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Resistance monitoring of Heliothis virescens to pyramided cotton varieties with a hydrateable, artificial cotton leaf bioassay
Crop Protection 30 (2011) 1196e1201
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Resistance monitoring of Heliothis virescens to pyramided cotton varieties with a hydrateable, artificial cotton leaf bioassay Ana R. Cabrera, Jaap Van Kretschmar, Jack S. Bacheler, Hannah Burrack, Clyde E. Sorenson, R. Michael Roe* North Carolina State University, Department of Entomology, Campus Box 7647, Raleigh, NC 27695-7647, USA
a r t i c l e i n f o
a b s t r a c t
Article history: Received 1 December 2010 Received in revised form 6 May 2011 Accepted 7 May 2011
Proof of concept was demonstrated for a practical, off-the-shelf bioassay to monitor for tobacco budworm resistance to pyramided Bt cotton using plant filtrates. The bioassay was based on a previously described feeding disruption test using hydrateable artificial diet containing a blue indicator dye, a diagnostic dose of insecticide and novel assay architecture. Using neonate larvae from a Bt-susceptible, laboratory reared tobacco budworm strain, a diagnostic dose for Bollgard II and WideStrike cotton was obtained that limited neonate blue fecal production to 0e2 pellets in 24 h (Bt-resistant larvae produced >2 fecal pellets). The bioassay was tested with three different field populations of tobacco budworm collected from tobacco in central North Carolina (USA) and shown to accurately diagnose susceptibility to Bt. The diagnostic doses were also successfully evaluated with two Bt-resistant, laboratory reared tobacco budworm strains. Shelf life studies showed the assay could be stored for at least 6 months at room temperature (longer storage times were not studied). The application of the bioassay as an easy to use monitoring tool is discussed. Ó 2011 Elsevier Ltd. All rights reserved.
Keywords: Bt Tobacco budworm Heliothis virescens Cotton Bollgard II WideStrike Resistance
1. Introduction The cultivation of transgenic crops expressing Bacillus thuringiensis Berliner (Bt) toxins to control insect pests provides both economical (Shelton et al., 2002) and environmental benefits from reductions in chemical insecticide use (Ferré and Van Rie, 2002), effective control of pests and minimal impact on non-target organisms (Sisterson et al., 2004). Since their introduction in the 1990s, the proportion of the U.S. acreage planted with Bt cotton cultivars has grown steadily. However, one of the major concerns is the development of insect resistance in the target pest due to their constant exposure to the toxin (Ferré and Van Rie, 2002; Gould, 1998). Possible resistance mechanisms to the Bt toxins include a reduction of proteolytic activity in the insect gut, Bt receptor modifications and midgut cell regeneration or replacement (Ferré and Van Rie, 2002). Before the commercialization of “pyramided” cultivars expressing multiple Bt toxins, Gould (1998) suggested that stacked genes could reduce the risk of resistance evolving based on field studies showing low frequency of resistance alleles. As an
* Corresponding author. Tel./fax: þ1 919 515 4325. E-mail address: [email protected] (R.M. Roe). 0261-2194/$ e see front matter Ó 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.cropro.2011.05.005
example, Zhao et al. (2003) found that Bt pyramided broccoli delayed the evolution of resistance in the diamondback moth, Plutella xylostella (L.), compared to broccoli expressing a single toxin. Pyramided cotton was introduced in the early 2000s, not only for resistance management, but also to improve the inconsistent control of pests like the bollworm, Helicoverpa zea (Boddie) (Jackson et al., 2007). Because changes in Bt binding sites in the insect gut seem to be the most common mechanism of resistance, pyramided cultivars were designed to express toxins that involved different receptors (Ferré and Van Rie, 2002). Although the likelihood of resistance development is reduced by pyramiding, the possibility of cross resistance between Bt toxins by a common mechanism is still possible. For example, selection in the lab for resistance to Bt produced tobacco budworm larvae that were 10,000-fold resistant to Cry1Ac (Gould et al., 1997); cross resistance from Cry1Ac to Cry1Ab and Cry1Fa was also observed (Ferré and Van Rie, 2002). Bioassays based on a diagnostic dose of the Bt toxin have been a useful tool for resistance diagnosis in lepidopteran larvae in cotton where the assay end point is mortality (Blanco et al., 2008). In addition, Bailey et al. (1998, 2001) and Roe et al. (2000, 2003, 2005) showed that a feeding disruption test (FDT) with Cry1Ac toxin incorporated into a hydrateable, artificial diet containing blue dye could also be used to monitor tobacco budworm resistance. In
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this assay, the end point was the absence or presence of blue feces, the latter being a diagnosis of resistance. Van Kretschmar et al. (2011) described a novel bioassay architecture for FDT to facilitate the observation of blue feces on a white background. With this new technology, diagnostic doses for tobacco budworm and bollworm were estimated for Cry1Ac, Cry1Ab and Cry1F toxins (Van Kretschmar et al., 2011). Most bioassays have been based on exposure of the target insect to diet containing a Bt toxin obtained from gene expression in transformed bacteria (Bailey et al., 1998, 2001; Blanco et al., 2008). Disadvantages of this approach include the limited availability of different Bt and other toxins currently found in commercial crops, variations among Bt batches in toxin production and insecticidal activity, and the challenges of high cost of production. A more direct approach would be to use the toxins directly as expressed in plants of commercial Bt crops. Thus, the goal of this study was to evaluate a feeding disruption bioassay for monitoring tobacco budworm resistance, using cotton leaf extracts incorporated into hydrateable, artificial insect meals, i.e., an artificial cotton leaf.
2. Materials and methods All leaves used to prepare leaf solutions were from 8-wk-old cotton plants. Cotton varieties used were non-Bt cotton PHY 425 RF (Dow AgroSciences, Indianapolis, Indiana, USA), Bollgard II cotton DP 161 B2RF expressing Cry1Ac and Cry2Ab toxins (Monsanto, St. Louis, Missouri, USA), and WideStrike cotton PHY 485 WRF expressing Cry1Ac and Cry1F (Dow AgroSciences, Indianapolis, Indiana, USA). The plants were grown under greenhouse conditions at 22e31 C under natural light. After harvest, the leaves were stored at 80 C until used. A leaf extract stock solution was prepared for each cotton variety using 4 ml of distilled water/gram leaf tissue or 0.25 mg leaf/ml. Concentrations that follow are expressed in units of the latter. The leaf tissue was homogenized for 5 min using a Polytron homogenizer (PCU, Kinematica, Switzerland) and then filtered through glass wool. The filtrates were stored at 80 C until further use. Assays were conducted in 16-well FDT plates (Van Kretschmar et al., 2011), each well of the plate designed to contain 200 ml of a hydrateable heliothine artificial diet (Burton, 1970) (called “diet pad” throughout the manuscript). The plates with diet pads were lyophilized, placed in vacuum-sealed bags and stored at 4 C until used. The diet pads were rehydrated for the studies that follow with 170 ml of leaf filtrate and kept at room temperature in open air for 2 h before use. The bioassay was then initiated by the addition of a single 0e24 h old (from egg hatch) neonate tobacco budworm to each well. The wells of the plates were then sealed with an adhesive cover and kept for 24 h at 27 C with a 14:10 L:D photoperiod. Numbers of blue fecal pellets produced by each insect were then determined under a dissecting scope. The diagnostic dose was determined as the concentration of leaf material that resulted in 0e2 fecal pellets per larva after 24 h (the end point used before for FDTs by Bailey et al., 2001). The bioassay described above was utilized to determine a diagnostic dose for Bollgard II and WideStrike leaf solutions which would minimize the production of blue feces by susceptible larvae to 0e2 fecal pellets while fecal production for resistant insects would be greater than that of the susceptible insects. The tobacco budworm larvae used in these experiments were from a Btsusceptible, laboratory reared strain (Hv02) from North Carolina State University (Raleigh, North Carolina, USA). The bioassays involved 0.7, 1.3, 2.6 and 5.3 mg of Bollgard II cotton leaf per diet pad and 5.3, 10.6, 21.2 and 42.5 mg of WideStrike cotton per diet pad using the solutions described earlier. The control in these
