Phenylalanine derived cyanogenic diglucosides from Eucalyptus camphora and their abundances in relation to ontogeny and tissue type

Article abstract of DOI:10.1016/j.phytochem.2011.08.022

The cyanogenic glucoside profile of Eucalyptus camphora was investigated in the course of plant ontogeny. In addition to amygdalin, three phenylalanine-derived cyanogenic diglucosides characterized by unique linkage positions between the two glucose moiet

Full text of DOI:10.1016/j.phytochem.2011.08.022

Phytochemistry 72 (2011) 2325–2334
Contents lists available at SciVerse ScienceDirect
Phytochemistry
journal homepage: www.elsevier.com/locate/phytochem
Phenylalanine derived cyanogenic diglucosides from Eucalyptus camphora
and their abundances in relation to ontogeny and tissue type
Elizabeth H. Neilson , Jason Q.D. Goodger , Mohammed Saddik Motawia b,c, Nanna Bjarnholt b,
a,
⇑
a
b
c
b,c
a
Tina Frisch , Carl Erik Olsen , Birger Lindberg Møller , Ian E. Woodrow
a
School of Botany, The University of Melbourne, Victoria 3010, Australia
Plant Biochemistry Laboratory, Department of Plant Biology and Biotechnology, University of Copenhagen, Copenhagen, Denmark
Villum Research Centre Pro-Active Plants, Copenhagen, Denmark
b
c
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 16 June 2011
Received in revised form 16 August 2011
Available online 24 September 2011
The cyanogenic glucoside profile of Eucalyptus camphora was investigated in the course of plant ontogeny.
In addition to amygdalin, three phenylalanine-derived cyanogenic diglucosides characterized by unique
linkage positions between the two glucose moieties were identified in E. camphora tissues. This is the first
time that multiple cyanogenic diglucosides have been shown to co-occur in any plant species. Two of
these cyanogenic glucosides have not previously been reported and are named eucalyptosin B and euca-
lyptosin C. Quantitative and qualitative differences in total cyanogenic glucoside content were observed
across different stages of whole plant and tissue ontogeny, as well as within different tissue types. Seed-
lings of E. camphora produce only the cyanogenic monoglucoside prunasin, and genetically based varia-
tion was observed in the age at which seedlings initiate prunasin biosynthesis. Once initiated, total
cyanogenic glucoside concentration increased throughout plant ontogeny with cyanogenic diglucoside
production initiated in saplings and reaching a maximum in flower buds of adult trees. The role of multi-
ple cyanogenic glucosides in E. camphora is unknown, but may include enhanced plant defense and/or a
primary role in nitrogen storage and transport.
Keywords:
Eucalyptus camphora
Myrtaceae
Cyanogenic glycoside
Prunasin
Amygdalin
Eucalyptosin
Chemical defense
Cyanide
Cyanogenesis
Eucalypt
Ó 2011 Elsevier Ltd. All rights reserved.
0
1
. Introduction
acid
L
-2-(2 -cyclopentenyl) glycine (Zagrobelny et al., 2008). Struc-
tural diversity may be limited in part by the channeled nature of
Plants produce a vast array of bioactive natural products. These
the biosynthetic pathway (Møller and Conn, 1980). In Sorghum
chemicals help defend plant tissues against herbivory and patho-
gens. Cyanogenic glucosides ( -hydroxynitrile glucosides) are an
bicolor, for example, conversion of L-tyrosine to the cyanogenic glu-
a
coside dhurrin is catalyzed by two cytochrome P450s and a UDPG-
glycosyltransferase, which are envisioned to form a multi-enzyme
complex (a metabolon) that prevents free diffusion of auto-toxic
intermediates (Jørgensen et al., 2005b; Møller and Conn, 1980;
Nielsen et al., 2008; Winkel, 2004). As a consequence, much of
the structural diversity of cyanogenic glucosides is achieved simply
by modification of the sugar moiety by additional glycosylation(s)
or galloylation (Fleming, 1999; Ling et al., 2002). Another factor
that is relevant to understanding the diversification in structure
of cyanogenic glycosides is their possible additional roles in pri-
mary metabolism (Møller, 2010), particularly in nitrogen storage
and transport (Bjarnholt and Møller, 2008; Jenrich et al., 2007;
Jones et al., 2000). For example, when Prunus serotina seedlings
germinate, the cyanogenic diglucoside amygdalin is transported
from the seeds and metabolized to supply nitrogen to the develop-
ing seedling without release of HCN to the surroundings (Swain
and Poulton, 1994). These and other data have prompted the sug-
gestion that cyanogenic diglucosides serve primarily as transport
forms (Sánchez-Pérez et al., 2008; Selmar et al., 1988; Swain and
important and widespread class of such defense chemicals, which
have been recorded in over 2500 plant species from diverse taxa.
When tissue of these plants is disrupted, HCN gas is enzymatically
released from cyanogenic glucosides, which thus act as phytoantic-
ipins affording an immediate chemical defense response to herbi-
vores and pathogens (Møller, 2010). Despite the importance of
cyanogenic glucosides to a wide array of plants, they constitute a
very small class of bioactive natural products with only 60 or so
different structures identified (Bjarnholt and Møller, 2008). This
contrasts with terpenoids, for example, where in excess of 40,000
different structures have been elucidated (Bohlmann and Keeling,
2
008).
Cyanogenic glucosides are biosynthesized from six different
building blocks: the five protein amino acids -valine, -isoleucine,
-tyrosine and the non-protein amino
L
L
L-leucine, L-phenylalanine, L
⇑
Corresponding author. Tel.: +61 3 83447165; fax: +61 3 93475460.
