Journal of Environmental Quality 31:241-247 (2002)
© 2002 American Society of Agronomy, Crop Science Society of America, and Soil Science Society of America
TECHNICAL REPORT
Organic Compounds in the Environment
Degradation of 14C-Atrazine Bound Residues in Brown Soil and Rendzina Fractions
C. Munier-Lamy*,a,
M.P. Feuvriera and
T. Chonéb
a Centre de Pédologie Biologique, CNRS, 17 rue Notre Dame des Pauvres, BP 5, 54501, Vand
uvre-lès-Nancy Cedex, France
b UMR-CNRS 5561 Biogéosciences, 6 bd Gabriel, 21000 Dijon, France
* Corresponding author (munier{at}cpb.cnrs-nancy.fr)
Received for publication December 29, 2000.
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ABSTRACT
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The remobilization and the fate of 14C-ring labeled atrazine (6-chloro-N2-ethyl-N4-isopropyl-1,3,5-triazine-2,4-diamine) bound residues was examined in relation with the turnover of natural soil organic matter. Soil fractions of a brown soil and a rendzina were incubated under controled laboratory conditions. The mineralization of natural organic matter and atrazine-bound residues was respectively estimated by the amounts of CO2 and 14CO2 evolved during the incubation. The remobilization and distribution of 14C residues among the soil organic fractions were achieved after physical–chemical extractions of the samples. Comparisons of samples in abiotic and biotic conditions allowed us to assess the influence of microbial activity on the fate of atrazine-bound residues. The mineralization curves showed that natural organic matter and atrazine-bound residues had similar decomposition patterns. After 100 d of incubation, 0.8 to 3.6% of total organic C was evolved as CO2, while only 0.1% of the initial radioactivity was mineralized as CO2, and 7 to 15% was becoming extractable with water and methanol. Few differences were observed in the distribution of residues within organic compounds for both fractions of the rendzina, except a decrease of the 14C radioactivity of the 50- to 5000-µm fraction and a slight increase of that of humin. For the 0- to 5000-µm brown soil fraction, increased radioactivity in humin at the expense of humic (HA) and fulvic (FA) acids was detected after incubation, while for the 0- to 50-µm fraction more radioactivity was recovered with FA.
Abbreviations: FA, fulvic acid • HA, humic acid
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INTRODUCTION
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ATRAZINE, the most common preemergent herbicide, is used for controlling weed growth in crops such as corn and sorghum. The fate of atrazine has been expressed in a number of publications. The degradation, movement, and persistence of atrazine and its degradation products depend on the soil characteristics (Barriuso and Calvet, 1992; Dousset et al., 1995; Koskinen and Clay, 1997) and agricultural practices (Graham et al., 1992; Barriuso and Houot, 1996). Although atrazine has been classified as a moderately persistent herbicide with a half-life of 20 to more than 100 d (Erickson and Lee, 1989), residues of both the parent compound and its degradation products were detected in soils years after application (Capriel and Haisch, 1983; Capriel et al., 1985; Schiavon, 1988; Demon et al., 1994).
Atrazine degradation in soils occurrs both via chemical and biological processes, resulting in the formation of metabolites that are more (i.e., deethylatrazine) or less (i.e., hydroxyatrazine) mobile than atrazine (Adams and Thurman, 1991; Sorenson et al., 1994).
