The Bacillus thuringiensis delta-endotoxin Cry1C as a potential bioinsecticide in plants

Plant Science 176 (2009) 315–324

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Plant Science journal homepage: www.elsevier.com/locate/plantsci

Review

The Bacillus thuringiensis delta-endotoxin Cry1C as a potential bioinsecticide in plants Dror Avisar 1, Haviva Eilenberg, Menachem Keller 2, Noam Reznik, Michal Segal 3, Baruch Sneh, Aviah Zilberstein * Department of Plant Sciences, The George S. Wise Faculty of Life Sciences, Tel Aviv University, Ramat Aviv 69978, Israel

A R T I C L E I N F O

A B S T R A C T

Article history: Received 5 October 2008 Received in revised form 11 December 2008 Accepted 11 December 2008 Available online 31 December 2008

The Cry1C group of Bacillus thuringiensis delta-endotoxins contains 10 highly homologous members of the Cry1Ca toxin sub-group and additional three members of the Cry1Cb sub-group that differ in domain III sequence. The Cry1Ca bioinsecticidal spectrum encompasses lepidopteran insects that are completely or partially tolerant to the current commercially used Bt crops. Plant-expressed Cry1Ca proteins successfully control specific lepidopteran pests, however, Bt crops expressing Cry1Ca have not been commercialized. This review summarizes the accumulating data in Cry1C research. Multiple sequence alignments of closely related Cry1Ca homologues show that the N-terminal half of the protein, comprising the ‘‘active toxin’’, is less conserved than the C-terminal part, which is involved in the assembly of the toxin-containing crystalline structure during the bacterial sporulation stage. Bioinformatics analyses predict high evolutionary diversity of amino acid residues in the regions identified as toxin–membrane interaction sites. All the three structural domains of Cry1Ca ‘‘active toxin’’ interact in vitro with membrane vesicles produced from epithelial cells of the larval gut. This multi-site-interaction depends on the normal assembly of membrane lipid raft domains, which is disturbed during cell division, when transient Cry1Ca insensitivity is observed. Cry1Ca interaction with the gut epithelial cells involves specific aminopeptidaseN receptors that differ from those described for other Cry1 toxins. The involvement of other membrane components in the interaction remains to be studied. Cry1A-resistant insect pests, such as the Cry1Actolerant mutants of diamondback moth, are sensitive to Cry1Ca, due to the involvement of different genetic loci. Hence pyramiding expression of Cry1Ca and other Cry toxins can broaden the bioinsecticidal spectrum of Bt crops and simultaneously delay the evolution of Cry-resistant insect populations. ß 2009 Elsevier Ireland Ltd. All rights reserved.

Keywords: Bt crops Cry1C Bacillus thuringiensis Delta-endotoxin Cry protein

Contents 1. 2. 3. 4. 5. 6. 7. 8. 9.

Introduction: Cry proteins as biopesticides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cry1C hypothesized structural features, based on amino acid homology and crystallographic analysis of Cry1Aa . . . . . . . . . . . . . . . . . . . . Identification of evolutionary conserved amino acid residues in Cry1Ca . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Receptors of Cry1 toxins and modes of interaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Specific interaction of Cry1Ca with putative cell membrane receptors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lipid rafts and Cry1Ca activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Overcoming possible selection for Cry-resistant insect populations in transgenic plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cry1C as a promising bioinsecticide . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

* Corresponding author. Tel.: +972 3 640 7410; fax: +972 3 640 6859. E-mail address: [email protected] (A. Zilberstein). 1 Present address: Plant Sciences Institute, Volcani Center, Bet Dagan 50250, Israel. 2 Present address: Hazera Genetics, Mivchor, Israel. 3 Present address: Department of Molecular Genetics, The Weizmann Institute of Science, Rehovot 76100, Israel. 0168-9452/$ – see front matter ß 2009 Elsevier Ireland Ltd. All rights reserved. doi:10.1016/j.plantsci.2008.12.010

315 316 316 317 319 320 321 321 322 322 322

1. Introduction: Cry proteins as biopesticides The rapidly increasing world population has imposed new demands on the agricultural community [1] urging the development of highly efficient pest control strategies to minimize crop

