Initial Steps of Ferulic Acid Polymerization by Lignin Peroxidase
2001; Elsevier BV; Volume: 276; Issue: 22 Linguagem: Inglês
10.1074/jbc.m009785200
ISSN1083-351X
AutoresGary E. Ward, Yitzhak Hadar, Itzhak Bilkis, Leonid Konstantinovsky, Carlos G. Dosoretz,
Tópico(s)Lignin and Wood Chemistry
ResumoThe major products of the initial steps of ferulic acid polymerization by lignin peroxidase included three dehydrodimers resulting from β-5′ and β-β′coupling and two trimers resulting from the addition of ferulic acid moieties to decarboxylated derivatives of β-O-4′- and β-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 performedin 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. The major products of the initial steps of ferulic acid polymerization by lignin peroxidase included three dehydrodimers resulting from β-5′ and β-β′coupling and two trimers resulting from the addition of ferulic acid moieties to decarboxylated derivatives of β-O-4′- and β-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 performedin 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. lignin peroxidase ionization potential ferulic acid high pressure liquid chromatography nuclear Overhauser effect gas chromatography-mass spectroscopy. Lignin peroxidase (LIP)1is considered to be one of the most important enzymes of the extracellular lignin degradation system secreted by the white rot fungus, Phanerochaete chrysosporium (1Hatakka A. FEMS Microbiol. Rev. 1994; 13: 125-135Crossref Scopus (895) Google Scholar). 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 (2Kersten D.J. Kalyanaraman B. Hammel K.E. Reinhammer B. Kent Kirk T. Biochem. J. 1990; 268: 475-480Crossref PubMed Scopus (312) Google Scholar, 3Hammel K.E. Kalyanaraman B. Kent Kirk T. J. Biol. Chem. 1996; 261: 16948-16952Abstract Full Text PDF Google Scholar). The high redox potential enables LIP to oxidize aromatic compounds with calculated ionization potential (IP) values of up to 9.0 eV (4ten Have R. Rietjens I.M.C.M. Hartmans S. Swarts H.J. Field J.A. FEBS Lett. 1998; 430: 390-392Crossref PubMed Scopus (24) Google Scholar). This has striking implications when considering the potential applications of peroxidases for useful biotransformations (5Colonna S. Gaggero N. Richelmi C. Pasta P. Trends Biotechnol. 1999; 17: 163-168Abstract Full Text Full Text PDF PubMed Scopus (225) Google Scholar, 6May S.W. Curr. Opin. Biotechnol. 1999; 10: 370-375Crossref PubMed Scopus (93) Google Scholar, 7Adam W. Lazarus M. Saha-Möller C.R. Weichold O. Hoch U. Häring D. Schreier P. Adv. Biochem. Eng. 1999; 63: 74-108Google Scholar). 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 (6May S.W. Curr. Opin. Biotechnol. 1999; 10: 370-375Crossref PubMed Scopus (93) Google Scholar, 7Adam W. Lazarus M. Saha-Möller C.R. Weichold O. Hoch U. Häring D. Schreier P. Adv. Biochem. Eng. 1999; 63: 74-108Google Scholar), as well as for the treatment of wastewater streams polluted with toxic phenols (8Aitken, M. D. (1993) Chem. Eng. J. B49–B58Google Scholar, 9Klibanov A.M. Tu T.-M. Scott K.P. Science. 1983; 221: 259-261Crossref PubMed Scopus (429) Google Scholar, 10Bumpus J.A. Tien M. Wright D. Aust S.D. Science. 1985; 228: 1434-1436Crossref PubMed Scopus (750) Google Scholar). Ferulic acid (FA), which is an extremely abundant and widespread cinnamic acid derivative (11Rosazza J.P.N. Huang Z. Dostal L. Volm T. Rousseau B. J. Ind. Microbiol. 1995; 15: 457-471Crossref PubMed Scopus (277) Google Scholar), 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 β-β′, β-5′, β-O-4′, 4-O-5′, and 5–5′ radical coupling (12Ralph J. Quideau S. Grabber J.H. Hatfield R.D. J. Chem. Soc. Perkin Trans. 