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experiments was non-Bt cotton at the rate of 42.5 mg per diet pad. For each dose of Bollgard II and WideStrike evaluated, 64 larvae were tested. The diagnostic doses for the larval FDT determined from laboratory reared insects were validated using field collected insects from three different sites in North Carolina, USA. Tobacco budworm eggs were collected from tobacco fields in Cameron in Moore County (25/08/2008), Clayton in Johnston County (30/06/2009), and Reidsville in Rockingham County (20/07/2009). Tobacco buds with one or more eggs were transported to the laboratory on the day of collection and most of the plant tissue was separated from the egg to minimize larval feeding after egg hatch. The eggs were placed in an 8-oz Styrofoam container (WinCup, Stone Mountain, Georgia, USA) fitted with a plastic lid to prevent larvae from escaping and kept at 27 C with a 14:10 L:D photoperiod at 65% RH until egg hatch. Newly hatched larvae (0e24 h old) were used for the evaluation of the diagnostic dose. The sample size for Bollgard II and the corresponding non-Bt control (1.3 mg/diet pad) was 60 and 63 larvae from Moore County, 27 and 30 from Johnston County, and 52 and 44 from Rockingham County, respectively. The sample size for WideStrike and the non-Bt control (42.5 mg/diet pad) was 60 and 61 from Moore County, 32 and 25 from Johnston County, and 51 and 45 from Rockingham County, respectively. The diagnostic doses for Bollgard II and WideStrike were also evaluated with two Bt-resistant, laboratory reared tobacco budworm strains provided by Dr. Fred Gould from North Carolina State University (Raleigh, North Carolina, USA). Strain YHD2 had been selected for resistance to Cry1Ac but is cross-resistant to Cry1A, Cry1Fa and Cry2A (Jackson et al., 2007). The strain CxC had been selected for resistance to Cry1A and Cry2Aa2 (Jackson et al., 2007). Bioassays were conducted using 64 larvae per treatment. A non-Bt cotton control was conducted alongside each cotton variety. The shelf life of ready-to-use FDTs was evaluated over a period of six months. Diet pads were rehydrated with either non-Bt or Bollgard II cotton leaf extracts (1.3 mg/diet pad). Afterward, the diet was lyophilized, plates were then transferred into vacuum-sealed plastic storage bags (RivalÒ Seal a Meal, The Holmes Group, Milford, Massachusetts, USA), and stored at room temperature in the laboratory (no light). At different times thereafter, plates were removed from the plastic bags, 150 ml of distilled water was added to each diet pad and the diet was allowed to fully rehydrate for 1 h at room temperature. Then, bioassays were conducted as previously described using susceptible tobacco budworm Hv02 neonates, each treatment replicated 64 times. Comparisons between each Bt cotton variety (Bollgard II and WideStrike) and each corresponding non-Bt control were conducted with KruskaleWallis nonparametric tests. Wilcoxon ranksum nonparametric tests were used for validation of the diagnostic dose with field collected tobacco budworm. For validation of the diagnostic dose with the resistant tobacco budworm strains, YHD2 and CxC, two-tail t-tests were used. For the evaluation of shelf life, Wilcoxon rank-sum nonparametric tests were conducted for each pair-wise comparison between the Bollgard II and non-Bt control for months 1e6. We set our Type I error rate in the above experiments to 0.05 and used the SAS program version 9.1.3 to conduct the analyses (SAS Institute, 2003). 3. Results and discussion 3.1. The diagnostic doses A discriminating leaf dose was investigated for Bollgard II and WideStrike cotton for proof of concept of using a feeding disruption bioassay to distinguish Bt-susceptible from Bt-resistant tobacco budworms. The end point was feeding disruption as measured by
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lack of blue feces produced. Results of the leaf dose response studies for Bollgard II are shown in Fig. 1 (top graph). There were significant differences in fecal pellet production between Bollgard II doses (H ¼ 221.07, p < 0.0001), but the lowest dose to produce 0e2 blue fecal pellets per larva after 24 h (the FDT end point used by Bailey et al. (2001)) for all of the insects tested in the current study was 1.3 mg of leaf material per diet pad (Fig. 1). In contrast, diet pads rehydrated with non-Bt cotton (42.5 mg/diet pad) produced 28.5 1.3 (1SE) fecal pellets (a range of 9e79 fecal pellets) after 24 h. Using the diagnostic dose for lab reared susceptible budworms, the resistance false positive rate was 0% while for the lower dose of 0.7, the false positive rate was 20.3% (Fig. 1). A typical assay diagnosis of susceptibility is shown in Fig. 2B. No fecal production is found at the diagnostic dose for Bollgard II (Fig. 2B) while the non-Bt control has abundant feces (Fig. 2A). The comparison with the non-Bt control is important in showing that the insects being tested are viable under the assay conditions and will feed in the absence of the Bt toxins.
Fig. 2. Individual wells with diet pads rehydrated with leaf extract from (A) non-Bt cotton (1.3 mg leaf material/diet pad), (B) Bollgard II (1.3 mg/diet pad), (C) non-Bt cotton (42.5 mg/diet pad) and (D) WideStrike (42.5 mg/diet pad). Fecal pellets (small dark spots on white background) were produced after 24 h for non-Bt cotton (A, C) but were minimal in Bollgard II (B) and WideStrike (D) by susceptible, laboratory reared tobacco budworm, Heliothis virescens, neonates.
Fig. 1. Mean number of fecal pellets (1 SE) produced per insect by susceptible, laboratory reared tobacco budworm, Heliothis virescens, neonates after 24 h of exposure to diet pads treated with leaf extract from non-Bt and different concentrations of Bt cotton. There were significant differences between Bollgard II (H ¼ 221.07, p < 0.0001) and WideStrike doses (H ¼ 198.16, p < 0.0001). The arrow points to the concentration of Bollgard II (top) or WideStrike (bottom) cotton selected as diagnostic dose. FP: false positives (for resistance); OR: observed range (of fecal pellets produced).
The determination of the diagnostic dose for WideStrike cotton is shown in Fig. 1 (bottom graph). There were significant differences in fecal pellet production between WideStrike doses (H ¼ 198.16, p < 0.0001), but the dose that produced 0e2 fecal pellets in all of the larvae tested was 42.5 mg of leaf material per diet pad. This diagnostic dose for lab reared susceptible budworms produced no resistance false positives while the lower doses of 21.2, 10.6 and 5.3 produced 4.6, 7.8 and 9.0% false positives respectively, (Fig. 1). Tobacco budworm neonates exposed to diet pads rehydrated with non-Bt cotton (42.5 mg/diet pad) produced 14.2 2.7 (1SE) fecal pellets. However, in this case the fecal pellet range between replicates was 0e46 fecal pellets; 1.6% of the larvae produced 0e2 fecal pellets (false negatives for susceptibility). Fig. 2 (C and D) shows a typical comparison between the non-Bt control and that for WideStrike at the diagnostic dose, respectively. The diagnostic dose for WideStrike cotton (42.5 mg/diet pad) was 32.7-fold greater than that for Bollgard II (1.3 mg/diet pad) (Fig. 1), suggesting that the latter may be more toxic. However, no research was conducted to determine whether this would be the case in a practical field application, and these results alone should not be used to infer differences in efficacy between these two commercial products. Differences in diagnostic doses can be affected by a number of parameters including the choice of plant age and tissue, efficiency of the Bt extraction for preparation of the filtrate used for diet hydration, assay end point (complete cessation of fecal production versus mortality), and the use of artificial diet. With respect to the latter two points, Gore et al. (2005) found that tobacco budworm and bollworm neonates avoided feeding on artificial diet containing Cry1Ac or Cry2Ab. Avoidance of feeding on Bt cotton was also demonstrated in the cabbage looper, Trichoplusia ni (Hübner) (Li et al., 2006). In addition, when using artificial diet treated with Bt, toxicity changes depend on whether the toxin is incorporated into the diet or applied on the diet surface (Blanco et al., 2008); therefore, direct comparison of our results with other studies using artificial diet is difficult. Also, baseline susceptibility estimates of tobacco budworm to Bt toxins based on diagnostic doses have been shown to
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vary between laboratory colonies, field populations, host plant and time of the year for collection (Ali and Luttrell, 2007). Although more research will be needed to fully understand our results for this bioassay for susceptible, lab reared tobacco budworms, it appears from the data presented (Figs. 1 and 2) that the assay effectively diagnosed susceptibility to Bt toxins expressed in Bollgard II and WideStrike cotton with an error rate of 0%. 3.2. Bioassay validation with field collected tobacco budworms The diagnostic doses for the bioassay were developed using insects reared in the laboratory on artificial diet. Since these insects were reared for generations in the laboratory on artificial diet, the question remains whether the assay will also diagnose tobacco budworm susceptibility to commercial Bt plants for field populations of the insect. To address this question, the diagnostic doses for Bollgard II and WideStrike cotton were evaluated against three field populations from the central region of North Carolina (USA). The results are shown in Fig. 3. Tobacco budworm larvae collected from the field and exposed to Bollgard II treated diet pads did not produce any fecal pellets after 24 h, while larvae that fed on the corresponding diet pads rehydrated with filtrate from non-Bt cotton produced an average of >2 fecal pellets/larva for all three
Fig. 3. Mean number of fecal pellets (1 SE) produced by neonate tobacco budworm, Heliothis virescens, collected as eggs from the field and exposed for 24 h to diet pads (1 larva/well) treated with non-Bt or Bt cotton at the diagnostic dose. Eggs were collected from tobacco fields in Cameron, Moore Co. (A), Clayton, Johnston Co. (B) and Reidsville, Rockingham Co. (C), North Carolina (USA). Bars with different letters represent a statistically significant difference between the treatment and its respective non-Bt control (P 0.05).
