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Applied and Environmental Microbiology, September 2000, p. 3905-3910, Vol. 66, No. 9
0099-2240/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.

Starvation Improves Survival of Bacteria Introduced into Activated Sludge

Kazuya Watanabe,* Mariko Miyashita, and Shigeaki Harayama

Marine Biotechnology Institute, Kamaishi Laboratories, Heita, Kamaishi City, Iwate 026-0001, Japan

Received 13 March 2000/Accepted 21 June 2000


    ABSTRACT
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References

A phenol-degrading bacterium, Ralstonia eutropha E2, was grown in Luria-Bertani (LB) medium or in an inorganic medium (called MP) supplemented with phenol and harvested at the late-exponential-growth phase. Phenol-acclimated activated sludge was inoculated with the E2 cells immediately after harvest or after starvation in MP for 2 or 7 days. The densities of the E2 populations in the activated sludge were then monitored by quantitative PCR. The E2 cells grown on phenol and starved for 2 days (P-2 cells) survived in the activated sludge better than those treated differently: the population density of the P-2 cells 7 days after their inoculation was 50 to 100 times higher than the population density of E2 cells without starvation or that with 7-day starvation. LB medium-grown cells either starved or nonstarved were rapidly eliminated from the sludge. The P-2 cells showed a high cell surface hydrophobicity and retained metabolic activities. Cells otherwise prepared did not have one of these two features. From these observations, it is assumed that hydrophobic cell surface and metabolic activities higher than certain levels were required for the inoculated bacteria to survive in the activated sludge. Reverse transcriptase PCR analyses showed that the P-2 cells initiated the expression of phenol hydroxylase within 1 day of their inoculation into the sludge. These results suggest the utility of a short starvation treatment for improving the efficacy of bioaugumentation.


    INTRODUCTION
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References

Introduction of exogenous microorganisms into environments (bioaugmentation) has been attempted to improve agricultural productivity (32) and to accelerate bioremediation (33). Although bioaugmentation has proven useful in some cases, many reports have documented that inoculated microorganisms survived poorly and had only minor influences on the total ecosystem functions (13, 16, 36). Scientists have thus investigated factors governing their fate in various environments, and several important environmental factors, including physicochemical parameters (4, 25), nutrient availability (5, 8, 36, 40), grazing pressure (8, 10), and the existence of microniches (21), have been elucidated. In addition to these environmental factors, the physiological characteristics of inoculated microorganisms seem to play a role in their colonization and survival in introduced environments. If an inoculum can use a specific substrate that is unavailable to a majority of indigenous microorganisms, it may have a selective advantage in environments containing that specific substrate (32). To cite an instance, Ogunseitan et al. demonstrated that the addition of salicylate to soil sustained the density of inoculated naphthalene degraders for more than 30 days (20). Devliegher et al. also showed that the treatment of soil with certain detergents resulted in 100- to 1,000-fold increases in the density of a detergent-degradative inoculum (1).

Natural environments contain a diverse array of microorganisms that exhibit kinetically different catabolic activities (9, 34). One may consider that microorganisms which exhibit higher catabolic activities, e.g., higher Vmax/Ks values in Haldene's equation, grow faster in such environments and hence are more suitable for bioaugmentation. However, this hypothesis has not necessarily been proven. For instance, it was demonstrated that a bacterial strain that exhibited a higher phenol-degrading activity was less competitive than strains which exhibited lower activities after their introduction into phenol-digesting activated sludge (35). There may be important factors other than the catabolic activity which determine the survivability of microorganisms in the natural environment.

It is desirable for bioaugmentation, if a microorganism exhibiting high catabolic activities can be sustained at a high population density in the introduced environment. Previous studies have examined effects of prestarvation on the survival of inoculant cells; an improved survivability has been reported (30), while another study has documented that prestarvation exerted no significant effects on the survival (31). In the present study, we further investigated this approach. A phenol-degrading bacterium, Ralstonia eutropha E2, was subjected to different prestarvation treatments, and its survivability after the introduction into activated sludge was examined. We found that cells starved for a short period of time showed a high survivability. Therefore, some physiological changes of the bacterium during the prestarvation treatment were also analyzed to obtain insights into mechanisms for the high survivability.


    MATERIALS AND METHODS
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References

Bacterial strain. R. eutropha E2 was isolated previously from the activated sludge of a wastewater treatment facility in an oil refinery (34). This bacterium is capable of growing on phenol as the sole carbon source. The genes of this strain encoding phenol hydroxylase (poxABCDEF) have been cloned and characterized (12).

