Proteomics in Insecticide Toxicology

  • Published : 2007.03.31

Abstract

Mechanisms of insecticide resistance found in insects may include three general categories. Modified behavioral mechanisms can let the insects avoid the exposure to toxic compounds. The second category is physiological mechanisms such as altered penetration, rapid excretion, lower rate transportation, or increased storage of insecticides by insects. The third category relies on biochemical mechanisms including the insensitivity of target sites to insecticides and enhanced detoxification rate by several detoxifying mechanisms. Insecticides metabolism usually results in the formation of more water-soluble and therefore more readily eliminated, and generally less toxic products to the host insects rather than the parent compounds. The representative detoxifying enzymes are general esterases and monooxygenases that catalyze the toxic compounds to be more water-soluble forms and then secondary metabolism is followed by conjugation reactions including those catalyzed by glutathione S-transferases (GSTs). However, a change in the resistant species is not easily determined and the levels of mRNAs do not necessarily predict the levels of the corresponding proteins in a cell. As genomics understands the expression of most of the genes in an organism after being stressed by toxic compounds, proteomics can determine the global protein changes in a cell. In this present review, it is suggested that the environmental proteomic application may be a good approach to understand the biochemical mechanisms of insecticide resistance in insects and to predict metabolomic changes leading to physiological changes of the resistant species.

Keywords

References

  1. Peter, A. et al. Mapping and identification of essential gene functions on the X chromosome of Drosophila. EMBO Rep. 3:34-38 (2002) https://doi.org/10.1093/embo-reports/kvf012
  2. Sparks, T. C. et al. The role of behavior in insecticide resistance. Pestic. Sci. 26:383-399 (1989) https://doi.org/10.1002/ps.2780260406
  3. Georghiou, G. P. The evolution of resistance to pesticides. Ann. Rev. Ecol. Syst. 3:133-168 (1972) https://doi.org/10.1146/annurev.es.03.110172.001025
  4. Plapp, Jr. F. W. The genetic basis of insecticide resistance in the housefly: evidence that a single locus plays a major role in metabolic resistance to insecticides. Pestic. Biochem. Physiol. 22:194-201 (1984) https://doi.org/10.1016/0048-3575(84)90089-0
  5. Lockwood, J. A., Sparks, T. C. & Story, R. N. Evolution of insect resistance to insecticides: a reevaluation of the roles of physiology and behavior. Bull. Entomol. Soc. Am. 30:41-50 (1984)
  6. Barson, G., Fleming, D. A. & Allan, E. Laboratory assessment of the behavioral reposnses of residual populations of Oryzaephilus surinamensis (L) (Coleoptera: silvanidae) to the contact insecticide pirimiphosmethyl by linear logistic modeling. J. Stored Prod. Res. 28:161-170 (1992) https://doi.org/10.1016/0022-474X(92)90036-P
  7. Little, E. J., McCaffery, A. R., Walker, C. H. & Parker, T. Evidence of an enhanced metabolism of cypermethrin by a monooxygenase in a pyrethroidresistant strain of the tobacco budworm (Heliothis virescens F.). Pestic. Biochem. Physiol. 34:58-68 (1989) https://doi.org/10.1016/0048-3575(89)90141-7
  8. Sawicki, R. M. & Farnham, A. W. Genetics of resistance to insecticides of the SKA strain of Musca domestica III. Location and isolation of the factors of resistance to dieldrin. Entomol. Exp. Appl. 11:133- 142 (1968) https://doi.org/10.1007/BF00305223
  9. Sawicki, R. M. & Farnham, A. W. Examination of the isolated autosomes of the SKA strain of houseflies (Musca domestica L.) for resistance to several insecticides with and without pretreatment with sesamex and TBPT. Bull. Entomol. Res. 59:409-421 (1968)
  10. Apperson, C. S. & Georghiou, G. P. Mechanisms of resistance to organophosphorus insecticides in Culex tarsalis. J. Econ. Entomol. 68:153-157 (1975) https://doi.org/10.1093/jee/68.2.153