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collection sites: Cameron, Moore Co. 0 vs 10.1 (z ¼ 10.05, p < 0.0001); Clayton, Johnston Co. 0 vs 24.7 (z ¼ 6.84, p < 0.0001); Reidsville, Rockingham Co. 0 vs 13.0 (z ¼ 9.01, p < 0.0001). However, a small percentage of non-feeding larvae (false negatives for susceptibility diagnosis) were observed in the non-Bt cotton treatment (3.3% of larvae collected from Cameron and 2.2% of the larvae collected from Reidsville). Similar results were observed with field collected tobacco budworm larvae tested with WideStrike-treated diet pads (Fig. 3). All the tested larvae produced 0e2 fecal pellets at 24 h on the WideStrike-treated diet pads while the larvae feeding on the corresponding non-Bt treatment produced an average of >2 fecal pellets/larva: Cameron, Moore Co. 0.1 vs 11.2 (z ¼ 9.95, p < 0.0001); Clayton, Johnston Co. 0.1 vs 17.2 (z ¼ 6.86, p < 0.0001); Reidsville, Rockingham Co. 0 vs 17.9 (z ¼ 8.88, p < 0.0001). In this case, the non-feeding larvae represented 9.5, 4.0 and 6.8% of tested larvae for Cameron, Clayton and Reidsville, respectively. All the larvae exposed to diet containing Bt cotton in these studies produced 0e2 fecal pellets while most (>90.5%) of the larvae feeding on non-Bt cotton treated diet produced >2 fecal pellets. This is consistent with a diagnosis of susceptibility and would be consistent with what would be expected for the insects collected; to date, there are no published reports of field resistance to Bt crops by tobacco budworm in the US (Bates et al., 2005) including the region where our samples were taken (J.S. Bacheler, personal communication). However, field collected tobacco budworm larvae in the non-Bt assays produced fewer fecal pellets with a larger proportion (9.5%) of the insects tested producing 0e2 fecal pellets compared to the laboratory reared susceptible strain (1.6%). Similar observations of non-feeding larvae exposed to non-Bt treatments were made by Bailey et al. (2001) when evaluating a Cry1Ac toxin diagnostic dose in artificial diet and using field collected tobacco budworm populations from Louisiana, North Carolina, Georgia and Mississippi (USA). These results may suggest an adaptation to feeding on an artificial diet in laboratory strains and/or reflect a higher rate of neonate mortality for field collected insects. Tobacco budworm populations also vary in the level of susceptibility to Bt toxins. For example, Ali et al. (2006) estimated LC50s for Cry1Ac for several tobacco budworm strains, including laboratory colonies and field populations. The combined estimation of the LC50 for laboratory colonies was almost 5-fold lower than the combined LC50 estimation for field populations (Ali et al., 2006). Variation in Bt toxin susceptibility has also been observed between tobacco budworm populations collected in different crops. A study by Ali and Luttrell (2007) showed that tobacco budworm larvae collected from tobacco fields had a higher LC50 than larvae collected from other crops, including cotton and corn. Tobacco budworm populations sampled in our study were collected from tobacco fields. Despite these issues, our lab derived diagnostic doses for resistance for Bollgard II and WideStrike cotton was reasonable (90.5% accurate) in the diagnosis of susceptibility of tobacco budworm from tobacco fields in central North Carolina. The small percentage of insects that did not feed on the non-Bt FDTs illustrates the importance of this control in the interpretation of the effects of the diagnostic dose. Additional studies will also be needed to better understand the variation possible in control results for field populations, the determination of the reason for some insects not producing blue feces, and what modifications might be possible to reduce the percentage of non-feeders (as determined by fecal production) to zero. Also, the issue of what would be the best non-Bt control for these assays should be considered. Although we used a cotton variety that was not engineered to produce Bt toxin as the control, there certainly must be other biochemical differences in addition to Bt between these plants and those of Bollgard II and WideStrike. However, it is unlikely that the non-feeders in our
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control are a result of the cotton solution only, since non-feeding of larvae was observed before in FDT tests with artificial diet without insecticide (Bailey et al., 2001). 3.3. Responses of Bt-resistant tobacco budworm strains Two Bt-resistant, laboratory reared tobacco budworm strains, YHD2 and CxC, were used to examine the ability of our assay to diagnose resistance to Bt. For both resistant strains, newly-hatched (0e24 h) larvae were fed diet pads rehydrated with the diagnostic doses of Bollgard II and WideStrike leaf filtrate solutions, and the fecal production of each was compared to that of larvae exposed to non-Bt meal pads (Figs. 4 and 5). The results for the YHD2 strain appear in Fig. 4. For both non-Bt controls, average fecal production was >2 fecal pellets/larva with an observed range of 10e60 fecal pellets/larva for Bollgard II and 5e77 fecal pellets/larva for WideStrike. The Bollgard II and WideStrike diagnostic doses were successful in diagnosing the resistance of both strains (average fecal production 2 fecal pellets/larva). For the diagnostic dose of Bollgard II, average fecal production was 12.4 fecal pellets/larva and for WideStrike was 21.5 fecal pellets/larva. For WideStrike, there was no difference in fecal production between the non-Bt control and the diagnostic dose (t ¼ 0.76, P ¼ 0.448), and the percentage of false negatives for resistance diagnosis (% of resistant larvae that produced 2 fecal pellets/larva) was zero. However, this was not the case for Bollgard II where mean fecal production for the diagnostic dose was significantly lower than that of the control (t ¼ 7.69, P < 0.0001). In this case, 21.9% of the larvae exposed to Bollgard II cotton produced 0e2 fecal pellets at the diagnostic dose and would have been incorrectly classified as susceptible (false negatives for resistance). The results for the CxC strain are shown in Fig. 5. Again, the resistant strain larvae were correctly diagnosed for resistance to both Bollgard II and WideStrike with mean fecal production greater than 0e2 fecal pellets produced per neonate. The mean fecal production for the Bollgard II diagnostic dose was significantly lower than for the corresponding non-Bt control (t ¼ 6.49, p < 0.0001); this was also the case for the mean fecal production for
Fig. 4. Mean number of fecal pellets (1 SE) produced per insect by YHD2 resistant, laboratory reared tobacco budworm, Heliothis virescens, neonates 24 h after exposure to diet pads treated with non-Bt and Bt cotton (Bollgard II and WideStrike) at the diagnostic dose. Bars with different letters represent a statistically significant difference between the treatment and its respective non-Bt control (P 0.05). FN: false negatives (for resistance); OR: observed range (of fecal pellets produced).
