Uptake and transformation of pesticide metabolites by duckweed (Lemna gibba)

Article abstract of DOI:10.1021/jf061301g

Uptake and transformation of ¹⁴C-labeled metabolites from several pesticides, 3-methyl-4-nitrophenol (1), 3,5-dichloroaniline (2), 3-phenoxybenzoic acid (3), (R,S)-2-(4-chlorophenyl)-3-methylbutanoic acid (4), and (1RS)-trans-3-(2,2-dichlorovin

Full text of DOI:10.1021/jf061301g

6286 J. Agric. Food Chem. 2006, 54, 6286−6293
Uptake and Transformation of Pesticide Metabolites by
Duckweed (Lemna gibba)
TAKUO FUJISAWA,* MOTOHIRO KUROSAWA, AND TOSHIYUKI KATAGI
Environmental Health Science Laboratory, Sumitomo Chemical Company, Ltd., 2-1, Takatsukasa
4-Chome, Takarazuka, Hyogo 665-8555, Japan
Uptake and transformation of 14C-labeled metabolites from several pesticides, 3-methyl-4-nitrophenol
(1), 3,5-dichloroaniline (2), 3-phenoxybenzoic acid (3), (R,S)-2-(4-chlorophenyl)-3-methylbutanoic acid
(4), and (1RS)-trans-3-(2,2-dichlorovinyl)-2,2-dimethylcyclopropanecarboxylic acid (5), were examined
by using duckweed (Lemna gibba) in Hoagland’s medium. More uptake into duckweed from the
exposure water at pH 7.0 was observed for non-ionized 1 and 2 than for 3-5 in an ionized form, and
their hydrophobicity accounted for these differences. While carboxylic acids 4 and 5 were scarcely
transformed in duckweed, 1-3 mainly underwent phase II conjugation with glucose for 1 and 2, malic
acid for 3, glutamic acid for 2, and malonylglucose for 3, the chemical identities of which were confirmed
by various spectrometric analyses (LC-MS, LC-MS/MS, and NMR) and/or HPLC cochromatography
with reference synthetic standards.
KEYWORDS: Uptake and metabolism in duckweed (Lemna gibba); metabolite of pesticide; phase II
conjugation
INTRODUCTION
Pesticides and their degradates reach various water bodies
mainly through direct application, spray drift, foliar washing
by precipitation, erosion, and runoff from agricultural land. A
critical aspect of environmental assessment is to determine their
distribution, transformation, and accumulation in water, sedi-
ment, and biota in relation to ecotoxicological impacts. Although
the fate of a pesticide and its metabolites has been extensively
investigated in a water-sediment system (1), neither their effects
on aquatic species nor their metabolic profiles therein have been
extensively examined. Aquatic plants are known to be habitat
and food for many species, and hence, the distribution and
metabolism of pesticides in them become important in studying
their effects on an aquatic ecosystem. Several investigators have
studied the toxicity and/or metabolism of polychlorinated
phenols or DDT using aquatic plants such as duckweed and
elodea (2-6). Information about phase I metabolism in aquatic
plants is very limited. It is reported that, as in higher plants,
oxidation, hydrolysis, and decarboxylation proceed in duck-
weeds with no mineralization of chlorinated phenols or aniline,
and phase II metabolism (conjugation) seems to occur rapidly
to form glucose, malonylglucose, and aspartic acid conjugates
as common pathways. However, information about its metabolic
profile is still scarce for phenolic, anilinic, and carboxylic
compounds which are main partial structures of pesticides and
have been reported to be primary degradates and metabolites
in the environment via photolysis, hydrolysis, and microbial
degradation. The possible toxicity of transformation products
Figure 1. Chemical structures of 14C-labeled 1
−5.
from these metabolites is a concern with respect to not only
aquatic plants themselves but also aquatic species via food
chains.
To collect the basic information about biotic transformations
of pesticide metabolites, the aquatic angiosperm Lemna gibba
(duckweed) was chosen as a model species for this study.
