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Canola engineered with a microalgal polyketide synthase-like system produces oil enriched in docosahexaenoic acid Terence A Walsh1, Scott A Bevan1, Daniel J Gachotte1, Cory M Larsen1, William A Moskal1, P A Owens Merlo1, Lyudmila V Sidorenko1, Ronnie E Hampton1, Virginia Stoltz1, Dayakar Pareddy1, Geny I Anthony1, Pudota B Bhaskar1, Pradeep R Marri1, Lauren M Clark1, Wei Chen1, Patrick S Adu-Peasah1, Steven T Wensing1, Ross Zirkle2 & James G Metz3,4 Dietary omega-3 long-chain polyunsaturated fatty acids (LC-PUFAs), docosahexaenoic acid (DHA, C22:6) and eicosapentaenoic acid (EPA, C20:5) are usually derived from marine fish. Although production of both EPA and DHA has been engineered into land plants, including Arabidopsis, Camelina sativa and Brassica juncea, neither has been produced in commercially relevant amounts in a widely grown crop. We report expression of a microalgal polyketide synthaselike PUFA synthase system, comprising three multidomain polypeptides and an accessory enzyme, in canola (Brassica napus) seeds. This transgenic enzyme system is expressed in the cytoplasm, and synthesizes DHA and EPA de novo from malonyl-CoA without substantially altering plastidial fatty acid production. Furthermore, there is no significant impact of DHA and EPA production on seed yield in either the greenhouse or the field. Canola oil processed from field-grown grain contains 3.7% DHA and 0.7% EPA, and can provide more than 600 mg of omega-3 LC-PUFAs in a 14 g serving. Fatty acids are usually synthesized in plants and algae by fatty acid synthases. The medium chain saturated fatty acid products of fatty acid synthase are elongated and desaturated (by oxygen-dependent desaturases) to form the diverse array of fatty acids found in these organisms. The omega-3 (n-3) fatty acids DHA and EPA are almost exclusively synthesized by marine microalgae and bacteria, and ascend the food chain to accumulate in marine fish, which are eaten to provide a dietary source of these ‘healthy’ fatty acids for human consumption1. However, some microorganisms can make LC-PUFAs using alternate oxygen-independent enzyme systems named PUFA synthases that have similarities to both the polyketide synthase (PKS) and fatty acid synthase (FAS)2,3 families. PUFA synthases make LC-PUFAs, such as DHA, directly from malonyl-CoA with no free intermediates. The remarkable biosynthetic capacity of some marine microalgae that use PUFA synthases to make oil has been harnessed to produce DHA-enriched oils by fermentation4. Land plants do not
contain the necessary enzymes to make DHA, but the introduction of elongase and desaturase genes (from algae or fungi) has enabled conversion of fatty acids into DHA in the model plant Arabidopsis5,6 and the minor oilseed crops Brassica juncea6 and Camelina sativa7,8. The engineered pathways require an abundant source of α-linolenic acid (ALA, C18:3), which is elongated and desaturated through a series of free intermediate fatty acids to form LC-PUFAs in seed oil. This requires the coordinate expression of up to seven transgenes in seeds and results in plant oils that contain DHA and EPA but also contain substantial quantities of other PUFAs (up to 30% of total oil fatty acids) that can potentially increase oxidative instability of the oil and the production of off-flavors9. Here, we report an alternate approach to provide a sustainable source of DHA that does not rely on sequential modification of native fatty acids (or a high level of precursor ALA) as in previous studies5–8. We introduce a microalgal PKS-like PUFA synthase system into a major oilseed crop plant to provide an orthogonal supplementary biosynthetic capacity for EPA and DHA in planta. Microalgal PUFA synthases usually comprise three large multidomain polypeptides (PFA1, PFA2 and PFA3) that contain all of the enzyme domain modules required to iteratively assemble C2 units from malonyl-CoA into LC-PUFAs, requiring only NADPH as a cofactor2,10, and carrying out more than 50 self-contained sequential enzymatic steps to produce one molecule of product. They are similar to type I or “megasynthase” PKS11 and FAS12 systems. The free fatty acid product is DHA together with varying amounts of docosapentaenoic acid, DPA(n-6) and EPA, depending on the species2,13. The microalgal proteins do not contain plastid-targeting transit peptides, so in our transgenic plant work the polypeptides were localized in the cytoplasm. Both malonyl-CoA and NADPH are abundant metabolites in the plant cytoplasm, and provide the required substrate and cofactors for the enzyme. The PFA1 polypeptide contains multiple tandem acyl-carrier protein (ACP) domains to which growing fatty acid chains are covalently tethered by a phosphopantetheinyl (PP) group. The PP modification is carried out by a dedicated
1Dow
AgroSciences LLC, Research and Development, Indianapolis, Indiana, USA. 2DSM Nutritional Products LLC, Columbia, Maryland, USA. 3DSM Nutritional Products LLC, Boulder, Colorado, USA. 4Present address: Metz Consultancy, Longmont, Colorado, USA. Correspondence should be addressed to T.A.W. (
[email protected]). Received 10 November 2015; accepted 27 April 2016; published online 11 July 2016; doi:10.1038/nbt.3585
nature biotechnology advance online publication
© 2016 Nature America, Inc. All rights reserved.
