Biocatalytic synthesis of polycatechols from toxic aromatic compounds

Article abstract of DOI:10.1021/es035458q

A process is described in which toxic aromatic compounds are converted by toluene dioxygenase and in turn toluene cis-dihydrodiol dehydrogenase to catechols which are further polymerized by peroxidase-catalyzed oxidation producing polycatechols. Three approaches for obtaining catechols were employed: (1) addition of halogenated aromatics to P. putida F1, resulting in the accumulation of halogenated catechols; (2) inhibition of catechol 2,3-dioxygenase of P. putida F1 by known aromatic and aliphatic inhibitors; and (3) overexpression of toluene dioxygenase and toluene cis-dihydrodiol dehydrogenase genes in E. coli JM109. The process is suitable for producing novel catechols that upon oxidation may yield polymers with unique properties, presenting a tool for producing tailor-made biopolymers. Formation of 3-chlorocatechol from chlorobenzene, 3,4-dichlorocatechol from 1,2-dichlorobenzene, and catechol from benzene and their subsequent oxidation and polymerization was demonstrated. Oxidation of catechol yielded polymers with molecular weights of up to 4000 Daltons. Their apparently high water solubility eliminates the need for water-miscible solvents. In aqueous solution oxidation of catechols was rapid, yet the presence of 20%, 30%, and 40% ethanol, resulted in a rate decrease of 31%, 95%, and 93%, respectively. The advantage is that significantly less peroxidase is required for performing the reactions if miscible solvents are not employed. Furthermore, water-soluble polymers may be desirable for many applications.

Full text of DOI:10.1021/es035458q

Environ. Sci. Technol. 2004, 38, 4753-4757
substituent, C2,3O is either inactivated by the halogenated
Biocatalytic Synthesis of
Polycatechols from Toxic Aromatic
Compounds
G A R Y W A R D , †
R E B E C C A E . P A R A L E S , ‡ A N D
C A R L O S G . D O S O R E T Z * , †
catechol itself (4) or its ring cleavage product, an acyl chloride
(5). As a result, accumulation of halogenated catechols results
when P. putida F1 acts on halogen-substituted aromatics
(6). Recombinant E. coli JM109(pDTG602), containing the
todC1C2BAD genes from P. putida F1 encoding TDO and
TDD has also been shown to yield catecholic transformation
products upon addition of aromatic substrates that are
substrates for TDO (7-9). Inhibition of C2,3O in P. putida
F1 would similarly be expected to result in the accumulation
of catecholic transformation products from aromatic sub-
strates. A range of aromatic and aliphatic compounds are
known be inhibitors of C2,3O (4, 10, 11).
Division of Environmental, Water and Agricultural
Engineering, Faculty of Civil and Environmental Engineering,
Technion-Israel Institute of Technology, Haifa 32000, Israel,
and Section of Microbiology, University of California,
Davis, California 95616
Peroxidases are heme enzymes, which catalyze the
oxidation of a variety of aromatic substrates utilizing H2O2
as an electron acceptor (12). Phenols are oxidized to generate
phenoxy radicals, which have a tendency to couple forming
oligomeric and polymeric products. This phenomenon has
been considered for the biocatalytic production of useful
oligomers and polymers (12, 13). The major advantages of
employing enzymes for the synthesis of oligomers and
polymers are as follows: (i) enzyme-catalyzed polymerization
of phenols proceeds under mild reaction conditions without
the use of toxic reagents (environmentally benign process);
(ii) phenol monomers having various substituents can be
polymerized to give new classes of functional polyaromatics;
(iii) the structure and solubility of the polymer can be
controlled by changing the reaction conditions; (iv) the
polymerization procedure as well as the polymer isolation
are very facile. Phenolic polymers synthesized by peroxidase-
catalyzed reactions possess electrical and optical properties
due to the conjugated backbone (14-16). These unique
properties could enable novel applications in the area of
adhesives, biosensors, electronics, molding, photonics, pho-
tolithography, rechargeable batteries, and thermoresistant
filters (12, 13, 17). Moreover, polymers formed by peroxidases
could be used as an alternative to phenolic-formaldehyde
resins (14). These resins show excellent toughness and
temperature-resistant properties. However, the toxic nature
of formaldehyde causes problems in their manufacture and
practical use.
