Nutrient Metabolism
Microbial Aromatic Acid Metabolites Formed in the Gut Account for a Major Fraction of the Polyphenols Excreted in Urine of Rats Fed Red Wine Polyphenols1 Marie-Paule Gonthier,2 Ve´ronique Cheynier,* Jennifer L. Donovan, Claudine Manach, Christine Morand, Isabelle Mila,† Catherine Lapierre,† Christian Re´me´sy and Augustin Scalbert Unite´ des Maladies Me´taboliques et Micronutriments, INRA Theix, 63122 Saint-Gene`s-Champanelle, France; *UMR Sciences pour l’Oenologie, INRA, 34060 Montpellier, France; and †Laboratoire de Chimie Biologique, INRA, Institut National Agronomique Paris-Grignon, 78850 Thiverval-Grignon, France ABSTRACT The health effects of dietary polyphenols might be explained by both intact compounds and their metabolites formed either in the tissues or in the colon by the microflora. The quantitative importance and biological activities of the microbial metabolites have seldom been examined in vivo. We measured the microbial metabolites formed in four groups of rats (n ⫽ 8) fed for 8 d a diet supplemented with 0.12 g/100 g catechin, 0.25 or 0.50 g/100 g red wine powder containing proanthocyanidins, phenolic acids, flavanols, anthocyanins and flavonols or an unsupplemented diet. Fourteen aromatic acid metabolites were assayed in urine collected for 24 h by an HPLC-electrospray ionization (ESI)-mass spectrometry (MS)-MS method. The three main metabolites formed from the catechin diet were 3-hydroxyphenylpropionic acid, 3-hydroxybenzoic acid and 3-hydroxyhippuric acid. Their total urinary excretion accounted for 4.7 g/100 g of the catechin ingested and that of intact catechins for 45.3 g/100 g. For wine polyphenols, the same microbial metabolites as observed for the catechin diet were identified in urine along with hippuric, p-coumaric, vanillic, 4-hydroxybenzoic and 3-hydroxyphenylacetic acids. All together, these aromatic acids accounted for 9.2 g/100 g of the total wine polyphenols ingested and intact catechins for only 1.2 g/100 g. The higher excretion of aromatic acids by rats fed wine polyphenols is likely due to their poor absorption in the proximal part of the gut. Some of the microbial metabolites still bear a reducing phenolic group and should also prevent oxidative stress in inner tissues. More attention should be given in the future to these microbial metabolites and their biological properties to help explain the health effects of polyphenols that are not easily absorbed through the gut barrier. J. Nutr. 133: 461– 467, 2003. KEY WORDS:
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polyphenols
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catechin
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wine
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gut microflora
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microbial metabolites
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rats
ability. The concentration of intact polyphenols (parent compounds and their tissular conjugated forms) in plasma rarely exceeds 1 mol/L, and their urinary recovery ranges from 1 to 25% of the ingested dose (2). However, some of the ingested polyphenols, not absorbed or excreted in the bile, reach the colon where they are extensively metabolized by the microflora into various aromatic acids (13–16). These metabolites are mainly derivatives of phenylpropionic, phenylacetic and benzoic acids with different hydroxylation patterns. Their microbial origin was established by in vitro degradation of various polyphenols with fecal microflora (17,18) and by in vivo experiments with rats administered antibiotics (19) or germfree rats (20). They are easily absorbed through the colonic barrier and can be further transformed in tissues by conjugation with glycine, glucuronic acid or sulfate groups (21,22). These different metabolites circulating in the body may also contribute to the health effects of dietary polyphenols. The yields of such microbial metabolites can be high, but their quantitative importance has seldom been assessed in vivo through comparison with intact phenolic precursors (22–24). In the present study, we fed rats diets supplemented with
Polyphenols are naturally occurring compounds widely distributed in higher plants, making them an integral part of the human diet. They are provided mainly by fruits, vegetables and some beverages (tea, coffee, fruit juices, red wine). The dietary intake of polyphenols has been estimated at 1 g/d (1,2). Dietary polyphenols are of interest due to their potential beneficial effects on human health. Polyphenols are the most abundant antioxidants in our diets and their consumption improves the antioxidant status in humans (3– 6). They may therefore contribute to the prevention of diseases associated with oxidative stress such as cancers and cardiovascular diseases. The results of a number of epidemiologic studies have shown an inverse correlation between the consumption of polyphenol-rich foods or beverages and the incidence of these diseases (7–12). Biological effects of polyphenols depend on their bioavail-
1 Supported by the European Community (POLYBIND contract QLK1–199900505). 2 To whom correspondence should be addressed. E-mail:
[email protected].