Duckweeds are adapted to a wide variety of geographic and
climatic zones and can be found in all but waterless deserts
and permanently frozen polar regions (7). Members of the
Lemnaceae family are ubiquitous, ecologically important, easily
grown, and readily and reliably manipulated. Also, its value as
a test species for toxicity assessment has been discussed
worldwide, and it has been adopted as a test material in
guidelines used for registration of agrochemicals (8). Given the
wide distribution and ecological importance of duckweed, it is
indispensable for aquatic risk assessment in determining its
uptake and metabolic response to typical pesticide metabolites.
However, there has not, to our knowledge, been any comparative
study on pesticide metabolites in duckweed, simultaneously.
From this standpoint, we have conducted an uptake and
* Corresponding author. E-mail: fujisawat1@sc.sumitomo-chem.co.jp.
10.1021/jf061301g CCC: $33.50
© 2006 American Chemical Society
Published on Web 08/02/2006

Uptake and Transformation of Pesticide Metabolites by Duckweed
J. Agric. Food Chem., Vol. 54, No. 17, 2006 6287
subjected to combustion. The 14CO2 that was produced was absorbed
into 9 mL of Packard Carb-CO2 absorber, mixed with 15 mL of Packard
Permafluor scintillator, and the radioactivity was quantified by LSC.
The efficiency of combustion was determined to be greater than 90%.
Chromatography. High-performance liquid chromatography (HPLC)
was carried out using a Shimadzu LC-20AT pump linked in series with
an SPD-20A UV-vis detector and a Perkin-Elmer Radiomatic610TR
radiodetector equipped with a 500 µL liquid cell. Ultima-Flo AP
(Packard) was utilized as a scintillator. A Sumipax ODS A-212 column
was employed for both analytical and preparative purposes with a flow
rate of 1.0 mL/min. The following gradient systems were used for
typical analysis, separation, and purification of the metabolites: 0.01%
trifluoroacetic acid (solvent A) and acetonitrile (solvent B), 90% A
and 10% B at 0 min, 10% A and 90% B at 40 min, 0% A and 100%
B at 40.1 min, and 0% A and 100% B at 50 min (method B); 1/33 M
phosphate buffer (solvent A) and acetonitrile (solvent B), 90% A and
10% B at 0 min and 10% A and 90% B at 40 min (method C). Retention
times of 1-8, the glutamate conjugate of 2 (9), and the malonylglucose
conjugate of 3 (10) in method B are 17.4, 31.5, 29.1, 29.8, 32.0, 9.5,
18.2, 25.2, 22.9, and 22.3 min, respectively.
Thin-layer chromatography (TLC) was conducted using silica gel
60 F254 thin-layer chromato plates (20 cm × 20 cm, 0.25 mm thick, E.
Merck). The cochromatography between 8 isolated from duckweeds
and its reference standard was carried out with a solvent system of
toluene, ethyl formate, and formic acid (5:7:1, v/v/v). An autoradiogram
was prepared by exposing the TLC plate to a BAS-IIIs Fuji Imaging
Plate for several hours. The radioactivity on the imaging plate was
detected by using a Typhoon (Amersham Bioscience Co., Ltd.), and
the non-radiolabeled reference standard was detected by exposing the
chromato plate to ultraviolet light. The typical Rf value of 8 was 0.35.
Plant Material, Maintenance, and Treatment. Duckweed (L.
gibba) was obtained from a paddy field located at the Kasai experi-
mental farm of Sumitomo Chemical Co., Ltd. (Hyogo, Japan). The
duckweed plants were maintained in pots filled with a water/sediment
system collected from the paddy field. The plants were grown in a
greenhouse equipped with a quartz glass ceiling, and the temperature
was kept at 25 °C. The plants were appropriately grown using Hyponex
liquid fertilizer with an N:P:K ratio of 20:20:20 (Hyponex Japan). Prior
to the experiments, sediment was removed from roots under running
tap water for 10 min and sterilized with 0.5% sodium hypochlorite for
1 min (15), and the sodium hypochlorite was washed off the plants by
dipping them into 100 mL of sterilized water. Hoagland’s medium [pH
adjusted to 7.0 with 1 N NaOH (16)] was sterilized by autoclaving the
medium for 20 min at 120 °C with an SS-325 autoclave (Tomy). 14C
test substance was then dissolved in the medium, and the plants were
added. The treated plants were grown in glass beakers without any
closing caps in a greenhouse at 25 °C.