A process is described in which toxic aromatic compounds
are converted by toluene dioxygenase and in turn
toluene cis-dihydrodiol dehydrogenase to catechols which
are further polymerized by peroxidase-catalyzed oxidation
producing polycatechols. Three approaches for obtaining
catechols were employed: (1) addition of halogenated
aromatics to P. putida F1, resulting in the accumulation of
halogenated catechols; (2) inhibition of catechol 2,3-
dioxygenase of P. putida F1 by known aromatic and aliphatic
inhibitors; and (3) overexpression of toluene dioxygenase
and toluene cis-dihydrodiol dehydrogenase genes in E. coli
JM109. The process is suitable for producing novel
catechols that upon oxidation may yield polymers with
unique properties, presenting a tool for producing tailor-
made biopolymers. Formation of 3-chlorocatechol from
chlorobenzene, 3,4-dichlorocatechol from 1,2-dichloroben-
zene, and catechol from benzene and their subsequent
oxidation and polymerization was demonstrated. Oxidation
of catechol yielded polymers with molecular weights of
up to 4000 Daltons. Their apparently high water solubility
eliminates the need for water-miscible solvents. In aqueous
solution oxidation of catechols was rapid, yet the presence
of 20%, 30%, and 40% ethanol, resulted in a rate decrease
of 31%, 95%, and 93%, respectively. The advantage is that
significantly less peroxidase is required for performing
the reactions if miscible solvents are not employed.
Furthermore, water-soluble polymers may be desirable for
many applications.
The current research tested the conception of a process
in which catechols, obtained upon transformation of mono-
aromatic compounds by TDO and in turn TDD, were further
polymerized by peroxidase-catalyzed oxidation to polycat-
echols as schematically represented in Figure 1. Three
approaches for producing catechols were studied. The
microbial-enzymatic process highlights the possibility of
effectively extending the range of substrates available for
peroxidase-catalyzed oxidation and offers a tool for designing
tailor-made biopolymer production.
Introduction
The toluene dioxygenase (TDO) of P. putida F1, which
initiates the degradation of aromatic compounds, is known
to oxidize on the order of 100 substrates (1). Dioxygenation
by TDO is followed by dehydrogenation by toluene cis-
dihydrodiol dehydrogenase (TDD) resulting in the formation
of a catechol (Figure 1). TDD can oxidize a wide range of
dihydrodiol substrates (2). The catechol 2,3-dioxygenase
(C2,3O) that catalyzes cleavage of the aromatic ring in the
next step of the pathway also has a broad substrate specificity
(3). However, when the aromatic ring contains a halogen
Materials and Methods
Strains. Pseudomonas putida F1 (18) and recombinant
Escherichia coli JM109(pDTG602) (19) were kindly provided
by D. T. Gibson.
Growth and Maintenance of Microorganism s. P. putida
F1 was maintained on nutrient agar and grown in minimal
medium (MSB) (20) containing 40 mM pyruvate as carbon
source at 30 °C in a rotary shaker at 200 rpm. TDO was induced
by toluene in the vapor phase. Filter paper, soaked in toluene,
was placed in a sidearm molded to the Erlenmeyer flask.
Recombinant E. coli JM109(pDTG602) carrying the
todC1C2BAD genes was maintained on agar solidified Luria-
Bertani (LB) medium containing ampicillin (200 mg/ L) (9)
and cultivated on LB medium, supplemented with 200 mg/ L
* Corresponding author phone: 972-4-8294962; fax: 972-4-
8228898; e-mail: carlosd@tx.technion.ac.il.
† Technion-Israel Institute of Technology.
‡ University of California.
9
10.1021/es035458q CCC: $27.50
Published on Web 08/06/2004
2004 Am erican Chem ical Society
VOL. 38, NO. 18, 2004 / ENVIRONMENTAL SCIENCE & TECHNOLOGY 4 7 5 3

HRP, varying concentrations of catechol, and 2.5 mM H2O2
in 50 mM Na-K phosphate buffer, pH 7.0 at 25 °C.
