Open Access
Review
Issue
OCL
Volume 23, Number 5, September-October 2016
Article Number D503
Number of page(s) 9
Section Dossier: New perspectives of European oleochemistry / Les nouvelles perspectives de l’oléochimie européenne
DOI https://doi.org/10.1051/ocl/2016023
Published online 15 June 2016

© J.-D. Faure and M. Tepfer, published by EDP Sciences, 2016

Licence Creative Commons
This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

1 Introduction

Over the past decade, worldwide production and distribution of plant oilseeds and their products have undergone remarkable expansion; the area devoted to growing oilseeds has expanded by 19%, while production has increased by 34% since 2005 (source USDA, 2016). The plant oil market is driven by the demand for higher yield and more sustainable production of the main crops (palm, soybean, rapeseed and sunflower), but also by the need for increased crop diversification. This increased demand will require oilseed crops to be adapted to more diversified markets, in particular to provide sources of novel feedstock sources for the petroleum-based chemical industry. However, traditional oilseed crops suffer from several disadvantages that have limited their use in diversifying oil production. Each oilseed crop has relatively low genetic diversity, and the length of plant growth cycles impedes the potential to create new varieties by conventional breeding. Furthermore, for many oilseed crops, genetic engineering is a difficult and lengthy process. Finally, potential problems must be faced concerning how large-scale cultivation of oilseed crops for industrial purposes, such as the production of novel lipids, will coexist with the continued cultivation of the same oilseed crop for human consumption. The last decade has seen the emergence of camelina (Camelina sativa (L.) Crantz) as an alternative species for diversifying oilseed production. Combining camelina’s very attractive agronomic traits with its unprecedented ease for genetic engineering, makes it an ideal plant chassis for biotechnology applications, in particular synthetic biology strategies (Napier et al., 2014; Vollmann and Eynck, 2015; Bansal and Durrett, 2016; Haslam et al., 2016). Also, since it remains a very minor crop in terms of human oil consumption, organizing co-existence should be easier than with the major oilseed food crops, such as rapeseed, soybean, or sunflower.

2 Camelina, the rebirth of an old crop

Camelina, also known as false flax, is an oilseed crop in the Brassicaceae family, and is closely related to well known wild species, including the intensively studied model species Arabidopsis thaliana and the widespread weed Capsella bursa-pastoris, shepherd’s purse (Al-Shehbaz et al., 2006). Under favorable conditions, camelina crops yield >2 t/ha, but lower yields (1.2 to 2.2 t/ha) are observed under conditions of limiting nutrients or water (Crowley and Frohlich, 1998; Gehringer et al., 2006). The history of camelina as a crop is quite unusual, with an ancient history interrupted by a lengthy period of neglect, followed by a renaissance of interest over the past decade. Camelina is thought to have been first domesticated in the steppes of southeastern Europe or southwestern Asia, and the earliest archeological traces of camelina cultivation date to the Neolithic (Toulemonde, 2010). Within the presumed region of domestication, Ukraine and adjacent parts of Russia are still a major center of camelina genetic diversity (Ghamkhar et al., 2010). Over the millennia following its domestication, camelina was widely grown in northern Europe, but starting at the end of the 19th century it was gradually replaced by higher yielding crops such as rapeseed. Nonetheless, during the 20th century, camelina continued to be cultivated on a small scale, essentially for production of oil for human consumption. Because of this century of neglect, camelina has undergone relatively little improvement by plant breeders, and thus the currently grown cultivars can be considered to be quite primitive, and should benefit greatly from the combined efforts of plant breeding and advanced techniques of modern biotechnology as described below.

thumbnail Fig. 1

Publications on camelina and acreage of camelina planted in Montana. Number of publications with camelina in the title referenced in Google Scholar (plain line) and the acreage of camelina grown in Montana (dashed line).

Over the past decade, there has been a remarkable increase in scientific interest in camelina, as shown by the increase in the numbers of publications with "camelina" in the title (Fig. 1), but this increase was driven by several quite different potential end uses. The high levels of omega-3 lipids in camelina oil are perceived as beneficial for human health (Zubr, 1997; Eidhin et al., 2003; Abramovič and Abram, 2005) and the high levels of tocopherols, including vitamin E, make camelina oil more stable to oxidation than other high omega-3 oils such as linseed oil (Abramovič et al., 2007; Hrastar et al., 2009). A further attractive feature is that the residual meal after pressing can be used for animal feed (Peiretti and Meineri, 2007; Pilgeram et al., 2007; Aziza et al., 2010; Bell et al., 2010). In addition to current food uses, there are several micro-niches for camelina oil in cosmetics, soaps, lubricants, etc. (Bonjean and Le Goffic, 1999; Pilgeram et al., 2007). Nevertheless, the intense recent activity in the USA and Canada regarding camelina has been driven primarily by camelina oil’s potential as a low input source of biofuel (Fröhlich and Rice, 2005), with a particularly favorable greenhouse-gas life-cycle assessment (Shonnard et al., 2010; Chen et al., 2015; Keshavarz-Afshar et al., 2015). As shown in Figure 1, this led to a sharp increase in the areas planted in camelina from 2005 to 2007 in Montana, but this was followed by a decline to levels even below the 2005 value, due to unexpectedly poor yields and the inability to compete with petroleum-based fuels (Mclaren and Sun, 2015). The current situation of low petroleum prices suggests that growing camelina for biofuel is unlikely to be economically viable in the near future, but that other uses, and particularly redesigning camelina oils to produce a variety of products including novel industrial feedstocks is a more realistic objective, and this will be the primary focus of this review.

3 Camelina is more than its oil

The increased interest in camelina oil occurred in parallel with increased interest in the crop because of its fundamental agronomic characteristics. Often cited as well adapted to growing on marginal soils, in fact camelina is remarkable adapted to a wide range of temperate climatic conditions, growing well in the semi-arid regions of western North America (Guy et al., 2014) and also in the distinctly humid environment of Ireland (Crowley and Frölich, 1998). It has been described as a low-input crop, requiring little or no fertilization, and since it appears at present to be resistant to many pests and pathogens that affect other Brassicaceae neither insecticides nor fungicides are routinely used on camelina (Seguin-Swartz et al.; Canadian Food Inspection Agency (CFIA), 2014; Vollmann and Eynck, 2015). Furthermore, the camelina life cycle is quite short; if planted in the spring, it can be harvested approximately three months later. Although spring-sown camelina is more widely grown, cultivars adapted to sowing in the fall to be harvested in the spring have also been developed (Bonjean and Le Goffic, 1999). Camelina’s short lifecycle opens particularly interesting possibilities for double cropping. For instance, in the northern US corn belt, winter camelina harvested in the spring can be followed immediately by soybean (Gesch et al., 2014), and winter wheat can be followed by spring camelina in the more arid Northern Great Plains. For northern Europe a fall-sown cereal crop could be followed by camelina sown in the spring (Groeneveld and Klein, 2014). These potential double-cropping systems are of great economic interest, but should also have a positive impact on protecting soils from erosion and increasing crop diversity. These strategies could be greatly facilitated if the camelina life cycle could be further shortened, although it could be of concern if life cycle shortening led to unacceptable loss of yield.

thumbnail Fig. 2

Camelina pipeline for modification of lipid composition. (A) Genetic transformation can be carried out by floral dip, but as shown here is more efficient using vacuum infiltration of flowers. (B) Fluorescent protein markers, such as GFP and DsRed shown here, allow simple screening of transformants among seeds produced by infiltrated plants. (C) A micropress for extracting oil from small seed samples has been developed for preliminary screening of transformed lines. (D) A sample of camelina seeds and the oil extracted using the micropress. (E) Kilogram quantities of seed can be produced in the greenhouse. (F) For larger amounts of seed and for assessment of agronomic traits, field trials are necessary.

