activation of lignin peroxidase in organic media by reversed micelles
TRANSCRIPT
Activation of Lignin Peroxidase in OrganicMedia by Reversed Micelles
Masayuki Kimura,1 Junji Michizoe,1 Shin-ya Oakazaki,1 Shintaro Furusaki,1
Masahiro Goto,1,2 Hiroo Tanaka,3 HiroyukiWariishi 3
1Department of Chemical Systems and Engineering, Graduate School ofEngineering, Kyushu University, 6-10-1, Hakozaki, Fukuoka 812-8581, Japan;telephone/fax:+81-92-642-3575; e-mail:[email protected] of Chemical Systems and Engineering, Kyushu University, JST,PRESTO, Fukuoka 812-8581, Japan3Department of Bioresources and Bioenvironmental Sciences, KyushuUniversity, Fukuoka 812-8581, Japan
Received 19 February 2004; accepted 16 July 2004
Published online 30 September 2004 in Wiley InterScience (www.interscience.wiley.com). DOI: 10.1002/bit.20277
Abstract: Activation of lignin peroxidase (LIP) in an organicsolvent by reversed micelles was investigated. Bis(2-ethylhexyl)sulfosuccinate sodium salt (AOT) was used asa surfactant to form a reversed micelle. Lyophilized LIPfrom an optimized aqueous solution exhibited no enzymaticactivity in any organic solvents examined in this study;however, LIP was catalytically active by being entrapped inthe AOT reversed micellar solution. LIP activity in thereversed micelle was enhanced by optimizing either thepreparation or the operation conditions, such as watercontent and pH in water pools of the reversed micelle andthe reaction temperature. Stable activity was obtained inisooctane because of the stability of the reversed micelle.The optimal pH was 5 in the reversed micellar system,which shifted from pH 3 in the aqueous solution. Thedegradation reaction of several environmental pollutantswas attempted using LIP hosted in the AOT reversedmicelle. Degradation achieved after a 1-h reaction reached81%, 50%, and 22% for p-nonylphenol, bisphenol A, and2,4-dichlorophenol, respectively. This is the first report onthe utilization of LIP in organic media. B 2004 WileyPeriodicals, Inc.
Keywords: lignin peroxidase; reversed micellar system;organic solvent; biodegradation
INTRODUCTION
Lignin is the most abundant renewable aromatic polymer
and is known as one of the most recalcitrant biomaterials
on Earth (Crawford, 1981; Sarkanen and Ludwig, 1971).
Its degradation plays a key role in the carbon cycle of
the biosphere (Crawford, 1981; Gold et al., 1989; Kirk
and Farrell, 1987). Only white-rot basidiomycetes are re-
sponsible for the complete mineralization of this polymer.
Phanerochaete chrysosporium, the best-studied white-rot
fungus, secretes two heme peroxidases, lignin peroxidase
(LIP) and manganese peroxidase, under ligninolytic condi-
tions (Gold et al., 1989; Kirk and Farrell, 1987; Tien, 1987).
Thus, these enzymes are believed to be involved in trig-
gering lignin biodegradation.
Nucleotide sequences of a number of LIP cDNAs and
genomic clones (Gold and Alic, 1993; Tien and Tu, 1987)
and crystal structures of LIP (Edwards et al., 1993; Piontek
et al., 1993; Poulos et al., 1993) have demonstrated that
important peroxidase catalytic residues are all conserved.
Therefore, LIP shares many structural and mechanistic
features with other peroxidases, yet it has several unique
properties (Dunford and Stillman, 1976; Gold et al., 1989).
The enzyme catalyzes the one-electron oxidation of non-
phenolic aromatic compounds with high redox potentials
via the formation of a substrate cation radical (Gold et al.,
1989; Kirk and Farrell, 1987; Tien, 1987). It has also been
reported that LIP oxidizes a wide range of environmentally
persistent aromatic compounds, such as polyaromatic hy-
drocarbons and polychlorinated dibenzo-p-dioxins as well
as lignin (Bumpus et al., 1985; Gold et al., 1993). Never-
theless, these pollutants are hydrophobic; the enzymatic
treatment has been attempted in aqueous media with a
limited concentration of pollutants, because the enzymes
including LIP are inactivated in organic media. If LIP is
utilized in organic media, the enzyme system could be ap-
plied to concentrated pollutant solutions. Therefore, the ac-
tivation of LIP in organic media is of great interest.
