Genetic Engineering of Crops for Insect Resistance

Gatehouse, John A.

doi:10.1007/978-1-4614-5797-8_239

John A. Gatehouse⁷

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Definition of the Subject

Genetic engineering of crops for insect resistance is the introduction of specific DNA sequences into crop plants to enhance their resistance to insect pests. The DNA sequences used usually encode proteins with insecticidal activity, so that in plants which contain introduced DNA, an insecticidal protein is present. However, other strategies to improve plant defenses against insects have been explored. Genetically engineered crops that are protected against major insect pests by production of insecticidal proteins from a soil bacterium, Bacillus thuringiensis , have become widely used in global agriculture since their introduction in 1996.

Introduction

Twenty years have elapsed since the first publications describing transgenic plants, which showed enhanced resistance to insect herbivores, as a result of the expression of a foreign gene encoding Bacillus thuringiensis (Bt) toxin [1–3]. In the intervening years, crops expressing these toxins have become widely used in global agriculture, and have led to reductions in pesticide usage and lower production costs [4] At the same time, the predictions made by lobby groups supporting “organic” crop production , that irreversible environmental damage would be caused by genetically engineered (GE) crops resistant to insect pests, have not been realized [5]. Despite all the controversy that GE crops have caused in many countries, it is difficult to dispute that the use of this technology to combat insect pests has had a positive impact on global agriculture.

This entry has two aims: first, to provide a summary of how and why Bt toxins have become the insect resistance genes of choice for commercial GE crop applications, and to anticipate some further developments of this technology; second, to consider some of the other approaches to engineering insect resistance in plants, and to assess their potential for future development in the development of sustainable agriculture.

Insecticidal Proteins from Bacillus thuringiensis

The presence of insecticidal toxins in the soil bacterium Bacillus thuringiensis (Bt) has enabled both the bacteria themselves, and genes derived from them, to be exploited as plant protectants. The toxicity is almost invariably based on proteins produced during sporulation of the bacteria, which form crystalline deposits associated with the spores. The insecticidal Bt proteins are encoded by genes present on plasmids, and the presence of these plasmids is the main feature which distinguishes Bt from other spore-forming bacilli [6]. Preparations of Bt spores have been used since the 1920s as a conventional, spray insecticide (and, as a “natural” product, are approved for use in organic agriculture), but their efficacy in the field is limited by inactivation and low persistence.

The ecological niche occupied by Bt appears to be simple to define. The life cycle starts with a spore and associated crystalline protein body which may be present in the soil. On being eaten by an insect, the protein deposit associated with the spore is dissolved and digested, converting the crystalline protoxin to an active toxin. The insect is then killed, and the carcass provides nutrients for the growing bacteria, which multiply rapidly. When the insect carcass is exhausted, the bacteria sporulate; the spores are dispersed, and the cycle recommences. However, this cycle is clearly too simplistic, as the target insects for Bt toxins are only rarely soil dwellers, and the dose of spores required to kill an insect larva is too large for dispersed spores to have much effect. Although Bt is widely distributed, levels of the bacterium in soils are generally too low to have any effect on insects, and spraying plants with spores does not result in persistent protection as a result of the establishment of a high bacterial population. The species has been described as an opportunistic pathogen, which has evolved the sporulation mechanism as a “backup” system to ensure its survival under unfavorable conditions [7]. Bt is naturally present in the phylloplane, as well as in soil, and has been detected on cabbage foliage [8], and in vegetative form on clover [9] at low levels, without any insecticidal effect. However, the insecticidal characteristic must be of benefit to the bacterium, since most of the insecticidal proteins are encoded by plasmids, and the plasmids are maintained in the Bt population as a whole, despite the obvious metabolic costs of producing large quantities of spore-associated proteins. Not only are toxin-encoding plasmids maintained, but there is also a huge reservoir of diversity in the toxins themselves, and much effort has been put into screening bacterial isolates for strains of Bt with novel pesticidal activities [10].

Bt toxins are now classified on the basis of amino acid sequence similarity (an earlier classification system based on pesticidal activity has been superseded), in a systematic hierarchical system [11]. For the purposes of this contribution, only the major distinctions need be considered. There are four types of insecticidal proteins produced by Bt:

1.
Proteins associated with Bt spores, usually as crystalline deposits; three domain structure; single toxins; designated by the symbol Cry
2.
Proteins associated with Bt spores, usually as crystalline deposits; binary toxins and other similar proteins, including truncated versions of three-domain toxins; also designated by the symbol Cry
3.
Proteins associated with Bt spores, usually as crystalline deposits; single domain structure; cytolytic; single toxins; designated by the symbol Cyt
4.
Proteins expressed vegetatively by Bt; single chain and binary toxins; designated by the symbol Vip

Each type of toxin is subdivided (on the basis of sequence similarity) into families (number; same number ≥ 45% sequence identity) and then further subdivided using capital letters (same letter ≥ 78% sequence identity), small letters (same letter ≥ 95% sequence identity) and numbers successively. The resulting system yields designations for specific toxins such as Cry1Aa. A single Bt strain can produce spores which contain only a single toxin, or a complex mixture, such as the Bt subspecies israelensis, whose spores contain Cry4Aa, Cry4Ba, Cry10Aa, Cry11Aa, Cyt1Aa, and Cyt2Ba toxins [12].

All four types of proteins have been proposed for use as crop protection agents, although Cyt toxins have not as yet been used in commercial insect-resistant transgenic plants, and three-domain Cry toxins are by far the most commonly used type. Cry and Cyt toxins belong to the class of proteins referred to as bacterial pore-forming toxins, and show structural similarity to the α-helical and β-barrel groups of toxins, respectively (where α-helical and β-barrel refer to the structures of the membrane-spanning parts of the toxin; reviewed by Parker and Feil [13]).These pore-forming toxins show common features of activity; they are produced as water-soluble proteins, and interact with specific receptors on cell surfaces, often after proteolytic activation by host proteinases. Binding to cell surfaces triggers a conformational change leading to oligomerization, which allows insertion into the cell membrane through promotion of a fluid, partially denatured structure. Insertion of the toxin into the membrane can either cause cell death directly, or result in effects on intracellular metabolism which lead to cell death.

How Do Bt Toxins Work?

Three-Domain Cry Toxins

The mechanism of action of the “conventional” three-domain Cry toxins is now well understood, and can be divided into four stages:

1.
Solubilization of the protoxin, and proteolytic activation by proteinases in the insect gut to produce active toxin
2.
Interaction of the toxin with one or more receptors on cell surfaces in the insect gut epithelium
3.
Oligomerization of the toxin
4.
Insertion of the oligomerized toxin into cell membranes, leading to the formation of open pores, and cell death (see Fig. 1)

**Genetic Engineering of Crops for Insect Resistance, Figure 1**

Following the pioneering work of Ellar’s group [14] tertiary structures of six different three-domain Cry toxins are known – Cry1Aa [15], Cry2Aa [16], Cry3Aa [14], Cry3Bb [17], Cry4Aa [18], and Cry4Ba [19]; whereas most structures are for the active form, the structure of Cry2Aa includes the N-terminal pro-region. These toxins all show a high degree of structural similarity, and thus the formulation of a general model for their mode of action is justified. The three domains present in the active forms of these proteins are designated I, II, and III, and are normally contained in a single polypeptide of approximately 600 amino acid residues (in some cases proteolytic cleavages are present within the active three-domain structure as a result of protoxin activation , resulting in multiple polypeptides making up the toxin, but the overall three-domain structure is conserved.). While conservation of structure and sequence is observed in the active forms of three-domain toxins, many toxins are synthesized with C-terminal extensions, which are variable in sequence between Bt strains, and in length between Cry families. The presence of C-terminal extensions leads to a large degree of heterogeneity in the size of the protoxins present in bacterial spores, with sizes ranging from approximately 600 amino acids (similar to the active toxin) to approximately 1,200 amino acids. These C-terminal extensions are not required for toxin function, and are removed during toxin activation, although their removal is not sufficient for toxicity to be shown. They are thought to play a role in the formation of crystalline inclusions in the bacterium during the spore-forming process .

The three domains of the active toxin are clearly distinguished in their structures.

1.
Domain I, approx. 260 aa, contains seven α-helices, of which six are amphipathic and one hydrophobic. This structure is typical of pore-forming toxins, with the hydrophobic and amphipathic helices being responsible for membrane insertion and pore formation. The hydrophilic sides of the amphipathic helices form the surface lining the pore, so that polar species such as ions are able to cross the membrane.
2.
Domain II, approx. 170 aa, forms a “β-prism” structure, with three β-sheets, and exposed loops on its surface.
3.
Domain III, approx. 160 aa, has a compact structure with two anti-parallel β-sheets in a “jellyroll” formation, and is structurally similar to carbohydrate-binding domains such as the cellulose-binding domain in cellulases [20]. A general model for three-domain toxins is shown in Fig. 2.

**Genetic Engineering of Crops for Insect Resistance, Figure 2**

The Proteolytic Activation Process

Ingestion of the Cry protoxins by the insect leads to solubilization of the proteins, and exposure to digestive proteinases in the insect gut. Although removal of the C-terminal protoxin region occurs at this stage, the essential step in protoxin activation is the proteolytic cleavage and removal of an N-terminal peptide, which varies from approx. 25–60 amino acids in different Cry proteins. A non-activatable Cry1Ac mutant toxin could not form pores in insect membrane vesicles derived from gut epithelial cells [21], and it is thought that the N-terminal peptide “masks” a region of the toxin involved with interaction with receptors [16]. The activated toxin is fairly resistant to further proteolytic cleavage, which enables it to survive long enough in the gut to reach its site of action, the gut epithelial surface (Fig. 1).

This summary overlooks a number of factors which contribute to toxicity. First, the location of the proteolysis may be important, since many insects, such as diptera (flies), carry out digestion in the foregut, which is chitin-lined and does not contain epithelial surfaces, or even outside the insect altogether, by secreted saliva or regurgitated gut contents. Under these circumstances, the toxin will need to be more resistant to proteolysis, or more effective, since the time between activation and reaching the site of action will be longer. Secondly, gut conditions vary significantly between insects from different orders, or even within orders; in general, larvae of lepidoptera (moths and butterflies) have a highly alkaline midgut environment (pH 10–11 in many major crop pests), whereas larvae of coleoptera (beetles) have an acidic gut environment (pH approx. 5 for many species). These differences in conditions will affect both the activation and survival of the protein, although they may be less relevant to steps taking place at the gut surface, where there is a separation from the gut lumen by the peritrophic membrane (a macroscopic porous chitin-based structure) and by lipids sloughed off from the gut surface. Finally, the nature of the digestive enzymes present in the insect gut differs considerably between different orders; whereas most insects use serine proteinases with an alkaline pH optimum as their major endoproteinases, many coleopteran larvae use cathepsin-type cysteine proteinases with an acidic pH optimum (similar to lysosomal proteinases). On the other hand, protoxin activation does not appear to be very sequence specific. Many lepidopteran-specific Cry proteins can be activated in vitro by mild treatment of the protoxin with bovine trypsin, yielding products that appear to be similar to those formed in vivo. This suggests that it is the three-dimensional structure of the protoxin that determines where proteolysis takes place, unless forcing conditions are used.

Interactions with Receptors

Proteins to which Cry proteins bind in the insect gut are termed “receptors,” although the specificity of interaction is determined by the Cry protein itself, and the ligands to which it binds do not show the properties of receptors as normally understood. Binding takes place on the microvillar membranes of the cells forming the midgut epithelium, and involves interactions with relatively abundant proteins, either attached to the cell membrane by glycosylphosphatidyl-inositol (GPI) -anchors, or integral to the membrane with large extracellular domains. The overall process is summarized in Fig. 3.

