Coexpression Analysis Identifies Two Oxidoreductases Involved in the Biosynthesis of the Monoterpene Acid Moiety of Natural Pyrethrin Insecticides in Tanacetum cinerariifolium^,

A set of dehydrogenases are involved in the synthesis of trans-chrysanthemic acid, the terpene moiety of the natural insecticide pyrethrins.

Abstract

Flowers of Tanacetum cinerariifolium produce a set of compounds known collectively as pyrethrins, which are commercially important pesticides that are strongly toxic to flying insects but not to most vertebrates. A pyrethrin molecule is an ester consisting of either trans-chrysanthemic acid or its modified form, pyrethric acid, and one of three alcohols, jasmolone, pyrethrolone, and cinerolone, that appear to be derived from jasmonic acid. Chrysanthemyl diphosphate synthase (CDS), the first enzyme involved in the synthesis of trans-chrysanthemic acid, was characterized previously and its gene isolated. TcCDS produces free trans-chrysanthemol in addition to trans-chrysanthemyl diphosphate, but the enzymes responsible for the conversion of trans-chrysanthemol to the corresponding aldehyde and then to the acid have not been reported. We used an RNA sequencing-based approach and coexpression correlation analysis to identify several candidate genes encoding putative trans-chrysanthemol and trans-chrysanthemal dehydrogenases. We functionally characterized the proteins encoded by these genes using a combination of in vitro biochemical assays and heterologous expression in planta to demonstrate that TcADH2 encodes an enzyme that oxidizes trans-chrysanthemol to trans-chrysanthemal, while TcALDH1 encodes an enzyme that oxidizes trans-chrysanthemal into trans-chrysanthemic acid. Transient coexpression of TcADH2 and TcALDH1 together with TcCDS in Nicotiana benthamiana leaves results in the production of trans-chrysanthemic acid as well as several other side products. The majority (58%) of trans-chrysanthemic acid was glycosylated or otherwise modified. Overall, these data identify key steps in the biosynthesis of pyrethrins and demonstrate the feasibility of metabolic engineering to produce components of these defense compounds in a heterologous host.

A small group of plants in the Asteraceae family, including Tanacetum cinerariifolium (formerly Chrysanthemum cinerariifolium), make insecticides known collectively as pyrethrins, with the highest levels of production observed in flowers (Casida, 1973; Casida and Quistad, 1995; Crombie, 1995). Due to their effective toxicity against a wide range of insect species, low toxicity to warm-blooded animals, and propensity for degradation by sunlight and oxidation, pyrethrins have been used for pest control since medieval times (Casida and Quistad, 1995; Katsuda, 1999). The commercial production of natural pyrethrins involves drying and grinding the flowers and dissolving the powder in an organic solvent, which can be used directly for spraying. However, the concentration of pyrethrins in the powder is low at about 2% (Casida, 1973). Synthetic pyrethrins, called pyrethroids, which are chemically similar but not identical to natural pyrethrins, are cheaper and used at much higher quantities worldwide. However, these synthetic products are generally less biodegradable and photolabile, persisting in the environment longer than natural pyrethrins and, thus, give rise to the emergence of resistance among insects. In addition, some pyrethroids are toxic to mammals and fish (Katsuda, 2012).

Natural pyrethrins comprise a group of six compounds. The type I pyrethrins (pyrethrin I, jasmolin I, and cinerin I) are esters of the monoterpenoid trans-chrysanthemic acid with one of the three respective fatty acid-derived alcohols pyrethrolone, jasmolone, and cinerolone. The type II pyrethrins (pyrethrin II, jasmolin II, and cinerin II) are esters of pyrethric acid with one of these respective three alcohols. Pyrethric acid is identical to trans-chrysanthemic acid, except that its C8 position has a methylated carboxyl group (Fig. 1A).

Figure 1.

A, Structures of pyrethrins. B, Proposed pathway for the biosynthesis of trans-chrysanthemic acid.

The synthesis of both the acid and alcohol moieties of pyrethrins is not fully understood. With respect to the acid moiety, Rivera et al. (2001) demonstrated that T. cinerariifolium flowers contain the enzyme trans-chrysanthemyl diphosphate synthase (CDS; EC 2.5.1.67), which condenses two dimethyl allyl diphosphate (DMAPP) molecules via what is known as an irregular C1′-2-3 linkage to form mostly trans-chrysanthemyl diphosphate (CDP; Fig. 1B) as well as a small amount of lavandulyl diphosphate. The identities of the enzymes responsible for the subsequent hypothetical steps are not clear. The conversion of CDP to trans-chrysanthemol could be catalyzed by a member of the terpene synthase family similar to those terpene synthases catalyzing the conversion of geranyl diphosphate to the alcohol monoterpenes geraniol and linalool in many species (Chen et al., 2011), or CDP could be hydrolyzed by phosphatases to give trans-chrysanthemol, similar to the conversion of geranyl diphosphate to geraniol in rose (Rosa spp.) flowers (Magnard et al., 2015). However, no such enzymatic activities were reported thus far in T. cinerariifolium. A recent study by Yang et al. (2014) reported that prolonged incubation of Escherichia coli-produced recombinant CDS protein with low concentrations of DMAPP led to the detection of trans-chrysanthemol in addition to CDP, suggesting that CDS can produce trans-chrysanthemol in vivo. Furthermore, the enzymes responsible for the subsequent oxidation reactions of trans-chrysanthemol to trans-chrysanthemal and then to trans-chrysanthemic acid (Fig. 1B) had not yet been reported.

To facilitate the characterization of pyrethrin biosynthesis, a transcriptome assembly of T. cinerariifolium was generated from RNA sequencing (RNAseq) analysis of leaf and flower tissues harvested at different stages of development. Candidate genes for trans-chrysanthemic acid biosynthesis were identified based on coexpression analysis with two previously functionally identified genes in pyrethrin biosynthesis, TcCDS and TcGLIP, the latter being the gene encoding a GDSL-family lipase that combines the acid and alcohol moieties into an ester (Kikuta et al., 2012). This analysis led to the identification of the genes TcADH2 and TcALDH1, encoding two flower-expressed oxidoreductases that respectively catalyze the two sequential oxidation steps from trans-chrysanthemol to trans-chrysanthemic acid. The discovery of these enzymes facilitated the reconstruction of trans-chrysanthemic acid biosynthesis from the precursor DMAPP, both in vitro and in vivo, the latter by transient expression of multiple genes in Nicotiana benthamiana.

RESULTS

Identification of Pyrethrins and Their Terpenoid Precursors in T. cinerariifolium Flowers

The flower heads of T. cinerariifolium consist of a collection of ray florets on the inside and disc florets on the outside, with both types set on a receptacle (Ramirez et al., 2013). The pyrethrin and terpenoid precursor contents in T. cinerariifolium leaves and flowers of different developmental stages (Fig. 2A) were determined by analysis of methyl tert-butyl ether (MTBE)-extracted macerated tissues by gas chromatography-mass spectrometry (GC-MS). As observed previously (Kikuta et al., 2012; Ramirez et al., 2012), pyrethrin content in flowers increased as they matured (Fig. 2B), and leaves contained negligible amounts of pyrethrins compared with the amounts observed in flowers. Also as described previously (Casida, 1973), pyrethrin I was the most abundant pyrethrin (Fig. 2C). In addition to pyrethrins, we also observed trans-chrysanthemol, trans-chrysanthemal, and trans-chrysanthemic acid in floral extracts, and quantitation of trans-chrysanthemic acid indicated that its concentration also increased as the flower matured (Fig. 2, D and E).

