Reconfigured cyanogenic glucoside biosynthesis in ...example of convergent evolution in the biosynthesis of cyanogenic . 67. glucosides in plants. 68. 69. Introduction . 70. Cyanogenic

1

Article title: 1

Reconfigured cyanogenic glucoside biosynthesis in Eucalyptus cladocalyx involves a cytochrome P450 2

CYP706C55 3

4

5

Short title (not exceeding 50 characters and spaces): 6

Aunique CYP706 in prunasin synthesis in Eucalyptus 7

All author names and affiliations: 8

Cecilie Cetti Hansen1, 2, Mette Sørensen1, 2, Thiago A. M. Veiga3, Juliane F. S. Zibrandtsen1,6 , Allison M. 9

Heskes1, Carl Erik Olsen1,2, Berin A. Boughton4, Birger Lindberg Møller1,2,5, Elizabeth H. J. Neilson1,2 10

Affiliations: 11 1 Plant Biochemistry Laboratory, Department of Plant and Environmental Sciences, University of 12

Copenhagen, Thorvaldsensvej 40, Frederiksberg C, Copenhagen, Denmark 13 2 VILLUM Center for Plant Plasticity, University of Copenhagen, Thorvaldsensvej 40, Frederiksberg C, 14

Copenhagen, Denmark 15 3 Department of Chemistry, Federal University of São Paulo, Diadema, Brazil 16 4 Metabolomics Australia, School of BioSciences, The University of Melbourne, Vic. 3010, Australia 17 5 Center for Synthetic Biology, University of Copenhagen, Thorvaldsensvej 40, Frederiksberg C, 18

Copenhagen, Denmark 19

Present addresses 20 6 Syngenta, Crescent House, Tower Business Park, M20 2JE Manchester, United Kingdom 21

Corresponding author: 22

Elizabeth H. J. Neilson 23

[email protected] 24

ORCID: 0000-0002-8279-9906 25

Plant Biochemistry Laboratory, Department of Plant and Environmental Sciences, University of 26

Copenhagen, Denmark 27

One sentence summary: 28

Cyanogenic glucoside biosynthesis in Eucalyptus cladocalyx requires an additional pathway step, involving 29

three CYPs from the CYP79, CYP706 and CYP71 families, and a glucosyltransferase UGT85. 30

31

List of author contributions: 32

EHJN conceived the study and research plans; CCH, MS, TAM, JFSZ and EHJN performed the experiments; EHJN, 33

BLM, AMH and BB supervised experimental work; CEO and BB provided technical assistance; CCH, MS and EHJN 34

analyzed the data; CCH wrote the draft with contributions from MS, EHJN, and BLM. All authors have reviewed 35

and approved the final manuscript. 36

Funding information: 37

Plant Physiology Preview. Published on October 8, 2018, as DOI:10.1104/pp.18.00998

Copyright 2018 by the American Society of Plant Biologists

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This research was supported by the “VILLUM Research Center for Plant Plasticity” (Project No. 7523), by the 38

UCPH Excellence Program for Interdisciplinary Research to Center for Synthetic Biology, and by an ERC 39

Advanced Grant (ERC-2012-ADG_20120314, Project No. 323034) to BLM. EHJN was supported by a Young 40

Investigator Program fellowship from the VILLUM Foundation (Project No. 13167), a grant from the 41

Carlsberg Foundation (Grant no. 2013_01_0908), and by a Danish Independent Research Council Sapere 42

Aude Research Talent Post Doctoral Stipend (Grant No. 6111-00379B). TAMV was supported by a visiting 43

professor fellowship from the Fundação de Amparo a Pesquisa do Estado de São Paulo (#FAPESP BPE 44

2014/11811-2). AMH was supported by a Marie Skłodowska Curie Individual Fellowship (Project No. 45

658677). Mass spectrometry imaging was conducted at Metabolomics Australia located at the School of 46

BioSciences, The University of Melbourne, Australia, which is an NCRIS initiative under Bioplatforms 47

Australia Pty Ltd. 48

Abstract 49

Cyanogenic glucosides are a class of specialized metabolites widespread in the plant kingdom. Cyanogenic 50

glucosides are α-hydroxynitriles, and their hydrolysis releases toxic hydrogen cyanide providing an effective 51

chemical defense against herbivores. Eucalyptus cladocalyx F. Muell. is a cyanogenic tree, allocating up to 52

20% of leaf nitrogen to the biosynthesis of the cyanogenic monoglucoside, prunasin. Here, mass 53

spectrometry analyses of E. cladocalyx tissues revealed spatial and ontogenetic variation in prunasin 54

content, as well as the presence of the cyanogenic diglucoside amygdalin in flower buds and flowers. 55

Identification and biochemical characterization of the prunasin biosynthetic enzymes revealed a unique 56

enzyme configuration for prunasin production in E. cladocalyx. Our result indicates that a multifunctional 57

cytochrome P450 (CYP), CYP79A125, catalyzes the initial conversion of L-phenylalanine into its 58

corresponding aldoxime, phenylacetaldoxime; a function consistent with other members of the CYP79 59

family. In contrast to the single multifunctional CYP known from other plant species, the conversion of 60

phenylacetaldoxime to the α-hydroxynitrile, mandelonitrile, is catalyzed by two distinct CYPs. CYP706C55 61

catalyzes the dehydration of phenylacetaldoxime, an unusual CYP reaction. The resulting phenylacetonitrile 62

is subsequently hydroxylated by CYP71B103 to form mandelonitrile. The final glucosylation step to yield 63

prunasin is catalyzed by an UDP-glucosyltransferase, UGT85A59. Members of the CYP706 family have not 64

previously been reported to participate in the biosynthesis of cyanogenic glucosides and the pathway 65

structure in E. cladocalyx represents an example of convergent evolution in the biosynthesis of cyanogenic 66

glucosides in plants. 67

68

Introduction 69

Cyanogenic glucosides are a class of specialized metabolites found widespread in the plant kingdom. They 70

are present in diverse taxa including species in the genera Pteridium (monilophyte), Taxus (gymnosperm), 71

Sorghum (monocot), Manihot, Trifolium, Prunus and Eucalyptus (eudicots) (Vetter 2017). Upon tissue 72

disruption, such as would be caused by herbivory, the cyanogenic glucoside is hydrolyzed by a specific β-73

glucosidase that cleaves off the sugar moiety. The resulting cyanohydrin dissociates, spontaneously or 74

catalyzed by an α-hydroxynitrile lyase, resulting in the release of toxic hydrogen cyanide. This process is 75

known as cyanogenesis (Gleadow and Møller 2014). Their ability to release hydrogen cyanide makes 76

cyanogenic glucosides chemical defense molecules against generalist herbivores and pathogens (Gleadow 77

and Woodrow 2002). Cyanogenic glucosides have also been shown to constitute a source of reduced 78



3

nitrogen that are mobilized by endogenous turnover pathways (Bjarnholt et al. 2018; Pičmanová et al. 79

2015; Schmidt et al. 2018a). 80

The biosynthetic pathway genes for a cyanogenic glucoside were first identified and cloned from sorghum 81

(Sorghum bicolor), which accumulates the tyrosine-derived dhurrin (Bak et al. 1998; Jones et al. 1999; Koch 82

et al. 1995). This provided a starting point for the identification of cyanogenic glucoside pathway genes in 83

other plant species. The biosynthetic pathway for the valine-derived linamarin and isoleucine-derived 84

lotaustralin has been elucidated from cassava (Manihot esculenta) (Andersen et al. 2000; Jørgensen et al. 85

2011; Kannangara et al. 2011) and Lotus japonicus (Forslund et al. 2004; Takos et al. 2011), and recently the 86

full pathway for the phenylalanine-derived diglucoside amygdalin from almond (Prunus dulcis) was 87

identified (Thodberg et al. 2018). In species with characterized cyanogenic glucoside biosynthetic pathways, 88

the production of monoglucosides involves two cytochrome P450s (CYPs) and a soluble UDP-89

glucosyltransferase (UGT). The first CYP is a member of the CYP79 family and catalyzes the conversion of an 90

amino acid to the corresponding aldoxime. A CYP71 or CYP736 family member subsequently converts the 91

aldoxime to a cyanohydrin. The labile cyanohydrin is stabilized by glucosylation to form the final cyanogenic 92

glucosides in a reaction catalyzed by a UGT from the UGT85 family. Work on dhurrin biosynthesis in S. 93

bicolor shows that the two CYPs and UGT of the pathway form a metabolon enabling efficient channeling 94

between enzymes that avoids the accumulation of toxic intermediates (Bassard et al. 2017; Laursen et al. 95

