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Microbiology and Biotechnology Letters

Review(총설)

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Environmental Microbiology  |  Biodegradation and Bioremediation

Microbiol. Biotechnol. Lett. 2023; 51(4): 374-389

https://doi.org/10.48022/mbl.2308.08011

Received: September 4, 2023; Revised: November 1, 2023; Accepted: November 3, 2023

Advanced Bioremediation Strategies for Organophosphorus Compounds

Anish Kumar Sharma1* and Jyotsana Pandit2

1Department of Biotechnology, School of Sciences, P.P. Savani University, Kosamba, Surat, Gujarat 394125, India
2Department of Environmental Science, School of Sciences, P.P. Savani University, Kosamba, Surat, Gujarat 394125, India

Correspondence to :
Anish Sharma,        anish.sharma@ppsu.ac.in

Abstract

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Organophosphorus (OP) pesticides, particularly malathion, parathion, diazinon, and chlorpyrifos, are widely used in both agricultural and residential contexts. This refractory quality is shared by certain organ phosphorus insecticides, and it may have unintended consequences for certain non-target soil species. Bioremediation cleans organic and inorganic contaminants using microbes and plants. Organophosphate-hydrolyzing enzymes can transform pesticide residues into non-hazardous byproducts and are increasingly being considered viable solutions to the problem of decontamination. When coupled with system analysis, the multi-omics technique produces important data for functional validation and genetic manipulation, both of which may be used to boost the efficiency of bioremediation systems. RNA-guided nucleases and RNA-guided base editors include zinc-finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs), and clustered regularly interspaced short palindromic repeats (CRISPR), which are used to alter genes and edit genomes. The review sheds light on key knowledge gaps and suggests approaches to pesticide cleanup using a variety of microbe-assisted methods. Researches, ecologists, and decision-makers can all benefit from having a better understanding of the usefulness and application of systems biology and gene editing in bioremediation evaluations.

Keywords: Bioremediation, organophosphates, biodegradation network, genome editing

Graphical Abstract


Introduction

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Owing to the widespread pollution of water and soil from unrestrained pesticide usage, research into pesticide degradation is crucial. For decades, farmers have used pesticides to lessen the spread of disease, protect crops from damage, enhance quality, and boost output [1]. When commercial pesticides are used in excess, only a small percentage kills the pests they are designed to kill, whereas the rest contaminate the environment. Chronic and acute consequences on non-target wildlife and human life are likely to result from this hazard to the soil and water [2]. Insects, plant diseases, weeds, mollusc’s, birds, and nematodes are examples of agricultural pests that may be controlled using chemical or biological pesticides. The most popular pesticides include organochlorines, organophosphates, carbamates, pyrethroids, and triazines. The biological mechanism function or their methods of application have also led to the categorization of pesticides as contact, systemic, specific, or nonspecific. Pesticides are categorized by target and help increase crop productivity and reduce pest-related agricultural losses [3]. Humans and other animals that are not intended to be exposed to pesticides can get their hands on them in several ways [4]. Direct contact (via the skin, mouth, or lungs), as well as indirect contact through contaminated water and food sources, can lead to absorption of chemical residues into the circulation. Approximately 45 percent of the world's pesticides are organophosphates, which are extremely harmful to the soil and water ecology. Over 200,000 fatalities per year are attributed to approximately 140 different varieties of organophosphate insecticides used worldwide.

Pesticides can be detected in the environment using several different techniques, including capillary electrophoresis, gas chromatography, liquid chromatography, bioassays, and immunoassays [5−7]. The food supply, from the farm to the table, is contaminated with pesticides. Pesticide contamination has been reported in various foods, including fruit juices, milk, and seaweeds [8]. The elimination of these chemical compounds poses an enormous ecological challenge to the sustainability of life on Earth. Long-term exposure to pesticides, either orally or topically, poses a threat to human health and can result in ailments linked to metabolic, growth, and developmental issues as well as cancer in humans. Several mechanisms, such as oxidation, reduction, isomerization, conjugation, hydrolysis, hydration, dehalogenation, methylation, and cyclization, have been reported to contribute to the physical, chemical, and biological degradation of pesticides. Bioremediation is used more often to deal with pollution because of recent developments in gene editing, systems biology, and omics technology. With the help of systems biology, we can learn more about microbial communities and how they adapt to various settings, even the most hostile ones. In addition, this review discusses gene editing methods, such as CRISPR Cas, TALEN, and ZFNs, all of which have the potential to enable the creation of a design microbe that has a functional gene of interest for the degradation of recalcitrant pesticides and further enhance bioremediation.

Overview of Organophosphorus Pesticides

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In terms of market share, organophosphorus (Ops) compounds account for almost 80% of all insecticides sold worldwide. Organophosphorus (OP) pesticides (malathion, parathion, diazinon, and chlorpyrifos) are among the most extensively used pesticides in both agricultural and residential settings. Every year, the United States uses approximately 80 million pounds of organophosphate (OP) pesticides, with agriculture accounting for 75 percent of that consumption. Poisoning from organophosphorus compounds is a worldwide health concern, resulting in approximately 3 million cases of poisoning and 200,000 fatalities per year [9, 10]. The ability of OPs to inhibit acetylcholinesterase, a critical enzyme in the control of the central and peripheral nervous systems, contributes to acute toxicity. OPs interfere with neurological signalling by inactivating acetylcholinesterase (AChE) via phosphorylation and irreversible adherence to the catalytic serine, which results in an increase in acetylcholine levels and cholinergic hyperactivity [11]. Salivation, vomiting, nausea, miosis, and convulsions can occur after exposure to large doses of OPs, leading to paralysis and death [12, 13]. Low-level, long-term exposition to OPs may also be harmful, with consequences that extend to the developing human brain and the health of children [14, 15].

Agrochemicals are eventually decomposed in soil by microorganisms, and it is well documented that only a fraction of the overall quantity of agrochemicals employed goes towards the prevention and treatment of pests and diseases, and the presence of agrochemicals and their breakdown products in the environment is unavoidable. A recalcitrant chemical does not break down in the environment because microorganisms lack the mechanism for utilizing it as a carbon or energy source. Some members of organ phosphorus pesticides have this refractory property and may consequently have adverse impacts on certain non-target soil organisms [2, 16]. Soil fertility and agricultural crop productivity rely on microbial transformations such as nitrification, ammonification, and organic matter decomposition, but these processes can be inhibited, eliminated, or altered by recalcitrant pesticides and their degradation products that accumulate in the topsoil [17]. Long-term exposure to pesticide-contaminated environments results in the evolution of microbial tolerance and degradation mechanisms against a particular pesticide; furthermore, they can be employed in the breakdown of pesticides [18]. When a soil microbe uses a pesticide as a carbon and energy source, catabolism and detoxification processes occur. Finding decontamination techniques that are both effective and practical is a serious challenge in light of the severe toxicity of OPs and the extensive usage of these chemicals. OP decontamination options have considered chemical, physical, and biological methods. These treatments are severe, unsuitable for decontaminating individuals, and unsuitable for large-scale environmental remediation. It is imperative that immediate action be taken to reduce the amount of environmental damage that is being brought about by the careless application of these chemical compounds in the field of agriculture.

Emerging Bioremediation Strategies

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Bioremediation is the technique of using microorganisms and plants for environmental cleanup of both organic and inorganic xenobiotics [19]. Since bioremediation eliminates, degrades, detoxifies, and immobilizes dangerous wastes and pollutants, microorganisms are crucial. Understanding the whole range of physiological, microbiological, ecological, biochemical, and molecular processes involved in pollutant transformation is essential for successful bioremediation employing microorganisms [20]. Ex situ or in situ bioremediation can be utilized, depending on a variety of criteria, such as cost, pollutant kinds, and concentration. Bioremediation is cheaper than incineration, and certain contaminants may be handled on site, minimizing exposure hazards for clean-up workers or broader exposure from transportation accidents. Bioremediation carried out in situ, including bioventing, biosparging, and bioaugmentation, decontaminates without removing soil from the site while ex situ treatments (such as land farming, biopiling, composting, bioreactors, and electrodialysis) treat the soil that is unearthed at the location [21].

There are a number of factors that can affect how quickly pesticides break down in the soil. These factors include the amount of pesticides that are already there, the activity level of microorganisms that break down pesticides, the bioavailability of the pesticides, and a range of soil factors such as pH, soil water content, and temperature [22]. By increasing the richness of its species or modifying its enzyme systems, a microbiome can help speed up the agrochemical's metabolism and subsequent removal from the environment. Organophosphates' severe mammalian toxicity and ubiquitous and widespread use make microbial breakdown of these substances of special interest.

Microbial degradation of organophosphorus compounds

Communities of microorganisms have developed remarkable defenses against the hazards of organophosphorus chemicals. Microorganisms are able to breakdown and detoxify organophosphorus chemicals because they possess particular genes which encode enzymes which degrade these chemicals. The bacteria ability to degrade contaminants suggests interesting directions for bioremediation technologies and the recovery of polluted ecosystems. The effectiveness and environmental fate of organophosphorus chemicals like glyphosate, chlorpyrifos, malathion and parathion have been the subject of much research. Degradation of these substances primarily occurs through hydrolysis, oxidation, alkylation, and dealkylation. Degradation by microorganisms, especially hydrolysis, is an important aspect of the detoxification process.

