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Spinosad selection of a laboratory strain of the tobacco budworm, Heliothis virescens (Lepidoptera: Noctuidae), and characterization of resistance
Crop Protection 22 (2003) 265–273
Spinosad selection of a laboratory strain of the tobacco budworm, Heliothis virescens (Lepidoptera: Noctuidae), and characterization of resistance Hugh P. Young, Woodward D. Bailey, R.M. Roe* Department of Entomology, North Carolina State University, Campus Box 7647, West Ligon Street Extension, Dearstyne-Entomology Building, Raleigh, NC 27695-7647, USA Received 11 November 2001; accepted 7 August 2002
Abstract The potential for insect resistance to the spinosyns, a novel class of insecticide chemistry, was examined using a laboratory strain of the tobacco budworm, Heliothis virescens (F.), originally collected from tobacco at sites in North Carolina. Technical grade spinosad (spinosyns A and D), was topically applied to third instars. Initially 533 third instars were used but one to two thousand larvae were treated per generation thereafter. Initially mortality ranged from 75% to 85% with doses of 0.044–0.088 mg per larva, until the fifth generation (G5) when mortality decreased. The selection dose was subsequently increased every generation from G5 to G11 in an attempt to restore mortality to >70%. After six generations of selection, the LD50 of the selected budworms was 1.68times that of the parental generation (G1) as estimated 15 d after treatment. By G14, the topical LD50 of the selected insects was 1068-fold greater than the parental generation. Four additional populations of the budworm from the southeastern US demonstrated similar LD50s to spinosad as our parental strain, suggesting that the parental budworms from North Carolina were representative of field populations elsewhere. The resistance ratio determined with spinosad (formulated as Tracers) in heliothine diet was 314-fold at 15 d after the start of exposure. Injection of spinosad into the larval hemocoel resulted in a >163-fold resistance ratio 15 d after injection, indicating that resistance could not be explained simply by altered penetration alone. Mortality was delayed in the resistant relative to the parental generation regardless of whether third instars were topically treated or exposed to treated diet. Spinosad resistance was also expressed in G14 adults, indicating that an adult vial test would be feasible for monitoring resistance. A feeding disruption assay was developed to monitor larval resistance in the field. r 2002 Elsevier Science Ltd. All rights reserved. Keywords: Spinosad; Tracer; Resistance; Tobacco budworm; Heliothis virescens
The availability and use of insecticides like the chlorinated hydrocarbons, cyclodienes, organophosphates, carbamates, pyrethroids and now Bt toxins for heliothine control have to a large degree been sequential. This has resulted in pests developing resistance to one intensively used insecticide after another (Eldefrawi and Eldefrawi, 1990). Resistance management is very important in a heavily treated crop such as cotton, and history has shown that the heliothine complex is one of the more problematic pest groups in this regard (Sparks, 1996; Sparks et al., 1993). To break this cycle there is a need for a number of different insecticides having different modes of action,
*Corresponding author. Tel./fax: +1-919-515-4325. E-mail address: michael [email protected] (R.M. Roe).
evaluation of the potential for insect resistance to these chemistries and resistance management. The spinosyns, derived from the actinomycete, Saccharopolyspora spinosa, were discovered in the 1980s; two of them, spinosyns A and D, have strong insecticidal activity with low levels of mammalian toxicity and relatively little toxicity to non-target insects (Sparks et al., 1998; Bret et al., 1997). Technical grade spinosad is the naturally occurring blend of spinosyns (Sparks et al., 1998; Bret et al., 1997). Spinosyns probably act as an agonist at the post-synaptic cholinergic ion channels and GABA-gated ion channels. In order to explore strategies for resistance management, if possible before any resistance is manifested in field populations, we began selecting a laboratory strain of the tobacco budworm, Heliothis virescens, with topically applied technical spinosad in November 1997.
0261-2194/03/$ - see front matter r 2002 Elsevier Science Ltd. All rights reserved. PII: S 0 2 6 1 - 2 1 9 4 ( 0 2 ) 0 0 1 4 7 - 3
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We herein present the results of our characterization of the level(s) of resistance of the selected strain, possible mechanisms of resistance and methods for resistance monitoring in the field.
1. Materials and methods 1.1. Chemicals Technical spinosad contains 88% (wt/wt) of active ingredient—mixture of spinosyns A and D while Tracers is a formulation containing 44.2% (wt/vol) of spinosyns A and D. Results are exclusively reported per unit weight of active ingredient (spinosyns A and D or spinosad). Hypochlorite solutions were a 50:1 (vol/vol) dilution of commercial laundry bleach. Trypan Blue (Direct Blue 14, Matheson Coleman and Bell (Norwood, OH, USA)) was used in the fecal production bioassay. 1.2. Insects and rearing A susceptible, parental strain of H. virescens was established in 1997 from egg collections from tobacco at several sites in North Carolina in 1996 and 1997. Fortyfour single pair matings from 1996 and 22 single pair matings from 1997 were the founding gene pool for the susceptible parental strain. Once the colony was established in the laboratory, they were routinely reared as follows. Pupae (40–60, unsexed) were placed in 2quart plastic containers with a gauze cover and provided a 20% (wt/vol) sucrose solution ad libtum. Eggs deposited on the gauze cover were soaked for 2 min in a 0.075% (wt/vol) hypochlorite solution to clean the eggs and release them from the cloth. Immediately after egg hatch, neonates were transferred to 10–12 ml of artificial heliothine diet (Burton, 1970) in 29.6 ml Souffle cups (Solo Cup Company, Urbana, IL). Insects were maintained at 27711C and 40–70% RH under a 14:10 (L:D) h photoperiod. The parental strain was reared in the absence of pesticides. 1.3. Selection process A stock solution of 8.8 and 17.6 mg of spinosyns A and D (active ingredient) per ml acetone was made from technical spinosad by diluting 8.8 and 17.6 mg active ingredient, respectively, in 1000 ml acetone in an Eppendorf tube. Serial dilutions were made to obtain appropriate doses in 1 ml acetone. Solutions were vortexed vigorously before use. Topical applications to insects were made with a 50 ml Hamilton syringe fitted with a 1 ml repeating dispenser. The dose was applied to the dorsal thorax of third instars. Preliminary experiments showed that a dose of 0.044 mg of spinosad (active
ingredient) per larva resulted in 75–85% mortality in the parental strain. Initially 533 larvae were selected in G1 and approximately 100 pupae were carried over into the breeding containers. Dosed larvae were placed singly on 10–12 ml of heliothine diet. Mortality was assessed at 11–13 d postdose. The criterion for death was failure to respond to a touch from a blunt probe within 10 s. Dead insects were discarded and viable pupae placed in oviposition containers as described earlier. The remaining larvae were examined at 5–7 d intervals, and pupae were setup in oviposition containers until 20–25 d had elapsed, when the selection round was terminated. 1.4. Estimation of topical dose response Third instars were topically dosed with technical spinosad as described earlier. The parental and selected generation 6 (G6) larvae were treated with five doses: 0.022, 0.044, 0.088, 0.176 and 0.352 mg spinosad (active ingredient) per larva. The selected G14 larvae were treated with 5.5, 11, 22, 44 and 88 mg of spinosad. The doses for the selected strain were applied in five 1 ml aliquots, because the saturation point of spinosad in acetone was approximately 17.6 mg/ml. G14 budworms were highly resistant, and doses >17.6 mg per larva were needed to obtain a dose–response and to estimate the LD50. In each replicate a solvent control was included, 1 ml acetone for each of the parental and G6 larvae and five 1 ml acetone aliquots for G14 larvae. Three replicates of 25 larvae per replicate were used per dose. Dosed larvae were placed individually in 29.6 ml souffle cups containing artificial diet as describe earlier. Mortality was assessed starting 3 d after treatment and daily through day 6, after which the insects were observed at 3 d intervals until all insects had either died or pupated. 1.5. Field collected strains of tobacco budworm Tobacco budworms from Johnston County, North Carolina (USA), were collected from tobacco as larvae; Washington County, Mississippi (USA) from geranium as eggs; Franklin Parish, Louisiana (USA) from velvetleaf as larvae; and Quitman, Georgia (USA) from cotton refugia (in an area of Bt cotton) as eggs. Insects from each collection site were reared in the laboratory for 2–4 generations before the susceptibility to topically applied technical spinosad was estimated. Spinosad (0.022–0.352 mg active ingredient per third instar) was applied to the dorsal thorax and mortality determined as described earlier. Three replicates of 25 third instars per replicate were used for insects from Johnston County and Franklin Parish. Two replicates of 25 larvae per replicate were used for the Washington and Quitman
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strains. An additional dose of 1.1 mg per larva was used on the Quitman strain. s
1.6. Estimation of toxicity by exposure to Tracer in diet Five-ml aliquots of diet were placed in 29.6 ml diet cups, frozen overnight at 201C, lyophilized for a minimum of 72 h (Bench Top 6, Virtis, Gardiner, NY; cold trap 701C, E200 mTorr, ambient temperature E231C) and stored in a dessicator over Drierite (Fisher Scientific, Pittsburgh, PA) until used (Roe et al., 2000). Eight to twelve cups were weighed before and after lyophilization to determine the water content of the diet (typically 80% (wt/vol) or 4 ml per cup). Dilutions of Tracers in distilled water were made as follows: 0.05, 0.5, 5, 50, 500 and 5000 mg of spinosad active ingredient/ ml. Diet was reconstituted with 4 ml of diluted Tracers or distilled water (the control) and allowed to completely rehydrate (1 h). The final concentrations of spinosad (active ingredient) in the insecticide treatments were 0.04, 0.4, 4, 40, 400 and 4000 mg of active ingredient/ml diet. Early fourth stadium larvae (parental and G14) were placed on the diet and allowed to feed for 48 h. Mortality was determined as described earlier, and the larvae transferred to fresh diet without insecticide. Larvae were then observed daily until all had died or pupated. Two replicates of 25 larvae per replicate were conducted for each dose.