Figure 2.

GC-MS analysis of pyrethrins and terpenoids from T. cinerariifolium leaves and flowers at different stages of development. A, Flowers of different stages of development and a leaf of T. cinerariifolium. B, Changes in relative concentrations of pyrethrin I during floral development. Pyrethrin I is the most abundant pyrethrin in the flower, and changes in the concentrations of other pyrethrins follow the same pattern as those of pyrethrin I. C, GC-MS chromatogram of the total ion mode of MTBE extracts from leaves and flowers harvested at stage 4. In each flower/leaf comparison, samples are shown with the same relative y axis scale, but the 7.2- to 15.2-min section is shown at a smaller scale to magnify the peaks. Peaks identified as terpenoids and internal standard (tetradecane) are labeled. D, GC-MS chromatogram (total ion mode) of MTBE extracts from leaves and flowers of different stages of development, showing the trans-chrysanthemic acid levels in each sample. E, Concentrations of trans-chrysanthemic acid in the leaf and in different stages of flowers. Quantification was achieved by normalization of the peaks in D to the tetradecane internal standard and comparison with a standard curve of authentic trans-chrysanthemic acid (n = 3; means ± sd). FW, Fresh weight.

Identification of Candidate Genes Involved in Trans-Chrysanthemic Acid Biosynthesis

To identify the genes encoding the enzymes responsible for the conversion of trans-chrysanthemol to trans-chrysanthemal and trans-chrysanthemal to trans-chrysanthemic acid, transcriptome assemblies were constructed from RNAseq libraries constructed from eight different T. cinerariifolium tissue samples: leaves, flowers at stage 1, flowers at stage 2, flowers at stage 3, ray florets at stage 4, disk florets at stage 4, ray florets at stage 5, and disk florets at stage 5 (Fig. 3A). Interrogating our database set (http://sativa.mcdb.lsa.umich.edu/blast/) for oxidoreductases with plant alcohol dehydrogenase (ADH) and aldehyde dehydrogenase (ALDH) sequences (see “Materials and Methods”) identified 12 transcripts encoding putative alcohol dehydrogenases (named TcADH1 to TcADH12) and three transcripts for putative aldehyde dehydrogenases (named TcALDH1 to TcALDH3). Four of the putative alcohol dehydrogenase genes, TcADH1, TcADH2, TcADH4, and TcADH6, and one putative aldehyde dehydrogenase gene, TcALDH1, showed the highest degree of coexpression with the known genes of pyrethrin biosynthesis, TcCDS or TcGLIP (Fig. 3B; Supplemental Table S1). However, the much lower relative transcript abundance of TcADH4 and TcADH6 compared with that of TcCDS and TcGLIP in flowers of T. cinerariifolium argued against their involvement in trans-chrysanthemic acid biosynthesis.

Figure 3.

Identification of candidate ADH and ALDH genes for trans-chrysanthemic acid biosynthesis. A, Images of T. cinerariifolium flowers of different stages and of leaves from which RNA samples were obtained for RNAseq analysis. B, Average-linkage hierarchical clustering of relative transcript abundance of putative ADHs and ALDHs with TcCDS and TcGLIP based on the number of reads of each transcript in each RNAseq library. The tree and heat map were generated by Cluster 3.0 software (see “Materials and Methods”). C, Verification of levels of expression of TcCDS, TcGLIP, TcADH1, TcADH2, and TcALDH1 by qRT-PCR. Transcript levels are expressed relative to that of GAPDH (glyceraldehyde-3-phosphate dehydrogenase) in each sample (n = 4; means ± sd). **, P < 0.01 and *, P < 0.05. The differences between leaf, stem, and root data points and any flower data points are all significant at P < 0.001.

The expression patterns of TcADH1, TcADH2, and TcALDH1 deduced from the RNAseq read frequencies were confirmed by quantitative reverse transcription (qRT)-PCR together with the transcript levels of TcCDS and TcGLIP (Fig. 3C). The transcripts of all five genes were significantly more abundant in floral tissues compared with leaves, roots, or stems. Generally, transcripts increased initially from floral stage 1 to floral stage 2 and then remained at similar levels or decreased somewhat, particularly in ray florets, a pattern consistent with the previously reported localization of pyrethrin accumulation (Kikuta et al., 2012; Ramirez et al., 2012).

Phylogenetic analysis that included T. cinerariifolium ADHs and other ADHs with assigned functions revealed that TcADH1 and TcADH2 are most closely related to terpene-modifying ADHs, including ADH1 from Artemesia annua, annotated in the National Center for Biotechnology Information (NCBI) as artemisinic alcohol dehydrogenase (accession no. AEI16475) and 8-hydroxygeraniol oxidoreductase from Catharanthus roseus (Miettinen et al., 2014; Fig. 4A). Similarly, TcALDH1 is most closely related (Fig. 4B) to ALDH1 from A. annua, an enzyme that converts artemisinic aldehyde and dihydroartemisinic aldehyde to artemisinic acid and dihydroartemisinic acid, respectively (Teoh et al., 2009). No prediction for organelle targeting was obtained for the inferred TcADH1, TcADH2, and TcALDH1 protein sequences using the TargetP 1.1 Server (http://www.cbs.dtu.dk/services/TargetP/) and WoLF PSORT (https://wolfpsort.hgc.jp/) programs (Nakai and Horton, 1999; Emanuelsson et al., 2007).

Figure 4.

Phylogenetic analysis of candidate T. cinerariifolium dehydrogenases for trans-chrysanthemic acid biosynthesis based on protein sequences. A, Phylogenetic tree for TcADH1 and TcADH2. B, Phylogenetic tree for TcALDH1. The protein sequences from other species are of functionally characterized enzymes whose sequences were identified by BLAST search to be most closely related to the T. cinerariifolium sequences. Phylogenetic analyses were conducted in MEGA7 (Kumar et al., 2016) with the following parameters: multiple sequence alignment with ClustalW, phylogenetic construction with the maximum likelihood method, and bootstrap tests of 1,000 replicates.

