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Purification and characterization of a p-coumarate decarboxylase and a vinylphenol reductase from Brettanomyces bruxellensis
International Journal of Food Microbiology 127 (2008) 6–11
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International Journal of Food Microbiology j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / i j f o o d m i c r o
Purification and characterization of a p-coumarate decarboxylase and a vinylphenol reductase from Brettanomyces bruxellensis Liliana Godoy a, Claudio Martínez a, Nelson Carrasco b, María Angélica Ganga a,⁎ a b
Departamento de Ciencia y Tecnología de los Alimentos, Facultad Tecnológica, Chile Departamento de Química de los Materiales, Facultad de Química y Biología, Universidad de Santiago de Chile, Alameda 3363, Estación Central, Santiago, Chile
A R T I C L E
I N F O
Article history: Received 5 December 2007 Received in revised form 9 May 2008 Accepted 11 May 2008 Keywords: Coumarate decarboxylase Vinylphenol reductase Brettanomyces sp. Wine Volatile phenols
A B S T R A C T The presence of Brettanomyces bruxellensis has been correlated with an increase of phenolic aromas in wine. The production of these aromas results from the metabolization of cinnamic acids, present in the wine, to their ethyl derivatives. Hence, the participation of two enzymes has been proposed: a p-coumarate decarboxylase (CD) and a vinylphenol reductase (VR). Both enzymes were purified and characterized from B. bruxellensis. In denaturing conditions, the CD enzyme had a molecular mass of 21 kDa, while in native conditions its mass was 41 kDa. The optimal activity was obtained at a temperature of 40 °C and a pH of 6.0. For p-coumaric acid, the Km value and Vmax were 1.22 ± 0.08 mM and 98 ± 0.15 µmol/min mg, respectively. The VR enzyme had a molecular mass of 37 kDa in SDS-PAGE, while in natural conditions its mass was 118 kDa. The Km value was N 3.37 ± 2.05 mM and its Vmax was 107.62 ± 50.38 µmol/min mg for NADPH used as a cofactor. Both enzymatic activities were stable at pH 3.4, but in the presence of ethanol the CD activity decreased drastically while the VR activity was more stable. This is the first report that shows the presence of a CD and a VR enzyme in B. bruxellensis. © 2008 Elsevier B.V. All rights reserved.
1. Introduction The aromatic components present in wine come from the grape and fermentation process giving a complex mixture of volatile organic compounds with different chemical nature and organoleptic characteristics (Cayot, 2007). Within this group of compounds the volatile phenols have an important influence on the aroma of the wine, where 4-vinylphenol, 4-vinylguaiacol, 4-ethylphenol and 4-ethylguaiacol are the most important molecules of this class (Heresztyn, 1986; Chatonnet et al., 1992). The high concentrations of these components are thought to be responsible for the “horsy”, “medicinal”, “animal”, “smoky” and “spicy” aromas (Chatonnet et al., 1992; Suárez et al., 2007). The phenolic acids, also called hydroxycinnamic acids, are the precursors of these volatile phenols and are usually present in small quantities in grapes (20 mg/L to 50 mg/L) (Baranowski et al., 1980; Golderg et al., 1998, Shinohara et al., 2000). One of the properties of these phenolic acids is to inhibit microbial growth (Baranowski et al., 1980; Theodorou et al., 1987; Stead, 1995; Said et al., 2004). It has been observed that those microorganisms that are capable of fermenting products of plant origin present enzymatic activities that would allow the rapid metabolization and transformation of these compounds to less toxic ones. The hydroxycinnamate decarboxylase activity would participate in this metabolization pathway described in numerous
⁎ Corresponding author. Tel.: +56 2 7184509; fax: +56 2 7764796. E-mail address: [email protected] (M.A. Ganga). 0168-1605/$ – see front matter © 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.ijfoodmicro.2008.05.011
microorganisms (Finkle et al., 1962; Huang et al., 1993; Clausen et al., 1994; Degrassi et al., 1995; Cavin et al., 1997a,b, 1998; Shinohara et al., 2000; Van Beek and Priest, 2000; Chamkha et al., 2001; Smit et al., 2003; Coghe et al., 2004). Subsequently, this enzymatic activity would only transform the hydroxycinnamic acids into their vinyl derivatives. However, some microorganisms have been described that could metabolize these vinyl derivatives into their respective ethyl derivatives (Chatonnet et al., 1992; Edlin et al., 1998; Chamkha et al., 2001). In particular, Chatonnet et al. (1992) showed that the presence of the yeast Brettanomyces sp. showed a correlation with the organoleptic detection of phenolic aromas in this product. Species of this genus have been mainly described during the maturity period of wines where its growth is correlated to the presence of 4-ethylphenol (Chatonnet et al., 1995; Stender et al., 2001; Dias et al., 2003). Chatonnet et al. (1992) showed that although many of the wine yeast species studied can produce 4-vinylphenol from p-coumaric acid, only species of the Brettanomyces genus could metabolize this latter cinnamic acid to 4ethylphenol. However, recent studies have described that Pichia guilliermondii is also capable of producing 4-ethylphenol in wine, although at lower concentrations when comparing its production to B. bruxellensis (Barata et al., 2006). Efforts have been made to define a possible metabolic pathway followed by Brettanomyces sp. for the transformation of hydroxycinnamic acids into their corresponding phenolic derivatives (Chatonnet et al., 1995). It has been proposed that this microorganism transforms p-coumaric acid to 4-vinylphenol through a coumarate decarboxylase activity similar to the one described for bacteria (Degrassi et al., 1995; Cavin et al., 1997b) and
L. Godoy et al. / International Journal of Food Microbiology 127 (2008) 6–11
S. cerevisiae (Clausen et al., 1994). This has been confirmed through the purification of this enzyme from Brettanomyces anomalus (Edlin et al., 1998). Subsequently, this vinyl derivative through a vinylphenol reductase activity would be metabolized to 4-ethylphenol (Chatonnet et al., 1992), however, this latter enzyme has not yet been purified from any of the species of this yeast genus. B. bruxellensis is recognized as an agent of phenolic taint in wine, where it has been isolated from the main wine-making countries (Chatonnet et al., 1992; Mitrakul et al., 1999; Rodrigues et al., 2001; Loureiro and Malfeito-Ferreira, 2003; Ganga and Martínez, 2004; Conterno et al., 2006) and it is therefore of interest to study the metabolization pathway that this yeast uses to produce 4-ethylphenol from p-coumaric acid. Therefore, the objective of this work has been to purify and characterize those enzymes involved in the metabolism of the hydroxycinnamic acids to their ethyl derivatives by this species. 2. Materials and methods 2.1. Microorganisms and culture conditions B. bruxellensis L-2480 was obtained from the strain collection maintained at the Laboratorio de Microbiología Aplicada y Biotecnología of the Universidad de Santiago de Chile. Strain S. cerevisae CECT 1451 was obtained from the Spanish Type Culture Collection. Both microorganisms were initially grown in 200 mL of media containing 20 g/L glucose and 6.7 g/L YNB on a shaker. To induce the enzymatic activities under study, 100 mg/L of p-coumaric acid or 100 mg/L of vinylphenol were added separately. The pH was adjusted to 5.0 with HCl. The culture was initially maintained at 30 °C for five days. The microorganism population was controlled using cell count in a Neubauer chamber. 2.2. Chemicals and reagents High Q anionic exchange column, gel filtration standard and protein standards of known isoelectric point (pI) were purchased from Bio-Rad while Sephacryl HR-200 molecular exclusion column was purchased from Amersham Biosciences. p-coumaric acid, 4-vinylphenol, caffeic acid, ferulic acid and m-coumaric acid were purchased from Sigma. All other chemicals and reagents were of analytical grade and were purchased from commercial sources. 2.3. Protein extraction and quantification The protein extraction method used was based on that described by Sambrook et al. (1998). Briefly, 200 mL of culture was centrifuged at 12,000 ×g for 1 min at 4 °C. The pellet was recovered and resuspended in phosphate-buffered saline. The cells were then centrifuged and the supernatant discarded. The pellet was resuspended in 200 mL of buffer A (1 M sorbitol, 10 mM MgCl2, 2 mM dithiothreitol, 50 mM potassium phosphate (pH 7.8) and 100 g/mL de phenylmethylsulfonyl fluoride). The suspension was incubated for 10 min at 30 °C and then centrifuged. The pellet was resuspended in 200 mL buffer B (1 M sorbitol, 10 mM MgCl2, 2 mM dithiothreitol, 25 mM potassium phosphate (pH 7.8), 25 mM sodium succinate (pH 5.5) and 100 µg/mL phenylmethylsulfonyl fluoride). The mixture was incubated for 2 min at 30 °C followed by the addition of 0.25 volume of a 10 mg/L solution of zymolase 100T (Seikagaku Corporation, Japan) and incubated at 30 °C for 30 min. The protoplasts obtained were recovered by centrifugation at 5000 ×g for 10 min and resuspended in 40 mL of lysis buffer (50 mM HEPES, 1% NP-40, 1 µg/mL aprotinin, 100 µg/mL PMSF) at 0 °C for 30 min. The suspension was centrifuged at 12,000 ×g for 10 min at 4 °C and the supernatant recovered was kept at 4 °C for subsequent studies. The protein concentration quantification was carried out using the method described by Bradford (1976), using bovine serum albumin as standard.
