Abstract
We tested toxins of Bacillus thuringiensis against larvae from susceptible, Cry1C-resistant, and Cry1A-resistant strains of diamondback moth (Plutella xylostella). The Cry1C-resistant strain, which was derived from a field population that had evolved resistance to B. thuringiensis subsp. kurstaki and B. thuringiensis subsp. aizawai, was selected repeatedly with Cry1C in the laboratory. The Cry1C-resistant strain had strong cross-resistance to Cry1Ab, Cry1Ac, and Cry1F, low to moderate cross-resistance to Cry1Aa and Cry9Ca, and no cross-resistance to Cry1Bb, Cry1Ja, and Cry2A. Resistance to Cry1C declined when selection was relaxed. Together with previously reported data, the new data on the cross-resistance of a Cry1C-resistant strain reported here suggest that resistance to Cry1A and Cry1C toxins confers little or no cross-resistance to Cry1Bb, Cry2Aa, or Cry9Ca. Therefore, these toxins might be useful in rotations or combinations with Cry1A and Cry1C toxins. Cry9Ca was much more potent than Cry1Bb or Cry2Aa and thus might be especially useful against diamondback moth.
Because of their safety to most nontarget organisms, spray formulations of insecticidal crystal proteins from Bacillus thuringiensis have been used widely to control insect pests (13). Transgenic crops producing B. thuringiensis toxins have also been grown on millions of hectares (4). Evolution of resistance by insects is the greatest threat to the continued success of B. thuringiensis.
Strains of more than 10 insect species have evolved resistance to B. thuringiensis toxins in laboratory selections (2, 14). Yet, so far, resistance has been reported in field populations of only the diamondback moth (2, 14), Plutella xylostella (L.) (Lepidoptera: Plutellidae), a worldwide pest of crucifers. Many populations of the diamondback moth have evolved resistance to spray formulations of B. thuringiensis toxins in the field (2, 14).
The first cases of resistance to B. thuringiensis in the field were to formulations of B. thuringiensis subsp. kurstaki containing Cry1A toxins, which had been used widely to control the diamondback moth (14). Strains of diamondback moth resistant to B. thuringiensis subsp. kurstaki and Cry1A toxins do not show cross-resistance to Cry1C (1, 18), a toxin present in spray formulations of B. thuringiensis subsp. aizawai but not of B. thuringiensis subsp. kurstaki (9). More recently, as the use of B. thuringiensis subsp. aizawai has increased, field-evolved resistance of the diamondback moth to this strain and to Cry1C has occurred (9, 11, 24, 25).
Knowledge of resistance to Cry1A and Cry1C toxins in the diamondback moth may be useful for managing resistance of the diamondback moth and other pests to B. thuringiensis. The most common type of resistance to Cry1A toxins in the diamondback moth, called mode 1 resistance, entails >500-fold resistance to at least one Cry1A toxin, recessive inheritance, little or no cross-resistance to Cry1C, and reduced binding of at least one Cry1A toxin to midgut membrane target sites (21). For example, the NO-QA strain of diamondback moth from Hawaii harbors an autosomal recessive gene that confers resistance to Cry1Aa, Cry1Ab, Cry1Ac, Cry1Fa, and Cry1Ja but not to Cry1B, Cry1C, and Cry1I (3, 19, 22). Although reduced binding of toxin to midgut membrane target sites is the only well-documented mechanism of resistance in diamondback moth, this mechanism does not account for all examples of diamondback moth resistance to Cry1A toxins (1, 20, 24). In laboratory studies of field-selected strains of the diamondback moth, resistance generally declined when exposure to B. thuringiensis subsp. kurstaki stopped (15).
Comparisons between diamondback moth resistance to Cry1A and Cry1C are beginning to reveal some key differences and similarities. Unlike resistance to Cry1A, reduced binding was not a major mechanism of resistance to Cry1C in field-selected strains from Malaysia, Hawaii, or Florida (10, 24, 25). In Hawaii and Florida strains, the dominance of resistance to Cry1C increased as concentration decreased and dominance was intermediate at the concentration killing 50% of the larvae tested (LC50) (7, 25). Like most cases of resistance to B. thuringiensis subsp. kurstaki, resistance to B. thuringiensis subsp. aizawai (24) and to Cry1C (25) declined when selection stopped.