Although, as mentioned above, camelina cultivars have not benefitted from intensive plant breeding efforts, there is good evidence for potentially useful genetic variation in the camelina gene pool for important characters, such as plant height, flowering time, seed size, and seed oil composition (Vollmann et al., 2005). Two collections in Germany and Canada are available with several hundred accessions (http://gbis.ipk-gatersleben.de/, http://pgrc3.agr.gc.ca/index_e.html). More recently, genetic maps based on RAPDs, SSRs AFLPs and SNPs have been created (Vollmann et al., 2005; Gehringer et al., 2006; Manca et al., 2013; Singh et al., 2015), and various transcriptome sequencing results have also been reported (Liang et al., 2013; Nguyen et al., 2013; Mudalkar et al., 2014; Singh et al., 2015). These efforts, as well as the draft complete genome sequence will be of great use in breeding improved camelina cultivars (Kagale et al., 2014).

There had been previous indications of the hexaploid nature of the camelina genome from studies of genes encoding key steps in lipid biosynthesis (Hutcheon et al., 2010), but this was fully demonstrated from the draft complete genome sequence (Kagale et al., 2014). In essence, the camelina genome is composed of three equivalents of the genome of its close relative Arabidopsis thaliana. Two of the camelina sub-genomes are extremely similar to each other, and may be derived from an event of autopolyploidy, which then would have been followed by addition of the third, slightly more divergent genome. Sub-genome-specific transcriptomic studies showed a remarkably low degree of gene loss and gene functional differentiation among the three sub-genomes, and the homeologues of all three sub-genomes were most often expressed, with only a slight advantage in expression level for the last-added sub-genome (Kagale et al., 2014). These features could make it difficult to create recessive mutants of interest, since in most cases all three homeologues would need to be mutated. The availability of new genome editing technologies like CRISPR/Cas9 should, however, eliminate these constraints, since CRISPR/Cas9 mutants have already obtained in polyploid species like wheat (Wang et al., 2014).

4 Camelina: a model crop for genetic engineering

Since it was shown that camelina can be genetically transformed with ease by floral dip, using protocols similar to those used for arabidopsis (Lu and Kang, 2008), camelina has become an essential proving ground for seed oil modification (Fig. 2). A remarkably simple pipeline for testing oil modification strategies includes the following steps: (1) gene discovery; (2) construction of transgenes in a vector with a fluorescent protein marker gene (DsRed or GFP); (3) transformation by floral dip or vacuum infiltration; (4) screening T1 seeds for DsRed or GFP fluorescence, screening for changes in lipid profile and/or yield in T1 or T2 seeds, using a newly developed micropress. The overall process from transgene conception to preliminary screening of seed lipids in T2 seeds can be carried out in less than a year. Final validation of the introduced trait must, however, be carried out in the field.

An important issue in the choice of camelina as a model crop for genetic engineering is its ability to cross with wild relatives. Some understanding of the potential for gene flow from genetically modified (GM) crops to wild relatives is necessary for authorization for GM crop field releases, and the ability to use wild relatives as a source of potentially valuable genes is obviously of great interest for classic plant breeding. For both reasons, the resurgence of interest in camelina has included re-examination of its ability to cross with wild relatives. Among the wild Camelina species, only C. microcarpa and C. alyssum are widespread, and with both, fertile hybrids with cultivated camelina can be obtained (Séguin-Swartz et al., 2013). These two species thus represent possible sources of genes of interest for future improvement of cultivated camelina. Since both are only occasionally observed in agricultural contexts, preventing pollen-mediated gene flow from GM camelina to C. microcarpa and C. alyssum in field trials should be easy to assure, as described by the Canadian Food Inspection Agency (CFIA 2012). In contrast to the relative rarity of the wild Camelina species, both Arabidopsis thaliana and Capsella bursa-pastoris are extremely abundant in agricultural environments, and occur in the margins of camelina fields (Tepfer, unpublished). Although both species flower primarily much earlier than camelina, they continue to flower throughout the summer, and are visited by the same potential pollinators (honeybees, bumblebees, syrphid flies (Groeneveld and Klein, 2014). In order to evaluate the potential for gene flow from camelina to these two wild relatives, crosses were made in the greenhouse (Julié-Galau et al., 2014; Martin et al., 2015). No F1 progeny seeds could be obtained with Arabidopsis, and with Capsella, very few F1 seeds were obtained, and the resulting F1 plants proved to be entirely male- and female-sterile (Julié-Galau et al., 2014; Martin et al., 2015). These results suggest that the likelihood of introgression of camelina transgenes in populations of Arabidopsis thaliana and Capsella bursa-pastoris is extremely low indeed.

5 Improving Camelina oil yield

Although, as described above, camelina has many attractive agronomic features, its relatively low oil yield compared to rapeseed is a real limitation to its agroindustrial use. Improving camelina oil yield is thus a priority for development of the crop for any large-scale uses.

One strategy for increasing yield would be to improve the efficiency of photosynthetic carbon fixation. CO2 fixation by Rubisco is limited by its oxygenase activity, initiating a photorespiration cycle leading to glycolate synthesis. Dalal et al. (2015) showed that expression in camelina of three E. coli enzymes that constitute a photorespiratory bypass led to a marked increase in vegetative biomass and and also increased seed yield by 57 to 73%. Although oil yield per seed was not changed, expression of the photorespiratory bypass should be reflected in important gains in yield/ha in the field (Dalal et al., 2015). Similarly, the expression of arabidopsis purple acid phosphatase AtPAP2 modified carbon assimilation and distribution from photosynthetsis and led to higher seed yields (Zhang et al., 2012). Increased photosynthesis efficiency associated with an increased seed mass and oil yield per plant was also achieved through the overexpression of the group III heterotrimeric Gγ-protein AGG3 (Roy Choudhury et al., 2014).

