Plant Growth Regulation

, Volume 83, Issue 2, pp 207–222 | Cite as

Genetic engineering approaches to enhance oil content in oilseed crops

  • Siddanna SavadiEmail author
  • Nemappa Lambani
  • Prem Lal Kashyap
  • Deepak Singh Bisht
Original paper


Oilseed crops play an important role in the agricultural economy. Apart from being an integral component of human diet and industrial applications, they are also gaining importance as replacement to fossil fuels for meeting the energy needs. The last two decades have been marked by several important events in genetic engineering and identification of gene targets for enhancing seed oil content in oilseed crops, and will aid the successful development of new generation high yielding oil crops. Specifically, genetic engineering has shown real breakthrough in enhancing oil content in oilseed rape, camelina, soybean and maize. Moreover, ongoing research efforts to decipher the possibilities of genetic modifications of key regulators of oil accumulation along with physiological and biochemical studies to understand lipid biosynthesis will set a platform to produce transgenic oilseed crops with enhanced oil content. In this review, we briefly describe different genetic engineering approaches explored by different researchers for enhancing oil content. Further, we discuss a few promising and potential approaches and challenges for engineering oil content in oilseed crops.


Oil content Oilseed Pathway Transgenics TAG 


Oilseed crops play an important role in the agricultural economy, next only to food grains in terms of area and production. Globally, the demand for vegetable oils is increasing due to the increasing per capita consumption of oil in our diets and its use as biofuels (Samarth and Mahanwar 2015). By 2050, the global demand for vegetable oils is expected to be more than twice the current production. Vegetable oils form an essential part of the human diet, as they provide good source of energy, and a variety of Fatty acids (FAs) needed for good health as well as helping in the assimilation of fat soluble vitamins (e.g. A, D, E, and K) into the body (DeLuca 2012). Oils are also used in industries as raw material for synthesis of various products such as soaps, cosmetics, polymers and pharmaceuticals (Elahi et al. 2016a). Apart from direct consumption and industrial applications, they are also gaining importance as a replacement for fossil fuels for meeting the energy needs (Issariyakul and Dalai 2014). Therefore, there is a need to enhance oil yields to meet the growing demands. Increasing the oil content in oilseed crops is one option for enhancing oil yield without increasing the area (~201.8 mha, OECD/FAO 2015) under oilseed crops and to save the inputs used to raise the extra crop required to meet the future global oil shortage.

Yield improvement in oilseeds can be addressed by increasing the quantity of oil per seed, by increasing the size of the seed, or by increasing the number of seeds per plant. Each of the steps from photosynthesis, metabolite partitioning to final conversion and deposition of different storage products in storage organs determines yield (Camp 2005; Bhat 2010). Accordingly, increasing source strength, sink strength and assimilate partitioning in favour of storage lipids is critical to achieving higher oil yields. Identification and strategic manipulation of the genetic factors and molecular networks controlling source strength, sink strength and assimilate partitioning would enable considerable increases in oil content. Oilseed crops are annual plants whose seeds are used mainly for extraction of culinary and industrial oils. All plant species accumulate oil, protein and carbohydrates in their seeds, but their proportions vary. Genetic and metabolic regulation determines the proportion of oil in seeds of different species. Oilseed crops have higher seed oil content than other agriculturally important crops. Even among oilseed crops, the oil content varies drastically for instance from 20% in soybean to 60% in sesame although the lipid biosynthetic pathway is similar in these species (Gupta 2008). This suggests that there is scope for engineering the levels of seed oils to be significantly higher than the present levels in the majority of the oilseed crops. However, the extent of success depends on the critical targets and control points in pathways that are species specific and the approaches employed for the manipulation of such targets.

Conventional plant breeding methods such as pure-line selection and mutation breeding have led to the development of oilseed crop varieties with increased seed oil. However, the scope for improving oil traits by these methods is limited by the genetic variation available in a crop species (Murphy 1995, 2014; Yadava et al. 2012; Hua et al. 2016). Further, oil yield is a quantitative trait controlled by many loci and is influenced by environment (Rahman et al. 2013). Several quantitative trait loci (QTLs) controlling seed oils have been identified in different oilseed crops (Eskandari et al. 2013; Pandey et al. 2014; Wang et al. 2015). However, there are no reports of their successful pyramiding in a suitable genetic background leading to significant improvement of oil content. Moreover, with the identification of genes involved in lipid biosynthesis and its regulation, generating transgenic oil plants capable of producing the desired oil characteristics has become possible with the help of molecular biology tools and transformation methods (Maheshwar and Kovalchuk 2014). Mounting evidence suggests that genetically engineered oil crops carry a great promise for the development of new and improved crop varieties with desirable traits, such as increased oil content. This article reviews regulation of oil biosynthesis, scope for increasing seed oil and recently employed as well as possible genetic engineering strategies for enhancing the seed oil content of oilseed crops.

Regulation of oil biosynthesis in plants

Generally, oils are chemically triaylglycerides (TAG) formed from sequential acylation of three FAs with a glycerol backbone. The basic lipid synthetic pathway and its components are similar among species (Harwood and Guschina 2013). However, the oil content of species varies, and this is attributed to differences in the regulatory mechanisms operating in different species (Murphy 1996; Guschina et al. 2014). Regulation of oil content is a complex process occurring at different stages of oil biosynthesis and storage (Elahi et al. 2016a; Fig. 1). In Arabidopsis, at least 120 enzymatic reactions and >600 genes are known to control the enzymatic and regulatory functions associated with lipid metabolism showing the complexity of the lipid metabolic pathways (Li-Beisson et al. 2013). Besides genetic factors, oil biosynthesis is also regulated by physiological and developmental factors (Baud and Lepiniec 2010; Fig. 1). Therefore, a comprehensive understanding of the regulatory framework of lipid biosynthesis in different species might address the underlying bottleneck in increasing the oil content of oilseed crops.

Fig. 1

Genetic and physiological factors governing seed oil biosynthesis, transport and storage in plants. These factors ultimately influence the seed oil content

Growing interest in enhancing oil yields of plants has prompted discovery of several molecular regulators of oil biosynthesis in Arabidopsis and other plant species. Principally, the molecular regulators of oil content include genes encoding transcription factors (TFs) and enzymes involved with FAs and TAG synthesis and carbon flux (Zou et al. 1999; Marillia et al. 2003; Jofuku et al. 2005; Shen et al. 2006; Wakao et al. 2008; Meyer et al. 2012; Kelly et al. 2013a, b; Chandran et al. 2014). Further, comparative genomics studies have revealed that allelic variations within the key genes can also be responsible for significant variation in the quality and quantity of oil in different oilseed crops (Sharma and Chauhan 2012; Teh and Möllers 2016). In addition, oil accumulation is also highly influenced by maternal effects as during the later stages of seed development the photosynthates from silique or pod walls are transported and accumulated as starch, protein, lipids and many secondary metabolites in seeds (Hua et al. 2012).

Regulation of the proportion of seed storage molecules including lipids among plant species can be measured by several biochemical and genetic techniques (Guschina et al. 2014; Elahi et al. 2016b). Recently, a metabolic control analysis (MCA) model has been used to study metabolite biosynthesis regulation (Zadran and Levine 2013). It provides a quantitative measure of the regulatory control of different pathways as well as the individual enzymes of each pathway. For instance, the group flux control coefficient of TAG biosynthesis is more than FA synthesis in oilseed rape but lower in soybean (Harwood and Guschina 2013). Thus, the complete understanding of the function and interaction among these molecular regulators will be critical for altering the level and composition of seed oil without any deleterious phenotypes, which is essential for success of any plant modification approaches.

Genetic engineering approaches to enhance oil content

Numerous studies have shown that genetic engineering can complement conventional plant breeding methods to break yield barriers in crop plants. Further, genetic engineering approaches bring about precise changes in the genome in short period of time with controlled expression of traits and without any linkage drag. In addition, recent advancements in the high-throughput technologies like genomics, transcriptomics, lipidomics and phenomics have led to the understanding of genes, gene networks, regulatory factors and their interactions that govern seed oil biosynthesis. Also, these advanced technologies have provided tools that allow devising of novel strategies for the genetic engineering of crop plants (Sanghera et al. 2011; Kashyap et al. 2011). In general, the process of GM crop development involves identification of suitable candidate genes, devising a suitable strategy for expressing the chosen candidates and development of transgenics followed by rigorous field evaluation of transgenic crops. Transgenics showing better agronomic performance would be incorporated into plant breeding programs or released for commercial cultivation (Fig. 2).

Fig. 2

Tools and strategies for genetic engineering of seed oil content in plants. Basic and advanced omics tools have facilitated discovery of gene targets and regulatory mechanisms governing the seed oil content. This information in turn has facilitated devising strategies for engineering higher seed oil content in plants

In the past, a number of transgenic strategies have been tested for the enhancement of seed oil content which include manipulation of TAG and/or FA synthesis, modulation of carbon flux towards TAG biosynthesis and or FA synthesis, manipulation of oil bodies, improving availability of limiting substrates like glycerol-3-phosphate (G3P) or oxygen and the introduction of novel pathways for TAG synthesis sourced from mammalian or yeast systems (Vigeolas et al. 2007, 2011; Petrie et al. 2012; Liu et al. 2013). Field trials of some of the transgenics developed for enhanced seed oil have shown promising results (Taylor et al. 2002, 2009; Sharma et al. 2008; Shen et al. 2010a). These methods are now being applied to a wide range of crops and have potential for oil crop improvement in terms of both yield and nutritional quality of the oils (Table S1). Recently, much attention has been paid to the deployment of new generation genetic engineering technologies such as monomeric transcription activator-like effectors based nucleases (mTALENs) and modified forms of the clustered regularly interspaced palindromic repeat (CRISPR)/Cas9 system for genetic manipulation of a wide range of crops including oilseed crops (Zhang et al. 2014; Chen et al. 2014). In the near future, with a better understanding of the regulation of oil biosynthesis along with application of modern as well as conventional plant breeding tools, a major breakthrough in the manipulation of oil crops is expected. In the following sections, the strategies followed by various researchers as well as potential ones for genetic engineering of plants for enhancing oil content are briefly reviewed.

Enhancement of oil yield by elevating rate of FA synthesis

FAs are synthesized in plastids and transported to cytosol forming a fatty acyl pool which is utilized for lipid synthesis in the endoplasmic reticulum (ER). Studies have suggested that oil synthesis may be limited by the production of FAs (Bao and Ohlrogge 1999; Sasaki and Nagano 2004). In turn, FAs biosynthesis is regulated by the activity of acetyl-CoA carboxylase (ACCase) and 3-ketoacyl-ACP synthase III (KAS III) (Shintani and Ohlrogge 1995; Ohlrogge and Jaworski 1997). In initial attempts, genes encoding enzymes involved in the FA biosynthetic pathway were the targets for genetic manipulation to enhance substrate accumulation for TAG synthesis and to ultimately increase oil content. Plastid-targeted expression of ACCase in Arabidopsis resulted in 5% increase in seed oil (Roesler et al. 1997) whereas reduction in ACCase activity lowered the FA content in the transgenic seeds (Thelen and Ohlrogge 2002) indicating ACCase activity is one of the limiting factors in the synthesis of FAs. Verwoert et al. (1995) overexpressed an Escherichia coli KAS III gene in transgenic rapeseed (B. napus), which resulted in 3–4 fold higher KAS III activity. However, total seed FA content was not significantly altered. Thus, transgenic studies employing genes encoding enzymes involved in FA biosynthesis resulted in either modest increase or no changes in seed oil content (Roesler et al. 1997; Klaus et al. 2004). Nevertheless, these initial attempts to enhance lipid content have shed light on the finer details of oil biosynthesis regulation and demonstrated the potential of genetic engineering application in oil crops improvement.