Cultivation and starvation conditions. Luria-Bertani (LB) medium (24) was inoculated with strain E2 and shaken at 100 rpm for 24 h at 30°C. Cells were collected by centrifugation at 10,000 × g for 5 min, and resuspended in MP medium containing (liter-1): 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. One liter of either LB medium or MP medium supplemented with 200 mg of phenol liter-1 (called MP200) was inoculated with approximately 107 E2 cells and shaken at 30°C. Cells were collected by centrifugation at a late-exponential-growth phase (with optical densities at 660 nm of 0.6 to 0.7 for the LB culture and 0.15 to 0.2 for the MP200 culture) and washed with MP before being suspended in MP at a cell concentration of 109 ml-1. The number of E2 cells was determined by epifluorescence microscopy after they were stained with 4',6'-diamidino-2-phenylindole (DAPI) (37).

Starvation of E2 cells was conducted by shaking the above cell suspensions at 100 rpm at 25°C.

Operation of laboratory activated-sludge units. Activated-sludge mixed liquor was obtained from the return sludge line of a municipal sewage treatment plant (Ohdaira, Kamaishi, Iwate, Japan). The activated sludge was infused into a laboratory unit composed of an aeration tank (3 liters) and a settling tank (2 liters) and acclimated to phenol by continuously supplying MP200 at a flow rate of 6 liters day-1. The hydraulic residence time (Trh) and phenol-loading rate were 0.5 day and 0.4 g liters-1 day-1, respectively. The concentration of mixed-liquor suspended solids in the aeration tank was maintained at between 1,800 and 2,000 mg liter-1 by wasting excess sludge from the aeration tank. The sludge residence time (Trs) was estimated to be approximately 10 days. Air was continuously supplied at a rate of 2 liters min-1, and the dissolved oxygen concentration was kept above 4 mg liter-1. The temperature was maintained at 25°C. Total direct counts of microorganisms in the sludge were determined by epifluorescence microscopy after flocs were disrupted with a blender in the presence of 5 mM sodium tripolyphosphate and the cells were stained with DAPI (37). The phenol concentration in the aeration tank was measured by a colorimetric assay with Phenol Test Wako (Wako Pure Chemicals) (36).

Quantitative PCR (qPCR). DNA was extracted from 5 ml of mixed liquor sampled from the aeration tank of the laboratory unit as described previously (36). The quantity and quality of the extracted DNA was checked by measuring the UV absorption spectrum of the DNA solution (24), and the DNA was finally dissolved in TE buffer (24) at a concentration of 100 µg ml-1.

Primers, P01f (5'-CACGCACCAGGAGACGCC-3') and P03r (5'-GTGCCCGGTGGTGCCTG-3'), were used in PCR to specifically detect the E2 population in activated sludge. These primers were designed from specific nucleotide sequences in the pox gene that were found by comparing the pox sequence (12) with the sequences of other phenol hydroxylase genes, namely, dmp from Pseudomonas sp. strain CF600 (19), phh from Pseudomonas putida P35X (18), phl from P. putida H (11), mop from Acinetobacter calcoaceticus NCIB8250 (3), and phc from Comamonas testosteroni R5 (29). A PCR with these primers amplifies a 260-bp fragment from strain E2.

The PCR amplification was performed with a Progene thermal cycler (Techne) by using a 50-µl mixture containing 1.25 U of Taq DNA polymerase (AmpliTaq Gold; PE Applied Biosystems), 10 mM Tris-HCl (pH 8.3), 50 mM KCl, 1.5 mM MgCl2, 0.001% (wt/vol) gelatin, 2% (wt/vol) formamide, each deoxynucleoside triphosphate at a concentration of 200 µM, 50 pmol of each primer, and 50 ng of activated-sludge DNA. The PCR conditions used were as follows: 10 min of the polymerase activation at 94°C followed by 35 cycles consisting of 1 min at 94°C, 1 min at 65°C, and 2 min at 72°C, and finally a 10-min extension at 72°C. If necessary, before PCR, the activated-sludge DNA solution was diluted with a negative-control DNA solution prepared from E2-noninoculated phenol-acclimated activated sludge. The PCR product (2 µl) was electrophoresed through 1.5% (wt/vol) agarose gel in TBE buffer and stained with SYBR green I (FMC Bioproducts). The band intensity was quantified using Gel Doc 2000 (Bio-Rad Laboratories) equipped with the Multianalyst Software (Bio-Rad Laboratories).