The bacterial genus Pseudomonas appears to be particularly effective when it comes to degrading pesticides, since it is able to break down approximately 90− 99 percent of them [23]. Bacillus pumilus strain C2A1, Bacillus aryabhattai, Streptomyces olivochromogenes, Pseudomonas resinovarans AST2.2 and P. indoloxydans has been shown to effectively degrade chlorpyrifos [24−28]. Evidence suggests that microorganisms may remove chlorpyrifos from the environment, albeit the rate of removal varies by strain. Complete removal of chlorpyrifos was obtained by Stenotrophomonas sp. after 28 h of treatment, by Enterobacter strain B-14 after 24 h, and by Sphingomonas sp. within 24 h [29]. Even more effective were Trichosporon sp. and Serratia sp., which were able to eliminate chlorpyrifos entirely in just 18 h [30]. Researchers in their work, used six wild-type microorganisms as prospective phosphotriesterases sources: Streptomyces phaeochromogenes, Streptomyces setonii, Nocardia corynebacterioides, Nocardia asteroides, and two Arthrobacter oxydans and then these were evaluated to hydrolyze paraoxon, methyl paraoxon, methyl parathion, coumaphos, dichlorvos, and chlorpyrifos [1].

The breakdown of pesticides into their component parts occurs in three distinct stages. Oxidation, reduction, or hydrolysis are some of the processes that are used to convert the potentially harmful forms of these chemicals into essences that are less damaging, watersoluble, and non-toxic. In the second act, the altered elixirs are combined with sugar or amino acids in order to produce an amalgam that is compatible with the other ingredients. In the third stage extracellular hydrolytic enzymes of bacteria and fungi conclude the degradation [31].

Organophosphate degradative genes

Degradative genes are often grouped or placed inside operons to enable the efficient degradation of organophosphorus compounds. Microbial communities in contaminated environments may acquire more degradation genes over time as they become more suited to the pollution. It is possible to enhance biodegradability by genetic engineering by recombining genes, which plays a role in biodegradability. That's why it's crucial to identify the genes responsible for resistance. It has been determined that several different microbial species are capable of degrading chlorpyrifos. Pseudomonas sp. WBC-3 is notable in that it possesses the oph gene, which is the gene responsible for the first step in the breakdown of chlorpyrifos. This oph gene is responsible for encoding the enzyme known as organophosphorus hydrolase (OPH), which is responsible for catalyzing the hydrolysis of chlorpyrifos into 3,5,6-trichloro-2-pyridinol (TCP). Many different kinds of microbes contribute to the breakdown of parathion. The phd gene, which encodes paraoxonase, hydrolyzes parathion to p-nitrophenol and is found in Pseudomonas aeruginosa and Pseudomonas sp. WBC-3 [32].

Support for enhancing organophosphorus biodegradation has been greatly bolstered by the discovery of many genes encoding organophosphorus degrading enzymes like opd, mpd, and oph [33−35]. P. diminuta's pCMS1 plasmid included the first known organophosphorus degrading (opd) gene. The gene was found to be 1.5 Kb and had restriction sites for SalI, PstI, XhoI-restricted and BamHI. In Flavobacterium sp., a plasmid designated pPDL2 had additional opd gene with an analogous restriction map. Widespread degrading ability against organophosphorus pesticides has been discovered after the isolation of the opd gene from Enterobacter sp. cons002. The breakdown power of altered OPH enzymes on P-S bonds of organophosphorus insecticides can be increased by genetic mutation and screening of OPH expressing genes. Parathion and fenitrothion are two pesticides that can be broken down by an enzyme encoded by the mpd gene, which is separate from the opd gene. Gene cloning has been used to effectively express the ophc2 gene, which has little similarity to other organophosphorus hydrolase genes, in Pichia pastoris and Escherichia coli [36, 37]. Table 1 summarizes the several bacterial genes that contribute to the breakdown of organophosphates. Toxic pesticides are degraded into less dangerous compounds by the enzymes produced by these genes, which aids in environmental cleanup.

Table 1 . Microbial genes playing role in organophosphorus degradation.

Sr No.OrganophosphateGeneSource MicroorganismReferences
1ChlorpyrifosmpdPlesiomonas sp. M6[38]
2Phosmet and FenthionopdAgrobacterium radiobacter P230[39]
3ChlorpyrifosmpdCupriavidus sp. DT-1[40]
4ChlorpyrifosopdDLactobacillus sakei WCP904[34]
5ChlorpyrifosmpdOchrobactrum sp. JAS2[41]
6ParathionphdPseudomonas aeruginosa[38]
7Methyl parathionophc2Stenotrophomonas sp. SMSP-1[42]
8Methyl parathion, Phorate, ParathionopdEEnterobacter sp.[43]
9Dimethoate, Paraoxon, Methyl parathionmphSerratia marcescens MEW06[44]
10Chlorpyrifos, Coumaphos, Diazinon, Methyl parathion, and ParathionopdBLactobacillus brevis WCP902[45]
11Triazophos, Methyl parathiontpdOchrobactrum sp., Pseudomonas putida KT2440[46, 47]


Enzymes degrading organophosphate compounds

Organophosphate insecticides (OPs) can be broken down by a wide variety of enzymes that can be found in both microorganisms and mammals. A few examples of these enzymes include the organophosphorus acid anhydrolase (OPAA), which was isolated from rabbit liver; the organophosphorus degrading hydrolases (OPHs), which involved phosphotriesterase (PTE) and organophosphorus degradation enzyme (OPD); and the methyl parathion hydrolase (MPH) and glycerophosphodiesterase (GpdQ), which were isolated from soil microorganisms [48].

The toxicity of organophosphorus compounds to animals including humans is much diminished once they have been hydrolyzed or breakdown. Organophosphate hydrolyzing enzymes are gaining popularity as potential solutions to the problem of decontamination since they can break down pesticides into residues that are harmless to people and the environment. Researchers have devised techniques to make use of the enzymes for biological remediation of OP chemicals, as microbial species have swiftly evolved enzymatic systems to breakdown these toxins. Enzymatic detoxification is an effective and environmentally friendly method for removing organophosphate pesticides and pollutants from contaminated sites [49]. Phosphatases, a type of hydrolase enzyme, have been widely used to eliminate phosphate groups from compounds, such as organophosphate pesticides, proteins, alkaloids, and nucleotides. Among these enzymes, phosphotriesterase (PTE) or organophosphorus hydrolase (OPH) was the earliest discovered enzyme with the ability to hydrolyze different types of organophosphate compounds by breaking P-S and P-O linkages. Microorganisms, particularly bacteria, are the primary source of these enzymes, including OPH, OP acid anhydrolases (OPAA), and methyl parathion hydrolase (MPH), which have proven to be highly effective for pesticide elimination.

S. phaeochromogenes entire cells converted 98% of chlorpyrifos into hydrolysis products at pH 8 and 40℃. Immobilized whole cells and enzyme extracts had similar hydrolytic activities, suggesting that both can be used depending on the application. Immobilization approaches, such as the utilization of genetically altered bacteria and bacterial consortia immobilized on diverse matrices, have been shown to be effective in pesticide biodegradation [50−52]. Compared with free cell cultures, studies have demonstrated that immobilized cells degrade and mineralize poisons more effectively. Immobilized cells showed superior breakdown rates compared to free cells when used for the treatment of chlorinated phenols and chlorpyrifos. Bio stimulation approaches, when used in conjunction with immobilized cells, represent a novel and promising approach to the bioremediation of pesticide-contaminated sites. Specific catalytic activity against a range of organophosphate pesticides has been observed in phosphotriesterases isolated from Flavobacterium sp. Another notable enzyme, OpdA, derived from Agrobacterium radiobacter, is an effective organophosphatedegrading enzyme that differs from OPH in substrate specificity because of differences in its active site structure [39, 53]. Immobilized OpdA has been demonstrated to efficiently remove methyl parathions from solution. Hydrolysis plays a crucial role in the breakdown of organophosphate compounds, making them more susceptible to microbial degradation. Organophosphorus hydrolase (OPH) and organophosphorus acid anhydrolase (OPAA) are two enzymes that have garnered the most attention. OPH is an enzyme found in a variety of bacteria; it has been the subject of much research due to its broad substrate specificity.

OpdB, a member of the OPD subclass, exhibits a hydrolase domain and displays a sequence identity of 52% with OPD. OpdB enzymes, in both their crude and their purified form, have an impressive capacity for the breakdown of organophosphates [54]. The discovery of this enzyme was kicked off by research conducted on a number of organophosphates (OPs) that were discovered in eight distinct types of pesticides. The capability of the Cupriavidus nantongensis X1T strain to hydrolyze OPs led to the discovery of the opdB gene, which is responsible for the breakdown of Ops [55].

Genetic engineering has been used to engineer microorganisms capable of the complete mineralization of organophosphorus compounds. Mutants with enhanced activity were obtained through directed evolution, leading to increased protein expression and improved degradation properties. Mutagenesis increases enzyme activity and substrate selectivity. More research is required to decipher the catalytic processes and evolutionary history of these enzymes. Table 2 summarizes various enzymes with potential to degrade organophosphorus compounds. Organophosphorus-degrading enzymes are increasingly used for prophylactic, therapeutic, and environmental purposes. Future research may improve the enzyme activity against difficult substrates and chemical combinations. New protein sequencing and structural determination will improve enzymatic destruction technologies, notably for chemical warfare weapons, and our understanding of evolutionary routes.

