INTRODUCTION

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Many scientists have used the rRNA approach (29, 30) to detect microbial populations and to describe the structures of microbial communities in various environments without isolating the component microorganisms. These studies have shown that most 16S ribosomal DNA (rDNA) sequences directly amplified from environmental samples are different from the sequences of comparable laboratory strains. Workers have concluded from such observations that many bacteria that are predominant in the natural environment have not been isolated in the laboratory yet and that the microbial diversity in the natural environment is much greater than the diversity of the bacteria that have been isolated (2, 7, 13, 25, 35, 36, 39, 40).

Currently, one important aspect of microbial ecology studies is functional dissection of microbial communities based on structural information obtained by the approach mentioned above. An analysis of a population shift accompanied by a change in the function of a community yields information useful for identifying functionally dominant populations (2, 3, 42), although information concerning the function (activity) of each population can never be obtained by this kind of approach. Hence, workers have emphasized that pure-culture experiments are indispensable for detailed analysis of the functions of each population and that isolation of the functionally dominant populations in a microbial community is quite important.

Phenol and its derivatives are some of the major hazardous compounds in industrial wastewater (1, 31, 43), and for this reason biodegradation of phenol has attracted keen attention (34, 46). However, since most studies of phenol biodegradation have been carried out under laboratory conditions with arbitrarily selected phenol-degrading bacteria, phenol biodegradation in the environment is not well understood yet. In the present study, to better understand phenol degradation in activated sludge, we isolated and characterized the phenol-degrading bacteria that were identified by the rRNA approach to be the dominant population in phenol-digesting activated sludge. Physiological and genetic differences between the dominant phenol-degrading bacteria isolated in this study and representative phenol-degrading bacteria characterized previously in several laboratories are discussed below.

	MATERIALS AND METHODS

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Acclimation of activated sludge to phenol. Samples of activated-sludge mixed liquor were obtained from the return sludge line of a municipal sewage treatment plant (Ohdaira, Kamaishi, Iwate, Japan) in July 1997. The concentration of mixed-liquor suspended solids was determined by weighing the dried sludge collected on a 0.22-µm-pore-size filter by Japan Industrial Standards method K0102 (20); the value obtained was 6,150 mg per liter. Approximately 1 liter of the mixed liquor was infused into a laboratory activated-sludge unit (Miyamoto) composed of an aeration tank (volume, 3 liters) and a settling tank (volume, 2 liters). 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 (pH 6.8 to 7.0) and supplemented with 200 mg of phenol per liter was continuously supplied to the aeration tank at a flow rate of 6 liters per day; the hydraulic residence time in the aeration tank was 0.5 day. The phenol-loading rate was calculated to be 0.4 g per liter per day. The mixed-liquor suspended solids concentration in the aeration tank was kept between 1,800 and 2,000 mg per liter by discarding the excess sludge from the aeration tank. The mean sludge residence time was calculated to be approximately 10 days. Air was continuously supplied at a rate of 2 liters per min, and the temperature was maintained at approximately 25°C. The total direct count of bacteria in the activated sludge was determined by a fluorescent-microscopy method after staining with 4',6-diamidino-2-phenylindole (DAPI) (39). The phenol concentration in the aeration tank was determined by a colorimetric assay performed with Phenol Test Wako (Wako Pure Chemicals) (42). The total organic carbon concentration in the aeration tank was determined with a total organic carbon concentration meter (model TOC-5000; Shimadzu) (42).

DNA extraction from the activated sludge. DNA was extracted from 5 ml of mixed liquor obtained from the aeration tank of the laboratory unit as described previously (44). The quantity and quality of the extracted DNA were checked by measuring the UV absorption spectrum of the DNA solution (33), and the DNA was finally dissolved in TE buffer (33) at a concentration of 100 µg per ml.