0022-3166/03 $3.00 © 2003 American Society for Nutritional Sciences. Manuscript received 8 July 2002. Initial review completed 5 August 2002. Revision accepted 5 November 2002. 461
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either a polyphenol-rich wine extract or pure catechin and quantified the main aromatic acid metabolites excreted in urine. Their relative abundance is compared to that of intact catechins (parent catechin and its tissular methylated forms), and their possible biological importance is discussed. MATERIALS AND METHODS Chemicals. (⫹)-Catechin hydrate, 3-hydroxybenzoic acid, 4-hydroxybenzoic acid, hippuric acid, vanillic acid, phenylacetic acid, 3-hydroxyphenylacetic acid, 3,4-dihydroxyphenylacetic acid, caffeic acid, ferulic acid, p-coumaric acid, syringic acid and Folin reagent were purchased from Sigma Chemical (St. Louis, MO); 3-hydroxyphenylpropionic acid and 3,4-dihydroxyphenylpropionic acid from Apin Chemicals Limited (Abingdon, UK) and taxifolin from Extrasynthese (Genay, France). 3-Hydroxyhippuric acid and 4-hydroxyhippuric acid were kindly provided by P.C.H. Hollman (RIKILT, Wageningen University, The Netherlands) and R. Scheline (University of Bergen, Norway), respectively. The 3⬘- and 4⬘-O-methylated conjugates of catechin were synthesized by in vitro methylation of catechin as previously described (25). Polyphenol-rich wine extract. Cabernet-Sauvignon wine (2500 L) was fractionated on a vinyl-divinyl benzene resin column (180 L). A water wash (400 L) was performed to discard residual sugars and other polar compounds (organic acids, proteins, polysaccharides). Polyphenols were eluted with ethanol. The water eluate was concentrated to a volume of 200 L and treated similarly on the same column. The two ethanol eluates were combined and spray-dried. The whole procedure was repeated 3 times. A total of 23 kg of polyphenol-rich wine powder was obtained. Phenolic acids, flavonols and anthocyanins were analyzed by HPLC-diode array detection (DAD)3 (26). Proanthocyanidins were characterized and quantified by thiolysis. Thioethers (formed from the upper and extension units) and flavanol monomers (end units) were analyzed by HPLC-DAD (26). The total amount of polyphenols identified accounted for one fifth of the wine powder and for two thirds of the total phenols as estimated by the Folin-Ciocalteu assay (Table 1). Animals and diets. Male Wistar rats (n ⫽ 32; Iffa Credo, L’Arbresle, Lyon, France) weighing 160 ⫾ 0.2 g at the beginning of the experiment were housed singly in metabolic cages in a temperature-controlled room (22°C) and maintained in a normal light:dark cycle (dark from 2000 to 0800 h) with free access to food from 1600 to 0800 h. After 14 d of adaptation to a semipurified diet (Table 2), rats were randomly divided into four groups of 8 rats and given the following four different diets for 8 d: the control semipurified diet or the same diet supplemented with 0.12 g/100 g catechin, 0.25 or 0.50 g/100 g wine powder. Animals were maintained and handled according to the recommendations of the Institutional Ethic Committee (INRA), in accordance to the decree N° 87– 848. Sampling procedure. Urine (24 h) was collected during the entire polyphenol diet period and stored at ⫺20°C. The excretion kinetics of metabolites over the 8-d diet period were determined from urine samples obtained by pooling urine aliquots collected from the 8 rats of each dietary group at a given time. The aliquot volumes were determined in proportion to the total daily volume of urine excreted by each rat. For plasma sampling, rats were anesthetized with pentobarbital (40 mg/kg body, intraperitoneally) 12 h after the beginning of the last meal. Blood was drawn from the abdominal aorta into heparinized tubes and centrifuged (10,000 ⫻ g, 2 min) to obtain plasma. Polyphenols are unstable when the pH of plasma is higher than 7.4; therefore, to prevent a drift of the pH due to gradual bicarbonate decomposition, plasma samples were immediately acidified with 10 mmol/L acetic acid (27). Aliquots of plasma were kept at ⫺20°C until analysis. Gas chromatography-mass spectrometric (GC-MS) identification of wine polyphenol and catechin metabolites. Urine samples (2 mL) were hydrolyzed by -glucuronidase (type H3, 125 kU/L, Sigma
3 Abbreviations used: DAD, diode array detection; ESI, electrospray ionization; GC-MS, gas chromatography-mass spectrometric;
TABLE 1 Polyphenol composition of the wine powder mg/g powder Flavanol monomers1 Catechin Epicatechin Flavonols1 Myricetin 3-O-glucoside Quercetin 3-O-glucoside Myricetin Quercetin Anthocyanins1 Delphinidin 3-O-glucoside Cyanidin 3-O-glucoside Petunidin 3-O-glucoside Peonidin 3-O-glucoside Malvidin 3-O-glucoside Malvidin 3-p-coumaroylglucoside Peonidin 3-O-p-glucoside Other acylated anthocyanins Phenolic acids1 t-Caftaric acid t-Coutaric acid Gallic acid Proanthocyanidins1,2 Total phenols3
8.4 7.0 0.2 0.5 1.3 2.2 1.5 0.4 1.3 1.2 7.2 1.9 3.1 2.5 6.9 2.0 5.6 146 299
1 Estimated by HPLC-diode array detection. 2 Average polymerization degree: 6.1; epigallocatechin/catechin
units: 3/17; extent of galloylation: 5.2%. 3 Measured by the Folin-Ciocalteu assay.
Chemical) and sulfatase (type V, 5 kU/L, Sigma Chemical) for 3 h at 37°C in a 0.1 mol/L sodium acetate buffer, pH 5. Methanol/water (5 mL; 80:20) was added to the samples and the mixture was centrifuged for 10 min at 2000 ⫻ g, 10°C. The supernatant was collected and methanol evaporated under vacuum. After acidification to pH 2 with 6 mol/L HCl, metabolites were extracted on a C18 solid phase cartridge (SepPak, Waters, Milford, MA) with methanol and acetonitrile. The extract was dried under vacuum and redissolved in acetonitrile. Polyphenol metabolites were silylated and analyzed by GC-MS as previously described (18). Compounds were identified by comparison of their MS spectra with those of the mass spectra library NIST92 (Varian, Les Ulis, France) and authentic reference standards. HPLC-electrospray ionization (ESI)-MS-MS analysis of aromatic acids. Urine samples (diluted in 0.1 mol/L sodium acetate buffer, pH 5, 175 L) containing syringic acid (3 mol/L) as an internal standard, were acidified to pH 4.9 with 20 L of 0.58 mol/L acetic acid and incubated at 37°C for 45 min in the presence of an Helix pomatia extract containing 1100 U -glucuronidase and 42 U sulfatase (Sigma Chemical). After acidification to pH 2 with 6 mol/L HCl, the urine was then extracted twice with ethyl acetate and centrifuged at 2400 ⫻ g for 10 min. The resulting supernatant was evaporated under nitrogen and then redissolved in 500 L of 25% aqueous methanol for HPLC-ESI-MS-MS analysis. HPLC-ESIMS-MS analyses were performed on a Hewlett-Packard HPLC system equipped with MS-MS detection (API 2000, Applied Biosystems, Toronto, Canada). The column was a Hypersil BDS C18 (5 m, 150 ⫻ 2.1 mm, Touzart & Matignon, Les Ulis, France) and the mobile phases consisted of 5% acetonitrile in 0.1% aqueous formic acid (solvent A) and 40% acetonitrile in 0.1% aqueous formic acid (solvent B). The following gradient was applied: 0 –15 min, linear gradient from 0% B to 100% B; 15–20 min: 100% B. The flow rate was 0.2 mL/min. Detection was carried out by using ESI conducted at 450°C in negative mode with a nebulizer pressure of 90 psi (630 kPa), a drying gas flow of 11 L/min, a fragmentor voltage of 20 V and a capillary voltage of 4000 V. The MS data were collected in multiple reaction monitoring mode, tracking the transition of parent and product ions specific to each compound with a dwell time of 500 ms. Aromatic acid metabolites and syringic acid (internal standard) were
BIOAVAILABILITY OF WINE POLYPHENOLS IN RATS
TABLE 2 Composition of diets Wine Wine Control powder powder Catechin diet 0.5 g/100 g 0.25 g/100 g 0.12 g/100 g g/kg dry feed Wheat starch Casein Peanut oil Mineral mixture1 Vitamin mixture2 Wine powder Catechin
755 150 50 35 10 — —
750 150 50 35 10 5 —
752.5 150 50 35 10 2.5 —
753.75 150 50 35 10 — 1.25
1 Mineral mixture AIN-93M (per kg of diet): CaHPO4, 18 g; K2HPO4, 3 g; KCl, 6 g; NaCl, 5 g; MgCl2, 2.5 g; Fe2O3, 3 mg, MnSO4, 150 mg; CuSO4, 125 mg; ZnSO4 䡠 7H2O, 120 mg; KI, 0.48 mg. 2 Vitamin mixture AIN-76A supplemented in choline (mg/kg of diet): thiamin, 15; riboflavin, 20; pyridoxine, 10; nicotinamide, 100; pantothenate, 70; folic acid, 5; biotin, 0.3; cyanocobalamin, 0.05; retinyl palmitate, 1.5; dl-␣-tocopheryl acetate, 125; cholecalciferol, 0.15; menadione, 1.5; ascorbic acid, 50; myo-inositol, 100; choline, 1360.
