Conversions of deoxyradicinin to radicinin and of radicinin to 3-epi-radicinin in the phytopathogenic fungus Bipolaris coicis

Article abstract of DOI:10.1016/j.phytochem.2011.11.010

Radicinin is a phytotoxic and antibiotic metabolite produced by some phytopathogenic fungi. Precursor administration and cell-free experiments with deoxyradicinin and radicinin were carried out in Bipolaris coicis H13-3. When deoxyradicinin was administered to the fungus, radicinin and 3-epi-radicinin formed. When radicinin administered, 3-epi-radicinin was formed. Their formation was confirmed by cell-free experiments. Deoxyradicinin 3-monooxygenase which catalyzes conversion of deoxyradicinin to radicinin showed the best activity at 35 °C and pH 7.0, and required NAD⁺ as co-enzyme. Its molecular weight was determined to be 130-184 kDa. Radicinin epimerase catalyzing the reaction of radicinin to 3-epi-radicinin was purified from a cell-free extract. Radicinin epimerase is a homodimer of a 28 kDa subunit, and its highest activity was achieved at 30-35 °C and pH 7.0-9.0.

Full text of DOI:10.1016/j.phytochem.2011.11.010

Phytochemistry 75 (2012) 14–20
Contents lists available at SciVerse ScienceDirect
Phytochemistry
journal homepage: www.elsevier.com/locate/phytochem
Conversions of deoxyradicinin to radicinin and of radicinin to 3-epi-radicinin
in the phytopathogenic fungus Bipolaris coicis
Masanobu Suzuki a, Emi Sakuno a, Atsushi Ishihara a, Jun-ichi Tamura b, Hiromitsu Nakajima a,
⇑
a Department of Agricultural Chemistry, Faculty of Agriculture, Tottori University Koyama, Tottori 680-8553, Japan
b Department of Environmental Sciences, Faculty of Education and Regional Sciences, Tottori University, Koyama, Tottori 680-8551, Japan
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 19 April 2011
Received in revised form 5 November 2011
Available online 16 December 2011
Radicinin is a phytotoxic and antibiotic metabolite produced by some phytopathogenic fungi. Precursor
administration and cell-free experiments with deoxyradicinin and radicinin were carried out in Bipolaris
coicis H13-3. When deoxyradicinin was administered to the fungus, radicinin and 3-epi-radicinin formed.
When radicinin administered, 3-epi-radicinin was formed. Their formation was confirmed by cell-free
experiments. Deoxyradicinin 3-monooxygenase which catalyzes conversion of deoxyradicinin to radici-
nin showed the best activity at 35 °C and pH 7.0, and required NAD+ as co-enzyme. Its molecular weight
was determined to be 130–184 kDa. Radicinin epimerase catalyzing the reaction of radicinin to 3-epi-rad-
icinin was purified from a cell-free extract. Radicinin epimerase is a homodimer of a 28 kDa subunit, and
its highest activity was achieved at 30–35 °C and pH 7.0–9.0.
Keywords:
Radicinin
Deoxyradicinin
3-epi-Radicinin
Bipolaris coicis
Conversion
Biosynthesis
Monooxygenase
Epimerase
Ó 2011 Elsevier Ltd. All rights reserved.
Fungus
1. Introduction
experiment with its 4-O-p-bromobenzoyl ester by Robeson et al.
(1982). The b-oxygenated
a-pyrone moiety of radicinin (3) is not
Radicinin (3) (Fig. 1) is a phytotoxic and antibiotic metabolite
produced by some phytopathogenic fungi. It was first isolated from
Stemphylium radicinum in the 1950s (Clarke and Nord, 1953), and
since then has been reported to be produced by several fungal spe-
cies, Cochliobolus lunatus (Nukina and Marumo, 1977), Alternaria
chrysanthemi (Robeson and Strobel, 1982), Alternaria helianthi
(Tal et al., 1985), Phoma andina (Noordeloos et al., 1993), Curvularia
sp. (Kadam et al., 1994), Alternaria radicina (Pryor and Gilbertson,
2002) and Alternaria petroselini (Pryor and Gilbertson, 2002). It
shows phytotoxicity, for example, killing roots of Lepidium sativum
(Hansen, 1954), browning and causing a loss of viability in Nicoti-
ana tabacum (Canning et al., 1992), as well as producing necrotic
lesions in Coix lachryma-jobi (Nakajima et al., 1997) and inhibiting
root growth in carrot seedlings (Solfrizzo et al., 2004).
