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The role of phenolic compounds during formation of turbidity in an aromatic bitter
Food Chemistry 123 (2010) 1035–1039
Contents lists available at ScienceDirect
Food Chemistry journal homepage: www.elsevier.com/locate/foodchem
The role of phenolic compounds during formation of turbidity in an aromatic bitter Anja Rødtjer, Leif H. Skibsted, Mogens L. Andersen * Food Chemistry, Department of Food Science, Faculty of Life Sciences, University of Copenhagen, Rolighedsvej 30, DK-1958 Frederiksberg C, Denmark
a r t i c l e
i n f o
Article history: Received 26 January 2010 Received in revised form 15 March 2010 Accepted 11 May 2010
Keywords: Alcoholic beverage Polyphenols Oxidation Haze
a b s t r a c t An aromatic bitter produced from alcoholic extracts of botanicals were studied over 2½ years. The concentrations of gallic acid and the flavonoids catechin, epicatechin, eriodictyol, and taxifolin decreased during the first year of storage in the presence of oxygen, whereas the turbidity stayed below the limit for visible detection. After one year the turbidity increased linearly with time and the total concentration of phenolic compounds decreased. In contrast, the concentrations of eugenol and phenolic acids with two or one hydroxyl groups remained constant or increased, and these compounds are probably not involved in formation of turbidity. The oxidative properties of the phenolic compounds were evaluated in an assay based on iron-catalysed Fenton reactions and spin trapping of radicals. The fresh bitter and bitter stored for one year increased the amount of radicals detected in the assay, demonstrating an overall prooxidative effect of the phenolic compounds. It is concluded that easily oxidised phenolics in the bitter are converted into soluble intermediates that slowly react with proteins and carbohydrates forming haze and deposits. Ó 2010 Elsevier Ltd. All rights reserved.
1. Introduction Formation of turbidity and deposits during storage is often observed for a number of beverages that contain high levels of plant derived phenolic compounds. The formation of haze in beer, apple juice, grape juice and wine are well described and are mainly caused by interactions between oxidised polyphenols and proteins (Siebert, 2006). Haze formed in beer is composed of phenolic compounds, protein and carbohydrate but the reactions between phenolics and proteins control the development of haze while carbohydrate appears to be passively complexed with the other materials during the haze formation and precipitation (Siebert, 1999). The precipitate formed during storage of an aromatic bitter consisted of mainly plant polyphenols and smaller amounts of proteins (16.5 wt.%) and carbohydrates (3.2 wt.%) (Refsgaard, Brockhoff, Meilgaard, Laursen, & Skibsted, 1996; Refsgaard, Schaumburg, & Skibsted, 1996). The formation of deposits was accelerated by addition of an azo compound that acted as a radical initiator or by a Fenton reagent (iron(III) salt and ascorbic acid), which suggested that the deposits were formed by oxidative reactions. It was proposed that formation of precipitates in alcoholic beverages of the bitter type involves an initial oxidative polymerisation of polyphenols followed by the coprecipitation of polymerised polyphenols, proteins and carbohydrates.
* Corresponding author. Tel.: +45 35333262; fax: +45 35333344. E-mail address: [email protected] (M.L. Andersen). 0308-8146/$ - see front matter Ó 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.foodchem.2010.05.056
To provide further basis for the discussion of the mechanism for the formation of deposit in a polyphenol-rich bitter and other alcoholic beverages containing plant extracts, this study examined the role of phenolics during the development of turbidity under conditions, which would be similar to storage at the consumer. The concentration of phenolics in the bitter was followed during storage of the bitter with or without oxygen in a 2½ years storage period and correlated with the development of turbidity in the bitter. 2. Materials and methods 2.1. Chemicals A commercially available bitter with 38 vol.% molasses spirit was obtained directly from the Danish spirit producer. The bitter was produced and bottled immediately before the start of the storage experiment. The bitter contained distillates and extracts of about thirty different seeds, drugs and herbs including rowanberry (Refsgaard, Brockhoff, et al., 1996). Folin–Ciocalteu phenol reagent, and analytical grade iron(II) sulphate heptahydrate were obtained from Merck (Darmstadt, Germany). Sodium carbonate heptahydrate was from Riedel-de Haën (Seelze, Germany). Gallic acid monohydrate and ()-epicatechin were from Aldrich Chemical Co. (Milwaukee, WI). (+)-Catechin, p-coumaric acid, eugenol, protocatechuic acid, sinapic acid and syringic acid were obtained from Sigma–Aldrich Chemie (Steinheim, Germany). Eriodictyol was from Roth (Karlsruhe, Germany). Taxifolin and vanillic acid was from Fluka Chemie AG (Buchs, Switzerland). Hydrogen peroxide (35%) was from Bie & Berntsen A/S (Rødovre, Denmark).
