Environmental Science and Pollution Research

, Volume 22, Issue 2, pp 1240–1249 | Cite as

Accelerated biodegradation of selected nematicides in tropical crop soils from Costa Rica

  • Juan Salvador Chin-Pampillo
  • Elizabeth Carazo-Rojas
  • Greivin Pérez-Rojas
  • Víctor Castro-Gutiérrez
  • Carlos E. Rodríguez-Rodríguez
Research Article

Abstract

Degradation and mineralization behavior of selected nematicides was studied in soil samples from fields cultivated with banana, potato, and coffee. Degradation assays in most of the studied soils revealed shorter half-lives for carbofuran (CBF) and ethoprophos (ETP) in samples with a history of treatment with these compounds, which may have been caused by enhanced biodegradation. A short half-life value for CBF degradation was also observed in a banana field with no previous exposure to this pesticide, but with a recent application of the carbamate insecticide oxamyl, which supports the hypothesis that preexposure to oxamyl may cause microbial adaptation towards degradation of CBF, an observation of a phenomenon not yet tested according to the literature reviewed. Mineralization assays for CBF and terbufos (TBF) revealed that history of treatment with these nematicides did not cause higher mineralization rates in preexposed soils when compared to unexposed ones, except in the case of soils from coffee fields. Mineralization half-lives for soils unexposed to these pesticides were significantly shorter than most reports in the literature in the same conditions. Mineralization rates for soils with a previous exposure to these pesticides were also obtained, adding to the very few reports found. This paper contributes valuable data to the low number of reports dealing with pesticide fate in soils from tropical origin.

Keywords

Enhanced biodegradation Degradation Mineralization Carbofuran Terbufos Ethoprophos Tropical soils 

Introduction

An important fraction of the total economic activity in tropical countries is based on agriculture; extensive areas are planted with numerous crops. Many of these areas are typically characterized by constant high temperatures and humidity, which favor an increase in populations of pests. These conditions often result in an increased usage of pesticides. Indeed, many tropical countries are ranked among the top pesticide users per cropland area in the world, even though this behavior cannot be fully generalized because large variations do exist (Arbeli and Fuentes 2010).

In Costa Rica, pesticide usage in potato, banana, and coffee fields was 37, 49, and 6.5 kg active ingredient (a.i.)/ha, respectively, in 2007. In that same year, an average of 26 kg a.i./ha of pesticides was used in the country (Vega 2012). Contrastingly, 11.4 kg a.i./ha were used in croplands in 2012, which represents a marked reduction compared to the total amount applied in 2007 (Servicio Fitosanitario del Estado 2014); however, this amount is still regarded as high in comparison with other countries in the region. In 2012, the potato-, banana-, and coffee-planted areas were 2,740, 41,426, and 93,774 ha (SEPSA 2013). Even with the aforementioned reduction in pesticide use at the national level, and probably at the crop level too, the large areas planted with coffee and banana represent a considerable consumption of pesticides.

Plant parasitic nematodes are serious pests in banana, potato, and coffee crops in Costa Rica, and application of nematicides is the most widely used control method for this problem (López and Azofeifa 1981; Chaves 2007). Between 2006 and 2009, insecticide-nematicide imports represented 6.8 % of the total pesticide imports. Terbufos (TBF) was the most imported product in this category. It is used mainly in banana, coffee, and corn crops. Ethoprophos (ETP) follows; it is used in banana, pineapple, melon, coffee, and sugarcane crops, vegetables, and ornamentals. Other insecticides-nematicides imported in lower quantities are oxamyl, carbofuran (CBF), and fenamiphos which are used also in banana, coffee, pineapple, and melon fields (Vega 2012).

Many of the aforementioned nematicides exhibit a high acute and chronic toxic potential for humans and animals, and some of them are severely regulated or completely banned in other countries, with CBF, ETP, and TBF being listed among them (Wesseling et al. 2003; Ruepert 2011). CBF has been banned in agricultural practice in the USA and the European Union, but it is still used in Costa Rica.

CBF, a carbamate, and TBF and ETP, organophosphates, exert their activity through the inhibition of acetylcholinesterase, an enzyme that degrades the neurotransmitter acetylcholine in synapses from vertebrates and invertebrates. CBF is highly toxic to mammals and reportedly embryotoxic and teratogenic (Gupta 1994; García 1997). Regarding organophosphates, even though considered biodegradable, they are highly toxic towards vertebrates and invertebrates, and their deleterious effects in wildlife in general are well documented (Galloway and Handy 2003). Given the high toxic potential of these compounds, the dynamics of their environmental degradation are of particular importance. Several cases involving intoxication and environmental effects derived from pesticide misuse in Costa Rica have been reported and compiled (García 1997; De la Cruz et al. 2004; Ruepert 2011).

