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Appl Environ Microbiol. 1998 April; 64(4): 1366–1371. | PMCID: PMC106156 |
Copyright © 1998, American Society for Microbiology Purification and Characterization of a Nylon-Degrading Enzyme Tetsuya Deguchi,1* Yoshihisa Kitaoka,1 Masaaki Kakezawa,1 and Tomoaki Nishida2 Environmental Technology Research Section, Chemical and Environmental Technology Laboratory, Kobe Steel, Ltd., Kobe 651-2271,1 and Department of Forest Resources Science, Faculty of Agriculture, Shizuoka University, Shizuoka 422,2 Japan Received August 20, 1997; Accepted January 25, 1998. A nylon-degrading enzyme found in the extracellular medium of a ligninolytic culture of the white rot fungus strain IZU-154 was purified by ion-exchange chromatography, gel filtration chromatography, and hydrophobic chromatography. The characteristics of the purified protein (i.e., molecular weight, absorption spectrum, and requirements for 2,6-dimethoxyphenol oxidation) were identical to those of manganese peroxidase, which was previously characterized as a key enzyme in the ligninolytic systems of many white rot fungi, and this result led us to conclude that nylon degradation is catalyzed by manganese peroxidase. However, the reaction mechanism for nylon degradation differed significantly from the reaction mechanism reported for manganese peroxidase. The nylon-degrading activity did not depend on exogenous H2O2 but nevertheless was inhibited by catalase, and superoxide dismutase inhibited the nylon-degrading activity strongly. These features are identical to those of the peroxidase-oxidase reaction catalyzed by horseradish peroxidase. In addition, α-hydroxy acids which are known to accelerate the manganese peroxidase reaction inhibited the nylon-degrading activity strongly. Degradation of nylon-6 fiber was also investigated. Drastic and regular erosion in the nylon surface was observed, suggesting that nylon is degraded to soluble oligomers and that nylon is degraded selectively. Nylon is a linear polymer containing the amide bond (—CONH—), which is also found in natural polymers, such as protein. However, nylon, with the exception of nylon-1, is believed to be resistant to attack by proteolytic enzymes, whereas protein is easily hydrolyzed by these enzymes. Recently, we reported that the white rot fungi strain IZU-154, Phanerochaete chrysosporium, and Trametes versicolor were able to degrade nylon-66 under ligninolytic conditions ( 5). Nuclear magnetic resonance (NMR) analysis of the degraded nylon revealed four end groups, —CHO, —NHCOH, —CH 3, and —CONH 2, that formed in degraded nylon, suggesting that nylon degradation was an oxidative process, not a hydrolytic process. White rot fungi are the best-known and most effective lignin-degrading microorganisms. Recently, these fungi have received worldwide attention because of their industrial use in biopulping ( 15), in biobleaching ( 7), in dye decolorization ( 26), and in detoxifying recalcitrant environmental pollutants, such as dioxins and chlorophenols ( 2, 14). The process of lignin degradation by these fungi is nonspecific and nonstereoselective, which explains why the fungi can mineralize lignin and various organic materials. Under ligninolytic conditions, many white rot fungi secrete extracellular enzymes. Among these enzymes are lignin peroxidase, manganese peroxidase (MnP), and laccase ( 21), which, together with an H 2O 2-generating system and cellulolytic and hemicellulolytic enzymes, may act synergistically during decay of wood. In this study, we purified and characterized the nylon-degrading enzyme produced by white rot fungus strain IZU-154. Interestingly, the protein purified and identified as the nylon-degrading enzyme is apparently MnP. However, the reaction system for nylon degradation differs significantly from the well-known MnP reaction system, especially with respect to the role of organic acid. Here we describe the enzymatic degradation of nylon and a new MnP reaction system. Organism. The white rot fungus strain IZU-154, which was isolated in our laboratory ( 20), was used in this study. IZU-154 has been deposited as strain NK-1148 under accession no. FERM BP-1859 in the National Institute of Bioscience and Human Technology of the Ministry of Industry