From the Department of Environmental Biotechnology,
MIGAL-Galilee Technology Center, South Industrial Zone, Kiryat
Shmona 10200, Israel, the § Department of Plant Pathology & Microbiology and the ¶ Institute of Biochemistry, Food Science and
Nutrition, The Faculty of Agricultural, Food and Environmental Quality
Sciences, The Hebrew University of Jerusalem, Rehovot 76100, Israel,
the
Department of Organic Chemistry, Weizmann Institute of
Science, Rehovot, 76100, Israel, and the ** Division of Environmental
and Water Resources Engineering, Faculty of Civil Engineering,
Technion-Israel Institute of Technology, Haifa 32000, Israel
Received for publication, October 26, 2000, and in revised form, January 12, 2001
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ABSTRACT |
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The major products of the initial steps of
ferulic acid polymerization by lignin peroxidase included three
dehydrodimers resulting from Lignin peroxidase (LIP)1
is considered to be one of the most important enzymes of the
extracellular lignin degradation system secreted by the white rot
fungus, Phanerochaete chrysosporium (1). Although LIP shares
spectral and kinetic features with other peroxidases, the enzyme has
several unique characteristics, including a redox potential higher than
those of other peroxidases (2, 3). The high redox potential enables LIP
to oxidize aromatic compounds with calculated ionization potential (IP)
values of up to 9.0 eV (4). This has striking implications when
considering the potential applications of peroxidases for useful
biotransformations (5-7). LIP can be expected to oxidize a wider range
of substrates and therefore have potential applications unsuitable for
less potent peroxidases.
Phenols are oxidized by peroxidases to generate phenoxy radicals, which
couple with other substrate molecules to form dimeric, oligomeric, and
polymeric products. This phenomenon can be exploited for the
biocatalytic production of useful oligomers and polymers (6, 7), as
well as for the treatment of wastewater streams polluted with toxic
phenols (8-10).
Ferulic acid (FA), which is an extremely abundant and widespread
cinnamic acid derivative (11), was chosen as a model substrate for
studying the initial steps of LIP-catalyzed polymerization of phenolic
compounds in vitro. In vivo, peroxidase-catalyzed oxidation
of FA esterified to primary plant cell wall polysaccharides results in
the formation of FA dehydrodimers, believed to enhance the rigidity and
strength of the cell wall. A range of regio-isomeric dehydrodimers
identified and quantified in several plant cell walls include products
of Of all the peroxidases LIP should theoretically be able to catalyze the
highest degree of FA polymerization, its high redox potential enabling
it to further oxidize dimers and oligomers with high IP values.
However, this may be dependent on the mechanism of polymerization,
because if polymers also arise by attack of FA radicals on preformed
dehydrodimers and oligomers, then all peroxidases should be capable of
achieving a similar degree of polymerization. This study evaluates the
mechanism of oligomer and polymer formation by LIP. In addition, the
identification of FA oligomers may prove fruitful, in light of the many
potential applications arising for FA and its derivatives in the
pharmaceutical and food industries (20).
Enzyme Purification
LIP isoenzyme H1 (LIP-H1) was produced from high nitrogen
cultures of P. chrysosporium Burds BKM-F-1767 as described
previously (21). The enzyme was purified in two steps by MonoQ HPLC,
first using a 0.01-1 M sodium acetate gradient at pH 6.0 (22) and then by employing a similar gradient at pH 4.7, equivalent to the pI value of H1. The purified enzyme had an Reinheitszahl
(A409/280) value > 4.0. LIP concentration
was determined at 409 nm using an extinction coefficient of 169 mM Oxidation of FA
For analytical HPLC, oxidation of 300 µM FA was
performed with 1 µM LIP-H1 in 50 mM sodium
tartrate buffer, pH 3.5, and varying concentrations of
H2O2 in a total reaction volume of 1 ml. To prevent H2O2-dependent enzyme
inactivation, H2O2 was added stepwise in
aliquots of 100 µM min For fractionation of the oxidation products, the reaction was carried
out on a larger scale. A total of 200 ml of 4 mM FA was
oxidized by 2 µM LIP and a total of 4 mM
H2O2, which was added in aliquots of 200 µM at 1-min intervals. 30 min after addition of the last
aliquot of H2O2, reaction products were
extracted with three volumes of ethyl acetate, evaporated to dryness,
and redissolved in a small amount of 50% (v/v) aqueous methanol before fractionation.
HPLC Analysis and Fractionation
HPLC analysis and fractionation were conducted using a Hewlett
Packard HPLC (HP1100 series) equipped with a diode array detector. All
solvents were of far UV quality HPLC grade purity where available.
Analytical Gel Permeation Chromatography
Gel permeation analysis was performed using a TSK gel G3000 HR
column (7.8 mm × 30 cm; particle size, 5 µM;
TosoHaas, Stuttgart, Germany). TSK polystyrene standards with molecular
weights of 300, 500, 1000, 2500, and 5000 were employed (TOSOH
Corporation, Tokyo, Japan). Elution was performed using tetrahydrofuran
as the mobile phase. The flow rate was maintained at 0.5 ml
min Analytical Reverse-phase Chromatography
Reverse-phase analysis was conducted using a Lichrospher 100 RP-C18 column (25 cm × 5 mm inner diameter; 5 µm; Merck).