In this study, the activation of LIP in organic solvents was
investigated by using a reversed micellar solution. Reversed
micelles are a thermodynamically stable molecular assem-
bly, which are formed by surfactant molecules in a hy-
drophobic organic solvent (Shield et al., 1986). Enzymes
entrapped in reversed micelles have recently been reported
to be active and stable (Ly et al., 1998; Okazaki et al., 2002;
Ono and Goto, 1997). However, there has been no report
describing the application of the reversed micellar system to
B 2004 Wiley Periodicals, Inc.
Correspondence to: M. Goto
Contract grant sponsors: Ministry of Education, Science, Sports and
Culture of Japan; Kyushu University; New Energy and Industrial Tech-
nology Development Organization (NEDO) of Japan
Contract grant number: B 12450322
biodegradation reactions to date. In the present study,
important parameters affecting the efficiency of the LIP
reaction were examined in detail, showing the optimal
conditions for activating LIP in organic solvents. Finally,
the oxidation of p-nonylphenol, bisphenol A, and 2,4-
dichlorophenol by LIP in organic solvents was performed.
MATERIALS AND METHODS
Chemicals
Bis(2-ethylhexyl)sulfosuccinate sodium salt (AOT) was
purchased from Kishida Chemical Ltd. (Osaka, Japan).
Veratryl alcohol was obtained from Aldrich (St. Louis,
MO) and purified by vacuum distillation before use. p-
Nonylphenol, bisphenol A, and 2,4-dichlorophenol were
purchased from Tokyo Chemical Industry Co., Ltd. (Tokyo,
Japan). All organic solvents used in this work were of
analytical grade. Sequentially distilled and deionized water
was employed throughout the experiments.
Lignin Peroxidase
LIP (isozyme 2) was isolated from the extracellular culture
medium of P. chrysosporium (ATCC 34541) and purified as
previously described (Johjima et al., 1999; Wariishi et al.,
1990). The concentration of LIP was determined by the
absorbance at 408 nm by using the extinction coefficient of
133 mM�1 cm�1 (Gold et al., 1989). The enzyme was
electrophoretically homogenous and has an RZ (A408/A280)
value >4.6. LIP activity in the aqueous solution was es-
timated from veratryl alcohol (VA) oxidation activity as
previously described (Gold et al., 1989; Kirk and Far-
rell, 1987).
Preparation of Reversed Micelles Containing LIP
Reversed micelles containing LIP were prepared by direct
injection of an aliquot of the LIP stock solution in 20 mM
potassium phosphate buffer, pH 2–8, into 100 mM AOT in
isooctane. The concentration of the LIP stock solution
varied in order to adjust the water content (Wo; [H2O]/
[Surfactant]) to 10–70 in the reversed micelles but the
final concentration of LIP was constant at 3 AM or other-
wise indicated.
To optimize the preparation conditions, the effect of the
concentration of AOT and the LIP final concentration on the
activity in isooctane were also examined as follows. AOT
concentration was varied from 50 to 400 mM at the Wo
value of 50, and the final LIP concentration of 3 AM. The
final concentration of LIP in the reversed micelles was also
varied from 0.3 to 5 AM at the Wo value of 50 and AOT
concentrations at either 100 or 400 mM. All the concen-
trations are based on the total volume of the reversed
micellar solution.
Reaction Conditions for LIP encapsuledin Reversed Micelles
To optimize either preparation or operation conditions, VA
was utilized as the reducing substrate. VA (100 mM toluene
stock solution) was added to the reaction system at the final
concentration of 1.0 mM. H2O2 reversed micellar solution
prepared separately in AOT/isooctane was added to initiate
reactions at the final H2O2 concentration of 0.01–1.0 mM.