The structure of radicinin (3) except for its stereochemistry, was
determined by Grove (1964) on the basis of chemical and spectro-
scopic evidence. Its absolute stereochemistry was inferred from the
CD spectrum of the 3,4-bis-O-p-chlorobenzoyl derivative of radicin-
ol by Nukina and Marumo (1977) based on the dibenzoate exciton
chirality rule, and this was supported by an X-ray crystallographic
unusual among natural products. Its biosynthesis has been studied
by some researchers. Radioactive tracer experiments by Grove
(1970) demonstrated that it is synthesized from two different
polyketide chains originating from acetate and malonate. Other re-
search groups confirmed this using 13C labeled compounds (Tanabe
et al., 1970; Seto and Urano, 1975). Although it has been assumed
that the direct precursor of radicinin (3) is deoxyradicinin (1), there
has been no experimental data to support this until now. Formation
of deoxyradicinin (1) through an uncommon condensation of two
polyketide chains, cyclization and ring-cleavage was demonstrated
by incorporation studies with 13C-labeled acetates and a 2H-labeled
one (Tal et al., 1988).
The Bipolaris coicis H13-3 used in this study is a plant pathogen
causing serious leaf blight on Job’s tears (Coix lachryma-jobi L.), and
it was reported that radicinin (3), 3-epi-radicinin (4), 3-epi-radicin-
ol (5) and its epoxide (6) were produced by this fungus (Nakajima
et al., 1997). Their structures suggested a biosynthetic relationship
between these metabolites shown in Fig. 1. In this scheme, both
radicinin (3) and 3-epi-radicinin (4) are synthesized from
deoxyradicinin (1), which was isolated from the plant pathogen
A. helianthi together with radicinin (3) (Robeson and Strobel,
1982); 3-epi-radicinin (4) is then reduced to 3-epi-radicinol (5),
which is oxidized to 3-epi-radicinol epoxide (6).
⇑
Corresponding author. Tel./fax: +81 857 31 5342.
E-mail address: nakajima@muses.tottori-u.ac.jp (H. Nakajima).
0031-9422/$ - see front matter Ó 2011 Elsevier Ltd. All rights reserved.
doi:10.1016/j.phytochem.2011.11.010

M. Suzuki et al. / Phytochemistry 75 (2012) 14–20
15
Fig. 1. Proposed processes for formation of radicinin (3) and 3-epi-radicinin (4) from deoxyradicinin (1) and transformation of 3-epi-radicinin (4) to metabolites (5 and 6) by
Bipolaris coicis H-13-3.
In this work, the hypothesis shown in Fig. 1 was examined by a
precursor administration experiment and a cell-free approach with
deoxyradicinin (1) and radicinin (3). The former was not commer-
cially available, and a sufficient amount of deoxyradicinin (1) could
not be obtained from B. coicis H13-3. We thus synthesized deoxy-
radicinin (1) according to the reported methods (Kato et al., 1969;
Suzuki et al., 1975).
the culture filtrate of the fungus B. coicis H13-3 grown on malt ex-
tract medium. Its optical purity was confirmed by analysis of the
NMR spectrum of its (ꢀ)-MTPA ester, in which no resonance due
to the (ꢀ)-MTPA ester of the enantiomer was detected. Thus, it
was concluded that radicinin (3) isolated from the fungus was opti-
cally pure.
Compounds 1 and 2 were administered to the fungus sepa-
rately, and the conversion products were analyzed by HPLC. As
shown in Fig. 3B, the amount of radicinin (3) (27.6 nmol/l) de-
tected, when compound 1 was administered, was about eight
times more than that of the control (3.4 nmol/l). Additionally, the
amount of 3-epi-radicinin (4) also increased as compared to the
control. There are two possible explanations for the increase in
the amount of 3-epi-radicinin (4). One is that (ꢀ)-deoxyradicinin
(1) was hydroxylated to be 3-epi-radicinin (4) directly and the
other is that radicinin (3) produced from (ꢀ)-deoxyradicinin (1)
was epimerized at C-3 to be 3-epi-radicinin (4). By contrast, admin-
istration of (+)-deoxyradicinin (2) caused no significant increase in
either the amounts of radicinin (3) or of 3-epi-radicinin (4)
compared with the control (Fig. 3C). This supports the fact that
compound 1 has the same stereochemistry at C-2 as radicinin (3)
produced by B. coicis, but compound 2 does not. To examine the
biogenetical origin of 3-epi-radicinin (4), radicinin (3) was admin-
istered to the fungus, and the conversion products were analyzed
by HPLC. As shown in Fig. 4, when radicinin (3) was administered,
there was about a 4-fold increase in the amount of 3-epi-radicinin
(4) (3.5 nmol/l) detected by HPLC compared with the control
(0.9 nmol/l). These results indicated that (ꢀ)-deoxyradicinin (1),
not (+)-deoxyradicinin (2), is a direct precursor of radicinin (3)
and also that the fungus have an epimerizing enzyme which
catalyzes the conversion of radicinin (3) to 3-epi-radicinin (4).
