INTRODUCTION

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A major constraint to the development of successful bioremediation technology is the limited ability to quantify bioavailable levels of contaminants to determine whether the concentrations are within the range for potential microbial degradation. There are presently no extraction techniques that are well correlated with bioavailability because current techniques remove some fraction of the sorbed or nonaqueous-phase contaminant which may be physically and chemically unavailable to microbial populations (9, 19, 21, 34). Thus, there is considerable interest in the development of assays that will determine contaminant bioavailability. One such assay uses bioluminescent reporter organisms. In these reporter organisms, the bioluminescence operon (lux) is inserted into biodegradation or resistance pathways of interest so that the lux genes are expressed concurrently and thus can be used to monitor the real-time genetic expression of the pathway of interest. Continued development of this bioluminescent reporter system can meet the urgent need in bioremediation research for tools to not only quantify bioavailable pollutants but also to perform in situ monitoring of degradation in the environment.

Luminescence is produced by the reporter bacterium in a luciferase-catalyzed reaction in response to the oxidation of reduced flavin mononucleotide (FMNH₂) and a long-chain aldehyde (15). A number of bioluminescent reporter bacteria have been engineered to quantify bioavailable concentrations of organic contaminants (1, 4, 12, 14, 26, 29, 30, 32) and heavy metals (22, 25). The majority of these organisms use either the luxAB genes encoding the bacterial luciferase or the complete lux operon (luxCDABE), which also encodes the fatty acid reductase complexes required to produce the aldehyde substrate (16, 24). Use of the complete lux operon allows the nondestructive tracking of a specific organism or monitoring of the presence or utilization of organic or heavy-metal compounds in environmental systems without the exogenous addition of the aldehyde substrate. The disadvantage in using the entire operon is that generation of the aldehyde is an ATP- and NADPH-dependent process that not only increases the metabolic load of the cell (5, 10) but also depends upon the channeling of fatty acids into the luminescence system. In addition, for all bioreporters, energy must be diverted to different components of the electron transport system for luminescence production (22). Therefore, the intensity of luminescence can reflect environmental and physiological changes that affect bioreporter metabolic activity.

A number of authors have observed the sensitivity of bioluminescence to various physiological and environmental factors (3, 4, 6, 8, 15, 17, 24). Previous work has also demonstrated the potential for utilization of lux genes for detection in static systems over a small range of concentrations where physiological and environmental conditions can be tightly controlled. For example, Heitzer et al. (7, 8) and Sticher et al. (30) observed a linear relationship between substrate concentration and luminescence, while Rattray et al. (24) and Meikle et al. (18) have looked at the effect of cell density on luminescence. However, no comprehensive studies have been conducted to evaluate the influence of a range of parameters on a single organism. Thus, there remains a need to evaluate physiological and environmental parameters on a broader scale in order to anticipate factors which might cause deviations from the predicted response for applications of the lux bioreporters in situ. In addition, for applications to dynamic systems, it is not enough to consider only the magnitude of luminescence at a specific sample time, as has been done in previous work. Rather, the maximum potential luminescence, the lag time to maximum luminescence, and the relationship of these values to expression of the genes of the regulatory pathway of interest must be understood.

The objective of this research was to conduct a comprehensive overview of the effect of a range of factors on several parameters with a single indicator organism to evaluate the potential use of lux bioreporters as analytical tools or metabolic indicators in a dynamic system such as a saturated flow column. The factors evaluated were cell growth stage, cell density, substrate concentration, oxygen tension, and the interference of potential background carbon substrates from a soil system. Parameters measured included the magnitude of luminescence, the lag time in attaining the maximum response, and the correlation of this response with comparable changes in substrate degradation rates and lag times.

The indicator bioreporter used was Pseudomonas putida RB1353, developed by Burlage et al. (4) and containing two plasmids, the NAH7 plasmid and the constructed nah-lux reporter plasmid, pUTK9. NAH7 is an 83-kb plasmid with genes for naphthalene catabolism in two operons referred to as the upper and lower pathways. The upper pathway degrades naphthalene to salicylate, while the lower pathway is responsible for salicylate metabolism (33). Plasmid pUTK9 is subcloned with a fusion between the promoter from the upper pathway of NAH7 and the luxCDABE genes of Vibrio fischeri (4). Strain RB1353 is an identical sister clone to RB1351 described by Burlage et al. (4a). Burlage et al. (4) demonstrated that light production from RB1351 was directly correlated with naphthalene catabolism and documented a 20-fold increase in light from colonies exposed to naphthalene vapors.

