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Applied and Environmental Microbiology, July 2002, p. 3502-3508, Vol. 68, No. 7
0099-2240/02/$04.00+0     DOI: 10.1128/AEM.68.7.3502-3508.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.

Solid-Phase Contact Assay That Uses a lux-Marked Nitrosomonas europaea Reporter Strain To Estimate Toxicity of Bioavailable Linear Alkylbenzene Sulfonate in Soil

Kristian K. Brandt,^* Anders Pedersen, and Jan Sørensen

Section of Genetics and Microbiology, Department of Ecology, Royal Veterinary and Agricultural University, Frederiksberg, Denmark

Received 27 November 2001/ Accepted 4 April 2002

Top Abstract Introduction Materials and Methods Results Discussion References

 
Information about in situ toxicity of the bioavailable pools of adsorptive soil pollutants is a prerequisite for proper ecological risk assessment in contaminated soils. Such toxicity data may be obtained by assays allowing for direct exposure of introduced test microorganisms to the toxicants, as they appear in solid solution equilibria in the natural soil. We describe a novel sensitive solid-phase contact assay for in situ toxicity testing of soil pollutants based on a recombinant bioluminescent reporter strain of Nitrosomonas europaea. A slurry of the reporter strain and soil sample was shaken for 1 h, after which bioluminescence was measured either directly (soil slurry protocol) or in the supernatant obtained after centrifugation (soil extract protocol). The assay was validated for both protocols by using linear alkylbenzene sulfonate (LAS) as a toxic and adsorptive model compound in the soil samples. Interestingly, LAS showed the same toxicity to the reporter strain with either soil incubation (both protocols) or pure culture, suggesting that adsorbed LAS pools contributed to the observed toxicity. The solid-phase contact assay that used the reporter strain of lux-marked N. europaea was slightly more sensitive for the detection of LAS toxicity in soil than activity-based assays targeting indigenous nitrifiers and much more sensitive than assays targeting indigenous heterotrophic microbes. We conclude that the new solid-phase contact assay, which is based on direct interaction of the test microorganisms with bioavailable pools of the toxicants in soil, provides a most sensitive and relevant method for evaluating the in situ toxicity and assessing the risks of soil contaminants.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Autotrophic ammonia-oxidizing bacteria (AOB) belonging to the ß-subclass of Proteobacteria are responsible for the initial rate-limiting step of nitrification activity in soils, thereby playing a key role for terrestrial N cycling (14, 31, 34). Furthermore, the bacteria have been found to be highly sensitive to a range of common soil pollutants (7, 10, 19, 36). As a result, AOB may be considered sensitive and relevant indicator bacteria for pollutant-induced toxicity effects on soil microbial communities (41, 44, 45). Recent developments of molecular techniques for studies of indigenous soil AOB (9, 30) have further made this group most suitable for in-depth ecological and ecotoxicological studies.

Despite the obvious relevance of ecotoxicological studies at the microbial community level, there is also a need for the development of bioassays that use selected microorganisms for toxicity testing of pollutants (11, 13, 40, 44). Unfortunately, previous assays that used test microorganisms were often based on exposure to soil extracts in water or organic solvent (12, 20, 32, 42, 43), thereby ignoring interactions of the test organisms with adsorptive soil pollutants. Furthermore, microbial toxicity testing in soil has also too often been based on irrelevant microorganisms (e.g., Microtox) or experimental conditions. Considerable attention is currently paid to the development of solid-phase contact assays in toxicity testing, thus allowing for a direct contact between the test microorganisms and the soil matrix (3, 38-40). As the inoculated test bacteria respond to bioavailable pools including both free and adsorbed subfractions of the soil pollutants, these solid-phase contact assays may provide a much improved determination of in situ toxicity for the many adsorptive contaminants present in soil.