Reverse transcriptase PCR. RNA was extracted from 5 ml of mixed liquor sampled from the aeration tank of the laboratory unit by the guanidine thiocyanate procedure (22) with slight modifications. Mixed liquor was centrifuged at 10,000 × g for 5 min at 4°C, and the precipitated cells were suspended in 5 ml of a lysing solution (4 M guanidine isothiacyanate, 25 mM sodium citrate [pH 7.0], 1% [wt/vol] sodium dodecyl sulfate, and 1% [vol/vol] 2-mercaptoethanol) before they were incubated at 70°C for 30 min. The cell suspension was extracted with 6 ml of a phenol-chloroform solution (24) after 1 ml of 2 M sodium acetate solution (pH 4) was added. It was centrifuged at 10,000 × g for 5 min at 4°C, and the aqueous phase was recovered. Then, 6 ml of 2-propanol was added, and after the suspension was gently mixed, it was centrifuged at 5,000 × g for 10 min at 4°C. Nucleic acids were dissolved in 10 ml of diethyl pyrocarbonate (DEPC)-treated TE buffer (24), and 5 ml of 7.5 M ammonium acetate and 30 ml of ethanol were added before the solution was incubated at -20°C for 12 h. The solution was centrifuged at 10,000 × g for 10 min at 4°C, and the precipitate was dissolved in 1 ml of DEPC-treated TE containing 0.5% (wt/vol) sodium dodecyl sulfate. After the solution was treated with 1 ml of chloroform, nucleic acids were recovered by centrifugation in the presence of 70% (vol/vol) ethanol and 10 mM sodium acetate (pH 6). Nucleic acids were dissolved in 0.1 ml of DEPC-treated TE and treated with 5 U of DNase (RNase-free DNase I; Takara) at 25°C for 30 min. Subsequently, the solution was extracted with 200 µl of the phenol-chloroform solution. RNA was precipitated by centrifugation at 15,000 × g for 10 min at 4°C in the presence of 70% (vol/vol) ethanol, washed with 70% ethanol, and dissolved in DEPC-treated TE. The quantity and quality of the RNA were checked by measuring the UV absorption spectrum (24), and it was finally dissolved at a concentration of 100 µg ml-1.

The extracted RNA was subjected to reverse transcription (RT) and subsequent PCR amplification using a Progene thermal cycler. An RT mixture (20 µl) contained 5 U of reverse transcriptase (XL; Takara), 50 mM Tris-HCl (pH 8.3), 40 mM KCl, 5 mM MgCl2, each deoxynucleoside triphosphate at a concentration of 1 mM, 50 pmol of random 9-mers, 1 U of RNase inhibitor (Takara), and 100 ng of the extracted RNA. RT was carried out at 30°C for 10 min, 50°C for 30 min, 99°C for 5 min, and 5°C for 5 min. The RT mixture was subsequently mixed with a PCR mixture (80 µl) containing 2.5 U of Taq DNA polymerase (AmpliTaq Gold), 10 mM Tris-HCl (pH 8.3), 50 mM KCl, 1.5 mM MgCl2, 50 pmol of primer P01f, and 50 pmol of primer P03r and then subjected to PCR amplification using the thermal cycles described above. The PCR (2 µl) product was electrophoresed through 1.5% (wt/vol) agarose gel in TBE buffer and stained with SYBR green I before being photographed.

Physiological analyses. Microscopic observation of E2 cells was conducted using an Optiphot phase-contrast microscope equipped with a Microflex automatic camera system (Nikon).

The cell surface hydrophobicity (i.e., the capacity to adhere to hydrophobic chromatography resins) and the floc-adhesion capacity (i.e., the capacity to adhere to sludge flocs) were measured at 25°C using the above-described cell suspensions. Activated sludge mixed liquor (1 ml) obtained from the aeration tank was centrifuged at 1,000 × g for 5 min and resuspended in 5 ml of MP medium containing cycloheximide at 250 µg ml-1. The cell suspension (0.5 ml) was mixed with 0.5 ml of either MP (control), the sludge suspension, or MP containing 10 mg of hydrophobic resins (Phenyl Sepharose 6 Fast Flow Lab Pack, low sub; Pharmacia LKB) per ml. The mixture was gently inverted for 1 min and settled for 5 min. The supernatant was recovered, and the cells in the supernatant were counted by the DAPI method. The number of bacteria in the supernatant of the sludge suspension was <107 cells ml-1. A percentage [Pad (%)] of cells adhering to hydrophobic resins or sludge flocs was estimated with the following equation: Pad(%) = 100 × (Nc - Na)/Nc, where Nc is the number of cells in the control supernatant, and Na is the number of cells in the supernatant after being mixed with hydrophobic resins or sludge flocs.