For investigation of the uptake of a chemical into duckweed, 0.1
µg/mL aqueous solutions (100 mL) of [14C]1-5 were individually
prepared in 200 mL glass beakers. 1-5 were isotopically diluted with
corresponding non-radiolabeled reference standards to give a total
radioactivity of ca. 83 kBq (5 000 000 dpm) in exposure water. A 3 g
sample of duckweed was exposed to each chemical, and sampling of
plants and exposure water was conducted on days 1, 2, and 4. During
the incubation, the mass of duckweed increased finally to 4.3-5.5 g.
To investigate metabolic profiles of each metabolite, 100 mL of
aqueous solutions of [14C]1-5 at the exaggerated concentration of 1
µg/mL were each prepared in a 200 mL beaker. The total radioactivity
in exposure water was set to be equal to those used for the plant uptake
experiments by isotopically diluting 14C labels with the corresponding
reference standards. Sampling was conducted during the fourth day of
exposure.
Extraction and Isolation of Metabolites. The harvested samples
were first divided into duckweed and exposure water by filtering them
through a glass wool plug. Sampled duckweeds and exposure water
were stored in a freezer (less than -20 °C) and a refrigerator (<4 °C),
respectively, until analysis was conducted. Duckweeds were cut into
small pieces by using scissors and then extracted using a homogenizer
(Nissei, AM-8) at 10 000 rpm for 10 min with an acetone/water mixture
(4:1, v/v) at a ratio of 5 mL/g of plant. The mixture was filtered through
a filter paper (pore size, 7 µm), and the residue was extracted two
metabolism study in L. gibba using one phenolic, one anilinic,
and three carboxylic compounds which are the primary degra-
dates from various kinds of agrochemicals (organophosphate,
pyrethroid, carbamate, etc.).
MATERIALS AND METHODS
Chemicals. The 14C-labeled test substances (Figure 1), 3-methyl-
4-nitrophenol (1), 3,5-dichloroaniline (2), 3-phenoxybenzoic acid (3),
(R,S)-2-(4-chlorophenyl)-3-methylbutanoic acid (4), and (1RS)-trans-
3-(2,2-dichlorovinyl)-2,2-dimethylcyclopropanecarboxylic acid (5), were
prepared in our laboratory according to the reported methods (9-12).
The chemical structures of radiolabeled compounds 1-5 were con-
firmed by LC-MS and by comparing their HPLC retention times with
those of the non-radiolabeled reference standards. The specific activities
of 1-4 uniformly labeled at the phenyl ring ([phenyl-14C]1, -2, and -3
and [phenoxyphenyl-14C]4) and 5 labeled at position 1 of the cyclo-
propane ring ([cyclopropane-1-14C]5) were ca. 5.0 and 2.0 GBq/mmol,
respectively. Their radiochemical purity was more than 97% as
determined by HPLC. The non-radiolabeled authentic standards, 1-3,
were purchased from Aldrich (Milwaukee, WI). 4 and 5 were
synthesized in similar manners to the corresponding radiolabels. Glucose
conjugates of 1 (6) and 2 (7) were synthesized by modifying the
procedure reported by Sinnott et al. and Mitts et al., respectively (13,
14). The acid chloride of 3, prepared by treatment with oxalyl chloride,
was reacted in tetrahydrofuran with dibenzyl 2-hydroxysuccinate,
prepared from DL-malic acid, to produce the malate conjugate of 3 (8)
with its two carboxylic acids protected by benzyl groups. The catalytic
reduction of this derivative by 10% Pd-C under a hydrogen atmosphere
yielded the authentic standard of 8. All intermediates and 8 were purified
by silica gel column chromatography. The chemical structures of 6-8
were confirmed by 1H NMR (δH vs TMSP, parts per million) and LC-
ESI-MS (m/z) spectrometries as follows. 6: 1H NMR δ 3.61 (s, 3H,
aromatic-CH3), 3.40-5.10 (m, 7H, sugar-H), 7.05-8.01 (d, 3H,
aromatic-H); MS m/z 316 ([M + H]+). 7: 1H NMR δ 3.30-4.60 (m,
7H, sugar-H), 6.75-6.80 (s, 3H, aromatic-H); MS m/z 324 ([M + H]+).