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losses worldwide [2,3]. The strict specificity of Bacillus thuringiensis (Bt) delta-endotoxins (Cry proteins) to certain insect species is considered as a major advantage for agricultural application because effects on non-target insects, including omnivorous predators [4]. Cry proteins are active against insects of the orders Lepidoptera, Coleoptera, Diptera, Hymenoptera as well as against nematodes [2,3]. More than 300 Cry proteins have been characterized and classified in 54 groups according to their amino acid sequences [5,6]. Bt delta-endotoxins have been used as biopesticides for the last halfcentury. These toxins provide an environmentally friendly alternative to about 5% of the 2 million tons of chemical insecticides. These chemicals often kill non-pathogenic insects and impose a high selection pressure on pest populations to evolve insecticideresistant lines. In addition, the misusage or careless usage of these chemicals causes 3 million human poisonings a year (http:// www.epa.gov/pesticides/biopesticides, last updated 12.9.2008). Presently Bt-corn and Bt-cotton are the only commercial crops expressing Bt toxins [7,8] and are grown over more than 160 million ha worldwide. During the first 5 years, since the introduction of Bt crops in 1995, Cry1Ab and Cry1Ac, designed to fit the plant transcription and translation machinery, were the predominate Cry toxins. Gradually, with accumulating information about the mode of action of Bt toxins and their epithelial gut membrane receptors, the deployment of pyramided advanced varieties, expressing two toxins with different receptor recognition modes, has been introduced (http://www.epa.gov/pesticides/biopesticides/pips/pip_list.htm, updated 24.7.2008). Recent reviews have profoundly dealt with various Bacillusderived toxins, including Vip toxins that are produced and secreted during the vegetative growth stage [9]. They describe the threedimensional structure analyses of Cry proteins and the proposed modes of action [10,11]. Others have estimated the possible evolution of Cry toxin-resistant insect populations due to the selective pressure imposed by the gradually increasing usage of Bt crops and discussed strategies to overcome this threat [3,7,12]. There are many insect species that are poorly affected by the commercialized Bt strains, as well as many Cry proteins, including those of the Cry1C group, whose specific interaction with the target cells of the larval gut epithelium remains to be unraveled. This review is focused on the Cry1C group of toxins, summarizing the presently known information regarding their structure, interaction with target cells and their possible application as toxins in Bt crops. Recently the pyramiding strategy has been strongly recommended, urging the generation and deployment of Bt crops expressing two different Cry toxins to minimize the evolution of resistance among insect populations exposed to Bt crops [7,13]. The pyramided toxins will only be effective if there are overlapping spectra of activity. Thus, defining Cry proteins that can act as an adjuvant to those that are currently in use may broaden the pyramiding options. 2. Cry1C hypothesized structural features, based on amino acid homology and crystallographic analysis of Cry1Aa The group of Cry1C proteins is divided, according to amino acid sequence homology, into two sub-groups: Cry1Ca and Cry1Cb. Currently, the Cry1Ca sub-group includes 10 highly homologous proteins that share 99% amino acid sequence identity with only very limited differences, as indicated in Fig. 1B. The three proteins comprising the Cry1Cb sub-group share only 84% identity with the proteins of the Cry1Ca group. The most prominent sequence variability between the two sub-groups is found in domain III of the active toxin (the domains are defined in Fig. 1A) that share only 40% amino acid sequence homology. The genes encoding Cry1Ca proteins were isolated mostly from Bt entomocidus 60.5, HD110 and Bt aizawai 729 and some from other strains such as K26-21, AF-

2 and G10-01A. Cry1Cb genes were isolated from Bt galleriae and Bt 087 strains (see the full list in [14]). Cry1C proteins belong to the family of three-domain-Cry toxins containing a N-terminal toxic half with three-domain structure (denoted herein as the ‘‘active toxin’’) and a C-terminal half, responsible for the assembly of the Cry proteins in a parasporal crystalline structure (inclusion) during sporulation, which terminates the Bt life cycle [5,15,16]. Upon ingestion of the spore inclusions by lepidopteran larvae, the C-terminal part (ca. 60– 70 kDa in Cry1 proteins) of the toxin, together with the first Nterminal 26–29 amino acid residues, are cleaved by trypsin-like gut proteases. The remaining part of the ‘‘active toxin’’ becomes suitably folded, allowing further oligomer formation and interaction with the apical cell membrane of the gut epithelium [2,17]. Cry1Ca active toxin shares 48% amino acid identity with the corresponding part of Cry1Aa, whose three dimensional structure has been analyzed [18]. Despite the amino acid sequence diversity, the three-domain structure of the active toxin is shared by Cry1Aa, Cry2Aa, Cry3Aa, Cry3Bb, Cry4Aa and Cry4Bb crystals [2,11,19] and by many other Cry proteins that belong to the three-domain toxin family. Hence, the conserved structural topology of Cry1Aa ‘‘active toxin’’, elucidated by crystallography, represents the general structure of Cry1C ‘‘active toxins’’. The amino acid sequence alignment of Cry1Ca5 and Cry1Aa, where domain borders and structural features are indicated according to the Cry1Aa analysis [18], are shown in Fig. 1A. Most members of the three-domain toxin family have a domain I composed of seven a helices (whose location is marked above Cry1Aa sequence in Fig. 1A). Domain I is involved in the formation of ion channels that are poorly selective to either cations or anions, depending on pH [20]. The intervening loops linking the b-sheets of domain II- and III are involved in very specific interaction/s with receptors present on the cell membrane of the gut epithelial cells in the insect larvae [11]. These interactions are a prerequisite for further conformational changes that lead to the insertion of a4 and a5 helices of domain I, from four toxin molecules, into the cell membrane, resulting in the formation of trans-membrane pores [17,21–23]. These pores act as non-selective ion channels causing cell swelling, lysis and death [2,11,24,25]. A recent revolutionary study shows that the whole active toxin, including all the three domains with the exception of domain I-a1 helix, is inserted into the cell membrane [26]. This study measured the interaction in vitro between different cysteine mutants of Cry1Aa- or Cry1Ab-active toxin and artificial phospholipid vesicles or brushborder membrane vesicles, derived from Manduca sexta larval midgut. These findings doubt the widely accepted models that proposed the insertion of certain a helices of domain I into the cell membrane and the spreading of domains II and III on the membrane surface [17,21–23]. Additional novel data, derived from unique 3D crystallography of the membrane-associated Cry4Ba complex, suggest the presence of two different trimeric conformations of Cry4Ba oligomers [27], rather than the tetrameric structures previously proposed and observed [17,21–23]. Interestingly, trimers of Cry4Ba were identified earlier in the rhombohedral crystals used to determine its 3D structure [17,21–23]. Thus, the whole concept of Cry toxin assembly in the cell membrane as well as the role of the three domains in this assembly require further verification. Recent Atomic Force Microscopy (AFM) imaging of apical membranes derived from M. sexta larval midgut epithelial cells has provided the first visual demonstration of specific toxin-membrane structures formed by the interaction of Cry1Aa, Cry1Ac and Cry1C with membrane vesicles [28]. 3. Identification of evolutionary conserved amino acid residues in Cry1Ca Cry1Ca toxins possess a quite different insecticidal spectrum when compared to other Cry proteins currently used in Bt crops

