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Applied and Environmental Microbiology, May 2000, p. 2052-2056, Vol. 66, No. 5
0099-2240/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
Redox Chemistry in Laccase-Catalyzed Oxidation of
N-Hydroxy Compounds
Feng
Xu,1,*
Juozas J.
Kulys,2,*
Kyle
Duke,1
Kaichang
Li,3
Kastis
Krikstopaitis,2
Heinz-Josef W.
Deussen,4
Eric
Abbate,1
Vilija
Galinyte,2 and
Palle
Schneider4,*
Novo Nordisk Biotech, Davis, California
956161; Institute of Biochemistry,
Mokslininku 12, 2600 Vilnius, Lithuania2;
Department of Forest Products, Oregon State University,
Corvallis, Oregon 973313; and Novo
Nordisk A/S, Novo Allé, DK-2880 Bagsværd,
Denmark4
Received 15 November 1999/Accepted 8 February 2000
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ABSTRACT |
1-Hydroxybenzotriazole, violuric acid, and
N-hydroxyacetanilide are three N-OH compounds capable of
mediating a range of laccase-catalyzed biotransformations, such as
paper pulp delignification and degradation of polycyclic
hydrocarbons. The mechanism of their enzymatic oxidation was studied
with seven fungal laccases. The oxidation had a
bell-shaped pH-activity profile with an optimal pH ranging from 4 to 7. The oxidation rate was found to be dependent on the redox potential difference between the N-OH substrate and laccase. A laccase with a
higher redox potential or an N-OH compound with a lower redox potential
tended to have a higher oxidation rate. Similar to the enzymatic
oxidation of phenols, phenoxazines, phenothiazines, and other
redox-active compounds, an "outer-sphere" type of
single-electron transfer from the substrate to laccase and proton
release are speculated to be involved in the rate-limiting step for
N-OH oxidation.
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INTRODUCTION |
Laccases (EC 1.10.3.2) are multi-Cu
oxidases that can catalyze the oxidation of a range of reducing
substances with the concomitant reduction of O2 (for recent
reviews, see reference 24 and references therein).
Because of their capability of catalyzing the oxidation of aromatic
compounds, laccases are receiving increasing attention as potential
industrial enzymes in various applications, such as pulp
delignification, wood fiber modification, dye or stain bleaching,
chemical or medicinal synthesis, and contaminated water or soil
remediation (15, 37).
Laccases contain one type 1 (T1) Cu center, one type 2 (T2) Cu center,
and one type 3 (T3) Cu center. The T2 and T3 sites form a trinuclear Cu
cluster onto which O2 is reduced. The T1 Cu oxidizes the
reducing substrate and transfers electrons to the T2 and T3 Cu. Laccase
is able to oxidize certain phenols with E0 values higher
than its own (0.5 to 0.8 V versus the normal hydrogen electrode
[NHE]) (36). However, many inorganic and organic compounds
with comparable E0 values (such as
1,2,3,5-tetramethoxybenzene [18]) are not laccase
substrates due to unfavorable kinetics. Under certain conditions,
however, these compounds can be indirectly oxidized by laccase
through the mediation of small, redox-active laccase
sub- strates. 2,2'-Azinobis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS)
was the first compound found capable of efficiently mediating the
laccase oxidation of high-E0, nonsubstrate lignin model
compounds (such as veratryl alcohol and nonphenolic lignin model
dimers) (8). Based on product structure analysis, it has
been proposed that laccase-oxidized ABTS can abstract an H atom from
the lignin model compounds, leading to indirect laccase catalysis upon
the oxidation of the compounds (25). To date, other types of
mediators, particularly phenoxazines and N-OH compounds, also have been
recognized for their mediation function in laccase catalysis (1,
6, 17, 29).
Mediated laccase catalysis has been applied to a wide range of
applications, such as pulp delignification (9, 10, 12, 22,
32), textile dye bleaching (31), polycyclic aromatic hydrocarbon degradation (16, 23), pesticide or insecticide degradation (1, 29), and organic synthesis (13,
28). For the paper and pulp industry, novel biological or
enzymatic bleaching technologies (including mediated laccase catalysis) have attracted increasing attention (9, 10, 12, 14, 22, 27,
32) because of concerns regarding the environmental impact of the
chlorine-based oxidants currently being used in delignification or bleaching.