PCR conditions. PCR primers P2 and P3 (containing 40 bp of GC clamp) (25) were used to amplify the variable V3 region of bacterial 16S rDNA (corresponding to positions 341 to 534 in the Escherichia coli sequence). Amplification was performed with a Progene thermal cycler (Techne) by using a 50-µl (total volume) mixture containing 1.25 U of Taq DNA polymerase (Amplitaq Gold; Perkin-Elmer), 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, 25 pmol of each primer, and 0.5 µl of the DNA solution. A modified form of the touchdown thermal profile technique (25) was used; this technique involved 10 min of activation of the polymerase at 94°C before two cycles consisting of 1 min at 94°C, 1 min at 65°C, and 2 min at 72°C. The annealing temperature was subsequently decreased by 1°C for every second cycle until it reached 55°C, at which point 20 additional cycles were carried out; finally, a 10-min extension step at 72°C was performed. Amplification of the PCR products of the proper size was confirmed by electrophoresis through a 1.5% (wt/vol) agarose gel (LO3 agarose; Takara Shuzo) in TBE buffer (33), followed by staining with ethidium bromide.

TGGE. A temperature gradient gel electrophoresis (TGGE) system (Taitec) was used as recommended by the manufacturer. Five microliters of a PCR-amplified mixture was subjected to electrophoresis in a 10% (wt/vol) polyacrylamide gel at 250 V for 3.5 h. Linear temperature gradients from 45 to 60°C were used to separate the 16S rDNA fragments, and linear temperature gradients from 55 to 70°C were used to separate the LmPH gene fragments; these gradients were applied parallel to the electrophoretic direction. After electrophoresis, the gel was stained with SYBR Green I (FMC Bioproducts) for 30 min as recommended by the manufacturer.

Sequencing of the TGGE bands. A gel slice containing a DNA band was excised and transferred into a sterile Eppendorf tube containing 100 µl of sterile TE buffer. The tube was gently shaken at 30°C for approximately 12 h, and after the gel slice was removed, 200 µl of ethanol was added. The tube was then incubated at 20°C for 1 h, and the DNA fragment was precipitated by centrifugation at 20,000 × g for 10 min. The precipitate was washed with 100 µl of a 70% (vol/vol) ethanol solution, and the resulting DNA was dissolved in 20 µl of sterile TE buffer. One microliter of this DNA solution was subjected to a second PCR, performed either under the same conditions as those for the first PCR to check the purity by TGGE or by using a modified procedure in which primers GC-2 (5'-GAAGTCATCATGACCGTTCTGGCACGGGGGGCCTA-3') and GC-2P (5'-GAAGTCATCATGACCGTTCTGGCACGGGGGGCGAT-3') were used instead of primers P3 and Phe149GC, respectively. The sequences of the last 15 bases of primers GC-2 and GC-2P are identical to sequences in the middle parts of primers P3 and Phe149GC, respectively, while the sequences of the first 21 bases of primers GC-2 and GC-2P are identical to the sequence of primer GC-1S (5'-GAAGTCATCATGACCGTTCTG-3'), which was used for the sequencing experiments described below. In the products of the second PCR that were amplified by using primers GC-2 and GC-2P, part of the GC clamp in the products of the first PCR was replaced by the sequence of primer GC-1S. The products of the second PCR were electrophoresed through a 1.5% (wt/vol) agarose gel in TBE buffer (33) and then purified with a QIAquick gel extraction kit (QIAGEN). The extracted DNA was quantified by measuring the absorbance at 260 and 320 nm (33). The nucleotide sequences of the PCR products were then determined in both orientations by using a DNA sequencing kit (Dye Terminator Cycle Sequencing kit; Perkin-Elmer) with primer P2 or GC-1S for 16S rDNA and with primer Phe212 or GC-1S for the LmPH gene. The products of the sequencing reactions were analyzed with a model 377 DNA sequencer (Perkin-Elmer).

Isolation of bacteria from the activated sludge. (i) Direct plating. Five milliliters of mixed liquor from the phenol-digesting activated sludge obtained from the aeration tank of the laboratory unit 20 days after phenol loading was begun 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) for 2 min. The resulting cell suspension was appropriately diluted with sterile MP medium containing 5 mM sodium tripolyphosphate and then spread onto agar plates containing dCGY medium, which was composed of (per liter) 0.5 g of Bacto Casamino Acids (Difco), 0.5 g of glycerol, and 0.1 g of Bacto Yeast Extract (Difco); the resulting plates were referred to as dCGY plates. The diluted cell suspension was also spread onto agar plates containing MP medium supplemented with 500 mg of phenol per liter (MP500 plates). The plates were incubated at 25°C for 7 days (dCGY plates) or for 14 days (MP500 plates). All of the colonies that appeared on one plate were picked and grown in 5 ml of dCGY medium, and the dCGY medium cultures were then restreaked onto dCGY plates. This purification procedure was repeated several times.