Salicylate was chosen as the substrate for this study because it is responsible for the induction of both nah operons and its high solubility allows investigations of the effects of a wide range of substrate concentrations. Possible inhibitory effects of the lux operon on expression of the nah genes were evaluated by comparison of RB1353 to the parent strain, Pseudomonas putida HK53 (4). Strain HK53 was formed by mating NAH7 into P. putida PB2440.

	MATERIALS AND METHODS

Top Abstract Introduction Materials and Methods Results and Discussion References

Bacterial strains and media. P. putida RB1353 with stable plasmids NAH7 and pUTK9 (kanamycin resistance) and Pseudomonas putida HK53 (rifampin resistance), were kindly supplied by Robert Burlage, Oak Ridge National Laboratories, Oak Ridge, Tenn., and Gary Sayler, Center for Environmental Biotechnology, University of Tennessee, Knoxville, Tenn., respectively. The strains were maintained in Luria broth (LB) containing 10 g of tryptone, 5 g of yeast extract, and 10 g of NaCl in 1 liter of deionized H₂O with the pH adjusted to 7.0. The medium was supplemented with either 100 mg of kanamycin sulfate liter¹ to select for plasmid pUTK9 or 50 mg of rifampin liter¹ for strain PB2440 as needed. Agar plates were made by amending LB with 1.5% Bacto Agar (Difco Laboratories, Detroit, Mich.). Mineral salts broth (MSB), used for growth on sodium salicylate, contained (in grams per liter) KH₂PO₄, 1.5; Na₂HPO₄, 0.5; MgSO₄ · 7H₂O, 0.2; NH₄Cl, 2.5; FeCl₃, 3 × 10⁴; and CaCl₂ · 2H₂O, 0.013. Sodium salicylate and antibiotics were purchased from Sigma Chemical Co., St. Louis, Mo. Bacterial strains were stored frozen in glycerol (12.5% glycerol), and fresh cultures were inoculated from these stocks for each experiment to avoid plasmid loss. Bacterial strains were grown at 24°C with constant shaking at 120 rpm. Cell density was determined spectrophotometrically at 550 nm with a U-2000 spectrophotometer (Hitachi Instruments, Inc., Fremont, Calif.) and confirmed by viable plate counts of serial dilutions.

Cell preparation and experimental design. Bacteria cells used for luminescence and salicylate degradation assays were grown in LB amended with antibiotics appropriate for the individual strain as specified above. Cultures were inoculated at a density of 10⁴ CFU ml¹ from a 30-h preculture in the same medium and allowed to grow until the desired growth stage according to a previously determined growth curve. Growth was characterized by an initial lag period, logarithmic growth, and a relatively long deceleration phase followed by stationary and death phases. Cells were grown in LB prior to transfer to salicylate because cells grown under these conditions produced a light signal at least twice the intensity of that produced by cells grown originally in salicylate. After removal from LB, the cells were washed twice in saline (0.85% NaCl), resuspended in MSB, and amended with sodium salicylate. Cultures for each treatment were prepared in triplicate in 250-ml Erlenmeyer flasks with 30 ml of culture per flask and placed on the shaker. Samples (1 ml) were removed from each flask at each sampling time and analyzed for luminescence or sodium salicylate.

Quantitation of luminescence and sodium salicylate. (i) Luminescence. Samples (1 ml) were analyzed for luminescence in 7-ml plastic scintillation vials in a 1600TR Tri Carb liquid scintillation analyzer (Packard Instrument Co., Meriden, Conn.). Samples were placed into a vial and immediately counted for 1 min in the single-photon mode, generating relative values expressed in counts per minute. Repeated counting of a single vial or prolonged incubation (longer than 30 min) of samples in plastic vials was avoided because such treatment was found to cause elevated counts compared to those of samples read immediately after removal from culture flasks. Luminescence values obtained at each sampling time were plotted as a function of time, and the peak luminescence value and peak time were recorded for each experiment. Peak time was defined as the time required to attain the maximum luminescence. Alternatively, total luminescence was calculated by integrating under the curve to evaluate the relationship between total luminescence generated and the salicylate degradation rate.

