1. INTRODUCTION:

A large number of pesticides are used on agricultural crops throughout the world ¹and majority of them have some toxic effects to both animals and humans. Moreover, excessive uses of pesticides adversely affect the soil fertility and can change the soil chemistry which might eventually become totally unfit for growing crops. The organophosphorous pesticides interfere with the activity of an enzyme acetylcholine sterase which plays an important role in the normal transmission of nerve impulse. Most of the organophosphorous pesticides have a chemical structure containing three phosphoester linkages.

The toxicity of the pesticides is greatly reduced when the hydrolysis of one of the phosphoester bonds takes place, thus eliminating their acetylcholine sterase inactivating properties².

Monocrotophos [Dimethyl (E)-1-methyl-2(methylcarbamoyl) vinyl phosphate], is an organo phosphorus pesticide which is being used extensively used in agriculture. It is a chlorinated organo phosphorous insecticide, acaricide and termicide against cutworms, gall midge, leaf folder, leaf hopper, etc. The half-life of monocrotophos varies from 10 to 120 days which largely depends upon environmental factors like temperature, pH, moisture content, pesticide formulation and organic carbon content. The major degradation product of monocrotophos is more water soluble than monocrotophos itself and thus it causes a high range of contamination in soil and in aquatic environment.

Different technologies have been implemented including physicochemical and biological treatments to solve the problem of soil and water pollution. It is worthy to mention that organophosphorus pesticides causes toxicity to humans and animals and greatly affects the central nervous system. Biological treatment using microorganisms has been confirmed as the best technique in reducing soil pollution as it is environment friendly and inexpensive. Many reports have shown the efficiency and potential of microorganisms to degrade xenobiotic compounds³. The degradation of pesticide was observed in alkaline soils and its phenomenon was related to its hydrolysis in high pH. However, complete hydrolysis of monocrotophos was observed with high pH in soil under sterile condition which indicated the involvement of soil microorganisms⁴. Previous works have reported that MCP could be degraded both in broth and in soil medium by bacteria such as Pseudomonas aeruginosa, Clavibacter michiganense, Arthrobacter atrocyaneus, Agrobacterium radiobacter, Bacillus megaterium and Pseudomonas mendocina ^2,6,5. However, very limited information is available regarding mycoremediation of pesticides.

In the present study, fungal strains were isolated possessing the capability of degrading monocrotophos completely. The biodegradation of monocrotophos and its major metabolite in mineral medium and soil were observed. This study actually aims the possible elucidation of the isolated fungal strains for the bioremediation of the monocrotophos contaminated environment.

2. MATERIALS AND METHODS:

2.1. Chemicals:

Certified analytical grade monocrotophos (99% purity) was purchased from Sigma-Aldrich. The technical grade monocrotophos, a 36% emulsifiable concentrate used in this study was procured from United Phosphorus Ltd, (Gujarat) India. All other reagents used in this study were of high purity and analytical grade.

2.2. Soil sample:

Soil sample was collected from the top layer (0-20 cm) of the paddy field which had been exposed to monocrotophos pesticide in Vellore district, Tamil Nadu, India. The soil sample was dried at the room temperature in the laboratory and sieved.

2.3. Enrichment procedure and isolation of fungal strain:

Monocrotophos degrading fungal strains were obtained by enrichment culture in the Czapek Dox broth containing (g/l) yeast extract 3; peptone 10; dextrose 2; and monocrotophos 100 mg/l. Approximately 5 g of soil sample was inoculated in 50 ml of Czapek Dox broth containing monocrotophos and flask was kept on a rotary shaker at 120 rpm, at room temperature. Following this it was incubated in rotary shaker for about 7 d, 10 fold dilutions of cultures were prepared and 100 µl of sample was spread on Czapek Dox agar medium containing 100 mg/l of monocrotophos. Isolated fungal strains were maintained on agar slopes of the same medium containing monocrotophos.