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(Table 1). Thus, it is possible that Cry1Ca differential interaction with the cell membrane of the midgut epithelium results from the less conserved regions that are more specific to the Cry1Ca active toxins. Although many dendrograms based on amino acid sequence homology of the members of the three-domain toxin family have already been published (e.g. [29,30]), the estimation of the evolutionary conserved amino acid residues has become more feasible after the development of the Conseq approach. This program calculates the substitution rate of each residue using the maximum likelihood paradigm [29,30]. As an initial step, we performed multiple sequence alignment of the ‘‘active toxin’’ part of 60 Cry proteins, all showing more than 30% identity with Cry1Ca5 sequence. The dendrogram shown in Fig. 1C is based on the comparison of the 32 ‘‘active toxin’’ amino acid sequences that showed the highest homology to Cry1Ca. From this non-rooted dendrogram and others (obtained by using ClastalW and T-Coffee that are based on amino acid-pairwise alignment and gap penalty), we could conclude that the Cry1J group is most closely related to the Cry1C group. Similar comparison of the C-terminal half of 25 out of these 32 proteins that possess this part (starting from residue 611 in Cry1Aa, Fig. 1D) showed that the C-terminal half is more conserved. Very small distances are seen between most of the toxins except of the distal Cry1B and Cry1K groups in the deduced dendrogram (Fig. 1D). This high homology suggests an important functional role of the Cterminal part. The most conserved element in this part is the arrangement of the cysteine residues, allowing the formation of symmetrical intermolecular disulphide bonds between different toxin molecules comprising the crystal [31,32]. The alkaline environment of the gut enables the cleavage of these S–S bonds and the solubilization of the crystal to individual protoxins after ingestion by lepidopteran larvae [31,32]. All the sequences included in the calculation of Fig. 1C dendrogram were used in the Conseq analysis [29] to elucidate the evolutionary conserved amino acid residues in the ‘‘active toxin’’. The results are displayed below the Cry1Aa–Cry1Ca5 alignment, demonstrating the high and low evolutionary conserved amino acid residues in the Cry1Ca5 sequence (Fig. 1A). Twelve evolutionary conserved residues were identified among the 28 amino acids that comprise the N-terminal part of the ‘‘active toxin’’. This part is cleaved at the earliest stage of the Cry1 toxin processing in the gut. The cleavage site (Arg28) is also conserved. The five relatively conserved regions, already identified before [32] (underlined in Fig. 1A) and domain I-a2b helix, contain many evolutionary conserved amino acid residues. The function of domain I-a2b helix remains to be elucidated. The most conserved component in domain I is a5 helix (Fig. 1A). This a-helix and the adjacent a4 helix of four Cry protein molecules were together proposed to form the amphipathic membrane structure of the nonselective ion channel [21,24]. However, as shown in Fig. 1A, the a4 counterpart and the intervening loop are less conserved. Another highly conserved region encompasses the C-terminal part of domain I-a7 helix and b1, which is the first b strand in sheet 3 of domain II. This region forms an essential linkage between domain I and II in all Cry proteins whose 3D structure has been determined [18,33]. A lower number of conserved amino acid residues is found in the loop that links b9 and b10 in domain III that interacts with domain I-a1 helix, despite its important role in the inter-domain interaction. This interaction may only hold during the first stage of the ‘‘active toxin’’ processing, because the a1 helix of Cry1A toxins is cleaved after the interaction with the cadherin-like-protein receptor. This is the second cleavage in the intoxication process and occurs between Val50 and Pro51 [2,34,35], which are also conserved residues (Fig. 1A). The significance of the highly conserved region identified downstream to domain III (Fig. 1A), including the C-terminal

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cleavage site that terminates the ‘‘active toxin’’ [36] and the very conserved C-terminal half, remains to be studied. The Conseq analysis suggests that there are evolutionary diverse motifs along all the three structural domains. These motifs are not limited to the loops of domain II and III containing the putative Cry protein recognition sites that specify the interaction with the cell membrane [11]. Thus, the dendrograms and Conseq calculations show two opposite evolutionary trends: high conservation of the C-terminal part and high amino acid diversity in the ‘‘active toxin’’. The first trend emphasizes the importance of maintaining the conserved ability of forming the spore crystalline structure that is only dissolved under alkaline conditions in the larval midgut of certain insects. Conversely, a second trend suggests that the ‘‘active toxin’’ part has been subjected to a completely different selection pressure. It might be speculated that frequent mutations and transpositions in the insect genome could change critical sites of the gut receptors and eventually enforce the evolution of correlative changes in single amino acid residues in the ‘‘active toxin’’, to allow further recognition of the rapidly mutated receptors. 4. Receptors of Cry1 toxins and modes of interaction The intoxication process of Cry1A proteins (Cry1Aa, Cry1Ab and Cry1Ac) is relatively well understood and begins with a specific interaction of the toxin with membrane receptors [2,11,37,38]. Cadherin-like-proteins (CLPs) were identified as the non-lipid-raft initial Cry1A interacting proteins [11], whereas aminopeptidase N (APN) and alkaline phosphatase (ALP) have been described as glycosylphosphatidylinositol (GPI)-Cry protein receptors, anchored to membrane lipid rafts. A sequential process of toxin-cell membrane interaction has been proposed, based on the initial interaction of a specific cadherin-like-protein with the monomeric version of Cry1A active toxins. This interaction induces a conformational change in the toxin that facilitates the cleavage of the N-terminal a1 helix by the membrane-bound proteases of the insect gut [34]. The toxin is then oligomerized to a tetrameric pre-pore structure that preferentially binds to aminopeptidase N as a result of an increase in binding affinity. Aminopeptidase N and/or alkaline phosphatase then direct/s the pre-pore tetramer to the lipid rafts, where membrane insertion occurs [2,17,37,39]. This possible mode of pore formation has been supported by the findings that modified Cry1A toxins, lacking the first a helix of domain I, do not require interaction with the specific cadherin-like-protein to undergo oligomerization [35,38]. These findings suggest that such modified Bt toxins can be applied in the future to overcome selective evolution of insect resistance resulting from mutations in the cadherin-like-protein [40]. Still, the expression and insect toxicity of the Domain I-a1-deleted-Cry1Ab and Cry1Ac in plants remain to be demonstrated. Another suggested mode of Cry1Ab toxicity is based on a Gprotein-induced signaling through cAMP formation and activation of a protein kinase A, a process initiated by the interaction of Cry1Ab with the membrane proximal repeat of the extra-cellular moiety of the cadherin-like-protein [41,42]. Combined effects of osmotic lysis by membrane pore formation and cell signaling were suggested to take place in Heliothis virescens during the intoxication process [37,43]. In this model, monomers of Cry1Ac-‘‘active toxin’’ bind to the binding domain of the cadherin-like-protein, leading to activation of an intracellular signaling pathway that is responsible for the toxic effect. Certain Cry protein interactions with the membrane involve binding to specific insect/nematode glycolipids whose glycosylation is central to the interaction process. Nematode mutants defective in defined lipid glycosylation steps were insensitive to