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a-(4-Pyridyl-1-oxide)-N-tert-butylnitrone (POBN) was from Sigma Chemical Co. (St. Louis, MO). All solvents were of HPLC grade. Water was purified through a Millipore Q-plus purification train (Millipore, Bedford, MA, USA). 2.2. Sample preparation and treatment Freshly produced bitter was poured into bottles and stored at room temperature in the dark with or without oxygen for 910 days. The presence of oxygen throughout the storage experiment was ensured by opening the bottles for 30 s once each week and thereafter turning the bottles upside down three times. Storage of the bitter without oxygen was carried out by storing the bitter in several bottles without air in the headspace. These bottles were all kept closed from the start of the experiment until they were taken out for analysis. 2.3. Measurement of turbidity The turbidity was measured using a Nephla Turbiditimeter (Hach Lange Ltd., Manchester, UK) and the turbidity was expressed as Formazin Nephelometric Units (FNU). 2.4. Quantification of the total amount of phenolic compounds The amount of total phenolics in the bitter was determined according to the Folin–Ciocalteu procedure (Slinkard & Singleton, 1977). Samples (200 ll, two replicates) were mixed with 1.0 ml of Folin–Ciocalteu’s reagent (diluted 1:10 with water) and 0.8 ml of a 7.5% solution of sodium carbonate were added. The absorption at 765 nm was measured after 30 min with a Cary 3 UV–vis spectrophotometer (Varian Techtron Pty. Ltd., Mulgrave, Victoria, Australia). The total phenolic content was expressed as gallic acid equivalents (GAE) in mg per litre. 2.5. Quantification of phenolics by HPLC A method previously developed for analysis of phenolic compounds in aromatic bitter products was used with minor modifications (Rødtjer, Skibsted, & Andersen, 2006a). Bitter samples were diluted in 38% ethanol, filtered, and 10 ll aliquots of the solutions were injected into the HPLC. Analysis was carried out with a gradient forming Hewlett Packard series 1050 HPLC system (Palo Alto, CA) that was coupled to an Agilent Technologies series 1100 autosampler (Palo Alto, CA). The HPLC system was equipped with a Hichrom Ltd. H1000DS-10C Guard column (Reading, UK) and the separation was performed on a 25 cm 4.6 mm (i.d.) MachereyNagel ET NucleosilÒ reversed phase 10 lm C18 stainless-steel column (Düren, Germany). The mobile phase consisted of 2.5% acetic acid (solvent A), and solvent B was a mixture of water, methanol and acetic acid (2.5:95:2.5). The gradient applied at a flow rate of 1.0 ml min1 was; 0–10 min, 100% A; and 10–110 min, 100% A to 50% A. An ESA Analytical Coulochem II dual-channel electrochemical detector equipped with an ESA Model 5010 analytical cell (Chelmsford, MA, USA) was used for detection. The settings for the electrochemical detector were as follows: Channel 1: Potential 0 mV; output range 20 lA, channel 2: Potential 600 mV; output range 2 lA, and the filter time was 2 s for both channels. Electronic data acquisition and peak integration were performed using a HP ChemStation for LC. Peak identification was confirmed by comparison with retention times of pure phenolic compounds. Standard curves based on pure phenolics were used for quantification. All samples and standard solutions contained sinapic acid as internal standard and HPLC analyses were performed in triplicate.
2.6. Fenton reaction model system with ESR detection of POBN spin adducts The assay was with minor modifications as described by Rødtjer, Skibsted, and Andersen (2006b). Four millilitre of 3.2 mM POBN dissolved in 1.0 M aqueous ethanol were mixed with 20 ll of 1.1–44 mM FeSO4 and 50 ll of either a bitter sample or 38% aqueous ethanol as a reference. Finally, 80 ll of 24 mM H2O2 were added. An aliquot of the reaction mixture was withdrawn directly into an ESR quartz capillary tube with an interior diameter of 0.75 mm (Wilmad, Buena, NJ, USA), and the ESR spectrum was recorded 2 min after the addition of the H2O2 solution on a Jeol JES-FR30 ESR spectrometer (JEOL Ltd., Tokyo, Japan). The measurements were carried out at room temperature with a microwave power of 4 mW, and a modulation width of 0.1 mT. The intensity of all signals were recorded relative to the intensity of a Mn(II)-marker (JEOL Ltd., Tokyo, Japan) attached to the cavity of the spectrometer. Measurements were carried out with varying concentrations of bitter and the degree of inhibition (IESR) was calculated from the height of the central peak of the ESR signal of the spin adduct by the following formula:
Peak heightsample 100% IESR ¼ 1 Peak heightreference
2.7. Statistical analysis Evaluation of statistical significance of differences was performed by using the Student’s t-test.
3. Results and discussion The development of turbidity and change in the total concentration of phenolic compounds in an aromatic bitter were studied over a period of 910 days (2½ years) (Fig. 1). One set of samples of the bitter were kept in bottles that were opened regularly, for 30 s every week and were thus repeatedly exposed to oxygen. The bottles were rotated three times every time they had been opened to ensure a uniform concentration of oxygen. This treatment intended to simulate the handling of the aromatic bitter by a typical consumer, where several small servings may be taken from the same bottle at intervals over a period for more than one
Fig. 1. Turbidity (circles) and concentration of total soluble phenolic compounds (squares) in an aromatic bitter product during storage. The bitter was stored in the dark with either periodical access to atmospheric oxygen (filled symbols) or without access to oxygen (open symbols). Standard deviation is indicated by error bars.
