Structure–Oxidative Metabolism Relationships of Substituted Flavones

Mar 21, 2017 - Aspergillus niger is a rich source of oxidative enzymes, which are important for many industrial applications. However, systematic eval...
0 downloads 0 Views 1MB Size
Download PDF
Article pubs.acs.org/JAFC

Structure−Oxidative Metabolism Relationships of Substituted Flavones by Aspergillus niger Da-Som Lim, Do-Hyung Lim, Ji-Ho Lee, Eun-Tae Oh, and Young-Soo Keum* Department of Crop Science, Konkuk University, 1 Hwayang-dong, Gwanjin-Gu, Seoul 143-701, Republic of Korea S Supporting Information *

ABSTRACT: Aspergillus niger is a rich source of oxidative enzymes, which are important for many industrial applications. However, systematic evaluation of their metabolic characteristics is limited. In this study, structure-dependent metabolism of flavones by Aspergillus niger were investigated with synthetic substrates. Metabolic inhibitor studies suggested that cytochrome P450s are the major enzymes in oxidative metabolism. The reactions include ring hydroxylation, O-demethylation, sulfone/ sulfoxide formation, and oxidation of alkyls to carboxy groups. Initial oxidative metabolism occurred almost exclusively at 4′substituents. 4′-Halogenated- and 3′,5′-dihalogenated analogues were stable against biodegradation. Hydrophilic flavones were more rapidly metabolized than lipophilic analogues. Molecular widths of the A and B ring were important determinants of the position of metabolic oxidation and biotransformation rate. The structure−metabolism relationship analysis indicates that the shape of the B ring was the most important parameter of biotransformation. The electrostatic environment of the same ring also affected the transformation. Additionally, the results showed that the B ring may preferentially be oriented toward the catalytic center. KEYWORDS: flavone, structure−metabolism, cytochrome P450, Aspergillus niger, regioselectivity

■

INTRODUCTION Aspergillus are ubiquitous fungi found in soils, plants, animals, and even in marine environments. Several Aspergillus are important for the production of fermented foods, antibiotics, and organic acids.1 Some extracellular enzymes are widely used in food processing (e.g., glycosidase, lipase, pectinases, etc.). According to the literature, the fungi can metabolize diverse synthetic chemicals. For example, some pharmaceuticals (e.g., naproxen) are extensively metabolized by A. niger.2 Aspergillus can also degrade several environmental contaminants, including pesticides, kerosene, and aromatics.3−5 A number of natural products are also subject to biotransformation. Plant cell wall polysaccharides can be depolymerized by Aspergillus glycosidases.6 Previous studies indicated that these fungi can metabolize numerous natural flavonoids.7,8 However, most studies were performed with a limited number of substrates. Tahara et al. reported that several enzymes may participate in fungal metabolism of prenylated flavones.9 The metabolic patterns of these xenobiotics suggest that oxidative enzymes (e.g., cytochrome P450s, CYP) have a major metabolic role. Recent genomic studies indicate that the Aspergillis genome contains more than 200 genes of xenobiotic metabolism and approximately 150 CYPs.10,11 However, structural aspects of fungal CYPs were uncovered only with some CYPs (e.g., CYP51). Because of the diverse metabolic enzymatic systems in these microorganisms, synthetic applications of the enzymes or whole organisms have recently been reported.12 For example, Mazzaferro et al. were able to prepare coumarin dimers from the monomeric units by a CYP-catalyzed reaction.13 Bioactive aurone derivatives can also be prepared from the same organism from chalcones.14 Weis et al. suggested the possible use of microbial CYPs for the production of drug metabolites.15 During the study with multiple substrates, it was proven that © XXXX American Chemical Society

some fungal enzymes, as with their mammalian counterparts, are specifically induced by synthetic chemicals.16 Overall, there are multiple xenobiotic catabolic enzymes in Aspergillus, of which the characteristics (e.g., substrate specificity, inducibility) differ from one another. Their properties can be determined directly by genomic methodology or inferred from the substrate structure-dependent activity relationships (SARs). In consideration of metabolic versatility, detailed understanding of structural characteristics of this microorganism is crucial for developing organisms/enzymes for industrial application. In addition, correlation analysis of physicochemical properties of substrates and microbial fates is also an important resource to characterize the bioavailability of substrates.17 The aims of this study were to (a) identify the major group of enzymes of flavone catabolism, (b) determine the metabolic characteristics of Aspergillus niger, and (c) construct structure− metabolism relationships of flavone.

■

MATERIALS AND METHODS

Chemicals. The following reagents were purchased from SigmaAldrich Korea Ltd. (Seoul, Korea): flavone, α-naphthoflavone, βnaphthoflavone, piperonylbutoxide, methimazole, ketoconazole, and N,O-bis(trimethylsilyl)trifluoroacetamide-trimethylsilyl chloride (BSTFA-TMCS). 4′-Hydroxyflavone was purchased from Indofine Chemical Ltd. (NJ, USA). Other reagents for the syntheses were obtained from Alfa Aesar Korea (Seoul, Korea). Potato dextrose broth (PDB) was purchased from BD Korea (Seoul, Korea). Solvents were HPLC grade or higher. Received: Revised: Accepted: Published: A

January 24, 2017 March 15, 2017 March 21, 2017 March 21, 2017 DOI: 10.1021/acs.jafc.7b00390 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Article

Journal of Agricultural and Food Chemistry Table 1. Experimental and Estimated Physicochemical Properties of Synthetic Flavones ID

logP

vol. (Å3)a

W (Å)b

L (Å)c

ID

logP

vol. (Å 3)a

W (Å)b

L (Å)c

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

3.27 3.76 3.20 3.80 3.74 4.41 3.85 3.76 3.29 3.37 3.80 4.56 3.95 4.06 3.33 4.00 3.31 3.70 4.89 3.29

203.32 219.76 227.98 238.42 217.73 232.14 220.03 220.14 227.90 206.13 217.88 231.87 220.12 227.57 228.06 238.60 206.30 217.67 269.41 227.96

4.31 5.88 7.07 7.12 4.85 5.13 5.53 4.32 6.43 4.52 4.82 5.05 4.32 4.32 4.30 4.41 4.30 4.31 4.33 4.31

