Insecticide resistance and management in diamondback moth (Plutella xylostella L.): A Review

Introduction

Excessive and continuous insecticide use has resulted in three major issues, known as the "3R's": resurgence, residue and resistance. Resurgence refers to the rapid re-increase of pest populations following repeated insecticide applications, while residues represent the persistent traces of insecticides in the environment and food systems. Resistance is defined as a heritable reduction in pest sensitivity evident when an insecticide repeatedly fails to provide effective control despite being used according to label recommendations (Anonymous, 2025a) [1] . Multiple factors contribute to resistance development in the diamondback moth (DBM), including improper insecticide use, high application frequency, enhanced detoxification mechanisms and rapid reproductive capacity. Globally, the APRD (Arthropod Pesticide Resistance Database) reports that up to 2024, 632 arthropod species have developed resistance to 364 chemical compounds, accounting for 18,871 resistance cases (Payumo et al., 2024) [40] . DBM ranks first among resistant species, exhibiting resistance to over 104 insecticidal compounds (Eslamloo and Ito, 2025) [10] . Within the genus Plutella, six species are recognized: P. xylostella, P. antiphona, P. australiana, P. armoraciae, P. hass and P. sera, of which P. xylostella is the most widely distributed worldwide (Kfir, 1998) [22] . The diamondback moth (Plutella xylostella L.), believed to have originated in the Mediterranean region, was first reported in India by Fletcher in 1914. It is now recognized as one of the most destructive pests of cruciferous crops across major states, including Punjab, Uttar Pradesh, Bihar, Maharashtra, Tamil Nadu, Karnataka and Gujarat. DBM is highly adaptable to diverse climatic conditions, exhibits rapid dispersal ability and has a short life cycle. Being oligophagous and multivoltine, it can complete 4 to 20 generations per year depending on temperature.

The gregarious larvae, equipped with chewing mouthparts, feed continuously on leaf tissues, causing characteristic windowing holes and severe defoliation. Damage may extend from seedlings to head formation stages, occurring year-round except during the rainy season (Mubashir and Seram, 2022) [35] . In the world diamondback moth first time developed resistance against DDT in 1953 at Lembang, Indonesia (Johnson, 1953) [20] . Later, in 1989 it also developed resistance to Bacillus thuringiensis var. kurstaki. In India, they first time they develop resistance against DDT and Parathion in 1968, while to from B. thuringiensis kurstaki in the year 1995 (Rabindra et al., 1995) [42] . [18] .  Example: DBM develop resistance against chlorantraniliprole and also they develop resistance against flubendiamide because both insecticide belong to same class diamide (IRAC, Group 28)  Multiple resistance: When insects develop resistance against more than one insecticide or more than one class of insecticide, it is called as multiple resistance.

Example: DBM develop resistance against many class of insecticide like, organochlorines, organophosphates, carbamates, synthetic pyrethroids and diamide.  Polygenic resistance: When insects develop resistance involving more than one gene, it is called as polygenic resistance.

Example: DBM has developed resistance to theCry1Ac and Cry2Ab genes, both genes are derived from B. thuringiensis.

Mechanisms of development of resistance

Insects develop resistance mainly ways reduced target site sensitivity increased detoxification by detoxifying enzymes (cytochrome P450s, esterases or glutathione S-transferases) and behavioural mechanism (Hubbard and Murillo, 2024) [15] . Target sites are the parts of the insect where insecticides act. When these sites change due to mutations, insecticides can not bind properly, so they cause less harm (Charaabi et al. 2018) [8] . Detoxification occurs when enzymes quickly break down insecticides into metabolites. Insects may also resist insecticides by reducing penetration through the cuticle or by changing their behaviour to avoid exposure to insecticide (Karaagac, 2012) [21] . Many mechanisms are responsible for the development of resistance in insects but, two major mechanisms are commonly observed in insects: behavioural mechanisms and physiological mechanisms (Sparks et al., 1989) [52] . [52] described behavioural resistance as adaptive changes in insect behaviour that reduce or prevent exposure to insecticides, enabling survival under otherwise lethal conditions. Such behaviours evolve under selective pressure and allow insects to detect, avoid or minimize contact with toxicants, for example, by ceasing feeding, moving to untreated leaf surfaces, shifting deeper into the crop canopy or dispersing from sprayed areas. In P. xylostella adults exhibit selective oviposition on safer sites while larvae actively migrate away from treated foliage, thereby limiting exposure to foliar insecticides. These avoidance behaviours contribute significantly to the diamondback moth's resistance to multiple insecticide classes, including organochlorines, organophosphates, carbamates and synthetic pyrethroids (Shehzad et al., 2023) [49] .

