Medires Publishers - Article

Abstract

The increasing use of herbicides has become worrisome as there have been reports on the harmful effects of herbicides on non-target species. This study monitored the fate and effects of three herbicides in pristine soil from Joseph Sarwuan Tarka University, Makurdi using GC-MS and laboratory culturing. Brevibacterium spp., Pseudomonas stutzeri, Bacillus flexus, Staphylococcus succinus, Paracoccus kawasakiensis, and Flavobacterium succinicans were found to degrade ButaForce yielding (1) 2,6-Diethylaniline; (2) 1 – tetradecene, (E); (3) 2-Sec-butyl-6-ethylaniline; (4) 2-Chloro-N-(2,6-diethylphenyl)-acetamide; (5) 1 – octadecene, alachlor and (6) 3 – Eicosene. On the other hand, PropaForce Plus was degraded by Pseudomonas viridiflava, Bacillus cereus, Flavobacterium columnare., and Staphylococcus saprophyticus producing (1) 4 – dimethylcumene; (2) 2,4 - Dichlorophenoxy methyl acetate and (3) Acetic acid (2,4-dichlorophenoxy)-2-ethylhexyl ester whereas force Up was degraded by Pseudomonas carboxydohydrogena, Bacillus flexus, Flavobacterium spp., and Phenilobacterium spp. yielding (1) 1 – Docosene; (2) 2-Hydroxy-1-(hydroxymethyl)-ethylhexyl ester and (3) 1,2,5-Oxadiazol-3-amine. Both Acinetobacter sp. and Lactobacillus sp. were inhibited by all three herbicides. There is need for moderation in the use of the current available herbicides while further research is advocated towards producing more ecofriendly herbicides.

Introduction

Herbicides are agents usually in the form of chemicals that are used to kill or inhibit weeds on agricultural lands, residential areas or walk ways. Technically, they are small molecular weight compounds used to kill weeds (Murray, 2019). According to Miller (2004), about 75% of all herbicides produced in the world is used in the developed countries, but the use of pesticides is increasing in developing countries too. China, the United States of America, and Argentina top as countries with the highest pesticide use globally. The use of herbicides to control weeds has increased tremendously in Nigeria in recent years. This has been linked to ease of application and effectiveness in weeds control in addition to the growing difficulty in hiring manual labour to carry out the traditional manual weeds removal (Best – Ordinioha, et al., 2017). The use of herbicides is also considered more profitable and less labour intensive compared to hand weeding (Anhwange, et al., 2013).

Nevertheless, the agrochemicals adversely affect soil biodiversity, agricultural sustainability and food safety, bringing in long term harmful effects on nutritional security, human and animal health. The irrational use of herbicides has led to the contamination of soil, water and air (Tahir et al., 2011). This impacts negatively on microbial functions and biochemical processes, affecting crops development and growth either by reducing nutrients availability or by increasing disease incidence. Several studies have reported the impact of numerous herbicides on subduing soil enzyme activities which affects the nutrient status of the soil and include hydrolases, nitrate reductase, urease, oxidoreductases, nitrogenase and dehydrogenase activities (Ram et al., 2020). Furthermore, the biological fixation of nitrogen with its biotransformation processes of nitrification, ammonification, denitrification, phosphorus solubilization and Sulphur -  oxidation is also affected by the application of herbicides (Monkiedge and Piteller, 2005).

Species of actinomycetes, bacteria, fungi, insects, nematodes, enchytraeids and other non – target groups of organisms that play vital roles in soil have been shown to be negatively affected by herbicides. Emurotu and Anyanwu (2016) reported that actinomycetes, Proteus sp., Staphylococcus aureus, and Leuconostoc sp., were eliminated by herbicides in a study carried out in Nigeria. Rhizopus sp., a fungus was also eliminated in the same study. In USA, Newman et al., (2016) report that the herbicide glyphosate caused a decline in populations of Acidobacteria which is important in the biogeochemical process, like the degradation of cellulose. Similarly, Zaller et al., (2018) found a 53% decrease in grapevine mycorrhization due to herbicide application in a study carried out in Austria.

The elimination of some microbes disrupts the ecological balance. This is dangerous as some microbes that hitherto were considered inconsequential might be favoured by certain herbicides and proliferate to dangerous levels. This is noteworthy because it might present some challenges that health workers, environmentalists and the entire human race might not be prepared for. Thus, it is important to monitor the fate of herbicides in the soil and their effects on the soil quality or health by in-depth studies on soil microbial activity (Chowdhurry, et al., 2008). It is against this backdrop that this study is necessitated.