TcADH2 Is a Trans-Chrysanthemol Dehydrogenase

Based on results of the coexpression and phylogenetic analyses that identified TcADH1 and TcADH2 as candidates for the conversion of trans-chrysanthemol to trans-chrysanthemal, we tested their activities in vitro. TcADH1 and TcADH2 proteins with a fused N-terminal His₆ tag were expressed in E. coli and purified. To obtain the trans-chrysanthemol substrate, which is not commercially available, for the enzymatic assays, we used recombinant TcCDS to convert DMAPP to CDP, which was hydrolyzed subsequently by the addition of commercial alkaline phosphatase to give trans-chrysanthemol (Fig. 5A). As reported previously (Rivera et al., 2001; Yang et al., 2014), the CDS-catalyzed reaction also produces small amounts of lavandulyl diphosphate, causing the trans-chrysanthemol preparation to contain a small amount of lavandulol (Fig. 5A). Under our assay conditions, we did not detect trans-chrysanthemol production by CDS for up to 24 h (Fig. 5A).

Figure 5.

Gas chromatography analyses of products obtained in in vitro biochemical assays of TcADH2 and TcALDH1. For all assays analyzed, reaction products were extracted with 100 µL of MTBE and run on an Rxi-5Sil column. Tetradecane was used as an internal standard. A, Synthesis of trans-chrysanthemol substrate. Top trace, 1 mm of a commercially available standard of a trans- and cis-chrysanthemol mixture; middle trace, reaction products obtained by incubating 30 μg of recombinant TcCDS with 2.5 mm DMAPP in a 50-μL reaction for 24 h at 30°C; bottom trace, reaction products obtained by incubating the products of the TcCDS-catalyzed condensation of DMAPP with 5 units of alkaline phosphatase (ALP) for 1 h at 37°C. B, In vitro production of trans-chrysanthemol. Reaction products obtained by incubating 0.64 mm trans-chrysanthemol and 1.5 mm NAD⁺ with 5 µL of eluted protein from empty vector (top trace) or 1.25 μg of purified TcADH2 (bottom trace) in a 60-μL reaction volume for 5 min. C, Production of trans-chrysanthemic acid from trans-chrysanthemol in a coupled assay containing 0.64 mm trans-chrysanthemol and 1.5 mm NAD⁺ with 1.25 μg of purified TcADH2 and 6 μg of purified TcALDH1 in a 60-μL reaction volume for 5, 10, 15, 25, and 45 min. A control reaction was performed using 5 µL of eluted protein from the empty vector. The bottom trace shows 0.3 mm of a commercial trans-chrysanthemic acid.

Recombinant TcADH1 and TcADH2 were initially tested with a variety of alcohol substrates at 0.3 mm concentration and 1 mm NAD⁺ or NADP⁺ (Table I). With both TcADH1 and TcADH2, NAD⁺ was a more effective cofactor; with NADP⁺, product yield with each substrate was less than 5% compared with NAD⁺. TcADH1, however, had no activity with trans-chrysanthemol, instead showing its highest level of activity with nerol and reduced activity with geraniol (48%; Table I), both monoterpene compounds. TcADH2 had the highest activity with trans-chrysanthemol, but it also had some activity (70% or less compared with its activity with trans-chrysanthemol) with several other monoterpene alcohols (Table I). GC-MS analysis of the reaction products showed that TcADH2 converted trans-chrysanthemol to trans-chrysanthemal but did not oxidize lavandulol (Fig. 5B). Kinetic analysis revealed that TcADH2 had a K_m value of 236 ± 5.8 µm and a turnover rate of 0.75 ± 0.0032 s⁻¹ for trans-chrysanthemol, while the K_m value for NAD⁺ was 192.6 ± 8.7 µm.

Table I. Relative activities of recombinant TcADH1, TcADH2, and TcALDH1 with selected substrates.

The activities of TcADH1 and TcADH2 were measured with 0.3 mm alcohols and 1 mm NAD⁺. The activities of TcALDH1 were measured with 0.04 mm aldehydes and 1 mm NADP⁺. Data are expressed as relative mean percentages from triplicate independent assays.

Alcohols	TcADH1	TcADH2	Aldehydes	TcALDH1
Cis-3-hexenol	N.D.a	N.D.	Trans-2-heptenal	47
Perillyl alcohol	N.D.	35	Octanal	90
Nerol	100b	42	Dodecanal	47
Geraniol	48	38	Citral	20
(S)-β-Citronellol	8	22	Perillaldehyde	40
Trans,trans-farnesol	N.D.	N.D.	Farnesal	54
Trans-chrysanthemol	N.D.	100c	Trans-chrysanthemal	100d
8-Hydroxygeraniol	N.D.	70	(S)-(−)-Citronellal	84
Benzyl alcohol	N.D.	N.D.	Benzaldehyde	90
Cinnamyl alcohol	N.D.	N.D.	Trans-cinnamaldehyde	8

^aN.D., Not detectable (less than 5% of highest activity).

^b100% relative activity, corresponds to 0.44 μmol min⁻¹ mg⁻¹ citral.

^c100% relative activity, corresponds to 0.5 μmol min⁻¹ mg⁻¹ trans-chrysanthemal.

^d100% relative activity, corresponds to 0.11 μmol min⁻¹ mg⁻¹ trans-chrysanthemic acid.

TcALDH1 Converts Trans-Chrysanthemal to Trans-Chrysanthemic Acid

Based on results of the coexpression and phylogenetic analyses that identified TcALDH1 as a candidate for the conversion of trans-chrysanthemal to trans-chrysanthemic acid, we tested its activity in vitro with trans-chrysanthemal and several other substrates. Trans-chrysanthemal was produced in a coupled enzymatic method employing TcCDS, alkaline phosphatase, and TcADH2 (Fig. 5A; see “Materials and Methods”). N-terminally tagged TcALDH1 purified from E. coli showed highest activity with trans-chrysanthemal substrate but also displayed substantial activity with several additional aliphatic and aromatic substrates (Table I). The enzyme had a K_m value of 4.6 ± 1.8 µm for trans-chrysanthemal when NAD⁺ was provided and a K_m value of 4.4 ± 2.2 µm for trans-chrysanthemal in the presence of NADP⁺ (Table II). Both NAD⁺ and NADP⁺ served as cofactors for the enzyme, with K_m of 20.4 µm for NAD⁺ and 68.6 µm for NADP⁺ (Table II). Incubation of trans-chrysanthemol with both TcADH2 and TcALDH1 led to the production of trans-chrysanthemic acid (Fig. 5C).

Table II. Kinetic properties of recombinant TcADH2 and TcALDH1.

Data are presented as means ± sd from triplicate independent assays. All assays were performed by GC-MS.

Enzyme	Substrate	Km	Kcat	Kcat/Km
		μm	s−1	m−1 s−1
TcADH2	Trans-chrysanthemola	236.0 ± 5.8	0.75 ± 0.0032	3,186.2
	NAD+b	192.6 ± 8.7	0.64 ± 0.0034	3,345.5
TcALDH1	Trans-chrysanthemalc	4.4 ± 2.2	0.11 ± 0.0049	25,122.4
	NADP+d	20.4 ± 7.1	0.090 ± 0.0013	4,391.8
	Trans-chrysanthemale	4. 6 ± 1.8	0.096 ± 0.0032	20,992.3
	NAD+f	68.6 ± 17.1	0.086 ± 0.0012	1,256.07

^aKinetic parameters were determined with 1 mm NAD⁺.