Elution was performed using a gradient system adapted from a previously described method (14), which increased the relative amounts of methanol
and acetonitrile present in aqueous 1 mM trifluoroacetic acid. The gradient profile consisted of solvent A (10%, v/v, aqueous acetonitrile plus trifluoroacetic acid to 1 mM), solvent B
(80%, v/v, aqueous methanol plus trifluoroacetic acid to 1 mM), and solvent C (80%, v/v, aqueous acetonitrile plus
trifluoroacetic acid to 1 mM) in the following program:
initially, 90% A, 5% B, and 5% C; linear gradient over 25 min to
26% A, 37% B, and 37% C; linear gradient over 5 min to 0% A, 50%
B, and 50% C; linear gradient over 15 min to 90% A, 5% B, and 5% C;
and held isocratically at 90% A, 5% B, and 5% C for a further 10 min. The flow rate was maintained at 1 ml min Fractionation of Oxidation Products
Oxidation products were fractionated using a semi-preparative
reverse-phase Lichrospher 100 RP-18 column (25 cm × 10 mm inner diameter; 10 µm; Lichrocart) employing the previously described gradient system. A flow rate of 6 ml min Analysis and Chemical Identification of Oxidation Products
GC-MS--
Dried products were silylated in 200 µl of dioxane
with 200 µl of N,O-bis
(trimethylsilyl)-acetamide for 30 min at 60 °C. Trimethylsilylated
derivatives were separated using a 0.25 mm × 30 m HP5 Phe Me
Silicone column on a Hewlett Packard 5972 series gas chromatograph with
helium as the carrier gas and detected with a Hewlett Packard
5972 mass selective detector. The column was ramped at 10 °C
min NMR Spectroscopy--
13C and 1H NMR
experiments were performed using a Bruker "Avance" DRX-400
instrument, operating at a frequency of 400.13 MHz for 1H
observation. The spectrometer was equipped with a 5-mm Bruker inverse
multinuclear resonance probe with a single-axis (z) gradient coil.
Spectra were measured at room temperature in CD3OD.
Chemical shifts (ppm) were given on the One-dimensional NOE--
One-dimensional NOE difference
experiments were acquired nonspinning in blocks of 40 on- and 40 off-resonance scans with a presaturation time of 2.5 s in an
interleaved manner.
Two-dimensional Gradient-enhanced Heteronuclear Multiple Quantum
Correlation Spectra--
Two-dimensional gradient-enhanced
heteronuclear multiple quantum correlation spectra were acquired
with a 17:20:25 gradient ratio (duration 1 ms), 1024-2048
points in F2, 128-256 complex increments in
F1, four to eight scans per increment. Apodization was with
a
Gradient-enhanced heteronuclear multiple-bond correlation
spectra were obtained with a 50:30:40.1 gradient ratio (duration 2 ms),
1024-2048 points in F2, 128-256 complex increments
in F1, and 40-scans per increment. The long range delay was
optimized to 60 ms. Spectra were obtained in magnitude mode and
transformed with a sine bell weighting function in both dimensions.
Quantum Chemical Calculations
The semi-empirical AM1 quantum chemical method was used for
calculating the optimal geometries and relative energies of the ferulic
acid trimers and their free radical precursors. All the calculations
were performed with the Gaussian 94 (25) and Spartan 5.1 programs.
Chemicals
H2O2 (a 30%, v/v, solution), FA,
and N,O-bis (trimethylsilyl)-acetamide were
obtained from Sigma. The concentration of stock solutions of
H2O2 was determined at 240 nm using an
extinction coefficient of 39.4 M HPLC Analysis--
Generation of phenoxy radicals from FA by
peroxidases can theoretically lead to a plethora of polymerization
products (11). To get an indication of the extent of polymerization
by LIP, gel permeation chromatography was performed on reactions that
had been frozen at
When the same reaction mixtures were subjected to reverse-phase HPLC,
numerous peaks were obtained corresponding to oxidation products. A
typical chromatogram obtained from large scale oxidation of FA is shown
(Fig. 2). Although numerous peaks were
obtained, only the major ones, labeled I-IV, were purified
and identified.
Product Identification--
To characterize the major products,
the reaction of LIP with FA was carried out on a larger scale and the
peaks of interest were fractionated. The structures of the peaks
labeled I-IV in Fig. 2 were primarily determined by
1H NMR, 13C NMR, COSY, one- and two-dimensional
NOE experiments. GC-MS was also employed.