Materials and Methods

Study Area

Study Area The study was carried out in the months of June, 2021 to January, 2022 using samples collected from the Joseph Sarwuan Tarka University Makurdi Research farm. The research farm is located in “North Core” section of the University. The soil is predominantly sandy loam in texture (Ibrahim and Idoga, 2015). Makurdi is one of the 23 Local Government Areas of Benue State and serves as the State Capital. It lies within latitude 7⁰301 to 8⁰001 N and longitude 8⁰301 to 9⁰001 E.

Makurdi is an agrarian city with two major seasons: rainy season (April to October) and dry season (November to March). Major crops cultivated include maize, tomatoes, cassava, potatoes, beniseed, melon, rice, among others. As a result of this, there is heavy dependence on herbicides to control weeds. Major herbicides used include glyphosate (Round up, Force up, bushfire), atrazine, butachlor (Buta Force), 2,4 – D, among others.

Sample Collection

Collection of Soil Sample.

Ten soil samples of 100 g were collected from a pristine soil site (7.808156, 8.618789) from a depth of 0 – 15 cm as described by Egbe et al., (2020) using a sterile soil auger and homogenized to produce a composite sample. The samples were packaged in sterile nylon bags as described by Tudararo – Aherobo and Ataikiru, (2020) and conveyed to Microbiology laboratory, Joseph Sarwuan Tarka University, Makurdi. The soil samples were air-dried at room temperature for two days, pulverized and sieved using a 2 mm mesh as described by Wibaya (2013) and Bulu et al. (2019).

Herbicides

A total of three herbicides were used in this study: PropaForce Plus (Active ingredients: 2,4 –D and Propanil) a selective Post – emergent herbicide; Force up (Active ingredient: Glyphosate), a non – selective systemic post-emergent herbicide and Butaforce (Active ingredient: Butachlor) a pre-emergent herbicide. They were bought with local suppliers in Makurdi. The application was according to the company’s recommended field rate, and twice the field rate, all in duplicates. A control was equally set up and no herbicide was added to it.

Equipment and Materials

Media

Nutrient agar was used for the cultivation of heterotrophic bacteria. It is a general purpose nutrient medium used for the cultivation of microbes and supports a wide range of microbes. It was prepared according to the manufacturers’ instruction of 28 g per 1000 ml.

Other Materials

Glass wares including measuring cylinder, beakers, test tubes, conical flasks were used. Other materials included autoclave, incubator, Gas chromatography – Mass Spectrometer, PCR machine, weighing balance, Petri dishes, microscope, wire loop, syringes, McCartney bottles. Other reagents are specified under the headings below.

Sterilization of Materials

Before the commencement of the work all materials were sterilized as adopted from Sebiomo et al. (2011). All glass wares were properly washed with detergent mixed with a solution of sodium hypochlorite, rinsed in clean water and sterilized in hot air oven at 170 ⁰C for 60 minutes. All media were sterilized by autoclaving at 121 ⁰C at 15 Psi. The work bench was disinfected with the aid of a solution of sodium hypochlorite.

Determination of Organic Matter Content of the Soil

The organic matter content of the soil was determined following the method described by Sebiomo et al. (2011). Soil samples collected were ground to pass through 0.5 mm sieve. One gram of each soil sample was weighed into 250 ml Erlenmeyer flasks and 10 ml of K2Cr2O7 solution was dissolved into each flask and swirled gently to disperse soil. Twenty milliliters of concentrated H2SO4 was rapidly added using automatic pipette and swirled gently until the soil and reagents were mixed, then the mixture was swirled more vigorously for one minute, the flasks were then rotated and allowed to stand in a sheet of asbestos for about 30 min. One hundred milliliters of distilled water was added to each flask, then 3 - 4 drops of indicator (ferroin) was added and filterated with 0.5 N ferrous sulphate solution to the end point, from greenish or dark green to red (maroon colour) and in reflected light against a white background. The organic matter was calculated according to the following formula: 

%Organic matter =

Where

 Correlation factor “ƒ” = 1.33

 me = normality of solution × milliliter of solution used and 1.729 = conversion. 

Determination of Soil Physicochemical Parameters

The other physicochemical parameters of the soil were determined as described by Tudararo-Aherobo and Ataikiru (2020) with slight modifications.