^bKinetic parameters were determined with 0.64 mm trans-chrysanthemol.

^cKinetic parameters were determined with 1 mm NADP⁺.

^dKinetic parameters were determined with 0.04 mm trans-chrysanthemal.

^eKinetic parameters were determined with 1 mm NAD⁺.

^fKinetic parameters were determined with 0.04 mm trans-chrysanthemal.

Transient Coexpression of TcADH2 and TcALDH1 with CDS in N. benthamiana results in Trans-Chrysanthemic Acid Production

To test the activities of TcADH2 and TcALDH1 in planta, N. benthamiana leaves were infiltrated with Agrobacterium tumefaciens strains harboring plasmids containing TcCDS, TcADH2, and TcALDH1. For controls, we infiltrated N. benthamiana leaves with TcCDS alone, TcCDS and TcADH2, or EGFP (Enhanced Green Fluorescent Protein; see “Materials and Methods”) as a control. In these experiments, the complete open reading frame of TcCDS was used, including the transit peptide that was shown to direct the protein to the plastids (Yang et al., 2014). Transformed leaves were harvested 10 d after infiltration, and the products were extracted and analyzed by GC-MS. As expected, transient expression of TcCDS alone resulted in the production of trans-chrysanthemol at 18.3 nmol g⁻¹ fresh weight and trace amounts of trans-chrysanthemic acid (Fig. 6). Notably, coexpression of TcCDS with TcADH2 in N. benthamiana leaves resulted in the production of trans-chrysanthemic acid at 328 nmol g⁻¹ fresh weight (Fig. 6), a 48-fold increase over the expression of TcCDS alone. Finally, when all three genes (TcCDS, TcADH2, and TcALDH1) were coexpressed in N. benthamiana leaves, the concentration of trans-chrysanthemic acid was 818.4 nmol g⁻¹ fresh weight, a 122-fold increase over the expression of TcCDS alone.

Figure 6.

Production of trans-chrysanthemol, trans-chrysanthemic acid, and related compounds in N. benthamiana leaves transiently expressing TcCDS, TcADH2, and TcALDH1 proteins. A, GC-MS chromatograms of MTBE extracts of N. benthamiana leaves expressing EGFP (control), TcCDS, TcCDS and TcADH2, and TcCDS with TcADH2 and TcALDH1. For terpenes, m/z = 123 was monitored, and for the internal control tetradecane, m/z = 198 was monitored. Peaks related to trans-chrysanthemol, trans-chrysanthemic acid, and internal standard are labeled. B and C, Concentrations of free trans-chrysanthemol (B) and free trans-chrysanthemic acid (C) in N. benthamiana leaves expressing the indicated constructs were determined by comparison with an authentic standard. FW, Fresh weight. D to F, Relative levels of Unknown 1 (D), Unknown 2 (E), and Unknown 3 (F) in N. benthamiana leaves expressing the indicated constructs. For each compound, the plant material expressing a specific construct that showed the highest levels (average of three biological replicates) was set at 100%. The data in B to F represent means ± sd from triplicate biological replicates. N.D., Not detected.

In addition to trans-chrysanthemic acid and its precursors, additional volatile derivatives of these metabolites were identified. A peak with a mass fragmentation spectrum similar to trans-chrysanthemol (Supplemental Fig. S1C) was found in N. benthamiana leaves expressing TcCDS; it eluted with a retention time of 30.16 min, compared with a retention time of 10.08 min for trans-chrysanthemol, and was labeled Unknown 1 in Figure 6. Despite the lack of an authentic standard for this compound, which precluded the determination of its actual concentration, we could measure changes in its relative concentrations. When TcCDS-expressing N. benthamiana leaf extract was treated with glycosidase, Unknown 1 was no longer observed (Supplemental Fig. S2A). When the same extract was treated with NaOH, there were no changes in Unknown 1 and trans-chrysanthemol (Supplemental Fig. S2B).

In leaves expressing both TcCDS and TcADH2, Unknown 1 concentration was reduced by 30-fold. In addition, a new compound judged to be related to trans-chrysanthemic acid by its mass fragmentation spectrum and designated Unknown 2 was detected with a retention time of 15.49 min (Fig. 6; Supplemental Fig. S1D). In leaves expressing TcCDS, TcADH2, and TcALDH1, the levels of Unknown 2 increased 7-fold over those observed in leaves expressing TcCDS and TcADH2 without TcALDH1. In addition, leaves expressing all three genes had a new trans-chrysanthemic acid-related compound, designated Unknown 3, with a retention time of 31.98 min (Fig. 6; Supplemental Fig. S1E). Unknown 2 and Unknown 3 were eliminated when the extract was treated with either glycosidase or NaOH (Supplemental Fig. S3, A and B), indicating that they are likely to be esters of trans-chrysanthemic acid. It is notable that the concentration of trans-chrysanthemal was always below detection levels in all N. benthamiana infiltrated leaves, whether expressing TcCDS by itself, TcCDS and TcADH2, or TcCDS, TcADH2, and TcALDH1.

To search for nonvolatile trans-chrysanthemol and trans-chrysanthemic acid conjugates, aliquots of the leaf MTBE extracts from these experiments were dried, dissolved in 70:30 acetonitrile:water (v/v) solution, and analyzed by liquid chromatography (LC)-quadrupole time of flight-mass spectrometry (MS). The sample from leaves expressing TcCDS alone contained a peak with mass-to-charge ratio (m/z) 803.3742 (Fig. 7B). The mass spectrum-mass spectrum (MS/MS) of this metabolite (Fig. 7E) was identical to that of a compound produced in tobacco (Nicotiana tabacum) expressing TcCDS (Yang et al., 2014) that these workers putatively identified as trans-chrysanthemyl malonylglucoside. In samples expressing TcCDS + TcADH2, a peak with m/z 831.3328 was detected with an MS/MS that closely matched the spectrum of another monoterpene acid glucoside, geranyl-6-O-malonyl-β-d-glucopyranoside (Yang et al., 2011). Based on this similarity, the m/z 831.3328 compound is putatively identified as the dimer ion (2M-H) of a trans-chrysanthemic acid malonylglucoside conjugate (Fig. 7, C and F). When TcCDS, TcADH2, and TcALDH1 were coexpressed in N. benthamiana, the level of a product with identical elution time and accurate mass increased ∼150-fold based on peak area compared with TcCDS and TcADH2 expression without TcALDH1 (Fig. 7, C and D). To determine the portion of trans-chrysanthemic acid present as malonylated glucoside in plants simultaneously expressing the three enzymes, we used GC-MS to compare the amount of free trans-chrysanthemic acid in extract hydrolyzed with 0.4 n NaOH at 80°C (which left no detectable trans-chrysanthemic acid malonylglucoside as determined by LC-quadrupole time of flight-MS; Supplemental Fig. S4) with that in nonhydrolyzed extract and found that 58% of the total trans-chrysanthemic acid in this extract was esterified, including with malonylated glucoside (Fig. 7G). Since the concentration of free trans-chrysanthemic acid in these leaves was determined to be 818.4 nmol g⁻¹ fresh weight (Fig. 6C), the total amount of trans-chrysanthemic acid produced in these leaves can be calculated to be 1,946.6 nmol g⁻¹ fresh weight.