Enhancement of oil yield by increasing TAG synthesis

Seed oil accumulates primarily in the form of triacylglycerol (TAG) and serves as an energy reserve for the germinating seed. TAGs are derived from sequential incorporation of FAs onto a glycerol backbone in the ER through the Kennedy pathway (Fig. S1). First, sn-glycerol-3-phosphate is acylated by the action of glycerol-3-phosphate acyltransferase (GPAT) and subsequently by lyso-phosphatidic acid acyltransferase (LPAAT) to produce lysophosphatidic acid (PA). PA is then dephosphorylated by phosphatidate phosphatase (PAP) to form sn-1,2-diacylglycerol (DAG) which is finally acylated by diacylglycerol acyltransferase (DGAT) to give TAG. Several transgenic and enzyme kinetics studies have shown that the Kennedy pathway enzymes regulate the TAG biosynthesis and as consequence the oil content of oil crops (Zou et al. 1997; Perry et al. 1999; Cao et al. 2006). Reaction catalyzed by GPAT is one of the rate limiting steps of TAG synthesis (Cao et al. 2006). Jain et al. (2000) demonstrated that the overexpression of safflower plastidial GPAT gene and E. coli GPAT in Arabidopsis resulted in up to 22 and 15% increase in seed oil content, respectively. Lysophosphatidate acyltransferase (LPAT) is the second enzyme of the Kennedy pathway, and it catalyzes the conversion of LPA into PA. Zou et al. (1997) obtained an oil increase of 8–48% by expressing mutant yeast LPAT in Brassica. Similarly, seed-specific overexpression of rapeseed microsomal LPAT and peanut LPAT2 in Arabidopsis increased oil content by 13 and 7.4% respectively (Maisonneuve et al. 2010; Chen et al. 2015). These plants had more unsaturated FAs and higher total FA content. In the siliques of homozygous transgenic lines, the relative expression levels of several genes involved in FA biosynthesis, TAG assembly, sucrose metabolism and glycolysis were also significantly increased suggesting a key role of LPAT in oil biosynthesis. Therefore, LPAT is of interest for the genetic engineering of seed oil and FA composition especially reducing the saturated FAs content in edible oils.

Interestingly among the Kennedy pathway enzymes, DGAT is the only enzyme that is solely committed to TAG biosynthesis as its activity diverts the DAG flux from phospholipids synthesis towards TAG synthesis. Further, activity of DGAT is relatively low compared to the activity of other enzymes in the pathway (Perry et al. 1999; Sharma et al. 2008). Thus, DGAT represents a key rate-limiting enzyme and acts as a check point in oil formation in the developing seeds. The regulatory role of DGAT in oil synthesis can be explained by the flux control model which demonstrates that this enzyme may exert considerable flux control under normal physiological conditions in oilseed crops (Ramli et al. 2005). Further, the key regulatory role of DGAT in TAG biosynthesis has been established through genetic and transgenic studies in several plant species (Tables S1, 1). Considering the key regulatory role in TAG synthesis, DGAT is the most studied acyltransferase of the Kennedy pathway enzymes. In Arabidopsis, there are two DGAT isoforms, DGAT1 and DGAT2, having differences in amino acid sequences but with acylation activity. Katavic et al. (1995) identified an Arabidopsis mutant, ASI1, with reduced seed oil content and later Zou et al. (1999) showed that ASI1mutant has a mutation in DGAT1 gene. RNAi-mediated silencing of DGAT1 in tobacco resulted in reduction of seed oil content by 9–49% in transgenic lines whereas protein and sugar content increased in the seeds of these lines (Zhang et al. 2005). A high-oil QTL affecting seed oil and oleic acid in maize encoded for DGAT and its ectopic expression in maize resulted in 41% increase in oil content (Zheng et al. 2008). Similarly, transgenic overexpression of DGAT1, either singly or in combination with other genes in different plant species has led to a significant increases in seed oil content (Tables S1, 1). Recently, overexpression of an allelic variant of DGAT1 in soybean resulted in an increase of 3% in the seed oil of transgenics (Roesler et al. 2016). Thus, the studies outlined above suggest that DGAT1 and its allelic variants provide suitable candidates for the engineering oilseed crops with high TAG levels.

Table 1

List of multigene approaches employed for improving oil content in plants




Oil phenotype





Increase in seed oil by 5%

van Erp et al. (2014)


Increase in seed oil by 3%


Increase in seed oil by 3.2%


Increase in seed oil by 6%

WRI1, DGAT1 and SDP1

Increase in seed oil by 7.4%


Mus musculus

Nicotiana sp.

Increases in leaf TAG 7.3 fold

Petrie et al. (2012)


Increases in leaf TAG 9.2 fold


Increases in leaf TAG 9.8 fold




Increase by 0.57% on a leaf dry weight

Vanhercke et al. (2013)


Increase by 0.45% on a leaf dry weight

WRI1 and DGAT1

Increase by 2.48% on a leaf dry weight


B. napus


B. napus

Increase in seed oil content by 14.46%

Liu et al. (2015)



Increase in seed oil content by 12.57%

tgd1-1 and sdp1

tgd1-1 and pxa1


Arabidopsis (tgd1-1)

Increase in leaf oil content by 9% of dry weight

Xu et al. (2016)



Arabidopsis (tdg5)

Increase oil content 8.5% of leaf dry weight

Xu et al. (2015)

Apart from the Kennedy pathway, TAGs are also derived from phospholipid:diacyglycerolacyltransferase (PDAT) catalyzed conversion of phosphatidylcholine (PC) to DAG (Dahlqvist et al. 2000; Stahl et al. 2004). A simultaneous down regulation of PDAT1 and DGAT1 decreased the seed oil content by 70–80%, thus establishing that PDAT1 and DGAT1 play partially redundant functions in Arabidopsis (Zhang et al. 2009). Recently, transcriptome analysis of sesame revealed that PDAT expression was 2–3.5 folds higher in the high-oil accession than the low-oil accessions at 10 days after pollination (DAP) whereas there was no significant difference in DGAT expression. This suggests that expression levels of PDAT during early seed development plays an important role in the determination of seed oil content (Wang et al. 2014a, b). Fan et al. (2013) showed that PDAT1 has a dual role in enhancing FAs and directing FAs from membrane lipids to TAG in Arabidopsis leaves. They also demonstrated that the combined expression of PDAT1 and OELOSIN1 (OLE1, gene encoding the most abundant seed oil droplet-specific protein of Arabidopsis) increases leaf TAG to 6.4% per dry weight in the wild type and 8.6% per dry weight in tgd1 without major negative growth consequences. Taking into consideration the potential benefits of maximizing the TAG content in seed as well as vegetative tissues of crops, PDAT1 provides an ideal candidate to design new strategies for genetic engineering oil in plants.

Recently, Nguyen et al. (2015) explored the possibility of enhancing omega-7 monounsaturated FAs in camelina (Camelina sativa) through a novel approach of redirecting metabolic flux. The omega-7 monounsaturated FAs content was enhanced up to 60–65% of the total FAs in camelina by seed-specific suppression of 3-keto-acyl-ACP synthase II and the FatB 16:0-ACP thioesterase genes to enhance substrate, 16:0-ACP for the ∆9-acyl-ACP desaturase and by blocking C18 FA elongation.

Enhancement of oil yield by increasing carbon flux towards oil biosynthesis

During seed maturation the seed storage compounds like starch, oil and protein are formed from photosynthates produced in silique wall and leaves (Baud et al. 2008; Meyer and Kinney 2010). Hence, in maturing seeds the carbon flux towards TAG synthesis is influenced by other metabolite pathways which are active during same period in seeds and compete for carbon. Accordingly increasing the carbon flux towards synthesis of TAGs is an important avenue for increasing seed oil content (Weselake et al. 2009). The mitochondrial pyruvate dehydrogenase complex (mtPDC) is shown to play a key role in the regulation of lipid biosynthesis by governing the generation and availability of acetyl CoA, the primary substrate of FA synthesis (Zou et al. 1999). mtPDC activity is negatively regulated by phosphorylation by pyruvate dehydrogenase kinase (PDHK) (Thelen et al. 1998). Down-regulation of AtPDHK via seed-specific antisense expression in Arabidopsis resulted in elevated mtPDC activity and enhanced energy (NADH) levels and acetyl CoA for FA synthesis. Correspondingly, the RNAi AtPDHK transgenic plants exhibited relative increases up to 50% in seed oil content with no changes in FA composition (Marillia et al. 2003). Further, glucose-6-phosphate dehydrogenase (G6PDH), the oxidative pentose phosphate pathway (OPPP) enzyme, plays an important role in supplying the energy needed for oil accumulation in developing seeds in which photosynthesis may be light limited (Wakao et al. 2008). Analyses of single and double mutants of cytosolic G6PDH isoforms (G6PD5 and G6PD6) showed that seeds of the double mutant but not of the single mutants had higher oil contents and increased weight compared to the wild type seeds without alteration in the protein content or FA composition. This could be the result of complete loss of cytosolic G6PDH activity in the double mutant leading to impairment in the OPPP pathway. Impairment in OPPP pathway increases the substrate availability for glycolysis and in turn, the increased glycolytic flux could provide more precursors for FA synthesis resulting in enhanced oil content in the double mutant (Focks and Benning 1998; Andre et al. 2007).

Several workers reported that glycolysis, a ubiquitous pathway, is essential for the production of oil and other storage compounds in developing seeds (Focks and Benning 1998; Andre et al. 2007; Wakao et al. 2008). Pyruvate kinase (PK) catalyzes the ADP-dependent conversion of PEP to pyruvate while producing ATP. Studies have suggested that plastidic PK (PPK) activity has a vital role with respect to accumulation of seed oil as PK activity provides ATP and pyruvate essential for sustained FA production in plastids (Ruuska et al. 2002; Baud et al. 2007). Further, there is no evidence for pyruvate transporters across the plastid membrane and hence, it is reasonable that the activity of plastidic PK rather than cytosolic PK provides precursors for FA synthesis. Andre et al. (2007) provided the direct experiential evidence for the role of PPK in the seed oil accumulation. In Arabidopsis, disruption of the gene encoding the β1-subunit reduced the plastidic pyruvate kinase (PPK) activity and a 60% reduction in seed oil content. This seed oil phenotype was fully restored by expression of the β1 subunit–encoding cDNA and partially by the β2 subunit–encoding cDNA. These results suggest that the PPK catalyzes a crucial step in the supply of carbon for FA biosynthesis. Further, the importance of glycolysis for lipid biosynthesis is shown by the fact that the glycolytic intermediate, glycerol-3-phosphate (G3P) is one of the primary substrates required for the synthesis of all the glycerolipids including seed storage lipids.