Determination of pH

Ten (10) gram of each of the soil sample was weighed into 50 ml beakers and 25 ml distilled deionized water was added to form 1:2.5 soil/water mixtures. The mixture was stirred for 30 minutes and allowed to stand for about 5 minutes. Two point calibrations were made with buffer solution having pH of 7 and 4 (buffer 7 and buffer 4). Finally, the pH meter electrodes (JENWA 3510) were immersed into the soil/water mixture and the pH was measured on the upper part of the suspension.

Determination of Organic Carbon

One gram of soil was weighed and transferred to 250ml Erlenmeyer flask and 10 ml of 1N K 2Cr2O7 solution was added into flask and swirled gently to disperse the soil. Then 20 ml concentrated H2SO4 was added rapidly using an automatic pipette. This was swirled vigorously for one minute and allowed to stand on a sheet of asbestos for about 30 minutes.  Then, 100ml of distilled water was added and left to stand for 30 minutes. Finally, 3-4 drops of O - phenanthroline indicator were added and titrated with 0.5N ferrous sulphate solution. At endpoint, ferrous sulphate was added drop by drop until the colour changed sharply from blue to red (maroon color in reflected light against a white background). The blank titration was made in the same manner but without soil to standardize the dichromate.

% Organic carbon was calculated using the formula below

% Organic Carbon =N(V1 - V2) X0.3F W

Where

N = Normality of ferrous sulphate solution

V1 = ml ferrous ammonium sulphate required for the blank

V2 = ml ferrous ammonium sulphate required for the sample

W = mass of sample in gram

F = correction factor = 1.33

Determination of Nitrate

Fifty ml of 2.5% acetic acid were used to shake and extract the nitrate from the soil samples. The solution was filtered into a clean beaker and 25 ml of Brucine reagent and 2 ml concentrated sulphuric acid were added. A yellow coloured solution was formed and read at 470 nm in the spectrophotometer (Hitachi 220 spectrophotometer) using water as blank. The standard concentration of nitrate in the sample was extrapolated from a standard nitrate graph.

Determination of phosphate

The available phosphate in the soil was extracted with Olsen’s extracting solution and analyzed using the ascorbic acid- molybdenum on a spectrophotometer. A 5 ml soil sample extract and 0.8 ml of the combined agent was measured into a clean beaker, and then left for 10 minutes. The bluish solution formed was read at 888 nm using a spectrophotometer (Hitachi 220 spectrophotometer) and distilled water as a blank. Potassium hydrogen orthophosphate and the combined reagent were used in preparing the standard phosphate concentration. This was absorbed at same wavelength. The concentration of phosphate in the sample was extrapolated from a standard graph of phosphate.

Design of Soil Microcosm Herbicide Exposure Experiment.

Determination of residual herbicides and their metabolites in Soil Microcosms was done on the first and last day of the study using GC – MS as adopted from Al - Ani et al. (2019) with little modifications. The air-dried, pre-stabilized soil was thoroughly mixed to homogenize. Thereafter, 35 g of it was transferred to individual sterile bottles. The soil microcosms shall then be exposed to the different herbicides at recommended field rate and twice the recommended field rate. The control had no herbicide treatment. The soil water content was adjusted to about 60% of the maximum water holding capacity for the soils using sterile water. The bottles were incubated at 30 ⁰C in a dark chamber. Sterile water was added every two days to maintain the water content of the soils. Samples were taken from these soil microcosms every seven days for bacterial analysis.

Determination of Residual Herbicides and Herbicides Metabolites in Soil Microcosms 

Determination of Residual Herbicides and herbicides metabolites in Soil Microcosms was done on the first and last day of the study using GC – MS as adopted from Egbe et al. (2020) with little modifications. Ten grams of soil samples already contaminated with herbicides were placed in a 250 ml beaker. Extraction was done using 10 ml n – hexane:dichloromethane (1:1) by shaking and allowing to stand for about 20 minutes. The organic phase was decanted and filtered through a glass funnel fitted with a filter paper. Sodium sulphate was added on the filter paper to absorb any moisture and the organic extract was concentrated to 1 µl by evaporating at room temperature. The concentrated extract was transferred to a GC vial for analysis. One µl of the extracted sample was injected into the GC column for analysis. The GC (Agilent 6890N) and MS (5975B MSD) was equipped with DB-5ms capillary column (30 m×0.25 mm; film thickness 0.25 µm). The initial temperature was set at 40 °C which increased to 150°C at the rate of 10 °C/min. The temperature was again increased to 230 °C at the rate of 5 °C/min. The process continued till the temperature reached 280 °C at the rate of 20 °C/min which was held for 8 minutes.