Major factors that contribute to pesticide degradation from soil include photodegradation, volatilization, and chemical and microbial degradation. Microbial degradation is regarded as the main mechanism through which pesticides in general (including CBF, TBF, and ETP) are eliminated from soil. The term accelerated biodegradation or enhanced biodegradation is applied to cases in which one or more previous applications of a pesticide cause a significant increase in the rate of biodegradation of such compound in a subsequent treatment. It may also be applied to a closely related phenomenon in which a rapid degradation of a pesticide occurs, caused by a previous application of a different pesticide with a similar chemical structure. Many pesticides, including herbicides, insecticides, nematicides, fungicides, and fumigants, have been proven to undergo this process (Arbeli and Fuentes 2007).

Research regarding pesticide fate in tropical environments is rather scarce; as a consequence, predictions related to the degradation of these compounds are less precise in these areas than in temperate zones (Arbeli and Fuentes 2010). Consequently, recommendations have been issued to promote pesticide degradation studies in these areas in order to adequately define specific guidelines for pesticide use (Racke et al. 1997).

Despite the existence of evident environmental, health-related, and economic implications of pesticide use in Costa Rica, very few reports dealing with enhanced biodegradation of these compounds have been carried out in the country (Anderson et al. 1998; Moens et al. 2004; Cabrera et al. 2010). This study aims to assess pesticide degradation and mineralization rates and to explore the phenomenon of enhanced biodegradation of nematicides in soil samples from different representative cropland areas in Costa Rica. Data obtained will aid in estimating the environmental risk and the possibility of enhanced biodegradation associated with the use of these nematicides in crops widely cultivated in the tropics.

Materials and methods

Chemicals

Formulated pesticides include CBF (Furadan® 10G 10 % w/w), TBF (Counter® 15G 15 %), ETP (Mocap® 15GR 15 % w/w). Primary standards are as follows: CBF (99.5 %, Chem Service), TBF (99.5 %, Chem Service), ETP (98.9 %, Chem Service). Radiolabeled compounds are 14C-CBF ([benzene ring-U-14C] carbofuran: specific activity 2.17 MBq/mg, radiochemical purity 100 %, chemical purity 97.4 %; Institute of Isotopes Co., Budapest, Hungary), 14C-TBF ([O-ethyl-1-10-C2] terbufos: specific activity 2.1 MBq/mg, radiochemical purity 96 %; Institute of Isotopes Co., Budapest, Hungary). Solvents include dichloromethane (GC grade, Merck), hexane (GC grade, SupraSolv®, Merck), acetone (GC grade, SupraSolv®, Merck), and methanol (LC grade, Merck). Physical characteristics of the pesticides used in this study that may affect their degradation are shown in Table 1.
Table 1

Physical properties of CBF, ETP, and TBF that may affect their degradation

 

Pesticide

CBF

ETP

TBF

Water solubility at 20 °C (mg/L)

322

1,300

4.5

Octanol-water partition coefficient (KOW) at 20 °C, pH 7.0

6.31 × 101

9.77 × 102

3.24 × 104

Vapor pressure at 25 °C (mPa)

0.08

78

34.6

Henry’s law constant at 20 °C (dimensionless)

2.09 × 10−8

6.10 × 10−6

1.09 × 10−3

Source: University of Hertfordshire (2013)

Soil samples

Soil samples were collected from fields cultivated with three different types of representative crops from Costa Rica, namely banana (Musa sp.), three fields: B1 (Siquirres, 10°05′54″N, 83°30′32″W, 62 m above sea level (masl)), B2 (San Alberto), and B3 (Guácimo, 10°12′46″N, 83°41′12″W, 114 masl) from Limón Province; potato (Solanum tuberosum), three fields: P1 (Pacayas, 9°54′39″N, 83°49′27″W, 1,935 masl), P2 (San Juan de Chicuá, 9°57′44″N, 83°49′57″W, 2,947 masl), and P3 (Tierra Blanca, 9°56′7″N, 83°52′57″W, 2,379 masl) from Cartago Province; and coffee (Coffea arabica), two fields: C1 (San Ramón, 10°04′39″N, 84°28′25″W, 1,065 masl) and C2 (Santa Ana, 9°55′24″N, 84°09′48″W, 1,165 masl) from the provinces of Alajuela and San José, respectively. Soil samples were randomly collected at a depth of 0–30 cm from the surface. Soil physical and chemical characteristics were determined and are shown in Table 2. Ca and Mg were extracted with a 1-N KCl solution, while K, P, Zn, Fe, Cu, and Mn were extracted with a modified Olsen solution. Cations were measured using an atomic absorption spectrophotometer. P was measured spectrophotometrically. To estimate the organic matter percentage, total carbon percentage was determined by dry combustion using a C/N autoanalyzer, and this value was multiplied by a factor of 1.43. Pesticide use history for selected nematicides analyzed in this study was recorded for each of the fields and is presented in Table 3.
Table 2