Two methods for delaying resistance to B. thuringiensis are to rotate or combine B. thuringiensis toxins that are unlikely to produce cross-resistance to each other. However, in contrast to the many published studies about cross-resistance associated with Cry1A resistance, little has been reported about cross-resistance associated with diamondback moth resistance to Cry1C. Thus, the primary objective of the present study was to determine the cross-resistance pattern of the Cry1C-resistant NO-95C strain of diamondback moth from Hawaii. We also examined the stability of Cry1C resistance.
MATERIALS AND METHODS
Insects.
We used three strains of diamondback moth: resistant strain NO-QA, resistant strain NO-95C, and susceptible strain LAB-PS (8). NO-QA was derived in 1989 and NO-95C was derived in 1995 from the NO field population in Hawaii, which had evolved resistance first to B. thuringiensis subsp. kurstaki and Cry1A toxins (16, 17) and later to B. thuringiensis subsp. aizawai and Cry1C (9). NO-QA was selected repeatedly with Dipel, a spore-crystal formulation of B. thuringiensis subsp. kurstaki, and was resistant to B. thuringiensis subsp. kurstaki, Cry1A toxins, Cry1F, and Cry1J (20). NO-95C was selected repeatedly with MYX833-4C1 (referred to hereafter as Cry1C), a liquid formulation containing Cry1C protoxin produced and encapsulated by transgenic Pseudomonas fluorescens (Dow Agrosciences, San Diego, Calif.). Larvae were reared on cabbage plants.
B. thuringiensis toxins.
We obtained from Ecogen powders containing spores and crystals of the following B. thuringiensis proteins: Cry1Aa, Cry1Ab, Cry1Ac, Cry1Bb, Cry1Ca, Cry1Fa, Cry1Ja, and Cry2Aa. Plant Genetic Systems provided powder containing purified Cry9Ca toxin. The powders were diluted with distilled water containing 0.2% Triton AG-98 (a surfactant; Rohm & Haas, Philadelphia, Pa.) for bioassays.
Bioassays.
We used leaf-residue bioassays (9). One week after eggs were placed on cabbage plants, larvae were used for bioassays. Ten third-instar larvae were placed on each treated leaf disk. For each bioassay, four replicates (40 larvae) were tested at each concentration for each toxin and strain. After 2 days, fresh untreated leaves were added to each petri dish. Mortality was recorded at 5 days. Bioassays and rearing were conducted at 28°C and a photoperiod of 14:10 (L:D) h.
Two sets of bioassays were conducted for most toxins. In the first set, we screened all toxins except Cry9Ca at two or three concentrations against susceptible strain LAB-PS and resistant strain NO-95C. For Cry1Bb, Cry1Ja, and Cry2Aa, resistant strain NO-QA was also tested in the first set of bioassays. In the second set of bioassays, we estimated LC50s for all toxins except Cry2Aa and Cry1Bb by testing at five concentrations, including a control against the NO-95C and LAB-PS strains. LC50s for Cry2Aa and Cry1Bb were not estimated because of their low toxicity and our limited quantities of these toxins.
Selection and crosses with susceptible strain.
Our goals were to increase the resistance to Cry1C and to reduce the frequency of the multitoxin resistance gene (3, 19) that confers resistance to Cry1A toxins, Cry1F, and Cry1J. To increase resistance to Cry1C, NO-95C was selected with Cry1C (MYX833-4C1) 22 times in 75 generations of laboratory rearing (Table 1). In each selection, 300 to 600 third-instar larvae were fed leaf disks that had been treated with Cry1C (2.5 to 10 ml/liter) by using the method described above for the bioassays. Mortality of Cry1C-selected larvae ranged from 41 to 99% (Table 1). Adult survivors from treated leaf disks were pooled to produce progeny for the next generation.