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1.9. Diagnosis of larval resistance to spinosad Trypan Blue was blended with artificial diet (described earlier) at 20 mg dye per 100 ml diet. Blue diet aliquots were dispensed (100 ml per well) into 8-well microtiter plate strips (Nalge Nunc, Naperville, IL). The blue dye served as a marker for feeding on the assay diet, with larvae feeding on colored diet producing blue feces, which were easily distinguished visually from feces derived from other food sources. Microtiter plate strips containing blue diet were lyophilized and stored until needed as described earlier. Diet was hydrated with Tracers at 0.8 mg active ingredient/ml of moist diet. Feeding studies (described earlier) indicated that this concentration would discriminate spinosad resistant from susceptible budworms. Neonates from the selected (G15) and the field-collected Franklin Parish, Louisiana strains were placed individually on the diet in each well. Strip caps (Nunc) were cut into single caps to seal each well of the microtiter plate strips, which facilitated observation within wells while not allowing neighboring larvae to escape. To reduce condensation within the wells and provide larvae with air, caps were punctured twice with a #3 insect pin. Blue fecal pellets were counted after 24 h. Forty neonates were bioassayed from the Franklin strain and 46 from the laboratory-selected colony. The neonates were less than 24 h from egg hatch when placed on the diet.
1.7. Injection of larvae with technical spinosad
1.10. Data analysis and statistics
Fourth-fifth instars were injected in the perivisceral sinus with technical spinosad in 1 ml acetone. The injections were made with a 10 ml Hamilton syringe fitted with a removable 2 cm long 22-guage needle which was inserted into the hemocoel, lateral to the first or second pair of prolegs, below the spiracles. Any larvae that bled were rejected. Doses of 0.088, 0.44, 0.88, 4.4 and 8.8 mg spinosad (active ingredient)/ml were used for both parental and selected (G19) strains. An additional dose of 0.044 mg/ml was used for the susceptible parental strain. Six replicates of 10 larvae per replicate were used for each dose. Insects were observed daily until all had either died or pupated.
Abbott’s correction (Abbott, 1925) was applied to all data in the dose–response experiments. The only exception was in the selection of the spinosad resistant strain, where the results are reported as uncorrected mortality. Median lethal doses were estimated plots of probit mortality versus log dose (Finney, 1971) using the method of least squares and inverse predictions of 95% fiducial ranges (Sokal and Rohlf, 1995). The variance– covariance matrix and 95% confidence intervals for the toxicity ratios were estimated by the methods of Steel and Torrie (1980) and Robertson and Preisler (1982), respectively. Calculations were made in Microsoft Excel spreadsheets (Microsoft, 1997).
1.8. Topical treatment of adult moths 2. Results and discussion Parental and G19 pupae were separated by sex. Oneday-old female moths were topically treated on the compound eye with 0.44, 0.88, 4.4 and 8.8 mg of spinosad (active ingredient) in 1 ml acetone. The selected strain was also tested with 17.6 and 35.2 mg, the latter being applied as two 1 ml aliquots. Treated insects and controls were observed 24 h post-dose. Four replicates of five moths per replicate were used at each dose.
2.1. Selection for spinosad resistance The mortality at different selection doses determined 11–14 d after application is shown in Fig. 1A. The total number of survivors each generation that were able to pupate and were transferred to oviposition containers is shown in Fig. 1B. From the parental strain, 533 larvae
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Fig. 1. History of the laboratory selection for spinosad resistance in the tobacco budworm, H. virescens, by the topical application of technical spinosad in acetone on third instars in successive generations: (A) examples of dose applied (spinosad active ingredient) per larva and the resulting uncorrected percent mortality each generation; and (B) the total number of viable pupae produced each generation after selection. See text for full details.
were selected in G1 and as much genetic diversity as possible was maintained from G2 through G13 by treating approximately 1000–2000 third instars per generation. Approximately 100 (parental generation) to greater than 700 pupae were produced every generation (Fig. 1B). More neonates were generated than were required for subsequent selection, but we attempted to use neonates from all ovipositional containers over a 1-week period to establish the next generation. Approximately 75% mortality occurred in G1 (Fig. 1A) at this dose and greater than 70% mortality was maintained at 0.044 mg of spinosad per larva from G1 through G4. However, in G5 this dose only produced 57% mortality (Fig. 1A) while 0.088 mg per larva produced approximately 87% mortality (data not shown). In G6, 0.066 mg per larva produced only 55% mortality (Fig. 1A), suggesting that a reduction in spinosad susceptibility had occurred. To examine this possible reduction of susceptibility further, the LD50 (15 d after treatment) for topically applied spinosad on unselected G6 third instars was estimated. The LD50 was
0.193 mg per larva (Table 1). The parental strain has a median lethal dose of 0.115 mg per larva. The resistance ratio was estimated as 1.68 with a 95% confidence interval of 1.12–2.51 (Robertson and Preisler, 1982) (Table 1). Apparently, reduced susceptibility had occurred in the selected budworms by the sixth generation and maybe even as early as G5 (Fig. 1A), but the reduction of susceptibility was small. The selection dose was increased to 0.176 mg per larva in G7 and to 0.264 in G8 in order to maintain >60% mortality (Fig. 1A). Multiple treatment levels were used in G9–G13 in an attempt to maintain the selection pressure without killing all of the insects and to determine a dose that would provide greater than 50% mortality. A dose of 1.4 mg per larva in G9 produced 61% mortality, 2.3 mg per larva in G10 produced only 14% mortality, and 52.8 mg per larva resulted in only 47 and 5.5% mortality in G11 and G13, respectively (Fig. 1A). The insects were not selected in G12. An estimate of the median lethal topical dose of spinosad was made in the selected strain in G14. After generation 13, the larvae were no longer routinely selected with spinosad. The G14 budworms had an extrapolated LD50 of 122.6 mg per larva (Table 1). An extrapolation was necessary because a dose of 88 mg (5 ml 17.6 mg/ml) per larva resulted in 47.1% mortality. The extrapolated LD50 for G14 selected budworms when compared to the parental strain, demonstrated a resistance ratio of 1068 and fiducial limits of 524–2177 (Table 1). The LD45 for G14 larvae was estimated without extrapolation as 103.8 mg per larva. The parental LD45 is 0.103 mg, a resistance ratio of 1008fold, which was in reasonable agreement with the ratio of 1068-fold calculated from our extrapolated LD50. The LD50 of our parental strain (0.115 mg of technical spinosad per larva) was in the range of the topical LD50 values obtained by Leonard et al. (1996) for tobacco budworms sampled from Texas, Louisiana, and Mississippi in 1991–1992. They reported LD50s from 0.40 to 8.55 mg/g budworm. Our median dose of 0.115 mg per 30 mg larva or 3.8 mg/g falls in the middle of the range of LD50s reported from the three states examined earlier. Interestingly, our G6 LD50 was 0.193 mg per larva (Table 1) or 6.4 mg/g, which is the upper end of the range found for budworms by Leonard et al. (1996). The G14 LD50 was 122.6 mg per larva or 4087 mg/g. 2.2. Comparison with field strains A comparison of the log-dose mortality (15 d after topical application) of our parental strain to tobacco budworm strains collected from Quitman (GA) and Franklin (LA), Washington (MS), and Johnston (NC) Counties in 1998 and the G14 selected strain, is shown in Fig. 2. The strain from Quitman, GA, demonstrated a resistance ratio of 2.9-fold (fiducial limits of 1.9–4.3) as
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Table 1 Tobacco budworm larval mortality 15 d after treatment for spinosad (active ingredient) applied topically, added to diet and injected Route of entry/strain
n
Median lethal dose/conca
95% CIb
Chi-square
Slope7SE
Resistance ratioc
95% CIb
Topical Parental Selected (G6) Selected (G14)
375 300 300
0.115 mg/larva 0.193 122.6
0.03–0.68 0.09–0.50 23.9–15, 11.8
7.59 1.66 2.34
2.6670.47 3.1070.31 1.7470.29
1.68 1068
1.12–2.51 5.24–2,177
Treated diet Parental Selected (G14)
200 200
0.103 mg/ml 32.3
NAd NA
5.04 3.56
0.8870.23 0.5170.16
316.6
24.5–4012
Injected Parental Selected (G19)
240 240
0.054 mg/larva >8.8
0.014–0.143
0.722
1.1770.14 >163
a
Results were corrected for control mortality. Dose or concentration is given in mg of active ingredient. CI, confidence interval. c Resistance ratio is the median lethal dose or concentration of the selected strain divided by that for the parental. d NA=not available, confidence interval could not be calculated. b
Spodoptera exigua, with natural variations of spinosad susceptibility as much as 24-fold. 2.3. Toxicity on treated diet
Fig. 2. Log dose (active ingredient) versus probit mortality for the topical application of technical spinosad in acetone on parental, generation (G) 14 selected, and four field strains of the tobacco budworm, H. virescens. The larvae at the time of treatment were third instars. The dotted line indicates 50% mortality.