2016). 96

Eucalyptus is a large genus with over 800 species (Coppen 2002). They are the world’s most widely planted 97

forest hardwood trees due to their fast growth, environmental plasticity and exceptional wood properties 98

(Myburg et al. 2014). The trees are grown for industrial purposes such as the production of pulp for paper, 99

firewood, charcoal, and essential oils. Eucalyptus plants synthesize a broad spectrum of specialized 100

metabolites including terpenes, phenolics, formylated phloroglucinols, and cyanogenic glucosides (Eschler 101

et al. 2000; Gleadow et al. 2008; Marsh et al. 2017; Padovan et al. 2014). Very little is known about the 102

biosynthesis of specialized metabolites in Eucalyptus. The genome of E. grandis published in 2014 (Myburg 103

et al. 2014) will potentially facilitate the identification of enzymes involved in these pathways. E. grandis 104

has many tandem duplicated genes and tandem expanded clusters as exemplified by the large number of 105

terpene synthase genes (113) and the expansion of some phenylpropanoid genes (Kulheim et al. 2015; 106

Myburg et al. 2014). The significant expansion of some gene families in Eucalyptus provides challenging 107

obstacles towards the identification of pathway genes due to the large number of possible candidates. 108

More than 400 Eucalyptus species have been screened for cyanogenesis and 23 cyanogenic species have 109

been identified, all containing prunasin as the major cyanogen (Fig. 1, B) (Gleadow et al. 2008). The South 110

Australian species E. cladocalyx (sugar gum) was first reported to be cyanogenic in 1928 (Finnemore and 111

Cox 1928) and a few years later the foliar hydrogen cyanide source was identified as prunasin (Finnemore 112

et al. 1935). In particular, the apical tips and young leaf tissue of E. cladocalyx can be highly cyanogenic with 113

up to 20% of leaf nitrogen allocated to prunasin biosynthesis (Gleadow et al. 1998; Gleadow and Woodrow 114

2000b). Cyanogenic Eucalyptus species display unique ontogenetic variation. For example, E. cladocalyx has 115

high levels of prunasin as a seedling compared to adult, while the opposite trend is observed for 116

E. yarraensis and E. camphora (Goodger et al. 2006; Neilson et al. 2011). As such, cyanogenic Eucalyptus 117

species are an excellent system to investigate the regulation of chemical defense (Møller 2010). 118



4

In this study, we investigate the spatial and ontogenetic variation and localization of cyanogenic glucoside 119

accumulation in E. cladocalyx. Furthermore, we report the identification and functional characterization of 120

the enzymes catalyzing the formation of prunasin from L-phenylalanine in E. cladocalyx. We show that the 121

conversion of phenylalanine into the corresponding cyanohydrin requires the sequential action of three 122

CYPs, and includes the production of a free nitrile intermediate. One of the CYPs belongs to the CYP706 123

family, a family not previously reported to be involved in the synthesis of cyanogenic glucosides. 124

Results 125

Spatial distribution of cyanogenic glucosides 126

Prunasin extracts of expanded juvenile leaves from young E. cladocalyx plants (4 months), along with 127

expanded adult leaves, immature flower buds, mature flower buds, flowers and mature fruit from mature 128

trees were analyzed by liquid chromatography-mass spectrometry (LC-MS) (Fig. 1). Prunasin was detected 129

in all tissues examined. The lowest levels of prunasin were detected in young and adult leaves (17 ± 2.8 130

μg/mgDW and 14 ± 4.2 μg/mgDW, respectively) and the highest levels were found in immature and mature 131

flower buds (62 ± 12 μg/mgDW and 53 ± 9 μg/mgDW, respectively) with intermediate levels detected in fruit 132

(43 ± 1 μg/mgDW), and flowers (40 ± 1 μg/mgDW). Additionally, the diglucoside amygdalin (Fig. 1, B) was 133

found in the mature flower buds (4 ± 1.8 μg/mgDW) and flowers (9 ± 0.6 μg/mgDW) accounting for 134

approximately 6% and 18% of total cyanogenic content, respectively (Fig. 1, A). 135

The use of matrix-assisted laser desorption ionization-mass spectrometry imaging (MALDI-MSI) to map the 136

spatial localization of prunasin and amygdalin within prepared tissues, revealed prunasin to be localized to 137

the mesophyll and vascular tissue of seedling and adult E. cladocalyx leaves (Fig. 2, A-B). Amygdalin was not 138

detected in the leaf material using this method. MALDI-MSI of an E. cladocalyx flower bud demonstrated 139

the presence of prunasin in the ground tissue of the ovary, hypanthium and operculum, as well as in 140

filaments and anthers. Amygdalin was observed to be co-localized with prunasin, specifically in the 141

filaments and staminal ring region (Fig. 2, C-D). 142

Phenylalanine is the precursor for prunasin synthesis in E. cladocalyx 143

Cyanogenic glucosides are known to be derived from amino acids (Gleadow and Møller 2014), and thus we 144

expected prunasin in E. cladocalyx to be derived from L-phenylalanine, based on its structure (Fig. 1, B). To 145

verify this, L-[14C]-phenylalanine was administered to apical tips, expanding leaves and fully expanded 146

leaves. All tested tissues demonstrated activity of the prunasin biosynthetic enzymes by the ability to 147

synthesize prunasin from L-phenylalanine (Fig. 3). 148

Identification and testing of three candidate genes involved in prunasin biosynthesis 149

A candidate CYP79 was identified by PCR, using cDNA generated from young leaves of E. cladocalyx and 150

primers designed to the E. yarraensis CYP79A34 (Neilson 2012). Based on characterized cyanogenic 151

glucoside biosynthetic pathways, candidate genes belonging to CYP71 and UGT85 families that showed high 152

expression in an E. cladocalyx transcriptome and no or very low expression in the acyanogenic E. globulus 153

transcriptome were identified (Spokevicius et al. 2017). Degenerate primers were designed to anneal to the 154

UTR regions of the CYP71 and UGT85 orthologs from the E. grandis genome available on Phytozome 155

(Goodstein et al. 2012) and used to isolate the two gene sequences from E. cladocalyx cDNA. The CYPs 156

where named CYP79A125 and CYP71B103 by David Nelson for the Standardized Cytochrome P450 157



5

Nomenclature Committee (Nelson 2009), and the UGT candidate was named UGT85A59 according to the 158

standard UGT Nomenclature Guidelines (Burchell et al. 1991; Mackenzie et al. 2005). 159

Testing of the three candidates by co-infiltration into Nicotiana benthamiana leaves only resulted in 160

formation of trace amounts of prunasin (Fig. 4, A; Supplemental Fig. S1). The data, however, confirmed that 161

CYP79A125 and UGT85A59 were functionally active in N. benthamiana. Expression of CYP79A125 resulted 162

in the production of glucosylated phenylacetaldoxime (Supplemental Fig. S2). This suggests that CYP79A125 163

can catalyze the conversion of L-phenylalanine into phenylacetaldoxime that subsequently becomes 164

glucosylated by endogenous N. benthamiana UGTs. Co-expression of UGT85A59 showed an increase in 165

glucosylated phenylacetaldoxime, demonstrating that UGT85A59 is active and able to glucosylate 166

phenylacetaldoxime. 167

CYP706C55 is involved in prunasin biosynthesis 168

To identify other potential candidates involved in prunasin biosynthesis, a transcriptome was prepared 169

from the highly cyanogenic and biosynthetically active apical tips of E. cladocalyx. CYP71B103 and 170

CYP79A125 were the second and third most highly expressed CYPs, respectively, and UGT85A59 was the 171

highest expressed UGT. The most highly expressed CYP in the transcriptome belonged to the CYP706 172

family. This CYP706 was named CYP706C55 (Nelson 2009) and selected for functional analysis. 173

CYP706C55 was tested by transient expression in N. benthamiana together with the previously identified 174

pathway candidates. Expression of CYP79A125, CYP706C55 and UGT85A59 resulted in small amounts of 175

prunasin (Fig. 4, A). However, combined expression of all three CYP candidate genes with UGT85A59 176

resulted in high accumulation of prunasin (Fig. 4, A). Additionally, a novel peak with m/z 404 was observed 177

and tentatively identified as a malonic acid ester of prunasin (Supplemental Fig. S3). Malonic acid 178

derivatives have previously been observed upon transient expression in Nicotiana sp. (Franzmayr et al. 179

2012; Tian and Dixon 2006; Ting et al. 2013), likely as a detoxification mechanism. No prunasin or 180

derivatives thereof were observed in the negative control where no E. cladocalyx genes were expressed. 181

