Transgenic crops producing insecticidal toxins from Bacillus thuringiensis (Bt) are commercially successful in reducing pest damage, yet knowledge of resistance mechanisms that threaten their sustainability is incomplete. Insect resistance to the pore-forming Cry1Ac toxin is correlated with the loss of high-affinity, irreversible binding to the mid-gut membrane, but the genetic factors responsible for this change have been elusive. Mutations in a 12-cadherin-domain protein confer some Cry1Ac resistance but do not block this toxin binding in in vitro assays. We sought to identify mutations in other genes that might be responsible for the loss of binding. We employed a map-based cloning approach using a series of backcrosses with 1,060 progeny to identify a resistance gene in the cotton pest Heliothis virescens that segregated independently from the cadherin mutation. We found an inactivating mutation of the ABC transporter ABCC2 that is genetically linked to Cry1Ac resistance and is correlated with loss of Cry1Ac binding to membrane vesicles. ABC proteins are integral membrane proteins with many functions, including export of toxic molecules from the cell, but have not been implicated in the mode of action of Bt toxins before. The reduction in toxin binding due to the inactivating mutation suggests that ABCC2 is involved in membrane integration of the toxin pore. Our findings suggest that ABC proteins may play a key role in the mode of action of Bt toxins and that ABC protein mutations can confer high levels of resistance that could threaten the continued utilization of Bt–expressing crops. However, such mutations may impose a physiological cost on resistant insects, by reducing export of other toxins such as plant secondary compounds from the cell. This weakness could be exploited to manage this mechanism of Bt resistance in the field.
Crystal toxin proteins from Bacillus thuringiensis (Bt) make ideal bioinsecticides because of their high potency against certain insects and lack of activity against most other species. Transgenic cotton and maize expressing pore-forming Cry1A Bt-toxins are now widely used in agriculture, enabling substantial reductions in the use of chemical insecticides. However this has greatly increased the selection pressure in pest populations for toxin resistance. Preventing or delaying the development of this resistance is a high priority, to avoid a replay of the onset of insecticide resistance brought on by dependency on chemical pesticides. Because the molecular details of Bt mode of action are still not fully understood, insect strains collected from the field and selected to high levels of resistance in the laboratory are useful in discovering the obstacles the toxin must overcome before it finally forms the pore and kills the insect. We used a genetic approach to explore a poorly understood step in the toxin mode of action, which is blocked in an extremely resistant strain of an important cotton pest. As well as providing the tools to diagnose this type of resistance when it appears in the field, this discovery suggests factors that may counteract its eventual spread.
Citation: Gahan LJ, Pauchet Y, Vogel H, Heckel DG (2010) An ABC Transporter Mutation Is Correlated with Insect Resistance to Bacillus thuringiensis Cry1Ac Toxin. PLoS Genet 6(12): e1001248. doi:10.1371/journal.pgen.1001248
Editor: Rodney Mauricio, University of Georgia, United States of America
Received: September 13, 2010; Accepted: November 16, 2010; Published: December 16, 2010
Copyright: © 2010 Gahan et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This research was supported by the Max-Planck-Gesellschaft and US National Science Foundation Grant 1517107. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Insecticidal protein toxins of the Cry1A family produced by certain strains of the gram-positive bacterium Bacillus thuringiensis (Bt) are highly active against many Lepidoptera but nontoxic to most other animal species. Transgenic cotton producing Cry1Ac and transgenic maize producing Cry1Ab have been grown commercially since 1996 and offer protection against some major pests, including species in the genera Heliothis, Helicoverpa, Ostrinia, and Pectinophora , . After ingestion and solubilization in the alkaline midgut lumen of the caterpillar, the protoxin is cleaved by digestive proteases to yield an active 60 kDa toxin which interacts with high-affinity binding sites on the brush border epithelium, eventually oligomerizing to form a transmembrane pore, leading to lysis of epithelial cells , . Additional mechanisms of toxicity involving an adenylyl cyclase/PKA signaling pathway have also been described . High toxin concentrations are lethal, lower toxin concentrations inhibit larval growth in a dose-dependent manner. The binding targets are critical in determining the range of species on which the toxin is active , and reduction or loss of binding is an important mechanism of genetically based resistance in the target pest species .
The most common type of Bt toxin resistance (“Mode 1”)  which has evolved in field populations of Plutella xylostella in response to sprays of formulated Bt toxins  and in laboratory-selected strains of other Lepidoptera – is characterized by recessive inheritance, >500-fold resistance to at least one Cry1A toxin, much less resistance to Cry1C, and greatly reduced binding of Cry1A toxins to target sites in the midgut membrane. Several cases of resistance to Bt crops in field populations of insect pests have also been reported, but the genetic basis of resistance has not been identified in any of these cases , , . Genetic mutations linked to Cry1A resistance have been identified in laboratory strains, but their role in Mode 1 resistance is still not fully understood. The mutations most commonly found in Cry1A-resistant strains inactivate a gene encoding a 12-cadherin-domain protein of about 1750 amino acids, expressed in the larval midgut –. These mutations confer resistance to Cry1A toxins including Cry1Ac, but do not block irreversible Cry1Ac binding to midgut membranes as measured by in vitro assays , , . Conversely, Mode 1 resistance in the NO-QA strain of P. xylostella which includes loss of Cry1Ac binding  is determined by a single gene that segregates independently from the 12-cadherin-domain protein gene .
What could account for the apparent independence of resistance-conferring cadherin mutations and resistance-conferring loss of irreversible membrane binding? There is evidence for a multi-step mechanism that could offer an explanation. Bravo et al.  have proposed that activated Cry1A toxin monomers first bind to an extracellular membrane-proximal domain of the 12-cadherin-domain protein. The toxin undergoes a conformational change, facilitating proteolytic cleavage of the Domain I helix α1 from the toxin N-terminus by a yet-uncharacterized protease. The resulting “clipped” toxin monomers subsequently assemble into a oligomeric pre-pore structure in solution, which binds reversibly to several other membrane-bound proteins, and finally inserts irreversibly into the membrane . Thus absence of the cadherin protein in resistant strains would slow the rate of monomer clipping and oligomerization of the active pore structure, but not directly affect the subsequent irreversible binding and insertion of the pore into the membrane. This would predict that even higher levels of resistance could be attained by interfering with the later binding steps in this sequential binding model. To test this idea, it would be useful to examine toxin binding to membranes of resistant strains, with or without the cadherin protein.
The first Bt-resistant cadherin mutation was identified  as the resistant allele 4r of the previously-mapped gene BtR-4  in the YHD2 strain of the cotton pest Heliothis virescens. This strain had evolved >10,000 fold resistance in response to laboratory selection by diet-incorporated Cry1Ab and Cry1Ac toxin over four years . Insertion of an LTR retrotransposon into the coding sequence of the 12-cadherin-domain protein defines the 4r allele, and resulted in a truncated 622-amino acid protein lacking the last 7 cadherin domains, membrane-proximal toxin binding region, transmembrane domain, and cytoplasmic domain. The absence of the 12-cadherin-domain protein from the midgut membranes of YHD2 was confirmed with antibodies . The first binding measurements on homozygous resistant 4r4r YHD2 published in 1995 showed greatly reduced binding of midgut epithelial brush border membrane vesicles (BBMV) to Cry1Aa, but surprisingly no reduction in Cry1Ab or Cry1Ac binding . YHD2 was subsequently selected to even higher levels of resistance, and later studies published in 2002 and 2004 showed a loss of membrane binding by Cry1Ab and Cry1Ac also, as well as a reduction in their pore-forming ability , . This suggested the existence of a second gene (which we named BtR-6) with a mutant allele 6r in the more resistant strain responsible for its increased resistance and decreased binding affinity to Cry1Ac.