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2.4. Enzyme assay The activity assays of the enzymes were carried out in phosphate potassium buffer 50 mM, pH 6.0. These were established as standard conditions. The detection method of coumarate decarboxylase (CD) activity was based on that described by Edlin et al. (1998) with some modifications. The reaction mixture contained 200 μL of protein extract, 50 mM phosphate buffer pH 6.0 and 2 mM of p-coumaric acid, and was incubated at 40 °C for 40 min. Subsequently, the mixture was diluted 50 times to avoid interference with the proteins. The decarboxylase activity was monitored by the decrease in absorbance at 285 nm. One unit (U) of enzymatic activity was defined as the amount of enzyme that consumes 1 µmol of p-coumaric acid per minute. The detection method of the vinylphenol reductase (VR) activity was based on that described by Li and Rosazza (2000) with some modifications. The reaction mixture was composed of 200 μL of protein extract, 50 mM phosphate buffer pH 6.0, 0.15 mM NADPH and 2 mM 4-vinylphenol. This mixture was incubated at 20 °C for 60 min, and the reaction stopped with 25 mM Tris–Cl, 0.3% SDS. The VR activity was monitored by the decrease in absorbance at 340 nm of NADPH, an oxidizable cofactor present in the reaction. One unit (U) of enzymatic activity was defined as the amount of enzyme that consumes 1 µmol of NADPH per minute. 2.5. Enzyme purification All the experiments described below were carried out at 15 °C in Bis–Tris–HCl 20 mM pH 6.0 buffer for both enzymes. Dias et al. (2003) showed that the maximum production of 4-ethylphenol was obtained during the logarithmic growth phase, time during which there could be a greater production of both enzymes due to a greater cellular mass. Based on these observations and with the purpose of purifying the CD and the VR enzymes from B. bruxellensis L-2480 after 3 days of growth, the cells were collected and the enzymes purified. 2.5.1. Purification of the CD enzyme A 10 mL volume of protein extract, obtained from a culture grown in the presence of 100 mg/L of p-coumaric acid was dialysed overnight in Bis–Tris–HCl 20 mM pH 6.0 buffer. Subsequently, the extract was applied to a High Q anionic exchange column (20 × 1.5 cm) and washed at a flow rate of 1.0 mL/min with 30 mL of Bis–Tris–HCI 20 mM pH 6.0 buffer. Using the same flow rate, an increasing NaCl gradient of between 0.0 and 1.0 M was carried out for 25 min. A total of 25 fractions, of 1.5 mL each, were collected and the CD and reductase activity on 4-vinylphenol were determined for each. The fraction containing the purified protein was stored at −20 °C for further analyses. 2.5.2. Purification of the VR enzyme A 10 mL volume of protein extract, obtained from a culture grown in the presence of 100 mg/L of 4-vinylphenol was dialysed overnight in Bis–Tris–HCl 20 mM pH 6.0 buffer. The extract was then applied to a Sephacryl HR-200 molecular exclusion column (80 × 10 cm) which was washed with a flow rate of 1.0 mL/min with 40 mL Bis–Tris–HCI 20 mM pH 6.0 buffer. Using the same flow rate, 40 fractions of 1.0 mL each were collected and the CD and VR activities were determined. The fractions that showed reductase activity were dialysed overnight in Bis–Tris–HCl 20 mM pH 6.0 buffer and concentrated by ultrafiltration in Ultrafree®-MC Centrifugal Filter Units with UF Membrane 10 NMWL (Millipore Corporation). The dialysed and concentrated solution was applied to a High Q anionic exchange column (20 × 0.2 cm). The column was washed with 30 mL of Bis–Tris–HCl 20 mM pH 6.0 buffer at a flow rate of 1.0 mL/min. Keeping the same flow velocity, an increasing NaCl gradient of between 0.0 and 1.0 M was applied for 20 min. A total of 28 fractions, of 1.0 mL each were
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L. Godoy et al. / International Journal of Food Microbiology 127 (2008) 6–11
collected and the reductase activity on 4-vinylphenol was determined for each. The fraction that contained the purified protein was kept at −20 °C for further analysis. 2.6. Polyacrylamide gel electrophoresis (PAGE), isoelectric point (pI) and silver staining Protein electrophoresis was carried out according to Sambrook et al. (1998), while isoelectric focusing (IEF) was performed on precast-IEF polyacrylamide gels containing Bio-Rad carrier ampholytes in the pH range 3.5–9.0. The sample was focused using Mini IEF Cell (Bio-Rad). The isoelectric point was determined by comparison with protein standards of known pI. In both cases (PAGE and IEF) the proteins were visualized using silver staining based on the methodology described by Switer et al. (1979). 2.7. Characterization of the purified CD and VR enzymes 2.7.1. Molecular mass determination of native enzymes The molecular mass of the native enzymes was estimated by Sephacryl HR-200 molecular exclusion (80 × 10 cm) which was eluted with a flow rate of 1.0 mL/min with Bis–Tris–HCI 20 mM pH 6.0 buffer. The column was calibrated using the gel filtration standard and a mixture of weight markers containing the following compounds: Vitamin B12 (1.3 kDa), myoglobin (horse) (17 kDa), ovalbumin (chicken) (44 kDa), γ-globulin (bovine) (158 kDa) and thyroglobulin (bovine) (670 kDa). 2.7.2. Determination of optimum pH and temperature Optimum pH was determined in the range of 3.0–9.0 using the following buffers: acetic/acetate buffer for the range of pH 3.0–5.5; potassium phosphate buffer for pH between 6.0 and 7.5; Tris–HCI for pH 7.5–9.0 and sodium boric/tetraborate acid for pH between 8.0 and 9.0. All the buffers had a final concentration of 50 mM. Once the optimum pH was determined for each enzyme, the temperature effect for each of the enzyme activities was determined in a range of 10– 60 °C. 2.7.3. Kinetic parameters The Michaelis–Menten constants, Km and Vmax, for the CD and VR activities were determined on 0.1–2.0 mM p-coumaric acid and 0.1– 1.4 mM NADPH, respectively. The numerical value of the kinetic constants was obtained by adjusting the specific activity values against the substrate concentration to the hyperbolic equation using the program Origin 5.0 (Microcal Software, Inc). 2.7.4. Substrate specificity 2.7.4.1. CD activity. The CD activity was monitored using p-coumaric acid, caffeic acid, ferulic acid and m-coumaric acid as substrate, each used at a final concentration of 2 mM. The reaction mixture as well as the determination of the enzymatic activity was carried out according to the methodology described above. 2.7.4.2. VR activity. The VR activity was monitored using NADPH and NADH as a second substrate at a final concentration of 0.15 mM. The reaction mixture as well as the enzymatic activity determination was carried out according to the methodology described above.
standard enzymatic assay (Section 2.4) at 1, 3, 5, 10, 30 and 60 min time intervals. To determine the enzymatic stability at pH 3.4, the purified enzymes were incubated in universal Teorell and Stenhagen buffer (Gallego et al., 2001) at 40 °C in the case of CD enzyme and at 25 °C room temperature in the case of VR enzyme. Aliquots were taken at 10, 30, 60, 120 min, 24 and 48 48 h time intervals. Control enzyme fractions were kept in both assays. In the first case, in the absence of ethanol in the incubation mixture, and in the second, the enzymes were kept at pH 6.0 in Teorell and Stenhagen buffer (Gallego et al., 2001). The enzyme activity obtained in each control sample was considered as 100% enzymatic activity. Another factor to consider was the effect of glucose concentration in the reaction media; therefore, glucose was added at a final concentration of 1% (w/v) and 2% (w/v). 2.8. Detection of 4-ethylphenol The detection and quantification of 4-ethylphenol was carried out in the Centro de Aromas of the Pontificia Universidad Católica de Chile using gas chromatography–mass spectrometry (GC–MS) (HP6890, MSH5972; Hewlett-Packard, Palo Alto CA). A DB Waxetr (J&W Scientific Inc., USA) polar column (60 m, 0.25 mm i.d., 0.25 mm film thickness) was used for the analysis with helium as carrier gas at a constant flow at 1.8 mL/min. When the enzymatic reaction was finished, the mixture was treated with dichloromethane to extract the aromatic compound. The temperature of the injected port was 180 °C and a 2 mL volume was injected. The GC–MS program was run for 25.04 min. Chromatogram compounds were identified comparing the mass spectra of each molecule with the NIST–EPA–NIH library as described by Ugarte et al. (2005). The quantitative determination was performed used a labeled molecule of 4-ethylphenol with deuterium atom as internal standard. 2.9. Statistical analysis Data were subjected to analysis of variance (ANOVA) and the mean values of the experiments were statistically analyzed using the LSD test. Differences were considered significant when the probability was b0.05. 3. Results With the purpose of using a S. cerevisiae strain as control for the enzymatic activities studied both the S. cerevisiae and B. bruxellensis strains were grown simultaneously in independent growth media containing p-coumaric acid and 4-vinylphenol. Clausen et al. (1994) described the gene that codes for the PAD enzyme in S. cerevisiae, the activity of which is responsible for metabolizing the cinnamic acids to their vinyl derivatives. With respect to the VR activity, it has not been described in this microorganism which makes it unable to produce ethylphenol from p-coumaric acid (Chatonnet et al.,1992). The CD and VR activities of both microorganisms are shown in Table 1. The S. cerevisiae strain showed a high CD capacity when compared to the results obtained with the B. bruxellensis strain. Additionally, in spite of growing the S. Table 1 CD and VR activities of cultures grown with or without p-coumaric acid and 4vinylphenol Total activity (U) CD activity
2.7.5. Influence of some enological factors on the enzymatic activity To determine how certain factors influence the stability of the enzymatic activities studied, the effect of ethanol concentration and a pH of 3.4 were assayed. In the case of ethanol, the pure enzymatic fractions were kept in potassium phosphate buffer pH 6.0 to which 10% (v/v) and 12% (v/v) ethanol were added separately. The samples were incubated at 25 °C and aliquots were taken to carry out the
Inducer (100 mg/L) Control (uninduced) p-coumaric acid 4-vinylphenol
S. cerevisiae 16.55a 18.80a 20.05a
VR activity B. bruxellensis 5.61a 13.55b 9.85c
S. cerevisiae Not detected Not detected Not detected
B. bruxellensis 10.85a 11.90a 16.55b
Values are the mean of three independent determinations. One unit of enzymatic activity (U) was defined as the amount of enzyme that consumes 1 µmol p-coumaric acid (CD activity) or NADPH (VR activity) per min. Means of the same column with the same superscript letters are not significantly different (p b 0.05).