No assay based on inoculation of the highly sensitive AOB as test microorganisms has so far been developed for soil toxicity testing. One suitable AOB for this purpose could be the recombinant luxAB-marked Nitrosomonas europaea strain ATCC 19718(pHLUX20) (25), which expresses the Vibrio harveyi luciferase enzyme, inducing a respiration-driven bioluminescence. Bioluminescence output in this strain is thus limited strictly by the amount of reducing power generated from ammonia oxidation (25), and the tight coupling represents a major advantage for a toxicity assay. Hence, when the reporter strain is used as a toxicity biosensor (28), it should respond immediately when respiration activity is affected in the test cells. In the solid-phase contact assay, the bioluminescent reporter strain should be particularly useful since light output can be recorded directly in the soil samples by luminometry with little or no background luminescence originating from indigenous members of the microbial soil community (35). In this report, we present the development of a novel solid-phase contact assay for toxicity testing in natural soil samples that uses the luxAB-marked N. europaea strain ATCC 19718(pHLUX20). The assay was used to determine the in situ toxicity in soil of the surfactant, linear alkylbenzene sulfonate (LAS), which is a common soil pollutant arising primarily from deposition of domestic sludge in agricultural soils.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Test bacterium and culture conditions.
A recombinant strain of N. europaea ATCC 19718(pHLUX20) containing the luxAB genes from V. harveyi downstream of a promoter for the hydroxylamineoxidoreductase gene (hao) from N. europaea was used (25). Expression of the luxAB genes is constitutive in this strain, and emission of light is strongly correlated to respiration (ammonia oxidation) activity (25). Cultures were routinely screened for contaminants and cultivated in 250-ml batches with autotrophic growth medium as described previously (10), except that kanamycin (25 mg liter^-1) was included.

Soils.
Lundgaard soil (pH_water, 6.3) was a loamy sand (U.S. Department of Agriculture nomenclature) with a humus content of 2.4% and cation exchange capacities of 41.9 meq kg^-1 for Ca²⁺ and 3.1 meq kg^-1 for Mg²⁺. Askov soil (pH_water, 6.8) was a sandy loam with a humus content of 2.8% and cation exchange capacities of 74.9 meq kg^-1 for Ca²⁺ and 4.7 meq kg^-1 for Mg²⁺. The fresh soils contained approximately 12% (wt/wt) soil moisture and were kept at 5°C. One day before experiments, the soils were sieved (2.5-mm pore size) and preincubated at 20°C.

Bioluminescence assay.
It was important to optimize the bioluminescence of the test bacterium with regard to NH₄⁺ and pH levels, as these factors are known to be cardinal to AOB activity. Early-stationary-phase cultures of the reporter strain, showing a density of approximately 7 · 10⁷ cells ml^-1 and approximately 7.5 mM NO₂^- in the spent medium, were used for all experiments. Optimization was carried out in both pure culture and air-dried Lundgaard soil.

For optimization of pure culture experiments, aliquots of 1 ml of culture were prepared in 2-ml transparent polypropylene microtubes (AxyGen, Inc., Union City, Calif.). Subsequently, different amounts of sterile NH₄Cl or NaOH were added (see Results) and the tubes were incubated horizontally on a rotary shaker (300 rpm) for 1 h at 22°C. After incubation, the luciferase substrate n-decanal (2.5 µl of a 10% [vol/vol] solution in 96% ethanol) was added to each tube. The samples were mixed for 3 s, and 5 s after the addition of the aldehyde, bioluminescence was measured by luminometry (BioOrbit 1253, StruersKebo Lab, Albertslund, Denmark) for 1 min.

For optimization of soil experiments, aliquots of 0.5 g of air-dried Lundgaard soil and 1.5 ml of culture were mixed in the microtubes described above. Following amendment with different amounts of sterile NH₄Cl or NaOH, the test bacteria were added by using the same incubation protocol as for the pure culture experiments. Subsequently, bioluminescence was measured in the luminometer by two different protocols: (i) the soil slurry protocol, which used direct measurement of the bioluminescence in the sample for 1 min after the addition of 3.8 µl of 10% decanal solution, and (ii) the soil extract protocol, which used measurement of the bioluminescence in supernatants obtained after centrifugation of the samples (500 x g, 1 min). In the latter protocol, 1 ml of supernatant was transferred to a new microtube and the bioluminescence was measured as described above for pure culture samples. For both soil protocols, the pH was finally measured in each microtube with a calibrated electrode (Radiometer, Copenhagen, Denmark). The light emission from microtubes with soil amended with sterile growth medium served as a negative control. The very low background levels (0.01 to 0.02 relative light units ml^-1 min^-1) obtained in the controls were subtracted from the bioluminescence measured in the natural soil samples.