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Fig. 1. Cry1C group summary: homology with other Cry proteins, estimation of structural domain components and bioinformatics prediction of evolutionary conserved amino acid residues shared by Cry1Ca and other homologous Cry proteins. (A) Demonstration of Cry1Ca5 putative structural components by alignment to Cry1Aa whose threedimensional structure analysis was published (PDB 1D-1ciy [18]). Cry1Ca evolutionary conserved or diverse amino acid residues are listed below the two-protein sequencealignment. Prediction of evolutionary conservation of residues is based on the calculated substitution rate of each residue using the maximum likelihood paradigm (ConSeq server http://conseq.tau.ac.il [29,30]). The analysis includes the amino acid sequence alignment of the ‘‘active toxin’’ of 12 representatives showing the highest homology to Cry1Ca5 in panel C. The evolutionary score scale is shown below, with dark green as the variable and dark violet as the highly conserved residues, respectively. Vertical blue lines show domain borders and red letters mark the essential cleavage sites, according to the two-step model of toxin–membrane interaction [32]. Cry1Ca domain III end is

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Cry toxins [19]. Notably, this lipid glycosylation process is specific to insect and nematode lipids and does not occur in vertebrates. Despite many intensive studies of Cry toxin mode of action, the coordinate action of Cry protein receptors and other membrane proteins, involved in the intoxication process, still requires further clarification (reviewed by [11]). Another unsolved question is the commonality of Cry-protein-receptors. Currently, only the receptors for Cry1A proteins have been identified, whereas less is known about receptors of other Cry toxins. 5. Specific interaction of Cry1Ca with putative cell membrane receptors Putative aminopeptidase N receptors for Cry1Ca have been detected in Spodoptera litura [44–46], M. sexta larvae [47], Spodoptera exigua [48] and Spodoptera frugiperda brush-border membrane vesicles [49,50]. There are five classes of membraneanchored aminopeptidase N proteins in insects, each sharing at least 56% sequence identity [11]. The involvement of aminopeptidase N in Cry1Ca binding was first shown in M. sexta midgut where Cry1Ca and Cry1Ac recognized functionally related, but structurally distinct, 106 kDa and 115 kDa isoforms of aminopeptidase N, respectively [47]. As yet, the gene encoding the M. sexta 106 kDa aminopeptidase N has not been identified. The role of Class 4aminopeptidase N as specific Cry1Ca receptors was elegantly shown in S. litura larvae by RNA interference silencing in vivo. An intra-hemocoelical injection of S. litura aminopeptidase N dsRNA into early 5th instar S. litura larvae reduced endogenous aminopeptidase N expression and decreased sensitivity to Cry1Ca toxin [46]. Loss of Cry1Ca binding was also observed when Sf21 cells expressing S. litura aminopeptidase N were transfected with aminopeptidase N siRNA [45]. Binding of Cry1Ca to S. litura aminopeptidase N, expressed in Sf21 cells, was shown by ligand blot interaction and immunoprecipitation [44]. This further confirmed the role of aminopeptidase N as a Cry1Ca receptor in S. litura. A Cry1Ca-resistant colony of S. exigua lacked the expression of a single Class 1-aminopeptidase N out of four defined aminopeptidase N genes that are expressed in susceptible larvae [48]. As yet, no direct evidence of Cry1Ca interaction with cadherin-like-proteins has been reported. Three different approaches have been undertaken to identify the Cry1Ca sites that interact with the cell membrane: measuring interaction of separate domains with membrane vesicles derived from gut epithelium; domain shuffling between different ‘‘active toxin’’s; and generation of single amino acid substitutions by sitedirected mutagenesis. All the three Cry1Ca structural domains interacted with Spodoptera littoralis brush-border membrane vesicles in binding assays performed in vitro [51]. Considering the findings that a1helix of Cry1A-domain I interacts with a cadherin-like receptor at the initial step of the binding process [35], the observed binding of Cry1Ca-domain I to BBMV [51] might reflect the initial interaction of domain I-a1 with, as yet, an undefined cadherin-like-protein.