The reactions were conducted at 20–50jC and monitored
by the formation of veratraldehyde using e310 = 9,470 M�1
cm�1 with a JASCO V-570 UV spectrophotometer (JASCO,
Tokyo, Japan). The extinction coefficient was separately
obtained in the present study. The enzyme reactions were
carried out at least three times under the same experimental
conditions, and the initial rates were plotted using the
average value.
Oxidation of Aromatic Pollutants by LIPin Organic Media
The oxidations of p-nonylphenol, bisphenol A, and 2,4-
dichlorophenol were employed in the reversed micellar
solution. The decrease of substrates was monitored by a
HPLC equipped with an ODS-3 column (GL Science,
Tokyo, Japan) eluted with 30% acetonitrile in water (0.1%
phosphoric acid) (0–10 min), followed by a linear gradient
from 30% to 90% acetonitrile (10–20 min) and then by
90% acetonitrile (20–25 min) at a flow rate of 1.0 mL/min
using a UV detector at 268 nm (p-nonylphenol), 276 nm
(bisphenol A), and 294 nm (2,4-dichlorophenol), respec-
tively. The experiments were repeated at least three times,
and we evaluated the degree of degradation for the
pollutants as the average value.
RESULTS AND DISCUSSION
Although it has been reported that LIP can oxidize aromatic
substrates in aqueous solutions containing water-miscible
organic solvents (Yoshida et al., 1997), there is no
successful report on LIP-catalyzed reactions in organic
media, especially hydrophobic solvents. Recently, laccase
and manganese peroxidase, well-characterized fungal lig-
ninolytic enzymes, were reported to be active in organic
solvents when coated with surfactant molecules (Okazaki
et al., 2000, 2002). The lyophilized enzymes covered with
surfactant molecules were soluble and active in various or-
ganic solvents (Kamiya et al., 2000; Maruyama et al., 2002,
2003; Okazaki et al., 2000, 2002). We then applied this
surfactant-coating method to LIP as previously reported;
however, surfactant–LIP complex showed no activity in
organic solvents (data not shown). Because the substrate-
binding site of LIP is known to exist on the surface of
the protein, which is one of very unique properties of this
enzyme (Doyle et al., 1998; Johjima et al., 1999; Wariishi
et al., 1994), the interaction of surfactant molecules and LIP
substrate oxidation site(s) may interfere with the catalytic
496 BIOTECHNOLOGY AND BIOENGINEERING, VOL. 88, NO. 4, NOVEMBER 20, 2004
action. Thus another technique was required to activate LIP
in organic media. A reversed micellar system seemed to be
advantageous because enzyme molecules are entrapped in
water pools of the reversed micelles. We can readily control
the size of reverse micelles (3–10 nm) by changing the
water content in the reversed micellar solution, which is the
same meaning as changing the Wo values. The longest
dimension of LIP is around 4 nm, therefore, the size of
reversed micelles is large enough for entrapping LIP.
Activation of LIP in Organic Media
VA oxidation by LIP hosted in the reversed micelles and
lyophilized LIP was performed, clearly showing that LIP in
the reversed micelles was active, while no activity was
observed with lyophilized LIP in isooctane (Fig. 1). LIP was
thought to be entrapped in water pools inside the reversed
micelles; therefore, the enzyme maintained a high activity
in isooctane. This is the first report indicating that LIP
catalyzes a reaction in organic media.
Effect of pH During Reversed Micelle Preparation
One of the most important factors affecting the enzymatic
activity is the pH in the reaction medium. The pH values of
the reaction mixtures in an aqueous system and the stock
solutions for reversed micelle preparation were varied. The
latter value was thought to be the pH of water pools in the
reversed micelles. The initial reaction rates were then
plotted against pH values (Fig. 2). The optimal pH was
observed at 3 for the aqueous system as previously reported
(Gold et al., 1989; Kirk and Farrell, 1987; Tien, 1987). On
the other hand, in the reversed micellar system, the enzyme
activity became maximal at around pH 5. The shift of
optimal pH might be explained by the negative charge of the
surfactant. AOT has an anionic sulfo group at the hydro-
philic head; therefore, the microenvironment for enzyme
surface becomes acidic due to the condensation of protons at
the oil–water interface (Barbaric and Luisi, 1981; Douzou
et al., 1979; Michizoe et al., 2001, 2003; Shield et al., 1986).
