Methods of Producing Insect Pheromones

Abstract

The present disclosure relates to methods of producing insect pheromone precursors and genetically modified plants capable of producing insect pheromone precursors. The genetically modified plants include a heterologous gene encoding at least one silencing suppressor protein and at least one enzyme selected from the group consisting of a fatty acyl desaturase, a fatty acyl elongase, a fatty acyl reductase, and an acyl-CoA oxidase.

Claims

1. A genetically modified plant having incorporated into the genome a heterologous gene encoding at least one silencing suppressor protein and at least one enzyme selected from the group consisting of a fatty acyl desaturase, a fatty acyl elongase, a fatty acyl reductase, and an acyl-CoA oxidase, wherein the plant produces at least one insect pheromone precursor.

2. The genetically modified plant of claim 1, wherein the at least one silencing suppressor protein is P19.

3. The genetically modified plant of claim 1, wherein the heterologous gene includes multiple copies of the same enzyme.

4. The genetically modified plant of claim 3, wherein the heterologous gene includes three copies of the same enzyme.

5. The genetically modified plant of claim 1, wherein the at least one enzyme is a fatty acyl desaturase.

6. The genetically modified plant of claim 5, wherein the fatty acyl desaturase is CpCPRQ.

7. The genetically modified plant of claim 1, wherein the plant is Camelina sativa.

8. The genetically modified plant of claim 1, wherein the at least one insect pheromone precursor is E8,E10-12:acid.

9. The genetically modified plant of claim 1, wherein the heterologous gene is expressed in seeds of the plant.

10. The genetically modified plant of claim 1, wherein the heterologous gene further includes at least one promoter.

11. The genetically modified plant of claim 10, wherein the promoter is napin.

12. A method of producing insect pheromone precursors, said method comprising: a) incorporating into the genome of a genetically modified plant a heterologous gene encoding at least one silencing suppressor protein and at least one enzyme selected from the group consisting of a fatty acyl desaturase, a fatty acyl elongase, a fatty acyl reductase, and an acyl-CoA oxidase; b) breed the genetically modified plant; and c) extract the insect pheromone precursors from the genetically modified plant.

13. The method of claim 12, wherein the at least one silencing suppressor protein is P19.

14. The method of claim 12, wherein the heterologous gene is expressed in seeds of the genetically modified plant.

15. The method of claim 12, wherein the genetically modified plant is Camelina sativa.

16. The method of claim 12, wherein the heterologous gene includes multiple copies of the same enzyme.

17. The method of claim 12, wherein the at least one enzyme is CpCPRQ.

18. The method of claim 12, wherein the heterologous gene further includes at least one promoter.

19. The method of claim 18, wherein the at least one promoter is napin.

20. A method of disrupting mating of a crop pest, said method comprising: a) incorporating into the genome of a genetically modified plant a heterologous gene encoding a crop pest pheromone attractant, wherein the genetically modified plant releases the crop pest pheromone attractant; and b) planting the genetically modified plant near a crop to be protected from the crop pest

Description

BRIEF DESCRIPTION OF THE DRAWINGS

[0059] These and other features and advantages of the various embodiments disclosed herein will be better understood with respect to the following description and drawings, in which like numbers refer to like parts throughout, and in which:

[0060] FIG. 1 depicts lab results of a genetically modified plant as described herein; and

[0061] FIG. 2 is a diagram of certain gene cassettes used to modifiy plants as described herein.

DETAILED DESCRIPTION

[0062] The detailed description set forth below is intended as a description of the presently preferred embodiment of the invention, and is not intended to represent the only form in which the present invention may be constructed or utilized. The description sets forth the functions and sequences of steps for constructing and operating the invention. It is to be understood, however, that the same or equivalent functions and sequences may be accomplished by different embodiments and that they are also intended to be encompassed within the scope of the invention.

[0063] Plant Platforms for Pheromone Production

[0064] In this disclosure, three different plant platforms were utilized: N. tabacum, N. benthamiana and Camelina sativa for pheromone production.

[0065] N. tabacum is also called cultivated tobacco which is an herbaceous plant and it is only found in cultivation. N. tabacum is the most commonly grown plant in the Nicotiana genus. It is commercially grown in many countries and the leaves are used to produce tobacco. The height of matured tobacco plants is between 1 and 2 meters. The leaves vary in size and the lower leaves are the largest with a length of up to 60 cm. N. benthamiana is a close relative of N. tabacum, and the mature plants show a large variation in height, ranging from as tall as 1.5 meters to shorter than 200 mm. In our greenhouse, the height of N. benthamiana was about 300 mm. The two Nicotiana species are both favourable to work with in metabolic engineering aiming at production of pheromone compounds as they have relatively short production times, large area of leaves to output volatiles and are relatively easy to grow in controlled growth conditions. In addition, there is less concern about contaminating food supplies as they are not food crops.

[0066] Camelina was chosen as the oilseed production platform for our studies because it is limited use as a food crop and is considered an ideal system for rapid introduction and evaluation of fatty acid and other oil-related traits (Iskandarov et al. 2014). Foremost, transgenes can easily be introduced into Camelina using a simple Agrobacterium-based method (Lu and Kang 2008), and it has a relatively short life cycle that allows up to three generations in a year for evaluation of engineered traits (Bansal and Durrett 2016). Camelina is also closely related to Arabidopsis thaliana, with a wealth of transgenic and genomic data for optimizing endogenous biosynthetic pathways for production of desired oil traits in seeds that typically are 30% to 40% oil by weight (Nguyen et al. 2013).

[0067] In many plants, trichomes are tiny specialized hair structures for secondary metabolite production and release. For instance, biosynthesis of the diterpenes takes place in trichome heads, where secretory vesicles and cells are located (Kandra and Wagner 1988; Duke 1994; Guo et al. 1995). CYP71D16 was confirmed as a trichome-specific promoter leading the downstream gene to be specifically expressed in plant trichome (Wang et al. 2002). To explore the possibility of releasing moth pheromones from Nicotiana leave's trichome cells, a N. tabacum trichome specific promoter pCYP71D16 was used for driving pheromone biosynthetic gene expression.

[0068] General Methodology

[0069] Labelling Experiment

[0070] Isotopic labelling was the technique used to identify moth pheromone biosynthetic pathways by tracking the passage of an isotope through the metabolic pathways in this disclosure. The deuterium-labelled precursors were separately dissolved in dimethylsulphoxide (DMSO) (Bjostad and Roelofs 1983; Yamaoka et al. 1984) and topically applied to the female abdominal tip where the pheromone gland is located. After incubation of a half to several hours, pheromone glands were excised and extracted (Bjostad and Roelofs 1984), and the samples could be analyzed by gas chromatography/mass spectrometry (GC/MS) (Christie 1998). The double bond positions in the fatty acid chain were confirmed by dimethyl disulfide (DMDS) derivatization (Dunkelblum et al. 1985).

[0071] Cloning and Plasmid Constructs for Assays

[0072] Amplification of genes was performed by PCR either from cDNA templates synthesized from total RNA or genome DNA extracted directly from plant materials (for trichome specific promoter pCYP71D16). Custom DNA synthesized by Invitrogen was used for some genes. All the genes contain the Gateway cloning site attB (Gateway cloning system, Invitrogen) and were subsequently cloned to Gateway entry vector in first step by BP reaction. For constructing co-expression clones, the Phusion PCR (Atanassov et al. 2009) was performed by putting two fragments that containing several dozens of homologue sequence bases and the DNA polymerase together. After hybridizing of the two fragments, the recombined sequence was cloned to Gateway vector. The expression clones in this disclosure were also constructed by using Gateway method (Katzen 2007). After the entry clones were confirmed by sequencing (Schuster 2007), all of them were sub-cloned to a destination vector by LR reaction with different multigene combinations for a variety of purposes.

[0073] Functional Assays

[0074] In this disclosure, we have used three platforms for gene functional characterization. One was the yeast expression system. The yeast expression vectors pYEX-CHT and pYES52 were used for the functional assays. The expression clones containing FADs were introduced into the double deficient olel/elol strain (MATa elol::HIS3 olel::LEU2 ade2 his3 leu2 ura3) of the yeast Saccharomyces cerevisiae, while the expression clones containing FARs were introduced into the INVSc strain of yeast S.c. (MATa HIS3 LEU2 trp1-289 ura3-52). The transformation of yeasts was carried out using the S.c. easy yeast transformation kit (Life technologies).

[0075] The second platform for gene functional assay was the plant transient expression system. In this, we used N. benthamiana as the plant platform for gene expression. The plant expression clones in pXZY393 vector containing the target genes were first introduced into Agrobacterium tumefaciens GV3101 strain (MP90RK) by electroporation (1700 V mm.sup.1, 5 ms, Eppendorf 2510). Meanwhile, a viral silencing suppressor protein P19 was introduced into the same A. tumefaciens strain as well in order to inhibit the transgene silencing of the host cells and extend transgene expression over a longer period of time with a higher degree of expression (Voinnet et al. 2003). Subsequently the transformed A. tumefaciens was incubated for several days until the culture concentration was high enough for infiltration of N. benthamiana. The infiltration experiment was carried out by using a 1 mL syringe without needle, containing the A. tumefaciens cells, to inject the lower side of a suitable four-week-old N. benthamiana leaf, with a gentle squeeze on the plunger and modest pressure on the leaf using a finger.

[0076] The last platform we used for gene functional assay was the insect cell expression system. The expression construct for candidate gene in the BEVS donor vector pDEST8 was made by LR reaction. Recombinant bacmids were made according to instructions for the Bac-to-Bac system given by the manufacturer Invitrogen using DH10MEmBacY (Geneva Biotech). Baculovirus generation was done using Sf9 cells (Invitrogen), Ex-Cell 420 medium (Sigma) and baculoFECTIN II (OET).

[0077] Leaf Disc Transformation Via Agrobacterium

[0078] The method of Agrobacterium-mediated leaf disc transformation (Clemente et al. 2006) was used for Nicotiana spp. stable transformation. First, the A. tumefaciens culture containing the construct was incubated at 30 C. in LB medium supplemented with suitable antibiotics, until the optical density (600 nm) can be adjusted to 0.9-1. Plant material was obtained from 4-5 weeks old Nicotiana plants grown under sterile conditions on MS medium (Murashige and Skoog 1962) in a climate chamber. Subsequently, the transgenic lines were obtained by Agrobacterium-mediated leaf-disc transformation. Leaf discs (20 mm20 mm) were cut out and incubated 5 min in an A. tumefaciens solution, dried with sterile napkin paper and transferred to Petri dishes with MS medium (Horsch el al. 1985). After 2-3 days incubation in darkness, leaf discs were transferred to selection medium. Then after 2-3 weeks of incubation, the callus produced on the leaf edges were transferred to shoot-inducing medium. After 2-3 weeks of incubation, the shoots were transferred to root-inducing medium. The shoots were finally transferred into soil and grown in greenhouse until maturity.