Detailed, comparative information on the interaction between mediator
and laccase remains to be reported (22), although various
physical and chemical characterizations have been performed on several
well-known laccase mediators (2, 4, 7, 11, 21, 35). For
N-OH-type mediators, it has not been clear whether their oxidation by
laccase involves H abstraction or electron transfer, similar to that
found with the oxidation of phenol (38). To better
understand the mechanism that governs the oxidation of these compounds
by laccase, we studied the interactions of three N-OH compounds (Fig.
1) with seven fungal laccases. The observed dependence of the reaction rate on 
E0 suggests
that the laccase-catalyzed oxidation of N-OH compounds is governed by a
mechanism similar to that reported for phenols, phenoxazines, and
phenothiazines.
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MATERIALS AND METHODS |
Materials.
The chemicals used were commercial products of at
least reagent grade. Botrytis cinerea laccase (BcL)
(22), Coprinus cinereus laccase-1 (CcL)
(30), Myceliophthora thermophila laccase (MtL) (5), Myrothecium verrucaria bilirubin oxidase
(MvBO) (39), Pycnoporus cinnabarinus laccase
(PcL) (22), Rhizoctonia solani laccase 4 (RsL)
(34), Scytalidium thermophilum laccase (StL) (39), and Trametes villosa (Polyporus
pinsitus) laccase 1 (TvL) (40) were purified as
previously reported. Violuric acid (VA) and 1-hydroxybenzotriazole
(HBT) were purchased from Aldrich. Promazine and chloropromazine were
purchased from Sigma. N-Hydroxyacetanilide (NHA) and
phenothiazine-10-propionic acid (PP) were synthesized as described
previously (26, 33). 10-Methyl phenothiazine, 3,10-dimethyl
phenothiazine, 10-ethyl phenothiazine, 10-(2-hydroxyethyl) phenothiazine, phenothiazine 10-methylpropionate, phenothiazine 10-propionamide, phenothiazine 10-propionitrile,
10-methyl-1-carboxylic acid phenothiazine, 10-methyl-2-carboxylic
acid phenothiazine, 10-methyl-3-carboxylic acid
phenothiazine, 10-ethyl-4-carboxylic acid phenothiazine,
10-(3-hydroxypropyl) phenothiazine,
10-(2-ethoxy-2'-hydroxyethyl) phenothiazine, 2-acetyl-10-methyl
phenothiazine, 10-methyl-3-(2-hydroxyethyl) phenothiazine,
2-chloro-10-methyl phenothiazine, 2-methoxy-10-methyl phenothiazine,
10-methyl phenoxazine, 10-(2-hydroxyethyl) phenoxazine, and phenoxazine
10-propionic acid were synthesized as described elsewhere
(20a).
Instruments.
UV-visible absorption spectroscopy (including
kinetic spectral measurements) was performed either on a
spectrophotometer (Shimadzu UV160U or Gilford Instruments 2600)
and a quartz cuvette or on a microplate reader (Molecular Devices
Thermomax) and 96-well microplates (Costar tissue culture plates).
Cyclic and differential pulse voltammetry analyses were performed on a
computer-controlled electroanalytical system (Cypress Systems),
with a glass carbon working electrode (Cypress Systems model
CS-1087), a KCl-saturated calomel reference electrode (Radiometer
model K-401), and a platinum wire counterelectrode (0.2-mm diameter,
4-cm length, mounted on the end of the reference electrode). Surface
cleansing of the working electrode was carried out by polishing with
alumina and washing with water.
Electrochemistry.
To determine the E0 of the
N-OH compounds, cyclic voltammetry was performed at 25°C in (aerobic)
solutions containing 1 mM N-OH compound, 0.1 M KCl, 33 mM sodium
phosphate, 33 mM sodium borate, and 33 mM sodium carbonate (pH 4 to
10). The scanning rate was 0.1 V/s. Measured potentials were compared
to the NHE by considering the E0 of the KCl-saturated
calomel reference electrode to be 0.242 V against the NHE.
