Molecular Mechanism for the Spread of Insecticide Resistance in Ades Aegyti
50Views & Citations
Aedes aegypti, is a mosquito that spread, dengue virus, chikungunya, Zika fever, Mayaro and yellow fever viruses. While the mosquito originated from the African continent, but has spread in tropical, subtropical and temperate regions, causing havoc to 100s of millions of people. Different control mechanism(s) are employed to curb the mosquito population including a repertoire of insecticide but certain limitations and biological resistance are often associated. The survival of A. aegypti is dependent on number of environmental factors, including temperature, humidity, pH and salinity and the eggs of can, nonetheless, remain viable in a desiccated state for up to one year. Factors that contribute for the spread of insecticide resistance are very complex and are subject of concern for vector Biologists. Chemical nature and mode of action of different insecticides, mechanisms of resistance to insecticides, environmental pollution and toxicity to humans, and role of miRNAs in insecticide resistance are explored. Many miRNAs are differentially expressed between insecticide sensitive and resistant state. A new paradigm in the control of A. aegypti can be obtained with this study.
Keywords: Aedes Aegypti, Insecticides, Biological Resistance, Environmental Factors
Massive urbanization and global warming have led to an explosion of vector-borne diseases in recent decades [1,2]. Several vector-borne diseases, such as Zika fever, dengue fever, yellow fever, and chikungunya, are transmitted by Aedes aegypti [3,4]. Most of these infections are not vaccine-preventable. Frequent genetic mutations in the antigenic regions of viral genomes and a wide diversity in viral serotypes pose elusive challenges in the development of vaccines [5-7]. Furthermore, a high cost of currently available vaccines also dissuades many third-world citizens from routine immunization. Therefore, pest management and control are presently the best options for the prevention of mosquito-borne diseases.
The lifespan of A. aegypti is brief, lasting from eight to ten days. However, it is dependent on a number of environmental factors, including temperature, humidity, pH and salinity [8,9]. The eggs of A. aegypti can, nonetheless, remain viable in a desiccated state for up to one year. This is an evolutionary advantage, the larvae emerge from the eggs as soon as extreme environmental conditions alleviate [10,11]. Only female A. aegypti mosquitoes are capable of biting since mouthparts of males cannot puncture the skin. Different chemical compounds present on human skin, for example, lactic acid, toluene, butanone, and octa decanoic acid, attract mosquitoes . Several odorant receptors in mosquitoes, such as Aaeg Or 4, draw them to the skin .
Mosquito repellents usually contain twenty to thirty per cent N, N-diethyl metatoluamide (DEET) . p-menthane-3, 8-diol, obtained from lemon eucalyptus (Corymbia citriodora), is also used in some insect repellents . Commercial insecticides used in the Kingdom of Saudi Arabia contain either organ phosphorous or pyrethroids Cypermethrin, in addition to twenty to thirty per cent DEET. Mosquitoes often develop resistance to these lower concentrations of DEET and higher concentrations could not be used since they are toxic to human beings and mammals [16,17]. Therefore, the evolution of insecticide-resistant mosquitoes is a serious health concern in many regions of the world.
Consequently, unconventional approaches to control pests are being studied and tried in different parts of the world. One such approach is the use of the bacterium Bacillus thuringiensis for mass-killing of mosquitoes . The spore-forming bacteria release crystal proteins (also known as Cry proteins or delta endotoxins) in the gut of mosquitoes. Cry proteins have broad-spectrum insecticidal activity; they can kill moths, butterflies, mosquitoes, flies, beetles, bees, wasps, and ants. Additionally, they are also effective against nematodes [19,20]. These toxins are encoded by the cry gene, which is usually carried on bacterial plasmids. The cry gene can, hence, be transferred from one strain to another by conjugation . Using B. thuringiensis, several groups have targeted the A aegypti in both lab and field trials.
Wolbachia is another bacterium being studied in this context. It is an endo symbiont that confers upon mosquitos the resistance to several viruses, such as dengue and Zika [22-24]. In several countries of the world, different strains of the viruses have been targeted using this approach [25,26].
Lately, a genetically engineered strain of A. aegypti, OX513A, was developed by a British biotech company. In these mosquitoes, a self-limiting gene is turned on, which diminishes survival in offspring. Male OX513A mosquitoes were released in the fields in Brazil and Panama since they are incapable of biting humans and animals. Ninety per cent reduction in the population of wild-type mosquitoes was observed. To further propagate OX513A in labs, the self-limiting gene is turned off using the tetracycline antidote. The mosquitoes are then released in fields. In the absence of the antidote in the environment, the self-limiting gene gets turned on again [27-29].
The most advanced technology is targeting microRNAs (miRNAs). Different miRNAs modulate different growth process in mosquitoes. For instance; miRNA-7, miR-8, miR9a, and miR124 regulate development; miR-124, miR-310–313 clusters, Bantam, Let-7, and miR1 regulate neurogenesis; miR278 and miR309 muscle and exoskeleton formation; and bantam and let7 wing development . The molecular mechanisms underlying these developmental processes are not clearly understood. However, miRNAs are known to have a strong association with the evolution of insecticide resistance [31,32].
Insects can potentially counter both the approaches, i.e., insecticide-based as well as bio control, by developing resistant strains. Therefore, it is imperative to look at the molecular and cellular mechanisms underlying the development of resistance.
CHEMICAL NATURE AND MODE OF ACTION OF DIFFERENT INSECTICIDES
Insecticides can be organic or inorganic and natural or synthetic. Organ chlorides, organophosphates, carbamates, pyrethroids, neonicotinoids, and ryanoids are the main classes of insecticides . Virtually all insecticides target the nervous system of insects, however, their mode of action slightly differ .