TABLE 1.
Response of diamondback moth strain NO-95C to selection with Cry1C
| Generation | Cry1C, concn (ml/liter) | n | % mortalitya |
|---|---|---|---|
| 1 | 5 | 300 | 86.0 |
| 3 | 5 | 600 | 62.2 |
| 3b | 5 | 450 | 82.4 |
| 4 | 5 | 445 | 75.5 |
| 6 | 5 | 610 | 88.0 |
| 7 | 5 | 598 | 88.1 |
| 9 | 10 | 600 | 78.3 |
| 10 | 10 | 142 | 74.6 |
| 14 | 10 | 273 | 49.1 |
| 22 | 5 | 450 | 65.3 |
| 27 | 2.5 | 450 | 94.7 |
| 28 | 2.5 | 450 | 74.0 |
| 29 | 2.5 | 450 | 79.1 |
| 31 | 2.5 | 450 | 78.7 |
| 35 | 5, 10 | 314 | 83.7 |
| 36 | 5 | 350 | 89.1 |
| 41 | 5 | 600 | 93.5 |
| 44 | 5 | 600 | 70.7 |
| 47 | 5 | 450 | 40.7 |
| 51 | 5 | 600 | 64.8 |
| 56 | 5 | 600 | 61.7 |
| 62 | 5 | 598 | 52.2 |
| 75 | 5 | 600 | 99.5 |
For generations 1 to 4, mortality was recorded at 2 days. For all other selections, mortality was recorded at 5 days. Mortality was adjusted for control mortality by using Abbott's method.
A second group of larvae were collected from the NO field population and selected with Cry1C. Offspring of survivors were added to NO-95C moths in generation 4 to increase genetic variation (7).
To reduce the frequency of the multitoxin resistance gene, we crossed NO-95C with susceptible strain LAB-PS at generations 19 and 31 of NO-95C. Mature larvae from each strain were sexed (6). We pooled about 50 LAB-PS males with about 50 NO-95C females for one cross and pooled about 50 LAB-PS females with about 50 NO-95C males for the reciprocal cross. F1 offspring from the two crosses were mixed and allowed to mate to produce F2 offspring. Selection with Cry1C resumed with the F2 from the crosses.
Stability of Cry1C resistance.
To evaluate the stability of Cry1C resistance, we compared mortality at single concentrations (Table 1) and resistance ratios (see below) during the course of selection. LC50s of Cry1C for NO-95C and a paired susceptible strain were estimated at generations 1 and 2 (9), generations 8 and 10 (7), generations 40 and 45 pooled (10), and generation 60. Between the first and second measurements of LC50 from generation 2 to 8, NO-95C was selected four times at 5 ml/liter. Between the last two measurements of LC50 from generation 45 to 60, NO-95C was selected with Cry1C three times at 5 ml/liter.
Data analysis.
Mortality was adjusted for mortality in controls using Abbott's method. We used probit analysis (12) to estimate LC50s and their 95% fiducial limits (FLs) and slopes of concentration-mortality lines and their standard errors (SE). LC50s were considered significantly different if their 95% FLs did not overlap. Resistance ratios were calculated as the LC50 for the resistant strain divided by the LC50 for LAB-PS.
RESULTS
Cross-resistance.
Compared with the susceptible LAB-PS strain, the Cry1C-selected NO-95C strain had a 11-fold resistance to Cry1Ca in powder and a 17-fold resistance to Cry1C in a liquid formulation (Table 2). NO-95C showed strong cross-resistance to Cry1Ab, Cry1Ac, and Cry1Fa, low to moderate cross-resistance to Cry1Aa and Cry9Ca, and no cross-resistance to Cry1Bb, Cry1Ja, and Cry2Aa (Tables 2 and 3). In contrast to NO-95C, the NO-QA strain selected with B. thuringiensis subsp. kurstaki showed strong cross-resistance to Cry1Ja (Table 3). NO-QA was not cross-resistant to Cry1Ba or Cry2Aa, which were least potent to susceptible larvae of any of the toxins tested (Table 3).
