Previous Article | Next Article 
Appl Environ Microbiol, April 1998, p. 1203-1209, Vol. 64, No. 4
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Population Dynamics of Phenol-Degrading Bacteria in
Activated Sludge Determined by gyrB-Targeted
Quantitative PCR
Kazuya
Watanabe,1,2,*
Satoshi
Yamamoto,2
Sanae
Hino,1 and
Shigeaki
Harayama2
Corporate Research and Development
Laboratories, Tonen Corporation, Ohi-machi, Iruma-gun,
Saitama,1 and
Marine Biotechnology
Institute, Kamaishi Laboratories, Kamaishi City, Iwate
026,2 Japan
Received 29 September 1997/Accepted 15 January 1998
 |
ABSTRACT |
A method for quantifying bacterial populations introduced into an
activated-sludge microbial community is described. The method involves
extraction of DNA from activated sludge, appropriate dilution of the
extracted DNA with DNA extracted from nonintroduced activated sludge,
PCR amplification of a gyrB gene fragment from the
introduced strain with a set of strain-specific primers, and quantification of the electrophoresed PCR product by densitometry. The
adequacy of the method was examined by analyzing the population dynamics of two phenol-degrading bacteria, Pseudomonas
putida BH and Comamonas sp. strain E6, that had been
introduced into phenol-digesting activated sludge. The density of each
of the two populations determined by the PCR method immediately after the introduction was consistent with the density estimated from a plate
count of the inoculum. This quantitative PCR method revealed different
population dynamics for the two strains in the activated sludge under
different phenol-loading conditions. The behavior of both of these
strains in the activated sludge reflected the growth kinetics of the
strains determined in laboratory axenic cultures.
| |
INTRODUCTION |
Bioaugmentation is a method for
enhancing in situ pollutant biodegradation by introducing exogenous
microorganisms with the desired catabolic traits. This method is
considered useful when effective pollutant-degrading populations are
not present at a polluted site. Successful bioaugmentation requires
that strains suitable for each polluted site be selected; for example,
at a highly contaminated site, a strain capable of rapidly degrading the pollutant present at a high concentration may be necessary, whereas
for thorough degradation a strain with high affinity for the pollutant
may be required. In this respect, it is desirable to establish
laboratory evaluation methods for selecting the strains to be
introduced. It has been found that phenol-degrading bacteria can be
classified into several kinetically different groups (46), and competition between two phenol-degrading bacteria with different growth kinetics has been investigated in an axenic sequencing fed-batch reactor (10). Similarly, the growth kinetics of
2,4-dichlorophenoxyacetic acid-degrading bacteria have been compared in
order to select strains suitable for bioaugmentation (15).
However, it has not yet been clarified whether laboratory data can be
extrapolated to predict and explain the growth and activity of strains
introduced into the environment. Ka et al. (22) analyzed the
competition among 2,4-dichlorophenoxyacetic acid-degrading bacteria in
soil and suggested that the lag time for growth of these strains
observed in laboratory batch culture experiments, rather than the
specific growth rate, is the principal determinant for competitiveness in soil.
To further investigate the relationships between the growth properties
of bacteria in test tubes and their behaviors in the natural
environment, quantitative monitoring of bacteria introduced into the
environment is a prerequisite. Selective plating has been used most
widely for this purpose (12, 29, 30, 32); however, this
method requires specific conditions under which only the introduced
strain can grow, so that it cannot be used to detect anonymous natural
isolates. In addition, it has recently been suggested that the plate
count method fails to detect bacteria that become unculturable (but are
still active) in response to environmental stress (20, 24,
34). Immunological methods have been used to detect nitrifying
Nitrosomonas populations (36) and
phenol-digesting populations (47, 48) in activated-sludge samples. Unfortunately, the sensitivity of these methods was relatively low; the detection limits were between 106 and
107 cells per ml. The use of gene probes in combination
with hybridization and/or PCR is a more attractive method, because it
provides higher specificity and higher sensitivity (33, 37,
41), although the success of this method when it is used for
specific detection is highly dependent on the specificity of the
nucleotide sequences used as the probes. The 16S rRNA sequence has been
the most commonly used sequence (6, 33); this sequence has
been used successfully to analyze overall bacterial community
structures at the genus level (5, 25, 44, 51). The DNA
sequences of genes encoding catabolic enzymes have also been used in
many cases (4, 19, 33); however, horizontal transfer of
these genes, either on the plasmid (9) or on the chromosome
(27) of the introduced strains, to indigenous populations
has been observed, especially when the environment contained substrates
for the catabolic genes. DNA fragments amplified by repetitive
sequence-based PCR have recently been used as strain-specific DNA
probes (28), but this method is thought to be somewhat
laborious. Use of the DNA sequence of the gyrB gene, in
combination with PCR, has been proposed as a method which could be used
for specific detection of bacterial strains in the natural environment
(49). The gyrB gene encodes the subunit B protein
of DNA gyrase (topoisomerase type II) (31). It has been
claimed that an advantage of using the gyrB sequence as a
strain-specific probe is the higher molecular evolution rate of the
gyrB gene and thus the greater diversity in the sequence of
this gene compared with the 16S rRNA sequence (16, 49). Thus, we decided to examine the use of the gyrB sequence for
specific detection of strains in a complex microbial community, such as activated sludge.
An aim of this study was to develop a gyrB-targeted
quantitative PCR method for analyzing the population dynamics of
bacterial strains introduced into an activated-sludge microbial
community. Quantitative PCR techniques have been shown to suffer from a
number of practical difficulties, including the different efficiencies of PCR amplification in different samples and the narrow linear response range (8). Hence, several modified forms of PCR,
such as a PCR coupled to limiting dilution (42) and a
competitive PCR (11, 23, 24), have been developed for
enumerating bacterial populations. However, these methods are laborious
and have resulted in only limited applications to analyses of
population dynamics (24). In this paper, we describe a less
laborious quantitative PCR method that revealed the different
population dynamics of two phenol-degrading strains introduced into
activated sludge under different phenol-loading conditions.
| |
MATERIALS AND METHODS |
Bacterial strains and culture conditions.
A phenol-degrading
strain, Pseudomonas putida BH, was isolated previously from
activated sludge after enrichment in batch cultures (17),
while Comamonas sp. strain E6 (previously identified as
Pseudomonas sp.) was obtained from another activated sludge after enrichment in a continuous culture (45). These two
strains were maintained on Luria-Bertani (LB) agar plates
(35) and were stored at 
80°C in the presence of 15%
(wt/vol) glycerol. The cell numbers of these strains in pure cultures
were determined by using LB plates. The synthetic media used were based
on MP medium containing (per liter) 2.75 g of
K2HPO4, 2.25 g of
KH2PO4, 1.0 g of
(NH4)2SO4, 0.2 g of
MgCl2 · 6H2O, 0.1 g of NaCl,
0.02 g of FeCl3 · 6H2O, and
0.01 g of CaCl2; the pH of this medium was between 6.8 and 7.0.