1H NMR spectroscopy indicated that peak I in
Fig. 2 corresponds to a product of FA dehydrodimerization, consisting
of two nonequivalent tri-substituted aromatic fragments, A and B, one
tri-substituted double bond, a saturated fragment, and two methoxy
groups. The protons of ring A were characterized by chemical shifts and
hyperfine structural patterns similar to those of the parent FA:
GC-MS analysis of peak I after silylation indicated
the formation of two isomeric tetrakis(trimethylsilylated)
Peak II in Fig. 2 is also a product of FA
dehydrodimerization. This compound consists of one tri-substituted
aromatic fragment A, one tetra-substituted aromatic fragment B, one
di-substituted double bond, a saturated fragment, two carboxylic groups
and two methoxy groups. The protons of ring A were characterized by the following chemical shifts and hyperfine structural patterns:
GC-MS analysis of peak II after silylation indicated the
presence of three different compounds with molecular masses of 674, 602 (the major component), and 558 (the minor component). These three
compounds may be assigned as follows: the 602 peak to the
tris(trimethylsilylated) derivative of 2a; the 674 peak
to the tetra(trimethylsilylated) derivative of 2b (the
possibility of partial disclosure of the furanoid ring in 2a
during the silylation procedure has been reported; Ref. 12), and the
558 peak to the tris(trimethylsilylated) derivative of the
decarboxylation product of 2b (see structure 2c and Ref. 12).
Peak III in Fig. 2 consisted of two major components. One of
them is a symmetric dehydrodimer of FA, consisting of two equivalent
tri-substituted aromatic rings and two equivalent saturated parts. The
aromatic protons were characterized by the following chemical shifts:
GC-MS analysis of peak III after silylation indicated the
formation of four major components: two having a molecular mass of 674, similar to those found for peak I, one with a molecular mass
of 602 (probably the tris(trimethylsilylated) product of lactone
1a), and the last with a molecular mass of 530 which fits
well with bis(trimethylsilylated) dilactone 1c.
The second major component in peak III has the structure of
a FA trimer. The 1H and 13C NMR spectra led us
to believe that there are three different tri-substituted aromatic
fragments in the molecule: two double bonds and one saturated fragment.
The
Structure 3 fits well with all of the aforementioned data
and conclusions (Fig. 5). Optimization of
structure 3 by molecular mechanics and semi-empirical
quantum chemical AM1 techniques yielded a geometry conforming to the
conclusions of the one-dimensional NOE experiment.
Peak IV in Fig. 2 consisted of one major component that is
probably also a FA trimer. It is composed of two tri-substituted aromatic fragments (A and B), one tetra-substituted aromatic fragment (C), two di-substituted double bonds, and a saturated fragment. The
1H and 13C chemical shifts as well as splitting
patterns are presented in Table V. The
assignment of the signals in the 1H and 13C NMR
spectra was based on the analysis of the splitting pattern, COSY
spectra, 13C-1H correlation spectra, and the
results of the two-dimensional NOE experiments. The two di-substituted
double bonds are connected to rings B and C. The aromatic ring A is
connected to the CH group (A
The aforementioned data are not enough for unambiguous determination of
the chemical structure of the major component of peak IV.
However, additional information was obtained from GC-MS analysis of
peak IV after silylation. The chromatogram indicated the
formation of two compounds. One of them was identified as
bis(trimethylsilylated) FA (M+ = 338). The molecular mass
of the second was found to be equal to 484, which fits well with the
bis(trimethylsilylated) product of the benzofuran structure
4 (Fig. 6). The addition of FA
to structure 4 will result in the formation of structure 5, the most probable major component of peak IV (Fig. 6).
Further Oxidation of Identified Products--
When the identified
products were isolated and further incubated with LIP in the presence
of H2O2, their peaks either disappeared completely or their intensity decreased as indicated by reverse-phase HPLC. This complies with other findings, in which the intensity of the
product peaks decreased at high H2O2
concentrations, indicating that these products were probably further
oxidized. Although no new peaks were observed when the mixtures were
analyzed by reverse-phase HPLC, gel permeation chromatography indicated
that higher molecular weight compounds may have resulted in some
instances. When peak III was incubated with LIP and
H2O2, a peak having a molecular weight
corresponding to that of a tetramer (molecular weight, 774) was
obtained (Fig. 7), although additional
techniques were not employed to determine its nature. It is possible
that even higher oligomers were formed, but because of lack of
solubility in tetrahydrofuran, lack of absorbance, or both, they went
unnoticed. These findings suggest that the products identified in this
study are but intermediates in the polymerization of FA, the limiting factors in the reaction being H2O2, and
probably also enzyme stability.
LIP-catalyzed oxidation of FA has been shown to follow
Michaelis-Menten kinetics.2
The Km and kcat values for FA
were dependent upon the concentration of H2O2
in the reaction mixture. The initial rate of FA oxidation at pH 3.5 reached saturation at an H2O2 concentration of
300 µM, and the corresponding Km and
kcat values for FA were calculated to be 116.8 µM and 41.7 s The results presented herein characterize products formed during
LIP-catalyzed oxidation of FA, including three dehydrodimers and two
novel trimers. Results also suggest that these products are further
oxidized, possibly leading to the formation of higher oligomers.
FA dehydrodimers produced both in vitro and in
vivo by plant peroxidases have been extensively characterized
(12-15). These regioisomers representing products of The three dehydrodimers produced during LIP-catalyzed oxidation of FA
included the benzofuran form of Interestingly, the second dehydrodimer identified in this study with
structure 1a (Fig. 3), resulting from The third dehydrodimer identified, appearing in peak III, is
also a product of Two novel trimers were identified from LIP-catalyzed oxidation of FA.