detected according to the respective m/z values of their parent and product ions: 3-hydroxybenzoic acid (137/93), 4-hydroxybenzoic acid (137/93), 3-hydroxyhippuric acid (194/150), 4-hydroxyhippuric acid (194/100), vanillic acid (167/123), phenylacetic acid (135/91), 3-hydroxyphenylacetic acid (151/107), 3,4-dihydroxyphenylacetic acid (167/123), 3-hydroxyphenylpropionic acid (165/121), 3,4-dihydroxyphenylpropionic acid (181/59), caffeic acid (179/135), ferulic acid (193/134), p-coumaric acid (163/119) and syringic acid (197/123). The inter- and intraday precision was 8.8 and 5.3%, respectively, and the mean accuracy was 2.3% (unpublished results). For quantification, calibration curves were prepared by spiking blank urine with aliquots of standard solutions and then plotting the peak areas of selected ions monitored against the corresponding calibration concentrations. The limit of detection (signal-to-noise ratio: 3) was in the range of 10 –200 nmol/L according to the nature of the compounds. HPLC-DAD analysis of hippuric acid. Urine and plasma samples (180 L) containing syringic acid (100 mol/L) as an internal standard, were acidified to pH 4.9 with 20 L of 0.58 mol/L acetic acid and incubated at 37°C for 45 min in the presence of an Helix pomatia extract containing 1100 U -glucuronidase and 42 U sulfatase (Sigma Chemical). Methanol/HCl (200 mmol/L, 500 L) was added to the samples and the mixture centrifuged for 4 min at 14,000 ⫻ g. The resulting supernatant was analyzed by reversed-phase HPLC on an Hypersil BDS C18 column (5 m, 150 ⫻ 4.6 mm, Life Sciences International, Cergy, France). The mobile phases consisted of 10% methanol in 0.5% aqueous formic acid (solvent A) and 80% methanol in 0.5% aqueous formic acid (solvent B). The separation was performed as follows: 0 –10 min, 0% B; 10 –25 min: linear gradient to 57% B; 25– 40 min: linear gradient to 100% B. The flow rate was 1 mL/min. Detection was carried out at 240 nm (hippuric acid) and 280 nm (internal standard) with a diode array detector (Kontron, Milan, Italy). The peaks were identified by comparison of retention times and UV spectra with those of authentic standards. The detection limit was 10 mol/L for all compounds. HPLC-ECD analysis of intact catechins. Both urine and plasma were assayed for intact catechins, i.e., parent catechin and its 3⬘-, and 4⬘-O-methylated derivatives, by HPLC analysis coupled to electrochemical detection (28). Detection was performed with an 8-electrode CoulArray Model 5600 system (Eurosep, Cergy-St-Christophe, France) with potentials set at 25, 100, 320, 400, 500, 700, 800 and 900 mV. The detection limit was at 20 nmol/L for all compounds. Folin-Ciocalteu determination of total phenols. The Folin reagent (diluted 1:10 in water, 750 L) and aqueous Na2CO3 (75 g/L, 600 L) were successively added to the wine powder extract dissolved in water (125 mg/L, 150 L). The mixture was kept in a water bath
463
at 50°C for 5 min, then chilled on ice before reading the absorbance at 760 nm. Catechin was used to establish the calibration curve and total phenol concentration was expressed as catechin equivalent (29). Aliquots (180 L) of urine collected over 24 h and pooled per group of animals as described above, were treated with 500 L methanol/200 mmol/L HCl and centrifuged for 4 min at 14,000 ⫻ g. The resulting supernatant (diluted 1:6 in water, 150 L) was then submitted to the Folin-Ciocalteu assay as described above. Data analysis. Data were entered into the Instat statistical analysis program (Instat, San Diego, CA). Comparisons of results were made by one-way ANOVA. Significant differences were determined by post-hoc analysis using Student-Newman-Keuls Multiple Comparison Test. Differences with P ⬍ 0.05 were considered significant. Numerical values are expressed as mean ⫾ SEM.