Estimation of G3P levels in developing seeds of rapeseed (Vigeolas and Geigenberger 2004) and Arabidopsis (Gibon et al. 2002) have demonstrated that the rate of G3P supply is not sufficiently rapid to maintain high G3P levels required during the peak oil accumulation phase. In plants, G3P is produced by two routes involving two different enzymes: cytosolic G3PDH and glycerol kinase (GK) (Lin 1976). Glycerol kinase catalyzes phosphorylation of glycerol to G3P mainly in germinating seeds whereas cytosolic G3PDH catalyzes the reduction of dihydroxyacetone phosphate (DHAP) to G3P in different plant parts including seeds (Huang 1975). Hence, cytosolic G3PDH serves as a major link between carbohydrate and lipid metabolism, and diverts carbon flux towards lipid biosynthesis. However, G3PDH is strongly inhibited by G3P resulting in feedback inhibition and as a consequence restriction on G3P production. Studies have shown that this sensitivity to feedback inhibition can be overcome by introduction of mutant version of G3PDH in Escherichia coli and Arabidopsis (Bell and Cronan 1975; Shen et al. 2010b). Further, the seed-specific expression of a yeast cytosolic G3PDH in oilseed rape (Brassica napus L.) led to a 3–4 fold increase in the G3P levels in developing seeds and as consequence a 40% increase in the final seed oil content without changes in the protein content (Vigeolas et al. 2007). These results show that G3P supply co-limits oil accumulation in developing seeds. A seed oil increase of 40% is a remarkable achievement because of the fact that a single transgene expression has such an impact on a quantitative trait like oil content. Therefore, employing the yeast G3PDH or any structurally different G3PDH encoding genes can have important implications for developing strategies to enhance the overall level of oil in oilseed crops.

Lower oxygen conditions exist in the maturing seeds and as a result ATP supply is low. Under such low ATP conditions cytosolic pyrophosphates support the glycolysis process. Therefore, pyrophosphates act as a vital source of energy for driving cytosolic reactions that provide precursors for oil biosynthesis in the maturing seeds. Pyrophosphatase is the enzyme catalyzing the hydrolysis of pyrophosphates; as a consequence it has strong negative influence on the rate of cytosolic glycolysis and supply of precursors for seed storage lipid biosynthesis (Meyer et al. 2012). Seed-specific expression of cytosolic pyrophosphatase encoding gene in Arabidopsis resulted in the reduction of seed oil whereas seed-specific RNAi silencing of cytosolic pyrophosphatases resulted in 1–4% increase in oil content at the expense of protein content in transgenic seeds (Meyer et al. 2012). These findings clearly indicated the central role cytosolic pyrophosphate pools in developing seeds and preference for lipid accumulation during seed filling. Thus, seed-specific RNAi of cytosolic pyrophosphatases provide an effective strategy for enhancing seed oil content.

Apart from the sugar metabolism (glycolysis, OPPP), sugar transportation to the sink is a key factor determining the accumulation of seed storage products including oil. Vacuolar sugar transporters and tonoplast monosaccharide transporters (TMTs) are involved in the transport of monosachcharide sugars (Wormit et al. 2006; Wingenter et al. 2010) and sucrose across the tonoplast membrane into the vacuoles (Schulz et al. 2011). Glucose transportation in double (tmt1/tmt2) and triple (tmt1/tmt2/tmt3) mutants of TMT genes is impaired and as result reduced seed yield in mutants (Wormit et al. 2006). Interestingly, overexpression of TMT1 in double tmt mutant rescued the mutant phenotype, and it resulted in larger plants with increased seed yield and increased oil and protein contents compared to wild type plants. It was hypothesized that the enhanced seed storage compounds in the TMT1 overexpressing mutants was the result of enhanced compartmentalization of sugars in favour of seed storage compounds in mutants (Wingenter et al. 2010) and also could be due to the enhanced sink strength effect (Ludwig and Flügge 2013). It will be interesting and informative to explore more metabolite transporters and study effects of their seed-specific expression on seed oil content in oilseed crops.

Enhancement of oil yield by altering expression of transcription factors (TFs)

Lipid biosynthesis and its regulation is complex due to the participation of several enzymes and regulatory proteins in a hierarchical manner. Therefore overexpression of genes encoding the individual enzymes of the FA and TAG biosynthetic pathways may not yield greater changes in seed oil content whereas altered expression of transcription factors (TFs) which regulate a number of reactions in the oil biosynthetic pathway may provide a novel strategy for engineering high levels of seed oil which could be exploited for engineering oil yields (Shen et al. 2010a; Tan et al. 2011). A number of TFs expressed during seed development that control oil biosynthesis have been identified (Santos-Mendoza et al. 2008; Verdier and Thompson 2008; Elahi et al. 2016a). However, while employing TFs for engineering plants their expression need to be optimized because TFs may regulate the expression of multiple genes involved in different metabolic pathways and their ectopic expression might result in unintended or pleiotropic phenotypes (Century et al. 2008).

Firstly, Leafy cotyledon1 (LEC1) and LEC1-LIKE (L1L) are nuclear transcription factor Y subunit beta (NFYB)-type TFs known to function as key regulators of oil accumulation. Overexpression of AtLEC1, BnLEC1 and BnL1L in Arabidopsis and ZmLEC1 in maize resulted in significantly increased levels of seed oil in the transgenic plants. However, overexpression of these genes produced pleiotropic phenotypes such as lethality, poor seed germination, and stunted growth with dark green and narrow leaves (Mu et al. 2008; Shen et al. 2010a). Such growth and developmental abnormalities were overcome by inducible and seed-specific expression of LEC1 and L1L. Seed-specific expression of BnLEC1 and BnL1L enhanced seed oil content by 6–22% in transgenic canola plants without any detectable abnormalities (Tan et al. 2011). Recently, the constitutive expression of BnLEC1 in B. napus increased seed oil content by 7–16%, while the down-regulation of BnLEC1 in B. napus reduced oil content by 9–12% (Elahi et al. 2016b). The results of above-mentioned genetic and transgenic studies suggest that the seed-specific expression of LEC1 and L1L genes would be expedient for engineering oil content in oilseed crops. Secondly, ABSCISIC ACID INSENSITIVE3 (ABI3), FUSCA3 (FUS3), LEAFY COTYLEDON2 (LEC2) are B3 domain containing TFs known to play role in embryo maturation and ABA-regulated gene expressions in seeds. Among these LEC2 is the main regulator associated with seed oil accumulation as it regulates several genes including ABI3, FUS3, LEC1 and WRINKLED1 (WRI1) during the seed oil accumulation. Acetaldehyde inducible expression of LEC2 increased total FAs content by 6.8% in the leaves of tobacco plants (Andrianov et al. 2010) whereas lec2 mutant of Arabidopsis has 30% less oil with increased levels of starch and sucrose than wild type plants (Angeles-Núñez and Tiessen 2011). Finally, WRI1 and APETALA2 (AP2) are AP2/EREBP domain containing TFs. WRI1 regulates metabolism particularly glycolysis in developing seeds (Baud et al. 2009). Mutation in the WRI1 gene in A. thaliana caused 80% reduction in seed oil content due to the decreased glycolytic efficiency (Focks and Benning 1998). Expression of AtWRI1 and two WRI1 orthologs from Zea mays were able to restore the phenotypes of wri1-1 and wri1-4 mutants (Cernac and Benning 2004; Pouvreau et al. 2011). Overexpression of BnWRI in Arabidopsis increased seed mass and oil content by 10–40% (Liu et al. 2010) and overexpression of ZmWRI1 increased maize seed oil by 46% without affecting seed germination and plant growth (Shenet al. 2010a). It is suggested that overexpression of WRI1 alters the expression of key enzymes of the glycolysis pathway and as a consequence, enhanced accumulation of seed oil. The other AP2/EREBP domain containing TF, AP2, previously known to play regulatory role in flower identity and development, is shown to be a negative regulator of seed development (Jofuku et al. 2005). The ap2 plants showed an increase in seed oil content by 25–113% compared to wild type plants. The negative role of AP2 in oil accumulation is possibly due to limitation of carbohydrates supply required for oil biosynthesis. Further, GLABRA2 (GL2) is a class IV homeodomain-ZIP TF found to regulate seed oil content and trichome development in Arabidopsis. Shen et al. (2006) obtained a high oil phenotype GL2 loss of function mutant, p777, having 8% more oil in seeds compared to the wild type. Further, overexpression and suppression of the BnaC.GL2, an ortholog of GL2 gene in Arabidopsis resulted in increase in the seed oil content by 3.5–5.0 and 2.5–6.1% respectively, and reduction in the leaf trichome number (Chai et al. 2010). It was surprising to note that overexpression of BnaC.GL2 also resulted in enhanced seed oil. It was hypothesized that BnaC.GL2 interrupted endogenous GL2 function due to a phenomenon known as ‘squelching’, in which overexpression of GL2 and GL2-like genes would affect function of GL2 due to reduction in the ratio of the heterodimers to the homodimer of GL2 (Guan et al. 2008). Furthermore, Shi et al. (2012) demonstrated that although GL2 is expressed in both the embryo as well as in the seed coat, the loss of activity of GL2 in the seed coat and as a consequence loss of seed coat mucilage was responsible for increased seed oil accumulation, possibly due to allocation of more carbon for oil biosynthesis in the embryo that was previously used for development of seed coat mucilage. These results suggest that GL2 is a negative regulator of seed oil accumulation. From these studies, it can be concluded that that RNAi suppression of negative regulators of seed oil content such as AP2 and GL2 could be useful strategy for engineering oil content in oilseed crops.

Enhancement of oil yield by reducing breakdown of stored lipids

Majority of the genetic engineering attempts aimed at enhancing seed oil yield have focused on boosting synthetic capacity. Recent studies have shown that suppression of TAG hydrolysis increased TAG contents in seed as well as vegetative tissues (Kelly et al. 2013a, b; van Erp et al. 2014). The oil content of developing seeds is reported to decline with maturation and desiccation due to lipid hydrolysis. For instance, the oil content of rapeseed (B. napus) has been reported to decline by about 10% in the final stage of seed development (Chia et al. 2005). Lipid hydrolysis is catalyzed by TAG lipase to yield glycerol and free FAs. Lipase activities have been reported in the oil bodies of oilseed crops (Lin and Huang 1983; Huang and Moreau 1978). In developing seeds of Arabidopsis, Sugar-Dependent 1 (SDP1) is the major lipase implicated in oil body degradation. Arabidopsis mutants deficient in SDP1 accumulate high levels of oils, probably due to a reduction in TAG degradation (Eastmond 2006). Recently, it was shown that accumulation of TAG in sdp1 roots increased as the plants attained maturity up to a 50-fold increase over the wild type (Kelly et al. 2013a). Further, RNAi suppression of SDP1 in Arabidopsis resulted in enhanced seed oil by 3.2% compared to wild type plants (van Erp et al. 2014). In addition, RNAi suppression of the SDP1 gene family during seed development enhanced seed oil up to 8 and 30% respectively in B. napus and Jatropa curcas (Kelly et al. 2013b; Kim et al. 2014). These findings suggest that suppression of lipolysis during seed maturation is a promising method of enhancing the seed oil and this strategy can be extended to other oil crops to yield higher seed oil.

Enhancement of oil yield by extending the period of oil biosynthesis

Oil and protein biosynthesis occurs sequentially during seed development; oil biosynthesis is followed by protein synthesis. The maximum synthesis of oils occurs during the mid-phase of the seed development process. Recently, Kanai et al. (2015) demonstrated that sequential activation of genes involved in oil or protein biosynthesis during the seed maturation phase determines the oil/protein ratio in Arabidopsis seeds. They employed a novel approach of extending the oil biosynthesis period in seeds as well as suppressing protein synthesis to enhance seed oil production. In Arabidopsis, they extended the expression time of WRI1 using the FUS3 promoter which resulted in enhanced seed oil content up to 140% compared to wild type. Further, the extended expression of WRI1 in the 12s1.4, the major seed storage protein knockout mutant, enhanced the seed oil by 170% compared to the wild type (Kanai et al. 2015). The results of this study suggest that length of the oil synthesis period during seed development is one of the key factors that determine the final oil content. Moreover, application of this strategy of combining extended oil biosynthesis period with suppression of protein synthesis during seed maturation can be effective for engineering major oilseed crops to achieve large increases in seed oil production.

