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Laccase-mediated biotransformation of dihydroquercetin (taxifolin)
Journal of Molecular Catalysis B: Enzymatic 123 (2016) 62–66
Contents lists available at ScienceDirect
Journal of Molecular Catalysis B: Enzymatic journal homepage: www.elsevier.com/locate/molcatb
Laccase-mediated biotransformation of dihydroquercetin (taxifolin) Maria Khlupova a , Irina Vasil’eva a , Galina Shumakovich a , Olga Morozova a , Vyacheslav Chertkov b , Alla Shestakova c , Alexander Kisin c , Alexander Yaropolov a,∗ a b c
Bach Institute of Biochemistry, Research Center of Biotechnology of the Russian Academy of Sciences, Leninsky Ave. 33, bld. 2, 119071 Moscow, Russia Department of Chemistry, Moscow State University, Leninskie Gory 1/3, 119992 Moscow, Russia State Research Institute of Chemistry and Technology of Organoelement Compounds, Shosse Entuziastov 38, 111123 Moscow, Russia
a r t i c l e
i n f o
Article history: Received 12 August 2015 Received in revised form 5 November 2015 Accepted 6 November 2015 Available online 11 November 2015 Keywords: Biocatalysis Laccase-mediated biotransformation Dihydroqurcetin (taxifolin) NMR investigation
a b s t r a c t A laccase (LC) from the basidial fungus Trametes hirsuta has been used for the first time as a catalyst in dihydroquercetin (DHQ) oxidative polymerization. The conditions selected enabled good yields of DHQ oligomers, which were then analyzed using UV–vis, FTIR, 1 Н and 13 C NMR spectroscopy. It was shown that the first stage of LC-mediated oxidation of DHQ is the removal of hydrogen from 3-OH group and formation of an alkoxy radical, which substitutes hydrogens mainly in positions C2 , C5 and C6 . The oligomers have the number average molecular weight of 1000 g/mol and the polydispersity index of 1.4. © 2015 Elsevier B.V. All rights reserved.
1. Introduction Biocatalysis, which is an environmentally friendly method, is increasingly becoming the alternative of chemical synthesis due to the mild conditions of the reaction. Oxidoreductases are an important tool in the oxidation of various natural compounds in order to produce novel compounds. Laccase (benzenediol: oxygen oxidoreductase, EC 1.10.3.2) and bilirubin oxidase (bilirubin: oxygen oxidareductase, EC 1.3.3.5) belong to multicopper oxidases and catalyze one-electron oxidation of inorganic and organic compounds by molecular oxygen, with the concomitant reduction of oxygen to water [1,2]. Aromatic compounds are oxidized in the presence of these enzymes by a radical mechanism with the subsequent coupling of intermediates, which finally results in the formation of oligomers and polymers [3–8]. Enzyme-catalyzed polymerization of natural physiologically active compounds, including various flavonoids, may enable the production of new derivatives for application as pharmaceuticals. As was reported in Refs. [9–12], high molecular weight natural polyphenols exhibit enhanced physiological properties as compared with their monomers.
Abbreviations: DHQ, dihydroquercetin; LC, laccase; BOD, bilirubin oxidase; LC-oligoDHQ, DHQ oligomers synthesized using laccase; BOD-oligoDHQ, DHQ oligomers synthesized using bilirubin oxidase; Mn, the number average molecular weight; PDI, the polydispersity index. ∗ Corresponding author. Fax: +7 495 954 27 32. E-mail address: [email protected] (A. Yaropolov). http://dx.doi.org/10.1016/j.molcatb.2015.11.010 1381-1177/© 2015 Elsevier B.V. All rights reserved.
Dihydroquercetin (taxifolin, (2R,3R)-2-(3,4-Dihydroxyphenyl)3,5,7-trihydroxy-2,3-dihydrochromen-4-one) belongs to natural flavonoids (Fig. 1, 1) and is close to quercetin (2) and rutin (3). As stated in Ref. [13], DHQ is not genotoxic. DHQ has a wide range of pharmaceutical effects, in particular antioxidant, angioprotective, hepatoprotective, antitumoral and other ones [14–17]. Furthermore, DHQ also shows regulatory properties and can control DNA reparation, apoptosis, and mitochondrial biogenesis, as well as regulate the activity of certain enzymes [14,18–21]. It is known that many flavonoids, including DHQ, decompose rather quickly in vivo, while their relatively high molecular weight derivatives have a longer lifespan [22]. The above data show promise for the synthesis of novel DHQ derivatives, which prompted us to investigate the potential of multicopper oxidases, namely a fungal laccase (LC) and bilirubin oxidase (BOD), for DHQ oxidative polymerization, as well as the structure and physicochemical properties of the products obtained. Earlier we reported the results of BOD-catalyzed polymerization of DHQ at neutral pH values of the reaction medium [23]. We showed that the reaction starts with the oxidation of the hydroxyl group in ring C (Fig. 1). Then the radical formed at this stage attacks another DHQ molecule via either the substitution of hydrogen in any possible positions in rings A and B (H2 , H5 , H6 , H6 и H8) or via the removal of H2 and H3 and the formation of double bond C2 = C3. This paper reports another part of our research in enzyme-catalyzed polymerization of DHQ, in which we have investigated the potential of a laccase obtained from Trametes hirsuta
M. Khlupova et al. / Journal of Molecular Catalysis B: Enzymatic 123 (2016) 62–66
and compared the reactions of BOD-catalyzed and LC-catalyzed polymerization of DHQ.