and Technology, Ibaraki, Japan. Since secondary mycelia were observed and the sexual cycle was not observed in our previous study, we propose that IZU-154 belongs to the family Deuteromycotina. Chemicals. The nylon-66 membrane used in this study was purchased from Sartorius. Catalase and superoxide dismutase (SOD) were purchased from Sigma Chemical Co. (St. Louis, Mo.) and Wako Pure Chemical Industries (Osaka, Japan), respectively. Nylon-6 fiber was kindly supplied by Toray Industries, Inc. (Tokyo, Japan). Culture conditions. To prepare an inoculum, agar cubes cut from IZU-154-colonized potato dextrose agar plates were incubated in CSL-glc medium (8 g of corn steep liquor per liter, 10 g of glucose per liter; pH 4.5) for 3 days at 30°C with shaking. Then the culture was homogenized in the same amount of distilled water and used to inoculate nitrogen-limited medium (300-ml portions in 5,000-ml Erlenmeyer flasks) by using a 5% (vol/vol) inoculum. The nitrogen-limited medium contained (per liter) 10 g of glucose, 0.1 g of ammonium tartrate, 1 g of KH2PO4, 0.2 g of NaH2PO4, 0.5 g of MgSO4 · 7H2O, 0.1 mg of thiamine-HCl, 0.1 mg of CaCl2, 0.1 mg of FeSO4 · 7H2O, 0.01 mg of ZnSO4 · 7H2O, 0.02 mg of CuSO4 · 5H2O, and 48 mg of MnSO4 · 5H2O. The flasks were incubated without shaking at 30°C. Purification of nylon-degrading enzyme. Culture fluids were centrifuged at 1,500 × g for 30 min to remove mycelia, and the resulting supernatants were subjected to anion-exchange adsorption with a Q-Sepharose Fast Flow column (Pharmacia). Briefly, additional purification procedures involved anion-exchange chromatography on a Mono Q column (type HR 5/5; Pharmacia), gel permeation chromatography on a Superdex 75 column (type HiLoad 26/60; Pharmacia), and hydrophobic chromatography on a Phenyl Superose column (type HR 5/5; Pharmacia). Details of the procedures used are described below. Detection of nylon-degrading activity. Nylon-degrading activity was qualitatively detected by observing the structural disintegration of a nylon-66 membrane (Fig. ). Culture fluid was passed through a 0.45-μm-pore-size filter and was concentrated fivefold with an Ultrafree-PFL filter (10,000-molecular-weight cutoff; Millipore). Then 1 mg of nylon-66 membrane (diameter, 5 mm) and 5 μl of 200 mM MnSO4 (final concentration, 1 mM) were added to 1 ml of the concentrated culture fluid in a 5-ml glass vessel. The reaction was allowed to proceed at 30°C for 2 days under aerobic conditions. The activity in the column eluate was detected in a manner similar to the manner described above, except that the reaction mixture contained 20 mM sodium acetate (pH 4.5), 10 mM KH2PO4, 1 mM MnSO4, 1 mg of nylon-66 membrane, and 5 to 10 μl of eluted sample. The composition and assembly of the reaction mixture are described below. Electrophoresis. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and isoelectric focusing (IEF) were performed with polyacrylamide gradient gels (10 to 20% polyacrylamide; Daiichi) and a 5% polyacrylamide gel (ampholine pH range, 4 to 6; Pharmacia), respectively, as recommended by the manufacturers. Proteins were visualized by staining with Coomassie blue R-250. Enzymatic nylon degradation. Each reaction mixture (1 ml) typically contained 20 mM acetate, 10 mM KH2PO4, 1 mM MnSO4, 1 mg of nylon-66 membrane, and purified enzyme. The pH was adjusted to 4.5 with NaOH. The reactions were performed in 5-ml glass vessels at 30°C for 2 days. After incubation, the nylon membranes were dissolved in hexafluoroisopropanol (HFIP) and were subjected to a gel permeation chromatography to determine the molecular weight distribution. The pH profile of nylon-degrading activity was determined with the reaction mixture described above adjusted to pH 3.5, 4.0, 4.5, 5.0, and 5.5 with NaOH. When the effects of organic acids and phosphate on nylon degradation were investigated, 20 mM acetate and 10 mM KH2PO4 were not included in the reaction mixture. Peroxidase activity of purified enzyme. Peroxidase activity was assayed by using 2,6-dimethoxyphenol (2,6-DMP). The reactions were initiated by adding H 2O 2 to a final concentration of 0.1 mM and were performed at room temperature. Oxidation rates were determined by monitoring the increase in absorbance at 496 nm ( 31). The pH profile