The existence of bound or nonextractable atrazine residues, which are not detected under standard extraction and analysis procedures, was revealed by field and laboratory experiments conducted with 14C-labeled atrazine. Following applications of 14C-atrazine to soil columns, 15 to 40% of the initial radioactivity was still present after 6 mo and the residues were mainly located in the 0- to 20-µm soil fractions, associated with humic compounds (Barriuso et al., 1991) as the result of their incorporation into the natural carbon pool. The atrazine residue amounts are closely related to the organic matter content of the soil (Calderbank, 1989; Bertin and Schiavon, 1989), but the nature of the residues and the bond involved is still unknown, partially due to the extraction and fractionation procedures. The release and the bioavailability of these nonextractable residues, via chemical and biological processes, are important aspects for the assessment of their potential to injure following crops (Frank et al., 1983) or to contaminate ground water. Many works deal with the degradation of atrazine by various pure or mixed cultures of microorganisms (Kaufmann and Blake, 1970; Behki and Khan, 1986; Behki et al., 1993; Mandelbaum et al., 1993; Radosevich et al., 1995), but to date only a few studies have dealt with the release and the biodegradability of atrazine-bound residues (Khan and Ivarson, 1982; Khan and Behki, 1990; Barriuso et al., 1994; Hayar et al., 1997). As bound residues are mainly located in the soil fractions smaller than 50 µm, which also contain 70 to 90% of total organic carbon as humified organic matter (Barriuso et al., 1991), their fate could be related to the humification processes. Such transformation processes are also mainly due to the activity of soil microflora. Thus, the first objective of our study deals with the influence of microbial activity on the release and mineralization of 14C-atrazine bound residues. Second, we attempt to compare their distribution with that of natural organic compounds in <50-µm soil fractions.
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MATERIALS AND METHODS
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Soil Fractions
The soil samples originated from an outdoor microlysimeter experiment (Demon et al., 1994). Briefly, the 0- to 20-cm upper layers of two soils representative of eastern France (a brown soil [Typic Eutrochrept] and a forest rendzina [Typic Rendoll]) were sieved at 5 mm and put in PVC columns. Then, 14C-ring labeled atrazine was applied to the top of the column at an agricultural dose of 2 kg ha-1. The columns were exposed during 365 d to outdoor conditions and then received a second application of atrazine. The soils were removed from the columns 6 mo after the second application and a part of each soil was sieved at 50 µm. The 0- to 5000- and 0- to 50-µm fractions were exhaustively extracted with distilled water, then with a mixture of methanol and water (80:20 [v/v] soil to solvent) until no radioactivity was recovered in the extracts. The fractions then were rinsed with distilled water. A part of each sample was then treated to examine the distribution of 14C among humic compounds with the analytical procedure described below. Before experiments, the other part was air-dried, then freeze-dried and analyzed for its 14C radioactivity, organic carbon, and nitrogen content (Table 1).
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Table 1. pH, organic carbon, nitrogen, and 14C-atrazine residue contents of the 0- to 5-mm and 0- to 50-µm soil fractions used for incubation.
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Incubation Device
The experiments were conducted in microcosm reactors and the experimental procedure was described previously (Metzger et al., 1996; Hayar et al., 1997). Twenty grams of the 0- to 5000-µm soil sample or eight grams of the 0- to 50-µm soil sample were placed respectively into 250- and 125-mL serum bottles and supplemented respectively with 5 and 2 mL of a mineral solution (KH2PO4, 0.5 g; NH4Cl, 0.4 g; MgSO4·7H2O, 0.1 g; in 1000 mL of distilled water). Because prior treatments destroyed the natural microbial community of the samples, the microbial community was reestablished, for biotic treatments, by adding 0.5 or 1 mL of a 10-1 suspension of fresh soil in 0.85% NaCl, respectively, to the 0- to 50- and the 0- to 5000-µm samples. To distinguish physico–chemical from biological processes, abiotic samples were also prepared and obtained after addition of a 0.1% solution of sodium ethyl mercurothiolate 2-benzoate. The solutions were evenly distributed over the soil samples. The total amount of liquid added brought each sample moisture to 80% field capacity, which was maintained during the incubation by introducing a small vial containing distilled water into the serum bottle. Biotic treatments were triplicated and abiotic ones were duplicated. The reactors were then connected to a closed incubation device immersed in a 24°C thermostated water bath and the incubation was conducted for 120 d.