The respiration rate was measured at 25°C using a Clark-type oxygen electrode (5/6 Oxygraph; Gilson). The reaction cuvette was filled with 1.9 ml of MP medium. After the signal was stabilized, 100 µl of the cell suspension was added, and the respiratory oxygen consumption was monitored. The respiration rate was normalized by the dry cell weight that was gravimetrically determined after collecting the cells on a 0.22-µm-pore-size membrane according to the method of Machado and Grady (15). One unit of the respiration rate is defined as 1 µmol of oxygen consumed per min, while the specific activity is defined as the activity per gram of dry cells.

The phenol-oxygenating activity was measured at 25°C using 100 µl of the cell suspension and 5/6 Oxygraph as described previously (34). One unit of the activity is defined as 1 µmol of oxygen consumed per min, while the specific activity is defined as the activity per gram of dry cells.

The dehydrogenase activity was determined by the method of Ryssov-Nielsen (23) using the above-described cell suspension and 2,3,5-triphenyltetrazolium chloride. One unit of activity is defined as 1 µmol of triphenylformazan produced per h. The specific activity is the activity per gram of dry cells.

Statistics. Data were statistically analyzed by the Student t test (P = 0.05).


    RESULTS
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References

qPCR. We previously demonstrated the utility of a qPCR method for analyzing the population dynamics of a specific strain introduced into activated sludge (36). In qPCR, a DNA template is appropriately diluted before PCR, so that the band intensity of the PCR product reflects the concentration of the template (i.e., the number of target cells in the activated sludge). A DNA sample from the introduced activated sludge was diluted with a solution of DNA similarly extracted from nonintroduced activated sludge to keep the amplification efficiency constant (36).

Figure 1 shows an example of qPCR where the population density of strain E2 in activated sludge 1 day after inoculation was determined. The population density on day 0 was calculated from the number of inoculated cells (the activated-sludge sample on day 0 was obtained 5 min after the inoculation). In Fig. 1, the density on day 1 was estimated to be 2.5 × 106 cells ml-1 from the ratio of the intensity of lane 5 and that of lane 9 and the dilution rates of these samples. The lower detection limit was approximately 104 cells ml-1. No fragment was amplified from DNA extracted from E2-noninoculated phenol-acclimated activated sludge (lane 11), demonstrating the specificity of the PCR to detect the E2 population.


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  		FIG. 1.   Quantitative PCR for determining the population density of E2 in activated sludge inoculated with the P-0 cells. Lane 1, 50- to 2,500-bp DNA size marker (FMC Corp.); lane 2, activated sludge immediately after inoculation (day 0), the population density being (3.1 ± 0.4) × 107 cells ml-1 (mean ± the standard deviation; n = 3); lanes 3 to 6, 10- to 10,000-fold dilutions of the DNA extracted from the activated sludge on day 0; lane 7, activated sludge 1 day after inoculation (day 1); lanes 8 to 10, 10- to 1,000-fold dilutions of the DNA extracted from the activated sludge on day 1; lane 11, E2-noninoculated activated sludge (day -1); lane 12, E2 pure culture.

Survival of E2 cells in activated sludge. Table 1 summarizes the inocula used in this study. The activated sludge was acclimated to phenol for 20 days before it was inoculated with one of these inocula on day 0. This acclimation period was needed to obtain a reproducible community structure of the activated sludge (analyzed as described previously [38]). The total cell counts in the aeration tank ranged from 3 × 109 to 6 × 109 cells ml-1 throughout the experiment. The population dynamics of E2 cells introduced into phenol-acclimated activated sludge were analyzed by the qPCR (Fig. 2). Among the inocula, only the P-2 cells did not show a rapid decrease in cell number during the initial several days, while the other inocula showed multiphasic declines, i.e., initial rapid declines followed by slower declines.