Behavioural mechanisms

Physiological mechanism

Refers to any internal change like reduce penetration, enhance detoxification and change target site shape by mutation is called physiological mechanism. It include three mechanisms, viz., penetration resistance, metabolic detoxification and altered target-site mutation (Siddiqui et al., 2023) [50] .

Penetration resistance

Source: https://encrypted-tbn0.gstatic.com/images?q=tbn:ANd9GcSa5fPJKXYkkBnxIigW-g_l0TlVjMjeI_iJ_1Z-Z2ZK_HGwHddg Penetration resistance refers to the ability of insect cuticles to prevent insecticides from penetrating, thereby reducing their effectiveness (Lin et al., 2012) [27] . For an insecticide to reach its target site, it must penetrate multiple cuticular layers, the epicuticle, exocuticle, endocuticle and epidermis before entering the haemolymph. In resistant insects, alterations in cuticle composition, increased cuticle thickness or reduced permeability slow insecticide penetration resulting in lower concentrations reaching the target site (Balabanidou et al., 2018) [5] . This reduced uptake allows detoxification enzymes to inactivate the insecticide efficiently, enabling the insect to survive and reproduce without toxic effects (Siddiqui et al., 2023) [50] . Although cuticular resistance in P. xylostella is less studied, recent findings indicate distinct differences in cuticle-associated microRNAs between resistant and susceptible populations.

MicroRNAs (miRNAs), small non-coding RNAs that regulate gene expression, can influence cuticle formation and modification, thereby contributing to the development of insecticide resistance (Shabbir et al., 2021) [48] .

Metabolic detoxification

Metabolic resistance is a major defence mechanism in insects, driven by enzymatic detoxification systems that neutralize insecticide molecules (Panini et al., 2016; Hilliou et al., 2021) [39,14] . Three key enzyme families are primarily responsible for resistance development: Cytochrome P450 monooxygenases (P450s), Glutathione S-transferases (GSTs) and Carboxyl/cholinesterases (CCEs) (Panicker, 2025) [38] . Metabolic detoxification occurs in two phases.

Phase I involves oxidation or hydrolysis of insecticides mainly mediated by P450s and esterases, producing reactive intermediates that become more water soluble (Zhou et al., 2019) [63] . Phase II consists of conjugation reactions catalyzed by GSTs, converting Phase I metabolites into highly water soluble, less toxic compounds that are readily excreted (Mak and Denisov, 2018) [31] . Through this mechanism, diamondback moth develops resistance to several insecticide groups, including carbamates, organophosphates and pyrethroids. Overall, metabolic detoxification relies on a diverse suite of enzymes capable of detoxifying multiple insecticide classes (Shehzad et al., 2023) [49] .

Table 1:

Major detoxification enzymes and the insecticide classes against which they confer metabolic resistance in insects.

Detoxify enzymes Classes Carboxylesterases

Carbamates, organophosphates, synthetic pyrethroids Glutathione-S-transferase Diamides, organophosphates, synthetic pyrethroids Cytochrome P450

Carbamates, organophosphates, synthetic pyrethroids

Altered target-site mutation

Target-site insensitivity is one of the most important and second most common mechanism (primary mechanism) of resistance (Liang et al., 2025; Karaagac, 2012) [25,21] . In this mechanism, the site where the toxin normally binds becomes modified, reducing the effectiveness of the insecticide (Charaabi et al., 2018; Anonymous, 2025b) [8,2] . Such modifications usually occur due to small genetic changes in insects, for example, a single nucleotide mutation in the DNA. These mutations alter the amino acid sequence of the target protein, changing its binding structure. As a result, the insecticide cannot attach properly to its target site, allowing the insect to survive the treatment. Several classes of insecticides, including diamides, organophosphates, organochlorines and synthetic pyrethroids, have developed resistance in P. xylostella through this target-site mechanism (Siddiqui et al., 2023) [50] .