This is the first time that FA trimers have been identified from
peroxidase-catalyzed oxidation of FA. The following reaction sequence
is proposed for the formation of trimer 5 (Fig. 8). It would appear to result from the
addition of phenoxy radical 6 to the decarboxylated
-5' and
-
'coupling and two
trimers resulting from the addition of ferulic acid moieties to
decarboxylated derivatives of
-O-4'- and
-5'-coupled
dehydrodimers. This is the first time that trimers have been identified
from peroxidase-catalyzed oxidation of ferulic acid, and their
formation appears to be favored by decarboxylation of dehydrodimer
intermediates. After initial oxidation, the coupling reactions appear
to be determined by the chemistry of ferulic acid phenoxy radicals,
regardless of the enzyme and of whether the reaction is performed
in vitro or in vivo. This claim is supported by
our finding that horseradish peroxidase provides a similar product
profile. Furthermore, two of the dehydrodimers were the two products
obtained from laccase-catalyzed oxidation (Tatsumi, K. S., Freyer,
A., Minard, R. D., and Bollag, J.-M. (1994) Environ. Sci.
Technol. 28, 210-215), and the most abundant dehydrodimer is the
most prominent in grass cell walls (Ralph, J., Quideau, S., Grabber,
J. H., and Hatfield, R. D. (1994) J. Chem. Soc.
Perkin Trans. 1, 3485-3498). Our results also indicate that the
dehydrodimers and trimers are further oxidized by lignin peroxidase,
suggesting that they are only intermediates in the polymerization of
ferulic acid. The extent of polymerization appears to be dependent on
the ionization potential of formed intermediates, H2O2 concentration, and, probably, enzyme stability.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-
',
-5',
-O-4', 4-O-5', and 5-5' radical coupling (12-15). Such dehydrodimers, along with FA, are also
believed to act as nucleation sites in the lignification process,
coupling with lignin monomers (16). This clearly indicates that FA
dehydrodimers can be further oxidized. Indeed, certain dehydrodimers
formed from oxidative coupling of FA have been reported to be more
effective antioxidants than FA itself (17, 18). Their antioxidant
activity appears to be related to the existence of a full conjugation
system in the molecule. Nevertheless, the formation of higher molecular
weight oligomers and polymers from FA is undefined. A recent report
indicates that ferulate trimers and larger coupling products are formed
in cultured maize cells, where they are believed to tighten the cell
wall (19), and higher oligomers of FA have been implicated during
polymerization with horseradish peroxidase under weakly basic
conditions (pH 8) (11). However, their structures were not
characterized, leaving much speculation surrounding their formation.
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
1 cm
1 (23). LIP activity
(units/liter) was assayed according to Tien and Kirk (24). The
catalytic activity of the stock enzyme solution was calculated to be
1.96 units/nmol heme protein. The enzyme was extensively dialyzed
against double-distilled water before use.
1. 30 min after
addition of H2O2, reactions were either frozen at
70 °C and then freeze-dried or extracted with three volumes of
ethyl acetate and evaporated to dryness. The dried reaction mixtures
were then redissolved in 300 µl of tetrahydrofuran for gel permeation
analysis or 300 µl of 50% (v/v) aqueous methanol for reverse-phase analysis.
1.
1.
1 ensured an
elution profile similar to that of the analytical column. Oxidation
products were collected using a fraction collector (Gilson model 203)
and fractions deemed pure by reanalysis were freeze-dried and stored
under nitrogen gas in a cool, dark place.
1 from 150 °C to 300 °C and held for 20 min. The
injector and detector were set at 300 °C.
scale; 1H NMR
spectra were referenced to internal tetramethylsilane, and 13C NMR spectra were referenced to the solvent.
/2-shifted square sine bell in both dimensions.
1
cm
1. Stock solutions of FA were prepared in 95% ethanol
and checked using a calculated extinction coefficient of 14,700 M
1 cm
1 at 320 nm.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
70 °C after 30 min, freeze-dried, and
redissolved in tetrahydrofuran (Fig. 1).
The oxidation of FA (molecular weight, 194) by LIP-H1 as a function of
the obligatory co-factor H2O2 led to the
formation of peaks of molecular weight corresponding to dehydrodimers
(molecular weight, 386) and trimers (molecular weight, 579). Increasing
H2O2 concentration, which was added stepwise in
aliquots of 100 µM min
1 to prevent
H2O2-dependent enzyme inactivation,
resulted in a decrease in the peak corresponding to FA, followed by an
increase in the peaks corresponding to dehydrodimers and trimers.
Increasing H2O2 concentration above 100 µM resulted in a decrease in the intensity of the peak
corresponding to dehydrodimers. Because identical profiles were
obtained when the same mixtures were left to react for 24 h before
freezing, the limiting factors in the polymerization reaction were
H2O2, IP values of the intermediate products,
and, probably, enzyme stability.
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Fig. 1.
Gel permeation chromatography of FA after
oxidation by LIP, depicting the formation of dimers and trimers.
Reactions containing 1 µM LIP, 300 µM FA,
and various amounts of H2O2, added stepwise at
100 µM min 1, were frozen at
70 °C
after 30 min, freeze dried, and subsequently redissolved in 300 µl of
tetrahydrofuran prior to analysis. The H2O2
concentrations were 0 (thick solid line), 100 (thin
solid line), 200 (dotted line), and 400 (dashed
line) µM. Polystyrene standards of known molecular
weight were employed as standards.
