Benzoic acid glucosinolate esters and other glucosinolates from Arabidopsis thaliana.

Article abstract of DOI:10.1016/S0031-9422(02)00014-6

The spectacular recent progress in Arabidopsis thaliana molecular genetics furnishes outstanding tools for studying the formation and function of all metabolites in this cruciferous species. One of the major groups of secondary metabolites in A. thaliana is the glucosinolates. These hydrophilic, sulfur-rich glycosides appear to serve as defenses against some generalist herbivores and pathogens, and as feeding and oviposition stimulants to specialist herbivores. To help study their biosynthesis and role in plant-insect interactions, we wanted to determine the complete glucosinolate content of A. thaliana. In previous studies, 24 glucosinolates had been identified from ecotype Columbia. We reinvestigated Columbia as well as additional ecotypes and mutant lines, and identified 12 further glucosinolates, including five novel compounds. Structures were elucidated by MS and NMR spectroscopy of their desulfated derivatives, and by enzymatic cleavage of the attached ester moieties. Four of the novel glucosinolates are benzoate esters isolated from the seeds. In all but one of these compounds, esterification is on the glucose moiety rather than the side chain, a very unusual feature for glucosinolates. Among additional glucosinolates identified were the first non-chain elongated, methionine-derived glucosinolate from A. thaliana and the first compounds that appear to be derived from leucine.

Full text of DOI:10.1016/S0031-9422(02)00014-6

Phytochemistry 59 (2002) 663–671
www.elsevier.com/locate/phytochem
Benzoic acid glucosinolate esters and other glucosinolates from
Arabidopsis thaliana
Michael Reichelt, Paul D. Brown1, Bernd Schneider, Neil J. Oldham, Einar Stauber,
Jim Tokuhisa, Daniel J. Kliebenstein2, Thomas Mitchell-Olds, Jonathan Gershenzon*
Max-Planck Institute for Chemical Ecology, Beutenberg Campus, Winzerlaer Strasse 10, D-07745 Jena, Germany
Received 1 October 2001; received in revised form 19 December 2001
Abstract
The spectacular recent progress in Arabidopsis thaliana molecular genetics furnishes outstanding tools for studying the formation
and function of all metabolites in this cruciferous species. One of the major groups of secondary metabolites in A. thaliana is the
glucosinolates. These hydrophilic, sulfur-rich glycosides appear to serve as defenses against some generalist herbivores and patho-
gens, and as feeding and oviposition stimulants to specialist herbivores. To help study their biosynthesis and role in plant-insect
interactions, we wanted to determine the complete glucosinolate content of A. thaliana. In previous studies, 24 glucosinolates had
been identified from ecotype Columbia. We reinvestigated Columbia as well as additional ecotypes and mutant lines, and identified
12 further glucosinolates, including five novel compounds. Structures were elucidated by MS and NMR spectroscopy of their
desulfated derivatives, and by enzymatic cleavage of the attached ester moieties. Four of the novel glucosinolates are benzoate
esters isolated from the seeds. In all but one of these compounds, esterification is on the glucose moiety rather than the side chain, a
very unusual feature for glucosinolates. Among additional glucosinolates identified were the first non-chain elongated, methionine-
derived glucosinolate from A. thaliana and the first compounds that appear to be derived from leucine. # 2002 Elsevier Science
Ltd. All rights reserved.
Keywords: Arabidopsis thaliana; Brassicaceae; Crucifers; Structure elucidation; Glucosinolates; Benzoic acid esters; Benzoyloxy glucosinolates;
Sinigrin; Myrosinase; Methionine
1. Introduction
One glucosinolate-producing species, Arabidopsis
thaliana (L.) Heynh., has become a model system for
studying many aspects of plant biology and biochem-
istry. The availability of extensive mutant collections,
mapping tools and the full genomic sequence for this
taxon provide extraordinary resources for studying the
formation and function of all metabolites present,
including glucosinolates. As part of our interest in the
biosynthesis of glucosinolates in A. thaliana and their
role in plant defense, we wanted to determine the com-
plete glucosinolate profile of this species.
Previous work on this species has identified 24 gluco-
sinolates (Hogge et al., 1988b; Agerbirk et al., 2001)
which can be divided into three classes based on their
likely amino acid precursor: methionine, phenylalanine
or tryptophan (Table 1). The methionine-derived glu-
cosinolates are the largest and structurally most variable
class, with methylthioalkyl, methylsulfinylalkyl, alkenyl
and hydroxyalkyl side chains being found. In the seeds,
the hydroxyalkyl glucosinolates often occur as benzoate
esters (Hogge et al., 1988b; Graser et al., 2001). The
Glucosinolates are sulfur-rich plant secondary meta-
bolites whose basic skeleton consists of a b-thioglucose
residue, an N-hydroxyiminosulfate moiety and a vari-
able side chain (Halkier, 1999). More than 120 different
glucosinolates have been identified (Fahey et al., 2001),
almost exclusively from species of the Brassicaceae (the
mustard family) and several closely related families.
These substances play prominent roles in plant-herbi-
vore and plant-pathogen interactions (Chew, 1988;
Halkier, 1999), and are attracting increasing attention
for their anticarcinogenic properties (Hecht, 2000).
* Corresponding author. Tel.: +49-3641-571300; fax: +49-3641-
571302.
E-mail address: gershenzon@ice.mpg.de (J. Gershenzon).
1
Current address: Department of Environmental Studies, Trinity
Western University, Langley, British Columbia V2Y 1Y1, Canada.
2
Current address: Department of Vegetable Crops, University of
California, Davis, CA 95616, USA.
0031-9422/02/$ - see front matter # 2002 Elsevier Science Ltd. All rights reserved.