Other strains of P. putida, including strains A10L
(39), FK715 (13), IFO14164 (40),
IFO14671 (1), and JCM6156 (2), Comamonas
terrigena IAM12052 (7), Comamonas
testosteroni IAM12419 (7), and Comamonas
acidovorans IFO13582 (14) were grown in LB medium at
30°C.
Sequencing of the gyrB gene.
Total DNA of each
bacterium was extracted by the Marmur procedure (26). The
gyrB gene was amplified from the extracted DNA by PCR with
primers UP-1 and UP-2r (49). The amplified product was
purified after electrophoresis on 1% low-melting-temperature agarose
(NucSieve GTG; FMC Bioproducts, Vallensbaek Strand, Denmark) as
described by Sambrook et al. (35). The sequence of each
amplified fragment was determined with a DNA sequencing kit (dye
terminator cycle sequencing kit; Perkin-Elmer, Foster City, Calif.) and
a model 373A DNA sequencer (Applied Biosystems, Foster City, Calif.) by
using the manufacturers' instructions.
Activated-sludge samples.
Activated-sludge samples were
obtained from two municipal sewage treatment plants, the Takinoshita
plant (Kawagoe, Saitama, Japan) in September 1996 and the Ohdaira plant
(Kamaishi, Iwate, Japan) in July 1997.
Laboratory activated-sludge experiments.
An activated-sludge
process was simulated in a laboratory unit (Miyamoto Corp., Osaka,
Japan) composed of an aeration tank (working volume, 3 liters) and a
settling tank (working volume, 2 liters). The laboratory unit was
inoculated with approximately 7 g (dry weight) of the activated
sludge obtained from the Takinoshita plant, and MP medium supplemented
with 200 ppm of phenol was continuously supplied at a flow rate of 6 liters per day. The phenol-loading rate was calculated to be 0.4 g
per liter per day. The mixed-liquor suspended solid (MLSS) content was
maintained between 1,800 and 2,000 ppm by discarding the excess sludge
from the aeration tank. The mean sludge residence time was calculated
to be approximately 10 days. Air was constantly supplied at a rate of 2 liters per min, and the temperature was maintained at approximately
25°C. The MLSS content was measured by weighing the dried sludge on a
0.22-µm-pore-size filter by using Japan Industrial Standards method
K0102 (21). The total cell count in the activated sludge was
determined by a fluorescent-microscopy method with acridine orange as
described previously (48). The phenol concentration in the
aeration tank was measured by a colorimetric assay with Phenol Test
Wako (Wako Pure Chemicals, Osaka, Japan) as described previously
(48).
Inocula added to the phenol-digesting activated sludge were prepared by
cultivating strains BH and E6 at 30°C in 500 ml of LB medium. Cells
in the late exponential growth phase were harvested by centrifugating
at 10,000 × g for 5 min and were then washed with 500 ml of MP medium. The cells were resuspended in 50 ml of MP medium and
used as inocula for the activated sludge in the laboratory unit.
To provide a shock load of phenol, MP medium supplemented with 5,000 ppm of phenol was supplied to the phenol-acclimated activated
sludge at
a flow rate of 6 liters per day (the phenol-loading
rate was calculated
to be 10 g per liter per day).
Extraction of DNA from activated sludge.
Five milliliters of
an activated-sludge suspension was mixed with 0.5 ml of 50 mM sodium
tripolyphosphate. In order to deflocculate the activated sludge, the
mixture was treated in a blender (Wheaton Instruments, Millville, N.J.)
for 2 min. The suspension was then centrifuged at 15,000 × g for 5 min, and the precipitate was resuspended in 0.5 ml
of a cell-suspending buffer (10 mM Tris-HCl [pH 8.0], 1 mM EDTA, 0.35 M sucrose, and 20 mg of lysozyme per ml). After incubation for 10 min
at 37°C, 0.75 ml of a lysing solution (100 mM Tris-HCl [pH 8.0],
0.3 M NaCl, 20 mM EDTA, 2% [wt/vol] sodium dodecyl sulfate, and 2%
[wt/vol] 2-mercaptoethanol) was added, and the suspension was
incubated at 55°C for an additional 30 min. Next, the suspension was
extracted four times with a phenol-chloroform solution (35),
and 0.8 ml of the aqueous solution was recovered. Then, 0.8 ml of
2-propanol was added to the aqueous solution, and after gently mixing,
the solution was incubated at 20°C for 30 min. Nucleic acids were
precipitated by centrifugation at 20,000 × g for 10 min, and, after the preparation was washed with 1 ml of an 80% ethanol
solution, the nucleic acids were dissolved in 0.5 ml of TE buffer
(35) containing 100 µg of RNase A. This solution was
gently shaken at 30°C for 12 h, and DNA was precipitated by
adding 0.5 ml of 2-propanol before the preparation was washed with 1 ml
of an 80% ethanol solution and dissolved in 0.2 ml of TE buffer. The
extracted DNA was quantified by measuring its UV absorption spectrum
(35) and was finally dissolved in TE buffer at a
concentration of 100 µg per ml.
DNA was also extracted from cells incorporated into activated-sludge
flocs. Five milliliters of an activated-sludge suspension
was
centrifuged at 500 ×
g for 5 min to precipitate the
activated-sludge
flocs. These flocs were resuspended in 5 ml of MP
medium, and
DNA was extracted from the suspension as described above.
Detection of strains BH and E6 by PCR.
The extracted DNA was
subjected to PCR amplification with a Trio-Thermoblock thermal cycler
(Biometra, Göttingen, Germany). The primers used were primers
BHS1 (5'-TGCTGAAGGATGAACGTAGT-3') and BHR1
(5'-TACCCTTCAACGGCAGGATT) for strain BH and primers SSS (5'-TGCGTGAACGCGCTCAGCAA-3') and SHR3
(5'-ACGCCGTTGTTCAGGAACGAG-3') for strain E6. The PCR for
strain BH was conducted in a reaction mixture (total volume, 100 µl)
containing 2.5 U of Taq DNA polymerase (AmpliTaq Gold;
Perkin-Elmer, Branchburg, N.J.), 10 mM Tris-HCl (pH 8.3), 50 mM KCl,
1.5 mM MgCl2, 0.001% (wt/vol) gelatin, each deoxynucleoside triphosphate at a concentration of 200 µM, 1 µg of
DNA, and 100 pmol of each primer. The PCR for strain E6 was performed
like the PCR for strain BH, except that 2.5 mM MgCl2 and 50 pmol of each primer were used. The thermal profile used for
amplification of the gyrB fragment of strain BH consisted of
10 min of activation of the polymerase at 94°C, followed by 30 cycles
consisting of 1 min at 94°C, 1 min at 60°C, and 2 min at 72°C,
and finally extension for 10 min at 72°C. The thermal profile used
for amplification of DNA from strain E6 consisted of activation of the
polymerase for 10 min at 94°C, followed by 30 cycles consisting of 1 min at 94°C, 1 min at 64°C, and 2 min at 72°C, and finally
extension for 10 min at 72°C.