In order to test the hypothesis that a separate mechanism affecting later steps in toxin binding existed in this more resistant strain, we sought to identify BtR-6 and the nature of the 6r allele by map-based cloning. We first isolated the two resistance mechanisms into separate strains and characterized their toxin-binding properties. We then used these strains in a series of backcrosses that were assayed for resistance using a sublethal, growth-inhibition bioassay. Fine-scale linkage mapping identified a cluster of ABC transporter genes, one of which showed an inactivating mutation in the most resistant strain. This implicates the ABC transporter family for the first time in the mode of action of Bt Cry1A toxins, and offers an explanation for Mode 1 resistance that is compatible with the sequential binding model.
Construction of Strains
In order to synthesize strains that were homozygous for different combinations of resistant and susceptible alleles at the BtR-4 and BtR-6 loci (4r vs 4s, 6r vs 6s), we used a combination of progeny testing on Cry1Ac-containing diet and marker-assisted selection of parents with a PCR (polymerase chain reaction) test diagnostic for 4r . These strains were then maintained on artificial diet containing the highest concentration of Cry1Ac that would allow the same larval growth rate as toxin-free diet. Strain YHD3 was homozygous resistant for both genes (4r4r/6r6r), had a resistance level similar to the newer YHD2, and was reared on 200 µg/ml Cry1Ac. YFO was 4r4r/6s6s and could be reared on at most 5 µg/ml Cry1Ac. YEE was 4s4s/6r6r and was reared on 50 µg/ml Cry1Ac. Fully susceptible strains CNW and JEN were 4s4s/6s6s and were reared on toxin-free diet; their growth rate was reduced 50% by only 0.064 µg/ml Cry1Ac.
Toxin Binding Measurements
Qualitative in vitro binding studies with Cry1Aa, Cry1Ab, and Cry1Ac using BBMV (Figure 1) showed that the doubly homozygous susceptible JEN strain (4s4s/6s6s) bound all three toxins as expected. The doubly homozygous resistant YHD3 strain (4r4r/6r6r) bound to none of the three, similar to YHD2 in 2002  and 2004 . The two intermediately resistant strains showed a complementary pattern: YFO (4r4r/6s6s, the hypothesized genotype of the older YHD2 strain) had lost only the ability to bind Cry1Aa, similar to YHD2 in 1995 . YEE (4s4s/6r6r) still bound Cry1Aa but failed to bind Cry1Ab and Cry1Ac, a pattern that has not been previously reported (Figure 1). Thus homozygosity for the 6r allele but not 4r is correlated with loss of Cry1Ac binding.
Qualitative binding of biotinylated Cry1Aa, Cry1Ab and Cry1Ac to brush border membrane vesicles (BBMV) of midguts from susceptible and resistant H. virescens strains. Biotinylated toxins (2.5 nM) were incubated with BBMV (20 µg protein) from the following strains: susceptible JEN (lane 1) and resistant YHD3 (lane 3), YEE (lane 4) and YFO (lane 5). BBMV were pelleted by centrifugation, and bound toxins were resolved by electrophoresis, blotted onto membranes and detected by a chemiluminescent-coupled streptavidin probe. Binding specificity was assessed by incubating biotinylated toxin with BBMV from the susceptible strain JEN in the presence of a 200-fold excess of unlabeled toxin (lane 2).
We explored the genetic basis of these resistance and binding differences by linkage mapping using a larval growth bioassay with Cry1Ac conducted on backcrosses as done previously . Backcrosses to YHD3 using F1 (YHD3 x YFO) mothers were first screened with a panel of RFLP markers to identify the linkage group containing BtR-6, by exploiting the absence of crossing-over during meiosis in female Lepidoptera. A probe previously mapped to linkage group 2 (LG2) with similarity to a microsomal glutathione transferase (GenBank HM150720) showed a highly significant association with resistance as measured by larval weight on the Cry1Ac diet. This confirmed that BtR-6 was genetically distinct from the two previously mapped resistance genes in this species, BtR-4 (the 12-cadherin-domain protein) on LG9  and BtR-5 on LG10 . Neither LG9 nor LG10 had a significant association with resistance in these crosses (in the backcross to YFO, all progeny are BtR-4r4r). The significant effect of LG2 was confirmed in backcrosses to YEE using F1 (YEE x YFO) mothers, which were also segregating at BtR-6. Ribosomal protein genes RpP0, RpS5, RpL8, RpL10A, and RpL30 also mapped to LG2 in H. virescens, indicating homology with Chromosome 15 (Chr15) of the domesticated silkmoth Bombyx mori, where these same genes had been mapped by recombinational  and cytogenetic  methods.
We localized BtR-6 relative to marker genes along LG2 using recombinational mapping in backcrosses with F1 males, which do undergo crossing-over during meiosis. In the first step, markers were chosen from H. virescens and Helicoverpa armigera cDNA clones homologous to genes that had been genetically mapped to Chr15 in B. mori. The second step at a finer scale used genes physically mapped to Chr15 after the assembled B. mori genome sequence was made available to the public in April 2008. The linkage map of LG2 in H. virescens was entirely collinear with the genetic and physical maps of Chr15 of B. mori (Figure 2). BtR-6 was localized within the interval between markers b7730 and b7793, showing zero recombinants out of a total of 1060 informative progeny from 3 sets of mapping families that had been reared on Cry1Ac-containing diet. The physical map of this region in B. mori contains 10 predicted genes, nine of which showed expression in B. mori larval midgut as indicated by microarray studies, and which also had homologs in cDNA libraries constructed from midgut tissue of larval H. armigera (Table S2).
Markers were mapped in 1,060 offspring from 3 sets of backcross families (cM = Haldane centimorgans). The linkage map is compared with the physical map of homologs on Bombyx mori Chromosome 15 (Mb = megabases of DNA). The scale of the middle portion of both maps is magnified 10-fold.
A PCR product corresponding to B. mori predicted gene BGIBMGA007793 was amplified from H. virescens midgut cDNA, and used to screen BAC libraries of H. virescens and its sister species H. subflexa. The latter library yielded a positive clone which was sequenced (GenBank Accession No. GQ332573, Figure 3A), revealing a cluster of three genes with high sequence similarity to ABC transporters (ABCC1, ABCC2, and ABCC3), in the same orientation as the corresponding region in B. mori (Figure 3B). Genomic sequence comparison of the ABCC2 gene from YHD3 (GenBank GQ332572) and YFO (GenBank GQ332571) strains revealed a 22-bp deletion in exon 2 occurring only in YHD3 (Figure S2). The same deletion was found in RT-PCR products from YHD3 larval midgut cDNA. The frameshift generated by this deletion predicts a truncated 99-residue protein from YHD3 mRNA. In contrast, the full-length ABCC2 protein of 1339 amino acids predicted from the YFO or H. subflexa sequence (97% amino acid identity) has all the features of the bipartite structure of ABC transporters, with six transmembrane segments and a large cytoplasmic ATP-binding domain in each half  (Figure 4, Figure 5). A PCR assay using primers flanking the exon 2 deletion region was used to determine genotypes of individual backcross progeny (Figure S1). Only those larvae with two copies of the exon 2 deletion allele grew rapidly on Cry1Ac-containing diet. The deletion was present in all YHD3 and YEE individuals tested; no YFO or CNW individuals had the deletion. This 22-bp deletion is taken to define the 6r allele of BtR-6 in YHD3.
H. subflexa BAC clone 89F08 (A) and corresponding region from Chromosome 15 of B. mori with BGI protein predictions (B). The asterisk marks exon 2 of ABCC2 containing the 22 bp deletion characterizing the 6r allele of BtR-6 in H. virescens.