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year. Reference samples were stored under identical conditions but without access to oxygen. The levels of turbidity were low in the aromatic bitter during the first 200 days of storage independently of the presence of oxygen. After this period the turbidity in the bottles that were regularly exposed to oxygen began to increase linearly with time during the rest of the storage, while the turbidity in the bottles without access to oxygen remained low and below the limit for visible detection (50 FNU) (Fig. 1). The low and practically constant level of turbidity during an initial phase (lag phase) followed by a phase where the turbidity increases linearly with time have been observed during storage of a number of polyphenol containing beverages (Fang, Zhang, Du, & Sun, 2007; Lee, Yusof, Hamid, & Baharin, 2007; Siebert, 2006; Tajchakavit, Boye, Bélanger, & Couture, 2001). The mechanism of haze formation is believed to involve an initial oxidation of polyphenolic compounds that generates oxidised polyphenols that can form haze-forming macromolecular complexes with other polyphenols or proteins. The insoluble deposits in the type of aromatic bitter used in this study have previously been shown to be a coprecipitate of polyphenols, proteins and carbohydrates (Refsgaard, Schaumburg, et al., 1996). Proteins accounted for 16.5% of the weight of the solid, and glucose, most likely bound as anthocyanins, was the dominating sugar component (3.2% of the weight of the solid). The involvement of polyphenols in the formation of haze was studied by following the total concentrations of phenols during the storage by the Folin–Ciocalteu method. The concentration of phenolic compounds was constant during the whole storage period in the bottles without access to oxygen (Fig. 1). The changes in the total concentration of soluble phenolic compounds in the presence of oxygen were observed to go through three distinct phases during the storage. The concentration of phenolic compounds was also constant during the first 250 days in the bottles exposed to oxygen, and it was not statistical significantly different from the concentration in the bottles without access to oxygen. A drop in the concentration of phenolic compounds was observed between days 250 and 400, which coincided with the period of time where the formation of turbidity accelerated. This shows that polyphenols are removed from the solution by the formation of the haze and
Fig. 2. HPLC chromatograms of phenolics compounds in bitter after storage. (a) 0 days, and (b) after 910 days with access to oxygen. The identified compounds are: (1) gallic acid, (2) protocatechuic acid, (3) (+)-catechin, (4) vanillic acid, (5) caffeic acid, (6) syringic acid, (7) ()-epicatechin, (8) p-coumaric acid, (9) taxifolin, (IS) sinapic acid as internal standard, (10) eriodictyol, and (11) eugenol. A–D, nonphenolic compounds.
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insoluble precipitates. However, the level of phenolic compounds only decreased slightly during the last 400 days of storage with access to oxygen, suggesting that all the polyphenols that are precursors for haze formation were depleted, and that the continued formation of haze must have been driven by formation of larger and larger deposit particles by complexation of non-phenolic compounds. The concentrations of specific phenolic compounds in the aromatic bitter were identified and quantified by HPLC analysis in order to examine the role of specific polyphenols as precursors for the formation of the turbidity. Eleven phenolic compounds were identified in the bitter including flavonoids, phenolic acids and eugenol (Rødtjer et al., 2006a) (Fig. 2). The concentration of five of these components, gallic acid, catechin, epicatechin, taxifolin and eriodictyol, decreased during storage of the bitter. Gallic acid was the only phenolic acid where the concentration decreased during storage (Fig. 3). All the gallic acid disappeared gradually during the first 500 days in bitter that was exposed to oxygen, whereas the level of gallic acid was almost constant during the first 400 days in bitter in the absence of oxygen, and a decay was only observed during the last period of the storage. The concentrations of all the identified flavonoids decreased during storage, and the decreases were accelerated by the presence of oxygen (Fig. 4). The major changes in concentrations took place during the first 300 days of storage, and the rates of disappearance in the presence of oxygen was: ()-epicatechin > (+)-catechin taxifolin eriodictyol.
Fig. 3. Change in concentration of phenolic benzoic acid derivates in bitter during storage. The bitter was stored in the dark either at the presence of oxygen (filled symbols) or without oxygen (un-filled symbols). Standard deviation is indicated by error bars.
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Fig. 4. Change in concentration of flavonoids in bitter during storage. The bitter was stored in the dark either at the presence of oxygen (filled symbols) or without oxygen (un-filled symbols). Standard deviation is indicated by error bars.
The disappearance of gallic acid, catechin, epicatechin, and taxifolin during the first 300 days of storage of the bitter in the presence of oxygen took place before the development of the turbidity. Furthermore, these four compounds also disappeared, albeit at a slower rate, in the absence of oxygen where turbidity did not develop in the bitter. This suggests that the initial soluble oxidation products presumably with a quinoidal structure must undergo further chemical and physical transformations before the formation of haze particles takes place. These subsequent transformations must be relatively slow, and accordingly rate determining for haze formation, since the oxidations of these four phenolic compounds were almost completed before the development of haze. It is noteworthy that nearly all gallic acid, catechin, epicatechin had disappeared before the formation of turbidity and the major decrease in the concentrations of the phenolic compounds. An alternative explanation could therefore be that the phenols, gallic acid, catechin, epicatechin, and taxifolin, act as antioxidants in the bitter and hinder oxidation during the initial storage phase. The initial oxidation of these compounds could protect other phenols, which upon oxidation form precipitates, from being oxidised. The antioxidative capacity of the bitter was therefore tested in an assay based on metal catalysed oxidation of ethanol (Graversen, Becker, Skibsted, & Andersen, 2008; Rødtjer et al., 2006b). The fresh bitter as well as the samples, which had been stored for 371 days, all accelerated the formation of radicals in the assay (Fig. 5). This suggests that the phenols in the bitter act as prooxidants and not
