Iwate Biotechnology Research Center, 22-174-4 Narita, Kitakami, Iwate 024-0003, Japan
Correspondence
Masaru Nagai
nagai{at}ibrc.or.jp
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ABSTRACT |
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INTRODUCTION |
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The mechanisms of mushroom browning have been investigated extensively in Agaricus bisporus (Burton, 1998; Espín et al., 1999
). Browning in this species is mainly due to DOPA and GDHB melanins (Jolivet et al., 1998
), and Tyr seems to play the most important role in their synthesis (Turner, 1974
). Burton (1988)
reported that epidermal tissues of A. bisporus had a greater activity of non-latent Tyr and a greater concentration of phenols than did the flesh. Previously, we also reported that Tyr activity of L. edodes fruit bodies increased during post-harvest storage and that gill browning increased with increasing Tyr activity (Kanda et al., 1996a
).
Laccases (Lcc; EC 1.10.3.2), catalyse the single-electron oxidation of phenols or aromatic amines to form different products via various pathways. Recently, Castro-Sowinski et al. (2002) reported that a strain of Sinorhizobium meliloti with an intracellular Lcc synthesized different types of melanin compared with strains without this Lcc. Ikeda et al. (2002)
also reported a correlation between melanin synthesis and intracellular Lcc in Cryptococcus neoformans. The melanins of Aspergillus conidia are formed from L-DOPA and DHN, and an Lcc gene of Aspergillus nidulans is specifically expressed in the conidia (Aramayo & Timberlake, 1990
). In addition, Clutterbuck (1972)
showed that yellow-spored mutants of Aspergillus nidulans are deficient in Lcc. Although there are many reports dealing with extracellular Lccs produced by white-rot basidiomycetes (Leonowicz et al., 2001
), there are few studies of the intracellular Lccs produced by these fungi (Burke & Cairney, 2002
; Schlosser et al., 1997
; Roy-Arcand & Archibald, 1991
). In A. bisporus, the biological significance of intracellular Lccs is considered to be very limited because of their low levels (Turner, 1974
), but their significance remains unclear.
In this paper, we continue in our attempt to clarify the relationship between gill browning and Lcc activity in L. edodes through the isolation and characterization of an Lcc from the mature gills and through the comparison of some properties of the purified enzyme with those of an extracellular Lcc (Lcc 1) purified previously from L. edodes (Nagai et al., 2002). To our knowledge, this is the first report of the purification of intracellular Lcc from a basidiomycete.
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METHODS |
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Organisms and culture conditions.
A commercial dikaryotic strain of L. edodes, strain H 600, was obtained from Hokken Sangyo and was used throughout this study. Mycelia were maintained on 1·5 % agar plates (diam. 90 mm) with 0·25x MYPG medium containing 0·25 % Bacto malt extract (Difco), 0·1 % Bacto yeast extract (Difco), 0·1 % tryptone peptone (Difco) and 0·5 % glucose.
For production of fruit bodies, mycelia were cultivated for 50 days in sawdust medium containing 3·7 kg sawdust, 1·3 kg Baideru (a nutrient supplement for mushroom production; Hokken Sangyo) and 7·6 l water according to the method of Matsumoto (1988). Fruit bodies were harvested immediately after the veil had broken.
Preparation of crude extract from fruit bodies.
Fruit bodies were separated into caps (pigmented rind and flesh), stipes and gills and frozen by liquid nitrogen. The frozen tissue was suspended in 10 times its volume of 10 mM sodium phosphate buffer (PB), pH 6·0, and homogenized using an Excel Auto Homogenizer (Nihon Seiki) at 10 000 r.p.m. for 1 min. The homogenate was centrifuged at 12 000 g for 20 min, after which the supernatant was collected as the crude extract.
Purification of Lcc from the browned gills.
All steps were carried out at 4 °C. Column chromatography was operated with a gradient controller (AC-5900 GradiCon III; ATTO) or with an FPLC system (Pharmacia).
A crude extract was prepared from 40 g sliced, fully browned gills. Powdered ammonium sulfate was then added to the extract to achieve 30 % saturation and the resulting precipitate was removed by centrifugation at 12 000 g for 20 min.