                              
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  		TABLE 1.   Inocula used in this study



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  		FIG. 2.   Population dynamics of E2 cells introduced into phenol-acclimated activated sludge determined by qPCR. E2 was grown in LB (a) or MP200 (b) medium. Symbols: open circle , not starved; black-lozenge , starved for 2 days; black-triangle, starved for 7 days. The dashed line represents a sludge washout curve (Trs = 10 days), while the dotted line represents a hydraulic washout curve (Trh = 0.5 day) (36). The means of three determinations are shown, and the error bars indicate the standard deviations. The results were reproducible in two independent inoculation experiments.

Expression of Pox in activated sludge. The lower detection limit of the RT-PCR assay for the pox mRNA in phenol-acclimated activated sludge was examined by experiments in which serial dilutions of an E2 culture growing exponentially on phenol were mixed with the activated sludge and subjected to the RT-PCR assay; the limit was found to correspond to a population density of between 105 and 106 cells per ml. In those experiments, no pox fragment was amplified from RT-negative control samples in which reverse transcriptase was not added.

The RT-PCR detected the pox mRNA in activated sludge inoculated with the P-0 cells on days 0 and 1 (Fig. 3a). Although the pox mRNA was not detected in activated sludge inoculated with the P-2 cells on day 0 (just after the inoculation), it was detected between day 1 and day 7 (Fig. 3b). The pox mRNA was not detected afterward; this might be due to the decrease in the population density. The pox mRNA was not detected in the activated sludge inoculated with the other inocula (data not shown).


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  		FIG. 3.   RT-PCR showing the expression of the pox mRNA in activated sludge inoculated with P-0 cells (a) and with P-2 cells (b). M, 50- to 2,500-bp DNA size marker.

Physiological changes during starvation. Microscopic observation of the inocula revealed a reduction in cell size, a change in cell shape, and a loss of the motility during starvation (Table 1).

When E2 cells were grown in MP200, most of the cells became adhered to hydrophobic resins and sludge flocs after the 2- and 7-day starvations, demonstrating that the increases in cell surface hydrophobicity and in floc adhesion capacity occurred due to starvation (Fig. 4). In contrast, the increases in these properties were less apparent in the LB-grown cells.


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  		FIG. 4.   Percentages of cells adhering to hydrophobic resins (dotted bar) and sludge flocs (cross-hatched bar) as indices for the cell surface hydrophobicity and floc adhesion capacity, respectively. The means of three determinations are shown, and the error bars indicate the standard deviations.

Metabolic activities were measured during the starvation of E2 cells (Fig. 5). The respiration rate and the dehydrogenase activity are considered to reflect the energy level (2) and the level of reducing equivalents such as NADH and NADPH (14), respectively, in the cell. These activities in LB-medium-grown cells were higher than those in MP200-grown cells. These activities in the LB-medium-grown cells were gradually decreased during the starvation for 10 days, while in the MP200-grown cells they were initially decreased and reached basal levels by day 7. The phenol-oxygenating activity was slightly detected in MP200-grown cells starved for 2 days, while it was completely lost after the 7-day starvation.


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  		FIG. 5.   Changes in metabolic activities during starvation. (a) Respiration rate. (b) Dehydrogenase activity. (c) Phenol-oxygenating activity. Symbols: open circle , LB-medium-grown cells; black-lozenge , MP200-grown cells. The phenol-oxygenating activities of the LB-medium-grown cells were not detected. The means of three determinations are shown, and the error bars indicate the standard deviations.


    DISCUSSION
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References

Multiphasic declines in the number of bacteria introduced into activated sludge have been observed in previous studies (16, 17, 26, 35), and this trend was also observed with the inocula used in this study except for the P-2 cells. The P-2 cells showed a significantly higher survivability after introduction into the phenol-acclimated activated sludge (Fig. 2), and this inoculum did not show the initial rapid decline in the population density. Since the decline curve of the P-2 cells was similar to the sludge washout curve (Fig. 2), we expected that these cells were sustained in the activated sludge due to the rapid adhesion of them to sludge flocs, which protected them from washout and protozoan grazing (16, 27). This inference was supported by the comparison of the physiological states of the P-0 and P-2 cells. Since the survivability of the P-0 cells was low but improved 2 days after the starvation, physiological states important for the survivability seemed to be induced by the starvation. In fact, the cell surface of strain E2 changed during the starvation; it became hydrophobic 2 days after the starvation, and a high sludge adhesion capacity was generated in P-2 cells (Fig. 4).