2. Materials and methods 2.1. Materials KH2 PO4 , NaOH, citric acid, 2,2 -azino-bis(3-ethyl benzthiazoline-6-sylfonate (ABTS) from Sigma–Aldrich, dimethylsulfoxide (DMSO) from Marbiopharm (Russia) were used without further purification. All the solutions were prepared using deionized water obtained in a Milli Q system (Millipore, USA). Dehydroquercetin (≥96%) was purchased from BioChemMak-ST (Russia). The laccase was isolated from a culture liquid of the basidial fungus T. hirsuta as described in Ref. [24]. The enzyme homogeneity was checked by SDS electrophoresis. The laccase activity was measured spectrophotometrically using 1 mM ABTS as chromogenic substrate ( = 420 nm, ε = 36000 M−1 cm−1 ) at 24 ◦ C in 0.1 M Na-citrate-phosphate buffer, pH 4.5. One unit of activity is defined as the amount of laccase oxidizing 1 m of substrate per min. The specific activity of the enzyme preparation was about 158 U/mg of protein.
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2.2. Methods 2.2.1. Enzyme-catalyzed DHQ polymerization DHQ oligomers were synthesized in buffer solutions containing 3% v/v ethanol at room temperature under aerobic conditions. LC-mediated DHQ polymerization was carried out in 0.1 M citratephosphate buffer, pH 4.5. in every experiment, 10 mg of DHQ was dissolved in 0.3 ml of ethanol, and then 10 ml of the corresponding buffer was added to the solution. The final DHQ concentration in the reaction mixture was 32 mM. DHQ polymerization initiated by adding the enzyme was performed in the air at room temperature (21–22 ◦ C) with constant stirring for 24 h until insoluble products were formed. The specific activity of the enzyme in the reaction mixture was 0.4 U/ml. DHQ oligomers were separated by centrifugation, washed many times with 3% ethanol, dried and used in experiments. 2.2.2. Characterization of DHQ oligomers The average molecular weight and polydispersity index of the DHQ oligomers produced in the enzyme-catalyzed reaction were determined by gel exclusion chromatography using a RI detector. The molecular weight of LC-oligoDHQ was determined on a PL-gel C column (“Phenomenex”) using THF as a mobile phase. The column was calibrated by polysterene standards. UV–vis spectra were recorded using a Shimadzu UV1240 mini spectrophotometer (Japan). FTIR spectra were obtained in KBr pellets using a Frontier FT-IR/FIR specrometer (PerkinElmer Inc.). Thermogravimetric analysis of the samples was carried out in N2 using a NETZSCH STA 409 PC/PG apparatus with a heating rate of 10 ◦ C/min. 1 Н and 13 C NMR spectra of DHQ and oligoDHQ were recorded for the solutions in DMSO-D6 using a Bruker AVANCE 600 spectrometer with an operating frequency of 600.03 MHz for 1 Н nuclei. 3. Results and discussion
Fig. 1. Structures of certain flavonoids.
All the data on the BOD-catalyzed reaction of DHQpolymerization used in this work for the comparison with the LC-catalyzed reaction are reported in detail in our previous paper [23]. In a preliminary experiment, we obtained pH dependences of LC activity in DHQ oxidation and found that Lc showed the optimal activity at pH 4.0–4.5 (see ESI). It is noteworthy that no insoluble oligomers are formed in LC-catalyzed oxidation of DHQ at pH > 6 and the maximum yield of insoluble products was observed at pH 4.5. On the contrary, in the BOD-catalyzed reaction [23], the maximum yield of insoluble oligomers was observed at pH 7.0 and no insoluble products appeared at pH < 6. The DHQ monomer shows intensive absorbance bands at 290 and 328 nm, which correspond to –* transitions of the aromatic moiety. The ratio of the intensity of the two bands depends on the pH of the solution, which is indicative of phenolic group ionization under pH changes. The absorbance band at 290 nm corresponds to nonionized DHQ, while that at 328 nm to its deprotonated form. Size exclusion chromatographic studies showed that LColigoDHQ had the number average molecular weight (Mn) of 1000 g/mol and the polydispersity index (PDI) of 1.4, while as was reported in Ref. [23] BOD-oligoDHQ had Mn = 2800 g/mol and PDI = 8.6. As compared with DHQ monomer spectrum, UV–vis spectrum of LC-oligoDHQ (Fig. 2) had a long wavelength tail indicating the presence of extended conjugation [25]. The thermogravimetric analysis (TGA) was used to study the thermal stability of the products of LCcatalyzed polymerization of DHQ (Fig. 3). The stability of DHQ and LC-oligoDHQ was approximately equal up to 240–250 ◦ C. At higher
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Fig. 2. UV–vis spectra of DMSO dissolved DHQ (1) and LC-oligoDHQ (2).