Analytical Methods
Analysis of Evolved 14CO2 and Total CO2
Total CO2 was measured from 1 mL of the gas collected with a syringe from the reactor atmosphere, using a LCA-2 infrared gas analyzer (ADC, Hoddesdon, Herts, England). Filter-sterilized CO2 free air was periodically flowed through the reactors to sweep evolved 14CO2, which was trapped into 10 mL of 0.5 M NaOH. One milliliter of solution from the trap was then pipeted into 20-mL polypropylene scintillation vials containing 10 mL of a scintillation cocktail (Ultima Gold; Packard BioScience B.V., Groningen, the Netherlands) and analyzed by liquid scintillation counting with a Packard (Meriden, CT) Model 4430 scintillation counter. Corrections were made for background and for quenching using external standards.
Soil Sample Analysis
Analyses were performed on the non-incubated and incubated samples. After incubation, three and two serum bottles were respectively collected for biotic and abiotic treatments. The replicates were then mixed to make up a homogeneous sample for analysis.
The total organic carbon, nitrogen, and 14C residue contents of the solid samples were determined, after combustion of 20 to 60 mg of powdered sample obtained after grinding in an agate mortar, at 1020°C in a Carlo–Erba (Milan, Italy) NA 1500 analyzer. The 14CO2 evolved during the combustion was trapped into 10 mL of 1 M NaOH before counting on a 1-mL aliquot as above.
Extractable 14C-Residues—Distribution of 14C-Residues among Humic Compounds
The release of 14C residues during incubation was evaluated, after exhaustive extractions by shaking the samples, for 16 h, first with distilled water (w/v), then with a mixture of methanol and water (80:20 [v/v]) and at last with distilled water. The extracts were separated from the samples by centrifugation at 4000 x g for 1 h and the radioactivity was determined in each supernatant. The procedure was repeated with each solvant until no appreciable radioactivity was detected in the sample extract. The 14C-nonextractable residue amount was evaluated as the difference between the total sample initial radioactivity and the sum of the 14C-CO2 and the total extractable 14C radioactivity.
Humic compounds were then extracted from the 0- to 50-µm fraction of the residual samples. For the 0- to 5000-µm samples, this was done after wet-sieving at 50 µm, and for the rendzine after decarbonation with 2 M HCl, to remove calcium carbonate that prevents the solubilization of humic substances in alkaline reagents (Bartoli and Burtin, 1979). Five grams of the fraction were shaken for 3 h with 80 mL of a 1% mixture of Na4P2O7 and 0.1 M NaOH. Undissolved material (humin) was removed by centrifugation at 13000 x g for 20 min. The process was repeated until colorless supernatant was obtained. The alkali extracts were adjusted to pH 1.5 with 2 M HCl to precipitate humic acids (HA), then stored overnight. The HA precipitate was separated from the soluble fulvic acids (FA) by centrifugation and then dissolved in 0.1 M NaOH. The radioactivity was counted in the FA and HA solutions.
The humin and 50- to 5000-µm fractions were air-dried and their 14C radioactivity determined by trapping the 14C-CO2 produced after combustion of 20 to 60 mg of sample. Distribution of 14C residues among humic compounds from incubated samples was compared with that of non-incubated samples (i.e., t = 0) treated in a similar manner.
Microflora Analysis
The aerobic microbial populations were enumerated from fresh soil samples, which have never been treated with atrazine, and from the incubated samples, by the dilution plate method using salt nutrient broth (Prolabo; Biokar, France), nutrient broth (Difco; Difco Laboratories, Detroit, MI), and potato dextrose agar (Difco). Enumerations of colony forming units (CFUs) for serial dilutions (10-3 to 10-8) were performed in triplicate. Colonies were counted after incubation of the plates in darkness at 24°C.