PII: S0031-9422(02)00014-6

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M. Reichelt et al. / Phytochemistry 59 (2002) 663–671
Table 1
Glucosinolates identified from Arabidopsis thaliana (Columbia and other ecotypes)
Systematic name
Common name
Structure
Glucosinolates derived from methionine
With methylthioalkyl side chains
3-Methylthiopropyl glucosinolate
4-Methylthiobutyl glucosinolate
5-Methylthiopentyl glucosinolate
6-Methylthiohexyl glucosinolate
7-Methylthioheptyl glucosinolate
8-Methylthiooctyl glucosinolate
Glucoiberverin
Glucoerucin
Glucoberteroin
Glucolesquerellin
With methylsulfinylalkyl side chains
2- Methylsulfinylethyl glucosinolatea,b
3-Methylsulfinylpropyl glucosinolate
4-Methylsulfinylbutyl glucosinolate
5-Methylsulfinylpentyl glucosinolate
6-Methylsulfinylhexyl glucosinolate
7-Methylsulfinylheptyl glucosinolate
8-Methylsulfinyloctyl glucosinolate
Glucoiberin
Glucoraphanin
Glucoalyssin
Glucohesperin
Glucoibarin
Glucohirsutin
With hydroxyalkyl side chains
3-Hydroxypropyl glucosinolate
4-Hydroxybutyl glucosinolate
With alkenyl side chains
2-Propenyl glucosinolate (allyl)a,c
3-Butenyl glucosinolate
Sinigrin
Gluconapin
Glucobrassicanapin
4-Pentenyl glucosinolatea,c
With other side chains
3-Benzoyloxypropyl glucosinolate
4-Benzoyloxybutyl glucosinolate
5-Benzoyloxypentyl glucosinolate
6-Benzoyloxyhexyl glucosinolate
Glucomalcomiin
(2R)-2-Hydroxy-3-butenylglucosinolatea,c
(2S)-2-Hydroxy-3-butenylglucosinolatea,c
(2S)-2-Benzoyloxy-3-butenylglucosinolatea,c
Progoitrin
Epiprogoitrin
With modifications on the glucose moiety
60-Benzoyl-4-methylthiobutyl glucosinolatea
60-Benzoyl-4-methylsulfinylbutyl glucosinolatea
60-Benzoyl-4-benzoyloxybutyl glucosinolatea
see Fig. 1
see Fig. 1
see Fig. 1
Glucosinolates derived from leucine
3-Methylbutyl glucosinolatea,c
4-Methylpentyl glucosinolatea,c
(continued on next page)

M. Reichelt et al. / Phytochemistry 59 (2002) 663–671
665
Table 1 (continued)
Systematic name
Common name
Glucobrassicin
Structure
Glucosinolates derived from tryptophan
Indol-3-ylmethyl glucosinolate
4-Methoxyindol-3-ylmethylglucosinolate
1-Methoxyindol-3-ylmethylglucosinolate
4-Hydroxyindol-3-ylmethylglucosinolate
4-Methoxyglucobrassicin
Neoglucobrassicin
4-Hydroxyglucobrassicin
Glucosinolates derived from phenylalanine
Benzyl glucosinolatea,c
Glucotropaeolin
Gluconasturtiin
2-Phenethyl glucosinolate
a
Identified for first time in this study. Other compounds first identified by Hogge et al. (1988b) or Agerbirk et al. (2001).
Isolated from mutant TU1.
Isolated from ecotypes other than Columbia.
b
c
structural diversity of the methionine-derived glucosi-
anion-exchange matrix. The HPLC-UV chromatogram
revealed many previously reported desulfated glucosi-
nolates (Hogge et al., 1988b; Agerbirk et al., 2001),
identified by comparison of their retention times and
UV spectra to those of standards. Especially prominent,
were compounds with benzoyloxy substituents, char-
acterized by their distinctive UV spectra (absorption
maxima at 195 and 230 nm, shoulder at 275 nm). The
most abundant of this group were desulfo-3-benzoyloxy-
propyl and desulfo-4-benzoyloxybutyl (d1) glucosino-
late. Four other desulfated benzoyloxyalkyl glucosinolates,
whose identities were unclear from the chromatogram,
were purified by preparative HPLC and subjected to
MS and NMR analyses. Three proved to be the novel
substances d3–d5 (described below), while one was
desulfo-5-benzoyloxypentyl glucosinolate (d2), pre-
viously reported by Hogge et al. (1988b) based on the
occurrence of the corresponding isothiocyanate hydro-
lysis products following treatment with myrosinase.
Spectral data for d2 are also presented here since this
was the first isolation of this compound.
nolates is increased further by the elongation of their
side chains with 1–6 additional methylene groups.
Here we report the isolation of 12 new glucosinolates
from A. thaliana. Of these, five (3–7) are novel, methio-
nine-derived compounds, all but one of which are
benzoate esters. Previous investigations have focused on
the Columbia ecotype (Col-0). We have reinvestigated
this ecotype (three of the five novel glucosinolates (3–5)
were isolated from Columbia), as well as searching for
novel constituents in additional ecotypes and mutants.
The complete distribution of glucosinolates in 39
Arabidopsis ecotypes has been reported separately
(Kliebenstein et al., 2001a).
In the present study, structure elucidation (UV, MS
and NMR) was carried out on the desulfated derivatives
of the isolated glucosinolates (d1–d7), because on-col-
umn desulfation is a very efficient step in separating
glucosinolates from other components of the plant
extracts (Thies, 1979). However, the naturally occurring
compounds are the sulfated glucosinolates (1–7) (Fig. 1).