2.4. Gradient plate:

The enrichment experiment which resulted in the three isolates was then furthered screened for monocrotophos tolerance capacity following gradient plate method. The monocrotophos concentration gradient was prepared by adding base layer of 20 ml of Czapek Dox agar without pesticide to the Petri plate tilted at 30° angle. The agar was allowed to solidify at room temperature and the other half of the Petri plate was subjected to the addition of 20 ml of same media containing monocrotophos (1,000 mg/l) to give monocrotophos gradient across the surface of the plate. Then, the isolated fungal strains were picked using a sterile cotton swab and were inoculated at the middle of the gradient plate agar. The inoculated plates were incubated at 30 ± 2 °C for 8 d. After which the length of the fungal growth along the gradient was recorded ⁷.

2.5. Minimum inhibitory concentration:

Minimum Inhibitory Concentration (MIC) and tolerance to monocrotophos were determined for the efficient fungal strain which was screened from gradient plate technique using broth assay. Erlenmeyer flasks (250 ml) containing 100 ml of M1 medium composed (g/l) of NaNO₃ 2; KCl 0.5; MgSO₄.7H₂O 0.5; glucose 10; FeCl₃ 10 mg; BaCl₂ 0.2; and CaCl₂ 0.5; per litre at pH 6.8 were taken with increasing concentration of monocrotophos. The flasks were then inoculated with fungal spore suspension and incubated at 30 ± 2 °C on a rotary shaker at 120 rpm. Mycelial growth obtained in the flask after 7 d of incubation was filtered using Whatman no.1 filter paper. The separated mycelial mass was washed with deionised water and the dry weight was checked by drying at constant weight for 80 °C in pre weighted aluminium foil cups. The MIC was noted as the concentration of monocrotophos resulting in the complete inhibition of mycelial growth in the flasks ⁷.

2.6. Growth kinetics:

In order to determine the growth pattern of fungal strain, 1 ml of spore suspension was inoculated in a series of flask containing Czapek Dox broth with and without monocrotophos (500 mg/l). The flasks were subjected at constant shaking in a rotary shaker at 120 rpm at 30 ± 2 °C. After incubation at regular time intervals, the mycelial mass from each series was separated by filtration using Whatman no.1 filter paper and washed with deionised water. Biomass determination was done by drying the fungal biomass for a constant weight at 80 °C in preweighed aluminium foil cups.

2.7. Taxonomic identification of fungal strain:

The isolated fungal strain was identified by 18S rRNA sequence analysis. The fungal genomic DNA was isolated by using AMpurE Fungal gDNA Mini kit. In this kit detergent and other non-corrosive chemicals are used to break open the cellulosic cell wall and plasma membrane to extract DNA from fungal cells. The 18S rRNA gene was amplified by polymerase chain reaction (PCR) using the universal primers CGW CGR AAN CCT TGT NAC GAS TTT TAC TN and AWG CTA CST GGT TGA TCC TSC CAG N. PCR reaction mix of 50 µl final volume contained: 50 ng sample gDNA, 100 ng forward primer, 100 ng reverse primer, 2 µl dNTP’s mixture (10 mm), 5 µl 10X Taq polymerase buffer, 3 U Taq polymerase enzyme and PCR grade water to make up the volume. Amplified PCR product was sequenced by using ABI3730xl genetic analyser (Amnion Biosciences Pvt. Ltd. Bangalore, India). The sequencing result was submitted to the Genbank National Centre for Biotechnology Information (NCBI) database.

2.8. Biodegradation of monocrotophos in mineral medium and soil:

The degradation of monocrotophos in liquid medium was carried out in Erlenmeyer flask containing 100 ml of M1 medium spiked with 500 mg/l of monocrotophos as the sole carbon source and 1 ml of fungal spore suspension of JPDA1 strain was inoculated. The flasks were incubated at 30 °C on a rotary shaker at 120 rpm and samples were taken at regular time intervals. The degradation of monocrotophos was carried out in the same soil sample from which fungal strain was isolated. Two trials were carried out: (1) Addition of pesticide (500 mg/kg), isolated fungal spore and nutrients (Carbon, Nitrogen, and Phosphorous), (2) Addition of pesticide (500 mg/kg) and isolated fungal spore without nutrients. The amount of Carbon, Nitrogen and Phosphorus were calculated using the relationship C/N/P 100:10:1. The sources of carbon, nitrogen and phosphorous were glucose, (NH₄)₂SO₄ and K₂HPO₄, respectively^8,9. The removal of monocrotophos was determined by High Performance Liquid Chromatography (HPLC).

