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U.S. Army ERDC-CRREL, 72 Lyme Rd., Hanover, NH 03755
* Corresponding author (David.B.Ringelberg{at}erdc.usace.army.mil)
Received for publication July 8, 2002.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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Abbreviations: ACN, acetonitrile • PLFAs, ester-linked polar lipid fatty acids • RDX, hexahydro-1,3,5-trinitro-1,3,5-triazine
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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Jenkins et al. (2001) have taken samples soon after munition detonation to determine the extent and nature of explosives distribution. They have shown that training activities can result in the presence of RDX (hexahydro-1,3,5-trinitro-1,3,5-triazine) in site soils at concentrations between 500 and 50000 µg kg-1. Although RDX solubility in water is only about 40 mg L-1, the organic C–normalized soil sorption coefficients (Koc) range from 63 to 270, which places RDX in a classification as a medium to high mobility compound (Spanggord et al., 1983; Layton et al., 1987; Price et al., 2001; Swarm et al., 1983). Jenkins et al. (2001) did detect RDX, at levels <1µg L-1, in monitoring wells located adjacent to one of their sites. Because RDX is toxic and its breakdown products, such as the nitrosamines, are suspected carcinogens, mutagens, or both, the potential for ground water aquifer contamination poses a human health risk to homeowners on private wells and to municipalities using ground water supply wells.
RDX attenuation is typically favored under reduced conditions where the formation of nitroso intermediates precedes ring cleavage and the formation of hydrazines (McCormick et al., 1981). However, on military training ranges, low-order, incomplete detonations deposit RDX into surface soils, where well-drained and unsaturated (oxidative) conditions often predominate. Hawari et al. (2000) have suggested that aerobic enzymatic attacks on the H–NO2 or C–H bonds in RDX may lead to the formation of unstable intermediates that then undergo spontaneous decomposition. As a result, aerobic biodegradation of RDX could serve as a basis for a viable bioremediation technology. However, in surface soils, soil water potential, especially matric potential, can have a significant effect on plant and microbial growth and the subsequent production of exogenous enzymes. The effect of different soil water potentials on soil microbial activity and subsequent biotransformation or biodegradation of RDX has not been well characterized.
Augmenting soils with contaminants to determine the fate and biodegradability of a pollutant is a common practice. Unfortunately, the influence of an organic solvent on the extant microbiota and how this in turn can affect contaminant fate is rarely taken into consideration. Brinch et al. (2002) recently showed that the use of dichloromethane, often used to spike soils with PAHs, had a marked effect on indigenous soil microorganisms. Dichloromethane applied to the entire soil sample had an initial inhibitory effect on all microorganisms, which was then followed by an increase in Pseudomonas sp. to a cell density approximately 1000x greater than that in the control soil. Such an artificial enrichment could have an overall effect on contaminant biodegradation, albeit an unrealistic effect in terms of natural soil attenuation.
In this study, we examined RDX attenuation in a military training range soil incubated with equal concentrations of RDX, but at three different acetonitrile concentrations and two different soil water potentials. Our objective was to compare the effects of soil water potential and acetonitrile concentrations on RDX attenuation. In a companion study, we examined the response of the extant microbiota to the differences in soil treatment and how community structure and function were related to RDX biodegradation.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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Soil moisture potential was determined by saturating soils, placing soils on porous ceramic plates, and applying pressure to the desired matric potential. Following equilibration, a water retention curve, relating soil water potential to gravimetric measurements, was developed (Klute, 1986).
Unsaturated or saturated conditions were established in each microcosm by the addition of 2 or 25 mL, respectively, of H2O–RDX or H2O–RDX–ACN. RDX was added at a concentration of 10 mg L-1. Acetonitrile was added at concentrations of 0, 0.1, and 1% (v/v). Saturated microcosms also received 100 µL of a 0.1% resazurin solution as a redox indicator. Control microcosms were autoclaved at 120°C for 20 min three consecutive times to attenuate the native microorganisms with an 8-h resting or growth period between autoclaving.
After RDX was added, microcosms were incubated for a total of 840 h in the dark without agitation at room temperature (21 ± 2°C). Three replicate vials were sacrificed at 0, 48, 168, 504, and 840 h to obtain contaminant concentrations. Contaminant concentrations were determined for both the aqueous and solid phases of the saturated microcosms. Following centrifugation at 2000 rpm for 20 min, the aqueous phase was decanted and 1 mL was diluted in 1 mL of ACN and 3 mL of Milli-Q H2O. The solution was then filtered through a Millipore Millex-FH hydrophobic PTFE 0.45-µm syringe filter (Bedford, MA) and analyzed by HPLC. The solid phase of the saturated and unsaturated microcosms were extracted with 10 mL of ACN for 18 h at 20°C with sonication; 2 mL of the supernatant was then filtered and diluted with 8 mL of Milli-Q H2O before HPLC analysis.