as antioxidants. A similar effect of phenols has been observed in wine, which has been explained by their ability to reduce iron and copper ions into the lower oxidation states that are needed for radical formation by the Fenton reaction (Danilewicz, 2003; Elias, Andersen, Skibsted, & Waterhouse, 2009a, 2009b; Li, Guo, & Wang, 2008). The concentration dependences of the three bitter samples tested in the assay were slightly different, which reflect their different profiles of phenolic compounds. The phenols in the sample stored for 371 days with access to oxygen gave slightly less prooxidative effects than the corresponding sample stored in a closed bottle (Fig. 5). This can be explained by the loss of the most easily oxidised phenolics in the former sample. Easily oxidised phenols are also better reductants, which are more likely to reduce Fe(III) and Cu(II) ions, and thereby exert a prooxidant effect. However, the assay also showed that the remaining phenols were also good prooxidants. The concentrations of the benzoic acids, protocatechuic acid, syringic acid and vanillic acid, increased during the storage (Fig. 3). It is possible that the increase in the concentrations of these compounds is caused by degradation of anthocyanidins. In neutral media protocatechuic acid has been shown to be formed by degradation of cyanidin, syringic acid by degradation of malvidin, and vanillic acid by degradation of peonidin (Fleschhut, Kratzer, Rechkemmer, & Kulling, 2006; Woodward, Kroon, Cassidy, & Kay, 2009). The corresponding anthocyanins are also degraded to the same hydroxybenzoic acids in neutral media, but the rates are much slower than the anthocyanidins (Woodward et al., 2009). Rowanberries are a likely source of cyanidin glycosides in the bitter (Koponen, Happonen, Mattila, & Törrönen, 2007). The presence of anthocyanins has also been demonstrated in deposits formed in the bitter (Refsgaard, Schaumburg, et al., 1996). The levels of the three hydroxybenzoic acids increased mostly in the presence of oxygen, which suggest that the release of these compounds is coupled to oxidation of their precursors. Eugenol, which gives an important clove flavour to the product, was the phenolic compound with the highest concentration among the identified phenolics in the bitter. However, the concentration of eugenol did not change during the storage, even in the presence of oxygen (Fig. 6). Similarly, the concentrations of the two cin-
Fig. 5. The effect of addition of increasing volume of bitter on the relative amount of radicals (spin adducts) produced in a Fenton/ethanol model system. The bitter samples were stored in 0 days (j), 371 days without oxygen (d), or 371 days with oxygen (N). The spin trap POBN (3 mM) and FeSO4 (106 lM) dissolved in 1 M aqueous ethanol was mixed with the bitter and H2O2 (463 lM).
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References
Fig. 6. Concentration of eugenol in bitter during storage. The bitter was stored in the dark either in the presence of oxygen (filled symbols) or without oxygen (unfilled symbols). Standard deviation is indicated by error bars.
namic acids, caffeic acid (3.5 ppm) and p-coumaric acid (0.9 ppm), did not change during the storage (results not shown). In conclusion, it was shown that before visible turbidity develops in an aromatic bitter some phenolic compounds have already been consumed due to oxidation. The easily oxidised phenolics in the bitter are converted into soluble intermediates most likely with a quinoidal structure, which slowly undergo polymerisation reactions with proteins and carbohydrates forming haze and deposits. Turbidity did not appear in bottles where oxygen was excluded during the storage. Other phenolic compounds proved to be stable against oxidation during the prolonged storage, and the concentration of some even increased, presumably due to break down of anthocyanidins and anthocyanins. Acknowledgements This research was sponsored by the Danish Ministry of Food, Agriculture, and Fisheries through LMC – Centre for Advanced Food Studies as a part of the collaboration project ‘‘Plant phenols in cherry liqueur and bitter. Product stability and utilization of pomace in food production” with Danish Distillers.
Danilewicz, J. C. (2003). Review of reaction mechanisms of oxygen and proposed intermediate reduction products in wine: Central role of iron and copper. American Journal of Enology and Viticulture, 54, 73–85. Elias, R. J., Andersen, M. L., Skibsted, L. H., & Waterhouse, A. L. (2009a). Identification of free radical intermediates in oxidized wine using electron paramagnetic resonance spin trapping. Journal of Agricultural and Food Chemistry, 57, 4359–4365. doi:10.1021/jf8035484. Elias, R. J., Andersen, M. L., Skibsted, L. H., & Waterhouse, A. L. (2009b). Key factors affecting radical formation in wine studied by spin trapping and ESR spectroscopy. American Journal of Enology and Viticulture, 60, 471–476. Fang, Z., Zhang, M., Du, W., & Sun, J. (2007). Effect of fining and filtration on the haze formation in bayberry (Myrica rubra Sieb. et Zucc.) juice. Journal of Agricultural and Food Chemistry, 55, 113–119. doi:10.1021/jf062699j. Fleschhut, J., Kratzer, F., Rechkemmer, G., & Kulling, S. E. (2006). Stability and biotransformation of various