The E
0 values of the phenoxazines and phenothiazines were
measured by differential pulse voltammetry in 50 mM sodium phosphate
(pH 7) at room temperature, with a 50-mV pulse height, a 2-mV
step
height, a 40-ms pulse width, an 0.8- to 0.3-V electrode potential
change, and a 12 to 21 µM concentration. E
0 values of
0.66 to
0.95 V were observed (
20a).
O2 electrode-based enzymatic assays.
Laccase
activity was measured in 10 mM morpholineethanesulfonic acid (MES)-NaOH
(pH 5.5) at 20°C with a Hansatech O2 cell (38). ABTS was used as a calibrator. The N-OH substrate
stock solutions were made in dimethylformamide (1 M for HBT, 0.5 M for VA, and 0.1 M for NHA). At the tested level (
10%), the
dimethylformamide introduced along with the substrate did not alter the
kinetic measurements (as tested by ABTS oxidation). The laccase
concentrations were 0.8 µM for TvL, 1 to 15 µM for RsL, 2 to 37 µM for MtL, 4 to 55 µM for StL, 3 to 19 µM for CcL, 0.4 to 4 µM
for PcL, and 1 to 3 µM for BcL.
The pH-activity profile was measured at 20°C in Britton-Robinson
buffer, made by mixing 0.1 M boric acid, 0.1 M acetic acid,
0.1 M
phosphoric acid, and 0.5 M NaOH. The substrate concentrations
were 40 to 100 mM for HBT, 33 to 58 mM for VA, and 6.7 mM for
NHA. The laccase
concentrations were 0.8 µM for TvL, 2 to 15 µM
for RsL, 2 to 11 µM for MtL, 2 to 16 µM for StL, 3 to 19 µM for
CcL, 0.7 µM for
PcL, and 3 to 6 µM for
BcL.
PP oxidation was performed with 0.01 to 1 mM PP in 10 mM MES (pH 5.5).
The concentrations of laccase were 0.1 µM for TvL,
2 µM for RsL, 2 µM for MtL, 5 µM for StL, and 1 µM for MvBO. The
stock solution
of PP (0.1 M) was made in 0.1 M
NaOH.
Spectrophotometric enzymatic assays.
Spectrophotometric
enzymatic assays were performed with 50 mM sodium acetate (pH 5.5) at
25°C. The oxidation of VA or NHA by laccase was monitored at 310 nm
with a molar absorption (
) value of 13.9 or 8.9 mM
1
cm1, respectively. Substrate concentration ranges were 10 to 120 µM for VA and 25 to 200 µM for NHA. Laccase concentration
ranges were 0.2 to 1 µM (when oxidizing VA) or 0.2 µM (when
oxidizing NHA) for TvL, 1 µM for CcL, and 1 µM (when oxidizing NHA)
or 10 µM (when oxidizing VA) for MtL. Third-order polynomial
[c = a + b(t) + c(t2) + d(t3)], where c is the concentration and
b is the initial rate constant) nonlinear regression was
applied (using the MathCad program) to the kinetic data to extract the
apparent rate constant.
For phenoxazines and phenothiazines, their oxidation (into cation
radicals) by TvL was monitored at 525 and 514 nm with
values of 16 and 8.9 mM
1 cm
1, respectively. The
reactions were carried out with 1.5 to 80
µM substrate and 2.5 to 40 nM TvL in 50 mM sodium acetate (pH
5.3) and 1% ethanol at 25°C.
Km values from 6 to 678 µM and
kcat (
Vmax/[laccase])
values from 120 to 8,580 min
1 were observed
(
20a).
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RESULTS |
Electrochemistry and redox potentials of N-OH compounds.
Under
the conditions used in this study, the cyclic voltammetry of HBT
exhibited irreversible oxidation, similar to the observation previously
reported (7, 19). Depending on pH, an anodic peak was
observed near a peak potential [Epa] of 1.1 to 1.2 V,
with a peak current intensity (Ipa) corresponding to 2.1 to
2.4 electrons transferred per HBT molecule. Within the scanning rate
range, only a small cathodic peak (with a peak potential
[Epc] near 0.54 V and a peak current intensity
[Ipc] 10% that of Ipa) was detected, indicating the residual reduction of oxidized HBT. As shown in Fig.
