Tài liệu Báo cáo khoa học: Cleavage of nonphenolic b-1 diarylpropane lignin model dimers by manganese peroxidase from Phanerochaete chrysosporium Evidence for a hydrogen abstraction mechanism docx

Cleavage of nonphenolic b-1 diarylpropane lignin model dimers
by manganese peroxidase from
Phanerochaete chrysosporium
Evidence for a hydrogen abstraction mechanism
G. Vijay B. Reddy
1
, Malayam Sridhar
2
and Michael H. Gold
Department of Biochemistry and Molecular Biology, OGI School of Science and Engineering at OHSU, Beaverton, Oregon, USA
Purified manganese peroxidase (MnP) from Phanerocha-
ete chrysosporium oxidizes nonphenolic b-1 diarylpropane
lignin model compounds in the presence of Tween 80, and
in three- to fourfold lower yield in its absence. In the presence
of Tween 80, 1-(3¢,4¢-diethoxyphenyl)-1-hydroxy-2-(4¢-
methoxyphenyl)propane (I) was oxidized to 3,4-diethoxy-
benzaldehyde (II), 4-methoxyacetophenone (III) and
1-(3¢,4¢-diethoxyphenyl)-1-oxo-2-(4¢-methoxyphenyl)pro-
pane (IV), while only 3,4-diethoxybenzaldehyde (II) and
4-methoxyacetophenone (III) were detected when the reac-
tion was conducted in the absence of Tween 80. In contrast
to the oxidation of this substrate by lignin peroxidase (LiP),
oxidation of substrates by MnP did not proceed under
anaerobic conditions. When the dimer (I) was deuterated at
the a position and subsequently oxidized by MnP in the
presence of Tween 80, yields of 3,4-diethoxybenzaldehyde,
4-methoxyacetophenone remained constant, while the yield
of the a-keto dimeric product (IV) decreased by approxi-
mately sixfold, suggesting the involvement of a hydrogen
abstraction mechanism. MnP also oxidized the a-keto di-
meric product (IV) to yield 3,4-diethoxybenzoic acid (V) and

4-methoxyacetophenone (III), in the presence and, in lower
yield, in the absence of Tween 80. When the reaction
was performed in the presence of
18
O
2
, both products,
3,4-diethoxybenzoic acid and 4-methoxyacetophenone,
contained one atom of
18
O. Finally, MnP oxidized the substrate
1-(3¢,5¢-dimethoxyphenyl)-1-hydroxy-2-(4¢-methoxyphenyl)
propane (IX) to yield 3,5-dimethoxybenzaldehyde (XI),
4-methoxyacetophenone (III) and 1-(3¢,5¢-dimethoxyphe-
nyl)-1-oxo-2-(4¢-methoxyphenyl)propane (X). In sharp
contrast, LiP was not able to oxidize IX. Based on these
results, we propose a mechanism for the MnP-catalyzed
oxidation of these dimers, involving hydrogen abstraction at
a benzylic carbon, rather than electron abstraction from an
aromatic ring.
Keywords: manganese peroxidase; hydrogen abstraction;
diarylpropane dimers; Mn(III); radical mediator.
Lignin is a complex, random, phenylpropanoid polymer
that constitutes 15–30% of woody plant cell walls [1].
White-rot basidiomycetous fungi are primarily responsible
for the initial decomposition of lignin in wood [2–5]. When
cultured under ligninolytic conditions, the best-studied
white-rot basidiomycete, Phanerochaete chrysosporium,
produces two extracellular peroxidases, lignin peroxidase
(LiP) and manganese peroxidase (MnP), which, along with

an H
2
O
2
-generating system, appear to be the major
components of its lignin degradation system [2,3,6–10].
LiP oxidizes a variety of lignin model compounds, including
the most prevalent nonphenolic b-aryl ether (b-O-4 type) as
well as diarylpropane (b-1 type) structures [10–14]. The
enzyme abstracts one electron from the aromatic ring to
form an aryl cation radical [3,11,15,16]. Chemical and ESR
spectroscopic evidence have confirmed the formation of
cation radical species in the LiP-catalyzed oxidation of
alkoxybenzenes [17–21]. In contrast, MnP oxidizes Mn(II),
its primary substrate, to Mn(III), which is chelated by
organic acids such as oxalate or malonate [9,22,23]. The
Mn(III)–organic acid complex, in turn, oxidizes monomeric
phenols and phenolic lignin models via formation of a
phenoxy radical [9,24–27]. MnP is also capable of oxidizing
nonphenolic lignin model dimers and veratryl alcohol, in the
presence of a radical mediator [28–30]. White-rot fungi,
which produce MnP and laccase but not LiP, are still able to
degrade lignin efficiently [31–33], suggesting that these fungi
may produce mediators, enabling MnP and/or laccase to
cleave nonphenolic lignin substructures. Both glutathione
[28] and Tween 80 have been examined as possible
mediators, and a peroxy radical has been implicated in the
Tween 80 reaction [29,30].
In this study, we show that MnP oxidizes nonphenolic
diarylpropane lignin models, in the presence and to a lesser

extent in the absence of Tween 80. Previously, two
mechanisms were considered for the oxidation and cleavage
of nonphenolic lignin dimers by MnP, electron abstraction,
and hydrogen abstraction [28–30]. In our current work,
we have attempted to differentiate between these two
Correspondence to M. H. Gold, Department of Biochemistry and
Molecular Biology, OGI School of Science and Engineering at OHSU,
20000 NW. Walker Road, Beaverton, OR 97006–8921, USA.
Fax: + 1 503 7481464; Tel.: + 1 503 6460957;
E-mail:
Abbreviations: LiP, lignin peroxidase; MnP, manganese peroxidase;
THF, tetrahydrofuran.
1
Present address: Merck, PO Box 2000, RY80L-109, Rahway, NJ
07065, USA.
2
Present address: Department of Chemistry & Biochemistry, Texas
Tech University, Lubbock, TX 79409, USA.
(Received 1 August 2002, revised 31 October 2002,
accepted 21 November 2002)
Eur. J. Biochem. 270, 284–292 (2003) Ó FEBS 2003 doi:10.1046/j.1432-1033.2003.03386.x
mechanisms. The products formed from the oxidation
and cleavage of several different diarylpropane substrates
under both aerobic and anaerobic conditions were exam-
ined. In addition, we have used
18
O
2
to follow the
incorporation of molecular oxygen into the products. Based

