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Appl Environ Microbiol, April 1998, p. 1366-1371, Vol. 64, No. 4
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
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 20 August 1997/Accepted 25 January 1998
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ABSTRACT |
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, 
-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.
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INTRODUCTION |
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, ---CH3, and
---CONH2, 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 H2O2-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.
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MATERIALS AND METHODS |
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. 1). 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.
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FIG. 1.
Morphological disintegration of nylon-66 membrane. (A)
Nylon-66 membrane (1 mg) and 5 µl of 200 mM MnSO4 were
added to 1 ml of fivefold-concentrated culture fluid on day 6. The
preparation was incubated for 2 days at 30°C. (B) Results obtained
when the concentrated culture fluid was used after it was boiled for 5 min.
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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 H2O2 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
MnSO4, 1 mM 2,6-DMP, and 0.1 mM
H2O2 (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 
Ni
Mi2/ Ni
Mi, where Ni is the number of
molecules and Mi is the molecular weight) and
the number average molecular weights (defined as Ni Mi/ 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 
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.
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RESULTS |
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 NH
4SO
4, 30 ml of 0.5 to 0.0 M
NH
4SO
4)
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.
2). This peak fraction had
nylon-degrading activity.
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FIG. 2.
Hydrophobic chromatography of a partially purified
nylon-degrading enzyme preparation from 6-day-old cultures of IZU-154.
Dotted line, absorbance at 270 nm; solid line, absorbance at 405 nm;
dashed line, (NH4)2SO4 gradient.
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The purified protein appeared to be homogeneous when active fractions
were analyzed by SDS-PAGE and IEF (Fig.
3). 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.
4).
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).
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FIG. 3.
Electrophoretic analysis of purified enzyme. (A)
SDS-PAGE analysis. Lane M contained molecular weight markers. (B) IEF
gel electrophoresis analysis. The arrows indicate the pI values of
markers.
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FIG. 4.
Absorption spectrum of purified enzyme. Protein was
dissolved in 20 mM sodium acetate (pH 4.5), and the spectrum was
determined at room temperature.
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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 H2O2. 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 -hydroxy acids, which are known to be essential for the MnP
reaction. Nylon degradation was observed in the absence of these
-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.
5. The optimum pH for both
reactions is
around 4.5.
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FIG. 5.
pH profiles of peroxidase activity and nylon-degrading
activity. Symbols:  , 2,6-DMP oxidation;  , weight average
molecular weight of nylon-66 membrane. Peroxidase activity was assayed
by using 2,6-DMP in a reaction mixture containing 20 mM acetate and 50 mM lactate.
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NMR analysis of enzymatically degraded nylon-66 membrane.
Three typical carbons resonating at 14, 166, and 210, which were
assigned to ---CH3, ---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 ---CH3 may be
caused by cleavage of a C-C bond in CH2-CH2
adjacent to a nitrogen atom. The formation of ---CHO may be caused by
cleavage of a C-N bond in NH-CH2, resulting in formation of
---CONH2. 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 ---CH3 group.
Degradation of nylon fiber.
Figure
6 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. 6B), indicating that the surface of the
fiber was stripped. Subsequently, many horizontal grooves were observed (Fig. 6C), and these grooves grew horizontally and became deeper (Fig.
6D). 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.
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FIG. 6.
Scanning electron micrographs of nylon-6 fiber. (A)
Unincubated control fiber. (B) Fiber after incubation for 1 day with
enzyme. (C) Fiber after 2 days of incubation with enzyme. (D) Fiber
after 4 days of incubation with enzyme.
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DISCUSSION |
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 -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 H2O2 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, -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 -hydroxy acids.
This suggests that in nylon degradation Mn(III) does not act as the
direct oxidizing agent. The inhibition by -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 H2O2, 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 H2O2 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 H2O2 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 H2O2 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
H2O2 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. 6B). Subsequently, grooves grow horizontally and become
deeper. The erosion obviously has a regularity (Fig. 6D). 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.
| |
ACKNOWLEDGMENTS |
We thank Kenneth Zahn of the Research Institute of Innovative
Technology for the Earth for helpful suggestions. We also thank H. Yasuda for the NMR analysis and H. Yoshida of Kobelco Research Institute, Inc., for scanning electron microscope observations.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Environmental
Technology Research Section, Chemical and Environmental Technology
Laboratory, Kobe Steel, Ltd., Takatsukadai l-chome, Nishi-ku, Kobe
651-22, Japan. Phone: 81-78-992-5733. Fax: 81-78-992-5547. E-mail:
te-deguchi{at}rd.kcrl.kobelco.co.jp.