Table 2 . Key enzymes in organophosphate pesticide breakdown.

S. No.EnzymeSource OrganismDegrading Organophosphorus PesticideReference
1Paraoxonase (PON1)HumanParathion, Malathion, Chlorpyrifos, Diazinon[56]
2Organophosphorus hydrolase (OPH)Pseudomonas indoloxydansChlorpyrifos[28]
3Organophosphorus hydrolase (OPH)Pseudomonas resinovaransChlorpyrifos[27]
4HydrolaseCladosporium cladosporioidesChlorpyrifos[57]
5HydrolasePlesiomonas sp.Methyl Parathion[38]
6HydrolaseFlavobacterium sp.Parathion[58]
7PhosphotriesterasePseudomonas monteilii C11Coroxon[39]
8CarboxylesterasePseudomonas sp. M-3Malathion[59]
9Organophosphorus hydrolase (OPH)Pseudomonas sp. BF1-3Chlorpyrifos[60]
10Methyl Parathion hydrolaseBurkholderia cepaciaMethyl Parathion[61]
11Esterase B1Bacillus strain C5Methyl Parathion[62]
12Organophosphorus phosphatasesBacillus thuringiensis MB497Chlorpyrifos, Triazophos and Dimethoate[63]
13Alkaline phosphataseSpirulina platensisChlorpyrifos[64]
14PhosphotriesterasePseudomonas diminutaMethyl Parathion, Diazinon, Cyanophos, Dursban, Parathion, Coumophos[65]
15Methyl Parathion hydrolaseSerratia marcescens MEW06.Methyl Parathion, Paraoxon, Dimethoate[44]


The multi omics approach

Microorganisms are essential for pesticide degradation in both soil and water. Important natural processes, such as nutrient recycling, also rely on these bacteria. However, researchers still need more data on the interplay between genes in these bacteria to accurately anticipate how pesticides will break down. Researchers seek to better understand how these bacteria degrade pesticides by integrating information from systems biology and biogeochemistry [66]. With this information, we can develop procedures and guidelines for the safe and effective management of areas polluted by pesticides [67]. Degradation networks, computational biology, and state-of-the-art high-throughput omics tools have helped researchers to examine how bacteria degrade pesticides [68]. Researchers should investigate how various microorganisms in a community interact with one another and learn how microbial systems impact other biological systems. Many researchers have found that modern multi-omics techniques, such as genomics, metagenomics, transcriptomics, metabolomics, and proteomic analysis, were used to collect data on the gene and protein synthesis stages in whole microbial taxa in hostile environments to determine how biodegradation works and how to use it effectively [69−71]. Recent advances in sequencing methods have also helped isolate genes involved in the degradation of persistent contaminants [72]. Owing to the rapid development of gene sequencing technology, particularly high-throughput sequencing (HTS), several bacteria with biodegradation capabilities have been discovered [73]. HTS and metagenomics work together to reveal the species of bacteria that interact with one another in a given ecosystem. We learned more about the traits of biodegrading bacteria through transcriptomics, proteomics, and metabolomics research into their genotypes and phenotypes. Genome-scale models (GEMs) that help choose the most efficient microbes for bioremediation may be developed with the use of this information. Modelling pesticide breakdown can be improved by using information gained from diverse microbial omics [74]. Microbial strains from genera Paenibacillus sp., Pseudomonas sp., Burkholderia sp., Rhodococcus sp., and Pencillium sp. have been demonstrated to be capable of degrading chlorpyrifos, parathion, dimethoate, deltamethrin, p-nitrophenol, chlorimuron-ethyl, and nicosulfuron by using a systems biology approach that accompanied multi-omics strategies [75]. Particularly promising for bioremediation of pesticides are bacterial strains for which the complete genome has been sequenced, such as Pseudomonas putida KT2440 and Rhodococcus sp. Integrated omics techniques, which include genomics, transcriptomics, metabolomics, and proteomics, are helpful in systems biology research on microorganisms to examine the genetic-level regulation of bioremediation processes [74]. In the study of the transcriptome, methods such as microarrays and, more recently, next-generation sequencing especially RNA-Seq have been utilized to evaluate the response of microorganisms to various types of pesticides [71]. The complex processes, metabolic activity, genetic regulation, and molecular biological variables involved in the breakdown of xenobiotics by microorganisms were investigated. To improve the efficacy of bioremediation systems, the multi-omics approach, when combined with system analysis, generates a useful output for functional validation and genetic manipulation.

Biodegradation network

An innovative strategy for pesticide bioremediation is the use of computational tools and bioinformatic resources. Information on microbial biodegradation pathways and the biocatalytic processes involved may be found in online biodegradative databases including UMBBD, Bionemo, PTID, MBGD, OxDBase, BioCyc, and MetaCyc [76]. These resources help find info on microorganisms degrading pesticides and also identify microbial degradation intermediates. The database UMBBD, maintained by the University of Minnesota, demonstrates how microorganisms utilise enzymes to degrade several pesticides into less toxic chemicals [77]. MetaRouter is a one-stop shop for biodegradation and bioremediation data mining [78], whereas MBGD makes it easier to do comparative genomic research [79, 80]. Biodegradative oxygenases are the main focus of OxDBases because of their importance in the degradation of persistent organic molecules. OxDBase is a powerful tool applicable to bioremediation studies, providing knowledge of oxygenases-catalyzed reactions and aiding in the understanding of degradation processes involved in bioremediation [81, 82]. Biodegradation gene sequences and data on their transcription and regulation can be found in Bionemo [76]. These records will help in the construction of reliable biodegradation systems. Utilizing various computational tools and resources from the field of bioinformatics will allow us to get an understanding of the process by which microorganisms in the environment break down harmful chemicals such as pesticides. With the provision of information on enzymes, genes, and reactions, the databases contribute to the process of bioremediation. To better clean the environment and lessen the effects of dangerous substances, scientists must understand these processes.

Way microbes interact with chemical substances and their potential use in bioremediation is being studied using state-of-the-art scientific tools and systems biology methodologies. Enzyme-based processes for enhanced bioremediation and soil health have been developed using computational approaches. Computational biology and other similar in-the-computer methods have made the study of genes, proteins, and cellular systems possible. Biodegradation and bioremediation rely on intricate metabolic pathways, which can only be fully comprehended with the aid of these computational methods. Data mining and comprehension of cellular metabolic networks, especially as they pertain to biodegradation and bioremediation, can be accomplished using a number of in silico methods [8]. Stoichiometric studies of metabolic networks are commonly performed using tools such as flux balance, metabolic flux, and metabolic route analyses. Increased consumption of pesticide chemicals and tinkering with the features of degrading bacteria are both possible with the use of computational techniques. To foretell how pesticide chemicals would interact with bioremediating bacteria, scientists employ quantitative structure-activity relationship (QSAR) and three-dimensional quantitative structure-activity relationship (3DQSAR) models [83]. For effective pesticide bioremediation, we can manipulate genes, introduce novel enzymes, and control metabolic pathways using tools such as OptKnock, OptStrain, and OptReg [8]. To further bioremediation studies, these computational tools will aid in the comprehension of genome-scale models, interacting genes, and genomic data.

Genome editing

Editing genes is a unique technique that allows precise manipulation of DNA through the use of designed nucleases. Meganucleases, also referred to as homing nucleases, were one of the earliest types of nucleases designed to target specific genomic regions for use in gene editing [84, 85]. Meganucleases have long nucleotide recognition sequences that may be present once in the genome, and these nucleases cause a double-strand break (DSB) when they reach their target. Protein engineering, structure-based design, and molecular evolution are some of the methods that may be used to reengineer meganucleases to target novel sequences, but the process is often time-consuming [86]. To edit DNA, one must first locate the target gene using a reference classification system created for that gene, then allow for a disruption that will be repaired by homologous recombination, and finally modify the desired sequence components [87]. Genome expurgation, also known as genome editing, is a form of clean gene technology that safely eliminates genetic markers that are utilized throughout the process of genetic modification; hence, plants are free of any genetic residue. This approach has the potential to lessen the risk caused by bacteria that have antibiotic resistance markers or intentionally chosen markers. Technological developments need to come in the form of more potent genome-editing technologies, which may allow for a broader range of comprehension. This includes creating novel strains that are genetically amenable to transformation and development of CRISPR-Cas-based methods for multiplex genome silencing and editing. A systems biology-based high-throughput strategy that combines the use of synthetic biology and genome modification with in silico simulation and prediction is expected to significantly accelerate the method of transformation in cells [88].

The eradication of xenobiotics, simplification of complex molecules, and breakdown of highly hazardous chemicals into less harmful compounds include all examples of biodegradation, and further genetic editing techniques can speed up the biodegradation process by changing the gene architecture of microbes [89]. Zinc finger nucleases (ZFNs), transcription activator-like effector nucleases (TALENs), and clustered regularly interspaced short palindromic repeat/CRISPR-associated protein (CRISPR/Cas) are commonly used tools for genome editing [90−92]. Techniques such as genome editing and systems biology hold great promise for use in the bioremediation of heavy metals, persistent organic pollutants, gasoline, acid drainage, and xenobiotics.