**Genetic Engineering of Crops for Insect Resistance, Figure 3**

Methods for identifying receptors to which Cry proteins bind have largely been based on immunoblotting of proteins prepared from brush border membrane vesicles (BBMV) . This method is not a good mimic of conditions in vivo, and may result in interactions with lower affinity, or which are dependent on protein conformations maintained by membranes, not being observed. Nevertheless, the major binding partners for Cry proteins which have been identified show binding when assayed as purified proteins, and as components of BBMVs, with binding constants in the range 1–100 nM.

The initial identification of membrane-anchored aminopeptidase N [23] and an integral membrane cadherin-like protein designated Bt-R1 [24] as Cry1A toxin receptors in lepidopteran insects has been supplemented more recently by identification of a 270 kDa glycoprotein [25] and alkaline phosphatase (membrane anchored; [26]) as additional potential receptors. Alkaline phosphatase appears to be the major receptor in mosquitoes [27]. A recent proteomic analysis has identified further potential receptors, such as V-ATP synthase subunit 1 [28]. However, this analysis also showed binding to actin, which could not be present at the cell surface, showing that results from blotting experiments need to be interpreted critically.

Functional roles as “receptors” for aminopeptidase N and cadherin Bt-R1 in Cry protein toxicity are supported by numerous studies. Strains of lepidopteran insects resistant to Cry1 toxins have been identified which show mutations in the gene encoding Bt-R1, leading to the production of a truncated cadherin lacking the extracellular domains [29, 30]. The correlation with loss of function of cadherin with loss of susceptibility to Cry toxins suggests that binding to the extracellular domains of cadherin is a necessary step for toxicity. Binding of Cry1A toxin to the cadherin extracellular domains has been demonstrated in vitro, and the binding regions have been identified in some detail [31]. Both gain-of-function and loss-of-function assays have been used to provide further evidence for involvement of cadherin in toxicity; when transiently expressed in mammalian cells that were not normally susceptible to Cry toxin, Bt-R1 genes from silkworm conferred sensitivity to Cry1A toxins [32]; whereas suppression of cadherin expression by RNA interference in tobacco hornworm (Manduca sexta) decreased sensitivity to Cry1Ab toxin [33]. In the case of aminopeptidase N, similar correlations between resistance to Bt toxin and lack of expression of specific isoforms of the protein have been observed [34], but more direct evidence has come from downregulation of aminopeptidase by RNA interference using double-stranded RNA. This technique has been carried out in lepidopteran larvae, giving decreased sensitivity to Cry1C toxin [35], and in lepidopteran cell cultures, giving decreased sensitivity to Cry1Ac [36]. A gain of function experiment in which transgenic fruit flies (Drosophila) expressing lepidopteran aminopeptidase N became sensitive to the lepidopteran-specific toxin Cry1Ac [37] showed elegantly and convincingly that this receptor plays a key role in toxicity. Binding to aminopeptidase N involves interaction of Cry toxins with the carbohydrate side-chains of the protein [38, 39], with specificity toward GalNAc residues being shown (this sugar can inhibit binding; [40]). Binding to carbohydrate facilitates subsequent protein–protein interactions, which are thought to be necessary for toxicity [41]. Functional evidence for alkaline phosphatase acting as a Cry toxin receptor has again been provided by correlative observations, in that insect lines resistant to Cry1Ac toxins have lower alkaline phosphatase levels than susceptible lines [26]. Interactions with protein-bound carbohydrate also seem to be involved in the binding of Cry toxins to alkaline phosphatase.

The roles of the different domains of Cry proteins in the interaction with receptors are clearly distinguished. Despite the presence of the N-terminal propeptide which must be removed for activity, domain I plays little or no role in the interaction with receptors, whereas domain II is responsible for most protein–protein interactions (see Fig. 2), and domain III is responsible for binding to carbohydrates. This division of roles is consistent with the observation that a single toxin can interact with more than one type of “receptor”; for example, Cry1Ac interacts with both Bt-R1 and aminopeptidase N [22]. The protein–protein interactions mediated by domain II have been localized to variable loop regions on the surface of the domain, whereas the carbohydrate-binding region of domain III is a typical binding site cleft, which is spatially well-separated from the domain II loops.

Oligomerization

Oligomerization is a common feature in bacterial pore-forming toxins, and Cry proteins appear to conform to the model, with the formation of oligomeric structures (probably tetramers) observed for toxins from the Cry1 and Cry3 families. Mutants of the Cry1Ab protein that have impaired oligomerization ability, but bind to the receptor, show much reduced toxicity or no toxicity toward lepidopteran larvae [42]. Similarly, monomeric Cry proteins have much lower intrinsic pore-forming abilities on synthetic membranes than oligomerized preparations [43]. Oligomerization is promoted by binding to a receptor; in the case of Cry1Ab protein binding to the cadherin Bt-R1 receptor, this process involves an additional proteolytic cleavage at the N-terminal end of the protein, in domain I [44]. The proteolytic cleavage, carried out by host enzymes, may aid the oligomerization process. The importance of oligomerization in promoting toxicity has been shown by two complementary studies. First, a peptide corresponding to the region of cadherin to which Cry1A binds has been shown to act as a synergist, increasing the toxicity of Cry1A toward lepidopteran larvae [45], presumably as a result of the binding between the peptide and Cry1A promoting oligomerization of the toxin prior to interaction with the gut epithelium. Secondly, mutants of Cry1Ab toxin have been produced which contain deletions corresponding to the proteolysis in helix 1 of domain I which occurs on binding to cadherin. These mutated toxins form oligomers in the absence of cadherin binding, and are effective against insects that have cadherin expression suppressed, or which have a cadherin mutation which leads to resistance to unmodified toxin [33]. These results have led to a current view that cadherin is the primary receptor for Cry toxins, since it is necessary to promote oligomerization, with other molecules taking the role of “secondary receptors” [33].

Insertion into the Cell Membrane

The oligomeric Cry protein must partially unfold in order for the pore-forming domains (domain I) to insert into the membrane. In the case of bacterial pore-forming toxins active against mammalian cells, this partial denaturation process is stimulated by acidic pH at the cell surface [13]. A similar mechanism could occur with Cry proteins active against lepidopteran insects, although the gut pH is very alkaline; the partial denaturation could still be triggered by a decrease in pH at the cell surface. The pH optimum for aminopeptidase N in lepidopteran larvae (8.0; [46]) is at least 2 pH units less than bulk gut content pH (>10), suggesting that a decrease in pH occurs near the cell surface. The involvement of lipid rafts, microdomains which are less fluid than the membrane as a whole, in pore formation has been suggested [47]. However, membrane-anchored proteins are selectively associated with these lipid rafts, and it is not clear whether lipid rafts are necessary for pore formation, or whether their involvement is a result of the presence of receptors. The trans-membrane cadherin-like Bt-R1receptor is not associated with lipid rafts.

A current model for pore formation by Cry1A toxins suggests that interaction with two receptors is necessary; an initial binding step with the cadherin-like Bt-R1 receptor leads to toxin oligomerization, followed by interaction of the oligomer with the aminopeptidase N receptor and insertion into the membrane [22, 48]. While this model is plausible, the details of the mechanism of toxicity must differ for different toxins, and a “two-receptor” model should not be assumed to be generally applicable. The gain of function experiments described above show that only one receptor is necessary for toxicity to be shown, and only a few lepidopteran-specific Cry toxins have been shown to interact with cadherin-like proteins [49]. If the major determinant of Cry protein toxicity is the assembly of oligomeric complexes at the surface of cells in the gut epithelium, then this requirement can be met in diverse ways, involving different “receptor” proteins to localize the toxin and promote oligomerization (although the interaction is always likely to involve the most abundant proteins at the cell surface). A “global” diversity of interactions is not inconsistent with specificity when interactions between specific toxins and hosts are considered.

Once the insertion of Cry toxin into the cell membrane leads to pore formation, the gut epithelial cell is unable to maintain its internal solute balance, as the open pore allows free exchange of ions and other small molecules between the gut lumen and the cytoplasm. The cytoplasm of gut cells has markedly different concentrations of ions (including H⁺) than the gut lumen; this difference in concentrations is used to drive active transport processes, such as amino acid transport [50]. Free movement of ions thus causes massive disruption to cell physiology, leading to death. The leakage of cell contents also causes proliferation of gut microflora, so that dying insects show massive bacterial infection of collapsing gut tissue. Cry proteins may also produce toxic effects through interference with signaling pathways. Binding of Cry1Ab to the transmembrane Bt-R1 receptor has been shown to activate a G-protein-mediated intracellular signaling pathway, resulting in the formation of cAMP by adenylyl cyclase, and activation of protein kinase A [51]. This process led to cytological changes typical of Bt toxin activity.

Binary Cry and Vip Toxins

The binary Cry toxins are exemplified by toxins active against corn rootworm [52]. These toxins are only active as a combination of two proteins, designated as families Cry34 (14 kDa protein) and Cry35 (44 kDa protein). The two proteins are the product of a single operon in the commonly used Bt strains. The binary toxin acts on the insect gut epithelium, and leads to swelling and vesicle production from epithelial cells, resulting in the disappearance of microvilli, and extensive disruption of the epithelium. However, it is not clear whether these symptoms are solely a result of open pore formation, or whether other modes of toxicity, such as ADP-ribosylation (see below) are occurring. No structural information on these proteins is available at present. There is evidence that the 44 kDa toxin protein Cry35 is evolutionarily related to an insecticidal toxin from Bacillus sphaericus [53]. The B. sphaericus toxins have received some attention due to their toxicity toward mosquitoes and other dipteran insects. They also bind to membrane-anchored receptors (α-glucosidase, in the case of the mosquito Culex pipiens [54]) and cause disruption of the gut epithelium [55]. However, their detailed mechanism of action is not known. Like Cry34/35, the B. sphaericus proteins are binary toxins, although in this case one component does show limited activity in the absence of the other. The designation of the corn rootworm binary Bt toxin by the symbol Cry obscures the fact that these toxins have little in common with the three-domain toxins, besides being found in crystalline deposits in Bt, and being insecticidal as a result of acting on the insect gut epithelium.

The Bt insecticidal Vip1/2 proteins (active against corn rootworm) are also binary toxins with similarity to the B. sphaericus toxins [56]. The mechanism of action of Vip1/2 toxins involves ADP-ribosylation by the active component, which disrupts actin polymerization in cellular microfilaments, similar to other bacterial ADP-ribosylating toxins such as botulinum toxin [57]. The inhibition of actin polymerization leads to massive disruption of cellular functions. The Vip1Ac binding component of the binary toxin interacts with membranes to form oligomeric channels, allowing the active component to gain access to the cell cytoplasm [58].

A further class of Vip proteins , Vip3, (active against lepidoptera) has been identified; these protein are single chain toxins which lyse insect gut cells by pore formation in membranes, and have no sequence similarity to Vip1/2 [59, 60]. Vip3 binds to brush border membrane vesicles prepared from target insect gut epithelial cells, but does not bind to the same receptors as Cry1 and Cry2 proteins [61]. Binding to 80 and 100 kDa membrane proteins is observed in ligand binding experiments [62], but these receptors have not been characterized. These proteins are promising candidates for further development; chimeric toxins containing regions from different Vip3 toxins have been produced and show extended ranges of toxicity toward lepidopteran pests [63].

