Environ. Sci. Technol. 2001, 35, 2734-2740
Biodegradation of Non-desorbable Naphthalene in Soils JEONG-HUN PARK, XIANDA ZHAO, AND THOMAS C. VOICE* Department of Civil and Environmental Engineering, Michigan State University, East Lansing, Michigan 48824
The degradation of naphthalene was studied in soilslurry systems, and a quantitative model was developed to evaluate the bioavailability of sorbed-phase contaminant. Four soils with different organic matter contents were used as sorbents. Two naphthalene-degrading organisms, Pseudomonas putida G7 and NCIB 9816-4, were also selected. Sorption isotherms and single and series dilution desorption studies were conducted to evaluate distribution coefficients, desorption parameters, and the amount of nondesorbable naphthalene. Biodegradation kinetics were measured in soil extract solutions and rate parameters estimated. Bioavailability assays involved establishing sorption equilibrium, inoculating the systems with organisms, and measuring naphthalene concentrations in both sorbed and dissolved phases over time. For all four soils, the sorption isotherms were linear, and desorption could be described by a model involving three types of sites: equilibrium, nonequilibrium, and non-desorption. Enhanced bioavailability, as evidenced by faster than expected degradation rates based on liquid-phase concentrations, were observed in soils with the higher sorption distribution coefficients. These observations could be described using model formulations that included solid-phase degradation. In all soils studied, degradation of non-desorbable naphthalene was observed.
Introduction Previous researchers have either shown (1) or made the assumption (2-10) that sorbed contaminants are not directly available for biodegradation. The most commonly accepted conceptual model is that sorbed contaminant first desorbs and then degrades. Recently, other studies have suggested that sorbed substrate may be directly available for degradation by attached cells (11-17). It should be noted that all of the evidence for direct availability is derived from mineralization studies. This approach, although useful for demonstrating the effects of sorbing materials on complete degradation, is somewhat limited in its potential to elucidate the underlying mechanisms. Among these limitations are that CO2 measurements do not indicate whether the mineralized material originates in the sorbed or dissolved phases, may not directly reflect substrate disappearance, and provide only an indicator of the combined effects of desorption and degradation, rather than of the individual processes. Central to this issue is that to independently evaluate a system controlled by multiple rate processes, multiple timeconcentration data sets are required. Finally, the availability of irreversibly sorbed contaminants to biodegradation has not been explicitly studied (9, 10, 18-22). * Corresponding author phone: (517) 355-8240; fax: (517) 3550250; e-mail:
[email protected]. 2734
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TABLE 1. Soil Characteristics soila SPCF Rubicon-BS1 Rubicon-A Kalkaska-A
organic matter sand content (%) (%) 1.9 1.4 2.0 3.9
78 87 89 91
silt (%) 17 8.2 8.4 7.7
clay distribution (%) coefficientb (Kd) 5 4.7 2.7 1.7
4.3 (0.23) 8.8 (0.18) 11.0 (0.23) 25.6 (2.0)
a SPCF was collected in 15-31 cm depth of forest in Michigan State University, East Lansing, MI. Rubicon-BS1 (subsurface in 15-31 cm depth), Rubicon-A (surface in 0-6 cm depth), and Kalkaska-A (surface in 0-6 cm depth) were collected at uncultivated forest in the northern half of Michigan’s lower peninsula (23). b Numbers in parentheses are the standard deviations.
In the present study, substrate depletion, instead of mineralization, was used to investigate the bioavailability of sorbed naphthalene. Sorption and degradation could be tracked independently by measuring naphthalene concentrations in both the sorbed and dissolved phases. Kinetic models were developed to quantitatively analyze the data. Naphthalene was selected as the test contaminant, because it has often been used as a model compound for polycyclic aromatic hydrocarbons (PAHs), and this class of contaminant poses a critical bioavailability issue at numerous petroleum contamination sites.