RESULTS The food intake and the weight gain of the rats were not different. Weight gain was 6.9 ⫾ 0.8 g/d. Rats consumed 19 ⫾ 2 g/d of the semipurified diets. Rats thus ingested 14.2 mg/d polyphenols (including 0.4 mg/d catechin) for the 0.25 g/100 g wine powder diet, 28.4 mg/d polyphenols (including 0.8 mg/d catechin) for the 0.50 g/100 g wine powder diet and 23.8 mg/d catechin for the 0.12 g/100 g catechin diet. Metabolites formed from the catechin diet. Several aromatic acid metabolites of catechin were identified in 24-h urine by GC-MS (Table 3) and quantified by HPLC-ESIMS-MS or HPLC-DAD (for hippuric acid) after enzymatic deconjugation. The major aromatic acid affected by the 8-d diet was 3-hydroxyphenylpropionic acid (Table 4). Urinary excretion of some benzoic acids (3-hydroxybenzoic acid, 3-hydroxyhippuric acid and to a lesser extent, 4-hydroxybenzoic acid) increased but not hippuric acid, the glycine conjugated form of benzoic acid. Rats fed the catechin diet had increased urinary excretion of ferulic acid and to a lesser extent of two dihydroxylated phenolic acids (3,4-dihydroxyphenylpropionic acid and 3,4-dihydroxyphenylacetic acid). The excretion of the various aromatic acids accounted for 4.7 g/100 g of the catechin ingested. GC-MS analysis enabled the identification of 3-hydroxyphenylvaleric acid and two lactones, 3,4-dihydroxyphenylvalerolactone and 3-methoxy-4-hydroxyphenylvalerolactone, in rat urine (Table 3). Traces of the opened forms of the lactones were also detected with 5-(4-hydroxy)-3-hydroxyphenylvaleric acid. However, the lack of corresponding standards did not allow their quantitation in the present study. Intact catechin and 3⬘-O-methylcatechin detected by GCMS (Table 3) were estimated by HPLC with multielectrode coulometric detection in both urine and plasma after enzymatic deconjugation. They were 5 and 8 times more abundant in urine than 3-hydroxyphenylpropionic acid, respectively (Table 4) and accounted together for 45.3 g/100 g of the catechin ingested. 4⬘-O-Methylcatechin was not detected. The urinary excretion of catechins was measured over the 8 d after the start of the catechin diet, in 24-h pooled urine (Fig. 1). It reached a plateau on d 1 of supplementation. Catechin and 3⬘-O-methylcatechin were also detected in plasma collected on the last day of the experiment, 12 h after the beginning of the 0.12 g/100 g catechin meal. Their respective concentrations were 1.3 ⫾ 0.3 and 8.0 ⫾ 1.1 mol/L. Metabolites formed from the wine polyphenol diets. The same aromatic acid metabolites as those formed with the catechin diet were observed in the urine of rats fed wine polyphenols: 3-hydroxyphenylpropionic acid, 3-hydroxybenzoic acid, 3-hydroxyhippuric acid, 4-hydroxybenzoic acid, feru-
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TABLE 3 Abbreviated mass spectra (electronic impact, 70 eV) used for the identification of metabolites (analyzed as their trimethylsilylated derivatives) present in urine of rats fed pure catechin or wine polyphenols1 Compounds
Mass
Benzoic acids 3-Hydroxybenzoic acid 4-Hydroxybenzoic acid Hippuric acid Vanillic acid Phenylacetic acids Phenylacetic acid 3-Hydroxyphenylacetic acid 3,4-Dihydroxyphenylacetic acid Phenylpropionic acids 3-Hydroxyphenylpropionic acid 3,4-Dihydroxyphenylpropionic acid Cinnamic acids Caffeic acid Ferulic acid Phenylpropanoic acid 3-Hydroxyphenylvaleric acid Phenylvalerolactones 3-Methoxy, 4-hydroxyphenylvalerolactone 3,4-Dihydroxyphenylvalerolactone Catechins Catechin 3⬘-O-Methylcatechin
Main ions (relative intensity) (M⫹, 19), 267 (100), 223 (20), 193 (23), 73 (24) (M⫹, 8), 267 (100), 223 (25), 193 (22), 73 (27) (M⫹ ⫺15, 32), 206 (95), 190 (15), 105 (100), 77 (65), 73 (60) (M⫹, 35), 297 (100), 282 (20), 267 (71), 253 (28), 223 (26), 73 (33)
282 282 251 312
282 282 236 312
208 296 384
193 (M⫹ ⫺15, 21), 164 (26), 137 (5), 118 (4), 91 (23), 73 (100) 296 (M⫹, 30), 281 (29), 252 (21), 179 (6), 164 (28), 147 (32), 105 (6), 73 (100) 384 (M⫹, 100), 369 (13), 340 (10), 267 (44), 237 (17), 179 (40), 73 (90)
310 398
310 (M⫹, 92), 205 (85), 192 (100), 177 (32), 73 (50) 398 (M⫹, 37), 341 (10), 280 (41), 267 (93), 191 (7), 147 (38), 73 (100)
396 338
396 (M⫹, 100), 381 (24), 307 (211), 219 (73), 73 (69) 338 (M⫹, 87), 323 (100), 308 (53), 293 (63), 279 (10), 249 (15), 219 (20), 73 (66)
338
338 (M⫹, 30), 306 (18), 224 (13), 248 (58), 411 (24), 73 (89)
294 352
294 (M⫹, 50), 279 (20), 264 (15), 247 (10), 209 (100), 205 (12), 179 (25), 149 (10), 73 (60) 352 (M⫹, 75), 337 (5), 267 (100), 247 (10), 205 (25), 179 (30), 149 (5), 147 (10), 73 (60)
650 592
650 (M⫹, 6), 383 (10), 368 (100), 355 (25), 73 (88) 592 (M⫹, 28), 383 (14), 355 (58), 310 (100), 280 (19), 209 (5), 73 (95)
1 All mass spectra were identified compared with mass spectral data banks or to authentic compounds, except for the valerolactone derivatives assigned from their mass fragmentation pattern only.