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Fig. 2.
A typical chromatogram obtained from
reverse-phase HPLC analysis of the reaction products obtained from
large scale oxidation of FA by lignin peroxidase. The peak for FA
was added for reference and is indicated. The major peaks are labeled
I-IV.
H(A5) = 7.40, d(J(A2,A6) = 2.0Hz);
H(A5) = 6.84, d(J(A5,A6) = 8.0Hz);
H(A6) = 7.18, dd(J(A2,A6) = 2.0Hz,
J(A5,A6) = 8.0Hz). The protons of ring B were shifted to the
strong field:
H(B2) = 6.92, broad s;
H(B5, 6) = 6.78, broad s. The tri-substituted double bond was connected to ring A because the two-dimensional NOE
experiment showed that the only vinylic proton
(
H(A
) = 7.54, d(J(A
, B
) = 2.1Hz)) is
located near the A6 proton. According to the same two-dimensional NOE
experiment, the methoxy group with
H(OCH3) = 3.92 belongs to ring A, and
the methoxy group with
H(OCH3) = 3.87 to ring B. Two nonequivalent protons were found in the saturated
fragment of the molecule. The first proton, (
H(B
) = 4.56), was characterized by weak
hyperfine interaction with two protons, A
and B
: dd(J(A
,
B
) = 2.1Hz, J(B
, B
) = 2.8Hz). The peak of the second
proton (
H(B
) = 5.64) was split by interaction
with the B
proton: J(B
, B
) = 2.8Hz. The chemical shifts
and hyperfine structures of these two peaks suggest that the saturated
part consists of two CH fragments. One of them is bound to the A
carbon atom of the tri-substituted double bond and probably also to a
CO2 group, and the second to aromatic ring B and an oxygen
atom. All of the aforementioned results led us to the conclusion that
the first isolated product of FA dehydrodimerization has the structure
1a (Fig. 3). It belongs to the
group of so-called
-
'-dehydrodimerization products also
comprising 1b, 1c and 1d. The
structure of 1a was also confirmed by comparison of its NMR
parameters with data previously published for this compound (12). The
primarily formed
-
'-dehydrodimer 1b may undergo an
intramolecular Michael addition of a carboxylic group from one of the
two FA moieties to the double bond of the second, leading to
1a or 1c (Fig. 3). Compound 1c was
identified as one of the major components of peak III, as
will be seen further on.
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Fig. 3.
,
'-diFA dimers. The
dimers with structures 1a and 1c were identified
as products of lignin peroxidase-catalyzed oxidation of ferulic acid.
1a corresponded to peak I, whereas 1c
was one of the products belonging to peak III in Fig. 2.
GC-MS analysis of the peak I after silylation indicated the
formation of isomers 1b and 1d.
-
'-dehydrodimers of FA (M+ = 674) (probably of
1b and 1d; Fig. 3 and Ref. 12). The expected
tris(trimethylsilylated) product of furanone 1a
(M+ = 602) was not obtained, probably because of fast
disclosure of the lactone ring under the silylation conditions
(12).
H(A2) = 6.95, d(J(A2,A6) = 1.8Hz);
H(A5) = 6.78, d(J(A5,A6) = 8.2Hz);
H(A6) = 6.83, dd(J(A2,A6) = 1.8Hz,
J(A5,A6) = 8.2Hz). The protons of ring B were shifted to the low
field:
H(B2) = 7.17, broad singlet;
H(B6) = 7.23, broad singlet. According to the
results of the two-dimensional NOE experiment, the methoxy group with
H(OCH3) = 3.81 belongs to ring A, and
the methoxy group with
H(OCH3) = 3.91 to ring B. The di-substituted double bond is connected to ring B. The
following chemical shifts and hyperfine interaction patterns were found
for the
- and
-vinylic protons:
H(B
) = 7.62, d(J(B
, B
) = 15.9Hz);
H(B
) = 7.62, d(J(B
, B
) = 15.9Hz). The saturated part of the
molecule consists of two CH fragments. One of them is bound to the
aromatic ring A and an oxygen atom (
C(A
) = 89.21), and the second to the aromatic ring
B(
C(A
) = 57.08). The proton of the CH group
connected to ring A was characterized by a chemical shift
H(A
) = 6.02 and by relatively strong hyperfine interactions with proton A
(d(J(A
, A
) = 7.7Hz) of the
second CH (
H(A
) = 4.27) group bound to ring B. Results of the two-dimensional NOE experiment confirmed the assignment:
proton A
is located near protons A2 and A6; proton A
is close to
B2 and B6. The NMR data led us to conclude that the second isolated
product of FA dehydrodimerization, peak II, has the
structure 2a (Fig. 4). It
belongs to the group of
-5'-dehydrodimers along with structure
2b in Fig. 4. The primarily formed
-5'-dehydrodimer 2b may undergo intramolecular addition of the phenolic hydroxyl group from ring B to the double bond connected to the B5
carbon atom leading to 2a (Fig. 4).
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Fig. 4.