Atmospheric pressure chemical ionization MS in a
positive ion mode resulted in an m/z of 430, representing
the expected [M+H]+ ion of d2 (C19H28NO8S). Colli-
sion induced dissociation (CID) of m/z 430 yielded pro-
duct ions at m/z 268, 146, 137 and 105 compared to
fragments of m/z 254, 132 and 105 for d1. The 1H NMR
spectrum of d2 closely resembled that of d1 except for
an additional multiplet for a more shielded methylene
group at ꢀ 1.46, which was assigned to H2–3 (Table 2).
2. Results and discussion
2.1. Novel benzoic acid glucosinolate esters from seeds
Glucosinolates were extracted from seeds of the eco-
type Columbia (Col-0) and separated from other con-
stituents of the extract by on-column desulfation on an

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M. Reichelt et al. / Phytochemistry 59 (2002) 663–671
1
Characteristic signals of the benzoyl moiety appeared at
ꢀ 7.97 (H-200/H-600), 7.66 (H-400) and 7.53 (H-300/H-500).
The chemical shift of H2–5 (ꢀ 4.27) indicated attachment
of the benzoyloxy unit to the terminal position of the
ybutyl side chain. The H NMR spectrum of d3 was
very similar to that of d1, but the additional set of ben-
zoyl signals in the aromatic part of the spectrum indi-
cated the occurrence of a second benzoyl moiety
(Table 2). The absence of a signal for one of the four
sugar hydroxyl groups provided evidence for the
attachment of the additional benzoyl function to the
glucose unit. The downfield shift of signals for H-60a
and H-60bby nearly one full ppm indicated attachment
at C-60, which was confirmed by corresponding 13C
NMR shift differences (observed in the HMQC spectra)
between the benzoylated and nonbenzoylated glucose
moieties of compounds d3 and d1, respectively (Table 3).
Thus, 3 is 60-benzoyl-4-benzoyloxybutyl glucosinolate.
As compound 3 has two benzoic acid groups that
might require cleavage before metabolic activation by
myrosinase (Rask et al., 2000), it was interesting to test
the behavior of its desulfated derivative (d3) upon
esterase treatment. Compound d3 completely dis-
appeared from the HPLC-UV chromatogram in the first
minute after application of porcine liver esterase, and
was replaced by an unknown substance assigned by
MS/MS experiments to be desulfo-60-benzoyl-4-hydro-
xybutyl glucosinolate ([M+H]+ at m/z 416, fragments
at m/z 267, 249, 150, 105). After one hour of esterase
treatment, the HPLC-UV trace finally showed a signal
for desulfo-4-hydroxybutyl glucosinolate, indicating
1
alkyl chain. The H–1H COSY spectrum confirmed the
1
assignment of H signals of the benzoyl ring and the
aliphatic chain, and was also used to identify the glu-
cose signals including the hydroxyl protons. As a final
confirmation of its structure, d2 was treated with ester-
ase to yield desulfo-5-hydroxypentyl glucosinolate,
identified by comparison of its HPLC retention time
and UV spectrum with that of a standard.
The desulfated derivative of compound 3 showed the
typical UV spectrum of a benzoyloxyalkyl glucosino-
late. However, the parent ion (m/z 520 for [M+H]+)
did not correspond to that of any known compound of
this type. The product ion spectrum of m/z 520 showed
a fragment at m/z 105 indicative of a benzoyl moiety,
but did not show any fragment at [M+H-162]+ repre-
senting the loss of the glucose moiety, as seen in the
mass spectrum of all typical desulfated glucosinolates.
Instead, a strong fragment at m/z 254 was present which
could be attributed to the loss of glucose and a benzoyl
moiety together [M+H-266]+, along with prominent
fragments at m/z 267, 249 and 132. The fragments at
m/z 254 and 132 were also observed for d1 (Graser et
al., 2001) and indicated the presence of a 4-benzoylox-

Table 2
1H NMR spectral data of desulfonated derivatives of compounds 1–5 and 7a
H
d1
d2
d3
d4
d5
d7
Aliphatic moiety 1
2
2.61 (m)
1.73 (m)
1.76 (m)
4.30 (t, J=7.7 Hz)
–
2.55 (m)
1.64 (m)
2.44 (m) / 2.58 (m)
1.55 (m) / 1.66 (m)
1.57 (m)
2.37 (m)
1.48 (m)
1.40 (m)
2.33 (m)
2.61 (m)
1.53 (m)
1.51 (m)
2.63 (m)
3.04 (m)
3.08 (m) / 3.25 (m)
3
1.46 (m)
1.74 (m)