Analysis of the PCR products.
Ten microliters of each PCR
product was subjected to electrophoresis on an agarose gel
(high-strength, analytical-grade agarose; Bio-Rad Laboratories,
Hercules, Calif.) containing agarose at concentrations of 1.5%
(wt/vol) for strain BH and 3.0% (wt/vol) for strain E6 in TAE buffer
(35). To quantitatively analyze the PCR products, 5 µl of
a DNA quantity standard (approximately 100 ng of DNA) was applied to
the gel; this standard was prepared by 10-fold dilution of the PCR
product amplified by the procedure described above from 1 µg of DNA
extracted from a pure culture of strain BH or E6. After
electrophoresis, the DNA in the gel was stained with SYBR Green I (FMC
Bioproducts) for 3 h as recommended by the manufacturer and then
photographed. The intensity of each PCR product was determined from a
negatively printed photograph of the gel with a model TIAS 100 densitometer (TEF Corporation, Tokyo, Japan).
Statistics.
Data were statistically analyzed by the
t test (P = 0.05).
Nucleotide sequence accession numbers.
The nucleotide
sequence data reported in this paper have been deposited in the GSDB,
DDBJ, EMBL, and NCBI nucleotide sequence databases under accession no.
AB002234, AB002235, AB002236, and AB002237.
| |
RESULTS AND DISCUSSION |
Design of PCR primers for specific probing.
A phylogenetic
analysis of the gyrB sequence of strain BH was conducted
previously (16), and this strain was classified as a
P. putida strain. PCR primers BHS1 and BHR1 for specific detection of strain BH were designed by comparing the gyrB
sequence of strain BH with the gyrB sequences of other
P. putida strains. The gyrB sequences of strains
BH, A10L, FK715, IFO14671, and JCM6156 have appeared in nucleotide
sequence databases under accession no. D86010, D86005, D86014, D86011,
and D37926, respectively. The gyrB sequence of strain
IFO14164 was identical to the sequence of P. putida PRS2000
(accession no. X54631). These primers allowed amplification of a 738-bp
fragment from the total DNA of strain BH (Fig. 1a, lane 11), but not
from the total DNAs of other P. putida strains (data not
shown).
The 805-bp nucleotide sequence in the 5' region of
gyrB of
strain E6 was determined. A phylogenetic analysis of this partial
gyrB sequence revealed the close relationship between this
strain
and
Comamonas strains. The E6
gyrB
sequence was 87, 83, and 83%
identical to the
gyrB
sequences of
C. terrigena IAM12052,
C. testosteroni IAM12419, and
C. acidovorans IFO13582,
respectively. PCR primers
SSS and SHR3 for specific detection of strain
E6 were designed
by comparing the
gyrB sequences of these
strains. These primers
allowed amplification of a 277-bp fragment from
the total DNA
of strain E6 (Fig.
1b, lane 11) but not from the total
DNAs of
other
Comamonas strains (data not shown).
Development of quantitative PCR.
Use of the primers designed
for specific amplification of the gyrB fragments of strains
BH and E6 allowed a quantitative PCR method to be developed. First,
experiments were conducted in which cultures of strains BH and E6 or
serial dilutions of cultures were mixed with phenol-digesting activated
sludge obtained from the laboratory unit (MLSS; 1,850 ppm; total cell
count, 3.8 × 109 cells per ml); this was followed by
DNA extraction and PCR amplification. The amount of DNA extracted from
each of the activated-sludge samples was 51 ± 6.1 µg
(n = 9). As shown in Fig.
1, only those fragments with the expected
molecular sizes were amplified by PCR, not only from DNA extracted from
each pure culture (lane 11), but also from DNA extracted from the
activated sludge mixed with the BH and E6 cultures (lane 2). In
addition, no fragments were amplified from DNA extracted from
noninoculated activated sludge (lane 10). These results suggest that
gyrB-targeted PCR can be used for specific detection of a BH
or E6 population in activated sludge.
View larger version (43K):
[in this window]
[in a new window]
|
FIG. 1.
PCR amplification of the gyrB fragments of
strains BH and E6. For the upper part of each gel 10-fold serial
dilutions of each pure culture were mixed with the activated sludge,
and DNA extraction from the mixed samples and PCR amplification of the
extracted DNA were carried out. (a) Strain BH. Lane 1, 50-2500 DNA size
marker (FMC Corporation); lane 2, 2.5 × 107 cells per
ml; lane 3, 2.5 × 106 cells per ml; lane 4, 2.5 × 105 cells per ml; lane 5, 2.5 × 104
cells per ml; lane 6, 2.5 × 103 cells per ml; lane 7, 2.5 × 102 cells per ml; lane 8, 2.5 × 101 cells per ml; lane 9, 2.5 × 100 cells
per ml; lane 10, uninoculated activated sludge; lane 11, pure BH
culture. (b) Strain E6. Lane 1, 50-2500 DNA marker; lane 2, 1.9 × 107 cells per ml; lane 3, 1.9 × 106 cells
per ml; lane 4, 1.9 × 105 cells per ml; lane 5, 1.9 × 104 cells per ml; lane 6, 1.9 × 103 cells per ml; lane 7, 1.9 × 102 cells
per ml; lane 8, 1.9 × 101 cells per ml; lane 9, 1.9 × 100 cells per ml; lane 10, uninoculated
activated sludge; lane 11, pure E6 culture. For the lower part of each
gel DNA was extracted from activated sludge containing strains BH and
E6 at densities of 2.5 × 107 and 1.9 × 107 cells per ml, respectively, before 10-fold serial
dilution of the extracted DNA with DNA extracted from the uninoculated
activated sludge. PCR amplification was carried out by using the
diluted samples. (a) Strain BH. (b) Strain E6. Lanes 1, 50-2500 DNA
size marker; lanes 2, undiluted DNA; lanes 3, 10-fold dilution; lanes
4, 102-fold dilution; lanes 5, 103-fold
dilution; lanes 6, 104-fold dilution; lanes 7, 105-fold dilution; lanes 8, 106-fold dilution;
lanes 9, 107-fold dilution; lanes 10, uninoculated
activated sludge; lanes 11, pure culture.
