Initial Steps of Ferulic Acid Polymerization by Lignin Peroxidase*

Gary WardDagger , Yitzhak Hadar§, Itzhak Bilkis, Leonid Konstantinovsky||, and Carlos G. Dosoretz**DaggerDagger

From the Dagger  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


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The major products of the initial steps of ferulic acid polymerization by lignin peroxidase included three dehydrodimers resulting from beta -5' and beta -beta 'coupling and two trimers resulting from the addition of ferulic acid moieties to decarboxylated derivatives of beta -O-4'- and beta -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

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 beta -beta ', beta -5', beta -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.

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).

    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

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-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.

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-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.

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-1.

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-1.

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-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.

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-1 from 150 °C to 300 °C and held for 20 min. The injector and detector were set at 300 °C.

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 delta  scale; 1H NMR spectra were referenced to internal tetramethylsilane, and 13C NMR spectra were referenced to the solvent.

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 pi /2-shifted square sine bell in both dimensions.

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-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

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 -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.


View larger version (18K):
[in this window]
[in a new window]
 
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.

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.


View larger version (15K):
[in this window]
[in a new window]
 
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.

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: delta H(A5) = 7.40, d(J(A2,A6) = 2.0Hz); delta H(A5) = 6.84, d(J(A5,A6) = 8.0Hz); delta 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: delta H(B2) = 6.92, broad s; delta 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 (delta H(Aalpha ) = 7.54, d(J(Aalpha , Bbeta ) = 2.1Hz)) is located near the A6 proton. According to the same two-dimensional NOE experiment, the methoxy group with delta H(OCH3) = 3.92 belongs to ring A, and the methoxy group with delta H(OCH3) = 3.87 to ring B. Two nonequivalent protons were found in the saturated fragment of the molecule. The first proton, (delta H(Bbeta ) = 4.56), was characterized by weak hyperfine interaction with two protons, Aalpha and Balpha : dd(J(Aalpha , Bbeta ) = 2.1Hz, J(Balpha , Bbeta ) = 2.8Hz). The peak of the second proton (delta H(Balpha ) = 5.64) was split by interaction with the Bbeta proton: J(Balpha , Bbeta ) = 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 Abeta 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 beta -beta '-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 beta -beta '-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.


View larger version (17K):
[in this window]
[in a new window]
 
Fig. 3.   beta ,beta '-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.

GC-MS analysis of peak I after silylation indicated the formation of two isomeric tetrakis(trimethylsilylated) beta -beta '-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).

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: delta H(A2) = 6.95, d(J(A2,A6) = 1.8Hz); delta H(A5) = 6.78, d(J(A5,A6) = 8.2Hz); delta 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: delta H(B2) = 7.17, broad singlet; delta H(B6) = 7.23, broad singlet. According to the results of the two-dimensional NOE experiment, the methoxy group with delta H(OCH3) = 3.81 belongs to ring A, and the methoxy group with delta 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 alpha - and beta -vinylic protons: delta H(Balpha ) = 7.62, d(J(Balpha , Bbeta ) = 15.9Hz); delta H(Bbeta ) = 7.62, d(J(Balpha , Bbeta ) = 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 (delta C(Aalpha ) = 89.21), and the second to the aromatic ring B(delta C(Abeta ) = 57.08). The proton of the CH group connected to ring A was characterized by a chemical shift delta H(Aalpha ) = 6.02 and by relatively strong hyperfine interactions with proton Abeta (d(J(Aalpha , Abeta ) = 7.7Hz) of the second CH (delta H(Abeta ) = 4.27) group bound to ring B. Results of the two-dimensional NOE experiment confirmed the assignment: proton Aalpha is located near protons A2 and A6; proton Abeta 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 beta -5'-dehydrodimers along with structure 2b in Fig. 4. The primarily formed beta -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).


View larger version (11K):
[in this window]
[in a new window]
 
Fig. 4.   beta ,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.