letters PP transferase enzyme (PPTase) that is required to make PFA1 enzymatically competent. Therefore a functional PUFA synthase system requires only four genes; three PUFA synthase polypeptides and a PPTase. We used the Sfp-like HetI PPTase from the blue-green alga Nostoc sp. PCC7120 (NoHetI)14 as this had previously been shown to function with the microalgal PUFA synthase15. For expression in plants, we tested a variety of seed-specific promoters to drive the genes and enable DHA to be available for assembly into seed triacylglycerols (Supplementary Table 1). We first tested the expression of a PUFA synthase system in plants by Agrobacterium tumefaciens–mediated transformation of the model oilseed plant Arabidopsis thaliana that allows for higher throughput and capacity than crop transformation. Another advantage was that selected vectors used for Arabidopsis transformation experiments could also be directly used for the resource-intensive transformation of canola. Agrobacterium transformation vectors encoding the three PUFA synthase component polypeptides and the NoHetI PPTase plus a selectable marker gene are very large (38–42 kb) and present a considerable challenge for efficient vector construction. We used MultiSite Gateway technology to assemble the large plasmid vectors in a modular fashion, enabling us to test constructs with different configurations. We also tested both native microalgal and canolarecoded versions of the PUFA synthase genes to assess whether plant codon bias conferred an advantage for plant expression16. The size and complexity of these multigenic constructs could pose problems for plant engineering including potential instability of Agrobacterium vectors due to recombination, reduced plant transformation efficiencies and fragmentation of the inserted transfer DNA (T-DNA) within the plant genome17,18. In addition, the choice of multiple regulatory elements, appropriate coding sequences, and their relative position and orientations can have considerable effects on the level of protein and trait expression and on the long-term stability of the trait19. We first tested PUFA synthase expression in plants using genes from the microalga Schizochytrium sp. ATCC 20888 (Schizo20888), which is used for commercial fermentation production of DHA oils2,4. The Schizo20888 PUFA synthase OrfA (8.7 kb), OrfB (6.2 kb) and hybrid OrfC (4.5 kb) genes (equivalent to PFA1, PFA2 and PFA3) and the NoHetI PPTase gene were synthesized using a canola codon bias. The OrfC polypeptide was also modified by replacing the DH2 dehydratase domain with one from Thraustochytrium sp. 23B to increase the proportion of DHA produced20. The genes were arranged in a sequential head-to-tail orientation in the T-DNA and were all expressed using the seed-specific Phaseolus vulgaris Dlec2 promoter21 (construct pDAB7369, Fig. 1a and Supplementary Table 2). We observed DHA levels of up to 0.88% of total fatty acids along with a relatively high proportion of DPA(n-6) (about 34% of new LC-PUFAs) in the resulting T2 seed of transformants (Supplementary Fig. 1). Although this was encouraging, we noted that the number of DHA-producing transgenic plants recovered was comparatively low (only 53% of events) even though all the transgenes were present. The DHA trait was also unstable, because DHA content declined in the subsequent T3 Arabidopsis seed generation, suggesting the presence of a high level of transgene silencing (Fig. 1b). We next used PUFA synthase genes from another marine microalga, Schizochytrium sp. strain ATCC PTA-9695 (Schizo9695), that produces an oil highly enriched in DHA with EPA as the predominant secondary PUFA13. The Schizo9695 PUFA synthase genes share 53–69% amino acid homology with those of Schizo20888 and offered several potential advantages for transgenic expression in plants. The oil produced by this microalga contains both DHA and EPA with low levels of DPA(n-6). Also the PUFA synthase genes are slightly
smaller, particularly PFA1, which contains only six ACP domain units versus nine in Schizo20888 OrfA13. In our multigene construct designs for this gene set, we tested both native microalgal and canola codon-optimized Schizo9695 PUFA synthase gene sequences. We used several different seed-specific promoters to express the genes to assess their relative effect on DHA content. To mitigate potential transgene silencing we also used different regulatory elements for the genes within each construct to minimize the presence of repeated promoter and terminator sequences in the T-DNA. We made constructs with different transcriptional orientations and with DNA spacers containing bidirectional termination sequences between gene cassettes to avoid potential read-through and generation of aberrant RNAs22 (Fig. 1a). We tested 21 multigene constructs in Arabidopsis that incorporated different combinations of these design elements using the Schizo9695 PUFA synthase genes. The relative performance of the constructs was evaluated in Arabidopsis by assessing the proportion of transgenic events that produced DHA, the amount (as % of total fatty acids) of DHA and other LC-PUFAs produced in the hemizygous seed from T1 plants, and the consistency of DHA production in subsequent homozygous seed generations (T2–T5). Using these criteria, the best-performing constructs (such as pDAB107961, Fig. 1a and Supplementary Table 2) contained different regulatory elements for each gene and the presence of one or more native algal coding sequences. These changes increased the proportion of low (one or two) copy number events producing LC-PUFAs to 66–95% (Supplementary Fig. 1), and these events contained up to 1.7% DHA in seeds from homozygous T2 plants (Fig. 1b). We found that the greatest improvement in maintaining DHA trait expression across generations came with the use of native microalgal coding sequences for all three PFA genes, rather than canola codon-optimized genes (compare pDAB101454 and pDAB109525 in Fig. 1b and Supplementary Table 2). This was somewhat surprising, as a common approach to improve expression of heterologous genes in plants is to recode the gene with a host plant codon bias16. In events that expressed a stable DHA trait across generations, the DHA content of Arabidopsis seed from homozygous T2 plants was consistently only slightly higher (by about 33%) than that of the seed from T1 hemizygous plants (Fig. 1b), indicating little or no effect of transgene copy number. This suggests a physiological (or temporal) limit to DHA accumulation by PUFA synthases in Arabidopsis seeds. One significant change in the fatty acid profile of the Arabidopsis seeds besides LC-PUFAs derived from PUFA synthase was a reduction in the amount of native elongated fatty acids from 27% to around 15% (Fig. 1c and Supplementary Table 3). This is consistent with the PUFA synthase system competing for the cytoplasmic pool of malonylCoA, which is also used for elongation of oleic acid to eicosenoic acid (C20:1) in Arabidopsis as each molecule of DHA made by PUFA synthase in the cytoplasm requires 11 malonyl-CoA units. Preliminary experiments expressing PUFA synthase in a fae1 Arabidopsis mutant background that lacks native cytoplasmic fatty acid elongase activity yielded no events with higher LC-PUFA content than in the wild-type background (Supplementary Table 4), suggesting that malonyl-CoA is not limiting for DHA production. We next transformed canola with construct pDAB107960 (Fig. 1a and Supplementary Table 2), which had conferred stable production of DHA in Arabidopsis seeds over four generations. Of 31 one- or two-copy canola events generated, 26 (84%) produced DHA in bulk seed from T0 plants, with an average DHA and EPA content of 1.16% and 0.31%, respectively, and maxima of 3.04% DHA and 1.16% EPA (Fig. 2a). To reduce downstream genetic and analytical complexity, we identified transgenic events that segregated as a single locus. We carried out advance online publication nature biotechnology
letters a
OrfB po
OrfA po
Hetl
OrfC po
SM
b
pDAB7369 7369–009
7369–016
pDAB101454
pDAB107961
pDAB109525
101454–039 101454–068 107961–115 107961–126 109525–020 109525–149
pDAB7369 (29.4 kb) At2S terminator PFA2 po
PFA3 po
CsVMV promoter SM Hetl
PFA2 nat
PFA3 nat
Hetl
AtuORF1 terminator
pDAB101454 (28.0 kb)
DHA (%)
1.5 Pvdlec2 5′ UTR promoter PFA1 po
1.0 0.5 0
SM
T2 Parent T3 Hemi T3 Homo T2 Parent T3 Hemi T3 Homo T2 Parent T3 Hemi T3 Homo T2 Parent T3 Hemi T3 Homo T2 Parent T3 Hemi T3 Homo T2 Parent T3 Hemi T3 Homo T2 Parent T3 Hemi T3 Homo T2 Parent T3 Hemi T3 Homo
PFA1 nat pDAB109525 (28.0 kb)
Hetl
PFA3 nat
SM
PFA2 nat
PvPhas
Atu 2x BnACP BoACP Pvdlec2 At2S BnNapin2 terminator
PFA1 nat
PFA3 nat
Hetl
SM
PFA2 nat
pDAB107960 (30.8 kb) Pvdlec2
PvPhas
At2S PFA1 po
PFA2 po
PFA3 nat
Pv∆Phas Hetl
SM
c Elongated fatty acids (%)
PFA1 nat pDAB107961 (31.6 kb)
30 25 20 15
© 2016 Nature America, Inc. All rights reserved.