Enhancement of oil yield by introducing novel TAG synthesis pathway

In animals, apart from de novo synthesis, the monoacylglycerol (MAG) pathway is also involved in TAG synthesis mediated by monoacylglycerol acyltransferase (MGAT). This pathway plays a predominant role in salvaging the large amounts of MAGs and FAs released from the digestion of dietary lipids in the intestine (Phan and Tso 2001). Although, there is evidence for the presence and de novo production of MGAT in plant tissues (Reddy et al. 2010), there is no evidence for MAGs contributing to lipid synthesis in plant tissues. Petrie et al. (2012) explored the feasibility of utilizing this MAG pool as a substrate for TAG synthesis. Constitutive expression of Musmusculus MGAT1 and MGAT2 in Nicotiana benthamiana resulted in 7.3- and 9.2-fold increase in leaf TAG levels in the MGAT1 and MGAT2 transgenics, respectively, compared to the control plants. Moreover, TAG levels increased by 9.8-fold in transgenics expressing both MGAT1 and DGAT1 (Petrie et al. 2012). Recently, El Tahchy et al. (2015) explored the feasibility of employing the mammalian MGAT pathway for increasing the seed oil content in Arabidopsis. Overexpression of mouse MGAT2 resulted in 1.32-fold increase in seed oil content of Arabidopsis. In vitro assays in MGAT2 transgenics showed up to 3.9-fold increase in radio labeled DAG, suggesting that DAG is re-synthesized by salvaging the lipid breakdown product, MAG, due to MGAT2 activity in developing transgenic seeds. Thus, expression of mammalian MGATs in plants introduces a novel and independent TAG biosynthesis pathway that is complementary to the endogenous Kennedy pathway and other glycerolipid synthesis pathways. In future, the potential of introducing this novel oil biosynthetic pathway in major oilseed crops need to be explored.

Enhancement of oil yield by improving physiological conditions for oil biosynthesis

Studies in oilseed crops have shown that the synthesis of seed storage molecules in the developing seeds is strongly limited by the low oxygen supply prevailing within developing seeds and in particular lipid synthesis is more affected than starch synthesis (Vigeolas et al. 2003; Rolletschek et al. 2007). Hemoglobins are oxygen-binding proteins ubiquitously expressed in plants and supply oxygen to different tissues. They are divided into two different classes Hb1 and Hb2 based on their gene expression pattern and oxygen-binding properties (Hunt et al. 2001; Hoy and Hargrove 2008). The feasibility of employing these hemoglobin proteins to overcome the hypoxic conditions during seed maturation to enhance the seed oil accumulation was explored by Vigeolas et al. (2011). Seed-specific overexpression of AtHb2 led to a 40% increase in the total oil content in developing and mature seeds. The increase in total oil content was largely due to an increase in the polyunsaturated C18:2 and C18:3 acids. Moreover, AtHb2 overexpression led to an increase in the C18:2/C18:1 and C18:3/C18:2 ratios and in the unsaturation/saturation index of total seed lipids. The increase in oil content was attributed to a threefold higher energy state and twofold higher sucrose content in the developing seeds leading to stimulation in the rate of TAG synthesis. In contrast to this, overexpression of class 1 hemoglobins did not display any significant increase in the metabolite content of the seeds. These results provide the evidence for unique role of class 2 hemoglobins in seed oil production and in promoting the accumulation of polyunsaturated FAs by facilitating oxygen supply in developing seeds. The success achieved in model plant Arabidopsis need to be investigated in major oilseed crops for enhanced seed oil production.

Enhancement of oil yield by increasing sink size for oil accumulation

Oil biosynthesis and accumulation is part of seed development. Several studies have revealed the influence of seed development genes, such as SHB1, KLUH, IKU2, GL2, and AP2, on metabolism and accumulation of the storage components (Jofuku et al. 2005; Shen et al. 2006, 2010a; Adamski et al. 2009; Liu et al. 2010; Fatihi et al. 2013). The increased oil in large sized seeds of Arabidopsis transgenics expressing seed development genes as well as high oil maize and rice lines were reported to be due to increase in embryo sizes or oil bearing seed tissues (Zheng et al. 2008; Adamski et al. 2009; Fatihi et al. 2013; Tian et al. 2016) suggesting that an increase in oil bearing tissues without much increase in the non-oil bearing tissues of seed can increase the seed oil (Fortescue and Turner 2007; Fig. S2). Further, source productions have been shown to be responsive to sink demand size (Paul and Foyer 2001). Increasing the seed oil content of endosperm in crops bearing endosperm could be a novel alternative approach to enhancing seed oil (Gu et al. 2012). In Arabidopsis, SHB1 is a key TF involved in embryo and endosperm proliferation through the activation of MINI3 and IKU2 genes. In SHB1-overexpressing transgenics and in shb1-D mutants, the seed size as well as the accumulation of seed storage components were significantly increased whereas in the loss of function mutant (shb1), seed mass and storage components were reduced compared to the wild type. In SHB1-overexpressing plants, seed protein content was increased by twofold and oil content by 1.5-fold in contrast to 20% reduction in lipid content of shb1 plants compared to the wild type (Zhou et al. 2009). Further, overexpression of another seed development TF known to play role in integument elongation, KLUH, resulted in bigger seeds with enhanced oil content by 9–12% in both wild type and klu-2 mutant background (Adamski et al. 2009). Similarly, overexpression of IKU2, a gene involved in embryo growth, increased seed size as well as oil content by 35% (Fatihi et al. 2013). Transgenic lines expressing ZmLEC1 showed 14.4 and 35% increases in embryo size and oil content, respectively, compared to null plants (Shen et al. 2010a). BnGRF2 is a gene identified in high oil line by differential expression analyses of lines with contrasting oil content in oilseed rape. Its overexpression increased photosynthesis, seed mass and oil content. The seed oil production was enhanced by 40–50% (Liu et al. 2012). Taken together, the results of altered expression of seed development genes suggest that increased embryo to seed volume should exert a positive effect on accumulation of seed storage compounds, in some way perhaps by signaling the increase in overall sink size.

Alternately, the seed oil sink strength can be enhanced by increasing the number of oil storing bodies of seeds (Fig. S2). Until now, there have been few reports on the effect of oleosin on seed lipid content. It has been suggested that increasing the oleosin protein level in developing seeds may increase the oil storage ability of embryonic cells and thus, stimulate more oil biosynthesis and storage in the oil bodies (Siloto et al. 2006; Froissard et al. 2009). In Arabidopsis, overexpression of a castor Oleosin gene resulted in enhanced oil content which mainly consisted of hydroxyl FA accumulation in transgenic seeds (Lu et al. 2006) whereas suppression of Oleosin in Arabidopsis seeds results in larger oil bodies and reduced oil content (Shimada et al. 2008). The expression level of Oleosin in high oil content B. napus line was found to be greater than that of a low oil content line (Hu et al. 2009). Recently, Liu et al. (2013) investigated the effect of embryo-specific overexpression of two types of soybean Oleosin (encoding 24 kDa proteins) on the lipid content and the morphology of oil bodies in rice. Overexpression of two types of soybean Oleosin resulted in an increase in seed oil content up to 36.9 and 46.1%, respectively, compared to the control. However, the overexpression of endogenous 16 and 18 kDa Oleosins in rice did not enhance the lipid content of rice seeds. It is noteworthy, that overexpression of soybean Oleosins did not produce any detrimental effects on growth, development, seed germination and other yield traits that could be attributed to the embryo-specific overexpression of transgenes. Further, overexpression of soybean Oleosin genes in rice endosperm did not significantly change seed oil. Previously, the ectopic expression of a sunflower Oleosin in Arabidopsis resulted in the accumulation of the oleosin protein in non-target tissues rather than oil bodies (Beaudoin and Napier 2000). These pleiotropic effects were expected, as oil body biogenesis is a result of the coordinated synthesis of oleosins and TAGs. Hence, embryo-specific overexpression of oleosins is essential for improving the seed lipid content without causing undesirable pleiotropic effects. Furthermore, recently, electron microscopy of the seed anatomy of B. napus cultivars revealed that oil bodies covered 50, 27–38 and 20–28% of total aleuronic cell area in ultrahigh, high and low oil content lines, respectively (Hu et al. 2013). These results raise the possibility that higher seed oil content could be engineered by increasing the embryo-specific expression of appropriate oleosin genes in oil seed crops.

Enhancement of oil yield by multigene approach

Transgenic pyramiding or stacking of genes is gaining importance especially for manipulation of metabolic pathways and yield traits that are controlled by multiple genes (Wan 2015). Lines with improved oil content have been developed in B. napus, soybean, maize and B. juncea through the use of a single transgene (Vigeolas et al. 2007; Rao and Hildebrand 2009; Shen et al. 2010a; Savadi et al. 2015, 2016). However, achieving larger increase in seed oil will require combinations of genes. Recently, the effect of pyramiding of lipid biosynthetic genes on oil biosynthesis in plants has been explored (Table 1). Further investigations are needed to study the effect of altered expressions of different genes controlling oil metabolism, accumulation and sink strength in different combinations on oil yield. Moreover, the success of multigene engineering depends on the specific combinations of genes that result in an additive effect and leads to enhanced oil yield.

An important strategy for engineering the oil biosynthetic pathway could be the altered co-expression of specific combinations of genes to know whether such a multigene strategy results in an additive enhancement of seed oil in plants compared to the single gene strategy (van Erp et al. 2014). The combination of genes could be the genes involved in carbon flux towards oil biosynthesis along with genes involved in synthesis and/or degradation of oils in maturing seeds providing a ‘Push–Pull effect’ or ‘Push–Pull-Protect effect’ to oil accumulation in seeds. For example, coexpression of G3PDH and DGAT1 provide Push–Pull effect while overexpression of WRI1 and DGAT1 along with suppression of Lipase gene provides Push–Pull-Protect effect on seed oil.

Although many studies have been conducted on using this kind of multigene strategy for manipulation of oils in vegetative parts (Winichayakul et al. 2013; Fan et al. 2013; Vanhercke et al. 2014), there is only one report on such manipulation of seed oils (van Erp et al. 2014). Transgenic Arabidopsis lines expressing WRI1, DGAT1 and RNAiSDP1, and lines expressing WRI1 and DGAT1 produced significantly higher seed oil compared to the single genes carrying lines. Interestingly, the extent of oil accumulation was even more in triple transgene carrying transgenic lines compared to two transgene carrying lines which demonstrates that the Push–Pull-Protect strategy with appropriate gene combinations is better than the just the Push–Pull strategy (van Erp et al. 2014). Recently, Liu et al. (2016) attempted multigene engineering for increasing oil content in potato tubers by over expression of WRI1, DGAT1 and OLEOSIN. The expression of WRI1 and OLEOSIN were under the control of a tuber-specific (PATATIN) promoter whereas DGAT1 expression was driven by a constitutive (CaMV-35S) promoter. It resulted in over a 100-fold increase in TAG accumulation of potato tube. In addition, both phospholipids and galactolipids were also significantly enhanced in the potato tubers. The increase of lipids in transgenic tubers was at the expense of starch content suggesting that cellular carbon flux was diverted in favour of lipid synthesis by the action of the three transgenes. Researchers in this study cleverly have chosen three classes of genes regulating carbon flux for synthesis of precursors of lipid synthesis, TAG biosynthesis and oil storing bodies which resulted in significant increases in tuber oil content.