L. Godoy et al. / International Journal of Food Microbiology 127 (2008) 6–11 Table 2 Purification stages of the CD and VR enzymes from B. bruxellensis L-2480 A) Purification of CD enzyme Step
Total activity (U)a
Protein concentration (µg/mL)
Specific activity (U/mg)
Yield (%)
Purification (fold)
Crude extract Ionic exchange
4.7 3.9
185 5
2.5 173.3
100 83
1.0 68.2
B) Purification of VR enzyme Step
Total activity (U)b
Protein concentration (µg/mL)
Specific activity (U/mg)
Yield (%)
Purification (fold)
Crude extract Gel filtration Ionic exchange
5.8 5.4 2.4
200 10 2
2.9 90.0 600.0
100 93 41
1.0 30.8 205.5
a One unit of enzymatic activity (U) was defined as the amount of enzyme that consumes 1 µmol p-coumaric acid per min. b One unit of enzymatic activity (U) was defined as the amount of enzyme that consumes 1 µmol NADPH per min.
cerevisiae strain in media containing p-coumaric acid or 4-vinylphenol, there was no significant increase in this activity when compared to the control media (absence of these compounds), as opposed to what was observed for B. bruxellensis where there was an increase in the CD activity when grown in media containing both compounds. In the case of the VR activity, this was not detected in S. cerevisiae. Having assayed different S. cerevisiae strains, Chatonnet et al. (1992) showed that this species is unable to transform p-coumaric acid into 4ethylphenol, which agrees with our results, since this microorganism lacks the VR activity responsible for the production of 4-ethylphenol. In the case of B. bruxellensis the VR activity also increased when grown in the presence of 4-vinylphenol, however the activity was unaffected when the microorganism was grown in the presence of p-coumaric acid. 3.1. Purification of enzymes The data of the different purification steps of the CD and VR enzymes are shown in Table 2. In the case of the CD enzyme once the cells were collected, these were lysed and the protein was purified following the steps described in materials and methods. The protein extract was loaded onto an ionic exchange column and then eluted in linear gradient of 0 to 1 M NaCl. The enzyme was purified 68.2-fold to a specific activity of 173.3 U/mg protein from cell with a yield of 83% (Table 2). The purified enzyme gave a single band in SDS-PAGE, indicating that the purified sample was electrophoretically homogeneous under the dissociating conditions. The molecular mass estimated by SDS-PAGE was 21 kDa. The relative molecular mass of the enzyme estimated by gel filtration on Sephacryl HR-200 was 41 kDa. The pI value was 8.0.
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In the case of the VR enzyme, once the yeasts were grown in the presence of 4-vinylphenol, these were collected and lysed. The protein extract was subjected to the purification process summarized in Table 2. The enzyme was purified 205-fold to a specific activity of 600 U/mg protein from cell with a yield of 41%. The purified enzyme gave a single band in SDS-PAGE with an estimated molecular mass of 37 kDa. The relative molecular mass of the native protein estimated after a gel filtration on Sephacryl HR-200 was 118 kDa with a pI of 6.5–7.0. Although it was suspected that this enzyme activity existed in yeast, until this study, it had not been purified. With the purpose of effectively confirming that the enzyme purified in our study was able to metabolize 4-vinylphenol to 4ethylphenol, besides the spectrophotometric tracking of the disappearance of NADPH (see materials and methods) with samples of pure protein, we also carried out the detection and quantification of 4-ethylphenol in the enzymatic reaction by GC–MS. The production of 837.5±27.5 µg/L of 4ethylphenol was observed in the enzymatic reaction. 3.2. Characterization of the purified CD and VR enzymes The activity of the purified CD enzyme from B. bruxellensis L-2480 had an optimum pH of 6.0 and an optimum temperature of 40 °C. The enzyme was active in a pH range of 3.0–8.0, while its activity decreases rapidly near 50 °C. The kinetic parameters were determined by adjusting the curve to the hyperbolic equation giving a Km of 1.22± 0.08 mM, a Vmax of 98±0.15 µmol/min mg and a Kcat of 2.5× 103 s− 1 for p-coumaric acid. On the other hand, the purified VR enzyme showed a pH and temperature optimum of 6.0 and 25 °C, respectively. The enzyme was active in a pH range of 3.0–10.0, while its activity declined rapidly near 50 °C. The kinetic parameters were determined by adjusting the curve to the hyperbolic equation obtaining a Km of N3.37 ± 2.05 mM, a Vmax of 107.62 ± 50.38 µmol/min mg and a Kcat of 1.1 × 103 s− 1 using NADPH as cofactor. The high deviations observed for the kinetic parameters of this enzyme resulted from the erratic behavior of the method used to measure this activity when high concentrations of substrate were used (NADPH). The conditions of initial velocity were very far from the enzyme saturation; therefore the adjustment of the curve to a Michaelis behavior generated Km and Vmax values with high deviations. To reduce the standard error of the samples, it would be necessary to quantify these kinetic parameters using 4-vinylphenol as the enzyme substrate. 3.2.1. Influence of some enological factors on the CD and VR activities The effect of ethanol and pH 3.4 were assayed to evaluate how some factors of the wine have an influence on the stability of the purified enzymes. Likewise, it was determined if the presence of glucose in the reaction mixture could have an influence on the activity of both enzymes. In the case of CD activity was possible to determine that after a few minutes of incubation in the presence of 10% (v/v) and 12% (v/v) ethanol this activity decreases drastically. After 10 min of incubation the enzyme loses 50% of its activity, and after 1 h the activity is completely lost (Table 3). On the other hand, the CD activity is very stable when incubated at pH 3.4 where after 48 h of incubation only a 25% decay was observed (Table 4). The presence of glucose at a
Table 3 Effect of ethanol on the stability of the CD and VR activities Time (min)
1 3 5 10 30 60
Relative activitya (%)
Relative activitya (%)
Enzyme incubated with 10% ethanol
Enzyme incubated with 12% ethanol
Table 4 Effect of pH 3.4 on the stability of the CD and VR activities Relative activitya (%)
Time (h)
CD
VR
CD
VR
90 83 74 50 24 0
95 90 86 80 40 20
90 80 70 48 18 0
90 84 76 66 36 12
a Values are the means of three independent determinations and expressed as the percentage obtained by incubating the enzyme in the absence of ethanol (100%).
0.16 0.5 1 2 24 48 a
CD
VR
96 96 94 90 84 75
100 100 96 96 90 85
Values are the means of three independent determinations and expressed as the percentage obtained by incubating the enzyme in Bis–Tris pH 6.0 buffer (100%).