E-mail address: neilson@unimelb.edu.au (E.H. Neilson).
0
031-9422/$ - see front matter Ó 2011 Elsevier Ltd. All rights reserved.
doi:10.1016/j.phytochem.2011.08.022

2
326
E.H. Neilson et al. / Phytochemistry 72 (2011) 2325–2334
Poulton, 1994). In addition, there is evidence that when available
nitrogen is in excess of that required for growth it can be stored
for future use as cyanogenic glucosides. For example, following a
rise in nitrate supply, the concentration of the cyanogenic gluco-
sides linamarin and lotaustralin was shown to increase in the shoot
apex of cuttings from six-week-old Manihot esculenta plants
(Fischer, 1885; Koo et al., 2005) and subjected to the same LC-
MS analysis as the plant extracts. Neoamygdalin did not co-elute
with any of the E. camphora constituents able to release HCN. This
would suggest that the unknown compounds are amygdalin iso-
mers in which the second glucoside moiety is linked differently
compared to the b(1 ? 6) linkage found in amygdalin and neo-
amygdalin. Two different approaches were undertaken to deter-
mine the nature of the glucosidic linkages present in the three
unknown structures.
The first approach involved spectroscopic comparison with
authentic standards of likely amygdalin isomers. An amygdalin iso-
mer (R)-mandelonitrile b-sophoroside in which the two glucose
residues are coupled by a b(1 ? 2) linkage has previously been iso-
lated from Eremophila maculata (Syah and Ghisalberti, 1996).
Therefore we isolated this compound from E. maculata and con-
firmed its structure by NMR. Comparison by GC–MS and LC-MS
analysis showed that (R)-mandelonitrile b-sophoroside co-eluted
with one of the unknown constituents (peak II, Fig. 1a) and gave
an identical fragmentation pattern. This confirmed the structural
assignment of the unknown as 5, which we have named eucalypt-
osin A (Fig. 2). Given there is no known plant source of the amyg-
dalin isomer (R)-mandelonitrile b-cellobioside in which the
glucose moieties are b(1 ? 4) linked, we chemically synthesized
this compound and confirmed its structure by NMR. This synthetic
standard was analyzed using LC-MS and GC–MS and found to co-
elute and possess an identical fragmentation pattern with one of
the other unknown cyanogenic constituents in the E. camphora ex-
tracts (peak III, Fig. 1a). We have assigned structure 6 to the second
unknown and named it eucalyptosin B (Fig. 2).
(Jørgensen et al., 2005a). Similarly, experiments with five-
week-old S. bicolor plants showed that the biosynthetic machinery
for synthesis of the cyanogenic glucoside dhurrin was transcrip-
tionally induced by application of nitrate fertilizer (Busk and
Møller, 2002).
Our understanding of the diverse roles of cyanogenic glucosides
may be advanced through studies of plants that contain multiple
cyanogenic glycosides, including mono- and diglucosides, espe-
cially where the relative abundance of these change through devel-
opment and in response to different environmental conditions.
Eucalyptus camphora is a plant species that fulfills these two crite-
ria. Firstly, there is evidence that its foliage contains at least five
different cyanogenic glucosides, including the monoglucoside
prunasin, its epimer sambunigrin, and the diglucoside of prunasin,
amygdalin (Neilson et al., 2006). Secondly, it displays strong onto-
genetic influence over cyanogenic glucoside biosynthesis with
seedlings significantly lower in total cyanogenic glucoside concen-
tration than their adult counterparts, with some seedlings incapa-
ble of releasing HCN after six months of growth (Neilson et al.,
2
006). In this paper, we report a detailed characterization of the
cyanogenic glucosides present in the foliar and reproductive tis-
sues of E. camphora, revealing three additional cyanogenic digluco-
sides designated eucalyptosin A, eucalyptosin B and eucalyptosin C
and their preferential occurrence in specific tissues. Whole-plant
and leaf ontogeny had unique influence on the overall content
and relative abundances of the different cyanogenic glucosides.
An alternative approach was required to determine the nature
of the glycosidic linkage in the third unknown amygdalin isomer
(peak IV, Fig. 1a). This was because its low abundance precluded
NMR analyses and a natural or chemical route to a synthetic stan-
dard was not known. The approach was based on theoretical pre-
dictions and empirical characterization of MS fragmentation of
oligosaccharides (see Domon and Costello, 1988; Mulroney et al.,
2
. Results
2
2.1. Identification of cyanogenic diglucosides in E. camphora
1995). Specifically, MS of the third unknown yielded a proportion-
ally high intensity of fragment ions at m/z 113 and 161, which is
diagnostic for the presence of a b(1 ? 3) linked laminaribiose
disaccharide (Mulroney et al., 1995). Thus structure 7 was assigned
to the third unknown, named here eucalyptosin C (Fig. 2). Indeed
using this approach alone, all four cyanogenic diglucosides present
in E. camphora could be successfully characterized as harboring
b(1 ? 2), b(1 ? 3), b(1 ? 4) and b(1 ? 6) linkages, respectively,
highlighting the utility of this technique. In a similar manner, a
number of MS studies of oligosaccharides have used fragments
representing opening and cleavage across the ring of sugar moie-
ties to provide important diagnostic information on linkage posi-
tions (Cmelík and Chmelík, 2010; Domon and Costello, 1988; Li
and Her, 1998; Mulroney et al., 1995; Wong et al., 1999).
Cyanogenic glucosides from E. camphora seedling and sapling
foliage and adult foliage, flower buds and fruits were extracted
and separated using reverse-phase HPLC. Fractionation and eluci-
dation of prunasin, sambunigrin and amygdalin from adult E. cam-
phora foliage has been described previously (Neilson et al., 2006).