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This initial interaction promotes Cry1A-toxin oligomerization [35]. A mutation in Gln374 residue of Cry1Ca-domain II (Fig. 1A) prevents toxin oligomerization [52], suggesting that Cry1Ca forms oligomers and that domain II is also involved in the initial interaction with the cell membrane and the subsequent oligomerization. Constructing various hybrids of active toxins showed the importance of Cry1Ca-domain III. For example, the hybrid containing Cry1Ea domain I and II and Cry1C domain III, formed by homologous recombination in E. coli (denoted as G27), was active against S. exigua, in contrast to the parental Cry1Ea [53]. Further construction of a hybrid toxin, comprising Cry1Ab domains I and II and Cry1Ca domain III, was more toxic to S. exigua than the parental Cry1Ab and Cry1Ca [54]. When domain III of Cry1Ab, Cry1Ac, Cry1Ba, Cry1Ea or Cry1Fa was exchanged with that of Cry1Ca, the hybrids became more active against S. exigua than the parental toxins [55]. Only the Cry1Da–Cry1Ca hybrid was less active against S. exigua than the parental proteins. The effect of Cry1Ca or H04 (a fusion between domains I and II of Cry1Ab with domain III of Cry1Ca) was tested in transgenic shallot (Allium cepa L.) plants harbouring H04 or Cry1Ca ‘‘active toxin’’s under the control of a Chrysanthemum RUBISCO small subunit promoter [56]. Both genes conferred resistance to S. exigua in transgenic shallots. Intra-domain III hybrids between Cry1E and G27 enabled identification of amino acid groups, as well as some individual Cry1Ca amino acid residues, which are involved in specific interaction in S. exigua [57]. The Trp479 (Trp476 in [57]) and the region between Thr537 to Val544 (Thr 534 to Val541 in [57]), in Cry1Ca domain III (Fig. 1A) are important for toxicity to S. exigua and increased G27 toxicity against M. sexta. Conseq analysis identified these residues as evolutionary diverse amino acids (Fig. 1A), supporting the speculation that the recently evolved amino acid residues/regions in Cry1Ca domain II and III play an important role in the specific interaction with each insect. This speculation is further supported by the evidence that mutations in the nonevolutionary conserved Cry1Ca-Gln374 (domain II-loop 2, Fig. 1A), prevented Cry1Ca oligomerization in the presence of S. exigua brush-border membrane vesicles (BBMV), but did not affect such oligomerization in the presence of M. sexta BBMV [52]. Sitedirected mutagenesis in loop 2 (between b6 and b7r) and loop 3 (between b10 and b11) of Cry1Ca domain II, differentially reduced toxicity against S. littoralis and Aedes aegypti larvae [58]. Both loops contain evolutionary diverse amino acid residues (Fig. 1A). Taken together, these observations indicate that both Cry1Cadomain II and III are involved in specific interactions with the cell membrane of the gut epithelial cells. We have demonstrated that both Cry1Ca interaction with the cell membrane and its toxicity require the presence of lipid rafts in the cell membrane [59]. Thus it may be assumed that at least a part of Cry1C-multi-stage interaction occurs in the lipid rafts. We have recently detected sequential binding of Cry1Ca to the cell membrane of Sf9 cells (derived from S. frugiperda), first to the non-lipid-raft membrane-soluble fraction and then to lipid rafts. After exposing Sf9 cells to low levels (1 mg/ml) of Cry1C, the toxin

marked in brown. The conserved amino acid blocks are underlined according to [32]. The amino acid sequence of Cry1Aa structural components are colored as follows: a helices in yellow, b sheets in gray and loops in magenta. The b sheets in domain II-sheet 1 are indicated in blue letters, in sheet 2—in light blue letters and in sheet 3 in orange letters. The b sheets of domain III are in black letters. There is an unnumbered b sheet between domains I and II, which is also marked in black letters. Green letters indicate salt bridges or formation of hydrogen bonds according to Cry1Aa topology, not always present in Cry1Ca. The amino acid residues of a5 helix, that are uncharged despite their polar side groups, are also indicated in green. Dark green background and white letters indicate residues required for toxicity against Spodoptera exigua, located in the outer b sheet of domain III [55,57]. Boxed regions indicate the interaction sites of a7 with b1b-a8 and the link between a1 and the intervening loop between b9 and b10. (B) Amino acid-homology dendrogram of all members of the Cry1C group. The length of each protein and the number of amino acid mis-matches (mism) are indicated in comparison to Cry1Ca5 sequence. (C) Amino acid-homology dendrogram of close homologues of the Cry1Ca5 ‘‘active toxin’’. BLASTp analysis was used to compare the amino acid sequence of Cry1Ca5 ‘‘active toxin’’ with other Cry protein ‘‘active toxin’’s. Proteins sharing above 30% identity with Cry1Ca5 were selected for further analyses. Multiple sequence alignments, using T-Coffee, MUSCLE and Clustal W2 were performed. Thereafter, 32 closely related sequences were chosen for constructing the dendrogram by MEGA 4 software. The Cry protein accession numbers can be identified in the NCBI protein database, according to the names mentioned in the dendrogram. AAD46139 is a yet noncharacterized Cry protein sequence. The accession number of Cry1-A32 (denoted as Cry1A32) is AAX86871. (D) Dendrogram of the C-terminal part, showing the homology between Cry1Ca5 and 25 closely related homologues. The multiple alignments were performed using proteins shown in C that possess the C-terminal part.

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Table 1 Insecticidal spectrum of Cry1Ca compared with other relatively efficient Cry proteins. Species

Common name

Effective Cry protein

ED50a

Ref.

Plant hosts

Spodoptera exigua

Beet armyworm

Cry1Ca

90

[93]

Alfalfa, corn, cotton, peanut, safflower, sorghum, soybean, sugarbeet, tobacco, turnip, asparagus, bean, beet, broccoli, cabbage, cauliflower, celery, chickpea, corn, cowpea, eggplant, lettuce, onion, pea, pepper, potato, radish, spinach, sweet potato, tomato http://creatures.ifas.ufl.edu/veg/leaf/beet_armyworm.htm

Cry1Da Cry1Fa Cry9Ca1

9.5 104 133

[93] [93] [94]

Cry1Ca4

93

[95]

Cry1Ea1 Cry9Ca1

62 65.5

[95] [94]

Cry1Ca1

31

[49]

Cry1Da1 Cry1Fa1

77 109

[49] [50]

Cry1Ca4

22 (L1)

[32]

Cry1Aa1 Cry1Ab5 Cry9Ca1

165 (L1) 162(L1) 79

[32] [32] [94]

Cry1C

45.3 ng/larva (L2)

[96]