(ii) Batch culture enrichment. MP liquid medium (500 ml) supplemented with 500 mg of phenol per liter (MP500 medium) in a 1-liter baffled flask was inoculated with 500 µl of the mixed liquor from the phenol-digesting activated sludge, and the culture was shaken at 100 rpm for 24 h at 25°C. The resulting culture (500 µl) was transferred into fresh MP500 medium and cultivated as described above. Cells from the fourth enrichment culture were then streaked onto dCGY plates, and all of the colonies that appeared on one-half of one plate were picked and purified.

(iii) Chemostat enrichment. One liter of MP medium (1 liter) in a model TBR-2 2-liter fermentor (Sakura Fine Technical) was inoculated with 500 ml of mixed liquor from the phenol-digesting activated sludge. MP medium containing 1,500 mg of phenol per liter was continuously supplied at a rate of 750 ml per day, and the culture volume was maintained at 1.5 liters. Air was supplied at a rate of 2 liters per min, and the temperature was kept at 25°C. The concentration of phenol in the culture was below the detection limit (<0.5 mg per liter) throughout the experiment. After 7 days of cultivation, the culture was appropriately diluted and streaked onto dCGY plates. The plates were incubated at 25°C for 7 days, and all of the colonies on one plate were picked and purified.

rep-PCR. Genomic fingerprints of the bacteria isolated were obtained by repetitive extragenic palindromic sequence PCR (rep-PCR) performed with primers REP1R-I and REP2-I (6). The PCR was performed in a 50-µl (total volume) mixture containing 1.25 U of Taq DNA polymerase (Amplitaq Gold; Perkin-Elmer), 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, 50 pmol of each primer, and a small amount of bacterial cells that were transferred by needle from a colony that developed on a dCGY plate. The PCR conditions used were as follows: 10 min of activation of the polymerase at 94°C, followed by 40 cycles consisting of 1 min at 94°C, 1 min at 40°C, and 8 min at 65°C and finally by 10 min of extension at 72°C. The PCR products were electrophoresed through a 1.5% (wt/vol) LO3 agarose gel in TBE and stained with SYBR Green I for 2 h. The rep-PCR analysis was repeated several times to determine the reproducibility of the method.

Analyses of the sequences of the bacteria isolated. A small amount of bacterial cells picked from a colony developed on a dCGY plate was subjected to PCR in order to amplify partial 16S rDNA fragments and the LmPH fragments by the methods used for the TGGE analyses described above. The nucleotide sequences of the fragments were then determined by the method used to sequence the TGGE fragments. The sequences determined were aligned by using ClustalW, version 1.7 (38). A neighbor-joining tree (32) was constructed by using the njplot software in ClustalW, version 1.7. A search of the GenBank database was conducted by using BLAST (23).

Physiological characterization of the bacteria isolated. (i) Growth on phenol. Each bacterial strain purified on a dCGY plate was transferred to 5 ml of dCGY medium and grown to the stationary phase. The cells in 50 µl of culture were washed with MP medium and used to inoculate MP500 medium (10 ml) in an L-shaped test tube. The test tube was shaken at 60 rpm at 25°C, and the bacterial growth in the tube was monitored every 10 min by automatically measuring the optical density at 660 nm (OD₆₆₀) with a model TN-2612 Bio-photometer (Advantec). The growth of isolates on phenol was also examined by using chemostat cultures; using these cultures, we determined whether a stable culture could be obtained under the chemostat conditions described previously (41), except that the temperature was 25°C. The phenol concentration in the feed and the dilution rate were 1,500 mg per liter and 0.67 per day, respectively.

(ii) Kinetics of phenol-oxygenating activity. Cells were grown at 25°C in a chemostat culture fed with phenol as the sole carbon source as described previously (41), and samples were obtained after the culture parameters (pH, dissolved oxygen concentration, phenol concentration, and OD₆₆₀) became stable (i.e., at least 3 days after the culture was begun). Cell samples were obtained from the chemostat culture immediately before the activity was measured in order to ensure that the physiological conditions were almost identical. The dry cell weight in the culture was determined gravimetrically by filtering the culture through a 0.22-µm-pore-size membrane by the method of Machado and Grady (24). The phenol-oxygenating activity was measured with a Clark type oxygen electrode (model 5/6 Oxygraph; Gilson) as described previously (41). One unit of activity was equivalent to 1 µmol of oxygen consumed per min, while the specific activity was defined as the activity per gram of dried cells. The apparent kinetic constants, K_s, K_SI, and V_max in Haldane's equation (41), were determined by the nonlinear regression method described previously (41).