dietary anthocyanins in vitro. European Journal of Nutrition, 45, 7–18. doi:10.1007/s00394-005-0557-8. Graversen, H. B., Becker, E. M., Skibsted, L. H., & Andersen, M. L. (2008). Antioxidant synergism between fruit juice and a-tocopherol. A comparison between high phenolic black chokeberry (Aronia melanocarpa) and high ascorbic blaccurrant (Ribes nigrum). European Food Research and Technology, 226, 737–743. doi:10.1017/s00217-007-0585-0. Koponen, J. M., Happonen, A. M., Mattila, P. H., & Törrönen, A. R. (2007). Contents of anthocyanins and ellagitannins in selected foods consumed in Finland. Journal of Agricultural and Food Chemistry, 55, 1612–1619. doi:10.1021/jf062897a. Lee, W. C., Yusof, S., Hamid, N. S. A., & Baharin, B. S. (2007). Effects of fining treatment and storage temperature on the quality of clarified banana juice. LWT – Food Science and Technology, 40, 1755–1764. doi:10.1016/j.lwt.2006.12.008. Li, H., Guo, A., & Wang, H. (2008). Mechanisms of oxidative browning in wine. Food Chemistry, 108, 1–13. doi:10.1016/j.foodchem.2007.10.065. Refsgaard, H. H. F., Brockhoff, P. M., Meilgaard, M. C., Laursen, P. F., & Skibsted, L. H. (1996). Light-induced sensory and chemical changes in aromatic bitters. Zeitschrift für Lebensmittel-Untersuchung und -Forschung, 203, 47–55. Refsgaard, H. H. F., Schaumburg, K., & Skibsted, L. H. (1996). Solid state C-13 NMR investigations of insoluble deposits in aromatic bitters. Zeitschrift für Lebensmittel-Untersuchung und -Forschung, 203, 287–292. Rødtjer, A., Skibsted, L. H., & Andersen, M. L. (2006a). Identification and quantification of phenolics in aromatic bitter and cherry liqueur by HPLC with electrochemical detection. European Food Research and Technology, 223, 663–668. doi:10.1007/s00217-005-0250-4. Rødtjer, A., Skibsted, L. H., & Andersen, M. L. (2006b). Antioxidative and prooxidative effects of extracts made from cherry liqueur pomace. Food Chemistry, 99, 6–14. doi:10.1016/j.foodchem.2005.07.011. Siebert, K. J. (1999). Effects of protein–polyphenol interactions on beverage haze, stabilization, and analysis. Journal of Agricultural and Food Chemistry, 47(2), 353–362. doi:10.1021/jf980703o. Siebert, K. J. (2006). Haze formation in beverages. Food Science and Technology, 39, 987–994. doi:10.1016/j.lwt.2006.02.012. Slinkard, K., & Singleton, V. L. (1977). Total phenol analysis: Automation and comparison with manual methods. American Journal of Enology and Viticulture, 28, 49–55. Tajchakavit, S., Boye, J. I., Bélanger, D., & Couture, R. (2001). Kinetics of haze formation and factors influencing the development of haze in clarified apple juices. Food Research International, 34, 431–440. Woodward, G., Kroon, P., Cassidy, A., & Kay, C. (2009). Anthocyanin stability and recovery: Implications for the analysis of clinical and experimental samples. Journal of Agricultural and Food Chemistry, 57, 5271–5278. doi:10.1021/ jf900602b.
Contents lists available at ScienceDirect
Food Chemistry journal homepage: www.elsevier.com/locate/foodchem
The role of phenolic compounds during formation of turbidity in an aromatic bitter Anja Rødtjer, Leif H. Skibsted, Mogens L. Andersen * Food Chemistry, Department of Food Science, Faculty of Life Sciences, University of Copenhagen, Rolighedsvej 30, DK-1958 Frederiksberg C, Denmark
a r t i c l e
i n f o
Article history: Received 26 January 2010 Received in revised form 15 March 2010 Accepted 11 May 2010
Keywords: Alcoholic beverage Polyphenols Oxidation Haze
a b s t r a c t An aromatic bitter produced from alcoholic extracts of botanicals were studied over 2½ years. The concentrations of gallic acid and the flavonoids catechin, epicatechin, eriodictyol, and taxifolin decreased during the first year of storage in the presence of oxygen, whereas the turbidity stayed below the limit for visible detection. After one year the turbidity increased linearly with time and the total concentration of phenolic compounds decreased. In contrast, the concentrations of eugenol and phenolic acids with two or one hydroxyl groups remained constant or increased, and these compounds are probably not involved in formation of turbidity. The oxidative properties of the phenolic compounds were evaluated in an assay based on iron-catalysed Fenton reactions and spin trapping of radicals. The fresh bitter and bitter stored for one year increased the amount of radicals detected in the assay, demonstrating an overall prooxidative effect of the phenolic compounds. It is concluded that easily oxidised phenolics in the bitter are converted into soluble intermediates that slowly react with proteins and carbohydrates forming haze and deposits. Ó 2010 Elsevier Ltd. All rights reserved.
1. Introduction Formation of turbidity and deposits during storage is often observed for a number of beverages that contain high levels of plant derived phenolic compounds. The formation of haze in beer, apple juice, grape juice and wine are well described and are mainly caused by interactions between oxidised polyphenols and proteins (Siebert, 2006). Haze formed in beer is composed of phenolic compounds, protein and carbohydrate but the reactions between phenolics and proteins control the development of haze while carbohydrate appears to be passively complexed with the other materials during the haze formation and precipitation (Siebert, 1999). The precipitate formed during storage of an aromatic bitter consisted of mainly plant polyphenols and smaller amounts of proteins (16.5 wt.%) and carbohydrates (3.2 wt.%) (Refsgaard, Brockhoff, Meilgaard, Laursen, & Skibsted, 1996; Refsgaard, Schaumburg, & Skibsted, 1996). The formation of deposits was accelerated by addition of an azo compound that acted as a radical initiator or by a Fenton reagent (iron(III) salt and ascorbic acid), which suggested that the deposits were formed by oxidative reactions. It was proposed that formation of precipitates in alcoholic beverages of the bitter type involves an initial oxidative polymerisation of polyphenols followed by the coprecipitation of polymerised polyphenols, proteins and carbohydrates.