11.29 11.23 11.25 11.28 11.24 11.25 11.30 11.31 11.93 11.30 11.27 11.30 12.10 12.21 13.41 13.56 11.48 11.86 13.44 12.21

21 22 23 24 25 26 27 28 29 30 31 32 33

3.80 3.51 4.35 2.50 3.73 3.38 2.54 2.98 4.17 3.45 4.14 2.73 4.36

217.64 206.09 224.90 253.00 231.70 242.19 252.10 225.70 236.82 208.50 231.72 276.73 248.87

11.88 11.30 11.31 11.81 11.25 14.01 13.26 12.14 11.66 11.32 11.31 13.22 11.66

34

4.40

248.07

35

4.79

248.71

36 37

4.66 2.05

249.11 198.77

4.30 4.31 4.30 6.27 4.85 4.32 6.49 5.06 6.60 4.72 5.32 8.32 4.30 8.22d 4.31 7.98d 4.31 6.02d 5.66 4.16

13.45 13.50 13.59 10.17

Van der Waals surface-based molar volume in cubic angstroms (A3). bMolecular width of the B ring of flavones (width1, W1) in angstroms (Å). The longest dimension was measured. dThe widths of naphthochromenone rings (width2, W2).

a c

Figure 1. Molecular dimensions of synthetic flavones and nomenclature of ring systems (a) and ketoconazole (b). Synthesis of Flavone. Among the 37 flavones in this study, 34 were prepared by chemical synthesis. Briefly, 2′-hydroxychalcone was obtained from benzaldehyde and 2′-hydroxyacetophenone under alkaline condition. Then, the desired flavone was synthesized by iodine-catalyzed cyclization. In some cases, flavone was prepared from benzoic acid and 2′-hydroxyacetophenone via β-diketones. Details of the synthetic procedures and analytical data are presented in the Supporting Information. Measurement and Estimation of Physicochemical Properties. For the correlations between the physicochemical properties of flavones and biotransformation to be demonstrated, some molecular properties were measured. Hydrophobicity (octanol−water partition coefficient, logP) was measured with a method from the literature.18 The torsional angles, lengths, and widths of flavones were calculated with HyperChemver 8.0.4 (Hypercube Inc.). Molar volumes of substrates and inhibitors were also calculated with the same software. The results are summarized in Table 1 and Figure 1. Detailed procedures and additional results are described in Figures S2 and S3. Kinetic Study of Flavone Biotransformation by Aspergillus niger. Aspergillus niger KACC 45093 (ATCC # 9029) was kindly provided by the National Agrobiodiversity center, RDA-genebank information center (Jeonju, Korea). The fungal seed culture was routinely grown on PDB for 3 days at 28 °C and 200 rpm. For the degradation kinetics study, the mycelium from seed culture (0.3 g, fresh weight) was added to freshly sterilized PDB (200 mL). Then,

aliquots (0.5 mL) of flavone solution (20 mg/10 mL in dimethyl sulfoxide, DMSO) were treated. The cultures were further cultivated at 28 °C and 200 rpm. For each flavone, triplicate samples were prepared, and the profiles of parent flavones and metabolites were analyzed at defined periods (0, 1, 3, 5, 7, and 14 days). For control experiments, 7 day cultures were sterilized at 110 °C for 30 min, and flavone solutions were treated as described above. After 7 days, the concentrations of flavones were analyzed. Solvent (DMSO, 0.5 mL) control was also prepared according to the same procedure. The degradation rates and half-lives were calculated from the pseudo-first-order reaction model (exponential decay model). Effects of Metabolic Inhibitors. For the metabolic inhibitor study, piperonyl butoxide and ketoconazole were selected as cytochrome P450 (CYP) inhibitors, and methimazole was used as a flavin-dependent monooxygenase (FMO) inhibitor. The mycelium from seed culture of A. niger KACC 45093 (0.3 g, fresh weight) was added to freshly sterilized PDB (200 mL) followed by stock solutions of metabolic inhibitors (500 mg/10 mL DMSO). The concentrations of inhibitors were set to 0.1, 1, and 10 mg/200 mL of PDB. After 6 h of preincubation at 28 °C and 200 rpm, a small amount (0.5 mL) of flavone solutions (20 mg/10 mL, DMSO) was added. Flavone (1) and 4′-methylthioflavone (16) were used as test chemicals. The mixture was incubated under the same conditions. The amount of parent and metabolites were measured after 7 days of incubation. B

DOI: 10.1021/acs.jafc.7b00390 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Article

Journal of Agricultural and Food Chemistry Extraction of Metabolites and Instrumental Analysis. The cultures, including both mycelia and medium, were treated with diluted HCl (1N, 15 mL). After maceration in a blender for 2 min, the media was filtered. The filter cake was extracted with MeOH (100 mL × 2). The aqueous filtrate and MeOH extracts were combined, and MeOH was removed under reduced pressure. The aqueous solution was saturated with NaCl. Parent flavones and metabolites were extracted with ethyl acetate (EA, 100 mL × 2). The EA extracts were dried over anhydrous Na2SO4 and concentrated to dryness. The residue was redissolved in EA (10 mL) and used for gas chromatography−mass spectrometry (GC-MS). For GC-MS, the extracts were derivatized by the following procedure. A small amount of extracts (1 mL) was evaporated under reduced pressure. The residue was dissolved in dry pyridine (0.9 mL) and mixed with BSTFA-TMCS (0.1 mL). The mixture was heated to 75 °C for 1.5h. The resulting mixture was analyzed by GC-MS. GC-MS Analysis of Parent Flavones and Metabolites. Metabolites were analyzed with a gas chromatograph−mass spectrometer (GC-MS, Shimadzu GC-2010 with GCMS-2010 SE) equipped with Rtx-5MS column (30 m, 0.25 μm film thickness, 0.25 nm i.d.; Resteck, USA). Helium was the carrier gas at a flow rate of 1 mL/min. The column temperature for metabolite analyses were programmed as follows: 160 °C (10 min) and raised to 295 °C at a rate of 2.5 °C/min and held for 30 min. The mass spectra (MS) of metabolites were obtained in full scan mode.