Figure 7.

LC-MS analysis of N. benthamiana leaves simultaneously expressing the three enzymes TcCDS, TcADH2, and TcALDH1. A to D, Extracted ion chromatograms of m/z 803.37 and 831.33 are shown for EGFP single expression control (A), TcCDS (B), TcCDS + TcADH2 (C), and TcCDS + TcADH2 + TcALDH1 (D). Chromatograms are all scaled the same, as indicated by the ion current (1.01e7) in the top right corner of each chromatogram. The sample for D was diluted 10-fold compared with the other three samples due to the high concentration of m/z 831.33 in the sample. E and F, MS/MS for ions m/z 803.37 (E) and m/z 831.33 (F) along with the proposed compound structures based on exact mass, fragmentation pattern, and similarity to previously published data. G, Relative levels of trans-chrysanthemic acid in N. benthamiana leaves expressing all TcCDS, TcADH2, and TcALDH1 with or without sodium hydroxide treatment.

DISCUSSION

Coexpression Analysis and Biochemical Assays Indicate That TcADH2 and TcALDH1 Catalyze Reactions in the Trans-Chrysanthemic Acid Biosynthetic Pathway

T. cinerariifolium is an important commercial source of the biodegradable natural pyrethrin insecticides, which are very efficient against flying insects and safe for humans and other vertebrates. It has been shown that pyrethrin biosynthesis begins at early stages in the developing achenes (dry fruits) in the inflorescence and reaches peak accumulation in the mature achene (Ramirez et al., 2012). Our observations on the pattern of accumulation of pyrethrins in the inflorescence (Fig. 2B) were consistent with these previous observations. Based on this information, we proceeded to perform transcriptomic profiling on RNA samples collected from five different stages of developing inflorescences as well as from leaf material. The last two stages of floral development, stages 4 and 5, afforded enough material to do separate analyses on ray florets (flowers on the outside perimeter, which have large petals) and disk florets (flowers on the inside, with small petals).

The transcriptomic profiling data (assembled transcripts and the number of reads from each transcript) were used to perform coexpression cluster analysis (Eisen et al., 1998) to identify candidate ADH and ALDH genes whose expression patterns most resembled that of TcCDS, the gene encoding the key enzyme in the pathway for trans-chrysanthemic acid (Rivera et al., 2001; Yang et al., 2014), as well as that of TcGLIP, encoding the enzyme that forms the final pyrethrin product (Kikuta et al., 2012). Both genes were shown to have peak transcript levels at the earliest developmental stage (stage 2) that could be examined (Ramirez et al., 2012). This analysis identified TcADH1, TcADH2, and TcALDH1 as the strongest candidate enzymes for the synthesis of the terpene moiety of pyrethrins. The results of the qRT-PCR analysis of the levels of transcripts of these three genes, as well as those of TcCDS and TcGLIP (Fig. 3C), were generally consistent with the read frequencies obtained in the transcriptomic profiling experiments (Supplemental Table S1). However, the transcript levels we measured for all five genes do not show a steep decline in later stages of floral development, as was reported previously for TcCDs and TcGLIP (Ramirez et al., 2012), although the discrepancy may be due to differences in the delineation of developmental stages and consequent differences in the actual ages of the materials examined.

Based on the identification of TcADH1, TcADH2, and TcALDH1 as the most likely candidates to be involved in the conversion of trans-chrysanthemol to trans-chrysanthemic acid, we proceeded to produce recombinant proteins from these three genes and test the catalytic activities of these proteins in in vitro assays. TcADH1 exhibited no activity with trans-chrysanthemol, but TcADH2 was able to catalyze the conversion of trans-chrysanthemol to trans-chrysanthemal, with a K_m value of 236 µm for trans-chrysanthemol (Table II). Similar in vitro assays demonstrated that TcALDH1 catalyzes the conversion of trans-chrysanthemal to trans-chrysanthemic acid, with a K_m value of 4.4 µm for trans-chrysanthemal. Transient coexpression of TcADH2 and TcALDH1 together with TcCDS in N. benthamiana leaves resulted in appreciable amounts of trans-chrysanthemic acid produced, 1,946.6 nmol g⁻¹ fresh weight, further demonstrating the ability of TcADH2 and TcALDH1 to catalyze the sequential oxidation of trans-chrysanthemol to trans-chrysanthemic acid.

Chrysanthemic Acid Can Be Made Efficiently by in Planta Heterologous Expression of Only TcCDS, TcADH2, and TcALDH1

Yang et al. (2014) reported that transgenic tobacco plants expressing only TcCDS produce trans-chrysanthemol and trans-chrysanthemyl malonylglucoside. Our analysis of N. benthamiana leaves transiently expressing TcCDS identified the production of these two metabolites as well as a compound by GC-MS that appears to be a modification of trans-chrysanthemol (Unknown 1). When extracts of N. benthamiana leaves transiently expressing TcCDS was treated with glycosidase, both trans-chrysanthemyl malonylglucoside and Unknown 1 disappeared, and the amount of trans-chrysanthemol increased by 35-fold, indicating that most of the trans-chrysanthemol in these leaves is in a glycone form. These results indicate (as also noted by Yang et al. [2014]) that trans-chrysanthemol is produced directly by TcCDS and/or that phosphatases present in the host plant can hydrolyze the two phosphate groups from trans-chrysanthemyl diphosphate to give trans-chrysanthemol. But in addition to trans-chrysanthemol and its derivatives, we also detected trace amounts of trans-chrysanthemic acid in the N. benthamiana leaves transiently expressing TcCDS (Fig. 6), showing that endogenous enzymes found in plant hosts are capable of further modifying the heterologously produced trans-chrysanthemol by additional oxidation reactions to give the corresponding acid. This observation is similar to the observed conversion of the monoterpene alcohol geraniol to geranial and geranic acid in transgenic tomato (Solanum lycopersicum) fruits (Davidovich-Rikanati et al., 2007).

The coexpression of TcCDS with TcADH2, and particularly the coexpression of these two genes with TcALDH1, greatly enhanced the production of trans-chrysanthemic acid in N. benthamiana leaves (Fig. 6). While trans-chrysanthemyl malonylglucoside and Unknown 1 (a modified trans-chrysanthemol) were no longer detected in plants expressing all three genes, the malonylglucoside ester of trans-chrysanthemic acid and two other esters of this acid, Unknown 2 and Unknown 3, were present at a combined concentration exceeding the levels of free trans-chrysanthemic acid by a factor of 1.5. These results indicate that, in addition to dehydrogenases that can act on trans-chrysanthemic acid precursors, N. benthamiana leaves also contain enzymes that can use trans-chrysanthemic acid as a substrate and modify it further. The observation that endogenous enzymes in a plant cell engineered to make compounds not previously present in the cell can react with such new compounds has been made before (Lewinsohn and Gijzen, 2009). For a successful genetic engineering attempt to reconstruct the pyrethrin biosynthetic pathway in a heterologous system, it will be necessary to counteract such nonproductive side reactions.