2. Results and discussion
2.1. Feeding experiment
To confirm the conversion of deoxyradicinin (1) to radicinin (3)
and the latter to 3-epi-radicinin (4), deoxyradicinin (1) and radicinin
(3) were administered to the fungus separately. The deoxyradicinin
(1) was synthesized from 4-methoxy-6-methyl-2H-pyran-2-one
(7) according to the reported literature (Kato et al., 1969; Suzuki
et al., 1975), and the synthetic scheme is shown in Fig. 2. Since
the final product was a mixture of deoxyradicinin (1) and its C-2
epimer (2), they were separated via chiral HPLC. The overall yield
of compounds 1 and 2 from compound 7 was 1.2% and 1.5%,
respectively. To determine which compound has the same stereo-
chemistry as the natural product, the optical rotation of each com-
pound was measured as ꢀ82° for compound 1 and +90° for
compound 2. To our knowledge, optical rotation of deoxyradicinin
(1) isolated from the filamentous fungus has not been reported,
and we therefore compared the optical rotation of the synthetic
compounds with [
a
]
ꢀ125° of radicinin (3) (Hansen, 1954), [
a
a
]
]
D
D
ꢀ105° of 3-epi-radicinin (4) (Nakajima et al., 1997) and [
D
ꢀ19° of 3-epi-radicinol (5) (Nakajima et al., 1997). From its nega-
tive optical rotation, (ꢀ)-deoxyradicinin (1) has the same stereo-
chemistry as radicinin (3) produced by this fungus. The optical
purity of compounds 1 and 2 was determined to be 94.4% ee and
96.6% ee, respectively, using chiral HPLC. The radicinin (3) used
in the precursor administration experiment was isolated from
2.2. Enzyme activity
To confirm the conversion of deoxyradicinin (1) to radicinin (3)
as indicated by precursor administration, the experiment using a
Fig. 2. Synthesis of deoxyradicinin (1 and 2). Reagents and conditions: (i) SeO2, dioxane, 160 °C; (ii) ethyl triphenyl phosphonium bromide, sodium bis(trimethylsilyl) amide,
DMF; (iii) TiCl4, crotonoyl chloride, CH2Cl2; (iv) TiCl4, CH2Cl2.

16
M. Suzuki et al. / Phytochemistry 75 (2012) 14–20
Fig. 5. Deoxyradicinin monooxygenase activity to convert (ꢀ)-deoxyradicinin (1) to
radicinin (3) in the cell-free extract, cytosolic and microsomal fractions. Data
presented is mean of three replicates and SD.
Fig. 3. HPLC profiles of products converted from deoxyradicinin by Bipolaris coicis
H13-3. (A) control, (B) (ꢀ)-deoxyradicinin (1), (C), (+)-deoxyradicinin (2).
enzyme activity was determined by comparing the reaction rates
at pH 4–9 at 35 °C. SD1 demonstrates that the reaction was cata-
lyzed most effectively by the monooxygenase at 35 °C, pH 7.0.
The monooxygenase prefers NAD+ to other co-enzymes (Fig. 6).
The molecular weight of the monooxygenase was determined to
be 130–184 kDa by gel filtration column chromatography.
Although we suggested in a previous paper (Nakajima et al.,
1997) that the enzyme catalyzing this reaction was cytochrome
P450 monooxygenase, the enzyme activity for the conversion of
deoxyradicinin (1) to radicinin (3) was distributed in the cytosolic
fraction and not in the microsomal fraction. The monooxygenase is
therefore, a soluble protein present in the cytoplasm. The enzyme
that catalyzes the hydroxylation like this was classified as oxidore-
ductase, and is activated in the presence of NAD+. Thus deoxyrad-
icinin 3-monooxygenase belongs to a monooxygenase group such
as EC 1.14.13.
To investigate the metabolism of radicinin (3) and the origin of
3-epi-radicinin (4), radicinin (3) was incubated with the cytosolic
or microsomal fraction in the presence of co-enzymes (NAD+,
NADP+, NADH and NADPH) for 30 min (Fig. 7). Incubation of radic-
inin (3) with the cytosolic fraction caused an increase in 3-epi-
radicinin (4), but no increase of 3-epi-radicinin (4) was observed
when incubating with the microsomal fraction. No 3-epi-radicinin
(4) was detected when incubating radicinin (3) with sterile water
in place of the cytosolic or the microsomal fraction, indicating that
radicinin (3) does not racemize at detectable rate without enzyme.