* Corresponding author. Tel.: +45 35333262; fax: +45 35333344. E-mail address: [email protected] (M.L. Andersen). 0308-8146/$ - see front matter Ó 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.foodchem.2010.05.056
To provide further basis for the discussion of the mechanism for the formation of deposit in a polyphenol-rich bitter and other alcoholic beverages containing plant extracts, this study examined the role of phenolics during the development of turbidity under conditions, which would be similar to storage at the consumer. The concentration of phenolics in the bitter was followed during storage of the bitter with or without oxygen in a 2½ years storage period and correlated with the development of turbidity in the bitter. 2. Materials and methods 2.1. Chemicals A commercially available bitter with 38 vol.% molasses spirit was obtained directly from the Danish spirit producer. The bitter was produced and bottled immediately before the start of the storage experiment. The bitter contained distillates and extracts of about thirty different seeds, drugs and herbs including rowanberry (Refsgaard, Brockhoff, et al., 1996). Folin–Ciocalteu phenol reagent, and analytical grade iron(II) sulphate heptahydrate were obtained from Merck (Darmstadt, Germany). Sodium carbonate heptahydrate was from Riedel-de Haën (Seelze, Germany). Gallic acid monohydrate and ()-epicatechin were from Aldrich Chemical Co. (Milwaukee, WI). (+)-Catechin, p-coumaric acid, eugenol, protocatechuic acid, sinapic acid and syringic acid were obtained from Sigma–Aldrich Chemie (Steinheim, Germany). Eriodictyol was from Roth (Karlsruhe, Germany). Taxifolin and vanillic acid was from Fluka Chemie AG (Buchs, Switzerland). Hydrogen peroxide (35%) was from Bie & Berntsen A/S (Rødovre, Denmark).
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a-(4-Pyridyl-1-oxide)-N-tert-butylnitrone (POBN) was from Sigma Chemical Co. (St. Louis, MO). All solvents were of HPLC grade. Water was purified through a Millipore Q-plus purification train (Millipore, Bedford, MA, USA). 2.2. Sample preparation and treatment Freshly produced bitter was poured into bottles and stored at room temperature in the dark with or without oxygen for 910 days. The presence of oxygen throughout the storage experiment was ensured by opening the bottles for 30 s once each week and thereafter turning the bottles upside down three times. Storage of the bitter without oxygen was carried out by storing the bitter in several bottles without air in the headspace. These bottles were all kept closed from the start of the experiment until they were taken out for analysis. 2.3. Measurement of turbidity The turbidity was measured using a Nephla Turbiditimeter (Hach Lange Ltd., Manchester, UK) and the turbidity was expressed as Formazin Nephelometric Units (FNU). 2.4. Quantification of the total amount of phenolic compounds The amount of total phenolics in the bitter was determined according to the Folin–Ciocalteu procedure (Slinkard & Singleton, 1977). Samples (200 ll, two replicates) were mixed with 1.0 ml of Folin–Ciocalteu’s reagent (diluted 1:10 with water) and 0.8 ml of a 7.5% solution of sodium carbonate were added. The absorption at 765 nm was measured after 30 min with a Cary 3 UV–vis spectrophotometer (Varian Techtron Pty. Ltd., Mulgrave, Victoria, Australia). The total phenolic content was expressed as gallic acid equivalents (GAE) in mg per litre. 2.5. Quantification of phenolics by HPLC A method previously developed for analysis of phenolic compounds in aromatic bitter products was used with minor modifications (Rødtjer, Skibsted, & Andersen, 2006a). Bitter samples were diluted in 38% ethanol, filtered, and 10 ll aliquots of the solutions were injected into the HPLC. Analysis was carried out with a gradient forming Hewlett Packard series 1050 HPLC system (Palo Alto, CA) that was coupled to an Agilent Technologies series 1100 autosampler (Palo Alto, CA). The HPLC system was equipped with a Hichrom Ltd. H1000DS-10C Guard column (Reading, UK) and the separation was performed on a 25 cm 4.6 mm (i.d.) MachereyNagel ET NucleosilÒ reversed phase 10 lm C18 stainless-steel column (Düren, Germany). The mobile phase consisted of 2.5% acetic acid (solvent A), and solvent B was a mixture of water, methanol and acetic acid (2.5:95:2.5). The gradient applied at a flow rate of 1.0 ml min1 was; 0–10 min, 100% A; and 10–110 min, 100% A to 50% A. An ESA Analytical Coulochem II dual-channel electrochemical detector equipped with an ESA Model 5010 analytical cell (Chelmsford, MA, USA) was used for detection. The settings for the electrochemical detector were as follows: Channel 1: Potential 0 mV; output range 20 lA, channel 2: Potential 600 mV; output range 2 lA, and the filter time was 2 s for both channels. Electronic data acquisition and peak integration were performed using a HP ChemStation for LC. Peak identification was confirmed by comparison with retention times of pure phenolic compounds. Standard curves based on pure phenolics were used for quantification. All samples and standard solutions contained sinapic acid as internal standard and HPLC analyses were performed in triplicate.
2.6. Fenton reaction model system with ESR detection of POBN spin adducts The assay was with minor modifications as described by Rødtjer, Skibsted, and Andersen (2006b). Four millilitre of 3.2 mM POBN dissolved in 1.0 M aqueous ethanol were mixed with 20 ll of 1.1–44 mM FeSO4 and 50 ll of either a bitter sample or 38% aqueous ethanol as a reference. Finally, 80 ll of 24 mM H2O2 were added. An aliquot of the reaction mixture was withdrawn directly into an ESR quartz capillary tube with an interior diameter of 0.75 mm (Wilmad, Buena, NJ, USA), and the ESR spectrum was recorded 2 min after the addition of the H2O2 solution on a Jeol JES-FR30 ESR spectrometer (JEOL Ltd., Tokyo, Japan). The measurements were carried out at room temperature with a microwave power of 4 mW, and a modulation width of 0.1 mT. The intensity of all signals were recorded relative to the intensity of a Mn(II)-marker (JEOL Ltd., Tokyo, Japan) attached to the cavity of the spectrometer. Measurements were carried out with varying concentrations of bitter and the degree of inhibition (IESR) was calculated from the height of the central peak of the ESR signal of the spin adduct by the following formula:
Peak heightsample 100% IESR ¼ 1 Peak heightreference
2.7. Statistical analysis Evaluation of statistical significance of differences was performed by using the Student’s t-test.