1994; 1: 3485-3498Crossref Scopus (428) Google Scholar, 13Micard V. Grabber H. Ralph J. Renard C.M.G.C. Thibault J.-F. Phytochemistry. 1997; 44: 1365-1368Crossref Scopus (66) Google Scholar, 14Waldron K.W. Parr A.J. Ng A. Ralph J. Phytochem. Anal. 1996; 7: 305-312Crossref Scopus (266) Google Scholar, 15Bartolome B. Faulds C.B. Kroon P.A. Waldron K.W. Gilbert H.J. Hazlewood G. Williamson G. Appl. Environ. Microbiol. 1997; 63: 208-212Crossref PubMed Google Scholar). Such dehydrodimers, along with FA, are also believed to act as nucleation sites in the lignification process, coupling with lignin monomers (16Ralph J. Hatfield R. Grabber J. Polyphénols Actualités. 1997; 17: 4-6Google Scholar). 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 (17Garcia-Conesa M.T. Plumb G.W. Kroon P.A. Wallace G. Williamson G. Redox Rep. 1997; 3: 239-244Crossref PubMed Scopus (44) Google Scholar, 18Garcia-Conesa M.T. Wilson P.D. Plumb G.W. Williamson G. J. Sci. Food Agri. 1999; 79: 379-384Crossref Scopus (69) Google Scholar). 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 (19Fry S.C. Willis S.C. Paterson A.E.J. Planta. 2000; 211: 679-692Crossref PubMed Scopus (155) Google Scholar), and higher oligomers of FA have been implicated during polymerization with horseradish peroxidase under weakly basic conditions (pH 8) (11Rosazza J.P.N. Huang Z. Dostal L. Volm T. Rousseau B. J. Ind. Microbiol. 1995; 15: 457-471Crossref PubMed Scopus (277) Google Scholar). 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 (20Kroon P.A. Williamson G. J. Sci. Food Agric. 1999; 79: 355-361Crossref Scopus (246) Google Scholar). LIP isoenzyme H1 (LIP-H1) was produced from high nitrogen cultures of P. chrysosporium Burds BKM-F-1767 as described previously (21Rothschild N. Hadar Y. Dosoretz C.G. Appl. Environ. Microbiol. 1997; 63: 857-861Crossref PubMed Google Scholar). The enzyme was purified in two steps by MonoQ HPLC, first using a 0.01–1 m sodium acetate gradient at pH 6.0 (22Kirk T.K. Croan A. Tien M. Murtagh K.E. Farrell R.L. Enzyme Microb. Technol. 1986; 8: 27-32Crossref Scopus (398) Google Scholar) and then by employing a similar gradient at pH 4.7, equivalent to the pI value of H1. The purified enzyme had an Reinheitszahl (A 409/280) value > 4.0. LIP concentration was determined at 409 nm using an extinction coefficient of 169 mm−1 cm−1 (23Tien M. Kirk T.K. Bull C. Fee J.A. J. Biol. Chem. 1986; 261: 1687-1693Abstract Full Text PDF PubMed Google Scholar). LIP activity (units/liter) was assayed according to Tien and Kirk (24Tien M. Kirk T.K. Methods Enzymol. 1988; 161: 238-249Crossref Scopus (1296) Google Scholar). 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. 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 mmH2O2, 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 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. 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. 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 (14Waldron K.W. Parr A.J. Ng A. Ralph J. Phytochem. Anal. 1996; 7: 305-312Crossref Scopus (266) Google Scholar), 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. 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. 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. 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 δ scale; 1H NMR spectra were referenced to internal tetramethylsilane, and13C NMR spectra were referenced to the solvent. 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 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 π/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. 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 (25Frisch 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. Pople J.A. Gaussian.in: Modern Nutrition in Health and Disease. Revision E.2. Vol. 94. Pittsburgh,Gaussian Inc., PA1995Google Scholar) and Spartan 5.1 programs. 