Biosynthesis of Pyrethric Acid

In three of the six pyrethrins that T. cinerariifolium synthesizes (pyrethrin II, cenerin II, and jasmolin II), pyrethric acid rather than trans-chrysanthemic acid constitutes the acid moiety. Pyrethric acid is identical to trans-chrysanthemic acid except that the C8 position (sometimes referred to as C10) has a methylated carboxyl group and, therefore, is likely derived from trans-chrysanthemic acid by a series of enzymatic reactions involving first the hydroxylation of C8, then two successive oxidations of C8 to give an aldehyde and then a carboxyl group, and finally a carboxylmethylation. The hydroxylation of C8 is likely to be catalyzed by an enzyme of the cytochrome P450 oxidoreductase family. The next two oxidation reactions, however, are equivalent to the reactions catalyzed by TcADH2 and TcALDH1, respectively. Our coexpression analysis identified only TcADH1, TcADH2, and TcALDH1 as likely candidate genes for involvement in the synthesis of the terpene moiety of pyrethrins. TcADH4 and TcADH6 were also identified as potential candidates, but the low levels of their transcripts in pyrethrin-producing tissue made them less likely to be involved in trans-chrysanthemic acid biosynthesis. Therefore, it is possible that the TcADH4 or TcADH6 proteins are involved in trans-chrysanthemic acid biosynthesis. Therefore, it is possible that the TcADH6 protein is involved in the biosynthesis of pyrethric acid by catalyzing the conversion of the C8 alcohol to the C8 aldehyde, as pyrethrins containing the pyrethric acid moiety are much less abundant that those containing trans-chrysanthemic acid. It is also possible that TcADH1, which had no activity with trans-chrysanthemol, catalyzes this reaction, and perhaps even TcADH2 possesses this activity. It is notable that all three of these ADHs are evolutionarily closely related to 8-hydroxygeraniol oxidoreductase from Catharanthus roseus, an enzyme that catalyzes the same C8 alcohol oxidation reaction on another monoterpenoid (Miettinen et al., 2014). Finally, it is possible that TcALDH1 catalyzes the conversion of the C8 aldehyde to give a carboxyl group. The lack of suitable substrates currently prevents the testing of these hypotheses, but the availability of transgenic plants producing some of the precursors in the pathway to pyrethric acid may ameliorate this problem.

MATERIALS AND METHODS

Plant Materials and Chemicals

Various tissues, including flowers, leaves, stems, and roots, were collected from Tanacetum cinerariifolium grown in a greenhouse with a 16/8-h day/night photoperiod. Harvested tissues were flash frozen in liquid nitrogen and stored at −80°C until use. All commercial chemicals were purchased from Sigma-Aldrich with the exception of 8-hydroxygeraniol, which was obtained from Santa Cruz Biotechnology, and trans-chrysanthemol and trans-chrysanthemal.

Trans-chrysanthemol was produced via an enzymatic method by TcCDS plus alkaline phosphatase from DMAPP, and its concentration was calculated according on GC-MS based on the standard curve of a commercial mixture of trans- and cis-chrysanthemol. Trans-chrysanthemal was produced by TcADH2 and NAD⁺ from trans-chrysanthemol, extracted by MTBE from the reaction volume, dried, and dissolved in water.

GC-MS Analysis of Pyrethrins and Terpenoids from T. cinerariifolium Leaves and Flowers of Different Stages of Development

Leaf and flower tissues of different stages of development were cut into small pieces, 100 mg of which was transferred into a tube containing 200 µL of MTBE with 0.01 ng µL⁻¹ tetradecane as an internal standard. The tube was vortexed for 3 min at maximum speed, incubated at room temperature for 2 h, and the MTBE phase was then collected. Sequential extractions with MTBE on test samples indicated that, in the initial extraction, more than 94% of the chrysanthemic acid partitioned into the MTBE phase. The MTBE extracts were analyzed by GC-MS. The measurement of trans-chrysanthemic acid was performed based on the corresponding standard curve.

RNAseq Analysis

RNAseq analysis was done essentially according to Moghe et al. (2017). Total RNA was extracted from leaf and flower parts at different stages of development using the Total RNA Isolation Kit from Omega. The quantity and quality of extracted RNA for sequencing were determined with Qubit (Thermo Fisher Scientific) and Bioanalyzer (Agilent Technologies). Libraries for all eight samples (flowers at stage 1, flowers at stage 2, flowers at stage 3, ray florets at stage 4, disk florets at stage 4, ray florets at stage 5, disk florets at stage 5, and leaf) were made using the KAPA stranded RNAseq library preparation kit and sequenced at the Michigan State University Genomics Core on the Illumina HiSeq 2500 with HiSeq SBS reagents in 2- × 150-bp format. Initial reads have been deposited in the NCBI (bioproject accession no. PRJNA399494).

To assemble the reads, paired end RNAseq reads were trimmed by quality scores using Trimmomatic version 0.32 (Bolger et al., 2014) with the following parameters (PE -threads 4 -phred33 ILLUMINACLIP:all_adapters_combined.fasta:2:30:10 LEADING:20 TRAILING:20 SLIDINGWINDOW:4:20 HEADCROP:10 MINLEN:50). Prior to generating a full-scale RNAseq assembly, we tested the impact of changing the k-mer value for de novo assembly using a set of normalized reads obtained using the insilico_read_normalization.pl function provided in Trinity version 2.2.0 (Haas et al., 2013) with the following options (–seqType fq–JM 100G–max_cov 50–pairs_together–SS_lib_type RF–CPU 10–PARALLEL_STATS–KMER_SIZE 25–max_pct_stdev 200). The assemblies were analyzed for completeness by screening the Metazoan and Eukaryote comparison databases of BUSCO version 1.22 (Simão et al., 2015) using default parameters. This analysis revealed k = 31 to be the most optimal value of k for assembling complete transcripts. This result also was supported by analysis of the length distribution of transcripts using the abyss_fac function in AbySS 1.9.0 (Simpson et al., 2009), with k = 31 producing the longest transcripts (N50 = 1,810 bp). Thus, we performed an assembly of all RNAseq reads with Trinity version 2.2.0 using the following parameters (–seqType fq–KMER_SIZE 31–max_memory 120G–SS_lib_type RF–CPU 16–min_kmer_cov 2). This assembly was used to identify ADH-like and ALDH-like transcripts using BLAST.

The expression levels of all transcripts were estimated using the align_and_estimate_abundance.pl function of the Trinity software using the following parameters (–transcripts Trinity.fasta–prep_reference–left PYR*_R1_*.fastq–right PYR*_R2_*.fastq–est_method RSEM–aln_method bowtie–SS_lib_type RF–thread_count 10–max_ins_size 1000–trinity_mode–seqType fq), which estimated transcript abundance using the RSEM function (Li and Dewey, 2011). These expression values were used to identify transcripts correlated with CDSases and GLIP transcripts across all tissues.