TABLE 2.
Responses of diamondback moth larvae to B. thuringiensis toxins
| Toxin | Strain | n | Slope ± SE | LC50 (95%FL) (mg/liter)a | RRb |
|---|---|---|---|---|---|
| Cry1Aa | LAB-PS | 200 | 2.4 ± 0.4 | 2.80 (1.94–4.26) | 3 |
| NO-95C | 200 | 1.8 ± 0.3 | 8.62 (5.47–14.61) | ||
| Cry1Ab | LAB-PS | 400 | 2.5 ± 0.2 | 1.60 (1.35–1.90) | 19 |
| NO-95C | 400 | 1.5 ± 0.2 | 30.72 (21.15–46.85) | ||
| Cry1Ac | LAB-PS | 200 | 1.9 ± 0.3 | 0.74 (0.50–1.16) | 20 |
| NO-95C | 200 | 2.1 ± 0.3 | 14.77 (11.16–20.64) | ||
| Cry1Ca | LAB-PS | 200 | 2.2 ± 0.3 | 10.55 (8.08–14.20) | 11 |
| NO-95C | 160 | 1.7 ± 0.7 | 116.57 (59.58–50,100) | ||
| Cry1Cc | LAB-PS | 200 | 2.8 ± 0.3 | 0.33 (0.26–0.41) | 17 |
| NO-95C | 200 | 2.2 ± 0.4 | 5.46 (4.25–7.40) | ||
| Cry1Fa | LAB-PS | 200 | 2.2 ± 0.3 | 2.43 (1.86–3.21) | 7 |
| NO-95C | 200 | 2.1 ± 0.3 | 17.94 (13.28–23.88) | ||
| Cry1Ja | LAB-PS | 200 | Not available | 3.53 (not available) | 1 |
| NO-95C | 200 | 4.0 ± 1.1 | 3.17 (2.00–4.31) | ||
| Cry9Ca | LAB-PS | 200 | 2.1 ± 0.3 | 1.03 (0.78–1.35) | 2 |
| NO-95C | 200 | 2.7 ± 0.3 | 2.12 (1.67–2.68) |
For all toxins except Cry1Ja, the LC50 for NO-95C was significantly greater than the LC50 for LAB-PS by nonoverlap of 95% FL.
RR, resistance ratio (LC50 for NO-95C/LC50 for LAB-PS).
The Cry1C liquid formulation was from Dow Agrosciences, with concentration in milliliters of Cry1C per liter.
TABLE 3.
Responses of diamondback moth larvae to single concentrations of B. thuringiensis toxins
| Toxin | Concn (mg/liter) | % Mortalitya
|
||
|---|---|---|---|---|
| LAB-PS | NO-95C | NO-QA | ||
| Cry1Aa | 1 | 18.4 | 0 | |
| 10 | 94.7 | 89.7 | ||
| Cry1Ab | 1 | 24.4 | 5.1 | |
| 10 | 97.5 | 44.9 | ||
| Cry1Ac | 1 | 74.3 | 19.9 | |
| 10 | 100 | 69.2 | ||
| Cry1Bb | 10 | 0 | 2.6 | |
| 100 | 7.9 | 10.3 | ||
| 500 | 81.6 | 71.8 | 76.9 | |
| Cry1Ca | 1 | 2.6 | 0 | |
| 10 | 65.8 | 2.6 | ||
| Cry1Fa | 1 | 39.5 | 7.7 | |
| 10 | 100 | 15.4 | ||
| Cry1Ja | 3 | 18.4 | 38.5 | 0 |
| 10 | 100 | 100 | 2.7 | |
| Cry2Aa | 10 | 0 | 12.8 | 2.6 |
| 100 | 25.2 | 15.4 | 10.6 | |
| Cry9Cab | 1 | 42.5 | 12.5 | |
| 10 | 97.5 | 95.0 | ||
Mortality was measured at 5 days and was corrected for control mortality by using Abbott's method.