|
|
Figure
1 also shows that the quantitative ranges of the PCR method for
the BH and E6 populations were 10
3 to 10
4 and
10
4 to 10
5 cells per ml, respectively. The
lower detection limit for strain
E6 was approximately 10 times higher
than the lower detection
limit for strain BH. This may have been due to
a lower efficiency
of PCR amplification for strain E6 than for strain
BH. The relative
efficiency of a PCR is influenced by a variety of
factors, including
the length and secondary structure of the target
nucleic acid
molecule (
8). In the case of the E6
gyrB sequence, its high
GC content may have reduced the
efficiency of the PCR. Selvaratnam
et al. (
38) reported that
the sensitivity of the reverse transcriptase-PCR
method for detecting a
dmpN-expressing pseudomonad population
in activated sludge
was 10
4 CFU per 10 mg of activated sludge, which is
identical to the
sensitivity of our method. The lower detection limit
of a PCR
for specific detection of
Azoarcus tolulyticus in
soil has been
reported to be 10
2 cells per ml
(
51). The differences in sensitivity values may
have been
caused by differences in the amounts of inhibitory substances
in the
extracted DNA samples or in the PCR conditions. Activated
sludge
contains a large amount of extracellular polysaccharide
which is
coextracted with DNA and is known to be inhibitory to
PCR
(
43). A number of modifications of the present method (for
example, further purification of the extracted DNA, an increase
in the
number of PCR cycles, and improvement in the detection
method
[
8]) may be possible and may increase the sensitivity
of the PCR assay, although our present PCR method was sensitive
enough
to monitor the population dynamics of strains BH and E6
in the
activated sludge.
One useful method for expanding the upper limit of detection of a
quantitative PCR assay is to use a diluted sample (
8),
so we
next tested the applicability of this method. DNA extracted
from
activated sludge containing BH and E6 cells was mixed with
DNA
extracted from uninoculated phenol-digesting activated sludge
in
different ratios so that the DNA concentration in each solution
was 100 µg per ml, and 1 µg of each DNA sample was then subjected
to PCR.
The DNA extracted from uninoculated activated sludge was
used to dilute
DNA from the activated sludge containing the BH
and E6 cells, because
it was found that the efficiency of the
PCR was strongly affected by
the amount of sludge-extracted DNA
subjected to the PCR (data not
shown). As shown in Fig.
1, the
band intensity of each PCR product
amplified from the diluted
DNA samples was similar to the band
intensity of each PCR product
amplified from DNA extracted from sludge
containing the same number
of cells of the strain. This result
indicates that DNA dilution
should allow quantitative PCR
determinations for broad ranges
of population densities of these
strains.
To determine more accurately the relationship between the amount of PCR
products and the number of cells in the activated
sludge, additional
experiments were conducted, in which dilutions
of BH and E6 cultures in
LB medium were mixed with the phenol-digesting
activated sludge; then
the DNA was extracted, and a PCR was performed.
The numbers of cells of
these strains after they were mixed with
the sludge were estimated from
plate counts of the pure cultures.
The PCR products were
electrophoresed together with a DNA quantity
standard prepared as
described in Materials and Methods, and the
relative intensity of the
PCR product compared with the intensity
of the DNA quantity standard
was determined by densitometry. Figure
2
shows that there was a linear relationship between the relative
intensity of each PCR product and the number of cells in the activated
sludge, indicating that the PCR method was quantitative within
the
ranges examined. The population density could be interpolated
from the
relative intensity of the PCR product with the equations
shown in Fig.
2.
View larger version (13K):
[in this window]
[in a new window]
|
FIG. 2.
Standard curves for determining the densities of BH and
E6 populations in laboratory unit sludge. A quantitative PCR was
carried out as described in the legend to Fig. 1, except that a DNA
quantity standard was used to compare the results obtained in different
gels. The ordinate indicates the intensity of the band of each PCR
product relative to the intensity of the DNA quantity standard. The
means of three determinations are shown, and the error bars indicate
standard deviations.
|
|
Finally, to examine the general applicability of the quantitative
PCR method, strain BH was introduced into the Ohdaira activated
sludge,
and its population density was determined by the quantitative
PCR
method. Cells of strain BH were mixed with the Ohdaira activated
sludge
(MLSS; 1,730 ppm; total cell count, 2.4 × 10
9 cells
per ml), and DNA was extracted from the inoculated sludge
and
uninoculated sludge and then subjected to PCR with the BH-specific
primers. The PCR conditions were the same as those developed for
detecting the BH population in the sludge of the laboratory unit
except
for the amount of DNA added to the PCR solution (0.2 µg)
and the
amplification cycle (40 cycles). The BH population in
the Ohdaira
sludge was specifically detected by this PCR method,
as shown in Fig.
3, with a sensitivity similar to that
observed
with the phenol-digesting activated sludge. A standard curve
for
enumerating the BH population in the Ohdaira sludge was produced
for a range of 1.3 × 10
3 to 7.8 × 10
3 cells per ml, a range similar to that observed for the
BH population
in the laboratory unit sludge. The standard curve could
be expressed
with the following equation:
R = 0.25 × 10
3 D + 0.19 (
r2 = 0.97), where
R is the relative
intensity and
D is the population
density (in cells per ml).
In addition, when the DNA extracted
from uninoculated sludge was used
for dilution, a BH population
density of 1.4 × 10
6 ± 0.3 × 10
6 cells per ml (
n = 3) in the
Ohdaira sludge was accurately determined
by the PCR method (data not
shown). These results suggest that
although the PCR conditions need to
be optimized depending on
the sludge, the method for PCR quantitation
developed in this
study is widely applicable for quantifying the
bacterial population
introduced into an activated-sludge microbial
community.
View larger version (113K):
[in this window]
[in a new window]
|
FIG. 3.
Specificity and sensitivity of the PCR method for
detecting the BH population in Ohdaira activated sludge. Lane 1, 50-2500 DNA size marker; lane 2, 3.5 × 106 cells per
ml; lane 3, 3.5 × 105 cells per ml; lane 4, 3.5 × 104 cells per ml; lane 5, 3.5 × 103
cells per ml; lane 6, 3.5 × 102 cells per ml; lane 7, 3.5 × 101 cells per ml; lane 8, 3.5 × 100 cells per ml; lane 9, uninoculated activated sludge;
lane 10, pure BH culture.
|
|
Population dynamics of strains BH and E6 in phenol-digesting
activated sludge.
