Insecticide Resistance in Diamondback Moth (Lepidoptera: Plutellidae) in Georgia

The diamondback moth, Plutella xylostella (L) (Lepidoptera: Plutellidae), has generally become resistant to any new insecticide mode of action used extensively for its control. Up to recent times, P. xylostella has developed resistance to 95 distinct insecticide active ingredients (Arthropod Pesticide Resistance Database, http://www.pesticideresistance.org/, accessed 19 June 2019) and has become one of the most difficult pests to control in cruciferous vegetables (Furlong et al. 2013, Annu. Rev. Entomol. 58: 517–541). Insecticide resistance levels in P. xylostella populations were evaluated in 2012, 2013, 2016, 2017, and 2018 at Tifton (Tift County), GA, for selected insecticides to provide some baseline data on insecticide efficacy. These data were assessed on P. xylostella populations in Tift County around which Georgia's acreage of curciferous crops is concentrated. The objective of this study was to establish baseline lethal concentration (LC) data for P. xyllostella to chlorantraniliprole (Coragen^®, E.I. du Pont de Nemours and Company, Wilmington, DE; a ryanodine receptor modulator, IRAC Group 28 https://www.irac-online.org/modes-of-action/, accessed 19 June 2019) and spinetoram (Radiant^®, Dow AgroSciences LLC, Indianapolis, IN; a nicotinic acetylcholine receptor allosteric activator, IRAC Group 5) in Tift County, GA.

Larval P. xylostella specimens were collected from the Tifton location (N 31.47°, E 83.53°), a University of Georgia (UGA) organic-designated research farm in Tift County, from April to June of 2012 for 50% lethal concentration (LC₅₀) bioassays. A susceptible P. xylostella population, which was not exposed to insecticides for several years and also from the Tifton location, was used as a check in 2012. Possible resistant P. xylostella populations which were exposed to insecticides in recent years were collected from commercial cabbage fields near Omega, GA, in April–June in 2013, 2016, 2017, and 2018. In addition, we had a susceptible laboratory population provided by Dr. T. Shelton (Cornell Univ., Ithaca, NY) as a Coragen-susceptible check in 2018. For Radiant, our laboratory colony was considered the susceptible check. For zeta-cypermethrin (Mustang Max^®, IRAC Group 3, FMC Corporation, Philadelphia, PA) and cyantraniliprole (Verimark^®, E.I. du Pont de Nemours and Company, Wilmington, DE), we only tested the Hort Hill Farm population, our “field susceptible” population. These P. xylostella larval field collections were placed in a Specimen Transfer Cage (1450TC, BioQuip Products, Rancho Dominguez, CA) with cabbage sprouts as larval feeding media. The resulting adults were fed on 10% (v/v) honey solution while in the cage and allowed to lay eggs continuously on cabbage seedlings. The resulting larvae were used in the following bioassays.

Formulated insecticides used for bioassays included zeta-cypermethrin (Mustang Max 600g ai/liter SC; concentrations mg/liter: 600, 6, 0.06, 0.015, and 0.00), spinetoram (Radiant 120g ai/liter SC; concentrations mg/liter: 120, 12, 1.2, 0.6, 0.12, 0.012, and 0.00), cyantraniliprole (Verimark 200g ai/liter SC; concentrations mg/liter: 50, 0.5, 0.25, 0.125, 0.0625, 0.03125, and 0.00), and chloranthraniliprole (Coragen 200g ai/liter SC; concentrations mg/liter: 50, 10, 1, 0.25, 0.0625, 0.015625, and 0.00). The other insecticides used in these studies included indoxacarb (Avaunt^®, IRAC Group 22A, Dupont Crop Protection, Newark, DE), cyclaniliprole (Harvanta^®, IRAC Group 28, Summit Agro USA, LLC, Durham, NC), cyantraniliprole (Exirel^®, IRAC Group 28, E. I. du Pont de Nemours and Company, Wilmington, DE), naled (Dibrom^®, IRAC Group 1B, AMVAC Chemical Corporation, Newport Beach, CA), emamectin benzoate (Proclaim^®, IRAC Group 6, Syngenta Crop Protection, Inc., Greensboro, NC), bifenthrin (Brigade^®, IRAC Group 3A, FMC Corporation, Philadelphia, PA) and Bacillus thuringiensis, subsp. kurstaki (Dipel^®, IRAC Group 28, Valent USA Corporation, Walnut Creek, CA) in similar dose ranges. Leaf dip bioassay was conducted using 1–2-mo-old cabbage plants which were grown in greenhouse condition. Cut, 6-cm diameter leaf discs were dipped in an insecticide solution with 1.8% (v/v) spreader sticker, Wetcit^® (ORO AGRI, INC., Fresno, CA), for 5 s. Control discs were treated with 1.8% Wetcit and water solution only. The leaf discs were dried at room temperature for approximately 1 h. One treated leaf disc with 10 second-to-third instars larvae were placed in a vented 100 × 15 mm Petri dish (VWR Corporation, Radnor, PA) with an extra 38-mm-diameter cut vent hole in the center of the top dish screened with nylon chiffon. For most LC₅₀ evaluations, each concentration tested consisted of three replications of 10 larvae or 30 larvae per seven insecticide concentrations (n =210; but where different, the n value is given in Table 1). The bioassay was conducted with an air-conditioned room temperature of 22.8–23.9°C and a room relative humidity of 44–46%. Mortality was monitored periodically (24, 48, 72, and 144 h). PROC PROBIT (SAS Institute 2003, SAS Institute, Cary, NC) was used for probit analysis for dose-response data and to estimate LC₅₀ values (concentration required to kill 50% of the test population). Only significant probit analyses during this time period are reported here.