Some of the oldest and most widely used insecticides are organ chlorides. Organ chlorides, as the name suggests, are chlorinated hydrocarbons. They have very low water solubility and resist degradation in the environment. Organ chlorides include dichlorodiphenyltrichloroethane (DDT), aldrin, dieldrin, and lindane among others. Organ chlorides are divided into two main subgroups: DDT-like compounds and chlorinated alicyclic . DDT-like compounds target the peripheral nervous system (PNS) of insects. In axons, they hamper the closure of sodium channels and, hence, membrane depolarization. Sodium ions keep leaking through the neurons and form a destabilizing negative after-potential, causing hyper excitability of the neurons . Chlorinated alicyclic, on the other hand, bind to gamma-Amino butyric acid (GABA) A receptor (GABAA receptor), inhibiting chloride ions to flow into the neurons. After 2-8 h of exposure, the activity of insects’ central nervous system is depressed. This follows hyper excitability, tremors, and seizures [35,37].
Organophosphates are esters of phosphoric acid. As much as fifty per cent of synthetic insecticides are organophosphates. Some of the commonly used organophosphates include parathion, malathion, chlorpyrifos, phosmet, and diazinon . Organophosphates disrupt the neuromuscular enzyme, acetyl cholinesterase, by binding to it via covalent irreversible inhibition . When the enzyme is disrupted, nerve impulses are inhibited. Cholinergic neurons use acetylcholine as a transmitter substance. Acetyl cholinesterase catalyses’ the hydrolysis of acetylcholine at the synaptic gap, thus controlling the transmission of nerve impulses. When acetyl cholinesterase is inhibited by organophosphates, acetylcholine accumulates. Consequently, the receptors get saturated with acetylcholine, making nerve impulses inoperative. Many vital systems are affected simultaneously. However, the respiratory system shuts down first, causing the insect to die of respiratory failure [40-42].
Carbamates are derivatives of carbamic acid. Carbamate’s insecticides have the functional group carbamate ester. Some of the most familiar carbamate insecticides are aldicarb, carbofuran, ethienocarb, and fenobucarb . The mode of action of carbamates and organophosphates is essentially the same. Carbamates also disrupt acetyl cholinesterase. The only difference is that organophosphates cause phosphorylation of acetyl cholinesterase while carbamates cause carbamylation [39,44].
Synthetic pyrethroids are structural derivatives of pyrethrins, which are naturally produced by the pyrethrum flowers (Chrysanthemum cinerariifolium and Chrysanthemum coccineum). Some of the commonly used pyrethroids include allethrin, imiprothrin, permethrin, and cypermethrin . Household insecticides and insect repellants are pyrethroids in nature. Pyrethroids are basically axonic excitotoxins. They target the voltage-gated sodium channels of axons, keeping them open and hence preventing the repolarization of neurons. Insects exposed to pyrethroids paralyze consequently [46,35].
Neonicotinoids are related to nicotine. Neonicotinoids have a nitro-methylene, nitro-imine or cyanoimine group. The use of these insecticides is getting increasingly common. Most common neonicotinoids are Imidacloprid, nithiazine, acetamiprid, and clothianidin. These are agonists at nicotinic acetylcholine receptors (nAChRs), interacting with both alpha and beta unit of the receptor [47,48]. In contrast to vertebrates, nAChRs exclusively occur in the central nervous system (CNS) of insects. In fact, the nervous system of insects is very rich with nAChRs. The neurotransmitter acetylcholine activates these receptors [49,50]. Low and moderate concentrations of acetylcholine trigger nerve impulses. However, high concentrations of acetylcholine or neonicotinoids block nAChRs. The organism dies of paralysis. Furthermore, while acetyl cholinesterase breaks down acetylcholine, it is incapable of degrading Nicotinoids . Some neonicotinoids, such as Imidacloprid, depolarizes the motor neurons .
Ryanoids are synthetic analogues of ryanodine, which is a diterpenoid found in Ryania speciose . Chlorantraniliprole is the most commonly used ryanoids . The mode of action of ryanoids and ryanodine is the same. Both the compounds bind with open ryanodine receptors with very strong affinity. Ryanodine receptors are a class of calcium receptors that occur in muscle cells. Ryanoids either partially or completely close the ryanodine receptors, depending upon their concentrations. Consequently, calcium ions are released from calcium stores in the muscle cells. First, massive muscle contractions occur and then paralysis ensues [54,52].
MECHANISMS OF RESISTANCE TO INSECTICIDES
In recent years, resistance to almost all insecticides has been observed. Resistance to an insecticide usually develops when expression of a xenobiotic-degrading enzyme gets elevated or the enzyme develops a better affinity for a xenobiotic compound. Alternatively, the receptor that an insecticide target may also undergo structural and compositional changes so that it no longer binds to that insecticide.
Esterase, Glutathione S-Transferases, and Monooxygenases are three main groups of enzymes associated with resistance to Organochlorines, organophosphates, carbamates, and pyrethroids. Esterase’s bound and turns over insecticides. They do not degrade insecticides instead, they sequester them [55,56]. The genes involved in esterase-based resistance are estα and estβ. In mosquitoes, most commonly elevated phenotypes are estα21 and estβ21 . Mutations in the regulatory elements of esterase’s, causing their up regulation, have also been reported . Furthermore, in resistant strains of mosquitoes, esterase’s that bind to organophosphates with very high affinity have been documented . Resistant strains are also frequently reported to have high levels of glutathione S-transferases [60,61]. This enzyme detoxifies a large number of xenobiotic compounds . They bring about the nucleophilic attack of reduced glutathione on electrophilic centers of lipophiles . Two glutathione S-transferases with elevated levels have been identified in DTT-resistant A. aegypti . Monooxygenases occur in numerous animal species, including insects [64-66]. These enzymes are involved in the degradation of xenobiotic and endogenous metabolism. One of the best-known monooxygenases is P450, which metabolizes virtually all insecticides. P450 first captures molecular oxygen and then it receives electrons from NADPH, finally incorporating an oxygen molecule into its substrate. Elevated levels of this enzyme are linked with resistance to pyrethroids. Certain monooxygenases also convert organophosphates into their respective Oxon analogues before organophosphates can inhibit acetyl cholinesterase [67,68].
