APPLIED AND ENVIRONMENTAL MICROBIOLOGY, May 1992, p. 1459-1465
0099-2240/92/051459-07$02.00/0 Copyright
)
Vol. 58, No. 5
1992, American Society for Microbiology
Effect of Inoculant Strain and Organic Matter Content on Kinetics of 2,4-Dichlorophenoxyacetic Acid Degradation in Soil LINDA E. GREERt AND DANIEL R. SHELTON* Pesticide Degradation Laboratory, Agricultural Research Service, U.S. Department of Agriculture, 10300 Baltimore Avenue, Beltsville, Maryland 20705 Received 16 September 1991/Accepted 22 January 1992
We monitored rates of degradation of soluble and sorbed 2,4-dichlorophenoxyacetic acid (2,4-D) in low-organic-matter soil at field capacity amended with 1, 10, or 100 ,ug of 2,4-D per g of wet soil and inoculated with one of two bacterial strains (MI and 155) with similar maximum growth rates (pmax) but significantly different half-saturation growth constants (Ks). Concentrations of soluble 2,4-D were determined by analyzing samples of pore water pressed from soil, and concentrations of sorbed 2,4-D were determined by solvent extraction. Between 65 and 75% of the total 2,4-D was present in the soluble phase at equilibrium, resulting in soil solution concentrations of ca. 8, 60, and 600 pg of 2,4-D per ml, respectively. Soluble 2,4-D was metabolized preferentially; this was followed by degradation of both sorbed (after desorption) and soluble 2,4-D. Rates of degradation were comparable for the two strains at soil concentrations of 10 and 100 ,ug of 2,4-D per g; however, at 1 ,ug/g of soil, 2,4-D was metabolized more rapidly by the strain with the lower Ks value (strain MI). We also monitored rates of biodegradation of soluble and sorbed 2,4-D in high-organicmatter soil at field capacity amended with 100 ,ug of 2,4-D per g of wet soil and inoculated with the low-Ks strain (strain MI). Ten percent of total 2,4-D was present in the soluble phase, resulting in a soil solution concentration of ca. 30 ,ug of 2,4-D per ml. Rates of degradation in the high-organic-matter soil were lower than in the low-organic-matter soil, presumably as a result of lower rates of desorption and microbial growth. The use of microorganisms for the bioremediation or bioreclamation of soils contaminated with toxic pollutants has become the focus of considerable attention because of the potential for restoring or reclaiming soils at a considerable cost savings compared with technologies such as incineration (13). The ability of soil microorganisms to degrade a variety of toxic pollutants, particularly pesticides, is well documented (2, 11). In fact, the phenomenon of accelerated degradation of soil-applied pesticides is essentially unwanted bioremediation (18). At sites where the appropriate indigenous strains are present, remediation may consist only of optimizing the soil conditions for microbial growth. However, there is no reason to believe that the appropriate strains will always be present, particularly where initial site conditions are deleterious to microbial survival and proliferation. Reliance on indigenous strains may therefore be inappropriate. An alternative approach would be inoculation of contaminated soils with microorganisms possessing the appropriate metabolic, physiological, and kinetic characteristics in conjunction with manipulation of the soil parameters to enhance the survival and proliferation of the inoculant strain(s) (10). Despite the apparent simplicity of such an approach, results of soil inoculation studies have been inconsistent. Several investigators have reported the failure of microbial inocula to stimulate rates of pollutant degradation in soils (9, 14, 15), whereas others have observed significantly enhanced rates of biodegradation directly attributable to soil inoculation (3, 6-8, 12). The reason(s) for the success or failure of soil inoculation is unclear. Most successful studies have been conducted in the laboratory under conditions
generally conducive to microbial growth and metabolism; the effectiveness of the same strains under field conditions, which are likely to be less than optimal, is uncertain. On the other hand, in instances in which soil inoculation has been unsuccessful, specific reasons for the failure of the inocula to survive or proliferate have generally not been identified. As with any new technology, successful development of soil inoculation requires that problems or limitations be accurately identified to determine whether viable solutions or alternatives can be found (9). A variety of factors are important in determining the success or failure of bioremediation. Environmental factors, such as soil pH, aeration, nutrients, and moisture content, are well understood and can usually be manipulated to enhance the survival and proliferation of any particular inoculant strain. Microbial factors, such as substrate range, longevity, and resistance to inhibition, may also be manipulated, in part by choosing an inoculant strain(s) with desirable metabolic and physiological characteristics (10). However, even in instances in which the soil parameters have been optimized for growth and one or more strains appropriate to the site have been selected, successful bioremediation is still dependent on achieving an acceptable rate and extent of degradation. Rates of biodegradation are dependent on the kinetic parameters of the inoculant strain(s) in conjunction with the bioavailability of the pollutant(s). The purpose of this study was to correlate rates of biodegradation with substrate bioavailability and microbial growth kinetics. Specifically, the goals of this research were (i) to compare the rates of metabolism of various concentrations of 2,4-dichlorophenoxyacetic acid (2,4-D) in soil inoculated with strains of bacteria that have significantly different Ks values and (ii) to examine the effect of soil organic matter on the partitioning of 2,4-D between soil solution and soil surfaces and on the kinetics of 2,4-D metabolism.