compared to the most susceptible Washington strain. Our parental strain demonstrated similar susceptibility to the field strains tested (Fig. 2) and to those studied by Leonard et al. (1996) discussed earlier. It appears that our parental strain from which we were able to obtain high levels of spinosad resistance by G14 (Fig. 2), was typical of budworms from the field with no apparent, natural resistance to spinosad. Moulton et al. (2000) found some field populations of the beet armyworm,
It was clear that G14 selected budworms were highly resistant to spinosad that was topically applied in acetone. However, in the field, budworms would most likely be exposed to the insecticide while crawling on treated plant surfaces and feeding. To more closely mimic field exposures, Tracers was added to artificial diet and the insects were allowed to feed on the treated diet for 2 d. After 2 d the insects were transferred to untreated diet. The LC50 of G14 selected budworms was 32.3 mg/ml diet 15 d after initial placement on the diet (13 d after being removed from the treated diet) compared to 0.103 mg/ml for the parental strain, a resistance ratio of 316.6-fold (Table 1). Three days after treatment (24 h after being removed from a 2 d exposure to the treated diet), the LC50 values were 0.200 and 70.7 mg/ml for parental and selected strains, respectively, a resistance ratio of 354. The G14 selected tobacco budworms are highly resistant to spinosad both when treated topically and allowed to crawl and feed on treated artificial diet. These studies also suggest that our laboratory selected budworms would likely be resistant to spinosad under field conditions where the insect might be crawling and feeding on treated plant material. 2.4. Injected toxicity of spinosad In order to determine if resistance was exclusively the result of reduce penetration of spinosad across the cuticle and digestive tract, the insecticide in acetone was injected directly into the hemolymph. At 15 d after injection, the selected strain LD50 was >8.8 mg per larva
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as compared to 0.054 mg per larva for the parental strain, a resistance ratio >163-fold (Table 1). The maximum dose tested on the resistant strain, 8.8 mg per insect, produced only 28.7% mortality. It is apparent from the injection studies that at least in part, resistance cannot be fully explained by just altered penetration either through the cuticle or digestive system. In addition, we can rule out behavioral resistance, since the pesticide is being delivered by injection. Therefore, one or more components of the resistance mechanism must be altered metabolism, sequestration, increased excretion and/or altered target site. 2.5. Delayed mortality In addition to high levels of spinosad resistance in the tobacco budworm, we also found that mortality was delayed in each strain and delayed more in the resistant as compared to the susceptible strain when larvae were either topically treated or exposed to treated diet. For example, Figs. 3A and B show the cumulative mortality of susceptible and resistant (G14) tobacco budworms, respectively, exposed for 2 d to spinosad incorporated into diet and then transferred to diet without the insecticide. In both strains, mortality increased with time in a dose- and strain-dependent fashion. In addition, mortality was also delayed more in the resistant as compared to the susceptible strain. To illustrate this, the median lethal time (LT50) was determined for susceptible and resistant budworms exposed to 50 mg of spinosad per ml diet (Fig. 4A). The LT50 is the elapsed time from when the insects were added to the treated diet and to when one-half of the larvae had died. From the plot of inverse time (1=t) versus probit mortality, the parental and selected strains demonstrated LT50s of 2.61 (95% confidence interval of 2.31–2.96) and 9.46 (7.30–13.49) d, respectively. Mortality in the selected strain was not only much lower than in the parental strain, but is delayed approximately 7 d. Again, note in both strains at the dose of 50 mg per larvae, mortality was linear with the inverse of time. Because mortality was delayed in each strain and to different degrees between strains and because in some cases the selected insects were highly resistant, median lethal doses had to be calculated in our studies at 15 d (Table 1). Also we found that delayed mortality in resistant as compared to susceptible budworms did not occur when spinosad was injected (data not shown). 2.6. Spinosad treatments delay development of survivors It was also apparent that sublethal intoxication of resistant budworms with spinosad delays their development. Fig. 4B shows that the time to 50% pupation (PT50) of the survivors exposed to 50 mg of spinosad per ml diet was 9.65 (95% confidence interval 7.77–11.38) d
Fig. 3. Cumulative percentage mortality of (A) parental and (B) generation (G) 14 selected tobacco budworms, H. virescens, allowed to feed on artificial diet containing spinosad. Late fourth stadium larvae were transferred to artificial diet containing different concentrations of Tracers and allowed to feed for 48 h. The concentrations indicated are spinosad (active ingredient) per ml artificial diet. After being allowed to feed for 48 h, mortality was determined and then the surviving larvae were transferred to diet without spinosad. Each point is the mean of 2 replicates. The dotted line indicates 50% mortality.
as compared to 7.72 (6.13–9.23) d for the control. Delayed pupation was more obvious at the PT90, 14.63 (12.87–16.80) d for the spinosad treatment as compared to 10.10 (8.60–11.77) d for the control. Liu et al. (1999) found that plants expressing the Bt toxin demonstrated sublethal effects on larval lepidopteran development, delaying pupation and adult eclosion. As a consequence, the life cycle of the resistant insect can become asynchronous with that of the susceptible
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our selected strain, larvae surviving high doses of spinosad feed at a reduced rate with minimum movement on or within the artificial diet, but successfully go through larval–larval and larval–pupal molts. These larvae will eventually pupate. The impact of this delayed development in the field may be an important factor in the promotion of spinosad resistance. However, it is also possible that delayed development, which under controlled laboratory conditions does not affect survivorship to the adult stage, might be a problem under field conditions, where nutrition, environmental stress, biological competition and other factors are not optimal. 2.7. Adult bioassay for spinosad resistance
Fig. 4. (A) Plot of inverse time versus probit mortality for parental and generation (G) 14 tobacco budworms, H. virescens. Late fourth stadium larvae of both strains were allowed to feed for 48 h on artificial diet containing Tracers (50 mg of active ingredient per ml diet) and then were transferred to diet without Tracers. See Fig. 3 for delayed mortality at other doses. Elapsed time is in days from the first exposure to the treated diet. (B) Plot of elapsed time from first exposure to spinosad in treated (50 mg of spinosad per ml) or untreated (control) diet and probit of the cumulative mortality of the larvae that pupated. The dotted line indicates 50% mortality. Data taken from the study shown in Fig. 3. The lines were fitted by least squares linear regression analysis.