pDAB101496 (28.3 kb) 10 Pvdlec2
At2S Pv∆Phas
AtuORF23
0
0.5
1.0
1.5
2.0
2.5
Total LC-PUFA (%)
Figure 1 Expression of PUFA synthase in Arabidopsis. (a) PUFA synthase constructs for plant transformation. 5′ untranslated regions (UTRs) are denoted as dark blue bars. pDAB7369 contains PUFA synthase genes from Schizo20888 whereas the other plasmids contain PUFA synthase genes from Schizo9695. SM indicates the selectable marker gene. (b) Stability of the DHA trait in Arabidopsis. The DHA content of the T2 seed from two representative events (black, T2 parent), and the T3 seed from 5–10 hemizygous (blue) and homozygous (red) progeny (T 3 Hemi and T3 Homo) are shown for each construct. Each symbol represents seed from an individual plant. The progeny of pDAB7369 and pDAB101454 show varying levels of trait silencing whereas the progeny of pDAB107961 and pDAB109525 exhibit trait stability. Constructs pDAB101454 and pDAB109525 are identical except that pDAB101454 contains canola codon-biased PUFA synthase coding sequences, whereas pDAB109525 has native microalgal coding sequences. (c) Elongated fatty acid (EFA) content is inversely correlated with PUFA synthase-derived LC-PUFA production in transgenic Arabidopsis seed. Each point represents the EFA content of the seed from an individual T 1, T2 or T3 plant. The line represents a linear regression model (y = −4.97× + 26.99) R2 = 0.85, n = 585; t-test of regression slope, P < 0.0001.
single-seed fatty acid methyl ester (FAME) analysis of 48 seeds from 17 events producing at least 1% DHA to identify 14 events that contained about 25% null DHA T1 seeds and were therefore likely to have the transgenes at a single locus by Mendelian segregation (Supplementary Fig. 2). Nine of these events were analyzed by quantitative real-time PCR of the transgenes in T1 plants. A selection of plants (up to ten) from each event that were homozygous for the transgenes was grown to maturity (with some hemizygous and null plants). SDS-PAGE and western immunoblot analyses of extracts from the resulting mature T2 seed showed that intact PUFA synthase polypeptides were highly expressed in the canola seed with little evidence of degradation of these large multidomain proteins (Fig. 2b, Supplementary Fig. 3 and Supplementary Table 5). Fatty acid analyses of T2 seed from plants across the nine selected events revealed average DHA contents of 2.87–3.43% (Fig. 2c), with a maximum of 3.87% DHA for an individual plant, and the average EPA content ranged from 0.34 to 0.80%. The PUFA synthase system produced more LC-PUFAs in the crop plant canola than in the model oilseed Arabidopsis, perhaps reflecting the greater headroom in oil biosynthetic capacity of canola. One of the breeder modifications to create modern canola in the 1960’s was to eliminate cytoplasmic elongase activity from traditional oilseed rape that previously directed 45% of fatty acids to erucic acid23. This has left a potentially useful orphaned precursor capacity of malonyl-CoA production in canola24 that we serendipitously exploited for LC-PUFA synthesis by the PUFA synthase system. The fatty acid profile from the transgenic canola seeds (Table 1) had no significant changes other than the addition of LC-PUFAs derived from PUFA synthase (Fig. 2d). These LC-PUFAs are mainly accommodated by a small reduction in the amount of oleic acid from nature biotechnology advance online publication
63 to 57% of total fatty acids. In addition to DHA and EPA there were smaller but detectable amounts of three omega-6 LC-PUFAs: DPA(n-6), arachidonic acid (C20:4, ARA) and γ-linolenic acid (C18:3n-6, GLA), that were present in plants expressing PUFA synthase. These side-products, which are absent from algal oil produced by Schizo9695, might indicate that the PUFA synthase system is less efficient in the plant cell environment compared with microalgal cells. Nevertheless, 75% of PUFA synthase-derived LC-PUFAs were the desired omega-3 LC-PUFAs, DHA and EPA. In the best canola transgenic lines, total PUFA synthase-derived PUFAs reached 6% of total seed fatty acids, showing the potential for the new extraplastidial lipid biosynthesis system to effectively compete with and supplement native fatty acids derived from plastidial lipid biosynthesis in the mature seed oil. We also expressed the PUFA synthase system in soybean by Agrobacterium-mediated transformation to produce up to 2.7% DHA and 1.5% EPA in homozygous T2 seed (data not shown). We anticipate that targeted modification of host plant biosynthetic pathways could further increase the levels of non-native fatty acids accumulated by PUFA synthase systems25. Agency deregulations (e.g., by the US Food and Drug Administration) are required for transgenic crop traits to enter the marketplace, so it is important that the inserted foreign DNA is intact with no extraneous pieces of T-DNA or fragments of the plasmid backbone. This is challenging to achieve with large complex T-DNAs such as the ones that we inserted. To characterize the nature of the 31-kb PUFA synthase T-DNA insertions in the canola genome, we sequenced the genomes of six selected pDAB107960 events. Two events contained one complete copy of the T-DNA and four events contained two adjacent copies in a tandem head-to-tail orientation
letters
GLA
EPA
PFA2 std., 100 ng
PFA3 std., 100 ng
Event-085, plant 16
Event-085, plant 31
DH12075 control
3
4
5
6
7
PFA1
268 238
PFA2
3.0
2.0
PFA3
171
1.0
117
3.0
71 DHA
31
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5.5
6
6.5
7
Ev-353
Ev-111
Ev-107
Ev-352
24:1
min
8.723 - C24:0
8.885 - C24:1
9
8.5
7.5 Elution time (min)
8
8.5
DHA
9.173 - C22:6 DHA
DPA(n-6)
8.885 - C24:1
EPA
8.834 - C22:5 DPA
ARA
8.724 - C24:0
40
7.404 - C18:3 GLA
60
6.900
5.962 - C16:1
80
GLA
7.019 - C18:1 VACC
100
6.782 - C18:0
5.748 - C16:0
120
6.243 - C17:0 SURR
140
8
8.326 - C20.5 EPA
160
7.5
8.075 - C20:4 8.118 - C22:0
180
8.117 - C22:0
7.573 - C20:0
6.778 - C18:0
7
6.5
6
7.872 - C20:2
5.5
6.995 - C18:1
pA
7.690 - C20:1
40
24:0
7.527 - C18:3 ALA
60
7.263 C18:2
5.961 - C16.1
80
16:1
6.241 - C17:0 SURR
5.746- C16.0
100
22:0
20:1
7.688 - C20:1
18:0
140 120
20:0 7.525 - C18:3 ALA
17:0 std.