As a result, in the reversed micellar system, the optimal pH
shifts toward a more alkaline value than that of LIP in the
aqueous solution. The water content in the reversed micelle
(Wo value) did not seem to affect the optimal pH.
Effect of Water Content and Sizeof Reversed Micelles
As described above, the interaction of the protein with
surfactant molecules should be considered to effectively
activate LIP in organic media. The Wo value defined by
[H2O]/[Surfactant] was used as the indicator of water
content in the reversed micelles. The Wo value has also been
known to correlate linearly with the size of the reversed
micelles (Pileni et al., 1985). The effect of the Wo value on
the initial rates of VA oxidation is shown (Fig. 3). The
enzymatic activity increased as the Wo value was increased,
and the activity attained a maximum at the Wo value of 50.
It is well known that the relationship between the Wo value
and enzymatic activity shows a bell-shaped tendency. An
entrapped enzyme often exhibits the highest activity when
Figure 1. LIP reaction in isooctane. Reactions were conducted at 25jC.
The reversed micelle was prepared by 100 mM AOT in isooctane and
3 AM LIP in 20 mM phosphate buffer, pH 3.0. Lyophilized LIP was pre-
pared from LIP in 20 mM succinate buffer (pH 3.0) and used in the
absence of AOT. The reaction was initiated by adding 0.1 mM H2O2.
Figure 2. pH dependency of LIP activity in aqueous system (o) and in
reversed micelle with Wo value of 50 (5) and 20 (D). The reactions were
conducted at 30jC.
Figure 3. Effect of Wo value on catalytic property of LIP in re-
versed micelle.
KIMURA ET AL.: ACTIVATION OF LIGNIN PEROXIDASE BY REVERSED MICELLES 497
the size of water pools coincides with that of an enzyme
(Khmelnitsky et al., 1989). The optimal Wo value for LIP
was substantially higher than the smaller protein a-
chymotrypsin and near that of the larger protein (laccase)
(Michizoe et al., 2001; Shield et al., 1986).
Effect of Reaction Temperature on the Activityof LIP in Reversed Micelles
Figure 4 shows the effect of the temperature on the reaction
rate in the reversed micellar system. Maximal activity was
observed at 40jC. The optimal temperature of LIP did not
depend on the water content in the reversed micelle. The
lignin peroxidase activity at the lower Wo dropped off more
quickly with temperature than that at the higher Wo. This
suggests that temperature has a larger effect on the stabil-
ity of smaller micelles. The tendency of the temperature
dependency for the reversed micellar system was similar to
that in an aqueous solution (Kirk and Farrell, 1987; Tien,
1987). It has been thought that thermal stability of enzymes
is enhanced by use in a dry organic solvent where the en-
zyme structure becomes rigid (or loose flexibility) (Zaks
and Klibanov, 1988). Because the enzyme exists in water
pools in the present system, the conformation of LIP is not
rigid in the reversed micellar solution. Therefore, when the
sizes of reversed micelles were large enough, the Wo values
exhibited little effect on the temperature dependency
(Fig. 4).
Effect of AOT Concentration in Isooctane
The effect of AOT concentration on the enzymatic activity
of LIP in the reversed micelles was examined and indicated
that the activity reached a maximum at 100 mM (Fig. 5).
The reversed micelles are a kinetic molecular assembly;
thus water molecules between the reversed micelles
mutually fuse and collapse at every collision. An exchange
rate constant (kex) in the AOT reversed micellar system has
been reported to be 106–108 M�1 s�1 at 10–30jC. The kex
has also been known to be increased upon increasing the
concentration of surfactant and temperature (Fletcher et al.,
1987). The catalytic rate of LIP prepared with 100 mM AOT
was improved compared to that with 50 mM AOT (Fig. 5).