Likewise, no prunasin was observed upon expression of CYP71B103, CYP706C55 and UGT85A59, confirming 182

that CYP79A125 is required for prunasin production in N. benthamiana. Small amounts of prunasin were 183

observed if the three CYPs were co-expressed, indicating that endogenous N. benthamiana 184

glycosyltransferases can glycosylate the mandelonitrile to some extent. However, the prunasin levels 185

increased 50-fold upon co-expression of UGT85A59 (Fig. 4, A). 186

To establish the order of the catalytic activity of CYP71B103 and CYP706C55 in prunasin biosynthesis, 187

microsomes were prepared from infiltrated N. benthamiana leaves and assayed using radioactively labelled 188

L-phenylalanine as substrate. Microsomes with CYP79A125 activity converted L-phenylalanine into a 189

product that co-migrated with phenylacetaldoxime (Fig. 4, B). Microsomes with CYP79A125 and CYP706C55 190

produced phenylacetaldoxime and phenylacetonitrile, and microsomes with all three CYPs showed 191

production of phenylacetaldoxime and mandelonitrile. Microsomes with CYP79A125 and CYP71B103 only 192

produced phenylacetaldoxime, and the negative control without E. cladocalyx CYPs did not show any 193

product formation. In summary, based on transient expression analysis and microsomal assays, we 194

demonstrate a route for prunasin biosynthesis in E. cladocalyx leaves. Specifically, CYP79A125 converts L-195

phenylalanine into phenylacetaldoxime, which is then dehydrated into phenylacetonitrile by CYP706C55, 196

hydroxylated by CYP71B103 to form mandelonitrile, and finally glucosylated by UGT85A59 to form prunasin 197

(Fig. 4, C). 198



6

Phylogenetic analysis of the CYP79, CYP706, CYP736 and CYP71 families 199

The amino acid sequences of CYP79A125, CYP706C55 and CYP71B103 were subjected to phylogenetic 200

analysis together with functionally characterized CYP members from the CYP79, CYP706, CYP71 and CYP736 201

families (Fig. 5). These CYP families all contain at least one member involved in cyanogenic glucoside 202

biosynthesis (Supplemental Table S1). All functionally characterized CYP79 members convert amino acids 203

into their corresponding oximes (Sørensen et al. 2018), including 12 CYP79 members involved in cyanogenic 204

glucoside biosynthesis. In contrast, members of the CYP71 family show broad functional diversity 205

(Supplemental Table S1). The cyanogenic glucoside CYP71 members are distributed across several 206

subfamilies, and this study introduces the first CYP71B involved in cyanogenic glucoside biosynthesis. Fewer 207

CYPs belonging to the CYP706 and CYP736 families have been functionally characterized, with only a single 208

member in each of the CYP706 and CYP736 families known to participate in cyanogenic glucoside 209

biosynthesis. 210

211

Prunasin accumulation through leaf ontogeny 212

The spatial variation of leaf prunasin was measured during plant development and prunasin was extracted 213

from apical tips, expanding leaves and fully expanded leaves in plants at 4 months, 7 months and 10 214

months of age. Quantitative variation in prunasin accumulation was observed throughout leaf ontogeny 215

(Fig. 6, A). The most cyanogenic tissue was the apical tips in 4-month-old plants. Accumulation of prunasin 216

decreased with leaf age; in 4-month-old plants prunasin accumulated to 45 ± 5 μg/mgDW in apical tips and 217

decreased to 25 ± 2 μg/mgDW in expanding leaves and 17 ± 3 μg/mgDW in fully expanded leaves. In older 218

plants at 7- and 10-months of age, prunasin accumulation in leaves was lower and showed less variation 219

between leaf types and plant age not exceeding 11 μg/mgDW. 220

Expression of CYP79A125 and CYP706C55 is positively correlated 221

Expression of the characterized biosynthetic pathway genes were analyzed in apical tips, expanding and 222

fully expanded leaves in plants at 4-, 7- and 10-months of age (Fig. 6, B). Expression of all genes was 223

observed in all tested tissues, consistent with their ability to synthesize prunasin (Fig. 3; Fig. 6, A). Biological 224

replicates showed a relatively large variation in expression, however, most individuals showed the same 225

trend across tissues and time. In particular, all plants except one showed the lowest CYP79A125 expression 226

in expanded leaves at the three different time points. A Pearson correlation analysis revealed that 227

expression of the CYPs correlated; expression of CYP79A125 correlated with CYP706C55 (P < 0.01) and 228

CYP71B103 (P < 0.01), and expression of CYP706C55 and CYP71B103 also correlated (P < 0.05) (Fig. 6, C). 229

UGT85A59 was generally highly expressed and did not correlate with the expression of the CYPs. 230

Discussion 231

Prunasin accumulation varies between tissue type and plant age 232

The accumulation of cyanogenic glucosides was investigated in different tissues of E. cladocalyx by LC-MS 233

and MALDI-MSI. At the overall spatial level, the highest prunasin concentration (62 ± 12 μg/mgDW) was 234

found in reproductive tissues (Fig. 1). In addition, E. cladocalyx leaf tissue was most cyanogenic in 4-month- 235

old plants and then decreased with plant age, and in particular prunasin accumulation was found to 236

decrease by leaf age with apical tips being the most cyanogenic followed by expanding leaves and 237

expanded leaves, respectively (Fig. 6). This is in agreement with a previous study observing relatively high 238

cyanide potentials in flower buds, flowers and fruits (Gleadow and Woodrow 2000b) and reports showing 239

that juvenile E. cladocalyx leaves are more cyanogenic than adult leaves, with the inverse relationship 240



7

between leaf age and cyanogenic glucoside concentration (Gleadow and Woodrow 2000b; Goodger et al. 241

2006). Only in young 1-week-old seedlings, prunasin content was found to be insignificant (Gleadow and 242

Woodrow 2000b). We also identified the presence of the cyanogenic diglucoside amygdalin E. cladocalyx 243

(Fig. 1), with the highest levels in the flower buds and flowers (4 ± 2 μg/mgDW and 9 ± 1 μg/mgDW, 244

respectively). Amygdalin has not previously been reported to be present in E. cladocalyx. Cyanogenic 245

diglucosides, including amygdalin, were previously identified in E. camphora, with the highest 246

concentration also identified in reproductive tissue (Neilson et al. 2011). 247

Juvenile E. cladocalyx plants tend to be more cyanogenic compared to adult plants (Goodger et al. 2006). 248

However, this is not the case in all cyanogenic Eucalyptus species and the ontogenetic trajectory of 249

cyanogenic glucoside production is known to differ between some species. In E. camphora, foliar 250

cyanogenic glucoside production was found to increase with plant age as prunasin accumulation increased 251

from seedling to sapling to adult (Neilson et al. 2011). A similar pattern has been observed for 252

E. polyanthemos (Goodger et al. 2004). A different trajectory was observed for E. yarraensis, where a 253

detailed study of prunasin concentration through ontogeny revealed that production gradually increased 254

with a maximum concentration at around 240 days after sowing; thereafter, prunasin accumulation 255

declined with age (Goodger et al. 2007). The species-specific differences in the onset of cyanogenic 256

glucoside production provide a unique experimental system for studying the role and regulation of 257

cyanogenic glucoside production. 258

The high concentration of cyanogenic glucosides in young plants, apical tips and reproductive tissue of 259

E. cladocalyx is in accordance with the optimal defense theory of plant defense, whereby the most 260

vulnerable and important structures are the most highly defended (Rhoades 1979). These results support 261

the general role of cyanogenic glucosides as chemical defense compounds against generalist herbivores 262

and pathogens (Gleadow and Woodrow 2002). In recent years, it has become apparent that cyanogenic 263

glucosides perform additional roles in plant metabolism, for example, as storage molecules or in quenching 264

reactive oxygen species (ROS) (Gleadow and Møller 2014; Schmidt et al. 2018a). Enzymes and 265

intermediates in endogenous recycling of auto-toxic cyanogenic glucosides, including prunasin and 266

amygdalin, have been identified (Bjarnholt et al. 2018; Jenrich et al. 2007; Pičmanová et al. 2015). Tissue 267

based localization studies using MALDI-MSI, provide important information to aid the understanding of 268

specialized metabolite multi-functionality in planta (Bjarnholt et al. 2014; Boughton et al. 2016). Using 269