16:0
7.015 - C18.1 VACC
160
18:3 (ALA)
18:2
7.576 - C20:0
6.991 - C18.1
180
7.249 - C18.2
18:1 (Vac) 18:1
pA
Response
Ev-106
Event
d © 2016 Nature America, Inc. All rights reserved.
Ev-100
41 0
Ev-085
0
55
1.0
Ev-702
2.0
Ev-646
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DPA(n-6) 4.0
2
4.0
460
ARA
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1
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LC-PUFAs (%)
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c
PFA1 std., 50 ng
b MW markers
a
9
min
Figure 2 Expression of PUFA synthase in canola. (a) LC-PUFA content of canola T1 seed from 31 T0 events transformed with pDAB107960. Each bar represents the PUFA synthase-derived LC-PUFA content of seed from an individual plant (a T 0 event). The events are sorted by DHA content. Five events produced no LC-PUFAs. Untransformed control plants contained no detectable LC-PUFAs. (b) SDS-PAGE of transgenic PUFA synthase-expressing T 2 canola seed extract. 20 µg of protein extracted from transgenic or control mature canola seeds were run on a 3–8% Tris-acetate SDS-PAGE gel and stained with Coomassie blue. The three large PUFA synthase polypeptides are readily visible in the upper portion of the gel. Purified individual PUFA synthase protein standards are in lanes to the left for reference. (c) LC-PUFA content of homozygous T2 transgenic canola seeds from nine transgenic events. The PUFA synthase-derived fatty acids are similar across nine independent homozygous single-locus events. LC-PUFAs are color-coded as in a. (d) GC-FAMEs profiles of T2 transgenic seed and DH12075 control. (Upper) Untransformed DH12075 canola seed. (Lower) Homozygous PUFA synthase-expressing T2 canola seed. Other than the PUFA synthase-derived fatty acids, there are no major changes in the fatty acid profiles of the transgenic canola seed and the untransformed parent seed.
(data not shown). Four of the events had no additional fragments derived from the T-DNA in the crop genome other than the desired transgenes, demonstrating that precise single-locus introduction of intact large T-DNAs encoding an entire biosynthetic system can be achieved in canola. The genomic location of single-copy event-085 is shown in Figure 3a.
For successful introgression into elite backgrounds for canola breeding, the PUFA synthase-based DHA trait needs to be stable in transgenic events for multiple generations. We selected lines from six pDAB107960 events and grew them in the greenhouse, selfing them to the T5 seed generation. In lines from four of these events, the DHA trait remained consistent across generations whereas lines advance online publication nature biotechnology
letters Table 1 Fatty acid composition (wt% of total fatty acids) of transgenic PUFA synthase-expressing canola T3 seeds from homozygous T2 plants No. of plants C16:0 C18:0 Event-085 Event-106 Event-107 Event-111 Event-352 Event-353 All events Null plants
8 8 10 10 5 5 46 7
3.63 3.82 3.75 3.65 4.79 3.99 3.86 4.03
3.53 3.17 3.28 3.37 2.67 3.12 3.24 3.27
C18:1
C18:1 Vacc.
C18:2
GLA 18:3
61.29 57.48 59.88 58.02 51.14 58.98 58.26 63.29
2.42 2.41 2.24 2.72 2.97 2.57 2.52 2.29
12.78 14.50 13.68 13.95 19.39 14.28 14.41 14.17
0.34 8.56 0.38 9.81 0.44 8.55 0.44 9.76 0.35 11.15 0.30 8.89 0.38 9.35 0 9.42
C18:3
ARA C20:0 C20:1 C20:4 1.09 1.08 1.16 1.08 0.74 0.83 1.03 0.96
1.14 1.19 1.13 1.07 0.99 0.99 1.10 1.32
EPA DPA C20:5 C24:0 C22:5
0.24 0.36 0.28 0.27 0.27 0.30 0.29 0
0.37 0.79 0.47 0.36 0.34 0.48 0.47 0
0.23 0.24 0.25 0.18 0.19 0.15 0.21 0.30
0.75 0.62 0.77 0.94 0.82 0.73 0.78 0
DHA C22:6
Minor DHA + Total FAs EPA LC-PUFAs
2.87 3.25 3.30 3.43 3.28 3.59 3.27 0
0.66 0.75 0.72 0.65 0.79 0.70 0.71 0.79
3.23 4.04 3.77 3.79 3.62 4.06 3.74 0
4.55 5.41 5.25 5.44 5.06 5.40 5.19 0
from two events showed significant decreases in later generations (P < 0.0001 for events 107 and 11; Fig. 3b). Thus the majority of events tested exhibited trait stability. To establish whether installation of the LC-PUFA biosynthetic capacity into canola could also function well in the field, seed from homozygous T2 plants from the six selected events was grown at two locations in the northern United States in 2014, and the harvested grain was analyzed for LC-PUFA content. The LC-PUFA content of the transgenic events was equivalent and in some cases higher than that observed for the same lines grown in the greenhouse (Fig. 3c and Supplementary Fig. 4). There was no difference in agronomic characteristics of the transgenic DHA-producing plants
N5 6
N
N7 N8
N9 N10
EPA
3.0
DHA
1.0
21N
b
Ev-085
Ev-106
Ev-352
Ev-353
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Field
DH12075 control
GH
Event 106
Field
N1
GH
N15
Field
4
N16
GH
N17
GH
8
Field
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0
N1
9
N1
Location of T-DNA insertion
4.0
2.0
11N
Inserted T-DNA region of plasmid
GLA ARA DPA(n-6)
Location Ev-107
Ev-111
e Grain yield (kg/ha)
N4
5.0
GH
N3
Ev-111 Ev-107 Ev-352 Ev-106 Ev-085 Ev-1353 6.0
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N2 N2
Reads coverage
d
c
LC-PUFA (%)
Canola chromosomes N1–N19
Backbone region
in comparison to the DH12075 non-transgenic controls (Fig. 3d). There was also no significant detrimental effect on yield, total seed oil content or protein content of the grain from transgenic plants relative to controls (Fig. 3e and Supplementary Table 6). The final product from canola is a refined, bleached, deodorized (RBD) oil that can be directly marketed as a bottled oil or used as a food ingredient, for example, in salad dressings. LC-PUFAs are intrinsically susceptible to oxidative degradation9, and we wanted to ensure that the PUFA-synthase-derived DHA was both stable in the canola grain for storage purposes and that DHA-containing grain could be processed to yield an RBD oil using conventional procedures and