8: 1H NMR δ 2.97 (dd, 2H, CHCH2), 5.54 (t, 1H, CHCH2), 7.05-
7.86 (m, 9H, aromatic-H); MS m/z 329 ([M - H]-).
Other reagents were of the purest grade commercially available.
1
Spectroscopy. H NMR and H-H COSY spectra were measured
with a Varian Unity 400 FT-NMR spectrometer operating at 400.45
MHz with a 5 mm PFG ATB probe, using trimethylsilylpropionate-
2,2,3,3-d4 (TMSP) as an internal standard (δ ) 0.0 ppm). Liquid
chromatography-atmospheric chemical ionization-mass spectrometry
(LC-APCI-MS) and liquid chromatography-electrospray ionization-
mass spectrometry (LC-ESI-MS) in positive and negative ion modes
were simultaneously performed using a Waters Micromass ZQ spec-
trometer equipped with Waters separation module model 2695 and
photodiode array detector model 2996 as a liquid chromatograph. Liquid
chromatography-electrospray ionization-tandem mass spectrometry
(LC-ESI-MS/MS) in positive and negative ion modes with a collision
energy of 10-40 V was conducted using a ThermoFinnigan TSQuan-
tum instrument attached with an Agilent 1100 series liquid chromato-
graph. Samples dissolved in methanol were manually injected into an
ionization source through a Sumipax ODS A-212 column (150 mm ×
6 mm inside diameter, 5 µm, SCAS Co., Ltd.) with a flow rate of 1.0
mL/min using a gradient system with acetonitrile (solvent A) and 0.1%
formic acid in water (solvent B). The composition of the mobile phase
was changed stepwise as follows: 10% A and 90% B at 0 min to 90%
A and 10% B at 40 min (method A).
Radioassay. Radioactivity in the plant extracts and exposure water
was determined by liquid scintillation counting (LSC). An aliquot of
the sample was mixed with 10 mL of Packard Scintillator Plus and
counted on a Packard model 1600TR and 2000CA liquid scintillation
counter equipped with an automatic external standard. The average
background of the LSC instrument was 30 dpm which was subtracted
from the measured sample disintegrations per minute. Radioactivity in
the unextractable residues from the treated plants was measured by
using a Packard model 306 sample oxidizer. Unextractable plant
residues were air-dried at room temperature overnight and weighed
with a Mettler model AE240 scale. An aliquot of each sample was

6288 J. Agric. Food Chem., Vol. 54, No. 17, 2006
Fujisawa et al.
Table 1. Physicochemical Properties of 1−5
a
compound
log Pa
2.48
2.69
3.91
3.38
0.5 (pH 7)
pKa
Henry’s law constantb
-8
-6
-8
-7
-6
1
2
3
4
5
8.33
2.37
3.95
4.65
5.02
1.61
8.11
5.27
3.81
4.20
×
×
×
×
×
10
10
10
10
10
−
a Experimental values from refs 32 36. b Calculated values using EPI-Suite, in
atmospheres per cubic meter per mole.
Table 2. Radioactivity and Metabolite Distribution of 3
% of the applied 14
C
day 1
day 2
day 4
duckweed 14
C
extractable
3
17.7
5.6
11.9
3.0
14.2
4.4
Figure 2. Radioactivity taken up by duckweed.
8
7.7
6.4
8.6
10
3.0
1.0
73.3
2.4
0.6
78.0
1.2
1.0
60.9
additional times in the same way. Aliquots of exposure water and plant
extracts were individually radioassayed by LSC. Plant extract from the
uptake experiment of 3 and plant extracts and exposure water from
the metabolic experiments of 1-5 were analyzed by HPLC cochro-
matography with authentic standards. Plant residues were air-dried in
open vessels at room temperature for 1 week, and subsamples of the
dried residues were subjected to combustion analysis to determine the
remaining radioactivity.
unextractable
exposure water 14
C
recovered 14
C
92.0
90.5
76.1
dependent on substituents at the phenyl ring (18). The physi-
cochemical properties of each compound tested such as dis-
sociation constant (pKa) and octa-1-nol/water partition coeffi-
cient (log P) were likely to account for these profiles. Since
the pH of the incubation medium was adjusted to 7.0, 1 and 2
existed completely in a non-ionized form as determined by their
pKa values (Table 1), while 3-5 would be mostly dissociated
as carboxylate ions. Therefore, the ionized species were
considered to be taken up much less by duckweeds, which
indicated that the uptake is sensitive to the chemical structure.