BH and E6 cells grown in LB medium were
introduced into the laboratory unit containing the phenol-digesting
Takinoshita activated sludge, and the fate of each of these strains was
investigated by the quantitative PCR method. Under stable operational
conditions (a phenol-loading rate of 0.4 g per liter per day), the
phenol concentration in the aeration tank never exceeded 0.5 ppm (the lower limit of detection of the phenol assay), and the total cell count
ranged from 3 × 109 to 5 × 109
cells per ml throughout this experiment. Based on plate counts determined for the inocula, the numbers of BH and E6 cells in the
aeration tank immediately after inoculation were estimated to be
5.2 × 107 ± 0.5 × 107 and 9.1 × 107 ± 0.4 × 107 cells per ml
(n = 3), respectively. One hour after inoculation, the
densities of the BH and E6 populations were determined by the
quantitative PCR to be 6.3 × 107 ± 1.2 × 107 and 7.9 × 107 ± 2.8 × 107 cells per ml (n = 3), respectively;
these values were not significantly different from the plate count
values described above.
Figure
4 shows the fate of the BH and E6
populations in the activated sludge. It was observed that the decline
in density
of each of these two populations was multiphasic; there was
an
initial rapid decline, followed by a slower decline. However,
the
time at which the phase shift occurred was different in these
two
populations; the BH population entered the slow-decline phase
on day
10, whereas the change occurred in the E6 population 5
days after
inoculation. In both populations, most of the cells
were incorporated
into activated-sludge flocs in the slow-decline
phase. In contrast, in
the initial rapid-decline phase, the tendencies
of the two populations
to be incorporated into flocs were found
to be different. For instance,
approximately 75% of the cells
in the E6 population were incorporated
into flocs on day 1, whereas
approximately 39% of the cells in the BH
population were in flocs.
These data indicate that the E6 population
was more rapidly incorporated
into the activated-sludge flocs than the
BH population, which
may have resulted in the earlier phase shift of
the E6 population.
View larger version (15K):
[in this window]
[in a new window]
|
FIG. 4.
Dynamics of the BH (a) and E6 (b) populations introduced
into phenol-digesting activated sludge under stable operational
conditions at a phenol-loading rate of 0.4 g per liter per day.
Symbols:  , population density in total activated sludge;  ,
population density in activated-sludge flocs (per milliliter of liquid
in the aeration tank); dashed line, theoretical washout curve at a
Tr of 10 days; dotted line, theoretical washout
curve at a Tr of 0.5 day. The theoretical curves
were drawn by using the following equation: St = S0 · exp[t/Tr],
where St is the population density at time
t and S0 is the initial population
density. The means of three determinations are shown, and the error
bars indicate standard deviations.
|
|
The E6 population survived in the activated sludge at a density that
was more than 10 times higher than that of the BH population.
Theoretical washout curves for dispersed cells (residence time
[
Tr], 0.5 day) and for cells incorporated into
activated-sludge
flocs (
Tr, 10 days) are also
shown in Fig.
4. The slope of the
E6 population curve in the
slow-decline phase was less steep than
the slope of the theoretical
curve (
Tr, 10 days), clearly indicating
that the
E6 population grew in the activated-sludge flocs during
this period. On
the other hand, judging from the slope, the growth
of the BH population
in the slow-decline phase was not as noticeable
as the growth of the E6
population. However, since predation must
have occurred, it is likely
that the BH population also grew at
a rate that compensated for the
decline in the population density
due to predation. From these
findings, we concluded that the E6
population grew more rapidly in the
activated-sludge flocs than
the BH population under stable operational
conditions.
On day 28, the activated sludge was subjected to shock loading of
phenol. One hour after the shock loading began, the phenol
concentration in the aeration tank reached 271 ppm, and the shock
loading was stopped at this moment. The phenol concentration gradually
decreased after that, and it was approximately 1 ppm at 19 h.
Phenol loading at a rate of 0.4 g per liter per day was restarted
at 23 h. During and after shock loading, the MLSS concentration
and the total cell counts ranged from 1,800 to 2,000 ppm and from
3 × 10
9 to 5 × 10
9 cells per ml,
respectively. Before and after the phenol shock
loading, the densities
of the BH and E6 populations were determined
by the PCR method (Fig.
5). It was found that the BH population
significantly increased (approximately 10-fold) after shock loading.
The increase in the BH population in the total activated sludge
seemed
to be greater than the increase in the activated-sludge
flocs, although
the difference was not statistically significant.
The E6 population
also increased after shock loading, although
the increase was small and
just exceeded the significant level.
After shock loading was stopped,
the BH population rapidly declined
and reached a level similar to the
level before shock loading.
However, the decrease in the E6 population
after shock loading
was as slow as the decrease observed during the
slow-decline phase
under the stable operational conditions shown in
Fig.
4. This
observation indicates that the BH population grew more
actively
than the E6 population during the shock loading period.
View larger version (37K):
[in this window]
[in a new window]
|
FIG. 5.
Dynamics of BH (a) and E6 (b) populations in
phenol-digesting activated sludge that was subjected to phenol shock
loading as described in the text. Dotted bars, population densities in
total activated sludge; cross-hatched bars, population densities in
activated-sludge flocs (per milliliter of liquid in the aeration tank).
The means of three determinations are shown, and the error bars
indicate standard deviations.
|
|
It has been suggested that several factors affect the growth of
bacteria in a microbial community (
3,
34); some of these
factors are nutrient availability, the presence of toxins, attachment
of cells to matrices, and physical parameters. One of these, nutrient
availability (i.e., the phenol concentration), was the only factor
that
apparently changed in the experiments described above, suggesting
that
the phenol concentration in the aeration tank influenced
the changes in
the growth tendencies of the BH and E6 populations.
The growth kinetics
of strains BH and E6 on phenol were determined
in a laboratory
pure-culture experiment by the method described
by Watanabe et al.
(
47). The three kinetic constants in Haldane's
equation
(
18,
50),
Ks,
Ki, and µ
max, determined were
22.2
± 2.2 ppm, 107 ± 10 ppm, and 1.52 ± 0.07 h
1, respectively, for BH and 10.5 ± 1.8 ppm,
46.4 ± 5.4 ppm, and
0.91 ± 0.07 h
1,
respectively, for E6 (estimated values ± standard errors). From
the kinetics, it could be estimated by extrapolation that the
specific
growth rate of strain E6 was higher than that of strain
BH in the
presence of phenol concentrations below 5.7 ± 0.7 ppm.