[0079] Floral Dip Transformation Via Agrobacterium

[0080] The method of Agrobacterium-mediated floral dip transformation was used for Camelina stable transformation. The constructed expression vectors were introduced into Agrobacterium tumefaciens strain GV3101 (MP90RK) by electroporation (1700 Vmm.sup.1, 5 ms, Eppendorf 2510). The transformed Agrobacterium cells were grown on solid LB medium supplemented with antibiotics (50 mg/L rifampicin, 50 mg/L gentamicin and 50 mg/L spectinomycin) after incubating at 30 C. for 36 h. Afterwards, a single clone from each expression clone was incubated in 2 mL liquid LB medium with suitable antibiotics as described above at 30 C. for 36 h. Then the Agrobacterium solution was transferred to 30 mL medium for a 36 h incubation, and after that, the solution was transferred to 1 L medium for a 24 h incubation. Subsequently, the five weeks old Camelina plants were then transformed (Lu and Kang 2008) by the floral dip/vacuum infiltration method as described in Liu et al. (2012). After eight to ten weeks, the seeds were collected and sown in soil, and herbicide was used to select the transformants.

[0081] Lipid Analysis

[0082] In order to analyze the gene functions in transformed yeast, insect cells and plants, as well as to detect the pheromone precursor production in Nicotiana spp. leaves and Camelina seeds, the lipid analysis was performed. For the lipid analysis, first the pheromone gland tissues, yeast cells, insect cells or plant leaves were extracted by chloroform:methanol (2:1 v:v) at room temperature overnight. For fatty acids (pheromone precursors) analysis, this was followed by base-methanolysis and transesterification as previously described (Dunkelblum and Kehat 1987) to convert all fatty acids to their corresponding methyl esters. The products were then recovered in n-heptane prior to GC/MS analysis. While the samples for pheromone compounds such as alcohol, aldehyde and acetate analysis were followed by adding n-heptane to re-dissolve the compounds after the total lipid dryness by nitrogen, which were then transferred to new glass vials for GC/MS analysis. The seed fatty acids extraction and methanolysis was performed directly by adding 2% sulphuric acid in methanol and incubated at 90 C. for 1 h, and then water and n-heptane were added to extract the fatty acid methyl ester products prior to GC/MS analysis.

[0083] Headspace Volatile Collection

[0084] To perform the experiment of plant headspace volatile collection, the experiment was carried out as described in Raguso and Pellmyr (1998). For the use of solid phase micro extraction (SPME) to collect volatile, the experiment was performed as described in Centini et al. (1996).

[0085] Results and Discussion

[0086] Labelling Experiments Reveal New FADs Activities

[0087] Tracking the pheromone biosynthetic pathways of S. exigua and L. botrana by isotope-labelling experiment, we confirmed an unusual 12 and a novel 7 desaturation activity to be involved, respectively. In the S. exigua, the in vivo labelling experiment showed that deuterium atoms from [16,16,16-.sup.2H.sub.3] hexadecanoic acid (D.sub.3-16:acid) were incorporated into all the detected acetates and alcohols, including (Z)-11-hexadecenyl acetate (Z11-16:OAc), (Z)-11-hexadecenol (Z11-16:OH), (Z)-9-tetradecenyl acetate (Z9-14:OAc), (Z)-9-tetradecenol (Z9-14:OH), (Z,E)-9,12-tetradecadienoic acetate (Z9,E12-14:OAc) and (Z,E)-9,12-tetradecadienol (Z9,E12-14:OH), as well as into the corresponding FAMEs, i.e., methyl (Z)-11-hexadecanoate (Z11-16:Me), methyl (Z)-9-tetradecenoate (Z9-14:Me), methyl (Z,E)-9,12-tetradecadienoate (Z9,E12-14:Me) and methyl myristate (14:Me). Notably, when non-labeled (Z)-9-tetradecenoic acid (Z9-14:acid) was applied to the pheromone gland, the amount of Z9,E12-14:OAc increased significantly. In contrast, when [14,14,14-.sup.2H3] tetradecanoic acid (D.sub.3-14:acid) and (E)-12-[14,14,14-.sup.2H3] tetradecenoic acid (D.sub.3-E12-14:acid) were applied, label incorporation was not detected in any of the above components. These results confirm that the production of major pheromone component Z9,E12-14:OAc in S. exigua starts from palmitic acid and a 11 FADs acts on palmitic acid to produce Z11-16:acid, which is then chain-shortened to Z9-14:acid, followed by the second desaturation at the 12 position to form Z9,E12-14:OAc. This pathway is in line with previous study of S. littoralis (Jurenka, 1997).

[0088] Deuterium labelled D.sub.3-16:acid, D.sub.3-14:acid, (Z)-11-[13,13,14,14,14,-.sup.2H3] tetradecenoic acid (D.sub.5-Z11-14:acid), (Z)-9-[12,12,12-.sup.2H3] dodecenoic acid (D.sub.3-Z9-12:acid) and [12,12,12-.sup.2H3]dodecanoic acid (D.sub.3-12:acid) were topically applied to the pheromone glands of L. botrana to track the biosynthetic pathway. The results showed that deuterium atoms from D.sub.3-16:acid were incorporated into all the pheromone components, including (Z)-9-dodecenyl acetate (Z9-12:OAc), (E,Z)-7,9-dodecadienol (E7,Z9-12:OH) and (E,Z)-7,9-dodecadienyl acetate (E7,Z9-12:OAc). In the fatty acyl compounds analysis, deuterium atoms from D.sub.3-16:acid were incorporated into methyl palmitate (16:Me), 14:Me, methyl (Z)-11-tetradecanoate (Z11-14:Me), methyl lauritate (12:Me) and methyl (E,Z)-7,9-dodecadienoate (E7,Z9-12:Me). Application of D.sub.3-14:acid showed the same incorporation results as observed for D.sub.3-16:acid, whereas no label incorporation was found from D.sub.3-14:acid into 16:acid. Labels from D.sub.5-Z11-14:acid were incorporated into Z9-12:OAc, E7,Z9-12:OH and E7,Z9-12:OAc, as well as into Z9-12:Me and E7,Z9-12:Me intermediates. Additionally, labels from D.sub.3-Z9-12:acid were extremely highly incorporated into E7,Z9-12:OAc and incorporated into Z9-12:OAc and E7,Z9-12:OH as well. In contrast, when D.sub.3-12:acid was applied, label incorporation was not detected in any of the above mentioned compounds. The incorporation of deuterium labels from D.sub.3-Z9-12:acid into E7,Z9-12:OAc, E7,Z9-12:OH, and E7,Z9-12:Me strongly suggests an elusive 7 desaturation on Z9-12:acid to form the major pheromone precursor E7,Z9-12:acid in L. botrana.

[0089] Functional Assays of Novel FAD Genes

[0090] Here is disclosed several novel FAD genes with a variety of functions. First, involving the 12 desaturation activity, two unusual 11/12 FAD genes SexiDes5 and SlitDes5 were characterized in S. exigua and the congeneric S. litura, with the same functions using palmitic acid and the subsequently chain-shortened product (Z)-9-tetradecenoic acid (Z9-14:acid) as substrates to produce (Z,E)-9,12-tetradecadienoic acid (Z9,E12-14:acid), respectively. A 11 FAD gene Lbo_PPTQ from L. botrana can produce important pheromone intermediate Z11-14:acid from myristic acid. C. suppressalis utilizes (Z)-11-hexadecenal (Z11-16:Ald) as its major pheromone component. By heterologous expression in yeast and plants, a 11 FAD gene CsupYPAQ presents very high substrate specificity to palmitic acid for production of (Z)-11-hexadecenoic acid (Z11-16:acid) with great activity. Another FAD gene CsupKPSE from C. suppressalis has preference for C.sub.16. It is interesting, however, that CsupKPSE switches the preference for C.sub.16 to C.sub.18 to form oleic acid when the culture nutrition was limited.

[0091] The predicted ER retention signature motif in 11/12 FADs SexiDes5, SlitDes5, and 11 FAD CsupYPAQ is LPAQ, LPSQ, and YPAQ respectively. It is noticed that the motif difference in FADs is to some extent related to the functions, e.g., FADs within the KPSE group are 9 desaturases having preference for C.sub.16, while the NPVE group are mainly modifying C.sub.18 (Rosenfield et al. 2001; Liu et al. 2002; Liu et al. 2004; Rodriguez et al. 2004). The xxxQ are most likely to be the 11, 10 and multi-functional FADs and a few exceptions are 9 FADs, which are exclusively involved in pheromone biosynthesis (Knipple et al. 1998; Hao et al. 2002; Serra et al. 2007; Xia et al. 2015). The 6 and 14 groups contain a mixture of different signature motifs from the 9 and 11 groups, and their biological functions are diverged from 9, and 11 FADs (Roelofs et al. 2002; Wang et al. 2010). On the other hand, there were some exceptions to the motif signal hypothesis. For example, Ding et al. (2016) reported that one amino acid at the cytosolic carboxyl terminus of the protein (258E), which is outside of the motif region, is critical for the Z activity of the Choristoneura rosaceana FAD.

[0092] In this disclosure, SexiDes5 and SlitDes5 were showing same functions in the yeast although with different ER motifs. The motif of CsupYPAQ has only one amino acid different from SexiDes5. However, CsupYPAQ is highly specific to palmitic acid, whereas SexiDes5 can form a variety of products with wide preferences for both saturated and unsaturated fatty acids. Thus, making mutations at different sites for these unique FADs could be a possible approach to explore the relationship between sequences and functions of FADs.

[0093] Production of Pheromone Precursors in Plants

[0094] Many functionally diversified FADs were characterized by previous studies, making it possible to design tailor-made pheromone precursor production in engineered plants. In this disclosure, N. tabacum, N. benthamiana and C. sativa were successfully transformed to obtain stable transgenic lines producing pheromone precursors.

[0095] The successful production of several insect sex pheromone precursors in N. tabacum and N. benthamiana by stable transformation was demonstrated. Levels of up to 17.6% Z11-16:acid in the total fatty acids was achieved. All of the transformed plastid FatBs and FADs were functionally active in producing pheromone precursor in both Nicotiana species. This is the first report of Z/E11-14:acid production in a plant by stable transformation. The performance of Z11-16:acid production in N. benthamiana was better than in N. tabacum. The average value of Z11-16:acid production in TO N. tabacum was 0.2% whereas in TO N. benthamiana it was 1.8%. The best N. benthamiana transgenic line #025 produced as high as 13.6% of Z11-16:acid of the total fatty acids in T2 plants, which is much higher than the production amount reported from Nenerov et al. 2004, claiming that 6% of Z11-16:acid of total fatty acids was produced in their transgenic N. tabacum NtD15B line. The quantity of Z11-16:acid in the form of methyl ester in N. benthamiana determined to be 335 g per gram fresh leaf is also higher, compared to 32 g per gram of N. tabacum fresh leaf in Nenerov et al. 2004. The results suggest that N. benthamiana has potential to be more efficient than N. tabacum as a plant factory for Z11-16:acid production. In the report of Ding et al. 2014, 381 g per gram fresh leaf of Z11-16:acid was produced in N. benthamiana by transient expression, which is a massive overexpression of exogenous genes over a few days that ignores the health of the plant. Here, the production of 335 g per gram leaf of Z11-16:acid by stable transformation shows the ability of vegetative material to function with the expression of CpuFatB1 and Atr11 and yield compounds over development, providing the potential for further commercial production.