Cyt Toxins

The cytolytic Cyt toxins, also found in crystalline inclusions in some Bt strains, are single polypeptides, of approx. 250 amino acids; the N-terminal region contains α-helices which wrap around a C-terminal β-sheet core in the three-dimensional structure [64]. Pore formation results from insertion of the β-sheet region into membranes [65]. Unlike the three-domain Cry toxins, this membrane insertion is not receptor-mediated [66]; the Cyt toxins insert directly into membranes, and are thus cytolytic to a wide range of cells. Like the three-domain Cry toxins, Cyt toxins are synthesized as inactive protoxins which are activated by proteolysis. Activation involves removal of propeptides from both the N- and C-termini of the protoxin; in the case of Cyt2Aa, 32 aa are removed from the N-terminus and 15 aa from the C-terminus to generate active toxin [67]. This process does not require specific proteinases.

The combination of Cry and Cyt toxins found in crystalline inclusions in some Bt strains, specifically in the strains of Bt subsp. israelensis active against mosquito larvae, is highly effective as a toxin due to synergistic interactions between its components. Not only are the three domain Cry protein components in these crystals more effective toxins in the presence of Cyt proteins, but the Cyt proteins also prevent resistance to Cry proteins from developing when insects are exposed to purified protein preparations under laboratory conditions [68]. This synergistic effect could result from the two types of toxin producing complementary disruption of the insect gut epithelial cell membranes, but evidence has been presented that Cry and Cyt toxins can interact directly. Specifically, Cry11Aa and Cyt1Aa bind strongly to each other, both in solution and in a membrane-bound state, and binding of Cry11Aa to mosquito gut epithelial cell membranes was enhanced by pretreating the membranes with Cyt1Aa [69]. The interaction with Cyt1Aa takes place through the loop region in Cry11Aa involved in protein–protein interactions with its “normal” receptor (membrane GPI-anchored alkaline phosphatase). Insertion of Cyt1Aa into gut cell membranes, which is not dependent on receptor mediation, thus generates additional “receptors” for Cry11Aa, increasing its toxicity, and preventing resistance developing by mutation of the insect-encoded “receptor.”

Expression of Genes Encoding Bt Insecticidal Proteins in Transgenic Plants

Expression of Three-Domain Cry Toxins from Transgenes in the Nuclear Genome

Almost all the insect-resistant transgenic crops currently in use express three-domain Cry proteins from Bt as their protective agent. The initial laboratory-based experiments expressed Cry1 toxins in plants to give protection against lepidopteran larvae, and this has remained the main focus of Bt gene utilization up to the present day. However, the three-domain Cry proteins pose a number of problems in terms of expression in plants. The technology involved in achieving sufficient levels of accumulation of these proteins to give adequate levels of protection was initially challenging, but developed rapidly, so that within 5 years of the initial reports of engineered resistance, the methodology for gene manipulation was essentially complete. The slower pace of transfer of this technology into major crop species observed subsequently has had much more to do with technical difficulties in plant transformation (particularly regenerating viable plants), than with any problems at the level of gene constructs. The minimum level of Cry protein expression in leaf tissue to give high levels of mortality of sensitive lepidopteran larvae under laboratory conditions is approximately 0.05% of total protein, but to give effective field protection against species which are less sensitive to Bt toxins, and to manage resistance to the toxin in pests (see later), levels of expression an order of magnitude higher (i.e., 0.5% of total protein) are desirable.

Engineering genes encoding three-domain Cry proteins for expression in transgenic plants has been extensively described (the review by Mazier et al. [70], gives a particularly comprehensive survey), but a short summary of the main considerations which had to be taken into account is relevant here. These were:

1.
How much of the protein coding sequence should be expressed in plants?
2.
Which promoters should be used to drive expression of the Cry protein coding sequence in plants?
3.
How should the coding sequence be altered to avoid poor expression?

Protein Coding Sequence

The C-terminal part of protoxins for three-domain Cry proteins is variable, and absent in some toxins. Its role in directing the formation of crystalline inclusions in Bt sporulation is not required when the proteins are expressed in plants (and might result in disruption of cells unless the protoxin was exported into intracellular spaces). All constructs which result in insecticidal activity have omitted this part of the molecule from the coding sequence expressed in plants. The initial research suggests that a complete protoxin accumulates in plant tissue at levels 10–50-fold less than a protoxin truncated so the C-terminal region is absent [3]. Since removal of the C-terminal region of the protoxin does not result in active toxin being produced, retention of the N-terminal activation peptide ensures that the initial protein product in transgenic plant tissue is not active, and proteolytic activation takes place as normal within the gut of insect herbivores. The coding sequence utilized thus corresponds to the three-domain structure shown in Fig. 2, plus the additional N-terminal propeptide.

Promoters

The problems experienced in achieving levels of expression of Cry proteins high enough to confer effective protection meant that the initial use of promoter sequences which only gave low levels of expression, such as those from Agrobacterium tumefaciens Ti plasmids, was rapidly superseded by strongly expressed promoters, most of which were based on the Cauliflower Mosaic Virus 35S RNA promoter (CaMV 35S). Constitutive expression of the Cry protein in all plant tissues does not appear to cause significant problems either in a yield penalty, or deleterious effects due to the accumulated protein. However, tissue-specific promoters have also been used, such as the ribulose-bisphosphate carboxylase small subunit promoter (e.g., [71]) or the phosphoenolpyruvate carboxylase promoter (e.g., [72]), both of which are specific for green tissue. The CaMV 35S promoter was initially considered to be specific for dicots, but further experience showed that it could also be functional in monocots, and, with suitable modification, could be used to direct Cry protein expression (e.g., [73]). However, many researchers have preferred to use promoters derived from constitutively expressed monocot genes in Cry protein expression constructs for use in cereal transformation (e.g., the maize ubiquitin-1 promoter; [74]). Root-expressed promoters have been used in constructs designed to protect cereals against corn rootworm [75].

Considerable research has also been undertaken on the use of promoters whose expression is only induced under specific conditions. The use of wound-induced promoters to direct Cry protein expression has the apparent advantage that production of Cry proteins in transgenic plants is, for the most part, only induced on attack by insect pests. Any potential deleterious effects on phenotype caused by production of the toxin in transgenic plants would therefore be minimized, and toxin residues in plant tissues would be reduced. A wound-inducible maize proteinase inhibitor gene promoter has been used to direct expression of Cry1B in transgenic rice, and has been shown to give effective protection against insect attack (against striped stem borer; [76]). However, the protection afforded by transgene constructs containing wound-inducible promoters is lower than when constitutive promoters are used, both in the laboratory and in the field [77].

While achievement of expression levels of Bt toxins sufficient to confer protection in transgenic plants is now considered routine, considerable technical problems may still need to be overcome when specific crop species are considered (e.g., soybean; [78]). These include the construction of the synthetic coding sequence for the toxin, choice of an appropriate promoter for the expression construct, developing protocols for efficient transformation and regeneration of the plant species, and production of homozygous progeny lines containing the transgene.

Engineering the Coding Sequence to Optimize Expression

The initial experiments in which Cry toxins were produced in transgenic plants showed that only low levels of Cry protein were accumulated, generally of the order of 0.01% of total protein, or less. Levels of Cry proteins were at least one order of magnitude lower than when plant proteins were expressed using similar promoters in expression constructs, leading to the deduction that the Cry protein coding sequence contained features which decreased protein production as a result of posttranscriptional events. Cry protein coding sequences are generally A-T rich compared to plants (coding%GC in Bacillus thuringiensis, 36%; in Arabidopsis thaliana, 45%; in Oryza sativa, 55%; Codon Usage Database, http://www.kazusa.or.jp/codon/) and codon usages thus differ significantly. Cry protein genes were reengineered, modifying the nucleotide sequence without altering the encoded amino acid sequence, to change the codon usage to one more appropriate for plants, resulting in either partially or wholly synthetic genes (reviewed by Mazier et al. [70]). Codon optimization for both dicots and monocots has been carried out. Codon-optimized synthetic genes show accumulation levels of Cry proteins of up to 1% of total protein in leaf tissue, which is adequate for complete protection of plants against pest insects [79].

The basis for poor expression of Cry proteins in transgenic plants has received comparatively little attention. Evidence suggests that the major problem is not codon usage, but instability of RNA transcripts [80, 81]. Expression of unmodified Cry protein coding sequences leads to accumulation of short, polyadenylated transcripts resulting from incorrect recognition of polyadenylation addition signal sequences within the protein coding sequence [82]. Specific modification of A-T-rich regions within the coding sequence of Cry1Ac toxin putatively responsible for transcription termination and polyadenylation (both AATAAA signal addition sequences and ATTTA upstream motifs) has been shown to lead to increased protein expression in transgenic tobacco [83]. Changing codon usage to increase GC content has eliminated these A-T-rich regions in synthetic Cry protein genes, which therefore can produce high levels of stable mRNA.

Expression of Three-Domain Cry Toxins from Transgenes in the Chloroplast Genome

The bacterial origin of the chloroplast is reflected in differences in both the genome composition and organization, and the biochemistry of transcription and translation within the organelle, compared to the nuclear genome and transcription and translation in the nucleus and cytoplasm. The bacterial origin of the genes encoding Cry proteins suggests that expression in the plastid, from transgene constructs introduced into the plastid genome, might result in high levels of protein production. This prediction was confirmed in 1995 with a report showing that incorporation of a construct containing a complete coding sequence for the Cry1Ac protoxin protein and the plastid rRNA operon promoter into the genome of tobacco chloroplasts led to accumulation of Cry1Ac protoxin (approx. 130 kDa – i.e., with the C-terminal crystal-forming region intact) in tobacco leaves to levels of 3–5% of total protein [84]. The high level of Cry protein accumulation meant that transformed plants were effectively protected against attack by several major lepidopteran pests, even beet armyworm (Spodoptera exigua), a species relatively insensitive to Bt toxins.

Despite this highly promising initial report, expression of Cry proteins via plastid transformation has not been widely adopted, and is not used in the current commercial crops. Reasons for this are difficult to pinpoint; there are significant technical problems in achieving stable transformation of plastids, since all of the copies of the plastid genome in the cell (up to 10,000) must be transformed [85], and plastid transformation has been problematic in species other than tobacco [86]. Nevertheless, methods exist to overcome these problems [87]. Cry1, Cry2, and Cry9 proteins have been expressed in plastids of tobacco [88–91], and Cry1Ab has been expressed in soybean plastids [92], all giving high levels of protection against lepidopteran pests to the resulting plants. Overexpression of the Cry2Aa2 operon is particularly effective in giving broad-spectrum protection against a range of pests.

Commercial introduction of transgenic insect-resistant crops based on plastid transformation is almost certainly feasible, but may as yet be restricted by economic considerations, or concern over long-term stability of the transgene phenotype. The maternal inheritance of plastid-encoded characteristics shown by most plants, which means that pollen cannot disperse the transgene to non-transgenic plant stocks, is a further advantage to the method, which could be used to overcome objections to coexistence of transgenic and “organic” agricultural practices by environmental pressure groups.

Expression of Other Genes Encoding Insecticidal Bt Toxins

Gene constructs for expression of other Bt toxins follow the same principles as those outlined above for three-domain Cry toxins. For example, corn expressing the binary Cry34/35 toxin (for protection against corn rootworm) was transformed with a construct containing a constitutive promoter (maize ubiquitin-1) and synthetic coding sequences for the 44 and 14 kDa polypeptides [52], giving expression levels of up to 0.9% and 0.2% respectively of total soluble proteins in plant tissues. Details of the constructs used for expressing these, and other Bt toxins, are apparently not reported in the scientific literature.

