Entomology, Ornithology & Herpetology: Current Research
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Fundamental Biological Mechanism and Resistance of Insect Repellent which Make Worse the Liability of Malaria in Emerging Nations

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Introduction

Malaria is a major health concern around the world has a profound socio-economic where it’s endemic [1]. In 2015, an estimated 429,000 deaths occurred through malaria to children below the age of 5 in Africa. Furthermore, an estimated 92% of malaria deaths occurred in Africa [2]. When taking the history of the dry zone of Sri Lanka, malaria was a leading cause of morbidity and mortality [3]. Yet, since October 2012 there has been no case of indigenous malaria reported in Sri Lanka. As a result, in 2016 Sri Lanka has been certified by the WHO as a country eliminated from malaria which was a life-threatening disease [2]. Furthermore, it is of vital importance for research to continue and to understand the impact of its problems. Also, find ways to manage the growth in insecticides resistance. This review showed the biological mechanism of insecticides that control malaria. For Indoor Residual Spraying (IRS), several strategies have proposed which might prevent or slow down resistance. Vector control remains one of the central components for malaria control through larval source reduction and adult vector control [4].

Insecticides

Insecticide resistance in pest populations affects both the economy and public health at a worldwide scale: it decreases crop yields (and thus profitability), induces the need to increase the quantity of insecticide and to develop new insecticides (thereby having a strong impact on costs and the environment), and finally it is responsible for an incidence of human or animal diseases [5,6]. Resistance is defined as a heritable decrease of the susceptibility to an insecticide [7].

Three categories of resistance can be distinguished: behavioral (avoidance of contact with insecticide), physiological (e.g., increased cuticle thickness), and biochemical (enhanced insecticide detoxification and sequestration and/or decreased insecticide target sensitivity). Few examples of behaviors such as Anopheles gambiae on Bioko Island and Senegal [8,9] or Anopheles funestus in Benin and Tanzania [10] and physiological resistances have been reported; whether they are heritable remains debated, and it is difficult to assess the level of protection they provide [11].

Biochemical resistances typically result in a relatively high level of protection and are genetically determined. Resistant individuals carry one or several genetic mutations that prevent insecticide disruption of the target functioning. As a result, the frequency of resistance gene(s)/allele(s) increases in the population over time. Insecticide resistance is confirmed by toxicological tests (bioassays) establishing resistance ratio (or RR corresponding to the number by which an insecticide dose must be multiplied to obtain the same mortality in resistant than in susceptible insects). It can be investigated at many levels, from the molecular characterization of genes/alleles conferring resistance and their biochemical products to the effect of these genes on the fitness (i.e., mean reproductive success) of the individuals carrying resistance alleles, to the dynamics and evolution of these resistance alleles in natural vector populations and their effect on disease control [12].

The first case of resistance has reported in 1908, in a population of San Jose scale (Aspidiotusperniciosus) resistant to lime sulfur [13]. A century later (2007), 553 arthropod species have reported as resistant to at least one insecticide, among many disease vectors. More than 100 mosquito species are resistant to at least one insecticide (including 56 Anopheles species, 39 Culicine species), Culexpipienspipiens and Anopheles albimanus are resistant to more than 30 different compounds [6].

Synthetic insecticides

Four major classes of organic (synthetic) insecticides are used to control insects: the organ chlorines (OCs), the organophosphates (OPs), the carbamates (CXs), and the pyrethroids (PYRs), with, 4429, 1375, 30, and 414 metric tons respectively, of active ingredient used annually for global vector control from 2000 to 2009 [14]. The use of synthetic insecticides was started back in 1943 for control of malaria. The first synthetic insecticide was DichloroDiphenylTrichloroethane (DDT) and later cyclodiene (CD) dieldrin was started to use. [15-19]. From the late 1970s, the PYRs class of vector control replaced OCs, and these became widely used in malaria vector control. They are today by far the most-used insecticides, with 81% of the World spray coverage [14]. PYR-based Indoor Residual Spraying (IRS) and Insecticide-Treated Nets and curtains (ITNs) are currently advocated as standard malaria vector control strategies. Finally, two other classes of synthetic insecticides are used at a large scale worldwide: the OPs and the CXs, which were first used in the 1940s and the 1950s, respectively [7,15].