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L. Godoy et al. / International Journal of Food Microbiology 127 (2008) 6–11
concentration of 1% and 2% in the reaction mixture had no influence on the CD activity (data not shown). The stability of the VR enzyme decreases gradually in time in the presence of 10% and 12% ethanol (Table 3). After a 1 h incubation in the presence of 10% ethanol, the enzyme lost 80% of its activity, while at a concentration of 12% ethanol its enzymatic activity decreased by 88%. On the other hand, the enzymatic stability studied by incubating the protein at pH 3.4, showed that this factor did not cause changes in its stability and even after 48 h of incubation, only 15% of its initial activity was lost (Table 4). The presence of 1% or 2% glucose in the enzymatic reaction mixture had no influence on the VR activity (data not shown). 3.2.2. Substrate specificity of CD activity and assay of VR activity using a different cofactor In order to determine if the CD activity is capable of decarboxilating different hydroxycinamic acids, three hydroxycinamic acid substrates were evaluated: caffeic acid, ferulic acid and m-coumaric acid. They gave relative decarboxylase activities (versus p-coumaric acid) of 120% and 80% for caffeic acid and ferulic acid, respectively. In the case of m-coumaric acid, the enzyme did not show activity with this substrate. To carry out the reduction of 4-vinylphenol, it is necessary to have an oxidizable cofactor. Therefore, the use of NADH as co-factor was evaluated. As a result, a 25% decrease in the enzyme activity of VR was observed when compared to the use of NADPH as cofactor.
proposed by Huang et al. (1994) where an unsubstituted hydroxy group at a para- position to the unsaturated side chain is necessary for the decarboxylation of cinnamic acid. On the other hand, the purified VR enzyme presents an optimum pH of 6.0 and optimum temperature of 25 °C. The ethylphenol aroma in wine is mainly detected during the period of maturity in barrels, which is also the time in which the B. bruxellensis population is greatest. The CD and VR enzymes are intracellular, therefore when B. bruxellensis reaches the stationary growth phase it would lyse and release both enzymes into the wine. This would allow the enzymes to act directly on the cinnamic acids. We studied the effect of glucose, ethanol and the pH 3.4 on the activity and stability of the enzymes. In the presence of alcohol, the CD activity is more sensitive that the VR activity. Dias et al. (2003) observed that B. bruxellensis varies in its capacity to produce volatile phenols since a greater production of these has been observed when the alcohol concentration is lower. This could occur because growth of the microorganisms is somehow affected as well as the enzymatic activities that participate in the formation of ethylphenols which are sensitive to ethanol. Both enzymes are quite stable when incubated at pH 3.4. Furthermore, it was determined that the presence of glucose at a concentration of 1% and 2% (w/v) in the reaction media does not affect the enzymatic activities either. Knowledge of the enzymes that participate in the formation of ethylphenols in B. bruxellensis will allow a greater understanding of the management of some technological variables to control the production of these unwanted phenolic acids during wine production.
4. Discussion Acknowledgment Those microorganisms that ferment natural products such as grapes must have the capacity to metabolize anti-microbial compounds into other less toxic ones (Clausen et al., 1994; Degrassi et al., 1995; Cavin et al., 1997a,b; Edlin et al., 1998). B. bruxellensis initially transforms hydroxycinamic acids into vinyl derivates by a CD activity, and then produces ethylphenol through a VR activity. We observed that the CD activity was induced by the presence of p-coumaric acid as well as 4-vinylphenol in the culture media, as opposed to what was observed for the VR activity which was only induced by the presence of 4-vinylphenol. In P. fluorescens the ferulic decarboxylase acid was found to be constitutive (Huang et al., 1994), contrary to what was observed for a ferulic acid and a p-coumarate decarboxylase from B. pumillus (Degrassi et al., 1995) and a p-coumarate decarboxylase from L. plantarum (Cavin et al., 1997b) which were induced by substrates. In this work, the purification of both enzymes allowed their characterization, providing important information about how these enzymes would act on some wine components. In brief, the CD and VR enzymes have a molecular mass of 21 and 37 kDa, respectively. Edlin et al. (1998) reported a hydroxycinnamate decarboxylase from B. anomalus with a molecular mass of 21.8 KDa, which is very close to the value obtained in this study. A ferulic decarboxylase acid from P. fluorescens had a molecular mass of 20.4 kDa (Huang et al., 1994), while in B. pumillus and L. plantarum its molecular mass was 23 and 23.5 kDa, respectively (Degrassi et al., 1995; Cavin et al., 1997b). Recently, Dhar et al. (2007) purified a vanillic acid decarboxylase that gave a molecular weight of 23.0 kDa. The optimum activity of the CD enzyme of B. bruxellensis L-2480 was at pH 6.0 and at a temperature of 40 °C; similar conditions were described for the hydroxycinnamate decarboxylase enzyme from B. anomalus (Edlin et al., 1998). These values are similar to those obtained for the purified enzymes from B. pumillus, and L. plantarum (Degrassi et al., 1995; Cavin et al., 1997b). The specificity assays showed that this enzyme is capable of decarboxylating a series of hydroxycinnamic acids, with a greater specificity for caffeic acid, a cinnamic acid present in wine. Furthermore, it was possible to determine that the paraposition of the substituent with respect to the hydroxyl group is essential for this activity. These results agree with the mechanism
This work was supported by Fondecyt—Chile 1050104 grant. References Baranowski, J., Davidson, P., Nagel, C., Branen, A., 1980. Inhibition of Saccharomyces cerevisiae by naturally occurring hydroxycinnamates. Journal of Food Science 45, 592–594. Barata, A., Nobre, A., Correia, P., Malfeito-Ferreira, M., Loureiro, V., 2006. Growth and 4ethylphenol production by yeast Pichia guillermondii in grape juice. American Journal of Enology and Viticulture 57, 133–138. Bradford, M., 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein dye binding. Analytical Biochemistry 72, 248–254. Cavin, J., Barthelmebs, L., Divies, C., 1997a. Molecular characterization of an inducible pcoumaric acid decarboxylse from Lactobacillus plantarum: gene cloning, transcriptional analysis, overexpression in Escherichia coli, purification and characterization. Applied and Environmental Microbiology 63, 1939–1944. Cavin, J., Barthelmebs, L., Guzzo, J., Van Beeumen, J., Samyn, B., Travers, J., Diviés, C., 1997b. Purification and characterization of an inducible p-coumaric acid decarboxylase from Lactobacillus plantarum. FEMS Microbiology Letters 147, 291–295. Cavin, J., Dartois, V., Diviés, C., 1998. Gene cloning, transcriptional analysis, purification and characterization of phenolic acid decarboxylase from Bacillus subtilis. Applied and Environmental Microbiology 64, 1466–1471. Cayot, N., 2007. Sensory quality of traditional food. Food Chemistry 102, 445–453. Chamkha, M., Garcia, J., Labat, M., 2001. Metabolism of cinnamic acids by some Clostridiales and emendation of the descriptions of Clostridium aerotolerans, Clostridium celerecrescens and Clostridium xylanolyticum. International Journal of Systematic and Evolutionary Microbiology 51, 2105–2111. Chatonnet, P., Dubourdieu, D., Boidron, J.-N., Pons, M., 1992. The origin of ethylphenols in wine. Journal of Science and Food Agriculture 60, 165–178. Chatonnet, P., Dubourdieu, D., Boidron, J., 1995. The influence of Brettanomyces/Dekkera sp. yeast and lactic acid bacteria on the ethylphenol contain of red wines. American Journal of Enology and Viticulture 46, 463–468. Clausen, M., Lamb, C., Menet, R., Doerner, P., 1994. PAD1 encodes phenylacrylic acid decarboxylase which confer resistance to cinnamic acid in Saccharomyces cerevisiae. Gene 142, 107–112. Coghe, S., Benoot, K., Delvaux, F., Vanderhaegen, B., Delvaux, F.R., 2004. Ferulic acid releases and 4-vinylguaiacol formation during brewing and fermentation: indication for feruloyl esterase activity in Saccharomyces cerevisiae. Journal of Agricultural and Food Chemistry 52, 602–608. Conterno, L., Joseph, C., Arvik, T., Henick-Kling, T., Bisson, L., 2006. Genetic and physiological characterization of Brettanomyces bruxellensis strains isolated from wines. American Journal of Enology and Viticulture 57, 139–147. Degrassi, G., Polverino de Laureto, P., Bruschi, C., 1995. Purification and characterization of ferulate p-coumarate decarboxylase from Bacillus pumilus. Applied and Environmental Microbiology 61, 326–332.
