Identification of dehydrogenases involved in the …...56 pyrethrolone, and cinerolone – that...

1

Coexpression analysis identifies two oxidoreductases involved in the biosynthesis of the 1

monoterpene acid moiety of natural pyrethrin insecticides in Tanacetum cinerariifolium1 2

3

Haiyang Xua, Gaurav D. Mogheb,2, Krystle Wiegert-Riningerc, Anthony L. Schilmillerd, 4

Cornelius S. Barry c, Robert L. Lastb,e, Eran Picherskya3 5

6 aDepartment of Molecular, Cellular, and Developmental Biology, University of Michigan, Ann 7

Arbor, MI, USA. 8 bDepartment of Biochemistry, Michigan State University, East Lansing MI, USA 9 cDepartment of Horticulture, Michigan State University, East Lansing, MI, USA. 10 dMass Spectrometry and Metabolomics Core Facility, Michigan State University, East Lansing, 11

MI, USA 12 eDepartment of Plant Biology, Michigan State University, East Lansing MI, USA 13

14

ORCID IDs: 0000000269540000 (H.X.); 0000-0002-8761-064X (GM); 0000-0001-7069-7120 15

(A.L.S); 0000-0003-4685-0273 (C.S.B.); 0000-0001-6974-9587 (R.L.L.) 0000-0002-4343-1535 16

(E.P.) 17

18 1This work was supported by the National Science Foundation collaborative research grants 19

1565355 to EP and 1565232 to RLL. 20

21 2Present address: Section of Plant Biology, School of Integrative Plant Sciences, Cornell 22

University, Ithaca, NY USA 23

24 3Address correspondence to: [email protected] 25

26

One Sentence Summary: A set of dehydrogenases are involved in the synthesis of trans-27 chrysanthemic acid, the terpene moiety of the natural insecticide pyrethrins. 28 29

Plant Physiology Preview. Published on November 9, 2017, as DOI:10.1104/pp.17.01330

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Running title: biosynthesis of trans-chrysanthemic acid 30

31

Key words: pyrethrins, trans-chrysanthemic acid, Tanacetum cinerariifolium, alcohol 32

dehydrogenase, aldehyde dehydrogenase, oxidoreductase, biosynthetic pathway. 33

34

Contact information for all authors: 35

Haiyang Xu: [email protected] 36

Gaurav Moghe: [email protected] 37

Krystle Wiegert-Rininger: [email protected] 38

Anthony Schilmiller: [email protected] 39

Cornelius S. Barry: [email protected] 40

Robert Last: [email protected] 41

Eran Pichersky: [email protected] 42

43

The author responsible for distribution of materials integral to the findings presented in this 44

article in accordance with the policy described in the Instructions for Authors 45

(www.plantphysiol.org) is: Eran Pichersky ([email protected]). 46

47

H.X, R.L.L and E.P. designed the experiments; H.X., G.M., A.L.S and K.W.R. conducted the 48

experiments; H.X., C.S.B, R.L.L. and E.P wrote the article; all authors edited the article. 49

50



3

ABSTRACT 51

Flowers of Tanacetum cinerariifolium produce a set of compounds known collectively as 52

pyrethrins, which are commercially important pesticides that are strongly toxic to flying insects 53

but not to most vertebrates. A pyrethrin molecule is an ester consisting of either trans-54

chrysanthemic acid or its modified form, pyrethric acid, and one of three alcohols – jasmolone, 55

pyrethrolone, and cinerolone – that appear to be derived from jasmonic acid. Chrysanthemyl 56

diphosphate synthase (CDS), the first enzyme involved in the synthesis of trans-chrysanthemic 57

acid, was previously characterized and its gene isolated. TcCDS produces free trans-58

chrysanthemol, in addition to trans-chrysanthemyl diphosphate, but the enzymes responsible for 59

the conversion of trans-chrysanthemol to the corresponding aldehyde and then to the acid have 60

not been reported. We used an RNAseq-based approach and co-expression correlation analysis to 61

identify several candidate genes encoding putative trans-chrysanthemol and trans-chrysanthemal 62

dehydrogenases. We functionally characterized the proteins encoded by these genes using a 63

combination of in vitro biochemical assays and heterologous expression in planta to demonstrate 64

that TcADH2 encodes an enzyme that oxidizes trans-chrysanthemol to trans-chrysanthemal, 65

while TcALDH1 encodes an enzyme that oxidizes trans-chrysanthemal into trans-chrysanthemic 66

acid. Transient co-expression of TcADH2 and TcALDH1 together with TcCDS in Nicotiana 67

benthamiana leaves results in the production of trans-chrysanthemic acid as well as several other 68

side-products. The majority (58%) of trans-chrysanthemic acid was glycosylated or otherwise 69

modified. Overall, these data identify key steps in the biosynthesis of pyrethrins and demonstrate 70

the feasibility of metabolic engineering to produce components of these defense compounds in a 71

heterologous host. 72

73

74



4

INTRODUCTION 75

76

A small group of plants in the Asteraceae family, including Tanacetum cinerariifolium (formerly 77

Chrysanthemum cinerariifolium), make insecticides known collectively as pyrethrins, with the 78

highest levels of production observed in flowers (Casida, 1973; Casida and Quistad, 1995; 79

Crombie, 1995). Due to their effective toxicity against a wide range of insect species, low 80

toxicity to warm-blooded animals and propensity for degradation by sunlight and oxidation, 81

pyrethrins have been used for pest control since medieval times (Casida and Quistad, 1995; 82

Katsuda, 1999). Commercial production of natural pyrethrins involves drying and grinding the 83

flowers and dissolving the powder in an organic solvent, which can be used directly for spraying. 84

However, the concentration of pyrethrins in the powder is low at about 2% (Casida, 1973). 85

Synthetic pyrethrins, called pyrethroids, which are chemically similar but not identical to natural 86

pyrethrins are cheaper and used at much higher quantities worldwide. However, these synthetic 87

products are generally less biodegradable and photolabile, persisting in the environment longer 88

than natural pyrethrins and thus give rise to the emergence of resistance among insects. In 89

addition, some pyrethroids are toxic to mammals and fish (Katsuda, 2012). 90

Natural pyrethrins comprise a group of six compounds. The type I pyrethrins – pyrethrin 91

I, jasmolin I, and cinerin I – are esters of the monoterpenoid trans-chrysanthemic acid with one 92

of the three respective fatty acid-derived alcohols pyrethrolone, jasmolone, and cinerolone. The 93

type II pyrethrins – pyrethrin II, jasmolin II, and cinerin II – are esters of pyrethric acid with one 94

of these respective three alcohols. Pyrethric acid is identical to trans-chrysanthemic acid, except 95

that its C8 position has a methylated carboxyl group (Figure 1A). 96

The synthesis of both the acid and alcohol moieties of pyrethrins is not fully understood. 97

With respect to the acid moiety, Rivera et al (2001) demonstrated that T. cinerariifolium flowers 98

contain the enzyme trans-chrysanthemyl diphosphate synthase (CDS, EC 2.5.1.67), which 99

condenses two dimethyl allyl diphosphate (DMAPP) molecules via what is known as an irregular 100

C1’-2-3 linkage to form mostly trans-chrysanthemyl diphosphate (CDP) (Figure 1B) as well as a 101

small amount of lavandulyl diphosphate. The enzymes responsible for the subsequent 102

hypothetical steps are unknown. The conversion of CDP to trans-chrysanthemol could be 103

catalyzed by a member of the terpene synthase (TPS) family similar to those TPS catalyzing the 104

conversion of geranyl diphosphate (GPP) to the alcohol monoterpenes geraniol and linalool in 105



5

many species (Chen et al., 2011), or CDP could be hydrolyzed by phosphatases to give trans-106

chrysanthemol, similar to the conversion of GPP to geraniol in rose flowers (Magnard et al., 107

2015). However no such enzymatic activities were reported thus far in T. cinerariifolium. A 108

recent study by Yang et al (2014) reported that prolonged incubation of E. coli–produced 109

recombinant CDS protein with low concentrations of DMAPP led to detection of trans-110

chrysanthemol in addition to CDP, suggesting that CDS can produce trans-chrysanthemol in vivo. 111