[0096] To investigate the feasibility of a plant factory production in Camelina, we established four different types of transformant lines by using different exogenous gene cassettes, as described below. In the regenerated Camelina seeds, the mono-unsaturated E9-12:acid with small amount of (Z)-9-dodecenoic acid (Z9-12:acid) and diunsaturated E8,E10-12:acid were produced in all the four types of transformant lines, as shown in FIG. 1. Camelina was genetically modified for production of E8,E10-12:acid via stable integration of gene cassettes using Agrobacterium-mediated floral dip transformation. This is the first report on production of di-unsaturated pheromone precursors in plants. Furthermore, the production amount of 6.3% of E8,E10-12:acid of total fatty acids is quite high. Because the oil content of the Camelina seeds, on a dry weight basis, is typically between 35 to 45% and the yields of Camelina are in a range of 336 to 2240 kg of seeds per hectare (Moser 2010). This means that 7.4 to 63.5 kg (minimum to maximum) of E8,E10-12:acid might be produced by cultivating our best Camelina line. Moreover, in this study we also investigated four strategies for optimization of the plant factory for production of E8,E10-12:acid. We demonstrated that co-expression of the desaturase with P19 and multiple gene copies can increase the production of C.sub.12 pheromone precursors significantly. Also, it was confirmed that stably expressing P19 regulated by the seed-specific napin promoter would not cause observable harm of plant development.

[0097] Our studies described here explore the use of an oilseed crop Camelina sativa (Camelina) as a production platform for C.sub.12 pheromone precursors. To optimize the seeds of oil crops for the production of these precursors requires redesigning fatty acid biosynthesis to generate maximal levels of 12:0, rather than the typical C.sub.16 and C.sub.18 fatty acids found in triacylglycerols (TAG) of these seeds. In plants, the biosynthesis of MCFA is a variation on typical de novo fatty acid synthesis that generates primarily C.sub.16 and C.sub.18 fatty acids. Chain-lengths of fatty acids are primarily determined by acyl-ACP thioesterases, including FatB thioesterases that typically release C.sub.16 acyl chains from de novo fatty acid biosynthesis (Li-Beisson et al. 2013). Variant forms of FatB, found in selected plant species, are able to release fatty acids of chain lengths shorter than C.sub.16, as demonstrated by transgenic expression in seeds (Pollard et al. 1991; Jones et al. 1995; Voelker 1996; Tjellstrm et al. 2013; Kim et al. 2015). In previous study, a FatB gene UcTE from California bay laurel (Umbellularia californica) was found to have high activity for production of 12:0 in rapeseed (Brassica napus) (Voelker et al. 1992). When the MCFAs are exported into the cytoplasm from the plastid in oilseeds, they become available for incorporation into TAG, which is formed most directly by the Kennedy pathway enzymes of glycerol-3-phosphate acyltransferase (GPAT), lysophosphatidic acid acyltransferase (LPAAT) and diacylglycerol acyltransferase (DGAT) continuously (Thelen and Ohlrogge, 2002; Cahoon et al. 2007; Dyer et al. 2008; Kim et al. 2015a). A recent report showed that the co-expression of a variant FatB thioesterase with LPAT in Camelina seeds, the MCFA accumulation was improved (Kim et al. 2015a). In addition, over-expression of the seed specific transcription factor WRINKLED1 from Arabidopsis was confirmed a useful approach to elevate the TAG concentration (Cernac and Benning 2004).

[0098] Raising the target compound in transgenic engineering plants by transformation of multiple transgene copies also have been reported in some studies (Carrier et al. 1998; Mansur et al., 2005; Nguyen et al., 2015; Schultz et al. 1987). This approach may be hampered by the observation that the silencing effect of the transgenes could be increased (Matzke et al. 1994; Tang et al. 2006). However, new biotechnological ways have been reported to avoid the gene silencing effect by using different promoter for each gene copy as well as codon optimization (Fath et al. 2011). Moreover, Naim et al. (2016) informed that stable expression of silencing-suppressor proteins (SSPs) such as P19, V2, PO could enhance the performance and longevity of a metabolic pathway.

[0099] In this study, we engineered the pathways towards to the production of the major fatty acyl pheromone precursor, E8,E10-12:acid, of the codling moth Cydia pomonella in Camelina seeds. Camelina was chosen as the oilseed production platform for our studies because it is limited use as a food crop and is considered an ideal system for rapid introduction and evaluation of fatty acid and other oil-related traits (Iskandarov et al. 2014). Foremost, transgenes can easily be introduced into Camelina using a simple Agrobacterium-based method (Lu and Kang 2008), and it has a relatively short life cycle that allows up to three generations in a year for evaluation of engineered traits (Bansal and Durrett 2016). Camelina is also closely related to Arabidopsis thaliana, with a wealth of transgenic and genomic data for optimizing endogenous biosynthetic pathways for production of desired oil traits in seeds that typically are 30% to 40% oil by weight (Nguyen et al. 2013).

[0100] To explore oilseed production of high amount of C.sub.12 pheromone precursors, we co-expressed the UcTE and CpCPRQ in Camelina seeds for production of (E)-9-dodecenoic acid (E9-12:acid) and (E,E)-8,10-dodecadienoic acid (E8,E10-12:acid). We investigated four strategies towards the production of these two compound in the plant factory: 1) co-expression of a seed-specific synthetic transcription factor WRINKLED1 from Arabidopsis thaliana enhancing fatty acid synthesis and a Kennedy pathway gene, lysophosphatidic acid acyltransferase gene CvLPAAT from Cuphea viscosissima; 2) transformation of only one gene copy of CpCPRQ into high lauric Camelina seeds; 3) transformation of multiple gene copies with different promoters and terminators; and 4) stable expression of the viral gene silencing suppressor protein P19 together with pheromone biosynthetic genes. Our findings demonstrated the feasibility of producing the codlemone precursor and other C.sub.12 pheromone precursors in stably transformed plant seeds.

[0101] Plant Material and Growth Conditions

[0102] For these studies, Camelina sativa cv. Suneson (Camelina) was used. The previously described high lauric (20 mol % of total seed fatty acids) Camelina line was used as our primary metabolic engineering platform (Kim et al. 2015a). Two plants per pot with soil (Krukvaxtjord lera & kisel) were grown under greenhouse conditions of 24 C., 14 h day/18 C., 10 h night, with supplemental lighting.

[0103] Gene CloningPreparation of Constructs

[0104] UcTE (Genebank access number: Q41635.1), codon optimized-CpCPRQ opening reading frames (ORFs), seed-specific promoters for the -subunit of -conglycinin gene (-con) (Chamberland et al. 1992), -phaseolin (-Phaseolin) (Geest and Hall 1996), oleosin gene (Oleosin) (Fan et al. 2013), nopaline synthase terminator (NOS), and nopaline synthase terminator (HSP) were synthesized by Invitrogen. ORF of CpCPRQ (AHW98354), AtWRINKLED1 (AY254038), CvLPAAT (ALM22867), P19 (P69516.1), and sequence of napin gene promoter (napin), octopine synthase terminator (OCS) were amplified from entry clones.

[0105] To compare different strategies toward production of high amounts of C.sub.12 pheromone precursors in Camelina, four plant expression vectors were constructed (FIG. 2). CPRQ1.0 contained four exogenous genes, which were UcTE controlled by Glycinin, CpCPRQ, AtWRINKLED1 and CvLPAAT controlled by napin (FIG. 2a). CPRQ1.1 contained one exogenous gene CpCPRQ codon optimized for Arabidopsis thaliana (Arabidopsis), controlled by Glycinin (FIG. 2b). CPRQ2.1 contained four exogenous genes controlled by different promoters, including two copies of CpCPRQ with or without codon optimized for Arabidopsis, a UcTE, and a virus silencing suppressor protein P19 (FIG. 2c). Similar to CPRQ2.1, CPRQ2.2 also contained four exogenous genes, which were three copies of CpCPRQ (one without codon optimized, one codon optimized for Arabidopsis, one codon optimized for Oryza sativa) controlled by different promoters, and a P19 (FIG. 2d). Except for CPRQ1.0 which was transformed into wild type Camelina, the other three vectors were transformed into high lauric acid type Camelina.

[0106] PCR amplification was performed using the entry clone as template with a pair of degenerate primers, on a Veriti Thermo Cycler, using Phusion Flash High-Fidelity PCR Master Mix (Thermo Scientific) under conditions as follows: start at 98 C. for 30 s, and 38 cycles at 98 C. for 5 s, 55 C. for 10 s and 72 C. for 50 s, followed by a final extension step at 72 C. for 10 mins. Subsequently, fusion PCR was performed using PhusionTaq (Thermo Scientific) (Atanassov et al., 2009) to do truncation and gene fusion for gene assembly, the same PCR programs as described before. All genes with promoters and terminators were cloned into the plant expression vector pBinGlyBar (Nguyen et al. 2013), which contained a bar marker gene for Basta selection of transformed plants, by using Multisite Gateway recombination cloning technology (Invitrogen). The constructed expression clones were confirmed by sequencing.

[0107] Floral Dip Transformation of Camelina by Agrobacterium-Media

[0108] The constructed expression vectors were introduced into Agrobacterium tumefaciens strain GV3101 (MP90RK) by electroporation (1700 Vmm.sup.1, 5 ms, Eppendorf 2510). The transformed Agrobacterium cells were grown on solid LB medium supplemented with antibiotics 50 mg/L rifampicin, 50 mg/L gentamicin and 50 mg/L spectinomycin after incubating at 30 C. for 36 h. Afterwards, a single clone from each expression clone was incubated in 2 mL liquid LB medium with antibiotics as described above at 30 C. for 36 h. The Agrobacterium solution was then transferred to 30 mL medium for a 36 h incubation, and after that, the solution was transferred to 1 L medium for a 24 h incubation. Subsequently, the 5 weeks old Camelina plants were transformed by the floral dip/vacuum infiltration method as described by Lu and Kang (2008) and Liu et al., 2012. Basta resistant gene was used as a selection marker (Nguyen et al. 2013).