In the system utilized in the present study, H2O2 was added
as the AOT reversed micelle solution. Therefore, the
exchange rate of water molecules in the LIP micelle and
the H2O2 micelle might be increased at a higher concen-
tration of AOT, which is reflected in a better activity with
100 mM AOT than with 50 mM.
However, beyond the AOT concentration of 100 mM, the
catalytic activity decreased (Fig. 5). The viscosity of the
reversed micelle solution drastically increased at an AOT
concentration of over 200 mM (Table I). This may result in
lowering the exchange rate of water molecules of LIP– and
H2O2–reversed micelles, which instantly causes a decrease
in enzymatic activity at a higher surfactant concentration.
However, we cannot explain the decrease in activity only by
the viscosity change because the decline of activity does not
correspond to the increase in viscosity with surfactant
concentration. This means that the rate-limiting process in
the enzyme reaction is not necessarily the exchange rate of
reversed micelles.
Effect of Enzyme Concentration in Reversed Micelles
Figure 6 shows the effect of LIP concentration in the
reversed micelles on the reaction rate. At the optimal AOT
concentration (100 mM), the enzymatic activity linearly
increased in the low enzyme concentration range and
attained a maximum value at the concentration of 3 AM. It
was also found that with a higher concentration of AOT
(400 mM), where the number of micelles increased, no
decrease of LIP activity occurred at the higher enzyme
concentration of 3 AM (Fig. 6). The addition of more LIP
into the reversed micelle may cause stronger repulsion
between the negatively charged enzyme (pI = 3.2) and the
Figure 4. Effect of reaction temperature on LIP activity in reversed
micellar system at Wo of 20 (D) and 50 (5).
Figure 5. Effect of AOT concentration during reversed micelle prep-
aration on LIP activity. Wo and pH were adjusted to 50 and 5.0, respec-
tively. The reaction was conducted at 40jC.
Table I. Viscosity of AOT isooctane solutions at 40jC.
AOT concentration (mM) 100 200 300 400
Viscosity (cP) 1.24 1.33 2.85 6.84
498 BIOTECHNOLOGY AND BIOENGINEERING, VOL. 88, NO. 4, NOVEMBER 20, 2004
anionic surfactant AOT. As a result, the reversed micelle
becomes unstable. Therefore, using more AOT molecules
helps maintain the LIP activity with a higher enzyme
concentration. As far as examined, the optimal activity was
seen with 100 mM AOT and 3 AM LIP.
Effect of H2O2 Concentration on LIP Activity
To oxidize an organic substrate by peroxidase, the resting
enzyme should be oxidized by two electrons coupled with
the reduction of H2O2 to H2O to form the oxidized
intermediate, compound I; compound I, in turn, is reduced
back to the resting state via the intermediate formation of
compound II, coupled with two successive one-electron
oxidations of organic substrates (Gold et al., 1989; Kirk and
Farrell, 1987; Tien, 1987). LIP compound II has also been
reported to readily react with H2O2 to form a catalytically
inactive intermediate, compound III, which further reacts
with H2O2, causing an irreversible heme bleach (Wariishi
and Gold, 1989, 1990). Therefore, the initial concentration
of H2O2 is crucial to optimize the LIP reaction. Figure 7
clearly indicates that the activity of LIP in the reversed
micellar system strongly depends on the H2O2 concen-
tration. The dependency of LIP activity with hydrogen
peroxide concentration shows a bell-shaped curve with the
optimal [H2O2] at 0.1 mM. This behavior is very similar to
the LIP reaction in the aqueous system (Kirk and Farrell,
1987; Tien, 1987), suggesting that the LIP reactivity with
H2O2 in water pools of the reversed micelles is almost
identical to that in aqueous solutions and that collision
efficiency of LIP–micelle and H2O2–micelle might not be a
rate-determining step under the conditions utilized.