MALDI-MSI, we show that prunasin is localized to the mesophyll and vascular tissue of seedling and adult E. 270

cladocalyx leaves (Fig. 2, A-B). The localization of prunasin in the vascular tissue of E. cladocalyx leaves 271

suggests the transport of cyanogenic glucosides within the plant. Transport of cyanogenic glucosides is 272

known in other cyanogenic species including M. esculenta (Jørgensen et al. 2005), Hevea brasiliensis 273

(Selmar 1993; Selmar et al. 1988) and S. bicolor (Darbani et al. 2016; Selmar et al. 1996). Once transported, 274

it is hypothesized that the cyanogenic glucoside is recycled and provides a source of reduced carbon and 275

nitrogen (Bjarnholt et al. 2018; Pičmanová et al. 2015). 276

MALDI-MSI of the E. cladocalyx flower bud revealed distinct patterns of prunasin and amygdalin 277

localization. Prunasin was widely distributed throughout the ground tissue of the ovary, hypanthium and 278

operculum, and present in the filaments and anthers (Fig. 2, C-D), consistent with a role in chemical 279

defense. Amygdalin co-localized with prunasin but was specifically restricted to the filaments and around 280

the staminal ring region (Fig. 2, C-D). In other species, cyanogenic diglycosides are suggested to be the 281



8

transport form (Neilson et al. 2011; Selmar 1993; Selmar et al. 1996), and the results here may suggest 282

transport of cyanogenic glucosides from the hypanthium to the stamens. The role of cyanogenic glucosides 283

in flower parts has received little attention (Lai et al. 2015). Cyanogenic glucosides have previously been 284

detected in the pollen of almond flowers (Del Cueto et al. 2017; London-Shafir et al. 2003), where they are 285

hypothesized to act as a chemical deterrent to inefficient pollinators. Other specialized metabolites, such as 286

flavonol glycosides, provide a critical role in pollen development, thermo tolerance and ROS protection 287

(Paupiere et al. 2014). With their multiple functions in plants, investigations into the role of cyanogenic 288

glucosides in Eucalyptus flowers would be an intriguing avenue of research. 289

Recruitment of CYP706C55 involves an additional free nitrile intermediate in prunasin synthesis 290

We have identified and characterized three CYPs and an UGT from E. cladocalyx catalyzing the formation of 291

prunasin. Similar to other known cyanogenic glucoside biosynthetic pathways, the first step of prunasin 292

biosynthesis in E. cladocalyx is catalyzed by a multifunctional CYP79 that converts an amino acid into an 293

aldoxime. The following conversion of the aldoxime into a cyanohydrin requires two CYPs in E. cladocalyx, 294

instead of a single multifunctional CYP. CYP706C55 converts phenylacetaldoxime into phenylacetonitrile, 295

which is subsequently hydroxylated by CYP71B103 to form mandelonitrile. This is in contrast to the well 296

characterized dhurrin biosynthesis in S. bicolor, where CYP71E1 catalyzes a rearrangement of (E)-p-297

hydroxyphenylacetaldoxime to (Z)-p-hydroxyphenylacetaldoxime, then a dehydration of (Z)-p-298

hydroxyphenylacetaldoxime to p-hydroxyphenylacetonitrile followed by a C-hydroxylation resulting in p-299

hydroxymandelonitrile (Bak et al. 1998; Clausen et al. 2015; Kahn et al. 1997; Kahn et al. 1999). In prunasin-300

producing Prunus species the conversion of phenylacetaldoxime to mandelonitrile is likewise catalyzed by a 301

single CYP71 (Yamaguchi et al. 2014). The small amount of prunasin observed from N. benthamiana leaves 302

expressing CYP79A125, CYP706C55 and UGT85A59 could imply that an endogenous N. benthamiana 303

enzyme can hydroxylate phenylacetonitrile or that CYP706C55 has minor hydroxylation activity. However, 304

mandelonitrile was not observed as a product in microsomes containing CYP79A125 and CYP706C55. 305

The involvement of an extra CYP in prunasin biosynthesis in E. cladocalyx supports the hypothesis that 306

cyanogenic glucoside biosynthesis has evolved independently in several plant lineages. This hypothesis is 307

exemplified by the identification and characterization of L. japonicus CYP736A2 in lotaustralin and linamarin 308

biosynthesis (Takos et al. 2011). Differences in the biosynthesis of hydrogen cyanide based defenses are 309

also known from outside the plant kingdom. In 2010, it was shown that cyanogenic glucoside biosynthesis 310

in plants and insects evolved independently; in the burnet moth (Zygaena filipendulae), linamarin and 311

lotaustralin are produced by two CYPs (CYP405A2 and CYP332A3) and a glucosyl transferase (UGT33A1) 312

(Jensen et al. 2011). The number of enzymes required for production of the cyanohydrin may also differ in 313

insects. The millipede Chamberlinius hualienensis produces mandelonitrile as a precursor substrate for its 314

hydrogen cyanide based defense, and recently a CYP (CYP320B1) was shown to hydroxylate 315

phenylacetonitrile to form mandelonitrile (Yamaguchi et al. 2017). This function is similar to that of 316

CYP71B103 reported here and indicates that the production of α-hydroxynitriles for hydrogen cyanide 317

defense in some insects might also involve three CYPs. 318

The three-CYP-system in prunasin biosynthesis in E. cladocalyx presented here comprises an additional 319

intermediate that needs to be channeled between enzymes for production of prunasin compared to the 320

two-CYP-system in other cyanogenic species. Interestingly, the volatile intermediate phenylacetonitrile is 321

produced and released in some plant species in response to herbivory. For example, production of 322



9

herbivore-induced volatiles including phenylacetonitrile has been reported for western balsam poplar 323

(Populus trichocarpa) (Irmisch et al. 2013), black poplar (P. nigra) (McCormick et al. 2014) and giant 324

knotweed (Fallopia sachalinensis) (Noge et al. 2011). It can be speculated whether some Eucalyptus species 325

might produce and release herbivore-induced nitriles. To our knowledge, however, emission of these 326

volatile compounds has not been reported for Eucalyptus. 327

Interestingly, CYPs from different families have acquired the ability to produce phenylacetonitrile from 328

phenylacetaldoxime. In E. cladocalyx, CYP706C55 from the CYP706 family catalyzes this reaction, whereas 329

P. trichocarpa and F. sachalinensis possess CYPs belonging to the large and functionally diverse CYP71 330

family that convert phenylacetaldoxime into phenylacetonitrile (Irmisch et al. 2014; Yamaguchi et al. 2016) 331

(Fig. 5, Supplemental Table S1). Phylogenetically, the E. cladocalyx CYP71B103 that hydroxylates 332

phenylacetonitrile group together with the P. trichocarpa CYP71s. These examples of functional diversity 333

within CYP families, together with the examples of independent evolution of cyanogenic glucoside 334

biosynthesis in plants, suggest that it is sometimes necessary to move away from phylogeny for 335

identification of CYP enzymes involved in the biosynthesis of interest. In contrast, the reactions catalyzed 336

by the CYP79 family are more conserved as they all convert an amino acid into its corresponding oxime 337

(Sørensen et al. 2018). 338

CYP706C55 catalyzes an atypical CYP reaction 339

CYP706C55 was identified as the second enzyme in the prunasin biosynthetic pathway in E. cladocalyx. 340

Three CYP706 members from other plant species have previously been functionally characterized and 341

demonstrated to catalyze oxygenation reactions: CYP706B1 from cotton (Gossypium arboruem L.) (Luo et 342

al. 2001), CYP706M1 from Alaskar cedar (Callitropsis nootkatensis), and a CYP706X from Erigeron 343

breviscapus (Liu et al. 2018). In contrast, CYP706C55 catalyzes an unusual CYP reaction, which is the 344

dehydration of phenyacetaldoxime to phenylacetonitrile. Although this is an unusual CYP reaction, CYP-345

catalyzed dehydration of an oxime to its corresponding nitrile have been characterized before for human 346

CYP3A4 (Boucher et al. 1994), Arabidopsis thaliana CYP71A13 (Nafisi et al. 2007), P. trichocarpa 347

CYP71B40v3 and CYP71B41v2 (Irmisch et al. 2014), and F. sachalinensis CYP71AT96 (Yamaguchi et al. 2016). 348

Dehydration of an aldoxime is also one of the partial reactions catalyzed by the multifunctional CYP71E1 in 349

dhurrin biosynthesis in sorghum as discussed above. Nonetheless, if CYP71E1 was administered with p-350

hydroxyphenylacetaldoxime in the absence of cytochrome P450 oxidoreductase (POR) and NAPDH, the 351

reaction resulted in accumulation of p-hydroxyphenylacetonitrile (Bak et al. 1998). 352