Changes in cell physiology due to starvation have been reported previously; these include changes in cell shape (6, 31), RNA content (6), protein expression pattern (7), motility (39), amounts of extracellular polymers (41), and stress resistance (31). Similar events may have happened in the E2 cells after they were subjected to starvation. Interestingly, some changes occurred in LB-medium- and MP200-grown E2 cells at different time points, e.g., MP200-grown cells rapidly ceased swimming (Table 1) and became hydrophobic (Fig. 4). It is conceivable that the onsets of these changes were influenced by the quantity of reserved materials (Fig. 5).

Zita and Hermansson have demonstrated that the cell surface hydrophobicity correlates with the ability of the cell to adhere to sludge flocs (42). This trend was also observed in the present study, but the correlation was weak in the L-2 cells (Fig. 4). This observation implies that factors other than the cell surface hydrophobicity may also be involved in the adhesion of bacterial cells to sludge flocs, flagella, and fimbriae.

A few studies have examined the effects of starvation treatment on the survival of bacteria after the introduction into soil ecosystems. Van Elsas et al. (30) reported an enhanced survival of starved cells, although the study of Van Overbeek et al. showed no effects of the starvation treatment (31). It has been suggested that this inconsistency may have been due to the different starvation conditions used in these studies (31). Our present study suggested that this hypothesis of Van Overbeek et al. might be correct. We demonstrated that the duration of the starvation treatment largely affects the survival rate of inocula in activated sludge and that there exists an optimum duration of the starvation treatment for an optimum survival of an inoculum. Since the importance of the cell surface hydrophobicity for the bacterial adhesion to soil particles has been demonstrated by Stenström (28), we expect that an optimum starvation treatment would increase the survivability of bacteria introduced into soil ecosystems.

In addition to cell surface hydrophobicity, the expression of the phenol-oxygenating activity (i.e., phenol-degradative enzymes) seems to be important for the survival of bacteria in the phenol-digesting activated sludge. LB-medium-grown E2 cells exhibiting no phenol-oxygenating activity showed a rapid decline in the activated sludge with or without the starvation treatment. Upon the growth on MP200, the phenol-oxygenating activity was induced in E2 cells, but this activity was reduced strongly during the starvation for 2 days, and the expression of the pox mRNA in the activated sludge inoculated by the P-2 cells was not detected on day 0. However, the pox mRNA was detected 1 day after the inoculation of the P-2 cells, suggesting that the recovery of the phenol-oxygenating activity in the P-2 cells was quick. In contrast to the P-2 cells, the pox mRNA expression was not detected in the sludge inoculated with the P-7 cells, although the population densities of strain E2 in these two sludge samples on day 1 were not significantly different. The data thus suggested that the E2 cells became unable to rapidly express phenol hydroxylase in activated sludge during the starvation treatment between days 2 and 7. It is also conceivable that the complete depletion of phenol-degrading activity may have caused the loss of E2 cells to survive in the activated sludge.

In conclusion, we suggest the utility of a short starvation treatment for improving the efficacy of a microbial agent in the environment. This suggestion is supported by the data showing a good survival of the P2 cells (Fig. 2) and the expression of Pox by P2 in the activated sludge (Fig. 3). This study also suggests that the survival of a microorganism after the introduction into the environment is affected by the physiological states of cells to be introduced. Further studies are needed in order to obtain general views on the cell physiology important for the survival in the environment.


    ACKNOWLEDGMENTS

We thank Ikuko Hiramatsu for technical assistance, Robert Kanaly for assistance in preparation of the manuscript, and Mitsuhiro Konno (Ohdaira Wastewater Treatment Plant, Kamaishi City, Japan) for help in the sampling of activated-sludge mixed liquor.

This work was performed as a part of the Industrial Science and Technology Project, Technological Development of Biological Resources in Bioconsortia, supported by the New Energy and Industrial Technology Development Organization (NEDO).


    FOOTNOTES

* Corresponding author. Mailing address: Marine Biotechnology Institute, Kamaishi Laboratories, 3-75-1 Heita, Kamaishi City, Iwate 026-0001, Japan. Phone: 81-193-26-5781. Fax: 81-193-26-6592. E-mail: kazuya.watanabe{at}kamaishi.mbio.co.jp.