Data came from the second set of bioassays.
Stability of Cry1C resistance.
Resistance to Cry1C in NO-95C declined when selection with Cry1C stopped for many generations (Tables 1 and 4). During the course of selection, variation in concentration and crosses with LAB-PS in generations 19 and 31 contributed to fluctuations in mortality (Table 1). However, after 13 generations without selection and without crossing to LAB-PS, mortality at 5 ml of Cry1C per liter increased from 52.2% at generation 62 to 99.5% at generation 75 (Table 1). NO-95C was selected four times at 5 ml of Cry1C per liter in six generations between the first and second measurements of LC50. Compared with a susceptible strain, the resistance of NO-95C to Cry1C increased from 22-fold (generations 1 and 2 pooled) to 76-fold (generation 8). In contrast, between generations 45 to 60, only three selections occurred and the resistance ratio declined from 48 to 17 (Table 4).
TABLE 4.
Stability of Cry1C resistance in diamondback moth strain NO-95C
DISCUSSION
Cross-resistance to B. thuringiensis toxins differs between the NO-95C and NO-QA strains of the diamondback moth. Both strains originated from the same watercress farm in Hawaii (9), but NO-95C was selected in the laboratory with Cry1C, while NO-QA was selected with B. thuringiensis subsp. kurstaki. NO-95C evolved resistance to Cry1C, but NO-QA did not (9). NO-95C had little cross-resistance to Cry1Aa, whereas NO-QA was extremely resistant to Cry1Aa (20). In contrast to the strong cross-resistance of NO-QA to Cry1Ja seen here (Table 3) and previously (18, 20, 22), NO-95C was not cross-resistant to Cry1Ja. Cross-resistance of NO-95C was 19-fold for Cry1Ab and 20-fold for Cry1Ac (Table 2), which is higher than the cross-resistance of NO-95C to Cry1Aa but much lower than the >1,000-fold resistance of NO-QA to these toxins (17, 20).
Previous work showed that NO-95C resistance to Cry1Ab was inherited independently from its resistance to Cry1C (7). Thus, resistance to Cry1Ab in NO-95C suggests that, despite two crosses with the susceptible LAB-PS strain, we did not completely eliminate a gene or genes conferring resistance to Cry1Ab (9) from NO-95C. Nonetheless, the lack of cross-resistance to Cry1Ja and relatively low cross-resistance to Cry1A toxins imply that NO-95C did not have a high frequency of the multitoxin resistance gene that confers resistance to Cry1Aa, Cry1Ab, Cry1Ac, Cry1Fa, and Cry1Ja in NO-QA (3, 19, 22).
Resistance to Cry1C in NO-95C was at most 76-fold greater, which is much lower than the over 60,000-fold-greater resistance to Cry1C in the Cry1C-Sel strain of diamondback moth from Florida (25). Nonetheless, resistance to Cry1C was unstable in both NO-95C (Tables 1 and 4) and Cry1C-Sel (25). Unstable resistance is a desirable characteristic for resistance management that can be exploited by rotating different B. thuringiensis toxins or by rotating B. thuringiensis toxins with other pest management agents.
Studies of cross-resistance in the NO-95C strain and in various other resistant strains of the diamondback moth (1, 5, 18, 23) suggest that resistance to Cry1A and Cry1C toxins confers little or no cross-resistance to Cry1Bb, Cry2Aa, or Cry9Ca. Therefore, these toxins might be useful in rotations or combinations with Cry1A and Cry1C toxins. Cry9Ca was much more potent than Cry1Bb or Cry2Aa and thus might be especially useful against the diamondback moth.
ACKNOWLEDGMENTS
We thank L. Anstine, J. Barnard, R. Biggs, S. Borgquist, W. Chang, M. Choo, N. Finson, B. Helvig, J. Riley, M. Silva, A. Taguchi, and J. Tuitele for technical assistance. We thank Juan Ferré for thoughtful comments on the manuscript. We also thank Dow Agrosciences, Ecogen, and Plant Genetics Systems for providing insecticidal materials for testing.