Physical and chemical characteristics of soil samples from potato, banana, or coffee fields

 

B1

B2

B3

P1

P2

P3

C1

C2

Sand (%)

26

57

59

60.1

59

62.5

23

58

Silt (%)

45

32

25

32.9

36.8

30

23

22

Clay (%)

29

12

17

7.1

4.2

7.5

54

20

Soil texture

Clay loam

Sandy loam

Sandy loam

Sandy loam

Sandy loam

Sandy loam

Clay

Sandy loam

Organic matter (%)

2.2

1.4

8.0

8.6

4.4

3.3

7.0

7.8

pH

5.4

6.3

5.5

5.8

5.6

5.6

4.6

5.6

Acidity (cmol (+) L−1)

0.72

0.47

0.45

0.19

0.4

0.22

1.96

0.27

Ca (cmol (+) L−1)

21.88

13.23

8.62

9.97

5.01

10.29

1.38

9.16

Mg (cmol (+) L−1)

7.01

2.9

2.63

3.81

1.13

2.59

0.44

2.13

K (cmol (+) L−1)

1.78

0.7

0.39

0.03

0.82

0.07

0.54

0.55

CEC (cmol (+) L−1)

31.39

18.3

12.09

14

7.36

13.17

4.32

12.11

SA (%)

2

3

4

1

5

2

45

2

P (mg L−1)

38

48

8

21

111

9

13

7

Zn (mg L−1)

5.4

7.3

4

7.4

5.6

4.6

9.6

4.4

Cu (mg L−1)

9

4

10

11

19

10

32

8

Fe (mg L−1)

29

104

192

134

348

95

270

209

Mn (mg L−1)

18

7

15

2

5

ND

46

11

CEC cation exchange capacity (acidity + Ca + Mg + K), SA saturation acidity (acidity / CEC) × 100

Table 3

Treatment history over the last four years for selected nematicides in banana, potato, and coffee fields sampled

Soil

Insecticide-nematicide

Pesticide application history

B1

CBF

Last application 10 months before sampling

B2

CBF

Last application 23 months before sampling

B3

CBF

None

P1

CBF

Last application 36 months before sampling

P2

CBF

None

P3

CBF

None

C1

CBF

Last application 12 months before sampling

C2

CBF

None

B1

ETP

Last application 10 months before sampling

B2

ETP

Last application 9 months before sampling

B3

ETP

Last application 20 months before sampling

P1

ETP

None

P2

ETP

None

P3

ETP

None

C1

ETP

None

C2

ETP

None

B1

TBF

Last application 7 months before sampling

B2

TBF

Last application 11 months before sampling

B3

TBF

Last application 4 months before sampling

P1

TBF

None

P2

TBF

None

P3

TBF

None

C1

TBF

Ongoing application when sampling

C2

TBF

None

Degradation assays

For degradation assays, the collected soil was air-dried for 1 day and homogenized. Degradation systems were prepared by placing 750 g of soil in 1-L open plastic containers. CBF or TBF was added (12 mg/kg), soil was homogenized manually, and the system was incubated for 128 days in a greenhouse room at ambient temperature (mean 20.3 °C, min. 14.1 °C, max. 28.3 °C). Humidity ranged from 79 to 91 %. The systems were not exposed to direct sunlight. Moisture was kept constant by regular addition of deionized water to the systems. The remaining pesticide concentration was determined by sacrificing triplicate systems at the following time points: 0, 4, 8, 16, 32, 64, and 128 days.