temperatures, all samples studied suddenly lost weight, the thermal stability of LC-oligoDHQ being higher than that of DHQ, i.e., LC-oligoDHQ lost 20% of the weight at 385 ◦ C, while DHQ showed the same weight lost at 306 ◦ C. The FTIR spectrum of LC-oligoDHQ resembles the DHQ vibration spectrum, but its peaks are wider and more smoothed (Fig. 4). The broad peak with the maximum at 3400 cm−1 corresponds to the vibration of the O H linkage of phenolic and hydroxyl groups [26], the peak at 1640 cm−1 can be attributed to the carbonyl groups [6,10], the peak at 1150 cm−1 is due to the vibration of Caryl O linkage [27], and the low peaks in the range 780–810 cm−1 can result from the vibrations of C C and C Н linkages of the aromatic ring [6]. It is noteworthy that the FTIR spectrum of Lc-oligoDHQ had no absorption bands at 870, 1200, 1240 cm−1 , which correspond to cooperative vibrations of the Caryl O Caryl linkages. However, these data on the DHQ oligomer structure can hardly be regarded as reliable. Therefore, we used 1 Н and 13 C NMR spectroscopy to study the structure of the LC-synthesized oligomers of DHQ in detail. The comparative analysis of 1 Н and 13 C NMR spectra of DHQ monomer and LC-oligoDHQ (Figs. 5 and 6, respectively) was performed in presumption that the DHQ carbon skeleton is rather stable [28] and remains unchanged in the course of enzymecatalyzed polymerization. The assignment of DHQ proton signals is shown in Fig. 5a. In order to reveal hydroxyl protons in 1 Н NMR spectra, the solutions
Fig. 3. Thermal stability of DHQ (1) and LC-oligoDHQ (2).
Fig. 4. FTIR-spectra of DHQ (a) and LC-oligoDHQ (b).
of oligoDHQ in DMSO-D6 were diluted 15 times with the solution of D2 O (10 l) in DMSO-D6 (630 l). As a result, the intensity of signals of hydroxyl protons decreases markedly due to the substitution of protons for deuterium atoms (Fig. 5c), which allowed us to recognize the peaks of the OH-groups. A comparison of areas at 8.3–13.5 ppm enables us to conclude that all the phenolic hydroxyls are conserved in the sample of LC-oligoDHQ (Fig. 5b and c). The hydroxyl protons 7-OH, 3 -OН and 4 -OН are significantly broadened due to fast exchange processes, while the protons of 5-OH groups in both DHQ monomer and LC-oligoDHQ participate in the formation of a very stable hydrogen bond with the carbonyl oxygen of ring C, which is confirmed by the presence of relatively narrow and clearly distinguished peaks at 11.3–12.3 ppm. It should be noted that the signal of the hydroxyl group at C3 is lacking in the spectra of oligoDHQ, which permits us to assume that LC-catalyzed DHQ polymerization is accompanied by oxidation of the hydroxyl group at C3. Broadened signals with a high integral intensity in the area 5.5–8.0 ppm of the proton spectra of LC-oligoDHQ can be assigned to the corresponding signals of DHQ monomer aromatic protons H2 , H5 , H6 , H6 и H8. However, the signals Н2 , Н5 and Н6 (Fig. 5b and c) have a markedly lower intensity as compared with the intensity of H6 and H8. There are also some broad signals in the area of H2 and H3 (4.5–5.8 ppm) in the spectrum of LC-oligoDHQ (Fig. 5b), but their total integral intensity is noticeably lower as compared with the intensity of the aromatic protons. Thus, the analysis of the proton spectra of DHQ monomer and LC-oligoDHQ allowed us to conclude that laccase-catalyzed polymerization of DHQ starts with the oxidation of the hydroxyl group in position 3 and the alkoxy radical formed at this stage attacks another DHQ molecule via radical or electrophilic substitution of ɑn accessible aromatic hydrogen in ring B. Analysis of 13 C NMR spectra (Fig. 6) provides some extra information on the structure of DHQ oligomers. A comparison of 13 C NMR spectra of DHQ monomer (Fig. 6a) with LC-oligoDHQ (Fig. 6b) revealed that all the peaks in the carbon spectrum of DHQ monomer also present in the spectrum of LC-oligoDHQ. For example, the broadened peaks of the carbonyl carbon of LC-oligoDHQ at 197–198 ppm correspond to the position of the carbonyl group in the monomer (197.7 ppm). However, of the most interest are the peaks lacking in the spectrum of DHQ monomer but present in the
M. Khlupova et al. / Journal of Molecular Catalysis B: Enzymatic 123 (2016) 62–66
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Fig. 5. 1 Н NMR spectra (DMSO-D6 , Bruker AV-600, 303 K): (ɑ) DHQ, (b) LC-oligoDHQ and (c) diluted solution of LC-oligoDHQ in DMSO-D6 (670 l) with addition of D2 O (10 l).
Fig. 6.