of peroxidase activity was determined after the pH was adjusted to 3.5, 4.0, 4.5, 5.0, and 5.5 with NaOH. One katal of peroxidase activity was defined as the amount of enzyme that formed 1 mol of the quinone dimer of 2,6-DMP per s at 30°C in a reaction mixture containing 50 mM sodium malate (pH 4.5), 0.5 mM MnSO 4, 1 mM 2,6-DMP, and 0.1 mM H 2O 2 ( 18). Determination of nylon molecular weight distribution. Nylon-66 membranes were washed with water, dried under a vacuum, dissolved in HFIP containing 10 mM trifluoroacetate, and subjected to gel permeation chromatography to determine changes in molecular weight distribution. An HFIP-80M column (Showa Denko), a mobile phase consisting of HFIP containing 10 mM trifluoroacetate and having a flow rate of 0.8 ml/min, and a refractive index detector were used. The weight average molecular weights (defined as Σ Ni Mi2/Σ Ni Mi, where Ni is the number of molecules and Mi is the molecular weight) and the number average molecular weights (defined as Σ Ni Mi/Σ Ni) were calculated based on results obtained with polymethylmethacrylate standards. NMR analysis. A nylon-66 membrane was incubated for 4 days at 30°C in a reaction mixture (0.5 ml) containing 20 mM sodium acetate (pH 4.5), 10 mM KH2PO4, 1 mM MnSO4, and purified enzyme. Then the nylon membrane was washed with water, dried under a vacuum, and dissolved in HFIP. The 13C NMR spectrum was determined with a Bruker model AC300P instrument in HFIP containing CDCl3 (3:1) at 300.13 MHz. Chemical shifts were given in the δ scale, and tetramethylsilane was used as the internal standard. Degradation of nylon fiber. Nylon-6 fiber was treated with purified enzyme to determine morphological variations. About 25 mm2 (30 mg) of nylon fiber was autoclaved for 15 min at 121°C with 10 ml of distilled water and then was placed in 1 ml of a reaction mixture containing 20 mM acetate (pH 4.5), 10 mM KH2PO4, 1 mM MnSO4, 0.1% Tween 80, and purified enzyme and incubated at 40°C. After incubation, the nylon fiber was washed with distilled water and then dried under a vacuum. Then the fiber was coated with Pt-Pd and was observed with a scanning electron microscope (model S-4000; Hitachi, Tokyo, Japan) at an acceleration voltage of 3 kV. Nylon degradation with an extracellular enzyme(s). In a static culture, IZU-154 grew as a mycelial mat on the surface of nitrogen-limited medium. Aliquots of culture fluid were removed daily, and nylon-degrading activity was monitored with a 2-day reaction after fivefold concentration. Nylon-degrading activity was observed in culture fluid obtained from days 4 to 8. When the concentrated active culture fluid was dialyzed against 20 mM acetate buffer (pH 4.5), nylon-degrading activity disappeared. The activity, however, was restored by redialysis against fresh nitrogen-limited medium, indicating that some components in the medium were necessary for nylon degradation. Table 1 shows the effects of the components on nylon degradation. Only KH 2PO 4 accelerated nylon degradation. The results of this experiment were used to determine the components of the reaction mixture used to assay for nylon-degrading activity in the column eluate; these components were 10 mM KH 2PO 4, 1 mM MnSO 4, and 20 mM acetate buffer (pH 4.5). We reported previously that nylon degradation by the fungus strain IZU-154 was significantly accelerated by adding manganese ( 5). In this study, manganese was found to induce production of nylon-degrading enzyme. The nylon-degrading activity was not apparent when the medium contained no MnSO 4 (data not shown). Thus, the role of manganese in this system is apparently identical to the role of manganese in the MnP system ( 10, 23). Purification of nylon-degrading enzyme. The cultures were harvested on day 6 and centrifuged at 1,500 × g for 30 min. About 1,100 ml of supernatant (7.6 mg of protein) was obtained from five Erlenmeyer flasks. To remove slime material, after the pH of the supernatant was adjusted to pH 6.5 with NaOH, the supernatant was loaded by using a peristaltic pump onto a Q-Sepharose Fast Flow column (Pharmacia) which was packed in a type HR 50/10 column (Pharmacia) and equilibrated with 20 mM phosphate buffer (pH 6.5). After extensive washing, absorbed protein was eluted with buffer A (200 mM NaCl