Genome editing tools. The discovery of novel genes involved in the degradation of various recalcitrant pollutants has been aided by high-throughput nextgeneration sequencing [93]. CRISPR-Cas, ZFN, and TALEN are the main gene editing techniques that can be applied to the aforementioned promises [91, 92]. ZFNs and TALENs merge transcription factor DNAbinding domains with the nuclease domain of FokI, creating an obligatory dimer. Zinc finger DNA-binding domains and TAL effector DNA-binding domains are responsible for this cleavage. These genome-editing techniques generate double-stranded breaks (DSBs) in the target gene sequence, which are subsequently repaired by homology-directed repair (HDR) and nonhomologous end joining (NHEJ) [94]. The goal of these genome-editing technologies is to produce microorganisms of the best possible quality, as well as better microorganisms with more complex genetic factors [95]. We can engineer new microorganisms with desirable traits by altering the genetic makeup of their wild ancestors [96]. Limitations of the use of the aforementioned genome editing procedures include off-target mutations, dangerous mutations, and the possibility of unintentional release of modified organisms into the natural environment [97].

Zinc finger nucleases. Zinc finger nucleases (ZFNs) are a specialized class of DNA-binding proteins created specifically for targeting and modifying particular DNA regions [98]. Zinc Finger Nucleases contain Zinc Finger Proteins (ZFPs) that are artificial restriction enzymes. Eukaryotic ZFPs are transcription factors that double the DNA-binding domains. The nucleotide cleavage domain (Folk I) from Flavobacterium okeanokoites is also present in the ZFNs. The cleavage domain is normally surrounded by four to six ZFPs; however, this number may vary depending on the target site. The ZFPs' 18-bp target specificity enables highly efficient targeted genome editing. ZFPs include 30 amino acids and are folded into a three-dimensional structure consisting of an alpha helix and two antiparallel beta sheets [8]. The programmable nuclease, ZFN, is composed of a sequence-specific zinc-finger protein with a nuclease domain obtained from the FokI restriction endonuclease. DSBs of ZFNs are induced by their DNA-cleavage domain once the DNA-binding domain recognizes the target DNA sequence. DNA damage is repaired by the cell's own mechanisms, which can lead to alterations or disruptions in the genes.

Although ZFNs have received much attention for their potential use in genome editing, their direct role in the bioremediation of pesticides is not very significant. ZFNs can be employed to create microbes or plants with enhanced ability to break down or detoxify pesticides. ZFNs can improve the bioremediation capacity of these species by altering genes involved in pesticide metabolism or resistance [8]. One strategy is to employ ZFNs to induce specific genetic alterations in pesticide-degrading microbes. ZFNs may improve the efficiency and efficacy of enzymes that assist in pesticide breakdown by precisely altering the genes essential for pesticide synthesis. This may lead to microorganisms with enhanced ability to break down or eliminate pesticide chemicals. ZFNs can also be used to create pesticide-resistant or detoxifying food crops. ZFNs can modify critical genes involved in pesticide sensitivity or metabolism to confer resistance or increase the plant's pesticide breakdown.

Transcription activator-like effector nucleases. When it comes to genome editing and modification, TALENS is state-of-the-art. TALENS contain TAL proteins that originate from the pathogenic bacterial species, Xanthomonas. TALEN comprises a DNA-binding domain and DNA-cleavage domain. The DNA-binding domain is similar to that of transcription activator-like effectors (TALEs). Tal proteins bind to sequences as short as 1−2 nucleotides hence making them extremely efficient. As a result of the presence of thirty-four tandem repetitions of amino acids, the nucleases implicated were extremely successful in binding. The TALE protein and nuclease domain of the FokI restriction endonuclease make up TALEN, a programmable nuclease. DSBs are created when the DNA cleavage domains of TALENs attach to their target DNA sequences. Although DSBs are very harmful, all existing organisms have developed repair mechanisms to restore the original sequence and preserve genomic function. TALENs were chosen to knock out the gene of interest (NHEJ) and knock out the target gene (HDR). The use of TALENs allows for the modification of crop plant genes that are involved in the metabolism or detoxification of pesticides, the introduction of new genes or metabolic pathways into microorganisms to breakdown particular pesticide compounds, and the modification of genes in microorganisms that encode specific enzymes that are involved in the degradation of pesticides [99].

DNA-binding and DNA-cleavage domains derived from TALE and FokI endonucleases, respectively, are combined to form TALEN. TALEN was initially used to disrupt genes and regulate transcription in the fungus Trichoderma reesei [100], and to modify genes in Rhizopus oryzae [101]. TALEN have several benefits over CRISPR, including a larger variety of target locations, fewer off-target effects, and greater editing effectiveness in heterochromatin areas. The creation of zinc finger nucleases and transcription activator-like effector nucleases requires considerable time and effort, further restricting their application in genome editing. ZFNs and TALENs are less common than CRISPR/Cas because of their complexity, high efficiency, and multiplexed genome editing capabilities [102, 103].

CRISPR-Cas. To perform precise genome editing or regulation at the DNA level, a molecular machine is required which consists of two primary components: a DNA-binding domain that recognizes and binds to specific DNA sequences, and an effector domain that enables DNA cleavage or regulates transcription near the binding site. CRISPR-Cas systems can be categorized into three distinct types: I, II, and III. Each system, including the model organisms, has its own distinct Cas [104]. CRISPR/Cas 9 has been widely studied and used for gene editing. This system is made up of endonuclease Cas9, CRISPR-derived RNA (crRNA), and trans-activating CRISPR RNA (tracer RNA) and has been the subject of numerous scientific investigations and developments [105]. Cas protein and guide RNAs (usually between 18 and 20 nucleotides in length) are two components used in CRISPR CAS technology. The advent of Cas9, an RNA-guided DNA endonuclease, has greatly simplified sequence-specific gene editing because it can be developed to target new sites by modifying its guide RNA sequence. Cas9, in its nuclease-inactive state, offers a flexible RNA-guided DNA-targeting platform for sequencespecific genome regulation, imaging, and epigenetic editing. Cleavage by Streptococcus pyogenes Cas9 (SpCas9) leads to blunt-end double-strand breaks (DSBs), whereas cleavage by Cas12a, which requires a crRNA to identify the T-rich PAM site, leads to staggered-end DSBs [106]. The complex, which consists of Cas9 and sgRNA, causes the nucleases RuvC and HNH to cleave the target DNA, resulting in breaks in the complementary and non-complementary strands, respectively. Specific guide RNAs (SgRNAs) are constructed to target locations three base pairs upstream of the protospacer adjacent motif (PAM) [107, 108]. The two endonuclease domains (RuvC and HNH) of the Cas enzyme cleave target DNA strands and generate double-stranded breaks (DSBs), which are subsequently repaired by processes such as non-homologous end joining (NHEJ) and homology directed repair (HDR).

Single-strand annealing (SSA) and microhomologymediated end joining (MMEJ) are two additional errorprone repair processes [109]. After a double-strand break (DSB), mutations such as indels and substitutions frequently result from NHEJ mechanisms [110]. NHEJ is recommended for gain/loss of functionality due to the mutagenic behaviour of likely insertions or deletions (indels) that result in changed reading frames and the development of mutations. In contrast, HR is a cell cycledependent event that takes place during the S/G2 phase and has a decreased efficiency of genome editing owing to its reliance on a template to carry out the repair process. In most cases, HR is the method of choice for gene knockout and insertion investigations. In addition to DNA, RNA can also be edited using CRISPR/Cas systems. Cas9, Cmr/Csm, and Cas13 are only a few CRISPR systems that can target and edit RNA molecules. A number of organisms, including fungi, have used these RNA-targeting tools to silence particular RNA, alter RNA in precise areas, and monitor RNA levels [111−113]. The key amino acids present in the nuclease domains of Cas9 and Cas12a have been changed to make their inactive variants [114]. These variants still have the capacity to attach to specific targets and hence allow for gene expression modification in various species. Repression of gene transcription by damaging RNA transcription factors is possible using inactive forms of Cas9 or Cas12a. Furthermore, these inactive variations can be used to stimulate gene expression via rearrangement of transcription activation regions. Cas12a, a novel endonuclease, cuts DNA without CRISPR RNA. Cas9 and Cas12a can create double-stranded breaks at several genomic target locations by using various crRNA guides.

Clustered Regularly Interspaced Short Palindromic Repeats technology has great potential to change many areas of biotechnology, and bioremediation is one area where it can make a significant difference. Conventional bioremediation methods are inefficient, slow, and difficult to target particular contaminants. Molecular biologists are warming up to CRISPR because this technique can be used with bacterial, fungal, and archaeal systems [115]. CRISPR technology, which permits genome editing, may help overcome these limitations. The bioremediation processes, such as the removal of xenobiotics, the transformation of harmful substances into less toxic substances, and the breakdown of pesticides into simple components, can all be significantly improved by the use of gene editing techniques [89, 95]. The end goal of these gene-editing methods is to develop microorganisms with the highest possible quality and to generate microbes with more complicated genes [95, 116]. Scientists can enhance the metabolic pathways, pollutant tolerance, and contaminant removal efficiency of microorganisms employed in bioremediation by modifying their genes employing CRISPR technology. This precise engineering improves the organisms' capabilities, leading to stronger and more efficient bioremediation procedures. Bioremediation and the degradation of pesticides by rhizospheric bacteria via CRISPR Cas mechanisms have attracted a lot of attention recently [117, 118].

