- Email: [email protected]
Field performance of terpene-producing Camelina sativa
Industrial Crops & Products 136 (2019) 50–58
Contents lists available at ScienceDirect
Industrial Crops & Products journal homepage: www.elsevier.com/locate/indcrop
Field performance of terpene-producing Camelina sativa a,b
a,c
a
T a,d
Jörg M. Augustin , Jordan R. Brock , Megan M. Augustin , Rachel L. Wellinghoff , ⁎ Matthew Shippa,e, Yasuhiro Higashia,f, Tadele T. Kumssag,h, Edgar B. Cahoong, Toni M. Kutchana, a
Donald Danforth Plant Science Center, 975 North Warson Rd, St. Louis, MO, 63132, United States Elemental Enzymes, 1685 Galt Industrial Blvd, St. Louis, MO, 63132, United States Washington University in St. Louis, Department of Biology, 1 Brookings Dr, St. Louis, MO, 63130, United States d Cofactor Genomics, 4044 Clayton Ave, St. Louis, MO, 63110, United States e KWS, 1005 N Warson Rd, St. Louis, MO, 63132, United States f RIKEN Center for Sustainable Resource Science, 1-7-22 Suehiro, Tsurumi, Yokohama, Kanagawa, 230-0045, Japan g University of Nebraska-Lincoln, 1400 R St, Lincoln, NE, 68588, United States h Noble Research Institute, 2510 Sam Noble Pkwy, Ardmore, OK 73401, United States b c
A R T I C LE I N FO
A B S T R A C T
Keywords: Terpene (+)-δ-cadinene (4S)-limonene (+)-5-epi-aristolochene Field trial Camelina sativa
Bioengineered lines of the low-input oil seed crop Camelina sativa (Camelina), augmented to accumulate the monoterpene (4S)-limonene, the sesquiterpene (+)-δ-cadinene, or the sesquiterpene (+)-5-epi-aristolochene in seed, were evaluated for two growing seasons under field conditions to determine performance of the introduced traits in an agricultural setup including the effects on overall plant fitness, and total seed yield. Field-grown Camelina plants were further compared to greenhouse-grown plants to evaluate commonalities and differences resulting from cultivation under either controlled or agriculturally relevant growth conditions. Morphological appearance and plant height differed marginally between transgenic and wild-type plants under both greenhouse and field conditions, indicating low impact of the terpene production traits toward overall plant fitness. Total seed yield, however, was independent of the growth conditions and was reduced in Camelina lines producing (4S)-limonene by 48% on average and by 30% on average for (+)-5-epi-aristolochene producing lines. Conversely, (+)-δ-cadinene producing Camelina seed yields remained wild-type equivalent. Additional investigations included seed terpene accumulation, seed oil amount, and seed fatty acid composition. Terpene accumulation was reduced up to 22% for field-grown plants as compared to greenhouse-grown plants. Seed oil amounts were similar under greenhouse and field conditions but were consistently lowered by 2–5% for terpene producing lines. Similarly, seed oil composition remained stable under both field and greenhouse conditions, but generally favored a more energy dense phenotype in terpene producing Camelina lines. Lastly, outcrossing of transgenic traits to adjacent wild-type Camelina plants was observed under field but not greenhouse conditions.
1. Introduction The re-emerging oilseed crop Camelina sativa (Camelina) is currently under consideration as a renewable, farmable source for the production of a variety of industrial and agricultural products due to its amendable properties in the field. Camelina is a low-input oil seed crop
that is thought to have been originally cultivated by man during the Iron Age (Larsson, 2013; Sostaric et al., 2009) as a food oil source, but was later abandoned due to the availability of higher yielding oil crops. In recent years Camelina has regained interest as a crop with minimal fertilization requirements and drought resistance traits making it suitable for growth in marginal soils and drier climates (Bansal and
Abbreviations: MEP, methylerythritol phosphate; GDS, geranyl diphosphate synthase; FDS, farnesyl diphosphate synthase; LS, (4S)-limonene synthase; CDNS, (+)-δcadinene synthase; EAS, (+)-5-epi-aristolochene synthase; DXS, 1-deoxy-D-xylulose-5-phosphate synthase; DXR, 1-deoxy-D-xylulose 5-phosphate reductoisomerase; HDS, (E)-4-hydroxy-3-methylbut-2-enyl diphosphate synthase; HDR, (E)-4-hydroxy-3-methylbut-2-enyl diphosphate reductase; DGL, transgenic Camelina sativa expressing DXS, GDS and LS; DFC, transgenic Camelina sativa expressing DXS, FDS and CDNS; FE, transgenic Camelina sativa expressing FDS and EAS; DDHHGL, transgenic Camelina sativa expressing DXS, DXR, HDS, HDR, GDS and LS; DDHHFC, transgenic Camelina sativa expressing DXS, DXR, HDS, HDR, FDS and CDNS ⁎ Corresponding author. E-mail addresses: [email protected] (J.M. Augustin), [email protected] (J.R. Brock), [email protected] (M.M. Augustin), rwellinghoff@gmail.com (R.L. Wellinghoff), [email protected] (M. Shipp), [email protected] (Y. Higashi), [email protected] (T.T. Kumssa), [email protected] (E.B. Cahoon), [email protected] (T.M. Kutchan). https://doi.org/10.1016/j.indcrop.2019.04.061 Received 15 March 2019; Received in revised form 19 April 2019; Accepted 28 April 2019 Available online 06 May 2019 0926-6690/ © 2019 Elsevier B.V. All rights reserved.
Industrial Crops & Products 136 (2019) 50–58
J.M. Augustin, et al.
Durrett, 2016; Putnam et al., 1993). Its compatibility with present day agricultural practices is expected to ease potential transiting into modern cultivation (Moser, 2012). Camelina has further shown resistance to a number of diseases and pests including blackleg, blackspot, Alternaria spp., and flea beetles that plague cruciferous oil crops such as rapeseed and its derivative Canola (Putnam et al., 1993; Vollmann and Eynck, 2015). Lastly, another reason for the renewed interest in Camelina is its high accumulation of unsaturated fatty acids including oleic (18:1), linoleic (18:2), linolenic (18:3), and eicosenoic (20:1) acid, which, compared to saturated fatty acids, have both health benefits for human consumption and lower melting points resulting, upon conversion into biodiesels, in fuel blends that remain liquid longer at cooler temperatures (Gunstone et al., 2007; Putnam et al., 1993). Genetic engineering of Camelina is feasible due to its ease of stable transformation, close relatedness to the well-studied model plant Arabidopsis thaliana, and the availability of genomic and transcriptomic datasets, which aid in relatively straightforward manipulability for specific industries (Kagale et al., 2014, 2016; Liang et al., 2013; Lu and Kang, 2008; Nguyen et al., 2013). Extensive work has been dedicated to alter Camelina’s lipid profile for production of wax esters, poly-3-hydroxybutyrate, cyclic fatty acids, and acetyl-triacylglycerols for, e.g., use as industrial lubricants, cosmetics, and plastics (Iven et al., 2015; Liu et al., 2015; Malik et al., 2015; Yu et al., 2017; Zhu et al., 2016). Additionally, modifications of the lipid profile of Camelina seed oil toward the production of omega-3 long-chain polyunsaturated fatty acids has shown great promise for replacement of fish oil in aquaculture (Betancor et al., 2016a, b; Betancor et al., 2015; Mansour et al., 2014; Petrie et al., 2014; Ruiz-Lopez et al., 2014). The oils of both natural accessions and genetically modified Camelina lines have further been highlighted as a potential biofuel source for biodiesel and Jet A fuel drop-ins with significantly lower particle emissions (Kim et al., 2015; Moore et al., 2017; Moser, 2012). Production of high value compounds in Camelina can increase the overall crop value. For example, using modified oil as a renewable source material for industrial or pharmaceutical applications increases the farmer’s income significantly as opposed to using the material as an addition to animal feed alone (Dai et al., 2017; Kim et al., 2017). Previously, we bioengineered Camelina lines to produce and accumulate the industrially relevant terpenes (4S)-limonene and (+)-δ-cadinene in seed by plastidial localization of 1) limonene synthase (LS), geranyl diphosphate synthase (GDS) and 1-deoxy-D-xylulose-5-phosphate synthase (DXS), or 2) (+)-δ-cadinene synthase (CDNS), farnesyl diphosphate synthase (FDS) and DXS; abbreviated as DGL and DFC lines, respectively (Augustin et al., 2015). Targeting enzymes to the plastids resulted in higher terpene accumulation due to the utilization of dimethylallyl pyrophosphate (DMAPP) and isopentenyl pyrophosphate (IPP) originating from the plastid-located methylerythritol phosphate (MEP) pathway (Fig. 1). Our previous results demonstrated that when these transgenic Camelina plants were grown under greenhouse conditions, the production of (4S)-limonene is stable for at least five consecutive generations. Viability of the terpene traits under field conditions in an agricultural setup, however, remained thus far uninvestigated. Here we present the evaluation of the DGL and DFC lines as well as a third line that produces the sesquiterpene (+)-5-epi-aristolochene, termed FE, under actual field conditions for two consecutive years and compare their morphology, yield, oil content, and terpene production to greenhouse grown DGL, DFC, and FE plants. During the second field season we further expanded the study to include two improved terpene producing lines, designated DDHHGL and DDHHFC (manuscript in preparation), that overexpress additional MEP pathway genes. Lastly, the observation of outcrossing events between the transgenic lines and adjacent wild-type plants during the first field season is discussed.
Fig. 1. Schematic of pathways leading to the biosynthesis of (4S)-limonene, (+)-δ-cadinene, and (+)-5-epi-aristolochene in transgenic Camelina plants of the DGL/DDHHGL, DFC/DDHHFC and FE lines, respectively. Isopentenyl pyrophosphate (IPP) and dimethylallyl pyrophosphate (DMAPP) derived from the plastid-localized MEP pathway are coupled by either geranyl diphosphate synthase (GDS) or farnesyl diphosphate synthase (FDS) to produce geranyl pyrophosphate (GPP). In DGL and DDHHGL plants, GPP is subsequently cyclized into (4S)-limonene by limonene synthase (LS). In DFC, DDHHFC, and FE plants, FDS catalyzes the linkage of another IPP molecule to GPP yielding farnesyl pyrophosphate (FPP), which is then cyclized by either cadinene synthase (CDNS) into (+)-δ-cadinene in DFC and DDHHFC plants or by epi-aristolochene synthase (EAS) into (+)-epi-aristolochene in FE plants.
2. Methods 2.1. Preparation of transgenic Camelina lines Generation of the Camelina sativa Suneson DFC and DGL lines was described previously (Augustin et al., 2015). For generation of the DDHHFC and DDHHGL lines, the transformation constructs used to create the DFC and DGL lines were stepwise extended with the MEP pathway genes encoding 1-deoxy-D-xylulose 5-phosphate reductoisomerase (DXR), (E)-4-hydroxy-3-methylbut-2-enyl diphosphate synthase (HDS), and (E)-4-hydroxy-3-methylbut-2-enyl diphosphate reductase (HDR) from Mentha piperita (manuscript in preparation). Similar to DXS, expression of DXR, HDS, and HDR is controlled by identically structured Glycine max glycinin expression cassettes. The transformation construct for the FE line derived from a precursor of the DFC transformation construct lacking the DXS expression cassette and carrying the (+)-5-epi-aristolochene synthase (EAS) ORF from Nicotiana sp. in place of the CDNS ORF. All transformation constructs were introduced into Agrobacterium tumefaciens GV3101:pMP90, which was subsequently used to transform wild-type Camelina sativa Suneson plants using the floral-dip method as described previously (Augustin et al., 2015; Lu and Kang, 2008). Transgenic seeds were screened using a green LED light in combination with a red filter to identify the presence of the selection marker DsRed2. The DsRed2 expression cassette was included in the TDNA section of
51
Industrial Crops & Products 136 (2019) 50–58
J.M. Augustin, et al.
2.4. FAME analysis
each plant transformation construct. Transgenic plants were propagated until homozygous plant lines, determined by ubiquitous presence of the DsRed2 trait in all derived seeds, were obtained.