It is thus
conceivable that the more rapid growth of the E6 population
than of the
BH population under stable operational conditions
and the reverse
growth trend after shock loading reflected the
growth kinetics of the
organisms. Additional experiments, such
as experiments to determine the
rates of incorporation into flocs
and predation, are needed to predict
the population dynamics in
activated sludge.
| |
ACKNOWLEDGMENTS |
We thank Nobuhiro Takahashi for continuing support of this work.
We also thank Ikuko Hiramatsu for technical assistance.
| |
FOOTNOTES |
*
Corresponding author. Mailing address: Marine
Biotechnology Institute, Kamaishi Laboratories, 3-75-1 Heita, Kamaishi
City, Iwate 026, Japan. Phone: 81 193 26 6537. Fax: 81 193 26 6584. E-mail: kazwata{at}kamaishi.mbio.co.jp.
| |
REFERENCES |
| 1.
|
Arima, K.,
K. Komagata, and S. Minoda.
1954.
Metabolism of aromatic compounds. Part 1. Bacterial oxidation of three isomers of monohydroxybenzoic acids.
Agric. Chem. Soc. Jpn.
28:629-635.
|
| 2.
|
Assinder, S. J., and P. A. Williams.
1990.
The TOL plasmids: determination of the catabolism of toluene and the xylenes.
Adv. Microb. Physiol.
31:1-69[Medline].
|
| 3.
|
Atlas, R. M., and R. Bartha.
1992.
Microbial communities and ecosystems, p. 130-162.
Microbial ecology: fundamentals and applications.
The Benjamin/Cummings Publishing Company Inc., Redwood City, Calif.
|
| 4.
|
Blackburn, J. W.,
R. K. Jain, and G. S. Sayler.
1987.
Molecular microbial ecology of a naphthalene-degrading genotype in activated sludge.
Environ. Sci. Technol.
21:884-890.
|
| 5.
|
Bond, P. L.,
P. Hugenholtz,
J. Keller, and L. L. Blackall.
1995.
Bacterial community structures of phosphate-removing and non-phosphate-removing activated sludge from sequencing batch reactors.
Appl. Environ. Microbiol.
61:1910-1916[Abstract].
|
| 6.
|
DeLong, E. F.,
G. S. Wickham, and N. R. Pace.
1989.
Phylogenetic strains: ribosomal RNA-based probes for the identification of single cells.
Science
243:1360-1363[Abstract/Free Full Text].
|
| 7.
|
de Vos, P.,
K. Kersters,
E. Falsen,
B. Pot,
M. Gillis,
P. Segers, and J. de Ley.
1985.
Comamonas Davis and Park 1962 gen. nov., nom. rev. emend., and Comamonas terrigena Hugh 1962 sp. nov., nom. rev.
Int. J. Syst. Bacteriol.
35:443-453[Abstract/Free Full Text].
|
| 8.
|
Diaco, R.
1995.
Practical considerations for the design of quantitative PCR assays, p. 84-107. In
M. A. Innis, D. H. Gelfand, and J. J. Sninsky (ed.), PCR strategies.
Academic Press, San Diego, Calif.
|
| 9.
|
Digiovanni, G. D.,
J. W. Neilson,
I. L. Pepper, and N. A. Sinclair.
1996.
Gene transfer of Alcaligenes eutrophus JMP134 plasmid pJP4 to indigenous soil recipients.
Appl. Environ. Microbiol.
62:2521-2526[Abstract].
|
| 10.
|
Dikshituru, S.,
B. C. Baltzis, and G. A. Lewandowski.
1993.
Competition between two microbial populations in a sequencing fed-batch reactor: theory, experimental verification, and implications for waste treatment applications.
Biotechnol. Bioeng.
42:643-656.
|
| 11.
|
Diviacco, S.,
P. Norio,
L. Zentilin,
S. Menzo,
M. Clementi,
G. Biamonti,
S. Riva,
A. Falaschi, and M. Giacca.
1992.
A novel procedure for quantitative polymerase chain reaction by coamplification of competitive templates.
Nucleic Acids Res.
122:313-320.
|
| 12.
|
Fujita, M.,
M. Ike, and K. Uesugi.
1994.
Operation parameters affecting the survival of genetically engineered microorganisms in activated sludge processes.
Water Res.
28:1667-1672.
|
| 13.
|
Furukawa, K.
1982.
Microbial degradation of polychlorinated biphenyls, p. 33-57. In
A. M. Chakrabarty (ed.), Biodegradation and detoxification of environmental pollutants.
CRC Press, Inc., Boca Raton, Fla.
|
| 14.
|
Gray, P. H. H.
1928.
The formation of indigotin from indol by soil bacteria.
Proc. R. Soc. Lond. B Biol. Sci.
102:263-279[Free Full Text].
|
| 15.
|
Greer, L. E.,
J. A. Joseph, and D. R. Shelton.
1992.
Kinetic comparison of seven strains of 2,4-dichlorophenoxyacetic acid-degrading bacteria.
Appl. Environ. Microbiol.
58:1027-1030[Abstract/Free Full Text].
|
| 16.
|
Harayama, S., and S. Yamamoto.
1996.
Phylogenetic identification of Pseudomonas strains based on a comparison of gyrB and rpoD sequences, p. 250-258. In
T. Nakazawa, K. Furukawa, D. Haas, and S. Silver (ed.), Molecular biology of pseudomonads.
American Society for Microbiology, Washington, D.C.
|
| 17.
|
Hashimoto, S., and M. Fujita.
1987.
Identification of three phenol-degrading microorganisms isolated from activated sludges and their characteristics.
J. Jpn. Sewage Works
9:655-660.
|
| 18.
|
Hill, G. A., and C. W. Robinson.
1975.
Substrate inhibition kinetics: phenol degradation by Pseudomonas putida.
Biotechnol. Bioeng.
17:1599-1615.
|
| 19.
|
Holben, W. E.,
B. M. Schroeter,
V. G. M. Calabrese,
R. H. Olsen,
J. K. Kukor,
V. O. Biederbeck,
A. E. Smith, and J. M. Tiedje.
1992.
Gene probe analysis of soil microbial populations selected by amendment with 2,4-dichlorophenoxyacetic acid.
Appl. Environ. Microbiol.
58:3941-3948[Abstract/Free Full Text].
|
| 20.
|
Islam, M. S.,
M. K. Hasan,
M. A. Miah,
G. C. Sur,
A. Felsenstein,
M. Venkatesan,
R. B. Sack, and M. J. Albert.
1993.
Use of the polymerase chain reaction and fluorescent-antibody methods for detection of viable but nonculturable Shigella dysenteriae type 1 in laboratory microcosms.
Appl. Environ. Microbiol.
59:536-540[Abstract/Free Full Text].
|
| 21.
|
Japanese Industrial Standard Committee.