They are usually used as larvicides (although some are now considered for ITN impregnation and IRS as an alternative to PYR, and are particularly well suited for species with delimited breeding sites [20]. OCs and PYRs are highly popular due to their very low toxicity to human and long half-life in the environment, which makes it more cost effective in vector control. On the other hand, OPs and CXs have a short half-life; and two to three rounds of IRS are needed per year. This significantly reduces the cost-effectiveness of the use of these two classes of insecticides. However, when compared to OCs and PYRs these two groups evidently have less potential of developing resistance [21]. Furthermore, the latter two groups have less environmental impacts due to their lower half-life in the environment than OCs and PYRs. Some new groups of insecticides such as neonicotinoids, phthalic acid diamides, or anthranilic acid diamides were introduced in the 2006-2008 period. However, these groups did not become much popular in disease vector control despite their popularity in agricultural pest control [6,22].

During the same period, another group of synthetic insecticides was introduced. These interfere with insect physiological processes by mimicking certain compounds produced in the endocrine system. Synthetic Insect Growth Regulators (IGR) is one of the best examples of this. Furthermore, synthetic products called juvenoids [23] that mimic the juvenile hormone (JH) and chitine inhibitors [24] are also some examples for this.

Mechanisms of resistance

The targets of most insecticides are critical proteins of the insect nervous system. Insecticides bind to specific sites on their targets and disrupt their function. Any mechanism that decreases the insecticide effect will lead to resistance. This encompasses reduced penetration of the insecticide, increased excretion or sequestration of the insecticide, increased metabolism of the insecticide, and finally target modification that limits the binding of the insecticide [8,11]. The first three mechanisms are poorly documented and do not seem to play a prominent role in resistance [25].

Metabolic resistance

Metabolic resistance includes the various mechanisms that lead to the degradation of the insecticide in less or nontoxic products, thus decreasing the number of toxic molecules that reach the target. The detoxification through enzymatic reactions is one of the major ways of metabolic resistance. There are several groups of enzymes involved in this type of resistance mechanisms. For instance, Glutathione S-Transferases (GSTs), carboxylesterases (COEs) and cytochrome P450 monooxygenases (P450s) are three of the major groups [26]. Among these GSTs catalyze the reaction between sulfhydryl group and electrophilic sites of xenobiotics and form conjugates that are more readily excreted and typically less toxic than the parent insecticide and this group associated with resistance to OCs, particularly DDT, and OPs COEs on the other hand, act by binding to an ester group on the xenobiotic molecule and then break the ester bound by a process of acylation, de-acylation [18,27]. The majority of insecticides, including almost all CXs and OPs, most PYRs, and some IGRs bear ester groups; hence resistance may develop due to the action of COEs [Pasteur N., 1984]. The other group, P450s involved in detoxification through monooxygenase activity and is responsible for the resistance to several groups of insecticides, particularly DDT, PYRs, and CXs [19,28].

Target-site modification

Resistance by target-site modification is due to point mutations in the insecticide target gene that results in reduced binding of insecticides. For an instance resistance to CDs may develop due to a decreased sensitivity to insecticide of the GABA receptor A, through a point mutation causing an amino acid change in the receptor-coding gene. The extensive use of CDs before their banning in the 1980s resulted in resistance in several insect species [29,30].

Voltage-gated sodium channels (vgscs)

VGSCs are glycoproteins with a pore for ion transport and can adopt three different states: resting, open, or inactivated; the Na + ions pass only when the channels are open [31]. VGSC are the targets of DDT and PYRs. When these insecticides bind to the VGSC, they slow their closing speed, prolonging the depolarization [32]. One major mechanism, named knockdown resistance (kdr), is responsible for PYR and DDT resistance, by reducing the sensitivity of the receptors (binding capacity) to these insecticides and modifying the action potential of the channel [23,32,33].

Acetylcholinesterase (Ache)

AChE is the target of OPs and CXs insecticides, which are competitive inhibitors of AChE when they bind to AChE; their very slow release prevents hydrolysis of the natural substrate. Consequently, AChE remains active in the synaptic cleft and the nervous influx is continued, leading to insect death by tetany. In most insects, there are two genes, ace-1 and ace-2, coding for AChE1 and AChE2, respectively. In these species, AChE1 is the main synaptic enzyme while the physiological role of AChE2 is still uncertain. Diptera of the Cyclorrapha group or “true” flies (such as D. melanogaster and M. domestica) possess a single AChE, which is encoded by the ace-2 gene and is the synaptic enzyme in that case. In mosquitoes where AChE1 is the synaptic enzyme, the most common resistance mutation (G119S) in the ace-1 gene is located just near the active site [34-36].

Other resistance mechanisms

Growth regulators: Juvenoids mimic JH and disrupt insect development. Few resistance cases have been described in various species. High resistance to methoprene has been described in the mosquito Ochlerotatus nigromaculis in California, potentially through target-site mutation [37]. While, 7.7-fold resistance have been reported to the same insecticide in C. pipiens from New York [38].