For determination of seed fatty acid profiles, triplicate fatty acid methyl ester (FAME) extractions were performed on a total of ten seeds per replicate. Seeds were combined, weighed, and transferred to screw top glass tubes followed by addition of 1.5 ml of 2.5% sulfuric acid in methanol and ground using a glass rod. Subsequently, 50 μg triheptadecanoin (per mg seed) was solubilized in a total volume of 500 μl toluene and added to each sample as internal standard. Samples were incubated at 95 °C for 50 min then cooled to room temperature prior to addition of 1 ml hexane and 1 ml 1 M NaCl. FAMEs were extracted into the hexane phase by rapid mixing. Subsequently, samples were centrifuged at 445 x g for 5 min to facilitate phase separation. Aliquots of the hexane phase were transferred to screw top autosampler vials and analyzed by GC-FID on a Thermoquest Trace GC 2000 system equipped with an Agilent HP-INNOwax column (30 m x 250 μm x 0.25 μm) using helium as the carrier gas. GC conditions began with a hold of 60 °C for 1 min, increasing to 185 °C at a rate of 40 °C min–1, then rising to 235 °C at a rate of 5 °C min–1 with a 5 min hold. FAME species identification was based on retention time comparison to known standards. Fatty acid abundances are reported as a percentage of each FAME peak area of the total peak area of all FAME peaks combined. Absolute abundances were further quantified relative to the triheptadecanoin internal standard and used to estimate total seed oil content.
2.2. Plant trial metrics Field trials were conducted in Mead, Nebraska, USA during April and June of 2014 and April and July of 2015. For the 2014 field season, seeds of homozygous T4 generation DFC, T4 DGL, T3 FE plants, and wild-type Camelina sativa Suneson were planted in single 2–3.5 m long rows. In the 2015 field season, in addition to cultivating seeds of homozygous T4 DFC, T4 DGL, T3 FE, and wild-type Camelina plants, T4 DDHHFC and T4 DDHHGL seeds were planted as well. In 2015, each line was planted in plots of four adjacent rows of approximately 2 m in length. The distance between rows within plots was approximately 0.15 m and the distance between individual plots 0.9 m. Greenhousepropagated DFC, DGL, and FE seed sowed in the 2015 field trial were prepared from the same T3 and T2 generation parental seed batches used to propagate seed for the 2014 field season. Lines grown in the greenhouse for comparison to the field were cultivated at the Donald Danforth Plant Science Center, St. Louis, Missouri, USA from July to October of 2014. Greenhouse conditions were 21 °C during the day and 20 °C in the night and within a humidity range of 40–90%. Seed used for the greenhouse comparison derived from the same T4 DFC, T4 DGL, and T3 FE line and wild-type Camelina sativa Suneson seed batches that were planted in the 2014 field trial. For both the field trials and the greenhouse comparison, ten individual plants were selected per line and used to determine average plant height, total seed yield, seed weight, seed terpene accumulation, and seed oil accumulation. Seed of each harvested plant were screened as described above for expression of DsRed2, indicative for the presence of transgenic traits.
2.5. NMR oil analysis Total seed oil content was quantified using a benchtop NMR analyzer MQC (Oxford Instruments). The instrument was tuned with a 40 °C standard of extracted Camelina oil before each set of runs. Weighed seeds (0.3 g — 0.5 g) were transferred to clean 10 mm glass sample tubes and incubated at 40 °C for at least 10 min prior to analysis. 3. Results and discussion
2.3. Terpene extraction and analysis
3.1. Phenotypic analysis
Typically, three terpene extracts were prepared and measured per plant. Ten individual seeds per extraction were combined and weighed. Each seed batch was subsequently placed in a glass screw top tube and submerged in 2 ml diethyl ether. After addition of 50 μl of an internal standard stock solution, seeds were ground to a fine powder using a glass rod. Internal standards were 400 ng/μl iso-butylbenzene for (4S)limonene analyses and 400 ng/μl hexadecane for (+)-δ-cadinene and (+)-5-epi-aristolochene analyses. Sealed tubes were incubated at room temperature for a minimum of 2 h while shaking at 150 rpm and inverted approximately every 15 min. Tubes were then centrifuged at 1000×g for 10 min and 150 μl supernatant was transferred to screw top autosampler vials, which were subsequently centrifuged again at 1000×g for 5 min. Finally, extracts were analyzed by GC–MS after injection of 1 μl sample onto an Agilent 7890 A GC equipped with a Restek Rtx-5MS column (30 m x 250 μm x 0.25 μm) connected to a 5975C inert XL MSD mass spectrometer using helium as carrier gas. For (4S)-limonene quantification, the oven temperature started with a hold at 30 °C for 3 min, then raised to 85 °C at a rate of 55 °C min–1, further raised to 120 °C at a rate of 10 °C min–1, and finally raised to 300 °C at 120 °C min–1 with a 3 min hold. Quantification of (+)-δ-Cadinene and (+)-5-epi-aristolochene containing samples began with an oven temperature of 50 °C for 3 min, then raised to 160 °C at a rate of 80 °C min–1, raised once more to 170 °C at a rate of 1.4 °C min–1, and finally raised to 300 °C at 120 °C min–1 and held for 3 min. (4S)Limonene concentrations were calculated based on a standard curve of commercial (4S)-limonene (Sigma-Aldrich). (+)-δ-Cadinene and (+)-5-epi-aristolochene concentrations were calculated relative to a standard curve of the sesquiterpene (+)-valencene (Sigma-Aldrich).
Terpene producing DFC, DGL, and FE lines of Camelina were compared to wild-type plants grown during field seasons in 2014 and 2015 as well as to plants grown under greenhouse conditions in 2014. In the 2015 field season, two additional terpene yield-improved lines designated DDHHFC and DDHHGL were investigated under field conditions as well. Notably, within each of the trials, adult plants at harvest did not show any obvious morphological differences between the terpene producing lines and wild-type plants (Fig. 2 A). In contrast, overall morphology of Camelina plants grown under field and greenhouse conditions differed substantially. Field grown plants developed thicker stems than greenhouse grown plants and lost all leaves in the field by the time of harvest (Fig. 2B). Similarly, average plant heights at harvest of all field grown Camelina lines were widely consistent throughout both seasons and no significant differences (p ≤ 0.05) were observed between terpene producing lines and wild-type plants while greenhouse grown plants were an average of 5–10 cm taller at harvest (Table 1). Differences in both overall morphology and plant height are likely the result of adaptions to the different physical constraints under the different growth conditions (i.e. exposure to wind and rain under field conditions favored the development of shorter, sturdier plants with stronger stems). Average seed yields per plant fluctuated strongly between the 2014 and 2015 field seasons, whereas average seed yields per plant in the 2015 field season and greenhouse grown plants resembled each other more closely (Table 1). The higher yield of the 2014 season may be due to increased physical space. In 2014, plants were grown in single rows as opposed to more compact patches in the 2015 field season, which more closely resembles growth in the greenhouse where plants are grown in trays. However, yield differences due to variable weather 52
Industrial Crops & Products 136 (2019) 50–58
J.M. Augustin, et al.
Fig. 2. Morphology of representative wild-type and terpene-producing transgenic Camelina plants either grown under greenhouse or field conditions. A) Field-grown Camelina plants from 2014 field season. B) Greenhouse-grown Camelina plants. Images were taken at day of harvest. Table 1 Phenotypic traits of wild-type (WT) and terpene producing transgenic Camelina lines (DFC, DGL, FE, DDHHFC, and DDHHGL) grown under greenhouse or field conditions. Each data point represents the average height (cm), seed yield (g), or individual seed weight (mg) of ten individual plants as indicated. For the individual seed weights, three samples of ten seeds were weighed per plant and the resulting weight of each sample divided by the number of seed. Error range represents standard error. Height (cm)
WT DFC DGL FE DDHHFC DDHHGL
Seed yield (g)
Individual seed weight (mg)
Greenhouse
Field 2014
Field 2015
Greenhouse
Field 2014
Field 2015
Greenhouse
Field 2014
Field 2015
82.2 81.6 81.3 80.2 – –
77.3 74.7 75.0 70.6 – –
72.1 74.4 74.4 69.5 74.6 74.2
1.36 1.50 0.91 1.07 – –
3.08 3.63 1.62 2.25 – –
1.56 1.48 0.75 0.92 1.57 0.81
1.09 1.10 0.90 1.06 – –
0.93 1.02 0.92 1.22 – –
0.73 0.98 0.99 1.06 1.09 1.03
± ± ± ±
0.7 1.6 2.1 2.7
± ± ± ±
2.0 1.1 1.8 1.6
± ± ± ± ± ±
2.3 2.6 2.2 1.9 2.1 1.6
± ± ± ±
0.08 0.10 0.10 0.10
conditions during the two field seasons cannot be excluded. Notably, the (+)-δ-cadinene producing Camelina lines, DFC and DDHHFC, had comparable per plant seed yields in all trials as concurrently grown wild-type plants, averaging 106% of wild-type level. In contrast, both (4S)-limonene producing Camelina lines, DGL and DDHHGL, with an average per plant seed yield throughout all trials of 52% of wild-type levels, as well as the (+)-5-epi-aristolochene producing FE line, averaging at 70% wild-type per plant seed yields, had consistently lower average yields than wild-type plants. This strongly indicates that the in planta production of (4S)-limonene and (+)-5-epi-aristolochene
± ± ± ±
0.65 0.42 0.22 0.30
± ± ± ± ± ±
0.37 0.59 0.14 0.30 0.42 0.17
± ± ± ±
0.02 0.02 0.03 0.01
± ± ± ±
0.03 0.02 0.01 0.02
± ± ± ± ± ±
0.02 0.03 0.03 0.07 0.04 0.06
negatively affects seed yield, which may render these lines less desirable as a field crop. However, the wild-type yield levels of the (+)-δcadinene producing plant lines contradict a general notion that seedspecific terpene production causes a detrimental effect on total seed yield. Thus, additional, independently generated DGL, DDHHGL, and FE lines should be evaluated to exclude line-specific low yield traits as plants were selected based upon terpene yield, not seed yield. The weight per seed was found to be generally consistent at 1 ± 0.1 mg seed−1 for all investigated lines throughout all trials. Exceptions were wild-type plants in the 2015 field season, which 53
Industrial Crops & Products 136 (2019) 50–58
J.M. Augustin, et al.
Fig. 4. Seed oil accumulation in wild-type and terpene producing transgenic Camelina plants either grown under greenhouse or field conditions. A) Oil accumulation in seed of wild-type (WT) Camelina and terpene producing transgenic DFC, DGL, FE, DDHHGL, and DDHHFC lines determined by FAME analysis. Three individual plants were investigated in three technical replicates per line and growth condition. B) Oil accumulation in seed of identical Camelina lines presented in panel A) determined by NMR analysis. Per line and growth condition, ten individual plants were investigated in one to five technical replicates (depending on seed availability). Error bars depict combined standard deviations.
Fig. 3. Seed terpene accumulation in transgenic Camelina plants either grown under greenhouse or field conditions. A) (+)-δ-Cadinene accumulation of DFC line grown under greenhouse conditions, under field conditions during 2014 field season (2014 FT) and 2015 field season (2015 FT), as well as DDHHFC line grown under field conditions during 2015 field season (DDHHFC). B) (4S)Limonene accumulation of DGL line grown under greenhouse conditions and field conditions (2014 FT, 2015 FT), as well as of the DDHHGL line grown under field conditions during 2015 field season (DDHHGL). C) (+)-5-epiAristolochene accumulation of FE line grown under greenhouse conditions and field conditions (2014 FT, 2015 FT). Per line and growth condition, ten individual plants were investigated in three technical replicates. Error bars depict combined standard deviation.
and were, thus, excluded from terpene quantitation. All investigated plants of the DFC line contained (+)-δ-cadinene in their seed, which averaged at 5.9 ( ± 0.7) mg g−1 in plants grown under greenhouse conditions, 4.9 ( ± 0.4) mg g−1 in seed of field trial plants from 2014, and 4.9 ( ± 1.3) mg g−1 in seed of field trial plants from 2015. The DDHHFC line, which was exclusively investigated during the 2015 field season, accumulated 7.4 ( ± 1.3) mg g−1 (+)-δ-cadinene in its seeds and, thus, showed substantially higher amounts as compared to the DFC line grown under identical conditions (Fig. 3A). Plants of the DGL line accumulated 10.5 ( ± 1.1) mg (4S)-limonene g−1 seed in the greenhouse, 9.1 ( ± 0.7) mg g−1 under field conditions in 2014, and 7.9 ( ± 1.3) mg g−1 under field conditions in 2015. In parallel grown DDHHGL plants, terpene accumulation averaged 9.3 ( ± 1.3) mg (4S)limonene g−1 seed during the 2015 field season (Fig. 3B). The FE line accumulated an average of 2.8 ( ± 1.2) mg (+)-5-epi-aristolochene g−1 seed under greenhouse conditions, 1.4 ( ± 0.4) mg g−1 under field condition in 2014, and only 0.3 ( ± 0.3) mg g−1 under field conditions in 2015 (Fig. 3C). As previously reported (Augustin et al., 2015), we detected glycosylated terpene products in both DGL and DFC lines, which was shown to increase the total terpene content by up to 30%. However, due to difficulties in quantitation of glycosylated (+)-δ-cadinene, the values reported here do not include glycosylated terpenes.
produced smaller seed with an average of 0.7 mg seed−1, and the FE line during the 2014 field season with slightly larger seeds that averaged 1.2 mg seed−1 (Table 1). Considering the very comparable (DFC) or even slightly higher weights per seed (DGL and FE) under field conditions, the reduced seed weight of wild-type Camelina in the field is unlikely a general trend for growing Camelina under field conditions as compared to cultivation in the greenhouse.