The injector port temperature remained constant at 280 °C and detector temperature was 250 °C. Helium was used as the carrier gas with a flow rate of 1 mL/min. Split ratio and ionization voltage were 110:1 and 70 eV respectively. To identify and quantify the target herbicides present in the extracted sample, their individual mass spectral peak values were compared with the database of National Institute of Science and Technology (2014); followed by obtaining the percent report from the equipment. The percent report showed the exact amount at which the targeted herbicides congeners were present.

Determination of Residual Herbicides and Herbicides Metabolites

The soil residual herbicide and herbicide metabolites in soil microcosms were determined using GC – MS was adopted from Egbe et al. (2020) with little modifications. Ten grams of soil sample already contaminated with herbicides were placed in a 250 ml beaker. Extraction was done using 10 ml n – hexane:dichloromethane (1:1) by shaking and allowing to stand for about 20 minutes. The organic phase was decanted and filtered through a glass funnel fitted with a filter paper. Sodium sulphate was added on the filter paper to absorb any moisture and the organic extract was concentrated to 1 µl by evaporating at room temperature. The concentrated extract was transferred to a GC vial for analysis.

Serial Dilution and Determination of Bacterial Communities

Nutrient agar was used to determine the presence of heterotrophic bacteria by pour plate method using dilution 107 as adopted Nkamigbo et al. (2020). The nutrient agar was incubated at 30⁰C for 24 - 48 hours following pour plating. Bacterial identification was by cultural characteristics, staining reactions and biochemical reactions.

Cultural and Biochemical Identification of Bacteria

Cultural identification of bacteria was done using colonial morphology such as shape, colour, translucency, elevation, edge, and surface texture followed by Gram staining and microscopic examination. The isolates were identified with reference to Bergey’s Manual of Determinative Bacteriology.

Gram Staining

Gram staining of the isolates was done as described by Ochei & Kolhatkar, (2000). A smear was made, allowed to dry and heat fixed by passing two to three times over Bunsen flame. Crystal violet was used to stain the smear for one minute and then it was washed with running water. Lugol’s iodine was applied for one minute and then washed with running water. Acetone or alcohol was used to decolourize the crystal violet (about 2 seconds for acetone, and 1 – 2 minutes for alcohol), and it was washed with running water. Safranin was used to counter stain the smear for one minute then it was washed with running water. A filter paper was used to blot the underside of the smear. The smear was allowed to air dry after which a drop of oil immersion was added to the stained slide before viewing under X 100 objective.

Biochemical Tests

The biochemical tests were carried out using the protocol described by Ochei & Kolhatkar, (2000). The tests carried out were: indole, coagulase, oxidase, citrate utilization test, urease and catalase tests. 

Molecular Identification

Molecular identification of the bacterial isolates was carried out at Inqaba Biotec Laboratory, Ibadan, Oyo State and the protocols were adopted from Illumina, (2015) and Meyer et al. (2008).

Molecular Identification: Adopted from Illumina, (2015) and Meyer et al. (2008)

Genomic DNA Isolation from Soil

After sample collection, DNA was isolated directly from the soil sample using the Meta-G-Nome DNA Isolation Kit (Epicentre®, an Illumina company). The protocol uses filtration technology and enzymatic lysis to isolate DNA from the soil sample. The kit is designed to isolate randomly sheared genomic DNA of high molecular weight.

Library Preparation

The Nextera XT DNA Library Prep Kit was used to construct libraries from the isolated DNA. Using a single enzymatic “tagmentation” reaction, the Nextera transposome simultaneously fragmented and tagged the DNA with unique adapter sequences. Limited-cycle PCR was used to amplify the tagged DNA and add sequencing indexes. Using this streamlined workflow, 11 DNA libraries were prepared for sequencing on the NextSeq 500 desktop sequencer in half a day.

Sequencing

All 11 libraries were pooled together for cluster generation and sequencing. Libraries were loaded onto a reagent cartridge and clustered on the NextSeq 500 System. A paired-end, 2 × 150 bp sequencing run was performed using the NextSeq 500 High-Output Kit. A single sequencing run generated 400 million reads in 29 hours, corresponding to an average of 40 million reads per sample after quality filtering. Base calls generated by the NextSeq 500 System were converted to FASTQ files for metagenomic analysis.