population. The formation of these sympatric populations in the presence of insecticide selection can favor the establishment of resistance. We have noticed that in
A common method for monitoring resistance in Lepidoptera collected from the field is the adult vial assay (Plapp et al., 1987) or a modification of this technique. The bioassay examines the effect of a diagnostic dose of an insecticide applied topically on the ability of the insect to fly and/or on mortality at some time period after treatment. Since the spinosad resistant tobacco budworm strain described above is the first to be resistant to this novel chemistry, it was important to validate whether resistance could be detected in the adult stage as is commonly done with the adult vial test. When applied topically to the compound eye of tobacco budworms, 0.44, 0.88, 4.4 and 8.8 mg of spinosad (active ingredient) per adult moth produced 11.178.0% (1SE), 20.078.0%, 9075.8% and 7575.0% mortality in the susceptible (parental) strain and no mortality in the resistant strain 24 h after treatment. In fact, no mortality was noted in the resistant strain even at 17.6 and 35.2 mg per adult. The doses of 0.88, 4.4 and 8.8 mg per insect successfully distinguished a homogenous population of susceptible budworms from the selected strain. The dose of 0.44 mg which produced 11.178.0% mortality in the susceptible strain was not statistically different from the 0% mortality at the same dose for the resistant budworms (t-test, P ¼ 0:05). The 4.4 mg per adult treatment appeared to be the minimum optimum dose for monitoring resistance in individuals since 90% of the susceptible strain was killed by this dose and no mortality occurred in the resistant strain. These studies also demonstrated that the mechanism of larval resistance to spinosad is expressed in the adult stage. 2.8. Larval feeding disruption assay for resistance monitoring Bailey et al. (1998), Roe et al. (2000) and Bailey et al. (2001) describe an alternative method for resistance detection, a larval feeding disruption bioassay. The advantages of this approach are that it measures resistance in the larvae (a target stage for spinosad),
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and it is a rapid neonate test for resistance specific to a particular field where the eggs were collected. In contrast, the adult bioassay assumes that resistance is expressed at the same level in both larvae and adults from the same generation and that moths collected in pheromone or light traps are an accurate representation of the level of resistance in a specific field. The adults are not genetically identical to the next generation of larval pests. In some applications of the larval feeding disruption test, the assay is non-lethal, permitting resistance detection for other insecticides on the same pest sample. The principle of the feeding disruption assay is the determination of the presence or absence of blue feces when neonates are placed on artificial diet containing a blue indicator dye and a diagnostic concentration of the insecticide. The production of blue feces indicates that the insect is resistant. Several concentrations of spinosad were examined. One useful diagnostic dose was 0.8 mg of spinosad (active ingredient) per ml of artificial, blue diet. At this concentration, 100% of the susceptible budworms (Franklin strain) produced 0–5 fecal pellets and 100% of G15 budworms produced greater than 5 fecal pellets after 24 h on the treated diet (Fig. 5). Only a small fraction (2.6%) of the susceptible insects produced 3–5 pellets. These studies demonstrate that the feeding disruption assay can be used for monitoring spinosad resistance in larvae of the tobacco budworm. The resolution of the assay can be enhanced by increasing
the incubation time on the diet, and the assay can also be used as a mortality assay (Bailey et al., 1998). In summary, the selection of tobacco budworms each generation with topically applied technical spinosad produced a laboratory strain which was highly resistant to the insecticide when exposed topically, by feeding on treated diet or by injection. Spinosad mortality is delayed in both susceptible and resistant budworms but to a greater extent in the latter. Resistance is expressed in both the larval and adult stage. A feeding disruption assay was also developed to monitor larval spinosad resistance in the field.
Acknowledgements We would like to acknowledge support for this work from Dow AgroSciences, Indianapolis, IN; Cotton Incorporated, Cary, NC; the National Science Foundation Industry Center for IPM, North Carolina State University, Raleigh, NC; a Southern Regional IPM Grant; and the NC Agricultural Research Service, Raleigh, NC. We would also like to thank Dr. G.D. Thompson and Dr. T.C. Sparks of Dow AgroSciences, Indianapolis, IN, for their valuable insights during the course of our research. We thank Drs. J.S. Bacheler and C.E. Sorenson and their collaborators for their assistance in obtaining field strains of the tobacco budworm. Technical spinosad was a gift from Dr. G.D. Thompson, Dow AgroSciences, Indianapolis, Indiana, USA. Tracers was a gift from Dr. Clyde Sorenson, Department of Entomology, North Carolina State University, Raleigh, North Carolina (USA).
References
Fig. 5. Feeding disruption assay for the detection of spinosad resistance in neonates of the tobacco budworm, H. virescens. The Franklin field collected and generation (G) 15 neonates were placed on artificial diet containing 0.8 mg of spinosad (active ingredient in Tracers) per diet, and blue feces production measured after 24 h.
Abbott, W.S., 1925. A method of computing the effectiveness of an insecticide. J. Econ. Entomol. 18, 265–267. Bailey, W.D., Zhao, G., Carter, L.M., Gould, F., Kennedy, G.G., Roe, R.M., 1998. Feeding disruption bioassay for species and Bacillus thuringiensis resistance diagnosis for Heliothis virescens and Helicoverpa zea in cotton (Lepidoptera: Noctuidae). Crop Prot. 17, 591–598. Bailey, W.D., Brownie, C., Bacheler, J.S., Gould, F., Kennedy, G.G., Sorenson, S.E., Roe, R.M., 2001. Species diagnosis and Bacillus thuringiensis resistance monitoring of Heliothis virescens and Helicoverpa zea (Lepidoptera: Noctuidae) field strains from the southern United States using feeding disruption bioassays. J. Econ. Entomol. 94, 76–85. Bret, B.L., Larson, L.L., Schoonover, J.R., Sparks, T.C., Thompson, G.D., 1997. Biological properties of spinosad. Down Earth 52, 6–13. Burton, R.L., 1970. A low-cost artificial diet for the corn earworm. J. Econ. Entomol. 63, 1969–1970. Eldefrawi, M.E., Eldefrawi, A.T., 1990. Nervous-system-based insecticides. In: Kuhr, R.J., Hodgson, E. (Eds.), Safer Insecticides: Development and Use. Marcel Dekker Inc., NY, pp. 155–208.
H.P. Young et al. / Crop Protection 22 (2003) 265–273 Finney, D.J., 1971. Probit Analysis, 3rd Edition. Cambridge University Press, Cambridge, UK. Leonard, B.R., Graves, J.B., Burris, E., Micinski, S.,Mascerenhas, V., 1996. Evaluation of selected commercial and experimental insecticides against lepidopteran cotton pests in Louisiana. In: Proceedings Beltwide Cotton Conferences, National Cotton Council, Memphis, TN, pp. 825–830. Liu, Y.-B., Tabashnik, B.E., Dennehy, T.J., Patin, A.L., Bartlett, A.C., 1999. Development time and resistance to Bt crops. Nature (London) 400, 519. Moulton, J.K., Pepper, D.A., Dennehy, T.J., 2000. Beet armyworm (Spodoptera exigua) resistance to spinosad. Pest Management Sci. 56, 842–848. Plapp, Jr., F.W., McWhorter, G.M., Vance, W.E., 1987. Monitoring for pyrethroid resistance in the tobacco budworm in Texas—1986. In: Proceedings Beltwide Cotton Conferences, National Cotton Council, Memphis, TN, pp. 324–326. Robertson, J.L., Preisler, H.K., 1982. Pesticide Bioassays with Arthropods. CRC Press Inc., Boca Raton, FL.
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Roe, R.M., Bailey, W.D., Gould, F., Kennedy, G.G., 2000. Insecticide Resistance Assay. United States Patent Number 6,060,039, May 9. Sokal, R.R., Rohlf, F.J., 1995. Biometry, 3rd Edition. W.H. Freeman and Sons, NY. Sparks, T.C., 1996. Toxicology of insecticides and acaricides. In: King, E.G., Phillips, J.R., Coleman, R.J. (Eds.), Cotton Insects and Mites: Characterization and Management, The Cotton Foundation Publisher, Memphis, TN, pp. 282–322. Sparks, T.C., Graves, J.B., Leonard, B.R., 1993. Insecticide resistance and the tobacco budworm: past, present and future. In: Roe, R.M., Kuhr, R.J. (Eds.), Reviews in Pesticide Toxicology Volume 2. Toxicology Communications Inc., Raleigh, NC, pp. 149–183. Sparks, T.C., Thompson, G.D., Kirst, H.A., Hertlein, M.B., Larson, L.L., Worden, T.V., Thibault, S.T., 1998. Biological activity of the spinosyns, new fermentation derived insect control agents, on tobacco budworm (Lepidoptera: Noctuidae) larvae. J. Econ. Entomol. 91, 1277–1283. Steel, R.G.D., Torrie, J.G., 1980. Principles and Procedures of Statistics: A Biometrical Approach, 2nd Edition. McGraw-Hill, New York.
