Previous Article | Next Article 
Applied and Environmental Microbiology, March 1999, p. 1152-1160, Vol. 65, No. 3
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
Anaerobic Degradation of Phthalate Isomers by
Methanogenic Consortia
Robbert
Kleerebezem,*
Look W. Hulshoff
Pol, and
Gatze
Lettinga
Subdepartment of Environmental Technology,
Department of Agricultural, Environmental, and Systems Technology,
Wageningen Agricultural University, 6703 HD Wageningen, The
Netherlands
Received 8 June 1998/Accepted 15 December 1998
 |
ABSTRACT |
Three methanogenic enrichment cultures, grown on
ortho-phthalate, iso-phthalate, or
terephthalate were obtained from digested sewage sludge or
methanogenic granular sludge. Cultures grown on one of the
phthalate isomers were not capable of degrading the other
phthalate isomers. All three cultures had the ability to degrade
benzoate. Maximum specific growth rates (µSmax) and
biomass yields (YXtotS) of the mixed
cultures were determined by using both the phthalate
isomers and benzoate as substrates. Comparable values for these
parameters were found for all three cultures. Values for
µSmax and YXtotS were
higher for growth on benzoate compared to the phthalate isomers.
Based on measured and estimated values for the microbial yield of the
methanogens in the mixed culture, specific yields for the phthalate
and benzoate fermenting organisms were calculated. A kinetic model,
involving three microbial species, was developed to predict
intermediate acetate and hydrogen accumulation and the final production
of methane. Values for the ratio of the concentrations of methanogenic
organisms, versus the phthalate isomer and benzoate fermenting
organisms, and apparent half-saturation constants (KS) for
the methanogens were calculated. By using this combination of measured
and estimated parameter values, a reasonable description of
intermediate accumulation and methane formation was obtained,
with the initial concentration of phthalate fermenting organisms
being the only variable. The energetic efficiency for growth of
the fermenting organisms on the phthalate isomers was calculated to
be significantly smaller than for growth on benzoate.
| |
INTRODUCTION |
All three phthalic acid isomers
(ortho, meta, and para benzene
dicarboxylic acid) are produced in massive amounts around the world and are primarily anthropogenic compounds. They are used in the
chemical production of a wide range of plastics. Diesters of
ortho-phthalic acid are mainly used as a plasticizer in the production of polyvinyl chloride (10).
para-Phthalic acid (terephthalic acid) and the corresponding
dimethyl ester are used in the production of polyester fibers and
polyethylene terephthalate. This latter compound is well known
for its application in bottles for carbonated drinks.
meta-Benzene dicarboxylic acid (isophthalic acid) is
produced in smaller amounts than its ortho- and
para-oriented isomers and is used in the production of
specialty chemicals.
Introduction of phthalic acid isomers into the environment may occur
through leaching from plastics (10). In addition to this
diffuse source of environmental pollution with phthalic acids, large
point sources are generated during production of phthalate isomers
from their corresponding xylenes (2). Both liquid and solid
wastestreams, generated during production of the phthalate isomers, contain high concentrations of these aromatic acids. Furthermore, heavily contaminated soil with high concentrations of
phthalic acid isomers can be found around chemical factories (20).
Microbial activity is the principal method for the removal of phthalic
acid isomers and their corresponding esters from the environment. The
metabolic pathway for anoxic degradation of phthalic acid isomers in a
denitrifying Pseudomonas sp. involves initial activation of ortho-phthalate by coenzyme A (CoA),
probably followed by decarboxylation resulting in the formation of
benzoyl CoA (21, 22, 30). No experimental evidence is
available about the exact location of the decarboxylation in
the degradation pathway, but the current view is that benzoyl-CoA is
the only central intermediate in the anoxic mineralization of a large
range of aromatic compounds (besides aromatic compounds with at least
two phenolic functional groups). Degradation of benzoyl-CoA
proceeds through initial reduction of the aromatic ring, followed by
ring cleavage (9, 26). Hardly any information is available
about the anaerobic mineralization of phthalate isomers under
sulfate reducing or methanogenic conditions. It has been suggested that
methanogenic degradation of ortho-phthalate proceeds
analogously to degradation under denitrifying conditions because
methanogenic enrichment cultures grown on
ortho-phthalate had the ability to degrade benzoate
(26).
In methanogenic environments, complex organic matter (including
aromatic compounds) is converted into a mixture of methane and carbon
dioxide in a complex network of various metabolic groups of bacteria.
Bacteria in these consortia depend entirely on each other to perform
the metabolic conversions observed (25, 28) and are
therefore referred to as syntrophic consortia. In these syntrophic
consortia, fermentative bacteria convert complex organic matter into a
mixture of acetate and hydrogen or formate, which are
substrates for methanogenic bacteria. Fermentative
conversions of substrates are often energetically unfavorable under
standard conditions (
G0' > 0), implying the need
for continuous removal of the fermentative products by methanogens.
In this study three methanogenic enrichment cultures degrading either
ortho-phthalate, isophthalate, or
terephthalate are described. Emphasis is put on the kinetics of
mineralization of the phthalate isomers. A method is described to
calculate the kinetic parameters of the individual trophic groups
involved in mineralization of the aromatic substrates. The
specific role of benzoate in the anaerobic degradation of
terephthalate is presented in a separate study (14).
| |
MATERIALS AND METHODS |
Source of the biomass and enrichment procedure.
Biodegradability experiments were performed with three different types
of biomass and a set of aromatic compounds that are present in phthalic
acid isomer production wastewaters (13). Sludge types used
for these experiments were digested sewage sludge and two types of
granular sludge from full-scale upflow anaerobic sludge bed (UASB)
reactors. The initial concentrations of the phthalate isomers were
2.1 mM. When, after a lag phase ranging from 15 to 150 days,
mineralization of the phthalate isomers was observed, the substrate
was replenished to a concentration of 6 mM. Substrate dosage was
repeated until a phthalate isomer degradation rate of approximately
1 mM · day
1 was reached. At this point 20% of the
culture liquid was transferred into prereduced fresh medium containing
6 to 12 mM phthalate isomer. Cultivation was performed in 300-ml
serum bottles sealed with butyl rubber septa by using a liquid volume
of 70 ml. Serum bottles were incubated statically at 37°C in the
dark. To monitor degradation of the phthalate isomers, the methane
concentration in the headspace of the serum bottles was measured at
least twice a week. This procedure was used for more than 18 months in
order to obtain stable and highly enriched cultures. Best growth on
ortho-phthalate was obtained with the culture initially
seeded with granular sludge from a UASB reactor treating wastewater
from a paper factory, while cultivation on isophthalate proceeded
most rapidly with cultures initially seeded with digested sewage
sludge. These two enrichment cultures were therefore used for the
experiments described here.