The radicinin epimerase that catalyzes the reaction of radicinin to
3-epi-radicinin (4) was purified with ammonium sulfate fraction-
ation and several chromatographic processes by monitoring
enzyme activity (Table 1). To characterize the epimerase, radicinin
(3) was incubated with this purified enzyme under several condi-
tions, demonstrating that the highest activity of the epimerase
Fig. 4. HPLC profiles of the products converted from radicinin by Bipolaris coicis
H13-3. (A) control, (B) radicinin (3).
cell-free system prepared from B. coicis H13-3 was carried out as
follows. Incubation of (ꢀ)-deoxyradicinin (1) with the crude cell
free extract, in the presence of co-enzymes (NAD+, NADP+, NADH
and NADPH 20 mM respectively), for 2 h gave rise to the enzymatic
formation of radicinin (3) (Fig. 5). Next, the crude cell free extract
was divided into cytosolic and microsomal fractions. When
(ꢀ)-deoxyradicinin (1) was incubated with the cytosolic fraction
in the presence of co-enzymes (NAD+, NADP+, NADH and NADPH),
the amount of radicinin (3) after 2-h incubation increased remark-
ably, but when incubated with the microsomal fraction, no signif-
icant formation of radicinin (3) was detected. (+)-Deoxyradicinin
(2) was also incubated with the cytosolic fraction or the micro-
somal fraction in the presence of co-enzymes (NAD+, NADP+, NADH
and NADPH), but, in both cases, no remarkable increases in the
amount of radicinin (3) in the 2-h incubation extracts were ob-
served. Deoxyradicinin 3-monooxygenase activity was measured
at various conditions with the cytosolic fractions. The optimum
temperature for the enzyme activity was determined by comparing
the reaction rates at 25–45 °C at pH 7.0. The optimum pH for the
Fig. 6. Effect of addition of co-enzyme to the enzyme assay solution on formation of
radicinin (3) from (ꢀ)-deoxyradicinin (1). The cytosolic fraction was used at 35 °C,
pH 7.0. Data presented is the mean of three replicates and SD.

M. Suzuki et al. / Phytochemistry 75 (2012) 14–20
17
substrates (Mazumder et al., 1962; Dahm et al., 1968; van der Drift
et al., 1975; Schmitz et al., 1994). However, no epimerization en-
zymes involved in secondary metabolism have been found until
now. The epimerase did not need any co-enzymes for the reaction,
indicating that the reaction proceeds through keto-enol tautomer-
ization. Thus, radicinin epimerase should belong to the tautomerase
group such as the EC number of 5.3.2. The toxicity of radicinin (3) for
Coix lachryma-jobi L. was ten times higher than 3-epi-radicinin (4)
(Nakajima et al., 1997), and hence it was assumed that radicinin epi-
merase regulated the pathogenicity of the fungus to the plants.
3. Conclusions
Fig. 7. Radicinin epimerase activity to epimerize radicinin (3) to 3-epi-radicinin (4)
in the cytosolic and microsomal fractions. Data presented is the mean of three
replicates and SD.
In our previous report (Nakajima et al., 1997), a biosynthetic
relationship was proposed between radicinin (3) and its analogues
based on their structural features. The present result obtained from
the feeding and cell-free experiments indicates that radicinin (3) is
synthesized from an anticipated precursor, deoxyradicinin (1). Fur-
thermore, a small amount of deoxyradicinin (1) was produced by
B. coicis, suggesting that the deoxyradicinin (1) biosynthesized is
rapidly converted to radicinin (3) or to the following biosynthetic
product, and thus ostensible amount of deoxyradicinin (1) is extre-
mely small in amount at any time. Previously, we proposed that
the same enzyme catalyzes the conversions of deoxyradicinin (1)
to radicinin (3) and also to 3-epi-radicinin (4). Actually, the precur-
sor administration experiment with deoxyradicinin (1) showed
formation of not only radicinin (3), but also 3-epi-radicinin (4).
At the same time, however, we established that the enzyme in
the cytosolic preparation from B. coicis H13-3 catalyzes reaction
of radicinin (3) to 3-epi-radicinin (4). This epimerization was
reversible and a new metabolic fate of radicinin (3). From these re-
sults, a biosynthesis and metabolism scheme for radicinin (3) was
deduced, as shown in Fig. 8. First, deoxyradicinin (1) is converted
to radicinin (3) by stereospecific hydroxylation at C-3. Then, radic-
inin epimerase catalyzes epimerization of radicinin (3) at C-3 to
3-epi-radicinin (4) reversibly. The direct conversion of deoxyradic-
inin (1) to 3-epi-radicinin (4) could not be confirmed in this study.
Finally, 3-epi-radicinin is probably converted to 3-epi-radicinol (5)
by stereospecific reduction at C-4, followed by epoxidation of the
side chain in 3-epi-radicinol (5). In this study, the key enzyme that
catalyzes the reaction of deoxyradicinin (1) to radicinin (3) could
not be isolated, probably because of enzyme instability.
Table 1
Purification of radicinin epimerase from Bipolaris coicis H 13-3. Total protein was
measured by Bradford protein assay. Total activity was calculated from the reaction
product monitoring by HPLC.