The efficiency of translocation of a chemical to shoots from
root has been explained by Shone and Wood (19), Briggs et al.
(20, 21), and Hsu et al. (22), using a concept of TSCF
(transpiration stream concentration factor) defined by (concen-
tration in transpiration stream)/(concentration in exposure water).
Briggs et al. (20, 21) have suggested that there is an optimum
lipid/water distribution for translocation, more polar or more
lipophilic compounds being less well translocated when TSCF
is investigated. These papers describe measurements of TSCF
of barley and soybean plants hydroponically grown, using 18
and 12 non-ionized compounds, respectively, with a wide range
of lipophilicity as measured by their log P values. The
translocation from root to shoot versus log P plot showed a
Gaussian curve with a maximum at a log P of 1.8 for barley
(21) and a log P of 3.1 for soybean (22). If this relationship
can be applied to the uptake of a chemical from root to tallus
in duckweed, the pesticide metabolite with its log P value of
2-3 would have a great advantage in being taken up (high
values of TSCF). These TSCF concepts seem to account for
the uptake profiles of 1-5 in duckweed. Under the test condition
at pH 7.0, 1 and 2 were efficiently taken up as determined by
their log P values of non-ionized forms (2.48 and 2.69,
respectively). The pKa values of 3-5 (3.9-5.0) indicate that
they are almost completely ionized in the incubation medium
at pH 7.0. For the ionized species, the log D value instead of
log P is used for evaluation of its partitioning ability. Log D is
defined as the ratio of the equilibrium concentrations of all the
species (non-ionized and ionized) of a molecule in octanol to
that of the same species in the water phase at a given
temperature, usually 25 °C. Generally, the log D value of
For the purpose of isolating unknown metabolites for spectroscopic
analysis, plant extracts at the exaggerated concentration were subjected
to preparative purification by the solid-phase absorbent Waters Sep-
Pack C18 cartridge. First, the concentrated extracts were reconstituted
in 5 mL of water, and the mixture was loaded onto the Sep-Pack
cartridge. The cartridge was successively washed with 30 mL of water
followed by 30 mL of methanol, and the water/methanol wash was
repeated three times. From LSC and HPLC analysis, most of the
unknown metabolites were eluted in the methanol fraction. Further
purification was conducted with HPLC using method B.
RESULTS AND DISCUSSION
Recovery of Radioactivity. From the results of uptake
experiments, a good recovery of [14C]1-5 from the test system
was observed during the incubation for up to 1 day: 90.0%
(1), 119.8% (2), 92.0% (3), 94.4% (4), and 92.4% (5) of the
applied 14C. However, it appears that the recovery of radioactiv-
ity decreased as the exposure period was extended. The 14C
recovery of [14C]1-5 after 4 days dropped to 78.0% (1), 80.8%
(2), 76.1% (3), 80.7% (4), and 86.5% (5) of the applied 14C,
possibly due to azeotropic vaporization. The insignificant
differences in the loss of 14C were observed, although Henry’s
law constants of 1-5 estimated by EPI-Suite (17) greatly varied
by a factor of 500: 1.61 × 10-8 (1), 8.11 × 10-6 (2), 5.27 ×
10-8 (3), 3.81 × 10-7 (4), and 4.20 × 10-6 atm m3 mol-1 (5).
Uptake of Test Compounds into Plants. The radioactivity
taken up by duckweed was estimated by combining extractable
and unextractable 14C, the latter of which was found to be less
than <1% of the total radioactive residues (TRR) in any case.
The characteristics of uptake of radioactivity into duckweed are
shown in Figure 2. A significant uptake of 14C was observed
for 1 and 2 (60.3% and 70.1% of the applied 14C, respectively)
and was close to reaching a plateau after 4 days. In contrast,
much less but similar amounts of carboxylic derivatives 3-5
with very different chemical structures were shown to be taken
up by duckweed, reaching a plateau after 1 day (14.2-20.7%
of the applied 14C). Phenoxyacetic and benzoic acid derivatives
have been reported to be taken up by Lemna minor at pH 5.1,
where the acids were partly undissociated to an extent that was

Uptake and Transformation of Pesticide Metabolites by Duckweed
J. Agric. Food Chem., Vol. 54, No. 17, 2006 6289
Figure 3. LC−ESI−MS/MS spectrum of 9.