Exogenous NADPH and POR are not necessarily needed in in vitro assays for CYP71A13 and CYP71B41v2 to 353

catalyze the formation of indole-3-acetacetonitrile and phenylacetonitrile, respectively (Irmisch et al. 2014; 354

Nafisi et al. 2007). Even though POR may not be required for CYP706C55 catalysis, it has been shown that a 355

reducing environment is needed for CYP catalyzed dehydration of oximes (Boucher et al. 1994; DeMaster et 356

al. 1992; Hart-Davis et al. 1998). A reducing environment established by, for example, addition of NADPH or 357

dithionite to an assay will bring the heme group of the CYP to the Fe(II) state. This enables binding of the 358

aldoxime nitrogen to the heme, which allows for a charge transfer from Fe(II) to the aldoxime C=N bond 359

favoring elimination of the hydroxy group (Boucher et al. 1994; Hart-Davis et al. 1998). 360

Regulation of prunasin biosynthesis 361

Prunasin accumulation and gene expression were investigated in 4-, 7- and 10-month-old E. cladocalyx 362

seedlings (Fig. 6). Overall, biological replicates showed a large variation in expression of the pathway genes. 363



10

Likewise, a large variation in prunasin content was also observed. A great variation in prunasin levels was to 364

be expected from the eight individuals as studies of a natural population of adult E. cladocalyx also showed 365

great variation with a difference of two orders of magnitude in hydrogen cyanide potential detected 366

(Gleadow and Woodrow 2000a). The expression pattern of the pathway genes in apical tips, expanding and 367

expanded leaves in 4-month-old plants showed less decrease with leaf age than expected based on the 368

observed pattern of prunasin accumulation. However, most plants showed the highest expression of the 369

CYPs in apical tips and expanding leaves at all three time points and the expression was found to correlate. 370

In contrast, the expression of UGT85A59 did not correlate with any of the CYPs. Similar to these findings, 371

expression levels of the two CYPs in cyanogenic glucoside biosynthesis in L. japonicus both decreased with 372

leaf age, and expression of the UGTs, which catalyze glucosylation of the cyanohydrins, did not followed the 373

CYP pattern (Takos et al. 2011). CYP79A125 was the lowest expressed pathway gene and showed the 374

largest relative variation between leaf types (Fig. 6), which may indicate that the first committed step in 375

prunasin biosynthesis controls the overall flux through the pathway. Similarly, the catalytic activity of 376

CYP79A1 constitutes the rate limiting step in dhurrin biosynthesis in sorghum (Busk and Møller 2002; 377

Nielsen et al. 2016). 378

Glucosylation by UGT85A59 results in prunasin formation 379

Transient expression of UGT85A59 in N. benthamiana showed that UGT85A59 catalyzes glucosylation of 380

the cyanohydrin, resulting in prunasin formation. Consistent with the function of stabilizing the labile 381

cyanohydrin to avoid toxic release of hydrogen cyanide, UGT85A59 was the highest expressed pathway 382

gene at most time points (Fig. 6). Since the diglucoside amygdalin was detected in E. cladocalyx metabolite 383

extracts (Fig. 1) but not upon transient expression of UGT85A59 in N. benthamiana, an additional UGT 384

remains to be identified that can catalyze a β(1→6)-O glycosylation of prunasin to yield amygdalin. In other 385

cyanogenic plant species, UGT85 members catalyze a mono-glucosylation of the cyanohydrins resulting in 386

the corresponding cyanogenic glucoside (Franks et al. 2008; Jones et al. 1999; Kannangara et al. 2011; 387

Takos et al. 2011). In contrast, less is known about UGTs that catalyze the addition of a second sugar. 388

Recently, two glucosyltransferases, UGT94AF1 and UGT94F2, were shown to glucosylate prunasin to yield 389

amygdalin in almond (Thodberg et al. 2018), and it can be speculated that a UGT94 also catalyzes the 390

glucosylation of prunasin in E. cladocalyx. 391

Conclusion 392

Our current study has characterized cyanogenic glucoside content and prunasin biosynthesis in 393

E. cladocalyx. The monoglucoside prunasin is the major cyanogen in all tissues, although significant levels of 394

the diglucoside amygdalin were detected in the reproductive organs. Until now, all characterized 395

biosynthetic pathways of cyanogenic glucosides in higher plants consist of two CYP-catalyzed reactions 396

followed by a final glucosylation reaction. Here, we show by biochemical characterization that prunasin 397

production in E. cladocalyx is catalyzed by three CYPs from different families, and a glucosyltransferase. 398

Notably, the second biosynthetic step is catalyzed by a CYP706, a CYP family not previously reported to 399

participate in the biosynthesis of cyanogenic glucosides. This finding represents a unique example of 400

convergent evolution in the biosynthesis of cyanogenic glucosides in plants. 401

Material and Methods 402

Plant material 403



11

Eucalyptus cladocalyx (Sugar Gum) seeds from a single maternal tree were purchased from CSIRO 404

Australian Tree Seed Centre (Clayton, Australia). Seedlings for the ontogenetic study were germinated in 405

the glasshouse at the University of Copenhagen on the 24th of August 2013 and grown at a minimum 22°C 406

during day (10 h light) and a minimum 19°C at night and harvested 4-, 7- and 10-months after sowing (110, 407

215 and 285 days after sowing, respectively). At each time point, eight biological replicates were sampled 408

and used for metabolite and RNA extraction. Plant material for transcriptomic sequencing were likewise 409

grown in glass houses at the University of Copenhagen. Apical tips from six individual 8-month-old 410

E. cladocalyx seedlings were harvested, combined and snap frozen in liquid nitrogen. Seedlings for MALDI-411

MSI imaging were germinated in the glasshouse at the University of Melbourne on September 10, 2014 and 412

were grown at an average temperature of 23.3 ± 1.3°C during day and 19.6 ± 1.5°C at night. E. cladocalyx 413

expanded adult leaves, immature and mature flower buds, flowers and young fruit were harvested from 414

three trees growing at the University of Melbourne Parkville Campus (37.7964° S, 144.9612° E, January 415

2015), the Royal Botanic Gardens (37.8304° S, 144.9796° E, February 2015), and from the University of 416

Melbourne Creswick Campus (37.4221° S, 143.8992° E, February 2015). Samples were embedded and 417

frozen on site for MALDI-MSI analysis. Samples for LC-MS were frozen in liquid nitrogen and stored at −80°C 418

until extraction. 419

Metabolite extraction and Liquid Chromatography Mass Spectrometry (LC-MS) 420

Metabolites from E. cladocalyx (15-50 mg homogenized tissue) were extracted with cold aqueous 80% 421

MeOH. Leaf metabolites from infiltrated N. benthamiana plants expressing different combinations of gene 422

candidates were extracted with cold aqueous 85% MeOH from leaf discs (1.5 cm in diameter) harvested 5 d 423

after infiltration. All plant tissue was incubated on ice in cold MeOH for 5 minutes and extracts were 424

subsequently centrifuged (3 min, 10,000 g, 4°C) to sediment plant tissue. Extracts were diluted 5 times in 425

MilliQ water and filtered through a Durapore membrane (22 μm pore size, Merck Millipore) before 426

injection into an Agilent 1100 Series LC system coupled to a Bruker HCT-Ultra ion trap mass spectrometer 427

with an electrospray ionization source run in positive mode. Metabolites were separated on a Zorbax SB-428

C18 column (2.1 x 50 mm, 1.8 µm, Agilent) at 35°C applying a flow rate of 0.2 mL min-1. The mobile phase A 429

consisted of 0.1% (v/v) HCOOH and 50 µM NaCl and phase B consisted of 0.1% HCOOH in MeCN. The first 430

interval was 0-0.5 min isocratic with 2% B followed by two consecutive linear gradients: 0.5-7.5 min, 2-40% 431

B and 7.5-8.5 min, 40-90% B, and a second isocratic interval 8.5-11.5 min, 90% B. The mobile phase was 432

returned to 2% B from 11.5-18 min. The flow rate was raised to 0.3 mL min-1 in the interval 11.5-13.5 min. 433

LC-MS data was analyzed using Bruker Data Analysis 4.0 software. 434

MALDI-MSI and sample preparation 435

MALDI-MSI was carried out according to (Schmidt et al. 2018b). In short, the first fully expanded leaf of 436

E. cladocalyx seedlings (aged 139 DAS), fully expanded leaves from adult trees, and flower buds were 437

embedded in denatured albumin (boiled egg-white) and gently frozen over a bath of liquid nitrogen. 438

Albumin embedded tissue was cryosectioned (30 µm) (Leica CM3050 S cryostat, Leica Microsystems, 439