13
C-{1 H} NMR spectra: DHQ (a) and LC-oligoDHQ (b).
spectrum of its oligomers. It is a group of peaks assigned to quaternary carbons at 191–193 ppm. Analysis of literature data enables us to assume that these peaks ought to be assigned to C4 carbonyl group, and the high field shift may be attributed to the gammaeffect caused by the replacement of the hydroxyl in position 3 with an alkoxy group [22,29–31]. The broad peaks in the spectrum of oligomers correspond to the signals of quaternary and CH carbons in rings A and B of monomeric DHQ (Fig. 6). At the same time, the peaks of C5–C10 in ring A of LC-oligoDHQ (Fig. 6b) remain virtually in the same position as in the monomer, while the signals of all the six carbons in ring B are noticeably broader and have a markedly lower intensity as compared with the carbons in ring A. Also numerous narrow peaks of quaternary carbons are observed at 123–147 ppm. This type of spectrum most likely indicates that LC-mediated polymerization of DHQ proceeds via substitution of hydrogen atoms for an alkoxy
radical in all three possible positions of ring B (Н2 , Н5 , Н6 ). Broad intensive signals are observed in LC-oligoDHQ spectrum in the region characteristic of aliphatic C2 and C3 of the parent monomer (83.0 and 71.5 ppm, respectively). Thus, the results of the comparative analysis of 1 Н and 13 C NMR spectra of DHQ monomer and LC-oligoDHQ, as well as some literature data on related compounds [22,29–31] allow us to assume that LC-catalyzed oxidation of DHQ starts with the removal of hydrogen from the hydroxyl group at C3 to produce an active radical, which can be predominantly attached to C2 , C5 or C6 carbons of another molecule of monomer. 4. Conclusions For the first time oxidative polymerization of dihydroquercetin (taxifolin) has been performed using a laccase from T. hirsuta and
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resultant LC-oligoDHQ have been studied using UV–vis, FTIR, TGA, 1 Н and 13 C NMR. It has been found that LC-oligoDHQ had the number average molecular weight of 1000 g/mol and the polydispersity index of 1.4, and DHQ oligomers were more thermostable than the monomer. We have also shown that the first stage of LC-mediated oxidation of DHQ at pH 4.5 is the removal of hydrogen from 3-OH group and formation of an alkoxy radical, which substitutes hydrogens mainly in positions C2 , C5 and C6 without the formation of oligoDHQ with double bond C2 = C3, in contrast to BOD-mediated oxidation of DHQ at pH 7.0. The comparison of the results of LC- and BOD-catalyzed polymerization of DHQ enables us to assume that DHQ oligomers formed at optimal pH values differ not only in the number average molecular weight and polydispersity index but also in structure. Acknowledgement The reported study was supported by RFBR, research project No. 14-04-00403a. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.molcatb.2015.11. 010. References [1] P. Baldrian, FEMS Microbiol Rev. 30 (2006) 215–242. [2] E.I. Solomon, U.M. Sundaram, T.E. Machonkin, Chem. Rev. 96 (1996) 2563–2606. [3] S. Witayakran, A.J. Ragauskas, Adv. Synth. Catal. 351 (2009) 1187–1209. [4] P. Walde, Z. Guo, Soft Matter 7 (2011) 316–331. [5] F. Hollmann, I.W.C.E. Arends, Polymers 4 (2012) 759–793. [6] L. Mejias, M.H. Reihmann, S. Sepulveda-Boza, H. Ritter, Macromol. Biosci. 2 (2002) 24–32.
[7] Y.-J. Kim, H. Uyama, Sh. Kobayashi, Macromolecules 36 (2003) 5058–5060. [8] M. Magharabi, M.A. Faramazzi, Adv. Synth. Catal. 356 (2014) 897–927. [9] A.E. Hagerman, K.M. Riedl, G.A. Jones, K.N. Sovik, N.T. Ritchard, P.W. Hartzfeld, T.L. Riechel, J. Agric. Food Chem. 46 (1998) 1887–1892. [10] M. Kurisawa, J.E. Chung, H. Uyama, S. Kobayashi, Biomacromolecules 4 (2003) 1394–1399. [11] M. Kurisawa, J.E. Chung, H. Uyama, S. Kobayashi, Macromol. Biosci. 3 (2003) 758–764. [12] J. Zhao, J. Wang, Y. Chen, R. Agarwa, Carcinogenesis 20 (1999) 1737–1745. [13] P.S. Makena, S.C. Pierce, K.-T. Chung, S.E. Sinclair, Environ. Mol. Mutagen. 50 (2009) 451–459. [14] A.E. Weidmann, Eur. J. Pharmacol. 684 (2012) 19–26. [15] G. Pantouris, Ch. G. Mowat, Biochem. Biophys. Res. Commun. 443 (2014) 28–31. [16] S.J. Polyak, Ch. Morishima, V. Lohmann, S. Pal, D.Y.W. Lee, Y. Liu, T.N. Graf, PNAS 107 (2010) 5995–5999. [17] M. Soto, K. Murakami, M. Uno, H. Ikubo, Y. Nakagama, S. Katayama, K. Akagi, K. Irie, Biosci. Biotechnol. Biochem. 