L. Godoy et al. / International Journal of Food Microbiology 127 (2008) 6–11 Dhar, A., Lee, K., Dhar, K., Rosazza, J., 2007. Nocardia sp. vanillic acid decarboxylase. Enzyme and Microbial Technology 41, 271–277. Dias, L., Dias, S., Sancho, T., Stender, H., Querol, A., Malfeito-Ferreira, M., Loureiro, V., 2003. Identification of yeast isolated from wine-related environments and capable of producing 4-ethylphenol. Food Microbiology 20, 567–574. Edlin, D., Narrad, A., Gasson, M., Dickinson, J., Lloyd, D., 1998. Purification and characterization of hydroxycinnamate decarboxylase from Brettanomyces anomalus. Enzyme and Microbial Technology 22, 232–239. Finkle, B., Lewis, J., Corse, J., Lindin, R., 1962. Enzyme reactions with phenolic compounds: formation of hydroxystyrenes through the decarboxylation of 4hydroxycinnamic acids by Aerobacter. Journal of Biological Chemistry 237, 2926–2931. Gallego, M., Piñaga, F., Ramón, D., Vallés, S., 2001. Purification and characterization of an a-L-rhamnosidase from Aspergillus terreus of interest in winemaking. Food and Chemical Toxicology 66, 204–209. Ganga, A., Martínez, C., 2004. Effect of wine yeast monoculture practice on the biodiversity of non-Saccharomyces yeast. Journal of Applied Microbiology 96, 76–83. Golderg, D., Tsang, E., Karumanchiri, A., Soleas, G., 1998. Quercetin and p-coumaric acid concentration in commercial wines. American Journal of Enology and Viticulture 49, 142–151. Heresztyn, T., 1986. Metabolism of volatile phenolic compounds from hydroxycinnamic acids by Brettanomyces yeast. Archives of Microbiology 146, 96–98. Huang, Z., Dostal, L., Rosazza, L., 1993. Microbial transformation of ferulic acid by Saccharomyces cerevisiae and Pseudomonas fluorescens. Applied and Environmental Microbiology 59, 2244–2250. Huang, Z., Dostal, L., Rosazza, J., 1994. Purification and characterization of a ferulic acid decarboxylase from Pseudomonas fluorescens. Journal of Bacteriology 176, 5912–5918. Li, T., Rosazza, J., 2000. The carboxylic acid reduction pathway in Nocardia. Purification and characterization of the aldehyde reductase. Journal of Industrial Microbiology and Biotechnology 25, 328–332. Loureiro, V., Malfeito-Ferreira, M., 2003. Spoilage yeasts in the wine industry. International Journal of Food Microbiology 86, 23–50. Mitrakul, C., Henick-Kling, T., Egli, C., 1999. Discrimination of Brettanomyces/Dekkera yeast isolates from wine by using various DNA fingerprinting methods. Food Microbiology 16, 3–14.
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Rodrigues, N., Goncalves, G., Malfeito-Ferreira, M., Loureiro, V., 2001. Development and use a differential medium to detect yeast of the genera Dekkera/Brettanomyces. Journal of Applied Microbiology 90, 588–599. Said, S., Neves, F., Griffiths, A., 2004. Cinnamic acid inhibits the growth of the fungus Neurospora crassa, but is eliminated as acetophenone. International Biodeterioration and Biodegradation 54, 1–6. Sambrook, J., Fritsch, F., Maniatis, T., 1998. Molecular Cloning: a Laboratory Manual, 2nd ed. Cold Spring harbor Laboratory, Cold Spring Harbor, N.Y. Shinohara, T., Kubodera, S., Yanagida, F., 2000. Distribution of phenolic yeast and production of phenolic off-flavors in wine fermentation. Journal of Bioscience and Bioengineering 90, 90–97. Smit, A., Cordero, R., Lambrechts, M., Pretorious, I., van Rensburg, P., 2003. Enhancing volatile phenol concentration in wine by expressing various phenol acid decarboxylase genes in Sacharomyces cerevisiae. Journal of Agricultural and Food Chemistry 51, 4909–4915. Stead, D., 1995. The effect of hydroxycinnamic acids and potassium sorbate on the growth of 11 strain of spoilage yeast. Journal of Applied Bacteriology 78, 82–87. Stender, H., Kutzman, C., Holding, J., Sorensen, D., Broomer, A., Oliveira, K., Perry, H., Sage, A., Young, B., Couli, J., 2001. Identification of Dekkera bruxellensis (Brettanomyces) from wine by fluorescence in situ hybridization using peptide nucleic acid probes. Applied and Environmental Microbiology 67, 938–941. Suárez, R., Suárez-Lepe, J., Morata, A., Calderón, F., 2007. The production of ethylphenols in wine by yeast of the genera Brettanomyces and Dekkera. A review. Food Chemistry 102, 10–21. Switer, R., Merril, C., Shifrin, S., 1979. A highly sensitive silver stain for detecting proteins and peptides in polyacrylamide gels. Analytical Biochemistry 98, 231–237. Theodorou, M., Gascoyne, D., Akin, D., Hartley, R., 1987. Effect of phenolic acids and phenolics from plant cell walls on rumenlike fermentation in consecutive batch culture. Applied and Environmental Microbiology 53, 1046–1050. Ugarte, P., Agosin, E., Bordeu, E., Villalobos, J., 2005. Reduction of 4 -ethylphenol and 4ethylguaiacol concentration in red wine using reverse osmosis and adsorption. American Journal of Enology and Viticulture 56, 30–36. Van Beek, S., Priest, F., 2000. Decarboxylation of substitutes cinnamic acid by lactic acid bacteria isolates during whisky fermentation. Applied and Environmental Microbiology 66, 5322–5328.
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International Journal of Food Microbiology j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / i j f o o d m i c r o
Purification and characterization of a p-coumarate decarboxylase and a vinylphenol reductase from Brettanomyces bruxellensis Liliana Godoy a, Claudio Martínez a, Nelson Carrasco b, María Angélica Ganga a,⁎ a b
Departamento de Ciencia y Tecnología de los Alimentos, Facultad Tecnológica, Chile Departamento de Química de los Materiales, Facultad de Química y Biología, Universidad de Santiago de Chile, Alameda 3363, Estación Central, Santiago, Chile
A R T I C L E
I N F O
Article history: Received 5 December 2007 Received in revised form 9 May 2008 Accepted 11 May 2008 Keywords: Coumarate decarboxylase Vinylphenol reductase Brettanomyces sp. Wine Volatile phenols
A B S T R A C T The presence of Brettanomyces bruxellensis has been correlated with an increase of phenolic aromas in wine. The production of these aromas results from the metabolization of cinnamic acids, present in the wine, to their ethyl derivatives. Hence, the participation of two enzymes has been proposed: a p-coumarate decarboxylase (CD) and a vinylphenol reductase (VR). Both enzymes were purified and characterized from B. bruxellensis. In denaturing conditions, the CD enzyme had a molecular mass of 21 kDa, while in native conditions its mass was 41 kDa. The optimal activity was obtained at a temperature of 40 °C and a pH of 6.0. For p-coumaric acid, the Km value and Vmax were 1.22 ± 0.08 mM and 98 ± 0.15 µmol/min mg, respectively. The VR enzyme had a molecular mass of 37 kDa in SDS-PAGE, while in natural conditions its mass was 118 kDa. The Km value was N 3.37 ± 2.05 mM and its Vmax was 107.62 ± 50.38 µmol/min mg for NADPH used as a cofactor. Both enzymatic activities were stable at pH 3.4, but in the presence of ethanol the CD activity decreased drastically while the VR activity was more stable. This is the first report that shows the presence of a CD and a VR enzyme in B. bruxellensis. © 2008 Elsevier B.V. All rights reserved.