LAS addition to soil.
A commercial LAS mixture (Marlon A375; Hüls AG, Marl, Germany) consisting of LAS homologues (C₁₀ to C₁₃ as sodium salts) was used. Marlon A is an aqueous paste containing approximately 75% LAS. Known amounts of LAS in 2 ml of solution were sprayed into 50-g soil samples kept in large glass petri dishes to obtain a homogenous distribution of LAS in the soils. All LAS treatments (0, 7.5, 37.5, 75, 150, 187.5, 225, 300, 375, or 750 mg of LAS kg [dry weightx of soil^-1) contained the same amount of liquid volume to ensure a comparable soil moisture content (14% [wt/wt] of fresh soil). For each LAS treatment, the soil was divided into two fractions, one of which was air dried at room temperature for 24 h and frozen at -18°C. This fraction was subsequently used for the LAS toxicity assay that used the N. europaea reporter strain (see below). The other fraction was incubated for 24 h in glass petri dishes mounted with lids for maintenance of soil moisture. This fraction was subsequently used for supplementing LAS toxicity assays based on potential nitrification activity (ammonia and nitrite oxidation) and soil respiration (basal respiration and substrate-induced respiration [SIR]) as described below.

Toxicity of LAS to N. europaea reporter strain tested in soil and pure culture.
An early-stationary-phase culture of the reporter strain (250 ml) was first adjusted to pH 7.8. Incubation (300 rpm for 1 h at 22°C, unless stated otherwise) was initiated by adding aliquots of 1.5 ml of culture and 22.5 µl of 0.5 M (NH₄)₂SO₄ (final NH₄⁺ concentration, 15 mM) to preweighed (0.5 g) and thawed soil samples in 2-ml transparent polypropylene microtubes. Bioluminescence and pH were subsequently measured by using both the soil slurry and the soil extract protocols as described above. A LAS toxicity test in pure culture was included for comparison. Briefly, different amounts of LAS were added to a series of sterile glass serum vials (one per LAS treatment) with LAS stock solutions and methanol as the solvent. All LAS treatments contained the same amount of methanol solvent, which was subsequently allowed to evaporate in a sterile hood (LaminAir; Holten, Allerød, Denmark). Incubation (300 rpm for 1 h at 22°C) was initiated by adding 20-ml aliquots of early-stationary-phase culture (pH 7.4) to each serum vial. At the end of the incubation, 1-ml aliquots of culture were transferred to 2-ml microtubes and bioluminescence was measured as described above.

Toxicity of LAS to microbial activity in soil.
Potential ammonia oxidation was measured by using the chlorate inhibition technique (5) as described previously (22). Briefly, triplicate samples of 1 g of soil were mixed with 9 ml of autotrophic growth medium (pH 7.5) containing 3.75 mM (NH₄)₂SO₄ and 10 mM KClO₃. The slurries were incubated in glass serum vials for 5 h at 22°C, and potential ammonia oxidation activity was determined from the nitrite accumulation curves. Potential nitrite oxidation was determined by the acetylene inhibition technique as described by Højberg et al. (23). Briefly, triplicate 1-g soil samples were amended with 9 ml of nitrifier medium in glass serum vials. NaNO₂ was added to a 40 µM concentration immediately before incubation, and 4 kPa of C₂H₂ was added to inhibit ammonia oxidation after capping the vials with Teflon-coated butyl rubber stoppers. Subsequently, the slurries were incubated for 5 h, and potential nitrite oxidation activities were calculated from the nitrite removal curves. Control experiments confirmed that the chosen inhibitor concentrations of KClO₃ and C₂H₂ for the two nitrification assays caused a total inhibition of the target populations in both soils (data not shown). Nitrite was determined spectrophotometrically in microtiter plates as described previously (47), except that samples were diluted (1:1) in 2 M KCl prior to measurement in order to prevent LAS interference with the nitrite analysis (A. Pedersen, unpublished data).