2, the pH dependence of Epa
for HBT was not significant.
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FIG. 2.
Formal redox potentials of the N-OH compounds as a
function of pH. Traces a, b, and c represent the pH dependence of
Epa of HBT, E1/2 of VA, and E1/2 of
NHA, respectively. For HBT, there was no significant acid-base
transition for its Epa; thus, no apparent pKa
was extracted from trace a. For VA, two apparent pKa values
of 6.4 and 8.6 were extracted from trace b. For NHA, two apparent
pKa values of 3.7 and 6.3 were extracted from trace c.
E1/2 values of 0.83 and 0.91 V at pH 4 have been reported
for NHA and VA, respectively, by R. Bourbonnais et al. (Oxidative
Enzymes for Lignocellulose Processing, Symp. Am. Chem. Soc. 217th Nat.
Meet., Anaheim, Calif., 21 to 25 March 1999).
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Unlike HBT, VA showed a well-shaped cathodic peak, indicating apparent
reversibility. The differences between the anodic and
cathodic peak
potentials (
E
p = E
pa 
E
pc) were ~70, 80, and 140
mV for pHs 4 to 8, 9 and 10, respectively. Based on the I
pa, ~1.4,
1.5, 1.6, 1.4, 1.3, and 1.1 electrons were transferred per VA
molecule during oxidation at
pHs 4, 5, 6 to 7, 8, 9, and 10,
respectively.
Like VA, NHA had a quasi-reversible cyclic voltammogram. The
differences (
E
p) were ~130, 110, and 80 mV for pHs 4 to 5, 6
to 7, and 8 to 10, respectively. Based on the I
pa,
~1.1, 1.0,
0.8, 0.9, and 1.1 electrons were transferred per NHA
molecule
during oxidation at pHs 4 to 6, 7, 8, 9, and 10, respectively.
As shown in Fig.
2, the formal redox potentials
{E
1/2 = [(E
pa + E
pc)/2]} of both VA and NHA were pH dependent. For pH
ranges
of 6 to 9 and 4 to 7, the E
1/2-pH plot of VA or NHA
had an apparent
slope of 50 or 56 mV per pH unit, respectively. For a
given pH,
the redox potentials of these three N-OH compounds were
on the
order of HBT > VA >
NHA.
Laccase-catalyzed oxidation of N-OH compounds.
Serving as a
reducing substrate for laccase, the three N-OH compounds exhibited
typical Michaelis-Menten kinetics, as monitored by concomitant
O2 reduction (Fig. 3). Table
1 shows the Km and kcat values observed in 10 mM MES (pH 5.5) for
the seven laccases and the three N-OH compounds. For MtL, up to 40 mM
NHA could not lead to saturation of the initial oxidation rate, thus
not allowing an accurate measurement of Km and
kcat values.
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FIG. 3.
O2 consumption in RsL-catalyzed oxidation of
NHA at pH 5.5. Plotted against the initial NHA concentration [NHA]
are the initial O2 consumption rate (as the output voltage
change rate) (A) and the final O2 consumed (as the
final output voltage change) (B). For graph clarity, two sets of
data, obtained with 0.05 and 0.3 mM NHA, are omitted from panel
B. Their symbols overlap with those for 0 and 0.33 mM NHA (shown), and
their values were included in the graph fitting. In panel A, the solid
curve shows the fit to the Michaelis-Menten equation
{v = Vmax[NHA]/(Km + [NHA])} with a Km of 2.0 ± 0.5 mM and
a Vmax of 0.39 ± 0.03 V
min1 or 0.19 ± 0.01 mM min1
(corresponding to a kcat of 150 ± 10 min1 (mean ± standard deviation). In panel B, the
horizontal broken line represents the voltage change (0.58 ± 0.03 V, averaged over the data obtained with 3.3, 5.0, and 6.7 mM NHA)
corresponding to maximal O2 consumption. Its cross point
with the other broken line (voltage change, 0.5 × [NHA];
r2, 0.92), obtained by fitting the data
obtained with 0, 0.05, 0.30, and 0.33 mM NHA, yielded a saturating
[NHA] of 1.2 mM. By dividing 1.2 mM by 0.28 mM, the dissolved
[O2] in water, we estimated an oxidation stoichiometry
number of 4.1.