Nucleotide sequence accession numbers. The nucleotide sequences reported in this paper have been deposited in the GSDB, DDBJ, EMBL, and NCBI nucleotide sequence databases under accession no. AB011551 to AB011580.

	RESULTS

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Acclimation of activated sludge to phenol. The laboratory unit was inoculated with activated sludge obtained from a municipal sewage treatment plant on day 0 and was fed phenol as the sole carbon source. Phenol was detected in the aeration tank on days 1 and 2 at concentrations of 42 and 19 mg per liter, respectively; however, on subsequent days, the phenol concentration was below the detection limit (<0.5 mg per liter). The total organic carbon concentration remained constant at 10 ± 3 mg per liter throughout the experiment except on days 1 and 2, when it was 38 and 15 mg per liter, respectively. The total direct counts ranged from 3 × 10⁹ to 5 × 10⁹ cells per ml. These observations indicate that the activated sludge had nearly completely digested the phenol several days after phenol feeding was begun.

Detection of dominant species by TGGE. The bacterial populations in the phenol-digesting activated sludge were detected by isolating DNA from the sludge every 2 or 3 days and then performing a TGGE analysis of the 16S rDNA fragments amplified from the DNA. It was found that several dominant populations were present after the phenol feeding was begun, and the population structure became stable after day 10. Figure 1a shows the TGGE profiles obtained on days 0 and 20. On day 20, several populations were visible; three of these populations were designated R1, R2, and R3, and R2 and R3 were the major bands. Database searches performed with the nucleotide sequences determined indicated that R2 was identical to a taxonomically unidentified member of the  subclass of the class Proteobacteria, strain LX1 (accession no. AJ001271), and R3 exhibited 96% homology with clone T70, an unidentified member of the  subclass of the Proteobacteria from activated sludge (36), 96% homology with strain HW1, another unidentified member of the  subclass of the Proteobacteria (22), and 94% homology with Variovorax paradoxus (17). No DNA sequence that exhibited a high level of homology with R1 was found in the database; the organism that exhibited the highest level of homology (86%) was Psychroserpens burtonensis (4).

View larger version (52K): [in this window] [in a new window]   FIG. 1.   TGGE profiles of the fragments of 16S rDNA (a) and the LmPH gene (b) after PCR amplification from activated sludge obtained before phenol feeding (day 0) and after acclimation to phenol (day 20).

Isolation of bacteria. A total of 103 colonies were isolated from the phenol-digesting activated sludge obtained on day 20 by the following four different methods: direct plating on dCGY plates, direct plating on MP500 plates, plating on dCGY plates after batch culture enrichment, and plating on dCGY plates after enrichment by chemostat culturing. The number of colonies isolated by each method is shown in Table 1. The purified colonies were subjected to a rep-PCR analysis to identify identical strains. As shown in Fig. 2, 27 distinct rep-PCR patterns were obtained from the 103 colonies; 10 of these patterns were observed in colonies obtained from direct plating on dCGY plates (designated patterns rN1 to rN10), 7 patterns were observed in colonies obtained from direct plating on MP500 plates (patterns rP1 to rP7), 2 patterns were observed in colonies obtained from the batch enrichment (patterns rB1 and rB2), and 8 patterns were observed in colonies obtained from the chemostat enrichment (patterns rC1 to rC8). The number of colonies that produced each rep-PCR pattern is also shown in Table 1. Below, each representative isolate is referred to by the designation of its rep-PCR pattern. The rep-PCR patterns were always different for strains obtained by different isolation methods.

View larger version (57K): [in this window] [in a new window]   FIG. 2.   Unique rep-PCR patterns of bacterial strains isolated from phenol-digesting activated sludge (day 20) by four different methods. The designations of the rep-PCR patterns are indicated below the lanes. Lanes M contained 50-2500 DNA size marker (FMC Corp.).