2.9. Analytical methods:

The liquid samples from the monocrotophos degradation ﬂasks were extracted with equal volume of ethyl acetate. 10 g of soil samples were collected from each treatment trails with and without amendment of nutrients for pesticide analysis. The soil samples were extracted with 20 ml of ethyl acetate to determine the pesticide concentration by HPLC. The isocratic mobile phase composed of methanol: water (70:30, V:V), which was pumped through the column at a flow rate of 1 ml/min. Monocrotophos and its metabolite were detected at 214 nm. Infrared spectra of the monocrotophos parent compound and sample after fungal degradation were recorded at room temperature in frequency range 4000-400cm^-1 with FTIR. Spectrophotometer (8400 Shimadza, Japan with Hyper IR-1.7 Software for windows) with helium neon laser lamp as a source of IR radiations. Pressed pellet were prepared by grinding the extract samples with potassium bromide in motor with 1:100 ratio immediately analyses in the region of 4000-400cm^-1at a resolution of 4 cm^-1.

3.RESULTS AND DISCUSSION:

In the present study, a selective enrichment method was used to isolate monocrotophos degrading fungal strains from paddy field and three distinct strains were obtained. Monocrotophos gradient plate assay was applied to screen the potential strain for highest tolerance to monocrotophos and the growth performance was recorded as the length of fungal growth (in cm) across the monocrotophos gradient. Among the three isolates, JPDA1 showed growth of > 5 cm along the surface of the gradient plate which was further affirmed using broth assay. The minimum inibitory concentration of monocrotophos of JPDA1 strain was in accordance with gradient plate assay. The isolae JPDA1, showed luxuriant growth up to 500 mg/l of monocrotophos, and hence was selected for further biodegradation.

Fig.1 Phylogenetic relationship of Aspergillus sojae JPDA1 based on 18S rRNA gene nucleotide sequences.

The molecular characterization based on 18S rRNA sequence analysis was used to identify JPDA1 strain. BLAST results of the 18S rRNA gene sequence of JPDA1 strain exhibited close relationship, possessing 99% similarity to that Aspergillus sojae. Multiple sequence alignments and phylogenetic tree (Fig. 1) revealed that the strain JPDA1 cluster with Aspergillus sp. Therefore, the JPDA1 isolate was designated as Aspergillus sojae JPDA1 and the sequence result was submitted to GenBank NCBI database and accession number KF175513 was obtained.

Fig. 2 Growth performance of Aspergillus sojae JPDA1 in the presence and absence of monocrotophos (500 mg/l).

Fig. 2., depicts the growth kinetics of Aspergillus sojae JPDA1 in the presence and absence of 500 mg/l monocrotophos. A visible increase of mycelial mass was observed with time which indicated the metabolism of monocrotophos by Aspergillus sojae JPDA1. Initially, the growth was found to be suppressed in the presence of monocrotophos but after acclamatisation to monocrotophos, the fungus showed a higher growth rate. Moreover, the amount of biomass produced in the medium containing monocrotophos was much higher compared to the growth in the absence of monocrotophos. This could be due to the availability of an additional carbon source upon degradation of monocrotophos in the medium. Previously, Bhalerao and Puranik^[7] reported that Aspergillus oryzae ARIFCC 1054 showed increased growth with addition of monocrotophos in the aqueous medium during the degradation of monocrotophos.The strain JPDA1 was efficient in degrading MCP in liquid medium which was the only available carbon and energy source. Aspergillus sojae JPDA1 degraded monocrotophos in the aqueous medium to an undetectable level in 72 h (Fig. 3.).

(a) The HPLC chromatogram of monocrotophos at standard condition

(b) HPLC chromatogram of biodegradation of monocrotophos in aqueous medium by Aspergillus sojae JPDA1.

Fig. 3

The degradation pattern of monocrotophos in the soil is presented in Fig. 4. The strain JPDA1 was inoculated in soil along with 500 mg/kg of monocrotophos and nutrients, such as carbon, nitrogen, and phosphorous, after 144 h of incubation the results confirmed 100% degradation of monocrotophos. There is no appreciable difference in the soil inoculated with JPDA1strain and 500 mg/kg monocrotophos without addition of nutrients, the 100% degradation was obtained within 168 h.