Basal respiration and SIR were measured in triplicate samples of approximately 0.3 g (dry weight) of soil in 3.5-ml glass vials (Venoject VT-030PX; Terumo, Leuven, Belgium). Potential respiration activity was determined by the SIR assay (4). In brief, 4 mg of glucose (priorly added to the soil from a 4:1 glucose-talcum mixture) was carefully incorporated into the soil and preincubated for 30 min. Incubations were initiated by out gassing the vials with CO₂-free air, and headspace CO₂ concentrations were measured after approximately 2 and 4 h of incubation at 22°C. A gas chromatograph (no. 6890; Hewlett-Packard Co., Palo Alto, Calif.) equipped with a Porapack Q column kept at 30°C and a thermal conductivity detector operated at 200°C (flow rate of 25 ml of He min^-1) was used. Actual respiration activity (basal respiration) was measured with the same protocol as for SIR but without the addition of glucose.

Estimation of toxicological test parameters.
EC₅₀s (50% effective concentrations) of LAS toxicity were estimated by nonlinear regression of log-transformed data as described previously (33).

Top Abstract Introduction Materials and Methods Results Discussion References

 
Optimization of toxicity assay.
Bioluminescence of the test bacterium could easily be detected in both solid-phase contact assays (soil slurry and soil extract protocols), although the detected light emission was approximately 10 to 50 times lower in the slurry assay. The centrifugation step associated with the soil extract protocol ensured the removal of soil particles, thus reducing the quenching and masking of emitted light. A standard centrifugation protocol (500 x g for 1 min) was chosen here as the best compromise, retaining as many of the test bacteria and removing as many of the soil particles as possible. Reducing the centrifugation speed to 100 x g thus resulted in increased turbidity of the soil extracts and only marginally increased the detected bioluminescence, whereas higher centrifugation speeds (1,000 to 4,000 x g) significantly decreased the detected bioluminescence (data not shown).

As shown in Fig. 1, the bioluminescence pattern was quite similar in pure cultures and soil extracts, thus including a high initial bioluminescence, which soon decreased to a constant level. In contrast, the pattern observed in soil slurries showed a significant increase in detectable bioluminescence after about 30 s, which was most likely due to reduced quenching and masking over time resulting from the rapid sedimentation of large suspended soil particles. Using the same batch culture of test bacteria, bioluminescence in soil extracts was found to be 48% ± 5% (Lundgaard soil; n = 10) and 38% ± 4% (Askov soil; n = 10) of the corresponding bioluminescence in pure cultures, indicating some loss of test bacteria during the centrifugation step of the soil extract protocol.

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FIG. 1. Representative progress curves for bioluminescence emission by N. europaea ATCC 19718(pHLUX20) in pure culture and soil samples following the addition of n-decanal to activate the luciferase enzyme. (A) Pure culture; (B) Lundgaard soil extract; (C) Lundgaard soil slurry. Data points represent single measurements. RLU, relative light units.

Figure 2 demonstrates that the bioluminescent N. europaea reporter strain responded similarly to changes in NH₄⁺ availability in pure culture and in soil. At low NH₄⁺ concentrations (up to 15 mM), bioluminescence was strongly correlated to NH₄⁺ availability, whereas bioluminescence was almost unaffected at higher NH₄⁺ concentrations. Likewise, as shown in Fig. 3, the test bacterium responded quite similarly to changes in pH in pure culture and in soil. Compared to the rather narrow pH optimum in the 7.3 to 7.5 range in pure culture, however, the soil-inoculated test bacteria demonstrated a slightly broader pH optimum in the 7.0 to 7.6 range. Based on these results, 15 mM NH₄⁺ and pH 7.4 were chosen as the standard conditions for the solid-phase contact toxicity assay. Due to the buffering capacity of the soils, it was found that the pH of the test culture should be adjusted to 7.8 prior to use in order to reach the standard pH of 7.4 for the assay.

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FIG. 2. Influence of ammonium availability on bioluminescence by N. europaea ATCC 19718(pHLUX20) in pure culture and soil. (A) Pure culture; (B) Lundgaard soil extract. Data points represent means ± standard deviations (n = 4). RLU, relative light units.

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FIG. 3. Influence of pH on the bioluminescence of N. europaea ATCC 19718(pHLUX20) in pure culture and soil. (A) Pure culture; (B) Lundgaard soil extract; (C) Lundgaard soil slurry. Data points for pure culture represent means ± standard deviations (n = 4) while data points for soil protocols represent single measurements. RLU, relative light units.