However, the enzymes responsible for the subsequent oxidation reactions of trans-112

chrysanthemol to trans-chrysanthemal and then to trans-chrysanthemic acid (Figure 1B) had not 113

yet been reported. 114

To facilitate characterization of pyrethrin biosynthesis, a transcriptome assembly of 115

Tanacetum cinerariifolium was generated from RNAseq analysis of leaf and flower tissues 116

harvested at different stages of development. Candidate genes for trans-chrysanthemic acid 117

biosynthesis were identified based on co-expression analysis with two previously functionally 118

identified genes in pyrethrin biosynthesis, TcCDS and TcGLIP, the latter being the gene 119

encoding a GDSL-family lipase that combines the acid and alcohol moieties into an ester (Kikuta 120



6

et al., 2012). This analysis led to the identification of genes TcADH2 and TcALDH1, encoding 121

two flower-expressed oxidoreductases that respectively catalyze the two sequential oxidation 122

steps from trans-chrysanthemol to trans-chrysanthemic acid. The discovery of these enzymes 123

facilitated the reconstruction of trans-chrysanthemic acid biosynthesis from the precursor 124

DMAPP, both in vitro and in vivo by transient expression of multiple genes in Nicotiana 125

benthamiana. 126

127

128



7

RESULTS 129

130

Identification of pyrethrins and their terpenoid precursors in Tanacetum cinerariifolium 131

flowers 132

133

The flower heads of Tanacetum cinerariifolium consist of a collection of ray florets on 134

the inside and disc florets on the outside, with both types set on a receptacle (Ramirez et al., 135

2013). The pyrethrin and terpenoid precursor content in Tanacetum cinerariifolium leaves and 136

flowers of different developmental stages (Figure 2A) were determined by analysis of methyl 137

tert-butyl ether (MTBE)-extracted macerated tissues by gas chromatography – mass 138

spectrometry (GC-MS). As previously observed (Kikuta et al., 2012; Ramirez et al., 2012), 139

pyrethrin content in flowers increased as they matured (Figure 2B), and leaves contained 140

negligible amounts of pyrethrins compared to the amounts observed in flowers. Also as 141

previously described (Casida, 1973), pyrethrin I was the most abundant pyrethrin (Figure 2C). In 142

addition to pyrethrins, we also observed trans-chrysanthemol, trans-chrysanthemal, and trans-143

chrysanthemic acid in floral extracts, and quantitation of trans-chrysanthemic acid indicated that 144

its concentration also increased as the flower matured (Figure 2D,E). 145

146

Identification of candidate genes involved in trans-chrysanthemic acid biosynthesis 147

148

To identify the genes encoding the enzymes responsible for the conversion of trans-149

chrysanthemol to trans-chrysanthemal and trans-chrysanthemal to trans-chrysanthemic acid, 150

transcriptome assemblies were constructed from RNASeq libraries constructed from eight 151

different T. cinerariifolium tissue samples: leaves, flowers at stage 1, flowers at stage 2, flowers 152

at stage 3, ray florets at stage 4, disk florets at stage 4, ray florets at stage 5 and disk florets at 153

stage 5 (Figure 3A). Interrogating our database set (http://sativa.mcdb.lsa.umich.edu/blast/) for 154

oxidoreductases with plant alcohol dehydrogenase (ADH) and aldehyde dehydrogenase (ALDH) 155

sequences (see Materials and Methods) identified 12 transcripts encoding putative alcohol 156

dehydrogenases (named as TcADH1 ~ TcADH12) and three transcripts for putative aldehyde 157

dehydrogenases (named as TcALDH1 ~ TcALDH3). Four of the putative alcohol dehydrogenase 158

genes, TcADH1, TcADH2, TcADH4 and TcADH6, and one putative aldehyde dehydrogenase 159



8

gene, TcALDH1, showed the highest degree of co-expression with the known genes of pyrethrin 160

biosynthesis TcCDS or TcGLIP (Figure 3B; Supplemental Table S1) . However, the much lower 161



9

relative transcript abundance of TcADH4 and TcADH6 compared with that of TcCDS and 162



10

TcGLIP in flowers of T. cinerariifolium argued against their involvement in trans-chrysanthemic 163

acid biosynthesis. 164

The expression patterns of TcADH1, TcADH2, and TcALDH1 deduced from the RNAseq 165

read frequencies were confirmed by qRT-PCR together with the transcript levels of TcCDS and 166

TcGLIP (Figure 3C). The transcripts of all five genes were significantly more abundant in floral 167

tissues compared with leaves, roots or stems. Generally, transcripts increased initially from floral 168

stage 1 to floral stage 2 and then remained at similar levels or decreased somewhat, particularly 169

in ray florets, a pattern consistent with the previously reported localization of pyrethrin 170

accumulation (Kikuta et al., 2012; Ramirez et al., 2012). 171

Phylogenetic analysis that included T. cinerariifolium ADHs and other ADHs with 172

assigned functions revealed that TcADH1 and TcADH2 are most closely related to terpene 173

modifying ADHs, including ADH1 from Artemesia annua, annotated in NCBI as artemisinic 174

alcohol dehydrogenase (accession AEI16475) and 8-hydroxygeraniol oxidoreductase from 175

Catharanthus roseus (Miettinen et al., 2014) (Figure 4A). Similarly, TcALDH1 is most closely 176

related (Figure 4B) to ALDH1 from Artemesia annua, an enzyme that converts artemisinic 177

aldehyde and dihydroartemisinic aldehyde to artemisinic acid and dihydroartemisinic acid, 178

respectively (Teoh et al., 2009). No prediction of organelle targeting was obtained for the 179

inferred TcADH1, TcADH2, and TcALDH1 protein sequences using the TargetP 1.1 Server 180

(http://www.cbs.dtu.dk/services/TargetP/) and WoLF PSORT (https://wolfpsort.hgc.jp/) 181

programs (Nakai and Horton, 1999; Emanuelsson et al., 2007). 182

183

TcADH2 is a trans-chrysanthemol dehydrogenase 184

185

Based on results of the co-expression and phylogenetic analyses that identified TcADH1 186

and TcADH2 as candidates for the conversion of trans-chrysanthemol to trans-chrysanthemal, 187

we tested their activities in vitro. TcADH1 and TcADH2 proteins with fused N-terminal HIS6 tag 188

were expressed in Escherichia coli and purified. To obtain the trans-chrysanthemol substrate ̶ 189

which is not commercially available ̶ for the enzymatic assays, we used recombinant TcCDS to 190

convert DMAPP to CDP, which was subsequently hydrolyzed by the addition of commercial 191

alkaline phosphatase to give trans-chrysanthemol (Figure 5A). As previously reported (Rivera et 192

al., 2001; Yang et al., 2014), the CDS-catalyzed reaction also produces small amounts of 193



11

lavandulyl diphosphate, causing the trans-chrysanthemol preparation to contain a small amount 194



12

of lavandulol (Figure 5A). Under our assay conditions, we did not detect trans-chrysanthemol 195



13

production by CDS for up to 24 h (Figure 5A). 196

Recombinant TcADH1 and TcADH2 were initially tested with a variety of alcohol 197

substrates at 0.3 mM concentration and 1mM NAD+ or NADP+ (Table 1). With both TcADH1 198

and TcADH2, NAD+ was a more effective co-factor; with NADP+, product yield with each 199

substrate was <5% compared with NAD+. TcADH1, however, had no activity with trans-200

chrysanthemol, instead showing its highest level of activity with nerol and reduced activity with 201

geraniol (48%, Table 1), both monoterpene compounds. TcADH2 had the highest activity with 202

trans-chrysanthemol, but it also had some activity (≤70% compared with its activity with trans-203

chrysanthemol) with several other alcohols (Table 1). GC-MS analysis of reaction products 204

showed that TcADH2 converted trans-chrysanthemol to trans-chrysanthemal, but did not 205

oxidize lavandulol (Figure 5B). Kinetic analysis revealed that TcADH2 had a Km value of 236.0 206

± 5.8 µM and a turnover rate of 0.75 ± 0.0032 s-1 for trans-chrysanthemol while the Km value for 207

NAD+ was 192.6 ± 8.7 µM. 208

209

TcALDH1 converts trans-chrysanthemal to trans-chrysanthemic acid 210

211

Based on results of the co-expression and phylogenetic analyses that identified 212