lic acid, 3,4-dihydroxyphenylpropionic acid and 3,4-dihydroxyphenylacetic acid (Table 4). The urinary excretion of several other aromatic acids not observed for the rats fed the catechin diet was induced by the wine polyphenol supplementation, i.e., vanillic acid, 3-hydroxyphenylacetic acid, p-coumaric acid, caffeic acid and hip-
puric acid. With the exception of caffeic acid, all of these metabolites showed a clear dose effect for the two (0.25 and 0.50 g/100 g) wine polyphenol diets. Hippuric acid was the major aromatic acid metabolite associated with wine polyphenol consumption. The kinetics of its excretion was also followed for the 8 d of the diet, in 24-h
TABLE 4 Urinary excretion of aromatic acids and catechins in rats after consuming a diet supplemented with a wine polyphenol extract or catechin for 8 d1 Control diet
0.25 g/100 g wine powder
0.50 g/100 g wine powder
0.12 g/100 g catechin
g/d 3-Hydroxybenzoic acid 4-Hydroxybenzoic acid Hippuric acid 3-Hydroxyhippuric acid 4-Hydroxyhippuric acid Vanillic acid Phenylacetic acid 3-Hydroxyphenylacetic acid 3,4-Dihydroxyphenylacetic acid 3-Hydroxyphenylpropionic acid 3,4-Dihydroxyphenylpropionic acid p-Coumaric acid Caffeic acid Ferulic acid Catechin 3⬘-O-Methylcatechin
ND2 59.3 ⫾ 4.6c 850.0 ⫾ 280c ND 72.6 ⫾ 12.8a 54.7 ⫾ 8.2c 280.6 ⫾ 70.1a 17.2 ⫾ 1.2c 7.4 ⫾ 0.5b 5.6 ⫾ 0.8d ND 3.4 ⫾ 0.9c ND 3.0 ⫾ 1.3c ND ND
4.3 ⫾ 0.4b 83.2 ⫾ 6.3b 1820.0 ⫾ 640b 12.8 ⫾ 1.5b 87.8 ⫾ 11.8a 111.6 ⫾ 15.4b 301.7 ⫾ 74.9a 39.1 ⫾ 3.7b 10.4 ⫾ 1.5b 89.8 ⫾ 9.8c 1.8 ⫾ 0.3b 46.4 ⫾ 3.7b 19.8 ⫾ 6.1a 14.3 ⫾ 1.1b ND ND
11.24 ⫾ 1.7b 134.2 ⫾ 4.6a 2790.0 ⫾ 230a 18.4 ⫾ 4.1b 98.7 ⫾ 10.8a 164.6 ⫾ 13.9a 382.8 ⫾ 74.4a 79.3 ⫾ 7.9a 13.0 ⫾ 1.1a 256.0 ⫾ 28.8b 4.7 ⫾ 1.1a 116.9 ⫾ 4.73a 20.0 ⫾ 1.5a 28.9 ⫾ 2.7a ND 330.0 ⫾ 41.2b
108.0 ⫾ 27.6a 88.9 ⫾ 9.2b 710.0 ⫾ 170c 133.4 ⫾ 44.6a 89.5 ⫾ 9.9a 57.7 ⫾ 7.3c 303.2 ⫾ 60.8a 23.9 ⫾ 1.8c 12.6 ⫾ 0.6a 823.5 ⫾ 222.1a 1.7 ⫾ 0.4b 3.6 ⫾ 0.7c ND 20.4 ⫾ 5.9b 3930.0 ⫾ 52.5 6870.1 ⫾ 91.2a
1 Values are means ⫾ SEM, n ⫽ 8. Means in a row without a common letter differ, P ⬍ 0.05. 2 ND, not detected (limits of detection for 3-hydroxybenzoic and 3-hydroxyhippuric acid: 0.2 g; 3,4-dihydroxyphenylpropionic acid: 0.1 g;
caffeic acid: 5 g; catechin and 3⬘-O-methylcatechin: 0.1 g).
BIOAVAILABILITY OF WINE POLYPHENOLS IN RATS
FIGURE 1 Urinary excretion of intact catechin metabolites in rats after consumption of wine powder and catechin diets. Values represent concentrations of catechin metabolites in 24-h urine pooled from 8 rats. C, catechin; 3⬘MC, 3⬘-O-methylcatechin.
pooled urine (Fig. 2). It differed from catechin in that it increased slowly, reaching a plateau after 5 d due to its microbial origin (30). The total excretion of aromatic acids accounted for 9.2 g/100 g of total wine polyphenols consumed, with a major contribution from hippuric acid (6.8 g/100 g). Hippuric acid was also determined in plasma. Its concentration increased significantly with the consumption of wine powder (Fig. 3). A significant enhancement of 29 and 50 mol/L was observed for rats fed the 0.25 and 0.50 g/100 g wine polyphenol diets, respectively, compared with the control or catechin-fed rats. Similar to rats fed the catechin diet, the same phenylvaleric acid and phenylvalerolactones were identified by GC-MS (Table 3) in rats fed wine polyphenols, but these were not quantified. Very low amounts of intact catechin, present essentially in its 3⬘-O-methylated form, were also detected in the urine of rats fed the 0.50 g/100 g wine polyphenol diet (Table 4). The excretion of 3⬘-O-methylcatechin accounted for 41.2 g/100 g of the catechin ingested, a value close to that determined for rats fed the catechin diet, and for only 1.2 g/100 g of the total wine polyphenols ingested. 3⬘-O-Methylcatechin was also detected in plasma (0.31 ⫾ 0.04 mol/L). The urinary and plasma concentrations of the 3⬘-O-methylcatechin in rats fed the 0.25 g/100 g wine polyphenol diet were too close to the limit of detection and could not be estimated reliably. Total reducing capacity of urine in rats fed the catechin and wine polyphenol diets. The Folin reagent was used to determine the total reducing capacity of 24-h pooled urine. This colorimetric assay is based on a redox reaction and can be used to estimate total polyphenols in foods, beverages or urine. In rats fed the catechin diet, there was a 1.5-fold increase in
FIGURE 2 Urinary excretion of hippuric acid in rats during 8 d of consumption of wine polyphenols or pure catechin. Values represent concentrations of hippuric acid in 24-h urine pooled from 8 rats.
465
FIGURE 3 Plasma concentrations of hippuric acid in rats fed wine polyphenols and catechin diets for 8 d. Values are means ⫾ SEM, n ⫽ 8. Means with different letters differ, P ⬍ 0.05.
the Folin value (⫹ 11.1 mg catechin equivalents/d) compared with that observed in rats fed the control diet (Fig. 4). The consumption of the wine polyphenol extract also induced a dose-dependent increase in the reducing capacity of the urine (⫹ 2.2 and 4.5 mg catechin equivalents/d in rats fed the 0.25 and 0.50 g/100 g wine polyphenol diets, respectively), although lower than for rats fed the catechin diet. DISCUSSION The quantitative importance and biological activities of microbial metabolites derived from dietary polyphenols have seldom been examined in vivo. In the present study, the urinary excretion and plasma levels of microbial aromatic acid metabolites were determined after feeding rats a diet supplemented with a polyphenol-rich wine extract or pure catechin. In rats fed the catechin diet, the main microbial aromatic acids excreted were 3-hydroxyphenylpropionic acid, and to a lesser extent 3-hydroxybenzoic acid and 3-hydroxyhippuric acid. These metabolites were previously identified but not quantified in the urine of rats and humans fed catechin (19,31). Traces of 4-hydroxybenzoic acid, 3,4-dihydroxyphenylpropionic acid, 3,4-dihydroxyphenylacetic acid and ferulic acid were also excreted in rats fed the catechin diet. 4-Hydroxylated metabolites have not often been described due to a preferential p-dehydroxylation by the microflora. The only 4-hydroxylated metabolite of catechin identified to date was 4-hydroxyphenylpropionic acid reported in an in vitro study with rat-cecal microflora (32). 3,4-Dihydroxyphenylpropionic
FIGURE 4 Total reducing capacity of urine in rats fed wine polyphenols or pure catechin. Values represent concentrations of total phenols in 24-h urine pooled from 8 rats.