on the substrates used, the products identified, and the
results of stable isotope studies, we propose a mechanism
for C
a
–C
b
cleavage of the dimers involving hydrogen
abstraction. Furthermore, our results do not support the
alternative mechanism [30], involving electron abstraction
from the aromatic ring. We also show that a-keto diaryl-
propane lignin dimeric compounds are degraded by a
hydrogen abstraction mechanism to produce benzoic acid
derivatives.
Materials and methods
Synthesis of substrates and products
The diarylethane model 1-(3¢,4¢-diethoxyphenyl)-1-oxo-2-
(4¢-methoxyphenyl)ethane was prepared as described previ-
ously [34,35]. This ketone was treated with methyl iodide in
the presence of potassium tertiary butoxide and anhydrous
dimethylsulfoxide to obtain the a-keto diarylpropane model
1-(3¢,4¢-diethoxyphenyl)-1-oxo-2-(4¢-methoxyphenyl)pro-
pane (IV) [12]. The ketone dimer IV was reduced with either
NaBH
4
or NaBD
4
to obtain 1-(3¢,4¢-diethoxyphenyl)-1-
hydroxy-2-(4¢-methoxyphenyl)propane (I) or [C
a
-

2
H
1
]I,
I(D), respectively. Crude products were purified by silica gel
preparative thin layer chromatography, using 5% ethyl
acetate in hexanes. 1-(3¢,4¢-Diethoxyphenyl)-1,3-dihydroxy-
2-(4¢-methoxyphenyl)propane (VI), 1-(3¢,4¢-diethoxyphe-
nyl)-1-oxo-2-(4¢-methoxyphenyl)-3-hydroxypropane (VIII)
[9,26], 1,2-dihydroxy-1-(4¢-methoxyphenyl)ethane, and
1-(4¢-methoxyphenyl)-1-oxo-2-hydroxyethane (VII) were
prepared as previously described [34,35].
1-(3¢,5¢-Dimethoxyphenyl)-1-hydroxy-2-
(4¢-methoxyphenyl)ethane
This product was prepared by the Barbier reaction [36].
To a mixture of magnesium turnings (1.1 mg atom) and
3,5-dimethoxybenzaldehyde (1.1 mmol) in anhydrous tetra-
hydrofuran (THF; 10 mL) was slowly added 4-meth-
oxybenzyl chloride (1 mmol; 0.14 mL) using a syringe at
room temperature under a nitrogen atmosphere and the
mixture was refluxed for 24 h. The reaction was quenched
with water (10 mL) and extracted with ether (3 · 5mL).
The organic layer was washed with water (1 · 10 mL),
dried over anhydrous sodium sulfate, rotary evaporated,
and the crude preparation was purified by column chro-
matography on neutral alumina, using 5% ethyl acetate in
hexanes.
1-(3¢,5¢-Dimethoxyphenyl)-1-oxo-2-
(4¢-methoxyphenyl)ethane
The pyridinium chlorochromate-on-alumina reagent [37]

(0.75 mmol; 0.8084 g) was added to a flask containing a
solution of 1-(3,5-dimethoxyphenyl)-1-hydroxy-2-(4¢-meth-
oxyphenyl)ethane (0.5 mmol) in hexanes (5 mL). After
stirring for 4 h at room temperature, the solution was
filtered, and washed with diethyl ether (3 · 5mL). The
combined filtrates were evaporated to obtain the product.
1-(3¢,5¢-Dimethoxyphenyl)-1-oxo-2-
(4¢-methoxyphenyl)propane (X)
n-BuLi (0.48 mmol, 0.30 mL of 1.6
M
solution in hexanes)
was added to a solution of diisopropylamine (0.48 mmol,
0.05 g) in dry THF (2 mL) under a nitrogen atmosphere at
0 °C and the mixture was stirred for 0.5 h [38]. A solution of
1-(3¢,5¢-dimethoxyphenyl)-1-oxo-2-(4¢-methoxyphenyl)-
ethane (0.4 mmol) in dry THF (3 mL) was added at 0 °C
and stirred for 1 h. Iodomethane (1.6 mmol) was added
at 0 °C, and the mixture was stirred for 8 h at room
temperature. The reaction was quenched with water (5 mL),
extractedwithether(3· 5 mL), dried over anhydrous
sodium sulfate, and concentrated by evaporation. The crude
preparation was purified by column chromatography on
silica gel using 3% ethyl acetate in hexanes.
1-(3¢,5¢-Dimethoxyphenyl)-1-hydroxy-2-
(4¢-methoxyphenyl)-1-propane (IX)
To a solution of 1-(3,5-dimethoxyphenyl)-1-oxo-2-(4-meth-
oxyphenyl)propane (X) (0.2 mmol, 0.06 g) in ethanol
(5 mL) was added an excess of sodium borohydride
(1.0 g) in three portions and the reaction mixture was
stirred for 4 h at room temperature. The reaction mixture

was neutralized with dilute hydrochloric acid, extracted with
ether (3 · 5 mL), dried over anhydrous sodium sulfate and
concentrated by rotary evaporation. The crude preparation
of IX was purified by column chromatography on neutral
alumina as described above.
Chemicals
3¢,4¢-Diethoxybenzaldehyde (II), 4¢-methoxyacetophenone
(III), 3¢,4¢-diethoxybenzoic acid (V) and 3¢,5¢-dime-
thoxybenzaldehyde (XI) were obtained from Aldrich.
18
O
2
gas (99%) was obtained from Isotec Inc. (Miamisburg,
OH, USA). Unless specified otherwise, other aromatic
compounds were purchased from Aldrich.
Enzymes
Manganese peroxidase (MnP) isozyme 1 and lignin peroxi-
dase (LiP) isozyme H8 were purified from the extracellular
medium of acetate-buffered, agitated, aerobic cultures of
P. chrysosporium OGC101 (ATCC 201542) as previously
reported [39,40]. Purified MnP and LiP were electrophore-
tically homogeneous and had an R
z
value of  5.0.
Enzyme reactions
Reactions with MnP were conducted at 28 °Cfor15hin
1mLof50m
M
sodium malonate, pH 4.5, containing the
dimeric substrate (180 l