Acetyl cholinesterase, GABA receptors, and sodium channels are known to develop resistance to insecticides by undergoing changes in their amino acid composition. Receptors and enzymes with altered amino acid composition either do not bind to insecticides at all or bind them with reduced affinity . For example, sodium channels in mosquitoes acquire ‘kdr’-like resistance to both DDT and pyrethroids. An example of kdr mutation is the substitution of phenylalanine instead of leucine in the sixth trans membrane segment of sodium channels [69,70]. Often single amino acid substitutions are involved. E3 esterase, for instance, develops resistance to malathion when tryptophan at the 251st position gets replaced with leucine . Likewise, in E3 esterase, glycine at the 137th position may also get replaced with aspartate conferring resistant to several organophosphates . In dieldrin-resistant A. aegypti, an alanine to serine substitution occurs in channel lining domain of GABA receptors .
ENVIRONMENTAL POLLUTION AND TOXICITY TO HUMANS
Each year 4.6 million tons of pesticides are introduced into the environment globally. Interestingly, 99 per cent of these pesticides come in contact with non-target soil, water bodies, and atmosphere . Consequently, these strayed chemicals are absorbed by organisms. Annually, three million cases of acute pesticide poisoning are documented, resulting in two hundred and fifty thousand deaths [75,76]. Even in developed countries like the United States, pesticides have been isolated from a majority of wells in rural areas. Ocean currents and atmospheric circulation have even conducted pesticides, such as DDT, to sheets of ice in Greenland and bodies of Antarctic penguins . Needless to say, residual pesticides are today detected in all soil types, including vegetable fields and forest lands and deep groundwater [77,78].
The adverse effects of pesticides on domesticated animals and wildlife are conspicuous. Organochlorines, such as DDT, cause thinning of eggshells in birds  and acute mortality in small animals due to inhibition of acetyl cholinesterase . Furthermore, DDT is also a well-known carcinogen and endocrine disruptor . Organophosphates cause immunotoxicity in chordates by inhibiting serine hydrolases and esterase’s . Parathion, for example, causes susceptibility to fungal infections. Organochlorines, such as Chlordane, also have adverse effects on the immune system of vertebrates . Nicotinoids affect all the major organ systems of mammals, for instance, cardiovascular, immune, and nervous system . Imidacloprid and pyrethroids severely affect foraging and growth rate in insect colonies . Thiamethoxam, another Imidacloprid, causes homing failure and thus mortality in worker bees .
Since there is an enormous similarity between the metabolism and organ systems of humans and insects, pesticides are frequently reported to have detrimental effects on human health. For example, organ chlorides have been implicated in several human cancers, including pancreatic cancer, non-Hodgkin’s lymphoma, and breast cancer. They are also associated with impaired lactation and reduced fertility in men . Organophosphates are also known to have adverse effects on the reproductive health of humans . Furthermore, they also affect fetal and infant development and cognitive development in children [88,89]. Pyrethroids are infamous for their neurotoxic reactions . Neonicotinoids is a relatively newer class of insecticides. Initially, they were thought to be safe. However, their neurotoxic and genotoxic effects in humans have also been documented recently .
ROLE OF miRNAS IN INSECTICIDE RESISTANCE
miRNAs are small noncoding RNA molecules that are involved in post-transcriptional regulation and RNA interference [92,93]. To do so, they base-pair with the complementary sequences on the target mRNA molecule. Consequently, the target mRNA is cleaved and/or its translation is impaired [94,95]. However, many recent studies have found that miRNAs also up regulate the translation of their target mRNAs [96-99]. Lately, miRNAs have been implicated in promoting and inhibiting the translation of various genes that resist insecticides.
The common fruit fly (Drosophila melanogaster) is a model organism for studying insecticide resistance. It is the DDT resistance that is usually studied in the fruit fly. Previously, detoxification genes, such as cytochrome P450, glutathione S-transferases, ATP binding cassette transporters, and esterase’s were implicated in the DDT resistance in the fly. Recently, however, a few studies have focused on miRNAs. Pittendrigh  found that ten miRNAs were differentially expressed between the DDT resistant and DDT sensitive strain of D. melanogaster . These miRNAs targeted transcripts encoding different detoxification genes. For example, miR-311-3p, miR-312-3p, and miR-313-3p which target cytochrome P450 monooxygenases were down regulated in resistant flies. Cytochrome P450 monooxygenases detoxify different insecticides, including DDT. MiR-986-5p, miR-995-3p, miR-312-3p, and miR-2a-3p, on the other hand, were found to interact as well as affect the transcriptional level of multidrug resistance-associated protein B7. MiR-986-5p was found to be the most highly expressed among all the differentially expressed miRNAs in the resistant strain. The function of miR-986-5p is not clearly understood, however, the study found that it interacts with the transcripts of multiple detoxification genes.
Cotton aphids (Aphis gossypii) parasite on dozens of edible plants, such as watermelons, squash, cantaloupes, and asparagus. Spirotetramat is a keto-enol insecticide that inhibits the lipid biosynthesis in sucking insects (including aphids) by inhibiting the Acetyl-CoA carboxylase enzyme . It was recently found that miR-276 and miR-3016 up regulate the Acetyl-CoA carboxylase gene post-transcriptionally, hence rendering the insects resistant to the insecticide .