TcALDH1 as a candidate for the conversion of trans-chrysanthemal to trans-chrysanthemic acid, 213

we tested its activity in vitro with trans-chrysanthemal and several other substrates. trans-214

chrysanthemal was produced in a coupled enzymatic method employing TcCDS, alkaline 215

phosphatase and TcADH2 (Figure 5A and see Materials and Methods). N-terminal tagged 216

TcALDH1was purified from E. coli showed highest activity with trans-chrysanthemal substrate 217

but also displayed substantial activity with several additional aliphatic and aromatic substrates 218

(Table 1). The enzyme had a Km value of 4.6 ± 1.8 µM for trans-chrysanthemal when NAD+ was 219

provided, and a Km value of 4.4 ± 2.2 µM for trans-chrysanthemal in the presence of NADP+ 220

(Table 2). Both NAD+ and NADP+ served as cofactors for the enzyme, with a Km of 20.4 µM for 221

NAD+ and 68.6 µM for NADP+ (Table 2). Incubation of trans-chrysanthemol with both 222

TcADH2 and TcALDH1 led to production of trans-chrysanthemic acid (Figure 5C). 223

224

Transient co-expression of TcADH2 and TcALDH1 with CDS in Nicotiana benthamiana 225

results in trans-chrysanthemic acid production 226



14

227

To test the activities of TcADH2 and TcALDH1 in planta, N. benthamiana leaves were 228

infiltrated with Agrobacterium tumefaciens strains harboring plasmids containing TcCDS, 229

TcADH2 and TcALDH1. For controls, we infiltrated N. benthamiana leaves with TcCDS alone, 230

TcCDS and TcADH2, or EGFP (Enhanced Green Fluorescent Protein, see Materials and 231

Methods) as control. In these experiments, the complete open reading frame of TcCDS was used, 232

including the transit peptide that was shown to direct the protein to the plastids (Yang et al., 233

2014). Transformed leaves were harvested 10 days after infiltration and the products were 234

extracted and analyzed by GC-MS. As expected, transient expression of TcCDS alone resulted in 235

the production of trans-chrysanthemol at 18.3 nmol/g FW and trace amounts of trans-236

chrysanthemic acid (Figure 6). Notably, co-expression of TcCDS with TcADH2 in N. 237

benthamiana leaves resulted in production of trans-chrysanthemic acid at 328.0 nmol/g FW 238

(Figure 6), a 48-fold increase over expression of TcCDS alone. Finally, when all three genes – 239

TcCDS, TcADH2 and TcALDH1 - were co-expressed in N. benthamiana leaves, the 240

concentration of trans-chrysanthemic acid was 818.4 nmol/g FW, a 122-fold increase over 241

expression of TcCDS alone. 242

In addition to trans-chrysanthemic acid and its precursors, additional volatile derivatives 243

of these metabolites were identified. A peak with a mass fragmentation spectrum (MS) similar to 244

trans-chrysanthemol (Supplemental Figure S1C) was found in N. benthamiana leaves expressing 245

TcCDS; it eluted with RT of 30.16 min, compared to a RT of 10.08 for trans-chrysanthemol, and 246

was labeled as “Unknown 1” in Figure 6. Despite the lack of an authentic standard for this 247

compound, which precluded determination of its actual concentration, we could measure changes 248

its relative concentrations. When TcCDS-expressing N. benthamiana leaf extract was treated 249

with glycosidase, Unknown 1 was no longer observed (Supplemental Figure S2A). When the 250

same extract was treated with NaOH, there were no changes in Unknown 1 and trans-251

chrysanthemol (Supplemental Figure S2B). 252

In leaves expressing both TcCDS and TcADH2, “Unknown 1” concentration was reduced 253

by 30-fold. In addition, a new compound judged to be related to trans-chrysanthemic acid by its 254

MS and designated as “Unknown 2” was detected with an RT of 15.49 (Figure 6; Supplemental 255

Figure S1D). In leaves expressing TcCDS, TcADH2 and TcALDH1, the levels of “Unknown 2” 256

increased 7-fold over those observed in leaves expressing TcCDS and TcADH2 without 257



15

TcALDH1. In addition, the leaves expressing all three genes had a new trans-chrysanthemic 258

acid-related compound, designated Unknown 3, with an RT of 31.98 (Figure 6, Supplemental 259

Figure S1E). Unknown 2 and Unknown 3 were eliminated when the extract was treated with 260

either glycosidase or NaOH (Supplemental Figure S3A,B), indicating that they are likely to be 261

esters of trans-chrysanthemic acid. It is notable that the concentration of trans-chrysanthemal 262



16

was always below detection levels in all N. benthamiana infiltrated leaves, whether expressing 263

TcCDS by itself, TcCDS and TcADH2, or TcCDS, TcADH2, and TcALDH1. 264

To search for non-volatile trans-chrysanthemol and trans-chrysanthemic acid conjugates, 265

aliquots of the leaf MTBE extracts from these experiments were dried, dissolved in 70% 266

acetonitrile, and analyzed by LC-qToF-MS. The sample from leaves expressing TcCDS alone 267

contained a peak with m/z 803.3742 (Figure 7B). The mass spectrum-mass spectrum (MS/MS) of 268

this metabolite (Figure 7E) was identical to that of a compound produced in N. tabacum 269

expressing TcCDS (Yang et al., 2014) that these workers putatively identified as trans-270

chrysanthemylmalonylglucoside. In samples expressing TcCDS + TcADH2, a peak with m/z 271

831.3328 was detected with an MS/MS spectrum that closely matched the spectrum of another 272

monoterpene acid glucoside, geranyl-6-O-malonyl-β-D-glucopyranoside (Yang et al., 2011). 273

Based on this similarity, the m/z 831.3328 compound is putatively identified as the dimer ion 274

(2M-H) of a trans-chrysanthemic acid malonylglucoside conjugate (Figure 7C,F). When TcCDS, 275

TcADH2, and TcALDH1 were co-expressed in N. benthamiana, the level of a product with 276

identical elution time and accurate mass increased ~150-fold based on peak area compared to 277

TcCDS and TcADH2 expression without TcALDH1 (Figure 7C,D). To determine the portion of 278

trans-chrysanthemic acid present as malonylated glucoside in plants simultaneously expressing 279

the three enzymes, we used GC-MS to compare the amount of free trans-chrysanthemic acid in 280

extract hydrolyzed with 0.4N NaOH at 80 0C (which left no detectable trans-chrysanthemic acid 281

malonylglucoside as determined by LC-qToF-MS, see Supplemental Figure S4) with that in non-282

hydrolyzed extract, and found that 58.0% of the total trans-chrysanthemic acid in this extract 283

was esterified, including with malonylated glucoside (Figure 7G). Since the concentration of 284

free trans-chrysanthemic acid in these leaves was determined to be 818.4 nmol/g FW (Figure 285

6C), the total amount of trans-chrysanthemic acid produced in these leaves can be calculated to 286

be 1946.6 nmol/g FW. 287

288

289



17

DISCUSSION 290

291

Co-expression analysis and biochemical assays indicate that TcADH2 and TcALDH1 292

catalyze reactions in the trans-chrysanthemic acid biosynthetic pathway 293

294

T. cinerariifolium is an important commercial source of the biodegradable natural 295

pyrethrin insecticides, which are very efficient against flying insects and safe for humans and 296

other vertebrates. It has been shown that pyrethrin biosynthesis begins at early stages in the 297

developing achenes (dry fruits) in the inflorescence and reaches peak accumulation in the mature 298

achene (Ramirez et al., 2012). Our observations on the pattern of accumulation of pyrethrins in 299

the inflorescence (Figure 2B) were consistent with these previous observations. Based on this 300



18

information, we proceeded to perform transcriptomic profiling on RNA samples collected from 301

five different stages of developing inflorescences as well as from leaf material. The last two 302

stages of floral development, stages 4 and 5, afforded enough material to do separate analysis on 303

ray florets (flowers on the outside perimeter, which have large petals) and disk florets (flowers 304

on the inside, with small petals). 305

The transcriptomic profiling data (assembled transcripts and the number of reads from 306

each transcript) were used to perform co-expression cluster analysis (Eisen et al., 1998) to 307

identify candidate ADH and ALDH genes whose expression patterns most resembled that of 308