Data Analysis

This was done using the publicly available MG-RAST program, an automated analysis platform that provides various tools for data visualization enabling users to assess species composition or functional abundance. FASTQ files generated by the NextSeq 500 System and sample metadata were uploaded to the MG-RAST server. Overlapping paired-end reads were joined and then processed in MG-RAST using the default trimming and filtering settings to allow comparison to other data sets in MG-RAST.

RESULTS

Studies on the production of metabolites from bacterial degradation of some selected herbicides in pristine soils of Makurdi, Benue State has been undertaken. The herbicides used were ButaForce (butachlor), PropaForce Plus (propanil and 2, 4 – D) and Force Up (glyphosate). Bacterial isolates with potential to utilize the herbicides as sole carbon sources for growth were isolated. For ButaForce, six (6) isolates which include Brevibacterium aureum, Pseudomonas stutzeri, Bacillus cereus, Staphylococcus sciuri, Paracoccus kawasakiensis, and Flavobacterium succinicans were obtained whereas for PropaForce Plus, Pseudomonas viridiflava, Bacillus cereus, Flavobacterium columnare., and Staphylococcus saprophyticus and for Force Up, Pseudomonas stutzeri, Bacillus flexus, Flavobacterium spp., and Phenilobacterium spp. were isolated after contamination of the soil with the herbicides (Table 1).

Table 2 shows the physicochemical parameters of the soil including pH, percentage nitrate, organic carbon, organic matter, nitrogen, phosphorous and cation exchangeable capacity. The organic matter percentage was 1.120, organic carbon 0.650 and cation exchangeable capacity of 7.050%.

Table 4 shows the degradates of bacterial metabolism of ButaForce, Force Up and PopaForce Plus in Pristine soil after 14 days of incubation. The metabolites produced by ButaForce degredation were: (1) 2,6-Diethylaniline; (2) 1 – tetradecene, (E); (3) 2-Sec-butyl-6-ethylaniline; (4) 2-Chloro-N-(2,6-diethylphenyl)-acetamide; (5) 1 – octadecene, alachlor and (6) 3 – Eicosene; the metabolites produced by for Force Up utilization were (1) 1 – Docosene; (2) 2-Hydroxy-1-(hydroxymethyl)-ethylhexyl ester and (3) 1,2,5-Oxadiazol-3-amine while the degradation of Propa Force Plus yielded (1) 4 – dimethylcumene; (2) 2,4 - Dichlorophenoxy methyl acetate and (3) Acetic acid (2,4-dichlorophenoxy)-2-ethylhexyl ester.

In all the results, none of the 3 herbicides was completely mineralized after 14 days of incubation as the indigenous bacteria were only able to transform the herbicides into less harmful forms.

Degradation and biotransformation of the selected herbicides by bacteria was observed as presented in Table 5. The percentage of the active ingredients remaining in the soil after bacterial activity was less than the initial percentage concentration of the herbicides. The rate of degradation of Buta Force spiked soil was higher (25.024%), followed by PropaForce spiked soil (9.739%). The least observed herbicide degraded by bacteria was in Force Up spiked soil (0.09%).

Table 6 shows the frequency distribution of genera of bacteria isolated from pristine soil. A total of 42

bacteria were isolated with Pseudomonas spp. as the dominant bacteria followed by Flavobacterium spp.

Table 1: Isolated Herbicide Utilizers

Table 4: Metabolites from bacterial Degradation of ButaForce, PropaForce Plus and Force Up in Pristine soil at Day 14

Table 5: Screening of the Active Ingredients of ButaForce, Force Up and PropaForce Plus in soil before and after Bacterial Action

Key:

NBF Before: ButaForce in Pristine Soil before bacterial action.

NBF After: ButaForce in Pristine Soil after bacterial action.

NFU Before: Force Up in Pristine Soil before bacterial action.

NFU After: Force Up in Pristine Soil after bacterial action.

NPFP Before: PropaForce Plus in Pristine Soil before bacterial action.

PFP After: PropaForce Plus in Pristine Soil after bacterial action.

Table 6: Frequency of Bacterial Genera Isolated from Pristine soil

Figure 1: shows the percentage rate of herbicide degradation. ButaForce degradation had the highest percentage rate (43%) of degradation as compared to PropaForce Plus (29%), while the lowest is Force Up (28%). This implies that ButaForce was more resisted by the bacterial community pristine soil, compared to either PropaForce Plus or Force Up.