Purification
step
Total
protein
(mg)
Total
Specific
activity
(nmol/min/
mg)
Purification Recovery
activity
(nmol/
min)
fold
(%)
Cytosol
DE-52
Phenyl
262
40.6
13.9
1035
230
140
4
5.7
10.1
1
1.4
2.6
100
22.3
14
Sepharose
Superdex
1st MonoQ
2nd MonoQ
2.12
0.22
0.04
38.7
13.7
9.1
18.3
62.3
228
4.5
15.8
57.7
3.7
1.3
0.9
was found at 30–35 °C and pH 7.0–9.0 (SD2), and that the epimerase
did not require any co-enzyme for this conversion. The molecular
weight of the epimerase was determined to be 52 kDa based on its
gel filtration chromatographic behavior. The fractions from the
2nd Mono Q HR 5/5 column chromatography were characterized
by SDS–PAGE analysis and enzyme assay. A major band correspond-
ing to 28-kDa on SDS–PAGE is in accordance with the enzyme activ-
ity, indicating that the epimerase is homodimeric in its native
condition (SD3). To ascertain whether the epimerase catalyzes the
inverse reaction, the isolated epimerase was incubated with 3-epi-
radicinin (4). HPLC analysis of the products indicated formation of
radicinin (3) from 3-epi-radicinin (4) (data not shown), demonstrat-
ing that this reaction was reversible. The enzyme activity was
inhibited by copper sulfate and iodoacetic acid (data not shown),
suggesting that radicinin epimerase is a SH enzyme. Most epimer-
ization enzymes that have been discovered so far utilize carbohy-
drates, amino acids, hydroxyl acids and their derivatives as
4. Experimental
4.1. Experimental procedures
NMR spectra were recorded in CDCl3 on a JEOL JNM-ECP 500
spectrometer. NMR chemical shifts were referenced to CDCl3 (dH
Fig. 8. Proposel biosynthesis of radicinin (3) from (–)-deoxyradicinin (1) and conversion of radicinin to 3-epi-radicinin (4). Solid arrows show the pathways established in this
research, and dashed arrows show the unproven pathways.

18
M. Suzuki et al. / Phytochemistry 75 (2012) 14–20
7.26, dC 77.0). Mass spectra were obtained with a JEOL AX-505
spectrometer. Optical rotations were determined with a Horiba
SEPA-200 high sensitive polarimeter. A Shimadzu LC-6A liquid
chromatography system was used for HPLC analysis. In the precur-
sor administration experiment and the enzyme assay for the con-
version of deoxyradicinin (1) to radicinin (3), HPLC was
performed using a DAISOPAK SP-120-5-ODS-AP column (DAISO
Co., Ltd., 150 ꢁ 6 mm), MeOH–H2O–AcOH (100:99:1, v/v/v) as
solvent at a flow rate of 0.5 ml/min, monitoring at 280 nm. In the
enzyme assay for the conversion of radicinin (3) to 3-epi-radicinin
(4), HPLC was carried out using a DAISOPAK SP-250-10-ODS-AP
column (DAISO Co., Ltd., 250 ꢁ 10 mm), MeOH–H2O–AcOH
(100:99:1, v/v/v) as solvent at a flow rate of 0.5 ml/min, and mon-
itoring at 280 nm. Silica gel flash column chromatography (CC) was
carried out by use of Wakogel FC-40 (Wako Pure Chemical Indus-
tries, Ltd.). Protein concentrations were determined with Bradford
reagent (Sigma–Aldrich) using BSA as a standard.
4.4.2. 4-Methoxy-6-[(E)-1-propenyl]-2H-pyran-2-one (9)
Ethyltriphenylphosphonium bromide (1.2 g, Wako Pure Chemi-
cal Industries, Ltd.) was dissolved into DMF (11 ml), and then
bis(trimethylsilyl) amide (3 ml, Sigma–Aldrich, Inc.) was added.
Compound 8 (120 mg) in DMF (4 ml) was added and the mixture
was stirred vigorously at r.t. for 3 h under N2. The reaction mixture
was poured into brine (70 ml) and extracted with EtOAc
(70 ml ꢁ 3). The combined EtOAc solubles were dried (Na2SO4)
and evaporated in vacuum to dryness. The residue was subjected
to Si gel flash CC (150 ꢁ 20 mm), which was developed succes-
sively with 200 ml each of Me2CO–n-hexane (5:95,1:9,2:8 and
3:7,v/v), respectively. Compound 9 and its Z-isomer were eluted
with Me2CO–n-hexane (1:9 and 1:4, v/v) fractions (61 mg). Heating
of the crystalline mixture at 130 °C in a sealed tube under N2
caused isomerization to give 57 mg of compound 9: dH 6.68 (1H,
dq, J = 15.5, 7.0 Hz, 8-H) 5.97 (1H, dq, J = 15.5, 1.5 Hz, 7-H), 5.74
(1H, d, J = 2.2 Hz, 5-H), 5.43 (1H, d, J = 2.2 Hz, 3-H), 3.80 (3H, s,
10-H), 1.88 (3H, dd, J = 7.0, 1.5 Hz, 9-H); EI-MS, 70 eV, m/z 166
(M+, 63%), 138 (100), and 69 (38).