* Corresponding author. t Present address: Natural Resources Defense Council, Washington, D.C. 20005.
1459
1460
APPL. ENvIRON. MICROBIOL.
GREER AND SHELTON
TABLE 1. Physical and chemical characteristics of soils Soil type
Texture
High organic matter
Sandy clay loam
Low organic matter
Sandy loam
a
Organic % matter
14 1.6
Particle size distribution
pH
Cation exchange capacity (meq)
Moisture content (%)a
63% sand, 17% silt, 20% clay 71% sand, 24% silt, 5% clay
4.2
16.6
57
4.9
3.4
10
At field capacity.
MATERIALS AND METHODS Inoculum. The two strains used in this study, MI and 155, have been previously described (10). These strains were chosen because of the statistically significant difference in their half-saturation growth constants (Ks). Soils. Two soils from the Maryland Sassafras series were collected from a single watershed in Salisbury, Md. Soils were crushed and sieved (pore size, 2 mm) to eliminate aggregates (24); they were then air dried and stored in plastic-lined trash barrels until used. Prior to use, the soil pH was adjusted to 6.8 to 7.0 with Ca(OH)2. Chemical and physical characteristics of the two soils were determined at the University of Maryland Soil Testing Laboratory, College Park (Table 1). The moisture content of the soils at field capacity was determined by using a ceramic plate extractor (Soil Moisture Equipment Co., Santa Barbara, Calif.). Soils were saturated with water and held at 40 kPa (-0.3 bar) in the extractor until they reached equilibrium (about 12 h). They were then dried at 100°C overnight to determine the moisture content. Incubations. Nonsterile soil was adjusted to field capacity with a mineral salts medium (22) containing 2,4-D by using a slurry technique (20). Final 2,4-D concentrations were 1, 10, or 100 ,ug/g (wet weight) of soil. Soils were incubated at 25 to 27°C for 48 h prior to inoculation with bacteria in order for partitioning of 2,4-D between the soluble and sorbed phases to achieve equilibrium. Preliminary experiments indicated that equilibrium was achieved in less than 6 h (data not shown). The soils were inoculated with approximately 106 cells per g soil. Bacteria were added to soils in a laminar flow hood with an atomizer; each soil sample was thoroughly mixed several times during the addition of cells. Concentrations of 2,4-D in soil solution were determined by using a Carver press (Fred S. Carver, Inc., Summit, N.J.) (21). A 30-g sample of the low-organic-matter soil or a 5-g sample of the high-organic-matter soil was pressed at 10,000 to 15,000 lb/in2 for 1 min or until water was no longer expressed from the soil. Soil pore water was transferred to microcentrifuge tubes by using a Pasteur pipette and centrifuged in a Microfuge 11 (Beckman Instruments Inc., Fullerton, Calif.) at high speed for 2 min to remove soil particles. The water was decanted and frozen until analysis. The total 2,4-D concentration was determined by soil extraction. Soil (10 or 20 g) and 100 ml of ethanol were combined in Teflon-lined glass jars. After ca. 48 h, the jars were shaken on a wrist-action shaker (Lab Line Instruments, Melrose Park, Ill.) for 1 h, the ethanol supernatant was filtered through glass fiber filters (no. 934-AH; Whatman International Ltd., Maidstone, England), and the ethanol was evaporated to near dryness by using a Rotavapor R110 (Brinkmann Instruments, Westbury, N.Y.). Residues were dissolved in 10 ml of ethanol and stored at 4°C until analysis. Extraction efficiencies for 2,4-D were 80% for the loworganic-matter soil and 54% for the high-organic-matter soil.