Taking Transgenic Plants Expressing Bt Toxins into the Field

Dealing with Pest Resistance to Bt Toxins

The development of successful strategies for commercial deployment of “first generation” insect-resistant crops expressing a single three-domain Cry toxin has focused on a single major potential problem, the development of resistance to the insecticidal compound by the targeted pest species. Development of resistance to exogenously applied chemical pesticides has occurred in over 500 insect species [93], and field resistance to Bt sprays has been observed in the lepidopteran pest diamondback moth (Plutella xylostella). Resistance to Bt toxins can be produced in the laboratory within a small number of generations of many pests, showing that resistance alleles are present in pest populations at a nonnegligible level, although resistance to high doses of specific toxins is only shown in individuals homozygous for the resistance allele. This topic has been ably reviewed in the context of the commercialization of Bt crops by [94]. The most common mechanism of resistance to Cry toxins in insects is mutation in a toxin receptor, leading to a failure to bind sufficient levels of toxin for lethal effects to be shown; however, the involvement of more than one “receptor” in current models for three-domain Cry toxin mechanisms of toxicity (see above) implies that multiple genetic loci for resistance in the pest are possible. Other mechanisms, such as altered proteolysis of toxins, have been proposed to account for the resistance to multiple toxins which can be produced in the laboratory.

The practical solution to prevent the development of resistance in pest populations, the “high-dose/refuge” strategy, has been extensively reviewed elsewhere [94]. In its simplest form, this strategy couples transgenic plants that are expressing sufficient levels of a specific toxin to kill all pest insects which are homozygous negative, or heterozygous, for a resistance allele, with a reservoir of untransformed plants which maintain a population of pests which have a normal frequency of resistance alleles. It assumes that the frequency of occurrence of resistance alleles is low (<10⁻³). Surviving pests on the transgenic plants will be almost all homozygous positives for the resistance alleles, but will be few in number due to the low frequency of occurrence of these alleles. The non-transformed plants will produce a large number of pest insects , most of which are homozygous negative for resistance alleles. Provided that transgenic and untransformed plants are not spatially separated, mating between resistant insects selected on transgenic plants will be a rare event, and most progeny will be homozygous negative or heterozygous for resistance alleles, and thus susceptible to the insecticidal activity of the transgenic plants. In this way, both the pest population is suppressed, and any increase in the frequency of resistance alleles in the population is minimized by the continuous “diluting out” effect.

This approach has been almost wholly successful in controlling pest resistance to Bt toxins in agricultural use of transgenic crops over 10 years. That it has been so successful may be a result of factors other than those originally considered, since the assumption that Bt toxin resistance alleles occur at a very low frequency in natural populations has been called into question. Although some insect populations show resistance allele frequencies in the 10⁻³ to 10⁻² range (e.g., tobacco budworm, Heliothis virescens in USA; [95]; Sesamia nonagrioides in Spain and Greece; [96]), estimates for pink bollworm (Pectinophora gossypiella) in Arizona, USA in 1997 were as high as 0.16 [97]. No evidence for selection for resistance was observed, since the frequency of resistance alleles did not increase over a 3-year monitoring period in which transgenic cotton expressing Bt toxins was extensively employed. A subsequent follow-up study [98] confirmed that frequencies of resistance alleles in this insect had not increased over an 8-year monitoring period, with values generally <10⁻², despite almost continuous exposure to Cry1Ac via transgenic cotton. The possibility that resistance alleles in the insect carry a significant fitness penalty is one additional factor that could account for these observations.

The success of the refuge strategy is dependent on farmers sacrificing part of their crop (untransformed plants) to maintain a pest population. This has been successfully enforced in the industrialized agriculture of developed countries, but may be more difficult to ensure when insect-resistant transgenic crops become available to rural farmers. Although greater agricultural diversity may play the same role as the refuge strategy in maintaining a pest population and decreasing selection pressure, emergence of resistance in pests to Bt crops has been delayed, not eliminated, and further strategies to manage it will be necessary.

Pests That Are Not Susceptible to Bt Toxins

As described above, most of the Bt toxins that have been investigated, and introduced into transgenic crops, are active against lepidopteran or coleopteran insect pests. This is partly a result of the practical requirements of agriculture, since these orders include most of the major pests. However, there are significant insect herbivores which remain outside the range of activity of Bt toxins that have been expressed in transgenic plants.

Dipteran pests , such as fruit flies and root flies, are serious pests in many crops, and Bt toxins active against diptera have been thoroughly investigated. A major problem with introducing protection against these pests into plants is that Bt strains active against dipteran insects usually contain a mixture of toxins, often including both Cry and Cyt proteins (see above). These toxins act synergistically, and individual components are only of low toxicity. Introduction of genes encoding the mixture of toxins found in a typical dipteran-active Bt strain into a transgenic plant has yet to be attempted, although it is not beyond the capacity of existing technology.

The major order of insect herbivores outside the range of Bt toxins is Hemiptera , which includes aphids, plant- and leafhoppers, whitefly, and other sap-suckers which feed directly on the contents of phloem and/or xylem vessels, predominantly sucrose and free amino acids. These insects are important pests and virus vectors. No Bt toxins with activity against them have been found. The reason for this is not clear; receptors similar to those in other insect orders are present in these insects [99], but generally they contain very low levels of digestive proteolytic activity, as a result of ingesting nitrogen in the form of amino acids rather than protein. This lack of digestive proteolytic activity may interfere with activation of Bt toxins, and prevent enough activated toxin to have effects on the insect being present in the gut.

Why Haven’t Plants Evolved Their Own Bt?

Despite the problems encountered in managing resistance of pests to Bt toxins, transgenic plants expressing these insecticidal proteins have proved their value in the field. However, the necessity for resistance management suggests that this solution to defense of plants against insect herbivores may not be viable on an evolutionary timescale. Endogenous expression of Bt toxins is not a “natural” method of defense against herbivores, since plants do not produce similar insecticidal proteins themselves. This failure on the part of plants to exploit a viable strategy for protection seems puzzling, and the obvious explanation, that plants lack the capacity to produce Bt toxin-like proteins, is not correct. Since introduction of suitably modified Bt genes gives adequate levels of protein expression for protection, there is no reason why plants could not have evolved a similar capacity. As discussed in the following section, plants have evolved a diverse array of defensive mechanisms, but make little use of proteins which are highly toxic to insect herbivores. Possibly, this is due to the relative ease with which insects can develop resistance to protein toxins which exert a very strong selection pressure on the population; although alternative hypotheses, such as the balance between investing plant resources into defense versus growth not favoring this strategy, or practical difficulties for a sessile organism in delivering toxins, should also be considered. Unfortunately, the experiments which would enable this issue to be investigated, namely, an evaluation of the “fitness” of Bt-expressing plants in a natural ecosystem in competition with varieties relying on endogenous defenses, and the persistence of Bt genes in a natural population, are unlikely to be carried out in the near future, due to obvious regulatory issues.

Whatever the reason for plants “in the wild” not using defensive proteins similar to Bt toxins, there is no reason to suppose that transgenic plants with engineered insect resistance will not continue to be useful in the artificial growing conditions of agriculture. Manipulation of crop plants by conventional breeding has successfully introduced characteristics such as large seed size, which were not present, and would not be viable, in their wild progenitors. Characteristics introduced into cultivated plants by plant genetic engineering do result from a process that is fundamentally different from selection, but both conventional breeding and genetic engineering are aiming for the same end results, agriculturally desirable phenotypes. Their products should be evaluated by similar criteria.

Developments to “First-Generation” Crops Expressing Bt Toxins

Plants Expressing Multiple Toxins (“Pyramiding ”)

The specificity of a single Cry toxin toward specific target pests can be a problem in the field where a secondary, minor pest species can replace the primary pest and cause serious damage to crops. An obvious method to counter this problem is to add or introduce a second Bt cry gene into the crop to extend the range of pests against which protection is afforded. The availability of a wide range of gene constructs encoding Cry toxins has made this a realistic possibility, with crossing singly transformed lines, or repeated transformation, or transformation with a construct containing two genes as alternative methods for introducing the genes into one line. Monsanto’s Bollgard transgenic cotton was improved by introducing a second Bt gene as early as 1999. Laboratory trials showed that cotton plants expressing both Cry1Ac and Cry2Ab proteins were more toxic to bollworms (Helicoverpa zea) and two species of armyworms (S. frugiperda and S. exigua) than cotton expressing Cry1Ac alone, even though doses in this trial were sublethal [100]. Subsequent evaluations in greenhouse and field trials [101] confirmed the superior insect resistance of plants expressing both toxins.

A further potential advantage of transgenic plants expressing two Cry proteins with differing specificities, that target different receptors in the insect, is in preventing the appearance of resistance in the pest, since multiple mutations are required to produce the loss of sensitivity to the toxins. This hypothesis was confirmed directly in work reported by [102], in which transgenic broccoli plants expressing either Cry1Ac, or Cry1C, or both proteins were produced. Plants were exposed to a population of diamondback moth (P. xylostella) which carried Bt resistance genes at a relatively low frequency in an extended greenhouse experiment, and results showed that selection over 24 generations led to a significant delay in the appearance of resistance in insects exposed to the pyramided two-gene plants. The success of these experiments has led to suggestions that the refuge approach to resistance management may be redundant for crops expressing multiple toxins [103]. However, some care is needed in the selection of genes in relation to potential pests, as resistance to multiple toxins has been observed in several cases. For example, a strain of the lepidopteran cotton pest H. virescens which has simultaneous resistance to Cry1Ac and Cry2Aa has been identified, in which the genetic bases of resistance to each toxin are different [104].

Many subsequent programs which have aimed to produce insect-resistant crops expressing Bt toxins have adopted the two-gene approach to broaden and improve protection against diverse pests, and to prevent resistance developing in insects (e.g., [105]). Although engineering to produce combinations of different three-domain Cry toxins is the most common approach, other potential resistance genes have been included also, such as those encoding Vip proteins [106], or even proteinase inhibitors (e.g., cowpea trypsin inhibitor; [107]). The “pyramiding ” or “stacking ” of resistance transgenes has been enthusiastically adopted by commercial organizations, and the recent announcement of a transgenic maize variety containing eight different transgenes by Monsanto and Dow Agrosciences [108] exemplifies this trend. This variety contains insect-resistance genes derived from both companies’ research programs, active against corn rootworm and lepidopteran pests (Herculex RW = Cry34Ab1 + Cry35Ab1, Herculex I = Cry1F; YieldGard VT Rootworm/RR2 = modified Cry3Bb1, YieldGard VT PRO = Cry1A.105 + Cry2Ab2), as well as two herbicide tolerance genes (giving resistance against glyphosate and glufosinate-ammonium), and is intended to be a “one-stop” solution to pest and weed problems.

Domain Exchange in Three-Domain Cry Toxins

The separate roles played by the different domains in the process of interaction of three-domain Bt toxins with their receptors, and their structural independence, suggested to investigators that hybrid toxins, in which domains from different naturally occurring toxins were grafted together, would be likely to be active, and could show novel specificities in their activity toward insects. This process can be made to occur in vivo in Bacillus thuringiensis, using a site-specific recombination vector [109], or can be carried out in vitro using conventional molecular biology techniques, followed by expression in a microbial host. Transfer of domain III between different Cry1 proteins led to identification of this domain as conferring primary specificity to different lepidopteran species, and the generation of hybrids with broader specificity than naturally occurring toxins [110]. Subsequent work generated a Cry1Ab-Cry1C hybrid, which was highly toxic to S. exigua, an insect resistant to Cry1A toxins [111], and identified Cry1Ca domain III as sufficient to confer toxicity toward Spodoptera in a variety of hybrids [112]. In contrast to the results obtained when exchanging domain III, exchange of domain I between different Cry1 toxins did not yield biologically active proteins [113].