The inoculum of the terephthalate degrading enrichment culture
was obtained from a laboratory-scale anaerobic hybrid reactor. Hybrid
reactors are upflow reactors equipped with carrier material in the top
of the reactor instead of the three-phase separator used in UASB
reactors. The hybrid reactor had been in continuous operation for a
period of 12 months with terephthalate as the sole carbon and
energy source. The specific terephthalate conversion rate of
the biomass grown in this reactor was approximately 1.7 mmol · (g · day)1. Similar cultivation methods were used
as described for the ortho-phthalate and
isophthalate degrading cultures for more than 1 year before the
experiments described here were carried out.
Medium and substrate preparation.
The basal medium used in
the experiments contained the following (in mg · liter of liquid
volume1): NaHCO3, 3,000; NH4Cl,
280; K2HPO4, 250; CaCl2 · 2H2O, 10; yeast extract, 18; and 1 ml of a trace element
stock solution according to the composition described by Huser et al.
(12). Stock solutions of disodium
ortho-phthalate, isophthalate, and
terephthalate were prepared in demineralized water.
Experimental procedure.
Experiments were performed in 300- or 120-ml serum bottles with liquid volumes of 70 or 25 ml,
respectively. Medium and substrate were added to the serum bottles. The
bottles were sealed with butyl rubber septa and aluminum screw or crimp
caps, and the headspace was flushed with a mixture of N2
and CO2 (70:30 [vol/vol]). After flushing,
Na2S · 7-9H2O was added from a
concentrated stock solution to obtain a final concentration of 150 mg · liter1. Serum bottles were preincubated at
37 ± 1°C in an orbital-motion shaker prior to inoculation by
syringe. Liquid samples (1 ml) were withdrawn from the serum bottles
for component analyses. Measured methane concentrations in the
headspaces of the bottles were corrected for the volumes of samples removed.
Analytical procedures.
The methane content of the headspace
was determined by gas chromatograph (Hewlett Packard 438/S). Samples
(100 µl) were injected by using a gas-lock syringe (Dynatech, Baton
Rouge, La.). A stainless-steel column (2 m by 2 mm) packed with Poropak
Q (80 to 100 mesh) was used. Nitrogen was used as a carrier gas. The
temperatures of the column, injection port, and flame ionization
detector were 60, 200, and 220°C, respectively.
The concentration of water-soluble aromatic acids was determined by
high- pressure liquid chromatography (HPLC). Centrifuged
liquid samples
(3 min at 10,000 ×
g) were diluted to concentrations
smaller than 50 mg · liter
1 by using a Meyvis
Dilutor (type no. 401), and a volume of 10
µl was injected by using
an autosampler (Marathon). Separation
of the aromatic acids was
obtained by using a Chromospher 5C18
column (100 by 3 mm). The solvent
used as a carrier was a mixture
of methanol and a 1% acetic acid
solution in water in a 40/60
ratio. The applied flow rate amounted 0.3 ml · min
1. The separated components were detected
by a UV detector (Spectroflow
773) at a wavelength of 230 nm. Typical
retention times for
ortho-phthalate,
terephthalate, isophthalate, and benzoate were 2.8, 3.6, 4.1,
and 6.8 min, respectively. Chromatograms were stored and
integrated
by using the software package
Minichrom.
The concentration and composition of volatile fatty acids in the medium
were determined with a gas chromatograph (Hewlett
Packard 5890A). A
glass column (2 m by 4 mm) packed with Supelcoport
(100 to 200 mesh),
coated with 10% Fluorad FC431, was used. The
temperatures of the
column, injection port, and flame ionization
detector were 130, 200, and 280°C, respectively. Nitrogen gas
saturated with formic acid was
used as a carrier gas at a flow
rate of 50 ml · min
1. Prior to analysis, the samples were diluted and
fixed with a
formic acid solution (3% [vol/vol]). After formic acid
addition
aromatic acids precipitate, and samples had to be centrifuged
(3 min, 10,000 ×
g).
Hydrogen was determined by gas chromatography (Hewlett Packard 5890).
The gas chromatograph was equipped with a stainless-steel
column (1.5 m
by 6.4 mm) packed with molecular sieve 25H (60 to
80 mesh). The
temperatures of the column, injection port, and
thermal
conductivity detector were 40, 110, and 125°C, respectively.
Argon was used as a carrier gas at a flow rate of 25 ml · min
1.
Determination of kinetic parameters.
The total biomass yield
of the mixed cultures growing on the phthalate isomers and benzoate
was determined in batch experiments. A known amount of biomass was
transferred into fresh medium containing a high concentration of
substrate (10 to 12 mM for benzoate and the phthalate isomers).
After complete mineralization of the substrate, the concentration of
volatile solids was determined and the biomass yield was calculated by
relating the increase of the biomass concentration to the amount of
substrate degraded. In case of degradation of a complex substrate by a
syntrophic culture, the sum of the yield factors of all the species
participating in the degradation is measured. Consequently, the
measured yield is referred to as YXtotS (g · mol-S1). For determination of the microbial
yield on acetate (YXAcMC2), several
portions of 5 mM acetate were supplied to avoid substrate inhibition.
The maximum specific growth rate (µ
Smax,
day
1) of the cultures was calculated from the exponential
part of substrate depletion
and/or product formation curves. In case of
zero-order kinetics
(C
S 
K
S), Monod-based
equations for substrate depletion and product
formation can be
integrated and the following equations apply
(neglecting maintenance
and/or decay):
|
(1)
|
|
(2)
|
where f
SP stands for the number of moles of product,
produced per mole of substrate (mol-P · mol-S
1)
according to the stoichiometry of the reaction (Table
1), and
XtotS is the electron yield as can be
calculated from the measured
yield according to equation A-4 (see
Appendix). By using measured
values for the microbial yield and
measured substrate concentrations
(C
S, mol · liter
1) and/or product concentrations (C
P,
mol · liter
1) as a function of time, the
initial biomass concentration [C
X(0),
g · liter
1] and µ
Smax can be estimated
with an optimization procedure as available
in most spreadsheet
programs. Optimization was based on minimizing
the absolute error
between measured and calculated values for
C
S and
C
P.
View this table:
[in this window]
[in a new window]
|
TABLE 1.
Chemical reaction equations for the individual steps in
mineralization of phthalate isomers and benzoate and the
standard Gibbs free-energy changes during the conversions, corrected
for a temperature of 37°C
|
|
Calculation of kinetic parameters.