A second strategy to improve oil yield is to globally enhance the levels of metabolic enzymes involved in triacylglycerol (TAG) synthesis by overexpression of specific transcription factors. WRINKLED1 (WRI1) was shown to be an essential transcription factor for TAG synthesis in many species, and its overexpression led to 10–30% increase in seed oil content in arabidopsis, rapeseed and even maize (Cernac and Benning, 2004; Baud et al., 2007; Liu et al., 2010; Pouvreau et al., 2011; Wu et al., 2014). Overexpression of arabidopsis AtWRI1 in camelina seeds led to an increase in seed weight and seed size, and as expected, to 8–31% increase in seed oil content (An and Suh, 2015). Although camelina already has three copies of endogenous WRI1, it is remarkable that significant yield gain could be achieved by overexpression of an orthologous gene.

A third strategy for improving seed oil content is to specifically target metabolic bottlenecks in TAG biosynthesis. Fatty acids produced by plastids are shuffled between phospholipid and TAG pools (Chapman and Ohlrogge, 2012). While phospholipids like phosphatidylcholine (PC) are essential for fatty acid desaturation, they represent an important pool of acyl chains not available for TAG production. Phospholipases A (PLA) are able to hydrolyse PC to lysophosphatidylcholine, releasing a free acyl chain available for TAG synthesis. Constitutive overexpression of several PLA genes in arabidopsis and camelina led to a significant increase in seed oil content, but at the expense of important developmental alterations (Li et al., 2013, 2015). Similar effects on seed oil content, albeit more modest and variable, were obtained by seed-specific overexpression of arabidopsis PLAIIIΔ,Δ but without impacting overall plant growth, stressing the importance of targetting the desired metabolic modifications uniquely to the seed during the maturation phase, while avoiding expression elsewhere and at other phases of the plant growth cycle (Li et al., 2015).

Although this has not yet been described, combining strategies that will increase seed yield with those that increase the proportion of oil in the seeds should make a major contribution to enhancing the economic viability of growing camelina.

6 Improving camelina oil composition for food and feed

The high levels of alpha-linolenic acid (C18:3, ALA) in camelina oil provide an ideal plant chassis for the synthesis of omega-3 long chain polyunsaturated fatty acids (LC-PUFAs) like eicosapentaenoic acid (C20:5, EPA) or docosahexaenoic acid (C22:6, DHA). Omega-3 LC-PUFAs are central dietary recommendations for fetal development and adult cardiovascular and cognitive health. The main dietary source of LC-PUFAs is oil-rich fish, such as atlantic salmon. Since fish do not synthesize LC-PUFAs efficiently, farmed fish are fed with fish meals and fish oil enriched in LC-PUFA extracted from wild-caught fish. A more sustainable solution would be to replace fish oil by vegetable oil enriched in omega-3 LC-PUFA. Land plants do not synthesize polyunsaturated fatty acids longer than 18 carbons and with more than 3 double bonds. Metabolic transformation of ALA (C18:3) into EPA (C20:5) and DHA (C22:6) requires successive fatty acyl desaturation and elongation steps. The efficiency of this metabolic conversion is impeded by the substrate dichotomy paradigm. Indeed, fatty acid elongation in higher plants relies on the acyl-CoA pool, while desaturation takes place principally on phosphatidylcholine, implying that successful synthesis of LC-PUFA requires continuous shuffling of acyl chains between the two acyl pools, acyl-CoA and phospholipids. Substrate dichotomy was thus proposed to be one of the main metabolic bottlenecks in LC-PUFA synthesis (Domergue et al., 2005a; Napier, 2007). A major breakthrough was the discovery that the acyl-CoA dependent Δ6 desaturase of the microalga Otrococcus tauri could convert ALA to stearidonic acid (C18:4, SDA) (Domergue et al., 2005b; Sayanova et al., 2012). The expression of the acyl-CoA dependent Δ6 desaturase in combination with Δ6 elongase and C20 Δ5 desaturase increased the accumulation of C20 intermediates of LC-PUFA biosynthesis in yeast and arabidopsis, demonstrating the potential for reducing substrate dichotomy (Sayanova et al., 2012). Endogenous levels of the LC-PUFA substrate, ALA, has a clear effect on Δ6 desaturase activity, since camelina expressing Δ6 desaturase accumulated three times more SDA (C18:4) than arabidopsis, which synthesizes lower levels of ALA (Sayanova et al., 2012). The complete expression of five enzymes of the LC-PUFA pathway including Δ12 desaturase, Δ15 desaturase, Δ9 elongase, Δ8 desaturase, Δ5 desaturase, for respectively the synthesis of linoleic acid (C18:2), ALA, eicosatrienoic acid (C20:3), eicosatetraenoic acid (ETA) and EPA, allowed efficient EPA synthesis in arabidopsis and camelina (Ruiz-Lopez et al., 2015). Camelina again proved to be a better host, with about 8% EPA accumulated in seeds compared to 3.6% for arabidopsis. An iterative approach of LC-PUFA enzyme combinations allowed respectively a four- and a ten-fold improvement in EPA and DHA accumulation in arabidopsis seeds (Ruiz-Lopez et al., 2013). Finally, the best constructs were validated in camelina, yielding LC-PUFA levels comparable to those found in fish, with 31% EPA and 14% DHA+12% EPA for the best EPA and DHA lines (Ruiz-Lopez et al., 2014). Similar results were obtained by optimizing the expression of the Δ6 desaturase in multiple gene stacking combinations (Petrie et al., 2014). The high potential of these lines to accumulate LC-PUFA in oil was confirmed in field trials (Usher, 2015). Three lessons can be learned from this success story. It is necessary to: (i) identify metabolic bottlenecks in the metabolic pathway (substrate dichotomy) (ii) systematically test all the possible enzyme combinations and (iii) choose an optimized plant host with the highest substrate availability (ALA). While arabidopsis and yeast have been valuable tools for identifying the appropriate enzymes, camelina was essential, not only for its oil profile, but also the ease of its genetic transformation, which facilitated the screening of a large number of transformants. Furthermore, EPA-enriched camelina oil was shown to be a suitable substitute for fish oil in aquaculture (Betancor et al., 2015a, b). These results suggest that LC-PUFA-enriched camelina oil could also represent an interesting alternative source for LC-PUFA in human nutrition. This is reinforced by results obtained using mice fed an EPA-enriched camelina oil diet, which was found to be as efficient as fish oil for providing EPA, thus opening the way to human feeding trials (Tejera et al., 2016).

Nervonic acid (C24:1Δ15) is a natural component of human breast milk fat, and is used in infant formula supplementation, but also in treatment of several neurological diseases, such as multiple sclerosis, adrenoleukodystrophy and Zellweger syndrome (Huai et al., 2015). This fatty acid is found in oil from several Brassicaceae, such as Lunaria annua, but the production of this species is insufficient to meet the demand. Production of nervonic acid in camelina was developed by the overexpression of L. annua keto acyl synthase (KCS), the first enzyme of the cyclic elongation reaction that provides fatty acid specificity of the elongase complex. To improve fatty acid elongation efficiency in camelina seeds, combinations of two elongase enzymes from arabidopsis were associated with LaKCS (Huai et al., 2015). Expression of LaKCS led to significant accumulation of nervonic acid in camelina seeds (up to 12% of lipids), but expression of the additional elongase genes did not improve the yield. Even if LaKCS expression allows nervonic acid synthesis, strategies could be implemented to improve its accumulation, such as: combine all four enzymes of the elongase complex rather than just two, use the elongase enzymes from camelina or L. annua directly, reduce the expression of endogenous KCS to minimize substrate competition by the different elongase complexes.