Spectrophotom etry. Spectral changes during oxidation
by HRP were monitored using a Hewlett-Packard HP8453
diode array spectrophotometer.
HPLC Analysis. HPLC analysis was conducted using a
Hewlett-Packard HPLC (HP1100 series) equipped with a diode
array detector. All solvents were of far-UV-quality HPLC-
grade purity where available.
FIGURE 1. Schematic representation of the biocatalytic synthesis
of polycatechols. The reaction is initiated by toluene dioxygenase
(TDO). Subsequent dehydrogenation by toluene cis-dihydrodiol
dehydrogenase (TDD) results in the formation of a catechol which
is oxidized by peroxidase (Pox) resulting in the formation of
polymeric products.
Analytical Reverse Phase HPLC. Analytical reverse phase-
HPLC was conducted as previously described (23), with a
few modifications. The mobile phase consisted of component
A (10%, v/ v, aqueous acetonitrile, 1 mM trifluoroacetic acid)
and component B (40%, v/ v, aqueous methanol, 40%, v/ v,
aqueous acetonitrile, 1 mM trifluoroacetic acid) in the
following program: initially 100% A; linear gradient over 15
min to 100% B; held isocratically at 100% B for 5 min; linear
gradient to 100% A over 5 min. The flow rate in all cases was
maintained at 1 mL/ min. Peak detection was at 210, 254,
and 280 nm. Catechol was quantified by integration of peak-
areas at 280 nm, with reference to calibrations, which were
made using known amounts of authentic catechol.
Semipreparative Reverse Phase HPLC. Transformation
products were fractionated using a semipreparative reverse
phase Lichrospher 100 reverse phase-C18 column (25 cm ×
10 mm i.d., 10 µm; Lichrocart) employing the previously
described gradient system. A flow rate of 6 mL min-1 ensured
an elution profile similar to that of the analytical column,
and transformation products were collected using a fraction
collector (Gilson, Model 203) (24).
ampicillin. Cells were grown to an OD ≈ 0.7 at 35 °C and 200
rpm and then induced for 2 h with 100 µM IPTG.
Resting Cell Studies with P. putida F1. P. putida F1 was
harvested by centrifugation (6000 rpm, 10 min) and washed
with 50 mM Na-K phosphate buffer, pH 7.0, before being
resuspended in the same buffer to give an OD of 6.4. Resting
cell experiments were then performed by adding varying
amounts of chlorobenzene (0, 0.2, 0.4, 0.6, 0.8, 1 mM) from
100 mM stock solutions in DMSO along with 0.2% pyruvate
as energy source. Aliquots of 25 mL were added to 125 mL
serum bottles, which were then sealed with butyl rubber
stoppers and incubated in the dark at 30 °C on a rotary shaker
at 200 rpm for 2 h. Cells were then precipitated by
centrifugation, and the supernatant was subjected to reverse
phase-HPLC analysis as described below.
Gel Permeation HPLC. Gel permeation analysis was
performed as previously described (24).
NMR Spectroscopy. 1H NMR experiments were performed
using a 500 MHz - Bruker Avance 500 with the broadband
inverse probe.
The influence of competitive inhibitors of C2,3O (1-
pentanol, 2-pentanone, benzyl alcohol, and 1,10-phenan-
throline) as well as the noncompetitive inhibitor, 4-chloro-
catechol on the accumulation of catechol from 1 to 2 mM
benzene was similarly studied.
Chem icals. H2O2 (30% v/ v solution) was obtained from
Sigma (Rehovot, Israel). The concentration of stock solutions
of H2O2 was determined at 240 nm using an extinction
coefficient of 39.4 M-1cm-1 (25).
Benzene, benzyl alcohol, chlorobenzene, 4-chlorocat-
echol, 1,2-dichlorobenzene, 2-pentanone, 1,10-phenanthro-
line, and phenol were all obtained from Aldrich (Rehovot,
Israel). Catechol and 1-pentanol were obtained from Fluka
(Rehovot, Israel).