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REFERENCES |
| 1.
|
Banci, L.,
I. Bertini,
T. Bini,
M. Tien, and P. Turano.
1993.
Binding of horseradish, lignin, and manganese peroxidases to their respective substrates.
Biochemistry
32:5825-5831[Medline].
|
| 2.
|
Bumpus, J. A.,
M. Tien,
D. Wright, and S. D. Aust.
1985.
Oxidation of present environmental pollutants by a white rot fungus.
Science
227:1434-1436.
|
| 3.
|
Chance, B.
1952.
Oxidase and peroxidase reactions in the presence of dihydroxymaleic acid.
J. Biol. Chem.
197:577-589[Free Full Text].
|
| 4.
|
Chiba, K., et al.
1988.
, p. 112.
Polyamide handbook
Nikkan Kogyo Shinbunsha, Tokyo, Japan. (In Japanese.)
|
| 5.
|
Deguchi, T.,
M. Kakezawa, and T. Nishida.
1997.
Nylon degradation by lignin-degrading fungi.
Appl. Environ. Microbiol.
63:329-331[Abstract].
|
| 6.
|
Farhangranzi, Z. S.,
B. R. Copeland,
T. Nakayama,
T. Amachi,
I. Yamazaki, and L. S. Powers.
1994.
Oxidation-reduction properties of compounds I and II of Arthromyces ramosus peroxidase.
Biochemistry
33:5647-5652[Medline].
|
| 7.
|
Fujita, K.,
R. Kondo,
K. Sakai,
Y. Kashino,
T. Nishida, and Y. Takahara.
1993.
Biobleaching of softwood kraft pulp with white rot fungus IZU-154.
TAPPI (Tech. Assoc. Pulp Pap. Ind.) J.
76:81-84.
|
| 8.
|
Glenn, J. K., and M. H. Gold.
1985.
Purification and characterization of an extracellular Mn(II)-dependent peroxidase from the lignin-degrading basidiomycete Phanerochaete chrysosporium.
Arch. Biochem. Biophys.
242:329-341[Medline].
|
| 9.
|
Glenn, J. K.,
L. Akileswaran, and M. H. Gold.
1986.
Mn(II) oxidation is the principal function of the extracellular Mn-peroxidase from Phanerochaete chrysosporium.
Arch. Biochem. Biophys.
251:688-696[Medline].
|
| 10.
|
Gold, M. H., and M. Alic.
1993.
Molecular biology of the lignin-degrading basidiomycete Phanerochaete chrysosporium.
Microbiol. Rev.
57:605-622[Abstract/Free Full Text].
|
| 11.
|
Halliwell, B., and J. D. Rycker.
1978.
Superoxide and peroxide-catalyzed reactions. Oxidation of dihydroxyfumarate, NADH and dithiothreitol by horseradish peroxidase.
Photochem. Photobiol.
27:751-761.
|
| 12.
|
Harris, R. Z.,
H. Wariishi,
M. H. Gold, and P. R. Ortiz de Montellano.
1991.
The catalytic site of manganese peroxidase.
J. Biol. Chem.
266:8751-8758[Abstract/Free Full Text].
|
| 13.
|
Johnson, F.,
G. H. Loew, and P. Du.
1994.
Homology models of two isozymes of manganese peroxidase: prediction of a Mn(II) binding site.
Proteins
20:312-319[Medline].
|
| 14.
|
Joshi, D. K., and M. H. Gold.
1993.
Degradation of 2,4,5-trichlorophenol by lignin-degrading basidiomycete Phanerochaete chrysosporium.
Appl. Environ. Microbiol.
59:1779-1785[Abstract/Free Full Text].
|
| 15.
|
Kashino, Y.,
T. Nishida,
Y. Takahara,
K. Fujita,
R. Kondo, and K. Sakai.
1993.
Biomechanical pulping using white rot fungus IZU-154.
TAPPI (Tech. Assoc. Pulp Pap. Ind.), J.
76:167-171.
|
| 16.
|
Kirk, T. K., and R. L. Farrell.
1987.
Enzymatic "combustion": the microbial degradation of lignin.
Annu. Rev. Microbiol.
41:465-505[Medline].
|
| 17.
|
Levantovskaya, I.,
B. M. Kovarskaya,
G. V. Dralyuk, and M. B. Neiman.
1965.