A measure of the potential for improvement in “natural” Bt toxins is shown by experiments reported by [114], in which a hybrid Cry protein, constructed by fusing domains I and III from Cry1Ba with domain II of Cry1Ia, was expressed in transgenic potato. Plants expressing the hybrid toxin at levels up to 0.3% of total soluble protein were produced, and not only showed resistance to the lepidopteran pest potato tuber moth (Phthorimaea operculella), but also had a high level of resistance to Colorado potato beetle. The “parental” Cry proteins have high toxicity towards lepidopterans, but only very limited toxicity towards coleopterans such as the potato beetle. The hybrid has effectively created a novel toxicity, which is suggested to be based on interaction with a novel receptor.

Mutagenesis of Three-Domain Cry Toxins

Modification of Bt toxins by site-directed mutagenesis to increase toxicity towards target pests has been employed as an alternative to the “domain swap” approach. Most mutagenesis experiments on Bt toxins have been carried out to explore structure-function relationships in these proteins (see above; reviewed by Dean et al. [115]), but the accumulated knowledge of which parts of the protein determine specificity of interactions with receptors in the insect have been exploited to produce variants with increased activity toward target pests.

The key role of domain II in three-domain Cry proteins in mediating interactions with insect receptors was shown by a mutagenesis experiment in which altering amino acid residues in the loop regions in this domain of Cry1Ab increased its toxicity toward larvae of gypsy moth (Lymantria dispar) by up to 40-fold, with a corresponding increase in binding affinity to brush border membrane vesicles [116]. These results were based on expression of the recombinant protein in microbial hosts. A similar strategy was used to increase the toxicity of Cry3A protein toward target coleopteran pests [117], and of Cry4Ba toxin [118, 119] and Cry19Aa toxin [120] toward mosquito larvae. The level to which rational design of toxins is possible is shown by the engineering of toxicity toward mosquito larvae into the lepidopteran-specific toxin Cry1Aa [121]. Alternatively, a directed evolution system based on phage display technology for producing toxins with improved binding to a receptor, and thus increased toxicity, has been described [122]. Mutagenesis of domain I has also been attempted, with claims that alteration of alpha helix 7 in Cry1Ac to resemble the corresponding helix in diphtheria toxin led to increased toxicity toward cotton bollworm (Helicoverpa armigera) larvae [123].

The impressive achievements of toxin engineering at the level of recombinant proteins, have led to the technology being used for gene constructs designed for expression in transgenic plants, although toxins with unmodified amino acid sequences continue to be widely used (largely as they give adequate protection). One example where toxin engineering has been successfully carried out is the current commercial transgenic corn variety with resistance to corn rootworm, MON863, which expresses a modified version of the Bt Cry3Bb1 toxin [75]. Unmodified Cry3Bb1 is active against a number of coleopteran species, including Colorado potato beetle and corn rootworm [124], but toxicity toward western corn rootworm (Diabrotica virgifera virgifera) was not sufficient to give adequate protection at levels of expression achievable in corn. Modifications to the amino acid sequence increased the toxicity of the protein toward corn rootworm approximately eightfold. The nature of the modifications has not been described in the scientific literature, and is only available through reference to a series of patents (see [75]).

Fusions

As a logical extension to the transformation of plants with separate gene constructs encoding two Cry proteins, some workers have chosen to produce a single construct containing a single translationally fused coding sequence encoding both proteins. This approach has been successfully demonstrated by producing a Cry1Ab-Cry1B translational fusion protein in transgenic maize [125], although there is no apparent advantage over simpler methods for introducing two genes. The Cry1Ab-Cry1B fusion protein has also been expressed in transgenic rice [126], which was fully resistant to yellow stem borer (Scirpophaga incertulas).

A more interesting possibility is the introduction of extra functionality into Cry toxins by addition of sequences from other proteins which could lead to binding interactions with more potential receptors in the insect gut, extending the range of toxicity and hindering development of resistance. In work reported by Mehlo et al. [127], the galactose-binding lectin domain (B-chain) from the ribosome-inactivating protein ricin was fused C-terminally to domain III of Cry1Ac, producing a Cry1Ac-ricin B-chain fusion protein. The fusion protein thus has the ability to bind to galactose residues in side chains of glycoproteins or glycolipids in the insect gut epithelium, as well as N-acetyl galactosamine residues which are bound by domain III. The fusion protein was expressed in transgenic maize and rice plants, and was shown to afford a high level of protection to larvae of stemborers (Chilo suppressalis) and leaf armyworm (Spodoptera littoralis), whereas plants expressing the unmodified Cry1Ac were susceptible to both insects. The transgenic maize plants were also resistant to a homopteran plant pest, the leafhopper Cicadulina mbila, although it is possible that this was an effect of the lectin domain in the fusion (see later section Lectins), since Bt toxins are not effective against homopteran insects.

The engineering of extended binding properties into three-domain Cry proteins to increase the range of toxicity toward insect pests is clearly possible, but needs to be approached with some caution. There is a risk that the extended range of activity will include mammalian toxicity, which would negate one of the major advantages of these insecticidal proteins.

Exploitation of Endogenous Plant Defensive Mechanisms Against Insect Herbivores

Plants have a range of endogenous mechanisms to defend themselves against insect herbivores, and use both static defense mechanisms based on the accumulation of pre-synthesized insecticidal compounds, and active defense mechanisms in which gene expression is induced as a result of insect damage (response to wounding, and responses to insect secretions), leading to the synthesis of insecticidal compounds [128]. Conventional breeding has sought to exploit endogenous insecticidal genes within a plant species, but the use of transgenic technology allows defensive compounds and mechanisms to be transferred between species, or allows the control of existing defensive systems to be altered to improve their effectiveness. The molecular biology involved in transfer of genes between plant species is technically straightforward, and does not involve the kind of reengineering necessary to make bacterial genes suitable for use in plants. This approach to increasing insect resistance in transgenic plants has almost as long a history as engineering for Bt Cry toxin expression, but to date has not resulted in a commercial product, or widescale adoption in agriculture. Some of the reasons for the lack of practical outcomes for this strategy will be discussed below.

Proteinase Inhibitors

Protein proteinase inhibitors (PIs) are ubiquitous in plant species. They are major components of both “static” and “active” defense in that they are accumulated in specific tissues (“static” defense), and are the major end-product in the induced response to wounding (“active” defense). They are generally small proteins, ranging in size from 4 to 25 kDa, with many different sequence families having been identified. They form tightly bound complexes with their target proteinases, which usually involve a “loop” on the inhibitor fitting into the enzyme active site (Fig. 4), blocking the site, and inactivating the enzyme. The observation that most of these inhibitors were active against digestive serine proteinases from higher animals, and not endogenous plant proteinases (where serine proteinases are comparatively rare, and not involved in protein digestion) suggested that they were defensive compounds, and bioassays in which purified PIs were fed in artificial diet confirmed that an antimetabolic effect was exerted on insect herbivores which relied on protein digestion for nitrogen supply, shown as a slower growth rate, retarded development, and increased mortality (reviewed by Garcia-Olmedo et al. and Ryan [130, 131]). Besides a direct effect on digestion of ingested proteins, PIs cause a loss of nitrogen to the insect by preventing the reabsorption of nitrogen used to produce digestive proteinases , which are normally (self)-degraded in the gut rather than excreted. The role of these proteins in induced defense against insects was shown by blocking the normal wounding response in transgenic tobacco plants by suppression of expression of the prosystemin gene, which produces the peptide hormone systemin , using antisense RNA. The transformed plants were unable to synthesize wound-induced PIs and were significantly more susceptible to herbivory by lepidopteran larvae [132]. The importance of the wounding response to plant defense in natural ecosystems has been extensively studied by Baldwin’s group (reviewed in [133]); this outstanding body of work has established a synthesis of responses in the plant under attack, responses in neighboring plants, and responses of natural enemies of insect herbivores , with communication via volatile signals produced by the plant under attack.

**Genetic Engineering of Crops for Insect Resistance, Figure 4**

A seed-expressed Bowman-Birk-type serine proteinase inhibitor from cowpea, which contained two inhibitory sites active against bovine trypsin (CpTI) was the first plant PI to be produced in another species [134], using a gene construct containing a CaMV 35S promoter. The resulting transgenic tobacco plants expressed CpTI at up to 1.0% of total soluble protein, and decreased growth and survival of tobacco budworm (H. virescens) by up to 50%, with similar effects on other lepidopteran larvae. Subsequent experiments carried out with wound-induced PIs showed that these also had similar effects when constitutively expressed in transgenic plants; for example, the tomato inhibitor II gene, when expressed in tobacco, was also shown to confer insect resistance [135], as did potato PI-II [136]. Both CpTi and PI-II were subsequently expressed in rice, where partial protection against stem borers was observed [137, 138]. The constitutive expression of foreign PIs could be mimicked in transgenic tomato plants by constitutive expression of the prosystemin gene (see above) leading to constitutive expression of wound-induced tomato PIs [139]. Tobacco plants modified in this manner show partial resistance to insect herbivores similar to that produced by expressing foreign PIs [140].

The problem with this strategy for producing insect-resistant plants soon became obvious; in contrast to the expression of Cry proteins, which, when optimized, routinely gave transgenic plants virtually complete protection against susceptible pests (mortality100%, damage minimal) expression of PIs only produced partial resistance. Investigation of the digestive biochemistry showed that exposure to PIs in the diets of lepidopteran and coleopteran herbivores resulted in the appearance of proteinase activities which were insensitive to the inhibitor(s) present [141, 142], or were able to degrade the ingested PIs [143]. These insects contain large families of genes encoding dietary proteinases, whose expression could be up- or downregulated by dietary inhibitors [144]. In effect, these insect herbivores were preadapted to be partially resistant to dietary PIs, as a result of similar or identical compounds being present routinely in their diet. Although expression of resistance to PIs in herbivorous insects has a fitness penalty, shown by reduced growth on diets to which inhibitors are added, or on plants which are expressing foreign PIs, or over-expressing endogenous PIs (see above), this is not sufficient to cause mortality at a level which affords more than partial protection. In some cases, low levels of expression of a foreign PI in transgenic plants can actually result in improved insect performance, as when tobacco and Arabidopsis plants expressing mustard trypsin inhibitor 2 were exposed to larvae of cotton worm (S. littoralis; [145]).

A number of investigators have attempted to select PIs for expression in transgenic plants which are optimally active against the dietary proteinases present in specific insect pests. Attempts to develop inhibitors active against specific lepidopteran digestive serine proteinases induced by dietary PIs have not been successful. On the other hand, not all pest insects rely on serine proteinases for digestion. Many herbivorous coleopteran larvae utilize cysteine proteinases, rather than serine proteinases , as their major digestive endoproteinases , and these proteinases can be inhibited by cystatins, a family of proteins present in all kingdoms of organisms. Enzyme assays in vitro were used to characterize digestive proteinases of a coleopteran pest, Chrysomela tremulae, as cysteine proteinases, and to show that a cystatin from rice, oryzacystatin, was an effective inhibitor. Transgenic poplar seedlings expressing oryzacystatin were produced, and leaves from these plants were shown to be toxic to larvae of the pest [146]. This promising result does not seem to have been followed up. Expression of oryzacystatin in transgenic potato only gave partial protection against larvae of Colorado potato beetle [147], suggesting preadaptation in this pest, which is known to employ a diverse range of digestive proteinases. In an attempt to use proteinase inhibitors which insects would not be preadapted to, synthetic multi-domain cysteine proteinase inhibitors based on domains found in animal and plant sources (kininogen, stefin, cystatin C, potato cystatin, and equistatin) were assembled and expressed in transgenic potato; the plants were deterrent to thrips, and gave partial resistance in greenhouse trials, but complete protection was not observed [148, 149]. Attempts to express the sea anemone cysteine/aspartic proteinase inhibitor equistatin itself in transgenic potato did not give significant levels of resistance to Colorado potato beetle, due to degradation of the inhibitor in the plant [150]. Multiple proteinase inhibitors (potato PI-II and PCI) active against two families of proteinases, serine proteinases and carboxypeptidases, have been expressed in transgenic tomato plants [151], but still only afforded partial protection against lepidopteran larvae due to adaptive mechanisms present in the insects.