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acid results from the incomplete dehydroxylation of catechin (32). The 3,4-dihydroxyphenylacetic acid should result from the ␣-oxidation of the previous compound. A similar ␣-oxidation, which is not a common pathway for plant polyphenols, has been reported for tyrosine (33). The origin of ferulic acid is less clear. Phenylvalerolactones and phenylvaleric acids are also well established metabolites of catechin. In humans, two derivatives of phenylvalerolactones, (-)-5-(3⬘,4⬘,5⬘-trihydroxyphenyl)-␥-valerolactone and (-)-5-(3⬘,4⬘-dihydroxyphenyl)-␥valerolactone, have been identified in urine and blood after tea consumption (34). Incubation of catechin with rat cecal microflora also led to the formation of traces of 3-hydroxyphenylvaleric and 3,4-dihydroxyphenylvaleric acids (35). In the present work, both phenylvalerolactones and phenylvaleric acids were identified by GC-MS analysis. A methylated derivative of phenylvalerolactone, 3-methoxy, 4-hydroxyphenylvalerolactone, was identified here as a metabolite of catechin for the first time. In rats fed the wine polyphenol diet, the same aromatic acids formed from catechin were identified in urine. However, the low catechin content of the wine polyphenol extract is not sufficient to account for the excretion levels of the different aromatic acids in urine. They most likely originate from the microbial metabolism of other flavonoids present in the wine polyphenol extract such as proanthocyanidins, flavonols or anthocyanins (Table 1). 3-Hydroxyphenylpropionic, 3-hydroxybenzoic, 3-hydroxyhippuric and 3,4-dihydroxyphenylpropionic acids likely originate from the metabolism of wine proanthocyanidins and to a lesser extent from that of catechin as suggested by previous in vitro studies with fecal microflora (15,18). 3,4-Dihydroxyphenylacetic acid likely results from the degradation of quercetin and 3-hydroxyphenylacetic acid from both proanthocyanidins and quercetin (16,18,24,36). Anthocyanins are also probably degraded into similar metabolites but their microbial metabolism has not yet been examined. Wine also contains caffeic and p-coumaric acids largely present as esters with tartaric acid. These are known as caftaric and coutaric acids, respectively (37). The caffeic and p-coumaric acids excreted in rats after wine polyphenol intake most likely derive from the hydrolysis of these esters in the gut. Microbial esterase activities have been reported in a human fecal slurry (38). Caffeic acid is further methylated into ferulic acid in the liver (39,40). Ferulic acid itself is further metabolized into vanillic acid (13,41). These cinnamic acids are also transformed by the microflora into 3-hydroxyphenylpropionic acid and 3-hydroxyhippuric acid, metabolites similar to those reported above for flavonoids (13). p-Coumaric acid liberated by the hydrolysis of coutaric acid is metabolized into 4-hydroxybenzoic, 4-hydroxyhippuric and hippuric acids, with the last-mentioned excreted in large amounts in the present experiment (42,43). Hippuric acid has often been described as a metabolite of polyphenols in rat and human studies using polyphenol-rich foods and beverages or crude polyphenol extracts (17,21,44 – 46), but the exact precursors were not determined. It cannot originate from catechin as previously suggested in a human study with black tea (44) because no increase in hippuric acid excretion was observed in the present work upon feeding pure catechin. The present work emphasizes the differences in bioavailability between a simple flavonoid (catechin) and a wine extract rich in high-molecular-weight polyphenols known to be poorly absorbed through the gut barrier (47– 49). The increase in urinary excretion of both aromatic acid metabolites and catechins (nonmethylated and methylated) after con-
sumption of the catechin or wine extract diets can be compared with the amount of polyphenol ingested. For the catechin diet, aromatic acids and intact catechins were 4.7 and 45.3 g/100 g, respectively, of the total catechin ingested. For the wine powder diet, they were 9.2 and 1.2 g/100 g, respectively, of the total wine polyphenols ingested. The major fraction of catechin, well absorbed through the small intestine, does not reach the distal part, and the yield of microbial metabolites is therefore limited. On the other hand, for the less easily absorbed wine polyphenols, a large fraction reaches the gut microflora and this explains the higher yields of microbial metabolites. The metabolism by the gut microflora may have a major effect on tissue exposure to polyphenols, particularly for higher-molecular-weight polyphenols such as proanthocyanidins or oxidized polymeric polyphenols abundant in many fruits, wine, tea or chocolate (50) and contribute to the antioxidant capacity of inner tissues. In rats fed the catechin diet, the increase in the reducing capacity of urine (⫹ 11.1 mg catechin equivalents per day) was essentially explained by the total excretion of catechin itself and its methylated derivative (10.8 mg/d). In rats fed the 0.50 g/100 g wine polyphenol diet, a smaller increase in the urine reducing capacity was observed (⫹ 4.5 mg catechin equivalents per day), explained not only by catechin excretion (0.33 mg/d) but also by the excretion of the aromatic acid metabolites bearing a phenolic group (total 0.68 mg/d), which also have reducing and antioxidant properties (51). These aromatic acids may thus make an important contribution to the protection against oxidative stress and explain some of the biological effects reported for proanthocyanidins and other high-molecular-weight polyphenols in animal and human studies (50). It has been shown that microbial metabolites such as 3,4-dihydroxyphenylacetic and 4-hydroxyphenylacetic acids were more effective than their rutin and quercetin precursors in inhibiting platelet aggregation in vitro (52). More attention should be given in the future to the microbial metabolites and their biological properties in particular to explain the health effects of polyphenols not easily absorbed through the gut barrier. ACKNOWLEDGMENTS We thank the Socie´ te´ Franc¸ aise de Distillerie and Michel Martin (Unite´ Expe´ rimentale de Pech Rouge, INRA, France) for providing us the polyphenol-rich wine extract and Jean-Marc Souquet (UMR Sciences pour l’Oenologie, INRA, France) for analyzing this sample. We gratefully acknowledge Catherine Besson, Marie-Anne Verny and Christine Cubizolles (Unite´ des Maladies Me´ taboliques et Micronutriments, INRA, France) for their contribution in animal handling. We thank Vanessa Crespy (Unite´ des Maladies Me´ taboliques et Micronutriments, INRA, France) for her careful reading of the manuscript.