In order to describe
intermediate accumulation and final methane production during growth of
the mixed cultures on the phthalate isomers and benzoate, a
mathematical model was derived (see Appendix). Nonmeasured parameter
values that were required as input for the model were estimated by
using the procedures described below.
The biomass yield of the organisms responsible for fermentation of the
phthalate isomers and benzoate
(Y
XFermS) was calculated
from the
(measured) total biomass yields (Y
XtotS)
and the biomass
yields of the acetoclastic and
hydrogenotrophic methanogens (Y
XAcM and
Y
XHyM) according to the following equation:
(3)
where f
PAC2 and f
PAH2 are, respectively,
the number of moles of acetate and of hydrogen formed per mole of
phthalate isomer
(mol-C2/H2 · mol-PA
1)
according to the chemical reaction equation for phthalate
fermentation
(Table
1, reaction 1), and
XFermPA is the electron yield of
the
phthalate isomer fermenting culture
(e-mol-X
Ferm · e-mol-PA
1).
Substitution of equation A-4 (
Appendix) into equation 1 allows
for
direct calculation of the biomass yield for the fermenting
bacteria.
Since during growth on the phthalate isomers the intermediate
concentrations of acetate and hydrogen were found to be relatively
constant (dC
C2/dt and dC
H2/dt equal 0;
Fig.
A1), the actual specific
growth rates of the different species in
the mixed culture must
be equal (
23). Definition of this
boundary condition allows
for calculation of the concentration ratio of
the acetoclastic
methanogens relative to the fermenting organisms
according to
the following equation:
|
(4)
|
From the average concentrations of acetate and hydrogen measured
during exponential growth on the phthalate isomers (see
Table
3)
and the observation that these concentrations remained
constant during
growth on the phthalate isomers, the apparent
half-saturation
constants for acetate and hydrogen (K
C2 and
K
H2)
can be estimated. At phthalate concentrations
significantly higher
than the half-saturation constant for
phthalate fermentation (K
PA),
the following
equation enables calculation of K
C2:
|
(5)
|
Expressions equivalent to equations 4 and 5 were used for
calculation of C
XFerm/ C
XHyM and
K
H2.
Gibbs free-energy changes.
Standard Gibbs
free-energy changes for the individual steps in anaerobic
mineralization of the phthalate isomers and benzoate were
calculated according to the method of Thauer et al. (31) (Table 1). Gf0 values for the phthalate
isomers were calculated from benzoate by using the group contribution
method described by Dimroth (5). Gf0 values were corrected for a temperature
of 37°C by using the Van't Hoff equation (4).
Microscopical observation.
The cultures were routinely
observed by using a phase-contrast microscope. Cultures grown on
ortho-phthalate and isophthalate were prepared for
scanning electron microscopy. Samples (10 ml) were filtered into 1-ml
observation chambers. The observation chambers were closed and fixed
with 2.5% gluteraldehyde. After fixation, samples were stored
overnight in a 0.5% osmium tetraoxide solution in a buffer of sodium
cacodylate (0.1 M, pH 7.1). After three rinses with demineralized
water, samples were dehydrated stepwise with ethanol. Samples were
mounted on stubs with carbon cement, critical point dried with
CO2, and sputter coated with 3 nm of platinum. The coated
specimens were observed in a Jeol JSM 6300F scanning microscope at 5 to
8 kV.
| |
RESULTS |
Enrichment cultures.
Three stable enrichment cultures with the
ability to degrade either ortho-phthalate,
isophthalate, or terephthalate were obtained through
numerous transfers into fresh medium throughout a period of more than 1 year. Multiple doses of substrate (6 to 12 mM phthalate isomer)
were necessary to obtain high conversion rates. Once the conversion
rate amounted to approximately 1 mM · day1, 20%
of the culture was transferred into fresh medium. No stable growth was
obtained when (i) smaller amounts of the cultures were transferred or
when (ii) the cultures were transferred when rates were significantly
lower. Moreover, when transferring the cultures after complete
depletion of the substrate, long lag phases and nonexponential growth
were observed. These observations were studied in more detail by using
the terephthalate-grown enrichment culture, and the results
obtained are described elsewhere (14). Due to their low
growth rate (Table 2), the cultures could
be transferred approximately once a month. A typical example of the
methane production profile found during several transfers of the
isophthalate-grown culture into fresh medium is shown in Fig.
1.
View this table:
[in this window]
[in a new window]
|
TABLE 2.
Maximum specific growth rate (µSmax)
and total biomass yield (YXtotS) for three
phthalate-isomer-degrading enrichment cultures, grown on the
phthalate isomers, benzoate, or acetatea
|
|
View larger version (19K):
[in this window]
[in a new window]
|
FIG. 1.
Calculated (solid lines) and measured ( ) methane
concentrations (CH4) and substrate levels (dashed lines)
during several transfers of the isophthalate (IF)-grown
culture.
|
|
Cultures grown on one of the phthalate isomers were not able to
degrade either of the other two phthalate isomers, suggesting
that specific organisms are responsible for the degradation of
each of
the phthalate isomers. All three cultures were able to
degrade
benzoate without a lag period at rates comparable to the
phthalate
isomer degradation
rates.
During exponential growth on the phthalate isomers, only small
amounts of acetate and hydrogen were found (approximately 1
to 4 mM and
3 to 5 Pa, respectively), indicating tight syntrophic
coupling between
the fermenting organisms and the methanogens
(Fig.
2). Intermediate accumulation of benzoate
was only observed
in the
ortho-phthalate-grown culture,
but the measured concentrations
were very low (1 to 3 µM). No other
intermediates were detected
during exponential growth on the
phthalate isomers.
View larger version (16K):
[in this window]
[in a new window]
|
FIG. 2.
Terephthalate (TA, ) degradation and intermediate
accumulation of acetate (C2,  ) and hydrogen (H2,  )
and final production of methane (CH4,  ). Markers
correspond to measured concentrations and lines were drawn by using the
mathematical model described in the Appendix.
|
|
Addition of Na
2SO
4 as an exogenous electron
acceptor (final concentration, 5 mM) did not affect the phthalate
isomer conversion
rate in any of the three cultures. The addition of
NaNO
3 (final
concentration, 5 mM) resulted in complete
inhibition of the degradation
of the phthalate
isomers.
Kinetic parameters.
The calculated values for the maximum
specific growth rate (µSmax) and the total yield
(YXtotS) of the three cultures are
summarized in Table 2. Values for these parameters were determined with the three phthalate isomers, with benzoate and acetate as
substrates. Both parameters were measured in exponentially growing
batch cultures.