  11. Patil, V. L. & Guthrie, F. E. Effect of anomalous cuticular phospholipids on penetration of insecticides in susceptible and resistant houseflies. Pestic. Biochem. Physiol. 11:13-19 (1979) https://doi.org/10.1016/0048-3575(79)90043-9
  12. Scott, J. G. & Georghiou, G. P. Mechanisms responsible for high levels of permethrin resistance in the housefly. Pestic. Sci. 17:195-206 (1986) https://doi.org/10.1002/ps.2780170302
  13. Sigfried, B. D. & Scott, J. G. Mechanisms reposible for propoxur resistance in the German cockroach, Blattella germinica. Pestic. Sci. 33:133-146 (1991) https://doi.org/10.1002/ps.2780330202
  14. Lee, S. E., Kim J. E. & Lee. H. S. Insecticide resistance in increasing interest. Agric. Chem. Biotech. 44: 105-112 (2002)
  15. Smissaert, H. R. Cholinesterase inhibition in spider mites susceptible and resistant to organophosphate. Science 143:129-131 (1964) https://doi.org/10.1126/science.143.3602.129
  16. Raymond, M. et al. Identification of resistance mechanisms in Culex pipiens (Diptera: Culicidae) from southern France: insensitive acetylcholinesterase and detoxifying oxidases. J. Econ. Entomol. 79:1452- 1458 (1986) https://doi.org/10.1093/jee/79.6.1452
  17. Yu, S. J. Insecticide resistance in the fall armyworm, Spodoptera frugiperda (Smith, J. E.). Pestic. Biochem. Physiol. 39:84-91 (1991) https://doi.org/10.1016/0048-3575(91)90216-9
  18. Karunaratne, K. M. & Plapp, Jr. F. W. Biochemistry and genetics of thiodicarb resistance in the housefly (Diptera: Muscidae). J. Econ. Entomol. 86:258-264 (1993) https://doi.org/10.1093/jee/86.2.258
  19. Wierenga, J. M. & Hollingworth, R. M. Inhibition of altered acetycholinesterases from insecticide-resistant Colorado potato beetles (Coleoptera: Chrysomelidae). J. Econ. Entomol. 86:673-679 (1993) https://doi.org/10.1093/jee/86.3.673
  20. Brown, T. M. & Bryson, P. K. Selective inhibitors of methylparathion-resistant acetylcholinesterase from Heliothis virescens. Pestic. Biochem. Physiol. 44:155 -164 (1992) https://doi.org/10.1016/0048-3575(92)90113-E
  21. Xu, G. & Brindley, W. A. Structure of populations of Lygus hesperus (Hemiptera: Miridae) with multiple resistance mechanism to trichlorfon. J. Econ. Entomol. 86:1656-1663 (1993) https://doi.org/10.1093/jee/86.6.1656
  22. Bisset, J. A. et al. The mechanisms of organophosphate and carbamate resistance in Culex quinquefasciatus (Diptera: Culicidae) from Cuba. Bull. Entomol. Res. 80:245-250 (1990) https://doi.org/10.1017/S0007485300050434
  23. Ayad, H. & Geoughiou, G. P. Resistance to organophosphates and carbamates in Anopheles albimanus based on reduced sensitivity to acetylcholinesterase. J. Econ. Entomol. 68:296-297 (1975)
  24. Chianliang, C. & Devonshire, A. L. Changes in membrane phospholipid, identified by Arrhenius plots of acetylcholinesterase and associated with pyrethroid resistance (kdr) in housefly. Pestic. Sci. 13:156-160 (1982) https://doi.org/10.1002/ps.2780130207
  25. Kasbekar, D. P. & Hall, L. M. A Drosophila mutation that reduces sodium channel number confers resistance to pyrethroid insecticides. Pestic. Biochem. Physiol. 32:135-145 (1998) https://doi.org/10.1016/0048-3575(88)90006-5
  26. Bull, D. L. & Pryor, N. W. Characteristics of resistance in houseflies subjected to long-term concurrent selection with malathion and permethrin. Pestic. Biochem. Physiol. 42:211-226 (1990)
  27. Pauron, D. et al. Pyrehtroid receptor in the insect Na+ channel: alteration of its properties in pyrethroid-resistant flies. Biochemistry 28:1673-1677 (1989) https://doi.org/10.1021/bi00430a037
  28. Amichot, M. et al. Target modification as a molecular mechanism of pyrethroid resistance in Drosohila melnogaster. Pestic. Biochem. Physiol. 44:183-190 (1992) https://doi.org/10.1016/0048-3575(92)90089-I