Hv-YHD3 is the predicted protein sequence of the YHD3 exon 2 frameshift mutant (GenBank GQ332572); HvABCC2 is from the YFO strain of Heliothis virescens (GQ332571); HsABCC2 is from Heliothis subflexa (GQ332573); BmABCC2 is from Bombyx mori; CG4562 is isoform A of CG4562 from Drosophila melanogaster (AAF44707); AAI50823 is ATP-binding cassette, sub-family C (CFTR/MRP) from Mus musculus (AAI50823). Sequence features determined for HvABCC2 include tm1 through tm12: transmembrane domains as predicted by Phobius . EC1 through EC6: predicted extracellular loops. Dashed line, cd03250: ABCC_MRP_domain1, Domain 1 of the ABC subfamily C (E = 5e-77); dotted line, cd03244: ABCC_MRP_domain2, Domain 2 of the ABC subfamily C (E = 2e-95). Boxed regions are A/P: Walker A motif/P-loop; Q: Q-loop/lid; C: ABC transporter signature motif; B: Walker B motif; D: D-loop; H: H-loop/switch region .
Temporal Allele Frequency Correlations with Toxin Binding
We used PCR analysis of archival DNA samples to investigate whether an increase of the 6r allele frequency occurred concomitantly with the decrease in Cry1Ac binding affinity of YHD2 over the years. DNA from parents of YHD2 backcrosses conducted in March 1993  yielded a 6r allele frequency estimate of 14% and an expected 2% frequency of 6r6r homozygotes; not high enough to appreciably reduce the binding to Cry1Ac . Thus the 6r allele was present although rare in the YHD2 strain as early as 1993. When we screened for 6r in DNA that had been isolated in December 2002 from the YHD2 larvae whose BBMV showed a loss of Cry1Ab and Cry1Ac binding , we found that the frequency of 6r had increased to 100%. Thus a loss of Cry1Ab and Cry1Ac binding was correlated with an increase in 6r within YHD2 over approximately 100 generations while the Cry1Ac resistance level as measured by bioassay also increased. This correlation also extended to other strains. DNA samples from the Cry1Ac-resistant KCBhyb strain had a 6r frequency of 5%; membranes from these larvae retained Cry1Ab and Cry1Ac binding, and binding of Cry1Aa only was dependent on the BtR-4 genotype . Both the Cry1Ac-resistant strain CxC with a 6r allele frequency of 0% and the Cry1Ac-susceptible strain YDK with 6% retained Cry1Aa, Cry1Ab and Cry1Ac binding  (Table S3).
Recent research has shown that the mode of action of Bt toxins is more complex than originally envisioned. Cry toxins may induce cell death by interacting with the 12-cadherin-domain protein without forming pores ; responses to Cry toxins may involve intracellular signal transduction pathways that protect cells against pore forming toxins , . Yet a major feature of Cry1A toxin action in Lepidoptera is the formation of pores in the plasma membrane leading to cell disruption by colloid-osmotic lysis . At high enough concentrations, Cry toxins can eventually insert and form pores in planar lipid bilayer membranes devoid of any other protein , . However, these toxins have evolved to interact with a series of host proteins in the midgut membrane to form pores much faster and at much lower concentrations. These interactions are toxin- and host-specific, e.g. Cry1A toxins are active against certain Lepidoptera, but not Diptera or Coleoptera. Interfering with one or more of these steps may confer resistance, such that higher concentrations of toxin are required to achieve the same mortality endpoint. Identifying the molecular changes that accompany resistance is a useful first step to posing hypotheses about the mode of toxin action. Based on the mapping results and binding correlations described here, we hypothesize that the ABCC2 protein participates in the mechanism of Cry1Ab and Cry1Ac toxicity by binding and facilitating insertion into the membrane, in an extension of the multi-step model of Bravo et al. .
In the first step of this model, reversible toxin binding to the 12-cadherin-domain protein accelerates the formation of clipped toxin monomers which are more competent to form the oligomeric pre-pore structure in solution. Evidence supporting this mechanism includes the enhanced toxicity of Cry1Ab or Cry1Ac toxin when fed to larvae along with a peptide fragment from the toxin-binding domain of the cadherin protein . This fragment itself binds to Cry1Ab and Cry1Ac , and accelerates the rate of formation of a 250 kDa oligomer of Cry1Ac . Additional evidence is provided by the elevated potency of “pre-clipped” Cry1Ab or Cry1Ac monomers generated by recombinant methods, which lack the α1 helix . These modified Cry1AbMod and Cry1AcMod toxin monomers rapidly form oligomers in the absence of the cadherin, and are more potent than unmodified toxins against Cry1Ac-resistant Pectinophora gossypiella with cadherin mutations , although possessing similar properties in most other respects .
According to this model, absence of the 12-cadherin-domain protein confers a certain level of resistance to Cry1Ab or Cry1Ac by slowing down the formation of oligomers, not by stopping it completely. Evidently oligomerization of these two toxins can occur in the absence of cadherin binding, but at a slower rate; since higher concentrations of Cry1Ab or Cry1Ac are still capable of killing resistant insects with cadherin mutations. Moreover, even if the cadherin functions in accelerating the “clipping” of Cry1Ab and Cry1Ac toxin monomers, this does not rule out a separate role in additional binding events. The cadherin appears to be the major binding protein for Cry1Aa; as BBMV from strains lacking it have also lost their ability to bind Cry1Aa , . Furthermore, presence of the cadherin appears to be necessary and sufficient for binding of BBMV to Cry1Aa, but not Cry1Ab or Cry1Ac (Figure 1). The 12-cadherin-domain protein from B. mori also binds to Cry1Aa , but experiments on the effect of the cadherin on oligomerization have not yet been conducted on Cry1Aa. Therefore, cadherin binding may play more than one role, depending on the toxin.
In the second binding step in the hypothesized mode of action , toxin oligomers bind to the soluble ectodomains of membrane-associated glycosylated proteins such as aminopeptidase N (APN) , , alkaline phosphatase , , P252 glycoprotein , or BTR-270 glycoprotein . These proteins are GPI-anchored and enriched in lipid rafts, and disruption of lipid rafts by cholesterol depletion reduces pore formation by Cry1Ab . Experimental cleavage of GPI anchors removes APN from the BBMV surface and reduces the amount of Cry1Ab toxin inserted into the membrane . Massive shedding of GPI-anchored proteins by the action of endogenous phospholipase C has been shown to occur in response to toxin consumption , which might be a defense mechanism against the second binding step, but so far this has not been observed to occur in any resistant strains.
Toxin binding to these glycoproteins appears to be predominantly reversible; e.g. binding of Cry1Ac to purified APN exhibits measurable on- and off-kinetics by surface plasmon resonance –. No single glycoprotein appears to be essential for Cry1A toxicity; e.g. mutants of Cry1Ac which eliminate binding to a 115 kDa APN only result in a two-fold decrease in toxicity . RNA interference directed against midgut APNs produces a measurable but slight decrease of toxicity , . Therefore the main significance of Cry1A toxin binding to these glycoproteins seems to be to increase the concentration of the pre-pore oligomer at the membrane surface, increasing the probability of eventual insertion by some other mechanism.
The final binding step proposed here involves interactions of the oligomeric toxin pre-pore structure with the ABCC2 protein. ABC transporters cycle between closed and open configurations as they transport small molecules out of the cell, driven by binding and hydrolysis of ATP by the intracellular nucleotide-binding domains. A recently determined structure for the ABCB1 P-glycoprotein shows that in the closed configuration, the extracellular loops between the transmembrane domains completely cover the channel opening, resulting in a large internal cavity facing the cytoplasm . In this pretransport state, the small molecule to be transported is located within the internal cavity. Binding of ATP by the two cytoplasmic nucleotide-binding domains causes their dimerization and a large conformational change resulting in the open state, in which several hydrophobic surfaces of the channel are transiently exposed to the outside of the cell while the small molecule is expelled . Hydrolysis of the ATP restores the ABC protein to the closed configuration. We hypothesize that Cry1Ab and Cry1Ac toxins, as pre-formed oligomers or possibly also as monomers, bind to the open configuration of ABCC2 and that this facilitates subsequent membrane insertion. The resistance conferred by BtR-6r would thus be due to the absence of this binding site for Cry1Ab and Cry1Ac. Direct toxin binding assays with the membrane-integrated ABCC2 protein would be required for evaluation of this hypothesis.