From the data presented in Table
2 it can be seen that the differences
in µ
Smax and Y
XtotS
among the three cultures grown on either phthalate,
isophthalate, or terephthalate are small. For all
enrichment cultures,
µ
Smax and
Y
XtotS are significantly smaller for the
phthalate isomer
than for
benzoate.
When the maximum specific growth rates on acetate and the phthalate
isomers are compared, it can be seen that higher values
were found for
growth on acetate. These data suggest that only
limited amounts of
acetate will accumulate during degradation
of the phthalate
isomers. This was confirmed by the low concentrations
of acetate,
measured during exponential growth on the phthalate
isomers (1 to 4 mM), as shown in Fig.
2 for growth on
terephthalate.
The situation is different for growth on benzoate. The calculated
maximum specific growth rates of the cultures grown on benzoate
are
either comparable (isophthalate and terephthalate) or
significantly
higher (
ortho-phthalate) than the maximum
specific growth rates
determined with acetate. These parameter
values indicate that
during incubation of the
ortho-phthalate-grown culture with benzoate,
large
amounts of acetate will accumulate. Smaller amounts of acetate
are
predicted to accumulate in the isophthalate- and
terephthalate-grown
cultures when incubated with benzoate.
These indications were
confirmed by measured acetate concentrations
during benzoate degradation,
as shown in Fig.
3.
View larger version (18K):
[in this window]
[in a new window]
|
FIG. 3.
Benzoate (BA, ) degradation and concomitant acetate
(C2, ) accumulation and methane (CH4, ) formation in
ortho-phthalate (A)-, isophthalate (B)-, and
terephthalate (C)-grown cultures. Markers correspond to
measured concentrations, and lines were calculated by using the
mathematical model described in the Appendix.
|
|
Product formation during degradation of the phthalate isomers
when incubated with BES.
In order to study product formation of
the fermenting organisms, the cultures were incubated with the
phthalate isomers and 10 mM bromoethanesulfonate (BES), a specific
inhibitor of methanogenesis. From the results shown in Fig.
4 it can be seen that trace amounts of
benzoate accumulated during incubation with BES of a
terephthalate-grown culture. No benzoate was detected in
exponentially growing cultures, except for the
ortho-phthalate-grown culture, where up to 3 µM benzoate was found.
View larger version (16K):
[in this window]
[in a new window]
|
FIG. 4.
Accumulation of acetate (C2, ), benzoate (BA, ),
and molecular hydrogen (H2, ) during incubation of the
terephthalate-grown culture with terephthalate and 20 mM BES (bottom graph). The actual Gibbs free energy for
terephthalate fermentation (Table 1, reaction 1) is indicated
in the top graph.
|
|
The methanogenic substrates acetate and hydrogen accumulated to
concentrations significantly higher than those observed in
exponentially growing cultures. According to the stoichiometry
of
the fermentation of the phthalate isomers (Table
1, reaction
1),
equimolar amounts of acetate and hydrogen should accumulate
during incubation with BES. However, the results show that much
higher
concentrations of acetate accumulate. Two reasons can be
distinguished
to explain this observation: (i) small amounts of
methane were formed
during the initial 2 days of incubation, suggesting
incomplete initial
inhibition of methanogenesis from hydrogen
by BES, and (ii) the
formation of another reduced product besides
molecular hydrogen was
observed. This product was identified as
cyclohexanecarboxylate by gas
chromatography.
Based on the measured concentrations of the phthalate isomer,
acetate, and hydrogen, the actual Gibbs free energy (
G') for
fermentation of the phthalate isomers (Table
1, reaction 1) can
be
calculated. From Fig.
4 it can be seen that fermentation of
terephthalate stopped at
G' values exceeding
69 kJ
· (mol-terephthalate)
1. As for the fermentation
of terephthalate, the degradation of
ortho-phthalate and isophthalate stopped when
G'
exceeded approximately
65 kJ · (mol- phthalate
isomer)
1.
Microscopical observation.
All three phthalate
isomer-grown cultures formed dense flocs of 0.5 to 2 mm in
statically incubated serum bottles. Methanosaeta-like organisms were identified as the predominant acetoclastic
methanogen in all three cultures.
In the
ortho-phthalate-grown culture (Fig.
5A), two dominant types of organisms were
observed other than the
Methanosaeta-like
organisms (arrow
1): short fat rods (0.6 to 0.8 by 1 to 2 µm)
with rounded ends (arrow
2) and very small rods (0.3 by 1.0 to
1.2 µm) (arrow 3). The fat rods
were found in large amounts and
are presumed to be responsible for
the fermentation of
ortho-phthalate.
The very small
rods were embedded in extracellular material and
always close to the
short fat rods and may be hydrogenotrophic
methanogens belonging to the
genus
Methanobacterium.
View larger version (136K):
[in this window]
[in a new window]
|
FIG. 5.
Scanning electron micrographs of the methanogenic
enrichment cultures grown on ortho-phthalate (A) and
isophthalate (B). The numbers are explained in the text.
|
|
The isophthalate-grown culture (Fig.
5B) was less well defined than
the
ortho-phthalate-grown culture, which may partly be
the result of the low conversion rates observed in this culture
at
the moment samples were drawn for electron microscopy. Dense
clusters
of different types of organisms, embedded in extracellular
material, were observed in a loose matrix of
Methanosaeta-like
organisms.
The terephthalate-grown culture was only examined with a light
microscope (not shown) and spore-forming rods of 3 to 4 µm
were
observed besides
Methanosaeta- and
Methanospirillum-like
organisms. The spore-forming rods were
probably involved in the
fermentation of
terephthalate.
| |
DISCUSSION |
General observations.
At least three different species
of bacteria are involved in the methanogenic degradation of
the phthalate isomers: fermentative organisms that convert
the phthalate isomers to a mixture of acetate and hydrogen (Table
1, reaction 1), acetoclastic methanogens that convert acetate
into a mixture of methane and bicarbonate (Table 1, reaction 4),
and hydrogenotrophic methanogens that reduce bicarbonate with hydrogen
under formation of methane (Table 1, reaction 3). Conversion of the
phthalate isomers into acetate and hydrogen is energetically
unfavorable under standard conditions (G0' = 38.6 kJ · mol-phthalate1), suggesting that the
phthalate-isomer-fermenting cultures strictly depend on the
presence of acetoclastic and hydrogenotrophic methanogens in the mixed
culture to maintain sufficiently low intermediate concentrations of
acetate and hydrogen. Methanogens depend on the
phthalate-isomer-fermenting bacteria for generation of their substrates. Due to their mutual dependency, the mixed cultures can be
designated as syntrophic cultures (25).