in 20 mM sodium acetate, pH 4.5). The nylon-degrading activity was not detected in breakthrough fractions, and 72% of the loaded protein was eluted from the column with buffer A. The eluate was concentrated and dialyzed against buffer B (20 mM sodium acetate, pH 4.5) in Amicon ultrafiltration cells by using a type YM10 membrane filter. Further chromatography was performed by using fast protein liquid chromatography at 4°C, and elution was simultaneously monitored at 280 and 405 nm. In step 2, the concentrated eluate was applied to a Mono Q column (type HR 5/5; Pharmacia) equilibrated with buffer B. After the column was washed at flow rate of 1 ml/min, chromatography was continued with a 200-ml linear gradient of buffer A. Three peaks with absorbance at 405 nm were observed; these peaks corresponded to NaCl concentrations of ca. 0.02, 0.04, and 0.05 M. Numerous peaks were observed when absorbance at 280 nm was monitored. The nylon-degrading activity was detected in all three peak fractions, but not in the other fractions. Only the third peak fraction, which contained the largest amount of protein in the three active fractions, was concentrated by ultrafiltration with a Centricon 10 microconcentrator (Amicon) and used for the next step. Step 3 consisted of gel permeation chromatography on a Superdex 75 column (type HiLoad 26/60; Pharmacia). Buffer A at flow rate of 0.5 ml/min was used as the mobile phase. A single symmetric peak with absorbance at 405 nm appeared, and nylon-degrading activity was detected in the peak fractions. However, when the eluate was monitored at 280 nm, the peak was found to be not symmetric. The active fractions, therefore, were subjected to another type of chromatography. Further purification was accomplished with hydrophobic chromatography (step 4). The active fractions were dialyzed against 100 mM acetate buffer (pH 4.2) containing 1.5 M ammonium sulfate and were applied to a Phenyl Superose column (type HR 5/5; Pharmacia). The proteins were eluted with a nonlinear gradient of ammonium sulfate (5 ml of 1.5 to 0.5 M NH4SO4, 30 ml of 0.5 to 0.0 M NH4SO4) at a flow rate of 0.5 ml/min. A symmetrical peak with absorbance at both 280 and 405 nm was observed after hydrophobic chromatography on the Phenyl Superose column (Fig. ). This peak fraction had nylon-degrading activity. The purified protein appeared to be homogeneous when active fractions were analyzed by SDS-PAGE and IEF (Fig. ). These analyses showed that the molecular weight and pI were 43,000 and 3.7, respectively. In addition, the enzyme was dialyzed against 20 mM acetate buffer (pH 4.5), and the absorption spectrum was recorded with a Hitachi model U-3200 spectrophotometer. The absorption spectrum had a maximum at 406 nm and smaller peaks at 502 and 632 nm (Fig. ). The absorption maximum at 406 nm shifted to an absorption maximum at 420 nm when 0.1 mM (final concentration) H 2O 2 was added (data not shown). These features (absorption spectrum and response to H 2O 2) were apparently identical to those of peroxidase ( 6, 22, 29). Comparison between peroxidase activity and nylon-degrading activity. Table 2 shows the requirements and inhibitors for peroxidase activity of the purified enzyme. This activity is completely dependent on manganese and lactate in addition to exogenous H 2O 2. This finding and all of the features described above are apparently identical to the features reported for MnP ( 8, 16, 18). In short, the purified nylon-degrading enzyme was MnP. However, as shown in Table 3, the requirements and inhibitors for nylon degradation differed from the requirements and inhibitors for peroxidase activity significantly. Two obvious differences between the reactions are the role of lactate and sensitivity to SOD. Nylon-degrading activity was strongly inhibited by lactate and SOD, but peroxidase activity required lactate and was quite insensitive to SOD. Table 4 shows the effects of organic acids and phosphate on nylon degradation. The first five organic acids shown in Table 4 (lactate, malate, glycolate, citrate, and tartrate) are α-hydroxy acids, which are known to be essential for the MnP reaction. Nylon degradation was observed in the absence of these α-hydroxy acids but not in their presence. These results indicate that the two reactions