3.2. Seed terpene accumulation To assess effectiveness of the introduced terpene traits under the different growth conditions, seeds of ten individual plants of each transgenic line grown in the greenhouse, in the field in 2014, and in the field in 2015 were analyzed for terpene content. Wild-type Camelina seeds are devoid of mono- and sesquiterpenes (Augustin et al., 2015) 54
Industrial Crops & Products 136 (2019) 50–58
J.M. Augustin, et al.
Fig. 5. Seed fatty acid composition of wild-type and terpene producing transgenic Camelina plants either grown under greenhouse or field conditions. A) Seed fatty acid composition of wildtype (WT) and transgenic DGL, DFC, and FE Camelina lines grown in the greenhouse. B) Seed fatty acid composition of WT and transgenic DGL, DFC, and FE Camelina plants grown under field conditions during 2014 field season. C) Seed fatty acid composition of WT and transgenic DGL, DFC, FE, DDHHGL, and DDHHFC Camelina plants grown under field conditions during 2015 field season. Per line and growth condition, three individual plants were investigated using three technical replicates. Error bars depict combined standard deviation.
construct. However, previous greenhouse comparisons of FE and FC lines, a DFC precursor line not overexpressing DXS (Augustin et al., 2015), also resulted in higher sesquiterpene accumulations for the (+)-δ-cadinene producing line than the (+)-5-epi-aristolochene producing line (data not shown). This indicates (+)-δ-cadinene is more efficiently produced, e.g. due to higher expression or activity rates of the involved sesquiterpene synthase, or more efficiently accumulated in Camelina seed than (+)-5-epi-aristolochene.
Therefore, the total terpene content in all lines analyzed may be significantly higher if the amount of glycosylated terpenes is taken into account. In summary, all three lines, DFC, DGL, and FE, accumulate more terpene under greenhouse conditions than under field conditions. Of the two field seasons, 2014 appeared more beneficial toward terpene production than 2015. Considering the generally higher seed yields in 2014 compared to 2015 (Table 1), it is possible that the higher terpene yield under 2014 field conditions resulted from generally elevated plant vigor. The two Camelina lines that carry, in addition, gene cassettes to overexpress DXR, HDS, and HDR (manuscript in preparation), i.e. DDHHFC and DDHHGL, both accumulate more cadinene and limonene, respectively, than the corresponding predecessor lines, DFC and DGL, under field conditions in 2015. This indicates that overexpression of additional MEP pathway enzymes in Camelina seed, which generates the terpene precursor molecules DMAPP and IPP in plastids, leads also to further increased terpene production under field conditions as previously seen in greenhouse studies (manuscript in preparation). Previously, limonene producing plants were found to accumulate more terpene (in mg g−1 seed weight) than (+)-δ-cadinene producing lines (Augustin et al., 2015). This observation holds up throughout all growth conditions included in this study. The reason is unknown but may be based on more efficient shuttling of DMAPP and IPP to the monoterpene precursor GPP by GDS in limonene producing Camelina plants than to FPP by FDS, required for sesquiterpene synthesis, in (+)-δ-cadinene producing plants. The (+)-5-epi-aristolochene producing FE line accumulates generally lower sesquiterpene amounts than the (+)-δ-cadinene producing lines used in this study. This may partially be due to the lack of any MEP pathway genes in the FE expression
3.3. Seed oil accumulation and fatty acid composition Total seed oil content was determined for ten plants of each line, under each growth condition, by NMR and three plants of each line, under each growth condition, by FAME analysis. Both benchtop NMR and FAME analysis showed significant (p ≤ 1E−5) oil accumulation reductions for both DGL and DFC lines compared to parallel grown wild-type plants under all investigated growth conditions, i.e. the greenhouse, the 2014 field trial, and the 2015 field trial (Fig. 4). Notably, the seed oil content of both DGL and DFC lines is equally lowered compared to wild-type Camelina. The two higher terpene yielding lines, DDHHGL and DDHHFC, showed a similar seed oil reduction as the preceding, lower terpene yielding DFC and DGL lines, rather than a further reduced oil accumulation correlating with their increased terpene production, as was expected. Moreover, the FE line, which produces only a small amount of terpene, has seed oil levels widely undistinguishable from parallel-grown wild-type plants. Taken together, corroborating previously reported findings (Augustin et al., 2015), terpene producing Camelina plants accumulate slightly less oil in seeds not only under greenhouse but also under field conditions. The stronger 55
Industrial Crops & Products 136 (2019) 50–58
J.M. Augustin, et al.
fatty acid profile is characterized by an increased ratio of fully saturated fatty acids (e.g. 18:0 and 20:0) as well as fatty acids with two double bonds such as 18:2 and 20:2, and a lowered ratio of fatty acids with three double bonds (e.g. 18:3, 20:3) and to a lesser extent fatty acids with one double bond (e.g. 18:1, 20:1) as compared to wild-type. Terpene production, thus, seems to predominantly impact accumulation of mono- and tri-unsaturated fatty acids, which could result from an altered fatty acid desaturase 2/ fatty acid desaturase 3 activity ratio as observed for Arabidopsis thaliana lines overexpressing the phospholipid flippase ALA10 (Botella et al., 2016). Considering the higher energy content of more saturated fatty acids, the observed shifts in the seed fatty acid composition of terpene producing plants are preferable for use as a biofuel crop.
Table 2 Ratios and terpene analysis of T2 seeds derived from DsRed2-expressing T1 seeds produced by wild-type Camelina plants harvested during the 2014 field season. DsRed2-expressing seed derived from field grown wild-type plants were planted in the greenhouse. For each plant, a sample of 100 representative seeds was used to determine the ratio of DsRed2-expressing seed to non-DsRed2-expressing seed. DsRed2-expressing seeds from each wild-type plant were further analyzed for terpene content. Seeds that did not survive to harvest are labeled with a dash. Plant
Seed type (Red:non-red)
Terpene detected
WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT
– 76:24 70:30 – – 68:32 77:23 75:25 72:28 74:26 77:23 75:25 71:29 74:26 75:25 72:28 – 76:24 76:24 72:28 71:29 78:22 74:26 70:30 – 75:25 66:33
– Epi-aristolochene Epi-aristolochene – – Cadinene Epi-aristolochene Cadinene Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene – Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene – Epi-aristolochene Epi-aristolochene
#1 seed 1 #2 seed 1 #2 seed 2 #2 seed 3 #2 seed 4 #2 seed 5 #3 seed 1 #3 seed 2 #3 seed 3 #4 seed 1 #4 seed 2 #4 seed 3 #4 seed 4 #4 seed 5 #5 seed 1 #5 seed 2 #6 seed 1 #6 seed 2 #7 seed 1 #7 seed 2 #8 seed 1 #8 seed 2 #8 seed 3 #8 seed 4 #9 seed 1 #10 seed 1 #10 seed 2
3.4. Confirmation of short distance outcrossing under field conditions Despite Camelina’s self-fertilizing nature and reported low outcrossing rate (Francis and Warwick, 2009; Walsh et al., 2012), low numbers, i.e. 1–5 per plant, of DsRed2-expressing seeds were found in the harvests of all investigated field grown wild-type plants from the 2014 field season. These wild-type plant-derived DsRed2-expressing seeds were propagated in the greenhouse to investigate whether they were homozygous or heterozygous with respect to the DsRed2 expression trait. All surviving plants, i.e. 22 out of 27, yielded both DsRed2and non-DsRed2-expressing seed in a ratio of approximately 3:1 (Table 2), thereby matching the expected segregation pattern of a transgenic trait from a heterozygous parent. Since all transgenic Camelina lines initially subjected to the field trial were homozygous for the DsRed2 expression trait, this strongly suggests that the DsRed2 expressing seeds identified in wild-type plant harvests originated from cross pollination, rather than resulting from contamination during harvest. In view of findings that Camelina sativa (L.) Crantz, as applied in this study, is able to hybridize with its North American relatives Camelina microcarpa, Camelina alyssum, and Camelina rumelica (SeguinSwartz et al., 2013), the ability of transgenic traits to spread to adjacent Camelina plants is crucial when considering appropriate gene-flow containment measures for large-scale field applications. However, as the outcrossing ability with non-Camelina Brassicaceae species is generally low (Julie-Galau et al., 2014) containment of pollen-mediated gene flow is likely to be feasible. To further investigate which transgenic Camelina lines were involved in the hybridization events, DsRed2 expressing seeds of the heterozygous greenhouse-propagated plants were investigated regarding terpene accumulation (Table 2). Most, i.e. 20 out of 22, hybridization-derived plants accumulated (+)-5-epi-aristolochene in their seeds, which is expected to derive from cross pollination with the FE line. The two remaining plants contained (+)-δ-cadinene thereby indicating origination due to cross pollination with the DFC line. The higher occurrence of crossings with the FE line is not surprising as
seed oil reduction for the more productive (+)-δ-cadinene and (4S)limonene producing lines compared to the low-terpene yield FE line suggests a redirection of resources away from fatty acid biosynthesis to terpene biosynthesis. However, this notion is somewhat contradicted by the absence of further seed oil reduction for the terpene yield-improved DDHHFC and DDHHGL lines as compared to their predecessor lines DFC and DGL, respectively. The seed fatty acid composition of each line remained stable throughout all investigated growth conditions (Supplementary Fig. 1), however, as previously observed in Augustin et al., 2015, differed between wild-type and terpene producing plants (Fig. 5). Notably, the altered seed fatty acid profile of terpene producing plants is generally more pronounced for the higher terpene yielding lines, i.e. DGL, DDHHGL, DFC, and DDHHFC, and less pronounced for the low terpene yielding (+)-5-epi-aristolochene producing FE line. This altered seed
Fig. 6. Schematic of 2014 Camelina field trial. Each horizontal line represents a row of Camelina plants of either the transgenic DGL, DFC, or FE line or wild-type (WT) plants. For each row, ten plants (1–10) were further investigated in regards of plant height, total seed yield, weight per seed, seed terpene accumulation, and total seed oil accumulation. Plants 1–3 and 8–10 of each line derived from opposing ends of each plot, whereas plants 4–7 were selected from the respective plot centers. For FAME analysis, seeds of plants 1, 4, and 8 of each plot were investigated. Plants for which at least one replicate was found to contain (+)-δ-cadinene during seed terpene analysis are indicated within circles. The number of DsRed2 expressing seeds found in the harvests of each wild-type plant is shown within diamonds above each corresponding plant. 56
Industrial Crops & Products 136 (2019) 50–58
J.M. Augustin, et al.