As proposed for denitrifying bacteria (
21,
22,
30), the
initial step in the degradation of phthalate isomers is
suggested
to be decarboxylation to benzoate because (i) all
the phthalate-isomer-grown
cultures were capable of benzoate
degradation without a lag phase
and (ii) small amounts of
benzoate accumulated in phthalate-isomer-degrading
cultures incubated with the methanogenic inhibitor
BES.
Modelling degradation of the phthalate isomers and
benzoate.
In order to describe the dynamic formation and
consumption of acetate and hydrogen during degradation of the
phthalate isomers and benzoate, a mathematical model was developed
(see Appendix). The model was based on the reaction stoichiometries
shown in Table 1. Even though no shift in microbial population can be
excluded to occur during benzoate degradation, we assumed that one
organism was responsible for the fermentation of the phthalate
isomers and benzoate. Additional kinetic parameter values that were
required as input for the model were calculated using equations 3 to 5.
The calculated biomass yields of the phthalate- and
benzoate-fermenting organisms (equation 3) are presented in Table
3.
For the biomass yield of the
acetoclastic methanogens in the mixed
culture, the measured
values as reported in Table
2 were used.
For the biomass yield of the
hydrogenotrophic methanogens we estimated
a value of 0.33 g
· mol-H
21 from the energetic
efficiency for growth under hydrogen-limiting
conditions
of
Methanobacterium bryantii (
27). With this
estimated
value, the maximum contribution of the biomass yield of the
hydrogenotrophic
methanogens to the total biomass yield during
growth on the phthalate
isomers or benzoate amounts to only 12%,
suggesting that the errors
introduced into the calculations of the
biomass yield of the fermenting
organisms are small.
View this table:
[in this window]
[in a new window]
|
TABLE 3.
Specific biomass yields, specific phthalate- and
benzoate-fermenting activities, and the specific biomass yield values
normalized to an energy quantum of 70 kJ · mol1 of
the fermentative organisms in the three mixed cultures grown on
benzoate or the phthalate isomers
|
|
Based on measured values for the maximum specific growth rate on the
phthalate isomers and benzoate (Table
2) and the corresponding
biomass yield values of the fermenting bacteria (Table
3), the
maximum
specific conversion rates for benzoate and the phthalate
isomers
(q
XFermSmax)
were calculated (Table
3). The results indicate that the maximum
specific conversion rates for benzoate and the phthalate isomers
are in the same order of magnitude, which confirms the observation
that
the initial rates of degradation of benzoate and the phthalate
isomers are comparable, as described
above.
The observation that acetate and hydrogen concentrations remained
constant during degradation of the phthalate isomers allowed
for
calculation of (i) the biomass concentration ratios
C
XFerm/C
XAcM and
C
XFerm/C
XHyM according to equation 4 and (ii) the half-saturation
constants for the methanogenic
substrates acetate and hydrogen
(K
C2 and
K
H2) according to Equation
5. To enable calculation of
K
H2, we estimated from the data reported by Seitz et
al. (
27)
a value of 0.60 mol-H
2 · g
1 · day
1 for
q
H2max. Calculated values for these
parameters are shown in Table
4.
View this table:
[in this window]
[in a new window]
|
TABLE 4.
Calculated specific biomass ratios and apparent
half-saturation constants for acetate and hydrogen for the
methanogens in the phthalate-isomer-degrading mixed cultures
|
|
By using the measured and estimated parameter values (shown in
Tables
2 to
4), the degradation of the phthalate isomers
and
benzoate can be described with the initial concentration of
fermenting
bacteria [X
Ferm(0)] as the only variable (and estimated
values for K
BA and K
PA). When the boundary
conditions are sufficiently
fulfilled, the derived model accurately
describes the accumulation
of acetate and hydrogen during the
degradation of the phthalate
isomers, as shown for
terephthalate degradation in Fig.
2.
The assumptions made to calculate the biomass ratios and the
half-saturation constants for the methanogens (Table
4) were
based on
measurements with the phthalate isomers as a substrate.
To validate
the model, intermediate accumulation and product formation
during
benzoate degradation were calculated. From the results
shown in Fig.
4
it can be seen that the intermediate formation
of acetate and final
production of methane are reasonably well
predicted for benzoate
degradation. If it is taken into account
that minor errors in,
for example, the measured values for
Y
XAcMC2 lead to large
differences in the estimated biomass ratio, the
results are
satisfactory.
In summary, we suggest that the developed three-species
model adequately describes the intermediate accumulation of acetate
and
hydrogen and the final production of methane during degradation
of the
phthalate isomers and benzoate. Description of the conversions
observed as a function of the kinetic properties of the individual
trophic groups in the mixed culture provides additional insight
into its metabolic
properties.
Kinetic parameter values.
Measured and estimated values of the
kinetic parameters for benzoate and acetate degradation
correspond well with previously reported values. A growth
yield on benzoate of 8.5 g · mol-benzoate1 has
been reported for strain BZ-2 cocultured with
Methanospirillum sp. strain PM-1 (assuming a protein content
of 60%) (6) and of 6.2 and 8.2 g · mol-benzoate1 for Syntrophus buswellii GA
cocultured with Methanospirillum hungatei or
Desulfovibrio sp., respectively (1). From our
data we calculated a similar average value of 9.3 ± 0.5 g · mol-benzoate1 if the growth of acetoclastic methanogens
was omitted from the calculation.
Maximum specific growth rates reported for two different
Syntrophus buswellii strains, cocultured with either
M. hungatei or
Desulfovibrio sp., are 0.10 to
0.29 and 0.17 to 0.37 day
1, respectively (
19,
32). The average value of 0.17 ± 0.04
day
1 we
obtained for the maximum growth rate on benzoate in our cultures
is in
the same order of
magnitude.
Methanosaeta-like organisms were observed in all three
phthalate isomers degrading mixed cultures by microscopical
observation.
The biomass yield, half-saturation constant, and maximum
growth
rate reported in the literature for
Methanosaeta
soehngenii grown
on acetate at 37°C are 1.47 g · mol-acetate
1, 0.47 mM, and 0.11 day
1,
respectively (
33). These values are in the same order of
magnitude
as the average values we measured or calculated for our
cultures
(Y
XC2 = 1.15 ± 0.28 g · mol-acetate
1, K
C2 = 0.55 ± 0.13 mM and
µ
C2max = 0.12 ± 0.02 day
1).
In the literature no kinetic parameter values for methanogenic cultures
degrading one of the phthalate isomers were found.