The diamondback both (Plutella xylostella) is an infamous pest of cruciferous vegetables. The resistance of this insect to synthetic insecticides and B. thuringiensis derived toxins is legendary . Chlorantraniliprole, a ryanoids insecticide, alters the expression of over one hundred miRNAs in the diamondback moth. A 2017 study by Zhu et al identified dozens of miRNAs differentially expressed between Chlorantraniliprole sensitive and Chlorantraniliprole resistant strains of the diamondback moth . The targets of these differentially expressed miRNAs were identified by miRanda and RNA hybrid. A majority of the miRNAs targeted genes encoding cytochrome P450, glutamate-gated chloride channel, glutathione S-transferases, ATP-binding cassette transporters, and cuticle proteins. For instance; pxy-miR-8533-3p targeted larval cuticle protein 30 ; pxy-miR-100-5p, glutamate-gated chloride channel ; pxy-miR-275-5p, multidrug resistance-associated protein 4 ; and pxy-miR-1175-5p, esterase FE4 . Etebari  found that enriching the diet of deltamethrin resistant P. xylostella larvae with miR-2b-3p increases mortality of the pest when it is exposed to deltamethrin. MiR-2b-3p down regulates the transcript levels of CYP9F2. CYP9F2 is a cytochrome P450 gene and it may have a role in metabolic resistance to insecticides .
The northern house mosquito (Culex pipiens), transmits a large number of infectious diseases, for example, West Nile fever, Japanese encephalitis, St Louis encephalitis, Sindbis fever, and lymphatic filariasis. Pyrethroids, such as deltamethrin, are used to eradicate the pest. However, the mosquito has developed a strong resistance to pyrethroids. Many recent studies implicated miRNAs in pyrethroids resistance in C. pipiens. For instance, in a 2016 study by Liu et al, the expression levels of miRNA-938 were found 1.8 times higher in deltamethrin resistant strains of C. pipiens pallens. Contrarily, the expression levels of CpCPR5, the target of miRNA-938, were 2.8 times lower in the resistant strains . The CpCPR5 protein plays an important role in the formation of the cuticle. It is postulated that the protein may render the cuticle permeable to deltamethrin . Likewise, miR-92a that targets CpCPR4, another cuticle protein, was found unregulated in deltamethrin resistant C. pipiens . Many other miRNAs are differentially expressed between deltamethrin sensitive and resistant C. pipiens strains. For instance, a 2014 study found that cpp-miR-71 is considerably under-expressed in the deltamethrin resistant strains. cpp-miR-7 targets CYP325BG3, a cytochrome P450 gene involved in detoxification . Lately, miR-278-3p  and miR-285  were implicated in pyrethroids resistance in C. pipiens via post-transcriptional regulation of CYP6AG11 and CYP6N23, respectively. Both these genes are also members of the cytochrome P450 family.
The Asian corn borer (Ostrinia furnacalis) is a pest of corn in South and South-East Asia. The pest has caused devastating loses in corn fields, such as eighty per cent in the Philippines, ninety-five per cent in Taiwan, and a hundred per cent in the Marianas . Asian corn borer feeds on almost all parts of the plant, especially kernels. Mostly, bio control methods, such as B. thuringiensis-based Cry toxins, are used to eradicate the pest. However, in recent decades, the pest developed strong resistance to them . Globally, Cry1Ab is the most widely commercialized Cry toxin for B. thuringiensis corn and Asian corn borer has developed a strong resistance to it. In a recent study, Xu et al found 22 miRNAs (21 known and 1 novel) that were differentially expressed in the resistant and sensitive pest strains. Half of the miRNAs were overexpressed in the resistant and half in the sensitive strains. The transcriptome profiling revealed that most of these miRNAs targeted B. thuringiensis toxin receptor genes, for example, amino peptidase N and cadherin-like protein. The other miRNAs targeted amino peptidases, chymotrypsin-like enzyme, alkaline phosphatases, ATP-binding cassette transporters, and trypsin-like enzyme. In the resistant strains, expression of amino peptidases 1 to 4 was unregulated and miRNAs targeting them, Ofu-miR-3851c–5p, Ofu-miR-963–3p, Ofu-miR-927–3p, and Ofu-miR-2731, was down regulated. Amino peptidases metabolize Cry toxins. Likewise, expression of trypsin-like serine protease and chymotrypsin-like protease was high in the resistant strains and Ofu-miR-6038 and Ofu-miR-3897–3p, which target them respectively, was low .
The project was funded by the Deanships of Scientific Research (DSR) at king Abdul-Aziz University, Jeddah under the grant no G-249-130-38. The authors, therefore acknowledge with thanks DSR for technical and financial support.
- Knudsen AB, Slooff R (1992) Vector-borne disease problems in rapid urbanization: New approaches to vector control. Bull World Health Organ 70(1): 1.
- Khasnis AA, Nettleman MD (2005) Global warming and infectious disease. Arch Med Res 36(6): 689-696.
- Tolle MA (2009) Mosquito-borne diseases. Curr Probl Pediatr Adolesc Health Care 39(4): 97-140.
- Kraemer MU, Sinka ME, Duda KA, Mylne AQ, Shearer FM, et al. (2015) The global distribution of the arbovirus vectors Aedes aegypti and Ae. albopictus. elife 4: e08347.
- Thomas SJ, Endy TP (2011) Critical issues in dengue vaccine development. Curr Opin Infect Dis 24(5): 442-450.
- Ishikawa T, Yamanaka A, Konishi E (2014) A review of successful flavivirus vaccines and the problems with those flaviviruses for which vaccines are not yet available. Vaccine 32(12): 1326-1337.
- Haug CJ, Kieny MP, Murgue B (2016) The zika challenge. N Engl J Med 374(19): 1801-1803.
- Christophers S (1960) Aedes aegypti (L.) the yellow fever mosquito: Its life history, bionomics and structure.