Fig. 5. Mean number of fecal pellets (1 SE) produced per insect by CxC resistant, laboratory reared tobacco budworm, Heliothis virescens, neonates 24 h after exposure to diet pads treated with non-Bt and Bt cotton (Bollgard II and WideStrike) at the diagnostic dose. Bars with different letters represent a statistically significant difference between the treatment and its respective non-Bt control (P 0.05). FN: false negatives (for resistance); OR: observed range (of fecal pellets produced). For the WideStrike Non-Bt control, 4.7% of the insects produced 2 fecal pellets in 24 h.
the WideStrike diagnostic dose compared to the corresponding non-Bt control (t ¼ 7.05, p < 0.0001). The percentage of CxC larvae exposed to the non-Bt controls that produced an average of 0e2 fecal pellets per neonate was low, 0% for Bollgard II and 4.7% for WideStrike. The percentage of false negatives for resistance to Bollgard II was 9.7% of tested CxC larvae and 48.4% of larvae exposed to the WideStrike diagnostic dose. Although diagnosis of resistance was correct for both the YHD2 and CxC strains based on average fecal production per larva, the correct diagnosis of individuals ranged for WideStrike from 100 to only 52% and for Bollgard II, from 78 to 90% for YHD2 versus CxC, respectively. These results demonstrated proof of concept for the feeding disruption assay using cotton leaf solutions but show that further research on the diagnostic dose will be needed, especially comparing field collected susceptible to field collected Bt-resistant insects if the diagnosis of resistant individual insects is to be achieved with low false negatives. Although the use of Bt-resistant laboratory strains with variable cross resistance to different Bt toxins (Ferré and Van Rie, 2002; Gahan et al., 2005; Jackson et al., 2007) has been helpful in our studies, at the same time the experiments are difficult to interpret since in the process of laboratory selection of these strains, insects like the YHD2 budworms no longer can successfully feed on cotton even in the absence of Bt toxin. Shaver et al. (1978) observed that gossypol-a polyphenol produced by cotton plants and other compounds found in cotton flower bud extracts can be detrimental to tobacco budworm larvae, particularly neonates; secondary plant compounds of any type may also have had an effect on the feeding response of the larvae in our assay and to varying degrees may contribute to feeding differences between field and laboratory reared insects. The use of field collected, Bt-resistant strains in the future will provide a more reasonable approach to better assess our discriminating doses and their use in the diagnosis of individual resistant insects. 3.4. Ready-to-use bioassay storage The shelf life at room temperature of the hydrateable diet pad has not been previously evaluated. Table 1 shows the performance
A.R. Cabrera et al. / Crop Protection 30 (2011) 1196e1201 Table 1 Shelf life at room temperature of the diet pad containing cotton leaf extract. Values reported are the mean number of fecal pellets (1 SE) produced per insect after 24 h by susceptible, laboratory reared Heliothis virescens neonates.a Month
Southern Regional IPM Program (USDA 2008-34103-19004). ARC was also supported by a graduate assistantship from the Department of Entomology at North Carolina State University.
Cotton variety Non-Bt
1 2 3 4 5 6
1201
% false positivesb
Bollgard II
mean
SE
mean
SE
14.2 23.5 20.1 38.7 35.5 35.1
1.5a 2.05a 1.62a 2.53a 1.60a 1.96a
0.1 0.2 0.2 0.3 0.1 0.5
0.03b 0.06b 0.07b 0.08b 0.10b 0.16b
0 0 0 0 0 4.6
a Different letter per row indicate a statistically significant difference between means (P < 0.05). b False positives: susceptible larvae producing >2 fecal pellets after 24 h at the diagnostic dose.
of the bioassay after 1e6 months of storage at room temperature for the diagnosis of susceptibility to Bollgard II at the diagnostic dose compared to that for non-Bt cotton for lab reared, susceptible tobacco budworm neonates. Similar studies with WideStrike were not conducted. Our results suggest that hydrateable diet pads containing cotton leaf extract stored at room temperature are diagnostic for 6 months. All susceptible, laboratory reared Heliothis virescens neonates, as expected, produced <2 fecal pellets in 24 h when exposed to diet pads containing the diagnostic dose of Bollgard II (1.3 mg cotton/diet pad) and stored for 1e5 months at room temperature. The mean fecal production per larva was also lower than that of the non-Bt cotton control (month 1: z ¼ 9.87, p < 0.0001; month 2: z ¼ 10.17, p < 0.0001; month 3: z ¼ 10.17, p < 0.0001; month 4: z ¼ 8.16, p < 0.0001; month 5: z ¼ 10.24, p < 0.0001). At 6 months, 4.6% of the larvae exposed to Bollgard II produced >2 fecal pellets. However, the Bollgard II mean fecal production was significantly lower than that for non-Bt (z ¼ 10.03, p < 0.0001). In summary, proof of concept was demonstrated for a feeding disruption bioassay for monitoring tobacco budworm resistance to pyramided Bt cotton using leaf filtrates. The bioassay was developed for Bollgard II and WideStrike, but should be useful for any commercially available cotton expressing one or more Bt toxins and/or other protein toxins. An important advantage of the bioassay is the use of the protein toxin obtained from plant extracts. The extracts can be prepared in advance in large quantities, stored frozen, and used over extended periods to produce a standardized, commercially relevant bioassay for larval resistance. The technique provides a practical, ready-to-use, off the shelf (just add water), easy to read 24 h bioassay. Acknowledgments The authors would like to thank Mariah Bock, Leonardo Magalhaes, Daniela Ramirez and Jiwei Zhu, for their assistance in field collections of tobacco budworms, Dan Mott for his help providing the cotton seeds and Dr. Consuelo Arellano for her advice on statistical analysis. This project was supported by grants to RMR from Cotton Inc., NC Biotechnology Center/Kenan Institute (2003CFG-8009), the NC Agricultural Research Service, the USDA Risk Assessment Program (USDA 2007-39211-18425) and the USDA
References Ali, M.I., Luttrell, R.G., 2007. Susceptibility of bollworm and tobacco budworm (Lepidoptera: Noctuidae) to Cry2Ab2 insecticidal protein. J. Econ. Entomol. 100, 921e931. Ali, M.I., Luttrell, R.G., Young, S.Y., 2006. Susceptibilities of Helicoverpa zea and Heliothis virescens (Lepidoptera: Noctuidae) populations to Cry1Ac insecticidal protein. J. Econ. Entomol. 99, 164e175. Bailey, W.D., Brownie, C., Bacheler, J.S., Gould, F., Kennedy, G.G., Sorenson, C.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, 76e85. Bailey, W.D., Zhao, G., Carter, L.M., Gould, F., Kennedy, G.G., Roe, R.M., 1998. Feeding disruption bioassays for species and Bacillus thuringiensis resistance diagnosis for Heliothis virescens and Helicoverpa zea in cotton (Lepidoptera: Noctuidae). Crop Prot. 17, 591e598. Bates, S.L., Zhao, J.Z., Rough, R.T., Shelton, A.M., 2005. Insect resistance management in GM crops: past, present and future. Nat. Biotech. 23, 57e62. Blanco, C.A., Storer, N.P., Abel, C.A., Jackson, R., Leonard, R., Lopez, J.D., Payne, G., Seigfried, B.D., Spencer, T., Terán-Vargas, A.P., 2008. Baseline susceptibility of tobacco budworm (Lepidoptera: Noctuidae) to Cry1F toxin from Bacillus thuringiensis. J. Econ. Entomol. 101, 168e173. Burton, R.L., 1970. A low cost artificial diet for the corn earworm. J. Econ. Entomol. 63, 1969e1970. Ferré, J., Van Rie, J., 2002. Biochemistry and genetics of insect resistance to Bacillus thuringiensis. Annu. Rev. Entomol. 47, 501e533. Gahan, L.J., Ma, Y.T., Coble, M.L.M., Gould, F., Moar, W.J., Heckel, D., 2005. Genetic basis of resistance to Cry1Ac and Cry2Aa in Heliothis virescens (Lepidoptera: Noctuidae). J. Econ. Entomol. 98, 1357e1368. Gore, J., Adamczyk, J.J., Blanco, C.A., 2005. Selective feeding of tobacco budworm and bollworm (Lepidoptera: Noctuidae) on meridic diet with different concentrations of Bacillus thuringiensis proteins. J. Econ. Entomol. 98, 88e94. Gould, F., 1998. Sustainability of transgenic insecticidal cultivars: Integrating pest genetics and ecology. Annu. Rev. Entomol. 