Biotransformation Rates of Flavones. The degradation rate constants and half-lives (T1/2) of flavones were 0.002− 0.521/day and 1.3−346.5 days, respectively (Table 2). In Table 2. Biomass of Aspergillus niger and Degradation Kinetic Parameters of Flavones

■

RESULTS Metabolic Inhibitors on the Degradation of Selected Flavones. A. niger KACC 45093 was preincubated with three metabolic inhibitors (piperonyl butoxide, ketoconazole, or methimazole) followed by flavone (1) and 4′-methylthioflavone (16). Ketoconazole, a fungicide, showed concentration-dependent growth inhibition, whereas piperonyl butoxide and methimazole did not inhibit the mycelial growth of A. niger (Figure 2). The biodegradations of flavones 1 and 16 were

Figure 2. Mycelial fresh weight of Aspergillus niger (a) and percent of remaining flavone (1) and 4′-methylthioflavone (16) (b) after 7 days of incubation with different concentrations of inhibitors in three replicates. The number indicates the concentration, i.e., P0.1 for 0.1 mg/200 mL of piperonylbutoxide. Different letters over each bar are significantly different at P < 0.05. C, solvent control; P, piperonylbutoxide; K, ketoconazole; M methimazole.

ID

substituents

C 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37

control all H 2′-Me 2′-MeO 2′-MeS 2′-Cl 2′-I 3′-Me 3-Me 3′-MeO 3′-F 3′-Cl 3′-I 4′-Me 4′-CF3 4′-MeO 4′-MeS 4′-F 4′-Cl 4′-tBu 5-MeO 6-Cl 7-F 7-Br 2′,3′-diMeO 2′,3′,-diCl 6-Cl-4′-MeO 3′,4′-diMeO 3′,4′-methylenedioxy 3′,5′-diMe 3′,5′-diF 3′,5′-diCl 3,4,5-triMeO α-naphthoflavone β-naphthoflavone γ-naphthoflavone 2-naphthyl analogue 4-pyridyl analogue

biomass (g)a 2.59 2.45 2.27 2.10 3.08 2.61 2.48 2.56 2.08 2.56 2.19 2.05 2.54 2.78 2.12 2.89 2.95 1.95 1.89 2.32 2.88 2.45 2.32 2.81 2.59 2.19 2.34 2.88 2.12 2.39 2.24 2.15 2.45 2.16 2.45 2.00 2.27 1.23

± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ± ±

0.89 1.12 1.05 0.54 1.21 1.02 0.98 0.45 0.89 1.12 1.13 0.75 0.97 1.42 0.44 1.08 1.18 0.42 1.02 0.88 1.54 1.07 0.79 1.05 1.25 1.27 1.45 0.32 1.27 0.85 1.31 1.12 0.74 1.12 0.45 1.45 1.11 0.45

rate constant (r2)b

T1/2 (day)

0.362 0.240 0.115 0.141 0.097 0.028 0.090 0.035 0.116 0.123 0.088 0.043 0.178 0.013 0.434 0.521 0.004 0.006 0.066 0.206 0.105 0.361 0.078 0.049 0.079 0.044 0.199 0.043 0.070 0.031 0.009 0.091 0.009 0.002 0.021 0.022 0.021

1.9 2.9 6 4.9 7.1 24.8 7.7 19.8 6 5.6 7.9 16.1 3.9 53.3 1.6 1.3 173.3 115.5 10.5 3.4 6.6 1.9 8.9 14.1 8.8 15.8 3.5 16.1 9.9 22.4 77 7.6 77 346.5 33 31.5 33

(0.875) (0.912) (0.937) (0.752) (0.887) (0.815) (0.943) (0.753) (0.751) (0.919) (0.694) (0.944) (0.915) (0.677) (0.792) (0.857) (0.579) (0.649) (0.793) (0.891) (0.922) (0.917) (0.795) (0.872) (0.887) (0.906) (0.818) (0.759) (0.844) (0.797) (0.714) (0.899) (0.715) (0.478) (0.736) (0.881) (0.815)

a

Fresh weight of mycelium after 7 days of culture. bValues in parentheses are regression coefficients (r2) from the exponential decay model.

general, nonsubstituted flavone (1) and flavones with alkyl, methoxy, and methylthio substituents were rapidly transformed to multiple metabolites. However, the rates of flavones with halogens (F, Cl, Br, or I) and trifluoromethyl (CF3) group were highly variable, depending on the position of substituents. For example, the T1/2 values of 5, 11, and 21 (2′-Cl, 3′-Cl, and 6-Cl flavones, respectively) were 7.1, 7.9, and 6.6 days, respectively. However, 4′-chloroflavone (18) was very stable to biodegradation (T1/2 = 115.5). Monofluorinated flavones (10, 17, and 22) have also shown similar position-dependent metabolism. Only a small portion of 4′-fluoroflavone (22) disappeared from cultures after 14 days, whereas 3′-fluoro- (10) and 7-

strongly inhibited by CYP inhibitors, whereas the inhibitory effect of methimazole, an FMO inhibitor, was evident only at high concentration. At the highest concentration (10 mg/200 mL media), the percent inhibitions of biodegradation (1 and 16) were 75−98% by CYP inhibitors, whereas the residual flavones in methimazole-treated cultures were 60−70% of the initial concentration. Among the flavones, the biotransformation of 1 was strongly inhibited by both CYP inhibitors, whereas the oxidation of 16 was more efficiently inhibited by ketoconazole (Figure 2). C

DOI: 10.1021/acs.jafc.7b00390 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Article

Journal of Agricultural and Food Chemistry

Figure 3. Total ion chromatogram (a) of 3-day culture extracts treated with flavone (1) after TMS-derivatization and mass spectra of selected metabolites 1-1 (b), 1-2 (c), and 1-3 (d). Asterisks (*) are diagnostic fragment ions.