TcCDS, the gene encoding the key enzyme in the pathway for trans-chrysanthemic acid (Rivera 309

et al., 2001; Yang et al., 2014), as well as that of TcGLIP, encoding the enzyme that forms the 310

final pyrethrin product (Kikuta et al., 2012). Both genes were shown to have peak transcript 311

levels at the earliest developmental stage (stage 2) that could be examined (Ramirez et al., 2012). 312

This analysis identified TcADH1, TcADH2, and TcALDH1 as the strongest candidate enzymes 313

for the synthesis of the terpene moiety of pyrethrins. The results of the qRT-PCR analysis of the 314

levels of transcripts of these three genes, as well as those of TcCDS and TcGLIP (Figure 3C), 315

were generally consistent with the read frequencies obtained in the transcriptomic profiling 316

experiments (Supplemental Table 1). However, the transcript levels we measured for all five 317

gene do not show a steep decline in later stages of floral development as was reported previously 318

for TcCDs and TcGLIP (Ramirez et al., 2012), although the discrepancy may be due to 319

differences in the delineation of developmental stages and consequent differences in the actual 320

ages of the materials examined. 321

Based on the identification of TcADH1, TcADH2, and TcALDH1 as the most likely 322

candidates to be involved in conversion of trans-chrysanthemol to trans-chrysanthemic acid, we 323

proceeded to produce recombinant proteins from these three genes and test the catalytic activities 324

of these proteins in in vitro assays. TcADH1 exhibited no activity with trans-chrysanthemol, but 325

TcADH2 was able to catalyze the conversion of trans-chrysanthemol to trans-chrysanthemal 326

with a Km value of 236 µM for trans-chrysanthemol (Table 2). Similar in vitro assays 327

demonstrated that TcALDH1 catalyzes the conversion of trans-chrysanthemal to trans-328

chrysanthemic acid, with a Km value of 4.4 µM for trans-chrysanthemal. Transient co-expression 329

of TcADH2 and TcALDH1 together with TcCDS in N. benthamiana leaves resulted in 330

appreciable amounts of trans-chrysanthemic acid produced - 1946.6 nmol/g FW - further 331



19

demonstrating the ability of TcADH2 and TcALDH1 to catalyze the sequential oxidation of 332

trans-chrysanthemol to trans-chrysanthemic acid. 333

334

Chrysanthemic acid can be made efficiently by in planta heterologous expression of only 335

TcCDS, TcADH2, and TcALDH1 336

337

Yang et al. (2014) reported that transgenic tobacco (N. tabacum) plants expressing only 338

TcCDS produce trans-chrysanthemol and trans-chrysanthemylmalonylglucoside. Our analysis of 339

N. benthamiana leaves transiently expressing TcCDS identified the production of these two 340

metabolites as well as a compound by GC-MS that appears to be a modification of trans-341

chrysanthemol (Unknown 1). When extracts of N. benthamiana leaves transiently expressing 342

TcCDS was treated with glycosidase, both trans-chrysanthemylmalonylglucoside and Unknown 343

1 disappeared, and the amount of trans-chrysanthemol increased by 35-fold, indicating that most 344

of the trans-chrysanthemol in these leaves is in a glycone form. These results indicate (as also 345

noted by Yang et al., 2014) that trans-chrysanthemol is produced directly by TcCDS, and/or that 346

phosphatases present in the host plant can hydrolyze the two phosphate groups from trans-347

chrysanthemyl diphosphate to give trans-chrysanthemol. But in addition to trans-chrysanthemol 348

and its derivatives, we also detected trace amounts of trans-chrysanthemic acid in the N. 349

benthamiana leaves transiently expressing TcCDS (Figure 6), showing that endogenous 350

enzymes found in plant hosts are capable of further modifying the heterologously produced 351

trans-chrysanthemol by additional oxidation reactions to give the corresponding acid. This 352

observation is similar to the observed conversion of the monoterpene alcohol geraniol to geranial 353

and geranic acid in transgenic tomato fruits (Davidovich-Rikanati et al., 2007). 354

The co-expression of TcCDS with TcADH2, and particularly the coexpression of these 355

two genes with TcALDH1, greatly enhanced the production of trans-chrysanthemic acid in N. 356

benthamiana leaves (Figure 6). While trans-chrysanthemylmalonylglucoside and Unknown 1 (a 357

modified trans-chrysanthemol) were no longer detected in plants expressing all three genes, the 358

malonylglucoside ester of trans-chrysanthemic and two other esters of this acid, Unknown 2 and 359

Unknown 3, were present at a combined concentration exceeding the levels of free trans-360

chrysanthemic acid by a factor of 1.5. These results indicate that in addition to dehydrogenases 361

that can act on trans-chrysanthemic acid precursors, N. benthamiana leaves also contain 362



20

enzymes that can use trans-chrysanthemic acid as a substrate and modify it further. The 363

observation that endogenous enzymes in a plant cell engineered to make compounds not 364

previously present in the cell can react with such new compounds has been made before 365

(Lewinsohn and Gijzen, 2009). For a successful genetic engineering attempt to reconstruct the 366

pyrethrin biosynthetic pathway in a heterologous system, it will be necessary to counteract such 367

non-productive side reactions. 368

369

Biosynthesis of pyrethric acid 370

371

In three of the six pyrethrins that T. cinerariifolium synthesizes (pyrethrin II, cenerin II, 372

and jasmolin II), pyrethric acid rather than trans-chrysanthemic acid constitute the acid moiety. 373

Pyrethric acid is identical to trans-chrysanthemic acid except that the C8 position (sometimes 374

referred to as C10) has a methylated carboxyl group, and is therefore likely derived from trans-375

chrysanthemic acid by a series of enzymatic reactions involving first an hydroxylation of C8, 376

then two successive oxidations of C8 to give an aldehyde and then a carboxyl group, and finally 377

a carboxylmethylation. The hydroxylation of C8 is likely to be catalyzed by an enzyme of the 378

cytochrome P450 oxidoreductase family. The next two oxidations reactions, however, are 379

equivalent to the reactions catalyzed by TcADH2 and TcALDH1, respectively. Our co-380

expression analysis identified only TcADH1, TcADH2 and TcALDH1 as likely candidate genes 381

for involvement in the synthesis of the terpene moiety of pyrethrins. TcADH6 was also identified 382

as potential candidates, but the low levels of its transcripts in pyrethrin-producing tissue made it 383

less likely to be involved in trans-chrysanthemic acid biosynthesis. Therefore, it is possible that 384

the TcADH6 protein is involved in biosynthesis of pyrethric acid by catalyzing the conversion of 385

the C8 alcohol to the C8 aldehyde, as pyrethrins containing the pyrethric acid moiety are much 386

less abundant that those containing trans-chrysanthemic acid. It is also possible that TcADH1, 387

which had no activity with trans-chrysanthemol, catalyzes this reaction, and perhaps even 388

TcADH2 possesses this activity. It is notable that all these three ADHs are evolutionarily closely 389

related to 8-hydroxygeraniol oxidoreductase from Catharanthus roseus, an enzyme that 390

catalyzes the same C8 alcohol oxidation reaction on another monoterpenoid (Miettinen et al., 391

2014). Finally, it is possible that TcALDH1 catalyzes the conversion of the C8 aldehyde to give 392

a carboxyl group. The lack of suitable substrates presently prevents the testing of these 393



21

hypotheses, but the availability of transgenic plants producing some of the precursors in the 394

pathway to pyrethric acid may ameliorate this problem. 395

396

397

MATERIALS AND METHODS 398

399

Plant materials and chemicals 400

401

The various tissues including flowers, leaves, stem, and roots were collected from 402

Tanacetum cinerariifolium grown in a greenhouse with a 16/8 hour day/night photoperiod. 403

Harvested tissues were flash frozen in liquid nitrogen and stored at -80°C until use. All 404

commercial chemicals were purchased from Sigma-Aldrich with the exception of 8-405

hydroxygeraniol which was obtained from Santa Cruz Biotechnology, and trans-chrysanthemol 406

and trans-chrysanthemal. 407

trans-chrysanthemol was produced via a enzymatic method by TcCDS plus alkaline 408

phosphatase (ALP) from DMAPP, and its concentration calculated according on GC-MS based 409

on the standard curve of commercial mixture of trans and cis-chrysanthemol. trans-410

chrysanthemal was produced by TcADH2 and NAD+ from trans-chrysanthemol, extracted by 411