Cry1Ab

29.8 ng/larva (L2)

[96]

Spodoptera littoralis

Spodoptera frugiperda

Mamestra brassicae

Mamestra configurata

Egyptian cotton leafworm

Fall armyworm

Cabbage moth

Bertha armyworm

Manduca sexta

Tobacco hornworm

Cry1Ca4 Cry1Aa1 Cry1Ab2 Cry2Ac1

74 5 7.5 12–16

[95] [95] [97] [98]

Bombyx mori

Silkworm

Cry1Ca2 Cry1Aa1 Cry1Aa6 Cry1Da1

30 ng/larva (L4) 2 ng/larva (L4) 3 ng/larva (L4) 6.1 ng/larva

[99] [100] [99] [99]

Plutella xylostella

Diamond back moth

Cry1Ca

5 (L3)

[101]

Cry1Ac Cry1Ac1 Cry1Ba2

0.9 (L3) 0.4 (L3) 1.2 (L3)

[101] [102] [90]

Cry1Ca

12

[103]

Cry1Ac Cry1Ab Cry2Ac1

1.1 3.4 2–5

[103] [103] [98]

Trichoplusia ni

Cabbage looper

Alfalfa, apples, cotton, flax, groundnuts, jute, grapes, maize, rice, soybean, tea, tobacco, cucurbit vegetables, potato, sweet potato http://extension.entm.purdue.edu/CAPS/pestInfo/egyptLeafworm.htm

Alfalfa, barley, Bermuda grass, buckwheat, cotton, clover, corn, oat, millet, peanut, rice, ryegrass, sorghum, sugarbeet, sudangrass, soybean, sugarcane, timothy, tobacco, wheat http://creatures.ifas.ufl.edu/field/fall_armyworm.htm

Cruciferaceae (cabbage, turnip), tobacco, red beet, flax, lettuce, chicory http://www.inra.fr/internet/Produits/HYPPZ/RAVAGEUR/ 6mambra.htm

Canola, rapeseed, mustard, alfalfa, flax, field peas, potato http://www.gov.mb.ca/agriculture/crops/insects/fad03s00.html

Tobacco, tomato http://creatures.ifas.ufl.edu/field/hornworm.htm

Cruciferae plants e.g. canola, mustard, broccoli, cabbage, cauliflower http://creatures.ifas.ufl.edu/veg/leaf/diamondback_moth.htm

Broccoli, cabbage, cauliflower, collards, kale, mustard, radish, rutabaga, turnip, watercress, beet, cantaloupe, celery, cucumber, lima bean, lettuce, parsnip, pea, pepper, potato, snap bean, spinach, squash, sweet potato, tomato, watermelon, cotton and tobacco http://creatures.ifas.ufl.edu/veg/leaf/cabbage_looper.htm

Instar age is neonate if not otherwise stated (L1, L2, L3 or L4—larval stage). Table based on the specificity database of Natural Resources Canada (http://www.glfc.forestry.ca/ bacillus/BtSearch.cfm). Additional insects sensitive to Cry1Ca: Pieris rapa (cabbage moth) (according to unpublished results (Keller M.)). Helicoverpa armigera (cotton bollworm) (according to unpublished results (Keller M.)). Phthorimaea operculella (potato tuber moth) (according to unpublished results (Keller M.)). Chrysodeixis chalcytes (tomato looper) (according to unpublished results (Keller M.)). Pectinophora gossypiella (pink bollworm) (Tabashnik et al. [104]). Heliothis virescens (tobacco budworm) (Hofte et al. [32]). a ED50 units are ng/cm2 if not otherwise stated.

could be located after 5 min only in the soluble membrane fraction, while after 15 min it was present in both soluble fraction and membrane lipid rafts (Segal, Eilenberg, Avisar and Zilberstein, unpublished data). S. frugiperda larvae change their expression of many midgut epithelial genes within 15 min after Cry1Ca challenge [60]. Hence, this gradual Cry1Ca interaction might resemble the sequential interaction of Cry1Ab with gut epithelium-derived vesicles (BBMV) observed in M. sexta and H. virescens [11,61].

6. Lipid rafts and Cry1Ca activity Membrane lipid rafts are involved in protein sorting and initiation of signaling pathways in eukaryotic cells. They also serve as the gate for various pathogen attacks and toxin interaction and intake [62,63]. Membrane lipid rafts are floating microdomains enriched in cholesterol, sphingolipids as well as glycosylphosphatidylinositol (GPI)-anchored proteins that remain insoluble after Triton X-100 detergent treatment in the cold. Sphingolipids

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and glycosphigolipids are synthesized in the Golgi apparatus. They assemble into lipid raft microdomains in the trans-Golgi network and are then transported to the plasma membrane via the endosomal vesicle pathway [64–66]. Proteins located in lipid rafts are also detergent-protected and can be separated from the other soluble membrane proteins by sucrose gradient centrifugation. Both insect aminopeptidase N and alkaline phosphate (ALP), that serve as receptors for the Cry1A toxins in gut epithelial cells of different lepidopteran larvae [2,37,61,67,68], are external GPIproteins anchored to the outer side of the membrane lipid rafts. Lipid rafts are essential for Cry1A toxin activity and are probably involved in the formation of lethal membrane pores by Cry1A oligomers [61]. Cry1Ca is toxic to Sf9 cells after binding to lipid rafts. The use of Sf9 cell line as a model system for studying Cry1Ca toxicity is based on their Cry1Ca-specific sensitivity [69–72]. These cells are very amenable for identifying genes and processes involved in the cellular response to Cry1Ca [70,73,74]. For example, changes in Sf9 cell membrane permeability and cellular shape were used for studying Cry1Ca-dependent pore formation [71,72,75,76]. The estimated LC50 value of Cry1Ca in Sf9 cells was 138 ng/ml medium [59]. Despite this high susceptibility, dividing cells showed a surprising transient lack of sensitivity to Cry1C and could complete their mitotic phase, while the neighboring nondividing cells swelled and died in response to Cry1Ca. Correlatively, when the cells were chemically arrested at the G2/M phase, a drastic reduction in Cry1Ca susceptibility (LC50 increased to about 715 ng/ml) and 10-fold decline in toxin binding were observed, in comparison to complete mortality of non-arrested cells of the control treatment. The observed Cry1Ca tolerance was transient and occurred only during G2/M arrest. Cells gradually regained Cry1C sensitivity upon release from G2/M blockage, interacted with Cry1Ca and died [59]. No lipid rafts were detected in the plasma membrane of the cells arrested in G2/M-phase [59], indicating that the intact organization of lipid rafts might be essential for Cry1Ca toxicity. In mammalian cells as well as in Drosophila S2 cells, Golgi stacks undergo complete fragmentation while entering G2 phase and this collapsing stage serves as a premitosis checkpoint [77,78]. The disruption of lipid raft organization observed in G2/M phase in lepidopteran cells is probably an outcome of Golgi stack fragmentation. Further elucidation of lipid raft components involved in the interaction with Cry proteins is required for comprehensive understanding of the Cry protein ingestion and will fill in the gaps that remain unsolved.