7 Developing new camelina oil profiles for industry

Due to its specific profile, camelina oil is already used in industrial applications. Highly unsaturated fatty acids are for instance more prone to epoxidation, which is of interest for adhesive properties (Kim et al., 2015a). Epoxidation could be followed by partial acrylation and dihydroxylation, leading to acrylic polyols, which are the source of numerous polymers (Li and Sun, 2015). Recent work demonstrated that alkyd resins used in the coating and paint industry could also be synthesized from camelina oil and polyglycerols (Nosal et al., 2015).

Camelina oil metabolism could also be modified to enhance the accumulation of lipids of industrial value. Jet fuels require medium-chain fatty acid (MCFA) of 8 to 14 carbon length. Camelina plastids elongate acyl-ACP by fatty acid synthase until C16 and C18 carbon length, and the resulting acyl CoAs are then released in the cytosol by acyl thioesterases. C16:0-ACP is hydrolyzed by FatB thioesterases, while FatA is more specific to C18:0- and C18:1-ACP. Cuphea species that accumulate MCFA in their TAG have specific FatB genes (Kim et al., 2015b). Expression of different Cuphea FatB genes associated with different MCFA profiles (C8, C10, C16) led to significant MCFA accumulation in camelina seeds. This effect was enhanced by the coexpression of coconut lysophosphatidic acid acyltransferase (LPAT) and the inhibition of the endogenous camelina plastidial beta-ketoacyl-ACP synthase CsKASII (Kim et al., 2015b).

A similar strategy was used to enrich camelina oil in omega-7 fatty acids, like palmitoleic acid (C16:1Δ9) and cis-vaccenic acid (C18:1Δ11), which have neutraceutical and industrial value for polyethylene production (Nguyen et al., 2015). Combined expression of six different transgenes allowed efficient redirection of fatty acid flux toward omega-7 lipids. First, plastidial C16:0-ACP was channelled toward C16:1Δ9-ACP by the combined inhibition of the 16:0-specific thioesterase (CsFatB) and the β-ketoacyl-ACP synthase (CsKASII) associated with the expression of C16:0-ACP Δ9 desaturase (COM25). To increase omega-7 fatty acid accumulation, the elongase FAE1 was also repressed by RNAi, and the cytosolic C16:0-CoA converted to omega-7 by the expression of C16:0-CoA Δ9 desaturase (Fat5). Altogether, this strategy successfully resulted in the accumulation of more than 60% palmitoleic acid and cis-vaccenic acid in camelina seeds.

Alternatively, the camelina oil profile could be modified simply by inhibiting the last steps of a metabolic pathway, such as for the synthesis of ALA. The down regulation of FAD2 desaturase by anti-sense or RNAi led to camelina oil with almost 50% oleic acid (Kang et al., 2011; Nguyen et al., 2013). Combined reduction of FAD2 with FAE1, the fatty acid elongase involved mainly in erucic acid synthesis in seeds, resulted in a further increase of oleic acid content up to 70% in camelina seeds (Nguyen et al., 2013). Interestingly, spatial analysis of TAG and PC in camelina embryos revealed unexpected heterogeneity (Horn et al., 2013). For instance, cotyledons were enriched in C20:1, while the embryonic axis accumulated more C18:2 in both lipid pools, suggesting tissue specificity in lipid metabolism within the embryo. The FAD2+FAE1 RNAi lines showed incomplete suppression of FAD2 in specific tissues, raising the question of either RNAi inefficiency or the presence of metabolic compartimentalization for oleic acid accumulation. These approaches, based on metabolic mapping by mass spectrometry, provide valuable information about the organization of lipid metabolism in the embryo that will help design further new strategies for modifying the camelina oil profile.

Camelina oil properties could also be profoundly changed by the accumulation of new lipids like acetyl-TAG. Acetyl-TAGs are unusual triacylglycerols in which the sn-3 position has an acetyl group instead of a fatty acyl group. This modification, which reduces viscosity and lowers the oil melting point compared to conventional TAG, is sought for lubricants, food emulsifiers and plasticizers. The main source of acetyl-TAG is the seeds of Euonymus alatus (Burning Bush) thanks to a specific acyltransferase (EaDAcT) that transfers the acetyl group of acetyl-CoA to the sn-3 position of DAG (Durrett et al., 2010). Overexpression of EaDAct in camelina led to an average of 50% acetyl-TAG in seeds, a value that could be increased to 80% when combined with down-regulation of DAGT1 by RNAi (Liu et al., 2015a). The effect of EaDAcT expression, combined or not with DGAT1 RNAi, was significantly higher in camelina transgenic lines compared to arabidopsis or soybean, confirming the particular value of camelina in oil engineering strategies. Interestingly, the high levels of acetyl-TAG accumulation in seeds did not impair seed yield, nor did it modify seed germination in several field studies (Liu et al., 2015a, b). As expected, camelina oil enriched in acetyl-TAG showed lower viscosity, a higher crystallization temperature, and higher caloric content, providing a direct alternative for biodegradable lubricants, hydraulic fluids and transformer oils. The fact that camelina acetyl-TAGs are also rich in oleic acid, and are thus less susceptible to oxidation, could open new markets in the food industry, such as for water retention on food surfaces, emulsifiers, foam stabilizers and packaging plasticizers.

8 Camelina as an alternative producer of non-TAG high value products

Wax esters are neutral lipids that have higher energy density compared to TAGs, and their refinement does not produce glycerol as a side product. They could thus represent a valuable source of biodiesel. They are also used as lubricants, since they have low melting points, an excellent resistance to oxidation, and yet are biodegradable. The desert shrub jojoba is a natural source of wax esters, but since it is not adapted to high yield cultivation, particularly in temperate regions, a more efficient and sustainable approach is required. Only two enzymes are necessary for wax ester synthesis: a fatty acyl-CoA reductase (FAR) and a wax ester synthase (WS). The possibility to accumulate wax esters in seeds was validated in arabidopsis with different variants of FAR and WS from mouse or the bacterium Marinobacter aquaolei (Lardizabal et al., 2000; Heilmann et al., 2012). A total of seven different novel enzyme combinations were tested first in arabidopsis, after which the best three were introduced into camelina (Iven et al., 2016). Similar types of wax esters were produced in arabidopsis and camelina, but the yield in camelina was half that in arabidopsis. Arabidopsis could accumulate 89–108 ng/seed (representing 43–59% neutral lipids) compared to 33–47 ng/seed (15–21%) for camelina. Since wax ester synthesis uses acyl-CoA as substrates, and thus competes directly with TAG synthesis, a possible improvement would be to favor substrate channelling toward wax ester biosynthesis instead of TAG. Recently, a relative 30% content in wax ester was achieved in camelina by the combined overexpression of jojoba FAR and WS with L. annua FAE1 and associated with FAD2 inhibition (Zhu et al., 2016). Nevertheless, camelina yields were lower than those of Crambe abyssinica transformed with the same constructs, perhaps because that the latter naturally accumulates high levels of very long chain fatty acids (Zhu et al., 2016).