Achieving gene editing successful is a major challenge since it depends on a wide variety of parameters, including the speed of the Cas enzyme, the design of the sgRNA, the number of copies of the gene, and many more. The form of sgRNA is a crucial aspect that is often overlooked by current sgRNA design methods. In order to better forecast the morphologies of sgRNAs and enhance the design process, we need more accurate computer models. The accessibility of the target gene and presence of additional substances in its immediate vicinity can potentially affect the success of gene editing. For instance, the Cas enzyme may have difficulty accessing the target gene if it passes through a stretch of the DNA. Improved sgRNA design requires the creation of methods for predicting the DNA structure around target gene. Off-target mutations, deadly mutations, and the potential for inadvertent or purposeful release of transformed organisms into the environment, among other things, restrict the use of the aforementioned gene editing methods with functional genes of interest [97].

The benefit of CRISPR technology is that it allows multiplexed genome editing, which means that numerous genes may be targeted at once. This allows researchers to fine-tune microbes for use in bioremediation of targeted contaminants and habitats. The effectiveness and adaptability of cleaning can be increased by focusing on several genes implicated in biodegradation. To detect environmental toxins, CRISPR-based biosensors integrate the CRISPR-Cas system with a reporter system. These biosensors enable effective monitoring of polluted areas by providing quick and sensitive detection of contaminants through the construction of appropriate gRNAs. This makes it easier to implement precise bioremediation solutions to clean up certain areas. Precise delivery of genetic alterations to microbes in contaminated settings is now possible using CRISPR technology. Researchers can improve the performance of preexisting microbial communities for bioremediation without having to replace them entirely by infusing Cas9, gRNA, and other components, or by employing viral vectors. To correct errors in the DNA code that trigger disease, researchers have developed base editors as a new tool. These base editors combine enzymes that can alter specific bases in the DNA code (nCas9) with the original Cas9 protein. Cytidine and adenosine base editors are the two main types of chromatin modifications. Base editors for cytidine alter C to T and G to A [119], whereas those for adenosine switch A to G and T to C [120]. These base editors have been successfully used in living organisms to make precise changes to their DNA at specific locations. These base editors have been successfully used in living organisms to induce precise changes in their DNA at specific locations [121, 122].

Bacteria that thrive in polluted environments are considered to be promising option for metabolic engineering and genome editing because of their capability to adapt and survive in a variety of harsh environments that include poisonous, refractory, and non-degradable xenobiotics [123]. CRISPR-Cas technology has been widely adopted for use in model organisms such as Pseudomonas and E. coli [124]. Recent research has shown that nonmodel bacteria notably Achromobacter sp. HZ01 and Comamonas testosteroni are potentially effective when used in bioremediation [125]. Because of advancements in our understanding of CRISPR tools including the invention of gene-targeting gRNA, these species now have the ability to express particular genes that are critical for ecological restoration. Researchers have genetically manipulated the Cupriavidus nantongensis X1T strain using the CRISPR/Cas9 system in conjunction with the red system. Following the construction of two plasmids, pACasN and pDCRH, and their subsequent transfer to the X1T strain, targeted deletion of the opdB gene, which is responsible for the catabolism of organophosphorus pesticides, was achieved. This work shed light on gene targeting and the breakdown process of organophosphorus insecticides in the Cupriavidus genus, which was the subject of the study [126]. CRISPR-Cas9 was recently utilized by researchers to remove the yvmC gene from Bacillus licheniformis, increasing biotransformation efficiency to 100%. CRISPR-Cas9 may be used to improve organism strains in the future. CRISPR-cas9 can create bacteria that bioremediate environmental OPs [127].

Conclusion and Future Perspective

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The application of CRISPR technology presents a significant opportunity to fundamentally alter bioremediation. Scientists are able to improve the metabolic capacities of microorganisms, construct contamination-sensing systems, and design synthetic microbes to target and efficiently degrade pollutants using precise and diverse genome editing capabilities. These innovations have the potential to dramatically increase the efficiency and speed of bioremediation operations, leading to cleaner and healthier ecosystems for humans.

Acknowledgments

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We are grateful to everyone who helped us finish this work. We also thank Dr. Parag Sanghani, Provost PP Savani University, for his encouragement, assistance, and generous resources, which helped us finish our research. Dr. Jyotsana Pandit's ideas and persistent effort in reviewing data and writing the report are also appreciated.