In all extracts except those from seedling foliage, the HPLC separa-
tion afforded additional fractions capable of releasing HCN upon
the addition of b-glucosidase.
Targeted LC-MS analysis of this fraction in negative mode for the
ꢀ
amygdalin pseudomolecular ion [M+C
calcd. 502.1555 for C20
resulted in three additional, analogous pseudomolecular ion peaks
1
H
1
O
2
]
at m/z 502.1585
(
H
27
N
1
O
11
C H
1 1
O
2
; k = ꢀ3.0 millimass units)
2
(
Fig. 1a). MS fragmentation of the formate adduct of each of these
parent ions yielded both glycosidic bond and cross ring bond cleav-
age. Major observed fragment ions in negative mode for amygdalin
and the three unknown diglucosides were m/z 323, 221, 179, 161,
2.2. Leaf- and whole-plant ontogeny and tissue specificity
Prunasin (1), amygdalin (3), eucalyptosin A (5), and the novel
compounds eucalyptosin B (6) and eucalyptosin C (7) were present
in all examined tissue types except for seedling leaves at 141 DAS,
which possessed only prunasin (Table 1, Fig. 3). Sambunigrin (2),
the epimer of prunasin, was not quantified in the different tissues
due to the inability to discriminate between epimers using LC-MS.
It is noteworthy that previous experimentation using GC–MS has
shown that 5% of the total cyanogenic glucosides in fully expanded
adult leaves of E. camphora can be present as sambunigrin (Neilson
et al., 2006). The relative abundances of the cyanogenic glucosides
varied across the different ontogenetic stages and in the different
tissue types. In adult expanding leaves (<50% of maximum size)
cyanogenic diglucosides constituted 6% of the total cyanogenic
1
31, 119, 113, 101, 89, 71 and 59 (Fig. 1c), corresponding to A
1 2
, A ,
B
1
, B , and C fragment ions using the nomenclature developed for
2
1
fragmentation of oligosaccharides (Domon and Costello, 1988;
Fig. 1b). Spectra recorded in positive mode were also recorded but
not as informative due to lack of diagnostic fragments derived from
cross ring bond cleavage.
Amygdalin has been reported to easily isomerise into neo-
amygdalin (4) especially when subjected to alkaline pH (Fig. 2;
Nahrstedt, 1975). To investigate whether one of the three un-
known cyanogenic glycosides could be neoamygdalin produced
as an artifact of the extraction procedure, a standard of neoamygd-
alin was synthesized by treatment of amygdalin with 5 mM NH
3

E.H. Neilson et al. / Phytochemistry 72 (2011) 2325–2334
2327
ꢀ
1 1 2
Fig. 1. Elucidation of cyanogenic diglucosides present in Eucalyptus camphora extracts using MS. (a) Extracted negative ion chromatograph of m/z 502.1585 [M+C H O ]
corresponding to the formate adduct of amygdalin reveals four separate peaks. (b) Nomenclature and fragmentation position of glycoconjugate product ions formed by
2
amygdalin (modified from Domon and Costello, 1988). (c) MS fragmentation of the parent ion of each peak (I–IV) yielded ions at m/z 323, 221, 179, 161, 131, 119, 113, 101,
8
9 and 59, corresponding to both glycosidic bond and cross ring cleavage, but with distinctly different abundances.
glucoside content. This increased to 17% in adult expanded leaves
with the concentration of eucalyptosin A, for example, increasing
from 3% to 7% as leaves developed. In flower buds and immature
fruits, cyanogenic diglucosides accounted for 27% and 31% of the
total cyanogenic glucoside concentration, respectively. In particu-
lar, high levels of eucalyptosin A were observed, accounting for
over 17% of total cyanogenic glucoside concentration in reproduc-
tive tissues. Adult fully expanded leaves and flower buds had sim-
ilarly high levels of total cyanogenic glucosides, but the total
concentration in fruits post-flowering was markedly lower with
levels intermediate between moderately cyanogenic saplings and
lowly cyanogenic seedlings (Table 1, Fig. 3).
Seedlings of E. camphora were repeatedly monitored for the
concentration of prunasin in foliage every few weeks until
339 days after sowing (DAS). The onset of prunasin biosynthesis
proceeded slowly and was first detected in three of the 57 seed-
lings at 115 DAS (Fig. 4). As the cohort of seedlings developed, they
progressively ‘‘switched on’’ prunasin production, until by 340 DAS
all individuals were cyanogenic (Fig. 4a). No relationship between
prunasin onset and plant height was observed (data not shown).

2
328
E.H. Neilson et al. / Phytochemistry 72 (2011) 2325–2334
Fig. 2. Structures of phenylalanine derived cyanogenic mono- and diglucosides present in Eucalyptus camphora. Prunasin is the most abundant cyanogenic glucoside in
foliage.
Once prunasin production was initiated, its foliar concentration in
each individual plant increased with time (Fig. 4b). Indeed, exam-
ination of the prunasin concentration in sapling leaves (1553 DAS)
and adult foliage (Table 1, Fig. 3) suggests prunasin concentration
continually increases as plants develop from seedlings, to saplings
and then to reproductively mature adult trees.
3. Discussion
3.1. Identification of multiple cyanogenic diglucosides
Three phenylalanine-derived cyanogenic diglucosides were
identified in E. camphora tissue: eucalyptosin A (5), eucalyptosin

E.H. Neilson et al. / Phytochemistry 72 (2011) 2325–2334
2329
Table 1
Total concentration and percentage abundance of the multiple cyanogenic glucosides present in different Eucalyptus camphora tissues.