Spinosad selection of a laboratory strain of the tobacco budworm, Heliothis virescens (Lepidoptera: Noctuidae), and characterization of resistance Hugh P. Young, Woodward D. Bailey, R.M. Roe* Department of Entomology, North Carolina State University, Campus Box 7647, West Ligon Street Extension, Dearstyne-Entomology Building, Raleigh, NC 27695-7647, USA Received 11 November 2001; accepted 7 August 2002
Abstract The potential for insect resistance to the spinosyns, a novel class of insecticide chemistry, was examined using a laboratory strain of the tobacco budworm, Heliothis virescens (F.), originally collected from tobacco at sites in North Carolina. Technical grade spinosad (spinosyns A and D), was topically applied to third instars. Initially 533 third instars were used but one to two thousand larvae were treated per generation thereafter. Initially mortality ranged from 75% to 85% with doses of 0.044–0.088 mg per larva, until the fifth generation (G5) when mortality decreased. The selection dose was subsequently increased every generation from G5 to G11 in an attempt to restore mortality to >70%. After six generations of selection, the LD50 of the selected budworms was 1.68times that of the parental generation (G1) as estimated 15 d after treatment. By G14, the topical LD50 of the selected insects was 1068-fold greater than the parental generation. Four additional populations of the budworm from the southeastern US demonstrated similar LD50s to spinosad as our parental strain, suggesting that the parental budworms from North Carolina were representative of field populations elsewhere. The resistance ratio determined with spinosad (formulated as Tracers) in heliothine diet was 314-fold at 15 d after the start of exposure. Injection of spinosad into the larval hemocoel resulted in a >163-fold resistance ratio 15 d after injection, indicating that resistance could not be explained simply by altered penetration alone. Mortality was delayed in the resistant relative to the parental generation regardless of whether third instars were topically treated or exposed to treated diet. Spinosad resistance was also expressed in G14 adults, indicating that an adult vial test would be feasible for monitoring resistance. A feeding disruption assay was developed to monitor larval resistance in the field. r 2002 Elsevier Science Ltd. All rights reserved. Keywords: Spinosad; Tracer; Resistance; Tobacco budworm; Heliothis virescens
The availability and use of insecticides like the chlorinated hydrocarbons, cyclodienes, organophosphates, carbamates, pyrethroids and now Bt toxins for heliothine control have to a large degree been sequential. This has resulted in pests developing resistance to one intensively used insecticide after another (Eldefrawi and Eldefrawi, 1990). Resistance management is very important in a heavily treated crop such as cotton, and history has shown that the heliothine complex is one of the more problematic pest groups in this regard (Sparks, 1996; Sparks et al., 1993). To break this cycle there is a need for a number of different insecticides having different modes of action,
*Corresponding author. Tel./fax: +1-919-515-4325. E-mail address: michael [email protected] (R.M. Roe).
evaluation of the potential for insect resistance to these chemistries and resistance management. The spinosyns, derived from the actinomycete, Saccharopolyspora spinosa, were discovered in the 1980s; two of them, spinosyns A and D, have strong insecticidal activity with low levels of mammalian toxicity and relatively little toxicity to non-target insects (Sparks et al., 1998; Bret et al., 1997). Technical grade spinosad is the naturally occurring blend of spinosyns (Sparks et al., 1998; Bret et al., 1997). Spinosyns probably act as an agonist at the post-synaptic cholinergic ion channels and GABA-gated ion channels. In order to explore strategies for resistance management, if possible before any resistance is manifested in field populations, we began selecting a laboratory strain of the tobacco budworm, Heliothis virescens, with topically applied technical spinosad in November 1997.
0261-2194/03/$ - see front matter r 2002 Elsevier Science Ltd. All rights reserved. PII: S 0 2 6 1 - 2 1 9 4 ( 0 2 ) 0 0 1 4 7 - 3
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We herein present the results of our characterization of the level(s) of resistance of the selected strain, possible mechanisms of resistance and methods for resistance monitoring in the field.
1. Materials and methods 1.1. Chemicals Technical spinosad contains 88% (wt/wt) of active ingredient—mixture of spinosyns A and D while Tracers is a formulation containing 44.2% (wt/vol) of spinosyns A and D. Results are exclusively reported per unit weight of active ingredient (spinosyns A and D or spinosad). Hypochlorite solutions were a 50:1 (vol/vol) dilution of commercial laundry bleach. Trypan Blue (Direct Blue 14, Matheson Coleman and Bell (Norwood, OH, USA)) was used in the fecal production bioassay. 1.2. Insects and rearing A susceptible, parental strain of H. virescens was established in 1997 from egg collections from tobacco at several sites in North Carolina in 1996 and 1997. Fortyfour single pair matings from 1996 and 22 single pair matings from 1997 were the founding gene pool for the susceptible parental strain. Once the colony was established in the laboratory, they were routinely reared as follows. Pupae (40–60, unsexed) were placed in 2quart plastic containers with a gauze cover and provided a 20% (wt/vol) sucrose solution ad libtum. Eggs deposited on the gauze cover were soaked for 2 min in a 0.075% (wt/vol) hypochlorite solution to clean the eggs and release them from the cloth. Immediately after egg hatch, neonates were transferred to 10–12 ml of artificial heliothine diet (Burton, 1970) in 29.6 ml Souffle cups (Solo Cup Company, Urbana, IL). Insects were maintained at 27711C and 40–70% RH under a 14:10 (L:D) h photoperiod. The parental strain was reared in the absence of pesticides. 1.3. Selection process A stock solution of 8.8 and 17.6 mg of spinosyns A and D (active ingredient) per ml acetone was made from technical spinosad by diluting 8.8 and 17.6 mg active ingredient, respectively, in 1000 ml acetone in an Eppendorf tube. Serial dilutions were made to obtain appropriate doses in 1 ml acetone. Solutions were vortexed vigorously before use. Topical applications to insects were made with a 50 ml Hamilton syringe fitted with a 1 ml repeating dispenser. The dose was applied to the dorsal thorax of third instars. Preliminary experiments showed that a dose of 0.044 mg of spinosad (active
ingredient) per larva resulted in 75–85% mortality in the parental strain. Initially 533 larvae were selected in G1 and approximately 100 pupae were carried over into the breeding containers. Dosed larvae were placed singly on 10–12 ml of heliothine diet. Mortality was assessed at 11–13 d postdose. The criterion for death was failure to respond to a touch from a blunt probe within 10 s. Dead insects were discarded and viable pupae placed in oviposition containers as described earlier. The remaining larvae were examined at 5–7 d intervals, and pupae were setup in oviposition containers until 20–25 d had elapsed, when the selection round was terminated. 1.4. Estimation of topical dose response Third instars were topically dosed with technical spinosad as described earlier. The parental and selected generation 6 (G6) larvae were treated with five doses: 0.022, 0.044, 0.088, 0.176 and 0.352 mg spinosad (active ingredient) per larva. The selected G14 larvae were treated with 5.5, 11, 22, 44 and 88 mg of spinosad. The doses for the selected strain were applied in five 1 ml aliquots, because the saturation point of spinosad in acetone was approximately 17.6 mg/ml. G14 budworms were highly resistant, and doses >17.6 mg per larva were needed to obtain a dose–response and to estimate the LD50. In each replicate a solvent control was included, 1 ml acetone for each of the parental and G6 larvae and five 1 ml acetone aliquots for G14 larvae. Three replicates of 25 larvae per replicate were used per dose. Dosed larvae were placed individually in 29.6 ml souffle cups containing artificial diet as describe earlier. Mortality was assessed starting 3 d after treatment and daily through day 6, after which the insects were observed at 3 d intervals until all insects had either died or pupated. 1.5. Field collected strains of tobacco budworm Tobacco budworms from Johnston County, North Carolina (USA), were collected from tobacco as larvae; Washington County, Mississippi (USA) from geranium as eggs; Franklin Parish, Louisiana (USA) from velvetleaf as larvae; and Quitman, Georgia (USA) from cotton refugia (in an area of Bt cotton) as eggs. Insects from each collection site were reared in the laboratory for 2–4 generations before the susceptibility to topically applied technical spinosad was estimated. Spinosad (0.022–0.352 mg active ingredient per third instar) was applied to the dorsal thorax and mortality determined as described earlier. Three replicates of 25 third instars per replicate were used for insects from Johnston County and Franklin Parish. Two replicates of 25 larvae per replicate were used for the Washington and Quitman
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strains. An additional dose of 1.1 mg per larva was used on the Quitman strain. s
1.6. Estimation of toxicity by exposure to Tracer in diet Five-ml aliquots of diet were placed in 29.6 ml diet cups, frozen overnight at 201C, lyophilized for a minimum of 72 h (Bench Top 6, Virtis, Gardiner, NY; cold trap 701C, E200 mTorr, ambient temperature E231C) and stored in a dessicator over Drierite (Fisher Scientific, Pittsburgh, PA) until used (Roe et al., 2000). Eight to twelve cups were weighed before and after lyophilization to determine the water content of the diet (typically 80% (wt/vol) or 4 ml per cup). Dilutions of Tracers in distilled water were made as follows: 0.05, 0.5, 5, 50, 500 and 5000 mg of spinosad active ingredient/ ml. Diet was reconstituted with 4 ml of diluted Tracers or distilled water (the control) and allowed to completely rehydrate (1 h). The final concentrations of spinosad (active ingredient) in the insecticide treatments were 0.04, 0.4, 4, 40, 400 and 4000 mg of active ingredient/ml diet. Early fourth stadium larvae (parental and G14) were placed on the diet and allowed to feed for 48 h. Mortality was determined as described earlier, and the larvae transferred to fresh diet without insecticide. Larvae were then observed daily until all had died or pupated. Two replicates of 25 larvae per replicate were conducted for each dose.