Field
a
N1
1,500
nature biotechnology advance online publication
Ev-353
Ev-352
Ev-111
Ev-107
Ev-106
Ev-085
DH12075
Figure 3 Characterization of 4.0 DHA-producing canola events. 1,000 (a) Genomic location of T-DNA 3.0 in a PUFA synthase canola event. Circos plot of a canola 500 DHA event (Event-085) with 2.0 a complete T-DNA integration 0 on chromosome N18. The 1.0 green arc represents the 19 canola chromosomes. The red Event arc represents the plasmid 0 T2 T3 T4 T5 T2 T3 T4 T5 T2 T3 T4 T5 T2 T3 T4 T5 T2 T3 T4 T5 T2 T3 T4 T5 sequence (the scale is expanded Seed generation relative to the chromosomes). The blue histogram above the T-DNA indicates the sequence coverage across the plasmid. The lack of reads for backbone indicates that the backbone is absent in the transgenic plant. The red lines connecting the ends of the transgene to N18 indicate those paired-end reads with one read mapped to the transgene and its pair mapped to the chromosome. (b) The PUFA synthase DHA trait is stable in canola over five seed generations. The DHA content of T2 through T5 seed remained relatively consistent across greenhouse-grown selfed generations for events 085, 106, 352 and 353, whereas it declined significantly (t-test of regression slope, P < 0.0001) with an increasing spread in the data in T4 and T5 seed for lines from events 107 and 111. The box plots show 25–50% and 50–75% quartiles (n = 5–30), and the whiskers extend to 1.5× the interquartile region. Outliers are shown as circles. The mean DHA content is shown by the wider bar. (c) Transgenic PUFA synthase canola plants in the field. Transgenic PUFA synthase-expressing plants producing DHA in the seed were phenotypically normal and robust relative to untransformed control plants (DH12075). Scale bars, 0.5 m. (d) LC-PUFA content of T4 grain from T3 plants grown in the field and greenhouse (GH). The PUFA synthase-derived fatty acids were similar across sites with the field-grown plants producing equivalent or slightly more DHA and less other PUFAs. (e) Grain yield of transgenic canola plants. There is no significant decrease in yield across transgenic events grown in the field compared to the untransformed DH12075 control plots. The box plots show 25–50% and 50–75% quartiles (n = 7–8) and the whiskers extend to 1.5× the interquartile region. Outliers are shown as circles. Mean grain yield is shown by the wider bar. Average DHA content (%)
© 2016 Nature America, Inc. All rights reserved.
Each value is the average of bulk seed samples from five to ten sibling homozygous T2 transgenic plants from six different canola events transformed with pDAB107960 compared to seven transgene null plants. Minor fatty acids make up the sum of C14:0, C16:1, C20:2, C22:0 and C22:1.
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letters without compromising DHA content or oil quality. A series of T2 lines from a canola event derived from construct pDAB101496 (Fig. 1a and Supplementary Table 2) was grown in the field and produced grain with up to 4.3% DHA and 0.6% EPA. Analysis of this fieldgrown T3 grain stored at room temperature for up to 9 months showed minimal losses (30%) seen with the elongase/desaturase transgene approach5–8 can also contribute to oxidation that can result in off-flavors9. The DHA and EPA content of canola seeds produced by the PUFA synthase system are lower than those reported using the elongase/desaturase approach in small-scale oilseed systems7,8 and more recently in canola32 (9–12% DHA from plants with low transgene copy numbers). However, the levels we report of up to 4.4% DHA + EPA in field-produced canola oil are commercially relevant, especially as we have demonstrated that
the PUFA synthase system is robust across the complete production cycle from introduction into a mainstream oilseed crop through field cultivation, grain storage and processing to a finished oil. The resulting fatty acid profile of the DHA-enriched canola oil is similar to an oil blend recently shown to lower predicted cardiovascular risk 33. Current typical dietary recommendations for consumption of omega3 LC-PUFAs are for 250–500 mg per day30 so one serving size (14 g) of the canola oil that we have produced (e.g., in a salad dressing), could provide over 600 mg of these healthful omega-3 fatty acids. Methods Methods and any associated references are available in the online version of the paper. Accession codes. The sequence reads from the six events characterized in this paper have been deposited in the SRA and can be found under NCBI BioProject PRJNA318522. Note: Any Supplementary Information and Source Data files are available in the online version of the paper. Acknowledgments We thank P. Roessler for early inspiration for this project. At Dow AgroSciences, we thank B. Martindale and L. Juberg for greenhouse support, M. Landes, A. Walker, B. Case, C. Ransom, R. Preuss, K. Ubayasena, S. Chennareddy, G. Booher and P. Nelson for technical assistance, M. Foster and A. Beach for providing recombinant protein standards, P. Graupner for triacylgycerol positional analysis and A. Wang for statistical analyses. The fae1/fad3 Arabidopsis mutant was obtained from M. Smith (NRC Canada, Saskatoon, Canada). AUTHOR CONTRIBUTIONS J.G.M. and R.Z. selected PUFA synthase genes, T.A.W., S.A.B., D.J.G., C.M.L., P.A.O.M. and L.V.S. were responsible for construct design strategy, plant transformation experiments and analyzed results. S.A.B. made the constructs. D.J.G., W.A.M., R.E.H. and V.S. performed technical analyses. D.P. and G.I.A. were responsible for canola transformation experiments. P.B.B. designed and analyzed field trials. P.R.M., L.M.C. and W.C. designed and conducted the canola whole genome sequencing experiments. P.S.A.