Toxicity of LAS to the N. europaea reporter strain.
Figure 4 shows that LAS amendment strongly inhibited bioluminescence emission by N. europaea strain ATCC 19718(pHLUX20) in both pure culture and soil. The shape of the dose-response curves indicated that changes in toxicity were most dramatic at low LAS doses (0 to 10 mg liter^-1), whereas higher LAS levels resulted in little additional inhibition of the bioluminescence. Remarkably, the dose-response curves were very similar for the reporter bacteria in pure culture and in soil, indicating a high bioavailability of the LAS compound in soil. The highest concentration tested (approximately 200 mg of LAS liter^-1) resulted in approximately 95% inhibition of N. europaea bioluminescence (data not shown). Increasing the incubation time from 1 to 4 and 24 h progressively increased the toxicity effect of LAS in pure culture (data not shown), indicating a rather slow action of the surfactant or slow response of the test bacterium. However, the standard incubation time of 1 h was still quite adequate to determine the toxicity response. Further, when comparing different soil experiments, longer incubation times also led to unacceptably high standard deviations (data not shown), suggesting that factors in addition to LAS toxicity may also affect the long-term activity response of the test cells in nonsterile soil.

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FIG. 4. Influence of LAS on bioluminescence by N. europaea ATCC 19718(pHLUX20) in pure culture and soil. (A) Pure culture; (B) Lundgaard soil extract; (C) Lundgaard soil slurry. Data points represent means ± standard deviations (n = 4). RLU, relative light units.

Toxicity of LAS to indigenous populations of soil microorganisms.
Figure 5 demonstrates the strong differences in LAS sensitivity for the different soil microbial activities tested. As judged from the strong inhibitions of potential ammonia and nitrite oxidation activities, AOB and nitrite-oxidizing bacteria were highly sensitive to LAS, whereas the heterotrophic soil microorganisms responsible for the most soil respiration activity were rather insensitive. Hence, SIR was only weakly inhibited at high LAS concentrations, whereas basal respiration even tended to be stimulated by LAS.

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FIG. 5. Influence of LAS on indigenous microbial activities in Lundgaard soil. (A) Potential ammonia oxidation (PAO); (B) Potential nitrite oxidation (PNO); (C) substrate-induced respiration (SIR); (D) Basal respiration (BR). Data points represent means ± standard deviations (n = 3).

The calculated EC₅₀s in Table 1 indicate that LAS was slightly more toxic in sandy Lundgaard soil than in the Askov soil with a higher clay content. The data also confirmed the order of toxicity effects by LAS among the investigated microbial activities. Sensitivity to LAS decreased in the following order: N. europaea strain ATCC 19718(pHLUX20) bioluminescence > potential ammonia oxidation > potential nitrite oxidation >> SIR > basal respiration.

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TABLE 1. LAS EC50s for microbial activities in two different soils

Top Abstract Introduction Materials and Methods Results Discussion References

 
Standard conditions.
The two protocols for solid-phase contact assays that used the lux-marked N. europaea reporter strain in soil were developed by proper testing of inoculum preparation, incubation time, centrifugation speed, medium pH and NH₄⁺ levels, and comparisons with results obtained in pure culture. In the initial testing, both soil protocols gave meaningful physiological responses when the test bacteria were exposed to different NH₄⁺ and pH levels in the soil samples. The results from the optimization experiments demonstrated a very broad working range for the ammonia concentration (i.e., the range of concentrations not affecting bioluminescence output) in the bioluminescence assay with the reporter strain (Fig. 2). In contrast, even small changes in soil pH caused large fluctuations in bioluminescence output (Fig. 3). Consequently, buffered media must be used and pH should be monitored routinely in the soil extracts or slurries, as only samples with nearly equal pH may be compared. Neutral to slightly alkaline pH was optimal for bioluminescence output, even if we found a slightly broader pH range for optimal activity and a higher bioluminescence at low pH in soil than in pure culture incubations (Fig. 3). The latter was probably due to soil heterogeneity and the presence of microhabitats maintaining a neutral pH even under acid soil conditions (14, 34).