- Centers for Disease Control and Prevention (2019) Mosquito Life-Cycle. Available online at: https://www.cdc.gov/dengue/resources/factsheets/mosquitolifecyclefinal.pdf
- Rezende GL, Martins AJ, Gentile C, Farnesi LC, Pelajo-Machado M, et al. (2008) Embryonic desiccation resistance in Aedes aegypti: Presumptive role of the chitinized serosal cuticle. BMC Dev Biol 8(1): 82.
- Faull KJ, Williams CR (2015) Intraspecific variation in desiccation survival time of Aedes aegypti (L.) mosquito eggs of Australian origin. J Vector Ecol 40(2): 292-300.
- Barnard DR (2002) Chemical analysis of human skin emanations: Comparison of volatiles from humans that differ in attraction of Aedes aegypti (Diptera: Culicidae). J Am Mosq Control Assoc 8(3): 186-195.
- McBride CS, Baier F, Omondi AB, Spitzer SA, Lutomiah J, et al. (2014) Evolution of mosquito preference for humans linked to an odorant receptor. Nature 515(7526): 222.
- Qiu H, McCall JW, Jun HW (1998) Formulation of topical insect repellent N, N-diethyl-m-toluamide (DEET): Vehicle effects on DEET in vitro skin permeation. Int J Pharm 163(1-2): 167-176.
- Trigg J, Hill N (1996) Laboratory evaluation of a ‐based repellent against four biting arthropods. Phytother Res 10(4): 313-316.
- Robbins PJ, Cherniack MG (1986) Review of the biodistribution and toxicity of the insect repellent N, N‐diethyl‐m‐toluamide (DEET). J Toxicol Environ Health 18(4): 503-525.
- Tenenbein M (1987) Severe toxic reactions and death following the ingestion of diethyltoluamide-containing insect repellents. JAMA 258(11): 1509-1511.
- Betz FS, Hammond BG, Fuchs RL (2000) Safety and advantages of Bacillus thuringiensis-protected plants to control insect pests. Regul Toxicol Pharmacol 32(2): 156-173.
- BravoA, Gill SS, Soberon M (2007) Mode of action of Bacillus thuringiensis Cry and Cyt toxins and their potential for insect control. Toxicon 49(4): 423-435.
- Pardo-Lopez L, Soberon M, Bravo A (2013) Bacillus thuringiensis insecticidal three-domain Cry toxins: Mode of action, insect resistance and consequences for crop protection. FEMS Microbiol Rev 37(1): 3-22.
- Vilas-Bôas GT, Lemos MVF (2004) Diversity of cry genes and genetic characterization of Bacillus thuringiensis isolated from Brazil. Can J Microbiol 50(8): 605-613.
- Bian G, Xu Y, Lu P, Xie Y, Xi Z (2010) The endosymbiotic bacterium Wolbachia induces resistance to dengue virus in Aedes aegypti. PLoS Pathog 6(4): e1000833.
- Hoffmann AA, Ross PA, Rašić G (2015) Wolbachia strains for disease control: Ecological and evolutionary considerations. Evol Appl 8(8): 751-768.
- Dutra HLC, Rocha MN, Dias FBS, Mansur SB, Caragata EP, et al. (2016) Wolbachia blocks currently circulating Zika virus isolates in Brazilian Aedes aegypti Cell Host Microbe 19(6): 771-774.
- Dutra HLC, da Silva VL, da Rocha Fernandes M, Logullo C, Maciel-de-Freitas R, et al. (2016) The influence of larval competition on Brazilian Wolbachia-infected Aedes aegypti Parasit Vectors 9(1): 282.
- Hancock PA, White VL, Ritchie SA, Hoffmann AA, Godfray HCJ (2016) Predicting Wolbachia invasion dynamics in Aedes aegypti populations using models of density-dependent demographic traits. BMC Biol 14(1): 96.
- Lacroix R, McKemey AR, Raduan N, Wee LK, Ming WH, et al. (2012) Open field release of genetically engineered sterile male Aedes aegypti in Malaysia. PloS One 7(8): e42771.
- Carvalho DO, Nimmo D, Naish N, McKemey AR, Gray P, et al. (2014) Mass production of genetically modified Aedes aegypti for field releases in Brazil. J Vis Exp 83: e3579.
- Carvalho DO, McKemey AR, Garziera L, Lacroix R, Donnelly CA, et al. (2015) Suppression of a field population of Aedes aegypti in Brazil by sustained release of transgenic male mosquitoes. PLoS Negl Trop Dis 9(7): e0003864.
- Feng X, Zhou S, Wang J, Hu W (2018) microRNA profiles and functions in mosquitoes. PLoS Negl Trop Dis 12(5): e0006463.
- Etebari K, Furlong MJ, Asgari S (2015) Genome wide discovery of long intergenic non-coding RNAs in Diamondback moth (Plutella xylostella) and their expression in insecticide resistant strains. Sci Rep 5: 14642.
- Zhu B, Li X, Liu Y, Gao X, Liang P (2017) Global identification of microRNAs associated with chlorantraniliprole resistance in diamondback moth Plutella xylostella (L.) Sci Rep 7: 40713.
- Laws Jr, ER (2013) Classes of pesticides, Elsevier.
- Costa LG (1997) Basic toxicology of pesticides. Occup Med 12(2): 251-268.
- Coats JR (1990) Mechanisms of toxic action and structure-activity relationships for organochlorine and synthetic pyrethroid insecticides. Environ Health Perspect 87: 255-262.
- Holan G (1969) New halo cyclopropane insecticides and the mode of action of DDT. Nature 221(5185): 1025.
- Kaushik P, Kaushik G (2007) An assessment of structure and toxicity correlation in organochlorine pesticides. J Hazard Mater 143(1-2): 102-111.