MTBE from the reaction volume, dried, and dissolved in water. 412

413

GC-MS analysis of pyrethrins and terpenoids from Tanacetum cinerariifolium leaves and 414

flowers of different stages of development 415

416

Leaf and flower tissues of different stages of development were cut into small pieces, 100 417

mg of which were transferred into a tube containing 200 µL MTBE with 0.01 ng/µl tetradecane 418

as internal standard. The tube was vortexed for 3 min at maximum speed, incubated at room 419

temperature for 2 hours, and the MTBE phase was then collected. Sequential extractions with 420

MTBE on test samples indicated that in the initial extraction >94% of the chrysanthemic acid 421

partitioned into the MTBE phase. The MTBE extracts were analyzed by GC-MS. The 422

measurement of trans-chrysanthemic acid was preformed based on the corresponding standard 423

curve. 424



22

425

RNAseq analysis 426

427

Total RNA was extracted from leaf and flower parts at different stages of development 428

using the Total RNA Isolation Kit from Omega. The quantity and quality of extracted RNA for 429

sequencing was determined with Qubit (Thermo Fisher Scientific, USA) and Bioanalyzer 430

(Agilent Technologies, Palo Alto, California). Libraries for all 8 samples (flowers at stage 1, 431

flowers at stage 2, flowers at stage 3, ray florets at stage 4, disk florets at stage 4, ray florets at 432

stage 5, disk florets at stage 5, and leaf) were made using the KAPA stranded RNA-seq library 433

preparation kit and sequenced at the Michigan State University Genomics Core on the Illumina 434

HiSeq 2500 with HiSeq SBS reagents in 2x150 bp format. Initial reads have been deposited in 435

NCBI (bioproject accession numbers PRJNA399494). 436

437

To assemble the reads, paired end RNA-seq reads were trimmed by quality scores using 438

Trimmomatic v0.32 (Bolger et al., 2014) with the following parameters (PE -threads 4 -phred33 439

ILLUMINACLIP:all_adapters_combined.fasta:2:30:10 LEADING:20 TRAILING:20 440

SLIDINGWINDOW:4:20 HEADCROP:10 MINLEN:50). Prior to generating a full-scale RNA-441

seq assembly, we tested the impact of changing the kmer value for de novo assembly using a set 442

of normalized reads obtained using insilico_read_normalization.pl function provided in Trinity 443

v2.2.0 (Haas et al., 2013), with the following options (--seqType fq --JM 100G --max_cov 50 --444

pairs_together --SS_lib_type RF --CPU 10 --PARALLEL_STATS --KMER_SIZE 25 --445

max_pct_stdev 200). The assemblies were analyzed for completeness by screening the Metazoan 446

and Eukaryote comparison databases of BUSCO v1.22 (Simao et al., 2015) using default 447

parameters. This analysis revealed k=31 to be the most optimal value of k for assembling 448

complete transcripts. This result was also supported by analysis of the length distribution of 449

transcripts using the abyss_fac function in AbySS 1.9.0 (Simpson et al., 2009), with k=31 450

producing the longest transcripts (N50=1810 bp). Thus, we performed an assembly of all RNA-451

seq reads with Trinity v2.2.0 using the following parameters: (--seqType fq --KMER_SIZE 31 --452

max_memory 120G --SS_lib_type RF --CPU 16 --min_kmer_cov 2). This assembly was used to 453

identify ADH-like and ALDH-like transcripts using BLAST. 454

455



23

The expression levels of all transcripts were estimated using the 456

align_and_estimate_abundance.pl function of the Trinity software, using the following 457

parameters (--transcripts Trinity.fasta --prep_reference --left PYR*_R1_*.fastq --right 458

PYR*_R2_*.fastq --est_method RSEM --aln_method bowtie --SS_lib_type RF --thread_count 10 459

--max_ins_size 1000 --trinity_mode --seqType fq), which estimated transcript abundance using 460

the RSEM function (Li and Dewey, 2011). These expression values were used to identify 461

transcripts correlated with CDSases and GLIP transcripts across all tissues. 462

463 464

Identifying ADH and ALDH sequences in the T. cinerariifolium transcriptome 465

466

To find candidate ADH and ALDH genes in our T. cinerariifolium transcriptome database, we 467

queried it on the publicly accessible site (http://sativa.mcdb.lsa.umich.edu/blast/) with various 468

plant ADH and ALDH sequences (e.g., cinnamyl alcohol dehydrogenase from Populous 469

trichocarpa, accession number ACC63874; geraniol dehydrogenase from Ocimum basilicum, 470

Q2KNL6; benzaldehyde dehydrogenase from Antirrhinum majus, ACM89738; aldehyde 471

dehydrogenase 1 from Artemisia annua, ACR61719.1 ) using the TBLASTN function. We also 472

screened the entire annotated database for the designation “alcohol dehydrogenase” and 473

“aldehyde dehydrogenase” and added transcripts identified in this way to our list of candidates. 474

475

Co-expression analysis 476

477

The number of reads from each transcript in the various RNAseq analyses of RNA samples from 478

different stages of development and parts of the plant were used to perform cluster analysis using 479

the freely available Cluster 3.0 software program 480

(http://bonsai.hgc.jp/~mdehoon/software/cluster/software.htm#ctv). This program is based on an 481

algorithm developed by Eisen et al., (1998), and the analysis results in a tree and heat map that 482

matches each gene with another gene whose expression pattern matches best with that of the first 483

one. 484

485



24

Quantitative RT-PCR analysis of TcCDS, TcGLIP, TcADH1, TcADH2 and TcALDH1 486

transcript Levels 487

488

For real-time RT-PCR analysis of transcripts in different tissues, RNA was isolated using 489

Total RNA Isolation Kit from Omega with a DNA digestion step. cDNAs were prepared using 490

High Capacity cDNA Reverse Transcription Kit from Thermo Fisher Scientific following the 491

manufacturer’s instructions. Primer design and real-time PCR were performed following the 492

manufacturer’s instructions. Assays were performed using four independent biological replicates. 493

The relative amounts of transcripts for different genes were normalized to glyceraldehyde-3-P 494

dehydrogenase (GAPDH) transcript levels using LinRegPCR software 495

(http://www.hartfaalcentrum.nl/index.php?main=files&fileName=LinRegPCR.zip&description=496

LinRegPCR:%20qPCR%20data%20analysis&sub=LinRegPCR). The CT mean values for 497

GAPDH are shown in Supplemental Figure S5. The statistics assay (unpaired t-test, two-tailed 498

option) was performed via software GraphPad Prism (https://www.graphpad.com/scientific-499

software/prism/). 500

501

Generation, expression, and purification of recombinant TcCDS, TcADH1, TcADH2 and 502

TcALDH1 503

504

The open reading frames of TcADH1, TcADH2 and TcALDH1 were obtained by RT-PCR 505

from prepared cDNA of flowers at stage 1 of T. cinerariifolium. The full-length cDNAs were 506

introduced into the expression vector pET28a+ or pHIS8, in each case generating a fusion gene 507

that encoded a “tag” of HIS6 residues at the N-terminus for expression in E. coli. To obtain 508

soluble proteins for expression in Escherichia coli, a truncated open reading frame of TcCDS, 509

missing the first 50 codons, was obtained by RT-PCR from prepared cDNA of flowers at stage 1 510

of T. cinerariifolium. The truncated TcCDS ORF was inserted into the pEXP5-CT/TOPO vector 511

following the manufacturer’s instructions, generating a fusion gene that encodes a “tag” of HIS6 512

residues at the C-terminus for expression in E. coli. The purification of recombinant proteins 513

were performed as described previously (Xu et al., 2013). All constructs were transformed into E. 514

coli BL21(+) cells. Transformed E.coli cells were grown in Luria-Bertani medium containing 515

appropriate antibiotics until optical density of the culture at 600 nm reached 0.6 and then 516



25

recombinant gene expression was induced with the addition of IPTG to a final concentration of 517