43, 701e726. Gould, F., Anderson, A., Jones, A., Sumerford, D., Heckel, D.G., Lopez, J., Micinski, S., Leonard, R., Laster, M., 1997. Initial frequency of alleles for resistance to Bacillus thuringiensis toxins in field populations of Heliothis virescens. PNAS (USA) 94, 3519e3523. Jackson, R.E., Marcus, M.A., Gould, F., Bradley, J.R., Van Duyn, J.W., 2007. Crossresistance responses of Cry1Ac selected Heliothis virescens (Lepidoptera: Noctuidae) to Bacillus thuringiensis protein Vip3A. J. Econ. Entomol. 100, 180e186. Li, Y.X., Greenberg, S.M., Liu, T.X., 2006. Effects of Bt cotton expressing Cry1Ac and Cry2Ab and non-Bt cotton on behavior, survival and development of Trichoplusia ni (Lepidoptera: Noctuidae). Crop Prot. 25, 940e948. Roe, R.M., Bailey, W.D., Gould, F., Kennedy, G.G., 2000. Insecticide Resistance Assay US Patent Number 6,060,039. Roe, R.M., Bailey, W.D., Gould, F., Kennedy, G.G., Sutula, C.L., 2003. Insecticide Resistance Assay U.S. Patent number: 6,517,856. Roe, R.M., Van Kretschmar, J., Thompson, D.M., Donohue, K.V., Sorenson, C.E., Thompson, G.D., Storer, N.P., Blanco, C., Lopez Jr., J.D., Leonard, B.R., Van Duyn, J., Kilpatrick, J., Hagerty, A., Brickle, D., 2005. Larval feeding disruption tests (FDT) for monitoring insect resistance to Cry 1Ac, Cry 1F and Cry 1Ab. In: Proceedings Cotton Beltwide Conferences. National Cotton Council, Memphis, TN, pp. 1651e1661. SAS Institute, 2003. SAS 9.1.3 Help and Documentation. SAS Institute, Cary, NC. Shaver, T.N., Dilday, R.H., Garcia, J.A., 1978. Interference of gossypol in bioassay for resistance to tobacco budworm in cotton. Crop Prot. 18, 55e57. Shelton, A.M., Zhao, J.Z., Rough, R.T., 2002. Economic, ecological, food safety, and social consequences of the deployment of Bt transgenic plants. Annu. Rev. Entomol. 47, 845e881. Sisterson, M.S., Biggs, R.W., Olson, C., Carriere, Y., Dennehy, T.J., Tabashink, B.E., 2004. Arthropod abundance and diversity in Bt and non-Bt cotton fields. Environ. Entomol. 33, 921e929. Van Kretschmar, J.B., Bailey, W.D., Arellano, C., Thompson, G.D., Sutula, C.L., Roe, R.M., 2011. Feeding disruption tests for monitoring the frequency of larval lepidopteran resistance to Cry1Ac, Cry1F, and Cry1Ab. Crop Prot. 30, 863e870. Zhao, J.Z., Cao, J., Li, Y.X., Collins, H.L., Roush, R.T., Earle, E.D., Shelton, A.M., 2003. Transgenic plants expressing two Bacillus thuringiensis toxins delay insect resistance evolution. Nat. Biotechnol. 21, 1493e1497.
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Resistance monitoring of Heliothis virescens to pyramided cotton varieties with a hydrateable, artificial cotton leaf bioassay Ana R. Cabrera, Jaap Van Kretschmar, Jack S. Bacheler, Hannah Burrack, Clyde E. Sorenson, R. Michael Roe* North Carolina State University, Department of Entomology, Campus Box 7647, Raleigh, NC 27695-7647, USA
a r t i c l e i n f o
a b s t r a c t
Article history: Received 1 December 2010 Received in revised form 6 May 2011 Accepted 7 May 2011
Proof of concept was demonstrated for a practical, off-the-shelf bioassay to monitor for tobacco budworm resistance to pyramided Bt cotton using plant filtrates. The bioassay was based on a previously described feeding disruption test using hydrateable artificial diet containing a blue indicator dye, a diagnostic dose of insecticide and novel assay architecture. Using neonate larvae from a Bt-susceptible, laboratory reared tobacco budworm strain, a diagnostic dose for Bollgard II and WideStrike cotton was obtained that limited neonate blue fecal production to 0e2 pellets in 24 h (Bt-resistant larvae produced >2 fecal pellets). The bioassay was tested with three different field populations of tobacco budworm collected from tobacco in central North Carolina (USA) and shown to accurately diagnose susceptibility to Bt. The diagnostic doses were also successfully evaluated with two Bt-resistant, laboratory reared tobacco budworm strains. Shelf life studies showed the assay could be stored for at least 6 months at room temperature (longer storage times were not studied). The application of the bioassay as an easy to use monitoring tool is discussed. Ó 2011 Elsevier Ltd. All rights reserved.
Keywords: Bt Tobacco budworm Heliothis virescens Cotton Bollgard II WideStrike Resistance
1. Introduction The cultivation of transgenic crops expressing Bacillus thuringiensis Berliner (Bt) toxins to control insect pests provides both economical (Shelton et al., 2002) and environmental benefits from reductions in chemical insecticide use (Ferré and Van Rie, 2002), effective control of pests and minimal impact on non-target organisms (Sisterson et al., 2004). Since their introduction in the 1990s, the proportion of the U.S. acreage planted with Bt cotton cultivars has grown steadily. However, one of the major concerns is the development of insect resistance in the target pest due to their constant exposure to the toxin (Ferré and Van Rie, 2002; Gould, 1998). Possible resistance mechanisms to the Bt toxins include a reduction of proteolytic activity in the insect gut, Bt receptor modifications and midgut cell regeneration or replacement (Ferré and Van Rie, 2002). Before the commercialization of “pyramided” cultivars expressing multiple Bt toxins, Gould (1998) suggested that stacked genes could reduce the risk of resistance evolving based on field studies showing low frequency of resistance alleles. As an
* Corresponding author. Tel./fax: þ1 919 515 4325. E-mail address: [email protected] (R.M. Roe). 0261-2194/$ e see front matter Ó 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.cropro.2011.05.005
example, Zhao et al. (2003) found that Bt pyramided broccoli delayed the evolution of resistance in the diamondback moth, Plutella xylostella (L.), compared to broccoli expressing a single toxin. Pyramided cotton was introduced in the early 2000s, not only for resistance management, but also to improve the inconsistent control of pests like the bollworm, Helicoverpa zea (Boddie) (Jackson et al., 2007). Because changes in Bt binding sites in the insect gut seem to be the most common mechanism of resistance, pyramided cultivars were designed to express toxins that involved different receptors (Ferré and Van Rie, 2002). Although the likelihood of resistance development is reduced by pyramiding, the possibility of cross resistance between Bt toxins by a common mechanism is still possible. For example, selection in the lab for resistance to Bt produced tobacco budworm larvae that were 10,000-fold resistant to Cry1Ac (Gould et al., 1997); cross resistance from Cry1Ac to Cry1Ab and Cry1Fa was also observed (Ferré and Van Rie, 2002). Bioassays based on a diagnostic dose of the Bt toxin have been a useful tool for resistance diagnosis in lepidopteran larvae in cotton where the assay end point is mortality (Blanco et al., 2008). In addition, Bailey et al. (1998, 2001) and Roe et al. (2000, 2003, 2005) showed that a feeding disruption test (FDT) with Cry1Ac toxin incorporated into a hydrateable, artificial diet containing blue dye could also be used to monitor tobacco budworm resistance. In
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this assay, the end point was the absence or presence of blue feces, the latter being a diagnosis of resistance. Van Kretschmar et al. (2011) described a novel bioassay architecture for FDT to facilitate the observation of blue feces on a white background. With this new technology, diagnostic doses for tobacco budworm and bollworm were estimated for Cry1Ac, Cry1Ab and Cry1F toxins (Van Kretschmar et al., 2011). Most bioassays have been based on exposure of the target insect to diet containing a Bt toxin obtained from gene expression in transformed bacteria (Bailey et al., 1998, 2001; Blanco et al., 2008). Disadvantages of this approach include the limited availability of different Bt and other toxins currently found in commercial crops, variations among Bt batches in toxin production and insecticidal activity, and the challenges of high cost of production. A more direct approach would be to use the toxins directly as expressed in plants of commercial Bt crops. Thus, the goal of this study was to evaluate a feeding disruption bioassay for monitoring tobacco budworm resistance, using cotton leaf extracts incorporated into hydrateable, artificial insect meals, i.e., an artificial cotton leaf.