- Bajgar J (2004) Organophosphates/nerve agent poisoning: Mechanism of action, diagnosis, prophylaxis, and treatment. Adv Clin Chem 38(1): 151-216.
- Main A (1979) Mode of action of anticholinesterases. Pharmacol Ther 6(3): 579-628.
- O'brien R (1963) Mode of action of insecticides, binding of organophosphates to cholinesterases. J Agric Food Chem 11(2): 163-166.
- Knowles C, Casida J (1966) Mode of action of organophosphate anthelmintics. Cholinesterase inhibition in Ascaris lumbricoides. J Agric Food Chem 14(6): 566-572.
- Eldefrawi M, Britten A, O'Brien R (1971) Action of organophosphates on binding of cholinergic ligands. Pestic Biochem Physiol 1(1): 101-108.
- Kuhr RJ, Dorough HW (1976) Carbamate insecticides: Chemistry, biochemistry, and toxicology, CRC Press, Inc.
- Fukuto TR (1990) Mechanism of action of organophosphorus and carbamate insecticides. Environ Health Perspect 87: 245-254.
- Elliott M, Janes N (1978) Synthetic pyrethroids-a new class of insecticide. Chem Soc Rev 7(4): 473-505.
- Casida JE, Gammon DW, Glickman AH, Lawrence LJ (1983) Mechanisms of selective action of pyrethroid insecticides. Ann Rev Pharmacol Toxicol 23(1): 413-438.
- Matsuda K, Buckingham SD, Kleier D, Rauh JJ, Grauso M, et al. (2001) Neonicotinoids: Insecticides acting on insect nicotinic acetylcholine receptors. Trends Pharmacol Sci 22(11): 573-580.
- Tomizawa M, Casida JE (2005) Neonicotinoid insecticide toxicology: Mechanisms of selective action. Ann Rev Pharmacol Toxicol 45: 247-268.
- Sargent PB (1993) The diversity of neuronal nicotinic acetylcholine receptors. Ann Rev Neurosci 16(1): 403-443.
- Tomizawa M, Casida JE (2001) Structure and diversity of insect nicotinic acetylcholine receptors. Pest Manag Sci 57(10): 914-922.
- Sattelle DB, Buckingham SD, Wafford K, Sherby S, Bakry N, et al. (1989) Actions of the insecticide 2 (nitromethylene) tetrahydro-1, 3-thiazine on insect and vertebrate nicotinic acetylcholine receptors. Proc Royal Soc B Biol Sci 237(1289): 501-514.
- Jefferies PR, Casida JE (1994) Ryanoid chemistry and action. ACS symposium series (USA).
- Brugger KE, Cole PG, Newman IC, Parker N, Scholz B, et al. (2010) Selectivity of chlorantraniliprole to parasitoid wasps. Pest Manag Sci 66(10): 1075-1081.
- Jefferies PR, Toia RF, Brannigan B, Pessah I, Casida JE (1992) Ryania insecticide: Analysis and biological activity of 10 natural ryanoids. J Agric Food Chem 40(1): 142-146.
- Kadous A, Ghiasuddin S, Matsumura F, Scott J, Tanaka K (1983) Difference in the picrotoxinin receptor between the cyclodiene-resistant and susceptible strains of the German cockroach. Pestic Biochem Physiol 19(2): 157-166.
- Hemingway J, Ranson H (2000) Insecticide resistance in insect vectors of human disease. Ann Rev Entomol 45(1): 371-391.
- Vaughan A, Hemingway J (1995) Mosquito carboxylesterase Estα21 (A2). Cloning and sequence of the full-length cDNA for a major insecticide resistance gene worldwide in the mosquito Culex quinquefasciatus. J Biol Chem 270(28): 17044-17049.
- Guillemaud T, Makate N, Raymond M, Hirst B, Callaghan A (1997) Esterase gene amplification in Culex pipiens. Insect Mol Biol 6(4): 319-328.
- Karunaratne SP, Hemingway J, Jayawardena KG, Dassanayaka V, Vaughan A (1995) Kinetic and molecular differences in the amplified and non-amplified esterases from insecticide-resistant and susceptible Culex quinquefasciatus mosquitoes. J Biol Chem 270(52): 31124-31128.
- Grant DF (1991) Evolution of glutathione S-transferase subunits in Culicidae and related Nematocera: Electrophoretic and immunological evidence for conserved enzyme structure and expression. Insect Biochem 21(4): 435-445.
- Grant DF, Dietze EC, Hammock BD (1991) Glutathione S-transferase isozymes in Aedes aegypti: Purification, characterization, and isozyme-specific regulation. Insect Biochem 21(4): 421-433.
- Prapanthadara LA, Koottathep S, Promtet N, Hemingway J, Ketterman AJ (1996) Purification and characterization of a major glutathione S-transferase from the mosquito Anopheles dirus (species B). Insect Biochem Mol Biol 26(3): 277-285.
- Grant DF, Hammock BD (1992) Genetic and molecular evidence for a trans-acting regulatory locus controlling glutathione S-transferase-2 expression in Aedes aegypti. Mol Gen Genet 234(2): 169-176.
- Hemingway J, Miyamoto J, Herath P (1991) A possible novel link between organophosphorus and DDT insecticide resistance genes in Anopheles: Supporting evidence from fenitrothion metabolism studies. Pestic Biochem Physiol 39(1): 49-56.
- Vulule J, Beach R, Atieli F, Roberts J, Mount D, et al. (1994) Reduced susceptibility of Anopheles gambiae to permethrin associated with the use of permethrin‐impregnated bed nets and curtains in Kenya. Med Vet Entomol 8(1): 71-75.
- Kasai S, Weerashinghe IS, Shono T (1998) P450 monooxygenases are an important mechanism of permethrin resistance in Culex quinquefasciatus Say larvae. Arch Insect Biochem Physiol 37(1): 47-56.