4.27 (t, J=6.6 Hz)
4
4.16 (m)
5
S(O)CH3
SCH3
N–OH
2.46 (s)b
2.68 (s)b
1.96 (s)b
10.95 (s)
10.93 (s)
10.90 (s)
10.98 (s)
10.98 (br s)
n.d.c
Glucose moiety 10
4.70 (d, J=9.9 Hz)
4.67 (d, J=9.9 Hz)
3.04 (dd, J=9.9, 9.2 Hz) 3.03 (dd, J=9.9, 9.2 Hz) 3.09 (m)
4.81 (d, J=9.9 Hz)
4.79 (d, J=9.9 Hz)
3.08 (m)
3.28 (obscured by HDO)
4.80 (d, J=9.5 Hz)
3.09 (m)
5.04 (d, J=9.8 Hz)
3.44 (dd, J=9.8, 9.8 Hz)
3.55 (dd, J=9.8, 9.8 Hz)
20
30
3.21 (dd, J=9.2, 8.4 Hz) 3.21 (dd, J=9.2, 8.4 Hz) 3.28 (obscured by HDO)
ꢀ3.3
(obscured by HDO)
3.21 (m)
40
50
60a
60b3.67 (
20-OH
30-OH
40-OH
60-OH
3.08 (dd, J=8.4, 9.2 Hz) 3.09 (dd, J=8.4, 9.2 Hz) 3.20 (m)
3.19 (m)
3.65 (ddd, J=8.8, 7.7, 1.2 Hz) 3.64 (ddd, J=8.8, 7.7, 1.5 Hz) 3.66 (m)
3.44 (dd, J=9.8, 9.8 Hz)
3.57 (dd, J=9.8, 2.2 Hz)
3.18 (m)
3.78 (br d, J=11.7 Hz)
dd, J=11.7, 6.2 Hz) 3.39 (dd, J=11.7, 6.0 Hz) 4.23 (dd, J=11.7,7.7 Hz)
3.17 (m)
3.66 (dd, J=11.7,5.1 Hz) 4.51 (dd, J=11.7,1.2 Hz)
4.62 (dd, J=11.7,1.5 Hz)
4.22 (dd, J=11.7,7.7 Hz)
5.46 (d, J=5.9 Hz)
5.26 (d, J=5.1 Hz)
5.37 (d, J=5.5 Hz)
–
4.61 (dd, J=11.4,5.1 Hz) 3.89 (dd, J=12.5,2.2 Hz)
4.24 (dd, J=11.4,7.7 Hz) 3.69 (dd, J=12.5,5.9 Hz)
5.36 (br s)
4.99 (br s)
5.09 (br s)
4.58 (br s)
5.34 (d, J=5.9 Hz)
4.99 (d, J=5.5 Hz)
5.10 (d, J=5.1 Hz)
4.57 (t, J=5.8 Hz)
5.49 (d, J=5.9 Hz)
5.27 (brs)
5.49 (d, J=7.0 Hz)
5.40 (d, J=5.1 Hz)
5.48 (d, J=5.1 Hz)
–
n.d.
n.d.
n.d.
n.d.
5.38 (d, J=4.8 Hz)
–
Benzoyl moiety 200/600
7.96 (dd, J=8.4, 1.5 Hz) 7.97 (d, J=8.4 Hz)
7.66 (tt, J=7.3, 1.5 Hz) 7.66 (t, J=7.7 Hz)
7.93 (d, J=8.4 Hz) /
7.95 (d, J=8.4 Hz)
7.60 (t, J=7.7 Hz) /
7.66 (t, J=7.7 Hz)
7.96 (d, J=8.4 Hz)
7.68 (t, J=7.3 Hz)
7.55 (dd, J=8.4,7.3 Hz)
7.95 (dd, J=8.4, 1.5 Hz)
7.68 (t, J=7.3 Hz)
400
300/500
7.53 (dd, J=7.3, 8.4 Hz) 7.53 (dd, J=7.7, 8.4 Hz) 7.50 (dd, J=8.4, 7.7 Hz)/
7.53 (dd, J=8.4, 7.7 Hz)
7.56 (d, J=8.4, 7.3 Hz)
a
b
c
Chemical shifts (ꢀ) were recorded at 500.13 MHz with DMSO-d6 (d1–d5) and D2O (d7) as solvents and TMS (d1–d5) and TMSP-2,2,3,3-d4 (d7) as internal standards.
Three proton intensity.
n.d.=Not detected.

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M. Reichelt et al. / Phytochemistry 59 (2002) 663–671
Table 3
13C NMR spectral data of desulfonated derivatives of compounds 1, 3,
4, and 7a
methylthiobutyl chain. 13C NMR data (Table 3),
obtained from an HMQC spectrum of d4, also con-
firmed that 4 is 60-benzoyl-4-methylthiobutyl glucosino-
late.
C
d1
d3
d4
d7
The desulfated derivative of compound 5 (d5) also
had a UV spectrum indicating the presence of a benzoyl
moiety, but eluted significantly earlier than d3 or d4 in
reversed-phase HPLC. Mass spectrometry of d5 gave a
molecular species ([M+H]+) at m/z 462. The MS/MS
experiment showed fragments at m/z 267 and 105,
which suggested benzoylation on the glucose moiety, in
analogy to d3 and d4. The base peak was at m/z 196,
consistent with a 4-methylsulfinylbutyl side chain, also
indicated by the formation of desulfo-4-methylsulfi-
nylbutyl glucosinolate upon treatment of d5 by esterase.
Aliphatic moiety
0
n.d.b
31.2
23.7
28.0
64.7
n.d.
30.5
22.9
27.3
64.1
n.d.
30.7
25.8
27.5
32.7
14.2
n.d.
30.1
54.2
1
2
3
4
SCH3
S(O)CH3
41.4
Glucose moiety
10
20
30
40
50
60
81.9
73.2
78.7
70.3
81.0
61.5
81.2
72.5
77.6
69.7
77.2
64.5
81.2
72.5
77.5
69.9
77.2
64.4
86.0
76.8
81.8
73.9
84.6
65.4
1
1H NMR (Table 2) and H–1H COSY experiments on
d5 proved the structure of compound 5 as 60-benzoyl-4-
methylsulfinylbutyl glucosinolate. In comparison with
Benzoyl moiety
COO
100
200/600
400
300/500
n.d.
n.d.
n.d.
n.d.
n.d.
n.d.
1
the H NMR spectrum of d4, major differences were
129.3 128.8 / 128.8 128.7
133.3 133.0 / 133.0 133.1
129.1 128.4 / 128.4 128.5
found only in the chemical shifts of the S-methyl signal
(ꢀ 2.46 vs. 1.96) and the methylene signal of H2–4 (ꢀ 2.63
vs. 2.33), which are both shifted downfield due to the
oxidation of the thiol function to a sulfoxide.
a
13C NMR data were obtained from HMQC spectra. Chemical
shifts (ꢀ) were recorded at 125.76 MHz with DMSO-d6 (d1–d4) and
D2O (d7) as solvents and TMS (d1–d4) or TMSP-2,2,3,3-d4 (d7) as
internal standards.