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The oxidation of VA and NHA by TvL, CcL, and MtL was also monitored
spectrophotometrically. Under laccase catalysis, the oxidation
of VA
led to a decrease of the absorbance centered at 310 nm.
The oxidation
of NHA increased the absorbances at 220 to 230 and
266 to 370 nm (with
maxima at 229, 283, and 308 nm) and decreased
the absorbance centered
at 245 nm (with two apparent isobestic
points at 230 and 266 nm).
Before the full formation of the apparently
stable product (peak
wavelengths [
max] at 229, 283, and 308 nm;
trough
wavelengths [
min] at 261 and 290 nm), a
transient product
seemed to be formed, as demonstrated by a spectrum
with
max at
245 and 323 nm and
min at 293 nm. A linear dependence of rate
on substrate concentration was observed
at the selected concentration
ranges. For VA, apparent rate constants
of ~350, 170, and 5.9
M
1 s
1 were observed
for TvL, CcL, and MtL, respectively. For NHA, apparent
rate constants
of ~2,100, 330, and 30 M
1 s
1 were
observed for TvL, CcL, and MtL,
respectively.
Dependence on E0 and pH.
Figure
4 shows the dependence of
kcat, Km, and
kcat/Km on
E0 {E0 [laccase (T1 Cu)] E0 [substrate]} at pH 5.5. For HBT, VA, and NHA,
the Epa, E1/2, and E1/2 determined
from cyclic voltammetry, respectively, were used to calculate
E0. For HBT, the use of Epa would slightly
overestimate E0, since the irreversible decay of the
immediately oxidized HBT would yield Epa greater than
or equal to E1/2 (~1.11 V at pH 4 [7]), according to the kinetic characteristics of a
homogeneous redox catalysis electrochemical reaction
(3). An apparently linear, positive correlation was observed between log(kcat) and
E0 as well as between
log(kcat/Km) and
E0, while an apparently linear, negative correlation was observed between log(Km) and E0.
All three N-OH substrates showed bell-shaped pH-rate profiles with an
optimal pH ranging from 4 to 7 (Table 1).
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FIG. 4.
Dependence of Km,
kcat, and
kcat/Km on
E0. Symbols:  , HBT;  , VA; ×, NHA. Correlation
lines: A, log(Km) = 1.3(E0) + 0.46 (r2, 0.26); B,
log(kcat) = 4.5(E0) + 2.5 (r2, 0.74); C,
log(kcat/Km) = 5.8(E0) + 5.0 (r2,
0.82). Units: A, Km, mM; B,
kcat, min1; C,
kcat/Km, M1
min1. E0 is reported in V. E0
values for laccase are taken from reference 36.
Error bars indicate standard deviations.
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Laccase-catalyzed oxidation of phenoxazines and
phenothiazines.
Serving as a reducing substrate for laccase,
the oxidation of PP exhibited typical Michaelis-Menten kinetics, as
monitored by concomitant O2 reduction. The
Km and kcat values
(mean ± standard deviation) observed in 10 mM MES (pH 5.5) were
120 ± 50 µM and 2,500 ± 400 min1 for TvL,
32 ± 5 µM and 8 ± 1 min1 for RsL, 120 ± 40 µM and 11 ± 4 min1 for MtL, 47 ± 5 µM and 4.7 ± 0.1 min1 for StL, and 30 ± 5 µM and 21 ± 1 min1 for MvBO, respectively. An
apparently linear correlation was observed between
log(kcat/Km) and
E0 (Fig. 5), similar to
the data obtained for the TvL-catalyzed oxidation of more than 20 phenothiazines and phenoxazines (20a).
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FIG. 5.
Correlations between
log(kcat/Km) and
E0 for phenothiazines and phenoxazines. Symbols: ,
oxidation by TvL of 3 phenoxazines and 20 phenothiazines (see Materials
and Methods for their formulas) (20a); , oxidation of PP
(E0, 0.71 V) by TvL, RsL, MtL, StL, and MvBO at pH 5.3 to
5.5. Correlation:
log(kcat/Km) = 5.1(E0) + 6.4 (r2,
0.47). Units: kcat/Km,
M1 min1; E0, V.