Source: https://pesticidestewardship.org/resistance/insecticide-resistance/insecticide-resistance-mechanisms/ In susceptible insects, insecticides effectively bind to their target sites, inhibiting chitin synthesis and disrupting the molting process. This leads to paralysis and eventual death, as the target site remains fully sensitive to the insecticide. This interaction follows a classic lock and key model where the insecticide (key) fits precisely into the target protein (lock). In resistant insects, mutations in the target site alter its structure, reducing or preventing insecticide binding. As a result, chitin synthesis continues normally, allowing the insect to grow, molt and reproduce despite insecticide exposure. The lock and key mechanism is compromised because the "lock" is modified and the insecticide no longer fits effectively. Such target-site alterations represent a major mechanism of resistance and can confer cross-resistance to multiple classes of insecticides. [43] .

Ryanodine receptor

Source: https://irac-online.org/documents/diamide-moa-rotation/ Because the toxic mode of action is effectively neutralized, resistant insects continue to feed, develop and reproduce without adverse effects from diamide exposure (Jeanguenat, 2013) [19] .

Reduced GABA receptor sensitivity notably the A2S (Alanine → Serine) and A2G (Alanine → Glycine) substitutions alter the insecticide-binding site, reducing binding affinity. Consequently, chloride ion flow through the GABA channel remains unaffected, normal hyperpolarization is maintained and nerve signaling proceeds normally. These structural changes enable resistant insects to survive insecticide exposure.

Methods for detection of insecticidal resistance

Resistance is determined by using different detection methods like conventional standard bioassay and biochemical, immunological and molecular methods. Among them, conventional standard bioassay and biochemical detection methods are widely used.

Conventional method

Conventional method published by International Resistance Action Committee (IRAC), in standard detection method of resistance insect collect from the field and rear them up to next generation. Larvae or adults are then exposed to different doses of an insecticide and their mortality is recorded. The LD₅₀ or LC₅₀ values for both susceptible and field populations are calculated using probit analysis (Karaagac, 2012) [21] . Total five steps are involved in conventional method.

 [33] reported that Dichlorvos 76 EC showed the highest resistance ratio against P. xylostella with a resistance level of 4.99 fold and LC₅₀ value is 15.63 mg/L.

Biochemical method

Biochemical assays can be used to measure the activity of detoxification enzymes (Esterase, Glutathione-S-transferase (GST) and CYTP450) enzymes that help to break down insecticides, and also identify which mechanism is involved in insecticide resistance. Once a population is well characterized, these assays can also be applied to monitor changes in the frequency of resistance related genes in field populations exposed to different levels of selection pressure (Karaagac, 2012) [21] . [11] . The mixture is incubated at 25-30 °C for 15-30 minutes to allow the enzymatic reaction to occur and produce measurable colour or fluorescence.  Step 3: During the biochemical detection assay each enzyme produces a characteristic colour after incubation with its specific substrate. Esterases produce blue colour, GST (Glutathione-S-transferase) produces yellow colour and cytochrome P450 produce red to pink colour. Intensity of the developed colour directly indicates the level of enzyme activity in the insect sample darker and stronger colour reflects higher enzymatic activity.  Step 4: After colour development, the intensity of the colour is measured by using a spectrophotometer or fluorometer. Higher absorbance (or fluorescence) values indicate higher enzyme activity reflecting a stronger metabolic detoxification capacity in the insect population.  Step 5: After measuring the color intensity, the enzyme activity level is calculated, which is then correlated with the presence and degree of insecticide resistance in the tested insect population. Enzyme activity levels are compared between susceptible and field populations. Higher activity of enzymes indicates the insects may have developed metabolic resistance to insecticides.

Eziah et al. (2009) [11] evaluated larval populations of P. xylostella highest ECOD activity was observed in the WiC strain (30.72; 5.75-fold higher than susceptible HaC strain), highest α-N-OH esterase activity occurred in the WiC strain (611.20; 1.88 fold increase) and highest GST CDNB conjugation activity was recorded in the YaC strain (448.11; 1.61-fold over than susceptible HaC strain) These results indicate that the WiC and YaC strains show strongest elevation in P450, esterase and GST enzymes respectively.