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: delta H(A2) = delta H(B2) = 6.96, delta H(A5) = delta H(B5) = 6.863, delta H(A6) = delta H(B6) = 6.857. Two pairs of equivalent CH fragments were found in the saturated part of the molecule. According to the chemical shift delta H(Aalpha ) = delta H(Balpha ) = 5.80, dd(J(Aalpha , Abeta ) = 1.1Hz, J(Aalpha , Bbeta ) = 1.1Hz), one of the two CH fragments is connected to the aromatic moiety and an oxygen atom. The second CH fragment with delta H(Abeta ) = delta H(Bbeta ) = 3.99, dd(J(Aalpha , Abeta ) = 1.1Hz, J(Abeta , Balpha ) = 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 alpha -proton is spatially close to the protons in positions 2 and 6 of the aromatic ring and (ii) the beta -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.

                              
View this table:
[in this window]
[in a new window]
 
Table I
NOE of the dimer with structure 1c

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 delta C, delta 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 delta 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 delta 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 delta 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 delta C = 106.29 is characteristic of the acetal carbon (26); (v) the second CH group (delta 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, (delta C = 75.80) resembles those of alcohol carbons (26).

                              
View this table:
[in this window]
[in a new window]
 
Table II
1H chemical shifts for CD3OD solutions of the trimer with structure 3

                              
View this table:
[in this window]
[in a new window]
 
Table III
13C chemical shifts for CD3OD solutions of the trimer with structure 3

                              
View this table:
[in this window]
[in a new window]
 
Table IV
NOE of the trimer with structure 3

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.


View larger version (15K):
[in this window]
[in a new window]
 
Fig. 5.   The trimer with structure 3, identified as one of the products found in peak III in Fig. 2.

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 (Aalpha ) of the saturated fragment through the A1 atom. This CH group is connected to the other CH group (Abeta ) and according to delta H(Aalpha ) and delta C(Aalpha ), 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 delta H(Abeta ) and delta C(Abeta ) 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.

                              
View this table:
[in this window]
[in a new window]
 
Table V
1H and 13C chemical shifts for CD3OD solutions of the trimer with structure 6

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).


View larger version (12K):
[in this window]
[in a new window]
 
Fig. 6.   The trimer with structure 5, identified as one of the products found in peak IV in Fig. 2 and the beta -O-4-diFA 4 obtained upon GC-MS analysis of peak IV after silylation.

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.


View larger version (14K):
[in this window]
[in a new window]
 
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

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-1, respectively, indicating that it is a very reactive substrate.

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 beta -beta ', beta -5', beta -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.

The three dehydrodimers produced during LIP-catalyzed oxidation of FA included the benzofuran form of beta -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 beta -5'-diFA, revealed the noncyclic form, with structure 2b, along with the decarboxylation product of 2b, structure 2c (Fig. 4).

Interestingly, the second dehydrodimer identified in this study with structure 1a (Fig. 3), resulting from beta -beta ' 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 gamma -lactone structure 1a results from intramolecular rearrangement of the primary formed beta -beta '-dehydrodimer 1b (Fig. 3).

The third dehydrodimer identified, appearing in peak III, is also a product of beta -beta ' 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 beta -beta ' radical coupling, it is possible that this mechanism of radical coupling is also favored, along with beta -5' radical coupling during oxidation. Similarly, the two products obtained during laccase-catalyzed oxidation were a result of beta -beta ' and beta -5' radical coupling (27). Although no dehydrodimers resulting from beta -O-4' radical coupling were identified, elucidation of the structure of trimer 3 indicates that such dehydrodimers were formed, as discussed further on.

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 beta -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.


View larger version (10K):
[in this window]
[in a new window]
 
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 beta -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, beta -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 beta -O-4'-diFA preferably undergoes coupling with FA radicals to form trimers. The mechanism governing the formation of decarboxylated beta -5'- and beta -O-4'-diFA is not clear.


View larger version (13K):
[in this window]
[in a new window]
 
Fig. 9.   The proposed mechanism for the formation of the trimer with structure 3.

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.

    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.

Dagger Dagger 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.

    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..

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

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[Abstract/Free Full Text]
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[Abstract/Free Full Text]
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


Copyright © 2001 by The American Society for Biochemistry and Molecular Biology, Inc.