For ETP, soil samples (50 g) were successively extracted with acetone (65 mL total) in an ultrasonic bath. The extract was then subjected to a liquid-liquid extraction using a dichloromethane-hexane mixture (1:1). Separation and quantification was done with a gas chromatograph and a FPD detector (limit of detection (LOD) 0.046 μg/kg, limit of quantification (LOQ) 0.089 μg/kg). Confirmation was carried out through single quadrupole mass spectrometry. Gas chromatography was performed in an Agilent 6890N GC equipped with a HP 5MS column ((5 %-phenyl-)-methylpolysiloxane; 30 m × 0.25 mm × 0.25 μm) and coupled to a FPD detector. The injector was operated in splitless mode, temperature of the injector was 260 °C, and injection volume was 1 μL. The column oven temperature was programmed as follows: 50 °C for 0.5 min, 25 °C min−1 to 140 °C, 5 °C min−1 to 159 °C, and 10 °C min−1 to 256 °C (2 min). Confirmation was done with a single quadrupole mass spectrometer (Hewlett-Packard), using the following ramp: 100 °C for 1 min, 25 °C min−1 to 210 °C, 7 °C min−1 to 240 °C, and 30 °C min−1 to 280 °C (5 min). Ions used for ETP in GC/MS were as follows: m/z = 157.9, 139.0, 126.0, and 200.0. The method is accredited under ISO/IEC 17025:2005. For quality control, spiked samples were used. Recoveries within 60–120 % were considered acceptable.

For CBF, soil samples (5 g) were extracted using methanol in an ultrasonic bath and a methanol-HCl 0.1 N (2:1) mixture. The extracts were then subjected to a liquid-liquid extraction with dichloromethane. High-pressure liquid chromatography was used for quantification, with post column derivatization and detection by fluorescence (Waters) (emission wavelength 445 nm, exCitation wavelength 339 nm) (LOD 5.3 μg/kg, LOQ 10.0 μg/kg). Chromatographic separation was done with a 3.9- × 150-mm column (Waters) at 25 °C with a mobile phase consisting of Milli-Q water (A), methanol (B), and acetonitrile (C) at 1.5 mL min−1. The following gradient was employed: elution started at 88 % A-12 % B (0–4 min), followed by a linear gradient from 68 % A-15 % B-16 % C to 30 % A-35 % B-35 % C (4.1–16 min) and finally returning to 88 % A-12 % B (16–35 min). The method is accredited under ISO/IEC 17025:2005. For quality control, spiked samples were used. Recoveries within 60–120 % were considered acceptable.

Pesticide degradation was modeled through a single first-order model: C = C0 ekt, where C and C0 represent pesticide concentration present at time t and time t = 0, respectively, and k is the removal rate.

Mineralization assays

Mineralization of CBF and TBF was determined through 14CO2 production in biometric systems. For the preparation of the systems, collected soil was air-dried, mortar-ground, and sieved (500 μm). Biometric systems were constructed using 400-mL sterile glass jars, soil (50 g), and commercial CBF or TBF (12 mg a.i./kg). 14C-CBF or 14C-TBF solution was also added to the soil to achieve a total activity of 3,000 dpm g−1 of soil. Sterile deionized water was added to the soil in each system in order to obtain 75 % field capacity. Soil was then homogenized manually in each system. 14CO2 traps were prepared by adding KOH (10 mL, 0.1 M) to 50-mL glass flasks suspended by copper wires inside the jars. A catheter was placed in the upper section of each jar and was used to remove and add the KOH solution when needed.

The systems were incubated at 19 ± 1 °C in the dark for a period of 64 days. Every 4 days, the entire KOH solution of three replicate biometric systems was removed through the catheter and replaced with the same volume of a fresh solution. At the same interval, systems were weighed and sterile deionized water was added to keep humidity constant. A 2-mL aliquot was taken from the removed solution, 8 mL of scintillantion fluid (Ultima Gold, PerkinElmer) was added, and the 14C activity was measured by liquid scintillation using a Beckman LS6000SC counter.

The total cumulative 14CO2 activity evolved (mineralized) and the initially added 14C-CBF and 14C-TBF activities were used to calculate the percentage of 14C-pesticide mineralized. Pesticide mineralization was modeled using a single first-order model, as described for degradation assays.