Soil extract protocol.
When based on the soil extract protocol, the solid-phase contact assay relies on the bioluminescence by a subpopulation of test cells retained in the supernatant after sample centrifugation. When comparing different samples, such a measurement could be problematic since the fraction of inoculated cells retained in the supernatant is unknown and could vary due to different cell adsorption properties of the soils (6, 37, 40). It has recently been proposed that bioluminescence data from inoculated reporter bacteria should be normalized, i.e., expressed per unit of viable test cell retained (40). However, this calculation is precluded in the present case of N. europaea inoculation in soil, since viability is even more sensitive to LAS than metabolic activity in N. europaea and other tested AOB strains (10) and bioluminescence output reflects respiration activity in the N. europaea ATCC 19718(pHLUX20) reporter strain (25; this study).

With some precaution, however, bioluminescence output and, thus, toxicity data obtained by the soil extract protocol may be compared in different soils. A prerequisite is that the extraction efficiency is preferably high, but at least similar, in the soils under comparison. Since the reporting cell population is inoculated only shortly (1 h) in the soil under the standard assay conditions, a rather high extraction efficiency may be expected, as is typically found for microbial inocula, including N. europaea (1, 21). Based on the optimization experiments for the soil extract protocol, bioluminescence was only slightly lower in the soil extracts than in pure culture with the same amount of inoculum (Fig. 1). Assuming that cell-specific activities, including bioluminescence (25) of N. europaea, were similar in the soil extract and pure culture (experiments were performed under identical nearly optimal conditions of temperature and NH₄⁺ and pH levels), at least 48% ± 5% and 38% ± 4% of the test cells were extracted in the supernatant after centrifugation of the Lundgaard and Askov soils, respectively. The extraction efficiency may actually have been slightly higher than this due to some inevitable masking of the emitted light in the soil extracts. The two agricultural soils characterized as loamy sand and sandy loam, respectively, could thus be suitably compared by using the soil extract protocol. Others have also reported that only soils of rather similar composition (clay, silt) may be compared when using solid-phase contact toxicity assays based on bioluminescent reporter bacteria (6, 37).

Soil slurry protocol.
The potential problem regarding extraction efficiencies and comparison of bioluminescence data obtained in different soils is precluded in the soil slurry protocol, in which the bioluminescence of all test bacteria, including both the free (pelagic) and particle-adsorbed (sessile) cells, is measured. During the recording of bioluminescence, the sedimentation kinetics of the soil particles inevitably affects the light signal with the soil slurry protocol (Fig. 1), but at least for soil samples of similar composition, this can be compensated for by a fixed time for recording (1 min, standard protocol). Another potential problem could be the dramatic quenching and masking of the light emission by soil particles, resulting in relatively low signal-to-noise ratios. This can also be compensated for, however, when using an adequate time for recording in the luminometer. Despite the severalfold-lower output of light from the soil slurries than from the supernatants in soil extracts, there was always an ample light signal for 1 min of recording in the slurries.

Toxicity of bioavailable LAS in soil.
LAS is known to have strong adsorptive properties in soil (15, 26, 27), and the bioavailability and in situ toxicity of such a pollutant to the soil microorganisms may thus be affected differently by dissolved and adsorbed pools in the soil. It has often been stated that only the free (dissolved) pool of an adsorptive pollutant is actually bioavailable and thus able to impose a toxic effect on the soil organisms (8, 24). On the other hand, many recent reports have suggested that the adsorbed pollutant pool may also be at least partially available for the microorganisms, both in terms of causing toxic effects (2, 3, 16, 38-40) and serving as substrates for biodegradation (26, 27). As amended LAS is primarily adsorbed to soil particles in the present soils (I. B. Kristiansen and H. de Jonge, personal communication; Pedersen, unpublished), our results provide strong evidence that both free and adsorbed LAS pools contribute to the toxicity affecting the inoculated reporter strain. Hence, the implication that N. europaea was indeed affected by both free and adsorbed LAS pools came from the similar dose-response curves (Fig. 4) and EC₅₀s (Table 1) in soil (with either of the protocols) and pure culture, respectively. In this experiment, the recorded LAS toxicity was thus similar whether based on (i) pelagic test cells exposed to LAS in solution (pure culture), (ii) pelagic test cells exposed to free and adsorbed LAS in soil (soil extract protocol), or (iii) pelagic and particle-bound test cells exposed to free and adsorbed LAS in soil (soil slurry protocol).

