Abstract Cyanogenic glycosides are ancient bio-

molecules found in more than 2,650 higher plant

species as well as in a few arthropod species.

Cyanogenic glycosides are amino acid-derived

b-glycosides of a-hydroxynitriles. In analogy to

cyanogenic plants, cyanogenic arthropods may use

cyanogenic glycosides as defence compounds.

Many of these arthropod species have been shown

to de novo synthesize cyanogenic glycosides by

biochemical pathways that involve identical

intermediates to those known from plants, while

the ability to sequester cyanogenic glycosides

appears to be restricted to Lepidopteran species.

In plants, two atypical multifunctional cyto-

chromes P450 and a soluble family 1 glycosyl-

transferase form a metabolon to facilitate

channelling of the otherwise toxic and reactive

intermediates to the end product in the pathway,

the cyanogenic glycoside. The glucosinolate path-

way present in Brassicales and the pathway for

cyanoalk(en)yl glucoside synthesis such as rhod-

iocyanosides A and D in Lotus japonicus exem-

plify how cytochromes P450 in the course of

evolution may be recruited for novel pathways.

The use of metabolic engineering using cyto-

chromes P450 involved in biosynthesis of cyano-

genic glycosides allows for the generation of

acyanogenic cassava plants or cyanogenic Arabid-

opsis thaliana plants as well as L. japonicus and

A. thaliana plants with altered cyanogenic,

cyanoalkenyl or glucosinolate profiles.

Keywords Metabolic engineering ÆMetabolons Æ CYP79 Æ Systems biology ÆPlant–insect interactions

Cyanogenic glycosides are ancient biomolecules

Cyanogenic glycosides (A1) are amino acid-

derived b-glycosides of a-hydroxynitriles. They

have been found in more than 2,650 higher plant

S. Bak (&) Æ M. Morant Æ A. V. Morant ÆS. Saito Æ N. Bjarnholt Æ M. Zagrobelny ÆK. Jørgensen Æ S. Osmani Æ H. T. Simonsen ÆR. S. Perez Æ T. B. van Heeswijck Æ B. L. MøllerPlant Biochemistry Laboratory, Department of PlantBiology, and Center for Molecular Plant Physiology(PlaCe), Royal Veterinary and AgriculturalUniversity, 40 Thorvaldsensvej, 1871 FrederiksbergCopenhagen, Denmarke-mail: bak@kvl.dk

S. M. PaquetteDepartment of Biological Structure, University ofWashington, HSB G-514/Box 357420, Seattle, WA98195-7420, USA

B. JørgensenResearch Group Cell Wall Biology and MolecularVirology, Department of Genetics andBiotechnology, Danish Institute of AgriculturalSciences, 40 Thorvaldsensvej, 1871 FrederiksbergCopenhagen, Denmark

Phytochem Rev (2006) 5:309–329

DOI 10.1007/s11101-006-9033-1

123

Cyanogenic glycosides: a case study for evolutionand application of cytochromes P450

Søren Bak Æ Susanne Michelle Paquette Æ Marc Morant Æ Anne Vinther Morant ÆShigeki Saito Æ Nanna Bjarnholt Æ Mika Zagrobelny Æ Kirsten Jørgensen ÆSarah Osmani Æ Henrik Toft Simonsen Æ Raquel Sanchez Perez ÆTorbjørn Bordier van Heeswijck Æ Bodil Jørgensen Æ Birger Lindberg Møller

Received: 24 May 2006 / Accepted: 22 September 2006 / Published online: 15 November 2006� Springer Science+Business Media B.V. 2006

species distributed among 130 families in pteri-

dophytes (ferns), gymnosperms and angiosperms

(flowering plants; Conn 1981; Siegler and Brinker

1993) implying that in plants the ability to

synthesize cyanogenic glycosides is at least

300 million years old. The widespread occurrence

of cyanogenic glycosides implies that they are

ancient biomolecules in terrestrial plants. Cyano-

genic glycosides are classified as phytoanticipins.

Upon disruption of plant tissue containing cya-

nogenic glycosides they are degraded by b-glyco-

sidases and a-hydroxynitrilases, resulting in

release of toxic hydrogen cyanide, an aldehyde

or ketone and glucose. This binary system—two

sets of components that separately are chemically

inert—provides plants with an immediate chem-

ical defence response to herbivores and patho-

gens that cause tissue damage (Møller and Seigler

1998; Jones et al. 2000; Morant et al. 2003; Nielsen

and Møller 2004; Poulton and Møller 1989).

Cyanogenic ferns and gymnosperm species con-

tain aromatic cyanogenic glycosides derived from

either tyrosine or phenylalanine, whereas angio-

sperms contain aliphatic as well as aromatic

cyanogenic glycosides derived from isoleucine/

valine/leucine or tyrosine/phenylalanine, respec-

tively (Fig. 1). In addition, a few plant species

such as Passiflora (Malphigiales, eurosids I) con-

tain cyanogenic glycosides derived from the

non-protein amino acid cyclopentenyl glycine

(Jaroszewski et al. 2002a, b). The specific pres-

ence of aromatic cyanogenic glycosides in ferns

and gymnosperms indicate that the cyanogenic

glycosides initially in nature were aromatic, and

that these served as progenitors for aliphatic

cyanogenic glycosides. This evolutionary path is

supported by the fact that ancestral angiosperms

like Magnoliales contain tyrosine-derived cyano-

genic glycosides. Within monocotyledons, Liliales

are known to contain aromatic cyanogenic glyco-

sides, and within Poales both aromatic and

aliphatic cyanogenic glycosides occur. In eudicots,

a wide distribution of aromatic as well as aliphatic

cyanogenic glycosides is observed, but the amino

acid precursor used within a given family is

generally conserved. The presence of aliphatic

cyanogenic glycosides in Poales and eudicots

raises the question of whether aliphatic

cyanogenic glycosides evolved independently at

least twice or whether they evolved before the

radiation of monocotyledons and eudicots. Typ-

ically, plants contain cyanogenic glycosides

derived from a single amino acid: tyrosine,

phenylalanine, leucine, valine or isoleucine,

except that valine- and isoleucine-derived cyano-

genic glycosides typically co-occur. It is only in

some polyploid species like hexaploid wheat

(Triticum aestivum, Poales) that aromatic and

aliphatic cyanogenic glycosides co-occur, but in

this case the co-occurrence is a result of the

allopolyploidity of the hexaploid wheat genome,

where the presence of either aliphatic or aromatic

cyanogenic glycosides in the diploid and tetra-

ploid ancestors have been retained during domes-

tication of wheat (our unpublished results).

Cyanogenic glycosides are not only found in

plants, but are also present in animals within a

limited number of anthropod clades (reviewed by

Zagrobelny et al. 2004; Fig. 2). A few species of

Diploda (millipedes), Chilopoda (centipedes),

Coleoptera (beetles) and Heteroptera (true bugs)

synthesize aromatic cyanogenic glycosides while

more than 200 species within Lepidoptera (but-

terflies and moths) synthesize aliphatic cyano-

genic glycosides. Accordingly, older animal

lineages and plant lineages contain aromatic

cyanogenic glycosides while relatively more

recent lineages as Lepidopteran species and

angiosperms have acquired the capacity to

contain aliphatic cyanogenic glycosides.