77 (2013) 1100–1103. [18] A. Bordoni, S. Hrelia, C. Angeloni, E. Giordano, C. Guarnieri, C.M. Caldarera, P.L. Bia, J. Nutr. Biochem. 13 (2002) 103–111. [19] D.R. Ferry, A. Smith, J. Malkhandi, D.W. Fyfe, P.G. Takats, D. Anderson, J. Baker, D.J. Kerr, Clin. Cancer Res. 2 (1996) 659–668. [20] K. Ono, H. Nakane, M. Fukushima, J.-C. Chermann, F. Barre-Sinoussi, Eur. J. Biochem. 190 (1990) 469–476. [21] Zh.R. Zhang, M.A. Zaharna, M.M.-K. Wong, S.-K. Chin, H.-Y. Cheng, PLoS One 8 (2013) e54577. [22] P.C. Hollman, M.B. Katan, Biomed. Pharmacother. 51 (1997) 305–310. [23] M.E. Khlupova, I.S. Vasil’eva, G.P. Shumakovich, O.V. Morozova, V.A. Chertkov, A.K. Shestakova, A.V. Kisin, A.I. Yaropolov, Biochemistry (Moscow) 80 (2015) 233–241. [24] E.S. Gorshina, T.V. Rusinova, V.V. Biryukov, O.V. Morozova, S.V. Shleev, A.I. Yaropolov, Appl. Biochem. Microbiol. 42 (2006) 558–563. [25] F.F. Bruno, A. Trotta, S. Fossey, S. Nagarajan, R. Nagarajan, L.A. Samuelson, J. Kumar, J. Macromol. Sci. Part A: Pure Appl. Chem. 47 (2010) 1191–1196. [26] S.Y. Park, Y.H. Kim, K. Won, B.K. Song, J. Mol. Catal. B: Enzym. 57 (2009) 312–316. [27] K. Won, Y.H. Kim, E.S. An, Y.S. Lee, B.K. Song, Biomacromol 5 (2004) 1–4. [28] E. Kiehlmann, E.P.M. Li, J. Nat. Prod. 58 (1995) 450–455. [29] E. Kiehlmann, P.W. Slade, J. Nat. Prod. 66 (2003) 1562–1566. [30] Y.A. Kolesnik, E.V. Titova, V.A. Chertkov, V.N. Tashlitskiy, V.P. Tichonov, D.A. Shmatkov, Planta Med. 77 (2011) 1266. [31] R. Sinha, A. Joshi, U.J. Joshi, S. Srivastava, G. Govil, Eur. J. Med. Chem. 80 (2014) 285–294.
Contents lists available at ScienceDirect
Journal of Molecular Catalysis B: Enzymatic journal homepage: www.elsevier.com/locate/molcatb
Laccase-mediated biotransformation of dihydroquercetin (taxifolin) Maria Khlupova a , Irina Vasil’eva a , Galina Shumakovich a , Olga Morozova a , Vyacheslav Chertkov b , Alla Shestakova c , Alexander Kisin c , Alexander Yaropolov a,∗ a b c
Bach Institute of Biochemistry, Research Center of Biotechnology of the Russian Academy of Sciences, Leninsky Ave. 33, bld. 2, 119071 Moscow, Russia Department of Chemistry, Moscow State University, Leninskie Gory 1/3, 119992 Moscow, Russia State Research Institute of Chemistry and Technology of Organoelement Compounds, Shosse Entuziastov 38, 111123 Moscow, Russia
a r t i c l e
i n f o
Article history: Received 12 August 2015 Received in revised form 5 November 2015 Accepted 6 November 2015 Available online 11 November 2015 Keywords: Biocatalysis Laccase-mediated biotransformation Dihydroqurcetin (taxifolin) NMR investigation
a b s t r a c t A laccase (LC) from the basidial fungus Trametes hirsuta has been used for the first time as a catalyst in dihydroquercetin (DHQ) oxidative polymerization. The conditions selected enabled good yields of DHQ oligomers, which were then analyzed using UV–vis, FTIR, 1 Н and 13 C NMR spectroscopy. It was shown that the first stage of LC-mediated oxidation of DHQ is the removal of hydrogen from 3-OH group and formation of an alkoxy radical, which substitutes hydrogens mainly in positions C2 , C5 and C6 . The oligomers have the number average molecular weight of 1000 g/mol and the polydispersity index of 1.4. © 2015 Elsevier B.V. All rights reserved.
1. Introduction Biocatalysis, which is an environmentally friendly method, is increasingly becoming the alternative of chemical synthesis due to the mild conditions of the reaction. Oxidoreductases are an important tool in the oxidation of various natural compounds in order to produce novel compounds. Laccase (benzenediol: oxygen oxidoreductase, EC 1.10.3.2) and bilirubin oxidase (bilirubin: oxygen oxidareductase, EC 1.3.3.5) belong to multicopper oxidases and catalyze one-electron oxidation of inorganic and organic compounds by molecular oxygen, with the concomitant reduction of oxygen to water [1,2]. Aromatic compounds are oxidized in the presence of these enzymes by a radical mechanism with the subsequent coupling of intermediates, which finally results in the formation of oligomers and polymers [3–8]. Enzyme-catalyzed polymerization of natural physiologically active compounds, including various flavonoids, may enable the production of new derivatives for application as pharmaceuticals. As was reported in Refs. [9–12], high molecular weight natural polyphenols exhibit enhanced physiological properties as compared with their monomers.