cannot proceed simultaneously, even though both reactions are catalyzed by a single enzyme. The pH profiles for both 2,6-DMP oxidation and nylon degradation are shown in Fig. . The optimum pH for both reactions is around 4.5. NMR analysis of enzymatically degraded nylon-66 membrane. Three typical carbons resonating at δ 14, 166, and 210, which were assigned to —CH 3, —NHCHO, and —CHO, respectively, were observed in the 13C NMR spectrum of enzymatically degraded nylon-66. This spectrum was identical to that of nylon-66 degraded by fungus strain IZU-154 ( 5), indicating that nylon degradation by fungus strain IZU-154 was essentially identical to nylon degradation by the purified enzyme. In a previous report, we suggested that the formation of the end groups described above can be explained by a thermal oxidative degradation mechanism in which methylene groups adjacent to a nitrogen atom are attacked by oxygen and then nylon is degraded further by a chain reaction ( 4, 17, 25). The formation of —NHCHO and the formation of —CH 3 may be caused by cleavage of a C-C bond in CH 2-CH 2 adjacent to a nitrogen atom. The formation of —CHO may be caused by cleavage of a C-N bond in NH-CH 2, resulting in formation of —CONH 2. In the thermal oxidative degradation process, thermal treatment is thought to be especially important for initiation. Similarly, the enzyme may play a role as an initiator, mainly because the reactions after initiation could automatically proceed and it is improbable that one enzyme can subtly catalyze all of these reactions, especially the formation of the —CH 3 group. Degradation of nylon fiber. Figure shows the variations in surface properties after degradation of nylon fibers. After 1 day of incubation, the smooth surface of the fiber became rough, but there was no change in diameter (Fig. B), indicating that the surface of the fiber was stripped. Subsequently, many horizontal grooves were observed (Fig. C), and these grooves grew horizontally and became deeper (Fig. D). Major grooves formed at regular intervals. The variation in the surface appearance indicates that nylon was solubilized, suggesting that the molecular weight of the nylon was significantly reduced since even the nylon-6 hexamer is almost insoluble. The variation observed was apparently promoted by adding Tween 80, possibly due to solubilization of low-molecular-weight nylon. In this paper we describe for the first time purification of an enzyme that catalyzes nylon degradation. The enzyme eluted as a single peak after four purification steps which included three different types of chromatography (anion-exchange chromatography, gel permeation chromatography, and hydrophobic chromatography). The characteristics of the purified protein (molecular weight, absorption spectrum, and requirements for peroxidase activity) were identical to those of MnP, and this led to the conclusion that nylon degradation is catalyzed by MnP. However, the reaction system for nylon degradation differed significantly from the reaction system reported for MnP. One of the most obvious differences was the role of organic chelators, such as α-hydroxy acids (Table 4). MnP is known to have a manganese-binding site in which Mn(II) is hexacoordinated to the carboxylate oxygens of Glu-35, Glu-39, Asp-179, a heme propionate oxygen, and two water oxygens ( 1, 12, 13, 27, 28). MnP has been shown to have a normal peroxidase catalytic cycle ( 6, 19, 29, 30). Resting MnP is oxidized by H 2O 2 in a single two-electron step to form MnP compound I, and the latter is reduced by Mn(II) back to the resting enzyme in two single-electron steps, with intermediate formation of MnP compound II ( 28). In each reduction step, one equivalent of Mn(III) is formed. Since MnP was first discovered in cultures of white rot fungi, α-hydroxy acids have been considered some of the key components in the MnP reaction system ( 8, 9, 30). These organic acids chelate Mn(III) that is generated and thus both facilitate the release of Mn(III) from the enzyme-manganese complex and stabilize this species in aqueous solutions. Then the released Mn(III) chelator, in turn, oxidizes various substrates. The MnP activity, therefore, can substitute for a nonenzymatically prepared Mn(III) chelator ( 9, 24). However, nylon