The percent 2,4-D in soil solution was calculated as follows: {[micrograms of 2,4-D per milliliter of pore water] x [fraction of soil moisture (milliliters of water per gram of wet soil)]/total 2,4-D (adjusted for extraction efficiency) (micrograms per gram of wet soil)} x 100. All values are the average of two experiments. Mineralization of 2,4-D was assessed by monitoring 14C02 production in biometer flasks (4) containing 30 g of inoculated soil amended with ca. 2 x 104 dpm of [U-nng-14C]2,4D. Flask side arms contained 10 ml of 0.1 M KOH trapping solution, which was periodically removed, subsampled (1 ml), and replaced with 10 ml of fresh KOH. Controls to determine background rates of 2,4-D mineralization consisted of uninoculated soil amended with [U-nng-14C]2,4-D. Initial population densities of the inoculum were determined by spread plating appropriate serial dilutions of the stock culture onto nutrient broth (0.2%) agar. Thereafter, 10 g of bulk soil was added to 90 ml of sterile water and blended for 2 min in an Osterizer blender. The soil slurry was serially diluted with mineral salts and plated onto 0.1% 2,4-Dmineral salts agar, and the plates were incubated at 27°C. Cells were counted after 72 h. The inoculum longevity was determined by a 30-day incubation of inoculated soil amended with 100 ppm of 2,4-D; soils were sampled periodically for viable CFU. Analytical. 2,4-D was analyzed by high-pressure liquid chromatography (HPLC) by using an HPLC system (Waters Associates, Inc., Milford, Mass.) consisting of M6000 A pumps and a Maxima 820 Chromatography Work Station radial compression module, with a Perkin-Elmer LC-95 UV-visible variable-wavelength detector set at 210 nm. Separations were achieved by using a C-18 Nova Pak (4,um) radially compressed cartridge. The mobile phase consisted of 40% dilute phosphoric acid (pH = 2) and 60% methanol, and the flow rate was 2.0 ml/min. Radiolabeled 2,4-D and 14C02 were counted by using an LS 6800 scintillation counter (Beckman Instruments, Irvine, Calif.). KOH trapping solution (1 ml) was counted in 10 ml of Beckman Ready-Solv HP scintillation cocktail. Beckman C-14 quench standards were used to determine counting efficiencies, and quench was corrected through a computer data-processing program in the scintillation counter. Chemicals. 2,4-D (96% purity) was purchased from Aldrich Chemical Co., Milwaukee, Wis.) and recrystallized from toluene to increase its purity. [U-_ing-14C]2,4-D (12 mCi/mmol) was obtained from California Bionuclear Corp., Sun Valley, Calif.). RESULTS We monitored rates of 2,4-D degradation in low-organicmatter soil at field capacity amended with 1, 10, or 100 pug of 2,4-D per g of wet soil and inoculated with either strain MI or 155 at ca. 106 bacteria/g of soil. The initial fraction of 2,4-D sorbed to soil surfaces at 10 and 100 ,ug/g varied from
VOL. 58, 1992
2,4-D DEGRADATION IN SOIL
1461
100
90
80
70 §
z
60
W
z
0)
c.
50
Li
L-
40 30 20 10
0 600
E
E )
-j
5
0
0)L'3
3
1 e+009
500
_5
0
400
u50-I
300
1 e+008 o D
CN w
LI-
0
3.