The high oil content of camelina seeds could also favor the accumulation of bioactive compounds such as terpenes, which are components of essential oils used in food additives, cosmetics, drugs, rubber and lubricants (Degenhardt et al., 2009). Terpenes also increase oil caloric value, which is an important parameter for biofuel applications, in particular for kerosene. As a proof of concept, the synthesis of the monoterpene (4S)-limonene and the sesquiterpene (+)-delta-cadinene was investigated in camelina seed. Interestingly, these two terpenes are produced via two different pathways: the cytosolic mevalonate pathway and the plastidial 2-C-methyl-D-erythritol 4-phosphate (MEP) pathway, respectively. Combinatorial association of the different enzymes expressed either in the cytosol or in the plastids allowed direct comparison of the most efficient pathways (Augustin et al., 2015). (4S)-limonene could only be accumulated in camelina seed via the expression of plastid-localized enzymes, while (+)-delta-cadinene was accumulated in camelina seeds expressing enzymes either in the cytosol or plastids. Interestingly, the ectopic localization of (+)-delta-cadinene biosynthetic enzymes in plastids was at least five times more efficient than that of the cytosol-localized enzymes. Globally, for both terpenes, the levels obtained were around 5–7 mg/g seed and about 22–29 mg/ml oil. The presence of terpenes in camelina oil increased its caloric value, but the ratio terpene/TAG was still too low for biofuel applications. The use of strong seed-specific promoters could improve terpene production in camelina seeds (Borghi and Xie, 2015). Nonetheless, the minimal loss of these volatile molecules during seed development, as well as their protection from possible degradation, makes camelina seed oil a potentially interesting alternative terpene source for cosmetics and pharmacological use.

There has been much interest in poly-3-hydroxybutyrate (PHB) for use as a biosourced biodegradable replacement for petroleum-derived plastics. Most plant-based PHB has been produced in leaves, but its accumulation in seeds of arabidopsis, rapeseed, tobacco or soybean has had limited success (7% PHB in rapeseed). Camelina was used in an attempt to alleviate this limitation, and the ectopic expression of the three biosynthetic enzymes with different combinations of seed-specific promoters allowed a modest increase to 15% for the best line (Malik et al., 2015). PHB accumulation was however toxic to the embryo, with cotyledon chlorosis and weak seed vigor. The challenge to come will be to develop strategies to minimize PHB toxicity in the embryo cells in order to achieve yields compatible with industrial use.

9 Conclusions

In many studies, camelina has been shown to be an efficient host for bioengineering strategies, with higher oil yields compared to arabidopsis, rapeseed or soybean. The fact that camelina has potentially valuable agronomical characteristics should also increase its interest, not only as as a new crop, but also as a convenient translational tool for arabidopsis research results. In conclusion, current strategies for modifying oil yield or changing the endogenous lipid profile in camelina have had some real success, and can now provide market-compatible products derived from camelina seeds like omega-3 L-PUFA- enriched oil. Exciting challenges remain to improve production yield of new lipids in camelina to levels that are economically viable. New strategies are currently being implemented that couple knowledge of the intricated metabolic fluxes over time and space with synthetic biology tools in order to fine-tune lipid metabolism for specific needs.

Acknowledgments

We thank Helen North for numerous helpful comments on the manuscript.