1986.
.
Testing methods for industrial wastewater, JIS K0102.
Japanese Standards Association, Tokyo, Japan.
|
| 22.
|
Ka, J. O.,
W. E. Holben, and J. M. Tiedje.
1994.
Analysis of competition in soil among 2,4-dichlorophenoxyacetic acid-degrading bacteria.
Appl. Environ. Microbiol.
60:1121-1128[Abstract/Free Full Text].
|
| 23.
|
Lee, S.-Y.,
J. Bollinger,
D. Bezdicek, and A. Ogram.
1996.
Estimation of the abundance of an uncultured soil bacterial strain by a competitive quantitative PCR method.
Appl. Environ. Microbiol.
62:3787-3793[Abstract].
|
| 24.
|
Leser, T. D.,
M. Boye, and N. Hendriksen.
1995.
Survival and activity of Pseudomonas sp. strain B13 (FR1) in a marine microcosm determined by quantitative PCR and an rRNA-targeting probe and its effect on indigenous bacterioplankton.
Appl. Environ. Microbiol.
61:1201-1207[Abstract].
|
| 25.
|
Manz, W.,
M. Wagner,
R. Amann, and K. Schleifer.
1994.
In situ characterization of the microbial consortia active in two wastewater treatment plants.
Water Res.
28:1715-1723.
|
| 26.
|
Marmur, J.
1961.
A procedure for the isolation of deoxyribonucleic acid from microorganisms.
J. Mol. Biol.
3:208-218.
|
| 27.
|
Matheson, V. G.,
L. J. Forney,
Y. Suwa,
C. H. Nakatsu,
A. J. Sexstone, and W. E. Holben.
1996.
Evidence for acquisition in nature of a chromosomal 2,4-dichlorophenoxyacetic acid/ -ketoglutarate dioxygenase gene by different Burkholderia spp.
Appl. Environ. Microbiol.
62:2457-2463[Abstract].
|
| 28.
|
Matheson, V. G.,
J. Munakata-Marr,
G. D. Hopkins,
P. L. McCarty,
J. M. Tiedje, and L. J. Forney.
1997.
A novel means to develop strain-specific DNA probes for detecting bacteria in the environment.
Appl. Environ. Microbiol.
63:2863-2869[Abstract].
|
| 29.
|
McClure, N. C.,
A. J. Weightman, and J. C. Fry.
1989.
Survival of Pseudomonas putida UWC1 containing cloned catabolic genes in a model activated-sludge unit.
Appl. Environ. Microbiol.
55:2627-2634[Abstract/Free Full Text].
|
| 30.
|
McClure, N. C.,
J. C. Fry, and A. J. Weightman.
1991.
Survival and catabolic activity of natural and genetically engineered bacteria in a laboratory-scale activated-sludge unit.
Appl. Environ. Microbiol.
57:366-373[Abstract/Free Full Text].
|
| 31.
|
McMacken, R.,
L. Silver, and C. Geogopoulos.
1987.
DNA replication, p. 578-580. In
F. C. Neidhardt, J. L. Ingraham, K. B. Low, B. Magasanik, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella typhimurium: cellular and molecular biology.
American Society for Microbiology, Washington, D.C.
|
| 32.
|
Nüblein, K.,
D. Maris,
K. Timmis, and D. F. Dwyer.
1992.
Expression and transfer of engineered catabolic pathways harbored by Pseudomonas spp. introduced into activated sludge microcosms.
Appl. Environ. Microbiol.
58:3380-3386[Abstract/Free Full Text].
|
| 33.
|
Pickup, R. W.
1991.
Development of molecular methods for the detection of specific bacteria in the environment.
J. Gen. Microbiol.
137:1009-1019.
|
| 34.
|
Roszak, D. B., and R. R. Colwell.
1987.
Survival strategy of bacteria in the natural environment.
Microbiol. Rev.
51:365-379[Free Full Text].
|
| 35.
|
Sambrook, J.,
E. F. Fritsch, and T. Maniatis.
1989.
.
Molecular cloning: a laboratory manual, 2nd ed.
Cold Spring Harbor Laboratory, Cold Spring Harbor, N.Y.
|
| 36.
|
Saraswat, N.,
J. E. Alleman, and T. J. Smith.
1994.
Enzyme immunoassay detection of Nitrosomonas europaea.
Appl. Environ. Microbiol.
60:1969-1973[Abstract/Free Full Text].
|
| 37.
|
Sayler, G. S., and A. C. Layton.
1990.
Environmental application of nucleic acid hybridization.
Annu. Rev. Microbiol.
44:625-648[Medline].
|
| 38.
|
Selvaratnam, S.,
B. A. Schoedel,
B. L. Mcfarland, and C. F. Kulpa.
1995.
Application of reverse transcriptase PCR for monitoring expression of the catabolic dmpN gene in a phenol-degrading sequencing batch reactor.
Appl. Environ. Microbiol.
61:3981-3985[Abstract].
|
| 39.
|
Shimao, M.,
T. Nakamura,
A. Okuda,
M. Abe, and S. Harayama.
1996.
Characterization of transposon insertion mutants of mandelic acid-utilizing Pseudomonas putida strain A10L.
Biosci. Biotechnol. Biochem.
60:1051-1055[Medline].
|
| 40.
|
Stanier, R. Y.,
N. J. Palleroni, and M. Doudoroff.
1966.
The aerobic pseudomonads: a taxonomic study.
J. Gen. Microbiol.
43:159-271[Abstract/Free Full Text].
|
| 41.
|
Steffan, R. J., and R. M. Atlas.
1991.
Polymerase chain reaction: application in environmental microbiology.
Annu. Rev. Microbiol.
45:137-161[Medline].
|
| 42.
|
Sykes, P. J.,
S. H. Neoh,
M. J. Brisco,
E. Hughes,
J. Condon, and A. A. Morley.
1992.
Quantitation of targets for PCR by use of limiting dilution.
BioTechniques
13:444-449.
[Medline] |
| 43.
|
Taylor, J. L.
1993.
A simple, sensitive, and rapid method for detecting seed contaminated with highly virulent Leptoshaeria maculans.
Appl. Environ. Microbiol.
59:3681-3685[Abstract/Free Full Text].
|
| 44.
|
Wagner, M.,
R. Amann,
H. Lemmer, and K. H. Schleifer.
1993.
Probing activated sludge with oligonucleotide specific for proteobacteria: inadequacy of culture-dependent methods for describing microbial community structure.
Appl. Environ. Microbiol.
59:1520-1525[Abstract/Free Full Text].
|
| 45.
|
Watanabe, K.,
S. Hino,
K. Onodera, and N. Takahashi.
1994.