This study was supported by a USDA Western Regional PIAP grant, USDA/CSRS Special Grant 95-34135-1771, USDA NRI grant 96-35302-3470, and the University of Arizona.
REFERENCES
- 1.Ferré J, Real M D, van Rie J, Jansens S, Peferoen M. Resistance to the Bacillus huringiensis bioinsecticide in a field population of Plutella xylostella is due to a change in a midgut membrane receptor. Proc Natl Acad Sci USA. 1991;88:5119–5123. doi: 10.1073/pnas.88.12.5119. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 2.Frutos R, Rang C, Royer M. Managing insect resistance to plants producing Bacillus thuringiensis toxins. Crit Rev Biotechnol. 1999;19:227–276. [Google Scholar]
- 3.Heckel D G, Gahan L G, Liu Y-B, Tabashnik B E. Genetic mapping of resistance to Bacillus thuringiensis toxins in diamondback moth using biphasic linkage analysis. Proc Natl Acad Sci USA. 1999;96:8373–8377. doi: 10.1073/pnas.96.15.8373. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 4.James C. Global status of commercialized transgenic crops: 1999. The International Service for the Acquisition of Agri-biotech Applications (ISAAA) Briefs no. 17. Ithaca, N.Y: ISAAA; 2000. [Google Scholar]
- 5.Lambert B, Buysse L, Decock C, Jansens S, Piens C, Bernadette S, Seurinck J, van Audenhove K, Van Rie J, van Vliet A, Peferoen M. A Bacillus thuringiensis insecticidal crystal protein with a high activity against members of the family Noctuidae. Appl Environ Microbiol. 1996;62:80–86. doi: 10.1128/aem.62.1.80-86.1996. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 6.Liu Y-B, Tabashnik B E. Visual determination of sex of diamondback moth larvae. Can Entomol. 1997;129:585–586. [Google Scholar]
- 7.Liu Y-B, Tabashnik B E. Inheritance of resistance to Bacillus thuringiensis toxin Cry1C in diamondback moth. Appl Environ Microbiol. 1997;63:2218–2223. doi: 10.1128/aem.63.6.2218-2223.1997. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 8.Liu Y-B, Tabashnik B E. Elimination of a recessive allele conferring resistance to Bacillus thuringiensis from a heterogeneous strain of diamondback moth (Lepidoptera: Plutellidae) J Econ Entomol. 1998;91:1032–1037. [Google Scholar]
- 9.Liu Y-B, Tabashnik B E, Pusztai-Carey M. Field-evolved resistance to Bacillus thuringiensis toxin Cry1C in diamondback moth (Lepidoptera: Plutellidae) J Econ Entomol. 1996;89:798–804. [Google Scholar]
- 10.Liu Y-B, Tabashnik B E, Masson L, Escriche B, Ferré J. Binding and toxicity of Bacillus thuringiensis protein Cry1C to susceptible and resistant diamondback moth (Lepidoptera: Plutellidae) J Econ Entomol. 2000;93:1–6. doi: 10.1603/0022-0493-93.1.1. [DOI] [PubMed] [Google Scholar]
- 11.Perez C J, Shelton A M. Resistance of Plutella xylostella (L.) (Lepidoptera: Plutellidae) to Bacillus thuringiensis Berliner in Central America. J Econ Entomol. 1997;90:87–93. [Google Scholar]
- 12.SAS Institute. SAS user's guide: statistics. 5th ed. Cary, NC: SAS Institute; 1985. [Google Scholar]
- 13.Schnepf E, Crickmore N, van Rie J, Lereclus D, Baum J, Feitelson J, Zeigler D R, Dean D H. Bacillus thuringiensis and its pesticidal crystal proteins. Microbiol Mol Biol Rev. 1998;62:775–806. doi: 10.1128/mmbr.62.3.775-806.1998. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 14.Tabashnik B E. Evolution of resistance to Bacillus thuringiensis. Annu Rev Entomol. 1994;39:47–79. doi: 10.1146/annurev.ento.54.110807.090518. [DOI] [PubMed] [Google Scholar]