In analogy to plants, cyanogenic animals store

the cyanogenic glycosides in their defensive

sections and use them in their own defence

against predators. Within Lepidoptera, several

species have been identified that are able to

sequester cyanogenic glycosides from their host

plants and in some cases also are able to carry out

de novo synthesis of cyanogenic glycosides when

the amount of cyanogenic glycosides in the host

plant is not sufficient to maintain desired levels in

the insect. This implies a close co-evolution of

Lepidopteran species with their preferred host

plants, and that the ability to synthesize cyano-

genic glycosides has been lost in some Lepidop-

teran species. The radiation of Lepidopteran

species beginning 210 million years ago on a

gymnosperm dominated flora precedes the

radiation of angiosperms beginning 140 million

310 Phytochem Rev (2006) 5:309–329

123

Pteridophytes~320 MYA(aromatic)

Gymnosperms~360 MYA(aromatic)

Angiosperms~140 MYA(aromatic and aliphatic)

Monocots Eucots

Core Eudicots

Rosids

Asterids

Eurosids I Eurosids II

Euasterids IEuasterids II

Poales

Liliales

Magnoliales

Alismatales

Bryophytes~425 MYA

Fig. 1 Schematicdiagram of the evolutionof higher plants and theoccurrence of aromaticand aliphatic cyanogenicglycosides. Only lineagesreferred to in the text arerepresented. Splits inmillion years ago betweenmajor taxonomic groupsare according to Willisand McElwain (2002)

Arthropoda<600 MYA

Hexapoda~500 MYA

Diplopoda(at least 46 species)(millipedes)~520 MYA

Chilopoda(at least 7 species)(centipedes)~550 MYA

Insecta~390 MYA

3 other groups

Pterygota

2 other groups

Neoptera

3 other groups

Endopterygota

15 othergroups

Lepidoptera(at least 200 species)(butterflies and moths)~210 MYA

10 other groups

Coleoptera(at least 3 species)(beetles)~300 MYA

Hemiptera

Heteroptera(at least 1 species)(true bugs)~220 MYA

3 other groups

Aromatic Cyanogenic glycosides

Aliphatic Cyanogenic glycosides

8 other groups

Fig. 2 Simplifiedevolutionary tree forArthropoda. Groupsincluding species thatcontain aromaticcyanogenic glycosides(red) and groupsincluding species thatcontain aliphaticcyanogenic glycosides(blue) are shown. Splits inmillion years ago betweenmajor taxonomic groupsare according toLabandeira and Sepkoski(1993) and Regier et al.(2005). Adapted fromZagrobelny et al. (2004)

Phytochem Rev (2006) 5:309–329 311

123

years ago (Labandeira 1993; Labandeira et al.

1994) implying that the ability to de novo

synthesize aliphatic cyanogenic glycosides in

some Lepidopteran species has been replaced by

sequestration as angiosperms containing aliphatic

cyanogenic glycosides proliferated.

The ability of Lepidoptera species to feed on

cyanogenic plants relies on the ability to metab-

olize toxic products formed by catabolism of

cyanogenic glycosides. While most animals rely

on detoxification of HCN by either b-cyanoala-

nine synthase or rhodanese, some Lepidopteran

species, like the neotropical butterfly (Heliconius

sara) have acquired an additional route to metab-

olize the cyanogenic glycosides via thiol

compounds. H. sara is able to sequester the

cyclopentenyl-derived cyanogenic glucoside,

epivolkenin (B4) from its host plant Passiflora

auriculata. In addition, it possesses the ability to

de novo synthesize the aliphatic cyanogenic

glycosides linamarin and lotaustralin. H. sara

metabolizes epivolkenin to the corresponding

thiol compound sarauriculatin by enzymatic

replacement of the nitrile group by a thiol. The

precise nature of this reaction is not known, but it

may enable H. sara to avoid release of free

hydrogen cyanide (Engler et al. 2000). The

replacement and possible ability to metabolize

the nitrile function into, e.g. ammonia and carbon

dioxide may constitute a nitrogen reservoir to

optimize the insect’s primary metabolism. This

adds yet another layer of complexity to the

co-evolution of cyanogenic glycoside metabolism

in insects and plants.

Biosynthesis of cyanogenic glycosides in higherplants and arthropods

In plants, the biosynthetic pathway for cyano-

genic glycosides is highly channelled (Møller and

Conn 1980) and catalysed by two unusual multi-

functional cytochromes P450 (P450) (Halkier and

Møller 1991; Sibbesen et al. 1994; Koch et al.

1995; Halkier et al. 1995; Sibbesen et al. 1995;

Kahn et al. 1997; Bak et al. 1998a) and a UDP-

glucosyl transferase (UGT; Jones et al. 1999;

Hansen et al. 2003; Fig. 3). The enzymes and the

corresponding structural genes were initially

identified from sorghum (Sorghum bicolor,

Poales) which synthesizes the tyrosine-derived

cyanogenic glucoside dhurrin (B3). While plant

cytochromes P450 are generally N-terminally

anchored membrane proteins, plant UGTs are

generally regarded as soluble cytosolic enzymes.

The first committed step in the dhurrin pathway

in sorghum is catalysed by the multifunc-

tional CYP79A1. This P450 catalyses two sequen-

tial N-hydroxylations of tyrosine followed by

a dehydration and a de-carboxylation reac-

tion to produce the corresponding aldoxime,

(Z)-p-hydroxyphenyl acetaldoxime. The second

cytochrome P450 in the pathway, CYP71E1,

catalyses a NADPH-dependent dehydration reac-

tion followed by a C-hydroxylation to produce the

cyanohydrin p-hydroxymandelonitrile, which is

HO

COOH

NH2

CYP79A1

HO

NOH

CYP71E1

HO

OH

CN

UGT85B1

HO

O

CN O

HO

OHOH

HO

Dhurrin

Fig. 3 The biosynthetic pathway of the tyrosine-derivedcyanogenic glucoside dhurrin in sorghum

312 Phytochem Rev (2006) 5:309–329

123

subsequently glycosylated by the family 1 glyco-

syltransferase UGT85B1 to produce dhurrin.

In vitro experiments involving simultaneous

administration of dual radiolabelled intermedi-

ates demonstrated that the dhurrin pathway is

highly channelled (Møller and Conn 1980). Our

working hypothesis is that CYP79A1 and

CYP71E1 and the otherwise soluble UGT85B1

form a metabolon, a multienzyme complex, to

facilitate channelling of the toxic and unstable

intermediates in the pathway (Nielsen and Møller

2004; Winkel 2004; Jørgensen et al. 2005b; Fig. 4).

Evidence for this has been obtained by two

independent approaches. Firstly, transient expres-

sion in sorghum epidermal cells of one or more of

the three biosynthetic enzymes tagged C-termi-

nally to spectral variants of green fluorescent

protein showed that each of the fusion proteins

were catalytically active. Nevertheless, when the

CYP79A1 and CYP71E1 were tagged simulta-

neously, no dhurrin production was observed

(Nielsen and Møller 2004). This suggested that

direct interaction was essential for dhurrin

production and was impeded due to the pres-

ence of bulky tags. The fluorescence shifted

from being cytosolic to ER located when GFP

tagged UGT85B1 was co-expressed with

native CYP79A1 and CYP71E1, showing that

UGT85B1 couples with CYP79A1 and CYP71E1

to form a multienzyme complex. In a second set

of experiments, co-expression of CYP79A1 and

CYP71E1 in Arabidopsis thaliana prevented

metabolic cross-talk with glucosinolate (A2) syn-

thesis whereas expression of CYP79A1 resulted

in efficient production of the tyrosine-

derived glucosinolate p-hydroxybenzylglucosino-

late (Kristensen et al. 2005). Again this suggests

that in vivo, CYP79A1, CYP71E1 and UGT85B1

form a tight complex that deprives the oxime

intermediate of access to the glucosinolate-pro-

ducing enzyme system. Evolution of a metabolon

for dhurrin synthesis incorporating the UGT85B1

enzyme would appear essential to ensure rapid

glucosylation of the toxic p-hydroxymandelonit-

rile intermediate and prevent its dissociation into

hydrogen cyanide and p-hydroxybenzaldehyde.