2. Materials and methods All leaves used to prepare leaf solutions were from 8-wk-old cotton plants. Cotton varieties used were non-Bt cotton PHY 425 RF (Dow AgroSciences, Indianapolis, Indiana, USA), Bollgard II cotton DP 161 B2RF expressing Cry1Ac and Cry2Ab toxins (Monsanto, St. Louis, Missouri, USA), and WideStrike cotton PHY 485 WRF expressing Cry1Ac and Cry1F (Dow AgroSciences, Indianapolis, Indiana, USA). The plants were grown under greenhouse conditions at 22e31 C under natural light. After harvest, the leaves were stored at 80 C until used. A leaf extract stock solution was prepared for each cotton variety using 4 ml of distilled water/gram leaf tissue or 0.25 mg leaf/ml. Concentrations that follow are expressed in units of the latter. The leaf tissue was homogenized for 5 min using a Polytron homogenizer (PCU, Kinematica, Switzerland) and then filtered through glass wool. The filtrates were stored at 80 C until further use. Assays were conducted in 16-well FDT plates (Van Kretschmar et al., 2011), each well of the plate designed to contain 200 ml of a hydrateable heliothine artificial diet (Burton, 1970) (called “diet pad” throughout the manuscript). The plates with diet pads were lyophilized, placed in vacuum-sealed bags and stored at 4 C until used. The diet pads were rehydrated for the studies that follow with 170 ml of leaf filtrate and kept at room temperature in open air for 2 h before use. The bioassay was then initiated by the addition of a single 0e24 h old (from egg hatch) neonate tobacco budworm to each well. The wells of the plates were then sealed with an adhesive cover and kept for 24 h at 27 C with a 14:10 L:D photoperiod. Numbers of blue fecal pellets produced by each insect were then determined under a dissecting scope. The diagnostic dose was determined as the concentration of leaf material that resulted in 0e2 fecal pellets per larva after 24 h (the end point used before for FDTs by Bailey et al., 2001). The bioassay described above was utilized to determine a diagnostic dose for Bollgard II and WideStrike leaf solutions which would minimize the production of blue feces by susceptible larvae to 0e2 fecal pellets while fecal production for resistant insects would be greater than that of the susceptible insects. The tobacco budworm larvae used in these experiments were from a Btsusceptible, laboratory reared strain (Hv02) from North Carolina State University (Raleigh, North Carolina, USA). The bioassays involved 0.7, 1.3, 2.6 and 5.3 mg of Bollgard II cotton leaf per diet pad and 5.3, 10.6, 21.2 and 42.5 mg of WideStrike cotton per diet pad using the solutions described earlier. The control in these
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experiments was non-Bt cotton at the rate of 42.5 mg per diet pad. For each dose of Bollgard II and WideStrike evaluated, 64 larvae were tested. The diagnostic doses for the larval FDT determined from laboratory reared insects were validated using field collected insects from three different sites in North Carolina, USA. Tobacco budworm eggs were collected from tobacco fields in Cameron in Moore County (25/08/2008), Clayton in Johnston County (30/06/2009), and Reidsville in Rockingham County (20/07/2009). Tobacco buds with one or more eggs were transported to the laboratory on the day of collection and most of the plant tissue was separated from the egg to minimize larval feeding after egg hatch. The eggs were placed in an 8-oz Styrofoam container (WinCup, Stone Mountain, Georgia, USA) fitted with a plastic lid to prevent larvae from escaping and kept at 27 C with a 14:10 L:D photoperiod at 65% RH until egg hatch. Newly hatched larvae (0e24 h old) were used for the evaluation of the diagnostic dose. The sample size for Bollgard II and the corresponding non-Bt control (1.3 mg/diet pad) was 60 and 63 larvae from Moore County, 27 and 30 from Johnston County, and 52 and 44 from Rockingham County, respectively. The sample size for WideStrike and the non-Bt control (42.5 mg/diet pad) was 60 and 61 from Moore County, 32 and 25 from Johnston County, and 51 and 45 from Rockingham County, respectively. The diagnostic doses for Bollgard II and WideStrike were also evaluated with two Bt-resistant, laboratory reared tobacco budworm strains provided by Dr. Fred Gould from North Carolina State University (Raleigh, North Carolina, USA). Strain YHD2 had been selected for resistance to Cry1Ac but is cross-resistant to Cry1A, Cry1Fa and Cry2A (Jackson et al., 2007). The strain CxC had been selected for resistance to Cry1A and Cry2Aa2 (Jackson et al., 2007). Bioassays were conducted using 64 larvae per treatment. A non-Bt cotton control was conducted alongside each cotton variety. The shelf life of ready-to-use FDTs was evaluated over a period of six months. Diet pads were rehydrated with either non-Bt or Bollgard II cotton leaf extracts (1.3 mg/diet pad). Afterward, the diet was lyophilized, plates were then transferred into vacuum-sealed plastic storage bags (RivalÒ Seal a Meal, The Holmes Group, Milford, Massachusetts, USA), and stored at room temperature in the laboratory (no light). At different times thereafter, plates were removed from the plastic bags, 150 ml of distilled water was added to each diet pad and the diet was allowed to fully rehydrate for 1 h at room temperature. Then, bioassays were conducted as previously described using susceptible tobacco budworm Hv02 neonates, each treatment replicated 64 times. Comparisons between each Bt cotton variety (Bollgard II and WideStrike) and each corresponding non-Bt control were conducted with KruskaleWallis nonparametric tests. Wilcoxon ranksum nonparametric tests were used for validation of the diagnostic dose with field collected tobacco budworm. For validation of the diagnostic dose with the resistant tobacco budworm strains, YHD2 and CxC, two-tail t-tests were used. For the evaluation of shelf life, Wilcoxon rank-sum nonparametric tests were conducted for each pair-wise comparison between the Bollgard II and non-Bt control for months 1e6. We set our Type I error rate in the above experiments to 0.05 and used the SAS program version 9.1.3 to conduct the analyses (SAS Institute, 2003). 3. Results and discussion 3.1. The diagnostic doses A discriminating leaf dose was investigated for Bollgard II and WideStrike cotton for proof of concept of using a feeding disruption bioassay to distinguish Bt-susceptible from Bt-resistant tobacco budworms. The end point was feeding disruption as measured by
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lack of blue feces produced. Results of the leaf dose response studies for Bollgard II are shown in Fig. 1 (top graph). There were significant differences in fecal pellet production between Bollgard II doses (H ¼ 221.07, p < 0.0001), but the lowest dose to produce 0e2 blue fecal pellets per larva after 24 h (the FDT end point used by Bailey et al. (2001)) for all of the insects tested in the current study was 1.3 mg of leaf material per diet pad (Fig. 1). In contrast, diet pads rehydrated with non-Bt cotton (42.5 mg/diet pad) produced 28.5 1.3 (1SE) fecal pellets (a range of 9e79 fecal pellets) after 24 h. Using the diagnostic dose for lab reared susceptible budworms, the resistance false positive rate was 0% while for the lower dose of 0.7, the false positive rate was 20.3% (Fig. 1). A typical assay diagnosis of susceptibility is shown in Fig. 2B. No fecal production is found at the diagnostic dose for Bollgard II (Fig. 2B) while the non-Bt control has abundant feces (Fig. 2A). The comparison with the non-Bt control is important in showing that the insects being tested are viable under the assay conditions and will feed in the absence of the Bt toxins.
Fig. 2. Individual wells with diet pads rehydrated with leaf extract from (A) non-Bt cotton (1.3 mg leaf material/diet pad), (B) Bollgard II (1.3 mg/diet pad), (C) non-Bt cotton (42.5 mg/diet pad) and (D) WideStrike (42.5 mg/diet pad). Fecal pellets (small dark spots on white background) were produced after 24 h for non-Bt cotton (A, C) but were minimal in Bollgard II (B) and WideStrike (D) by susceptible, laboratory reared tobacco budworm, Heliothis virescens, neonates.