-P. and S.T.W. designed and conducted oil processing and analyses. T.A.W. wrote the manuscript. All authors discussed results and reviewed the manuscript. COMPETING FINANCIAL INTERESTS The authors declare competing financial interests: details are available in the online version of the paper. Reprints and permissions information is available online at http://www.nature.com/ reprints/index.html. 1. Abedi, E. & Sahari, M.A. Long-chain polyunsaturated fatty acid sources and evaluation of their nutritional and functional properties. Food Sci. Nutr. 2, 443–463 (2014). 2. Metz, J.G. et al. Production of polyunsaturated fatty acids by polyketide synthases in both prokaryotes and eukaryotes. Science 293, 290–293 (2001). 3. Shulse, C.N. & Allen, E.E. Widespread occurrence of secondary lipid biosynthesis potential in microbial lineages. PLoS One 6, e20146 (2011). 4. Barclay, W., Weaver, C. & Metz, J.G. in Single Cell Oils (eds. Z. Cohen & C. Ratledge) (AOCS Press, Urbana, IL, 2005). 5. Petrie, J.R. et al. Metabolic engineering plant seeds with fish oil-like levels of DHA. PLoS One 7, e49165 (2012). 6. Wu, G. et al. Stepwise engineering to produce high yields of very long-chain polyunsaturated fatty acids in plants. Nat. Biotechnol. 23, 1013–1017 (2005). 7. Ruiz-Lopez, N., Haslam, R.P., Napier, J.A. & Sayanova, O. Successful high-level accumulation of fish oil omega-3 long-chain polyunsaturated fatty acids in a transgenic oilseed crop. Plant J. 77, 198–208 (2014). 8. Petrie, J.R. et al. Metabolic engineering Camelina sativa with fish oil-like levels of DHA. PLoS One 9, e85061 (2014). 9. Frankel, E.N. Lipid Oxidation 2nd edn. (The Oily Press, Bridgwater, UK, 2005). 10. Metz, J.G. et al. Biochemical characterization of polyunsaturated fatty acid synthesis in Schizochytrium: release of the products as free fatty acids. Plant Physiol. Biochem. 47, 472–478 (2009). 11. Staunton, J. & Weissman, K.J. Polyketide biosynthesis: a millennium review. Nat. Prod. Rep. 18, 380–416 (2001).
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letters 12. Leibundgut, M., Maier, T., Jenni, S. & Ban, N. The multienzyme architecture of eukaryotic fatty acid synthases. Curr. Opin. Struct. Biol. 18, 714–725 (2008). 13. Apt, K.E., Richter, L., Simpson, D. & Zirkle, R. Polyunsaturated fatty acid synthase nucleic acid molecules and polypeptides, compositions, and methods of making and uses thereof. US patent application 20100266564A1 (2010). 14. Copp, J.N. & Neilan, B.A. The phosphopantetheinyl transferase superfamily: phylogenetic analysis and functional implications in Cyanobacteria. Appl. Environ. Microbiol. 72, 2298–2305 (2006). 15. Hauvermale, A. et al. Fatty acid production in Schizochytrium sp.: Involvement of a polyunsaturated fatty acid synthase and a type I fatty acid synthase. Lipids 41, 739–747 (2006). 16. Ullrich, K.K., Hiss, M. & Rensing, S.A. Means to optimize protein expression in transgenic plants. Curr. Opin. Biotechnol. 32, 61–67 (2015). 17. Peremarti, A. et al. Promoter diversity in multigene transformation. Plant Mol. Biol. 73, 363–378 (2010). 18. Naqvi, S. et al. When more is better: multigene engineering in plants. Trends Plant Sci. 15, 48–56 (2010). 19. Dietz-Pfeilstetter, A. Stability of transgene expression as a challenge for genetic engineering. Plant Sci. 179, 164–167 (2010). 20. Weaver, C.A., Zirkle, R., Doherty, D.H. & G, M.J. Chimeric PUFA polyketide synthase systems and uses thereof. US patent application 8309796B2 (2012). 21. Hoffman, L.M. & Donaldson, D.D. Characterization of two Phaseolus vulgaris phytohemagglutinin genes closely linked on the chromosome. EMBO J. 4, 883–889 (1985). 22. Gudynaite-Savitch, L., Johnson, D.A. & Miki, B.L.A. Strategies to mitigate transgenepromoter interactions. Plant Biotechnol. J. 7, 472–485 (2009).
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23. Fourmann, M. et al. The two genes homologous to Arabidopsis FAE1 co-segregate with the two loci governing erucic acid content in Brassica napus. Theor. Appl. Genet. 96, 852–858 (1998). 24. Schwender, J. et al. Quantitative multilevel analysis of central metabolism in developing oilseeds of oilseed rape during in vitro culture. Plant Physiol. 168, 828–848 (2015). 25. Napier, J.A., Haslam, R.P., Beaudoin, F. & Cahoon, E.B. Understanding and manipulating plant lipid composition: Metabolic engineering leads the way. Curr. Opin. Plant Biol. 19, 68–75 (2014). 26. Lau, W. & Sattely, E.S. Six enzymes from mayapple that complete the biosynthetic pathway to the etoposide aglycone. Science 349, 1224–1228 (2015). 27. Yalpani, N., Altier, D.J., Barbour, E., Cigan, A.L. & Scelonge, C.J. Production of 6-methylsalicylic acid by expression of a fungal polyketide synthase activates disease resistance in tobacco. Plant Cell 13, 1401–1409 (2001). 28. Hertweck, C. The biosynthetic logic of polyketide diversity. Angew. Chem. Int. Ed. Engl. 48, 4688–4716 (2009). 29. Weissman, K.J. & Leadlay, P.F. Combinatorial biosynthesis of reduced polyketides. Nat. Rev. Microbiol. 3, 925–936 (2005). 30. Salem, N. Jr. & Eggersdorfer, M. Is the world supply of omega-3 fatty acids adequate for optimal human nutrition? Curr. Opin. Clin. Nutr. Metab. Care 18, 147–154 (2015). 31. Xue, Z. et al. Production of omega-3 eicosapentaenoic acid by metabolic engineering of Yarrowia lipolytica. Nat. Biotechnol. 31, 734–740 (2013). 32. Petrie, J.R. & Singh, S.P. Lipid comprising docosapentaenoic acid. US patent application 20150374654A1 (2015). 33. Jones, P.J. et al. DHA-enriched high-oleic acid canola oil improves lipid profile and lowers predicted cardiovascular disease risk in the canola oil multicenter randomized controlled trial. Am. J. Clin. Nutr. 100, 88–97 (2014).