Prunasin
Amygdalin
Eucalyptosin A
Eucalyptosin B
Eucalyptosin C
Total
ꢀ
1
ꢀ1
ꢀ1
ꢀ1
ꢀ1
ꢀ1
l
g g fw
%
l
g g fw
%
l
g g fw
%
l
g g fw
%
l
g g fw
%
lg g fw
Seedling expanded leaves (141 DAS)
Sapling expanded leaves (1553 DAS)
Adult expanding leaves
Adult expanded leaves
Flower buds
7
100
85
94
83
73
69
n.d.
36
19
97
45
20
n.d.
4
1
5
2
n.d.
41
57
143
367
87
n.d.
4
3
7
17
19
n.d.
58
29
93
147
29
n.d.
6
2
5
7
n.d.
n.d.
7
778
1806
1619
1548
312
<1
<1
<1
<1
913
1912
1952
2107
451
Immature fruits
4
6
3
1
Fig. 3. Cyanogenic glucoside composition at various developmental stages of Eucalyptus camphora analyzed in different tissues. Total cyanogenic glucoside concentration and
the relative abundances of the mono- and diglucosides differed in the various tissue types. Eucalyptosin C was detected in all sapling and adult tissues examined, but only at
ꢀ
1
very low levels (<1 lg g fw).
B (6) and eucalyptosin C (7), in which the glucose moieties are
combined by b(1 ? 2), b(1 ? 4) and b(1 ? 3) linkages, respec-
tively. This is in addition to the previous identification of the phen-
ylalanine-derived monoglucosides prunasin (1), and sambunigrin
Her, 2002; Daikoku et al., 2009) and anomeric configurations of
the glycosidic bond (Mulroney et al., 1995; Yamagaki and Sato,
2009).
In general, plant species containing aromatic cyanogenic gluco-
sides contain a single cyanogenic glucoside and in limited in-
stances, the corresponding b(1 ? 6) linked diglucoside. For
example, prunasin and amygdalin have been reported to co-occur
in the seeds of Rosaceaeous species (Dicenta et al., 2002; Miller
et al., 2004; Vetter, 2000). Similarly dhurrin, has been reported to
co-occur with its gentiobioside in S. bicolor (Poaceae; Selmar
et al., 1996). Although gentiobiosides are the most common disac-
charide associated with aromatic cyanogenic glucosides, other su-
gar moieties such as xylose and apiose have been reported in
cyanogenic diglycosides in Clerodendrum grayi (Lamiaceae; Miller
et al., 2006), Xeranthemum cylindraceum (Asteraceae; Schwind
et al., 1990), and in Davallia species (Davalliaceae; Kofod and
Eyjólfsson, 1969). In all the above instances, a b(1 ? 6) linkage ex-
ists between the two sugar moieties of the diglycosides except for
the aforementioned 5 isolated from E. maculata and Perilla frutes-
cens (Aritomi et al., 1985; Syah and Ghisalberti, 1996). It is there-
fore unprecedented that E. camphora was found to produce four
cyanogenic diglucosides with the glucose moieties coupled by
b(1 ? 6), b(1 ? 2), b(1 ? 3) and b(1 ? 4) linkages.
(2), and the b(1 ? 6) linked diglucoside amygdalin (3) in adult
leaves of this species (Neilson et al., 2006). Diglucoside 5 has pre-
viously been reported from the E. maculata (Myoporaceae; Syah
and Ghisalberti, 1996) and Perilla frutenscens var. acuta (Lamiaceae)
plants (Aritomi et al., 1985), whereas 6 and 7 are novel compounds.
Notably, neoamygdalin (4), the epimer of 3, was not identified in
any E. camphora tissue.
Structural elucidation of 5 and 6 was based on spectroscopic
comparison with authentic standards that were either isolated
from other plant species or chemically synthesized. The structural
assignment of the lowly abundant 7 was based on the consistency
2
of its MS fragmentation pattern with the empirical criteria for the
presence of a b(1 ? 3) linkage (Domon and Costello, 1988; Mulro-
ney et al., 1995). In retrospect, it would have been possible to iden-
tify the linkage types present within all four E. camphora
cyanogenic diglucosides using mass spectrometry alone, without
the need for NMR analyses. This knowledge should facilitate anal-
ysis and structural elucidation of cyanogenic diglucosides from
other plant species, especially when they are present in low abun-
dance. Similar approaches based on MS fragmentation of different
oligosaccharides have been used to distinguish between different
sugar residues (Garozzo et al., 1990; Verardo et al., 2009), stereo-
chemistry (Fang and Bendiak, 2007), linkage positions (Li and
Her, 1998; Spengler et al., 1990), branching patterns (Cheng and
The relative abundance of these diglucosides vary with plant
and tissue age and with tissue type. A similar structural diversity
of cyanogenic glucoside esters has been found in leaves of Phyllaga-
this rotundiflora (Melastomataceae) where seven different mono-,
di-, tri- and tetra-galloylated forms of prunasin positioned at the

2
330
E.H. Neilson et al. / Phytochemistry 72 (2011) 2325–2334
Fig. 4. The time dependent production of foliar prunasin in individually analyzed Eucalyptus camphora seedlings. (a) All 57 seedlings were initially incapable of synthesizing
prunasin. Prunasin biosynthesis was detected at differing starting points but all seedlings produced prunasin 339 days after sowing. (b) Once each individual plant began to
synthesize prunasin, its foliar concentration increased with time. Bars represent mean (±1 s.e.) of all cyanogenic seedlings at each harvest date.
0
0
0
0
C-2 , C-3 , C-4 and C-6 carbons were detected (Ling et al., 2002).