Fig. 4 Model of the metabolon responsible for dhurrinsynthesis in sorghum and a molecular model of the UDP-glucosyl transferase UGT85B1. (A) The metabolon isthought to include the two membrane-anchored cyto-chrome P450s, CYP79A1 and CYP71E1, the membrane-anchored cytochrome P450 reductase and the solubleUDP-glucosyl transferase, UGT85B1. (B) In the model ofUGT85B1, a-helices are shown in purple and b-sheets inturquoise. N and C termini are indicated. Substrates are

shown as ball-and-stick presentations in the catalyticpocket of UGT85B1 modelled with the donor moleculeUDP-Glc and the acceptor p-hydroxymandelonitrile.Green star position of removed loop A, yellow starsbeginning and ending of removed loop B, purple starsbeginning and ending of removed loop C, blue star positionof removed loop D. The loop positions are defined inThorsoe et al. (2005, Fig. 1)

Phytochem Rev (2006) 5:309–329 313

123

Currently, crystal structures of plant cytochromes

P450 are not available hampering the elucidation

of how CYP79A1 and CYP71E1 couple. How-

ever, a structural model of sorghum UGT85B1

was built based on hydrophobic cluster analysis

and the crystal structures of two bacterial GTs,

GtfA and GtfB, which each showed approxi-

mately 15% overall amino acid sequence identity

to UGT85B1 (Thorsoe et al. 2005). Despite the

overall low sequence identity, the UGT85B1

model enabled predictions about amino acid

residues important for catalysis and sugar donor

specificity. Furthermore, the UGT85B1 model

identified a hyper variable loop that contained a

hydrophobic patch that may mediating binding of

UGT85B1 to CYP79A1, and CYP71E1 within the

dhurrin metabolon. The mechanism that enables

docking of UGT85B1 into a metabolon is now

being investigated by molecular modelling and

site-directed mutagenesis (Thorsoe et al. 2005)

combined with reconstitution experiments con-

ducted using nanodisc technology (Bayburt and

Sligar 2003) where CYP79A1 and CYP71E1 have

been assembled into nanodiscs together with

sorghum cytochrome P450 reductase.

Many cyanogenic arthropod species have been

shown to de novo synthesize cyanogenic

glycosides by biochemical pathways that involve

similar or identical intermediates to those known

from plants, indicating that a homologous

pathway is present in animals (Duffey 1981;

Davis and Nahrstedt 1987; Zagrobelny et al.

2004; Feyereisen 2005). Older studies have shown

that all of the radiolabelled amino acid, N-

hydroxy amino acid, oxime and nitrile precursors

injected into Heliconius and Zygaena larvae

species or millipedes (Duffey and Towers 1978;

Holzkamp and Nahrstedt 1994) were not metab-

olized into cyanogenic glycosides. Most of these

precursors are now known to be highly unstable

compounds that are readily metabolized and

detoxified in planta (Bak et al. 2000). In addition,

the detection methods used were indirect, and

less sensitive than the LC-MS methods used in

the more recent plant studies. Feeding experi-

ments with Zygaena filipendulae larvae following

application of radiolabelled valine and isoleucine

directly to leaf surfaces of the host plant showed

that in the larvae the radiolabelled valine and

isoleucine ingested with the leaves are not

detected in intermediates but primarily ends up

in the cyanogenic glycosides linamarin and

lotaustralin. This supports a channelled pathway

in insects (Zagrobelny et al. in press) as seen in

plants.

Phylogeny of CYP79s in higher plants

CYP79 orthologs have been identified and iso-

lated from a number of cyanogenic angiosperms

(Fig. 5) using bioinformatics approaches, and

their function confirmed by reconstitution exper-

iments with heterologously expressed enzymes

(Halkier et al. 1996; Bak et al. 1999, 2000;

Andersen et al. 2000; Nielsen and Møller 2000).

No clear orthologs of sorghum CYP71E1 and

UGT85B1 have been reported. Nor have any

genes involved in cyanogenesis been identified in

arthropods. Accordingly, information on evolu-

tion of cyanogenic glycosides at the molecular

level is currently restricted to the CYP79 family in

plants. Phylogenetic reconstruction, using CYP79

sequences and selected clan 71 family members

Fig. 5 Neighbour-join phylogenetic tree illustrating thephylogenetic relationships of the P450s involved incyanogenesis against a background of A-type cytochromeP450 sequences. Sequences are from monocots (blue),eudicots (red), gymnosperms (purple) and moss (green).Protein sequences were taken from GenBank and http://drnelson.utmem.edu/biblioD.html and aligned usingClustalX 1.83, and phylogenetic analysis was carried outusing MEGA 3.1 (Kumar et al. 2004). Bootstrap values areindicated in % and based on 1,000 iterations and the treehas been rooted with CYP51s as the outgroup. Bootstrapvalues less than 50% are excluded from the tree. Thealignment for this sequence can be accessed in PDF formatat http://www.p450.kvl.dk/Review2006/Figure5_Align-ment.pdf. Ao Asparagus officinalis, At Arabidopsis thali-ana, Bs Berberis stolonifera, Cj Coptis japonica, EcEschscholzia californica, Ga Gossypium arboreum, GeGlycyrrhiza echinata, Gm Glycine max, Hv Hordeumvulgare, Le Lycopersicon esculentum, Lr Lolium rigidum,Ls Liquidambar styraciflua, Ma Musa acuminata, MeManihot esculenta, Ms Medicago sativa, Mt Medicagotruncatula, Nt Nicotiana tabacum, Os Oryza sativa, PaPersea americana, Pc Pyrus communis, Ph Petunia xhybrida, Pp Physcomitrella patens, Pr Pinus radiata, PsPisum sativum, Psa Pastinaca sativa, Pt Pinus taeda, PtrPopulus trichocarpa, Sb Sorghum bicolor, So Saccharumofficinarum, Ta Triticum aestivum, Tar Thlaspi arvense,Tm Triglochin maritima