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ONLINE METHODS
PUFA synthase plasmids for plant transformation. Canola codonoptimized OrfA, OrfB and OrfC genes from Schizo20888 and the NoHetI gene were synthesized by GeneArt AT (Regensburg, Germany; acquired by Life Technologies, Carlsbad, CA). OrfC was modified by replacing the 1.55 kb DH2 dehydratase domain with one from a Thraustochytrium sp. 23B (ATCC 20892) microalga that reduces the proportion of DPA(n-6) produced. Canola codon-optimized PFA1, PFA2 and PFA3 genes from Schizo9695 were synthesized by DNA2.0 (Menlo Park, CA). The canola codon-optimized genes were mobilized into Gateway entry vectors using conventional cloning techniques. As the native microalgal PFA1, PFA2 and PFA3 genes13 contained internal restriction sites that interfered with conventional cloning, they were amplified by PCR and mobilized into entry vectors using In-Fusion reactions (Clontech Laboratories, Mountain View, CA). Plant transformation plasmids were assembled using MultiSite Gateway Pro LR technology (Life Technologies, Carlsbad, CA). The regulatory elements used in plasmid construction are described in Supplementary Table 1 (refs. 21,34–40). Single promoter-gene-terminator cassettes were placed into Gateway entry vectors. A plant selectable marker gene driven by the cassava vein mosaic virus (CsVMV) promoter and A. tumefaciens ORF1 terminator, and sequences required for Agrobacterium-mediated plant transformation were placed on a Gateway destination vector, and the five vectors were recombined through a Gateway Pro LR reaction. Initial entry vectors for the PUFA synthase and NoHetI genes were synthesized by Blue Heron (Bothell, WA) to contain a promoter and terminator flanked by Gateway att sites. Unique restriction sites were included in the synthetic vectors to facilitate exchanging promoter, terminator and att elements by conventional cloning techniques. Constructs are illustrated in Figure 1 and described in Supplementary Table 2. Plant transformation. Transgenic Arabidopsis events were generated using Agrobacterium-mediated floral dip transformation41 of wild-type Col-0 plants. Some experiments were also performed with a fae1/fad3 Arabidopsis mutant42 obtained from M. Smith (NRC Canada, Saskatoon, Canada). RecA− Agrobacterium strains were used to minimize recombination within large T-DNAs. Transgenic T1 plants were selected by spraying with appropriate herbicide at 280 g/ha seven and 9 days after planting. Surviving resistant plants were then transplanted to individual pots and grown in a greenhouse (16-h day, 20 °C). After copy number analysis, one- to two-copy T 1 events were grown to maturity and the T2 seed from individual plants harvested and cleaned of plant debris for analysis. Canola transformation experiments were performed by Agrobacteriummediated transformation of hypocotyl segments. Binary plasmid vectors were introduced into A. tumefaciens strain, Z707S RecA− (ref. 43) by electroporation and used to transform hypocotyl segments from 5 to 6-day old seedlings of canola line DH12075 (ref. 44). Regeneration of herbicide-resistant T0 plants was performed according to a method adapted from De Block et al.45. T0 plants were bagged before flowering, grown to maturity in the greenhouse and the resulting T1 seed harvested for analysis. Transgene copy number determination. Genomic DNA was isolated from young Arabidopsis or canola leaf tissue using a BioSprint 96 magnetic particle automation platform and the BioSprint 96 DNA Plant Kit (Qiagen, Valencia, CA) following the manufacturer’s protocol. Extracted genomic DNA was diluted 1:5 with water before use as a template in quantitative real time PCR reactions. Real-time qPCR assays for copy number detection were designed against PUFA synthase coding sequences using the Roche Assay Design Center (http://www. universalprobelibrary.com). These target assays used hydrolysis probes from the Roche Universal Probe library (Roche, Indianapolis, IN). For the native version of PFA1, a custom hydrolysis probe assay was designed using VisualOMP (DNA Software, Ann Arbor, MI). This assay, as well as reference assays for the HMG1 gene from canola46, or the TAFFII15 gene of Arabidopsis (AT4G31720) were synthesized by Integrated DNA Technologies (Coralville, IA). Sequences for target and reference assays are in Supplementary Table 10. Real-time PCR reactions were run on a Roche LightCycler480 II using standard protocols. A comparative Ct method was used to calculate Ct value target-to-reference ratios. Zygosity and copy number estimates were
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generated by normalizing to equivalent data from events containing transgenes with verified copy number. Fatty acid analysis. Seed samples (5 to 10 mg) were homogenized by milling with a steel ball in 1.2 ml heptane containing 33 p.p.m. triheptadecanoin (Nu-Chek Prep, Elysian, MN) as a triacylglycerol internal standard using a Geno/Grinder (Spex SamplePrep, Metuchen, NJ) at 40 °C with constant shaking. Prior to homogenization, 0.2 ml of freshly prepared 0.25 M sodium methoxide (Sigma-Aldrich, St. Louis, MO) in methanol was added. The extract was diluted to a level suitable for GC analysis to bring the internal standard concentration to 3.3 p.p.m. Heptane extraction of fatty acid methyl esters (FAMEs) was repeated three times and the heptane layers were pooled before analysis. Fatty acid recoveries were normalized to recovery of methylated heptadecanoic acid. Resulting FAMEs were analyzed by Agilent 6890 GC-FID using a capillary column BPX 70 (15 m × 0.25 mm × 0.25 µm) from SGE Analytical Science (Austin, TX). Each FAME was identified by retention time and quantified by the injection of a rapeseed oil reference mix from Matreya LLC (Pleasant Gap, PA) as a calibration standard containing additional LC-PUFA standards of DHA, EPA, γ-linolenic acid, arachidonic acid and DPA (n-6) methyl esters (Nu-Chek Prep). Peak identification of novel PUFA synthase-derived LC-PUFAs in transgenic plant samples was confirmed using an Agilent 6890 GC equipped with an Agilent EI/CI-MSD 6975c mass spectrometer with the same separation conditions described above. Mass spectral fragmentations were compared to a library of pure standards under chemical mediated ionization with methane to increase parent ion intensity. Immunoanalysis of transgenic proteins. Quantitative western blot methods were developed to detect each PUFA synthase polypeptide and NoHetI in seed samples. Antigens for full length PFA1 and PFA3 were recombinantly expressed in E. coli ArcticExpress (DE3)RIL cells (Agilent Technologies, Santa Clara, CA) with an N-terminal hexahistidine tag and purified via cobalt affinity chromatography. Full length PFA2 was isolated directly from inclusion bodies. An N-terminal PFA2 fragment (amino acid residues 1-204) and a PFA3 fragment (residues 1114-1281) that overlaps