Preparative Biocatalytic Synthesis of Dead End Products
by P. putida F1. 1.5 L of toluene induced culture was equally
divided into three 1 L Erlenmeyer flasks. The cultures were
then incubated with 1 mM of chlorobenzene at 30 °C and
200 rpm for 2 h. Subsequently, cells were removed by
centrifugation at 6000 rpm for 10 min, and the supernatant
was acidified with a few drops of concentrated HCl and
extracted with ethyl acetate in a ratio of 1:5 (solvent:
supernatant). After evaporation to dryness, the products were
redissolved in 2 mL of methanol and fractionated by
semipreparative reverse phase-C18 HPLC. Peaks corre-
sponding to dead end products were collected, acidified with
a few drops of concentrated HCl, and re-extracted with ethyl
acetate (ratio 1:5). After evaporation to dryness, the products
were stored in the dark at -20 °C before being examined by
1H NMR or oxidized by HRP.
Results
Analysis by reverse phase HPLC of the cell-free supernatant
obtained after incubation of chlorobenzene with resting cells
of P. putida F1 previously induced with toluene revealed a
peak with a retention time of 11.0 min, the intensity of which
increased with increasing chlorobenzene concentration. The
1H NMR spectrum of the fractionated peak presenting
chemical shifts at H6: 6.75 ppm, 1H, dd, 8.1 and 1.6 Hz; H5:
6.62 ppm, 1H, t, 8.0 Hz; H4: 6.69 ppm, 1H, dd, 8.1 and 1.6
Hz is identical to that of 3-chlorocatechol (26).
When cell free supernatants containing 3-chlorocatechol
were incubated with HRP and varying amounts of H2O2,
reverse phase HPLC analysis revealed a gradual decrease in
the area of the peak corresponding to the 3-chlorocatechol
with increasing H2O2 concentration. In the absence of either
peroxidase or H2O2, no decrease in peak area was observed,
demonstrating that the disappearance of 3-chlorocatechol
was a result of enzymatic oxidation. The oxidation of
3-chlorocatechol by HRP was accompanied by instantaneous
formation of pink-colored products.
Whole-Cell Biotransform ations with Recom binant E.
coli JM109 (pDTG602). After induction with 100 µM IPTG
for 2 h, the pH was adjusted to 7.0 with KOH. Then, 1 mM
transformation substrate was added, and the cultures were
incubated overnight at 35 °C and 200 rpm. Cells were then
precipitated by centrifugation, and the supernatant was
subjected to reverse phase-HPLC analysis as described below.
Enzym atic Reactions. HRP, type I with an RZ (A409/ A280
)
of 1.0 and an activity of 78 U/ mg solid, was purchased from
Sigma (Rehovot, Israel), and its concentration was measured
at 403 nm using an extinction coefficient of 100 mM-1 cm-1
(21).
HRP reactions were carried out in 50 mM Na-K phosphate
buffer, pH 7.0 at 25 °C. Conditions, including enzyme,
reducing substrate, and H2O2 concentrations are given in
the figure legends.
Gel permeation analysis of the same reactions revealed
an increase in the molecular weight upon oxidation (Figure
2). As can be seen, a gradual decrease in intensity of the peak
corresponding to a molecular weight of 144 (3-chlorocat-
echol) was noticed upon addition of increasing amounts of
Reaction rates for the oxidation of catechol were measured
as previously described (22). Reactions contained 0.02 µM
9
4 7 5 4 ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 38, NO. 18, 2004

FIGURE 2. Gel permeation analysis of cell free supernatant
containing 3-chlorocatechol after incubation with 0.5 µM HRP and
varying amounts of H O in 50 mM phosphate buffer, pH 7.0. Initial
FIGURE 4. Gel permeation analysis of 2 mM catechol following
oxidation by 0.5 µM HRP and varying concentrations of H O in 50
2
2
2
2
mM phosphate buffer, pH 7.0. Initial H O concentrations were 0 (s,
2
2
H O concentrations were 0 mM (s), 0.25 mM (- -), and 0.5 mM (-
2
2
thin line), 0.25 (- -), 0.5 (- -EnDash), and 2 (s, thick line) mM. One
hour after the reactions were initiated, they were acidified to pH
2.5, frozen at -78 °C, lyophilized, redissolved in tetrahydrofuran,
and subjected to gel permeation analysis.