c

314 Phytochem Rev (2006) 5:309–329

123

CYP71F1 TaCYP71K1 Os

CYP71V LrCYP71D11 Lj

CYP726A1 ElCYP71A1 Pa

CYP71C12 OsCYP71J1 AoCYP71G1 Ao

CYP71N1 MaCYP71E1 SbCYP83E1 Ms

CYP71B1 TarCYP83A1 At

CYP750A1 PtCYP736A1 Pc

CYP84A1 AtCYP84A3 LsCYP92A6 Ps

CYP92C1 OsCYP706A1 AtCYP706C1 Os

CYP706B1 GaCYP80A1 Bs

CYP80B1v2 EcCYP76B7 Psa

CYP76A6 LeCYP75A1 Ph

CYP75A14 PtCYP75B1 At

CYP75B25 SbCYP81D1 At

CYP81A1 ZmCYP82B1 Ec

CYP82E1 NtCYP82A2 Gm

CYP705A1 AtCYP712A1 At

CYP93A1 GmCYP93B1 Ge

CYP73 Pp1CYP73A20 Pt

CYP73A5 AtCYP73A17 Ta

CYP89A2 AtCYP89B1 OsCYP77A9 Os

CYP77B1 AtCYP723A2 OsCYP719A1 Cj

CYP701A3 AtCYP701A6 Os

CYP78A5 AtCYP78B5 Os

CYP78A4 PrCYP98 Pp1CYP98A1 Sb

CYP98A3 AtCYP98A19 Pt

CYP703A2 AtCYP703A3 Os

CYP703B1 PpCYP703B2 Pp

CYP79A10 OsCYP79A9 Os

CYP79A5 TaCYP79A6 Ta

CYP79A7 OsCYP79E1 TmCYP79E2 Tm

CYP79D1 MeCYP79D2 Me

CYP79A1 SbCYP79A So

CYP79A12 HvCYP79A8 HvCYP79A Mt1

CYP79A Mt2CYP79A Mt3

CYP79D3 LjCYP79D4 LjCYP79D8 Ptr

CYP79D5 PtrCYP79D7 PtrCYP79D6v1 PtrCYP79D6v2 Ptr

CYP51 Pp1CYP51G1 At

CYP51G1 Sb

98

85

100

100

100

100100

100

100

100

99100

100

99

7098

85

53

100

100

66100

100

100

10060

98100

100

100

100100

100

8499

9899

99

99

96

99

98

94

5592

78

7187

84

83

83

82

76

71

69

55

54

51

75

72

69

55

58

98

Ao – A.officinalisAt – A. thalianaBs – B. stoloniferaCj – C. japonicaEc – E. californicaGa – G. arboreumGe – G. echinataGm – G. maxHv – H. vulgareLe – L. esculentaLr – L. rigidumLs – L. styracifluaMa – M. acuminataMe – M. esculentaMs – M. sativaMt – M. truncatulaNt – N. tabacumOs – O. sativaPa – P. americanaPc – P. communisPh – P. hybridaPp – P. patensPr – P. radiataPs – P. sativumPsa – P. sativaPt – P. taedaPtr – P. trichocarpaSb – S. bicolorSo – S. officinarumTa – T. aestivumTar – T. arvenseTm – T. maritima

MonocotsEudicotsGymnospermsPhyscomitrella

0.2

Phytochem Rev (2006) 5:309–329 315

123

from a number of angiosperms, showed that at

least within angiosperms the first committed step

is catalysed by an orthologous enzyme that

predates the segregation of angiosperms and

gymnosperms (Fig. 5). The tree is rooted with

ubiquitous obtusifoliol 14a-demethylase CYP51,

from Physcomitrella, Sorghum as well as Arabid-

opsis, to represent a bryophyte, a monocotyledon

and a eudicotyledon, respectively. The clan 71

constitutes a monophyletic clade predicted to

primarily encompass P450s involved in natural

products biosynthesis (secondary metabolism).

This clade proliferated vastly during the

course of plant land colonization over the last

430 million years through multiple gene, genome

segment and genome duplications and rearrange-

ments (Paquette et al. 2003; Nelson et al. 2004;

Galbraith and Bak 2005) in the continuous

processes of neo- and subfunctionalizations. The

CYP79 family branches off as a discrete clade

relatively deep in the tree which reflects the

ancient nature of cyanogenic glycosides. Deep in

the tree, the CYP73 and CYP98 families involved

in biosynthesis of lignin and other phenylpropa-

noids (Humphreys and Chapple 2002) branch off,

as well as the CYP701 family involved in biosyn-

thesis of gibberellins (Helliwell et al. 1998); the

occurrence of these pathways most likely predate

cyanogenic glucoside biosynthesis in terrestrial

plants. As CYP79 sequences from cyanogenic

gymnosperms and pteridophytes are as yet not

available, the analysis is restricted to angiosperms.

This may influence the position of the individual

CYP79s in the 79 clade as well as the position of

the CYP79 clade in relation to the CYP73, CYP98

and CYP703 clades from which sequences are

indeed available from bryophytes, gymnosperms

as well as angiosperms. From monocotyledons,

CYP79 sequences encoding enzymes with aro-

matic as well as aliphatic amino acids as substrates

are available from both Poales and Alismatales

(triglochin). The only known CYP79 sequence

from eudicotyledons is restricted to eurosids I in

Rosids and have aliphatic amino acids as sub-

strates, though aromatic cyanogenic glycosides are

present in eurosides. In contrast to most other

plant P450s, members of the CYP79 family are

characterized by unique amino acid substitutions

in the otherwise generally conserved ‘‘PERF’’ and

heme-binding domain (Werck-Reichhart et al.

2002; Bak et al. 1998b). These substitutions have

facilitated the isolation of cDNAs from a number

of plants using family CYP79 specific primers

(Andersen et al. 2000; Nielsen and Møller 2000)

and has provided a tool to readily identify CYP79s

among other cytochrome P450 sequences in

databases (Nielsen and Møller 2000; Forslund

et al. 2004). In the PERF region of the CYP79

family, the otherwise conserved apolar and aro-

matic phenylalanine residue has been replaced by

the positively charged amino acid residue histidine

giving the CYP79 family consensus PERH.

Similarly, in the heme-binding consensus se-

quence PFGXGRRXCXG, three substitutions of

hydrophobic amino acids with polar serine or thre-

onine residues are observed, affording a CYP79

consensus of (S/T)F(S/T)TGRRGCXG (see full-

sequence alignment at http://www.p450.kvl.dk/

Review2006/Figure5_Alignment.pdf and see

sequence logos at http://www.p450.kvl.dk//

p450.shtml#logos). The identification of CYP79s

from eudicots, other than rosids, and especially

from gymnosperms and pteridophytes will reveal

how far back these unique substitutions can be

tracked. The PERF and heme-binding regions are

part of the active site and the significance of more

polar amino acid residue in the PERF and heme-

binding region is currently not understood. How-

ever, the substitutions may be related to the

unusual catalytic activities of the CYP79 family

and/or to the fact that amino acid substrates as

well at the generated intermediates are unusually

hydrophilic compared to most other cytochrome

P450 substrates. The availability of crystal struc-

tures of a number of mammalian P450s (Williams

et al. 2003; Scott et al. 2003; Williams et al. 2004)

may facilitate homology modelling and docking of

substrates to further the understanding of the

impact of these changes, analogously to the

information gained by the sorghum UGT85B1

model (Fig. 4; Thorsoe et al. 2005).

Glucosinolates constitute another subclass of

N-containing natural products. They are present

in Brassicales within eurosides II (Bak et al.

1998b; Halkier and Gershenzon 2006). They are

derived from a subset of protein amino acids that

partly overlap with those used for synthesis of

cyanogenic glycosides as well as from chain

316 Phytochem Rev (2006) 5:309–329

123

elongated derivatives of these amino acids. As in

cyanogenic glucoside synthesis, the conversion of

these parent amino acids into the corresponding

oximes is catalysed by CYP79’s (Bak et al.

1998b). CYP79s in glucosinolate containing spe-

cies are clearly phylogenetically related to

CYP79s from cyanogenic species, and this most

likely represents recruitment of genes from the

cyanogenic glucoside pathway to the more re-

cently evolved glucosinolate pathway (Bak et al.

1998b). Interestingly, glucosinolates are also

found in Drypetes belonging to Malphigiales,

eurosides I. This has given rise to the hypothesis

of a parallel evolution of glucosinolate biosyn-

thesis in rosids from a cyanogenic ancestor

(Rodman et al. 1998). The isolation of genes

encoding enzymes in glucosinolate biosynthesis

from Drypetes should resolve this issue.

In the glucosinolate pathway in Arabidopsis,

the oximes generated by CYP79s are further

metabolized by CYP83A1 or CYP83B1 (Bak and

Feyereisen 2001; Bak et al. 2001; Hansen et al.

2001; Naur et al. 2003). In sorghum, the oxime is

metabolized by CYP71E1 to the cyanohydrin.

CYP83A1 and CYP83B1 share more than 55%

sequence identity at the amino acid level. Accord-

ing to the generally approved nomenclature

system (Nelson et al. 1996), they should then

have been placed in the same subfamily. How-

ever, it has been decided to retain the two

subfamilies to prevent confusion in the literature.