Insecticide Resistance Management (IRM)

Insecticide Resistance Management (IRM) involves designing and implementing pest control strategies that minimize selection pressure on insect populations, thereby slowing the development of resistance and preserving the long-term effectiveness of insecticides (Sarangi et al., 2025) [46] . Effective IRM includes rotation of insecticides with different modes of action, integration of non-chemical methods and use of synergists. Insecticides should be applied only when pest populations reach or exceed the threshold, ensuring that unnecessary sprays are avoided. This approach reduces excessive selection pressure, slows the development of resistance and supports sustainable resistance management.

Window approach

The window approach is a simple resistance management strategy in which the entire crop period is divided into different "windows" such as the transplanting stage (W1), cupping stage (W2), head-formation stage (W3) and harvesting stage (W4) and each window is assigned a different class of insecticide with a different mode of action. By spraying only one insecticide group within a window and use other group in next window continuous rotation is achieved, reducing selection pressure on any single chemistry. The insect generations (e.g., DBM Gen-1, Gen-2 and Gen-3) occur during different crop windows and by changing the insecticide group in each window every new generation is exposed to a different mode of action. This prevents the same population from getting the same insecticide again and again, which helps slow down resistance development (Chaudhari, 2020) [9] . Chaudhari (2020) [9] concluded that resistance development in P. xylostella to chlorantraniliprole was delayed after the use of different groups of insecticides in sequential windows. In lab CH-resistance population resistance is reduced in treatment 1 (1132.15 to 99.56), in Sabarkantha CH-Resistant population also resistance ratio is reduced in treatment 1from 212.19 to 13.14 and in Rajkot CH-Resistant again resistance is decrease from 169.64 to 6.90. This indicates that the sequential use of different classes of insecticides effectively delayed and reduced resistance in P. xylostella.

Use of synergists

Synergists have been used commercially for about 50 years and played a significant role in improving insecticide performance (Bernard and Philogene, 1993) [7] . Synergists is nontoxic chemicals added to insecticides to enhance their effectiveness against insect-pests and also help to insecticide work normally by blocking detoxification enzymes; They do not actually increase the inherent potency of the insecticides. By inhibiting detoxification enzymes synergists can make resistant populations susceptible again and help prevent or delay the development of insecticide resistance. [29] concluded that the use of synergists DEF, DEM and PBO significantly reduced the LC₅₀ values of cyantraniliprole in all three strains indicating their ability to enhance insecticide toxicity. The strongest synergistic effect was observed with DEM in both the Cya-SEL strain (SR=3.2) and UNSEL strain (SR=2.1) followed by PBO and DEF. In LAB strain PBO produced highest synergism (SR=1.5). Overall, data suggested that metabolic detoxification enzymes particularly esterases and cytochrome P450s play a major role in cyantraniliprole resistance and synergists can partially restore susceptibility.

Another study conducted by Chaudhari (2020) [9] on the effect of sesame oil on cyantraniliprole resistance in P. xylostella, reported that after the use sesame oil with cyantraniliprole, reduced the LC₅₀ value and increased the synergistic ratio in all tested populations, indicating that the synergist enhanced the effectiveness of insecticide and delayed resistance.

Insecticidal mixture

Insecticide mixtures are considered an essential strategy for managing resistance in many insect pests (South and Hastings, 2018) [51] . Insecticide mixtures increase the level of target pest control and broadening the range of pests controlled; however, they may offer benefits for IRM when appropriately incorporated into rotation strategies with additional mode(s) of action, but generally a single mixture should not be relied upon alone (IRAC, 2012) [17] . Shabbir et al. (2021) [48] The results show that the FOH-DBM strain was the most susceptible baseline for both chlorantraniliprole and Bt, while the other three strains (Bt-DBM, CL-DBM and BtC-DBM) exhibited high levels of resistance, particularly to chlorantraniliprole, where CL-DBM showed the highest resistance ratio (31.

Future prospects

Future research should prioritize the development of rapid, high-throughput assays for detecting resistance across multiple insecticide classes in diverse DBM populations, alongside detailed investigations into behavioral mechanisms that facilitate avoidance and reduced exposure to insecticides. Additionally, expanding genome-editing targets and enhancing the delivery efficiency of advanced molecular tools such as RNAi and CRISPR/Cas9 will be crucial for designing next-generation strategies to manage resistance in P. xylostella.