References

  • Abramovič H, Abram V. 2005. Physico-chemical properties, composition and oxidative stability of camelina sativa oil. Food Technol. Biotechnol. 43: 63–70. [Google Scholar]
  • Abramovič H, Butinar B, Nikoliè V. 2007. Changes occurring in phenolic content, tocopherol composition and oxidative stability of Camelina sativa oil during storage. Food Chem. 104: 903–909. [CrossRef] [Google Scholar]
  • Al-Shehbaz IA, Beilstein MA, Kellogg EA. 2006. Systematics and phylogeny of the Brassicaceae (Cruciferae): An overview. Plant System. Evol. 259: 89–120. [CrossRef] [Google Scholar]
  • An D, Suh MC. 2015. Overexpression of Arabidopsis WRI1 enhanced seed mass and storage oil content in Camelina sativa. Plant Biotechnol. Rep. 9: 137–148. [CrossRef] [Google Scholar]
  • Augustin JM, Higashi Y, Feng X, Kutchan TM. 2015. Production of mono- and sesquiterpenes in Camelina sativa oilseed. Planta 242: 693–708. [CrossRef] [PubMed] [Google Scholar]
  • Aziza AE, Quezada N, Cherian G. 2010. Antioxidative effect of dietary Camelina meal in fresh, stored, or cooked broiler chicken meat. Poultry Sci. 89: 2711–2718. [CrossRef] [Google Scholar]
  • Bansal S, Durrett TP. 2016. Camelina sativa: An ideal platform for the metabolic engineering and field production of industrial lipids. Biochimie 120: 9–16. [CrossRef] [PubMed] [Google Scholar]
  • Baud S, Mendoza MS, To A, Harscoet E, Lepiniec L, Dubreucq B. 2007. WRINKLED1 specifies the regulatory action of LEAFY COTYLEDON2 towards fatty acid metabolism during seed maturation in Arabidopsis. Plant J. 50: 825–838. [CrossRef] [PubMed] [Google Scholar]
  • Bell JG, Pratoomyot J, Strachan F, et al. 2010. Growth, flesh adiposity and fatty acid composition of Atlantic salmon (Salmo salar) families with contrasting flesh adiposity: Effects of replacement of dietary fish oil with vegetable oils. Aquaculture 306: 225–232. [CrossRef] [Google Scholar]
  • Betancor MB, Sprague M, Sayanova O, et al. 2015a. Evaluation of a high-EPA oil from transgenic Camelina sativa in feeds for Atlantic salmon (Salmo salar L.): Effects on tissue fatty acid composition, histology and gene expression. Aquaculture 444: 1–12. [CrossRef] [PubMed] [Google Scholar]
  • Betancor MB, Sprague M, Usher S, Sayanova O, Campbell PJ, Napier JA, Tocher DR. 2015b. A nutritionally-enhanced oil from transgenic Camelina sativa effectively replaces fish oil as a source of eicosapentaenoic acid for fish. Sci. Rep. 5: 8104. [CrossRef] [PubMed] [Google Scholar]
  • Bonjean A, Le Goffic F. 1999. La cameline – Camelina sativa (L.) Crantz ? : une opportunité pour l’agriculture et l’industrie européennes. Oilseeds and fats. Crops and Lipids 6: 28–35. [Google Scholar]
  • Borghi M, Xie DY. 2015. Tissue-specific production of limonene in Camelina sativa with the Arabidopsis promoters of genes BANYULS and FRUITFULL. Planta 243: 549–561. [CrossRef] [PubMed] [Google Scholar]
  • Canadian Food Inspection Agency (CFIA). 2012. Addendum II: Terms and Conditions for Confined Research Field Trials of Camelina (Camelina sativa). Available at: http://www.inspection.gc.ca/plants/plants-with-novel-traits/approved-under-review/field-trials/terms-and-conditions/camelina/eng/1384464422223/1384464422879. [Google Scholar]
  • Canadian Food Inspection Agency (CFIA). 2014. The Biology of Camelina sativa (L.) Crantz (Camelina). Available at: http://www.inspection.gc.ca/plants/plants-with-novel-traits/applicants/directive-94-08/biology-documents/camelina-sativa-l-/eng/1330971423348/1330971509470. [Google Scholar]
  • 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: 575–585. [CrossRef] [PubMed] [Google Scholar]
  • Chapman KD, Ohlrogge JB. 2012. Compartmentation of triacylglycerol accumulation in plants. J. Biol. Chem. 287: 2288–2294. [CrossRef] [PubMed] [Google Scholar]
  • Chen C, Bekkerman A, Afshar RK, Neill K. 2015. Intensification of dryland cropping systems for bio-feedstock production: Evaluation of agronomic and economic benefits of Camelina sativa. Ind. Crops Prod. 71: 114–121. [CrossRef] [Google Scholar]
  • Crowley JG, Frohlich A. 1998. Factors affecting the composition and use of camelina. Oak Park, Carlow, Ireland: Teagasc publication 1 901138 66 6. [Google Scholar]
  • Dalal J, Lopez H, Vasani NB, et al. 2015. A photorespiratory bypass increases plant growth and seed yield in biofuel crop Camelina sativa. Biotechnology for Biofuels 8: 175. [CrossRef] [PubMed] [Google Scholar]
  • Degenhardt J, Köllner TG, Gershenzon J. 2009. Monoterpene and sesquiterpene synthases and the origin of terpene skeletal diversity in plants. Phytochemistry 70: 1621–1637. [CrossRef] [PubMed] [Google Scholar]
  • Domergue F, Abbadi A, Heinz E. 2005a. Relief for fish stocks: Oceanic fatty acids in transgenic oilseeds. Trends Plant Sci. 10: 112–116. [CrossRef] [PubMed] [Google Scholar]
  • Domergue F, Abbadi A, Zähringer U, Moreau H, Heinz E. 2005b. In vivo characterization of the first acyl-CoA Delta6-desaturase from a member of the plant kingdom, the microalga Ostreococcus tauri. Biochem. J. 389: 483–490. [CrossRef] [PubMed] [Google Scholar]
  • Durrett TP, Mcclosky DD, Tumaney a. W, Elzinga D a., Ohlrogge J, Pollard M. 2010. A Distinct DGAT ith sn-3 Acetyltransferase Activity that Synthesizes Unusual, Reduced-Viscosity Oils in Euonymus and Transgenic Seeds. Proc. Natl. Acad. Sci. USA 107: 9464–9469. [CrossRef] [Google Scholar]
  • Eidhin DN, Burke J, O’Beirne D. 2003. Oxidative stability of omega3-rich camelina oil and camelina oil-based spread compared with plant and fish oils and sunflower spread. J. Food Sci. 68: 345–353. [CrossRef] [Google Scholar]
  • Fröhlich A, Rice B. 2005. Evaluation of Camelina sativa oil as a feedstock for biodiesel production. Ind. Crops Prod. 21: 25–31. [CrossRef] [Google Scholar]
  • Gehringer A, Friedt W, Luhs W, Snowdon RJ, Lu W. 2006. Genetic mapping of agronomic traits in false flax (Camelina sativa subsp. sativa). Genome 49: 1555–1563. [CrossRef] [PubMed] [Google Scholar]
  • Gesch RW, Archer DW, Berti MT. 2014. Dual cropping winter camelina with soybean in the northern corn belt. Agronomy J. 106: 1735–1745. [CrossRef] [Google Scholar]
  • Ghamkhar K, Croser J, Aryamanesh N, Campbell M, Kon’kova N, Francis C. 2010. Camelina (Camelina sativa (L.) Crantz) as an alternative oilseed: molecular and ecogeographic analyses. Genome 53: 558–567. [CrossRef] [PubMed] [Google Scholar]
  • Groeneveld JH, Klein AM. 2014. Pollination of two oil-producing plant species: Camelina (Camelina sativa L. Crantz) and pennycress (Thlaspi arvense L.) double-cropping in Germany. GCB Bioenergy 6: 242–251. [CrossRef] [Google Scholar]