fluoroflavone (17) were rapidly transformed to various metabolites. In general, methyl-substituted flavones were susceptible to metabolism. However, the position of the methyl group is still an important factor for determining the rate. For example, 3-methylflavone (8) was degraded more slowly than others (2, 7, and 13). Flavone with a bulky t-butyl group (19) was also rapidly metabolized. In comparison with methyl flavones, a halogenated alkyl group (4′-CF3) of flavone (14) was not oxidized. MS of its major metabolite indicated hydroxylation on the A or B ring with intact CF3 group. In addition to the physicochemical properties of individual substituents, the degradation of substituted flavones seemed to be dependent on the three-dimensional shapes of substrates. The T1/2 values of flavones with multiple methyl or methoxy groups (24, 27, 29, and 32) were 3.5−14.1 days. In comparison with the 3′,5′-dimethyl analogue (29), dihalogenated flavones (30 and 31) were stable to biotransformation. Interestingly, 3′,4′-methylenedioxyflavone (28) was more resistant to biodegradation than its closely related analogue 3′,4′dimethoxyflavone (27). The transformations of naphthoflavones (33−36) were very slow: only 1−10% transformation after 14 days. 2-(4-Pyridinyl)-chromen-4-one (37), a pyridinyl analogue of flavone (1), was also rather stable to biotransformation. Although the concentration of parent gradually decreased, notable metabolites were not found. In comparison with others, 37 has shown interesting biological activity. The result of mycelial biomass indicated that this compound has a strong growth inhibitory effect (Table 2). It is noteworthy that 4′-halogenated flavones (17 and 18) have also shown mild growth inhibition. Metabolite Profiles. Numerous metabolites were found during the metabolism of flavones by A. niger KACC 45093 (Tables S3 and S5). The number of metabolites ranged from 1 to 11 depending on the substrates. Approximately six metabolites were found from the flavone (1)-treated cultures; from detailed mass spectral analysis, these were found to be

flavone derivatives with 1-3 hydroxy groups (Figure 3). Metabolite 1-1 was identified as 4′-hydroxyflavone with synthetic standard. Metabolites 1-2 and 1-3 were regioisomers with two hydroxy groups on flavone rings. MS of metabolite 13 contains a diagnostic fragment (m/z 208), which indicates the TMS-derivatized hydroxychromene ring, whereas the same fragment was not observed in MS of 1-2 (Figure 3). These findings indicate that the two hydroxyl groups of 1-2 are located in ring A, whereas the same functional groups are introduced to each ring (rings A and B) in metabolite 1-3. Similar diagnostic ions of metabolites were found from many flavones (Figure 4 and Table S3). Most methoxy-flavones (3, 9, 15, 20, 27, and 32) were rapidly metabolized into their hydroxyl metabolites. Among these, 15 and 27 (flavones with 4′-methoxy group) metabolized

Figure 4. Proposed ion fragmentations of TMS-derivatized flavone metabolites (a) 4′,6-dihydroxyflavone from 1, (b) 4′-chloro-6hydroxyflavone from 18, (c) 4′-hydroxy-a-naphthoflavone from 33, and (d) 7-hydroxy-2-(naphthalen-2-yl)-4H-chromen-4-one from 36. D

DOI: 10.1021/acs.jafc.7b00390 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Article

Journal of Agricultural and Food Chemistry

Figure 5. Total ion chromatogram of 3-day culture extracts treated with 4′-methylflavone (13, a) and 4′-methylthioflavone (16, b) after TMS derivatization and mass spectra of selected metabolites 13-1 (c), 13-2 (d), 16-1 (e), and 16-2 (f). Asterisks (*) are for parent flavones.

to their O-demethylated products as main metabolites (Tables S3 and S4). However, O-demethylation product was not observed from the cultures of 3′,4′,5′-trimethoxyflavone (32). MS of the most abundant metabolite of 32 has shown monohydroxychromene ring fragments. O-Demethylation was also a minor metabolism of 2′/3′-monomethoxyflavones (3 and 9) in which ring hydroxylation was the dominant reaction. Similarly, the methoxy group of 5-methoxyflavone (20) was not oxidized during the culture. 2′,3′-Dimethoxyflavone (24) was transformed to multiple metabolites, where the large portion of metabolites contains two methoxy groups (Tables S3 and S4). In the case of another 4′-methoxyflavone with chlorine (26), Odemethylated flavone was one of the most abundant metabolites. According to the spectral analysis, several interesting metabolites were observed during the metabolism of methylflavones (2, 7, 8, 13, and 29) and methylthio analogues (4 and 16) (Figure 5). For example, two metabolites (13-1 and 13-2) from 13 were characterized as 4′hydroxymethyl- and 4′-carboxy-flavones, respectively. Although there were several minor products, 16-1 and 16-2 were the major metabolites of 16. In consideration of molecular weights (284 and 300) and fragment ions (m/z 165 and 181), these metabolites were identified as 4′-methylsulfinylflavone (16-1) and 4′-methylsulfonylflavone (16-2), respectively. The metabolite profiles of other methylflavones (2, 7, 8, and 29) were

different from those of the 4′-methyl analogue (13). Flavones with a hydroxymethyl or carboxy group were not found from the cultures of 2′-methyl- and 3-methylflavone (2 and 8). In the case of 3′-methyl and 3′,5′-dimethylflavones (7 and 29), the major metabolites were hydroxymethylflavones (Tables S3 and S4). However, carboxy products were minor constituents in both flavones, and some metabolites with an intact methyl group were also found. It has to be mentioned that oxidation of both methyl groups of 29 was not observed even after 14 days of incubation. Two major metabolites (19-1 and 19-2) of 4′-tbutylflavone (19) were characterized as flavone with hydroxylated t-butyl group (19-1) and the ring hydroxylated product of 19-1. The MS of some minor metabolites (19-2 and 19-5) indicated that the products contain a carboxy group (Tables S3 and S4). In comparison with other alkyl flavones, 4′trifluoromethylflavone (14) was much more stable to biodegradation. A trace metabolite (14-1) was tentatively identified as B-ring monohydroxylated product by MS. In this study, 14 ring-halogenated flavones were tested (5, 6, 10−12, 17, 18, 21−23, 25, 26, 30, and 31). Their biodegradation rates were highly dependent on the position of substituents. As mentioned above, 4′-halogenated flavones were resistant to metabolism. In addition, the structures of major metabolites were also different from those of other analogues. For example, the MS of metabolites 17-1 and 18-1 from 17 and 18 have E