4.2. Fungal strain
The strain H13-3 of B. coicis was used in the experiments (Nak-
ajima et al., 1997). B. coicis H13-3 was maintained on potato dex-
trose agar slants.
4.4.3. 3-[(E)-2-butenoyl]-4-methoxy-6-[(E)-1-propenyl]-2H-pyran-2-
one (10)
To a mixture of compound 9 (80 mg) and TiCl4 (260
Pure Chemical Industries, Ltd.) in CH2Cl2 (1 ml), crotonoyl chloride
(70 l, Wako Pure Chemical Industries, Ltd.) was added slowly. The
ll, Wako
4.3. Isolation of radicinin
l
solution was stirred at r.t. for 20 min, and then at 45 °C for 6 h. It
was then poured into brine (30 ml), and extracted with EtOAc
(30 ml x 3). The EtOAc solubles were combined, dried over Na2SO4,
and evaporated to dryness. The resulting residue was purified by Si
gel flash CC (150 ꢁ 20 mm) developed successively with 100 ml
each of Me2CO in n-hexane (1:4,1:3,3:7 and 35:65 v/v). Compound
10 (40 mg) was eluted with Me2CO–n-hexane (1:4, v/v) whereas
compounds 1 and 2 were eluted using Me2CO–n-hexane (3:7, v/
v). Separation of compounds 1 and 2 by chiral HPLC used a CHIR-
ALPAK OD column (Daicel Chemical Industries, Ltd.,
4.6 ꢁ 250 mm), eluted with EtOH-n-hexane (1:1, v/v) at a flow rate
of 0.5 ml/min, and monitoring at 280 nm to afford compounds 1
(2.5 mg) and 2 (3.0 mg). The retention times of compounds 1 and
2 were 26.2 and 30.8 min, respectively. Compound 10: dH 1.85
(3H, dd, J = 7.0, 1.4 Hz, 9-H), 1.87 (3H, dd, J = 7.0, 1.4 Hz, 140-H),
3.82 (3H, s, 10-H), 5.94 (1H, s, 5-H), 5.97 (1H, dq, J = 15.5, 1.4 Hz,
7-H), 6.40 (1H, dq, J = 15.5, 1.4 Hz, 12-H), 6.82 (2H, m, 8, 13-H);
EI-MS, 70 eV, m/z 234 (M+, 100%), 193 (80), 165 (48), and 138 (32).
The fungus was grown without shaking at 24 °C for 14 days in
the dark in a 500 ml conical flask containing liquid medium
(200 ml ꢁ 5) made up of glucose (30 g/l), peptone (3 g/l) and an ex-
tract from 50 g/l of malt and H2O. The culture filtrate was acidified
to pH 2.0 with HCl, and the metabolites in the culture filtrate were
extracted with EtOAc (500 ml ꢁ 3). The EtOAc extract was dried
over Na2SO4 and evaporated. The residue (270 mg) was applied
to
a silica gel column (Daisogel IR-60, DAISO, Co., Ltd.,
180 ꢁ 18 mm), and the column was washed with 1500 ml of
Me2CO–n-hexane (1:9,v/v), then developed successively with
750 ml each of Me2CO–n-hexane (2:8, 3:7 and 4:6, v/v). Fractions
3 and 4 eluted by Me2CO–n-hexane (3:7, v/v) were combined
and evaporated. Recrystallization of the residue (34 mg) from
MeOH afforded radicinin (3) as colorless needles (6 mg): dH 6.93
(1H, dq, J = 15.5, 7.0 Hz, 20-H), 6.02 (1H, dq, J = 15.5, 1.7 Hz, 10-H),
5.83 (1H, s, 8-H), 4.36 (1H, dq, J = 12.4, 6.0 Hz, 2-H), 3.97 (1H, d,
J = 12.4 Hz, 3-H), 1.96 (3H, dd, J = 7.0, 1.7 Hz, 3’-H), 1.64 (3H, d,
J = 6.0 Hz, 2-Me); dC (CDCl3) 188.7, 176.0, 164.4, 156.8, 141.0,
122.6, 98.1, 97.2, 80.0, 72.0, 18.8, 18.1, CI-MS (iso-butane, probe),
200 eV, m/z 237 ([M+H]+, 100%).