M
), MnSO
4
(0.5 m
M
), MnP (5 lg)
and Tween 80 (polyoxyethylenesorbitan monooleate)
(0.1%). LiP reactions were conducted at 28 °Cfor5min
in 1 mL of 20 m
M
succinate, pH 3.0, containing the
substrate (180 l
M
) and enzyme (5 lg). The reactions were
initiated by the addition of 100 l
M
H
2
O
2
and were
Ó FEBS 2003 Mn peroxidase oxidation of diarylpropane dimers (Eur. J. Biochem. 270) 285
conducted under either aerobic or anaerobic conditions.
For anaerobic experiments, reaction mixtures were evacu-
ated and flushed with argon twice to ensure removal of
oxygen. H
2
O
2
was evacuated and purged with argon

separately before addition. Reaction products and unreact-
ed substrates were extracted with ethyl acetate, dried over
anhydrous sodium sulfate, evaporated under nitrogen, and
analyzed directly by GC or as their (acetyl or trimethylsilyl)
derivatives by GC or GC-MS, as described previously [18].
Reaction mixtures were also analyzed directly by HPLC.
New substrates were also analyzed by NMR.
Incorporation of
18
O from
18
O
2
Reactions were carried out in 2-mL vials fitted with rubber
septa. All components of the reaction mixture, except H
2
O
2
,
were added. The vials were evacuated, flushed with argon,
reevacuated, and finally equilibrated with
18
O
2
. Reactions
were initiated with the addition of a deoxygenated solution
of H
2
O
2

, and incubated for 5 min (LiP reactions) or 15 h
(MnP reactions) as described [16,27]. Reaction products
were extracted and prepared for analysis as described above.
Chromatography and mass spectrometry
GC-MS was performed at 70 eV on a Finnigan 4500 mass
spectrometer using a Galaxy data system and fitted with a
Hewlett Packard (HP) 5790 A gas chromatograph and a
30-m fused silica column (DB-5; J & W Scientific, Folsom,
CA, USA). The oven temperature was increased from 70 to
320 °Cat10°Cmin
)1
. HPLC analysis of products was
conducted with an HP Lichrospher 100 RP8 column, using
a linear gradient of 0–100% acetonitrile in 0.05% phos-
phoric acid over 10 min, with a flow rate of 1 mLÆmin
)1
.
Products were detected at 285 nm. Product yields on HPLC
were quantitated using calibration curves obtained with
standards.
Results
Oxidation of diarylpropane substrates by MnP
Time courses for the oxidation of diarylpropanes I and VI
by MnP, in the presence and absence of Tween 80, are
shown in Fig. 1. After 12 h of incubation, over 90% of the
added diarylpropane I was oxidized in the presence of
Tween 80, while in the absence of Tween 80, 24% of the
diarylpropane I was oxidized. In addition, while about 30%
of the diarylpropane VI was oxidized in the presence of
Tween 80 during the 12 h incubation, in the absence of

Tween 80, only about 5% of the VI was oxidized.
The products of the oxidation reactions are shown in
Table 1 and Fig. 2. The products and percent yields of the
MnP oxidation of the diarylpropane I in the presence of
Tween 80 included the a-keto diarylpropane (IV, 58%),
3,4-dimethoxybenzaldehyde (II, 25%) and 4-methoxyaceto-
phenone (III, 21%) (Fig. 2A, Table 1). A small amount
( 2%) of 3,4-diethoxybenzoic acid (V) was also detected
(not shown). The oxidation products for diarylpropane I in
the absence of Tween 80 included 3,4-diethoxybenzaldehyde
(II, 20%) and 4-methoxyacetophenone (III, 14%) (Table 1).
No detectable amount of a-keto diarylpropane (IV) was
produced in the absence of Tween 80. For the oxidation of
the diarylpropane VI, the products included the corres-
ponding a-keto diarylpropane (VIII, 12%), 3,4-diethoxy-
benzaldehyde (II, 13%) and 4-methoxyphenyl-ketol (VII,
8%) (Fig. 2C, Table 1). In the absence of Tween 80, only
about 5% of the added diarylpropane VI was oxidized to
yield 3,4-diethoxybenzaldehyde (II, 2.5%) and 4-methoxy-
phenyl-ketol (VII, 1.5%) (Table 1). Again, no detectable
amount of a-keto diarylpropane VIII was formed in the
absence of Tween 80 (Table 1). To further examine the
mechanism of diarylpropane oxidation by the MnP system,
the diarylpropane IX was prepared. The products of the
MnP oxidation of IX in the presence of Tween 80 included
the a-keto diarylpropane (X, 32%), 3,5-diethoxybenzalde-
hyde (XI, 9.7%) and 4-methoxyacetophenone (III, 11.5%)
(Fig. 2E, Table 1). The products of the MnP oxidation of
the diarylpropane IX in the absence of Tween 80 included
3,5-dimethoxybenzaldehyde (XI, 11.2%) and 4-methoxya-

cetophenone (III, 14.4%). No detectable amount of the
a-keto diarylpropane (X) was produced in the absence of
Tween 80 (Table 1). To determine possible pathways for the
oxidation of the diarylpropanes, the oxidation of the
intermediate a-keto diarylpropanes, described above, also
were examined. Oxidation of the a-keto diarylpropane IV by
MnP, in the presence of Tween 80, yielded 4-methoxyac-
etophenone (III, 22%) and 3,4-diethoxybenzoic acid
(V, 13%) (Table 1). When the oxidation of the a-keto
diarylpropane (IV) was carried out in the absence of Tween
80, yields of the products 4-methoxyacetophenone (III) and
3,4-diethoxybenzoic acid (V) were decreased by approxi-
mately threefold (Table 1).
In the presence of Tween 80, the a-keto diarylpropane
(VIII) was oxidized to 3,4-diethoxybenzoic acid (V, 1.2%)
and 4-methoxyphenyl-ketol (VII, 3%) (Table 1). Less than
1% yield of these products occurred when the reaction was
Fig. 1. Oxidation of the diarylpropanes I (d,s)andVI(m,n)byMnP,
in the presence (d,m) and absence (s,n)ofTween80.Enzyme reac-
tions were carried out in 50 m
M
malonate for 12 h, extracted with ethyl
acetate, and the amount of substrate remaining was quantitated by
HPLC as described in the text and in the legend to Table 1.
286 G. V. B. Reddy et al. (Eur. J. Biochem. 270) Ó FEBS 2003
conducted in the absence of Tween 80. Finally, in the
presence of Tween 80, oxidation of a-keto diarylpropane (X)
resulted in the formation of 3,5-dimethoxybenzoic acid (XII,
5.0%) and the 4-methoxyacetophenone (III, 6.7%). When
the reaction was conducted in the absence of Tween 80, the