The supernatant was applied to a TOYOPEARL Butyl-650 M (Tosoh) column (25x80 mm) equilibrated with PB containing 30 % saturated ammonium sulfate. The column was washed with the same buffer and adsorbed proteins were eluted by a linear concentration gradient of ammonium sulfate (300 ml, 300 % saturation) in PB, at a flow rate of 2 ml min-1. The fractions containing Lcc activity were collected, dialysed against 20 mM sodium acetate buffer, pH 4·0, and applied to a TOYOPEARL CM-650 M (Tosoh) column (10x50 mm) equilibrated with 20 mM sodium acetate buffer, pH 4·0. After washing the column with the same buffer, the adsorbed proteins were eluted by a linear concentration gradient of NaCl (40 ml, 0500 mM) at a flow rate of 1 ml min-1. The Lcc active fractions were pooled and concentrated to about 250 µl by ultrafiltration using a Centricon-30 concentrator (30 kDa cut-off; Amicon). The concentrated enzyme solution was applied to a Superdex 75 HR 10/30 column (1x30 cm; Pharmacia) equilibrated with PB containing 100 mM NaCl. The enzyme was eluted with the same buffer at a flow rate of 250 µl min-1. Fractions exhibiting Lcc activity were pooled and dialysed against PB, and powdered ammonium sulfate was added to achieve 20 % saturation. The enzyme solution was then applied to a Phenyl Superose HR 5/5 column (5x50 mm, Pharmacia) equilibrated with PB containing 20 % saturated ammonium sulfate. After washing the column with the same buffer, the final elution was with a linear concentration gradient of ammonium sulfate (20 ml, 200 % saturation) in PB, at a flow rate of 500 µl min-1.
Enzyme assay.
To determine Lcc activity, 2,2'-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid)diammonium salt (ABTS) (Sigma) was used as the substrate. The reaction mixture for the standard assay contained 1 mM ABTS, McIlvaine buffer (pH 3·0) and the enzyme solution in a total volume of 100 µl. After incubation at 30 °C for 20 min, the reaction was stopped by adding 100 µl 5 % trichloroacetic acid. The formation of the cation radical was detected by measuring the absorbance increase at 420 nm (420=36 000 M-1 cm-1). One unit of Lcc activity was defined as the amount of enzyme that catalysed the oxidation of 1 µmol ABTS in 100 µl reaction mixture at 30 °C in 1 min. Tyr activity was measured at pH 6·0 using 1 mM catechol as the substrate. The formation of the cation radical was detected by measuring the absorbance increase at 450 nm (
450=2211 M-1 cm-1). One unit of Tyr activity was defined as the amount of enzyme that catalysed the oxidation of 1 µmol catechol in 100 µl reaction mixture at 30 °C in 1 min.
Protein assay.
Protein concentration was measured with BCA Protein Assay Reagent (Pierce) using BSA (Sigma) as the standard. During Lcc purification steps, protein concentration was monitored spectrophotometrically by A280.
Electrophoresis.
Native PAGE was performed according to the method of Davis (1965) using a 520 % polyacrylamide gradient gel (NPG-520L PAGEL; ATTO) and Premixed 10xTris/Glycine Buffer (Bio-Rad). SDS-PAGE was performed according to the method of Laemmli (1970)
using a 10 % polyacrylamide gel (NPU-10L PAGEL; ATTO) and Premixed 10xTris/Glycine/SDS Buffer (Bio-Rad). The samples were boiled in 2 % SDS and 5 % 2-mercaptoethanol for 10 min and then applied to the gel. The isoelectric point (pI) of the enzyme was measured in an isoelectric focusing gel between pH 3·5 and 9·5 (Ampholine PAG plate; Pharmacia) and Multiphore II system (Pharmacia) using an isoelectric focusing calibration kit, pH 3·59·3 (Pharmacia). Proteins were stained with Coomassie brilliant blue R 250 (PAGE Blue 83; Daiichi Chemicals). Activity staining was carried out by incubating the gel after native PAGE at room temperature in McIlvaine buffer (pH 3·0) with 1 mM ABTS.
Estimation of molecular mass.
The molecular mass of the enzyme was estimated by two methods: (1) gel filtration on a Superdex 75 HR 10/30 column with Gel filtration standards (Bio-Rad); and (2) SDS-PAGE as described above with Precision Protein Standards (Bio-Rad). The molecular mass of the enzyme was calculated from the mobility versus molecular mass plots of the marker proteins.
Determination of carbohydrates.
Carbohydrate molecules in the purified Lcc were determined by endoglycosidase treatment. The purified enzyme (1 µg) was boiled with 5 % 2-mercaptoethanol for 10 min, then incubated with 5 mU Endoglycosidase-H (Roche Diagnostics) in PB at 37 °C for 16 h. After this treatment, the molecular mass of the protein was calculated by SDS-PAGE.
Effect of pH and temperature on Lcc activity and stability.
The effect of pH on Lcc activity was examined at pH values from 1·0 to 6·0, using 0·1 M KCl/HCl buffer at pH values from 1·0 to 2·0, 0·1 M Glycine/HCl buffer at values from 2·0 to 4·0 and McIlvaine buffer at values from 4·0 to 6·0.