To our knowledge, ABC transporters have not yet been suggested as binding targets for Bt toxins. Failure to detect them may be due to the under-representation or absence of integral membrane proteins in 1-D or 2-D gels used in ligand binding studies with labelled toxin , . Failure to isolate them could be due to the general difficulty of isolating membrane proteins. The midgut proteins from Lepidoptera previously isolated on the basis of binding to Cry1A-toxin-immobilized affinity columns , immunoprecipitation ,  or preparative gel electrophoresis  all have a large ectodomain projecting into the lumen available for binding, and are readily solubilized, being attached to the membrane by a GPI anchor or a single transmembrane domain. The predicted structure of ABCC2, however, presents only 6 small loops (of 19, 5, 5, 43, 5, and 5 residues respectively) projecting into the lumen, which connect the 6 α-helices of each of the two transmembrane domains buried in the lipid bilayer (Figure 4, Figure 5). Cry1A toxins are known to bind to carbohydrate residues of glycoproteins, but none of the 6 loops of ABCC2 have predicted glycosylation sites. If the toxin binds primarily to the hydrophobic interior of the channel, then methods stringent enough to solubilize the ABC protein would likely disrupt this interaction.
If confirmed, the role of an ABC transporter in Bt toxin action proposed here could have implications for the management of Cry1Ac resistance in field populations of H. virescens and other lepidopteran pests currently controlled by Bt-cotton or Bt-maize. We emphasize that as no attention has been paid to ABC transporters in Bt resistance previously, we do not know whether this or similar mutations occur in the field in H. virescens or any other species. However, the genetic basis of field-evolved resistance to Bt sprays by Plutella xylostella ,  and Trichoplusia ni , and to Bt crops by Helicoverpa zea, Spodoptera frugiperda, and Busseola fusca  has not yet been identified, and these strains should be examined for ABC transporter mutations. We do not know whether H. virescens larvae homogozygous for the ABCC2 mutation can survive on cotton, with or without Cry1Ac toxin. Developmental arrest in the last larval instar of the YHD2 strain feeding on non-transgenic cotton was observed prior to 1993 , when BtR-4r was nearly fixed, indicating a strong fitness cost to the cadherin mutation; but at that time BtR-6r was still at a very low frequency. We do not know how ABCC2 mutations would respond to selection for Cry1A-toxin resistance in the field. In India, China, and many other countries, the predominant varieties of Bt-cotton still produce the single toxin Cry1Ac, thus selection for Cry1Ac resistance is strong. The Bt-cotton currently used in the USA and Australia produces Cry2Ab in addition to Cry1Ac; the different modes of action of these two toxins are thought to produce a “redundant killing” effect whereby selection for resistance to either single toxin is greatly weakened. However, we do not know whether ABC transporter mutations confer cross-resistance to Cry2Ab. The binding targets of Cry2Ab are unknown and ABC proteins have not yet been investigated as candidates. Moreover, Cry2Ab resistance is detectable using F2 screens in Australian populations of Helicoverpa armigera  and H. punctigera , and the molecular basis of the resistance mechanism involves binding site alterations in both species .
The biological function of ABCC2 is unknown, but its similarity to multidrug resistance proteins suggests that it could export small hydrophobic toxins from midgut epithelial cells for eventual elimination in the feces. Homozygous deletions of ABCC2 as seen in the YEE and YHD3 strains have no obvious effect on insects consuming artificial toxin-free diet in the laboratory. However, plant secondary compounds that deter herbivory or poison the herbivore would be encountered by larvae consuming plants in nature, affecting Bt-susceptible and resistant insects in different ways. If exported by an active ABCC2 in Bt-susceptible insects, they could potentiate the Bt-toxin by increasing the proportion of time the channel is in the open state, exposing the hydrophobic inner surfaces to toxin binding. There is evidence for an effect of different plant tissues with different amounts of secondary compounds on the potency of Cry1Ac . Additionally, by imposing a fitness cost on Bt-resistant insects they could select against resistance alleles encoding defective variants of the ABCC2 protein that fail to export them. For example, Bt-resistant Pectinophora gossypiella is more sensitive to the cotton secondary compound, gossypol . Even a slight fitness cost of ABCC2 mutations would be effective in delaying the increase of resistance alleles, the goal of the high-dose/refuge strategy mandated by the US Environmental Protection Agency. PCR-based DNA diagnostics for specific ABCC2 mutants shown to be present in the field could be useful in supporting the continued success of this strategy by monitoring resistance alleles in field populations of insect pests.
Materials and Methods
Marker-Assisted Selection to Develop Resistant Strains
All crosses used virgin adults of Heliothis virescens in single-pair matings. Resistant strain YHD2 was crossed to the susceptible strain CNW (July 2001) and F1 offspring were intercrossed. F2 progeny were reared on artificial diet  containing 0.2 µg/ml Cry1Ac toxin for 10 days, individually weighed, and transferred to toxin-free diet for rearing to adulthood. The 10-day weights were used as an indication of the ability to resist the growth-inhibiting effect of this sublethal Cry1Ac concentration, due to the presence of different combinations of resistant and susceptible alleles at the two resistance genes BtR-4 and BtR-6. F2 adults from the top third of the weight distribution were intercrossed to form the YHD3 strain, which was subjected to selection on Cry1Ac-containing diet over 25 generations, eventually attaining the same resistance level as the parent YHD2. It was maintained on artificial diet with 200 µg/ml Cry1Ac. To develop the YFO strain, F2 adults from the middle third of the weight distribution were repeatedly backcrossed in single-pair matings to the susceptible CNW strain. Parents were scored for the presence of the BtR-4r allele by PCR using the primers SF1, SR2, and RR3 (Figure S1) , after collection of fertile eggs. Only progeny of parents that still carried the BtR-4r allele were retained for subsequent matings. Larvae of these generations were reared on toxin-free diet to avoid any toxin-based selection of resistance alleles. After 6 generations of backcrossing and PCR screening, YFO adults were intercrossed and subsequent generations made homozygous for BtR-4r, after which the strain was raised on 5 µg/ml Cry1Ac. The YEE strain was developed by intercrossing the F2 from the lower third of the weight distribution and subsequent generations, and keeping only progeny of parents with the lowest frequency of BtR-4r alleles as detected by PCR. Larvae of this strain were reared on diet with 5 µg/ml Cry1Ac toxin to select for BtR-6r alleles. After no parents were found to carry BtR-4r alleles, the YEE strain was maintained on 50 µg/ml Cry1Ac. As YFO was homozygous BtR-4r4r and YEE was subsquently shown to be homozygous BtR-6r6r, the ABCC2 mutation permits larvae to consume 10 times as much Cry1Ac without growth retardation as does the cadherin mutation. All strains showed equivalent growth in the laboratory on artificial diet with no toxin.
Backcross larval progeny were tested by rearing on a sublethal concentration of Cry1Ac in artificial diet , allowing normal growth (i. e. equivalent to susceptible individuals on non-Bt diet over the same time period) in individuals homozygous resistant for the gene segregating in the cross, but suppressing growth in heterozygotes. Larvae were weighed to the nearest 1 mg after 7 days; backcross size distributions were strongly bimodal consistent with segregation of a single major resistance gene (Figure S3). All larvae were then transferred to toxin-free diet and reared to adults for DNA extraction. Polymorphisms at genetic marker loci were scored using RFLPs (restriction fragment length polymorphisms) visualized by Southern blots of restriction-digested genomic DNA, or scored by screening for intron size polymorphisms by PCR using primers placed in adjacent exons.