during the 2014 field season wild-type plants were grown adjacent to a row of FE plants in the field (Fig. 6). In addition to cross pollination between wild-type Camelina plants and transgenic Camelina plants, hybridization was further indicated to occur between different transgenic lines, as discovered during the seed terpene analysis (Fig. 3). Individual replicates of three DGL plants and two FE plants were found to contain (+)-δ-cadinene in addition to (4S)-limonene and (+)-5-epiaristolochene, respectively. None of the investigated DFC plant samples appeared to contain other terpenes in addition to (+)-δ-cadinene. This may result from dominance of the (+)-δ-cadinene production trait over other terpene production traits, as was observed in attempts to generate transgenic Camelina lines that simultaneously produce both (4S)-limonene and (+)-δ-cadinene. Though not exclusive to (+)-δ-cadinene, these lines seemed to favor the production of (+)-δ-cadinene over that of (4S)-limonene (manuscript in preparation). None of the wild-type plants from the greenhouse study gave rise to DsRed2 expressing seed, and none of the seed samples of transgenic lines from the greenhouse study were found to contain any additional terpenes besides the expected terpene resulting from the intentionally introduced trait. These consistent observations indicate the absence of any cross pollination events under typical greenhouse conditions. This notion is further corroborated by the non-observation of spreading of selection markers or introduced traits from transgenic to non-transgenic Camelina lines or between transgenic lines during years of Camelina greenhouse studies by these authors, involving dozens of incidences of growing various transgenic Camelina lines in close proximity to each other and to wild-type Camelina. Thus, occurrence of Camelina cross pollination is indicated to require environmental factors that are only available during growth under field conditions, e.g., wind and/or insects. For the 2014 field season, the average percent of DsRed2 expressing seeds per wild-type plant was found to be 0.15%, or approximately 3 seeds per 2,000, which is consistent with previously reported results (Walsh et al., 2012). No cross pollination investigations were conducted during the 2015 field season.
views and opinions of authors expressed herein do not necessarily state or reflect those of the United States Government or any agency thereof. We would like to thank the William H. Danforth Fellowship in Plant Sciences awarded to JRB. Appendix A. Supplementary data Supplementary material related to this article can be found, in the online version, at doi:https://doi.org/10.1016/j.indcrop.2019.04.061. References Augustin, J.M., Higashi, Y., Feng, X., Kutchan, T.M., 2015. Production of mono- and sesquiterpenes in Camelina sativa oilseed. Planta 242, 693–708. Bansal, S., Durrett, T.P., 2016. Camelina sativa: an ideal platform for the metabolic engineering and field production of industrial lipids. Biochimie 120, 9–16. Betancor, M.B., Sprague, M., Usher, S., Sayanova, O., Campbell, P.J., Napier, J.A., Tocher, D.R., 2015. A nutritionally-enhanced oil from transgenic Camelina sativa effectively replaces fish oil as a source of eicosapentaenoic acid for fish. Sci. Rep. 5, 8104. Betancor, M.B., Sprague, M., Montero, D., Usher, S., Sayanova, O., Campbell, P.J., Napier, J.A., Caballero, M.J., Izquierdo, M., Tocher, D.R., 2016a. Replacement of marine fish oil with de novo omega-3 oils from transgenic Camelina sativa in feeds for gilthead sea bream (Sparus aurata L.). Lipids 51, 1171–1191. Betancor, M.B., Sprague, M., Sayanova, O., Usher, S., Metochis, C., Campbell, P.J., Napier, J.A., Tocher, D.R., 2016b. Nutritional evaluation of an EPA-DHA oil from transgenic Camelina sativa in feeds for post-smolt Atlantic Salmon (Salmo salar L.). PLoS One 11, e0159934. Botella, C., Sautron, E., Boudiere, L., Michaud, M., Dubots, E., Yamaryo-Botte, Y., Albrieux, C., Marechal, E., Block, M.A., Jouhet, J., 2016. ALA10, a phospholipid flippase, controls FAD2/FAD3 desaturation of phosphatidylcholine in the ER and affects chloroplast lipid composition in Arabidopsis thaliana. Plant Physiol. 170, 1300–1314. Dai, X.X., Weimer, P.J., Dill-McFarland, K.A., Brandao, V.L.N., Suen, G., Faciola, A.P., 2017. Camelina seed supplementation at two dietary fat levels change ruminal bacterial community composition in a dual-flow continuous culture system. Front. Microbiol. 8. Francis, A., Warwick, S.I., 2009. The biology of Canadian weeds. 142. Camelina alyssum (Mill.) Thell.; C. microcarpa andrz. Ex DC.; C. sativa (L.) Crantz. Can. J. Plant Sci. 89, 791–810. Gunstone, F.D., Harwood, J.L., Dijkstra, A.J., 2007. The Lipid Handbook With CD-ROM. CRC Press, Boca Raton. 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. 14, 252–259. Julie-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. Transgen. Res. 23, 67–74. Kagale, S., Koh, C., Nixon, J., Bollina, V., Clarke, W.E., Tuteja, R., Spillane, C., Robinson, S.J., Links, M.G., Clarke, C., Higgins, E.E., Huebert, T., Sharpe, A.G., Parkin, I.A., 2014. The emerging biofuel crop Camelina sativa retains a highly undifferentiated hexaploid genome structure. Nat. Commun. 5, 3706. Kagale, S., Nixon, J., Khedikar, Y., Pasha, A., Provart, N.J., Clarke, W.E., Bollina, V., Robinson, S.J., Coutu, C., Hegedus, D.D., Sharpe, A.G., Parkin, I.A., 2016. The developmental transcriptome atlas of the biofuel crop Camelina sativa. Plant J. 88, 879–894. Kim, H.J., Silva, J.E., Vu, H.S., Mockaitis, K., Nam, J.W., Cahoon, E.B., 2015. 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. Bot. 66, 4251–4265. Kim, J.W., Koo, B., Nyachoti, C.M., 2017. Digestible, metabolizable, and net energy of camelina cake fed to growing pigs and additivity of energy in mixed diets. J. Anim. Sci. 95, 4037–4044. Larsson, M., 2013. Cultivation and processing of Linum usitatissimum and Camelina sativa in southern Scandinavia during the Roman Iron Age. Veg. Hist. Archaeobot. 22, 509–520. Liang, C., Liu, X., Yiu, S.M., Lim, B.L., 2013. De novo assembly and characterization of Camelina sativa transcriptome by paired-end sequencing. BMC Genomics 14, 146. Liu, J., Rice, A., McGlew, K., Shaw, V., Park, H., Clemente, T., Pollard, M., Ohlrogge, J., Durrett, T.P., 2015. Metabolic engineering of oilseed crops to produce high levels of novel acetyl glyceride oils with reduced viscosity, freezing point and calorific value. Plant Biotechnol. J. 13, 858–865. 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. Malik, M.R., Yang, W., Patterson, N., Tang, J., Wellinghoff, R.L., Preuss, M.L., Burkitt, C., Sharma, N., Ji, Y., Jez, J.M., Peoples, O.P., Jaworski, J.G., Cahoon, E.B., Snell, K.D., 2015. Production of high levels of poly-3-hydroxybutyrate in plastids of Camelina sativa seeds. Plant Biotechnol. J. 13, 675–688. Mansour, M.P., Shrestha, P., Belide, S., Petrie, J.R., Nichols, P.D., Singh, S.P., 2014. Characterization of oilseed lipids from "DHA-producing Camelina sativa": a new transformed land plant containing long-chain omega-3 oils. Nutrients 6, 776–789. Moore, R.H., Thornhill, K.L., Weinzierl, B., Sauer, D., D’Ascoli, E., Kim, J., Lichtenstern,
4. Conclusions This study evaluated the performance of previously generated terpene producing Camelina lines over two growing seasons under field conditions and compared them to greenhouse grown plants to evaluate suitability of the transgenic lines as farmable crops. Our results, in conjunction with those found previously (Usher et al., 2015), support phenotypic stability of transgenic Camelina in the field and therefore further validate its use for the production of agricultural and industrial goods. Terpene accumulation under field conditions was, however, consistently found to be slightly lower when compared to greenhousegrown plants. Of the bioengineered Camelina lines investigated in this study, the (+)-δ-cadinene producing DDHHFC line appears most applicable as a terpene-producing crop, since its wild-type equivalent seed yield outweighs the 25% higher terpene accumulation of the highest (4S)-limonene producing line (DDHHGL), whose seed yield is approximately halved compared to wild-type level. The higher energy density of the seed oil of bioengineered terpene producing Camelina lines renders it more suitable for biofuel applications. Concurrently, this change is expected to lower its value as a food oil. Outcrossing of transgenic traits was observed to occur under field conditions, demonstrating that proper precautions should be taken to prevent cross pollination between transgenic and wild-type Camelina plants. However, the outcrossing rate is low, making appropriate containment feasible. Acknowledgements The information, data, or work presented herein was funded in part by the Advanced Research Projects Agency-Energy (ARPA-E), U.S. Department of Energy, under Award Number DE-AR0000202. The 57
Industrial Crops & Products 136 (2019) 50–58
J.M. Augustin, et al.
Camelina species. Plant Breed. 132, 390–396. Sostaric, R., Alegro, A., Hrsak, V., Stancic, Z., Kuster, H., 2009. Plant remains from an Early Iron Age well at Hajndl, Slovenia. Coll. Antropol. 33, 1295–1301. Usher, S., Haslam, R.P., Ruiz-Lopez, N., Sayanova, O., Napier, J.A., 2015. Field trial evaluation of the accumulation of omega-3 long chain polyunsaturated fatty acids in transgenic Camelina sativa: making fish oil substitutes in plants. Metab. Eng. Commun. 2, 93–98. Vollmann, J., Eynck, C., 2015. Camelina as a sustainable oilseed crop: contributions of plant breeding and genetic engineering. Biotechnol. J. 10, 525–535. Walsh, K.D., Puttick, D.M., Hills, M.J., Yang, R.C., Topinka, K.C., Hall, L.M., 2012. Short Communication: first report of outcrossing rates in camelina [Camelina sativa (L.) Crantz], a potential platform for bioindustrial oils. Can. J. Plant Sci. 92, 681–685. Yu, X.H., Cahoon, R.E., Horn, P.J., Shi, H., Prakash, R.R., Cai, Y., Hearney, M., Chapman, K.D., Cahoon, E.B., Schwender, J., Shanklin, J., 2017. Identification of bottlenecks in the accumulation of cyclic fatty acids in camelina seed oil. Plant Biotechnol. J. Zhu, L.H., Krens, F., Smith, M.A., Li, X., Qi, W., van Loo, E.N., Iven, T., Feussner, I., Nazarenus, T.J., Huai, D., Taylor, D.C., Zhou, X.R., Green, A.G., Shockey, J., Klasson, K.T., Mullen, R.T., Huang, B., Dyer, J.M., Cahoon, E.B., 2016. Dedicated industrial oilseed crops as metabolic engineering platforms for sustainable industrial feedstock production. Sci. Rep. 6, 22181.