    REFERENCES
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References

1. Devliegher, W., M. A. S. Arif, and W. Verstraete. 1995. Survival and plant growth promotion of detergent-adapted Pseudomonas fluorescens ANP15 and Pseudomonas aeruginosa 7NSK2. Appl. Environ. Microbiol. 61:3865-3871[Abstract].
2. Eckenfelder, W. W., and D. J. O'Conner. 1961. Biological waste treatment. Pergamon, Oxford, England.
3. Ehrt, S., F. Schirmer, and W. Hillen. 1995. Genetic organization, nucleotide sequence and regulation of expression of genes encoding phenol hydroxylase and catechol 1,2-dioxygenase in Acinetobacter calcoaceticus NCIB8250. Mol. Microbiol. 18:13-20[CrossRef][Medline].
4. Evans, J., C. Wallace, and N. Dobrowolski. 1993. Interaction of soil type and temperature on the survival of Rhizobium leguminosarum bv. viciae. Soil Biol. Biochem. 25:1153-1160[CrossRef].
5. Fujita, M., M. Ike, and K. Uesugi. 1994. Operation parameters affecting the survival of genetically engineered microorganisms in activated sludge processes. Wat. Res. 28:1667-1672[CrossRef].
6. Givskov, M., L. Eberi, S. Møller, L. K. Poulsen, and S. Molin. 1994. Responses to nutrient starvation in Pseudomonas putida KT2442: analysis of general cross protection, cell shape, and macromolecular content. J. Bacteriol. 176:7-14[Abstract/Free Full Text].
7. Givskov, M., L. Eberl, and S. Molin. 1994. Responses to nutrient starvation in Pseudomonas putida KT2442: two-dimensional electrophoretic analysis of starvation- and stress-induced proteins. J. Bacteriol. 176:4816-4824[Abstract/Free Full Text].
8. Goldstein, R. M., L. M. Mallory, and M. Alexander. 1985. Reasons for possible failure of inoculation to enhance bioremediation. Appl. Environ. Microbiol. 50:977-983[Abstract/Free Full Text].
9. 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].
10. Habte, M., and M. Alexander. 1975. Protozoa as agents responsible for the decline of Xanthomonas campestris in soil. Appl. Microbiol. 29:159-164[Medline].
11. Herrmann, H., C. Moller, I. Schmidt, J. Mahnke, L. Petruschka, and K. Hahnke. 1995. Localization and organization of phenol degradation genes of Pseudomonas putida strain H. Mol. Gen. Genet. 247:240-246[CrossRef][Medline].
12. Hino, H., K. Watanabe, and N. Takahashi. 1998. Phenol hydroxylase cloned from Ralstonia eutropha strain E2 exhibits novel kinetic properties. Microbiology 144:1765-1772[Abstract/Free Full Text].
13. Ho, W. C., and W. H. Ko. 1985. Soil microbiostasis: effects of environmental and edaphic factors. Soil Biol. Biochem. 17:167-170[CrossRef].
14. Jørgensen, K. P. 1984. Determination of the enzyme activity of activated sludge by methylene blue reduction. J. Wat. Pollut. Control Fed. 56:89-93.
15. Machado, R. J., and L. Grady, Jr. 1989. Dual substrate removal by an anexic bacterial culture. Biotechnol. Bioeng. 33:327-337[CrossRef].
16. 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].
17. 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].
18. Ng, L. C., V. Shingler, C. C. Sze, and C. L. Poh. 1994. Cloning and sequencing of the first eight genes of the chromosomally encoded (methyl) phenol degradation pathway from Pseudomonas putida P35X. Gene 151:29-36[CrossRef][Medline].
19. Nordlund, I., J. Powlowski, and V. Shingler. 1990. Complete nucleotide sequence and polypeptide analysis of multicomponent phenol hydroxylase from Pseudomonas sp. strain CF600. J. Bacteriol. 172:6826-6833[Abstract/Free Full Text].
20. Ogunseitan, O. A., I. L. Delgado, Y. L. Tsai, and B. H. Olson. 1991. Effect of 2-hydroxybenzoate on the maintenance of naphthalene-degrading pseudomonads in seeded and unseeded soil. Appl. Environ. Microbiol. 57:2873-2879[Abstract/Free Full Text].
21. Postma, J., C. H. Hok-A-Hin, and J. A. van Veen. 1990. Role of microniches in protecting introduced Rhizobium leguminosarum biovar trifolii against competition and predation in soil. Appl. Environ. Microbiol. 56:495-502[Abstract/Free Full Text].
22. Puissant, C., and L. M. Houdebine. 1990. An improvement of the single-step method of RNA isolation by acid guanidinum thiocyanate-phenol-chloroform extraction. BioTechniques 8:148-149[Medline].