Wetzler, Germany), mounted onto double-sided carbon tape attached to glass slides and freeze-dried 440

overnight. 2,5-Dihydroxybenzoic acid (DHB) was used as the matrix and sublimed onto tissue sections using 441

a custom-built sublimation apparatus. MS images were acquired in positive ion mode using a Bruker SolariX 442

XR 7-Tesla Hybrid ESI/MALDI-FT-ICR-MS (Bruker Daltonik, Bremen, Germany). 443



12

A mass range of 100-2000 m/z was employed with the instrument set to broadband mode with a time 444

domain for acquisition of 2M providing an estimated resolving power of 130,000 at m/z 400. The laser was 445

set to 30-45% power using the minimum spot size (laser spot size approximately 10 × 15 µm), smart-walk 446

selected and random raster enabled resulting in ablation spots of approximately 35-40 µm in diameter, a 447

total 1500-2000 shots were fired per spectra at a frequency of 2 kHz within a 40 × 40 µm array. Optical 448

images of tissue sections were acquired using an Epson Photosmart 4480 flatbed scanner using a minimum 449

setting of 4800 d.p.i. Optical images are presented as an inverted image, generated in Adobe Photoshop 450

CS2 (Adobe). The data were analyzed using Compass FlexImaging 4.1 (Bruker), data was normalized using 451

RMS normalization and individual ion images were scaled as a percentage of maximum signal intensity to 452

enhance visualization. Potassium adducts of prunasin ([M + K]+ m/z 334.0691, calculated 334.0687, 1.2 ppm 453

error) and amygdalin ([M + K]+ m/z 496.1202, calc. 496.1216, 2.6 ppm error) were compared to authentic 454

standards, spotted and dried onto a separate glass slide. 455

Characterization of the E. cladocalyx cyanogenic glucoside biosynthetic system 456

To test the substrate, product and biosynthetic capacity, E. cladocalyx leaves of different developmental 457

stages, newly emerged apical tips, expanding and expanded leaves were harvested from 5-month-old 458

seedlings (biological duplicates) and incubated in L-[14C(U)]-phenylalanine (1.25 mCi, 321 mCi mmol-1; 459

Amersham Biosciences, GE Healthcare, England) and NADPH (0.1 mM) in a total volume of 165 μL for 12 h 460

at room temperature. At the end of incubation, cyanogenic glucosides were extracted in boiling MeOH 461

(80%, 1.5 mL, 5 min), defatted with two extractions using pentane (700 μL), filtered (0.22 µm low-binding 462

Durapore membrane, Millipore, United States), and 5 μL aliquots were spotted onto a TLC plate (Silica 463

gel60F254 0.2 mm thickness, Merck, Germany), which was developed in EtOAc:HOAc:MeOH:H2O (16:5:5:2, 464

v/v/v/v). Radioactively labeled cyanogenic glucosides were visualized using a STORM 840 PhosporImager 465

(Molecular Dynamics, United States) and compared to the position of an unlabeled, co-applied prunasin 466

standard (Møller et al. 2016) visualized by UV absorption. 467

RNA extraction 468

Total RNA was extracted from E. cladocalyx leaves (apical tips, expanding and expanded leaves) at three 469

different ages in biological triplicates using the SpectrumTM Plant Total RNA Kit (Sigma) according to the 470

manufacturer’s instruction with a few modifications: approximately 20 mg homogenized tissue was used 471

for extraction and centrifugation was carried out at 4°C. DNAse treatment was performed using RNAse-free 472

DNAse I from Omega Bio-tek. The integrity of RNA was verified on an Agilent 2100 Bioanalyzer instrument 473

using a RNA 6000 Nano Kit (Agilent Technologies) according to the manufacturer’s instructions and quantity 474

was estimated using a NanoDrop ND-1000 spectrophotometer (Saveen Werner, Sweden). 475

cDNA synthesis and quantitative PCR 476

cDNA was synthesized from 1.5 μg RNA using SuperScript IV VILO Master Mix (Invitrogen) according to the 477

manufacturer’s instructions. The cDNA was diluted 30 times and 4 μL was used as template in the qPCR 478

reactions (total volume 10 μL) using SeniFAST SYBR® No-ROX Kit (Bioline) and gene specific primers (final 479

concentration: 0.4 μM). Reactions were carried out in technical triplicates and triplicate non-template 480

controls were included for every primer pair. A serial dilution from a pool of cDNA was also included for all 481

primer pairs to correct for primer efficiency. Reactions were carried out on a CFX384 Touch Real-Time PCR 482

Detection System (Bio-Rad) using two-step cycling parameters: 95°C for 2 min, followed by 45 cycles of 483

95°C for 5 sec and then 60°C for 20 sec. Melt curve analysis was performed by gradually increasing the 484



13

temperature from 65°C to 95°C with 0.5°C raises. Data was analyzed using Bio-Rad CFX Manager 3.1 and 485

qbase+ version 3.1. Eight candidate reference genes previously used for gene expression analysis in 486

Eucalyptus species were selected from literature and primers were designed to match nucleotide 487

sequences from our transcriptome (Supplemental Table S2. Eight candidate reference genes were tested 488

for stability on 10 different samples using geNorm (Vandesompele et al. 2002). tRNA and MDH showed the 489

highest expression stability (Supplemental Fig. S4) and were thus used to normalize the expression of the 490

prunasin pathway genes. Primers specific for prunasin pathway transcripts are listed in Supplemental Table 491

S3. Normalized relative quantities were calculated according to Hellemans et al. (2007) by setting Cqreference 492

to 0. 493

Transcriptome analysis 494

RNA extracted from apical tips of E. cladocalyx was sent for transcriptomic sequencing (Hiseq2500, 100bp 495

paired-end) by Macrogen Inc. (Seoul, Republic of Korea). The transcriptome was de novo assembled, 496

functionally classified and expression levels were quantified by Sequentia Biotech (Bellaterra, Spain). In 497

brief, reads were quality checked and low quality reads were removed using Trimmomatic (v 0.33) (Bolger 498

et al. 2014) with a minimum length setting of 30 bp and a minimum quality score of 25. The high quality 499

reads were de novo assembled using the Trinity pipeline (Haas et al. 2013). Transcripts were translated 500

using Transdecoder with a minimum length of 50 amino acids and annotated by two different approaches: 501

by a BLAST search against a database of A. thaliana amino acid sequences and gene ontology (GO) 502

annotated using the InterproScan software (v 5.13.52.0) (Jones et al. 2014). To quantify the expression 503

levels, reads were mapped to the assembled transcriptome using STAR (Dobin et al. 2013) and the 504

expression levels were calculated with the software eXpress (v1.5.1) (Roberts and Pachter 2013). 505

Cloning of genes involved in prunasin biosynthesis 506

CYP79A125 was isolated from E. cladocalyx cDNA using primers that were designed based on CYP79A34 507

from E. yarraensis. CYP71B103 and UGT85A59 were isolated from E. cladocalyx cDNA using degenerate 508

primers that were designed based on the orthologous sequences in the E. grandis genome available on 509

Phytozome (Goodstein et al. 2012). Gene-specific primers with attB overhangs were subsequently used to 510

amplify the open reading frames for Gateway recombination into pDONR207 (Invitrogen). Primer 511

sequences are listed in Supplemental Table S4. CYP706C55 was ordered as a synthetic gene (GenScript) 512

with a C-terminal strep-tag. In addition, the full open reading frame of CYP706C55 was amplified from 513

cDNA. Candidate genes were cloned into the pJAM1502 expression vector (Luo et al. 2007) by Gateway 514

recombination. Candidate gene sequences in expression constructs were verified by sequencing (EZ-Seq, 515

Macrogen, The Netherlands). 516

Transient expression in Nicotiana benthamiana 517

Plasmids were introduced into Agrobacterium tumefaciens AGL1 (Lazo et al. 1991) by electroporation. 518

Electroporated cells were recovered in YEP (yeast extract peptone) media for 2 h at 28°C and selected on 519

YEP agar media containing 25 μg/mL rifampicin, 50 μg/mL carbenicillin and 50 μg/mL kanamycin. 520

Transformants were grown over night at 28°C, 225 rpm. Agrobacteria cells were harvested and re-521

suspended in MilliQ water to a final OD600 of 1. Equal volumes of agrobacteria suspensions were mixed and 522

introduced into N. benthamiana leaves (approximately 4-weeks-old) using a blunt syringe. An expression 523

construct carrying the ORF of p19 (Hearne et al. 1990) was always co-infiltrated to prevent post-524

transcriptional gene silencing (Lakatos et al. 2004; Voinnet et al. 1999). Two plants were infiltrated per 525