References

Go to
  1. Santillan JY, Muzlera A, Molina M, Lewkowicz ES, Iribarren AM. 2020. Microbial degradation of organophosphorus pesticides using whole cells and enzyme extracts. Biodegradation 31: 423-433.
    Pubmed CrossRef
  2. Fosu-Mensah B, Okoffo E, Darko G, Gordon C. 2016. Organophosphorus pesticide residues in soils and drinking water sources from cocoa producing areas in Ghana. Environ. Syst. Res. 5: 10.
    CrossRef
  3. Allmaras RR, Wilkins DE, Burnside OC, Mulla DJ. 2018. Agricultural Technology and adoption of conservation practices, pp. 99-158. In: Advances in Soil and Water Conservation (Eds.) FJ Pierce & WW Frye. Routledge, New York.
    CrossRef
  4. Eskenazi B, Bradman A, Castorina R. 1999. Exposures of children to organophosphate pesticides and their potential adverse health effects. Environ. Health Perspect. 107(Suppl 3): 409-419.
    Pubmed KoreaMed CrossRef
  5. Anastassiades M, Lehotay S, Stajnbaher D, Schenck F. 2003. Fast and easy multiresidue method employing acetonitrile extraction/partitioning and "dispersive solid-phase extraction" for the determination of pesticide residues in produce. J. AOAC Int. 86: 412-431.
    Pubmed CrossRef
  6. Aldas VA, Poursat B, Sutton N. 2022. Potential and limitations for monitoring of pesticide biodegradation at trace concentrations in water and soil. World J. Microbiol. Biotechnol. 38: 240.
    Pubmed KoreaMed CrossRef
  7. Lekshmi C. 2023. Graphene quantum dots derived from honey and mangostin as sustainable materials to construct fluorescence turn on molecular switches for pesticide detection. ChemistrySelect 8: e202204869.
    CrossRef
  8. Jaiswal S, Singh D, Shukla P. 2019. Gene editing and systems biology tools for pesticide bioremediation: A review. Front. Microbiol. 10: 87.
    Pubmed KoreaMed CrossRef
  9. Karalliedde L, Senanayake N. 1999. Organophosphorus insecticide poisoning. J. Int. Fed. Clin. Chem. 11: 4-9.
  10. Sogorb MA, Vilanova E, Carrera V. 2004. Future application of phosphotriesterases in the prophylaxis and treatment of organophosphorus insecticide and nerve agent poisoning. Toxicol. Lett. 151: 219-233.
    Pubmed CrossRef
  11. Beauregard G, Lum J, Roufogalis BD. 1981. Effect of histidine modification on the aging of organophosphate-inhibited acetylcholinesterase. Biochem. Pharmacol. 30: 2915-2920.
    Pubmed CrossRef
  12. Zwiener RJ, Ginsburg CM. 1988. Organophosphate and carbamate poisoning in infants and children. Pediatrics 81: 121-126.
    Pubmed CrossRef
  13. Lessenger JE, Reese BE. 1999. Rational use of cholinesterase activity testing in pesticide poisoning. J. Am. Board Fam. Pract. 12: 307-314.
    Pubmed CrossRef
  14. De Silva HJ, Samarawickrema NA, Wickremasinghe AR. 2006. Toxicity due to organophosphorus compounds: what about chronic exposure. Trans. R. Soc. Trop. Med. Hyg. 100: 803-806.
    Pubmed CrossRef
  15. Ray DE, Richards PG. 2001. The potential for toxic effects of chronic, low-dose exposure to organophosphates. Toxicol. Lett. 120: 343-351.
    Pubmed CrossRef
  16. Mkenda P, Ndakidemi P, Mbega E, Stevenson P, Arnold S, Gurr G, et al. 2019. Multiple ecosystem services from field margin vegetation for ecological sustainability in agriculture: scientific evidence and knowledge gaps. PeerJ. 7: e8091.
    Pubmed KoreaMed CrossRef
  17. Agnihotri NP, Pandey SY, Jain HK, Srivastava KP. 1981. Persistence, leaching and movement of chlorfenvinphos, chlorpyrifos, disulfoton, fensulfothion, monocrotophos and tetrachlorvinphos in soil. Indian J. Agric. Chem. 24: 27-31.
  18. Farhan M, Khan AU, Wahid A, Ahmad M, Ahmad F. 2012. Biodegradation of chlorpyrifos using indigenous Pseudomonas sp. isolated from industrial drain. Pak. J. Nutr. 11: 1183-1189.
    CrossRef
  19. Abatenh E, Gizaw B, Tsegaye Z, Wassie M. 2017. The role of microorganisms in bioremediation- a review. Open J. Environ. Biol. 22: 038-046.
    CrossRef
  20. Nayak P, Solanki H. 2022. Impact of agriculture on environment and bioremediation techniques for improvisation of contaminated site. Int. Assoc. Biol. Comput. Digest. 1: 145-156.
    CrossRef
  21. Parween T, Bhandari P, Sharma R, Jan S, Siddiqui ZH, Patanjali PK. 2018. Bioremediation: a sustainable tool to prevent pesticide pollution. pp. 215-227. In Modern Age Environmental Problems and their Remediation, eds Mohammad O, Mohammad ZK, Iqbal MII. Cham: Springer.
    CrossRef
  22. Parkin TB, Daniel RS. 1994. Modeling environmental effects on enhanced carbofuran degradation. Pestic. Sci. 40: 163-168.
    CrossRef
  23. Amani F, Safari Sinegani AA, Ebrahimi F, Nazarian S. 2018. Biodegradation of chlorpyrifos and diazinon organophosphates by two bacteria isolated from contaminated agricultural soils. Biol. J. Microorg. 7: 27-39.
  24. Anwar S, Liaquat F, Khan QM, Khalid ZM, Iqbal S. 2009. Biodegradation of chlorpyrifos and its hydrolysis product 3, 5, 6-trichloro2-pyridinol by Bacillus pumilus strain C2A1. J. Hazard. Mater. 168: 400-405.
    Pubmed CrossRef
  25. Briceno G, Fuentes MS, Rubilar O, Jorquera M, Tortella G, Palma G, et al. 2015. Removal of the insecticide diazinon from liquid media by free and immobilized Streptomyces sp. isolated from agricultural soil. J. Basic Microbiol. 55: 293-302.
    Pubmed CrossRef
  26. Pailan S, Gupta D, Apte S, Krishnamurthi S, Saha P. 2015. Degradation of organophosphate insecticide by a novel Bacillus aryabhattai strain SanPS1, isolated from soil of agricultural feld in Burdwan, West Bengal, India. Int. Biodeterior. Biodegrad. 103: 191-195.
    CrossRef
  27. Sharma A, Pandit J, Sharma R, Shirkot P. 2016. Biodegradation of chlorpyrifos by Pseudomonas resinovarans strain AST2.2 isolated from enriched cultures. Curr. World Environ. 11: 267-278.
    CrossRef
  28. Sharma AK, Kasture A, Pandit J, Shirkot P. 2017. Isolation, characterization and identification of chlorpyrifos tolerant Pseudomonas strain exhibiting extracellular Organo-Phosphorus Hydrolase (OPH) activity from apple orchard soils of himachal pradesh. Asian J. Microbiol. Biotechnol. Environ. Sci. 19: 935-944.
  29. Li XH, He J, Li SP. 2007. Isolation of a chlorpyrifos-degrading bacterium, Sphingomonas sp. strain Dsp-2, and cloning of the mpd gene. Res. Microbiol. 158: 143-149.
    Pubmed CrossRef
  30. Xu G, Li Y, Zheng W, Peng X, Li W, Yan Y. 2007. Mineralization of chlorpyrifos by co-culture of Serratia and Trichosporon sp. Biotechnol. Lett. 29: 1469-1473.
    Pubmed CrossRef
  31. Van Eerd LL, Hoagland RE, Zablotowicz RM, Hall JC. 2003. Pesticide metabolism in plants and microorganisms. Weed Sci. 51: 472-495.
    CrossRef
  32. Singh BK. 2009. Organophosphorus-degrading bacteria: ecology and industrial applications. Nat. Rev. Microbiol. 7: 156-164.
    Pubmed CrossRef
  33. Gothwal A, Dahiya M, Beniwa P, Hooda V. 2014. Purification and kinetic studies of organophosphorus hydrolase from B. diminuta. Int. J. Pharm. Pharmaceut. Sci. 6: 341-344.
  34. Haque MA, Hong SY, Hwang CE, Kim SC, Cho KM. 2018. Cloning of an organophosphorus hydrolase opdD gene of Lactobacillus sakei WCP904 isolated from chlorpyrifos-impregnated kimchi and hydrolysis activities of its gene product for organophosphorus pesticides. Appl. Biol. Chem. 61: 643-651.
    CrossRef
  35. Ying W, YaPing W, Can H, Lixin M, Hong Y, Yong M, et al. 2021. High-level extracellular production and immobilization of methyl parathion hydrolase from Plesiomonas sp. M6 expressed in Pichia pastoris. Protein Expr. Purif. 183: 105859.
    Pubmed CrossRef
  36. Yu H, Yan X, Shen W, Hong Q, Zhang J, Shen Y, Li S. 2009. Expression of methyl parathion hydrolase in Pichia pastoris. Curr. Microbiol. 59: 573-578.
    Pubmed CrossRef
  37. Zhongli C, Shunpeng L, Guoping F. 2001. Isolation of methyl parathion-degrading strain M6 and cloning of the methyl parathion hydrolase gene. Appl. Environ. Microbiol. 67: 4922-4925.
    Pubmed KoreaMed CrossRef
  38. Cui ZL, Li SP, Fu GP. 2001. Isolation of methyl parathion degrading strain M6 and cloning of the methyl parathion hydrolase gene. Appl. Environ. Microbiol. 67: 4922-4925.
    Pubmed KoreaMed CrossRef
  39. Horne I, Sutherland TD, Harcourt RL, Russell RJ, Oakeshott JG. 2002. Identification of an opd (organophosphate degradation) gene in an Agrobacterium isolate. Appl. Environ. Microbiol. 68: 3371-3376.
    Pubmed KoreaMed CrossRef
  40. Lu P, Li Q, Liu H, Feng Z, Yan X, Hong Q, et al. 2013. Biodegradation of chlorpyrifos and 3,5,6-trichloro-2-pyridinol by Cupriavidus sp. DT-1. Bioresour. Technol. 27: 337-342.
    Pubmed CrossRef
  41. Abraham J, Silambarasan S. 2016. Biodegradation of chlorpyrifos and its hydrolysis product 3,5,6-trichloro-2-pyridinol using a novel bacterium Ochrobactrum sp. JAS2: A proposal of its metabolic pathway. Pestic. Biochem. Physiol. 26: 13-21.
    Pubmed CrossRef
  42. Shen YJ, Lu P, Mei H, Yu HJ, Hong Q, Li SP. 2010. Isolation of a methyl parathion-degrading strain Stenotrophomonas sp. SMSP-1 and cloning of the ophc2 gene. Biodegradation 21: 785-792.
    Pubmed CrossRef