  • Guy SO, Wysocki DJ, Schillinger WF, et al. 2014. Camelina: Adaptation and performance of genotypes. Field Crops Res. 155: 224–232. [CrossRef] [Google Scholar]
  • Haslam RP, Sayanova O, Kim HJ, Cahoon EB, Napier JA. 2016. Synthetic Redesign of Plant Lipid Metabolism. Plant J., DOI: 10.1111/tpj.13172. [Google Scholar]
  • Heilmann M, Iven T, Ahmann K, Hornung E, Stymne S, Feussner I. 2012. Production of wax esters in plant seed oils by oleosomal cotargeting of biosynthetic enzymes. J. Lipid Res. 53: 2153–2161. [CrossRef] [PubMed] [Google Scholar]
  • Horn PJ, Silva JE, Anderson D, et al. 2013. Imaging heterogeneity of membrane and storage lipids in transgenic Camelina sativa seeds with altered fatty acid profiles. Plant J. 76: 138–150. [PubMed] [Google Scholar]
  • Hrastar R, Petrisic MG, Ogrinc N, Kosir IJ. 2009. Fatty acid and stable carbon isotope characterization of Camelina sativa oil: implications for authentication. J. Agric. Food Chem. 57: 579–585. [CrossRef] [PubMed] [Google Scholar]
  • Huai D, Zhang Y, Zhang C, Cahoon EB, Zhou Y. 2015. Combinatorial effects of fatty acid elongase enzymes on nervonic acid production in Camelina sativa. PLoS ONE 10: 1–16. [CrossRef] [PubMed] [Google Scholar]
  • Hutcheon C, Ditt RF, Beilstein M, et al. 2010. Polyploid genome of Camelina sativa revealed by isolation of fatty acid synthesis genes. BMC Plant Biol. 10: 233. [CrossRef] [PubMed] [Google Scholar]
  • Iven T, Hornung E, Heilmann M, Feussner I. 2015. Synthesis of oleyl oleate wax esters in Arabidopsis thaliana and Camelina sativa seed oil. Plant Biotechnol. J. 252–259. [Google Scholar]
  • Julié-Galau S, Bellec Y, Faure J-D, Tepfer M. 2014. Evaluation of the potential for interspecific hybridization between Camelina sativa and related wild Brassicaceae in anticipation of field trials of GM camelina. Transgenic Res. 23: 67–74. [CrossRef] [PubMed] [Google Scholar]
  • Kagale S, Koh C, Nixon J, et al. 2014. The emerging biofuel crop Camelina sativa retains a highly undifferentiated hexaploid genome structure. Nature Commun. 5: 3706. [CrossRef] [PubMed] [Google Scholar]
  • Kang J, Snapp AR, Lu C. 2011. Identification of three genes encoding microsomal oleate desaturases (FAD2) from the oilseed crop Camelina sativa. Plant Physiol. Biochem. 49: 223–229. [CrossRef] [PubMed] [Google Scholar]
  • Keshavarz-Afshar R, Mohammed YA, Chen C. 2015. Energy balance and greenhouse gas emissions of dryland camelina as influenced by tillage and nitrogen. Energy 91: 1057–1063. [CrossRef] [Google Scholar]
  • Kim N, Li Y, Sun XS. 2015a. Epoxidation of Camelina sativa oil and peel adhesion properties. Ind. Crops Prod. 64: 1–8. [CrossRef] [Google Scholar]
  • Kim HJ, Silva JE, Vu HS, Mockaitis K, Nam JW, Cahoon EB. 2015b. Toward production of jet fuel functionality in oilseeds: Identification of FatB acyl-acyl carrier protein thioesterases and evaluation of combinatorial expression strategies in Camelina seeds. J. Exp. Botany 66: 4251–4265. [CrossRef] [Google Scholar]
  • Lardizabal KD, Metz JG, Sakamoto T, Hutton WC, Pollard MR, Lassner MW. 2000. Purification of a jojoba embryo wax synthase, cloning of its cDNA, and production of high levels of wax in seeds of transgenic arabidopsis. Plant Physiol. 122: 645–655. [CrossRef] [PubMed] [Google Scholar]
  • Li M, Bahn SC, Fan C, et al. 2013. Patatin-Related Phospholipase pPLAIII d Increases Seed Oil Content with Long-Chain Fatty Acids. Plant Physiol. 162: 39–51. [CrossRef] [PubMed] [Google Scholar]
  • Li M, Wei F, Tawfall A, Tang M, Saettele A, Wang X. 2015. Overexpression of patatin-related phospholipase AIII d altered plant growth and increased seed oil content in camelina. Plant Biotech. J. 13: 766–778. [CrossRef] [Google Scholar]
  • Li Y, Sun XS. 2015. Camelina oil derivatives and adhesion properties. Ind. Crops Prod. 73: 73–80. [CrossRef] [Google Scholar]
  • Liang C, Liu X, Yiu S-M, Lim BL. 2013. De novo assembly and characterization of Camelina sativa transcriptome by paired-end sequencing. BMC Genomics 14: 146. [CrossRef] [PubMed] [Google Scholar]
  • 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 Brassica napus. Plant Physiol. Biochem. 48: 9–15. [CrossRef] [PubMed] [Google Scholar]
  • Liu J, Rice A, Mcglew K, Shaw V, 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. 858–865. [Google Scholar]
  • Liu J, Tjellström 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. [CrossRef] [Google Scholar]
  • Lu C, Kang J. 2008. Generation of transgenic plants of a potential oilseed crop Camelina sativa by Agrobacterium-mediated transformation. Plant Cell. Rep. 27: 273–278. [CrossRef] [PubMed] [Google Scholar]
  • Malik MR, Yang W, Patterson N, et al. 2015. Production of high levels of poly-3-hydroxybutyrate in plastids of Camelina sativa seeds. Plant Biotechnol. J. 13: 675–688. [CrossRef] [PubMed] [Google Scholar]
  • Manca A, Pecchia P, Mapelli S, Masella P, Galasso I. 2013. Evaluation of genetic diversity in a Camelina sativa (L.) Crantz collection using microsatellite markers and biochemical traits. Genet. Resour. Crop Evol. 60: 1223–1236. [CrossRef] [Google Scholar]
  • Martin SL, Sauder CA, James T, Cheung KW, Razeq FM, Kron P, Hall L. 2015. Sexual hybridization between Capsella bursa-pastoris (L.) Medik (♀) and Camelina sativa (L.) Crantz (♂) (Brassicaceae). Plant Breeding 134: 212–220. [CrossRef] [Google Scholar]
  • Mclaren JS, Sun XS. 2015. Can camelina compete as a feedstock for biobased products? INFORM 26: 632–634. [Google Scholar]
  • Mudalkar S, Golla R, Ghatty S, Reddy AR. 2014. De novo transcriptome analysis of an imminent biofuel crop, Camelina sativa L. using Illumina GAIIX sequencing platform and identification of SSR markers. Plant Mol. Biol. 84: 159–171. [CrossRef] [PubMed] [Google Scholar]
  • Napier JA. 2007. The production of unusual fatty acids in transgenic plants. Ann. Rev. Plant Biol. 58: 295–319. [CrossRef] [PubMed] [Google Scholar]
  • Napier JA, Haslam RP, Beaudoin F, Cahoon EB. 2014. Understanding and manipulating plant lipid composition: Metabolic engineering leads the way. Curr. Opin. Plant Biol. 19: 68–75. [CrossRef] [PubMed] [Google Scholar]
  • Nguyen HT, Park H, Koster KL, Cahoon RE, Nguyen HTM, Shanklin J, Clemente TE, Cahoon EB. 2015. Redirection of metabolic flux for high levels of omega-7 monounsaturated fatty acid accumulation in camelina seeds. Plant Biotechnol. J. 13: 38–50. [CrossRef] [PubMed] [Google Scholar]