1. Introduction The aromatic components present in wine come from the grape and fermentation process giving a complex mixture of volatile organic compounds with different chemical nature and organoleptic characteristics (Cayot, 2007). Within this group of compounds the volatile phenols have an important influence on the aroma of the wine, where 4-vinylphenol, 4-vinylguaiacol, 4-ethylphenol and 4-ethylguaiacol are the most important molecules of this class (Heresztyn, 1986; Chatonnet et al., 1992). The high concentrations of these components are thought to be responsible for the “horsy”, “medicinal”, “animal”, “smoky” and “spicy” aromas (Chatonnet et al., 1992; Suárez et al., 2007). The phenolic acids, also called hydroxycinnamic acids, are the precursors of these volatile phenols and are usually present in small quantities in grapes (20 mg/L to 50 mg/L) (Baranowski et al., 1980; Golderg et al., 1998, Shinohara et al., 2000). One of the properties of these phenolic acids is to inhibit microbial growth (Baranowski et al., 1980; Theodorou et al., 1987; Stead, 1995; Said et al., 2004). It has been observed that those microorganisms that are capable of fermenting products of plant origin present enzymatic activities that would allow the rapid metabolization and transformation of these compounds to less toxic ones. The hydroxycinnamate decarboxylase activity would participate in this metabolization pathway described in numerous
⁎ Corresponding author. Tel.: +56 2 7184509; fax: +56 2 7764796. E-mail address: [email protected] (M.A. Ganga). 0168-1605/$ – see front matter © 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.ijfoodmicro.2008.05.011
microorganisms (Finkle et al., 1962; Huang et al., 1993; Clausen et al., 1994; Degrassi et al., 1995; Cavin et al., 1997a,b, 1998; Shinohara et al., 2000; Van Beek and Priest, 2000; Chamkha et al., 2001; Smit et al., 2003; Coghe et al., 2004). Subsequently, this enzymatic activity would only transform the hydroxycinnamic acids into their vinyl derivatives. However, some microorganisms have been described that could metabolize these vinyl derivatives into their respective ethyl derivatives (Chatonnet et al., 1992; Edlin et al., 1998; Chamkha et al., 2001). In particular, Chatonnet et al. (1992) showed that the presence of the yeast Brettanomyces sp. showed a correlation with the organoleptic detection of phenolic aromas in this product. Species of this genus have been mainly described during the maturity period of wines where its growth is correlated to the presence of 4-ethylphenol (Chatonnet et al., 1995; Stender et al., 2001; Dias et al., 2003). Chatonnet et al. (1992) showed that although many of the wine yeast species studied can produce 4-vinylphenol from p-coumaric acid, only species of the Brettanomyces genus could metabolize this latter cinnamic acid to 4ethylphenol. However, recent studies have described that Pichia guilliermondii is also capable of producing 4-ethylphenol in wine, although at lower concentrations when comparing its production to B. bruxellensis (Barata et al., 2006). Efforts have been made to define a possible metabolic pathway followed by Brettanomyces sp. for the transformation of hydroxycinnamic acids into their corresponding phenolic derivatives (Chatonnet et al., 1995). It has been proposed that this microorganism transforms p-coumaric acid to 4-vinylphenol through a coumarate decarboxylase activity similar to the one described for bacteria (Degrassi et al., 1995; Cavin et al., 1997b) and
L. Godoy et al. / International Journal of Food Microbiology 127 (2008) 6–11
S. cerevisiae (Clausen et al., 1994). This has been confirmed through the purification of this enzyme from Brettanomyces anomalus (Edlin et al., 1998). Subsequently, this vinyl derivative through a vinylphenol reductase activity would be metabolized to 4-ethylphenol (Chatonnet et al., 1992), however, this latter enzyme has not yet been purified from any of the species of this yeast genus. B. bruxellensis is recognized as an agent of phenolic taint in wine, where it has been isolated from the main wine-making countries (Chatonnet et al., 1992; Mitrakul et al., 1999; Rodrigues et al., 2001; Loureiro and Malfeito-Ferreira, 2003; Ganga and Martínez, 2004; Conterno et al., 2006) and it is therefore of interest to study the metabolization pathway that this yeast uses to produce 4-ethylphenol from p-coumaric acid. Therefore, the objective of this work has been to purify and characterize those enzymes involved in the metabolism of the hydroxycinnamic acids to their ethyl derivatives by this species. 2. Materials and methods 2.1. Microorganisms and culture conditions B. bruxellensis L-2480 was obtained from the strain collection maintained at the Laboratorio de Microbiología Aplicada y Biotecnología of the Universidad de Santiago de Chile. Strain S. cerevisae CECT 1451 was obtained from the Spanish Type Culture Collection. Both microorganisms were initially grown in 200 mL of media containing 20 g/L glucose and 6.7 g/L YNB on a shaker. To induce the enzymatic activities under study, 100 mg/L of p-coumaric acid or 100 mg/L of vinylphenol were added separately. The pH was adjusted to 5.0 with HCl. The culture was initially maintained at 30 °C for five days. The microorganism population was controlled using cell count in a Neubauer chamber. 2.2. Chemicals and reagents High Q anionic exchange column, gel filtration standard and protein standards of known isoelectric point (pI) were purchased from Bio-Rad while Sephacryl HR-200 molecular exclusion column was purchased from Amersham Biosciences. p-coumaric acid, 4-vinylphenol, caffeic acid, ferulic acid and m-coumaric acid were purchased from Sigma. All other chemicals and reagents were of analytical grade and were purchased from commercial sources. 2.3. Protein extraction and quantification The protein extraction method used was based on that described by Sambrook et al. (1998). Briefly, 200 mL of culture was centrifuged at 12,000 ×g for 1 min at 4 °C. The pellet was recovered and resuspended in phosphate-buffered saline. The cells were then centrifuged and the supernatant discarded. The pellet was resuspended in 200 mL of buffer A (1 M sorbitol, 10 mM MgCl2, 2 mM dithiothreitol, 50 mM potassium phosphate (pH 7.8) and 100 g/mL de phenylmethylsulfonyl fluoride). The suspension was incubated for 10 min at 30 °C and then centrifuged. The pellet was resuspended in 200 mL buffer B (1 M sorbitol, 10 mM MgCl2, 2 mM dithiothreitol, 25 mM potassium phosphate (pH 7.8), 25 mM sodium succinate (pH 5.5) and 100 µg/mL phenylmethylsulfonyl fluoride). The mixture was incubated for 2 min at 30 °C followed by the addition of 0.25 volume of a 10 mg/L solution of zymolase 100T (Seikagaku Corporation, Japan) and incubated at 30 °C for 30 min. The protoplasts obtained were recovered by centrifugation at 5000 ×g for 10 min and resuspended in 40 mL of lysis buffer (50 mM HEPES, 1% NP-40, 1 µg/mL aprotinin, 100 µg/mL PMSF) at 0 °C for 30 min. The suspension was centrifuged at 12,000 ×g for 10 min at 4 °C and the supernatant recovered was kept at 4 °C for subsequent studies. The protein concentration quantification was carried out using the method described by Bradford (1976), using bovine serum albumin as standard.
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2.4. Enzyme assay The activity assays of the enzymes were carried out in phosphate potassium buffer 50 mM, pH 6.0. These were established as standard conditions. The detection method of coumarate decarboxylase (CD) activity was based on that described by Edlin et al. (1998) with some modifications. The reaction mixture contained 200 μL of protein extract, 50 mM phosphate buffer pH 6.0 and 2 mM of p-coumaric acid, and was incubated at 40 °C for 40 min. Subsequently, the mixture was diluted 50 times to avoid interference with the proteins. The decarboxylase activity was monitored by the decrease in absorbance at 285 nm. One unit (U) of enzymatic activity was defined as the amount of enzyme that consumes 1 µmol of p-coumaric acid per minute. The detection method of the vinylphenol reductase (VR) activity was based on that described by Li and Rosazza (2000) with some modifications. The reaction mixture was composed of 200 μL of protein extract, 50 mM phosphate buffer pH 6.0, 0.15 mM NADPH and 2 mM 4-vinylphenol. This mixture was incubated at 20 °C for 60 min, and the reaction stopped with 25 mM Tris–Cl, 0.3% SDS. The VR activity was monitored by the decrease in absorbance at 340 nm of NADPH, an oxidizable cofactor present in the reaction. One unit (U) of enzymatic activity was defined as the amount of enzyme that consumes 1 µmol of NADPH per minute. 2.5. Enzyme purification All the experiments described below were carried out at 15 °C in Bis–Tris–HCl 20 mM pH 6.0 buffer for both enzymes. Dias et al. (2003) showed that the maximum production of 4-ethylphenol was obtained during the logarithmic growth phase, time during which there could be a greater production of both enzymes due to a greater cellular mass. Based on these observations and with the purpose of purifying the CD and the VR enzymes from B. bruxellensis L-2480 after 3 days of growth, the cells were collected and the enzymes purified. 2.5.1. Purification of the CD enzyme A 10 mL volume of protein extract, obtained from a culture grown in the presence of 100 mg/L of p-coumaric acid was dialysed overnight in Bis–Tris–HCl 20 mM pH 6.0 buffer. Subsequently, the extract was applied to a High Q anionic exchange column (20 × 1.5 cm) and washed at a flow rate of 1.0 mL/min with 30 mL of Bis–Tris–HCI 20 mM pH 6.0 buffer. Using the same flow rate, an increasing NaCl gradient of between 0.0 and 1.0 M was carried out for 25 min. A total of 25 fractions, of 1.5 mL each, were collected and the CD and reductase activity on 4-vinylphenol were determined for each. The fraction containing the purified protein was stored at −20 °C for further analyses. 2.5.2. Purification of the VR enzyme A 10 mL volume of protein extract, obtained from a culture grown in the presence of 100 mg/L of 4-vinylphenol was dialysed overnight in Bis–Tris–HCl 20 mM pH 6.0 buffer. The extract was then applied to a Sephacryl HR-200 molecular exclusion column (80 × 10 cm) which was washed with a flow rate of 1.0 mL/min with 40 mL Bis–Tris–HCI 20 mM pH 6.0 buffer. Using the same flow rate, 40 fractions of 1.0 mL each were collected and the CD and VR activities were determined. The fractions that showed reductase activity were dialysed overnight in Bis–Tris–HCl 20 mM pH 6.0 buffer and concentrated by ultrafiltration in Ultrafree®-MC Centrifugal Filter Units with UF Membrane 10 NMWL (Millipore Corporation). The dialysed and concentrated solution was applied to a High Q anionic exchange column (20 × 0.2 cm). The column was washed with 30 mL of Bis–Tris–HCl 20 mM pH 6.0 buffer at a flow rate of 1.0 mL/min. Keeping the same flow velocity, an increasing NaCl gradient of between 0.0 and 1.0 M was applied for 20 min. A total of 28 fractions, of 1.0 mL each were
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collected and the reductase activity on 4-vinylphenol was determined for each. The fraction that contained the purified protein was kept at −20 °C for further analysis. 2.6. Polyacrylamide gel electrophoresis (PAGE), isoelectric point (pI) and silver staining Protein electrophoresis was carried out according to Sambrook et al. (1998), while isoelectric focusing (IEF) was performed on precast-IEF polyacrylamide gels containing Bio-Rad carrier ampholytes in the pH range 3.5–9.0. The sample was focused using Mini IEF Cell (Bio-Rad). The isoelectric point was determined by comparison with protein standards of known pI. In both cases (PAGE and IEF) the proteins were visualized using silver staining based on the methodology described by Switer et al. (1979). 2.7. Characterization of the purified CD and VR enzymes 2.7.1. Molecular mass determination of native enzymes The molecular mass of the native enzymes was estimated by Sephacryl HR-200 molecular exclusion (80 × 10 cm) which was eluted with a flow rate of 1.0 mL/min with Bis–Tris–HCI 20 mM pH 6.0 buffer. The column was calibrated using the gel filtration standard and a mixture of weight markers containing the following compounds: Vitamin B12 (1.3 kDa), myoglobin (horse) (17 kDa), ovalbumin (chicken) (44 kDa), γ-globulin (bovine) (158 kDa) and thyroglobulin (bovine) (670 kDa). 2.7.2. Determination of optimum pH and temperature Optimum pH was determined in the range of 3.0–9.0 using the following buffers: acetic/acetate buffer for the range of pH 3.0–5.5; potassium phosphate buffer for pH between 6.0 and 7.5; Tris–HCI for pH 7.5–9.0 and sodium boric/tetraborate acid for pH between 8.0 and 9.0. All the buffers had a final concentration of 50 mM. Once the optimum pH was determined for each enzyme, the temperature effect for each of the enzyme activities was determined in a range of 10– 60 °C. 2.7.3. Kinetic parameters The Michaelis–Menten constants, Km and Vmax, for the CD and VR activities were determined on 0.1–2.0 mM p-coumaric acid and 0.1– 1.4 mM NADPH, respectively. The numerical value of the kinetic constants was obtained by adjusting the specific activity values against the substrate concentration to the hyperbolic equation using the program Origin 5.0 (Microcal Software, Inc). 2.7.4. Substrate specificity 2.7.4.1. CD activity. The CD activity was monitored using p-coumaric acid, caffeic acid, ferulic acid and m-coumaric acid as substrate, each used at a final concentration of 2 mM. The reaction mixture as well as the determination of the enzymatic activity was carried out according to the methodology described above. 2.7.4.2. VR activity. The VR activity was monitored using NADPH and NADH as a second substrate at a final concentration of 0.15 mM. The reaction mixture as well as the enzymatic activity determination was carried out according to the methodology described above.