same products were formed but in lower yield (Table 1).
Effect of deuterium on the oxidation of the
diarylpropane (I)
When the a-deuterated diarylpropane I(D) was oxidized by
MnP in the presence of Tween 80, the yields of the products,
3,4-diethoxybenzaldehyde (II, 28%), and 4-methoxyaceto-
phenone (III, 21%), were similar to those obtained with the
undeuterated substrate I, whereas the a-keto diarylpropane
product (IV, 9%) was formed in a sixfold lower amount
(Table 1).
Oxidation of diarylpropane substrates by LiP
Oxidation of diarylpropane I by LiP yielded 3,4-diethoxy-
benzaldehyde (II, 70%) and 4-methoxyacetophenone (III,
61%) in approximately 5 min (Fig. 2G). The a-keto
diarylpropane IV product was not detected in this reaction.
Identical results were obtained with a-deuterated diarylpro-
pane I(D). When the diarylpropane (IX) was incubated with
LiP, no products were formed and the amount of substrate
remained unchanged (Table 1). Finally, when either of the
a-keto diarylpropanes (IV or X) was incubated with LiP, no
products were observed and the amount of the substrates
remained unchanged (Table 1).
Diarylpropane oxidation under either
18
O
2
or argon
When the oxidation of the diarylpropane I, by MnP, was
performed under
18

O
2
, 0.8 atoms of
18
O was incorporated
into the 4-methoxyacetophenone (III), whereas 3,4-dieth-
oxybenzaldehyde (II) did not contain detectable amounts of
18
O (Table 2). When the oxidation of a-keto diarylpropane
(IV) by MnP was carried out under
18
O
2
,
18
Owas
incorporated into both products. Approximately 0.8 atoms
of
18
O were incorporated into 3,4-diethoxybenzoic acid (V)
and 0.75 atoms into the 4-methoxyacetophenone (III)
(Table 2). The percentage of
18
O incorporated was estima-
ted by the ratio of the M
+
/(M
+
+2)peaksinthemass
spectrum of the products. Neither the diarylpropane (I) nor

the a-keto diarylpropane (IV) was oxidized by MnP when
the reaction was carried out under argon, either in the
presence or absence of detergent (data not shown).
Discussion
Under ligninolytic conditions, P. chrysosporium secretes
two extracellular peroxidases, MnP and LiP, which are
mainly responsible for the initial depolymerization of lignin
in wood [2,3,22,41]. While both enzymes are iron heme-
containing peroxidases, their detailed reaction mechanisms
differ considerably. LiP abstracts an electron from the
Table 1. Products obtained from the oxidation of nonphenolic diarylpropane substrates I, I(D),VI, IX and a-keto diarylpropane substrates IV, VIII, X
by MnP
a
or LiP
b
. t ¼ trace.
Tween 80
(+/–)
Products formed (mol %)
Dimeric substrate Enzyme II III IV V VII VIII X XI X
Nonphenolic diarylpropane substrates I, I(D),VI, IX
I MnP – 20 14
IMnP+2521582
I(D) MnP + 28 21 9 1
VI MnP – 2.5 1.5
VI MnP + 13 8 12
IX MnP – 14.4 11.2
IX MnP + 11.5 32 9.7
I LiP – 70 61
IX LiP – no reaction

a-Keto diarylpropane substrates IV, VIII, XII by MnP
a
or LiP
b
IV MnP – 7 3.5
IV MnP + 22 13
VIII MnP – t t
VIII MnP + 1.2 3
X MnP – 4.4 3.5
X MnP + 6.7 5.0
IV LiP – no reaction
X LiP – no reaction
a
MnP reactions were conducted at 28 °C for 15 h in 50 m
M
malonate, pH 4.5, containing enzyme, substrate, MnSO
4
, and H
2
O
2
in the
presence or absence of Tween 80, and the products were analyzed by HPLC and GC-MS as described in the text. Amount of products
formed (mol percentage) are shown. Each reaction was run in triplicate; the results are the mean values.
b
Enzyme reactions were conducted
at 28 °C for 5 min in 20 m
M
succinate, pH 3.0, containing enzyme, H
2

O
2
, and substrate, and the products were analyzed by HPLC and
GC-MS as described in the text. Amount of products formed (mol percentage) are shown.
Ó FEBS 2003 Mn peroxidase oxidation of diarylpropane dimers (Eur. J. Biochem. 270) 287
substrate aromatic ring, generating an aryl cation radical,
which decomposes further by enzymatic and nonenzymatic
processes [2,3,11–13,15,16,41,42]. In contrast, MnP oxidizes
Mn(II) to Mn(III) and the latter oxidizes the aromatic
substrate [2,22,24,39].
In the absence of radical mediators, MnP mainly oxidizes
phenolic lignin substructures [9,25–27]. However, in the
presence of mediators, MnP is able to oxidize nonphenolic
lignin substructures [28,29]. In vitro experiments demon-
strate that MnP cleaves nonphenolic b-arylether dimers in
the presence of Tween 80, an unsaturated fatty acid
containing detergent [29,30,43]. In contrast, Tween 20,
which contains a saturated fatty acid, does not act as a
mediator [29]. These studies suggest the involvement of
lipid-derived peroxy radicals as mediators in the oxidation
of nonphenolic lignin model compounds. However, the
detergent-mediated mechanism of oxidation is not clearly
understood. It has been proposed that fatty acid-based
peroxy radicals can oxidize b-aryl ether lignin model
compounds by abstracting either a hydrogen from the C1
position to produce a carbon-centered radical, or an
electron from the aromatic ring to produce an aryl cation
radical intermediate [29,30]. The C1-centered radical,
resulting from a-hydrogen abstraction, has been proposed
to undergo two subsequent reactions. The addition of