Results and discussion

Degradation assays

For all soils, a single first-order model adequately described CBF and ETP removal behavior. Calculated removal rate constants and half-lives are shown in Table 4. The amount of pesticide (CBF and ETP) remaining was plotted against incubation time for representative selected crop soils (Fig. 1). Since light exposure of the soil systems was homogeneous, photodegradation is not expected to have caused important differences in pesticide degradation half-lives between tested soils.
Table 4

Correlation coefficients (r), removal rate constants (k), and half-lives (t1/2) for first-order fitting of CBF and ETP degradation in soil

Soil

Insecticide-nematicide

r

k (day−1)

t1/2 (days)

B1

CBF

0.997

0.0513 ± 0.0021

13.5 ± 0.6

B2

CBF

0.935

0.118 ± 0.022

5.9 ± 1.1

B3

CBF

0.987

0.0780 ± 0.0064

8.9 ± 0.7

P1

CBF

0.987

0.1056 ± 0.0099

6.6 ± 0.6

P2

CBF

0.881

0.0232 ± 0.0072

29.9 ± 9.3

P3

CBF

0.946

0.0235 ± 0.0046

29.5 ± 5.8

B1

ETP

0.880

0.102 ± 0.028

6.8 ± 1.8

B2

ETP

0.888

0.051 ± 0.015

13.7 ± 4.0

B3

ETP

0.929

0.085 ± 0.017

8.2 ± 1.6

P1

ETP

0.974

0.0445 ± 0.0046

15.6 ± 1.6

P2

ETP

0.928

0.0446 ± 0.0080

15.5 ± 2.8

P3

ETP

0.955

0.0413 ± 0.0065

16.8 ± 2.6

Fig. 1

Removal of CBF and ETP in different crop soils: CBF in soil B1 (open upright triangles), CBF in soil P1 (filled inverted triangles), ETP in soil B3 (open circles), and ETP in soil P2 (filled circles)

CBF

For CBF, volatilization is generally considered a minor route of elimination from soil (Howard 1991). Chemical hydrolysis and microbial degradation appear to be the most important degradation processes for CBF in soil (Howard 1991). Chemical hydrolysis is expected to occur more rapidly in alkaline soil as compared to neutral or acidic soils, in which microbial degradation plays a more important role (Getzin 1973). In this study, CBF degradation half-lives ranged from 5.9 to 29.9 days. Since all studied soils had an acid pH (Table 2), differences in degradation half-lives are considered to be mostly due to the microbial degradation component.

The longest CBF degradation half-lives corresponded to soil samples from two potato fields (P2 and P3) in which no prior CBF exposure was documented (29.9 and 29.5 days, respectively). Comparatively shorter degradation half-lives (13.5, 5.9, and 6.6 days) were observed in soils from banana and potato fields (B1, B2, and P1) with a history of previous CBF treatments 10, 23, and 36 months earlier. These results are overall consistent with previous studies in which enhanced biodegradation of CBF has been proven in different soil types (Felsot et al. 1981; Harris et al. 1984; Read 1987) even when the last CBF application in the field had occurred 4 years (Karpouzas et al. 1999) or 5 years earlier (Suett et al. 1993). Moens et al. (2004) observed enhanced biodegradation of CBF in nematode count bioassays with soil samples from the same province as the banana fields in this study. Data obtained are also in accordance with previous studies which found CBF half-lives ranging from 26.1 to 44.1 days in topsoil samples with no previous exposure to CBF and half-lives ranging from 1.5 to 14.4 days in topsoil samples pretreated with 10 mg/kg of the pesticide (sandy loam, organic matter (O.M.) 3.46 %, pH 6.6, water holding capacity 40 %, 10 mg/kg, 15 °C) (Karpouzas et al. 2001).

Soil samples from field B3 showed a rapid degradation of CBF (half-life 8.9 days) even though no previous exposure to CBF was recorded. Field B3 had a previous treatment with oxamyl, a carbamate family pesticide, less than a month before soil samples were collected from the field. Oxamyl, as is the case for CBF, belongs to the N-methyl carbamate family of nematicides. This oxamyl exposure may have predisposed the soil microflora for a rapid degradation of CBF when it was subsequently applied in the laboratory. Cross-enhancement of CBF biodegradation in soil samples previously treated with other carbamate pesticides has been observed several times (Read 1987; Racke and Coats 1988a; Morel-Chevillet et al. 1996) and is regarded as a well-known phenomenon. However, no reports were found in the literature describing the extent of CBF degradation after soil treatment with oxamyl. In a related observation, the effect of CBF treatment in soil on the degradation of oxamyl was studied by Harris et al. (1984). They found that in a soil sample, oxamyl was actively degraded after the sample was treated four times with a 10-mg/kg dose of CBF, but not after a single treatment or no treatment at all. Therefore, cross-acclimation caused by oxamyl treatment constitutes a plausible explanation for the results obtained; nevertheless, specific assays should be carried out to confirm this hypothesis.