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RESULTS AND DISCUSSION
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Organic Matter Mineralization
Cumulative C-CO2 evolution values from samples during 100 to 120 d of incubation are shown in Fig. 1
. After 100 d of incubation, the total amounts of C-CO2 released from the soil reached 3.6 and 1.8% of the total organic carbon content for the brown soil 0- to 5000- and 0- to 50-µm fractions, respectively, and only 1.0 and 0.8% for the rendzina 0- to 5000- and 0- to 50-µm fractions, respectively. Very low mineralization rates were observed in the absence of microflora (i.e., for the abiotic samples), even after 100 d of incubation. The total CO2 evolution from abiotic samples occurred at essentially linear rates and was a result of the presence of a microbial strain resistant to thimerosal.
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Fig. 1. Cumulative evolution of total C-CO2 released from incubated samples. Boxes = biotic assays; circles = abiotic assays; open symbols = 0- to 5000-µm fractions; filled symbols = 0- to 50-µm fractions.
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For both brown soil fractions, the results obtained for biotic samples showed a similar decomposition pattern. The rate of CO2 evolution was initially very rapid, as 55 and 38% of the total CO2 evolved in 100 d of incubation was measured during the first 25 d for the 0- to 5000- and 0- to 50-µm samples, respectively. Higher mineralization of organic matter in the 0- to 5000-µm fraction was attributed to the presence of fresh and more labile materials in the coarse fraction (Berthelin et al., 1999).
For the rendzina, although more organic C was evolved as milligrams of C-CO2, the rate of CO2 evolution was relatively constant after 25 d and the amounts of organic C mineralized during the initial stages of incubation were small compared with the brown soil. This suggests that the readily decomposable organic C fractions in brown soil are greater than those in rendzina. Indeed, the presence of calcium carbonate in the rendzina samples prevented organic matter degradation (Duchaufour, 1976).
Bound Residue Mineralization
No 14C-CO2 was released from the abiotic samples compared with the biotic ones. Thus, microbial activity promoted the release, although very slight, of atrazine-bound residues (Fig. 2)
. For the brown soil a sort of disruption was observed at Day 50, similar to that observed for the total CO2 evolution. For the rendzina the curves of 14C-CO2 evolved were more linear and presented also some similarities with those of total CO2. Such results suggested that bound residue mineralization followed an identical pathway to that of endogenous organic matter mineralization. But contrary to the total CO2 evolution, for all samples a lag phase was observed at the beginning. It accounted either for the adaptation of the microflora, which was reestablished from a fresh soil suspension, or for the release of 14C-CO2 that occurred only when the natural compounds, where residues were incorporated, were degraded.
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Fig. 2. Cumulated 14C-CO2 released from samples incubated in biotic conditions. Open symbols = 0- to 5000-µm fractions; filled symbols = 0- to 50-µm fractions.
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For the brown soil, although the degradation of fresh labile organic matter present in the coarse fraction (>50 µm) led to a release of twice as much CO2 as the 0- to 50-µm fraction, few differences between the 14C-CO2 amount evolved from the two fractions were observed. The residue mineralization rate reached approximately 0.10% of the initial radioactivity. This suggests that the availability of labile organic matter in the coarse fraction enhanced its potential degradation by the microflora, but seemed to have no consequence on the residue mineralization, probably because residues were concentrated in the 0- to 50-µm fraction (Table 1), where organic matter is more humified.
In the 0- to 50-µm fraction of the rendzina, the mineralization was slow. It reached only 0.06% at the end of incubation, half of that obtained from the 0- to 5000-µm soil samples. According to Levanon (1993), the atrazine ring mineralization involved both bacterial and fungal soil populations. Hence, the low amounts of 14C-CO2 evolved may be due to the low availability of residues, but also to the low ring-cleaving microbial population as observed by Jayachandran et al. (1998), who found that N-ethyl-dealkylating populations in soil were more numerous than ring-cleaving microorganisms.