Although the authors showed that these compounds were absent
from root and stem tissues, no further tissue specificity or ontoge-
netic variation was examined.
Morrow and Lucas, 1986), thus forming a protective physical bar-
rier around the site of tissue damage (Stone and Clarke, 1992).
Regardless of whether hydrolysis occurs sequentially or simulta-
neously, it is likely that E. camphora possess multiple b-glucosi-
dases, each capable of acting upon a different glucosidic linkage
type and potentially specific to each of the different cyanogenic
diglucosides. High affinity towards particular disaccharide moie-
ties has been observed for other plant b-glucosidases. For example,
a b-primeverosidase from Camellia sinensis shows high selective
activity towards a b-primeveroside moiety, will hydrolyze other
3
.2. Possible diversification of cyanogenic diglucoside function
The combined presence of the two monoglucosides prunasin
and sambunigrin together with four cyanogenic diglucosides in
adult E. camphora tissues may provide a measure of defense supe-
rior to that achieved in plants containing a single cyanogenic glu-
coside. The cleavage of the cyanogenic diglucosides may occur
sequentially or simultaneously depending on whether the two su-
gar residues are removed in turn or as a disaccharide unit (Poulton,
disaccharides containing an
a(1–6) or b(1–6) linkage to some de-
gree, but does not hydrolyze
a(1–4) or b(1–4) linked disaccharide
moieties (Sakata et al., 2003). Similarly, a vicianin hydrolase from
Vicia angustifolia hydrolyzes its native substrate, the cyanogenic
glycoside vicianin with a b-vicianoside moiety, but not amygdalin
with a b-gentbioside moiety (Ahn et al., 2007). Future work should
focus on characterizing the enzyme(s) that hydrolyze the multiple
cyanogenic glucosides in E. camphora.
Although it is likely that the mono-glucosides in E. camphora
play a role in defense against herbivory, the cyanogenic digluco-
sides in this species may play additional roles related to nitrogen
storage and transport, pollinator attraction and possibly even seed-
ling germination. For example, it is known from studies on adult
1
990; Selmar, 1993). If hydrolysis of the four different cyanogenic
diglucosides present in E. camphora was to follow a simultaneous
pathway, the diglucosides gentiobiose, cellobiose, sophorose and
laminaribiose would be formed. Therefore, in addition to the toxic
cyanide and deterrent benzaldehyde released, the diglucosides
formed may serve as signals that elicit other strong plant defense
responses (Field, 2009; Shibuya and Minami, 2001; Morrow and
Lucas, 1986). For example, disaccharide formation may stimulate
(
1–3)-b-glucan synthase activity and callose production (see

E.H. Neilson et al. / Phytochemistry 72 (2011) 2325–2334
2331
Hevea brasiliensis trees that the cyanogenic diglucoside linustatin is
translocated away from its site of synthesis in leaves, whereas the
monoglucoside linamarin remains stored in leaves. Linustatin is
translocated to inner bark of the trunk where it is endogenously
catabolized and used for latex regeneration and rubber production
et al., 2006). The availability of eucalypt species exhibiting differ-
ent onset of cyanogenic glucoside production offers a unique mod-
el system to identify transcription factors involved in regulation of
cyanogenic glucoside formation.
(Kongsawadworakul et al., 2009). The diglucosides found in E. cam-
4
. Concluding remarks
phora may play a similar role in transport to organs such as devel-
oping leaves or flowers. Given the highest levels of diglucosides
were found in flower buds and expanded leaves of adult E. campho-
ra trees, it is tempting to suggest that the diglucosides are synthe-
sized in the fully expanded leaves and transported to the
developing flower buds. The much lower level of diglucosides
and indeed total cyanogenic glucosides in immature fruits suggests
nitrogen may have been remobilized from these compounds and
possibly used in flower development or incorporated into pollen
and nectar as pollinator cues. With respect to this latter possibility,
the honey bee (Apis mellifera) has been shown to be able to detect
and distinguish between different levels of amygdalin in floral
parts of almonds (London-Shafir et al., 2003).
Adult E. camphora foliage and reproductive tissues contain two
cyanogenic glucosides and four cyanogenic diglucosides. E. cam-
phora displays strong ontogenetic and tissue-specific control over
both the relative abundance of these compounds and total cyano-
genic glucoside content. These attributes make E. camphora an
excellent model species for studying the role of multiple cyano-
genic glucosides in plants, to identify the UDP-glucosyltransferases
responsible for diglucoside formation and to study the differential
degradation of the cyanogenic diglucosides by b-glucosidases.
5
. Experimental
Finally, the cyanogenic diglucosides may also function as regu-
lators of seed germination. For example, the cyanogenic digluco-
side eucalyptosin A (5) described here in E. camphora tissues has
previously been linked to germination inhibition in E. maculata
5
.1. General
LC-MS analyses were performed on an Agilent 1200 series LC
system (Agilent, Santa Clara, USA) connected to a quadrupole-
orthoganal time-of-flight (Q-TOF) mass spectrometer (Agilent
6
1
(
Richmond and Ghisalberti, 1994; Syah and Ghisalberti, 1996).
E. maculata grows in arid regions throughout central Australia
and chemical inhibitors in the fruit wall prevent germination until
significant rainfall leaches them out, thereby enabling seed germi-
nation to occur (Chinnock, 2007; Richmond and Ghisalberti, 1994).
In particular, 5 has been shown to be a component of the fruit wall
and indeed when this diglucoside was extracted and applied di-
rectly to seeds, germination was completely inhibited (Syah and
Ghisalberti, 1996). In a similar way, the cyanogenic diglucosides
in E. camphora may also signal, delay or inhibit regulatory path-
ways such as seed germination.