oxygen at C1 followed by loss of HOO
Æ
yields an uncleaved
a-keto dimer, which has been proposed to undergo homo-
lytic C2–O fission to expel a phenoxy radical [30].
Alternatively, it has been proposed that a cation radical
intermediate is formed, which subsequently undergoes
C
a
–C
b
cleavage, similar to LiP-catalyzed oxidations [30].
However, the evidence in these studies for electron abstrac-
tion by a peroxy radical is not convincing. Reduced
glutathione also acts as a mediator in the oxidation of
veratryl alcohol and nonphenolic b-ether structures by
MnP. Enzymatically generated Mn(III) oxidizes the thiol to
thiyl radical, which initiates substrate oxidation through
hydrogen, but not electron, abstraction [28].
To determine the relative importance of a-hydrogen vs.
electron abstraction, we studied the mechanism of oxidation
of nonphenolic diarylpropane lignin model compounds by
MnP, in the presence and absence of the unsaturated fatty
acid-based detergent Tween 80. As the initial products of
b-ether dimer cleavage undergo further reactions [30],
elucidating the initial cleavage mechanism is difficult.
Therefore, we selected b-1-type lignin model substructures
as substrates, because they produce relatively stable primary
products, which do not undergo extensive subsequent MnP-
catalyzed oxidation.

The diarylpropane I is oxidized by MnP in the presence
and absence of the radical mediator Tween 80; however, the
oxidation rates are approximately fourfold greater in the
presence of Tween 80 than in its absence. In the presence of
Tween 80, the diarylpropane I undergoes a–b cleavage to
produce 3,4-diethoxybenzaldehyde (II) and 4-methoxy-
acetophenone (III) as well as C
a
oxidation to produce the
corresponding a-keto diarylpropane (IV), which is the
dominant product. In contrast, in the absence of Tween 80,
diarylpropane (I) is oxidized at a slower rate to produce only
Table 2. Incorporation of
18
O during the oxidation of the nonphenolic
diarylpropane I and the a-keto diarylpropane IV by MnP.
Substrate Product m/z
18
O incorporated (%)
a
I II 196 (M
+
+2) –
III 152 (M
+
+2) 70
I V 212 (M
+
+2) 80
III 152 (M

+
+2) 75
a18
O incorporation divided by the total oxygen incorporation
· 100. Enzyme reactions were carried out in duplicate under
18
O
2
in
50 m
M
malonate, containing enzyme, substrate, Tween 80, MnSO
4
,
and H
2
O
2
for 15 h at 28 °C. Products were extracted and quanti-
tated by GC-MS as described in the text.
Fig. 2. Products identified during the oxidation of nonphenolic diaryl-
propanes (reactions A, C, and E) and a-keto diarylpropanes (reactions B,
D, and F) by homogeneous MnP and LiP (G). MnP reactions were
carried out in 50 m
M
malonate buffer in the presence or absence of
Tween80for15hat28°C as described in the text. LiP reactions were
carried out in 50 m
M
succinate at 28 °C for 5 min as described in the

text. Products were extracted and analyzed by HPLC and GC-MS, as
described in the text. Yields of products are presented in Table 1.
288 G. V. B. Reddy et al. (Eur. J. Biochem. 270) Ó FEBS 2003
the a–b cleavage products. Under these conditions, the
a-keto diarylpropane product (IV) is not observed, suggest-
ing that C1 oxidation does not occur. Although only about
20% of the diarylpropane (I) is oxidized by MnP in the
absence of Tween 80, this finding is surprising, because it was
previously thought that MnP is incapable of attacking
nonphenolic lignin models in the absence of radical media-
tors [28–30]. However, recent evidence suggests that metal–
oxo complexes as well as Mn(III) complexes are able to
abstract an H atom from certain aromatic compounds
[44,45].
Corresponding products are formed during the oxidation
of the diarylpropane VI, although at a lower rate. The lower
reactivity of VI, when compared to I, may be due to the
stearic hindrance offered by the hydroxyl group at C3,
which may inhibit hydrogen abstraction at C2. When the
MnP reaction was conducted under anaerobic conditions,
in the presence or absence of the detergent, oxidation of
these diarylpropanes does not occur, indicating that
molecular oxygen is required.
The LiP oxidation of diarylpropane substructures has
been well studied and proceeds through an aryl cation
radical intermediate [11,15,16]. Therefore, we compared the
LiP and MnP oxidation of the diarylpropane I under a
variety of conditions. LiP oxidizes over 70% of the
diarylpropane I in 5 min to produce the C
a

–C
b
cleavage
products 3,4-diethoxybenzaldehyde (II) and 4-methoxy-
acetophenone (III). Furthermore, unlike the MnP reaction,
the LiP-catalyzed oxidation of the diarylpropane I proceeds
efficiently under both aerobic and anaerobic conditions [16].
Finally, the a-keto diarylpropane is not formed in the LiP-
catalyzed oxidation of the diarylpropane I [12,16], whereas
it is the major product in the MnP reaction. As the LiP
oxidation of the diarylpropane I proceeds by the formation
of an aryl cation radical [16], these results suggest that a
cation radical is not an intermediate in the MnP-catalyzed
reactions reported here.
To pursue this question further, we examined the oxida-
tion of 1-(3¢,5¢-dimethoxyphenyl)-1-hydroxy-2-(4¢-methoxy-
phenyl)-1-propane (IX) by MnP and LiP. As expected, LiP
was not able to oxidize the diarylpropane IX, owing to the
lack of an electron-donating methoxy group at the para
position of the A ring. This strongly suggests that a cation
radical is difficult to produce with this substrate. In contrast,
in the presence or absence of Tween 80, MnP oxidizes the
diarylpropane (IX)-producing products corresponding to
those produced during the oxidation of the diarylpropane
(I). These results support our view that electron abstraction
is an unlikely mechanism for the MnP-catalyzed oxidation
of these diarylpropane substrates.
The presence of deuterium at the C1 position of the
diarylpropane (I) (Fig. 2A) had a strong influence on the
yield of the a-keto diarylpropane (IV) formed. When