References

1. Anonymous. Insecticide Resistance Action Committee (IRAC) [Internet]. 2025 [cited 2026 Feb 4]. Available from: https://irac-online.org/
2. Asad M, Chang Y, Liao J, Yang G. CRISPR/Cas9 genome editing in the diamondback moth: Current progress, challenges, and prospects. Int J Mol Sci. 2025;26(4):1515.
3. Baker GJ, Kovaliski J. Detection of insecticide resistance in Plutella xylostella (L.) (Lepidoptera: Plutellidae) populations in South Australian crucifer crops. Aust J Entomol. 1999 Jun;38(2):132-134.
4. Balabanidou V, Grigoraki L, Vontas J. Insect cuticle: A critical determinant of insecticide resistance. Curr Opin Insect Sci. 2018;27:68-74.
5. Bautista MA, Miyata T, Miura K, Tanaka T. RNA interference-mediated knockdown of a cytochrome P450, CYP6BG1, from the diamondback moth, Plutella xylostella, reduces larval resistance to permethrin. Insect Biochem Mol Biol. 2009;39(1):38-46.
6. Bernard CB, Philogène BJ. Insecticide synergists: Role, importance, and perspectives. J Toxicol Environ Health A. 1993;38(2):199-223.
7. Charaabi K, Bouhachem BS, Makni M, Denholm I. Occurrence of target-site resistance to neonicotinoids in the aphid, Myzus persicae in Tunisia, and its status on different host plants. Pest Manag Sci. 2018;74(6):12971301.
8. Chaudhari JD. Diamide insecticides resistance in diamondback moth, Plutella xylostella (Linnaeus) [dissertation]. Anand (Gujarat): Anand Agricultural University; 2020.
9. Eslamloo G, Ito T. Insecticidal efficacy of non-Bt bacterial strains against diamondback moth, Plutella xylostella (L.). Front Sustain Food Syst. 2025 Jun 25;9:1591114.
10. Eziah VY, Rose HA, Wilkes M, Clift AD. Biochemical mechanisms of insecticide resistance in the diamondback moth (DBM), Plutella xylostella L. (Lepidoptera: Yponomeutidae), in the Sydney region, Australia. Aust J Entomol. 2009;48(4):321-327.
11. Food and Agriculture Organization of the United Nations. The state of food and agriculture. Rome: FAO; 1967.
12. Gong W, Yan HH, Gao L, Guo YY, Xue CB. Chlorantraniliprole resistance in the diamondback moth (Lepidoptera: Plutellidae). J Econ Entomol. 2014;107(2):806-814.
13. Hilliou F, Chertemps T, Maïbèche M, Le Goff G. Resistance in the genus Spodoptera: Key insect detoxification genes. Insects. 2021;12(6):544.
14. Hubbard CB, Murillo AC. Behavioral resistance to insecticides: current understanding, challenges, and future directions. Curr Opin Insect Sci. 2024;63:101177.
15. Insecticide Resistance Action Committee. Management of insecticide resistance in vectors of public health importance. IRAC; 2011.
16. IRAC. IRAC international insecticide mixture statement; 2012.
17. Ishadi A, Mazlan N. Insecticide use impacts on pest resistance: An evidence from diamondback moth. Int J Sci Basic Appl Res. 2015;22:131-150.
18. Jeanguenat A. The story of a new insecticidal chemistry class: The diamides. Pest Manag Sci. 2013;69(1):7-14.
19. Johnson DR. Plutella maculipennis resistance to DDT in Java. J Econ Entomol. 1953;46(1):176.
20. Karaagac SU. Insecticide resistance. London: IntechOpen; 2012.
21. Kfir R. Origin of the diamondback moth (Lepidoptera: Plutellidae). Ann Entomol Soc Am. 1998;91(2):164167.
22. Knight SC, Elias J. Managing insecticide resistance in Asia. In: VIII International Conference on Management of the Diamondback Moth and Other Crucifer Insect Pests; 2019, p. 90.
23. Li Z, Feng X, Liu SS, You M, Furlong MJ. Biology, ecology, and management of the diamondback moth in China. Annu Rev Entomol. 2016;61(1):277-296.