Toxin receptors: Bacillus thuringiensis (Bt) toxins have a complex mode of action not clearly understood. Bt resistance is increasing in the field in several pests [39]. Presently, the only report of field resistance in the mosquito is a 33-fold resistance to Bti (Bacillus thuringiensis var. israelensis) the only Bt variety active on mosquitoes) detected in a natural population of C. p. pipiens from New York. However, the mechanism of this resistance was not investigated [38]. Genomic studies suggested several candidates for Bti resistance in Ae. Aegypti, but they are not yet validated [28] finally; it appears that depending on the environmental conditions, some of the four Bti toxins may be inactivated, which could favor the emergence of full Bti resistance through intermediate bouts of selection to each toxin independently [40]. For Bacillus sphaericus (Bs) toxins, resistance has been described essentially in mosquitoes of the C. p. pipiens complex, due to a mutation in the toxin receptor. It developed very rapidly within the first year of treatment in India [41] and in Tunisia [42]. Similarly, control using Bs toxins started in the early 1990s in Southern France and first failure was reported in 1994 in Port-Louis (near Marseille). This resistance was due to a recessive sex-linked gene, named sp-1. In 1996, Bs resistance was reported close to the Spain border it was due to a second gene, sp-2, which was recessive and sex-linked [43]. Now Bs resistance has been observed worldwide in the C. p. pipiens complex [41]. Two of the alleles identified (sp-2R and an allele selected in a laboratory strain from California [44] change the toxin receptor binding properties and were found to be due to “ stop ” mutations or mobile element insertion in the toxin receptor [45,46].

Conclusion

Vector control remains a powerful and accessible tool to control these diseases and upgrade the socioeconomic burden they cause in developing countries. However, any disease control studies taken can influence its success (vector control failures) and as a result, may have a direct effect on pathogen transmission. This includes first establishing a continuous survey of resistance at a local scale by implicating the local population, a difficult but essential task to set goals and evaluate success. Several survey sites in different conditions are required for sentinel purposes, together with some baseline information, to rapidly detect resistance, identify the mechanisms, and change the policies adequately. These local surveys should then be integrated at a more global scale for vector control coordination, allowing informed decisions for using alternative tools to insecticides and preserving the remaining insecticides by carefully planning their use to minimize resistance selection.

Variation in insecticide resistance mainly depends upon the type of insecticide and frequency of use. Although various mechanisms of insecticide resistance in insects such as metabolic resistance (i.e. esterases, monooxygenase or glutathione-stransferase), resistance due to reduced penetration or behavioral resistance are reported in several vectors, generally it is governed by either involvement of metabolic mechanisms or alterations at target sites. Revealing the mechanism of resistance is equally important to that of monitoring resistance in mosquito vectors. An Anopheles species are highly resistant to DDT. Insecticide resistance is a serious emerging problem in Developing countries.

Currently, the national program has no alternative insecticide for effective vector control or for insecticide resistance management. There are some insecticides available for vector control, an approach focused on the rotational use of insecticides or a mosaic strategy can be adapted to delay development of resistance in malaria vectors. Also, highlighting needs to be given to other eco-friendly methods of vector control, such as bio control with larvivorous fish and biolarvicides especially Bacillus thuringiensis var. israelensis included in the integrated vector management program. Effective resistance management mainly depends upon early detection of the status of resistance, therefore monitoring of insecticide resistance at regular intervals is necessary so that an effective management strategy can be designed.

Consent for Publication

We certify this manuscript has not been published elsewhere and is not submitted to another Journal.

Competing Interests

The author(s) declare that they have no competing interests.

Authors Contribution

LS and AD conceived the study design, interpreted and manuscript preparation. SMT and MS are drafting of the article details with an edit the manuscripts. All authors read and approved the final manuscript.

Acknowledgement

I wish to express my gratitude to Ms. S. Amarakoon from Department of Animal Science, Faculty of Agricultural & Food Sciences, University of Manitoba,Winnipeg, Canada for her valuable comments on the manuscript.