In conclusion, the expression of suitable PIs in transgenic plants can give protection against lepidopteran and coleopteran pests, but has not been able to produce results comparable with those achieved by use of Bt toxins.

Amylase Inhibitors

The widespread occurrence of protein inhibitors of mammalian amylases in plants has become accepted as another defensive mechanism against herbivores (reviewed by Franco et al. [152]). Like proteinase inhibitors, these are generally small proteins, resistant to proteolysis, ranging in size from approx. 8–30 kDa. Although they are also active against insect amylases , it is not clear to what extent these proteins contribute to insect resistance in most cases, since the relatively low nitrogen content of plant tissues compared to insects means that most herbivorous insects are nitrogen limited, not carbon limited, and starch digestion is unlikely to be a limiting factor in growth. However, in the case of coleopteran herbivores whose larvae attack seeds specifically, such as seed weevils (bruchids), there is good evidence for α–amylase inhibitors from legume seeds being highly insecticidal [153], and in being causative factors in the resistance of specific varieties of legumes to bruchids [154]. These proteins belong to a different sequence family than the more common types of α–amylase inhibitors found in cereals, and are similar to legume lectins in sequence [155].

Like proteinase inhibitors, amylase inhibitors form tightly bound complexes with their target amylase (Fig. 5), although the same interaction of a loop on the inhibitor with the active site of the enzyme is not possible, since the enzyme substrate is a polysaccharide, not a polypeptide. The mechanism of toxicity clearly involves inhibition of starch digestion, since bruchid larvae exposed to the α–amylase inhibitor from French bean (Phaseolus vulgaris) show induction of amylase enzymes [157], although other mechanisms of toxicity may also be present, since these proteins can cause 100% mortality in susceptible insect species at levels of <1.0% of total protein. Alternatively, these highly specialized herbivores may lack the adaptive mechanisms to plant defensive proteins shown by species that feed on a wide range of plant foodstuffs [128]. High levels of toxicity toward insects have not been observed in general with amylase inhibitors. For example, α–amylase inhibitors are not strongly toxic to lepidopteran larvae, where the alkaline environment of the gut may interfere with the formation of inhibitor-enzyme complexes. The α–amylase inhibitor from French bean is inactivated by high pH.

**Genetic Engineering of Crops for Insect Resistance, Figure 5**

The isolation of a lectin-like α–amylase inhibitor gene from P. vulgaris [155] stimulated research in this area, and in a ground-breaking series of experiments, this gene was assembled into a construct with a strong seed-specific promoter (from the P. vulgaris seed lectin gene), and expressed in seeds of transgenic garden pea. The resulting seeds contained up to 3% of the foreign protein, and were highly resistant to larvae of cowpea and Azuki bean weevils [158], which do not normally attack garden peas in the field, but are stored product pests, and to larvae of the pea weevil Bruchus pisorum [159], which is a field pest of garden pea. In all cases larval development from eggs laid on seeds was halted at a very early stage, and damage to the crop was minimal. Subsequent experiments showed that transgenic azuki beans could also be protected against bruchid storage pests [160], and that transgenic garden pea was protected against pea weevil under field conditions [161]. The success of this strategy led to hopes that the Phaseolus α–amylase inhibitor gene could be incorporated into a range of crops, particularly other grain legumes such as lentils, mungbean, groundnuts, and chickpeas to give protection against a variety of bruchids. Technical problems with transformation of some of these crop species have delayed this goal being achieved, but transgenic chickpeas expressing high levels of the Phaseolus α–amylase inhibitor have been successfully produced [162].

Despite the success of this strategy, full agricultural deployment of transgenic crops expressing the Phaseolus α–amylase inhibitor gene has not taken place. Commercial reasons have played a major part in preventing widescale adoption, but safety concerns have also arisen. The protein product of the Phaseolus α–amylase inhibitor gene expressed in pea shows minor structural differences to the native product (i.e., expressed in P. vulgaris) as a result of differences in posttranslational processing (differences in the extent of glycosylation, and in minor components resulting from proteolysis). Whereas consumption of the native form of the Phaseolus α–amylase inhibitor by mice did not result in immunological responses, consumption of transgenic peas expressing this protein led to the presence of circulating antibodies directed against it, and systemic immunological responses including inflammatory responses (i.e., allergic responses) to inhaled or injected protein [163]. In contrast to some earlier work claiming that consumption of transgenic plant material was harmful, this study has been published in a fully peer-reviewed journal and the quality of the research has not been disputed. Further research will be necessary to identify, and remove, the cause of this increased antigenicity. An additional potential drawback was revealed by feeding trials of transgenic peas expressing Phaseolus α–amylase inhibitor with pigs and chickens. These trials did not show immunological effects on animal health, but did show that starch utilization by the animals was significantly decreased due to the presence of the inhibitor in the transgenic peas when compared to non-transgenic peas, consistent with the effect of the protein on higher animal amylases [164, 165]. This factor would limit the utility of transgenic peas as animal feed.

Lectins

Lectins , or carbohydrate-binding proteins , occur throughout the plant kingdom, and in many species are accumulated in plant tissues as defensive proteins, being particularly abundant in seeds and other storage tissues, where they can account for up to 1% or more of total protein (reviewed by Peumans and van Damme and van Damme et al. [166, 167]). They are multimeric proteins containing polypeptides which range from 10 to 35 kDa in size. The insecticidal activity of lectins was first observed in assays with larvae of coleopteran species (e.g., LE QA done [168, 169]), where retardation of development, and in some cases, mortality, was observed when lectins were incorporated into diets at 1–5% of total protein. Lectins have only relatively low antimetabolic effects on lepidopteran larvae when fed in diet [170], possibly as a result of high gut pH inactivating the carbohydrate-binding activity. The mechanism of toxicity of these proteins remains obscure, but is dependent on carbohydrate binding.

Although transgenic tobacco and potato plants expressing lectins from garden pea [171] and snowdrop [172] have been produced by standard transformation techniques, and have been shown to confer partial resistance to lepidopteran larvae (>50% reductions in plant damage, with increased larval mortality and decreased growth), the availability of better insecticidal genes specific for these pests has directed this approach toward different targets. Homopteran plant pests , which are not affected by known Bt toxins, were shown to be susceptible to lectin toxicity when the proteins were delivered via artificial diet [173]. Susceptibility varied between species, and between lectins, but LC₅₀ values as low as 6 μM have been estimated (for snowdrop lectin fed to rice brown planthopper (Nilaparvata lugens); [174]). Expression of the mannose-specific snowdrop lectin (Galanthus nivalis agglutinin; GNA) in transgenic rice plants was carried out, using both a phloem-specific (rice sucrose synthase) and a constitutive (maize ubiquitin-1) promoter [175]. The resulting plants were partially resistant to rice brown planthopper, with reductions of up to 50% in survival of immature insects to adulthood, and reduced development and fertility of survivors. Results were confirmed by independent transformations of indica rice varieties [176]. GNA-expressing rice was also resistant to other homopteran plant pests, such as green leafhopper (Nephotettix virescens; [177]) and whitebacked planthopper (Sogatella furcifera; [178]). Plants expressing both GNA and Cry1Ac were protected against both brown planthopper and striped stem borers (C. suppressalis), but no synergistic effects between the two insecticidal proteins was observed [179]. Further progress on this research has been limited, due to concerns about possible adverse consequences to higher animals of ingesting snowdrop lectin. While earlier data must be regarded as unreliable, a recent study found that no adverse effects of consumption of transgenic rice expressing GNA by rats, although significant differences in some parameters to a control group were observed [180]. GNA expression has also been engineered into potato [181] and maize [182], to give partial resistance to peach–potato aphid (Myzus persicae) and corn leaf aphid (Rhopalosiphum maidis), respectively. However, these insects are insensitive to lectin toxicity, and only marginal effects on fecundity were observed.

Introduction of foreign lectin genes into plants has become established as a potential method for engineering insect resistance, although with the lectins tested at best only partial protection against homopteran pests is conferred, and some species are relatively insensitive to the effects of lectins. As is the case with PIs, it is likely that plant pests are preadapted to the presence of lectins as defensive compounds, and are able to tolerate the toxic effects to varying degrees, although responses induced in insects by ingested lectins have not been characterized. Attempts have been made to select lectins which are the most effective toxins against target insect pests; a mannose-specific lectin expressed specifically in garlic leaves (ASA-L) was observed to show a high level of toxicity toward homopteran pests [183]. A gene encoding this lectin has been engineered into a variety of transgenic plant species, including tobacco [184] and Indian mustard [185], in both cases producing partial resistance to aphid species, with reduced survival and fecundity. Expression of this lectin in transgenic rice using constitutive [186] or phloem-specific promoters [187] gave protection against homopteran pests comparable to, or slightly better than, earlier transformations using gene constructs encoding GNA. The transgenic rice plants expressing ASA-L were shown to decrease transmission of Rice Tungro Virus by its insect vector (green leafhopper), presumably by causing decreased feeding by the pest [188].

Despite these encouraging results, widescale adoption of transgenic crops expressing lectins will probably not occur unless a major commercial company is able to gain exclusive marketing rights, and invests in pushing the transgenic varieties through the regulatory process. This is unlikely to happen, as the technology is not readily protectable by patenting.

Oxidative Enzymes

Induction of polyphenol oxidase (PPO) synthesis is one of the end-results of the plant wounding response [189], and it would seem reasonable to suppose that increased levels of this enzyme would lead to enhanced resistance to insect attack. PPO activity leads to tissue browning, which has been correlated with enhanced insect resistance. The oxidative cross-linking of tannins to proteins catalyzed by PPO decreases protein digestibility, and limits nitrogen availability [190]. However, there is little or no evidence that PPO levels are correlated with insect resistance (e.g., [191]). High-level, constitutive over expression of a poplar PPO gene in transgenic poplar seedlings led to levels of PPO up to 50x higher than normal in plant tissues [192], but these plants had only marginal effects on larvae of the lepidopteran insect pest forest tent caterpillar (Malacosoma disstria). No feeding deterrence was observed, and there was no effect on larval growth or survival except under conditions where larval survival was poor on controls. PPO activity was detected in insect gut and frass, so the negative results were not due to enzyme inactivation. The conclusion that herbivorous insects are preadapted to be able to deal with PPO activity, as a result of exposure to the wounding response on an evolutionary timescale (in a similar manner to preadaptation to PIs – see above) is difficult to avoid.

Peroxidase activity is also induced when plants are stressed, or attacked by pathogens, as part of a lignification response, and several attempts have been made to over-express peroxidases in transgenic plants to enhance insect resistance, despite a lack of clear-cut evidence that peroxidase activity in plant tissues is toxic to insect herbivores. Initial results using tobacco as the host plant, with over-expression of tobacco anionic peroxidase, showed only marginal effects [193], although limited broad-range protection against a variety of pests was observed in the field [194]. The limited protection afforded by this technique argues against further development.