14

combination and infiltrations were repeated to confirm results. After infiltration, the plants were kept in 526

the greenhouse (21°C day/19°C night) until leaf material was harvested. 527

Microsomal preparation 528

Leaves from transiently transformed N. benthamiana were harvested 5 d after infiltration. Approximately 6 529

g of leaves were homogenized with 0.1 g polyvinylpolypyrrolidone/g fresh weight and 30 mL buffer (100 530

mM Tricine (pH 7.9), 250 mM sucrose, 50 mM NaCl, 2mM DTT, 1 tablet cOmpleteTM protease inhibitor 531

cocktail (Roche) per 200 mL buffer) using a mortar and pestle at 4°C. The homogenate was filtered through 532

two layers of nylon cloth (22 μm pore size). Samples were centrifuged at 15,000 g for 15 min at 4°C. The 533

supernatant was subsequently applied to ultracentrifugation for 1 h at 164244 g at 4°C. The microsomal 534

pellets were washed in buffer and re-suspended in 200 µL re-suspension buffer (50 mM Tricine (pH 7.9), 20 535

mM NaCl, 2 mM DTT). 536

Microsomal assays 537

The assay reactions (total volume 25 uL) consisted of 10 uL microsomes, L-[14C(U)]-phenylalanine (0.25 μCi, 538

specific activity 487 mCi mmol-1, Perkin-Elmer), 1 mM NADPH and buffer (50 mM Tricine (pH 7.5), 100 mM 539

NaCl). Reaction mixtures were incubated for 30 min at 30°C, 300 rpm. The reactions were stopped by direct 540

application to a TLC plate (Silica gel 60F254 0.2 mm thickness, Merck, Germany). Substrate and products 541

were separated using a mobile phase consisting of n-pentane: EtOAc: MeOH (16:2:1, v/v/v). Migration of 542

phenylacetaldoxime, phenylacetonitrile and mandelonitrile were determined by application of unlabeled 543

authentic standard, which were visualized by their UV absorbance. Analysis of radiolabeled products was 544

carried out by exposure of the TLC plate to a Storage Phosphor Screen (GE Healthcare) for 8 d followed by 545

visualization using a Typhoon TRIO Variable Mode Imager (GE Healthcare). 546

Phylogenetic analysis 547

Named and functionally characterized CYPs from literature were subjected to phylogenetic analysis 548

together with the CYPs characterized here. Accession numbers and references are listed in Supplemental 549

Table S1. Amino acid sequences were aligned in MEGA7 (Kumar et al. 2016) using MUSCLE with the default 550

setting. A Neighbor Joining tree was generated with n = 1000 bootstrap replicates. 551

Statistical Analyses 552

Metabolite data was analyzed in SigmaPlot (ver. 13.0) using one-way ANOVA. Normality (Kolmogorov-553

Smirnov) and equal variance (Brown-Forsythe) was tested to a cutoff of P > 0.02 and P > 0.04, respectively, 554

and when assumptions were not met, the data was transformed by natural logarithm. Pearson correlation 555

was used to analyze gene expression. 556

Accession numbers 557

Sequence data for the prunasin biosynthetic genes identified in this study can be found in Genbank under 558

following accession numbers: CYP79A125 (MH974543), CYP706C55 (MH974544), CYP71B103 (MH974545) 559

and UGT85A59 (MH974546). 560

Supplemental Data 561

Supplemental Figure S1. Additional LC-MS data for prunasin production in N. benthamiana 562

Supplemental Figure S2. LC-MS data for glucosylated phenylacetaldoxime 563



15

Supplemental Figure S3. LC-MS data for malonic acid derivative of prunasin 564

Supplemental Figure S4. geNorm analysis of candidate reference gene stabilities 565

Supplemental Table S1. Accession numbers and references for sequences used in the phylogenetic analysis 566

Supplemental Table S2. Primer sequences and references for tested candidate reference genes 567

Supplemental Table S3. Primer sequences used for gene expression analysis of the prunasin pathway genes 568

Supplemental Table S4. Cloning primers 569

Acknowledgements 570

The authors gratefully acknowledge Dr. Antanas Spokevicius for pre-publication access to the 571

transcriptomic results of E. cladocalyx and E. globulus. Ian Woodrow is thanked for hosting JSFZ during her 572

stay at The University of Melbourne. David Vernon is heartily thanked for his generous support of wildlife 573

research and encouragement to early career scientists. 574

575

Figure legends 576

Figure 1. Prunasin and amygdalin accumulation in E. cladocalyx. A. Prunasin (dark grey) and amygdalin 577

(light grey) content in expanded leaves from seedling and adult trees, immature and mature flower buds, 578

flowers and fruits. Bars represent mean with standard error. The low amounts of amygdalin present are 579

shown in more detail in the insert. Letters denote significance at P < 0.05 by one-way ANOVA analysis. B. 580

Structures of prunasin and amygdalin. 581

Figure 2. Localization of cyanogenic glucosides in E. cladocalyx tissue. Cross section of seedling dorsiventral, 582

A, and adult isobilateral, B, leaves prepared for matrix-assisted laser desorption ionization-mass 583

spectrometry imaging (MALDI-MSI), with corresponding ion maps of prunasin (yellow; m/z 334.0687; 584

[M+K]+) localizing to the mesophyll and vascular tissue. Longitudinal section of a flower bud, C, prepared 585

for MALDI-MSI with corresponding ion maps of prunasin and amygdalin (pink; m/z 496.1216; [M+K]+). 586

Enlarged portions of the flower bud, D, with the section overlaid are shown in the corresponding boxes 1 587

and 2. Prunasin ions distributed in the ground tissue within the hypanthium, stamens, and in the 588

operculum. Co-localization with amygdalin occurs in the filaments and around the staminal ring (indicated 589

by arrows). All images are root mean square (RMS) normalized, with internal scaling of 100% for prunasin 590

and 25% for amygdalin. Scale bar = 500 μM. Abbreviations: EG, embedded gland; PM, palisade; SM, spongy 591

mesophyll; VB, vascular bundle. 592

Figure 3. E. cladocalyx leaves of different developmental stages can synthesize prunasin from 593

phenylalanine. A. E. cladocalyx seedling. The tissues used for administration with L-[14C]-phenylalanine are 594

indicated. B. TLC plate showing the production of prunasin from L-[14C]-phenylalanine in apical tips, 1st 595

and 2nd expanding leaves and fully expanded leaves. 596

Figure 4. Prunasin biosynthesis is catalyzed by three CYPs and a UGT. A. LC-MS extracted ion 597

chromatograms for prunasin ([M+Na]+ m/z 318) of extracts from N. benthamiana leaves transiently 598

expressing candidate genes. B. Microsomes prepared from N. benthamiana leaves transiently expressing 599



16

candidate CYP genes were assayed with 14C-phenylalanine and products were separated by TLC. C. 600

Proposed pathway for prunasin synthesis in E. cladocalyx. 601

Figure 5. Phylogenetic analysis of functionally characterized CYPs belonging to the CYP79, CYP71, CYP706 602

and CYP736 families, including members involved in cyanogenic glucoside biosynthesis (indicated by * ). 603

Phylogeny was inferred using the Neighbour-Joining method with n = 1000 bootstrap replicates. Full 604

species names, Genbank accession numbers and references are listed in Supplemental Table S1. 605

Figure 6. Prunasin accumulation and expression of the biosynthetic genes in 4, 7 and 10 months old 606

E. cladocalyx plants (n = 6-8). A. Prunasin accumulation in apical tips, expanding leaves and expanded 607

leaves. Bars represent mean +/- standard error. Letters denote significance at P < 0.05 by one-way ANOVA 608

analysis. B. Normalized gene expression of CYP79A125, CYP706C55, CYP71B103 and UGT85A59 in apical 609

tips, expanding leaves and expanded leaves. Symbols represent biological replicates (n = 3) C. Spearman 610

correlation analysis of gene expression showing co-regulation of some genes. 611

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Figure 1. Prunasin and amygdalin accumulation in E. cladocalyx. A. Prunasin (dark grey) and amygdalin (light grey) content in expanded leaves from seedling and adult trees, immature and mature flower buds, flowers and fruits. Bars represent mean with standard error. The low amounts of amygdalin present are shown in more detail in the insert. Letters denote significance at P < 0.05 by one-way ANOVA analysis. B. Structures of prunasin and amygdalin.

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ab

ac

c

bc

ab

a a



Bo

x 1

. B

ox

2.