In addition, based on their amino acid sequence

identity and by their clustering within the CYP71

family in phylogenetic trees, the CYP83 family

clearly belongs to the CYP71 family (Fig. 5).

Mainly due to historical reasons as mentioned for

CYP83B1, the CYP83 family has been kept.

Though CYP83s in Arabidopsis and CYP71E1

in sorghum both have oximes as substrates, the

catalytic mechanism does not appear to be

conserved. In cyanogenic plants, oximes are

dehydrated to a nitrile and subsequently hydrox-

ylated to produce an a-hydroxynitrile. It has not

been resolved whether both of these partial

reactions require NADPH or whether the overall

requirement for NADPH is restricted to the

second partial reaction which is a classical cyto-

chrome P450 C-hydroxylation reaction. In the

glucosinolate pathway, oximes are by a yet

unknown mechanism converted to a thiohydroxy-

mate via a proposed reactive aci-nitro compound

(Ettlinger and Kjær 1968; Bak et al. 2001; Bak

and Feyereisen 2001). If the mechanistics of the

CYP71E1 catalysed dehydration reaction in cya-

nogenic glucoside synthesis was resolved, this

may serve to shed new light on the identity of the

intermediates in glucosinolate synthesis. How-

ever, it remains an open question whether the

CYP83s in glucosinolate biosynthesis have

evolved from a CYP71E1 ortholog, or whether

glucosinolate-producing plants have recruited

other CYP71 homologs to catalyse this step.

Indole-3-acetaldoxime, derived from CYP79B2

and/or CYP79B3 metabolism of tryptophan, is

apart from being an intermediate in indole

glucosinolate biosynthesis an intermediate in

biosynthesis of the main phytoalexin camalexin

in Arabidopsis. CYP71B15, which is a relatively

close homolog to both CYP71E1 and CYP83A1/

B1 (Fig. 5) catalyses the conversion of indole-

3-acetaldoxime-derived dihydrocamalexic acid to

camalexin (A3; Schuhegger et al. 2006), implying

that the CYP71 family is a major player in the

recruitment of enzyme catalysts for evolution of

novel natural products biosynthetic pathways in

plants.

The CYP71 family is by far the most

expanded cytochrome P450 family in plants,

and this combined with the apparent lack of

unique sequence substitutions as seen for the

CYP79 family has despite the increasing num-

ber of plant genome and EST sequencing

projects prevented the identification of clear

CYP71E1 orthologs in other species apart from

the closely related sugar cane and rice. Simi-

larly, no clear UGT85B1 orthologs have as yet

been identified.

Cyanoalk(en)yl glycosides: a subclass of natural

products related to cyanogenic glycosides

Lotus japonicus contains high levels of rhodiocy-

anosides A (C1) and D as well as of the

cyanogenic glycosides linamarin (B1) and

lotaustralin (B2; Forslund et al. 2004). Rhodiocy-

anosides A and D belong to a group of

cyanoalkenyl glycosides structurally related to

Phytochem Rev (2006) 5:309–329 317

123

cyanogenic glycosides. In contrast to the

cyanogenic glycosides, cyanoalk(en)yl glycosides

are not cyanogenic, i.e. they do not release HCN

by treatment with b-glucosidases. This reflects

that these glycosides are not derived from

cyanohydrins and that they are hydrolysed into

stable nitriles by b-glucosidases. Rhodiocyano-

sides A and D were first isolated from Rhodiola

species. Rhodiola extracts are used in Chinese

traditional medicine, and the rhodiocyanosides

have been proposed to function as inhibitors of

histamine release in allergic responses (Yoshika-

wa et al. 1995, 1996a, b). As cyanogenic glycosides

are ancient, several animals and fungi have

evolved to be able to use the otherwise toxic

HCN as a source of nitrogen. Accordingly, for the

plant the presence of cyanogenic glycosides may

in fact be disadvantageous. The transition to

cyanoalk(en)yl glycosides may counteract the

adaptation of herbivores and microbes to HCN.

Rhodiocyanosides are derived from the amino

acid isoleucine (Forslund et al. 2004). This implies

that the biosynthesis of rhodiocyanosides is

catalysed by the very same multienzyme complex

involved in lotaustralin synthesis. However, when

rhodiocyanosides are synthesized, the putative

nitrile intermediate undergoes two hydroxyla-

tions and a dehydration reaction (Forslund et al.

2004). The precise mechanism is not understood,

but we suggest that the initial hydroxylation

produces a product that in contrast to the cyano-

hydrin is not easily released from the active site.

This affords the possibility of a dehydration

reaction to proceed while the intermediate

remains bound in the catalytic site. The rhodio-

cyanosides may then be formed by a subsequent

hydroxylation reaction. Metabolite profiling of

Lotus, Rhodiola and also Ribes species by LC-MS

demonstrated that the rhodiocyanosides co-occur

with lotaustralin (Bjarnholt et al. our unpublished

results). Hydroxylated cyanoalkenyl glycosides

such as sarmentosin (C2) have been identified in

Sedum sarmentosum (Fang et al. 1982; Lechten-

berg and Nahrstedt 1999). A similar complex

profile is known from barley, which in addition

to the leucine-derived cyanogenic glucoside

epi-heterodendrin contains the cyanoalkenyl

glycosides osmaronin, dihydroosmaronin (C4),

epidermin and sutherlandin (Nielsen et al.

2002). Our current working hypothesis is that in

species containing cyanoalk(en)yl glycosides,

CYP71E1 orthologs have been duplicated and

subsequently evolved to catalyse two sequential

hydroxylations accompanied by a dehydration

reaction. The availability of model plants like L.

japonicus containing rhodiocyanoside A and D

will allow the elucidation of how this new

pathway has evolved from a cyanogenic glucoside

predisposition on the molecular level. Again,

elucidation of these reaction events may serve

to cast new light on the intermediates involved in

the conversion of oximes to thiohydroxymates in

glucosinolate synthesis.

The proposed evolution of glucosinolates,

camalexin and cyanoalk(en)yl glycosides from a

cyanogenic glucoside predisposition illustrates

how the ancient pathway for cyanogenic gluco-

side synthesis has given rise to new classes and

subclasses of natural products (Fig. 6).

Localization of CYP79s and cyanogenic

glycosides

In L. japonicus, the cyanogenic glycosides accu-

mulate in aerial tissues (Forslund et al. 2004).

CYP79D3 transcripts are primarily found in

young and developing tissues where the concen-

tration of cyanogenic glycosides per mg fresh

weight is also the highest. In a similar manner,

sorghum CYP79A1 is predominantly expressed in

the young developing seedling (Busk and Møller

2002). The level of cyanogenic glycosides per mg

fresh weight decreases in sorghum and L. japo-

nicus as a function of developmental age. This

indicates that synthesis primarily occurs in young

and developing tissues and that the levels found

in older plant parts decreases because de novo

synthesis proceeds at a lower rate than catabolic

turnover. The presence of high amounts of

cyanogenic glycosides in young and developing

tissues supports the proposed function as defence

compounds. In cassava, sorghum and eucalyptus,

the level of cyanogenic glycosides increase under

high nitrogen levels (Gleadow and Woodrow

2000; Busk and Møller 2002; Gleadow and

Woodrow 2002; Jørgensen et al. 2005a) indicating

that cyanogenic glycosides may also serve a

function as storage compounds for reduced

318 Phytochem Rev (2006) 5:309–329

123

nitrogen in at least some plant species. In addition

to CYP79D3, L. japonicus contains a paralogous

P450 CYP79D4 which is ~95% identical to

CYP79D3 at the amino acid level, but shares

limited identity within the promoter region.