Abbreviations: DHQ, dihydroquercetin; LC, laccase; BOD, bilirubin oxidase; LC-oligoDHQ, DHQ oligomers synthesized using laccase; BOD-oligoDHQ, DHQ oligomers synthesized using bilirubin oxidase; Mn, the number average molecular weight; PDI, the polydispersity index. ∗ Corresponding author. Fax: +7 495 954 27 32. E-mail address: [email protected] (A. Yaropolov). http://dx.doi.org/10.1016/j.molcatb.2015.11.010 1381-1177/© 2015 Elsevier B.V. All rights reserved.
Dihydroquercetin (taxifolin, (2R,3R)-2-(3,4-Dihydroxyphenyl)3,5,7-trihydroxy-2,3-dihydrochromen-4-one) belongs to natural flavonoids (Fig. 1, 1) and is close to quercetin (2) and rutin (3). As stated in Ref. [13], DHQ is not genotoxic. DHQ has a wide range of pharmaceutical effects, in particular antioxidant, angioprotective, hepatoprotective, antitumoral and other ones [14–17]. Furthermore, DHQ also shows regulatory properties and can control DNA reparation, apoptosis, and mitochondrial biogenesis, as well as regulate the activity of certain enzymes [14,18–21]. It is known that many flavonoids, including DHQ, decompose rather quickly in vivo, while their relatively high molecular weight derivatives have a longer lifespan [22]. The above data show promise for the synthesis of novel DHQ derivatives, which prompted us to investigate the potential of multicopper oxidases, namely a fungal laccase (LC) and bilirubin oxidase (BOD), for DHQ oxidative polymerization, as well as the structure and physicochemical properties of the products obtained. Earlier we reported the results of BOD-catalyzed polymerization of DHQ at neutral pH values of the reaction medium [23]. We showed that the reaction starts with the oxidation of the hydroxyl group in ring C (Fig. 1). Then the radical formed at this stage attacks another DHQ molecule via either the substitution of hydrogen in any possible positions in rings A and B (H2 , H5 , H6 , H6 и H8) or via the removal of H2 and H3 and the formation of double bond C2 = C3. This paper reports another part of our research in enzyme-catalyzed polymerization of DHQ, in which we have investigated the potential of a laccase obtained from Trametes hirsuta
M. Khlupova et al. / Journal of Molecular Catalysis B: Enzymatic 123 (2016) 62–66
and compared the reactions of BOD-catalyzed and LC-catalyzed polymerization of DHQ.
2. Materials and methods 2.1. Materials KH2 PO4 , NaOH, citric acid, 2,2 -azino-bis(3-ethyl benzthiazoline-6-sylfonate (ABTS) from Sigma–Aldrich, dimethylsulfoxide (DMSO) from Marbiopharm (Russia) were used without further purification. All the solutions were prepared using deionized water obtained in a Milli Q system (Millipore, USA). Dehydroquercetin (≥96%) was purchased from BioChemMak-ST (Russia). The laccase was isolated from a culture liquid of the basidial fungus T. hirsuta as described in Ref. [24]. The enzyme homogeneity was checked by SDS electrophoresis. The laccase activity was measured spectrophotometrically using 1 mM ABTS as chromogenic substrate ( = 420 nm, ε = 36000 M−1 cm−1 ) at 24 ◦ C in 0.1 M Na-citrate-phosphate buffer, pH 4.5. One unit of activity is defined as the amount of laccase oxidizing 1 m of substrate per min. The specific activity of the enzyme preparation was about 158 U/mg of protein.
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2.2. Methods 2.2.1. Enzyme-catalyzed DHQ polymerization DHQ oligomers were synthesized in buffer solutions containing 3% v/v ethanol at room temperature under aerobic conditions. LC-mediated DHQ polymerization was carried out in 0.1 M citratephosphate buffer, pH 4.5. in every experiment, 10 mg of DHQ was dissolved in 0.3 ml of ethanol, and then 10 ml of the corresponding buffer was added to the solution. The final DHQ concentration in the reaction mixture was 32 mM. DHQ polymerization initiated by adding the enzyme was performed in the air at room temperature (21–22 ◦ C) with constant stirring for 24 h until insoluble products were formed. The specific activity of the enzyme in the reaction mixture was 0.4 U/ml. DHQ oligomers were separated by centrifugation, washed many times with 3% ethanol, dried and used in experiments. 2.2.2. Characterization of DHQ oligomers The average molecular weight and polydispersity index of the DHQ oligomers produced in the enzyme-catalyzed reaction were determined by gel exclusion chromatography using a RI detector. The molecular weight of LC-oligoDHQ was determined on a PL-gel C column (“Phenomenex”) using THF as a mobile phase. The column was calibrated by polysterene standards. UV–vis spectra were recorded using a Shimadzu UV1240 mini spectrophotometer (Japan). FTIR spectra were obtained in KBr pellets using a Frontier FT-IR/FIR specrometer (PerkinElmer Inc.). Thermogravimetric analysis of the samples was carried out in N2 using a NETZSCH STA 409 PC/PG apparatus with a heating rate of 10 ◦ C/min. 1 Н and 13 C NMR spectra of DHQ and oligoDHQ were recorded for the solutions in DMSO-D6 using a Bruker AVANCE 600 spectrometer with an operating frequency of 600.03 MHz for 1 Н nuclei. 3. Results and discussion
Fig. 1. Structures of certain flavonoids.