520 system) and fitted with a Zorbax SB-Aq column (2.1 mm ꢁ
ꢀ1
50 mm, 3.5 lm, 40 °C). Elution (0.5 mL min ) was carried out
using a MeCN (0.1% formic acid) gradient from 10% to 20% over
min, increased to 80% over 3.5 min, followed by 1 min at 80%.
ESI-MS was carried out in negative-ion mode (nebuliser pressure
5
4
1
0 psi, gas temperature 320 °C, capillary voltage 4000 V, fragmentor
50 V, skimmer 65 V and collision energy 25 V). The instrument was
operated in the extended dynamic range mode and data collected in
from 40–1700 amu with a scan rate of 2 scans s
GC–MS analysis was performed using a 7890A Agilent gas chro-
matograph coupled to a 5975C Agilent quadrupole mass spectrom-
eter (Agilent, Santa Clara, USA) fitted with a VF-5MS column (30 m
ꢀ
1
.
3.3. Distribution of cyanogenic glucosides at different ontogenetic
stages
with 0.2
lm film thickness; Agilent) and an Integra guard column
(
10 m; SGE, Ringwood, Australia). Samples were dried in vacuo and
Quantitative and qualitative variation in total cyanogenic gluco-
ꢀ1
derivatised for 120 min at 37 °C in 10
amine hydrochloride in pyridine followed by treatment for 30 min
at 37 °C with 20 L of BSTFA and 2 L of a retention time standard
mixture (0.029% (v/v) n-dodecane, n-pentadecane, n-nonadecane,
n-docosane, n-octacosane, n-dotriacontane, n-hexatriacontane dis-
solved in pyridine). Aliquots (1
umn using a hot needle technique at 250 °C and separated using a
temperature program from 70 to 325 °C (7 °C min , flow rate
0
lL of 30 mg mL methoxy-
side abundance was observed throughout different ontogenetic
stages in E. camphora. In particular, the onset of cyanogenic gluco-
side biosynthesis was highly variable with individuals ‘‘switching
on’’ prunasin biosynthesis for the first time between 115 and 339
DAS (Fig. 4a). By 340 DAS, all half-sibling family members were
cyanogenic, an observation consistent with the lack of acyanogenic
plants in two previously screened adult populations (Neilson et al.,
006). This is the first time such ontogenetic polymorphism has
been demonstrated, whereby genetically based variation in the
timing of chemical defense initiation is apparent.
Once prunasin production was initiated, cyanogenic capacity
increased through time with the biosynthesis of cyanogenic diglu-
cosides initiated during the developmental stage between seed-
lings 141 DAS and saplings 1553 DAS (Figs. 3 and 4, Table 1).
Total foliar cyanogenic glucoside concentration continually in-
creased from seeding to sapling to adult in a similar manner to that
observed for prunasin in Eucalyptus polyanthemos (Goodger et al.,
l
l
lL) were injected onto the GC col-
ꢀ1
2
ꢀ1
.8 mL min ). Authentic 3, 5 and 6 were used as standards.
1
NMR spectra ( H, cosydq, jres, noesy, tocsy) of 5 and 6 were re-
1
corded in methanol-d4 on a Bruker Avance 400 instrument ( H
NMR at 400 MHz, C NMR at 100.6 MHz). H shifts are relative
to internal TMS; C shifts are based on d (C6 ) = 62.6 ppm. All re-
ported C shifts were extracted from the 2D spectra hsqc or hmbc.
1
3
1
1
3
0
1
3
5.2. Plant material
2
004), where maximum prunasin concentrations were also ob-
E. camphora subsp. humeana L.A.S. Johnson & K.D. Hill (moun-
tain swamp gum; henceforth referred to as E. camphora) is a small
to medium-sized tree of southeastern Australia (Brooker and Klei-
nig, 2006). A population of adult trees was sampled from the Bux-
ton region, Victoria, Australia (37°25 22.8 S, 145°42 32.4 E) and
bulk samples of foliage (separated into expanding and expanded
leaves), flower buds, and immature fruit (non-dehisced) harvested
for cyanogenic glucoside compositional analysis in March, 2009.
Seed was also collected at this time and sown directly into individ-
ual pots and placed in a glasshouse (see Goodger et al. (2007) for
served in reproductively mature adults. Nevertheless this ontoge-
netic strategy appears to differ from that of other cyanogenic
eucalypts such as Eucalyptus yarraensis, where prunasin concentra-
tion reaches a maximum at 240 DAS (Goodger et al., 2007), and
Eucalyptus cladocalyx where prunasin concentration is greatest at
an even earlier stage (100 DAS; Goodger et al., 2006). Such variable
ontogenetic strategies in cyanogenic eucalypts are likely to reflect
a balance between growth and defense in environments with dif-
fering nutrient availability and herbivore pressures (Goodger
0
00
0
00

2
332
E.H. Neilson et al. / Phytochemistry 72 (2011) 2325–2334
germination and growth conditions). A bulk sample of fully ex-
panded leaves was harvested from the seedlings 141 DAS as part
of the compositional analysis. A previous cohort of 60 seedlings de-
rived from the same population was germinated in the same glass-
house in February 2005. A single leaf was harvested from each
seedling every few weeks until 339 DAS as part of the ontogenetic
study of cyanogenic capacity. At each harvest, plant height was
determined as a measure of biomass. Twenty-four of these plants
acetyl-b-
pyranose, 2,3,4,6-tetra-O-acetyl-b-
6-tri-O-acetyl- -glucopyranose, 2,3,4,6-tetra-O-acetyl-b-
pyranosyl-(1 ? 4)-2,3,6-tri-O-acetyl- -glucopyranosyl fluoride, 2-
phenyl-2-trimethylsilyoxyacetonitrile and 2-(S,R)-[(2,3,4,6-tetra-
O-acetyl-b- -glucopyranosyl)-(1 ? 4)-2,3,6-tri-O-acetyl-b- -gluco-
D
-glucopyranosyl-(1 ? 4)-1,2,3,6-tetra-O-acetyl-
-glucopyranosyl-(1 ? 4)-2,3,
-gluco-
D-gluco-
D
D
D
D
D
D
pyranosyloxy]-2-phenylacetonitrile as intermediates. Detailed
chemical synthesis and experimental data are to be published else-
0
00
0
00
were relocated to Alberton, Victoria (38°36 45 S, 146°39 58 E) on
2 October, 2006 and left to grow under natural environmental
where (Motawia et al., unpublished data). Structural characteriza-
1
2
tion: UV kmax (MeCN) 208 nm; H NMR (400 MHz, CD
3
OD) d: 7.62–
conditions. A bulk sample of fully expanded leaves was harvested
from these reproductively immature saplings as part of the compo-
sitional analysis in May, 2009 (1553 DAS). Adult E. maculata subsp.