Effect of Solvents on LIP Activity
The reversed micellar solutions were prepared using various
organic solvents and the enzymatic activity was measured in
each organic solvent (Fig. 8). When decane was used for the
solvent, the initial rate was increased about 4-fold compared
to that in isooctane. However, when the reaction temper-
ature increased to 50jC, the phase separation was observed
in the decane system. Furthermore, when the preparation
temperature was at 30jC or more, it was difficult to generate
a stable reversed micelle in decane. All the solvents used,
except for isooctane, are straight-chain hydrocarbons. The
phase separation observed when using these solvents may
suggest alignment of the linear chains with the hydrophobic
moiety of AOT, thereby disrupting the reversed micelles.
On the basis of the results in Figure 8, we conclude that
isooctane is a more suitable solvent to prepare stable
reversed micelles for the LIP reaction system.
Oxidation of Organic Pollutants
The oxidation of aromatic pollutants was conducted using
LIP hosted in the AOT reversed micelle under the optimized
conditions described above. A decrease in substrates was
observed for all three pollutants, p-nonylphenol, bisphenol
A, and 2,4-dichlorophenol, utilized in this study (Fig. 9). On
Figure 7. H2O2 dependency of LIP activity in reversed micelle. Prep-
aration and operation conditions are the same as described in Figure 6,
but 100 mM AOT was utilized.
Figure 6. Effect of LIP concentration in water pools of the reversed
micelle on reactivity. Wo and pH were adjusted to 50 and 5.0, respectively.
The micelles were prepared using 100 mM (o) or 400 mM (5) AOT. The
reaction was conducted at 40jC.
Figure 8. LIP activity in several organic solvents. LIP – reversed
micelles were prepared in the solvents listed, and the activity was
examined in the same solvent. Preparation and operation conditions are the
same as described for Figure 7, except for the solvents. The activity in
isooctane was defined as one unit.
KIMURA ET AL.: ACTIVATION OF LIGNIN PEROXIDASE BY REVERSED MICELLES 499
the other hand, no reaction was observed with the
lyophilized LIP in isooctane. The decrease of bisphenol A
and 2,4-dichlorophenol was less compared to that of p-
nonylphenol; however, these reactions were confirmed to be
reactivated by adding H2O2 as the reversed micellar solution
(data not shown). Although a large amount of H2O2 allows
LIP denaturation, intermittent addition of H2O2 is effective
to improve the degradation efficiency of pollutants. We
investigated the solubility of the substrates into the reaction
solvent isooctane. 2,4-Dichlorophenol was shown to be
more soluble in isooctane than p-nonylphenol, which was
most completely degraded. This result means that the
solubility of substrates could not be an indicator for
evaluating the degree of degradation of the pollutants by
lignin peroxidase. When the substrate has a surface-active
property, the degree of degradation tends to be high. The
development of an effective procedure using the LIP–
reversed micellar system to degrade environmentally per-
sistent hydrophobic pollutants, including the hydrophobic
phenols used in this study, is now under way as well as
product identification.
CONCLUSIONS
The activation of LIP in an organic solvent was achieved by
using a reversed micellar system. LIP exhibited a high and
stable peroxidative activity in isooctane by being entrapped
in the anionic AOT reversed micelle. This is the first report
that LIP shows a high activity in organic media. The
reversed micellar system was actually very efficient to
activate LIP in organic media, since LIP in the reversed
micelle exhibited only one-tenth of the LIP activity in a
physiological aqueous solution. The enzymatic performance
of LIP in the reversed micelles was influenced by the water
content in the micelle and by pH and the LIP concentration
of water pools of the micelles. The size of the water pool
was also suggested to be larger than the LIP protein for
better performance. Some unique properties for optimizing
LIP catalytic action in organic media were derived from the
unique catalytic mechanism of the enzyme; that is, the
substrate binding and oxidizing site exists at the surface of
the LIP protein. Finally, a possible application of LIP for
degrading environmentally persistent hydrophobic pollu-
tants in organic media is described for the first time.