Identifying ADH and ALDH Sequences in the T. cinerariifolium Transcriptome

To find candidate ADH and ALDH genes in our T. cinerariifolium transcriptome database, we queried it on the publicly accessible site (http://sativa.mcdb.lsa.umich.edu/blast/) with various plant ADH and ALDH sequences (e.g. cinnamyl alcohol dehydrogenase from Populous trichocarpa, accession no. ACC63874; geraniol dehydrogenase from Ocimum basilicum, Q2KNL6; benzaldehyde dehydrogenase from Antirrhinum majus, ACM89738; aldehyde dehydrogenase1 from Artemisia annua, ACR61719.1) using the TBLASTN function. We also screened the entire annotated database for the designations alcohol dehydrogenase and aldehyde dehydrogenase and added transcripts identified in this way to our list of candidates.

Coexpression Analysis

The number of reads from each transcript in the various RNAseq analyses of RNA samples from different stages of development and parts of the plant were used to perform cluster analysis using the freely available Cluster 3.0 software program (http://bonsai.hgc.jp/∼mdehoon/software/cluster/software.htm#ctv). This program is based on an algorithm developed by Eisen et al. (1998), and the analysis results in a tree and heat map that matches each gene with another gene whose expression pattern matches best with that of the first one.

qRT-PCR Analysis of TcCDS, TcGLIP, TcADH1, TcADH2, and TcALDH1 Transcript Levels

For real-time RT-PCR analysis of transcripts in different tissues, RNA was isolated using the Total RNA Isolation Kit from Omega with a DNA digestion step. cDNAs were prepared using the High Capacity cDNA Reverse Transcription Kit from Thermo Fisher Scientific following the manufacturer’s instructions. Primer design and real-time PCR were performed following the manufacturer’s instructions. Assays were performed using four independent biological replicates. The relative amounts of transcripts for different genes were normalized to GAPDH transcript levels using LinRegPCR software (http://www.hartfaalcentrum.nl/index.php?main=files&fileName=LinRegPCR.zip&description=LinRegPCR:%20qPCR%20data%20analysis&sub=LinRegPCR). The C_T mean values for GAPDH are shown in Supplemental Figure S5. The statistical assay (unpaired Student’s t test, two-tailed option) was performed via GraphPad Prism software (https://www.graphpad.com/scientific-software/prism/).

Generation, Expression, and Purification of Recombinant TcCDS, TcADH1, TcADH2, and TcALDH1

The open reading frames of TcADH1, TcADH2, and TcALDH1 were obtained by RT-PCR from prepared cDNA of flowers at stage 1 of T. cinerariifolium. The full-length cDNAs were introduced into the expression vector pET28a+ or pHIS8, in each case generating a fusion gene that encoded a tag of His₆ residues at the N terminus for expression in Escherichia coli. To obtain soluble proteins for expression in E. coli, a truncated open reading frame of TcCDS, missing the first 50 codons, was obtained by RT-PCR from prepared cDNA of flowers at stage 1 of T. cinerariifolium. The truncated TcCDS open reading frame was inserted into the pEXP5-CT/TOPO vector following the manufacturer’s instructions, generating a fusion gene that encodes a tag of His₆ residues at the C terminus for expression in E. coli. The purification of recombinant proteins was performed as described previously (Xu et al., 2013). All constructs were transformed into E. coli BL21(+) cells. Transformed E. coli cells were grown in Luria-Bertani medium containing appropriate antibiotics until optical density of the culture at 600 nm reached 0.6, and then recombinant gene expression was induced with the addition of isopropylthio-β-galactoside to a final concentration of 0.15 mm and cells were grown overnight at 16°C. The resulting His-tagged fusion proteins were purified using Ni-NTA affinity columns.

Enzymatic Assays of Recombinant TcCDS

The enzymatic assay for TcCDS broadly followed the protocol described by Rivera et al. (2001) as follows. The reaction was initiated by adding 30 µg of affinity-purified His-tagged enzyme in a final volume of 50 μL of assay buffer (pH 7.5) containing 50 mm Tris-HCl, 2 mm DTT, 5 mm MgCl₂, and 2.5 mm DMAPP. The assay was incubated at 30°C for 24 h. To analyze the production of CDP in this assay by GC-MS, hydrolysis of CDP to trans-chrysanthemol by 5 units of Roche rAPid alkaline phosphatase (Sigma-Aldrich) was preformed following the manufacturer’s instructions at 37°C for 2 h. Reaction products were extracted with 100 µL of MTBE, and the MTBE extract was injected and analyzed by GC-MS.

Enzymatic Assays of Recombinant TcADH1 and TcADH2

Dehydrogenase activity assays were done essentially according to Iijima et al. (2006). To assay the substrate specificity of TcADH1 and TcADH2, reactions were initiated by adding 1.25 µg of affinity-purified His-tagged enzyme in a final volume of 50 μL of assay buffer (pH 8) containing 50 mm Tris-HCl, 2 mm DTT, 1 mm NAD⁺, and 0.3 mm selected alcohols. The assays were incubated at 30°C for 10 min, after which reaction products were extracted with 100 µL of MTBE. The MTBE extract was analyzed by GC-MS for product and remaining substrate.

To determine the kinetic parameters of TcADH2, a similar protocol was followed. The K_m value for NAD⁺ was determined by using 0.64 mm trans-chrysanthemol, whereas the K_m value for trans-chrysanthemol was determined with 1 mm NAD⁺. K_m and k_cat values were calculated from initial rate data by using the hyperbolic regression analysis method in Hyper32 software (version 1.0.0; http://hyper32.software.informer.com/).

Enzymatic Assays of Recombinant TcALDH1

To test the substrate specificity of TcALDH1, reactions were initiated by adding 2.5 µg of affinity-purified His-tagged enzyme in a final volume of 50 μL of assay buffer (pH 8.5) containing 50 mm Tris-HCl, 2 mm DTT, 1 mm NADP⁺, and 40 µm selected aldehydes. The assays were incubated at 30°C for 10 min, after which reaction products were extracted with 100 µL of MTBE. The MTBE extract was analyzed by GC-MS. The reaction rate was calculated according to the decrease of substrate based on the corresponding standard curve.

To determine the kinetic parameters of TcALDH1, a similar protocol was followed. The K_m value for NAD⁺ and NADP⁺ was determined by using 40 µm trans-chrysanthemal, whereas the K_m value for trans-chrysanthemal was determined with 1 mm NAD⁺ or NADP⁺. The reaction rate was calculated according to the production of trans-chrysanthemic acid based on the corresponding standard curve. K_m and k_cat values were calculated as described above.

Plasmid Construction for Transient Expression in N. benthamiana Leaves

The complete open reading frame of EGFP was amplified from pSAT6-EYFP-N1 vector and used as a control gene in this study. The EGFP, TcCDS, TcADH2, and TcALDH1 genes were spliced into pEAQ-HT binary vector between AgeI and XhoI restriction sites using the NEBuilder HiFi DNA Assembly Cloning Kit (https://www.neb.com/products/e5520-nebuilder-hifi-dna-assembly-cloning-kit; NEB) according to the manufacturer’s instructions to express them under the control of the 35S promoter.