3. Results and discussion The development of turbidity and change in the total concentration of phenolic compounds in an aromatic bitter were studied over a period of 910 days (2½ years) (Fig. 1). One set of samples of the bitter were kept in bottles that were opened regularly, for 30 s every week and were thus repeatedly exposed to oxygen. The bottles were rotated three times every time they had been opened to ensure a uniform concentration of oxygen. This treatment intended to simulate the handling of the aromatic bitter by a typical consumer, where several small servings may be taken from the same bottle at intervals over a period for more than one
Fig. 1. Turbidity (circles) and concentration of total soluble phenolic compounds (squares) in an aromatic bitter product during storage. The bitter was stored in the dark with either periodical access to atmospheric oxygen (filled symbols) or without access to oxygen (open symbols). Standard deviation is indicated by error bars.
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year. Reference samples were stored under identical conditions but without access to oxygen. The levels of turbidity were low in the aromatic bitter during the first 200 days of storage independently of the presence of oxygen. After this period the turbidity in the bottles that were regularly exposed to oxygen began to increase linearly with time during the rest of the storage, while the turbidity in the bottles without access to oxygen remained low and below the limit for visible detection (50 FNU) (Fig. 1). The low and practically constant level of turbidity during an initial phase (lag phase) followed by a phase where the turbidity increases linearly with time have been observed during storage of a number of polyphenol containing beverages (Fang, Zhang, Du, & Sun, 2007; Lee, Yusof, Hamid, & Baharin, 2007; Siebert, 2006; Tajchakavit, Boye, Bélanger, & Couture, 2001). The mechanism of haze formation is believed to involve an initial oxidation of polyphenolic compounds that generates oxidised polyphenols that can form haze-forming macromolecular complexes with other polyphenols or proteins. The insoluble deposits in the type of aromatic bitter used in this study have previously been shown to be a coprecipitate of polyphenols, proteins and carbohydrates (Refsgaard, Schaumburg, et al., 1996). Proteins accounted for 16.5% of the weight of the solid, and glucose, most likely bound as anthocyanins, was the dominating sugar component (3.2% of the weight of the solid). The involvement of polyphenols in the formation of haze was studied by following the total concentrations of phenols during the storage by the Folin–Ciocalteu method. The concentration of phenolic compounds was constant during the whole storage period in the bottles without access to oxygen (Fig. 1). The changes in the total concentration of soluble phenolic compounds in the presence of oxygen were observed to go through three distinct phases during the storage. The concentration of phenolic compounds was also constant during the first 250 days in the bottles exposed to oxygen, and it was not statistical significantly different from the concentration in the bottles without access to oxygen. A drop in the concentration of phenolic compounds was observed between days 250 and 400, which coincided with the period of time where the formation of turbidity accelerated. This shows that polyphenols are removed from the solution by the formation of the haze and
Fig. 2. HPLC chromatograms of phenolics compounds in bitter after storage. (a) 0 days, and (b) after 910 days with access to oxygen. The identified compounds are: (1) gallic acid, (2) protocatechuic acid, (3) (+)-catechin, (4) vanillic acid, (5) caffeic acid, (6) syringic acid, (7) ()-epicatechin, (8) p-coumaric acid, (9) taxifolin, (IS) sinapic acid as internal standard, (10) eriodictyol, and (11) eugenol. A–D, nonphenolic compounds.
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insoluble precipitates. However, the level of phenolic compounds only decreased slightly during the last 400 days of storage with access to oxygen, suggesting that all the polyphenols that are precursors for haze formation were depleted, and that the continued formation of haze must have been driven by formation of larger and larger deposit particles by complexation of non-phenolic compounds. The concentrations of specific phenolic compounds in the aromatic bitter were identified and quantified by HPLC analysis in order to examine the role of specific polyphenols as precursors for the formation of the turbidity. Eleven phenolic compounds were identified in the bitter including flavonoids, phenolic acids and eugenol (Rødtjer et al., 2006a) (Fig. 2). The concentration of five of these components, gallic acid, catechin, epicatechin, taxifolin and eriodictyol, decreased during storage of the bitter. Gallic acid was the only phenolic acid where the concentration decreased during storage (Fig. 3). All the gallic acid disappeared gradually during the first 500 days in bitter that was exposed to oxygen, whereas the level of gallic acid was almost constant during the first 400 days in bitter in the absence of oxygen, and a decay was only observed during the last period of the storage. The concentrations of all the identified flavonoids decreased during storage, and the decreases were accelerated by the presence of oxygen (Fig. 4). The major changes in concentrations took place during the first 300 days of storage, and the rates of disappearance in the presence of oxygen was: ()-epicatechin > (+)-catechin taxifolin eriodictyol.
Fig. 3. Change in concentration of phenolic benzoic acid derivates in bitter during storage. The bitter was stored in the dark either at the presence of oxygen (filled symbols) or without oxygen (un-filled symbols). Standard deviation is indicated by error bars.