Other Plant Proteins

Ribosome inactivating proteins (RIPs) and chitinases have also been viewed as defensive proteins in plants, although it is not clear that they are part of defense against insect herbivores. Both types of proteins have been expressed in transgenic plants, with variable results in conferring insect resistance. Expression of a maize RIP in transgenic tobacco resulted in very low levels of protection against corn earworm (H. zea), which were barely statistically significant [195]. Plant chitinases in general show low toxicity toward insects, but a poplar chitinase, designated WIN6, was selected on the basis that its expression was induced by insect attack. Expression of WIN6 in trangenic tomato plants led to partial protection against larvae of Colorado potato beetle, with retardation of larval development observed [196]. Expression of the chitin-degrading enzyme N-acetylhexosaminidase from Arabidopsis in various transgenic plant tissues also gave some protective effects against lepidopteran larvae [197], but it is difficult to see what advantages over other strategies this approach could give. Orally ingested insect chitinases are strongly toxic to lepidopteran larvae (e.g., [198]). However, expression in transgenic plants gave only partial protection against insect herbivores [199], or, in one case, increased susceptibility to attack [200]. Expression of chitinase A from baculovirus AcMNPV in transgenic tobacco gave similar results, with only small effects on lepidopteran larvae and aphids [201].

Engineering Secondary Metabolism for Plant Defense

Compounds synthesized as the end-products of secondary metabolism play major roles in both constitutive and induced defense against insect herbivores in many plant species (reviewed by Wittstock and Gershenon [202]). The idea that these compounds could be used as insecticides has been a part of agriculture for thousands of years, and has been exploited successfully by synthetic chemistry in the production of classes of insecticides such as pyrethroids, based on terpenoid esters produced by flowers of pyrethrums (Chrysanthemums). Although the concept of synthesizing a foreign, insecticidal secondary metabolite in a transgenic plant developed concurrently with plant transformation technology, the biosynthesis of most secondary compounds was poorly understood, and the necessity of cloning and introducing a series of genes expressing biosynthetic enzymes to produce a secondary metabolite was considered beyond the techniques available at the time. Anticipation of problems in ensuring controlled co-expression of a series of biosynthetic genes has proved to be over-pessimistic, and plants containing multiple expressing transgenes have been produced without difficulty.

The explosion of knowledge brought about by large-scale cDNA sequencing programs and the Arabidopsis genome program has resulted in a much better understanding of secondary metabolism, with many biosynthetic pathways now reasonably well understood, and clones encoding biosynthetic enzymes available. The first successful demonstration that a foreign secondary compound could confer insect resistance in a transgenic plant [203] exploited a biosynthetic pathway for cyanogenic glycosides. The cereal Sorghum bicolor produces a cyanogenic glycoside , dhurrin, by a biosynthetic pathway starting from the amino acid tyrosine, a product of primary metabolism. Two oxidation reactions catalyzed by cytochrome P450 oxidases generate p-hydroxymandelonitrile, which is then glycosylated by a UDP-glycosyltransferase to form dhurrin. The three sorghum enzymes responsible were cloned and assembled into expression constructs using constitutive (CaMV 35S) promoters [204], and Arabidopsis plants were successively transformed with a construct containing both P450 oxidase sequences, and the glycosyl transferase sequence. All the enzymes were localized correctly (to endoplasmic reticulum membranes) and functioned properly. Surprisingly, little disruption to endogenous metabolism was observed in the transgenic plants expressing medium levels of dhurrin, and accumulation of pathway intermediates was not observed. The implication is that the plastic nature of plant metabolism can accommodate and regulate activity in new biosynthetic pathways that are introduced. The resulting plants included individuals producing levels of dhurrin similar to sorghum plants in leaf tissue (up to 4 mg/g fresh weight) and produced hydrogen cyanide on tissue damage (due to the hydrolysis of dhurrin by an endogenous Arabidopsis enzyme). The dhurrin-expressing plants showed enhanced resistance to attack by the flea beetle Phyllotreta nemorum, a specialist feeder on crucifers; adult beetles avoided feeding on dhurrin-expressing leaves when offered a choice, and larvae under no-choice conditions either failed to initiate feeding, or on initiating feeding showed a significant level of mortality. These initial results clearly imply that production of high levels of dhurrin in transgenic Arabidopsis caused phenotypic abnormalities, but subsequent refining of the technology allowed accumulation of dhurrin at up to 4% dry weight in Arabidopsis tissues without deleterious effects on plant growth [205]; expression levels of the UPD-glycosyl transferase must be high enough to prevent accumulation of the p-hydroxymandelonitrile intermediate.

Although these results represent science of the highest quality, this method is of marginal usefulness for crop protection as it stands, due to the dhurrin end product being toxic to higher organisms, due to the production of hydrogen cyanide when it is hydrolyzed. Worse, many insect herbivores, particularly those which have a polyphagous feeding habit, can detoxify cyanide [206]. However, the feasibility of engineering secondary metabolism in crop plants has now been established. Expression of the cassava cyanogenic glycosides, linamarin and lotaustralin (derived from valine and isoleucine respectively), has also been achieved in Arabidopsis [207], and grape vine root cultures have been engineered to produce dhurrin [208], although in this case no protection against root aphids was observed. Other types of secondary metabolites have also been exploited; production of the alkaloid caffeine from its precursor xanthosine in tobacco was achieved by the introduction of three genes encoding N-methyl transferases [209]. The resulting plants contained up to 5 μg/g fresh weight caffeine in leaves, and showed a strong feeding deterrent effect toward a generalist lepidopteran herbivore, Spodoptera litura. An alternative approach to modifying secondary metabolism was taken by [210], who introduced a gene encoding β-glucosidase from Aspergillus niger into tobacco plants, and demonstrated that transgenic plants expressing the enzyme had insecticidal activity toward whiteflies (Bemisia spp.) and dipterans (flies), putatively due to hydrolysis of unidentified glycosides in the plant (although the greater density of secretory trichomes observed in transgenic plants may also have been significant). Further developments in this area can be expected.

Besides engineering, secondary metabolism to produce defensive compounds normally present in other plant species, the biosynthetic capacity of plants can be used to produce a variety of volatile secondary compounds used for communication. Better understanding of the terpenoid biosynthesis pathways has led to the production of a number of transgenic plants with altered volatile composition (reviewed by Aharoni et al. [211]). Suppression of expression of a cytochrome P450 oxidase gene expressed in trichomes by RNAi led to transgenic tobacco plants which deterred aphid colonization [212], due to the final step in production of the diterpenoid cembratriene-diol being blocked, resulting in accumulation of the precursor, cembratriene-ol. These compounds are both volatile and components of trichome secretions. Transgenic Arabidopsis plants constitutively over-expressing a dual linalool/nerolidol synthase in plastids produced significant amounts of linalool , both as a free alcohol (volatile) and as glycosylated derivatives, and were repellent to aphids (M. persicae) when tested in a choice experiment [213]. Modifications to isoprenoid synthesis in Arabidopsis have also been shown to attract predatory mites, which could protect plants by destroying pests [214]. This strategy of attracting natural enemies to pests has also been exemplified by transforming Arabidopsis with the maize terpene synthase gene TPS10, which is responsible for producing sesquiterpene volatiles emitted by maize. The resulting plants emitted the volatiles normally produced in maize and attracted parasitoid wasps which attack maize pests [215]. A different approach to utilizing terpene production in transgenic plants exploits the activity of the sesquiterpene (E)-β-farnesene as an alarm pheromone in aphids, which causes cessation of feeding and avoidance, as well as acting as an attractant for aphid predators and parasitoids [216]. Arabidopsis was transformed with an (E)-β-farnesene synthase gene from mint, under control of a constitutive promoter (CaMV 35S); resulting plants produced (E)-β-farnesene as a volatile. The transgenic plants showed significant levels of aphid deterrence in choice experiments, and were attractive to the aphid parasitoid Diaeretiella rapae. Experiments which engineer the volatiles emitted by plants are an exciting area of research at present, which has established the role that volatiles emitted by plants play in the interactions between plants, herbivores, and natural enemies at the tritrophic level. This technology has yet to show that it is a practical method for crop protection in the field, but practical applications look likely to follow.

Some Novel Approaches

Many other approaches to engineering insect resistance in transgenic plants have been proposed, and progressed to varying degrees. The following section gives an overview of some of the most promising of these approaches, which have been taken forward to the stage of demonstrating feasibility by producing insect-resistant plants. Of necessity, many other interesting ideas have had to be omitted, such as transformation of plants with transcription factors which alter gene expression [217, 218], or the use of transgenic plants expressing potentially toxic proteins from insects [219] or insect peptide hormones [220]. Despite the lack of commercial deployment of any of the insect-resistant transgenic plant other than those expressing proteins derived from Bt, this field of research is active and new approaches will continue to be put forward and evaluated.

Photorhabdus luminescens Insecticidal Proteins

Photorhabdus luminescens is an enterobacterial symbiont of entomophagous (insecticidal) nematodes of Heterorhabditis species, used for small-scale biological control of insect pests. The bacteria are present in the nematode gut, and when nematodes enter an insect host, bacterial cells are released into the insect circulatory system. The bacterial cells release toxins which cause cell death, leading to a lethal septicemia, providing a substrate for both bacteria and nematodes to grow on [221, 222]. The toxins are present as high-molecular-weight (M_r approx. 10⁶) complexes, which are toxic when injected or fed to insects from four major orders of agricultural pests. The complex has been separated into four components, encoded by genetic loci tca, tcb, tcc, and tcd; the products of tca and tcd are toxic individually when fed to lepidopteran larvae. The mechanism of action of the toxins remains unresolved. Subsequent investigation has shown that Photorhabdus contains a large number of potentially insecticidal components, some of which are only toxic by injection, whereas others are orally toxic (reviewed by ffrench-Constant [223]); a variety of mechanisms of toxicity, including promotion of apoptosis, seems to be exploited by the bacterium. This presence of a reservoir of redundant insecticidal activities, reminiscent of the situation in Bacillus thuringiensis , led to Photorhabdus being put forward as a successor to Bt as a source of insecticidal genes for expression in transgenic plants.

In order to be able to exploit insecticidal genes, investigators have sought to isolate single toxic proteins from Photorhabdus. Two proteins, designated toxin A and toxin B, were isolated from culture supernatant and shown to be orally toxic [224]. They exist as high-molecular-weight complexes (approx. 860 kDa) in solution, and each consist of two polypeptides, 201 and 63 kDa molecular weight. The mature polypeptides are produced from single precursor protoxin polypeptides of 283 kDa by proteolysis by endogenous bacterial proteinases. The 283 kDa protoxin A is the product of a gene designated tcdA in Photorhabdus, which has been cloned and assembled into expression constructs for use in transgenic plants. Expression levels of mRNA and protein were improved by adding 5′ and 3′ UTR sequences from a tobacco osmotin gene, but the coding sequence was not reengineered. Expression in transgenic Arabidopsis gave plants that contained intact protoxin, with a range of expression levels [225]; expression of toxin A at levels above 0.07% of total soluble protein in leaves gave almost complete protection against larvae of the lepidopteran tobacco hornworm (M. sexta). The toxin is not species specific, and leaf extracts were also toxic to the coleopteran corn rootworm (Diabrotica undecimpunctata). Commercial development of this technique is highly likely.

Entomophagous nematodes of Steinernema species also contain mutualistic bacteria, of Xenorhabdus species, which produce insecticidal toxins. These proteins could also be exploited to produce insect resistance in transgenic plants, but have not yet received as much attention as Photorhabdus toxins [223].