References

1. Gallup JL, Sachs JD. The economic burden of malaria. Am J TropHedHyg. 2001;64(suppl): 85-96.
2. GunathilakaN, Denipitiya T, Hapugoda M, Abeyewickreme W, Wickremasinghe R. Determination of the foraging behaviour and blood meal source of malaria vector mosquitoes in Trincomalee District of Sri Lanka using a multiplex real time polymerase chain reaction assay. Malar J. 2016;15(1): 242.
3. Greenwood B. Can malaria be eliminated? Trans R Soc Trop Med Hyg. 2009;103(Suppl 1):S2-S5.
4. Georghiou GP, Lagunes-Tejeda A. The occurrence of resistance to pesticides in arthropods. 1991.
5. Whalon ME, Mota-Sanchez D, Hollingworth RM (Eds) Global pesticide resistance in arthropods. Oxford University Press, Oxford, UK. 2008;169.
6. Nauen R. Insecticide resistance in disease vectors of public health importance. Pest Manag Sci. 2007; 63(7):628-633.
7. Reddy MR, Overgaard HJ, Abaga S, Reddy VP, Caccone A, Kiszewski AE, et al. Outdoor host seeking behaviour of Anopheles gambiae mosquitoes following initiation of malaria vector control on Bioko Island, Equatorial Guinea. Malar J.2011;10(1):184.
8. Ndiath MO, MazenotC, Sokhna C, Trape JF. How the malaria vector Anopheles gambiae adapts to the use of insecticide-treated nets by African populations. PloS One. 2014; 9(6):e97700.
9. Moiroux N, Gomez MB, Pennetier C, Elanga E, Djènontin A, Chandre F, et al. Changes in Anopheles funestus biting behavior following universal coverage of long-lasting insecticidal nets in Benin. J Infect Dis. 2012;206(10):1622-1629.
10. Sougoufara S, Diédhiou SM, Doucouré S, Diagne N, Sembène PM, Harry M, et al. Biting by Anopheles funestus in broad daylight after use of long-lasting insecticidal nets: a new challenge to malaria elimination. MalarJ. 2014;13(1):125.
11. Wood RJ. In: Bishop JA, Cook LM (eds). Insecticide resistance: genes and mechanisms. Academic Press, London, UK. 1981;53e96.
12. Melander AL. Can insects become resistant to sprays. J Econ Entomol. 1914;7(1):167.
13. World Health Organization. Global insecticide use for vector-borne disease control: a 10-year assessment [2000-2009] (5th edn). 2011. https://apps.who.int/iris/handle/10665/44670.
14. World Health Organization. Pesticides and their Application for the Control of Vectors and Pests of Public Health Importance (6th edn). 2006. https://apps.who.int/iris/handle/10665/69223.
15. Roberts DR, Andre RG. Insecticide resistance issues in vector-borne disease control. Am J Trop Med Hyg. 1994;50(Suppl 6):21-34.
16. Hemingway J, Ranson H. Insecticide resistance in insect vectors of human disease. AnnuRev Entomol. 2000;45(1):371-391.
17. Oakeshott JG, Devonshire AL, Claudianos C, Sutherland TD, Horne I, Campbell PM, et al. Comparing the organophosphorus and carbamate insecticide resistance mutations in cholin-and carboxyl-esterases. Chem-Biol Interact. 2005;157:269-275.
18. Feyereisen R, Dermauw W, Van Leeuwen T. Genotype to phenotype, the molecular and physiological dimensions of resistance in arthropods. PesticBiochem Physiol. 2015;121:61-77.
19. Oxborough RM, Mosha FW, Matowo J, Mndeme R, Feston E, Hemingway J, et al. Mosquitoes and bednets: testing the spatial positioning of insecticide on nets and the rationale behind combination insecticide treatments. Ann Trop Med Parasitol. 2008;102(8):717-727.
20. Coleman M, Casimiro S, Hemingway J, Sharp B. Operational impact of DDT reintroduction for malaria control on Anopheles arabiensis in Mozambique. J Med Entomol. 2008;45(5):885-890.
21. Nauen R. Insecticide mode of action: return of the ryanodine receptor. Pest Manag Sci.2006;62(8):690-692.
22. Hollingworth RM, Dong K. The biochemical and molecular genetic basis of resistance to pesticides in arthropods. In: Whalon ME, Mota-Sanchez D (Eds). Global pesticide resistance in arthropods.CABI,Wallingford, UK. 2008:40.
23. Hirose T, Sunazuka T, ŌmuraS. Recent development of two chitinase inhibitors, Argifin and Argadin, produced by soil microorganisms. ProcJpnAcadSer B PhysBiol Sci. 2010:86(2):85-102.