23. Ryssov-Nielsen, H. 1975. Measurement of the inhibition of respiration in activated sludge by a modified determination of the TTC-dehydrogenase activity. Wat. Res. 9:1179-1185[CrossRef].
24. 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.
25. Shonnard, D. R., R. T. Taylor, M. L. Hanna, C. O. Boro, and A. G. Duba. 1994. Injection-attachment of Methylosinus trichosporium OB3b in a two-dimensional miniature sand-filled aquifer simulator. Wat. Resource Res. 30:25-35.
26. Soda, S., M. Ike, and M. Fujita. 1998. Effects of inoculation of a genetically engineered bacterium on performance and indigenous bacteria of a sequencing batch activated sludge process treating phenol. J. Ferment. Bioeng. 86:90-96[CrossRef].
27. Soda, S., M. Ike, and M. Fujita. 1999. Adsorption of bacterial cells onto activated sludge flocs. J. Ferment. Bioeng. 87:513-518.
28. Stenström, T. A. 1989. Bacterial hydrophobicity, an overall parameter for the measurement of adhesion potential to soil particles. Appl. Environ. Microbiol. 55:142-147[Abstract/Free Full Text].
29. Teramoto, M., H. Futamata, S. Harayama, and K. Watanabe. 1999. Characterization of a high-affinity phenol hydroxylase from Comamonas testosteroni R5 by gene cloning, and expression in Pseudomonas aeruginosa PAO1c. Mol. Gen. Genet. 262:552-558[CrossRef][Medline].
30. Van Elsas, J. D., A. C. Wolters, C. D. Clegg, H. M. Lappin-Scott, and J. M. Anderson. 1994. Fitness of genetically modified Pseudomonas fluorescens in competition for soil and root colonization. FEMS Microbiol. Ecol. 13:259-272.
31. Van Overbeek, L. S., L. Eberi, M. Givskov, S. Molin, and J. D. van Elsas. 1995. Survival of, and induced stress resistance in, carbon-starved Pseudomonas fluorescens cells residing in soil. Appl. Environ. Microbiol. 61:4202-4208[Abstract].
32. Van Veen, J. A., L. S. van Overbeek, and J. D. van Elsas. 1997. Fate and activity of microorganisms introduced into soil. Microbiol. Mol. Biol. Rev. 61:121-135[Abstract].
33. Vogal, T. M. 1996. Bioaugmentation as a soil bioremediation approach. Curr. Opin. Biotechnol. 7:311-316[CrossRef][Medline].
34. 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.
35. 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[CrossRef].
36. Watanabe, K., S. Yamamoto, S. Hino, and S. Harayama. 1998. Population dynamics of phenol-degrading bacteria in activated sludge determined by gyrB-targeted quantitative PCR. Appl. Environ. Microbiol. 64:1203-1209[Abstract/Free Full Text].
37. Watanabe, K., M. Teramoto, H. Futamata, and S. Harayama. 1998. Molecular detection, isolation, and physiological characterization of functionally dominant phenol-degrading bacteria in activated sludge. Appl. Environ. Microbiol. 64:4396-4402[Abstract/Free Full Text].
38. Watanabe, K., M. Teramoto, and S. Harayama. 1999. An outbreak of nonflocculating catabolic populations caused the breakdown of a phenol-digesting activated-sludge process. Appl. Environ. Microbiol. 65:2813-2819[Abstract/Free Full Text].
39. Wei, X., and W. D. Bauer. 1998. Starvation-induced changes in motility, chemotaxis, and flagellation of Rhizobium meliloti. Appl. Environ. Microbiol. 64:1708-1714[Abstract/Free Full Text].
40. Wilson, M., and S. E. Lindow. 1995. Enhanced epiphytic coexistence of near-isogenic salicylate-catabolizing and non-salicylate-catabolizing Pseudomonas putida strains after exogenous salicylate application. Appl. Environ. Microbiol. 61:1073-1076[Abstract].
41. Wrangstadh, M., U. Szewzyk, J. Ostling, and S. Kjelleberg. 1990. Starvation-specific formation of a peripheral exopolysaccharide by a marine Pseudomonas sp., strain S9. Appl. Environ. Microbiol. 56:2065-2072[Abstract/Free Full Text].
42. Zita, A., and M. Hermansson. 1997. Effects of bacterial cell surface structures and hydrophobicity on attachment to activated sludge flocs. Appl. Environ. Microbiol. 63:1168-1170[Abstract].

Applied and Environmental Microbiology, September 2000, p. 3905-3910, Vol. 66, No. 9
0099-2240/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.


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