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−1cm−1. Stock solutions of FA were prepared in 95% ethanol and checked using a calculated extinction coefficient of 14,700m−1 cm−1 at 320 nm. Generation of phenoxy radicals from FA by peroxidases can theoretically lead to a plethora of polymerization products (11Rosazza J.P.N. Huang Z. Dostal L. Volm T. Rousseau B. J. Ind. Microbiol. 1995; 15: 457-471Crossref PubMed Scopus (277) Google Scholar). 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. 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. 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 by1H 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: δH(A5) = 7.40, d(J(A2,A6) = 2.0Hz); δH(A5) = 6.84, d(J(A5,A6) = 8.0Hz); δH(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: δH(B2) = 6.92, broad s; δH(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 (δH(Aα) = 7.54, d(J(Aα, Bβ) = 2.1Hz)) is located near the A6 proton. According to the same two-dimensional NOE experiment, the methoxy group with δH(OCH3) = 3.92 belongs to ring A, and the methoxy group with δH(OCH3) = 3.87 to ring B. Two nonequivalent protons were found in the saturated fragment of the molecule. The first proton, (δH(Bβ) = 4.56), was characterized by weak hyperfine interaction with two protons, Aα and Bα: dd(J(Aα, Bβ) = 2.1Hz, J(Bα, Bβ) = 2.8Hz). The peak of the second proton (δH(Bα) = 5.64) was split by interaction with the Bβ proton: J(Bα, Bβ) = 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 Aβ 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 structure1a (Fig. 3). It belongs to the group of so-called β-β′-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 (12Ralph J. Quideau S. Grabber J.H. Hatfield R.D. J. Chem. Soc. Perkin Trans. 1994; 1: 3485-3498Crossref Scopus (428) Google Scholar). The primarily formed β-β′-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 to1a or 1c (Fig. 3). Compound 1c was identified as one of the major components of peak III, as will be seen further on. GC-MS analysis of peak I after silylation indicated the formation of two isomeric tetrakis(trimethylsilylated) β-β′-dehydrodimers of FA (M+ = 674) (probably of1b and 1d; Fig. 3 and Ref. 12Ralph J. Quideau S. Grabber J.H. Hatfield R.D. J. Chem. Soc. Perkin Trans. 1994; 1: 3485-3498Crossref Scopus (428) Google Scholar). 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 (12Ralph J. Quideau S. Grabber J.H. Hatfield R.D. J. Chem. Soc. Perkin Trans. 1994; 1: 3485-3498Crossref Scopus (428) Google Scholar). 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: δH(A2) = 6.95, d(J(A2,A6) = 1.8Hz); δH(A5) = 6.78, d(J(A5,A6) = 8.2Hz); δH(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: δH(B2) = 7.17, broad singlet; δH(B6) = 7.23, broad singlet. According to the results of the two-dimensional NOE experiment, the methoxy group with δH(OCH3) = 3.81 belongs to ring A, and the methoxy group with δH(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 α- and β-vinylic protons: δH(Bα) = 7.62, d(J(Bα, Bβ) = 15.9Hz); δH(Bβ) = 7.62, d(J(Bα, Bβ) = 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 (δC(Aα) = 89.21), and the second to the aromatic ring B(δC(Aβ) = 57.08). The proton of the CH group connected to ring A was characterized by a chemical shift δH(Aα) = 6.02 and by relatively strong hyperfine interactions with proton Aβ (d(J(Aα, Aβ) = 7.7Hz) of the second CH (δH(Aβ) = 4.27) group bound to ring B. Results of the two-dimensional NOE experiment confirmed the assignment: proton Aα is located near protons A2 and A6; proton Aβ 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 β-5′-dehydrodimers along with structure2b in Fig. 4. The primarily formed β-5′-dehydrodimer2b 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). 