4.4.4. Deoxyradicinin (1)
A mixture of compound 10 (40 mg), TiCl4 (90 ll) and CH2Cl2
(1 ml) was stirred at 45 °C for 3 h. It was then poured into brine
(30 ml), extracted with EtOAc (30 ml ꢁ 3). The organic layer was
washed with brine (100 ml ꢁ 3). The combined EtOAc extracts
were dried (Na2SO4) and evaporated to dryness, with the residue
was subjected to Si gel flash CC (150 ꢁ 20 mm) developed succes-
sively with 100 ml each of Me2CO-n-hexane (1:4,1:3,3:7 and
35:65,v/v). Compounds 1 and 2 were eluted with Me2CO-n-hexane
(3:7, v/v), and separated by chiral HPLC as above to afford com-
pounds 1 (1.8 mg) and 2 (2.5 mg). Compound 1, 2: dH 6.86 (1H,
m, 20-H), 5.95 (1H, dq, J = 15.5, 1.8 Hz, 10-H), 5.76 (1H, s, 8-H),
4.67 (1H, m, 2-H), 2.59 (2H, m, 3–H), 1.88 (3H, dd, J = 7.3, 1.4 Hz,
30-H), 1.46 (3H, d, J = 5.6 Hz, 2-Me); dC 186.2, 176.6, 163.8, 156.7,
138.9, 124.0, 100.7, 99.1, 77.6, 44.3, 20.4, 18.6; EI-MS, 70 eV, m/z
4.4. Synthesis of deoxyradicinin
4.4.1. 6-Formyl-4-methoxy-2H-pyran-2-one (8)
The mixture of 4-methoxy-6-methyl-2H-pyran-2-one (7;
500 mg, Sigma–Aldrich, Inc.), SeO2 (1.2 g, Kanto Chemical Co.,
Inc.) and anhydrous dioxane (5 ml) in a sealed tube was heated
at 160 °C (outside) and stirred vigorously. After 3 h, the precipitate
was removed by filtration and washed with dioxane. The latter was
removed by evaporation in vacuo to afford a residue, which was
poured into brine (100 ml). The resulting product was extracted
with EtOAc (100 ml ꢁ 3) with the EtOAc solubles combined. After
drying over Na2SO4, the extract was evaporated to dryness. The
residue was purified by Si gel flash CC. The column
(150 ꢁ 20 mm) was developed successively with 300 ml of
Me2CO–n-hexane (3:7, 4:6 and 1:1 v/v), respectively. Compound
8 (362 mg) was eluted with Me2CO–n-hexane (4:6, v/v): dH 9.48
(1H, s, 7-H), 6.63 (1H, d, J = 2.3 Hz, 5-H), 5.70 (1H, d, J = 2.3 Hz, 3-
H), 3.82 (3H, s, 8-H); EI-MS, 70 eV, m/z 154 (M+, 22%), 125 (100),
69 (20), and 59 (15).
220 (M+, 100%), 205 (50), and 177 (59). Compound 1: [
0.1, CHCl3). Compound 2: [
ꢀ82° (c 0.1, CHCl3).
a] +90°(c
D
a]
D
4.5. Precursor administration
The fungus was grown on medium (500
malt extract broth (Difco Laboratories Inc.) in
l
l) containing 15 g/l of
test tube
a

M. Suzuki et al. / Phytochemistry 75 (2012) 14–20
19
(75 ꢁ 8 mm) without shaking at 24 °C for 7 days in the dark. Then
the medium was removed aseptically from the tube with fungal
mats washed with 100 mM K-Pi buffer, pH 7.0. The fungus was
incubated in 500 ll of the same buffer at 24 °C for 3 days in the
dark. After incubation, the buffer was removed from the tube asep-
tically and the fungal mats were washed with buffer. Then, new
buffer (500 ll) was introduced into the tube and DMSO (50 ll)
containing precursor (0.05 mg) was added to the buffer. After
5 days incubation, the fungal mat was removed and the remaining
The column was washed with 80 ml of buffer, followed by a linear
gradient elution of 0–0.5 M NaCl in the buffer, at a flow rate of
0.6 ml/min, and each 3.0 ml was collected as one fraction. Active
fractions eluted between 0.19 and 0.31 M NaCl were combined,
and an equivalent amount of buffer containing 1.6 M (NH4)2SO4
was added. The solution was loaded onto a Phenyl Sepharose CL-
4B (Sigma–Aldrich) column (90 ꢁ 9 mm). The column was washed
with 0.8 M (NH4)2SO4 buffer (80 ml), followed by a linear gradient
elution of 0.8–0 M (NH4)2SO4 in the buffer, at a flow rate of 0.6 ml/
min, and each 2.0 ml fraction was collected. Active fractions eluting
between 0.56 and 0.32 M were combined, loaded onto a Superdex
200 10/300 GL (GE Healthcare) column, eluted with 20 mM Tris–
HCl buffer (pH 8.0) at a flow rate of 0.6 ml/min, and each 0.5 ml
was collected as one fraction. Active fractions were combined and
loaded onto a Mono Q HR 5/5 (Amersham) column (100 ꢁ 38 cm)
equilibrated with 20 mM Tris–HCl buffer (pH 8.0). The column
was washed with 80 ml of the same buffer, followed by a linear gra-
dient elution of 0–0.4 M NaCl in the buffer, at a flow rate of 0.5 ml/
min, and each 0.3 ml was collected as one fraction. The active frac-
tions eluted between 0.18 and 0.23 M NaCl were combined, and
the same purification with a Mono Q HR 5/5 (Amersham) column
was repeated. Radicinin epimerase was eluted at around 0.21 M
NaCl (see Table 1 and SD3B).
buffer was extracted with EtOAc (100
l
l ꢁ 3). The EtOAc extracts
were combined, air-dried overnight, redissolved in MeOH and ana-
lyzed by HPLC as described in Section 4.1. Retention times of com-
pounds 1, 3 and 4 were 15.8, 13.5 and 11.8 min, respectively.