The maximum
growth rates we calculated for the phthalate isomers
(µ
PAmax) were low: 0.091 ± 0.003 day
1. Comparable values have been found for methanogenic
enrichment
cultures degrading the poorly degradable substrates toluene
and
ortho-xylene (0.11 and 0.07 day
1,
respectively [
7]).
Energetic efficiency for growth on the phthalate isomers and
benzoate.
Correlations between thermodynamics and biomass yields
have been developed by several researchers (11, 17, 29).
Stouthamer (29) stated that the biomass yield of anaerobic
bacteria is in the range of 5 to 12 g · mol-ATP1, with an average value of 10.5 g · mol-ATP1. Under physiological conditions, an
average amount of 70 kJ · mol-ATP1 was estimated
to be needed for irreversible ATP synthesis (25). With
average values for the substrate and product concentrations for
fermentation of the phthalate isomers and benzoate, the Gibbs free
energy change of the fermentations and the biomass yield normalized to
an energy quantum of 70 kJ · mol1
(YXFerm
G) can be calculated from the
measured specific biomass yields
(YXFermS), as shown in Table 3.
The average value we calculated for
Y
XFermG for benzoate fermentation is
11.2 ± 1.9 g · 70-kJ
1, which is close
to the average value of 10.5 g · mol-ATP
1 reported by
Stouthamer (
29). Values for
Y
XFermG for fermentation
of the
phthalate isomers are much lower (3.3 ± 0.3 g · 70-kJ
1), suggesting that the energetic
efficiency for growth on the
phthalate isomers is much lower
compared to benzoate. The observation
that no degradation of the
phthalate isomers was observed at Gibbs
free-energy changes
exceeding
65 kJ · mol
1 confirms that an
energetic inefficiency in degradation of the
phthalate isomers may
exist. Assuming that the phthalate isomers
and benzoate are
degraded by the same organism and that benzoate
is the first
intermediate in the degradation of the phthalate
isomers, the
postulated energetic inefficiency during growth on
the
phthalate isomers should manifest within the initial steps
of
the phthalate isomer
degradation.
A possible reason for the postulated energetic inefficiency
in fermentation of the phthalate isomers can be found in
the mechanism
of substrate transport across the microbial
membrane. The pK
a1,2 values for the phthalate isomers
(pK
a1,2 for
ortho-, iso-, and
terephthalate are 3.0, 3.6, and 3.5, respectively) are
considerably
lower than the pK
a value of 4.2 for
benzoate (
8). To enable
comparable uptake rates for the
phthalate isomers and benzoate
as suggested by the comparable
values for
q
XFermPAmax and
q
XFermBAmax
(Table
3), active uptake under expense of Gibbs
free-energy
may be required for the phthalate isomers,
whereas no (or less)
energy is required for the uptake of
benzoate.
Another explanation for the postulated energetic inefficiency can be
found in the decarboxylation of the phthalates to benzoate
(or
their CoA analogues). Taylor and Ribbons (
30) suggested
that
decarboxylation of phthalate may proceed after the initial
partial
reduction of the aromatic ring (in a one- or two-electron
mechanism),
followed by oxidative decarboxylation. The initial
reduction of
the aromatic ring is endergonic and requires the
investment of energy
in the form of ATP (
3). If this amount
of energy cannot (or
can only partially) be regained during oxidative
decarboxylation, this
may explain the net energy consumption during
decarboxylation of the
phthalate
isomers.
In summary, it is postulated that the calculated low energetic
efficiency for growth on the phthalate isomers can be due to
the
need for energy consumption (i) for active uptake of the phthalate
isomers across the microbial membrane or (ii) to initiate the
decarboxylation of the phthalate isomers to
benzoate.
| |
APPENDIX |
A mathematical model was developed for predicting the
intermediate accumulation of acetate and hydrogen and for production of
methane during degradation of the phthalate isomers and benzoate. The model is based on Monod kinetics (18) and, consequently, the volumetric rate of substrate consumption (A-1), biomass growth (A-2), and product formation (A-3) in a batch reactor can be described by using the following equations (neglecting maintenance and/or decay):
|
(A-1)
|
|
(A-2)
|
|
(A-3)
|
where f
SP stands for the number of moles of product
formed per mole of substrate according to the chemical reaction
equation
mol-S · mol-P
1 and where
XS is the biomass yield for growth of biomass X on
substrate S, expressed as electron yield (e-mol-X · e-mol-S
1). This parameter can be calculated from the
measured (or calculated)
biomass yield according to the following
equation:
|
(A-4)
|
where
is the degree of reduction (e-mol · C-mol
1) as defined by Heijnen et al. (
11),
C-length is the carbon chain length
(C-mol · mol
1), and MW is the molecular weight
(g · mol
1). We used an average biomass composition of
C
4H
7.2O
2N
0.8 (
11,
16) and NH
4+ as the nitrogen source. By
using these kinetic equations and
mass balances based on the
stoichiometry of the conversion reactions
shown in Table
1,
differential equations for all substrates,
products, and different
types of biomass were derived (Fig.
A1).
During the derivation of these
equations the following assumptions
were made: (i) the phthalate
isomers and benzoate are fermented
by the same organism
(X
Ferm); (ii) kinetic parameters for fermentation
of the
phthalate isomers and benzoate may be different; and (iii)
the
ratio of fermentation products equals the ratio predicted
by the
chemical conversion reactions described in Table
1 and
is therefore
independent of biomass growth.
View larger version (15K):
[in this window]
[in a new window]
|
FIG. A1.
Differential equations describing terephthalate
and benzoate degradation with concomitant production and consumption of
acetate and hydrogen and production of methane.
|
|
The differential equation describing the change in hydrogen
concentration over time is expressed in atm · (liter · day)1 by correction of the molar rates for the liquid
volume (Vliquid, liter), the headspace volume
(Vg, liter), of the serum bottle, and the volume of 1 mole
of gas (Vmg, liter · mol1). To include
the observed threshold concentrations of hydrogen in the model, an
additional term was included in the differential equation describing
hydrogen consumption due to hydrogenotrophic methanogenesis. This
term is based on reversible enzyme kinetics as described by Labib
et al. (15) and avoids hydrogen consumption if
hydrogenotrophic methanogenesis is endergonic
(G'H2ox > 0). No product inhibition terms for the
fermentation of the phthalate isomers and benzoate were included in
the model because in general only small amounts of acetate and hydrogen
accumulated during our experiments.
The differential equations A-5 to A-12 (Fig. A1) were integrated
by using a fourth-order Runge-Kutta algorithm with adaptive step-size control (24). Optimization of parameter values or initial conditions was performed by using the downhill
simplex method in multidimensions as described by Press et al.