The elevated organic matter content in soil from field B3 (8.0 %) may have also contributed to the rapid degradation of CBF in soil from the previously unexposed field B3 by enhancing microbial activity related to pesticide degradation (Perucci et al. 2000). Even though no correlation was determined between organic matter content and half-lives when all the soils were considered, when preexposed and unexposed soils were considered separately, a correlation was found among these variables only in the case of unexposed soils (close to linear, r = 0.971). This finding might suggest that organic matter content could be important at enhancing unspecific microbial activity; meanwhile, in preexposed soils, the efficiency of an adapted microbiota seems to overshadow the effect of organic matter.

ETP

The main mechanism of dissipation for ETP in soils is degradation through microbial metabolism (Cochran 1995). ETP degradation half-life values ranged from 6.8 to 16.8 days (Table 4). Mean values were significantly shorter (p = 0.041) in soils from banana fields (6.8–13.7 days) when compared to soils from potato fields (15.5–16.8 days). All three banana fields have had at least one previous exposure to ETP in the last 2 years, while ETP applications were not recorded for potato fields in recent years (Table 3).

As with CBF, enhanced biodegradation of ETP may be responsible for its shorter half-life in soil from previously exposed fields. Accelerated biodegradation has been observed for ETP in the past (Smelt et al. 1996), including a report using nematode count assays with soil samples from the same region as in this study (Moens et al. 2004). Karpouzas et al. (1999) found that a single application of the pesticide in the previous year allowed an enhanced degradation of ETP in soil samples from Greece. Further study on soils from Northern Greece led to the conclusion that enhanced biodegradation of ETP can be very stable once established, being evident even 2 years after the last ETP treatment (Karpouzas and Walker 2000). Other investigators have stated that accelerated biodegradation of ETP can persist for at least 3 years, returning to normal when measured after 5 years (Smelt et al. 1996).

Degradation half-lives are in accordance with previous studies. Smelt et al. (1996) found degradation half-lives ranging from 26 to 41 days for previously untreated soils, while in soils with a recent history of ETP application (≤3 years), calculated half-lives ranged from 1.3 to 41 days (sandy and loamy soils, O.M. 1.3–2.3 %, pH 5.6–7.7, moisture 10–17 %, 4–8 mg a.i./kg of dry soil, 15 °C).

In the past, various organophosphate nematicides different from ETP (such as phorate and chlorpyrifos) had been periodically applied to the potato fields studied in this investigation. However, cross-acclimation between these pesticides and ETP, leading to shorter half-lives, is not evident from the results obtained. According to previous investigations, it appears that cross-enhanced biodegradation is not as frequent with organophosphorous pesticides as it is with carbamates (Forrest et al. 1981; Arbeli and Fuentes 2007). Anderson et al. (1998), in a study using soil samples from Australia, Costa Rica, Honduras, and Germany, pointed out that microorganisms adapted to degrade a particular organophosphate pesticide are quite specific for that particular compound. Similarly, Karpouzas and Walker (2000) found no cross-enhancement leading to the rapid degradation of various organophosphorus nematicides in soils previously treated with ETP.

As described for CBF, a linear correlation (r = 0.996) was found between organic matter content and ETP half-lives only when unexposed soils were considered.

Mineralization assays

Single first-order model adequately described CBF and TBF mineralization behavior for soils from banana and potato fields, while data from coffee fields adjusted less accurately to the model. Calculated rate constants and half-lives are shown in Table 5. The amount of 14CO2 evolved shown as a percentage of the initially added 14C-CBF or 14C-TBF activity was plotted against time (Figs. 2 and 3).
Table 5

Correlation coefficients (r), removal rate constants (k), and half-lives (t1/2) for first-order fitting of CBF and TBF mineralization in soil

Soil

Insecticide-nematicide

r

k (day−1)

t1/2 (days)