The effect of pH on enzyme stability was investigated by measurement of the activity remaining after incubation for 16 h at 30 °C in various buffers with 50 µg BSA ml-1. The buffers were 0·1 M KCl/HCl (pH 1·02·0), 0·1 M Glycine/HCl (2·04·0), McIlvaine buffer (4·06·0), 0·1 M sodium acetate (5·07·0), 0·1 M Tris/HCl (7·08·0) and CAPS (8·010·0). The effect of temperature on enzyme activity was determined at pH 3·0, with reactions performed by incubating at each temperature and pH 3·0 for 10 min. The thermal stability of Lcc was investigated by incubating preparations in PB with 50 µg BSA ml-1 for 30 min at various temperatures. After incubation, the activity remaining was determined.
Substrate specificity.
Spectrophotometric measurement of substrate oxidation by purified Lcc was carried out in a 100 µl reaction mixture containing the test substrates in McIlvaine buffer (pH 3·06·0). Activity against ABTS, p-phenylenediamine, 2,6-dimethoxyphenol, catechol, guaiacol, ferulic acid and L-DOPA was assayed at concentrations of between 0·1 and 1 mM. Activity against pyrogallol was assayed between 1 and 10 mM. All reactions were conducted at 30 °C for 10 min. The rate of substrate oxidation was determined by measuring the absorbance increase, with the molar extinction coefficient () obtained from the literature (Eggert et al., 1996
; Shin & Lee, 2000
). Michaelis constants (Km) were calculated from LineweaverBurk plots at the optimum pH in each case.
The oxidative products of L-DOPA and catechol were also analysed by HPLC. The analysis was carried out using a reverse phase HPLC cartridge and a Tsk gel ODS-8TM (150 mmx4·6 mm i.d.; Tosoh), radially compressed by a separations module (Waters 2960) at 25 °C. The mobile phases (flow rate 1 ml min-1) consisted of 5 ml PIC B8 (Waters 84283) in 1 l 0·05 % acetic acid for L-DOPA, and CH3CN and 5 % acetic acid (12 : 88) for catechol. Detection was performed between 220 and 400 nm with a photodiode array detector (Waters 996) connected to a Millennium Chromatography Manager (Waters).
In vitro gill browning experiments.
Gills (4 g) were cut from the fruit bodies of L. edodes, frozen and homogenized in an Excel Auto Homogenizer at 10 000 r.p.m. for 1 min with 10 ml McIlvaine buffer, pH 4·0. Tyr (0·1 U ml-1), Lcc 1 (0·4 U ml-1) or Lcc 2 (0·4 U ml-1) was added to 80 µl of the supernatant of the homogenate and the reaction mixture was incubated at 30 °C for 60 min. An absorbance increase at 460 nm, showing the synthesis of L-DOPA quinone, was measured at the end of the incubation period.
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RESULTS |
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Effect of pH and temperature
The pH profile for Lcc 2 activity against ABTS showed a single peak of maximum activity at pH 3·0 (Fig. 3a). When the effect of pH on enzyme stability was tested at 30 °C for 16 h, the enzyme was stable over a pH range of 4·07·0 (Fig. 3b
). The optimum temperature of Lcc 2, determined at pH 3·0, was 40 °C (Fig. 4
a). The thermal stability of Lcc 2 was determined by incubating the enzyme at pH 6·0 (Lcc 2 was stable at 30 °C for 16 h at this pH) for 30 min. No loss of activity was observed after incubation at 50 °C, but incubation at 60 °C resulted in a 33 % loss of Lcc activity (Fig. 4b
). Thus, we concluded that Lcc 2 is stable up to 50 °C.
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DISCUSSION |
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The purified intracellular Lcc, Lcc 2, was characterized and compared with an extracellular Lcc, Lcc 1, previously purified from the culture filtrate of L. edodes. The molecular mass of Lcc 2 (58 kDa) was lower than Lcc 1 (72·2 kDa) and other Lccs so far characterized from basidiomycetes (Thurston, 1994). The molecular masses of both Lccs after treatment with endoglycosidase-H were also determined. The molecular mass of deglycosylated Lcc 2 (53 kDa) was lower than that of Lcc 1 (55 kDa). The isoelectric point (pI) of Lcc 2 (pH 6·9) was higher than that of Lcc 1 (around pH 3·0). To our knowledge, the pI values of most Lccs secreted in culture media are acidic (Eggert et al., 1996
; Dedeyan et al., 2000
; Galhaup et al., 2002
; Saparrat et al., 2002
), and the pI of Lcc 2 is the highest of all Lccs reported previously, except for an Lcc from Coriolus hirsta (pH 7·4; Shin & Lee, 2000
). Thus, the neutral pI of Lcc 2 might be very specific. Although the N-terminal amino acid sequence of Lcc 2 was analysed by the methods described previously (Nagai et al., 2002
), the clear sequence was not detected because the N terminus was probably blocked. The facts that the molecular mass of the deglycosylated Lcc 2 was lower than that of Lcc 1 and that the ion charge of the surface of these proteins was different, suggest that the two Lccs are encoded individually.