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DISCUSSION |
Redox chemistry of N-OH compounds.
It is known that
the oxidation of HBT generates a highly unstable intermediate,
putatively an N-O· radical, that quickly decays into
catalytically inactive secondary product(s), including
benzotriazole (21). An apparent E1/2 of ~1.1 V
has been reported for a two-electron electrochemical oxidation of HBT
at pH 4 (7). In our study, instability of the putative HBT
radical was observed over the pH range of 4 to 10. The better stability
observed for the immediate oxidation products (likely
N-O· in nature) of VA and NHA could be related to their
E1/2 values, which were 0.2 to 0.3 V lower than that of
HBT. The reduction in E1/2 might decrease the oxidative potency or activity of N-O
thus enhancing stability.
As shown in Fig.
2, the E
1/2 of VA and NHA decreased when
pH increased. Since phenyl-N-OH is a heteroatomic homolog of phenol,
the oxidation of an aromatic N-OH compound could lead
to H
+ release (N-OH
N-O
· + e
+ H
+), as for phenol (C-OH
C-O
· + e
+ H
+). According to the Nernst
equation, E
0 = E
0 + {RT/F}
ln {[N-O
·][H
+]/[N-OH]} = E
0 + {RT/F} ln
{[N-O
·]/[N-OH]}
RT/{F log(e)}pH, such
H
+ release would lead to a lower potential at a
higher pH (~0.06-V
reduction per pH unit at room temperature),
reflecting the fact
that the quenching of H
+ by
OH
at a higher pH facilitates thermodynamically the N-OH
oxidation.
Thus, the decrease in E
1/2 of VA and NHA with an
increase in pH
indicated the involvement of H
+ release and
the concomitant production of N-O· during the
oxidation.
Electron transfer from N-OH compounds to laccases.
At steady
state, the rate-limiting step for phenol oxidation by laccase involves
the Marcus "outer-sphere" mechanism. In this mechanism,
E0 (together with reorganization energy and
transmission coefficient) determines the electron transfer rate,
distinguishing it from other oxidation mechanisms (i.e., H
abstraction), where energetic factors related to covalent bond are most
important (i.e., homolytic O---H bond dissociation energy). As shown in
Fig. 4, a linear correlation existed between
log(kcat) or
log(kcat/Km) (in which
kcat/Km could be
approximated as the second-order rate constant of the oxidation) and
E0 (the driving force for electron transfer from the
N-OH compound to laccase) for laccase-catalyzed oxidation of the N-OH
compounds. When the data for a wide variety of phenols, phenothiazines,
phenoxazines, N-OH compounds, and other inorganic and organic
redox-active molecules are analyzed together, a common linear
correlation between
log(kcat/Km) and E0 can be found (Fig. 6).
Thus, as for other laccase substrates, the rate-limiting step of
laccase-catalyzed N-OH oxidation involves electron transfer from the
substrate to the T1 Cu site in laccase. It is E0 that
dominates the oxidation rate. The higher E0 (laccase) or
the lower E0 (N-OH) is, the faster the oxidation rate tends to be. Other factors (such as the composition, structure, or
pKa of the substrate) seem to be minor, but they could
fine-tune the activity for a given E0 (an effect that
might contribute to the scattering shown in Fig. 6).
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FIG. 6.
Correlations between
log(kcat/Km) and
E0 for laccase catalysis. Symbols: , oxidation of 24 phenols by 10 fungal on plant laccases (36);  , oxidation
of 3 phenoxazines and 22 phenothiazines by TvL, as well as oxidation of
PP by 5 fungal laccases (data from Fig. 5); , oxidation of three
N-OH compounds by 7 fungal laccases (data from Table 1 and Fig. 4C); +,
oxidation of ABTS, K4Fe(CN)6 and
morpholinoaniline by up to 10 fungal or plant laccases (20,
36). Other conditions: pH, 5.3 to 5.5; temperature, 20 to 25°C.