It is likely that incubation conditions during the toxicity testing in soil (suspension in a 3:1 [wt/wt] culture-to-soil ratio) may increase the free (dissolved) pool of some adsorptive pollutants relative to conditions in the natural soil (46). Our solid phase-contact assay for toxicity testing may therefore overestimate the bioavailable fraction of such pollutants compared to conditions in native agricultural soils. This is unimportant for the toxicity testing of LAS, however, since the reporter cells seemed to react similarly to free and adsorbed pools of the pollutant. In the case of LAS, the data obtained by the solid-phase contact assay based on the above suspension therefore also reflected the toxicity effect of bioavailable LAS in the soils.

Interestingly, the respiration activity (bioluminescence) of the N. europaea ATCC 19718(pHLUX20) reporter strain appeared to be slightly more sensitive to LAS than the potential respiration activity of indigenous AOB in the soils (Table 1). The difference was not apparent within the N. europaea ATCC 19718(pHLUX20) reporter strain per se, since EC₅₀s for bioluminescence (12 mg liter^-1) (this study) and ammonia oxidation activity (16 mg liter^-1) (10) were nearly identical. One explanation could be that the indigenous AOB are relatively tolerant of the pollutant, but actually the N. europaea species used as a reporter strain appears to be more tolerant of LAS (10) than the Nitrosospira spp. which occur more commonly in soil (9, 29, 30). Alternatively, indigenous AOB are mostly sessile (1, 21) and the surface-associated cells could be less exposed to the LAS amendment or represent different cell types with a higher LAS tolerance. We propose that the solid phase-contact assay with the reporter strain N. europaea ATCC 19718(pHLUX20) is more sensitive in measuring the toxicity effects of soil pollutants than the commonly used assay of potential ammonia oxidation activity.

Comparing several microbial activities in the soils, both the respiration activity (bioluminescence) of the N. europaea ATCC 19718(pHLUX20) reporter strain and the potential respiration activities of indigenous ammonia and nitrite oxidizers were much more sensitive to LAS than the potential respiration activity of the heterotrophic microorganisms (Fig. 5; Table 1). This is consistent with other recent data, indicating that nitrifying bacteria, and ammonia oxidizers in particular, are particularly sensitive to LAS and more so than the heterotrophs in soil (10, 17, 18). Interestingly, basal (actual) soil respiration by the heterotrophs appeared to be stimulated by LAS amendment (Fig. 5). Even assuming a very short half-life of 7 days for complete LAS mineralization (15), our data indicate an approximately six-times-larger increase of basal respiration at low LAS concentrations (10 mg kg^-1) than can be accounted for solely by mineralization of the applied amount of LAS. This in turn suggests that the increase of basal soil respiration was largely due to an acute stress effect (i.e., an increase of the metabolic quotient by LAS) on the heterotrophic microbial community.

In conclusion, we propose that the solid-phase contact assay with N. europaea ATCC 19718(pHLUX20) constitutes a most useful method to estimate the toxicity effects on microorganisms in soil. First, the assay reflects the toxicity of bioavailable pools of contaminants in soil. Second, the assay is highly sensitive and targets the functioning of a key group of important microorganisms in soil. Third, the use of a reporter strain with known responses to a range of environmental conditions (as opposed to tests relying on indigenous microbial populations) allows for detailed investigations of factors modifying the toxicity of environmental contaminants in soil. In addition, the assay may quite easily be modified for use in other environmental matrices or for other purposes, e.g., probing the general conditions for nitrification activity.

 
This work was supported by the Center for Sustainable Land Use and Management of Contaminants, Carbon, and Nitrogen (Denmark) and by the Danish Veterinary and Agricultural Research Council (grant no. 9600638).

We thank Tarou Iizumi for providing the recombinant N. europaea strain used and Bente Østergaard for excellent technical assistance.

 
* Corresponding author. Present address: Center for Microbial Ecology, Michigan State University, 540 Plant and Soil Science Building, East Lansing, MI 48824. Phone: (517) 353-7858. Fax: (517) 353-2917. E-mail: kkb{at}kvl.dk.

Top Abstract Introduction Materials and Methods Results Discussion References

 

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Applied and Environmental Microbiology, July 2002, p. 3502-3508, Vol. 68, No. 7
0099-2240/02/$04.00+0     DOI: 10.1128/AEM.68.7.3502-3508.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.

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