Fig. 1. Mean number of fecal pellets (1 SE) produced per insect by susceptible, laboratory reared tobacco budworm, Heliothis virescens, neonates after 24 h of exposure to diet pads treated with leaf extract from non-Bt and different concentrations of Bt cotton. There were significant differences between Bollgard II (H ¼ 221.07, p < 0.0001) and WideStrike doses (H ¼ 198.16, p < 0.0001). The arrow points to the concentration of Bollgard II (top) or WideStrike (bottom) cotton selected as diagnostic dose. FP: false positives (for resistance); OR: observed range (of fecal pellets produced).
The determination of the diagnostic dose for WideStrike cotton is shown in Fig. 1 (bottom graph). There were significant differences in fecal pellet production between WideStrike doses (H ¼ 198.16, p < 0.0001), but the dose that produced 0e2 fecal pellets in all of the larvae tested was 42.5 mg of leaf material per diet pad. This diagnostic dose for lab reared susceptible budworms produced no resistance false positives while the lower doses of 21.2, 10.6 and 5.3 produced 4.6, 7.8 and 9.0% false positives respectively, (Fig. 1). Tobacco budworm neonates exposed to diet pads rehydrated with non-Bt cotton (42.5 mg/diet pad) produced 14.2 2.7 (1SE) fecal pellets. However, in this case the fecal pellet range between replicates was 0e46 fecal pellets; 1.6% of the larvae produced 0e2 fecal pellets (false negatives for susceptibility). Fig. 2 (C and D) shows a typical comparison between the non-Bt control and that for WideStrike at the diagnostic dose, respectively. The diagnostic dose for WideStrike cotton (42.5 mg/diet pad) was 32.7-fold greater than that for Bollgard II (1.3 mg/diet pad) (Fig. 1), suggesting that the latter may be more toxic. However, no research was conducted to determine whether this would be the case in a practical field application, and these results alone should not be used to infer differences in efficacy between these two commercial products. Differences in diagnostic doses can be affected by a number of parameters including the choice of plant age and tissue, efficiency of the Bt extraction for preparation of the filtrate used for diet hydration, assay end point (complete cessation of fecal production versus mortality), and the use of artificial diet. With respect to the latter two points, Gore et al. (2005) found that tobacco budworm and bollworm neonates avoided feeding on artificial diet containing Cry1Ac or Cry2Ab. Avoidance of feeding on Bt cotton was also demonstrated in the cabbage looper, Trichoplusia ni (Hübner) (Li et al., 2006). In addition, when using artificial diet treated with Bt, toxicity changes depend on whether the toxin is incorporated into the diet or applied on the diet surface (Blanco et al., 2008); therefore, direct comparison of our results with other studies using artificial diet is difficult. Also, baseline susceptibility estimates of tobacco budworm to Bt toxins based on diagnostic doses have been shown to
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vary between laboratory colonies, field populations, host plant and time of the year for collection (Ali and Luttrell, 2007). Although more research will be needed to fully understand our results for this bioassay for susceptible, lab reared tobacco budworms, it appears from the data presented (Figs. 1 and 2) that the assay effectively diagnosed susceptibility to Bt toxins expressed in Bollgard II and WideStrike cotton with an error rate of 0%. 3.2. Bioassay validation with field collected tobacco budworms The diagnostic doses for the bioassay were developed using insects reared in the laboratory on artificial diet. Since these insects were reared for generations in the laboratory on artificial diet, the question remains whether the assay will also diagnose tobacco budworm susceptibility to commercial Bt plants for field populations of the insect. To address this question, the diagnostic doses for Bollgard II and WideStrike cotton were evaluated against three field populations from the central region of North Carolina (USA). The results are shown in Fig. 3. Tobacco budworm larvae collected from the field and exposed to Bollgard II treated diet pads did not produce any fecal pellets after 24 h, while larvae that fed on the corresponding diet pads rehydrated with filtrate from non-Bt cotton produced an average of >2 fecal pellets/larva for all three
Fig. 3. Mean number of fecal pellets (1 SE) produced by neonate tobacco budworm, Heliothis virescens, collected as eggs from the field and exposed for 24 h to diet pads (1 larva/well) treated with non-Bt or Bt cotton at the diagnostic dose. Eggs were collected from tobacco fields in Cameron, Moore Co. (A), Clayton, Johnston Co. (B) and Reidsville, Rockingham Co. (C), North Carolina (USA). Bars with different letters represent a statistically significant difference between the treatment and its respective non-Bt control (P 0.05).
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collection sites: Cameron, Moore Co. 0 vs 10.1 (z ¼ 10.05, p < 0.0001); Clayton, Johnston Co. 0 vs 24.7 (z ¼ 6.84, p < 0.0001); Reidsville, Rockingham Co. 0 vs 13.0 (z ¼ 9.01, p < 0.0001). However, a small percentage of non-feeding larvae (false negatives for susceptibility diagnosis) were observed in the non-Bt cotton treatment (3.3% of larvae collected from Cameron and 2.2% of the larvae collected from Reidsville). Similar results were observed with field collected tobacco budworm larvae tested with WideStrike-treated diet pads (Fig. 3). All the tested larvae produced 0e2 fecal pellets at 24 h on the WideStrike-treated diet pads while the larvae feeding on the corresponding non-Bt treatment produced an average of >2 fecal pellets/larva: Cameron, Moore Co. 0.1 vs 11.2 (z ¼ 9.95, p < 0.0001); Clayton, Johnston Co. 0.1 vs 17.2 (z ¼ 6.86, p < 0.0001); Reidsville, Rockingham Co. 0 vs 17.9 (z ¼ 8.88, p < 0.0001). In this case, the non-feeding larvae represented 9.5, 4.0 and 6.8% of tested larvae for Cameron, Clayton and Reidsville, respectively. All the larvae exposed to diet containing Bt cotton in these studies produced 0e2 fecal pellets while most (>90.5%) of the larvae feeding on non-Bt cotton treated diet produced >2 fecal pellets. This is consistent with a diagnosis of susceptibility and would be consistent with what would be expected for the insects collected; to date, there are no published reports of field resistance to Bt crops by tobacco budworm in the US (Bates et al., 2005) including the region where our samples were taken (J.S. Bacheler, personal communication). However, field collected tobacco budworm larvae in the non-Bt assays produced fewer fecal pellets with a larger proportion (9.5%) of the insects tested producing 0e2 fecal pellets compared to the laboratory reared susceptible strain (1.6%). Similar observations of non-feeding larvae exposed to non-Bt treatments were made by Bailey et al. (2001) when evaluating a Cry1Ac toxin diagnostic dose in artificial diet and using field collected tobacco budworm populations from Louisiana, North Carolina, Georgia and Mississippi (USA). These results may suggest an adaptation to feeding on an artificial diet in laboratory strains and/or reflect a higher rate of neonate mortality for field collected insects. Tobacco budworm populations also vary in the level of susceptibility to Bt toxins. For example, Ali et al. (2006) estimated LC50s for Cry1Ac for several tobacco budworm strains, including laboratory colonies and field populations. The combined estimation of the LC50 for laboratory colonies was almost 5-fold lower than the combined LC50 estimation for field populations (Ali et al., 2006). Variation in Bt toxin susceptibility has also been observed between tobacco budworm populations collected in different crops. A study by Ali and Luttrell (2007) showed that tobacco budworm larvae collected from tobacco fields had a higher LC50 than larvae collected from other crops, including cotton and corn. Tobacco budworm populations sampled in our study were collected from tobacco fields. Despite these issues, our lab derived diagnostic doses for resistance for Bollgard II and WideStrike cotton was reasonable (90.5% accurate) in the diagnosis of susceptibility of tobacco budworm from tobacco fields in central North Carolina. The small percentage of insects that did not feed on the non-Bt FDTs illustrates the importance of this control in the interpretation of the effects of the diagnostic dose. Additional studies will also be needed to better understand the variation possible in control results for field populations, the determination of the reason for some insects not producing blue feces, and what modifications might be possible to reduce the percentage of non-feeders (as determined by fecal production) to zero. Also, the issue of what would be the best non-Bt control for these assays should be considered. Although we used a cotton variety that was not engineered to produce Bt toxin as the control, there certainly must be other biochemical differences in addition to Bt between these plants and those of Bollgard II and WideStrike. However, it is unlikely that the non-feeders in our
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control are a result of the cotton solution only, since non-feeding of larvae was observed before in FDT tests with artificial diet without insecticide (Bailey et al., 2001). 3.3. Responses of Bt-resistant tobacco budworm strains Two Bt-resistant, laboratory reared tobacco budworm strains, YHD2 and CxC, were used to examine the ability of our assay to diagnose resistance to Bt. For both resistant strains, newly-hatched (0e24 h) larvae were fed diet pads rehydrated with the diagnostic doses of Bollgard II and WideStrike leaf filtrate solutions, and the fecal production of each was compared to that of larvae exposed to non-Bt meal pads (Figs. 4 and 5). The results for the YHD2 strain appear in Fig. 4. For both non-Bt controls, average fecal production was >2 fecal pellets/larva with an observed range of 10e60 fecal pellets/larva for Bollgard II and 5e77 fecal pellets/larva for WideStrike. The Bollgard II and WideStrike diagnostic doses were successful in diagnosing the resistance of both strains (average fecal production 2 fecal pellets/larva). For the diagnostic dose of Bollgard II, average fecal production was 12.4 fecal pellets/larva and for WideStrike was 21.5 fecal pellets/larva. For WideStrike, there was no difference in fecal production between the non-Bt control and the diagnostic dose (t ¼ 0.76, P ¼ 0.448), and the percentage of false negatives for resistance diagnosis (% of resistant larvae that produced 2 fecal pellets/larva) was zero. However, this was not the case for Bollgard II where mean fecal production for the diagnostic dose was significantly lower than that of the control (t ¼ 7.69, P < 0.0001). In this case, 21.9% of the larvae exposed to Bollgard II cotton produced 0e2 fecal pellets at the diagnostic dose and would have been incorrectly classified as susceptible (false negatives for resistance). The results for the CxC strain are shown in Fig. 5. Again, the resistant strain larvae were correctly diagnosed for resistance to both Bollgard II and WideStrike with mean fecal production greater than 0e2 fecal pellets produced per neonate. The mean fecal production for the Bollgard II diagnostic dose was significantly lower than for the corresponding non-Bt control (t ¼ 6.49, p < 0.0001); this was also the case for the mean fecal production for
Fig. 4. Mean number of fecal pellets (1 SE) produced per insect by YHD2 resistant, laboratory reared tobacco budworm, Heliothis virescens, neonates 24 h after exposure to diet pads treated with non-Bt and Bt cotton (Bollgard II and WideStrike) at the diagnostic dose. Bars with different letters represent a statistically significant difference between the treatment and its respective non-Bt control (P 0.05). FN: false negatives (for resistance); OR: observed range (of fecal pellets produced).