Figure 2 shows the relative abundance of the bacterial genera isolated from pristine soil. Pseudomonas had the highest species abundance (19), followed by Flavobacterium (9), Staphylococcus (4), Bacillus (3), Paracoccus (2) while Lactobacillus, Acinetobacter, Phenilobacterium and Brevibacterium were each represented by one species. There was a significant difference between the relative abundance of Pseudomonas spp. and Flavobacterium spp. and the other genera, while no significant difference was observed between the rest of the genera isolated.

In figure 6, the chromatogram of the degradation of PropaForce Plus in Pristine Soil at Day 14 is shown. The identified metabolites were: (1) 4 – dimethylcumene; (2) 2,4 - Dichlorophenoxy methyl acetate and (3) Acetic acid (2,4-dichlorophenoxy)-2-ethylhexyl ester.

Fig 1: Percentage of Bacterial Herbicide Degraders

Fig 6: Chromatogram of degradation of PropaForce Plus in Pristine Soil at Day 14

7.328: 4 – dimethylcumene. 9.291: 2,4 - Dichlorophenoxy methyl acetate. 11.752: Acetic acid (2,4-dichlorophenoxy)-2-ethylhexyl ester.

The chromatogram for Bacterial Degradation of Buta Force in Pristine Soil at Day 14 is shown in Figure 7. The degredates were: (1) 2,6-Diethylaniline; (2) 1 –

tetradecene, (E); (3) 2-Sec-butyl-6-ethylaniline; (4) 2-Chloro-N-(2,6-diethylphenyl)-acetamide; (5) 1 – octadecene, alachlor and (6) 3 – Eicosene.

Figure 8 is the chromatogram for Bacterial Degradation of Force Up in Pristine Soil on Day 14. The identified compounds were: (1) 1 – Docosene; (2) 2-Hydroxy-1-(hydroxymethyl)-ethylhexyl ester and (3) 1,2,5-Oxadiazol-3-amine.

The result of the metagenomics for the top class classification for bacterial genomic sequences obtained from pristine soil presented in Figure 11 shows that Betaproteobacteria dominated with 42.28%, followed by alphaproteobacteria (16.45%) and gammaproteobacteria (7.04%) while 17.57% represented unidentified bacterial classes. The metagenomics analysis of 16s gene amplicons on the Sequel system using PacBio generated 6,140 Operational Taxonomic Units (OTUs) falling into 29 phyla were identified from the classifiable sequences. Betaproteobacteria (42.28%), alphaproteobacteria (16.45%), gammaproteobacteria (7.04%), actinobacteria (4.63%) and planctomycetia (4.38%) predominated while coriobacteria, fibrobacteria and bacteroidia accounted for 0.02, 0.03 and 0.03% of the total identifiable bacterial community.

Fig 7: Chromatogram for Bacterial Degradation of Buta Force in Pristine Soil on Day 14

2,6-Diethylaniline, 1 – tetradecene, (E), 2-Sec-butyl-6-ethylaniline, 2-Chloro-N-(2,6-diethylphenyl)-acetamide, 1 – octadecene, alachlor and 3 – Eicosene.

Fig 8: Chromatogram for Bacterial Degradation of Force Up in Pristine Soil on Day 14

11.620: 1 – Docosene. 14.269: 2-Hydroxy-1-(hydroxymethyl)-ethylhexyl ester. 18.807: 1,2,5-Oxadiazol-3-amine.

Discussion and Conclusion

Discussion

The surge in global population, as well as weed menace has necessitated the use of herbicides to control weeds in order to boost food production. The herbicides applied on farmlands to control weeds eventually get into the soil, hence its impact on beneficial non-target organisms (which maintain soil health), its fate in the soil and its impact on other components of the ecosystem needs to be ascertained. It is therefore imperative to detoxify contaminated soils in order to restore soil health, productivity and sustained ecological system. Bacteria have been widely applied in the remediation of contaminated environments. Several genera of bacteria abound in the soil, but not all are culturable using available media.

The present study monitored the production of metabolites from bacterial degredation of three selected herbicides: Butaforce (butachlor), Force Up (glyphosate) and PropaForce Plus (propanil and 2,4 – D) in pristine soil and their effects on bacterial community structure using laboratory culturing and metagenomics, and phylogenetic analysis was conducted based on the results. Furthermore, bacterial degradation of the herbicides was also monitored by Gas Chromatography – Mass Spectroscopy.