As an alternative to ‘Push–Pull’ and ‘Push–Pull-Protect’ strategies, the ‘Push–Pull-Pile more’ strategy may also be an important rational approach, because increases in seed size are also associated with increases in seed oil in transgenics expressing seed development genes like KLUH and IKU2 (Adamski et al. 2009; Fatihi et al. 2013). The majority of genetic and transgenic efforts have focused on increasing either seed mass or oil content but there are no reported efforts of improving both simultaneously. Since oil yield is a function of both seed mass and oil content, combined expression of genes controlling seed size and oil content could lead to increase in oil yield that is higher than the individual gene expressions. For instance, combined expression of SHB1 (a gene involved in seed development) and G3PDH (key gene in oil biosynthesis) genes in plants by co-transformation or transformation as gene cassette might result in higher oil yield per plant.

Precise engineering of oilseed crops to avoid unintended effects

Oil content is controlled by multiple enzymes, and the overexpression of these enzymes at the same level and in all tissues can be detrimental and result in abnormal phenotypes (Shen et al. 2010a; Liu et al. 2012; Farré et al. 2015; Li et al. 2015). Studies have shown that stage- and tissue-specific promoters driving transgene expression at specific stages and in specific tissues could minimize or avoid abnormal phenotypes in transgenics (Guo et al. 2014; Li et al. 2015). In seed oil content engineering, transgenes linked with seed-specific promoters such as Oleosin, Napin, Phaseolin, FA Elongase1, FUSCA3 and LEA gene promoters or synthetic ones driving expression during the later stages of seed development or seed maturation phase can be used to produce high seed oil containing transgenics without yield reductions or growth and development abnormalities. However, in a study expressing BnLEC1 and BnL1L in B. napus using the Napin promoter, a seed-specific promoter, resulted in pleiotropic effects. The modified version of Napin driving the expression of these genes in conditional manner produced normal phenotype like the non-transformed plants (Tan et al. 2011). So it may require fine tuning or modification of promoters to achieve controlled transgene expression and thus have optimized oil yields without detrimental effects. Further, in addition to the controlled spatial expression, the temporal expression of genes involved in oil biosynthesis is needed to avoid unintended effects in transgenics. For instance, the duration of TAG synthesis at rapid pace occurs during the mid-phase of seed development [6–12 days after flowering (DAF)] and protein synthesis occurs during late phase of seed development (10–14 DAF). Overexpression of WRI1 with the FUSCA3 promoter active during mid-phase of seed development, resulted in 40% increase in oil and 30% increase in protein whereas WRI1 expression with constitutive promoter 35S and late phase active LEA promoter did not change either oil or protein contents, suggesting that temporal tuning of the transgene expression is required for increasing oil content in plants without pleiotropic effects (Kanai et al. 2015).

In multigene engineering of the lipid metabolic pathway, a synchronized expression of transgene cassettes that form the metabolic pathway is needed to have high oil phenotypes without abnormalities. Multigene cassettes may comprise of both silencing and expression gene cassettes. However, multigene engineering often suffers from gene silencing and position effects resulting in less than expected yields and/or with unintended effects in transgenic plants (Halpin 2005; Zorrilla-López et al. 2013). There are a few approaches to overcome the challenges encountered in multigene engineering. One such approach is construction of gene cassette with a series of genes encoding a polyprotein containing different enzymes of a metabolic process (Hunt and Maiti 2001). A second approach is the insertion of transgene cassettes in targeted regions of the genome. These approaches may aid in coordinating the expression and/or silencing genes. The precise insertion and removal of genes in higher plants can be achieved using the emerging technologies like site-specific recombination technologies such as Cre/lox, FLP/FRT and R/RS systems (Kerbach et al. 2005; Li et al. 2009; Khan et al. 2011) and genome editing/gene targeting technologies like zinc finger nucleases and transcription activator-like (TAL) effector proteins (Gaj et al. 2013).


Energy needs of the world are escalating with the increasing population, urbanization and industrialization. In particular the demand for vegetable oils is increasing rapidly due to increasing per capita consumption and the need for production of biofuels to replace the conventional fossil fuels, which are predicted to exhaust in the near future. Increasing oil yields of oilseed crops is a promising option. However, this task is difficult due to the systematic regulation of metabolic pathways in plants during their evolution. Recent understanding of these regulatory mechanisms would enable us to harness the potential of the modern plant modification technologies like transgenic technology. Recent superfast developments in high throughput technologies like genomics, proteomics and metabolomics along with new genetic engineering strategies such as altered expression of transcription factors, introduction of novel pathway genes, seed development genes, mutligene engineering and extending length of seed oil biosynthesis will usher tailoring oilseed crops to meet the future energy needs of the world.



We acknowledge Dr. S. R. Bhat, ICAR-NRCPB, New Delhi, for useful discussion on the topic and Indian Council of Agricultural Research for the financial assistance.


The funding was provided by Indian Council of Agricultural Research (ICAR).

Compliance with ethical standards

Conflict of interest

The authors have no conflict of interest.

Supplementary material

10725_2016_236_MOESM1_ESM.doc (180 kb)
Supplementary material 1 (DOC 179 KB)
10725_2016_236_MOESM2_ESM.docx (27 kb)
Supplementary material 2 (DOCX 26 KB)