  • Nguyen HT, Silva JE, Podicheti R, et al. 2013. Camelina seed transcriptome: a tool for meal and oil improvement and translational research. Plant Biotechnol. J. 11: 759–769. [CrossRef] [PubMed] [Google Scholar]
  • Nosal H, Nowicki J, Warzała M, Nowakowska-Bogdan E, Zarêbska M. 2015. Synthesis and characterization of alkyd resins based on Camelina sativa oil and polyglycerol. Progress in Organic Coatings 86: 59–70. [CrossRef] [Google Scholar]
  • Peiretti PG, Meineri G. 2007. Fatty acids, chemical composition and organic matter digestibility of seeds and vegetative parts of false flax (Camelina sativa L.) after different lengths of growth. Anim. Feed Sci. Technol. 133: 341–350. [CrossRef] [Google Scholar]
  • Petrie JR, Shrestha P, Belide S, et al. 2014. Metabolic engineering Camelina sativa with fish oil-like levels of DHA. PLoS ONE 9: 1–8. [CrossRef] [PubMed] [Google Scholar]
  • Pilgeram AL, Sands DC, Boss D, et al. 2007. Camelina sativa, A Montana Omega-3 and Fuel Crop. Proceedings of the sixth National Symposium, Creating Markets for Economic Development of New Crops and New Uses, pp. 129–131. [Google Scholar]
  • Pouvreau B, Baud S, Vernoud V, et al. 2011. Duplicate Maize Wrinkled1 Transcription Factors Activate Target Genes Involved in Seed Oil Biosynthesis. Plant Physiol. 156: 674–686. [CrossRef] [PubMed] [Google Scholar]
  • Roy Choudhury S, Riesselman AJ, Pandey S. 2014. Constitutive or seed-specific overexpression of Arabidopsis G-protein γ subunit 3 (AGG3) results in increased seed and oil production and improved stress tolerance in Camelina sativa. Plant Biotechnol. J. 12: 49–59. [CrossRef] [PubMed] [Google Scholar]
  • Ruiz-Lopez N, Haslam RP, Usher SL, Napier JA, Sayanova O. 2013. Reconstitution of EPA and DHA biosynthesis in Arabidopsis: Iterative metabolic engineering for the synthesis of n-3 LC-PUFAs in transgenic plants. Metab. Eng. 17: 30–41. [CrossRef] [PubMed] [Google Scholar]
  • Ruiz-Lopez N, Haslam RP, Napier JA, Sayanova O. 2014. Successful high-level accumulation of fish oil omega-3 long-chain polyunsaturated fatty acids in a transgenic oilseed crop. Plant J. 77: 198–208. [CrossRef] [PubMed] [Google Scholar]
  • Ruiz-Lopez N, Haslam RP, Usher S, Napier JA, Sayanova O. 2015. An alternative pathway for the effective production of the omega-3 long-chain polyunsaturates EPA and ETA in transgenic oilseeds. Plant Biotechnol. J. 1264–1275. [Google Scholar]
  • Sayanova O, Ruiz-Lopez N, Haslam RP, Napier JA. 2012. The role of Delta6-desaturase acyl-carrier specificity in the efficient synthesis of long-chain polyunsaturated fatty acids in transgenic plants. Plant Biotechnol. J. 10: 195–206. [CrossRef] [PubMed] [Google Scholar]
  • Seguin-Swartz G, Eynck C, Gugel RK, Strelkov SE, Olivier CY, Li JL, Klein-Gebbinck H, Borhan H, Caldwell CD, Falk KC. 2009. Diseases of Camelina sativa (false flax). Canadian J. Plant Pathol.-Revue Canadienne de Phytopathologie 31: 375–386. [CrossRef] [Google Scholar]
  • Seguin-Swartz G, Nettleton JA, Sauder C, Warwick SI, Gugel RK. 2013. Hybridization between Camelina sativa (L.) Crantz (false flax) and North American Camelina species. Plant Breed. 132: 390–396. [CrossRef] [Google Scholar]
  • Shonnard DR, Williams L, Kalnesc TN. 2010. Camelina-Derived Jet Fuel and Diesel: Sustainable Advanced Biofuels. Environ. Progress Sustain. Energy 29: 382–392. [CrossRef] [Google Scholar]
  • Singh R, Bollina V, Higgins EE, et al. 2015. Single-nucleotide polymorphism identification and genotyping in Camelina sativa. Mol. Breed. 35. [Google Scholar]
  • Tejera N, Vauzour D, Betancor MB, et al. 2016. A Transgenic Camelina sativa Seed Oil Effectively Replaces Fish Oil as a Dietary Source of Eicosapentaenoic Acid in Mice 1–3. J. Nutr. 227–235. [Google Scholar]
  • Toulemonde F. 2010. Camelina sativa, l ’or végétal du Bronze et du Fer. Anthropobotanica 1.1: 3–14. [Google Scholar]
  • Vollmann J, Eynck C. 2015. Camelina as a sustainable oilseed crop: Contributions of plant breeding and genetic engineering. Biotechnol. J. 10: 525–535. [CrossRef] [PubMed] [Google Scholar]
  • Vollmann J, Grausgruber H, Stift G, Dryzhyruk V, Lelley T. 2005. Genetic diversity in camelina germplasm as revealed by seed quality characteristics and RAPD polymorphism. Plant Breed. 124: 446–453. [CrossRef] [Google Scholar]
  • Wang Y, Cheng X, Shan Q, Zhang Y, Liu J, Gao C, Qiu J-L. 2014. Simultaneous editing of three homoeoalleles in hexaploid bread wheat confers heritable resistance to powdery mildew. Nat. Biotechnol. 32: 1–6. [CrossRef] [PubMed] [Google Scholar]
  • Wu XL, Liu ZH, Hu ZH, Huang RZ. 2014. BnWRI1 coordinates fatty acid biosynthesis and photosynthesis pathways during oil accumulation in rapeseed. J. Integr. Plant Biol. 56: 582–593. [CrossRef] [PubMed] [Google Scholar]
  • Zhang Y, Yu L, Yung K-FF, Leung DYC, Sun F, Lim BL. 2012. Over-expression of AtPAP2 in Camelina sativa leads to faster plant growth and higher seed yield. Biotechnol. Biofuels 5: 19. [CrossRef] [PubMed] [Google Scholar]
  • Zhu L-H, Krens F, Smith MA, et al. 2016. Dedicated Industrial Oilseed Crops as Metabolic Engineering Platforms for Sustainable Industrial Feedstock Production. Sci. Rep. 6: 22181. [CrossRef] [PubMed] [Google Scholar]
  • Zubr J. 1997. Oil-seed crop: Camelina sativa. Ind. Crops Prod. 6: 113–119. [CrossRef] [Google Scholar]

Cite this article as: Jean-Denis Faure, Mark Tepfer. Camelina, a Swiss knife for plant lipid biotechnology. OCL 2016, 23(5) D503.

All Figures

thumbnail Fig. 1

Publications on camelina and acreage of camelina planted in Montana. Number of publications with camelina in the title referenced in Google Scholar (plain line) and the acreage of camelina grown in Montana (dashed line).

In the text
thumbnail Fig. 2

Camelina pipeline for modification of lipid composition. (A) Genetic transformation can be carried out by floral dip, but as shown here is more efficient using vacuum infiltration of flowers. (B) Fluorescent protein markers, such as GFP and DsRed shown here, allow simple screening of transformants among seeds produced by infiltrated plants. (C) A micropress for extracting oil from small seed samples has been developed for preliminary screening of transformed lines. (D) A sample of camelina seeds and the oil extracted using the micropress. (E) Kilogram quantities of seed can be produced in the greenhouse. (F) For larger amounts of seed and for assessment of agronomic traits, field trials are necessary.

In the text

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