,i
0
cr
0
0
200
1 e+007
1-100
hWM, Ii'
D 0
10
20
30
40
50
1e+006 60
0
HOURS
FIG. 1. Degradation of 100 ,ug of 2,4-D per g of soil by strain MI in low-organic-matter soil at field capacity (total = soluble + sorbed).
approximately 25 to 35%, with the remainder in soil solution (Fig. 1 to 4). Initial pore water concentrations of 2,4-D were ca. 60 and 600 ,ug/ml at soil concentrations of 10 and 100 ,ug/g, respectively. Rates of degradation were generally comparable for the two strains at 10 and 100 ,ug/g, with pore water 2,4-D metabolized preferentially for the first 20 h, followed by comparable rates of degradation for both soluble and sorbed 2,4-D. Pore water 2,4-D was metabolized below 1 ,ug/ml within 30 h. Log numbers of cells (CFU) increased from ca. 106 to 108 or 109 cells per g of soil at soil concentrations of 10 and 100 ,ug/g, respectively. Partitioning between soluble and sorbed phases at 1 ,g/g is not shown because concentrations of 2,4-D in ethanol extracts (sorbed 2,4-D) were below the detection limit. Pore water 2,4-D was metabolized more rapidly and to a greater extent by strain MI than by strain 155 (Fig. 5). Cell numbers of both strains increased ca. 10-fold; however, CFUs also increased ca. 10-fold in the absence of 2,4-D. We monitored rates of 2,4-D degradation in high-organicmatter soil at field capacity amended with 100 pg of 2,4-D per g of wet soil and inoculated with ca. 106 of MI (Fig. 6). The initial fraction of 2,4-D sorbed to soil surfaces was 90%. Initial pore water concentrations were ca. 30 ,g/ml. Soluble and sorbed 2,4-D appeared to be metabolized simultaneously, although, 70 h after inoculation, pore water 2,4-D had been metabolized below the detection limit while ca. 5% of the sorbed 2,4-D had not been metabolized. Cell numbers are not reported because of difficulties in quantitative recoveries from the high-organic-matter soil.
1 e+006
0
10
20
30
40
50
60
HOURS
FIG. 2. Degradation of 100 pLg of 2,4-D per g of soil by strain 155 in low-organic-matter soil at field capacity (total = soluble +
sorbed). There was negligible mineralization of 2,4-D in either lowor high-organic-matter soil during the time course of incubations in the absence of inoculation. Significant mineralization of 2,4-D (.10%) by indigenous microorganisms was observed in low-organic-matter soil after a lag of 10 days; initial population densities were
0099-2240/92/051459-07$02.00/0 Copyright
)
Vol. 58, No. 5
1992, American Society for Microbiology
Effect of Inoculant Strain and Organic Matter Content on Kinetics of 2,4-Dichlorophenoxyacetic Acid Degradation in Soil LINDA E. GREERt AND DANIEL R. SHELTON* Pesticide Degradation Laboratory, Agricultural Research Service, U.S. Department of Agriculture, 10300 Baltimore Avenue, Beltsville, Maryland 20705 Received 16 September 1991/Accepted 22 January 1992
We monitored rates of degradation of soluble and sorbed 2,4-dichlorophenoxyacetic acid (2,4-D) in low-organic-matter soil at field capacity amended with 1, 10, or 100 ,ug of 2,4-D per g of wet soil and inoculated with one of two bacterial strains (MI and 155) with similar maximum growth rates (pmax) but significantly different half-saturation growth constants (Ks). Concentrations of soluble 2,4-D were determined by analyzing samples of pore water pressed from soil, and concentrations of sorbed 2,4-D were determined by solvent extraction. Between 65 and 75% of the total 2,4-D was present in the soluble phase at equilibrium, resulting in soil solution concentrations of ca. 8, 60, and 600 pg of 2,4-D per ml, respectively. Soluble 2,4-D was metabolized preferentially; this was followed by degradation of both sorbed (after desorption) and soluble 2,4-D. Rates of degradation were comparable for the two strains at soil concentrations of 10 and 100 ,ug of 2,4-D per g; however, at 1 ,ug/g of soil, 2,4-D was metabolized more rapidly by the strain with the lower Ks value (strain MI). We also monitored rates of biodegradation of soluble and sorbed 2,4-D in high-organicmatter soil at field capacity amended with 100 ,ug of 2,4-D per g of wet soil and inoculated with the low-Ks strain (strain MI). Ten percent of total 2,4-D was present in the soluble phase, resulting in a soil solution concentration of ca. 30 ,ug of 2,4-D per ml. Rates of degradation in the high-organic-matter soil were lower than in the low-organic-matter soil, presumably