In the course of analyzing the seeds of 39 different
ecotypes of A. thaliana by HPLC-diode array detection
(Kliebenstein et al., 2001a), several ecotypes (e. g., Pog-
0, Tac) were observed to have a compound (d6) that
eluted only 0.2 min earlier than d1 and also gave a ben-
zoyloxy-type UV spectrum. The occurrence of this new
glucosinolate (6) in the sample was correlated with the
occurrence of (2S)-2-hydroxy-3-butenyl glucosinolate.
In fact, esterase treatment of isolated d6 gave desulfo-
(2S)-2-hydroxy-3-butenyl glucosinolate suggesting that
the structure of 6 is (2S)-2-benzoyloxy-3-butenyl gluco-
sinolate. The mass spectrum (molecular species at m/z
414) and CID-MS/MS (fragments at m/z 252 and 130)
of d6 were also consistent with the proposed structure.
Further fragmentation of the species at m/z 130 showed
the same pattern (m/z 78, 85, and 97) as that produced
by fragmentation of the m/z 130 arising from authentic
desulfo-(2S)-2-hydroxy-3-butenyl glucosinolate.
The seeds of A. thaliana are rich in benzoic acid glu-
cosinolate esters, and the present investigation has
added four new structures to the four reported pre-
viously (Hogge et al., 1988b). These substances are less
polar than their non-benzoylated counterparts, and may
represent adaptations for storage in the low water
environment of the seed. Three of the four new sub-
stances described here are esterified at C-6 of the glucose
moiety rather than on the terminal hydroxyl group of
the side chain. Other glucosinolates have been reported
that are also esterified to aromatic acids through C-6 of
the glucose moiety, including 60-sinapoyl-4-methylsulfi-
nyl-3-butenyl glucosinolate from Rhaphanus sativus
(Linscheid et al., 1980) and various isoferulic and sina-
pic acid esters of 4-methylsulfinyl-3-butenyl glucosino-
b
n.d.=Not detectable by means of HMQC.
that cleavage of the ester bond in the aliphatic side
chain is much faster than that in the glucose moiety.
From its UV spectrum, d4 was also likely to contain a
benzoyl moiety. However, the product of esterase
cleavage of d4 was not a desulfated hydroxyalkyl glu-
cosinolate but desulfo-4-methylthiobutyl glucosinolate.
Mass spectrometry of d4 showed a molecular species
([M+H]+) at m/z 446. The product ion spectrum of m/z
446 showed some of the same fragments as the product
ion spectrum of the [M+H]+ of compound d3, m/z 267,
249 and 105, suggesting a similar benzoate esterification
of the glucose moiety. However, the fragments from the
4-benzoyloxybutyl side chain (m/z 254 and 132) were
absent. Instead, a fragment at m/z 180 was found indi-
cative of a 4-methylthiobutyl side chain. This fragment,
which represents the aliphatic side chain remaining after
glucose cleavage, was also observed during the MS/MS
analysis of a standard sample of desulfo-4-methylthio-
butyl glucosinolate. The 1H NMR spectrum of d4
exhibited signals for an aliphatic chain of four methyl-
ene groups, a glucose moiety and, in contrast to d3, only
one benzoyl unit (Table 2). The chemical shifts of the
glucose signals of d4, especially those of H-50, H-60a,
and H-60b, closely resembled those of d3, indicating
attachment of the benzoyl unit to C-6 of the sugar. The
chemical shift of the methylene signal H2–4 at ꢀ 2.33,
which was assigned by means of a H–1H COSY spec-
trum, and the methyl signal (ꢀ 1.96) were indicative of a
1

M. Reichelt et al. / Phytochemistry 59 (2002) 663–671
669
late (Bjerg and Sørensen, 1987). In considering the
physiological significance of these substituents, we were
intrigued by the possibility that esterification on the glu-
cose moiety might impede hydrolysis by myrosinase (a
prerequisite to formation of the biologically-active hydro-
lysis products). However, in a preliminary experiment the
60-benzoyl glucosinolates from the seeds of ecotype
Columbia were hydrolyzed as readily as all of the other
glucosinolates in this tissue by a commercial preparation
of myrosinase from Sinapis alba (results not shown).
the literature, but not yet reported from this species,
including allyl (2-propenyl), 4-pentenyl, (2R)-2-
hydroxy-3-butenyl, (2S)-2-hydroxy-3-butenyl, benzyl
(see also Kliebenstein et al., 2001b), 3-methylbutyl and
4-methylpentyl glucosinolate (Table 1). Most of these
compounds are probably methionine-derived, but leu-
cine is the likely precursor of the last two. Curiously,
(2R)-2-hydroxy-3-butenyl glucosinolate is always accom-
panied by its (2S)-epimer, with the two compounds
always occurring in a ratio (2R:2S) of approximately 1:3
(listed incorrectly in Kliebenstein et al., 2001a). All of
these substances were identified by comparison of UV
and mass spectra with those reported in the literature
(Hogge et al., 1988a; Griffiths et al., 2000; Tolra et al.,
2000; Kiddle et al., 2001), and by comparison of HPLC
retention times and spectral data with those of stan-
dards isolated from other species. Mass spectra were
obtained by atmospheric pressure chemical ionization-
MS and by electron impact-MS of the TMSi-derivatives
(Brown et al., 1991). In addition, the structures of the
last two compounds listed were confirmed by GC–MS
analysis of their myrosinase hydrolysis products using
methods previously described (Brown et al., 1994).
