https://doi.org/10.1074/jbc.M607986200.

Swarbrick CMD, Nanson JD, Patterson EI, Forwood JK. 2020. Structure, function, and regulation of thioesterases. *Progr Lipid Res* 79. https://doi.org/10.1016/j.plipres.2020.101036.

Tarazona P, Feussner K, Feussner I. 2015. An enhanced plant lipidomics method based on multiplexed liquid chromatography-mass spectrometry reveals additional insights into cold- and drought-induced membrane remodeling. *Plant J* 84(3): 621–33. https://doi.org/10.1111/tpj.13013.

Ucciani E. 1994. Nouveau dictionnaire des huiles végétales: compositions en acides gras. Technique et Documentation. Paris: Lavoisier.

van de Loo FJ, Broun P, Turner S, Somerville C. 1995. An oleate 12-hydroxylase from *Ricinus communis* L. is a fatty acyl desaturase homolog. *Proc Natl Acad Sci USA* 92(15): 6743–7. https://doi.org/10.1073/pnas.92.15.6743.

van der Vossen HAM, Mkamilo GS, Eds. 2007. PROTA 14 Oléagineux, ressources végétales de l'Afrique de l'ouest [Transl. by Chauvet JM and Siemonsma JS]. In: PROTA. Wageningen: Fondation PROTA, Backhuys Publishers.

Vandeputte J. 2012. Les agro-tensio actifs. *OCL* 192(2): 133–137.

Vanhercke T, El Tahchy A, Liu Q, et al. 2014. Metabolic engineering of biomass for high energy density: Oilseed-like triacylglycerol yields from plant leaves. *Plant Biotechnol J* 12(2): 231–9. https://doi.org/10.1111/pbi.12131.

Vega-Morales T, Mateos-Diaz C, Perez-Machin R, et al. 2019. Chemical composition of industrially and laboratory processed Cyperus esculentus rhizomes. *Food Chem* 297: 124896. https://doi.org/10.1016/j.foodchem.2019.05.170.

Vigeolas H, Waldeck P, Zank T, Geigenberger P. 2007. Increasing seed oil content in oil-seed rape (*Brassica napus* L.) by over-expression of a yeast glycerol-3-phosphate dehydrogenase under the control of a seed-specific promoter. *Plant Biotechnol J* 5(3): 431–441. https://doi.org/10.1111/j.1467-7652.2007.00252.x.

Vrinten P, Hu Z, Munchinsky MA, Rowland G, Qiu X. 2005. Two FAD3 desaturase genes control the level of linolenic acid in flax seed. *Plant Physiol* 139(1): 79–87. https://doi.org/10.1104/pp.105.064451.

Wang G, Lin Q, Xu Y. 2007. Tetraena mongolica Maxim can accumulate large amounts of triacylglycerol in phloem cells and xylem parenchyma of stems. *Phytochemistry* 68(15): 2112–2117. https://doi.org/10.1016/j.phytochem.2007.04.040.

Wang X, Yu C, Liu Y, et al. 2019. GmFAD3A, a omega-3 fatty acid desaturase gene, enhances cold tolerance and seed germination rate under low temperature in rice. *Int J Mol Sci* 20(15). https://doi.org/10.3390/ijms20153796.

Wang L, Jing M, Wang Y, et al. 2020. Integrative analysis of lipidomics and transcriptomics revealed dynamic details of lipids metabolism and accumulation in developing tiger nut (Cyperus Esculentus) tubers. *Res Square*. https://doi.org/10.21203/rs.3.rs-115598/v1.

Wiberg E, Banas A, Stymne S. 1997. Fatty acid distribution and lipid metabolism in developing seeds of laureate-producing rape (*Brassica napus* L.). *Planta* 203(3): 341–8. https://doi.org/10.1007/s004250050200.

Winnacker M, Rieger B. 2016. Biobased polyamides: Recent advances in basic and applied research. *Macromol Rapid Commun* 37(17): 1391–1413. https://doi.org/10.1002/marc.201600181.
Xu C, Shanklin J. 2016. Triacylglycerol metabolism, function, and accumulation in plant vegetative tissues. *Annu Rev Plant Biol* 67: 179–206. https://doi.org/10.1146/annurev-arplant-043015-111641.

Zanetti F, Monti A, Berti MT. 1993. Challenges and opportunities for new industrial oilseed crops in EU-27: A review. *Ind Crops Prod* 50: 580–595. https://doi.org/10.1016/j.indcrop.2013.08.030.

Zeng F, Roslinsky V, Cheng B. 2017. Mutations in the promoter, intron and CDS of two FAD2 generate multiple alleles modulating linoleic acid level in yellow mustard. *Sci Rep* 7(1): 8284. https://doi.org/10.1038/s41598-017-08317-y.

Zhai Z, Liu H, Shanklin J. 2021. Ectopic Expression of OLEOSIN 1 and inactivation of GBSS1 have a synergistic effect on oil accumulation in plant leaves. *Plants (Basel)* 10(3). https://doi.org/10.3390/plants10030513.

Zhang M, Fan J, Taylor DC, Ohlrogge JB. 2009. DGAT1 and PDAT1 acyltransferases have overlapping functions in Arabidopsis triacylglycerol biosynthesis and are essential for normal pollen and seed development. *Plant Cell* 21(12): 3885–901. https://doi.org/10.1105/tpc.109.071795.

Zhang L, Jia B, Tan X, et al. 2014. Biodiesel production from tung (Vernicia montana) oil and its blending properties in different fatty acid compositions. *PLoS One* 9(8): e105298. https://doi.org/10.1371/journal.pone.0105298.

**Cite this article as:** Chardot T. 2022. A comparison of natural and induced diversity in plant oils. *OCL* 29: 34.