brevifolia (Benth.) Chinnock (spotted emu bush or native fuchsia;
henceforth referred to as E. maculata) plants possessing fruit were
purchased from Wail Nursery (Wail, Victoria).
7.33 (5H, m, H-arom), 6.02, 5.90 (1H, 2s, CHCN, epimers), 4.74
(J = 8.0 Hz), 4.43 (J = 8.0 Hz), 4.42 (J = 8.0 Hz), 4.34 (J = 7.6 Hz) (4d,
b b 3
2H, H-1 & H-1 , epimers); C NMR (100 MHz, CD OD) d: 135.2,
0
13
134.9, 131.1, 130.8, 130.2, 130.2, 130.0, 130.0, 129.0, 129.0,
128.8, 128.8 (C-arom), 119.4, 118.5 (CN, epimers), 104.6, 104.6
0
(C-1, epimers), 102.2, 101.9 (C-1 , epimers), 80.5, 80.5,78.2, 78.2,
7
7.9, 77.9, 77.9, 77.1,76.9, 76.3, 76.3, 74.9, 74.9, 74.5, 71.4, 71.4
0
0
0
0
5.3. Structural elucidation of 5, 6 and 7
(C-2, C-3, C-4, C-5, C-2 , C-3 , C-4 , C-5 , cellobiose), 68.8, 68.7
0
(CHCN, epimers), 62.5, 62.5, 61.8, 61.6 (2C-6, 2C-6 , cellobiose epi-
ꢀ
The bulk samples of leaf and reproductive tissues were homog-
mers). MS (ESI ) m/z: 502.1533 (calcd. 502.1555 for C20
C H O ). MS fragmentation of [M+C H O ] (rel. int.): 161.0453
27
H N
1
O
11
2
ꢀ
enized in liquid N
2
using a mortar and pestle. Freshly homogenized
1 1 2 1 1 2
samples (20 g) of E. camphora leaves, flower buds and fruits as well
as E. maculata fruits were defatted using petroleum ether (sol-
vent:tissue, 10:1 v/w, four extractions), and then twice extracted
(100), 89.0242 (58), 101.0243 (46), 113.0241 (34), 59.0143 (29),
71.0142 (29), 456.1522 (29) and 119.0356 (25).
with cold MeOH. The MeOH filtrate was concentrated under N
and an equivalent volume of CHCl added with sufficient water
2
5
.3.3. Mandelonitrile b-laminaribioside, eucalyptosin C (7)
3
Cyanogenic diglucoside 7 was elucidated based on LC-MS data
to enable phase separation. The aqueous phase was collected, con-
centrated to dryness in vacuo, and the residual material re-sus-
pended in water, filtered and fractionated using an Alltech Maxi-
Clean C18 SPE cartridge (900 mg; Deerfield, USA) using gradient
2
and MS fragmentation patterns and fragment abundance. UV
ꢀ
kmax (MeCN) 208 nm; MS (ESI ) m/z: 502.1506 (calcd. 502.1555
2
ꢀ
for C20
H
27
N
1
O
11
1
C H
O
1 2
). MS fragmentation of [M+C
1 1
H O
2
]
(
rel. int.): 161.0448 (100), 323.0974 (85), 113.0237 (79),
ꢀ
1
2
elution (1 mL min , 0–100% MeOH–H 0). The fractions obtained
1
1
01.0246 (76), 89.0243 (66), 59.0141 (59), 71.0131 (33) and
19.0366 (30).
were concentrated in vacuo and cyanogenic glucoside derived
HCN release was monitored following addition of a commercial
preparation of emulsin, the cyanogenic glucoside cleaving b-gluco-
sidase from almond (Prunus amygdalis; b-D-glucoside glucohydro-
lase; EC 3.2.1.21, Sigma–Aldrich). Cyanogenic glucoside-
containing fractions were analyzed by HPLC using a Phenomenex
5.4. Quantitative determination of HCN potential and individual
cyanogenic glucosides
5.4.1. Individual cyanogenic glucosides
Luna C18 (2) column (250 mm ꢁ 10 mm, 5
lm particles; Torrance,
USA) and eluted with 20% MeCN (2 mL min ).