The heterotrophic bacteria isolated in our study were predominantly Pseudomonas spp., followed by Flavobacterium spp. and Staphylococcus spp. Seven genera of bacteria were isolated from the soil contaminated with the different herbicides and identified as herbicide utilizers. The various species were: Brevibacterium aureum, Pseudomonas stutzeri, Bacillus cereus, Staphylococcus sciuri, Paracoccus kawasakiensis, and Flavobacterium succinicans. These could utilize ButaForce, while Pseudomonas stutzeri, Bacillus flexus, Flavobacterium spp., and Phenilobacterium spp., could utilize Force Up and PropaForce Plus was utilized by Pseudomonas viridiflava, Bacillus cereus, Flavobacterium columnare., and Staphylococcus saprophyticus. It was observed that the genera Pseudomonas, Flavobacterium and Bacillus could utilize all the three herbicides, while Staphylococcus could utilize only ButaForce and PropaForce Plus. The genera Brevibacterium and Phenilobacterium could only utilize ButaForce and Force Up respectively. Two genera of bacteria were however, inhibited by all the three herbicides which were Lactobacillus and Acinetobacter. Force Up and PropaForce Plus inhibited Brevibacterium spp., and Paracoccus spp. while Phenilobacterium sp. could only tolerate Force Up. The inhibited bacteria can serve as bacterial bio indicators of soil contamination with the various herbicide causing the inhibition, while the herbicide utilizers can serve in bioremediation of soils contaminated with the herbicides.

These bacteria have been reported as important biofertilizers for crop cultivation, improving the quality of crop products, securing soil health (Devi and Sumathy, 2018). Pseudomonas spp. have been generally reported as nutritionally versatile (OECD, 1997), able to utilize a wide array of substrates for growth hence their predominant isolation from environmental samples.

The results agree with earlier works reported by Sebiomo and Banjo (2020) and Emurotu and Anyanwu (2016) who found that Pseudomonas spp. and Bacillus spp. occurred in all herbicides treated soils in a study carried out in Ibadan using atrazine, Xtravest, gramoxone and glyphosate. This research also consolidates the results Emurotu and Anyanwu (2016) who reported that Pseudomonas spp., Bacillus spp. and Flavobacterium spp. were the most frequently isolated bacteria from herbicide treated soils. The herbicides in their study were atrazine and butachlor. Similarly, Singh et al. (2020) also reported the utilization of glyphosate by Pseudomonas sp., Bacillus sp., and Flavobacterium sp. as phosphorus and carbon sources. In addition, Singh et al. (2018) also reported the degradation of butachlor by Pseudomonas spp. in 30 hours. Li et al. (2019) further consolidate our findings when they reported the degradation of butachlor by Paracoccus spp. in single – chamber microbial fuel cells.

The physicochemical properties of a soil indicate its nutrient status and based on the chemistry of the herbicides, some are more solubilized at higher pH, making them available for plant intake (Bulu et al., 2019) and microbial degradation. The low acidity of the soil implies that less herbicides will be sorbed onto soil particles as herbicide adsorption negatively correlates with pH (Rigi et al., 2015). This means that less quantity of herbicides will be required to control weeds, posing less threat to the ecosystem. The nitrogen content of the soil was low (< 0.15%) based on the rating of Singh (2002). This might be as a result of the decline in the activities of nitrogen fixing bacteria in the soil. This is in harmony with the work of Bulu et al. (2019) who reported low to medium values of nitrogen in their studies.

Based on the rating of Landon (1991), the organic carbon content of the soil was very low. Low organic contents were also reported by Bulu et al. (2019). The low organic carbon content could be due to the shallow root system of the crops / weeds growing in the soil. Soil organic matter is reported to improve soil quality by reducing the negative effects of pesticides, providing carbon and energy for soil microbes, stabilize and hold soil particles together and by storing and supplying plant nutrients as well as increasing the CEC of the soil (Cooperband and Wisconsin, 2002). This leads to an increase in microbial growth leading to disease suppression and pollution remediation. Organic matter content of a soil enhances the sorbing of herbicides and hence, their persistence in the soil (Thukur, 2018). Fertility, water availability, susceptibility to erosion, soil compaction and even resistance to insects and disease all depend on soil organic matter (Cooperband and Wisconsin, 2002).  However, the quantity of herbicides needed to effectively control weeds in a soil with higher organic matter will be increased, leading to environmental pollution.