degradation is apparently inhibited by the α-hydroxy acids. This suggests that in nylon degradation Mn(III) does not act as the direct oxidizing agent. The inhibition by α-hydroxy acids may be related to the possibility that this inhibition facilitates the release of Mn(III) from the enzyme-manganese complex. Another difference between MnP activity and nylon-degrading activity has to do with the varieties of active oxygen involved in nylon degradation. Unlike 2,6-DMP oxidation, nylon degradation does not require exogenous H 2O 2, although it is inhibited by the addition of catalase (Tables 2 and 3). Furthermore, SOD inhibits only nylon-degrading activity. These results suggest that both H 2O 2 and the superoxide anion radical are involved in nylon degradation. Horseradish peroxidase is known to catalyze the peroxidase-oxidase reaction in addition to the peroxidase reaction ( 28). The peroxidase-oxidase reaction also does not require exogenous H 2O 2 and is inhibited by both catalase and SOD ( 3, 11). Yokota and Yamazaki proposed a mechanism for this reaction, in which a catalytic amount of H 2O 2 is necessary for initiation and the superoxide anion radical is an active intermediate in a chain reaction ( 32). This mechanism may also explain the roles of H 2O 2 and the superoxide anion radical in nylon degradation. We also tried to degrade nylon-6 fiber with the enzyme described here. The first step of degradation appears to be a stripping off of the surface (Fig. B). Subsequently, grooves grow horizontally and become deeper. The erosion obviously has a regularity (Fig. D). Since nylon is a crystalline polymer like cellulose, this regularity may be related to the nylon structure. The well-known MnP reaction system in which Mn(III) acts as the direct oxidizing agent is very efficient for oxidation of polymeric substrates, such as lignin, because Mn(III) is mobile in polymeric substrates which may be inaccessible to polymeric enzymes. In this paper we describe a new MnP reaction system which may perhaps be grouped with the peroxidase-oxidase reaction mechanism which has been reported to be one of the horseradish peroxidase-catalyzed reactions. Further work is needed to clarify the mechanism of this reaction system and its activity with substrates other than nylon.
TABLE 1 Effect of medium components on nylon  degradation | Additiona | Nylon disintegration |
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| Glucose | − | | Ammonium tartrate | − | | KH2PO4 | + | | MgSO4 | − | | CaCl2 | − | | Metal solutionb | − |
TABLE 2 Requirements of 2,6-DMP oxidation | Reaction mixturea | % of activity with complete mixture |
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| Complete mixture | 100 | | Complete mixture − enzyme | <1 | | Complete mixture − Mn(II) | <1 | | Complete mixture − lactate | <1 | | Complete mixture − H2O2 | <1 | | Complete mixture + catalase (650 U/ml) | 4 | | Complete mixture + SOD (330 U/ml) | 105 | | Complete mixture + NaN3 (1 mM) | <1 |
TABLE 3 Requirements of nylon degradation | Reaction mixturea | Mol wt of nylon
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| Weight average | Number average |
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| Complete mixture | 38,206 | 12,477 | | Complete mixture − enzyme | 89,845 | 49,423 | | Complete mixture − Mn(II) | 90,110 | 50,854 | | Complete mixture + lactate (50 mM) | 90,059 | 49,882 | | Complete mixture + catalase (650 U/ml) | 75,334 | 33,002 | | Complete mixture + SOD (330 U/ml) | 88,203 | 45,264 | | Complete mixture + NaN3 (1 mM) | 89,300 | 49,639 |
TABLE 4 Effects of organic acids and phosphate on nylon degradation | Compounda | Concn (mM) | Mol wt of nylon
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| Weight average | Number average |
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| α-Hydroxy acids |  Lactate | 50 | 88,394 | 49,590 | | Malate | 50 | 89,177 | 50,161 | | Glycolate | 50 | 89,895 | 49,709 | | Citrate | 50 | 90,931 | 50,068 | | Tartrate | 50 | 88,964 | 49,178 | | Other compounds | | Acetate | 50 | 48,410 | 16,781 | | 10 | 71,101 | 31,460 | | Succinate | 50 | 44,828 | 14,961 | | 10 | 51,513 | 20,545 | | Phosphate | 50 | 84,589 | 40,003 | | 10 | 39,300 | 12,768 |
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