M., Scheibe, M., Beaton, B., Beyersdorf, A.J., Barrick, J., Bulzan, D., Corr, C.A., Crosbie, E., Jurkat, T., Martin, R., Riddick, D., Shook, M., Slover, G., Voigt, C., White, R., Winstead, E., Yasky, R., Ziemba, L.D., Brown, A., Schlager, H., Anderson, B.E., 2017. Biofuel blending reduces particle emissions from aircraft engines at cruise conditions. Nature 543, 411–415. Moser, B.R., 2012. Biodiesel from alternative oilseed feedstocks: camelina and field pennycres. Biofuels 2012 (3), 17. Nguyen, H.T., Silva, J.E., Podicheti, R., Macrander, J., Yang, W., Nazarenus, T.J., Nam, J.W., Jaworski, J.G., Lu, C., Scheffler, B.E., Mockaitis, K., Cahoon, E.B., 2013. Camelina seed transcriptome: a tool for meal and oil improvement and translational research. Plant Biotechnol. J. 11, 759–769. Petrie, J.R., Shrestha, P., Belide, S., Kennedy, Y., Lester, G., Liu, Q., Divi, U.K., Mulder, R.J., Mansour, M.P., Nichols, P.D., Singh, S.P., 2014. Metabolic engineering Camelina sativa with fish oil-like levels of DHA. PLoS One 9, e85061. Putnam, D.H., Budin, J.T., Field, L.A., BW, M., 1993. Camelina: A Promising Low-Input Oilseed. Wiley, New York. Ruiz-Lopez, N., Haslam, R.P., Napier, J.A., 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. Seguin-Swartz, G., Nettleton, J.A., Sauder, C., Warwick, S.I., Gugel, R.K., 2013. Hybridization between Camelina sativa (L.) Crantz (false flax) and North American
58
Contents lists available at ScienceDirect
Industrial Crops & Products journal homepage: www.elsevier.com/locate/indcrop
Field performance of terpene-producing Camelina sativa a,b
a,c
a
T a,d
Jörg M. Augustin , Jordan R. Brock , Megan M. Augustin , Rachel L. Wellinghoff , ⁎ Matthew Shippa,e, Yasuhiro Higashia,f, Tadele T. Kumssag,h, Edgar B. Cahoong, Toni M. Kutchana, a
Donald Danforth Plant Science Center, 975 North Warson Rd, St. Louis, MO, 63132, United States Elemental Enzymes, 1685 Galt Industrial Blvd, St. Louis, MO, 63132, United States Washington University in St. Louis, Department of Biology, 1 Brookings Dr, St. Louis, MO, 63130, United States d Cofactor Genomics, 4044 Clayton Ave, St. Louis, MO, 63110, United States e KWS, 1005 N Warson Rd, St. Louis, MO, 63132, United States f RIKEN Center for Sustainable Resource Science, 1-7-22 Suehiro, Tsurumi, Yokohama, Kanagawa, 230-0045, Japan g University of Nebraska-Lincoln, 1400 R St, Lincoln, NE, 68588, United States h Noble Research Institute, 2510 Sam Noble Pkwy, Ardmore, OK 73401, United States b c
A R T I C LE I N FO
A B S T R A C T
Keywords: Terpene (+)-δ-cadinene (4S)-limonene (+)-5-epi-aristolochene Field trial Camelina sativa
Bioengineered lines of the low-input oil seed crop Camelina sativa (Camelina), augmented to accumulate the monoterpene (4S)-limonene, the sesquiterpene (+)-δ-cadinene, or the sesquiterpene (+)-5-epi-aristolochene in seed, were evaluated for two growing seasons under field conditions to determine performance of the introduced traits in an agricultural setup including the effects on overall plant fitness, and total seed yield. Field-grown Camelina plants were further compared to greenhouse-grown plants to evaluate commonalities and differences resulting from cultivation under either controlled or agriculturally relevant growth conditions. Morphological appearance and plant height differed marginally between transgenic and wild-type plants under both greenhouse and field conditions, indicating low impact of the terpene production traits toward overall plant fitness. Total seed yield, however, was independent of the growth conditions and was reduced in Camelina lines producing (4S)-limonene by 48% on average and by 30% on average for (+)-5-epi-aristolochene producing lines. Conversely, (+)-δ-cadinene producing Camelina seed yields remained wild-type equivalent. Additional investigations included seed terpene accumulation, seed oil amount, and seed fatty acid composition. Terpene accumulation was reduced up to 22% for field-grown plants as compared to greenhouse-grown plants. Seed oil amounts were similar under greenhouse and field conditions but were consistently lowered by 2–5% for terpene producing lines. Similarly, seed oil composition remained stable under both field and greenhouse conditions, but generally favored a more energy dense phenotype in terpene producing Camelina lines. Lastly, outcrossing of transgenic traits to adjacent wild-type Camelina plants was observed under field but not greenhouse conditions.
1. Introduction The re-emerging oilseed crop Camelina sativa (Camelina) is currently under consideration as a renewable, farmable source for the production of a variety of industrial and agricultural products due to its amendable properties in the field. Camelina is a low-input oil seed crop
that is thought to have been originally cultivated by man during the Iron Age (Larsson, 2013; Sostaric et al., 2009) as a food oil source, but was later abandoned due to the availability of higher yielding oil crops. In recent years Camelina has regained interest as a crop with minimal fertilization requirements and drought resistance traits making it suitable for growth in marginal soils and drier climates (Bansal and
Abbreviations: MEP, methylerythritol phosphate; GDS, geranyl diphosphate synthase; FDS, farnesyl diphosphate synthase; LS, (4S)-limonene synthase; CDNS, (+)-δcadinene synthase; EAS, (+)-5-epi-aristolochene synthase; DXS, 1-deoxy-D-xylulose-5-phosphate synthase; DXR, 1-deoxy-D-xylulose 5-phosphate reductoisomerase; HDS, (E)-4-hydroxy-3-methylbut-2-enyl diphosphate synthase; HDR, (E)-4-hydroxy-3-methylbut-2-enyl diphosphate reductase; DGL, transgenic Camelina sativa expressing DXS, GDS and LS; DFC, transgenic Camelina sativa expressing DXS, FDS and CDNS; FE, transgenic Camelina sativa expressing FDS and EAS; DDHHGL, transgenic Camelina sativa expressing DXS, DXR, HDS, HDR, GDS and LS; DDHHFC, transgenic Camelina sativa expressing DXS, DXR, HDS, HDR, FDS and CDNS ⁎ Corresponding author. E-mail addresses: [email protected] (J.M. Augustin), [email protected] (J.R. Brock), [email protected] (M.M. Augustin), rwellinghoff@gmail.com (R.L. Wellinghoff), [email protected] (M. Shipp), [email protected] (Y. Higashi), [email protected] (T.T. Kumssa), [email protected] (E.B. Cahoon), [email protected] (T.M. Kutchan). https://doi.org/10.1016/j.indcrop.2019.04.061 Received 15 March 2019; Received in revised form 19 April 2019; Accepted 28 April 2019 Available online 06 May 2019 0926-6690/ © 2019 Elsevier B.V. All rights reserved.
Industrial Crops & Products 136 (2019) 50–58
J.M. Augustin, et al.
Durrett, 2016; Putnam et al., 1993). Its compatibility with present day agricultural practices is expected to ease potential transiting into modern cultivation (Moser, 2012). Camelina has further shown resistance to a number of diseases and pests including blackleg, blackspot, Alternaria spp., and flea beetles that plague cruciferous oil crops such as rapeseed and its derivative Canola (Putnam et al., 1993; Vollmann and Eynck, 2015). Lastly, another reason for the renewed interest in Camelina is its high accumulation of unsaturated fatty acids including oleic (18:1), linoleic (18:2), linolenic (18:3), and eicosenoic (20:1) acid, which, compared to saturated fatty acids, have both health benefits for human consumption and lower melting points resulting, upon conversion into biodiesels, in fuel blends that remain liquid longer at cooler temperatures (Gunstone et al., 2007; Putnam et al., 1993). Genetic engineering of Camelina is feasible due to its ease of stable transformation, close relatedness to the well-studied model plant Arabidopsis thaliana, and the availability of genomic and transcriptomic datasets, which aid in relatively straightforward manipulability for specific industries (Kagale et al., 2014, 2016; Liang et al., 2013; Lu and Kang, 2008; Nguyen et al., 2013). Extensive work has been dedicated to alter Camelina’s lipid profile for production of wax esters, poly-3-hydroxybutyrate, cyclic fatty acids, and acetyl-triacylglycerols for, e.g., use as industrial lubricants, cosmetics, and plastics (Iven et al., 2015; Liu et al., 2015; Malik et al., 2015; Yu et al., 2017; Zhu et al., 2016). Additionally, modifications of the lipid profile of Camelina seed oil toward the production of omega-3 long-chain polyunsaturated fatty acids has shown great promise for replacement of fish oil in aquaculture (Betancor et al., 2016a, b; Betancor et al., 2015; Mansour et al., 2014; Petrie et al., 2014; Ruiz-Lopez et al., 2014). The oils of both natural accessions and genetically modified Camelina lines have further been highlighted as a potential biofuel source for biodiesel and Jet A fuel drop-ins with significantly lower particle emissions (Kim et al., 2015; Moore et al., 2017; Moser, 2012). Production of high value compounds in Camelina can increase the overall crop value. For example, using modified oil as a renewable source material for industrial or pharmaceutical applications increases the farmer’s income significantly as opposed to using the material as an addition to animal feed alone (Dai et al., 2017; Kim et al., 2017). Previously, we bioengineered Camelina lines to produce and accumulate the industrially relevant terpenes (4S)-limonene and (+)-δ-cadinene in seed by plastidial localization of 1) limonene synthase (LS), geranyl diphosphate synthase (GDS) and 1-deoxy-D-xylulose-5-phosphate synthase (DXS), or 2) (+)-δ-cadinene synthase (CDNS), farnesyl diphosphate synthase (FDS) and DXS; abbreviated as DGL and DFC lines, respectively (Augustin et al., 2015). Targeting enzymes to the plastids resulted in higher terpene accumulation due to the utilization of dimethylallyl pyrophosphate (DMAPP) and isopentenyl pyrophosphate (IPP) originating from the plastid-located methylerythritol phosphate (MEP) pathway (Fig. 1). Our previous results demonstrated that when these transgenic Camelina plants were grown under greenhouse conditions, the production of (4S)-limonene is stable for at least five consecutive generations. Viability of the terpene traits under field conditions in an agricultural setup, however, remained thus far uninvestigated. Here we present the evaluation of the DGL and DFC lines as well as a third line that produces the sesquiterpene (+)-5-epi-aristolochene, termed FE, under actual field conditions for two consecutive years and compare their morphology, yield, oil content, and terpene production to greenhouse grown DGL, DFC, and FE plants. During the second field season we further expanded the study to include two improved terpene producing lines, designated DDHHGL and DDHHFC (manuscript in preparation), that overexpress additional MEP pathway genes. Lastly, the observation of outcrossing events between the transgenic lines and adjacent wild-type plants during the first field season is discussed.
Fig. 1. Schematic of pathways leading to the biosynthesis of (4S)-limonene, (+)-δ-cadinene, and (+)-5-epi-aristolochene in transgenic Camelina plants of the DGL/DDHHGL, DFC/DDHHFC and FE lines, respectively. Isopentenyl pyrophosphate (IPP) and dimethylallyl pyrophosphate (DMAPP) derived from the plastid-localized MEP pathway are coupled by either geranyl diphosphate synthase (GDS) or farnesyl diphosphate synthase (FDS) to produce geranyl pyrophosphate (GPP). In DGL and DDHHGL plants, GPP is subsequently cyclized into (4S)-limonene by limonene synthase (LS). In DFC, DDHHFC, and FE plants, FDS catalyzes the linkage of another IPP molecule to GPP yielding farnesyl pyrophosphate (FPP), which is then cyclized by either cadinene synthase (CDNS) into (+)-δ-cadinene in DFC and DDHHFC plants or by epi-aristolochene synthase (EAS) into (+)-epi-aristolochene in FE plants.
2. Methods 2.1. Preparation of transgenic Camelina lines Generation of the Camelina sativa Suneson DFC and DGL lines was described previously (Augustin et al., 2015). For generation of the DDHHFC and DDHHGL lines, the transformation constructs used to create the DFC and DGL lines were stepwise extended with the MEP pathway genes encoding 1-deoxy-D-xylulose 5-phosphate reductoisomerase (DXR), (E)-4-hydroxy-3-methylbut-2-enyl diphosphate synthase (HDS), and (E)-4-hydroxy-3-methylbut-2-enyl diphosphate reductase (HDR) from Mentha piperita (manuscript in preparation). Similar to DXS, expression of DXR, HDS, and HDR is controlled by identically structured Glycine max glycinin expression cassettes. The transformation construct for the FE line derived from a precursor of the DFC transformation construct lacking the DXS expression cassette and carrying the (+)-5-epi-aristolochene synthase (EAS) ORF from Nicotiana sp. in place of the CDNS ORF. All transformation constructs were introduced into Agrobacterium tumefaciens GV3101:pMP90, which was subsequently used to transform wild-type Camelina sativa Suneson plants using the floral-dip method as described previously (Augustin et al., 2015; Lu and Kang, 2008). Transgenic seeds were screened using a green LED light in combination with a red filter to identify the presence of the selection marker DsRed2. The DsRed2 expression cassette was included in the TDNA section of
51
Industrial Crops & Products 136 (2019) 50–58
J.M. Augustin, et al.
2.4. FAME analysis
each plant transformation construct. Transgenic plants were propagated until homozygous plant lines, determined by ubiquitous presence of the DsRed2 trait in all derived seeds, were obtained.