B1

CBF

0.998

0.0129 ± 0.0002

53.7 ± 0.9

B2

CBF

0.997

0.0129 ± 0.0003

53.7 ± 1.1

B3

CBF

0.998

0.0152 ± 0.0003

45.6 ± 0.8

P1

CBF

0.999

0.0133 ± 0.0002

52.1 ± 0.7

P2

CBF

0.981

0.0196 ± 0.0010

35.4 ± 1.8

P3

CBF

0.998

0.0120 ± 0.0002

57.8 ± 1.0

C1

CBF

0.907

0.0102 ± 0.0012

68.0 ± 7.7

C2

CBF

0.979

0.0056 ± 0.0015

123 ± 32

B1

TBF

0.984

0.0117 ± 0.0007

59.2 ± 3.3

B2

TBF

0.993

0.0101 ± 0.0004

68.6 ± 2.5

B3

TBF

0.987

0.0150 ± 0.0007

46.2 ± 2.1

P1

TBF

0.991

0.0120 ± 0.0004

57.8 ± 2.2

P2

TBF

0.987

0.0226 ± 0.0010

30.7 ± 1.4

P3

TBF

0.988

0.0117 ± 0.0005

59.2 ± 2.5

Fig. 2

Mineralization of CBF at spiking level of 3,000 dpm/g of in different soils, as determined by the cumulative 14CO2 produced from 14C-CBF. (A) Soils from potato crops: P1 (filled circles), P2 (open circles), and P3 (filled inverted triangles). (B) Soils from banana crops: B1 (filled circles), B2 (open circles), and B3 (filled inverted triangles). (C) Soils from coffee crops: C1 (filled circles) and C2 (open circles). Values plotted are means of triplicates; maximum relative standard deviations (RSD) for each plot are 16 % (P1), 3 % (P2), 17 % (P3), 17 % (B1), 13 % (B2), 41 % (B3), 5 % (C1), and 3 % (C2)

Fig. 3

Mineralization of TBF at spiking level of 3,000 dpm/g in different soils, as determined by the cumulative 14CO2 produced from 14C-TBF. (A) Soils from potato crops: P1 (filled circles), P2 (open circles), and P3 (filled inverted triangles). (B) Soils from banana crops: B1 (filled circles), B2 (open circles), and B3 (filled inverted triangles). Values plotted are means of triplicates; maximum relative RSD for each plot are 5 % (P1), 21 % (P2), 36 % (P3), 50 % (B1), 43 % (B2), and 26 % (B3)

CBF

CBF mineralization half-lives ranged from 35.4 days (P2) to 123.8 days (C2). Mineralization half-life values for CBF were in all cases longer than degradation half-life values (1.2 to 9.1 times longer, 4.9 on average). For mineralization assays, a complete transformation of the parent compound to 14CO2 is required for detection, while for degradation assays, any change in the parent compound is significant. Considerably longer mineralization half-lives (compared to degradation half-lives) suggest that the initial transformation of the parent compound is not the rate-limiting step, but instead, degradation of CBF metabolites is probably responsible of limiting the mineralization rate.

The half-life from the unexposed coffee field almost doubled that from the preexposed field (123.8 vs 68.0 days). On the other hand, and contrary to degradation rates, there is no apparent correlation between higher mineralization rates for CBF and previous exposure in soils from banana and potato fields. CBF mineralization half-lives from unexposed soils (35.4–123.8 days) were significantly shorter when compared to most reports in the same condition. Smith et al. (2006) calculated half-lives ranging from 87 to 622 days on four different unexposed tropical soils from Sri Lanka (clay content <1 to 40 %, organic C (O.C.) 0.15–7.6 %, pH 4.5–6.9, max. water holding capacity 75 %, unlabeled pesticide 5 mg/kg, labeled pesticide per sample 1,850 Bq or 0.05 μCi, 28 °C, day/night light cycle). Benicha et al. (2013) tested a surface soil from Northwestern Morocco and found that after a 63-day incubation, only 20.3 % of the initially added 14C-CBF was mineralized (clayey soil, O.M. 2.2 %, pH 7.2, water holding capacity 60 %, 10-mg/kg unlabeled pesticide, 0.25-μCi-labeled pesticide per sample, 25 °C, in the dark). Ou et al. (1982) found mineralization half-lives for CBF in six different unexposed soils from the USA ranging from 216 to 944 days (clay content 6–35 %, O.C. 0.8–3.9 %, pH 5.6–7.7, soil/water tension 0.1 to 15 bar, unlabeled pesticide 10 mg/kg, labeled pesticide 0.1 μCi/g, 15–35 °C).