Mechanism of thermal oxidative degradation of polyamides.
Polymer Sci. USSR
6:2089-2095.
|
| 18.
|
Matsubara, M.,
J. Suzuki,
T. Deguchi,
M. Miura, and Y. Kitaoka.
1996.
Characterization of manganese peroxidase from the hyperlignolytic fungus IZU-154.
Appl. Environ. Microbiol.
62:4066-4072[Abstract].
|
| 19.
|
Mino, Y.,
H. Wariishi,
N. J. Blackburn,
T. M. Loehr, and M. H. Gold.
1988.
Spectral characterization of manganese peroxidase, an extracellular heme enzyme from the lignin-degrading basidiomycete, Phanerochaete chrysosporium.
J. Biol. Chem.
263:7029-7036[Abstract/Free Full Text].
|
| 20.
|
Nishida, T.,
Y. Kashino,
A. Mimura, and Y. Takahara.
1988.
Lignin biodegradation by wood-rotting fungi. I. Screening of lignin-degrading fungi.
Mokuzai Gakkaishi
34:530-536.
|
| 21.
|
Orth, A. B.,
D. J. Royse, and M. Tien.
1993.
Ubiquity of lignin-degrading peroxidases among various wood-degrading fungi.
Appl. Environ. Microbiol.
59:4017-4023[Abstract/Free Full Text].
|
| 22.
|
Pasczynski, A. V.,
B. Huynh, and R. Crawford.
1986.
Comparison of ligninase-I and peroxidase-M2 from the white-rot fungus Phanerochaete chrysosporium.
Arch. Biochem. Biophys.
244:750-765[Medline].
|
| 23.
|
Perie, F. H., and M. H. Gold.
1991.
Manganese regulation of manganese peroxidase expression and lignin degradation by the white rot fungus Dichomitus squalens.
Appl. Environ. Microbiol.
57:2240-2245[Abstract/Free Full Text].
|
| 24.
|
Popp, J. L., and T. K. Kirk.
1991.
Oxidation of methoxybenzenes by manganese peroxidase and by Mn3+.
Arch. Biochem. Biophys.
278:145-148.
|
| 25.
|
Sharkey, W. H., and W. E. Mochel.
1959.
Mechanism of the photooxidation of amides.
J. Am. Chem. Soc.
81:3000-3005.
|
| 26.
|
Spadaro, J. T.,
M. H. Gold, and V. Renganathan.
1992.
Degradation of azo dyes by lignin-degrading fungus Phanerochaete chrysosporium.
Appl. Environ. Microbiol.
58:2397-2401[Abstract/Free Full Text].
|
| 27.
|
Sundaramoorthy, M.,
K. Kishi,
M. H. Gold, and T. L. Poulos.
1994.
The crystal structure of manganese peroxidase from Phanerochaete chrysosporium at 2.06-Å resolution.
J. Biol. Chem.
269:32759-32767[Abstract/Free Full Text].
|
| 28.
|
Swedin, B., and H. Theorell.
1940.
Dioximaleic acid oxidase action of peroxidases.
Nature
145:71-72.
|
| 29.
|
Wariishi, H.,
L. Akileswaran, and M. H. Gold.
1988.
Manganese peroxidase from the basidiomycete Phanerochaete chrysosporium: special characterization of the oxidized states and the catalytic cycle.
Biochemistry
27:5365-5370[Medline].
|
| 30.
|
Wariishi, H.,
H. B. Dunford,
I. D. MacDonald, and M. H. Gold.
1989.
Manganese peroxidase from the lignin-degrading basidiomycete Phanerochaete chrysosporium.
J. Biol. Chem.
264:3335-3340[Abstract/Free Full Text].
|
| 31.
|
Wariishi, H.,
K. Valli, and M. H. Gold.
1992.
Manganese(II) oxidation by manganese peroxidase from the basidiomycete Phanerochaete chrysosporium.
J. Biol. Chem.
267:23688-23695[Abstract/Free Full Text].
|
| 32.
|
Yokota, K., and I. Yamazaki.
1965.
Reaction of peroxidase with reduced nicotinamide-adenine dinucleotide and reduced nicotinamide-adenine dinucleotide phosphate.
Biochim. Biophys. Acta
105:301-312[Medline].
|
Appl Environ Microbiol, April 1998, p. 1366-1371, Vol. 64, No. 4
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