4.6. Preparation of cell-free extract
The grown mycelia were homogenized with sea sand, a mortar
and pestle in 50 mM K-Pi buffer, pH 7.0 at 4 °C. The homogenate
was centrifuged at 2500g for 10 min at 4 °C. The supernatant was
centrifuged at 19,000g for 15 min at 4 °C to yield the cell-free ex-
tract. The extract thus obtained was then subjected to ultracentri-
fugation at 30,000g for 180 min at 4 °C to afford cytosolic and
microsomal fractions. The microsomal precipitates were sus-
pended in 50 mM K-Pi buffer, pH 7.0, and used for the experiments.
Glycerol was added to each fraction up to 15% (v/v), and fractions
were maintained at ꢀ80 °C until use.
4.9. Molecular weights of deoxyradicinin monooxygenase and
radicinin epimerase
4.7. Enzyme activity
To determine the molecular weights of monooxygenase and
epimerase, Superdex 200 10/300 GL CC as described above was
performed with the standard proteins, thyroglobulin (670 kDa),
gamma globulin (158 kDa), ovalbumin (44 kDa), myoglobin
(17 kDa) and vitamin B-12 (1.35 kDa).
The following method was used to detect deoxyradicinin mono-
oxygenase or radicinin epimerase activity in the cell extract, cyto-
solic and microsomal fractions. The crude enzyme preparation
(10
20 mM substrate in MeOH, 3
20 mM each of NAD+, NADP+, NADH and NADPH), 35
K-Pi buffer, pH 7.0. During purification of epimerase, 3
was used in place of 3 l of co-factor solution in the enzyme assay.
After incubation, the reaction mixture was extracted with EtOAc
(100
l ꢁ 3). The combined EtOAc solution was air-dried overnight,
l
l) was incubated at 35 °C for 2 h or 30 min with 2
l of co-factor solution (containing
l of 70 mM
l of buffer
ll of
l
4.10. SDS–PAGE
l
l
After reduction with 2-mercaptoethanol, the relative molecular
mass of the purified enzyme was determined by sodium dodecyl
sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) on 10%
polyacrylamide gels at 75 mA per slab with Tris–glycine, pH 8.3,
using 0.1% SDS as running buffer. Coomassie brilliant blue stain
solution (CBB R-25 1 g, MeOH 100 ml, AcOH 30 ml, in D.W.
400 ml) was used to stain the enzyme. After decolorization, the
gel was stained with a silver staining kit (Silver Staining II kit,
Wako Pure Chemical Industries, Ltd.). LMW Marker Kit (GE Health-
care) was used as molecular marker (SD3A).
l
l
dissolved in the MeOH, and analyzed by HPLC described in Sec-
tion 4.1. In the HPLC analysis for the conversion of deoxyradicinin
(1) to radicinin (3), their retention times were 15.8 and 13.5 min
and in the HPLC analysis for conversion of radicinin (3) to 3-epi-
radicinin (4), those were 28.5 and 24.0 min. The crude enzyme
solution usually contained radicinin (3) and 3-epi-radicinin (4).
Thus EtOAc was added first to the enzyme cocktail prior to sub-
strate and then substrate was added. The mixture was extracted
with EtOAc without incubation. The EtOAc extract was analyzed
by HPLC to afford the initial amount of the products in the enzyme
solution. The true amount of the product formed by the enzyme
preparation was obtained by subtracting initial amount from
amount after enzyme reaction. To determine the effect of pH on
enzyme activity and stability, 0.2 M NaOAc buffer was used for
pH 4.0 to 5.0, 0.2 M K-Pi buffer for pH 6.0 to 8.0 and 0.2 M Tris–
HCl buffer for pH 7.0 to 9.5. One milli molar of phenylmethylsul-
fonyl fluoride in 50 mM K-Pi buffer (pH 7.0) was used for the opti-
mum temperature determination of the monooxygenase enzyme
(SD1), and 0.4 mM EDTA, 10 mM MgCl2, 10% glycerol in 20 mM
Tris–HCl buffer (pH 8.0) was used for the optimum temperature
determination of the epimerase enzyme (SD2).
Acknowledgments
We would like to thank Ms. N. Inoue and Mr. S. Matsunaga for
technical assistance. This work was supported by a Global COE Pro-
gram (J-09) from the Japan Society for the Promotion of Science.
Appendix A. Supplementary data
Supplementary data associated with this article can be found, in
the online version, at doi:10.1016/j.phytochem.2011.11.010.
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