  43. Chino-Flores C, Dantán-González E, Vázquez-Ramos A, Tinoco-Valencia R, Díaz-Méndez R, Sánchez-Salinas E, et al. 2012. Isolation of the opdE gene that encodes for a new hydrolase of Enterobacter sp. capable of degrading organophosphorus pesticides. Biodegradation 23: 387-397.
    Pubmed CrossRef
  44. Wang Y, Liu C, Wan J, Sun X, Ma W, Ni H. 2018. Molecular cloning and characterization of a methyl parathion hydrolase from an organophosphorus-degrading bacterium, Serratia marcescens MEW06. FEMS Microbiol. Lett. 365. doi: 10.1093/femsle/fny279.
    CrossRef
  45. Islam SMA, Math RK, Cho KM, Lim WJ, Hong SY, Kim JM, et al. 2010. Organophosphorus hydrolase OpdB of Lactobacillus brevis WCP902 from kimchi is able to degrade organophosphorus pesticides. J. Agric. Food Chem. 58: 5380-5386.
    Pubmed CrossRef
  46. Zhong W, Chen J. 2006. Environmental biotechnology in China: Progress and prospect. Biotechnol. J. 1: 1241-1252.
    Pubmed CrossRef
  47. Gu LF, He J, Huang X, Jia KZ, Li SP. 2006. Construction of a versatile degrading bacteria Pseudomonas putida KT2440-DOP and its degrading characteristics. Wei Sheng Wu Xue Bao. 46: 763-766.
  48. Schenk G, Mateen I, Ng TK, Pedroso MM, Miti'c N, Jafelicci M, Marques RFC, et al. 2016. Organophosphate degrading metallohydrolases: Structure and function of potent catalysts for applications in bioremediation. Coord. Chem. Rev. 317: 122-131.
    CrossRef
  49. Sidhu GK, Singh S, Kumar V, Dhanjal DS, Datta S, Singh J. 2019. Toxicity, monitoring and biodegradation of organophosphate pesticides: A review. Crit. Rev. Environ. Sci. Technol. 49: 1135-1187.
    CrossRef
  50. Moreno-Medina DA, Sanchez-Salinas E, Ortiz-Hernandez L. 2014. Removal of methyl parathion and coumaphos pesticides by a bacterial consortium immobilized in Luffa cylindrica. Rev. Int. Contam. Ambient. 30: 51-63.
  51. Mulchandani A, Kaneva I, Chen W. 1999. Detoxification of organophosphate nerve agents by immobilized Escherichia coli with surface-expressed organophosphorus hydrolase. Biotechnol. Bioeng. 63: 216-223.
    CrossRef
  52. Talwar MP, Ninnekar HZ. 2015. Biodegradation of pesticide profenofos by the free and immobilized cells of Pseudoxanthomonas suwonensis strain HNM. J. Basic. Microbiol. 55: 1094-1103.
    Pubmed CrossRef
  53. Gresham C, Rosenbaum C, Gaspari RJ, Jackson CJ, Bird SB. 2010. Kinetics and efficacy of an organophosphorus hydrolase in a rodent model of methyl-parathion poisoning. Acad. Emerg. Med. 17: 736-740.
    Pubmed KoreaMed CrossRef
  54. Fang LC, Shi QY, Xu LY, Shi TZ, Wu XW, Li QX, Hua RM. 2020. Enantioselective uptake determines degradation selectivity of chiral profenofos in Cupriavidus nantongensis X1T. J. Agric. Food Chem. 68: 6493-6501.
    Pubmed CrossRef
  55. Shi TZ, Fang LC, Qin H, Wu XW, Li QX, Hua RM. 2019. Minutespeed biodegradation of organophosphorus insecticides by Cupriavidus nantongensis X1T. J. Agric. Food Chem. 67: 13558-13567.
    Pubmed CrossRef
  56. Furlong CE, Richter RJ, Seidel SL, Costa LG, Motulsky AG. 1989. Spectrophotometric assays for the enzymatic hydrolysis of the active metabolites of chlorpyrifos and parathion by plasma paraoxonase/arylesterase. Anal. Biochem. 180: 242-247.
    Pubmed CrossRef
  57. Gao Y, Chen SH, Hu MY, Hu QB, Luo JJ, Li YN. 2012. Purification and characterization of a novel chlorpyrifos hydrolase from Cladosporium cladosporioides Hu-01. PLoS One 7: e38137.
    Pubmed KoreaMed CrossRef
  58. Mulbry WW, Karns JS. 1989. Purification and characterization of three parathion hydrolases from gram-negative bacterial strains. Appl. Environ. Microbiol. 55: 289-293.
    Pubmed KoreaMed CrossRef
  59. Singh AK, Srikanth NS, Malhotra OP, Seth PK. 1989. Characterization of carboxylesterase from malathion degrading bacterium: Pseudomonas sp. M-3. Bull. Environ. Contam. Toxicol. 42: 860-867.
    Pubmed CrossRef
  60. Barman DN, Haque MA, Islam SM, Yun HD, Kim MK. 2014. Cloning and expression of ophB gene encoding organophosphorus hydrolase from endophytic Pseudomonas sp. BF1-3 degrades organophosphorus pesticide chlorpyrifos. Ecotoxicol. Environ. Saf. 108: 135-141.
    Pubmed CrossRef
  61. Ekkhunnatham A, Jongsareejit B, Yamkunthong W, Wichitwechkarn J. 2012. Purification and characterization of methyl parathion hydrolase from Burkholderia cepacia capable of degrading organophosphate insecticides. World J. Microbiol. Biotechnol. 28: 1739-1746.
    Pubmed CrossRef
  62. Hao J, Liu J, Sun M. 2014. Identification of a marine Bacillus strain C5 and parathion-methyl degradation characteristics of the extracellular esterase B1. Biomed. Res. Int. 2014: 863094.
    Pubmed KoreaMed CrossRef
  63. Ambreen S, Yasmin A, Aziz S. 2020. Isolation and characterization of organophosphorus phosphatases from Bacillus thuringiensis MB497 capable of degrading Chlorpyrifos, Triazophos and Dimethoate. Heliyon 6: e04221.
    Pubmed KoreaMed CrossRef
  64. Thengodkar RRM, Sivakami S. 2010. Degradation of chlorpyrifos by an alkaline phosphatase from the cyanobacterium Spirulina platensis. Biodegradation 21: 637-644.
    Pubmed CrossRef
  65. Dumas DP, Caldwell SR, Wild JR, Raushel FM. 1989. Purification and properties of the phosphotriesterase from Pseudomonas diminuta. J. Biol. Chem. 264: 19659-19665.
    Pubmed CrossRef
  66. Bhatt P. 2019. Smart bioremediation technologies: microbial enzymes. Amsterdam: Elsevier Science.
  67. Chakraborty R, Wu CH, Hazen TC. 2012. Systems biology approach to bioremediation. Curr. Opin. Biotechnol. 23: 483-490.
    Pubmed CrossRef
  68. Otwell AE, Lopez Garcıa de Lomana A, Gibbons SM, Orellana MV, Baliga NS. 2018. Systems biology approaches towards predictive microbial ecology. Environ. Microbiol. 20: 4197-4209.
    Pubmed CrossRef
  69. Bharagava RN, Purchase D, Saxena G, Mulla S. 2019. Applications of metagenomics in microbial bioremediation of pollutants: from genomics to environmental cleanup. pp. 459-477. In: Microbial Diversity in the Genomic Era. Academic Press.
    CrossRef
  70. Marco DE, Abram F. 2019. Using genomics, metagenomics and other "Omics" to assess valuable microbial ecosystem services and novel biotechnological applications. Front. Microbiol. 10: 151.
    Pubmed KoreaMed CrossRef
  71. Rodriguez A, Castrejón-Godinez ML, Salazar-Bustamante E, Gama-Martinez Y, Sánchez-Salinas E, Mussali-Galante P, et al. 2020. Omics approaches to pesticide biodegradation. Curr. Microbiol. 77: 545-563.
    Pubmed CrossRef
  72. Bhatt P, Gangola S, Chaudhary P, Khati P, Kumar G, Sharma A, et al. 2019. Pesticide induced up-regulation of esterase and aldehyde dehydrogenase in indigenous Bacillus spp. Bioremediat. J. 23: 42-52.
    CrossRef
  73. Techtmann SM, Hazen TC. 2016. Metagenomic applications in environmental monitoring and bioremediation. J. Ind. Microbiol. Biotechnol. 43: 1345-1354.
    Pubmed CrossRef
  74. De Sousa CS, Hassan SS, Pinto AC, Silva WM, De Almeida SS, Soares SDC, et al. 2018. Microbial omics: applications in biotechnology. pp. 3-20. In Omics Technologies and Bio-Engineering, eds D. Barh and V. Azevedo. Cambridge, MA: Academic Press.
    CrossRef
  75. Hassan S, Ganai BA. 2023. Deciphering the recent trends in pesticide bioremediation using genome editing and multi-omics approaches: A review. World J. Microbiol. Biotechnol. 39: 151.
    Pubmed CrossRef
  76. Arora PK, Bae H. 2014. Integration of bioinformatics to biodegradation. Biol. Proc. Online 6: 8.
    Pubmed KoreaMed CrossRef
  77. Erythropel HC, Zimmerman JB, de Winter TM, Petitjean L, Melnikov F, Lam CH, et al. 2018. The green chemisTREE: 20 years after taking root with the 12 principles. Green Chem. 20: 1929-1961.
    CrossRef
  78. Korjus H. 2014. Polluted soils restoration. pp.411-480. In: Numben ST, Almaraz RA, Eswaran H, editors. Climate Change and Restoration of Degraded Land. Madrid: Colegio de Ingenieros de Montes.
  79. Bhatt P. 2018. In silico tools to study the bioremediation in microorganisms. pp. 389-395. In: Handbook of Research on Microbial Tools for Environmental Waste Management. Harrisburg: IGI Global.
    CrossRef
  80. Selzer PM, Marhöfer RJ, Koch O. 2018. Comparative genome analyses. pp. 123-140. In: Applied Bioinformatics. Cham: Springer.
    CrossRef
  81. Jaiswal S, Shukla P. 2020 Alternative strategies for microbial remediation of pollutants via synthetic biology. Front. Microbiol. 11: 808.
    Pubmed KoreaMed CrossRef
  82. Shah V, Jain K, Desai C, Madamwar D. 2012. Molecular analyses of microbial activities involved in bioremediation. pp. 221-247. In: Microorganisms in Environmental Management. Dordrecht: Springer.
    CrossRef
  83. Almeida L, Moraes L, Trigo J, Omoto C, Consoli F. 2017. The gut microbiota of insecticide-resistant insects houses insecticidedegrading bacteria: a potential source for biotechnological exploitation. PLoS One 12: e0174754.
    Pubmed KoreaMed CrossRef
  84. Rouet P, Smih F, Jasin M. 1994. Expression of a site-specific endonuclease stimulates homologous recombination in mammalian cells. Proc. Natl. Acad. Sci. USA 91: 6064-6068.