(ii) Salicylate. Salicylate samples (1 ml) were added to 0.5 ml of 1 M NaOH to inhibit further degradation. Before analysis, the samples were centrifuged at 16,000 × g for 10 min to remove cell debris. The sodium salicylate concentrations were then determined from a standard curve by using the U-2000 spectrophotometer at 296 nm and plotted as a function of the sampling time. Degradation curves were characterized by an initial lag followed by a period of increasing degradation rate until the maximum degradation rate (V_max) was attained with approximately 80% of the initial substrate remaining. A decrease in the degradation rate typically began with 35% of the original substrate remaining. Thus, V_max was defined as the regression of the linear portion of the degradation curve between 35 and 80% of the original sodium salicylate concentration. Maximum degradation was repeatedly found within this interval regardless of initial substrate concentration. Induction or lag time for salicylate degradation was defined as the time required to degrade the initial 20% of substrate before V_max was attained.

Growth stage, cell density, and substrate experiments. Three series of experiments were conducted with RB1353(pUTK9, NAH7) to evaluate the influence of growth stage, cell density, and substrate concentration on the following four parameters: magnitude of luminescence, peak luminescence time, maximum salicylate degradation rate (V_max), and degradation induction time. Growth stage experiments were conducted with cells from the log, deceleration, stationary, and death phases obtained from a continuously growing LB culture as described previously. Cells removed at each growth stage were washed twice in saline, diluted to a standard concentration of 10⁷ CFU ml¹ in MSB, amended with 20 mg of sodium salicylate liter¹, and sampled to determine the luminescence and degradation rate. The influence of cell density was evaluated by using 10⁶, 10⁷, and 10⁸ CFU of RB1353 ml¹ obtained from a stationary-phase LB culture. Cells were exposed to 20 mg of sodium salicylate liter¹, and luminescence and salicylate degradation data were determined. Finally, substrate concentration experiments were conducted in the same way with 10⁷ CFU of stationary-phase RB1353 cells ml¹. Sodium salicylate was supplied at a range of concentrations from 0 to 40 mg liter¹, and data were collected as described previously.

Interference from background carbon substrates. Luminescence produced in response to nonspecific carbon substrates was measured to evaluate potential false signals produced in a dynamic soil system. Stationary-phase cells were used for all experiments to maximize potential interference, since cells of this growth phase were found to produce the strongest luminescence response (see Results). Triplicate flasks containing 4.5 × 10⁷ CFU of stationary-phase RB1353 cells ml¹ were exposed to either 20 mg of sodium salicylate liter¹, 50% LB, or soil extract broth. Soil extract broth was prepared as described in the Handbook of Microbiological Media (2) with a sandy loam from an oak and pine forest in Rose Canyon (Santa Catalina Mountains, Tucson, Az.) and was diluted 1:1 with MSB. Soil extract was used to evaluate potential interference produced in response to the presence of soil organic material as a substrate, and LB was chosen as the rich substrate preferred for growth of this organism. Luminescence was monitored for 5 h.

Influence of lux genes on degradation rates. Comparisons were done between P. putida RB1353(NAH7, pUTK9) and the parent strain, P. putida PB2440(NAH7), to evaluate the influence of the lux gene plasmid pUTK9 on salicylate degradation behavior. Simultaneous salicylate degradation assays were conducted as described above with the same cell density and growth stage for both strains. In addition, degradation rates for PB2440 were determined for both deceleration- and stationary-phase cells to evaluate whether changes in cell growth phase had similar effects on V_max for both RB1353 and PB2440.