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1.9. Diagnosis of larval resistance to spinosad Trypan Blue was blended with artificial diet (described earlier) at 20 mg dye per 100 ml diet. Blue diet aliquots were dispensed (100 ml per well) into 8-well microtiter plate strips (Nalge Nunc, Naperville, IL). The blue dye served as a marker for feeding on the assay diet, with larvae feeding on colored diet producing blue feces, which were easily distinguished visually from feces derived from other food sources. Microtiter plate strips containing blue diet were lyophilized and stored until needed as described earlier. Diet was hydrated with Tracers at 0.8 mg active ingredient/ml of moist diet. Feeding studies (described earlier) indicated that this concentration would discriminate spinosad resistant from susceptible budworms. Neonates from the selected (G15) and the field-collected Franklin Parish, Louisiana strains were placed individually on the diet in each well. Strip caps (Nunc) were cut into single caps to seal each well of the microtiter plate strips, which facilitated observation within wells while not allowing neighboring larvae to escape. To reduce condensation within the wells and provide larvae with air, caps were punctured twice with a #3 insect pin. Blue fecal pellets were counted after 24 h. Forty neonates were bioassayed from the Franklin strain and 46 from the laboratory-selected colony. The neonates were less than 24 h from egg hatch when placed on the diet.
1.7. Injection of larvae with technical spinosad
1.10. Data analysis and statistics
Fourth-fifth instars were injected in the perivisceral sinus with technical spinosad in 1 ml acetone. The injections were made with a 10 ml Hamilton syringe fitted with a removable 2 cm long 22-guage needle which was inserted into the hemocoel, lateral to the first or second pair of prolegs, below the spiracles. Any larvae that bled were rejected. Doses of 0.088, 0.44, 0.88, 4.4 and 8.8 mg spinosad (active ingredient)/ml were used for both parental and selected (G19) strains. An additional dose of 0.044 mg/ml was used for the susceptible parental strain. Six replicates of 10 larvae per replicate were used for each dose. Insects were observed daily until all had either died or pupated.
Abbott’s correction (Abbott, 1925) was applied to all data in the dose–response experiments. The only exception was in the selection of the spinosad resistant strain, where the results are reported as uncorrected mortality. Median lethal doses were estimated plots of probit mortality versus log dose (Finney, 1971) using the method of least squares and inverse predictions of 95% fiducial ranges (Sokal and Rohlf, 1995). The variance– covariance matrix and 95% confidence intervals for the toxicity ratios were estimated by the methods of Steel and Torrie (1980) and Robertson and Preisler (1982), respectively. Calculations were made in Microsoft Excel spreadsheets (Microsoft, 1997).
1.8. Topical treatment of adult moths 2. Results and discussion Parental and G19 pupae were separated by sex. Oneday-old female moths were topically treated on the compound eye with 0.44, 0.88, 4.4 and 8.8 mg of spinosad (active ingredient) in 1 ml acetone. The selected strain was also tested with 17.6 and 35.2 mg, the latter being applied as two 1 ml aliquots. Treated insects and controls were observed 24 h post-dose. Four replicates of five moths per replicate were used at each dose.
2.1. Selection for spinosad resistance The mortality at different selection doses determined 11–14 d after application is shown in Fig. 1A. The total number of survivors each generation that were able to pupate and were transferred to oviposition containers is shown in Fig. 1B. From the parental strain, 533 larvae
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Fig. 1. History of the laboratory selection for spinosad resistance in the tobacco budworm, H. virescens, by the topical application of technical spinosad in acetone on third instars in successive generations: (A) examples of dose applied (spinosad active ingredient) per larva and the resulting uncorrected percent mortality each generation; and (B) the total number of viable pupae produced each generation after selection. See text for full details.
were selected in G1 and as much genetic diversity as possible was maintained from G2 through G13 by treating approximately 1000–2000 third instars per generation. Approximately 100 (parental generation) to greater than 700 pupae were produced every generation (Fig. 1B). More neonates were generated than were required for subsequent selection, but we attempted to use neonates from all ovipositional containers over a 1-week period to establish the next generation. Approximately 75% mortality occurred in G1 (Fig. 1A) at this dose and greater than 70% mortality was maintained at 0.044 mg of spinosad per larva from G1 through G4. However, in G5 this dose only produced 57% mortality (Fig. 1A) while 0.088 mg per larva produced approximately 87% mortality (data not shown). In G6, 0.066 mg per larva produced only 55% mortality (Fig. 1A), suggesting that a reduction in spinosad susceptibility had occurred. To examine this possible reduction of susceptibility further, the LD50 (15 d after treatment) for topically applied spinosad on unselected G6 third instars was estimated. The LD50 was
0.193 mg per larva (Table 1). The parental strain has a median lethal dose of 0.115 mg per larva. The resistance ratio was estimated as 1.68 with a 95% confidence interval of 1.12–2.51 (Robertson and Preisler, 1982) (Table 1). Apparently, reduced susceptibility had occurred in the selected budworms by the sixth generation and maybe even as early as G5 (Fig. 1A), but the reduction of susceptibility was small. The selection dose was increased to 0.176 mg per larva in G7 and to 0.264 in G8 in order to maintain >60% mortality (Fig. 1A). Multiple treatment levels were used in G9–G13 in an attempt to maintain the selection pressure without killing all of the insects and to determine a dose that would provide greater than 50% mortality. A dose of 1.4 mg per larva in G9 produced 61% mortality, 2.3 mg per larva in G10 produced only 14% mortality, and 52.8 mg per larva resulted in only 47 and 5.5% mortality in G11 and G13, respectively (Fig. 1A). The insects were not selected in G12. An estimate of the median lethal topical dose of spinosad was made in the selected strain in G14. After generation 13, the larvae were no longer routinely selected with spinosad. The G14 budworms had an extrapolated LD50 of 122.6 mg per larva (Table 1). An extrapolation was necessary because a dose of 88 mg (5 ml 17.6 mg/ml) per larva resulted in 47.1% mortality. The extrapolated LD50 for G14 selected budworms when compared to the parental strain, demonstrated a resistance ratio of 1068 and fiducial limits of 524–2177 (Table 1). The LD45 for G14 larvae was estimated without extrapolation as 103.8 mg per larva. The parental LD45 is 0.103 mg, a resistance ratio of 1008fold, which was in reasonable agreement with the ratio of 1068-fold calculated from our extrapolated LD50. The LD50 of our parental strain (0.115 mg of technical spinosad per larva) was in the range of the topical LD50 values obtained by Leonard et al. (1996) for tobacco budworms sampled from Texas, Louisiana, and Mississippi in 1991–1992. They reported LD50s from 0.40 to 8.55 mg/g budworm. Our median dose of 0.115 mg per 30 mg larva or 3.8 mg/g falls in the middle of the range of LD50s reported from the three states examined earlier. Interestingly, our G6 LD50 was 0.193 mg per larva (Table 1) or 6.4 mg/g, which is the upper end of the range found for budworms by Leonard et al. (1996). The G14 LD50 was 122.6 mg per larva or 4087 mg/g. 2.2. Comparison with field strains A comparison of the log-dose mortality (15 d after topical application) of our parental strain to tobacco budworm strains collected from Quitman (GA) and Franklin (LA), Washington (MS), and Johnston (NC) Counties in 1998 and the G14 selected strain, is shown in Fig. 2. The strain from Quitman, GA, demonstrated a resistance ratio of 2.9-fold (fiducial limits of 1.9–4.3) as
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Table 1 Tobacco budworm larval mortality 15 d after treatment for spinosad (active ingredient) applied topically, added to diet and injected Route of entry/strain
n
Median lethal dose/conca
95% CIb
Chi-square
Slope7SE
Resistance ratioc
95% CIb
Topical Parental Selected (G6) Selected (G14)
375 300 300
0.115 mg/larva 0.193 122.6
0.03–0.68 0.09–0.50 23.9–15, 11.8
7.59 1.66 2.34
2.6670.47 3.1070.31 1.7470.29
1.68 1068
1.12–2.51 5.24–2,177
Treated diet Parental Selected (G14)
200 200
0.103 mg/ml 32.3
NAd NA
5.04 3.56
0.8870.23 0.5170.16
316.6
24.5–4012
Injected Parental Selected (G19)
240 240
0.054 mg/larva >8.8
0.014–0.143
0.722
1.1770.14 >163
a
Results were corrected for control mortality. Dose or concentration is given in mg of active ingredient. CI, confidence interval. c Resistance ratio is the median lethal dose or concentration of the selected strain divided by that for the parental. d NA=not available, confidence interval could not be calculated. b
Spodoptera exigua, with natural variations of spinosad susceptibility as much as 24-fold. 2.3. Toxicity on treated diet
Fig. 2. Log dose (active ingredient) versus probit mortality for the topical application of technical spinosad in acetone on parental, generation (G) 14 selected, and four field strains of the tobacco budworm, H. virescens. The larvae at the time of treatment were third instars. The dotted line indicates 50% mortality.