2.2. Other new glucosinolates from A. thaliana
Nearly all of the glucosinolates derived from methio-
nine are modified by the addition of at least one methy-
lene group to the side chain via a cyclic, three-step
pathway (Chapple et al., 1988; Graser et al., 2000). The
few such compounds without any side chain elongation
include 2-methylthioethyl glucosinolate (Groband
Matile, 1980) and 2-hydroxyethyl glucosinolate (Hu et
al., 1989). The TU1 and TU5 mutant lines of A. thaliana
ecotype Col-0 exhibit a block in the chain elongation
process such that 3-methylsulfinylpropyl glucosinolate
(the product of one chain elongation cycle) is a principal
metabolite instead of 4-methylsulfinylbutyl glucosino-
late (the product of two elongation cycles) (Haughn et
al., 1991; Kroymann et al., 2001). In the process of
characterizing these mutants for our biosynthetic stud-
ies, we observed an early-eluting peak in the HPLC
chromatogram of the TU1 leaf extract that was not
present in the extract from Col-0 wildtype leaves. The
UV spectrum was similar to that of the desulfated
methylsulfinylalkyl glucosinolates and the chemical
ionization mass spectrum of the isolated substance gave
a [M+H]+ parent ion at m/z 330 (C10H20NO7S2), in
accordance with that expected for the heretofore unre-
ported desulfo-2-methylsulfinylethyl glucosinolate (d7).
CID-MS/MS resulted in fragments at m/z 168 and 104,
in analogy with the fragmentation pattern known for
desulfo-3-methylsulfinylpropyl and desufo-4-methylsul-
finylbutyl glucosinolate (Hogge et al., 1988a; Kiddle et
3. Experimental
3.1. Plant material
Arabidopsis thaliana ecotype Columbia (Col-0) seeds
were purchased from Lehle Seeds (Round Rock, Texas,
USA). Seeds of other ecotypes and the TU1 and TU5
mutant lines were obtained from the Nottingham Ara-
bidopsis Stock Centre, Nottingham, UK (http://nasc.-
nott.ac.uk/) and the Arabidopsis Biological Resource
Center, Columbus, Ohio, USA (http://aims.cps.m-
su.edu/aims/index.html/). Plants were grown under a
mixture of 60 W cool white and broad spectrum bulbs
for a 16 h light/8 h dark cycle in a standard soil/vermi-
culite mixture. [See Kliebenstein et al. (2001a) for com-
plete details.]
1
al., 2001). The H NMR spectrum, measured in D2O,
(Table 2) showed signals for a glucose unit, two meth-
ylene groups (ꢀ 3.04 and 3.08/3.25), and a methyl signal
at ꢀ 2.68, with the downfield chemical shift of the latter
signal indicative of the adjacent sulfoxide function. Fur-
ther assignment of the proton signals by means of 1H–1H
COSY, and determination of 13C NMR shifts (Table 3)
from the HMQC spectrum of d7 helped confirm the
structure of 7 to be 2-methylsulfinylethyl glucosinolate,
the first non-chain elongated methionine-derived gluco-
sinolate to be isolated from A. thaliana. The configura-
tion at the sulfinyl group was not determined.
3.2. Glucosinolate extraction and desulfation on an
anion exchange matrix
Extracts of desulfated glucosinolates were prepared
using methods adapted from previously published pro-
tocols (Thies, 1979; Graser et al., 2000). For the bulk
extract of ecotype Columbia seeds, 16 g seeds were
immersed in 300 ml boiling water containing 8 ml 0.3 M
Pb(OAc)2–0.3 M Ba(OAc)2 solution for 10 min. After
grinding with an Ultraturrax homogenizer, the homo-
genate was centrifuged at 4000 g for 10 min. The pellet
was re-extracted with 150 ml water and centrifuged
again, and the combined supernatants loaded onto a
column of DEAE Sephadex A25 (10 g, Sigma, swelled
During our study of the various A. thaliana ecotypes
(Kliebenstein et al., 2001a), we identified seven other
glucosinolates that have been previously described in

670
M. Reichelt et al. / Phytochemistry 59 (2002) 663–671
in water). The column was rinsed exhaustively with 67%
(aq.) MeOH and subsequently with 20 mM MES buffer
(pH 5.2). The column was then treated with 4 ml sulfa-
tase solution (prepared as described by Graser et al.,
2001) and left to stand overnight. The resulting desul-
foglucosinolates were eluted from the column with 40
ml 60% (aq.) MeOH and 40 ml deionized water, and the
combined eluent evaporated to dryness with a rotary
evaporator. For HPLC separation, samples were redis-
solved in 8 ml water. Extracts of leaf samples were pre-
pared in the same manner except that, in the initial step,
100 mg of freeze-dried tissue were immersed in 4 ml
boiling water (Reintanz et al., 2001).
(4000 g, 10 min), the supernatant was loaded onto a
column of DEAE Sephadex. After washing with water
and formic acid/2-propanol/water solution, the intact
glucosinolate fraction was eluted from the column with
0.5 M K2SO4 directly into ethanol. This fraction was
centrifuged, the supernatant evaporated to dryness and
the residue redissolved in 1 ml of water. A portion of the
extract (100 ml) was treated with myrosinase (thioglucosi-
dase from Sinapis alba, EC 3.2.3.1, 859 units gꢂ1, Sigma)
b y mixing with 700ml 50 mM Tris (pH 7.0) and 100 ml
aqueous myrosinase solution (20 mg mlꢂ1). After 5 min at
room temperature, the mixture was boiled for 5 min to
denature the enzyme and then applied to a DEAE Sepha-
dex column. Elution of desulfoglucosinolates followed the
protocol described above. The resulting sample was ana-
lyzed by HPLC-UV and compared with the HPLC trace
of a non-hydrolyzed control sample (subjected to the
identical procedure but without myrosinase).