-). One hour after the reactions were initiated, they were acidified
to pH 2.5, frozen at -78 °C, lyophilized, redissolved in tetrahydrofuran,
and subjected to gel permeation analysis.
FIGURE 3. Influence of catechol 2,3-dioxygenase inhibitors on the
accumulation of catechol by P. putida F1 following oxidation of
benzene (1-2 mM).
H2O2, which was accompanied by the appearance of peaks
corresponding to higher molecular weight products. Those
corresponding to molecular weights of ∼250 Da and ∼400
Da could possibly be dimers and trimers of 3-chlorocatechol,
respectively. No difference was noticed if HRP was incubated
with purified 3-chlorocatechol or with cell free supernatants
containing 3-chlorocatechol, indicating that the cell free
supernatant did not contain components that interfered with
the peroxidase-catalyzed oxidation of 3-chlorocatechol.
Similar findings were witnessed when 1,2-dichloroben-
zene was added to resting cells, and the supernatant was
subsequently treated with HRP and H2O2. 1H NMR analysis
revealed the formation of 3,4-dichlorocatechol upon trans-
formation of 1,2-dichlorobenzene, which was oxidized by
HRP to form colored products accompanied by an increase
in molecular weight (data not shown). Furthermore, the
enzymes lignin peroxidase and soybean peroxidase were also
efficient at oxidizing the formed catechols.
Clearly these findings demonstrate that halogenated
aromatics can be specifically transformed into oligomeric
and possibly polymeric products in a two stage process, the
first involving dihydroxylation and dehydrogenation by TDO
and TDD of P. putida F1, respectively, resulting in the
formation of dead-end catechol products, which are sub-
sequently oxidized and polymerized in the second stage.
Another approach for transforming toxic aromatics to
catechols involves inhibiting C2,3O of P. putida F1. A range
of aromatic and aliphatic compounds are known be com-
petitive inhibitors of C2,3O (4, 10, 11). When 1-pentanol,
FIGURE 5. Photos demonstrating the fate of the polymeric products
obtained upon oxidation of 2 mM phenol and 2 mM catechol by 1
µM HRP and 5 mM H O in 50 mM phosphate buffer, pH 7.0. One
hour after the reactions were initiated, the pH was either reduced
to pH 2.0 or not adjusted (pH 7.0), and samples were subjected to
centrifugation at 14000 rpm for 5 min.
2
2
2-pentanone (K ) 0.15 mM (10)) and 1,10-phenanthroline
I
were added to resting cells at a concentration of 1 mM, little
if any catechol was detected. On the other hand, in the
presence of concentrations of between 0.2 and 1 mM of the
noncompetitive inhibitor, 4-chlorocatechol, considerably
catechol accumulated in the extracellular fluid. C2,3O is either
inactivated by the halogenated catechol itself (4) or its ring
cleavage product, an acyl chloride (5).
The most promising approach for producing catechols
involves the use of recombinant E. coli JM109(pDTG602),
which overexpresses TDO and TDD. This organism has been
shown to give high effective mass yields of catechols from
a number of aromatic compounds (7-9). We demonstrated
the effective production of 3-chlorocatechol, 3,4-dichloro-
catechol, and catechol, upon addition of either chloroben-
zene, 1,2-dichlorobenzne, or benzene, respectively, to grow-
ing cells of E. coli JM109(pDTG602) (data not shown).