Fig. 4 (a) Biodegradation of monocrotophos in soil with nutrients and soil without nutrients.

Comparison of FTIR spectrum of control (Fig., 5a) with extracted metabolites after complete degradation of monocrotophos by Aspergillus sojae JPDA1 (Fig. 5b) clearly confirmed the degradation of monocrotophos.

(a) FTIR spectrum of monocrotophos at standard condition

Fig. 5 (b) FTIR spectrum of biodegradation of monocrotophos in aqueous medium.

The infrared spectrum of monocrotophos degraded sample showed a band at 3460 cm^-1 corresponds to N-H stretch. The band was presented at 1643 cm^-1 which was the characteristics of C=C stretch. A strong reduction was observed in the band intensity at 1443 cm^-1 corresponding to N-H deformation of monocrotophos and this band have indicated the presence of N-methyl acetamide, formic acid, and inorganic phosphate as intermediate products of monocrotophos degradation⁷. Aspergillus oryzae ARIFCC 1054, exhibited the highest tolerance which was conﬁrmed by growing the isolates in broth cultures with varying monocrotophos concentrations of 100 to 1000 mg/l. Monocrotophos degradation by A. oryzae ARIFCC 1054 confirmed that there was a rapid depletion in monocrotophos concentration (around 70%) in the ﬁrst 50 h. The monocrotophos concentration reached undetectable levels (<1 mg/l) at 168 h of incubation. Rapid disappearance of monocrotophos in the initial phase did not correlate with a slow increase in biomass. This could be due to the degradation of monocrotophos by the enzymes secreted by the fungal mass growing initially on the readily available, but limited, carbon source, glucose. Zidan and Ramadan^[10]reported 75% and 50% degradation of monocrotophos within 4 days at the concentration 200 mg/l by Aspergillus and Penicillium sp., respectively. Monocrotophos is characterized by a P-O-C linkage and amide bond, and has been reported to be degraded as a sole carbon or phosphorus source in liquid media by Pseudomonas aeruginosa sp., Clavibacter michiganense sp.⁶, Arthrobacter atrocyaneus sp., Bacillus megaterium sp. and Pseudomonas mendocina⁵. Metabolic reactions, such as N-demethylation, O-demethylation, hydroxylation of N-methyl groups and cleavage of the phosphate-crotanamide linkage, occur during the metabolism of monocrotophos by microbial cultures and in soils ^5,11,12, with the formation of O-desmethylmonocrotophos monomethyl phosphate, dimethyl phosphate, N-methylacetoacetamide and N-methylbutyramide. Although aerobic degradation is more effective for the microbial removal of hydrocarbons, completely aerobic conditions are hard to implement in the ﬁeld, as an uneven distribution of water ﬂow, nutrients and microbial populations creates a dynamic spectrum of aerobic and anaerobic conditions¹³. In our study, Aspergillus sojae JPDA1 utilized monocrotophos even without addition of nutrients in the soil and its degrading ability was positively influenced by the presence and absence of nutrient sources. This might be due to secretion of monocrotophos degrading enzymes present in Aspergillus sojae JPDA1 which were expressed even in the absence of readily available nutrient sources.

4. CONCLUSIONS:

In this work, monocrotophos degrading fungal strain Aspergillus sojae JPDA1 with the unique potential to degrade monocrotophos in liquid medium and soil was isolated. It is worthy to mention that the JPDA1 strain had the ability to degrade monocrotophos in soil without addition of nutrient after 168 h of incubation. The results of monocrotophos degradation in soil confirms that Aspergillus sojae JPDA1 could be used as a potential candidate for the removal of monocrotophos from contaminated soils.

5. ACKNOWLEDGMENT:

Authors acknowledge the support from DST (Department of Science and Technology, Govt. of India, New Delhi) research grant, sanction no. DST/TSG/NTS/2009/67.

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Received on 02.08.2016 Modified on 21.08.2016

Research J. Pharm. and Tech 2016; 9(12):2155-2160.

DOI: 10.5958/0974-360X.2016.00437.6