Studies on population dynamics of bacteria in a wastewater treatment process by using enzyme immunoassay, abstr. 3-F-9-35, p. 720-721.
Abstracts of the 28th Annual Meeting of the Japan Society on Water Environment.
The Japan Society on Water Environment, Tokyo, Japan.
|
| 46.
|
Watanabe, K.,
S. Hino,
K. Onodera,
S. Kajie, and N. Takahashi.
1996.
Diversity in kinetics of bacterial phenol-oxygenating activity.
J. Ferment. Bioeng.
81:562-565.
|
| 47.
|
Watanabe, K.,
S. Hino, and N. Takahashi.
1996.
Effects of exogenous phenol-degrading bacteria on performance and ecosystem of activated sludge.
J. Ferment. Bioeng.
82:291-298.
|
| 48.
|
Watanabe, K., and S. Hino.
1996.
Identification of a functionally important population in phenol-digesting activated sludge with antisera raised against isolated bacterial strains.
Appl. Environ. Microbiol.
62:3901-3904[Abstract].
|
| 49.
|
Yamamoto, S., and S. Harayama.
1995.
PCR amplification and direct sequencing of gyrB genes with universal primers and their application to the detection and taxonomic analysis of Pseudomonas putida strains.
Appl. Environ. Microbiol.
61:1104-1109[Abstract].
|
| 50.
|
Yang, R. D., and A. E. Humphrey.
1975.
Dynamics and steady state studies of phenol degradation in pure and mixed cultures.
Biotechnol. Bioeng.
17:1211-1235[Medline].
|
| 51.
|
Zhou, J.,
A. V. Palumbo, and J. M. Tiedje.
1997.
Sensitive detection of a novel class of toluene-degrading denitrifiers, Azoarcus tolulyticus, with small-subunit rRNA primers and probes.
Appl. Environ. Microbiol.
63:2384-2390[Abstract].
|
Appl Environ Microbiol, April 1998, p. 1203-1209, Vol. 64, No. 4
0099-2240/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
This article has been cited by other articles:
-
Vilchez, R., Pozo, C., Gomez, M. A., Rodelas, B., Gonzalez-Lopez, J.
(2007). Dominance of sphingomonads in a copper-exposed biofilm community for groundwater treatment. Microbiology
153: 325-337
[Abstract]
[Full Text]
-
Kasai, Y., Takahata, Y., Manefield, M., Watanabe, K.
(2006). RNA-Based Stable Isotope Probing and Isolation of Anaerobic Benzene-Degrading Bacteria from Gasoline-Contaminated Groundwater.. Appl. Environ. Microbiol.
72: 3586-3592
[Abstract]
[Full Text]
-
Futamata, H., Nagano, Y., Watanabe, K., Hiraishi, A.
(2005). Unique Kinetic Properties of Phenol-Degrading Variovorax Strains Responsible for Efficient Trichloroethylene Degradation in a Chemostat Enrichment Culture. Appl. Environ. Microbiol.
71: 904-911
[Abstract]
[Full Text]
-
Kasai, Y., Kishira, H., Harayama, S.
(2002). Bacteria Belonging to the Genus Cycloclasticus Play a Primary Role in the Degradation of Aromatic Hydrocarbons Released in a Marine Environment. Appl. Environ. Microbiol.
68: 5625-5633
[Abstract]
[Full Text]
-
Iwabuchi, N., Sunairi, M., Urai, M., Itoh, C., Anzai, H., Nakajima, M., Harayama, S.
(2002). Extracellular Polysaccharides of Rhodococcus rhodochrous S-2 Stimulate the Degradation of Aromatic Components in Crude Oil by Indigenous Marine Bacteria. Appl. Environ. Microbiol.
68: 2337-2343
[Abstract]
[Full Text]
-
Watanabe, K., Watanabe, K., Kodama, Y., Syutsubo, K., Harayama, S.
(2000). Molecular Characterization of Bacterial Populations in Petroleum-Contaminated Groundwater Discharged from Underground Crude Oil Storage Cavities. Appl. Environ. Microbiol.
66: 4803-4809
[Abstract]
[Full Text]
-
Fuller, M. E., Streger, S. H., Rothmel, R. K., Mailloux, B. J., Hall, J. A., Onstott, T. C., Fredrickson, J. K., Balkwill, D. L., DeFlaun, M. F.
(2000). Development of a Vital Fluorescent Staining Method for Monitoring Bacterial Transport in Subsurface Environments. Appl. Environ. Microbiol.
66: 4486-4496
[Abstract]
[Full Text]
-
Watanabe, K., Miyashita, M., Harayama, S.
(2000). Starvation Improves Survival of Bacteria Introduced into Activated Sludge. Appl. Environ. Microbiol.
66: 3905-3910
[Abstract]
[Full Text]
-
Boon, N., Goris, J., De Vos, P., Verstraete, W., Top, E. M.
(2000). Bioaugmentation of Activated Sludge by an Indigenous 3-Chloroaniline-Degrading Comamonas testosteroni Strain, I2gfp. Appl. Environ. Microbiol.
66: 2906-2913
[Abstract]
[Full Text]
-
Michotey, V., Méjean, V., Bonin, P.
(2000). Comparison of Methods for Quantification of Cytochrome cd1-Denitrifying Bacteria in Environmental Marine Samples. Appl. Environ. Microbiol.
66: 1564-1571
[Abstract]
[Full Text]
-
Mesarch, M. B., Nakatsu, C. H., Nies, L.
(2000). Development of Catechol 2,3-Dioxygenase-Specific Primers for Monitoring Bioremediation by Competitive Quantitative PCR. Appl. Environ. Microbiol.
66: 678-683
[Abstract]
[Full Text]
-
Kasai, H., Ezaki, T., Harayama, S.
(2000). Differentiation of Phylogenetically Related Slowly Growing Mycobacteria by Their gyrB Sequences. J. Clin. Microbiol.
38: 301-308
[Abstract]
[Full Text]
-
Watanabe, K., Teramoto, M., Harayama, S.
(1999). An Outbreak of Nonflocculating Catabolic Populations Caused the Breakdown of a Phenol-Digesting Activated-Sludge Process. Appl. Environ. Microbiol.
65: 2813-2819
[Abstract]
[Full Text]
-
Watanabe, K., Teramoto, M., Futamata, H., Harayama, S.
(1998). Molecular Detection, Isolation, and Physiological Characterization of Functionally Dominant Phenol-Degrading Bacteria in Activated Sludge. Appl. Environ. Microbiol.
64: 4396-4402
[Abstract]
[Full Text]
Top
Abstract
Introduction