Figure 4. LC−ESI−MS spectrum of 8.
carboxylate ion has been reported to be ∼4.1 log units lower
than the log P value of the corresponding undissociated form
(23). Therefore, the fact that the log D values of 3-5 are likely
to be in the range of -0.7 to 0.5 implies less favorable uptake
of these compounds into duckweed, which is in accordance with
the observation.
Distribution of Metabolites. The formation and declines of
metabolites were examined for [14C]3 in detail through the

6290 J. Agric. Food Chem., Vol. 54, No. 17, 2006
Fujisawa et al.
Figure 5. 1H NMR spectrum of 8.
Figure 6.
H−H COSY spectrum of 8.
uptake experiment at 0.1 µg/mL. The product distribution is
summarized in Table 2. After reaching a plateau at day 1, the
amount of extractable fraction was almost constant during the
incubation time. The amount of 3 remained almost constant
around 30% of the extractable 14C with metabolites 8 and 10,
and the amount of the latter gradually decreased with time. On
the basis of this product distribution, the incubation period in
the study examining metabolic profiles of [14C]1-5 at 1 µg/
mL was taken to be 4 days after treatment of each chemical
where uptake reached a plateau. The chemical identification of
[14C]1-5 was confirmed by HPLC cochromatography with the
corresponding authentic standard. With regard to phenolic
compound 1, 1 and 6 were the major components of the
extractable 14C after 4 days, amounting to 20.4% and 64.9%
TRR, respectively. The remaining radioactivity (15% TRR)
consisted of minor peaks, each of which was less than 4% TRR.
Aniline derivative 2 was efficiently metabolized to 7 (51% TRR)
and 9 (24% TRR), and 2 remained unchanged as 25% TRR. In
the case of carboxylic acid derivatives 3-5, clear differences
were observed in the metabolic profiles. 3 existed as 29% TRR
after a 4 day incubation period with metabolites 8 and 10
amounting to 57% and 8% TRR, respectively (Table 2), while
4 and 5 were scarcely metabolized (<4% TRR) in duckweed.
Simultaneously, HPLC analysis of radioactivity in exposure
water showed that any degradation of [14C]1-5 did not occur
in the exposure water during the incubation period.
Identification of Metabolites. The chemical structure of
metabolite 6 from 1 was confirmed to be an O-glucoside
conjugate by HPLC cochromatography with the synthetic
standard. This type of conjugation is well-known for the various

Uptake and Transformation of Pesticide Metabolites by Duckweed
J. Agric. Food Chem., Vol. 54, No. 17, 2006 6291
Figure 7. LC−MS/MS spectrum of the natural component portion constituting 8.
Figure 8. LC−ESI−MS spectrum of 10.
types of phenols not only in terrestrial plants (24) but also in
aquatic plants (5). In the case of 2, N-glucoside 7 was formed
as a major metabolite. The chemical structure of 7 was also
confirmed by HPLC cochromatography with the synthetic
standard. As an additional metabolic pathway of 2, the conjuga-
tion with glutamic acid at the amino group to form metabolite
9 was confirmed. To examine the chemical identity of 9, we
first conducted LC-APCI-MS and LC-ESI-MS analyses in
positive and negative ion modes. Metabolite 9 was shown to
have a molecular weight of 291 ([M + H]+) detected only by
LC-ESI-MS in a positive ion mode. From an increase in the
molecular weight of 129, some natural component was consid-
ered to attach to 2 and the chemical structure of 9 was assumed
to be a glutamate conjugate of 2. To further investigate the
chemical nature of 9, it was subjected to LC-ESI-MS/MS
analysis in a positive ion mode, and a daughter ion pattern at
m/z 291 was investigated (Figure 3). The mass fragment profile
was totally consistent with the results reported by Mutlib et al.