deuterated diarylpropane I(D) was used as the substrate, the
yield of the a-keto diarylpropane (IV) was decreased by
about 6 fold, suggesting the involvement of H-abstraction.
However, yields of the C
a
–C
b
cleavage products 3,4-
diethoxybenzaldehyde (II) and 4-methoxyacetophenone
(III) remained almost identical, for the deuterated and
undeuterated substrates. This result is in contrast to that
with the LiP-catalyzed oxidation of I(D), which is not
affected by the presence of deuterium.
Oxidation of the diarylpropane I to 3,4-diethoxybenzal-
dehyde (II), 4-methoxyacetophenone (III), and a-keto
diarylpropane (IV) can be explained on the basis of an
initial hydrogen abstraction reaction at one of two posi-
tions. A benzylic radical can be generated at C1, which is
resonance stabilized by aromatic ring A, or at C2, which is
stabilized by ring B, as well as by the adjacent methyl group
through hyperconjugation. However, the presence of a
hydroxyl group at C1 renders the C1–H bond more labile
than the C2–H bond. A radical generated at C1 would add
to O
2
to form a peroxyl radical, which would then eliminate
HOO
Æ
to yield the a-keto diarylpropane IV, as shown in
Fig. 3. This mechanism is similar to that reported earlier for

the thiyl radical-mediated oxidation of nonphenolic lignin
dimers by MnP [28]. Alternatively, a radical generated at C2
would add to O
2
and the resulting peroxy radical could
abstract a hydrogen atom to form an unstable hydroper-
oxide. The latter would undergo C
a
–C
b
cleavage with the
elimination of H
2
O to form 3,4-diethoxybenzaldehyde (II)
and 4-methoxyacetophenone (III) (Fig. 3). Addition of
deuterium at the C1 position would slow the hydrogen
abstraction at C1, but would have no effect on the
formation of a radical at C2 as we observed.
Fig. 3. Proposed hydrogen abstraction mechanism for the oxidative
cleavage of the diarylpropane (I) by MnP in the presence of Tween 80.
Ó FEBS 2003 Mn peroxidase oxidation of diarylpropane dimers (Eur. J. Biochem. 270) 289
When the oxidation of the diarylpropane I is performed
in the presence of
18
O
2
,
18
O is incorporated only into the
acetophenone (III). The benzaldehyde (II) did not contain

any
18
O. This is similar to the LiP-catalyzed oxidation
(electron abstraction), where
18
O is incorporated only into
benzylic radical-derived products [16]. Therefore, hydrogen
abstraction at C2 and electron abstraction from the
aromatic ring result in C
a
–C
b
cleavage and
18
O incorpor-
ation only into the acetophenone (III). The crucial differ-
ence is that the hydrogen abstraction reaction catalyzed by
MnP, unlike the LiP-catalyzed oxidation, does not proceed
in the absence of O
2
[16].
The a-keto diarylpropane (IV) also is oxidized by MnP in
the presence of Tween 80. The reaction products included
4-methoxyacetophenone (III) and 3,4-diethoxybenzoic acid
(V). The benzaldehyde (II) is not detected as a product of
the reaction. When the reaction is carried out in the absence
of Tween 80, the oxidation rate is about 3-fold lower, but
again, 4-methoxyacetophenone (III) and 3,4-diethoxyben-
zoic acid (V) were detected. The a-keto dimer (IV) was not
oxidized under anaerobic conditions, indicating that oxygen

is required for this reaction as well.
In contrast to the results with MnP, LiP is not able to
oxidize the dimer (IX), nor any of the a-keto diarylpropane
dimers. Furthermore, LiP oxidation of the diarylpropane (I)
does not yield an a-keto diarylpropane product. Finally, the
LiP oxidation of diarylpropanes occurs in the absence of
molecular oxygen. Because LiP oxidizes these substrates by
electron abstraction to form an aryl cation radical, these
results strongly suggest electron abstraction is not occurring
in the MnP reactions.
The mechanism we propose for the MnP oxidation of
the a-keto diarylpropane (IV) is shown in Fig. 4. The
C2-centered radical generated after C2–H abstraction
would add to oxygen to form a peroxy radical. The latter
can react with the carbonyl group, producing an unstable
dioxetane, which decomposes to 4-methoxyacetophenone
(III) and 3,4-diethoxybenzoic acid (V) (Fig. 4).
When the oxidation of a-keto diarylpropane (IV) by
MnP is conducted in the presence of
18
O
2
,
18
O is incorpor-
ated into 4-methoxyacetophenone (III) and 3,4-diethoxy-
benzoic acid (V), supporting the mechanism shown in
Fig. 4. MnP slowly oxidizes the a-keto diarylpropane (IV)
even in the absence of Tween 80. Formation of aromatic
acids from nonphenolic lignin model compounds has not

been reported previously. In this study we also identified a
small amount of 3,4-diethoxybenzoic acid during the
oxidation of diarylpropane I by MnP in the presence of
Tween 80. This is most likely due to the further oxidation of
either 3,4-diethoxybenzaldehyde (II) or the a-ketodiaryl-
propane (IV) produced during the oxidation of I. We have
confirmed this secondary oxidation process in a separate
reaction using 3,4-diethoxybenzaldehyde (II) as a substrate
(data not shown). The oxidation of a benzaldehyde to
benzoic acid by MnP has been discussed recently [29].
According to the mechanisms shown in Fig. 4, products
formed from the C
a
–C
b
cleavage should be in equal molar
ratio. However, we observe that yields of the acetophenone
(III) are slightly lower than for the benzaldehyde (II). It is
likely that some of the products are degraded further by the
enzyme. We are investigating the nature of the oxidant in
the reaction of the diarylpropane I by MnP in the absence of
unsaturated detergent (Tween 80). However, both metal
oxo complexes and Mn(III) complexes are capable of
abstracting a hydrogen atom from organic compounds
[44,45]. In addition, a malonic acid-generated free radical
may be involved in the process [46].
In conclusion, the present study shows that MnP can
catalyze C
a
oxidation and C