Oxidation of the resulting catechol by HRP leads to formation
of polymeric products (Figure 4). The molecular weight of
the products obtained increased with increasing H2O2
concentration, and products possessing molecular weights
of up to 4000 Daltons were obtained in aqueous reactions,
without the need for water-miscible organic solvents. In
Figure 5, it is evident that the polymeric products obtained
from catechol at pH 7.0 remained in solution, whereas those
obtained from oxidation of phenol under identical conditions
clearly precipitated. The polycatechol products precipitated
only after the reaction mixture was acidified to pH 2.0.
previously shown to have a K of 0.58 mM for C2,3O (10), was
I
added at concentrations between 0.2 and 1 mM to resting
cells in the presence of 1-2 mM benzene, less than 10 µM
catechol accumulated at all concentrations of 1-pentanol
(Figure 3). In similar experiments in which the alleged
competitive inhibitors benzyl alcohol (K ) 1.4 mM (10)),
I
9
VOL. 38, NO. 18, 2004 / ENVIRONMENTAL SCIENCE & TECHNOLOGY 4 7 5 5

enable novel applications in the area of biosensors, elec-
tronics, photonics, photolithography, and rechargeable bat-
teries.
Previous reports indicate that enzymatic production of
polymers from phenols in the aqueous phase does not appear
to be practical for two main reasons (29): (i) Most phenols
are poorly soluble in water, thus necessitating working with
dilute solutions, which leads to low productivities. (ii) The
initially formed phenolic dimers and trimers are insoluble
in water and readily precipitate, preventing further polym-
erization. The findings reported here indicate that polycat-
echols exhibit high water solubility (Figures 4 and 5), a
property that could eliminate the need for the addition of
water miscible solvents to obtain the desired polymeric
products. Such solvents are detrimental not only to the
environment but also to peroxidase activity as demonstrated
here (Figure 6). The advantage is that significantly less
peroxidase is required for performing the reactions if miscible
solvents are not required. Furthermore, water soluble
polymers may be desirable for many applications.
FIGURE 6. Influence of initial substrate concentration on the rate
of oxidation of catechol by 0.02 µM HRP and 2.5 mM H O in 50 mM
2
2
phosphate buffer, pH 7.0. Inset: Influence of ethanol on the time
dependent oxidation of 2 mM catechol by 0.5 µM HRP and 6 mM
H O , monitored spectrophotometrically at 500 nm.
2
2
The apparent high water solubility of the polymers
obtained upon oxidation of catechol presumably eliminates
the need for water miscible solvents to obtain high molecular
weight polymeric products (as is the case for oxidation of
phenol). Because water miscible solvents are detrimental to
peroxidase activity, a significant increase in the amount of
enzyme is required. In aqueous solution oxidation of
catechols by peroxidase proceeds rapidly. To highlight this,
the influence of substrate concentration on the rate of
oxidation of catechol by 0.02 µM HRP and 2.5 mM H2O2 in
50 mM phosphate buffer, pH 7.0 is shown in Figure 6. The
overall rate of oxidation is significantly decreased by the
addition of small amounts of water-miscible solvents, and
the inset of Figure 6 shows the influence of ethanol on the
time dependent oxidation of 2 mM catechol by 0.5 µM HRP
and 6 mM H2O2 in 50 mM phosphate buffer, pH 7.0,
monitored spectrophotometrically at 500 nm. From the initial
slope, the presence of 20%, 30%, and 40% ethanol resulted
in a decrease of 31%, 95%, and 93% in the rate of oxidation,
respectively. Addition of ethanol at 5% or 10% had little or
no effect on product formation.
Acknowledgments
This work was funded by the Fund from the Grand Water
Research Institute. We are grateful to Martin Burger and Anat
Kiviti for their technical assistance.
Literature Cited
(1) Wackett, L. P.; Hershberger, C. D. Microbial diversity: Catabo-
lism of organic compounds is broadly distributed. In Biocatalysis
and Biodegradation: Microbial Transformation of Organic
Compounds; ASM Press: Washington, DC, 2001; pp 39-69.
(2) Rogers, J. E.; Gibson, D. T. Purification and properties of cis-
toluene dihydrodiol dehydrogenase from Pseudmonas putida.
J. Bacteriol. 1977, 130, 1117-1124.
(3) Furukawa, K.; Hirose, J.; Suyama, A.; Zaiki, T.; Hayashida, S.
Gene components responsible for discrete substrate specificity
in the metabolism of biphenyl (bph operon) and toluene (tod
operon). J. Bacteriol. 1993, 175, 5224-5232.
(4) Klecka, G. M.; Gibson, D. T. Inhibition of catechol 2,3-
dioxygenase from Pseudomonas putida by 3-chlorocatechol.