Three series of interstrain crosses were used to generate backcross families (BRX) segregating at BtR-6. In BRX28 (February 2006) and BRX35 (December 2007), F1 progeny from crosses between YHD3 (4r4r 6r6r) and YFO (4r4r 6s6s) were backcrossed to YHD3. Backcross progeny were expected to be 4r4r 6r6r or 4r4r 6r6s; they were tested on 25 µg/ml Cry1Ac. In BRX36 (June 2008), F1 progeny from crosses between YFO and YEE (4s4s 6r6r) were backcrossed to YEE. Backcross progeny were expected to be 4s4s 6r6r, 4r4s 6r6r, 4s4s 6r6s, or 4r4s 6r6s. To minimize the effect of segregation of the 4r allele, which is recessive at high concentrations, backcross progeny were tested on 50 µg/ml Cry1Ac and otherwise treated as in the other two series.
The linkage analysis strategy exploited the absence of crossing-over during meiosis in female Lepidoptera . Female-informative backcrosses (with F1 mothers) were examined first to verify that segregation of LG2 markers correlated with larval weight. Male-informative backcrosses (with F1 fathers in which crossing-over occurs) were then used to estimate linkage relationships among LG2 markers and resistance as measured by larval weight on Cry1Ac-containing diet.
For RFLP analysis, DNA was isolated from adults using phenol and chloroform, digested with HindIII or PstI, electrophoresed on 0.8% agarose gels, and transferred to Hybond N+ filters for probing with 32P-labelled probes. RFLP probes for LG2 markers were generated from H. virescens or Helicoverpa armigera cDNA probes previously mapped to LG2, or from genes mapped to Bombyx mori Chromosome 15. These were used to search EST databases of H. virescens and H. armigera by BLAST, or to design degenerate PCR primers for amplification and sequencing from H. virescens cDNA or gDNA. Intron size polymorphisms in some markers were scored by agarose gel electrophoresis of PCR products generated using primers positioned in adjacent exons.
Three strategies were used to screen B. mori Chromosome 15 for markers that could be used in mapping, in a sequential approach to narrow the interval containing BtR-6. First, sequence information from the RAPD-based linkage map of Yasukochi et al. ,  was used in BLASTN searches of the wgs section of GenBank (http://www.ncbi.nlm.nih.gov) to identify whole-genome-shotgun contigs produced by the first  and second  genome assemblies, and these in turn were screened for conserved coding sequences present in the H. armigera and H. virescens cDNA libraries. This approach was limited by small contig size and frequent occurrence of chimeric contigs. Second, a BAC-walking strategy was employed using BAC-end sequences deposited in the gss section of GenBank . BAC ends occuring in contigs were identified by BLASTN to gss, the other end was obtained by a text search using the BAC clone name, and used to identify the contig in which it occurred by BLASTN to wgs. Third, when the third genome assembly  was made available to the public on-line on SilkDB (http://silkworm.swu.edu.cn/silkdb/)  and Kaikobase (http://sgp.dna.affrc.go.jp/index.html) , predicted genes in the genome browser view were used. Serial numbers of BGI predicted genes are represented here by the last four digits; e.g. b7795 for BGIBMGA007795. These approaches were successful because of the high degree of evolutionary conservation of gene order among Bombyx and Heliothis for this linkage group.
Recombinants were identified by reference to parental and grandparental genotypes and tallied by hand in order to guide the direction of search for additional markers. The final linkage map was constructed using 20 markers and 1060 offspring using the program Mapmaker3  with Haldane centimorgans. A Macintosh PowerBook running the MacPort implementation of the unix version was used, as the MS-DOS version of this program running under Windows crashed with our dataset.
After BtR-6 was localized within the interval between markers b7730 and b7793 showing zero recombinants, the linkage map of B. mori Chr15 was examined and found to also have zero recombinants out of 190 informative progeny in the corresponding region . The physical map of this region in B. mori contains 10 predicted genes , nine of which showed expression in B. mori larval midgut as indicated by microarray studies  and also had homologs in cDNA libraries constructed from midgut tissue of larval H. armigera (Table S2).
Preparation of Brush Border Membrane Vesicles and Binding Assays
Actively feeding early fifth-instar H. virescens larvae were chilled on ice and dissected, (May 2007). Tracheae, Malpighian tubules, peritrophic matrix and food bolus were removed and the midgut tissue was rinsed briefly in ice-cold phosphate-buffered saline (PBS). Brush border membrane vesicles (BBMV) were prepared by the Mg2+ precipitation method according to Wolfersberger et al. . The final BBMV pellet was resuspended at a protein concentration of 1 mg/ml in PBS (determined by the BCA protein assay with BSA as standard, Bio-Rad) and stored at −80°C until use. Brush border membrane enrichment was estimated by measuring the aminopeptidase activity using L-leucine-p-nitroanilide as a substrate. Typical enrichment of the leucyl-aminopeptidase activity in the BBMV preparation was between 5 and 6 fold compared to the initial midgut homogenate.
E. coli strains harboring individual Cry1Aa, Cry1Ab, or Cry1Ac genes cloned into pKK223-3 were obtained from the Bacillus Genetic Stock Center (Ohio State University). Cry1A protoxins were prepared according to Lee et al. , and were activated by trypsin at a trypsin/protoxin ratio of 1/50 (w/w) at 37°C for 1 h. Activated toxins were further purified by anion exchange chromatography using a 1 ml RESOURCE Q column (GE Healthcare). For toxin biotinylation, 0.5 mg of purified toxins was incubated (1∶30 molar ratio) with NHS-Biotin (Sigma) for 30 min at room temperature. To remove excess biotin, samples were run through a 5 ml HiTrap desalting column (GE healthcare).
Qualitative binding assays were performed by incubating 2.5 nM of each biotinylated Cry1A toxin with BBMV (containing 20 µg protein) for 1 h at room temperature. Then, BBMV were pelleted by centrifugation (13,000 g, 10 min, 4°C) and washed three times with PBS to remove unbound toxin. The final pellet was resuspended in SDS-PAGE sample buffer, boiled for 5 min, and proteins were resolved on a 10% SDS-PAGE gel. Toxin binding was revealed by western blot using streptavidin-HRP (Sigma) and ECL (GE Healthcare). The homologous competition experiment was performed as described above except that biotinylated toxin and BBMV were incubated in the presence of a 200-fold excess of the corresponding unlabeled Cry1A toxin.
BAC Library Screening and Sequencing
High-density filters for a BAC library of H. virescens  were obtained from the Texas A&M BAC Center (http://hbz7.tamu.edu), and high-density filters for a BAC library of H. subflexa were obtained from the Clemson University Genomics Institute (CUGI, http://www.genome.clemson.edu). These were screened by hybridization using a 32P-labelled 236-bp PCR product amplified from H. virescens larval midgut cDNA using primers Ha-ABC2-U14-F1 (5′ AACAA TCGTT ACCTG ATGGC GT) and Ha-ABC2-U14-R2 (5′ AGGAT TGGTA TCGAA AAATC TCATT AC) for the H. subflexa filters, and a 252-bp PCR product from the nachbar gene using primers Ha-bgi07733-F7 (5′ GAACT TGGGA CCTAC AGGTG GTAT) and Ha-bgi07733-R10 (5′ GCAGC ATTAC GGATA TTAAT TTCAA C). The H. virescens filters yielded two positive clones, and the H. subflexa filters 20 positive clones, which were obtained from CUGI and re-screened by PCR with primers Hs-BACscr02-F1 (5′-CACCG GCTCA ACACC ATCAT) and Hs-BACscr02-R2 (5′-GTCCT TGGCC ATGCT GTAGAA). Clone HS_Ba 89F08 was chosen and shot-gun sequenced at the Max Planck Institute for Chemical Ecology, Department of Entomology and deposited in GenBank as GQ332573. Primers designed from the H. subflexa sequence (Table S1) were used to amplify the ABCC2 gene in overlapping fragments from genomic DNA; sequence from YHD3 was deposited as GQ332572 and from YFO as GQ332571. Alignment of exon 2 of the YHD3 and YFO sequence revealed a 22-bp deletion in the former, causing a frameshift and resulting in a predicted stop codon after residue 99 (Figure S2).