[0109] Fatty Acid Analysis of Seed Oils

[0110] Seeds harvested from the floral-dipped plants were sown in soil for Basta selection, and the surviving plants were T1 plants and considered as transformants. T2 seeds were harvested from matured T1 plants for fatty acid analysis. Fatty acids were analyzed as fatty acid methyl esters (FAMEs), which were generated from putative transformants either by grinding 25 pooled T2 seeds (for production analysis of one transformant) or by grinding 15 T2 seeds individually (for variation analysis within one transformant) from each T1 plants in 1 mL 2% H.sub.2SO.sub.4 in methanol in a 4 mL glass vial. After grinding, the samples were incubated for 1 h at 90 C. After cooling, 1 mL water and 1 mL heptane were added and the vial was vortexed. Then, the heptane phase containing the FAMEs was transferred to a new Agilent vial for GC/MS analysis on an Agilent 5975 mass-selective detector, coupled to an Agilent 6890 series gas chromatograph equipped with a polar column (HP-INNOWax, 30 m0.25 mm, 0.25 m), and helium was used as carrier gas. The oven temperature was set at 80 C. for 1 min, then increased to 230 C. at a rate of 10 C./min and held for 10 min. Fatty acid compounds were identified by comparison of retention times and mass spectra with those of reference compounds. To determine the position of double bonds in unsaturated fatty acids, DMDS derivatization was performed according to Dunkelblum et al. (1985). The DMDS-adducts were analyzed by GC/MS on a non-polar column (HP-5MS, 30 m0.25 mm, 0.25 m) under the following oven temperature program: 80 C. for 2 min, then increased at a rate of 15 C./min to 140 C., and then increased at a rate of 5 C./min to 260 C., and held for 30 min.

[0111] Assembly of Pathways for Moth Pheromone Precursor Biosynthesis and Accumulation in Camelina Seeds

[0112] The thioesterase encoded by UcTE taking the laurate plastid acyl carrier protein (ACP) from chain elongation, to form the corresponding lauric (12:0) acid. The free fatty acid is acylated to 12:CoA when transported out of the plastid into the cytosol. The 12:CoA is subsequently converted into corresponding pheromone precursors Z/E9-12:CoA, and E8,E10-12:CoA by the desaturase encoded by CpCPRQ.

[0113] C.sub.12 Pheromone Precursors were Produced in Camelina Transformants

[0114] GC/MS analysis of C.sub.12 to C.sub.18 chain length fatty acids showed that wild type Camelina seeds contained the highest amount of linolenic acid (18:3), followed by linoleic acid (18:2) and oleic acid (18:1). In addition, small fractions of lauric acid, myristic acid, and arachidic acid were found in the seeds. In the high lauric acid (12:0) Camelina seeds, the amount of 12:0 was as high as 18:3, and 18:1 was the second most abundant fatty acid.

[0115] The C.sub.12 pheromone precursors were produced under every strategy in the present study. By analyzing 25 pooled seeds from each transformant, we found that among the nine transformants produced by introduction of gene cassette CPRQ1.0, the most productive Camelina produced 1.7% of E9-12:acid and 0.15% of E8,E10-12:acid from the total fatty acids. The percentage of fatty acids here was calculated as weight percent of total fatty acids. CPRQ1.1 was transformed into high lauric acid Camelina, but the high concentration of 12:0 did not lead to higher production of the unsaturated products. In total, 85 transformants were obtained from CPRQ1.1, of which the best line produced 1.6% of E9-12:acid and 0.15% of E8,E10-12:acid. Similar to CPRQ1.1, the vectors of CPRQ2.1 and CPRQ2.2 were separately transformed into high lauric acid Camelina. In total, we obtained 96 and 30 transformants of CPRQ2.1 and CPRQ2.2, respectively. The results of fatty acids analysis showed that about 83 of CPRQ2.1 transformants lost a lot of the content of 12:0, which dropped from 20% to 3% and no unsaturated pheromone precursor was produced. The other 13 transformants of CPRQ2.1 produced a higher amount of 12:0, increasing up to 31.6%, and produced up to 2.3% of E9-12:acid and 0.20% of E8,E10-12:acid. In the CPRQ2.2 transformants, abundance of E9-12:acid and E8,E10-12:acid in 25 pooled seed was 6.4% and 3.7%, respectively from the most productive plant.

[0116] Comparison of Pheromone Precursors Production Between Four Strategies

[0117] Comparing the use ratio of 12:0 in CPRQ1.0 transformants to CPRQ1.1 and CPRQ2.1 showed that the CpCPRQ in the former transformants converted a higher percentage of 12:0 into unsaturated pheromone precursors. The CPRQ2.2 transformants converted highest percentage of 12:0 into corresponding pheromone precursors.

[0118] Compared to CPRQ1.0 and 1.1, the CPRQ2.1 and 2.2 transformants which contained multiple gene copies and P19 increased the production of pheromone precursors. The most productive Camelina harboring CPRQ2.1 contained up to 2.3% of E9-12:acid and 0.2% of E8,E10-12:acid, which is higher than the best ones in CPRQ1.0 and CPRQ1.1. The Camelina expressing gene cassette of CPRQ2.2 which contained three copies of the 9 desaturase CpCPRQ produced much higher E9-12:acid and E8,E10-12:acid than CPRQ2.1 that had two copies of CpCPRQ. In addition, no abnormal plant development was observed among CPRQ2.1 and CPRQ2.2 transformants.

[0119] Increase of Oleic Acid in CPRQ2.1 Transformants

[0120] The CPRQ2.1 transformants showed two different fatty acid profile groups. In the first profile group, the 12:0 or 18:3 is the dominant fatty acid, followed by 18:2 and 18:1, which is similar to other transformants (CPRQ1.0, CPRQ1.1, CPRQ2.2), the wild type and the high lauric type. In the second profile group, oleic acid (18:1) is the dominant fatty acid species, which was shown in the 13 of the CPRQ2.1 transformants with a decreased amount of 12:0. Moreover, the ones that lost the 12:0 did not produce any unsaturated pheromone precursors.

[0121] The successful production of mono- and di-unsaturated C.sub.12 pheromone precursors in Camelina seeds confirmed the in-planta function of the 9 desaturase derived from C. pomonella. When CpCPRQ was expressed in Camelina it could produce E9-12:acyl from lauric acid as well as a small amount of the corresponding Z isomer. Furthermore, it had the activity to convert the monoene intermediate into E8,E10-12:acid with conjugated double bonds. This gene bifunctionality is consistent with the previous study of Ding and Lfstedt, unpublished using the sf9 insect cell expression system. Our studies substantiate the application of the plant expression system for functional characterization of insect pheromone biosynthetic genes, as well as demonstrated the feasibility of production of diene moth sex pheromone precursors in plant.

[0122] We established four different types of transformant lines by using different exogenous gene cassettes. In the regenerated Camelina seeds, the mono-unsaturated E9-12:acid with small amount of (Z)-9-dodecenoic acid (Z9-12:acid) and diunsaturated E8,E10-12:acid were produced in all the four types of transformant lines. Comparing the fatty acid profiles of transformants from CPRQ1.0 and CPRQ1.1 suggests that co-expression with the transcription factor WRINKLED1 from Arabidopsis AtWRI1 and one of the Kennedy pathway gene lysophosphatidic acid acyltransferase from C. viscosissima CvLPAAT can improve the substrate utilization of 12:0. For example, the mean use ratio of 12:0 in CPRQ1.0 was 7.5%, while in CPRQ1.1 transformant it was 2.9%. Also, this implied that the bottleneck of production of pheromone precursors were due to the activity of CpCPRQ, but not the amount of 12:0. Among 85 transformants from CPRQ1.1, containing ca. 20% of 12:0, the most productive plant produced even lower E9-12:acid and same level of E8,E10-12:acid as compared to CPRQ1.0, containing at the most 10% of 12:0 in nine transformants.

[0123] Nevertheless, CpCPRQ showed high activity towards E9-12:acid and E8,E10-12:acid production in insect cells (Ding and Lfstedt, unpublished). Why did CpCPRQ show a lower activity in Camelina CPRQ1.0 and 1.1 transformants? One explanation could be the silencing effect of transgenes in Camelina, i.e., small RNAs-guided silencing pathways in plants may have the ability to shut down the exogenous expression (Hagan et al. 2003; Schubert et al. 2004). By expressing two other gene cassettes CPRQ2.1 and CPRQ2.2 in Camelina seeds, we concluded that the low activity of CpCPRQ might indeed be caused by transgene silencing. Both CPRQ2.1 and CPRQ2.2 contained the gene encoding P19 which is a viral silencing suppressor protein (VSP) from Tomato bushy stunt virus (TBSV), was reported to have the ability to suppress the silencing effect of transgenes in previous studies (Naim et al. 2016). Naim et al. 2016 demonstrated that expression of silencing-suppressor protein can protect and enhance stable transgene performance. Here, the expression of CPRQ2.1 and CPRQ2.2 did increase the production of E/Z9-12:acid and E8,E10-12:acid significantly.

[0124] Additionally, the VSPs have been widely used together with transgene cassettes to enhance their expression during transient expression experiments. But for the stable transformation the VSP is rarely utilized and explored as they may interfere with endogenous microRNA-regulated process and lead to abnormal plant development (Dunoyer et al. 2004; Fusaro et al. 2012). In the present study, we investigated the use of fully functional P19 expressed only in seeds and controlled by the napin promoter. We demonstrated that stably expressed P19 in Camelina seeds with napin promoter did not cause any abnormal plant development. The germination rate of the most productive seeds from CPRQ2.2 was as high as 95%. This may be due to the native napin promoter driving the seed storage proteins like napin, whose physiological role is to provide the growing seedling with essential nutrients prior to the establishment of the photosynthetic capacity (Rask et al. 1998). Thus, using the napin promoter to control the expression of P19 would not disrupt the small RNA driven regulatory pathways towards the development of vegetative shoot in Camelina (Wong et al. 2011).

[0125] Additionally, it was surprising to find that the best tested individual seed from CPRQ2.1 #010 transformant only produced 0.5% of E8,E10-12:acid, while the seed from CPRQ2.2 #005 transformant produced 6.3% of E8,E10-12:acid. This means that one more copy only of CpCPRQ contributed to a more than ten times higher production amount of this di-unsaturated pheromone precursor. Thus, it demonstrated that multiple gene copies are performing excellently in Camelina towards the di-unsaturated pheromone precursors production. Also, we observed a large variation in the production among the different individual seeds, which means that the production of these C.sub.12 pheromone precursors in seeds are still unstable and it is also implied that there is a potential to obtain improved Camelina lines producing even higher concentration of the target compounds.