compared to the most susceptible Washington strain. Our parental strain demonstrated similar susceptibility to the field strains tested (Fig. 2) and to those studied by Leonard et al. (1996) discussed earlier. It appears that our parental strain from which we were able to obtain high levels of spinosad resistance by G14 (Fig. 2), was typical of budworms from the field with no apparent, natural resistance to spinosad. Moulton et al. (2000) found some field populations of the beet armyworm,
It was clear that G14 selected budworms were highly resistant to spinosad that was topically applied in acetone. However, in the field, budworms would most likely be exposed to the insecticide while crawling on treated plant surfaces and feeding. To more closely mimic field exposures, Tracers was added to artificial diet and the insects were allowed to feed on the treated diet for 2 d. After 2 d the insects were transferred to untreated diet. The LC50 of G14 selected budworms was 32.3 mg/ml diet 15 d after initial placement on the diet (13 d after being removed from the treated diet) compared to 0.103 mg/ml for the parental strain, a resistance ratio of 316.6-fold (Table 1). Three days after treatment (24 h after being removed from a 2 d exposure to the treated diet), the LC50 values were 0.200 and 70.7 mg/ml for parental and selected strains, respectively, a resistance ratio of 354. The G14 selected tobacco budworms are highly resistant to spinosad both when treated topically and allowed to crawl and feed on treated artificial diet. These studies also suggest that our laboratory selected budworms would likely be resistant to spinosad under field conditions where the insect might be crawling and feeding on treated plant material. 2.4. Injected toxicity of spinosad In order to determine if resistance was exclusively the result of reduce penetration of spinosad across the cuticle and digestive tract, the insecticide in acetone was injected directly into the hemolymph. At 15 d after injection, the selected strain LD50 was >8.8 mg per larva
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as compared to 0.054 mg per larva for the parental strain, a resistance ratio >163-fold (Table 1). The maximum dose tested on the resistant strain, 8.8 mg per insect, produced only 28.7% mortality. It is apparent from the injection studies that at least in part, resistance cannot be fully explained by just altered penetration either through the cuticle or digestive system. In addition, we can rule out behavioral resistance, since the pesticide is being delivered by injection. Therefore, one or more components of the resistance mechanism must be altered metabolism, sequestration, increased excretion and/or altered target site. 2.5. Delayed mortality In addition to high levels of spinosad resistance in the tobacco budworm, we also found that mortality was delayed in each strain and delayed more in the resistant as compared to the susceptible strain when larvae were either topically treated or exposed to treated diet. For example, Figs. 3A and B show the cumulative mortality of susceptible and resistant (G14) tobacco budworms, respectively, exposed for 2 d to spinosad incorporated into diet and then transferred to diet without the insecticide. In both strains, mortality increased with time in a dose- and strain-dependent fashion. In addition, mortality was also delayed more in the resistant as compared to the susceptible strain. To illustrate this, the median lethal time (LT50) was determined for susceptible and resistant budworms exposed to 50 mg of spinosad per ml diet (Fig. 4A). The LT50 is the elapsed time from when the insects were added to the treated diet and to when one-half of the larvae had died. From the plot of inverse time (1=t) versus probit mortality, the parental and selected strains demonstrated LT50s of 2.61 (95% confidence interval of 2.31–2.96) and 9.46 (7.30–13.49) d, respectively. Mortality in the selected strain was not only much lower than in the parental strain, but is delayed approximately 7 d. Again, note in both strains at the dose of 50 mg per larvae, mortality was linear with the inverse of time. Because mortality was delayed in each strain and to different degrees between strains and because in some cases the selected insects were highly resistant, median lethal doses had to be calculated in our studies at 15 d (Table 1). Also we found that delayed mortality in resistant as compared to susceptible budworms did not occur when spinosad was injected (data not shown). 2.6. Spinosad treatments delay development of survivors It was also apparent that sublethal intoxication of resistant budworms with spinosad delays their development. Fig. 4B shows that the time to 50% pupation (PT50) of the survivors exposed to 50 mg of spinosad per ml diet was 9.65 (95% confidence interval 7.77–11.38) d
Fig. 3. Cumulative percentage mortality of (A) parental and (B) generation (G) 14 selected tobacco budworms, H. virescens, allowed to feed on artificial diet containing spinosad. Late fourth stadium larvae were transferred to artificial diet containing different concentrations of Tracers and allowed to feed for 48 h. The concentrations indicated are spinosad (active ingredient) per ml artificial diet. After being allowed to feed for 48 h, mortality was determined and then the surviving larvae were transferred to diet without spinosad. Each point is the mean of 2 replicates. The dotted line indicates 50% mortality.
as compared to 7.72 (6.13–9.23) d for the control. Delayed pupation was more obvious at the PT90, 14.63 (12.87–16.80) d for the spinosad treatment as compared to 10.10 (8.60–11.77) d for the control. Liu et al. (1999) found that plants expressing the Bt toxin demonstrated sublethal effects on larval lepidopteran development, delaying pupation and adult eclosion. As a consequence, the life cycle of the resistant insect can become asynchronous with that of the susceptible
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our selected strain, larvae surviving high doses of spinosad feed at a reduced rate with minimum movement on or within the artificial diet, but successfully go through larval–larval and larval–pupal molts. These larvae will eventually pupate. The impact of this delayed development in the field may be an important factor in the promotion of spinosad resistance. However, it is also possible that delayed development, which under controlled laboratory conditions does not affect survivorship to the adult stage, might be a problem under field conditions, where nutrition, environmental stress, biological competition and other factors are not optimal. 2.7. Adult bioassay for spinosad resistance
Fig. 4. (A) Plot of inverse time versus probit mortality for parental and generation (G) 14 tobacco budworms, H. virescens. Late fourth stadium larvae of both strains were allowed to feed for 48 h on artificial diet containing Tracers (50 mg of active ingredient per ml diet) and then were transferred to diet without Tracers. See Fig. 3 for delayed mortality at other doses. Elapsed time is in days from the first exposure to the treated diet. (B) Plot of elapsed time from first exposure to spinosad in treated (50 mg of spinosad per ml) or untreated (control) diet and probit of the cumulative mortality of the larvae that pupated. The dotted line indicates 50% mortality. Data taken from the study shown in Fig. 3. The lines were fitted by least squares linear regression analysis.