Fig. 5. Mean number of fecal pellets (1 SE) produced per insect by CxC resistant, laboratory reared tobacco budworm, Heliothis virescens, neonates 24 h after exposure to diet pads treated with non-Bt and Bt cotton (Bollgard II and WideStrike) at the diagnostic dose. Bars with different letters represent a statistically significant difference between the treatment and its respective non-Bt control (P 0.05). FN: false negatives (for resistance); OR: observed range (of fecal pellets produced). For the WideStrike Non-Bt control, 4.7% of the insects produced 2 fecal pellets in 24 h.
the WideStrike diagnostic dose compared to the corresponding non-Bt control (t ¼ 7.05, p < 0.0001). The percentage of CxC larvae exposed to the non-Bt controls that produced an average of 0e2 fecal pellets per neonate was low, 0% for Bollgard II and 4.7% for WideStrike. The percentage of false negatives for resistance to Bollgard II was 9.7% of tested CxC larvae and 48.4% of larvae exposed to the WideStrike diagnostic dose. Although diagnosis of resistance was correct for both the YHD2 and CxC strains based on average fecal production per larva, the correct diagnosis of individuals ranged for WideStrike from 100 to only 52% and for Bollgard II, from 78 to 90% for YHD2 versus CxC, respectively. These results demonstrated proof of concept for the feeding disruption assay using cotton leaf solutions but show that further research on the diagnostic dose will be needed, especially comparing field collected susceptible to field collected Bt-resistant insects if the diagnosis of resistant individual insects is to be achieved with low false negatives. Although the use of Bt-resistant laboratory strains with variable cross resistance to different Bt toxins (Ferré and Van Rie, 2002; Gahan et al., 2005; Jackson et al., 2007) has been helpful in our studies, at the same time the experiments are difficult to interpret since in the process of laboratory selection of these strains, insects like the YHD2 budworms no longer can successfully feed on cotton even in the absence of Bt toxin. Shaver et al. (1978) observed that gossypol-a polyphenol produced by cotton plants and other compounds found in cotton flower bud extracts can be detrimental to tobacco budworm larvae, particularly neonates; secondary plant compounds of any type may also have had an effect on the feeding response of the larvae in our assay and to varying degrees may contribute to feeding differences between field and laboratory reared insects. The use of field collected, Bt-resistant strains in the future will provide a more reasonable approach to better assess our discriminating doses and their use in the diagnosis of individual resistant insects. 3.4. Ready-to-use bioassay storage The shelf life at room temperature of the hydrateable diet pad has not been previously evaluated. Table 1 shows the performance
A.R. Cabrera et al. / Crop Protection 30 (2011) 1196e1201 Table 1 Shelf life at room temperature of the diet pad containing cotton leaf extract. Values reported are the mean number of fecal pellets (1 SE) produced per insect after 24 h by susceptible, laboratory reared Heliothis virescens neonates.a Month
Southern Regional IPM Program (USDA 2008-34103-19004). ARC was also supported by a graduate assistantship from the Department of Entomology at North Carolina State University.
Cotton variety Non-Bt
1 2 3 4 5 6
1201
% false positivesb
Bollgard II
mean
SE
mean
SE
14.2 23.5 20.1 38.7 35.5 35.1
1.5a 2.05a 1.62a 2.53a 1.60a 1.96a
0.1 0.2 0.2 0.3 0.1 0.5
0.03b 0.06b 0.07b 0.08b 0.10b 0.16b
0 0 0 0 0 4.6
a Different letter per row indicate a statistically significant difference between means (P < 0.05). b False positives: susceptible larvae producing >2 fecal pellets after 24 h at the diagnostic dose.
of the bioassay after 1e6 months of storage at room temperature for the diagnosis of susceptibility to Bollgard II at the diagnostic dose compared to that for non-Bt cotton for lab reared, susceptible tobacco budworm neonates. Similar studies with WideStrike were not conducted. Our results suggest that hydrateable diet pads containing cotton leaf extract stored at room temperature are diagnostic for 6 months. All susceptible, laboratory reared Heliothis virescens neonates, as expected, produced <2 fecal pellets in 24 h when exposed to diet pads containing the diagnostic dose of Bollgard II (1.3 mg cotton/diet pad) and stored for 1e5 months at room temperature. The mean fecal production per larva was also lower than that of the non-Bt cotton control (month 1: z ¼ 9.87, p < 0.0001; month 2: z ¼ 10.17, p < 0.0001; month 3: z ¼ 10.17, p < 0.0001; month 4: z ¼ 8.16, p < 0.0001; month 5: z ¼ 10.24, p < 0.0001). At 6 months, 4.6% of the larvae exposed to Bollgard II produced >2 fecal pellets. However, the Bollgard II mean fecal production was significantly lower than that for non-Bt (z ¼ 10.03, p < 0.0001). In summary, proof of concept was demonstrated for a feeding disruption bioassay for monitoring tobacco budworm resistance to pyramided Bt cotton using leaf filtrates. The bioassay was developed for Bollgard II and WideStrike, but should be useful for any commercially available cotton expressing one or more Bt toxins and/or other protein toxins. An important advantage of the bioassay is the use of the protein toxin obtained from plant extracts. The extracts can be prepared in advance in large quantities, stored frozen, and used over extended periods to produce a standardized, commercially relevant bioassay for larval resistance. The technique provides a practical, ready-to-use, off the shelf (just add water), easy to read 24 h bioassay. Acknowledgments The authors would like to thank Mariah Bock, Leonardo Magalhaes, Daniela Ramirez and Jiwei Zhu, for their assistance in field collections of tobacco budworms, Dan Mott for his help providing the cotton seeds and Dr. Consuelo Arellano for her advice on statistical analysis. This project was supported by grants to RMR from Cotton Inc., NC Biotechnology Center/Kenan Institute (2003CFG-8009), the NC Agricultural Research Service, the USDA Risk Assessment Program (USDA 2007-39211-18425) and the USDA
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