Materials and Methods Experimental Procedures. Four sandy-loam soils with organic matter contents from 1 to 4% were collected from forest environments in Michigan (23). Soil characteristics are summarized in Table 1. Soil samples were air-dried and sieved through a U.S. Sieve Series No. 20 sieve to remove larger components (>650 µm). Prior to use, the soils were sterilized by γ-irradiation (60Co source) at a dosage of 2 Mrad. Following sterilization, sealed containers were maintained at room temperature. Before the soils were used, 0.1 g of each soil was placed on a nutrient agar plate and incubated at 30 °C for 3 days to verify sterility. No colony-forming units (CFU) were observed. Two naphthalene-degrading organisms, Pseudomonas putida G7 and Pseudomonas sp. strain NCIB 9816-4, were selected because these organisms have been reported to be motile and chemotactic to naphthalene (24, 25). G7 and NCIB 9816-4 were grown in 100 mL of minimal mineral salt media (26). Solid naphthalene was added to the sterilized liquid at a final concentration of 200 mg/L. The liquid medium was inoculated by adding 1 mL of starved liquid culture (cell density of ∼107 CFU/mL) to 100 mL of medium and stirring. Growth was monitored by absorbance at 600 nm, and cells were harvested at a point determined to correspond to early stationary phase based on a full growth curve. Cells were separated by centrifugation (1900g, 20 min) and resuspended in a chemotaxis buffer (CB) (26) before use. The CB contained 13.6 g/L potassium phosphate and 7.4 mg/L EDTA and was adjusted by 10 N NaOH to pH 7. This procedure was repeated three times to ensure the removal of the remaining naphthalene from the cell growth medium. All of the cell suspensions were kept at room temperature and used within 3 h. Naphthalene soil/water distribution coefficients were measured experimentally. An aliquot of each sterile soil (0.68 g of SPCF, 0.57 g of Rubicon-BS1, 0.39 g of Rubicon-A, and 0.18 g of Kalkaska-A) and 4.2 mL of CB containing [14C]naphthalene stock (in methanol) were prepared in 5 mL screw-cap vials with Teflon-lined septa. The soil/water ratios 10.1021/es0019326 CCC: $20.00
2001 American Chemical Society Published on Web 05/30/2001
were carefully selected to achieve approximately equal masses of naphthalene in both liquid and solid phases at the end of the sorption period. The headspace was 98% of the amount calculated by difference for all soils. All experiments were performed at room temperature (24 ( 1 °C) unless otherwise specified. The desorption assay utilized batch soil slurries that had been equilibrated with an initial naphthalene concentration of 800 µg L-1 using the same procedure as in the sorption isotherms. Following 2 days of equilibration, each vial was centrifuged for 5 min at 1200g to separate soil, and the supernatant was sampled. The final concentration of naphthalene in the liquid phase was determined by LSC, and the amount of sorbed naphthalene was calculated by difference. The supernatant was then decanted to the extent possible and the residual water determined by weight. The desorption process was initiated by adding naphthalene-free CB to make up the original volume. The vials were then tumbled again at 6 rpm, and each vial was removed and the liquid phase periodically sampled for analysis. The amount remaining on the soil was calculated by difference, with confirmation of mass balance by soil extraction for the final samples. Series dilution desorption assays were conducted to measure the amount of non-desorbable naphthalene. Triplicate soil slurries were prepared for each soil using the desorption procedure described above. The slurries were tumbled for 24 h, the supernatant was decanted, sampled, and analyzed for naphthalene, and the bottles were refilled with naphthalene-free CB to the original volume. This procedure was repeated for a total of six successive desorption periods. The non-desorbable naphthalene was calculated by difference, with confirmation of mass balance by methanol extraction of the soil following this series dilution procedure. Bioavailability assays were performed using soil slurries in 5 mL screw-cap vials with Teflon-lined septa. Using the same soil to water ratio as in the desorption assays, naphthalene was added at an initial liquid-phase concentration of ∼800 µg/L, and the vials were tumbled for 2 days in the dark. Sterility of the soil slurries was checked by plating out the suspension on nutrient agar plates. The vials were then inoculated with 0.05 mL of cell suspension harvested during the early stationary phase to initiate biodegradation of naphthalene. Cell density in the serum vials was ∼107 CFU/mL. Inoculated vials were tumbled at 6 rpm. At predetermined time intervals, vials were removed and centrifuged at 1200g for 5 min to separate soil. One milliliter of supernatant was transferred to a vial containing 0.01 mL of 10 N NaOH to stop naphthalene degradation and analyzed by HPLC. The remaining supernatant was decanted, and the soil was extracted with methanol as described above to determine sorbed-phase concentrations by HPLC. Biodegradation rates were measured in soil extract solutions to be representative of those expected in systems
with soil present. Soil extract solutions were the supernatants of systems prepared by tumbling the soil slurries for 2 days at the same soil to water ratio as in the desorption and bioavailability experiments. Kinetic data were obtained using the same procedure as in the bioavailability assay, except that no soil was present in the vials. Both sterile CB and soil extract controls were also used to monitor for abiotic losses of naphthalene during the sorption and degradation periods. Naphthalene was analyzed by HPLC using a C18 reversedphase column, an 80% acetonitrile/20% water mobile phase, and fluorescence detection (280 nm excitation, 340 nm emission). The analytical detection limit was 0.2 µg/L. The detection limit for LSC was 15% in the present study, this could occur through direct partitioning to the cell membrane or via degradation by extracellular enzymes. The present study was not designed to discriminate between these explanations, and further research in this area may be warranted. Nonetheless, it can be concluded that material that was nondesorbable in these experimental systems was bioavailable.
Acknowledgments We thank Dr. Harwood at the University of Iowa for providing the bacteria. This work was supported by the U.S. Department of Agriculture National Research Initiative Competitive Grants Program, the Center for Microbial Ecology, and the Great Lakes and Mid-Atlantic Center for Hazardous Substance Research (Environmental Protection Agency, Michigan Department of Environmental Quality).
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Received for review December 4, 2000. Revised manuscript received March 28, 2001. Accepted April 23, 2001. ES0019326