For determination of seed fatty acid profiles, triplicate fatty acid methyl ester (FAME) extractions were performed on a total of ten seeds per replicate. Seeds were combined, weighed, and transferred to screw top glass tubes followed by addition of 1.5 ml of 2.5% sulfuric acid in methanol and ground using a glass rod. Subsequently, 50 μg triheptadecanoin (per mg seed) was solubilized in a total volume of 500 μl toluene and added to each sample as internal standard. Samples were incubated at 95 °C for 50 min then cooled to room temperature prior to addition of 1 ml hexane and 1 ml 1 M NaCl. FAMEs were extracted into the hexane phase by rapid mixing. Subsequently, samples were centrifuged at 445 x g for 5 min to facilitate phase separation. Aliquots of the hexane phase were transferred to screw top autosampler vials and analyzed by GC-FID on a Thermoquest Trace GC 2000 system equipped with an Agilent HP-INNOwax column (30 m x 250 μm x 0.25 μm) using helium as the carrier gas. GC conditions began with a hold of 60 °C for 1 min, increasing to 185 °C at a rate of 40 °C min–1, then rising to 235 °C at a rate of 5 °C min–1 with a 5 min hold. FAME species identification was based on retention time comparison to known standards. Fatty acid abundances are reported as a percentage of each FAME peak area of the total peak area of all FAME peaks combined. Absolute abundances were further quantified relative to the triheptadecanoin internal standard and used to estimate total seed oil content.
2.2. Plant trial metrics Field trials were conducted in Mead, Nebraska, USA during April and June of 2014 and April and July of 2015. For the 2014 field season, seeds of homozygous T4 generation DFC, T4 DGL, T3 FE plants, and wild-type Camelina sativa Suneson were planted in single 2–3.5 m long rows. In the 2015 field season, in addition to cultivating seeds of homozygous T4 DFC, T4 DGL, T3 FE, and wild-type Camelina plants, T4 DDHHFC and T4 DDHHGL seeds were planted as well. In 2015, each line was planted in plots of four adjacent rows of approximately 2 m in length. The distance between rows within plots was approximately 0.15 m and the distance between individual plots 0.9 m. Greenhousepropagated DFC, DGL, and FE seed sowed in the 2015 field trial were prepared from the same T3 and T2 generation parental seed batches used to propagate seed for the 2014 field season. Lines grown in the greenhouse for comparison to the field were cultivated at the Donald Danforth Plant Science Center, St. Louis, Missouri, USA from July to October of 2014. Greenhouse conditions were 21 °C during the day and 20 °C in the night and within a humidity range of 40–90%. Seed used for the greenhouse comparison derived from the same T4 DFC, T4 DGL, and T3 FE line and wild-type Camelina sativa Suneson seed batches that were planted in the 2014 field trial. For both the field trials and the greenhouse comparison, ten individual plants were selected per line and used to determine average plant height, total seed yield, seed weight, seed terpene accumulation, and seed oil accumulation. Seed of each harvested plant were screened as described above for expression of DsRed2, indicative for the presence of transgenic traits.
2.5. NMR oil analysis Total seed oil content was quantified using a benchtop NMR analyzer MQC (Oxford Instruments). The instrument was tuned with a 40 °C standard of extracted Camelina oil before each set of runs. Weighed seeds (0.3 g — 0.5 g) were transferred to clean 10 mm glass sample tubes and incubated at 40 °C for at least 10 min prior to analysis. 3. Results and discussion
2.3. Terpene extraction and analysis
3.1. Phenotypic analysis
Typically, three terpene extracts were prepared and measured per plant. Ten individual seeds per extraction were combined and weighed. Each seed batch was subsequently placed in a glass screw top tube and submerged in 2 ml diethyl ether. After addition of 50 μl of an internal standard stock solution, seeds were ground to a fine powder using a glass rod. Internal standards were 400 ng/μl iso-butylbenzene for (4S)limonene analyses and 400 ng/μl hexadecane for (+)-δ-cadinene and (+)-5-epi-aristolochene analyses. Sealed tubes were incubated at room temperature for a minimum of 2 h while shaking at 150 rpm and inverted approximately every 15 min. Tubes were then centrifuged at 1000×g for 10 min and 150 μl supernatant was transferred to screw top autosampler vials, which were subsequently centrifuged again at 1000×g for 5 min. Finally, extracts were analyzed by GC–MS after injection of 1 μl sample onto an Agilent 7890 A GC equipped with a Restek Rtx-5MS column (30 m x 250 μm x 0.25 μm) connected to a 5975C inert XL MSD mass spectrometer using helium as carrier gas. For (4S)-limonene quantification, the oven temperature started with a hold at 30 °C for 3 min, then raised to 85 °C at a rate of 55 °C min–1, further raised to 120 °C at a rate of 10 °C min–1, and finally raised to 300 °C at 120 °C min–1 with a 3 min hold. Quantification of (+)-δ-Cadinene and (+)-5-epi-aristolochene containing samples began with an oven temperature of 50 °C for 3 min, then raised to 160 °C at a rate of 80 °C min–1, raised once more to 170 °C at a rate of 1.4 °C min–1, and finally raised to 300 °C at 120 °C min–1 and held for 3 min. (4S)Limonene concentrations were calculated based on a standard curve of commercial (4S)-limonene (Sigma-Aldrich). (+)-δ-Cadinene and (+)-5-epi-aristolochene concentrations were calculated relative to a standard curve of the sesquiterpene (+)-valencene (Sigma-Aldrich).
Terpene producing DFC, DGL, and FE lines of Camelina were compared to wild-type plants grown during field seasons in 2014 and 2015 as well as to plants grown under greenhouse conditions in 2014. In the 2015 field season, two additional terpene yield-improved lines designated DDHHFC and DDHHGL were investigated under field conditions as well. Notably, within each of the trials, adult plants at harvest did not show any obvious morphological differences between the terpene producing lines and wild-type plants (Fig. 2 A). In contrast, overall morphology of Camelina plants grown under field and greenhouse conditions differed substantially. Field grown plants developed thicker stems than greenhouse grown plants and lost all leaves in the field by the time of harvest (Fig. 2B). Similarly, average plant heights at harvest of all field grown Camelina lines were widely consistent throughout both seasons and no significant differences (p ≤ 0.05) were observed between terpene producing lines and wild-type plants while greenhouse grown plants were an average of 5–10 cm taller at harvest (Table 1). Differences in both overall morphology and plant height are likely the result of adaptions to the different physical constraints under the different growth conditions (i.e. exposure to wind and rain under field conditions favored the development of shorter, sturdier plants with stronger stems). Average seed yields per plant fluctuated strongly between the 2014 and 2015 field seasons, whereas average seed yields per plant in the 2015 field season and greenhouse grown plants resembled each other more closely (Table 1). The higher yield of the 2014 season may be due to increased physical space. In 2014, plants were grown in single rows as opposed to more compact patches in the 2015 field season, which more closely resembles growth in the greenhouse where plants are grown in trays. However, yield differences due to variable weather 52
Industrial Crops & Products 136 (2019) 50–58
J.M. Augustin, et al.
Fig. 2. Morphology of representative wild-type and terpene-producing transgenic Camelina plants either grown under greenhouse or field conditions. A) Field-grown Camelina plants from 2014 field season. B) Greenhouse-grown Camelina plants. Images were taken at day of harvest. Table 1 Phenotypic traits of wild-type (WT) and terpene producing transgenic Camelina lines (DFC, DGL, FE, DDHHFC, and DDHHGL) grown under greenhouse or field conditions. Each data point represents the average height (cm), seed yield (g), or individual seed weight (mg) of ten individual plants as indicated. For the individual seed weights, three samples of ten seeds were weighed per plant and the resulting weight of each sample divided by the number of seed. Error range represents standard error. Height (cm)
WT DFC DGL FE DDHHFC DDHHGL
Seed yield (g)
Individual seed weight (mg)
Greenhouse
Field 2014
Field 2015
Greenhouse
Field 2014
Field 2015
Greenhouse
Field 2014
Field 2015
82.2 81.6 81.3 80.2 – –
77.3 74.7 75.0 70.6 – –
72.1 74.4 74.4 69.5 74.6 74.2
1.36 1.50 0.91 1.07 – –
3.08 3.63 1.62 2.25 – –
1.56 1.48 0.75 0.92 1.57 0.81
1.09 1.10 0.90 1.06 – –
0.93 1.02 0.92 1.22 – –
0.73 0.98 0.99 1.06 1.09 1.03
± ± ± ±
0.7 1.6 2.1 2.7
± ± ± ±
2.0 1.1 1.8 1.6
± ± ± ± ± ±
2.3 2.6 2.2 1.9 2.1 1.6
± ± ± ±
0.08 0.10 0.10 0.10
conditions during the two field seasons cannot be excluded. Notably, the (+)-δ-cadinene producing Camelina lines, DFC and DDHHFC, had comparable per plant seed yields in all trials as concurrently grown wild-type plants, averaging 106% of wild-type level. In contrast, both (4S)-limonene producing Camelina lines, DGL and DDHHGL, with an average per plant seed yield throughout all trials of 52% of wild-type levels, as well as the (+)-5-epi-aristolochene producing FE line, averaging at 70% wild-type per plant seed yields, had consistently lower average yields than wild-type plants. This strongly indicates that the in planta production of (4S)-limonene and (+)-5-epi-aristolochene
± ± ± ±
0.65 0.42 0.22 0.30
± ± ± ± ± ±
0.37 0.59 0.14 0.30 0.42 0.17
± ± ± ±
0.02 0.02 0.03 0.01
± ± ± ±
0.03 0.02 0.01 0.02
± ± ± ± ± ±
0.02 0.03 0.03 0.07 0.04 0.06
negatively affects seed yield, which may render these lines less desirable as a field crop. However, the wild-type yield levels of the (+)-δcadinene producing plant lines contradict a general notion that seedspecific terpene production causes a detrimental effect on total seed yield. Thus, additional, independently generated DGL, DDHHGL, and FE lines should be evaluated to exclude line-specific low yield traits as plants were selected based upon terpene yield, not seed yield. The weight per seed was found to be generally consistent at 1 ± 0.1 mg seed−1 for all investigated lines throughout all trials. Exceptions were wild-type plants in the 2015 field season, which 53
Industrial Crops & Products 136 (2019) 50–58
J.M. Augustin, et al.
Fig. 4. Seed oil accumulation in wild-type and terpene producing transgenic Camelina plants either grown under greenhouse or field conditions. A) Oil accumulation in seed of wild-type (WT) Camelina and terpene producing transgenic DFC, DGL, FE, DDHHGL, and DDHHFC lines determined by FAME analysis. Three individual plants were investigated in three technical replicates per line and growth condition. B) Oil accumulation in seed of identical Camelina lines presented in panel A) determined by NMR analysis. Per line and growth condition, ten individual plants were investigated in one to five technical replicates (depending on seed availability). Error bars depict combined standard deviations.
Fig. 3. Seed terpene accumulation in transgenic Camelina plants either grown under greenhouse or field conditions. A) (+)-δ-Cadinene accumulation of DFC line grown under greenhouse conditions, under field conditions during 2014 field season (2014 FT) and 2015 field season (2015 FT), as well as DDHHFC line grown under field conditions during 2015 field season (DDHHFC). B) (4S)Limonene accumulation of DGL line grown under greenhouse conditions and field conditions (2014 FT, 2015 FT), as well as of the DDHHGL line grown under field conditions during 2015 field season (DDHHGL). C) (+)-5-epiAristolochene accumulation of FE line grown under greenhouse conditions and field conditions (2014 FT, 2015 FT). Per line and growth condition, ten individual plants were investigated in three technical replicates. Error bars depict combined standard deviation.
and were, thus, excluded from terpene quantitation. All investigated plants of the DFC line contained (+)-δ-cadinene in their seed, which averaged at 5.9 ( ± 0.7) mg g−1 in plants grown under greenhouse conditions, 4.9 ( ± 0.4) mg g−1 in seed of field trial plants from 2014, and 4.9 ( ± 1.3) mg g−1 in seed of field trial plants from 2015. The DDHHFC line, which was exclusively investigated during the 2015 field season, accumulated 7.4 ( ± 1.3) mg g−1 (+)-δ-cadinene in its seeds and, thus, showed substantially higher amounts as compared to the DFC line grown under identical conditions (Fig. 3A). Plants of the DGL line accumulated 10.5 ( ± 1.1) mg (4S)-limonene g−1 seed in the greenhouse, 9.1 ( ± 0.7) mg g−1 under field conditions in 2014, and 7.9 ( ± 1.3) mg g−1 under field conditions in 2015. In parallel grown DDHHGL plants, terpene accumulation averaged 9.3 ( ± 1.3) mg (4S)limonene g−1 seed during the 2015 field season (Fig. 3B). The FE line accumulated an average of 2.8 ( ± 1.2) mg (+)-5-epi-aristolochene g−1 seed under greenhouse conditions, 1.4 ( ± 0.4) mg g−1 under field condition in 2014, and only 0.3 ( ± 0.3) mg g−1 under field conditions in 2015 (Fig. 3C). As previously reported (Augustin et al., 2015), we detected glycosylated terpene products in both DGL and DFC lines, which was shown to increase the total terpene content by up to 30%. However, due to difficulties in quantitation of glycosylated (+)-δ-cadinene, the values reported here do not include glycosylated terpenes.
produced smaller seed with an average of 0.7 mg seed−1, and the FE line during the 2014 field season with slightly larger seeds that averaged 1.2 mg seed−1 (Table 1). Considering the very comparable (DFC) or even slightly higher weights per seed (DGL and FE) under field conditions, the reduced seed weight of wild-type Camelina in the field is unlikely a general trend for growing Camelina under field conditions as compared to cultivation in the greenhouse.