- Ayad H, Georghiou GP (1975) Resistance to organophosphates and carbamates in Anopheles albimanus based on reduced sensitivity of acetylcholinesterase. J Econ Entomol 68(3): 295-297.
- Hemingway J, Georghiou GP (1983) Studies on the acetylcholinesterase of Anopheles albimanus resistant and susceptible to organophosphate and carbamate insecticides. Pestic Biochem Physiol 19(2): 167-171.
- Williamson MS, Denholm I, Bell CA, Devonshire AL (1993) Knockdown resistance (kdr) to DDT and pyrethroid insecticides maps to a sodium channel gene locus in the housefly (Musca domestica). Mol Gen Genet 240(1): 17-22.
- Williamson MS, Martinez-Torres D, Hick CA, Devonshire AL (1996) Identification of mutations in the housefly para-type sodium channel gene associated with knockdown resistance (kdr) to pyrethroid insecticides. Mol Gen Genet 252(1-2): 51-60.
- Newcomb R, East P, Russell R, Oakeshott J (1996) Isolation of a cluster esterase genes associated with organophosphate resistance in Lucilia cuprina. Insect Mol Biol 5(3): 211-216.
- Newcomb RD, Campbell PM, Russell RJ, Oakeshott JG (1997) cDNA cloning, baculovirus-expression and kinetic properties of the esterase, E3, involved in organophosphorus resistance in Lucilia cuprina. Insect Biochem Mol Biol 27(1): 15-25.
- Thompson M, Shotkoski F, ffrench-Constant R (1993) Cloning and sequencing of the cyclodiene insecticide resistance gene from the yellow fever mosquito Aedes aegypti: Conservation of the gene and resistance associated mutation with Drosophila. FEBS Lett 325(3): 187-190.
- Zhang W, Jiang F, Ou J (2011) Global pesticide consumption and pollution: With China as a focus. Proc Int Acad Ecol Environ Sci 1(2): 125.
- Lee WJ, Cha ES, Park ES, Kong KA, Yi JH, et al. (2009) Deaths from pesticide poisoning in South Korea: Trends over 10 years. Int Arch Occup Environ Health 82(3): 365-371.
- Langley RL, Mort SA (2012) Human exposures to pesticides in the United States. J Agromedicine 17(3): 300-315.
- Wang X, Xue Y, Gong P, Yao T (2014) Organochlorine pesticides and polychlorinated biphenyls in Tibetan Forest soil: Profile distribution and processes. Environ Sci Pollut Res 21(3): 1897-1904.
- Zaki MH, Moran D, Harris D (1982) Pesticides in groundwater: The aldicarb story in Suffolk County, NY. Am J Public Health 72(12): 1391-1395.
- Rattner BA (2009) History of wildlife toxicology. Ecotoxicology 18(7): 773-783.
- Fleischli MA, Franson J, Thomas N, Finley D, Riley W (2004) Avian mortality events in the United States caused by anticholinesterase pesticides: A retrospective summary of National Wildlife Health Center records from 1980 to 2000. Arch Environ Contam Toxicol 46(4): 542-550.
- Turusov V, Rakitsky V, Tomatis L (2002) Dichlorodiphenyltrichloroethane (DDT): Ubiquity, persistence, and risks. Environ Health Perspect 110(2): 125-128.
- Galloway T, Handy R (2003) Immunotoxicity of organophosphorus pesticides. Ecotoxicology 12(1-4): 345-363.
- Galloway TS, Depledge MH (2001) Immunotoxicity in invertebrates: Measurement and ecotoxicological relevance. Ecotoxicology 10(1): 5-23.
- Lin PC, Lin HJ, Liao YY, Guo HR, Chen KT (2013) Acute poisoning with neonicotinoid insecticides: A case report and literature review. Basic Clin Pharmacol Toxicol 112(4): 282-286.
- Gill RJ, Ramos-Rodriguez O, Raine NE (2012) Combined pesticide exposure severely affects individual-and colony-level traits in bees. Nature 491(7422): 105.
- Henry M, Beguin M, Requier F, Rollin O, Odoux JF, et al. (2012) A common pesticide decreases foraging success and survival in honey bees. Science 336(6079): 348-350.
- Longnecker MP, Rogan WJ, Lucier G (1997) The human health effects of DDT (dichlorodiphenyltrichloroethane) and PCBS (polychlorinated biphenyls) and an overview of organochlorines in public health. Ann Rev Public Health 18(1): 211-244.
- Peiris-John RJ, Wickremasinghe R (2008) Impact of low-level exposure to organophosphates on human reproduction and survival. Trans Royal Soc Trop Med Hyg 102(3): 239-245.
- Engel SM, Wetmur J, Chen J, Zhu C, Barr DB, et al. (2011) Prenatal exposure to organophosphates, paraoxonase 1, and cognitive development in childhood. Environ Health Perspect 119(8): 1182-1188.
- Aldridge W (1990) An assessment of the toxicological properties of pyrethroids and their neurotoxicity. Crit Rev Toxicol 21(2): 89-104.
- Han W, Tian Y, Shen X (2018) Human exposure to neonicotinoid insecticides and the evaluation of their potential toxicity: An overview. Chemosphere 192: 59-65.
- Ambros V (2004) The functions of animal microRNAs. Nature 431(7006): 350.
- Bartel D P (2004) MicroRNAs: Genomics, biogenesis, mechanism, and function. Cell 116(2): 281-297.
- Bartel D P (2009) MicroRNAs: Target recognition and regulatory functions. Cell 136(2): 215-233.
- Fabian MR, Sonenberg N, Filipowicz W (2010) Regulation of mRNA translation and stability by microRNAs. Ann Rev Biochem 79: 351-379.