Transient Expression in Leaves of N. benthamiana

Agrobacterium tumefaciens strain GV3101 infiltration (agroinfiltration) was performed as described previously (Sainsbury et al., 2009). Briefly, A. tumefaciens strain GV3101 was grown at 28°C at 200 rpm for 24 h in Luria-Bertani medium containing kanamycin (50 mg L⁻¹), rifampicin (50 mg L⁻¹), and gentamycin (25 mg L⁻¹). Cells were collected by centrifugation for 15 min at 3,000g at 20°C and then resuspended in 10 mm MES buffer containing 10 mm MgCl₂ and 100 μm acetosyringone (4′-hydroxy-3′,5′-dimethoxyacetophenone) to a final OD₆₀₀ of 0.4, followed by incubation at 20°C for 3 h. For coinfiltration, equal numbers of cells from each of the cultures of strains harboring different binary plasmids were mixed together, collected by centrifugation, and resuspended as above to a final OD₆₀₀ of 0.4 per strain. N. benthamiana plants were grown from seeds on soil in a greenhouse with a 16/8-h day/night photoperiod at 25°C. Leaves of 4-week-old N. benthamiana plants were infiltrated using a 2-mL syringe without a needle. N. benthamiana plants transformed with the binary vector harboring EGFP alone were used as a control. The infiltrated leaves were collected 10 d after infiltration.

To analyze the compounds produced in the leaves, the harvested plant materials were flash frozen and ground into a fine power in liquid nitrogen. Three grams of powder was extracted with 4 mL of MTBE. The extracts were briefly vortexed for 3 min at maximum speed and then incubated at room temperature with shaking at 50 rpm for 3 h, followed by centrifugation for 15 min at 8,000g. The MTBE layer was transferred to a fresh vial, dehydrated using anhydrous Na₂SO₄, and concentrated by evaporating the solvent to a final volume of about 0.3 mL. Analysis of the samples was performed with an Rxi-5Sil column on a Shimadzu QP-2010 GC-MS system.

GC-MS and LC-MS Analyses

Analytes from 1-µL samples were separated by the Shimadzu QP-2010 GC-MS system equipped with the Rxi-5Sil column (30 m × 0.25 mm × 0.25 μm film thickness; Restek) using helium as the carrier gas at a flow rate of 1.4 mL min⁻¹. The injector was used in split mode at a ratio of 1:2 with the inlet temperature set to 240°C. The initial oven temperature of 50°C was increased after holding for 3 min to 110°C at a rate of 10°C min⁻¹, then increased to 150°C at a rate of 5°C min⁻¹, held for 3 min at 150°C, increased to 300°C at a rate of 10°C min⁻¹, and finally held for 3 min at 300°C. Compounds were identified by comparison of mass spectra and retention times with those of the authentic standards, when available, or with known retention indices and mass fragmentation from the literature and NIST library.

Each MTBE extract (4 mL) was evaporated with the BUCHI Rotavapor and dissolved in 0.5 mL of 70% acetonitrile and 30% water for analysis using a Waters Xevo G2-XS quadrupole time of flight-mass spectrometer interfaced with a Waters Acquity binary solvent manager and 2777c autosampler. Samples (5 µL each) were injected onto an Acquity BEH C18 ultra-performance liquid chromatography column (2.1 × 100 mm, 1.7 μm particle size; Waters) at 40°C. Initial conditions were 0.3 mL min⁻¹ 99% solvent A (water + 0.1% formic acid) and 1% solvent B (acetonitrile). Following injection, solvent B was increased in a linear gradient over 16 min to 99%, followed by a hold at 99% B for 2 min, then return to 99% A at 18.01 min and equilibration for 2 min before starting the next sample. Ions were generated by electrospray ionization in negative-ion mode. Capillary voltage was 2 kV, sample cone voltage was 40 V, and source temperature was 100°C. Desolvation temperature was 350°C, and desolvation gas flow was 600 L h⁻¹. Data were acquired using a data-independent method providing both nonfragmenting and fragmenting conditions for each run, and lock mass correction was performed using a Leu enkephalin standard. MS/MS analysis was performed for selected ions in separate runs.

Hydrolysis of Modified Trans-Chrysanthemol and Trans-Chrysanthemic Acid

Acylglucosides such as chrysanthemic acid glucoside can typically be hydrolyzed by base, but chrysanthemyl glucosides cannot. Some nonspecific glycosidases can hydrolyze both classes of compounds. Therefore, we used both and glycosidase treatments to hydrolyze volatile and nonvolatile derivatives of trans-chrysanthemol and trans-chrysanthemic acid produced in N. benthamiana leaves. Samples were prepared by flash freezing leaves and grinding them into a fine power in liquid nitrogen. For base hydrolysis, homogenates (0.5 g per sample) were mixed completely with 50 µL of 4 n NaOH and incubated at 80°C for 20 min, followed by neutralization with 50 µL of 4 n HCl, and then adding 1 mL of MTBE containing 0.01 ng µL⁻¹ tetradecane as an internal standard and vortexing at maximum speed for 4 min. MTBE extracts were transferred to a fresh vial, dehydrated, and concentrated to a final volume of about 0.2 mL for GC-MS analysis or dried and dissolved in 0.3 mL of 70% acetonitrile and 30% water for LC-MS analysis. For glycosidase treatment, homogenates were treated with 4 mg of β-glucosidase (EC 3.2.1.21; purchased from Sigma-Aldrich) at 37°C for 1.5 h and then extracted and analyzed as described above. GC-MS analysis was used to measure volatile compounds as described above. LC-MS analysis was performed to check for disappearance of the nonvolatile trans-chrysanthemic acid conjugates also as described above.

Accession Numbers

The sequence data used in this study can be obtained from the NCBI with the following GenBank accession numbers: TcADH1, MF497443; TcADH2, MF497444; TcALDH1, MF497445. The bioproject accession number for the RNAseq data is PRJNA399494.

Supplemental Data

The following supplemental materials are available.

• Supplemental Figure S1. Mass spectral analysis of Unknown 1, Unknown 2, and Unknown 3 produced in N. benthamiana leaves expressing TcCDS, TcADH2, and TcALDH1.

• Supplemental Figure S2. GC-MS analysis of hydrolysis assays of tissues of N. benthamiana leaves expressing TcCDS.

• Supplemental Figure S3. GC-MS analysis of hydrolysis assays of tissues of N. benthamiana leaves expressing TcCDS, TcADH2, and TcALDH1.

• Supplemental Figure S4. LC-MS analysis of hydrolysis assays of tissues of N. benthamiana leaves expressing TcCDS, TcADH2, and TcALDH1.

• Supplemental Figure S5. The C_T mean values of TcGAPDH.

• Supplemental Table S1. Transcript abundance for selected candidates in the RNAseq database of leaves and flowers at different stages from T. cinerariifolium.