as a result of lower rates of desorption and microbial growth. The use of microorganisms for the bioremediation or bioreclamation of soils contaminated with toxic pollutants has become the focus of considerable attention because of the potential for restoring or reclaiming soils at a considerable cost savings compared with technologies such as incineration (13). The ability of soil microorganisms to degrade a variety of toxic pollutants, particularly pesticides, is well documented (2, 11). In fact, the phenomenon of accelerated degradation of soil-applied pesticides is essentially unwanted bioremediation (18). At sites where the appropriate indigenous strains are present, remediation may consist only of optimizing the soil conditions for microbial growth. However, there is no reason to believe that the appropriate strains will always be present, particularly where initial site conditions are deleterious to microbial survival and proliferation. Reliance on indigenous strains may therefore be inappropriate. An alternative approach would be inoculation of contaminated soils with microorganisms possessing the appropriate metabolic, physiological, and kinetic characteristics in conjunction with manipulation of the soil parameters to enhance the survival and proliferation of the inoculant strain(s) (10). Despite the apparent simplicity of such an approach, results of soil inoculation studies have been inconsistent. Several investigators have reported the failure of microbial inocula to stimulate rates of pollutant degradation in soils (9, 14, 15), whereas others have observed significantly enhanced rates of biodegradation directly attributable to soil inoculation (3, 6-8, 12). The reason(s) for the success or failure of soil inoculation is unclear. Most successful studies have been conducted in the laboratory under conditions
generally conducive to microbial growth and metabolism; the effectiveness of the same strains under field conditions, which are likely to be less than optimal, is uncertain. On the other hand, in instances in which soil inoculation has been unsuccessful, specific reasons for the failure of the inocula to survive or proliferate have generally not been identified. As with any new technology, successful development of soil inoculation requires that problems or limitations be accurately identified to determine whether viable solutions or alternatives can be found (9). A variety of factors are important in determining the success or failure of bioremediation. Environmental factors, such as soil pH, aeration, nutrients, and moisture content, are well understood and can usually be manipulated to enhance the survival and proliferation of any particular inoculant strain. Microbial factors, such as substrate range, longevity, and resistance to inhibition, may also be manipulated, in part by choosing an inoculant strain(s) with desirable metabolic and physiological characteristics (10). However, even in instances in which the soil parameters have been optimized for growth and one or more strains appropriate to the site have been selected, successful bioremediation is still dependent on achieving an acceptable rate and extent of degradation. Rates of biodegradation are dependent on the kinetic parameters of the inoculant strain(s) in conjunction with the bioavailability of the pollutant(s). The purpose of this study was to correlate rates of biodegradation with substrate bioavailability and microbial growth kinetics. Specifically, the goals of this research were (i) to compare the rates of metabolism of various concentrations of 2,4-dichlorophenoxyacetic acid (2,4-D) in soil inoculated with strains of bacteria that have significantly different Ks values and (ii) to examine the effect of soil organic matter on the partitioning of 2,4-D between soil solution and soil surfaces and on the kinetics of 2,4-D metabolism.
* Corresponding author. t Present address: Natural Resources Defense Council, Washington, D.C. 20005.
1459
1460
APPL. ENvIRON. MICROBIOL.
GREER AND SHELTON
TABLE 1. Physical and chemical characteristics of soils Soil type
Texture
High organic matter
Sandy clay loam
Low organic matter
Sandy loam
a
Organic % matter
14 1.6
Particle size distribution
pH
Cation exchange capacity (meq)
Moisture content (%)a
63% sand, 17% silt, 20% clay 71% sand, 24% silt, 5% clay
4.2
16.6
57
4.9
3.4
10
At field capacity.