- Vasudevan S, Tong Y, Steitz JA (2007) Switching from repression to activation: microRNAs can up-regulate translation. Science 318(5858): 1931-1934.
- Ørom UA, Nielsen FC, Lund AH (2008) MicroRNA-10a binds the 5′ UTR of ribosomal protein mRNAs and enhances their translation. Mol Cell 30(4): 460-471.
- Vasudevan S (2012) Posttranscriptional upregulation by microRNAs. Wiley Interdiscip Rev RNA 3(3): 311-330.
- Orang AV, Safaralizadeh R, Kazemzadeh-Bavili M (2014) Mechanisms of miRNA-mediated gene regulation from common downregulation to mRNA-specific upregulation. Int J Genomics 2014: 1-15.
- Seong KM, Coates BS, Kim DH, Hansen AK, Pittendrigh BR (2018) Differentially expressed microRNAs associated with changes of transcript levels in detoxification pathways and DDT-resistance in the Drosophila melanogaster strain 91-R. PLoS One 13(4): e0196518.
- Lümmen P, Khajehali J, Luther K, Van Leeuwen T (2014) The cyclic keto-enol insecticide spirotetramat inhibits insect and spider mite acetyl-CoA carboxylases by interfering with the carboxyltransferase partial reaction. Insect Biochem Mol Biol 55: 1-8.
- Wei X, Zheng C, Peng T, Pan Y, Xi J, et al. (2016) miR-276 and miR-3016-modulated expression of acetyl-CoA carboxylase accounts for spirotetramat resistance in Aphis gossypii Glover. Insect Biochem Mol Biol 79: 57-65.
- Furlong MJ, Wright DJ, Dosdall LM (2013) Diamondback moth ecology and management: Problems, progress, and prospects. Ann Rev Entomol 58: 517-541.
- Pan Y, Peng T, Gao X, Zhang L, Yang C, et al. (2015) Transcriptomic comparison of thiamethoxam-resistance adaptation in resistant and susceptible strains of Aphis gossypii Glover. Comp Biochem Physiol Part D Genomics Proteomics 13: 10-15.
- Ikeda T, Zhao X, Kono Y, Yeh JZ, Narahashi T (2003) Fipronil modulation of glutamate-induced chloride currents in cockroach thoracic ganglion neurons. Neurotoxicology 24(6): 807-815.
- Bariami V, Jones CM, Poupardin R, Vontas J, Ranson H (2012) Gene amplification, ABC transporters and cytochrome P450s: Unraveling the molecular basis of pyrethroid resistance in the dengue vector, Aedes aegypti. PLoS Negl Trop Dis 6(6): e1692.
- Hsu PK, Huang LH, Geib SM, Hsu J-C (2016) Identification of a carboxylesterase associated with resistance to naled in Bactrocera dorsalis (Hendel). Pestic Biochem Physiol 131: 24-31.
- Etebari K, Afrad M, Tang B, Silva R, Furlong M, et al. (2018) Involvement of microRNA miR‐2b‐3p in regulation of metabolic resistance to insecticides in Plutella xylostella. Insect Mol Biol 27(4): 478-491.
- Liu B, Tian M, Guo Q, Ma L, Zhou D, et al. (2016) MiR-932 regulates pyrethroid resistance in Culex pipiens pallens (Diptera: Culicidae). J Med Entomol 53(5): 1205-1210.
- Fang F, Wang W, Zhang D, Lv Y, Zhou D, et al. (2015) The cuticle proteins: A putative role for deltamethrin resistance in Culex pipiens pallens. Parasitol Res 114(12): 4421-4429.
- Ma K, Li X, Hu H, Zhou D, Sun Y, et al. (2017) Pyrethroid-resistance is modulated by miR-92a by targeting CpCPR4 in Culex pipiens pallens. Comp Biochem Physiol Part B Biochem Mol Biol 203: 20-24.
- Hong S, Guo Q, Wang W, Hu S, Fang F, et al. (2014) Identification of differentially expressed microRNAs in Culex pipiens and their potential roles in pyrethroid resistance. Insect Biochem Mol Biol 55: 39-50.
- Lei Z, Lv Y, Wang W, Guo Q, Zou F, et al. (2015) MiR-278-3p regulates pyrethroid resistance in Culex pipiens pallens. Parasitol Res 114(2): 699-706.
- Tian M, Liu B, Hu H, Li X, Guo Q, et al. (2016) MiR-285 targets P450 (CYP6N23) to regulate pyrethroid resistance in Culex pipiens pallens. Parasitol Res 115(12): 4511-4517.
- Nafus D, Schreiner I (1991) Review of the biology and control of the Asian corn borer, Ostrinia furnacalis (Lep: Pyralidae). Int J Pest Manag 37(1): 41-56.
- Xu L, Wang Z, Zhang J, He K, Ferry N, et al. (2010) Cross‐resistance of Cry1Ab‐selected Asian corn borer to other Cry toxins. J Appl Entomol 134(5): 429-438.
- Xu L-N, Ling Y-H, Wang Y-Q, Wang Z-Y, Hu B-J, et al. (2015) Identification of differentially expressed microRNAs between Bacillus thuringiensis Cry1Ab-resistant and-susceptible strains of Ostrinia furnacalis. Sci Rep 5: 15461.
- International Journal of Medical and Clinical Imaging (ISSN:2573-1084)
- Journal of Infectious Diseases and Research (ISSN: 2688-6537)
- Journal of Neurosurgery Imaging and Techniques (ISSN:2473-1943)
- International Journal of Internal Medicine and Geriatrics (ISSN: 2689-7687)
- Journal of Pathology and Toxicology Research
- Journal of Oral Health and Dentistry (ISSN:2638-499X)
- Journal of Cancer Science and Treatment (ISSN:2641-7472)