LITERATURE CITED 1. Ku¨ hnau, J. (1976) The flavonoids: a class of semi-essential food components: their role in human nutrition. World Rev. Nutr. Diet. 24: 117–191. 2. Scalbert, A. & Williamson, G. (2000) Dietary intake and bioavailability of polyphenols. J. Nutr. 130: 2073S–2085S. 3. Maxwell, S., Cruickshank, A. & Thorpe, G. (1994) Red wine and antioxidant activity in serum. Lancet 344: 193–194. 4. Serafini, M., Ghiselli, A. & Ferro-Luzzi, A. (1996) In vivo antioxidant effect of green and black tea in man. Eur. J. Clin. Nutr. 50: 28 –32. 5. Serafini, M., Maiani, G. & Ferro-Luzzi, A. (1998) Alcohol-free red wine enhances plasma antioxidant capacity in humans. J. Nutr. 128: 1003–1007. 6. Young, J. F., Nielsen, S. E., Haraldsdottir, J., Daneshvar, B., Lauridsen, S. T., Knuthsen, P., Crozier, A., Sandstrom, B. & Dragsted, L. O. (1999) Effect of fruit juice intake on urinary quercetin excretion and biomarkers of antioxidative status. Am. J. Clin. Nutr. 69: 87–94. 7. Hertog, M. G. L., Kromhout, D., Aravanis, C., Blackburn, H., Buzina, R.,
BIOAVAILABILITY OF WINE POLYPHENOLS IN RATS Fidanza, F., Giampaoli, S., Jansen, A., Menotti, A., Nedeljkovic, S., Pekkarinen, M., Simic, B. S., Toshima, H., Feskens, E. J. M., Hollman, P. C. H. & Katan, M. B. (1995) Flavonoid intake and long-term risk of coronary heart disease and cancer in the Seven Countries Study. Arch. Intern. Med. 155: 381–386. 8. Knekt, P., Jarvinen, R., Reunanen, A. & Maatela, J. (1996) Flavonoid intake and coronary mortality in Finland: a cohort study. Br. Med. J. 312: 478 – 481. 9. Rimm, E. B., Katan, M. B., Ascherio, A., Stampfer, M. J. & Willett, W. C. (1996) Relation between intake of flavonoids and risk for coronary heart disease in male health professionals. Ann. Intern. Med. 125: 384 –389. 10. Steinmetz, K. A. & Potter, J. D. (1996) Vegetables, fruits, and cancer prevention: a review. J. Am. Diet. Assoc. 96: 1027–1039. 11. Hollman, P. C. H., Hertog, M. G. L. & Katan, M. B. (1996) Role of dietary flavonoids in protection against cancer and coronary heart disease. Biochem. Soc. Trans. 24: 785–789. 12. Adlercreutz, H. & Mazur, W. (1997) Phyto-oestrogens and Western diseases. Ann. Med. 29: 95–120. 13. Booth, A. N., Emerson, O. H., Jones, F. T. & DeEds, F. (1957) Urinary metabolites of caffeic and chlorogenic acids. J. Biol. Chem. 229: 51–59. 14. Das, N. P. (1969) Studies on flavonoid metabolism. Degradation of (⫹)-catechin by rat intestinal contents. Biochim. Biophys. Acta 177: 668 – 670. 15. Groenewoud, G. & Hundt, H.K.L. (1986) The microbial metabolism of condensed (⫹)-catechins by rat-caecal microflora. Xenobiotica 16: 99 –107. 16. Scheline, R. R. (1991) CRC Handbook of Mammalian Metabolism of Plant Compounds. CRC Press, Boca Raton, FL. 17. Harmand, M. F. & Blanquet, P. (1978) The fate of total flavanolic oligomers (OFT) extracted from Vitis vinifera L. in the rat. Eur. J. Drug Metab. Pharmacokinet. 3: 15–30. 18. De´ prez, S., Bre´ zillon, C., Rabot, S., Philippe, C., Mila, I., Lapierre, C. & Scalbert, A. (2000) Polymeric proanthocyanidins are catabolized by a human colonic microflora into low molecular weight phenolic acids. J. Nutr. 130: 2733– 2738. 19. Griffiths, L. A. (1964) Studies on flavonoid metabolism. Identification of the metabolites of (⫹)-catechin in rat urine. Biochem. J. 92: 173–179. 20. Goodwin, B. L., Ruthven, C. R. J. & Sandler, M. (1994) Gut flora and the origin of some urinary aromatic phenolic compounds. Biochem. Pharmacol. 47: 2294 –2297. 21. Quick, A. J. (1931) The conjugation of benzoic acid in man. J. Biol. Chem. 92: 65. 22. Rechner, A. R., Kuhnle, G., Bremner, P., Hubbard, G. P., Moore, K. P. & Rice-Evans, C. A. (2002) The metabolic fate of dietary polyphenols in humans. Free Radic. Biol. Med. 33: 220 –235. 23. Das, N. P. & Griffiths, L. A. (1969) Studies on flavonoid metabolism. Metabolism of (⫹)-[14C]catechin in the rat and guinea pig. Biochem. J. 115: 831– 836. 24. Sawaı¨, Y., Kohsaka, K., Nishiyama, Y. & Ando, K. (1987) Serum concentrations of rutinoside metabolites after oral administration of a rutinoside formulation to humans. Arzneimittelforschung 37: 729 –732. 25. Donovan, J. L., Bell, J. R., Kasim-Karakas, S., German, J. B., Walzem, R. L., Hansen, R. J. & Waterhouse, A. L. (1999) Catechin is present as metabolites in human plasma after consumption of red wine. J. Nutr. 129: 1662– 1668. 26. Fulcrand, H., Remy, S., Souquet, J. M., Cheynier, V. & Moutounet, M. (1999) Study of wine tannin oligomers by on-line liquid chromatography electrospray mass spectrometry. J. Agric. Food Chem. 47: 1023–1028. 27. Manach, C., Morand, C., Crespy, V., Demigne´ , C., Texier, O., Re´ ge´ rat, F. & Re´ me´ sy, C. (1998) Quercetin is recovered in human plasma as conjugated derivatives which retain antioxidant properties. FEBS Lett. 426: 331–336. 28. Donovan, J. L., Crespy, V., Manach, C., Morand, C., Besson, C., Scalbert, A. & Re´ me´ sy, C. (2000) Catechin is metabolized by both the small intestine and the liver in rats. J. Nutr. 131: 1753–1757. 29. Scalbert, A., Monties, B. & Janin, G. (1989) Tannins in wood: comparison of different estimation methods. J. Agric. Food Chem. 37: 1324 –1329. 30. Adamson, R. H., Bridges, J. W., Evans, M. E. & Williams, R. T. (1970) Species differences in the aromatization of quinic acid in vivo and the role of gut bacteria. Biochem. J. 116: 437– 443. 31. Das, N. P. (1971) Studies on flavonoid metabolism. Absorption and metabolism of (⫹)-catechin in man. Biochem. Pharmacol. 20: 3435–3445.