- 15.Tabashnik B E. Transgenic crops for the Pacific Basin: prospects and problems. In: Zalucki M P, Drew R A I, White G G, editors. Proceedings of the Sixth Australasian Applied Entomology Research Conference. University of Queensland, Brisbane, Australia. 1:161–168. 1998. [Google Scholar]
- 16.Tabashnik B E, Cushing N L, Finson N, Johnson M W. Field development of resistance to Bacillus thuringiensis in diamondback moth (Lepidoptera: Plutellidae) J Econ Entomol. 1990;83:1671–1676. [Google Scholar]
- 17.Tabashnik B E, Finson N, Johnson M W, Moar W J. Resistance to toxins from Bacillus thuringiensis subsp. kurstaki causes minimal cross-resistance to B. thuringiensis subsp. aizawai in the diamondback moth (Lepidoptera: Plutellidae) Appl Environ Microbiol. 1993;59:1332–1335. doi: 10.1128/aem.59.5.1332-1335.1993. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 18.Tabashnik B E, Malvar T, Liu Y-B, Finson N, Borthakur D, Shin B-S, Park S-H, Masson L, de Maagd R A, Bosch D. Cross-resistance of the diamondback moth indicates altered interactions with domain II of Bacillus thuringiensis toxins. Appl Environ Microbiol. 1996;62:2839–2844. doi: 10.1128/aem.62.8.2839-2844.1996. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 19.Tabashnik B E, Liu Y-B, Finson N, Masson L, Heckel D G. One gene in diamondback moth confers resistance to four Bacillus thuringiensis toxins. Proc Natl Acad Sci USA. 1997;94:1640–1644. doi: 10.1073/pnas.94.5.1640. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 20.Tabashnik B E, Liu Y-B, Malvar T, Heckel D G, Masson L, Ballester V, Granero F, Mensua J L, Ferré J. Global variation in the genetic and biochemical basis of diamondback moth resistance to Bacillus thuringiensis. Proc Natl Acad Sci USA. 1997;94:12780–12785. doi: 10.1073/pnas.94.24.12780. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 21.Tabashnik B E, Liu Y-B, Malvar T, Heckel D G, Masson L, Ferre J. Insect resistance to Bacillus thuringiensis: uniform or diverse? Philos Trans R Soc Lond B. 1998;353:1751–1756. [Google Scholar]
- 22.Tabashnik B E, Johnson K W, Engleman J T, Baum J A. Cross-resistance to Bacillus thuringiensis toxin Cry1Ja in a strain of diamondback moth adapted to artificial diet. J Invert Pathol. 2000;76:81–83. doi: 10.1006/jipa.2000.4941. [DOI] [PubMed] [Google Scholar]
- 23.Tang J D, Shelton A M, Van Rie J, de Roeck S, Moar W J, Roush R T, Peferoen M. Toxicity of Bacillus thuringiensis spore and crystal protein to resistant diamondback moth (Plutella xylostella) Appl Environ Microbiol. 1996;62:564–569. doi: 10.1128/aem.62.2.564-569.1996. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 24.Wright D J, Iqbal M, Granero F, Ferré J. A change in the single receptor in the diamondback moth (Plutella xylostella) is only in part responsible for field resistance to Bacillus thuringiensis subsp. kurstaki and B. thuringiensis subsp. aizawai. Appl Environ Microbiol. 1997;63:1814–1819. doi: 10.1128/aem.63.5.1814-1819.1997. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 25.Zhao J-Z, Collins H L, Tang J D, Cao J, Earle E D, Roush R T, Herrero S, Escriche B, Ferré J, Shelton A M. Development and characterization of diamondback moth resistance to transgenic broccoli expressing high levels of Cry1C. Appl Environ Microbiol. 2000;66:3784–3789. doi: 10.1128/aem.66.9.3784-3789.2000. [DOI] [PMC free article] [PubMed] [Google Scholar]