standard enzymatic assay (Section 2.4) at 1, 3, 5, 10, 30 and 60 min time intervals. To determine the enzymatic stability at pH 3.4, the purified enzymes were incubated in universal Teorell and Stenhagen buffer (Gallego et al., 2001) at 40 °C in the case of CD enzyme and at 25 °C room temperature in the case of VR enzyme. Aliquots were taken at 10, 30, 60, 120 min, 24 and 48 48 h time intervals. Control enzyme fractions were kept in both assays. In the first case, in the absence of ethanol in the incubation mixture, and in the second, the enzymes were kept at pH 6.0 in Teorell and Stenhagen buffer (Gallego et al., 2001). The enzyme activity obtained in each control sample was considered as 100% enzymatic activity. Another factor to consider was the effect of glucose concentration in the reaction media; therefore, glucose was added at a final concentration of 1% (w/v) and 2% (w/v). 2.8. Detection of 4-ethylphenol The detection and quantification of 4-ethylphenol was carried out in the Centro de Aromas of the Pontificia Universidad Católica de Chile using gas chromatography–mass spectrometry (GC–MS) (HP6890, MSH5972; Hewlett-Packard, Palo Alto CA). A DB Waxetr (J&W Scientific Inc., USA) polar column (60 m, 0.25 mm i.d., 0.25 mm film thickness) was used for the analysis with helium as carrier gas at a constant flow at 1.8 mL/min. When the enzymatic reaction was finished, the mixture was treated with dichloromethane to extract the aromatic compound. The temperature of the injected port was 180 °C and a 2 mL volume was injected. The GC–MS program was run for 25.04 min. Chromatogram compounds were identified comparing the mass spectra of each molecule with the NIST–EPA–NIH library as described by Ugarte et al. (2005). The quantitative determination was performed used a labeled molecule of 4-ethylphenol with deuterium atom as internal standard. 2.9. Statistical analysis Data were subjected to analysis of variance (ANOVA) and the mean values of the experiments were statistically analyzed using the LSD test. Differences were considered significant when the probability was b0.05. 3. Results With the purpose of using a S. cerevisiae strain as control for the enzymatic activities studied both the S. cerevisiae and B. bruxellensis strains were grown simultaneously in independent growth media containing p-coumaric acid and 4-vinylphenol. Clausen et al. (1994) described the gene that codes for the PAD enzyme in S. cerevisiae, the activity of which is responsible for metabolizing the cinnamic acids to their vinyl derivatives. With respect to the VR activity, it has not been described in this microorganism which makes it unable to produce ethylphenol from p-coumaric acid (Chatonnet et al.,1992). The CD and VR activities of both microorganisms are shown in Table 1. The S. cerevisiae strain showed a high CD capacity when compared to the results obtained with the B. bruxellensis strain. Additionally, in spite of growing the S. Table 1 CD and VR activities of cultures grown with or without p-coumaric acid and 4vinylphenol Total activity (U) CD activity
2.7.5. Influence of some enological factors on the enzymatic activity To determine how certain factors influence the stability of the enzymatic activities studied, the effect of ethanol concentration and a pH of 3.4 were assayed. In the case of ethanol, the pure enzymatic fractions were kept in potassium phosphate buffer pH 6.0 to which 10% (v/v) and 12% (v/v) ethanol were added separately. The samples were incubated at 25 °C and aliquots were taken to carry out the
Inducer (100 mg/L) Control (uninduced) p-coumaric acid 4-vinylphenol
S. cerevisiae 16.55a 18.80a 20.05a
VR activity B. bruxellensis 5.61a 13.55b 9.85c
S. cerevisiae Not detected Not detected Not detected
B. bruxellensis 10.85a 11.90a 16.55b
Values are the mean of three independent determinations. One unit of enzymatic activity (U) was defined as the amount of enzyme that consumes 1 µmol p-coumaric acid (CD activity) or NADPH (VR activity) per min. Means of the same column with the same superscript letters are not significantly different (p b 0.05).
L. Godoy et al. / International Journal of Food Microbiology 127 (2008) 6–11 Table 2 Purification stages of the CD and VR enzymes from B. bruxellensis L-2480 A) Purification of CD enzyme Step
Total activity (U)a
Protein concentration (µg/mL)
Specific activity (U/mg)
Yield (%)
Purification (fold)
Crude extract Ionic exchange
4.7 3.9
185 5
2.5 173.3
100 83
1.0 68.2
B) Purification of VR enzyme Step
Total activity (U)b
Protein concentration (µg/mL)
Specific activity (U/mg)
Yield (%)
Purification (fold)
Crude extract Gel filtration Ionic exchange
5.8 5.4 2.4
200 10 2
2.9 90.0 600.0
100 93 41
1.0 30.8 205.5
a One unit of enzymatic activity (U) was defined as the amount of enzyme that consumes 1 µmol p-coumaric acid per min. b One unit of enzymatic activity (U) was defined as the amount of enzyme that consumes 1 µmol NADPH per min.
cerevisiae strain in media containing p-coumaric acid or 4-vinylphenol, there was no significant increase in this activity when compared to the control media (absence of these compounds), as opposed to what was observed for B. bruxellensis where there was an increase in the CD activity when grown in media containing both compounds. In the case of the VR activity, this was not detected in S. cerevisiae. Having assayed different S. cerevisiae strains, Chatonnet et al. (1992) showed that this species is unable to transform p-coumaric acid into 4ethylphenol, which agrees with our results, since this microorganism lacks the VR activity responsible for the production of 4-ethylphenol. In the case of B. bruxellensis the VR activity also increased when grown in the presence of 4-vinylphenol, however the activity was unaffected when the microorganism was grown in the presence of p-coumaric acid. 3.1. Purification of enzymes The data of the different purification steps of the CD and VR enzymes are shown in Table 2. In the case of the CD enzyme once the cells were collected, these were lysed and the protein was purified following the steps described in materials and methods. The protein extract was loaded onto an ionic exchange column and then eluted in linear gradient of 0 to 1 M NaCl. The enzyme was purified 68.2-fold to a specific activity of 173.3 U/mg protein from cell with a yield of 83% (Table 2). The purified enzyme gave a single band in SDS-PAGE, indicating that the purified sample was electrophoretically homogeneous under the dissociating conditions. The molecular mass estimated by SDS-PAGE was 21 kDa. The relative molecular mass of the enzyme estimated by gel filtration on Sephacryl HR-200 was 41 kDa. The pI value was 8.0.