[0126] In conclusion, in the present study we investigated the possibility of producing di-unsaturated moth pheromone precursors in transgenic plants. Camelina was genetically modified for production of E8,E10-12:acid via stable integration of gene cassettes using Agrobacterium-mediated floral dip transformation. This is the first report on production of di-unsaturated pheromone precursors in plants. Furthermore, the production amount of 6.3% of E8,E10-12:acid of total fatty acids is quite high. Because the oil content of the Camelina seeds, on a dry weight basis, is typically between 35 to 45% and the yields of Camelina are in a range of 336 to 2240 kg of seeds per hectare (Moser 2010). This means that 7.4 to 63.5 kg (minimum to maximum) of E8,E10-12:acid might be produced by cultivating our best Camelina line. Moreover, in this study we also investigated four strategies for optimization of the plant factory for production of E8,E10-12:acid. We demonstrated that co-expression of the desaturase with P19 and multiple gene copies can increase the production of C.sub.2 pheromone precursors significantly. Also, it was confirmed that stably expressing P19 regulated by the seed-specific napin promoter would not cause observable harm of plant development.

[0127] New Platform for Functional Assay of Pheromone Biosynthetic Genes

[0128] In this disclosure, a new concept for molecular mechanism exploration of pheromone biosynthesis is provided, which is using N. benthamiana transient expression platform. The idea emerged from the principle of plant factories. Transient expression of various insect genes in plant leaves as factories for pheromone production has been demonstrated for several years (Ding et al. 2014), but utilizing this platform for gene function studies has never been explored. In this study, we confirm that the plant transient expression system is efficient and useful.

[0129] Plants were used as a platform for functional characterization of the C. suppressalis biosynthetic gene candidates in order to compare the results to the yeast expression system. The observed functions of these genes in yeast and plant are similar and complementary. For instance, it is interesting that CsupKPSE showed C.sub.18>C.sub.16 substrate preference when oleic acid was absent in olel elol yeast. However, in the plant, CsupKPSE shows its preference on C.sub.16>C.sub.18 because the leaf does not lack oleic acid. The phylogenetic analysis indicated that CsupKPSE belongs to the C.sub.16>C.sub.18 clade. This implies that CsupKPSE can adjust its function to produce oleic acid similar to the function of ancestral metabolic desaturase when the nutrition is limited. Therefore, expression of desaturase genes in plant leaves for identification of different functions has the advantage of avoiding the problem of supplemented nutrition inference, especially for 9 desaturase identification. A FAR gene CsupFAR2 expressed low activity in yeast, while in plants it shows very high activity. Because the plant lipids are different from yeast, it offers an alternative way for gene function studies. The pgFAR CsupFAR2 showed additional minor activities on polyunsaturated fatty acids of linoleic acid and -linolenic acid in N. benthamiana, which has never been reported in previous FAR function studies. Moreover, the algae ELO IgalASE1 elongated linoleic acid (18:2) and -linolenic acid (18:3) to eicosadienoic acid (20:2) and eicosatrienoic acid (20:3) in N. benthamiana, show the same function as reported in yeast S. cerevisiae expression system (33), indicating that heterologous expression of ELO in plant leaves for functional characterization is reliable and feasible.

[0130] New ACO, ELO and FAR Findings

[0131] The specific skeleton of an unsaturated fatty acyl chain is generally produced by combination of FAD and ELO or ACO. Among the functionally characterized pheromone biosynthetic genes in L. botrana and C. suppressalis, there were two novel ACO genes Lbo_31670, Lbo_49602 and a first reported ELO gene CsupELO4 showing functions. The former two genes encoding two acyl-CoA oxidases that may produce Z9-12:acid by chain shortening of Z11-14:acid. The later gene encoding an elongase may produce Z13-18:acid, an immediate precursor of a pheromone component in C. suppressalis.

[0132] We also functionally characterized a new FAR gene from C. suppressalis, CsupFAR2 that shows very high activity to Z11-16:acid, producing corresponding Z11-16:OH. It also has other minor activity with C.sub.16-C.sub.18 fatty acids, similar to HarFAR from Helicoverpa armigera (Hagstrm et al. 2012; 2013), but having more specific activities on Z11-16:acid. The high activity and substrate specificity of CsupFAR2 suggests it to be a valuable candidate for further pheromone production in both yeast and plant factories.

[0133] Increase the Production Amount of Pheromone Compounds-Trichome Specific Promoter

[0134] A long-term vision is to produce genetically modified plants that eventually can be used in intercropping as natural dispensers of pheromones and as part of a push-pull (Cook et al. 2007) strategy, providing an innovative and environmentally friendly approach for pest management. Production of a high yield of pheromones in plants by stable transformation is still challenging. With the purpose of producing fatty alcohols or acetates especially for releasing these compounds from the leaf, a trichome specific promoter derived from N. tabacum was cloned and used for FAR and ATF gene expression.

[0135] In moths, after the immediate acyl-CoA precursors are produced, FAR can catalyze the reduction of acyl-CoA into fatty alcohols, which either are used as pheromones for some moth species, or in some other moths, converted to corresponding acetates after trans-acetylation by ATFs. Therefore, in order to explore the possibility of releasing pheromone alcohol and acetate in transformed plants, the FAR gene HarFAR and the ATF gene ATF1 was constructed with a trichome specific promoter CYP71D16, respectively, producing the pheromones Z11-16:OH and Z11-16:OAc.

[0136] CYP71D16 was confirmed as a trichome-specific promoter leading the downstream gene to be specifically expressed in plant trichomes (Wang et al. 2002). After 3-5 days, the infiltrated N. benthamina plant expressing HarFAR controlled by CYP71D16 promoter produced 70 g Z11-16:OH in per gram fresh leaf, while the plant expressing HarFAR derived by CaMv35S promoter only produced a tiny amount of Z11-16:OH. The same combination of CpFatB1, Atr11 and HarFAR derived by 35S promoter were also tested in the study of Ding et al. 2014, which reported 18 g Z11-16:OH in per gram fresh leaf was produced. Also, ATF1 controlled by CYP71D16 expressing in the plant produced much more Z11-16:OAc than 35S. This result suggested that CYP71D16 promoter can increase the pheromone production amount significantly compared to 35S promoter.

[0137] Conclusions and Perspectives

[0138] Initially, we characterized several key pheromone biosynthetic genes in the serious pest species, including two beet armyworm Spodoptera species, European grapevine moth L. botrana and the rice stem borer C. suppressalis. These findings will not only help understanding the mechanisms of pheromone biosynthesis, but will also provide many putative candidates for pheromone production in bio-factories.

[0139] In the perspective of utilizing results from basic science for application, the second contribution of this disclosure is that we established several stable transgenic plant lines for pheromone precursors production, ranging from the carbon chain length of 12 to 16, either in the leaves or seeds. This is the first report on an extended production of pheromone precursors (Cu and C.sub.14) over generations in plants. We confirm that N. benthamiana is a suitable platform for stable production of C.sub.16 pheromone precursors. We also established productive Camelina line for di-unsaturated C.sub.12 pheromone precursor production. Up to 6.3% of E8,E10-12:acid of total fatty acids was achieved. Because the oil content of the Camelina seeds, on a dry weight basis, is typically between 35 to 45% and the Camelina yields anywhere are from 336 to 2240 kg of seeds per hectare, which means 7.4 to 63.5 kg (minimum to maximum) of E8,E10-12:acid might be produced by cultivating our best Camelina line.

[0140] A further aim is to enable stable production of final pheromone components in bio-factories, which remains to be further explored. First, increasing the pheromone precursor production to provide more substrate for pheromone biosynthesis is one of the possible approaches. How to increase the precursor production, for instance by elevating the TAG accumulation in plant cells, needs further efforts. Our long-term goal is to design the tailor-made production of any moth pheromone in stably transformed plants. However, much remains unknown about the molecular mechanisms underlying the pheromone biosynthesis. The identification of genes encoding pheromone biosynthesis enzymes is a crucial step in ensuring the compatibility of bio-factories. Prior to this disclosure, only FAD and FAR genes have been functionally characterized, but we now have found several functional genes encoding ACOs and ELO that were involved in pheromone biosynthesis. We also developed the plant platform for insect gene functional characterization, contributing an efficient way for further fatty acid metabolic and pheromone biosynthesis studies.

[0141] Ultimately, plant factory for pheromone production could be used in a push-pull strategy for pest control, which is using pheromones that act to make the protected resource unattractive to the pests (push) while luring them toward an attractive source (pull) from where the pests are subsequently removed. This requires that the plants can release pheromones into the atmosphere which is challenging. We intended to accomplish this by using a trichome-specific promoter CYP71D16 to drive pheromone biosynthetic genes. It was surprising to find that this strategy increased the pheromone production amount quite substantially.

[0142] The above description is given by way of example, and not limitation. Given the above disclosure, one skilled in the art could devise variations that are within the scope and spirit of the invention disclosed herein. Further, the various features of the embodiments disclosed herein can be used alone, or in varying combinations with each other and are not intended to be limited to the specific combination described herein. Thus, the scope of the claims is not to be limited by the illustrated embodiments.