population. The formation of these sympatric populations in the presence of insecticide selection can favor the establishment of resistance. We have noticed that in
A common method for monitoring resistance in Lepidoptera collected from the field is the adult vial assay (Plapp et al., 1987) or a modification of this technique. The bioassay examines the effect of a diagnostic dose of an insecticide applied topically on the ability of the insect to fly and/or on mortality at some time period after treatment. Since the spinosad resistant tobacco budworm strain described above is the first to be resistant to this novel chemistry, it was important to validate whether resistance could be detected in the adult stage as is commonly done with the adult vial test. When applied topically to the compound eye of tobacco budworms, 0.44, 0.88, 4.4 and 8.8 mg of spinosad (active ingredient) per adult moth produced 11.178.0% (1SE), 20.078.0%, 9075.8% and 7575.0% mortality in the susceptible (parental) strain and no mortality in the resistant strain 24 h after treatment. In fact, no mortality was noted in the resistant strain even at 17.6 and 35.2 mg per adult. The doses of 0.88, 4.4 and 8.8 mg per insect successfully distinguished a homogenous population of susceptible budworms from the selected strain. The dose of 0.44 mg which produced 11.178.0% mortality in the susceptible strain was not statistically different from the 0% mortality at the same dose for the resistant budworms (t-test, P ¼ 0:05). The 4.4 mg per adult treatment appeared to be the minimum optimum dose for monitoring resistance in individuals since 90% of the susceptible strain was killed by this dose and no mortality occurred in the resistant strain. These studies also demonstrated that the mechanism of larval resistance to spinosad is expressed in the adult stage. 2.8. Larval feeding disruption assay for resistance monitoring Bailey et al. (1998), Roe et al. (2000) and Bailey et al. (2001) describe an alternative method for resistance detection, a larval feeding disruption bioassay. The advantages of this approach are that it measures resistance in the larvae (a target stage for spinosad),
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and it is a rapid neonate test for resistance specific to a particular field where the eggs were collected. In contrast, the adult bioassay assumes that resistance is expressed at the same level in both larvae and adults from the same generation and that moths collected in pheromone or light traps are an accurate representation of the level of resistance in a specific field. The adults are not genetically identical to the next generation of larval pests. In some applications of the larval feeding disruption test, the assay is non-lethal, permitting resistance detection for other insecticides on the same pest sample. The principle of the feeding disruption assay is the determination of the presence or absence of blue feces when neonates are placed on artificial diet containing a blue indicator dye and a diagnostic concentration of the insecticide. The production of blue feces indicates that the insect is resistant. Several concentrations of spinosad were examined. One useful diagnostic dose was 0.8 mg of spinosad (active ingredient) per ml of artificial, blue diet. At this concentration, 100% of the susceptible budworms (Franklin strain) produced 0–5 fecal pellets and 100% of G15 budworms produced greater than 5 fecal pellets after 24 h on the treated diet (Fig. 5). Only a small fraction (2.6%) of the susceptible insects produced 3–5 pellets. These studies demonstrate that the feeding disruption assay can be used for monitoring spinosad resistance in larvae of the tobacco budworm. The resolution of the assay can be enhanced by increasing
the incubation time on the diet, and the assay can also be used as a mortality assay (Bailey et al., 1998). In summary, the selection of tobacco budworms each generation with topically applied technical spinosad produced a laboratory strain which was highly resistant to the insecticide when exposed topically, by feeding on treated diet or by injection. Spinosad mortality is delayed in both susceptible and resistant budworms but to a greater extent in the latter. Resistance is expressed in both the larval and adult stage. A feeding disruption assay was also developed to monitor larval spinosad resistance in the field.
Acknowledgements We would like to acknowledge support for this work from Dow AgroSciences, Indianapolis, IN; Cotton Incorporated, Cary, NC; the National Science Foundation Industry Center for IPM, North Carolina State University, Raleigh, NC; a Southern Regional IPM Grant; and the NC Agricultural Research Service, Raleigh, NC. We would also like to thank Dr. G.D. Thompson and Dr. T.C. Sparks of Dow AgroSciences, Indianapolis, IN, for their valuable insights during the course of our research. We thank Drs. J.S. Bacheler and C.E. Sorenson and their collaborators for their assistance in obtaining field strains of the tobacco budworm. Technical spinosad was a gift from Dr. G.D. Thompson, Dow AgroSciences, Indianapolis, Indiana, USA. Tracers was a gift from Dr. Clyde Sorenson, Department of Entomology, North Carolina State University, Raleigh, North Carolina (USA).
References
Fig. 5. Feeding disruption assay for the detection of spinosad resistance in neonates of the tobacco budworm, H. virescens. The Franklin field collected and generation (G) 15 neonates were placed on artificial diet containing 0.8 mg of spinosad (active ingredient in Tracers) per diet, and blue feces production measured after 24 h.
Abbott, W.S., 1925. A method of computing the effectiveness of an insecticide. J. Econ. Entomol. 18, 265–267. Bailey, W.D., Zhao, G., Carter, L.M., Gould, F., Kennedy, G.G., Roe, R.M., 1998. Feeding disruption bioassay for species and Bacillus thuringiensis resistance diagnosis for Heliothis virescens and Helicoverpa zea in cotton (Lepidoptera: Noctuidae). Crop Prot. 17, 591–598. Bailey, W.D., Brownie, C., Bacheler, J.S., Gould, F., Kennedy, G.G., Sorenson, S.E., Roe, R.M., 2001. Species diagnosis and Bacillus thuringiensis resistance monitoring of Heliothis virescens and Helicoverpa zea (Lepidoptera: Noctuidae) field strains from the southern United States using feeding disruption bioassays. J. Econ. Entomol. 94, 76–85. Bret, B.L., Larson, L.L., Schoonover, J.R., Sparks, T.C., Thompson, G.D., 1997. Biological properties of spinosad. Down Earth 52, 6–13. Burton, R.L., 1970. A low-cost artificial diet for the corn earworm. J. Econ. Entomol. 63, 1969–1970. Eldefrawi, M.E., Eldefrawi, A.T., 1990. Nervous-system-based insecticides. In: Kuhr, R.J., Hodgson, E. (Eds.), Safer Insecticides: Development and Use. Marcel Dekker Inc., NY, pp. 155–208.
H.P. Young et al. / Crop Protection 22 (2003) 265–273 Finney, D.J., 1971. Probit Analysis, 3rd Edition. Cambridge University Press, Cambridge, UK. Leonard, B.R., Graves, J.B., Burris, E., Micinski, S.,Mascerenhas, V., 1996. Evaluation of selected commercial and experimental insecticides against lepidopteran cotton pests in Louisiana. In: Proceedings Beltwide Cotton Conferences, National Cotton Council, Memphis, TN, pp. 825–830. Liu, Y.-B., Tabashnik, B.E., Dennehy, T.J., Patin, A.L., Bartlett, A.C., 1999. Development time and resistance to Bt crops. Nature (London) 400, 519. Moulton, J.K., Pepper, D.A., Dennehy, T.J., 2000. Beet armyworm (Spodoptera exigua) resistance to spinosad. Pest Management Sci. 56, 842–848. Plapp, Jr., F.W., McWhorter, G.M., Vance, W.E., 1987. Monitoring for pyrethroid resistance in the tobacco budworm in Texas—1986. In: Proceedings Beltwide Cotton Conferences, National Cotton Council, Memphis, TN, pp. 324–326. Robertson, J.L., Preisler, H.K., 1982. Pesticide Bioassays with Arthropods. CRC Press Inc., Boca Raton, FL.
273
Roe, R.M., Bailey, W.D., Gould, F., Kennedy, G.G., 2000. Insecticide Resistance Assay. United States Patent Number 6,060,039, May 9. Sokal, R.R., Rohlf, F.J., 1995. Biometry, 3rd Edition. W.H. Freeman and Sons, NY. Sparks, T.C., 1996. Toxicology of insecticides and acaricides. In: King, E.G., Phillips, J.R., Coleman, R.J. (Eds.), Cotton Insects and Mites: Characterization and Management, The Cotton Foundation Publisher, Memphis, TN, pp. 282–322. Sparks, T.C., Graves, J.B., Leonard, B.R., 1993. Insecticide resistance and the tobacco budworm: past, present and future. In: Roe, R.M., Kuhr, R.J. (Eds.), Reviews in Pesticide Toxicology Volume 2. Toxicology Communications Inc., Raleigh, NC, pp. 149–183. Sparks, T.C., Thompson, G.D., Kirst, H.A., Hertlein, M.B., Larson, L.L., Worden, T.V., Thibault, S.T., 1998. Biological activity of the spinosyns, new fermentation derived insect control agents, on tobacco budworm (Lepidoptera: Noctuidae) larvae. J. Econ. Entomol. 91, 1277–1283. Steel, R.G.D., Torrie, J.G., 1980. Principles and Procedures of Statistics: A Biometrical Approach, 2nd Edition. McGraw-Hill, New York.