3.2. Seed terpene accumulation To assess effectiveness of the introduced terpene traits under the different growth conditions, seeds of ten individual plants of each transgenic line grown in the greenhouse, in the field in 2014, and in the field in 2015 were analyzed for terpene content. Wild-type Camelina seeds are devoid of mono- and sesquiterpenes (Augustin et al., 2015) 54
Industrial Crops & Products 136 (2019) 50–58
J.M. Augustin, et al.
Fig. 5. Seed fatty acid composition of wild-type and terpene producing transgenic Camelina plants either grown under greenhouse or field conditions. A) Seed fatty acid composition of wildtype (WT) and transgenic DGL, DFC, and FE Camelina lines grown in the greenhouse. B) Seed fatty acid composition of WT and transgenic DGL, DFC, and FE Camelina plants grown under field conditions during 2014 field season. C) Seed fatty acid composition of WT and transgenic DGL, DFC, FE, DDHHGL, and DDHHFC Camelina plants grown under field conditions during 2015 field season. Per line and growth condition, three individual plants were investigated using three technical replicates. Error bars depict combined standard deviation.
construct. However, previous greenhouse comparisons of FE and FC lines, a DFC precursor line not overexpressing DXS (Augustin et al., 2015), also resulted in higher sesquiterpene accumulations for the (+)-δ-cadinene producing line than the (+)-5-epi-aristolochene producing line (data not shown). This indicates (+)-δ-cadinene is more efficiently produced, e.g. due to higher expression or activity rates of the involved sesquiterpene synthase, or more efficiently accumulated in Camelina seed than (+)-5-epi-aristolochene.
Therefore, the total terpene content in all lines analyzed may be significantly higher if the amount of glycosylated terpenes is taken into account. In summary, all three lines, DFC, DGL, and FE, accumulate more terpene under greenhouse conditions than under field conditions. Of the two field seasons, 2014 appeared more beneficial toward terpene production than 2015. Considering the generally higher seed yields in 2014 compared to 2015 (Table 1), it is possible that the higher terpene yield under 2014 field conditions resulted from generally elevated plant vigor. The two Camelina lines that carry, in addition, gene cassettes to overexpress DXR, HDS, and HDR (manuscript in preparation), i.e. DDHHFC and DDHHGL, both accumulate more cadinene and limonene, respectively, than the corresponding predecessor lines, DFC and DGL, under field conditions in 2015. This indicates that overexpression of additional MEP pathway enzymes in Camelina seed, which generates the terpene precursor molecules DMAPP and IPP in plastids, leads also to further increased terpene production under field conditions as previously seen in greenhouse studies (manuscript in preparation). Previously, limonene producing plants were found to accumulate more terpene (in mg g−1 seed weight) than (+)-δ-cadinene producing lines (Augustin et al., 2015). This observation holds up throughout all growth conditions included in this study. The reason is unknown but may be based on more efficient shuttling of DMAPP and IPP to the monoterpene precursor GPP by GDS in limonene producing Camelina plants than to FPP by FDS, required for sesquiterpene synthesis, in (+)-δ-cadinene producing plants. The (+)-5-epi-aristolochene producing FE line accumulates generally lower sesquiterpene amounts than the (+)-δ-cadinene producing lines used in this study. This may partially be due to the lack of any MEP pathway genes in the FE expression
3.3. Seed oil accumulation and fatty acid composition Total seed oil content was determined for ten plants of each line, under each growth condition, by NMR and three plants of each line, under each growth condition, by FAME analysis. Both benchtop NMR and FAME analysis showed significant (p ≤ 1E−5) oil accumulation reductions for both DGL and DFC lines compared to parallel grown wild-type plants under all investigated growth conditions, i.e. the greenhouse, the 2014 field trial, and the 2015 field trial (Fig. 4). Notably, the seed oil content of both DGL and DFC lines is equally lowered compared to wild-type Camelina. The two higher terpene yielding lines, DDHHGL and DDHHFC, showed a similar seed oil reduction as the preceding, lower terpene yielding DFC and DGL lines, rather than a further reduced oil accumulation correlating with their increased terpene production, as was expected. Moreover, the FE line, which produces only a small amount of terpene, has seed oil levels widely undistinguishable from parallel-grown wild-type plants. Taken together, corroborating previously reported findings (Augustin et al., 2015), terpene producing Camelina plants accumulate slightly less oil in seeds not only under greenhouse but also under field conditions. The stronger 55
Industrial Crops & Products 136 (2019) 50–58
J.M. Augustin, et al.
fatty acid profile is characterized by an increased ratio of fully saturated fatty acids (e.g. 18:0 and 20:0) as well as fatty acids with two double bonds such as 18:2 and 20:2, and a lowered ratio of fatty acids with three double bonds (e.g. 18:3, 20:3) and to a lesser extent fatty acids with one double bond (e.g. 18:1, 20:1) as compared to wild-type. Terpene production, thus, seems to predominantly impact accumulation of mono- and tri-unsaturated fatty acids, which could result from an altered fatty acid desaturase 2/ fatty acid desaturase 3 activity ratio as observed for Arabidopsis thaliana lines overexpressing the phospholipid flippase ALA10 (Botella et al., 2016). Considering the higher energy content of more saturated fatty acids, the observed shifts in the seed fatty acid composition of terpene producing plants are preferable for use as a biofuel crop.
Table 2 Ratios and terpene analysis of T2 seeds derived from DsRed2-expressing T1 seeds produced by wild-type Camelina plants harvested during the 2014 field season. DsRed2-expressing seed derived from field grown wild-type plants were planted in the greenhouse. For each plant, a sample of 100 representative seeds was used to determine the ratio of DsRed2-expressing seed to non-DsRed2-expressing seed. DsRed2-expressing seeds from each wild-type plant were further analyzed for terpene content. Seeds that did not survive to harvest are labeled with a dash. Plant
Seed type (Red:non-red)
Terpene detected
WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT WT
– 76:24 70:30 – – 68:32 77:23 75:25 72:28 74:26 77:23 75:25 71:29 74:26 75:25 72:28 – 76:24 76:24 72:28 71:29 78:22 74:26 70:30 – 75:25 66:33
– Epi-aristolochene Epi-aristolochene – – Cadinene Epi-aristolochene Cadinene Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene – Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene Epi-aristolochene – Epi-aristolochene Epi-aristolochene
#1 seed 1 #2 seed 1 #2 seed 2 #2 seed 3 #2 seed 4 #2 seed 5 #3 seed 1 #3 seed 2 #3 seed 3 #4 seed 1 #4 seed 2 #4 seed 3 #4 seed 4 #4 seed 5 #5 seed 1 #5 seed 2 #6 seed 1 #6 seed 2 #7 seed 1 #7 seed 2 #8 seed 1 #8 seed 2 #8 seed 3 #8 seed 4 #9 seed 1 #10 seed 1 #10 seed 2
3.4. Confirmation of short distance outcrossing under field conditions Despite Camelina’s self-fertilizing nature and reported low outcrossing rate (Francis and Warwick, 2009; Walsh et al., 2012), low numbers, i.e. 1–5 per plant, of DsRed2-expressing seeds were found in the harvests of all investigated field grown wild-type plants from the 2014 field season. These wild-type plant-derived DsRed2-expressing seeds were propagated in the greenhouse to investigate whether they were homozygous or heterozygous with respect to the DsRed2 expression trait. All surviving plants, i.e. 22 out of 27, yielded both DsRed2and non-DsRed2-expressing seed in a ratio of approximately 3:1 (Table 2), thereby matching the expected segregation pattern of a transgenic trait from a heterozygous parent. Since all transgenic Camelina lines initially subjected to the field trial were homozygous for the DsRed2 expression trait, this strongly suggests that the DsRed2 expressing seeds identified in wild-type plant harvests originated from cross pollination, rather than resulting from contamination during harvest. In view of findings that Camelina sativa (L.) Crantz, as applied in this study, is able to hybridize with its North American relatives Camelina microcarpa, Camelina alyssum, and Camelina rumelica (SeguinSwartz et al., 2013), the ability of transgenic traits to spread to adjacent Camelina plants is crucial when considering appropriate gene-flow containment measures for large-scale field applications. However, as the outcrossing ability with non-Camelina Brassicaceae species is generally low (Julie-Galau et al., 2014) containment of pollen-mediated gene flow is likely to be feasible. To further investigate which transgenic Camelina lines were involved in the hybridization events, DsRed2 expressing seeds of the heterozygous greenhouse-propagated plants were investigated regarding terpene accumulation (Table 2). Most, i.e. 20 out of 22, hybridization-derived plants accumulated (+)-5-epi-aristolochene in their seeds, which is expected to derive from cross pollination with the FE line. The two remaining plants contained (+)-δ-cadinene thereby indicating origination due to cross pollination with the DFC line. The higher occurrence of crossings with the FE line is not surprising as
seed oil reduction for the more productive (+)-δ-cadinene and (4S)limonene producing lines compared to the low-terpene yield FE line suggests a redirection of resources away from fatty acid biosynthesis to terpene biosynthesis. However, this notion is somewhat contradicted by the absence of further seed oil reduction for the terpene yield-improved DDHHFC and DDHHGL lines as compared to their predecessor lines DFC and DGL, respectively. The seed fatty acid composition of each line remained stable throughout all investigated growth conditions (Supplementary Fig. 1), however, as previously observed in Augustin et al., 2015, differed between wild-type and terpene producing plants (Fig. 5). Notably, the altered seed fatty acid profile of terpene producing plants is generally more pronounced for the higher terpene yielding lines, i.e. DGL, DDHHGL, DFC, and DDHHFC, and less pronounced for the low terpene yielding (+)-5-epi-aristolochene producing FE line. This altered seed
Fig. 6. Schematic of 2014 Camelina field trial. Each horizontal line represents a row of Camelina plants of either the transgenic DGL, DFC, or FE line or wild-type (WT) plants. For each row, ten plants (1–10) were further investigated in regards of plant height, total seed yield, weight per seed, seed terpene accumulation, and total seed oil accumulation. Plants 1–3 and 8–10 of each line derived from opposing ends of each plot, whereas plants 4–7 were selected from the respective plot centers. For FAME analysis, seeds of plants 1, 4, and 8 of each plot were investigated. Plants for which at least one replicate was found to contain (+)-δ-cadinene during seed terpene analysis are indicated within circles. The number of DsRed2 expressing seeds found in the harvests of each wild-type plant is shown within diamonds above each corresponding plant. 56
Industrial Crops & Products 136 (2019) 50–58
J.M. Augustin, et al.