MATERIALS AND METHODS Inoculum. The two strains used in this study, MI and 155, have been previously described (10). These strains were chosen because of the statistically significant difference in their half-saturation growth constants (Ks). Soils. Two soils from the Maryland Sassafras series were collected from a single watershed in Salisbury, Md. Soils were crushed and sieved (pore size, 2 mm) to eliminate aggregates (24); they were then air dried and stored in plastic-lined trash barrels until used. Prior to use, the soil pH was adjusted to 6.8 to 7.0 with Ca(OH)2. Chemical and physical characteristics of the two soils were determined at the University of Maryland Soil Testing Laboratory, College Park (Table 1). The moisture content of the soils at field capacity was determined by using a ceramic plate extractor (Soil Moisture Equipment Co., Santa Barbara, Calif.). Soils were saturated with water and held at 40 kPa (-0.3 bar) in the extractor until they reached equilibrium (about 12 h). They were then dried at 100°C overnight to determine the moisture content. Incubations. Nonsterile soil was adjusted to field capacity with a mineral salts medium (22) containing 2,4-D by using a slurry technique (20). Final 2,4-D concentrations were 1, 10, or 100 ,ug/g (wet weight) of soil. Soils were incubated at 25 to 27°C for 48 h prior to inoculation with bacteria in order for partitioning of 2,4-D between the soluble and sorbed phases to achieve equilibrium. Preliminary experiments indicated that equilibrium was achieved in less than 6 h (data not shown). The soils were inoculated with approximately 106 cells per g soil. Bacteria were added to soils in a laminar flow hood with an atomizer; each soil sample was thoroughly mixed several times during the addition of cells. Concentrations of 2,4-D in soil solution were determined by using a Carver press (Fred S. Carver, Inc., Summit, N.J.) (21). A 30-g sample of the low-organic-matter soil or a 5-g sample of the high-organic-matter soil was pressed at 10,000 to 15,000 lb/in2 for 1 min or until water was no longer expressed from the soil. Soil pore water was transferred to microcentrifuge tubes by using a Pasteur pipette and centrifuged in a Microfuge 11 (Beckman Instruments Inc., Fullerton, Calif.) at high speed for 2 min to remove soil particles. The water was decanted and frozen until analysis. The total 2,4-D concentration was determined by soil extraction. Soil (10 or 20 g) and 100 ml of ethanol were combined in Teflon-lined glass jars. After ca. 48 h, the jars were shaken on a wrist-action shaker (Lab Line Instruments, Melrose Park, Ill.) for 1 h, the ethanol supernatant was filtered through glass fiber filters (no. 934-AH; Whatman International Ltd., Maidstone, England), and the ethanol was evaporated to near dryness by using a Rotavapor R110 (Brinkmann Instruments, Westbury, N.Y.). Residues were dissolved in 10 ml of ethanol and stored at 4°C until analysis. Extraction efficiencies for 2,4-D were 80% for the loworganic-matter soil and 54% for the high-organic-matter soil.
The percent 2,4-D in soil solution was calculated as follows: {[micrograms of 2,4-D per milliliter of pore water] x [fraction of soil moisture (milliliters of water per gram of wet soil)]/total 2,4-D (adjusted for extraction efficiency) (micrograms per gram of wet soil)} x 100. All values are the average of two experiments. Mineralization of 2,4-D was assessed by monitoring 14C02 production in biometer flasks (4) containing 30 g of inoculated soil amended with ca. 2 x 104 dpm of [U-nng-14C]2,4D. Flask side arms contained 10 ml of 0.1 M KOH trapping solution, which was periodically removed, subsampled (1 ml), and replaced with 10 ml of fresh KOH. Controls to determine background rates of 2,4-D mineralization consisted of uninoculated soil amended with [U-nng-14C]2,4-D. Initial population densities of the inoculum were determined by spread plating appropriate serial dilutions of the stock culture onto nutrient broth (0.2%) agar. Thereafter, 10 g of bulk soil was added to 90 ml of sterile water and blended for 2 min in an Osterizer blender. The soil slurry was serially diluted with mineral salts and plated onto 0.1% 2,4-Dmineral salts agar, and the plates were incubated at 27°C. Cells were counted after 72 h. The inoculum longevity was determined by a 30-day incubation of inoculated soil amended with 100 ppm of 2,4-D; soils were sampled periodically for viable CFU. Analytical. 