467
32. Groenewoud, G. & Hundt, H.K.L. (1984) The microbial metabolism of (⫹)-catechin to two novel diarylpropan-2-ol metabolites in vitro. Xenobiotica 14: 711–717. 33. Curtius, H. C., Mettler, M. & Ettlinger, L. (1976) Study of the intestinal tyrosine metabolism using stable isotopes and gas chromatography-mass spectrometry. J. Chromatogr. 126: 569 –580. 34. Li, C., Lee, M. J., Sheng, S., Meng, X., Prabhu, S., Winnik, B., Huang, B., Chung, J. Y., Yan, S., Ho, C. T. & Yang, C. S. (2000) Structural identification of two metabolites of catechins and their kinetics in human urine and blood after tea ingestion. Chem. Res. Toxicol. 13: 177–184. 35. Scheline, R. R. (1970) The metabolism of (⫹)-catechin to hydroxyphenylvaleric acids by the intestinal microflora. Biochim. Biophys. Acta 222: 228 – 230. 36. Aura, M. A., O’Leary, K. A., Williamson, G., Ojala, M., Bailey, M., Puupponen-Pimia¨ , R., Nuutila, A. M., Aksman-Caldentey, K. M. & Poutanen, K. (2002) Quercetin derivatives are deconjugated and converted to hydroxyphenylacetic acids but not methylated by human fecal microflora in vitro. J. Agric. Food Chem. 50: 1725–1730. 37. Baderschneider, B. & Winterhalter, P. (2001) Isolation and characterization of novel benzoates, cinnamates, flavonoids, and lignans from Riesling wine and screening for antioxidant activity. J. Agric. Food Chem. 49: 2788 –2798. 38. Plumb, G. W., Garcia-Conesa, M. T., Kroon, P. A., Rhodes, M., Ridley, S. & Williamson, G. (1999) Metabolism of chlorogenic acid by human plasma, liver, intestine and gut microflora. J. Sci. Food Agric. 79: 390 –392. 39. Masri, M. S., Robbins, D. J., Emerson, O. H. & DeEds, F. (1964) Selective para- or meta-O-methylation with catechol O-methyl transferase from rat liver. Nature (Lond.) 202: 878 – 879. 40. Gumbinger, H. G., Vahlensieck, U. & Winterhoff, H. (1993) Metabolism of caffeic acid in the isolated perfused rat liver. Planta Med. 59: 491– 493. 41. Teuchy, H. & Van Sumere, C. F. (1971) The metabolism of (1-14C) phenylalanine, (3-14C)cinnamic acid and (2-14C) ferulic acid in the rat. Arch. Int. Physiol. Biochem. 79: 589 – 618. 42. Booth, A. N., Masri, M. S., Robbins, D. J., Emerson, O. H., Jones, F. T. & DeEds, F. (1960) Urinary phenolic acid metabolites of tyrosine. J. Biol. Chem. 235: 2649. 43. Teuchy, H., Quatacker, J., Wolf, G. & Van Sumere, C. F. (1971) Quantitative investigation of the hippuric acid formation in the rat after administration of some possible aromatic and hydroaromatic precursors. Arch. Int. Physiol. Biochem. 79: 573–587. 44. Clifford, M. N., Copeland, E. L., Bloxsidge, J. P. & Mitchell, L. A. (2000) Hippuric acid as a major excretion product associated with black tea consumption. Xenobiotica 30: 317–326. 45. Graefe, E. U. & Veit, M. (1999) Urinary metabolites of flavonoids and hydroxycinnamic acids in humans after application of a crude extract from Equisetum arvense. Phytomedicine 6: 239 –246. 46. Pietta, P. G., Gardana, C. & Mauri, P. L. (1997) Identification of Gingko biloba flavonol metabolites after oral administration to humans. J. Chromatogr. B Biomed. Sci. Appl. 693: 249 –255. 47. Donovan, J. L., Manach, C., Rios, L., Morand, C., Scalbert, A. & Re´ me´ sy, C. (2002) Procyanidins are not bioavailable in rats fed a single meal containing a grape seed extract or the procyanidin dimer B3. Br. J. Nutr. 87: 299 –306. 48. De´ prez, S., Mila, I., Huneau, J.-F., Tome´ , D. & Scalbert, A. (2001) Transport of proanthocyanidin dimer, trimer and polymer across monolayers of human intestinal epithelial Caco-2 cells. Antioxid. Redox Signal. 3: 957–967. 49. Mulder, T. P., van Platerink, C. J., Wijnand Schuyl, P. J. & van Amelsvoort, J. M. (2001) Analysis of theaflavins in biological fluids using liquid chromatography-electrospray mass spectrometry. J. Chromatogr. B Biomed. Sci. Appl. 760: 271–279. 50. Santos-Buelga, C. & Scalbert, A. (2000) Proanthocyanidins and tannin-like compounds: nature, occurrence, dietary intake and effects on nutrition and health. J. Sci. Food Agric. 80: 1094 –1117. 51. Singleton, V. L. & Rossi, J. A. J. (1965) Colorimetry of total phenolics with phosphomolybdic-phosphotungstic acid reagents. Am. J. Enol. Viticult. 16: 144 –158. 52. Kim, D.-H., Jung, E.-A., Sohng, I.-S., Han, J.-A., Kim, T.-H. & Han, M. J. (1998) Intestinal bacterial metabolism of flavonoids and its relation to some biological activities. Arch. Pharm. Res. 21: 17–23.