,5'-diFA dimers. The dimer
with structure 2a was identified as a product of lignin
peroxidase-catalyzed oxidation of ferulic acid, corresponding to peak
II in Fig. 2. GC-MS analysis of the peak II after
silylation, indicated the formation of isomers 2b and
2c.
H(A2) =
H(B2) = 6.96,
H(A5) =
H(B5) = 6.863,
H(A6) =
H(B6) = 6.857. Two
pairs of equivalent CH fragments were found in the saturated part of
the molecule. According to the chemical shift
H(A
) =
H(B
) = 5.80, dd(J(A
, A
) = 1.1Hz, J(A
, B
) = 1.1Hz), one of the
two CH fragments is connected to the aromatic moiety and an oxygen
atom. The second CH fragment with
H(A
) =
H(B
) = 3.99, dd(J(A
, A
) = 1.1Hz,
J(A
, B
) = 1.1Hz) is connected to a carboxylic group. The
results of the one-dimensional NOE experiments (Table
I) led us to the conclusion that (i) the
-proton is spatially close to the protons in positions 2 and 6 of
the aromatic ring and (ii) the
-proton is located near the proton in
position 2. These facts enabled us to assign the dilactone structure
1c to the dehydrodimer found in peak III.
NOE of the dimer with structure 1c
C,
H, and splitting pattern are
presented in Tables II and
III. The assignment of the signals in
1H and 13C NMR spectra is based on the analysis
of the splitting pattern, COSY spectra, and
13C-1H correlation spectra. The data presented
in Tables II and III, along with the results of the one-dimensional NOE
experiment (Table IV), led us to conclude
that: (i) the double bond with the chemical shift of the vinylic proton
H = 6.37 is connected to ring C, because this proton is located
near proton C2; (ii) the double bond with the chemical shift of the
vinylic proton
H = 6.33 is connected to ring A, because this
proton is spatially close to proton A2; (iii) aromatic fragments A and
C are connected to the CH group of the saturated part with
H = 6.01, because this proton is located near both protons A5 and C5; (iv)
the aromatic fragments A and C are connected to the aforementioned CH
group through the phenolic oxygen atoms, because the chemical shift of
the corresponding carbon atom
C = 106.29 is characteristic of
the acetal carbon (26); (v) the second CH group (
H = 4.95) is
connected to aromatic fragment B, because the corresponding proton is
located near protons B2 and B6; and (vi) the second CH group is
also connected to an oxygen atom, because the chemical shift of the
corresponding carbon atom, (
C = 75.80) resembles those of
alcohol carbons (26).
1H chemical shifts for CD3OD solutions of the trimer
with structure 3
13C chemical shifts for CD3OD solutions of the trimer
with structure 3
NOE of the trimer with structure 3
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Fig. 5.
The trimer with structure 3, identified as
one of the products found in peak III in Fig. 2.
) of the saturated fragment through the
A1 atom. This CH group is connected to the other CH group (A
) and
according to
H(A
) and
C(A
), it is
also connected to an oxygen atom. The second CH group is connected to
position C5 of the tetra-substituted ring C and according to
H(A
) and
C(A
) also to an oxygen
atom. Obviously, the tri-substituted aromatic ring B is connected to one of the two CH fragments through the O-4 oxygen atom.
1H and 13C chemical shifts for CD3OD solutions
of the trimer with structure 6
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Fig. 6.
The trimer with structure 5, identified as
one of the products found in peak IV in Fig. 2 and the
-O-4-diFA 4 obtained upon GC-MS
analysis of peak IV after silylation.
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Fig. 7.
Gel permeation chromatography of isolated
Peak III after further oxidation by LIP, indicating the formation of
higher oligomers. The reaction contained 1 µM LIP
and 800 µM H2O2 (dotted
line), added stepwise in aliquots of 200 µM
min 1. The control was without
H2O2 (solid line).
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
1, respectively, indicating
that it is a very reactive substrate.
',
-5',
-O-4', 4-O-5', and 5-5' radical
coupling, are found in plant cell walls, where they cross-link
polymers, providing strength and rigidity.
-5'-diFA 2a (Fig. 4),
which has been identified in the cell walls of several plants (12-14).
In grass cell walls, it has been identified as the most abundant
dehydrodimer (12). Interestingly, our studies also indicate it to be
the major dehydrodimer formed in vitro during LIP-catalyzed
oxidation of FA. The peak corresponding to this structure was always
the major component, as obtained by analytical reverse-phase HPLC (Fig.
2). During preparative chromatography as well, a large amount of this
compound was isolated relative to the other compounds. Because this was
also one of two dehydrodimers identified during laccase-catalyzed
oxidation of FA (27), these findings would suggest that the chemistry
of FA favors the formation of this dehydrodimer irrespective of which
enzyme is employed to oxidize the substrate and irrespective of whether
the reaction is performed in vitro or in vivo.
GC-MS analysis of the benzodihydrofuran form of
-5'-diFA,
revealed the noncyclic form, with structure 2b, along with
the decarboxylation product of 2b, structure 2c
(Fig. 4).
-
' coupling, was
the other dehydrodimer identified during laccase-catalyzed oxidation of
FA (27). In other words, the two dehydrodimers identified from
laccase-catalyzed oxidation of FA were identified during LIP-catalyzed
oxidation. The
-lactone structure 1a results from
intramolecular rearrangement of the primary formed
-
'-dehydrodimer 1b (Fig. 3).