DOI: 10.1021/acs.jafc.7b00390 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Article

Journal of Agricultural and Food Chemistry shown the presence of m/z 208 fragment ion as well as ions with m/z 121 and 136 (Tables S3 and S4). These ions are of TMS-hydroxychromene and halogenated phenyl rings (Figure 4). The characteristics suggested that these metabolites are Aring hydroxylation products. From the detailed analysis of chromatograms, dehalogenated products were not found from any halogenated flavones. The transformation of naphthoflavones (33−36) occurred very slowly (Table 1). Only 1-2 metabolites were observed at trace levels from the culture extracts (33-1, 34-1, 35-1, 36-1, and 36-2; Table S3). GC-MS of their MS have shown an interesting difference in hydroxylation position between 36 and others (Figure S4). A characteristic TMS-hydroxychromene fragment (m/z 208) was found from the MS of metabolite 36-1, whereas the MS of 33-1 and 35-1 has shown a fragment ion (m/z 170) derived from the nonsubstituted naphthochromene ring (Figure 4). In general, parent flavones were transformed to more polar metabolites by A. niger. However, several additional metabolisms, other than the oxidative mechanism, were also observed. Those were O-methylation and glycosidation. For example, small amounts of methoxy metabolites were observed during the metabolism of two flavones (3 and 5) (Table S3). Sack et al. reported that A. niger can introduce methoxy groups to aromatic hydrocarbons.19 Ten flavones (1, 4, 5, 9, 14, 15, 17, 18, 21, and 22) gave carbohydrate-conjugated metabolites (Table S3). Their MS indicated that the conjugates are comprised of hexose and flavones with 1-2 hydroxy groups (Table S3). Although A. niger is a well-known species with extracellular glycosidases, the results indicated that this organism can produce flavonoid glycosides. Metabolites from hydrolytic cleavage of flavone rings or chalcones were not observed in all flavone treatments.

Among several physicochemical properties of substrates, bioabsorption (translocation into the cell) is one of the most important determinants of intracellular biotransformation. LogPs of synthetic flavones were measured by a method presented in the literature (Table 1).18 Experimental results have shown that lipophilic flavones were transformed more slowly than that of their hydrophilic analogues. For example, the T1/2s of flavones with 3′-F (10), 3′-Cl (11), and 3′-I (12) were 5.6, 7.9, and 16.1 days, of which the logPs were 3.37, 3.80, and 4.56, respectively. Flavones with 2′- and 7′-halogens (5, 6, 22, and 23) also showed similar results. However, the relationships seemed to be limited to some analogues. 4′-tButylflavone (19) was much more lipophilic than βnaphthoflavone (34). However, its degradation was approximately 35 times faster than that of 34. These results indicate that several additional properties are also important for determining the degradation rates (e.g., geometic shapes, three-dimensional properties of enzymes).32 According to the instrumental analysis, the initial metabolism of flavone by A. niger occurs preferentially on the 4′ carbon of the B ring or the substituents on it. The oxidative reactions include ring hydroxylation, O-demethylation, methylsulfone/ sulfoxide formation, and oxidation of alkyls to hydroxyalkyl/ carboxy groups. Flavones with halogens or trifluoromethyl (CF3) at the same position were metabolically stable, whereas the 2′- and 3′-halogenated analogues rapidly disappeared from the cultures (Table 2). It is noteworthy that their metabolites still contain halogens and CF3 groups. Microbial dehalogenation is frequently observed during the biodegradation of chlorinated alkenes and aromatics. For example, pentachlorophenol was converted to its tetrachlorinated metabolite by Phanerochaete chrysosporium.33 CYP-catalyzed reductive dehalogenation was also reported from some Pseudomonad.34 However, the experimental results suggested that strain KACC 45093 does not have dehalogenase activity on ring halogens and the fluoroalkyl group. In addition to oxidative transformation, the nonoxidative metabolism of several flavonoids has been reported, including ring opening to chalcones and hydrolytic cleavage to benzoic acids.35 However, detailed analysis of culture extracts with strain KACC 45093 did not show any trace of such metabolites. The major initial metabolites of 2′-, 3′-, and 5-methoxyflavones (3, 9, and 20) were those with one hydroxy and one methoxy group, whereas 4′-hydroxyflavone was the predominant product from 4′-methoxyflavone (15). 4′-Methylthioflavone (16) was completely oxidized to 4′-methylsulfinyl- and 4′-methylsulfonylflavone. However, its 2′-analogue (4) was partially oxidized to give a complex mixture (2′-methylsulfinyl, 2′-methylsulfonyl, and many ring hydroxylated metabolites with an intact 2′-methylthio group). These findings suggested that there is a preferred metabolic position of flavone substrates. In other words, the 4′-carbon of the B ring may preferentially be directed to the catalytic site of oxidative enzymes. Similar metabolic selectivity on the 4′-position was also reported with several Aspergillus species and Cunninghamella elegans.35−37 The selectivity seemed to be preferential rather than exclusive. For example, the 2-naphthyl analogue (36) was transformed mainly to the A-ring hydroxyl product. Minor reactions in 2′-, 3′methoxy-, or methylthioflavone (3, 4, and 9) were additional examples of limited but possible metabolism at these positions. In addition to the substrate structures, the regioselectivity is also governed by the properties of enzymes.