Filament

Filament

Anther

Operculum

EG

1.

2.

Prunasin Amygdalin Overlay C

D

B

A

Placenta

Operculum

Stamens (detached)

1.

2.

Ovules

Pedicle

Staminal ring

Upper Epidermis

PM

PM

SM

VB

EG

VB Cuticle

Epidermis

Lower Epidermis

Prunasin

Figure 2. Localization of cyanogenic glucosides in Eucalyputs cladocalyx tissue. Cross section of seedling dorsiventral, A, and adult isobilateral, B, leaves prepared for matrix-assisted laser desorption ionisation-mass spectrometry imaging (MALDI-MSI), with corresponding ion maps of prunasin (yellow; m/z 334.0687; [M+K]+) localizing to the mesophyll and vascular tissue. Longitudinal section of a flower bud C prepared for MALDI-MSI with corresponding ion maps of prunasin and amygdalin (pink; m/z 496.1216; [M+K]+). Enlarged portions of the flower bud, D, with the section overlaid are shown in the corresponding boxes 1 and 2. Prunasin ions distributed in the ground tissue within the hypanthium, stamens, and in the operculum. Co-localisation with amygdalin occurs in the filaments and around the staminal ring (indicated by arrows). All images root mean square (RMS) normalized, with internal scaling of 100% for prunasin and 25% for amydgalin. Scale bar = 500 µM. Abbreviations: EG, embedded gland; PM, palisade; SM, spongy mesophyll; VB, vascular bundle.



Figure 3. E. cladocalyx leaves of different developmental stages can synthesize prunasin from phenylalanine. A. E. cladocalyx seedling. The tissues used for administration with L-[14C]-phenylalanine are indicated. B. TLC plate showing the production of prunasin from L-[14C]-phenylalanine in apical tips, 1st and 2nd expanding leaves and fully expanded leaves.

Prunasin

Phenylalanine

Tip

1st expanding leaf

2nd expanding leaf

Expanded leaf

B A



Figure 4. Prunasin biosynthesis is catalyzed by three CYPs and a UGT. A. LC-MS extracted ion chromatograms for prunasin ([M+Na]+ m/z 318) of extracts from N. benthamiana leaves transiently expressing candidate genes. B. Microsomes prepared from N. benthamiana leaves transiently expressing candidate CYP genes were assayed with 14C-phenylalanine and products were separated by TLC. C. Proposed pathway for prunasin synthesis in E. cladocalyx.

A

phenylacetonitrile

phenylacetaldoxime

CYP79A125 + + + + + + +

+ - - -

-

B

- - -

mandelonitrile

benzoic acid

CYP706C55

CYP71B103

C

Phenylalanine

Prunasin

Mandelonitrile

Phenylacetonitrile

Phenylacetaldoxime

CYP79A125

CYP706C55

CYP71B103

UGT85A59

CYP706C55 + CYP71B103 + UGT85A59

CYP79A125 + CYP706C55 + CYP71B103 + UGT85A59

Inte

nsity

3x10

8 (E

IC m

/z 3

18

)

Negative control

Prunasin standard

CYP79A125 + CYP71B103 + UGT85A59

CYP79A125 + CYP706C55 + UGT85A59

CYP79A125 + CYP706C55 + CYP71B103

4 6 Time [min]



Figure 5. Phylogenetic analysis of functionally characterized CYPs belonging to the CYP79, CYP71, CYP706 and CYP736 families, including members involved in cyanogenic glucoside biosynthesis (indicated by * ). Phylogeny was inferred using the Neighbour-Joining method with n = 1000 bootstrap replicates. Full species names, Genbank accession numbers and references are listed in Supplemental Table S1.

CY

P71A

V3 L

. sativa

CYP79E1 T. maritima

0.10

* *

* *

* *

*

*

*

*

*

*

*

*

*

*

*

* *

* *

*



CYP71B103

No

rma

lize

d R

ela

tive

ex

pre

ss

ion

0

2

4

6

8

10

12UGT85A59

CYP79A125

No

rma

lize

d R

ela

tive

ex

pre

ss

ion

0

1

2

3

4

5

6

CYP706C55

Tip

s

Exp

an

din

g

Exp

an

de

d

Tip

s

Exp

an

din

g

Exp

an

de

d

Tip

s

Exp

an

din

g

Exp

an

de

d

Pru

na

sin

(u

g/m

gd

w)

0

10

20

30

40

50

60

4 months 7 months 10 months 4 months 7 months 10 months

*Significant at P < 0.05 **Significant at P < 0.01

CY

P79A

125

CY

P706C

55

CY

P71B

103

UG

T85A

59

CYP79A125 - 0.552** 0.489** 0.205

CYP706C55 - 0.475* 0.0148

CYP71B103 - -0.0950

UGT85A59 -

TipsExpanding

Expanded

Figure 6. Prunasin accumulation and expression of the biosynthetic genes in 4, 7 and 10 months old E. cladocalyx plants (n = 6-8). A. Prunasin accumulation in apical tips, expanding leaves and expanded leaves. Bars represent mean +/- standard error. Letters denote significance at P < 0.05 by one-way ANOVA analysis. B. Normalized gene expression of CYP79A125, CYP706C55, CYP71B103 and UGT85A59 in apical tips, expanding leaves and expanded leaves. Symbols represent biological replicates (n = 3) C. Spearman correlation analysis of gene expression showing co-regulation of some genes.

B

A C 4 months 7 months 10 months

a

ab

b

c c c c c c



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Bassard, J. E., Møller, B. L., and Laursen, T. (2017). "Assembly of Dynamic P450-Mediated Metabolons-Order Versus Chaos." Curr MolBiol Rep, 3(1), 37-51.


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Bjarnholt, N., Neilson, E. H. J., Crocoll, C., Jørgensen, K., Motawia, M. S., Olsen, C. E., Dixon, D. P., Edwards, R., and Møller, B. L.(2018). "Glutathione transferases catalyze recycling of auto-toxic cyanogenic glucosides in sorghum." Plant J, 94(6), 1109-1125.


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Boughton, B. A., Thinagaran, D., Sarabia, D., Bacic, A., and Roessner, U. (2016). "Mass spectrometry imaging for plant biology: areview." Phytochem Rev, 15, 445-488.


Burchell, B., Nebert, D. W., Nelson, D. R., Bock, K. W., Iyanagi, T., Jansen, P. L. M., Lancet, D., Mulder, G. J., Chowdhury, J. R., Siest, G.,Tephly, T. R., and Mackenzie, P. I. (1991). "The UDP Glucuronosyltransferase Gene Superfamily: Suggested Nomenclature Based onEvolutionary Divergence." DNA Cell Biol, 10(7), 487-494.


Busk, P. K., and Møller, B. L. (2002). "Dhurrin synthesis in sorghum is regulated at the transcriptional level and induced by nitrogenfertilization in older plants." Plant Physiol, 129(3), 1222-31.


Clausen, M., Kannangara, R. M., Olsen, C. E., Blomstedt, C. K., Gleadow, R. M., Jorgensen, K., Bak, S., Motawie, M. S., and Møller, B. L.(2015). "The bifurcation of the cyanogenic glucoside and glucosinolate biosynthetic pathways." Plant J, 84(3), 558-73.


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Darbani, B., Motawia, M. S., Olsen, C. E., Nour-Eldin, H. H., Møller, B. L., and Rook, F. (2016). "The biosynthetic gene cluster for thecyanogenic glucoside dhurrin in Sorghum bicolor contains its co-expressed vacuolar MATE transporter." Sci Rep, 6, 37079.


Del Cueto, J., Ionescu, I. A., Pičmanová, M., Gericke, O., Motawia, M. S., Olsen, C. E., Campoy, J. A., Dicenta, F., Møller, B. L., andSanchez-Perez, R. (2017). "Cyanogenic Glucosides and Derivatives in Almond and Sweet Cherry Flower Buds from Dormancy toFlowering." Front Plant Sci, 8, 800.


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https://www.ncbi.nlm.nih.gov/pubmed?term=Noge%2C K%2E%2C Abe%2C M%2E%2C and Tamogami%2C S%2E %282011%29%2E %22Phenylacetonitrile from the giant knotweed%2C Fallopia sachalinensis%2C infested by the Japanese beetle%2C Popillia japonica%2C is induced by exogenous methyl jasmonate%2E%22 Molecules&dopt=abstract

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Reconfigured cyanogenic glucoside biosynthesis in ...example of convergent evolution in the biosynthesis of cyanogenic . 67. glucosides in plants. 68. 69. Introduction . 70. Cyanogenic