24. Nkya TE, Akhouayri I, Kisinza W, David JP. Impact of environment on mosquito response to pyrethroid insecticides: facts, evidences and prospects. Insect BiochemMolec.2013;43(4):407-416.
25. Strode C, Wondji CS, David JP, Hawkes NJ, Lumjuan N, Nelson DR, et al. Genomic analysis of detoxification genes in the mosquito Aedesaegypti. Insect BiochemMolec. 2008;38(1):113-123.
26. Brooke BD, KlokeG, Hunt RH, Koekemoer LL, Tem EA, Taylor ME, et al. Bioassay and biochemical analyses of insecticide resistance in southern African Anopheles funestus (Diptera: Culicidae). BullEntomol Res. 2001;91(4):265-272.
27. Despres L, David JP, Gallet C. The evolutionary ecology of insect resistance to plant chemicals. Trends EcolEvol. 2007; 22(6):298-307.
28. Hemingway J, Hawkes NJ, McCarroll L, RansonH. The molecular basis of insecticide resistance in mosquitoes. Insect BiochemMol Biol. 2004;34:653-665.
29. ffrench-Constant RH, Anthony N, Aronstein K, Rocheleau T, Stilwell G. Cyclodiene insecticide resistance: from molecular to population genetics. AnnuRevEntomol. 2000;45(1):449-466.
30. Lund AE. Pyrethroid modification of sodium channel: current concepts. PesticBiochem Physiol. 1984;22(2):161-168.
31. Soderlund DM, Knipple DC. The molecular biology of knockdown resistance to pyrethroid insecticides. Insect BiochemMol Biol. 2003;33(6):563-577.
32. Dong K, Du Y, Rinkevich F, Nomura Y, Xu P, Wang L, et al. Molecular biology of insect sodium channels and pyrethroid resistance. Insect BiochemMol Biol. 2014;50:1-17.
33. Weill M, Lutfalla G, Mogensen K, Chandre F, Berthomieu A, Berticat C, et al. Comparative genomics: Insecticide resistance in mosquito vectors. Nature. 2003;423(6936):136.
34. Weill M, Berthomieu A, Berticat C, Lutfalla G, Nègre V, Pasteur N, et al. Insecticide resistance: a silent base prediction. Curr Biol. 2004;14(14):R552-R553.
35. Labbé P, Berthomieu A, Berticat C, Alout H, Raymond M, Lenormand T, et al. Independent duplications of the acetylcholinesterase gene conferring insecticide resistance in the mosquito Culexpipiens. MolBiolEvol. 2007;24(4):1056-1067.
36. Cornel AJ, Stanich, MA, McAbee RD, Mulligan III FS. High level methoprene resistance in the mosquito Ochlerotatusnigromaculis (Ludlow) in Central California. Pest Manag Sci. 2002;58(8):91-798.
37. Paul A, Harrington LC, Zhang L, Scott JG. Insecticide resistance in Culexpipiens from New York. J Am Mosquito Contr. 2005;21(3):305-310.
38. Tabashnik BE, Brévault T, CarrièreY. Insect resistance to Bt crops: lessons from the first billion acres. Nature biotechnol. 2013;31(6):510.
39. Tetreau G, Alessi M, Veyrenc S, Périgon S, David JP, Reynaud S, et al. Fate of Bacillus thuringiensis subsp. israelensis in the field: evidence for spore recycling and differential persistence of toxins in leaf litter. Appl Environ Microbiol.2012;78(23):8362-8367.
40. Mittal PK. Biolarvicides in vector control: challenges and prospects. J Vector Borne Dis. 2003;40(1-2):20-32.
41. Nielsen-Leroux C, Pasteur N, Prètre J, Charles JF, Ben Sheikh H, ChevillonC. High resistance to Bacillus sphaericus binary toxin in Culexpipiens (Diptera: Culicidae): the complex situation of west Mediterranean countries. J Med Entomol.2002;39(5):729-735.
42. Chevillon C, Bernard C, Marquine M, Pasteur N. Resistance to Bacillus sphaericus in Culexpipiens (Diptera: Culicidae): interaction between recessive mutants and evolution in southern France. J Med Entomol. 2001;38(5):657-664.
43. Nielsen‐Leroux C, Charles JF, Thiéry I, Georghiou GP. Resistance in a laboratory population of Culexquinquefasciatus (Diptera: Culicidae) to Bacillus sphaericus binary toxin is due to a change in the receptor on midgut brush‐border membranes. EurJ biochemi. 1995;228(1):206-210.
44. Darboux I, Pauchet Y, Castella C, Silva-Filha MH, Nielsen-LeRoux C, Charles  JF, et al. Loss of the membrane anchor of the target receptor is a mechanism of bioinsecticide resistance. ProcNatlAcadSci USA.. 2002;99(9):5830-5835.
45. Darboux I, Charles JF, Pauchet Y, Warot S, PauronD. Transposon‐mediated resistance to Bacillus sphaericus in a field‐evolved population of Culexpipiens (Diptera: Culicidae). Cell Microbiol.2007;9(8):2022-2029.