0.15 mM and cells grown for overnight at 16°C. The resulting His-tagged fusion proteins were 518

purified using Ni-NTA affinity columns. 519

520

Enzymatic assays of recombinant TcCDS 521

522

The enzymatic assay for TcCDS broadly followed the protocol described by Rivera et al. 523

(2001) as follows: The reaction was initiated by adding 30 µg of affinity-purified His-tagged 524

enzyme in a final volume of 50 μL of assay buffer (pH 7.5) containing 50 mM Tris-HCl, 2 mM 525

DTT, 5 mM MgCl2, and 2.5 mM DMAPP. The assay was incubated at 30°C for 24 hours. To 526

analyze the production of CDP in this assay by GC-MS, hydrolysis of CDP to trans-527

chrysanthemol by 5 units of Roche rAPid alkaline phosphatase (Sigma) was preformed following 528

the manufacturer’s instructions at 37°C for 2 h. Reaction products were extracted with 100 µL 529

MTBE and the MTBE extract was injected and analyzed by GC-MS. 530

531

Enzymatic assays of recombinant TcADH1 and TcADH2 532

533

For assaying of substrate specificity of TcADH1 and TcADH2, reactions were initiated 534

by adding 1.25 µg of affinity-purified His-tagged enzyme in a final volume of 50 μL of assay 535

buffer (pH 8.0) containing 50 mM Tris-HCl, 2 mM DTT, 1 mM NAD+ and 0.3 mM of selected 536

alcohols. The assays were incubated at 30°C for 10 min, after which reaction products were 537

extracted with 100 µL MTBE. The MTBE extract was analyzed by GC-MS for product and 538

remaining substrate. 539

To determine the kinetic parameters of TcADH2, a similar protocol was followed. The Km 540

value for NAD+ was determined by using 0.64 mM trans-chrysanthemol whereas the Km value 541

for trans-chrysanthemol was determined with 1 mM NAD+. Km and kcat values were calculated 542

from initial rate data by using the hyperbolic regression analysis method in Hyper32 software 543

(version 1.0.0, http://hyper32.software.informer.com/). 544

545

Enzymatic assays of recombinant TcALDH1 546

547



26

For testing the substrate specificity of TcALDH1, reactions were initiated by adding 2.5 548

µg of affinity-purified His-tagged enzyme in a final volume of 50 μL of assay buffer (pH 8.5) 549

containing 50 mM Tris-HCl, 2 mM DTT, 1 mM NADP+ and 40 µM selected aldehydes. The 550

assays were incubated at 30°C for 10 min after which reaction products were extracted with 100 551

µL MTBE. The MTBE extract was analyzed by GC-MS. The reaction rate was calculated 552

according to the decrease of substrate based on the corresponding standard curve. 553

To determine the kinetic parameters of TcALDH1, a similar protocol was followed. The 554

Km value for NAD+ and NADP+ was determined by using 40 µM trans-chrysanthemal whereas 555

the Km value for trans-chrysanthemal was determined with 1mM NAD+ or NADP+. The reaction 556

rate was calculated according to production of trans-chrysanthemic acid based on the 557

corresponding standard curve. Km and kcat values were calculated as described above. 558

559

Plasmid construction for transient expression in N. benthamiana leaves 560

561

The complete open reading frame of EGFP was amplified from pSAT6-EYFP-N1 vector 562

and used as control gene in this study. The EGFP, TcCDS, TcADH2 and TcALDH1 genes were 563

spliced into pEAQ-HT binary vector between AgeI and XhoI restriction sites using the 564

NEBuilder HiFi DNA Assembly Cloning Kit (https://www.neb.com/products/e5520-nebuilder-565

hifi-dna-assembly-cloning-kit, NEB) according to manufacturer’s instructions to express them 566

under the control of 35S promoter. 567

568

Transient expression in leaves of N. benthamiana 569

570

Agrobacterium tumefaciens strain GV3101 infiltration (agro-infiltration) was performed 571

as described previously (Sainsbury et al., 2009). Briefly, A. tumefaciens strain GV3101 was 572

grown at 28°C at 200 rpm for 24 hours in LB medium containing kanamycin (50 mg L-1), 573

rifampicin (50 mg L-1) and gentamycin (25 mg L-1). Cells were collected by centrifugation for 15 574

min at 3000 g at 20°C, and then resuspended in 10 mM 2-(N-morpholino)ethanesulfonic acid 575

(MES) buffer containing 10 mM MgCl2 and 100 μM acetosyringone (4’-hydroxy-3’,5’-576

dimethoxyacetophenone) to a final OD600 of 0.4, followed by incubation at 20°C for 3 hours. For 577

co-infiltration, equal number of cells from each of the cultures of strains harboring different 578



27

binary plasmids were mixed together, collected by centrifugation, and resuspended as above to a 579

final OD600 of 0.4 per strain. N. benthamiana plants were grown from seeds on soil in a 580

greenhouse with 16/8 hours day/night photoperiod at 25°C. Leaves of 4-week-old N. 581

benthamiana plants were infiltrated using a 2 ml syringe without a needle. N. benthamiana plants 582

transformed with the binary vector harboring EGFP alone were used as control. The infiltrated 583

leaves were collected 10 days after infiltration. 584

To analyze the compounds produced in the leaves, the harvested plant materials were flash-585

frozen and ground into a fine power in liquid nitrogen. Three grams of powder was extracted 586

with 4 ml MTBE. The extracts were briefly vortexed for 3 min at maximum speed and then 587

incubated at room temperature with shaking at 50 rpm for 3 h, followed by centrifugation for 15 588

min at 8000 g. The MTBE layer was transferred to a fresh vial, dehydrated using anhydrous 589

Na2SO4 and concentrated by evaporating the solvent to a final volume of about 0.3 ml. Analysis 590

of the samples was performed with an Rxi-5Sil column on a Shimadzu QP-2010 GC-MS system. 591

592

GC-MS and LC-MS analyses 593

594

Analytes from 1 µl samples were separated by Shimadzu QP-2010 GC-MS system 595

equipped with the Rxi-5Sil column (30 m × 0.25 mm × 0.25 μm film thickness, RESTEK, USA) 596

using helium as the carrier gas at a flow rate of 1.4 ml min-1. The injector was used in split mode 597

at ratio of 1:2 with the inlet temperature set to 240°C. The initial oven temperature of 50°C was 598

increased after holding for 3 min to 110°C at a rate of 10°C min-1, then increased to 150 °C at a 599

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 600

finally held for 3 min at 300°C. Compounds were identified by comparison of mass spectra and 601

retention time with those of the authentic standards, when available, or with known retention 602

indices and mass fragmentation from the literature and NIST library. 603

Each MTBE extract (4 mL) was evaporated by BUCHI Rotavapor and dissolved in 0.5 604

ml of 70% acetonitrile/30% water for analysis using a Waters Xevo G2-XS Q-Tof mass 605

spectrometer interfaced with a Waters Acquity binary solvent manager and 2777c autosampler. 606

Samples (5 µL each) were injected onto an Acquity BEH C18 UPLC column (2.1 x 100 mm, 1.7 607

μm particle size; Waters Corp) at 40°C. Initial conditions were 0.3 ml/min of 99% solvent A 608

(water + 0.1% formic acid) and 1% solvent B (acetonitrile). Following injection, solvent B was 609