When the effect of metal ions on enzyme activity was tested, 10 mM Cu2+ activated Lcc 1, but not Lcc 2 (Table 2; Nagai et al., 2002
). The absorption spectrum of Lcc 1 showed a peak absorption at around 610 nm, suggesting the existence of type-I copper ions. However, no secondary peak absorption at around 310 nm was observed to indicate a type-II copper signal. Thus, we speculated that the activation of Lcc 1 by Cu2+ may be due to the filling of type-II copper-binding sites with copper ions (Nagai et al., 2002
). In this study the absorption spectrum of Lcc 2 showed a peak for a type-II copper signal as well as the principle peak at 610 nm indicating a type-I copper signal.
When substrate specificities of Lcc 2 were tested, the optimum pH of Lcc 2 for each substrate was different (Table 3). Such differences have been observed by several authors and have been ascribed to variable degrees of substrate protonation under different pH conditions (D'Annibale et al., 1996
; Fukushima & Kirk, 1995
). The optimum pHs of Lcc 2 for many substrates were lower than those of Lcc 1. Schlosser et al. (1997)
have also reported that the optimum pH of the intracellular Lccs from Trametes versicolor is lower than that of the extracellular Lccs. In addition, they have reported that the substrate specificity of the Lcc contained in the culture filtrate was different from the Lcc contained in the crude extract of the mycelia.
The Km values of both enzymes for typical Lcc substrates ABTS, p-phenylenediamine, 2,6-dimethoxyphenol, ferulic acid or guaiacol were similar, but those of Lcc 2 for catechol and L-DOPA (1·72 and 1·22 mM, respectively), were noticeably lower than those of Lcc 1 (22·4 mM for catechol and zero oxidation for L-DOPA). Also, the activities (Kcat/Km values) of Lcc 2 against catechol and L-DOPA were significantly higher than those of Lcc 1. Because dihydroxyphenols are known to be oxidized by Tyr for synthesis of melanin in vivo (Bell & Wheeler, 1986), we analysed the activities of Tyr and Lcc 2 for oxidation of L-DOPA and catechol. As shown in Figs 5 and 6
, both enzymes oxidized L-DOPA and catechol to generate the same products. Both enzymes generated three products from catechol. It is established that catechol is oxidized by Tyr to o-benzoquinone and three types of semiquinone radical, and that these quinones are polymerized to catechol dimers in vivo (Bell & Wheeler, 1986
). The three peaks obtained by HPLC analysis in the present study probably correspond to these quinones or catechol dimers. Thus, Lcc 2 appears to oxidize L-DOPA and catechol to the same products as generated by Tyr.
Lcc 2 had oxidative activity for the phenolic compounds extracted from the gill of L. edodes fruit bodies (Fig. 7). The slight browning that occurred during incubation of gill extracts without enzymes may have been caused by native enzyme activity or by non-enzymic oxidation reactions. Although the addition of Lcc 1 to the extract did not effect browning, addition of Lcc 2 and Tyr did, suggesting that the phenolic compounds (perhaps especially dihydroxyphenols) in the gills of L. edodes fruit bodies can be oxidized by Lcc 2 but not by Lcc 1. Because the maximum absorbance change was obtained by the addition of Tyr, we speculated that Tyr can oxidize a wider variety of compounds than Lcc 2. However, increased oxidation following the addition of Lcc 2 to the Tyr reaction mixture, especially in the early stages, suggests either synergistic interactions between Tyr and Lcc 2 or perhaps the existence of phenolic compounds which can be oxidized by Lcc 2 but not Tyr. In this study, the reactions were carried out with a ratio of 4 Lcc 2 : 1 Tyr. The corresponding enzyme ratio in gill homogenates prepared from the fruit bodies of L. edodes preserved for 3 days was 22 : 1 (data not shown). Thus, we conclude that Lcc 2 has an important role in gill browning.
Jolivet et al. (1995) reported that the amount of phenols (tyrosine, GHB and GDHB) in the epidermal tissue was related to susceptibility to browning in A. bisporus. Kanda et al. (1996a)
showed a dependency of gill browning in L. edodes on de novo synthesis of a latent-type Tyr and increasing Tyr activity. Our results suggested that not only Tyr, but also an intracellular Lcc (Lcc 2) is produced during post-harvesting preservation and functions in the oxidation of phenolic compounds to turn gills brown. Considering the fact that purified Lcc 2 oxidized both L-DOPA and catechol, Lcc 2 might have a strong activity against dihydroxyphenols. Investigations into the phenol content of the gills of L. edodes fruit bodies are currently under way. We are also analysing the Lcc 2 gene with a view to its repression or disruption to construct a strain which can be preserved for longer periods without gill browning.
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Received 10 April 2003;
revised 13 June 2003;
accepted 13 June 2003.