Predicted Structure of ABCC2 Protein
Conceptual translations of the ABCC2 coding sequence from H. subflexa and the YFO allele of H. virescens were subjected to analysis for conserved domains by blastp to the Conserved Domain Database of NCBI (http://www.ncbi.nlm.nih.gov/cdd) and for transmembrane topology by the server (http://phobius.sbc.su.se/) for the prediction program Phobius . Potential glycosylation sites were screened for using the CBS Prediction Servers (http://www.cbs.dtu.dk/services/); none were found in the sequences examined. Conserved domains, predicted transmembrane domains and extracellular loops are depicted on a sequence alignment of ABCC2 from H. virescens, H. subflexa, B. mori, and homologues from Drosophila melanogaster and Mus musculus (Figure 4, Figure 5).
Estimation of Allele Frequencies from Archival DNA Samples
PCR with primers eU02-F1 and eiT02-R10 (Table S1) flanking the region containing the 22-bp deletion in the 6r allele were used to genotype individuals (Figure S1) used in previous mapping crosses and binding studies. YHD2 strain individuals from March 1993 are the adults used in crosses to map BtR-4 from which DNA was still available; no binding data are available from that generation. No DNA was available from individuals in the binding studies of Lee et al.  in 1995. All other samples come from binding studies of Jurat-Fuentes et al. in 2004  in which midguts were dissected from individual larvae in December 2002, the genotypes at BtR-4 were determined by PCR, and midguts from individuals with the same BtR-4 genotypes were pooled for binding analysis as shown in Figure 2 of that publication  (Table S3).
PCR assays for Hel-1 insertion in BtR-4, and exon 2 deletion in BtR-6.
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ABCC2 exon 2 sequence of susceptible YFO (GenBank GQ332571) and resistant YHD3 (GenBank GQ332572) strains showing the BtR-6 mutation.
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Frequency histograms of larval weight at 10 days for the three backcrosses and a susceptible strain.
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List of PCR and sequencing primers used.
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Predicted B. mori genes and H. armigera homologs in the nonrecombining region of B. mori Chr. 15 and H. virescens LG 2.
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Estimates of 6r allele frequency of BtR-6 from archival DNA samples.
(0.07 MB DOC)
We thank Joan LeGloahec, Chelsea Buscher, Regina Seibt, and Jan Scheirs for rearing insects; Domenica Schnabelrauch and Henriette Ringys-Beckstein for DNA sequencing; Bianca Ulitzsch for assistance in the binding studies; and Dr. Bruce Webb for making available EST sequences from H. virescens prior to deposition in GenBank.
Conceived and designed the experiments: LJG YP DGH. Performed the experiments: LJG YP. Analyzed the data: LJG YP DGH. Contributed reagents/materials/analysis tools: HV. Wrote the paper: LJG DGH.
- 1. James C (2008) Global Status of Commercialized Biotech/GM Crops: 2008. Ithaca, NY: International Service for the Acquisition of Agri-biotech Applications.
- 2. Tabashnik BE, Van Rensburg JBJ, Carrière Y (2009) Field-evolved insect resistance to Bt crops: Definition, theory, and data. J Econ Entomol 102: 2011–2025.
- 3. Pietrantonio PV, Gill SS (1996) Bacillus thuringiensis toxins: Action on the insect midgut. In: Lehane MJ, Billingsley PF, editors. Biology of the Insect Midgut. London: Chapman & Hall. pp. 345–372.
- 4. Soberón M, Gill SS, Bravo A (2009) Signaling versus punching hole: How do Bacillus thuringiensis toxins kill insect midgut cells? Cell Mol Life Sci 66: 1337–1349.
- 5. Zhang XB, Candas M, Griko NB, Taussig R, Bulla LA (2006) A mechanism of cell death involving an adenylyl cyclase/PKA signaling pathway is induced by the Cry1Ab toxin of Bacillus thuringiensis. Proc Natl Acad Sci USA 103: 9897–9902.
- 6. Pigott CR, Ellar DJ (2007) Role of receptors in Bacillus thuringiensis crystal toxin activity. Microbiol Mol Biol Rev 71: 255–281.
- 7. Ferré J, Van Rie J (2002) Biochemistry and genetics of insect resistance to Bacillus thuringiensis. Annu Rev Entomol 47: 501–533.
- 8. Tabashnik BE, Liu YB, Malvar T, Heckel DG, Masson L, et al. (1998) Insect resistance to Bacillus thuringiensis: uniform or diverse? Philos Trans R Soc Lond, Ser B: Biol Sci 353: 1751–1756.
- 9. Tabashnik BE (1994) Evolution of resistance to Bacillus thuringiensis. Annu Rev Entomol 39: 47–97.
- 10. Gould F, Anderson A, Reynolds A, Bumgarner L, Moar W (1995) Selection and genetic analysis of a Heliothis virescens (Lepidoptera: Noctuidae) strain with high levels of resistance to Bacillus thuringiensis toxins. J Econ Entomol 88: 1545–1559.
- 11. Akhurst RJ, James W, Bird LJ, Beard C (2003) Resistance to the Cry1Ac delta-endotoxin of Bacillus thuringiensis in the cotton bollworm, Helicoverpa armigera (Lepidoptera: Noctuidae). J Econ Entomol 96: 1290–1299.
- 12. González-Cabrera J, Escriche B, Tabashnik BE, Ferré J (2003) Binding of Bacillus thuringiensis toxins in resistant and susceptible strains of pink bollworm (Pectinophora gossypiella). Insect Biochem Mol Biol 33: 929–935.
- 13. Carrière Y, Crowder DW, Tabashnik BE (2010) Evolutionary ecology of insect adaptation to Bt crops. Evolutionary Applications 3: 561–573.
- 14. Storer NP, Babcock JM, Schlenz M, Meade T, Thompson GD, et al. (2010) Discovery and characterization of field resistance to Bt maize: Spodoptera frugiperda (Lepidoptera: Noctuidae) in Puerto Rico. J Econ Entomol 103: 1031–1038.
- 15. Xu XJ, Yu LY, Wu YD (2005) Disruption of a cadherin gene associated with resistance to Cry1Ac delta-endotoxin of Bacillus thuringiensis in Helicoverpa armigera. Appl Environ Microbiol 71: 948–954.
- 16. Morin S, Biggs RW, Sisterson MS, Shriver L, Ellers-Kirk C, et al. (2003) Three cadherin alleles associated with resistance to Bacillus thuringiensis in pink bollworm. Proc Natl Acad Sci USA 100: 5004–5009.
- 17. Gahan LJ, Gould F, Heckel DG (2001) Identification of a gene associated with Bt resistance in Heliothis virescens. Science 293: 857–860.
- 18. Lee MK, Rajamohan F, Gould F, Dean DH (1995) Resistance to Bacillus thuringiensis CryIA delta-endotoxins in a laboratory-selected Heliothis virescens strain is related to receptor alteration. Appl Environ Microbiol 61: 3836–3842.
- 19. Jurat-Fuentes JL, Gahan LJ, Gould FL, Heckel DG, Adang MJ (2004) The HevCaLP protein mediates binding specificity of the Cry1A class of Bacillus thuringiensis toxins in Heliothis virescens. Biochemistry 43: 14299–14305.
- 20. Tabashnik BE, Liu YB, Malvar T, Heckel DG, Masson L, et al. (1997) Global variation in the genetic and biochemical basis of diamondback moth resistance to Bacillus thuringiensis. Proc Natl Acad Sci USA 94: 12780–12785.
- 21. Baxter SW, Zhao JZ, Gahan LJ, Shelton AM, Tabashnik BE, et al. (2005) Novel genetic basis of field-evolved resistance to Bt toxins in Plutella xylostella. Insect Mol Biol 14: 327–334.