Few reports dealing with CBF mineralization in preexposed soils are found in the literature. Half-lives of previously exposed soils in this study (52.1–68.0 days) were shorter when compared to some reports but longer when compared to others. The mineralization behavior of CBF was assayed by Zayed et al. (2001) on previously treated soil samples from Egypt (clay loam, O.M. 1.1 %, pH 7.7, max. water holding capacity 55 %, 10-mg/kg labeled pesticide, 25 °C), who found that after 90 days, only 12 % of the initially added 14C-CBF was mineralized. However, in assays performed by Trabue et al. (1997) using surface soil samples from Hastings, Florida (fine sand, pH 5.9, O.C. 4.2 g/kg, moisture 99 mL/kg, 10-mg/kg unlabeled pesticide, 1.7-KBq-labeled pesticide per sample), more than 90 % mineralization of 14C-CBF was observed after 3 days. The elevated percentage of sand in the soil used in their study may have limited pesticide absorption, therefore promoting biodegradation.

TBF

Hydrolysis and microbial degradation are the primary degradation processes for TBF in the environment when it is applied to soil (US EPA 2003). TBF mineralization half-lives in the present study ranged from 30.6 days (P2) to 68.4 days (B2), but no apparent correlation between TBF use history (Table 3) and shorter TBF mineralization half-lives (Table 5) can be inferred from the data. Enhanced biodegradation of TBF has been observed in several studies (Chapman and Harris 1990; Pattison and Versteeg 2000) including a report with soil samples from banana plantations in Limón Province of Costa Rica (Cabrera et al. 2010). However, other investigators failed to find enhanced biodegradation of TBF even after eight and five applications of this compound in banana plantations from the country (Behm et al. 1991; Moens et al. 2004).

TBF mineralization in unexposed soils from this study (P1, P2, and P3) seems to occur at faster rates when compared to most studies with soils in the same condition. Anderson et al. (1998) found that only 5.7 and 1.1 % of the applied 14C-TBF was mineralized to 14CO2 in soils with no history of TBF treatment from Germany and Costa Rica, respectively (O.C. 0.8 %, pH 5.3/O.C. 0.9 %, pH 3.6) after a 4-week incubation (40–55 % water holding capacity, 10 mg a.i./kg dry soil labeled pesticide, 20 ± 2 °C, in the dark). A study by Racke and Coats (1988b) in six different soils found that 12.2–18.5 % of 14C-TBF was mineralized to 14CO2 after 4 weeks (clay 22.9–36.4 %, O.M. 2.5–4.8 %, pH 6.9–7.4, field capacity moisture, 5 mg/kg labeled pesticide, 25 °C, in the dark). Comparatively, in unexposed soils from our study, after 4 weeks, 33.0 to 55.5 % of 14C-TBF was mineralized to 14CO2.

Very few reports in which the mineralization of TBF is analyzed in previously exposed soils are found in the literature. Racke and Coats (1990) found 12.5 % mineralization of 14C-TBF in pretreated soil samples after one week incubation (0.3 bar soil moisture tension, 5 μg/g labeled pesticide, 25 °C, in the dark). These results are consistent with those found in this study, in which mineralization in previously exposed soils ranged from 10.9 to 20.9 % after 8 days.

Conclusions

Degradation assays performed in this study provide further evidence that enhanced biodegradation of CBF and ETP, previously reported mostly in studies from temperate zones, also occurs in cultivated soils from tropical areas. Cross-acclimation between carbamate pesticides is a well-known phenomenon, but evidence of a probable cross-enhanced biodegradation of CBF following oxamyl treatment was found, being a previously unreported observation in the literature reviewed. Mineralization rates for CBF and TBF in unexposed soils found in this study were faster than previous reports from temperate and tropical areas in the same condition. The same type of studies using preexposed soils are very scarce; therefore, valuable data on the environmental fate of these pesticides, particularly in tropical soils, was obtained. In addition, half-life values estimated in this study may be helpful for future design of pesticide use policies that take into consideration degradation and mineralization behavior of the tested nematicides in the tropical context.

Notes

Acknowledgments

This work was supported by the International Atomic Energy Agency (partial results of TC/COS5026), and Centro de Investigaciones Agronómicas (CIA), Universidad de Costa Rica.

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Copyright information

© Springer-Verlag Berlin Heidelberg 2014

Authors and Affiliations

  • Juan Salvador Chin-Pampillo
    • 1
  • Elizabeth Carazo-Rojas
    • 1
  • Greivin Pérez-Rojas
    • 1
  • Víctor Castro-Gutiérrez
    • 1
    • 2
  • Carlos E. Rodríguez-Rodríguez
    • 1
  1. 1.Centro de Investigación en Contaminación AmbientalUniversidad de Costa RicaSan JoséCosta Rica
  2. 2.Facultad de MicrobiologíaUniversidad de Costa RicaSan JoséCosta Rica

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