(24). Optimization was based on minimizing the
absolute error between measured and calculated values for the
individual concentrations. The software program incorporating the
model was written in Turbo Pascal 6.0 on a personal computer
(80486-DX33).
An example of the model is shown in Fig.
A2 for the degradation of
terephthalate and benzoate. This figure clearly demonstrates the differences in intermediate accumulation during
terephthalate and benzoate degradation. The parameter values
used for calculating Fig. A2 are presented in Tables 2 to 4.
View larger version (19K):
[in this window]
[in a new window]
|
FIG. A2.
Terephthalate (TA) and benzoate (BA) degradation
and concomitant intermediate accumulation of acetate (C2) and hydrogen
(H2) and final production of methane (CH4) as
predicted by the model.
|
|
| |
ACKNOWLEDGMENTS |
We thank A. E. van Aelst and G. Gonzalez for help with
electron microscopy. R.K. wishes to thank Hervé Macarie and
Alfons J. M. Stams for critical review of the manuscript.
This project was supported through IOP Milieubiotechnologie
(Innovative Research Program Environmental Biotechnology, The Hague, The Netherlands).
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Wageningen
Agricultural University, Department of Agricultural, Environmental and
Systems Technology, Subdepartment of Environmental Technology,
"Biotechnion" Bomenweg 2, 6703 HD Wageningen, The Netherlands.
Phone: (31-317) 483798. Fax: (31-317) 482108. E-mail:
robbert.kleerebezem{at}algemeen.mt.wau.nl.
| |
REFERENCES |
| 1.
|
Auburger, G., and J. Winter.
1995.
Isolation and physiological characterization of Syntrophus buswellii strain GA from a syntrophic benzoate-degrading, strictly anaerobic coculture.
Appl. Microbiol. Biotechnol.
44:241-248.
|
| 2.
|
Bemis, A. G.,
J. A. Dindorf,
B. Horwood, and C. Samans.
1982.
Phthalic acids and other benzenepolycarboxylic acids, p. 732-777.
In
H. F. Mark, D. F. Othmer, C. G. Overberg, G. T. Seaborg, M. Grayson, and D. Eckroth (ed.), Kirk Othmer encyclopedia of chemical technology, vol. 17. John Wiley & Sons, New York, N.Y.
|
| 3.
|
Boll, M., and G. Fuchs.
1995.
Benzoyl-coenzyme A reductase (dearomatising), a key enzyme of anaerobic aromatic metabolism. ATP dependence of the reaction, purification and some properties of the enzyme from Thauera aromatica strain K172.
Eur. J. Biochem.
234:921-933[Medline].
|
| 4.
|
Conrad, R., and B. Wetter.
1990.
Influence of temperature on energetics of hydrogen metabolism in homoacetogenic, methanogenic, and other bacteria.
Arch. Microbiol.
155:94-98.
|
| 5.
|
Dimroth, K.
1983.
Thermochemische daten organischer verbindungen, p. 997-1038.
In
D'ans-Lax, Taschenbuch fur chemiker und Physiker, vol. 2. Springer-Verlag, Berlin, Germany.
|
| 6.
|
Dolfing, J., and J. M. Tiedje.
1988.
Acetate inhibition of methanogenic, syntrophic benzoate degradation.
Appl. Environ. Microbiol.
54:1871-1873[Abstract/Free Full Text].
|
| 7.
|
Edwards, E. A., and D. Grbic-Galic.
1994.
Anaerobic degradation of toluene and o-xylene by a methanogenic consortium.
Appl. Environ. Microbiol.
60:313-322[Abstract/Free Full Text].
|
| 8.
|
Fieser, L. F., and M. Fieser.
1956.
Organic chemistry, 3rd ed.
Reinhold Publishing Corp., New York, N.Y.
|
| 9.
|
Fuchs, G.,
M. E. S. Mohamed,
U. Altenschmidt,
J. Koch,
A. Lack,
R. Brackmann,
C. Lochmeyer, and B. Oswald.
1994.
Biochemistry of anaerobic biodegradation of aromatic compounds, p. 513-553.
In
C. Ratledge (ed.), Biochemistry of microbial degradation. Kluwer Academic Publishers, Dordrecht, The Netherlands.
|
| 10.
|
Giam, C. S.,
E. Atlas,
M. A. Powers, Jr., and J. E. Leonard.
1984.
Phthalic acid esters, p. 67-140.
In
O. Hutzinger (ed.), Anthropogenic compounds, vol. 3, part C. Springer-Verlag, Berlin, Germany.
|
| 11.
|
Heijnen, J. J.,
M. C. M. van Loosdrecht, and L. Tijhuis.
1992.
A black box mathematical model to calculate auto- and heterotrophic biomass yields based on Gibbs energy dissipation.
Biotechnol. Bioeng.
40:1139-1154.
|
| 12.
|
Huser, B. A.,
K. Wuhrmann, and A. B. J. Zehnder.
1980.
Methanothrix soehngenii gen. nov. sp. nov., a new acetotrophic non-hydrogen-oxidizing methane bacterium.
Arch. Microbiol.
132:1-9.
|
| 13.
| Kleerebezem, R., L. W. Hulshoff Pol, and G. Lettinga. Anaerobic biodegradability of aromatic acids and esters.
Biodegradation, in press.
|
| 14.
|
Kleerebezem, R.,
L. W. Hulshoff Pol, and G. Lettinga.
1999.
The role of benzoate in anaerobic degradation of terephthalate.
Appl. Environ. Microbiol.
65:1161-1167[Abstract/Free Full Text].
|
| 15.
|
Labib, F.,
J. F. Ferguson,
M. M. Benjamin,
M. Merigh, and N. L. Ricker.
1993.
Mathematical modelling of an anaerobic butyrate degrading consortium: predicting their response to organic overloading.
Environ. Sci. Technol.
27:2673-2683.
|
| 16.
|
McCarty, P. L.
1971.
Energetics and bacterial growth, p. 495-512.
In
S. D. Faust, and J. V. Hunter (ed.), Organic compounds in aquatic environments. Marcel Dekker, Inc., New York, N.Y.
|
| 17.
|
McCarty, P. L.
1972.
Energetics of organic matter degradation, p. 91-118.
In
R. Mitchell (ed.), Water pollution microbiology. John Wiley & Sons, New York, N.Y.
|
| 18.
|
Monod, J.
1949.
The growth of bacterial cultures.
Annu. Rev. Microbiol.