  1. Adamski NM, Anastasiou E, Eriksson S, O′Neill CM, Lenhard M (2009) Local maternal control of seed size by KLUH/CYP78A5-dependent growth signaling. Proc Nat Acad Sci USA 106:20115–20120PubMedPubMedCentralCrossRefGoogle Scholar
  2. Andre C, Froehlich JE, Moll MR, Benning C (2007) A heteromeric plastidic pyruvate kinase complex involved in seed oil biosynthesis in Arabidopsis. Plant Cell 19:2006–2022PubMedPubMedCentralCrossRefGoogle Scholar
  3. Andrianov V, Borisjuk N, Pogrebnyak N, Brinker A, Dixon J, Spitsin S, Flynn J, Matyszczuk P et al (2010) Tobacco as a production platform for biofuel: overexpression of Arabidopsis DGAT and LEC2 genes increases accumulation and shifts the composition of lipids in green biomass. Plant Biotechnol J 8:277–287PubMedCrossRefGoogle Scholar
  4. Angeles-Núñez JG, Tiessen A (2011) Mutation of the transcription factor LEAFY COTYLEDON 2 alters the chemical composition of Arabidopsis seeds, decreasing oil and protein content, while maintaining high levels of starch and sucrose in mature seeds. J Plant Physiol 168:1891–1900PubMedCrossRefGoogle Scholar
  5. Bao X, Ohlrogge JB (1999) Supply of FA is one limiting factor in the accumulation of triacylglycerol in developing embryos. Plant Physiol 120:1057–1062PubMedPubMedCentralCrossRefGoogle Scholar
  6. Baud S, Lepiniec L (2010) Physiological and developmental regulation of seed oil production. Prog Lipid Res 49:235–249PubMedCrossRefGoogle Scholar
  7. Baud S, Wuilleme S, Dubreucq B, De Almeida A, Vuagnat C, Lepiniec L et al (2007) Function of plastidial pyruvate kinases in seeds of Arabidopsis thaliana. Plant J 52(3):405–419PubMedCrossRefGoogle Scholar
  8. Baud S, Dubreucq B, Miquel M, Rochat C, Lepiniec L (2008) Storage reserve accumulation in Arabidopsis: metabolic and developmental control of seed filling. Arabidopsis Book 6:e0113PubMedPubMedCentralCrossRefGoogle Scholar
  9. Baud S, Wuillème S, To A, Rochat C, Lepiniec L (2009) Role of WRINKLED1 in the transcriptional regulation of glycolytic and FA biosynthetic genes in Arabidopsis. Plant J 60:933–947PubMedCrossRefGoogle Scholar
  10. Beaudoin F, Napier JA (2000) The targeting and accumulation of ectopically expressed oleosin in non-seed tissues of Arabidopsis thaliana. Planta 210:439–445PubMedCrossRefGoogle Scholar
  11. Bell RM, Cronan JEJr (1975) Mutants of Escherichia coli defective in membrane phospholipid synthesis phenotypic suppression of sn-glycerol-3-phosphate acyltransferase Km mutants by loss of feedback inhibition of the biosynthetic sn-glycerol-3-phosphate dehydrogenase. J Biol Chem 250:7153–7158PubMedGoogle Scholar
  12. Bhat SR (2010) Transgenics for increasing productivity of crops. J Plant Biochem Biotechnol 19:1–7CrossRefGoogle Scholar
  13. Cao Z, Gao H, Liu M, Jiao P (2006) Engineering the acetyl-CoA transportation system of Candida tropicalis enhances the production of dicarboxylic acid. Biotechnol J 1(1):68–74PubMedCrossRefGoogle Scholar
  14. Century K, Reuber TL, Ratcliffe OJ (2008) Regulating the regulators: the future prospects for transcription-factor-based agricultural biotechnology products. Plant Physiol 147:20–29PubMedPubMedCentralCrossRefGoogle Scholar
  15. Cernac A, Benning C (2004) WRINKLED1 encodes an AP2/EREB domain protein involved in the control of storage compound biosynthesis in Arabidopsis. Plant J 40:575–585PubMedCrossRefGoogle Scholar
  16. Chai G, Bai Z, Wei F, King GJ, Wang C, Shi L, Dong C, Chen H, Liu S (2010) Brassica GLABRA2 genes: analysis of function related to seed oil content and development of functional markers. Theor Appl Genet 120:1597–1610PubMedCrossRefGoogle Scholar
  17. Chandran D, Sankararamasubramanian HM, Kumar MA, Parida A (2014) Differential expression analysis of transcripts related to oil metabolism in maturing seeds of Jatropha curcas L. Physiol Mol Biol Plants 20(2):181–190PubMedPubMedCentralCrossRefGoogle Scholar
  18. Chen L, Hao L, Parry MA, Phillips AL, Hu YG (2014) Progress in TILLING as a tool for functional genomics and improvement of crops. J Integrative Plant Biol 56:425–443CrossRefGoogle Scholar
  19. Chen S, Lei Y, Xu X, Huang J, Jiang H, Wang J, Li Y (2015) The peanut (Arachis hypogaea L.) gene AhLPAT2 increases the lipid content of transgenic Arabidopsis seeds. PloS One 10(8):e0136170PubMedPubMedCentralCrossRefGoogle Scholar
  20. Chia TY, Pike MJ, Rawsthorne S (2005) Storage oil breakdown during embryo development of Brassica napus (L.) J Exp Bot 56(415):1285–1296PubMedCrossRefGoogle Scholar
  21. Dahlqvist A, Stahl U, Lenman M, Banas A, Lee M, Sandager L, Ronne H, Stymne S (2000) Phospholipid: diacylglycerolacyltransferase: an enzyme that catalyzes the acyl-CoA-independent formation of triacylglycerol in yeast and plants. Proc Nat Acad Sci USA 97:6487–6492PubMedPubMedCentralCrossRefGoogle Scholar
  22. DeLuca H (2012) (ed) The fat-soluble vitamins, vol 2. Springer Science & Business Media, New YorkGoogle Scholar
  23. Eastmond PJ (2006) SUGAR-DEPENDENT1 encodes a patatin domain triacylglycerol lipase that initiates storage oil breakdown in germinating Arabidopsis seeds. Plant Cell 18(3):665–675PubMedPubMedCentralCrossRefGoogle Scholar
  24. El Tahchy A, Petrie JR, Shrestha P, Vanhercke T, Singh SP (2015) Expression of mouse MGAT in Arabidopsis results in increased lipid accumulation in seeds. Front Plant Sci 6:1180PubMedPubMedCentralCrossRefGoogle Scholar
  25. Elahi N, Duncan RW, Stasolla C (2016a) Molecular regulation of seed oil accumulation. J Adv Nutri Human Metabol 2:1–11Google Scholar
  26. Elahi N, Duncan RW, Stasolla C (2016b) Modification of oil and glucosinolate content in canola seeds with altered expression of Brassica napus LEAFY COTYLEDON1. Plant Physiol Biochemi 100:52–63.CrossRefGoogle Scholar
  27. Eskandari M, Cober ER, Rajcan I (2013) Genetic control of soybean seed oil: I QTL and genes associated with seed oil concentration in RIL populations derived from crossing moderately high-oil parents. Theor Appl Genet 126(2):483–495PubMedCrossRefGoogle Scholar
  28. Fan J, Yan C, Zhang X, Xu C (2013) Dual role for phospholipid: diacylglycerol acyltransferase: enhancing FA synthesis and diverting FAs from membrane lipids to triacylglycerol in Arabidopsis leaves. Plant Cell 25(9):3506–3518PubMedPubMedCentralCrossRefGoogle Scholar
  29. Farré G, Twyman RM, Christou P, Capell T, Zhu C (2015) Knowledge-driven approaches for engineering complex metabolic pathways in plants. Curr Opin Biotechnol 32:54–60PubMedCrossRefGoogle Scholar
  30. Fatihi A, Zbierzak AM, Dormann P (2013) Alterations in seed development gene expression affect size and oil content of Arabidopsis seeds. Plant Physiol 163:973–985PubMedPubMedCentralCrossRefGoogle Scholar
  31. 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 118:91–101PubMedPubMedCentralCrossRefGoogle Scholar
  32. Fortescue JA, Turner DW (2007) Changes in seed size and oil accumulation in Brassica napus L. by manipulating the source–sink ratio and excluding light from the developing siliques. Crop Pasture Sci 58:413–424.CrossRefGoogle Scholar
  33. Froissard M, D’Andrea S, Boulard C, Chardot T (2009) Heterologous expression of AtClo1, a plant oil body protein, induces lipid accumulation in yeast. FEMS Yeast Res 9:428–438PubMedCrossRefGoogle Scholar
  34. Gaj T, Gersbach CA, Barbas CF (2013) ZFN TALEN and CRISPR/Cas-based methods for genome engineering. Trends Biotechnol 31:397–405PubMedPubMedCentralCrossRefGoogle Scholar
  35. Gibon Y, Vigeolas H, Tiessen A, Geigenberger P, Stitt M (2002) Sensitive and high throughput metabolite assays for inorganic pyrophosphate, ADPGlc, nucleotide phosphates, and glycolytic intermediates based on a novel enzymic cycling system. Plant J 30:221–235PubMedCrossRefGoogle Scholar
  36. Gu K, Yi C, Tian D, Sangha JS, Hong Y, Yin Z (2012) Expression of FA and lipid biosynthetic genes in developing endosperm of Jatropha curcas. Biotechnol Biofuels 5(1):47PubMedPubMedCentralCrossRefGoogle Scholar
  37. Guan XY, Li QJ, Shan CM, Wang S, Mao YB, Wang LJ, Chen XY (2008) The HD-Zip IV gene GaHOX1 from cotton is a functional homologue of the Arabidopsis GLABRA2. Physiol Plant 134:174–182PubMedCrossRefGoogle Scholar
  38. Guo L, Ma F, Wei F, Fanella B, Allen DK, Wang X (2014) Cytosolic phosphorylating glyceraldehyde-3-phosphate dehydrogenases affect Arabidopsis cellular metabolism and promote seed oil accumulation. Plant Cell 26:3023–3035PubMedPubMedCentralCrossRefGoogle Scholar
  39. Gupta PK (2008) Molecular biology and genetic engineering. Deep and Deep Publications, New DelhiGoogle Scholar
  40. Guschina IA, Everard JD, Kinney AJ, Quant PA, Harwood JL (2014) Studies on the regulation of lipid biosynthesis in plants: application of control analysis to soybean. Biochim Biophys Acta (BBA) Biomembr 1838:1488–1500CrossRefGoogle Scholar
  41. Halpin C (2005) Gene stacking in transgenic plants–the challenge for 21st century plant biotechnology. Plant Biotechnol J 3:141–155PubMedCrossRefGoogle Scholar
  42. Harwood JL, Guschina IA (2013) Regulation of lipid synthesis in oil crops. FEBS Lett 587:2079–2081PubMedCrossRefGoogle Scholar
  43. Hoy JA, Hargrove MS (2008) The structure and function of plant hemoglobins. Plant Physiol Biochem 46:371–379PubMedCrossRefGoogle Scholar
  44. Hu Z, Wang X, Zhan G, Liu G, Hua W, Wang H (2009) Unusually large oilbodies are highly correlated with lower oil content in Brassica napus. Plant Cell Rep 28:541–549PubMedCrossRefGoogle Scholar
  45. Hu Z-Y, Hua W, Zhang L, Deng L-B et al (2013) Seed structure characteristics to form ultrahigh oil content in rapeseed. PLoS One 8:e62099PubMedPubMedCentralCrossRefGoogle Scholar
  46. Hua W, Li R, Zhan G, Liu J, Li J, Wang X, Liu, Wang H (2012) Maternal control of seed oil content in B. napus: the role of silique wall photosynthesis. Plant J 69:432–444PubMedCrossRefGoogle Scholar
  47. Hua W, Liu J, Wang H (2016) Molecular regulation and genetic improvement of seed oil content in Brassica napus L. Front Agr Sci Eng 3:186–194CrossRefGoogle Scholar
  48. Huang AH (1975) Enzymes of glycerol metabolism in the storage tissues of fatty seedlings. Plant Physiol 55:555–558PubMedPubMedCentralCrossRefGoogle Scholar
  49. Huang AH, Moreau RA (1978) Lipases in the storage tissues of peanut and other oil seeds during germination. Planta 141:111–116PubMedCrossRefGoogle Scholar
  50. Hunt AG, Maiti IB (2001) Strategies for expressing multiple foreign genes in plants as polycistronic constructs. In Vitro Cell Dev Biol Plant 37:313–320CrossRefGoogle Scholar
  51. Issariyakul T, Dalai AK (2014) Biodiesel from vegetable oils. Renew Sustain Energy Rev 31:446–471CrossRefGoogle Scholar
  52. Jain RK, Coffey M, Lai K, Kumar A, MacKenzie SL (2000) Enhancement of seed oil content by expression of glycerol-3- phosphate acyltransferase genes. Biochem Soc Trans 28:958–961PubMedCrossRefGoogle Scholar
  53. Jofuku KD, Omidyar PK, Gee Z, Okamuro JK (2005) Control of seed mass and seed yield by the floral homeotic gene APETALA2. Proc Nat Aca Sci USA 102:3117CrossRefGoogle Scholar
  54. Kanai M, Mano S, Kondo M, Hayashi M, Nishimura M (2015) Extension of oil biosynthesis during the mid-phase of seed development enhances oil content in Arabidopsis seeds. Plant Biotechnol J. doi: 10.1111/pbi.12489 PubMedGoogle Scholar