This research was supported by a Grant-in-Aid for Scientific
Research (B 12450322) from the Ministry of Education, Science,
Sports and Culture of Japan, Kyushu University Interdisciplinary
Programs in Education and Projects in Research Development (to
M.G. and H.W.), and the Proposal-Based New Industry Creative
Type Technology R&D Promotion Program from the New Energy
and Industrial Technology Development Organization (NEDO) of
Japan (to M.G., S.F., H.W., and H.T.).
References
Barbaric S, Luisi PL. 1981. Micellar solubilization of biopolymers in
organic solvents. 5. Activity and conformation of a-chymotrypsin in
isooctane– AOT reversed micelles. J Am Chem Soc 103:4239– 4244.
Bumpus JA, Tien M, Wright D, Aust SD. 1985. Oxidation of persist-
ent environmental pollutants by a white rot fungus. Science 228:
1434–1436.
Crawford RL. 1981. Lignin biodegradation and transformation. New York:
John Wiley & Sons.
Douzou P, Keh E, Balny C. 1979. Cryoenzymology in aqueous me-
dia: micellar solubilized water clusters. Proc Natl Acad Sci USA 76:
681–684.
Doyle WA, Blodig W, Veitch NC, Piontek K, Smith AT. 1998. Two
substrate interaction sites in lignin peroxidase revealed by site-
directed mutagenesis. Biochemistry 37:15097– 15105.
Dunford HB, Stillman JS. 1976. On the function and mechanism of action
of peroxidases. Coord Chem Rev 19:187– 251.
Edwards SL, Raag R, Wariishi H, Gold MH, Poulos TL. 1993. Crys-
tal structure of lignin peroxidase. Proc Natl Acad Sci USA 90:
750–754.
Fletcher PDI, Howe AM, Robinson BH. 1987. The kinetics of solubilisate
exchange between water droplets of a water-in-oil microemulsions. J
Chem Soc, Faraday Trans I 83:985– 1006.
Gold MH, Alic M. 1993. Molecular biology of the lignin-degrading
basidiomycete Phanerochaete chrysosporium. Microbiol Rev 57:
605–622.
Gold MH, Joshi D, Valli K, Wariishi H. 1994. Degradation of chlorinated
phenols and chlorinated dibenzo-p-dioxins by Phanerochaete chrys-
osporium. In: Hinchee RE, Leeson A, Semprini L, Ong SK, editors.
Bioremediation of chlorinated and polycyclic aromatic hydrocarbon
compounds. Boca Raton, FL: Lewis Publishers. p 231– 238.
Gold MH, Wariishi H, Valli K. 1989. Extracellular peroxidases involved in
lignin degradation by the white rot basidiomycete Phanerochaete
chrysosporium in biocatalysis in agricultural biotechnology. In:
Whitaker JR, Sonnet P, editors. ACS Symposium Series 389.
Washington, DC: American Chemical Society. p 127– 140.
Johjima T, Itoh N, Kabuto M, Tokimura F, Nakagawa T, Wariishi H,
Tanaka H. 1999. Direct interaction of lignin and lignin peroxidase
from Phanerochaete chrysosporium. Proc Natl Acad Sci USA
96:1989– 1994.
Kamiya N, Inoue M, Goto M, Naruta Y. 2000. Catalytic and structural
properties of surfactant-horseradish peroxidase complex in organic
media. Biotechnol Prog 16:52– 58.
Khmelnitsky YL, Kananov AV, Klyachko NL, Levashov AV, Martinek K.
1989. Enzymatic catalysis in reversed micelles. In: Pileni MP, editors.
Figure 9. Oxidative conversion of environmental pollutants by LIP–
reversed micelle system in isooctane. p-Nonylphenol (.), bisphenol A (E),
and 2,4-dichlorophenol (n) was added as the substrate (0.1 mM). Prep-
aration and operation conditions are the same as described for Figure 7.
500 BIOTECHNOLOGY AND BIOENGINEERING, VOL. 88, NO. 4, NOVEMBER 20, 2004
Structure and reactivity in reverse micelles. Amsterdam: Elsevier.
p 230– 261.
Kirk TK, Farrell RL. 1987. Enzymatic ‘‘combustion’’: the microbial
degradation of lignin. Annu Rev Microbiol 41:465– 505.