Concentrations of RDX, MNX (hexahydro-1-nitroso-3,5-dinitro-1,3,5-triazine), DNX (hexahydro-1,3-dinitroso-5-nitro-1,3,5-triazine), and TNX (hexahydro-1,3,5-trinitroso-1,3,5-triazine) were determined by HPLC. For the primary separation, a 15-cm by 3.9-mm (4-µm) Nova Pak C8 (Waters Millipore) column was eluted with 1.4 mL min-1 15:85 isopropanol–water. For the confirmation separation, a 25-cm by 4.6-mm (5-µm) cyano (Supelco LC-CN) column was eluted with 1.2 mL min-1 12:23:62 methanol–acetonitrile–water. The injection volume for each separation was 100 µL. Following these HPLC separations, we recorded absorbance at 254 nm on a Spectra Physics Spectra 100 variable wavelength UV detector.
Rates of RDX loss were calculated as described in Miyares and Jenkins (2000). The observed concentrations for RDX were plotted as ln(C/C0) vs. time (t), where C is the RDX concentration at time t and C0 is the RDX concentration at time 0. Lines were then fitted to the data to determine that a first-order rate process was apparent. Using the following rate equation, ln(C/C0) = kt (where k is the rate constant equal to the slope of the line), we calculated the half-life for RDX under the varying scenarios by dividing the natural log of 0.5 (-0.693) by the rate constant k.
Total microbial biomass was estimated from the concentration of ester-linked polar lipid fatty acids (PLFAs) recovered from the soils. The PLFAs were quantitatively recovered as described in White and Ringelberg (1998) and PLFA concentrations were converted to cell numbers as described in Balkwill et al. (1988).
Statistical evaluations were made using the Statistica software package, version 6.0 (StatSoft, Inc. Tulsa, OK). Significant differences between treatments were determined by an analysis of variance using the Scheffe test in a post hoc analysis at an 
level of 0.05.
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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In the saturated treatments, there was no significant difference in RDX loss as a result of ACN concentration. Although the first-order rate coefficient for the 1% exposure was slightly lower than that of the 0 and 0.1% treatments, the significance of the decrease was minor (Table 2).
RDX loss (as well as mean RDX concentrations) between the active and control microcosms were significantly different (Fig. 2). First-order rate coefficients for the control microcosms averaged 0.001 ± 0.0002 across the two acetonitrile treatments, six times less than that observed in the three active microcosms (Table 2). The differences in rates and extents of RDX loss between the active and control microcosms indicated that RDX degradation was, at least partially, attributable to biotic processes. Conversely, the observed loss of RDX in the control microcosms could not be entirely attributed to abiotic processes, as viable bacteria were observed in the control microcosms at study termination (data not shown). It is acknowledged that autoclaving does not completely sterilize a soil, but does attenuate the bacterial signal. In this study, the attenuation was significant. It is also acknowledged that abiotic degradation of RDX by thermolysis, photolysis, or reduction by zero valent metals can occur. In each of these abiotic processes as well as in certain biotic degradation pathways, an initial attack of the cyclic nitramine results in ring cleavage and the formation of the intermediate methylenenitramine that is then further degraded to N2O and HCHO. Based on a comparison to the autoclaved controls, biotic process were typically two to three times greater than abiotic processes at the 168, 504, and 840 h sampling periods. In addition, samples were incubated in the dark, at room temperature with no supplemental metals.
Singh et al. (1998) have shown that Fe0 can increase the rate and extent of RDX degradation. However, their experiments involved amendments with Fe0 so that final Fe0 concentrations were 10 to 50 g kg-1 soil. Extractable Fe present in the Ft. Greely soils used in this study were 0.25 mg kg-1 and Fe was not added. In addition, Singh et al. found that RDX activity remained unchanged at aqueous Fe0 concentrations of <2 g L-1.
The high organic matter content of the soil (approximately 10%) suggests that available C other than RDX was abundant for microbial metabolism. Given the availability of organic C, it seems unlikely that RDX would be a desirable substrate. However, in other studies, RDX serves as a source of N and not C. In fact, Rhodococcus rhodochrous strain 11Y is capable of using RDX as a sole source of N but not a s a sole source of C (Seth-Smith et al., 2002). In addition, high concentrations of soil organic matter promote the formation of reduced conditions due to high levels of microbial activity and RDX biodegradation is favored under a reduced environment.