2,4-D was analyzed by high-pressure liquid chromatography (HPLC) by using an HPLC system (Waters Associates, Inc., Milford, Mass.) consisting of M6000 A pumps and a Maxima 820 Chromatography Work Station radial compression module, with a Perkin-Elmer LC-95 UV-visible variable-wavelength detector set at 210 nm. Separations were achieved by using a C-18 Nova Pak (4,um) radially compressed cartridge. The mobile phase consisted of 40% dilute phosphoric acid (pH = 2) and 60% methanol, and the flow rate was 2.0 ml/min. Radiolabeled 2,4-D and 14C02 were counted by using an LS 6800 scintillation counter (Beckman Instruments, Irvine, Calif.). KOH trapping solution (1 ml) was counted in 10 ml of Beckman Ready-Solv HP scintillation cocktail. Beckman C-14 quench standards were used to determine counting efficiencies, and quench was corrected through a computer data-processing program in the scintillation counter. Chemicals. 2,4-D (96% purity) was purchased from Aldrich Chemical Co., Milwaukee, Wis.) and recrystallized from toluene to increase its purity. [U-_ing-14C]2,4-D (12 mCi/mmol) was obtained from California Bionuclear Corp., Sun Valley, Calif.). RESULTS We monitored rates of 2,4-D degradation in low-organicmatter soil at field capacity amended with 1, 10, or 100 pug of 2,4-D per g of wet soil and inoculated with either strain MI or 155 at ca. 106 bacteria/g of soil. The initial fraction of 2,4-D sorbed to soil surfaces at 10 and 100 ,ug/g varied from
VOL. 58, 1992
2,4-D DEGRADATION IN SOIL
1461
100
90
80
70 §
z
60
W
z
0)
c.
50
Li
L-
40 30 20 10
0 600
E
E )
-j
5
0
0)L'3
3
1 e+009
500
_5
0
400
u50-I
300
1 e+008 o D
CN w
LI-
0
3.
,i
0
cr
0
0
200
1 e+007
1-100
hWM, Ii'
D 0
10
20
30
40
50
1e+006 60
0
HOURS
FIG. 1. Degradation of 100 ,ug of 2,4-D per g of soil by strain MI in low-organic-matter soil at field capacity (total = soluble + sorbed).
approximately 25 to 35%, with the remainder in soil solution (Fig. 1 to 4). Initial pore water concentrations of 2,4-D were ca. 60 and 600 ,ug/ml at soil concentrations of 10 and 100 ,ug/g, respectively. Rates of degradation were generally comparable for the two strains at 10 and 100 ,ug/g, with pore water 2,4-D metabolized preferentially for the first 20 h, followed by comparable rates of degradation for both soluble and sorbed 2,4-D. Pore water 2,4-D was metabolized below 1 ,ug/ml within 30 h. Log numbers of cells (CFU) increased from ca. 106 to 108 or 109 cells per g of soil at soil concentrations of 10 and 100 ,ug/g, respectively. Partitioning between soluble and sorbed phases at 1 ,g/g is not shown because concentrations of 2,4-D in ethanol extracts (sorbed 2,4-D) were below the detection limit. Pore water 2,4-D was metabolized more rapidly and to a greater extent by strain MI than by strain 155 (Fig. 5). Cell numbers of both strains increased ca. 10-fold; however, CFUs also increased ca. 10-fold in the absence of 2,4-D. We monitored rates of 2,4-D degradation in high-organicmatter soil at field capacity amended with 100 pg of 2,4-D per g of wet soil and inoculated with ca. 106 of MI (Fig. 6). The initial fraction of 2,4-D sorbed to soil surfaces was 90%. Initial pore water concentrations were ca. 30 ,g/ml. Soluble and sorbed 2,4-D appeared to be metabolized simultaneously, although, 70 h after inoculation, pore water 2,4-D had been metabolized below the detection limit while ca. 5% of the sorbed 2,4-D had not been metabolized. Cell numbers are not reported because of difficulties in quantitative recoveries from the high-organic-matter soil.
1 e+006
0
10
20
30
40
50
60
HOURS
FIG. 2. Degradation of 100 pLg of 2,4-D per g of soil by strain 155 in low-organic-matter soil at field capacity (total = soluble +
sorbed). There was negligible mineralization of 2,4-D in either lowor high-organic-matter soil during the time course of incubations in the absence of inoculation. Significant mineralization of 2,4-D (.10%) by indigenous microorganisms was observed in low-organic-matter soil after a lag of 10 days; initial population densities were