Appl. Environ. Microbiol. 1981, 41, 1159-1165.
(5) Bartels, L.; Knackmuss, H.-J.; Reineke, W. Suicide inactivation
of catechol 2,3-dioxygenase from Pseudomonas putida mt-2 by
3-halocatechols. Appl. Environ. Microbiol. 1984, 47, 500-505.
(6) Reineke, W.; Knackmuss, H.-J. Construction of haloaromatics
utilising bacteria. Nature 1979, 277, 385-386.
(7) Bui, V. P.; Hansen, T. V.; Stenstrom, Y.; Hudlicky, T. Direct
biocatalytic synthesis of functionalized catechols: a green
alternative to traditional methods with high effective mass yield.
Green Chem. 2000, 2, 263-265.
(8) Bui, V. P.; Hudlicky, T.; Hansen, T. V.; Stenstrom, Y. Direct
biooxidation of arenes to corresponding catechols with E. coli
JM109(pDTG602). Application to synthesis of combretastatins
A-1 and B-1. Tetrahedron Lett. 2002, 43, 2839-2841.
(9) Endoma, M. A.; Bui, V. P.; Hansen, J.; Hudlicky, T. Medium-
scale preparation of useful metabolites of aromatic compounds
via whole-cell fermentation with recombinant organisms. Org.
Process Res. Dev. 2002, 6, 525-532.
(10) Bertini, I.; Briganti, F.; Scozzafava, A. Aliphatic and aromatic
inhibitors binding to the active site of catechol 2,3-dioxygenase
from Pseudomonas putida mt-2. FEBS Lett. 1994, 343, 56-60.
(11) Arciero, D. M.; Orville, A. M.; Lipscomb, J. D. [17O]Water and
nitric oxide binding by protocatechuate 4,5-dioxygenase and
catechol 2,3-dioxygenase. Evidence for binding of exogenous
ligands to the active site Fe2+ of extradiol dioxygenases. J. Biol.
Chem. 1985, 14035-14044.
Discussion
The current research demonstrates the potential of employing
TDO and in turn TDD for obtaining novel catechols, which
are subsequently polymerized by peroxidase-catalyzed oxi-
dation to polycatechols in aqueous solution.
Three approaches may be adopted for obtaining cat-
echols: (1) addition of halogenated aromatics to P. putida
F1, which results in the accumulation of halogenated
catechols due to inactivation of C2,3O (4, 5); (2) inhibition
of C2,3O of P. putida F1 by known inhibitors of C2,3O (4, 10,
11); and (3) overexpression of TDO and TDD in E. coli JM109
(7-9).
Oxidation of the accumulating catechols results in the
formation of polymers. The formation of colored products
in this stage is the basis for a recently reported bioassay for
the detection of bioavailable benzene, toluene, ethyl benzene,
and xylenes (BTEX) (27). The intensity of the colored products
was correlated with the concentration of the BTEX com-
pounds. However, the formation of polymers during the HRP-
catalyzed oxidation of the catechols formed from BTEX was
overlooked. In fact, there is very little information on the
pattern of polymerization of catechols by peroxidases,
although a recent study does relate to the production of
polycatechol by peroxidase in two types of solvent systems
(28).
(12) Adam, W.; Lazarus, M.; Saha-Moller, C. R.; Weichold, O.; Hoch,
U.; Haring, D.; Schreier, P. Biotransformations with peroxidases.
Adv. Biochem. Eng./Biotechnol. 1999, 63, 74-108.
(13) May, S. W. Applications of oxidoreductases. Curr. Opin. Bio-
technol. 1999, 10, 370-375.
(14) Dordick, J.; Marletta, M. A.; Klibanov, A. M. Polymerization of
phenols catalyzed by peroxidase in nonaqueous media. Bio-
technol. Bioeng. 1987, 30, 31-36.
(15) Premachandran, R. S.; Banerjee, S.; Wu, X.-K.; John, V. T.; Akkara,
J. A.; Ayyagari, M.; Kaplan, D. L. The enzymatic synthesis of
Polyphenols synthesized by peroxidase-catalyzed reac-
tions possess electrical and optical properties due to the
conjugated backbone (14-16). These unique properties could
9
4 7 5 6 ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 38, NO. 18, 2004

fluorescent naphthol based polymers. Macromolecules 1996,
29, 6452-6460.