a
–C
b
cleavage of nonphenolic
diarylpropane model compounds in the presence of Tween
80 and at a lower rate in its absence. Based on the products
formed under various conditions, a mechanism based on
electron abstraction can be ruled out; rather, these
compounds apparently are oxidized solely via hydrogen
abstraction mechanisms. This study also shows that the
a-keto-1,2-diarylpropane is oxidized via hydrogen abstrac-
tion to produce an aromatic acid and an acetophenone
product.
Acknowledgments
This research was supported by Grants MCB-9808430 from the
National Science Foundation and DE-FG03–96ER30325 from the
Division ofEnergy Biosciences, U.S.Department of Energy (to M.H.G).
Fig. 4. Proposed hydrogen abstraction mechanism for the oxidative
cleavage of the a-keto diarylpropane (IV) by MnP in the presence of
Tween 80.
290 G. V. B. Reddy et al. (Eur. J. Biochem. 270) Ó FEBS 2003
References
1. Sarkanen, K.V. & Ludwig, C.H. (1971) Lignins. Occurrence,
Formation, Structure and Reactions. Wiley-Interscience, New
York, USA.
2. Gold, M.H. & Alic, M. (1993) Molecular biology of the lignin-
degrading basidiomycete Phanerochaete chrysosporium. Microbiol.
Rev. 57, 605–622.
3. Kirk, T.K. & Farrell, R.L. (1987) Enzymatic ÔcombustionÕ:the
microbial degradation of lignin. Annu.Rev.Microbiol.41, 465–

505.
4. Crawford, R.L. (1981) Lignin Biodegradation and Transformation.
Wiley and Sons, New York, USA.
5. Buswell, J.A. & Odier, E. (1987) Lignin biodegradation. CRC Crit.
Rev. Biotechnol. 6, 1–60.
6. Leisola, M.S.A., Kozulic, B., Meussdoerffer, F. & Fiechter, A.
(1987) Homology among multiple extracellular peroxidases from
Phanerochaete chrysosporium. J. Biol. Chem. 262, 419–424.
7. Kuwahara, M., Glenn, J.K., Morgan, M.A. & Gold, M.H. (1984)
Separation and characterization of two extracellular H
2
O
2
-
dependant oxidases from the ligninolytic cultures of Phaner-
ochaete chrysosporium. FEBS Lett. 169, 247–250.
8. Tien, M. & Kirk, T.K. (1984) Lignin-degrading enzyme from
Phanerochaete chrysosporium: purification, characterization,
and catalytic properties of a unique H
2
O
2
-requiring oxygenase.
Proc. Natl Acad. Sci. USA 81, 2280–2284.
9. Glenn, J.K. & Gold, M.H. (1985) Purification and characteriza-
tion of an extracellular Mn(II)-dependent peroxidase from the
lignin-degrading basidiomycete, Phanerochaete chrysosporium.
Arch. Biochem. Biophys. 242, 329–341.
10. Gold, M.H., Kuwahara, M., Chiu, A.A. & Glenn, J.K. (1984)
Purification and characterization of an extracellular H

2
O
2
-
requiring diarylpropane oxygenase from the white rot basidio-
mycete, Phanerochaete chrysosporium. Arch. Biochem. Biophys.
234, 353–362.
11. Schoemaker, H.E. (1990) On the chemistry of lignin biodegrada-
tion. Recl. Trav. Chim. Pays-Bas 109, 255–272.
12. Miki, K., Renganathan, V. & Gold, M.H. (1986) Mechanism of
b-aryl ether dimeric lignin model compound oxidation by lignin
peroxidase of Phanerochaete chrysosporium. Biochemistry 25,
4790–4796.
13. Miki, K., Kondo, R., Renganathan, V., Mayfield, M.B. & Gold,
M.H. (1988) Mechanism of aromatic ring cleavage of a b-biphenyl
ether dimer catalyzed by lignin peroxidase of Phanerochaete
chrysosporium. Biochemistry 27, 4787–4794.
14. Hammel, K.E., Tien, M., Kalyanaraman, B. & Kirk, T.K. (1985)
Mechanism of oxidative C
a
–C
b
cleavage of a lignin model dimer
by Phanerochaete chrysosporium ligninase. Stoichiometry and
involvement of free radicals. J. Biol. Chem. 260, 8348–8353.
15. Higuchi, T. (1990) Lignin biochemistry: biosynthesis and bio-
degradation. Wood Sci. Technol. 24, 23–63.
16. Renganathan, V., Miki, K. & Gold, M.H. (1986) Role of
molecular oxygen in lignin peroxidase reactions. Arch. Biochem.
Biophys. 246, 155–161.

17. Kersten, P.J., Tien, M., Kalyanaraman, B. & Kirk, T.K. (1985)
The ligninase of Phanerochaete chrysosporium generates cation
radicals from methoxybenzenes. J. Biol. Chem. 260, 2609–2612.
18. Joshi, D.K. & Gold, M.H. (1996) Oxidation of dimethoxylated
aromatic compounds by lignin peroxidase from Phanerochaete
chrysosporium. Eur. J. Biochem. 237, 45–57.
19. Khindaria, A., Grover, T.A. & Aust., S.D. (1995) Evidence for
formation of the veratryl alcohol cation radical by lignin per-
oxidase. Biochemistry 34, 6020–6025.
20. Candeias, L.P. & Harvey, P.J. (1995) Lifetime and reactivity of the
veratryl alcohol radical cation. Implications for lignin peroxidase
catalysis. J. Biol. Chem. 270, 16745–16748.
21. Teunissen, P.J.M., Sheng, D., Reddy, G.V.B., Moenne-Loccoz,
P., Field, J.A. & Gold, M.H. (1998) 2-Chloro-1,4-dimethoxy-
benzene cation radical: formation and role in the peroxidase
oxidation of anisyl alcohol. Arch. Biochem. Biophys. 360, 233–
238.
22. Gold, M.H., Youngs, H.L. & Sollewijn Gelpke, M.D. (2000)
Manganese peroxidase. In Manganese and its Role in Biological
Processes (Sigel, A. & Sigel, H., eds), pp. 559–586. Marcel Dekker,
New York, USA.
23. Kuan, I.C. & Tien, M. (1993) Stimulation of Mn peroxidase
activity: a possible role for oxalate in lignin biodegradation. Proc.
NatlAcad.Sci.USA90, 1242–1246.
24. Glenn, J.K., Akileswaran, L. & Gold, M.H. (1986) Mn(II) oxi-
dation is the principal function of the extracellular Mn-peroxidase
from Phanerochaete chrysosporium. Arch. Biochem. Biophys. 251,
688–696.
25. Paszczyn
˜