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 2aduring the silylation procedure has been reported; Ref. 12Ralph J. Quideau S. Grabber J.H. Hatfield R.D. J. Chem. Soc. Perkin Trans. 1994; 1: 3485-3498Crossref Scopus (428) Google Scholar), and the 558 peak to the tris(trimethylsilylated) derivative of the decarboxylation product of 2b (see structure 2cand Ref. 12Ralph J. Quideau S. Grabber J.H. Hatfield R.D. J. Chem. Soc. Perkin Trans. 1994; 1: 3485-3498Crossref Scopus (428) Google Scholar). 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: δH(A2) = δH(B2) = 6.96, δH(A5) = δH(B5) = 6.863, δH(A6) = δH(B6) = 6.857. Two pairs of equivalent CH fragments were found in the saturated part of the molecule. According to the chemical shift δH(Aα) = δH(Bα) = 5.80, dd(J(Aα, Aβ) = 1.1Hz, J(Aα, Bβ) = 1.1Hz), one of the two CH fragments is connected to the aromatic moiety and an oxygen atom. The second CH fragment with δH(Aβ) = δH(Bβ) = 3.99, dd(J(Aα, Aβ) = 1.1Hz, J(Aβ, Bα) = 1.1Hz) is connected to a carboxylic group. The results of the one-dimensional NOE experiments (TableI) led us to the conclusion that (i) the α-proton is spatially close to the protons in positions 2 and 6 of the aromatic ring and (ii) the β-proton is located near the proton in position 2. These facts enabled us to assign the dilactone structure1c to the dehydrodimer found in peak III.Table INOE of the dimer with structure 1cIrradiated atomAtoms with enhanced intensityα2, 6, ββ2, 6, αH(OCH3)25, 6α, β2α, β, OCH3 Open table in a new tab 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 lactone1a), 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 δC, δH, and splitting pattern are presented in Tables II andIII. The assignment of the signals in1H and 13C NMR spectra is based on the analysis of the splitting pattern, COSY spectra, and13C-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 δH = 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 δH = 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 δH = 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 δC = 106.29 is characteristic of the acetal carbon (26Kalinowski H.-O. Berger S. Braun S. Carbon-13 NMR Spectroscopy. John Wiley & Sons, Inc., Chichester, UK1988Google Scholar); (v) the second CH group (δH = 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, (δC = 75.80) resembles those of alcohol carbons (26Kalinowski H.-O. Berger S. Braun S. Carbon-13 NMR Spectroscopy. John Wiley & Sons, Inc., Chichester, UK1988Google Scholar).Table II1H chemical shifts for CD3OD solutions of the trimer with structure 3Atom1H chemical shiftMultiplicityJHzAα7.57d16.0Aβ6.37d16.0A27.17d2.0A67.04dd2.0/8.4A56.95d8.4A(OCH3)3.79Bα7.53d16.0Bβ6.33d16.0B27.09d2.0B66.97dd2.0/8.4B56.75d8.4B(OCH3)3.69sCα6.01d5.6Cβ4.95d5.6C27.14d2.0C66.96dd2.0/8.0C56.78d8.0C(OCH3)3.84s Open table in a new tab Table III13C chemical shifts for CD3OD solutions of the trimer with structure 3Atom13C chemical shiftAα145.88Aβ118.32Aγ(CO2H)170.63A2112.57A3152.24A5120.72A6122.78A1132.94A(CH3O)56.49Bα145.88Bβ118.22Bγ(CO2H)170.63B2112.63B3152.08B5120.17B6122.68B(OCH3)56.52Cα106.29Cβ75.80C2112.57C3148.36C5115.72C6121.71C(OCH3)56.41A1, B1, C1131.18, 131.49, 131.67A4, B4, C4148.45, 149.11, 149.35 Open table in a new tab Table IVNOE of the trimer with structure 3Irradiated atomAtoms with enhanced intensityA2Aα, Aβ, A(OCH3)AβA2B2Bα, Bβ, B(OCH3)BβB2C2C(OCH3)CαCβ, C2, C6CβA5, B5, C2 Open table in a new tab 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
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