  55. Kashyap PL, Sanghera GS, Wani SH, Shafi W, Kumar S et al (2011) Genes of microorganisms: paving way to tailor next generation fungal disease resistant crop plants. Not Sci Biol 3:147–157Google Scholar
  56. Katavic V, Reed DW, Taylor DC, Giblin EM, Barton DL, Zou J et al (1995) Alteration of seed FA composition by an ethyl methane sulfonate–induced mutation in Arabidopsis thaliana affecting diacylglycerol acyltransferase activity. Plant Physiol 108:399–409PubMedPubMedCentralCrossRefGoogle Scholar
  57. Kelly AA, Erp HV, Quettier AL, Shaw E, Menard G, Kurup S, Eastmond PJ (2013a) The SUGAR-DEPENDENT1 lipase limits triacylglycerol accumulation in vegetative tissues of Arabidopsis. Plant Physiol 162:1282–1289PubMedPubMedCentralCrossRefGoogle Scholar
  58. Kelly AA, Shaw E, Powers SJ, Kurup S, Eastmond PJ (2013b) Suppression of the SUGAR-DEPENDENT1 triacylglycerol lipase family during seed development enhances oil yield in oilseed rape (Brassica napus L.). Plant Biotechnol J 11:355–361PubMedCrossRefGoogle Scholar
  59. Kerbach S, Lörz H, Becker D (2005) Site-specific recombination in Zea mays. Theor Appl Genet 111:1608–1616PubMedCrossRefGoogle Scholar
  60. Khan RS, Nakamura I, Mii M (2011) Development of disease-resistant marker-free tomato by R/RS site-specific recombination. Plant Cell Rep 30:1041–1053PubMedCrossRefGoogle Scholar
  61. Kim MJ, Yang SW, Mao HZ, Veena SP, Yin JL, Chua NH (2014) Gene silencing of sugar-dependent1 (JcSDP1) encoding a patatin-domain triacylglycerol lipase enhances seed oil accumulation in Jatropha curcas. Biotechnol Biofuels 7:36PubMedPubMedCentralCrossRefGoogle Scholar
  62. Klaus D, Ohlrogge JB, Neuhaus HE, Dörmann P (2004) Increased FA production in potato by engineering of acetyl-CoA carboxylase. Planta 219:389–396PubMedCrossRefGoogle Scholar
  63. Li Z, Xing A, Moon BP, McCardell RP, Mills K, Falco SC (2009) Site-specific integration of transgenes in soybean via recombinase-mediated DNA cassette exchange. Plant Physiol 151:1087–1095PubMedPubMedCentralCrossRefGoogle Scholar
  64. Li M, Wei F, Tawfall A, Tang M, Saettele A, Wang X (2015) Overexpression of patatin related phospholipase AIIIδ altered plant growth and increased seed oil content in camelina. Plant Biotechnol J 13:766–778PubMedCrossRefGoogle Scholar
  65. Li-Beisson Y, Shorrosh B, Beisson F, Andersson MX, Arondel V, Bates PD et al (2013) Acyl-lipid metabolism. Arabidopsis Book 11:e0161PubMedPubMedCentralCrossRefGoogle Scholar
  66. Lin ECC (1976) Glycerol dissimilation and its regulation in bacteria. Ann Rev Microbiol 30:535–578CrossRefGoogle Scholar
  67. Lin YH, Huang AH (1983) Lipase in lipid bodies of cotyledons of rape and mustard seedlings. Arch Biochem Biophys 225:360–369PubMedCrossRefGoogle Scholar
  68. Liu J, Hua W, Zhan G, Wei F, Wang X, Liu G, Wang H (2010) Increasing seed mass and oil content in transgenic Arabidopsis by the overexpression of wri1-like gene from B. napus. Plant Physiol Biochem 48:9–15PubMedCrossRefGoogle Scholar
  69. Liu J, Hua W, Yang HL, Zhan GM, Li RJ, Deng LB et al (2012) The BnGRF2 gene (GRF2-like gene from Brassica napus) enhances seed oil production through regulating cell number and plant photosynthesis. J Exp Bot 63:3727–3740PubMedPubMedCentralCrossRefGoogle Scholar
  70. Liu WX, Liu HL, Qu LQ (2013) Embryo-specific expression of soybean oleosin altered oil body morphogenesis and increased lipid content in transgenic rice seeds. Theor Appl Genet 126:2289–2297PubMedCrossRefGoogle Scholar
  71. Liu F, Xia Y, Wu L, Fu D, Hayward A, Luo J et al (2015) Enhanced seed oil content by overexpressing genes related to triacylglyceride synthesis. Gene 557:163–171PubMedCrossRefGoogle Scholar
  72. Liu Q, Guo Q, Akbar S, Zhi Y, El Tahchy A, Mitchell M et al (2016) Genetic enhancement of oil content in potato tuber (Solanum tuberosum L.) through an integrated metabolic engineering strategy. Plant Biotechnol J. doi: 10.1111/pbi.12590 Google Scholar
  73. Lu C, Fulda M, Wallis JG, Browse J (2006) A high-throughput screen for genes from castor that boost hydroxy FA accumulation in seed oils of transgenic Arabidopsis. Plant J 45:847–856PubMedCrossRefGoogle Scholar
  74. Maheshwar P, Kovalchuk I (2014) Genetic engineering of oilseed crops. Biocatal Agricult Biotechnol 3:31–37Google Scholar
  75. Maisonneuve S, Bessoule J, Lessire R, Delseny M, Roscoe TJ (2010) Expression of rapeseed microsomal Lysophosphatidic acid acyltransferase isozymes enhances seed oil content in Arabidopsis. Plant Physiol 152:670–684PubMedPubMedCentralCrossRefGoogle Scholar
  76. Marillia E, Micallef BJ, Micallef M, Weninger A, Pedersen KK, Zou J, Taylor DC (2003) Biochemical and physiological studies of Arabidopsis thaliana transgenic lines with repressed expression of the mitochondrial pyruvate dehydrogenase kinase. J Exp Bot 54:259–270PubMedCrossRefGoogle Scholar
  77. Meyer K, Kinney AJ (2010) Biosynthesis and biotechnology of seed lipids including sterols carotenoids and tocochromanols. In: Lipids in photosynthesis, Springer Netherlands, pp 407–444Google Scholar
  78. Meyer K, Stecca KL, Ewell-Hicks K, Allen SM, Everard JD (2012) Oil and protein accumulation in developing seeds is influenced by the expression of a cytosolic pyrophosphatase in Arabidopsis. Plant Physiol 159:1221–1234PubMedPubMedCentralCrossRefGoogle Scholar
  79. Mu J, Tan H, Zheng Q, Fu F, Liang Y, Zhang J, Yang X, Wang T, Chong K, Wang XJ, Zuo J (2008) LEAFY COTYLEDON1 is a key regulator of FA biosynthesis in Arabidopsis. Plant Physiol 148:1042–1054PubMedPubMedCentralCrossRefGoogle Scholar
  80. Murphy DJ (1995) The use of conventional and molecular genetics to produce new diversity in seed oil composition for the use of plant breeders—progress problems and future prospects. In: The methodology of plant genetic manipulation: criteria for decision making, Springer Netherlands, pp 433–440Google Scholar
  81. Murphy DJ (1996) Engineering oil production in rapeseed and other oil crops. Trends Biotechnol 14:206–213CrossRefGoogle Scholar
  82. Murphy DJ (2014) Using modern plant breeding to improve the nutritional and technological qualities of oil crops. OCL 21:D607CrossRefGoogle Scholar
  83. Nguyen HT, Park H, Koster KL, Cahoon RE, Nguyen HTM et al (2015) Redirection of metabolic flux for high levels of omega-7monounsaturated FA accumulation in camelina seeds. Plant Biotechnol J 13:38–50PubMedCrossRefGoogle Scholar
  84. OECD/FAO (2015) OECD-FAO agricultural outlook. OECD agriculture statistics (database). doi: 10.1787/agr-outl-data-en
  85. Ohlrogge JB, Jaworski JG (1997) Regulation of fatty acid synthesis. Annu Rev Plant Physiol Plant Mol Biol 48:109–136PubMedCrossRefGoogle Scholar
  86. Pandey MK, Wang ML, Qiao L, Feng S, Khera P, Wang H et al (2014) Identification of QTLs associated with oil content and mapping FAD2 genes and their relative contribution to oil quality in peanut (Arachis hypogaea L.). BMC Genet 15:133PubMedPubMedCentralCrossRefGoogle Scholar
  87. Paul MJ, Foyer CH (2001) Sink regulation of photosynthesis. J Exp Bot 52:1383–1400PubMedCrossRefGoogle Scholar
  88. Perry HY, Bligny R, Gout E, Harwood JL (1999) Changes in Kennedy pathway intermediates associated with increased triacylglycerol synthesis in oilseed rape. Phytochem 52:799–804CrossRefGoogle Scholar
  89. Petrie JR, Vanhercke T, Shrestha P, El-Tahchy A, White A, Zhou XR, Liu Q, Mansour M, Nichols PD, Singh SP (2012) Recruiting a new substrate for triacylglycerol synthesis in plants: the monoacylglycerolacyltransferase pathway. PLoS One 7:e35214PubMedPubMedCentralCrossRefGoogle Scholar
  90. Phan CT, Tso P (2001) Intestinal lipid absorption and transport. Front Biosci 6:D299–D319PubMedCrossRefGoogle Scholar
  91. Pouvreau B, Baud S, Vernoud V, Morin V, Py C, Gendrot G, Pichon J-P, Rouster J, Paul W, Rogowsky PM (2011) Duplicate maize Wrinkled1 transcription factors activate target genes involved in seed oil biosynthesis. Plant Physiol 156:674–686PubMedPubMedCentralCrossRefGoogle Scholar
  92. Rahman H, Harwood JL, Weselake R (2013) Increasing seed oilcontent in Brassica species through breeding and biotechnology. Lipid Technol 25:182–185CrossRefGoogle Scholar
  93. Ramli US, Salas JJ, Quant PA, Harwood JL (2005) Metabolic control analysis reveals an important role for diacylglycerol acyltransferase in olive but not in oil palm lipid accumulation. FEBS J 272:5764–5770PubMedCrossRefGoogle Scholar
  94. Rao SS, Hildebrand D (2009) Changes in oil content of transgenic soybeans expressing the yeast SLC1 gene. Lipids 44:945–951PubMedCrossRefGoogle Scholar
  95. Reddy VS, Rao DKV, Rajasekharan R (2010) Functional characterization of lysophosphatidic acid phosphatase from Arabidopsis thaliana. Biochim Biophys Acta 1801:455–461PubMedCrossRefGoogle Scholar
  96. Roesler K, Shintani D, Savage L, Boddupalli S, Ohlrogge J (1997) Targeting of the Arabidopsis homomeric acetyl-coenzyme A carboxylase to plastids of rapeseeds. Plant Physiol 113:75–81PubMedPubMedCentralCrossRefGoogle Scholar
  97. Roesler K, Shen B, Bermudez E, Li C, Hunt J, Damude HG, Feng L (2016) An improved variant of soybean type 1 diacylglycerol acyltransferase increases the oil content and decreases the soluble carbohydrate content of soybeans. Plant Physiol 171:878–893PubMedPubMedCentralGoogle Scholar
  98. Rolletschek H, Borisjuk L, Sánchez-García A, Gotor C, Romero LC, Martínez-Rivas JM, Mancha M (2007) Temperature-dependent endogenous oxygen concentration regulates microsomal oleate desaturase in developing sunflower seeds. J Exp Bot 58:3171–3181PubMedCrossRefGoogle Scholar
  99. Ruuska SA, Girke T, Benning C, Ohlrogge JB (2002) Contrapuntal networks of gene expression during Arabidopsis seed filling. Plant Cell 14:1191–1206PubMedPubMedCentralCrossRefGoogle Scholar
  100. Samarth NB, Mahanwar PA (2015) Modified vegetable oil based additives as a future polymeric material-review. Open J Org Pol Mat 5:1–22CrossRefGoogle Scholar
  101. Sanghera GS, Kashyap PL, Singh G, da Silva JAT (2011) Transgenics: fast track to plant stress amelioration. Transgenic Plant J 5:1–26Google Scholar
  102. Santos-Mendoza M, Dubreucq B, Baud S, Parcy F, Caboche M, Lepiniec L (2008) Deciphering gene regulatory networks that control seed development and maturation in Arabidopsis. Plant J 54:608–620PubMedCrossRefGoogle Scholar
  103. Sasaki Y, Nagano Y (2004) Plant acetyl-CoA carboxylase: structure, biosynthesis, regulation, and gene manipulation for plant breeding. Biosci Biotechnol Biochemi 68:1175–1184CrossRefGoogle Scholar
  104. Savadi S, Naresh V, Kumar V, Bhat SR (2015) Effect of overexpression of Arabidopsis thaliana SHB1 and KLUH genes on seed weight and yield contributing traits in Indian mustard (Brassica juncea L. (Czern.). Indian J Genet 75:349–356Google Scholar
  105. Savadi S, Naresh V, Kumar V, Bhat SR (2016) Seed-specific overexpression of Arabidopsis DGAT1 in Indian mustard (Brassica juncea) increases seed oil content and seed weight. Botany 94:177–184CrossRefGoogle Scholar
  106. Schulz A, Beyhl D, Marten I, Wormit A, Neuhaus E, Poschet G et al (2011) Proton-driven sucrose symport and antiport are provided by the vacuolar transporters SUC4 and TMT1/2. Plant J 68:129–136PubMedCrossRefGoogle Scholar
  107. Sharma A, Chauhan RS (2012) In silico identification and comparative genomics of candidate genes involved in biosynthesis and accumulation of seed oil in plants. Comparat Funct Genom 2012:914843Google Scholar