DO concentration experiments. Two series of experiments were designed to determine the effects of dissolved-oxygen (DO) tension on the salicylate degradation rate and the corresponding luminescence response. Stationary-phase cells were used for all experiments, prepared as described previously, and added to the MSB salicylate (20 mg liter¹) broth to give an approximate cell density of 10⁷ CFU ml¹ for each assay unless stated otherwise. DO concentrations were determined by using a micro-oxygen electrode and oxygen meter (Microelectrodes, Inc. Bedford, N.H.). Calibration was done with buffer sparged with N₂ gas and ambient air for 0- and 8.5-ppm values, respectively.

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	RESULTS AND DISCUSSION

Top Abstract Introduction Materials and Methods Results and Discussion References

Growth phase. The growth phase was found to have a significant impact on both the magnitude of the luminescence response and the salicylate degradation rate for strain RB1353 (Fig. 1). Luminescence produced by stationary-phase cells was 1 order of magnitude greater than that produced by deceleration phase cells and 2 orders of magnitude greater than that produced from either log-phase or late-stationary-phase cells. The implications of these results are twofold. First, maximum sensitivity for detection of low substrate concentrations or low cell numbers will be attained in a static system with early-stationary-phase RB1353 cells. As a corollary to this, these results suggest that the effect of growth phase on the lux response should be evaluated for all new lux strains to achieve maximum sensitivity. Second, quantitation of carbon substrate, based on the light response, must take into consideration the growth potential of the cells in a dynamic system. As such, alternate standard curves must be developed for actively growing as opposed to stationary-phase cells. The maximum salicylate degradation rate was also found to vary with growth phase, with maximum rates being detected from logarithmic- and deceleration-phase cells (Fig. 1). The rates decreased with stationary-phase cells and continued to decline as cells entered the death phase. The shortest lag times for both V_max and peak luminescence corresponded to the growth phase associated with maximum V_max values (Fig. 1).

View larger version (20K): [in this window] [in a new window]   FIG. 1.   Effect of cell growth stage on peak luminescence, Vmax, and the associated lag times of RB1353.

Cell density. A positive correlation was observed with increasing cell density for both peak luminescence and V_max (Fig. 2). A particularly good linear correlation was found between peak luminescence and cell number (r² = 0.9973). Lag times were also affected by changes in biomass. An inverse relationship was observed between cell density and lag times associated with both peak luminescence and V_max (Fig. 2). Significant shifts in lag time, from 60 to >200 min, were observed in response to 1-log-unit changes in cell counts. Thus, in dynamic systems, changes in cell number will have significant effects on both the magnitude and timing of luminescence and V_max.

View larger version (19K): [in this window] [in a new window]   FIG. 2.   Effect of cell density on peak luminescence, Vmax, and the associated lag times of RB1353.

Substrate concentration. A strong positive linear correlation was found between substrate concentration and both luminescence (r² = 0.98) and V_max (r² = 0.98) (Fig. 3). Lag times increased for both peak luminescence and V_max in response to increasing substrate concentration (Fig. 3). Thus, both the magnitude and lag time of peak luminescence and V_max are affected by substrate concentration.

View larger version (18K): [in this window] [in a new window]   FIG. 3.   Effect of sodium salicylate concentration on peak luminescence, Vmax, and the associated lag times of RB1353.

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Influence of background carbon substrates. Experiments were conducted to evaluate possible false signals generated by nonspecific carbon sources for application of the luminescence assay to in situ experiments such as saturated flow soil columns. MSB with no carbon substrate served as a negative control to indicate background luminescence levels, and 20 mg of sodium salicylate liter¹ was the positive control. Maximum luminescence produced by 4.6 × 10⁷ stationary-phase cells in response to the nonspecific carbon sources evaluated was less than 0.1% of the signal produced by the same number of cells in response to 20 mg of sodium salicylate liter¹ (Table 1). No significant luminescence was detected in response to a 5-h incubation with the soil extract broth, and a maximum increase of 15 times background (from 48 to 749 cpm) was observed over the 5-h incubation period in the more complex medium, 50% LB. The maximum LB response (749 cpm) was still only half the luminescence peak generated by 55% the number of cells in response to 1 mg of sodium salicylate liter¹ (1,500 cpm [Fig. 3]). Thus, no significant interference from nonspecific carbon sources is anticipated during a typical 5-h assay period when analyzing substrate levels of 1 mg liter¹ or greater.