REFERENCES

[0143] Agaba M., Tocher D. R., Dickson C. A., Dick J. R., and Teale A. J. (2004). Zebrafish cDNA encoding multifunctional fatty acid elongase involved in production of eicosapentaenoic (20: 5n-3) and docosahexaenoic (22: 6n-3) acids. Marine Biotechnol. 6, 251-261. [0144] Anderson C. M., and Stahl A. (2013). SLC27 fatty acid transport proteins. Mol. Aspects. MED. 34, 516-528. [0145] Ando T., Inomata S. and Yamamoto M. (2004). Lepidopteran Sex Pheromones. Top Curr. Chem. 239, 51-96. [0146] Arsequell G., Fabrias G. and Camps F. (1990). Sex pheromone biosynthesis in the processionary moth Thaumetopoea pityocampa by delta-13 desaturation. Arch. Insect. Biochem. Physiol. 14, 47-56. [0147] Atanassov I. I., Atanassov I. I., Etchells J., and Turner, S. R. (2009). A simple, flexible and efficient PCR-fusion/Gateway cloning procedure for gene fusion, site-directed mutagenesis, short sequence insertion and domain deletions and swaps. Plant methods. 5, 14. [0148] Bansal S., and Durrett T. P. (2016). Camelina sativa: an ideal platform for the metabolic engineering and field production of industrial lipids. Biochimie. 1, 9-16. [0149] Beale M. H., Birkett M. A., Bruce T. J., Chamberlain K., Field L. M., et al. (2006). Aphid alarm pheromone produced by transgenic plants affects aphid and parasitoid behavior. Proc. Nat. Acad. Sci. 103, 10509-10513. [0150] Behrouzian B. and Buist P. H. (2002). Fatty acid desaturation: variations on an oxidative theme. Curr. Opin. Chem. Biol. 6, 577-582. [0151] Bjostad L. B., and Roelofs W. L. (1983). Sex Pheromone Biosynthesis in Trichoplusia ni: Key Steps Involve Delta-11 Desaturation and Chain-Shortening. Science 220, 1387-1389. [0152] Bjostad L. B., and Roelofs W. L. (1984). Sex pheromone biosynthetic precursors in Bombyx mori. Insect biochem. 14, 275-278. [0153] Bjostad L. B., and Roelofs, W. L. (1981). Sex pheromone biosynthesis from radiolabeled fatty acids in the redbanded leafroller moth. J. Biol. Chem. 256, 7936-7940. [0154] Black P. N., and DiRusso C. C. (2007). Vectorial acylation: linking fatty acid transport and activation to metabolic trafficking. In Fatty Acids and Lipotoxicity in Obesity and Diabetes: Novartis Foundation Symposium, John Wiley and Sons, Ltd. 286, 127-141. [0155] Boehm R. (2007). Bioproduction of therapeutic proteins in the 21st century and the role of plants and plant cells as production platforms. Ann. NY. Acad. Sci. 1102, 121-134. [0156] Brittain C., and Potts S. G. (2011). The potential impacts of insecticides on the life-history traits of bees and the consequences for pollination. Basic Appl. Ecol. 12, 321-331. [0157] Buek A., Matoukov P., Vogel H., ebesta P., Jahn U., et al. (2015). Evolution of moth sex pheromone composition by a single amino acid substitution in a fatty acid desaturase. Proc. Nat. Acad. Sci. 112, 12586-12591. [0158] Burton M., Rose T. M., Faergeman N. J., and Knudsen J. (2005). Evolution of the acyl-CoA binding protein (ACBP). Biochem J. 392, 299-307 [0159] Butenandt V. A. (1959). Uber den sexsual-lockstoff des seidenspinners Bombyx mori. Reindarstellung und konstitution. Z. Naturforschg. B. 14, 283. [0160] Cahoon E. B., Shockey J. M., Dietrich C. R., Gidda S. K., Mullen, R. T., et al. (2007). Engineering oilseeds for sustainable production of industrial and nutritional feedstocks: solving bottlenecks in fatty acid flux. Curr. Opin. Plant Biol. 10, 236-244. [0161] Centini F., Masti A., and Comparini I. B. (1996). Quantitative and qualitative analysis of MDMA, MDEA, MA and amphetamine in urine by head-space/solid phase micro-extraction (SPME) and GC/MS. Forensic Sci. Int. 83, 161-166. [0162] Cernac A., and Benning C. (2004). WRINKLED1 encodes an AP2/EREB domain protein involved in the control of storage compound biosynthesis in Arabidopsis. Plant J 40, 575-585. [0163] Chertemps T., Duportets L., Labeur C., Ueda R., Takahashi K., et al. (2007). A female-biased expressed elongase involved in long-chain hydrocarbon biosynthesis and courtship behavior in Drosophila melanogaster. Proc. Nat. Acad. Sci. 104, 4273-4278. [0164] Chertemps T., Duportets L., Labeur C., and Thomas, C. W. (2005). A new elongase selectively expressed in Drosophila male reproductive system. Biochem. Biophys. Res. Commun. 333, 1066-1072 [0165] Christie W. W. (1998). Gas chromatography-mass spectrometry methods for structural analysis of fatty acids. Lipids 33, 343-353. [0166] Clemente T. (2006). Nicotiana (Nicotiana tobaccum, Nicotiana benthamiana). In Agrobacterium protocols. Humana Press. 143-154. [0167] Clinkenbeard K. D., Sugiyama T., Moss J., Reed W. D., and Lane, M. D. (1973). Molecular and catalytic properties of cytosolic acetoacetyl coenzyme A thiolase from avian liver. J. Biol. Chem. 248, 2275-2284. [0168] Cloney R. (2016). Synthetic biology: Automating genetic circuit design. Nat. Rev. Genetics 17, 314. [0169] Conner W. E., Eisner T., Vander Meer R. K., Guerrero A., Ghiringelli D., et al. (1980). Sex attractant of an arctiid moth (Utetheisa ornatrix): a pulsed chemical signal. Behav. Ecol. Sociobiol. 7, 55-63. [0170] Cook S. M., Khan Z. R., and Pickett J. A. (2007). The use of push-pull strategies in integrated pest management. Annu. Rev. Entomol. 52, 375-400. [0171] Dehesh K., Edwards P., Hayes T., Cranmer A. M., and Fillatti J. (1996a). Two novel thioesterases are key determinants of the bimodal distribution of acyl chain length of Cuphea palustris seed oil. Plant Physiol. 110, 203-210. [0172] Ding B. J., Carraher C., and Lfstedt C. (2016). Sequence variation determining stereochemistry of a 11 desaturase active in moth sex pheromone biosynthesis. Insect Biochem. Mol. Biol. 74, 68-75. [0173] Ding B. J., Hofvander P., Wang H. L., Durrett T. P., Stymne S., et al. (2014). A plant factory for moth pheromone production. Nat. commun. 5, 3353. [0174] Ding B. J., Linard M. A., Wang H. L., Zhao C. H., and Lfstedt C. (2011). Terminal fatty-acyl-CoA desaturase involved in sex pheromone biosynthesis in the winter moth (Operophtera brumata). Insect Biochem. Mol. Biol. 41, 715-722. [0175] DiRusso C. C., Li H., Darwis D., Watkins P. A., Berger J., et al. (2005). Comparative biochemical studies of the murine fatty acid transport proteins (FATP) expressed in yeast. J. Biol. Chem. 280, 16829-16837. [0176] Duke S. O. (1994). Glandular trichomes-a focal point of chemical and structural interactions. Intern. J. Plant Sci. 155, 617-620. [0177] Dunkelblum E., Tan S. H., and Silk P. J. (1985). Double-bond location in monounsaturated fatty acids by dimethyl disulfide derivatization and mass spectrometry: Application to analysis of fatty acids in pheromone glands of four lepidoptera. J. Chem. Ecol. 11, 265-277. [0178] Dyer J. M., Stymne S., Green A. G., and Carlsson A. S. (2008). High-value oils from plants. Plant J. 54, 640-655. [0179] Eccleston V. S., and Ohlrogge J. B. (1998). Expression of lauroyl-acyl carrier protein thioesterase in Brassica napus seeds induces pathways for both fatty acid oxidation and biosynthesis and implies a set point for triacylglycerol accumulation. Plant Cell 10, 613-621. [0180] Frgeman N. J., Wadum M., Feddersen S., Burton M., Kragelund B. B., et al. (2007). Acyl-CoA binding proteins; structural and functional conservation over 2000 MYA. Mol. Cellular Biochem. 299, 55-65. [0181] Filichkin S. A., Slabaugh M. B., and Knapp S. J. (2006). New FATB thioesterases from a high laurate Cuphea species: functional and complementation analyses. Eur. J. Lipid Sci. Tech. 108, 979-990. [0182] Foster S. P. (2005). Lipid analysis of the sex pheromone gland of the moth Heliothis virescens. Arch. Insect. Biochem. Physiol. 59, 80-90. [0183] Foster S. P., and Roelofs W. L. (1988). Sex pheromone biosynthesis in the leafroller moth Planotortrix excessana by 10 desaturation. Arch. Insect. Biochem. Physiol. 8, 1-9. [0184] Foster S. P., and Roelofs W. L. (1996). Sex pheromone biosynthesis in the tortricid moth, Ctenopseustis herana (Felder and Rogenhofer). Arch. Insect. Biochem. Physiol. 33, 135-147. [0185] Ghosh S. K., Bhattacharjee A., Jha J. K., Mondal A. K., Maiti M. K., et al. (2007). Characterization and cloning of a stearoyl/oleoyl specific fatty acyl-acyl carrier protein thioesterase from the seeds of Madhuca longifolia (latifolia). Plant Physiol. Biochem. 45, 887-897. [0186] Gibbs W. W. (2004). Synthetic life. Sci. Am. 290, 74-81 Graham S. A. (1998). Revision of Cuphea section Diploptychia (Lythraceae). Syst. Bot. 53, 1-96. [0187] Graham S. A., and Kleiman R. (1992). Composition of seed oils in some Latin American Cuphea (Lythraceae). Ind. Crop. Prod. 1, 31-34. [0188] Green C. D., and Ozguden-Akkoc C. G., Wang Y., Jump D. B., and Olson L. K. (2010). Role of fatty acid elongases in determination of de novo synthesized monounsaturated fatty acid species. J. Lipid Res. 51, 1871-1877. [0189] Grimberg A., Carlsson A. S., Marttila S., Bhalerao R., and Hofvander P., 2015. Transciptional transitions in Nicotiana benthamiana leaves upon induction of oil synthesis by WRINKLED1 homologs from diverse species and tissues. BMC Plant Biol. 15, 192. [0190] Guo Z., and Wagner G. J. (1995). Biosynthesis of labdenediol and sclareol in cell-free extracts from trichomes of Nicotiana glutinosa. Planta 197, 627-632. [0191] Hagstrm . K., Albre J., Tooman L. K., Thirmawithana A. H., Corcoran J., et al. (2014). A novel fatty acyl desaturase from the pheromone glands of Ctenopseustis obliquana and C. herana with specific Z5-desaturase activity on myristic acid. J. Chem. Ecol. 40, 63-70. [0192] Hagstrm . K., Linard M. A., Groot A. T., Hedenstrm E., and Lfstedt C. (2012). Semi-selective fatty acyl reductases from four heliothine moths influence the specific pheromone composition. PLoS One, 7, e37230. [0193] Hagstrm . K., Wang H. L., Linard M. A., Lassance J. M., Johansson, T., et al. (2013). A moth pheromone brewery: production of (Z)-11-hexadecenol by heterologous co-expression of two biosynthetic genes from a noctuid moth in a yeast cell factory. Microb. Cell Fact. 12, 125. [0194] Hao G., Liu W., O'Connor M., and Roelofs W. L. (2002). Acyl-CoA Z9- and Z10-desaturase genes from a New Zealand leafroller moth species, Planotortrix octo. Insect Biochem. Mol. Biol. 32, 961-966. [0195] Hashimoto K., Yoshizawa A. C., Okuda S., Kuma K., Goto S., et al. (2008). The repertoire of desaturases and elongases reveals fatty acid variations in 56 eukaryotic genomes. J. Lipid Res. 49, 183-191. [0196] Hawkins D. J., and Kridl J. C. (1998) Characterization of acyl-ACP thioesterases of mangosteen (Garcinia mangostana) seed and high levels of stearate production in transgenic canola. Plant J. 13, 743-752. [0197] Heppner J. B. (1991). Faunal regions and the diversity of Lepidoptera. Tropical Lepidoptera 2, 1-85. [0198] Horsch R. B., Rogers S. G., and Fraley R. T. (1985) Transgenic plants, Cold Spring Harb. Symp. Quant. Biol. 50, 433-437. [0199] Iskandarov U., Kim H. J., and Cahoon E. B. (2014). Camelina: an emerging oilseed platform for advanced biofuels and bio-based materials. In Plants and bioenergy, McCann M. C., Ed, Springer, New York Press, pp, 131-140. [0200] Jeong H., Nda Z., and Barabsi A. L. (2003). Measuring preferential attachment in evolving networks. Europhys. Lett. 61, 567. [0201] Jones A., Davies H. M., and Voelker T. A. (1995). Palmitoyl-acyl carrier protein (ACP) thioesterase and the evolutionary origin of plant acyl-ACP thioesterases. Plant Cell 7, 359-371. [0202] Jump D. B. (2009). Mammalian fatty acid elongases In Lipidomics, Totowa N. J., Ed, Humana Press, pp, 375-389. [0203] Jurenka R. (2004). The Chemistry of Pheromones and Other Semiochemicals I. In Insect pheromone biosynthesis. Springer Berlin Heidelberg Press, pp, 97-132. [0204] Jurenka R. (1997). Biosynthetic pathway for producing the sex pheromone component (Z,E)-9,12-tetradecadienyl acetate in moths involves a 12 desaturase. Cell. Mol. Life. Sci. 53, 501-505. [0205] Kandra L., and Wagner G. J. (1988). Studies of the site and mode of biosynthesis of tobacco trichome exudate components. Arch. Biochem. Biophysic. 265, 425-432. [0206] Karg S. R., and Kallio P. T. (2009). The production of biopharmaceuticals in plant systems. Biotechnol. Adv. 27, 879-894. [0207] Katzen F. (2007). Gateway recombinational cloning: a biological operating system. Expert. Opin. DrugDis. 2, 571-589. [0208] Kim H. J., Silva J. E., Vu H. S., Mockaitis K., Nam J. W., et al. (2015a). 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. [0209] Knaut J., and Richtler H. (1985). Trends in industrial uses of palm and lauric oils. J Am. Oil Chem. 'Soc. 62, 317-327. [0210] Knipple D. C., Rosenfield C. L., Miller S. J., Liu W., and Tang J., et al. (1998). Cloning and functional expression of a cDNA encoding a pheromone gland-specific acyl-CoA 11-desaturase of the cabbage looper moth, Trichoplusia ni. Proc. Nat. Acad. Sci. 95, 15287-15292. [0211] Knipple D. C., Rosenfield C. L., Nielsen R., You K. M., and Jeong S. E. (2002). Evolution of the integral membrane desaturase gene family in moths and flies. Genetics 162, 1737-1752. [0212] Knutzon D. S., Bleibaum J. L., Nelsen J., Kridl J. C. and Thompson G. A. (1992) Isolation and characterization of 2 safflower oleoyl-acyl carrier protein thioesterase cDNA clones. Plant Physiol. 100, 1751-1758. [0213] Knutzon D. S., Hayes T. R., Wyrick A., Xiong H., Davies H. M., et al. (1999). Lysophosphatidic acid acyltransferase from coconut endosperm mediates the insertion of laurate at the sn-2 position of triacylglycerols in lauric rapeseed oil and can increase total laurate levels. Plant Physiol. 120, 739-746. [0214] Lamprecht I., Schmolz E., and Schricker B. (2008). Pheromones in the life of insects. Eur. Biophys. J. 37, 1253-1260. [0215] Lassance J. M., Groot A. T., Linard M. A., Antony B., Borgwardt C., et al. (2010). Allelic variation in a fatty-acyl reductase gene causes divergence in moth sex pheromones. Nature 466, 486. [0216] Leonard J. M., Slabaugh M. B., and Knapp S. J. (1997). Cuphea wrightii thioesterases have unexpected broad specificities on saturated fatty acids. Plant Mol. Biol. 34, 669-679. [0217] Li-Beisson Y., Shorrosh B., Beisson F., Andersson M. X., Arondel V., et al. (2013). Acyl-lipid metabolism. Arabidopsis Book 11, e0161. [0218] Linard D., Sourrouille C., Gomord V., and Faye L. (2007). Pharming and transgenic plants. Biotech. Ann. Rev. 13, 115-147. [0219] Linard M. A., Hagstrm A. K., Lassance J. M., and Lfstedt C. (2010). Evolution of multicomponent pheromone signals in small ermine moths involves a single fatty-acyl reductase gene. Proc. Nat. Acad. Sci. 107, 10955-10960. [0220] Linard M. A., Lassance J. M., Wang H. L., Zhao C. H., Piskur J., et al. (2010). Elucidation of the sex-pheromone biosynthesis producing 5, 7-dodecadienes in Dendrolimus punctatus (Lepidoptera: Lasiocampidae) reveals 11- and 9-desaturases with unusual catalytic properties. Insect Biochem. Mol. Biol. 40, 440-452. [0221] Liu W., Jiao H., Murray N. C., O'Connor M., and Roelofs W. L. (2002). Gene characterized for membrane desaturase that produces (E)-11 isomers of mono- and diunsaturated fatty acids. Proc. Nat. Acad. Sci. 99, 620-624. [0222] Liu W., Jiao H., O'Connor, M., and Roelofs W. L. (2002). Moth desaturase characterized that produces both Z and E isomers of 11-tetradecenoic acids. Insect Biochem. Mol. Biol. 32, 1489-1495. [0223] Liu W., Ma P. W. K., Marsella-Herrick P., Rosenfield C. L., Knipple D. C., et al. (1999) Cloning and functional expression of a cDNA encoding a metabolic acyl-CoA Delta 9-desaturase of the cabbage looper moth, Trichoplusia ni. Insect Biochem. Mol. Biol. 29, 435-43. [0224] Liu W., Rooney A. P., Xue B., and Roelofs W. L. (2004). Desaturases from the spotted fireworm moth (Choristoneura parallela) shed light on the evolutionary origins of novel moth sex pheromone desaturases. Gene 342, 303-311. [0225] Lfstedt C., and Bengtsson M. (1988). Sex pheromone biosynthesis of (E,E)-8,10-dodecadienol in codling moth Cydia pomonella involves E9 desaturation. J. Chem. Ecol. 14, 903-915. [0226] Lfstedt C., Hansson B. S., Petersson E., Valeur P., and Richards A. (1994). Pheromonal secretions from glands on the 5th abdominal sternite of hydropsychid and rhyacophilid caddisflies (Trichoptera). J Chem. Ecol. 20, 153-170. [0227] Lfstedt C., and Kozlov M. (1997). A phylogenetic analysis of pheromone communication in primitive moths., In Insect pheromone research, Boston, M. A. Ed. Springer Press, pp, 473-489. [0228] Lfstedt C., Wahlberg N., and Millar J. M. (2016). Evolutionary patterns of pheromone diversity in Lepidoptera. In Pheromone communication in moths: evolution, behavior and application, Allison J. D. Ed. University of California Press, pp, 43-82. [0229] 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. [0230] Ma J. K., Chikwamba R., Sparrow P., Fischer R., Mahoney R., et al. (2005). Plant-derived pharmaceuticalsthe road forward. Trends Plant Sci. 10, 580-585. [0231] Ma P. W., Ramaswamy S. B., Blomquist G. J., and Vogt R. G. (2003). Biology and ultrastructure of sex pheromone-producing tissue. In Insect pheromone biochemistry and molecular biology, Blomquist G. Ed, Academic Press, pp, 19-51. [0232] Mandal M. N. A., Santha I. M., Lodha M. L., and Mehta S. L. (2000). Cloning of acyl-acyl carrier protein (ACP) thioesterase gene from Brassica juncea. Biochem. Soc. Trans. 28, 967-969. [0233] Martinez T., Fabrias G., and Camps F. (1990). Sex pheromone biosynthetic pathway in Spodoptera littoralis and its activation by a neurohormone. J. Biol. Chem. 265, 1381-1387. [0234] Matoukov P., Pichov I., and Svato A. (2007). Functional characterization of a desaturase from the tobacco hornworm moth (Manduca sexta) with bifunctional Z11- and 10, 12-desaturase activity. Insect Biochem. Mol. Biol. 37, 601-610. [0235] Mett V., Farrance C. E., Green B. J., and Yusibov V. (2008). Plants as biofactories. Biologicals 36, 354-358. [0236] Michaelis S., and Herskowitz I. (1988). The -factor pheromone of Saccharomyces cerevisiae is essential for mating. Mol. Cellu. Biol. 8, 1309-1318. [0237] Mogensen I. B., Schulenberg H., Hansen H. O., Spener F., and Knudsen J. (1987). A novel acyl-CoA-binding protein from bovine liver. Effect on fatty acid synthesis. Biochemical J. 241, 189-192. [0238] Moser B. R. (2010). Camelina (Camelina sativa L.) oil as a biofuel's feedstock: Golden opportunity or false hope? Lipid Technol. 22, 270-273. [0239] Moto K. I., Suzuki M. G., Hull J. J., Kurata R., Takahashi S., et al. (2004). Involvement of a bifunctional fatty-acyl desaturase in the biosynthesis of the silkmoth, Bombyx mori, sex pheromone. Proc. Nat. Acad. Sci. 101, 8631-8636. [0240] Moto K. I., Yoshiga T., Yamamoto M., Takahashi S., Okano, K., et al. (2003). Pheromone gland-specific fatty-acyl reductase of the silkmoth, Bombyx mori. Proc. Nat. Acad. Sci. 100, 9156-9161. [0241] Murashige T., and Skoog F. (1962). A revised medium for rapid growth and bio assays with tobacco tissue cultures. Physiol. Plant. 15, 473-497. [0242] Nenrov P., ebek P., Macek T., and Svato A. (2004). First semi-synthetic preparation of sex pheromones. Green Chem. 6, 305-307. [0243] Nguyen H. T., Silva J. E., Podicheti R., Macrander J., Yang W., et al. (2013). Camelina seed transcriptome: a tool for meal and oil improvement and translational research. Plant Biotech. J. 11, 759-769. [0244] Oh C. S., Toke D. A., Mandala S., and Martin C. E. (1997). ELO2 and ELO3, Homologues of the Saccharomyces cerevisiae ELO1 Gene, Function in Fatty Acid Elongation and Are Required for Sphingolipid Formation. J. Biol. Chem. 272, 17376-17384 [0245] Ohlrogge J. B., Kuhn D. N., and Stumpf P. K. (1979). Subcellular localization of acyl carrier protein in leaf protoplasts of Spinacia oleracea. Proc. Nat. Acad. Sci. 76, 1194-1198. [0246] Othman A., Lazarus C., Fraser T. and Stobart K. (2000). Cloning of a palmitoyl-acyl carrier protein thioesterase from oil palm. Biochem. Soc. Trans. 