Cholesterol Oxidase

The identification of a protein from Streptomyces that was highly insecticidal to larvae of the coleopteran pest cotton boll weevil (Anthonomus grandis) resulted from a screening program assaying culture filtrates of different bacterial species [226]. The protein, which was toxic at levels comparable to a Bt three-domain Cry protein, was identified as a cholesterol oxidase. It was able to lyse the midgut epithelium in the insect. The mechanism of action involves the activity of the enzyme, since no activity is seen in lepidopteran larvae where the gut pH is high, and the enzyme has low activity, but may also involve effects on membrane-bound alkaline phosphatase [227]. Oxidation of membrane sterols such as cholesterol in the insect gut epithelium can destabilize membranes, leading to cell lysis as observed. However, expression of this protein in transgenic plants could prove problematic, since it is equally capable of oxidizing sterols in plant cell membranes. The encoding gene for the cholesterol oxidase was isolated, and assembled into expression constructs containing either the complete coding sequence, the mature protein coding sequence, or the coding sequence fused to a chloroplast targeting peptide from the Arabidopsis ribulose bisphosphate carboxylase (RuBisCO) small subunit gene [228]. No codon optimization was carried out. Transgenic tobacco plants were produced by transformation of the nuclear genome, and all constructs were shown to result in synthesis and accumulation of active enzyme. The constructs which omitted the chloroplast targeting peptide caused protein to accumulate in the cytoplasm, and these plants were developmentally abnormal, possibly as a result of interference with plant sterol hormone signaling pathways. Plants in which the enzyme was localized in chloroplasts were phenotypically normal. Leaf tissue from all transgenic plants was toxic to boll weevil larvae when fed as a component of an artificial diet.

This work does not seem to have been progressed beyond the stage of a demonstration of concept, and no further references to it are present in the scientific literature. This gene would seem a good candidate for introduction into the chloroplast genome to engineer insect resistance, although potential effects on chloroplast membrane systems would remain a drawback.

Avidin as an Insecticidal Protein

Exploitation of the biotin-binding properties of the avian egg white protein avidin (and its bacterial functional homologue, streptavidin ) in a variety of biochemical techniques has obscured its role as a defensive protein, which is toxic to bacteria. The antibacterial activity is based on its essentially irreversible binding of biotin, leading to this essential enzyme cofactor being unavailable. The insecticidal activity of avidin was recognized in 1993, when assays carried out in artificial diet showed toxicity to coleopteran and lepidopteran larvae at levels as low as 10 ppm in diet (estimated as of the order of 0.01% of total protein), although the level necessary to show toxicity was up to 100x higher for other pest species. The toxic effect was eliminated by addition of biotin to diets, suggesting that the mechanism of avidin insecticidal activity is also through biotin sequestration. Both growth reduction and mortality were observed, and the suggestion was made that gene constructs expressing avidin could provide protection against insect pests in transgenic plants [229]. Subsequent assays confirmed that susceptibility to avidin as an insecticide varies widely between different insect species, and that biotin carried over in the egg between generations had a significant effect on subsequent avidin toxicity [230].

Initial reports of expression of avidin in transgenic maize were focused on producing the protein as a high-value product [231]. An expression construct containing a codon-optimized avidin coding sequence with an N-terminally fused signal peptide from barley α-amylase, driven by the maize ubiquitin-1 promoter, resulted in expression levels of avidin of >2.0% of total protein in seed. Seed from these plants was subsequently bioassayed for resistance to larvae of three different coleopteran storage pests, including red flour beetle (Tribolium castaneum), with 100% mortality at avidin levels above 100 ppm of seed (approx. 0.1% of total protein). However, not all pests were as susceptible; larvae of the larger grain borer, Prostephanus truncatus, were effectively insensitive to avidin, whether added to artificial diet or expressed in transgenic plant material. The engineered maize was nontoxic to mice over 21 days [232]. Subsequent reports confirmed the insecticidal effects of avidin expressed in transgenic plants: these include protection of tobacco against noctuid lepidopterans [233], using vacuolar targeting sequences from potato proteinase inhibitors to direct avidin accumulation in the vacuole at levels up to 1.5% of total leaf protein [234]; protection of apple against lepidopteran pests [235]; and protection of rice against coleopteran stored grain pests, using a similar approach to that used for maize [236]. Targeting of the foreign protein to vacuolar or similar compartments is important; expression of streptavidin in tomato using plant and bacterial signal peptides and strong promoters led to developmental abnormalities in the plants, which could be corrected by topical application of biotin, suggesting that sequestration of cellular biotin is equally detrimental for plants as well as insects [237].

Despite many promising results, this technology appears to have failed to gain any acceptance for agricultural crops, as illustrated by a recent study in which seed meal from transgenic avidin-expressing maize was tested as an insecticide for topical application to stored maize [238]. Studies have shown that avidin can increase the protection afforded by Bt expression in transgenic plants against insect pests which have limited susceptibility to the toxin (e.g., potato expressing Cry3A; [239]), but it is clear that little further development in this area is taking place.

RNA Interference Using Double-Stranded RNA

Downregulation of gene expression by double-stranded RNA (dsRNA) corresponding to part or all of a specific gene transcript has been used as a research technique in insect genetics since 1998. The method has been based on delivery of synthetic dsRNA produced in vitro by injection into insect cells or tissues, which is clearly not practical for applications in crop protection. However, recent results have shown that dsRNA can be introduced into insects as a component of artificial diet, and is effective in downregulating genes normally expressed in gut tissue. This technique has been used to downregulate the production of a gut carboxylesterase in larvae of the lepidopteran Epiphyas postvittana (light brown apple moth; [240]), leading to suppression of mRNA in the insect. More significantly, two recent papers show that dsRNA can be delivered to insect pests by expression in plant material, and that this can lead to an insecticidal effect when pests are exposed to plants. Transgenic tobacco and Arabidopsis plant material expressing dsRNA directed against a cotton bollworm detoxification enzyme (cytochrome P450 gene CYP6AE14) for gossypol suppressed expression of the gene, and caused the insect to become more sensitive to gossypol in the diet, leading to reduced performance compared to controls [241]. A similar technique was used to suppress expression of a V-type ATPase in larvae of the coleopteran Diabrotica virgifera virgifera (Western corn rootworm); transgenic corn plants producing dsRNA directed against this gene showed protection against feeding damage by the insect [242]. The feasibility of using dsRNA in crop protection strategies has thus been demonstrated. This approach holds great promise for future development, as it allows a wide range of potential targets for suppression of gene expression in the insect to be exploited.

Insect-Resistant Genetically Engineered Crops and Sustainability

The success of Bt-expressing crops in the field has been a direct result of taking “sustainability ” into account in their introduction, particularly with respect to managing the emergence of pest resistance to the toxins through the refuge strategy, as described earlier. Even organizations hostile to Genetic Engineering technology, such as organic growers in the USA, have reported that Bt cotton and corn have reduced insecticide usage significantly (by up to 0.2 kg/ha/year), showing that these crops are compatible with the goals of “sustainable” agriculture [243].

The “sustainability ” of transgenic insect-resistant crops has also been examined in terms of potential effects on the wider ecosystem in which the plants are grown. Numerous studies have been carried out to effects on predators and parasites at the third trophic level, and on nontarget insects and other invertebrates. Some initial reports which did report negative effects were based on dubious assumptions, or used experimental designs which had little relevance to field conditions (e.g., the supposed threat to monarch butterflies posed by transgenic Bt corn; reviewed by Gatehouse et al. [244]). Nevertheless, it must be the case that if a pest population is decreased as a result of endogenous resistance in crops, then there will be a “knock on” effect to the wider ecosystem, and particularly to predators and parasites of the pest species, when the resistant crop is compared to a nonresistant one that is not treated with pesticide. However, this is not a realistic comparison, since in agricultural practice a crop that does not have endogenous resistance is treated with exogenous insecticides. The use of the refuge strategy allows significant pest populations to be present, and thus can support both beneficial insects which attack the pest, and a wider ecosystem, which would be destroyed by exogenous insecticide application.

Looking to the future, wider use of insect-resistant transgenic crops could contribute positively to “sustainability” in agriculture in general, by further decreasing insecticide usage and thereby decreasing energy inputs. However, the “sustainability” of the insect-resistant crops themselves is going to come under increasing pressure, as less controlled deployment of insect-resistant plants evades the present compulsory use of the refuge strategy, and use of crop varieties with multiple Bt toxins renders the refuge strategy apparently less necessary to prevent pest resistance to Bt toxins developing. Field resistance to Bt crops has been observed recently (reviewed by Tabashnik et al. [245]), but is manageable using existing practices, or modifications of them. The sustainability of relying on one mechanism of crop protection can be questioned, especially as plants in general have evolved mixed defense strategies [246]. In the longer term, a wider range of strategies for producing insect-resistant plants is going to be necessary, not only to deal with the potential for nonspecific resistance to Bt toxins, but to extend the range of crop pests that can be targeted, and further reduce the application of pesticides.

Future Directions

After 20 years, insect-resistant transgenic crops have been a greater success in some ways than the early experiments suggested, but have failed to meet all the hopes that were initially raised. The success is self-evident when the widescale adoption of the technology in certain crops such as cotton and maize is considered, and documented evidence of reductions in damage to human health and the environment as a result of decreases in the use of exogenously applied pesticides. The failure does not lie in any technical shortcomings in the science, although improvements and new strategies are always possible; it lies in a failure to disseminate the technology as widely as should have been the case, so that it remains largely in the hands of commercial organizations, and is limited to the major crops. Is it an unrealistic hope to anticipate that after another 20 years, amateur gardeners in developed countries will be able to choose to buy seed to grow genetically engineered cabbages, which will be resistant to cabbage white butterfly larvae, in their allotments and gardens? Or that rural farmers in developing countries will have free access to engineered rice varieties, suitable for their growth conditions, that are resistant to pests such as stemborers? Both these aims have been scientifically achievable for at least the last 10 years, and it is surely about time that a more rational approach, which cuts through both the largely futile debate about the rights and wrongs of plant genetic engineering, and the protectionism of agrochemical companies, was taken to address the looming problem of producing enough crops to meet humanity’s needs.

Abbreviations

Coding sequence:: The part of a gene which determines the sequence of the protein product
Domain:: A region of a protein which forms a distinct 3-D structure, and will often form this structure even when separated from the rest of the protein
Genetic engineering:: Introduction of a specific DNA sequence into an organism by artificial means
Insect orders:: Lepidoptera = butterflies and moths; diptera = flies; coleoptera = beetles; hemiptera/homoptera = sucking insects such as aphids
Mutagenesis:: Alteration to a DNA sequence, often resulting in alteration to the sequence of a protein which the DNA specifies
Oligomerization:: Formation of polymers containing a relatively low number of repeating units
Proteolysis:: Introduction of breaks in the chain of amino acids making up a protein by a proteinase
Transgenic:: Organism into which a gene has been introduced by genetic engineering technology

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School of Biological and Biomedical Sciences, Durham University, South Road, Durham, UK
John A. Gatehouse

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John A. Gatehouse
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Correspondence to John A. Gatehouse .

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Department de Produccio Vegetal i Ciencia Forestal, Universitat de Lleida/ICREA, Lleida, Spain
Paul Christou
Institució Catalana de Recerca i Estudis Avançats (ICREA), Barcelona, Spain
Paul Christou
Department of Crop and Forest Sciences, University of Lleida, Lleida, Spain
Roxana Savin
University of New England, Marine Science Center, Biddeford, ME, USA
Barry A. Costa-Pierce
Department of Animal and Dairy Science Breeding and Genetics, University of Georgia, Athens, GA, USA
Ignacy Misztal
The Roslin Institute and Royal (Dick) School of Veterinary Studies, Division of Developmental Biology, University of Edinburgh, Midlothian, Scotland, UK
C. Bruce A. Whitelaw

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Gatehouse, J.A. (2013). Genetic Engineering of Crops for Insect Resistance . In: Christou, P., Savin, R., Costa-Pierce, B.A., Misztal, I., Whitelaw, C.B.A. (eds) Sustainable Food Production. Springer, New York, NY. https://doi.org/10.1007/978-1-4614-5797-8_239

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DOI: https://doi.org/10.1007/978-1-4614-5797-8_239
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Online ISBN: 978-1-4614-5797-8
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