Influence of lux genes on nah gene expression. Potential impacts of the lux operon on expression of the nah genes were evaluated to assess possible negative aspects to its use as a bioreporter system. In a simultaneous salicylate degradation assay, the V_max for RB1353 was found to be slightly lower than for the parent strain, PB2440 (2.7 and 3.1 mg liter¹ h¹, respectively), but the difference was insignificant compared to changes caused by factors such as growth stage, cell number, and substrate concentration. Growth phase experiments with the parent strain revealed a similar pattern to RB1353, with a higher V_max observed from deceleration-phase cells than from stationary-phase cells (data not shown). Thus, the presence of the pUTK9 plasmid bearing the lux genes does not appear to have a significant effect on the degradative behavior of the engineered RB1353 strain under ambient conditions.

DO tension. Results from the first oxygen experiment (Fig. 4A) show a significant drop in DO tension during incubation for flasks filled to 90% of capacity, while flasks filled to 12% maintained a fairly constant oxygen level. The lower oxygen levels in the former flasks had no effect on salicylate degradation behavior (Fig. 4B). Limited oxygen availability was most apparent when consumption was highest, most probably due to reduced diffusion rates through the liquid media. Diffusion through water is in the range of 10⁵ cm² s¹, compared to 10³ cm² s¹ for air (31). In contrast, the luminescence response was enhanced by the reduced oxygen availability (Fig. 4C). This response was in contrast to anticipated behavior based on the observation that for two different luminescent Photobacterium spp., the luciferase reaction required as much as 10 to 20% of the total available oxygen (6). This competition for oxygen between luciferase and other metabolic enzymes has been previously identified as a possible source of complex luminescence behavior (12). A similar enhancement was documented previously in Vibrio fischerii, the donor organism for the lux genes, but a simultaneous inhibition of glucose metabolism was also observed. These results led to the hypothesis that luciferase has a lower K_m value for oxygen and therefore was functioning as a substitute for cytochrome as the terminal carrier of electrons to oxygen under microaerophilic conditions (6). However, for RB1353, substrate degradation was unaffected. Thus, the enhanced luminescence may be because the slightly reduced conditions in this system enhanced FMNH₂ or aldehyde availability.

View larger version (22K): [in this window] [in a new window]   FIG. 4.   Effect of limited oxygen diffusion on DO concentration (A), sodium salicylate degradation (B), and luminescence (C) for RB1353 in response to 20 mg of sodium salicylate liter1.

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View larger version (29K): [in this window] [in a new window]   FIG. 5.   DO concentrations during incubation for assays with different initial DO levels. (A) Unsparged controls; (B) initial DO concentrations of 2.5 to 6.2 mg liter; (C) initial DO concentrations less than or equal to 1.5 mg liter1.

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View larger version (20K): [in this window] [in a new window]   FIG. 6.   Effect of initial DO levels on Vmax (A), total luminescence (B), and the ratio of luminescence to Vmax, (C) for RB1353 in response to 20 mg of sodium salicylate liter1.

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Regulation of biodegradation and luminescence expression. One possible explanation for the observed effects of substrate and oxygen concentration on salicylate degradation and luminescence behavior can be found through an evaluation of the nahR regulatory system in combination with general bacterial global regulatory mechanisms. Bacterial cells must be able to adapt rapidly to a wide range of fluctuations in environmental conditions in order to survive. An important survival strategy involves the meticulous control of numerous operons to avoid the waste of energy resulting from the synthesis of excess mRNA or enzymes (28). Expression of the sal and lux genes in RB1353 is controlled by the lower-pathway promoter, P_sal, and the upper-pathway promoter, P_nah, respectively (4). Both promoters have a site at bp 70 preceding the transcription start site that is recognized by the nahR gene regulatory protein (4). The nahR gene is transcribed constitutively, but the NahR protein is activated (NahRa) only following binding to the inducer, salicylate. Subsequent NahRa-concentration-dependent 10- to 50-fold increases in production of nah enzymes have been observed (33). Thus, it is logical that a linear relationship was observed between substrate concentration and both V_max and luminescence.

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