with the predicted enoyl reductase domain were also expressed in E. coli as antigens. All were submitted as SDS-PAGE gel slices for polyclonal production in rabbits (Covance, Denver, PA and SDIX, Newark, DE). Antigen for full-length NoHetI PPTase with an N-terminal hexahistidine tag was overexpressed in E. coli Bl21(DE3) cells and purified via cobalt affinity chromatography. Antigen was submitted as soluble protein at about 2 mg/mL in Tris-buffered saline for polyclonal production in rabbits (Covance, Denver, PA). All antisera were purified by Protein G affinity chromatography. Recombinant protein reference standards of PFA1, PFA2, PFA3, and HetI for gel quantifications were also produced in E. coli ArcticExpress (DE3)RIL cells and purified via CoMAC His-tagged purification. Protein concentrations were determined by densitometry using a bovine serum albumin standard curve for in-gel quantitation with SDS-PAGE and Coomassie blue staining. NoHetI standard was also quantified by amino acid analysis. Canola seed extracts were prepared for quantitative protein analysis from delipidated cakes after hexane extraction for FAMEs analysis. Extraction buffer (50 mM Tris pH 8.5, 10 mM EDTA, 2% SDS) was added (1 ml per 50 mg bulk seed) and sample tubes were rocked gently for 15–30 min, centrifuged for 30 min at 3,000g and the supernatant collected. Total soluble protein in the extract was determined by Pierce 660 nm Protein Assay (Thermo Scientific, Rockford, IL). Samples were normalized to 1.55 mg/ml total soluble protein in extraction buffer before dilution into lithium dodecyl sulfate sample buffer (Invitrogen, Carlsbad, CA) containing 40 mM dithiothreitol. Twenty µg total soluble protein was loaded per lane and electrophoresed in 3–8% Tris acetate gels (Invitrogen, Carlsbad, CA) for PFA1, PFA2, and PFA3 or 4–12% Bis Tris gels (Invitrogen, Carlsbad, CA) for HetI then transferred to nitrocellulose or PVDF membranes. Blots were blocked in 1% skim milk in phosphate-buffered saline with Tween-20 and probed with antibodies against the PUFA synthase PFA1, PFA2 and PFA3 polypeptides and NoHetI using an anti-rabbit fluorescentlabeled secondary antibody (Goat Anti-Rabbit AF 633 (Invitrogen, Carlsbad, CA)) for detection. Blots were visualized on a Typhoon Trio Plus fluorescent
doi:10.1038/nbt.3585
imager (GE Healthcare, New Brunswick NJ) and proteins quantitated by calibration with reference standards run simultaneously.
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Whole genome sequencing of transgenic canola plants. DNA was isolated using a modified cetyltrimethylammonium bromide extraction method47 for whole genome sequencing (WGS). WGS genomic DNA libraries (800 bp) were made following Illumina’s Truseq DNA Sample Prep v2 Low Throughput Protocol (Illumina Inc., San Diego, CA) and were sequenced with paired-end chemistry according to Illumina’s SBS HiSeq protocol. Initial quality control of the sequenced reads was done using the CASAVA software. The reads were then trimmed for adaptor sequences and Q30. The trimmed reads were mapped to the canola DH12075 and pDAB107960 reference T-DNA sequences using BWA48. Sequence coverage for genome and transgenes was obtained using BEDTools49 to detect the integrity of the transgene. Diagrams illustrating the transgene integrations in the genome were made using Circos50. Accession code NCBI BioProject PRJNA318522. Field evaluation of transgenic DHA-producing canola events. Field testing was performed for several DHA-producing canola events derived from constructs pDAB101496 and pDAB107960 in Minnesota and North Dakota. In 2013, pooled T2 seed from several homozygous T1 plants from six pDAB101496 events producing moderate to high levels of DHA (2 to 3.5% DHA) were used for field evaluations at two locations. In 2014, pooled T3 seed from several homozygous T2 plants derived from six pDAB107960 events producing moderate to high levels of DHA (2 to 4% DHA) were used at two locations. Untransformed DH12075 canola plants were used as controls and commercial varieties were used as performance checks. A Randomized Complete Block design was used with every entry replicated four times at each location. Plots were 6 m long and 1.5 m wide with six rows in each plot. Seeding rates ranged from 7 to 11 plants/ft2 reflecting local planting practices. Standard spring canola cultivation and management practices were followed throughout the growing season. Seed from each plot was harvested at maturity and analyzed for yield, LC-PUFA content and other seed quality characteristics. The LC-PUFA content of the harvested canola grain from each experimental plot was determined from FAME extractions of three 10-seed aliquots sampled from the bulk grain from each field plot. The total oil content of grain samples was determined with a Bruker mq10 MiniSpec NMR Analyzer with a PA102 probe using AOCS Recommended Practice Ak 5-01 (ref. 51) and also by quantitative extraction of total FAMEs. Triacylgycerol positional analysis was performed by band-selective heteronuclear single quantum coherence (HSQC) 13C NMR spectroscopy using a 600 MHz Bruker Avance III NMR spectrometer operating at 600.13 MHz and equipped with a 5 mm inverse RT probe. Samples were dissolved in 0.6 ml CDCl3 and spectra acquired at 25 °C. Selective pulses were used in the carbon dimension to excite a six p.p.m. band centered at 34 p.p.m., to give signals from the carbons attached to the acyl end of the triacylglyceride. Authentic triacylglycerides with DHA at the sn-2 or sn-1/3 positions (Larodan Fine Chemicals, Malmo, Sweden) were used for validation. DHA stability in stored grain. Canola grain was stored at ambient temperatures (22 ± 1 °C) for 12 months. A sample was taken for oil analysis each month. Oil was prepared by grinding the seeds, extracting with multiple batches of hexane in an explosion-proof Waring blender and vacuumfiltering the resulting slurry through a Buchner funnel with Whatman #4 filter paper. The filtered hexane/oil mixture was rotary-evaporated to recover the oil fraction and analyzed for FAME content, free fatty acids, peroxide value, and P-anisidine value. Refined, bleached and deodorized (RBD) oil. Lab scale processes that simulate commercial scale equipment were used to extract and refine canola oil for characterization and stability testing. In brief, grains were heated to 80 °C for 20 min and mechanically pressed to extract approximately two-thirds of the oil. Residual oil in the presscakes was then extracted with hexane and the solvent stripped from the oil. The solvent-extracted oil was then combined with the oil from mechanical pressing. The total crude oil was then sequentially treated with dilute phosphoric acid to remove phospholipids, caustic to remove free fatty acids as soaps, and bleaching clay to remove chlorophyll
doi:10.1038/nbt.3585
and other pigments. The oil was steam-stripped at 210 °C for 60 min under