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Fig. 4. Change in concentration of flavonoids in bitter during storage. The bitter was stored in the dark either at the presence of oxygen (filled symbols) or without oxygen (un-filled symbols). Standard deviation is indicated by error bars.
The disappearance of gallic acid, catechin, epicatechin, and taxifolin during the first 300 days of storage of the bitter in the presence of oxygen took place before the development of the turbidity. Furthermore, these four compounds also disappeared, albeit at a slower rate, in the absence of oxygen where turbidity did not develop in the bitter. This suggests that the initial soluble oxidation products presumably with a quinoidal structure must undergo further chemical and physical transformations before the formation of haze particles takes place. These subsequent transformations must be relatively slow, and accordingly rate determining for haze formation, since the oxidations of these four phenolic compounds were almost completed before the development of haze. It is noteworthy that nearly all gallic acid, catechin, epicatechin had disappeared before the formation of turbidity and the major decrease in the concentrations of the phenolic compounds. An alternative explanation could therefore be that the phenols, gallic acid, catechin, epicatechin, and taxifolin, act as antioxidants in the bitter and hinder oxidation during the initial storage phase. The initial oxidation of these compounds could protect other phenols, which upon oxidation form precipitates, from being oxidised. The antioxidative capacity of the bitter was therefore tested in an assay based on metal catalysed oxidation of ethanol (Graversen, Becker, Skibsted, & Andersen, 2008; Rødtjer et al., 2006b). The fresh bitter as well as the samples, which had been stored for 371 days, all accelerated the formation of radicals in the assay (Fig. 5). This suggests that the phenols in the bitter act as prooxidants and not
as antioxidants. A similar effect of phenols has been observed in wine, which has been explained by their ability to reduce iron and copper ions into the lower oxidation states that are needed for radical formation by the Fenton reaction (Danilewicz, 2003; Elias, Andersen, Skibsted, & Waterhouse, 2009a, 2009b; Li, Guo, & Wang, 2008). The concentration dependences of the three bitter samples tested in the assay were slightly different, which reflect their different profiles of phenolic compounds. The phenols in the sample stored for 371 days with access to oxygen gave slightly less prooxidative effects than the corresponding sample stored in a closed bottle (Fig. 5). This can be explained by the loss of the most easily oxidised phenolics in the former sample. Easily oxidised phenols are also better reductants, which are more likely to reduce Fe(III) and Cu(II) ions, and thereby exert a prooxidant effect. However, the assay also showed that the remaining phenols were also good prooxidants. The concentrations of the benzoic acids, protocatechuic acid, syringic acid and vanillic acid, increased during the storage (Fig. 3). It is possible that the increase in the concentrations of these compounds is caused by degradation of anthocyanidins. In neutral media protocatechuic acid has been shown to be formed by degradation of cyanidin, syringic acid by degradation of malvidin, and vanillic acid by degradation of peonidin (Fleschhut, Kratzer, Rechkemmer, & Kulling, 2006; Woodward, Kroon, Cassidy, & Kay, 2009). The corresponding anthocyanins are also degraded to the same hydroxybenzoic acids in neutral media, but the rates are much slower than the anthocyanidins (Woodward et al., 2009). Rowanberries are a likely source of cyanidin glycosides in the bitter (Koponen, Happonen, Mattila, & Törrönen, 2007). The presence of anthocyanins has also been demonstrated in deposits formed in the bitter (Refsgaard, Schaumburg, et al., 1996). The levels of the three hydroxybenzoic acids increased mostly in the presence of oxygen, which suggest that the release of these compounds is coupled to oxidation of their precursors. Eugenol, which gives an important clove flavour to the product, was the phenolic compound with the highest concentration among the identified phenolics in the bitter. However, the concentration of eugenol did not change during the storage, even in the presence of oxygen (Fig. 6). Similarly, the concentrations of the two cin-
Fig. 5. The effect of addition of increasing volume of bitter on the relative amount of radicals (spin adducts) produced in a Fenton/ethanol model system. The bitter samples were stored in 0 days (j), 371 days without oxygen (d), or 371 days with oxygen (N). The spin trap POBN (3 mM) and FeSO4 (106 lM) dissolved in 1 M aqueous ethanol was mixed with the bitter and H2O2 (463 lM).
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References
Fig. 6. Concentration of eugenol in bitter during storage. The bitter was stored in the dark either in the presence of oxygen (filled symbols) or without oxygen (unfilled symbols). Standard deviation is indicated by error bars.
namic acids, caffeic acid (3.5 ppm) and p-coumaric acid (0.9 ppm), did not change during the storage (results not shown). In conclusion, it was shown that before visible turbidity develops in an aromatic bitter some phenolic compounds have already been consumed due to oxidation. The easily oxidised phenolics in the bitter are converted into soluble intermediates most likely with a quinoidal structure, which slowly undergo polymerisation reactions with proteins and carbohydrates forming haze and deposits. Turbidity did not appear in bottles where oxygen was excluded during the storage. Other phenolic compounds proved to be stable against oxidation during the prolonged storage, and the concentration of some even increased, presumably due to break down of anthocyanidins and anthocyanins. Acknowledgements This research was sponsored by the Danish Ministry of Food, Agriculture, and Fisheries through LMC – Centre for Advanced Food Studies as a part of the collaboration project ‘‘Plant phenols in cherry liqueur and bitter. Product stability and utilization of pomace in food production” with Danish Distillers.
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