Correlation:
log(kcat/Km) = 6.4(E0) + 6.4 (r2,
0.65). Units: kcat/Km,
M1 min1; E0, V.
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The apparent negative correlation between
log(
Km) and
E
0 suggests
that substrate affinity tends to increase when
E
0
increases
(realized by either E
0 [laccase] increase or
E
0 [substrate] decrease)
(Fig.
4A), a phenomenon also
observed for phenolic substrates
(
36,
39). Prior to
electron transfer, the filled (valence)
molecular orbitals of N-O
in the N-OH compounds (or the phenoxy-O
in phenols) overlap with the
half-occupied molecular orbitals
(HOMO) of T1 Cu when the
substrate is bound to laccase. A larger
E
0 could create
a transitional energy state more favorable for
the molecular
orbital interaction, resulting in better substrate
binding and
consequently faster electron
transfer.
Dependence of activity on pH.
When oxidizing a phenolic
substrate, laccase generally possesses a bell-shaped pH-activity
profile. Two opposing factors, E0 (involving
substrate and laccase T1 Cu) and OH inhibition
(involving T2 Cu in laccase), are suggested to play important roles in
determining the pH-activity profile (38). Like phenols, HBT,
VA, and NHA have redox potentials that decrease when pH increases (Fig.
2). Since the E0 of laccase is often quite insensitive to
pH change (38), the decrease in the E0 of N-OH as pH increases would increase E0, which in turn would
enhance the oxidation rate through the correlation shown in Fig. 4.
However, the OH inhibition of laccase would become
overwhelming at an alkaline pH. The combination of these two effects
might contribute to the bell-shaped pH-activity profiles of N-OH compounds.
The speculation of an H
+ release step during
laccase-catalyzed N-OH oxidation, together with the observation that
the reduction
of one O
2 was accompanied by the oxidation of
about four N-OH
groups, indicates that the reaction N-OH
N-O· + e
+ H
+ might be
involved in the rate-limiting step, similar to the reaction
C-OH
C-O· + e
+ H
+, which is
involved in laccase-catalyzed phenol oxidation (
36).
Overall remarks.
The results of this study suggest that the
initial oxidation of a phenol (aryl C-OH) compound by laccase is quite
similar to the oxidation of an aryl N-OH (phenol homolog) compound in terms of the dependence of the initial rate on E0 and pH.
In general, phenol is first oxidized to a highly unstable phenoxy
radical (aryl C-O·), which then surrenders an
additional e (at a rate faster than that of the first
e transfer) to yield a stable, but much less active,
quinone. Oxidation of N-OH compounds also involves a single
e transfer at the initial oxidation step.
N-O· could be less active but more stable than a phenoxy
radical. In laccase-catalyzed pulp delignification, a desirable redox
mediator should be a good laccase substrate, have a half-life at its
oxidized form long enough to permit diffusion to heterogeneous lignin,
and possess high oxidation potency to effectively oxidize lignin. In
comparison with those of a phenoxy radical, the activity and stability
of N-O· seem to be better balanced, which could
contribute to the better performance of the latter as a mediator for
laccase-based delignification.
| |
ACKNOWLEDGMENTS |
We thank Alan V. Klotz and Henrik Bisgård-Frantzen of
Novo Nordisk for critical reading of the manuscript and helpful suggestions.
| |
FOOTNOTES |
*
Corresponding author. Mailing address for Feng Xu: Novo
Nordisk Biotech, 1445 Drew Ave., Davis, CA 95616. Phone: (530)
757-8100. Fax: (530) 758-0317. E-mail: fengxu{at}nnbt.com.
Mailing address for Juozas J. Kulys: Institute of Biochemistry,
Mokslininku 12, 2600 Vilnius, Lithuania. Phone: 370-2-729176. Fax:
370-2-729196. E-mail: jkulys{at}bchi.lt. Mailing address for
Palle Schneider: Novo Nordisk A/S, Novo Allé, 6BS.99, DK-2880
Bagsværd, Denmark. Phone: 45-44422261. Fax: 45-44422202. E-mail:
ps{at}novo.dk.
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Applied and Environmental Microbiology, May 2000, p. 2052-2056, Vol. 66, No. 5
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Abstract
Introduction