ski, A., Huynh, V.B. & Crawford, R. (1986) Comparison
of ligninase-I and peroxidase-M2 from the white-rot fungus
Phanerochaete chrysosporium. Arch. Biochem. Biophys. 244, 750–
765.
26. Wariishi, H., Valli, K. & Gold, M.H. (1989) Oxidative cleavage of
a phenolic diarylpropane lignin model dimer by manganese per-
oxidase from Phanerochaete chrysosporium. Biochemistry 28,
6017–6023.
27. Tuor, U., Wariishi, H., Schoemaker, H.E. & Gold, M.H. (1992)
Oxidation of phenolic arylglycerol b-aryl ether lignin model
compounds by manganese peroxidase from Phanerochaete chrys-
osporium: oxidative cleavage of an alpha-carbonyl model com-
pound. Biochemistry 31, 4986–4995.
28. Wariishi, H., Valli, K., Renganathan, V. & Gold, M.H. (1989)
Thiol-mediated oxidation of nonphenolic lignin model com-
pounds by manganese peroxidase of Phanerochaete chrysospor-
ium. J. Biol. Chem. 264, 14185–14191.
29. Bao, W., Fukushima, Y., Jensen, K.A. Jr, Moen, M.A. &
Hammel, K.E. (1994) Oxidative degradation of non-phenolic lig-
nin during lipid peroxidation by fungal manganese peroxidase.
FEBS Lett. 354, 297–300.
30. Kapich, A.N., Jensen, K.A. & Hammel, K.E. (1999) Peroxyl
radicals are potential agents of lignin biodegradation. FEBS Lett.
461, 115–119.
31. Pe
´
rie
´
, F.H. & Gold, M.H. (1991) Manganese regulation of man-
ganese peroxidase expression and lignin degradation by the white

rot fungus Dichomitus squalens. Appl. Environ. Microbiol. 57,
2240–2245.
32. Hatakka, A. (1994) Lignin-modifying enzymes from selected
white-rot fungi: production and role in lignin degradation. FEMS
Microbiol. Rev. 13, 125–135.
33. Srebotnik, E., Jensen, J.K. & Hammel, K. (1994) Fungal
degradation of recalcitrant nonphenolic lignin structures without
lignin peroxidase. Proc. Natl Acad. Sci. USA 91, 12794–12797.
34. Enoki, A., Goldsby, G.P. & Gold, M.H. (1981) b-Ether cleavage
of the lignin model compound 4-ethoxy-3-methoxyphenyl-
glycerol-b-guaiacyl ether and derivatives by Phanerochaete
chrysosporium. Arch. Microbiol. 129, 141–145.
35. Enoki, A. & Gold, M.H. (1982) Degradation of the diarylpropane
lignin model compound 1-(3,4-diethoxyphenyl)-1,3-dihydroxy-2-
(4-methoxyphenyl)-propane and derivatives by the basidiomycete
Phanerochaete chrysosporium. Arch. Microbiol. 132, 123–130.
36. Blomberg, C. & Hartog, F.A. (1977) Barbier Reaction, Synthesis,
18.
37. Vogel, A.I. (1989) Oxidation of pyridinium dichromate. Vogel’s
Textbook of Practical Organic Chemistry, 5th edn, p. 610. Wiley,
New York, USA.
38. Larock, R.C. (1989) Alkylation using lithium diisopropylamide.
Comprehensive Organic Transformations, a Guide to Functional
Group Preparations, p. 738. VCH Publishers, New York, USA.
Ó FEBS 2003 Mn peroxidase oxidation of diarylpropane dimers (Eur. J. Biochem. 270) 291
39. Wariishi, H., Akileswaran, L. & Gold, M.H. (1988) Manganese
peroxidase from the basidiomycete Phanerochaete chrysosporium:
spectral characterization of the oxidized states and the catalytic
cycle. Biochemistry 27, 5365–5370.
40. Sheng, D.W. & Gold, M.H. (1999) Oxidative polymerization of

ribonuclease A by lignin peroxidase from Phanerochaete chrysos-
porium. Role of veratryl alcohol in polymer oxidation. Eur. J.
Biochem. 259, 626–634.
41. Eriksson, K E., Blanchette, R.A. & Ander, P. (1990) Biode-
gradation of lignin. Microbial and Enzymatic Degradation of Wood
and Wood Components, pp. 225–333. Springer-Verlag KG, Berlin,
Germany.
42. Kirk, T.K., Tien, M., Kersten, P.J., Mozuch, M.D. &
Kalyanaraman, B. (1986) Ligninase of Phanerochaete chrysos-
porium. Mechanism of its degradation of the non-phenolic
arylglycerol b-aryl ether substructure of lignin. Biochem. J. 236,
279–287.
43. Kapich, A.N., Hofrichter, M., Vares, T. & Hatakka, A. (1999)
Coupling of manganese peroxidase-mediated lipid peroxidation
with destruction of nonphenolic lignin model compounds
and
14
C-labeled lignins. Biochem. Biophys. Res. Commun. 259,
115–119.
44. Mayer, J.M. (1998) Hydrogen atom abstraction by metal-
oxo complexes: understanding the analogy with organic radical
reactions. Acc. Chem. Res. 31, 441–450.
45. Bryant, J.R., Taves, J.E. & Mayer, J.M. (2002) Oxidations of
hydrocarbons by manganese(III) tris(hexafluoroacetylacetonate).
Inorg. Chem. 41, 2769–2776.
46. Hofrichter, M., Ziegenhagen, D., Vares, T., Friedrich, M.,
Jager, M.G., Fritsche, W. & Hatakka, A. (1998) Oxidative
decomposition of malonic acid as basis for the action of
peroxidase in the absence of hydrogen peroxide. FEBS Lett. 434,
362–366.

292 G. V. B. Reddy et al. (Eur. J. Biochem. 270) Ó FEBS 2003