Ly I, Ono T, Kamiya N, Goto M, Nakashio F. 1998. Catalytic properties of
novel reversed micellar system on trans-esterification by a-chymo-
trypsin in organic media. Biochem Eng J 2:29– 33.
Maruyama T, Nagasawa S, Goto M. 2002. Enzymatic synthesis of sugar
amino acid esters in organic solvents. J Biosci Bioeng 94:357–361.
Maruyama T, Noda S, Kamiya N, Goto M. 2003. Ring-opening
polymerization of lactones catalyzed by surfactant-coated lipases in
organic solvents. J Chem Eng Jpn 36:307– 312.
Michizoe J, Goto M, Furusaki S. 2001. Catalytic performance of laccase
hosted in reversed micelles. J Biosci Bioeng 92:67– 71.
Michizoe J, Okazaki S, Goto M, Furusaki S. 2001. Catalytic properties of
lignin peroxidase ALIP-P3 hosted in reversed micelles. Biochem Eng
J 8:129– 134.
Michizoe J, Uchimura Y, Maruyama T, Kamiya N, Goto M. 2003. Con-
trol of water content by reverse micellar solutions for peroxidase
catalysis in a water-immiscible organic solvent. J Biosci Bioeng 95:
425– 427.
Okazaki S, Goto M, Wariishi H, Tanaka H, Furusaki S. 2000. Character-
ization and catalytic property of surfactant– laccase complex in
organic media. Biotechnol Prog 16:583–588.
Okazaki S, Michizoe J, Goto M, Furusaki S, Wariishi H, Tanaka H. 2002.
Oxidation of bisphenol A catalyzed by laccase in reversed micelles in
organic media. Enzyme Microb Technol 31:227–232.
Ono T, Goto M. 1997. Application of reversed micelles in bioengineering.
Curr Opin Colloid Interface Sci 2:397–401.
Pileni M-P, Zemb T, Petit C. 1985. Solubilization by reverse micelles:
solute localization and structure perturbation. Chem Phys Lett
118:414– 415.
Piontek K, Glumoff T, Winterhalter KH. 1993. Low-pH crystal structure
of glycosylated lignin peroxidase from Phanerochaete chrysosporium
at 2.5-A resolution. FEBS Lett 315:119 – 124.
Poulos TL, Edwards SL, Wariishi H, Gold MH. 1993. Crystallographic
refinement of lignin peroxidase at 2 A. J Biol Chem 268:4429– 4440.
Sarkanen KV, Ludwig CH. 1971. Lignins: occurrence, formation, structure
and reactions. New York: John Wiley & Sons.
Shield JW, Ferguson HD, Bommarius AS, Hatton TA. 1986. Enzyme in
reversed micelles as catalyst for organic-phase synthesis reaction. Ind
Eng Chem Fundam 25:603– 612.
Tien M. 1987. Properties of ligninase from Phanerochaete chrysosporium
and their possible applications. CRC Crit Rev Microbiol 15:141– 168.
Tien M, Tu C-PD. 1987. Cloning and sequencing of a cDNA for a
ligninase from Phanerochaete chrysosporium. Nature 326:520– 523.
Wariishi H, Gold MH. 1989. Lignin peroxidase compound III. Formation,
inactivation, and conversion to the native enzyme. FEBS Lett
243:165– 168.
Wariishi H, Gold MH. 1990. Lignin peroxidase compound III. Mechanism
of formation and decomposition. J Biol Chem 265:2070– 2077.
Wariishi H, Sheng D, Gold MH. 1994. Oxidation of ferrocytochrome c by
lignin peroxidase. Biochemistry 33:5545– 5552.
Yoshida S, Watanabe T, Honda Y, Kuwahara M. 1997. Effects of water-
miscible organic solvents on the reaction of lignin peroxidase of
Phanerochaete chrysosporium. J Mol Catal B 2:243– 251.
Zaks A, Klibanov AM. 1988. Enzymatic catalysis in nonaqueous solvents.
J Biol Chem 263:3194– 3201.
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