All the data on the BOD-catalyzed reaction of DHQpolymerization used in this work for the comparison with the LC-catalyzed reaction are reported in detail in our previous paper [23]. In a preliminary experiment, we obtained pH dependences of LC activity in DHQ oxidation and found that Lc showed the optimal activity at pH 4.0–4.5 (see ESI). It is noteworthy that no insoluble oligomers are formed in LC-catalyzed oxidation of DHQ at pH > 6 and the maximum yield of insoluble products was observed at pH 4.5. On the contrary, in the BOD-catalyzed reaction [23], the maximum yield of insoluble oligomers was observed at pH 7.0 and no insoluble products appeared at pH < 6. The DHQ monomer shows intensive absorbance bands at 290 and 328 nm, which correspond to –* transitions of the aromatic moiety. The ratio of the intensity of the two bands depends on the pH of the solution, which is indicative of phenolic group ionization under pH changes. The absorbance band at 290 nm corresponds to nonionized DHQ, while that at 328 nm to its deprotonated form. Size exclusion chromatographic studies showed that LColigoDHQ had the number average molecular weight (Mn) of 1000 g/mol and the polydispersity index (PDI) of 1.4, while as was reported in Ref. [23] BOD-oligoDHQ had Mn = 2800 g/mol and PDI = 8.6. As compared with DHQ monomer spectrum, UV–vis spectrum of LC-oligoDHQ (Fig. 2) had a long wavelength tail indicating the presence of extended conjugation [25]. The thermogravimetric analysis (TGA) was used to study the thermal stability of the products of LCcatalyzed polymerization of DHQ (Fig. 3). The stability of DHQ and LC-oligoDHQ was approximately equal up to 240–250 ◦ C. At higher
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Fig. 2. UV–vis spectra of DMSO dissolved DHQ (1) and LC-oligoDHQ (2).
temperatures, all samples studied suddenly lost weight, the thermal stability of LC-oligoDHQ being higher than that of DHQ, i.e., LC-oligoDHQ lost 20% of the weight at 385 ◦ C, while DHQ showed the same weight lost at 306 ◦ C. The FTIR spectrum of LC-oligoDHQ resembles the DHQ vibration spectrum, but its peaks are wider and more smoothed (Fig. 4). The broad peak with the maximum at 3400 cm−1 corresponds to the vibration of the O H linkage of phenolic and hydroxyl groups [26], the peak at 1640 cm−1 can be attributed to the carbonyl groups [6,10], the peak at 1150 cm−1 is due to the vibration of Caryl O linkage [27], and the low peaks in the range 780–810 cm−1 can result from the vibrations of C C and C Н linkages of the aromatic ring [6]. It is noteworthy that the FTIR spectrum of Lc-oligoDHQ had no absorption bands at 870, 1200, 1240 cm−1 , which correspond to cooperative vibrations of the Caryl O Caryl linkages. However, these data on the DHQ oligomer structure can hardly be regarded as reliable. Therefore, we used 1 Н and 13 C NMR spectroscopy to study the structure of the LC-synthesized oligomers of DHQ in detail. The comparative analysis of 1 Н and 13 C NMR spectra of DHQ monomer and LC-oligoDHQ (Figs. 5 and 6, respectively) was performed in presumption that the DHQ carbon skeleton is rather stable [28] and remains unchanged in the course of enzymecatalyzed polymerization. The assignment of DHQ proton signals is shown in Fig. 5a. In order to reveal hydroxyl protons in 1 Н NMR spectra, the solutions
Fig. 3. Thermal stability of DHQ (1) and LC-oligoDHQ (2).
Fig. 4. FTIR-spectra of DHQ (a) and LC-oligoDHQ (b).