Table 1 Median lethal concentrations (LC₅₀) for selected insecticides against P. xylostella populations collected at Tifton, GA. (—) indicates no ratio was calculated because no susceptible check was available.

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Table 1 Continued.

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The LC₅₀ values of the insecticides tested (Table 1) show a distinct increase in LC₅₀ values (i.e., a reduction in the efficacy) of chlorantraniliprole and spinetoram in 2016–2018 compared to 2012 and the Ithaca susceptible check from Cornell University. In 2016, control failures with both of these products were observed in Tift and Colquitt counties (D.G.R., unpubl. data). This was in spite of recommended insecticide rotations (Riley 2013, J. Entomol. Sci. 49: 130–143) purported to reduce insecticide resistance selection pressure (Zhao et al. 2010, Pest Manag. Sci. 66: 1101–1105). Both chlorantraniliprole and spinetoram were highly effective against P. xylostella at the time of commercial labeling, which led to over-use of these products for control of this pest. Given the resistance-prone nature of P. xylostella and the over-use of these materials, resistance development was a certainty.

Additional insecticides tested were observed to have higher than susceptible LC₅₀ values based on previous insecticide resistance documentation for P. xylostella for the organophosphates, carbamates, organochlorines, and pyrethroids (Sun et al. 1986, Pp. 359–371 In Talekar (ed.), Proceedings 1st International Workshop, Shanhua, Taiwan) and B. thurengiensis (Heckel et al. 2004, Pp. 27–36 In Ridland and Endersby (eds.), Proceedings 4th International Workshop, Victoria, Australia) spinosyns (Sparks et al. 2012, Pest. Biochem. Physiol. 102: 1–10.), indoxacarb (Sayyed and Wright 2006, Pest Manag. Sci. 62: 1045–1051), emamectin benzoate (Zhao et al. 2006, J. Econ. Entomol. 99: 176–181), and other diamides (Troczka et al. 2012, Insect Biochem. Mol. Biol. 42: 873–880).

One of the reasons for this widespread adaptation to insecticides is the range of resistance mechanisms occurring in P. xylostella populations (Cheema et al. 2011, Pesticide Res. J. 23: 123–134). Interestingly, lepidopteran pests have shown more development of resistance to phytochemicals (Bhandari et al. 2018, Agric. Environ. Letters 3:180037) than have hymenopteran pollinators (Bhandari et al. 2018, Crop Sci. 58: 2665–2671) and dipteran pests (Bhandari et al. 2018, Texas J. Agric. Nat. Res. 31: T1–T5). In contrast, some hymenopteran pests, such as red imported fire ants, Solenopsis invicta Buren, were found to be very susceptible to phytochemicals (Bhandari et al. 2018, Crop, Forage & Turfgrass Manag. 4:180005) due to the presence of some well-known insect-deterring compounds in the plants (Bhandari et al. 2019, Indust. Crops Prod. 133: 1–9).