3.3. Desulfoglucosinolate fractionation and analysis
The desulfoglucosinolate fraction was separated by
HPLC on a Hewlett Packard HP 1100 Series system
with autosampler, diode-array detector and fraction
collector. The procedure employed a C-18, fully-end-
capped, reversed phase column (LiChrospher RP-18,
250ꢁ4.6 mm i.d., 5 mm particle size, Chrompack) oper-
ated at 1 ml minꢂ1 and 25 ꢃC. Elution was accomplished
with a gradient (solvent A: H2O, solvent B: MeCN) of
1.5–5% (v/v) B (6 min), 5–7% B (2 min), 7–21% B (10
min), 21–29%B (5 min), and 29–57% B (14 min), fol-
lowed by a cleaning cycle (57–93% B in 2 min, 5 min
hold, 93 to 1.5% B in 3 min, 6 min hold). Eluent was
monitored at 229 nm and desulfoglucosinolates were
collected with a fraction collector. Fractions were eva-
porated by use of a rotary evaporator and redissolved in
1 ml water. Purity of the fractions was checked by
HPLC-UV, and if necessary fractions were purified fur-
ther by HPLC using smaller fraction sizes.
3.6. NMR and MS analyses
NMR measurements were carried out on a Bruker
DRX 500 spectrometer at 500.13 MHz (1H), and 125.76
MHz (13C). DMSO-d6 was used as a solvent and TMS
as internal standard, except for compound 7, which was
measured in D2O (int. std. TMSP-2,2,3,3-d4). 1H NMR,
1H–1H COSY, and HMQC experiments were recorded
in a 2.5 mm inverse detection microprobe head.
Mass spectrometry was performed on a Quattro II
(Micromass Ltd, Altrincham, UK) tandem quadrupole
mass spectrometer (quadrupole-hexapole-quadrupole
geometry) equipped with an atmospheric pressure che-
mical ionization (APCI) source. The corona pin was
operated at 3.5 kV and the sample cone at 16 V. A
mixture of acetonitrile and water (1:1) was used as the
solvent, at a flow rate of 0.5 ml minꢂ1. Vaporization
was achieved with a nitrogenꢃsheath gas (300 l hꢂ1) and
drying gas (150 l hꢂ1) at 400 C. Standard mass spectra
were measured using the first quadrupole only. Product
ion mass spectra were recorded by setting the first
quadrupole to transmit the parent ion of interest and
scanning the second quadrupole from m/z 50 to 10 amu
above the [M+H]+ of the chosen precursor ion. Argon
was used as the collision gas at 2ꢁ10ꢂ3 mbar, and a
collision energy of 10–20 eV was employed to achieve
fragmentation. For LC/MS analysis, a Hewlett Packard
HP 1100 LC was connected to the mass spectrometer. A
flow rate of 1 ml minꢂ1 was used with 1:1 post-column
splitting. The solvent program was identical to that used
in the HPLC analysis described above.
3.4. Hydrolysis of benzoate esters
Putative benzoylated desulfoglucosinolates were trea-
ted with porcine liver esterase (EC 3.1.1.1, 3.2 M
(NH4)2SO4 suspension, Sigma). Desulfoglucosinolates
(ca. 5–10 mg in 20 ml from purified HPLC fractions)
were mixed with 78 ml 50 mM Tris (pH 8.0) and 2 ml
esterase suspension (ca. 10 units of activity) in a glass
vial and incubated for one hour at room temperature.
The total incubation mixture was loaded onto a DEAE
Sephadex column and both the flow-through and a 1 ml
60% MeOH eluent were collected and evaporated to
dryness. The sample was then resuspended in water and
analyzed by HPLC.
3.5. Hydrolysis of intact glucosinolates
Extracts of intact glucosinolates from A. thaliana eco-
type Columbia seeds were prepared according to Thies
(1988). Briefly, 2 g of seed were immersed in 40 ml of
boiling water. After homogenization and centrifugation
Acknowledgements
We thank John Allen for technical assistance, an
anonymous reviewer for valuable comments on the

M. Reichelt et al. / Phytochemistry 59 (2002) 663–671
671
manuscript, and the Deutsche Forschungsgemeinschaft
(grant FOR 383 to J.G.) and the Max-Planck Gesell-
schaft for financial support.
Hecht, S.S., 2000. Inhibition of carcinogenesis by isothiocyanates.
Drug Metabolism Reviews 32, 395–411.
Hogge, L.R., Reed, D.W., Underhill, E.W., 1988a. The identification
of desulfoglucosinolates using thermospray liquid chromatography-
mass spectrometry. Journal of Chromatographic Science 26, 348–
351.
References
Hogge, L.R., Reed, D.W., Underhill, E.W., Haughn, G.W., 1988b.
HPLC separation of glucosinolates from leaves and seeds of Arabi-
dopsis thaliana and their identification using thermospray liquid
chromatography-mass spectrometry. Journal of Chromatographic
Science 26, 551–556.
Agerbirk, N., Petersen, B.L., Olsen, C.E., Halkier, B.A., Nielsen, J.K.,
2001. 1,4-Dimethoxyglucobrassicin in Barbarea and 4-hydro-
xyglucobrassicin in Arabidopsis and Brassica. Journal of Agri-
cultural and Food Chemistry 49, 1502–1507.
Hu, Z., Lewis, J.A., Hanley, A.B., Fenwick, G.R., 1989. 2-Hydro-
xyethyl glucosinolate from Capparis masaikai of Chinese origin.
Phytochemistry 28, 1252–1254.