24. Liang J, Xiao F, Ojo J, Chao WH, Ahmad B, Alam A, International Journal of Advanced Biochemistry Research https://www.biochemjournal.com et al. Insect resistance to insecticides: Causes, mechanisms, and exploring potential solutions. Arch Insect Biochem Physiol. 2025;118(2):e70045.
25. Lima JE, Siqueira HA. Selection of Plutella xylostella (L.) (Lepidoptera: Plutellidae) to chlorfenapyr resistance: Heritability and the number of genes involved. Rev Caatinga. 2017;30(4):1067-1072.
26. Lin Y, Jin T, Zeng L, Lu Y. Cuticular penetration of βcypermethrin in insecticide-susceptible and resistant strains of Bactrocera dorsalis. Pestic Biochem Physiol. 2012 Jul 1;103(3):189-193.
27. Liu N. Insecticide resistance in mosquitoes: Impact, mechanisms, and research directions. Annu Rev Entomol. 2015;60:537-559.
28. Liu X, Ning Y, Wang H, Wang K. Cross-resistance, mode of inheritance, synergism, and fitness effects of cyantraniliprole resistance in Plutella xylostella. Entomol Exp Appl. 2015;157(3):271-278.
29. Ma R, Haji-Ghassemi O, Ma D, Jiang H, Lin L, Yao L, et al. Structural basis for diamide modulation of ryanodine receptor. Nat Chem Biol. 2020;16(11):12461254.
30. Mak PJ, Denisov IG. Spectroscopic studies of the cytochrome P450 reaction mechanisms. Biochim Biophys Acta Proteins Proteom. 2018;1866(1):178-204.
31. McFarlane GR, Whitelaw CB, Lillico SG. CRISPRbased gene drives for pest control. Trends Biotechnol. 2018;36(2):130-133.
32. Meghana C, Jayappa J, Reddy NA, Devappa V, Sridhar V, Jyothi K. Assessing susceptibility of diamondback moth, Plutella xylostella (Lepidoptera: Plutellidae) population of different geographic region to selected newer insecticides. J Entomol Zool Stud. 2018;6(1):320-327.
33. Mohan M, Gujar GT. Susceptibility pattern and development of resistance in the diamondback moth, Plutella xylostella L, to Bacillus thuringiensis Berl var kurstaki in India. Pest Manag Sci. 2000;56(2):189-194.
34. Mubashir S, Seram D. Insecticidal resistance in diamondback moth (Plutella xylostella L.). Pharma Innov J. 2022;11(7):958-962.
35. Oliveira AC, Siqueira HÁ, Oliveira JV, Silva JE, Michereff Filho M. Resistance of Brazilian diamondback moth populations to insecticides. Sci Agric. 2011;68:154-159.
36. Oplopoiou M, Elias J, Slater R, Bass C, Zimmer CT. Characterization of emamectin benzoate resistance in the diamondback moth, Plutella xylostella (Lepidoptera: Plutellidae). Pest Manag Sci. 2024;80(2):498-507.
37. Panini M, Manicardi GC, Moores GD, Mazzoni EJ. An overview of the main pathways of metabolic resistance in insects. Invertebr Surviv J. 2016;13(1):326-335.
38. Payumo J, Bello-Bravo J, Chennuru V, Mercene SA, Yim C, Duynslager L, et al. An assessment model for agricultural databases: the arthropod pesticide resistance database as a case study. Insects. 2024;15(10):747.
39. Pu X, Yang Y, Wu S, Wu Y. Characterisation of abamectin resistance in a field-evolved multiresistant population of Plutella xylostella. Pest Manag Sci. 2010;66(4):371-378.
40. Rabindra RJ, Gailce Leo Justin C, Jayaraj S. Relative susceptibility of two larval populations of diamondback moth (Plutella xylostella) to insecticides and Bacillus thuringiensis. Indian J Agric Sci. 1995;65:152.
41. Ranganathan M, Narayanan M, Kumarasamy S. Importance of metabolic enzymes and their role in insecticide resistance. In: New and future development in biopesticide research: biotechnological exploration. Singapore: Springer Nature Singapore; 2022, p. 243260.
42. Ribeiro LM, Siqueira HA, Teixeira WV, Ferreira HN, Silva WM, Silva JE, et al. Field resistance of Brazilian Plutella xylostella to diamides is not metabolismmediated. Crop Prot. 2017;93:82-88.