Microbial activity was also expressed by a decrease of sample pH. Whereas for abiotic treatment the pH stayed constant, for biotic samples, it decreased 0.5 to 1 unit, respectively, for the rendzina and the brown soil. The enumeration of microbial population showed an increase in the number of colonies after incubation (Table 2), whereas the type of colonies was less diversified. Comparing fresh soils and incubated soils, in the brown soil fractions all colony forming units increased. For the rendzina, if the number of bacteria increased, that of fungi decreased for the 0- to 5000-µm sample, while it increased in the 0- to 50-µm fraction. The population size was significantly larger in the rendzina samples than in the brown soil samples. Thus, there is no clear relationship between total plate count and the release of CO2.
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Table 2. Enumeration of microorganisms in colony forming units (CFU) (x 105) g-1 dry sample. Numbers in brackets correspond to standard deviation for triplicate.
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By comparison of the evolution of the ratio between cumulated rates of 14C-CO2 and C-CO2 (Fig. 3)
, the shape of the curves was similar for both brown soil fractions. The ratio increased up to 50 d of incubation, then the mineralization of atrazine residues occurred at the same rate as that of organic matter. As the same rate of 14C-CO2 evolved was obtained for both samples, the higher value of the ratio observed for the 0- to 50-µm fraction is clearly due to a more active mineralization of labile organic matter in the 0- to 5000-µm fraction than in the 0- to 50-µm fraction. For the rendzina the ratio increased with incubation time, suggesting that the atrazine residues became more available than in brown soil. Contrary to brown soil, for both rendzina fractions, few differences were observed between the total C-CO2 evolved. Hence, the higher ratio values for the 0- to 5000-µm fraction must be explained by a lower availability of the residues in the 0- to 50-µm fraction than the 0- to 5000-µm fraction.
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Fig. 3. Evolution of the ratio between cumulated rates of 14C-CO2 and C-CO2 during the sample incubation. Boxes = brown soil; circles = rendzina; open symbols = 0- to 5000-µm fractions; filled symbols = 0- to 50-µm fractions.
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Extractable Residues
Regardless of treatment (abiotic or biotic), a proportion of the 14C residues (i.e., 7–15%) was extractable after incubation. However, differences were observed between the amounts of radioactivity recovered by water and methanol (Fig. 4)
. For the 0- to 5000-µm fractions of the brown soil, more radioactivity was extracted with water than with methanol from abiotic samples, while it was the opposite for the biotic ones. Such a difference was less pronounced for the 0- to 50-µm fraction, where the total of 14C extractable residues reached up to twice that of the 0- to 5000-µm fraction. The presence of microflora resulted in a decrease in the amount of extractable residues and, especially for the 0- to 5000-µm samples, the amount of easily water-extractable residues. This increase in the residue stability was probably related to fresh organic matter degradation, by enhancing the sorption of atrazine residues on decomposing organic material.
For both rendzina fractions, with regard to the total extractable residues and the amounts recovered with water or methanol–water, no differences were observed between the biotic and abiotic samples. Thus, the residues in the soil organic matrix of the rendzina appear to have a high stability toward microbial remobilization. But, the 14C residues from the 0- to 5000-µm fraction were more easily extractable (i.e., with water) than those from the 0- to 50-µm fraction. Hence, organic compounds, especially humic compounds of the 0- to 50-µm fraction, seem to retain residues more strongly by hydrophobic bonds, while in the 0- to 5000-µm fraction, a part of the residues could be weakly sorbed on fresh organic material, becoming more extractable.
For both soil samples, the differences obtained between the amounts of water and water–methanol extractable residues confirmed those observed between the mineralization ratios presented in Fig. 3.
The release of extractable 14C-residues from abiotic samples resulted probably from physico–chemical processes owing to the sequence of drying and remoistening of the samples, although Shelton et al. (1995) observed the opposite (i.e., decreased extraction efficiencies as a consequence of drying and rewetting). Following exhaustive extractions to remove extractable residues, our samples were air-dried prior incubation and then remoistened at 80% of field capacity. Such treatments led to physical disruption of the soil structure, substrate desorption, and liberation of organic compounds entrapped in the internal voids (Van Gestel et al., 1991; Ladd et al., 1996).