28

increased in a linear gradient over 16 min to 99%, followed by a hold at 99% B for 2 min, then 610

return to 99% A at 18.01 min and equilibrate for 2 min before starting the next sample. Ions were 611

generated by electrospray ionization in negative-ion mode. Capillary voltage was 2.0 kV, sample 612

cone voltage was 40 V and source temperature was 100°C. Desolvation temperature was 350°C 613

and desolvation gas flow was 600 L/hr. Data were acquired using a data-independent MSe 614

method providing both non-fragmenting and fragmenting conditions for each run, and lock mass 615

correction was performed using a leucine enkephalin standard. MS/MS was performed for 616

selected ions in separate runs. 617

618

Hydrolysis of modified trans-chrysanthemol and trans-chrysanthemic acid 619

620

Acylglucosides such as chrysanthemic acid glucoside can typically be hydrolyzed by 621

base, but chrysanthemyl glucosides cannot. Some non-specific glycosidases can hydrolyze both 622

classes of compounds. We therefore used both and glycosidase treatments to hydrolyze volatile 623

and non-volatile derivatives of trans-chrysanthemol and trans-chrysanthemic acid produced in N. 624

benthamiana leaves. Samples were prepared by flash-freezing leaves and grounding them into a 625

fine power in liquid nitrogen. For base hydrolysis, homogenates (0.5 g per sample) were mixed 626

completely with 50 µL 4N NaOH and incubated at 80°C for 20 min, followed by neutralization 627

with 50 µL 4N HCl, and then adding 1 mL MTBE containing 0.01 ng/µl tetradecane as internal 628

standard and vortexing at maximum speed for 4 min. MTBE extracts were transferred to a fresh 629

vial, dehydrated and concentrated to a final volume of about 0.2 mL for GC-MS analysis, or 630

dried and dissolved in 0.3 ml of 70% acetonitrile/30% water for LC-MS analysis. For 631

glycosidase treatment, homogenates were treated with 4 mg of β-glucosidase (EC 3.2.1.21, 632

purchased from Sigma) at 37°C for 1.5 hour, and then extracted and analyzed as described above. 633

GC-MS analysis was used to measure volatile compounds as described above. LC-MS analysis 634

was performed to check for disappearance of the non-volatile trans-chrysanthemic acid 635

conjugates also as described above. 636

637

Accession Numbers 638

639



29

The sequence data used in this study can be obtained from NCBI with the following 640

GenBank accession numbers: TcADH1 – MF497443; TcADH2 – MF497444; TcALDH1 – 641

MF497445. The bioproject accession numbers for the RNAseq data is PRJNA399494 642

643

Supplemental Data 644

645

The following supplemental materials are available. 646

647

Supplemental Figure S1. Mass spectral analysis of Unknown 1, Unknown 2 and Unknown 648

3 produced in N. benthamiana leaves expressing TcCDS, TcADH2 and TcALDH1. 649

Supplemental Figure S2. GC-MS analysis of hydrolysis assays of tissues of N. benthamiana 650

leaves expressing TcCDS. 651

Supplemental Figure S3. GC-MS analysis of hydrolysis assays of tissues of N. benthamiana 652

leaves expressing TcCDS, TcADH2 and TcALDH1. 653

Supplemental Figure S4. LC-MS analysis of hydrolysis assays of tissues of N. benthamiana 654

leaves expressing TcCDS, TcADH2 and TcALDH1. 655

Supplemental Figure S5. The CT mean values of TcGAPDH. 656

Supplemental Table S1. Transcript Abundance for Selected Candidate in the RNASeq 657

database of leaves and flowers at different stages from Tanacetum cinerariifolium. 658

659

ACKNOWLEDGMENTS 660

661

We thank Prof. Daniel Jones (Michigan State University) for help with the analysis of 662

non-volatiles compounds in N. benthamiana leaves transiently expressing various T. 663

cinerariifolium genes. 664



30

665

666

667

668

669

670



31

Figure Legends: 671

672

Figure 1. A. Structures of pyrethrins. B. Proposed pathway for the biosynthesis of trans-673

chrysanthemic acid. 674

675

Figure 2. GC-MS analysis of pyrethrins and terpenoids from Tanacetum cinerariifolium leaves 676

and flowers at different stages of development. 677

A. Flowers of different stages of development and a leaf of T. cinerariifolium. 678

B. Changes in relative concentrations of pyrethrin I during floral development. Pyrethrin I is the 679

most abundant pyrtherin in the flower, and changes in concentrations of other pyrethrins follow 680

the same pattern as those of pyrethrin I. 681

C. GC-MS chromatogram of total ion mode of MTBE extracts from leaves and flowers harvested 682

at stage 4. In each flower/leaf comparison, samples are shown with the same relative y-axis scale, 683

but the 7.2-15.2 min section is shown at a smaller scale to magnify the peaks. Peaks identified as 684

terpenoids and internal standard (tetradecane) are labeled. 685

D. GC-MS chromatogram (total ion mode) of MTBE extracts from leaves and flowers of 686

different stages of development, showing the trans-chrysanthemic acid levels in each sample. 687

E. Concentrations of trans-chrysanthemic acid in the leaf and in different stages of flowers. 688

Quantification was achieved by normalization of the peaks in (D) to the tetradecane internal 689

standard and comparison to a standard curve of authentic trans-chrysanthemic acid (n = 3; means 690

± SD). 691

692

Figure 3. Identification of candidate ADH and ALDH genes for trans-chrysanthemic acid 693

biosynthesis. 694

A. Images of T. cinerariifolium flowers of different stages and of leaves from which RNA 695

samples were obtained for RNAseq analysis. 696

B. Average-linkage hierarchical clustering of relative transcript abundance of putative ADHs and 697

ALDHs with TcCDS and TcGLIP based on number of reads of each transcript in each RNASeq 698

library. Tree and heat map generated by the Cluster 3.0 software (see Materials and Methods). 699

C. Verification of levels of expression of TcCDS, TcGLIP, TcADH1, TcADH2, and TcALDH1 by 700

qRT-PCR. Transcript levels are expressed relative to that of GAPDH (glyceraldehyde-3-701



32

phosphate dehydrogenase) in each sample (n = 4; means ± SD). ** p<0.01, * p<0.05. The 702

differences between leaf, stem and root datapoints and any flower datapoints are all significant at 703

p<0.001. 704

705

Figure 4. Phylogenetic analysis of candidate T. cinerariifolium dehydrogenases for trans-706

chrysanthemic acid biosynthesis based on protein sequences. 707

A. Phylogenetic tree for TcADH1 and TcADH2. 708

B. Phylogenetic tree for TcALDH1. 709

The protein sequences from other species are of functionally characterized enzymes whose 710

sequences were identified by BLAST search to be most closely related to the T. cinerariifolium 711

sequences. Phylogenetic analyses were conducted in MEGA7 (Kumar et al., 2016) with the 712

following parameters; multiple sequence alignment with ClustalW, phylogenetic construction 713

with the Maximum Likelihood method and bootstrap tests of 1000 replicates. 714

715

Figure 5. Gas chromatography analyses of products obtained in in vitro biochemical assays of 716

TcADH2 and TcALDH1. For all assays analyzed, reactions products were extracted with 100 µl 717

MTBE and run on Rxi-5Sil column. Tetradecane was used as internal standard. 718

A. Synthesis of trans-chrysanthemol substrate. Top chromatograph trace: 1 mM of a 719

commercially available standard of trans- and cis-chrysanthemol mixture. Middle trace: 720

Reaction products obtained by incubating 30 μg recombinant TcCDS with 2.5 mM DMAPP in 721

50 μL reaction for 24 hours at 30°C. Bottom chromatograph trace: Reaction products obtained 722

by incubating the products of the TcCDS-catalyzed condensation of DMAPP with 5 units of 723

alkaline phosphatase (ALP) for 1 hour at 37°C. 724

B. In vitro production of trans-chrysanthemol. Reaction products obtained by incubating 0.64 725

mM trans-chrysanthemol and 1.5 mM NAD+ with 5 µl eluted protein from empty vector (top 726

chromatograph trace) or 1.25 μg purified TcADH2 (bottom chromatograph trace) in 60 μL 727

reaction volume for 5 min. 728

C. Production of trans-chrysanthemic acid from trans-chrysanthemol in a coupled assay 729

containing 0.64 mM trans-chrysanthemol and 1.5 mM NAD+ with 1.25 μg purified TcADH2 and 730

6.00 μg purified TcALDH1 in 60 μL reaction volume for 5, 10, 15, 25, and 45 min. A control 731



33

reaction was performed using 5µl eluted protein from empty vector. Bottom chromatograph trace: 732

0.3 mM of commercial trans-chrysanthemic acid. 733

734

Figure 6. Production of trans-chrysanthemol, trans-chrysanthemic acid, and related compounds 735

in N. benthamiana leaves transiently expressing TcCDS, TcADH2, and TcALDH1 proteins. 736

A. GC-MS chromatograms of MTBE extracts of N. benthamiana leaves expressing EGFP 737

(control), TcCDS, TcCDS and TcADH2, and TcCDS with TcADH2 and TcALDH1. For terpenes, 738

m/z =123 was monitored, and for the internal control tetradecane, m/z =198 was monitored. 739