  29. Pepper, D. R. & Osborne, M. P. Electrophysiological identification of site-insensitive mechanisms in knockdown- resistant strains (kdr, super-kdr) of the housefly larva (Musca domestica). Pestic. Sci. 39:279-286 (1993) https://doi.org/10.1002/ps.2780390405
  30. Dong, K. & Scott, J. G. Linkage of kdr-type resistance and the para-homologous sodium channel gene in German cockroaches (Blatella germanica). Insect Biochem. Mol. Biol. 24:647-654 (1994) https://doi.org/10.1016/0965-1748(94)90051-5
  31. Williamson, M. S., Denholm, I., Bell, C. A. & Devonshire, A. L. Knockdown resistance (kdr) to DDT and pyrethroid insecticides maps to a sodium channel gene locus in the housefly (Musca domestica). Mol. Gen. Genet. 240:17-22 (1994)
  32. Taylor, M. F., Heckel, D. G., Brown, T. M. & Kreitman, M. E. Linkage of pyrethroid insecticide resistance to a sodium channel locus in the tobacco budworm. Insect Biochem. Mol. Biol. 23:763-775 (1993) https://doi.org/10.1016/0965-1748(93)90064-Y
  33. Takada, Y., Imahase, T., Hirano, M. & Hiroyoshi, T. Mapping of the third chromosomal recessive resistance gene to permethrin in two strains of the housefly, Musca domestica L. (Diptera: Muscidae). Appl. Entomol. Zool. 25:333-338 (1990) https://doi.org/10.1303/aez.25.333
  34. Lin, H., Bloomquist, J. R., Beeman, R. W. & Clack, J. M. Mechanisms underlying cyclodiene resistance in the red flour beetle, Tribolium castaneum (Herbst). Pestic. Biochem. Physiol. 45:154-165 (1993) https://doi.org/10.1006/pest.1993.1018
  35. Krijgsveld, J. et al. Metabolic labeling of C. elegans and D. melanogaster for quantitative proteomics. Nature Biotechnol. 21:927-931 (2003) https://doi.org/10.1038/nbt848
  36. Weill, M. G. et al. Comparative genomics: Insecticide resistance in mosquito vectors. Nature 423:136-137 (2003)
  37. Sharma, R., Komatsu, S. & Noda, H. Proteomic analysis of brown planthopper: application to the study of carbamate toxicity. Insect Biochem. Mol. 34:425-432 (2004) https://doi.org/10.1016/j.ibmb.2004.01.004
  38. Heinrichs, E. A. Impact of insecticides on the resistance and resurgence of rice planthoppers. In: Denno, R. F., Perfect, T. J. (Eds.), Planthoppers: Their Ecology and Management. Chapman and Hall Press, New York, 571-614 (1994)
  39. Schmutterer, H. Properties and potential of natural pesticides from the neem tree Azadirachta indica. Annu. Rev. Entomol. 35:271-297 (1990) https://doi.org/10.1146/annurev.en.35.010190.001415
  40. Hu, M. Y., Zhao, S. H., Wang, L. C. & Kuang, X. W. Studies on the effectiveness and growth inhibitory effect of margosan-O on Diamondback Moth (Plutella xylostella). J. South China Agric. Uni. 17:549-554 (1996)
  41. Immaraju, J. A. The commercial use of azadirachtin and its integration into viable pest control programmes. Pestic. Sci. 54:277-284 (1998) https://doi.org/10.1002/(SICI)1096-9063(1998110)54:3<277::AID-PS801>3.0.CO;2-I
  42. Huang, Z., Shi, P., Dai, J. & Du, J. Protein metabolism in Spodoptera litura (F.) is influenced by the botanical insecticide azadirachtin. Pestic. Biochem. Physiol. 80:85-93 (2004) https://doi.org/10.1016/j.pestbp.2004.07.001
  43. Rong, X. D., Xu, H. H. & Chiu, S. F. Progressing on botanical insecticide-neem research. Chin. J. Pestic. Sci. 2:9-14 (2000)
  44. Li, X. D., Chen, W. K. & Hu, M. Y. Studies on the effects and mechanisms of azadirachtin and rhodojaponin- on Spodoptera litura (F.). J. South China Agric. Univ. 16:80-85 (1995)
  45. Annadurai, R. S. & Rembold, H. Azadirachtin A modulates the tissue-specific 2D polypeptide patterns of the desert locust, Schistocerca gregaria. Naturwissenschaften 80:127-130 (1993) https://doi.org/10.1007/BF01131015
  46. Li, X. D., Chen, W. K. & Hu, M. Y. Studies on the effects and mechanisms of azadirachtin and rhodojaponin- on Spodoptera litura (F.). J. South China Agric. Univ. 16:80-85 (1995)
  47. Rao, G. V. R., Wightman, D. V. & Rao, R. World review of the natural enemies and diseases of Spodoptera litura (F.) (Lepidoptera: Noctuidae), Insect Sci. Appl. 14:273-284 (1993)