43. Santos VC, De Siqueira HA, Da Silva JE, De Farias MJ. Insecticide resistance in populations of the diamondback moth, Plutella xylostella (L.) (Lepidoptera: Plutellidae), from the state of Pernambuco, Brazil. Neotrop Entomol. 2011;40:264270.
44. Sarangi S, Bhavana P, Padhan EK. Insecticide resistance and management strategies. In: Dash SS, Pradhan PP, Sahoo S, Nag YK, Editors. Crop Protection; 2025, p. 184-208.
45. Shabbir MZ, He L, Shu C, Yin F, Zhang J, Li ZY. Assessing the single and combined toxicity of chlorantraniliprole and Bacillus thuringiensis (GO33A) against four selected strains of Plutella xylostella (Lepidoptera: Plutellidae), and a gene expression analysis. Toxins. 2021;13(3):227.
46. Shabbir MZ, Yang X, Batool R, Yin F, Kendra PE, Li ZY. Bacillus thuringiensis and chlorantraniliprole trigger the expression of detoxification-related genes in the larval midgut of Plutella xylostella. Front Physiol. 2021;12:780255.
47. Shehzad M, Bodlah I, Siddiqui JA, Bodlah MA, Fareen AG, Islam W. Recent insights into pesticide resistance mechanisms in Plutella xylostella and possible management strategies. Environ Sci Pollut Res. 2023;30(42):95296-95311.
48. Siddiqui JA, Fan R, Naz H, Bamisile BS, Hafeez M, Ghani MI, et al. Insights into insecticide-resistance mechanisms in invasive species: Challenges and control strategies. Front Physiol. 2023;13:1112278.
49. South A, Hastings IM. Insecticide resistance evolution with mixtures and sequences: A model-based explanation. Malar J. 2018;17(1):80.
50. Sparks TC, Lockwood JA, Byford RL, Graves JB, Leonard BR. The role of behavior in insecticide resistance. Pestic Sci. 1989;26(4):383-399.
51. Sunitha V, Singh TV, Supriya GB, Satyanarayana J. Insecticide resistance monitoring of diamond back moth, Plutella xylostella (Linn.) in Delhi population. J Entomol Zool Stud. 2020;8(5):1706-1712.
52. Tabashnik BE, Cushing NL, Finson N, Johnson MW. Field development of resistance to Bacillus thuringiensis in diamondback moth (Lepidoptera: Plutellidae). J Econ Entomol. 1990;83(5):1671-1676.
53. Tamilselvan R, Kennedy JS, Suganthi A. Monitoring the resistance and baseline susceptibility of Plutella xylostella (L.) (Lepidoptera: Plutellidae) against International Journal of Advanced Biochemistry Research https://www.biochemjournal.com spinetoram in Tamil Nadu, India. Crop Prot. 2021;142:105491.
54. Athens (GA): University of Georgia; 2018.
55. Truong KM, Pessah IN. Comparison of chlorantraniliprole and flubendiamide activity toward wild-type and malignant hyperthermia-susceptible ryanodine receptors and heat stress intolerance. Toxicol Sci. 2019;167(2):509-523.
56. Verma AN, Sandhu GS. Chemical control of diamondback-moth, Plutella maculipennis (Curtis). 1968.
57. Wang M, Zhu B, Zhang L, Xiao Y, Liang P, Wu K. Influence of seasonal migration on evolution of insecticide resistance in Plutella xylostella. Insect Sci. 2022;29(2):496-504.
58. Wang X, Wu Y. High levels of resistance to chlorantraniliprole evolved in field populations of Plutella xylostella. J Econ Entomol. 2012;105(3):10191023.
59. Zhang S, Zhang X, Shen J, Li D, Wan H, You H, et al. Cross-resistance and biochemical mechanisms of resistance to indoxacarb in the diamondback moth, Plutella xylostella. Pestic Biochem Physiol. 2017;140:85-89.
60. Zhang S, Zhang X, Shen J, Mao K, You H, Li J. Susceptibility of field populations of the diamondback moth, Plutella xylostella, to a selection of insecticides in Central China. Pestic Biochem Physiol. 2016;132:3846.
61. Zhou Y, Fu WB, Si FL, Yan ZT, Zhang YJ, He QY, et al. UDP-glycosyltransferase genes and their association and mutations associated with pyrethroid resistance in Anopheles sinensis (Diptera: Culicidae). Malar J. 2019;18(1):62.