■

DISCUSSION Metabolic inhibitor studies can provide valuable information to specify the enzymes in substrate biotransformation. Azolyl fungicides (e.g., ketoconazole) are competitive inhibitors of various CYPs, including mammalian and fungal CYPs, whereas piperonyl butoxide is a noncompetitive CYP inhibitor.20,21 The metabolisms of two flavones (flavone 1 and 4′-methylthioflavone 16) were almost completely inhibited by these compounds, whereas the inhibition by methimazole was limited. Methimazole is reported as a specific inhibitor of mammalian FMOs.22 Previous studies with the same set of inhibitors indicated that CYPs are major catabolic enzymes in xenobiotic biotransformation and that the contribution of FMO is limited.23−25 Similar results were also provided with many other inhibitors.26,27 In addition to CYPs and FMOs, several fungi (e.g., Aspergillus, Paecilomyces, and white rot fungi) are known to produce additional oxidative enzymes (e.g., laccase and peroxidases).28,29 Many recalcitrant compounds could be decomposed by these enzymes. For example, mushroom peroxidase can oxidize dibenzothiophene to sulfone/sulfoxide metabolites.30 Organic inhibitors of fungal laccase/lignin peroxidases are generally good metal chelators to deplete the metal ions (e.g., copper) from the catalytic center.31 However, there were no reports regarding metal chelation by the current set of inhibitors. In addition, oxidation of the sulfur atom in flavone 16 was strongly inhibited by CYP/FMO-specific inhibitors. These findings indicated that CYP/FMOs play a major role in flavone metabolism by A. niger rather than other oxidative enzymes. F

DOI: 10.1021/acs.jafc.7b00390 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Article

Journal of Agricultural and Food Chemistry

Figure 6. Summary of Aspergillus niger-catalyzed oxidative metabolism of synthetic flavones. The black bold line is for the major site of metabolism. Black dotted- and gray lines are sites of minor metabolism and no metabolic transformation, respectively.

and lengths were closely related. However, instrumental analysis showed that the primary metabolic reaction of 32 is chromenone hydroxylation, whereas O-demethylation was dominant in 24 and 27. This regioselectivity may suggest that the CYP catalytic center is not complementary with large width ligands. Compound 32 showed the largest (8.32 Å) width of ring B (W1), and the W 1s of 24 and 27 were 6.27 and 6.49 Å, respectively. The molecular widths (W1) of other readily metabolized flavones were within the range of 4.31(1) to 7.12 Å (4) (Figure 1 and Table 1). These results suggest that the catalytic center has limited geometric space, where ring B of 32 cannot be directed for metabolic conversion (Figure 6). In addition to the geometric shape, experimental results also provide information on electrostatic requirements for biotransformation. For example, geometric properties (e.g., logP, molar volumes, length, W1, and W2) of 3′,5′-dimethyl- and 3′,5′-dichloro-flavone (29 and 31) were very similar. 3′,5′Difluoroflavone (30) confers highly similar physicochemical properties with those of flavone (1). However, 30 and 31 were much more stable than 1 and 29 (Tables 1 and 2). The finding suggests that multiple electronegative substituents in the 3′and 5′-positions are detrimental to biotransformation. It is wellknown that there are strong correlations between the compound lipophilicities and their bioavailability-parabolic dependence with optimal logP for absorption.41 Because of the strong lipophilicity (and consequent low bioavailability), limited biodegradation was expected for the naphthoflavones (33−36) (Table 2). The experimental results supported such assumptions. The width of ring B (W1) of naphthoflavones (33−35) was 4.3 Å, which is quite similar to that of unsubstituted flavone (1). However, the width of the naphthochromenone ring (W2) of α-naphthoflavone (33) was much larger than that of others (Table 2). Van der Waals volumes of these were also much larger than 1. From these differences, it can be suggested that the limited biodegradation of these analogues results from the low bioavailability and structural limited complementarity with enzymes. In addition, it has to be mentioned that α-naphthoflavone (33) is a wellknown CYP inhibitor.42 Slow conversion of naphthoflavones may be attributed to both limited bioavailability and inhibitory effects. The results of naphthoflavone metabolism also give interesting information related to structural differences of

As mentioned above, approximately 150 CYPs were reported from the Aspergillus genome.10,11 However, detailed investigations of them were limited. In this study, the general features of these enzymes were postulated through structure− metabolism relationship analysis. Although isolation or functional identification of specific enzymes was not performed, the analysis will give outlines of CYPs, which are related to flavone oxidation. In general, steric characteristics of the binding pocket are one of the most important factors to cover the substrate diversities. For example, human CYP3A4 can metabolize an extremely large number of xenobiotics than other CYPs (e.g., CYP 1As and 2As). Their solvent accessible volumes were estimated as 1000−2000, 390, and 230−307 cubic angstroms (Å3) for CYP3A4, CYP1A2, and CYP2As, respectively.38 Among fungal CYPs, lanosterol 14-demethylases (CYP51s) are the most well-defined oxidative enzymes. A recent study proves that the binding site of CYP51 of A. flavus is large enough to fit many bulky azole fungicides.39,40 For example, the molar volume of ketoconazole was estimated as 457.53 Å3 by molecular simulation, whereas the values of substrates were 203.32 and 238.60 Å3 for 1 and 16, respectively (Table 1). It has to be mentioned that the inhibitory effects of ketoconazole result from the strong binding of this drug on the catalytic center of CYPs. The estimated molecular lengths of ketoconazole and piperonyl butoxide were approximately 19− 20 Å. According to the molecular simulation of ketoconazole, the width between the imidazole pharmacophore and 2,4dichlorophenyldioxolanyl group was 8.91 Å (Figure 1). The lengths and widths of flavones were smaller than those of ketoconazole at 10.17−14.01 and 4.16−8.32 Å, respectively (Table 1). In consideration of the common binding conformations of ketoconazole with various CYPs and the inhibitory activity against flavone oxidation, some conclusions can be proposed (Figure 6). The first structural requirements are related to the geometric shape of flavones (e.g., molecular width and length). For example, the lipophilicities of 15 and 26 are very similar. However, the half-life of 26 was 10-times longer than that of 15. Notable differences between 26 and 15 were the molecular lengths of 14.01 vs 13.41 Å, respectively (Tables 1 and 2). The metabolic characteristics of 2′,3′-, 3′,4′dimethoxy-, and 3′,4′,5′-trimethoxyflavones (24, 27, and 32) were expected to be very similar because their lipophilicities G

DOI: 10.1021/acs.jafc.7b00390 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Article