  48. Redfern, R. E., Kelly, T. J., Borkovec, A. B. & Hayes, D. K. Ecdysteroid titers and moulting aberrations in last stage Oncopeltus nymphs treated with insect growth regulations. Pestic. Biochem. Physiol. 18:351 -356 (1982) https://doi.org/10.1016/0048-3575(82)90076-1
  49. Smith, S. L. & Mitchell, M. J. Effects of azadirachtin on insect cytochrome P-450 dependent ecdysone 20- monooxygenase activity. Biochem. Biophys. Res. Commun. 154:559-563 (1988) https://doi.org/10.1016/0006-291X(88)90176-3
  50. Chiu, S. F., Zhang, X., Liu, S. K. & Huang, D. P. Growth-disruptive effects of azadirachtin on the larvae of the Asiatic corn borer (Ostrinia furnacalis G.). Acta Entomol. Sin. 27:241-247 (1984)
  51. Kumar, M. B. et al. A single point mutation in ecdysone receptor leads to increased ligand specificity: implications for gene switch applications. Proc. Natl. Acad. Sci. 99:14710-14715 (2002)
  52. Localization of ecdysone receptor protein during colour pattern formation in wings of the butterfly precis coenia (Lepidoptera: Nymphalidae) and co-expression with Distal-less protein. Dev. Genes Evol. 212:571- 584 (2003)
  53. Schnepf, E. et al. Bacillus thuringiensis and its pesticidal crystal proteins. Microbiol. Mol. Rev. 62:775- 806 (1998)
  54. Knowles, B. H. Mechanism of actin of Bacillus thuringiensis $\delta$-endotoxins. Adv. Insect Physiol. 24:275- 308 (1994) https://doi.org/10.1016/S0065-2806(08)60085-5
  55. Sangadala, S., Walters, F. S., English, L. H. & Adang, M. J. A mixture of Manduca sexta aminopeptidase and phosphatase enhances Bacillus thuringiensis insecticidal CryIA (c) toxin binding and 86Rb $(^+)-K^+$ efflux in vitro. J. Biol. Chem. 269:10088-10092 (1994)
  56. Knight, P. J., Knowles, B. H. & Ellar, D. J. The receptor for Bacillus thuringiensis CryA (c) delta-endotoxin in the brush border membrane of the lepidopteran Manduca sexta is aminopeptidase N. Mol. Microbiol. 11:429-436 (1994) https://doi.org/10.1111/j.1365-2958.1994.tb00324.x
  57. Carroll, J., Wolfersberger, M. G. & Ellar, D. J. The bacillus thuringiensis Cry1 Ac toxin-induced permeability change in Manduca sexta midgut brush border membrane vesicles proceeds by more than one mechanism. J. Cell. Sci. 110:3099-3104 (1997)
  58. Vadlamudi, R. K. et al. Cloning and expression of a receptor for an insecticidal toxin of Bacillus thuringiensis. J. Biol. Chem. 270:5490-5494 (1995) https://doi.org/10.1074/jbc.270.10.5490
  59. Nagamatsu, Y. et al. The cadherin-like protein is essential to specificity determination and cytotoxic action of the Bacillus thuringiensis insecticidal Cry1Aa toxin. FEBS Lett. 460:385-390 (1999) https://doi.org/10.1016/S0014-5793(99)01327-7
  60. Dorsch, J. A. et al. Cry1A toxins of Bacillus thuringiensis bind specifically to a region adjacent to the membrane-proximal extracellular domain of BT-R (1) in Manduca sexta: involvement of a cadherin in the entomopathogenicity of Bacillus thuringiensis. Insect Biochem. Mol. Biol. 32:1025-1036 (2002) https://doi.org/10.1016/S0965-1748(02)00040-1
  61. Candas, M. et al. Insect resistance to Bacillus thuringiensis: alterations in the indianmeal moth larval gut proteome. Mol. Cell. Proteomics. 2:19-28 (2003) https://doi.org/10.1074/mcp.M200069-MCP200
  62. McNall, R. J. & Adang, M. J. Identification of novel Bacillus thuringiensis Cry1Ac binding proteins in Manduca sexta midgut through proteomic analysis. Insect. Biochem. Mol. 33:999-1010 (2003) https://doi.org/10.1016/S0965-1748(03)00114-0
  63. Mooseker, M. S. Organization, chemistry, and assembly of the cytoskeletal apparatus of the intestinal brush border. Ann. Rev. Cell Biol. 1:209-241 (1985) https://doi.org/10.1146/annurev.cb.01.110185.001233
  64. Schwartz, L. M., Jones, M. E. E., Kosz, L. & Kuah, K. Selective repression of actin and myosin heavy chain expression during the programmed death of insect skeletal muscle. Dev. Biol. 158, 448-455 (1993) https://doi.org/10.1006/dbio.1993.1202