Whereas CYP79D3 is preferentially expressed in

Amino acids

CYP79A1orthologs CYP71B15

CYP83A1/B1

Camalexin (A3)

Oximes

Cyanogenic glycosides(A1 + B1-B4)

Non-cyanogeniccyanoalk(en)yl glycosides

(C1-C4)

Glucosinolates(A2)

CYP79E1orthologs

Cyanohydrins

UGT85B1orthologs

CH3CN

CH3

O

OHOH

OH

OH

O

B1. Linamarin(L-valine)

B2. 2R = Lotaustralin(L-isoleucine)

B3. 2S = Dhurrin(L-tyrosine)

B4. 1S, 4R = Epivolkenin(2-(2'-cyclopentenyl) glycine)

H

OH

OCN

OHOH

OH

OH

O

HOH

OCN

OHOH

OH

OH

O

CH3

CNCH3

O

OHOH

OH

OH

O

a

SO3OGlc

S N

R

-Sugar R1

CN

R2

O

A1. Cyanogenic glycoside A2. Glucosinolate

N

S

NH

A3. Camalexin

b

c

CN

CH3

CH3

O

OHOH

OH

OH

O

C3. Epidermin(L-leucine)

C4. 3S = Dihydroosmaronin(L-leucine)

C1. Rhodiocyanoside A(L-isoleucine)

C2. Sarmentosin(L-isoleucine)

CNO

CH3

OHOH

OH

OH

O CN

OH

O

OHOHOH

OH

O

H

NC

CH3O

OHOHOH

OH

O

Fig. 6 Biosynthetic relationship between cyanogenicglycosides, cyanoalkenyl glucosides, glucosinolates andthe indole alkaloid camalexin. General structures ofcyanogenic glycosides, glucosinolates and camalexin

(a), cyanogenic glycosides derived from aliphatic, aromaticand non-protein amino acids (b) and non-cyanogeniccyanoalk(en)yl glycosides (c) are shown

Phytochem Rev (2006) 5:309–329 319

123

leaves, CYP79D4 is preferentially expressed in

root tissue although at an apparent relatively

lower level. The two enzymes possess the same

kinetic parameters when heterologously ex-

pressed in yeast cells (Forslund et al. 2004). The

role of expression of a fully functional paralog

in an otherwise apparent acyanogenic tissue is

currently not understood. However, as a legume

L. japonicus interacts with Rhizobium bacteria to

form nitrogen-fixing nodules under low nitrogen

levels. Our analyses of CYP79D3 and CYP79D4

expression as well as the distribution of cyano-

genic and cyanoalkenyl glycosides in L. japonicus

has not included Rhizobium infected roots but

only tissue from hydroponically grown plants

supplemented with high nitrogen levels. The

possibility that cyanogenic or cyanoalkenyl gly-

cosides may serve as signalling compounds in

symbiotic nitrogen fixation or as transporters of

reduced nitrogen regulated by the nitrogen supply

and Rhizobium interaction remains an intriguing

question. Generation of transgenic L. japonicus

lines with CYP79D3 and CYP79D4 promoters

fused to reporter genes is in progress and will aid

to elucidate the functions of cyanogenic and

cyanoalkenyl glycosides in interactions with

micro-organisms and herbivores (Saito et al.,

our unpublished results).

In cassava, cyanogenic glycosides are prefer-

entially synthesized in the aerial parts and subse-

quently transported to the roots for storage

(Jørgensen et al. 2005a). The mechanism of

transport and the possible existence of a specific

transport form remain elusive. Studies in Hevea

cotyledons suggest that transport of the cyano-

genic glucoside linamarin requires an initial

conversion into the diglucoside linustatin (Selmar

et al. 1988). In contrast to the monoglucoside, the

diglucoside is not hydrolysed by the apoplastic

b-glucosidases and thus fulfil the prerequisite of a

non-degradable transport form. In L. japonicus

and sorghum, the localization of CYP79s at the

tissue level has been determined by RT-PCR and

northern-blotting analyses. In cassava, the local-

ization of CYP79D1 and CYP79D2 transcripts at

the cellular level has been visualized by FITC

labelling by in-tube in situ PCR in 3-month-old

plants (Jørgensen et al. 2005a). The first unfolded

leaf and its petiole have the highest cyanogenic

glucoside biosynthetic activity. In these two

tissues, the transcripts were localized within the

epidermis and the adjacent two cortex cell layers,

and in the endodermis and pericycle cell layers

(Fig. 7). CYP79D1 and CYP79D2 are also highly

expressed in tissues surrounding laticifers and

phloem cells, in regions between vascular bun-

dles, in parenchymatic cells in the vascular tissue

and especially in between the protoxylem and

metaxylem cells. The expression of CYP79D1

Fig. 7 Cellular localization of CYP79D1 in the petiolefrom the first fully unfolded cassava leaf as monitored byin-tube in situ RT-PCR analysis using light microscopy of100 lm transverse sections. (A) Petiole control to visualizethe different cell types. (B) Petiole labelling backgroundcontrol with FITC as observed without primers in the RTreaction. (C) Expression of CYP79D2 in petiole asmonitored by FITC labelling. (D) Expression of CYP79D1in petiole as monitored by FITC labelling. (E) Expressionof CYP79D1 in petiole as monitored by alkaline phospha-tase labelling. e Epidermis, p phloem, l laticifers, enendodermis. Bars = 200 lm. Adapted from Jorgensenet al. (2005)

320 Phytochem Rev (2006) 5:309–329

123

and CYP79D2 in epidermal and cortex cells and

around laticifers represents the function of cya-

nogenic glycosides as biochemical protectants as

they are expressed in conjunction with cassava

linamarin b-glucosidases apoplastically localized

in laticifers. Expression around the phloem and

vascular bundles could represent sites of biosyn-

thesis targeted for transport to the tubers.

Both L. japonicus and cassava possess two

CYP79Ds. In L. japonicus, the two paralogs

represent a clear case of subfunctionalization

where the promoters have diverged while the

coding regions are highly conserved as are their

catalytic parameters. No ESTs for CYP79D4 has

been identified, but RT-PCR experiments reveal

expression in roots (Forslund et al. 2004). In

contrast, the two cassava paralogs are only ~85%

identical at the amino acid level, have the same

catalytic parameters and are co-expressed

(Andersen et al. 2000; Jørgensen et al. 2005a).

The presence of two cassava paralogs is most

likely a consequence of the allotetraploid genetic

background of cassava with one allele originating

from each of the parental lines.

Metabolic engineering of cyanogenic glycosides

The pathway for conversion of a parent amino

acid into the corresponding cyanogenic glucoside

involves six intermediates of which several are

labile. From a genetic point of view, the cyano-

genic glucoside pathway is quite simple as two of

the enzymes responsible for these conversions are

multifunctional. Accordingly, the entire pathway

is encoded by just three structural genes. These

three genes are available from sorghum (Tatter-

sall et al. 2001; Kristensen et al. 2005). This gives

cyanogenic glycosides a pioneering position in

metabolic engineering of natural products in

plants because most other pathways for complex

natural product, e.g. alkaloids have not yet been

fully elucidated and because a substantially larger

number of genes are involved (Kutchan 2005).

The ability to engineer up to 4% dry weight of the

tyrosine-derived cyanogenic glucoside dhurrin

into transgenic A. thaliana plants by ectopic

expression of sorghum CYP79A1, CYP71E1 and

UGT85B1 (Tattersall et al. 2001; Kristensen et al.