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[48], confirming the involvement of Cry1Ca-specific amonopeptidase N. Furthermore, silencing of Class 4-aminopeptidase N expression by RNA interference also caused reduced sensitivity to Cry1Ca in S. litura [46]. Resistance to Cry1Ca and Cry1Ac was genetically analyzed in a field collected colony of Plutella xylostella [83,84] and further confirmed by AFLP markers. A single linkage group was associated with Cry1Ac resistance whereas two other linkage groups are positively associated with Cry1Ca resistance, indicating that the resistance to the two different Cry toxins is conferred by independent genes localized to separate loci [85]. The introduction into crops of multiple (‘‘pyramided’’) Cry genes, including the Bt vip3A gene, has recently become the strategy of choice for minimizing Cry-specific selection pressure [13,86]. This approach relies on the very low probability of multiple mutation occurrence that would cause a general loss of sensitivity. Therefore, finding additional Bt toxins that interact with different receptors in the epithelial cells of the insect gut of the same target insect species is of primary importance. Bollguard II cotton expressing both Cry1Ac and Cry2Ab was first introduced in 1999. These plants were more toxic to bollworms (H. zea) and also to S. frugiperda and S. exigua as secondary pests, in comparison to cotton plants expressing Cry1Ac alone [87]. Recently ‘‘pyramided’’ Indian mustard (Brassica juncea) plants co-expressing Cry1Ac and Cry1Ca were shown to effectively control Cry1A- or Cry1Ca-resistant larvae of P. xylostella (diamondback moth) [88]. However, it should be emphasized that the concurrent use of transgenic plants in adjacent fields, expressing a single and two Bt genes, including an overlapped one, was shown to speed up P. xylostella resistance to pyramided plants [13]. Thus, although the genetic reason for this elevated resistance still remains to be elucidated, such concurrent use should be avoided. Therefore, other toxins, such as members of Cry1Ca group that probably interact with a different aminopeptidase N [11,44,45] and other as yet unknown membrane proteins, are preferred candidates. The recently proposed Cry1AbMod and Cry1AcMod modified ‘‘active toxin’’s [35], both lacking the first a helix of domain I that interacts with the insect cadherin-like-protein receptor, would only partially solve the problem of mutagenized CLP that does not interact with Cry1A toxins [89]. These two toxins still interact with the same aminopeptidase N receptors as their wild type unmodified counterparts. 8. Cry1C as a promising bioinsecticide

7. Overcoming possible selection for Cry-resistant insect populations in transgenic plants The intensive use of crops expressing a single Cry toxin can impose selection pressure towards evolution of insect populations tolerant to a certain Cry protein. Recent analysis of 10-year data of six major insect pests in Australia, China, Spain and USA, has identified an increase in resistant alleles only in some field populations of Helicoverpa zea but not in other pests [7]. In contrast, another recent report has doubted this reported increase in Cry1Ac-resistant alleles of H. zea [6], claiming that the estimation of field efficacy in Bt-cotton areas is missing [79]. However, selections for resistance against Cry proteins under laboratory conditions have shown that many species are able to evolve high degree of resistance but the organisms are highly unfit and cannot survive in the wild [79,80]. Cry1Ca-resistant populations of S. exigua and S. littoralis were selected by continuous feeding on Cry1Ca-containing diet [81,82]. In the S. exiguaresistant mutant, no change in binding site concentration but a fivefold decrease in Cry1Ca binding affinity was observed [81]. This resistant mutant lacks the expression of Class-1 aminopeptidase N

Cry1Ca is essentially a lepidopteran-specific insecticidal protein with toxicity spectrum different from that of the Cry1A ‘‘active toxin’’s. The insecticidal range of Cry1Ca is compared to other relatively effective Cry proteins in Table 1. Cry1Ca is toxic to a wide spectrum of insect larvae including Spodoptera spp., H. virescens, Mamestra configurata, Phthorimaea operculella, Pectinophora gossypiella and P. xylostella (M. Keller unpublished results), that are not well controlled by Cry1A. These insects cause extreme losses to major crops, such as corn, cotton, potato, canola, and soybean worldwide. Consequently bacterial formulations containing Cry1Ca have been developed and used for controlling pests essentially belonging to Spodoptera spp. A synthetic cry1Ca5 with plant codon usage and without RNA pre-termination signals was synthesized [36] and constitutively expressed as a plant nuclear gene in alfalfa, Arabidopsis, cotton, lettuce, potato, tobacco and tomato (Fig. 2 [36] and M. Keller, B. Sneh and A. Zilberstein unpublished results). All the Cry1Ca expressing plants were resistant to S. littoralis larvae, including the fifth instar larvae, which are the least susceptible to most of the chemical insecticides (Fig. 2A–C and [36]) and are not controlled by Cry1A. These Cry1C