Although abiotic processes of RDX degradation were undoubtedly present in the saturated and unsaturated microcosms of this study, the extent of abiotic degradation appeared to be far less than that attributable to biotic processes. Precise descriptions of the nature of any abiotic processes present were outside the scope of this research effort.
Loss of RDX in the saturated microcosms coincided with a gradual increase in the anaerobicity of the system, which was indicated by a color loss in the added resazurin dye. The increase in anaerobicity also coincided with the occurrence of nitroso intermediates in both the active and control microcosms (Fig. 3) . The MNX and DNX concentrations reached a maximum 168 h into the time course and were transient in nature in the active microcosms, falling to below detection limits by study termination. The concentration of TNX reached a maximum at 840 h into the time course (Fig. 3). No other known intermediates (of RDX biodegradation) were targeted for detection. The results described here for RDX loss, nitroso intermediate formation, and subsequent nitroso intermediate disappearance in conditions that are microaerophilic to anaerobic are consistent with the findings of Light et al. (1997), Shen et al. (1997), Young et al. (1997), Boopathy et al. (1998), and Guiot et al. (1999). No nitroso intermediates were detected in the soil extract following centrifugation (i.e., sorbed to the soil), which is consistent with the finding of Sheremata et al. (2001).
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No nitroso-intermediates were detected in the unsaturated microcosms. Although Sheremata and Hawari (2000) reported the presence of MNX as an RDX intermediate during the aerobic biodegradation of RDX by the white rot fungi, Phanerochaete chrysosporium, this organism was not present in this study.
The ACN had a significant effect on RDX biodegradation in the unsaturated soil only. The 1% ACN addition (10000 ppm) inhibited RDX degradation by approximately 25% (Fig. 2). Because microorganisms were present, approximately 4 x 1011 cells kg-1 at 48 h, the inhibition of RDX catabolism suggests either that: (i) the acetonitrile acted as a selective toxicant, or (ii) that the acetonitrile served as an alternative C source. Acetonitrile has a low toxicity to bacteria, cyanobacteria, green algae, and protozoans, with thresholds above 500 µL L-1 (Hashimoto et al., 1993; USEPA, 1985). In this study acetonitrile final concentrations ranged from 1000 (0.1%) to 10000 µL L-1 (1.0%), which are well above the presumed threshold. However, there was no significant loss in viable cell numbers over the time course of the experiment as a result of the 1% ACN addition (Table 1). If it is assumed that toxicity would result in a decrease in viable cell numbers, then these data suggest that acetonitrile was not toxic at the levels administered.
Acetonitrile has been shown to be readily degraded by the aerobic heterotroph, Pseudomonas putida (Nawaz et al., 1989). In addition, available N, such as NH3, has been shown to interfere with RDX biodegradation under aerobic conditions (Binks et al., 1995; Coleman et al., 1998; Yang et al., 1983). Acetonitrile may function in a similar manner to NH3 by serving as an alternative source of N. Although not presented here, results show that a shift in microbial community structure occurred as a result of the 1% acetonitrile addition. The shift in community structure is hypothesized to be an enrichment of organisms capable of utilizing N or C from acetonitrile for metabolism.
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CONCLUSIONS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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The results of this study demonstrated that RDX and its nitroso-derivatives can be biodegraded by an indigenous microbiota in a cold region soil with relatively high organic content. No distinction is made between biotransformation or biomineralization. For this soil, RDX loss rates were significantly greater when the soil was saturated and was rapidly becoming reduced. Although it is acknowledged that autoclaving does not completely sterilize soils, it does attenuate the in situ microbiota. Therefore, the significantly greater loss of RDX in the active microcosms, as compared with the autoclaved controls, suggested that biodegradation was a significant component to the observed RDX loss. The transient nature of nitroso-intermediates was viewed as a further indication of probable microbial activity (Walker and Kaplan 1992).
The fact that organic solvents can influence microbial degradation activity is well established, but nevertheless, these solvents are often used to introduce organic substrates into soils. The manner in which these solvents interfere with the degradation process is not well understood. In this study, the C or N in acetonitrile appears to have interfered with the aerobic biodegradation of RDX when present at 1% of the total aqueous volume, but had no detectable effect on the anaerobic catabolic process. The differential response of the microbiota to the presence of the solvent highlights the dynamic and intricate nature of an indigenous microbial community. These results emphasize that caution may be needed when extrapolating laboratory-based results in the making of far reaching land management decisions.
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ACKNOWLEDGMENTS |
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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