2005) has been recently reviewed (Jørgensen

et al. 2005b; Kutchan 2005; Morant et al. 2003;

Memelink 2005). The transfer of the entire high

flux dhurrin pathway from sorghum to Arabidop-

sis by genetic engineering proceeded essentially

without inadvertent effects on the metabolome

and transcriptome (Kristensen et al. 2005; Fig. 8).

This demonstrates that effective encapsulation of

toxic intermediates by metabolon formation is

also achievable after heterologous expression in a

plant species that would not in nature produce the

same class of natural product (Tattersall et al.

2001; Kristensen et al. 2005). Insertion of an

incomplete pathway (sorghum CYP79A1 and

CYP71E1) resulted in stunted plants, transcrip-

tome alterations, accumulation of numerous new

glycosides derived from detoxification of inter-

mediates in the dhurrin pathway, and in loss of

the Brassicaceae specific UV protectants sinapoyl

glucose and sinapoyl malate (Bak et al. 2000;

Tattersall et al. 2001; Kristensen et al. 2005).

When separately introduced into Arabidopsis,

sorghum CYP79A1 established a highly efficient

interaction with the down-stream glucosinolate-

producing enzymes and resulted in accumulation

of large amounts of p-hydroxybenzylglucosinolate

(up to 3% dry weight; Bak et al. 1999). This gluc-

osinolate is not normally present in Arabidopsis

and thereby changes the overall glucosinolate

profile of Arabidopsis (Petersen et al. 2001). The

feasibility to redirect tyrosine utilization in

Arabidopsis into glucosinolate or cyanogenic

glucoside production without loss of plant fitness

(Bak et al. 1999; Tattersall et al. 2001; Mikkelsen

et al. 2002; Mikkelsen and Halkier 2003; Kristen-

sen et al. 2005) demonstrates the existence of

inherent routes for transport and storage of new

natural product classes introduced into plants by

genetic engineering and an inherent ability to

redirect and optimize the flux of intermediates to

counteract imbalances in primary and secondary

metabolism in plants. The ability to accommodate

altered levels of intermediates, however, depends

on the type of metabolic cross-talk introduced. In

Arabidopsis, tryptophan-derived oximes are key

intermediates in formation of the phytohormone

indole acetic acid (Woodward and Bartel 2005) as

well as in biosynthesis of glucosinolates (Bak

et al. 2001) and the indole alkaloid camalexin

Phytochem Rev (2006) 5:309–329 321

123

(Glawischnig et al. 2004). CYP83A1 and

CYP83B1 convert oximes into the corresponding

thiohydroximates (Bak and Feyereisen 2001; Bak

et al. 2001; Hansen et al. 2001; Naur et al. 2003)

which are further metabolized by SUR1 (Mikkel-

sen et al. 2004) and UGT74B1 to desulphogluc-

osinolates (Douglas et al. 2004) which are finally

sulphurylated (Piotrowski et al. 2004; Klein et al.

2006). Knock-out of these enzyme activities

disturbs auxin homeostasis and results in pheno-

types reminiscent of auxin overproduction. Thus,

channelling of tryptophan-derived intermediates

into natural product synthesis may serve an

important function to balance primary metabo-

dhurrin

sinapoylglucose sinapoylmalate

B Intens. x106

5 10 15 20 25 30 min

7 8 9 1212

12 14

13

12

12

12

14

13

1212

1214 13

5

1212 12

14 136

p-hydroxybenzylglucosinolate

Wt

1x

2x

3x

A

2

4

6

6

4

2

6

4

2

6

4

2

EIC 316

EIC 309

EIC 363

EIC 337

EIC 368

EIC 409

EIC 485

EIC 601

EIC 617

EIC 763

EIC 334

EIC 323 TIC

C

322 Phytochem Rev (2006) 5:309–329

123

lism. A disturbance of oxime metabolism affects

phenylpropanoid metabolism and the monomer

composition of lignin, but the nature of this

interaction is not yet understood (Hemm et al.

2003). The ability to redirect aromatic amino

acids into new metabolic pathways also reflects

the plasticity of the shikimate pathway. In plants,

this pathway accommodates up to 20% of the flux

of fixed carbon and is primarily regulated by feed-

back inhibition to facilitate metabolic alterations

of aromatic compound in plants (Herrmann

1995).

Ectopic expression of cassava CYP79D2 in

transgenic L. japonicus plants resulted in a ~20-

fold up-regulation of the valine-derived cyano-

genic glucoside linamarin (Forslund et al. 2004;

Fig. 9A). A similar increase of isoleucine-derived

lotaustralin and the cyanoalkenyl glycosides rhod-

iocyanoside A and D was not observed. Accord-

ingly, the relative differences in catalytic

efficiency (Kcat/Km) of cassava CYP79D2 as

compared to L. japonicus CYP79D3 towards the

substrates valine and isoleucine is a major deter-

minator of the ratio of linamarin and lotuastralin

obtained. The cassava CYP79D2 enzyme has

about the same catalytic efficiency towards valine

and isoleucine, and accordingly generates an

approximately 1:1 ratio of linamarin to lotaustr-

alin in this species (Andersen et al. 2000). In

contrast, L. japonicus CYP79D3 has a ~6-fold

higher catalytic efficiency towards isoleucine as

compared to valine (Forslund et al. 2004). This

results in a correspondingly lower relative level of

linamarin to lotaustralin. Accordingly, the trans-

genic L. japonicus lines accumulated much more

linamarin and approached the linamarin to

lotaustralin ratio in cassava. A surprising result

of the ectopic expression of cassava CYP79D2 in

L. japonicus was the accumulation of linamarin

and lotaustralin in roots, demonstrating that roots

indeed do have the capacity to synthesize cyano-

genic glycosides and that presence of the CYP79

step exerts quantitative and qualitative control of

the flux through the pathway. The cyanoalkenyl

glycosides rhodiocyanoside A and D did not

co-accumulate in roots, which is a strong indica-

tion of the presence of two separate biosynthetic

pathways for cyanogenic and alkenyl glucoside

synthesis in roots and aerial tissues in L. japoni-

cus. The possibility that the root content of

linamarin and lotaustralin was derived from

translocation from the aerial parts to the roots

cannot be ruled out. However, the presence in

L. japonicus of two CYP79D paralogs with

different expression patterns supports the pres-

ence of two independent pathways. Unfortu-

nately, the CYP71E1 orthologs in L. japonicus

have not yet been identified and this hampers the

understanding of how L. japonicus organizes the

synthesis of cyanogenic and cyanoalkenyl glyco-

sides in aerial tissues; either as parallel metabo-

lons or as a single promiscuous metabolon

that synthesizes both sets of glycosides simulta-

neously.

Ectopic expression of cassava CYP79D2

has also been used to alter the glucosinolate

profile in Arabidopsis (Mikkelsen and Halkier

2003). In analogy to metabolic cross-talk

observed by expression of sorghum CYP79A1,

the CYP79D2 expressing transgenic lines accu-

mulate up to 0.3% dry weight of the valine- and

isoleucine-derived isopropyl- and 1-methylpro-

pyl-glucosinolates, two glucosinolates that

Fig. 8 Metabolic engineering of dhurrin synthesis intransgenic Arabidopsis plants with marginal inadvertenteffects on the metabolome and transcriptome. Impact onvisual phenotype (A), metabolome (B) and transcriptomeas a result of expression of sorghum CYP79A1 either alone(1·) or combined with sorghum CYP71E1 (2·) or withsorghum CYP71E1 and UGT85B1 (3·) as compared towild-type (WT). (B) LC-MS analysis of the metabolitecontent of the plants shown in (A) as monitored by thetotal ion chromatograms (TIC, black lines) and extractedion chromatograms (EIC, coloured lines). Metabolitesreferred to in the text are high lighted: Sinapoyl glucose(14), sinalpoylmalate (13), dhurrin (6) and p-hydroxy-benzylglucosinolate (7 and 8), additional labelling as inKristensen et al. (2005). c Scatter plot of trancriptomeanalyses of wild-type (WT) versus dhurrin-producing A.thaliana plants (3· plants). The global arrays contains~29,000 probes while the focussed array contains probesfor 453 selected genes in secondary metabolism, aromaticamino acid biosynthesis, all 246 full-length A. thalianacytochromes P450, except CYP84A4, and all 112 family 1UGT except for members of the UGT80 and UCT81families. Phenylpropanoid marker genes are shown in blue,glucosinolate marker genes in pink and sorghum transg-enes in yellow (not present on global array). Genes scoredas up-regulated are shown in red and genes down-regulated in green. Signals below two time’s averagebackground are boxed. Blue lines indicate twofold orfourfold up- or down-regulated expression levels. Adaptedfrom Kristensen et al. (2005)