16S rDNA and LmPH genes in the isolates. The partial 16S rDNA sequences of the 27 isolates were determined, and these sequences were compared with the sequences of the dominant populations in the activated sludge that were detected by the TGGE analysis. This comparison revealed that the sequences of two isolates obtained from the chemostat enrichment were identical to the sequence of the R2 band, while the sequences of four isolates (two isolates obtained by direct plating on dCGY plates, one isolate obtained by direct plating on a MP500 plate, and one isolate obtained by the chemostat enrichment) were identical to the sequence of the R3 band (Table 1). No isolate was found to have a sequence identical to the sequence of the R1 band. Table 1 also shows the results of the database searches; a closely related bacterial strain in the databases is shown for each isolate.

Phylogeny of LmPHs. To examine phylogenetic relationships among the LmPHs, an unrooted neighbor-joining tree was constructed by using the partial amino acid sequences of the subunits (Fig. 3). LmPHs in rC4 and rC8 had the same amino acid sequence, although the nucleotide sequences of the structural genes were not identical. Likewise, the amino acid sequence of LmPH in rN9 was identical to the amino acid sequences of LmPHs in rC7, rP2, and rN7, although the nucleotide sequence of the LmPH gene in rN9 was different from the nucleotide sequences of the LmPH genes in the other three isolates. The tree suggests that there are three distinct groups of LmPHs. LmPHs encoded by the major TGGE bands from the phenol-digesting activated sludge, bands P2 and P3, belong to groups II and I, respectively. Most of the previously identified LmPHs were affiliated with group III; in particular, LmPHs from the Pseudomonas strains formed a peculiar cluster in group III (designated the Dmp family), which also contained LmPH of rB1, the dominant isolate after batch enrichment. TbmD, the toluene/benzene 2-monooxygenase from Pseudomonas sp. strain JS150 (21), was placed in group I. 

View larger version (16K): [in this window] [in a new window]   FIG. 3.   Unrooted neighbor-joining tree based on the partial amino acid sequences of LmPHs. TbmD, the largest subunit of toluene/benzene 2-monooxygenase from Pseudomonas sp. strain JS150 (21); DmpN, LmPH from Pseudomonas sp. strain CF600 (27); PhhN, LmPH from P. putida P35X (26); PhlD, LmPH from P. putida H (15); MopN, LmPH from A. calcoaceticus NCIB8250 (10); PoxD, LmPH from R. eutropha E2 (16); PheA4, LmPH from P. putida BH (37). C. testosteroni R5 (previously assigned to the genus Alcaligenes) and B. cepacia E1 (previously assigned to the genus Achromobacter) are phenol-degrading bacteria that were isolated from another activated sludge after chemostat enrichment (41). Nucleotide sequences of the partial LmPH gene sequences of strains R5 and E1 were also determined in this study. Boxes indicate amino acid sequences encoded by the dominant TGGE bands in Fig. 1. Bar = 0.019 substitution per amino acid site.

Physiological analyses of the isolates. Growth in MP500 medium was not a common phenotypic trait among the isolates, because many isolates harboring mPH could not grow in MP500 medium (Table 1). One isolate, rB1, which was the predominant strain after batch enrichment, exhibited the fastest growth in MP500 medium. Some isolates which could not grow in MP500 medium, including rN7, rC1, rC4, and rC7, could thrive when phenol was the sole carbon source in chemostat cultures.

View larger version (20K): [in this window] [in a new window]   FIG. 4.   Phenol-oxygenating activities of phenol-digesting activated sludge and of bacterial isolates as a function of phenol concentration. AS, phenol-digesting activated-sludge sample obtained on day 20.

	DISCUSSION

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In this study, TGGE of PCR-amplified 16S rDNA was used to identify the dominant phenol-degrading bacteria in phenol-digesting activated sludge. This technique has frequently been used to analyze the structures of microbial communities (25). Although the bias in PCR could interfere with accurate estimation of the sizes of bacterial populations, we recently demonstrated that such bias was minimal in a PCR-TGGE analysis of 16S rDNA (45).