(25) for the glutamate conjugate of 1-[3-(aminomethyl)phenyl]-
N-[3-fluoro-2-(methylsulfonyl)(1,1-biphenyl)-4-yl]-3-(trifluoro-

6292 J. Agric. Food Chem., Vol. 54, No. 17, 2006
Fujisawa et al.
and chromatographic evidence together with information about
conjugation forms of other pesticides (27-29), we proposed
the chemical structure of 8 to be the malate conjugate of 3.
Finally, the chemical identity of 8 was definitely confirmed by
both HPLC and TLC cochromatographies with the reference
standard synthesized in our laboratory. Several researchers
suggested the presence of malate conjugates in barley, plant
cell cultures, and rat (27-29) on the basis of mass spectrometry.
The study presented here gave the first concrete evidence of
the malate conjugate by the definite identification using HPLC
cochromatography with synthetic standards together with LC-
ESI-MS and NMR analyses.
The minor pathway was malonylglucoside conjugation which
has been reported for many phenol metabolites of pesticides
(24). For identification of 10, LC-APCI-MS and LC-ESI-
MS analyses in positive and negative ion modes were conducted.
The m/z value of 462 for 10 showed that the natural component
with a high molecular weight was linked to 3 as a conjugate
(Figure 8). The profiles of the daughter ions corresponding to
[M - COOH]- at m/z 417 and [M - malonylglucose - H]- at
m/z 213 were compared in detail with those of the phenol
metabolite from tolclofos-methyl (30). From these results, 10
was identified as the malonylglucose conjugate of 3.
Interestingly, although phase I metabolism such as oxidation
and reduction (3, 31) was not detected, 1-3 underwent a
different phase II metabolism which was possibly dependent
on their chemical structure and functional group. 1-3 were all
conjugated by glucose, but differences were observed in the
conjugation with small organic acid derivatives (Figure 9). In
contrast, simple acids 4 and 5 were scarcely transformed in
duckweed, and therefore, either reactivity or steric hindrance
at the carboxyl group may result in these differences versus 3.
This phenomenon will be an interesting research area to be
pursued.
Figure 9. Proposed metabolic pathways of 1−3.
methyl)-1H-pyrazole-5-carboxamide detected in rat, using LC-
MS/MS, 1H NMR, 13C NMR, and HMBC. In conclusion,
metabolite 9 was identified as the glutamate conjugate of 2.
This type of conjugation has been reported for phenoxyacetic
acid herbicides in the experiments using tissue culture (24) and
indole-3-acetic acid (26). Recently, Mitsou et al. (2) have
reported N-acetylation of 3,4-dichloroaniline in the metabolism
of propanil in L. minor, but the corresponding metabolite could
not be detected in our study. Either the substituted positions of
chlorine atoms in the phenyl ring or species differences may
yield the different metabolic patterns.
LITERATURE CITED
(1) Katagi, T. Behavior of pesticides in water-sediment systems. ReV.
EnViron. Contam. Toxicol. 2006, 187, 133-252.
Benzoic acid derivative 3 was found to undergo two metabolic
transformations. The major pathway was formation of metabolite
8 which was subjected to LC-APCI-MS and LC-ESI-MS
analyses in positive and negative ion modes. The molecular
weight of 8 isolated from the extracts was most likely to be
330, as demonstrated by the LC-ESI-MS peak in the negative
ion mode. The [M - H]- peak was observed at m/z 329 together
with the daughter ion corresponding to [M - malate - H]- at
m/z 213 (Figure 4). The 1H NMR spectrum of isolated 8 (Figure
5) showed the nine aromatic protons observed at 7.05-7.86
ppm very similar to those of 3, indicating that no structural
change occurred in the phenoxyphenyl moiety of 3. The methine
proton observed at 5.21 ppm (triplet) and methylene ones at
2.85 ppm (double doublet) were demonstrated to be correlated
by the H-H COSY spectrum (Figure 6), indicating the presence
of the CH-CH2 structure in 8. To investigate the chemical
identity attached to 8 in more detail, the corresponding daughter
ion to the natural component (115 amu) was subjected to LC-
ESI-MS/MS analysis in a negative ion mode. The decrease of
44 amu from the parent ion strongly suggested the existence of
a COOH group (Figure 7). Furthermore, the existence of a
carboxyl group was indirectly shown when 8 was analyzed with
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Received for review May 8, 2006. Revised manuscript received June
22, 2006. Accepted June 26, 2006.
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