The soil samples analyzed using GC-MS for bacterial degradation of ButaForce (Butachlor) in pristine soil yielded 2,6-Diethylaniline, 1 – tetradecene, (E), 2-Sec-butyl-6-ethylaniline, 2-Chloro-N-(2,6-diethylphenyl) -acetamide, 1 – octadecene, alachlor and 3 – Eicosene. These compounds were identified by comparing their mass spectral values with the database of National Institute of Science and Technology (2014) library. The results consolidated previous works reported in literature. Singh et al. (2018) identified three residues of butachlor degradation by Ammoniphilus sp. JF isolate from agricultural fields of Punjab, India. These included acetamide, 2-chloro-N, N-diethyl, N-hydroxymethyl-2-chloro-N-(2,6-diethyl-phenuyl)-acetamide and N-(2,6-diethyl-phenyl) –N- hydroxyl-methy-acetamide. Also, 1,2-benzenedicarboxylic acid, bis (2-methylpropyl) ester and 2,4-bis (1,1-dimethylethyl)-phenol. Similarly, Olukanni et al. (2020) identified 1 – docosene, 1 – octadecene and 3 – eicosene as metabolites of the degradation of butaforce (butachlor) by Aspergillus niger.

Eicosene and 2-tetradecene have been reportedly suspected to possess anticancer, antimicrobial and antioxidant ability (Tiloke et al., 2018). Similarly, 1-octadecene has been reported to exhibit both anticancer and antimicrobial activity (Renukadevi et al., 2011). Furthermore, there are reports on significant antimicrobial activity of 3-eicosene, (E)- and 5-eicosene, (E) against pathogenic Escherichia coli, Aspergillus flavus and A. niger (Lulamba et al., 2021), Penicillium chrysogenum and Alternaria alternata (Talie et al., 2020). While the inhibition of pathogens such as Aspergillus, Alternaria and E. coli is plausible as it will ensure the protection of crops, animals and humans from infectious diseases, the inhibitory activity of 3-eicosene, (E)- and 5-eicosene, (E) on P. chrysogenum is much regretted as it promotes crop growth and induces defence related genes and downy mildew disease resistance in pearl millet (Murali, et al., 2013).

Seven genera of bacteria were isolated and identified of which 6 (43%) could degrade Buta Force, 4 (28%) Force Up and 4 (29%) PropaForce Plus. This agrees with the results of Singh, et al. (2020) who reported the utilization of glyphosate by Pseudomonas sp. for growth. Also, Emuroru & Anyanwu (2016) comparing bacterial degradation of atrazine and butachlor reported that butachlor is more readily degraded by bacteria as found in our study. In their study however, Staphylococcus spp. could not degrade butachlor contrary to our findings.

Herbicides have harmful effects on non-target organisms in the soil such as bacteria. Our study found that of the 9 bacterial genera isolated and identified, Force Up inhibited 5, PropaForce Plus 6 and Buta Force 3. This shows that of the 3 herbicides studied, Butaforce has the least toxicological impact on soil bacteria. This agrees with Emuroru & Anyanwu (2016) where butachlor has less toxicological effect compared to atrazine, and Sebiomo, et al. (2011) where glyphosate was reported to have nearly similar toxicity on bacteria as atrazine.

All the herbicides in our study showed various levels of bacterial degradation under the study conditions with Buta Force and PropaForce Plus with the highest and least degradability respectively. All the herbicides showed varying levels of biotransformation as none of the herbicides was completely mineralized by the indigenous bacteria within the period under study.

Conclusion

The fate and effects of three selected herbicides studied on both paddy and pristine soil had a significant inhibitory effect on the bacterial community structure, although some bacteria were able to resist and degrade the herbicides. This corroborates previous research findings which reported on the metabolic versatility of bacteria in the removal of xenobiotics from the environment. Even though the indigenous bacterial community in paddy and pristine soils could not completely mineralize the herbicides, they successfully transformed all of them into less harmful compounds. As seen from the cultural and molecular results using metagenomics confirmed the bacterial isolates obtained by conventional laboratory culturing. This study also showed a significant difference of cation exchangeable capacity, nitrogen, organic matter, pH and organic carbon between paddy soil and pristine soils while there was no significant difference in the nitrate content of both soil types.

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