28, 619-622. [0247] Percy-Cunningham J. E., and MacDonald J. A. (1987). Biology and Ultrastructure of Sex Pheromone-Producing Glands. In Pheromone biochemistry, Prestwich G. D. Ed, Academic Press, pp, 27-75. [0248] Pollard M. R., Anderson L., Fan C., Hawkins D. J., and Davies H. M. (1991). A specific acyl-ACP thioesterase implicated in medium-chain fatty acid production in immature cotyledons of Umbellularia californica. Arch. Biochem. Biophy. 284, 306-312. [0249] Raguso R. A., and Pellmyr O. (1998). Dynamic headspace analysis of floral volatiles: a comparison of methods. Oikos 81, 238-254. [0250] Raina A. K., Wergin W. P., Murphy C. A., and Erbe E. F. (2000). Structural organization of the sex pheromone gland in Helicoverpa zea in relation to pheromone production and release. Arthropod Struct. Dev. 29, 343-353. [0251] Rasmussen L. E. L., Lee T. D., Zhang A., Roelofs W. L., and Daves G. D. (1997). Purification, identification, concentration and bioactivity of (Z)-7-dodecen-1-yl acetate: sex pheromone of the female Asian elephant, Elephas maximus. Chem. Senses 22, 417-437. [0252] Reddy G. V. P., and Guerrero A. (2010). New pheromones and insect control strategies. In Vitamins and Hormones, Massachusetts L. G. Ed. Elsevier Press, pp, 493-519. [0253] Reynolds K. B., Taylor M. C., Cullerne D. P., Blanchard C. L., Wood C. C., et al. (2017). A reconfigured Kennedy pathway which promotes efficient accumulation of medium-chain fatty acids in leaf oils. Plant Biotechnol. J. 15, 1397-1408. [0254] Rodriguez S., Hao G., Liu W., Pia B., Rooney A., et al. (2004). Expression and evolution of 9 and 11 desaturase genes in the moth Spodoptera littoralis. Insect Biochem. Mol. Biol. 34, 1315-1328 [0255] Rodriguez-Rodriguez M. F., Salas J. J., Garces R., and Martinez-Force E. (2014). Acyl-ACP thioesterases from Camelina sativa: cloning, enzymatic characterization and implication in seed oil fatty acid composition. Phytochemistry 107, 7-15. [0256] Raclot T. (1997). Selective mobilization of fatty acids from white fat cells: evidence for a relationship to the polarity of triacylglycerols. Biochemical J. 322, 483-489. [0257] Roelofs W. L., Liu W., Hao G., Jiao H., Rooney A. P., et al. (2002). Evolution of moth sex pheromones via ancestral genes. Proc. Nat. Acad. Sci. 99, 13621-13626. [0258] Rosenfield C. L., You K. M., Marsella-Herrick P., Roelofs W. L., and Knipple D. C. (2001). Structural and functional conservation and divergence among acyl-CoA desaturases of two noctuid species, the corn earworm, Helicoverpa zea, and the cabbage looper, Trichoplusia ni. Insect Biochem. Mol. Biol. 31, 949-964. [0259] Salas J. J., and Ohlrogge J. B. (2002) Characterization of substrate specificity of plant FatA and FatB acyl-ACP thioesterases. Arch. Biochem. Biophys. 403, 25-34. [0260] Sanchez-Garcia A., Moreno-Perez A. J., Muro-Pastor A. M., Salas J. J., Garces R. et al. (2010) Acyl-ACP thioesterases from castor (Ricinus communis L.): an enzymatic system appropriate for high rates of oil synthesis and accumulation. Phytochemistry 71, 860-869. [0261] Schultz D. J., and Ohlrogge J. B. (2001). Metabolic engineering of fatty acid biosynthesis. In Lipid Biotech. Kuo T. M. Ed, Marcel Dekker Inc. [0262] Schuster S. C. (2007). Next-generation sequencing transforms today's biology. Nat. Methods 5, 16. [0263] Serra M., Pia B., Abad J. L., Camps F., and Fabrias G. (2007). A multifunctional desaturase involved in the biosynthesis of the processionary moth sex pheromone. Proc. Nat. Acad. Sci. 104, 16444-16449. [0264] Serra M., Pia B., Bujons J., Camps F., and Fabrias G. (2006). Biosynthesis of 10, 12-dienoic fatty acids by a bifunctional 11 desaturase in Spodoptera littoralis. Insect Biochem. Mol. Biol. 36, 634-641. [0265] Serrano-Vega M. J., Garces R., and Martinez-Force E. (2005) Cloning, characterization and structural model of a FatA-type thioesterase from sunflower seeds (Helianthus annuus L.). Planta 221, 868-880. [0266] Shanklin J., Whittle E., and Fox B. G. (1994). Eight histidine residues are catalytically essential in a membrane-associated iron enzyme, stearoyl-CoA desaturase, and are conserved in alkane hydroxylase and xylane monooxygenase, Biochemistry 33, 12787-12794. [0267] Simmons G. S., Suckling D. M., Carpenter J. E., Addison M. F., Dyck V. A., et al. (2010). Improved quality management to enhance the efficacy of the sterile insect technique for lepidopteran pests. J. Appl. Entomol. 134, 261-273. [0268] Slabaugh M. B., Leonard J. M., and Knapp S. J. (1998). Condensing enzymes from Cuphea wrightii associated with medium chain fatty acid biosynthesis. Plant J. 13, 611-620. [0269] Stahl S. S. (2004). Palladium oxidase catalysis: selective oxidation of organic chemicals by direct dioxygen-coupled turnover. Angew. Chemie. Intern. Ed. 43, 3400-3420. [0270] Strandh M., Johansson T., Ahrn D., and Lfstedt C. (2008). Transcriptional analysis of the pheromone gland of the turnip moth, Agrotis segetum (Noctuidae), reveals candidate genes involved in pheromone production. Insect. Mol. Biol. 17, 73-85. [0271] Ternes P., Franke S., Zahringer U., Sperling P., and Heinz E. (2002). Identification and characterization of a sphingolipid delta 4-desaturase family. J. Biol. Chem. 277, 25512-25518. [0272] Thelen J. J., and Ohlrogge J. B. (2002). Metabolic engineering of fatty acid biosynthesis in plants. Metab. Eng. 4, 12-21. [0273] Tjellstrm H., Strawsine M., Silva J., Cahoon E. B., and Ohlrogge J. B. (2013). Disruption of plastid acyl:acyl carrier protein synthetases increases medium chain fatty acid accumulation in seeds of transgenic Arabidopsis. FEBS Letters 587, 936-942. [0274] Tocher D. R., Leaver M. J., and Hodgson P. A. (1998). Recent advances in the biochemistry and molecular biology of fatty acyl desaturases. Prog. Lipid Res. 37, 73-117. [0275] Toke D. A., and Martin C. E. (1996). Isolation and characterization of a gene affecting fatty acid elongation in Saccharomyces cerevisiae. J. Biol. Chem. 271, 18413-18422. [0276] Tupec M., Buek A., Valterov I., and Pichov I. (2017). Biotechnological potential of insect fatty acid-modifying enzymes. Zeitschrift Fr Naturforschung C. 72, 387-403. [0277] Tvrdik P., Westerberg R., Silve S., Asadi A., Jakobsson A., et al. (2000). Role of a new mammalian gene family in the biosynthesis of very long chain fatty acids and sphingolipids. J. Cell Biol. 149, 707-718. [0278] van Nieukerken E. J., Kaila L., Kitching I. J., Kristensen N. P., Lees D. C., et al. (2011). Order Lepidoptera Linnaeus, 1758. Zootaxa, 3148, 212-221. [0279] Vanhercke T., El Tahchy A., Liu Q., Zhou X. R., Shrestha P., et al. (2014b). Metabolic engineering of biomass for high energy density: oilseed-like triacylglycerol yields from plant leaves. Plant Biotechnol. J. 12, 231-239. [0280] Visser J. H., and Thiery D. (1986). Effects of feeding experience on the odour-conditioned anemotaxes of Colorado potato beetles. Entomol. Exp Appl. 42, 198-200. [0281] Voelker T., and Kinney A. J. (2001). Variations in the biosynthesis of seed-storage lipids. Ann. Rev. Plant Biol. 52, 335-361. [0282] Voelker T. A., Hayes T. R., Cranmer A. M., Turner J. C., and Davies H. M. (1996). Genetic engineering of a quantitative trait: Metabolic and genetic parameters influencing the accumulation of laurate in rapeseed. Plant J. 9, 229-241. [0283] Voelker T. A., Worrell A. C., Anderson L., Bleibaum J., Fan C., et al. (1992). Fatty acid biosynthesis redirected to medium chains in transgenic oilseed plants. Science 257, 72-74. [0284] Voinnet O., Rivas S., Mestre P., and Baulcombe D. (2003). An enhanced transient expression system in plants based on suppression of gene silencing by the p19 protein of tomato bushy stunt virus. Plant J. 33, 949-956. [0285] Wang E. M., Gan S. S., and Wagner G. J. (2002). Isolation and characterization of the CYP71D16 trichome-specific promoter from Nicotiana tabacum L. J. Exp. Bot. 53, 1891-1897. [0286] Wang H. L., Linard M. A., Zhao C. H., Wang C. Z., and Lfstedt C. (2010). Neofunctionalization in an ancestral insect desaturase lineage led to rare 6 pheromone signals in the Chinese tussah silkworm. Insect Biochem. Mol. Biol. 40, 742-751. [0287] Weatherston I., and Stewart R. (2002). Regulatory issues in the commercial development of pheromones and other semiochemicals. IOBC wprs Bulletin, 25, 1-10. [0288] Wyatt T. D. (2003). Pheromones and animal behaviour: communication by smell and taste. Cambridge University Press. [0289] Xia Y. H., Zhang Y. N., Ding B. J., Wang H. L., and Lfstedt C. (2019). Multi-Functional Desaturases in Two Spodoptera Moths with 11 and 12 Desaturation Activities. J. Chem. Ecol. 45, 378-387. [0290] Xia Y. H., Zhang Y. N., Hou X. Q., Li F., and Dong S. L. (2015). Large number of putative chemoreception and pheromone biosynthesis genes revealed by analyzing transcriptome from ovipositor-pheromone glands of Chilo suppressalis. Sci. Rep. 5, 7888. [0291] Yamaoka R., Taniguchi Y., and Hayashiya K. (1984). Bombykol biosynthesis from deuterium-labeled (Z)-11-hexadecenoic acid. Experientia 40, 80-81. [0292] Young M. D., Wakefield M. J., Smyth G. K., and Oshlack A. (2010). Gene ontology analysis for RNA-seq: accounting for selection bias. Genome Biol. 11, 14. [0293] Zale J., Jung J. H., Kim J. Y., Pathak B., Karan R., et al. (2016). Metabolic engineering of sugarcane to accumulate energy-dense triacylglycerols in vegetative biomass. Plant Biotechnol. J 14, 661-669. [0294] Zhao C., Lfstedt C., and Wang X. (1990). Sex pheromone biosynthesis in the Asian corn borer Ostrinia furnicalis (II): biosynthesis of (E)- and (Z)-12-tetradecenyl acetate involves 14 desaturation. Arch. Insect. Biochem. Physiol. 15, 57-65.