- 22. Bravo A, Gómez I, Conde J, Muñoz-Garay C, Sánchez J, et al. (2004) Oligomerization triggers binding of a Bacillus thuringiensis Cry1Ab pore-forming toxin to aminopeptidase N receptor leading to insertion into membrane microdomains. Biochim Biophys Acta Biomemb 1667: 38–46.
- 23. Heckel DG, Gahan LC, Gould F, Anderson A (1997) Identification of a linkage group with a major effect on resistance to Bacillus thuringiensis Cry1Ac endotoxin in the tobacco budworm (Lepidoptera: Noctuidae). J Econ Entomol 90: 75–86.
- 24. Jurat-Fuentes JL, Gould FL, Adang MJ (2002) Altered glycosylation of 63- and 68-kilodalton microvillar proteins in Heliothis virescens correlates with reduced Cry1 toxin binding, decreased pore formation, and increased resistance to Bacillus thuringiensis Cry1 toxins. Appl Environ Microbiol 68: 5711–5717.
- 25. Gahan LJ, Gould F, López JD, Micinski S, Heckel DG (2007) A polymerase chain reaction screen of field populations of Heliothis virescens for a retrotransposon insertion conferring resistance to Bacillus thuringiensis toxin. J Econ Entomol 100: 187–194.
- 26. Gahan LJ, Ma YT, Coble MLM, Gould F, Moar WJ, et al. (2005) Genetic basis of resistance to Cry1Ac and Cry2Aa in Heliothis virescens (Lepidoptera: Noctuidae). J Econ Entomol 98: 1357–1368.
- 27. Yasukochi Y, Ashakumary LA, Baba K, Yoshido A, Sahara K (2006) A second-generation integrated map of the silkworm reveals synteny and conserved gene order between lepidopteran insects. Genetics 173: 1319–1328.
- 28. Yoshido A, Bando H, Yasukochi Y, Sahara K (2005) The Bombyx mori karyotype and the assignment of linkage groups. Genetics 170: 675–685.
- 29. Aller SG, Yu J, Ward A, Weng Y, Chittaboina S, et al. (2009) Structure of P-glycoprotein reveals a molecular basis for poly-specific drug binding. Science 323: 1718–1722.
- 30. Bellier A, Chen CS, Kao CY, Cinar HN, Aroian RV (2009) Hypoxia and the hypoxic response pathway protect against pore-forming toxins in C. elegans. PLoS Pathog 5: e1000689. doi:10.1371/journal.ppat.1000689.
- 31. Chen CS, Bellier A, Kao CY, Yang YL, Chen HD, et al. (2010) WWP-1 Is a novel modulator of the DAF-2 insulin-like signaling network involved in pore-forming toxin cellular defenses in Caenorhabditis elegans. PLoS ONE 5: e9494. doi:10.1371/journal.pone.0009494.
- 32. Knowles BH, Ellar DJ (1987) Colloid-osmotic lysis is a general feature of the mechanism of action of Bacillus thuringiensis delta-endotoxins with different insect specificity. Biochim Biophys Acta 924: 509–518.
- 33. Schwartz JL, Garneau L, Savaria D, Masson L, Brousseau R, et al. (1993) Lepidoperan-specific crystal toxins from Bacillus thuringiensis form cation-selective and anion-selective channels in planar lipid bilayers. J Membr Biol 132: 53–62.
- 34. Slatin SL, Abrams CK, English L (1990) Delta-endotoxins form cation-selective channels in planar lipid bilayers. Biochem Biophys Res Commun 169: 765–772.
- 35. Bravo A, Gill SS, Soberón M (2007) Mode of action of Bacillus thuringiensis Cry and Cyt toxins and their potential for insect control. Toxicon 49: 423–435.
- 36. Chen J, Hua G, Jurat-Fuentes JL, Abdullah MA, Adang MJ (2007) Synergism of Bacillus thuringiensis toxins by a fragment of a toxin-binding cadherin. Proc Natl Acad Sci USA 104: 13901–13906.
- 37. Peng DH, Xu XH, Ye WX, Yu ZN, Sun M (2010) Helicoverpa armigera cadherin fragment enhances Cry1Ac insecticidal activity by facilitating toxin-oligomer formation. Appl Microbiol Biotechnol 85: 1033–1040.
- 38. Soberón M, Pardo-López L, López I, Gómez I, Tabashnik BE, et al. (2007) Engineering modified Bt toxins to counter insect resistance. Science 318: 1640–1642.
- 39. Muñoz-Garay C, Portugal L, Pardo-López L, Jiménez-Juárez N, Arenas I, et al. (2009) Characterization of the mechanism of action of the genetically modified Cry1AbMod toxin that is active against Cry1Ab-resistant insects. Biochim Biophys Acta 1788: 2229–2237.
- 40. Nagamatsu Y, Toda S, Koike T, Miyoshi Y, Shigematsu S, et al. (1998) Cloning, sequencing, and expression of the Bombyx mori receptor for Bacillus thuringiensis insecticidal CryIA(a) toxin. Biosci Biotechnol Biochem 62: 727–734.
- 41. Sangadala S, Walters FS, English LH, Adang MJ (1994) A mixture of Manduca sexta aminopeptidase and phosphatase enhances Bacillus thuringiensis insecticidal CryIA(c) toxin binding and (Rb+-K+)-Rb-86 efflux in vitro. J Biol Chem 269: 10088–10092.
- 42. Knight PJK, Crickmore N, Ellar DJ (1994) The receptor for Bacillus thuringiensis CrylA(c) delta-endotoxin in the brush border membrane of the lepidopteran Manduca sexta is aminopeptidase N. Mol Microbiol 11: 429–436.
- 43. Jurat-Fuentes JL, Adang MJ (2004) Characterization of a Cry1Ac-receptor alkaline phosphatase in susceptible and resistant Heliothis virescens larvae. Eur J Biochem 271: 3127–3135.
- 44. Arenas I, Bravo A, Soberón M, Gómez I (2010) Role of alkaline phosphatase from Manduca sexta in the mechanism of action of Bacillus thuringiensis Cry1Ab toxin. J Biol Chem 285: 12497–12503.
- 45. Pandian GN, Ishikawa T, Togashi M, Shitomi Y, Haginoya K, et al. (2008) Bombyx mori midgut membrane protein P252, which binds to Bacillus thuringiensis Cry1A, is a chlorophyllide-binding protein, and the resulting complex has antimicrobial activity. Appl Environ Microbiol 74: 1324–1331.
- 46. Valaitis AP, Jenkins JL, Lee MK, Dean DH, Garner KJ (2001) Isolation and partial characterization of gypsy moth BTR-270, an anionic brush border membrane glycoconjugate that binds Bacillus thuringiensis Cry1A toxins with high affinity. Arch Insect Biochem Physiol 46: 186–200.
- 47. Zhuang MB, Oltean DI, Gómez I, Pullikuth AK, Soberón M, et al. (2002) Heliothis virescens and Manduca sexta lipid rafts are involved in Cry1A toxin binding to the midgut epithelium and subsequent pore formation. J Biol Chem 277: 13863–13872.
- 48. Valaitis AP (2008) Bacillus thuringiensis pore-forming toxins trigger massive shedding of GPI-anchored aminopeptidase N from gypsy moth midgut epithelial cells. Insect Biochem Mol Biol 38: 611–618.
- 49. Masson L, Lun YJ, Mazza A, Brousseau R, Adang MJ (1995) The CryIA(c) receptor purified from Manduca sexta displays multiple specificities. J Biol Chem 270: 20309–20315.
- 50. Luo K, Sangadala S, Masson L, Mazza A, Brousseau R, et al. (1997) The Heliothis virescens 170kDa aminopeptidase functions as Receptor A by mediating specific Bacillus thuringiensis Cry1a delta-endotoxin binding and pore formation. Insect Biochem Mol Biol 27: 735–743.