Journal of Agricultural and Food Chemistry

(10) Galagan, J. E.; Clavo, S. E.; Cuomo, C.; Ma, L. J.; Wortman, J. R.; Batzoglou, S.; Lee, S. I.; Baturkmen, M.; Spevak, C. C.; Clutterbuck, J.; Kapitonov, V. Sequencing of Aspergillus nidulans and comparative analysis with A. f umigatus and A. oryzae. Nature 2005, 438, 1105−1115. (11) Sun, J.; Lu, X.; Rinas, U.; Zeng, A. P. Metabolic peculiarities of Aspergillus niger disclosed by comparative metabolic genomics. Genome Biol. 2007, 8, R182. (12) Durairaj, D.; Hur, J.-S.; Yun, H. Versatile biocatalysis of fungal cytochrome P450 monooxygenases. Microb. Cell Fact. 2016, 15, 125. (13) Mazzaferro, L. S.; Hüttel, W.; Fries, A.; Müller, M. Cytochrome P450-catalyzed regio- and stereoselective phenol coupling of fungal natural products. J. Am. Chem. Soc. 2015, 137, 12289−12295. (14) Sanchez-Gonzalez, M.; Rosazza, J. P. N. Biocatalyticsynthesis of butein and sulfuretin by Aspergillus alliaceus. J. Agric. Food Chem. 2006, 54, 4646−4650. (15) Weis, R.; Winkler, M.; Schittmayer, M.; Kambourakis, S.; Vink, M.; Rozzell, J. D.; Glieder, A. A diversified library of bacterial and fungal bifunctional cytochrome P450 enzymes for drug metabolite synthesis. Adv. Synth. Catal. 2009, 351, 2140−2146. (16) Dutta, D.; Ghosh, D. K.; Mishra, A. K.; Samanta, T. B. Induction of benzo(a)pyrene hydroxylase in Aspergillus ochraceus TS: evidences of multiple forms of cytochrome P-450. Biochem. Biophys. Res. Commun. 1983, 115, 692−699. (17) Gharasoo, M.; Centler, F.; Van Cappellen, P.; Wick, L. Y.; Thullner, M. Kinetics of substrate biodegradation under the cumulative effects of bioavailability and self-Inhibition. Environ. Sci. Technol. 2015, 49, 5529−5537. (18) Rothwell, J. A.; Day, A. J.; Morgan, M. R. A. Experimental determination of octanol-water partition coefficients of quercetin and related flavonoids. J. Agric. Food Chem. 2005, 53, 4355−4360. (19) Sack, U.; Heinze, T. M.; Deck, J.; Cerniglia, C. E.; Cazau, M. C.; Fritsche, W. Novel metabolites in phenanthrene and pyrene transformation by Aspergillus niger. Appl. Environ. Microbiol. 1997, 63, 2906−2909. (20) Franklin, M. R. Inhibition of hepatic oxidative xenobiotic metabolism by piperonyl butoxide. Biochem. Pharmacol. 1972, 21, 3287−3299. (21) Greenblatt, D. J.; Zhao, Y.; Venkatakrishnan, K.; Duan, S. X.; Harmatz, J. S.; Parent, S. J.; Court, M. H.; von Moltke, L. L. Mechanism of cytochrome P450−3A inhibition by ketoconazole. J. Pharm. Pharmacol. 2011, 63, 214−221. (22) Nace, C. G.; Genter, M. B.; Sayre, L. M.; Crofton, K. M. Effect of methimazole, an FMO substrate and competitive inhibitor, on the neurotoxicity of 3,3′-iminodipropionitrile in male rats. Toxicol. Sci. 1997, 37, 131−140. (23) Buisson, D.; Quintin, J.; Lewin, G. Biotransformation of polymethoxylatedflavonoids: access to their 4′-O-demethylated metabolites. J. Nat. Prod. 2007, 70, 1035−1038. (24) Chen, Y.; Zhang, L.; Qin, B.; Zhang, X.; Jia, X.; Wang, X.; Jin, D.; You, S. An insight into the curdione biotransformation pathway by Aspergillus niger. Nat. Prod. Res. 2014, 28, 454−460. (25) Schlenk, D.; Bevers, R. J.; Vertino, A. M.; Cerniglia, C. E. P450 catalysed S-oxidation of dibenzothiophene by Cunninghamella elegans. Xenobiotica 1994, 24, 1077−1083. (26) Svobodová, K.; Mikesková, H.; Petráčková, D. Fungal microsomes in a biotransformation perspective:protein nature of membrane-associated reactions. Appl. Microbiol. Biotechnol. 2013, 97, 10263−10273. (27) Zhu, Y. J.; Keum, Y. S.; Yang, L.; Lee, H.; Park, H.; Kim, J. H. Metabolism of a fungicide mepanipyrim by soil fungus Cunninghamella elegans ATCC36112. J. Agric. Food Chem. 2010, 58, 12379−12384. (28) Tamayo-Ramos, J. A.; van Berkel, W. J.; de Graaff, L. H. Biocatalytic potential of laccase-like multicopper oxidases from Aspergillus niger. Microb. Cell Fact. 2012, 11, 165. (29) Viswanath, B.; Rajesh, B.; Janardhan, A.; Kumar, A. P.; Narasimha, G. Fungal laccases and their applications in bioremediation. Enzyme Res. 2014, 2014, 1.

mammalian and fungal CYPs. For example, mammalian CYPs commonly oxidize the chromone rings rather than phenyl substituents. However, a recent study with Aspergillus and Verticillium indicates that the phenyl ring was more easily oxidized than chromone rings.42,43 In conclusion, structure−flavone metabolism relationship analysis was performed with Aspergillus niger. The results provided several important structural features of flavone metabolism by this microorganism, including the preference of the 4′-position of ring B, various oxidative reactions, except dehalogenation, and structural outlines of fungal CYP, which is responsible for flavone metabolism. In consideration of the industrial importance of this microorganism, the results may give valuable information on structural studies of CYPs and possible industrial applications.

■

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jafc.7b00390. Synthetic methods, physicochemical properties of flavones, and GC-MS total ion chromatograms and MS of flavones and their microbial metabolites (PDF)

■

AUTHOR INFORMATION

Corresponding Author

*Tel: 82-2-450-3758. Fax: 82-2-450-3726. E-mail: rational@ konkuk.ac.kr. Funding

This work was supported by Konkuk University, research fund # 2014-A019-0044. Notes

The authors declare no competing financial interest.