Bound Residue Distribution within the Organic Fractions
The amounts of bound 14C among humic compounds are presented in Table 3. Except for the brown soil 0- to 50-µm sample, the distribution of 14C-nonextractable residues among humic compounds was somewhat similar, that is, the main part of the radioactivity was associated with the humin as observed by Capriel et al. (1985).
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Table 3. Distribution of 14C-atrazine bound residues among organic compartments, before and after incubation of soil samples. NI, non-incubated; A, abiotic; B, biotic; HA, humic acid; FA, fulvic acid. Results in percent of residual non-extractable radioactivity.
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For the rendzina, few changes were observed in the amounts and distribution of 14C residues after incubation. The radioactivity decreased in the 50- to 5000-µm fraction of the 0- to 5000-µm samples after incubation and that of humin showed a slight increase. The 0- to 50-µm organic matter fraction was slowly degraded and presented no changes in the distribution of natural humic compounds and 14C atrazine residues. Therefore, the easily or water-extractable fraction of the 0- to 5000-µm sample can be attributed to the 50- to 5000-µm fraction and to the FA compartment where radioactivity decreased with time.
For the brown soil, the distribution of residues was different between the two fractions. The main part of radioactivity (38%) was in humin for the 0- to 5000-µm samples and increased to 60% after incubation, while that of the coarse fraction (i.e., 50–5000 µm) and that of FA and HA decreased. Thus, the increase of 14C in humin must be related to the microbial degradation of labile organic matter and of alkaline-soluble humic compounds. By contrast, more radioactivity was associated with FA for the 0- to 50-µm sample and that in humin seemed to decrease slightly in the presence of microorganisms.
However, for the brown soil 0- to 5000-µm and the rendzina 0- to 50-µm samples, microbial activity led to an increase of radioactivity of the humin and a decrease of that of FA and therefore could participate in the formation of more stable residues. Such an evolution of the distribution of residues with time was also observed by Bertin and Schiavon (1989).
The decrease of FA radioactivity could result from a higher residue availability toward degradation from these organic compounds, as shown for other organic pollutants (Scheunert et al., 1992; Andreux et al., 1993).
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CONCLUSION
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The influence of microbial activity on the fate of bound residues was assessed by comparison of samples incubated under biotic and abiotic conditions with the corresponding non-incubated soil samples.
As observed from the results obtained in abiotic conditions, mechanical disruption and rearrangement of soil components, caused by soil desiccation and rewetting, make organic compounds available for subsequent mineralization. The amounts of these organic compounds varied with the aggregation state of the samples.
In the presence of microorganisms, 14C-atrazine residue mineralization occurs after a lag phase and seems to follow an identical pathway to that of endogenous organic matter decomposition. For both brown soil fractions, differences in total organic matter mineralization were observed, and the same rate of 14C residue mineralization was obtained. Bound residues from the 0- to 5000-µm fraction appeared to be less available toward biodegradation than those of the 0- to 50-µm fraction. This fact was also expressed by the lowest extractable residue amounts from the 0- to 5000-µm fraction and by the important proportion of 14C recovered with FA in the 0- to 50-µm fraction. In addition, the decomposition of less humified organic matter in the coarse fraction seemed to increase the formation of bound residues.
For rendzina it is the opposite: the mineralization rate of organic C from both fractions was quite similar, but residue seems to be more available from the 0- to 5000-µm fraction as also shown by the presence of more easily water-extractable residues.
By comparison of biotic and abiotic assays, it was difficult to attribute the amounts of extractable residues to microbial activity. In presence of microorganisms, 14C-residues are (except for the 0- to 50-µm brown soil fraction) less easily extractable, and hence microbial activity can be involved not only in the residue mineralization but can also lead simultaneously to the stabilization of residues in soil.
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