-
' coupling having the structure 1c, which results from intramolecular rearrangement of structures 1a and 1b (Fig. 3). Because two of the three
dehydrodimers identified result from
-
' radical coupling, it is
possible that this mechanism of radical coupling is also favored, along
with
-5' radical coupling during oxidation. Similarly, the two
products obtained during laccase-catalyzed oxidation were a result of
-
' and
-5' radical coupling (27). Although no dehydrodimers
resulting from
-O-4' radical coupling were identified,
elucidation of the structure of trimer 3 indicates that such
dehydrodimers were formed, as discussed further on.
-5'-diFA 2c that was identified as one of the major
dehydrodimers (Fig. 4). Two free radicals, 7a and
7b, are formed in the first step of the reaction. According
to AM1 semi-empirical quantum chemical calculations, 7a is
about 3 Kcal/mol less stable than 7b, but its ionization
potential is 0.32 eV (about 7 Kcal/mol) lower than the ionization
potential of 7b. This means that carbocation 7c
(the result of the one-electron oxidation of the less stable radical
7a) will be the major intermediate of the reaction if the
formation of 7b is reversible. Intramolecular addition of
the phenolic hydroxyl group to the positively charged carbon atom in
7c leads to trimer 5.
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Fig. 8.
The proposed mechanism for the formation of
the trimer with structure 5.
The trimer with structure 3 also contains a decarboxylated
moiety, and its formation can be explained by the addition of phenoxy
radical 6 to the decarboxylated derivative of -O-4'-diFA 8 (Fig.
9). Two free radicals, 9a and 9b, are formed in this step of the reaction. According to
the AM1 semi-empirical quantum chemical calculations, 9a is
about 15 Kcal/mol more stable than 9b and can be considered a major intermediate. One-electron oxidation of 9a leads to
the corresponding carbocation 9c. The latter then reacts with a molecule of water, giving trimer 3. Interestingly,
-O-4'-diFA and its decarboxylated derivative 8 were not identified, indicating that they were present only as minor
peaks. Because trimer 3 was considered one of the major
products, this indicates that once formed, the decarboxylated
derivative of
-O-4'-diFA preferably undergoes coupling
with FA radicals to form trimers. The mechanism governing the formation
of decarboxylated
-5'- and
-O-4'-diFA is not
clear.
|
It has been suggested that different peroxidases catalyze the formation
of FA oxidation products in different ways (11). However, this may be a
misconception resulting from incomplete characterization of all of the
reaction products. Our findings would suggest that product formation is
independent of the enzyme employed. When we conducted the oxidation of
FA with horseradish peroxidase, products were similar to those obtained
for LIP, as indicated by similar reverse-phase HPLC profiles and
partial identification (data not shown). Moreover, the main
dehydrodimers obtained are also predominant in the cell walls of plants
(12-15), and two of them have also been identified from
laccase-catalyzed oxidation (27). Although the mechanism of phenol
oxidation is different for laccases and peroxidases, both result in the
formation of a phenoxy radical. All this would suggest that after
formation, the phenoxy radicals vacate the active site of the enzyme,
and the subsequent coupling of radicals to form dimeric and oligomeric products is nonenzymatic and is probably determined by the chemistry of
FA phenoxy radicals. Therefore, the products identified here are
certainly not exclusive to LIP-catalyzed oxidation. It would appear,
however, that such dehydrodimers and trimers are but intermediates in
the oxidation of FA. Not only did the intensity of the peaks corresponding to these products decrease with increasing concentrations of H2O2 (added stepwise to prevent
H2O2-dependent enzyme
inactivation), they also decreased with further incubation of isolated
dehydrodimers and trimers with LIP, indicating that they were further
oxidized, possibly resulting in the formation of higher oligomers in
certain instances (Fig. 7). The high redox potential of LIP enables it to oxidize substrates with high IP values that other peroxidases cannot
oxidize (2, 3). Therefore, if the ionization potential of primary
products determines the extent of polymerization (4), then
LIP-catalyzed oxidation may provide the highest degree of polymerization. Work is currently being undertaken to clarify this and
to further understand the factors governing enzymatic polymerization of phenolics.
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FOOTNOTES |
---|
* This research was supported by the Israel Science Foundation.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
To whom correspondence should be addressed. Tel.:
972-4-829-4962; Fax: 972-4-8228898; E-mail:
carlosd@techunix.technion.ac.il.
Published, JBC Papers in Press, February 13, 2001, DOI 10.1074/jbc.M009785200
2 Ward, G., Hadar, Y., and Dosoretz, C. G. (2001) Enzyme Microb. Technol., in press.
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ABBREVIATIONS |
---|
The abbreviations used are: LIP, lignin peroxidase; IP, ionization potential; FA, ferulic acid; HPLC, high pressure liquid chromatography; NOE, nuclear Overhauser effect; GC-MS, gas chromatography-mass spectroscopy..