Since it has been suggested that mPHs are predominant in the environment (28, 31), a set of primers for PCR amplification of the genes for LmPHs was designed. PCR performed with these primers resulted in amplification of the LmPH genes from 17 of 27 isolates. Thus, our method did not detect all of the phenol hydroxylase genes present in the sludge populations. Nevertheless, the major LmPH genes detected by PCR and TGGE (bands P2 and P3) may correspond to the dominant genes in the sludge populations because they are encoded by the R2 and R3 bacteria, which were shown to be dominant in the activated sludge by the PCR-TGGE analysis of 16S rDNA.

Due to its simplicity, the batch culture enrichment method is most commonly used to isolate microbes that are capable of degrading a variety of hydrocarbons. However, it has recently been pointed out that this enrichment method is highly selective, resulting in the isolation of a few microbial species from diverse natural microbial populations (8, 9). Alternative methods, such as direct plating (8, 9) and enrichment in chemostat cultures, (41) have been proposed. Since so far no study has been conducted to rigorously compare the isolation efficiencies of these methods, we examined the differences in the genetic and phenotypic traits of bacteria isolated by four different methods. As expected, the batch enrichment method was highly selective compared with the other methods and resulted in isolation of only two types of strains, one of which (rB1) was predominant in the batch culture and grew most rapidly in MP500 medium (Table 1). It should be noted that the batch enrichment technique was the only method with which the populations that were dominant in the activated sludge were not isolated. Therefore, this method should be considered for selecting peculiar bacteria with a particular catabolic trait (e.g., rapid growth in a batch culture).

The bacteria isolated by the different methods produced different rep-PCR patterns. This observation implies that each isolation method had its own bias. The strains belonging to and related to the rN7 group were obtained at the highest frequency by direct plating on dCGY plates. However, rC4, representing the second most dominant population in the activated sludge, could be isolated only after enrichment in a chemostat culture. This result suggests that it is important to use multiple isolation methods to recover the desired strains from a complex microbial community. In particular, direct plating and enrichment in chemostat cultures are the two methods recommended for isolating dominant bacterial populations in the environment.

Bacteria possessing group I LmPHs, including Comamonas testosteroni R5, Burkholderia cepacia E1 (41), and rN7, exhibited phenol-oxygenating activities characterized by low K_s values. Similarly, bacteria possessing group II LmPHs, including R. eutropha E2 (41) and rC4, exhibited low-K_s phenol-oxygenating activities. In contrast, the phenol-oxygenating activities in bacteria possessing group III LmPHs were characterized by high K_s values. It has been reported that the K_s and K_SI values for phenol-oxygenating activities in intact cells are determined by the characteristics of mPH (16). Therefore, it is likely that the K_s values for phenol-degrading bacteria containing group I or II LmPHs are low, while the K_s values for bacteria containing group III LmPHs are high.

All of the strains closely related to the genera Acinetobacter and Pseudomonas in the  subclass of the Proteobacteria harbored LmPHs affiliated with group III, while bacteria harboring group I LmPHs were members of the  subclass of the Proteobacteria. This observation suggests that horizontal transfer of the mPH genes between members of two different subclasses of Proteobacteria does not occur often.

It has been suggested that the  subclass of the Proteobacteria is the dominant subclass in activated sludge (36, 39), and the Rhodocyclus group of the  subclass of the Proteobacteria has been considered important for phosphate removal in a sequential batch reactor (2). The importance of members of the  subclass of the Proteobacteria for degradation of phenol in activated sludge was suggested by the results of this study.

Phenol-degrading strains harboring the Dmp family of enzymes have been isolated from a variety of habitats; Pseudomonas sp. strain CF600 was isolated from activated sludge in England (12), P. putida P35X was isolated from river mud in England (18), P. putida H was isolated from river water in Germany (19), and P. putida BH was isolated from activated sludge in Japan (14). Hence, it is surprising that mPHs of these strains exhibit such high levels of homology (34). However, all of these strains were isolated after enrichment in batch cultures containing phenolic compounds at concentrations higher than 1 mM, as was rB1 in this study. This observation suggests that conventional enrichment in a batch culture selects bacteria with specific genotypes for a catabolic enzyme, even if different environmental samples are used as the sources for isolation.

We concluded that the genotypes and phenotypes of the functionally dominant phenol-degrading populations in activated sludge were much different from the genotypes and phenotypes of the representative phenol-degrading bacteria characterized previously in several laboratories. This conclusion should be true for other catabolic populations in the natural environment, and thus our findings could shed light on the bacterial populations responsible for degrading various compounds in the natural environment.