b

Phytochem Rev (2006) 5:309–329 323

123

normally do not accumulate in the Columbia

ecotype (Mikkelsen et al. 2002). The relative

lower levels of aliphatic glucosinolate levels

compared to tyrosine-derived glucosinolates

obtained, most likely relicts the differences in

plasticity of the isoleucine and valine biosynthetic

pathways as compared to the shikimate pathway.

Cassava is one of the most important root

crops in the world. However, the high amounts of

the cyanogenic glycosides linamarin and lotaus-

tralin which accumulate in the starchy tubers

constitute a major health risk to the people that

daily rely on this stable crop due to the ability

of these compounds to release toxic HCN.

Accordingly, careful processing is necessary to

remove the cyanogenic glycosides from the tubers

before consumption. A major drawback of this

processing is the concurrent loss of most of the

proteins, vitamins and minerals, leaving behind a

product rich in starch, but with a very low

nutritional value. Traditional breeding has not

provided acyanogenic cassava lines. The inability

to select such lines by traditional breeding may be

a consequence of the presence of two unlinked

CYP79D paralogs and the low seed set in cassava.

Accordingly, down-regulation of the pathway by

RNAi technology is an attractive alternative.

Lines with acyanogenic (<1% of wild-type) leaves

and near-acyanogenic tubers (<8% of wild-type)

have been obtained using a single RNA interfer-

ence construct that simultaneously blocks

CYP79D1 and CYP79D2 (Jørgensen et al.

2005a; Fig. 9B). When grown in vitro, such

down-regulated lines displayed a distinct

0

500

1000

1500

2000

2500

3000

3500

4000

wt 10 4 19 8 12 #6 16 6 1 20 9 3 13 7 17 2 11 5 18 15 14 #5

% o

fthe

linam

arin

con

tent

inw

ildty

pe A

0

10

20

30

40

50

60

70

80

90

100

wt 83 98 91 84 110 108 78 94 103 90 102100 79 97 109 86 104101 93 85 99 89

% o

fthe

linam

arin

con

tent

inw

ildty

pe B

Fig. 9 Metabolic engineering of linamarin content in L.japonicus and cassava. (A) In L. japonicus, the synthesis oflinamarin has been enhanced by over-expression ofCYP79D2 from cassava under the control of the 35Spromoter as measured in apical shoots. A similar increaseis not observed for lotaustralin, because CYP79D2preferentially utilizes valine as substrate, the parent aminoacid for linamarin formation. (B) In cassava, the content of

linamarin is reduced by expression of an RNAi constructdirected against CYP79D1 and CYP79D2 under thecontrol of the enhanced 35S promoter as measured inthe first unfolded leaf. A similar reduction is observed forlotaustralin. The content of linamarin is measured by LC-MS and the change in linamarin content is calculated as %of the wild-type content

324 Phytochem Rev (2006) 5:309–329

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morphological phenotype with long, slender

stems and long internodes as compared to in

vitro grown wild-type plants. The phenotype was

partially restored by increasing the nitrogen

concentration in the media, and completely

restored by transplanting the lines to soil. The

apparent interactions between the levels of

endogenous nitrogen and cyanogenic glycosides

are not understood. The observed conditional

phenotype indicates that cyanogenic glycosides

may possess other roles than simply being phyto-

anticipins involved in plant defence. These may

include roles as, e.g. signalling compounds or an

involvement in balancing nitrogen levels. In a

parallel approach using anti-sense technology and

the leaf-specific CAB1 promoter, transgenic

plants with reduced cyanogenic glucoside content

in leaves was also observed (Siritunga and Sayre

2003, 2004).

In conclusion, CYP79 activity regulates the flux

of intermediates through the cyanogenic gluco-

side pathway and the substrate specificity of the

CYP79s determines the cyanogenic glucoside

profile. Furthermore, CYP79s from cyanogenic

plants may be used to up- or down-regulate the

level of cyanogenic glycosides as well as to

introduce new glucosinolate profiles. Likewise,

insertion of CYP79s from the glucosinolate path-

way into cyanogenic plants may enable synthesis

of cyanogenic glycosides derived from new parent

amino acids. However, the level achieved of such

new cyanogenic glycosides would be expected to

be low because of the predictable strong interac-

tion between the endogenous cyanogenic CYP79

with the down-stream enzymes in the cyanogenic

glucoside pathway.

Concluding remarks and perspectives

The availability of P450 sequences involved in

biosynthesis of cyanogenic glycosides from an

increasing number of cyanogenic plants has

allowed a first glimpse of the molecular phylog-

eny of this ancient pathway. The addition of more

genomic, EST and cDNA sequences will help to

further our understanding at the molecular level

of how this pathway has evolved and how novel

biosynthetic pathways like the cyanoalk(en)yl,

camalexin and glucosinolate pathways have

evolved from a cyanogenic predisposition. The

poplar genome has revealed the presence of five

closely related CYP79D paralogs (Fig. 5) and the

availability of an EST corresponding to either

CYP79D5 or CYP79D7 documents that at least

some of these genes are transcribed. As cassava,

poplar belongs to Malpighiales (eurosids I) and it

is therefore tempting to suggest that poplar

contain linamarin and lotaustralin. The availabil-

ity of the genome of a putatively cyanogenic tree

model provides the necessary tools to study the

impact of cyanogenesis in a tree. Homology

modelling of cytochromes P450 will facilitate an

understanding of the importance of the unique

substitutions discovered in the PERF and heme-

binding regions of the CYP79s and contribute to

the elucidation of the unusual chemistries of the

multifunctional P450s in this pathway. Compara-

tive modelling of CYP71E1 homologs in cya-

noalkenyl and cyanogenic glucoside pathways will

provide novel information as to how new natural

products evolved at the molecular level. The

availability of genomic sequences from rice,

L. japonicus and poplar constitute important

resources for studying regulation of cyanogenic

glycosides under abiotic and biotic stress, modu-

lated by transcription factors and non-coding

RNAs. Technology platforms as untargeted trans-

criptomics and metabolomics will serve to unravel

the regulatory and metabolic networks and hubs

in interactions between primary and secondary

metabolism. This is important as compelling data

suggest that cyanogenic and cyanoalkenyl glyco-

sides may have evolved to serve other functions

than merely as phytoanticipins. The use of met-

abolic engineering of natural product profiles

using P450s provides a powerful tool to elucidate

the delicate plant–insect/microbe interactions as

well as to exploit the use of transgenic plants as

green factories for production of natural products

for the biotechnic and pharmaceutical industries.

Acknowledgements Steen Malmmose, Susanne Jensenand Charlotte Sørensen are thanked for excellent technicalassistance in the greenhouses and laboratories. All formermembers of the Cyanogenic Glycosides group are thankedfor their contributions.

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