■

REFERENCES

(1) Samson, R. A.; Visagie, C. M.; Hong, S. B.; Hubka, V.; Klaassen, C. H. W.; Perrone, G.; Seifert, K. A.; Susca, A.; Tanney, J. B.; Varga, J.; Kocsube, S.; Szigeti, G.; Yaguchi, T.; Frisvad, J. C. Phylogeny, identification and nomenclature of the genus Aspergillus. Stud. Mycol. 2014, 78, 141−173. (2) Domaradzka, D.; Guzik, U.; Wojcieszynska, D. Biodegradation and biotransformation of polycyclic non-steroidal anti-inflammatory drugs. Rev. Environ. Sci. Bio/Technol. 2015, 14, 229−239. (3) Bhalerao, T. S.; Puranik, P. R. Biodegradation of organochlorine pesticide, endosulfan, by a fungal isolate. Int. Biodeterior. Biodegrad. 2007, 59, 315−321. (4) Saratale, G.; Kalme, S.; Bhosale, S.; Govindwar, S. Biodegradation of kerosene by Aspergillus ochraceus NCIM-1146. J. Basic Microbiol. 2007, 47, 400−405. (5) Ye, J. S.; Yin, H.; Qiang, J.; Qin, H. M.; Zhang, N.; He, B. Y. Biodegradation of anthracene by Aspergillus f umigatus. J. Hazard. Mater. 2011, 185, 174−181. (6) De Vries, R. P.; Visser, J. Aspergillus enzymes involved in degradation of plant cell wall polysaccharides. Microbiol. Mol. Biol. Rev. 2001, 65, 497−522. (7) Parshikov, I. A.; Sutherland, J. B. Biotransformation of steroids and flavonoids by cultures of Aspergillus niger. Appl. Biochem. Biotechnol. 2015, 176, 903−923. (8) Sakai, S. Degradation of the plant flavonoid phellamurin by Aspergillus niger. Appl. Environ. Microbiol. 1977, 34, 500−505. (9) Tahara, S.; Ingham, J. L.; Mizutani, J. Substrate specificity in the fungal metabolism of prenylated flavonoids. Z. Naturforsch C 1991, 46, 341−348. H

DOI: 10.1021/acs.jafc.7b00390 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Article

Journal of Agricultural and Food Chemistry (30) Aranda, E.; Kinne, M.; Kluge, M.; Ullrich, R.; Hofrichter, M. Conversion of dibenzothiophene by the mushrooms Agrocybe aegerita and Coprinellus radians and their extracellular peroxygenases. Appl. Microbiol. Biotechnol. 2009, 82, 1057−1066. (31) Call, H. P.; Mücke, I. History, overview and applications of mediated lignolytic systems, especially laccase-mediator-systems (Lignozym®-process). J. Biotechnol. 1997, 53, 163−202. (32) Athersuch, T. J.; Wilson, I. D.; Keun, H. C.; Lindon, J. C. Development of quantitative structure-metabolism (QSMR) relationships for substituted anilines based on computational chemistry. Xenobiotica 2013, 43, 792−802. (33) Mileski, G. J.; Bumpus, J. A.; Jurek, M. A.; Aust, S. D. Biodegradation of pentachlorophenol by the white rot fungus Phanerochaete chrysosporium. Appl. Environ. Microbiol. 1988, 54, 2885−2889. (34) Castro, C. E.; Yokoyama, W. H.; Belser, N. O. Biodehalogenation; reductive reactivities of microbial and mammalian cytochrome P450 compared with heme and whole-cell models. J. Agric. Food Chem. 1988, 36, 915−919. (35) Das, S.; Rosazza, J. P. N. Microbial and enzymatic transformations of flavonoids. J. Nat. Prod. 2006, 69, 499−508. (36) Cao, H.; Chen, X.; Jassbi, A. R.; Xiao, J. Microbial biotransformation of bioactive flavonoids. Biotechnol. Adv. 2015, 33, 214−223. (37) Ibrahim, A. R. S. Biotransformation of chrysin and apigenin by Cunninghamella elegans. Chem. Pharm. Bull. 2005, 53, 671−673. (38) Gay, S. C.; Roberts, A. G.; Halpert, J. R. Structural features of cytochromes P450 and ligands that affect drug metabolism as revealed by x-ray crystallography and NMR. Future Med. Chem. 2010, 2, 1451− 1468. (39) Hargrove, T. Y.; Wawrzak, Z.; Lamb, D. C.; Guengerich, F. P.; Lepesheva, G. I. Structure-functional characterization of cytochrome P450 sterol alpha-demethylase (Cyp51B) from Aspergillus f umigatus and molecular basis for the development of antifungal drugs. J. Biol. Chem. 2015, 290, 23916−23934. (40) Monk, B. C.; Tomasiak, T. M.; Keniya, M. V.; Huschmann, F. U.; Tyndall, J. D.; O’Connell, J. D.; Cannon, R. D.; McDonald, J. G.; Rodriguez, A.; Finer-Moore, J. S.; Stroud, R. M. Architecture of a single membrane spanning cytochrome P450 suggests constraints that orient the catalytic domain relative to a bilayer. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 3865−3870. (41) Tute, M. S. Lipophilicity: a history. In Lipophilicity in drug action and toxicology; Pliska, V., Testa, B., van de Waterbeemd, H., Eds.; VCH: New York, NY, 1996; Vol. 4, pp 7−24. (42) Koley, A. P.; Buters, J. T.; Robinson, R. C.; Markowitz, A.; Friedman, F. K. Differential mechanisms of cytochrome P450 inhibition and activation by alpha-naphthoflavone. J. Biol. Chem. 1997, 272, 3149−3152. (43) Poplonski, J.; Sordon, S.; Tronina, T.; Bartmanska, A.; Huszcza, E. Fungal metabolism of naphthoflavones. J. Mol. Catal. B: Enzym. 2015, 117, 1−6.

I

DOI: 10.1021/acs.jafc.7b00390 J. Agric. Food Chem. XXXX, XXX, XXX−XXX