Peaks related to trans-chrysanthemol, trans-chrysanthemic acid and internal standard are labeled. 740

B. Concentrations of free trans-chrysanthemol and (C) free trans-chrysanthemic acid in N. 741

benthamiana leaves expressing the indicated constructs were determined by comparison with an 742

authentic standard. 743

D-F. The relative levels of Unknown 1 (D), Unknown 2 (E), and Unknown 3 (F) in N. 744

benthamiana leaves expressing the indicated constructs. For each compound, the plant material 745

expressing a specific construct that showed the highest levels (average of three biological 746

replicates) was set at 100%. 747

The data in B–F represent mean ± SD from triplicate biological replicates. N.D. - not detected. 748

749

Figure 7. LC-MS analysis of N. benthamiana leaves simultaneously expressing the three 750

enzymes TcCDS, TcADH2, and TcALDH1. Extracted ion chromatograms of m/z 803.37 and 751

831.33 are shown for: A. EGFP single expression control, B. TcCDS, C. TcCDS + TcADH2, D. 752

TcCDS + TcADH2 + TcALDH1. Chromatograms are all scaled the same, as indicated by the ion 753

current (1.01e7) in the upper right corner of each chromatogram. The sample for panel D was 754

diluted 10-fold compared to the other three samples due to high concentration of m/z 831.33 in 755

the sample. MS/MS spectra are shown for ions m/z 803.37 (E) and m/z 831.33 (F) along with the 756

proposed compound structures based on exact mass, fragmentation pattern, and similarity to 757

previously published data. G. Relative levels of trans-chrysanthemic acid in N. benthamiana 758

leaves expressing all TcCDS, TcADH2 and TcALDH1 with or without sodium hydroxide 759

treatment. 760

761



34

Table 1: Relative activities of recombinant TcADH1, TcADH2 and TcALDH1 with selected 762

substrates 763

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 100e

8-Hydroxygeraniol N.D. 70 (S)-(-)-Citronellal 84

Benzyl alcohol N.D. N.D. Benzaldehyde 90

Cinnamyl acohol N.D. N.D. trans-

Cinnamaldehyde

8

The activities of TcADH1 and TcADH2 were measured with 0.3 mM alcohols and 1mM NAD+. 764

The activities of TcALDH1 were measured with 0.04 mM aldehydes and 1mM NADP+. Data are 765

expressed as relative mean percentages from triplicate independent assays. 766

a N.D., not detectable (< 5% of highest activity) 767

b100% relative activity, corresponds to 0.44 μmol min-1 mg-1 of citral. 768

c100% relative activity, corresponds to 0.50 μmol min-1 mg-1 of trans-chrysanthemal. 769

e100% relative activity, corresponds to 0.11 μmol min-1 mg-1 of trans-chrysanthemic acid.770



35

Table 2: Kinetic properties of recombinant TcADH2 and TcALDH1 771

Enzyme Substrate Km(μM) Kcat(s-1) Kcat/Km (M-1

s-1)

TcADH2 trans-chrysanthemola 236.0 ± 5.8 0.75 ± 0.0032 3186.2

NAD+b 192.6 ± 8.7 0.64 ± 0.0034 3345.5

TcALDH1

trans-chrysanthemalc 4.4 ± 2.2 0.11 ± 0.0049 25122.4

NADP+d 20.4 ± 7.1 0.090 ± 0.0013 4391.8

trans-chrysanthemale 4. 6 ± 1.8 0.096 ± 0.0032 20992.3

NAD+f 68.6 ± 17.1 0.086 ± 0.0012 1256.07

Data are presented as mean ± SD from triplicate independent assays. All assays were performed 772

on GC-MS. 773 a Kinetic parameters were determined with 1 mM NAD+. 774 b Kinetic parameters were determined with 0.64 mM trans-chrysanthemol. 775 c Kinetic parameters were determined with 1 mM NADP+. 776 d Kinetic parameters were determined with 0.040 mM trans-chrysanthemal. 777 e Kinetic parameters were determined with 1 mM NAD+. 778 f Kinetic parameters were determined with 0.040 mM trans-chrysanthemal. 779

780

781 782 783 784 785 786 787 788 789 790 791 792 793



36

794



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CrossRef: Author and TitleGoogle Scholar: Author Only Title Only Author and Title

Teoh KH, Polichuk DR, Reed DW, Covello PS (2009) Molecular cloning of an aldehyde dehydrogenase implicated in artemisininbiosynthesis in Artemisia annua. Botany 87:635–642


Xu H, Zhang F, Liu B, Huhman DV, Sumner LW, Dixon RA, Wang G (2013) Characterization of the formation of branched short-chain fattyacid:CoAs for bitter acid biosynthesis in hop glandular trichomes. Mol Plant 6: 1301-1317


Yang T, Stoopen G, Yalpani N, Vervoort J, de Vos R, Voster A, Verstappen FW, Bouwmeester HJ, Jongsma MA (2011) Metabolicengineering of geranic acid in maize to achieve fungal resistance is compromised by novel glycosylation patterns. Metab Eng 13: 414-425


Yang T, Gao L, Hu H, Stoopen G, Wang C, Jongsma MA (2014) Chrysanthemyl diphosphate synthase operates in planta as abifunctional enzyme with chrysanthemol synthase activity. J Biol Chem 289: 36325-36335



http://search.crossref.org/?page=1&rows=2&q=Simpson JT, Wong K, Jackman SD, Schein JE, Jones SJ, Birol I (2009) ABySS: a parallel assembler for short read sequence data. Genome Res 19: 1117-1123

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http://scholar.google.com/scholar?as_q=Molecular cloning of an aldehyde dehydrogenase implicated in artemisinin biosynthesis in Artemisia annua.&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

http://scholar.google.com/scholar?as_q=Molecular cloning of an aldehyde dehydrogenase implicated in artemisinin biosynthesis in Artemisia annua.&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Teoh&as_ylo=2009&as_allsubj=all&hl=en&c2coff=1

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http://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Xu&hl=en&c2coff=1

http://scholar.google.com/scholar?as_q=Characterization of the formation of branched short-chain fatty acid:CoAs for bitter acid biosynthesis in hop glandular trichomes&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

http://scholar.google.com/scholar?as_q=Characterization of the formation of branched short-chain fatty acid:CoAs for bitter acid biosynthesis in hop glandular trichomes&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Xu&as_ylo=2013&as_allsubj=all&hl=en&c2coff=1

http://www.ncbi.nlm.nih.gov/pubmed?term=Yang T%2C Stoopen G%2C Yalpani N%2C Vervoort J%2C de Vos R%2C Voster A%2C Verstappen FW%2C Bouwmeester HJ%2C Jongsma MA %282011%29 Metabolic engineering of geranic acid in maize to achieve fungal resistance is compromised by novel glycosylation patterns%2E Metab Eng 13%3A 414%2D425&dopt=abstract

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http://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Yang&hl=en&c2coff=1

http://scholar.google.com/scholar?as_q=Metabolic engineering of geranic acid in maize to achieve fungal resistance is compromised by novel glycosylation patterns&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

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http://www.ncbi.nlm.nih.gov/pubmed?term=Yang T%2C Gao L%2C Hu H%2C Stoopen G%2C Wang C%2C Jongsma MA %282014%29 Chrysanthemyl diphosphate synthase operates in planta as a bifunctional enzyme with chrysanthemol synthase activity%2E J Biol Chem 289%3A 36325%2D36335&dopt=abstract

http://search.crossref.org/?page=1&rows=2&q=Yang T, Gao L, Hu H, Stoopen G, Wang C, Jongsma MA (2014) Chrysanthemyl diphosphate synthase operates in planta as a bifunctional enzyme with chrysanthemol synthase activity. J Biol Chem 289: 36325-36335

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http://scholar.google.com/scholar?as_q=Chrysanthemyl diphosphate synthase operates in planta as a bifunctional enzyme with chrysanthemol synthase activity&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Yang&as_ylo=2014&as_allsubj=all&hl=en&c2coff=1


Identification of dehydrogenases involved in the …...56 pyrethrolone, and cinerolone – that...

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