  108. Sharma N, Anderson M, Kumar A, Zhang Y, Giblin EM, Abrams SR et al (2008) Transgenic increases in seed oil content are associated with the differential expression of novel Brassica-specific transcripts. BMC Genom 9:619CrossRefGoogle Scholar
  109. Shen B, Sinkevicius KW, Selinger DA, Tarczynski MC (2006) The homeobox gene GLABRA2 affects seed oil content in Arabidopsis. Plant Mol Biol 60:377–387PubMedCrossRefGoogle Scholar
  110. Shen B, Allen WB, Zheng P, Li C, Glassman K, Ranch J, Nubel D, Tarczynski MC (2010a) Expression of ZmLEC1 and ZmWRI1 increases seed oil production in maize. Plant Physiol 153:980–987PubMedPubMedCentralCrossRefGoogle Scholar
  111. Shen W, Li JQ, Dauk M, Huang Y, Periappuram C, Wei Y, Zou J (2010b) Metabolic and transcriptional responses of glycerolipid pathways to a perturbation of glycerol-3-phosphate metabolism in Arabidopsis. J Biol Chem 285:22957–22965PubMedPubMedCentralCrossRefGoogle Scholar
  112. Shi L, Katavic V, Yu Y, Kunst L, Haughn G (2012) Arabidopsis glabra2 mutant seeds deficient in mucilage biosynthesis produce more oil. Plant J 69:37–46PubMedCrossRefGoogle Scholar
  113. Shimada TL, Shimada T, Takahashi H, Fukao Y, Hara-Nishimura I (2008) A novel role for oleosins in freezing tolerance of oilseeds in Arabidopsis thaliana. Plant J 55:798–809PubMedCrossRefGoogle Scholar
  114. Shintani DK, Ohlrogge JB (1995) Feedback inhibition of FA synthesis in tobacco suspension cells. Plant J 7:577–587CrossRefGoogle Scholar
  115. Siloto RMP, Findlay K, Lopez-Villalobos A, Yeung EC, Nykiforuk CL, Moloney MM (2006) The accumulation of oleosins determines the size of seed oilbodies in Arabidopsis. Plant Cell 18:1961–1974PubMedPubMedCentralCrossRefGoogle Scholar
  116. Stahl U, Carlsson AS, Lenman M, Dahlqvist A, Huang B, Banas W et al (2004) Cloning and functional characterization of a phospholipid:diacylglycerol acyltransferase from Arabidopsis. Plant Physiol 135:1324–1335PubMedPubMedCentralCrossRefGoogle Scholar
  117. Tan HL, Yang XH, Zhang FX, Zheng X, Qu CM, Mu JY, Fu FY, Li JN, Guan RZ, Zhang HS, Wang GD, Zuo JR (2011) Enhance seed oil production in canola by conditional expression of B. napus LEAFY COTYLEDON1 (BnLEC1) and LEC1-LIKE (BnL1L) in developing seeds. Plant Physiol 156:1577–1588PubMedPubMedCentralCrossRefGoogle Scholar
  118. Taylor DC, Katavic V, Zou J, MacKenzie SL, Keller WA (2002) Field testing of transgenic rapeseed cv Hero transformed with a yeast sn-2 acyltransferase results in increased oil content erucic acid content and seed yield. Mol Breed 8:317–322CrossRefGoogle Scholar
  119. Taylor DC, Zhang Y, Kumar A, Francis A, Giblin EM, Barton D, Ferrie JR, Laroche A, Shah S, Zhu W et al (2009) Molecular modification of triacylglycerol accumulation by overexpression of DGAT1 to produce canola with increased seed oil content under field conditions. Botany 87:533–543CrossRefGoogle Scholar
  120. Teh L, Möllers C (2016) Genetic variation and inheritance of phytosterol and oil content in a doubled haploid population derived from the winter oilseed rape Sansibar × Oase cross. Theor Appl Genet 129:181–199PubMedCrossRefGoogle Scholar
  121. Thelen JJ, Ohlrogge JB (2002) Both antisense and sense expression of Biotin carboxyl carrier protein isoform 2 inactivates the plastid acetyl coenzyme-A carboxylase in Arabidopsis thaliana. Plant J 32:419–431PubMedCrossRefGoogle Scholar
  122. Thelen JJ, Miernyk JA, Randall DD (1998) Partial purification and characterization of the maize mitochondrial pyruvate dehydrogenase complex. Plant Physiol 116:1443–1450PubMedPubMedCentralCrossRefGoogle Scholar
  123. Tian Y, Zhang M, Hu X, Wang L, Dai J, Xu Y, Chen F (2016) Overexpression of CYP78A98 a cytochrome P450 gene from Jatropha curcas L., increases seed size of transgenic tobacco. Electron J Biotechnol 19:15–22CrossRefGoogle Scholar
  124. Van Camp W (2005) Yield enhancement genes: seeds for growth. Curr Opi Biotechnol 16:147–153CrossRefGoogle Scholar
  125. van Erp H, Kelly AA, Menard G, Eastmond PJ (2014) Multigene engineering of triacylglycerol metabolism boosts seed oil content in Arabidopsis. Plant Physiol 165:30–36PubMedPubMedCentralCrossRefGoogle Scholar
  126. Vanhercke T, El Tahchy A, Shrestha P, Zhou XR, Singh SP, Petrie JR (2013) Synergistic effect of WRI1 and DGAT1 coexpression on triacylglycerol biosynthesis in plants. FEBS Lett 587:364–369PubMedCrossRefGoogle Scholar
  127. Vanhercke T, El Tahchy A, Liu Q, Zhou XR, Shrestha P, Divi UK et al (2014) Metabolic engineering of biomass for high energy density: oilseed-like triacylglycerol yields from plant leaves. Plant Biotechnol J 12:231–239PubMedCrossRefGoogle Scholar
  128. Verdier J, Thompson RD (2008) Transcriptional regulation of storage protein synthesis during dicotyledon seed filling. Plant Cell Physiol 49:1263–1271PubMedCrossRefGoogle Scholar
  129. Verwoert II, van der Linden KH, Walsh MC, Nijkamp HJJ, Stuitje AR (1995) Modification of Brassica napus seed oil by expression of the Escherichia coli fabH gene encoding 3-ketoacyl-acyl carrier protein synthase III. Plant Mol Biol 27:875–886PubMedCrossRefGoogle Scholar
  130. Vigeolas H, Geigenberger P (2004) Increased levels of glycerol-3-phosphate lead to a stimulation of flux into triacylglycerol synthesis after supplying glycerol to developing seeds of B. napus L. in planta. Planta 219:827–835PubMedCrossRefGoogle Scholar
  131. Vigeolas H, van Dongen JT, Waldeck P, Hühn D, Geigenberger P (2003) Lipid storage metabolism is limited by the prevailing low oxygen concentrations within developing seeds of oilseed rape. Plant Physiol 133:2048–2060PubMedPubMedCentralCrossRefGoogle Scholar
  132. Vigeolas H, Waldeck P, Zank T, Geigenberger P (2007) Increasing seed oil content in oil-seed rape (B. napus L.) by overexpression of a yeast glycerol-3-phosphate dehydrogenase under the control of a seed-specific promoter. Plant Biotechnol J 5:431–441PubMedCrossRefGoogle Scholar
  133. Vigeolas H, Huhn D, Geigenberger P (2011) NonsymbioticHemoglobin-2 leads to an elevated energy state and to a combined increase in polyunsaturated FAs and total oil content when overexpressed in developing seeds of transgenic Arabidopsis plants. Plant Physiol 155:1435–1444PubMedPubMedCentralCrossRefGoogle Scholar
  134. Wakao S, Andre C, Benning C (2008) Functional analyses of cytosolic glucose-6-phosphate dehydrogenases and their contribution to seed oil accumulation in Arabidopsis. Plant Physiol 146:277–288PubMedPubMedCentralCrossRefGoogle Scholar
  135. Wan B (2015) Transgenic pyramiding for crop improvement. In: Advances in plant breeding strategies. Breeding Biotechnology and Molecular Tools Springer International Publishing, Chicago, pp 369–396Google Scholar
  136. Wang Z, Huang W, Chang J, Sebastian A, Li Y, Li H et al (2014a) Overexpression of SiDGAT1 a gene encoding acyl-CoA: diacylglycerol acyltransferase from Sesamum indicum L. increases oil content in transgenic Arabidopsis and soybean. Plant Cell Tissue Organ Cult 119:399–410CrossRefGoogle Scholar
  137. Wang Y, Han Y, Teng W, Zhao X, Li Y, Wu L, Li D, Li W (2014b) Expression quantitative trait loci infer the regulation of isoflavone accumulation in soybean (Glycine max L Merr.) seed. BMC Genom 15:680CrossRefGoogle Scholar
  138. Wang ML, Khera P, Pandey MK, Wang H, Qiao L, Feng S et al (2015) Genetic mapping of QTLs controlling FAs provided insights into the genetic control of FA synthesis pathway in peanut (Arachis hypogaea L.). PLoS One 10:e0119454PubMedPubMedCentralCrossRefGoogle Scholar
  139. Weselake RJ, Taylor DC, Rahman MH, Shah S, Laroche A, McVetty PB, Harwood JL (2009) Increasing the flow of carbon into seed oil. Biotechnol Adv 27:866–878PubMedCrossRefGoogle Scholar
  140. Wingenter K, Schulz A, Wormit A, Wic S, Trentmann O, Hoermiller II et al (2010) Increased activity of the vacuolar monosaccharide transporter TMT1 alters cellular sugar partitioning sugar signaling and seed yield in Arabidopsis. Plant Physiol 154:665–677PubMedPubMedCentralCrossRefGoogle Scholar
  141. Winichayakul S, Scott RW, Roldan M, Hatier JHB, Livingston S, Cookson R et al (2013) In vivo packaging of triacylglycerols enhances Arabidopsis leaf biomass and energy density. Plant Physiol 162:626–639PubMedPubMedCentralCrossRefGoogle Scholar
  142. Wormit A, Trentmann O, Feifer I, Lohr C, Tjaden J, Meyer S et al (2006) Molecular identification and physiological characterization of a novel monosaccharide transporter from Arabidopsis involved in vacuolar sugar transport. Plant Cell 18:3476–3490PubMedPubMedCentralCrossRefGoogle Scholar
  143. Xu C, Fan J, Yan C, Shanklin J (2015) U.S Patent No20150337017. U.S Patent and Trademark Office, WashingtonGoogle Scholar
  144. Xu C, Fan J, Yan C, Shanklin J (2016) U.S Patent No20160002651. U.S Patent and Trademark Office, WashingtonGoogle Scholar
  145. Yadava DK, Vasudev S, Singh N, Mohapatra T, Prabhu KV (2012) Breeding major oil crops: present status and future research needs. In: Gupta SK (ed) Technological innovations in major world oil crops, vol 1: breeding. Springer Science and Business Media, LLC, New York, pp 17–51CrossRefGoogle Scholar
  146. Zadran S, Levine RD (2013) Perspectives in metabolic engineering: understanding cellular regulation towards the control of metabolic routes. Appl Biochemi Biotechnol 169:55–65CrossRefGoogle Scholar
  147. Zhang FY, Yang MF, Xu YN (2005) Silencing of DGAT1 in tobacco causes a reduction in seed oil content. Plant Sci 169:689–694CrossRefGoogle Scholar
  148. 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 21:3885–3901PubMedPubMedCentralCrossRefGoogle Scholar
  149. Zhang H, Zhang J, Wei P, Zhang B, Gou F, Feng Z, Zhu JK (2014) The CRISPR/Cas9 system produces specific and homozygous targeted gene editing in rice in one generation. Plant Biotechnol J 12:797–807PubMedCrossRefGoogle Scholar
  150. Zheng P, Allen WB, Roesler K, Williams ME, Zhang S, Li J et al (2008) A phenylalanine in DGAT is a key determinant of oil content and composition in maize. Nat Genet 40:367–372PubMedCrossRefGoogle Scholar
  151. Zhou Y, Zhang XJ, Kang XJ, Zhao XY, Zhang XS, Ni M (2009) SHORT HYPOCOTYL UNDER BLUE1 associates with MINISEED3 and HAIKU2 promoters in vivo to regulate Arabidopsis seed development. Plant Cell 21:106–117PubMedPubMedCentralCrossRefGoogle Scholar
  152. Zorrilla-López U, Masip G, Arjó G, Bai C, Banakar R, Bassie L et al (2013) Engineering metabolic pathways in plants by multigene transformation. Int J Dev Biol 57:565–576PubMedCrossRefGoogle Scholar
  153. Zou JT, Katavic V, Giblin EM, Barton DL, MacKenzie SL, Keller WA, Hu X, Taylor DC (1997) Modification of seed oil content and acyl composition in the Brassicaceae by expression of a yeast sn-2 acyltransferase gene. Plant Cell 9:909–923PubMedPubMedCentralCrossRefGoogle Scholar
  154. Zou J, Wei Y, Jako C, Kumar A, Selvaraj G, Taylor DC (1999) The Arabidopsis thaliana TAG1 mutant has a mutation in a diacylglycerolacyltransferase gene. Plant J 19:645–653PubMedCrossRefGoogle Scholar

Copyright information

© Springer Science+Business Media Dordrecht 2016

Authors and Affiliations

  • Siddanna Savadi
    • 1
    Email author
  • Nemappa Lambani
    • 2
  • Prem Lal Kashyap
    • 1
  • Deepak Singh Bisht
    • 2
  1. 1.ICAR-Indian Institute of Wheat and Barley Research (IIWBR)ShimlaIndia
  2. 2.ICAR-Indian Agricultural Research Institute (IARI)New DelhiIndia

Personalised recommendations