during the 2014 field season wild-type plants were grown adjacent to a row of FE plants in the field (Fig. 6). In addition to cross pollination between wild-type Camelina plants and transgenic Camelina plants, hybridization was further indicated to occur between different transgenic lines, as discovered during the seed terpene analysis (Fig. 3). Individual replicates of three DGL plants and two FE plants were found to contain (+)-δ-cadinene in addition to (4S)-limonene and (+)-5-epiaristolochene, respectively. None of the investigated DFC plant samples appeared to contain other terpenes in addition to (+)-δ-cadinene. This may result from dominance of the (+)-δ-cadinene production trait over other terpene production traits, as was observed in attempts to generate transgenic Camelina lines that simultaneously produce both (4S)-limonene and (+)-δ-cadinene. Though not exclusive to (+)-δ-cadinene, these lines seemed to favor the production of (+)-δ-cadinene over that of (4S)-limonene (manuscript in preparation). None of the wild-type plants from the greenhouse study gave rise to DsRed2 expressing seed, and none of the seed samples of transgenic lines from the greenhouse study were found to contain any additional terpenes besides the expected terpene resulting from the intentionally introduced trait. These consistent observations indicate the absence of any cross pollination events under typical greenhouse conditions. This notion is further corroborated by the non-observation of spreading of selection markers or introduced traits from transgenic to non-transgenic Camelina lines or between transgenic lines during years of Camelina greenhouse studies by these authors, involving dozens of incidences of growing various transgenic Camelina lines in close proximity to each other and to wild-type Camelina. Thus, occurrence of Camelina cross pollination is indicated to require environmental factors that are only available during growth under field conditions, e.g., wind and/or insects. For the 2014 field season, the average percent of DsRed2 expressing seeds per wild-type plant was found to be 0.15%, or approximately 3 seeds per 2,000, which is consistent with previously reported results (Walsh et al., 2012). No cross pollination investigations were conducted during the 2015 field season.
views and opinions of authors expressed herein do not necessarily state or reflect those of the United States Government or any agency thereof. We would like to thank the William H. Danforth Fellowship in Plant Sciences awarded to JRB. Appendix A. Supplementary data Supplementary material related to this article can be found, in the online version, at doi:https://doi.org/10.1016/j.indcrop.2019.04.061. References Augustin, J.M., Higashi, Y., Feng, X., Kutchan, T.M., 2015. Production of mono- and sesquiterpenes in Camelina sativa oilseed. Planta 242, 693–708. Bansal, S., Durrett, T.P., 2016. Camelina sativa: an ideal platform for the metabolic engineering and field production of industrial lipids. Biochimie 120, 9–16. Betancor, M.B., Sprague, M., Usher, S., Sayanova, O., Campbell, P.J., Napier, J.A., Tocher, D.R., 2015. A nutritionally-enhanced oil from transgenic Camelina sativa effectively replaces fish oil as a source of eicosapentaenoic acid for fish. Sci. Rep. 5, 8104. Betancor, M.B., Sprague, M., Montero, D., Usher, S., Sayanova, O., Campbell, P.J., Napier, J.A., Caballero, M.J., Izquierdo, M., Tocher, D.R., 2016a. Replacement of marine fish oil with de novo omega-3 oils from transgenic Camelina sativa in feeds for gilthead sea bream (Sparus aurata L.). Lipids 51, 1171–1191. Betancor, M.B., Sprague, M., Sayanova, O., Usher, S., Metochis, C., Campbell, P.J., Napier, J.A., Tocher, D.R., 2016b. Nutritional evaluation of an EPA-DHA oil from transgenic Camelina sativa in feeds for post-smolt Atlantic Salmon (Salmo salar L.). PLoS One 11, e0159934. Botella, C., Sautron, E., Boudiere, L., Michaud, M., Dubots, E., Yamaryo-Botte, Y., Albrieux, C., Marechal, E., Block, M.A., Jouhet, J., 2016. ALA10, a phospholipid flippase, controls FAD2/FAD3 desaturation of phosphatidylcholine in the ER and affects chloroplast lipid composition in Arabidopsis thaliana. Plant Physiol. 170, 1300–1314. Dai, X.X., Weimer, P.J., Dill-McFarland, K.A., Brandao, V.L.N., Suen, G., Faciola, A.P., 2017. Camelina seed supplementation at two dietary fat levels change ruminal bacterial community composition in a dual-flow continuous culture system. Front. Microbiol. 8. Francis, A., Warwick, S.I., 2009. The biology of Canadian weeds. 142. Camelina alyssum (Mill.) Thell.; C. microcarpa andrz. Ex DC.; C. sativa (L.) Crantz. Can. J. Plant Sci. 89, 791–810. Gunstone, F.D., Harwood, J.L., Dijkstra, A.J., 2007. The Lipid Handbook With CD-ROM. CRC Press, Boca Raton. 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. 14, 252–259. Julie-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. Transgen. Res. 23, 67–74. Kagale, S., Koh, C., Nixon, J., Bollina, V., Clarke, W.E., Tuteja, R., Spillane, C., Robinson, S.J., Links, M.G., Clarke, C., Higgins, E.E., Huebert, T., Sharpe, A.G., Parkin, I.A., 2014. The emerging biofuel crop Camelina sativa retains a highly undifferentiated hexaploid genome structure. Nat. Commun. 5, 3706. Kagale, S., Nixon, J., Khedikar, Y., Pasha, A., Provart, N.J., Clarke, W.E., Bollina, V., Robinson, S.J., Coutu, C., Hegedus, D.D., Sharpe, A.G., Parkin, I.A., 2016. The developmental transcriptome atlas of the biofuel crop Camelina sativa. Plant J. 88, 879–894. Kim, H.J., Silva, J.E., Vu, H.S., Mockaitis, K., Nam, J.W., Cahoon, E.B., 2015. 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. Bot. 66, 4251–4265. Kim, J.W., Koo, B., Nyachoti, C.M., 2017. Digestible, metabolizable, and net energy of camelina cake fed to growing pigs and additivity of energy in mixed diets. J. Anim. Sci. 95, 4037–4044. Larsson, M., 2013. Cultivation and processing of Linum usitatissimum and Camelina sativa in southern Scandinavia during the Roman Iron Age. Veg. Hist. Archaeobot. 22, 509–520. Liang, C., Liu, X., Yiu, S.M., Lim, B.L., 2013. De novo assembly and characterization of Camelina sativa transcriptome by paired-end sequencing. BMC Genomics 14, 146. Liu, J., Rice, A., McGlew, K., Shaw, V., Park, H., Clemente, T., Pollard, M., Ohlrogge, J., Durrett, T.P., 2015. Metabolic engineering of oilseed crops to produce high levels of novel acetyl glyceride oils with reduced viscosity, freezing point and calorific value. Plant Biotechnol. J. 13, 858–865. 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. Malik, M.R., Yang, W., Patterson, N., Tang, J., Wellinghoff, R.L., Preuss, M.L., Burkitt, C., Sharma, N., Ji, Y., Jez, J.M., Peoples, O.P., Jaworski, J.G., Cahoon, E.B., Snell, K.D., 2015. Production of high levels of poly-3-hydroxybutyrate in plastids of Camelina sativa seeds. Plant Biotechnol. J. 13, 675–688. Mansour, M.P., Shrestha, P., Belide, S., Petrie, J.R., Nichols, P.D., Singh, S.P., 2014. Characterization of oilseed lipids from "DHA-producing Camelina sativa": a new transformed land plant containing long-chain omega-3 oils. Nutrients 6, 776–789. Moore, R.H., Thornhill, K.L., Weinzierl, B., Sauer, D., D’Ascoli, E., Kim, J., Lichtenstern,
4. Conclusions This study evaluated the performance of previously generated terpene producing Camelina lines over two growing seasons under field conditions and compared them to greenhouse grown plants to evaluate suitability of the transgenic lines as farmable crops. Our results, in conjunction with those found previously (Usher et al., 2015), support phenotypic stability of transgenic Camelina in the field and therefore further validate its use for the production of agricultural and industrial goods. Terpene accumulation under field conditions was, however, consistently found to be slightly lower when compared to greenhousegrown plants. Of the bioengineered Camelina lines investigated in this study, the (+)-δ-cadinene producing DDHHFC line appears most applicable as a terpene-producing crop, since its wild-type equivalent seed yield outweighs the 25% higher terpene accumulation of the highest (4S)-limonene producing line (DDHHGL), whose seed yield is approximately halved compared to wild-type level. The higher energy density of the seed oil of bioengineered terpene producing Camelina lines renders it more suitable for biofuel applications. Concurrently, this change is expected to lower its value as a food oil. Outcrossing of transgenic traits was observed to occur under field conditions, demonstrating that proper precautions should be taken to prevent cross pollination between transgenic and wild-type Camelina plants. However, the outcrossing rate is low, making appropriate containment feasible. Acknowledgements The information, data, or work presented herein was funded in part by the Advanced Research Projects Agency-Energy (ARPA-E), U.S. Department of Energy, under Award Number DE-AR0000202. The 57
Industrial Crops & Products 136 (2019) 50–58
J.M. Augustin, et al.
Camelina species. Plant Breed. 132, 390–396. Sostaric, R., Alegro, A., Hrsak, V., Stancic, Z., Kuster, H., 2009. Plant remains from an Early Iron Age well at Hajndl, Slovenia. Coll. Antropol. 33, 1295–1301. Usher, S., Haslam, R.P., Ruiz-Lopez, N., Sayanova, O., Napier, J.A., 2015. Field trial evaluation of the accumulation of omega-3 long chain polyunsaturated fatty acids in transgenic Camelina sativa: making fish oil substitutes in plants. Metab. Eng. Commun. 2, 93–98. Vollmann, J., Eynck, C., 2015. Camelina as a sustainable oilseed crop: contributions of plant breeding and genetic engineering. Biotechnol. J. 10, 525–535. Walsh, K.D., Puttick, D.M., Hills, M.J., Yang, R.C., Topinka, K.C., Hall, L.M., 2012. Short Communication: first report of outcrossing rates in camelina [Camelina sativa (L.) Crantz], a potential platform for bioindustrial oils. Can. J. Plant Sci. 92, 681–685. Yu, X.H., Cahoon, R.E., Horn, P.J., Shi, H., Prakash, R.R., Cai, Y., Hearney, M., Chapman, K.D., Cahoon, E.B., Schwender, J., Shanklin, J., 2017. Identification of bottlenecks in the accumulation of cyclic fatty acids in camelina seed oil. Plant Biotechnol. J. Zhu, L.H., Krens, F., Smith, M.A., Li, X., Qi, W., van Loo, E.N., Iven, T., Feussner, I., Nazarenus, T.J., Huai, D., Taylor, D.C., Zhou, X.R., Green, A.G., Shockey, J., Klasson, K.T., Mullen, R.T., Huang, B., Dyer, J.M., Cahoon, E.B., 2016. Dedicated industrial oilseed crops as metabolic engineering platforms for sustainable industrial feedstock production. Sci. Rep. 6, 22181.
M., Scheibe, M., Beaton, B., Beyersdorf, A.J., Barrick, J., Bulzan, D., Corr, C.A., Crosbie, E., Jurkat, T., Martin, R., Riddick, D., Shook, M., Slover, G., Voigt, C., White, R., Winstead, E., Yasky, R., Ziemba, L.D., Brown, A., Schlager, H., Anderson, B.E., 2017. Biofuel blending reduces particle emissions from aircraft engines at cruise conditions. Nature 543, 411–415. Moser, B.R., 2012. Biodiesel from alternative oilseed feedstocks: camelina and field pennycres. Biofuels 2012 (3), 17. Nguyen, H.T., Silva, J.E., Podicheti, R., Macrander, J., Yang, W., Nazarenus, T.J., Nam, J.W., Jaworski, J.G., Lu, C., Scheffler, B.E., Mockaitis, K., Cahoon, E.B., 2013. Camelina seed transcriptome: a tool for meal and oil improvement and translational research. Plant Biotechnol. J. 11, 759–769. Petrie, J.R., Shrestha, P., Belide, S., Kennedy, Y., Lester, G., Liu, Q., Divi, U.K., Mulder, R.J., Mansour, M.P., Nichols, P.D., Singh, S.P., 2014. Metabolic engineering Camelina sativa with fish oil-like levels of DHA. PLoS One 9, e85061. Putnam, D.H., Budin, J.T., Field, L.A., BW, M., 1993. Camelina: A Promising Low-Input Oilseed. Wiley, New York. Ruiz-Lopez, N., Haslam, R.P., Napier, J.A., 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. Seguin-Swartz, G., Nettleton, J.A., Sauder, C., Warwick, S.I., Gugel, R.K., 2013. Hybridization between Camelina sativa (L.) Crantz (false flax) and North American
58