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Fig. 2. Synthetic Cry1C expressing plants are tolerant to Spodoptera littoralis (A–C), Chrysodeixis chalcytes (D) and Phthorimaea operculella (E and F). Potato (A, D, E), lettuce (B) and tomato (C and F) constitutively over expressing the synthetic Cry1Ca5 [36] (left) and wild type (WT) plants (right) were challenged with 3rd to 5th instar larvae and photographed when a significant damage was observed in the wild type plants. Cry1A toxins do not control these insects.

expressing plants were also resistant to Chrysodeixis chalcytes (Fig. 2D) and potato tuber moth (Fig. 2E and F). The potato tuber moth P. operculella is the most destructive insect pest of potato in warm climate countries, including the Middle Eastern countries. Conventional breeding has not been successful in developing resistant cultivars to the P. operculella larvae due to lack of reliable resistance sources in available potato germplasm. Although Cry1Ba is more toxic than Cry1Ca to P. operculella larvae, transgenic potato and tomato plants, expressing Cry1Ca, provided the plants with sufficient resistance (Fig. 2E and F). The same cry1Ca5-based synthetic gene [36] was also introduced into broccoli and Indian mustard and rendered the plants resistant to diamondback moth [13,88], which was always hard to control with Cry1A. Larvae of Spodoptera spp., which are relatively resistant to Cry1A toxins, show high susceptibility to Cry1Ca. Insects that have evolved resistance to Cry1Ac [88], remain sensitive to Cry1Ca, further indicating that these toxins interact with different membrane receptors [90,91]. Correlatively, binding assays to various aminopeptidase N receptors, specific for Cry1A recognition, showed that Cry1Ca only interacts with unique aminopeptidase N proteins isolated from Cry1Ca susceptible larvae (summarized by [11,46,50,53]). Comparable effects of Cry1Ca and Cry1Cb were observed in Trichplusia ni. However, Cry1Cb was less toxic to S. exigua than Cry1Ca [92]. Thus, the Cry1Ca group of almost identical toxins may serve as promising bioinsecticides to broaden the spectrum of genes introduced to Bt crops. Further understanding of the interaction between Cry1Ca and its specific receptor/s and also with other membrane components will eventually result in extending its utility in the pyramiding approach. This could lead to improved Bt crops with activity towards a wider spectrum of insects and with more resilience against the forces of evolution. 9. Conclusion The members of the Cry1Ca sub-group of proteins are highly similar and their insecticidal effects have been partially investigated. The present Cry1Ca studies show that Cry1Ca acts via more than one specific receptor that are different from Cry1A receptors, although still sharing the general dependence on intact membrane lipid rafts. These data strongly support the usage of Cry1Ca in

widening the anti-lepidopteran-protein spectrum of Bt crops. It would better cover target insect pests that are widely scattered in moderate and warm climates, such as various Spodoptera species. Since Cry1Ca interacts with more than one membrane target, it may be useful as an efficient component in the pyramiding strategy to avoid evolution of resistant insect populations in the coming generations of Bt crops.

Acknowledgements We thank Prof. M. Adang, University of Georgia. Athens, Ga 30605, USA and Prof. Raj Bhatnagar, Dr. Neema Agrawal and Bindiya Sachdev, The International Centre for Genetic Engineering and Biotechnology, New Delhi, India, for their collaboration and mutual exchange of ideas and know-how. The help of Dr. Mohamad Abu-Abied, Institute of Plant Sciences, Volcani Center, Israel and Dr. Ron Vunch, Department of Plant Sciences, the Weizmann Institute of Science, Rehovot, Israel, in generating Cry1C expressing tomato and lettuce plants is highly appreciated. This study was supported by the U.S.–Israel Binational Science Foundation, Grant No. 2001-235 and by Joint India–Israel Research Grant 411-4161. References [1] C. Zhu, S. Naqvi, S. Gomez-Galera, A.M. Pelacho, T. Capell, P. Christou, Transgenic strategies for the nutritional enhancement of plants, Trends Plant Sci. 12 (2007) 548–555. [2] A. Bravo, S.S. Gill, M. Soberon, Mode of action of Bacillus thuringiensis Cry and Cyt toxins and their potential for insect control, Toxicon 49 (2007) 423–435. [3] J.A. Gatehouse, Biotechnological prospects for engineering insect-resistant plants, Plant Physiol. 146 (2008) 881–887. [4] J.B. Torres, J.R. Ruberson, Interactions of Bacillus thuringiensis Cry1Ac toxin in genetically engineered cotton with predatory heteropterans, Transgenic Res. 17 (2008) 345–354. [5] R.A. de Maagd, A. Bravo, N. Crickmore, How Bacillus thuringiensis has evolved specific toxins to colonize the insect world, Trends Genet. 17 (2001) 193–199. [6] C. Zhong, D.J. Ellar, A. Bishop, C. Johnson, S. Lin, E.R. Hart, Characterization of a Bacillus thuringiensis delta-endotoxin which is toxic to insects in three orders, J. Invertebr. Pathol. 76 (2000) 131–139. [7] B.E. Tabashnik, A.J. Gassmann, D.W. Crowder, Y. Carriere, Insect resistance to Bt crops: evidence versus theory, Nat. Biotechnol. 26 (2008) 199–202. [8] J. Clive, Global status of commercialized biotech/GM crops: 2007, ISAAA Brief No. 37 ISAAA: Ithaca, NY, 2007. [9] R.A. de Maagd, A. Bravo, C. Berry, N. Crickmore, E. Schnepf, Structure, diversity, and evolution of protein toxins from spore-forming entomopathogenic bacteria, Annu. Rev. Genet. 37 (2003) 409–433.

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