Bjerg, B., Sørensen, H., 1987. Isolation of intact glucosinolates by
column chromatography and determination of their purity. In:
Wathelet, J.-P. (Ed.), World Crops: Production, Utilization,
Description, Vol. 13: Glucosinolates in Rapeseed: Analytical
Aspects. Martinus Nijhoff Publishers, Dordrecht/Boston/Lancaster,
pp. 59–75.
Kiddle, G., Bennett, R.N., Botting, N.P., Davidson, N.E., Robertson,
A.A.B., Wallsgrove, R.M., 2001. High-performance liquid chroma-
tographic separation of natural and synthetic desulphoglucosino-
lates and their chemical validation by UV, NMR and chemical
ionisation–MS methods. Phytochemical Analysis 12, 226–242.
Kliebenstein, D.J., Kroymann, J., Brown, P., Figuth, A., Pedersen, D.,
Gershenzon, J., Mitchell-Olds, T., 2001a. Genetic control of natural
variation in Arabidopsis glucosinolate accumulation. Plant Physiol-
ogy 126, 811–825.
Brown, P.D., Morra, M.J., Borek, V., 1994. Gas chromatography of
allelochemicals produced during glucosinolate degradation in soil.
Journal of Agricultural and Food Chemistry 42, 2029–2034.
Brown, P.D., Morra, M.J., McCaffrey, J.P., Auld, D.L., Williams, L.,
1991. Allelochemicals produced during glucosinolate degradation in
soil. Journal of Chemical Ecology 17, 2021–2034.
Chapple, C.C.S., Decicco, C., Ellis, B.E., 1988. Biosynthesis of 2–20
methylthioethylmalate in Brassica carinata. Phytochemistry 27,
3461–3464.
Kliebenstein, D.A., Gershenzon, J., Mitchell-Olds, T., 2001b. Com-
parative quantitative trait loci mapping of aliphatic, indolic and
benzylic glucosinolate production in Arabidopsis thaliana leaves and
seeds. Genetics 159, 359–370.
Chew, F. S., 1988. Biological effects of glucosinolates. In: Cutler, H. G.
(Ed.), Biologically Active Natural Products. ACS Symposium Series
380, American Chemical Society, pp. 155–181.
Kroymann, J., Textor, S., Tokuhisa, J., Falk, K., Bartram, S., Ger-
shenzon, J., Mitchell-Olds, T., 2001. A gene controlling variation in
Arabidopsis thaliana glucosinolate composition is part of the
methionine chain elongation pathway. Plant Physiology 127, 1077–
1088.
Fahey, J.W., Zalcmann, A.T., Talalay, P., 2001. The chemical diver-
sity and distribution of glucosinolates and isothiocyanates among
plants. Phytochemistry 56, 5–51.
Linscheid, M., Wendisch, D., Strack, D., 1980. The structures of
sinapic acid esters and their metabolism in cotyledons of Raphanus
sativus. Zeitschrift fur Naturforschung C 35, 907–914.
Graser, G., Oldham, N.J., Brown, P.D., Temp, U., Gershenzon, J.,
2001. The biosynthesis of benzoic acid glucosinolate esters in Ara-
bidopsis thaliana. Phytochemistry 57, 23–32.
¨
Rask, L., Andreasson, E., Ekbom, B., Eriksson, S., Pontoppidan, B.,
Meijer, J., 2000. Myrosinase: gene family evolution and herbivore
defense in Brassicaceae. Plant Molecular Biology 42, 93–113.
Reintanz, B., Lehnen, M., Reichelt, M., Gershenzon, J., Kowalczyk,
M., Sandberg, G., Godde, M., Uhl, R., Palme, K., 2001. Bus, a
bushy Arabidopsis CYP79F1 knockout mutant with abolished
synthesis of short-chain aliphatic glucosinolates. Plant Cell 13, 351–
367.
Graser, G., Schneider, B., Oldham, N.J., Gershenzon, J., 2000. The
methionine chain elongation pathway in the biosynthesis of gluco-
sinolates in Eruca sativa (Brassicaceae). Archives of Biochemistry
and Biophysics 378, 411–419.
Griffiths, D.W., Bain, H., Deighton, N., Botting, N.P., Robertson,
A.A.B., 2000. Evaluation of liquid chromatography-atmospheric
pressure chemical ionisation-mass spectrometry for the identifica-
tion and quantification of desulphoglucosinolates. Phytochemical
Analysis 11, 216–225.
Thies, W., 1979. Detection and utilization of a glucosinolate sulfohy-
drolase in the edible snail, Helix pomatia. Naturwissenschaften 66,
364–365.
Grob, K., Matile, P., 1980. Capillary GC of glucosinolate-derived
horseradish constituents. Phytochemistry 19, 1789–1793.
Halkier, B. A. 1999. Glucosinolates. In: Ikan, R. (Ed.), Naturally
Occurring Glycosides: Chemistry Distribution and Biological Prop-
erties. John Wiley & Sons, pp. 193–223.
Thies, W., 1988. Isolation of sinigrin and glucotropaeolin from cruci-
ferous seeds. Fat Science and Technology 90, 311–314.
Tolra, R.P., Alonso, R., Poschenrieder, C., Barcelo, D., Barcelo, J.,
2000. Determination of glucosinolates in rapeseed and Thlaspi
caerulescens plants by liquid chromatography-atmospheric pressure
chemical ionization mass spectrometry. Journal of Chromatography
889, 75–81.
Haughn, G.W., Davin, L., Giblin, M., Underhill, E.W., 1991. Bio-
chemical genetics of plant secondary metabolites in Arabidopsis
thaliana: The glucosinolates. Plant Physiology 97, 217–226.