of oligoDHQ in DMSO-D6 were diluted 15 times with the solution of D2 O (10 l) in DMSO-D6 (630 l). As a result, the intensity of signals of hydroxyl protons decreases markedly due to the substitution of protons for deuterium atoms (Fig. 5c), which allowed us to recognize the peaks of the OH-groups. A comparison of areas at 8.3–13.5 ppm enables us to conclude that all the phenolic hydroxyls are conserved in the sample of LC-oligoDHQ (Fig. 5b and c). The hydroxyl protons 7-OH, 3 -OН and 4 -OН are significantly broadened due to fast exchange processes, while the protons of 5-OH groups in both DHQ monomer and LC-oligoDHQ participate in the formation of a very stable hydrogen bond with the carbonyl oxygen of ring C, which is confirmed by the presence of relatively narrow and clearly distinguished peaks at 11.3–12.3 ppm. It should be noted that the signal of the hydroxyl group at C3 is lacking in the spectra of oligoDHQ, which permits us to assume that LC-catalyzed DHQ polymerization is accompanied by oxidation of the hydroxyl group at C3. Broadened signals with a high integral intensity in the area 5.5–8.0 ppm of the proton spectra of LC-oligoDHQ can be assigned to the corresponding signals of DHQ monomer aromatic protons H2 , H5 , H6 , H6 и H8. However, the signals Н2 , Н5 and Н6 (Fig. 5b and c) have a markedly lower intensity as compared with the intensity of H6 and H8. There are also some broad signals in the area of H2 and H3 (4.5–5.8 ppm) in the spectrum of LC-oligoDHQ (Fig. 5b), but their total integral intensity is noticeably lower as compared with the intensity of the aromatic protons. Thus, the analysis of the proton spectra of DHQ monomer and LC-oligoDHQ allowed us to conclude that laccase-catalyzed polymerization of DHQ starts with the oxidation of the hydroxyl group in position 3 and the alkoxy radical formed at this stage attacks another DHQ molecule via radical or electrophilic substitution of ɑn accessible aromatic hydrogen in ring B. Analysis of 13 C NMR spectra (Fig. 6) provides some extra information on the structure of DHQ oligomers. A comparison of 13 C NMR spectra of DHQ monomer (Fig. 6a) with LC-oligoDHQ (Fig. 6b) revealed that all the peaks in the carbon spectrum of DHQ monomer also present in the spectrum of LC-oligoDHQ. For example, the broadened peaks of the carbonyl carbon of LC-oligoDHQ at 197–198 ppm correspond to the position of the carbonyl group in the monomer (197.7 ppm). However, of the most interest are the peaks lacking in the spectrum of DHQ monomer but present in the
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Fig. 5. 1 Н NMR spectra (DMSO-D6 , Bruker AV-600, 303 K): (ɑ) DHQ, (b) LC-oligoDHQ and (c) diluted solution of LC-oligoDHQ in DMSO-D6 (670 l) with addition of D2 O (10 l).
Fig. 6.
13
C-{1 H} NMR spectra: DHQ (a) and LC-oligoDHQ (b).
spectrum of its oligomers. It is a group of peaks assigned to quaternary carbons at 191–193 ppm. Analysis of literature data enables us to assume that these peaks ought to be assigned to C4 carbonyl group, and the high field shift may be attributed to the gammaeffect caused by the replacement of the hydroxyl in position 3 with an alkoxy group [22,29–31]. The broad peaks in the spectrum of oligomers correspond to the signals of quaternary and CH carbons in rings A and B of monomeric DHQ (Fig. 6). At the same time, the peaks of C5–C10 in ring A of LC-oligoDHQ (Fig. 6b) remain virtually in the same position as in the monomer, while the signals of all the six carbons in ring B are noticeably broader and have a markedly lower intensity as compared with the carbons in ring A. Also numerous narrow peaks of quaternary carbons are observed at 123–147 ppm. This type of spectrum most likely indicates that LC-mediated polymerization of DHQ proceeds via substitution of hydrogen atoms for an alkoxy
radical in all three possible positions of ring B (Н2 , Н5 , Н6 ). Broad intensive signals are observed in LC-oligoDHQ spectrum in the region characteristic of aliphatic C2 and C3 of the parent monomer (83.0 and 71.5 ppm, respectively). Thus, the results of the comparative analysis of 1 Н and 13 C NMR spectra of DHQ monomer and LC-oligoDHQ, as well as some literature data on related compounds [22,29–31] allow us to assume that LC-catalyzed oxidation of DHQ starts with the removal of hydrogen from the hydroxyl group at C3 to produce an active radical, which can be predominantly attached to C2 , C5 or C6 carbons of another molecule of monomer. 4. Conclusions For the first time oxidative polymerization of dihydroquercetin (taxifolin) has been performed using a laccase from T. hirsuta and
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resultant LC-oligoDHQ have been studied using UV–vis, FTIR, TGA, 1 Н and 13 C NMR. It has been found that LC-oligoDHQ had the number average molecular weight of 1000 g/mol and the polydispersity index of 1.4, and DHQ oligomers were more thermostable than the monomer. We have also shown that the first stage of LC-mediated oxidation of DHQ at pH 4.5 is the removal of hydrogen from 3-OH group and formation of an alkoxy radical, which substitutes hydrogens mainly in positions C2 , C5 and C6 without the formation of oligoDHQ with double bond C2 = C3, in contrast to BOD-mediated oxidation of DHQ at pH 7.0. The comparison of the results of LC- and BOD-catalyzed polymerization of DHQ enables us to assume that DHQ oligomers formed at optimal pH values differ not only in the number average molecular weight and polydispersity index but also in structure. Acknowledgement The reported study was supported by RFBR, research project No. 14-04-00403a. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.molcatb.2015.11. 010. References [1] P. Baldrian, FEMS Microbiol Rev. 30 (2006) 215–242. [2] E.I. Solomon, U.M. Sundaram, T.E. Machonkin, Chem. Rev. 96 (1996) 2563–2606. [3] S. Witayakran, A.J. Ragauskas, Adv. Synth. Catal. 351 (2009) 1187–1209. [4] P. Walde, Z. Guo, Soft Matter 7 (2011) 316–331. [5] F. Hollmann, I.W.C.E. Arends, Polymers 4 (2012) 759–793. [6] L. Mejias, M.H. Reihmann, S. Sepulveda-Boza, H. Ritter, Macromol. Biosci. 2 (2002) 24–32.
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