- 51. Lee MK, Jenkins JL, You TH, Curtiss A, Son JJ, et al. (2001) Mutations at the arginine residues in alpha 8 loop of Bacillus thuringiensis delta-endotoxin Cry1Ac affect toxicity and binding to Manduca sexta and Lymantria dispar aminopeptidase N. FEBS Lett 497: 108–112.
- 52. Jenkins JL, Lee MK, Sangadala S, Adang MJ, Dean DH (1999) Binding of Bacillus thuringiensis Cry1Ac toxin to Manduca sexta aminopeptidase N receptor is not directly related to toxicity. FEBS Lett 462: 373–376.
- 53. Rajagopal R, Sivakumar S, Agrawal N, Malhotra P, Bhatnagar RK (2002) Silencing of midgut aminopeptidase N of Spodoptera litura by double-stranded RNA establishes its role as Bacillus thuringiensis toxin receptor. J Biol Chem 277: 46849–46851.
- 54. Yang YL, Zhu YC, Ottea J, Husseneder C, Leonard BR, et al. (2010) Molecular characterization and RNA interference of three midgut aminopeptidase N isozymes from Bacillus thuringiensis-susceptible and -resistant strains of sugarcane borer, Diatraea saccharalis. Insect Biochem Mol Biol 40: 592–603.
- 55. Candas M, Loseva O, Oppert B, Kosaraju P, Bulla LA (2003) Insect resistance to Bacillus thuringiensis - Alterations in the indianmeal moth larval gut proteome. Mol Cell Proteomics 2: 19–28.
- 56. Krishnamoorthy M, Jurat-Fuentes JL, McNall RJ, Andacht T, Adang MJ (2007) Identification of novel CrylAc binding proteins in midgut membranes from Heliothis virescens using proteomic analyses. Insect Biochem Mol Biol 37: 189–201.
- 57. Vadlamudi RK, Ji TH, Bulla LA (1993) A specific binding protein from Manduca sexta for the insecticidal toxin of Bacillus thuringiensis subsp Berliner. J Biol Chem 268: 12334–12340.
- 58. Nagamatsu Y, Toda S, Yamaguchi F, Ogo M, Kogure M, et al. (1998) Identification of Bombyx mori midgut receptor for Bacillus thuringiensis insecticidal CryIA(a) toxin. Biosci Biotechnol Biochem 62: 718–726.
- 59. Baxter SW, Zhao JZ, Shelton AM, Vogel H, Heckel DG (2008) Genetic mapping of Bt-toxin binding proteins in a Cry1A-toxin resistant strain of diamondback moth Plutella xylostella. Insect Biochem Mol Biol 38: 125–135.
- 60. Wang P, Zhao JZ, Rodrigo-Simón A, Kain W, Janmaat AF, et al. (2007) Mechanism of resistance to Bacillus thuringiensis toxin Cry1Ac in a greenhouse population of the cabbage looper, Trichoplusia ni. Appl Environ Microbiol 73: 1199–1207.
- 61. Mahon RJ, Olsen KM, Downes S, Addison S (2007) Frequency of alleles conferring resistance to the Bt toxins Cry1Ac and Cry2Ab in Australian populations of Helicoverpa armigera (Lepidoptera: noctuidae). J Econ Entomol 100: 1844–1853.
- 62. Downes S, Parker T, Mahon R (2010) Incipient resistance of Helicoverpa punctigera to the Cry2Ab Bt toxin in Bollgard II (R) cotton. PLoS ONE 5: e12567. doi:10.1371/journal.pone.0012567.
- 63. Caccia S, Hernández-Rodríguez CS, Mahon RJ, Downes S, James W, et al. (2010) Binding site alteration is responsible for field-isolated resistance to Bacillus thuringiensis Cry2A insecticidal proteins in two Helicoverpa species. PLoS ONE 5: e9975. doi:10.1371/journal.pone.0009975.
- 64. Olsen KM, Daly JC (2000) Plant-toxin interactions in transgenic Bt cotton and their effect on mortality of Helicoverpa armigera (Lepidoptera: Noctuidae). J Econ Entomol 93: 1293–1299.
- 65. Carrière Y, Ellers-Kirk C, Biggs R, Higginson DM, Dennehy TJ, et al. (2004) Effects of gossypol on fitness costs associated with resistance to Bt cotton in pink bollworm. J Econ Entomol 97: 1710–1718.
- 66. Joyner K, Gould F (1985) Developmental consequences of cannibalism in Heliothis zea (Lepidoptera: Noctuidae). Ann Entomol Soc Am 78: 24–28.
- 67. Robinson R (1971) Lepidoptera Genetics. Oxford: Pergamon Press.
- 68. Yasukochi Y (1998) A dense genetic map of the silkworm, Bombyx mori, covering all chromosomes based on 1018 molecular markers. Genetics 150: 1513–1525.
- 69. Mita K, Kasahara M, Sasaki S, Nagayasu Y, Yamada T, et al. (2004) The genome sequence of silkworm, Bombyx mori. DNA Res 11: 27–35.
- 70. Xia QY, Zhou ZY, Lu C, Cheng DJ, Dai FY, et al. (2004) A draft sequence for the genome of the domesticated silkworm (Bombyx mori). Science 306: 1937–1940.
- 71. Yamamoto K, Nohata J, Kadono-Okuda K, Narukawa J, Sasanuma M, et al. (2008) A BAC-based integrated linkage m0ap of the silkworm Bombyx mori. Genome Biology 9: R21.
- 72. Xia QY, Wang J, Zhou ZY, Li RQ, Fan W, et al. (2008) The genome of a lepidopteran model insect, the silkworm Bombyx mori. Insect Biochem Mol Biol 38: 1036–1045.
- 73. Wang J, Xia QY, He XM, Dai MT, Ruan J, et al. (2005) SilkDB: a knowledgebase for silkworm biology and genomics. Nucleic Acids Res 33: D399–D402.
- 74. Shimomura M, Minami H, Suetsugu Y, Ohyanagi H, Satoh C, et al. (2009) KAIKObase: An integrated silkworm genome database and data mining tool. BMC Genomics 10: 486.
- 75. Lander ES, Green P, Abrahamson J, Barlow A, Daly MJ, et al. (1987) MAPMAKER: An interactive computer package for constructing primary genetic linkage maps of experimental and natural populations. Genomics 1: 174–181.
- 76. Xia QY, Cheng DJ, Duan J, Wang GH, Cheng TC, et al. (2007) Microarray-based gene expression profiles in multiple tissues of the domesticated silkworm, Bombyx mori. Genome Biology 8: R162.
- 77. Wolfersberger MG, Luethy P, Maurer P, Prenti P, Sacchi VF, et al. (1987) Preparation and partial characterisation of amino acid transporting brush border membrane vesicles from the larval midgut of the cabbage butterfly (Pieris brassicae). Comp Biochem Physiol 86A: 301–308.
- 78. Lee MK, Milne RE, Ge AZ, Dean DH (1992) Location of a Bombyx mori receptor binding region on a Bacillus thuringiensis delta-endotoxin. J Biol Chem 267: 3115–3121.
- 79. Wu CC, Proestou D, Carter D, Nicholson E, Santos F, et al. (2009) Construction and sequence sampling of deep-coverage, large-insert BAC libraries for three model lepidopteran species. BMC Genomics 10: 283.
- 80. Kall L, Krogh A, Sonnhammer ELL (2007) Advantages of combined transmembrane topology and signal peptide prediction - the Phobius web server. Nucleic Acids Res 35: W429–W432.
- 81. Kall L, Krogh A, Sonnhammer ELL (2004) A combined transmembrane topology and signal peptide prediction method. J Mol Biol 338: 1027–1036.
- 82. Gaudet R, Wiley DC (2001) Structure of the ABC ATPase domain of human TAP1, the transporter associated with antigen processing. EMBO J 20: 4964–4972.