9
In the case of the VR enzyme, once the yeasts were grown in the presence of 4-vinylphenol, these were collected and lysed. The protein extract was subjected to the purification process summarized in Table 2. The enzyme was purified 205-fold to a specific activity of 600 U/mg protein from cell with a yield of 41%. The purified enzyme gave a single band in SDS-PAGE with an estimated molecular mass of 37 kDa. The relative molecular mass of the native protein estimated after a gel filtration on Sephacryl HR-200 was 118 kDa with a pI of 6.5–7.0. Although it was suspected that this enzyme activity existed in yeast, until this study, it had not been purified. With the purpose of effectively confirming that the enzyme purified in our study was able to metabolize 4-vinylphenol to 4ethylphenol, besides the spectrophotometric tracking of the disappearance of NADPH (see materials and methods) with samples of pure protein, we also carried out the detection and quantification of 4-ethylphenol in the enzymatic reaction by GC–MS. The production of 837.5±27.5 µg/L of 4ethylphenol was observed in the enzymatic reaction. 3.2. Characterization of the purified CD and VR enzymes The activity of the purified CD enzyme from B. bruxellensis L-2480 had an optimum pH of 6.0 and an optimum temperature of 40 °C. The enzyme was active in a pH range of 3.0–8.0, while its activity decreases rapidly near 50 °C. The kinetic parameters were determined by adjusting the curve to the hyperbolic equation giving a Km of 1.22± 0.08 mM, a Vmax of 98±0.15 µmol/min mg and a Kcat of 2.5× 103 s− 1 for p-coumaric acid. On the other hand, the purified VR enzyme showed a pH and temperature optimum of 6.0 and 25 °C, respectively. The enzyme was active in a pH range of 3.0–10.0, while its activity declined rapidly near 50 °C. The kinetic parameters were determined by adjusting the curve to the hyperbolic equation obtaining a Km of N3.37 ± 2.05 mM, a Vmax of 107.62 ± 50.38 µmol/min mg and a Kcat of 1.1 × 103 s− 1 using NADPH as cofactor. The high deviations observed for the kinetic parameters of this enzyme resulted from the erratic behavior of the method used to measure this activity when high concentrations of substrate were used (NADPH). The conditions of initial velocity were very far from the enzyme saturation; therefore the adjustment of the curve to a Michaelis behavior generated Km and Vmax values with high deviations. To reduce the standard error of the samples, it would be necessary to quantify these kinetic parameters using 4-vinylphenol as the enzyme substrate. 3.2.1. Influence of some enological factors on the CD and VR activities The effect of ethanol and pH 3.4 were assayed to evaluate how some factors of the wine have an influence on the stability of the purified enzymes. Likewise, it was determined if the presence of glucose in the reaction mixture could have an influence on the activity of both enzymes. In the case of CD activity was possible to determine that after a few minutes of incubation in the presence of 10% (v/v) and 12% (v/v) ethanol this activity decreases drastically. After 10 min of incubation the enzyme loses 50% of its activity, and after 1 h the activity is completely lost (Table 3). On the other hand, the CD activity is very stable when incubated at pH 3.4 where after 48 h of incubation only a 25% decay was observed (Table 4). The presence of glucose at a
Table 3 Effect of ethanol on the stability of the CD and VR activities Time (min)
1 3 5 10 30 60
Relative activitya (%)
Relative activitya (%)
Enzyme incubated with 10% ethanol
Enzyme incubated with 12% ethanol
Table 4 Effect of pH 3.4 on the stability of the CD and VR activities Relative activitya (%)
Time (h)
CD
VR
CD
VR
90 83 74 50 24 0
95 90 86 80 40 20
90 80 70 48 18 0
90 84 76 66 36 12
a Values are the means of three independent determinations and expressed as the percentage obtained by incubating the enzyme in the absence of ethanol (100%).
0.16 0.5 1 2 24 48 a
CD
VR
96 96 94 90 84 75
100 100 96 96 90 85
Values are the means of three independent determinations and expressed as the percentage obtained by incubating the enzyme in Bis–Tris pH 6.0 buffer (100%).
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L. Godoy et al. / International Journal of Food Microbiology 127 (2008) 6–11
concentration of 1% and 2% in the reaction mixture had no influence on the CD activity (data not shown). The stability of the VR enzyme decreases gradually in time in the presence of 10% and 12% ethanol (Table 3). After a 1 h incubation in the presence of 10% ethanol, the enzyme lost 80% of its activity, while at a concentration of 12% ethanol its enzymatic activity decreased by 88%. On the other hand, the enzymatic stability studied by incubating the protein at pH 3.4, showed that this factor did not cause changes in its stability and even after 48 h of incubation, only 15% of its initial activity was lost (Table 4). The presence of 1% or 2% glucose in the enzymatic reaction mixture had no influence on the VR activity (data not shown). 3.2.2. Substrate specificity of CD activity and assay of VR activity using a different cofactor In order to determine if the CD activity is capable of decarboxilating different hydroxycinamic acids, three hydroxycinamic acid substrates were evaluated: caffeic acid, ferulic acid and m-coumaric acid. They gave relative decarboxylase activities (versus p-coumaric acid) of 120% and 80% for caffeic acid and ferulic acid, respectively. In the case of m-coumaric acid, the enzyme did not show activity with this substrate. To carry out the reduction of 4-vinylphenol, it is necessary to have an oxidizable cofactor. Therefore, the use of NADH as co-factor was evaluated. As a result, a 25% decrease in the enzyme activity of VR was observed when compared to the use of NADPH as cofactor.
proposed by Huang et al. (1994) where an unsubstituted hydroxy group at a para- position to the unsaturated side chain is necessary for the decarboxylation of cinnamic acid. On the other hand, the purified VR enzyme presents an optimum pH of 6.0 and optimum temperature of 25 °C. The ethylphenol aroma in wine is mainly detected during the period of maturity in barrels, which is also the time in which the B. bruxellensis population is greatest. The CD and VR enzymes are intracellular, therefore when B. bruxellensis reaches the stationary growth phase it would lyse and release both enzymes into the wine. This would allow the enzymes to act directly on the cinnamic acids. We studied the effect of glucose, ethanol and the pH 3.4 on the activity and stability of the enzymes. In the presence of alcohol, the CD activity is more sensitive that the VR activity. Dias et al. (2003) observed that B. bruxellensis varies in its capacity to produce volatile phenols since a greater production of these has been observed when the alcohol concentration is lower. This could occur because growth of the microorganisms is somehow affected as well as the enzymatic activities that participate in the formation of ethylphenols which are sensitive to ethanol. Both enzymes are quite stable when incubated at pH 3.4. Furthermore, it was determined that the presence of glucose at a concentration of 1% and 2% (w/v) in the reaction media does not affect the enzymatic activities either. Knowledge of the enzymes that participate in the formation of ethylphenols in B. bruxellensis will allow a greater understanding of the management of some technological variables to control the production of these unwanted phenolic acids during wine production.
4. Discussion Acknowledgment Those microorganisms that ferment natural products such as grapes must have the capacity to metabolize anti-microbial compounds into other less toxic ones (Clausen et al., 1994; Degrassi et al., 1995; Cavin et al., 1997a,b; Edlin et al., 1998). B. bruxellensis initially transforms hydroxycinamic acids into vinyl derivates by a CD activity, and then produces ethylphenol through a VR activity. We observed that the CD activity was induced by the presence of p-coumaric acid as well as 4-vinylphenol in the culture media, as opposed to what was observed for the VR activity which was only induced by the presence of 4-vinylphenol. In P. fluorescens the ferulic decarboxylase acid was found to be constitutive (Huang et al., 1994), contrary to what was observed for a ferulic acid and a p-coumarate decarboxylase from B. pumillus (Degrassi et al., 1995) and a p-coumarate decarboxylase from L. plantarum (Cavin et al., 1997b) which were induced by substrates. In this work, the purification of both enzymes allowed their characterization, providing important information about how these enzymes would act on some wine components. In brief, the CD and VR enzymes have a molecular mass of 21 and 37 kDa, respectively. Edlin et al. (1998) reported a hydroxycinnamate decarboxylase from B. anomalus with a molecular mass of 21.8 KDa, which is very close to the value obtained in this study. A ferulic decarboxylase acid from P. fluorescens had a molecular mass of 20.4 kDa (Huang et al., 1994), while in B. pumillus and L. plantarum its molecular mass was 23 and 23.5 kDa, respectively (Degrassi et al., 1995; Cavin et al., 1997b). Recently, Dhar et al. (2007) purified a vanillic acid decarboxylase that gave a molecular weight of 23.0 kDa. The optimum activity of the CD enzyme of B. bruxellensis L-2480 was at pH 6.0 and at a temperature of 40 °C; similar conditions were described for the hydroxycinnamate decarboxylase enzyme from B. anomalus (Edlin et al., 1998). These values are similar to those obtained for the purified enzymes from B. pumillus, and L. plantarum (Degrassi et al., 1995; Cavin et al., 1997b). The specificity assays showed that this enzyme is capable of decarboxylating a series of hydroxycinnamic acids, with a greater specificity for caffeic acid, a cinnamic acid present in wine. Furthermore, it was possible to determine that the paraposition of the substituent with respect to the hydroxyl group is essential for this activity. These results agree with the mechanism
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