ꢀ
1
Quantification of individual cyanogenic glucosides present in
different E. camphora tissues was performed by LC-MS. Homoge-
nized plant material (25 mg) was boiled in 80% MeOH (1 mL,
5
hydrophobic components present in the sample, the supernatant
was diluted to a final concentration of 50% MeOH and one volume
Amygdalin standard was purchased from Sigma–Aldrich (Syd-
ney, Australia). Neoamygdalin was synthesized from the amygda-
lin standard (100 mg) by addition of 10 mL of 5 mM ammonia
and incubation (room temperature, 2 h). During this time, equilib-
rium between amygdalin and neoamygdalin was established
min), vortexed (15 s) and centrifuged (10 min, 5000g). To remove
of CHCl
tained following vortexing (15 s) and centrifugation (5 min,
000g) was diluted 1:20 and filtered (0.22 m low-binding Dura-
3
added to enable phase separation. The MeOH phase ob-
(Fischer, 1885; Koo et al., 2005).
5
l
5.3.1. (R)-mandelonitrile b-sophoroside, eucalyptosin A (5)
pore membrane). Quantification was performed by comparing
the extracted ion chromatogram of the different cyanogenic gluco-
sides to an external standard series of authentic prunasin (Sigma,
St. Louis, USA) and amygdalin of known concentrations (2–
200 ng). Duplicate samples were extracted and analyzed for each
tissue type and the resultant values are the means of the
duplicates.
Cyanogenic diglucoside 5 was compared to a standard isolated
from mature fruits of E. maculata (Syah and Ghisalberti, 1996).
Structural characterization: UV kmax (MeCN) 208 nm; 1H NMR
1
(
4
3
(
2
(
[
(
400 MHz, CD
3
OD) H NMR d: 7.65 (H-2), 7.45 (H-3), 5.91 (H-7),
00
0
0
.61 (J = 7.8 Hz, H-1 ), 4.51 (J = 7.1 Hz, H-1 ), 3.61 (H-2 ), 3.53 (H-
0
00 13
), 3.21 (H-2 ). C NMR (100 MHz, CD
C-2), 130.2 (C-3), 69.1 (C-7), 119.5 (C-8), 101.5 (C-1 ), 82.7 (C-
), 77.9 (C-3 ), 62.6 (C-6 ), 105.3 (C-1 ). MS (ESI ) m/z: 502.1525
calcd. 502.1555 for C20
3
OD) d: 135.1 (C-1), 128.6
0
0
0
0
00
ꢀ
H
27
N
1
O
11
1 1
C H O
2
). MS2 fragmentation of
5.4.2. Total cyanogenic capacity
Freeze-dried plant tissue (40 mg) was placed in a glass vial
with a 0.5 mL tube containing 100 lL 1 M NaOH. The vial tissue
ꢀ
M+C H
1 1
O
2
]
(rel. int.): 89.0245 (100), 161.0455 (87), 59.0142
65), 101.0243 (55), 71.014 (47), 456.1512 (45), 119.0347 (43),
13.0241 (38) and 131.0368 (30).
1
was hydrated with 1 mL of 0.1 mM citrate buffer (pH 5.5) and
the vial immediately sealed. Following incubation (18 h, 37 °C),
cyanide trapped in the NaOH solution was determined using a
miniaturized colorimetric (590 nm) microtiterplate based cya-
nide assay (Brinker and Seigler, 1989; Goodger et al., 2002). To
quantify the amount of cyanide each plant tissue can produce
naturally, i.e. cyanogenic capacity, no exogenous b-glucosidase
enzyme was added.
5
-
.3.2. Mandelonitrile b-cellobioside 2-(S,R)-[(4-O-b-
b- -glucopyranosyloxy]-2-phenylacetonitrile, eucalyptosin B (6)
Cyanogenic diglucoside 6 was chemically synthesized as an epi-
meric mixture (in the ratio of 2:1) in six steps using cellobiose and
benzaldehyde as starting materials and proceeding with b- -gluco-
pyranosyl-(1 ? 4)- -glucopyranose (cellobiose), 2,3,4,6-tetra-O-
D-glucopyranosyl)
D
D
D

E.H. Neilson et al. / Phytochemistry 72 (2011) 2325–2334
2333
nitrile metabolism. Proceedings of the National Academy of Sciences of the
United States of America 104, 18848–18853.
Acknowledgements
Jones, P.R., Andersen, M.D., Nielsen, J.S., Hoj, P.B., Møller, B.L., 2000. The
biosynthesis, degradation, transport and possible function of cyanogenic
glucosides. In: Romeo, J.T., Ibrahim, R., Varin, L., DeLuca, V. (Eds.), Evolution of
Metabolic Pathways, vol. 34, pp. 191–247.
Jørgensen, K., Bak, S., Busk, P.K., Sorensen, C., Olsen, C.E., Puonti-Kaerlas, J., Møller,
B.L., 2005a. Cassava plants with a depleted cyanogenic glucoside content in
leaves and tubers. Distribution of cyanogenic glucosides, their site of synthesis
and transport, and blockage of the biosynthesis by RNA interference technology.
Plant Physiology 139, 363–374.
E.H.N. was supported by the Holsworth Wildlife Research
Endowment (managed by ANZ Trustees). J.Q.D.G., B.L.M. and
I.E.W. were supported by a Grant from the Australian Research
Council (Project DP1094530). B.L.M. acknowledges financial sup-
port from the Villum Foundation to the Research Centre ‘‘Pro-ac-
tive Plants’’ and a Research Grant from the Danish Research
Council for Independent Research/Technology and Production Sci-
ences E.H.N. acknowledges Metabolomics Australia for advice.
Jørgensen, K., Rasmussen, A.V., Morant, M., Nielsen, A.H., Bjarnholt, N., Zagrobelny,
M., Bak, S., Møller, B.L., 2005b. Metabolon formation and metabolic channeling
in the biosynthesis of plant natural products. Current Opinion in Plant Biology 8,
2
80–291.
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