(16) Akkara, J. A.; Aranda, F. A.; Rao, D. V. G. L. N.; John, V. T.
Nonlinear optical properties of bioengineered materials. In
Electrical and Optical Polymer Systems; Wise, D. L., Wnek, G.
E., Trantolo, D. J., Cooper, T. M., Gresser, J. D., Eds.; Marcel
Dekker: New York, 1998; pp 453-465.
(24) Ward, G.; Hadar, Y.; Bilkis, I.; Konstantinovsky, L.; Dosoretz, C.
G. Initial steps of ferulic acid polymerization by lignin peroxidase.
J. Biol. Chem. 2001, 276, 18734-18741.
(25) Nelson, D. P.; Kiesow, L. A. Enthalpy of decomposition of
hydrogen peroxide by catalase at 25 degrees C (with molar
extinction coefficients of H2O2 solutions in the UV). Anal.
Biochem. 1972, 49, 474-478.
(17) Kobayashi, S.; Uyami, H.; Kimura, S. Enzymatic polymerization.
Chem. Rev. 2001, 101, 3793-3818.
(18) Gibson, D. T.; Koch, J. R.; Kallio, R. E. Oxidative degradation of
aromatic hydrocarbons by microorganisms. I. Enzymatic for-
mation of catechol from benzene. Biochemistry 1968, 7, 2653-
2662.
(19) Zylstra, G. J.; Gibson, D. T. Toluene degradation by Pseudomonas
putida F1: nucleotide sequence of the todC1C2BADE genes
and their expression in E. coli. J. Biol. Chem. 1989, 264, 14940-
14946.
(20) Stanier, R. Y.; Palleroni, N. J.; Doudoroff, M. The aerobic
pseudomonads; a taxonomic study. J. Gen. Microbiol. 1966, 43,
159-271.
(26) Held, M.; Suske, W.; Schmid, A.; Engesser, K.-H.; Kohler, H.-P.
E.; Witholt, B.; Wubbolts, M. G. Preparative scale production of
3-substituted catechols using a novel monooxygenase from
Pseudomonas azelaica HBP 1. J. Mol. Catal. B-Enzyme. 1998, 5,
87-93.
(27) Xu, Z.; Mulchandani, A.; Chen, W. Detection of benzene, toluene,
ethyl benzene, and xylenes (btex) using toluene dioxygenase-
peroxidase coupling reactions. Biotechnol. Prog. 2003, 19, 1812-
1815.
(28) Dubey, S.; Singh, D.; Misra, R. A. Enzymatic synthesis and various
properties of poly(catechol). Enzyme Microb. Technol. 1998,
23, 432-437.
(29) Wolnak, B.; Associates. Enzymes in commercial polymer pro-
duction: An in-depth technical and economic analysis. A
multiclient study; Report III. Specific enzymatic polymerizations;
Chicago, IL, 2001.
(21) Dunford, H. B. In Peroxidases in Chemistry and Biology; Everse,
J., Everse, K. E., Grisham, M. B., Eds.; CRC Press: Boca Raton,
FL, 1991; Vol. II, pp 1-24.
(22) Ward, G.; Hadar, Y.; Bilkis, I.; Dosoretz, C. G. Mechanistic features
of lignin peroxidase catalyzed oxidation of substituted phenols
and 1,2-dimethoxyarenes. J. Biol. Chem. 2003, 278, 39726-39734.
(23) Ward, G.; Hadar, Y.; Dosoretz, C. G. Inactivation of lignin
peroxidase during oxidation of the highly reactive substrate
ferulic acid. Enzyme Microb. Technol. 2001, 29, 34-41.
Received for review December 24, 2003. Revised manuscript
received June 11, 2004. Accepted June 30, 2004.
ES035458Q
9
VOL. 38, NO. 18, 2004 / ENVIRONMENTAL SCIENCE & TECHNOLOGY 4 7 5 7