    Pubmed KoreaMed CrossRef
  85. Silva G, Poirot L, Galetto R, Smith J, Montoya G, Duchateau P, et al. 2011. Meganucleases and other tools for targeted genome engineering: perspectives and challenges for gene therapy. Curr. Gene Ther. 11: 11-27.
    Pubmed KoreaMed CrossRef
  86. Grizot S, Epinat JC, Thomas S, Duclert A, Rolland S, Pâques F, et al. 2010. Generation of redesigned homing endonucleases comprising DNA-binding domains derived from two different scaffolds. Nucleic Acids Res. 38: 2006-2018.
    Pubmed KoreaMed CrossRef
  87. Bier E, Harrison MM, O'Connor-Giles KM, Wildonger J. 2018. Advances in engineering the fly genome with the CRISPR-Cas system. Genetics 208: 1-18.
    Pubmed KoreaMed CrossRef
  88. Campbell K, Xia J, Nielsen J. 2017. The impact of systems biology on bioprocessing. Trends Biotechnol. 35: 1156-1168.
    Pubmed CrossRef
  89. Hussain I, Aleti G, Naidu R, Puschenreiter M, Mahmood Q, Rahman MM, et al. 2018. Microbe and plant assisted-remediation of organic xenobiotics and its enhancement by genetically modified organisms and recombinant technology: a review. Sci. Total Environ. 628-629: 1582-1599.
    Pubmed CrossRef
  90. Singh V, Gohil N, Ramírez García R, Braddick D, Fofié CK. 2018. Recent advances in CRISPR-Cas9 genome editing technology for biological and biomedical investigations. J. Cell. Biochem. 119: 81-94.
    Pubmed CrossRef
  91. Waryah CB, Moses C, Arooj M, Blancafort P. 2018. Zinc fingers, TALES, and CRISPR systems: a comparison of tools for epigenome editing. Epigenome Editing 1767: 19-63.
    Pubmed CrossRef
  92. Wong DWS. 2018. Gene targeting and genome editing. pp. 187-197. In The ABCs of Gene Cloning, ed Wong DWS. Cham: Springer.
    CrossRef
  93. Suenaga H. 2012. Targeted metagenomics: a high-resolution metagenomics approach for specific gene clusters in complex microbial communities. Environ. Microbiol. 14: 13-22.
    Pubmed CrossRef
  94. Yadav R, Kumar V, Baweja M, Shukla P. 2018. Gene editing and genetic engineering approaches for advanced probiotics: A review. Crit. Rev. Food Sci. Nutr. 58: 1735-1746.
    Pubmed CrossRef
  95. Basu S, Rabara RC, Negi S, Shukla P. 2018. Engineering PGPMOs through gene editing and systems biology: a solution for phytoremediation. Trends Biotechnol. 36: 499-510.
    Pubmed CrossRef
  96. Dai Z, Zhang S, Yang Q, Zhang W, Qian X, Dong W, et al. 2018. Genetic tool development and systemic regulation in biosynthetic technology. Biotechnol. Biofuels 11: 152.
    Pubmed KoreaMed CrossRef
  97. Canver MC, Joung JK, Pinello L. 2018. Impact of genetic variation on CRISPR-Cas targeting. CRISPR J. 1: 159-170.
    Pubmed KoreaMed CrossRef
  98. Meyer M, Angelis M, Wurst W, Kühn R. 2010. Gene targeting by homologous recombination in mouse zygotes mediated by zinc-finger nucleases. Proc. Natl. Acad. Sci. USA 107: 15022-15026.
    Pubmed KoreaMed CrossRef
  99. Lillico S, Proudfoot C, Carlson D, Stverakova D, Neil C, Blain C, et al. 2013. Live pigs produced from genome edited zygotes. Sci. Rep. 3: 2847.
    Pubmed KoreaMed CrossRef
  100. Liu P, Wang W, Wei D. 2017. Use of transcription activator-like effector for efficient gene modification and transcription in the filamentous fungus Trichoderma reesei. J. Ind. Microbiol. Biotechnol. 44: 1367-1373.
    Pubmed CrossRef
  101. Tsuboi Y, Sakuma T, Yamamoto T, Horiuchi H, Takahashi F, Igarashi K, et al. 2022. Gene manipulation in the Mucorales fungus Rhizopus oryzae using TALENs with exonuclease overexpression. FEMS Microbiol. Lett. 369: fnac010.
    Pubmed KoreaMed CrossRef
  102. Banerjee A, Banerjee C, Negi S, Chang J-S, Shukla P. 2018. Improvements in algal lipid production: a systems biology and gene editing approach. Crit. Rev. Biotechnol. 38: 369-385.
    Pubmed CrossRef
  103. Shah T, Andleeb T, Lateef S, Noor MA. 2018. Genome editing in plants: advancing crop transformation and overview of tools. Plant Physiol. Biochem. 131: 12-21.
    Pubmed CrossRef
  104. Cooper LA, Stringer AM, Wade JT. 2018. Determining the specificity of cascade binding, interference, and primed adaptation In Vivo in the Escherichia coli type IE CRISPR-Cas system. mBio 9: e02100-17.
    Pubmed KoreaMed CrossRef
  105. Swartjes T, Staals RHJ, van der Oost J. 2020. Editor's cut: DNA cleavage by CRISPR RNA-guided nucleases Cas9 and Cas12a. Biochem. Soc. Trans. 48: 207-219.
    Pubmed KoreaMed CrossRef
  106. Zetsche B, Gootenberg JS, Abudayyeh OO, Slaymaker IM, Makarova KS, Essletzbichler P, et al. 2015. Cpf1 is a single RNAguided endonuclease of a class 2 CRISPR-Cas system. Cell 163: 759-771.
    Pubmed KoreaMed CrossRef
  107. Barrangou R, Birmingham A, Wiemann S, Beijersbergen RL, Hornung V, Smith AB. 2015. Advances in CRISPR‐Cas9 genome engineering: Lessons learned from RNA interference. Nucleic Acids Res. 43: 3407-3419.
    Pubmed KoreaMed CrossRef
  108. Liu D, Hu R, Palla KJ, Tuskan GA, Yang X. 2016. Advances and perspectives on the use of CRISPR/Cas9 systems in plant genomics research. Curr. Opin. Plant Biol. 30: 70-77.
    Pubmed CrossRef
  109. Huang J, Cook DE. 2022. The contribution of DNA repair pathways to genome editing and evolution in filamentous pathogens. FEMS Microbiol. Rev. 46: fuac035.
    Pubmed KoreaMed CrossRef
  110. Barakate A, Stephens J. 2016. An overview of crispr based tools and their improvements: New opportunities in understanding plant-pathogen interactions for better crop protection. Front. Plant Sci. 7: 765.
    Pubmed KoreaMed CrossRef
  111. Abudayyeh OO, Gootenberg JS, Essletzbichler P, Han S, Joung J, Belanto JJ, et al. 2017. RNA targeting with CRISPR-Cas13. Nature 550: 280-284.
    Pubmed KoreaMed CrossRef
  112. Terns MP. 2018. CRISPR-based technologies: Impact of RNA-targeting systems. Mol. Cell 72: 404-412.
    Pubmed KoreaMed CrossRef
  113. Zhang Y, Ge H, Marchisio MA. 2022. A mutated Nme1Cas9 Is a functional alternative RNase to both LwaCas13a and RfxCas13d in the yeast S. cerevisiae. Front. Bioeng. Biotechnol. 10: 922949.
    Pubmed KoreaMed CrossRef
  114. Swarts DC, Jinek M. 2018. Cas9 versus cas12a/cpf1: structurefunction comparisons and implications for genome editing. Wiley Interdiscip. Rev. RNA. 9: e1481.
    Pubmed CrossRef
  115. Greene AC. 2018. CRISPR-based antibacterials: transforming bacterial defense into offense. Trends Biotechnol. 36: 127-130.
    Pubmed CrossRef
  116. Dangi AK, Sharma B, Hill RT, Shukla P. 2019. Bioremediation through microbes: systems biology and metabolic engineering approach. Crit. Rev. Biotechnol. 39: 79-98.
    Pubmed CrossRef
  117. Saxena P, Singh NK, Singh AK, Pandey S, Thanki A, Yadav TC. 2020. Recent advances in phytoremediation using genome engineering CRISPR-Cas9 technology. pp.125-141. In: Bioremediation of Pollutants. New York.
    CrossRef
  118. Stein HP, Navajas-Pérez R, Aranda E. 2018. Potential for CRISPR genetic engineering to increase xenobiotic degradation capacities in model fungi. pp. 61-78. In: Approaches in Bioremediation. Cham: Springer.
    CrossRef
  119. Gehrke JM, Cervantes O, Clement MK, Wu Y, Zeng J, Bauer DE, et al. 2018. An APOBEC3A-Cas9 base editor with minimized bystander and off-target activities. Nat. Biotechnol. 36: 977-982.
    Pubmed KoreaMed CrossRef
  120. Gaudelli NM, Komor AC, Rees HA, Packer MS, Badran AH, et al. 2017. Programmable base editing of A·T to G·C in genomic DNA without DNA cleavage. Nature 551: 464-471.
    Pubmed KoreaMed CrossRef
  121. Ryu SM, Koo T, Kim K, Lim K, Baek G, Kim ST, et al. 2018. Adenine base editing in mouse embryos and an adult mouse model of Duchenne muscular dystrophy. Nat. Biotechnol. 36: 536-539.
    Pubmed CrossRef
  122. Zafra MP, Schatoff EM, Katti A, Foronda M, Breinig M, Schweitzer AY, et al. 2018. Optimized base editors enable efficient editing in cells, organoids and mice. Nat. Biotechnol. 36: 888-893.
    Pubmed KoreaMed CrossRef
  123. Patel H, Shakhreliya S, Maurya R, Pandey VC, Gohil N, Bhattacharjee G, et al. 2022. CRISPR-assisted strategies for futuristic phytoremediation. Assisted Phytoremediation 2022: 203-220.
    CrossRef
  124. Okoli AS, Blix T, Myhr AI, Xu W, Xu X. 2021. Sustainable use of CRISPR/Cas in fish aquaculture: the biosafety perspective. Transgenic Res. pp. 1-21.
    Pubmed KoreaMed CrossRef
  125. Zhang J, Zhang H, Li S, Li J, Yan L, Xia L. 2021. Increasing yield potential through manipulating of an ARE1 ortholog related to nitrogen use efficiency in wheat by CRISPR/Cas9. J. Integr. Plant Biol. 63: 1649-1663.
    Pubmed CrossRef
  126. Zhang Y, Geng Y, Li S, Shi T, Ma X, Hua R, et al. 2023. Efficient knocking out of the organophosphorus insecticides degradation gene opdB in Cupriavidus nantongensis X1T via CRISPR/Cas9 with red system. Int. J. Mol. Sci. 24: 6003.
    Pubmed KoreaMed CrossRef
  127. Qian S, Chen Y, Xu X, Peng C, Wang X, Wu H, et al. 2022. Advances in amplification-free detection of nucleic acid: CRISPR/Cas system as a powerful tool Anal. Biochem. 643: 114593.
    Pubmed CrossRef

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