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REFERENCES |
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---|
1. | Hatakka, A. (1994) FEMS Microbiol. Rev. 13, 125-135[CrossRef] |
2. | Kersten, D. J., Kalyanaraman, B., Hammel, K. E., Reinhammer, B., and Kent Kirk, T. (1990) Biochem. J. 268, 475-480[Medline] [Order article via Infotrieve] |
3. |
Hammel, K. E.,
Kalyanaraman, B.,
and Kent Kirk, T.
(1996)
J. Biol. Chem.
261,
16948-16952 |
4. | ten Have, R., Rietjens, I. M. C. M., Hartmans, S., Swarts, H. J., and Field, J. A. (1998) FEBS Lett. 430, 390-392[CrossRef][Medline] [Order article via Infotrieve] |
5. | Colonna, S., Gaggero, N., Richelmi, C., and Pasta, P. (1999) Trends Biotechnol. 17, 163-168[CrossRef][Medline] [Order article via Infotrieve] |
6. | May, S. W. (1999) Curr. Opin. Biotechnol. 10, 370-375[CrossRef][Medline] [Order article via Infotrieve] |
7. | Adam, W., Lazarus, M., Saha-Möller, C. R., Weichold, O., Hoch, U., Häring, D., and Schreier, P. (1999) Adv. Biochem. Eng. 63, 74-108 |
8. | Aitken, M. D. (1993) Chem. Eng. J. B49-B58 |
9. | Klibanov, A. M., Tu, T.-M., and Scott, K. P. (1983) Science 221, 259-261 |
10. | Bumpus, J. A., Tien, M., Wright, D., and Aust, S. D. (1985) Science 228, 1434-1436[Medline] [Order article via Infotrieve] |
11. | Rosazza, J. P. N., Huang, Z., Dostal, L., Volm, T., and Rousseau, B. (1995) J. Ind. Microbiol. 15, 457-471[Medline] [Order article via Infotrieve] |
12. | Ralph, J., Quideau, S., Grabber, J. H., and Hatfield, R. D. (1994) J. Chem. Soc. Perkin Trans. 1, 3485-3498 |
13. | Micard, V., Grabber, H., Ralph, J., Renard, C. M. G. C., and Thibault, J.-F. (1997) Phytochemistry 44, 1365-1368[CrossRef] |
14. | Waldron, K. W., Parr, A. J., Ng, A., and Ralph, J. (1996) Phytochem. Anal. 7, 305-312[CrossRef] |
15. | Bartolome, B., Faulds, C. B., Kroon, P. A., Waldron, K. W., Gilbert, H. J., Hazlewood, G., and Williamson, G. (1997) Appl. Environ. Microbiol. 63, 208-212[Abstract] |
16. | Ralph, J., Hatfield, R., and Grabber, J. (1997) Polyphénols Actualités 17, 4-6 |
17. | Garcia-Conesa, M. T., Plumb, G. W., Kroon, P. A., Wallace, G., and Williamson, G. (1997) Redox Rep. 3, 239-244 |
18. | Garcia-Conesa, M. T., Wilson, P. D., Plumb, G. W., and Williamson, G. (1999) J. Sci. Food Agri. 79, 379-384 |
19. | Fry, S. C., Willis, S. C., and Paterson, A. E. J. (2000) Planta 211, 679-692[CrossRef][Medline] [Order article via Infotrieve] |
20. | Kroon, P. A., and Williamson, G. (1999) J. Sci. Food Agric. 79, 355-361[CrossRef] |
21. | Rothschild, N., Hadar, Y., and Dosoretz, C. G. (1997) Appl. Environ. Microbiol. 63, 857-861[Abstract] |
22. | Kirk, T. K., Croan, A., Tien, M., Murtagh, K. E., and Farrell, R. L. (1986) Enzyme Microb. Technol. 8, 27-32[CrossRef] |
23. |
Tien, M.,
Kirk, T. K.,
Bull, C.,
and Fee, J. A.
(1986)
J. Biol. Chem.
261,
1687-1693 |
24. | Tien, M., and Kirk, T. K. (1988) Methods Enzymol. 161, 238-249 |
25. | Frisch, M. J., Trucks, G. W., Schlegel, H. B., Gill, P. M. W., Johnson, B. G., Robb, M. A., Cheeseman, J. R., Keith, T., Petersson, G. A., Montgomery, J. A., Raghavachari, K., Al-Laham, M. A., Zakrzewski, V. G., Ortiz, J. V., Foresman, J. B., Cioslowski, J., Stefanov, B. B., Nanayakkara, A., Challacombe, M., Peng, C. Y., Ayala, P. Y., Chen, W., Wong, M. W., Andres, J. L., Replogle, E. S., Gomperts, R., Martin, R. L., Fox, D. J., Binkley, J. S., Defrees, D. J., Baker, J., Stewart, J. P., Head-Gordon, M., Gonzalez, C., and Pople, J. A. (1995) Gaussian, Vol. 94, Revision E.2., Gaussian Inc., Pittsburgh, PA |
26. | Kalinowski, H.-O., Berger, S., and Braun, S. (1988) Carbon-13 NMR Spectroscopy , John Wiley & Sons, Inc., Chichester, UK |
27. | Tatsumi, K. S., Freyer, A., Minard, R. D., and Bollag, J.-M. (1994) Environ. Sci. Technol. 28, 210-215 |