3:371-394.
|
| 19.
|
Mountfort, D. O.,
W. J. Brulla,
L. R. Krumholz, and M. P. Bryant.
1984.
Syntrophus buswellii gen. nov., sp. nov.: a benzoate catabolizer from methanogenic ecosystems.
Int. J. Syst. Bacteriol.
34:216-217[Abstract/Free Full Text].
|
| 20.
|
Naumov, A. V.,
A. B. Gafarov, and A. M. Boronin.
1996.
A novel ortho-phthalate utilizer an acetic acid bacterium Acetobacter sp.
Microbiology
65:182-186.
|
| 21.
|
Nozawa, T., and Y. Maruyama.
1988.
Anaerobic metabolism of phthalate and other aromatic compounds by a denitrifying bacterium.
J. Bacteriol.
170:5778-5784[Abstract/Free Full Text].
|
| 22.
|
Nozawa, T., and Y. Maruyama.
1988.
Denitrification by a soil bacterium with phthalate and aromatic compounds as substrates.
J. Bacteriol.
170:2501-2505[Abstract/Free Full Text].
|
| 23.
|
Powell, G. E.
1984.
Equalisation of specific growth rates for syntrophic associations in batch cultures.
J. Chem. Technol. Biotechnol.
34B:97-100.
|
| 24.
|
Press, W. H.,
B. P. Flannery,
S. A. Teukolsky, and W. T. Vetterling.
1989.
Numerical recipes in Pascal: the art of scientific computing.
Cambridge University Press, Cambridge, United Kingdom.
|
| 25.
|
Schink, B.
1992.
Syntrophism among prokaryotes, p. 276-299.
In
A. Balows, H. G. Trüper, M. Dworkin, W. Harder, and K. H. Schleifer (ed.), The prokaryotes, vol. I. Springer-Verlag, New York, N.Y.
|
| 26.
|
Schink, B.,
A. Brune, and S. Schnell.
1992.
Anaerobic degradation of aromatic compounds, p. 219-242.
In
G. Winkelmann (ed.), Microbial degradation of natural products. VCH, Weinheim, Germany.
|
| 27.
|
Seitz, H. J.,
B. Schink,
N. Pfennig, and R. Conrad.
1990.
Energetics of syntrophic ethanol oxidation in defined chemostat cocultures: energy sharing in biomass production (2).
Arch. Microbiol.
155:89-93.
|
| 28.
|
Stams, A. J. M.
1994.
Metabolic interactions between anaerobic bacteria in methanogenic environments.
Antonie Leeuwenhoek
66:271-294[Medline].
|
| 29.
|
Stouthamer, A. H.
1979.
The search for correlation between theoretical and experimental growth yields.
Int. Rev. Biochem.
21:1-47.
|
| 30.
|
Taylor, B. F., and D. W. Ribbons.
1983.
Bacterial decarboxylation of o-phthalic acids.
Appl. Environ. Microbiol.
46:1276-1281[Abstract/Free Full Text].
|
| 31.
|
Thauer, R. K.,
K. Jungermann, and K. Decker.
1977.
Energy conservation in chemotrophic anaerobic bacteria.
Bacteriol. Rev.
41:100-179[Free Full Text].
|
| 32.
|
Wallrabenstein, C., and B. Schink.
1994.
Evidence of reversed electron transport in syntrophic butyrate or benzoate oxidation by Syntrophomonas wolfei and Syntrophus buswellii.
Arch. Microbiol.
162:136-142.
|
| 33.
|
Zehnder, J. B.,
B. A. Huser,
T. D. Brock, and K. Wuhrmann.
1980.
Characterization of an acetate-decarboxylating, non-hydrogen-oxidizing methane bacterium.
Arch. Microbiol.
124:1-11[Medline].
|
Applied and Environmental Microbiology, March 1999, p. 1152-1160, Vol. 65, No. 3
0099-2240/99/$04.00+0
Copyright © 1999, American Society for Microbiology. All rights reserved.
This article has been cited by other articles:
-
Qiu, Y.-L., Hanada, S., Ohashi, A., Harada, H., Kamagata, Y., Sekiguchi, Y.
(2008). Syntrophorhabdus aromaticivorans gen. nov., sp. nov., the First Cultured Anaerobe Capable of Degrading Phenol to Acetate in Obligate Syntrophic Associations with a Hydrogenotrophic Methanogen. Appl. Environ. Microbiol.
74: 2051-2058
[Abstract]
[Full Text]
-
Chen, C.-L., Macarie, H., Ramirez, I., Olmos, A., Ong, S. L., Monroy, O., Liu, W.-T.
(2004). Microbial community structure in a thermophilic anaerobic hybrid reactor degrading terephthalate. Microbiology
150: 3429-3440
[Abstract]
[Full Text]
-
Qiu, Y.-L., Sekiguchi, Y., Imachi, H., Kamagata, Y., Tseng, I-C., Cheng, S.-S., Ohashi, A., Harada, H.
(2004). Identification and Isolation of Anaerobic, Syntrophic Phthalate Isomer-Degrading Microbes from Methanogenic Sludges Treating Wastewater from Terephthalate Manufacturing. Appl. Environ. Microbiol.
70: 1617-1626
[Abstract]
[Full Text]
-
Voget, S., Leggewie, C., Uesbeck, A., Raasch, C., Jaeger, K.-E., Streit, W. R.
(2003). Prospecting for Novel Biocatalysts in a Soil Metagenome. Appl. Environ. Microbiol.
69: 6235-6242
[Abstract]
[Full Text]
-
Wu, J.-H., Liu, W.-T., Tseng, I-C., Cheng, S.-S.
(2001). Characterization of microbial consortia in a terephthalate-degrading anaerobic granular sludge system. Microbiology
147: 373-382
[Abstract]
[Full Text]
-
Entcheva, P., Liebl, W., Johann, A., Hartsch, T., Streit, W. R.
(2001). Direct Cloning from Enrichment Cultures, a Reliable Strategy for Isolation of Complete Operons and Genes from Microbial Consortia. Appl. Environ. Microbiol.
67: 89-99
[Abstract]
[Full Text]
-
Scholten, J. C. M., Conrad, R.
(2000). Energetics of Syntrophic Propionate Oxidation in Defined Batch and Chemostat Cocultures. Appl. Environ. Microbiol.
66: 2934-2942
[Abstract]
[Full Text]
-
Kleerebezem, R., Pol, L. W. H., Lettinga, G.
(1999). The Role of Benzoate in Anaerobic Degradation of Terephthalate. Appl. Environ. Microbiol.
65: 1161-1167
[Abstract]
[Full Text]
Top
Abstract
Introduction
